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Equine Reproduction
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Companion CD The book includes a companion CD that contains 713 color illustrations.
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Equine Reproduction Second edition Volume 1 Edited by
Angus O. McKinnon BVSc, MSc, Diplomate ACT and ABVP Goulburn Valley Equine Hospital Congupna Victoria 3632 Australia
Edward L. Squires MS, PhD, Diplomate ACT (hon) Executive Director of the Gluck Equine Research Foundation Research Professor, Department of Veterinary Science Gluck Equine Research Center University of Kentucky Lexington, KY 40546-0099 USA
Wendy E. Vaala VMD, Diplomate ACVIM Sr. Equine Technical Service Veterinarian Intervet Schering-Plough Animal Health Alma, WI 54610 USA
Dickson D. Varner DVM, MS, Diplomate ACT Professor of Theriogenology and Pin Oak Stud Chair of Stallion Reproductive Studies Department of Large Animal Clinical Sciences, College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX 77843-4475 USA
A John Wiley & Sons, Ltd., Publication
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This edition first published 2011
C 1993 Blackwell Publishing Ltd C 2011 Blackwell Publishing Ltd C 2011 C.J. (Kate) Savage Chapter 57 C 2011 Robert P. Franklin Chapter 92 C 2011 Dee L. Cross Chapter 249
BlackwellPublishing was acquired by John Wiley & Sons in February 2007. Blackwell’s publishing program has been merged with Wiley’s global Scientific, Technical, and Medical business to form Wiley-Blackwell. First edition published 1993 Second edition published 2011 Registered office John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, United Kingdom Editorial offices 2121 State Avenue, Ames, Iowa 50014-8300, USA The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, United Kingdom 9600 Garsington Road, Oxford, OX4 2DQ, United Kingdom For details of our global editorial offices, for customer services, and for information about how to apply for permission to reuse the copyright material in this book, please see our Website at www.wiley.com/wiley-blackwell. Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Blackwell Publishing, provided that the base fee is paid directly to the Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license by CCC, a separate system of payments has been arranged. The fee code for users of the Transactional Reporting Service is ISBN-13: 978-0-8138-1971-6/2011. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Disclaimer The contents of this work are intended to further general scientific research, understanding, and discussion only and are not intended and should not be relied upon as recommending or promoting a specific method, diagnosis, or treatment by practitioners for any particular patient. The publisher and the author make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of fitness for a particular purpose. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of medicines, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each medicine, equipment, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. Readers should consult with a specialist where appropriate. The fact that an organization or Website is referred to in this work as a citation and/or a potential source of further information does not mean that the author or the publisher endorses the information the organization or Website may provide or recommendations it may make. Further, readers should be aware that Internet Websites listed in this work may have changed or disappeared between when this work was written and when it is read. No warranty may be created or extended by any promotional statements for this work. Neither the publisher nor the author shall be liable for any damages arising herefrom Library of Congress Cataloging-in-Publication Data Equine reproduction / edited by Angus O. McKinnon . . . [et al.]. – 2nd ed. p. ; cm. Includes bibliographical references and index. ISBN 978-0-8138-1971-6 (hardcover : alk. paper) 1. Horses–Reproduction. 2. Horses–Breeding. I. McKinnon, A. O. [DNLM: 1. Horses–physiology. 2. Reproduction. SF 768.2.H67] SF768.2.H67E68 2011 636.1089’82–dc22 2010038949 A catalog record for this book is available from the U.S. Library of Congress. This book is published in the following electronic format: ePDF 9781444397635 R , Inc., New Delhi, India Set in 7.8/9.8pt Palatino by Aptara
1 2011
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Contents Contributors, xvii Preface to the second edition, xxxiv Preface to the first edition, xxxv Acknowledgments, xxxvi Color plates appear between pages 812 and 813
Volume 1 Part I: The Foal
10
Section (a) Prepartum assessment of the foal 1
Identification of the high-risk pregnancy, 5 Elizabeth M. Santschi and Wendy E. Vaala
2
Monitoring the high-risk pregnancy, 16 Wendy E. Vaala
3
Management of the high-risk pregnancy, 25 Elizabeth M. Santschi and Wendy E. Vaala
4 5
Peri-parturient management of the mare and neonate, 111 T. Douglas Byars and Barry W. Simon
Section (d) Problems of the immediate postpartum period 11
Foal rejection, 117 Wendy E. Vaala
12
Prematurity, dysmaturity and assessment of maturity, 121 Guy D. Lester
Ultrasonographic monitoring of the fetus, 39 Stefania Bucca
13
Sedation and anesthesia of the pregnant mare, 55 John A.E. Hubbell
Resuscitation (foal and birth), 128 Niamh M. Collins, Jane E. Axon and Jonathan E. Palmer
14
Shock, 136 Niamh M. Collins, Jane E. Axon and Jonathan E. Palmer
Section (b) Foal adaptation to parturition 6
The normal post partum foal, 63 Sarah J. Stoneham
15
Perinatal asphyxia syndrome, 147 Pamela A. Wilkins
7
Endocrinological adaptation, 69 Jennifer C. Ousey
16
8
The placenta, 84 Peter R. Morresey
Sepsis, 154 L. Chris Sanchez
Section (e) Specific diagnostic and management techniques
Section (c) Postpartum fetal assessment 9
Examination of the placenta, 99 Donald H. Schlafer
17
v
Critical care – assessment, 167 Jane E. Axon
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Critical care – monitoring, 177 Darien J. Feary
35
Neonatal isoerythrolysis, 353 Jill R. Johnson
19
Critical care – treatment, 189 Jane E. Axon
36
Immunodeficiencies in foals, 361 M. Julia B. Felippe Flaminio
20
Complications of neonatal infection, 198 Mary Rose Paradis
37
Severe combined immunodeficiency, 369 Melissa T. Hines
38
21
Fluid therapy, 205 Robert P. Franklin
Thrombocytopenia and selective IgM deficiency, 373 Rodney L. Belgrave
22
Respiratory assistance, 215 Jonathan E. Palmer
Section (g) Gastrointestinal system
23
Sedation, anesthesia and analgesia, 221 Lori A. Bidwell
39
Meconium impaction, 379 Wendy E. Vaala
24
Stabilization and preparation for transport, 230 Robert P. Franklin and Jennifer R. Read
40
Neonatal diarrhea, 385 K. Gary Magdesian
41
25
Transfusion techniques, 238 John F. Freestone
Gastroduodenal ulceration: The neonatal perspective, 394 L. Chris Sanchez
26
Diagnostic ultrasonography of the abdomen, 243 Virginia B. Reef and JoAnn Slack
42
Surgical gastrointestinal conditions, 399 Brett J. Woodie
27
Antibiotic therapy, 253 K. Gary Magdesian
43
28
Enteral and parenteral nutrition for the neonatal foal, 263 Wendy E. Vaala
Other gastrointestinal disorders (chyloperitoneum, atresia, lethal white), 405 K. Gary Magdesian
44
Hepatic disease, 409 Thomas J. Divers and T. Douglas Byars
Growth of horses, 280 Clarissa Brown-Douglas, Peter Huntington and Joe Pagan
Section (h) Hemopoetic system 45
Important gastrointestinal parasites, 292 Eugene T. Lyons, Mariana Ionita and Sharon C. Tolliver
Anemia, 419 Jane E. Axon
46
Coagulation disorders, 426 Fairfield T. Bain
29
30
31
Vaccination of mares, foals and weanlings, 302 W. David Wilson
Section (f) Immune system
Section (i) Musculoskeletal system 47
Fetal ossification and normal joint development, 433 Elwyn C. Firth
48
Development of the foal immune system, 331 M. Julia B. Felippe Flaminio and Rebecca L. Tallmadge
Flexural deformities, 441 Stephen B. Adams and Timothy B. Lescun
49
Fractures commonly seen in foals, 446 Angus R. Adkins
33
Colostrum: Assessment of quality and artificial supplementation, 342 Guy D. Lester
50
Infectious arthritis and osteomyelitis, 457 Fairfield T. Bain
34
Assessment and modification of passive transfer, 346 Sally L. Vivrette
51
Muscle disorders, 463 Erica C. McKenzie and Jennifer M. MacLeay
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66
Bacterial and viral respiratory disease of the neonate, 601 Renaud L´eguillette
67
Neurological evaluation, 477 Robert J. MacKay
Congenital abnormalities of the respiratory system, 609 Eric J. Parente
68
Neurological disorders in the neonatal foal, 483 Robert J. MacKay
Genetic basis of respiratory disorders, 615 Eliane Marti, Caroline Tessier and Vincent Gerber
Section (p) Urogenital system
Tendon and ligament disorders, 469 Timothy B. Lescun and Stephen B. Adams
Section (j) Nervous system 53 54
69
Techniques in diagnosis and monitoring of cardiovascular disease, 499 Anne Desrochers
Disorders of the bladder, ureters and urethra, 625 Thomas J. Divers
70
Congenital abnormalities of the cardiovascular system, 507 Celia M. Marr
Disorders of the umbilicus and urachus, 632 Kirsten M. Neil
71
Renal problems, 646 Jonathan E. Palmer
Acquired abnormalities of the cardiovascular system, 511 C.J. (Kate) Savage
Section (q) System disorders
Section (k) Cardiovascular system 55
56
57
72
Congenital problems of foals, 663 Antonio M. Cruz
73
Inherited problems of foals, 673 Sharon J. Spier, Carrie J. Finno and Stephanie J. Valberg
Section (l) Endocrinology system 58
Endocrine abnormalities, 523 Melissa T. Hines
Section (m) Ocular system 59
Ophthalmic disorders of the foal, 535 Caryn E. Plummer and Dennis E. Brooks
Section (r) Problems of older foals, weanlings and yearlings 74
Gastroduodenal ulcer complex of the older foal, 683 Bonnie Barr
75
Rhodococcus pneumonia: Epidemiology and prophylaxis, 688 Noah D. Cohen
76
Rhodococcus pneumonia: Pathogenesis, diagnosis and therapy, 695 Steeve Gigu`ere
77
Strangles, 704 John F. Timoney
78
Other bacterial respiratory diseases of the older foal, 710 Steeve Gigu`ere
79
Viral respiratory diseases of the older foal, 720 Lutz S. Goehring and D. Paul Lunn
80
Neurological conditions (EPM, EMND wobblers, trauma, etc), 728 Guy D. Lester
Section (n) Integument system 60
Diseases of the skin, 555 Derek C. Knottenbelt
Section (o) Respiratory System 61
Diagnostic approach to respiratory disorders, 575 Guy D. Lester
62
Acute respiratory distress syndrome (ARDS), 582 Pamela A. Wilkins
63
vii
Respiratory diseases specific to the neonate, 586 Guy D. Lester
64
Aspiration pneumonia, 592 Mary Rose Paradis
65
Other disorders of breathing, 597 Pamela A. Wilkins
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81
Diarrhea, 734 Kirsten M. Neil
98
Oxidative stress in sperm, 991 Barry A. Ball
82
Colic, 749 Joanne Hardy
99
83
Weight loss, 758 Kirsten M. Neil
Endocrine–paracrine–autocrine regulation of reproductive function in the stallion, 996 Janet F. Roser
84
Angular limb deformities, 768 Earl M. Gaughan and Sarah E.T. Hanna
85
Developmental orthopedic disease, 772 C. Wayne McIlwraith
86
Feeding the growing horse to avoid developmental orthopedic disease (DOD), 782 Paul D. Siciliano
87 88 89
Presale endoscopy, 794 Rolf M. Embertson Presale radiographs, 801 James P. Morehead and Brent N. Cassady Presale radiographic interpretation and significance, 809 C. Wayne McIlwraith
Section (s) Appendix
100 Puberty, 1015 Noah L. Heninger 101 Spermatogenesis, 1026 Larry Johnson, Cleet E. Griffin and Michael T. Martin 102 Spermatozoal function, 1053 Rupert P. Amann and James K. Graham 103 Sperm-oviduct interactions, 1085 Barry A. Ball 104 Sperm–uterine interactions, 1092 Terttu Katila 105 Testicular descent, 1099 Mimi Arighi
Section (b) Abnormalities of the stallion’s reproductive tract
90
Development of a NICU, 821 Nathan M. Slovis and Lynne M. Hewlett
106 Developmental abnormalities of the male reproductive tract, 1109 Mimi Arighi
91
Diagnostic laboratory resources and critical care supplies, 830 Nathan M. Slovis and Wendy E. Vaala
107 Abnormalities of the accessory sex glands, 1113 Dickson D. Varner and James Schumacher
92
Formulary, 838 Robert P. Franklin
108 Abnormalities of the ejaculate, 1119 Regina M.O. Turner
93
Hematology and biochemical reference values, 851 Robert P. Franklin and Joan Palmero
109 Abnormalities of the penis and prepuce, 1130 James Schumacher and Dickson D. Varner
94
SI Conversion Table, 857
110
Abnormalities of the spermatic cord, 1145 James Schumacher and Dickson D. Varner
111
Epididymal abnormalities, 1156 Aime K. Johnson and Charles C. Love
112
Abnormalities of the testicles, 1161 Warren Beard
113
Neoplasia of the reproductive tract, 1166 John F. Edwards
Part II: The Stallion Section (a) Anatomy, physiology and endocrinology 95
Functional anatomy of the adult male, 867 Rupert P. Amann
96
Physiology and endocrinology, 881 Rupert P. Amann
97
From a sperm’s eye view: Revisiting our perception of this intriguing cell, 909 Dickson D. Varner and Larry Johnson
Section (c) Management 114
Drugs that adversely affect spermatogenesis, 1175 Rupert P. Amann
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Contents 115
Dual hemisphere breeding programs, 1184 Norman W. Umphenour
116
Immunocastration, 1191 Tom A.E. Stout and Ben Colenbrander
117
118
119
Management of stallions in artificial insemination, 1198 H. Steve Conboy Management of stallions in natural-service programs, 1208 Norman W. Umphenour, Phillip McCarthy and Terry L. Blanchard Nutrition and exercise for breeding stallions, 1228 Stephen G. Jackson
120 Management of subfertile stallions under natural-mating conditions, 1240 Terry L. Blanchard
132 Semen extenders for cooled semen (North America), 1341 Steven P. Brinsko 133 Factors affecting sperm production and output, 1344 Edward L. Squires and Bill W. Pickett 134 Reproductive parameters of draft horse, friesian and warmblood stallions, 1362 Tom A.E. Stout and Ben Colenbrander 135 Reproductive parameters from light horse stallions, 1367 Edward L. Squires 136 Reproductive parameters from miniature stallions, 1377 Dale L. Paccamonti and Paul J. De Vries Section (e) Sexual behavior
121 Venereal disease, 1250 Elizabeth S. Metcalf
137 Normal sexual behavior, 1385 Sue M. McDonnell
Section (d) Semen collection, evaluation and artificial insemination
138 Handling the breeding stallion, 1391 Dickson D. Varner
122 Historical perspectives of artificial insemination, 1261 John M. Bowen
139 The novice breeding stallion, 1396 H. Steve Conboy
123 Semen collection techniques and insemination procedures, 1268 Steven P. Brinsko 124 Evaluation of semen, 1278 Julie Baumber-Skaife 125 Spermatozoal motility, 1292 Aaron D.J. Hodder and Irwin K.M. Liu 126 Spermatozoal morphology, 1297 D.N. Rao Veeramachaneni 127 Principles of cooled semen, 1308 James K. Graham 128 Breeding with cooled transported semen, 1316 Juan C. Samper 129 Experiences with a large-scale cooled – transported semen program, 1323 Simon J. Robinson
140 Training a stallion to the phantom, 1402 Glenn P. Blodgett 141 Abnormal sexual behavior, 1407 Sue M. McDonnell 142 Pharmacological manipulation of ejaculation, 1413 Sue M. McDonnell 143 Pharmacological manipulation of stallion behavior, 1415 Sue M. McDonnell 144 Specific erection and ejaculatory dysfunctions, 1419 Sue M. McDonnell 145 Stereotypic behavior of stallions, 1423 Katherine A. Houpt Section (f) Specific techniques used in reproductive examination of the stallion
130 Containers for transport of equine semen, 1330 Terttu Katila
146 Historical information, 1429 Charles C. Love
131 Semen extenders for cooled semen (Europe), 1336 Christine Aurich
147 Diagnostics and therapeutics for stallions with declining fertility: An endocrine– paracrine–autocrine approach, 1435 Janet F. Roser
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148 Endoscopy of the internal reproductive tract, 1448 Carla L. Carleton
Section (h) Surgical procedures of the male reproductive tract 156 Clinical uses of testicular biopsy, 1517 Terry L. Blanchard
149 Examination of external genitalia, 1458 Patricia L. Sertich
157 Testicular biopsy, 1523 Walter R. Threlfall
150 Cytogenetic evaluation, 1462 Keith W. Durkin, Terje Raudsepp and Bhanu P. Chowdhary
158 Cryptorchid castration, 1531 Thomas M. Russell and Patrick J. Pollock
151 Ultrasonography of the genital tract, 1469 Regina M.O. Turner
159 Inguinal hernia, 1540 Patrick J. Pollock and Thomas M. Russell
Section (g) Specific techniques used in semen evaluation of the stallion
160 Laparoscopic surgery, 1546 David G. Wilson
152 Acrosomal function, 1491 Stuart A. Meyers 153 Evaluation of the plasma membrane, 1498 Ben Colenbrander and Tom A.E. Stout 154 Sperm chromatin structure assay, 1506 Aime K. Johnson and Charles C. Love 155 Spermatozoal viability, 1510 Stuart A. Meyers
161 Laser assisted castration, 1553 Scott E. Palmer 162 Normal field castration, 1557 Glenn P. Blodgett 163 Surgery of the penis and prepuce, 1562 James Schumacher and Dickson D. Varner Index, 1573
Volume 2 Part III: The Mare Section (a) Anatomy 164 External reproductive anatomy, 1577 John J. Dascanio 165 Internal reproductive anatomy, 1582 Robert A. Kainer
Section (b) Physiology and endocrinology 166 How hormones work, 1601 Peter R. Morresey 167 GnRH, 1608 Susan L. Alexander and Clifford H.G. Irvine 168 FSH and LH, 1619 Susan L. Alexander and Clifford H.G. Irvine 169 Estrogens, 1631 Bruce W. Christensen 170 Progesterone, 1637 Dirk K. Vanderwall
171 Prostaglandins, 1642 Tom A.E. Stout 172 Equine chorionic gonadotropin (eCG), 1648 Amanda M. de Mestre, Douglas F. Antczak and W.R. (Twink) Allen 173 Adrenal steroids, 1665 Valerie J. Linse 174 Melatonin, 1669 Daniel C. Sharp 175 Oxytocin, inhibin, activin, relaxin and prolactin, 1679 Peter R. Morresey Section (c) The estrous cycle 176 Puberty, 1689 Bruce E. Eilts 177 Anestrus, 1696 Donald L. Thompson, Jr. 178 Vernal transition into the breeding season, 1704 Daniel C. Sharp
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Contents 179 Estrus, 1716 Patrick M. McCue, Charles F. Scoggin and Alicia R.G. Lindholm
198 New techniques in hormonal delivery, 1879 Patrick J. Burns
180 Diestrus, 1728 Robert M. L¨ofstedt
199 Contraception, 1888 Jay F. Kirkpatrick and John W. Turner
181 Autumnal transition out of the breeding season, 1732 Sheryl S. King 182 The abnormal estrous cycle, 1754 Patrick M. McCue and Ryan A. Ferris Section (d) The transitional mare 183 Photoperiod, 1771 Daniel C. Sharp 184 Progesterone, 1778 Edward L. Squires 185 GnRH, 1782 John Hyland 186 Dopamine antagonists, 1788 Ahmed Tibary Section (e) Pharmacological manipulation of reproduction 187 Prostaglandins, 1797 Simon A. Staempfli 188 Human chorionic gonadotropin, 1804 John R. Newcombe 189 Progestagens and progesterone, 1811 Carlos R.F. Pinto 190 Gonadotropin-releasing hormones, 1820 Edward L. Squires
Section (f) Techniques in reproductive examination 200 History, 1897 Walter W. Zent 201 Vaginal examination, 1900 Walter W. Zent and John V. Steiner 202 Direct rectal palpation, 1904 Thomas R. Bowman, Jr. 203 Ultrasonography, 1914 Jonathan F. Pycock 204 Uterine cytology, 1922 Michelle M. LeBlanc 205 Endometrial biopsy, 1929 Charles C. Love 206 Endoscopic examination, 1940 Claire E. Card 207 Cytogenetic evaluation of mares and foals, 1951 Teri L. Lear and Daniel A.F. Villagomez 208 Uterine and clitoral cultures, 1963 Sidney W. Ricketts 209 Use of chromogenic agar to diagnose reproductive pathogens, 1979 Angus O. McKinnon and David P. Beehan
191 Estrogen therapy, 1825 Ahmed Tibary
210 Evaluation of uterine tubal patency, 1988 William B. Ley
192 Oxytocin, 1830 Michelle M. LeBlanc
211
Laparoscopy, 1991 Dean A. Hendrickson and David G. Wilson
193 Superovulation, 1836 Edward L. Squires and Patrick M. McCue 194 Estrus suppression, 1845 Dirk K. Vanderwall and Gary J. Nie 195 Inhibin immunotherapy, 1854 Patrick M. McCue 196 Induction of ovulation, 1858 Angus O. McKinnon and Patrick M. McCue 197 Synchronization of ovulation, 1870 Etta A. Bradecamp
Section (g) Diagnostic ultrasonography 212 Normal anatomy, 2003 Elaine M. Carnevale and Lawrence M. Olsen 213 Folliculogenesis, 2009 Mohd A. Beg and Don R. Bergfelt 214 Ovulation: Part 1. Follicle development and endocrinology during the periovulatory period, 2020 Eduardo L. Gastal
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215 Ovulation: Part 2. Ultrasonographic morphology of the preovulatory follicle, 2032 Eduardo L. Gastal 216 Luteal development, 2055 Don R. Bergfelt and Gregg P. Adams 217 Pregnancy, 2065 Don R. Bergfelt and Gregg P. Adams 218 Equine fetal sex determination between 55 and 150 days, 2080 Richard D. Holder 219 Fetal gender determination from mid to advanced gestation, 2094 Stefania Bucca 220 Management of twins, 2099 Angus O. McKinnon 221 Early embryonic loss, 2118 Dirk K. Vanderwall 222 Ovarian abnormalities, 2123 Patrick M. McCue and Angus O. McKinnon 223 Uterine abnormalities, 2137 Angus O. McKinnon and Patrick M. McCue 224 Mammary gland, 2162 Bryan M. Waldridge Section (h) Pregnancy, parturition and the puerperal period 225 Embryo morphology, growth and development, 2167 Keith J. Betteridge 226 Fetal membrane differentiation, implantation and early placentation, 2187 W.R. (Twink) Allen, Susan Gower and Sandra Wilsher 227 Maternal recognition of pregnancy, 2200 Karen J. McDowell and Daniel C. Sharp
233 Parturition, 2268 Bruce W. Christensen 234 Lactation, 2277 Patrick M. McCue and Sofie Sitters 235 Uterine involution, 2291 Mary E. Stanton 236 Breeding mares on foal heat, 2294 Terry L. Blanchard and Margo L. Macpherson 237 Inter and extraspecies equine pregnancies, 2302 W.R. (Twink) Allen, Julia H. Kydd, Roger V. Short and Douglas F. Antczak 238 Induction of abortion, 2320 Claire E. Card Section (i) Problems of pregnancy 239 Embryonic loss, 2327 Barry A. Ball 240 Abortions and stillbirths: A pathologists overview, 2339 Katherine E. Whitwell 241 Origin and outcome of twin pregnancies, 2350 Angus O. McKinnon 242 Placentitis, 2359 Mats H.T. Troedsson and Margo L. Macpherson 243 Hydrops, 2368 Rudolf O. Waelchli 244 Fetal maceration and mummification, 2373 Claire E. Card 245 Equine herpesvirus, 2376 Peter J. Timoney 246 Equine viral arteritis, 2391 Peter J. Timoney 247 Contagious equine metritis, 2399 Peter J. Timoney
228 Sex determination and differentiation, 2211 Bruce W. Christensen and Vicki N. Meyers-Wallen
248 Mare reproductive loss syndrome, 2410 David G. Powell
229 Endocrinology of pregnancy, 2222 Jennifer C. Ousey
249 Fescue toxicosis, 2418 Dee L. Cross
230 Development and morphology of the placenta, 2234 Sandra Wilsher and W.R. (Twink) Allen
250 Prepubic and abdominal wall rupture, 2428 Dwayne H. Rodgerson
231 Pregnancy examination, 2245 Patrick M. McCue and Angus O. McKinnon
251 Uterine prolapse, 2431 Michael A. Spirito and Kim A. Sprayberry
232 Induction of parturition, 2262 Margo L. Macpherson and Dale L. Paccamonti
252 Uterine torsion, 2435 James R. Vasey and Tom Russell
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Contents 253 Abnormalities of pregnancy, 2441 Robert M. L¨ofstedt 254 Maintenance of pregnancy, 2455 Angus O. McKinnon and Jonathan F. Pycock 255 Dystocia management, 2479 Grant Frazer 256 Fetotomy, 2497 Grant Frazer 257 Cesarian section, 2505 David E. Freeman 258 Referral dystocias, 2511 Rolf M. Embertson 259 Periparturient hemorrhage, 2517 T. Douglas Byars and Thomas J. Divers
273 Fungal endometritis, 2643 Marco A. Coutinho da Silva and Marco A. Alvarenga 274 Pyometra, 2652 Kristina G. Lu 275 Treatment of fluid accumulation, 2655 Jonathan F. Pycock 276 Uterine cysts, 2665 Mary E. Stanton 277 Uterine abnormalities, 2669 John P. Hurtgen 278 Laboratory methods for isolation and evaluation of bacteria, fungi and yeast, 2674 David L. Hartman and Shalyn Bliss
260 Retained fetal membranes, 2520 Walter R. Threlfall
Section (l) Diseases of the ovary and oviduct
261 Postpartum metritis, 2530 Terry L. Blanchard
279 Disorders of ovulation, 2685 Patrick M. McCue
Section (j) Female urogenital surgery 262 Pneumovagina, 2537 Etta A. Bradecamp 263 Surgery of the caudal reproductive tract, 2545 Angus O. McKinnon and Sarah L. Jalim 264 Cervical surgery, 2559 Patrick J. Pollock and Thomas M. Russell
280 Disorders of the oviduct, 2692 Irwin K.M. Liu 281 Non-neoplastic abnormalities, 2697 Donald H. Schlafer 282 Ovarian neoplasia, 2707 Claire E. Card Section (m) Problems of the cervix
265 Ovariectomy, 2564 Dwayne H. Rodgerson and Dawn A. Loesch
283 Abnormalities of cervical and vaginal development, 2719 John P. Hurtgen
266 Hysterectomy, 2574 Anna K. Roetting and David E. Freeman
284 Cervix adhesions, 2721 Patricia L. Sertich
267 Rectal tears, 2578 David E. Freeman
285 Failure to dilate, 2724 Ahmed Tibary Section (n) The mammary gland
Section (k) Disease of the uterus 268 Immunological considerations, 2587 Sara K. Lyle 269
Abnormalities of contractility, 2597 Terttu Katila
270 Abnormalities of development, 2607 John P. Hurtgen 271 Endometritis, 2608 Mats H.T. Troedsson 272 Breeding the problem mare, 2620 Michelle M. LeBlanc and Angus O. McKinnon
286 Pre-foaling mammary secretions, 2733 William B. Ley 287 Mastitis, 2738 Erica K. Gee and Patrick M. McCue 288 Mammary neoplasia, 2742 Jack R. Snyder and Omar Maher Section (o) Specific diagnostic and management techniques 289 Preventative medicine for broodmare farms, 2747 John B. Chopin
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Section (p) Nutrition 290 Dietary support for geriatric broodmares and stallions, 2755 Sarah L. Ralston 291 Nutrition for the broodmare, 2760 Laurie M. Lawrence Section (q) Behavior 292 Stereotypic behavior, 2771 Paul McGreevy Section (r) General interest 293 Reproductive efficiency, 2779 Laura C. Nath 294 Pituitary pars intermedia dysfunction (Equine Cushing’s syndrome), 2790 Philip J. Johnson, Nat T. Messer and Venkataseshu K. Ganjam 295 Epidemiology applied to horse reproduction, 2796 Nigel R. Perkins 296 Management of the geriatric mare, 2803 Scott Madill 297 Parentage testing, 2820 Ernest Bailey 298 Relevance of genomics to equine reproduction, 2827 Ernest Bailey
304 Cooled transported embryo technology, 2880 Patrick M. McCue, Catherine A. DeLuca, and Jillian J. Wall 305 Freezing of embryos, 2887 Jean-Fran¸cois Bruyas 306 Import and export of embryos, 2921 Erin C. Bishop 307 Nuclear Transfer, 2924 Katrin Hinrichs Section (b) Oocytes 308 Immature oocyte collection and maturation, 2931 Katrin Hinrichs 309 Mature oocyte collection, 2936 Elaine M. Carnevale 310 Oocyte transfer, 2941 Elaine M. Carnevale 311
Gamete intrafallopian transfer (GIFT), 2945 Marco A. Coutinho da Silva
312 Intracytoplasmic sperm injection (ICSI), 2948 Young-Ho Choi and Katrin Hinrichs 313 Oocyte cryopreservation, 2953 Lisa J. Maclellan Section (c) Semen
Section (s) Other equids
314 Principles of cryopreservation, 2959 James K. Graham
299 Donkey reproduction, 2835 Robyn R. Wilborn and David G. Pugh
315 Semen extenders for frozen semen, 2964 Harald Sieme
300 Miniature pony reproduction, 2839 Paul J. De Vries and Dale L. Paccamonti
316 Freezing semen, 2972 Harald Sieme
301 Asian wild horse reproduction: Equus ferus przewalskii, 2846 C. Wynne Collins, Lee E. Boyd and Katherine A. Houpt
317 Freezing epididymal spermatozoa, 2983 Jason E. Breummer
302 Zebra reproduction, 2851 Cassandra M.V. Nunez, ˜ Cheryl S. Asa and Daniel I. Rubenstein
318 Breeding with frozen semen, 2987 Sandro Barbacini 319 Techniques for evaluating frozen semen, 2994 Ben Colenbrander and Tom A.E. Stout
Part IV: Assisted Reproductive Technologies (ART)
320 Storage management and distribution of frozen semen, 3005 Paul R. Loomis
Section (a) Embryos
321
303 Embryo transfer, 2871 David L. Hartman
Diseases potentially transmitted with frozen or cooled semen, 3015 Peter J. Timoney
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Contents 322 Import and export of frozen semen, 3029 Elizabeth S. Metcalf 323 Low dose insemination, 3036 Lee H.A. Morris and Sara K. Lyle
Section (d) Future technologies 325 Future reproductive technology, 3051 George E. Seidel Jr Index, 3057
324 Sex-sorted spermatozoa, 3042 Lee H.A. Morris
Companion CD The book includes a companion CD that contains 713 color illustrations.
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Contributors Gregg P. Adams DVM, PhD
Rupert P. Amann PhD
Diplomate ACT Professor Department of Veterinary Biomedical Sciences Western College of Veterinary Medicine University of Saskatchewan Saskatoon, SK Canada
Professor Emeritus Department of Biomedical Sciences College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO USA
Douglas F. Antczak
Stephen B. Adams DVM, MS
James A. Baker Institute for Animal Health College of Veterinary Medicine Cornell University Ithaca, NY USA
Diplomate ACVS Professor Veterinary Clinical Sciences Purdue University School of Veterinary Medicine West Lafayette, IN USA
Mimi Arighi DVM, MSc Diplomate ACVS Director, Associate Professor Veterinary Teaching Hospital School of Veterinary Medicine Purdue University West Lafayette, IN USA
Angus R. Adkins BVSc, FACVSc Director, Senior Surgeon Scone Equine Hospital Department of Surgery Scone, NSW Australia
Cheryl S. Asa PhD
Susan L. Alexander PhD
Director of Research Research Department, Saint Louis Zoo; Adjunct Professor, Biology Department Saint Louis University Saint Louis, MO USA
(Retired) Equine Research Unit Lincoln University Canterbury New Zealand
Christine Aurich DVM, PhD
W.R. (Twink) Allen CBE, ScD, FRCVS
Diplomate ECAR Professor in Reproduction Graf Lehndorff Institute for Equine Science University for Veterinary Sciences Vienna Austria
Director The Paul Mellon Laboratory of Equine Reproduction Newmarket Suffolk United Kingdom
Jane E. Axon BVSc (Hons), MACVSc., Marco A. Alvarenga
Diplomate ACVIM, CMAVA Director Clovelly Intensive Care Unit Scone Equine Hospital Scone, NSW Australia
Associate Professor Department of Animal Reproduction Sao Paulo State University Botucatu, SP Brazil
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Contributors
Ernest Bailey PhD
Mohd A. Beg BVSc, MVSc, PhD
Professor Department of Veterinary Science University of Kentucky Gluck Equine Research Center Lexington, KY USA
Research Scientist Department of Pathobiological Sciences School of Veterinary Medicine University of Wisconsin-Madison Madison, WI USA
Fairfield T. Bain DVM, MBA
Rodney L. Belgrave DVM, MS
Diplomate ACVIM, ACVP, ACVECC Internist/Pathologist Equine Sports Medicine and Surgery Weatherford, TX USA
Barry A. Ball DVM, PhD Diplomate ACT Professor and Albert G. Clay Endowed Chair in Equine Reproduction Gluck Equine Research Center Lexington, KY USA
Diplomate ACVIM Staff Internist Mid-Atlantic Equine Medical Center Ringoes, NJ USA
Don R. Bergfelt MS, PhD Honorary Fellow Office of Science Coordination and Policy US Environmental Protection Agency Washington, DC USA
Keith J. Betteridge BVSc, MVSc, PhD, FRCVS
Select Breeders Services Italia San Daniele Po-Cremona Italy
University Professor Emeritus Department of Biomedical Sciences Ontario Veterinary College University of Guelph Guelph, ON Canada
Bonnie Barr VMD
Lori A. Bidwell DVM
Diplomate ACVIM Internist Medicine Department Rood and Riddle Equine Hospital Lexington, KY USA
Diplomate ACVA Assistant Professor Department of Anesthesia Ross University School of Veterinary Medicine Basseterre Saint Kitts West Indies
Sandro Barbacini DMV
Julie Baumber-Skaife PhD Director of Quality Assurance Select Breeders Service Inc. Colora, MD USA
Erin C. Bishop MS
Warren Beard DVM, MS
Terry L. Blanchard DVM, MS
Diplomate ACVS Professor Department of Clinical Sciences Kansas State University Manhattan, KS USA
David P. Beehan MVB Theriogenology Resident Department of Theriogenology Veterinary Teaching Hospital and Clinics Louisiana State University Baton Rouge, LA USA
General Manager Select Breeders Southwest Inc., TX USA
Diplomate ACT Professor Department of Large Animal Clinical Sciences College of Veterinary Medicine Texas A&M University, TX USA
Shalyn Bliss DVM Associate Veterinarian Hartman Equine Reproduction Center; Resident, Theriogenology Large Animal Clinical Sciences Texas A&M University, TX USA
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Contributors Glenn P. Blodgett DVM
Jean-Franc¸ois Bruyas DVM, MSc, PhD
Horse Division Manager & Veterinarian Four Sixes Ranch Guthrie, TX USA
Diplomate ECAR Professor Biotechnology and Pathology Reproduction Unit Department of Clinical Sciences College of Veterinary Medicine Food Science and Engineering Nantes-Atlantique ONIRIS Nantes France
John (Jim) M. Bowen FRCVS Professor Emeritus Department of Large Animal Clinical Sciences Virginia Maryland Regional College of Veterinary Medicine Virginia Polytechnic Institute and State University Blacksburg, VA USA
Thomas R. Bowman, Jr. DVM 10395 Rileys Mill Road Chestertown, MD USA
Lee E. Boyd PhD Professor Department of Biology Washburn University Topeka, KS USA
Etta A. Bradecamp DVM Diplomate ACT, ABVP Warrenton, VA USA
Steven P. Brinsko DVM, MS, PhD Diplomate ACT Associate Professor and Chief of Theriogenology Department of Large Animal Clinical Sciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University Texas USA
Dennis E. Brooks DVM, PhD Diplomate ACVO Professor of Ophthalmology University of Florida Florida USA
Clarissa Brown-Douglas PhD Research Fellow Kentucky Equine Research Mulgrave, Victoria Australia
Jason E. Bruemmer PhD Associate Professor Department of Animal Sciences Colorado State University Fort Collins, CO USA
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Stefania Bucca DVM Somerton Equine Hospital/SBS Ireland Kildare Co Kildare Ireland
Patrick J. Burns PhD Vice President, CSO BioRelease Technologies/Bet Pharm Lexington, KY USA
T. Douglas Byars DVM Diplomate ACVIM, ACVECC Owner Byars Equine Advisory, LLC Georgetown, KY USA
Claire E. Card DVM, PhD Diplomate ACT Professor Department Large Animal Clinical Sciences Western College of Veterinary Medicine University of Saskatchewan Saskatoon, SK Canada
Carla L. Carleton DVM, MS Diplomate ACT Equine Theriogenologist Department of Large Animal Clinical Sciences Michigan State University College of Veterinary Medicine East Lansing, MI USA
Elaine M. Carnevale DVM, PhD Associate Professor Animal Reproduction and Biotechnology Laboratory Colorado State University Fort Collins, CO USA
Brent N. Cassady DVM Equine Medical Associates, PSC Lexington, KY USA
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Contributors
Young-Ho Choi DVM, PhD
H. Steve Conboy DVM
Research Scientist Department of Veterinary Physiology & Pharmacology College of Veterinary Medicine and Biomedical Sciences Texas A&M University, TX USA
Maple Lane Farm Lexington, KY USA
John B. Chopin BVSc, PhD FACVSc Resident Veterinarian Coolmore Australia, NSW Australia
Marco A. Coutinho da Silva DVM, PhD Diplomate ACT Assistant Professor Theriogenology and Reproductive Medicine Department of Veterinary Clinical Sciences The Ohio State University Columbus, OH USA
Bhanu P. Chowdhary BVSc&AH, MVSc, VMD, PhD
Dee L. Cross MS, PhD
Associate Dean, Research and Graduate Studies; Professor and Faculty Fellow Department of Veterinary Integrative Biosciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University, TX USA
Professor Emeritus Animal and Veterinary Sciences Clemson University Clemson; President/CEO Equi-Tox Inc., SC USA
Bruce W. Christensen DVM, MS
Antonio M. Cruz DVM, MVM, MSc, DMV
Diplomate ACT Assistant Professor Iowa State University Veterinary Clinical Sciences, IA USA
Diplomate ACVS and ECVS Equine Surgeon and Hospital Director Paton and Martin Veterinary Services British Columbia Canada
Noah D. Cohen VMD, MPH, PhD
John J. Dascanio VMD
Diplomate ACVIM Professor Large Animal Clinical Sciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University, TX USA
Diplomate ACT and ABVT Professor, Theriogenology Ross University School of Veterinary Medicine Basseterre St. Kitts West Indies
Catherine A. DeLuca DVM, MS Professor Emeritus of Male Fertility Department of Equine Sciences Utrecht University Utrecht The Netherlands
Diplomate ACT Resident, Equine Reproduction Department of Clinical Sciences Colorado State University Fort Collins, CO USA
C. Wynne Collins MVB
Amanda M. de Mestre BVSc, PhD, MRCVS
Diplomate ACT Post-doctoral Fellow Department of Reproductive Sciences Center for Species Survival Royal Smithsonian Institution’s National Zoological Park, VA USA
Lecturer, Reproductive Biology Department of Veterinary Basic Sciences The Royal Veterinary College University of London London United Kingdom
Niamh M. Collins MVB, MSc
Diplomate ACVIM Clinical Assistant Professor in Equine Internal Medicine Marion DuPont Scott Equine Medical Center Virginia-Maryland Regional College of Veterinary Medicine
Ben Colenbrander DVM, PhD
Clovelly Intensive Care Unit Scone Equine Hospital Scone, NSW Australia
Anne Desrochers DVM
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Contributors Leesburg, VA USA
Fort Collins, CO USA
Paul J. De Vries DVM, SER (RNVA)
Carrie J. Finno DVM
De Veluwe Equine Clinic Heerde The Netherlands
Resident 3 College of Veterinary Medicine University of California-Davis Davis, CA USA
Thomas J. Divers DVM Diplomate ACVIM and ACVECC Professor of Medicine Department of Clinical Sciences College of Veterinary Medicine Cornell University Ithaca, NY USA
Keith W. Durkin PhD Research Assistant Department of Veterinary Integrative Biosciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University, TX USA
John F. Edwards Professor of Veterinary Pathology Department of Veterinary Pathobiology College of Veterinary Medicine and Biomedical Sciences Texas A&M University, TX USA
Bruce E. Eilts DVM, MS Diplomate ACT Professor of Theriogenology Department of Veterinary Clinical Sciences School of Veterinary Medicine Louisiana State University, LA USA
Rolf M. Emberston DVM Diplomate ACVS Equine Surgeon, Partner Rood and Riddle Equine Hospital Lexington, KY USA
Darien J. Feary BVSc, MS Diplomate ACVIM, ACVECC Lecturer in Equine Medicine Department of Veterinary Science University of Sydney University Veterinary Center Camden Campus, NSW Australia
Ryan A. Ferris DVM Clinical Instructor Department of Clinical Sciences Colorado State University Equine Reproduction Lab
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Elwyn C. Firth BVSc, MS, PhD, DSc Diplomate ACVS Professor Massey Equine, Institute of Veterinary Animal and Biomedical Sciences Massey University Palmerston North New Zealand
M. Julia B. Felippe Flaminio DVM, MS, PhD Diplomate ACVIM Assistant Professor of Medicine Department of Clinical Sciences College of Veterinary Medicine Cornell University Ithaca, NY USA
Robert P. Franklin DVM Diplomate ACVIM Chief of Internal Medicine Weatherford Equine Medical Center Weatherford, TX USA
Grant Frazer BVSc, MS Diplomate ACT, MBA Professor of Veterinary Reproduction Director, Veterinary Clinical & Diagnostic Services Veterinary Medical Center School of Veterinary Science University of Queensland Gatton, QLD Australia
David E. Freeman MVB, PhD Diplomate ACVS Professor and Interim Department Chair Chief of Large Animal Surgery Department of Large Animal Clinical Sciences University of Florida College of Veterinary Medicine Gainesville, FL USA
John F. Freestone BVSc, FRCVS, MACVSc Diplomate ACVIM Coolmore Australia, NSW Australia
Venkataseshu K. Ganjam BSc, BVSc, MS, PhD Professor Department of Biomedical Sciences College of Veterinary Medicine
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Contributors
University of Missouri Columbia, MO USA
Suffolk United Kingdom
James K. Graham PhD Eduardo L. Gastal DVM, MS, PhD Assistant Professor Department of Animal Science, Food and Nutrition Southern Illinois University Carbondale, IL USA
Earl M. Gaughan DVM Diplomate ACVS Surgeon Littleton Equine Medical Center Littleton, CO USA
Professor Department of Biomedical Sciences, Animal Reproduction and Biotechnology Laboratory Colorado State University Fort Collins, CO USA
Cleet E. Griffin DVM Clinical Assistant Professor Department of Large Animal Clinical Sciences College of Veterinary Medical and Biomedical Sciences Texas A&M University, TX USA
Sarah E.T. Hanna DVM Erica K. Gee BVSc, PhD Diplomate ACT Senior Lecturer Massey University Veterinary Teaching Hospital Massey Equine Institute of Veterinary, Animal and Biomedical Sciences Palmerston North New Zealand
Vincent Gerber DMV, PhD Diplomate ACVIM, Diplomate ECEIM Equine Clinic Department of Clinical Veterinary Medicine Vetsuisse Faculty University of Bern Bern Switzerland
` DVM, PhD Steeve Giguere Diplomate ACVIM Professor of Large Animal Internal Medicine Marguerite Thomas Hodgson Chair in Equine Studies University of Georgia Athens, GA USA
Lutz S. Goehring DVM, MS, PhD Diplomate ACVIM Assistant Professor in Equine Medicine Department of Clinical Sciences James L. Voss Veterinary Teaching Hospital College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO USA
Susan Gower I.S.T. SNR Immunohistochemist Department of Histology Paul Mellon Laboratory Newmarket
Associate Veterinarian Bristol Veterinary Service Union Grove, WI USA
Joanne Hardy DVM, PhD Diplomate ACVS, ACVECC Clinical Associate Professor Department of Veterinary Large Animal Clinical Sciences Texas A&M University, TX USA
David L. Hartman DVM Owner Hartman Equine Reproduction Center Whitesboro Gainesville, TX USA
Dean A. Hendrickson DVM, MS Professor of Surgery Department of Clinical Sciences College of Veterinary Medicine and Biomedical Sciences James L. Voss Veterinary Teaching Hospital Colorado State University Fort Collins, CO USA
Noah L. Heninger MS, PhD, DVM Associate Veterinarian Beard-Navasota Veterinary Hospital Los Lunas Animal Clinic Los Lunas, NM USA
Lynne M. Hewlett CHT, AS Technical Coordinator Department of Medicine Hagyard Equine Medical Institute Lexington, KY USA
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Contributors Melissa T. Hines DVM, PhD Diplomate ACVIM Associate Professor Veterinary Clinical Sciences Washington State University College of Veterinary Medicine Washington USA
Katrin Hinrichs DVM, PhD Diplomate ACT Professor and Patsy Link Chair in Mare Reproductive Studies Department of Veterinary Physiology and Pharmacology College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX USA
Aaron D.J. Hodder BVSc Diplomate ACT Edgewater Equine Clinic Santa Cruz, CA USA
Richard D. Holder DVM Hagyard Equine Medical Institute Lexington, KY USA
Katherine A. Houpt VMV, PhD Diplomate ACVB Professor Animal Behavior Clinic College of Veterinary Medicine Cornell University Ithaca, NY USA
John A.E. Hubbell DVM, MS, ACVA Professor of Veterinary Anesthesiology College of Veterinary Medicine The Ohio State University Columbus, OH USA
Peter Huntington BVSc Director of Nutrition Kentucky Equine Research Mulgrave, Victoria Australia
Avenel Victoria Australia
Mariana Ionita PhD, DVM Lecturer, Parasitology and Parasitic Diseases and Animal Biology Faculty of Veterinary Medicine of Bucharest University of Agronomical Sciences and Veterinary Medicine of Bucharest Bucharest Romania
Clifford H.G. Irvine∗ FACVSc DSc, DSc (hc), DVSc Professor Emeritus (Retired) Lincoln University Canterbury New Zealand
Stephen G. Jackson PhD President Bluegrass Equine Nutrition Versailles, KY USA
Sarah L. Jalim BVM&S, MACVSc Goulburn Valley Equine Hospital Congupna Victoria Australia
Aime K. Johnson DVM Diplomate ACT Assistant Professor of Theriogenology Department of Clinical Sciences College of Veterinary Medicine Auburn University Auburn, AL USA
Jill R. Johnson DVM, MS Diplomate ACVIM, ABVP Professor of Equine Medicine Department of Veterinary Clinical Sciences School of Veterinary Medicine Louisiana State University Baton Rouge, LA USA
Larry Johnson PhD
Diplomate ACT NANDI Veterinary Associates, LLC New Freedom, PA USA
Professor Department of Integrative Biosciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University, TX USA
John Hyland BVSc, PhD, FACVSc
Philip J. Johnson BVSc(Hons), MS, MRCVS
John P. Hurtgen∗ DVM, PhD
Avenel Equine Veterinary Surgery P/L
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Diplomate ACVIM, ECVIM
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Contributors
Professor Veterinary Medicine & Surgery College of Veterinary Medicine University of Missouri Columbia, MO USA
Teri L. Lear PhD Research Associate Professor Department of Veterinary Science Gluck Equine Research Center University of Kentucky Lexington, KY USA
Robert A. Kainer DVM, MS Professor Emeritus Department of Anatomy and Neurobiology Colorado State University Fort Collins, CO USA
Michelle M. LeBlanc DVM
Terttu Katila DVM, PhD, ECAR Diplomate
´ Renaud Leguillette DVM, PhD
Professor Faculty of Veterinary Medicine Department of Production Animal Medicine University of Helsinki Saarentaus Finland
Diplomate ACVIM Assistant Professor Department of Veterinary Clinical and Diagnostic Sciences Faculty of Veterinary Medicine University of Calgary Calgary, AB Canada
Sheryl S. King PhD, PAS Professor and Director of Equine Science Department of Animal Science, Food and Nutrition Southern Illinois University Carbondale, IL USA
Jay F. Kirkpatrick PhD Director The Science & Conservation Center ZooMontana Billings, MT USA
Derek C. Knottenbelt OBE, BVM&S, DVM&S, DipECEI, MRCVS Professor of Equine Internal Medicine Department of Veterinary Clinical Science University of Liverpool Leahurst Campus Neston Wirral United Kingdom
Julia H. Kydd PhD University Lecturer School of Veterinary Medicine & Science Sutton Bonington Campus University of Nottingham Loughborough Leicestershire United Kingdom
Laurie M. Lawrence PhD Professor Department of Animal and Food Sciences University of Kentucky Lexington, KY USA
Diplomate ACT Rood and Riddle Equine Hospital Lexington, KY USA
Timothy B. Lescun BVSc, MS Diplomate ACVS Associate Professor Department of Veterinary Clinical Sciences Purdue University West Lafayette, IN USA
Guy D. Lester School of Veterinary and Biomedical Science Division of Health Science Murdoch University Murdoch Australia
William B. Ley DVM, MS Diplomate ACT Horse-Repro.Com, PLLC Rectortown, VA USA
Alicia R.G. Lindholm DVM Equine Reproduction Laboratory Department of Clinical Sciences Colorado State University Fort Collins, CO USA
Valerie J. Linse MS, DVM Hagyard Equine Medical Institute Lexington, KY USA
Irwin K.M. Liu DVM, MPVM, PhD Professor Emeritus (Retired) Department of Population Health and Reproduction School of Veterinary Medicine
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Contributors University of California Davis, CA USA
Dawn A. Loesch DVM Diplomate ACVS 1653 Pinewood Lake Court, Greenacres West Palm Beach, FL USA
¨ Robert M. Lofstedt BVSc, MS Diplomate ACT Professor of Theriogenology Department of Health Management Atlantic Veterinary College at UPEI Charlottetown Prince Edward Island Canada
Paul R. Loomis MS Founder, CEO Select Breeders Service, Inc. 1088 Nesbitt Rd. Colora, MD USA
Charles C. Love DVM, PhD Associate Professor Section of Theriogenology Department of Large Animal Clinical Sciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University, TX USA
Eugene T. Lyons PhD Professor Department of Veterinary Science Gluck Equine Research Center University of Kentucky Lexington, KY USA
Robert J. MacKay BVSc, PhD Diplomate ACVIM Professor Alec P. and Louise H. Courtelis Equine Teaching Hospital University of Florida Gainesville, FL USA
Jennifer M. MacLeay DVM, PhD Diplomate ACVIM Medical Director Research Hills Pet Nutrition Inc, KS USA
Lisa J. Maclellan MS, PhD Research Scholar Equine Reproduction Laboratory Department of Veterinary and Biomedical Sciences Colorado State University, Fort Collins, Colorado Seven Creeks Equine Veterinary Clinic, Victoria Australia
Margo L. Macpherson DVM, MS
Diplomate ACT Associate Veterinarian Hagyard Equine Medical Institute Lexington, KY USA
Diplomate ACT Associate Professor Service Chief, Theriogenology Large Animal Clinical Sciences College of Veterinary Medicine University of Florida Gainesville, FL USA
D. Paul Lunn BVSc, MS, PhD MRCVS
Scott Madill BVSc, DVSc,
Diplomate ACVIM Professor and Head Department of Clinical Sciences James L. Voss Veterinary Teaching Hospital College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO USA
Diplomate ACT Assistant Clinical Specialist Veterinary Population Medicine University of Minnesota St. Paul, MN USA
Kristina G. Lu VMD
Sara K. Lyle DVM, PhD Diplomate ACT Assistant Professor of Theriogenology Department of Veterinary Clinical Sciences Louisiana State University Baton Rouge, LA USA
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K. Gary Magdesian DVM Diplomate ACVIM, ACVECC, ACVCP Associate Professor Equine Medicine and Critical Care Department of Veterinary Medicine and Epidemiology School of Veterinary Medicine University of California Davis, CA USA
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Contributors
Omar Maher DV Diplomate ACVS Surgeon New England Equine Medical & Surgical Center Dover, NH USA
Celia M. Marr BVMS, MVM, PhD, MRCVS Diplomate EIM, ECEIM Specialist in Equine Internal Medicine Rossdale Equine Hospital Newmarket Suffolk United Kingdom
Eliane Marti Associate Professor Division of Clinical Research Department of Clinical Veterinary Medicine Vetsuisse Faculty, University of Berne Berne Switzerland
Michael T. Martin DVM Associate Professor Department of Large Animal Clinical Sciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX USA
Phillip McCarthy∗ DVM Lexington, KY USA
Patrick M. McCue DVM, PhD Diplomate ACT Professor Animal Reproduction and Biotechnology Laboratory Colorado State University Fort Collins, CO USA
Sue M. McDonnell MS, PhD Cert Applied Animal Behaviorist Adjunct Professor Section of Reproduction Department of Clinical Studies New Bolton Center School of Veterinary Medicine University of Pennsylvania Kennett Square, PA USA
Karen J. McDowell PhD Associate Professor Department of Veterinary Science University of Kentucky
Lexington, KY USA
Paul McGreevy BVSc, PhD, MACVSc Associate Professor Faculty of Veterinary Science University of Sydney Sydney, NSW Australia
C. Wayne McIlwraith BVSc, PhD, DSc, FRCVS Diplomate ACVS University Distinguished Professor Barbara Cox Anthony University Chair in Orthopedics; Director of Orthopedic Research Colorado State University Fort Collins, CO USA
Erica C. McKenzie BSc, BVMS, PhD Diplomate ACVIM College of Veterinary Medicine Oregon State University Corvallis, OR USA
Angus O. McKinnon BVSc, MSc Diplomate ACT and ABVP Goulburn Valley Equine Hospital Congupna, Victoria Australia
Nat T. Messer DVM Professor Department of Veterinary Medicine & Surgery College of Veterinary Medicine University of Missouri Columbia, MO USA
Elizabeth S. Metcalf DVM, MS Diplomate ACT CEO Honahlee Reproduction PC, OR USA
Stuart A. Meyers DVM, PhD Professor Department of Anatomy, Physiology and Cell Biology School of Veterinary Medicine University of California Davis, CA USA
Vicki N. Meyers-Wallen VMD, PhD Diplomate ACT Associate Professor Baker Institute for Animal Health College of Veterinary Medicine Cornell University
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Contributors Ithaca, NY USA
James P. Morehead DVM Equine Medical Associates, PSC Lexington, KY USA
Peter R. Morresey BVSc, MACVSc
Jennifer C. Ousey PhD Post-Doctoral Scientist Rossdale & Partners, Beaufort Cottage Laboratories Newmarket Suffolk United Kingdom
Dale L. Paccamonti DVM, MS
Diplomate ACT and ACVIM Associate Department of Internal Medicine Rood and Riddle Equine Hospital Lexington, KY USA
Diplomate ACT Professor and Head School of Veterinary Medicine Louisiana State University Baton Rouge, LA USA
Lee H.A. Morris BVSc, DVSc
Joe Pagan PhD
Diplomate ACT Director EquiBreed NZ Ltd Cambridge New Zealand
President Kentucky Equine Research Versailles, KY USA
Laura C. Nath BVSc (Hons) Cert EM (Stud Medicine), FACVSc Associate Veterinarian Flemington Equine Clinic Melbourne Australia
Kirsten M. Neil BVSc (Hons), MACVSc, MS Diplomate ACVIM Specialist in Equine Medicine Sporthorse Veterinary Specialists Berwick Victoria Australia
John R. Newcombe BVM, MRCVS Principal, Equine Fertility Clinic Newcombe and East, Veterinary Surgeons Brownhills West Midlands United Kingdom
Gary J. Nie DVM, MS, PhD Diplomate ACT, ABVP, ACVIM, VMA Owner, Veterinarian Word Wide Veterinary Consultants Springfield, MO USA
˜ PhD Cassandra M.V. Nuηez Associate Research Scholar Department of Ecology & Evolutionary Biology Princeton University Princeton, NJ USA
Lawrence M. Olsen DVM, PhD Owner/Veterinarian Newman, GA USA
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Jonathan E. Palmer VMD Diplomate ACVIM Director of Perinatology/Neonatology Programs; Chief, Neonatal Intensive Care Service Connelly Intensive Care Unit New Bolton Center University of Pennsylvania Kennett Square, PA USA
Scott E. Palmer VMD Diplomate ABVP Equine Practice Hospital Director New Jersey Equine Clinic, NJ USA
Joan Palmero DVM Resident Large Animal Internal Medicine School of Veterinary Medicine University of California Davis, CA USA
Mary Rose Paradis DVM, MS Diplomate ACVIM-LAIM Associate Professor Department of Clinical Sciences Tufts Cummings School of Veterinary Medicine Tufts University North Grafton, MA USA
Eric J. Parente DVM Diplomate ACVS Associate Professor of Surgery New Bolton Center Department of Clinical Sciences
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Contributors
University of Pennsylvania, PA USA
Nigel R. Perkins BSVc (Hons), MS, PhD Diplomate ACT, FACVSc Director AusVet Animal Health Services Program Co-ordinator, Australian Biosecurity CRC; Research Manager Horse R&D Program RIRDC, QLD Australia
Bill W. Pickett PhD Professor Emeritus Animal Reproduction and Biotechnology Laboratory College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO USA
Carlos R.F. Pinto MedVet, PhD Diplomate ACT Associate Professor Section Head Theriogenology and Reproductive Medicine Department of Veterinary Clinical Sciences The Ohio State University College of Veterinary Medicine Columbus, OH USA
Caryn E. Plummer DVM Diplomate ACVO Assistant Professor, Comparative Ophthalmology College of Veterinary Medicine University of Florida Departments of Large and Small Animal Clinical Sciences Gainesville, FL USA
Patrick J. Pollock BVMS CertES(Soft Tissue), Diplomate ECVS, MRCVS European and Royal College Recognised Specialist in Equine Surgery; Senior Lecturer in Equine Surgery Weipers Centre for Equine Welfare Faculty of Veterinary Medicine University of Glasgow Glasgow Scotland United Kingdom
David G. Powell BVSc, FRCVS 48, Sarno Square Abergavenny Wales United Kingdom
David G. Pugh DVM, MS Diplomate ACT, ACVN
Veterinary Consultant Professional Services Fort Dodge Animal Health Waverly, AL USA
Jonathan F. Pycock BVM, PhD, DESM, MRCVS RCVS Specialist in Equine Medicine (Reproduction) Equine Reproductive Services Ryton Yorkshire United Kingdom
Sarah L. Ralston VMD, PhD Diplomate ACVN Associate Professor Department of Animal Science Rutgers University New Brunswick, NJ USA
Terje Raudsepp PhD Associate Professor of Molecular Cytogenetics Department of Veterinary Integrative Biosciences College of Veterinary Medicine Texas A&M University College Station, TX USA
Jennifer R. Read BVMS (Hons) Resident Large Animal Medicine Department of Clinical Studies University of Pennsylvania School of Veterinary Medicine, PA USA
Virginia B. Reef DVM Diplomate ACVIM - LAIM Mark Whitner and Lila Griswold Allam Professor of Medicine Department of Clinical Studies New Bolton Center University of Pennsylvania, PA USA
Sidney W. Ricketts LVO, BSc, BVSc, DESM, DipECEIM, FRCPath, FRCVS RCVS Recognised Specialist in Equine Medicine (Stud Medicine) Rossdale and Partners Newmarket Suffolk United Kingdom
Simon J. Robinson BVSc, BScAgr, MACVSc Associate Veterinarian Goulburn Valley Equine Hospital Congupna Victoria Australia
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Contributors Dwayne H. Rodgerson DVM, MS
Elizabeth M. Santschi DVM
Diplomate ACVS Surgeon Davidson Surgery Center Hagyard Equine Medical Institute Lexington, KY USA
Diplomate ACVS Clinical Associate Professor Department of Veterinary Clinical Sciences The Ohio State University Columbus, OH USA
Anna K. Roetting DMV, PhD
C.J. (Kate) Savage BVSc(hons), MS, PhD
Diplomate ACVS, ECVS Section Head, Equine Surgery Clinic for Horses University of Veterinary Medicine Hannover Hannover Germany
Diplomate ACVIM Specialist in Equine Medicine Equine Centre University of Melbourne Victoria Australia
Janet F. Roser MS, PhD
Donald H. Schlafer DVM, MS, PhD
Professor Department of Animal Science University of California Davis, CA USA
Daniel I. Rubenstein PhD Class of 1877 Professor and Chair Department of Ecology and Evolutionary Biology Princeton University Princeton, NJ USA
Thomas M. Russell BVMS, MACVSc, Dip ECVS Head of Complicated Surgery Goulburn Valley Equine Hospital Gongupna Victoria Australia
Tom Russell Diplomate ECVS Senior Surgeon Goulburn Valley Equine Hospital Congupna Victoria Australia
Juan C. Samper DVM, MSc, PhD Diplomate ACT Owner JCS Veterinary Reproductive Services Ltd Langley British Columbia Canada
L. Chris Sanchez DVM, PhD Diplomate ACVIM Associate Professor Large Animal Clinical Sciences College of Veterinary Medicine University of Florida Gainesville, FL USA
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Professor of Theriogenologic Pathology Department of Biomedical Sciences Section of Anatomic Pathology College of Veterinary Medicine Cornell University Ithaca, NY USA
James Schumacher Department of Large Animal Clinical Sciences College of Veterinary Medicine University of Tennessee Knoxville, TN USA
Charles F. Scoggin DVM, MS Resident Veterinarian Claiborne Farm Lexington, KY USA
George E. Seidel, Jr. PhD Professor Department of Biomedical Sciences Colorado State University Animal Reproduction and Biotechnology Laboratory Fort Collins, CO USA
Patricia L. Sertich MS, VMD Diplomate ACT Associate Professor - Clinical Educator Section of Reproduction School of Veterinary Medicine University of Pennsylvania Kennett Square, PA USA
Daniel C. Sharp PhD Professor Emeritus Department of Animal Sciences University of Florida; Consultant
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Contributors
Peterson & Smith Equine Reproduction Center Gainesville, FL USA
Davis, CA USA
Sharon J. Spier DVM, PhD
Professor University of Melbourne The Dean’s Ganglion Melbourne Victoria Australia
Diplomate ACVIM Professor Department of Medicine and Epidemiology College of Veterinary Medicine University of California Davis, CA USA
Paul D. Siciliano PhD
Michael A. Spirito DVM
Roger V. Short PhD
Associate Professor Department of Animal Science North Carolina State University Raleigh, NC USA
Harald Sieme DVM
Surgeon Department of Surgery Hagyard Equine Medical Institute Lexington, KY USA
Kim A. Sprayberry DVM
Professor of Equine Reproduction Unit for Reproductive Medicine - Clinic for Horses University of Veterinary Medicine Hannover Hannover Germany
Diplomate ACVIM Internist McGee Medical Center Hagyard Equine Medical Institute Lexington, KY USA
Barry W. Simon DVM
Edward L. Squires MS, PhD
COO, CreoSalus Inc. Thorn BioScience LLC Lexington, KY USA
Sophie Sitters DVM Graduate, Department of Equine Sciences Faculty of Veterinary Medicine Utrecht University Utrecht The Netherlands
JoAnn Slack DVM, MS Diplomate ACVIM, LAIM Assistant Professor of Cardiology and Ultrasound Department of Clinical Studies New Bolton Center University of Pennsylvania, PA USA
Nathan M. Slovis DVM Diplomate ACVIM, CHT Director McGee Medical Center Hagyard Equine Medical Institute Lexington, KY USA
Jack R. Snyder DVM, PhD Diplomate ACVS Professor Surgical and Radiological Sciences University of California
Diplomate ACT (hon) Executive Director Gluck Equine Research Foundation; Research Professor Department of Veterinary Science Gluck Equine Research Center University of Kentucky Lexington, KY USA
Simon A. Staempfli DMV, DACT, DABVP, MRCVS The Arundel Equine Hospital LLP West Sussex United Kingdom
Mary E. Stanton DVM Diplomate ACT Equine Veterinary Reproduction Specialists of Ocala Ocala, FL USA
John V. Steiner∗ DVM Diplomate ACT Rhinebeck Equine Rhinebeck, NY USA
Sarah J. Stoneham BVSc Cert ESM, MRCVS Director Lone Oak Stud Newmarket Suffolk United Kingdom
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Contributors Tom A.E. Stout MA, Vet MB, PhD, MRCVS Diplomate ECAR Professor Professor of Equine Medicine and Reproduction Department of Equine Sciences-Utrecht University Utrecht The Netherlands
Rebecca L. Tallmadge PhD Research Associate Department of Clinical Sciences Cornell University College of Veterinary Medicine Ithaca, NY USA
Caroline Tessier DVM, MS Diplomate ACVS, Diplomate ECVS Equine Clinic Department of Clinical Veterinary Medicine Vetsuisse Faculty University of Bern Bern Switzerland
Donald L. Thompson, Jr. PhD Professor School of Animal Sciences Louisiana State University Agricultural Center Louisiana State University Baton Rouge, LA USA
Walter R. Threlfall DVM, MS, PhD Diplomate ACT Director Veterinary Continuing Education College of Veterinary Medicine The Ohio State University Veterinary Medical Center Columbus, OH USA
Ahmed Tibary DMV, MS, DSc, PhD Diplomate ACT Professor Department of Veterinary Clinical Sciences College of Veterinary Medicine Washington State University Pullman, WA USA
John F. Timoney DSc, MVB, MS, PhD, MRCVS Keeneland Chair of Infectious Diseases Gluck Equine Research Center University of Kentucky Lexington, KY USA
Peter J. Timoney MVB, MS, PhD, FRCVS Professor Frederick Van Lennep Chair in Equine Veterinary Science
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Department of Veterinary Science Gluck Equine Research Center University of Kentucky Lexington, KY USA
Sharon C. Tolliver MS Department of Veterinary Science Gluck Equine Research Center University of Kentucky, KY USA
Mats H.T. Troedsson DVM, PhD, DACT, DECAR Director Gluck Equine Research Center; Professor and Chair Department of Veterinary Science University of Kentucky Lexington, KY USA
John W. Turner Professor Department of Physiology and Pharmacology University of Toledo College of Medicine Toledo, OH USA
Regina M.O. Turner VMD, PhD Diplomate ACT Assistant Professor of Large Animal Reproduction Department of Clinical Studies New Bolton Center University of Pennsylvania School of Veterinary Medicine Kennett Square, PA USA
Norman W. Umphenour DVM Resident Veterinarian Ashford Stud Lexington, KY USA
Wendy E. Vaala VMD Dipomate ACVIM Sr. Equine Technical Service Veterinarian Intervet Schering-Plough Animal Health Alma, WI USA
Stephanie J. Valberg DVM, PhD Diplomate ACVIM Professor College of Veterinary Medicine University of Minnesota St. Paul, MN USA
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Contributors
Dirk K. Vanderwall DVM, PhD
Bryan M. Waldridge DVM, MS
Diplomate ACT Associate Professor and Chief Section of Reproduction Department of Clinical Studies New Bolton Center School of Veterinary Medicine University of Pennsylvania Kennett Square, PA USA
Diplomate ABVP (Equine Practice), ACVIM Kentucky Equine Research Versailles, KY USA
Dickson D. Varner DVM, MS Diplomate ACT Professor of Theriogenology Pin Oak Stud Chair of Stallion Reproductive Studies Department of Large Animal Clinical Sciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX USA
James R. Vasey Partner Goulburn Valley Equine Hospital Congupna Victoria Australia
D.N. Rao Veeramachaneni BVSc, MScVet, PhD
Jillian J. Wall BS Research Associate Department of Biomedical Sciences Colorado State University Fort Collins, CO USA
Katherine E. Whitwell BVSc, Dipl ECVP, FRCVS RCVS Recognised Specialist in Veterinary Pathology (Equine) Equine Pathology Consultancy Newmarket Suffolk United Kingdom
Robyn R. Wilborn DVM Diplomate ACT Department of Clinical Sciences College of Veterinary Medicine Auburn University Auburn, AL USA
Professor Animal Reproduction and Biotechnology Laboratory Department of Biomedical Sciences College of Veterinary Medicine and Biomedical Science Colorado State University Fort Collins, CO USA
Pamela A. Wilkins DVM, MS, PhD
Daniel A.F. Villagomez PhD, DVM
Sandra Wilsher BSc (Hons), PhD
Departmento de Produccion Animal Universidad de Guadalajava Zapopan Mexico
Sally L. Vivrette DVM, PhD Diplomate ACVIM Triangle Equine Mobile Veterinary Services Cary, NC USA
Rudolf O. Waelchli DMV, MSc Dr. habil Research Associate Biomedical Sciences Ontario Veterinary College University of Guelph, Guelph; Associate Veterinarian Dufferin Veterinary Services Orangeville, ON Canada
Diplomate ACVIM (LA), Diplomate ACVECC Professor of Equine Internal Medicine and Emergency/Critical Care Department of Veterinary Clinical Medicine College of Veterinary Medicine University of Illinois, IL USA
Senior Research Associate The Paul Mellon Laborabory of Equine Reproduction Cheveley Newmarket United Kingdom
David G. Wilson DVM Diplomate ACVS Professor of Large Animal Surgery Department of Large Animal Clinical Sciences Western College of Veterinary Medicine University of Saskatchewan Saskatoon Saskatchewan Canada
W. David Wilson BVMS, MS Professor Director of the Veterinary Medical Teaching Hospital and Associate Dean for Clinical Programs Department of Medicine and Epidemiology
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Contributors School of Veterinary Medicine University of California Davis, CA USA
Brett J. Woodie DVM, MS Diplomate ACVS Surgeon Rood and Riddle Equine Hospital Lexington, KY USA
Walter W. Zent DVM Diplomate ACT (hon) Partner Hagyard Equine Medical Institute Theriogenology Department Lexington, KY USA ∗
Deceased
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Preface to the second edition This second edition of Equine Reproduction is, like the first edition, dedicated to ‘the horse.’ Our unique bond, which eludes simple definition, comes from a recognition that horses are truly different from any other domestic animal. Horses are unique not just in their reproductive physiology but in their place in our society. The horse can be a companion, a work animal, a performance animal, or simply something ethereal for its owners. All of the editors have spent a considerable part of their lives riding and educating young horses as well as managing foal and breeding farm problems. We all share a common bond of gratitude to the horse for the gifts it has provided. Readers are referred to the preface from the first edition, as many of the principles, ideals, and reasons for commissioning this text are still fundamentally relevant. Most notable is the continual education opportunities for our clients and the recognition that we must remain current. In addition we feel that a broad base of knowledge is important for veterinarians working on breeding farms. To this end we question the training of veterinarians as theriogenologists who do not understand how to evaluate a sick foal or a lame broodmare. To be competitive in the employment market, young veterinarians must be proficient in more than just reproductive management. This has resulted in an increase in scope of this edition. The expansion in chapter numbers from 117 to 325 resulted in a logistic challenge to edit accurately and expediently. A total of 250 authors have contributed. The huge investment in time and energy from the authors has been repaid not just with an increase in our knowledge but also the satisfaction that the new edition is an accurate reflection of the current level of knowledge in equine reproduction. Only time will tell how long this book remains current. Certain deficiencies in the previous edition were recognized by us, most specifically the level of information on the foal. We have aimed to remedy that by attempt-
ing to make the foal section the most comprehensive treatise of foal diseases and management written to date. The recognition that veterinarians working on breeding farms need to provide advice on non-reproductive management, and the requirement on many farms to radiograph pre-sale and examine the upper respiratory system with endoscopy, has resulted in specific chapters to address those issues. Working with authors has been a rewarding experience. The level of dedication and professionalism continues to surprise and inspire us. There were no complaints when the editors suggested that further work (or, on occasion, less work) should be included. Although authorship is often dictated by our affiliations, all chapters are written by recognized leaders in their field. Special efforts in authorship are gratefully noted from Jane Axon, Pat McCue, and Ahmed Tibary. These authors prepared extra chapters, in addition to their initial chapter requests, and often at short notice, when it became apparent that others had been waylaid. The retirement of Jim Voss meant that his unique editing style has been greatly missed. His retirement presented us with an opportunity to bring in three more editors to share responsibility and rebalance the team. We hope that everything we have strived for meets your expectations as readers. The editorial process has been a long and sometimes difficult one, but in the final analysis we feel it was well worth the effort. As with the previous edition, all proceeds due to editors and contributors have been allocated, by common agreement, to equine reproductive research, the distribution of which will be determined by the individual editors.
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A.O. McKinnon, E.L. Squires, W.E. Vaala and D.D. Varner
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Preface to the first edition The text was generated by our desire to create a useful addition to the available literature on all aspects of equine reproduction. Our idea was that practising equine veterinarians needed a text devoted exclusively to equine reproduction, which while showing due respect to historical perspectives, should collate current information in a clear, concise, and easily accessible form. In addition, a perceived disproportionate interest in reproductive physiology of the mare should be addressed by inclusion of sections on the stallion and neonate. The practice of veterinary medicine is progressing rapidly toward species specialization; within this, discipline specialization now occurs. However, the nature of our service necessitates talented general practitioners, and for the foreseeable future, they will remain the backbone of the profession. At the same time, education of the client has improved through greater access to information, and many clients have become more demanding and critical of veterinary services. This has placed many of us in unenviable situations. How do we remain cognizant of all the latest and pertinent research findings? Publication of most current research, at best, is scattered widely through many research and veterinary educational journals. Researchers without a veterinary educational background have contributed greatly to recent discoveries and have published reports, albeit unwittingly, in journals beyond our usual access. How many practicing veterinarians routinely scrutinize journals such as Biology of Reproduction, Animal Science, or even Reproduction and Fertility, apart from special supplements? Similarly, how many basic and applied equine researches read the Proceedings of the American Associations of Equine Practitioners or articles in popular equine magazines and have a real understanding of the relative importance of problems of the equine industry? A need seemed to exist to combine recent information within the discipline of equine reproduction into a format useful for practicing equine veterinar-
ians. Veterinarians in mixed practice, veterinary students, basic and applied scientists, and graduate students may benefit from the breadth and specificity of the information presented her in 118 well referenced chapters. The text is divided into three main sections: the mare, the stallion, and the neonate. Each is equally important to veterinarians involved in equine reproductive practice. Too often, interest in the mare wanes once her pregnancy is diagnosed. Monitoring of pregnancy and neonatal care are areas of prime importance, because presumably the aim of any breeding program is to produce viable offspring. In addition, education regarding stallion reproductive physiology and management is alarmingly sparse, and the stallion is often neglected despite being half the fertility equation. The discussions of the mare and stallion are divided into the following topics: anatomy, physiology, and endocrinology; breeding management; diseases of the reproductive tract; and reproductive surgery. Additional topics in the section on the mare are pregnancy, parturition, and the puerperal period. Discussion of the neonate is divided into neonatal care and diseases according to body systems. Ninety-five authors and co-authors have contributed to this work. The number of North American authors reflects out affiliations; however, readers should not chapters by numerous internationally recognized educators from both Northern and Southern Hemispheres. Editing the contributors has been an enriching experience for both of us. The satisfaction from learning something new in each chapter has been incalculable. We are pleased to note that all proceeds that would normally be due contributors of this text will be allocated by common agreement to equine reproductive research.
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Sheppartown, Victoria Australia Angus O. McKinnon Fort Collins, Colarado, USA James. L. Voss
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Acknowledgments The Editors would like to acknowledge the assistance of friends, colleagues and family who either knowingly or unwittingly contributed to the completion of the book through generous giving of time or tacit approval of our editorial absences. The time spent editing and writing necessitated more than our own commitments and we are deeply grateful for the understanding that was shown to us all.
The assistance of the staff of Wiley-Blackwell was always available and the team assembled to guide us through the maze of time schedules and editorial commitments was most approachable and dedicated. There were the issues of the number of color plates and if a full color book could be financially viable that were a source of frustration, however these were offset by the wonderful help from Antonia Seymour, Mirjana Misina and Teresa Netzler.
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PART I
The Foal
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Section (a): Prepartum Assessment of the Foal
1.
Identification of the High-Risk Pregnancy Elizabeth M. Santschi and Wendy E. Vaala
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2.
Monitoring the High-Risk Pregnancy Wendy E. Vaala
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3.
Management of the High-Risk Pregnancy Elizabeth M. Santschi and Wendy E. Vaala
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4.
Ultrasonographic Monitoring of the Fetus Stefania Bucca
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5.
Sedation and Anesthesia of the Pregnant Mare John A.E. Hubbell
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Chapter 1
Identification of the High-Risk Pregnancy Elizabeth M. Santschi and Wendy E. Vaala
Introduction Very few sights are simultaneously as placid and potentially unnerving to an equine clinician as contemplating a group of valuable broodmares in late-term pregnancy. While the majority of pregnancies end with a viable mare and foal, a small but very real percentage of pregnancies end in death or disability for dam or neonate. These high-risk pregnancies often do not show premonitory signs before real trouble begins, and are a challenge for horse owners, animal care personnel and veterinarians. Considerable time and money are spent manipulating the mare’s reproductive cycle to ensure conception and early foaling dates. Numerous sonographic examinations are performed early in the reproductive process to detect ovulation, coordinate insemination or breeding, implant embryos, confirm pregnancy, and rule out twinning. But after the first 45 to 60 days the typical broodmare does not receive close, routine scrutiny again until the last 4–8 weeks of gestation. Consequently many causes of abortion and fetal/neonatal morbidity and mortality during the latter two-thirds of pregnancy remain poorly understood. Several large retrospective necropsy studies1, 2 have examined the causes of abortion, stillbirth, and perinatal death in horses. Placentitis, due to bacterial, viral and fungal causes, and delivery complications associated with dystocia, birth trauma, and asphyxia represented the most common causes of fetal and neonatal foal mortality. Conditions that cause high-risk pregnancy can originate in the mare, fetus or placenta. While some situations defy identification before crisis, many can be detected, and intervention can be instituted. As we learn more, treatment is improving the outcomes of
equine high-risk pregnancies. Successful intervention in high-risk pregnancy will require identification of risk at the earliest possible time, and is the goal of the equine perinatologist. Conditions that cause risk to the mare and foal can be subdivided into conditions of the mare, the foal, and the placenta (including umbilical cord) (Tables 1.1–1.3). While these categorizations are not absolute, analyzing them in this way can demonstrate trends in diagnosis and prognosis. In general, maternal conditions are discovered by the clinical signs a mare exhibits and by physical examination. Pain, whether abdominal or lameness, is a frequent component. Assuming appropriate therapy, the risk to the mares’ health for maternal conditions is largely determined by the severity of the disease. The fetus is affected in parallel by the maternal systemic condition and potentially by the drugs and therapies administered to the dam. For lameness, unless it is debilitating, the impact is usually minimal, but for conditions with a systemic maternal component, the risk to the foal is roughly an order of magnitude greater for the foal than the mare. Our experience is that most preterm deliveries are the result of fetal death, and the fatal insult usually involves interference with the transfer of nutrients and oxygen and the removal of waste at the placenta. Placental and umbilical conditions range from clinically silent to those easily detected by an experienced clinician. The most common premonitory signs of placentitis are premature lactation and vulvar discharge, but pregnancies affected with hydrops allantois will present with an abnormally enlarged abdomen, a small fetus, and some degree of abdominal or respiratory distress. Most placental conditions pose little risk to the mare (hydrops being the notable exception) but all pose significant risk to the fetus.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Prepartum Assessment of the Foal
Table 1.1 Maternal conditions causing high-risk pregnancy, most common clinical signs, and risk estimate to mare and foal Maternal condition
Premonitory clues
Risk to mare
Risk to foal
Visceral colic Uterine torsion Endotoxemia Laminitis Other severe lameness Abdominal wall injury Infectious disease Misshapen pelvis Poor quality endometrium
Abdominal pain Abdominal pain, rectal palpation of torsion Fever, elevated heart rate, dehydration Lameness Lameness Pain, swelling of ventral or lateral abdomen Fever, renal, respiratory signs Rectal exam reveals compromise to pelvic inlet None
Varies with severity of disease Mild to moderate Varies with severity of disease Varies with severity of disease Varies with severity of disease Varies with severity of disease Usually minimal Minimal, but can be severe from dystocia None
Mild to moderate Moderate to severe Moderate to severe Mild Usually mild Moderate to severe Moderate to severe Moderate to severe Moderate to severe
The detection or prediction of fetal conditions causing high-risk pregnancy fall into two categories, those predictable by historical information about the previous pregnancy or suspicious signs during the current pregnancy and those rarely detected or not detectable at this stage of our knowledge. Maternal risk from these conditions is either none or very serious, and the risk to the foal is often severe. Any mare at risk for a complicated pregnancy or delivery should have a minimum data base established (Table 1.4) that includes a thorough history, general physical examination, and reproductive tract evaluation. Sonographic evaluation and serial monitoring of the fetus, uterus, and placenta is also recommended. Additional tests are performed as indicated by the primary underlying condition.
Conditions that can cause high-risk pregnancy Maternal conditions
Gastrointestinal causes of colic Most colic episodes that occur during pregnancy are mild and resolve with no or minimal treatment.
These episodes pose little danger to either the mare or her pregnancy, although frequent episodes, even if mild, can be a cause for concern. Pregnancy can complicate the determination of the best course of therapy for a mare with colic in late pregnancy, as the large uterus and fetus make effective rectal palpation impossible. The pregnant uterus can similarly make obtaining a diagnostic abdominal fluid sample difficult. Transabdominal ultrasonography can be helpful in location of a site for abdominocentesis and for the diagnosis of a cause for colic, but the large uterus also complicates this diagnostic test, and in a late pregnant mare the decision for surgery is often made based on pain and the response to analgesics, presence of reflux, and lack of fecal passage. Colic that requires surgery but is swiftly resolved is likely to have minimal impact on fetal outcome during most of gestation.3, 4 An exception is a mare with colic in late-term pregnancy, when placental oxygenation is both critical and vulnerable.3 Hypoxia in the last 60 days of pregnancy results in poorer fetal outcomes for those dams than mares without hypoxia.3 Short surgical times, appropriate ventilation, and oxygen insufflation can help to promote delivery of oxygen to the fetus; however, in some cases, maintaining appropriate oxygenation is not possible.
Table 1.2 Placental or umbilical conditions causing high-risk pregnancy, most common clinical signs, and risk estimate to mare and foal Conditions of placenta or umbilicus
Premonitory clues
Risk to mare
Risk to foal
Infectious placentitis
None, premature lactation, uterine discharge
No direct risk. Indirect risk from dystocia
Moderate to severe
Premature placental separation
Occasionally premature lactation, rarely hemorrhage
No direct risk. Indirect risk from dystocia
Moderate to severe
Fescue toxicosis Umbilical abnormalities Hydrops allantois
Prolonged gestation, failure of lactation None Severe abdominal enlargement due to uterus
No direct risk. Indirect risk from dystocia No direct risk. Indirect risk from dystocia Severe
Severe Severe Severe
Placental insufficiency
Possibly prolonged gestation
None
Moderate to severe
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Identification of the High-Risk Pregnancy
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Table 1.3 Fetal conditions causing high-risk pregnancy, most common clinical signs, and risk estimate to mare and foal Conditions of the foal
Premonitory clues
Risk to mare
Risk to foal
Neonatal isoerythrolysis
Historical production of NI foal; testing for antibodies
None
Moderate to severe
Twins Fetal malformation Arthrogryposis Fetal malpresentation Mare reproductive loss syndrome
Twins detected post breeding None None None None
Moderate to severe Moderate to severe Moderate to severe Moderate to severe Minimal
Moderate to severe Severe Severe Moderate to severe Severe
Pregnancies continuing after any surgery should be considered as at-risk and should receive extra scrutiny, but those in which intraoperative oxygenation was poor should be considered at ‘high risk.’ This determination necessitates more aggressive antepartum evaluation of the fetus and placenta in the postoperative treatment period, which is discussed in detail in Chapters 2 and 4. Table 1.4 Data base for high-risk pregnancies Signalment of mare Age Breed Parity Past reproductive history Outcome of past pregnancies; if abnormal, was the cause prepartum, intrapartum, or postpartum in origin Length of gestation(s) Description of periparturient behavior Results of last uterine biopsy/culture Current general health and reproductive history Vaccination history, deworming regimen, diet Description of current medical/surgical conditions List of medications administered during pregnancy Date last bred; 340-day due date Presence of vulvar discharge, premature udder development or lactation Prepartum evaluation and monitoring Physical examination with vital signs Bodyweight Rectal palpation to assess cervix, gravid uterus Vaginal examination (only if clinically indicated) Transrectal ultrasonography to evaluate pericervical utero-placental integrity, fetal fluid clarity, fetal activity Transabdominal ultrasonography to evaluate fetal heart rate (range and reactivity), breathing movements, tone, position and size, fetal fluid clarity and volumes, utero-placental integrity Maternal complete blood count/serum biochemistries Serial monitoring of progestagen (progesterone) concentrations Mammary secretion electrolyte concentrations Frequent evaluation of pelvic ligament relaxation, udder development, vulva elongation Regular observation for signs of parturition
Endotoxemia Endotoxemia resulting from colic is a separate but very important concern. Pregnant mares that show clinical or laboratory signs of endotoxemia such as fever, tachycardia, dehydration, leukopenia, and peripheral vasodilatation/hypotension or cyanosis are at great risk of fetal compromise. Quick resolution of the primary condition is paramount and, when that is not possible, mares should be closely monitored for fetal distress or death. Clinical signs of endotoxemia in pregnant mares with colic are associated with poor fetal outcomes3, 4 and all pregnant mares, after treatment of colic, should undergo fetal ultrasonography periodically while hospitalized. Similarly, endotoxemia resulting from other conditions such as pleuritis, pneumonia, colitis, and peritonitis will have a negative impact on the fetus and should engender similar concern and scrutiny. It is the author’s (EMS) observation that pregnancies in mid-gestation are the least susceptible to the effects of endotoxin. An explanation may be that endogenous prostaglandin release as a result of experimental endotoxemia can cause luteolysis and abortion in early pregnancy,5 and in late pregnancy may compromise uterine perfusion by initiating excessive uterine contractions.
Uterine torsions Uterine torsions occur in mid to late gestation and the primary clinical sign is abdominal pain. Mares with uterine torsions usually exhibit mild to moderate pain, and some will continue to pass feces despite discomfort.6, 7 Diagnosis of uterine torsion is made by rectal palpation. On rectal palpation, affected mares will have a tight broad ligament that runs vertically on the side that the uterus is twisted towards, and a tight broad ligament dorsal to the uterus on the side from which the uterus is twisted away. Vaginal palpation is of minimal diagnostic value since the cervix and vagina are rarely involved in uterine torsion in mares.
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When the diagnosis of uterine torsion is made, a high-risk pregnancy is identified. Survival rates for mares (97%) and foals (72%) are better if the torsion occurs at less than 320 days of gestation, and worse (mares 65%, foals 32%) when the torsion occurs at 320 or more days of gestation.7 Torsions greater than 180◦ increase the risk of compromised utero-placental blood flow and subsequently fetal oxygenation. If torsion occurs at term, it is usually associated with dystocia. Uterine rupture with subsequent septic peritonitis is the leading cause of maternal fatality.8
Lameness Laminitis and other lamenesses are fairly common in broodmares, and the impact of the musculoskeletal problem on the pregnancy is thought to be relatively minimal. However, some indirect factors associated with the management and treatment of severe lameness may have an effect. The administration of medications is often implicated in the delivery of small or weak foals, but the effect of drugs is largely speculative. Less tangible factors such as poor appetite, a lack of exercise and social herd interaction, and the stress of disability probably contribute to poor fetal outcomes.
Abdominal wall injuries Disruptions of the abdominal tunic most commonly occur in late pregnancy, and presumably are at least partially the result of the expanding uterine mass and volume. Tears occur in the prepubic tendon and the muscular portions of the abdomen9, 10 and are more common in older, multiparous mares. The condition is more prevalent in draft and heavier breeds and unfit horses, although prepubic tendon rupture and/or defects in the rectus abdominis musculature have been reported in lighter breeds including Arabians and ponies. Conditions that cause severe distention of the body wall such as hydrops, twinning, severe ventral edema or body wall trauma may result in rupture of the prepubic tendon or abdominal wall hernia. This is a serious injury to the mare that results in pain, difficulty moving, and warm, painful edema and swelling of the affected portion of the ventral abdomen. Severe cases are depressed, tachycardic, tachypneic, and off-feed. Affected mares frequently assume a ‘sawhorse’ stance and are reluctant to move. Signs of shock and colic may develop if peritonitis and/or internal hemorrhage develop. If injury to the prepubic tendon is severe, the iliosacral joint dorsiflexes, resulting in an elevated tail head and tuber ischii, and lordosis of the thoracolumbar spine. The ventral aspect of the abdomen extends more ven-
trally, the mammary gland flattens, and the teats tip caudally. There can be hemorrhage from the teats. The diagnosis of hernias and thinned areas of the abdominal wall can be appreciated by external and rectal palpation. Transabdominal ultrasound can confirm rents in the abdominal wall. Abdominal tunic injuries result in a high-risk pregnancy largely because of the risk to the mare. The abdominal disruption itself is painful, and further colic can result from viscera becoming entrapped in the abdominal wall hernia with the development of intra-abdominal adhesions and peritonitis. Direct risk to the fetus is moderate, and can be evaluated by ultrasonic examination of the fetus and placenta. All mares with abdominal wall tears should be evaluated for hydrops of the fetal membranes, a condition that worsens the fetal prognosis. A small retrospective study reported a 75% short-term survival rate for mares and about a 50% survival rate for foals.10 This is impressive, and is superior to the authors’ experience. Successful treatment is directly related to the amount of abdominal wall damage and internal abdominal trauma; when damage is extensive the risk for both mare and foal is significant.
Miscellaneous infectious agents Several infectious agents have been shown to cause equine abortion and include equine herpesvirus 1 (EHV-1), leptospirosis,1, 11 and equine viral arteritis (EVA). Selected causes of infectious equine abortion are listed in Table 1.5. Equine herpesvirus 1 (EHV-1) is capable of causing abortion during the last 4 months of pregnancy, as well as respiratory disease and meningoencephalomyelitis. EHV-1 produces a latent infection in at least 80% of the horses it infects. Latency is established in circulating lymphocytes and the trigeminal ganglion. Broodmares may become infected from new arrivals shedding virus in their respiratory secretions, or from stress-induced reactivation of their own latent herpesvirus within the blood vessels of the uterus or placenta, resulting in endothelial cell infection and thrombo-ischemia in those organs.12–14 This cell-associated viremia has an affinity for endothelial cells and has been shown to cause abortion in one of two ways: (i) infection of the placental vasculature, resulting in ischemic insult and placental insufficiency, premature placental detachment, and abortion of a virus-negative fetus; and (ii) infection of the fetus resulting in fetal demise and abortion of a virus-positive fetus. Mares affected by EHV1 can abort during late pregnancy or deliver weak, compromised foals prematurely or at term.15 Stress from shipping or overcrowding is considered to be a contributory factor to the occurrence of abortion.
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Table 1.5 Selected infectious causes of equine abortions Infectious agent
Clinical signs
Diagnostics
EHV-1
No premonitory signs; abortion during last trimester; aborted fetus is fresh with minimal autolysis; hydrothorax, hydroperitoneum; multifocal hepatic necrosis Mild disease in mares; uveitis; abortion follows mild illness by 1–3 weeks; abortions usually occur ≥ 6 months of gestation
Fetal tissues (liver, lung, thymus): histopathological findings of intranuclear inclusions; fluorescent antibody test (FAT) positive; virus isolation
Leptospirosis
Potomac horse fever
Mare experiences varying degrees of illness due to PHF (fever, colitis, laminitis) 2–4 months prior to abortion
Nocardioform
Sporadic abortions usually during late gestation (8–11 months); premature lactation; gross changes in portions of placenta occupying base of uterine horns or cranial uterine body Rare outbreaks of respiratory disease with subsequent abortion; abortion can occur with no visible illness in mare
Equine viral arteritis
Aborting mares often do not show any premonitory signs, although the EHV-1 strain responsible for abortion is also implicated in herpesviral myelitis outbreaks. The fetus and placenta are usually grossly normal. High levels of virus can be found in fetal tissues and placenta, and infected mares can shed EHV1 virus in their reproductive tract secretions for as long as 3 weeks following abortion. Confirmation of EHV-1 as the abortigenic agent often is by demonstration of the virus by culture or polymerase chain reaction (PCR) and immunohistochemical identification of intranuclear inclusion bodies in fetal and placental tissues. Leptospirosis can be a significant cause of equine abortion, with recent studies claiming 1.5–5.9% of equine abortions are related to leptospirosis.11, 16 Although multiple serovars have been associated with abortion, among the most common serovar is Pomona type kennewicki.17 Horses are considered incidental hosts for most serovars of Leptospira, but may act as maintenance hosts for L. interrogans serovar Bratislava.18–22 Leptospiremia, resulting in vasculitis, precedes infection of the reproductive organs, and in pregnant mares can cause fetal resorption, abortion, stillbirth or full-term delivery of weak neonates. Leptospiral abortions typically occur in middle to late gestation. Placentas are frequently thickened, edematous, and hemorrhagic with the chorionic surface appearing brown and mucoid.16 Aborting mares may not show any premonitory signs of disease and yet
Leptospires in fetal fluid or blood; dark-field or phase-contrast microscopy; FAT; PCR; MAT; paired sera titers; L. pomona kennewicki is the most prominent pathogenic serovar in horses Fetuses have increased volume of feces within small and large intestines; liver discoloration; histopathological findings: lymphohistiocytic enterocolitis, myocarditis, periportal hepatitis; N. risticii recovered by cell culture from fetal bone marrow, spleen, colon, liver; PCR Culture placenta, fetus and uterine discharge; bacteria grow slowly on blood agar
Fetus often autolytic; necrosis in media of small arteries; fetal tissues – FAT, virus isolation; rising VN titers in mares obtained 1–2 weeks apart
still mount a high antibody titer. When present, systemic signs in the dam can include fever, anorexia, listlessness, icterus and laboratory evidence of renal disease.11 Uveitis may develop a variable period of time post-exposure. Most aborting mares have titers to multiple leptospiral serovars and these titers do not necessarily correspond with the cause of the abortion.23 Many mares will have titers indicating exposure to these infectious agents and not abort, but rising titers or extremely high titers (≥ 1 : 100 000) to L. interrogans18 indicates infection. Confirmation of the etiologic agent is made on examination of aborted fetal tissues, and in the case of Leptospira, can be made by detecting the agent in mare urine. Culture, dark-field microscopy, immunohistochemical staining, PCR, and fluorescent antibody testing (FAT) are some of the techniques used to identify Leptospira in fetal tissues and fluids.11 Equine viral arteritis (EVA), another potentially abortigenic virus, is transmitted between horses primarily through either respiratory or venereal routes.24 Abortion, when it occurs, is usually not preceded by any premonitory signs in the broodmare and may occur any time from 3 months to over 10 months of gestation.24, 25 Clinical signs, when they occur, include pyrexia, depression, anorexia, conjunctivitis and rhinitis, periorbital edema, edema of the ventrum, mammary gland and hindlimbs, and urticaria.24, 25 Diagnosis is based on virus isolation, indirect immunohistochemistry, and PCR performed on
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fetal and placental tissues or nasopharyngeal swabs from the affected dam.24, 26 Another pathogen that may not be thought of as an abortigenic agent is Neorickettsia risticii, the causative agent of Potomac horse fever (PHF). PHF can cause colitis and laminitis in horses, and occasionally causes abortion in mares around 7 months of gestation.27 In experimentally infected mares, abortion occurred 65–111 days after inoculation,28 which might allow this pathogen to be forgotten as a possible cause of pregnancy loss by the time a mare aborts following initial infection. Transplacental transmission of N. risticii has been reported in natural infections, and the organism can induce fetal resorption, abortion, or cause the delivery of weak foals.29 Any pregnancies exposed (airspace for the viruses, water, feed and urine for leptospirosis) to horses affected by these conditions should be considered at high risk. Risk to the mare infected with these agents is variable. Mares affected with EHV-1 and EVA experience mild risk, but systemic effects of leptospirosis raise the risk to moderate levels due to possibility of renal insult.
Pelvic abnormalities Mares that have had pelvic injuries that result in a diminution of the pelvic inlet can have serious difficulty in delivery. The diagnosis of the condition is made during rectal palpation, and risk is assessed by the amount of encroachment on the pelvic inlet. Dystocia would result in severe risk to the foal and moderate to severe risk to the mare.
Endometrial insufficiency A poor-quality endometrium can result in a high-risk pregnancy because of poor delivery of nutrients and disposal of fetal waste. This can result from uterine scarring, endometrial atrophy, or surgical removal of portions of the uterus. Foals gestating in this environment can be poorly nourished, and grow poorly, and often have prolonged gestations. In severe cases, abortion can result if a fetus outgrows its nutrition. Premonitory signs are most commonly a previous history of small foals and prolonged gestation, but could include treatment of uterine infection and severe dystocia. There is little risk to the mare, but fetal risk with a poor-quality endometrium is moderate to severe.
Miscellaneous The most serious insults to the fetus are those that interfere with perfusion of the placenta or the delivery of nutrients and oxygen and removal of waste prod-
ucts. A healthy placenta is critical to delivery of a healthy foal.30 Conditions that result in hypoxemia, hypovolemia, hypotension or toxemia (including endotoxemia) can all have a negative effect on the fetus. Examples of uncommon conditions that would affect the pregnancy include heart failure, severe chronic obstructive pulmonary disease, upper respiratory obstructions, severe hemorrhage, and renal failure. Diagnosis of these conditions in a pregnant mare should increase scrutiny of the fetus. Risk to both mare and foal will depend on the severity of the underlying disease.
Placental conditions
Placentitis Infection of the placenta is a common condition, and can be caused by bacteria, viruses or fungal elements.1 Bacteria are the most common cause of placentitis with Streptococcus equi subspecies zooepidemicus, Escherichia coli, Klebsiella spp, Pseudomonas spp, and Staphylococcus aureus most often implicated in disease. Mares rarely show clinical or laboratory evidence of systemic infection. The condition is usually well-established before diagnosis, and can be clinically silent until abortion. More frequently, mares will show premature mammary gland development, lactation, and vulvar discharge.31–33 Most infections are ascending,34 although hematogenous infection can occur. Mares are typically affected during the last trimester of pregnancy. Infecting pathogens disrupt the contact between allantochorion and endometrium, resulting in placental separation that begins at the cervical star and dissects cranially along the body of the uterus. A speculum examination of the caudal reproductive tract will often reveal loss of the cervical plug and uterine discharge, which will sometimes be apparent at the vulva. Transrectal or transabdominal ultrasound of the uterus and conceptus will often reveal thickening of the caudal segment of the placenta and uterus, separation of the placenta from the endometrium, and pus between uterus and placenta.35, 36 Gram stains and culture of uterine discharge can establish the infectious agent. Placentitis that involves more cranial segments of the placenta can be difficult to image. Several fungal elements classified as nocardioform invade the base of the gravid horn,37 and can be difficult to detect. The limitation of transabdominal ultrasound is that not all of the placenta can be imaged. In these cases, placentitis is suspected, based on the presence of premature lactation. In mares with precocious mammary development, it is important to exclude the presence of twins, using ultrasound. Twin pregnancies will often present similarly, but require different risk
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tation alone. Once human error and miscalculation of the foaling date and/or inaccurate breeding date have been ruled out, other causes of prolonged gestation length include fescue toxicosis, delayed embryonic development, severe maternal malnutrition, and other ill-defined hormonal imbalances including a syndrome described in Western Canada associated with congenital hypothyroidism and dysmaturity. Duration of gestation is partly controlled by a number of factors. Colts tend to have longer gestation lengths than fillies. Mares due to foal earlier in the year may have longer gestation lengths due to the shorter photoperiod. Older mares tend to have longer gestations.
Fescue toxicosis
Figure 1.1 Speculum examination of a mare with a hemorrhagic vaginal discharge associated with diffuse premature placental separation.
assessment and therapy. There is little risk to the mare affected with placentitis, and fetal risk is difficult to estimate. Anecdotal evidence suggests that approximately 75% of pregnancies end with delivery of a live foal.38, 39 It is the authors’ experience that better outcomes occur when clinical signs occur later in gestation and when there is a quick reversal of mammary development in response to appropriate therapy.
Premature placental separation Occasionally, premature placental separation can occur without evidence of placental infection. Clinical signs are similar to placentitis, and in addition, occasionally uterine hemorrhage can be seen (see Figure 1.1). The diagnosis is made on clinical signs and ultrasonic examination of the uterus. Risk to the mare is low, and fetal risk is related to the amount of placenta affected.
Prolonged gestation The mare’s reported range of ‘normal’ gestation lengths is quite elastic, with some references describing normal pregnancies lasting for up to 374 days or longer. Therefore it is difficult to define what constitutes a post-term pregnancy by days of ges-
Fescue toxicosis, an intriguing model of maternal and fetal disease, is the result of mares eating fescue grass infested with a fungus (Acremonium coenophialum) that elaborates a mycotoxin that interacts with dopamine receptors. Dopaminergic stimulation results in decreased prolactin secretion. Prolactin plays an active role in the endocrine regulation of steroidogenesis and lactogenesis.40 The initiation of parturition in the mare involves proper maturation and function of the fetal hypothalamic–pituitary–adrenal (HPA) axis.41 Prolonged gestation length associated with fescue toxicosis may be due to hypoprolactinemia-induced changes in utero-feto-placental steroid metabolism or may be due to inhibition of D2 -dopamine receptors on corticotrophs in the fetal anterior pituitary gland.40, 41 In the mare, fescue toxicosis is characterized by an unreadiness-for-parturition leading to prolonged gestation, abortion, dystocia and agalactia accompanied by placental abnormalities that include premature placental separation, thickening and edema. 42 In the foal an unreadiness-for-birth is associated with an increased incidence of sepsis, peripartum asphyxia syndrome, failure of passive transfer, dysmaturity, and birth of weak foals with failure ‘to adapt’ following delivery.42–45 Late-term mares grazing endophyte-infected tall fescue demonstrate decreased concentrations of prolactin and relaxin43,45–47 and fail to exhibit the normal late gestational surge in progestins.43, 46, 48 Some studies report increased levels of estrogens and decreased levels of thyroxine (T4 )50 in exposed mares. Foals exposed to ergopeptine alkaloids during late gestation have decreased plasma levels of immunoassayable progestins, cortisol, T4 , and tri-iodothyronine (T3 ).43, 51 Mares should not be exposed to fescue infected with the endophyte at all, but at a minimum should be removed from infected pastures for the last 30 to
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60 days of gestation. Mares exhibiting toxicity are at great risk from dystocia, as is the fetus. Identification of the condition is made by determining the grass in pasture.
Umbilical cord abnormalities Approximately 3% of aborted and foals born dead at term are associated with abnormalities of the umbilical cord that include edema, torsion, sacculations, strangulations, and over-long cords.1 These disorders are virtually impossible to predict before delivery, and risk is assessed in retrospect. Average cord length in Thoroughbreds is between 36 and 83 cm. Unusually long cords (>80 cm) have been associated with hypoperfusion of the allantochorion and may be prone to cord torsion, which may contribute to fetal strangulation, hypoxia, death, and abortion. Excessively short cords place increased traction on the placenta during delivery and may predispose to premature placental separation. Not all abnormalities cause fetal compromise, but should increase scrutiny of an affected live foal so that any necessary intervention can occur without delay.
Hydrops of the fetal membranes Hydrops refers to the overproduction of fluid into either the amniotic or allantoic cavities (hydrallantois or hydramnion). Both conditions are rare, but hydrallantois is the more common of the two. Clinical signs associated with hydrops typically develop in the last trimester after an otherwise uneventful pregnancy. Although the pathophysiology of this condition is poorly understood, there are multiple causes that can result in either increased production or decreased clearance of fetal fluids, and include fetal defects, placental disease, and umbilical cord abnormalities. The increased volume of fetal fluids results in maternal anorexia, tachycardia and tachypnea, dyspnea, depression, colic, decreased manure production, difficulty walking, and marked ventral edema. Advanced cases may develop ventral abdominal hernia or prepubic tendon rupture. Suspicion of the condition is raised when there is a dramatic increase in abdominal distention in a later-term mare, that results in the mare having difficulty moving, eating or breathing.31 Diagnosis is made by rectal palpation of the uterus, which will be grossly fluid-filled, and the fetus will be difficult to palpate. Transabdominal ultrasound of a mare affected by placental hydrops will also reveal a massively fluid-filled uterus. Hydrops when severe can result in damage to the maternal abdominal tunic, causing uterine rupture, abortion, and retained placenta. At delivery, the expulsion of fluid
can cause hypovolemic shock. For most patients, severe risk would be assigned to both mare and foal, although two reports of hydrops record live foal deliveries (one long-term survival) and mare survival after treatment of hypovolemic shock after delivery.52, 53 Clinical signs of hydrops allantois include an enormous increase in abdominal size over 10 to 14 days due to rapid accumulation of allantoic fluid. Mares with hydramnion develop less dramatic abdominal distention over a more prolonged time course of weeks to months. Palpation per rectum reveals a large, fluiddistended uterus. It is often difficult or impossible to feel the fetus. The condition is confirmed via transabdominal ultrasonography. A large volume of clear fetal fluids is observed. Hydrops allantois tends to be more life-threatening for the mare due to its rapid onset and the excessive volume of fluids that accumulate (sometimes in excess of 100 L). The fetus may be normal in some of these pregnancies, but fetal survival depends on how late in the pregnancy the condition develops and how successfully the mare is supported during labor. Subsequent pregnancies can be normal.
Placental insufficiency Placental insufficiency is a poorly defined condition and is often a histological diagnosis of exclusion. The result of placental insufficiency is inadequate fetal nutrition, and the cause is believed to be a compromised endometrium that does not allow attachment of a robust placenta. The inability to attach to healthy endometrium results in a small and poorly developed (avillous) placenta. Fetuses can be aborted due to placental insufficiency, but are also born small, underweight even for their small size, and dysmature (fine hair coats, weak musculoskeletal systems, immature cardiovascular and gastrointestinal systems). This diagnosis is often made in retrospect, although historical information can be useful, as mares tend to deliver dysmature foals repeatedly. Risk to the mare from placental insufficiency is minimal, but is moderate to severe for the foal. Fetal conditions
Neonatal isoerythrolysis Hemolytic anemia occurs in horse and mule foals that absorb maternal immunoglobulins in colostrum that contains antibodies that bind to the foal’s erthrocytes.54 Mares at greatest risk for producing foals affected by neonatal isoerythrolysis (NI) lack certain red blood cell antigens. Aa and Qa are most commonly implicated alloantigens, but others are also
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Twins Twinning is a common condition in the Thoroughbred mare, and also occurs in other light horse breeds. Twinning results in high-risk pregnancy because the fetuses are often aborted and because, if carried to term, dystocia can occur and result in maternal reproductive tract trauma and even death. The mortality rate for twins is almost 90%.56 Most twins are diagnosed early in pregnancy and either one or both vesicles are eliminated by veterinarians or naturally by the mare. This early pregnancy loss causes no risk to the mare. Twin pregnancies become a greater issue once past the stage of possible natural reduction (about 40 days of gestation),57 although natural reductions at a later stage of gestation can occur. The diagnosis of twin pregnancy can be made by detecting a large uterus by rectal palpation, and is confirmed by transrectal or transabdominal ultrasound. At the later stages of gestation, diagnosis can be challenging, because the hearts of both fetuses cannot be obtained in one image, and care must be taken to assure that there truly are two fetuses present. If not aborted when small, twin pregnancies pose a moderate risk to the mare and a severe risk to both foals. The risk is due to the possibility of preterm birth and dystocia.
Fetal malformations, arthrogryposis, and fetal malpresentations Fetal abnormalities such as schistosomas reflexus, arthrogryposis, and hydrocephalus cause high-risk pregnancy due to risk to the foal because these conditions are inconsistent with life, and to the mare because they cause dystocia. These conditions are almost never diagnosed before delivery. Possibly these conditions could be diagnosed antepartum by transabdominal ultrasound, but that level of scrutiny, routinely given to human fetuses, is rarely afforded to
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the horse. These conditions are diagnosed during the second stage of parturition when they result in dystocia.
Fetal malpresentations Malpositioning of the fetus causes high-risk pregnancy to the fetus because of the potential for asphyxia due to a prolonged stage 2 labor, and the potential for damage to the mare’s reproductive tract.58 Most malpositionings occur during delivery, so could not be diagnosed before delivery began. An exception is a breech presentation, which is rare, but can be diagnosed via ultrasound in late pregnancy, by observing a caudal presentation after the time a fetus can no longer turn in the uterus. Malpresentations are diagnosed by direct fetal palpation through an open cervix, although the entire fetus cannot always be palpated. Some malpresentations such as ‘dog-sitter’s’ (dorso-sacral presentation with flexed hindlimbs that cause impingement of the hind fetlocks in or on the pelvis) only become apparent as the fetus is in the process of delivery.
Mare reproductive loss syndrome During the spring of 2001, veterinarians reported an outbreak of early fetal losses at about 77 days of gestation (approximately 25% of mares),59 later-term abortions, and birth of weak foals in central Kentucky. The cause of the syndrome was obscure, and is still not totally elucidated, but a risk factor was access to pasture.59 Pathological examination of aborted fetuses was not pathognomonic and revealed a range of findings, most commonly pulmonary lesions, placentitis, and funisitis (umbilical inflammation).60 Actinobacillus and Streptococcus could commonly be cultured from tissues. The syndrome could be reproduced by feeding the Eastern tent caterpillar (ETC) to pregnant mares between 40 and 80 days of gestation, 61 but the specific toxic principle has not been elucidated. A recent review of the syndrome supports the hypothesis that, following ingestion of the ETC, the setae (bristles on the skin) inbed into the submucosa of the horse’s alimentary tract and create microgranulomtous lesions that permit translocation of gut bacteria into the bloodstream.62 The bacteria (most commonly incriminated are Streptococcus, Actinobacillus and Enterococcus spp) establish infections in distant locations, including the fetus and placenta, where the immune surveillance of the mare is reduced.62 Identification of the syndrome can be made immediately before abortion by observing hyperechoic allantoic fluid (J.P. Morehead, personal communication, 2001). The strongest risk factor is access to the Eastern tent caterpillar,63 which is not confined to central
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Kentucky. Risk to the mare is minimal, but risk to the fetus is severe.
References 1. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith MS, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases ( 1986–1991). J Am Vet Med Assoc 1993;203(8):1170–5. 2. Hong CB, Donahue JM, Giles RC, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Equine abortion and stillbirth in central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5(4):560–6. 3. Santschi EM, Slone DE, Gronwall R, Juzwiak JS, Moll HD. Types of colic and frequency of postcolic abortion in pregnant mares: 105 cases (1984–1988). J Am Vet Med Assoc 1991;199(3):374–7. 4. Boening KJ, Leendertse. Review of 115 cases of colic in the pregnant mare. Equine Vet J 1993;25(6):518–21. 5. Kindahl H, Daels P, Odensvik K, Daunt D, Fredricksson G, Stabenfeldt G, Hughes JP. Experimental models of endotoxemia related to abortion in the mare. J Reprod Fertil Suppl 1991;44;509–16. 6. Pascoe JR, Meagher DM, Wheat JD. Surgical management of uterine torsion in the mare: a review of 26 cases. J Am Vet Med Assoc 1981;179(4):351–4. 7. Chaney KP, Holcombe SJ, LeBlanc MM, Hauptman JG, Embertson RM, Mueller PO, Beard WL. The effect of uterine torsion on mare and foal survival: a retrospective study, 1985–2005. Equine Vet J 2007;39(1):33–6. 8. Steele CM, Gibson KT. Colic in the pregnant and periparturient mare. Equine Vet Educ 2001;13(2):94–104. 9. Hanson RR, Todhunter RJ. Herniation of the abdominal wall in pregnant mares. J Am Vet Med Assoc 1986;189(7):790–3. 10. Ross J, Palmer JE, Wilkins PA. Body wall tears during late pregnancy in mares: 13 cases (1995–2006). J Am Vet Med Assoc 2008;232(2):257–61. 11. Loynachan AT, Slovis NM. Leptospirosis: Fundamental principles of disease. Proceedings of ACVIM Forum and Canadian Veterinary Medical Convention, 2009; pp. 124–5. 12. Smith KC, Borchers K. A study of the pathogenesis of equid herpesvirus-1 (EHV-1) abortion by DNA in-situ hybridization. J Comp Pathol 2001;125(4):304–10. 13. Smith KC, McGladdery AJ, Binns MM, Mumford JA. Use of transabdominal ultrasound-guided amniocentesis for detection of equid herpesvirus 1-induced fetal infection in utero. Am J Vet Res 1997;58(9):997–1002. 14. Smith KC, Mumford JA, Lakhani K. A comparison of equid herpesvirus-1 (EHV-1) vascular lesions in the early versus late pregnant equine uterus. J Comp Pathol 1996;114(3):231–47. 15. Allen GP. Epidemic disease caused by Equine Herpesvirus1: recommendations for prevention and control. Equine Vet Educ 2002; pp. 177–84. 16. Hines MT. Leptospirosis. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 301–9. 17. Donahue JM, Smith BJ, Poonacha KB, Donahoe JK, Rigsby CL. Prevalence and serovars of leptospira involved in equine abortions in central Kentucky during the 1991–1993 foaling seasons. J Vet Diagn Invest 1995;7:87–91.
18. Bernard WV, Bolin C, Riddle T, Durando M, Smith BJ, Tramontin RR. Leptospiral abortion and leptospiruria in horses from the same farm. J Am Vet Med Assoc 1993;202:1285–6. 19. Ellis WA, O’Brien JJ, Cassells JA, Montgomery J. Leptospiral infection in horses in Northern Ireland: serological and microbiological findings. Equine Vet J 1983;12:317–20. 20. Kiston-Piggot AW, Prescott JF. Leptospirosis infection in horses in Ontario. Can J Vet Res 1987;51:448–51. 21. Van Den Ingh TS, Hartman EG, Bercovich Z. Clinical Leptospira interrogans serogroup Australis serovar Lora infection in a stud farm in The Netherlands. Vet Q 1989;11:175– 82. 22. Williams DM, Smith BJ, Donahue JM, Poonacha KB. Serological and microbiological findings on 3 farms with equine leptospiral abortions. Equine Vet J 1994;26:105–8. 23. Poonacha KB, Donahue JM, Giles RC, Hong CB, PetritesMurphy MB, Smith BJ, Swerczek TW, Tramonton RR, Tuttle PA. Leptospirosis in equine fetuses, stillborn foals, and placentas. Vet Pathol 1993;30:362–9. 24. Timoney PJ, McCollum WH. Equine viral arteritis. Vet Clin North Am Equine Pract 1993;9:295–309. 25. Doll ER, Knappenberger RE, Bryans JT. An outbreak of abortion caused by the equine arteritis virus. Cornell Vet 1957;47:69–75. 26. Chirnside ED, Spaan WJ: Reverse transcription and cDNA amplification by the polymerase chain reaction of equine arteritis virus (EAV). J Virol Methods 1990;30:133–40. 27. Long MT, Goetz TE, Whiteley HE, Lock TE, Holland CJ, Foreman JH, Baker GJ. Identification of Ehrlichia risticii as the causative agent of two equine abortions following natural maternal infection. J Vet Diagn Invest 1995;7:201–5. 28. Long MT, Goetz TE, Kakoma I, Lock TE. Evaluation of fetal infection and abortion in pregnant ponies experimentally infected with Ehrlichia risticii. Am J Vet Res 1995;56:1307–16. 29. Pusterla N, Madigan JE. Neorickettsia risticii. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 357–62. 30. Cottrill CM, Jeffers-Lo J, Ousey JC, McGladdery AJ, Ricketts SW, Silver M, Rossdale PD. The placenta as a determinant of fetal well-being in normal and abnormal equine pregnancies. J Reprod Fertil Suppl 1991;44:591–601. 31. Santschi EM, LeBlanc MM. Fetal and placental conditions that cause high-risk pregnancy in mares. Comp Cont Educ Pract Vet 1995;17(5):710–21. 32. Macpherson ML. Identification and management of the high-risk pregnant mare. Proc Am Assoc of Equine Pract, 2007; pp. 293–304. 33. LeBlanc MM, Macpherson M, Sheerin P. Ascending placentitis: What we know about pathophysiology, diagnosis and treatment. Proc Am Assoc Equine Pract 2004;50:127–43. 34. Platt H. Infection of the horse fetus. J Reprod Fertil Suppl 1975;23:605–10. 35. Renaudin CD, Troedsson MHT, Gillis CL, King VL, Bodena A. Ultrasonographic evaluation of the equine placenta by transrectal and transabdominal approach in the normal pregnant mare. Theriogenology 1997;47(2):559–73. 36. Morris S, Kelleman AA, Stawicki RJ, Hansen, PJ, Sheerin PC, Sheerin BR, Paccamonti DL, LeBlanc MM. Transrectal ultrasonography and plasma progestin profiles identifies feto-placental compromise in mares with experimentally induced placentitis. Theriogenology 2007;67(4): 681–91. 37. Christensen BW, Roberts JF, Pozor MA, Giguerre S, Sells SF, Donahue JM. Nocardioform placentitis with isolation
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38. 39.
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of Amycolatopsis spp in a Florida-bred mare. J Am Vet Med Assoc 2006;228(8):1234–9. Macpherson ML. Treatment strategies for mares with placentitis. Theriogenology 2005;64:528–34. Macpherson ML, Bailey CS. A clinical approach to managing the mare with placentitis. Theriogenology 2008; 70(3):435–40. Redmond LM, Cross DL, Strickland JR and Kennedy SW. Efficacy of domperidone and sulpiride as treatments for fescue toxicosis in horses. Am J Vet Res 1994;55(5):722–9. Brendemeuhl JP, Williams MA, Boosinger TR, Ruffin DC. Plasma progestagens, triiodothyronine and cortisol concentrations in post date gestation foals exposed in utero to tall fescue endophyte Acremonium coenophialum. Biol Reprod Mono 1995;1:53–9. Putnam MR, Bransby DI, Schumacher J, Boosinger TR, Bush L, Shelby RA, Vaughn JT, Ball D, Brendemeuhl JP. Effects of the fungal endophyte Acremonium coenophialum in fescue on pregnant mares and foal viability. Am J Vet Res 1991;52(12):2071–4. Brendemeuhl JP. Reproductive aspects of fescue toxicosis. In: Robinson NE (ed.) Current Therapy in Equine Medicine 4. Philadelphia: Saunders, 1997; pp. 571–3. Evans TJ, Rottinghaus GE, Casteel SW. Ergopeptine alkaloid toxicoses in horses. In: Robinson NE (ed.) Current Therapy in Equine Medicine 5. Philadelphia: Saunders, 2003; pp. 796–8. Green EM, Raisbeck MF. Fescue toxicosis. In: Robinson NE (ed.) Current Therapy in Equine Medicine 4. Philadelphia: Saunders, 1997; pp. 670–3. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995;73(3):899– 908. Ryan PL, Bennet-Wimbush K, Vaala WE, Bagnell CA. Systemic relaxin in pregnant pony mares grazed on endophyte-infected fescue: effects of fluphenazine treatment. Theriogenology 2001;56:471–83. Evans TJ, Constantinescu GM, Ganjam VK. Clinical reproductive anatomy and physiology of the mare. In: Youngquist RS (ed.) Current Therapy in Large Animal Theriogenology. Philadelphia: Saunders, 1997; pp. 43–70. Brendemeuhl JP. Effects of Acremonium coenophialuminfected tall fescue on progestagen production in pregnant mares. In Proceedings of Society of Theriogenology, 1992; pp. 108–12. Messer NT, Riddle T, Traub-Dargatz JL, Dargatz DA, Refsal KJ, Thompson DL. Thyroid hormone levels in Thoroughbred mares and their foals at parturition. Proc Am Assoc Equine Pract 1998;44:248–51.
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51. Boosinger TR, Brendemeuhl JP, Bransby DL, Wright JC, Kemppainen RJ, Kee DD. Prolonged gestation, decreased tri-iodothyronine concentration and thyroid gland histomorphologic features in newborn foals of mares grazing Acremonium coenophialum infected fescue. Am J Vet Res 1995;56:66–9. 52. Smith JM. Hydroallantois in the mare. Mod Equine Med 1983;2:7–12. 53. Christensen BW, Troedsson MHT, Murchie TA, Pozor MA, Macpherson ML, Estrada AH, Carrillo NA, Mackay RJ, Roberts GD, Langlois J. Management of hydrops amnion in a mare resulting in birth of a live foal. J Am Vet Med Assoc 2006;228(8):1228–33. 54. Boyle AG, Magdesian KG, Ruby RE. Neonatal isoerythrolysis in horse foals and a mule foal: 18 cases (1988–2003). J Am Vet Med Assoc 2005;227(8):1276–83. 55. MacLeay JM. Neonatal isoerythrolysis involving the Qc and Db antigens in a foal. J Am Vet Med Assoc 2001;219:79–81. 56. Pascoe R. Methods for the treatment of twin pregnancy in the mare. Equine Vet J 1983;15:40–2. 57. Ginther OJ. The nature of embryo reduction in mares with twin conceptuses: Deprivation hypothesis. J Am Vet Res 1989;50(1):45–53. 58. Lu KG, Barr BS, Embertson R, Dallap-Schaer B. Dystocia: a true emergency. Clin Tech Equine Pract 2006;5(2):145–53. 59. Morehead JP, Blanchard TL, Thompson JA, Brinsko SP. Evaluation of early fetal losses on four equine farms in central Kentucky: 73 cases (2001). J Am Vet Med Assoc 2002;220(12):1828–830. 60. Williams NM, Bolin DC, Donahue JM, Giles RC, Harrison LR, Hong CB, Poonacha KB, Roberts JF, Sebastian MM, Smith BJ, Smith RA, Swerczek TW, Tramontin RR, Vickers ML. Gross and histopathological correlates of mare reproductive loss syndrome. Proceedings of 1st Workshop Mare Reproductive Loss Syndrome, 2002; pp. 24–5. 61. Bernard WV, LeBlanc MM, Webb BA, Stromberg AJ. Evaluation of early fetal loss induced by gavage with eastern tent caterpillars in pregnant mares. J Am Vet Med Assoc 2004;225(5):717–21. 62. McDowell KJ, Webb BA, Williams NM, Donahue JM, Newman KE, Lindemann MD, Horohov DW. Invited review: the role of caterpillars in mare reproductive loss syndrome: a model for environmental causes of abortion. J Anim Sci 2010;88(4):1379–87. 63. Webb BA, Barney WE, Dahlman DL, DeBorde SN, Weer C, Williams NM, Donahue JM, McDowell KJ. Eastern tent caterpillars (Malacosoma americanum) cause mare reproductive loss syndrome. J Insect Physiol 2004;50:185–93.
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Chapter 2
Monitoring the High-Risk Pregnancy Wendy E. Vaala
Introduction Considerable attention has been focused on how to improve conception rates in the mare and how to save the critically ill foal. Once the mare is confirmed pregnant and there is no evidence of twinning, veterinary attention frequently wanes until parturition. The three periparturient events that have the most devastating effect on foal survival are hypoxia, infection, and derangement of in utero development. Periparturient monitoring should focus on detection of these events to permit timely prepartum therapy for the mare and ensure attended delivery and early evaluation of the foal. Periparturient conditions associated with high-risk pregnancy and fetal/neonatal compromise are listed in Table 2.1. Most clinical trials designed to examine hormone changes in the at-risk mare, sonographic changes in the placenta, fetal fluids or fetal activity and heart rate, or to evaluate the efficacy of various treatment modalities on pregnancy outcome are underpowered statistically due to the small number of clinical cases with similar problems. Such studies often permit only anecdotal observations and conclusions to be inferred. Until larger multicentric, collaborative trials validate the usefulness of monitoring specific maternal hormones or fetal biophysical parameters, or the efficacy of various therapies designed to manage and maintain complicated pregnancies, it is the author’s recommendation that no single monitoring aid or therapy be relied upon exclusively. Any mare receiving supportive therapy to prolong an abnormal pregnancy should receive frequent assessment in case silent or undetected fetal demise occurs without ensuing abortion, or in the event parturition occurs without the customary warning signs. Prepartum diseases or disorders affecting the dam, utero-placental unit or fetus may result in a high-
risk pregnancy. Adverse events associated with delivery itself may also compromise an otherwise normal pregnancy. Examples of such conditions are listed in Table 2.1. Periparturient complications during late pregnancy may include severe maternal illness, lameness or colic, reproductive tract disease or fetal/placental anomalies. The challenge is early recognition of mares with jeopardized pregnancies at the earliest possible stage in gestation or in the disease process, to permit timely and effective intervention with the goal of improving maternal and neonatal morbidity and mortality. Candidates for more aggressive monitoring during pregnancy include mares with a history of past complications such as premature placental separation, placentitis, premature delivery, and post-term pregnancies, and mares experiencing a problem with their current pregnancy due to severe maternal illness, colic, surgery and/or reproductive tract disease (e.g., uterine torsion, uterine artery hemorrhage, abdominal wall rupture). Any prepartum or intrapartum event that jeopardizes utero-placental perfusion has the potential to disrupt delivery of oxygen and nutrients to the fetus. Certain conditions such as ascending placentitis have the added potential of introducing sepsis into the intrauterine environment. Any local or systemic maternal infection has the potential to activate proinflammatory mediators that may contribute to endogenous prostaglandininduced delivery. Table 2.2 lists specific adverse periparturient events and complications to anticipate.
Evaluation of feto-placental well-being: biochemical indices Maternal hormones The search continues for the hormone or group of hormones that most accurately predict feto-placental
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Monitoring the High-Risk Pregnancy Table 2.1 Periparturient conditions associated with high-risk equine pregnancya Maternal factors Severe systemic disease: colitis, hepatic disease, pneumonia/pleuritis Severe neurological disease: equine protozoal myelitis, cervical cord compression, equine herpesvirus myelitis Endocrinopathy: equine Cushing disease (pituitary pars intermedia dysfunction), fescue toxicosis Endotoxemia Abdominal surgery/general anesthesia Fever Anemia Severe hypoproteinemia Severe maternal malnutrition Premature udder development/vaginal discharge/precocious lactation Hydrops allantois/amnii Prepubic tendon rupture/rectus abdominus hernia Pelvic injuries Uterine torsion Severe endometrial fibrosis Premature labor Agalactia/poor-quality colostrum Prolonged gestation Placental factors Small placenta Placentitis: bacterial, fungal, nocardioform Premature placental separation Umbilical cord complications: umbilical cord torsion, excessively short cord associated with premature placental separation Vasculitis, edema Thrombosis, infarction Placental insufficiency Uterine artery hemorrhage Delivery factors Induced delivery Dystocia Cesarean section Premature placental separation Prolonged labor, uterine inertia Fetal/neonatal factors Congenital anomalies Chromosomal anomalies Cardiopulmonary disease Infection/sepsis Prematurity/dysmaturity Twinning Neonatal isoerythrolysis Failure of passive transfer a Adapted from: Vaala WE, Sertich PL. Perinatology. In: Higgins AJ, Snyder JR (eds) The Equine Manual, 2nd edn. London: Elsevier Saunders, 2006; pp. 765–868.
well-being and impending parturition. Progestagens and estrogens remain the most likely candidates, but the specific endocrine mediators responsible for maintenance of pregnancy during late gestation and stimulation of the final stages of fetal maturation and
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initiation of parturition remain the focus of considerable research and controversy. In many compromised equine pregnancies the prepartum changes of these hormones and their metabolites may be too acute to provide advanced warning of impending fetal demise. Non-lethal forms of fetal compromise associated with infection, hypoxia or disrupted patterns of fetal development may be even harder to detect using the hormonal assays currently available.
Progestagens The prepartum hormonal changes in the mare are different from those of other large animal species. In most other species progesterone (P4) is the dominant progestagen during pregnancy, and concentrations of P4 decline prior to spontaneous labor suggesting that P4 withdrawal is conducive to active labor. In the mare, P4 is produced by the ovaries until days 120–150 of gestation, at which time the feto-placental tissues begin to synthesize P4 from pregnenolone (P5) supplied by the fetal adrenal glands. After days 180–240 of pregnancy, P4 concentrations are negligible in both maternal and fetal circulations. Instead, P4 is metabolized into a number of other progestagens that include saturated pregnanes and unsaturated pregnenes.1, 2 In the mare, progestagens other than P4 remain relatively constant until 3 weeks prepartum, after which time progestagen levels rise gradually before parturition.1, 3, 4 In particular, the concentration of 5α-pregnane,3,20-dione (5α-DHP), a direct metabolite of P4, increases gradually during the last few weeks of gestation and then declines precipitously within days or hours of delivery.5, 6 Interestingly, in women, 5α-DHP rather than P4 undergoes a prepartum decline.7 Commercial radio-immunoassays (RIAs) and enzyme-linked immunosorbent assays (ELISAs) used to measure ‘progesterone’ concentrations in pregnant mares can be used to measure some of these other progesterone metabolites that predominate during late pregnancy. This is possible because the antibodies used in these assays cross-react with many of the other progestagens, particularly the α-pregnanes. Values may vary between laboratories due to the different levels of cross-reactivity between the different assay technologies.2, 8, 9 Using a validated ELISA, progestagen values reported for mares between 180 and 310 days of gestation ranged from 2 to 6 ng/mL.10 In healthy, late-term mares, progestagens are produced by the utero-placental tissues from the precursor, P5, supplied by the fetal adrenals.2, 11 Fetalderived P5 is then converted to P4 by utero-placental metabolism and secreted into the umbilical circulation or metabolized further to 5α-DHP in the
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Table 2.2 Adverse periparturient events and fetal/neonatal complications to anticipate Periparturient event
Potential fetal/neonatal complication
Precocious udder development, vulvar discharge, premature lactation
Placentitis, twinning; premature delivery/abortion, peripartum asphyxia
Excessive abdominal enlargement
Twinning, hydrops allantois/amnii, abdominal wall rupture; premature delivery/abortion, peripartum asphyxia
Prolonged gestation, agalactia, dystocia, premature placental separation, placental edema
Fescue toxicosis, maladapted foal with increased risk for FPT and endocrine abnormalities
Prolonged gestation, in utero growth retardation
Placental insufficiency, dysmature foal, chronic in utero asphyxia
Premature placental separation
Peripartum asphyxia, hypoxic-ischemic encephalopathy (HIE)
Dystocia
Hypoxic ischemic encephalopathy, birth trauma: scleral hemorrhage, fractured ribs, physeal damage, ruptured bladder, soft tissue injuries
Colitis in the mare
Ascending placentitis, fetal/neonatal sepsis, premature delivery
Aged broodmare
Placental insufficiency associated with chronic endometrial changes, low birthweight foal, poor-quality colostrum with increased risk of failure of passive transfer (FPT)
Colic in the mare
Utero-placental insufficiency associated with reduced perfusion due to abdominal distention, endotoxemia; premature delivery
Heavy (> 10% foal’s birthweight) and/or discolored placenta
Placentitis, premature placental separation, placental edema associated with endometrial changes; peripartum asphyxia, HIE, sepsis, neonatal dysmaturity
Uterine torsion
Varying degrees of placental vascular compromise; peripartum asphyxia
endometrium or other progestagens within the fetus or utero-placental tissues. At least 10 progestagens have been identified and measured in the late-term mare.1 Seven of these metabolites increase with advancing gestational age, while levels of P4 remain virtually undetectable. Little is known about the biological activity of most of these compounds, although 5α-DHP is known to inhibit myometrial contractility in women12 and binds more strongly than P4 to the uterine progesterone receptor in mares.13 During the last week of gestation, an unknown stimulus results in an increase in fetal adrenocorticotropic hormone (ACTH) and the appearance of the enzyme 17α-hydroxylase. This enzyme is responsible for the conversion of progesterone to cortisol.14 This prepartum surge in fetal cortisol is associated with the initiation of a critical cascade of endocrine events in the fetus that result in readiness for birth and an ability to adapt to extrauterine life. Cortisol stimulates an increase in the neutrophil : lymphocyte (N : L) ratio and a rise in thyroid hormones that, in turn, have a positive impact on skeletal muscle tone, body temperature regulation, and glucose homeostasis. Maturation of the fetal hypothalamic–pituitary–adrenal (PHA) axis is essential for triggering parturition and ensuring the final phase of fetal maturation. It is appealing to surmise that pregnancies jeopardized by placental anomalies or fetal compromise
will have altered concentrations of progestagens due to disrupted or altered metabolic pathways within the feto-placental unit. A number of trials have followed mares with either naturally occurring highrisk pregnancies or with experimentally induced placental disease, and have documented wide variations in maternal concentrations of plasma progestagens, including premature elevations and precipitous declines in hormone concentrations.1, 6, 10, 15, 16 Several patterns of progestagen activity during late pregnancy begin to emerge. The worst scenario is a pattern of rapidly declining progestagen concentrations with values approaching zero. In pregnancies complicated by acute fetal distress there was a dramatic decline in progestagen production, which re-emphasized the importance of a healthy fetoplacental unit for normal production and metabolism of progestagens.1 Most of these pregnancies were associated with fetal death and abortion. The second pattern is one of precociously elevated levels of nearly all progestagen metabolites that is often associated with bacterial placentitis, enhanced fetal adrenocortical activity, and promising neonatal survival.1, 6, 10 A few pregnancies compromised by placental anomalies other than bacterial placentitis, such as placental edema or villous atrophy, exhibited a slightly different pattern. Levels of certain progestagens such as P4 were elevated, but other metabolites including P5 were decreased, possibly reflecting
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Monitoring the High-Risk Pregnancy a lack of functional placenta.1 A third pattern of progestagen activity occurs when maternal progestagens fail to show the normal prepartum rise. Mares experiencing prolonged gestations due to ingestion of ergot alkaloids associated with fescue toxicosis or delayed parturition due to other causes may fall into this category.8, 17, 18
Estrogens There are two groups of estrogens in the mare: the common phenolic estrogens, estrone and estradiol 17β, and the ring unsaturated estrogens, equilin and equilenin, which are unique to the horse. The fetal gonads provide the precursors that are ultimately converted by the utero-placental tissues to estrogens. Plasma estrogen concentrations peak during months 7 and 8 of gestation, followed by a gradual decline thereafter, which parallels the rise and fall in the size of fetal gonads. Between 150 and 280 days of gestation, estrogen levels greater than 1000 ng/mL are considered normal and levels less than 1000 ng/mL are indicative of fetal stress.10 Estrogen levels below 500 ng/mL are associated with severely compromised or dead fetuses. The significance of elevated estrogen concentrations has not received much attention, although higher than normal levels of estradiol17β have been reported in late-term mares exposed to endophyte-infected tall fescue.19 Higher than normal concentrations of estradiol-17β may contribute to the agalactia observed in mares suffering from fescue toxicosis. The SI conversion table in the appendix of this book serves as a resource to convert and compare hormone values reported in different units by various laboratoriess. The role of estrogen in the late-term mare remains uncertain. One frequently referenced study evaluated the effect on pregnancy outcome of removing estrogen precursors by gonadectomizing horse fetuses in mid to late gestation.20 Mares experienced dramatic decreases in plasma estrogen concentrations, accompanied by prolonged, ineffective labor, decreased prostaglandin production, and the birth of dysmature foals.20 In other species, estrogen has an effect on uterine contractility and uterine blood flow.21 Proposed mechanisms of action for estrogen include promoting formation of uterine oxytocin receptors, increasing uterine gap junction formation, and alteration of uterine blood flow,22 which may help explain the weak labor in estrogen-deficient mares. Limited studies in late-term mares, stressed by medical or surgical disease or induced abortion, concluded that maternal serum concentrations of estrone sulfate (ES) were not a sensitive indicator of fetal compromise or death.15, 23 Maternal ES concentrations declined only after severe fetal stress or abortion had occurred.
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There are no controlled clinical trials (only anecdotal observations) examining the effects of estrogen supplementation in the late gestational mare. Therefore, no specific recommendations regarding the form of estrogen, the dosage or route of administration can be offered for its use in the late-term mare. Further discussion about the rationale behind estrogen supplementation during early pregnancy can be found in Chapter 254.
Relaxin In the horse, the peptide hormone relaxin is produced primarily by the placenta.24 Relaxin promotes uterine and cervical growth and remodeling that is essential for normal fetal growth and parturition. Relaxin decreases the collagen content in the extracellular matrices of the pubic symphysis and cervix, inhibits uterine contractility, and plays a role in mammary gland development. Relaxin has angiogenic and vasodilatory effects on endometrial and mammary tissues. This effect may be mediated through upregulation of vascular endothelial growth factor (VEGF). The maternal concentration of relaxin begins to increase after day 75 of gestation and peaks at day 175 and then remains elevated until foaling, followed by a precipitous decline postpartum.25, 26 Low maternal concentrations of relaxin during late gestation have been associated with high-risk pregnancies and complicated deliveries. Some of the periparturient problems associated with low relaxin levels have included premature placental separation, placentitis, hydrops, symptomatic maternal pituitary neoplasia, oligohydramnios, and fescue toxicosis.26–28 Although low relaxin concentrations have been associated with a variety of late-term uteroplacental complications, to date there has not been good correlation between a clinical response to therapy and an increase in relaxin levels.28 Currently there are no commercially available assays for equine relaxin. Perhaps with the recent production of recombinant equine prorelaxin28 there will be new research re-evaluating relaxin as a marker of utero-placental function in the mare.
Pre-foaling mammary secretion electrolytes Pre-foaling mammary secretion electrolytes are related to fetal readiness for birth in normal pregnancies.29 Calcium concentrations increase, accompanied by an inversion of sodium and potassium concentrations.30 Mature, term foals are usually produced from deliveries associated with the following mammary electrolyte profile: calcium ≥ 40 mg/dL, sodium ≤ 30 mEq/L, and potassium ≥ 35 mEq/L.29, 30
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Table 2.3 Scoring system for electrolyte concentrations in pre-foaling mammary secretions. From Ousey et al.29 Calcium mg/dL ≥40 ≥28 ≥20
Sodium
Potassium
mEq/L
SI
mg/dL
mEq/L
SI
mg/dL
mEq/L
SI
Points for each electrolyte in specified range
≥20 ≥14 ≥10
≥10 ≥7 ≥5
≤69 ≤115 ≤184
≤30 ≤50 ≤80
≤30 ≤50 ≤80
≥136.5 ≥117 ≥78
≥35 ≥30 ≥20
≥35 ≥30 ≥20
15 10 5
A total score ≥35 for all three electrolytes suggests fetal maturity and probability of a safe induction.
Mammary secretion electrolyte concentrations, once diluted, can be measured using commercial serum chemistry analyzers. Table 2.3 presents a scoring system for fetal maturity based on pre-foaling mammary secretion electrolyte concentrations. Stall-side test kits that measure calcium concentrations alone are commercially available (FoalWatchTM test kit, CHEMetrics Inc, Calverton, VA; Predict-aFoalTM , Animal Healthcare Products, Vernon, CA). Water hardness test strips can also be used. All of these tests are semi-quantitative and measure calcium and magnesium. Impending parturition is predicted based on the calcium concentrations in the milk. These assays are not foolproof. Mares with precocious udder development and lactation due to placentitis may have early, and misleading, elevation of mammary secretion calcium concentration.6 These tests are best utilized to help forestall premature induction of delivery rather than to guarantee fetal maturity.
Biophysical indices Transrectal ultrasonography During late gestation, transrectal linear ultrasonography is useful for evaluating placental integrity at the cervical star and fetal fluid clarity while providing a limited view of the fetus (e.g., fetal orbit, fetal activity). Transrectal ultrasonography of the caudal allantochorion provides a good image of the cervical star.31, 32 Since this often is the first region of the placenta to undergo changes associated with ascending placentitis and fescue toxicity, transrectal ultrasonography is an effective tool to detect placental thickening and/or premature placental separation associated with these conditions.31–33 The technique is described in detail in Chapter 4. Normal measurements for the combined thickness of the utero-placental junction (CTUP), obtained at a site just cranial and lateral to the cervical–placental junction, are presented in Table 2.4. CTUP measurements may be slighter higher in Warmbloods and lower in ponies.
Transrectal ultrasonography can also be used to identify other abnormal conditions including premature placental separation and hyperechoic changes in fetal fluids. Although palpation per rectum alone is often enough to confirm the presence of excessive volumes of fetal fluids, transrectal ultrsonography can be used to document excessive volumes of amniotic and/or allantoic fluids in the case of pregnancies affected by hydrops.
Transabdominal ultrasonography Transabdominal ultrasonography is an excellent tool for evaluating the fetus and placenta in late-term mares.34–38 Transabdominal ultrasonography can be used to evaluate a variety of parameters including fetal position, fetal activity and tone, fetal heart rate (range and variability), allantoic and amniotic fluid volume and clarity, and placental thickness and integrity. The technique is described in detail in several excellent references34, 37 and in Chapter 4. During late pregnancy, ultrasonographic warning signs of utero-placental disease and potential fetal compromise include persistent bradycardia (FHR < 50 bpm), sustained tachycardia (> 120 bpm) in the absence of fetal movement, loss of fetal movement and tone, increased or decreased volumes of allantoic or amniotic fluid (Figure 2.1), marked increases in fetal fluid echogenicity, large and/or increasing areas of placental detachment, and increased uteroplacental thickness (≥ 15 mm). The transabdominal approach is unlikely to detect the pericervical
Table 2.4 Normal values for combined thickness of the uterus and placenta (CTUP). From Renaudin et al.31, 32 Gestational day Day 271–300 Day 301–220 >Day 330
CTUP (mm) <8 <10 <12
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Figure 2.1 Ultrasonographic appearance of hydrops allantois in a late-term mare.
placental changes around the cervical star that are frequently observed in cases of ascending placentitis. However, this modality is useful for detecting placental thickening and/or separation in mares with placentitis originating from a hematogenous infection which results in a global placental insult, or those mares suffering from nocardioform placentitis that produces placental separation and accumulation of purulent debris at the base of the gravid horn and junction of the uterine body. Measurement of fetal eye orbit, femur length, aortic diameter, and biparietal diameter can also be used to estimate fetal size in pregnancies where gestation length is uncertain or in utero growth retardation is suspected.39 Normal and abnormal transabdominal sonographic findings are listed in Table 2.5. Serial transabdominal examinations should be performed to verify and monitor fetal well-being or distress. Daily ultrasonographic assessments are commonly performed in high-risk mares. Fetuses experiencing distress are often evaluated several times a day to assess heart rate and activity level. Fetal electrocardiography Fetal electrocardiography (ECG) can be used to monitor fetal viability after day 150.40, 41 Fetal ECGs can be measured continuously using telemetry or can be monitored using more conventional techniques. Patch electrodes are better tolerated by the mare than traditional alligator clamps. Since the late-term fetus changes position, multiple sites of electrode attachment may be needed to obtain a suitable recording.42 As a starting point, the limb lead electrode (LA) is placed on the dorsal midline of the mare in the midlumbar or sacral region. The other limb electrodes are placed on either side of the flank. One electrode
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is often relocated to approximately 15–20 cm cranial to the mare’s udder on the ventral midline. The hair should be clipped prior to lead placement and alcohol or ECG gel applied to improve skin contact with the electrodes. Maternal ECG deflections are larger and slower and usually have a different polarity than those of the fetus, which are of lower amplitude (0.05–0.1 mV). Fetal ECG polarity is variable and depends on fetal position. A poor fetal signal may result from fetal movement, improper electrode placement or electronic noise. The normal fetal heart rate during late pregnancy has a regular rhythm and averages between 65 and 110 bpm. Twinning produces two separate heart rate patterns. Fetal cardiac arrhythmias occur with a variety of abnormal pregnancies. Bradycardia is considered a fetal adaptation to stress, most likely hypoxia. Marked bradycardia often indicates severe hypoxemia and impending fetal demise. Tachycardia occurs in response to fetal activity. Persistent tachycardia in the absence of fetal movement is suggestive of severe fetal distress. Doppler velocimetry Another diagnostic tool that has had minimal application in the pregnant mare is Doppler velocimetry.43 This technique is used in women to examine feto-placental circulation in an attempt to identify pregnancies at risk for fetal intrauterine growth retardation and maternal toxemia. Using a hand-held Doppler transducer, umbilical and uterine arteries can be evaluated and displayed as wave forms. Different vessels have different waveform profiles based on the dynamics of blood flow during systole and diastole. To evaluate relative flow velocity and placental perfusion, the systolic to diastolic ratio (S/D) in the umbilical and uterine vessels is measured. In normal pregnancies the S/D ratio decreases with advancing gestation, reflecting decreasing placental vascular resistance and increasing umbilical blood flow required by the growing fetus.44 McGladdery and co-workers used this technique to examine a small group of pregnant pony mares. Preliminary results showed a similar decline in S/D ratio with advancing gestation.43 Umbilical and uterine circulations are high-flow, low-resistance systems and their wave forms are characterized by a persistent diastolic component. Disappearance of flow during diastole is abnormal and indicates increasing vascular resistance. Changes in flow dynamics may precede, by many weeks, changes in fetal well-being detected by transabdominal ultrasonography. Increases in placental vascular resistance have been associated with infants
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Table 2.5 Fetal and uteroplacental parameters assessed via transabdominal ultrasonography during late gestation (>300 days). From Reef, Vaala et al.34–37, 40 Parameter
Normal range
Abnormal findings/significance
Fetal number
One
>1 fetus; increased risk of abortion; birth hypoxia affecting one or both neonates; disparity in size and degree of maturity of twins
Fetal presentation
Anterior
Transverse or posterior presentations associated with increased risk of dystocia
Fetal activity and tone
Fetus should exhibit some activity during a 30-minute examination
Prolonged periods of fetal inactivity and loss of tone are suggestive of fetal stress, usually hypoxia
Fetal breathing
Rhythmic breathing present
Absence of fetal breathing indicative of fetal stress/hypoxia
Fetal heart rate and reactivity
75 ± 7 bpm; rate should increase by 15–20 bpm with bouts of fetal activity; Normal cardiac rhythm
Persistent bradycardia (FHR <55–60 bpm) or tachycardia (FHR >125 bpm) in the absence of fetal activity are suggestive of fetal stress/hypoxia. FHRs that do not increase with fetal activity and arrhythmias are abnormal
Fetal aortic diameter
22.8 ± 2.15 mm
<18.5 mm (horse fetus) is suggestive of intrauterine growth retardation (IUGR) or twins
Fetal thoracic diameter
10 times the aortic diameter
Less reliable than aortic diameter; smaller thoracic diameter suggestive of IUGR.
Fetal fluids: maximum vertical pocket depth
Amniotic: 7.9 ± 3.5 cm Allantoic: 13.4 ± 4.4 cm
Amniotic: <0.8 cm or >18.5 cm Allantoic: <4.4 cm or >22.1 cm Inadequate fluid volumes may be associated with prolonged fetal hypoxia or premature rupture of fetal membranes; Increased volume of amniotic fluid may be associated with abnormal fetal swallowing
Fetal fluid clarity
Occasional particles observed in both fluid pockets
Large clouds of particles in either fluid compartment is abnormal and may be due to increase in cells (WBC, RBC); large particulate debris in amniotic fluid is indicative of fetal stress and may indicate presence of meconium
Utero-placental thickness
1.15 ± 0.24 cm in the fetal horn; UP thickness is often increased when measured in the non-fetal horn
>2.4 cm; <3.9 mm; Significant placental thickening is suggestive of placentitis or placental edema; Thinner than normal placentas are suggestive of uterine fibrosis and/or placental insufficiency. The surface area of the placenta deemed to be abnormal may be more important than absolute values
Utero-placental contact/integrity
Occasional small anechoic spaces are imaged between the uterus and placenta; placental folding may be more prominent at junction of non-fetal horn and uterine body
Large areas of placental disruption are abnormal and indicative of premature placental separation
suffering from IUGR or small-size-for-gestational age, as well as maternal hypertension and toxemia, and other perinatal complications.44–47 Doppler velocimetry has also been used to evaluate the effect of various treatment strategies, including low-dose aspirin therapy, to improve utero-placental perfusion by reducing vascular resistance. This technique still offers promise for clinical application in equine perinatology.
References 1. Ousey JC, Houghton E, Grainger L, Rossdale PD, Fowden AL. Progestagen profiles during the last trimester of gestation in Thoroughbred mares with normal or compromised pregnancies. Theriogenology 2005;63:1844–56.
2. Holtan DW, Houghton E, Silver M, Fowden AL, Ousey J, Rossdale PD. Plasma progestagens in the mare, fetus and newborn foal. J Reprod Fertil 1991;44(Suppl):511–19. 3. Barnes RJ, Nathanielsz PW, Rossdale PD, Comline RS, Silver M. Plasma progestagens and oestrogens in fetus and mother in late pregnancy. J Reprod Fertil Suppl 1975;23:617–22. 4. Pashen RL, Maternal and foetal endocrinology during late pregnancy and parturition in the mare. Equine Vet J 1984;16:233–6. 5. Hamon M, Clarke SW, Houghton E, Fowden AL, Silver M, Rossdale PD, Ousey JC, Heap RB. Production of 5αdihydroprogesterone during late pregnancy in the mare. J Reprod Fertil 1991;44(Suppl):529–35. 6. Rossdale PD, Ousey JC, Cottrill CM, Chavatte P, Allen WR, McGladdery AJ. Effects of placental pathology on maternal plasma progestagen and mammary secretion calcium concentrations and on neonatal adrenocortical function in the horse. J Reprod Fertil 1991;44(Suppl):579–90.
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Monitoring the High-Risk Pregnancy 7. Lofgren M, Backstrom T, Joelsson I. Decline in serum concentration of 5α-pregnane-3,20-dione prior to spontaneous labor. Acta Obstet Gynecol Scand 1988;67:467–70. 8. Brendemeuhl JP, Williams MA, Boosinger TR. Plasma progestagens, triiodothyronine and cortisol concentrations in post date gestation foals exposed in utero to tall fescue endophyte Acremonium coenophialum. Biol Reprod Mono 1995;1:53–9. 9. Evans TJ, Constantinescu GM, Ganjam VK. Clinical reproductive anatomy and physiology of the mare. In: Youngquist RS (ed.) Current Therapy in Large Animal Theriogenology. Philadelphia: WB Saunders; 1997;43–70. 10. LeBlanc MM, Macpherson M, Sheerin P. Ascending placentitis: What we know about pathophysiology, diagnosis and treatment. Proc Am Assoc Equine Pract 2004;50:127–43. 11. Ousey JC, Forhead AJ, Rossdale PD, Grainger L, Houghton E, Fowden AL. The ontogeny of uteroplacental progestagen production in pregnant mares during the second half of gestation. Biol Reprod 2003;69:540–8. 12. Lofgren M, Host J, Backstrom T. Effects in vitro of progesterone and two 5α-reduced progestins, 5α-pregnan,3,20dione and 5α-pregnan-3a-ol-20-one, on contracting human myometrium at term. Acta Obstet Gynecol Scand 1992;71:28–33. 13. Chavatte-Palmer P, Duchamp G, Palmer E. Progesterone, oestrogen and glucocorticoid receptors in the uterus and mammary glands for mares from mid- to late-gestation. J Reprod Fertil 2000;56(Suppl):661–72. 14. Mason JI, Hinshelwood MM, Murray BA. Tissue-specific expression of steroid 17 α-hydroxylase/C-17,20 lyase, 3βhydroxysteroid dehydrogenase and aromatase in the fetal horse. In: Proceedings Society for Gynecological Investigation Meeting 1993; p. 357. 15. Santschi EM, LeBlanc MM, Weston PG. Progestogen, oestrogen sulphate and cortisol concentrations in pregnant mares during medical and surgical disease. J Reprod Fertil 1991;44(Suppl):627–34. 16. Macpherson ML. Diagnosis and treatment of equine placentitis. Vet Clin North Am Equine Pract 2006;22(3):763– 76. 17. Brendemeuhl JP. Effects of Acremonium coenophialuminfected tall fescue on progestagen production in pregnant mares. In: Proceedings Society of Theriogenology 1992; pp. 108–12. 18. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995;73(3):899–908. 19. Cross DL. Fescue toxicosis in horses. In: Bacon CW, Hill NS (eds) Neotyphodium/Grass Interactions. New York: Plenum Press, 1997; pp. 289–309. 20. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J Reprod Fertil 1979;Suppl27: 499–509. 21. Silver M. Prenatal maturation, the timing of birth and how it may be regulated in domestic animals. Exp Physiol 1990;75:285–307. 22. Garfield RE, Kannan MS, Daniel ME. Gap junction formation in the myometrium: control by estrogens, progesterone and prostaglandins. Am J Physiol 1980;238:C81 23. Madej A, Kindahl H, Nydahl C, Edqvist LE, Stewart DR. Hormonal changes associated with induced late abortions in the mare. J Reprod Fertil 1987;Suppl35:479. 24. Klonisch T, Mathias S, Cambridge G, Hombach-Klonisch S, Ryan PL, Allen WR. Placental localization of relaxin in the pregnant mare. Placenta 1997;18(2):121–8.
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25. Stewart DR, Addiego LA, Pascoe DR, Haluska GJ, Pashen R. Breed differences in circulating equine relaxin. Biol Reprod 1992;46(4):648–52. 26. Ryan PL, Vaala WE, Bagnell C. Relaxin as an indicator of placental insufficiency. Proc Am Assoc Equine Pract 1998;(44):62–3. 27. Ryan PL, Bennet-Wimbush K, Vaala WE, Bagnell CA. Systemic relaxin in pregnant pony mares grazed on endophyte-infected fescue: effects of fluphenazine treatment. Theriogenology 2001;56(3):471–83. 28. Ryan PL, Christiansen DL, Hopper RM, Bagnell CA, Vaala WE, Leblanc MM. Evaluation of systemic relaxin blood profiles in horses as a means of assessing placental function in high-risk pregnancies and responsiveness to therapeutic strategies. Ann NY Acad Sci 2009;1160:169–78. 29. Ousey JC, Dudan F, Rossdale PD. Preliminary studies of mammary secretions in the mare to assess foetal readiness for birth. Equine Vet J 1984;16:259–63. 30. Leblanc MM. Induction of parturition in the mare: Assessment of readiness for birth. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 34–9. 31. Renaudin CD, Troedsson MHT, Gillis CL, King VL, Bodena A. Ultrasonographic evaluation of the equine placenta by transrectal and transabdominal approach in the normal pregnant mare. Theriogenology 1997;47:559. 32. Renaudin CD, Troedsson MHT, Gillis CL. Transrectal ultrasonographic evaluation of the normal equine placenta. Equine Vet Educ 1999;11:75. 33. Renaudin CD, Liu IKM, Troedsson MHT, Schrenzel MD. Transrectal ultrasonographic diagnosis of ascending placentitis in the mare: a report of two cases. Equine Vet Educ 1999;11:69. 34. Reef VB, Vaala WE, Worth LT, Sertich PL, Spencer PA. Ultrasonographic assessment of fetal well-being during late gestation: Development of an equine biophysical profile. Equine Vet J 1996;28(3):200–8. 35. Reef VB, Vaala WE, Worth LT. Transabdominal ultrasonographic assessment of fetal well-being during late gestation: a preliminary report of the development of an equine biophysical profile. Equine Vet J 1995;28:200–8. 36. Reef VB, Vaala WE, Worth LT. Transabdominal ultrasonographic evaluation of the fetus and intrauterine environment in healthy mares during late gestation. Vet Radiol Ultrasound 1995;36:533–41. 37. Reef VB. Late term pregnancy monitoring. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 410–16. 38. Adams-Brendemuehl C, Pipers FS. Antepartum evaluation of the equine fetus. J Reprod Fert 1987;35(suppl.):565–73. 39. Pantaleon LG, Bain FT, Zent W, Powell DG. Equine fetal growth and development. Comp Contin Educ 2003;25(6):470–6. 40. Vaala WE, Sertich PL. Management strategies for mares at risk for periparturient complications. Vet Clin North Am Equine Pract 1994;10(1):237–65. 41. LeBlanc MM. Identification and treatment for the compromised equine fetus: A clinical perspective. Equine Vet J Suppl 1997;24:100–3. 42. Wilkins PA. High risk pregnancy. In: Paradis MR (ed.) Equine Neonatal Medicine. Philadelphia: Elsevier Saunders, 2006; pp. 13–29. 43. McGladdery AJ, Ousey JC, Rossdale PD. Serial Doppler ultrasound studies of the umbilical artery during
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equine pregnancy. In: Proceedings Third Conference of the International Veterinary Perinatology Society, 1993; p. 37. 44. Schulman H, Fleischer A, Stern W, Farmakides G, Jagani N, Blattner P. Umbilical velocity wave ratios in human pregnancy. Am J Obstet Gynecol 1984;148:985. 45. Ghosh G, Bregorowicz A, Brasert M, Maczkiewicz M, Kobelski M, Dubiel M, Gudmundsson S. Evaluation of third trimester uterine artery flow
velocity indices in relationship to perinatal complications. J Matern Fetal Neonatal Med 2006;19(9):551–5. 46. Gudmundsson S, Korszum P, Olofsson P, Dubiel M. New score indicating placental vascular resistance. Acta Obstet Gynecol Scand 2003;82(9):807–12. 47. Pietryga M, Brazert J, Wender-Oegowska E, Biczysko R, Dubiel M, Gudmundsson S. Abnormal uterine Doppler is related to vasculopathy in pregestational diabetes mellitus. Circulation 2005;112:2496–500.
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Chapter 3
Management of the High-Risk Pregnancy Elizabeth M. Santschi and Wendy E. Vaala
Introduction The treatment of high-risk pregnancy has two main objectives: the most important is treatment of the primary condition, and the second is therapy directed at promoting gestation. Sometimes the therapies are the same, and in other instances the therapies run concurrently. This chapter will not deal in great detail on how to treat the primary conditions that affect pregnant mares, but will provide information on how management of the primary condition can affect pregnancy and overall mare health. Specifically, additional detail about treatments to promote gestation is warranted as therapies are sometimes controversial, and usually poorly understood. An additional concern in the treatment of highrisk pregnancy is determining which patient (mare or foal) is the main concern of the owner. Of course, sometimes there is no choice to be made, but if there is, weighting patient priorities to the owner can be important. A final general consideration is the detection of fetal death. If the fetus dies, the mare will often deliver the fetus, although one of the authors (WV) has had several late-term pregnant mares present with dead fetuses and no signs of impending parturition. One case was an older dam in her last trimester that presented with hydrops allantois and a small dead fetus. There is another report of a retained mummified fetus in a mare that had received altrenogest for a significant portion of gestation.1 Therefore, any high-risk mare should receive regular transabdominal sonograms to assess feto-placental well-being. In early to mid-pregnancy, mares can usually deliver dead fetuses without problem, but in late pregnancy, dystocia is a common result. The dystocia is due to the dead fetus’s inability to extend its head, neck, and forelimbs to properly present itself for de-
livery. The dystocia is usually poll and carpal flexion, and can be easily corrected if detected early. Mares can still control the delivery to some extent, and most dead foals will be delivered at night, necessitating 24 h monitoring. Determination of fetal death is done by transabdominal ultrasound detection of asystole. If a fetus is dead and cannot be monitored, induction of parturition should be considered (Chapter 232). Pro-gestational therapies
Progestagens Progesterone is the hormone essential to pregnancy maintenance in many species, and has many active metabolites, called progestagens. The mare has a complex progestagen profile during gestation, and the compounds and their sources change during gestation.2 Probably, not all the compounds have been identified or characterized, and neither is their activity fully understood. What appears clear is that it is not common for a mare to lose a pregnancy due to disease simply due to ‘progesterone insufficiency.’ However, experimental endotoxemia can cause pregnant mares to abort due to luteolysis if endotoxin is administered before day 55 of pregnancy.3, 4 This may also occur in natural disease, and would explain early pregnancy loss in cases of colic and endotoxemia. We do know that progestagen concentrations in the maternal circulation vary widely during high-risk pregnancy,2, 5, 6 and these changes are more likely the result of the primary condition causing the high-risk pregnancy, rather than the cause of the risk to the pregnancy. Advocates of progestagen supplementation cite its ability to promote uterine quiescence and inhibit prostaglandin-mediated abortion.7, 8 Anecdotal reports abound regarding the apparent ability of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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progestagen supplementation to prevent pre-term labor in late gestational mares with placentitis. Critics of such therapy question the benefit of using progestagen supplementation in mares that may already have elevated concentrations of progesterone metabolites.2, 9 Most mares at risk for premature delivery are also receiving other therapy including antibiotics, non-steroidal medication such as flunixin meglumine, and anti-cytokine drugs such as pentoxifylline. Until recently, it was difficult to separate the impact of altrenogest administration from the effects of other concurrently administered medications. Research using mares with experimentally induced bacterial placentitis evaluated the impact of administering or withholding altrenogest supplementation in addition to antibiotics and anticytokine medications.10, 11 Ten of 12 mares with experimentally induced placentitis delivered live foals when altrenogest (ReguMateTM , 0.088 mg/kg orally q 24 h) was added to the treatment regimen that included trimethoprim sulfa (30 mg/kg orally q 12 h) and pentoxifylline (8.5 mg/kg orally q 8 h) in comparison to a previous study, using the same experimental infection model, 5/6 infected pony mares aborted when they were treated with only trimethoprim sulfa and pentoxifylline.12 Pre-term birth is a leading cause of neonatal infant morbidity and mortality in the developed world. Progesterone supplementation has been used in women with a history of pre-term birth since the 1970s. In reviews of multicentric trials, women receiving progesterone were less likely to experience recurrent miscarriage and premature delivery, to have an infant of low birthweight or to have an infant diagnosed with interventricular hemorrhage.13–17 However, the use of progesterone in pregnant women remains controversial and its precise mechanism of action, optimal formulation, route of administration, dose, and optimal gestational age at initiation have yet to be determined.15 Because of the activity of progestagens in pregnancy maintenance, it has been debated whether supplementing progestagens during pregnancy is beneficial. We cannot solve that controversy; however, experiments with mares treated with abortigenic doses of prostaglandins and treated with altrenogest do suggest that the drug acts to support pregnancy.7, 18 Progestagens are suggested to quiet the uterus, and placentitis has been shown to affect the uterine myoelectrical activity in mares.19 It has been suggested that altering the uterine activity may improve utero-placental perfusion, and improve the fetal condition. Progestagen therapy has been suggested to promote uterine quiescence,20 which could improve uterine perfusion and improve fetal health. The authors favor the use of progestational compounds in the treatment of high-risk pregnancy. Dur-
ing a crisis, altrenogest is used at 0.88 mg/kg, p.o., s.i.d. This dose is reduced by 50% once clinical resolution of the crisis has occurred. If hospitalized, mares are kept on altrenogest for a minimum of a week after returning to their home farm. Many managers prefer to treat the mare for a longer duration. There appears to be little downside to treatment in this fashion, other than expense. However, determination of fetal viability is recommended before, and periodically during, therapy. Progestagen supplementation is available as oral altrenogest or injectable progesterone. Oral altrenogest, a synthetic progestin, has been administered at doses of 0.088 mg/kg once daily to help prevent abortion and premature delivery in mid- to lateterm mares.7, 11, 21, 22 Alternatively, and anecdotally, some clinicians prefer using injectable progesterone (300 mg, q 24 h, i.m.). One author (WV) recommends using a short-acting form of progestin supplementation in late-term mares rather than preparations claiming longer duration of action, to permit daily adjustments in therapy as warranted. It has been the author’s experience that mares can deliver live foals as well as abort while receiving oral altrenogest at doses up to 0.044 mg/kg orally q 24 h. If mares on progestagen supplementation do reach their anticipated foaling dates then it is recommended to decrease the dose of altrenogest gradually, rather than discontinuing therapy abruptly.
Non-steroidal anti-inflammatories Non-steroidal anti-inflammatories (NSAIDs) are used in the treatment of many conditions that affect pregnant mares. The most commonly use drugs, such as phenylbutazone and flunixin meglumine, are nonselective inhibitors of the cyclo-oxygenase pathway. Recently COX-2 selective inhibitors such as firocoxib have become available. For pregnancy maintenance, these anti-inflammatories are used to reduce placental and uterine inflammation and, more specifically, to inhibit prostaglandin production. This is believed especially important in cases of placentitis.19 Flunixin meglumine has also been shown experimentally to protect corpus luteum dependent pregnancies from the effect of prostaglandins.23 Concerns have been raised that chronic use of NSAIDs may impair the ability of the foal to adapt its cardiovascular system after birth, but this does not appear to be a clinical concern.
Steroids If medical conditions warrant induction, or premature delivery appears imminent despite therapy, then the delivery of as mature a foal as possible becomes the next focus of therapy. Fetal organ maturation is regulated by the hypothalamic–pituitary–adrenal
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Management of the High-Risk Pregnancy (HPA) axis and a rise in fetal plasma cortisol concentrations during the last several days before delivery.24 Administration of adrenocorticotropic hormone (ACTH) directly to the equine fetus in utero promotes precocious fetal maturation and early delivery, but is too invasive a technique to use safely in practice.25 ACTH administration to late-term mares stimulated fetal maturation in some, but not all, mares.26 In 1975, Alm and co-workers27 used high doses of dexamethasone (100 mg, i.m., q 24 h for 4 days) to induce mares to foal. Maternal glucocorticoid administration significantly reduced gestation length and resulted in the birth of foals that were small but reportedly mature. Another study performed using the same prepartum dose of steroids resulted in dystocia and fetal death.28 Anecdotal reports from clinical practice suggested that varying doses of dexamethasone administered prepartum to mares resulted in improved fetal survival. Evaluation of the potential benefits of maternal steroid therapy was revisited recently by Ousey and co-workers.29 Five Thoroughbred mares received 100 mg of dexamethasone once daily from 315 to 317 days of gestation. Dexamethasone-treated mares delivered significantly earlier than controls. Duration of second and third stage labor did not differ between treated and control mares. One steroid-treated mare experienced premature placental separation, but the allantochorion ruptured spontaneously before manual intervention. All of the foals appeared healthy and mature and exhibited normal postpartum behavior. Steroid-treated mares had poorerquality colostrum at the time of delivery, and four foals in the treated group required colostrum supplementation. Administration of dexamethasone to prepartum mares remains an attractive and inexpensive method of inducing precocious fetal development. However, caution is urged before using this therapy since the number of mares treated under controlled conditions has been small, and the few studies examining this induction strategy have differed in their results. The optimal dose of dexamethasone and the frequency and timing of administration require further research. In particular, the time of gestation at which this therapy is initiated may be critical in terms of fetal endocrine development and response to treatment. Antenatal steroid administration for women at risk for premature delivery has emerged as an effective intervention for prevention of respiratory distress syndrome, reduction of neonatal morbidity and mortality.30,31
Pentoxifylline Pentoxifylline is a xanthine derivative drug that is aimed at improving peripheral blood flow by re-
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laxing smooth muscle, and improving red blood cell flexibility and platelet deaggregation. It also is an inhibitor of TNF-α, and therefore has antiinflammatory potential.32 Recent studies using microdialysis technology in pregnant pony mares have shown that, following oral administration, pentoxifylline penetrates placental membranes and is detectable in allantoic fluid.33 The use of pentoxifylline seems to be one of clinician preference, and efficacy in the promotion of pregnancy has not been established.
Presumptive tocolytics Clenbuterol is a beta-2-adrenergic agonist and is therapeutically used to relax smooth muscle in the respiratory tract. When used intravenously in cattle during dystocia, it facilitates manipulation, presumably due to uterine relaxation. In pregnant mares, intravenous clenbuterol causes transient tachycardia in both mare and fetus and causes a reduction in palpable uterine tone.34 However, intravenous clenbuterol has also been found to be ineffective in delaying the onset of normal parturition in healthy mares,35 and was not believed to be effective in preventing abortion in pregnant mares with colic.36 The effect of clenbuterol on the uterus in the horse when used orally is unknown, and its use as a tocolytic to forestall preterm labor is questionable. Another theoretical tocolytic is isoxsuprine hydrochloride. Isoxsuprine is a beta adrenoreceptor antagonist and agonist that has the potential to contribute to uterine relaxation. However, isoxsuprine has a short half-life when given intravenously, and has poor bioavailability when given orally.37 It probably has little therapeutic effect on the tone of the uterus.
Anti-dopamine compounds Chronic stimulation of dopamine release in the pregnant mare impairs both fetal and maternal readiness for birth. The dopaminergic receptors are the sites of binding for the ergot alkaloid, ergovaline, found in fescue grass. Fescue toxicosis causes a plethora of reproductive maladies in pregnant mares. When the toxin interacts with receptors, dopamine is released and inhibits fetal maturation by preventing proper maturation of the fetal adrenal–cortical axis.38 Improper maturation of the fetus results in a lack of fetal and maternal readiness for birth. Dopamine also directly inhibits maternal readiness for birth by inhibiting the secretion of prolactin which stimulates mammary gland development. An important compound in the treatment of fescue toxicosis is domperidone, which works by blocking dopaminergic receptors. Domperidone R ; 1.1 mg/kg orally q 24 h) is used to help (Equidone mature pregnancies in mares exposed to fescue39 and
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is used in other mares to treat agalactia due to other causes. For example, domperidone is useful to treat late-term mares with pituitary pars intermedia dysfunction (PPID; e.g., equine Cushing syndrome) that have been treated during pregnancy with pergolide. Pergolide acts as a dopamine agonist and, while it alleviates many of the clinical signs associated with PPID, it also results in agalactia, prolonged gestation lengths, and difficult deliveries if mares are not removed from the drug at least 2 to 3 weeks prior to anticipated delivery. Domperidone is often used to hasten lactation and help prepare these mares for delivery once the pergolide has been discontinued.
Vitamin E (tocopherol) Vitamin E is administered orally as an antioxidant for the treatment of placental inflammation and dysfunction, and as a neuroprotective agent for the fetus. When administered prior to traumatic brain injury, vitamin E administration improves neurological outcome in experimental models and may be useful in the treatment of neonatal hypoxic-ischemic encephalopathy.40, 41 It has been extrapolated that high doses of vitamin in the high-risk mare and her fetus may ameliorate the effects of peripartum hypoxia. Daily doses of 1000–8000 IU of vitamin E have been used in mares with suspected utero-placental compromise; however, there is no evidence that this treatment is useful in the horse.
Supplemental oxygen Oxygen exchange between maternal and fetal circulations in the horse is very effective42, 43 and is flowlimited due to the high diffusing capacity of oxygen.44 The countercurrent system of maternal and fetal blood flow within the capillary architecture of the equine feto-placental unit also contributes to optimal oxygen exchange between dam and fetus.44 Oxygen therapy for pregnant mares with compromised placental function has been advocated to reduce the risk of fetal hypoxia prior to and during parturition. Nasal insufflation with oxygen has been demonstrated to raise the Pa o2 in horses with respiratory disease,45 and is recommended for late-term mares at risk for poor oxygen delivery to the uterus.46–48 At flow rates of 10–20 L/min, Pa o2 concentrations can be increased by two- to threefold. Increasing the delivery of oxygen to the placenta should be helpful to fetuses experiencing hypoxia, and can be considered for late-term mares after colic surgery, and those affected by placentitis, premature placental separation, and dystocia. Mares with advanced recurrent airway obstruction are also candidates for oxygen therapy. Supplemental oxygen should also be administered into any accessible foal airway during the correc-
Figure 3.1 Nasotracheal intubation of a foal during correction of dystocia.
tion of dystocia. Nasotracheal intubation and positive pressure ventilation can be very effective in supporting cardiopulmonary function in foals trapped within the birth canal during severe dystocia (see Chapter 13; Figure 3.1; Chapters 255 and 258).
Nutritional support The equine placenta is 5–7 times more efficient in nutrient transfer per unit area than either cow or sheep44 and this may be attributed to the countercurrent blood flow exchange mechanism in the equine placenta. The high transplacental glucose gradient that exists between dam and fetus across the equine placenta results in efficient glucose transfer between mare and fetus.42, 44 During late gestation, glucose utilization by the equine placenta decreases, so that proportionately more of the transferred glucose taken up by the uterus is available to the equine fetus.44 Therefore, the fetal foal is expected to benefit nutritionally from efforts made to maintain or enhance glucose concentrations in the mare. Fasting of the late-term mare for even 12 to 30 hours has been shown to increase production of uterine prostaglandin F metabolites (PGFM).49 Likewise, enteral re-feeding or intravenous infusion of glucose produces a rise in plasma glucose and
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Management of the High-Risk Pregnancy immediate fall in uterine PGFM concentrations. This fasting-induced surge in PGFM was greatest in late gestational mares. In one study, five out of eight fasted mares delivered prematurely within 1 week of the period of food withdrawal (<48 hr).49 Hypoglycemia in the pregnant mare can adversely affect placental metabolism and glucose delivery to the fetus. Based on the experience of one of the authors (WV), it is recommended that late-term mares that must be fasted for >12–24 hours, as is often prescribed following abdominal surgery, should be maintained on glucose infusions to prevent hypoglycemia and possible hyperlipemia, optimize glucose delivery to the fetus, and reduce the risk of prostaglandin-mediated pre-term labor. Intravenous infusion of 2.5% or 5.0% dextrose in 0.45% saline administered at fluid rates of 1–2 mg/kg/min of dextrose are recommended. Pregnant mares that are anorexic due to systemic illness should be supplemented via nasogastric intubation or with partial parenteral nutrition, to prevent a similar cascade of prostaglandin-mediated events.
Induction techniques In an elegant study using direct radiography to monitor the position, posture and presentation of the equine fetus during late gestation and delivery,50 the important sequence of events during stage 1 labor was clearly demonstrated. During naturally occurring stage 2 labor the full-term fetus undergoes purposeful movements to rotate the head and forelimbs into the dorsal position, followed by extension of the forelimbs and neck into the pelvic canal. The stimulus for initiation of these movements is not known but, given the relatively low incidence of dystocia in spon-
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taneously foaling mares, the mechanism that controls this in utero fetal ‘dance’ is normally quite efficient. The increased incidence of dystocia and premature placental separation associated with various methods of induction28, 51, 52 stresses the importance of limiting chemical induction of the mare to a select group of periparturient emergencies. Questions to consider and discuss with the owner are presented in Table 3.1. The optimal dose of oxytocin and route of administration for induction remains controversial. Some researchers and clinicians promote the use of low doses of oxytocin as being more physiological and less likely to induce untoward delivery complications.53, 54 Whenever possible, induction should only be undertaken in late-term mares that are close to their own physiological foaling dates. Indications for induction should be limited to conditions that seriously threaten maternal or fetal health should the pregnancy be allowed to continue, or should unsupervised, spontaneous delivery occur. Examples of such conditions include hydrops, prepubic tendon rupture, imminent death of the mare due to colic or other systemic illness, and maternal history of severe dystocia requiring mandatory assistance during delivery. Criteria used to assess readiness to foal include normal udder development, the presence of colostrum in the teats, a favorable electrolyte profile in prepartum mammary secretions,55 and a dilated or softening cervix. If induction is imperative and the cervix is unfavorable, topical instillation of prostaglandin E2 gel to the cervix should be considered. Intracervical application of prostaglandin E2 has been used in women56 and mares57 to induce cervical ripening prior to induction of parturition.
Table 3.1 Questions to address when considering induction in the mare Question
Implication
Is the mare or foal (or both) the owner’s primary concern?
If foal survival is critical then prolonging the pregnancy to allow maximal fetal maturation is important and induction should be delayed as long as possible
If a dystocia occurs and can not be resolved with manual manipulation, is a cesarean section an option?
If a C-section is an option then induction should be performed near a surgical facility
Does the owner understand the inherent risks associated with induction (e.g., cervical trauma due to insufficient cervical dilation, premature placental separation, dystocia, delivery of a foal suffering from prematurity/dysmaturity and/or peripartum asphyxia, fetal/neonatal death) and is aggressive neonatal care an option?
Personnel and supplies should be available to provide neonatal resuscitation/oxygen supplementation and initial stabilization of a high-risk foal
Have physiological criteria been met prior to induction, suggesting that the mare is close to her ‘own’ due date? Is colostrum present in udder? Are mammary secretion electrolytes consistent with fetal maturity? Is the cervix soft and dilated?
Alternate source of colostrum should be available; if mammary secretion electrolytes are not compatible with term pregnancy then supportive care for a premature foal should be available and the potential cost of such intensive treatment.
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Management of conditions that can cause high-risk pregnancy Maternal conditions
Gastrointestinal causes of colic The general treatment of colic in the pregnant mare does not differ from the non-pregnant horse, but late-term pregnancy can complicate diagnosis (as mentioned in Chapter 1) and surgery. The overall pregnancy loss after colic surgery is about 20%, with approximately half of those abortions strongly linked to the colic episode or surgery.36, 58 Most losses were associated with hypoxia in late-term mares or persistent endotoxemia. Occasionally, there is a temptation to remove a fetus in a mare during an exploratory for colic. There can be compelling reasons to do so, such as an inability to access a lesion where the mare is the priority patient (e.g., distal small colon lesions), or a mare that is being euthanized; however, fetal removal before term should be avoided in most cases. Maturity in the foal by dates is just an estimate, and most fetuses stand the best chance of survival if left in place. Maturation of the fetal hypothalamic–pituitary–adrenal axis is a relatively late event believed to occur 3 to 5 days prior to delivery.59 Therefore, delivery of a neonate seemingly close to term, based on calculated gestational age, can result in a foal poorly adapted for extrauterine survival due to marked endocrine and metabolic immaturity. If a foal is removed, a neonatal intensive care unit is a must, as rigorous supportive care and monitoring will likely be required. During surgery and recovery, avoidance of hypoxemia is important to the health of the fetus.46, 58 All possible attempts should be made to keep the maternal Pa o2 above 80 mmHg, and oxygen insufflation should be considered in the recovery stall. During the perioperative management of colic in pregnant mares, consideration should be given to administering flunixin meglumine and supplemental progestagens. Flunixin should be continued for as long as uterine inflammation from surgery or signs of clinical endotoxemia exist.
Endotoxemia The management of endotoxemia in pregnant mares requires treatment of the primary condition, and supportive therapy to minimize the deleterious effects of the systemic inflammatory response. Most of the primary conditions will be gastrointestinal, but any Gram-negative infection has the potential to elaborate endotoxin. Once in the circulation, endotoxemia induces a systemic inflammatory response syndrome mediated by the release of inflammatory
cytokines, resulting in deleterious alterations in perfusion and coagulation that can lead to death.60 Supportive therapy can include the intravenous administration of isotonic and hypertonic fluids, colloids, and plasma. Drugs administered include cyclooxygenase inhibitors such as flunixin meglumine, and endotoxin-binding agents such as polymyxin B. 61 Specific therapies directed at the pregnancy include the use of progestagens, and perhaps supplemental oxygen during an endotoxic crisis. Therapy should be continued for several days or more after clinical signs resolve and feto-placental monitoring is reassuring.
Colitis Colitis is particularly hazardous to maintenance of pregnancy and delivery of a non-septic foal when the condition is accompanied by profuse diarrhea, leukopenia/neutropenia, hypoproteinemia, and dehydration. Not only do the hemodynamic events associated with endotoxemia and dehydration jeopardize utero-placental perfusion and increase the risk of fetal hypoxia, but voluminous diarrhea results in persistent fecal contamination of the perineum and increases the risk of ascending placentitis prepartum and infection of the fetus during delivery or immediately postpartum. Affected mares may also experience periodic episodes of bacteremia leading to hematogenous infection of the placenta and fetus. Broad-spectrum, bacteriocidal antibiotic therapy should be considered for late-term mares with severe colitis. If clostridial pathogens are suspected or confirmed, then metronidazole should be added to the treatment regimen. Additional supportive therapy for colitis often includes intravenous fluid therapy with crystalloids, colloid therapy with plasma and/or hetastarch, and antidiarrheal medications including oral intubation with d-smectite. Progestagen supplementation with oral altrenogest is advised for mares showing concurrent signs of endotoxemia. Applying a tail wrap coupled with frequent cleaning of the mare’s perineum and udder prior to delivery and during the immediate postpartum period may help reduce the foal’s exposure to intestinal pathogens. Regardless of aggressive peripartum precautions, foals born to mares experiencing infectious diarrhea should be assumed to be septic until proven otherwise, and should be evaluated.
Uterine torsions Uterine torsions can be corrected either by a rolling procedure or surgically.62 The rolling procedure is inexpensive, but requires general anesthesia, cannot address concurrent gastrointestinal lesions or uterine tears, and is not always effective. It is recommended
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Management of the High-Risk Pregnancy only for mid-gestation torsions or when owners lack the resources to pay for surgery. Surgical treatment of uterine torsions can be performed either via flank laparotomy standing or via a ventral midline celiotomy under general anesthesia.62, 63 Flank laparotomies are preferred for mid-gestation torsions and may improve fetal outcome as compared to general anesthesia.62 Late-gestation uterine torsions are a challenge, and it is the author’s (EMS) preference to correct these torsions via a ventral midline celiotomy under general anesthesia. These torsions are often difficult to correct, and this approach provides best control of the dam and allows for an abdominal exploratory. After correction, these pregnancies should be closely monitored via ultrasound and treated with NSAIDs and progestagens.
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Figure 3.2 Mare with hydrops, wearing a commercially available belly bandage for added support. Photo courtesy of University of Florida School of Veterinary Medicine.
Lameness Lameness conditions do not usually affect pregnancy in the horse unless they result in exercise restriction. Pregnancy may impact on the lameness condition if the increasing weight of the pregnancy is a factor. Pregnant mares eat better, have less gastrointestinal problems, and develop less peripheral edema if they have access to pasture and gentle exercise, so this should be allowed wherever possible. Lamenesses should be treated appropriately for the specific condition. Laminitis is common, and frequently frustrating disease due to the severe pain and the common progression. The use of NSAIDs is common for all lameness issues, and should be used appropriately. Other analgesic regimens can be considered when appropriate such as opiates and intravenous lidocaine, but their impact on the fetus is unknown.
Abdominal wall injuries The successful management of abdominal wall injuries can be very challenging. It is often difficult to distinguish abdominal wall hernia and prepubic tendon rupture, and clinically that may be irrelevant. Treatment of the mare is symptomatic, and includes pain control, restriction of exercise, and abdominal wall support. If the mare has a predisposing condition such as hydrops, this condition must be managed concurrently with efforts to prevent further abdominal wall deterioration. Effective delivery of abdominal support can be difficult to achieve and requires strong support materials. Several options for abdominal support are available, but most are ineffective. Heavy canvas panels that can be laced together and adjusted are preferred by the author (EMS), and require disposable cotton pads underneath as well as padding over the back to reduce the risk of pressure necrosis.
Circumferential wrapping with tape over absorbable padding can also be applied, but is not adjustable and needs to be reapplied, often daily. Mares also sweat profusely under impermeable tapes. Several sources exist for commercially available belly bandages (see Figure 3.2) constructed of canvas or stretchy fabric with Velcro bands used to secure and adjust the bandage to provide different levels of comfort. These bandages can be reused and laundered as needed. Examples of commercially produced bandages used for abdominal wall defects include CM Heal Hernia Belt and Post Surgical Pressure Bandage (CM Equine Products, www.cmequineproducts.com) and The BoaTM (Rewrap Products, www. wire2wirevetproducts.com/rewrapBOA.htm). The outcome of these cases is variable and difficult to predict. Some mares do well with induction of parturition and controlled vaginal delivery, while others may spontaneously herniate and eviscerate during the induction. Elective cesarean section is not an attractive option since abdominal closure is difficult and at increased risk for dehiscence, and there may be peritonitis associated with the abdominal wall rent. A small retrospective study indicates that foal survival was better in conservatively managed mares; however, the extent of abdominal injury was not described.64 The most severe injuries are more likely to be those treated by intervention, which may confound the results. However, intervention should be used cautiously, if at all. Mares with severe injuries that involve the prepubic tendon and muscular portions of the abdominal wall are very difficult to manage. Mares with edema and pain from stretching (rather than tearing) of the abdominal tunic can be successfully managed with conservative treatment. Whether parturition is induced or is spontaneous, delivery must be attended since considerable
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assistance is usually required to extract the fetus due uterine inertia and the lack of an effective abdominal press by the mare.
Miscellaneous infectious agents Management of mares exposed to equine herpesvirus (EHV-1) or equine arteritis virus should include isolating them from known carriers, and if sufficiently debilitating, treatment of fever with NSAIDs. Acyclovir has been used prophylactically in EHV-1 outbreaks, and the pharmacokinetics described.65 However, the effect of acyclovir on a developing fetus is unknown, so that possible risk should be weighed against the risk of abortion. More recently, research has documented the use of valacyclovir (loading dose: 27 mg/kg p.o., q 8 h for 2 days; maintenance dose: 18 mg/kg p.o., q 12 h) to reduce the days of nasal shedding and viremia and ameliorate the neurological signs following challenge with a neuropathic mutant strain of EHV-1.66 Mares with high or rising titers to leptospirosis should be treated with antibiotics (usually penicillin)67 and isolated from wet areas where the organism may live.
Pelvic and cervical abnormalities The vaginal delivery of foals from mares with pelvic and cervical abnormalities raises both fetal and maternal risk. In the case of pelvic injuries, the internal pelvic canal is diminished by callus formation after fracture healing, and can cause dystocia. The most common cervical abnormality is scarring after repair of previous damage, and a failure of cervical dilation can cause dystocia, and will likely re-tear and result in permanent infertility. Elective hysterotomy for fetal delivery will avoid difficulties with vaginal delivery. One report indicates that mare (100%) and foal (88%) survival was good in a case series of eight pregnancies.68 However, others believe that the physics of vaginal delivery confers advantages to foal survival and indicate lower foal survival rates (60%) than previously reported (M.M. LeBlanc, 2008, personal communication). The most difficult decision to make is the timing of delivery, and is made on the basis of breeding dates and, more importantly, maternal signs of readiness for birth including cervical softening and the presence of colostrum in the udder. Mammary secretion electrolytes (sodium, potassium, and calcium) can be used as supplementary information about fetal maturity,69 and are best used, not to determine when to deliver, but rather when to postpone delivery. Mares hospitalized for elective hysterotomy should be on ‘foaling watch’ as, despite best pre-
dictive efforts, mares may start to deliver with little premonitory signs, and surgery should occur before the foal has a chance to engage the pelvic inlet. In one retrospective study of eight elective cesarean sections, foals experienced reduced anesthetic depression and improved survival if the time elapsed from induction of anesthesia to delivery of the foal was ≤ 20 minutes.68 Foals delivered by elective hysterotomy should be treated as high risk, and also should be removed from the hospital as soon as practical after delivery, to avoid nosocomial infections.
Endometrial insufficiency There is no proven effective treatment for endometrial insufficiency. Treatments that could be considered include altrenogest, pentoxyphylline, and insufflation with oxygen. Of greatest importance in the management of these mares is to attend to their overall health, and to be patient when the mare’s gestation appears over-long. Equine pregnancy is not considered overlong until 365 days, and in mares with placental insufficiency and poor-quality endometrium, the fetus should stay in the uterus until it send the signal that it is ready for birth. Placental conditions
Placentitis The foundation of the management of placentitis is the administration of antimicrobials to eliminate the causative organisms. The most common organisms isolated from either vulvar discharge in the dam prepartum or the placentas after abortion or stillbirth are Streptococcus equi subspecies zooepidemicus, Escherichia coli, Pseudomonas aeruginosa, Klebsiella and nocardioform species. Most bacteria are believed to ascend through the cervix and affect the caudal portion of the placenta in the region of the cervical star,70 although some (especially nocardioform organisms) have an effect further cranial in the uterus along the base of the uterine horns.71 Early termination of pregnancy may be the direct result of fetal infection or may be secondary to the proinflammatory cytokine cascade triggered by chorionic inflammation. Therapy is often multifaceted and is aimed at treating infection with antimicrobials, blocking proinflammatory cytokines with NSAIDs and pentoxifylline, promoting uterine quiescence and cervical tone with altrenogest, and minimizing hypoxic tissue damage with antioxidants and maternal oxygen therapy. Medications and dosages commonly used to treat placentitis are presented in Table 3.2. Many mares affected with placentitis are treated with a combination of a beta lactam and an aminoglycoside (in North America, usually penicillin and
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Table 3.2 Therapeutic agents commonly used to treat placentitis in the mare Proposed action/rationale for therapy
Drug/therapy
Dose, route, frequency of administration
Antimicrobial
Trimethoprim sulfamethoxazole Potassium penicillin Procaine penicillin G Ceftiofur Cefazolin Gentocin Flunixin meglumine Phenylbutazone Pentoxifylline Altrenogest Vitamin E (tocopherol) Intranasal oxygen
15–30 mg/kg, orally q 12 h 22 000 units/kg, i.v. q 6 h 22 000 units/kg, i.m. q 12 h 2.2 mg/kg, i.v. or i.m. q 12 h 20 mg/kg, i.v. q 6 h 6.6 mg/kg, i.v. or i.m. q 24 h 0.5–1.0 mg/kg i.v. q 12 h 2–4 mg/kg i.v. or p.o. q 12–24 h 8.5 mg/kg p.o. q 8 h 0.088 mg/kg p.o. q 24 h 1000–8000 IU p.o. q 24 h 10–20 L/min
Anti-inflammatory (anti-prostaglandin, anti-cytokine)
Tocolytic/uterine quiescence Anti-oxidant Oxygen delivery
gentamicin) to apparent good effect. Penetration of both drugs into the allantoic fluid has been demonstrated,72 although the optimal dosing regimen for treatment of the conceptus is not known. It may be sufficient to get the drugs to the utero-placental interface, which is the primary site of the pathology. Similarly, trimethoprim and sulfamethoxazole given orally at 30 mg/kg q 12 h have anecdotally been effective in controlling placentitis and have been detected in allantoic fluid,10 but the pharmacokinetics are unknown. Adjunctive therapy for placentitis such as NSAIDs and progestins are also believed to be valuable. There is an increase in the detectable concentrations of cytokines in allantoic fluid of mares with induced placentitis,73 and these events are associated with preterm birth in women, so an attempt to ameliorate their effects may be beneficial. Many clinicians will use other progestational compounds, especially pentoxifylline. In a retrospective study of placentitis diagnosed by transrectal ultrasound alone, treatment with antimicrobials, pentoxifylline, and tocolytics resulted in an approximately 75% live foal birth rate.74 A detailed discussion of placentitis is presented in Chapter 242.
coenophialum is strategic withdrawal of mares from infected pasture or hay, 60 to 90 days prepartum, to allow time for the dam to prepare for delivery and the foal to mature. If this is not possible or practical, the administration of D2 -dopamine receptor antagonists including domperidone (1.1 mg/kg p.o., q 24 h), sulpiride (3.3 mg/kg p.o., q 24 h) or perphenazine (0.3–0.5 mg/kg p.o., q 12 h) have been used successfully, beginning on day 300 of gestation, to prevent the endocrine alterations and clinical signs of fescue toxicosis in late-term mares.39, 75 The authors prefer the use of domperidone which, unlike the other D2 antagonists, does not cross the blood–brain barrier. Therapy is often continued at least 1 to 2 weeks after delivery, to support lactation.39 If presented with a mare in dystocia that is affected with fescue toxicosis, a strong consideration should be given to hysterotomy, as manipulation and extraction can cause serious damage to the mare’s reproductive tract and may cause severe fetal asphyxia.
Umbilical cord abnormalities
There is no direct management of premature placental separation. General progestational therapies are often used, and nasal oxygen insufflation may be helpful to raise the fetal oxygenation. Restricting exercise to stall rest with hand walking or small paddock turnout may often be helpful.
There is no effective management of umbilical abnormalities prepartum, and they are rarely diagnosed. Because these abnormalities are associated with fetal loss,76 any foal with umbilical disorders should be treated as a high-risk neonate. Excessively long umbilical cords may be more prone to excessive twisting on themselves (see Figure 3.3) or around the fetal body parts with secondary vascular compromise (see Figure 3.4), while short cords may predispose to excessive intrapartum traction on the fetal membranes leading to partial premature placental detachment.
Fescue toxicosis
Hydrops of the fetal membranes
The ideal management strategy for mares consuming tall fescue infected with the endophyte Acremonium
Reports of successful management of equine fetal hydrops are rare,77, 78 especially if the mare is not near
Premature placental separation
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Figure 3.5 Ventral abdominal herniation associated with hydrallantois.
Figure 3.3 Excessively long umbilical cord with multiple twists and obvious vascular compromise.
term with a well-developed mammary gland that contains colostrum. Maintaining a hydropsical pregnancy is risky due to the danger of further trauma to the mare’s uterine and abdominal walls. If the mare is in distress and parturition is not imminent, abortion
Figure 3.4 An aborted fetus delivered with an excessively long umbilical cord wrapped around its neck, resulting in vascular compromise. The mare had sonographic evidence of hydrops amnii.
is recommended. Hydrops creates intra-abdominal hypertension and contributes to respiratory distress and the potential for hypovolemic shock during delivery. Mares are at increased risk for developing ventral body wall hernias, prepubic tendon rupture, and/or uterine rupture (see Figure 3.5). In advanced cases of hydrops, delivery should be induced to relieve pressure on the mare’s internal organs. Sudden expulsion of large volumes of fetal fluids, sometimes in excess of 100 to 200 L, results in hypotensive shock in the mare. To reduce the risk of shock, the mare should be pre-treated with intravenous crystalloids (e.g., 20 L bolus) and hypertonic saline (4 mL/kg) or other colloid (hetastarch 10 mL/kg) followed by an additional infusion of crystalloids (10–40 mL/kg) to maintain blood pressure. Controlled drainage of fetal fluids can be attempted using a 24–32 Fr trocar catheter attached to a large-bore extension set. The bulging allantochorionic membrane is identified manually per vagina and the trocar is introduced slowly through the cervix and then popped through the allantochorion. In some instances, large volumes of fetal fluids can be removed in a semi-controlled fashion prior to induction. In many mares, once the cervix and placenta are breeched, the mares often strain and push a large volume of fluid out at once. Oxytocin is then given to stimulate uterine contractions, but poor uterine tone and potential uterine inertia should be anticipated, and delivery assisted to reduce birth stress for mare and foal and minimize peripartum hypoxia. Pre-treating the cervix with prostaglandin E2, using misoprostil, may be necessary if there is inadequate cervical softening prior to induction. Delivery of the fetus is usually
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Management of the High-Risk Pregnancy not eventful as it is often small. Unless at term, fetuses should be euthanized as survival is very unlikely. Fetal abnormalities have been reported with hydrops, so any surviving neonate should be examined quickly for serious congenital defects prior to pursuing aggressive intensive care. After fetal delivery, mares should be closely monitored for signs of distress and given oxytocin to promote passage of the placenta. Retained fetal membranes are common and uterine involution is often delayed. A successful outcome requires adequate personnel prepared to assist both the mare and the foal if both are to be given optimal chance for survival. If the mare is comfortable and fetal viability is reasonable, milder cases of hydrops allantois can be supported with intravenous and oral fluids and nutritional support, laxative diets, judicious use R (0.088 mg/kg orally of NSAIDS and Regu-Mate q 24 h) to help maintain the pregnancy long enough to allow adequate fetal maturation. Frequent transabdominal fetal monitoring is recommended to monitor for fetal distress. Stall rest is advised. Since mares with hydrops are at increased risk for developing abdominal wall hernias or prepubic tendon rupture, serum creatine kinase (CK) concentrations can be monitored to help identify early signs of muscle damage. It is the author’s (WV) experience that mares recovering from hydrops, uncomplicated by abdominal wall trauma, have normal fertility after the season of occurrence and the condition is unlikely to recur in subsequent pregnancies.
Placental insufficiency This condition is often undetected prepartum. Therapy is similar to that for premature placental separation. Fetal conditions
Neonatal isoerythrolysis Prevention of neonatal isoerythrolysis is best, which requires preventing ingestion of the dam’s colostrum and providing an alternative source. Treatment of neonatal isoerythrolysis is directed at the fetus and is discussed in Chapter 35. Specific treatment of the mare is unnecessary, although the condition may be avoided in future by selecting a stallion that is compatible with the mare’s blood type.
Twins Any multiple pregnancy in the horse has the potential to be high risk; however, the risk of maternal disability is low early in pregnancy. Many twin preg-
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nancies naturally self-reduce to singletons, but if the pregnancy reaches over 90 days, the likelihood of naturally reducing to one fetus and carrying it to term is low. Most clinicians recommend not allowing mares to attempt to go to term with two live fetuses. While there are exceptions and two live foals can be born, the risk to both mare and fetuses is real, and one or both foals often suffer from varying degrees of dysmaturity and peripartum asphyxia. When owners select this option, the mares should be closely watched for signs of parturition, in case dystocia results. Elimination strategies for twin pregnancies either abort both fetuses or attempt to salvage one. Abortion of both fetuses can be performed with the use of prostaglandins early in pregnancy or the use of oxytocin later (see Chapter 238). If an attempt is made to maintain one pregnancy, there are several methods of selective fetal reduction. In early pregnancy, manual vesicle reduction can be performed and is discussed in Chapter 220. In later pregnancy, a single twin can be removed surgically,79 or injected with potassium chloride,80 or decapitated by manual methods via a laparotomy81 (see Chapters 220 and 255 for more detail). After selective fetal reduction, the use of NSAIDs and progestagens are common.
Fetal malformations and malpresentations Fetal abnormalities such as schistosomas reflexus, arthrogryposis, and hydrocephalus cannot be treated during pregnancy and are almost always fatal to the fetus. Such abnormalities are often discovered during the second stage of labor, usually because they cause dystocia, and are covered in Chapter 255.
Mare reproductive loss syndrome It appears likely that the occurrence of mare reproductive loss syndrome (MRLS) in mares is caused by exposure to the Eastern tent caterpillar (ETC), so the best management of this condition is to prevent the exposure of mares and their forage to this insect. This is accomplished by the use of appropriate insecticides, and the depletion of the caterpillars’ preferred environment. As the exact toxic principle is unknown, a recommendation about specific treatments cannot be made. The most popular theory regarding the cause of abortion due to MRLS is bacterial translocation across the mare’s intestinal mucosa secondary to focal trauma caused by the ETC setae with subsequent infection of the fetal and placental tissues.82, 83 Therefore, prophylactic administration of broad-spectrum antibiotics and progestational drugs intuitively would seem to be a reasonable treatment option for mares suspected of ingesting large numbers of ETC during gestation.
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References 1. Barber JA, Troedsson MHT. Mummified fetus in a mare. J Am Vet Med Assoc 1996;208(9):1438–40. 2. Ousey JC, Houghton E, Grainger L, Rossdale PD, Fowden AL. Progestagen profiles during the last trimester of gestation in Thoroughbred mares with normal or compromised pregnancies. Theriogenology 2005;63:1844–56. 3. Kindahl H, Daels P, Odensvik K, Daunt D, Fredricksson G, Stabenfeldt G, Hughes JP. Experimental models of endotoxemia related to abortion in the mare. J Reprod Fertil Suppl 1991;44;509–16. 4. Daels PF, Stabenfeldt GH, Hughes JP, Odensvik K, Kindahl H. Evaluation of progesterone deficiency as a cause of fetal death in mares with experimentally induced endotoxemia. Am J Vet Res 1991;52(2):282–8. 5. Santschi EM, LeBlanc MM, Weston PG. Progestogen, oestrone sulfate and cortisol concentrations in pregnant mares during medical and surgical disease. J Reprod Fertil Suppl 1991;44:627–34. 6. Morris S, Kelleman AA, Stawicki RJ, Hansen PJ, Sheerin PC, Sheerin BR, Paccamonti DL, LeBlanc MM. Transrectal ultrasonography and plasma progestin profiles identifies feto-placental compromise in mares with experimentally induced placentitis. Theriogenology 2007;67(4):681–91. 7. Daels PF, Besognet B, Hansen B, Mohammed H, Odensvik K, Kindahl H. Effect of progesterone on prostaglandin F-2 alpha secretion and outcome of pregnancy during cloprostenol-induced abortion in mares. Am J Vet Res 1996;57:1331–4. 8. LeBlanc MM, Macpherson M, Sheerin P. Ascending Placentitis: What we know about pathophysiology, diagnosis and treatment. Proc Am Assoc Equine Pract 2004;50:127–43. 9. Holtan DW. Progestin therapy in mares with pregnancy complications: necessity and efficacy. Proc Am Assoc Equine Pract 1993;39:165–6. 10. Macpherson ML. Treatment strategies for mares with placentitis. Theriogenology 2005;64:528–34. 11. Macpherson ML, Bailey CS. A clinical approach to managing the mare with placentitis. Proceedings of Society for Theriogenology 2008;70:435–40. 12. Bailey CS, Macpherson ML, Pozor MA, Troedsson MH, Benson S, Giguerre S, Sanchez LC, Le Blanc MM, Vickroy TW. Treatment efficacy of Trimethoprim sulfamethoxazole, pentoxifylline and altrenogest in experimentally induced equine placentitis. Theriogenology, 2010, April 21 (Epub ahead of print). 13. Rode L, Langhoff-Roos J, Andersson C, Dinesen J, Hammerum MS, Mohapeloa H, Tabor A. Systematic review of progesterone for the prevention of preterm birth in singleton pregnancies. Acta Obstet Gynecol Scand 2009;88(11):1180–9. 14. Lamont RF, Jaggat AN. Emerging drug therapies for preventing spontaneous preterm labor and preterm birth. Expert Opin Investig Drugs 2007: 16(3):337–45. 15. Vidaeff AC, Ramin SF. From concept to practice: the recent history of preterm delivery prevention. Part II. Subclinical infection and hormonal effects. Am J Perinatol 2006;23(2):75–84. 16. Di Renzo GC, Mattei A, Gojnic M, Gerli S. Progesterone and pregnancy. Curr Opin Obstet Gynecol 2005;17(6):598–600. 17. DaFonseca EB, Bittar RE, Carvalho MHB, Zugaib M. Prophylactic administration of progesterone by vaginal suppository to reduce the incidence of spontaneous preterm birth in women at increased risk: a randomized placebo-
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19.
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21.
22.
23.
24.
25.
26.
27.
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30.
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controlled double-blinded study. Am J Obstet Gynecol 2003;188:419. Daels PF, Mohammed HO, Odensvik K, Kindahl H. Effect of flunixin meglumine on endogenous prostaglandin F2 alpha secretion during cloprostenol-induced abortion in mares. Am J Vet Res 1995;56(12):1603–10. McGlothlin JA, Lester GD, Hansen PJ, Thomas M, Pablo L, Hawkins DL, LeBlanc MM. Alteration in uterine contractility in mares with experimentally induced placentitis. Reproduction 2004;127(1):57–66. Chavatte-Palmer P, Duchamp G, Palmer E, Ousey JC, Rossdale PD, Lombes M. Progesterone, oestrogen and glucocorticoid receptors in the uterus and mammary glands of mares from mid- to late-gestation. J Reprod Fertil Suppl 2000;5:661–72. Macpherson ML. Identification and management of the high-risk pregnancy mare. Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007; 293–304. Troedsson MHT. Myometrial activity and progestin supplementation. In: Proceedings of Society for Theriogenology, 2000; pp. 17–24. Daels PF, Stabenfeldt GH, Hughes JP, Odensvik K, Kindahl H. Effects of flunixin meglumine on endotoxin-induced prostaglandin F2α secretion during early pregnancy in mares. Am J Vet Res 1991;52(2):276–81. Silver M. Prenatal maturation, the timing of birth and how it may be regulated in domestic animals. Exp Physiol 1990;75:285–307. Ousey JC, Rossdale PD, Dudan FE. The effects of intrafetal ACTH administration on the outcome of pregnancy in the mare. Reprod Fertil Dev 1998;10:359–67. Ousey JC, Rossdale PD, Palmer L, Grainger L, Houghton E. Effects of maternally administered Depot ACTH 1–24 on fetal maturation and the timing of partuition n the mare. Equine Vet J 2000;32:489–96. Alm CC, Sullivan JJ, First NL. The effect of a corticosteroid (dexamethasone), progesterone, oestrogen and prostaglandin F2α on gestation length in normal and ovariectomized mares. J Reprod Fertil Suppl 1975;23:637– 40. Jeffcott LB, Rossdale PD. A critical review of methods for induction of parturition in the mare. Equine Vet J 1977;9:208–15. Ousey JC, Kolling M, Allen WR. The effects of maternal dexamethasone treatment on gestation length and foal maturation in Thoroughbred mares. Anim Reprod Sci 2006;94:436–8. Roberts D, Dalziel S. Antenatal corticosteroids for accelerating fetal lung maturation for women at risk of preterm birth. Cochrane Database Syst Rev 2008;3:CD004454. Mwansa-Kambafwile J, Cousens S, Hansen T, Lawn J. Antenatal steriods in preterm labour for the prevention of neonatal deaths due to complications of preterm birth. International J of Epidemiology 2010; 39:i122–i133. Milam SB, MacKay RJ, Skelley LA. Secretion of tumor necrosis factor by endotoxin-treated equine mammary exudate macrophages: effect of dexamethasone and pentoxifylline. Cornell Vet 1992;82(4):435–46. Rebello S, Macpherson M, Murchie T, LeBlanc M, Vickroy T. The detection of placental drug transfer in equine allantoic fluid. Theriogenology 2005;64:776–7 (abstr). Card CE, Wood MR. Effects of acute administration of clenbuterol on uterine tone and equine fetal and maternal heart rates. Biol Reprod Mono 1995;1:7–11.
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Management of the High-Risk Pregnancy 35. Palmer E, Chavette-Palmer P, Duchamp G, Levy I. Lack of effect for clenbuterol for delaying parturition in late pregnant mares. Theriogenology 2002;58:797–9. 36. Boening KJ, Leendertse IP. Review of 115 cases of colic in the pregnant mare. Equine Vet J 1993;25(6):518–21. 37. Erkert RS, Macallister CG. Isoxsuprine hydrochloride in the horse: a review. J Vet Pharmacol Ther 2002;25(2):81–7. 38. Putnam MR, Bransby DI, Schumacher J, Boosinger TR, Bush L, Shelby RA, Vaughan JT, Ball B, Brendemuehl JP. Effects of the fungal endophyte Acremonium coenophialum in fescue on pregnant mares and foal viability. Am J Vet Res 1991;52(12):2071–4. 39. Redmond LM, Cross DL, Strickland JR, Kennedy SW. Efficacy of domperidone and sulpiride as treatments for fescue toxicosis in horses. Am J Vet Res 1994 55(5):722–9. 40. Inci S, Ozcan OE, Kilinc K. Time-level relationship for lipid peroxidation and the protective effect of alpha-tocopherol in experimental mild and severe brain injury. Neurosurgery 1998;43:330–5. 41. Daneyemez M, Kurt E, Cosar A, Yuce E, Ide T. Methylprednisone and vitamin E therapy in perinatal hypoxic-ischemic brain damage in rats. Neuroscience 1999;92:693–7. 42. Silver M, Comline RS. Transfer of gases and metabolites in the equine placenta: a comparison with other species. J Reprod Fertil Suppl 1975;23:589–94. 43. Comline RS, Silver M. PO2 levels in the placental circulation of the mare and ewe. Nature 1968;217:76–7. 44. Wooding FBD, Fowden AL. Nutrient transfer across the equine placenta: correlation of structure and function. Equine Vet J 2006;38(2):175–83. 45. Wilson DV, Schott HC, Robinson NE, Berney CE, Eberhart SW. Response to nasopharyngeal oxygen administration in horses with lung disease. Equine Vet J 2006;38(3):219–23. 46. Dolente B. Critical peripartum disease in the mare. Vet Clin Equine 2004;20:151–65. 47. Wilkins PA. High-risk pregnancy. In: Paradis MR (ed.) Equine Neonatal Medicine. Philadelphia: Elsevier Saunders, 2006; pp. 13–29. 48. Wilkins PA, Seahorn TL. Intranasal oxygen therapy in adult horses. J Vet Emerg Crit Care 2000;10:221. 49. Silver M, Fowden AL. Uterine prostaglandin F metabolite production related to glucose availability in late pregnancy and a possible influence of diet on time of delivery in the mare. J Reprod Fertil Suppl 1982;32:511–19. 50. Jeffcott LB, Rossdale PD. A radiographic study of the fetus in late pregnancy and during foaling. J Reprod Fertil Suppl 1979;27:563–9. 51. Rossdale PD, Jeffcott LB. Problems encountered during induced foaling in Pony mares. Vet Rec 1975;97:371–2. 52. Macpherson ML, Chaffin MK, Carroll GL, Jorgensen J, Arrott C, Varner DD, Blanchard TL. Three methods of oxytocin- induced parturition and their effects on foals. J Am Vet Assoc 1997;210(6):799–803. 53. Paschen RL. Low doses of oxytocin can induce foaling at term. Equine Vet J 1980;12:85–7. 54. Ley WB, Holyoak GR. Normal prefoaling mammary secretions. In: Samper LC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders, 2007; pp. 446–51. 55. Ley WM. Pre-foaling management of the mare and induction of parturition. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 3rd edn. Philadelphia: Saunders, 1992; 664–668. 56. Jackson GM, Sharp HT, Varner MW. Cervical ripening before induction of labor: a randomized trial of prostaglandin
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58.
59.
60.
61.
62.
63.
64.
65.
66.
67.
68.
69.
70. 71.
72.
73.
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E2 gel versus low-dose oxytocin. Am J Obstet Gynecol 1994;171(14):1092–6. Rigby S, Love C, Carpenter K, Varner D, Blanchard T. Use of prostaglandin E2 to ripen the cervix of the mare prior to induction of parturition. Theriogenology 1998;50(6): 897–904. Santschi EM, Slone DE, Gronwall R, Juzwiak JS, Moll HD. Types of colic and frequency of postcolic abortion in pregnant mares: 105 cases (1984–1988). J Am Vet Med Assoc 1991;199(3):374–7. Silver M, Fowden AL. Prepartum adrenocortical maturation in the fetal foal: responses to ACTH. J Endocrinol 1994;142(3):417–25. Werners AH, Bull S, Fink-Gremmels J. Endotoxemia: a review with implications for the horse. Equine Vet J 2005;37(4):371–83. Barton MH, Parviainen A, Norton N. Polymyxin B protects horses against induced endotoxaemia in vivo. Equine Vet J 2004;36(5):397–401. Chaney KP, Holcombe SJ, LeBlanc MM, Hauptman JG, Embertson RM, Mueller PO, Beard WL. The effect of uterine torsion on mare and foal survival: a retrospective study, 1985–2005. Equine Vet J 2007;39(1):33–6. Pascoe JR, Meagher DM, Wheat JD. Surgical management of uterine torsion in the mare: a review of 26 cases. J Am Vet Med Assoc 1981;179(4):351–4. Ross J, Palmer JE, Wilkins PA. Body wall tears during late pregnancy in mares: 13 cases (1995–2006). J Am Vet Med Assoc 2008;232(2):257–61. Wilkins PA, Papich M, Sweeney RW. Pharmacokinetics of Acyclovir in adult horses. J Vet Emerg Crit Care 2005;15(3):174–8. Maxwell LK, Bentz BG, Gilliam LL, Ritchey JW, Eberle RW, Holbrook TC, Mcfarlane D, Rezabek GB, MacAllister CG, Goad CL, Allen GP. Efficacy of valacyclovir against disease following EHV-1 challenge. Proc Am Coll Vet Intern Med, 2009; p. 176. Bernard WV, Bolin C, Riddle T, Durando M, Smith BJ, Tramontin RR. Leptospiral abortion and leptospiruria in horses from the same farm. J Am Vet Med Assoc 1993;202(8):1285–6. Watkins JP, Taylor TS, Day WC, Varner DD, Schumacher J, Baird AN, Welch RD. Elective cesarean section in mares: eight cases (1980–1989). J Am Vet Med Assoc 1990;197(12):1639–45. Leadon DP, Jeffcott JB, Rossdale PD. Mammary secretions in normal, spontaneous, and induced premature parturition in the mare. Equine Vet J 1984;16:256–9. Platt H. Infection of the horse fetus. J Reprod Fertil Suppl 1975;23:605–10. Christensen BW, Roberts JF, Pozor MA, Giguere S, Sells SF, Donahue JM. Nocardioform placentitis with isolation of Amycolatopsis spp in a Florida-bred mare. J Am Vet Med Assoc 2006;228(8):1234–9. Murchie TA, Macpherson ML, LeBlanc MM, Luznar S, Vickroy TW. Continuous monitoring of penicillin G and gentamicin in allantoic fluid of pregnant pony mares by in vivo microdialysis. Equine Vet J 2006;38(6): 520–5. LeBlanc MM, Giguere S, Brauer K, Paccamonti DL, Horohov DW, Lester GD, O’Donnell LJ, Sheerin BR, Pablo L, Rodgerson DH. Premature delivery in ascending placentitis is associated with increased expression of placental cytokines and allantoic fluid prostaglandins e2 and F2α. Theriogenology 2002;58:841–4.
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74. Troedsson MHT, Zent WW. Clinical ultrasonic evaluation of the equine placenta as a method to successfully identify and treat mares with placentitis. Workshop on the equine placenta, Gluck Equine Research Foundation, 2003; pp. 66–7. 75. Brendemeuhl JP. Reproductive aspects of fescue toxicosis. In: Robinson NE (ed.) Current Therapy in Equine Medicine 4. Philadelphia: Saunders, 1997; pp. 571–3. 76. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203(8):1170–5. 77. Smith JM. Hydroallantois in the mare. Mod Equine Med 1983;2:7–12. 78. Christensen BW, Troedsson MHT, Murchie TA, Pozor MA, McPhereson ML, Estrada AH, Carillo NA, Mackay RJ, Roberts GD, Langlois J. Management of hydrops amnion in a mare resulting in birth of a live foal. J Am Vet Med Assoc 2006;228(8):1228–33.
79. Pascoe DR, Stover SM. Surgical removal of one conceptus from fifteen mares with twin concepti. Vet Surg 1989;18(2):141–5. 80. Rantanen N, Kincaid B. Ultrasound guided fetal cardiac puncture: a method of twin reduction in the mare. Proc Ann Conv Assoc Am Equine Pract 1988;29:173–9. 81. Wolfsdorf KE, Rodgerson D, Holder R. How to manually reduce twins between 60 and 120 days gestation using cranio-cervical dislocation. Proc Am Assoc Equine Pract 2005;51:284–7. 82. McDowell KJ, Webb BA, Williams NM, Donahue JM, Newman KE, Lindermann MD, Orohov DW. Invited review: the role of caterpillars in mare reproductive loss syndrome: a model for environmental causes of abortion. J Anim Sci 2010;88(4):1379–87. 83. Sebastian MM, Bernard WV, Riddle TW, Latimer CR, Fitzgerald TD, Harrison LR. Review paper: mare reproductive loss syndrome. Vet Pathol 2008;45(5):710– 22.
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Chapter 4
Ultrasonographic Monitoring of the Fetus Stefania Bucca
Introduction The equine pregnancy can be successfully explored by ultrasonography (US), from about 12 days gestation to term, by combining transrectal and transabdominal techniques. Intensive investigation of feto-placental health and development is traditionally carried out by clinicians in the field, up to 60–70 days gestation, with standard, transrectal US technique. Monitoring of the early equine pregnancy aims at preventing implantation of multiple embryonic vesicles,1 detecting early signs of impending embryonic loss, and determining fetal gender.2 Following that time, the equine pregnancy ceases to be the focus of veterinary attention until delivery time approaches, unless obvious complications arise. Multiple circumstances, related to the physiology of equine pregnancy, support this undesirable lack of supervision. The most common motivations for discontinuing pregnancy monitoring past this time are: presence of endometrial cups, limiting options of intervention; size of the gravid uterus extending over the brim of the pelvis and a disproportionate increase in the volume of fetal fluid compared with fetal mass between days 70 and 100 of gestation, which makes the fetus sonographically inaccessible per rectum and transabdominally; inadequacy of fetal visualization per rectum for appropriate assessment of fetal gender and viability; and a lower fetal loss incidence after 2 months gestation. Concurrent farm management necessities may contribute to reduced access to pregnant mares after that time, as in general mares in foal are moved to better pasture or off the farm, typically away from the farm’s veterinary examination facilities. Following a transient phase of inaccessibility,3 the equine pregnancy gradually regains entrance to the
pelvis, after 3 months gestation, making the fetus once more visible by US per rectum, at around 100 days. As gestation progresses and fetal growth provides more detailed anatomic imaging, prolonged times of US observation are needed for in-depth fetoplacental assessment, making US per rectum a rather unsuitable approach for this purpose. Percutaneous, transabdominal US allows extended periods of observation with minimal discomfort to the mare and a wide acoustic window to assess the ventrolateral aspects of the gravid uterus.4 Although the equine fetus may be visualized by this approach as early as 60 days, detailed assessment of the pregnant uterus content can realistically be obtained from 4 months gestation onwards. The most caudal aspects of the gravid uterus can only be explored, throughout gestation, by US per rectum. Indications for feto-placental US imaging include fetal gender determination,2, 5, 6 detection and management of multiple fetuses,7, 8 guiding of needle puncture for collection of fetal fluid (amnio-allantocentesis),9 and assessment of feto-placental well-being. US of the equine pregnancy in advanced gestation is a discipline of relatively recent acquisition, as most of the US parameters available nowadays have been generated by studies conducted in the last two decades. Furthermore, most of the initial studies were confined to the late gestational mare and a relative paucity of data were available to assess the equine fetus at different stages of development. The wealth of available information has increased in recent years, but remains far from being complete, compared with human medicine standards. The US aspects of basic fetal anatomy and physiology have been reported, 10–15 but subtle signs of fetal distress and, ultimately, feto-placental compromise need further imaging clarification.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Other means of assessing fetal viability include fetal electrocardiography and Doppler technology. Fetal electrocardiography allows constant monitoring of fetal cardiac activity and is still used in hospitalized mares carrying high-risk pregnancies. In recent years, the technique has been replaced by US monitoring, as the size of the mare represents a limiting factor for appropriate tracing recordings, in particular during episodes of fetal activity. Doppler technology may offer a promising approach to the identification of disrupted feto-maternal exchange pathways, particularly when cord blood flow compromise is suspected.16 Complexity of the placental circulatory network and difficulty in consistently accessing cord vascular structures pose major obstacles to the routine application of this technology. Further investigation and characterization of blood flow patterns are required. Currently, US remains the most frequently used imaging technique for feto-placental assessment and monitoring.
Examination technique Ideally, ultrasonographic examination of the gravid uterus should be carried out with the mare restrained in stocks, in quiet and relaxed surroundings. Excessive maternal anxiety may have fetal implications in terms of heart rate accelerations, sometimes persistent, with or without accompanying bouts of activity. The transient, mother-dependent nature of this type of fetal distress can be demonstrated by moving the mare to more familiar surroundings for reassessment. Sedation of mares for US assessment of fetal wellbeing should be avoided as it depresses fetal cardiac activity.17 A bucket of feed and/or hay net strategically placed in the examination room may provide sufficient diversion to allow for a full examination to take place, uneventfully. US examination of the equine pregnancy per rectum employs a standard reproductive technique and relies on linear-array or convex technology, mounted with a 5-to 7.5-MHz frequency transducer or higher. Excellent imaging can be obtained by this route, owing to the close proximity of the US probe to the gravid uterus. The narrow acoustic window obtainable allows a complete investigation of the uterine content in early pregnancy, but imaging in advanced gestation is limited to the cervix, the most caudal aspects of the utero-placental unit, and the contents of the posterior body of the uterus. Transabdominal US examination relies on adequate skin contact with the transducer for diagnostic imaging. Clipping, shaving of the ventral abdomen, and liberal application of alcohol and US coupling gel may all optimize image quality. Clipping should extend cranially as far as the umbilicus and laterally to
both flank folds, for pregnancies up to 5 months gestation. Further clipping should be obtained as gestation advances, to reach the sternum cranially and mid-flank laterally in term pregnancies. In the author’s experience, proper skin preparation requires a hard hair brush to remove dust and dirt, hair saturation with alcohol, and a generous amount of US coupling gel, over the transducer footprint, reapplied as needed. Additional clipping, shaving, and use of hair-removing creams should be restricted to mares in need of repeated assessments, mares with a substantial layer of abdominal fat or subcutaneous edema, and mares with a thick and matted coat over the ventral abdomen. Linear-array, convex, or sector technology may be used by this route; it is a matter of preference and availability. Linear-array and convex technologies offer good imaging, but transducer contact may be difficult at times; on the contrary, sectorscanner probes, with a smaller footprint, can negotiate a snug skin contact even in the most inaccessible areas of the ventrocaudal abdomen; however, some image distortion may occur. Transducers with a frequency of 3.5–5 MHz achieve scanning depths of 10–20 cm and adequate imaging of pregnancies in light breed mares up to 8 months gestation, depending on maternal and fetal size; 2–2.5 MHz probes are needed for more advanced gestations or heavier breeds4, 13, 18, 19 to obtain maximal penetration and reach scanning depths up to 25–30 cm. Finally, the ultrasonographer should be provided with comfortable seating, good eye contact with the US monitor, and free access to the mare’s abdomen within a safe environment.
Scanning technique US assessment of the equine pregnancy over 4 months gestation should always include a combined transrectal and transabdominal examination, as described above. A systematic and consistent approach should be adopted, both for completeness of examination and for comprehensive record-keeping. Rectal palpation of the equine pregnancy should be performed first to assess cervical competence and tone and presence of palpable fetal parts and to estimate fetal fluid volume/pressure and assess fetal activity. Transrectal ultrasonography visualizes the cervix; evaluates continuity and combined thickness of the utero-placental unit at the cervical pole; assesses fetal presentation and orbital size; fetal peripheral pulses and episodes of minor activity; quality and quantity of fetal fluid in the caudal compartments of the uterine body; and presence of free-floating particles (vernix) within both compartments of fetal fluids. Fetal fluid vernix consists of desquamated epithelial cells, cellular debris, and fetal waste products.
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Ultrasonographic Monitoring of the Fetus Free-floating particles are commonly seen swirling within the fetal fluids, during periods of fetal activity, stirred around by fetal parts in motion. Owing to the closer proximity to the fetal body, free-floating particles are generally more evident within the amniotic fluid. Occasionally, the cardiac area may be visualized per rectum, and fetal gender determination may be possible up to 8 months gestation, when the fetus lies in a posterior presentation. Transabdominal ultrasonography allows prolonged periods of examination for a thorough appraisal of the intrauterine milieu and for assessment of fetal behavior. Examinations lasting up to 1 hour, or even longer, may be needed when the fetus lies in a dormant phase. The latest generation of US units are lightweight and portable, equipped with multifrequency transducers, capable of storing images, and make the ideal candidate for transabdominal assessment of the equine pregnancy in the field. A comprehensive transabdominal US examination of the equine pregnancy includes: number of fetuses; identification of fetal horn; fetal presentation and position; multiple fetal heart rate (FHR) recordings, taken at rest and during activity; FHR reactivity; level of activity; depth and quality of fetal fluids; aortic diameter; depth of chest; assessment of fetal anatomy, gender, and umbilical cord; and continuity and combined thickness of the utero-placental unit. During the course of a transabdominal US examination, each half of the abdomen is scanned from the ipsilateral side, starting over the ventral midline, just in front of the mammary gland, and travelling cranially as far as the umbilicus or the sternum, depending on the stage of gestation, mare’s size, and abdomen conformation. The scan head is then moved along parasagittal planes in a similar fashion (caudal to cranial), and then over perpendicular transverse planes until the fetus is visualized and orientation is determined. The fetal chest is usually identified first, as the acoustic shadowing, produced by ribs and vertebral bodies, confers a distinctive zebra stripe effect. The cardiac silhouette is easily recognized in the proximal aspect of the thorax. Alignment of the US transducer with the fetal longitudinal plane helps in identifying presentation, whereas position is detected by locating the fetal spinal cord in relation to the mare’s pelvic bones. Left and right lateral recumbency will be identified by the proximity of the fetal gastric silhouette to the scan head, as the stomach indicates the left cranial abdomen and the transducer footprint the mare’s lower abdominal wall. Forward scanning within the fetal chest identifies the cardiac area and the aorta; FHR recordings, aortic diameter, and thoracic depth measurements may be obtained. Scanning the fetus in a caudal direction, along the sagittal plane, displays views of the diaphragm and abdominal organs and the pelvic structure, whereby
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fetal gender may be determined and the umbilical cord evaluated. Fetal activity is estimated, as fetal membranes and fluids are assessed. Several measurements are taken and assessments recorded during a single examination, for biophysical profiling and prospective interpretation of fetal variables. With the exception of fetal gender determination and puncture guiding for fetal fluid collection, identification of feto-placental compromise remains the primary objective for US investigations of the equine pregnancy. Even when multiple fetuses are detected, US helps choose the less viable fetus, in which a reduction technique is likely to be more successfully applied. Assessment of feto-placental well-being by US includes the evaluation of multiple parameters grouped as follows: adequate growth and development, appropriate levels of activity and fetal responsive patterns, and adequate environment. It is useful to have a chart or form to follow to prevent omission of certain parts of the evaluation (Figure 4.1).
Ultrasonographic findings Fetal growth and development Detailed anatomical descriptions of the equine fetus are reported in the literature for different stages of gestation.10, 11, 13, 15 Structures that have been measured sequentially to estimate fetal size include orbital11, 15, 20, 21 and aortic diameters13, 15, 21 (Table 4.1). Other parameters used to assess fetal growth include biparietal diameter, thoracic diameter, and femur length,21 but the size and orientation of the equine fetus and the bidimensional nature of ultrasound imaging make thoracic diameter and femur length not consistently measurable in late gestation. Fetal aortic diameter is significantly correlated with neonatal foal weight as well as girth and hip height in normal pregnancies,10 although fetuses found to be emaciated at delivery were thought to be normal in size, from measurements of the aorta. Abnormal aortic diameters were defined as those greater or less than four times the standard error (5.038) from the aortic diameter predicted from the weight of the pregnant term mare.13 Aortic diameter was predicted from the following regression equation (Y = 0.00912 × X + 12.46), where Y is the predicted aortic diameter and X is the pregnant mare’s weight in pounds. The high prevalence of anterior presentations in advanced gestation makes the fetal orbit easy to access by US examination per rectum (Figure 4.2). Combined measurements of orbital diameters have been reported for different stages of gestation and demonstrate a significant relationship with gestational age. Unfortunately, the eye is a poor indicator of fetal age, and orbital size
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Mare Name: ______________________
Vulva _____________________________
Owner Name: ______________________
Discharge : ________________________
Premises _____________Date_________
Caslick ___________________________
Breed ________________________
MM gland: ________________________
Age __________
Abdominal volume __________________
Parity ___________________________
Sacro-ischiatic ligament ______________
Gestation_____________________
Foalert ____________________________
Recent history______________________
TMT __________________________
Physical examination ________________ Transrectal examination: Time: ________:________ SCORE: Palpation: CX tightness ____________________ Length_______________ Volume of FF_____________/5 Fetal parts________________
US: CXP_________________________ Edema ___________________
Detachment ____________________________ ______ ↔ _Mid________
_________ ↔ _Cran_______
_________ ↔ __
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Echotexture ______________________________________________________________________________ Edema________________________________________ Content ____________________________________ Amnion _________________________ RUA ____________________________ LUA _______________________ DOPPLER ___________________ Presentation ___________________________ Orbit ____________________________ Fetal parts seen _____________________________________________________________________________ Activity ______________________________________ FF __________________________________________ Vernix: ______________________________________ Fetal carotid pulse __________________________________________________________________________ Figure 4.1 Mare feto-placental assessment. MM gland, mammary gland; TMT, treatment; CX, cervix; CXP, cervical pole; FF, fetal fluid; RUA, right uterine artery; LUA, left uterine artery; UC, umbilical cord; MUA, middle branch of uterine artery; CTUPU, combined thickness of utero-placental unit; ALL, allantoic; AMN, amniotic; AOØ, aortic diameter.
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FHR ________________________________________ AOØ__________________________ Chest___________________ Stomach______________________________ Gonad _____________________ UC __________________ Bladder_______________________________ Kidneys____________________________________________ Sexual organs _____________________________________________________________________________ Comment: ________________________________________________________________________________ _________________________________________________________________________________________ DOPPLER US (per rectum): SCORE: RUA: _____________________________________________________________________________________ LUA _____________________________________________________________________________________ MUA _____________________________________________________________________________________ Comments _________________________________________________________________________________ __________________________________________________________________________________________ Transabdominal examination: Start ________:________ End ________:________ SCORE: Presentation Time______________ Time _________________ Time _______________Time______________ Position Time__________________ Time _________________ Time _______________ Time _____________ FHR Time__________________ Time _________________ Time _______________ Time ________________ Time_______________________ Time _________________ Time ______________ Time ________________ Activity Time _______________
Time__________________ Time ______________
Time _____________ Time ________________ Time_______________________ Time _________________ Figure 4.1 (Continued)
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Activity Level:________ /3 AO Ø ___________________________________________________________________________________ Chest ___________________________________________________________________________________ Head: Orbit _________________________________________ Skull ________________________________ Primary sex organs _______________________________________________________________________ Other organs ____________________________________________________________________________ Comments
Transabdominal examination: Fetal abdomen SCORE: Stomach __________________________________________________________________________________ Intestine __________________________________________________________________________________ Gonad ___________________________________________________________________________________ Bladder __________________________________________________________________________________ UC ______________________________________________________________________________________ Kidney: L ______________________________________________ R ________________________________ Comments ________________________________________________________________________________ _________________________________________________________________________________________ _________________________________________________________________________________________ Transabdominal examination: CTUPU and FF SCORE: RCD ________________________ ALL __________________ AMN _________________ MCD _______________________ ALL _________________ AMN _________________ LCD ________________________ ALL ___________________ AMN ________________ RM _________________________ ALL ____________________ AMN ________________ Figure 4.1 (Continued)
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MM _________________________ ALL ___________________ AMN ________________ LM __________________________ ALL ___________________ AMN ________________ RCR ________________________ ALL____________________ AMN ________________ MCR ________________________ ALL ___________________ AMN ________________ LCR _________________________ ALL ___________________ AMN _______________ Hippomane ___________________ Vernix __________________ NPH_________________ Cystic structures ____________________________________________________________________________ Excessive amniotic fluid _____________________________________________________________________ Excessive allantoic fluid _____________________________________________________________________ Mammary gland: SCORE: Development: _____/3_ Waxed______________ Premature lactation _________________ Pre-colostral electrolytes: Time: Ca_______ Na_______ K________ Readiness:___________ Time: Ca_________ Na_________ K________ Readiness:_______________ Time: Ca_________ Na_________ K________ Readiness:_______________ Colostrum specific gravity ____________________________________________________________________ Comments ________________________________________________________________________________ _________________________________________________________________________________________ Discharge/cervical culture: SCORE: Collection: ___________________________ Transport medium: _______________________________ Laboratory: _________________________________ Results: No growth/Positive culture (circle one) Organism # 1: _________________ Growth: _______ Sensitivity: _____________________ Organism # 2: _________________ Growth: _______ Sensitivity: _____________________ Organism # 3: _________________ Growth: _______ Sensitivity: _____________________ Comments ________________________________________________________________________________ Laboratory report attached Figure 4.1 (Continued) Ca, Calcium; K, Potassium; LCD; Left Caudal; LCR, Left Cranial; LM, Left Mid; MCD, Mid Caudal; MCR, Mid Cranial; MM, Mid Mid; RCD, Right Caudal; Na, Sodium; RCR, Right Cranial; RM, Right Mid.
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Table 4.1 Fetal aortic and orbital diameters throughout gestation
Gestation
Aortic diameter
Sum of orbital diameters
4.43 ± 0.62 5.4 ± 0.8 7.82 ± 1.29 11.1 ± 1.75 13.93 ± 1.23 17.32 ± 1.75 20.24 ± 2.67 23.49 ± 1.9 26.83 ± 1.69 27.47 ± 0.71
20.34 ± 1.27 31.26 ± 3.75 39.76 ± 2.79 46.31 ± 3.47 52.68 ± 4.0 58.76 ± 3.99 61.68 ± 2.97 65.76 ± 3.43 69.4 ± 1.71 70.06 ± 0.11
80 70 60 50 40
Mean and SD aortic and orbital diameter values recorded from day 60 gestation to term, in normal, light breed mares.
offers only a rough estimate of trends in fetal growth20 (Figure 4.3) in advanced gestation. However, a recent study of pregnant small pony mares employed measurement of the fetal eye (vitreous body) for prediction of parturition date.22 Most anatomical structures of the head and cranial neck may be identified on transrectal scans of fetuses in anterior presentation. Microphthalmia may be diagnosed antepartum by US evaluation of the fetal orbit (Figure 4.4). The fetal external carotid artery may be visualized as it courses over the external compartment of the guttural pouch. Pulse tracings of the external carotid artery may be obtained, with M-mode or Doppler US per rectum and pulse frequency calculated. A direct correlation with FHR has been established. This technique provides a rapid field assessment of fetal viability23 (Figure 4.5). Stomach size increases as gestation advances15 (Figure 4.6). A direct correlation between stomach
Figure 4.2 Transrectal scan of a fetus in anterior presentation at 275 days gestation; orbital diameters are measured. Right of sonogram: mare’s tail.
30 20 10
27 0 27 1– 30 0 30 1– 33 0 33 1– 36 0
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1– 21
18
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91
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Figure 4.3 Sum of the fetal orbital diameters on the y-axis; gestation on x-axis.15
Figure 4.4 Transrectal scan of a term fetus in anterior presentation. Disruption of ocular architecture is evident; both eye globes were shrunken for gestational age, with a hyperechoic texture. Bilateral microphthalmus was confirmed at birth. Left of sonogram: mare’s tail.
Figure 4.5 Doppler image and spectral graph of the external carotid artery (ECA), as it courses over the lateral compartment of the guttural pouch. The internal carotid branch (ICA) is also identified. Transrectal scan of a fetus in anterior presentation at 245 days gestation. Left of sonogram: mare’s tail. (See color plate 1.)
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Figure 4.6 Transabdominal longitudinal scan of a fetus in anterior presentation and right lateral recumbency at 205 days gestation. Detail of the abdominal content. Right sonogram: mare’s tail.
size and fetal swallowing activity has been demonstrated.24, 25 Fetal swallowing represents one of the mechanisms regulating the volume of amniotic fluid and requires integrity of anatomic structures and neurological pathways of the fetal upper digestive tract. Occasionally, abnormal patterns of segmental intestinal distension may be observed. Congenital abnormalities of the gastrointestinal (GI) tract may support this condition and may be diagnosed antepartum. Severe small intestinal distension was observed in a filly fetus, from 120 days gestation to term, with congenital ileal obstruction (Figures 4.7 and 4.8) owing to the presence of an occluding membrane. Additional studies are needed to further characterize the US appearance of the GI tract in the equine fetus. Fetal bladder measurements should be obtained, particularly when mechanical obstruction of the umbilical flow is suspected within the amniotic portion of
Figure 4.7 Transrectal scan of a fetus in anterior presentation at 120 days gestation. Several dilated loops of intestine are visible within the abdomen. Left of sonogram: mare’s tail.
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Figure 4.8 Transabdominal, longitudinal scan of a term fetus (see Figure 4.4) in anterior presentation, presenting severe dilatation of the small intestine. Relevant structures (from right to left): fetal chest, diaphragmatic curvature, cross-sectional and longitudinal views of small intestinal loops (arrow), with swirling content; the gastric profile can be identified on the left lower corner of the sonogram. Right of sonogram: mare’s tail.
the cord (Figure 4.9). Ossified or mineralized remnants of the yolk sac may be detected from early gestation to term, when the allantoic portion of the cord is identified47 (Figure 4.10). Generally, the US appearance is of a round, hollow structure, with hyperechoic walls, 5–10 cm in diameter, attached to the cord by a stalk. On rare occasions, these masses may compromise blood flow through the cord and result in abortion. Chronic bladder distension may be associated with umbilical cord pathology. The author has observed the condition to occur more notably in the colt fetus, where a longer urethra may further
Figure 4.9 Transabdominal scan of a 7-month-old female fetus in posterior presentation. The sonogram displays views of the caudal abdomen, pelvis, and pelvic limbs. Relevant structures: gonad upper left; bladder and umbilical arteries in crosssectional view on the caudal border of the bladder (RUA, right umbilical artery; LUA, left umbilical artery). Right of sonogram: mare’s tail.
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Figure 4.10 Transabdominal scan at 264 days gestation, displaying views of the umbilical cord with attached remnant of the yolk sac in its allantoic portion.
Figure 4.12 Transabdominal scan of the cardiac area of an 8month-old fetus in transverse presentation. An M-mode tracing was obtained at rest and FHR calculated. Right of sonogram: mare’s tail.
Fetal viability: cardiac function, activity, and responsive patterns delay urine discharge. The urethra is clearly identified as it courses over the male perineum and penile shaft, and primary sexual organs and fetal gonads may be visualized from 100 days up to 8 months gestation.5, 6, 11 The fetal scrotum displays a distinctive composite echodensity, with two hypoechoic, oval-shaped, scrotal compartments, attributable to the location of paired gubernaculums testis (Figure 4.11). Male fetal gender determination can easily and indisputably be accomplished when US imaging of the perineal urethra and scrotum are obtained at their meeting point.
The most significant US parameters indicating fetal viability include FHR and activity. US assessment of FHR and rhythm has been described by Reef et al.13, 14 (Figure 4.12). FHR decreases as gestational age progresses and accelerations occur in response to activity (FHR reactivity). Mean FHR values, at rest and post activity, have been reported for different gestational ages.11, 13, 15 (Table 4.2). Unusually low heart rates, below 50 beats/min, may occasionally be observed in term fetuses and may not be cause for alarm, providing they represent short-lived episodes. Transient elevation of FHR >120 beats/min, during the last weeks of pregnancy, is not threatening either, unless it fails to return to baseline. Maternal anxiety or pain may sometimes be the underlying cause. Abnormal patterns of FHR reactivity represent the most sensitive indicator of fetal compromise. Abnormal FHR patterns include persistent fetal tachycardia, bradycardia, and cardiac arrhythmias. Table 4.2 Fetal heart rate throughout gestation Gestation
Figure 4.11 Transabdominal scan of a male fetus in posterior presentation at 205 days gestation. The sonogram shows views of the pelvis, perineum, and fetal buttocks. Relevant structures: the pelvic urethra (right of sonogram), as it courses between the buttocks; the scrotal compartments, as a pair of hypoechoic, oval, hypoechoic structure. Right of sonogram: mare’s tail.
Day 60–90 Day 91–120 Day 121–150 Day 151–180 Day 181–210 Day 211–240 Day 241–270 Day 271–300 Day 301–330 Day 330–360
FHR at rest
FHR during activity
135.1 ± 6.35 124.53 ± 7.24 117.47 ± 5.62 109.8 ± 5.58 104.11 ± 6.14 99.65 ± 6.82 88.18 ± 11.42 75.9 ± 7.44 67.25 ± 11.66
176.71 ± 18.33 155.22 ± 11.18 142.72 ± 8.52 139.79 ± 11.72 129.68 ± 7.08 125.15 ± 11.54 130.29 ± 13.66 111.86 ± 13.53 97.35 ± 13.62 87.8 ± 13.31
Fetal heart rate (FHR) recordings at rest and during activity taken from light breed mares throughout gestation
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Ultrasonographic Monitoring of the Fetus Fetal tachycardia has been observed preceding abortion14, 26 and stillbirth.10, 14 It has also been documented during abnormal deliveries.14, 25, 26 Fetal bradycardia that is inappropriate for gestational age has also been detected,4, 10, 14, 26 associated with a negative outcome. It is regarded, by some investigators, as the most reliable indicator of impending fetal demise.27 Fetal asystole ultimately identifies fetal demise.7, 14 Cardiac accelerations in response to fetal activity have been recorded using fetal ultrasonography, electrocardiography, and Doppler transabdominal US.7, 10, 12 Spontaneous fetal activity generally resulted in accelerations of FHR of 25–40 beats/min in amplitude and 20–40 seconds in duration10 Equine fetal movements without FHR accelerations were observed routinely, while accelerations in the absence of detectable stimuli occurred only 5% of the time.10, 13 Stress testing has also been performed in the pregnant mare by administration of oxytocin or the use of a vibroacoustic device placed in close proximity to the fetal head, during rectal palpation.28 Preliminary findings suggest that stress testing may be of use in the assessment of fetal well-being, but further studies are needed to determine whether it can be used as an effective means of discriminating between the normal and compromised fetus. Fetal activity is a reflection of central nervous system (CNS) function and development; depression of CNS function results in a decreased level of activity. Fetal mobility is at its highest early in gestation, when fetal mass is relatively small compared with fluid volume, providing a great freedom of movement, restricted only by the length of the umbilical cord. As gestation progresses, the space available decreases, but fetal activity becomes more complex and articulated. From 4 months of gestation, the range of movement observed includes flexion and extension of extremities, rotations about the short (spinning) and the long (rolling) axis and vertical and horizontal whole body shifting (translation).4, 10,13–15 Rolling is detected when changes in fetal position are identified. The resting equine fetus may be found in the dorso-pubic position, but more commonly appears to lie in left or right lateral recumbency, within the uterus. Scanning over the fetal abdomen may discriminate fetal position, by identifying which half of the abdomen lies closer to the US transducer. The gastric silhouette identifies the left half of the abdomen. Spinning occurs when a change of presentation is detected. Presentation may change up to 9 months gestation.15, 29 The normal size equine fetus is unable to rotate about its short axis in advanced gestation. Spinning may occur in late gestation, only when small, usually compromised, fetuses are present and fetal fluids are plentiful; most
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commonly this condition occurs when hydroallantois is diagnosed. Fine-tuned movements of the head, tail, and extremities may be observed from midto late gestation, including ear twitching, blinking, nostril flaring, and suckling activity. Rhythmical breathing movements may also be detected from 8 months gestation to term,13, 15 when synchronous excursion of the diaphragm and thoracic walls are demonstrated. Breathing activity is intermittent and is not consistently detected. Fetal movements are required to ensure satisfactory muscular development and the proper function of skeletal structures. The equine neonate is particularly dependent on a high level of development of these anatomical parts to ensure successful postnatal adaptation. Recording of fetal activity may be reported as a subjective impression of level of activity, scored 1–3,14 or as the number of major and finetuned movements detected in time.15 The most vigorous episodes of fetal activity and exaggerated FHR responses reportedly occur 48–72 hours prior to parturition and may arise in response to early oxytocin release. Dormant (inactive) phases are observed in fetuses of all ages, but appear to be more frequent in late gestation, when they may last ≤10 minutes.13, 30 However, 30–60 minutes or longer dormant phases have been observed on occasion.30 Therefore, caution should be exercised in interpreting fetal activity and reassessments are advisable. During a resting phase, in advanced gestation, most of the fetal trunk frequently shifts towards the cranial abdomen, gently curved just caudal to the dam’s diaphragm. Under those circumstances, attempts to arouse the fetus by ballottement of the gravid uterus per rectum may be unsuccessful. The equine fetus is usually very active and long periods of true inactivity should be regarded as significant. Lack of fetal movements has been associated with a negative outcome in a number of studies.4, 10, 12, 13, 31 However, sudden bouts of excessive activity followed by abrupt cessation have also been recorded in fetuses that subsequently died. 31 The detection of either prolonged inactivity or hyperactivity should be regarded as suspicious for fetal compromise and may suggest poor fetal outcome. Fetal environment Assessment of fetal environment includes evaluation of the utero-placental unit and fetal fluids. The average combined thickness of the utero-placental unit has been reported for different stages of gestation32, 33 and different areas of the mare’s ventral abdomen (Table 4.3). Thickness should be assessed only on those areas showing no contact with the fetus. Measurements of the utero-placental unit differ significantly between fetal and non-fetal horn, with the
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Table 4.3 Mean values and SD of utero-placental unit combined thickness taken at nine different sections of the mare’s ventral abdomen Gestation (days)
LCD MCD RCD LMD MMD RMD LCR MCR RCR
150–180
181–210
211–240
241–270
271–300
301–330
331–360
7.08 ± 2.5 5.2 ± 1.04 5.6 ± 2.24 5.6 ± 1.1 5.1 ± 1.0 5.17 ± 0.8 6.0 ± 0.9 3.5 ± 0.6 2.9 ± 1.2
6.1 ± 2.0 6.57 ± 1.38 6.22 ± 1.65 6.25 ± 1.61 5.07 ± 1.49 5.15 ± 1.55 5.02 ± 1.11 5.17 ± 1.65 5.06 ± 0.8
7.29 ± 1.6 7.97 ± 2.34 7.8 ± 1.68 6.98 ± 1.24 6.46 ± 0.5 5.98 ± 1.39 7.08 ± 2.5 7.84 ± 2.5 6.01 ± 1.8
7.33 ± 2.23 8.07 ± 1.47 8.22 ± 1.86 7.73 ± 2.79 7.48 ± 2.1 5.85 ± 1.69 7.08 ± 2.5 5.86 ± 1.15 6.8 ± 2.5
7.67 ± 1.63 7.8 ± 2.0 7.22 ± 1.28 8.66 ± 3.37 8.7 ± 3.37 7.6 ± 3.43 7.08 ± 2.5 6.7 ± 2.4 7.2 ± 3.8
10.0 ± 5.14 9.01 ± 2.37 9.34 ± 2.91 9.89 ± 4.01 9.44 ± 3.37 7.06 ± 2.73 7.08 ± 2.5 6.75 ± 2.3 7.53 ± 2.99
10.1 ± 4.39 10.75 ± 5.9 9.5 ± 4.8 10.7 ± 3.5 11.1 ± 5.56 7.58 ± 3.8 7.08 ± 2.5 5.95 ± 1.66 6.38 ± 2.76
LCD, left caudal; MCD, mid-caudal; RCD, right caudal; LM, left mid; MM, mid-mid; RM, right mid; LCR, left cranial; MCR, mid-cranial; RCR, right cranial.
latter displaying a highly corrugated surface and a much greater thickness. Combined thickness of the utero-placental unit at the cervical pole has been the focus of attention in recent years, as abnormalities detected in this area may identify early signs of ascending placentitis. Specific measurements have been reported, for different gestational stages. Renaudin et al.32 reported combined measurements of 4 mm between 4 and 9 months, with progressive increases of 1.5–2 mm for each consecutive month. The values recently reported by Colon34 for the combined uteroplacental thickness appear to be considerably more elevated than Renaudin’s findings, probably due to the greater resolution and imaging capacity of later technology. Measurements taken in this area should be carefully evaluated and interpreted with caution, as forward shifting of the fetus and fetal fluids over
the pelvis may cause transient increases in thickness and the appearance of a corrugated pattern, especially in older mares, with a pendulous abdomen. Furthermore, allanto-chorial edema may frequently be observed in this area in term pregnancies close to delivery and may persist for days, without any cause for concern15, 34 (Figure 4.13a,b). Abnormally thick or thin utero-placental units have been associated with a poor fetal outcome. However, not all mares at risk for premature labor exhibit changes in the ultrasonographic appearance of a diseased placenta. Sheerin and co-workers35 demonstrated that only 60% of experimentally infected mares had ultrasonographic evidence of placental thickening or separation before delivery. Large areas, where utero-placental contact is lost, are also associated with a negative outcome (Figure 4.14). Under normal circumstances, no
(a)
(b)
Figure 4.13 Transrectal scans of the utero-placental unit at the cervical pole at different times of the day (a,b). The two sonograms show the variability in aspect and thickness of the utero-placental unit at the cervical pole, according to fluid dynamics and proximity of fetal parts.
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Figure 4.14 Transrectal scan of the cervical pole of a term mare with ascending placentitis, displaying a large area of allantochorial (AC) detachment. UT, uterine wall.
structural distinction should be detected on ultrasonography between allantochorion and uterine wall. The need for adequate utero-placental contact is highlighted by the difficulty mares have in maintaining pregnancies involving multiple fetuses. However, the extent of endometrio-placental contact required to ensure fetal survival to term is not known and cannot be appreciated in vivo. Large and/or progressively enlarging areas of placental separation, as observed by Reef et al.,14 may lead to inefficient fetomaternal exchange pathways and adversely affect fetal well-being. Fetal fluid depth and quality are also evaluated; the two fluid compartments are separated by the amnion and fetal urine is voided completely into the allantoic side.36 Reef et al.13 reported minimal and maximal allantoic fluid depths of 47 and 221 mm respectively, and 8 and 149 mm were considered normal values, by the same authors, for minimal and maximal amniotic fluid depths in term mares. Pathological increases in amniotic fluid have been reported.37, 38 Markedly reduced amounts of amniotic fluid have also been observed in mares with severe systemic illness.39 Excessive accumulation of allantoic fluid has been recognized in several instances. 39–41 Reduced amounts of allantoic fluid have also been observed in mares, associated with a poor fetal outcome. The condition could be sustained by circumstances similar to those responsible for oligohydramnios in human pregnancies. Chronic hypoxia decreases renal perfusion and lower volumes of urine are produced by the fetal kidney. In the equine pregnancy, fetal urine is voided directly into the allantois, by means of the urachus; hence, a lower urine output may influence total volume of allantoic fluid. Freefloating particles (vernix) may be detected within both fetal fluid compartments, from mid-gestation
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to term. Amniotic fluid has been described to originate from amniotic epithelium, saliva, and nasopharyngeal secretions of the fetus.42, 43 As pregnancy advances, the equine fetal skin releases desquamated cells and debris, which increase the cloudiness of the amniotic fluid. The free-floating particles observed in allantoic fluid may reflect fetal kidney function, with a high concentration of calcium in the allantoic fluid.44 The hippomane, frequently identified as free-floating, multilayered, allantoic structures, is considered to be an allantoic calculus, made up of urine sediment and proteinaceous material. Its significance and purpose are not fully understood. The echogenicity of fetal fluids increases with fetal activity, as sedimented particles are actively stirred around by fetal parts in motion. This accounts for the greater echogenicity of amniotic versus allantoic fluid, as previously reported,32 owing to the closer proximity of amniotic fluid to the fetus. Generally, amniotic free-floating particles are more clearly visualized transrectally, as larger volumes of amniotic fluid tend to accumulate around the cranial parts of the fetus. Passage of meconium within the amniotic fluid or presence of blood or inflammatory debris may occasionally be observed in severely compromised pregnancies, with marked increase of fetal fluid echogenicity Biophysical profile A scoring system evaluating the biophysical parameters described above has been designed to objectively identify fetal compromise resulting from acute or chronic insults. A fetal biophysical profile was originally developed by Adams Brendemuehl and Pipers,10 adapted from a human model, for use in late gestational mares. The initial five-parameter scoring system was later revised and integrated with additional traits.14 A dynamic US technique has also been proposed, employing provocative testing, for a more comprehensive evaluation of fetal responses.45 Parameters used in the revised scoring system include: FHR, aortic diameter, level of activity, uteroplacental thickness and contact, and maximal allantoic fluid depth (Table 4.4). Each parameter is given a score value from 0 to 2. Normalcy scores for these six variables are summed to give a biophysical profile value that ranges from 0 to 12. A score of 8 or less ensures a negative fetal outcome, but a maximal score does not guarantee a positive result,14 because unpredictable periparturient events may occur. Applicability of this scoring system throughout gestation requires validation. Further studies should be carried out to enhance the predictive value of a potentially very useful tool for the assessment of feto-placental well-being.
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Foal: Prepartum Assessment of the Foal Table 4.4 Equine biophysical profile (EBP)14 Parameter
Abnormal
∗
Fetal heart rate (FHR) Rhythm Low FHR < 320 days gestation (bpm) Low FHR 320–360 days gestation (bpm) Low FHR > 360 days gestation (bpm) High FHR (post-activity) (bpm) FHR range ∗ Fetal aortic diameter Y = 0.00912 × X +12.46 ∗ Fetal activity and tone Fetal activity Fetal tone ∗ Fetal fluid depths Min/maximal allantoic depth (mm) Min/maximal amniotic depth (mm) ∗ Utero-placental thickness Uterus + chorioallantois (mm) ∗ Utero-placental contact Areas of discontinuity
Irregular or absent <57 <50 <41 >126 >50 or <5 Y ± 4 × SE (5.038)
> or
221 <8 or >185 <3.9 or >2.1 Large
To calculate EBP assign 2 points to each category marked with an (∗ ) if all evaluations are normal, assign 0 points at each category if one of the evaluations is abnormal. Y, predicted aortic diameter; X, pregnant mare’s weight in pounds; SE, standard error.
Applications Pregnant mares experiencing medical or surgical conditions during the course of gestation may pose a threat to fetal or neonatal well-being. The opposite may also be true, as a compromised or dead fetus may complicate the clinical course of a critically ill dam and pregnancy termination maybe desirable. Identification of pregnancies carrying an elevated risk of an unfavorable outcome prompts close supervision of maternal parameters and intensive monitoring of feto-placental traits. Pregnant mares with a history of previous complicated gestations warrant supervision too, as some problems tend to recur in subsequent pregnancies. A history of abortion, placental pathology, recurrent dystocia, abnormal neonate, uterine artery hemorrhage, or premature birth should be investigated at appropriate times during the course of gestation, to allow for corrective strategies to be implemented. Mares with a remote reproductive history of ascending placentitis should be scanned at regular intervals from 4–5 months gestation and transrectal US parameters of the continuity and combined thickness of utero-placental unit recorded.33, 34, 46 Association of hormonal profiling for plasma progestin assessment may add further diagnostic and prognostic value to the US technique, prior to day 300 of gestation.48 Management of a high-risk pregnancy requires a team effort in most instances,
and the desires of the owner should be made clear, in terms of preferential survival of the dam or the foal. Upon initial diagnosis, whereby a comprehensive ultrasonographic feto-placental database is collected, a plan on sequential examinations over time is made, to allow the clinician to identify changes as they occur and assess response to treatment. The frequency of examination depends on the severity of the initial condition and acuteness of feto-placental compromise. Vital fetal parameters will need to be evaluated several times daily in advanced gestation and term mares, when premature placental separation is suspected, a uterine torsion is detected, pre-foaling hemorrhage into the broad ligament is diagnosed, or a ventral body wall tear is identified, particularly if signs of premature lactation are present. Initially, fetal hyperactivity, tachycardia, and an increased range of heart rate reactivity are almost invariably present, indicating acute fetal distress. Intensive US monitoring, on a 24-hour-basis will recognize sudden fetal demise or rapidly declining fetal parameters, becoming an essential tool when elective induction of parturition or cesarean section are available options. Frequent determination of colostral electrolyte concentrations should be performed throughout the 24 hours to identify fetal readiness for birth.49 The equine fetus is extremely dependent on maternal oxygen and glucose for survival and acute oxygen and/or glucose deprivation may rapidly
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Ultrasonographic Monitoring of the Fetus induce fetal compromise. Acute and chronic fasting should always be avoided and, when mares become acutely anorectic, glucose should be provided intravenously to support fetal energy balance and prevent placental production of prostaglandins and the risk of preterm delivery. The risk of preterm delivery increases within one week of an anorectic episode. Maternal intranasal oxygen insufflation should be considered in all conditions where reduced oxygen delivery to the fetus is suspected. Uterine torsion, decreased placental perfusion, placental dysfunction, maternal hypoxemia, maternal anemia, and hypovolemia may all result in various degrees of fetal oxygen deprivation and potentially predispose the fetus to disease conditions caused by hypoxic–ischemic asphyxia. Beneficial effects of maternal glucose administration and oxygen insufflation may be observed ultrasonographically in terms of increased levels of fetal activity and appropriate fetal heart rate reactivity. Life-threatening maternal medical and surgical conditions will also require frequent examinations. In compromised pregnancies in which clinical signs of early delivery do not regress with treatment, administration of exogenous corticosteroid therapy may be beneficial in advancing fetal maturity and increasing the chances of neonatal survival.50
10.
11.
12.
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17.
References 1. Jeffcott LB, Whitwell KE. Twinning as a cause of foetal and neonatal loss in the thoroughbred mare. J Comp Pathol 1973;83:91–105. 2. Curran S, Ginther OJ. Ultrasonic fetal gender diagnosis during months 5 to 11 in mares. Theriogenology 1993;40:1127–35. 3. Ginther OJ. Equine pregnancy: physical interactions between the uterus and conceptus. In: Proceedings of the 44th Annual Convention of the American Association of Equine Practitioners, 1998, pp. 73–103. 4. Pipers FS, Adams Brendemuehl CS. Techniques and application of transabdominal ultrasonography in the pregnant mare. J Am Vet Med Assoc 1984;185:766–71. 5. Holder RD. Fetal sex determination in the mare between 55 and 150 days gestation. In: Proceedings of the 47th Annual Convention of the American Association of Equine Practitioners, 2001, pp. 321–4. 6. Bucca S. Fetal gender determination from mid to advanced gestation by ultrasound. Theriogenology 2005;64:558–63. 7. Rantanen NW, Kincaid B. Ultrasound guided fetal cardiac puncture: a method of twin reduction in the mare. In: Proceedings of the 34th Annual Convention of the American Association of Equine Practitioners, 1988, pp. 173–9. 8. Wolfsdorf KE, Rodgerson D, Holder R. How to manually reduce twins between 60 and 120 days gestation using cranio-cervical dislocation. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners, 2005, pp. 284–7. 9. Holdstock NB, McGladdery AJ, Ousey JC, Rossdale PD. Assessing methods of collection and changes in selected biochemical constituents of amniotic and allan-
18. 19.
20.
21. 22.
23.
24.
25. 26. 27.
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toic fluid throughout equine pregnancy. Biol Reprod Mono 1995;1:21–38. Adams Brendemuehl CS, Pipers FS. Antepartum evaluation of the equine foetus. J Reprod Fertil 1987;35(Suppl): 565–73. Renaudin CD, Gillis CL, Tarantal AF, Coleman DA. Ultrasonographic evaluation of equine fetal growth from 100 days gestation to parturition. In: Proceedings of the 7th International Symposium on Equine Reproduction, 1998, pp. 171–2. Adams Brendemuehl CS. Fetal assessment. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 16–33. Reef VB, Vaala WE, Worth LT, Spencer PL, Hammett B. Ultrasonographic evaluation of the foetus and intrauterine environment in healthy mares during late gestation. Vet Radiol Ultrasound 1995;36:533–41. Reef VB, Vaala WE, Worth LT, Sertich PA, Spencer PL. Ultrasonographic assessment of fetal well-being during late gestation: development of a biophysical profile. Equine Vet J 1996;28:200–8. Bucca S, Fogarty U, Collins A, Small V. Assessment of foeto-placental well-being in the mare from mid-gestation to term: transrectal and trans-abdominal ultrasonographic features. Theriogenology 2005;64; 542–57. McGladdery AL, Ousey JC, Rossdale PD. Serial Doppler ultrasound studies of the umbilical artery during equine pregnancy. In: Proceedings of the 3rd Conference of the International Veterinary Perinatology Society, 1993, p. 37. McGladdery AJ, Cottrill CM, Rossdale PD. Abstract. Effects upon the foetus of sedative drugs administered to the mare. In: Rossdale PD (ed.) Proceedings of the Second International Conference on Veterinary Perinatology, Cambridge, UK: International Society of Veterinary Perinatology, 1990, p. 14. Reef VB. The use of diagnostic ultrasound in the horse. Ultrasound Q 1991;9:1–34. Allen KA, Stone LR. Equine diagnostic ultrasonography: equipment selection and use. Comp Equine Educ Pract Vet 1990;12:1307–11. McKinnon AO, Voss JL, Squires EL, Carnevale EM. Diagnostic ultrasonography. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993, pp. 266–302. Pantaleon LG, Bain FT, Zent W, Powell DG. Equine fetal growth and development. Comp Vet Educ 2003;25:470–6. Turner RM, McDonnell SM, Feit EM, Grogan EH, Foglia R. Real-time ultrasound measure of the fetal eye (vitreous body)for prediction of parturition date in small ponies. Theriogenology 2006;66:331–7. Bucca S, Carli A, Fogarty U. How to assess fetal viability by transrectal ultrasound evaluation of fetal peripheral pulses. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2001, pp. 321–4. Wintour AM, Barnes A, Brown AJ, Hardy KJ, Horacek I, McDougall JM, Scoggins BA. The role of deglutition in the production of hydramnios. Theriogenology 1977;8: 160. Bucca S, Romano G. Un caso di idramnios in una fattrice in gestazione avanzata. Ippologia 2000;11:35–8. Yamamoto K, Yasuda J, Too K. Arrhythmias in the newborn thoroughbred foals. Equine Vet J 1992;24:169–73. Colles CM, Parkes RD, May CJ. Foetal electrocardiography in the mare. Equine Vet J 1978;10:32–7.
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28. McGladdery AJ, Rossdale PD. Response of the equine foetus to maternal drug administration and acoustic stimulation. In: Proceedings of the 37th Annual Convention of the American Association of Equine Practitioners, 1991, pp. 223–8. 29. Ginther OJ, Griffin PG. Equine fetal kinetics; presentation and location. Theriogenology 1993;40:1–11. 30. Fraser AF, Keith NW, Hastie H. Summarized observations on the ultrasonic detection of pregnancy and foetal life in the mare. Vet Rec 1973;92:20–1. 31. Fraser AF, Hastie H, Callicott RB, Brownlie S. An exploratory ultrasonic study on quantitative foetal kinesis in the horse. Appl Anim Ethol 1975;1:395–489. 32. Renaudin CD, Troedsson MHT, Gillis CL, King VL, Bodena A. Ultrasonographic evaluation of the equine placenta by transrectal and transabdominal approach in the normal pregnant mare. Theriogenology 1997;47:559–73. 33. Troedsson MHT, Renaudin CD, Zent WW, Steiner JV. Transrectal ultrasonography of the placenta in normal mares and mares with pending abortion: a field study. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 256–8. 34. Colon JL. Trans-rectal ultrasonographic appearance of abnormal combined utero-placental thickness in late-term gestation and its incidence during routine survey in a population of Thoroughbred mares. In: Proceedings of the 54th Annual Convention of the American Association of Equine Practitioners, 2008, pp. 279–85. 35. Sheerin PC, Morris S, Kellerman A. Diagnostic efficiency of transrectal ultrasonography and plasma progestins profiles in identifying mares at risk of premature delivery. In: Proceedings of the Focus on Equine Reproduction Meeting, 2003, pp. 22–3. 36. Arthur GH. The foetal fluids in domestic animals. J Reprod Fertil Suppl 1969;9:45–52. 37. Allen WE. Two cases of abnormal equine pregnancy associated with excess foetal fluid. Equine Vet J 1986;18: 220–2. 38. Sertich PL, Reef VB, Oristaglio-Turner RM, Habecker PL, Maxson AD. Hydrops amnii in a mare. J Am Vet Med Assoc 1994;204:1481–2.
39. Reimer JM. Use of transcutaneous ultrasonography in complicated latter-middle to late gestation on pregnancies in the mare: 122 cases. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 259–61. 40. Vandeplassche M, Bouters R, Spincemaille M, Bonte P. Dropsy of the foetal sacs in mares: induced and spontaneous abortions. Vet Rec 1976;99:67–9. 41. Oppen TV, Bartmann CP. Two cases of hydroallantois in the mare. Pferdeheilkunde 2001;17:593–6. 42. Verpoest MJ, Seelen JC, Westerman CF. Changes in the appearance of the amniotic fluid during pregnancy: the macroscore. J Perinat Med 1976;4:12–25. 43. Vaala WE, Sertich PL. Management strategies for mares at risk for periparturient complications. Vet Clin North Am Equine Pract 1994;10:237–65. 44. . Jainudeen MR, Hafez ESE. Gestation, prenatal physiology and parturition. In: Hafez ESE (ed.) Reproduction in Farm Animals, 6th edn. Philadelphia: Lea & Febiger, 1993, pp. 213–36. 45. Smith LJ, Schott H. Abstract. Xylazine induced foetal bradycardia. In: Rossdale PD (ed.) Proceedings of the 2nd International Conference on Veterinary Perinatology, Cambridge, UK: International Society of Veterinary Perinatology, 1990, p. 36. 46. Renaudin CD, Liu IKM, Troedsson MT, Schrenzel MD. Transrectal ultrasonographic diagnosis of ascending placentitis in the mare: a report of two cases. Equine Vet Educ 1999;11:75. 47. Cassar TIY, Fallon LH, Martinez EH, Schlafer DH. Segmental ossification of involuted yolk sacs in equine umbilical cords. Anim Reprod Sci 2006;94:439–42. 48. LeBlanc MM, McPherson M, Sheerin PC. Ascending placentitis: what we know about pathophysiology, diagnosis and treatment. In: Proceedings of the 50th Annual Convention of the Annual American Association of Equine Practitioners, 2004, pp. 127–43. 49. Paccamonti DL. Milk electrolytes and induction of parturition. Pferdeheilkunde 2001;17:616–18. 50. Ousey JC. Hormone profiles and treatments in the late pregnant mare. Vet Clin North Am Equine Pract 2006;22:727–47.
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Chapter 5
Sedation and Anesthesia of the Pregnant Mare John A.E. Hubbell
Anesthesia of the horse carries the greatest risk of all of the common domestic species with incidences of mortality placed as high as 4.3% in horses undergoing emergency procedures for colic.1, 2 The source of this increased incidence of mortality is multifactorial and includes temperament, the cardiopulmonary changes associated with anesthesia and recumbency, and the horse’s apparent primordial need to regain its feet rapidly after anesthesia. Pregnancy, particularly in its advanced stages, can cause exacerbation of the cardiopulmonary changes associated with anesthesia and recumbency primarily because of decreases in respiratory reserve due to cranial displacement of the diaphragm. The health of the fetus can be a primary or secondary consideration when anesthetizing the mare. The greatest risks to the mare and the fetus are maternal hypotension and hypoxia. Hypotension (mean arterial blood pressures less than 60 mmHg) occurs in over 50% of anesthetized horses.3 In the author’s practice, approximately 15% of horses anesthetized and placed in dorsal recumbency have arterial partial pressures of oxygen below 100 mmHg, approaching hypoxemia. Thus, pregnant mares are prone to hypotension and hypoxia at any stage of pregnancy and steps to minimize the incidence and duration of hypotension and hypoxemia should be taken. Historically, much discussion has occurred with regard to the optimal timing for ‘elective’ surgery during pregnancy. The first trimester has been thought to be best avoided because of the potential for teratogenesis but no link to any commonly used anesthetic drug has been established.4 Most have suggested that the ‘middle’ trimester (4 to 7 months) presents the least risk although there is little supportive evidence. Third-trimester anesthesia may pose
the greatest risk because the risk of hypoxemia increases as the belly becomes rounded.5 Despite the noted risks, the equine fetus is remarkably resilient. Horses requiring exploratory abdominal surgery due to symptoms of colic are among the highest-risk patients anesthetized. In one retrospective study, 80% of mares surviving exploratory surgery for colic delivered a live foal.6 Neither the stage of pregnancy nor the duration of anesthesia had an effect on outcome but endotoxemia and hypoxia in mares anesthetized during the last 60 days of pregnancy were associated with abortion or a compromised foal at birth.6 In another study, 30% of pregnant mares presented for colic either aborted or delivered a dead foal.7 Pregnant mares requiring surgery were 3.5 times more likely than those treated medically to abort or deliver a dead foal.7 Additional factors increasing risk included anesthetic durations of greater than 3 hours and hypotension during the anesthetic period.7 The reported incidence of abortion in Thoroughbred mares due to any cause in Australia was 19%, thus the risk of anesthesia would not appear to be extraordinary.8 The total number of equine pregnancies in a given year is not recorded, thus it is impossible to determine either the percentage of pregnant mares requiring anesthesia yearly or the incidence of complications. In a large study of pregnant Swedish women approximately 0.75% required surgery for non-obstetrical reasons during their pregnancy.9 Anesthesia and surgery were not associated with an increased incidence of congenital abnormalities but there was an increased incidence of low birth weight (7.47% compared to 5.13% in the general population) and early infant mortality.9
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Prepartum Assessment of the Foal
The effect of pregnancy on anesthesia The components of the physiological response to pregnancy that affect anesthesia include the presence of the enlarging fetus in the abdomen, the hormonal changes that result from pregnancy, and the changes in serum electrolytes associated with parturition and the onset of lactation. Other events associated with parturition (exhaustion, hemorrhage) may further complicate the presentation. The presence of the fetus in the abdomen does not become a factor in the anesthetic management of the mare until the period of rapid fetal growth in the last months of gestation. While the author could find no studies on the cardiopulmonary effects of pregnancy in the mare, there are a number of references from other species, particularly women.10, 11 Tidal volume and minute ventilation increase early on during pregnancy in women, with greater than 50% increases seen at term.12 Further, at term, a woman’s functional residual capacity (the volume of air present in the lungs at the end of expiration) decreases by 20%.12 These changes reduce respiratory reserve and support the use of supplemental oxygen prior to induction of anesthesia, the use of increased inspired oxygen concentrations and mechanical ventilation during anesthesia with continued oxygen supplementation during recovery. Cardiac output and blood volume increase during pregnancy in women. Arterial blood pressures do not change dramatically but near term women can become acutely hypotensive when placed on their back due to aortocaval compression.12 Aortocaval compression has not been demonstrated in animals but heart rate, respiratory rate, and mean uterine artery blood pressure increased and arterial oxygenation decreased in cows in the third trimester of pregnancy positioned in dorsal recumbency.13 Hormonal changes during pregnancy may also affect anesthesia. The administration of progesterone to rabbits lowers the concentration of halothane required to produce anesthesia.14 Pregnancy also lowers the requirements for anesthetics in women and makes them more sensitive to the effects of local anesthetics. In one mare study, serum calcium levels decreased and parathyroid concentrations increased immediately prior to foaling and remained decreased for 4 to 6 days.15 In another study in normal Arabian mares no changes in calcium, sodium, potassium, or magnesium were seen.16 Once parturition begins, heart rate and respiratory rate increase and the potential for exhaustion, hypovolemia, and lactic acidosis due to exertion could complicate anesthesia for correction of dystocia.
The fetus as a patient Virtually all anesthetic drugs used in horses are lipophilic, thus most readily cross the placental barrier and affect the fetus.17, 18 Further evidence of placental transfer of drug is found in two abstracts that note decreased fetal heart rates in mares given detomidine19 and xylazine.20 Cardiac output in the foal is dependent on heart rate at birth so it is assumed that the decreases in heart rate reflect decreases in cardiac output, but the author could find no definitive reports on the subject. In addition to their effects on heart rate, virtually all anesthetic drugs reduce cardiac output and respiratory function in the mare by a variety of mechanisms, potentially leading to fetal compromise. Studies in pregnant pony mares show that, compared to the dam, the fetus has lower blood oxygen tensions and higher blood carbon dioxide tensions and is acidotic (pH 7.44 in the dam, 7.35 in the fetus).21 Fetal plasma lactate values are higher and plasma glucose levels lower than in the dam.21 During anesthesia fetal blood pH and oxygen tensions were lower and carbon dioxide tensions higher than when the dam was awake.21 Lower fetal pH may cause drugs, like detomidine, that are weak bases to accumulate.22 Oxygen uptake by uterine tissues increases two- to threefold in late gestation.23
Approach to anesthesia and sedation of the pregnant mare All anesthetic agents used in the horse cross the placenta and affect the foal, with the exception of neuromuscular blocking drugs like atracurium. There are no absolute contraindications for any sedative or anesthetic drug. The emphasis in providing sedation, anesthesia or analgesia to the pregnant mare should be to use the lowest possible dose of any drug that allows the procedure to be performed with minimal stress to the mare. Anesthetic techniques should be chosen that optimize cardiopulmonary function. Vigilant monitoring is paramount so that early intervention can be accomplished if necessary. The chronic use of non-steroidal anti-inflammatory drugs during pregnancy may cause drug accumulation in the fetus because of delayed metabolism.24
Sedation of the pregnant mare Drugs used for sedation of the horse include phenothiazine tranquilizers, alpha-2 adrenoreceptor agonists, and opioid agonists and partial agonists (see Table 5.1). Phenothiazines block the action of neurotransmitters centrally and peripherally and cause alphaadrenergic blockade leading to arterial hypotension.
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Table 5.1 Drugs frequently used for standing chemical restraint Drug
Dose and route
Onset of effect
Comments
Acepromazine Xylazine
0.02–0.05 mg/kg, i.m., i.v. 0.5–1.0 mg/kg, i.v. 1.0–2.2 mg/kg, i.m.
30 to 40 min, i.v., i.m. 3 to 5 min, i.v. 10 to 20 min, i.m.
Use cautiously in stressed or hypotensive horses Ataxia produced, head-down posture Start with low dose and repeat as needed
Detomidine
0.01–0.02 mg/kg, i.v. 0.02–0.04 mg/kg, i.m. 0.06 mg/kg, p.o.
3 to 5 min, i.v. 10 to 20 min, i.m. 30 to 45 min, p.o.
Ataxia produced, head-down posture Start with low dose, repeat as needed Can be given orally
Romifidine
0.04–0.12 mg/kg, i.v.
2 to 5 min, i.v.
Reduced ataxia, reduced analgesia
Butorphanol Buprenorphine Morphine
0.01–0.03 mg/kg, i.v. 0.006 mg/kg, i.v. 0.3–0.5 mg/kg, i.v.
3 to 5 min 10 to 20 min 3 to 5 min
Usually used in combination with a sedative or tranquilizer
Phenothiazines do not significantly affect respiration but respiratory rate frequently decreases. The duration of sedation depends on the dose administered but frequently lasts for 6 to 10 hours. Acepromazine does not produce analgesia but may make analgesic drugs more effective. Increasing the dose of acepromazine does not usually produce a greater effect but the duration of action is increased. Hypovolemic horses administered acepromazine may become acutely hypotensive, faint, and become recumbent. Treatment of hypotension should include large volumes of intravenous fluids. No specific antagonist for phenothiazine tranquilizers is known. Alpha-2 agonists produce sedation with muscle relaxation, ataxia, and analgesia when given orally, intravenously, or intramuscularly to horses. Xylazine, detomidine, and romifidine are approved for use in the horse in the United States. Arterial blood pressure is initially increased due to drug-induced increases in peripheral vascular resistance. Hypertension may be sustained (20 to 60 minutes), particularly when detomidine is used. Heart rate decreases and sinus arrhythmia, and first- and second-degree atrioventricular blockade are common. Decreases in heart rate produce significant decreases in cardiac output, often to levels 50% of predrug values. Respiratory rate is usually decreased but tidal volume increases. Of interest in the non-pregnant mare, detomidine, romifidine, and xylazine cause increases in intrauterine pressure after intravenous administration.25, 26 The repeated administration of alpha-2 agonists to pregnant mares does not appear to have untoward effects on pregnancy.27 Potentially, the administration of alpha-2 agonists for sedation could make obstetrical manipulations during dystocia more difficult, due to increases in tone of the uterine muscle, but abdominal press is probably an overriding factor. The level of sedation produced by administration of an alpha-2 agonist is more pronounced than that produced
Sedate with xylazine or detomidine prior to administering morphine. Potential for excitement. Reversible with naloxone
by phenothiazine administration. Horses assume a ‘head-down’ or ‘saw horse’ stance and frequently shift their weight from side to side. Combining alpha-2 agonists with other agents may permit the use of reduced doses, lessening the head-down posture and causing the horse to stand more squarely on all four feet. Opioids are used to produce analgesia and augment the effects of sedatives and tranquilizers. Butorphanol is a synthetic opioid agonist/antagonist that is approved for use in the horse for the treatment of abdominal pain.28, 29 Butorphanol is less prone to cause excitement than morphine or fentanyl, but ataxia can be severe at high doses. Butorphanol is frequently combined with alpha-2 agonists. When combined with xylazine, reduced doses of xylazine are usually used resulting in a horse that is less ataxic than with full doses of xylazine. Butorphanol is also used in combination with detomidine and romifidine. It has been suggested that the inclusion of butorphanol reduces the incidence of unexpected arousal. Other opioids including morphine, buprenorphine, and pentazocine have been used to produce standing chemical restraint.30, 31 The combination of xylazine and morphine produces a profoundly obtunded animal. Alpha-2 agonist administration should precede administration of morphine to prevent opioid-induced excitement. The head should be elevated and a nose twitch placed, if necessary, because the horse will tend to lean forward. Horses remain sensitive to touch so local anesthesia should be incorporated with the technique. The combination of acepromazine with an alpha-2 agonist allows for a reduction in the dose, reducing cardiopulmonary side effects and ataxia. Typically, 0.02 to 0.03 mg/kg of acepromazine is combined with 0.2 to 0.5 mg/kg of xylazine. The drugs can be combined in the same syringe and are usually given intravenously.
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Foal: Prepartum Assessment of the Foal
Anesthesia of the pregnant mare Anesthesia of the pregnant mare for any reason must be accomplished with a minimum of cardiorespiratory depression, paying particular attention to arterial blood pressures and oxygenation. All but the briefest and most urgent of field anesthesias should be discouraged because supplementation of the fraction of inspired oxygen may not be possible and increased levels of monitoring and cardiovascular support may be unavailable. The use of sedation and local anesthesia should be considered. Short-term (less than 30 minutes) field anesthesia is most easily accomplished using an alpha-2 agonist, a muscle relaxant such as diazepam or guaifenesin, and ketamine.32 The dose of alpha-2 should be minimized in order to produce the required level of sedation while minimizing decreases in cardiac output and ventilation. The required level of sedation is greater when diazepam is used as a muscle relaxant than when guaifenesin is used because guaifenesin can more easily be given to effect. Ketamine is administered intravenously once muscle relaxation is apparent. Short-term anesthetic techniques are presented in Table 5.2. Longer-term anesthesia should not be attempted unless oxygen can be supplemented. Oxygen can be supplemented with a nasal oxygen catheter with flow rates in excess of 15 L/min to effect a demonstrable change in arterial oxygen tension. More efficient methods of delivering oxygen can be achieved by using a demand valve which supplies oxygen at high flow rates during the inspiratory phase of respiration.33 In addition, a demand valve allows a breath to be delivered, usually by pressing a button that triggers air flow. The button is pressed until a normal tidal volume (as judged by movement of the chest wall) is delivered. The button is released and exhalation occurs. Endotracheal intubation is preferred but
breaths can be delivered by advancing a 10 to 14 mm diameter tube into one nostril. The tube is adapted to the demand valve and both nares are occluded. The breath is delivered and the nostrils released to allow exhalation. A source of oxygen is also useful when resuscitating foals. Anesthesia can be extended by continuing the administration of parenteral agents such as guaifenesin recipes (triple-drip) but oxygen should be supplemented.34, 35 If available, an anesthetic machine should be used to increase inspired oxygen tensions and provide for delivery of inhalants. Isoflurane and sevoflurane are the primary inhalant anesthetics used in the horse. Isoflurane and sevoflurane produce rapid induction to and recovery from anesthesia. Cardiovascular function is depressed in a dosedependent manner leading to decreases in cardiac output and arterial blood pressure.36 Respiratory depression should be anticipated with measures taken to assure that significant hypercapnia does not occur. The level of monitoring required for extended anesthesia in the pregnant mare includes measurement of arterial blood pressure and some index of respiratory function. Direct monitoring of arterial blood pressure is accomplished by catheterizing a peripheral artery. The facial, transverse facial, and dorsal metatarsal arteries are all easily palpated and catheterized in horses. The pressure in the artery can be measured using a simple aneroid manometer. Commercial units use pressure transducers and display the pressure waveform on a screen or compute the pressure and display it digitally. Mean arterial blood pressure should be maintained between 60 and 90 mmHg. Respiratory function is best assessed using arterial blood gas analysis. Additional methods of assessing ventilation include end-tidal carbon dioxide tension measurements and pulse oximetry. Hypotension is treated by decreasing the delivery of anesthetics (if possible), administering intravenous
Table 5.2 Drugs used for induction and maintenance of anesthesia in sedated horses Drug
Dose
Comments
Ketamine
1.5–2.0 mg/kg, i.v.
Horses must be maximally relaxed prior to administration. Relaxation can be produced with xylazine, diazepam, or guaifenesin
Guaifenesin
50 mg/kg, i.v., to effect
Primarily a muscle relaxant. Do not use alone. Use in sedated horses in combination with ketamine
Diazepam
0.06–0.1 mg/kg, i.v.
Primarily a muscle relaxant. Used primarily with ketamine
Tiletamine-zolazepam
0.7–1.0 mg/kg, i.v.
Horse should be fully sedated before administration. Produces 30 minutes of anesthesia with good muscle relaxation. Hypoventilation may occur
‘Triple-drip’ (mixture of guaifenesin 5%, ketamine 0.1%, and xylazine 0.05%)
2 mL/kg/h
Not used for induction in adult horses. Used to extend xylazine-ketamine or xylazine-diazepam-ketamine anesthesia. Monitor ventilation and the degree of muscle relaxation
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Sedation and Anesthesia of the Pregnant Mare fluids, and administering vasoactive drugs. If the anesthetic plane is appropriate, the maintenance rate of fluid administration (5–10 mL/kg/h) should be increased. A packed cell volume (PCV) and serum total protein (TP) should be monitored to assess the horse’s fluid status and prevent hemodilution. Hypovolemia in the presence of low PCV and TP must be corrected using blood or blood products. Hypotension that does not respond to fluid therapy is treated with vasoactive agents such as dobutamine or ephedrine. Dobutamine is a catecholamine drug that increases cardiac output and arterial blood pressure by increasing the strength of myocardial contractions.3 Dobutamine (1–5 µg/kg bwt/min) is given slowly (infused) and to effect and then titrated to maintain a desired arterial blood pressure. Ephedrine (0.05 to 0.1 mg/kg bwt) is given as an intravenous bolus and produces effects that persist for 20 to 30 minutes or longer.37 The administration of calcium solutions may produce increases in arterial blood pressure, particularly if ionized calcium levels are marginal or decreased such as occurs near foaling.38
Anesthesia for cesarean section or fetal manipulation for dystocia Anesthesia for cesarean section or fetal manipulation during dystocia is frequently an emergency procedure.39 In addition to the aforementioned concerns with hypotension and hypoxemia, the additional stressors of time and the viability of the foal are present. The time span from the administration of anesthetic drugs to the delivery of the foal is a critical component of fetal viability, in part because of drug transfer across the placenta and in part because fetal mortality increases with 30 to 40 minutes of labor.40, 41 In addition, if labor has begun, the mare may be exhausted (hypoglycemic and/or hypovolemic) or hyocalcemic. Elective cesarean section in the mare carries a good to excellent prognosis for both the mare and the foal.42 Initial examination of mares with dystocia should be conducted quickly. If sedation is required, alpha2 agonists (xylazine, detomidine) provide rapid onset of action. Xylazine may be preferred because its duration of action is shorter than that of detomidine. This is of particular importance if the fetus is compromised. Epidural anesthesia may be of limited value in an emergency situation because of the time it takes to see significant effects and its lack of effect on abdominal contractions. Epidural anesthesia/analgesia with or without catheter placement is of value for surgeries of the perineum and pain control.43 A catheter should be placed in a jugular vein and intravenous fluids begun as diagnostics and discus-
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sions with the owner take place. The mare should be moved to the induction area, preferably equipped with a hoist. In rapid order, the mare should be sedated with xylazine (1.0 mg/kg) or detomidine (0.02–0.04 mg/kg) intravenously followed within 3 to 5 minutes by diazepam (0.1 mg/kg, i.v.) and ketamine (2 mg/kg, i.v.). Upon the assumption of lateral recumbency, an orotracheal tube should be placed and connected to a large animal anesthetic machine or other oxygen source (demand valve). Fetal manipulation is facilitated by raising the hindlimbs using hobbles and a hoist or other lifting apparatus. Surgical preparation of the abdomen should be performed concurrent with fetal manipulation in case the manipulation is unsuccessful. Inhalant anesthetic administration should be instituted if delivery is not rapidly accomplished. Procedures for fetal manipulation and cesarean section are presented elsewhere in this text (see Chapters 235, 257 and 258) and reviewed in the literature.44 Either procedure can be associated with significant hemorrhage so attentive monitoring as described elsewhere in this chapter is important. Intravenous fluid support should be continued through the recovery period with calcium and glucose supplementation. Respiratory depression in the foal born to an anesthetized mare may be significant, thus provision for the placement of an endotracheal tube (10 to 14 mm internal diameter, 30 cm in length). Short-term ventilation can be accomplished using a standard small animal anesthetic machine, demand valve, or ‘ambu’ bag. Support of ventilation is preferred to the administration of antagonist drugs or analeptics.
References 1. Mee AM, Cripps PJ, Jones RS. A retrospective study of mortality associated with general anesthesia in horses: emergency procedures. Vet Rec 1998;142:307–9. 2. Johnston GM. Findings from the Confidential Enquiry into Perioperative Equine Fatalities. Proc 50th Ann Meet Am Assoc Eq Pract 2004;50:281–6. 3. Donaldson LL. Retrospective assessment of dobutamine therapy for hypotension in anesthetized horses. Vet Surg 1988;17(1):53–7. 4. Mhuireachtaigh RN, O’Gorman DA. Anesthesia in pregnant patients for nonobstetric surgery. J Clin Anes 2002;18:60–6. 5. Moens Y, Lagerweij E, Gootjes P, Poortman J. Distribution of inspired gas to each lung in the anaesthetized horse and influence of body shape. Equine Vet J 1995;27(2):110–16. 6. Santschi EM, Slone DE, Gronwall R, Juzwiak JS, Moll HD. Types of colic and frequency of postcolic abortion in pregnant mares: 105 cases (1984–1988). J Am Vet Med Assoc 1991;199(3):374–7. 7. Chenier TS, Whitehead AE. Foaling rates and risk factors for abortion in pregnant mares presented for medical or
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Foal: Prepartum Assessment of the Foal surgical treatment of colic: 153 cases (1993–2005). Can Vet J 2009;50:481–5. Bain AM. Foetal losses during pregnancy in the Thoroughbred mare: a record of 2,562 pregnancies. N Z Vet J 1969;17:155–8. Mazze RI, Kallen B. Reproductive outcome after anesthesia and operation during pregnancy: a registry study of 5405 cases. Am J Obstet Gynecol 1989;161:1178–85. Rosen MA. Management of anesthesia for the pregnant surgical patient. Anesthesiology 1999;91(4):1159–63. Littleford J. Effects on the fetus and newborn of maternal analgesia and anesthesia: a review. Can J Anesth 2004;51(6):586–609. McAuliffe F, Kametas N, Costello J, Rafferty GF, Greenough A, Nicolaides K. Respiratory function in singleton and twin pregnancy. Br J Obstet Gynaecol 2002;109:765–9. Dunlop CI, Hodgson DS, Smith JA, Chapman PL, Tyler LM. Cardiopulmonary effects of positioning pregnant cows in dorsal recumbency during the third trimester. Am J Vet Res 1994;55(1):147–51. Datta S, Migliozzi RP, Flanagan HL, Krieger NR. Chronically administered progesterone decreases halothane requirements in rabbits. Anes Analg 1989;68(1):46–50. Martin KL, Hoffman RM, Kronfeld DS, Ley WB, Warnick LD. Calcium decreases and parathyroid hormone increases in serum of periparturient mares. J Anim Sci 1996;74:834–9. Rook JS, Braselton WE, Nachreiner RF, Lloyd JW, Shea ME, Shelle JE, Hitzler PR. Multi-element assay of mammary secretions and sera from periparturient mares by inductively coupled argon plasma emission spectroscopy. Am J Vet Res 1997;58(4):376–8. Hubbell JAE, Muir WW, Sams RA. Guaifenesin: cardiopulmonary effects and plasma concentrations in horses. Am J Vet Res 1980;41:1751–5. Bidwell LA, Embertson R, Bone NL, Ryu MH. Diazepam levels in foals after dystocia birth. Proc 54th Ann Meet Am Assoc Eq Pract 2008;54:286–97. McGladdery AJ, Cottrill CM, Rossdale PD. Effects upon the fetus of sedative drugs administered to the mare. Proc 2nd Int Conf Vet Perinatol 1990;2:14. Smith LJ, Schott H. Xylazine-induced fetal bradycardia. Proc 2nd Int Conf Vet Perinatol 1990;2:36. Taylor PM, Silver M, Fowden AL. Intravenous catheterisation of foetus and mare in late pregnancy: management and respiratory, circulatory and metabolic effects. Equine Vet J 1992;24(5):391–6. Hubbell JAE, Sams RA, Schmall LM, Robertson JT, Hinchcliff KW, Muir WW. Pharmacokinetics of detomidine administered to horses at rest and after maximal exercise. Equine Vet J 2009;41:419–22. Fowden AL, Forhead AJ, White KL, Taylor PM. Equine uteroplacental metabolism at mid and late gestation. Exp Physiol 2000;85:539–45. Crisman MV, Wilcke JR, Sams RA, Gerken DF. Concentrations of phenylbutazone and oxyphenbutazone in postparturient mares and their neonatal foals. J Vet Pharm Ther 1991;14:330–4. Schatzmann U, Josseck H, Stauffer JL, Goosens L. Effects of alpha-2 agonists on intrauterine pressure and sedation in horses: comparison between detomidine, romifidine and xylazine. J Vet Med 1994;41:523–9. Von Reitzenstein M, Callahan MA, Hansen PJ, LeBlanc MM. Aberrations in uterine contractile patterns in mares
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with delayed uterine clearance after administration of detomidine and oxytocin. Theriogenology 2002;58:887– 98. Katila T, Oijala M. The effect of detomidine on the maintenance of equine pregnancy and foetal development: ten cases. Equine Vet J 1988;20:323–6. Robertson JT, Muir WW. Cardiopulmonary effects of butorphanol tartrate in horses. Am J Vet Res 1981;42:41–4. Robertson JT, Muir WW. A new analgesic drug combination in the horse. Am J Vet Res 1983;44:1667–9. Clarke KW, Paton BS. Combined use of detomidine with opiates in the horse. Equine Vet J 1988;20:331–4. Corletto F, Raisis AA, Brearley JC. Comparison of morphine and butorphanol as pre-anesthetic agents in combination with romifidine for field castration in ponies. Vet Anaes Analg 2005;32:16–22. Brock N, Hildebrand SV. 1990. A comparison of xylazinediazepam-ketamine and xylazine-guaifenesin-ketamine in equine anesthesia. Vet Surg 1990;19:468–74. Riebold TW, Evans AT, Robinson NE. Evaluation of the demand valve for resuscitation of horses. J Am Vet Med Assoc 1980;176:623–6. Greene SA, Thurmon JC, Tranquilli WJ, Benson GJ. Cardiopulmonary effects of continuous intravenous infusion of guaifenesin, ketamine, and xylazine in ponies. Am J Vet Res 1986;47:2364–8. McCarty JE, Trim CM, Ferguson D. 1990. Prolongation of anesthesia with xylazine, ketamine, and guaifenesin in horses: 64 cases (1986–1989). J Am Vet Med Assoc 1990;197:1646–50. Daunt DA, Steffey EP, Pascoe JR, Willits N, Daels PF. Actions of isoflurane and halothane in pregnant mares. J Am Vet Med Assoc 1992;201(9):1367–74. Grandy JL, Hodgson DS, Dunlop CI, Chapman PL, Heath RB. Cardiopulmonary effects of ephedrine in halothaneanesthetized horses. J Vet Pharm Ther 2008;12:389– 96. Grubb TL, Benson GJ, Foreman JH, Constable PD, Thurmon JC, Olson WO, Tranquilli WJ, Davis LE. Hemodynamic effects of ionized calcium in horses anesthetized with halothane or isoflurane. Am J Vet Res 1999;60:1430–5. Johnston GM, Taylor PM. Perioperative care of mares subjected to caesarean section. Part 1. Anesthesia. Equine Vet Educ 2002;Manual 5:65–9. Norton JL, Dallap BL, Johnston JK, Palmer JE, Sertich PL, Boston R, Wilkins PA. Retrospective study of dystocia in mares at a referral hospital. Equine Vet J 2007;39: 37–41. Freeman DE, Hungerford LL, Schaeffer D, Lock TF, Sertich PL, Baker GJ, Vaala WE, Johnston JK. Caesarean section and other methods for assisted delivery: comparison of effects on mare mortality and complications. Equine Vet J 1999;31:203–7. Watkins JP, Taylor TS, Day WC, Varner DD, Schumacher J, Baird AN, Welch RD. Elective cesarean section in mares: eight cases (1980–1989). J Am Vet Med Assoc 1990;197(12):1639–45. Gold JR. How to use an epidural in a field situation for analgesia or local anesthesia. Proc 54th Ann Meet Am Assoc Eq Pract 2008;54:290–4. Byron CR, Embertson RM, Bernard WV, Hance SR, Bramlage LR, Hopper SA. Dystocia in a referral hospital setting: approach and results. Equine Vet J 2002;35(1):82–5.
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Section (b): Foal Adaptation to Parturition
6. The Normal Post Partum Foal Sarah J. Stoneham
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Chapter 6
The Normal Post Partum Foal Sarah J. Stoneham
Introduction The first 2 weeks of a foal’s life carry the highest risk of mortality and many of the problems that occur are unique to this period. As a prey species, foals as the most vulnerable age group have evolved to exhibit minimal signs of weakness until disease processes are relatively advanced. Consequently signs of disease in the neonatal foal are often subtle and similar. Neonatal foals cannot be considered as scaleddown mature horses. Familiarity with the significant physiological differences and changes that occur in the first few days is essential to enable successful early diagnosis and treatment of disease.
Behavior of the postpartum foal After delivery of the hips, foals start the first gasping attempts to breathe which inflate the lungs. The foal struggles to right itself and sit on its sternum where it remains for a few minutes. At this time the mare usually nickers and starts looking round at the foal (see Figure 6.1). Most foals exhibit sucking movements of the mouth and tongue within a few minutes of birth. As the foal moves around towards the mare’s head the umbilical vessels stretch and break usually 2–3 cm from the ventral abdominal wall (see Figure 6.2). If the mare is disturbed she may stand, hastening this process. As the mare starts to lick the foal it extends its forelimbs as it attempts to stand. Most foals will stand within an hour of birth; however, some of the larger breeds and individuals may take longer. Foals that take more than 2 hours to stand should be considered abnormal. When the foal first stands it adopts a widebased stance and attempts exaggerated movements. As the foal’s coordination increases it will start udder seeking, moving along the mare’s ventral abdomen towards the udder. Many mares will gently nuzzle the foal towards the udder.
A few maiden mares will be reluctant to let the foal suckle. It is important not to interfere in the mare–foal bonding processes, but a few individual mares will require varying degrees of physical or chemical restraint to allow the foal to suckle and the mare–foal bond to become established. Most foals will suckle within 2 hours; more than 3 hours is considered abnormal. After their first full feed of colostrum the foal will often lie down and sleep. Neonatal foals usually sleep in full lateral recumbency with the dam standing close by. When resting they are usually in sternal recumbency. Some foals, often the larger, long-legged individuals, may take 8 hours or more to lie down, and only when exhausted or assisted are they able to coordinate their movements to lie down. These foals often develop some edema of the distal hindlimbs that resolves once they establish a normal pattern of activity and recumbency. During the first week the foal will suckle 5–7 times an hour for periods of about 2 minutes. Frequency of suckling bouts decreases during the night when the foal will sleep for longer periods. Foals butt the udder to encourage milk letdown and switch between teats during each suckling bout.1 Foals will consume approximately 23% of their bodyweight by day 2.2 Mares that become irritable or even aggressive towards their foals after behaving normally for the first day or two may have inadequate milk to meet the foal’s needs. These foals require careful management with supplementary feeding until the mare reaches full lactation. Careful observation of the foal suckling and examination of the udder will help to establish whether the mare is suffering from poor milk letdown or partial agalactia. If the mare is suffering poor letdown the use of small doses of oxytocin (2.5–5 IU) will help milk letdown. In some cases, when poor letdown is associated with pain or discomfort, the use of analgesics
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Foal Adaptation to Parturition
Figure 6.1 The newborn foal rapidly sits in sternal recumbency.
may be useful. Maiden mares may require some reassurance and restraint until the mare–foal bond is fully established. Mares with agalactia can be treated with domperidone (1.1 mg/kg daily per os). Domperidone is used as a therapeutic agent to treat endophyte-infected fescue toxicosis and enhances prolactin release. An increasing plane of nutrition and access to grass and exercise are also important. Over the first 12 hours the foal becomes increasingly coordinated, active, and inquisitive.
Figure 6.3 The Brix refractometer can be used to estimate colostral IgG.
Colostrum Colostrum is produced as a unique event under hormonal influence during the last few weeks of pregnancy. It contains immunoglobulins (IgG), complement, lactoferrin, lysozyme, and lymphocytes that are essential to enhance the foal’s immune system. The range of antibodies transferred to the foal is dependent on the antigens to which the mare has been vaccinated or exposed. Thoroughbred mares usually produce 1–2 litres (see Chapter 33). There is considerable variation in the IgG concentration between individual mares’ colostrum; some individuals repeatedly produce colostrum with a low IgG concentration (< 40 g/L). Colostral quality should be assessed immediately after foaling. The use of a Brix refractometer (Bellingham & Stanley, Kent, UK) (see Figures 6.3 and 6.4) has been reported to provide a good measure of colostral IgG concentration.3 Alternative methods for
120 100 80 60 40 20 0 0
Figure 6.2 The umbilical vessels break naturally as they are stretched when the foal starts to struggle to move.
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Figure 6.4 Interpretation of Brix refractometer percentage readings for assessing colostral quality.
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The Normal Post Partum Foal assessment of colostral quality include the colostrometer, which measures specific gravity. Good-quality colostrum has a specific gravity > 1.060.4 If colostrum is poor quality the foal should be supplemented with good-quality donor colostrum (IgG > 60 g/L). Efficacy of passive transfer of colostral immunity declines rapidly after birth. Immunoglobulin absorption has been shown to be 51% efficient 1 hour postpartum, declining to 1% 22 hours postpartum. If foals fail to suckle within 3–4 hours of birth, goodquality colostrum should be administered without delay by stomach tube or bottle to maximize efficacy of absorption. Specialized short-lived cells in the epithelium of the small intestine pinocytose macromolecules including immunoglobulins. The macromolecules are transported through the cell in membrane-bound vesicles that exocytose their contents into the lamina propria from where immunoglobulins pass into the lymphatics and into the circulation via the lymphatic ducts. IgG is detected in the foal’s blood 6–8 hours after ingestion of colostrum.
Routine treatment of the newborn foal Routine veterinary examination of the neonatal foal should include a methodical physical examination to detect any early signs of disease or congenital abnormality. The placenta should also be examined for evidence of in utero disease. A healthy placenta should weigh approximately 10–11% of the foal’s bodyweight. Observation of the foal undisturbed allows behavior and mare–foal interaction to be assessed. Physical appearance can provide information on maturity. Immature foals may exhibit a silky mole-like coat, have a domed forehead, tendon and ligamentous laxity, generalized weakness, and may be underweight and small in size. Poor body condition may also indicate intrauterine growth retardation. Normal parameters for heart and respiratory rates change over the first 24 hours (see Table 6.1). Mucous membranes are normally pale pink and moist. Tem-
Table 6.1 Normal heart and respiratory rate of the neonatal foal
Age
Heart rate (beats/minute)
Respiratory rate (breaths/minute)
1 minute 15 minute 12 hours 24 hours
60–80 120–160 80–120 80–100
Gasping 40–60 30–40 30
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perature range is 37–39◦ C (99–102◦ F). Hypothermia or pyrexia are often indicative of disease. However, pyrexia in the neonate is not a consistent sign of bacterial infection. Auscultation of the lungs is a less reliable indicator of disease than in the adult. The thin body wall with little subcutaneous tissue contributes to increased, slightly harsh lung sounds in the normal newborn foal. Rib fractures can be detected by careful palpation of the thorax although ultrasonographic examination is the most reliable method of confirming the diagnosis.5 An audible end-inspiratory clicking may also be ausculted over the fracture site. Umbilical hernias and abnormalities of the umbilical remnant (e.g., patent urachus, omphalitis) can be detected by palpation and colts should be examined for scrotal and inguinal hernias. The umbilical stump should be treated daily for the first few days until it dries and seals. Studies have shown 0.5% chlorhexidine solution to be most persistent and effective in reduction of bacterial contamination of the stump.6 The use of antibiotic spray is common and has proved effective in practice. Solutions such as > 2% iodine should be avoided as they can induce tissue necrosis. Meconium is usually passed within 2–12 hours of birth. The quantity varies considerably between individuals. In most foals the appearance of soft, pale, tan-colored milk dung indicates complete passage of meconium. Some stud farms routinely administer a phosphate enema or a gravity-flow soapy water enema after birth to hasten the passage of meconium. There is no evidence to suggest this reduces the incidence of meconium impaction. Observation of the foal walking and standing on a firm level surface allows detection of angular and flexural limb deformities that may require careful management or veterinary intervention. Routine screening blood samples taken at 12–24 hours for hematology, proteins, IgG, plasma fibrinogen, and serum amyloid A may provide information on efficacy of passive transfer and early signs of disease such as neonatal isoerythrolysis, or sepsis.
The transition from intrauterine to extrauterine life Thermoregulation Healthy newborn foals are able to thermoregulate effectively. They are dependent on high levels of activity and frequent ingestion of colostrum and then milk to maintain body temperature. Shivering is also a mechanism used to increase body temperature. Young foals are susceptible to hypothermia as they have poorly developed mechanisms of
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thermogenesis, relatively large surface area to volume ratio, and little subcutaneous fat. This is particularly significant in immature, sick or anesthetized young foals. Respiratory system At birth the foal is hypoxemic and hypercapnic. This stimulates the first few gasping breaths, the lungs expand and Pa o2 rises. The chest wall of the newborn foal is very compliant and the lungs are relatively inelastic, which limits tidal volume and increases the effort required to breathe. Low functional reserve capacity is counteracted to some extent by a high respiratory rate. Lateral recumbency reduces Pa o2 levels by up to 20 mmHg.7 During the first 7 days of life tidal volume, Pa o2 , and alveolar ventilation increase. By the end of the first week there is a significant increase in the foal’s functional respiratory reserve.7, 8 Alveolar development begins at about 260 days of gestation and by 320 days surfactant is present. However in foals delivered before full term, surfactant activity in the lungs may be reduced.9 In the Thoroughbred, bronchiolar duct endings continue to increase in number over the first year, a feature that appears not to occur in ponies.10 Cardiovascular system The first few breaths that expand the lungs result in a marked decrease in pulmonary vascular resistance, increasing blood flow through the lungs. This increases flow into the left atrium and results in closure of the foramen ovale. The cessation of venous return from the umbilical vein reduces right atrial pressure. Constriction of the ductus arteriosus produces physiological closure at about 24 hours of age. Anatomical closure occurs around 3 days. Over the first few days of life hypoxemia, hypercapnia, and acidosis can result in left-to-right shunting with reopening of the ductus arteriosus and foramen ovale and rising pulmonary vascular resistance. This reversion to transitional or fetal circulation results in severe atelectasis and worsening hypoxemia that rapidly becomes irreversible without aggressive intervention. Left-sided systolic murmurs can be detected in many foals for the first few days and can persist for more than 35 days. In a few foals these systolic murmurs can be associated with turbulent flow through the ductus or, more often, the murmurs are physiological.11 As stroke volume of the neonatal foal is limited, cardiac output is rate-dependent.
Nervous system The nervous system of the newborn foal is mature, allowing the foal to stand, suckle, and gallop within hours of birth. There are several significant differences in the neurological response of the newborn foal compared to the adult. Neurological examination of the newborn foal has been described12 (see Chapter 53). It is important to be aware of the differences in normal responses that in the adult may be abnormal and associated with pathology. Particular points to remember include:
r
r r r r r r
Until about 3 weeks of age, foals have a marked crossed extensor reflex of the forelimbs, with marked resting extensor tone when in lateral recumbency. These reflexes are present to a lesser extent in the hindlimbs. Reflexes of the hindlimbs are hyperflexic, e.g., patellar reflex. Gaits are hypermetric. Movements of the head and neck are rather jerky and exaggerated. The menace response is incomplete until about 2 weeks of age.13 Foals have a biphasic pupillary light response (PLR) and for the first few days the PLR is slower than in the adult.13 Pupil position is ventromedial but should become dorsomedial by about 1 month of age; however, pupil position may change with brain dysfunction.13
Urinary system Renal function is relatively mature. However, at birth, mechanisms of tubular secretory function may not be fully mature and the affected neonate will experience increased sodium, potassium, and chloride losses until tubular secretory function matures. Glomerular filtration rate and renal plasma blood flow are close to adult values. Foals usually urinate for the first time within 8–12 hours postpartum. Foals produce large volumes (148 mL/kg/day) of hypotonic urine (specific gravity 1.008–1.012).14 There is transient proteinuria due to the non-specific transfer of macromolecules into the circulation over the first 24 hours postpartum.
Gastrointestinal system The foal is able to digest colostrum and milk immediately after birth. Digestion of mare’s milk is approximately 98% efficient.2 Growth hormones and trophic
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The Normal Post Partum Foal factors in colostrum and milk are important for the normal development of the gastrointestinal tract. Meconium is usually passed 2–12 hours after birth. Some foals, particularly those with longer gestational ages, may have considerable quantities of meconium. Meconium is usually firm to hard in consistency, is dark brown, and often glistening in appearance. The passage of pale, soft, tan milk feces indicates that all meconium has been passed.
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Complement levels and lymphocyte numbers are reduced in the newborn; there is considerable individual variation in the timing of the increase in lymphocyte subset (that is, different types of lymphocytes divided into subsets based on differences in structure and function, e.g., T lymphocytes CD+4, CD+8) numbers.
Musculoskeletal system Immune system At birth all the components of the immune system are present; however, the system is naive. Due to the epitheliochorial structure of the equine placenta the newborn foal is born agammaglobulinemic and depends on colostral transfer of passive immunity for immunoglobulins. Passively derived antibodies have a relatively short half-life: IgGa = 176 days, IgGb = 32 days, IgG(T) = 21 days, and IgA = 3.4 days. Endogenous antibody production rises slowly, with endogenous IgE and IgGb not being detected until the foal is about 2 months of age.15 This inevitably results in trough antibody levels occurring at 1–2 months of age. In addition to immunoglobulins, colostrum contains complement, lactoferrin, cytokines, growth factors, hormones, and maternal lymphocytes. Recent work in the calf has shown the transfer of these lymphocytes to the newborn has a positive effect, enhancing rapid development and activation of the calf’s own lymphocytes.16 The opsonizing capacity of foal serum is enhanced following ingestion of colostrum.17, 18 Maintenance of pregnancy requires some degree of suppression of the maternal immune system to prevent rejection of the fetus as a foreign body. This is modulated through soluble factors produced by the placenta that switch the balance of maternal immune response in favour of Th2-type responses.19 Inevitably there is a ‘hangover’ effect on the immune system of the newborn that skews immune response in favor of Th2 immunity. T-helper cell responses are classified based on the pattern of cytokines they secrete. Th1 response in general is a proinflammatory response associated with IL2 and IFN gamma. It favours cell-mediated immunity and is involved in autoimmune and rejection response. Th2-type response is associated with IL4, IL10, and IL 13 secretion and an anti-inflammatory, atopic/allergic response. It favours humoral immunity. A successful immune response to challenge involves a balance between these two types of T-helper cell responses. This bias also affects the mucosal immune system. Newborn foals have been shown to be gamma interferon deficient,20 suggesting their ability to mount Th1type responses is limited.
The musculoskeletal system is well developed in the full-term newborn foal, with well-ossified bones. Ossification of the cuboidal bones occurs relatively late in gestation and incomplete ossification of the carpal and tarsal bones is a frequent problem in premature foals which can be challenging to manage effectively. Limbs of the newborn foal are approximately 68% of mature length. As the legs have been non-weightbearing in utero, tendons and ligaments are lax to varying degrees. This relative laxity produces an increased range of joint movement. Weight bearing and increasing levels of exercise increase tendon and ligament tension and muscular strength. Newborn foals usually have a toe-out conformation with a mild degree of carpal valgus (< 5◦ ) that self-corrects over the first month of life. At a month of age, < 3◦ carpal valgus is considered normal. Growth rates during the first month are rapid, with daily weight gains of 1.5–2 kg/day in Thoroughbred foals. Most foals will double their birthweight in the first month with significant increases in subcutaneous fat and muscle.
References 1. Carson K, Wood-Gush DGM. Behaviour of Thoroughbred foals during nursing. Equine Vet J 1983;15:257–62. 2. Ousey JC, McArthur AJ, Rossdale PD. How much energy do sick neonatal foals require compared to healthy ones? Pferdeheilkunde 1996;12:231–7. 3. Cash RSG. (1999) Colostral quality determined by refractometry. Equine Vet Educ 1999;11:36–8. 4. LeBlanc MM, McLaurin BI, Boswell R. Relationships among serum immunoglobulin concentration in foals, colostral specific gravity, and colostral immunoglobulin concentration. J Am Vet Med Assoc 1986;189:57. 5. Jean D, Picador V, Maier S, Beauregard G, Danto MA, Beauchamp G. Detection of rib trauma in newborn foals in equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 2007; 39(2):158–63. 6. Lavan RP, Madigan JE, Walker R, Muller N. Effects of disinfectant treatments on the bacterial flora of the umbilicus of neonatal foals. In: Proceedings of 40th Annual Convention of the American Association of Equine Practitioners, 1994; pp. 37–8.
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7. Stewart JH, Rose RJ, Barko AM. Respiratory studies in foals from birth to seven days. Equine Vet J 1984;16:329–31. 8. Rose RJ. Cardiorespiratory adaptations in neonatal foals. Equine Vet J Suppl 1988;5:11–13. 9. Pattle RE, Rossdale PD, Schock C, Creasey MJ. The development of the lung and its surfactant in the foal and in other species. J Reprod Fert Suppl 1975;2:651. 10. Beech DJ, Sibbons PD, Rossdale PD, Ousey JC, Holdstock NB, Chavatte P, Ansari T. Organogenesis of lung and kidney in thoroughbreds and ponies. Equine Vet J 2001;33: 438–45. 11. Livesey LC, Marr CM, Boswood A, Freeman S, Bowen IM, Corley KTT. Auscultation and two dimensional, M mode, spectral and colour flow Doppler findings in pony foals from birth to seven weeks of age. J Vet Intern Med 1998;12:255. 12. Mayhew IG. Large Animal Neurology: A Handbook for Veterinary Clinicians. Philadelphia: Lea & Febiger, 1989. 13. Enzerink E. The menace response and papillary light reflex in neonatal foals. Equine Vet J 1998;30:546–8. 14. Brewer BD, Clements SF, Lots WS, Gronwall R. Renal clearance: urinary secretion of endogenous substances and urinary diagnostic indices in healthy neonatal foals. J Vet Intern Med 1991;5:28–33.
15. Wagner B, Flaminio JB, Hillegas J, Leibold W, Erb HN, Antczak DF. Occurrence of IgE in foals: evidence for transfer of maternal IgE by the colostrum and late onset of endogenous IgE production in the horse. Vet Immunol Immunopathol 2006;110(3–4):269–78. 16. Reber AJ, Donovan DC, Gabbard J, Galland K, AcevesAvila M, Holbert KA, Marshall L, Hurley DJ. Transfer of maternal colostral leukocytes promotes development of the neonatal immune system. Part II Effects on neonatal lymphocytes. Vet Immunol Immunopathol 2008;15(123):305–13. 17. Leblanc MM, Pritchard EL. Effect of bovine colostrum, foal serum immunoglobulin concentration and intravenous plasma transfusion on chemiluminescence of foal neutrophils. Anim Genet 1988;19(4):435–45. 18. McTaggart C, Penhale J, Raidal SL. Effect of plasma transfusion on neutrophil function in healthy and septic foals. Aust Vet J 2005;83(8):499–505. 19. Morein B, Blomqvist G, Hu K. Immune responsiveness in the neonatal period. J Comp Pathol 2007;137(Suppl 1):S27–31. 20. Breathnach CC, Sturgill-Wright T, Stiltner JL, Adams AA, Lunn DP, Horohov DW. Foals are interferon gamma deficient at birth. Vet Immunol Immunopathol 2006;112(3–4): 199–209.
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Chapter 7
Endocrinological Adaptation Jennifer C. Ousey
The foal has an active endocrine system from early in fetal life. Together with the placenta, the equine fetus synthesizes precursors for the major endocrine products of pregnancy, namely progestagens and estrogens, which maintain the pregnancy up to parturition. Once delivered and independent from the homeostatic mechanisms of the mare and the placenta, the foal has to respond dynamically to the extrauterine environment in order to survive. The transition from life as a fetus in utero to a free-living foal ex utero is reflected in a series of endocrine changes that can be measured within minutes or hours of birth. In some situations, this endocrine transition is not made and the foal remains in a semi-fetal state after birth, often with fatal consequences. This chapter will describe the normal endocrine changes that occur in healthy newborn foals during the perinatal period and explain why they are important in terms of postnatal adaptation. The hormones of clinical significance in the neonatal foal and their functions are summarized in Table 7.1.
The hypothalamo–pituitary– adrenocortical axis A functional hypothalamo–pituitary–adrenocortical (HPA) axis is essential for postnatal survival. Activation of the fetal HPA axis drives many of the endocrine changes that occur in the periparturient period by inducing enzyme systems and cellular changes that lead to structural and functional maturity of many fetal tissues.1 It may also provide the signal for impending parturition.2 The HPA axis is regulated by the classic negative feedback system, and the major components of this endocrine pathway and their enzymes are shown in Figure 7.1. For most of gestation, the primary glucocorticoid produced by the fetal adrenal cortex is corticosterone; but close to
parturition and after birth, increasing quantities of cortisol are synthesized, associated with induction of the adrenal enzyme, 17α-hydroxylase (17α−OH).3, 4 This is the key steroidogenic enzyme responsible for producing cortisol from its precursor, pregnenolone (P5), via the 17α-hydroxyprogesterone (17αOH P4) pathway. Following induction of 17α-OH, fetal cortisol concentrations rapidly rise. This occurs during the last 24–48 h of gestation in the horse and is proportionally later in gestation compared to other species.5 Cortisol levels continue to increase in the newborn foal, peaking at 1 h before declining to baseline by 4–6 h postpartum (Figure 7.2a). The perinatal rise in cortisol is stimulated by rising adrenocorticotropin (ACTH) concentrations in the fetus (Figure 7.2b) through upregulation of the whole HPA system. ACTH concentrations peak at parturition and then decline rapidly to baseline.6, 7 There are differences between breeds, with Thoroughbred foals achieving higher postnatal cortisol and ACTH concentrations than pony foals.9, 10 This may arise from the stress of delivery which, normally, lasts longer in Thoroughbred compared to pony mares. The changes that occur in the adrenocortical hormones in the fetus are linked to morphological maturation of the adrenal gland in late gestation. The equine fetal adrenal gland increases in size from 60 to 100 mg/kg bodyweight between 300 days of gestation and term.7 At the same time, a distinct band of cells develops between the cortex and medulla (the zona reticularis) where expression of 17α-OH activity is localized.4, 11 Both immunocytochemical staining and radiolabeled incubation studies have demonstrated that there is little 17α-OH activity in the fetal adrenal glands until birth.3, 4 Consequently, for much of gestation the fetal adrenal glands are relatively unresponsive to ACTH stimulation and unable to produce cortisol. Instead, they synthesize pregnenolone,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 7.1 A summary of clinically relevant hormones and their actions during the perinatal period in the foal Typical plasma concentrations
Hormone
Source
Primary role
Cortisol
Adrenal cortex
Stimulates organ maturation (particularly lung) and tissue differentiation (fetus) and physiological adaptations after birth (foal): nutrient provision (gluconeogenesis), catecholamine release, cardiac contractility, vascular responses (increase in blood pressure). Cortisol is also important for stress responses, anti-inflammatory actions and immune suppression
Birth: > 80 ng/mL; neonate: 20–30 ng/mL (basal) or 35–60 ng/mL (stimulated)
Adrenocorticotrophin (ACTH)
Anterior pituitary
Essential component of the HPA axis which is upregulated during the perinatal period. ACTH stimulates adrenal activity leading to pregnenolone production (fetus) via P450scc (side chain cleavage) activity, and biosynthesis of cortisol (foal) via 17α-hydroxylase activity
Birth: > 250 pg/mL; neonate: 10–30 pg/mL (basal) or 180 pg/mL (stimulated)
Catecholamines: epinephrine (adrenaline), norepinephrine (noradrenaline), and dopamine
Adrenal medulla and sympathetic nervous system
Increase heart rate/contractility, peripheral vasoconstriction, increase systemic vascular resistance and pressure, decrease pulmonary resistance and pressure (hypotension stimulates norepinephrine release; hypoxia stimulates release of both epinephrine and norepinephrine). Catecholamines initiate closure of fetal shunts. Increase circulating glucose concentrations via glycogenolysis and gluconeogenesis (hypoglyaemia stimulates a marked epinephrine response). Stimulate fatty acid metabolism and non-shivering thermogenesis
Adrenaline: birth: 3 ng/mL; neonate: 0.2–0.3 ng/mL (basal) or > 2 ng/mL (stimulated); Norepinephrine: birth 4–6 ng/mL; neonate: 0.09–0.3 ng/mL (basal) or > 0.6 ng/mL (stimulated)
Insulin
Pancreatic beta cells Pancreatic alpha cells (islets of Langerhans)
Essential for glucose and nutrient metabolism in the fetus and foal. Insulin lowers blood glucose concentrations after feeding by stimulating glucose uptake via glycogenesis, (glycogen storage in the liver and muscles), glycolysis (energy production), and synthesis of fatty acids. Glucagon stimulates glucose production from hepatic glycogen stores, hence high glucagon concentrations after birth
Insulin: birth: 70–170 pmol/L; day 1: unfed 20–40 pmol/L, fed 140 pmol/L; day 7: unfed 130 pmol/l, fed 250 pmol/L; Glucagon: birth: 1500 pg/mL
Thyroxine (T4) and tri-iodothyronine (T3)
Thyroid (follicular cells)
Important for fetal bone ossification, tendon development, neurodevelopment, locomotor skills, and lung maturation; T3 and T4 act synergistically with cortisol and somatomedin. Thyroid hormones have widespread effects by stimulating growth, energy metabolism (increasing basal metabolic rate and cardiac output), and thermogenesis
Total T3: birth: 8–12 nmol/L; neonate: 14 nmol/L; Total T4: birth: 400–500 nmol/L; neonate: 140 nmol/L
Renin–angiotensin– aldosterone system (RAAS) Vasopressin
Kidneys and other organs (see Figure 7.6) Posterior pituitary
RAAS and vasopressin control fluid and electrolyte homeostasis and maintain blood pressure. They minimize urinary sodium losses during the neonatal period when urine production is high. RAAS/vasopressin are also important for increasing blood pressure, reducing heart rate, and stimulating the onset of pulmonary circulation and occlusion of placental-umbilical circulation, after birth
Aldosterone: birth: 400–500 pg/mL; neonate: 50–150 pg/mL Vasopressin: birth: 20–25 pg/mL; older foal: 2–4 pg/mL
Gastrin
Parietal cells (stomach)
Promotes protein catabolism by initiating release of gastric acid and pepsinogen into the stomach. Also stimulates gut development hence high concentrations in the neonate
Gastrin: neonate: unfed: 15–25 pmol/L; fed: > 30 pmol/L
Pancreatic polypeptide
Pancreatic P cells
Inhibits pancreatic exocrine secretions and stimulates gastric emptying. Low concentrations in the neonate permit maximum digestion of milk
Enteroglucagon
Ileum and colon
Stimulates growth of the small intestine
Glucagon
Note: Stimulated concentrations refer to published values obtained either following organ stimulation tests, feeding, during sickness or stress situations.
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Figure 7.1 The major endocrine pathways associated with the HPA axis in the feto-placental unit and in the newborn foal. Single or double arrows indicate endocrine pathways that predominate in the fetus or newborn foal, respectively. Dashed arrows indicate positive (+) or negative (-) feedback. Adrenal pathway (dark gray). Endometrial/placental pathway (light gray). CRH, corticotrophic releasing hormone; POMC, proopiomelanocortin; P4, progesterone; P5, pregnenolone; 5αDHP, 5α-dihydroprogesterone; 11-DOC, 11-deoxycorticosterone; 17αOHP5, 17α-hydroxypregnenolone; 3βHSD, 3β-hydroxysteroid dehydrogenase. For all other abbreviations, see text.
through P450 side-chain cleavage (P450scc) of cholesterol, which is the major precursor for production of progestagens by the feto-placental unit (Figure 7.1).12, 13 Administration of exogenous ACTH to the fetus will elicit an increase in (maternal) progestagen concentrations but has little effect on fetal cortisol concentrations before 300 days of gestation.14, 15 After 300 days, the cortisol response to ACTH becomes proportionally greater with gestation and postnatal age. At 2 to 3 weeks before birth, 1–2 µg/kg ACTH i.v. stimulates a 50–60% increase in fetal plasma cortisol, while after birth the same dose elicits a 300 to 400% increase in mature, healthy foals.6, 15 These results support the concept of gradually enhanced fetal adrenocortical activity, with significant cortisol production and induction of 17α-OH only at birth. Maturation of the HPA axis continues to develop postpartum because adrenocortical responsiveness to stressful stimuli (e.g., hypoglycemia, hypotension) gradually increases in foals during the first few weeks after birth.16, 17
The lack of adrenal cortisol response to ACTH in late gestation is limited not only by 17α-OH activity, but also by high concentrations of fetal cortisolbinding globulin (CBG) which controls the availability of free cortisol. Fetal CBG concentrations are approx 50 ng/mL but decline substantially during the last 2–3 days of gestation to reach about 10 ng/mL at birth (Figure 7.2c).8 As CBG declines in the perinatal period, the availability of bioactive, free cortisol rises, thereby enhancing the physiological effects of cortisol and highlighting the importance of measuring free, rather than total, cortisol concentrations in foals. The concentrations of CBG are very low in newborn foals compared to neonates of other species, although an additional, unidentified, protein has also been detected in foal plasma which binds equally as much cortisol as CBG.18 After birth, CBG concentrations gradually rise over several months until higher, adult levels are achieved. The importance of cortisol in promoting fetal maturation and survival postpartum is demonstrated by foals delivered before the prepartum cortisol rise,
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their cortisol concentrations remain low after birth, similar to fetal concentrations in late gestation (Figure 7.2).6 The pituitary pathway is active in these foals because plasma ACTH concentrations are significantly higher in premature than mature foals after birth, indicating poor adrenal responsiveness, similar to that observed in the fetus (see above). Moreover such foals continue to produce high concentrations of progestagens (pregnenolone and its metabolite pregnenediol, P5ββ) after birth, supporting the concept of adrenal dysfunction and a persistent fetal state.20, 21 Other clinical problems, for example, perinatal asphyxia syndrome (PAS) or septicemia, stimulate release of high concentrations of ACTH and other stress hormones (arginine vasopressin [AVP], catecholamines) including cortisol, and are associated with higher mortality rates.21–23 In contrast, foals born from mares fed fescue grass infected with fungal endophytes have suppressed HPA function, with low ACTH and cortisol concentrations, because the ergot alkaloids act centrally on the D2 dopamine receptors to block corticotrophs in the fetal anterior pituitary and inhibit ACTH release.24, 25 Those foals with persistently low cortisol concentrations have high mortality rates. Collectively, these results demonstrate the importance of a functional HPA axis after birth, and how disturbances at any level in the endocrine pathway may have serious, if not fatal, consequences for the health of the newborn foal.
Other adrenocortical responses
30
The adrenal cortex also secretes mineralocorticoids from the zona glomerulosa, the most important of which is aldosterone. This regulates blood pressure and electrolyte balance. Secretion of aldosterone is controlled by the kidneys and is discussed below.
20 10 0 -15
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Figure 7.2 (a) Ontogenic changes in plasma cortisol concentrations during the perinatal period in full-term and premature foals. (b) Ontogenic changes in plasma adrenocorticotropin (ACTH) concentrations during the perinatal period in full-term and premature foals. (c) Ontogenic changes in cortisol-binding globulin (CBG) concentrations during the perinatal period in full-term foals. Adapted from Silver et al. (1984),6 Fowden and Silver (1995),7 and Cudd et al. (1995).8
such as spontaneously delivered premature foals (see Chapter 12) or those delivered early by cesarean section due to maternal disease. These foals display a range of clinical problems associated with organ immaturity and have high mortality rates.5, 19 Typically
Adrenal medulla In addition to synthesizing glucocorticoids and mineralocorticoids, the adrenal gland also synthesizes catecholamines (epinephrine, norepinephrine, and dopamine) from the medullary (inner) layer. Epinephrine is produced from norepinephrine via the enzyme phenylethanolamineN-methyltransferase (PNMT), which is particularly evident in the medullary cells in late gestation, and its activity is regulated by glucocorticoids.4 Norepinephrine is released from the adrenal medulla and from the sympathetic nervous system and, therefore, any factor which affects the whole sympathoadrenal system will cause significant catecholamine release.
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Endocrinological Adaptation Catecholamines regulate intermediary metabolism and play an important role in the circulatory adjustments to stress; they act via α and β adrenergic receptors located in a variety of tissues. Innervation of the adrenal medulla is not complete by birth, but it becomes progressively more effective at releasing catecholamines in response to direct and indirect stimuli over the perinatal period.26 Direct (electrical) stimulation of the splanchnic nerve causes a small rise in circulating catecholamine concentrations, a response which increases with age in fetuses from late gestation continuing into the postnatal period. However, anoxia or asphyxia over the same period will elicit a much greater catecholamine release. Basal levels of epinephrine and norepinephrine in the fetus are < 200 pg/mL and 2 ng/mL, respectively, until the last few days before delivery when concentrations increase substantially (to 400–600 pg/mL and 4–6 ng/mL, respectively) and are positively correlated with the rise in fetal cortisol concentrations.27 These high levels are maintained at birth, associated with the hypoxic stress of delivery, and are necessary to induce the circulatory changes (increased heart rate/contractility, peripheral vasoconstriction, increased systemic vascular resistance/pressure, decreased pulmonary resistance/pressure, closure of fetal shunts) that occur after birth.28, 29 There is a negative correlation between umbilical blood pH and catecholamine concentrations in newborn foals, reflecting the degree of stress and hypoxemia experienced during delivery.6 Catecholamine concentrations decline immediately after birth but may rise again over the next 2 h, associated with epinephrineinduced glycogenolysis and gluconeogenesis in response to the high calorie (glucose) requirements of the newborn foal. Mature foals tend to have higher concentrations of epinephrine than premature foals at birth, associated with higher cortisol concentrations, whereas premature foals that are stressed and acidotic have much higher norepinephrine concentrations than term foals.6, 30 Hypoglycemia is a major stimulus for catecholamine release and exogenous insulin administration elicits a significant increase in plasma norepinephrine, a response that also increases between late gestation and after birth. This adrenomedullary response is linked to circulating cortisol concentrations and maturation of the adrenal gland pre- and postpartum.17, 31 The catecholamine response to hypoglycemia in newborn foals (aged 4–6 h), although greater than in the fetus, is not so apparent compared to older foals (7 days) which have a more rapid onset of hypoglycemia and greater release of epinephrine.17 The diminished adrenomedullary sensitivity in newborn foals is associated with a de-
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gree of insulin resistance (see below) and has been attributed to the high sympathoadrenal stimulation that occurs during delivery. The catecholamine stress response increases with postnatal age as the sympathoadrenal system continues to mature over the first few weeks after birth, but basal concentrations of catecholamines change little over this time.16, 17
Thyroid gland Thyroid hormones affect many physiological functions by regulating oxidative metabolic pathways such as ATP synthesis and metabolism and, therefore, play a key role in energy metabolism and thermogenesis. They are also involved in the differentiation and maturation of the nervous system, muscular and skeletal growth, and lung maturation. Consequently, a functional thyroid gland after birth is essential for postnatal adaptation and survival. The thyroid gland synthesizes mainly thyroxine [T4] and triiodothyronine [T3], with lesser quantities of reverse T3 [rT3] (Figure 7.3); all thyroid hormones contain iodine, which is essential for normal thyroid function. Most of the circulating T3 originates from peripheral monodeiodination of T4 controlled by three deiodinating enzymes (types I, II, and III) which are preferentially expressed in different tissues, notably the liver, kidneys and brain, but also the placenta (Table 7.2).32, 33 These enzymes are selenoproteins which require selenium for normal activity. More than 99% of the circulating T4 and T3 is protein bound, primarily to thyroid-binding globulin (TBG) but also to transthyretin and albumin. Concentrations of TBG are substantially higher in the foal compared to the adult horse and to other species, so that total plasma thyroid hormone concentrations in the neonatal foal are some of the highest reported.34, 35 Thyroid hormone concentrations are regulated by a negative feedback loop with thyroid-releasing hormone from the hypothalamus, and thyroidstimulating hormone (TSH) from the pituitary, being released when thyroid hormone concentrations are low. In general, thyroid hormone concentrations are lower during gestation compared to after birth, because metabolism is lower and thermoregulation is controlled primarily by the dam. However, at key stages during fetal and postnatal life, there are surges in thyroid hormones and their receptor concentrations in various tissues, the timing of which correlates with the level of maturity achieved by a particular species at birth.32 In the equine fetus, plasma T3 and T4 concentrations gradually rise from 1.0 to 3.5 nmol/L (0.65 to 2.3 ng/mL) and from 40 to 350 nmol/L (31 to 272 ng/mL), respectively, between 5 and 11 months
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Figure 7.3 Synthesis, transport, and action of thyroid hormones in the neonatal foal.
of gestation, associated with the tissue maturational effects, musculoskeletal development, and a rise in the absolute rate of oxygen consumption.36, 37 The major surge in T3 concentrations in foals occurs just prior to parturition, concurrent with the rise in fetal plasma cortisol concentrations; moreover, infusion with exogenous ACTH in late gestation stimulates a rise in both fetal cortisol and T3.38 At birth, foal umbilical cord concentrations of total T4 and T3 are approximately 400–500 nmol/L (311–389 ng/mL) and 8–12 nmol/L (5.2–7.8 ng/mL) respectively, and
T3 concentrations continue to rise for about 10 h after birth.34, 38 These values are higher than in neonates of other species in any physiological state.34 Although T3 and T4 gradually decline from birth, levels remain higher than the adult for up to 4 months of age, associated with a higher resting metabolic rate per kg body mass.35,39–41 The high concentrations of thyroid hormones at birth are important for many of the metabolic changes that take place in the early postnatal period. The metabolic rate rises from about 7 mL/kg/min
Table 7.2 Distribution, role, and activity of the thyroid deiodinating enzymes during fetal and neonatal life Deiodinase type 1
Deiodinase type 2
Tissue distribution
Liver, kidney, thyroid
Brain, pituitary, brown fat, thyroid, Placenta, fetal, tissues, uterus, skeletal muscle developing brain, skin
Primary role
Production of T4 from rT3, and T3 from T4, mainly for the circulating T3 pool
Production of T3 from T4, primarily for intracellular use
Inactivation of T4 and T3 to rT3 and T2 (inactivation by hepatic sulfation also important)
Activity during fetal life
Mostly low
Increases locally when intracellular T3 required
High activity. Decreases when more intracellular T3 required
Activity during neonatal life
Upregulated by hyperthyroidism
Upregulated by hypothyroidism, downregulated by hyperthyroidism
Decreases after birth so plasma T3 increases. Upregulated by hyperthyroidism
T4, thyroxine; T3, triiodothyronine; rT3, reverse T3; T2, 3,5-diiodo-L-thyronine.
Deiodinase type 3
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in the fetus to 19.5 mL/kg/min at birth, associated with thermoregulation and respiratory work. Thyroid hormones stimulate heat production (thermogenesis), which is essential for survival in the colder environment ex utero. Exposure to cold after birth is a major stimulus for thyroid production42 as is separation from the placenta, because of placental factors (adenosine, PGE) which inhibit thyroid hormones and thermogenesis in utero.43, 44 Thyroid activity is also influenced by mode of delivery, for example higher T3 concentrations are observed in lambs delivered per vaginam than those born by cesarean section, which clearly may affect the viability of the neonate.45 These factors have not been investigated in foals, but the rise in T3 during the first 30 min after spontaneous vaginal delivery is certainly correlated with high metabolic rates, which increase the foal’s rectal temperature by > 1◦ C (Figure 7.4).38, 41 Heat is produced by oxidative metabolic pathways in the mitochondria of various tissues, but
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this process is most efficient in brown adipose tissue (BAT) due to the action of a unique uncoupling protein (UCP1) which short-circuits the proton electrochemical gradient to produce heat. This process is termed non-shivering thermogenesis (NST) and is stimulated by thyroid hormones and norepinephrine.46, 47 NST predominates in the neonate of other species. However, the foal appears to have little BAT, while UCP1 concentrations are low in white adipose tissue, suggesting that NST may not be the primary thermogenic mechanism in the foal (Ousey and Symmonds, unpublished). Instead, shivering thermogenesis can increase the foal’s metabolic rate by twofold and is observed in wet, newborn foals and in older, dry foals exposed to cool (< 10◦ C) air temperatures.41, 48 Thyroid hormones are also important for musculoskeletal development, lung maturation, and neural development, as demonstrated by congenitally hypothyroid foals and thyroidectomized equine fetuses.49, 50 These foals, delivered at term or post-term, have low T3 and T4 concentrations with incomplete ossification of the skeleton, tendon laxity, poor mentation, and poor locomotor skills.49–51 Although respiratory distress is not the primary clinical feature of hypothyroid foals, it has been reported in septic foals and in premature foals associated with low T3 concentrations.19, 52 The importance of thyroid hormones for lung maturation has been demonstrated in sheep fetuses infused with T3, which increases pulmonary blood flow by 90% and increases fetal lung gas exchange surface area and lung compliance.53, 54 Similarly, infants with birth asphyxia and hypoxicischemic encephalopathy (HIE) have lower concentrations of T3, T4, and TSH at birth than healthy infants.55 Thyroid hormones are also essential for normal brain development and, in many species including humans, an iodine-deficient diet during early pregnancy causes low maternal T4 concentrations and neurological disabilities in the neonate.56 Irvine34 suggests that the degree of myelination is a good indicator of brain maturity, and certainly the foal which is precocious at birth has higher concentrations of thyroid hormones and a greater degree of myelination than infants, pigs, and rodents which are relatively immature at birth. The locomotor and neurological skills required to stand and suck indicate the advanced state of neural (and hence thyroid) development needed for postnatal survival in healthy, neonatal foals.57
The endocrine pancreas Pancreatic β-cells release insulin in order to facilitate the utilization of glucose. Glucose is an essential metabolic fuel for the brain, liver, and other
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tissues. Before birth, circulating glucose concentrations in the fetus are primarily dependent on the maternal glucose supply and its transport across the placenta.58 This is by facilitated diffusion using two placental glucose transporter enzymes, GLUT 1 and GLUT 3, which act in sequence, and requires a concentration gradient between the maternal (glucose ≈ 6 mmol/L or 108 mg/dL [fed]) and fetal (≈ 3 mmol/L or 54 mg/dL) circulations.31, 59 The main role of the fetal endocrine pancreas is to match the rate of fetal glucose utilization with the glucose supply. Therefore, basal insulin concentrations (40–50 pmol/L or 232–290 pg/mL) are related to glucose concentrations and normally do not change with gestational age.60 Approximately 70% of the glucose supplied to the uterus is utilized within the utero-placental tissues (on a weight-specific basis), which decreases the proportion of glucose reaching the fetus. Because its own glucogenic capacity is low, the fetus relies heavily on maternal glucose, and is vulnerable to hypoglycemia when maternal nutrient intake is impaired.61 Under such conditions, the fetus becomes less able to maintain plasma glucose concentrations as gestation proceeds. As the fetus grows, it has an increased requirement for glucose. Moreover, it needs to store energy in preparation for birth. In order to meet these increased nutritional demands, a number of morphological and physiological changes (increased nutrient delivery, increased uterine blood flow) occur within the uteroplacental tissues to ensure that more glucose reaches the fetus, which doubles its body mass during the last trimester of gestation.62, 63 Despite these adjustments, the maximum rate at which the placenta can supply glucose to the growing fetus is reached by about 300 days of gestation so that, after this time, the proportion of glucose taken up by the fetus decreases.37, 64 The glucogenic capacity of the equine fetus develops late in gestation as hepatic and renal concentrations of glucose-6-phosphatase (G-6-Pase), the key enzyme for gluconeogenesis and glycogenolysis, are induced by rising fetal cortisol concentrations.65 However, there is little excess glucose available for storage, and consequently hepatic glucose concentrations are relatively low in the newborn foal compared to other species. Once delivered and separated from its placental glucose supply, endogenous glucose is provided by glycogenolysis from hepatic stores.66 In healthy foals, glucose concentrations rise above fetal levels and reach 4–5 mmol/L (72–91 mg/dL) within 30 min of birth, accompanied by an increase in plasma insulin (from 70 pmol/L to 170 pmol/L; 406–987 pg/mL). The importance of carbohydrate as a metabolic fuel is demonstrated by the high non-protein respiratory quotient, RQ (> 0.9), immediately after birth.41 RQ
measures the inherent composition and utilization of carbohydrates, fats, and proteins as they are converted to energy substrate units that can be used by the body as energy. The rise in plasma glucose is stimulated by high plasma concentrations of catecholamine and/or glucagon (> 1500 pg/mL) released from the pancreatic α-cells, at birth.67 Glycogen stored in heart, lung, and muscles cannot contribute to the circulating glucose pool because these tissues do not possess G-6-Pase; instead, this substrate provides energy for local use within the tissue, for example, shivering thermogenesis in muscles. Because the hepatic glycogen stores are low and the foal’s metabolic rate is high, stored carbohydrate is rapidly depleted, within an hour after birth. However, plasma glucose levels are maintained in healthy foals because fatty acids are used for oxidative metabolism and replace carbohydrate as a metabolic fuel, demonstrated by a decline in the RQ (< 0.8) after 30 min postpartum. In healthy foals, there is sufficient non-structural body fat to provide energy for more than 24 h and, even in the absence of feeding, blood glucose concentrations are maintained.68, 69 Feeding is a major stimulus for pancreatic endocrine activity after birth. The postnatal change in nutrient delivery from continuous, parenteral nutrients to intermittent, enteral nutrition requires functional pancreatic β-cells to respond to the fluctuating glucose supply (from milk lactose). Pancreatic β-cell activity develops gradually during the last trimester; the plasma insulin response to intravenous glucose becomes progressively greater in older fetuses, a process which correlates more with the circulating cortisol concentrations than with gestational age.70 At birth, β-cell sensitivity is well developed and, therefore, the neonatal foal increases its circulating insulin following feeding (Figure 7.5).71, 72 Enteral feeding elicits a gradual (over 15–30 min) but significant rise in both plasma glucose and insulin concentrations in healthy foals, whereas intravenous glucose causes a more rapid response (5–15 min). The increment in postprandial insulin concentrations is evident within a few hours of birth. However, glucose clearance is slower compared to subsequently, suggesting that newborn foals may be insulin resistant, possibly because of the high concentrations of glucagon and catecholamines released at birth, which promote gluconeogenesis and reduce clearance of glucose from the circulation.71 This metabolic adjustment may help to maintain plasma glucose concentrations at a time when sucking patterns have not been fully established, and endogenous glucose stores have been depleted. The ontogenic increase in β-cell sensitivity continues to develop gradually after birth, as postprandial
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insulin concentrations are greater in 1- and 3-monthold foals and adult horses compared to foals aged 1 day and 1 week.66, 71, 73, 74
Gastrointestinal tract The gastrointestinal (GI) tract is a major endocrine organ producing a number of regulatory peptides from the mucosal cells lining the tract and from the enteric nervous system. These gut hormones are released in response to food entering the stomach or lumen of the GI tract, and may act in a systemic, paracrine, or neurocrine manner. During fetal life, there is gradual morphological maturation of the GI tract, with a cluster of tissue maturational changes that occur in the perinatal period.75 The specific nature of these changes is dependent upon the stage of development at which the animal is born, its postnatal feeding pat-
terns, and the nutrient and non-nutrient factors supplied in the milk.76 In the human fetus the majority of nutrients are delivered parenterally, but it is estimated that amniotic fluid contributes about 10–20% of the fetal energy requirements in late gestation and is essential for growth and differentiation of the GI tract and overall fetal growth.77, 78 A similar process is likely to occur in the horse fetus because equine amniotic fluid contains 10–20 mg/dL glucose (total amniotic and allantoic fluid volume 8–41 L) and the fetal stomach, which is filled with amniotic fluid, grows with gestational age.79–81 In the perinatal period, the GI tract undergoes enhanced growth and morphological (proliferation and migration of crypt cells) and functional (production of digestive enzymes) differentiation, stimulated by increasing circulating cortisol concentrations.82
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A greater stimulus to GI development postnatally is the onset of enteral feeding, which promotes proportionately greater growth of the GI tract compared to the increase in total body mass.83, 84 Feeding with colostrum increases the intestinal mass and length and height of the villi, changes which are diminished in neonatal animals receiving water, lactose solution or milk formula, whilst an absence of enteral feeding causes villous atrophy.85–87 These morphological changes are stimulated by the abundant bioactive factors (growth factors, hormones, and other factors) found in colostrum and milk.87, 88 Potent mitogens such as insulin-like growth factor (IGF) and epidermal growth factor (EGF) and other immunoprotective factors (immunoglobulins, lysozyme, lactoferrin) have been detected in equine mammary secretions, with the highest concentrations found in colostrum compared to milk produced later in lactation.89–91 Preliminary evidence in healthy foals suggests that formula feeding or feeding colostrum substitute devoid of these growth factors, can affect GI development by reducing digestible energy levels for up to 4 days postpartum.92 Gut hormones Gut hormones circulate in the bloodstream of foals and are released from different locations within the GI tract in response to feeding.93 They alter many aspects of GI function including motility, gastric acid secretion, gastric emptying, inhibition or secretion of enzymes, GI growth, smooth muscle function, and neural activity. Basal levels of plasma gut hormone concentrations are modified by diet and postnatal age associated with maturation of the GI tract.94–97 For example, gastrin promotes trophic development of the GI tract and, therefore, concentrations are highest at birth (neonatal hypergastrinemia), but decline with postnatal age.74, 98, 99 Feeding stimulates transient changes in gut hormones (Figure 7.5). Gastrin, which is released from the parietal cells of the stomach into the bloodstream, stimulates release of gastric acid (HCl) and pepsinogen for protein degradation.74 Consequently, gastrin concentrations rise following feeding. However, after the first feed (2–3 h after birth), gastrin concentrations are low compared to older foals (Figure 7.5) and are associated with a more alkaline gastric pH.72, 100 These low gastrin levels are important because, together with antitrypsin factor found in colostrum, they help prevent degradation of immunoglobulins within the stomach.101 Similar ontogenic changes are observed in pancreatic polypeptide (PP) concentrations. This hormone is released by the PP cells of the endocrine pancreas and is a potent inhibitor of pancreatic exocrine secretions (trypsin, amylase, lipase) and stimulates gastric emptying. PP
concentrations are significantly higher following the first feed in newborn foals compared to subsequent feeds during the first week postpartum.72 High PP concentrations at birth may promote the rapid transit of immunoglobulins intact into the small intestine. Plasma enteroglucagon (or glicentin) concentrations also tend to increase in foals with postnatal age. Enteroglucagon, released from the ileum and colon, is an important trophic factor for the gut and probably stimulates GI tract growth in the early neonatal period.102 From a clinical viewpoint, much interest has centered upon GI hormones and ‘minimal enteral feeding’ or ‘trophic feeding’ whereby micro-quantities of milk (12–24 mL/kg/day) are introduced into the GI tract of premature infants or those with compromised GI function, to promote cellular changes and release of gut hormones, to provide growth factors, and to support optimal digestion of nutrients without overloading the GI tract.103 Studies in infants have demonstrated that those individuals receiving trophic feeds in combination with parenteral nutrition have increased body growth, reduced parenteral intakes, tolerate full milk feeds earlier, and are discharged from hospital earlier than controls receiving TPN (total parenteral nutrition) only.104 In premature infants significant increases in plasma gut hormones (enteroglucagon, gastrin, gastric inhibitory peptide [GIP]) were observed after a cumulative volumes of 12 mL/kg of enteral milk feeding, effects that were absent in infants receiving only TPN.105 In healthy piglets, enteral feed intakes of 40 to 60% total daily requirement stimulate mucosal proliferation and growth of the GI tract and increases in several gut hormones (glucagon-like peptide 2, GIP, and peptide YY) which are important mediators of intestinal growth.106 These feed volumes may be higher than those needed for trophic effects on the GI tract because the piglets were fed an enteral nutrient solution rather than milk. There is little information about the effects of trophic feeding on gut hormones in foals, but intakes of 25 mL/h for a 50 kg foal have been recommended as a supportive measure for sick foals.107
Kidneys The kidneys control water and electrolyte homeostasis and have marked effects on blood pressure, via the renin–angiotensin–aldosterone system (RAAS; see Figure 7.6). Renin, released by the juxtaglomerular cells, is stimulated by a decrease in renal perfusion pressure through systemic hypotension. Renin stimulates conversion of angiotensin 1 to angiotensin 2 in the liver via angiotensin-converting enzyme (ACE)
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Figure 7.6 Hormone pathways involved in the renin–angiotensin–aldosterone system (RAAS) controlling electrolyte balance and water homeostasis. + indicates positive/stimulatory response; – indicates negative or inhibitory response; Na+ sodium; K+ potassium; Cl- chloride; ↑ increase; ↓ decrease.
which is located mainly in the lungs but also in the kidneys and, during pregnancy, the placenta.108 Angiotensin 2 stimulates the release of aldosterone from the adrenal cortex and vasopressin (antidiuretic hormone) from the pituitary. Aldosterone is important for sodium and hence water resorption in the kidneys, whilst vasopressin causes vasoconstriction and retention of water and sodium. This leads to improved renal perfusion which then has a negative feedback effect and suppresses renin release. Angiotensin 2 also causes vasoconstriction and a rise in blood pressure but this effect is moderated, in part, by intra-renal release of prostaglandin E2 and prostacyclin which are potent vasodilators.109 Collectively these mechanisms ensure that renal perfusion and glomerular filtration rate (GFR) are maintained within the normal range. Blood volume and vessel tone also affect GFR and hence blood pressure and these are regulated by many factors including atrial natriuretic peptide (ANP) which causes diuresis (water loss), natriuresis (sodium loss) and reduces blood volume/pressure. In healthy neonatal foals, average GFR and renal plasma flow are approximately 2.15 mL/kg/min and 16 mL/kg/min, respectively, values which are similar to or even higher than those of adult horses, suggesting that renal function is mature in neonatal foals.110–112 In pony fetuses the maximum number of glomeruli is achieved before birth, and fetal urine is produced from mid gestation, indicated by rising
concentrations of creatinine in the allantoic fluid with gestational age.81,113–115 Allantoic fluid is the major reservoir for fetal urine, excreted via the urachus, and reflects, to a large extent, the functional development of the fetal kidneys. For example, allantoic sodium concentrations decline during the last month of gestation as high fetal aldosterone concentrations stimulate renal retention of sodium (see below). These observations indicate that the fetal kidneys and RAAS are functional during gestation and become increasingly active towards term. Aldosterone concentrations are 400–500 pg/mL in late-gestation fetuses and foals at birth, but then decline. By 4–10 days of age, aldosterone concentrations are approximately 50–150 pg/mL, similar to adult values.116 The high aldosterone concentrations at birth help to maintain electrolyte concentrations and minimize sodium loss, although this system may not be completely developed at birth (see below). Fetal plasma vasopressin concentrations are low for much of gestation (< 5 pg/mL), but increase substantially after 300 days (20–25 pg/mL), peaking at birth and then declining to 2–4 pg/mL during the first 2 weeks postpartum.16, 27 The rise in vasopressin is associated with maturation of the fetal HPA axis and is positively correlated with rising fetal plasma cortisol concentrations and fetal blood pressure.27, 117 The high levels at birth contribute to the hemodynamic response of the newborn foal to rising fetal blood pressure and hypoxemia/stress during delivery;
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vasopressin may also help to decrease lung liquid production and promote lung liquid reabsorption.28, 118 Plasma ACE concentrations are low (< 80 U/L) until the last few weeks or days before delivery, when concentrations rise (100–150 U/L) in conjunction with the increase in fetal cortisol.117 It is likely that, for much of gestation, the circulating ACE originates from the placenta, as in other species. However, the prepartum rise in plasma ACE probably reflects the onset of pulmonary ACE production because pulmonary tissue concentrations of ACE are approximately five- to tenfold greater than renal ACE concentrations in the perinatal period.119 High pulmonary ACE concentrations may compensate, in part, for the loss of placental ACE and help to stimulate vascularization of the lungs at birth,120 while the high circulating ACE concentrations may contribute to the rise in fetal blood pressure through activity of the fetal RAAS, which also secretes renin and angiotensin 2 (Figure 7.6).117 The high pulmonary ACE concentrations, together with aldosterone and vasopressin at birth, help to regulate normal RAAS function in the early postnatal period, while the subsequent decline in these hormones may indicate downregulation of the RAAS once the major renal and cardiovascular adaptations at parturition have occurred. In healthy foals, the RAAS appears to be fully functional by birth. Neonatal foals usually produce about 6–8 liters/day of dilute urine (SG < 1.008) and certainly challenge with furosemide (which blocks chloride and sodium reabsorption in the ascending loop of Henle, leading to sodium and water loss) stimulates the expected rise in both plasma renin substrate and aldosterone concentrations, in the short term.121 However, after 6 h, serum sodium concentrations decline, reaching significantly lower concentrations than baseline. Therefore, although the hormonal responses of the RAAS are active after birth, there is some evidence of immaturity of tubular function even in healthy foals. The normal fractional excretion rates of electrolytes (i.e., those that are ultimately excreted in urine and lost from the body) for neonatal foals are higher for potassium, calcium, and phosphorus and lower for sodium compared to the adult horse, also suggesting that foals are susceptible to electrolyte imbalances.110 Certainly, over 24 h, there is increased renal clearance of sodium (hypernatremia) with a net potassium loss during intravenous isotonic electrolyte infusion.68, 122 Sick foals are susceptible to renal/circulatory disorders.122 Foals diagnosed with hypovolemia show a significant decline in aldosterone with fluid resuscitation, but the expected decline in vasopressin does not
occur whilst ANP concentrations are unchanged.123 These responses to fluid therapy differ to those observed in adult horses. Stereological studies also have shown that the kidney (cortical and glomerular) volume continues to increase after birth, and renal ACE concentrations are low for up to 2 weeks postpartum, suggesting that the physiological responses of the kidney may not be fully mature in newborn foals and may continue to develop for some time after birth.115, 119
Conclusion This chapter has highlighted some of the key hormonal events that occur in the perinatal period that are important for postnatal survival of the foal. Many of the major physiological adjustments that take place are related to the endocrine activity of the HPA axis, which is the driving force for organ maturation and the successful transition from fetal to neonatal life. Other chapters in this book will describe the clinical outcomes associated with abnormal endocrine events after birth and the devastating effects these can have on the health of the neonatal foal.
References 1. Fowden AL, Forhead AJ. Endocrine mechanisms of intrauterine programming. Reproduction 2004;127:515–26. 2. Fowden AL, Forhead AJ, Ousey JC. The endocrinology of equine parturition. Exp Clin Endocrinol Diabetes 2008;116:393–403. 3. Chavatte P, Rossdale PD, Tait AD. Corticosteroid synthesis by the equine fetal adrenal. Biol Reprod Mono 1995;1:13–20. 4. Han X, Fowden AL, Silver M, Holdstock N, McGladdery AJ, Ousey JC, Allen WR, Rossdale PD, Challis JR. Immunohistochemical localisation of steroidogenic enzymes and phenylethanolamine-N-methyl-transferase (PNMT) in the adrenal gland of the fetal and newborn foal. Equine Vet J 1995;27:140–6. 5. Silver M. Prenatal maturation, the timing of birth and how it may be regulated in domestic animals. Exp Physiol 1990;75:285–307. 6. Silver M, Ousey JC, Dudan FE, Fowden AL, Knox J, Cash RS, Rossdale PD. Studies on equine prematurity 2: Postnatal adrenocortical activity in relation to plasma adrenocorticotrophic hormone and catecholamine levels in term and premature foals. Equine Vet J 1984;16:278–86. 7. Fowden AL, Silver M. Comparative development of the pituitary-adrenal axis in the fetal foal and lamb. Reprod Dom Anim 1995;30:170–7. 8. Cudd TA, LeBlanc M, Silver M, Norman W, Madison J, Keller-Wood M, Wood CE. Ontogeny and ultradian rhythms of adrenocorticotropin and cortisol in the lategestation fetal horse. J Endocrinol 1995;144:271–83. 9. Ousey JC, Rossdale PD, Fowden AL, Palmer L, Turnbull C, Allen WR. Effects of manipulating intrauterine growth on postnatal adrenocortical development and other parameters of maturity in neonatal foals. Equine Vet J 2004;36:616–21.
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Endocrinological Adaptation 10. Rossdale PD, Silver M, Ellis L, Frauenfelder H. Response of the adrenal cortex to tetracosactrin (ACTH1–24 ) in the premature and full-term foal. J Reprod Fertil Suppl 1982;32:545–53. 11. Webb PD, Steven DH. Development of the adrenal cortex in the fetal foal: an ultrastructural study. J Dev Physiol 1981;3:59–73. 12. Chavatte P, Holtan D, Ousey JC, Rossdale PD. Biosynthesis and possible biological roles of progestagens during equine pregnancy and in the newborn foal. Equine Vet J Suppl, 1995;(24):89–95. 13. Holtan DW, Houghton E, Silver M, Fowden AL, Ousey J, Rossdale PD. Plasma progestagens in the mare, fetus and newborn foal. J Reprod Fertil Suppl 1991;44:517–28. 14. Rossdale PD, McGladdery AJ, Ousey JC, Holdstock N, Grainger L, Houghton E. Increase in plasma progestagen concentrations in the mare after foetal injection with CRH, ACTH or betamethasone in late gestation. Equine Vet J 1992;24:347–50. 15. Silver M, Fowden AL. Prepartum adrenocortical maturation in the fetal foal: responses to ACTH. J Endocrinol 1994;142:417–25. 16. O’Connor SJ, Gardner DS, Ousey JC, Holdstock N, Rossdale P, Edwards CM, Fowden AL, Giussani DA. Development of baroreflex and endocrine responses to hypotensive stress in newborn foals and lambs. Pflugers Arch 2005;450:298–306. 17. Silver M, Fowden AL, Knox J, Ousey JC, Franco R, Rossdale PD. Sympathoadrenal and other responses to hypoglycaemia in the young foal. J Reprod Fertil Suppl 1987;35:607–14. 18. Irvine CH, Alexander SL. Measurement of free cortisol and the capacity and association constant of cortisolbinding proteins in plasma of foals and adult horses. J Reprod Fertil Suppl 1987;35:19–24. 19. Lester G. Maturity of the neonatal foal. Vet Clin North Am Equine Pract 2005;21:333–55. 20. Houghton E, Holtan D, Grainger L, Voller BE, Rossdale PD, Ousey JC. Plasma progestagen concentrations in the normal and dysmature newborn foal. J Reprod Fert Suppl 1991;44:609–17. 21. Rossdale PD, Ousey JC, McGladdery AJ, Prandi S, Holdstock N, Grainger L, Houghton E. A retrospective study of increased plasma progestagen concentrations in compromised neonatal foals. Reprod Fertil Dev 1995;7:567–75. 22. Gold JR, Divers TJ, Barton MH, Lamb SV, Place NJ, Mohammed HO, Bain FT. Plasma adrenocorticotropin, cortisol, and adrenocorticotropin/cortisol ratios in septic and normal-term foals. J Vet Intern Med 2007;21:791–6. 23. Hurcombe SD, Toribio RE, Slovis N, Kohn CW, Refsal K, Saville W, Mudge MC. Blood arginine vasopressin, adrenocorticotropin hormone, and cortisol concentrations at admission in septic and critically ill foals and their association with survival. J Vet Intern Med 2008;22:639– 47. 24. Brendemeuhl JP, Williams MA, Boosinger TR, Ruffin DC. Plasma progestagen, trioiodothyronine and cortisol concentrations in postdate gestation foals exposed in utero to the tall fescue endophyte Acremonium coenophialum. Biol Reprod Mono 1995;1:53–9. 25. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995;73:899–908. 26. Comline RS, Silver M. Catecholamine secretion by the adrenal medulla of the foetal and new-born foal. J Physiol 1971;216:659–82.
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27. Giussani DA, Forhead AJ, Fowden AL. Development of cardiovascular function in the horse fetus. J Physiol 2005;565:1019–30. 28. Birk E, Iwamoto HS, Heymann MA. Hormonal effects on circulatory changes during the perinatal period. Bailli`ere’s Clin Endocrinol Metab 1989;3:795–815. 29. Thomas WP, Madigan JE, Backus KQ, Powell WE. Systemic and pulmonary haemodynamics in normal neonatal foals. J Reprod Fertil Suppl 1987;35:623–8. 30. Rose RJ, Rossdale PD, Leadon DP. Blood gas and acid–base status in spontaneously delivered, terminduced and induced premature foals. J Reprod Fertil Suppl 1982;32:521–8. 31. Silver M, Fowden AL. Sympathoadrenal and other endocrine and metabolic responses to hypoglycaemia in the fetal foal during late gestation. Exp Physiol 1995;80:651–62. 32. Fisher DA, Polk DH. Development of the thyroid. Bailli`ere’s Clin Endocrinol Metab 1989;3:627–57. 33. Silvestri E, Schiavo L, Lombardi A, Goglia F. Thyroid hormones as molecular determinants of thermogenesis. Acta Physiol Scand 2005;184:265–83. 34. Irvine C. Hypothyroidism in the foal. Equine Vet J Suppl 1984;16:302–6. 35. Irvine C, Evans M. Postnatal changes in total and free thyroxine and triiodothyronine in foal serum. J Reprod Fertil Suppl 1975;23:709–15. 36. Allen AL, Doige CE, Haas SD, Card CE, Fretz PB. Concentrations of triiodothyronine and thyroxine in equine fetal serum during gestation. Biol Reprod Mono 1995;1:49–52. 37. Fowden AL, Forhead AJ, White KL, Taylor PM. Equine uteroplacental metabolism at mid- and late gestation. Exp Physiol 2000;85:539–45. 38. Silver M, Fowden AL, Knox J, Ousey J, Cash R, Rossdale PD. Relationship between circulating tri-iodothyronine and cortisol in the perinatal period in the foal. J Reprod Fertil Suppl 1991;44:619–26. 39. Chen CL, Riley AM. Serum thyroxine and triiodothyronine concentrations in neonatal foals and mature horses. Am J Vet Res 1981;42:1415–17. 40. Murray MJ, Luba NK. Plasma gastrin and somatostatin, and serum thyroxine (T4), triiodothyronine (T3), reverse triiodothyronine (rT3) and cortisol concentrations in foals from birth to 28 days of age. Equine Vet J 1993;25:237–9. 41. Ousey JC, McArthur AJ, Rossdale PD. Metabolic changes in thoroughbred and pony foals during the first 24 h post partum. J Reprod Fertil Suppl 1991;44:561–70. 42. Clarke L, Darby CJ, Lomax MA, Symonds ME. Effect of ambient temperature during 1st day of life on thermoregulation in lambs delivered by cesarean section. J Appl Physiol 1994;76:1481–8. 43. Gunn TR, Gluckman PD. The endocrine control of the onset of thermogenesis at birth. Bailli`ere’s Clin Endocrinol Metab 1989;3:869–86. 44. Sack J, Beaudry M, DeLamater PV, Oh W, Fisher DA. Umbilical cord cutting triggers hypertriiodothyroninemia and nonshivering thermogenesis in the newborn lamb. Pediatr Res 1976;10:169. 45. Clarke L, Heasman L, Firth K, Symonds ME. Influence of route of delivery and ambient temperature on thermoregulation in newborn lambs. Am J Physiol 1997;272:R1931–9. 46. Silva JE. The thermogenic effect of thyroid hormone and its clinical implications. Ann Intern Med 2003;139:205–13. 47. Symonds ME, Bird JA, Clarke L, Gate JJ, Lomax MA. Nutrition, temperature and homeostasis during perinatal development. Exp Physiol 1995;80:907–40.
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48. Ousey JC, McArthur AJ, Murgatroyd PR, Stewart JH, Rossdale PD. Thermoregulation and total body insulation in the neonatal foal. J Therm Biol 1992;17:1–10. 49. Allen AL, Fretz PB, Card CE, Doige CE. The effects of partial thyroidectomy on the development of the equine fetus. Equine Vet J 1998;30:53–9. 50. Allen, A.L., Townsend, H.G., Doige, C.E. and Fretz, P.B. (1996) A case-control study of the congenital hypothyroidism and dysmaturity syndrome of foals. Can Vet J 1996;37:349–51; 354–8. 51. McLaughlin BG, Doige CE, McLaughlin PS. Thyroid hormone levels in foals with congenital musculoskeletal lesions. Can Vet J 1986;27:264–7. 52. Murray MJ. Hypothyroidism and respiratory insufficiency in a neonatal foal. J Am Vet Med Assoc 1990;197:1635–8. 53. Chan L, Miller TF, Yuxin J, Farina C, Chander A, Shaffer TH, Wolfson MR. Antenatal triiodothyronine improves neonatal pulmonary function in preterm lambs. J Soc Gynecol Invest 1998;5:122–6. 54. Lorijn RH, Nelson JC, Longo LD. Induced fetal hyperthyroidism: cardiac output and oxygen consumption. Am J Physiol 1980;239:H302–7. 55. Pereira DN, Procianoy RS. Effect of perinatal asphyxia on thyroid-stimulating hormone and thyroid hormone levels. Acta Paediatr 2003;92:339–45. 56. Morreale de Escobar G, Obregon MJ, Escobar del Rey F. Role of thyroid hormone during early brain development. Eur J Endocrinol 2004;151 Suppl 3:U25–37. 57. MacKay RJ. Neurologic disorders in neonatal foals. Vet Clin North Am Equine Pract 2005;21:387–406. 58. Fowden AL, Ousey JC, Forhead AJ. Comparative aspects of prepartum maturation: provision of nutrients. Pferdeheilkunde 2001;17:653–8. 59. Wooding FB, Fowden AL. Nutrient transfer across the equine placenta: correlation of structure and function. Equine Vet J 2006;38:175–83. 60. Fowden AL, Barnes RJ, Comline RS, Silver M. Pancreatic beta-cell function in the fetal foal and mare. J Endocrinol 1980;87:293–301. 61. Fowden AL, Taylor PM, White KL, Forhead AJ. Ontogenic and nutritionally induced changes in fetal metabolism in the horse. J Physiol 2000;528(Pt 1):209–19. 62. Fowden AL, Ousey JC, Forhead AJ. Nutritive and endocrine functions of the equine placenta. In: Workshop on the Equine Placenta. Powell DG, Hale FD, Hale G (eds). Kentucky Agricultural Experiment Station, Kentucky, 2004; pp. 36–41. 63. Platt H. Growth of the equine foetus. Equine Vet J 1984;16:247–52. 64. Macdonald AA, Chavatte P, Fowden AL. Scanning electron microscopy of the microcotyledonary placenta of the horse (Equus caballus) in the latter half of gestation. Placenta 2000;21:565–74. 65. Fowden AL, Mundy L, Ousey JC, McGladdery A, Silver M. Tissue glycogen and glucose 6-phosphatase levels in fetal and newborn foals. J Reprod Fertil Suppl 1991;44:537–42. 66. Fowden AL, Comline RS, Silver M. Insulin secretion and carbohydrate metabolism during pregnancy in the mare. Equine Vet J 1984;16:239–46. 67. Fowden AL, Forhead AJ, Bloomfield M, Taylor PM, Silver M. Pancreatic alpha cell function in the fetal foal during late gestation. Exp Physiol 1999;84:697–705. 68. Buchanan BR, Sommardahl CS, Rohrbach BW, Andrews FM. Effect of a 24-hour infusion of an isotonic electrolyte replacement fluid on the renal clearance of elec-
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trolytes in healthy neonatal foals. J Am Vet Med Assoc 2005;227:1123–9. Ousey JC. Thermoregulation and the energy requirement of the newborn foal, with reference to prematurity. Equine Vet J Suppl 1997;(24):104–8. Fowden AL, Gardner DS, Ousey JC, Giussani DA, Forhead AJ. Maturation of pancreatic beta-cell function in the fetal horse during late gestation. J Endocrinol 2005;186:467–73. Holdstock NB, Allen VL, Bloomfield MR, Hales CN, Fowden AL. Development of insulin and proinsulin secretion in newborn pony foals. J Endocrinol 2004;181:469–76. Ousey JC, Ghatei M, Rossdale PD, Bloom SR. Gut hormone responses to feeding in healthy pony foals aged 0 to 7 days. Biol Reprod Mono 1995;1:87–96. Smyth GB, Young DW, Duran SH. Maturation of insulin and glucose responses to normal feeding in foals. Aust Vet J 1993;70:129–32. Smyth GB, Young DW, Schumacher J. Postprandial serum gastrin concentrations in normal foals. Equine Vet J 1989;21:285–7. Sangild PT. Gut responses to enteral nutrition in preterm infants and animals. Exp Biol Med (Maywood) 2006;231:1695–711. Weaver LT. Breast and gut: the relationship between lactating mammary function and neonatal gastrointestinal function. Proc Nutr Soc 1992;51:155–63. Mulvihill SJ, Stone MM, Debas HT, Fonkalsrud EW. The role of amniotic fluid in fetal nutrition. J Pediatr Surg 1985;20:668–72. Trahair JF. Is fetal enteral nutrition important for normal gastrointestinal growth? A discussion. J Parenter Enteral Nutr 1993;17:82–5. Adams-Brendemuehl C, Pipers FS. Antepartum evaluations of the equine fetus. J Reprod Fertil Suppl 1987;35:565–73. Bucca S, Fogarty U, Collins A, Small V. Assessment of feto-placental well-being in the mare from mid-gestation to term: transrectal and transabdominal ultrasonographic features. Theriogenology 2005;64:542–57. Paccamonti D, Swiderski C, Marx B, Gaunt S, Blouin D. Electrolytes and biochemical enzymes in amniotic and allantoic fluid of the equine fetus during late gestation. Biol Reprod Mono 1995;1:39–48. Trahair JF, Sangild PT. Systemic and luminal influences on the perinatal development of the gut. Equine Vet J Suppl 1997;24:40–50. Bjornvad CR, Schmidt M, Petersen YM, Jensen SK, Offenberg H, Elnif J, Sangild PT. Preterm birth makes the immature intestine sensitive to feeding-induced intestinal atrophy. Am J Physiol Regul Integr Comp Physiol 2005;289:R1212–22. Zabielski R, Le Huerou-Luron I, Guilloteau P. Development of gastrointestinal and pancreatic functions in mammalians (mainly bovine and porcine species): influence of age and ingested food. Reprod Nutr Dev 1999;39:5–26. Berseth CL. Breast-milk-enhanced intestinal and somatic growth in neonatal rats. Biol Neonate 1987;51:53–9. Widdowson EM. Development of the digestive system: comparative animal studies. Am J Clin Nutr 1985;41:384–90. Xu RJ. Development of the newborn GI tract and its relation to colostrum/milk intake: a review. Reprod Fertil Dev 1996;8:35–48. Koldovsky O. Hormonally active peptides in human milk. Acta Paediatr Suppl 1994;402:89–93.
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Endocrinological Adaptation 89. Hess-Dudan F, Vacher PY, Bruckmaier RM, Weishaupt MA, Burger D, Blum JW. Immunoreactive insulin-like growth factor I and insulin in blood plasma and milk of mares and in blood plasma of foals. Equine Vet J 1994;26:134–9. 90. Murray MJ, Schaudies RP, Cavey DM. Epidermal growth factor-like activity in mares’ milk. Am J Vet Res 1992;53:1729–31. 91. Schweigert FJ. Milk more than a nutrient: hormones, growth factors, and bioactive factors. Pferdeheilkunde 2001;17:665–8. 92. Ousey JC, Prandi S, Zimmer J, Holdstock N, Rossdale PD. Effects of various feeding regimens on the energy balance of equine neonates. Am J Vet Res 1997;58:1243–51. 93. Aynsley-Green A. The endocrinology of feeding in the newborn. Bailli`ere’s Clin Endocrinol Metab 1989;3:837– 68. 94. Aynsley-Green A, Lucas A, Bloom SR. The effect of feeds of differing composition on entero-insular hormone secretion in the first hours of life in human neonates. Acta Paediatr Scand 1979;68:265–70. 95. Guilloteau P, Le Huerou-Luron I, Chayvialle JA, Mouats A, Bernard C, Cuber JC, Burton J, Puigserver A, Toullec R. Plasma and tissue levels of digestive regulatory peptides during postnatal development and weaning in the calf. Reprod Nutr Dev 1992;32:285–96. 96. Huang CG, Eng J, Yalow RS. Ontogeny of immunoreactive cholecystokinin, vasoactive intestinal peptide, and secretin in guinea pig brain and gut. Endocrinology 1986;118:1096–101. 97. Lucas A, Bloom SR, Aynsley-Green A. Development of gut hormone responses to feeding in neonates. Arch Dis Child 1980;55:678–82. 98. Sangild PT, Hilsted L, Nexo E, Fowden AL, Silver M. Vaginal birth versus elective caesarean section: effects on gastric function in the neonate. Exp Physiol 1995;80:147– 57. 99. Zarate A, Ochoa R, Fonseca E, Moran C. Hypergastrinemia of newborns and infants during breast-feeding. J Soc Gynecol Invest 1995;2:531–4. 100. Baker SJ, Gerring EL. Gastric pH monitoring in healthy, suckling pony foals. Am J Vet Res 1993;54:959–64. 101. Saikku A, Koskinen E, Sandholme M. Sequential changes of IgG and antitrypsin in different compartments during colostral-intestinal transfusion of immunity to the newborn foal. J Vet Med (B) 1989;36:391–6. 102. Tadokoro R, Shimizu T, Hosaka A, Kaneko N, Satoh Y, Yamashiro Y. Postnatal and postprandial changes in plasma concentrations of glicentin in term and preterm infants. Acta Paediatr 2003;92:1175–9. 103. Burrin DG, Stoll B. Key nutrients and growth factors for the neonatal gastrointestinal tract. Clin Perinatol 2002;29:65–96. 104. McClure RJ, Newell SJ. Randomised controlled study of clinical outcome following trophic feeding. Arch Dis Child Fetal Neonatal Ed 2000;82:F29–33. 105. Lucas A, Bloom SR, Aynsley-Green A. Gut hormones and ‘minimal enteral feeding’. Acta Paediatr Scand 1986;75:719–23. 106. Burrin DG, Stoll B, Jiang R, Chang X, Hartmann B, Holst JJ, Greeley GH Jr, Reeds PJ. Minimal enteral nutrient requirements for intestinal growth in neonatal piglets: how much is enough? Am J Clin Nutr 2000;71:1603–10.
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107. Buechner-Maxwell VA. Nutritional support for neonatal foals. Vet Clin North Am Equine Pract 2005;21:487–510. 108. Poisner AM. The human placental renin-angiotensin system. Front Neuroendocrinol 1998;19:232–52. 109. Verlander JW. Renal Physiology: Glomerular filtration, 2nd edn. Philadelphia: Saunders, 1997; pp. 511–21. 110. Brewer BD, Clement SF, Lotz WS, Gronwall R. Renal clearance, urinary excretion of endogenous substances, and urinary diagnostic indices in healthy neonatal foals. J Vet Intern Med 1991;5:28–33. 111. Gonda KC, Wilcke JR, Crisman MV, Ward DL, Robertson JL, Finco DR, Braselton WE. Evaluation of iohexol clearance used to estimate glomerular filtration rate in clinically normal foals. Am J Vet Res 2003;64:1486–90. 112. Holdstock NB, Ousey JC, Rossdale PD. Glomerular filtration rate, effective renal plasma flow, blood pressure and pulse rate in the equine neonate during the first 10 days post partum. Equine Vet J 1998;30:335–43. 113. Beech DJ, Sibbons PD, Rossdale PD, Ousey JC, Holdstock NB, Chavatte P, Ansari T. Organogenesis of lung and kidney in Thoroughbreds and ponies. Equine Vet J 2001;33:438–45. 114. Holdstock NB, McGladdery AJ, Ousey JC, Rossdale PD. Assessing methods of collection and changes in selected biochemical constituents in amniotic and allantoic fluid throughout equine pregnancy. Biol Reprod Mono 1995;1:21–38. 115. Holdstock NB, Rossdale PD, Beech DJ, Ansari T, Sibbons PD. Microanatomical development of the equine kidney and defects associated with intra-uterine growth retardation. Pferdeheilkunde 2001;17:659–61. 116. Giry J, Safwate A, Barlet JP. Control of aldosterone secretion in domestic mammals during the perinatal period. Reprod Nutr Dev 1985;25:993–1005. 117. Forhead AJ, Broughton Pipkin F, Taylor PM, Baker K, Balouzet V, Giussani DA, Fowden AL. Developmental changes in blood pressure and the renin-angiotensin system in pony fetuses during the second half of gestation. J Reprod Fertil Suppl 2000;56:693–703. 118. Perks AM, Kindler PM, Marshall J, Woods B, Craddock M, Vonder Muhll I. Lung liquid production by in vitro lungs from fetal guinea pigs: effects of arginine vasopressin and arginine vasotocin. J Dev Physiol 1993;19:203–12. 119. O’Connor SJ, Fowden AL, Holdstock N, Giussani DA, Forhead AJ. Developmental changes in pulmonary and renal angiotensin-converting enzyme concentration in fetal and neonatal horses. Reprod Fertil Dev 2002;14:413–17. 120. Morrell NW, Grieshaber SS, Danilov SM, Majack RA, Stenmark KR. Developmental regulation of angiotensin converting enzyme and angiotensin type 1 receptor in the rat pulmonary circulation. Am J Respir Cell Mol Biol 1996;14:526–37. 121. Broughton Pipkin F, Ousey JC, Wallace CP, Rossdale PD. Studies on equine prematurity 4: Effect of salt and water loss on the renin-angiotensin-aldosterone system in the newborn foal. Equine Vet J 1984;16:292–7. 122. Corley KT. Monitoring and treating haemodynamic disturbances in critically ill neonatal foals. Part 1: Haemodynamic monitoring. Equine Vet Educ Manual 2003;6:68–77. 123. Hollis AR, Boston RC, Corley KT. Plasma aldosterone, vasopressin and atrial natriuretic peptide in hypovolaemia: a preliminary comparative study of neonatal and mature horses. Equine Vet J 2008;40:64–9.
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Chapter 8
The Placenta Peter R. Morresey
Introduction The placenta can be considered to have four major functions, all of which are essential for the maintenance of pregnancy. First, the placenta is responsible for nutrient synthesis. Second, the placenta facilitates nutrient transfer to the fetus and elimination of waste products by removal to the maternal circulation. Third, the placenta acts as an immunological barrier, between the fetus and the maternal immune system, as well as protecting the fetus from infectious agents. Fourth, the placenta acts as a temporary endocrine organ, producing and modifying hormones affecting both the fetus and the mare. Placental adequacy is vital to fetal viability, with normal development and function of the placenta dependent on maternal health, reproductive competence, and freedom from infectious or other pathologies. Diminished placental competency may result from deficiencies on either the maternal or fetal side of the uteroplacental interface. The importance of systematic and comprehensive placental evaluation (see Chapter 9) cannot be overemphasized. Invaluable clinically relevant information about the reproductive competency of the mare, the in utero environment of the fetus, and the perinatal health of the foal can be gained.
Historical aspects The placenta has been ascribed many properties throughout history, ranging from spiritual to bestowing protection from disease.1 The Egyptians viewed the placenta as essential to well-being, with great effort expended to preserve it throughout the life of the owner. Literature dating from the time of the ancient Greeks describes placental anatomy in detail, with information derived from mainly animal sources. The term placenta, originally used to describe the human discoid placenta by Columbus in 1559, did not acquire its current connotation until the late seventeenth century.
The equine placenta has been researched throughout history in a discontinuous fashion.2 Early observations date from Ruini’s Anatomia del Cavallo, published in 1598, where the diffuse nature of the equine placenta was first illustrated. The microcotyledons of the mature equine placenta were later described by Fabricius in De Formatu Foetu, published 1604. Separation of the maternal and fetal circulations (during human studies) was definitively proven by John Hunter in the eighteenth century. Advances in microscopy, beginning in the early nineteenth century, allowed the direct visualization of the microscopic vascular arrangement of the fetal maternal interface, the concept of deciduate and adeciduate placentation (loss and conservation of maternal tissue, respectively), and placental form as an evolutionary marker.3 The twentieth century saw the advent of endocrinological studies leading to the discovery of what was then known as pregnant mare serum gonadotropin (PMSG, now equine chorionic gonadotropin, eCG). Activity was first discovered in the serum of pregnant mares in contrast to the discovery of gonadotropins in the urine of pregnant human females. Further studies revealed that PMSG could be recovered from the endometrial cups formed apposing the allantochorionic girdle, demonstrating a placental origin.4 Considerable evaluation of equine placentas and investigation of abnormalities occurred in the 1970s.5, 6 Normal values for placental weight and component dimensions were established. Later, fetoplacental metabolism was elucidated by the use of chronically placed intrafetal catheters.7, 8 Placental efficiency as measured by weight of fetus produced per unit of placenta has also been calculated. More recent investigations have evaluated the microscopic structure of the maternal and fetal interface using electron microscopy.9 Influence of maternal size on conceptus development has been elegantly demonstrated by the use of between-breed embryo transfer.10 Functional
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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The Placenta exchange capacity between the endometrium and placenta has been further refined by the use of stereology to more fully assess the degree of association between the endometrium and chorionic epithelium.11 This has enabled investigation into other factors affecting maternal influence on placental development such as endometrial integrity.12
Anatomy The equine placenta is described as adeciduate, diffuse, microcotyledonary, and epitheliochorial, referring to the preservation of maternal tissue, shape of the fetal and maternal tissue apposition, method of placental attachment, and degree of invasiveness of the fetal tissues, respectively. Expulsion of the adeciduate equine placenta does not result in loss of maternal endometrial tissue. This is similar to cattle and pigs, but in contrast to the situation in dogs and cats where maternal tissue is eroded by placental formation. The equine placenta is further classified with respect to the shape of the area where physiological exchange occurs between the fetus and dam. The diffuse attachment of the equine allantochorion to the endometrial surface is similar to that of pigs and camelids, where nearly the entire allantochorion is involved. This is in contrast to ruminants (caruncular), primates (discoid), and carnivores (zonary). The microplacentome of the equine placenta is composed of fetal cotyledon and maternal caruncle, the fetal component being an extension of the villous projections of the allantochorion. These structures resemble the larger placentomes of ruminant species, but on a much smaller scale. Between these attachments, well-defined areolae or spaces, similar to those present in the porcine placenta, are present. Uterine glands deliver protein-rich secretions into the areolae for absorption by specialized overlying trophoblastic cells, thus allowing a second histiotrophic pathway for fetal nutrition.13 The equine placenta is classified as epitheliochorial. All three extraembryonic tissue layers derived from the fetus are present, being fetal vascular endothelium, mesodermal connective tissue, and epithelial-derived chorionic ectoderm.14 The corresponding maternal tissue layers remain intact, resulting in fetally derived chorionic epithelium apposed to maternal endometrial epithelium.
Embryological origins and development The capsule The early conceptus is surrounded by the capsule, a thin acellular layer deposited between the zona pellu-
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cida and the trophoblast at the time of blastocyst formation (around day 6). The capsule consists of glycoprotein. Both the trophoderm and the endometrium were thought to contribute to the origin of the capsule as it was absent during in vitro culture of precapsular embryos;15 however, xenografts of trophoderm were alone able to stimulate its formation.16 The capsule dissolves around day 21–22, exposing trophoblastic cells to the uterine environment, allowing them to migrate over the choriovitelline membrane and protrude into the adjacent endometrial glands.17 By allowing the early conceptus to maintain a spherical shape, the high level of mobility necessary for the extensive endometrial contact vital for pregnancy recognition is facilitated.
Choriovitelline placentation Aggregation of the embryonic endoderm on the inside of the trophoblast around day 8 completes the formation of the yolk sac. Encirclement of the blastocele and attachment of the yolk sac to the chorion completes around day 11 and constitutes the genesis of the choriovitelline placenta. This structure is responsible for maternal–fetal exchange via the vitelline veins during the initial period of gestation, ending approximately day 40.14 The vitelline artery and multiple blood vessels develop within the mesodermal tissue present between the chorion and yolk sac. Unlike the allantois, the yolk sac blood vessels are located on the endodermal surface.18 Rapid growth of the conceptus occurs between days 11 and 16, with a steady increase in volume of the yolk sac. Following fixation around day 16–17, growth plateaus until loss of the capsule around day 21–22.19, 20
Allantochorionic placentation The allantois emerges from the hindgut at approximately day 20–21 and grows rapidly to envelop the embryo from its dorsal aspect by day 22. Fusion of the allantois to the chorion begins formation of the allantochorionic placenta by approximately day 25.21 Trophoblastic cells invade the maternal endometrium around day 35 as a first step in the formation of the endometrial cups.20 By day 40, functional replacement of the choriovitelline placenta approaches completion, and following expansion the allantois occupies the majority of the embryonic vesicle.21 The allantoic sac, having now displaced the embryo to the opposite pole of the vesicle from where it began forming, and the yolk sac meet at the dorsal pole of the embryonic vesicle, forming the attachment of the umbilical cord.
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Non-invasive trophoblastic cells around day 40–45 begin to make stable, microvillous attachments to the endometrium. Over the next 20 days, blunt allantochorionic projections continue to engage with the endometrium. Beyond day 60, branching continues to increase in magnitude, becoming longer and deeper until the microcotyledon is formed.22 Placental interdigitation and growth is initiated by steroid hormones in concert with local growth factors.23 Fetal chorionic girdle cells were shown to rapidly proliferate by days 30–32 of gestation but stop dividing after invasion of the endometrium. Trophoblastic cells of the allantochorion increase in mitotic activity after day 38. Endometrial luminal epithelium proliferates only after the primary villi of the epitheliochorial placenta have been formed, with this effect seen only in the pregnant horn where placentation is further advanced by days 58–70.24 High concentrations of insulin-like growth factor-II messenger RNA (IGF-II mRNA) are present in the fetal-derived mesoderm, chorionic girdle and mature endometrial cup tissue.25 Transforming growth factor β1 (TGF β1) expression is increased in the maternal endometrium at the time of implantation, regulating endometrial and/or trophoblast growth and differentiation during placentation.26 Upregulation of endometrial epithelial growth factor (EGF) occurs during pregnancy in mares, with EGF receptors found in the fetal allantochorion and maternal endometrium, suggesting a role for EGF in the substantial growth of these two tissues during placentation.27 Endometrial glands remain functional throughout gestation and secrete nutrients in an exocrine fashion into the areola-like spaces between the microcotyledonary attachments. By day 45, the allantois has replaced the yolk sac’s contribution to placentation.21
Placental components Membranes
Allantochorion The allantochorion forms as a fusion of fetal allantois with the chorion. However, this has been shown not to be a homogenous structure. A basement membrane develops under the allantoic endoderm, isolated from the vascular layer of the allantois by a mesothelial layer and an exocelom-like space.18 This exocelom-like space persists until approximately the stage of villous formation (around day 45), after which it is diminished by collagen deposition with remnants remaining over larger allantoic vessels.20 The basement membrane supports the allantoic endoderm while the underlying space allows move-
ment between this endoderm and the allantoic vascular layer, which becomes more closely associated with the endometrial and exocelomic surfaces as pregnancy progresses.18 The allantochorion is also not uniform throughout its entire area. The tip of the pregnant horn differs histologically from the remainder of the pregnant horn. Growth retardation is evidenced by widespread remnants of the extraembryonic celom and hypoplastic villi caused by disordered formation of the microcotyledon. Resulting placental hypoxia and ischemia lead to trophoblast necrosis, squamous metaplasia, thickened basement membrane, and fibrosis of the villi.28
Amnion The amnion arises from the somatopleure in the embryo, attaching to the stalk of the yolk sac, which reduces to an umbilical ring as the fetus develops; however, the amnion is associated with the umbilical cord throughout the amnionic cavity. As the allantois develops it comes to envelop the amnion and fuses with it to form the allantoamnion. The fetus is then protected from the allantoic fluid, which contains urinelike fetal waste products.20 As pregnancy progresses, the epithelial linings of the allantoamnion progress from squamous to cuboidal, reflecting an increase in cellular activity. The basement membrane thickens, decreasing permeability and minimizing transfer between the allantoic and amnionic cavities. Plaques of glycogen appear on the fetal side of the membrane beginning around day 70. The vascularity of the allantoamnion also increases greatly during gestation.20 Microcotyledon The microplacentome is a union of the fetal microcotyledon and the maternal microcaruncle. This arrangement maximizes surface area contact between the fetal and maternal epithelial layers. Both maternal and fetal blood vessels indent the base of the adjacent epithelial layer.2 The microcotyledon is developed from several primary folds of trophoblast which become elaborately subdivided as gestation proceeds.22 Each microcotyledon can be considered a collection of villi, each supplied by a central artery and drained by two peripheral veins, with numerous branched communicating vessels. Microcotyledons become more regular and globular in shape as gestation progresses, increasing in height, breadth, and complexity. Trophoblastic cell height, opposing endometrial cell height, and distance between maternal and fetal capillaries all decreased with advancing gestational age.29
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The Placenta Microcotyledons have been previously considered to be fully formed by day 150 of gestation;30 however, recent observations showed lengthening and branching of microcotyledonary villi throughout gestation.9 Changes also occur in the structure of the maternal caruncles. Height of the maternal epithelium reduces between days 60 and 150 of gestation, with no corresponding decrease on the fetal side of the placenta. However, between days 100 and 250, indentation of maternal epithelium by fetal capillaries reduces the effective thickness of the placental barrier.22 Throughout the second half of gestation, high levels of mitotic activity are still present at the periphery of the microcotyledons, where growth is observed to continue.29 Vascular structures From early gestation to near term, considerable morphological changes occur in the vasculature of the microcotyledons. Additionally, integrity of the maternal endometrium affects development. In endometrotic mares, most of the microplacentomes were irregular in shape, of lesser length, and of larger diameter when compared to primiparous mares. Vascular density was also greater in the endometrotic mares, with bulb-like capillary networks at the base of the stem of each villus. In primiparous mares, the capillaries were arranged simply, mostly in straight lines along the axis of the villus, and with communications visible at irregular intervals. Simple and slightly more complicated side capillary loops could be seen along the whole length of the villi and at the top of the terminal villi. Most of the capillaries were characterized by zones containing dilated sinusoids, which increased the surface area for materno-fetal exchange. Therefore, age-related endometrial degenerative changes on microplacentome development include extent and intimacy of physical and hematological contact, thereby affecting fetal growth.29
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onic girdle is initially a band of tall columnar epithelial cells that change to stratified columnar and become indented by clefts and pits. Binucleate chorionic girdle cells invade the maternal endometrium 36–38 days post-ovulation.34 Adhesion to and penetration through the endometrial luminal epithelium occurs rapidly, but is initially in very limited areas with strands of trophoblast cells invading through the basal lamina into the stroma. Here the cells aggregate, giving rise to the endometrial cups. Only chorionic girdle cells adjacent to the basal lamina or that have entered the endometrial stroma hypertrophy and differentiate into cup cells. The luminal epithelium contains numerous lymphocytes at the initiation of trophoblastic invasion; however, by day 42 lymphocytes are largely confined to the periphery of the cup. Areas of girdle cells that do not attach, and superficial portions of areas of invasion, undergo necrosis.35 After day 70 the endometrial cups become pale due to degenerative changes. Necrotic surface tissue is sloughed, allowing regional endometrial glands to expel their content which is rich in eCG. Leukocytes invade the area and destroy the remaining cup cells. At around 100–120 days of gestation, the cup material is sloughed off the surface of the endometrium.34
Allantochorionic pouches Following maternal rejection of the endometrial cups they are sloughed into adjacent invaginations of the overlying allantochorion, the allantochorionic pouches. The lumen of this pouch can be demonstrated to communicate with the endometrial surface. Grossly, these appear as pedunculated structures forming a ring around the site of umbilical cord attachment at the dorsal wall of the pregnant horn. Remnants of the endometrial cup appears as brown viscous material within the pouch.6
Hippomane Fluids and compartments Normal allantoic fluid volume in the Thoroughbred mare ranges from 8 to 18 liters.31 An increase indicates a placental or histological endometrial abnormality. Normal amnionic fluid volumes range from 3 to 8 liters.31 Amniotic fluid differs from allantoic fluid in that it resembles an ultrafiltrate of serum.32 Specialized structures
Chorionic girdle and the endometrial cups The chorionic girdle forms around the conceptus, always in the region of contact between the expanding allantois and regressing yolk sac.33 The chori-
The hippomane, also known as the allantoic calculus, is uniformly present in the allantoic fluid. They are reported to not be found until around day 85 of gestation.32 Morphology has been extensively studied.36 Early hippomanes are small, white, and ovoid in shape. As gestation advances, hippomane color changes to match the changes of the allantoic fluid, becoming brown-olive. It is suspected that desquamated epithelial cells accumulating in the dependent portion of the uterus provide a nidus for the later formation of the hippomane. Increase in size is due to the deposition of an amorphous ground substance and cellular debris from the allantoic fluid. Chemical composition has also been investigated.32 The predominant substance is mucoprotein,
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with deposition of calcium phosphate. Small quantities of sodium, potassium, and magnesium are also present. The various concentrations reflect those found in the allantoic fluid. Umbilical cord The umbilical cord anchors the fetus to the original implantation site at the base of the gravid horn on the dorsal wall of the uterus. With expansion of the allantois and diminishing of the yolk sac during early development, the umbilical cord increases in length. As a result, the urachus also lengthens. As the yolk sac collapses it is reduced to a band of tissue adjacent to the umbilical vessels. The umbilical cord can be divided into two sections with regard to the attachment of the amnion near its midsection. The amnionic portion is attached nearest to the fetus. Contained within the amnionic portion, two umbilical arteries carry less-well oxygenated blood away from the fetal heart, one vein carries better-oxygenated blood towards the fetal heart (formed from an anastomosis of two veins proximally), the urachus connects the fetal bladder to the allantoic cavity for the expulsion of fetal urine, and the remnants of the yolk sac. By convention, vessels leading blood away from the heart are considered arteries, and vessels leading blood towards the heart are considered veins, regardless of their state of oxygenation. The allantoic portion attaches to the allantochorion and contains two umbilical arteries and two umbilical veins (which anastomose to form the single vein present in the amnionic portion). The two umbilical arteries perfuse separate areas of the allantochorion: one supplies the pregnant horn and cranial body, and the other supplies the non-pregnant horn, middle, and distal body. The urachus opens into the allantoic cavity near the amnionic attachment of the allantoic section of the umbilical cord. In addition to the vascular structures noted above, yolk sac remnants can be noted. These may be ossified and attached by a vascular stalk; however, they are often found as a thin band of connective tissue parallel to the blood vessels.
Ultrastructure of the interface between placenta and endometrium The placenta, during the early stages of pregnancy, lies in direct apposition to the endometrium but is without substantive physical attachment until formation of microcotyledons between 75 and 100 days of gestation. Remaining epitheliochorial throughout the duration of gestation, progressive indentation of the chorionic epithelium by fetal capillaries and thinning of the maternal epithelium decreases the distance nu-
trients and gases must diffuse over and decreases the width of placental tissue separating fetal and maternal blood vessels. The smooth endoplasmic reticulum undergoes considerable development following the first trimester, with the rough endoplasmic reticulum increasing until day 200, indicating a considerable increase in protein synthetic activity. The trophoblastic cells also display a high level of pinocytotic activity, indicating that a large amount of nutrient transfer is occurring.37 The maternal microcaruncle consists of approximately 16 uterine crypts grouped in a connective tissue sheath. During the second half of pregnancy the depth of the microcaruncle increases. The fetal microcotyledon is formed by a clustering of fetal villi. The villi lengthen and branch considerably during the last 2–3 months of gestation. Branches are seen arising from close to the base of a villous stem, with branching also occurring along secondary and tertiary branches arising from the villus stem. Branching at the tips of the villi continues throughout gestation, even in the last few days before birth.9 Ultrastructural studies of the uterine glands at intervals during pregnancy show special trophoblast regions located between the bases of the microcotyledons (areolae), allowing uptake of endometrial gland secretions by pino- and phagocytosis.13, 38 The trophoblast cells become pseudo-stratified and are specially adapted to take up the exocrine secretions of the regional endometrial glands. This constitutes a second, histiotrophic form of nutrition for the developing fetus.
Physiology and function of the placenta Blood flow Placental blood vessels are oriented so that maternal blood flows countercurrent to fetal blood in the capillary networks, producing a very efficient exchange system.39, 40 Mimicking the structure of the microcotyledon (discussed above), the surface of the microcaruncle receives its arterial supply as branches of subendometrial vessels. They branch around the microcaruncle rim before draining through a capillary network to a solitary vein at the base. The complexity of this arrangement increases throughout gestation.9 Gas exchange Exchange of oxygen and carbon dioxide is by simple diffusion down a concentration gradient. In the equine, there is a relatively small oxygen gradient between the dam and fetus.40 Also, as pregnancy progresses placental oxygen demand increases in proportion with the increase in uteroplacental mass.41
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The Placenta Nutritive placental functions The equine placenta is a highly metabolic tissue, with considerable production of phospholipids to meet fetal needs, as well as modification of essential fatty acids.42 Compared to other species, metabolism of amino acids is of lesser importance with respect to energy balance. Nutrient transport and waste elimination The diffuse epitheliochorial placenta is a more efficient deliverer of nutrients than other types of placentas, as evidenced by the greater delivery of glucose per unit mass of placenta and the greater mass of fetus produced per unit mass of placenta, when compared to other species. As the fetus increases in size throughout gestation, the nutrient demands increase exponentially to term. To meet these requirements, the placenta, as the only channel for the transport of nutrients, continues to grow in mass and size. Total area of microscopic fetal–maternal contact also increases, with areolae and microcotyledons increasing considerably. Glucose transport through the placental cell cytoplasm is by diffusion of the free molecule and facilitated diffusion, with consumption per unit uteroplacental weight decreasing during pregnancy.7 Calcium ion transport is also likely by facilitated diffusion, in contrast to iron in uteroferrin molecules within membrane-bound vesicles.43 Amino acids are actively transported in quantities sufficient for protein deposition only, leaving no excess for oxidative metabolism; hence the equine fetus produces lower levels of urea than other species. Glandular secretions are transported by pino/phagocytosis.38 Endocrine functions Feto-placental production of estrogens occurs as the result of placental aromatization of androstenedione and dihydroepiandrosterone produced by the fetal gonads.44, 45 The total amount of estrogens produced varies throughout pregnancy in line with size changes in the gonads, and may also be dependent on the breed of the fetus, as evidenced by higher levels in pregnancies carrying Thoroughbred versus pony fetuses. This may be the result of the Thoroughbred fetus having larger gonads and producing more C19 precursor steroids that are aromatized by the placenta to estrogens.11 Progestagen production utilizes maternal cholesterol46 and begins around day 70 of gestation.47 For progesterone synthesis, pregnenolone from first the mare, then the fetal adrenal, is used.48 Progestagens released into the fetal circulation are further metabolized by fetal tissue, and return to the placenta before
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release into the maternal circulation.49 Maternal progestagen level may be considered a measure of fetoplacental well-being.50 Changes in production have effects on onset of parturition.49 Fetal stress in the final weeks of pregnancy increases mean plasma progestagen concentrations much earlier and to a significantly higher level compared to normal pregnancies, reflecting the premature maturation of the fetal adrenal gland. In turn this results in increased secretion of pregnenolone by the adrenal cortex for conversion to progestagens by the placenta.11 Prostaglandins (PG) are also produced by the utero-placental unit. They accumulate in the amnionic and allantoic fluids, and are released into both maternal and fetal circulations.51 Levels increase with advancing gestational age.52 Functions include patency of fetal vasculature, stimulation of fetal cortisol production, and control of the onset of labor.53
Immunological functions Epitheliochorial placentation, as in the mare, is the least intimate association between maternal and fetal tissues. To create the endometrial cups, fetally derived trophoblastic cells invade the uterine stroma, only to be destroyed by maternal leukocytes midway through gestation.54 Co-dominant maternally and paternally derived major histocompatibility (MHC) class I antigens are expressed at high levels on the invasive trophoblast cells of the chorionic girdle between days 32 and 36 of pregnancy,55 stimulating production of the anti-paternal alloantibodies that are observed during early gestation;56 however, they are not on the adjacent non-invasive trophoblast of the chorion and allantochorion membranes.55 Downregulation of MHC genes by trophoblast cells, making these cells largely non-antigenic, is considered to be a primary mechanism preventing maternal immune rejection of the fetal-placental unit. Downregulation of MHC class I antigen expression may be controlled at the transcriptional level.55 The invasive MHC class I-positive trophoblast cells demonstrate continuous levels of mRNA compatible with adult horse lymphoid tissue, compared to non-invasive MHC class I-negative trophoblast cells which display MHC class I at levels similar to adult horse non-lymphoid tissue.57
Factors affecting placental development and function Maternal factors The chorionic surface of the placenta can be considered a marker of the endometrial health of the
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mare, with mare age and parity also affecting placental development. Maternal influences on placental development are covered in detail in Chapters 226 and 230. Placental factors Fetal development is adversely affected by restrictions on placental formation and attachment, with foal weight statistically related to both effective allantochorionic surface area and allantochorionic weight.29, 58 The most common cause of diminished placental function (via a loss of effective contact between chorion and endometrium) is the occurrence of a twin pregnancy. The direct apposition of placentas in utero results in diminished endometrial surface contact, decreasing nutrient and waste exchange ability. Chronic nutrient deprivation of one or, usually, both fetuses results. Additionally, uterine space is not allocated evenly, one fetus usually being confined to a single uterine horn only, with the second fetus occupying the remaining horn and uterine body. Chronic in utero deprivation leading to death of the more confined fetus can cause widespread placental separation. Death and expulsion of both fetuses may result. Other causes of decreased maternal–fetal exchange include placental edema, chronic placentitis, and premature separation of placenta. Feto-placental infections are considered to cause up to one-third of all equine abortions; however, in approximately one-quarter of these cases no causative agent was found.59 Bacteria, equine herpesvirus, fungi, or placentitis of unknown etiological agent were the most common causes of abortion determined in one retrospective study. In another review, major pathogens identified included Streptococcus zooepidemicus, Leptospira spp., Escherichia coli, Pseudomonas aeruginosa, Streptococcus equisimilis, Enterobacter agglomerans, Klebsiella pneumoniae, alphahemolytic Streptococcus, nocardioform actinomycete, and fungi.60 Mixed bacterial growth was also found to occur. Umbilical cord pathology can restrict blood flow, decreasing venous return from the placenta to the fetus. Thrombosis of the fetal circulation within the placenta may occur, leading to venous stasis and congestion, with acidemia and anoxia damaging both the fetus and the placenta. Decreased perfusion of the chorionic villus decreases functional placental exchange area. Functional parenchyma of fetal vital organs may be decreased. A self-reinforcing cycle of feto-placental pathology can develop. In fact, the health of the placenta appears dependent upon a normal healthy fetus.61 Inflammation of the umbilical cord is known as funisitis, being associated with infectious causes of
abortion.62 This appears as a fibrinous deposit on the surface of the umbilical cord with inflammatory changes histologically. There appears to be a strong association with Leptospira spp. infections, with funisitis also being reported in other causes of placentitis, especially mare reproductive loss syndrome (MRLS). Fetal factors Developmental interaction between the placenta and fetus is shown to occur in both directions. In cases of malpresentation or musculoskeletal defects in viable foals, placental separation and involution of the uterus appear to proceed normally. However, in cases of abortion, stillbirth, or dysmaturity, damage to both fetal and maternal tissues is apparent, suggesting placental health is dependent upon association with a normal healthy fetus.61
Unique features of the equine placenta There is no physical attachment between the allantoamnion and the allantochorion in the equine. The fetus, encased by the allantoamnion and tethered by the umbilical cord, is highly mobile and regularly traverses between the pregnant and non-pregnant horns in early pregnancy. This continues until increasing size confines it to the pregnant horn. This is in contrast to other large domesticated species, such as ruminants, where attachment of the allantoamnion to the allantochorion throughout gestation limits fetal excursions. Although rare, it is possible for the fetus to develop in the opposite horn from the umbilical cord attachment (the designated pregnant horn). The umbilical cord is relatively long in the equine (95% are between 36 and 84 cm in normal Thoroughbred foals) with approximately four twists evenly distributed along its length.6 In conjunction with the mobility noted above, excessive torsion of the cord can occur, leading to urachal compromise, poor placental perfusion, and vascular compromise of the fetus and placenta. Thrombosis of fetal and placental vasculature can result. Fetal vital organ parenchymal mass can be decreased. Ischemic necrosis of placental villi vasculature that decreases functional fetomaternal exchange area can occur. Decreased umbilical cord length may result in premature rupture during delivery, or chronic in utero hypoxia due to excessive fetal traction. Excessive traction on the allantochorion may also lead to placental necrosis. Body wall defects may result from chronic traction to the umbilical area. Five avillous sites normally occur in the equine placenta: the chorion apposed to the internal os of
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The Placenta the cervix (known as the cervical star), the chorion adjacent to the utero-tubule junction, placental gathering at the site of umbilical cord attachment (due to normal choronic fetal umbilical cord traction), sites corresponding to the location of the endometrial cups, and along allantochorionic folds or large blood vessels. Pale or fibrotic areas other than these indicate sites of placental detachment or lack of microcotyledon formation.
Pathology Placentitis and twinning will be extensively reviewed in Chapters 9 and 242. However, non-infectious abnormalities of placental development leading to diminished functional competence result in placental insufficiency. Age-related degenerative endometrial changes decrease functional placental exchange by decreasing the area of normal placental development and apposition to the underlying endometrium. Endometrial degenerative changes greatly affect microplacentome development, diminishing both the extent and intimacy of physical and hematological contact at the feto-maternal interface.29 Stromal fibrosis, mononuclear cellular infiltration, glandular degeneration, and lymphatic stasis can occur. These changes have been called endometrosis.63
Implications of placental changes on fetal development and health Degenerative and developmental placental lesions have been shown to interfere with both placental function and efficiency, hence optimal fetal health and growth do not occur1.29, 58 Foal weight has been shown to be related to both allantochorionic weight and surface area. Gestational age can be related to placental surface area per unit foal weight. Significantly, the occurrence of a histological abnormality of the placenta has more effect on the foal than the gross surface area actually involved in the abnormality.58 Fetal stresses and insults resulting from placental incompetence are cumulative. Cellular damage and derangements of endocrine, metabolic, and cardiovascular processes over time produce a syndrome described as intrauterine growth retardation (IUGR).64 The outcome is dependent on the severity and duration of changes, as well as concurrent fetal developmental events. The fetus may experience an inappropriate gestational length, and may be delivered underweight with an inappropriate level of maturity. Fetal death with expulsion or failure of the fetus to survive parturition may also result. In less severe cases, fetal distress may be manifest as meconium
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passage in utero with subsequent aspiration, predisposing the fetus to respiratory distress syndrome and sepsis. Early in the neonatal period, intrauterine stresses can result in significant morbidity and mortality. In utero growth retardation impairs postnatal adaptive responses and can affect the adrenocortical, cardiovascular, and metabolic systems. Subtle growthretarding effects in utero may lead to deficits not identified until a period of metabolic stress is imposed, or may limit future health and athletic performance as is seen in other species, including humans.64
Interpretation of the placental evaluation A comprehensive review of the systematic evaluation of the equine placenta is found in Chapter 9. When interpreting this examination, mare signalment and reproductive history should be taken into consideration. Events relating to the current pregnancy (gestational age at parturition, duration of stages II and III of labor) are of major importance regarding the foal’s readiness for birth. Deviations in the appearance of the placenta from that considered normal provides information regarding both the mare and the foal. Of note, the presence of any histological abnormality, not the actual proportion of the placenta involved in the pathology, is indicative of potential compromise to the foal.58
Placental weight The placenta has been determined to be 11% of the foal’s birthweight in healthy Thoroughbred mares. A linear relationship exists between the birthweight of the foal and the allantochorionic weight.58 An increased weight of the placenta over that expected is therefore significant and may indicate edema, such as from Neotyphodium coenophialum (fescue toxicosis) or placentitis. Fetal nutrition and gas transfers are diminished due to the increased distance required to effect exchange. Neonatal compromise is possible. With fescue toxicosis, the foal may be presented dysmature with a long hair coat and be underweight following a possible prolonged gestation. The placenta itself may appear grossly normal but be markedly enlarged with fluid. Of further problem to the neonate, the mare may exhibit agalactia and have diminished colostrum quality. Placental weight may also be increased over that expected in cases of placentitis. The foal may be delivered underweight due to in utero malnutrition or infectious challenge
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transferred from the placenta. In these cases, neonatal hematology values will often indicate ongoing sepsis. Allantochorion
Color When appropriately formed, the entirety of the chorionic membrane appears as a rich red velvet-like surface, this being the macroscopic appearance of the villi. Prominent, deeply red engorged areas may be present, but are of unknown significance with no known associated pathology. However, five avillous sites normally occur in the equine placenta: the cervical star, the uterotubule junction, placental gathering corresponding to the umbilical cord attachment (due to normal chronic fetal umbilical cord traction), sites corresponding to the location of the endometrial cups, and along allantochorionic folds or large blood vessels. Any pale or fibrotic chorionic areas other than those noted above indicate sites of placental detachment or lack of microcotyledon formation. This indicates a loss of functional exchange area between fetus and dam. As the metabolic demands of both fetus and placenta are high, substantial avillous chorionic areas indicate compromised in utero nutrition. With normal endometrium being required for microcotyledon development, avillous chorion indicates significant degenerative endometrial change. This suggests compromised maternal reproductive health and potentially decreased future reproductive performance. The allantoic surface should appear clear, glistening and smooth and be free of excessive gross hemorrhage. The placenta and fetal fluids are likely grossly discolored in cases of placentitis or as a result of meconium passage due to in utero stress. When present, substantial hemorrhage indicates traumatic fetal delivery or vascular stasis involving the umbilical cord. This may precipitate hypoxic insult and anemia in the neonate. Dilated blood vessels and gross edema of the allantois indicates abnormal fluid balance within the feto-placental unit, most dramatically evident in dropsical conditions.
Thickness The non-pregnant horn is generally thinner in crosssection, has a more puckered wall, and is smaller in size when compared to the pregnant horn. While both horn tips are edematous, the pregnant horn is more prominently so in the normal placenta. Should the non-pregnant horn contain areas of overt thickness, focal placentitis may be the cause with potential to inoculate the fetus.
Cervical star thickening Intrauterine infections most often ascend through an incompetent cervix, although hematogenous and direct spread also occur. Causes of inappropriate cervical patency include pre-existing cervical trauma, vaginal urine pooling, and poor perineal conformation allowing aspiration of air and irritant debris into the cranial vagina. Inoculation of infectious agents, chiefly resident vaginal flora, through the cervix and eventually to the fetus is possible. When an ascending placentitis present, gross thickening, copious exudate, and a line of demarcation between the affected area and the adjacent placental body are characteristic findings. Detachment and deviation of the cervical star from the internal os often occurs, preventing the fetus exiting the allantochorion at the weakest point. This premature placental separation during parturition delays fetal expulsion, markedly increasing chances of meconium aspiration and hypoxic insult to the fetus. The neonate compromised secondary to ascending placentitis may have an insidious onset of difficulties and experience a rapid period of deterioration without warning. Placental evaluation will allow earlier detection of neonatal conditions. In such cases laboratory data at delivery is likely compatible with sepsis. Abnormalities may include abnormal white blood cell count and in chronic cases elevated fibrinogen levels, indicating fetal response to in utero derived infectious challenge. Pneumonic changes as evidenced by elevated respiratory rate and effort are likely. Neurological compromise can range from decreased activity levels through overt seizure activity. Prompt institution of appropriate antimicrobial therapy is required. Culture and sensitivity of the affected placenta may yield information useful to any subsequent treatment of the foal. As in utero deprivation from placental detachment is likely, supportive therapies and treatments for hypoxic insult should not be delayed.
Allantoamnion The allantoamnion should appear as an opaque to translucent white membrane without thickening or edema. Multiple thin branching blood vessels should be present, without signs of dilation. Small white plaques can be present; these are deposits of glycogen and not to be considered a pathological finding. The presence of discoloration, hemorrhage, edema or abnormal inclusions on the surface or within the allantoamnion may indicate an abnormal in utero environment. Allantoamnionic thickening may result from chronic fetal diarrhea, infection, inflammation, or dropsical conditions of the placenta. Perforation of
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The Placenta the allantoamnion by fetal movement, or rupture due to external trauma, have been reported. Fetal aspiration pneumonia, or entrapment of a limb with subsequent ischemia, are possible should damage to the allantoamnion be discovered.
Umbilical cord Increased length is the most common umbilical cord abnormality, predisposing the fetus to potentially lethal complications such as fetal strangulation, excessive torsion, and necrosis of the cord. Additionally, decreased umbilical cord length may result in premature rupture during delivery. Hemorrhage and tearing of intimal vessels, or excessive hemorrhage within the umbilical cord, is indicative of excessive torsion (up to four twists of the umbilical cord is normal)6 or traction in utero or during delivery. Nonreducible kinks in the cord indicate in utero torsion. Chronic excessive traction on the allantochorion may also lead to persistent in utero hypoxia and focal placental necrosis. Umbilical body wall defects may also result from chronic traction to that area. The umbilical cord of the mature fetus normally ruptures soon after parturition a short distance from the body wall; however, immaturity or pathology of this area can lead to excessive blood loss or result in abnormal separation allowing entry for infectious agents. Hemorrhage and tearing of intimal vessels, or excessive hemorrhage within the umbilical cord is indicative of excessive torsion or traction in utero or during delivery. With umbilical cord pathology, the neonatal foal can be considered at risk for neurological and metabolic problems due to chronic in utero deprivation. The neonate should also be evaluated for hypovolemia should excessive volumes of blood be present in association with the umbilical cord.
Placental fluids Abnormal quantities of fetal fluids indicate problems with either the dam or fetus. Hydrops allantois is an abnormal accumulation of fluid within the allantoic cavity. Normal allantoic fluid volume in the Thoroughbred mare ranges from 8 to 18 liters. Up to 200 liters has been reported in cases of hydrops allantois. Severe stresses are placed on the maternal cardiovascular system, body wall, and supporting ligamentous structures. Occurrence usually indicates a placental or histological endometrial abnormality that similarly affects nutrient and gas exchange. Abnormal accumulation within the amniotic cavity is called hydrops amnion, or hydramnion. Normal amniotic fluid volumes range from 3 to 8 liters.
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Increases indicate a fetal abnormality, with facial deformity often associated with this condition. Discoloration of fetal fluids may be the result of fetal distress causing meconium release, accumulation of inflammatory debris in utero, or occult hemorrhage from placental or umbilical lesions. In these cases, evaluation of the neonatal foal for pneumonic changes and consideration of pre-existing sepsis are prudent.
The mare The reproductive history of the mare should be considered when assessing placental health. Abnormal vulvar discharge, premature mammary development, or overt loss of colostrum prepartum should alert the clinician to the potential for placental compromise, fetal distress, and neonatal complications. In situations where placental dysfunction has led to premature lactation, the colostrum should be evaluated for immunoglobulin content at parturition. Prepartum maternal endocrinological measurements (progestins, total estrogens) are thought to give an insight into the metabolism of the feto-placental unit.49, 50 Prematurely elevated progestagen (progesterone metabolite) levels in late pregnancy (greater than 3 weeks before term, therefore preceding the normal rise) are considered to indicate placentitis or other placental compromise, although no substantive link has been demonstrated with maintenance of pregnancy. Decreased levels are considered to indicate fetal compromise. Similarly, total estrogen levels are also thought to reflect fetal viability, with a normal rise and fall during gestation coincident with the size of the fetal gonad.
The foal Placental compromise leading to an inappropriate level of maturity at birth (prematurity, dysmaturity) may be grossly obvious (underweight for gestational age, abnormal hair coat, orthopedic abnormalities); however, compromise to organ and metabolic function due to chronic in utero deprivations may not be overt and may lead to unknown effects later in life.65 Non-invasive assessment of feto-placental health can be achieved by ultrasonographic evaluation of the preterm fetus for size, activity level, heart rate, placental dimensions (both transabdominal and transrectal), and fetal fluid characteristics. Increased placental thickness suggests placentitis or vascular compromise. Placental detachment and folding may also be visualized. In the initial period following delivery, the foal that has experienced chronic placental dysfunction
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may appear to have normal behavior and physical examination findings. Similarly, aspiration of meconium may not immediately cause detectable pulmonary compromise. Onset of infection and inflammation following vital organ insult is usually progressive. Timely placental evaluation to detect abnormalities attunes the clinician to the requirement for closer monitoring of the neonate allowing earlier detection of those subtle variations from normal that indicate neonatal compromise.
Summary Diagnosis of placental abnormality has the potential to decrease perinatal morbidity and mortality by alerting the clinician to possible in utero and peripartum fetal stress. Assessment of the appropriateness of placental size and anatomy give insights into the fetal environment. Abnormal findings revealed by a closer postpartum placental evaluation can help indicate the potential for a less than optimal neonatal period for the foal. In addition, as the chorion is a reflection of endometrial health, considerable insight can be gained into the reproductive competency of the mare.
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References 1. Steven DH. Placenta depicta: Illustrations and Ideas. In: Steven DH (ed.) Comparative Placentation: Essays in Structure and Function. London: Academic Press, 1975; pp. 1–24. 2. Steven DH, Samuel CA. Anatomy of the placental barrier in the mare. J Reprod Fertil Suppl 1975;23:579–82. 3. Pijnenborg R, Vercruysse L. Shifting concepts of the fetal–maternal interface: a historical perspective. Placenta 2008; 29 Suppl A: S20–5. 4. Betteridge KJ. Comparative aspects of conceptus growth: a historical perspective. Reproduction 2001;122(1):11–19. 5. Whitwell KE. Morphology and pathology of the equine umbilical cord. J Reprod Fertil Suppl 1975;23:599–603. 6. Whitwell KE, Jeffcott LB. Morphological studies on the fetal membranes of the normal singleton foal at term. Res Vet Sci 1975;19(1):44–55. 7. Fowden AL, Forhead AJ, White KL, Taylor PM. Equine uteroplacental metabolism at mid- and late gestation. Exp Physiol 2000;85(5):539–45. 8. Fowden AL, Silver M. Glucose and oxygen metabolism in the fetal foal during late gestation. Am J Physiol 1995;269(6 Pt 2):R1455–61. 9. Macdonald AA, Chavatte P, Fowden AL. Scanning electron microscopy of the microcotyledonary placenta of the horse (Equus caballus) in the latter half of gestation. Placenta 2000;21(5–6):565–74. 10. Allen WR, Wilsher S, Stewart F, Ousey J, Fowden A. The influence of maternal size on placental, fetal and postnatal growth in the horse. II. Endocrinology of pregnancy. J Endocrinol 2002;172(2):237–46. 11. Allen WR, Wilsher S, Turnbull C, Stewart F, Ousey J, Rossdale PD, Fowden AL. Influence of maternal size on placen-
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
tal, fetal and postnatal growth in the horse. I. Development in utero. Reproduction 2002;123(3):445–53. Wilsher S, Allen WR. The effects of maternal age and parity on placental and fetal development in the mare. Equine Vet J 2003;35(5):476–83. Samuel CA, Allen WR, Steven DH. Studies on the equine placenta. III. Ultrastructure of the uterine glands and the overlying trophoblast. J Reprod Fertil 1977;51(2):433–7. Noden DM, de Lahunta A. Extraembryonic membranes and placentation. In: Noden DM, de Lahunta A (eds) The Embryology of the Domestic Animals. Baltimore, MD: Williams & Wilkins, 1985; pp. 47–69. Betteridge KJ. The structure and function of the equine capsule in relation to embryo manipulation and transfer. Equine Vet J Suppl 1989;8:92–100. Albihn A, Waelchli RO, Samper J, Oriol JG, Croy BA, Betteridge KJ. Production of capsular material by equine trophoblast transplanted into immunodeficient mice. Reproduction 2003;125(6):855–63. Allen WR, Stewart F. Equine placentation. Reprod Fertil Dev. 2001;13(7–8):623–34. Enders AC, Liu IK. A unique exocelom-like space during early pregnancy in the horse. Placenta 2000;21(5–6):575–83. Betteridge KJ. Equine embryology: an inventory of unanswered questions. Theriogenology 2007;68(1):9–21. Ginther OJ. Embryology and placentation. In: Ginther OJ (ed.) Reproductive Biology of the Mare: Basic and Applied Aspects. Cross Plains, WI: Equiservices, 1992; pp. 345–418. van Niekerk CH, Allen WR. Early embryonic development in the horse. J Reprod Fertil Suppl 1975;23:495–8. Samuel CA, Allen WR, Steven DH. Ultrastructural development of the equine placenta. J Reprod Fertil Suppl 1975;23:575–8. Gerstenberg C, Allen WR, Stewart F. Factors controlling epidermal growth factor (EGF) gene expression in the endometrium of the mare. Mol Reprod Dev 1999;53(3):255–65. Gerstenberg C, Allen WR, Stewart F. Cell proliferation patterns during development of the equine placenta. J Reprod Fertil 1999;117(1):143–52. Lennard SN, Stewart F, Allen WR. Insulin-like growth factor II gene expression in the fetus and placenta of the horse during the first half of gestation. J Reprod Fertil 1995;103(1):169–79. Lennard SN, Stewart F, Allen WR. Transforming growth factor beta 1 expression in the endometrium of the mare during placentation. Mol Reprod Dev 1995;42(2):131–40. Lennard SN, Gerstenberg C, Allen WR, Stewart F. Expression of epidermal growth factor and its receptor in equine placental tissues. J Reprod Fertil 1998;112(1):49–57. Oikawa M, Yoshihara T, Kaneko M, Yoshikawa T. Morphology of equine allantochorion at the tip of the pregnant horn. J Comp Pathol 1990;103(3):343–9. Abd-Elnaeim MM, Leiser R, Wilsher S, Allen WR. Structural and haemovascular aspects of placental growth throughout gestation in young and aged mares. Placenta 2006;27(11–12):1103–13. Samuel CA, Allen WR, Steven DH. Studies on the equine placenta. I. Development of the microcotyledons. J Reprod Fertil 1974;41(2):441–5. Roberts SJ. Gestation period – embryology, fetal membranes and placenta – teratology. In: Roberts SJ (ed.) Veterinary Obstetrics and Genital Diseases (Theriogenology). Woodstock: Roberts, 1986; pp. 38–92. Dickerson JW, Southgate DA, King JM. The origin and development of the hippomanes in the horse and zebra. II.
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The chemical composition of the foetal fluids and hippomanes. J Anat 1967;101(Pt 2):285–93. Allen WR, Moor RM. The origin of the equine endometrial cups. I. Production of PMSG by fetal trophoblast cells. J Reprod Fertil 1972;29(2):313–16. Allen WR, Hamilton DW, Moor RM. The origin of equine endometrial cups. II. Invasion of the endometrium by trophoblast. Anat.Rec. 1973;177(4):485–501. Enders AC, Liu IK. Trophoblast-uterine interactions during equine chorionic girdle cell maturation, migration, and transformation. Am J Anat 1991;192(4):366–81. King JM. The origin and development of the hippomanes in the horse and zebra. I. The location, morphology and histology of the hippomanes. J Anat 1967;101(Pt 2):277–84. Samuel CA, Allen WR, Steven DH. Studies on the equine placenta II. Ultrastructure of the placental barrier. J Reprod Fertil 1976;48(2):257–64. Wooding FB, Fowden AL. Nutrient transfer across the equine placenta: correlation of structure and function. Equine Vet J 2006;38(2):175–83. Silver M, Barnes RJ, Comline RS, Burton GJ. Placental blood flow: some fetal and maternal cardiovascular adjustments during gestation. J Reprod Fertil Suppl 1982;31:139–60. Silver M, Comline RS. Transfer of gases and metabolites in the equine placenta: a comparison with other species. J Reprod Fertil Suppl 1975; (23):589–94. Comline RS, Silver M. PO2, PCO2 and pH levels in the umbilical and uterine blood of the mare and ewe. J Physiol 1970;209(3):587–608. Stammers JP, Hull D, Silver M, Fowden AL. Fetal and maternal plasma lipids in chronically catheterized mares in late gestation: effects of different nutritional states. Reprod Fertil Dev 1995;7(5):1275–84. Wooding FB, Morgan G, Fowden AL, Allen WR. Separate sites and mechanisms for placental transport of calcium, iron and glucose in the equine placenta. Placenta 2000;21(7):635–45. Bhavnani BR, Short RV, Solomon S. Formation of estrogens by the pregnant mare. I. Metabolism of 7–3Hdehydroisoandrosterone and 4–14C-androstenedione injected into the umbilical vein. Endocrinology 1969;85(6): 1172–9. Tait AD, Santikarn S, Allen WR. Identification of 3 betahydroxy-5,7-pregnadien-20-one and 3 beta-hydroxy-5,7androstadien-17-one as endogenous steroids in the fetal horse gonad. J Endocrinol 1983;99(1):87–92. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J Reprod Fertil Suppl 1979;( 27): 499–509. Holtan DW, Squires EL, Lapin DR, Ginther OJ. Effect of ovariectomy on pregnancy in mares. J Reprod Fertil Suppl 1979;( 27): 457–63. Chavatte P, Holtan D, Ousey JC, Rossdale PD. Biosynthesis and possible biological roles of progestagens during equine pregnancy and in the newborn foal. Equine Vet J Suppl 1997; (24): 89–95. Ousey JC, Forhead AJ, Rossdale PD, Grainger L, Houghton E, Fowden AL. Ontogeny of uteroplacental progestagen production in pregnant mares during the second half of gestation. Biol Reprod 2003;69(2):540–8.
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50. Ousey JC. Hormone profiles and treatments in the late pregnant mare. Vet Clin North Am Equine Pract 2006;22(3):727–47. 51. Silver M, Barnes RJ, Comline RS, Fowden AL, Clover L, Mitchell MD. Prostaglandins in maternal and fetal plasma and in allantoic fluid during the second half of gestation in the mare. J Reprod Fertil Suppl 1979;( 27): 531–9. 52. Silver M, Fowden AL. Uterine prostaglandin F metabolite production in relation to glucose availability in late pregnancy and a possible influence of diet on time of delivery in the mare. J Reprod Fertil Suppl 1982;32:511–19. 53. Fowden AL, Ralph MM, Silver M. Nutritional regulation of uteroplacental prostaglandin production and metabolism in pregnant ewes and mares during late gestation. Exp Clin Endocrinol 1994;102(3):212–21. 54. Engelhardt H, King GJ. Uterine natural killer cells in species with epitheliochorial placentation. Nat Immun 1996;15(1):53–69. 55. Maher JK, Tresnan DB, Deacon S, Hannah L, Antczak DF. Analysis of MHC class I expression in equine trophoblast cells using in situ hybridization. Placenta 1996;17(5–6):351–9. 56. Donaldson WL, Oriol JG, Pelkaus CL, Antczak DF. Paternal and maternal major histocompatibility complex class I antigens are expressed co-dominantly by equine trophoblast. Placenta 1994;15(2):123–35. 57. Bacon SJ, Ellis SA, Antczak DF. Control of expression of major histocompatibility complex genes in horse trophoblast. Biol Reprod 2002;66(6):1612–20. 58. Cottrill CM, Jeffers-Lo J, Ousey JC, McGladdery AJ, Ricketts SW, Silver M, Rossdale PD. The placenta as a determinant of fetal well-being in normal and abnormal equine pregnancies. J Reprod Fertil Suppl 1991;44:591– 601. 59. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203(8):1170–5. 60. Hong CB, Donahue JM, Giles RC Jr, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Etiology and pathology of equine placentitis. J Vet Diagn Invest 1993;5(1):56–63. 61. Steven DH, Jeffcott LB, Mallon KA, Ricketts SW, Rossdale PD, Samuel CA. Ultrastructural studies of the equine uterus and placenta following parturition. J Reprod Fertil Suppl 1979; (27):579–86. 62. Sebastian M, Giles R, Roberts J, Poonacha K, Harrison L, Donahue J, Benirschke K. Funisitis associated with leptospiral abortion in an equine placenta. Vet Pathol 2005;42(5):659–62. 63. Allen WR. Proceedings of the John P. Hughes International Workshop on Equine Endometritis. Equine Vet J 1993;25(3):184–93. 64. Rossdale PD. The maladjusted foal: influences of intrauterine growth retardation and birth trauma. In: Proceedings of the 50th conference of the American Association of Equine Practitioners, 2004; pp. 75–126. 65. Rossdale PD, Ousey J. Fetal programming for athletic performance in the horse: potential effects of IUGR. Equine Vet Educ 2002;14(2):98–112.
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9.
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Examination of the Placenta Donald H. Schlafer
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Peri-parturient Management of the Mare and Neonate T. Douglas Byars and Barry W. Simon
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Chapter 9
Examination of the Placenta Donald H. Schlafer
Introduction What is a placenta? This may seem an odd question, but terms are best clarified early in any discussion. Variation in placentation between species is amazing and the medical terminology can be daunting. Tissues delivered with the foal are technically ‘the fetal membranes’, extraembryonic tissues derived from embryonic endoderm, mesoderm and trophectoderm. ‘The placenta’, in purest form, is reserved to describe the combined fetal and maternal tissues that form the complex tissue interface responsible for the transport of nutrient and wastes and performance of many other functions. In practical terms, clinicians, horsemen, farmers, most animal scientists and diagnostic pathologists, through common usage, refer to those tissues normally expelled with the fetus at birth as ‘the placenta’. This is the convention that will be used here. This chapter provides an overview of how one can perform a thorough placental examination using a standardized procedure. It is predicated on the assumption that the reader has some knowledge of basic embryology. Interpretation of findings requires inclusion of a brief review of equine placental anatomy and discussion of how to discriminate lesions of diagnostic significance from normal anatomical structures or tissue artifacts.
Placental examination Assessment of placental tissues for lesions of diagnostic significance and interpretation of the significance of changes contained within placental tissue relies on a working knowledge of many aspects of placental development, function and disease. It is also helpful to have an appreciation of basic tissue response to infection. Many chapters in this textbook contain important pertinent information about
placental embryogenesis, histology, gross anatomy, physiology, the role of the placenta as an endocrine gland (including development of role of endometrial cups), control of placental function and basic tenets of placental pathology. There are also many helpful references in the literature that address placental development and anatomy to which the reader is referred.1–9 Other useful references discuss a broad spectrum of infectious diseases that can infect the pregnant mare.10–20 Although many articles on equine placental pathology were published some time ago, they are still of great value.21–28 Under what circumstances should placentas undergo detailed gross examination? When should busy clinicians undertake placental examination on the farm? Detailed placental examinations should be done when there are abortions, stillbirths, delivery of small-for-gestational-age fetuses or those from weak or otherwise ill-thrifty newborns. Placental tissues often contain lesions that help explain why the newborns are ill, poorly grown, or delivered early and can help guide treatment plans or preventive measures if more than one mare has pregnancy failure. There are clear advantages to sending these tissues to a fully equipped laboratory for diagnostic examination, sampling and testing, especially if they come from a mare that aborted or delivered a dead, ill, or developmentally abnormal foal. Diagnostic facilities usually have higher success rates at arriving at a specific diagnosis if the examination and collection of samples are done within the laboratory. These advantages must be weighed against the additional costs, time delays, and the effort associated with preparing and shipping the tissues. Other reasons for opting to send tissue to a laboratory include inherent difficulties associated with a field necropsy on someone’s farm. These include
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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commitment of time, preparation of equipment and the facilities where the examination will be done, and, most importantly, there is the possibility of spreading infectious agents that may have caused the abortion around the premises to other horses, to people working there, or to oneself. It can be difficult to charge appropriately for your professional time, yet there are also clear advantages to doing this work yourself. Perhaps foremost is that one becomes directly involved and feeback is immediate. Committing to do the placental (and in cases of abortion, fetal) necropsy raises interest in the case and each placental examination enhances one’s clinical acuity and knowledge that is also helpful in interpretation of ultrasound images during examination of the uteruses of gravid mares. Doing the examination may yield a quick diagnosis, as there are many conditions that do have gross lesions that are specific for different causes of fetal loss or compromise to fetal growth. Samples are much fresher than when the placenta has been shipped or taken to a diagnostic laboratory. Time is of the essence for isolating causative microbial agents and for preservation of tissues in fixatives for histopathology. When the decision is made to submit placental tissues to a diagnostic laboratory, it is prudent to take some swabs and place them in a commercial transport medium to be sent with fresh tissues. This will improve the probability that causative microbes will be identified. One’s ability to identify gross lesions of diagnostic significance relies on knowledge and experience gained by carefully examining as many normal placentas as possible. By taking advantage of the owner’s and/or farm manager’s concerns and interests, a professionally conducted necropsy examination with the owner or farm manager attending provides the opportunity to explain what you are doing, what the tissues are, how they function, and so on. This can be an excellent client educational opportunity and can be a practice builder. On larger farms, the help of knowledgeable farm managers or owners to identify abnormal placentas can be very helpful as they can be trained to collect, weigh and do a cursory examination of placentas from every foaling and alert the attending veterinarian of potential problems.
Figure 9.1 The three major tissues of the equine fetal placenta as shown in this photograph are: the umbilical cord, the amnion and the large chorioallantois. The shape of the chorioallantois is a mirror image of the inside of the gravid mare’s uterus in which it developed and grew to meet the functional demands of the rapidly growing fetus. The ‘arms’ of the ‘Y’-shaped placenta correspond to the two uterine horns and equally large uterine body. Note that the amnion is only attached at the middle of the cord and not at all to the chorioallantois. Reproduced from Schlafer29 with permission of the American Association of Equine Practitioners.
niotic blood vessels which can appear even more impressive when congested (Figure 9.3). Their presence frequently causes concern, but these are normal anatomic features. Most equine placentas are inverted during delivery (like a glove turning inside out as the hand is pulled away). Therefore, the allantoic surface of delivered placentas is usually outermost (Figure 9.4). The allantoic surface of the chorioallantois is smooth, shiny and whitish. Branches of the umbilical blood vessels course over its surface. The red, ‘velvety’ microcotyledonary surface of the chorioallantois will be on the inner surface (Figures 9.5 and 9.6).
Performing an equine placental examination Figure 9.1 is a photograph of a normal equine fetus, its umbilical cord and attached amnion and chorioallantois. In Figure 9.2 the amnion has not been ruptured and the fetus and amniotic fluid are contained in a normal amnion. Note the very prominent am-
Figure 9.2 Mid-gestational equine fetus contained within its highly vascularized amnion. The prominence of these amniotic vessels is a common concern, but they are always normal unless there is clouding of the amniotic membrane (amnionitis).
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Cross section of uterine wall and attached placenta Choriollantois fetal allantoic cavity
Endometrium Myometrium
Allantonic surface
serosa
Microplacentomes Microcotyledons
Figure 9.3 Amniotic membrane with prominent congested blood vessels and uniformly distributed amniotic plaques. Amniotic plaques are routinely found on the fetal side of the amniotic membrane. They are uniformly distributed and most dense closer to the attachment of the umbilical cord. The amniotic membranes also extend over the amniotic segment of the umbilical cord so amniotic plaques are commonly found on the amniotic surface of the cord. There is progressive reduction in size up the cord towards its attachment to the fetus.
Placental examination is best done on a clean, large, flat surface in an area with plenty of light and that can be thoroughly cleaned and disinfected. Protective clothing, instruments and containers for collection of samples should be assembled before the examination is done. Many diagnostic laboratories provide ‘abortion kits’ that contain the necessary forms, fixatives, syringes, instruments and containers. These are a great help and most will also have a detailed listing of samples that should be taken. Following a brief gentle rinse with a stream of cold water from a hose using moderate pressure, bedding material can be removed. Rinsing should
Figure 9.4 Thorough gross examination of the fetal membranes requires a clear, well-lit, flat surface in an area that can be thoroughly cleaned after the examination and collection of samples is done. Following a quick gentle rinse with a stream of cold water from a hose to remove bedding debris, the membranes are laid out so that both uterine horns extend perpendicular to the body region, forming an ‘F’-shaped pattern. All folds are gently smoothed out so that small lesions are not missed.
Chorionic surface
Figure 9.5 The upper panel is a tissue section from the uterine wall of a pregnant mare in which care was taken not to disrupt the attachment of the fetal chorioallantoic membrane. When the chorioallantois is pulled free, delicate villi protruding from the chorionic surface (microcotyledonary villi) that had been interlocked in furrows and crypts in the endometrium are freed. Their presence all over the chorionic surface of the chorioallantois gives it a deep red color (lower panel) due to the presence of engorged capillaries within the villi. The middle panel is a photograph of the smooth allantoic surface of the chorioallantoic membranes. Reproduced from Schlafer29 with permission of the American Association of Equine Practitioners.
Figure 9.6 A photomicrograph of a single microplacentome formed by the interdigitation of microcotyledonary villi of the fetal chorioallantois with crypts called microcaruncles in the remodeled endometrial surface. The larger round structures at the bottom edge of the photomicrograph are distended endometrial glands that are producing uterine secretions that are released at the endometrial surface and taken up by specialized trophoblast cells covering the chorioallantois over domedshaped small chambers at the openings of the endometrial glands (areolae).
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be limited to a short time as the tissues will tend to imbibe water, which may potentially complicate histopathology. This will appear as allantoic edema which is an important lesion in many placental infections. It is helpful to first find the umbilical cord and to then spread the chorioallantoic membranes. This takes some time, but ensures that the entire allantoic surfaces are examined and no lesions missed. Examination of the chorionic surfaces is equally important and requires that the chorioallantois be everted so that the chorionic surface is outermost (as it would be in the uterus). Examination of each end of the three arms of the chorioallantoic membrane will reveal a tear at the tip of the ‘body’ segment that will have pale avillous radiating bands of the cervical star (Figure 9.7). Normal deliveries will have the chorioallantois ruptured in the area of the cervical star (Figure 9.4). Any areas of thickening, discoloration or abnormal texture should be noted and recorded. Swabs for culture or cytology can be taken, but removal of a piece for fixation is delayed until both sides of the chorioallantois have been thoroughly examined. For every fetal placenta, all four ‘surfaces’ of the chorioallantois must be examined. Infections of the equine placenta most commonly develop in the chorioallantois that covers the inner aspect of the cervical canal (Figure 9.8). These ascending infections produce prominent thickening, necrosis and fibrosis in the area of the cervical star, thus making this area the most important for sample collection and careful examination.
Figure 9.7 Cervical star. The chorionic surface is covered with tens of thousands of small tufts of highly vascularized villi of the microcotyledons. This gives a gross appearance of a uniform red, finely velvety surface. In areas where the microcotyledonary villous tufts are lost, the surface of the chorion appears whitish, like that shown in the picture of this cervical star. Other areas that will also be devoid of microcotyledonary villi are where the papilla of uterine tubes enter the uterus, over areas where endometrial cups are, at the base of one uterine horn, where the chorioallantois is folded on itself, where there is severe endometrial damage or where the chorioallantois is against the chorioallantoic membrane of a co-twin.
Figure 9.8 Chronic necrotizing placentitis surrounding the cervical star area. There is a radiating pattern of increased placental thickening covered by tan necrotic debris. This pattern is very characteristic of a chronic ascending infection either by fungal or bacterial organisms.
The uterine horn and uterine body are of about similar lengths, and are shaped in a ‘Y’. Because there is a pronounced ventral bulge in the uterine body, one cannot lay the fetal membranes flat. In order to do so both uterine horns extend perpendicular to the body region, forming an ‘F’-shaped pattern (Figure 9.4). The allantois surface is smooth, shiny and light colored with various sized branches of umbilical arteries and veins on its surface. Examination continues by turning the membranes over to complete the examination of the allantoic surfaces on the ‘down side’. To examine the chorionic surface, the chorioallantois must be turned inside out and spread out again so the chorionic surface can be examined. The entire chorioallantois can be turned inside out by reaching through the torn cervical star tissue, gently grasping a fold of the wall of the body region and pulling it out to start the inversion process. Once eversion of the membranes has begun and the dark red color of the chorionic surface of the chorioallantois is present, the horns can be inverted by using gravity to gently ‘shake’ them out. The inverted membranes will need to once again be flipped over to examine the chorionic surface that was facing the table. All folds are gently smoothed out so that small lesions are not missed. Any areas of thickening, discoloration or abnormal texture should be noted, measured, described and sampled. Digital photographs are very helpful and offer a great way to present areas you feel are important. These can be sent with the samples as prints or as digital files to the diagnostic laboratory. Careful examination and sampling of chorioallantoic tissues in the region around the cervical star is especially important. Ascending infections commonly occur through the cervix, therefore placentitis is most
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Examination of the Placenta commonly found involving the chorioallantois that abuts the inner os of the cervix. Microvilli that cover the surface of the chorioallantois give it a velvet-like appearance (Figures 9.5–9.7). The villi can be seen with the naked eye if the tissues are placed under water and gently moved. The presence of artifacts can cause confusion. Like all other tissues, placentas autolyze with time and lesions become more difficult to appreciate. Placentas should be kept cool and moist until they can be examined to minimize autolytic changes. Storage will alter the gross appearance of placental tissues even for short periods as the microcotyledonary villi are so highly vascularized. The chorionic surface will normally have a uniform deep red color. This is because the capillaries in the microcotyledonary villi in very fresh placentas are uniformly filled with blood that has been retained after delivery. When the placentas are placed in a bag or box, or placed in a pile, pressure from their own weight forces blood from one area of the chorioallantois to another, producing patterns of dark and pallor that can greatly complicate assessment and interpretation of findings. Placental infections result in acute or chronic changes. These are usually focal or locally extensive and disrupt the normally uniform red chorionic surface (Figure 9.8) Acute and subacute placental inflammation cause placental edema, necrosis of chorionic villi and accumulation of inflammatory debris. Mycotic infections cause chronic fibrosis and necrotizing placentitis that leaves the chorioallantois firm, tan and thick (Figure 9.7).
Structures and features of the normal equine placenta that can be confused with lesions of diagnostic significance There are many normal anatomic structures that can be mistaken for lesions. The most important of these include: thickened ‘gravid’ uterine horn (Figures 9.9 and 9.10), plush villi on the chorionic surface of the ‘non-gravid’ horn, focal area of chorioallantoic edema and degeneration where the hind feet of the foal press against the placenta and uterine wall [fetal foot PAD (placental area of degeneration) lesion] (Figures 9.10 and 9.11), chorioallantoic vesicles (Figure 9.12), chorioallantoic pouches (Figures 9.13 and 9.14), hippomanes (Figures 9.15 and 9.16), mild congestion (Figure 9.17), avillous areas on the chorionic surface of the chorioallantois [cervical star (Figure 9.17), over endometrial cups (Figure 9.13) over linear areas overlying large branches of the umbilical vessels, areas of twin placental apposition
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Figure 9.9 The apical ends of both horns of a normal equine placenta. Note the dramatic difference between the gross appearance of the upper chorioallantois (the ‘gravid horn’ chorioallantois) that was attached to the uterine horn in which the embryo attached and to which the umbilical cord would be attached, and the lower, thinner, ‘non gravid’ horn. (These terms are confusing, but have become embedded in clinical nomenclature.) The edematous ‘gravid horn’ chorioallantois is much thicker.
(Figure 9.20) and over the papillae of the uterine tubes] and amniotic plaques (Figure 9.3). Thickened ‘gravid’ uterine horn and plush villi on the chorionic surface of the ‘non-gravid’ horn There is a dramatic difference between the thickness and gross appearance of the thicker and edematous chorioallantois of the ‘gravid horn’ and the thinner ‘non-gravid’ horn. Distention of the membranes tends to make the villi on the surface of the gravid horn less dense and shorter, in marked contrast to the tall, dense plush appearance of the chorionic surface of the non-gravid horn. Fetal foot, placental area of degeneration, (PAD) lesion The hind feet of the foal press against the placenta and uterine wall during the last trimester of
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Figure 9.10 Cut sections of chorioallantoic membrane shown intact in Fig. 9.9 comparing a fetal foot PAD lesion (left) within the gravid chorioallantoic membrane to a section normal nongravid membrane (right) taken from an area the same distance from its apex. The foot PAD lesion in the chorioallantois on the left is ten times thicker (approximately 2.0 cm vs 0.2 cm). It is markedly edematous and the microcotyledonary villi on its surface are much shorter, their relative density less, and histopathology revealed degeneration, mineralization and squamous metaplasia of trophoblast cells.
Figure 9.12 Chorioallantoic vesicles: focal areas of allantoic edema typically located around blood vessels. These areas vary greatly in size.
filled vesicles. These are most prominent around blood vessels as shown in Figure 9.12. gestation, causing chronic locally extensive damage to chorioallantoic membrane. The degenerative tissue is grossly visible a short distance from the tip of the gravid horn (Figures 9.10 and 9.11). Chorioallantoic vesicles Focal accumulation of fluid within the allantoic stroma of the chorioallantoin sometimes forms fluid-
Chorioallantoic pouches Endometrial cups grow in the endometrium from specialized trophoblast cells that migrated from the chorionic girdle of the outer trophectoderm of the embryo and live for about 2 months. They are then destroyed by a targeted maternal immune reaction. Their necrotic remains are sloughed from the surface of the endometrium into an infolding pocket of the overlying chorioallantoic membrane (Figures 9.13 and 9.14). The resulting chorioallantoic pouches are found in an area near the attachment of the umbilical cord to the chorioallantois. They are usually, but not always, present and vary in size and number. Those shown in Figure 9.13 are especially large. The necrotic contents within one of these chorioallantoic pouches is seen in Figure 9.14. Hippomanes
Figure 9.11 ‘Fetal foot PAD’. A common lesion in normal placentas consisting of a locally extensive area of chorioallantoic degeneration in the ‘gravid horn’ chorioallantois. It is caused by chronic repetitive trauma from pressure at the point of contact of hooves of the hind feet of the fetus. The chorioallantoic in these ‘fetal foot PADs’ is thickened and edematous, and the microcotyledons are degenerate or even necrotic and mineralized. The PAD lesions tend to be rectangular and are found a short distance from the tip of the gravid chorioallantic horn; but not at the tip as one would expect. The fetal foot PAD lesion in this photograph is more prominent than usual because of the degree of focal degeneration and the marked congestion of the entire gravid horn.
Hippomanes, also known as allantoic calculi, are found only within the allantoic cavity and are usually present as single, soft cream to dark tan, free-floating masses (Figures 9.15 and 9.16). They form as a precipitate of lipids and cellular debris with laminar bands forming as mineral is deposited. They grow in size as more material is deposited on their surface. Congestion of chorioallantoic capillaries Congestion of vascular beds can be associated with inflammation (placentitis), mechanical compression limiting vascular return of blood (cord torsion or
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Figure 9.13 Chorioallantic pouches. These structures contain necrotic endometrial cup material (see Figure 9.14). They are always found near the base of the ‘gravid’ chorioallantic horn overlying the endometrium that contained the endometrial cups. Cups are destroyed by a maternal immune reaction leading to coagulative necrosis followed by extrusion into an infolding of the overlying chorioallantoic membrane, forming a pouch. The pale white band seen in the photograph is the area devoid of microcotyledonary villi on the chorion surface where the chorioallantois covered the endometrial cups growing within the endometrium, preventing development of the microcotyledonary villi. Endometrial cups are more frequently present than not, but are not found in all placentas.
cord incarceration or compression), or when blood is compressed from one area of the membranes and pooled in another. The latter can occur from uneven pressure on the membranes at delivery or as an artifact when the different area of the delivered placenta are compressed as it lies in a mass. Subgross mi-
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Figure 9.15 Hippomane (‘allantoic calculus’). These soft, flat, almond-shaped bodies are composed of cellular debis, lipids and mineral deposits. Hippomanes frequently folded upon themselves (see cut section in Fig. 9.16). There is usually only one hippomane. They are found only in the allantoic cavity. Occasionally they are attached to the allantoic membrane or similar material is occasionally found attached to fibrin strands on the allantoic surface. Reproduced from Schlafer29 with permission of the American Association of Equine Practitioners.
croscopic tissue sections comparing normal and congested chorioallantoic membranes are shown in Figure 9.17. Normal avillous areas on the chorionic surface of the chorioallantois These are found at cervical star (Figure 9.7), over endometrial cups (Figure 9.13), over linear areas overlying large branches of the umbilical vessels, areas of twin placental apposition, and over the papillae of the uterine tubes. Amniotic plaques Amniotic plaques are focal areas of squamous metaplasia uniformly scattered on the fetal side of the amniotic membrane. They tend to be most dense closer to the attachment of the umbilical cord. Because the amniotic membranes also extend over the umbilical
Figure 9.14 Histological tissue section of a chorioallantoic pouch containing necrotic endometrial cup material. Microcotyledons line the chorionic surface of the chorioallantois (along the upper surface).
Figure 9.16 The cut surface of a hippomane. Note the white laminated bands of mineralization and the folded pattern. Hippomanes are typically almond shaped, occur singularly, and can be up to 15 cm long.
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Identification of placental lesions One of the simplest ways to tell if a placenta may be abnormal is to examine its weight. If the placental weight is either lighter or heavier than normal it should be referred to a veterinarian for further examination. A rule of thumb is that at term the entire chorioallantois, cord and amnion should normally weight approximately 11% of the foal’s body weight.7 Weight and measurement parameters for normal placentas Data on normal weights and measurements of basic placental physical parameters are available in the literature. Most data sets are from term Thoroughbred placentas,7, 8, 30 but limited data on Standardbred placentas has been reported.7 Morphometric techniques have been used to quantitatively study the microscopic attachment of the chorioallantois to the endometrium.6, 9
Length of the umbilical cord
Figure 9.17 Stained tissue sections of pieces of chorioallantois taken from placentas of three different mares. All were photographed at the same magnification. The tissue on top was taken from a normal placenta. The microcotyledonary tufts of villi form a uniform covering on the lower surface and the smooth allantoic surface is on the upper side. Compare this to the tissue in the middle from a placenta of a fetus that aborted with a torsed umbilical cord. Blood vessels are very congested and the chorioallantoic membrane appears thickened. The chorioallantoic membrane at the bottom is massively necrotic and thickened. There is extensive inflammation, necrosis and reactive allantoic cystic proliferation (cystic adenomatous allantoic dysplasia). This lesion was caused by chronic fungal infection. Reproduced from Schlafer29 with permission of the American Association of Equine Practitioners.
cord, amniotic plaques are commonly found on the surface of the umbilical cord with progressive reduction in size of those further up the cord towards its attachment to the fetus. In some placentas, they are very small, and in others they are a prominent feature. If the surface of the plaques becomes keratinized, they will appear as small horn-like rugose growths. If the fetus was stressed and meconium was released into the amniotic fluids, the amniotic plaques will be stained yellow.
The length of umbilical cords of Standardbred mares (most, but not all, were term) were 58.58 ± 17.57 cm in a study carried out in Canada.31 Eighty Thoroughbred cords measured after foaling were 66.9 + 22.0 cm.30 Another larger study of 143 Thoroughbred mares found a surprisingly broad range of lengths from 32 to 90 cm with a mean of 55 cm (SEM 0.93).7 Cord length has been found to correlate to weight of the chorioallantoic membranes.8
Weight of the chorioallantois and amnion The most detailed study is that of Whitwell and Jeffcott (1975)7 who reported total placental weight of 5.7 ± 0.08 (SEM) kg in 139 Thoroughbred mare placentas in Newmarket. The chorioallantois alone (n = 144 measurements) was 3.6 ± 0.05 kg (SEM) and the weight of the amnion recorded from 140 of these same mares was 1.85 ± 0.04 kg (SEM) holding from 3 to 5 liters of amniotic fluid. This same report described the surface area of the chorioallantois as 16,700 ± 150 (SEM) square cm and the surface area of the amnion as 22 600 ± 824 (SEM) square cm. The authors suggested that the amnion had a larger surface area than the chorioallanois because it followed the contours of the surface of the fetus. They also reviewed measurement data from nine other studies from the time period 1924–1970. Data from a more recent study from mares foaling in Kentucky found the average weight of the chorioallantois to be 4.40 kg (9.7 lb),30 slightly more than the 3.6 kg reported by Whitwell and Jeffcott in 1975.7
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Examination of the Placenta Placentitis Areas of placental inflammation and necrosis usually lose their red color. Affected areas are tan to brown, thickened, and may have inflammatory exudates on their surface (Figure 9.8). Placentas with areas of placentitis are usually heavy. Bacterial and mycotic placentitis in the mare usually develop because infection ascends to the placenta (cervical star area) through the cervix. The most prominent tissue changes in ascending bacterial placentitis occur on the chorionic surface (Figure 9.17). Bacterial infections tend to spread from the cervix along the plane between the endometrium and chorioallantois, causing placental separation as the infection spreads away from the cervix. It is critical to include samples of the chorioallantoic membranes taken from the cervical star area in all tissue sample sets submitted for culture and histopathology (Figure 9.18). Maternal inflammatory cells from the endometrium drawn through chemotactic mechanisms accumulate on the chorionic surface and become admixed with necrotizing villi (Figure 9.17). This material can be identified during gross placental examination. There are only a few infectious dis-
Figure 9.18 The arrows in this line drawing indicate areas from which tissues should be collected for fixation in formalin and submission for histopathology (see also Figures 9.1 and 9.2). It is especially important to include sections from the cervical star area, even if no lesions are grossly evident. Tissue or swabs from the chorioallantois at or near the cervical star should also be collected for submission for culture. Impression smears are also frequently helpful. Reproduced from Schlafer29 with permission of the American Association of Equine Practitioners.
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eases that produce diffuse placentitis (leptospirosis, candidiasis) and some produce placentitis in specific locally extensive areas (such as the chronic suppurative placentitis at the base of the uterine horns and body typically seen in nocardioform infections). Placental separation Another reason for changes in placental color is placental separation. If at delivery the chorioallantois at the cervical start does not tear, uterine contractions will force the intact chorioallantoic membranes through the open cervix, tearing it away from its attachment to the endometrium. The pathogenesis is unclear, but may relate to failure of maturation of the chorioallantoic tissues to weaken or there may be lesions from prior inflammation at this site from infections that ascended through the cervical canal. The condition is called ‘red bag’ because the separated chorioallantoic membrane becomes very congested. The congested vessels in the microcotyledonary villi become so congested that they give the surface of the chorioallantois a bright red color in contrast to the areas of the membrane that stay attached until delivery (Figure 9.19).
Figure 9.19 ‘Red bag placenta’. This is the chorioallantois from a placenta that had partially separated prior to birth of the foal. The cervical star is at the very bottom edge. The very red area in the bottom half of the chorioallantois shown is very congested and would correspond in appearance to the sectioned tissues in the middle of Figure 9.8. The condition is called ‘red bag’. Reproduced from Schlafer29 with permission of the American Association of Equine Practitioners.
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Figure 9.21 This torsed umbilical cord decreased blood flow to the placenta resulting in acute placental insufficiency and abortion. Note the attachment of the cord to the ventral abdomen of the fetus at the lower right. Abortion due to torsion of the umbilical cord is common. Excessively long umbilical cords are more prone to become torsed. There are normally only about four 380◦ twists to the umbilical cord. Note also the segmental distention of the amniotic segment of the cord near its attachment to the ventral wall of the fetus. The distention was caused by constricted flow of urine from the bladder through the urachus. Commonly torsion will cause hemorrhage in the wall of the cord and congestion of the chorioallantois.
Figure 9.20 The matching pale areas on the chorionic surface of the chorioallantois of each of these aborted equine twins are devoid of microcotyledonary villi where they touched. Note that the fetus on the right is smaller due to growth retardation that was likely a result of restricted functional placental contact area with its dam’s endometrium.
In addition to the color changes described above (the tan to cream color associated with necrotizing placentitis, the bright red from marked congestion in ‘red bag’, and the regionally mottled red areas that develop with storage after delivery), the absence of villi causes the normally red velvety chorionic surface to appear smooth and gray to silver, as seen in the photograph of the large cervical star in Figure 9.7. The large ‘matching’ avillous lesions on the chorionic membranes of the twin fetuses in Figure 9.20 are typical of those found on all placentas of twins.
Lesions of the umbilical cord The most common lesion of the equine umbilical cord is excessive length. Abnormally long umbilical cords are commonly associated with non-infectious abortions. Placental examination should include recording umbilical cord length. Torsion of the umbilical cord is a common cause of abortion in mares and the diagnosis can be made based on the presence of more than four or five tight
twists in the cord with associated hemorrhage in the cord. Frequently there is associated distention of the urachus (Figure 9.21). Another lesion found in the umbilical cord is ossification of remnants of the yolk sac (Figure 9.22).32 The involuted remnant of the yolk sac can be found in the central cavity of the allantoic segment of the umbilical cord. For unknown reasons, small areas of osseous metaplasia develop within the middle segment of the wall of the thin cord-like yolk sac tissues. These can expand, usually developing a fluidfilled central cavity surrounded by a wall composed of plates of lamellar bone, forming a round mass with some characteristics similar to the bony plates of the calvarium. As they enlarge, they become displaced from the inner cord and enveloped by the allantoic membranes. They extend from the cord tethered by a round band of tissue containing the proximal and distal segments of the involuted yolk sac. On rare occasions, these pedunculated structures can compromise blood flow through the umbilical vessels and cause abortion.
Sample collection and documentation of findings A standard set of tissues should be taken from each placenta.29 Areas from which tissues should be collected for fixation in formalin and submission for
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ciation of Equine Practitioners in 2004.29 Several images originally published in those proceedings are included in this chapter with permission of the AAEP.
References
Figure 9.22 An ossified remnant of a yolk sac. These bony structures are found within or attached to the allantoic segment of the umbilical cord. They have a fluid-filled center and thin wall containing multiple lamellar bony plates. They somewhat resemble the cranium of a fetus, and have been mistakenly considered to be blighted embryos, like the amorphous globosus found in bovine placentas. Larger ossified yolk sacs are usually attached by a cord of connective tissue, adding to the impression the mass has its own umbilical cord. The ‘cord’, however, is proximal and distal linear aspects of the yolk sac that continue within the umbilical cord with normal attachment points.
histopathology are indicated in Figure 9.18. It is especially important to include sections from the cervical star area, even if lesions are not grossly evident. Tissue or swabs from the chorioallantois at or near the cervical star should also be collected for submission for culture. Impression smears are also frequently helpful as cytology may reveal inflammatory cells or the causative microorganisms. A simple gross pathology report form is readily available that provides guidance and an easy way to record placental necropsy findings.29 One of the most common problems associated with submissions to diagnostic laboratories is lack of adequate histories. Any suspected lesion should be photographed. Digital cameras capable of recording high-quality images are relatively inexpensive and digital files can be recorded on discs or sent over the Internet. Photographs sent with tissue samples are of great help to diagnosticians. They also serve as an important part of the pathology record. Practitioners are encouraged to establish a close working relationship with personnel in each of the various laboratories to which they send samples for diagnostic testing. The rewards for such effort can be substantial. Note This chapter is based on a manuscript by this author published in the Proceeding of the American Asso-
1. Carluccio A, Panzani S, Tosi U, Riccaboni P, Contri A, Veronesi MC. Morphological features of the placenta at term in the Martina Franca donkey. Theriogenology 2008;69:918–24. 2. Flood PF. Fertilization, early development, and the establishment of the placenta. In: McKinnon AO, Voss, JL (eds) Equine Reproduction, Lea & Febiger: [place of publication], 1993; pp. 473–85. 3. Ginther OJ. Embryology and placentation. In: Reproductive Biology of the Mare, 2nd edn. Equiservices: Cross Plains, WI, 1992; pp. 345–418. 4. Mcdonald AA, Chavatte P, Fowden AL. Scanning electron microscopy of the microcotyledonary placenta of the horse (Equus caballus) in the latter half of gestation. Placenta 2000;21:565–74. 5. Schlafer DH. The umbilical cord – lifeline to the outside world: structure, function, and pathology of the equine umbilical cord. In: Powell DG, Furry D, Hale G (eds) Proceedings of the workshop on the equine placenta. Lexington, KY: University of Kentucky, 2004; pp. 92–8. 6. Villani M, Wilsher S, Carluccio A, Contri A, Veronesi MC. Stereology in the term placenta of the donkey, Thoroughbred and pony: a comparative study. Anim Reprod Sci 2006;94:411–12. 7. Whitwell KE, Jeffcott LB. Morphological studies on the fetal membranes of the normal singleton foal at term. Res Vet Sci 1975;19:44–55. 8. Whitwell KE. Morphology and pathology of the equine umbilical cord. J Reprod Fertil Suppl 1975;23:599–603. 9. Wilsher S, Allen WR. The effects of maternal age and parity on placental and fetal development in the mare. Equine Vet J 2003;35:476–83. 10. Donahue JM, Williams NM, Sells SF, Labeda DP. Crossiella equi sp. Nov., isolated from equine placentas. Int J System Evolut Microbiol 2002;52:2169–73. 11. Ellis WA. Leptospirosis as a cause of reproductive failure. Vet Clin North Am Food Anim Pract 1994;10:463–78. 12. Gerst S, Borchers K, Gower SM, Smith KC. Detection of EHV-1 and EHV-4 in placental sections of naturally occurring EHV-1- and EHV-4-related abortions in the UK: use of the placenta in diagnosis. Eq Vet J 2003;35:430–3. 13. Kenney RM, Doig PA. Equine endometrial biopsy. In: Morrow DA (ed.) Current Therapy in Theriogenology. W.B. Saunders Company, Philadelphia, PA: Saunders, 1986; pp. 723–29. 14. Ostlund EN. The equine herpesviruses. Vet Clin North Am Equine Pract 1994;9:283–94. 15. Palmer JE. Potomac horse fever. Vet Clin North Am Equine Pract 1993;9:399–410. 16. Paradis MR. Update on neonatal septicemia. Vet Clin North Am Equine Pract 1994;10:109–35. 17. Poonacha KB, Donahue JM, Giles RC, Hong CB, PetritesMurphy MB, Smith BJ, Swerczek TW, Tramontin RR, Tuttle PA. Leptospirosis in equine fetuses, stillborn foals, and placentas. Vet Pathol 1993;30:362–69. 18. Schlafer DH, Miller R. Pathology of the female reproductive system. In: Maxie MG (ed.) Jubb, Kennedy and Palmer’s
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Pathology of Domestic Animals, 5th edn. [place of publication]: Elsevier Saunders, 2007; pp. 429–564. Smith KC, Whitwell KE, Blunden AS, Bestbier ME, Scase TJ, Geraghty RJ, Nugent J, Davis-Poynter NJ, Cardwell JM. Equine herpesvirus-1 abortion: atypical cases with lesions largely or wholly restricted to the placenta. Eq Vet J 2004;36:79–82. Szeredi L, Aupperle H, Steiger K. Detection of equine herpesvirus-1 in the fetal membranes of aborted equine fetuses by immunohistochemical and in-situ hybridization techniques. J Comp Path 2003;129:147–53. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203:1170–5. Hong CB, Donahue JM, Giles RC, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Etiology and pathology of equine placentitis. J Vet Diagn Invest 1993;5:156–63. Hong CB, Donahue JM, Giles RC, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Equine abortion and stillbirth in central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. Jeffcott LB, Whitwell KE. Twinning as a cause of foetal and neonatal loss in the thoroughbred mare. J Comp Path 1973;83:91–106. Sebastian MM, Bernard WV, Riddle TW. Review paper: mare reproductive loss syndrome. Vet Pathol 2008;45:710–22.
26. Smith KC, Blunden AS, Whitwell KE, Dunn KA, Wales AD. A survey of equine abortion, stillbirth and neonatal death in the UK from 1988 to 1997. Equine Vet J 2003;35:496– 501. 27. Tengelsen LA, Yamini B, Mullaney TP, Bell TG, Render JA, Patterson JS, Steficek BA, Fitzgerald SD, Kennedy FA, Slanker MR, Ramos-Vara JA. A 12-year retrospective study of equine abortion in Michigan. J Vet Diagn Invest 1997;9:303–6. 28. Whitwell KE. Investigations into fetal and neonatal losses in the horse. Vet Clin North Am 1980;12:313–31. 29. Schlafer DH. Postmortem examination of the equine placenta, fetus, and neonate: methods and interpretaton of finding. In: Proceedings of the American Association of Equine Practitioners, 2004;50:1–19. 30. Oulton RA, Fallon LH, Meyer H, Zent WW. Observations of the foaling data from two central Kentucky thoroughbred operations during the spring of 2003;. In: Powell DG, Furry D, Hale G (eds) Proceedings of the workshop on the equine placenta. Lexington, KY: University of Kentucky, 2003 pp. 68–71. 31. Whitehead AE, Foster R, Chenier T. Placental characteristics of Standardbred mares. In: Powell DG, Furry D, Hale G (eds) Proceedings of the workshop on the equine placenta. Lexington, KY: University of Kentucky, 2003; pp. 71–6. 32. Cassar TIY, Fallon LH, Martinez EH, Schlafer DH. Segmental ossification of involuted yolk sacs in equine umbilical cords. Ninth International Symposium on Equine Reproduction. Kerkrade, The Netherlands, August 7–11, 2006. Anim Reprod Sci 2006;94:439–42.
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Chapter 10
Peri-parturient Management of the Mare and Neonate T. Douglas Byars and Barry W. Simon
Routine management of the periparturient mare and neonatal foal reflects an appreciation of normal mare and foal behavior and abnormalities that signal disease, abnormal development, or traumatic insults relating to birthing. There is no substitute for the experience gained by the observation of many normal pregnancies, deliveries, and foals. The recognition of problems begins with an appreciation for a normal pregnancy devoid of premature lactation, vaginal discharge, and normal mare behavior without clinical compromise. The average gestation period in the mare is 11 months (330–345) days as normal range since the last stallion cover or impregnation by artificial insemination. An additional period of 10 days is often added to the 11 months and considered a more representative timeframe with variation due to previous gestations, maiden mare status, or exposure to environmental factors that affect the gestational period. These factors may include fescue toxicosis, mare reproductive loss syndrome (MRLS), or infectious diseases such as leptospirosis, rhinopneumonitis (equine herpesvirus type I), and uteroplacental infections either by ascending cervical insults or systemic diseases that affect the uterus and a vulnerable pregnancy. High-risk mares include those with dubious previous pregnancies that have produced dead, maladjusted or septic foals, and dysmature foals with abnormal birth weights and stature. Mares that have produced previous jaundiced foals due to neonatal isoerythrolysis (NI) may be screened by agglutination or hemolytic antigenic tests for blood incompatibilities. After foaling compatibility can be tested by having colostrum and newborn foal blood examined for agglutination prior to nursing (‘button test’ or jaundiced foal agglutination; JFA). Foals at risk for NI should be muzzled and given supplemental compatible colostrum
followed by milk replacer for approximately 48 hours post-foaling while the dam is stripped or milked of her colostrum over that time period. Core vaccinations are routinely given to the mare approximately a month prior to foaling to boost passive colostral antibodies and provide passive immunity for the neonate. Vaccinations that are recommended for most broodmares include tetanus, Eastern and Western encephalomyelitis, West Nile virus, rabies, equine herpesvirus 1 and 4, and influenza. Additional vaccinations that are administered based on risk of disease include botulism, rotavirus, strangles, and rarely an autogenous Salmonella. Parasite control in the mare is usually provided by a routine deworming program and additional treatments are not indicated prior to delivery. Most mares are dewormed shortly after foaling to reduce the risk of transmammary passage of Strongyloides westeri and to reduce fecal egg passage in the manure during the immediate postnatal period. However, with the most recent research concerning the developing resistance of equine parasites to the available battery of equine anthemintics, new deworming schedules are being utilized based on quantitative fecal egg counts. These schedules include all individuals on a farm basis and provide a more diagnostic basis for worming programs. (See Chapter 30) The broodmare diet is influenced by the available feed sources and time of year. As is customary, clean water and hay or grazing free choice is a minimum. Supplementary feeding of grain is normal and is at the discretion of owners and managers. The excessive use of supplements in addition to a balanced grain ration may compromise the overall ration and predispose to toxicities by certain vitamins, minerals, and herbal concoctions. Selenium- or iodine-deficient geographic areas can be supplemented as directed
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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by a nutritionist or agricultural extention service agent. Exercise or free movement in fields and paddocks should be provided as much as possible. Excessive time spent confined in the barn may allow for more disease exposure. Confined mares such as those being laminitic or enduring other orthopedic problems are at higher risk to produce stillbirths or foals of small stature. Overweight mares should be assessed for metabolic syndrome as a potential for infertility. The evaluation of the mare includes the assessment of any premature mammary development and vaginal discharge. Mucopurulent vaginal discharge would be the most common notable clinical sign although urine or hemorrhage can also be observed. The former is usually associated with bacterial placentitis and the latter is usually from vaginal varicosities in middle-aged to older mares. Evaluation of mares with a vaginal discharge would include vaginal evaluation that may extend to a speculum examination of the cervix and transrectal sonographic examination of the uteroplacental thickness and integrity adjacent to the cervical star as well as the volume and character of fetal fluids. Microbiological techniques, both bacterial and fungal, should be performed by culture and cytology of any uterine discharge observed by speculum. In addition to transrectal ultrasonographic examination, a transabdominal fetal ultrasonogram may be performed to assess the fetal heart rate, placental thickness and separation, foal position and development, and to look for abnormal fluid accumulation. Hydrops mares can be identified with rectal determination of a non-palpable fetus due to excessive allantoic and/or amniotic fluid accumulation or ultrasonographic assessment of fluid volume. Mares with ruptured prepubic tendons will require abdominal truss support (see Chapter 259). Impending foaling is noted by mammary enlargement, waxing of the teats, edema, and relaxation of the vulva and relaxation of the pelvic ligaments. Mares with a Caslick’s vulva procedure performed earlier in the pregnancy should be opened within a few days or hours prior to parturition. Parturition is a relatively rapid act in the mare, divided into stages. Stage 1 is the mare’s behavioral activities indicating the onset of labor. Stage 2 is the rupture of the chorioallantoic membranes and delivery of the foal. Stage 3 is the post-foaling expulsion of the placenta and fetal membranes. Stage 1 labor is characterized by excessive walking and frequent defecation. Some mares may begin to sweat and most look at their sides in either the standing or recumbent position. Active labor will be noted by the mare beginning to push or ‘squint’ her abdomen with increasing
pressure associated with rupture of the chorioallantois (‘breaking of water’). Foal delivery should occur in approximately 20 minutes with the forefeet and head presented following rupture of the amniotic sac. The chorioallantois should not arrive first, but if so it should be manually ruptured to prevent suffocation of the foal. Assistance during foaling is prompted by abnormal presentation and position of the foal and/or the recognition of a prolonged time of labor. The placenta should be passed within 2– 3 hours of delivery. (See Chapter 9) Mares may be foaled in a variety of environments; however, if episodes of sequential and severe diarrheas are consistently noted in newborns, foaling outside the barn in pens or lighted paddocks is recommended. A common foaling stall or area is often a significant factor in spreading infectious diseases. Stall washings and disinfection are often inadequate in problem barns. Cleaning the mare’s perineum and udder just before delivery may also help reduce the neonate’s exposure to pathogenic bacteria prior to ingestion of colostrum. Close observation of the foal by foaling personnel should begin immediately after delivery, beginning with the assessment of the oral mucous membranes to be assured of pink, oxygenated mucosa. The foal should be able to sit sternally in 1– 2 minutes, have a suckle reflex in 2–20 minutes, stand in 1–2 hours and nurse within 2–4 hours.1, 2 The foal’s heart rate should be 80–120 beats per minute and respirations should be 30–40 breaths per minute. The average rectal temperature is 99–101.5◦ F (37.3–38.5◦ C). Septic foals are often hypothermic. External palpation of the thoracic cavity is performed to determine if fractured ribs may be present. The navel is usually treated with an appropriate topical astringent or disinfectant. Neurological evaluation in newborns is considered difficult since reflexes such as the menace and spinal reflexes are slow or absent. Obvious orthopedic issues should be evaluated although the recumbent foal is significantly less amenable to musculoskeletal evaluation than the standing and ambulatory foal. Meconium should pass within 1–2 hours and enemas are optional unless straining or abdominal tympany are present. Urination is variable and usually noted within 24 hours of foaling regardless of the foal’s gender. Neonatal isoerythrolysis-positive foals (positive JFA to colostrum or serum agglutination against the foal’s RBCs) should be muzzled. The foal should receive compatible colostrum by bottle or nasogastric tube. Colostrum supplementation should be followed by milk replacer at a volume of approximately 20% of the foal’s weight per day administered at 2- to 4-hour intervals.
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Peri-parturient Management of the Mare and Neonate The routine use of antibiotics in the newborn should be considered discretionary. Farms with infectious diseases or individual foals with abnormal clinical signs may receive prophylactic antibiotics.2 A complete physical examination, including hematology, should be routinely performed approximately 24 hours after birth with an assessment of the IgG concentration. Most insurance companies will accept an IgG concentration > 400 mg/dL, although some companies may require at least 800 mg/dL. If less than these values passive immunoglobulin transfer will require intravenous plasma infusion to the foal followed by a retest of the IgG level obtained. The increase in IgG is frequently relatively poor (about 200 mg/dL per liter of plasma administered) and this may reflect intravascular dilution by increasing the plasma oncotic gradients within the foal’s circulation from the 50% extracellular fluid in the neonate. Abnormalities in blood counts or serum chemistry should be considered in relation to clinical signs in the foal. The complete physical examination should include examination of the eyes, oral examination for cleft palate and checking the navel and external urogenitals.3 Assuming medical or orthopedic compromise is not present, mares and their foals may be turned out even in most inclement weather conditions within hours of parturition unless fractured ribs were noted either by physical examination or ultrasound evaluation. For foals with fractured ribs4 approximately 2-3 weeks of stall confinement is recommended prior to initial turn-out. The mare should receive brief periods of exercise during the time of the foal’s confinement. Exercise for the mare will improve uterine involution or outflow of uterine fluid discharge and may lessen excessive stall activity in the mare that could potentiate trauma to the foal. The neonatal foal will become a perinate within a few days, then being referred to as a suckling in a couple of weeks. Farm management practices have historically included an initial anthelmintic treatment as the first worming during this timeframe with re-
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peat wormings at approximately 2-month intervals. However, special attention is given to roundworms which take 8–12 weeks to mature. Attention should be paid to anthelmintic resistance, with an avoidance of ivermectin and routine doses of the benzimidazoles. In conformance with the current recommendations, deworming treatment based on fecal egg counts should be considered from the start. Larvicidal dosing with fenbendazole (i.e., 10 mg/kg p.o. q 24 h for 5 days) or moxidectin have been recommended. Moxidectin should not be used in foals under 6 months of age as per label directions. Vaccination with a series of core vaccines commences at 3–4 months of age although colostral antibodies may reduce the efficacy of some vaccines such as influenza. Weaning of foals usually is performed at 4–6 months of age although some breeding programs will wean as early as 2 months of age. Orphaned foals should be placed onto a foster mare (nurse mare) or, if raised entirely by humans, they will often require behavior adjustment prior to group introductions. Routine foal management is the result of experience and the cooperative efforts of the owner, manager and the veterinarian. The direct or indirect reliance upon veterinary services is pivotal to the success of the pregnancy and production of a live viable foal.
References 1. Barton, M. Neonatal sepsis. DVM Magazine, Advanstar Com., N. Olmstead, Ohio, 2E-5E, May 2008. 2. Traub-Dargatz JL. Postnatal care of the foal. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadephia: Lea & Febiger, 1993; p. 982. 3. Magdesian G, Wilkins P. Neonatology, Section 11. In: Orsini JA, Divers TJ (eds) Equine Emergencies, 3rd edn. Philadephia: Saunders, 2008; pp. 486–521. 4. Sprayberry KA, Bain FT, Seahorn T et al. 56 cases of rib fractures in neonatal foals hospitalized in a referral center intensive care unit from 1997–2001. Proceedings of the 47th Annual Meeting of the American Association of Equine Practitioners, 2001; pp. 395–9.
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Foal Rejection Wendy E. Vaala
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Prematurity, Dysmaturity and Assessment of Maturity Guy D. Lester
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Resuscitation (Foal and Birth) Niamh M. Collins, Jane E. Axon and Jonathan E. Palmer
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Shock Niamh M. Collins, Jane E. Axon and Jonathan E. Palmer
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Perinatal Asphyxia Syndrome Pamela A. Wilkins
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Sepsis L. Chris Sanchez
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Chapter 11
Foal Rejection Wendy E. Vaala
Foal rejection is not a common occurrence, but it can have extensive consequences for the foal, ranging from delayed or inadequate colostrum ingestion to severe trauma or even death. Foals that must be removed from their unaccepting dams face the added stress associated with hand-rearing or bonding onto a foster mare. Foals failing to receive adequate colostral antibodies are at increased risk for bacterial infection during the early neonatal period. Owners of rejected foals may be faced with the labor-intensive task of hand-rearing and socializing the orphan or the added expense of finding a suitable foster dam
Maternal behavior The physiological control of normal maternal behavior in the mare has not been investigated in great detail. One theory is that the abrupt decline in both estrogen and progesterone, accompanied by a transient increase in oxytocin, sensitizes the central nervous system to sight, sound, and particularly the smell and taste of a newborn foal.1 Mare-to-foal bonding is believed to begin during the delivery process when the mare begins sniffing the chorioallantoic fluid and placenta. This helps explain why some mares will lick and nicker to human attendants who have been soaked in fetal fluids during the delivery process as though mistaking them for their foal based on olfactory cues. Mares often display a flehmen response while smelling the fetal fluids or fetal membranes. Soon after delivery the mare begins licking the foal and fetal membranes. Once this olfactory recognition has occurred, the mare is able to discriminate between her foal and other foals and will reject attempts of foreign foals to nurse.2 Within minutes of delivery the mare will become distressed if her foal is removed from sight. This need for dams to smell and taste their foals and fetal fluids soon after delivery may explain why mare–foal bonding can be disrupted by periparturient events such as excessive human interference
soon after delivery, a change in a sick foal’s smell following administration of certain medications such as DMSO, or following an abnormal delivery such as a dystocia or cesarean section when the mare may be deprived of the natural sequence of smelling and licking her foal and placenta. Some experts believe that the critical period for maternal bond formation is within the first 6 hours following delivery.2 Mares may lose the mothering instinct within several days of delivery if they do not remain in direct physical contact with their foals. Other early signs of good mothering in the dam include attending to the foal within minutes of birth by licking, nuzzling, and nickering to the newborn, avoiding stepping on the foal when moving around the stall or lying down, protecting the foal from perceived threats, and allowing or even assisting the foal to nurse.3 Some mares actively facilitate nursing by repositioning themselves and even raising a hind leg to make it easier for the foal to locate the udder. The foal also plays a role in stimulating and enhancing the bonding process. Healthy foals begin vocalizing in response to their dam’s nickers and will nuzzle and nudge their dam even while they are still recumbent. Once standing, a normal foal will begin searching for the udder and display udder-bumping behavior. After the foal has found and suckled from the teats it should be able to relocate the udder with fewer and fewer misdirected advances towards other body parts on the dam. The foal should also begin to identify with its dam and prefer to remain in close proximity to her. Foals that may not be able to fulfill their role in the bonding process include neonates that fail to develop normal suckle reflexes, udderseeking behavior or the ability to stand and ambulate due to disease conditions that include sepsis, dysmaturity, and hypoxic ischemic encephalopathy. Mares may also have a difficult time receiving the normal bonding cues from debilitated newborns that require
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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constant nursing care and high-volume stall traffic, supportive therapy that necessitates the placement of ‘foreign’ intravenous catheters, feeding tubes, oxygen cannulas, and the administration of medications that alter the smell of the foal’s urine and/or feces. Twinning may lead to different outcomes. The mare may accept both twins or may select one twin and reject the other. In one instance, a multiparous Standardbred mare selected the smaller and weaker filly and rejected the larger and more active colt. During the first few days of life the mare and foal spend 95–100% of their time within 5 m of each other. 2 Mares also exhibit a protective response to their sleeping foals, described as the recumbency response. 2 Whenever a foal lies down the normal mare will disrupt her random grazing pattern or eating behavior to stand over her foal or graze near her foal. This protective response, when displayed while on pasture and or in the stall, is an encouraging sign in a foalrejecting mare and offers promise that she is beginning to bond with her newborn. Mares also demonstrate a protective response by positioning themselves between their foal and any external threat.
Types of foal rejection Foal rejection can be grouped into several broad categories: (i) avoidance of the foal, (ii) refusal of the mare to allow the foal to nurse, and (iii) overt, unprovoked maternal aggression towards the foal.8 One behavorist describes additional categories of aberrant maternal behavior, including: (i) ambivalence, (ii) extreme protectiveness, (iii) fear of the foal, (iv) avoidance of the foal, and (v) savage attack of the foal.3 Regardless of the classification system used, there seem to be two general groups of foal-rejecting dams. One group includes mares that may simply show ambivalence toward their foal owing to inexperience, disrupted bonding, or an abnormal foal, and others that are afraid or anxious around the foal, particularly during nursing attempts. Some mares may prevent suckling because they are feeling discomfort associated with a full and tender udder, strong postpartum uterine contractions, or other underlying disease condition. Periparturient events that may disrupt early maternal bonding include excessive human intervention surrounding delivery, premature removal of the placenta, abnormal deliveries such as dystocia or cesarean section that delay mare and foal contact. The second broad category includes mares that demonstrate overt hostility toward the foal that may or may not be triggered by attempts to nurse and mares that are misdirecting aggression towards other horses to their foals. Savage attacks on the foal often involve biting and even shaking or throwing the foal.3 A mare demonstrating this behavior should be separated im-
mediately from her foal and the foal should be placed on a surrogate dam. Within each category there may be many different contributing factors. It is important to try to characterize the type of behavior observed and attempt to determine the underlying cause, since it will affect not only what therapy is pursued, but whether treatment for the mare is even attempted. Maternal behavior that should not be mistaken for foal rejection includes occasional squealing when a foal first begins nursing or nipping at her foal when it becomes too vigorous in its suckling attempts. Mares experiencing varying degrees of agalactia may also nip or kick at their foals to terminate prolonged or unproductive nursing bouts owing to udder discomfort. This is not foal rejection. Is there a ‘psychological profile’ of a mare that is more likely to reject her foal? Based on a limited number of retrospective surveys on selected populations of broodmares, some risk factors have been suggested by noted equine behavorists.
r r r r
Lack of experience of being a mother plays a role since maiden mares are overrepresented in the group of foal-rejecting mares. Older mares reject foals more commonly than younger ones.1 Dams raised in isolation from other foals during their youth are more likely to reject their foals than mares that have been exposed to other foals during their development.1 Arabian mares are more likely to reject their foals than mares of other breeds and, within this breed, Arabian mares of Egyptian lineage may be even more likely to show poor mothering instincts.1, 4,8
Another review of the topic included the following risk factors for foal rejection: (i) primiparous mares of any breed, (ii) Arabian mares, especially those whose paternal grandmother rejected her foal, (iii) multiparous mares that have rejected two or more foals.5 In the same review, it was noted that there was no association with method of breeding: 82% of rejecting mares and 89% of accepting mares were bred by natural cover. Mares that rejected their foals were more likely not to have licked their foals during the early postnatal period. A review of 23 cases for foal rejection noted that premature removal of the placenta or excessive human contact during the first 24 hours after delivery may increase the likelihood of foal rejection.5
Treatment and management of foal rejection The first goal of therapy is to ensure the safety of the foal while trying to determine the cause and potential reversibility of the abnormal maternal behavior.
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Foal Rejection If unprovoked maternal aggression is the problem then prompt removal of the foal is paramount and attempts to reunite the mare with her foal should be discontinued to avoid trauma not only to the foal but to those working with the pair. If the mare’s behavior is potentially amenable to correction, but will likely require hours or days of intervention, then the foal should be given colostrum from its own dam or another mare to ensure adequate absorption of colostral antibodies within the first 6–8 hours following delivery. While working to reunite neonate with dam, the foal’s minimal nutritional requirements should continue to be met with mare’s milk or a suitable mare milk replacer. It is often helpful to keep the foal slightly hungry so that it will continue to do its part in searching for the udder and attempting to nurse. Providing 10% of the foal’s bodyweight in milk per day is a reasonable level of nutrition to provide necessary calories and help maintain hydration. Bottle feeding is the preferred route of milk administration to encourage suckling and udder-bumping behavior. Simple maternal ambivalence towards the foal may respond to quiet restraint of the mare while quietly encouraging the foal to nurse. Reintroducing the placenta to certain mares may stimulate the olfactory cues to initiate the bonding process, especially those that underwent an unusual or prolonged delivery that required maternal sedation. Some mares may be preoccupied with or distracted by their own discomfort. A careful physical examination may reveal excessive discomfort due to stage III uterine contractions, colic due to intestinal disease, or udder tenderness associated with edema. Therapy with analgesic and anti-inflammatory drugs such as flunixin meglumine (Banamine, Intervet Schering Plough, Millsboro, DE; 0.5–1.0 mg/kg intravenously (i.v.) or per os (p.o.) q.12–24 hours) may alleviate pain and allow the mare to refocus on her foal. Mares that seem to accept the foal but will not let it nurse may have a tender udder or may simply lack experience and self-confidence when approached by the foal. If the udder is tender to palpation, cool compresses can be applied and flunixin meglumine can be administered. A low dose of oxytocin (5 IU i.v. or intramuscularly (i.m.)) helps facilitate milk let down and gentle hand-milking will relieve some of the mammary gland’s distension. If the mare is afraid of the foal then two handlers are needed. One individual restrains and reassures the mare and the other handler helps guide the foal toward the udder while protecting it from aggressive actions by the mare. Positive reinforcement when the mare tolerates the foal is preferred to punishing her whenever she tries to misbehave; otherwise, the anxious or nervous mare may quickly begin to associate the
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foal with excessive punishment. Allow the mare to sniff the foal’s hindquarters as it attempts to nurse. Most normal foals, when allowed to nurse independently, naturally align themselves with their heads towards the mare’s tail and their tails towards the mare’s head. This posturing encourages the mare to sniff and nuzzle the foal’s hindquarters and often stimulates the dam to lick around the foal’s tail and perineum. These maternal actions may play a role in helping the dam identify her foal and the licking often stimulates more vigorous nursing attempts. Handlers should avoid the tendency to force the mare to nuzzle the foal’s face or head since this action seems to be more threatening for some mares. If the mare tends to bite at the foal, a grazing muzzle can be applied to protect the newborn and decrease the need to verbally or physically reprimand the mare. Hobbles, if tolerated, can be used for mares that tend to kick at the foal. The dam can be restrained manually or tied in the stall behind a protective bar placed at a height level with her stifles that allows the foal to access the udder while preventing the mare from turning and biting or kicking. Twitching should be avoided. If simple restraint of the dam, with or without oxytocin administration, is not enough, then tranquilization with acepromazine (0.03–0.066 mg/kg i.v./i.m.) may be required to calm the mare without causing ataxia. This dopamine antagonist also has the potential to stimulate lactation through enhanced prolactin release. Xylazine and other α-adrenergic agonists should be used conservatively, since this drug class may be more likely to render the mare ataxic and yet still able to kick or strike at the foal. A combination of detomidine and butorphenol can be used to sedate particularly difficult mares. The effect that long-term use of these sedatives and analgesics may have on the mare’s ability to bond with her foal has not been reported. Some rejecting mares, especially those with a history of foal rejection, may benefit from hormonal therapy using altrenogest (ReguMate, Intervet Schering Plough, Millsboro, DE; 0.044 mg/kg orally q.12–24 hours) to raise the threshold for aggression.2,8 If the mare shows promise in accepting her foal, a few behavioral tests can be used to validate her maternal interest. The foal can be taken out of the stall and out of the dam’s sight. If this act induces separation anxiety and vocalization in the mare it is an encouraging sign. Likewise, if orchestrated threats to the foal elicit protective behavior in the mare, then there is hope that she will eventually develop a normal maternal bond with her foal. A mare that stands over her sleeping foal is often on the road to accepting her maternal duties. Some mares and foals benefit from spending time out of the stall and in a small
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paddock or pasture that provides a more natural setting.
Prevention of foal rejection Several prepartum management strategies are recommended to reduce the risks of foal rejection. Allow the expectant dam several weeks to become accustomed to the foaling stall and surroundings by moving her into the foaling environment at least a few weeks prior to her anticipated delivery date. Ensure that the foaling stall is quiet and away from high traffic flow areas in the barn and prevents direct visual contact with other horses. Avoid unnecessary human intervention during the first 12–24 hours following parturition and limit the number of different individuals entering the foaling stall. Primiparous mares that are unaccustomed to having their flanks or udders touched should be desensitized prior to foaling. Two people are required to perform this procedure – one to hold and reward the mare and the other to gently accustom her to palpation of the flank and udder. The mare can be offered food rewards as positive reinforcement whenever she accepts udder and teat contact without squealing or kicking. If the mare fails to respond favorably within a reasonable amount of time then the desensitization procedure should be discontinued to avoid aggravating the mare’s intolerance. Mares with a history of rejecting previous foals may benefit from initiation of prophylactic hormone therapy immediately following delivery, rather than waiting for the first signs of rejection to appear. The author has had good results using 3–5 days of a
combination drug protocol that includes the administration of altrenogest (0.044 mg/kg orally q.12–24 hours), estradiol benzoate (10 mg i.m. q.24 hours) and domperidone (Equidone, Equi-Tox, Central, SC; 1.1 mg/kg orally q.12–24 hours). This hormone protocol is similar to the one developed to induce lactation in mares as well as to create surrogate dams from non-lactating mares.6, 7
References 1. Houpt KA. Foal rejection. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia: WB Saunders Co., 1987; pp. 126–8. 2. Barber JA. Perinatal behavior of the mare and foal. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 4th edn. Philadelphia: WB Saunders Co., 1997; pp. 562–5. 3. McDonnell SM. Mare behavior problems. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. Philadelphia: WB Saunders Co., 2003; pp. 264–5. 4. Juarbe-Diaz SV, Houpt KA, Kusunose R. Prevalence and characteristics of foal rejection in Arabian mares. Equine Vet J 1998;5:424–8. 5. Houpt KA, Olm D. Equine behavior: foal rejection: a review of 23 cases. Equine Pract 1984;6:38–40. 6. Daels PF, Bowers-Lepore J. How to induce lactation in a mare and make her adopt an orphan foal: what 5 years of experience have taught us. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007, pp. 349–53. 7. Porter RH, Duchamp F, Nowak R, Daels PF. Induction of maternal behavior in non-parturient adoptive mares. Physiol Behav 2002;77:151–4. 8. Houpt KA. Foal rejection. In: Robinson NE, Sprayberry K (eds) Current Therapy in Equine Medicine, 6th edn. Philadelphia: WB Saunders Co., 2009:116–118.
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Chapter 12
Prematurity, Dysmaturity and Assessment of Maturity Guy D. Lester
The normal gestational period The mean gestational length, calculated from the day of insemination to the day of parturition, has been consistently reported between 340 and 342 days in the Thoroughbred breed.1–5 However, in contrast to most other domesticated species, the range of the ‘normal’ gestational period in horses is very wide. For example, data from England indicated that 95% of Thoroughbred mares foaled between 327 and 357 days of pregnancy.1 The mean duration of pregnancy is relatively consistent across most breeds. In the Fresian breed the mean length of pregnancy was reported to be 332 days6 and, in a second study, 338 days.7 The mean gestational age for Arabian, Dutch Freiberger, and Draught foals was 332,8 336,9 and 343 days,7 respectively. Gestational length was reportedly shorter in United Kingdom pony mares with mean ages of 32510 and 333 days, with a range of 315 to 350 days.5 However, similar values to Thoroughbreds were reported for Haflinger ponies (341 days), Fjord ponies (342 days), and Shetland ponies (337 days) in the Netherlands.7 Caspian Miniature horses have a mean gestational period of 335 days,11 and similar values have been suggested for American Miniatures. Donkeys have a longer gestational period with mean duration of 371 days in Martina Franc and Ragusana jennies.12 Gestational length for mule foals is believed to be between that of donkeys and mares. There are several factors that influence the length of gestation, including foal gender and timing of breeding. Colts on average have a longer gestation than fillies, are heavier than fillies, have a heavier placenta, and take slightly longer to stand.4 The reported difference in gestational age between colts and fillies is small, around 1.5 to 2.5 days.2–4,6 The time of conception within the breeding season directly in-
fluences the duration of gestation. Mares conceiving early in the breeding season will have longer pregnancies than those bred towards the end of the season. The difference may be as much as 10 days1, 2 and appears to be related to photoperiod as, on average, gestational lengths can be reduced by 10 days with an extended 16-hour photoperiod that was begun on 1 December and maintained until foaling.13 The year of breeding may influence the duration of pregnancy, but specific year-to-year factors such as changes in weather patterns or feed have not been determined.3 A role for genetic factors in determining gestational length is unclear. An influence of mare, sire, and the sire of the dam was reported in Friesians,6 and an effect of sire was reported in Freibergers.9 A study of Thoroughbred mares concluded that dam, but not sire, had an impact on pregnancy duration.2 The age or parity of the mare does not influence the length of gestation.2, 3
Terminology The variability in gestational length in horses makes strict definitions of maturity difficult. Retrospective studies often report wide variability in gestational age, but do not report physical characteristics or survival of foals. Ranges of 305 to 365 days,3 315 to 387 days,2 and 286 to 370 days14 have been reported in Thoroughbreds from different sites around the world. Given this wide variability, any definitions determined solely on gestational age would falsely classify a small number of appropriately mature animals. The term ‘premature’ describes the birth of an infant or animal after a gestational period shorter than normal. A premature human infant is defined according to gestational age, i.e., delivered 21 or more days
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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before the mean pregnancy duration of 266 days. A definition of prematurity in the foal has included foals born prior to 320 days gestation.5 This value was determined, in part, through studies reporting significantly lower birthweights and poor outcomes of foals born before this time.15 An alternative definition of prematurity is foals born outside the lower 95% confidence interval of the normal gestational period. This would be less than 325 days3 or 327 days.1 Using similar logic, foals that are born after a gestational period outside of the upper 95% confidence interval limits should be regarded as ‘post-term’. This would be either 3563 or 357 days.1 There is an important distinction between post-term and post-mature, the latter describing a condition of increased neonatal morbidity as a consequence of failing placental function. ‘Dysmaturity’ is used to describe foals that have experienced some degree of intrauterine growth retardation. Affected animals often demonstrate signs of physical immaturity despite having a ‘normal’ gestational age.
Physical characteristics The physical characteristics of prematurity include a low birthweight, small body frame, a short and silky hair coat, a prominent domed head, periarticular laxity, and droopy ears. Flexural laxity is common with elevation of the toe, although some premature foals will show fetlock contracture. Muscle development is usually poor. Most will demonstrate generalized weakness and hypotonia and will have difficulty in standing. Post-mature foals usually have an acceptable birthweight but a large frame with poor muscle development, creating a ‘lanky’ appearance. Post-term or post-mature foals will often have long tails, a long hair coat, and erupted incisors. In term foals the central incisors normally erupt during the first 5–7 days of postnatal life. Fetlock contracture is common, although laxity can also be seen.
Causes Exogenous oxytocin or prostaglandins are important causes of pre-term birth. The chemical induction of parturition in 49 mares before 300 days gestation and in 31 mares between 300 and 320 days produced a foal survival rate of only 5%, with the youngest survivor delivered at 318 days gestation.16 There are several circumstances where premature induction of labor may occur. These include incorrect breeding dates, life-threatening maternal disease, evidence of significant in utero stress, or misinterpretation of in-
testinal colic as ineffective labor. Premature birth can occur as a sequelae to placental problems, such as infection, edema, and/or placental separation. The nature and duration of the stimulus can hasten fetal maturation and therefore affect postnatal survival. Placental insufficiency due to twinning is another cause of intrauterine growth retardation, premature parturition, or abortion. Consumption of tall fescue pasture infected with the endophyte, Neotyphodium coenophialum, by pregnant mares can produce a range of signs that include prolongation of gestation, perinatal mortality, and agalactia.17 The large skeletal frame of these post-term/post-mature foals predispose mares to dystocia. The delay in parturition may be linked to interference with fetal corticotropinreleasing hormones and delay in maturation of the hypothalamic–pituitary–adrenal (HPA) axis. Problems with cortisol-induced maturation of thyroid function are reflected by reduced tri-iodothyronine (T3) levels compared to control foals.18 Congenital hypothyroidism has been reported in foals in Western Canada.19 Affected foals have a prolonged gestation, dysmaturity with a range of musculoskeletal abnormalities including flexural deformities, delayed ossification, and mandibular prognathia. Consumption of diets that contain nitrate or that are deficient in iodine are postulated causes.20
Maturation of the fetal hypothalamic–pituitary–adrenal axis The final stage in maturation of several key organ systems is related to changes in the fetal HPA axis that occur in the latter stages of pregnancy.21 While fetal cortisol is critical for the final stages of organ maturation, early or excessive levels have been associated intrauterine growth restriction in some species. Mechanisms are in place in the normal pregnancy to protect the fetus from cortisol during much of gestation. The enzyme, 11β-hydroxysteroid dehydrogenase (11β-HSD) type 2 converts biologically active cortisol into the inactive cortisone in the placenta, reducing the exposure of the fetus to glucocorticoids during pregnancy. An increase in prostaglandins before parturition results in upregulation of 11β-HSD type 1, an isoform which favors the production of cortisol from cortisone.22 The relative proportions of these isoforms have yet to be determined in the equine placenta.23 The postnatal adrenal gland, under the influence of ACTH from the pituitary, can readily synthesize cortisol from cholesterol and pregnenolone. The enzymes required for this conversion, 3β-HSD, P450scc, and P450C17 (also referred to as 17α-hydroxylase),
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Prematurity, Dysmaturity and Assessment of Maturity are either inhibited or deficient during most of pregnancy, again protecting the developing fetus from excessive cortisol during development. During the majority of gestation the principal products of steroidogenesis are progesterone and the 5α-reduced progestagens.23 Foals undergo enhanced adrenal activity just before birth such that the healthy term newborn foal will have high plasma cortisol and ACTH concentrations in the first hours after birth.24 This coincides with substantial change in the amount and localization within the late term foal adrenal gland of 3β-HSD, P450scc, and P450C17.25 Based on work in other species, it is likely that the rise in fetal cortisol has a direct effect on the placenta to increase prostaglandin H synthase 2, leading to secretion of prostaglandins.21, 22, 26 Prostaglandins further stimulate the fetal HPA axis, stimulate placental 11β-HSD1 (which favors the production of cortisol from cortisone), and facilitate the conversion of estrogen from pregnenolone. In contrast to other species the mare foals in the face of a declining concentrations of estrogen. The production of fetal cortisol occurs during the final 48–72 hours of pregnancy in mares.27 There are a number of key events that are tightly associated with the prepartum increase in ACTH and cortisol,23 including changes in red and white blood cell parameters, most notably a large increase in the neutrophil to lymphocyte ratio.27, 28 In other species the prepartum rise in plasma cortisol induces deiodination of the outer ring of T4 to produce the biologically active T3.29 Normal term foals have very high levels of thyroid hormones, including T3, at the time of birth.30 These levels decline over the initial weeks or months of postnatal life. Adequate levels of T3 are required for a number of biological functions including postnatal thermogenesis and respiratory function, particularly the normal post-delivery reabsorption of lung liquid.31 There is a close relationship between T3 levels and cortisol in premature, dysmature, and mature foals.32
Accelerated maturation of the fetal HPA axis There are several known inducers of premature maturation of the fetal HPA axis. Hypoxemia is a potent stimulator of the axis in sheep with rises in both fetal ACTH and cortisol.33 The HPA axis can also be manipulated using exogenous glucocorticoids; betamethasone is commonly administered to women in danger of pre-term birth to hasten HPA maturation and improve postnatal survival. Poor nutrition before and after conception in sheep produces a shortened gestational period and accelerated maturation
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of the axis.26 Placental and/or fetal infection can exert a similar effect. Cytokines induced by infection increase prostaglandin synthesis, which in turn exerts a range of actions along with promoting fetal cortisol production. The precise stimuli leading to precocious HPA axis maturation in foals are not well described. The exception is placentitis, where foals are often delivered preterm with laboratory findings consistent with HPA axis maturation. Spatial and nutritional deprivation resulting from placement of a Thoroughbred foal in a pony uterus also resulted in premature maturation of the fetal adrenal gland and increased plasma cortisol.10, 34 Unfortunately many problems in the late mare do not have a significant effect on foal maturity. It is not uncommon for mares to deliver pre-term foals after a stressful incident, such as the hypotension and hypoxemia-associated general anesthesia. Colic surgery in mares during the final 60 days of pregnancy results in a high rate of abortion or pre-term delivery of compromised foals with a high mortality rate.35 It is likely that the insult in these cases was so severe that the interval between surgery and delivery was inadequate for maturation of the axis to occur.
The late pregnant mare Management of the high-risk pregnant mare is aimed at improving fetal well-being and preventing preterm birth. This not only increases the likelihood of HPA axis maturation but also effective ossification and maturation of other body systems. There are circumstances where maintenance of the pregnancy may not be possible. Consequently, any treatment that could hasten fetal maturity and improve postnatal survival would be advantageous. In contrast to human antenatal therapy, the administration of corticosteroids to pregnant mares appears to have little effect on maturation of the HPA axis in foals, at least at doses used commonly in clinical practice.5 Relatively high doses of dexamethasone (100 mg intramuscularly once daily) administered to Thoroughbred mares between 315 and 317 days’ gestation did produce premature parturition when compared to saline-treated control mares (322 and 335 days, respectively).36 The treated mares had less mammary development, produced lower-quality colostrum, and delivered lighter and smaller foals. However, there were no differences in time taken to stand or suck, total white blood cell counts, peripheral neutrophil : lymphocyte ratios, or mean red cell volumes between foals from the two groups. Administration closer to the time of parturition may lead to problems with dystocia.
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Experimentally the intramuscular injection of ACTH1–24 to the fetus in utero led to measurable increases in fetal cortisol. The effect was dependent on gestational age, with maximal responses around 313 days and no response when administered before day 295 of gestation.27 Similarly, direct injection of CRH or betamethasone into fetal muscle under ultrasound guidance increased maternal progestagen levels, a finding consistent with maturation of the fetal adrenal gland.37, 38 The clinical application of these techniques have not been widely practiced due to the small, but real, risk of abortion attributable to the procedure itself. Exogenous ACTH1–24 (4–5 mg) administered to normal late pregnant pony mares at days 300, 301, and 302 days produced a shortened gestational period and smaller foal, but with evidence of accelerated fetal maturation.39 Management of the post-term pregnancy can also be very difficult, particularly in view of the concerns expressed by many owners when the anticipated ‘due date’ has long passed. Given the wide variation in gestational range it is usually in the best interests of the foal to let the pregnancy continue. Ideally ultrasonographic assessment of the fetus and chorioallantois should be made, looking for evidence of fetal stress or thickening or detachment of the fetal membranes. Any induction of parturition should be based on appropriate changes in milk electrolytes, in the knowledge that electrolyte concentrations may be unreliable predicators of foaling.40 Mares grazing endophyte-infected tall fescue can be medicated with dopamine receptor antagonists, such as domperidone.17
Laboratory assessment Premature or dysmature foals that struggle for postnatal survival typically have minimal cortisol secretion in the face of adequate levels of ACTH. The expected increase in plasma cortisol in response to exogenous ACTH1–24 (0.125 mg IM) is typically inadequate.24 Low cortisol levels coupled with increased progestagens would provide further evidence that effective maturation of HPA axis has not occurred in the foal before birth.41 Total white blood cell count, neutrophil and lymphocyte counts, and the neutrophil to lymphocyte ratio (N:L) are influenced by the degree of HPA axis maturity and, as such, are good predictors of survival. Inadequate adrenal maturation will be reflected by a low total white cell and neutrophil count, and a N:L characteristically less than 1:1.28 Neutropenia is also a common finding in neonatal sepsis. Evidence of shifting towards immature cell types and neutrophil toxicity indicate primary sepsis or prema-
turity complicated by sepsis. Premature foals that fail to improve their total white blood cell and neutrophil counts over the initial 48 hours of treatment usually do not survive. An elevated MCV is reported in non-stressed preterm foals.28 Hyperfibrinogenemia and mature neutrophilia are positive prognostic factors in premature foals as it reflects prepartum exposure to bacterial infection and hastened maturation of the HPA axis. Often the higher the WBC count the better the prognosis, assuming any lingering infection responds to antimicrobial therapy without complication. Hypoglycemia is a common finding in premature or dysmature foals and reflects poor glycogen reserves and reduced glucose-6-phosphatase levels.42, 43 Plasma creatinine levels are often elevated in newly born pre-term or dysmature foals due to placental dysfunction. This increase is independent of foal renal function.44
Clinical course The clinical course will be dictated by the extent of endocrinological maturity, the presence of additional perinatal problems, and the degree of physical maturity. Foals born prematurely, but exposed to an appropriate in utero stress, will typically be weak and depressed in the immediate postpartum period and may require resuscitation. After a longer than normal period of postural adaptation they will usually manage to stand, but will often require assistance. Suckle reflex and appetite may be reduced or absent and many will need to be fed initially via nasogastric tube. They will frequently have trouble maintaining body temperature and blood glucose. However, after 24 hours many will improve in physical strength, appetite, and mentation. Foals with incomplete maturation of the HPA axis may mimic the clinical course described above up until 6–18 hours of age, after which a range of progressive abnormalities develop. Owners or attendants are often buoyed by an initial period of alertness in the immediate postpartum period, but foals rarely stand unassisted during this time and the level of alertness commonly drops over the ensuing hours. Common clinical problems include weakness, depression, intolerance to milk, seizures, respiratory failure, and cardiovascular collapse. The signs of circulatory problems include a reduction in peripheral pulse pressure, cool extremities, darkening of the oral mucous membranes, reduced urine flow, and the development of subcutaneous edema. This coincides with worsening neurological function. Death will occur without aggressive support, and even with intensive care, mortality rates are high.
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Management Successful management of the premature or dysmature foal requires anticipation and early recognition of problems. Most will experience some degree of pulmonary insufficiency characterized by reduced ventilation capacity, tachypnea, hypoxemia, and varying levels of hypercapnia, the severity of which is linked closely to maturation of the HPA axis. Arterial blood gas analysis and measurement of O2 saturation of hemoglobin are the most sensitive clinical indicators of respiratory function. Reduced O2 tensions in premature foals are due to several factors including extrapulmonary shunts45 and ventilation/perfusion mismatching as a result of a poorly reactive pulmonary vasculature and dependent atelectasis. Deficiency of lung surfactant is unlikely to play a major role in most premature foals as surfactant activity is fully developed in most foals by 300 days, although it may be delayed in some foals until after 340 days.46 Treatment is dependent on the severity of dysfunction, but most foals will benefit from intranasal humidified oxygen. Initial flow rates of 5 liters/minute are recommended, with adjustment in flow dictated by positive changes in arterial blood gas analyses or improvement in ventilation rate and depth. Long periods of lateral recumbency should be avoided, to minimize the dependent atelectasis. If the foal is unable to stand then placement in sternal recumbency is recommended. This is made easier by use of a V-pad. The most severe form of respiratory dysfunction is neonatal respiratory distress syndrome (RDS). The condition is often linked to delayed HPA axis maturation and is characterized by progressive respiratory difficulty and failure, severe hypoxemia and hypercapnia, coma, and death. A common clinical scenario involves foals delivered prematurely by emergency cesarean section due to acute maternal illness, such as a colonic volvulus. Intervention requires mechanical ventilation, but outcomes are poor irrespective of the level of care. The respiratory complications associated with human pre-term infants are often managed using exogenous surfactant and glucocorticoids, such as hydrocortisone. The use of glucocorticoids in premature or dysmature foals is compelling. There has also been some interest in using multiple injections of Depot ACTH1–24 in postpartum premature foals to accelerate maturation. Exogenous surfactant has been administered to high-risk foals, with mixed success in the author’s experience. The expense of the product, coupled with the difficulty of administration and mixed outcomes has resulted in limited uptake in equine practice. Ataxic or periodic breathing is a feature of several neonatal problems, including prematurity.
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This can occasionally produce a poorly compensated respiratory acidosis. Treatment may include intravenous infusion of doxapram hydrochloride (0.02–0.05 mg/kg/h) or oral caffeine (10 mg/kg p.o. loading dose, then 2.5–5.0 mg/kg p.o. s.i.d.). Management of a failing cardiovascular system is particularly challenging. This is often complicated by inconsistent responses to standard vasoreactive therapy. Early recognition is critical. This is through physical examination, recognition of urine output, and changes in blood pH, lactate, and anion gap. Indirect or direct measurement of mean blood pressure does not accurately reflect tissue perfusion, although may help assess response to treatment. An initial approach to the treatment of failing perfusion may involve intravenous plasma followed, if necessary, by dopamine (3–10 µg/kg/min) and/or dobutamine infusion (5–20 µg/kg/min). Constant rate infusion (CRI) of epinephrine or norepinephrine can be used if additional pressor support is required. Vasopressin (0.25–0.50 mU/kg/min) has been used successfully in combination with dopamine and dobutamine to manage resistant hypoperfusion in neonates. It is important to avoid fluid overloading, particularly given that many foals have abnormal renal function. Urine output should be appropriate for the volume of fluids administered, and anuria or oliguria treated aggressively. This may include dopamine or fenoldopam infusion, furosemide boluses or CRI, or mannitol infusion. Establishment of urine flow is critical. The over-administration of replacement solutions such as lactated Ringer’s solution often results in significant hypernatremia. Limited gluconeogenic enzyme activity and glycogen stores predispose to hypoglycemia. This is managed acutely by infusion of 10 mL/kg bodyweight of a 10% glucose/dextrose solution over several minutes, followed by a constant infusion at about 6 mg/kg/minute. This equates to around 200 mL/hour of a 5% dextrose solution to a 30 kg foal. Blood glucose should be monitored regularly. Gastrointestinal tract problems usually relate to overfeeding, with signs that include abdominal distention, reduced fecal passage, gastric reflux, and colic. Feeding should be restricted to small volumes (e.g., 10–50 mL hourly) until the foal appears to be systemically stable. Concurrent parenteral nutrition is indicated to prevent weight loss. Foals should be assessed for frequency of fecal passage, changes in abdomen size (using a soft measuring tape), and frequent ultrasonographic measurement of intestinal diameter and gastric size. The combination of prolonged asphyxia and prematurity is a risk factor for the development of necrotizing enterocolitis. Premature and dysmature foals are more susceptible to hypothermia than term foals. Body
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temperature needs careful management as rapid warming may result in peripheral vasodilatation and possible cardiovascular collapse. Initially, the foal should be covered by blankets and removed from any drafts. Fluids should be warmed before use. Once the foal begins to improve, heat lamps and circulating warm-water blankets can be used. Skeletal maturity is assessed by radiography of both carpus and tarsus for evidence of ossification. Accelerated maturation of the HPA axis is critical for development of several key organ systems but does not appear to influence the rate of ossification. This, coupled with periarticular laxity, predisposes the premature foal to long-term skeletal problems. Restriction of exercise is recommended to minimize collapse of developing carpal or tarsal bones, but forced recumbency may exacerbate other problems, such as lung disease. Some weight bearing is important to encourage ossification, but if there is periarticular laxity the resultant load bearing is abnormal and further increases the risk of cuboidal bone crush injury. Laxity and exercise will also predispose to angular limb deformity. Splinting and attention to hoof care is recommended if angular limb deviation develops. Flexural deformities and laxities will improve over time. Dorsal splints are recommended for flexural deformities involving the fetlock, and heel extensions are helpful to foals with flexural laxity. Most premature or dysmature foals will experience some degree of failure of passive transfer for a variety of reasons. Plasma transfusion is often indicated even in foals less than 18–24 hours of age. Foals born prematurely due to placental infection are often septic, often with bacterial lung disease. This, coupled with the fact that many foals have an impaired immune system, warrants the use of broad-spectrum antimicrobial therapy.
Formulating a prognosis The prognosis for short-term and long-term survival is formulated from several factors. These include the reason for early delivery, additional problems during the perinatal period, laboratory data, what resources are available, financial considerations, and a knowledge of intended use. In a study of foals with a gestational age of 320 days or less, short-term survival was in part predicted by total white blood cell count, neutrophil count, lymphocyte count, and the neutrophil to lymphocyte (N:L) ratio at admission. 47 The N:L ratio of surviving premature foals (12.5 ± 1.7:1) was well above that reported for both normal term foals and that of non-surviving premature foals. Many of the surviving animals were exposed to confirmed or suspected placental infection. Outcome was not affected by gestational age (survivors
311 days and non-survivors 307 days). These data confirm that in utero stress, with probable maturation of the HPA axis, is a good prognostic factor for survival in foals delivered pre-term. Most premature or dysmature foals will be smaller than their peers for the initial 12–18 months of life, but this physical difference becomes less noticeable after that time. Surviving premature Thoroughbred foals are less likely to be successful athletes than their siblings.47 There are no data on short- or long-term outcomes for postmature foals, but it is likely that the outlook would be very good once the foal has survived the immediate postpartum period.
References 1. Rossdale PD, Short RV. The time of foaling of thoroughbred mares. J Reprod Fertil 1967; 13(2):341–3. 2. Ropiha RT, Matthews RG, Butterfield RM, Moss FP, McFadden WJ. The duration of pregnancy in thoroughbred mares. Vet Rec 1969;84(22): 552–5. 3. Hintz HF, Hintz RL, Lein DH, Van Vleck LD. Length of gestation periods in Thoroughbred mares. J Equine Med Surg 1979; 3:289–92. 4. Campitelli S, Carenzi C, Verga M. 1982. Factors which influence parturition in the mare and development of the foal. Appl Anim Ethol 1982; 9:7–14. 5. Rossdale PD. Clinical view of disturbances in equine foetal maturation. Equine Vet J Suppl 1993; 14:3–7. 6. Sevinga M, Barkema HW, Stryhn H, Hesselink JW. Retained placenta in Friesian mares: incidence, and potential risk factors with special emphasis on gestational length. Theriogenology 2004; 61(5):851–9. 7. Bos H, Van Der Mey GJW. Length of gestation periods of horses and ponies belonging to different breeds. Livest Prod Sci 1980; 7:181–7. 8. El-Wishy AB, El-Sayed MAI, Seida AA, Ghoneim IM, Serur BH. Some aspects of reproductive performance in Arabian mares in Egypt. Reprod Domest Anim 1990; 25(5):227–34. 9. Giger R, Meier HP, Kupfer U. Gestation length of Freiberger mares with mule and horse foals. Schweiz Archiv Tierheilkunde 1997; 139(7):303–7. 10. Allen WR, Wilsher S, Stewart F, Ousey J, Fowden A. The influence of maternal size on placental, fetal and postnatal growth in the horse. II. Endocrinology of pregnancy. J Endocrinol 2002; 172(2):237–46. 11. Dardari S, Mirian J, Mohammad D, Saeedabadi M. ‘Duration of the Caspian Miniature Horse Gestation Period.’ Paper presented at the 37th Annual Meeting of the Society for the Study of Reproduction, Vancouver, Canada, 2004; p. 812. 12. Carluccio A, De Amicis I, Panzani S, Tosi U, Faustini M, Veronesi MC. Electrolytes changes in mammary secretions before foaling in jennies. Reprod Domest Anim 2008; 43(2):162–5. 13. Hodge SL, Kreider JL, Potter GD, Harms PG, Fleeger JL. Influence of photoperiod on the pregnant and postpartum mare. Am J Vet Res 1982; 43(10):1752–5. 14. Sato K, Miyaki M, Sugiyama K, Yoshikawa T. An analytical study of the duration of gestation in horses. Jap J Zoo Tech Sci 1973; 44:375–9. 15. Koterba AM. Definitions of equine perinatal disorders: problems and solutions. Equine Vet Educ 1993; 5(5):271–3.
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Prematurity, Dysmaturity and Assessment of Maturity 16. Leadon DP, Jeffcott LB, Rossdale PD. Behavior and viability of the premature neonatal foal after induced parturition. Am J Vet Res 1986; 47(8):1870–3. 17. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995; 73(3):899– 908. 18. Boosinger TR, Brendemuehl JP, Bransby DL, Wright JC, Kemppainen RJ, Kee DD. 1995. Prolonged gestation, decreased triiodothyronine concentration, and thyroid gland histomorphologic features in newborn foals of mares grazing Acremonion coenophialum-infected fescue. Am J Vet Res 1995; 56(1):66–9. 19. Allen AL, Doige CE, Fretz PB, Townsend HG. Hyperplasia of the thyroid gland and concurrent musculoskeletal deformities in western Canadian foals: reexamination of a previously described syndrome. Can Vet J 1994; 35(1):31–8. 20. Allen AL, Townsend HG, Doige CE, Fretz PB. A casecontrol study of the congenital hypothyroidism and dysmaturity syndrome of foals. Can Vet J 1996; 37(6):349–58. 21. Challis JR, Sloboda D, Matthews SG, Holloway A, Alfaidy N, Patel FA, Whittle W, Fraser M, Moss TJ, Newnham J. The fetal placental hypothalamic-pituitary-adrenal (HPA) axis, parturition and postnatal health. Mol Cell Endocrinol 2001; 185(1–2):135–44. 22. Alfaidy N, Xiong ZG, Myatt L, Lye SJ, MacDonald JF, Challis JR. Prostaglandin F2alpha potentiates cortisol production by stimulating 11beta-hydroxysteroid dehydrogenase 1: a novel feedback loop that may contribute to human labor. J Clin Endocrinol Metab 2001; 86(11):5585–92. 23. Ousey JC. Peripartal endocrinology in the mare and foetus. Reprod Domest Anim 2004; 39(4):222–31. 24. Silver M, Ousey JC, Dudan FE, Fowden AL, Knox J, Cash RS, Rossdale PD. Studies on equine prematurity 2: Post natal adrenocortical activity in relation to plasma adrenocorticotrophic hormone and catecholamine levels in term and premature foals. Equine Vet J 1984; 16(4):278–86. 25. Han X, Fowden AL, Silver M, Holdstock N, McGladdery AJ, Ousey JC, Allen WR, Rossdale PD, Challis JR. Immunohistochemical localisation of steroidogenic enzymes and phenylethanolamine-N-methyl-transferase (PNMT) in the adrenal gland of the fetal and newborn foal. Equine Vet J 1995; 27(2):140–6. 26. Challis JR, Sloboda DM, Alfaidy N, Lye SJ, Gibb W, Patel FA, Whittle WL, Newnham JP. Prostaglandins and mechanisms of preterm birth. Reproduction 2002; 124(1):1–17. 27. Silver M, Fowden AL. Prepartum adrenocortical maturation in the fetal foal: responses to ACTH. J Endocrinol 1994; 142(3):417–25. 28. Jeffcott LB, Rossdale PD, Leadon DP. Haematological changes in the neonatal period of normal and induced premature foals. J Reprod Fertil Suppl 1982; 32:537–44. 29. Nathanielsz PW, Berghorn KA, Derks JB, Giussani DA, Docherty C, Unno N, Davenport A, Kutzlers M, Koenen S, Visser GH, Nijland MJ. Life before birth: effects of cortisol on future cardiovascular and metabolic function. Acta Paediatr 2003; 92(7):766–72. 30. Irvine CH, Evans MJ. Postnatal changes in total and free thyroxine and triiodothyronine in foal serum. J Reprod Fertil Suppl 1975; 23:709–15. 31. Wallace MJ, Hooper SB, Harding R. Role of the adrenal glands in the maturation of lung liquid secretory mechanisms in fetal sheep. Am J Physiol 1996; 270(1 Pt 2):R33–40.
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32. Silver M, Fowden AL, Knox J, Ousey J, Cash R, Rossdale PD. Relationship between circulating tri-iodothyronine and cortisol in the perinatal period in the foal. J Reprod Fertil Suppl 1991; 44:619–26. 33. Matthews SG, Challis JR. Developmental regulation of preproenkephalin mRNA in the ovine paraventricular nucleus: effects of stress and glucocorticoids. Brain Res Dev Brain Res 1995; 86(1–2):259–67. 34. Ousey JC, Rossdale PD, Fowden AL, Palmer L, Turnbull C, Allen WR. Effects of manipulating intrauterine growth on post natal adrenocortical development and other parameters of maturity in neonatal foals. Equine Vet J 2004; 36(7):616–21. 35. Santschi EM, Slone DE, Gronwall R, Juzwiak JS, Moll HD. Types of colic and frequency of postcolic abortion in pregnant mares: 105 cases (1984–1988). J Am Vet Med Assoc 1991; 199(3):374–7. ¨ 36. Ousey JC, Kolling M, Allen WR. The effects of maternal dexamethasone treatment on gestation length and foal maturation in Thoroughbred mares (Abstract). Anim Repro Sci 2006; 94:436–8. 37. Rossdale PD, McGladdery AJ, Ousey JC, Holdstock N, Grainger L, Houghton E. Increase in plasma progestagen concentrations in the mare after foetal injection with CRH, ACTH or betamethasone in late gestation. Equine Vet J 1992; 24(5):347–50. 38. Ousey JC, Rossdale PD, Dudan FE, Fowden AL. The effects of intrafetal ACTH administration on the outcome of pregnancy in the mare. Reprod Fertil Dev 1998; 10(4): 359–67. 39. Ousey JC, Rossdale PD, Palmer L, Grainger L, Houghton E. Effects of maternally administered depot ACTH(1–24) on fetal maturation and the timing of parturition in the mare. Equine Vet J 2000; 32(6):489–96. 40. Brown-Douglas CG, Perkins NR, Stafford KJ, Hedderley DI. Prediction of foaling using mammary secretion constituents. N Z Vet J 2002; 50(3):99–103. 41. Houghton E, Holtan D, Grainger L, Voller BE, Rossdale PD, Ousey JC. Plasma progestagen concentrations in the normal and dysmature newborn foal. J Reprod Fertil Suppl 1991; 44:609–17. 42. Fowden AL, Silver M, Ellis L, Ousey J, Rossdale PD. Studies on equine prematurity 3: Insulin secretion in the foal during the perinatal period. Equine Vet J 1984; 16(4):286–91. 43. Fowden AL, Mundy L, Ousey JC, McGladdery A, Silver M. Tissue glycogen and glucose 6-phosphatase levels in fetal and newborn foals. J Reprod Fertil Suppl 1991; 44:537–42. 44. Brewer BD. Renal Disease. In: Koterba et al. (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 446–58. 45. Rose RJ, Stewart J. ‘Basic Concepts of Respiratory and Cardiovascular Function and Dysfunction in the Full Term and Premature Foal.’ Paper presented at the Proceedings of the annual convention of the American Association of Equine Practitioners, 1983; pp. 167–78. 46. Pattle RE, Rossdale PD, Schock C, Creasey JM. The development of the lung and its surfactant in the foal and in other species. J Reprod Fertil Suppl 1975; 23:651–7. 47. Lester GD. ‘Outcomes in Foals With a Gestational Age Less Than 320 Days.’ Paper presented at the Dorothy Havermeyer Neonatal Septicemia Workshop, Talliores, France, 2001; pp. 42–4.
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Chapter 13
Resuscitation (Foal and Birth) Niamh M. Collins, Jane E. Axon and Jonathan E. Palmer
Resuscitation of the neonatal foal may be required immediately after birth or during hospitalization. Birth is a high-risk period, and neonates may fail to make the transition from fetal to neonatal physiology. Foals born by cesarean section or with assisted vaginal deliveries, and foals from mares with placentitis, premature placental separation, or maternal systemic illness, are at increased risk of peripartum complications. However, foals from healthy mares with apparently normal births can also arrest. In neonatal foals, respiratory arrest usually precedes cardiac arrest as primary cardiac disease is rare in the foal. This is in contrast to humans, in whom primary cardiac arrest is most common because of the high incidence of coronary artery disease. Respiratory arrest can be more easily corrected with oxygen supplementation, intubation, and positive pressure ventilation than cardiac arrest and therefore carries a better prognosis. However, if left untreated, respiratory arrest (with progressive hypoxemia and respiratory and metabolic acidosis) can rapidly result in bradycardia, then asystole and death. Cardiopulmonary arrest in hospitalized foals is usually secondary to other serious conditions such as septic shock, respiratory failure, or severe metabolic disorders. Before cardiopulmonary arrest, several changes may be observed, including obtundation, hypothermia, bradycardia, hypotension, and dilated, unresponsive pupils. The definitive clinical signs of cardiopulmonary arrest include loss of consciousness, absence of spontaneous ventilation and/or absence of heart sounds on auscultation. Patient selection is critical to successful resuscitation. Foals with end-stage organ failure are unsuitable candidates for resuscitation given the poor short- and long-term prognosis for survival. Cardiopulmonary cerebral resuscitation (CPCR) is the restoration of spontaneous breathing and circulation. Careful planning is essential for successful foal resuscitation. A cardiopulmonary cerebral resuscita-
tion protocol must be in place in each hospital to allow prompt intervention in critical situations. Ideally, the resuscitation team should consist of at least three people: (i) primary clinician in charge of intubating and ventilating the foal, directing the other members of the team and making therapeutic drug decisions; (ii) a person in charge of monitoring heart rate if spontaneous circulation is present and instituting chest compressions if necessary; and (iii) a person to dry the foal, administer drugs as directed by the primary clinician, and relieve the operator performing cardiac compressions. It is important that veterinarians are trained and proficient in providing CPCR to minimize the stress and maximize the efficiency of CPCR. It is also essential to have emergency equipment and drugs readily available for easy transport (Figure 13.1a,b). The resuscitation kit must be kept readily available in a central location and fully stocked at all times (Table 13.1). It should also contain an information sheet which details drug dosages in terms of the amount in milliliters needed for the average 50-kg foal to avoid incorrect dosing during CPCR (Table 13.2).
Birth resuscitation The foal should be placed in lateral recumbency on a flat, hard surface immediately after birth. An uneven or soft surface results in excessive movement of the foal and reduces the force of thoracic compressions. The foal should be placed on intranasal oxygen insufflation at 8–10 L/min while initial assessment is being performed. The initial 20 seconds after birth should be used to assess the foal’s breathing (spontaneous, gasping, or apnea) and effectiveness of spontaneous circulation (asystole, heart rate 40–60 beats/min, or heart rate greater than 60 beats/min). Fetal membranes should be cleared away from the foal’s nose. The foal’s head and thorax should be vigorously towel dried to stimulate spontaneous breathing. The
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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(a)
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(b)
Figure 13.1 (a and b) An oxygen bag containing equipment for ventilation and a carry box for the drugs required for resuscitation are ideal for easy transport.
foal’s ear canal or nasal passages can be stimulated to encourage breathing. The thorax can be gently compressed. If the foal has sustained fractured ribs during the birthing process, the side with the fractured ribs should be placed closest to the ground. The foal should be assessed for severe congenital defects (e.g., severe arthrogryposis, hydrocephalus) that may make resuscitation undesirable. If the foal remains apneic, has an irregular, gasping respiratory pattern and/or has a non-perfusing bradycardia (less than 60 beats/min), rapid establishment of an airway and institution of positive pressure ventilation is immediately required (Figure 13.2).
Ventilation Nasotracheal intubation should be attempted initially with the foal positioned in lateral recumbency.
Raising the head should be avoided because this action may further compromise cerebral circulation. A cuffed 9- to 10-mm internal diameter endotracheal tube is suitable for the average Thoroughbred or Standardbred foal. It should be approximately 55 cm in length, which situates the cuff in the midtracheal region. The foal’s head should be in a straight line with the neck to increase the likelihood of the endotracheal tube being placed in the trachea. The tube should be directed medially and ventrally so it enters the ventral meatus. When the tube is at the level of the arytenoids, twisting the tube may aid the beveled end to spread the arytenoids, maximizing the chance of entering the trachea. When the tube is being advanced into the trachea, minimal resistance is palpated. Palpating the proximal esophagus as the tube is advanced allows retraction of the tube without delay if it enters the esophagus. Inadvertent
Table 13.1 Essential and non-essential items of resuscitation kit Resuscitation kit contents Essential items
Non-essential items
Endotracheal tubes 8–10 mm in diameter and 55 cm in length Self-inflating resuscitation bag Oxygen cylinder, flowmeter and tubing (oxygen needs to be humidified if using for longer term) Syringe to inflate cuff of endotracheal tube Drugs, minimum requirement: Epinephrine (adrenaline), 3 syringes containing dose for 50-kg foal Vasopressin, 1 syringe containing dose for 50-kg foal Intravenous catheter (16-gauge, 8.3 cm (3.25 inches) and 13.3 cm (5.25 inches) over-the-needle catheters)a
Electrocardiogram Non-invasive blood pressure monitor Capnograph Defibrillator
2-ml syringes and 20-g needles Intravenous fluids and fluid administration sets Towels Pen light a
E.G. Angiocath, Becton Dickinson Infusion Therapy Systems Inc., Sandy, UT.
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Table 13.2 Dosages of drugs commonly used in foal resuscitation Drug
Dose (per kg)
Supplied
mL/kg
mL/50 kg
Epinephrine, low dose Epinephrine, high dose Vasopressin Atropine Lidocaine
0.01–0.02 mg 0.1 mg 0.6 U 0.02 mg 1.5 mg
1 mg/mL 1 mg/mL 20 U/mL 20 mg/mL 20 mg/mL
0.01–0.02 0.1 0.03 0.001 0.075
0.5–1 5 1.5 1.8 3.75
Clear airway of mucus Provide tactile stimulation
YES
Repeat doses 3–5 min apart Repeat doses 3–5 min apart Once only Maximum of two times Every 5 mins, max 3.3 mg/kg
Vigorously dry and warm Evaluate if CPCR appropriate
BREATHING?
Below 60
Evaluate HR
Notes
NO or gasping
PPV with 100% oxygen
15 - 30 sec
Evaluate HR
Above 60 > 60
40-60
< 40
Evaluate color and oxygenation HR increasing? Continue PPV YES Blue
Pink
Provide INO2
Watch for spontaneous respiration then discontinue ventilation
Continue ventilation
NO
Start chest compressions
If nonperfusing HR after 30 sec
Administer drugs Observe and monitor
Continue PPV Continue chest compressions
Figure 13.2 Overview of resuscitation of the foal at birth. HR, heart rate; PPV, positive ventilation; INO2 , intranasal oxygen insufflation. (Adapted from Palmer JE. Foal cardiopulmonary resuscitation. In: Orsini JA, Divers TJ (eds) Manual of Equine Emergencies, 2nd edn. Philadelphia: Saunders, 2002; p. 606.)
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Figure 13.3 Cardiopulmonary cerebral resuscitation of a neonatal foal following a dystocia and respiratory arrest. An endotracheal tube has been placed and positive pressure ventilation is being administered. One clinician is monitoring the heart rate while a second clinician secures intravenous access.
inflation of the stomach is clearly harmful. The tube should be advanced until only the adaptor is protruding at the foal’s nasal passageways to minimize dead space during ventilation (Figure 13.3). The cuff should be inflated until no air leak is heard when a breath is delivered to ensure a tight seal during positive pressure ventilation. As silicone tubes warm to body temperature, the cuff may require further inflation to maintain the seal. Overinflation of the cuff should be avoided to prevent obstructing the tube. The thoracic wall should be seen to rise when the first breath is given. If attempts at nasotracheal intubation are not immediately successful, the operator should attempt alternate methods such as oral intubation, the use of an airway exchange catheter, or, as a last resort, tracheostomy. Oral intubation can result in damage to the endotracheal tube as the foal regains consciousness and the respiratory tract is contaminated with oral bacteria. However, in foal resuscitation, the rapid establishment of an airway is of paramount importance. An airway exchange catheter can be used to act as a stylet, potentially facilitating a difficult intubation. Administration of oxygen through the airway exchange catheters diminishes the potential for hypoxia while attempts are made at intubation.
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A self-inflating resuscitation bag (e.g., Ambu SPUR adult disposable resuscitator with no mask; Ambu Inc., Glen Burnie, MD) designed for human adult resuscitation (1600 mL with a 2600-mL oxygen reservoir bag) connected to the endotracheal tube is a safe and easy to use method for providing ventilation.1, 2 If attached to an oxygen source via the gas inflow port, high concentrations of oxygen can be delivered. Resuscitation using 100% oxygen is recommended; however if unavailable, resuscitation with room air can be effective.3 The optimum breathing rate in neonatal foals is not known. Rates of 10–20 breaths/min have been suggested;1 however, rates on the lower end of this range (10 breaths/min) are currently being recommended when CPCR is being initiated2 and a faster rate (20 breaths/min) initiated once return of spontaneous circulation has occurred. Studies in adult pigs have shown that a rate of 30 breaths/min results in dramatically lower survival rates than 12 breaths/min.4 Increased intrathoracic pressure induced by positive pressure ventilation interferes with cardiac return and decreases coronary perfusion.5 Cardiac output during cardiopulmonary resuscitation is approximately 70–75% lower than normal cardiac output.6 Intrapulmonary shunting, which occurs during cardiopulmonary resuscitation, further decreases effective alveolar perfusion. This results in a dramatic reduction in the required minute volume from normal values,7 supporting the use of a low respiratory rate during resuscitation. Careful observation of chest excursion is the best way to guide tidal volume. The goal is not a deep breath. Also, a short inspiratory time should be used to minimize the negative effect on coronary perfusion.2 If an endotracheal tube is not immediately available during foal resuscitation, mouth-to-nose ventilation can be effective. This is facilitated by foals being obligate nasal breathers. The fingers of one hand are placed just behind the foal’s chin with the palm and thumb occluding the down nostril and placing the other hand on the foal’s poll so that the head can be maximally dorsiflexed, straightening the airway, and thus decreasing the risk of aerophagia.2 While breathing into the up nostril, the chest can be observed to rise, ensuring the proper tidal volume. The fractional inspired oxygen content of exhaled air is approximately 16–18%, but this may be adequate when perfusion is failing. A face mask and resuscitation bag or pump can also be utilized and is useful on stud farms without resident veterinarians. The use of oxygen demand valves is not recommended in foal resuscitation, as there is an increased risk of barotrauma, particularly with inexperienced operators. Doxapram has been traditionally recommended for stimulation of respiration; however, doxapram administration is not recommended in human infants because the drug has been shown
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Figure 13.4 Ex utero intrapartum treatment (EXIT) procedure. A nasotracheal tube is placed if the nares of the foal can be palpated in the mare’s pelvic canal and positive pressure ventilation is started. This procedure improves the chances of a live foal from a dystocia or cesarean section.
to decrease cerebral blood flow and increase cerebral oxygen consumption and requirement.8–10 In secondary apnea (which is most common in foals which have postnatal respiratory arrest), the respiratory center is not responsive to doxapram; thus, use of this drug is no longer recommended in foal CPCR.1 If a dystocia is present and it is possible to gain access to the foal’s nares, intubation and positive pressure ventilation can be performed as the dystocia is corrected.11 This EXIT (ex utero intrapartum treatment) procedure minimizes the duration of hypoxemia during the birthing process and decreases the chance of the foal dying during a prolonged dystocia (Figure 13.4). It can also be used in an attempt to maintain fetal viability during transport of a dystocia to a referral center. Correct placement of the endotracheal tube in the trachea is clearly essential and can be verified by palpating the cranial esophagus and ensuring a tube cannot be palpated. A capnograph, if available, can be used to non-invasively determine end-tidal carbon dioxide (ETCO2 ). An absent ETCO2 reading indicates esophageal intubation.
of 80–120 beats/min are considered appropriate with complete chest recoil between compressions. The foal should be in lateral recumbency on a hard surface (remove any blankets or bedding from under the foal). There is no difference in effectiveness of chest compressions from either side of the thorax as it is believed that it is the negative intrathoracic pressure generated during recoil of the chest wall after release of the cardiac compression which is important in producing venous return to the heart.2 The operator should kneel beside the foal and place one hand over the other hand to deliver the compressions over the widest point of the foal’s thorax, just caudal to the foal’s triceps. The elbows should remain straight and the operator’s shoulders should be directly above the hands in order to deliver maximum force to the chest compressions.2 This rapid rate results in tiring of the operator, so, ideally, the operator should be replaced every 2–5 minutes in order to maintain adequate force and rate. If an airway is secured, coordination between ventilation and chest compression is not needed. Chest compressions result in no more than 25–30% of normal cardiac output. To maximize cardiac output, half of the cycle should be compression and half relaxation. Chest compressions should not be interrupted for any reason for longer than 10 seconds during an arrest. The mechanism of blood flow during chest compressions is not clear. The thoracic pump theory suggests that application of pressure to the chest wall in a rhythmic manner creates blood flow by increasing intrathoracic pressure which closes the thin-walled intrathoracic veins. When combined with one-way venous valves, retrograde blood flow is minimal. At the same time, the thick-walled arteries tend to remain open, facilitating forward blood flow. When chest compressions are released, the veins reopen and a pressure gradient develops across the thoracic inlet, increasing vascular flow from the extrathoracic to intrathoracic veins and in turn enhancing return of blood to the heart.12 During chest compressions, coronary blood flow is restricted to the diastolic period. The difference between aortic diastolic pressure and right atrial pressure determines myocardial perfusion. Studies in humans and animals have shown that successful return of spontaneous circulation corresponds with maintenance of sufficiently high myocardial perfusion pressures.13, 14
Circulation In foals with an absent heart beat, a non-perfusing cardiac rhythm, heart rates less than 40, or heart rates between 40 and 60 that fail to increase to greater than 60 with poor perfusion after 60–90 seconds of positive pressure ventilation, chest compressions should be initiated without delay. The optimum rate of chest compressions in foals is not known but rates
Drugs Publications supporting the use of specific drugs in the CPCR of neonatal foals are not currently available. All information about drugs and doses is based on clinical experience and extrapolation from other species. In human infants requiring CPCR, it has been suggested that, if drugs are necessary after
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Resuscitation (Foal and Birth) positive pressure ventilation and thoracic compressions, the prognosis is poor.15 The same is likely similar in neonatal foals. Drugs should be considered if the heart rate remains below 40 and is not increasing after 30 seconds of thoracic compressions and adequate ventilation.1, 2 Drugs are usually administered intravenously and the jugular vein is usually identifiable and can be injected relatively easily in even the most severely compromised neonatal foal. A 16-gauge over-the-needle catheter can be quickly placed to secure venous access. Alternatively, intraosseous administration (e.g., medial tibial plateau) can also be used if vascular access is not available.2 Drugs injected into the bone marrow are absorbed almost immediately into the systemic circulation as the marrow cavity does not collapse in the presence of hypovolemia or circulatory shock.2 The pharmacodynamics of intraosseous infused drugs are almost the same as those for intravenous infusion.16 In mature newborn foals, standard intraosseous needles designed for human adults can be used (12-gauge, 2.3-cm intraosseous infusion needle; Sur-Fast Intraosseous Infusion Needle, Cook Critical Care, Bloomington, IN).2 Drugs administered by the transtracheal route are not reliably absorbed. In order to be absorbed, the drug must be delivered beyond the endotracheal tube into the bronchial tree. Drugs should only be administered by this route if no other options are available. Intracardiac injection should be avoided. In addition to the difficulty of injecting blindly into the left ventricle, intracardiac injections carry significant associated hazards. These include laceration of the coronary artery, injection of the drug into the myocardium resulting in fibrillation, cardiac tamponade, or pneumothorax.17 Epinephrine Epinephrine is the most effective drug in CPCR. Epinephrine is an endogenous catecholamine with both α- and β-adrenergic effects, but it is the αadrenergic effects that are most useful in cardiac resuscitation.18 The α-adrenergic effects of this drug cause peripheral vasoconstriction, elevation of arterial blood pressure, and a reduction in perfusion of the splanchnic, renal, and dermal circulation. The βadrenergic effects include an increase in myocardial contractility. As a result, there is an increase in aortic diastolic pressure due to an increase in peripheral arterial constriction and an increase in aortic tone. This increases blood flow through the coronary arteries and myocardium during the relaxation phase of thoracic compressions. The short half-life of the drug necessitates frequent administration (every 3 minutes) until spontaneous circulation returns. The optimal dose of epinephrine is not known in foals. High-
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dose epinephrine in human studies has been associated with a higher defibrillation rate and a lower survival rate.19 Low dose rates (0.01–0.02 mg/kg intravenously) are currently recommended for routine use in neonatal foals rather than high dose rates (0.1 mg/kg intravenously). Anecdotal experience shows that some foals which are refractory to resuscitation efforts may respond after receiving high-dose epinephrine and recover without detectable sequela. However, the high dose should not be used except where other resuscitation efforts have failed. Indications for the use of epinephrine in neonatal foals include asystole, idioventricular rhythms, pulseless electrical activity, and symptomatic bradycardia non-responsive to ventilation with oxygen. Complications associated with the use of epinephrine include post-resuscitation hypertension, tachyarrhythmias, and myocardial necrosis.
Vasopressin The use of vasopressin has gained popularity in human and small animal CPCR. Vasopressin is a nonadrenergic endogenous stress hormone which is a very potent peripheral vasoconstrictor (more marked in cardiac arrest), resulting in centralization of the blood volume. Studies in humans and dogs have shown it to be as effective as epinephrine for treatment of ventricular fibrillation, ventricular tachycardia, and pulseless electrical activity.6 There is some evidence in humans that, when the non-perfusing rhythm is asystole, the use of vasopressin or vasopressin in combination with epinephrine may be more effective than epinephrine alone in returning spontaneous circulation.20, 21 The suggested dose of vasopressin is a total of 0.6 U/kg, given as a single dose or divided.2 Repeated doses are not necessary as the drug effect lasts 10–20 minutes. Post-resuscitation hypertension or arrhythmias are not likely.
Atropine Atropine is a parasympatholytic drug that accelerates sinus or atrial pacemakers and atrioventricular conduction. High vagal tone (except secondary to hypoxia) is an uncommon cause of bradycardia in neonatal foals. Therefore, treatment with atropine is rarely indicated. Instead, the administration of oxygen, positive pressure ventilation, and epinephrine (if required) is more likely to have a positive therapeutic effect. Additionally, if atropine does result in an increase in heart rate, it may increase myocardial oxygen consumption and myocardial workload in the face of inadequate oxygen delivery, exacerbating the hypoxic insult to the foal.1, 2
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Defibrillation Defibrillators (manual or automated) with monophasic or biphasic waveforms are now available in many equine hospitals. Automated external defibrillators are sophisticated computerized devices with voice prompts and have the ability to evaluate the rhythm and recommend defibrillation when required.6 Biphasic waveforms allow less energy and thus may be safer. Although shockable rhythms (ventricular fibrillation and pulseless ventricular tachycardia) are rare initial rhythms in foal cardiac arrest patients, they may develop in approximately onethird of cases during resuscitation therapy. Electrical defibrillation is the only useful therapy to correct these arrhythmias. Defibrillation should not be used to manage asystole or bradyarrhythmias. Ventilation and thoracic compressions should be continued until the moment of defibrillation. Cardiac compressions should be resumed as soon after the shock as possible. Defibrillation results in depolarization of a critical mass of myocardial cells to allow spontaneous organized myocardial depolarizations to resume.2 When defibrillating, the initial charge should be 2 J/kg, delivered as soon after diagnosing a shockable rhythm as possible. Subsequent defibrillations should be 4 J/kg delivered 30–60 seconds after treatment with epinephrine or lidocaine.2 There should be at least 2 minutes of chest compressions between each shock to minimize the interruptions in chest compressions.2, 6 Defibrillation is potentially dangerous to the operator and other personnel. In particular, improper placement of the paddles can cause serious burns and use of alcohol before defibrillation is a fire hazard.
Intravenous fluids Intravenous fluids are not usually required in the initial phase of neonatal foal resuscitation and may be detrimental to coronary and cerebral blood flow. Most foals immediately after birth have normal circulating blood volumes unless there has been excessive loss of blood (e.g., umbilical cord hemorrhage, fractured ribs with hemothorax, or hemoabdomen). In foals that arrest during hospitalization, fluids may also not be required unless severe hypovolemia and dehydration are present. The administration of intravenous fluids will increase venous return and elevate right atrial pressure. As the coronary artery perfusion pressure is dependent on the difference between aortic diastolic pressure and right atrial diastolic pressure, the administration of large volumes of fluid will reduce the aortic to right atrial diastolic pressure gradient. The resultant decreased coronary perfusion pressure and decreased coronary blood flow is detri-
mental.22 Similarly, the major determinant of cerebral blood flow is cerebral perfusion pressure, which is determined by the difference between mean arterial pressure and intracranial pressure. Aggressive intravenous fluid therapy in CPCR increases the intracranial pressure more than the mean arterial pressure, therefore decreasing cerebral perfusion pressure and cerebral blood flow.22 Hyperglycemia and hyperosmolality secondary to rapid glucose infusions during resuscitation have been associated with poor neurological outcomes in humans.23 Fluid therapy, including glucose-containing fluids, is indicated in the postresuscitation phase to maintain cardiac output. The use of sodium bicarbonate during cardiopulmonary arrest remains controversial. There is no strong evidence that sodium bicarbonate improves survival rates in cardiopulmonary resuscitations and there are many contraindications to its use. Bicarbonate compromises cerebral perfusion pressure by reducing systemic vascular resistance. It can create extracellular alkalosis that shifts the oxyhemoglobin saturation curve and inhibits oxygen release. It can also produce hypernatremia that results in hyperosmolality. It can result in the production of excess carbon dioxide, which diffuses into myocardial and cerebral cells, producing intracellular acidosis.24 Thus, the use of sodium bicarbonate is rarely indicated in cardiopulmonary resuscitation of the neonatal foal.
Monitoring during CPCR If a spontaneous heart beat is present, the heart rate and rhythm should be monitored. An electrocardiogram may be useful to correctly identify arrhythmias, but the leads are easily displaced during resuscitations. Although palpation of an arterial pulse has traditionally been used to determine the effectiveness of CPCR, palpation of a central arterial pulse (carotid or femoral artery) can be difficult during the motion caused by thoracic compressions. Thoracic compressions should not be interrupted in order to palpate for pulses. On the other hand, it is useful to monitor the size and position of the pupils with a pen light during resuscitation. When perfusion becomes inadequate, the pupils become fixed and dilated. When perfusion is re-established, lessening of the pupillary dilation may be observed.2 The use of capnography to measure ETCO2 is the most effective measurement of cardiac output currently available during foal resuscitations.2 If there is no cardiac output, no carbon dioxide will be delivered to the lungs and the ETCO2 will be 0 mmHg. If cardiac output is present, carbon dioxide will be successfully delivered to the lung and the ETCO2 will increase. As cardiac output increases with effective
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Resuscitation (Foal and Birth) chest compressions, the ETCO2 will increase to a level of 12–18 mmHg. Notably, the use of some drugs [e.g. epinephrine (0.1–0.2 mg/kg)] can temporarily decrease the ETCO2 concentrations independent of cardiac output. Neither arterial blood gas analysis nor pulse oximetry is useful during CPCR.
7.
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When to stop and post-resuscitation care If spontaneous cardiac output is not re-established within 10–15 minutes of initiation of CPCR, the chance of a successful outcome and a functional foal are minimal. CPCR can be stopped when return of spontaneous circulation occurs. As a guide, this occurs when the foal’s heart rate is greater than 60 beats/min and spontaneous respirations (with a normal rate and effort) have resumed. The heart rate, however, may not reflect the cardiac output in the early post-arrest period. The more severe the hypoxic insult, the longer the time interval before commencement of spontaneous ventilation. At this stage, the endotracheal tube can be removed and humidified intranasal oxygen can be administered at rates of 5–10 L/min. The foal should be kept warm and monitored closely for at least 30 minutes to ensure a stable condition. Fluid therapy and/or inotropic and vasopressor support may be required in the postresuscitation period. Following CPCR resuscitation, the foal is at high risk of repeat cardiopulmonary arrest and neurological, gastrointestinal, cardiopulmonary, and urinary complications over the next 24–48 hours, so, ideally, should be transferred to a neonatal intensive care unit for further monitoring and treatment.
10. 11.
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References 1. Corley KTT, Axon JE. Resuscitation and emergency management for neonatal foals. Vet Clin North Am Equine Pract 2005;21:431–55. 2. Palmer JE. Neonatal foal resuscitation. Vet Clin North Am Equine Pract 2007;23:159–82. 3. International Guidelines for Neonatal Resuscitation. An excerpt from the Guidelines 2000 for Cardiopulmonary Resuscitation and Emergency Cardiovascular Care: International Consensus on Science. Pediatrics 2000;106:1–16. 4. Aufderheide TP, Lurie KG. Death by hyperventilation: a common and life-threatening problem during cardiopulmonary resuscitation. Crit Care Med 2004;32:S345–51. 5. Yannopoulos D, Tang W, Ruossos C, Aufderheide TP, Idris AH, Lurie KG. Reducing ventilation frequency during cardiopulmonary resuscitation in a porcine model of cardiac arrest. Respir Care 2005;50:628–35. 6. Anonymous. American Heart Association Guidelines for Cardiopulmonary Resuscitation and Emergency Cardio-
20.
21.
22.
23.
24.
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vascular Care, part 7.2: Management of Cardiac Arrest. Circulation 2005;112 (Suppl. IV):58–66. Aufderheide TP. The problem with and benefits of ventilations: should our approach be the same in cardiac and respiratory arrest? Curr Opin Crit Care 2006;12:207–12. Dani C, Bertini G, Pezzati M, Pratesi S, Filippi L, Tronchin M, Rubaltelli FF. Brain hemodynamic effects of doxapram in preterm infants. Biol Neonate 2006;89:69–74. Ebihara S, Ogawa H, Sasaki H, Hida W, Kikuchi Y. Doxapram and perception of dyspnea. Chest 2002; 121:1380–1. Roll C, Horsch S. Effect of doxapram on cerebral blood flow velocity in preterm infants. Neuropediatrics 2004;35:126–9. Palmer JE, Wilkins PA. How to use EXIT (ex-utero intrapartum treatment) to rescue foals during dystocia. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners, 2005, pp. 281–3. Guerci AD, Halperin HR, Beyar R, Beattie C, Tsitlik JE, Wurmb EC, Chandra NC, Weisfeldt ML. Aortic diameter and pressure-flow sequence identify mechanism of blood flow during external chest compression in dogs. J Am Coll Cardiol 1989;14:790–8. Paradis NA, Martin GB, Goetting MG, Rosenberg JM, Rivers EP, Appleton TJ, Nowak RM. Simultaneous aortic, jugular bulb and right atrial pressures during cardiopulmonary resuscitation in humans. Insights into mechanisms. Circulation 1989;80:361–8. Niemann JT, Criley JM, Rosborough JP, Niskanen RA, Alferness C. Predictive indices of successful cardiac resuscitation after prolonged arrest and experimental cardiopulmonary resuscitation. Ann Emerg Med 1985;14:521–8. Sims DG, Heal CA, Bartle SM. Use of adrenaline and atropine in neonatal resuscitation. Arch Dis Child Fetal Neonatal ED 1994;70:F3–10. Berg RA. Emergency infusion of catecholamines into bone marrow. Am J Dis Child 1984;138:810–11. Jespersen HF, Granborg J, Hansen U, Torp-Pedersen C, Pedersen A. Feasibility of intracardiac injection of drugs during cardiac arrest. Eur Heart J 1990;11:269–74. Zhong JQ, Dorian P. Epinephrine and vasopressin during cardiopulmonary resuscitation. Resuscitation 2005;66:263– 9. Hilwig RW, Kern KB, Berg RA, Sanders AB, Otto CW, Ewy GA. Catecholamines in cardiac arrest: role of alpha agonists, beta-adrenergic blockers and high-dose epinephrine. Resuscitation 2000;47:203–8. Wenzel V, Krismer AC, Arntz HR, Sitter H, Stadlbauer KH, Lindner KH. A comparison of vasopressin and epinephrine for out-of-hospital cardiopulmonary resuscitation. N Engl J Med 2004;350:105–13. Guyette FX, Guimond GE, Hostler D, Callaway CW. Vasopressin administered with epinephrine is associated with a return of a pulse in out-of-hospital cardiac arrest. Resuscitation 2004;63:277–82. Ditchey RV, Lindenfeld J. Potential adverse effects of volume loading on perfusion of vital organs during closedchest resuscitation. Circulation 1984;69:181–9. Kliegel A, Losert H, Sterz F, Holzer M, Zeiner A, Havel C, Laggner AN. Serial lactate determinations for prediction of outcome after cardiac arrest. Medicine (Baltimore) 2004;83:274–9. Graf H, Leach W, Arieff AI. Evidence for a detrimental effect of bicarbonate therapy in hypoxic lactic acidosis. Science 1985;227:754–6.
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Chapter 14
Shock Niamh M. Collins, Jane E. Axon and Jonathan E. Palmer
The primary function of the cardiovascular system is to maintain an adequate delivery of oxygen to the tissues to ensure the continued production of energy via oxidative phosphorylation. Oxygen delivery is primarily dependent on cardiac function and is therefore sensitive to changes in myocardial contractility, preload, and afterload. Shock is a condition of inadequate delivery of oxygen and nutritive substrates to the tissues owing to cardiovascular instability. Characteristically, shock-induced hypotension results in hypoperfusion of organs and organ dysfunction. Inadequate removal of cellular waste products from the tissues will also occur concurrently. The hypotension may be refractory to fluid resuscitation alone. Shock is often classified on the basis of etiology. The three major etiologies in neonatal foals are hypovolemia, cardiac disease, and sepsis/systemic inflammatory response syndrome. Minor etiologies would include severe anemia, anaphylaxis, and neurogenic shock (Table 14.1). Hypovolemic shock can result from numerous etiologies in the neonatal foal, including reduced fluid intake, hemorrhage, or severe fluid and/or plasma loss or sequestration. Hypovolemia will decrease venous return and myocardial preload. Because of the reduction in myocardial preload, there is a resultant decrease in stroke volume, cardiac output, and blood pressure. Cardiogenic shock can occur with any cardiac abnormality that decreases the ability of the heart to pump blood; thereby producing a marked reduction in cardiac output and hypotension. This would include conditions such as myocarditis, cardiac tamponade secondary to pericardial effusion, severe congenital heart defects, and arrhythmias. Systemic inflammatory response syndrome (SIRS), which represents a common phase of the inflammatory response characterized by malignant global activation of multiple proinflammatory pathways, is defined by the presence of two or more of the following abnormalities: fever or hypothermia (rectal temper-
ature greater than 39.2◦ C or < 37.2◦ C); tachycardia (heart rate greater than 120 beats/min); tachypnea (respiratory rate greater than 30 breaths/min); leukocytosis (peripheral white blood cell count > 12.5 × 109 /l)/leukopenia (peripheral white blood cell count < 4 × 109 /l), or greater than 10% immature (‘band’) neutrophils.1–4 The SIRS is a non-specific innate inflammatory response to an array of severe clinical insults such as bacterial infections, viral infections, endotoxemia, trauma, burns, ischemia, and hypoxia.5 Many mediators are involved in SIRS,2 and can result in the development of shock. SIRS caused by infection is termed sepsis. SIRS-induced shock due to sepsis and perinatal asphyxial syndrome would be the two most common causes of hemodynamic disturbances in the neonatal foal. Anaphylactic shock is uncommon in neonatal foals but can be due to drug administration or the administration of blood or plasma. The release of histamine from basophils in the blood and mast cells in the tissues can cause diffuse peripheral vasodilation, increased permeability of capillaries, and systemic hypotension. Severe brain or spinal injury can rarely cause neurogenic shock and loss of vasomotor tone, resulting in the development of severe hypotension. The neonatal foal has a larger reserve of interstitial fluid than the adult horse, allowing it to respond more rapidly and completely than the adult horse to hypovolemic challenges by rapid mobilization of this reserve. However, once these reserves are expended, the foal is at increased risk of developing hypovolemic shock. Additionally, it has previously been noted that neonatal foals do not consistently mount the typical mature physiological response to hypotension (i.e., an increase in heart rate and cardiac output) commonly observed in adult horses.6, 7 Once distress is detected in the neonatal foal, global homeostasis is likely severely perturbed. Additionally, the failure to consistently exhibit classic clinical signs of shock such as tachycardia can result in a delay of identification of shock in the earlier stages, resulting
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 14.1 Major and minor causes of shock in neonatal foals Major Etiologies of Shock Hypovolemia Water and electrolyte losses
Severe enteritis/diarrhea/peritonitis Polyuria due to renal disease Uroperitoneum Simple obstructive or strangulating gastrointestinal lesions Inadequate milk intake e.g. due to HIS, sepsis Severe burns
Severe hemorrhage
Trauma Hemoabdomen Hemothorax Bleeding from umbilical remnants Gastrointestinal bleeding Congenital hemostatic defects Severe disseminated intravascular coagulation
Plasma losses (third spacing)
Peritonitis Pleural effusion
Cardiac Disease Congenital heart disease Cardiac tamponade Myocarditis
Pericarditis Hemopericardium Hypoxic or ischemic damage Sepsis
Arrhythmias Sepsis/SIRS Sepsis
HIS Severe trauma Endotoxemia
Bacterial infections Viral infections Fungal infections
Strangulating intestinal lesions Peritonitis
Severe burns Severe Anemia Neonatal isoerythrolysis Severe hemorrhage Minor Etiologies of Shock Allergy or Anaphylaxis Administration of drugs, plasma or blood transfusion Snake-bite Neurogenic Shock Neurological injury (brain or spinal cord) Miscellaneous Tension pneumothorax Pulmonary embolism
in a delay in instituting treatment and increased morbidity and mortality.6
Pathophysiology of the clinical signs of shock Cardiac output falls as a result of a combination of reduced blood volume, decreased venous pressure, decreased cardiac preload, and/or a reduction
in myocardial contractility. Stretch receptors in the great vessels and atria detect the reduced preload and stimulate sympathetic outflow from the vasomotor center in the brain. Sympathetic venoconstriction reduces the capacitance of the peripheral circulation, resulting in an increased venous return which initially increases cardiac output. Arterial vasoconstriction initially preserves the blood pressure through an
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Foal: Problems of the Immediate Postpartum Period
increase in resistance alone, but, as blood volume loss continues, perfusion is diverted from non-essential areas such as the skin and kidney. As a result, the peripheral extremities become cool, the capillary refill time lengthens, and oliguria develops. Blood is also diverted from the splanchnic circulation, resulting in ileus with reduced gut sounds and/or mild abdominal distension. Moderate to severe gastrointestinal distension due to ileus should be differentiated from the possibility of uroperitoneum, intestinal obstruction, or gastrointestinal perforation which may also result in shock. In cases with SIRS and/or sepsis, widespread endothelial activation occurs. Increased vascular permeability results in extravasation of fluids into the interstitial space; this further contributes to the hypotension and hypovolemia. Accumulation of fibrin and aggregation of platelets and red blood cells secondary to activation of the coagulation system results in occlusion of the vasculature. Venous vasodilation in large-capacitance vessels promotes the decreased venous return to the heart. Loss of peripheral vasomotor tone may also occur in shock. Sepsis is the most frequent cause of vasodilatory shock, although other conditions such as hemorrhagic shock, cardiogenic shock, and anaphylaxis can cause vasodilatory shock. The underlying mechanism for the loss of peripheral vasomotor tone is not fully understood. Present evidence suggests that at least three pathways are involved: activation of ATP-sensitive potassium channels, increased synthesis of nitric oxide, and vasopressin deficiency.8
Clinical signs and diagnostic findings Although some foals may still be standing while in shock, most foals will usually be collapsed and recumbent owing to hypotension and a progressive reduction in cerebral perfusion. The foals are generally depressed, not nursing, and poorly responsive. Changes in mentation may be apparent only 2–3 hours after the onset of cardiovascular shock.9 Clinical signs of hypoperfusion include cold extremities as blood is shunted centrally. In early compensated shock, the more distal areas are colder than the proximal areas with ice-cold hooves, cold lower legs, and cool proximal legs, nose, and ears. There is a significant core to periphery temperature gradient. As shock progresses, this peripheral to central gradient becomes less distinct. Other physical examination findings that help assess perfusion are pulse quality, arterial tone, and arterial fill. Careful assessment of pulse quality can help determine pulse pressure (the difference between systolic and diastolic pressures) (Figure 14.1). Arterial tone is assessed by the amount of finger pres-
Figure 14.1 Palpating the pulse in the metatarsal artery. The temperature of the extremities can be felt at the same time.
sure required to feel the pulse and the amount required to eliminate the pulse. In foals with very poor arterial tone, a pulse can be felt with only a very light touch. Arterial fill is assessed by feeling the size of the artery’s lumen as finger pressure is applied. Arterial fill relates to the blood volume on the arterial side of the circulation. In foals with shock, pulse pressure, arterial tone, and arterial fill will likely be significantly reduced, and, in severely shocked foals, palpation of a peripheral pulse may not be possible. The association between abnormal mucous membrane findings and specific hemodynamic alterations are not as strong in the neonatal foal as in the adult horse. Cyanotic mucous membranes with a prolonged capillary refill time, symptomatic of low cardiac output and poor peripheral perfusion, are indicative of a poor prognosis. Injected mucous membranes have been associated with peripheral vasodilation. Pale mucous membranes may indicate peripheral vasoconstriction or anemia (Figure 14.2).10 Severe jaundice with anemia in the neonate would be very highly suggestive of neonatal isoerythrolysis.11 Prolonged capillary refill time suggests hypovolemia. Evaluation of the inner surface of the pinna or mucous membranes may reveal petechial or ecchymotic hemorrhages suggestive of altered coagulation function and/or disseminated intravascular coagulation. Foals with sepsis may have altered endothelial permeability with resultant edema formation, particularly if intravenous fluids have been administered. Tachycardia (heart rate > 120 beats/min) may be identified in foals in shock but frequently the heart rate may be within the normal range (70–103 beats/min) despite significant hypoperfusion.12 Bradycardia (heart rate < 60 beats/min) is usually associated with hypoxia13 and is likely a failure of the baroreceptor response secondary to autonomic failure.7 While flow murmurs are commonly identified in the neonatal foal, persistently loud murmurs with wide radiation and those with precordial thrills warrant further diagnostic investigation to rule
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Figure 14.3 Urine collection system used for monitoring urine output. Specific urine collection bags can be used (as demonstrated here) or empty fluid bags.
Figure 14.2 Pale mucous membranes seen with acute hemorrhagic shock, as seen with this foal which had a hemothorax.
out the presence of congenital heart malformations. Large-volume pericardial effusions may result in reduced cardiac output and shock due to a reduction in diastolic filling. Muffled heart sounds may be audible on cardiac auscultation in these cases. Transient cardiac arrhythmias are not unusual during the first hours after birth. However, foals with persistently elevated or persistently irregular heart rates should be investigated further with an electrocardiogram. Persistent atrial and ventricular arrhythmias may be due to myocardial damage secondary to hypoxia, sepsis, or trauma (e.g., contusions or lacerations secondary to fractured ribs). Tachypnea (respiratory rate > 40 breaths/min) may be present in response to hypoxemia and also as a compensatory respiratory response to metabolic acidosis. Rectal temperature may be hyperthermic (rectal temperature > 39.2◦ C), normothermic, or hypothermic (rectal temperature < 37.2◦ C). As mentioned previously, shock from any cause is likely to result in ileus, which can result in mild to moderate abdominal distension. Diarrhea staining of the tail and hind limbs may be evident with foals with enteritis. Palpation of the thorax may reveal fractured ribs with or without associated plaques of subcutaneous edema overlying the ribs or ventral thorax, especially behind the elbows. The foal may flinch when the ribs are palpated, the fractured ends may be directly palpable, and audible or palpable crepitus may be present. Displacement of the fracture ends may result in hemoabdomen, hemothorax, hemoperi-
cardium, and laceration of intercostal vessels. Many cases of rib fractures sustained during parturition are overlooked unless a careful physical examination is performed. Rarely, tension pneumothorax in the foal will result in increased resistance to venous return, resulting in impairment to cardiac filing, a reduction in cardiac output, and ensuing shock. In these foals, auscultation of the thorax will reveal lack of air movement dorsally. Urine output is a useful indirect indicator of endorgan perfusion and fluid balance in neonatal foals (Figure 14.3). It is typically markedly reduced in foals with shock. Foals normally first urinate when they are 12–18 hours of age. Delayed urination may be caused by many factors, including failure of fetal–neonatal renal transition, dehydration, hypovolemia, or, rarely, anuric renal failure.7 Indirect measurement of blood pressure by oscillometry can be extremely useful in neonatal foals presenting in shock. Only in rare cases with severe hypotension is direct blood pressure measurement necessary. Arterial blood pressure is the product of cardiac output (flow) and the resistance to flow in the vasculature (systemic vascular resistance). Measurement of blood pressure forms a useful way of identifying some cardiovascular disturbances, and serial blood pressure monitoring can be used to guide treatment.1 Studies in humans have shown that mean arterial pressure (rather than systolic or diastolic arterial pressure) is the true driving pressure for systemic blood flow and organ perfusion and, therefore, it has been recommended that the mean arterial blood pressure is used to guide therapeutic decisions.14 The mean (and diastolic) arterial pressure obtained by indirect oscillometry has been shown to have good agreement with that obtained by direct blood pressure measurement in neonatal foals.15, 16 Indirect oscillometry is performed most commonly over the middle coccygeal artery at the base of the
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tail. Other sites of cuff placement that have been recommended for indirect blood pressure measurement include the dorsal metatarsal or median arteries,6 but suitable sites for cuff placement depend on the monitor used.16 The use of appropriately sized cuff bladder widths is important to avoid either underestimation (cuff bladder width too wide) or overestimation (cuff bladder width too narrow) of indirect blood pressure measurements. Bladder width to tail girth ratios of 1:1.9 to 1:2.8 have been suggested as appropriate for neonatal foals.15 The same size cuff should be consistently used to minimize variation. The blood pressure should be measured during a quiet or sleep state. Values obtained during activity, and especially when foals are standing or heavily restrained, likely have limited relevance to perfusion. Maintenance of mean arterial pressures greater than 60–65 mmHg are believed to be important to maintain particularly cerebral, renal, and coronary blood flow in man.17 Similar desired values have been suggested for neonatal foals and therefore the institution of cardiovascular support has been advised when the mean arterial pressure falls below 60–65 mmHg.18 However, some foals will have good tissue perfusion at mean arterial pressures < 60 mmHg; thus, the mean arterial blood pressure should only be used as a guide and should be interpreted in conjunction with physical examination findings such as peripheral extremity temperature, pulse quality, arterial fill, and tone. The presence of oliguria, metabolic acidosis, or other signs of organ hypoperfusion will support therapeutic intervention. Human preterm infants have been shown to have mean arterial pressures below those of fullterm babies because of the presence of low-resistance vessels.19 The same may be true of some premature neonatal foals.
Therapeutic management of shock in foals Early goal-directed therapy is essential in foals in shock. Shock results in inadequate perfusion of organs leading to hypoxic (potentially irreversible) damage. The longer the shock state persists, the more difficult it becomes to treat effectively. The treatment of shock is aimed at ensuring perfusion of critical vascular beds (coronary, cerebral, hepatic, and renal) and preventing or correcting metabolic abnormalities arising from cellular hypoperfusion. Foals with clinical signs of shock should be transported to a hospital facility without delay (see Chapter 24). This often involves transporting the foal ahead of the mare and bringing the mare subsequently once the foal has been stabilized. Keeping the foal warm during the journey and administering
intranasal oxygen is useful. If a delay in referral is unavoidable, intranasal oxygen insufflation and intravenous fluid therapy should be instituted. Initially on admission to the hospital, attempts are made to stabilize the foal’s cardiovascular system and respiratory system, and then a thorough physical examination is performed to determine the underlying etiology of the shock. The most important goal of early therapy is assuring tissue oxygen delivery by maximizing pulmonary loading of hemoglobin, guaranteeing sufficient blood oxygen content, and returning lung and tissue perfusion to normal. Intranasal oxygen is initially administered at rates of 5–10 L/min to maximize blood oxygen loading in order to compensate for potential ventilation–perfusion mismatching and/or hypoventilation. Intravenous access is secured by placing an intravenous catheter, most commonly in a jugular vein. Saphenous, cephalic, and lateral thoracic veins can also be used. A 16-gauge 8-inch (20 cm) over-thewire catheter (Arrow Central Venous Catheterization Set, Arrow International, Reading, PA) is used most commonly as it is less likely to kink at the catheter insertion site, prolonging its duration of usefulness. For initial fluid resuscitation, a 16-gauge over-the-needle catheter can also be used (Milacath-Extended Use, Mila International, Erlanger, KY). Heat loss should be minimized by hand-drying the foal with towels, careful bandaging of limbs, and covering the foal with rugs. Active external warming will cause cutaneous vasodilation, which can lead to further hypoperfusion. It should be avoided until the foal’s condition is stable. In order to increase ventricular preload, stroke volume, and, therefore, cardiac output in hypovolemic foals, intravenous fluids (crystalloids and colloids) are commonly administered. If hypotension persists despite aggressive intravenous fluid therapy, antibiotics, blood products, and other treatments appropriate for the disease process, the use of vasoactive agents should be considered. The ultimate goal is to restore tissue perfusion and oxygenation. Crystalloid fluids Hypovolemia, as occurs in septic shock, high-volume diarrhea, or acute hemorrhage, demands rapid volume repletion of the extracellular fluid. Fluids should ideally be warmed and can be administered via a pressurized cuff to increase the administration rate (Figure 14.4). Consideration should be given to the bodyweight of the foal. Pony cross foals may only be 25–30 kg at birth while draft foals may weigh in excess of 65–70 kg. Ideally, the foal’s weight is recorded on arrival to the intensive care unit to allow accurate determination of a fluid plan.
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Figure 14.4 A pressure bag is used for rapid fluid administration.
Balanced ionic solutions such as Hartmann’s, lactated Ringer’s or Normosol-R solution are the most suitable fluid for administration in shock as they have a strong ion balance approximating the foal’s plasma. Since large volumes of fluid will be administered, there will be a tendency for the plasma osmolarity and strong ion balance to move toward that of the fluids being administered. Use of large volumes of saline (0.9% NaCl) with no strong ion difference has a significant acidifying effect and should be avoided. Rapid administration of dextrose-containing fluids can result in electrolyte derangements, hyperglycemia, and osmotic diuresis. Balanced electrolyte solutions may not be the most suitable choice in foals with hyperkalemia due to uroperitoneum or renal failure. In such cases, a combination of 0.9% NaCl and 5–10% dextrose is recommended. Hypertonic saline is unsuitable for fluid resuscitation in the neonatal foal. Its use can cause deleterious effects because of the high sodium concentration and subsequent rapid change in plasma osmolarity. This may result in central nervous system (CNS) damage. Administration of fluid boluses of 20 mL/kg over 10–20 minutes with re-evaluation of perfusion after each bolus is most efficient.20 Administration of a bolus of fluid in foals will increase venous return; therefore, augmenting the ventricular end-diastolic volume. The myocardium will be stretched by this volume; and, as in Starling’s isolated heart tissue
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model, will respond by a greater force of contraction and increased cardiac output. Evidence of successful therapy includes improved pulse quality, warm legs, return of borborygmi, urine production, improved mental status, and improved blood pressure. Repeat fluid boluses are usually required. If greater than 60–80 mL/kg is required, inopressor therapy may be necessary. Monitoring perfusion status after each 20mL/kg fluid bolus helps to limit the amount of fluids given to the amount required to satisfy the immediate needs but avoiding excessive fluid delivery. Rapid redistribution of crystalloid fluids from the intravascular space is a common problem of fluid therapy in the neonatal foal. Fluid administration in the neonatal foal with SIRS can be even more challenging. These foals may have additional complicating factors, such as increased endothelial permeability and reduced oncotic pressure with consequent increased risk of interstitial edema formation. Interstitial edema formation is undesirable as it may interfere with normal organ function and perfusion. Furthermore, neonatal foals are prone to sodium overload and have difficulty in maintaining normal sodium homeostasis, especially when treated with sodium-based crystalloids such as Hartmann’s or lactated Ringer’s solution. Sodium handling is further confused when the neonate suffers a hypoxic ischemic event. These foals often have renal tubular damage and may develop significant sodium wasting. Although uncommon, pulmonary edema is also a possibility with overzealous fluid administration. This may be recognized by increasing respiratory effort and the presence of crackles on auscultation of the lungs. Ideally, a fluid plan should be used that minimizes the risk of these adverse complications. Colloids There is no compelling evidence that shows a clear advantage of using crystalloids or colloids when treating shock.21 There is also debate as to whether synthetic colloids such as modified gelatins (Gelofu R R sine ) or hydroxyethyl starches (Hetastarch and R Pentastarch ) have a role in volume resuscitation.22 Apart from conflicting views in human medical fields, one major concern is the increased movement of the colloid molecules through damaged endothelial barriers into the interstitium. If this occurs, the synthetic colloids may exacerbate the retention of fluids by holding fluid in the interstitial space and may not be cleared as rapidly as natural colloids. In the acute stages of shock in foals, the authors do not routinely use synthetic colloids. On the other hand, commercially available hyperimmune plasma is a very useful colloid which is
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routinely administered in foals presenting with shock. In addition to improving colloidal oncotic pressure, it also provides immunoglobulins for immunotherapy, clotting factors, complement, fibronectin, antithrombin, and other beneficial components. The plasma should be administered using an intravenous blood administration set containing filtration devices to remove fibrin and particulate debris and the patient should be carefully monitored for transfusion reactions (e.g., tachycardia, tachypnea, shivering, restlessness, urticarial wheals, eyelid, or muzzle edema). While plasma may be administered safely at more rapid rates in healthy foals, in critically ill neonatal foals rates of 20 mL/kg/h are more appropriate. Critically ill neonatal foals receiving 2–3 liters of plasma should be monitored for urine output and circulatory overload.
Blood Foals, similar to adult horses, are tolerant of a low packed cell volume if the loss has been incremental and gradual, particularly if they have a normal circulating blood volume. Blood loss is more often acute and catastrophic in nature in neonatal foals, and affected foals may also have an additional serious illness such as sepsis or perinatal asphyxia syndrome. Packed cell volume less than 14% and hemoglobin concentration less than 5 mg/dL can be used as broad indicators for transfusion.23 Elevations in blood lactate concentration or oxygen extraction ratios are also useful indicators of impaired tissue oxygenation,24 although the etiopathogenesis of elevations in blood lactate concentrations are not restricted to anaerobic metabolism in neonatal foals. The oxygen extraction ratio is calculated as (Sa O2 – Sv O2 /Sa O2 ) × 100; where Sa O2 and Sv O2 represent arterial and mixed (or central) venous oxygen saturation respectively. The technique requires placement of a central venous catheter. When oxygen delivery is reduced, as occurs in severe anemia, oxygen extraction by the tissues increases in an attempt to maintain constant uptake of oxygen by tissues. A normal oxygen extraction ratio in five healthy foals aged 30–46 hours was found to be approximately 18%.6 Oxygen extraction ratios ≥ 50% have been suggested for use as a transfusion trigger in human medicine.25 Similar values likely apply to foals. In foals with neonatal isoerythrolysis (Chapter 35) that present in shock due to acute severe anemia and hypoxemia, the use of cross-matched whole blood from a healthy gelding or washed red blood cells from the mare can be life-saving. In an emergency situation, cross-matching for compatibility may not always be possible. Administration of a hemoglobinbased oxygen-carrying solution such as oxyglobin
(10–20 mL/kg; Biopure, Cambridge, MA) may be considered if a transfusion is not immediately available. Oxyglobin consists of chemically stabilized bovine hemoglobin in a saline solution. The product has a long shelf life (up to 3 years) and crossmatching is not necessary. The bovine hemoglobin has a short half-life in the recipient (approximately 18–26 hours), thus severely anemic foals will still require a blood transfusion as the bone marrow will not have been allowed adequate time to respond. In one recent study,11 administration of oxyglobin was associated with non-survival in foals with neonatal isoerythrolysis, although the number of foals treated with this product in the case series was small (seven of 72 foals). Inotropes and vasopressors If there is inadequate return of perfusion after fluid boluses or if the patient is deteriorating despite the fluid loading, inotropic or vasopressor therapy should be initiated. Inotropes increase myocardial contractility, thereby increasing cardiac stroke volume. Vasopressor agents increase vascular smooth muscle tone principally in the arterioles. There is currently a paucity of clinical studies reporting the use of inotropes and vasopressors in the treatment of shock in the critically ill neonatal foal; therefore, the choice of the most appropriate inotrope or vasopressor in each patient can be difficult. The most commonly used vasoactive agents in foals are dobutamine, norepinephrine, and vasopressin. With inotropic and vasopressor therapy, the optimal drug or combination of drugs, optimal dose, and duration of therapy must be tailored to each individual foal by monitoring for signs of improved perfusion during continuous rate infusions and adjusting the drugs and doses accordingly. Owing to variation in many factors, including plasma half-life of these drugs, receptor density, receptor affinity, receptor reactivity, and the effect of plasma pH on all of these factors, the response to inotropic and vasopressor therapy cannot be easily predicted. The lowest dose of each drug should be instituted initially and the dose can be increased as necessary depending on the observed response. Typically, refractory systemic hypotension is managed initially with an adrenergic agonist (dobutamine). If the hypotension does not improve, vasopressor therapy is considered. The goal is to use the lowest possible dose for the shortest period of time. Because of the short half-life, the effect of new doses is readily evident within 10–15 minutes. The individual may change with time depending on many confounding factors, so the delivered dose may also need to be adjusted. If the foal is dependent on inotropic or vasopressor support or has been on this support for
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Shock some time, the drugs should be weaned slowly to determine that the foal is capable of sustaining a normal blood pressure. While the use of inotropes and vasopressors can prove invaluable treatment in the neonatal foal with shock, the use of these agents in foals is not without hazard. It is vital that adequate volumes of intravenous fluids have been administered to restore the foal’s circulating blood volume prior to instituting inotropic and vasopressor therapy. Vasopressors and inotropes will not be effective without adequate volume resuscitation. Dobutamine may cause tachycardia and increase myocardial oxygen consumption and can damage myocardial cells, particularly if used in high doses or with prolonged use.26 It can also occasionally cause arrhythmias.7 Vasopressors, such as norepinephrine and vasopressin, can increase mean arterial pressure by increasing systemic vascular resistance, but may increase cardiac afterload, augmenting myocardial workload. They may also decrease cardiac output and oxygen delivery by baroreceptor-mediated reflex reduction in cardiac output.9 Whenever the cardiovascular system is supported by pharmacological intervention, there may be both an increase in perfusion and, simultaneously, an increase in the maldistribution of that perfusion. There is a delicate balance between improved perfusion and exaggerated malperfusion.27 When instituting cardiovascular support, the goal is to return perfusion to minimally acceptable levels and not to try to achieve normal or supranormal perfusion as the latter may have catastrophic consequences.7 The short half-life of all these drugs means that the drugs can be titrated to give a therapeutic effect, but also means the drugs must be administered via a constant rate infusion. Inotropes and vasopressors should be administered via an electronic infusion pump to ensure accurate dosing.10 All inotropes and vasopressors need to be diluted prior to administration. Dobutamine can be diluted in isotonic saline, 5% dextrose or Hartmann’s solution; norepinephrine is diluted in 5% dextrose, and vasopressin is diluted in isotonic saline or 5% dextrose.9 Dobutamine is a synthetic catecholamine that is used extensively to support cardiovascular function in critically ill foals18 and is often the initial agent of choice when fluid therapy has failed to restore cardiovascular function, particularly in cases where more advanced cardiovascular monitoring such as cardiac output monitoring is not available. Dobutamine is a β 1 -adrenergic agonist with weak β 2 and α-adrenergic affinity that acts predominantly as a positive inotrope.18, 28 Therefore, the administration of dobutamine may result in increased cardiac contractility, cardiac output, heart rate, and mean arterial pressure.10 Initial starting dose is usually
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2 µg/kg/min and doses up to 20 µg/kg/min may be required. High doses may lead to tachycardia and occasionally arrhythmias.7 In foals with SIRS and shock, part of the pathophysiology of peripheral vasodilation can be impaired responsiveness to circulating catecholamines. 10 Vasopressors may improve mean arterial pressure by increasing vascular smooth muscle tone, principally in the arterioles. This increases the pressure gradient across a tissue capillary bed, allowing perfusion.18 Norepinephrine and vasopressin are the two most commonly used vasopressors in critically ill neonatal foals. Norepinephrine is a potent vasopressor with α and to a lesser extent β 1 receptor agonist activity.28, 29 A starting dose of 0.1 µg/kg/min with further titration to an effective dose has been suggested.18 The addition of low-dose dobutamine (5 µg/kg/min) to norepinephrine has been demonstrated to result in better splanchnic perfusion in human septic patients, possibly because of a vasodilating effect of dobutamine on splanchnic mucosal microcirculation.30 Norepinephrine used in combination with low-dose dobutamine (5 µg/kg/min) was found to be effective in increasing mean arterial pressures in six of seven critically ill foals that were refractory to treatment with dobutamine and/or dopamine.1 All foals in this study showed an increase in urine output coincident with the start of the norepinephrine infusion. Three of the seven foals survived to compete as racehorses.1 Cardiac output monitoring was not performed in this study. Studies in pharmacologically induced hypotensive, anesthetized healthy neonatal foals have found dobutamine, norepinephrine, and vasopressin to increase mean arterial pressure.28 However, only dobutamine and norepinephrine improved cardiac output. Craig et al.31 reported an increase in mean arterial pressure but a decrease in cardiac output in anaesthetized, normotensive, healthy neonatal foals receiving a combined infusion of norepinephrine and dobutamine. In another study in conscious, healthy, normotensive neonatal foals,29 a norepinephrine infusion was found to increase mean arterial blood pressure and systemic vascular resistance. A combined norepinephrine and dobutamine infusion resulted in higher arterial pressure than a norepinephrine infusion alone. Both infusions resulted in a reduction in cardiac output. The pressor effect of vasopressin is a result of its action on peripheral V1a receptors found in the vascular smooth muscle; this causes intense nonadrenergic vasoconstriction. Vasopressin preferentially constricts arterioles in extracerebral tissues, with relatively less constriction in renal and coronary blood vessels, and may dilate the cerebral vasculature improving cerebral perfusion. It also acts on
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V2 receptors in the collecting tubule of the nephron to cause water resorption.10, 28 A starting dose of 0.25-0.5 mU/kg/min has been suggested for foals.32 Hypotension is a potent stimulus for vasopressin release. In septic shock in human patients, plasma vasopressin levels are initially elevated in order to maintain adequate blood pressure and organ perfusion. However, vasoactive levels of vasopressin become rapidly depleted with continued stimulation.8 Vasopressin deficiency may contribute to the pathogenesis of irreversible shock. Autonomic dysfunction with the loss of baroreceptor function, decreased endogenous vasopressin production, and exhausted vasopressin stores may account for this observed dysfunction of the vasopressinergic system in critically ill patients.33 Vasopressin in low, physiological doses can restore vascular tone in refractory vasodilatory shock states due to closure of ATP-gated potassium channels, inhibitory action on nitric oxide, and potentiation of an endogenous and exogenous vasoconstrictors.34 Vasopressin increased mean arterial pressure in healthy foals anaesthetized with isoflurane.28 There are currently no studies describing its use in critically ill neonatal foals. Anecdotally, the authors have observed a consistent clinically apparent positive effect of vasopressin on perfusion and have begun to use vasopressin as a first-line treatment for neonatal foals with hypotension refractory to fluid resuscitation alone. Terlipressin is a synthetic long-acting analog of vasopressin. Terlipressin has a higher affinity for vascular receptors and a longer duration of action than vasopressin. Terlipressin in intermittent intravenous dosing has been used for the treatment of hypotension in shock states. It does have powerful vasoconstrictive effects that may be of concern in relation to ischemia. It is currently being used as a rescue therapy in human patients with refractory septic shock.35 There is no current information on its use in the veterinary patient. Continuous evaluation of response to therapy is imperative in the neonatal foal. It is important to assess inotropes and vasopressors in light of multiple indicators of perfusion rather than relying on mean blood pressure or cardiac output measurements alone to direct pharmacological therapy. A critical review should be urgently performed of a therapeutic agent that results in a reduction in urine production or an increase in serum lactate.18 Other therapeutic interventions for the foal in shock Intravenous broad-spectrum bactericidal antimicrobial therapy is indicated in critically ill neonatal foals in shock. Antimicrobial selections will depend on the
clinician’s experience and geographic location; however, combinations such as penicillin and an aminoglycoside (gentamicin, amikacin) or third- or fourthgeneration cephalosporin have been recommended. The aminoglycosides can cause renal tubular damage and should be used with caution in the azotemic foal. Therapeutic drug monitoring is recommended when using these antimicrobials. Fluoroquinolones, such as enrofloxacin, have been associated with arthropathy in foals, thus are not recommended as a first choice antimicrobial in foals.36 Use of polymyxin B (1000–5000 iu/kg i.v.) in foals with septic shock to bind endotoxin in the bloodstream prior to activation of cellular inflammatory cascades may be useful. Durando et al.37 showed that pretreatment with polymyxin B attenuated both the clinical signs and the activities of inflammatory mediators (specifically tumor necrosis factor and interleukin 6) in foals with experimentally induced endotoxemia. Clinical studies scientifically evaluating the use of polymyxin B in foals with endotoxemia are currently not available. Polymyxin B has nephrotoxic potential and should be used with caution in the neonatal foal with shock and renal hypoperfusion.38 Positive pressure ventilation may be required in foals with severe hypoxemia that does not respond to oxygen therapy. Such cases of severe hypoxemia are almost always due to pulmonary parenchymal disease such as pneumonia, pulmonary edema, or pulmonary hemorrhage. Positive pressure ventilation can be effective as it can re-expand previously collapsed alveoli and better ventilate poorly functioning alveoli to improve the efficiency of the lung. In addition, it takes over the work of breathing, preventing patient exhaustion and subsequent cardiopulmonary arrest. Nutritional support, in the form of exogenous glucose supplementation and/or total parenteral nutrition, is also indicated. Owing to the diversion of blood from the splanchnic vascular beds in shock and the high risk of ileus, enteral therapy is rarely suitable in the early stages of treatment. Although use of physiological doses of corticosteroids in shock may improve the vascular smooth muscle response to endogenous catecholamines, the benefits of the use of corticosteroids in severe sepsis and septic shock remain controversial in human medicine.39 Low-dose corticosteroid therapy may have a beneficial effect on the short-term mortality rate in human patients with septic shock,39 but has not been shown to alter longer term (> 28 days) mortality rates.40 Recent studies in foals suggest that hypothalamic–pituitary–adrenal (HPA) axis dysfunction occurs in septic foals and these foals are more likely to have shock and multiple organ failure.41 Further studies into the use of corticosteroid replacement
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Shock therapy in septic foals with HPA axis dysfunction will be beneficial to evaluate its use as a treatment in foals with shock.
Monitoring of response to therapy in neonatal shock Routine monitoring for the shocked neonatal foal should include repeated physical examinations (with particular attention placed on mentation, quality of peripheral pulses, temperature of extremities), monitoring of urine output, indirect arterial blood pressure, blood lactate concentrations, and arterial blood gas analyses. For foals that fail to respond to therapy, more advanced techniques such as cardiac output monitoring, central venous pressure monitoring, end-tidal carbon dioxide monitoring, and (rarely) pulmonary arterial occlusion pressures may be employed. The goal of hemodynamic monitoring is to ensure adequate oxygen delivery to tissues in an attempt to maintain normal organ function. Early recognition of metabolic or physiological derangements allows prompt medical intervention and may contribute to increased foal survival.
Complications With appropriate and aggressive therapy, cardiac output, organ perfusion, and tissue oxygenation and nutrition can be restored and the foal can make a recovery. Shock cannot always be simply corrected by treating the etiology. If shock has resulted in activation of inflammatory pathways, release of cytokines, and organ injury, these processes may proceed despite aggressive treatment and restoration of global perfusion. Tissue injury may be exacerbated by reperfusion injury when blood flow is returned. Pulmonary dysfunction can occur owing to a combination of mechanisms such as persistent hypoxia, increased pulmonary vascular permeability, pulmonary edema, microthrombi formation, pulmonary epithelial injury, and decreased surfactant production. Acute renal failure may be evidenced by oliguria and azotemia; this can be due to the altered renal blood flow in hypotension resulting in hypoxic damage and microthrombi formation leading to renal infarcts. Gastrointestinal dysfunction can result in ileus, loss of mucosal barrier function with bacterial translocation, and endotoxin absorption, further perpetuating the severity of the shock. Liver failure can also occur with depression of its many metabolic and detoxification functions. Dysfunction of the central nervous system can be initially manifested as depression but can progress to septic encephalopathy with extensive neuronal injury.
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Widespread activation of the coagulation system can result in disseminated intravascular coagulation. In all forms of shock, myocardial and vasomotor failure can take place, leading to cardiovascular failure unless judicious and timely therapy is offered. If organ perfusion and function cannot be restored, death may ensue.
References 1. Corley KT, McKenzie HC, Amoroso LM, Furr MO. Initial experience with norepinephrine infusion in hypotensive critically ill foals. J Vet Emerg Crit Care 2000;10: 267–76. 2. McKenzie HE, Furr MO. Equine neonatal sepsis: the pathophysiology of severe inflammation and infection. Comp Cont Educ Pract Vet 2001;23:661–72. 3. Furr M. Systemic inflammatory response syndrome, sepsis and antimicrobial therapy. Clin Tech Equine Pract 2003;2:3–8. 4. Corley KT, Donaldson LL, Furr MO. Arterial lactate concentration, hospital survival, sepsis and SIRS in critically ill neonatal foals. Equine Vet J 2005;37:53–9. 5. Bone RC, Balk RA, Cerra FB, Dellinger RP, Fein AM, Knaus WA, Schein RM, Sibbald WJ. Definitions for sepsis and organ failure and guidelines for the use of innovative therapies in sepsis. Chest 1992;101:1644–55. 6. Corley KTT. Monitoring and treating haemodynamic disturbances in critically ill neonatal foals. Part 1. Haemodynamic monitoring. Equine Vet Educ 2002;5:345–48. 7. Palmer J. Recognition and resuscitation of the critically ill foal. In: Paradis MR (ed.) Equine Neonatal Medicine: A CaseBased Approach. Philadelphia: Elsevier Saunders, 2006; pp. 121–34. 8. Landry DW, Oliver JA. The pathogenesis of vasodilatory shock. N Engl J Med 2001;345:558–95. 9. Corley K. Treatments aimed at improving tissue perfusion. In: Corley K, Stephen J (eds) The Equine Hospital Manual. Oxford: Blackwell Publishing, 2008; pp. 446–56. 10. Corley KTT. Monitoring and treating the cardiovascular system in neonatal foals. Clin Tech Equine Pract 2003;2:42–55. 11. Polkes AC, Giguere S, Lester GD, Bain FT. Factors associated with outcome in foals with neonatal isoerythrolysis (72 cases, 1988-2003). J Vet Intern Med 2008;22:1216–22. 12. Divers T. Monitoring tissue oxygenation in the ICU patient. Clin Tech Equine Pract 2003;2:138–44. 13. Corley KTT, Axon JE. Resuscitation and emergency management for neonatal foals. Vet Clin North Am Equine Pract 2005;21:431–55. 14. Hollenberg SM, Ahrens TS, Astiz M, Chalfin D, Dasta J, Heard S, Martin C, Napolitano L, Susla G, Totaro R, Vincent J, Zanotti-Cavazzoni S. Practice parameters for haemodynamic support of sepsis in adult patients: 2004 update. Crit Care Med 2004;32:1928–48. 15. Nout YS, Corley KTT, Donaldson LL, Furr MO. Indirect oscillometric and direct blood pressure measurements in anesthetized and conscious neonatal foals. J Vet Emerg Crit Care 2002;12:75–80. 16. Giguere S, Knowles HA, Valverde A, Bucki E, Young L. Accuracy of indirect measurement of blood pressure in neonatal foals. J Vet Intern Med 2005;19:571–6. 17. LeDoux D, Astiz ME, Carpati CM, Rackow EC. Effects of perfusion pressure on tissue perfusion in septic shock. Crit Care Med 2000;28:2729–32.
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18. Corley KT. Inotropes and vasopressors in adults and foals. Vet Clin North Am Equine Pract 2004;20:77–106. 19. Gregory GA. Resuscitation of the newborn. In: Grenvik A, Ayres SM, Holbrook PR, Shoemaker WC (eds). Textbook of Critical Care, 4th edn. Philadelphia: WB Saunders, 2000; pp. 27–37. 20. Carcillo JA, Davis AL, Zaritsky A. Role of early fluid resuscitation in pediatric septic shock. J Am Med Assoc 1991;266:1242–5. 21. Rizoli SB. Crystalloids and colloids in trauma resuscitation: a brief overview of the current debate. J Trauma 2003:54:S82–8. 22. Magdesian KG. Monitoring the critically ill equine patient. Vet Clin North Am Equine Pract 2004;20:11–39. 23. Sprayberry KA. Neonatal transfusion medicine: the use of blood, plasma, oxygen-carrying solutions, and adjunctive therapies in foals. Clin Tech Equine Pract 2003;2:31– 41. 24. Giguere S, Polkes AC. Immunologic disorders in neonatal foals. Vet Clin North Am Equine Pract 2005;21:241–72. 25. Sehgal LR, Zebala LP, Takagi I, Curran RD, Votapka TV, Caprini JA. Evaluation of oxygen extraction ratio as a physiologic transfusion trigger in coronary artery bypass graft surgery patients. Transfusion 2001;41:591–5. 26. Todd GL, Baroldi G, Pieper GM, Clayton FC, Eliot RS. Experimental catecholamine-induced myocardial necrosis. I. Morphology, quantification and regional distribution of acute contraction band lesions. J Mol Cell Cardiol 1985;17:317–38. 27. Stopfkuchen H. What is sufficient blood pressure in the preterm newborn? Klin Padiatr 2003;215:16–21. 28. Valverde A, Giguere S, Sanchez LC, Shih A, Ryan C. Effects of dobutamine, norepinephrine, and vasopressin on cardiovascular function in anesthetized neonatal foals with induced hypotension. Am J Vet Res 2006;67:1730–7. 29. Hollis AR, Ousey JC, Palmer L, Stoneham SJ, Corley KT. Effects of norepinephrine and a combined norepinephrine and dobutamine infusion on systemic hemodynamics and indices of renal function in normotensive neonatal Thoroughbred foals. J Vet Intern Med 2006;20:1437–42. 30. Duranteau J, Sitbon P, Teboul JL, Vicaut E, Anquel N, Richard C, Samii K. Effects of epinephrine, nore-
31.
32.
33.
34.
35.
36.
37.
38. 39.
40.
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pinephrine, or the combination of norepinephrine and dobutamine on gastric mucosa in septic shock. Crit Care Med 1999;27:893–900. Craig CA, Haskins SC, Hildebrand SV. The cardiopulmonary effects of dobutamine and norepinephrine in isoflurane-anesthetized foals. Vet Anaesth Analg 2007;34: 377–87. Palmer JE. Foal cardiopulmonary resuscitation. In: Orsini JA, Divers TJ (eds) Manual of Equine Emergencies, 2nd edn. Philadelphia: Saunders, 2003; pp. 581–614. Holmes CL, Landry DW, Granton JT. Science review: vasopressin and the cardiovascular system. Part 1. Receptor physiology. Crit Care 2003;7:427–34. Holmes CL, Landry DW, Granton JT. Science review: vasopressin and the cardiovascular system. Part 2. Clinical physiology. Crit Care 2004;8:15–23. O’Brien A, Clapp L, Singer M. Terlipressin for norepinephrine-resistant septic shock. Lancet 2002;359: 1209–10. Davenport CL, Boston RC, Richardson DW. Effects of enrofloxacin and magnesium deficiency on matrix metabolism in equine articular cartilage. Am J Vet Res 2001;52:160–6. Durando MM, MacKay RJ, Linda S, Skelley LA. Effect of polymyxin B and Salmonella typhimurium antiserum on horses given endotoxin intravenously. Am J Vet Res 1994;55:921–7. Barton MH. Use of polymyxin B for treatment of endotoxaemia in horses. Comp Cont Educ Pract Vet 2000;11:1056–9. Annane D, Bellissant E, Bollart PE, Briegel J, Confalonieri M, De Gaudio R, Keh D, Kupfer Y, Oppert M, Meduri GU. Corticosteroids in the treatment of severe sepsis and septic shock in adults: a systematic review. J Am Med Assoc 2009;301:2362–2375. Sligl WI, Milner DA, Sundar S, Mphatswe W, Majumdar SR. Safety and efficacy of corticosteroids for the treatment of septic shock: a systematic review and meta-analysis. Clin Infect Dis 2009;49:93–101. Hart KA, Barton, MH. Stressed out? Relative adrenal insufficiency in critically ill foals. In: Proceedings of the 26th American College of Veterinary Internal Medicine Forum, 2008, pp. 201–3.
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Chapter 15
Perinatal Asphyxia Syndrome Pamela A. Wilkins
Introduction Hypoxic ischemic encephalopathy (HIE), also referred to as neonatal encephalopathy (NE), is the most common clinical presentation of a broader syndrome of perinatal asphyxia syndrome (PAS). Management of foals presenting with signs consistent with a diagnosis of NE requires the clinician to fully examine other body systems and provide therapy directed at treating other involved systems.1 Although PAS is most clinically obvious as NE, the gastrointestinal tract and kidneys are frequently also affected and complications associated with these systems should be expected. Cardiovascular, respiratory, and endocrine disorders may also be present. Neonatal encephalopathy is one of the most common diseases of the equine neonate and is also described as dummy foal syndrome and neonatal maladjustment syndrome (NMS).2–4 A wide spectrum of clinical signs are associated with NE ranging from mild depression with loss of the suck reflex to grand mal seizure activity. The majority of affected foals are normal at birth but show signs of central nervous system (CNS) abnormalities within a few hours following birth while some will not show signs until 24 hours of age.
Predisposing factors Conditions associated with PAS in foals include dystocia, induced parturition, cesarean sections, placentitis (Figures 15.1 and 15.2), premature placental separation, meconium aspiration, twin foals, fetal infection, severe maternal illness, maternal surgery, post-term pregnancies, and normal parturition. A fair number of foals have no known peripartum period of hypoxia, suggesting that these foals result from unrecognized in utero hypoxia. Severe maternal illness may also result in foals born with PAS. There is increasing evidence that cytokinemia, resulting from
placental infection or insult, is a major contributor to NE in infants. The incidence of NE is increased with the presence of maternal fever, something veterinarians have suspected for the last decade or so.5, 6
Clinical signs Foals with PAS/NE may appear healthy at birth but exhibit central nervous system (CNS) abnormalities within hours of birth to 1–2 days of age. Clinical signs of PAS/NE in the foal are extremely variable and can include one or more of the following: weakness (hypotonia), mental depression (stupor, somnolence, difficult to arouse, lethargy, coma), mild to severe seizure activity, tremors, and hypertonia. Other clinical signs such as inability to find the udder, inappropriate nursing behavior, loss of suckle reflex, loss of affinity for the dam, loss of recognition of environment, abnormal vocalization, dysphagia, weak tongue tone, central blindness, irregular respiratory pattern (apnea, abnormally slow respiratory rate), and proprioceptive deficits may also be encountered. It is again important to recognize that while the earliest and most obvious manifestation of PAS is NE, other organ systems are generally affected to varying degrees. Repetitive detailed examination of other systems is a requirement of management of foals with PAS/NE. Differential diagnoses for PAS/NE include severe sepsis, meningitis, cerebral trauma, cerebral vascular accidents, and developmental abnormalities such as hydrocephalus.
Pathophysiology The underlying pathophysiological details of PAS and NE in the foal are unknown and likely multifactorial, and equine neonatologists have long looked to human studies and models of the human disease
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 15.1 Placenta from a mare where the foal suffered severe perinatal asphyxia syndrome secondary to both placentitis and probable in utero cytokinemia and premature delivery.
for understanding of the syndrome in the equine neonate. Perinatal brain damage in the mature human fetus is thought to result from severe uterine asphyxia due to an acute reduction of uterine or umbilical circulation. If the hypoxic insult continues, cardiac output falls and cerebral circulation diminishes with concomitant decreased energy production.7 Eventually, ionic gradients across neurons are lost, leading to calcium overload of neurons. Calcium overload of the neuron leads to cell damage by activation of calcium-dependent proteases, lipases, and endonucleases. Protein biosynthesis is halted. Calcium also enters the cells by glutamate-regulated ion channels as glutamate, an excitatory neurotransmitter, is released from presynaptic vesicles following anoxic cellular depolarization. Once the anoxic event is over, protein synthesis remains inhibited in specific areas of the brain and returns to normal in less vulnerable
areas of the brain. Loss of protein synthesis appears to be an early indicator of cell death due to the primary hypoxic/anoxic event.8 A second wave of neuronal cell death occurs during the ‘reperfusion’ phase and is thought to be similar to classically described ‘post-ischemic reperfusion injury’ in that damage is due to production of and release of oxygen radicals, synthesis of nitric oxide (NO), and inflammatory reactions.9 Part of the secondary cell death that occurs is thought to be due to apoptosis while additional secondary cell death is thought be secondary to the neurotoxicity of glutamate and aspartate.10, 11 There is evidence that the excitotoxic cascade that evolves during NE/PAS extends over several days from the time of insult and is modifiable.10, 11 The activation of the N-methyl-D-aspartate (NMDA) subtype of glutamate receptors is implicated in the pathophysiology of traumatic brain injury and is suspected to play a role in NE/PAS.10–12 Magnesium has been demonstrated to block NMDA receptors and, while clinical trials investigating the efficacy of magnesium treatment following hypoxia in infants are under way, few reports currently exist. Magnesium sulfate was used to treat nine infants after perinatal asphyxia in one study, there was no control group, and all children were neurologically normal at 1 year of age. Seizures did not occur in any of these children nor were any adverse side effects noted.13 Magnesium sulfate administration failed to delay the global impairment in energy metabolism after hypoxia-ischemia, characteristic of severe brain damage, in newborn piglets and at 48 hours post-HI no difference could be found in the severity of injury in magnesium-treated piglets compared to placebo-treated piglets, suggesting magnesium may not be protective with severe acute injury.14
Treatment
Figure 15.2 Transrectal ultrasonographic examination of caudal reproductive tract of mare with placentitis. Note thickened placenta and fluid accumulation between placenta and cervix. Placentitis such as this can be associated with the development of perinatal asphyxia syndrome in the fetus postpartum.
Therapy for the various manifestations of PAS involves control of seizures, general cerebral support, correction of metabolic abnormalities, maintenance of normal arterial blood gas values, maintenance of tissue perfusion, maintenance of renal function, treatment of gastrointestinal dysfunction, prevention/ recognition/early treatment of secondary infections, and general supportive care (see Table 15.1). It is important that seizures be controlled as cerebral oxygen consumption increases fivefold during seizures. Diazepam and midazolam can be used for emergency control of seizures. If seizures are not readily stopped with diazepam (0.1–0.2 mg/kg or 5–10 mg to a 50 kg foal i.m. or i.v.) or midazolam (0.1–0.2 mg/kg or 5–10 mg to a 50 kg foal i.m. or i.v.), or more than two seizures are recognized, then diazepam should
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Table 15.1 Drugs and dosages used in the treatment of perinatal asphyxial syndrome and neonatal encephalopathy Drug
Indication
Dose
Potential complications
Diazepam
Single or short-term seizure control
0.1–0.2 mg/kg (5–10 mg to a 50 kg foal i.m. or i.v.) NOT water soluble
Rapid i.v. administration may result in respiratory depression
Midazolam
Single or short-term seizure control
0.1–0.2 mg/kg (5–10 mg to a 50 kg foal i.m. or i.v.).
Rapid i.v. administration may result in respiratory depression
Midazolam CRI
Longer-term control for repetitive seizures. Mild sedation for hyper-responsive or ‘jittery’ foals
3–6 mg/h. Midazolam is water soluble and CRI is administered in isotonic crystalloid fluid using a 0.5 mg/mL concentration
Rapid i.v. administration may result in respiratory depression. Higher doses may be used if necessary. Advantage is ability to titrate to effect and reversibility if needed
Phenobarbital
Longer-term seizure control
2–5 mg/kg i.v.IV slowly over 20 minutes. Start with lower dose and monitor to effect. Maximal effect expected after 45 minutes
Respiratory depression, hypothermia, hypotension and pharyngeal collapse particularly at higher doses or in more severely affected foals
Thiamine
Metabolic support
1–20 mg/kg q 12 h added to i.v. fluids (protect from light)
None
Mannitol
Intercellular edema: osmotic diuretic
0.25–1.0 g/kg i.v. as a 20% solution rapidly over 15–20 minutes
Dehydration. May result in significant hyperosmolarity with repeated administration. May exacerbate cerebral bleeding
Dimethylsulfoxide
Anti-inflammatory, Intercellular edema: osmotic agent
0.1–1 g/kg i.v. as a 10% solution
Odor; OSHA restrictions in some areas, hemolysis
Gabapentin
Neuroprotection: GABA receptor agonist
10–15 mg/kg/day divided equally and given orally 3 to 4 times a day
None described: an uninvestigated drug in equine neonates
Magnesium CRI
Neuroprotection: NMDA receptor antagonist
See below. Can precipitate other infusates, test compatibility or discontinue when administering intravenous antimicrobial drugs
In very high doses (> 10×) muscular weakness can occur as can hypotension
CRI: constant rate infusion. Magnesium CRI: Remove 20 mL from 100 mL bag sterile isotonic saline (0.9%). Add 20 mL 50% MgSO4 for a final volume of 100 mL. Administer loading dose at 25 mL/h for a 50 kg foal for 1 hour, decrease infusion to 12.5 mL/h thereafter. This is approximately 50 mg/kg loading dose followed by 25 mg/kg maintenance dose. Infusion can be maintained for 24–48 hours.
be replaced with either phenobarbital (2–5 mg/kg i.v. slowly over 20 minutes) given to effect or a midazolam constant rate infusion (CRI) (3–6 mg/h). The half-life of phenobarbital can be quite long in the foal (more than 200 hours) and this should be kept in mind when monitoring neurological function in these cases after phenobarbital administration (JE Palmer, personal communication).15 Complications commonly reported with phenobarbital include hypothermia, hypotension, loss of respiratory drive, and decreased pharyngeal tone with consequent airway obstruction, even when used at relatively low dosages, in severely affected foals. Midazolam may be more ‘user friendly’ – titratable with fewer side effects – but requires the ability to administer and maintain a constant rate infusion. Severe cases may require a combination of antiseizure medications.
In cases of NE, ketamine and xylazine should be avoided because of their association with increased intracranial pressure. It is important to protect the foal from injury during a seizure and also to ensure the patency of their airway to prevent the onset of negative pressure pulmonary edema or aspiration pneumonia.16 Probably the most important therapeutic interventions are aimed at maintaining cerebral perfusion. Cerebral interstitial edema is only truly present in the most severe cases and, if suspected, can be treated with mannitol administration. In most cases the lesion is intracellular edema and both mannitol and hypertonic saline infusions are minimally effective in treating cellular edema.17 Thiamine supplementation (10–20 mg/kg q 12 h added to i.v. fluids and protected from light) can be administered to support metabolic
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Figure 15.3 Dorsal view of brain from foal with severe perinatal asphyxia syndrome/neonatal encephalopathy. In this case, severe edema of the brain resulted in cerebellar herniation; note ‘coning’ of the caudal cerebellum.
processes, specifically mitochondrial metabolism and membrane Na+/K+/ATPases, involved in maintaining cellular fluid balance.18, 19 Only if cellular necrosis and vasogenic edema are present are drugs such as mannitol (0.25–1.0 g/kg i.v. as a 20% solution rapidly) and dimethylsulfoxide (DMSO)(0.1–1 g/kg i.v. as a 10% solution) indicated, and again, these cases are usually the most severely affected and most likely to suffer the severe complication of cerebellar herniation (see Figure 15.3). This author rarely uses DMSO and has recognized no change in outcome over more than 13 years by discontinuing its use. Other practitioners continue to use this drug, with many using it now at the lower end of the dose range. Naloxone (0.01–0.02 mg/kg i.v.) has been advocated for the treatment of NE in humans and in foals,20, 21 perhaps based on a study suggesting that post-asphyxia blood–brain barrier disruption was causally related to poor neurological outcome in a lamb model of NE, and that naloxone prevented both disruption and neurological dysfunction among those survivors with intact blood–brain barrier.21 There is also evidence suggesting that naloxone increases ischemic injury in neonatal rats, making use of this compound controversial.22 Gammaaminobutyric acid (GABA)ergic agonists such as gabapentin are being used by some practitioners in the management of PAS/NE in foals, based on evidence showing neuroprotection when used in ischemia, both alone and in combination with NMDA antagonists such as magnesium. No specific dose is reported for use in foals; however the human pediatric dose is 10–15 mg/kg/day divided equally and given orally 3 to 4 times a day.23–25
Despite a lack of consensus regarding the use of magnesium in the treatment of infants with NE, magnesium sulfate constant rate infusion has been used in the treatment of PAS/NE. The rationale is based primarily in the evidence demonstrating protection in some studies and a failure of any one study to demonstrate significant detrimental effects. Other potentially neuroprotective drugs used by other practitioners include ascorbic acid (50–100 mg/ kg/day orally), alpha-tocopherol (500–1000 U/day orally), and allopurinol. Foals with PAS often have a variety of metabolic problems including hypo- or hyperglycemia, hypoor hypercalcemia, hypo- or hyperkalemia, hypo- or hyperchloremia, and varying degrees of metabolic acidosis. While these problems need to be addressed, the normal period of hypoglycemia that occurs postpartum should not be forgotten and should not be treated aggressively for fear of worsening the neurological injury. Foals suffering from PAS will also have frequent and recurrent bouts of hypoxemia and occasional bouts of hypercapnia. Intranasal oxygen at 3–5 L/min (INO2) is generally needed in these cases both as a preventative therapy and as direct treatment, as the appearance of the abnormalities can be sporadic and unpredictable. Repeated arterial blood gas analyses should be performed to ensure Pao2 remains less than ∼ 120 mmHg and greater than ∼ 80 mmHg. While pulse oximetry is useful in recognizing hemoglobin desaturation events, it is not useful for recognizing either mild to moderate hypoxemia or hyperoxia. Additional respiratory support, particularly in those foals with centrally mediated hypoventilation and periods of apnea or abnormal breathing patterns, include caffeine (10 mg/kg per os or per rectum loading dose, 2.5 mg/kg maintenance dose prn) or doxapram CRI (0.02–0.05 mg/kg/h) and positive pressure ventilation. Recent studies have compared the efficacy of caffeine and doxapram in healthy anesthetized foals and also retrospectively in sick foals, but there currently is no definitive recommendation as to which to choose under which conditions in order to optimize ventilation.26, 27 Mechanical ventilation of patients with central hypoventilation can be very rewarding and is generally required for less than 48 hours. It is important that blood pH be monitored and maintained within the normal range but metabolic alkalosis in some of these cases may require the clinician to tolerate some degree of hypercapnia. When evaluating these cases and considering alternatives for treatment, pH is important. If the respiratory acidosis is not so severe as to adversely affect the patient (generally > 70 mmHg), and the pH is within normal limits, hypercapnia may be tolerated.28
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Perinatal Asphyxia Syndrome Maintaining tissue perfusion and oxygen delivery to tissues is a cornerstone of therapy for PAS in order to avoid additional injury. Oxygen-carrying capacity of the blood should be maintained; some foals will require transfusions to maintain a PCV > 20%. Adequate vascular volume is important, but care should be taken to avoid fluid or sodium overloading. Early evidence of fluid overload is subtle accumulation of ventral edema between the front legs and over the distal limbs. Fluid overload can result in cerebral edema, pulmonary edema, and edema of other tissues, including the gastrointestinal tract. This edema interferes with normal organ function and may worsen the condition of the patient. Perfusion is maintained by supporting cardiac output and blood pressure by judicious use of intravenous fluid support and inotrope/pressor support. We do not aim for any ‘magic’ systolic, mean or diastolic pressure. Instead we monitor urine output, mentation, limb perfusion, gastrointestinal function, and respiratory function as indicators that perfusion is acceptable. The kidney is a target for injury in these patients and it is not unusual for renal compromise to play a significant role in the demise of PAS foals. Clinical signs of renal disease are generally referable to disruption of normal control of renal blood flow and tubular edema, leading to tubular necrosis and renal failure. These foals present with signs of fluid overload and generalized edema. It is important that urine output and fluid therapy be balanced in these cases to prevent additional organ dysfunction associated with edema. Although evidence has accumulated that neither dopamine nor furosemide play a role in either protecting the kidney or reversing acute renal failure, these agents can be useful in managing volume overload in these cases.29–31 Overzealous use of diuretics and pressors in these cases can result in diuresis requiring increased intravenous fluid support and can be counterproductive. Low doses of dopamine, administered as a constant rate infusion (CRI) of 2–5 µg/kg/min, are usually effective in establishing diuresis by natriuresis. Bolus (0.25–1.0 mg/kg) or CRI (0.25–2.0 mg/kg/h) furosemide can administered but once furosemide diuresis is established electrolyte concentrations and blood gas tensions must be evaluated frequently as potassium, chloride, and calcium losses can be considerable and significant metabolic alkalosis can develop due to strong ion imbalances. The aim is urine output that is appropriate to fluid administration. The urine of normal newborn foals is quite dilute, reflecting the large free water load they incur by their diet. Expecting critically ill foals to produce large volumes of urine, particularly those on restricted diets or receiving total parenteral nutrition (TPN), is an exercise in futility and ma-
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nipulating fluid, pressor and/or diuretic therapy in an attempt to meet an artificial goal is, quite simply, poor medicine. One final caveat regarding renal dysfunction in PAS is that therapeutic drug monitoring should be performed when it is available. Many antimicrobial agents used in the management of these cases, most notably the aminoglycosides, depend on renal clearance. Aminoglycoside toxicity occurs in the equine neonate and will exacerbate or, at the least, complicate the management of renal failure originally due to primary hemodynamic causes. Foals with PAS, as in human infants, suffer from a variety of problems associated with abnormalities within the gastrointestinal tract.32 Commonly they present with ileus, recurrent excessive gastric reflux, and gas distention. These problems are exacerbated by constant feeding in the face of continued dysfunction and continued hypoxia. Frequently, enteral feeding cannot meet their nutritional requirements and partial (PPN) or total parenteral nutrition (TPN) is required. Special attention should be paid to passive transfer of immunity status and glucose homeostasis in these cases. Clinical signs of injury to the gastrointestinal tract can be subtle and lag behind other abnormalities for days to weeks. Any evidence of bloody gastric contents should be taken as an indication of significant intestinal injury. Low-grade colic, decreased gastrointestinal motility, decreased fecal output, and low weight gain are amongst the most common clinical signs of gastrointestinal dysfunction in these case, but more severe problems, including necrotizing enterocolitis and intussusception, have been associated with PAS as well. The return to enteral feeding must be slow in many of these cases. Sucralfate, 1–2 g q 6 h, has been used by the author in cases where feeding is delayed and gastrointestinal injury is suspected. Foals with PAS are also susceptible to secondary infection and many have the complication of failure of passive transfer. Intravenous plasma as treatment for FPT is indicated as is broad-spectrum antimicrobial prophylaxis. Renal damage is frequently present in these cases and the use of nephrotoxic drugs, such as the aminoglycoside antimicrobials and nonsteroidal anti-inflammatory agents, should be minimized. If infection is recognized in these patients after hospitalization, attention should be given to the likelihood of nosocomial infection and antimicrobial therapy should be directed, until culture and sensitivity results become available, based on known nosocomial pathogens in the neonatal intensive care unit (NICU) or hospital/clinic and their susceptibility patterns. Repeat determination of IgG concentration should be made and additional intravenous plasma therapy may be required. Nosocomial infections can be rapidly overwhelming and any acute
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deterioration in the condition of a foal with PAS indicates a need for further evaluation for possible sepsis. The prognosis for foals with PAS is good to excellent when it is recognized early and aggressively treated in term foals. Up to 80% of these neonates survive and go on to lead productive and useful athletic lives.33–35 Prognosis decreases with delayed or insufficient treatment and concurrent problems such as prematurity and sepsis, and evidence is accumulating that failure to resolve hyperlactatemia within 48 hours of presentation in foals with PAS indicates a poorer prognosis for ultimate survival.36, 37
References 1. Whitelaw A. Systematic review of therapy after hypoxicischaemic brain injury in the perinatal period. Semin Neonatol 2000;5(1):33–40. 2. Rossdale Peter D. Clinical studies on 4 newborn thoroughbred foals suffering from convulsions with special reference to blood gas chemistry and pulmonary ventilation. Res Vet Sci 1969;10(3):279–91. 3. Palmer AC, Rossdale PD. Neuropathology of the convulsive foal syndrome. J Reprod Fertil Suppl 1975;23:691–4. 4. Kitchen H, Rossdale PD. Metabolic profiles of newborn foals. J Reprod Fertil Supplement 1975;23:705–7. 5. Nelson KB, Willoughby RE. Infection, inflammation and the risk of cerebral palsy. Curr Opin Neurol 2000;13(2): 133–9. 6. McGlothlin JA, Lester GD, Hansen PJ, Thomas M, Pablo L, Hawkins DL, LeBlanc MM. Alteration in uterine contractility in mares with experimentally induced placentitis. Reproduction 2004;127(1):57–66. 7. Goetzman BW, Itskovitz J, Rudolph AM. Fetal adaptations to spontaneous hypoxemia and responses to maternal oxygen breathing. Biol Neonate 1984;46(6):276–84. 8. Evrard P. Pathophysiology of perinatal brain damage. Dev Neurosci 2001;23(3):171–4. 9. Andine P, Jacobson I, Hagberg H. Enhanced calcium uptake by CA1 pyramidal cell dendrites in the postischemic phase despite subnormal evoked field potentials: excitatory amino acid receptor dependency and relationship to neuronal damage. J Cereb Blood Flow Metab 1992;12(5):773– 83. 10. Sebastiao AM, de Mendonca A, Moreira T, Ribeiro JA. Activation of synaptic NMDA receptors by action potentialdependent release of transmitter during hypoxia impairs recovery of synaptic transmission on reoxygenation. J Neurosci 2001;21(21):8564–71. 11. Vexler ZS, Ferriero DM. Molecular and biochemical mechanisms of perinatal brain injury. Semin Neonatol 2001;6(2):99–108. 12. D’Souza SW, McConnell SE, Slater P, Barson AJ. Glycine site of the excitatory amino acid N-methyl-D-aspartate receptor in neonatal and adult brain. Arch Dis Child 1993;69(2):212–15. ´ 13. Maroszynska I, Sobolewska B, Gulczynska E, Zylinska L, ´ Lerch E, Kicinska M, Rudecka M. Can magnesium sulfate reduce the risk of cerebral injury after perinatal asphyxia? Acta Polon Pharma 1999;56(6):469–73.
14. Greenwood K, Cox P, Mehmet H, Penrice J, Amess PN, Cady EB, Wyatt JS, Edwards AD. Magnesium sulfate treatment after transient hypoxia-ischemia in the newborn piglet does not protect against cerebral damage. Pediatr Res 2000;48(3):346–345. 15. Spehar AM, Hill MR, Mayhew IG, Hendeles L. Preliminary study on the pharmacokinetics of phenobarbital in the neonatal foal, Equine Vet J 1984;16(4):368–71. 16. Tute AS, Wilkins PA, Gleed RD, Credille K, Murphy D, Ducharme NG. Negative pressure pulmonary edema as a post-anesthetic complication associated with upper airway obstruction in a horse. Vet Surg 1996;25(6):519– 23. 17. Kempski O. Cerebral edema. Semin Nephrol 2001;21(3):303– 7. 18. Watanabe I, Tomita T, Hung KS, Iwasaki Y. Edematous necrosis in thiamine-deficient encephalopathy of the mouse. J Neuropath Exp Neurol 1981;40(4):454–71. 19. Wilkins PA, Vaala WE, Zivotofsky Dm Twitchell ED. A herd outbreak of equine leukoencephalomalacia. Cornell Vet 1994;84(1):53–9. 20. Chernick V, Craig RJ. Naloxone reverses neonatal depression caused by fetal asphyxia. Science 1982;216(4551):1252– 3. 21. Ting P, Pan Y. The effects of naloxone on the post-asphyxic cerebral pathophysiology of newborn lambs. Neurol Res 1994;16(5):359–64. 22. Young RS, Hessert TR, Pritchard GA Yagel SK. Naloxone exacerbates hypoxic-ischemic brain injury in the neonatal rat. Am J Obst Gynec 1984;150(1):52–6. 23. Bain FT. Neurologic disorders in foals other than hypoxicischemic encephalopathy. Proceedings of the International Veterinary Emergency Critical Care Symposium, San Antonio, TX, 1998, pp. 691–2. 24. Lyden PD, Lonzo L. Combination therapy protects ischemic brain in rats: a glutamate antagonist plus a gammaaminobutyric acid agonist. Stroke 1994;25(1):189–96. 25. Madden KP. Effect of gamma-aminobutyric acid modulation on neuronal ischemia in rabbits. Stroke 1994;25(11):2271–4. 26. Gigu`ere S, Sanchez LC, Shih A, Szabo NJ, Womble AY, Robertson, SA. Comparison of the effects of caffeine and doxapram on respiratory and cardiovascular function in foals with induced respiratory acidosis. Am J Vet Res 2007;68(12):1407–16. 27. Gigu`ere S, Slade JK, Sanchez LC. Retrospective comparison of caffeine and doxapram for the treatment of hypercapnia in foals with hypoxic-ischemic encephalopathy. J Vet Intern Med 2008;22(2):401–5. 28. Ambalavanan N, Carlo WA. Hypocapnia and hypercapnia in respiratory management of newborn infants. Clin Perinatol 2001;28(3):517–31. 29. Filler G. Acute renal failure in children: aetiology and management. Paediatric Drugs 2001;3(11):783–92. 30. Rudis MI. Low-dose dopamine in the intensive care unit: DNR or DNRx? Critl Care Med 2001;29(8):1638–9. 31. Kellum JA, M Decker J. Use of dopamine in acute renal failure: a meta-analysis. Crit Care Med 2001;29(8):1526– 31. ˜ F, Bur32. Martin-Ancel A, Garcia-Alix A, Gaya F, Cabanas gueros M, Quero J. Multiple organ involvement in perinatal asphyxia. J Pediatr 1995;127(5):786–93. 33. Baker SM, Drummond WH, Lane TJ, Koterba AM. Followup evaluation of horses after neonatal intensive care. J Am Vet Med Assoc 1986;189(11):1454–7.
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Perinatal Asphyxia Syndrome 34. Axon J, Palmer J, Wilkins PA. Short-term and long-term athletic outcome of neonatal intensive care unit survivors. Proceedings of the American Association of Equine Practitioners 1999;45:224–5. 35. Freeman L, Paradis MR. Evaluating the effectiveness of equine neonatal intensive care. Vet Med 1992;87:921– 6.
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36. Henderson ISF, Franklin RP, Boston RC, Wilkins PA. Association of hyperlactataemia with age, diagnosis and survival in equine neonates. J Vet Emerg Crit Care 2008;18(5):496–502. 37. Wotman K, Wilkins PA, Boston RC, Palmer JE. Association of blood lactate concentration and outcome in foals. J Vet Intern Med 2009;23(3):598–605.
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Chapter 16
Sepsis L. Chris Sanchez
Pathophysiology Much of the clinical syndrome classically associated with sepsis is caused by a non-specific inflammatory response. The systemic inflammatory response syndrome (SIRS) refers to a systemic inflammatory response, regardless of the inciting cause, which results in at least two of the following four clinical manifestations: fever; tachycardia; tachypnea or hyperventilation; leukocytosis, leukopenia or relative increase of circulating immature neutrophils. When SIRS occurs in response to a confirmed infectious process, the process is termed sepsis. The presence of viable bacteria in the blood is termed bacteremia, and the presence of other viable organisms in the blood is termed similarly (i.e., viremia, fungemia). Septic shock is defined as sepsis-induced hypotension which persists despite adequate fluid therapy.1 Endotoxin plays a critical role in the pathophysiology of septic shock in Gram-negative sepsis,2, 3 which is common in equine neonates.4, 5 Septic foals can have decreased gene expression of TNF-α and TNF-β and increased expression of Il-8 relative to healthy foals6 or increased gene expression of Il-4 and TLR4.7
ure of passive transfer of immunoglobulins. A number of studies have documented a close relationship between the foal serum IgG concentration and incidence of disease.10–13 The route and timing of transfer are also likely relevant, along with the potential for bacterial challenge. For example, colostrum administration via nasogastric tube was associated with disease in one report.9 Farm management is particularly important, and one study demonstrated that foals with partial failure of passive transfer were not at greater risk of disease than those with adequate transfer on a well-managed farm.14 Postnatal routes of infection include the umbilicus, gastrointestinal tract, and respiratory tract, with the intestinal tract emphasized.15 The other main postnatal factors beyond failure of passive transfer include gestational age and environmental conditions. Foals with exceptionally short or long gestational lengths are at risk for development of sepsis.16 Unsanitary environmental conditions can result in an increased bacterial load to the neonatal gastrointestinal tract, especially during the initial periods of udder seeking.
Clinical signs and diagnosis Predisposing factors and routes of infection Many events can predispose an equine neonate to infection, such as maternal illness, alterations in gestational length, partial or complete failure of passive transfer, poor sanitary conditions, and others, but none are completely necessary for development of infection. Maternal factors include dystocia, premature placental separation, placentitis, and other maternal illness such as colic. Such problems were reported in 24% of bacteremic foals in one study8 and birth complications (i.e., dystocia) were associated with disease in another.9 Most postnatal risk factors correlate with possible routes of infection. The main exception to this is fail-
Physical examination findings Initial clinical signs can be vague and vary widely but frequently include depression, decreased or absent suckling from the mare, and lethargy, which may progress to recumbency. Dehydration becomes a more significant problem as time progresses; tachycardia and tachypnea are common. The mucous membranes often develop a bright or injected appearance (with or without petechiation noted on oral or conjunctival mucous membranes or the internal pinnae of the ears) or hyperemia along the coronary bands, and the capillary refill time may be rapid (see Figure 16.1). Rectal temperature may be normal
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Sepsis
Figure 16.1 Oral mucous membranes from a 2-day-old Thoroughbred colt presenting in septic shock. Note the overall dark color and injected vessels. Small petechiae are also seen in the upper right corner.
or mildly increased. Hypothermia can be associated with hypovolemia or moderate to severe prematurity. Other signs of hypovolemia include decreased borborygmi, poor peripheral pulses, cool extremities, and decreased or absent urine output. Diarrhea is a common early localizing signs in foals with sepsis and no other evidence of enteric pathogens and can be the primary presenting complaint. Other localizing signs include uveitis, seizures, joint effusion, lameness, respiratory disease or distress, subcutaneous abscesses, patent urachus, and omphalitis. Physical examination of the mare and placenta, if available, should be considered an extension of the foal’s physical examination. Any indication of placental abnormality, including increased placental weight, or maternal illness should immediately increase one’s index of suspicion for neonatal illness. Although a thorough discussion of placental examination is discussed elsewhere in this text (see Chapters 8 and 9), one should note that not all placental abnormalities evident upon histological examination are evident upon gross inspection. Thus, if other factors suggest in-utero infection, a grossly normal-appearing placenta should not exclude this possibility.
Clinicopathological findings Clinical signs and historical information alone are often sufficient for suspicion of sepsis. However, in addition to physical examination findings, laboratory data may be helpful in diagnosing early sepsis. Leukopenia, characterized by neutropenia, is the most common hematological finding associated with acute sepsis. During the first week of life, normal values for total leukocyte and neutrophil counts
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can vary, ranging from 6900–14 400/µL and 5550–12 380/µL, respectively (<12 hours of age), to roughly 5000–11 000/µL and 3200–9000/µL, respectively (1–3 days of age), to 6300–13 600/µL and 4350–10 550/µL, respectively (1 week of age).17 In one study, septic foals less than 1 week of age had lower total WBC, neutrophils, and lymphocytes and higher bands and monocytes than healthy age-matched controls.18 One should note that premature or dysmature foals will also commonly have a decreased neutrophil count in the absence of sepsis. However, septic foals typically have a degenerative left shift and evidence of toxicity (Doehle bodies, toxic granulation, and vacuolization), whereas these findings are not typical of uncomplicated prematurity. In older septic foals (8–14 days), the total WBC, neutrophils, and bands were higher than in age-matched controls.18 A high fibrinogen concentration at or shortly after birth should raise the suspicion of in-utero infection.19 Hypoglycemia (<100 mg/dL) is common initially, especially in foals less than 24 hours of age.19 While hypoglycemia is predominantly related to decreased intake, endotoxemia can contribute to hypoglycemia by decreasing hepatic gluconeogenesis and increasing peripheral glucose uptake. In the initial phase of treatment, one should carefully monitor the rate of supplementation, as many foals subsequently develop hyperglycemia in response to dextrose infusion. Other biochemical abnormalities common in septic foals include azotemia and hyperbilirubinemia.4 Septic foals were reported to have higher ACTH and cortisol concentrations and ACTH/cortisol ratios than similarly aged normal foals in one report20 and increased arginine vasopressin, ACTH, and cortisol concentrations in another.21 Common abnormalities on arterial blood gas analysis include acidemia and hyperlactemia.19, 22 Arterial hypoxemia (<70 mmHg) has been reported in as many as 83% of septic foals in some populations.19 However, other populations of septic foals reportedly have arterial oxygen tensions of 90 mmHg (mean)23 and 64 mmHg (median).8 One should take care when interpreting arterial oxygen tension in foals, as a decrease in Pa O2 is common following a change from standing or sternal recumbency to lateral recumbency.24 The coagulation and fibrinolytic systems of the septic newborn are often abnormal, with clinically relevant decreases in antithrombin III and elevations in prothrombin time (PT), activated partial thromboplastin time (APTT), fibrinogen, fibrin degradation products,18 and D-dimer concentrations.25 Some patients may develop active hemorrhage and/or thrombosis. This can include thrombosis of major arteries, such the aorta, iliac, femoral, or brachial arteries.
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Definitive diagnosis Blood culture is the gold standard for diagnosis of systemic bacterial infection. Identification of a causative organism allows for directed antimicrobial therapy as well as determination of patterns in infection. Samples for culture should be collected from a large vein (jugular, cephalic, or saphenous) after surgical clip and aseptic preparation. The sample should be collected into a sterile syringe without anticoagulant and immediately placed into appropriate media (such as BBLTM SEPTI-CHEKTM Thioglycollate and Tryptic soy broth, Becton Dickinson & Company, Sparks, MD). Sterile sample collection from a venous catheter at the time of placement is also acceptable. For those foals receiving antimicrobial therapy prior to sample collection, use of an appropriate medium with resins may improve microbial recovery. Care should be taken to infuse the recommended volume of blood to promote optimum recovery. Two main factors limit the usefulness of blood cultures from a diagnostic standpoint. First, positive results are usually not available for at least 48 hours following submission. Second, a positive blood culture, while extremely specific, does not confer optimal sensitivity. Many foals with histological evidence of sepsis have historical evidence of a negative blood culture. In one study, only 40% of E. coli infections were successfully identified by blood culture relative to those organisms obtained from culture at necropsy.26 Thus, a means of identifying at-risk foals would be a valuable tool for the attending clinician. The modified ‘sepsis score’ currently used in many hospitals is calculated based upon a number of historical, physical, and laboratory findings with reported sensitivity and specificity of 92.8% and 85.9%, respectively.27 The sepsis score has not been as accurate at other institutions. Recent data have shown a false-negative rate of 48% in blood culture-positive foals in Ohio.8 In a study at the University of Georgia examining foals with either a sepsis score ≥ 11, a positive blood culture, or ≥ 3 foci of infection, 43/247 had a sepsis score <11 but at least one of the other criteria and 46/250 had a sepsis score ≥ 11 without either of the other criteria.28 In Virginia, the modified27 and original sepsis scores29 each produced a positive predictive value of 84% with negative predictive values of 55 and 53%, respectively.23 Results from these studies stress the importance of regional and institutional variability in the use of such scoring systems and their relatively low specificity and negative predictive value. Thus, while a ‘positive’ score is supportive of sepsis in a suspected animal, a ‘negative’ score alone should not be used to withhold antibiotic therapy from an at-risk foal. Similarly, the use of a positive score alone, without complementary culture results or necropsy find-
ings, should be used cautiously to confirm a diagnosis of sepsis for retrospective studies.
Causative organisms Several retrospective studies have evaluated the most common organisms isolated from both blood culture and necropsy specimens in septic foals over the years. Whereas Gram-positive organisms predominated in the 1940s-1950, E. coli has remained the predominant organism isolated from septic foals regardless of clinic location or methodology since that time.5, 8, 19, 26, 28,30–32 Era and geographic location appear to play a major role in prevalence of other causative organisms. In the late 1990s, Gram-positive bacteria (Enterococcus, Streptococcus, Staphylococcus spp.) cumulatively played a major role in disease pathogenesis in Pennsylvania,5 whereas Actinobacillus spp. accounted for approximately 30% of all isolates in Ohio.8 Recently, Gram-positive organisms were isolated from 40.3% of all blood cultures from neonatal foals in Australia.32 A Georgia study reported a dramatic decrease in the percentage of E. coli isolates between 1986–1990 and subsequent 5-year sampling periods.28 The predominant organisms with increased percentages over the same period were Enterococcus and Staphylococcus spp. A Florida study documented a decrease in Gram-negative enteric organisms in the 2000s relative to the 1980s, a decrease in Salmonella spp. in the 2000s relative to earlier years, and a decrease in Actinobacillus spp. in later years relative to the 1980s.31 Systemic fungal infections can also occur in neonatal foals. Clinical signs include persistent fever and thrush (white plaques on the lingual surface). The most commonly implicated organism is Candida albicans.33, 34 Risk factors include prolonged hospitalization34 and immunodeficiency.33 Foals infected with equine herpesvirus 1 can initially be mistaken for those with uncomplicated sepsis. However, viremic foals are typically icteric and extremely leukopenic (<3000 total WBC/µL) and rarely survive despite aggressive therapy.35
Therapy and monitoring Because specifics of critical care monitoring, fluid therapy, antimicrobial therapy, anti-acid therapy, enteral and parenteral nutrition and infections of specific body systems are covered in detail elsewhere in this text (see Section (e) Specific diagnostic and management techniques), they will be discussed only briefly in this and the following sections. Antibiotics provide the basis of therapy for septic foals. Initially a broad-spectrum bactericidal approach must be used, based on previous experiences
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Sepsis and costs. Antimicrobial therapy should begin immediately in any foal in which sepsis is suspected then altered if necessary when blood culture sensitivity data become available. A minimum therapeutic course of 2 weeks is recommended for bacteremic foals without localizing clinical signs. If localizing signs, such as pneumonia or septic arthritis, are present, a minimum course of 4 weeks is recommended.16 Based upon available data, combining amikacin with penicillin or ampicillin is an excellent initial therapeutic approach in a foal with adequate hydration. In the presence of severe hypovolemia and azotemia, a third-generation cephalosporin is a safer initial choice. With aminoglycoside therapy, serial creatinine monitoring every 2–3 days is recommended to monitor for potential renal side-effects. Cefotaxime is a good choice for foals with Gramnegative meningitis36 or those with unresponsive pneumonia. Due to significant resistance, trimethoprim/sulfa combinations should not be used in septic foals without documented sensitivity, and then only as a long-term option following initial parenteral therapy. In a recent Australian study, overall antimicrobial susceptibility was lower than previous reports, multi-drug resistance was common, and Gram-positive isolates demonstrated unpredictable antimicrobial susceptibility profiles.32 However, despite these findings, survival to hospital discharge was comparable to other reports. Attempted medical treatment options for systemic candidiasis include fluconazole, itraconazole, miconazole, voriconazole, and amphotericin B. Fungal sensitivity profiles, if available, may help direct therapy. Pharmacokinetic analyses of these drugs have not been established in foals. Amphotericin B has been administered but can cause potentially lifethreatening nephrotoxicity. Fluconazole has broadspectrum activity and is effective for many Candida spp. with the exception of all C. krusei and up to 15% of C. glabrata isolates. A dose of 5 mg/kg p.o. s.i.d. is recommended.37 Because many septic foals have concurrent failure of passive transfer, plasma transfusion is often clinically warranted. Plasma administration can increase neutrophil function8 and in one study specific anti-endotoxin antibody-enriched plasma improved survival relative to one commercially available hyperimmunized plasma in septic foals.39 Not surprisingly, many septic foals have detectable plasma endotoxin concentrations18 and upregulation of toll-like receptor 4 (TLR-4) gene expression in peripheral blood mononuclear cells.7 Recent in vitro work has also shown that beta-lactam antimicrobials appear more likely than aminoglycosides (alone or in combination with ampicillin) to in-
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duce endotoxemia and TNF-α activity during treatment of E. coli sepsis.3 Agents commonly used for the treatment of endotoxemia include flunixin meglumine, pentoxifylline, and polymixin B sulfate.40, 41 None of these agents has been scientifically evaluated for the treatment of endotoxemia in foals. Flunixin and polymixin B are potentially nephrotoxic, thus should be used with caution. Flunixin also has the potential for gastric ulceration. Pentoxifylline has been shown to reduce mortality without adverse effects in septic human neonates42 but not adults.43 Intravenous fluid therapy is often indicated in the treatment of neonatal sepsis. Fluid resuscitation is critical to correct hypovolemia associated with severe sepsis or septic shock. Initial therapy with a balanced ionic solution (such as lactated Ringer’s, NormosolR or Plasmalyte) avoids the acidifying effect of large volume replacement with no strong ion difference, such as saline.44 Initial provision of 20 mL/kg over 10–20 minutes followed by re-evaluation of perfusion allows for an efficient start to therapy, ideally with avoidance of fluid overload. When hypovolemia is severe or accompanied by hypoproteinemia, provision of an initial synthetic colloid (such as hydroxyethyl starch in sodium chloride) bolus (up to 10 mL/ kg) can be administered and followed with additional crystalloid replacement. Subsequent maintenance needs are then provided with either a replacement or maintenance crystalloid solution. For maintenance fluid requirements, one must take care to avoid sodium overload. One method of accomplishing this goal is to provide a combination of 5% dextrose with a balanced ionic solution. The total daily fluid requirement can vary depending upon the foal’s metabolic rate. Some sources suggest 100– 120 mL/kg/day as a starting point for most foals once plasma volume has been restored.45 For severely ill foals likely to have renal compromise, use of a ‘dry’ maintenance rate has been suggested.44 This formula is based upon a decreasing amount of fluid required per kg as the total bodyweight increases, which equates to approximately half of the previously suggested amount (2.5 L/day versus 5 L/day) for a 50 kg foal.44 Provision of exogenous glucose helps prevent catabolism. One can target 4 mg/kg/min of glucose, which equates to 240 mL/h of 5% dextrose to a 50 kg foal. Blood glucose concentration should be monitored periodically, and the rate of glucose supplementation can be increased up to 8 mg/kg/min if necessary, or therapy with insulin can be initiated for foals that become hyperglycemic.44 Additional electrolyte replacement can be provided if necessary. Restoration of plasma volume will typically resolve mild to moderate lactic acidosis; hence bicarbonate replacement should be reserved for those foals with
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ongoing bicarbonate loss, such as those with diarrhea, and instituted only when metabolic acidosis persists despite volume replacement. Arterial or venous lactate concentration and acid–base status, indirect arterial blood pressure, pulse oximetry, cardiac output, central venous pressure, and central venous oxygen saturation can provide additional information regarding volume status and estimation of tissue perfusion (see Figure 16.2). While extremely useful, many of these procedures are not very practical in field situations. Thus, physical parameters should be examined carefully and frequently. Of particular importance are development of edema, urine output, vital signs, and temperature of the distal limbs. Careful monitoring of urine output and development of edema, along with careful provision of maintenance fluids and sodium, will typically prevent overhydration. In the event of persistent hypotension (MAP < 60 mmHg) despite normovolemia, administration of inopressor therapy (dopamine, dobutamine, vasopressin, or norepinephrine) may be indicated. In such circumstances, indirect assessment of cardiac output (via ultrasonography) can be especially useful to monitor need for and response to therapy. The level of nursing care required for an individual septic foal depends upon the severity of disease. For those foals remaining predominantly recumbent, appropriate body positioning and padding is critical. Maintenance of sternal recumbency allows improved ventilation; appropriate padding and maintenance of well-positioned dry diapers and frequent turning will help prevent development of decubiti. Provision of appropriate complex caloric intake is critical to a successful outcome. Due to generalized and/or pharyngeal weakness or ineffective suckling frequently associated with sepsis, affected foals may not suckle from the mare efficiently enough to support their caloric requirement. Thus, intervention via the enteral or parenteral route is often necessary. Placement of an indwelling nasogastric or nasoesophageal tube allows administration of enteral nutritional support; bottle feeding should be avoided in sick foals with an ineffective suckle response. In foals with a functional gastrointestinal tract, supplementation with mare’s milk or replacer should begin at a somewhat conservative rate (such as a daily total of 5–10% of bodyweight divided into 12–24 feedings). If tolerated well, this amount can be gradually increased to a daily total of 25% of bodyweight over several days.46 Premature foals may require an even more conservative start. For those individuals with moderate to severe diarrhea or ileus, provision of parenteral nutrition allows for a period of gastrointestinal rest while maintaining appropriate caloric intake; alternatively, a combination of enteral and parenteral
Figure 16.2 Captured screen from a critical care monitor attached to a 36-hour-old foal. The foal was diagnosed with septic shock and neonatal encephalopathy and had been placed on mechanical ventilation. Spirometry data are noted along the left side of the screen. These include a pressure/volume loop (graph), peak airway pressure (Ppeak), plateau pressure (Pplat), mean airway pressure (Pmean), positive end expiratory pressure (PEEP), inspired and expired tidal volume (TVinsp, TVexp), inspired and expired minute ventilation (MVinsp, MVexp), inspiratory to expiratory ratio (I:E), compliance (Compl), and airway resistance (Raw). The traces shown include (top to bottom) direct arterial pressure, central venous pressure, oxygen saturation, and end tidal CO2 . End tidal (35) and inspired (44) percentages of oxygen are seen in the bottom right of the screen.
nutrition can allow a gradual increase in enteral feeding. Once the gastrointestinal tract becomes functional, the enteral route of feeding is again preferred. Careful monitoring of urine output and specific gravity, abdominal size, and daily bodyweight are useful to ensure appropriate intake; frequent monitoring of blood glucose concentration is indicated in foals receiving parenteral nutrition.
Focal infection and potential sequelae Pneumonia is a common focal infection in the septic foal, with a reported incidence ranging from 28%8 to 50%.47 Typically infection is thought to occur via hematogenous or inhalational spread of bacteria or via aspiration due to either bottle feeding or suckling from the mare despite generalized or pharyngeal weakness. Clinical signs may include fever, increased respiratory rate and effort, and abnormal thoracic auscultation. Cough and nasal discharge are rare in neonatal foals with pulmonary disease, and thoracic auscultation can be misleading. Cyanosis typically does not occur until Pa o2 decreases to a level of or below approximately 40 mmHg. Respiratory function is best assessed in septic foals via arterial blood gas analysis4, 48 while thoracic radiographs
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Sepsis and/or ultrasonography can provide an estimation of disease severity and distribution. In neonates with pneumonia, causative organisms are nearly identical to those associated with bacteremia discussed previously. Thus, antimicrobial therapy is the major form of therapy. If a given individual does not respond to initial therapy, identification of a causative organism via culture of a sample obtained via a guarded brush or transtracheal aspirate is indicated. Supplemental intranasal insufflation of humidified oxygen should be considered in foals with severe (Pa o2 < 55 mmHg or Sa o2 < 90% if older than 12–24 hours of age) or mild hypoxemia combined with evidence of impaired tissue oxygenation (hyperlactatemia, increased oxygen extraction ratio, or decreased oxygen uptake). Initially, arterial blood gas should be used to monitor response to therapy, with a target Pa o2 of 70–105 mmHg. Foals with severe or persistent hypercapnia (>65–77 mmHg), especially if accompanied by acidemia, may require mechanical ventilation. If hypercapnea is accompanied by hypoventilation, pharmacological stimulation of ventilation with doxapram may be effective.49, 50 One should note that Pa co2 is approximately 3–5 mmHg higher in laterally recumbent foals relative to those standing or in sternal recumbency,24 and hypercapnia can be exacerbated by fever or the administration of bicarbonate Diarrhea is also common in septic foals, with reported incidence between 16% and 38%.8, 16, 47, 51 In one study, foals with Actinobacillus sp. bacteremia were 6 times more likely to have diarrhea than those with other isolates.8 Conversely, 50% of foals <30 days of age with a primary presenting complaint of diarrhea were bacteremic at the time of admission in one report52 and a similar percentage (49%) of foals <10 months of age either presenting with diarrhea or developing diarrhea within the first 48 hours following admission were bacteremic if a blood culture was performed.53 In those studies, although the inclusion criteria differed substantially, the median age of both populations at the time of blood culture was 4 days. But, Gram-negative bacteria in general, and E. coli specifically, were more commonly isolated in the Florida study, whereas Enterococcus spp. were isolated more frequently in the Pennsylvania study.52, 53 In the Florida study, an association between a positive blood culture and the type of infectious agent identified in the feces was not evident.53 One must note that diarrhea may be associated with many problems other than bacteremia or sepsis in neonatal foals. Commonly identified infectious agents include Clostridium perfringens, Clostridium difficile, Salmonella spp., rotavirus, and various parasites. In one report, foals <1 month of age were more likely to have C. perfringens detected or to have
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negative fecal diagnostics relative to older foals; conversely, foals > 1 month of age were more likely to have Salmonella spp., rotavirus, and parasites detected in feces relative to younger foals.53 With or without enteritis, septic foals can also display signs of ileus and/or colic. Most of these problems resolve with symptomatic treatment and systemic improvement. In addition to broad-spectrum antimicrobial therapy, intravenous fluids and supportive care are typically indicated in septic foals with diarrhea. Additional therapy depends upon the identity (if any) of infectious organisms identified in feces of affected individuals. Development of diarrhea does not appear to have a negative impact upon survival and, in one report, was positively associated with survival. Neonates with diarrhea (whether or not they have bacteremia) typically carry a favorable prognosis.52, 53 In one report, foals with C. perfringens type C carried a more guarded prognosis,54 but in another report survival was not affected by the type of infectious agent (or lack thereof) identified in feces.53 Umbilical remnant infections are considered to be a common source of continued bacterial shedding. Ultrasonographic evaluation of these structures is critical, as external signs (pain, heat, and swelling) are frequently absent. The recommended frequency of repeat ultrasonographic examinations is dependent upon the individual foal’s clinical progression. Treatment options include long-term antibiotic therapy or surgical resection. Patent urachus can develop without involvement of other structures, with a reported incidence of 21%,51 and will often resolve with antimicrobial therapy. Uroperitoneum has been reported to occur in 2.5% of hospitalized neonates.55 Presumptively septic foals receiving fluid therapy appear to be older and less likely to have the classic electrolyte abnormalities associated with uroperitoneum than foals diagnosed at the time of admission, suggesting that ill foals are typically diagnosed earlier, but the condition occurred later in life.55, 56 Orthopedic infections represent one of the most important life-threatening and performance-limiting complications associated with sepsis. The reported incidence of infective synovitis ranges from 26% to 33%8, 51, 57 and that of osteomyelitis from 11% to 12%.51, 57 Clinical signs include lameness, joint effusion, and pain on palpation of physes; any such abnormality in a neonate should be considered septic until proven otherwise. A suspected diagnosis can be confirmed via arthrocentesis (synovitis) or imaging (septic physitis or osteomyelitis). Ultrasonography, radiography, and cross-sectional imaging (computed tomography or magnetic resonance imaging) can provide additional information regarding lesion extent and severity. Aggressive lavage of an affected
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joint and regional perfusion with appropriate antimicrobials can provide adjunctive therapy. Development of orthopedic infections appears to worsen both the prognosis for short-term survival31 and the potential for athletic performance.58, 59 Meningitis is a rare but extremely serious complication. Major clinical signs include seizures and severe depression, although the latter is somewhat difficult to assess in a severely compromised foal. Other signs include head tilt, strabismus, nystagmus, and extensor rigidity, depending upon the areas of involvement. CSF culture and cytology will confirm the diagnosis, as affected foals will typically have a neutrophilic pleocytosis. Prognosis is poor to grave, but if therapy is attempted, a third-generation cephalosporin (such as cefotaxime and ceftriaxone but not ceftiofur) has been recommended.36 The major differential diagnosis for neurological signs in a septic neonate is hypoxic ischemic encephalomyelopathy (HIE). Typically foals with HIE present within 24–48 hours following birth, whereas the age of foals with meningitis is more variable. The most common ocular complications in the septic foal are corneal ulceration and uveitis. Ulceration can occur due to entropion or trauma. Because foals often do not display blepharospasm or epiphora despite ulceration, routine daily ophthalmological examination with fluorescein staining is important for diagnosis. In foals remaining recumbent for prolonged periods of time, ocular lubrication may prevent development of corneal ulceration. When uveitis occurs, it is typically an ocular extension of the systemic disease process and may be immune-mediated or infectious. Clinical signs may include miosis, hypopyon, corneal edema or deep vascularization, and changes in iris color (often a yellow-green hue). Severely affected foals may have hypotony (softness of the globe). Systemic therapy should include antimicrobial, analgesic, and anti-inflammatory medication; topical ocular therapy is dependent upon whether or not the cornea is intact, and may include mydriatics (e.g., atropine), antimicrobials with good corneal penetration (e.g., chloramphenicol or triple antibiotic), corticosteroids (e.g., prednisolone acetate), non-steroidal anti-inflammatory drugs, osmotic agents, or anticollagenase therapy (e.g., serum or EDTA).60 Disorders of coagulation can occur in septic foals, manifested clinically by either hemorrhage or thromosis. The most common abnormality is jugular venous thrombosis, but brachial, digital, metatarsal, and metacarpal arterial and diffuse vascular thromboses throughout the distal limb (see Figure 16.3), aortic termination, lungs, and colon have been reported.18,61–64 In a given individual, manual platelet count, PT/PTT, or thromboelastography may pro-
Figure 16.3 Dissected forelimb of a neonatal foal previously diagnosed with sepsis. Note thrombosis of the medial palmar artery as it branches into the medial and lateral palmar digital arteries, which are also thrombosed.
vide additional information regarding coagulation status.
Prognosis/outcomes Most retrospective reports cite short-term survival in the range of 45–60%,8, 28, 31, 39, 51, 65 but it has ranged from as low as 25%19 to 32%18 to as high as 70–72%.30, 32, 47 Various factors have been associated with shortterm survival including a negative association between infection with Gram-negative organisms and survival,18 an inverse correlation between duration of illness prior to admission and survival but a positive correlation between a foal’s ability to stand on admission and survival.51 At the University of Georgia, sepsis score <11, negative blood culture at admission, blood glucose ≥ 60 mg/dL, body temperature ≥ 100◦ F, Tco2 > 15 mmol/L, and a low or normal fibrinogen were positive predictors of survival.28 Another study found that normal arterial lactate concentration (<2.5 mmol/L) either at admission or 18–36 hours later was a good predictor of survival, whereas elevated lactate was not a good predictor of non-survival.22 Increasing age at admission, septic arthritis, band neutrophils, and creatinine were negatively associated with survival in bacteremic foals in a Florida study, whereas year of admission, diarrhea, temperature, segmented neutrophils, and arterial pH were positively associated with survival.31 Interestingly, bacteremia was not associated with survival in two reports of foals with diarrhea.52, 53 Few studies have addressed the long-term survival and performance of neonatal intensive care unit survivors, much less septic foals specifically. In one study of bacteremic foals, racing performance data were evaluated for 102 surviving Thoroughbred foals
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Sepsis and 194 of their maternal siblings and no significant difference was demonstrated between foals and siblings in percent starters, percent winners, or number of starts, but affected foals had significantly lower number of wins, total earnings, and standard starts index (SSI) than siblings.31 This is somewhat in contrast to data reported for overall neonatal intensive care unit survivors, where the percentage of starters was lower than the control population, but performance over a 2-year period was not different in those animals able to make at least two starts.66 Similar findings have been reported for foals with neurological disease67 and those with R. equi pneumonia.68
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Clearly, given the wide range of potentially devastating problems associated with sepsis, prevention will always outweigh treatment, but no one program will ever totally eliminate the risk of sepsis. For example, in a recent survey of 36 stud farms (1031 foals) in Newmarket, UK, the practice of prophylactic administration of antimicrobials for 3 days following birth did not significantly alter the incidence of infectious or non-infectious disorders.69 Thus, the most important recommendations include the following. A clean environment should be maintained for the foal during udder seeking. In order to facilitate this goal, foaling stalls should be thoroughly cleaned and disinfected between mares and the mare’s hindquarters, perineum, and udder should be thoroughly cleaned then dried prior to the foal’s introduction. Ideally, this should be done just outside the stall to prevent environmental contamination. The foal should suckle quickly (i.e., within the first 2 hours). If a given foal is too weak to accomplish this task on its own, it should be fed via nasogastric tube. Weak foals should not be force-fed or syringe-fed, as this practice may result in aspiration pneumonia. In addition, failure to suckle quickly should indicate a potential problem. Adequate transfer of passive immunity should be ensured by measurement of IgG concentration in serum, plasma, or whole blood and administration of plasma, if necessary. Finally, appropriate umbilical care should be ensured. In a review of published studies in human neonates, 4% chlorhexidine consistently reduced the risk of umbilical and periumbilical infections,70 but such an evaluation is not currently available for the foal.
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References 1. Bone RC, Balk RA, Cerra FB, Dellinger RP, Fein AM, Knaus WA, Schein RM, Sibbald WJ. Definitions for sepsis and organ failure and guidelines for the use of innovative therapies in sepsis. The ACCP/SCCM Consensus Conference
20.
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Committee. American College of Chest Physicians/Society of Critical Care Medicine. Chest 1992;101:1644–55. Moore JN, Morris DD. Endotoxemia and septicemia in horses: experimental and clinical correlates. J Am Vet Med Assoc 1992;200:1903–14. Bentley AP, Barton MH, Lee MD, Norton NA, Moore JN. Antimicrobial-induced endotoxin and cytokine activity in an in vitro model of septicemia in foals. Am J Vet Res 2002;63:660–8. Paradis MR. Update on Neonatal Septicemia. Vet Clin N Am Equine Pract 1994;10:109–35. Marsh PS, Palmer JE. Bacterial isolates from blood and their susceptibility patterns in critically ill foals: 543 cases (1991–1998). J Am Vet Med Assoc 2001;218:1608–10. Pusterla N, Magdesian KG, Mapes S, Leutenegger CM. Expression of molecular markers in blood of neonatal foals with sepsis. Am J Vet Res 2006;67:1045–9. Gold JR, Perkins GA, Erb HN, Ainsworth DM. Cytokine profiles of peripheral blood mononuclear cells isolated from septic and healthy neonatal foals. J Vet Intern Med 2007;21:482–8. Stewart AJ, Hinchcliff KW, Saville WJA, Jose-Cunilleras E, Hardy J, Kohn CW, Reed SM, Kowalski JJ. Actinobacillus sp. bacteremia in foals: clinical signs and prognosis. J Vet Intern Med 2002;16:464–71. Wohlfender FD, Barrelet FE, Doherr MG, Straub R, Meier HP. Diseases in neonatal foals. Part 2: potential risk factors for a higher incidence of infectious diseases during the first 30 days post partum. Equine Vet J 2009;41:186–91. Raidal SL. The incidence and consequences of failure of passive transfer of immunity on a thoroughbred breeding farm. Aust Vet J 1996;73:201–6. Tyler-McGowan CM, Hodgson JL, Hodgson DR. Failure of passive transfer in foals: incidence and outcome on four studs in New South Wales. Aust Vet J 1997;75:56–9. Robinson JA, Allen GK, Green EM, Fales WH, Loch WE, Wilkerson CG. A prospective study of septicaemia in colostrum-deprived foals. Equine Vet J 1993;25:214–19. Clabough DL, Levine JF, Grant GL, Conboy HS. Factors associated with failure of passive transfer of colostral antibodies in Standardbred foals. J Vet Intern Med 1991;5:335–40. Baldwin JL, Cooper WL, Vanderwall DK, Erb HN. Prevalence (treatment days) and severity of illness in hypogammaglobulinemic and normogammaglobulinemic foals. J Am Vet Med Assoc 1991;198:423–8. Madigan JE. Method for preventing neonatal septicemia, the leading cause of death in the neonatal foal. Proc Am Assoc Equine Pract 1997;43:17–19. Brewer BD. Neonatal infection. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 295–316. Harvey JW, Asquith RL, McNulty PK, Kivipelto J, Bauer JE. Haematology of foals up to one year old. Equine Vet J 1984;16:347–53. Barton MH, Morris DD, Norton N, Prasse KW. Hemostatic and fibrinolytic indices in neonatal foals with presumed septicemia. J Vet Intern Med 1998;12:26–35. Koterba AM, Brewer BD, Tarplee FA. Clinical and clinicopathological characteristics of the septicaemic neonatal foal: review of 38 cases. Equine Vet J 1984;16:376–82. Gold JR, Divers TJ, Barton MH, Lamb SV, Place NJ, Mohammed HO, Bain FT. Plasma adrenocorticotropin, cortisol, and adrenocorticotropin/cortisol ratios in septic and normal-term foals. J Vet Intern Med 2007;21:791–6.
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21. Hurcombe SD, Toribio RE, Slovis N, Kohn CW, Refsal K, Saville W, Mudge MC. Blood arginine vasopressin, adrenocorticotropin hormone, and cortisol concentrations at admission in septic and critically ill foals and their association with survival. J Vet Intern Med 2008;22:639–47. 22. Corley KT, Donaldson LL, Furr MO. Arterial lactate concentration, hospital survival, sepsis and SIRS in critically ill neonatal foals. Equine Vet J 2005;37:53–9. 23. Corley KTT, Furr MO. Evaluation of a score designed to predict sepsis in foals. J Vet Emerg Crit Care 2003;13:149–55. 24. Koterba AM. Respiratory disease: approach to diagnosis. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 153–76. 25. Armengou L, Monreal L, Tarancon I, Navarro M, Rios J, Segura D. Plasma D-dimer concentration in sick newborn foals. J Vet Intern Med 2008;22:411–17. 26. Wilson WD, Madigan JE. Comparison of bacteriologic culture of blood and necropsy specimens for determining the cause of foal septicemia: 47 cases (1978–1987). J Am Vet Med Assoc 1989;195:1759–63. 27. Brewer BD, Koterba AM, Carter RL, Rowe ED. Comparison of empirically developed sepsis score with a computer generated and weighted scoring system for the identification of sepsis in the equine neonate. Equine Vet J 1988;20:23–4. 28. Henson S, Barton MH. Bacterial isolates and antibiotic sensitivity patterns from septicemic neonatal foals: a 15 year retrospective study (1986–2000). In: Proceedings of Dorothy Havemeyer Foundation Neonatal Workshop 3 2001; pp. 50–2. 29. Brewer BD, Koterba AM. Development of a scoring system for the early diagnosis of equine neonatal sepsis. Equine Vet.J. 1988;20: 18–22. 30. Raisis AL, Hodgson JL, Hodgson DR. Equine neonatal septicaemia: 24 cases. Aust Vet J 1996;73:137–40. 31. Sanchez LC, Giguere S, Lester GD. Factors associated with survival of neonatal foals with bacteremia and racing performance of surviving Thoroughbreds: 423 cases (1982–2007). J Am Vet Med Assoc 2008;233:1446–52. 32. Russell C, Axon J, Blishen A, Begg A. Blood culture isolates and antimicrobial sensitivities from 427 critically ill neonatal foals. Aust Vet J 2008;86:266–71. 33. McClure JJ, Addison JD, Miller RI. Immunodeficiency manifested by oral candidiasis and bacterial septicemia in foals. J Am Vet Med Assoc 1985;186:1195–7. 34. Reilly LK, Palmer JE. Systemic candidiasis in four foals. J Am Vet Med Assoc 1994;205:464–6. 35. Perkins G, Ainsworth DM, Erb HN, Del Piero F, Miller M, Wilkins PA, Palmer J, Frazer M. Clinical, haematological and biochemical findings in foals with neonatal Equine herpesvirus-1 infection compared with septic and premature foals. Equine Vet J 1999;31:422–6. 36. Morris DD, Rutkowski J, Lloyd KC. Therapy in two cases of neonatal foal septicaemia and meningitis with cefotaxime sodium. Equine Vet J 1987;19:151–4. 37. Giguere S. Antifungal chemotherapy. In: Giguere S, Prescott JF, Baggot JD, Walker RD, Dowling PM (eds) Antimicrobial Therapy in Veterinary Medicine. Ames, IA: Blackwell, 2006; pp. 301–22. 38. McTaggart C, Penhale J, Raidala SL. Effect of plasma transfusion on neutrophil function in healthy and septic foals. Aust Vet J 2005;83:499–505. 39. Peek SF, Semrad S, McGuirk SM, Riseberg A, Slack JA, Marques F, Coombs D, Lien L, Keuler N, Darien BJ. Prognostic value of clinicopathologic variables obtained at admission
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and effect of antiendotoxin plasma on survival in septic and critically ill foals. J Vet Intern Med 2006;20:569–74. Baskett A, Barton MH, Norton N, Anders B, Moore JN. Effect of pentoxifylline, flunixin meglumine, and their combination on a model of endotoxemia in horses. Am J Vet Res 1997;58:1291–9. Durando MM, MacKay RJ, Skelley LA. Effects of polymyxin B and Salmonella typhimurium antiserum on horses given endotoxin intravenously. Am J Vet Res 1994;55:921–7. Haque K, Mohan P. Pentoxifylline for neonatal sepsis. Cochrane Database Syst Rev 2003; CD004205. Staubach KH, Schroder J, Stuber F, Gehrke K, Traumann E, Zabel P. Effect of pentoxifylline in severe sepsis: results of a randomized, double-blind, placebo-controlled study. Arch Surg 1998;133:94–100. Palmer JE. Fluid therapy in the neonate: not your mother’s fluid space. Vet Clin N Am Equine Pract 2004;20:63–75. Spurlock SL, Furr M. Fluid therapy. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 671–700. Buechner-Maxwell VA. Nutritional support for neonatal foals. Vet Clin N Am Equine Pract. 2005;21:487–510, viii. Freeman L, Paradis MR. Evaluating the effectiveness of equine neonatal care. Vet Med 1992;87:921–6. Magdesian KG. Monitoring the critically ill equine patient. Vet Clin N Am Equine Pract 2004;20:11–39. Giguere S, Sanchez LC, Shih A, Szabo NJ, Womble AY, Robertson SA. Comparison of the effects of caffeine and doxapram on respiratory and cardiovascular function in foals with induced respiratory acidosis. Am J Vet Res 2007;68:1407–16. Giguere S, Slade JK, Sanchez LC. Retrospective comparison of caffeine and doxapram for the treatment of hypercapnia in foals with hypoxic-ischemic encephalopathy. J Vet Intern Med 2008;22:401–5. Gayle JM, Cohen ND, Chaffin MK. Factors associated with survival in septicemic foals: 65 cases (1988–1995). J Vet Intern Med 1998;12:140–6. Hollis AR, Wilkins PA, Palmer JE, Boston RC. Bacteremia in equine neonatal diarrhea: a retrospective study (1990–2007). J Vet Intern Med 2008;22(5):1203–9. Frederick J, Giguere S, Sanchez LC. Infectious agents detected in the feces of diarrheic foals: a retrospective study of 233 cases (2003–2008). J Vet Intern Med 2009;23: 1254–60. East LM, Savage CJ, Traub-Dargatz JL, Dickinson CE, Ellis RP. Enterocolitis associated with Clostridium perfringens infection in neonatal foals: 54 cases (1988–1997). J Am Vet Med Assoc 1998;212:1751–6. Kablack KA, Embertson RM, Bernard WV, Bramlage LR, Hance S, Reimer JM, Barton MH. Uroperitoneum in the hospitalised equine neonate: retrospective study of 31 cases, 1988–1997. Equine Vet J 2000;32:505–8. Dunkel B, Palmer JE, Olson KN, Boston RC, Wilkins PA. Uroperitoneum in 32 foals: influence of intravenous fluid therapy, infection, and sepsis. J Vet Intern Med 2005;19:889–93. Paradis MR. Septic arthritis in the equine neonate: a retrospective study. Proc ACVIM Forum 1991;9:458–9. Smith LJ, Marr CM, Payne RJ, Stoneham SJ, Reid SW. What is the likelihood that Thoroughbred foals treated for septic arthritis will race? Equine Vet J 2004;36:452–6. Steel CM, Hunt AR, Adams PL, Robertson ID, Chicken C, Yovich JV, Stick JA. Factors associated with prognosis for
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survival and athletic use in foals with septic arthritis: 93 cases (1987–1994). J Am Vet Med Assoc 1999;215:973–7. Dwyer AE. Beginning to see the light: pediatric equine ophthalmology. In: Proceedings of the American Association of Equine Practitioners: focus on the first year of life. 2008. Brianceau P, Divers TJ. Acute thrombosis of limb arteries in horses with sepsis: five cases (1988–1998). Equine Vet J 2001;33:105–9. Forrest LJ, Cooley AJ, Darien BJ. Digital arterial thrombosis in a septicemic foal. J Vet Intern Med 1999;13:382–5. Moore LA, Johnson PJ, Bailey KL. Aorto-iliac thrombosis in a foal. Vet Rec 1998;142:459–62. Triplett EA, O’Brien RT, Wilson DG, Steinberg H, Darien BJ. Thrombosis of the brachial artery in a foal. J Vet Intern Med 1996;10:330–2. Brewer BD, Koterba AM. Bacterial isolates and susceptibility patterns in foals in a neonatal intensive-care unit. Comp Cont Educ Pract Vet 1990;12:1773–81. Axon J, Palmer J, Wilkins P. Short- and long-term athletic outcome of neonatal intensive care unit survivors. Proc Am Assoc Equine Pract 1999;45:224–5.
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67. Bryant JE, Bernard W, Wilson WD, Gardner I. Race earnings as an indicator of future performance in neonatal foals treated for neurological disorders. Proc Am Assoc Equine Pract 1994;40:198. 68. Ainsworth DM, Eicker SW, Yeagar AE, Sweeney CR, Viel L, Tesarowski D, Lavoie JP, Hoffman A, Paradis MR, Reed SM, Erb HN, Davidow E, Nalevanko M. Associations between physical examination, laboratory, and radiographic findings and outcome and subsequent racing performance of foals with Rhodococcus equi infection: 115 cases (1984–1992). J Am Vet Med Assoc 1998;213: 510–15. 69. Wohlfender FD, Barrelet FE, Doherr MG, Straub R, Meier HP. Diseases in neonatal foals. Part 1: the 30 day incidence of disease and the effect of prophylactic antimicrobial drug treatment during the first three days post partum. Equine Vet J 2009;41:179–85. 70. Mullany LC, Darmstadt GL, Tielsch JM. Role of antimicrobial applications to the umbilical cord in neonates to prevent bacterial colonization and infection: a review of the evidence. Pediatr Infect Dis J 2003;22:996– 1002.
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Section (e): Specific Diagnostic and Management Techniques
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Critical Care – Assessment Jane E. Axon
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Critical Care – Monitoring Darien J. Feary
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Critical Care – Treatment Jane E. Axon
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Complications of Neonatal Infection Mary Rose Paradis
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Fluid Therapy Robert P. Franklin
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Respiratory Assistance Jonathan E. Palmer
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Sedation, Anesthesia and Analgesia Lori A. Bidwell
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Stabilization and Preparation for Transport Robert P. Franklin and Jennifer R. Read
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Transfusion Techniques John F. Freestone
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Diagnostic Ultrasonography of the Abdomen Virginia B. Reef and JoAnn Slack
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Antibiotic Therapy K. Gary Magdesian
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Enteral and Parenteral Nutrition for the Neonatal Foal Wendy E. Vaala
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Growth of Horses Clarissa Brown-Douglas, Peter Huntington and Joe Pagan
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Important Gastrointestinal Parasites Eugene T. Lyons, Mariana Ionita and Sharon C. Tolliver
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Vaccination of Mares, Foals and Weanlings W. David Wilson
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Chapter 17
Critical Care – Assessment Jane E. Axon
Early recognition of the subtle changes that are seen in the critically ill neonate is paramount for a successful outcome. The examination technique should always follow the same routine so it is complete and no abnormalities are missed. Many foals examined in the emergency setting are in cardiovascular and respiratory collapse, thus initially a cursory evaluation is performed while treatment is being instituted. A thorough physical examination is performed after the foal has been stabilized. Examination of the neonate is incomplete without obtaining the mare’s periparturient and previous reproductive history and performing a physical examination of the mare with particular emphasis on her udder, reproductive tract, and placenta. In a non-emergency setting, watching the foal’s behavior unattended can assist with evaluating the degree of compromise. Healthy foals are bright and inquisitive. The first signs of neonatal weakness are a lower head carriage with ears drooping, and falling asleep standing. Sicker foals spend an increased time recumbent, less time nursing, and stand under the mare not nursing so the dam’s udder becomes distended and the inside of the mare’s legs and the foal’s face becomes covered with milk that then dries. The affinity of the foal for the dam and the foal’s nursing capability should be assessed. Foals may look as though they are nursing, but on closer inspection they are not nursing effectively and do not have a proper tongue seal around the teat (Figure 17.1). Loss of affinity for the mare in an otherwise bright foal is often seen with hypoxic ischemic syndrome (HIS). Disinterest in nursing and loss of udder seeking can be seen with meconium impaction. Inability to nurse may be associated with neurological dysfunction (most commonly HIS), other systemic illness, or oral or physical problems. The shape of the teats should also be evaluated; very small teats in maiden mares with an edematous udder can be the cause of a foal’s inability to nurse. Maiden mares may
also be less willing to let the foal nurse or let milk down, thus may predispose the foal to failure of passive transfer. Rectal temperature is not a reliable indicator of infection in neonates. Foals with acute sepsis are often hypothermic and premature foals have poor thermoregulatory control.
General body appearance A cursory visual examination of the foal can help determine if the foal is premature or has had intrauterine growth retardation (IUGR). Premature foals classically are small with floppy ears, silky hair coat, domed forehead, and increased tendon and joint laxity. Foals with IUGR are usually thin, small for gestational age, have a long hair coat, and may have increased tendon and joint laxity. Skin abrasions associated with increased recumbency and attempts to stand are often present on the elbows, around the orbits and musculoskeletal prominences and are often first detected as discrete areas of moisture (Figure 17.2). The ventrum, axilla region, and perineum should be assessed for the presence of edema associated with fluid overload and possible renal dysfunction. Swelling under the axilla can also be seen with fractured ribs, cellulitis or trauma; ventral swelling can be seen with urine accumulation from a torn urachus or cellulitis from overzealous umbilical disinfection. Skin tenting as an assessment of hydration is not as reliable in foals as in adults. Sunken eyes may be a sign of dehydration but also may be associated with IUGR.
Cardiovascular system Normal mucous membranes should be pink, moist and have a capillary refill time of 1–2 seconds. Hyperemic and injected mucous membranes accompanied by scleral injection and hyperemic coronary bands
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 17.1 A foal with a normal tongue seal around the mare’s teat.
Figure 17.2 Discrete areas of moisture on the skeletal prominences of the hind leg indicative of increased recumbency.
are classic findings in early sepsis. Jaundiced mucous membranes are most commonly associated with sepsis rather than neonatal isoerythrolysis and can also be seen with internal hemorrhage, meconium retention, equine herpesvirus-1 infection or liver disease. Petechial hemorrhages on mucosal surfaces or within the pinnae are seen with sepsis or thrombocytopenia with disseminated intravascular coagulation or alloimmune thrombocytopenia. Pale mucous membranes suggest anemia which can be due to umbilical cord hemorrhage, internal hemorrhage from fractured ribs or torn internal umbilical remnants, gastrointestinal or urinary tract hemorrhage, or neonatal isoerythrolysis. Gray or cyanotic membranes are indicative of shock, poor peripheral perfusion, or hypoxemia. Cyanotic mucous membranes should prompt further investigation of the cardiovascular and respiratory systems. The heart rate of a foal at birth is 40–80 bpm. This should increase with attempts to stand and will increase to 120–150 bpm or higher over the next few hours and then stabilize at around 80–100 bpm within the first week of life.1 The causes of persistent tachycardia in the neonate are extensive; however, it usually reflects adaptive mechanisms to hypovolemic or septic shock, severe anemia, pain, hypoxemia, and/or stress. Persistent bradycardia can be due to hypoxemia, electrolyte abnormalities, hypoglycemia and/or hypothermia, and is usually indicative of more serious compromise. Transient arrhythmias occur relatively frequently in the postpartum period. In a study of 101 foals, 91% had a transient arrhythmia at birth.2 Atrial premature contraction and sinus arrhythmia were the most common arrhythmias. The longest duration of an arrhythmia (atrial fibrillation) was just over 2 hours; however, most disappeared within 15 minutes. The Pa O2 value in foals with severe arrhythmias was comparatively lower than that of foals with mild arrhythmias. It was suggested that hypoxemia played an important role in neonatal arrhythmias.2 The clinical significance of a persistent arrhythmia should take into consideration clinical, metabolic, and hemodynamic findings and can be associated with hypoxemia, electrolyte abnormalities, and sepsis. Persistently abnormal heart rates and arrhythmias should be investigated further with an electrocardiogram. Murmurs are not uncommon in the newborn period. A continuous machinery or systolic murmur associated with the closing ductus arteriosus may be heard up to 4 days of age over the left heart base.3 A study on 10 pony foals found that continuous murmurs were not detected after 3 days of age and systolic murmurs were still detected in 7-week-old foals.4 Color flow Doppler echocardiography in these foals found no consistent association between flow
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Critical Care – Assessment through the foramen ovale and ductus arteriosus and the presence of a systolic murmur, thus suggesting that the murmurs auscultated over the left heart base are attributed to left ventricular ejection.5 Murmurs that are present after 7 days of age or that are associated with hypoxemia or exercise intolerance need further evaluation. Serious heart abnormalities may not be accompanied by any murmur. Echocardiography can be used to evaluate murmurs, any suspected cardiac causes of cyanosis or congenital anomalies.5 Arterial pulses are usually detected in the brachial and great metatarsal artery. No jugular pulse should be visible regardless of state of recumbency. Distal extremities should be warm, and if cold are often associated with poor pulses and perfusion. Hypotension is often, but not always, associated with palpably weak pulses. Conversely, poor pulse quality does not necessarily indicate hypotension.6 The normal mean arterial blood pressure (MAP) of a thoroughbred foal is 69–111 mmHg;7 however, normal tissue perfusion can occur at lower values. Decreased MAP can result from hypovolemia, decreased systemic vascular resistance, or reduced cardiac output. A clinical impression of tissue perfusion can be obtained from urine output, mental attitude, and lactate concentration and should be used in conjunction with interpretation of the MAP.
Respiratory system A cough, nasal discharge, and abnormal lungs sounds on auscultation are not commonly seen in neonates with early pneumonia; thus subtle changes in effort and rate and other diagnostic aids become important in evaluating the extent of lower airway disease. Severe respiratory distress can occur shortly after birth associated with congenital respiratory or cardiac abnormalities. It can be seen with a dorsally displaced soft palate from anatomic or neurological dysfunction. The respiratory rate in the newborn foal is 60–80 bpm and decreases to 30 bpm within the first hour of birth. There are many causes of tachypnea, which can be pulmonary or non-pulmonary in origin. The pattern of breathing should be regular when the foal is awake. It is normal when the foal is asleep to have irregular breathing patterns. These irregularities occur in rapid eye movement sleep and are often accompanied by rapid eye movements, body twitching, and vocalization. If however the foal is not asleep irregular patterns, in particular periodic apnea and slow rates, can be associated with CNS disease, metabolic and electrolyte abnormalities, prematurity, and
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Figure 17.3 Paradoxical respiration in a premature foal. The soft ribs and weak musculature result in the ribs being drawn in during increased inspiratory effort.
hypothermia. Increased respiratory effort is often the first sign a foal with respiratory disease will exhibit. As the respiratory disease progresses and the effort increases, the foal may develop paradoxical breathing due to respiratory muscle fatigue (Figure 17.3). This is a common complication in premature foals with pneumonia and weak respiratory musculature. Thoracic auscultation is not a reliable indicator of lung disease in the neonate. Severe lung pathology is often present before changes on auscultation are heard. Often referred upper airway noise is auscultated or an upper respiratory stridor is heard with neurological dysfunction (due to HIS) of the soft palate or less commonly with congenital anomalies such as subepiglottic cyst, stenotic nares or tracheal collapse. The thorax should be palpated to determine if fractured ribs or costochondral dislocations are present. Crepitus is usually, but not always, felt on palpation. Sometimes the first indication of a fractured rib is a faint ‘clicking’ that is heard on thoracic auscultation, or edema around the fracture site. Affected foals may grunt with expiration and may prefer to lie on the unaffected side. Arterial blood gas Arterial blood gas analysis is the most sensitive indication of respiratory function that is readily available to the practitioner.8 Hypoxemia (Pa O2 < 70 mmHg) can result from hypoventilation, ventilation/perfusion mismatch, or right-to-left shunting of blood (intrapulmonary or intracardiac).8 Hypoxemia can occur with hypo-, normo- or hypercapnia. Hypocapnia or normocapnia can occur with hyperventilation, impaired gas diffusion (pulmonary
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edema, pneumonia) or right-to-left shunting. Hypercapnia with a decreased Pa O2 is usually indicative of respiratory failure. The Pa CO2 results, however, need to be interpreted with consideration of the pH and bicarbonate concentrations as hypercapnia can occur in response to metabolic alkalosis.9 Diagnostic imaging The ultrasonographic appearance of pathological changes in the foal’s thorax is similar to adults. Fractured ribs can be identified by a disruption of the cortical surface of the bone and surrounding edema. Pleural fluid may appear cellular with an infectious process or a hemothorax which may exhibit the classic ‘smoke whirls’. Anechoic fluid can be seen with uroperitoneum, infectious process, or congenital anomaly. Ultrasonographic findings in pulmonary parenchymal disease are usually non-specific and, like radiographs, are not indicative of the specific etiology. Radiographs are useful in the diagnosis of diaphragmatic hernias, fractured ribs, and pneumothorax. Although radiographic findings are not specific for pulmonary disease the images do provide information regarding the type and extent of disease. Transtracheal aspiration The author does not routinely perform transtracheal aspiration (TTA) on a neonate with suspected pneumonia until the neonate has been stabilized, as the excessive stress of the procedure on an already compromised neonate can have fatal consequences. Care must also be taken if the procedure is performed in a foal with an elevated respiratory rate as subcutaneous emphysema and subsequent pneumomediastinum can occur.
Gastrointestinal system The mouth should be examined for presence of a cleft palate, prognathism, or brachygnathism, and the tongue examined for the presence of Candida which varies in appearance from small white plaques to an extensive thick tan discoloration over the tongue (Figure 17.4). The presence of Candida can be an indication of an ill-thrifty foal or systemic infection in the critically ill neonate, and can be a cause of increased salivation and reduced nursing. Auscultation of the abdomen provides an assessment of the gastrointestinal motility; however, the presence or absence of sounds is not diagnostic. The changes in the auscultation findings over time are more useful in assisting with a diagnosis. A silent abdomen which progresses to hypermotile sounds can
Figure 17.4 Severe candidiasis. Note the area of hyperemia at the edge of the plaque which can result in salivation and reluctance to nurse.
be indicative of enteritis; audible sounds which then become silent can suggest a surgical lesion. Transcutaneous palpation of the abdomen can be performed on recumbent, relaxed neonates. Meconium impactions, intussusceptions, and hematomas of the umbilical remnants can be palpated. The inguinal rings, scrotum, and umbilicus should be palpated for any evidence of enlargement, herniation, or infection. Ballottement of the abdomen can detect fluid waves which occur with excessive peritoneal fluid. A tympanic sound on percussion is indicative of gas accumulation. Abdominal distension can be monitored using a tape to measure the abdominal circumference, marking the site of measurement with a small shaved area. A ‘sprung’ appearance to the left caudal ribcage can be detected in neonatal foals with an enlarged stomach, which may be due to ileus or very recent nursing. Digital examination of the rectum with a welllubricated finger is very useful to detect a distal meconium impaction or intestinal atresia. However, not all neonates with a meconium impaction will have digitally palpable meconium. If there is no evidence of fecal material and there is clear or white cloudy mucus on the glove, intestinal atresia should be suspected. Digital examination is also useful in older foals which may be impacted from gravel or sand ingestion or a perirectal abscess. It is important to attempt to obtain nasogastric reflux on all critically ill neonates and foals with colic. Decompression assists with pain relief and the presence of reflux is indicative of a mechanical or functional small intestinal obstruction or gastric emptying disorder. Mechanical obstructions are more common in older foals. Functional obstructions
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Critical Care – Assessment are common in younger foals associated with ileus from sepsis, HIS, prematurity, enteritis, overfeeding, and electrolyte abnormalities. Diagnostic imaging Ultrasonographic examination is an essential part of the diagnostic evaluation of the foal’s abdomen.10, 11 Some conditions such as uroperitoneum, hemoperitoneum, intussusception, strangulating obstruction, meconium impaction, hernias, and enteritis can be tentatively diagnosed with the use of ultrasound. The ultrasound findings should always be interpreted in conjunction with clinical signs. Abdominal radiography may assist in determining the location, not necessarily the etiology, of gas and fluid distension and identify the composition of ingesta in the gastrointestinal tract. The reliance on radiographic examination has decreased with the increasing use of ultrasonographic evaluation of the neonatal abdomen. The author however still utilizes contrast radiography to assist with documenting gastric outflow obstructions, intestinal atresia, and evaluating the urinary system. Abdominocentesis Abdominocentesis should be performed with extreme caution in a foal as an enterocentesis is more likely to occur. Ultrasonographic examination can be used to assist with locating pockets of fluid for sampling. If no fluid is seen during an ultrasonographic examination, an abdominocentesis should not be performed.
Neurological system The neurological examination should be evaluated with respect to other physical examination findings. Many critically ill neonates are depressed, recumbent, and unable to suckle. These findings can be associated with a primary neurological disease but are more commonly seen secondary to underlying disease processes such as sepsis. Examination of the cranial nerves in neonates is similar to an adult, though some normal variations are present.12 The pupillary light reflex (direct and consensual) is present shortly after birth; however, it may be sluggish in excited foals. The menace response is not developed in foals until several weeks of age; however, the foal should withdraw its head away from the menacing gesture. Nystagmus should not be present when the head is still. Vision in a foal is difficult to evaluate due to adaptation of the foal and use of the mare as its ‘eyes’. It can be assessed by observation of the foal’s behavior, navigation through
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obstacles (especially when the foal is removed from the mare’s side), and menace response. The ability to swallow can be evaluated by observing the foal nursing. Milk coming out of the mouth or nostrils can be associated with a swallowing deficit due to HIS or less commonly a cleft palate, generalized weakness, subepiglottic cyst, or choke. Pelvic and thoracic limb reflexes can be evaluated with the foal in lateral recumbency. Foals are hyperreflexic up to several weeks of age and have strong resting extensor tone. A cross-extensor reflex, accompanying a withdrawal reflex, may be present in normal foals up to 4 weeks of age. A central recognition of pain should occur with the withdrawal reflex; however, this may be absent in systemically ill foals as well as those with neurological disease. The gait is examined for weakness or ataxia. Peripheral neuropathies, in particular radial nerve and brachial plexus abnormalities, are not uncommon from trauma at parturition. Hindlimb paresis may be seen with congenital anomalies, birth trauma, and HIS and needs to be differentiated from musculoskeletal causes. Generalized muscle weakness is seen with HIS, sepsis, prematurity, and less commonly with botulism, vitamin E/selenium deficiency, or heritable muscle disorders. The seizure in foals Seizures can vary from mild localized signs of facial grimacing, twitching, abnormal eye movements, chewing, abnormal breathing patterns, head and neck rigidity to generalized tonic-clonic convulsions with dorsiflexion and paddling. By far the most common cause of neonatal seizures is HIS; the onset can be at birth or over the next 48 hours and clinical signs are variable. The diagnosis is made by exclusion of other diseases, supportive history, and laboratory findings. Hyponatremia also causes neonatal seizures and is frequently associated with ‘chewing’ type of seizure activity. Other causes such as meningitis, metabolic derangements, hepatic encephalopathy, trauma, idiopathic epilepsy, and congenital anomalies are uncommon and are differentiated by serum chemistries, electrolytes, and CSF analysis.
Urogenital system Initial physical examination includes evaluation of the external genitalia and umbilicus. Preputial edema and paraphimosis can be the result of tenesmus due to meconium impaction, scrotal hernias, or fluid overload. The umbilicus should be palpated for swellings associated with hematomas, urine leakage through a torn urachus, or body wall defects. Acquired patent urachus is common in recumbent
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foals. It is also associated with excessive straining or an infection of the umbilicus and coincides with the sloughing of the necrotic stump. Stranguria can be seen with urachitis, cystitis, or uroperitoneum and frequent posturing to urinate is often confused with signs of abdominal pain. Hematomas of the internal umbilical remnants or within the bladder can be palpated through the abdominal wall in a relaxed recumbent neonate. Distended bladders due to outflow obstruction can occur with blood clots, a ‘kinked’ penis or occasionally with no apparent reason, possibly from a loss of coordinated neuromuscular control due to HIS, or in utero twisting of the umbilicus.
Ultrasonographic evaluation Ultrasonographic evaluation of the internal umbilical remnants and kidneys in foals has been described in detail.10, 11 The kidneys are evaluated for size, appearance of the parenchyma, perirenal edema, or cysts, though clinical signs and laboratory changes seen in early renal failure are often present without obvious ultrasonographic changes. Enlarged umbilical vessels with hypoechoic to anechoic cores indicate infection. Hematomas in the umbilical remnants and blood clots in the bladder can be visualized with ultrasonography. Defects in the bladder wall can be imaged; however, a full bladder does not eliminate the possibility of a leak. The urachal remnant should be examined for tears which can lead to uroperitoneum or subcutaneous urine accumulation.
can be seen associated with dystocias or trauma after parturition. Ruptured gastrocnemius tendon or muscle can occur during or shortly after foaling and can be a cause of recumbency in a foal. Patella luxations can occur unilaterally or bilaterally. Tendon contracture or laxity should be assessed and can be so severe that the foal is unable to stand. Foals with mildly contracted limbs are sometimes presented with a markedly elevated respiratory and heart rate and muscle tremors associated with the pain from the affected legs (usually forelimbs). The extensor tendons should be evaluated in foals with flexor contracture as ruptures can occur which are seen clinically as a soft fluctuant swelling on the lateral aspect of the carpus. All joints and physes should be palpated for heat, pain, swelling or edema, and if lameness or other abnormalities are found they should be considered infected until proven otherwise.
Diagnostic imaging Radiographs should be taken to assess cuboidal bone ossification, and evaluate affected areas for fractures or evidence of infection. Care needs to be taken interpreting the radiographs as there is variability of the appearance of the centers of ossification; radiographs of the opposite leg provide a useful comparison. Ultrasonography can be utilized to evaluate fractured ribs, ruptured gastrocnemius tendon, and infectious orthopedic disease.
Eyes (see also Chapter 59) Musculoskeletal system The head is examined for evidence of trauma around the orbits associated with the use of eyehooks, a domed calvarium which may be indicative of prematurity or hydrocephalus, or wry nose. Deviation of the vertebrae may be seen alone or with other musculoskeletal defects. The thorax should be palpated to determine if fractured ribs or costochondral dislocations are present. A flail chest may be present if multiple fractures are present. In one study up to 20% of foals less than 3 days of age had physical evidence of thoracic trauma.13 Whilst in this study there were no reported clinical problems associated with the rib fracture, in other studies rib fractures have contributed to morbidity and mortality.14, 15 Ultrasonographic examination is more reliable than radiography in detecting the fracture and allows visualization of the fracture site, displaced extremities, and evaluation of the adjacent structures.14, 16 Fractures and dislocation of joints are not common but
The most common cause of corneal edema and ulceration is entropion. Corneas are less sensitive in foals, thus ulceration can be present without evidence of pain. Fluorescein staining should be performed if there is suspicion of an ulcer or routinely as part of the physical examination. Hypopyon, uveitis, and iriditis are frequently associated with septicemia and can be present at birth if the foal is born with sepsis. The most common reported congenital anomalies are cataracts.17 Focal retinal hemorrhages, believed to be associated with HIS, have been reported. Scleral hemorrhages are usually associated with birth trauma though can be an incidental finding. Scleral injection is usually associated with sepsis.
Laboratory data Hematology and serum biochemistry should be performed if there is any suspicion of neonatal disease or in-utero compromise. Foals can be born with laboratory abnormalities reflecting the abnormal
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Table 17.1 Normal hemogram reference values for foals ≤ 7 days of age.18 Reference ranges determined by mean ± 2 SD Age
RBC PCV Hb MCV MCHC WBC Neutrophils Lymphocytes Monocytes Eosinophils Basophils Platelets
× 1012 /L L/L g/dL FL g/dL × 109 /L ×109 /L × 109 /L × 109 /L × 109 /L × 109 /L × 109 /L
< 1 h presuck
< 12 h postsuck
24 hours
7 days
9.3–12.9 0.40–0.52 13.4–19.9 37–45 33–39 — — — — — — —
9.0–12.0 0.37–0.49 12.6–17.4 36–45 32–40 6.9–14.4 5.55–12.38 0.46–2.54 0.04–0.43 0 0–0.02 105–446
8.2–11.0 0.32–0.46 12.0–16.6 36–46 32–40 4.9–11.7 3.36–9.57 0.67–2.12 0.07–0.39 0–0.02 0–0.03 129–409
7.4–10.6 0.28–0.43 10.7–15.8 35–44 35–40 6.3–13.6 4.35–10.55 1.43–2.28 0.03–0.54 0–0.09 0–0.18 111–387
in-utero environment, thus helping the veterinarian identify a foal at increased risk for developing clinical problems. Veterinarians should become familiar with the results from their laboratory and the individual variations that can occur in the reported reference ranges (Tables 17.1, 17.2 and 17.3). Glucose and lactate concentrations can be easily measured with handheld analyzers. Arterial blood gas analysis can be performed with handheld analyzers to evaluate respiratory function and acid–base abnormalities. A brief outline of the common abnormalities detected on frequently used clinicopathological data in foals are listed below. Packed cell volume The packed cell volume (PCV) of a newborn foal is 40–52%. This decreases with the ingestion of colostrum and blood volume expansion to 32–46% after 24 hours.18 An elevated PCV may be associated
with chronic in-utero hypoxia or dehydration. A decreased PCV can be due to internal or external blood loss (fractured ribs, umbilical remnant or gastrointestinal hemorrhage), prematurity, hemolysis associated with neonatal isoerythrolysis, or DIC due to sepsis. Hemolysis is usually associated with icterus. Decreased red blood cell production and anemia associated with chronic disease is usually not seen until later in the neonate’s disease process. Total protein Plasma protein levels vary from 4–6 g/dL at birth to 5–8 g/dL after 24 hours.18 Total protein concentrations increase with colostral absorption, dehydration, and hyperglobulinemia from chronic infection which may also occur from in-utero infections. Decreased levels can occur with blood loss, renal and gastrointestinal disease, and failure of passive transfer.
Table 17.2 Electrolyte reference values for foals ≤ 7 days of age.19 Reference ranges determined by mean ± 2 SD Age Analyte Sodium Potassium Chloride Carbon dioxide Phosphorus Calcium Magnesium Anion gap
mEq/L mEq/L mEq/L mEq/L mg/dL mg/dL mg/dL mEq/L
< 1 h pre-suck
< 12 h post-suck
1 day
7 days
— — — — — — — —
148 ± 15 4.4 ± 1.0 105 ± 12 25 ± 5 4.7 ± 1.6 12.8 ± 2.0 1.5 ± 0.8 21 ± 12
141 ± 18 4.6 ± 1.0 102 ± 12 27 ± 6 5.6 ± 1.8 11.7 ± 2.0 2.4 ± 1.8 16 ± 8
142 ± 12 4.8 ± 1.0 102 ± 8 28 ± 4 7.4 ± 2.0 12.5 ± 1.2 2.0 ± 0.6 17 ± 8
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Table 17.3 Reference values for serum biochemistry in foals ≤ 7 days of age19–21 Age < 1 h pre-suck
< 12 h post-suck
1 day
7 days
ALP
IU/L
—
1729 ± 537 (152–2835)
1487 ± 452 (861–2671)
652 ± 253 (137–1169)
GGT
IU/L
18.9 ± 7.4 (8.0–33.0)
—
—
48.4 ± 28.5 (16.0–98.0)
IU/L
—
22 ± 7 (13–39)
24 ± 7 (18–43)
42 ± 31 (14–64)
SDH
IU/L
9.4 ± 5.5 (1.2–21.2)
—
—
5.1 ± 5.1 (1.1–18.2)
Total bilirubin
mg/dL
2.1 ± 0.6 (0.9–2.8)
—
—
2.4 ± 0.5 (1.9–3.3)
mg/dL
—
2.0 ± 0.6 (0.9–2.8)
2.9 ± 1.3 (1.3–4.5)
1.7 ± 0.5 (0.8–3.0)
mg/dL
0.1 ± 0.04 (0.1–0.2)
—
—
0.22 ± 0.1 (0.1–0.4)
mg/dL
—
0.5 ± 0.1 (0.3–0.6)
0.5 ± 0.2 (0.3–0.7)
0.5 ± 0.1 (0.3–0.7)
Bile acids
µmol/L
41.4 ± 178.3 (21.7–81.7)
—
—
22.5 ± 4.7 (16.7–29.4)
Triglycerides
mg/dL
24 ± 8 (11–35)
—
—
95 ± 56 (35–200)
mg/dL
—
44 ± 13 (24–88)
82 ± 42 (30–193)
124 ± 58 (30–239)
Cholesterol
mg/dL
—
185 ± 76 (111–432)
246 ± 116 (110–562)
227 ± 71 (139–445)
AST
IU/L
—
97–315
146–340
237–620
Direct bilirubin
CK
IU/L
—
65–380
40–909
52–143
Creatinine
mg/dL
—
1.7–4.2
1.2–4.3
1.0–1.7
BUN (urea)
mg/dL
—
12–27
9–40
4–20
Glucose
mg/dL
—
108–109
121–223
121–192
White blood cell count
Platelets
At birth the white blood cell (WBC) count is similar to an adult; however, there is considerable variation in WBC, neutrophil and lymphocyte numbers. Neutropenia with toxic changes and increased number of immature cells is associated with sepsis. A neutrophil : lymphocyte ratio < 2–3 without signs of sepsis is associated with prematurity. Premature foals with a neutropenia and N : L ratio < 1.5 in the first 24 hours of life have been reported to have a poor prognosis.18 An elevated WBC at birth may represent the fetal inflammatory response to the infected placenta rather than primary sepsis in the foal. Persistently low lymphocyte counts may be associated with combined immunodeficiency disease.
Platelet counts are relatively constant during the first year of life.18 Thrombocytopenia can be seen with DIC due to sepsis, viral infection (EHV-1, equine viral arteritis), alloimmune thrombocytopenia, or ulcerative dermatitis.
Fibrinogen Fibrinogen concentration increases gradually after birth.18 Values greater than 4 g/L are indicative of an infection or inflammation; those of 3–4 g/L raise suspicion. Hyperfibrinogenemia at birth is indicative of in-utero sepsis and inflammation.
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Immunoglobulin G
Alkaline phosphatase
Foals that have absorbed adequate colostrum have an immunoglobulin G (IgG) concentration greater than 8 g/L. Those with complete failure of passive transfer have an IgG concentration less than 2 g/L; those with partial failure of passive transfer have values between 4 and 8 g/L. Foals can be born with an IgG concentration greater than 8 g/L with chronic in-utero infection. Foals which are septic catabolize their immunoglobulins, resulting in a foal which originally had an IgG concentration greater than 8 g/L subsequently having a lower IgG concentration.
Alkaline phosphatase activity is high in the first week of life due to increased bone development, intestinal pinocytosis, and/or hepatic maturation.23 Foals with incomplete ossification have a prolonged elevation of alkaline phosphatase.22
Creatinine and blood urea nitrogen concentrations Elevated creatinine and urea concentrations can be due to prerenal, renal, and postrenal causes. In newborn foals markedly elevated creatinine concentrations can occur associated with fetoplacental problems and is an indicator of a potentially compromised foal. If the increase is due to fetoplacental problems, the creatinine concentration decreases quickly over the following days. If the creatinine concentration remains elevated or decreases slowly, concurrent renal problems or other causes of azotemia should be considered. Urea is not generally elevated with creatinine in placental dysfunction. It is often elevated in neonates associated with catabolism.
Bilirubin Newborn foals often have elevated bilirubin concentrations as a result of an increased total and unconjugated bilirubin. The cause is unclear but is attributed to an accelerated breakdown of neonatal RBCs or immature hepatic metabolism of bilirubin.24 Other hepatic enzymes, gamma glutamyl transferase, sorbitol dehydrogenase and aspartate aminotransferase, are also transiently increased. Elevated bilirubin concentrations are also seen with liver disease, hemolysis, sepsis, meconium impaction, and from RBC absorption after internal hemorrhage.
Electrolytes Causes for serum electrolyte changes in foals are similar to adults, although changes in the newborn foal also reflect placental dysfunction and altered in-utero life. If the renal function of the neonate is normal, the electrolytes should normalize over the following 48 hours.
Lactate Glucose Newborn foals have a relatively low glucose concentration which increases after nursing. Hypoglycemia can be seen with: lack of milk intake; increased metabolic demands (sepsis, SIRS); reduced gluconeogenesis which can occur with sepsis, SIRS, prematurity or in-utero stress. Hyperglycemia can be seen with excessive glucose administration, increased catecholamine production, prematurity, HIS, sepsis and SIRS which can cause slow insulin responses or inappropriate gluconeogenesis.
Lactate concentrations are used to evaluate peripheral perfusion and tissue oxygenation. Newborn foals have slightly higher levels that are similar to adult levels by 24 hours; a value of less than 2.5 mmol/L in a neonate is considered normal.25 Increased values are seen in foals in association with tissue hypoxia and poor perfusion. Increased values are also seen hypermetabolic states (sepsis, seizures, increased muscle activity), activation of inflammatory cells and mediators with sepsis and SIRS, and impaired hepatic clearance. (See also Chapter 18).
Conclusion Creatine kinase Increased concentrations may be seen with trauma at parturition, prolonged recumbency, glycogen storage disease IV, and white muscle disease. Increased concentrations have also been seen with HIS and have been attributed to global damage to the cellular membranes resulting in ‘sick cell syndrome’.22
The findings from the clinical assessment and clinicopathological results will help the clinician make a diagnosis, provide direction for further diagnostic evaluation, and help formulate the treatment and management of the critically ill neonate. Early detection of these changes allows for prompt therapeutic intervention and ultimately an increased chance for survival of the neonate.
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References 1. Rossdale PD. Clinical studies of the newborn thoroughbred foal: Heart rate, auscultation, and electrocardiogram. Br Vet J 1967;123(12):521–32. 2. Yamamoto K, Yasuda J, Too K. Electrocardiographic findings during parturition and blood gas tensions immediately after birth in thoroughbred foals. Jpn J Vet Res 1991;39:143–57. 3. Reef VB. Cardiovascular disease in the equine neonate. Vet Clin N Am Equine Prac 1985;1:117–29. 4. Livesey LC, Marr CM, Boswood A, Freeman S, Bowen IM, Corley KTT. Auscultation and two-dimensional, M mode, spectral and colour flow Doppler findings in pony foals from birth to seven weeks of age. J Vet Intern Med 1998;12:255. 5. Marr CM. Cardiac murmurs: congenital heart disease. In: Marr CM (ed.) Cardiology of the Horse. London: Saunders, 1999; pp. 210–32. 6. Corley KTT. Monitoring and treating haemodynamic disturbances in critically ill neonatal foals. Part 1: Haemodynamic monitoring. Equine Vet Educ 2002;14:270–9. 7. Franco RM, Ousey JC, Cash RS, Rossdale PD, Silver M. Study of arterial blood pressure in newborn foals using an electric sphygmomanometer. Equine Vet J 1986:18:475–8. 8. Wilkins PA. Lower respiratory problems of the neonate. Vet Clin N Am Equine Prac 2003;19:19–33. 9. Palmer JE. Ventilatory support of the critically ill foal. Vet Clin N Am Equine Prac 2005;21(2):457–86. 10. Reef VB. Pediatric abdominal ultrasonography. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia: Saunders, 1998; pp. 364–403. 11. Porter MB, Ramirez S. Equine neonatal thoracic and abdominal ultrasonography. Vet Clin N Am Equine Prac 2005;21:407–29. 12. MacKay, RJ. Neurological disorders of neonatal foals. Vet Clin N Am Equine Prac 2005;21:387–406. 13. Jean D, Laverty S, Halley J, Hannigan D, Leveille R. Thoracic trauma in newborn foals. Equine Vet J 1999;31(2):149– 52.
14. Sprayberry KA, Bain FT, Seahorn TL, Slovis NM, Byars TD. 56 cases of rib fractures in neonatal foals hospitalized in a referral center intensive care unit from 1997–2001. In: Proceedings of the American Association of Equine Practitioners, San Diego, 2001; pp. 395–9. 15. Schambourg MA, Laverty S, Mullim S, Fogarty UM, Halley J. Thoracic trauma in foals: post mortem findings. Equine Vet J 2003;35:78–81. 16. Jean D, Picandet V, Macieira S, Beauregard G, D’Anjou MA, Beauchamp G. Detection of rib trauma in newborn foals in an equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 207;39(2):158–63. 17. Whitley RD. Neonatal equine ophthalmology. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 531–57. 18. Harvey JW. Normal hematologic values. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 561–70. 19. Bauer JE. Normal blood chemistry. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 602–14. 20. Bauer JE, Asquith RL, Kivipelto J. Serum biochemical indicators of liver function in neonatal foals. Am J Vet Res 1989;50(12):2037–41. 21. Barton MH, LeRoy BE. Serum bile acids concentrations in healthy and clinically ill neonatal foals. J Vet Intern Med 2007;21:508–13. 22. Axon JE, Palmer JE. Clinical pathology of the foal. Vet Clin N Am Equine Prac 2008;24(2):356–86. 23. Bauer JF, Harvey JW, Asquith RL, McNulty PK, Kivipelto J. Clinical chemistry reference values of foals during the first year of line. Equine Vet J 1984;16(4):361–3. 24. Hawthorne TB. Neonatal hyperbilirubinemia. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 589– 601. 25. Corley KTT, Donaldson LL, Furr MO. Arterial lactate concentration, hospital survival, sepsis and SIRS in critically ill neonatal foals. Equine Vet J 2005;37(1):53–9.
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Chapter 18
Critical Care – Monitoring Darien J. Feary
Introduction Critically ill neonatal foals are evaluated using basic physical examination skills to enable the veterinarian to make an initial assessment of the foal’s clinical status and the necessity and urgency for therapeutic intervention. Technology, in the form of various monitoring devices, can be used to augment the clinical picture. Point-of-care monitoring equipment for analysis of variables such as lactate, glucose, electrolytes, blood pressure, blood gas, and renal parameters have become practical, affordable, accurate, and increasingly available for use in equine practice. Familiarity with these basic monitoring techniques facilitates early identification of homeostatic imbalances that may not already be clinically identifiable, and provides a quantitative guide to titration and end points of therapy. The limitations of physical examination are particularly highlighted in neonates. Foals with significant cardiorespiratory disturbances may initially have normal heart rates, pulse pressure, mucous membranes, respiratory rates, and mentation, and in some instances have reduced ability to compensate for dysfunction in the normal manner. Progression to the next step of performing a thorough quantitative evaluation through point-of-care diagnostics is relatively simple. More information can be acquired within minutes following collection of a 3-mL heparinized venous or arterial blood sample, a free catch urine sample, and the use of a centrifuge, a refractometer, an i-STAT portable clinical analyzer, and a handheld lactate handheld (Heska Corporation). Further assessment of arterial oxygenation, blood pressure, and electrocardiography can be readily achieved stall-side with various portable monitoring devices (Figure 18.1). The intensity of monitoring required in any individual foal will vary depending on the foal’s clinical status. Ambulatory foals may only require frequent
physical examination every 4 to 6 hours, basic assessment of hydration status such as packed cell volume (PCV), total protein (TP), and lactate every 12 to 24 hours, and analysis of urine specific gravity performed on frequently collected urine samples. Weak, recumbent foals require more intensive care and are more likely to be hemodynamically unstable and have compromised oxygen delivery to tissues. These foals typically do not respond adequately to standard fluid resuscitation, and close attention to less obvious sources of homeostatic dysregulation is often required in order to identify correctable physiological deficits.
Packed cell volume (PCV) Packed cell volume can be measured directly from a small amount of freshly collected whole blood, or blood stored in an EDTA (ethylene diamine tetra-acetic acid) anticoagulant tube (Figures 18.2 and 18.3). The measurement of PCV is not as sensitive a method of evaluating volume and hydration status in neonatal foals as it can be in adult horses, and hypovolemic foals will often have a PCV that measures within the normal range. Age-related changes in PCV occur in foals, such that prior to nursing newborn foals normally have a higher PCV (40–52%), which gradually decreases over the next 5 days (PCV = 32–44%) to 3 weeks (PCV = 28–43%).1 In addition, foals that are anemic due to hemolysis or hemorrhage may have inadequate circulating blood volume and have a low PCV. A single PCV measurement in a sick foal can provide useful information in some instances of severe volume depletion, and is useful to detect neonatal isoerythrolysis upon initial examination. However, it is preferable to monitor trends in PCV over time to indicate more subtle changes of under- or overhydration. Relying on PCV alone probably should
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 18.1 Practical, stall-side use of portable monitoring equipment for intermittent or continuous measurement of indirect arterial blood pressure and pulse oximetry in a critically ill newborn foal.
not be regarded as a reliable laboratory method of assessing circulatory status in compromised neonatal foals. Rather, PCV should be evaluated in combination with other more specific indicators of cardiovascular volume such as blood lactate, urine specific gravity, and urine output.
Figure 18.3 Measurement of packed cell volume following centrifugation.
Total protein (TP) Measurement of plasma protein in the centrifuged blood sample using a refractometer is simple to perform and should be part of routine laboratory evaluation of the compromised neonatal foal (Figure 18.4). Plasma protein is influenced by age-related changes in foals and is normally low at birth (4.4–5.9 g/dL) prior to nursing, and increases following ingestion of immunoglobulin in colostrum (5.1–7.6 g/dL).1 Compared with adults, TP is far less likely to increase significantly in foals that are hypovolemic. Conversely, a TP measurement below the normal range in foals over 24 hours of age should alert the veterinarian to conditions such as inadequate passive transfer of immunoglobulin, protein-losing enteropathy secondary to diarrhea, recent hemorrhage, or sepsis, all of which warrant further investigation.
Colloid osmotic pressure (COP) Oncotic pressure refers to the osmotic (oncotic) effect of macromolecules, mostly albumin, that
Figure 18.2 Bench-top centrifuge for separation of blood and plasma using a microhematocrit tube to facilitate measurement of packed cell volume (PCV) and total plasma protein (TP).
Figure 18.4 Measurement of total plasma protein with a handheld refractometer.
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Critical Care – Monitoring
Figure 18.5 Colloid osmometer for direct laboratory measurement of plasma colloid oncotic pressure.
cannot permeate the vascular endothelium and thus act to retain fluid within the vasculature. In hypoproteinemic foals receiving synthetic colloids (hydroxyethylstarch, pentastarch, human albumin, dextrans, or modified gelatins) COP can be monitored by either direct measurements using a commercially available, bench-top colloid osmometer (Figure 18.5) or indirect calculated estimations using the Landis–Pappenheimer equation: COP = 2.1 TP + (0.16 TP2 ) + (0.009 TP3 ) where TP = total protein. Direct measurements are the preferred method as indirect calculations are inaccurate in critically ill neonatal foals.2 The normal range of COP in neonatal foals (13–22 mmHg) is lower than in adult horses (20–30 mmHg).2, 3 COP measurement can be used as a trigger point for initiating colloid therapy in foals with albumin loss, and for monitoring response to therapy in foals receiving synthetic colloids. This may be necessary as the contribution of the synthetic colloids to COP cannot be detected by refractometry.
Lactate Blood lactate can be measured in whole blood or plasma by gold-standard laboratory analyzers, or by means of various point-of-care (POC) monitors – iSTAT (Abbott Laboratories, IL, USA), Accutrend (Roche Diagnostics, Mannheim, Germany), Accusport (Accusport International Inc., NC, USA), Lactate Scout (SensLab, Germany), and Lactate Pro (Arkray
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Global Business Inc., Japan). POC lactate monitors are inexpensive, portable, simple to operate, and accurate; are readily available to equine practitioners worldwide; and can provide reliable, clinically useful information in neonatal foals (Figure 18.6). The accuracy and reliability of some commercially available stall-side lactate monitors have been evaluated against laboratory analyzers in normal horses4, 5 and in abnormal adult horses with colic and other emergency-type conditions,6 and have demonstrated clinical utility.7, 8 Although lactate measurement in equine plasma samples is slightly different and more accurate than that for whole blood,6 whole blood samples are more convenient and the results remain sufficiently reliable to be clinically useful. Results are less reliable at high PCVs (> 53%) and high lactate concentrations (> 10 mmol/L)4, 6 in horses. Lactate analysis involves collection of a venous or arterial blood sample into a heparinized syringe or lithium heparin tube, placing a drop of blood onto a test strip, and waiting up to 60 seconds for a result to be displayed. Because erythrocytes continue to produce lactate following collection, it is recommended that if there is a delay of 5 minutes or more in sample analysis then samples should be separated or stored in ice water. Venous blood lactate concentrations reported in newborn foals immediately postpartum (< 3.5 mmol/L)9 and within the first 30–46 hours of life (< 2.5 mmol/L)10 are slightly higher than in adults (< 2.0 mmol/L),11 but should decrease to adult levels within the first 24 to 48 hours in a normal resting foal. Lactate is the normal end product of glycolysis (anaerobic metabolism) and is a clinically useful biochemical marker of the adequacy of global oxygen delivery to tissues. Elevations in blood lactate concentration occur most commonly in sick foals as a result of reduced cardiac output (stroke volume) due to hypovolemia and hypoperfusion, thus therapeutic interventions such as intravenous fluid therapy that result in decreasing blood lactate concentrations generally indicate improved tissue perfusion and a beneficial response.11 However, elevations in blood lactate can occur with relative hypovolemia due to vasodilation associated with sepsis or perinatal asphyxia, and for a variety of other reasons (Table 18.1), and these should be urgently investigated and addressed in foals with increasing or persistently elevated lactate concentrations despite adequate correction of deficits in circulating blood volume. The observation that blood lactate changes are more often associated with tissue perfusion than hypoxemia or anemia should be emphasized.12 Serial monitoring of blood lactate is clinically useful as a therapeutic trigger or end point, and
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(a)
(b)
Figure 18.6 (a) Point-of-care lactate monitor, test strips, and (b) application using whole blood collected in a heparinized syringe.
for titration of specific interventions such as intravenous fluid therapy, inopressor therapy, oxygen therapy, or blood transfusion. Blood lactate measurement is also useful as an indicator of severity of disease and prognosis for survival in critically ill neonatal foals. Mean blood lactate concentrations in critically ill foals aged less than 7 days at admission and at 18–36 hours were significantly lower
in foals that survived to hospital discharge (admission lactate 4.37 ± 0.55 mmol/L; 18–36-hour lactate 3.23 ± 0.53 mmol/L) compared with non-survivors (admission lactate 9.31 ± 0.86 mmol/L; 18–36-hour lactate 9.12 ± 0.71 mmol/L).13 Equine veterinarians should be cautioned against interpreting a single lactate concentration as a means of differentiating survivors from non-survivors upon
Table 18.1 Causes of elevated blood lactate concentration in horses Tissue hypoxia
Tissue hypoxia not present
1. Reduced O2 delivery Hypovolemia/hypoperfusion (global or local): (a) circulating blood volume deficit (b) acute blood loss (c) sepsis/endotoxemia (d) heart failure (e) arterial thromboembolism Severe anemia (Hct < 15%) Severe hypoxemia (P a o2 < 40 mmHg) Hemoglobin dysfunction (methemoglobinemia/CO toxicity) 2. Increased O2 consumption Strenuous exercise Muscle fasciculations/trembling Seizures
Liver hypoperfusion or failure Increased glycolysis: (a) catecholamines (b) glucose or HCO3 − infusion, parenteral nutrition Malignancy Sepsis Endotoxemia
Hct hematocrit; P a o2 , artertal oxygen tension.
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Critical Care – Monitoring initial examination of sick foals. However, single lactate determinations can provide useful information, as in most situations the higher the blood lactate concentration of the foal, the greater the degree of cardiovascular compromise, and the greater the severity of underlying disease. The correlation between the magnitude of elevation in blood lactate concentration and degree of hypoperfusion has been defined in human patients as mild (2–5 mmol/L), moderate (5–7 mmol/L), and severe (> 7 mmol/L).14 Although not defined as such in horses, clinical experience would suggest a similar correlation. More important than the magnitude of elevated blood lactate concentration at initial examination is the serial monitoring of blood lactate over time (every 4 to 6 hours) in response to therapy and as an indicator of prognosis. In general, failure of appropriate therapeutic intervention to lower significantly lactate concentrations over the first 12 to 48 hours is a poor prognostic indicator in critically ill foals. Blood lactate can return to normal values in adult horses within 2 hours following appropriate therapy,13 in the absence of liver or kidney failure. A similar response is probably true for neonatal foals, but this has not been documented in critically ill neonatal foals. Lactate can also be measured in body fluids other than blood. Measuring peritoneal fluid lactate and the disparity between it and peripheral blood lactate values has demonstrated clinical utility in differentiating strangulating from non-strangulating intestinal obstructions, at least in adult horses with colic.7, 15, 16 Analysis of lactate concentrations in peritoneal, pleural, synovial, and cerebrospinal fluid is also of diagnostic value in differentiating septic and non-septic effusions. Lactate concentrations are believed to be elevated higher in septic effusions, compared with non-septic inflammatory effusions, due to additional lactate production by bacterial metabolites and increased neutrophil glycolysis. Body fluid lactate should be evaluated in comparison with peripheral blood lactate values, as well as simultaneous determination of a blood-to-fluid glucose difference of > 20 mg/dL (1.1 mmol/L). However, there is considerable overlap in the magnitude of lactate concentration between inflammatory conditions and septic conditions in body fluids, and there is no single published lactate value or blood-to-fluid gradient that is reliably diagnostic for septic effusions in horses. Therefore, other laboratory and clinical findings must also be considered.
Glucose Derangements in blood glucose concentrations are common in compromised neonatal foals. The reported normal range for blood glucose values in
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neonatal foals is variable, and subject to age-related changes in the first week of life. Logically, presuckle glucose concentrations [> 35–40 mg/dL (1.92– 2.20 mmol/L)] are normally lower than post-suckle glucose concentrations.17 Some neonatal intensive care units (NICUs) report a wide range of normally acceptable blood glucose values in neonatal foals, with reported ranges of 76–131 mg/dL (4.18–7.20 mmol/L)18 and 60–180 mg/dL (3.3–9.9 mmol/L).19 Critically ill neonatal foals can be hypoglycemic or hyperglycemic at the time of initial assessment. Hypoglycemia [glucose < 60 mg/dL (< 3.30 mmol/L)] can be life threatening if severe and is manifest clinically by neurological derangements (weakness, coma, seizure), which are usually associated with glucose concentrations below 30–40 mg/dL (1.65– 2.20 mmol/L). In addition, hypoglycemia in presuckle foals is associated with placental insufficiency and perinatal asphyxia syndrome.20 Although not regarded as directly life threatening per se, hyperglycemia [glucose > 180 mg/dL (> 9.90 mmol/L)] is also detrimental via a number of mechanisms and contributes to hypoxic-ischemic brain injury.18 The beneficial effect of tight glucose control via insulin infusion in critically ill human patients has been well described.21 Studies in equine patients have demonstrated an association between hypoglycemia and survival (in foals), and between hyperglycemia and survival (in adults).18 Clearly, monitoring blood glucose in sick neonatal foals is important because glycogen stores at birth are limited, body fat stores are very low, and unfed foals that are not nursing are prone to hypoglycemia. Rapid and repeated assessment of blood glucose for foals receiving dextrose-containing fluids, parenteral nutrition, or insulin therapy is often required. Blood glucose monitoring can be performed using a variety of techniques including bench-top laboratory chemistry analyzers, portable iSTAT blood gas analyzers, and point-of-care glucometers. Stall-side glucose monitors are the most convenient means for frequent glucose measurement and are widely used in NICUs throughout the world. There are at least 25 different glucometers designed for use in human patients that are readily available, inexpensive, simple, provide rapid results, and require only a single drop of blood, making them attractive for use in foals (Figure 18.7). However, the accuracy of these monitors has been questioned. Large variations in glucose readings can occur in association with the hematocrit of the patient (high or low), the vessel from which the blood is drawn, whether whole blood or plasma/serum is measured, and reagent strip storage and handling; there can also be marked variation between individual monitors and reagent
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Figure 18.7 Measurement of blood glucose using a portable glucometer.
strip batches. Comparison between point-of-care glucometers, the laboratory standard, and blood gas analysis has been investigated in critically ill neonatal foals and has demonstrated a less than ideal agreement between methods, with glucometry being the least accurate.19 It is important to recognize the limitations of glucometry and to minimize inaccuracies by using the same method of blood collection for each sample, and performing recalibration for each new batch of reagent strips; also, each facility should evaluate its glucometer by frequent comparision with the laboratory standard. Glucometry remains clinically useful in horses providing its inaccuracies are recognized and unexpected or aberrant results are re-evaluated using laboratory standards. Newer glucometers with improved accuracy for equine patients may become commercially available in the future. Continuous glucose monitors have also been developed and are available for use in small animals. These devices continuously measure glucose concentrations in the interstitial space via a small flexible sensor inserted into the subcutaneous tissues. They appear to be well tolerated in small animal patients and may become clinically useful in unstable critically ill neonatal foals in the future.
Electrolytes and clinical chemistries Bench-top laboratory and portable iSTAT monitors can be used to measure electrolytes and creatinine and blood urea nitrogen at initial presentation, often in combination with blood gas, glucose, lactate, coagulation markers, cardiac isoenzymes, and other measurements, depending on the cartridges selected. Electrolytes most commonly of interest in a sick foal include sodium, chloride, potassium, and ionized calcium. The serum sodium concentration is a reflection of water balance and distribution in the body and is tightly regulated. Severe hyponatremia and hypernatremia, or the consequence of
inappropriate correction of these disturbances, can have life-threatening neurological sequelae. Clinical conditions resulting in serum sodium and chloride derangements that occur commonly in neonatal foals are often associated with hypovolemia (diarrhea, intestinal ileus, gastric reflux, colitis, blood loss) and third-space sequestration (bladder rupture, peritonitis), or are of renal origin (renal failure, diuretic therapy). In addition, iatrogenic hypernatremia may be due to incorrect formulation of milk replacer, or iatrogenic hyponatremia may arise from overzealous administration of 5% glucose in water. Hyperkalemia can also result in life-threatening muscle weakness and cardiac arrhythmia in foals with uroabdomen or hyperkalemic periodic paralysis. Although uncommon, severe hypocalcemia causing muscle tetany, ileus, and tachycardia also occurs in foals and is most often idiopathic. It is important to remember that plasma calcium can be measured as total calcium or ionized calcium concentration. It is the ionized form that is physiologically the most important, and it is measured by most point-of-care electrolyte monitors. Total calcium concentrations are decreased in hypoalbuminemia, and it is important to correct for this using the following equation: corrected calcium (mg/dL) = measured serum total calcium − serum albumin (g/dL) + 3.5 Early detection and ongoing monitoring of electrolyte abnormalities and renal function guides appropriate intravenous fluid therapy, determines the appropriate rates to avoid the detrimental affects of too rapid correction or overcorrection, and identifies inorganic causes of acid–base abnormalities. Knowing the concentrations of sodium and glucose in a patient also allows calculation of plasma osmolality and fractional excretion of electrolytes. Normal plasma osmolality in healthy neonatal foals is 280–300 mOsm/kg, and is highest at birth.22 Measurement of osmolality is useful for differentiating hypotonic dehydration (most common) from hypertonic or isotonic dehydration, and for monitoring during hyperosmolar intravenous fluid therapy for renal failure or conditions where cerebral edema is suspected.
Urine output and urine specific gravity (USG) Monitoring urine output in foals is a useful means of assessing fluid balance, end organ perfusion, and renal function. Normal foals ingest a predominantly fluid diet and should produce approximately 6 mL/kg/h of urine, and have a USG that is normally hyposthenuric (< 1.008) but within the range of 1.001 to 1.010.22
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Critical Care – Monitoring Specific gravity of the very first urine voided is usually concentrated (> 1.035), however, and this is normally rapidly diluted as the foal begins ingesting milk. When intensive monitoring of urine output becomes necessary in foals with cardiovascular dysregulation (i.e., prematurity, dysmaturity, perinatal asphyxia, sepsis, oliguric/anuric renal failure) or for postoperative bladder repair management, indwelling urinary catheterization can be achieved with well secured 8- to 12-Fr, 33-cm (fillies) to 64-cm (colts) long Foley catheters.23 Urine can be collected via a closed sterile collection system. The advantages of placement of a continuous closed urine collecting system must be weighed against the risks of urinary tract infection, which can be nosocomial and life threatening in some instances. Urine output in hospitalized foals varies according to fluid input, which includes intravenous fluid therapy, parenteral, and enteral nutrition, but should amount to no less than approximately two-thirds of total fluid input.24 Reduced urine production, or inappropriately concentrated urine, may be due to circulatory shock, hypovolemia, hypotension, third-space fluid losses, renal failure, or urinary tract obstruction or leakage. Increased urine production is often iatrogenic, as a result of diuretic therapy or prolonged hyperglycemia and osmotic diuresis. When intensive urine output monitoring is not required or practical in recumbent foals, a crude estimation of urine output can be obtained by weighing urine collected onto pre-weighed absorbent pads (such as a diaper). This method assumes that all urine is voided onto the pads and that 1 mL of urine weighs 1 g. In ambulatory foals the simple observation of the frequency of urination and urine specific gravity and osmolality of free catch urine samples determined at regular intervals (at least every 12 hours) may be adequate for basic monitoring of fluid balance. Urine specifc gravity depends on the number of particles in solution as well as their molecular weight. Normally the relationship between USG and urine osmolality is linear (for small solutes): each 0.0001 of specific gravity = 30 mOsm/kg (so that a USG of 1.010 = urine 300 mOsm/kg, by multiplying the last two digits of USG × 30). Urine specific gravity and osmolality are primarily determined by the actions of antidiuretic hormone (ADH), which is secreted in response to hyperosmolality and hypovolemia. A dehydrated/hypovolemic or hypernatremic patient should have a high USG and osmolality. Conversely, a hyponatremic patient should have a low USG and osmolality because secretion of ADH is not stimulated. The observation that the maximum urine osmolality achieved in dehydrated foals (up to 760 mOsm/kg) is much less than that measured in dehydrated adult horses (up to 1540 mOsm/kg) is
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most likely attributed to the limited capability of foal kidneys to produce a highly concentrated urine.22 The pH of neonatal foal urine is normally acidic compared with that of the adult horse, and a transient but marked proteinuria is normally observed in the first 48 hours of life due to ingestion of colostrum.22 In addition, daily abdominal ultrasonographic examination of recumbent critically ill foals can be useful to confirm urine production based on the presence of urine in the bladder, and to monitor for urinary tract rupture. Hospitalized recumbent foals, particularly those with infection, sepsis, and perinatal asphyxia, appear to have an increased risk for rupture of the urinary bladder and require close monitoring.25
Bodyweight The use of serial bodyweight measurements as a routine daily estimation of fluid balance and nutritional status, is a useful monitoring tool. Body weight determination is also important for accurate drug dosing. A normal 40–50 kg neonatal foal should gain approximately 1 ± 0.5 kg/day. Sick foals often lose weight due to their underlying illness, reduced appetite, and perhaps the limitations of or reluctance to provide nutritional support as part of hospital management. A realistic goal of nutritional management of hospitalized foals should be at the very least, maintenance of bodyweight.24 An unexpected or acute change in bodyweight should alert the veterinarian to investigate possible causes of fluid imbalance, such as underhydration, overhydration, fluid retention (as seen with renal disease, heart failure, syndrome of inappropriate ADH secretion), or third-space fluid accumulation (i.e., uroperitoneum).
Pulse oximetry Pulse oximetry is a non-invasive, indirect method of measuring the arterial oxygen saturation of hemoglobin. The technique relies on placement of a human fingertip probe over an area of pulsatile arterial flow and detects the difference in absorption of reflected light at two wavelengths: red (oxyhemoglobin) (660 nm) and infrared (deoxyhemoglobin) (940 nm). Based upon the ratio of changing absorbance of the red and infrared light caused by the difference in color between oxygen-bound (bright red) and oxygen unbound (dark red or, in severe cases, blue) blood hemoglobin, a measure of oxygenation (the percent of hemoglobin molecules bound with oxygen molecules) can be made. Pulse oximetry is a useful technique for continuous or intermittent monitoring of the oxygenation
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status of foals and can limit the need for frequent invasive arterial sample collection. Measured arterial oxygen saturation (Sp o2 ) values of ≥ 96% measured by pulse oximetry correlate with a Pa o2 of at least 80 mmHg based on extrapolation from a normal oxyhemoglobin dissociation curve.26 Therefore, recorded values < 96% generally indicate hypoxemia (Pa o2 < 80 mmHg) and warrant further investigation and treatment. Pulse oximetry is also useful for calculation of oxygen extraction ratio (O2 ER) in combination with venous blood values as a trigger for blood transfusion in anemic foals (see below). Pulse oximetry depends on detection of peripheral arterial pulsatile flow, therefore accuracy is limited by hypotension, peripheral vasoconstriction, hypothermia, hairy, thick, or pigmented skin, motion, ambient lighting, and low actual arterial oxygen saturation (Sa o2 < 80%). Veterinarians can maximize the accuracy of pulse oximetry in clinical patients by:
r r r r
placing the probe at a site that is non-pigmented, warm and thin (foal tongue, ear, lip, or flank fold; Figure 18.8); minimizing movement of the patient during readings; ensuring that the digital and audible pulse rate displayed by the pulse oximeter correlates with the patient’s actual pulse rate at the time; taking multiple readings, and confirming with arterial blood gas analysis any readings that do not correlate with the patient’s clinical picture.
Standard pulse oximetry monitors do not differentiate carboxyhemoglobin or methemoglobin, and are of limited value in rare cases of carbon monoxide toxicity or in the presence of oxidants such as in
Figure 18.8 Non-invasive monitoring of arterial oxygen saturation (pulse oximetry) in a neonatal foal. The probe is placed on the lip or tongue.
red maple leaf toxicity. In these situations the use of co-oximetry, which employs a laboratory-based instrument able to measure the wavelengths of all four hemoglobin species, is indicated. Pulse oximetry is more reliable for evaluating foals breathing room air than those on supplemental oxygen. When supplemental oxygen is administered significant deterioration in lung function can occur without a decrease in Sp o2 below 99–100%, based on the relationship between the fraction of inspired oxygen (Fi o2 ) and the expected Pa o2 at sea level: Pa o2 = 5 × Fi o2 .27 Veterinarians should also be aware that significantly anemic patients should have maximum saturation of hemoglobin with oxygen and normal Sp o2 readings if pulmonary function is normal, thus arterial blood gas analysis is more reliable in detecting hypoxemia in anemic foals.
Blood gas analysis Blood gas analysis is the determination of the pH, and partial pressures of oxygen (Po2 ) and carbon dioxide (Pco2 ) in arterial or venous blood. Many laboratory (Figure 18.9) and portable blood gas analyzers also measure oxygen saturation (So2 ) and bicarbonate (HCO3 − ), thus the pulmonary function and acid–base status of a patient can be readily assessed in compromised foals. Additionally, most analyzers or specific cartridges enable simultaneous measurement of electrolytes, thus providing more information regarding the origin of the acid–base abnormality. An arterial blood sample is required for accurate assessment of respiratory function. This can be obtained most commonly from the dorsal metatarsal
Figure 18.9 A laboratory bench-top blood gas analyzer enables results of a variety of biochemical and blood gas parameters to be obtained almost immediately from a 3-mL heparinized whole blood sample.
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Figure 18.10 Collection of arterial blood from the metatarsal artery of a laterally restrained neonatal foal.
artery in recumbent restrained foals (Figure 18.10). Alternative sites of collection include the facial, transverse facial, and the brachial arteries. The latter site is almost always palpable and is a particularly useful site for sampling in foals that have poor peripheral perfusion. Ideally, an arterial sample is obtained at initial evaluation of critically ill foals, and if pulmonary oxygenation is adequate based on clinical assessment and a normal Pa o2 value, then venous blood gas analysis may be sufficient for ongoing monitoring of ventilatory and acid–base status. The normal Pa o2 of a standing horse breathing room air at sea level is 80 to 120 mmHg. However, both the postnatal age and position of the foal during sample collection can dramatically affect blood gas interpretation. Normal arterial blood gas values reported for newborn foals in lateral recumbency are less than in adults; ranging between 50 and 60 mmHg during the first 60 minutes postpartum, and increasing to 70–80 mmHg within the first 24 to 48 hours.28, 29 Lateral recumbency results in a lower Pa o2 than in the standing position, with the reduction being up to 12 mmHg.28, 30 Samples obtained from the dorsal metatarsal artery from hypotensive, poorly perfused neonates may have a lower Pa o2 compared to those obtained from the larger, more centrally located brachial artery. These points should be taken into consideration when interpreting results of blood gas analysis in foals, and an effort made to obtain samples from the same location if possible. Critically ill foals in which the Pa o2 measurement is < 60 mmHg may be severely hypoxemic, therefore intranasal oxygen therapy is always indicated. Pa o2 values of <80 mmHg in foals aged 48 hours or more indicates hypoxemia and may warrant oxygen therapy.
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The Pco2 is a measure of ventilatory status and can be assessed from arterial or venous samples. The normal Pco2 in newborn, laterally recumbent foals is reported to be between 42 and 48 mmHg.24, 31 Venous Pco2 is approximately 3–6 mmHg higher than arterial Pco2 ,24 as it also reflects tissue metabolism. Hypoventilation is common in critically ill neonatal foals, and is also a cause of hypoxemia. Arterial or venous Pco2 values greater than 60 mmHg that are not a compensation for metabolic alkalosis, warrant further investigation and possible intervention, such as pharmacological stimulation or mechanical ventilation. Sample handling can significantly affect blood gas analysis results. Samples should be collected in heparinized syringes, either with a commercially available blood gas syringe, or using a 1- or 3-mL syringe coated with 1:1000 heparin solution. Care should be taken to ensure heparin accumulates only in the hub of the needle after coating, as heparin dilution of samples falsely decreases pH and Pco2 and increases Po2 . Exposure to air will falsely increase pH and Pco2 and decrease Pa co2 .32 Once the samples have been collected, any air bubbles should be evacuated and the sample sealed, either with a cap or by placing the needle into a blood collection tube stopper. Samples are ideally analyzed immediately, but if not they may be stored in an ice-water bath for up to 30 min (1-mL sample) or 2 h (3-mL sample).32 Samples that are left at room temperature will have a decreased pH and Pa o2 , and an increased Pco2 , due to ongoing erythrocyte metabolism (anaerobic glycolysis). Traditional blood gas analysis adjusts the sample to body temperature. The importance of temperature correction of samples remains controversial. One theory suggests that the unadjusted results are more useful as a guide to therapy, as the blood pH will vary physiologically with the patient’s temperature in order to maintain normal ionization of molecules.33, 34 However, normal values of blood pH for extremes in temperature have not been established, and rectal temperature can vary significantly from core body temperature. In the majority of clinical equine neonatal cases the difference between corrected and uncorrected values is probably not significant enough to affect the course and response to treatment. The theory and practice of interpreting blood gas results based on uncorrected values is widely accepted.33
Venous oxygen tension (P v o2 ) Venous oxygen tension Pv o2 provides clinically useful information in assessing only one variable – the adequacy of oxygen delivery to tissue. It does not provide information pertaining to pulmonary
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function. A jugular venous sample provides an estimate of global tissue oxygenation, which is normally in the range of 40 to 50 mmHg in animals breathing room air. The venous oxygen tension may be low in situations of either reduced tissue oxygen delivery or increased tissue oxygen demand. A Pv o2 value of less than 30 mmHg warrants further investigation into the cause of inadequate tissue oxygenation and appropriate therapy instituted. Consideration of the foal’s hematocrit, adequacy of cardiac output, and arterial oxygenation are all important in order to treat a low Pv o2 appropriately.24 Other methods of assessing tissue oxygenation include comparison of the difference between arterial and venous oxygen (normally < 50 mmHg), and arterial and venous saturation (normally < 30%).12 Blood gas analysis of simultaneous arterial and venous blood samples is required for such comparisons, and can be particularly useful in determining the need for whole blood transfusions in foals. The oxygen extraction ratio (O2 ER) represents the fraction of oxygen delivered to tissues in relation to the oxygen utilized by the tissues. It may be estimated by the following equation: O2 ER = Sa o2 − Sv o2 /Sa o2 where Sa o2 is the arterial oxygen saturation of hemoglobin and Sv o2 is the venous oxygen saturation of hemoglobin. An oxygen extraction ratio greater than 30% is a marker of significant tissue oxygen debt, and values greater than 50% indicate clinically severe tissue hypoxia.26
Arterial blood pressure Arterial blood pressure (ABP) monitoring is routine practice in many NICUs, allowing recognition of cardiovascular system derangements and titration of therapies such as intravenous fluids, colloids, and vasopressor and inotropic agents. Direct ABP can be monitored accurately and continuously via cannulation of a peripheral artery, most commonly the dorsal metatarsal artery. Direct methods are less practical for monitoring non-anesthetized foals, although recumbent semi-comatose foals may be managed for days with an arterial catheter in place. Several cost-effective, indirect, automated oscillometric blood pressure devices are available to veterinarians. The device used should report mean arterial pressure (MAP), in addition to systolic and diastolic pressure, as MAP is more important for organ and tissue perfusion. Bladder cuff size and position are important factors influencing the accuracy of indirect
Figure 18.11 Determination of indirect arterial blood pressure using a portable, non-invasive monitor. The cuff is placed snugly around the coccygeal artery.
ABP measurement in foals. Indirect oscillometry using a cuff placed over the coccygeal artery (Figure 18.11) or dorsal metatarsal artery has been shown to be an acceptable method for measuring MAP in foals.35 The cuff bladder width to appendage circumference ratio should be between 25 and 50%. Loose cuff placement, or those with bladder widths that are too small, are more likely to result in falsely high blood pressure readings, while cuffs that are too tight or those with bladder widths that are too wide may produce falsely low readings.24, 35 To improve accuracy, the cuff should also be placed at the level of the heart, with minimal to no movement, the measured heart rate should be correct, and MAP measurements repeated three times and the average value recorded. Indirect blood pressure reportedly ranges between 60 and 120 mmHg for MAP for normal neonatal foals.24 Lower values for MAP may be observed in premature foals and in the immediate postpartum period. Critically ill foals with MAP values less than 65 mmHg may require urgent therapeutic intervention to increase cardiac output and tissue oxygenation. Unfortunately, the correlation between cardiac output and blood pressure in neonatal foals is poor.36 MAP is also a less accurate indicator of blood flow when systemic vascular resistance is altered, such as during sepsis, shock, and perinatal hypoxia. Attempting to obtain an indirect blood pressure in a hypothermic, vasoconstricted neonate with signs of poor perfusion is unlikely to produce reliable results. MAP monitoring should always be assessed in conjunction with other markers of global tissue perfusion, such as venous oxygen saturation (Sv o2 ), blood lactate, urine output, and physical examination findings.
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Critical Care – Monitoring Table 18.2 Normal range of selected variables for neonatal foals ≤ 2 weeks of age 3. Monitoring variable
Units
Normal range
Packed cell volume Plasma protein Colloid oncotic pressure Blood glucose Venous blood lactate Blood gas analysis: pH P a o2 P a co2 HCO3 − Pulse oximetry Mean arterial blood pressure Urine specific gravity
% g/dL (g/L) mmHg mg/dL (mmol/L) mmol/L
28–44a 4.0–8.1 (40–81)a 13–22b 65–180 (3.6–10.0)c < 2.5d
mmHg mmHg mmol/L % mmHg
7.35–7.45 60–90e 43–48e 22–29 > 96 69–111f
4.
5.
6.
7.
1.001–1.010
P a o2 , arterial oxygen tension; P a co2 , arterial carbon dioxide tension. Sources: a Koterba et al.37 b Magdesian et al.2 c Russell et al.19 d Magdesian and Madigan3 e Wilkins31 f Corley10
Conclusions With some or all of the monitoring equipment discussed, the equine veterinarian can perform a thorough and accurate evaluation of the hemodynamic, oxygenation, and metabolic status of the critically ill foal using portable, practical techniques within the first 20 to 30 minutes of examination. Table 18.2 lists the normal range of data in neonatal foals (< 2 weeks old) for the specific monitoring techniques discussed. In conjunction with a complete physical examination, a quantitative assessment provides significantly more information from which further, informed management decisions can be made.
8.
9.
10.
11.
12. 13.
14.
15.
16.
Acknowledgement 17.
The author would like to thank Dr Jenni Read at the Equine Medical Centre of Ocala, Florida, USA for her assistance with the critically ill foal monitoring photography in this chapter.
References 1. Harvey JW, Asquith RL, McNulty PK, Kivipelto J, Bauer JE. Haematology of foals up to one year old. Equine Vet J 1984;16:347–53. 2. Magdesian KG, Fielding CL, Madigan JE. Measurement of plasma colloid osmotic pressure in neonatal foals under intensive care: comparison of direct and indirect methods and
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the association of COP with selected clinical and clinicopathological variables. J Vet Emerg Crit Care 2004;14:108–14. Magdesian KG, Madigan JE. Volume replacement in the neonatal ICU: crystalloids and colloids. Clin Tech Equine Pract 2003;2:20–30. Evans DL, Golland LC. Accuracy of Accusport for measurement of lactate concentrations in equine blood and plasma. Equine Vet J 1996;28:398–402. van Oldruitenborgh-Oosterbaan M, van den Broek E, Spierenburg A. Evaluation of the usefulness of the portable device Lactate Pro for measurement of lactate concentrations in equine whole blood. J Vet Diagn Invest 2007;20:83–5. Tennent-Brown B, Wilkins PA, Lindborg S, Russell G, Boston R. Assessment of a point-of-care lactate monitor in emergency admissions of adult horses to a referral hospital. J Vet Int Med 2007;21:1090–8. Saulez MN, Cebra CK, Dailey M. Comparative biochemical analyses of venous blood and peritoneal fluid from horses with colic using a portable analyser and an in-house analyzer. Vet Rec 2005;157:217–23. Schulman ML, Nurton JP, Guthrie AJ. Use of the Accusport semi-automated analyzer to determine blood lactate as an aid in the clinical assessment of horses with colic. J S Afr Vet Assoc 2001;72:12–17. Magdesian KG. Blood lactate levels in neonatal foals: Normal values and temporal effects in the post-partum period. J Vet Emerg Crit Care 2003;13:174 (abstract). Corley KTT. Monitoring and treating haemodynamic disturbances in critically ill neonatal foals. Part 1: Haemodynamic monitoring. Equine Vet Educ 2002;14:270–9. Moore JN, Owen RR, Lumsden JH. Clinical evaluation of blood lactate levels in equine colic. Equine Vet J 1976;8:49–54. Divers TJ. Monitoring tissue oxygenation in the ICU patient. Clin Tech Equine Pract 2003;2:138–44. Corley KTT, Donaldson LL, Furr MO. Arterial lactate concentration, hospital survival, sepsis and SIRS in critically ill neonatal foals. Equine Vet J 2005;37:53–9. Boysen S. Lactate and the critically ill patient. Monitoring glucose in the ICU/NICU. In: Proceedings of the 13th Annual International Veterinary Emergency and Critical Care Symposium, 17–21 September, San Antonio, Texas, 2006. Latson KM, Nieto JE, Beldomenico PM, Snyder JR. Evaluation of peritoneal fluid lactate as a marker of intestinal ischemia in equine colic. Equine Vet J 2005;37:342–6. Delesalle C, Dewulf J, Lefebvre RA, Schuurkes JAJ, Proot J, Lefere L, Deprez P. Determination of lactate concentrations in blood plasma and peritoneal fluid in horses with colic by an Accusport analyzer. J Vet Int Med 2007;21:293–301. Vaala W. How to stabilize a critical foal prior to and during referral. In: Proceedings of the 45th American Association of Equine Practitioners Annual Convention, November 26–29, San Antonio, Texas, 2000. Corley KTT. Tank half full/half empty? Monitoring glucose in the ICU/NICU. In: Proceedings of the 13th Annual International Veterinary Emergency and Critical Care Symposium, 17–21 September, San Antonio, Texas, 2006. Russell C, Palmer JE, Boston RC, Wilkins PA. Agreement between point-of-care glucometry, blood gas and laboratory-based measurement of glucose in an equine neonatal intensive care unit. J Vet Emerg Crit Care 2007; 17:236–42. Vaala W. Peripartum asphyxia syndrome in foals. In: Proceedings of the 45th American Association of Equine
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Practitioners Annual Convention, December 5–8, Albuquerque, New Mexico, 1999. Van den Berghe G, Wouters P, Weekers F, Verwaest C, Bruyninckx F, Schetz M, Vlasselaers D, Ferdinande P, Lauwers P, Bouillon R. Intensive insulin therapy in critically ill patients. N Engl J Med 2001;345:1359–67. Edwards DJ, Brownlow MA, Hutchins DR. Indices of renal function: values in eight normal foals from birth to 56 days. Aust Vet J 1990;67:251–4. Corley KTT. Monitoring and treating the cardiovascular system in neonatal foals. Clin Tech Equine Pract 2003;2:42–55. Magdesian KG. Monitoring the critically ill equine patient. Vet Clin N Am Equine Pract 2004;20:11–39. Dunkel B, Palmer JE, Olson KN, Boston RC, Wilkins PA. Uroperitoneum in 32 foals: influence of intravenous fluid therapy, infection, and sepsis. J Vet Intern Med 2005;19:889–93. Orton CE. The respiratory system. In: Wingfield WE, Raffe MR (eds) The Veterinary ICU Book. Wyoming: Teton NewMedia, 2002; pp. 281–97. Hopper K. Assessment of oxygenation. In: Proceedings of the 12th Annual International Veterinary Emergency and Critical Care Symposium, 17–21 September, San Antonio, Texas, 2006. Stewart JH, Rose RJ, Barko AM. Respiratory studies in foals from birth to seven days old. Equine Vet J 1984;16:323–8. Rose RJ, Rossdale PD, Leadon DP. Blood gas and acidbased status of spontaneously delivered, term induced
30.
31.
32.
33.
34.
35.
36.
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and induced premature foals. J Reprod Fertil Suppl 1982: 521–8. Madigan JE, Thomas WP. Cardiopulmonary function in normal newborn foals from 1-14 days of age. In: Proceedings of the 4th Annual American College of Veterinary Internal Medicine Forum, Washington DC, 1984. Wilkins PA. Acute respiratory failure: Diagnosis, monitoring techniques, and therapeutics. Clin Tech Equine Pract 2003;2:56–66. Smarick S. Blood gas analysis for beginners. In: Proceedings of the 13th Annual International Veterinary Emergency and Critical Care Symposium, 26–30 September, New Orleans, Louisiana, 2007. Wingfield WE. Accidental hypothermia. In: Wingfield WE, Raffe MR (eds) The Veterinary ICU Book. Jackson, Wyoming: Teton NewMedia, 2002; pp. 1117–29. Hardy J. Monitoring the equine patient: Pulse oximetry and more. In: Proceedings of the 12th Annual International Veterinary Emergency and Critical Care Symposium, 17–21 September, San Antonio, Texas, 2006. Giguere S, Knowles HA, Valverde A, Bucki E, Young L. Accuracy of indirect blood pressure measurement of blood pressure in neonatal foals. J Vet Int Med 2005;19:571–6. Corley KTT. Monitoring and treating the cardiovascular system in neonatal foals. Clin Tech Equine Pract 2003;2: 42–55. Koterba AM, Drummond WH, Koseh PC. Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990.
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Chapter 19
Critical Care – Treatment Jane E. Axon
Emergency treatment of the critically ill neonate is aimed at stabilizing the cardiovascular and respiratory system to ensure good tissue perfusion and oxygenation. The underlying diseases are then treated; however, some have no specific treatment thus therapy is aimed at supporting the neonate to allow damaged tissues to repair and the final stages of transition from fetus to foal to occur. Many treatments are able to be carried out on the farm by experienced dedicated personnel; however, some do require the expertise, equipment, and personnel that are available only at an intensive care unit (ICU).
Respiratory support The majority of critically ill neonates need some form of respiratory support. They spend more time recumbent and may have a weak ribcage, less compliant lungs, poor perfusion, and/or poor central control of respiration. Foals which have an elevated respiratory rate, cyanotic mucous membranes, increased respiratory effort, Pa O2 of < 60 mmHg, or Sa O2 < 90% should be started on intranasal oxygen (INO2 ) insufflation (Figure 19.1). Recumbent foals also benefit from INO2 insufflation as lateral recumbency can reduce Pa O2 and the stress of manipulating an already compromised neonate can be fatal.1 Therapy is aimed at maintaining the Pa O2 between 80–110 mmHg, Sa O2 > 92%, decreasing respiratory effort and rate and improving mucous membrane color and demeanor of the foal.2 The foal begins on 5 L/min and the rate is adjusted according to response. Oxygen should be run through a water-filled bubble humidifier (using sterile water not saline) to avoid drying of the airways. If a response is not seen other treatments are instigated. Other respiratory treatments Respiratory stimulants are useful in foals that hypoventilate secondary to a blunted central CO2 recep-
tor sensitivity which occurs most commonly in foals with hypoxic ischemic syndrome (HIS).2 This is seen clinically as irregular breathing patterns with periodic apnea or inappropriately low respiratory rates. Caffeine is the most commonly used pharmacological respiratory stimulant; however, continuous intravenous infusion of doxapram is also utilized.3 Mechanical or assisted ventilation is usually indicated when the Pa CO2 is greater than 70 mmHg. The Pa CO2 , however, needs to be interpreted with consideration of the pH and bicarbonate concentrations to ensure the hypercapnia is not buffering a metabolic alkalosis. Temporary assisted ventilation can be achieved with manual ventilation using a selfinflating resuscitation bag with an attached source of humidified oxygen. Sustained ventilation is achieved with mechanical ventilation.2 The use of mechanical ventilation requires careful continual monitoring of the foal’s respiratory function with the frequent use of arterial blood gas and capnograph results, and the foal’s clinical response to ventilating and the different ventilator settings.2 Nitric oxide and sildenafil have been used in conjunction with mechanical ventilation to provide smooth muscle relaxation of the pulmonary vessels in foals with pulmonary hypertension.2, 4 Nebulization therapy can be utilized to promote secretion removal and administer local antibiotic therapy.5 Ultrasonic nebulizers that disperse droplets less than 5 µm in diameter should be used. Saline alone is very useful at moistening secretions and, together with coupage of the chest, promotes secretion removal. Bronchodilators should be used with caution in critically ill neonates due to potential cardiac effects and alterations that can be caused in the ventilation : perfusion ratio. Methylxanthines are not recommended due to their narrow therapeutic index. Aerosolized β 2 -agonist bronchodilators (salbutamol, clenbutarol) can be administered 5–10 minutes prior to nebulizing to assist with bronchodilation and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Specific Diagnostic and Management Techniques sodium is also different.6 The foal’s diet (milk) is low in sodium and so the neonatal foal has developed renal mechanisms to conserve sodium. Thus the neonatal foal is prone to sodium overloading, especially when treated with sodium–based crystalloids. However, in a neonate with renal tubular damage there may be excessive sodium loss, thus ideally urinary excretion of sodium should be evaluated. Types of fluids
Figure 19.1 INO2 insufflation. The tubing is measured from the medial canthus of the eye to the nares. It is then taped around half a tongue depressor (arrowed) and connected to tubing supplying humidified oxygen. The tongue depressor is taped to the foal’s muzzle.
delivery of the antimicrobials to the lower airways. The aerosol route allows the use of smaller doses thus decreases the risks of systemic toxicity. Foals that have significant upper airway obstruction may require temporary nasotracheal or pharyngeal tube placement until the obstruction has resolved. This however may increase the chance of secondary lung infection, thus broad-spectrum prophylactic antimicrobial therapy should be instigated.
Balanced electrolyte solutions are the best fluids for correcting volume deficits and to improve hypoperfusion caused by hypovolemia except in conditions with hyperkalemia, hyponatremia, and hypochloremia.6 For these foals 0.9% NaCl or 1.8% NaCl are best for correcting hypovolemia. Hypertonic saline (7–7.5% NaCl) has no role in resuscitation of neonates due to the rapid change in plasma osmolarity it causes and resultant central nervous system (CNS) damage; 0.9% NaCl is also not recommended for correcting hypovolemia as it is an acidifying solution and has a large sodium load. Synthetic colloids have a questionable role in volume resuscitation in the neonate.6, 7 One major concern is the increased movement of the colloid molecules through damaged endothelial barriers into the interstitium which promotes edema formation. Plasma is by far the most commonly used colloid and may be utilized to improve oncotic pressure in the treatment of hypovolemia, as immunotherapy, for coagulation factors as well as other benefits. As anaphylactic reactions may occur it should be given slowly; thus it is not recommended to be used in acute resuscitation but is given after the foal’s condition has stabilized. Initial fluid therapy
Intravenous fluid therapy Intravenous fluid therapy is essential in the supportive care of the critically ill neonate. Deciding what volume and type of fluid to use requires an understanding of the neonate’s physiology and interpretation of clinical signs and laboratory data. Hypovolemia in most critically ill neonates is due to poor vascular tone unless extra fluid losses such as diarrhea or decreased intake are present.6 Many hypovolemic, critically ill neonates presented in the first 48 hours are actually overhydrated since neonates subjected to intrauterine stresses undergo intrauterine fluid shifts which overload the interstitial space.6 Critically ill neonates also have an increased vascular permeability and are more predisposed to edema formation as the vessels are more permeable to plasma proteins and colloids.6 The neonate’s ability to handle
Initial fluid therapy is aimed at improving tissue perfusion by correcting hypovolemia, providing glucose support and passive immunity, and correcting electrolyte abnormalities. The foal initially is administered fluids to correct hypovolemia, and is then maintained on glucose therapy which provides both glucose and fluid support. Plasma is used to provide passive immunity and to assist with improving oncotic pressure.
Fluid therapy for hypovolemia Fluid boluses of warmed balanced electrolyte solutions at 20 mL/kg over 20 minutes with re-evaluation of perfusion after each bolus are recommended.6, 7 Improved perfusion, thus hypovolemia, is seen clinically as improved pulse quality, improved blood
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Critical Care – Treatment pressure, warm legs, return of borborygmi, urine production, and improved mentation. A maximum 4 L/50 kg is given. If no improvement is seen, inotrope and pressor therapy needs to be commenced. Foals which have uncontrolled bleeding should have the fluids given cautiously to avoid exacerbation of the bleeding. Colloid therapy needs to be considered as an adjunct to the crystalloids if the protein concentration is less than 35 g/L.
Fluid therapy to improve hydration After hypovolemia has been treated, dehydration should be evaluated. An estimation of fluid replacement for dehydration is similar to adults; however, parameters such as skin tenting, tear film production, sunken eyes, and mucous membrane moisture which are used in adults are not as reliable in foals. Half the volume to be replaced should be replaced over the first 6 hours and the remainder over the next 6 hours with maintenance requirements and ongoing losses.
Fluid therapy to provide glucose support All critically ill neonates benefit from exogenous glucose support. There are two approaches to glucose therapy.6, 8 1. Supply glucose at a rate equal to that supplied by the placenta which is 6.8 mg/kg/min (3.78 mmol/kg/min). 2. Supply glucose to meet the sick and recovering foal’s resting energy requirements which have been estimated at 44 kcal/kg/day. The author utilizes the glucose supplementation based on placental supply. The initial rate of supplementation begins at 5 mg/kg/min which is altered depending on the neonate’s response (Table 19.1). If the neonate has severe hypoglycemia (< 2.8 mmol/L; < 50 mg/dL), supplementation begins at a higher rate. As the volume of fluid required for the glucose supplementation often exceeds the fluid requirement of the foal, 10% glucose is used (Table 19.1). The response to glucose supplementation varies with each individual’s physiological condition. Glucose supplementation may result in hyperglycemia which can resolve with decreasing the rate of supplementation; however, insulin therapy maybe necessary in neonates with inappropriate insulin responses. High rates of insulin therapy may be needed in foals with sepsis. It is critical that blood glucose concentration is monitored closely (every 1–2 hours) until the glucose concentrations have stabilized. Glucose solutions should not be bolus dosed as the resultant hyperglycemia may result in glucosuria with loss of
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Table 19.1 Initial fluid rates for the critically ill neonate Fluid therapy for hypovolemia 20 mL/kg × 50 kg = 1 L Hartmann’s over 20 minutes Reassess Repeat up to 3 times if needed. Maximum of 80 mL/kg (4 L for 50 kg foal) Glucose supplementation 5 mg/kg/min 5 mg × 50 kg × 60 min = 15 000 mg/h = 15 g/h = 150 mL/h of 10% glucose Fluid therapy for maintenance 1. 2–4 mL/kg/h 2. Holliday–Segar formula (‘dry’ maintenance formula) 100 mL/kg/day for first 10 kg of body weight = 100 mL/kg/day + 50 mL/kg/day for second 10 kg of body weight = 50 mL/kg/day + 25 mL/kg/day for the remainder of the body weight Thus for a 50 kg foal: (100 mL/kg × 10) + (50 mL/kg × 10) + (25 mL/kg × 30)/day Total: 2250 mL/day or 94 mL/h
infused electrolyte, fluids, and glucose. In the field situation, 20 mL of 50% glucose can be added to a liter of fluids and administered over an hour and, in suspected cases of severe hypoglycemia, 50 mL of 50% glucose can be added.8
Electrolytes Electrolyte replacement or restriction should be considered if life-threatening abnormalities are present; however, many abnormalities will correct after appropriate fluid therapy and improved tissue perfusion. A newborn neonate’s electrolyte abnormalities are a reflection of the in-utero environment and usually do not require correction as they will correct over the next 24 hours if renal function is normal. Oral supplementation is utilized in adjunct to intravenous supplementation if the gastrointestinal tract is functional. Hyperkalemia most commonly occurs with uroperitoneum in neonates. It may also be seen during a clinical episode of hyperkalemic periodic paralysis in a homozygote foal. If hyperkalemia is suspected, non-potassium fluids such as 0.9% NaCl and 5% glucose should be used. Hypokalemia is not uncommon in critically ill neonates. If potassium is < 2.5 mEq/L intravenous supplementation should be given. Potassium supplementation can be estimated by: potassium replacement = 0.4 × body weight (kg) × potassium deficit. The potassium can be added to the glucose as long as the rate of administration is no greater than 0.5 mEq/kg/h (25 mEq/h for 50 kg foal). Both hypo- and hypernatremia can have fatal consequences if not corrected properly. The
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underlying cause and change to the extracellular fluid volume should be determined so that correct therapy is instigated.9 Sodium concentrations can be corrected fairly rapidly up to 115 mEq/L in acute cases, especially if accompanied by seizures, and then corrected at approximately 0.5 mEq/h.10 This can be done using polyionic replacement solutions, 0.9% NaCl, or hypertonic solutions of NaCl. In cases of hypernatremia the sodium concentration should also be decreased slowly (no greater than 0.5 mEq/h). Sodium levels should be monitored closely during correction and fluids administered by an infusion pump. It is difficult to accurately predict the source of an acid–base disturbance in the neonate without blood gas analysis. The bicarbonate measurements are influenced by strong ions (sodium, chloride, potassium, lactate, and unidentified anions) and may compensate respiratory abnormalities. Thus correction of the acid–base disturbance is aimed at correcting the primary problem. In cases of compensatory metabolic alkalosis, improving ventilation will help reduce the metabolic compensation. In cases of diarrhea, correction of electrolyte concentrations and improving tissue perfusion will help reduce the metabolic acidosis. Maintenance fluid therapy The volume of fluid required to maintain the neonate will vary with each individual depending on the water losses, systemic disease, and renal function. So there is no formula that fits all. There are currently two different approaches. 1. Higher rates: 3–5 mL/kg/h9 or 4–6 mL/kg/h.11 These rates may predispose to overhydration and edema formation if there is poor renal function. 2. A ‘dry’ fluid rate (Holliday–Segar formula). This formula provides the maintenance needs of neonates and avoids overhydration.6 The author uses the Holliday–Segar formula in neonates that have poor renal function or initially in neonates less than 3 days old where renal function is not known. A higher fluid rate of 2–4 mL/kg is used in foals with adequate renal function as determined by appropriate urine specific gravity, volume of urine, lack of edema formation, and normal renal laboratory values. Estimated ongoing losses of fluid that can occur with diarrhea or reflux are then added to the maintenance volume. The type of maintenance fluid used is very important, especially in the neonate less than 3 days old as sodium overloading can occur. A liter of balanced electrolytes or plasma contains a day’s sup-
ply of sodium for a foal. The author sodium-restricts critically ill neonates less than 3 days old and uses urine specific gravity, volume, and frequency of urine to determine continuing therapy. Maintenance formations such as Normosol-M and Plasma-Lyte 56 have low sodium concentrations. The author uses a 50:50 combination of 10% glucose with a balanced electrolyte solution. Foals with diarrhea usually have associated sodium loss, thus should initially not be sodium restricted.
Inotrope and pressor therapy Inotrope and pressor therapy are utilized in the critically ill neonate to improve tissue perfusion if fluid loading fails. The drugs should be administered as constant rate infusions and clinical responses monitored closely. Dobutamine is a commonly used inotrope and is often combined with norepinephrine if tissue perfusion has not been obtained and pressor therapy is required.12 Dopamine, norepinephrine, and vasopressin are also used in inopressor therapy.
Immune system support This can be given by colostrum or plasma. There are a variety of other products available, such as lyophilized IgG or bovine colostrum, which should not be solely relied upon for immune support. Colostrum is ideal if the neonate has a functional gastrointestinal tract and is less than 12 hours old. When gastrointestinal function is questionable, intravenous plasma should be used.
Nutrition Adequate nutritional support of the critically ill neonate is essential. If the gastrointestinal tract (GIT) is functional and there is no medical contraindication for oral feeding, enteral nutrition is the preferred route. Feeding is withheld until the foal’s temperature is greater than 37.5◦ C, borborygmi are present, meconium is passed, and tissue perfusion has improved. If foals have an effective suckle reflex, nursing should be encouraged and closely monitored. If the foal has an effective suckle but is not nursing from the mare, a bottle or bucket can be used. If the foal does not have an effective suckle the foal is fed via an indwelling nasogastric tube (NGT) (Figure 19.2). The foal is begun on a milk intake of 5% of the bodyweight/day. This is divided into feeds that are given every 2 hours. If this is tolerated the amount is gradually increased to reach a volume of 15–20% of their bodyweight/day. If the GIT is not functional or there is a medical condition which prevents enteral feeding,
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therapies described for HIS; however, their efficacy in neonates has not been proven14, 15 (Table 19.2). Diazepam is the most useful drug for emergency control of seizures; however, its effect is short-lived and repeated dosing can cause respiratory depression. If seizure activity continues, phenobarbital or midazolam can be used (Table 19.2). The half-life of phenobarbital is about 100 hours and it can cause marked respiratory depression and hypothermia and may exacerbate hypoperfusion. Midazolam has the advantage over phenobarbital as it has a shorter half-life and is given in a continuous infusion that can be decreased according to the foal’s response. It is especially useful in hyperactive foals from cesarean section delivery, where the side effects of phenobarbital are of concern. In some cases, the author combines phenobarbital with the midazolam which can provide better seizure control at lower doses of each drug.
Gastrointestinal system support
Figure 19.2 Feeding a foal via a nasogastric tube. The foal should be refluxed prior to feeding. Milk is fed in by gravity flow and the foal is fed, supported in sternal recumbency.
parenteral nutrition (PN) is required. Part or all the foal’s nutritional requirements are provided intravenously by a constant rate infusion. Different formulations are utilized that have a mixture of glucose, amino acids, lipids, vitamins, and/or trace minerals.13
Antimicrobial coverage Antimicrobial therapy is essential in septic and compromised foals. Therapy should be aimed at both Gram-negative and Gram-positive organisms and should be instigated pending culture results. Bactericidal drugs are favored due to the immaturity of the neonatal immune system (Table 19.2). The drug choice should be based on the sensitivity of isolates from the surrounding geographical area. Nephrotoxic drugs should be avoided in cases where renal function is not normal or is unknown.
Central nervous system support Maintaining cerebral perfusion with fluid replacement therapy and maintaining adequate blood pressure is the most important therapy for central nervous system (CNS) support. There are a variety of
Many of the therapies for gastrointestinal support are discussed in detail elsewhere in this text. Supportive therapy in correcting fluid and electrolyte deficits and providing pain relief is similar regardless of the etiology, and is often instigated prior to the primary cause being identified. Meconium impactions can often be relieved with gentle digital manipulation; however, usually a warm soapy-water enema or a commercial phosphate-based enema is required. If the impaction continues and abdominal distension develops, other therapies such as acetylcysteine enema, oral mineral oil, and feed restriction need to be utilized. Ileus and dysmotility are commonly seen as a secondary complication with sepsis, prematurity, HIS, or enteritis. Correction of the primary problem will result in improvement in gastrointestinal motility. ‘A tincture of time’ is the most important factor in improving gastrointestinal function in the critically ill neonate with HIS, sepsis, or prematurity. Prokinetics have been used but were often associated with intussusceptions, thus are not currently recommended. Lidocaine constant rate infusions have been used by the author to assist with providing pain relief and resolving the ileus in foals with severe enteritis (Table 19.2). There are many therapeutic interventions for foals with enteritis, including intravenous fluids and electrolyte supplementation, correction of hypoproteinemia, alleviation of pain and provision of caloric needs, which are similar regardless of the etiology. There are also a large variety of anti-diarrheal agents which have varying degrees of success. Broadspectrum antimicrobials should be administered to neonates with enteritis due to the high risk of
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Table 19.2 Common drug dosages used in the treatment of the critically ill neonatal foal14–16 Drug
Dose, route, and frequency
Comments
Foals < 14 days old: 25–30 mg/kg i.v. s.i.d. Foals > 14 days old: 20–25 mg/kg i.v. s.i.d. High doses may be required. Therapeutic drug monitoring should be performed
Ensure adequate hydration. Nephrotoxic. TDM: 30 min peak: 50–60 µg/mL 8 h trough: < 15–20 µg/mL 12 h trough: < 5–7 µg/mL
Ceftiofur
5 mg/kg i.v. b.i.d. 10 mg/kg i.v. q.i.d. (broader Gram-negative spectrum) CRI 1.5 mg/kg/h
3rd gen. cephalosporin, not typical action: no CNS penetration. Suitable for i/m
Cefuroxime
30 mg/kg/day p.o. b.i.d./t.i.d. 200–240 mg/kg/day: meningitis 10 mg/kg p.o. q.i.d. 25 mg/kg i.v. b.i.d. 10 mg/kg p.o. t.i.d./b.i.d. 1 mg/kg i.v. b.i.d. 50 mg/kg p.o. q.i.d. 10 mg/kg p.o. b.i.d. 7.5 mg/kg p.o. s.i.d.; 5.5 mg/kg i.v. s.i.d. Foals < 14 days old: 11–15 mg/kg i.v. s.i.d. Foals > 14 days old: 8.8 mg/kg i.v. s.i.d. High doses may be required. Therapeutic drug monitoring should be performed
2nd gen. cephalosporin
Antimicrobials Amikacin sulfate
Cephalexin Ceftriaxone Cefpodoxime Cefquinome Chloramphenicol palmitate Doxycycline Enrofloxacin Gentamicin
1st gen. cephalosporin 3rd gen. cephalosporin 3rd gen. cephalosporin 4th gen. cephalosporin Public health concerns May cause tendon laxity Can cause arthropathy in foals Ensure adequate hydration, nephrotoxic. TDM: 30 min peak: 32–40 µg/mL 8 h trough: < 4–5 µg/mL 12 h trough: < 2–3 µg/mL
Metronidazole
10–15 mg/kg p.o. or i.v. t.i.d. (q.i.d. if severe clostridiosis) 25 mg/kg p.o. b.i.d.
Oxytetracycline
Osteomyelitis: 10 mg/kg i.v. b.i.d. Contracted tendon: 44 mg/kg i.v. s.i.d. for maximum of 3 days
Nephrotoxic. Can cause problems when given quickly so give slowly
Potassium/sodium penicillin Rifampin Ticarcillin and clavulonic acid R (Timentin )
20 000–50 000 IU/kg i.v. q.i.d. 5 mg/kg p.o. b.i.d. 50–100 mg/kg i.v. q.i.d. 2–4 mg/kg/h CRI
Use upper dose Use with other antibiotics
Trimethoprim potentiated sulfa
30 mg/kg p.o./i.m./i.v. b.i.d.
Dose/kg is combined trimethoprim and sulpha concentration
Diazepam
0.05–0.4 mg/kg i.v. (5–10 mg bolus for 50 kg foal)
Sedation. Short-acting seizure control. Respiratory depression with repeated use
Midazolam
Loading dose: 0.05–0.2 mg/kg i.v. (5 mg/50 kg) Maintenance: 0.02–0.1 mg/kg/h
Usual therapeutic range 0.03–0.06 mg/kg/h
Phenobarbital
2–10 mg/kg i.v. s.i.d./b.i.d. (usually 2–4 mg/kg) Over 20 min to effect
Long T1/2 . Causes respiratory depression, hypothermia, hypoperfusion
Phenytoin
5–10 mg/kg p.o./i.m./p.o. q 2–4 hr
Anticonvulsants
Gastrointestinal Acetylcysteine retention enemas
8 g acetylcysteine with 200 mL water
Ensure foal sedated, lateral recumbency, not straining, elevated behind. 30 cc balloon Foley catheter
Bismuth subsalicylate Omeprazole
0.5–2 mL/kg p.o. q 4–6 h Treatment need 4 mg/kg p.o. s.i.d. Prevention: 2 mg/kg p.o. s.i.d. 2 mg/kg p.o. q.i.d. 6.6 mg/kg p.o. t.i.d. 3000 IU p.o. q 4 h
Causes dark feces Proton pump inhibitor
Loading dose: 1.3 mg/kg over 5–10 min Maintenance: 0.05 mg/kg/min 10–20 mg/kg p.o. t.i.d./q.i.d.
CNS signs with toxicity which resolve with discontinuing CRI Mucosal protectant
Ranitidine Lactase Lidocaine Sucralfate
H2 receptor antagonist
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Table 19.2 (Continued) Drug Inotropes/Pressors Dobutamine Dopamine Noradrenaline Vasopressin
Dose, route, and frequency
Comments
2–20 µg/kg/min Starting dose: 2 µg/kg/min 5–25 µg/kg/min Starting dose: 5 µg/kg/min 0.1 µg/kg/min – 5 µg/kg/min Starting dose: 0.1 µg/kg/min 0.25–1.0 mU/kg/min
Respiratory stimulants Caffeine
Loading dose: 10 mg/kg p.o./p.r. Maintenance: 2.5 mg/kg p.o./p.r. s.i.d./b.i.d.
Doxapram infusion Miscellaneous Furosemide Insulin (regular insulin)
400 mg total dose infused at 0.05 mg/kg/min
Nitric oxide Sildenafil 7.5% glucose (75 mg/mL) 10% glucose (100 mg/mL) 25% glucose (250 mg/mL) Fluid drip rates Medications used with hypoxic ischemic encephalopathy
Respiratory stimulant. Side effects: tachycardia, general stimulation
1 mg/kg CRI: 0.5–2.5 mg/kg/h CRI: 0.01 IU/kg/h starting dose, check BG q 2–4 h then double dose until effect
Use glass bottle. Prime lines for 20 min. Attach to TPN line
5–40 ppm 0.5–2.5 mg/kg p.o. up to q 4 h 50 mL 50% glucose to 1 L 5% glucose 111 mL 50% glucose to 889 mL 5% glucose 445 mL 50% glucose to 555 mL 5% glucose 100 mL/h = 8 drops per 15 sec 20 drop/mL infusion set Dose
Action
Allopurinol
44 mg/kg p.o. within 4 hours of birth
Xanthine oxidase inhibitor, prevention of free radical formation
Ascorbic acid Dimethyl sulfoxide (DMSO) Magnesium sulfate Mannitol
10–100 mg/kg/day p.o. 0.5–1 g/kg p.o. or i.v. as 10% solution q 12–24 h 50 mg/kg/h infusion for 1st hour then 25 mg/kg/h 0.25–1 g as 20% solution q 6–12 h as i.v. bolus over 20 min 5–10 mg/kg p.o. or i.v. s.i.d. 5000 IU p.o. s.i.d. 68 IU/50 kg p.o. s.i.d.
NMDA-receptor blockade, antioxidant Free oxygen radical scavenger NMDA-receptor blockade Osmotic agent for cerebral edema
Thiamine Vitamin E
Supports cerebral energy metabolism Antioxidant
BG, blood glucose. TDM, therapeutic drug monitoring.
translocation of bacteria across the compromised intestinal wall. Enteritis due to Clostridium spp is specifically treated with penicillin and metronidazole. Foals with salmonellosis should be treated with systemic antimicrobials according to sensitivity results of the organism. There is no specific treatment for cryptosporidia in foals. Due to the autoinfective stage of the life cycle, anti-diarrheal agents which prolong transit time should be avoided. Foals need to be monitored closely after the diarrhea has resolved. It is not uncommon for them to remain dull and ill thrifty as a result of transient malabsorption, continued electrolyte abnormalities or gastroduodenal ulcer syndrome. Anti-ulcer medication is not routinely used in critically ill neonates as it is believed that the gastric ul-
cers are secondary to poor perfusion, hypoxic damage and bacterial colonization. The acid environment also forms a natural barrier against pathogens.17 Sucralfate offers an alternative to use in neonates as it does not alter the gastric acidity and has mucosal healing properties. Anti-ulcer medication is however used in foals older than 7 days of age, proton pump inhibitors such as omeprazole being commonly utilized.
Renal support Many of the neonatal diseases result in altered renal blood flow patterns or acute tubular damage, thus paying particular detail to fluid therapy and sodium balance will decrease the work load on the
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kidneys. Often diligent fluid therapy and time alone is successful and avoids the potential problems associated with the use of diuretics. Dopamine and furosemide do not reverse acute renal failure but are useful in managing the fluid volume overload.14, 18
Umbilical treatment The majority of infected umbilical remnants are treated medically with broad-spectrum antimicrobial therapy. Fibrinogen concentrations are monitored and ultrasonographic examinations are performed to ensure resolution. If there is no response an empirical antimicrobial change may be necessary. Occasionally when there is a discrete large (greater than 2.5 cm) abscess or there is a concern the remnant is a source of infection, surgical removal is recommended. A patent urachus is common occurrence and usually resolves with medical therapy. The foal is kept in a clean dry environment and continued on oral broad-spectrum antimicrobial therapy. Ultrasonographic examination is used to evaluate the umbilical remnants. Very rarely does the urachus remain patent for a prolonged period (greater than 4 weeks) and need surgical removal.
Musculoskeletal treatments There are a variety of methods of treating angular limb deformities and infectious orthopedic disease which can be found elsewhere in this text.
Ocular therapies Eyelid contusions are treated with topical ointment (avoiding contact with the eye) and the eye lavaged with warm saline to remove the debris. Topical antibiotic ointment is then applied until a complete examination is possible. Warm compressing may also assist in reducing the swelling. Recumbent neonates should have artificial tears placed in their eyes every 4–6 hours. Superficial corneal ulceration is treated initially with topical antibiotic ointment and if no response is seen or the eye is deteriorating corneal cytology and culture should be performed. Uveitis is usually associated with sepsis in the neonate and once the sepsis is under control the uveitis resolves. Atropine can be used to dilate the pupil. Systemic and topical corticosteroid therapy is not routinely recommended in the critically ill neonate and may predispose the neonate to developing corneal ulceration thus topical non-steroidal antiinflammatory medication can be considered.
Lower lid entropion is common in the critically ill neonate, either through dehydration or poor muscle tone. The author uses vertical mattress sutures along the entire lower lid with 4/0 polygalactin 910. The sutures are placed 3–5 mm from the eyelid margin ensuring the sutures extend to the medial and lateral canthus as often these corners continue to roll in.
Referral and transport The increasing awareness of owners and insurance companies to the level of care that can be provided at an intensive care unit that is not possible on a farm makes referral an important issue to discuss with owners and breeding farm managers. If the decision is made to refer the foal for intensive care, the foal should be referred as soon as possible. If recumbent, the foal is best transported in a car and the mare transported separately. The foal should be kept in sternal recumbency, on INO2 insufflation and wrapped in blankets to keep warm and protect it from self-trauma. Intravenous fluids with glucose may be recommended to be given if the referral center is several hours away. Early referral of the foal to the ICU improves the likelihood of survival.
References 1. Kosch PC, Koterba AM, Coons TJ, Webb AI. Developments in management of the newborn foal in respiratory distress. 1. Evaluation. Equine Vet J 1984;16(4):312–18. 2. Palmer JE. Ventilatory support of the critically ill foal. Vet Clin N Am Equine Pract 2005;21(2):457–86. 3. Giguere S, Sanchez LC, Shih A, Szabo NJ, Womble AY, Robertson SA. Comparison of the effects of caffeine and doxapram on respiratory and cardiovascular function in foals with induced respiratory acidosis. Am J Vet Res 2007; 68(12):1407–16. 4. Lester GD, DeMarco VG, Norman WM. Effect of inhaled nitric oxide on experimentally induced pulmonary hypertension in neonatal foals. Am J Vet Res 1999;60(10):1207– 12. 5. McKenzie HC. Treating foal pneumonia. Comp Cont Educ Pract Vet Equine 2006;Spring:47–53. 6. Palmer JE. Fluid therapy in the neonate: not your mother’s fluid space. Vet Clin N Am Equine Pract 2004;20(1): 63–75. 7. Magdesian KG, Madigan JE. Volume replacement in the neonatal ICU: crystalloids and colloids. Clin Tech Equine Pract 2003;2(1):20–30. 8. Corley KTT, Axon JE. Resuscitation and emergency management for neonatal foals. Vet Clin N Am Equine Pract 2005;21(2):431–55. 9. Spurlock SL, Furr M. Fluid therapy. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 671–700. 10. Magdesian KG. Neonatal foal diarrhea. Vet Clin N Am Equine Pract 2005;20(1):295–312.
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Critical Care – Treatment 11. Vaala WE. Supportive care of the abnormal newborn. In: Smith BP (ed.) Large Animal Internal Medicine, 4th edn. St Louis: Mosby, 2009; pp. 325–32. 12. Corley KTT. Ionotropes and vasopressors in adults and foals. Vet Clin N Am Equine Pract 2004;20(1):77–106. 13. Hansen TO. Nutritional support: parenteral feeding. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 747–62. 14. Bain FT. Management of the foal from the mare with placentitis: A clinician’s approach. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, Denver, CO, 2004; pp. 162–4.
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15. McKenzie HC, Furr MO. Aminoglycoside antibiotics in neonatal foals. Comp Cont Edu 2003;25(6):457–69. 16. Corley K. Appendix. In: Corley K, Stephen J (eds) The Equine Hospital Manual. Oxford: Blackwell Publishing, 2008; pp. 655–669. 17. Barr BS, Wilkins PS, DelPiero F, Palmer JE. Is prophylaxis for gastric ulcers necessary in critically ill equine neonates? A retrospective study of necropsy cases 1995–1999. In: Proceedings of the 18th Annual Meeting of the Veterinary Medicine Forum, 2000; p. 705. 18. Wilkins PA. Disorders of foals. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine. St. Louis, MO: Saunders, 2004; p. 1399.
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Chapter 20
Complications of Neonatal Infection Mary Rose Paradis
Introduction Perinatal infection is cited as one of the most common reasons for morbidity and mortality in the neonatal foal. In the general population, the morbidity ranges from 2.7% in the UK to 28% on a Standardbred farm in New York.1, 2 The incidence of infection in the equine neonate at tertiary care referral centers ranges from 26% to approximately 50% of the presenting newborn foals.3–5 Mortality studies have a similar finding, with a range of 26–33% of deaths in foals ranging from 24 hours to 10 days of age.6, 7 It is difficult to compare studies involving infections in the newborn because of different methods of diagnosis and description. An infection may be localized with very few systemic effects or it may be generalized with severe consequences of multiple organ dysfunction and death. The terms septicemia and sepsis are often used to describe foals where a positive culture is obtained from blood or there are multiple sites of infection in an animal. Systemic inflammatory response syndrome (SIRS) is a more encompassing term that is used describe the body’s exaggerated reaction to infection or other insults such as trauma, hypo/hyperthermia, or intense hypoxia. Foals may present with signs of SIRS but have a negative blood culture. Sepsis is SIRS with a positive blood culture.
Risk factors In utero, the equine fetus has some immunological capacity to respond to infection but it is not as responsive as an adult’s immune system.8 Diseases such as equine herpesvirus 1 infection or bacterial placentitis in the dam may result in prenatal infections that more seriously affect the foal than the mare.
Postnatal infections are often associated with inadequate colostral antibody transfer, otherwise known as complete or partial failure of passive transfer (FPT). The mare’s epitheliochorial placenta effectively prevents any in utero transfer of maternal antibodies to the foal. Consequently, the foal is born with no circulating immunoglobulin G (IgG) and only a small amount of its own IgM. It has only its innate immune system to prevent microbial colonization. Although the foal is able to mount an adaptive response to pathogens, there is a timelag that makes a newborn foal more susceptible to microbial invasion. Without some sort of ‘immunological assistance’ the foal is unprotected against environmental microorganisms that it will encounter at birth. Gastrointestinal absorption of colostrum from the dam is the primary source of this assistance. Any factor that may delay receipt of colostrum by a foal should be considered a risk factor for neonatal infection. Some causes for delayed colostral ingestion include prematurity, dysmaturity, dystocia, musculoskeletal abnormalities, neonatal encephalopathy, and perinatal asphyxia. Certain factors in the mare also put the foal at risk for FPT. Premature lactation, agalactia, premature delivery, induction of labor, and advanced maternal age may lower the amount of IgG in a mare’s colostrum. In utero infections in the mare are associated with an increased risk of infection in the foal. Even receiving the highest quality colostrum at birth will not prevent an infection that has established itself before birth. Delayed foal acceptance by maiden mares or occasional foal rejection may result in inadequate or delayed colostrum ingestion by the foal and is also a risk factor. Excessive environmental contamination may also increase the incidence of neonatal infection.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Complications of Neonatal Infection
Routes of entry There are several routes of entry for bacteria to invade the newborn foal. The three most common sites are the umbilicus, the gastrointestinal system, and the respiratory system. Any skin trauma or wounds can also allow entry of bacteria. When the umbilicus breaks naturally, the umbilical arteries and vein retract internally, and this may be important in decreasing the incidence of omphalophlebitis. Current ‘standard of care’ for the umbilicus involves disinfecting it with 0.5% chlorhexidine solution or 2% iodine/betadine solution.9 Gastrointestinal entry of pathogens is likely due to the ‘openness’ of the intestine in the first 12 hours of life. It is during this time that the gut allows absorption of colostral antibodies and other large-molecular-weight compounds. Hypothetically, this would also allow bacteria to cross the intestinal barrier.9 The respiratory tract is one of the most frequently affected sites for infection of the newborn. The innate immune system of the respiratory tract helps to prevent infection through the mucosal barrier with numerous ‘filters’ such as the nasal turbinates, the long trachea, and the ciliated respiratory tract lining cells in the adult.10 These barriers are less competent in the foal. An immature ciliary apparatus and the presence of fewer alveolar macrophages in neonates may lead to decreased bacterial clearance.11 Infections in the respiratory tract can be the result of direct inoculation, aspiration, or hematogenous spread from other sites.
Pathogens The predominant organisms involved in neonatal sepsis are Gram-negative bacteria, with Escherichia coli being the most frequently cultured.12, 13 Also prevalent are Klebsiella spp., Actinobacillus spp., Salmonella spp., Enterobacter spp., Citrobacter spp., and Pseudomonas aeruginosa.3–5,12–14 Gram-positive organisms have also been cultured from septic foals with a 30% incidence. The most common Gram-positive organisms cultured include Streptococcus spp., Staphylococcus spp., and Enterococcus spp.14 Most Gram-positive infections are complicated with a Gram-negative infection at the same time. Pure Gram-positive cultures are infrequent in neonatal sepsis.15
Clinical signs and diagnosis General signs of sepsis Foals with neonatal infection can present with a wide spectrum of clinical signs. It is not difficult to diag-
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nose sepsis in a foal that presents in septic shock. The problem arises in trying to diagnose the foal that is only showing early, subtle signs of infection. The variety of presentations is related to the host’s immune system, the duration of the illness, and the severity of the infection. Fever is not always present in infected foals. Some foals may be normal or hypothermic. Early in the septic process, the sick foal may only present with lethargy, decreased suckle, tachycardia, and tachypnea. The mucous membranes may be injected and the extremities are warm. As the infection progresses, the foal moves to a hypodynamic situation where it presents with severe depression, hypovolemia, prolonged capillary refill time, hypotension, and thready pulses. Petechiation of the mucous membranes, injected scleral vessels, cold limbs, and hyperemic coronary bands may also be present on physical examination. These foals are in septic shock. The ‘gold standard’ for diagnosis of sepsis in the foal is a positive blood culture. Though considered the most definitive of the diagnostic tools for sepsis, blood cultures have been reported to be positive in only 60–81% of confirmed septic foals. False negatives from necropsy-confirmed sepsis range from 12% to 37%.4, 16, 17 There is a lag time of several days from taking of the blood culture and the reporting of the results. Because of this and the concern about false negatives, blood cultures may have limited value when developing a therapeutic plan. The importance of blood culture is in the reporting of sensitivity patterns, which may be useful later if the foal is not responding to the initial antimicrobial treatment. Pertinent blood work can help direct diagnosis and treatment. A leukopenia with a neutropenia is the most common white cell aberration in the septic foal. A high band to segmented neutrophil ratio and toxic changes are highly suggestive of sepsis. Fibrinogen levels begin to rise 48–72 h after inflammation. High levels of fibrinogen at less than 24 hours of age may be helpful in determining whether the infection began in utero. Hypoglycemia is common in foals under 24 hours of age and may be profound. This is probably secondary to the foal’s failure to nurse and lack of glycogen reserves. Because FPT is the most common risk factor in septic foals, IgG levels should be determined. IgG levels of less than 800 mg/dL indicate partial or complete failure of passive transfer.16 A sepsis scoring system has been developed to address the limitations of blood cultures in the diagnosis of sepsis in the foal. Brewer and Koterba developed the Sepsis Score in the 1980s (Table 20.1). It is a system that awards points for some of the above historical, clinical, and laboratory abnormalities found in the suspect foal. The more points, the more likely the foal has sepsis. A foal with a score exceeding 11 is considered highly suspicious of being septic. With the exception of a complete blood count (CBC), much
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Table 20.1 Sepsis Score System. Each of the parameters is evaluated and a score is assigned.17 Barton M. Septicemia, In: Paradis MR (ed) Equine Neonatal Medicine: A Case Based Approach. Philadelphia: Elsevier, 2006; pp. 87. SCORE
PARAMETER
4
History: Placentitis, vulvar discharge, dystocia, long transportation of mare, mare sick, induced, prolonged gestation (> 365 days)
2
1
0
Yes
Premature (days of gestation) Clinical signs: Petechiae, scleral injection Fever Hypotonia, coma, depression, seizure Anterior uveitis, diarrhea, respiratory distress, swollen joints, open wounds
< 300
300–310
311–330
> 330
Severe
Moderate > 102◦ F Marked
Mild < 100◦ F Mild
None
2000–4000 or > 12 000 50–200 Mild > 600 < 50 400–800 40–50
8000–12 000
Normal
< 2000
Marked
< 200
> 200 Moderate
200–400 < 40
Score for this case
No
Yes
Laboratory data: Neutrophil count (per µL) Bands (per µL) Any toxic change in neutrophils Fibrinogen (mg/dL) Blood glucose (mg/dL) IgG (mg/dL) Arterial oxygen (mmHg)a Metabolic acidosisa
3
400–600 50–80 50–69 Yes
Normal None
None None < 400 > 80 > 800 > 70 No
Total points for this case A score of > 11 predicts sepsis. a If these two parameters are included, the positive cut-off value is 12.
of the data can be collected stall side, making this system more practical for early diagnosis.17, 18 Specific body system involvement Septicemia is the presence of bacteria or their toxins in the blood. As the microorganisms circulate through the body they may settle in a variety of body systems. In particular, one may associate pneumonia, septic arthritis or osteomyelitis, omphalophlebitis, septic meningitis, and anterior uveitis with the hematogenous spread of the bacteria. So in addition to the general signs of sepsis, you may identify clinical problems that are specific to one of these body systems. The respiratory system is particularly vulnerable to infection. In one study 50% of foals presented to a referral hospital had some form of pulmonary involvement. Early in the disease, clinical signs of pulmonary involvement may be subtle. Auscultation of the chest is not always helpful in determining the degree of lung involvement. The presence of abnormal sounds generally coincides with abnormal lungs but the absence of abnormal sounds does not rule out
lung involvement. Pneumonia is one of the biggest causes of mortality in the newborn foal.18, 19 Diagnostically, the best tests to determine the extent of pneumonia in the foal are arterial blood gases, chest radiographs, and thoracic ultrasound. Arterial blood gases gives some idea of how the lungs are functioning – determining oxygenation and ventilation. In the normal foal, oxygen partial pressures are low at birth (Pa o2 = 55–60 mmHg) but increase quickly to normal levels over the first 24 hours of life (Pa o2 = 70–90 mmHg). Because of a pliable chest wall, foals that are in lateral recumbency will have lower than normal arterial oxygen. Partial pressures of carbon dioxide are generally elevated at birth (Pa co2 = 50–55 mmHg) and decline to normal ranges within 24 hours (Pa co2 = 42–50 mmHg). Impaired diffusion of gases can occur with pneumonia, resulting in hypoxemia. Impaired ventilation or the shunting of blood through non-ventilated lungs will result in hypercapnea. Thoracic radiology is often helpful in determining the type and extent of pulmonary disease. Abnormalities can range from a mild bronchial pattern to
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Figure 20.2 Lateral radiograph of a septic physitis of the stifle. An area of lysis is noted on the cranial, distal femoral physis. Figure 20.1 Lateral radiograph of the chest of a septic foal. A diffuse alveolar and interstitial pattern is suggestive of a hematogenous spread of bacteria.
severe consolidation. Repeat radiographs over time will help to determine if the treatment is effective (Figure 20.1). Ultrasonography of the thorax can assess the equine neonatal lung and pleural space. Consolidation, pleural effusion, abscesses and rib fractures can be seen.20 Septic arthritis/osteomyelitis is also a manifestation of bacteremia in the foal. Approximately 25% of foals that presented to one referral hospital with sepsis scores greater than 11 developed evidence of septic arthritis.21 Septic arthritis was recognized as a cause of death in 12.5% of foals on farms in Texas.22 In the young foal the blood to a joint is supplied through a main arteriole that branches into the synovial membrane and the epiphysis. In the newborn foal there is a branch of the epiphyseal blood supply that crosses the physis.23, 24 Hematogenous bacteria can settle into these three areas creating septic foci. The clinical hallmarks of septic arthritis/ osteomyelitis are joint effusion and lameness. The lameness seen in these foals may vary from subtle to severe depending on the location of the infection. Lesions on weight-bearing structures elicit the most pain. In early septic physitis, joint effusion may be absent but swelling and pain elicited on palpation over the physis may be noted. Single or multiple joints can be affected in a foal. If multiple joints are involved, the foal may not exhibit a specific lameness because it is painful on multiple limbs. These foals will tend to be recumbent for long periods of time and the development of pressure sores may be the first indication of orthopedic pain. The joint prevalence varies but the tarsus, stifle, fetlock, and carpus were the most common joints noted in one study.25 The age range of affected animals is from birth to 21 days. Clinically the younger neonatal foal (birth to 36 hours) presents with joint
effusion as a part of the general signs of septicemia. Alternatively the slightly older foal (> 36 hours) may appear relatively normal at first and develops a lameness over a period of days or weeks.26 The presence of a septic joint is usually confirmed by sampling the synovial fluid. Normal joints usually have a cell count less than 200 cells/mm3 with approximately 25% neutrophils and a protein level of less than 2 g/dL. Infected joints will have an increased number of white blood cells with a predominance of neutrophils and an increased protein concentration. Radiographic imaging can be helpful in the presence of bone involvement but there can be a lag period in which bone demineralization goes unrecognized. Computed tomography has been useful in detecting bone involvement in some foals before radiographic changes have occurred27 (Figures 20.2 and 20.3). One of the points of possible entry of bacteria into the foal’s bloodstream is the umbilicus. Fifty percent
Figure 20.3 Lytic lesion in the physis of the front fetlock seen on computed tomography.
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of foals with evidence of an ongoing infectious process have demonstrated ultrasonographic evidence of omphalitis or omphalophlebitis.28, 29 One study found that foals with septic arthritis/osteomyelitis were more likely to have umbilical infections than were other foals with evidence of infection.30 Umbilical infections may or may not have external clinical signs. Ultrasonography is the most favored method of diagnosing a problem of the umbilicus. Bacterial meningitis is an infrequent sequela of perinatal infections. Koterba found that approximately 10% of septic foals presented with septic meningitis. Clinically the foals are depressed to the point of somnolence. Neck stiffness may be indicative of neck pain and meningitis. Seizures may be a presenting sign. The central nervous system of the neonate has an incomplete blood–brain barrier. Open tubulocisternal endoplasmic reticulum, components of the cerebral endothelial and choroid plexus epithelial cells, are thought to be sites of bacterial seeding in the neonate.31 Septic meningitis must be differentiated from other causes of central nervous system dysfunction in foals, such as trauma, hydrocephalus, and neonatal encephalopathy. The most useful diagnostic test for septic meningitis is cerebral spinal fluid (CSF) analysis and culture. Normal CSF should have very few cells present, typically 1 to 5 cells/µL. These cells should consist of monocytes and/or lymphocytes. Any neutrophils are considered abnormal. CSF protein levels have been reported to be elevated (138 ± 50 mg/dL) in normal pony foals less than 40 hours of age compared with adults.32 Systemic illness in neonatal foals frequently results in an anterior uveitis. This uveitis can be secondary to ocular invasion by the microorganism or result from an immune-mediated or endotoxic event. Clinical signs of uveitis include photophobia, lacrimation, miosis, aqueous flare, and blepharospasm. As the problem progresses, hypopyon, hyphema, fibrin in the anterior chamber, posterior synechia, and cataract formation may develop.33 Diagnosis is made with a thorough ophthalmic examination (Figure 20.4).
Treatment Treatment of the septic foal is directed to many fronts simultaneously. Supportive care of the foal in septic shock should be directed at correcting the problems of hypovolemia, hypotension, hypoglycemia, hypoxia, and hypothermia (see Chapter 14 for details). Immunological support for these foals includes plasma transfusion to address the FPT, polymixin B sulfate to bind endotoxin, and broad-spectrum
Figure 20.4 Fibrin in the anterior chamber of a septic foal.
antibiotic coverage to address the bacterial infection until culture results are available. Commercially available equine plasma with increased levels of IgG can be obtained from several Food and Drug Administration (FDA) approved sources.34 It comes frozen and has a shelf life of several years. It is administered intravenously through an in-line filter to remove fibrin clumps. The plasma should be administered slowly (0.5 mL/kg over 10–20 min) initially, and the foals should be carefully observed for any signs of a transfusion reaction, such as increased respiratory and heart rates, muscle fasciculation, and fever. If there are no reactions then the rate can be increased to 40 mL/kg/h. IgG levels should be evaluated periodically in the septic foal even after they have reached the desired level of 800 mg/dL as in the septic foal there is evidence that IgG is consumed by the process.35 Polymixin B sulfate has demonstrated endotoxinneutralizing properties in foals by ameliorating endotoxin-induced fever and cytokine production.36 The recommended dose is 1000–5000 IU/kg i.v. every 8–12 h. Antibiotic choice should reflect the likelihood of dealing with a Gram-negative organism and possibly a mixed Gram-negative and -positive infection. The combination of an aminoglycoside with a penicillin derivative has been advocated. Due to reports of developing resistance to gentamicin, the aminoglycoside of choice is now amikacin. The combination of amikacin and penicillin or ampicillin is projected to be effective against 90% of foal isolates. Other drugs that have been used include imipenem, enrofloxacin, ceftriaxone, ceftazidime, chloramphenicol, ceftiofur and tetracycline in decreasing order of efficacy.5, 14, 37 Foals presented in septic shock require intensive fluid resuscitation. Hypoperfusion in the critical foal is generally secondary to hypovolemia. Many of these foals are hypoglycemic as well as hypovolemic,
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Complications of Neonatal Infection especially if they are less than 24 hours of age. An iso-osmotic crystalloid fluid with added dextrose is the fluid of choice, given as boluses. The volume and rate of fluid resuscitation depend on the degree of hypovolemia. If the foal is recumbent and you are unable to feel a pulse in the distal metatarsal artery, or the blood pressure measurement is low, then the foal should receive an intravenous bolus of 20 mL/kg over a 20-min timespan. The foal should be evaluated after the bolus for pulse pressure, warming of the limbs, and urine production. If these parameters are still abnormal the bolus can be repeated several times with a re-evaluation of the clinical signs after each. Some foals may not respond to fluids alone. Inopressor therapy, such as dobutamine (5 µg/kg/min), dopamine (5 µg/kg/min), norepinephrine (0.3–0.5 µg/kg/min), or vasopressin (0.25–1.0 mU/kg/min), can be started.38 Finally, specific therapy for some of the manifestations of septicemia in the foal should be considered. Foals that present with pneumonia are often hypoxic and sometime hypercarbic. Hypoxemia is often responsive to intranasal oxygen administration. This can be delivered through a small indwelling tube inserted through the nasal passage into the pharynx. The tube is then secured to the nose with suture or tape. Hypercarbia is the result of blood being shunted through unventilated parts of the lung. A rising Pa co2 is often a sign of respiratory failure. Support in the form of mechanical ventilation may be necessary. If septic arthritis is a complication of sepsis, then the affected joints or physis require special attention. Systemic antibiotics alone are not effective in eliminating deep-seated synovitis or osteomyelitis. Aggressive local therapy with joint lavage will decrease the tension within the joint capsule and dilute the inflammatory products in the joint fluid. Intra-articular antibiotics or regional limb perfusion will increase the antibiotic concentration to much higher levels than can be achieved systemically.39 Treatment for infected umbilical structures depends on the degree of involvement and abscessation. Many foals will respond, with ultrasonographic improvement, after 3–7 days of systemic antibiotics. Foals with large abscesses in the umbilical stalk or vein may require surgical resection of the infected structures. If anterior uveitis is present in the septic foal and it is uncomplicated by corneal ulceration, then topical corticosteroids are indicated. The preferred steroids are 0.2% dexamethasone or 1% prednisolone acetate.33 Topical mydriatics, antibiotics, and nonsteroidal anti-inflammatory drugs are typically part of the regime. In cases where a large fibrin clot is visi-
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ble in the anterior chamber, an intracameral injection of tissue plasminogenase (TPA) administered under anesthesia can help to clear the anterior chamber. Care should be taken because of the high risk of complications.33
Prognosis Because of the many different presentations of sepsis in the foal, the reported overall short-term survival rate is variable, ranging from 10% to 70%. It appears that the survival rate has been improving over the past 30 years.40–42 More studies are needed to determine the long-term prognosis of septic foals.19
References 1. Stoneham S, Wingfield Digby N, Ricketts S. Failure of passive transfer of colostral immunity in the foal: incidence, and the effect of stud management and plasma transfusions. Vet Rec 1991;128:416–19. 2. Baldwin J, Cooper W, Vanderwall D, Erb HN. Prevalence (treatment days) and severity of illness in hypogammaglobulinemic and normoglobulinemic foals. J Am Vet Med Assoc 1991;198:423–428. 3. Brewer B, Koterba A. Bacterial isolates and susceptibility patterns in foals in a neonatal intensive care unit. Comp Cont Educ Pract Vet 1990;12:1773–81. 4. Wilson WD, Madigan JE. Comparison of bacteriologic culture of blood and necropsy specimens for determining the cause of foal septicemia: 47 cases (1978–1987). J Am Vet Med Assoc 1989;195:1759–63. 5. Henson S, Barton M. 2001. Bacterial isolates and antibiotic sensitivity patterns from septicemic neonatal foals: one 15 year retrospective study (1986–2000). In Dorothy Havemeyer Foundation Third Neonatal Septicemia Workshop Talloires, France. pp. 50–52. http://www. havemeyerfoundation.org/Neonatal%20Septicemia.htm 6. Haas S, Bristol F, Card C. Risk factors associated with the incidence of foal mortality in an extensively managed mare herd. Can Vet J 1996;37:91–5. 7. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek, TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203:1170–5. 8. Tizard I. Veterinary Immunology, 7th edn. Philadelphia: Elsevier Saunders, 2004. 9. Madigan J. Method for preventing neonatal septicemia, the leading cause of death in the neonatal foal. In: Proceedings of the 43rd Annual American Association of Equine Practitioners Convention, 1997; pp. 17–19. 10. Barton M. Septicemia. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case Based Approach. Philadelphia: Elsevier, 2006; pp. 75–95. 11. Zink MC, Johnson JA. Cellular constituents of clinically normal foal bronchoalveolar lavage fluid during postnatal maturation. Am J Vet Res 1984;45:893–7. 12. Corley KTT, Pearce G, Magdesian KG, Wilson WD. Bacteremia in neonatal foals: clinicopathological differences between Gram-positive and Gram-negative
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infections and single organism and mixed infections. Equine Vet J 2007;39:84–9. Sanchez LC, Giguere S, Lester GD. Factors associated with survival of neonatal foals with bacteremia and racing performance of surviving Thoroughbreds:423 cases (1982–2007). J Am Vet Med Assoc 2008;233:1446–52. Marsh P, Palmer J. Bacterial isolates from blood and their susceptibility patterns in critically ill foals: 543 cases (1991–1998). J Am Vet Med Assoc 2001;218:1608–10. Paradis MR. A retrospective review of equine neonatal septicemia. In: Proceedings of the American College of Veterinary Internal Medicine Forum, Washington DC, 1990; pp. 458–9. Koterba AM, Brewer BD, Tarpee FA. Clinical and clinicopathological characteristics of the septicaemic neonatal foal: review of 38 cases. Equine Vet J 1984;16:376–83. Brewer B, Koterba A. Development of a scoring system for the early diagnosis of equine neonatal sepsis. Equine Vet J 1988;20:18–22. Brewer B, Koterba A, Rowe E. Comparison of empirically developed sepsis score with a computer generated and weighted scoring system for the identification of sepsis in the equine neonate. Equine Vet J 1988;20:23–4. Uri L, Paradis MR. Evaluation of the effectiveness of equine neonatal care. Vet Med 1992;87:921–6. Bedenice D. Foal with septic pneumonia. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case Based Approach. Philadelphia: Elsevier. 2006; pp. 99–111. Paradis MR. Septic arthritis in the equine neonate: a retrospective study. In: Proceedings of the American College of Veterinary Internal Medicine Forum, 1992; pp. 458–459. Cohen N. Causes of and farm management factors associated with disease and death in foals. J Am Vet Med Assoc 1994;204:1644–51. Bennett D. Pathological feature of multiple bone infections in the foal. Vet Rec 1978;103:482–5. Firth EC. Current concepts of infectious polyarthritis in foals. Equine Vet J 1983;15:5–9. McCoy J, Paradis MR. 1998. An assessment of neonatal septic arthritis at Tufts University School of Veterinary Medicine, 1990–1998. Dorothy Havemeyer Neonatal Septicemia Workshop 2, Boston, MA. pp. 29. http://www. havemeyerfoundation.org/Neonatal%20Septicemia.htm Paradis MR. Foal with septic arthritis. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case Based Approach. Philadelphia: Elsevier, 2006; pp. 112–20. Paradis MR, Solano M, Tidwell A, Maranda L. The use of computed tomography in the diagnosis of septic arthritis/osteomyelitis in the neonatal foal. J Vet Intern Med 2009;24(3):719.
28. Reimer JM. Ultrasonography of umbilical remnant infections in foals. In: Proceedings of the 39th Annual Convention of the AAEP, 1993; p. 247. 29. White SL, Huff T. 1998. Retrospective study of surgical vs. medical management of umbilical remnant infections in neonates. Dorothy R. Havemeyer Neonatal Septicemia Workshop 2, Boston, MA, pp. 48. http://www. havemeyerfoundation.org/Neonatal%20Septicemia.htm 30. Paradis MR. Update on neonatal septicemia. In: Vaala W (ed.) Vet Clin N Am-Equine 1994;10(1):109–35. 31. Foreman JH. Considerations in the diagnosis and management of infectious neonatal neurological disease. In: Proceedings of the 8th American College of Veterinary Internal Medicine Forum, Washington DC, 1990; pp. 607–10. 32. Rossdale PD, Cash RS, Leadon DP, Jeffcott LB. Biochemical constituents of cerebrospinal fluid in premature and full term foals. Equine Vet J 1982;14:134–8. 33. Jurk I. Septic foal with hypopyon. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case Based Approach. Philadelphia: Elsevier, 2006; pp. 268–70. 34. American Association of Equine Practitioners. White paper: Information on equine plasma and serum products for the equine practitioner. Developed by the Biological and Therapeutic Agents Committee of the AAEP. 2009; URL: http://www.aaep.org/press room.php?term=2009&id= 340. 35. Sellon DC. Foal with failure of passive transfer. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case Based Approach. Philadelphia: Elsevier, 2006; pp. 31–9. 36. Durando MM, MacKay RJ, Linda S, Skelley LA. Effects of polymixin B and salmonella typhimurium antiserum on horses given endotoxin intrevenously. Am J Vet Res 1994;55:921–7. 37. Madigan J. Manual of Equine Neonatal Medicine, 3rd edn. Woodland, CA: Liveoak Publishing, 1997. 38. Palmer J. Foal in hemodynamic shock. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case Based Approach. Philadelphia: Elsevier, 2006; pp. 121–34. 39. Werner LA, Hardy J, Bertone AL. Bone gentamicin concentration after intra-articular injection or regional intravenous perfusion in the horse. Vet Surg 2003;32:559. 40. Hoffman A, Staempfli H, Willan A. Prognostic variables for survival of neonatal foal under intensive care. J Vet Intern Med 1992;6:89–95. 41. Raisis A, Hodgson J, Hodgson D. Equine neonatal septicemia: 24 cases. Aust Vet J 1996;73:137–40. 42. White SL, Barton M. 2001. The sepsis score revisited. Dorothy R. Havemeyer neonatal Septicemia Workshop 3, Talloires, France, pp. 13–15. http://www.havemeyer foundation.org/Neonatal%20Septicemia.htm
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Chapter 21
Fluid Therapy Robert P. Franklin
Intravenous fluid therapy is a crucial part of neonatal care both in the field and at the hospital. Fluids are required in cases requiring volume replacement, patients needing ongoing maintenance fluid therapy, and in foals that require specialized electrolyte replacement. In many situations the need for a response of fluid therapy is governed by clinical parameters alone. Where practical, biochemical values, electrolytes, and packed cell volume (PCV) and total protein (TP) levels should be used to dictate the needs and types of fluids administered and to monitor the response to such efforts.
Normal fluid requirements When organizing a fluid plan three variables should be considered. The first is the foal’s maintenance fluid requirements. Neonates require 10% of their body weight per day in fluids (∼4 mL/kg/h). A normal foal will consume 20–25% of its body weight per day in milk by the first week of life; therefore, a normal foal will maintain its hydration very well on a milk diet and will consequently have dilute urine (specific gravity < 1.008) owing to the relative water excess.1, 2 Juvenile patients have a smaller fluid requirement in the order of 7.5% of body weight per day (∼3 mL/kg/h). The distinction between neonate and juvenile is often made at 30 days old. The second consideration in the fluid plan is fluid deficits. Foals will tolerate mild levels of dehydration up to 5%. Thus, a foal with notable signs of volume depletion should be considered to be at least 6% dehydrated. A foal with mild to moderate dehydration (6%) will have signs of depression, tachycardia, dry mucous membranes, and concentrated urine. More severe dehydration (8%) will be noted by prerenal azotemia, hyperlactatemia, decreased capillary refill
time, and stuporous behavior. Shock is noted in the most severe levels of dehydration (≥ 10%), and these patients often present in lateral recumbency with severe derangements of many of the previously noted features. It is important to assign a level, not a range, of dehydration to foals in order to make basic calculations for fluid needs. It is also notable that these calculations are meant to provide a starting point for resuscitative efforts. Therefore, differences in assessment by clinicians, whereby one assigns a level of 6% to a foal whereas another assigns 8%, do not result in any significant problems when prescribing fluid therapy. Both clinicians will be providing a close level of fluid resuscitation; for example, in a 50-kg foal clinician A will estimate a volume deficit of 3 liters whereas clinician B determines the deficit to be 4 liters. Fluid deficits, once calculated, are administered to severely dehydrated patients by giving bolus infusions of 10–20 mL/kg until symptoms of dehydration improve, or by providing the deficit as a constant infusion over 6 hours to less severely affected patients. Any residual fluid volume after boluses is then carried over to the daily fluid plan. The last determination, and often the most difficult to quantify, is ongoing fluid losses. Patients with gastrointestinal reflux can be charted to determine their hourly and daily losses. However, foals with diarrhea are more difficult to assess. When planning for these cases the clinician must add an estimate to the equation and then use clinical monitoring to determine whether losses are being remedied by the current fluid rates. Serial body weight measurements are of value when monitoring for ongoing fluid losses, but often this process is impractical outside of an intensive care unit. Table 21.1 provides an example of a 24-hour fluid plan for a 50-kg Thoroughbred foal with enteritis and moderate dehydration (6%).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 21.1 Fluid schedule for a 50-kg foal with enteritis. Maintenance fluid requirements, deficit (degree of initial dehydration), and ongoing losses are figured and combined to determine the total number of liters needed in a 24-hour period. A constant rate infusion hourly rate can then be calculated Maintenance
10% of 50 kg = 5 liters/day
Deficit (residual)
6% of 50 kg = 3 liters – 1.5 liters given by bolus on presentation = 1.5 liters
Ongoing losses
Estimated at 2 liters/day
Total daily fluid requirements
8.5 liters
Hourly fluid rate
8.5 liters/24 hours = 354 mL/h
the equine adult, but because of high sodium loads and the relative inefficiency of the neonatal kidney in dealing with such quantities, this type of solution should not be routinely used as maintenance fluids for foals.2 That being said, foals with diarrhea or cerebral disease (head trauma, meningitis, neonatal encephalopathy) may be more appropriately treated with replacement fluids to replace electrolyte losses or prevent cerebral edema, respectively. Examples of replacement fluids include: lactated Ringer’s solution, 0.9% sodium chloride, Normosol-R, PlasmaLyte 148, and Hartmann’s solution. Bicarbonate infusions
Fluid types Maintenance fluids Maintenance fluids are those fluid types that are meant to provide physiological levels of electrolytes and water in order to hydrate the intracellular and extracellular fluid spaces. These solutions are not to be given via bolus infusion or to provide resuscitation to acutely hypovolemic foals. Instead, maintenance fluids are used in euvolemic patients that require ongoing fluid support to provide daily requirements of water and electrolytes when they cannot eat or drink.3 The electrolyte levels of maintenance fluids are most notably different from replacement fluids in that they contain approximately one-half to one-third of the sodium load. The relative deficiency in sodium provides more free water to hydrate the intracellular spaces as needed. Clinicians should exercise caution when caring for foals with cerebral disease as an increase in free water may lead to cerebral edema. Maintenance fluids also often have increased levels of potassium and are commonly adjusted to be near isotonic by the addition of dextrose. Examples of maintenance fluids include: Plasma-Lyte 56 with 5% dextrose, Normosol-M with 5% dextrose, and 0.45% saline with 5% dextrose (Table 21.2). Replacement fluids Replacement fluids are solutions designed to match physiological constituents of the extracellular fluid compartment and, by doing so, are confined to those spaces. This property makes these fluids especially useful in bolus administration aimed at restoring acute volume loss and directly improving blood pressure and organ perfusion. It is should be noted that the effects of replacement crystalloid fluids are short lived and the intravascular distribution is limited to 25% (75% to the interstitial space).3 Replacement fluids are commonly used as maintenance fluids in
Sodium bicarbonate fluids are selectively used to correct normovolemic metabolic acidosis or to drive potassium to the intracellular space in uremic patients with hyperkalemia. Foals with diarrhea or uroperitoneum most commonly require bicarbonate therapy. Blood gas analysis or measurement of total carbon dioxide (Tco2 ; 95% of the Tco2 value represents bicarbonate and therefore is a functional estimate for calculations) on a biochemical profile are used to calculate bicarbonate deficits. Hypovolemic patients should be volume replaced prior to calculating and correcting base deficits as restored perfusion corrects or improves many of these cases. The formula to determine milliequivalents (mEq) of bicarbonate needed is: mEq HCO3 = base deficit (25 − HCO3 or TCO2 ) × 0.6 × BW where BW is bodyweight in kilograms. Volume replenished foals with a pH < 7.25 and with base deficits > 10 mEq are suitable candidates for bicarbonate administration. Traditional administration of bicarbonate dictates that one-half of the calculated dosage should be given over several hours followed by a reassessment of pH and base deficit. Experience has suggested that foals with severe diarrhea and metabolic acidosis may be given their entire deficit in order to correct the derangement. Table 21.3 provides an example of bicarbonate administration. Extreme caution should be used in patients with hypokalemia as bicarbonate administration will worsen this syndrome and potentially lead to fatal arrhythmias. Thus, hypokalemia should be corrected before bicarbonate administration and concomitant administration of potassium should occur during treatment of patients predisposed to hypokalemia. Caution should also be used, as bicarbonate infusions will generate excessive amounts of carbon dioxide. Foals with central nervous disorders, such as neonatal encephalopathy, will not respond appropriately to this
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LRS, lactated Ringer’s solution.
– –
– –
–
Acetate
–
– 1232
Gluconate
27
Bicarbonate
–
–
–
–
103
Chloride
–
–
1232
–
3
Magnesium
Lactate
5
142
5
Sodium
Potassium
Calcium
–
Plasma
–
–
–
–
154
–
–
–
154
0.9% saline
–
–
29
–
109
–
3
4
130
LRS
23
27
–
–
98
3
–
5
140
–
–
29
–
111
–
2
5
131
Hartmann’s solution
–
16
–
–
40
3
–
13
40
–
–
–
–
77
–
–
–
77
0.45% saline with 5% dextrose
–
–
–
1000
–
–
–
–
1000
8.4% sodium bicarbonate
–
–
–
595
–
–
–
–
595
5% sodium bicarbonate
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5% dextrose
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Table 21.2 Electrolyte constituents of available intravenous fluids
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Table 21.3 A 66-kg foal with rotaviral diarrhea leading to metabolic acidosis. Potassium level 3.8 mEq/L, normal respiratory function. Blood gas taken after volume restoration with lactated Ringer’s solution pH Bicarbonate level Base deficit Body weight Bicarbonate dose Plan A
7.18 10 mEq (25–10) = 15 66 kg 66 × 15 × 0.6 = 594 mEq Administer 500 mL of 5% sodium bicarbonate (297.5 mEq) over 2 hours and recheck blood gas
Plan B
Administer 1000 mL 5% sodium bicarbonate (595 mEq) over 1 hour and recheck blood gas in 12 hours
rise in carbon dioxide and a respiratory acidosis will be generated. Bicarbonate infusions will also contain high sodium loads. This may lead to osmolarity problems and hypernatremia. Bicarbonate should not be given with calcium- or lactate-containing fluids owing to the risk of precipitate development. Bicarbonate infusions are available as hypertonic sodium solutions in either 5% or 8.4%. A 5% sodium bicarbonate solution contains 595 mEq bicarbonate per liter whereas 8.4% sodium bicarbonate contains 1000 mEq bicarbonate per liter. Each gram of sodium bicarbonate contains 12 mEq sodium and 12 mEq bicarbonate. Isotonicity may be achieved by adding 4.6 mL sterile water to each milliliter of 8.4% sodium bicarbonate.4 Persistently acidemic patients may also require the administration of oral bicarbonate in the form of baking soda. One tablespoon (15 mL) of baking soda contains 62.5 mEq bicarbonate. Experience suggests that two tablespoons (30 mL) can be given every 12–24 hours. Bicarbonate status should be monitored during this course to adjust dosing. Colloids Intravenous fluids are defined as colloids or crystalloids based on their ability to permeate through the vascular endothelium. Colloid fluids are composed of larger molecules, which prevent diffusion through the endothelium. This property allows much of the administered fluid to remain within the intravascular space and therefore colloids have a direct effect on restoring circulating volume.3 This preference for the intravascular space also poses a potential problem for volume overload when large doses are used in foals. In contrast, crystalloid fluids readily distribute evenly throughout the extracellular and intracellular fluid spaces based on their electrolyte content. Colloid fluids are indicated for treatment of severe hypovolemic shock as part of a low-volume resuscitation effort, failure of passive transfer, sepsis, anemia,
and hypoproteinemia. Available colloids for the neonate include the synthetic amylopectin hetastarch (hetastarch sodium chloride injection 6% hetastarch in 0.9% sodium chloride; Baxter), a bovine-derived hemoglobin-based oxygen carrier OxyglobinTM [Oxyglobin hemoglobin glutamer 200 (bovine); Biopure Corp, Bellevue, WA], human albumin ([Flexbumin 25% (albumin (human)), USP, 25% solution; Baxter], and equine origin plasma. A recent White Paper has been established by the American Association of Equine Practitioners to assist practitioners in the selection of equine origin plasma and serum products.5 Low-volume resuscitation involves the combined or singular use of hypertonic saline at 2–4 mL/kg and/or a synthetic colloid such as hetastarch at 2–10 mL/kg. There are differing opinions on the use of colloids versus crystalloids for rapid resuscitation.6 The author believes that low-volume resuscitation efforts do afford a positive clinical response in older foals with shock, but cautions against the use of such efforts to resuscitate neonatal foals owing to their immature blood–brain barriers and the potential to induce sudden changes in osmolarity. Normal oncotic pressure in foals is > 15 mmHg. Colloids are used to treat the effects of hypoproteinemia when colloid oncotic pressure (COP) falls below 15 mmHg, when measured by colloid osmometry R (Colloid Osmometer, model 4420, Wescor). Indirect COP can be calculated by the Landis–Pappenheimer equation: COP = 2.1(serum total protein) + 0.16(serum total protein)2 + 0.009(serum total protein)3 However, the accuracy of indirect measurement decreases when applied to critically ill neonates.7 The use of colloids may also be indicated when total protein levels are less than 4.5 g/dL when direct COP measurement is not available. Protein levels in newborn foals are often below this threshold and failure of passive transfer should be considered and managed before using hetastarch or other products aimed at solely managing oncotic pressure. Colloid fluids act to retain fluid within the intravascular space while counteracting the forces of hydrostatic pressure. COP is often compromised in patients with protein-losing enteropathies such as foals with diarrhea or those infected with Lawsonia intracellularis. Crystalloid fluids administered to foals with low COP will promote the development of edema in the interstitial tissues. Preloading with a colloid fluid may prove especially useful in preventing edema when large doses of crystalloids are given to maintain hydration.
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Table 21.4 Examples for dosing hetastarch, equine origin plasma, or human albumin for a 200-kg weanling with proliferative enteropathy due to infection by Lawsonia intracellularis. Targeted albumin concentration is 3.0 g/dL Approximate cost in US$ Body weight Albumin Total protein Oncotic pressure Hetastarch Equine plasma Flexbumin 25%
200 kg 1.0 g/dL 2.7 g/dL 7.01 mmHg 10ml × 200kg = 2.0 L 10ml × 200kg = 2.0 L 3.0(Alb des) – 1.0(Alb cur) × 200kg = 400 g albumin
Hetastarch, a low-cost synthetic colloid consisting of the starch amylopectin with molecules of an average of 130 000 kDa, is administered as a bolus dose of 10 mL/kg to improve COP with doses repeated daily until the patient is stabilized. Hetastarch gives a reading of 2.6 g/dL on refractometry and will slightly raise total protein levels in patients with protein levels < 2.6 g/dL and slightly lower total protein levels in patients with protein levels > 2.6 g/dL, although this has no dilutional effect on the true COP. Therefore, total protein levels become less reliable indicators of COP after hetastarch administration. Equine origin plasma is a readily available option for managing low COP. Plasma may be obtained from donor horses or commercial sources. Plasma is administered at a bolus dose of 10 mL/kg and repeated daily until the patient is stabilized. Human albumin is particularly effective in restoring COP and improving albumin levels. In adult horses, the effects of human albumin were superior to those of hetastarch.8 The recommended dose is: Alb = (Albdesired − Albcurrent ) × BW where Alb is albumin in grams and BW is bodyweight in kilograms. Table 21.4 provides dosing examples for Hetastarch, equine plasma, and albumin. Severe anemia requires blood transfusion techniques to be used (see Chapter 25). However, as a lifesaving bridge technique, Oxyglobin can be administered to dysoxic foals. Oxyglobin is given at a dose of 30 mL/kg. The effects of this treatment are increased oxygen delivery capacity along with an increase in COP.9 Equine origin plasma is ideally suited for treatment of septic patients and failure of passive transfer. Plasma should be given at a dose of 10–40 mL/kg. Foals that have complete failure of passive transfer often require 40–60 mL/kg of plasma. A dose of 60 mL/kg (or 3 liters for a 50-kg foal) requires supplementation over the course of 2 days to avoid hypertension, so that not more than 40 mL/kg are given in a 24-hour period. Plasma is also a useful treatment in cases of sepsis regardless of passive transfer
80 300 1280
status. Lower doses of 10–20 mL/kg provide clotting factors, non-specific antibodies, and albumin to these patients.
Electrolytes, glucose, and bicarbonate Electrolyte abnormalities are common in the sick neonate and in the older foal with diarrhea. Disorders of sodium, chloride, potassium, calcium, and glucose present most commonly. Neonates also require a cognizant approach of supplementation of electrolytes when being managed as nil per os. Sodium
Hyponatremia Hyponatremia, sodium < 130 mEq/L with serum plasma osmolality < 285 mmol/L, is always due to a relative excess of water. Hyponatremia should be categorized by duration of illness, patient hydration status, and the presence of neurological disease. Hypovolemic hyponatremia accompanies gastrointestinal disease or renal fluid loss with antidiuretic hormone acting to retain water. Euvolemic hyponatremia is noted with adrenal insufficiency, syndrome of inappropriate antidiuretic hormone secretion, hypothyroidism, and excess water consumption. Hypervolemic hyponatremia is seen with renal failure, liver failure, and nephrotic syndrome. The author most commonly notes hyponatremia in foals with renal failure, often due to high-dose oxytetracycline administration for contracted tendons, and additionally in foals with diarrhea, uroperitoneum, or foals that have been administered large levels of low-sodium fluids, such as elevated rates of administration of maintenance fluids or 5% dextrose in water. Clinical signs of acute hyponatremia are related to neurological disorders secondary to brain edema due to osmotic shifts of water towards the hyperosmolar neurons. The brain compensates during chronic hyponatremia by eliminating intracellular solutes to prevent edema. This compensation event requires
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Table 21.5 Two scenarios of hyponatremia in 1-week-old foals and recommended treatment plans. Patient 1 has acute, symptomatic, hypovolemic hyponatremia. Patient 2 is asymptomatic and euvolemic with chronic hyponatremia Hydration status
Neurological status
Duration of illness
Sodium level (mEq/L)
Patient 1: diarrhea, 60 kg
8% dehydrated
Stupor, head pressing
2 days
113
Goal is to restore volume and correct sodium as quickly and safely as possible (1 mEq/L/h). Address dehydration with 2-liter 0.9% saline bolus, reevaluate electrolyte concentrations, continue constant rate infusions or intermittent bolusing of replacement fluid with goal of raising sodium to 125 mEq/L in 12 hours)
Patient 2: premature with suspected adrenal insufficiency, 25 kg
Normal
Normal
4 days
118
Goal is to reduce water intake and correct sodium levels slowly (12 mEq/day). Plasma-Lyte 56 maintenance fluids are changed to Plasma-Lyte 148 replacement fluids and the foal’s daily fluid rate is confirmed not to exceed 10% of bodyweight, accounting for all sources of water intake (intravenous medicines, milk, etc.). Sodium levels monitored twice daily
Patient
24–48 hours after the onset of hyponatremia. Patients may be noted to be depressed, disoriented, blind, or seizuring. Pseudohyponatremia, which typically does not cause clinical signs or require sodiumdirected therapy, is found when plasma osmolality levels are normal (285–295 mmol/L) and high levels of lipids or proteins are present, or when plasma osmolality levels are high (> 295 mmol/L) owing to hyperglycemia or mannitol in the blood. The last two situations lead to a dilution of sodium in laboratory samples, but do not imply a deficiency in sodium. Care should be taken with both symptomatic and asymptomatic patients to not increase sodium levels too fast. Electrolyte monitoring should be performed at least daily and may be required as often as 12 times a day when correcting severe sodium aberrations. Potassium levels are always monitored when using loop diuretics such as furosemide. Acute cases (< 48 hours’ duration) of hyponatremia are corrected at a rate of up to 24 mEq/L/day (1 mEq/L/h) to attempt to establish a sodium level of 125 mEq/L as quickly as possible. Chronic cases are corrected at a slower rate of 12 mEq/L/day (0.5 mEq/L/h). Overzealous correction of sodium can lead to brain dehydration, specifically central pontine myelonolysis, which also manifests as severe neurological impairment.10 The goal of treatment of the symptomatic hypovolemic, hyponatremic patient is restoring plasma volume. Ideally, 0.9% saline is given as a bolus followed by maintenance of the daily fluid plan (residual deficit, ongoing losses, maintenance) with a replacement fluid. Treatment goals for the acute symptomatic
Treatment plan
euvolemic, hyponatremic patients are restoring plasma sodium levels quickly. These patients suffer from water excess and are treated by small doses of 3% hypertonic saline (1 mL/kg q. 6–12 hours) and furosemide [0.5 mg/kg intravenously (i.v.) q. 6–12 hours]. Asymptomatic, chronic, euvolemic patients have adjusted their cerebral osmolytes to the hyponatremia and must have their sodium levels returned to normal slowly. These cases are treated by decreasing total water intake, normally by reducing i.v. infusions and switching to a replacement fluid with a higher sodium concentration. Hypervolemic hyponatremia is typically chronic and asymptomatic. Treatment goals are aimed at managing the underlying disorder. Table 21.5 provides examples of managing foals with syndromes of hyponatremia.
Hypernatremia Hypernatremia, sodium > 158 mEq/L and osmolality > 300 mmol/L, is encountered with salt excess or volume loss. Hypernatremia should be categorized by duration of illness, patient hydration status, and the presence of neurological disease. Hypervolemic hypernatremia suggests sodium excess. Hypertonic saline, sodium bicarbonate infusions, replacement solutions given as maintenance fluids, and improper formulation of milk replacers are the most likely reasons for hypervolemic hypernatremia in the neonate. Hypovolemic hypernatremia is indicative of volume loss. Neonatal foals that have renal (mannitol, hyperglycemia, diuretics) or intestinal (osmotic diarrhea)
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Fluid Therapy losses of hypotonic fluids may have difficulty in replacing free water owing to the inability to drink in a debilitated state. Clinical signs of acute hypernatremia are neurological in origin and are related to cellular dehydration of the brain due to osmotic fluid shifts towards the hyperosmolar extracellular fluid compartment. The brain compensates during chronic hypernatremia by the addition of intracellular solutes that draw water back into the cells. This compensation event occurs 24–48 hours after the onset of hypernatremia. Similar to hyponatremia, acute hypernatremia requires prompt correction to avoid or rectify neurological problems, whereas chronic hypernatremia calls for a slower response so that neurological problems are not encountered in response to treatment (rapid water replacement will cause cerebral edema). Acute cases are corrected at a rate of up to 36–48 mEq/L/day (1.5–2 mEq/L/h). Chronic cases are corrected at a slower rate of 12 mEq/L/day (0.5 mEq/L/h). Electrolyte monitoring should be performed at least daily and may be required as often as 12 times a day when correcting severe sodium aberrations. Hypovolemic hypernatremia is treated in a similar way to any patient with volume depletion by correcting fluid deficits using replacement solutions such as 0.9% saline, lactated Ringer’s solution, etc. These solutions restore volume and actually have a lower osmolality than the patient’s blood (Table 21.2). After volume is restored, additional fluid therapy is given in the form of free water: oral tap water or low-sodium maintenance fluids such as 5% dextrose in water, Plasma-Lyte 56 with 5% dextrose, or 0.45% saline with 5% dextrose at an appropriate rate to correct the deficiency and neurological signs (1.5–2.0 mEq/L/h). Hypervolemic hypernatremia requires furosemide treatment (0.5 mg/kg i.v. q. 6–12 hours) to remove excess salt and fluids with 5% dextrose in water given intravenously to replace urinary losses at an appropriate rate to correct hypernatremia safely. Proper formulation of milk replacer and access to tap water are then used to maintain hydration and electrolyte status. Potassium Milk and foodstuffs are generally quite high in potassium. Patients that are nil per os will require potassium supplementation as most fluid preparations are deficient in potassium levels to ensure safety in bolus situations. Maintenance fluids are spiked with potassium chloride at 40 mEq/L and are administered at rates not to exceed 0.5 mEq/kg/h (50-kg foal at < 25 mEq/h or < 625 mL/h of the 40-mEq/L supplemented fluid). Bolus and high-rate replacement infusions are not spiked in order to avoid hyperkalemia.
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Hypokalemia Hypokalemia, potassium <3 mEq/L, that is encountered with diarrhea, insulin therapy, ileus, bicarbonate administration, or diuretic treatment is also treated with potassium chloride. Potassium levels should be corrected as quickly as safely possible, especially in cases < 2 mEq/L. Deficits are difficult to measure owing to the majority of potassium residing in the intracellular fluid space. In the case of severe hypokalemia, potassium < 2 mEq/L, replacement fluids are spiked with 0.5–1 mEq/kg and administered over 1 hour. This treatment is continued for 2–3 hours and potassium levels are then rechecked and additional treatments are given if hypokalemia persists. Intravenous infusions of dextrose or bicarbonate and insulin administration should be discontinued (or at least slowed if a medical condition prevents stopping altogether) as these treatments may worsen hypokalemia. Oral supplementation is warranted in addition to i.v. potassium in refractory cases. Moderate hypokalemia, potassium 2–3 mEq/L, is treated by providing additional potassium in constant rate infusions or by supplementing with Lite SaltTM given orally as a paste with water. One teaspoon, 5 mL, of Lite SaltTM contains 45 mEq potassium. A dose of 1 teaspoon/50 kg twice daily is given and potassium levels are monitored closely.
Hyperkalemia Renal failure and post-renal azotemia (e.g., uroperitoneum) are the most common causes of hyperkalemia. Clinical features of hyperkalemia, potassium > 6 mEq/L, include generalized muscle weakness and cardiac arrhythmias. Treatment of hyperkalemia is accomplished by discontinuing all potassium supplementation, stimulating intracellular shifts of potassium, and increasing renal excretion of potassium. Additionally, calcium is given to prevent cardiac arrhythmias. Treatment plans in the intensive care unit that may include high levels of potassium include penicillin therapy, i.v. infusions, and oral supplements. Insulin, glucose, and bicarbonate will stimulate intracellular shifts of potassium. A common therapy used for hyperkalemia involves administering 50 mL calcium borogluconate 23% in 1 liter of 0.9% saline followed by the addition of 100 mL 50% dextrose and 100 mL 5% bicarbonate to a further 1 liter of 0.9% saline (other replacement fluids often contain some potassium; see Table 21.2). This treatment is given over 30–60 minutes to a 50-kg foal followed by monitoring potassium levels for 30 minutes afterwards. Insulin may also be given at 0.1 U/kg i.v. at the beginning of treatment. Cation exchange resins R ; sodium polystyrene sulfonate) absorb (Kayexalate
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potassium when given orally or via enema. A stanR with 100 mL of 20% dard dose is 20 g Kayexalate sorbitol to prevent impaction. Albuterol can lower potassium levels quickly and may be used during anesthesia, although arrhythmia generation must be monitored closely with this drug.3 Once potassium levels have stabilized, the treatment plan is continued with a maintenance rate of either 0.45% saline with 5% dextrose or, if there are ongoing fluid losses, 0.9% saline with additional 100 mL 50% dextrose.
Total serum calcium exists in protein-bound (45%), free, or ionized calcium (iCa) (50%) and calcium complexed to anions (5%). iCa levels reflect available calcium to cells and are independent of protein levels. Alkalosis decreases iCa, while acidosis increases iCa. Calcium supplementation is required in foals that are nil per os. Replacement and maintenance fluids contain low levels of calcium. Calcium borogluconate 23% (1 mL contains 230 mg or 1.069 mEq calcium) is added to maintenance fluids at a rate of 2.5 mL/kg/day (2.67 mEq/kg/day) or 20 mL (21.38 mEq) per liter.1
biologically active form of glucose. Neonatal foals either that are unable to nurse or that are withheld from nursing require glucose supplementation to their fluid plan. A detailed description on intravenous nutrition can be found in Chapter 28. However, short treatment periods, such as when foals are being transported to hospital facilities, can be addressed with dextrose infusions alone. Dextrose is given as a 5–10% solution during constant rate infusions. Glucose should not be given by rapid bolus administration to foals as this may induce glucose swings that worsen hypoglycemia. Maintenance fluids often contain 5% dextrose and are adequate to provide short-term i.v. nutrition. Replacement fluids are spiked with 100–200 mL 50% dextrose per liter to make a 5–10% dextrose solution, respectively. This fluid preparation should also be given as an infusion if necessary (increased sodium needed to replace residual deficits or ongoing losses). A solution of 5% dextrose in water is a poor choice for glucose supplementation owing to the amount of free water that is delivered once the dextrose is metabolized. This fluid should be reserved for specialized drug dilutions or management of severe hypernatremia, both of which circumstances are uncommon.
Hypocalcemia
Hypoglycemia
Idiopathic hypocalcemia, calcium < 10 mg/dL or iCa < 5 mg/dL, is encountered occasionally in neonates. Common clinical complaints consist of tachycardia, tachypnea, diarrhea, spasticity or weakness, seizures, or hyperhidrosis.11 Calcium borogluconate 23% (1 mL/kg) and magnesium sulfate (10 mg/kg) should be given in 1 liter of replacement fluids over 30 minutes. Treatment may need to be repeated based on reevaluation of calcium levels. Ongoing maintenance levels are then given intravenously in addition R to oral therapy with calcium citrate tablets (Citracal 1 or 2 tablets/50 kg once or twice a day). Many of these foals require several weeks of oral treatment after initial stabilization with i.v. therapy. A poor prognosis is given for foals unable to maintain calcium levels > 8 mg/dL.
Hypoglycemia, glucose < 100 mg/dL (< 5.55 mmol/ L), is common in the neonate and causes signs of depression, lethargy, hypothermia, or seizures. Treatment goals are to correct hypoglycemia and to regulate glucose levels of 100–180 mg/dL thereafter. Bolus infusions of glucose, especially hypertonic doses of 50%, should be discouraged because of potential damage to the central nervous system and rebound hypoglycemia.1 Instead, an infusion of 10% dextrose is recommended. Most patients are also hypovolemic and require replacement fluid therapy. Lactated Ringer’s solution is supplemented with 200 mL of 50% dextrose to achieve a 10% solution and is given over 30–60 minutes. Regulation of glucose levels then requires the use of a dextrose infusion. At this point, a 5% dextrose solution, either maintenance fluid containing dextrose or replacement fluid supplemented with 100 mL 50% dextrose per liter, is given until the foal is either able to nurse successfully or until it has arrived at a hospital facility. A slow drip of such an infusion will preserve the glucose levels once the foal has had a rescue dose of dextrose.
Calcium
Hypercalcemia Hypercalcemia is uncommon in neonatal foals, although it is sometimes noted with prematurity and is a negative prognostic marker.12 Diuresis is the treatment of choice.
Hyperglycemia Glucose Glucose is a monosaccharide that has two sterioisomers (l- and d-glucose). d-Glucose (for dextrorotatory glucose) is known as dextrose and is the
Hyperglycemia, glucose > 180 mg/dL (> 9.9 mmol/ L), is most often the result of overzealous glucose infusions. Critically ill neonates are also predisposed to hyperglycemia and this represents the potential for
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Table 21.6 A general guide to fluid choices when treating foals with various medical conditions Condition
Resuscitative fluid choice
Maintenance fluid choice
Fluid additives
Septic shock
LRS or other replacement fluid; or if severe, hypertonic saline with hetastarch (caution in newborns or those with neonatal encephalopathy)
Plasma-Lyte 56 with 5% dextrose
100–200 mL 50% dextrose in resuscitative fluids if hypoglycemic, 40 mEq KCl/L + 20 mL calcium borogluconate 23%/L in maintenance fluids
Diarrhea/ileus with ongoing losses
LRS or other replacement fluid +/– sodium bicarbonate
LRS if sodium deficient or maintenance fluid such as Plasma-Lyte 56 with 5% dextrose if sodium levels stable
40 mEq KCl/L + 20 mL calcium borogluconate 23%/L in maintenance fluids
Uroabdomen/renal failure
0.9% saline
0.45% saline with 5% dextrose as CRI; or if using fluid boluses 0.9% saline
If hyperkalemic: 50 mL calcium borogluconate 23% in 1 liter 0.9% saline followed by 100 mL 50% dextrose, 100 mL 5% bicarbonate in 1 liter 0.9% saline
Neonatal encephalopathy
LRS or other replacement fluid
Plasma-Lyte 56 with 5% dextrose or LRS if concerned about cerebral edema
100–200 mL 50% dextrose in resuscitative fluids if hypoglycemic, 40 mEq KCl/L + 20 mL calcium borogluconate 23%/L in maintenance fluids
Premature
LRS or other replacement fluid
Plasma-Lyte 56 with 5% dextrose
100–200 mL 50% dextrose in resuscitative fluids if hypoglycemic, 40 mEq KCl/L + 20 mL calcium borogluconate 23%/L in maintenance fluids
LRS, lactated Ringer’s solution; CRI, constant rate infusion.
a worsened outcome in children. Information gained through goal-directed therapy has shown the benefits of controlling glucose levels in adult surgical intensive care patients, although these data have not been completely supported in more recent studies of adults and children.13, 14 The first step in regulating glucose is to decrease, but not discontinue, the dose of any dextrose infusions. Insulin therapy is used to control hyperglycemia when minimal, but necessary, dextrose infusions are not tolerated. It is important to note that insulin therapy may result in a hypoglycemic crisis or in cardiac arrhythmias generated by hypokalemia secondary to its effects. Careful monitoring of serum potassium levels during insulin R therapy is critical. A low dose of insulin [Vetsulin (porcine insulin zinc suspension), 40 IU/mL lente preparation, Intervet; 0.1 U/kg subcutaneously (s.c.)] is given first and the response to therapy is measured by glucometry after 1 hour. Serial doses may be given if glucose regulation does not occur. However, multiple doses do pose more risk of developing severe hypoglycemia, especially if discontinuation of dextrose infusions occurs. A constant rate infusion of inR sulin (Vetsulin ) may be a more appropriate choice in difficult cases (0.00125–0.05 IU/kg/h i.v. constant
rate infusion (CRI)). Modest levels of dextrose supplementation should be maintained during glucose titration to prevent hypoglycemia. Table 21.6 lists common scenarios requiring fluid supplementation and appropriate fluid choices and supplements.
Assessing the need for fluids Fluid therapy is often dictated after clinical examination alone. Estimation of dehydration is covered above. In addition to clinical examination, several laboratory data can be used to guide fluid treatment plans. The most basic forms of testing are urine specific gravity (USG) and handheld field lactate monitors (Accutrend Lactate Analyzer; www.lactate.com). Foals that are nursing normally will void large volumes of hyposthenuric urine (USG < 1.008). Concentrating the urine compensates mild dehydration. Fluid supplementation should be considered when USG rises above 1.020. This may be in the form of milk given via a nasogastric tube, increasing constant rate infusions, or administration of replacement fluid boluses.
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Lactate monitoring is easily performed in the field. Hyperlactatemia, > 2.0 mmol/L, occurs with circulatory, pulmonary, or anemic conditions that limit oxygen delivery and with systemic inflammatory response syndrome, hepatic dysfunction, hyperglycemia, and increased catecholamine levels.15, 16 Lactate levels will correspond to the severity of dehydration and are also used to monitor a response to fluid therapy. Therefore, serial lactate measurements are used on foals treated on the farm to gauge a clinical course and the necessity for further intervention or hospitalization. PCV and TP levels are commonly used to guide fluid administration. However, there are some notable limitations to PCV/TP utility. Hemoconcentration, PCV 40–48%, is initially seen in the 1-day-old foal before it has nursed adequate amounts of milk. Total protein levels at this time (< 12 hours old, range 4.0–7.9 g/dL) are low owing to hypoglobulinemia. Normal foals, post-colostrum intake, will retain a PCV between 28% and 32% and will have a TP between 5 and 6 g/dL. With these values in mind, a dehydrated patient may hemoconcentrate, but not to the degree that adult patients do: PCV > 50%, TP > 8 g/dL. Using this information, PCV/TP are useful data in monitoring foals but they should not be the only information used to dictate fluid needs. Dehydration may lead to prerenal azotemia. The placenta is responsible for removing nitrogenous waste in utero; therefore, elevations in blood urea nitrogen (BUN) and creatinine do not always indicate dehydration during the first few days of life. Clinical examination, including assessment for uroabdomen, should be performed in a 1-day-old foal to determine the significance of azotemia. This distinction between in utero accumulation of nitrogenous wastes and prerenal or renal is sometimes difficult and one should err on tailoring treatment plans to avoid nephrotoxic medication and provide some degree of fluid support in nebulous cases. Azotemia noted in foals after the immediate postpartum period should be considered specific for dehydration, renal failure, or uroperitoneum.1 Central venous pressure (CVP) is a useful tool in regulating fluid therapy in foals instrumented with a central line that approximates the level of the vena cava or right atrium. A 20-cm i.v. catheter inserted half-way down the jugular vein will allow the tip to be positioned within the thoracic cavity and is suitable for CVP measurement. A water manometer is used to measure the pressure and should be placed at the level of the sternum in a recumbent foal and at the same height as the shoulder in a standing
patient. Normal CVP in foals less than 14 days old is 2–9 mmHg. Elevations in CVP are seen with volume overload. Decreases in CVP indicate hypovolemia.17
References 1. Spurlock SL, Furr M. Fluid therapy. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 671–700. 2. Palmer JE. Fluid therapy in the neonate: not your mother’s fluid space. Vet Clin North Am Equine Pract 2004;20:63–75. 3. Faubel S, Topf J. The Fluid, Electrolyte and Acid-Base Companion. Chelsea, MI: Sheridan Books, 1999. 4. Plumb DC. Veterinary Drug Handbook, 3rd edn. Ames: Iowa State University Press, 1998. 5. AAEP. White Paper: Information on equine plasma and serum products for the equine practitioner, 2009. http://www.aaep.org/press room.php?id=340 6. Perel P, Roberts I, Pearson M. Colloids versus crystalloids for fluid resuscitation in critically ill patients. Cochrane Database Syst Rev 2008;17(4):CD000567. www.cochrane. org/reviews/en/ab000567.html 7. Magdesian KG, Fielding CL, Madigan JE. Measurement of plasma colloid osmotic pressure in neonatal foals under intensive care: comparison of direct and indirect methods and the association of COP with selected clinical and clinicopathologic variables. J Vet Emerg Crit Care 2004;14:108–14. 8. Dewitt SF, Paradis MR. Use of human albumin as a colloidal therapy in the hypoproteinemic equine. J Vet Emerg Crit Care 2005;15:S15. 9. Belgrave RL, Hines MT, Keegan RD, Wardrop KJ, Bayly WM, Sellon DC. Effects of a polymerized ultrapurified bovine hemoglobin blood substitute administered to ponies with normovolemic anemia. J Vet Intern Med 2000;16:396–403. 10. Soupart A, Decaux G. Therapeutic recommendations for management of severe hyponatremia: current concepts on pathogenesis and prevention of neurologic complications. Clin Nephrol 1996;46:149–69. 11. Beyer MJ, Freestone JF, Reimer JM, Bernard WV, Rueve ER. Idiopathic hypocalcemia in foals. J Vet Intern Med 1997;11:356–60. 12. Lester GD. Maturity of the neonatal foal. Vet Clin North Am Equine Pract 2005;21:333–55. 13. Branco RG, Tasker RC, Garcia PCR, Piva JP, Xavier LD. Glycemic control and insulin therapy in sepsis and critical illness. J Pediatr (Rio J) 2007;83(5 Suppl): S128–36. 14. Van den Berghe G, Wilmer A, Hermans G, Meersseman W, Wouters PJ, Milants I, Van Wijngaerden E, Bobbaers H, Bouillon R. Intensive insulin therapy in the medical ICU. N Engl J Med 2006;354:449–61. 15. Marino PL. 1998. The ICU Book, 2nd edn. Philadelphia: Williams & Wilkins. 16. Franklin RP, Peloso JG. Review of the clinical use of lactate. In: Proceedings of the 52nd Annual Convention of the American Association of Equine Practitioners, 2006, pp. 305–9. 17. Corley KTT. Fluid therapy. In: Bertone J, Horspool LJI (eds) Equine Clinical Pharmacology. Philadelphia: Elsevier Health Sciences, 2004; p. 330.
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Chapter 22
Respiratory Assistance Jonathan E. Palmer
Respiratory support including mechanical ventilation has been used successfully in neonatal foals for more than 30 years and has been recently reviewed.1, 2 Modern mechanical ventilators designed for human medicine have made delivery of this intensive care modality to neonatal foals more successful, and has considerably simplified its application. Ventilating foals without respiratory failure, such as those suffering from botulism or neonatal encephalopathyassociated central hypercapnia, has become very successful.1, 3 However, foals with respiratory failure secondary to severe sepsis, acute respiratory distress syndrome or primary pneumonia remain a major challenge, and successful outcomes in many cases are only achieved utilizing all the accumulated knowledge and skills available.
Indications for respiratory support The most useful and, simultaneously, the most dangerous drug in intensive care is oxygen. Because of this duality, oxygen therapy should not be universally applied, but based on careful monitoring. Neonates are uniquely suited to low oxygen levels because of hypoxic preconditioning as a fetus. Because of the recognition of the possible toxicity and lack of evidence that high oxygen concentrations are necessary, the recommendations for trigger levels for initiating therapy and therapeutic target levels are changing, especially in the presence of lung inflammation.4 Currently the author recommends oxygen therapy in any neonate with a Pa o2 < 60 mmHg (< 8 kPa (kiloPascals)) or a Sa o2 (oxygen saturation of hemoglobin in arterial blood) < 88%, with a goal of oxygen therapy to maintain the Pa o2 > 60 but < 100 mmHg (> 8 but < 13.3 kPa) and a Sa o2 > 90% but < 94%. Overshooting these goals is common with oxygen therapy, but the aim should be to continually titrate to these goals. Oxygen content should also be monitored, but care must be taken that calculations
are based on measured saturation and the patient’s hemoglobin level. Many blood gas analyzers use standard hemoglobin levels and calculated saturation when calculating oxygen content in limited analysis modes, which results in erroneous reassurance. Neonates with a high resting PCV may have enough oxygen-carrying capacity that delivery is adequate despite a low Pa o2 and poor Sa o2 . Neonates with mild anemia resulting in decreased oxygen content will still have enough reserve content to meet tissue needs as long as lung loading is normal and Pa o2 and Sa o2 are normal, allowing for adequate unloading at the tissues. However, the combination of marginal anemia and poor lung loading, producing a low Pa o2 and Sa o2 , will result in failure to deliver enough oxygen for tissue needs. Since neonates frequently have pulmonary problems which lead to periods of hypoxemia, in the face of mild anemia (PCV < 20%), it is important to consider a blood transfusion as an important therapeutic option. Just as the trigger for oxygen therapy has been dropping, there is growing uncertainty about how much of the hypercapnic acidosis which occurs in some critical patients should be corrected. Permissive hypercapnia is a term that describes the technique of ventilation in acute lung injury which allows a degree of hypercapnic acidosis, so that gentler ventilation settings can be used to spare the lungs mechanical trauma.5 As experience with permissive hypercapnia has broadened, it has become evident that the positive effect goes beyond the decrease in mechanical damage and, in fact, the concept of therapeutic hypercapnic acidosis has developed.6–9 Hypercapnic acidosis decreases lung inflammation by suppressing the inflammatory response, reducing nitric oxide production, modifying neutrophil-mediated mechanisms, protecting lung barrier function, and has complex effects on reactive oxygen species.5, 7, 8 The protective effect appears to occur even after the inflammatory response has been initiated.7, 9
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Despite the fact that hypercapnia will decrease the contractility of cardiac smooth muscle (an effect mediated by acidosis), cardiac output will actually increase because hypercapnia will also cause an increased preload, heart rate and coronary blood flow and decreased afterload (due to systemic vasodilatation).5, 10 Hypercapnia will also cause improved ventilation/perfusion matching and reduce intrapulmonary shunting, resulting in improved oxygen loading and improved microcirculatory blood flow, reduced tissue oxygen demand and release of oxygen from hemoglobin at lower oxygen tensions, leading to improvements in tissue aerobic function.5, 10 So oxygen loading, delivery and tissue utilization can all be improved by hypercapnia. Hypercapnic acidosis has been shown to have both beneficial effects (improved cerebral blood flow, oxygen delivery, and anti-inflammatory effects) and adverse effects on neonates suffering from neonatal encephalopathy.5, 10 Hypercapnic acidosis can have other systemic negative effects. It appears that mild hypercapnic acidosis may be beneficial whereas extreme hypercapnic acidosis is detrimental. Not knowing where the tipping point is makes firm recommendations difficult, and the multitude of effects means that that point may vary between individuals, depending on their problems. My current practice is to tolerate a pH as low as 7.30 and Pco2 as high as 65 mmHg as long as the foal appears to be stable, and when I correct hypercapnic acidosis to watch for, not only an improvement in blood gas values, but also deterioration of perfusion or end-organ function, with the willingness to be less aggressive in reversing the hypercapnic acidosis if I see a deterioration in the systemic stability. (see Chapter 18).
Oxygen therapy The usual technique for delivering oxygen insufflation involves the use of a nasal cannula with the tip in the pharynx at the level of the medial canthus of the eye (Figure 22.1). Prior to entering the patient, the oxygen should travel through a water-filled humidifier controlled by a flowmeter. The usual flows necessary to maintain an adequate Pa o2 range from 2 to 15 liters per minute, with common flows between 4 and 8 liters per minute. The Fi o2 resulting from intranasal oxygen cannot be predicted easily because it will vary with placement of the nasal cannula, patency of the cannula openings, tidal volume, minute volume, and size of the foal’s nares. Complications of intranasal oxygen insufflation include oxygen toxicity and nasal irritation, rhinitis, and airway drying resulting in excessive tracheal and nasal discharge. Nasal irritation resulting in a rhinitis is the most common complication. This will be mani-
Figure 22.1 Foal receiving intranasal oxygen insufflation. Note intranasal line entering nostril, secured with a taped stent.
fested by teeth grinding, upper airway moisture, and the development of an occasionally profuse nasal discharge. This nasal irritation is in part due to the physical presence of the cannula and in part due to the drying effect of the oxygen flow despite its humidification. Tracheal drying will also result in tracheal mucosal irritation, leading to proliferation of mucusproducing goblet cells contributing to the discharge. The higher the flow rate, the more severe the irritation and copious the discharge. Ensuring adequate cardiac output is important not only in oxygen delivery to the tissues, but also in providing pulmonary perfusion allowing for adequate ventilation/perfusion matching necessary for effective gas transport in the lungs. This often means the use of inopressor therapy.11 Our recent experience also suggests that the combination of dobutamine and vasopressin therapy can be quite helpful in maintaining perfusion.11, 12 Another key to maintaining ventilation/perfusion matching in certain cases is maintaining the neonate in sternal recumbency. Maintaining sternal recumbency can increase the Pa o2 by 10 to 20 mmHg.12 However, with adequate lung oxygen loading with the help of intranasal oxygen insufflation, many foals will not benefit from forced sternal recumbency and, indeed, if the foal fights or shifts position in a sternal support frame it may be counterproductive. In general, the need to keep all neonatal foals in sternal recumbency is overrated in equine neonatal practice.
Chemical adjuncts Although mechanical ventilation is the definitive treatment for hypoventilation, if the hypoventilation is secondary to blunted central carbon dioxide receptor sensitivity, as often occurs with neonatal encephalopathy, then the use of caffeine or doxapram as a pharmacological respiratory stimulant is indicated. As mentioned above, mild hypercapnia usually has more advantages than disadvantages and
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Respiratory Assistance usually does not require therapy. Severe hypercapnia is dangerous and should be treated by assisted ventilation when possible. But when this is not possible, respiratory stimulants are a reasonable option although they are less likely to be effective in the face of severe hypercapnia. Caffeine (10 mg/kg per os or per rectum) is the safest methylxanthine for use in foals with central respiratory center depression. The effect is usually seen within 2 hours of dosing by a decrease in the hypercapnic acidosis. The dose may need to be repeated s.i.d. or b.i.d. until the respiratory center sensitivity improves. It will only be effective in lowering the Pa co2 when there is a primary respiratory acidosis. When the Pa co2 is increased in an attempt to balance the pH secondary to a metabolic alkalosis, caffeine has less effect. It will, however, act as a general CNS stimulant and transform a somnolent, poorly responsive neonate into a more active patient.12 Doxapram (CRI 0.02 and 0.05 mg/kg/h i.v.) has also been used successfully to treat hypercapnia; however, an increase in metabolic acidosis was associated with its use.13 Several conditions of neonates can lead to the development of pulmonary hypertension resulting in right to left shunting through the still patent foramen ovale and ductus arteriosus. Actually, the ability to shunt blood in the face of pulmonary hypertension is an adaptive advantage, insuring adequate systemic cardiac output and thus avoiding systemic ischemia which would potentiate the damage caused by the hypoxemia. The neonate’s unique ability to exist in a hypoxemic state, and its ability to maintain systemic cardiac output, allows compensation and the ability to survive pulmonary hypertension without generalized systemic ischemic episodes. The advent of inhaled nitric oxide (NO) therapy has revolutionized the treatment of neonatal pulmonary hypertension, and appears to be effective in the large animal species.2, 14, 15 Inhaled nitric oxide at rates as low as 5 to 10 ppm can reverse pulmonary hypertension. But NO can also cause significant pulmonary toxicity through the production of free radicals in the presence of high oxygen concentrations. Therefore, inhaled concentrations of NO must be carefully controlled.16 Phosphodiesterase inhibitors such as sildenafil (0.5–2.5 mg/kg p.o.) may also be useful in pulmonary hypertension in foals.2 Although it is too early to predict the success or anticipate the complications of this therapy, limited anecdotal experience is promising.
Mechanical ventilation Mechanical positive pressure ventilation can address a number of key aspects of respiratory failure (Figure 22.2). Mechanical ventilation can sup-
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Figure 22.2 Foal receiving positive pressure ventilation.
port and allow manipulation of pulmonary gas exchange, increase lung volume returning normal functional residual capacity, decrease the work of breathing allowing ventilatory muscles to rest when fatigued, and decrease the oxygen, energy and perfusion needed to support the work of breathing, thereby freeing these resources for other vital organs. Clinical indications for mechanical ventilation in the neonate include persistent pulmonary hypertension, acute respiratory failure, neonatal encephalopathyassociated weakness or central respiratory center failure, weakness associated with prematurity, neurologically mediated or sepsis-induced hypotension, septic shock, and botulism. Mechanical ventilation is the most effective therapeutic intervention for acute respiratory failure as defined as failure of gas exchange. Causes of acute respiratory failure in neonates include acute respiratory distress syndrome, pulmonary dysfunction secondary to sepsis, infectious pneumonia (viral, bacterial or aspiration pneumonia), non-infectious pneumonia (meconium aspiration, interstitial pneumonia, aspiration pneumonia), and trauma secondary to fractured ribs. Typically the goal is to provide respiratory support while therapies for underlying causes of the acute event are initiated.17–19 Mechanical ventilation may improve gas exchange by increasing ventilation, improving ventilation/ perfusion (V/Q) matching, decreasing intrapulmonary shunting, and decreasing the work of breathing in pulmonary failure and septic shock. Respiratory support through mechanical ventilation can be an important therapeutic modality in treating septic shock.20, 21 The most useful ventilator modes in most foals include synchronized intermittent mandatory ventilation (SIMV) and pressure support ventilation (PSV) both with positive end expiratory pressure/continuous positive airway pressure (PEEP/CPAP). PSV is an example of flow-cycled mechanical ventilation. A preset airway pressure is
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applied once the machine is triggered and is cycled off after the inspiratory flow decreases to a predetermined percentage of its peak value. This mode is readily accepted by most foals and is very useful as long as the foal does not suffer from central respiratory center insensitivity. Some ventilators have a variation of this mode called volume support ventilation, where the ventilator adjusts the target pressure support on each breath based on the last breath’s tidal volume, to attempt to meet a target tidal volume.2, 22 SIMV is a combination of spontaneous ventilation and assist/control ventilation. The delivery of the mechanical breath is synchronized to support the patient’s spontaneous breaths at a preset rate. If spontaneous breathing occurs at a rate faster than the ventilator-set SIMV rate, the patient takes a breath with no preset volume or pressure. If the patient’s spontaneous efforts slow or stop, the ventilator delivers breathes by default at the SIMV rate. SIMV is well tolerated, since breaths above the preset rate are completely controlled by the patient (timing, depth, duration). The preset breaths are still at a fixed and unforgiving volume or pressure to insure a minimum minute volume. PEEP refers to the maintenance of positive end expiratory pressure in the airways between ventilatorinduced positive pressure inspirations (during exhalation and between breaths) so that the airway pressure never falls below the set PEEP value. CPAP refers to the maintaining continuous positive airway pressure throughout spontaneous respiration (during inspiration, exhalation, and between breaths). The primary physiological effect of PEEP/CPAP is to increase functional residual capacity by maintaining patency of alveoli at the end of exhalation. PEEP/CPAP is often effective in treating hypoxemia by decreasing intrapulmonary shunting and ventilation/perfusion mismatch. Beneficial physiological effects of PEEP/CPAP are created by an increased transpulmonary pressure resulting in an increased functional residual capacity (FRC) and stabilization of an unstable chest wall. PEEP/CPAP affects pulmonary mechanics, cardiovascular stability, and pulmonary vascular resistance. With the upper airway bypassed by tracheal intubation, sufficient heat and moisture must be added to the inspired gas mixtures to prevent mucosal injury secondary to drying and cooling. The simplest and most effective way to precondition the gases is by utilizing a passive humidifier such as a heat-moisture exchange device (HME filter) that utilizes heat and moisture trapped from expired gases. Many HME filters act as a very efficient antimicrobial filter, excluding nosocomial bacteria and viruses from the patient and confining the patient’s pathogens to the endotracheal tube and airway.
Monitoring during ventilation is important to allow dynamic adjustment of mechanical ventilation parameters, to understand and correct underlying pathophysiology, to prevent damage from the act of mechanical ventilation, and to decide if it is time to discontinue mechanical ventilation. The most useful parameters which can be monitored include: arterial blood gas, end-tidal carbon dioxide (capnography), fraction of inspired oxygen, tidal volume/minute volume, airway pressure, compliance/resistance, and patency of the endotracheal tube. Since the need to use mechanical ventilation is often emergent, having a plan in place before the need arises is essential. Having a ventilator setup checklist, dry runs with the staff practicing setting up the equipment, and sessions covering equipment troubleshooting procedures can minimize confusion and uncertainty when the need for ventilation arises. Typical equipment needed includes: the ventilator, access to oxygen (with minimal interruption of the patient’s oxygen insufflation), access to medical grade compressed air or a compressed air generator, interface lines, gas blender (often built into the ventilator), capnograph with lines and adaptor, humidifying device (often an HME filter), ventilator circuit, endotracheal tube, sterile gloves, sterile lubrication, means of securing the endotracheal tube, stethoscope, selfinflating bag (in case of an emergency) and, most importantly, adequate trained help to restrain the foal, intubate the foal, secure the endotracheal tube, and begin adjusting the ventilator settings as soon as the foal is intubated. As part of setting up the ventilator, the circuit and other attachments should be inspected to be certain that everything is in proper working order and the circuit should be checked for leaks. This can be done by attaching an artificial lung to the circuit, occluding the exhalation port and charging the circuit with a manual breath or checking the ventilator’s ability to maintain PEEP. It is convenient to select the initial ventilator setting after checking for leaks (see Table 22.1). A selection of 55 cm long, 7 to 10 mm internal diameter nasotracheal tubes should be available. The largest tube which will fit should be used to minimize the resistance generated by the tube. It is also important to check the integrity of the endotracheal tube’s cuff before intubation. Leaking endotracheal cuffs are a very common problem during ventilation. Sterile endotracheal tubes should be used to minimize introduction of nosocomial bacteria during intubation. While checking the cuff, care should be taken to maintain sterility of the tube beyond the cuff inflation port. During setup, choice of heating and humidifying the delivered gas should be made and set up. When placing the foal on the ventilator,
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Table 22.1 Common initial ventilator settings Parameter
Setting
Comments
Tidal volume
6–10 mL/kg
Set for low airway pressure but adequate ventilation
Inspiratory pause/plateau
0–0.2 sec (range 0–2 sec) 0–5% inspiratory time (range 0–20%)
Goal to improve distribution of ventilation, especially when airway resistance is an issue
Breath rate
20–30 bpm
Adjust with capnography and ABG for adequate ventilation
Peak flow
60–90 lpm
Highly dependent on breath rate, inspiratory pause, I/E ratio (initially I/E ∼ 1:2)
Fio2
0.5 (range 0.3–1.0)
Initial setting based on pre-ventilation response to oxygen therapy; titrated with ABG; goal < 0.5
PEEP/CPAP
4–5 cmH2 O (range 0–20)
Should be adjusted based on compliance and P a o2 values
Trigger sensitivity
2–3 cmH2 O
Do not set so low that auto-triggering occurs
Pressure support
8–12 cmH2 O
Set higher with low compliance; increase if tidal volume is too small (tidal volume may not respond to pressure support level)
I/E ratio, inspiratory/expiratory ratio; F i o2 , fractional inspired oxygen; ABG, arterial blood gas; PEEP, positive end expiratory pressure; CPAP, continuous positive airway pressure.
adequate restraint for intubation is needed but, in the author’s experience, sedation is not needed while establishing or maintaining ventilation. Use of sedatives could unnecessarily complicate the clinical course of the case. Having an uncooperative foal usually indicates poor choices of ventilatory mode or parameters. Complications of mechanical ventilation include oxygen toxicity, nosocomial infection (pneumonia or bacteremia), volutrauma/barotrauma (secondary to high levels of PEEP, excessive tidal volumes, high peak airway pressures, and coexistence of significant lung disease), laryngotracheal injury (very rare with modern endotracheal tubes), and irritation of the upper airway resulting in excessive respiratory secretion production. The most common and serious of these are secondary infections and airway irritation. It is the author’s routine to change the endotracheal tube once a day, both to avoid increased resistance caused by the collection of dried respiratory secretions in the tube lumen and to monitor for the development of respiratory infection. Finding excessive tenacious secretion adherent to the endotracheal lumen is characteristic of improper preconditioning of the respiratory gases (heating and humidification), which should be addressed. Changes of character of the respiratory secretions accumulating in the endotracheal tube lumen (yellow or dark red color, odor, etc.) should alert the clinician of developing infection. It is also the author’s routine to periodically culture the lumen of the removed endotracheal tube. The bacteria grown may not be established in the respiratory tract but culture results forewarn the clinician of those pathogens which may become established and also give insight on the antimicrobial sensitivity of
potential pathogens, helping in the antimicrobial selection if infection develops. Critically ill foals often have respiratory failure and benefit from respiratory support. Some only require intranasal oxygen insufflation whereas others need full respiratory support with mechanical ventilation. Modern mechanical ventilators developed for human medicine are easily adapted to foals, and make mechanical ventilation of foals not only possible but frequently successful. Conventional mechanical ventilation using SIMV or PSV along with PEEP/CPAP is useful in many foals. As soon as the foal is begun on mechanical ventilation, the clinician should ask himself if the foal is ready to be weaned. Early weaning is as important as timely initiation of ventilation. Monitoring also aids in deciding when it is appropriate to wean the foal.
References 1. Esteban A, Anzueto A, Frutos F, Alia I, Brochard L, Stewart TE. Characteristics and outcomes in adult patients receiving mechanical ventilation: a 28-day international study. J Am Med Assoc 2002;287:345–55. 2. Fenstermacher D, Hong D. Mechanical ventilation: What have we learned? Crit Care Nurs Q 2004;27(3):258–294. 3. Gali B, Goyal DG. Positive pressure mechanical ventilation. Emerg Med Clin North Am 2003;21(2):453–73. 4. Giguere S, Slade JK, Sanchez LC. Retrospective comparison of caffeine and doxapram for the treatment of hypercapnia in foals with hypoxic-ischemic encephalopathy. J Vet Intern Med 2008;22:401–5. 5. Gross I. Recent advances in respiratory care of the term neonate. Ann NY Acad Sci 2000;900:151–8. 6. Jankov RP, Tanswell AK. Hypercapnia and the neonate. Acta Pædiatr 2008;97:1502–9.
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7. Kavanagh BP, Laffey JG. Hypercapnia: permissive and therapeutic. Mikerva Anestesiol 2006;72:567– 76. 8. Laffey JG, Kavanagh BP. Carbon dioxide and the critically ill – too little of a good thing? Lancet 1999;354:1283– 6. 9. Lester GD, DeMarco VG, Norman WM. Effect of inhaled nitric oxide on experimentally induced pulmonary hypertension in neonatal foals. Am J Vet Res 1999;60(10):1207– 12. 10. Michelet P. Hypercapnic acidosis: How far? Anesthesiology 2008;109:771–2. 11. Palmer JE. Ventilatory support of the neonatal foal. Vet Clin North Am Equine Pract 1994;10(2):167–86. 12. Palmer JE. Use of inhaled nitric oxide in improving pulmonary gas exchange in persistent pulmonary hypertension and pulmonary failure secondary to septic shock in the neonatal foal and calf. In: Proceedings of the 2nd Dorothy Havemeyer International Workshop on Neonatal Septicemia, Boston: 1998; pp. 36–7. 13. Palmer JE. When fluids are not enough: Inopressor therapy. In: Proceedings of 8th International Veterinary and Critical Care Symposium, San Antonio, TX: 2002; pp 220–4. 14. Palmer JE. Foal cardiopulmonary resuscitation. In: Orsini JA, Divers TJ (eds) Manual of Equine Emergencies: Treat-
15. 16.
17.
18.
19.
20. 21. 22.
ment and Procedures, 2nd edn. Philadelphia: Saunders, 2003; pp. 581–614. Palmer JE. Ventilatory support of the critically ill foal. Vet Clin North Am Equine Pract 2005;21(2):457–86. Perl M, Radermacher P. Hypercapnic acidosis in acute lung injury: Inevitable side effect or unexpected benefit? Crit Care Med 2008;36(12):3268–9. Pollack CV. Mechanical ventilation and noninvasive ventilatory support. In: Marx JA, Hockberger RS, Walls RM (eds) Marx-Rosen’s Emergency Medicine: Concepts and Clinical Practice, 5th edn. St Louis, MO: Mosby, 2002; pp. 21–8. Randolph AG. Management of acute lung injury and acute respiratory distress syndrome in children. Crit Care Med 2009;37:2448–54. Slutsky AS. Mechanical ventilation (American College of Chest Physicians’ Consensus Conference). Chest 1993;104:1833–59. Swenson ER. Therapeutic hypercapnic acidosis: pushing the envelope. Am J Respir Crit Care Med 2004;169:8–9. Tobin MJ. Medical progress: advances in mechanical ventilation. N Engl J Med 2001;344:1986–96. Wilkins PA, Palmer JE. Mechanical ventilation in foals with botulism: 9 cases (1989–2002). J Vet Intern Med 2003;17(5):708–12.
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Chapter 23
Sedation, Anesthesia and Analgesia Lori A. Bidwell
Introduction Anesthesia and sedation of foals introduces the challenge of minimizing depression of a sensitive cardiovascular system while avoiding unnecessary stress on an immature metabolic system. In terms of anesthesia, foals can be divided into neonates (typically less than 1 week, but can include foals up to 1 month of age) and metabolically or physiologically mature foals (typically those older than 4–6 weeks of age).1 Foals older than 6 months of age can be treated like adults in regards to anesthetic protocols. Unhealthy foals have physiological characteristics similar to a neonate and should be treated as such when determining sedative and anesthetic protocols. During the first few days of life, a neonate is more sensitive to anesthetics due to increased permeability of the blood–brain barrier, an immature sympathetic nervous system that cannot respond to anestheticinduced vasodilation resulting in hypotension and hypothermia, a lower percentage of body fat to assist in redistribution of anesthetic, higher body water content altering drug distribution, immature hepatic metabolism, immature renal function, and a higher metabolic rate.2 Due to alterations in liver enzyme activity and relative hepatic mass, until foals reach 1 to 2 weeks of age, they should be treated as if they are incapable of complete hepatic metabolism.3, 4 Because of an immature hepatic system, neonatal patients have lower albumin concentrations that can result in increased free portions of drugs that are usually protein bound. Use of drugs that are highly protein-bound like phenylbutazone and barbiturates and drugs that require significant hepatic metabolism like opioids, ketamine, lidocaine and alpha-2 agonists should be used with discretion in neonates and unhealthy foals. Evidence has shown that there are differences in diazepam clearance
between 4 days of age and 21, 42, and 84 days of age. Clearance of drug was found to be significantly less in foals 4 days of age versus 21 days and older, therefore caution should be used in re-dosing diazepam in neonates as this drug can accumulate.5 An immature sympathetic nervous system can result in the inability to respond to the vasodilatory and cardio-depressant effects of anesthetics. Healthy foals have evidence of cardiac right-to-left shunt for the first 3 days to week of life; this can persist in unhealthy foals.6 Sensitivity of foals to the deleterious effects of anesthetic often results in vasodilation, bradycardia, and resulting hypotension. There appears to be a fine line between enough anesthetic and too much in foals. Therefore, a balance between preoperative and intraoperative analgesics and sedation and inhalant or total intravenous anesthesia is imperative as less of each drug minimizes side effects. Otherwise, the anesthetist will need to increase inhalant or intravenous anesthetic in response to arousal and the result is cardiovascular depression that must then be dealt with using positive inotropes, chronotropes, and/or fluid therapy. In adult horses, total intravenous anesthesia (TIVA) has shown promise as a safe means of anesthesia for procedures of less than 1 hour7, 8 but caution must be used when administering TIVA combinations such as triple drip (guaifenesin, xylazine, and ketamine) to foals as decreased metabolism can result in prolonged and rough recoveries. Nasotracheal intubation provides an alternative to intravenous techniques in neonates and compromised foals. Nasotracheal intubation avoids the excitement associated with mask induction and minimizes personnel exposure to the anesthetic. The most commonly used inhalant anesthetics offer the advantage of having minimal metabolic
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requirements, therefore allowing quick recovery at the termination of the anesthetic.9 The technique is most appropriate when a foal is recumbent and offers little resistance to placement of a nasotracheal tube via the ventral meatus. Endotracheal tubes used for nasotracheal intubation are smaller in internal diameter (9 mm or 10 mm) but longer (50–55 cm) than those used for oral intubation. Active neonates or foals can be nasally intubated with physical restraint but this can be facilitated by using 3–5 mL of 2% lidocaine sprayed into the nostril. If the foal is active enough to make endotracheal tube placement difficult, then a pre-anesthetic sedative/analgesic and intravenous induction are more appropriate. Additional complications that must be considered with neonates and foals are hypothermia, hypercapnia, and dependent lung atalectasis. Increased ratio of surface area to body mass results in rapid loss of body heat. This should be taken into account when selecting a location for procedures. A warm stall with soft bedding and a blanket will assist in maintaining normothermia. Circulating hot water blankets are recommended for longer procedures as well as appropriately placed diapers or towels to prevent wetness from urination or lavage fluids. Hypercapnia occurs in sedated and anesthetized foals in lateral or dorsal recumbency. Although the foal appears to be ventilating adequately, Pa co2 can rise to levels well above 60 mmHg. Acute elevations in the arterial partial pressure of carbon dioxide above 90 mmHg have been found to result in narcosis in humans.10 Therefore, high elevations in carbon dioxide should be prevented as elevations could prolong and enhance the effects of injectable or inhalant anesthetics. Dependent lung atalectasis from prolonged lateral recumbency can be deleterious in foals that have respiratory disease or have suffered from a hypoxic episode. Oxygen supplementation by nasal insufflation can decrease the risk associated with prolonged recumbency. Corneal abrasions leading to ulceration can occur in any foal that is sedated or anesthetized. Ocular lubrication is inexpensive and should always be placed in both eyes when a foal is sedated for a procedure or anesthetized. The expense of a tube of sterile ophthalmic lubrication is much less than the treatment of a corneal ulcer. Additionally, the mare should remain with the foal during sedation and until complete induction has occured when a procedure requires general anesthesia. Removal of the mare during sedation or induction will produce excitement resulting in poor sedation or induction.
Sedation The sedatives used in adult horses can produce deleterious effects in foals; therefore certain drugs should be avoided or doses minimized to prevent complica-
tions. Alpha-2 adrenergic agonists xylazine and detomidine have been associated with mild complications although the high-quality sedation and analgesia often outweigh any negative effects. Xylazine has been reported to produce cardiovascular and respiratory depression although no significant changes were noted in pH or blood gas tensions.11–13 In foals 10 and 28 days old, 1.1 mg/kg intravenous xylazine decreased rectal temperature at 30 min to 120 min. Unlike adults, xylazine did not produce hypoinsulinemia and hyperglycemia.11 Detomidine, the alpha-2 adrenergic agonist, was studied in foals between the ages of 14 to 94 days. Doses of 10, 20, and 40 µg/kg were administered intravenously. All doses produced profound sedation that was mildly increased between the low and high dose. Analgesia was minimal at the low dose and considered good at the high dose. All doses resulted in bradycardia and in some foals ataxia, labored respiration, arrhythmia, sweating, and frequent urination were also noted.14 Cardiac output is dependent on heart rate and stroke volume, the latter being less likely to compensate in foals when heart rate is decreased by drugs such as alpha2 adrenergic agonists. Therefore, sedatives selected for neonates should have minimal cardiovascular depressant effects. Benzodiazepines (valium or midazolam), ketamine, and the opioid agonist-antagonist butorphanol are common choices for sedation in foals (see Table 23.1). These drugs can be administered alone or in combination and given intravenously or intramuscularly. Butorphanol has been studied in 3and 12-day-old pony foals at a dose of 0.05 mg/kg i.v. and i.m. Both the intravenous and intramuscular dosing produced sedation with minimal effects on the cardiovascular system and increased time spent nursing rather than increased locomotor activity as is often observed in adults.15 Administration of phenothiazines (e.g., acepromazine) can result in significant non-reversible vasodilation in foals; therefore its use should be avoided in pediatric patients. Foals less than 1 week of age will lie down when they feel tired or sleepy; therefore a small amount Table 23.1 Pre-anesthetic, sedative and an opioid reversal dose for foals∗ Drug
Dose
Xylazine Detomidine Romifidine Butorphanol Morphine Ketamine Nalaxone (opioid reversal)
0.5–1.0 mg/kg i.v.; 1–2 mg/kg i.m. 0.005–0.01 mg/kg i.v.; 0.01–0.03 mg/kg i.m. 0.04–0.06 mg/kg i.v. 0.02–0.04 mg/kg i.v., i.m., s.c. 0.1–0.3 mg/kg i.v., i.m., s.c. 1–2 mg/kg i.m. 0.002–0.021 mg/kg i.v.
∗
Doses are extrapolated from adult equine references.
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Sedation, Anesthesia and Analgesia Table 23.2 Constant rate infusions∗ Drug
Dose
Reference
Propofol Lidocaine Ketamine Midazolam
0.05–0.2 mg/kg/min i.v. 0.05 mg/kg/min i.v. 0.002–0.01 mg/kg/min i.v. 0.02–0.08 mg/kg/h i.v.
Matthews et al. 199916 Valverde et al. 200517 Muir & Sams 199218 Yamashita et al. 2007;19 Kushiro et al. 200520
∗
Doses extrapolated from adult equine references.
of sedative will usually result in recumbency. This allows for easy handling for procedures such as bandage changing, radiographs, catheter placement, ultrasonographic examination, or procedures to correct entropion. In a 30 to 40 kg neonate, a dose of 2–5 mg of butorphanol with 2.5–5 mg midazolam or diazepam administered intravenously will result in recumbency that lasts 15–30 minutes. With the addition of 100–150 mg ketamine i.v., a more invasive procedure can be performed like a joint lavage or urinary catheter placement. Healthy foals older than 2 weeks might require the addition of an alpha-2 agonist such as xylazine (30–80 mg i.v.) alone or combined with a short-acting opioid like butorphanol, followed by 150–200 mg ketamine i.v. alone or combined with intravenous diazepam or midazolam for more invasive procedures. Older foals (> 1 week of age) will typically not become recumbent when sedated; therefore these foals may need to be placed in lateral recumbency after administration of ketamine/diazepam. Procedures performed in a stall or involving intravenous sedation and anesthesia without the inclusion of oxygen insufflation can result in mild to severe hypoxia. Humidified oxygen delivered at a rate of at least 3–5 L/min through a nasal cannula is recommended. Standing procedures can be performed in foals greater than 1 month of age after sedation with either an alpha-2 agonist or butorphanol. For longer sedation in foals, particularly vigorous dummy foals that need to be quiet enough to be acclimated to or maintained on a ventilator or to protect fluid lines, a urinary catheter or a nasal cannula, intramuscular ketamine combined with midazolam is effective for sedation lasting 2–4 hours. Constant rate infusions of midazolam, ketamine or propofol are other alternatives for sedation lasting 4 to 12 hours (see Table 23.2).
General anesthesia General anesthesia in neonates presents a unique complication as they respond to pre-anesthetics and inhalant anesthetics with vasodilation and decreased cardiac contractility.21 This is a particular problem in sick foals. Pre-anesthetic examination should in-
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clude palpation of the pulse, temperature, auscultation of the heart and lungs, and a minimum data base that includes packed cell volume, total protein, white blood cell count, and fibrinogen. When presented with a sick neonate it is important to evaluate glucose and electrolyte concentrations prior to anesthesia, particularly sodium, potassium, and calcium.22–24 Common elective procedures include umbilical hernia repair and leg straightening procedures like periosteal transection and transphyseal fixation or bridging. Emergency procedures include surgical treatment of ruptured bladder, inguinal hernia, flail chest, and colic. Induction should be performed when the foal is sedate and relaxed with the dam standing within viewing distance. There are various intravenous induction techniques that have been used successfully: the combination of ketamine/diazepam or midazolam, ketamine alone, or propofol. A study comparing induction and maintenance techniques in foals 7 to 15 days old found that the combination of diazepamketamine-isoflurane produced less cardiopulmonary depression than xylazine-ketamine-isoflurane during abdominal surgery.12 Monitoring during any surgical procedure should include electrocardiogram, invasive or non-invasive arterial blood pressure, visually monitoring respiration, mucous membrane color, and pulse quality. Pulse oximetry, capnography, or arterial blood gases to evaluate Pa co2 are recommended to determine oxygenation and quality of ventilation but are not necessary for short elective procedures. It is not uncommon for foals to have Pa co2 levels at 60 mmHg or higher with spontaneous ventilation during elective procedures. Assisting ventilation at 1–2 breaths per minute aids in opening atelectic alveoli and preventing hypercapnia. Maintenance fluids should be administered at a rate of 10–15 mL/kg/h (some foals may require a reduced maintenance fluid rate based on their other underlying problems) but can be administered up to 90 mL/kg/h in emergency or shock situations. It is wise to calculate the required hourly fluid volume prior to anesthesia and use 1 L fluid bags rather than 3 L or 5 L bags in order to minimize excessive fluid administration. Because renal function is immature, fluid administration should be dosed appropriately to avoid the production of pulmonary edema. The minimum alveolar concentration (MAC) of isoflurane in neonates is lower than adults; 0.84% vs. 1.3 to 1.6% for adults.25 Neonates are most sensitive to volatile anesthetics but MAC values reach a maximum in older foals/weanlings around 6 months of age. Table 23.3 compares adult MAC values to available foal values. Foals are sensitive to the vasodilatory effects of volatile anesthetics; therefore it is important to minimize the anesthetic.
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Table 23.3 Minimum alveolar concentration (MAC) values for inhalational anesthetics
Halothane Isoflurane Sevoflurane Desflurane
Foal or neonate MAC
Adult MAC
0.84%
0.88% 1.31% 2.31% 7.02%
Reference Steffey et al. 197726 Dunlop et al. 1989;25 Steffey et al. 197726 Aida et al. 199427 Tendillo et al. 199728
It is typical to begin inhalant anesthesia with the vaporizer settings at 2% isoflurane or 3% sevoflurane and then to turn these concentrations down to 1.5% and 2.5% respectively within 5 minutes of initiating gas anesthesia. The anesthetic concentration required should be determined by clinical signs indicating depth. Eye position, blink reflex and strap muscle tone in the neck are helpful determinants of anesthetic level in adults and foals around 1–2 months of age and older but are not predictable in neonates. Functional residual capacity of the lung is smaller in neonates and therefore induction of anesthesia is faster. It should be assumed that within 5 minutes of introducing inhalant anesthetic, concentrations can be decreased. Blood pressure is helpful in determining depth of anesthesia in foals but vasodilation can occur rapidly despite the fact that a surgical plane of anesthesia has not been achieved. When attempting to place an arterial catheter in a sick or healthy neonate; it is important to place the catheter as quickly as possible as the development of vasodilation makes placement progressively more difficult. Administering a positive inotrope or vasopressor might aid in the placement of the arterial catheter by improving cardiac output or vascular tone but neonates have incomplete beta innervation in the myocardium. Foals are often fluid depleted, therefore a bolus of isotonic fluids or colloids and turning the vaporizer down may be more effective. Mucociliary clearance is impeded by volatile anesthetics; therefore foals with respiratory disease should be monitored postoperatively for worsening of clinical signs like increased cough, wheezing or respiratory distress. Cilia in the airways are paralyzed for up to 3 hours post-anesthesia as a result of the presence of volatile anesthetic. Cilia are responsible for mucus clearance at a rate of 1 mm/min in small airways and up to 20 mm per minute in the trachea.29, 30 Therefore it is best to avoid inhalant anesthesia in foals with significant respiratory disease for procedures of short duration. The alternative to inhalant anesthesia in these cases is the use of injectable anesthetics while delivering 100% humidified oxygen through nasal insufflation or oro-
tracheal/nasotracheal intubation and using oxygen from a vigorously flushed anesthesia machine circle system at a flow rate of 1–2 L/min. Additional boluses of ketamine or a slow administration of triple drip can be adequate for short procedures like periosteal transaction or screw removals. Hypotension Intraoperative adjuncts include positive inotropes (dobutamine, dopamine, ephedrine, and calcium gluconate), pressors such as phenylephrine or vasopressin, anticholinergics, crystalloids, and colloids. Cardiac effects of dobutamine, norepinephrine, and vasopressin have been compared in foals 1–5 days old with induced vasodilation from isoflurane anesthetic. The result of the study determined that infusions of dobutamine at 4 or 8 µg/kg/min and norepinephrine at 0.3 or 1.0 µg/kg/min improved cardiac index and maintained splanchnic circulation better than infusions of 0.3 or 1.0 mU/kg/min of vasopressin.31 In foals 1–2 weeks old anesthetized with isoflurane, norepinephrine, and dobutamine were compared as to their effect on cardiac index and blood pressure. The study found that norepinephrine increased blood pressure but decreased cardiac index and dobutamine increased cardiac index while only producing a mild increase in blood pressure. Both drugs were determined to be useful for treatment of hypotension in foals.32 Although normal mean blood pressure is typically lower in neonates up to 1 week of age (47–50 mmHg), mean arterial blood pressure should be maintained above 55 mmHg in young foals. There has been a case report of a neonatal foal developing fatal postoperative myositis after having a mean arterial blood pressure between 45 and 65 mmHg for a procedure lasting 4 hours.33 Older foals should be treated as adults regarding blood pressure, with a goal of maintaining the mean arterial pressure above 70 mmHg. Inotropes, pressors, and fluids should be administered as needed while minimizing volatile anesthetic administration. Normal heart rate in neonates is between 70–90 bpm and can range between 40 and 60 bpm in older foals.34 Heart rate can have a significant impact on cardiac output in foals, as they have less ability to increase stroke volume in response to hypotension. Therefore, the use of anticholinergics like glycopyrrolate and atropine may be necessary, although the clinical effect may be variable in neonates. Atropine is effective in increasing the heart rate of newborns although its effect is more significant with increasing age.35 Hypothermia and hypoglycemia Hypothermia and hypoglycemia are common in anesthetized foals due to their large surface area to body mass ratio, small percentage of body fat,
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Sedation, Anesthesia and Analgesia decreased ability to compensate for vasodilation, and little to no energy reserves. Shivering increases metabolic rate and oxygen requirements; this is a concern in patients that are hypoventilating or have fatigued muscles of ventilation. Circulating hot water blankets or heated hot water bottles aid in maintaining normothermia. Caution should be taken with hot water bottles in that they can lose their warming capacity quickly and can produce the opposite effect as they cool. Young foals still nursing a mare should not be fasted prior to anesthesia as hypoglycemia can occur. Typically, a weaned foal can be treated like an adult and fasted at least 3–5 hours prior to surgery. Glucose should be measured at regular intervals while a neonate is anesthetized (typically every 30–45 minutes). Hypoglycemia should be treated with intravenous infusions of dextrose-containing fluids while continuing to monitor glucose levels.
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recovery stall. It is helpful to have a blanket to cover the foal while it recovers, to maintain or achieve normothermia. Warm blown air or circulating hot water blankets are ideal to warm the foal as it wakes. All foals should be assisted in recovery, often only requiring one person to hold the foal down until it is ready to stand and then assisting with a hand on the tail for stability. Checking glucose levels in the recovery stall is recommended; slow recoveries allow treatment with dextrose if needed. In the case of elective procedures, it is important that the foal return to the dam and resume nursing as soon as possible. Emergency cases often involve foals that will remain recumbent or will require supportive care but it is still important that these foals be vigorous during recovery rather than remaining in a drug stupor.
Analgesics Ventilation Neonatal foals have a higher respiratory rate (30–40 bpm) than adults. Older foals have a respiratory rate of 12–14 bpm.34 Assisted ventilation is often unnecessary in short elective procedures but assisting one breath per minute will help open airways and reduce dependent lung atelectasis. Longer procedures or emergency cases typically require intermittent positive pressure ventilation, although assisted ventilation is just as, if not more, effective but requires time and energy to perform. Tidal volume is estimated at 10–15 mL/kg. Airway pressure should be maintained below 20 cmH2 O. A mechanical ventilator allows for more thorough patient monitoring when staff is limited. Ventilator size can be a limiting factor when deciding between assisted or controlled ventilation as many of the ventilators appropriate for neonates were designed for humans with maximum tidal volumes of around 1 L. These ventilators are appropriate for foals less than 100 kg but are inadequate for anything larger. Respiratory rates should start at 16–17 bpm in neonates and 12–14 bpm in older foals. Pulmonary function should be monitored with capnography (normal range end tidal CO2 35–60 mmHg) and/or blood gases. It is important to remember that a compliant ribcage allows the lungs to be prone to barotrauma with aggressive assisted ventilation. Serial measurement of blood gases is recommended to monitor the quality of ventilation. Blood gases should be evaluated every 30 minutes in critical cases. The reader is referred to see Respiratory Assistance, Chapter 22 for a more in-depth review of ventilation techniques in foals. Recovery from anesthesia Recovery should be in a warm environment; neonates in the stall with their dam; older foals in a
Minimizing respiratory and cardiovascular depression while providing relief from pain can prove challenging in the neonate and foal. Very few studies have examined analgesia in pediatric equine patients; therefore most drug doses are extrapolated from adult equine, human pediatric, lamb, calf, and companion animal dosages. Pain has been studied extensively in human neonates but has not been analyzed in foals. In human neonates, early pain experiences are associated with alteration in responses to pain as adults. There is some evidence that human neonates that undergo circumcision without adequate analgesia develop pain memory that increases the neurological response to pain later in life.36 Drug choices in neonates can include alpha-2 agonists, opioids, ketamine, tramadol, gabapentin, and non-steroidal anti-inflammatory drugs (NSAIDs). All of these choices can have deleterious effects and most have not been thoroughly studied in the neonate and foal, therefore dosing should be adjusted to the patient’s physiological status (Table 23.4). Intravenous, oral, intramuscular, and subcutaneous administration of many analgesics is acceptable in foals. Constant rate infusion of lidocaine and/or ketamine and/or butorphanol or pure mu opioid agonists can be beneficial in foals for painful procedures under anesthesia and for postoperative care (Table 23.4).
Epidural and intra-articular analgesics Epidural analgesia using opioids, local anesthetics, and alpha-2 agonists is an often-overlooked method of pain management in neonates and foals but can produce benefits with minimal respiratory and cardiovascular depressant effects. From personal experience, epidural placement of 1 mL of 2% lidocaine aids in the treatment of a prolapsed rectum due
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Table 23.4 Analgesic doses for foals
Drug
Intravenous
Intramuscular
Epidural
Constant rate infusion or other
Buprenorphine
0.005–0.01 mg/kg
0.005–0.01 mg/kg
0.005 mg/kg
0.006 mg/kg (sublingual)
Butorphanol
0.02–0.04 mg/kg
0.02–0.04 mg/kg
Not effective
Fentanyl
0.001–0.002 mg/kg
NA
Not effective
Morphine
0.1 mg/kg
0.1–0.3 mg/kg
0.1 mg/kg
Tramadol Phenylbutazone Flunixin meglumine Ketoprofen Lidocaine
0.2–1.6 mg/kg 2–4 mg/kg q 12 h 1 mg/kg q 12–24 h 2–3 mg/kg q 24 h 0.5–2 mg/kg
NA NA Not recommended NA NA
1mg/kg NA NA NA 0.2–0.3 mg/kg
0.01 mg/kg i.v., then 0.02–0.04 mg/kg/hr i.v. 2µg/kg i.v. followed by 1–5 µg/kg/h Patch available: 5–10 mg/ 150 kg 0.16 mg/kg i.v., then 0.1 mg/kg/h NA Oral 2–4 mg/kg Oral 1 mg/kg
Xylazine Gabapentin
0.5–1 mg/kg
1–2 mg/kg
0.17 mg/kg
∗
References Messenger et al.54 Fischer et al. 200937 Arguedas et al. 200815 Eberspacher et al. 200838
Bennett & Steffey 200239 Natalini & Robinson 200040
1.3 mg/kg i.v. followed by 0.004–0.1 mg/kg/min 2.5 mg/kg p.o.
Davis et al. 200741
Many of these values are based on adult doses. NA, not available.
to straining from diarrhea and 0.1 mg/kg morphine with or without 0.25 mg/kg xylazine can be placed in the epidural space for thoracic or hindlimb pain.42 Tramadol and buprenorphine have also been examined in adults for epidural use. Tramadol produced analgesia within 30 minutes of placement that lasted 4–5 hours.40 Buprenorphine combined with detomidine in the epidural space produced analgesia similar to the combination of morphine and detomidine.37 Intra-articular morphine has not been evaluated in foals but has shown promise in adults with no deleterious effects to the joint surface and synovium at a dose of 0.1 mg/kg.43, 44 Transdermal patches Transdermal fentanyl patches have been shown to be a safe means of analgesia although variable in duration and plasma concentrations in adult horses.45, 46 A study in foals found that detectable plasma levels result within 20 minutes of placement but are variable and produce an elevation in body temperature that does not coincide with elevated fentanyl concentrations.38 From personal experience, fentanyl patches produce analgesic effects within 2–4 hours of placement on foals with few systemic side effects and several days duration from a single patch application. The patches are sold as 2.5, 5.0, 7.5, and 10.0 mg sizes that deliver 25, 50, 75 and 100 µg/h doses, respectively, and the recommended dose is 10 mg per 150 kg bodyweight although conservative doses are clinically accepted, particularly in neonates.47 If only
higher concentration patches are available, part of the patch can be covered with tape to minimize the surface contact area before application. Fentanyl patches are dependent on systemic uptake, therefore location over a major vessel (e.g., cephalic vein) will aid in the effectiveness. Lidocaine patches are available in a 5% concentration situated on a 10 × 14 cm felt backing that can be cut to sizes appropriate to the location. An unpublished study carried out by the author found that the patch was effective for arthritis pain (objectively studied using a force plate) and produced 95% improvement in lameness within 30 minutes of patch application over a septic joint in a foal (subjectively determined). The patch should be placed directly over the area that requires analgesia after prepping the site by clipping with a number 10 blade. If a bandage can be placed over the area, it is recommended. Otherwise, tissue glue assists in maintaining placement of the patch over a joint or irregular surface area. Lidocaine patches have been shown to be dependent on a local effect rather than systemic uptake.48 Non-steroidal anti-inflammatory drugs Non-steroidal anti-inflammatory drugs are an important part of an analgesic protocol for most equine cases. The NSAIDs ketoprofen (available for intravenous administration) and flunixin meglumine (intravenous and oral suspension) have been studied in neonates less than 24 hours old and were
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Sedation, Anesthesia and Analgesia found to have larger volumes of distribution than older foals and adults but clearance was significantly lower. Consequently, doses should be increased to as much as 1.5 times an adult dose to achieve equivalent analgesia but dosing intervals should be prolonged to avoid toxicity.49, 50 Phenylbutazone studies in neonates 5–17 hours old demonstrated an increased volume of distribution, longer half-life and lower total clearance than adults.51 Other options include the new coxib class of NSAIDs meloxicam R IV and oral suspension) and firocoxib (Metacam R (Equioxx oral suspension only) but these drugs have not been studied in neonates or foals. Hydration status, renal function, and stress levels are important to consider when administering an NSAIDs to foals. NSAID administration often results in ulceration of the glandular portion of the stomach. Gastric ulceration and/or rupture can occur within a few days of initiating treatment.52 Therefore, concurrent administration of an H2 antagonist such as cimetadine or the proton pump inhibitor omeprazole is recommended and adequate hydration is imperative. Abdominal pain There are several options available for treatment of abdominal pain in the neonate and foal including NSAIDs, alpha-2 agonists, opioids, and the combination of an NSAID and an anticholinergic, N-butylscopolammonium bromide (BuscopanTM ). Flunixin meglumine is an effective NSAID for abdominal pain and for minimizing the hemodynamic effects associated with endotoxin release,53 although some clinicians favor ketoprofen. Caution should be used when administering an NSAID to a fluiddepleted foal due to the increased risk of altered renal perfusion. Buscopan is indicated for the treatment of spasmodic and gas colic as well as mild impaction colic. Buscopan has not been studied in foals but clinical evidence has shown that 2 mL i.v. given for meconium impactions can aid in relieving discomfort from spasm. The recommended dose is 1.5 mL/100 kg i.v. The kappa opioid agonist butorphanol is recommended for treatment of gastrointestinal pain. There are a large concentration of kappa receptors in the abdominal viscera; therefore doses of 2–5 mg i.v. butorphanol are effective in relieving discomfort of mild abdominal pain in foals. Alpha-2 adrenergic agonists are also effective in treating abdominal pain and the sedation associated with the analgesia might be beneficial while transporting a foal to a surgical facility. Lidocaine Lidocaine is a very inexpensive drug that has many uses in equine medicine. Constant rate infusions can
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be used for abdominal pain or any pain state, during general anesthesia to minimize minimum alveolar concentration of anesthetics, and for standing procedures that might require desensitization of an airway. Boluses of lidocaine can be used under general anesthesia in neonates and foals to extend the duration of anesthesia. Typically, 1–3 mL of 2% lidocaine given as a bolus to a neonate or foal result in an additional 2–5 minutes of anesthesia if given during or at the end of a procedure when a small amount of additional relaxation is needed. Lidocaine does not replace the administration of ketamine when a foal is aroused during a surgical procedure but it does act as a mild analgesic that does not appear to affect the quality of recovery.
Alternative analgesics Tramadol is a centrally acting analgesic drug that has mild effect at mu opioid receptors. Although not a commonly used drug and more expensive than many other analgesics, there is potential for this drug in foals. Gabapentin is an anti-epilepsy medication that acts at voltage gated N-type calcium ion channels in the central nervous system. The drug has been used extensively in human medicine for neuropathic pain and is developing popularity in small animal medicine as well as equine. It is an inexpensive drug and has shown clinical use in chronic pain states in adults (laminitis, fracture, arthrodesis). Morphine, buprenorphine, and fentanyl are all mu opioid agonists that vary in duration and expense but are additional options for analgesia in the neonate and foal.
Summary Anesthesia and sedation in neonates and foals requires an understanding of the physiological and metabolic differences associated with immaturity. Minimizing injectable and inhalant anesthetics decreases the potential for build-up of drug in the body. By using drugs that are reversible or minimally metabolized, the anesthetist can avoid unwanted complications. Although there have been limited studies regarding analgesia in equine pediatrics it is imperative that proper peri-anesthetic analgesia be included in protocols.
References 1. Tranquilli WJ, Thurman JC. Management of anesthesia in the foal. Vet Clin N Am Equine Pract 1990;6:651–63. 2. Dunlop CL. Anesthesia and Sedation in Foals. Vet Clin N Am Equine Pract Perinatol 1994;10(1):67–85.
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3. Vaala WE. Aspects of pharmacology in the neonatal foal. Vet Clin N Am Equine Pract 1985;1:51–75. 4. Gossett KA, French DD. Effect of age on liver enzyme activity in serum of healthy Quarter Horses. Am J Vet Res; 1984;45:354–6. 5. Normal WM, Court MH, Greenblatt DJ. Age-related changes in the pharmacokinetic disposition of diazepam in foals. Am J Ves Res 1997;58:878–80. 6. Thomas WP, Madigan JE, Backus KQ, Powell WE. Systemic and pulmonary hemodynamics in normal neonatal foals. J Reprod Fertil 1987;35:623–8. 7. Bidwell LA, Bramlage LR, Rood WA. Equine perioperative fatalities associated with general anaesthesia at a private practice – a retrospective case series. Vet Anaesth Analg 2007;34:23–30. 8. Johnston GM, Eastment JK, Wood JLN, Taylor PM. The confidential enquiry into perioperative equine fatalities (CEPEF): mortality results of phases 1 and 2. Vet Anaesth Analg 2002;29:159–70. 9. Webb AI. Nasal intubation in foals. J Am Vet Med Assoc 1984;185:48–51. 10. Refsum HE. Relationship between state of consciousness and arterial hypoxemia and hypercapnia in patients with pulmonary insufficiency, breathing air. Clin Sci 1963;25:361–7. 11. Robertson SA, Carter SW, Donovan M, Steele CJ. Effects of intravenous xylazine hydrochloride on blood glucose, plasma insulin and rectal temperature in neonatal foals. Equine Vet J 1990;22:43–7. 12. Kerr CL, Bouri LP, Pearce SG, McDonnell WN. Cardiopulmonary effects of diazepam-ketamine-isoflurane or xylazine-ketamine-isoflurane during abdominal surgery in foals. Am J Vet Res 2009;70:574–80. 13. Carter SW, Robertson SA, Steel CJ, Jourdinais DA. Cardiopulmonary effects of xylazine sedation in foals. Equine Vet J 1990;22:384–8. 14. Oijala M, Katila T. Detomidine (Dormosedan) in foals: sedative and analgesic effects. Equine Vet J 1988;20:327–30. 15. Arguedas MG, Hine MT, Papich MG, Farnsworth KD, Sellon DC. Pharmacokinetics of butorphanol and evaluation of physiologic and behavioral effects after intravenous and intramuscular administration to neonatal foals. J Vet Intern Med 2008;22:1417–26. 16. Matthews NS, Hartsfield SM, Hague B, Carroll GI, Short CE. Detomidine-propofol anesthesia for abdominal surgery in horses. Vet Surg 1999;28:196–201. 17. Valverde A, Gunkelt C, Doherty TJ, Giguere S, Pollak AS. Effect of a constant rate infusion of lidocaine on the quality of recovery from sevoflurane or isoflurane general anesthesia in horses. Equine Vet J 2005;37:559–64. 18. Muir WW, Sams R. Effects of ketamine infusion on halothane minimal alveolar concentration in horses. Am J Vet Res 1992;53:1802–6. 19. Yamashita K, Wijayathilaka TP, Kushiro T, Umar MA, Taguchi K, Muir WW. Anesthetic and cardiopulmonary effects of total intravenous anesthesia using a midazolam, ketamine and medetomidine drug combination in horses. J Vet Med Sci 2007;69:7–13. 20. Kushiro T, Ymashita K, Umar MA, Maehara S, Wakaiki S, Abe R, Seno T, Tsuzuki K, Izumisawa Y, Muir WW. Anesthetic and cardiovascular effects of balanced anesthesia using constant rate infusion of midazolamdetamine-medetomidine with inhalation of oxygensevoflurane (MKM-OS anesthesia) in horses. J Vet Med Sci 2005;67:379–84.
21. Friedman WF, George BL. Treatment of congestive heart failure by altering loading conditions of the heart. J Pediatr 1985;106:697–706. 22. Perkins G, Valberg SJ, Madigan JM, Carlson GP, Jones SL. Electrolyte disturbances in foals with severe rhabdomyolysis. J Vet Intern Med 1998;12:173–7. 23. Lakritz J, Madigan J, Carlson GP. Hypovolemic hyponatremia and signs of neurologic disease associated with diarrhea in a foal. J Am Med Assoc 1992;200:1114–16. 24. Behr MJ, Hackett RP, Bentinck-Smith J, Hillman RB, King JM, Tennant BC. Metabolic abnormalities associated with rupture of the urinary bladder in neonatal foals. J Am Vet Med Assoc 1981;178:263–6. 25. Dunlop CS, Hodgson DS, Grandy JL, Chapman PL, Heath RB. The MAC of isoflurane in foals. Vet Surg 1989;18:247–54. 26. Steffey EP, Howland DJ, Giri S, Eger EI. Enflurane, halothane and isoflurane potency in horses. Am J Vet Res 1977;38:1037–9. 27. Aida H, Mizuno Y, Hobo S, Yoshida K., Fujinaga T. Determination of the minimum alveolar concentration (MAC) and physical response to sevoflurane inhalation in horses. J Vet Med Sci 1994;56:1161–5. 28. Tendillo FJ, Mascias A, Santos M, Lopez-Sanroman J, De Rossi R, Sanroman F, Gomez de Segura IA. Anesthetic potency of desflurane in the horse: Determination of the minimum alveolar concentration. Vet Surg 1997;26:354–7. 29. Dixon PM. Respiratory mucociliary clearance in the horse in health and disease, and its pharmaceutical modification. Vet Rec 1992;131:229–35. 30. West JB. Environmental and other diseases. In: West JB. Pulmonary Pathophysiology, The Essentials, 6th edn. Baltimore: Lippincott Williams & Wilkins, 2003; pp. 123–41. 31. Valverde, A, Giguere S, Sanchez LC, Shih A, Ryan C. Effect of dobutamine, norepinephrine and vasopressin on cardiovascular function in anesthetized neonatal foals with induced hypotension. Am J Vet Res 2006;67:1730–7. 32. Craig CA, Haskins SC, Hildebrand SV. The cardiopulmonary effect of dobutamine and norepinephrine in isoflurane anesthetized foals. Vet Anesth Analg 2007;34:377–87. 33. Manning M., Dubielzig R., McGuirk S. Postoperative myostitis in a neonatal foal: a case report. Vet Surg 1995;24:69–72. 34. Rossdale PD. Some parameters of respiratory function in normal and abnormal newborn foals with specific reference to levels of Pa o2 during air and oxygen inhalation. Res Vet Sci 1970;11:270–6. 35. Matsui K, Sugano S, Masuyama I, Amada A, Kano Y. Alteration in the heart rate of Thoroughbred horse, pony and Holstein cow through pre- and post-natal stages. Jpn J Vet Sci 1984;46:505–10. 36. Taddio A, Katz J, Illersich AL, Koren G. Effect of neonatal circumcision on pain response during subsequent routine vaccination. Lancet 1997;349:599–603. 37. Fischer BL, Ludders JW, Asakawa M, Fortier LA, Fubini SL, Nixon AJ, Radcliffe RM, Erb HN. A comparison of epidural buprenorphine plus detomidine with morphine plus detomidine in horses undergoing bilateral stifle arthroscopy. Vet Anaesth Analg 2009;36:67–76. 38. Eberspacher E, Stanley SD, Rezende M, Steffey EP. Pharmacokinetics and tolerance of transdermal fentanyl administration in foals. Vet Anaesth Analg 2008;35:249–55. 39. Bennett RC, Steffey EP. Use of opioids for pain and anesthetic management in horses. Vet Clin N Am Equine Pract 2002;18:47–60. 40. Natalini CC, Robinson EP. Evaluation of the analgesic effects of epidurally administered morphine, alfentanil,
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butorphanol, tramadol, and U50488H in horses. Am J Vet Res 2000;61:1579–86. Davis JL, Posner LP, Elce Y. Gabapentin for the treatment of neuropathic pain in a pregnant horse. J Am Vet Med Assoc 2007;231:755–8. Skarda RT, Muir WW. Comparison of antinociceptive, cardiovascular, and respiratory effects, head ptosis, and position of pelvic limbs in mares after caudal epidural administration of xylazine and detomidine hydrochloride solution. Am J Vet Res 1996;57:1338–48. Nixon AJ. Perioperative considerations. In: Nixon AJ (ed.) Equine Fracture Repair. Philadelphia: Saunders, 1996; pp. 43–51. Santos LC, de Moraes AN, Saito ME. Effects of intraarticular ropivacaine and morphine on lipopolysaccharideinduced synovitis in horses. Vet Anaesth Analg 2009;36:280–6. Thomasy SM, Slovis N, Maxell NK, Kollias-Baker C. Transdermal fentanyl combined with nonsteroidal antiinflammatory drugs for analgesia in horses. J Vet Intern Med 2004;18:550–4. Orsini JA, Moate PJ, Kuersten K, Soma LF, Boston RC. Pharmacokinetics of fentanyl delivered transdermally in healthy adult horses – variability among horses and its clinical implication. J Vet Pharmacol Ther 2006;29:539–46. Wegner K, Franklin RP, Long MT, Robertson SA. How to use fentanyl transdermal patches for analgesia in horses.
48.
49.
50.
51.
52.
53.
54.
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In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, Orlando, FL, 2002, pp. 291–4. Bidwell LA, Wilson DV, Caron JP. Lack of systemic absorption of lidocaine from 5% patches placed on horses. Vet Anaesth Analg 2007;34:443–6. Crisman MV, Wilcke JR, Sams RA. Pharmacokinetics of flunixin meglumine in healthy foals less than twenty-four hours old. Am J Vet Res 1996;57:1759–61. Wilcke JR, Crisman MV, Scarratt WK, Sams RA. Pharmacokinetics of ketoprofen in healthy foals less than twenty-four hours old. Am J Ves Res 1998;59: 290–2. Wilcke JR, Crisman MV, Sams RA, Gerken DF. Pharmacokinetics of phenylbutazone in neonatal foals. Am J Vet Res 1992;54:2064–7. Traub-Dargatz JL, Bertone JJ, Gould DH, Wrigley RH, Weiser MG, Forney SD. Chronic flunixin meglumine therapy in foals. Am J Vet Res 1988;49:7–12. Baskett A, Barton MH, Norton N, Anders B, Moore JN. Effect of pentoxifylline, flunixin meglumine, and their combination on a model of endotoxemia in horses. Am J Vet Res 1997;58:1291–9. Messenger KM et al. The Pharmacokinetics of intravenous and sublingual buprenorphine in horses. From the Proceedings of the Fifteenth International Veterinary Emergency and Critical Care Symposium in Chicago, IL.
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Chapter 24
Stabilization and Preparation for Transport Robert P. Franklin and Jennifer R. Read
The compromised neonatal patient requires some basic provisions prior to or during transportation to an intensive care unit. Often a significant amount of time will elapse during this period. A comprehensive approach to stabilizing the patient may positively affect the outcome in many cases.
Fluids The administration of intravenous fluids in the field prior to and during transport can be life-saving, as valuable time when volume replacement is essential can be lost during this period, especially if transportation is delayed or there is a long trip to a referral facility. The basics of initial fluid resuscitation are covered in this chapter. Refer to Chapter 000 for more detail. An intravenous catheter should be placed in the jugular vein and should be long enough (ideally at least 4 inches or longer) and well secured so that the risk of dislodgement during transport is minimized. If the foal is struggling during catheter placement it can be sedated with butorphanol (0.05 mg/kg intravenously (i.v.)) or diazepam (0.1–0.2 mg/kg i.v.). The catheter should be well secured by using suture and may be bandaged based on clinician preference. In the time period between initial assessment at the farm and arrival at a referral facility, calculation of fluid deficits often needs to be done in the absence of laboratory data, unless a handheld lactate analyzer is available. Dysoxic tissues undergoing anaerobic metabolism produce lactate. Dysoxia can be caused by hypoxemia or poor oxygen delivery due to hypovolemia. A lactate of greater than 2.5 mmol/L together with any other clinical sign of hypovolemia is indicative of the need for volume replacement.1 Neonatal foals will not always show the classic clinical signs of
hypovolemia as an adult horse would, such as tachycardia, prolonged capillary refill time, weak pulses, cool extremities, and poor jugular filling. Heart rate may be normal, bradycardic, or tachycardic. Therefore, the foal’s nursing history and potential fluid losses such as diarrhea become very important. Any neonatal foal that has not nursed for the previous 4 hours or is showing any one of the clinical signs of a hypovolemic adult should be suspected of having some degree (at least 6%) of dehydration.2 Moderately (8%) to severely (10%) dehydrated foals are more easily identified and will usually have sunken eyes, depression, or collapse. Foals immediately postpartum (e.g., postdystocia) or those that are well enough to still be nursing and do not have ongoing fluid losses (e.g., neonatal isoerythrolysis) generally are not hypovolemic and do not require massive amounts of intravenous fluid therapy. Fluid replacement is not typically used in resuscitation of human neonates unless they are hypotensive and the hypotension is unresponsive to cardiopulmonary resuscitation.3 However, compromised foals that are unable to nurse during long transportation or those that may be of questionable hydration status will benefit from a moderate fluid bolus (20 mL/kg). These foals typically do not require ongoing replacement or maintenance fluid therapy during transit. Once the need for fluid intervention has been determined, a replacement volume is calculated. Bodyweight in kilograms is multiplied by the percentage of dehydration to give a volume deficit in liters. This calculation can be performed using rough estimations in the field and should not be overscrutinized. Half of the deficit is replaced quickly with a bolus and the remainder can be given during transport and in the following 12–24 hours.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Stabilization and Preparation for Transport A polyionic replacement fluid is the most appropriate fluid choice in the absence of blood electrolyte values. Lactated Ringer’s solution (LRS), Hartmann’s solution, Plasma-Lyte 148, and Normosol-R are the most readily available polyionic replacement fluids; 0.9% sodium chloride is also a readily available replacement fluid. These fluids are safe to be given as a bolus. Foals with suspected electrolyte disturbances such as hyponatremia (e.g., diarrhea, uroperitoneum) or hyperkalemia (e.g., uroperitoneum) may benefit more from a potassium-free, sodium-rich fluid such as 0.9% sodium chloride. Plain 5% dextrose should not be given as a resuscitative fluid, as the glucose is rapidly metabolized leaving the foal with a large hypotonic free water bolus, which poorly restores circulating volume. As previously mentioned, a 20-mL/kg bolus will be of benefit to any dehydrated foal, and, if dehydration is mild and transport time is short, may be all that is required to stabilize the foal sufficiently until more fluids can be administered at the referral facility. The on-farm clinician can always reassess the patient’s hydration status after a resuscitation bolus is given to determine whether more fluids are indicated. Ongoing or maintenance fluid rates should also be calculated for transport. The maintenance rate for foals is 10% of body weight per day plus residual deficits. Therefore a 50-kg foal will require 5 liters of fluids per day (50 kg × 10% = 5 liters) plus the residual from the deficit calculation. If a residual is present the foal can simply go on a twice maintenance rate, or one could calculate the residual and add it to the maintenance rate. As a fluid pump is likely to be unavailable in the field or during transport, the drip rate should be estimated for ongoing fluid therapy after administration of a bolus. Administration sets display the number of drops/mL on the packaging, usually 10 or 20 drops/mL. To calculate a rate in drops/10 s, divide the mL/h rate by 360 (the number of 10-second intervals in 1 hour) then multiply by the number of drops/mL. For example, to calculate the rate in drops/10 s for a 50-kg foal with a maintenance requirement of 208 mL/h, using a 20 drops/mL i.v. administration set: 208/360 = 0.58 mL/10 s 0.58 × 20 = 11.6 drops/10 s
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If the rate is to be twice maintenance, the drops/s rate will be 11.6 × 2 = 23.2 drops/10 s. Blood glucose measurements are also of value, as knowing whether a foal is hypoglycemic can mean more accurate administration of glucose in fluids. Usually, a value of less than 80 mg/dL (4.4 mmol/L) will indicate glucose supplementation is required.2 A 5–10% glucose concentration in a polyionic isotonic solution is most commonly used. Glucose administered at a rate of 4–8 mg/kg/min is going to be of benefit to most critical neonates;4, 5 see Table 24.1 for concentrations and rates that will achieve this. Again, 5% dextrose in water should not be used as a resuscitation fluid because of the large amount of free water that will be left once the dextrose is absorbed. Based on these concentrations, the 5% concentration should be used for foals requiring a higher fluid rate, and the 10% concentration can be used for those requiring closer to maintenance rates. Foals with neonatal isoerythrolysis (NI) that are showing signs of severe anemia may require a blood transfusion prior to a long or delayed transport period. In the field, a blood lactate analyzer will be of greatest assistance in determining whether a transfusion is required, as an elevated lactate (>2.2 mmol/L) will indicate insufficient oxygen delivery to tissues.6 Oxyglobin (30 mL/kg i.v. bolus) may be administered as an emergency treatment for hemolytic or blood loss anemia. Alternatively, a simplified method of delivering red cells can be performed on the farm if the foal’s condition is so severe that it is unlikely to survive transport, or if transport is delayed or prolonged, as prompt delivery of red blood cells can prevent life-threatening cerebral hypoxia. Whole blood is collected from the mare in sterile bottles, blood collection bags, or empty 1-liter fluid bags, with 125 mL sodium citrate or 5,000 IU heparin added per liter of blood collected. The cells are allowed to settle for approximately 1 hour, after which the red cells are transferred by gravity into an empty fluid bag. A minimum 1 cm buffer should be left between the plasma and red cells. Two suspensions may be optimal in removing alloantibodies and should be performed if time allows. The maximum dose for whole blood that avoids hypertension in the majority of foals is 40 mL/kg, which is usually sufficient to provide
Table 24.1 Compounding and administration protocol for dextrose-containing infusions. Reproduced from Wilkins PA, 2003, ‘Disorders of Foals’ in Reed SM, Bayly WM & Sellon DC, Equine Internal Medicine p. 1389. Courtesy of Elsevier. Amount of 50% dextrose added to 1-liter bag fluids
Final concentration of dextrose in bag
Approximate rate to administer
Approximate rate to administer to average 50-kg foal
100 mL 200 mL
5% 10%
5–10 mL/kg/h 2.5–5 mL/kg/h
250–500 mL/h 125–250 mL/h
Adapted from Wilkins.5
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adequate red blood cells.7 Packed red blood cells may be resuspended in 0.9% saline at a ratio of 1:1, avoiding the use of calcium-containing fluids, to ease flow through administrations sets and catheters. A specific blood product administration set should be used.
Antibiotics Antibiotic therapy is of extreme importance in many neonatal disorders, especially to treat sepsis or prevent it from developing in high-risk foals. Potent antibiotic therapy should be started as soon as possible in any ill or high-risk foal, as it is likely that valuable ground is lost in the time taken to administer the first dose of antibiotics.8 Foals that are weak, recumbent, have not nursed, have failure of passive transfer, are hypothermic, hypovolemic, endotoxemic, premature, or where placentitis was present during gestation generally require antibiotics. If in doubt, antibiotics should be given. If blood culture bottles are available, a blood culture should be obtained prior to antibiotic administration. If no suitable medium for blood culture collection is available, antibiotic therapy should not be delayed for the sake of blood collection for culture. Broad-spectrum antibiotics should be used in the initial treatment, with special consideration given to ensuring Gram-negative organisms are covered.8–10 Intravenous antibiotics are preferable and an intravenous catheter should be placed for their administration if it is likely that more than one dose will need to be given prior to arrival at the referral facility. Consideration must be made for renal compromise when selecting antibiotics. A fundamental selection of antibiotics for foals that have normal renal function (no signs of hypovolemia or ischemic/hypoxic damage to the kidneys) includes the combination of potassium penicillin (44 000 IU/kg i.v. q. 6 hours), procaine penicillin (22 000 IU/kg intramuscularly (i.m.) q. 12 hours) or ampicillin (22 mg/kg i.v. q. 6 hours) with amikacin (25 mg/kg i.v. q. 24 hours).10 If renal damage is likely, a third-generation cephalosporin such as cefotaxime (40 mg/kg i.v. q. 6 hours) can be selected as a broad-spectrum antibiotic with minimal renal toxicity. Ceftiofur (5–10 mg/kg i.v. q. 6–12 hours) is another third-generation cephalosporin that can be given as the pharmacokinetic data have recently become available for the intravenous route. In cases where intestinal clostridiosis is suspected, metronidazole (15 mg/kg per os (p.o.) q. 12 hours) should be given as ileus is often noted and precludes the absorption of oral medication. Antibiotic therapy should be continued at the required dosing interval during transportation if possible.
Neuroprotectants Foals that are suspected of having neurological disorders, such as neonatal encephalopathy (NE), can be started on neuroprotectant drugs before transport to a referral facility. It is important to note that none of these recommended protocols or agents have been subject to clinical trials in neonatal foals. They comprise vitamins (most importantly thiamine, vitamin E, and vitamin C), magnesium, flunixin meglumine, dimethylsulfoxide (DMSO), and mannitol. Thiamine (1–5 mg/kg q. 24 hours i.v., i.m., subcutaneously (s.c.), or p.o., or 10 mg/kg in i.v. fluids q. 24 hours) can be given as a bolus dose or added to the i.v. fluids after initial resuscitation. Thiamine plays a role in many essential central nervous system pathways, including the breakdown of lactate, and is also essential in energy pathways in the brain.5 Vitamin C (10–30 mg/kg in i.v. fluids q. 24 hours) is believed to be a neuroprotectant owing to its reactive oxygen species (ROS)-scavenging abilities. Vitamin E (20 IU/kg s.c. for first dose then 20 IU/kg p.o. q. 24 hours; ‘Vital E 300’, Schering-Plough Animal Health, d-alpha-tocopherol 300 IU/mL) is also given as an ROS scavenger. Magnesium (50 mg/kg/h constant rate infusion (CRI) i.v. over the first hour and then 25 mg/kg/h CRI i.v. for the next 23 hours) acts as an N-methyld-aspartate (NMDA) receptor antagonist and therefore may help prevent brain injury by reducing influx of excitatory electrolytes, specifically calcium, into brain cells. This may reduce the incidence of seizures caused by NE.5 This protocol has also not been subject to clinical trials in foals but has been evaluated extensively in laboratory rats and human infants.11–14 Flunixin (1.1 mg/kg i.v. q. 12 hours) may be given to neonatal foals prior to transport as an antiinflammatory agent for the brain. Again the benefits of this drug in NE have not been studied. Care must be taken if renal compromise is suspected as can occur secondary to hypoxic injury to the kidneys. DMSO (0.25–1.0 g/kg i.v. q. 12–24 hours maximal concentration of 10%) is a ROS scavenger and is widely and empirically applied to a host of medical conditions; however, its use remains controversial in the treatment of foals with NE. Mannitol (0.5–1.0 g/kg q. 12 hours) is usually reserved for those foals with severe cerebral edema. It is an osmotic diuretic and therefore its main purpose is to decrease excess fluid in cerebral cells. It also has some scavenging action against ROS. Mannitol should be diluted to a 20% solution and given i.v. over 30 minutes. Hypothermic body temperatures have recently been determined in human medicine to be
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Stabilization and Preparation for Transport neuroprotective when hypoxic damage has occurred to the brain.15, 16 The best course of action remains controversial;17 however, it may be recommended that neonatal foals with NE are not actively warmed.7 This is acceptable as long as the foal is not shivering, as this process increases oxygen consumption.
Respiratory support Respiratory support may be essential in the form of resuscitation, stimulation, or oxygen supplementation. If the foal is in respiratory arrest and fails to respond to physical stimulation (rubbing with towels), the airway should be drained of fluid by postural drainage and a nasotracheal tube passed, and breaths given with an Ambu bag at 10 breaths/min.7 The chest should be observed with each breath to ensure the appropriate tidal volume is being administered. For a 50-kg foal, a complete compression of a human-sized (1 liter) Ambu bag is usually appropriate. Oxygen is not essential but can be used if available; however, recent human neonate studies suggest room air may be preferable to 100% oxygen for resuscitation.18–20 Breaths can be given via mouth-totube or mouth-to-nose if an endotracheal (ET) tube or Ambu bag is not available, or if an ET tube cannot be passed easily. Constant monitoring is essential during resuscitation so that changes can be made accordingly. Ventilation should be continued if necessary during transport. Independent supplemental oxygen may be required once the foal is breathing on its own. Foals that are premature, have suffered a hypoxic insult, are septic, or are in respiratory distress owing to primary respiratory disease are also likely to benefit from oxygen therapy. Anemic foals may benefit from oxygen therapy, especially when ventilation is also compromised; however, these foals primarily require red blood cells in order to improve oxygenation. Foals that are tachypneic or in obvious respiratory distress, have injected or cyanotic mucous membranes, or have a history of a hypoxic episode should be administered supplemental oxygen. If arterial oxygen saturation or blood lactate can be measured, these can also be valuable indicators of oxygen deprivation. Arterial oxygen saturation levels (Sp o2 ) of less than 90% and/or high blood lactate (> 2.2 mmol/L) concentrations are strong indicators for the need for supplemental oxygen. If a portable oxygen cylinder is available, an indwelling nasal oxygen catheter can be secured and humidified oxygen delivered at 5–8 L/min. An indwelling nasal oxygen tube can be placed using any flexible plastic tubing; however, intranasal oxygen catheters (18 Fr Suction Catheter Without Control Valve, Covi-
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dien, Massachusetts, USA) are preferable. If not available, a shortened feeding tube or an oxygen mask (McCulloch Medical, O2 Recovery Mask) can be used. Nasal catheters should be inserted into the ventral nasal meatus to the level of the medial canthus. The catheter can be sutured in place using a Chinese finger trap, and finally secured with adhesive bandage around the muzzle, taking care to ensure the bandage is not too tight and does not cover the lips. Oxygen extension tubing can then be attached from the humidifier to the nasal catheter. Efforts should be made to keep the foal positioned in sternal recumbency as this will prevent atelectasis of the dependent lung and therefore maximize gas exchange. This maneuver alone can increase partial pressure of oxygen in arterial blood (Pa o2 ) by 10–20 mmHg.21 Foals with neurological disease often have central respiratory depression, resulting in hypoventilation and hypercapnea with subsequent respiratory acidosis and hypoxemia. Drugs that stimulate respiration can be used in an attempt to improve ventilation. The safest drug available is caffeine (loading dose of 10 mg/kg p.o., and then at 2.5–3.0 mg/kg p.o. q. 24 hours). Caffeine is readily available and is given either orally or via a nasogastric feeding tube. If the foal is unable to tolerate enteral administration, caffeine can also be given per rectum at the same dose.21 The use of doxapram has not been recommended as a respiratory stimulant during resuscitation owing to its effects of reducing cerebral blood flow and increasing myocardial oxygen demands;5, 21, 22 however, it has been found to be significantly more effective than caffeine in reducing partial pressure of carbon dioxide in arterial blood (Pa co2 ) in hypoventilatory foals.23, 24
Warmth Active warming of a hypothermic patient should not be commenced until fluid deficits have been corrected, as vasodilation of the periphery will further decrease arterial pressure. As mentioned earlier, it may be preferable not to actively warm a foal with hypoxic brain injury, as hypothermia has the potential to be neuroprotective. Where hypothermia is present with normal hydration status, efforts to warm should include wrapping the foal in blankets, moving the foal to a more sheltered location if possible, wrapping the distal limbs, and using heating devices such as heat lamps, warmed fluid bags, and forced air patient-warming systems. Care should be taken to ensure that thermal burns from a direct radiant heating source, such as a heating blanket, are avoided. If the foal is normothermic before transport, efforts should be made to ensure that it does not become hypothermic
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during transport. Many of the above measures can be instituted on the trailer. Intravenous fluids can be warmed when possible. Ensuring the foal has nutritional support and normal blood glucose levels will aid in its own ability to thermoregulate.25, 26 Whether this is best achieved through enteral feeding or via i.v. fluid therapy will depend on the ability of the foal to cope with enteral feeding.
Seizure control Seizure episodes are common in foals with NE and hypoglycemia and should be controlled immediately. Seizure activity can range from small abnormal head and jaw movements to grand mal seizures. Seizures will increase oxygen consumption of skeletal muscle, increase cardiac oxygen demand, and also increase energy consumption in the brain that can lead to further seizure activity.5, 22, 27 Seizures can be controlled initially with diazepam (0.1–0.2 mg/kg i.v. pro re nata (p.r.n.) for two or three doses). If seizures are refractory to this, phenobarbital (5–15 mg/kg administered slowly i.v. as a loading dose) may be given. Phenobarbital can be continued at the hospital if required. Midazolam given as a constant rate infusion (0.1–0.2 mg/kg/h) is the next choice if phenobarbital is ineffective. A foal that is seizuring should be physically protected from causing traumatic injury to itself. Therefore, wrapping legs, supporting the foal on padded mats or deep bedding, and having someone restrain the foal until medications can be administered are important.
Pain control Analgesia is required for foals that are colicky, have septic arthritis, or have suffered a traumatic injury. Butorphanol (0.01–0.05 mg/kg i.v.) is the opioid of choice for neonatal foals. It has analgesic as well as sedative effects that can also be beneficial in preventing excessive motion and thrashing. Butorphanol has also been shown to increase hunger in neonatal foals. 28 Flunixin (1.1 mg/kg i.v. q. 12 hours) can be used cautiously in neonates with normal renal function. Care must be taken where renal or gastrointestinal compromise is likely to be present, such as in hypovolemic or hypoxemic patients.29 Fentanyl patches (10.2 mg, 100 µg/h patch for a 50-kg foal) have anecdotally been useful in treating post-operative pain in foals; however, in the immediate period prior to referral, intravenous analgesia is likely to have a faster onset of action and therefore greater efficacy.30 Prevention of further traumatic injury from thrashing movements, incoordination, and failed at-
tempts to stand is important before and during transportation. The foal should be supported on deep bedding, legs wrapped where possible, and should have someone restraining it. This is important during transport, as considerable injury can occur. Butorphanol (0.05 mg/kg i.v.) or diazepam (0.1 mg/kg i.v.) can be given as a sedative in fractious neonates.
Endotoxemia or failure of passive transfer Foals that have not nursed by the time they are seen at the farm will be at risk of failure of passive transfer (FPT). FPT is a factor in increasing the risk of sepsis, which can lead to endotoxemia, systemic inflammatory response syndrome (SIRS), multiple organ dysfunction syndrome (MODS), and death.5, 31, 32 Endotoxemia is important in neonatal foal sepsis, as the majority of organisms are Gram negative. Foals that are endotoxemic and septic may show a wide range of clinical signs but are typically depressed, have lost interest in nursing, and may be recumbent. Mucous membranes may be bright or injected with petechial hemorrhages and have a decreased capillary refill time; fever may or may not be present.5, 8 If the foal is able to tolerate enteral feeding, it should be fed colostrum stripped from the mare (provided the foal does not have NI) either by bottle if it has a good suckle reflex or via nasogastric tube. Ideally, a 50-kg foal will receive at least 1 liter of highquality colostrum (Brix > 23% or specific gravity >1.060) divided into feedings appropriate for the foal’s condition. The volume that is safe to give will depend on the foal’s ability to tolerate enteral feeding and may range from 0 to 500 mL per feeding in a newborn. If the foal is severely hypothermic, hypovolemic, hypoxemic, comatose, or semicomatose, or has abdominal distension with or without diarrhea, enteral feeding should be withheld. These foals are administered plasma and i.v. nutritional support via an intravenous catheter, which can be given in the field prior to transport. One liter (or 20 mL/kg) of plasma may be given intravenously as a bolus over 10–30 minutes, taking care to monitor for plasma reactions, to address passive transfer concerns, and to play a part of a resuscitative fluid plan. If rapid volume resuscitation is not indicated, the plasma transfusion can be given at a slower rate of 1 L over 1–2 hours. Giving a larger volume of colloid may create fluid overloading, which can progress to pulmonary edema. In the authors’ opinion the safe dose for blood products is ≤ 40 mL/kg/day to prevent volume overload. Foals with sepsis and endotoxemia may be hyperglycemic or hypoglycemic; therefore, a hand-held glucometer
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Stabilization and Preparation for Transport should be used to measure blood glucose levels. In cases of hypoglycemia, dextrose should be added to isotonic replacement fluids to make up a 5% solution (100 mL of 50% dextrose in 1 liter of replacement fluid) to be administered as a drip during transport. Plain 5% dextrose should not be used for the reasons already mentioned. Hyperglycemia should not be treated with insulin in the field as this may lead to severe hypoglycemia during transport. Antibiotics are likely the single most important treatment of sepsis. The antibiotic or combination of antibiotics chosen should be broad spectrum, especially encompassing Gram-negative organisms, as these are the most common involved in sepsis. If renal compromise is suspected, a third-generation cephalosporin should be used. See the section on antibiotics for more detail. The use of anti-endotoxic drugs may also be instituted before transport. The use of flunixin for endotoxemia in foals has not been studied but may be of benefit. Owing to likely concurrent inflammation, the anti-inflammatory dose can be used (1.1 mg/kg i.v. q. 12 hours). Care must be taken with the use of this drug if severe renal or gastrointestinal compromise is suspected.
Nutrition Some foals will be able to tolerate some enteral nutrition, and this can be commenced prior to and during transport to the hospital. Foals that are immediately postpartum and able to tolerate enteral feeding, but cannot nurse, will certainly benefit from administration via nasogastric tube of colostrum stripped from the mare, as glycogen stores in the foal provide energy for only approximately the first hour of life.25 This initial high-energy meal can be of great benefit to the foal, as well as beginning the process of passive transfer of antibodies from the mare to the foal. Foals that are safe to feed are usually those that have not suffered a hypoxic insult where the gastrointestinal tract has been compromised. Therefore, any foal that is alert and able to stand and walk is likely to be able to tolerate feeding. Foals that are recumbent, depressed, hypothermic, unresponsive, have abdominal distension, are colicky, or have severe diarrhea should not be fed enterally. Some recumbent foals will be able to tolerate enteral feeding; however, caution should be used with the amount given and foals should always be fed in sternal recumbency. Starting at 100 mL/kg/24 h (10% of bodyweight per day) divided into 12 feedings given every 2 hours is a conservative start. The foal should be monitored for signs of colic and abdominal distension, and refluxed if these occur. Feeding should then be stopped.
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If the foal is able to nurse from the mare it should be allowed to continue to do so, with the exception of foals with NI in the first 36 hours of life, and foals with severe diarrhea. If the foal cannot nurse but has a good suckle reflex, bottle-feeding can be attempted. Milk stripped from the mare is best; however, if this is not available, milk from another mare or a specific mare milk formula can be used. A goat teat should be used to allow best nursing from the bottle. For foals that are unable to suckle, bottle-feeding should not be attempted. Syringe-feeding should never be attempted owing to the high risk of aspiration pneumonia. If available, a nasogastric feeding tube can be placed and milk administered via this directly into the stomach. The tube should be passed through the ventral nasal meatus, with care taken to ensure the guidewire does not protrude from any of the openings in the end of the tube. The foal’s head should be flexed when the tube is at the level of the pharynx to maximize the likelihood of passing the tube into the esophagus. The tube should be able to be palpated dorsal to the trachea just behind the pharynx, and this should be confirmed before advancing the tube completely into the stomach and administering milk or colostrum. The main risks of commencing enteral feeding are intolerance to feeding due to ileus, and aspiration pneumonia. The foal should be monitored for abdominal distension. If the foal becomes colicky it should be refluxed; if a nasogastric tube is not already in place, a stallion urinary catheter passed into the stomach and a dose syringe are effective in removing stomach contents. If enteral feeding cannot be administered, dextrose can be administered in fluids once any necessary fluid bolus has been given. The aim is not to provide parenteral nutrition but rather to prevent or treat hypoglycemia in the time taken to arrive at the referral hospital. A blood glucose level can be obtained using a hand-held glucometer to determine whether the foal is hypoglycemic. Foals that are stressed may be hyperglycemic and this needs to be taken into consideration; these foals may still benefit from glucose supplementation as the hyperglycemia is not persistent. Any foal that has not nursed or has had an extended period of not nursing will likely be hypoglycemic. Dextrose can be administered as a 5–10% solution in an isotonic fluid at a rate based on fluid deficits (Table 24.1), but such concentrations should not be given by bolus infusion and therefore should be given concurrently or after any required fluid bolus has been given. Severe hypoglycemia is best treated by a slow infusion of 10% dextrose (200 mL of 50% dextrose in 1 liter of replacement fluids) over 30– 60 minutes.
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Transportation The transportation of the foal to the referral center will vary in each individual case depending on available means of transport, distance to travel, and urgency of arrival at the referral center. All possible scenarios cannot be anticipated and covered here; however, the main objective is to transport the foal and mare as quickly and safely as possible. To minimize trauma from the mare, the foal should preferably be separated from the mare but still within close proximity to her so that she can see the foal. Commercial horse vans may offer an attendant to accompany the foal. The authors cannot recommend farm assistants to ride in trailers as this poses many risks to said individuals, although this practice is common in many horse areas. The trailer should be deeply bedded and padded barriers added to the walls surrounding the foal where possible. Leg wraps on the foal may also assist in minimizing trauma to the legs. The eyes may also be lubricated. Any hyperactive or seizure activity should be controlled prior to transportation with butorphanol or diazepam sedatives, respectively. If a trailer is not available or adverse weather conditions prevent transport in a large vehicle, the foal can be transported in the back of a sport utility vehicle that is deeply bedded with blankets. The mare may be sedated and transported at a separate time. If this approach is taken, efforts must be made to minimize the time the mare is separated from the foal.
References 1. Corley KT, Donaldson LL, Furr MO. Arterial lactate concentration, hospital survival, sepsis and SIRS in critically ill neonatal foals. Equine Vet J 2005;37:53–9. 2. Corley KT, Axon JE. Resuscitation and emergency management for neonatal foals. In: Sanchez LC (ed.) Veterinary Clinics of North America Equine Practice Neonatal Medicine and Surgery. Philadelphia: Saunders Elsevier, 2005; pp. 431–55. 3. Wyckoff MH, Perlman JM, Laptook AR. Use of volume expansion during delivery room resuscitation in near-term and term infants. Pediatrics 2005;115:950–5. 4. Spurlock SL, Furr M. Fluid therapy. In: Koterba AM, Drummond WH, Koch PC (eds) Equine Clinical Neonatology. Lea & Febiger Philadelphia, 1990; pp. 671–700. 5. Wilkins PA. Disorders of foals. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine. St. Louis, MO: Saunders Elsevier, 2004; pp. 1381–431. 6. Magdesian KG, Fielding CL, Rhodes DM, Ruby RE. Changes in central venous pressure and blood lactate concentration in response to acute blood loss in horses. J Am Vet Med Assoc 2006;229:1458–62. 7. Franklin RP. Identification and treatment of the high-risk foal. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007, pp. 320–8. 8. Corley KT, Pearce G, Magdesian KG, Wilson WD. Bacteraemia in neonatal foals: clinicopathological differences be-
9.
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tween Gram-positive and Gram-negative infections, and single organism and mixed organism infections. Equine Vet J 2007;39:84–9. Sanchez LC. Equine neonatal sepsis. In: Sanchez LC (ed.) Veterinary Clinics of North America Equine Practice Neonatal Medicine and Surgery. Philadelphia: Saunders Elsevier, 2005; pp. 273–93. Henson S, Barton M. Bacterial isolates and antibiotic sensitivity patterns from septicemic neonatal foals: a 15 year retrospective study (1986–2000). In: Proceedings of the Dorothy Havemeyer Foundation Neonatal Septicemia Workshop 3, Talloires, France, 2001, pp. 350–2. Degos V, Loron G, Mantz J, Gressens P. Neuroprotective strategies for the neonatal brain. Anesth Analg 2008;106:1670–80. Spandou E, Soubasi V, Papoutsopoulou S, AugoustidesSavvopoulou P, Loizidis T, Pazaiti A, Karkavelas G, GuibaTziampiri O. Neuroprotective effect of long-term MgSO4 administration after cerebral hypoxia-ischemia in newborn rats is related to the severity of brain damage. Reprod Sci 2007;14:667–77. Ichiba H, Yokoi T, Tamai H, Ueda T, Kim TJ, Yamano T. Neurodevelopmental outcome of infants with birth asphyxia treated with magnesium sulphate. Pediatr Int 2006;48:70–5. Berger R, Garnier Y, Jensen A. Perinatal brain damage: underlying mechanisms and neuroprotective strategies. J Soc Gynecol Invest 2002;9:319–28. Jacobs S, Hunt R, Tarnow-Mordi W, Inder T, Davis P. Cooling for newborns with hypoxic ischaemic encephalopathy. Cochrane Database Syst Rev 2007;17(4):CD003311. Shankaran S, Laptook AR, Ehrenkranz RA. Whole body hypothermia for neonates with hypoxic-ischemic encephalopathy. N Engl J Med 2005;353:1574–84. ¨ Gunn AJ, Hoehn T, Hansmann G, Buhrer C, Simbruner G, Yager J, Levene M, Hamrick SE, Shankaran S, Thoresen M. Hypothermia: an evolving treatment for neonatal hypoxic ischemic encephalopathy. Pediatrics 2008;121:648–9. Robertson NJ. Air or 100% oxygen for asphyxiated babies? Time to decide. Crit Care 2005;9:128–30. Tan A, Schulze A, O’Donnell CP, Davis PG. Air versus oxygen for resuscitation of infants at birth. Cochrane Database Syst Rev 2004;3:CD002273. Vento M, Sastre J, Asensi MA, Vina J. Room-air resuscitation causes less damage to heart and kidney than 100% oxygen. Am J Respir Crit Care Med 2005;172:1393–8. Palmer JE. Ventilatory support of the critically ill foal. In: Sanchez LC (ed.) Veterinary Clinics of North America Equine Practice Neonatal Medicine and Surgery. Philadelphia: Saunders Elsevier, 2005; pp. 457–86. Lu KG, Barr BS, Embertson R, Dallap Schaer B. Dystocia: a true equine emergency. Clin Tech Equine Pract Emerg Med 2006;5:145–53. Gigu`ere S, Slade JK, Sanchez LC. Retrospective comparison of caffeine and doxapram for the treatment of hypercapnea in foals with hypoxic-ischemic encephalopathy. J Vet Intern Med 2008;22:401–5. Gigu`ere S, Sanchez LC, Shih A, Szabo NJ, Womble AY, Robertson SA. Comparison of the effects of caffeine and doxapram on respiratory function in foals with induced respiratory acidosis. Am J Vet Res 2007;68:1407–15. Buechner-Maxwell VA. Nutritional support for neonatal foals. In: Sanchez LC (ed.) Veterinary Clinics of North America Equine Practice Neonatal Medicine and Surgery. Philadelphia: Saunders Elsevier, 2005; pp. 487–510.
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Stabilization and Preparation for Transport 26. Mellor DJ, Cockburn F. A comparison of energy metabolism in the new-born infant, piglet and lamb. Q J Exp Physiol 1986;71:361–79. 27. Mackay RJ. Neurologic disorders of neonatal foals. In: Sanchez LC (ed.) Veterinary Clinics of North America Equine Practice Neonatal Medicine and Surgery. Philadelphia: Saunders Elsevier, 2005; pp. 387–406. 28. Arguedas MG, Hines MT, Papich MG, Farnsworth KD, Sellon DC. Pharmacokinetics of butorphanol and evaluation of physiologic and behavioral effects after intravenous and intramuscular administration to neonatal foals. J Vet Intern Med 2008;22:1417–26.
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29. Crisman MV, Wilcke JR, Sams RA. Pharmacokinetics of flunixin meglumine in healthy foals less than twenty-four hours old. Am J Vet Res 1996;57:1759–61. 30. Ebersp¨acher E, Stanley SD, Rezende M, Steffey EP. Pharmacokinetics and tolerance of transdermal fentanyl administration in foals. Vet Anesth Analg 2008;35:249–55. 31. Tyler-McGowan CM, Hodgson JL, Hodgson DR. Failure of passive transfer in foals: incidence and outcome on four studs in New South Wales. Aust Vet J 1997;75:56–9. 32. Robinson JA, Allen GK, Green EM, Fales WH, Loch WE, Wilkerson CG. A prospective study of septicaemia in colostrum-deprived foals. Equine Vet J 1993;25:214–19.
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Chapter 25
Transfusion Techniques John F. Freestone
There have been dramatic advances in our ability to collect and administer blood and plasma. At present, red blood cells and plasma remain the most commonly transfused blood products in foals, although over time it may become possible to harvest and transfuse additional blood factors and blood-like products.
Blood transfusion Administration of whole blood is used to treat acute or chronic blood loss (Table 25.1). In adults it has been suggested that, following acute blood loss, a packed cell volume (PCV) less than 20% maintained for over 24 hours is an indication for a blood transfusion.1 In foals, it may be more appropriate in acute blood loss to rely on clinical examination findings rather than PCV measurements as these are highly variable but will be decreased. A foal with acute blood loss will often present in recumbency or with profound weakness, lethargy, and a loss of interest in nursing. Physical examination can identify very pale pink to white or severely jaundiced mucous membranes, tachycardia, and tachypnea. Further diagnostics may confirm (hematology), or identify and confirm, the site and cause of the blood loss (e.g., ultrasonography of the chest (hemothorax, hemopericardium) or abdomen (hemoperitoneum)) (Table 25.1). In these cases of acute blood loss, oxygen-carrying support is rapidly needed. In the past, this has been addressed by administration of whole red cells, but now hemoglobin-based oxygen carriers (HBOCs) are also available (Oxyglobin, Biopure Corporation, Cambridge, MA). Chronic anemia allows the patient more time to adapt to the decreased red cell volume but it is accepted that a PCV of less than 10% warrants blood transfusion. A low but stable PCV may not require transfusion, especially if red cell regeneration is evi-
dent. It is very rare to need to transfuse a foal with anemia of chronic disease (Table 25.1). In severe acute blood loss in the foal the clinician often needs to improve oxygen-carrying capacity immediately for the survival of the patient. HBOCs are readily available and easily administered, followed by whole blood administration. For whole blood administration a donor, collection equipment, and administration techniques need to be readily at hand. Laboratory facilities may not be available for complete crossmatching. Agglutination crossmatching as a quick test is advised but simply may not be available in the field during an emergency. In this situation, the first transfusion is usually safe as natural equine alloantibodies are rare and react only weakly.2 Clinicians should not hesitate to collect and administer whole blood in a blood loss emergency. A young gelding with no prior history of receiving a transfusion is the best initial choice, with Standardbreds and Quarter Horses preferred.3 If subsequent transfusions are needed then crossmatching is indicated. Calculations can be made on the required volume of donor blood needed to restore a patient’s PCV to a desired level:4 (bodyweight (kg) × blood volume (mL/kg) × (PCV desired − PCV observed)) /PCV of donor blood The blood volume expressed as a function of bodyweight is high at birth and rapidly decreases over the first months of life. Guideline blood volumes for different aged foals are 150 mL/kg for a 2-dayold foal, 99 mL/kg at 2 weeks, 93 mL/kg at 4 weeks, 82 mL/kg at 12 weeks, and 69 mL/kg at 24 weeks.5 The benefit of crossmatched compatible red cells is short with a half-life of only 4 days but is often critical in the face of ongoing hemorrhage.3
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 25.1 Anemia in foals Cause Acute anemia Fractured ribs
Neonatal isoerythrolysis
Trauma Prolonged umbilical hemorrhage Neonatal thrombocytopenia Coagulopathies Chronic anemia Low-grade ongoing blood loss Gastric ulceration Parasites NSAID toxicity Anemia of chronic disease
Result
Hemothorax Hemopericardium Hemoperitoneum Rapid destruction of red cells with low PCV (< 12) Progressive red cell destruction with decreasing PCV over 3–7 days of age, may or may not need transfusion Damage to a blood vessel Laceration of lung, spleen, kidney, or liver Bleeding disorders
PCV, packed cell volume; NSAID, non-steroidal anti-inflammatory drug.
In chronic anemia a more measured approach to blood transfusion is undertaken as it is rarely an emergency procedure. Complete crossmatching is indicated. Red blood cells can be collected from a suitable donor, centrifuged, and washed so that only red cells are transfused. This will minimize the likelihood of transfusion reactions and limit the administration of foreign proteins. Neonatal isoerythrolysis (NI) most commonly presents with clinical signs of acute blood loss, but at times there is an ongoing slow decline in PCV resembling chronic blood loss (Table 25.1). Here, maternal red blood cells can be easily collected but must be washed to remove erythrocyte antibodies present in the mare’s plasma. Thorough washing of the maternal red cells can often be performed by human hospitals or blood banks. Collected maternal red cells can be washed by allowing the red cells to sit for 2 hours and removing as much plasma as possible followed by washing in normal saline. This is a useful technique but not as effective as centrifugation as some maternal plasma-containing alloantibodies invariably remain.
Hemoglobin-based oxygen carrier: Oxyglobin Oxyglobin is an acellular hemoglobin solution that is able to deliver oxygen to the tissues and is indicated in the treatment of any type of anemia. Oxyglobin has been successfully used to treat cases of acute hemorrhage in the horse.6 Oxyglobin is readily available for administration in the face of acute blood
loss and is ideal as a therapeutic bridge while preparing for a whole blood transfusion. The half-life of administered Oxyglobin is shorter than red blood cells at 18–43 hours.7 Oxyglobin has a long shelf life of 36 months without refrigeration and no blood typing or crossmatching is required owing to removal of red cell membranes during ultrapurification. In foals care is needed in the rate of administration as the large polymers of hemoglobin result in increased plasma oncotic pressure and can predispose to volume overload and pulmonary edema.8 A conservative administration rate in foals is 7.5 mL/kg/h, which equates to three 125-mL bags for a 50-kg foal (F.T. Bain, personal communication). At present, our knowledge of the safety and efficacy of Oxyglobin in the foal is ongoing, but it appears safe and highly beneficial as a single emergency bridging procedure.
Plasma transfusion The administration of plasma has now become a routine procedure in equine medicine. It is widely used in a number of clinical situations for protein replacement to restore plasma oncotic pressure and provide clotting factors or immunological support. The two most common conditions in the foal are for protein replacement secondary to gastrointestinal protein losses (acute enteritis/colitis, e.g., salmonellosis, Lawsonia intracellularis, Clostridium perfringens) and for the treatment of failure of passive transfer in both healthy and sick newborn foals. Owing to its increased clinical use, plasma is now readily available commercially. All large equine clinics maintain a
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bank of frozen plasma collected either via their own plasmapheresis equipment or from a commercial supplier. Commercial plasma is collected via plasmapheresis in a sterile fashion and is free of red cells. In addition, the commercial plasma donors are blood typed and lack the common red cell alloantigens Aa and Qa and serum alloantibodies, forgoing the need for compatibility testing. Donors are also screened for communicable diseases such as equine infectious anemia (EIA), piroplasmosis (Babesia equi and Babesia caballi), dourine (Trypanosoma equiperdum), glanders (Burkholderia mallei), brucellosis, and equine erhlichiosis. It should be noted that not all plasma, serum, and blood products sold are licensed. The differences for licensed and non-licensed blood products are addressed by the Biological and Therapeutic Agents Committee of the American Association of Equine Practitioners (AAEP) and a list of North American licensed agents noted.9 During plasmapheresis whole blood is separated into its components, allowing the plasma fraction to be removed and donor red cells to return, which enables more frequent use of the donors for plasma collections. The volume of donor plasma to correct a total protein deficit can be estimated using: ((70 − PPR )(0.05 BWR ))/PPD where PPR and PPD are the recipient and donor’s plasma protein concentration (g/L), respectively, and BWR is the recipient’s bodyweight (kg). In general, the volume of plasma to administer to a patient is dictated by the clinical scenario and aims to replace deficits and ongoing losses. In the foal the most common use of plasma is to support foals with failure of passive transfer. A healthy foal with partial failure of passive transfer (immunoglobulin (Ig) G < 800 mg/dL) often receives 1 liter of plasma and then the IgG concentration is reassessed. Alternatively, a septic neonate with complete failure of passive transfer may require repeated administration of plasma to replace a low IgG concentration and to meet ongoing losses. Two specific and commonly available commercial plasma products contain antibodies against Rhodococcus equi obtained from donors vaccinated with R. equi antigens and administered to foals to reduce the risk of R. equi pneumonia or J-5 anti-endotoxin antibodies for the treatment of endotoxemia and sepsis. It is common for high-risk farms with virulent R. equi to establish a program where all foals receive R. equi plasma in the first week of life, and possibly repeated administration in 21–28 days to help prevent infection in immunologically naive foals. Plasma can also be produced by collecting whole blood, centrifuging it, or allowing it to sit for over 2 hours. Plasma can then be collected from the top
of the collection bags or bottles. This procedure is not ideal as it runs the risk of non-sterile collection and contamination with red cells resulting in recipient sensitization.
Donor selection Horse blood typing is complex. There have been at least 30 blood factors identified and these are grouped into seven blood systems: A, C, D, K, P, Q, and U with the possibility of 400 000 equine blood groups.3 For this reason the universal blood donor probably does not exist. The red cell alloantigens Aa and Qa are the most immunogenic and donors should be negative. Ideally, crossmatches should be performed prior to blood administration to find the most suitable donor as alloantibody production still results in over 50% of cases.4 Routine crossmatching utilizes washed red cells from the donor (major) and recipient (minor) with incubated serum from the other. Gross and microscopic examination for clumping identifies serum agglutinins. Hemolysins must be detected by adding complement and, for this reason, this assay is generally performed by a specialized laboratory. Donor selection for plasma collection is not as critical as it is for red blood cell or whole blood administration. Only alloantibodies to recipient red blood cells need to be considered. As a general guideline, horses negative for the Aa and Qa alloantigens are suitable plasma donors. Multiparous mares should not be used as donors.
Blood collection and administration Blood is collected from a suitable donor from the surgically prepared jugular vein via a large-gauge catheter (10–12 G) or needle (14 G) with adherence to strict asepsis. The donor can be tranquilized as needed. Blood is collected into bottles or bags containing acid–citrate–dextrose (ACD) solution or sodium citrate. Citrate binds calcium and prevents clotting. The dextrose in ACD solution provides an energy source for the collected red cells and is therefore the preferred anticoagulant for transfusion of red cells or whole blood, whereas sodium citrate is used for plasma collection and storage. Plastic bags are superior to bottles as they do not activate platelets or clotting factors. Plastic bags come in a variety of sizes from 500 mL to 3 liters. Blood collection into 1-liter or 2-liter Baxter evacuated glass containers (Baxter Health Care Corporation, Deerfield, IL) is quick and easy.10 ACD solution is added to the bottle prior to collection (200 mL per 2-liter bottle). Care is needed following ACD instillation to prevent the loss of vacuum by
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Table 25.2 Transfusion reactions Hemolytic
Stop transfusion, replace blood with normal saline. Maintain renal blood flow. Furosemide 1 mg/kg i.m. or i.v.
Non-hemolytic febrile Anaphylactic
Administer flunixin meglumine 1 mg/kg i.v. Stop transfusion. Support circulation and airway as necessary. Administer epinephrine 1 mL of 1:1000/100 kg i.v. Antihistamines (pyrilamine maleate 1 mg/kg i.v.). Prednisolone sodium succinate 1–2.5 mg/kg i.v. Flunixin meglumine (1 mg/kg i.v.)
Urticarial or allergic reactions
Antihistamines or hydroxyzine (1 mg/kg orally b.i.d.) and anti-inflammatory agents. Consider corticosteroids if no improvement
Transfusion-related acute lung injury
Support respiratory distress by administering O2 and anti-inflammatory agents
Bacterial contamination
Antibiotics and anti-inflammatory agents
i.v., intravenously; i.m., intramuscularly; b.i.d., bis in die (twice a day).
using a single stick and careful removal of the needle from the vacutainer bottle first. Following venipuncture, blood fills the collection tubing which is then introduced into the evacuated bottle containing ACD solution. The collection bottle is continuously swirled during collection. Once the bottle is filled, it can be administered immediately to the patient via administration sets equipped with appropriate filters to remove fibrin and other debris. High-flow Mila (Mila International Inc., Florence, KY) blood collection and administration sets are available allowing 8 liters of blood to be collected over 20 minutes.10
Complications of administration of blood products All clinicians who routinely administer blood products or plasma fear a severe transfusion reaction. For this reason, transfusions should be started slowly and patients should be closely monitored, assessing heart rate and respiration rate throughout the procedure. Common clinical signs of a transfusion reaction include tachypnea, tachycardia, dyspnea, sweating, muscle fasiculations, fever, urticaria, and sudden death.3 The transfusion should be stopped immediately at the first signs of a problem. Transfusion reactions can be further classified as hemolytic reactions, non-hemolytic febrile reactions, anaphylactic reactions, urticarial or allergic reactions, allosensitization to blood components, transmission of disease, and reactions associated with the presence of endotoxin or other blood contaminants. It is advisable to have epinephrine (1 mL of 1:1000 solution/100 kg intravenously (i.v.)), flunixin meglumine (1.1 mg/kg i.v.), antihistamines (pyrilamine maleate 1 mg/kg i.v., intramuscularly (i.m.) or subcutaneously), corticosteroids (prednisolone sodium succinate 1.0–2.5 mg/kg i.v.) and calcium gluconate (slowly i.v. to effect) available to address an emergency situation (Table 25.2). Corticosteroid administration
is helpful in decreasing inflammation and allergic reactions by suppressing neutrophil migration and capillary permeability but will take 4–6 hours for onset of action. Antihistamines have shown little benefit as a sole medication in equine anaphylaxis but may be useful as part of a broader treatment plan.11 Although citrate in the ACD solution can bind calcium and induce signs of hypocalcemia, it rarely occurs. Following a hemolytic transfusion reaction, isotonic polyionic fluids should be administered to protect renal blood flow and renal function. Fluids should be administered slowly with care and may be contraindicated with signs of dyspnea secondary to pulmonary edema. Transfusion of incompatible plasma will result in red cell hemolysis and bilirubin production leading to anemia and jaundice respectively. Particular care is needed if additional transfusions are required and more than 4 days has elapsed since the initial transfusion, as alloantibodies may have developed, although all transfusions have an element of risk.
References 1. Carlson GP. Diseases of the hematopoietic and hemolymphatic systems. In: Smith BP (ed.) Large Animal Internal Medicine, 2nd edn. St. Louis: Mosby, 1996; pp. 1198–213. 2. Sellon DC. Diseases of the hematopoietic system: blood and blood component therapy. In: Kobluk CN, Ames TR, Goer RJ (eds) The Horse: Diseases and Clinical Management. Philadelphia: W.B. Saunders, 1995; pp. 1103–10. 3. Morris DD. The haemolymphatic system: blood and plasma therapy. In: Higgins AJ, Snyder JR (eds) The Equine Manual, 2nd edn. London: Elsevier Saunders,1995; pp. 525–7. 4. Williamson L. Blood and plasma therapy. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 3rd edn. Philadelphia: W.B. Saunders, 1992; pp. 517–20. 5. Spensley MS, Carlson GP, Harrold D. Plasma, red blood cell, total blood and extracellular fluid volumes in healthy horse foals during growth. Am J Vet Res 1987;48: 1703–7.
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6. Maxson AD, Giger U, Sweeney CR, Tomasic M, Saik JE, Donawick WJ, Cothran EG. Use of bovine hemoglobin preparation in the treatment of cyclic ovarian hemorrhage in a miniature horse. J Am Vet Med Assoc 1994;204:509–11. 7. Hohenhaus AE. Editorial: Oxyglobin: A transfusion solution? J. Vet Intern. Med. 2002;16:394–5. 8. Belgrave RL, Hines MT, Keegan RD, Warsdrop KJ, Bayly WM, Sellon DC. Effects of a polymerized ultrapurified bovine haemoglobin blood substitute administered to ponies with normovolemic anemia. J Vet Intern Med 2002;16:396–403.
9. American Association of Equine Practitioners. White Paper: Information on equine plasma and serum products for the equine practitioner. Biological and Therapeutic Agents Committee. Lexington, KY: AAEP, 2009 (www.AAEP.org). 10. Slovis NM, Murray G. How to approach whole blood transfusions in horses. In: Proceedings of the 47th Annual Convention of the American Association of Equine Practitioners, 2001, pp. 266–9. 11. Swiderski C. Hypersensitivity. In: Kobluk CN, Ames TR, Goer RJ (eds) The Horse: Diseases and Clinical Management. Philadelphia: W.B. Saunders, 1995; pp. 1065–72.
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Chapter 26
Diagnostic Ultrasonography of the Abdomen Virginia B. Reef and JoAnn Slack
Ultrasonography is a non-invasive diagnostic technique that provides the veterinary clinician with a window through which the intra-abdominal structures and organs can be evaluated. The foal abdomen is better suited to ultrasonographic investigation than the adult abdomen because of its overall smaller size, better visualization of the liver and bladder during the neonatal period, relatively small size of the gasfilled large colon, and ability to position the foal (lateral recumbency or standing).1–3 The preponderance of small intestinal disorders in foals presenting with colic also enables the veterinary ultrasonographer to assess many different gastrointestinal lesions.1–5 Sedation is usually not necessary for patient preparation except in unusually fractious foals. The foal should be prepared by clipping the hair over the portion of the abdomen under investigation with a #40 surgical clipper blade, cleaning the skin, and applying an ultrasonographic coupling jelly. Transducer selections should be based on use of the highest frequency transducer that will penetrate adequately to the area under investigation and yield optimal image quality. Ultrasonographic evaluation of the foal abdomen and gastrointestinal tract should begin with a 15.0–10.0-MHz transducer for optimal resolution of the intestinal wall. Edematous compromised loops of intestine are heavier, fall to the ventral aspect of the abdomen, and are readily imaged. After thorough investigation of the abdomen with this probe, a 10.0–5.0-MHz microconvex transducer may be used to image the deeper abdominal structures, followed by a 5.0–3.0-MHz large convex transducer, if needed, for additional penetration in larger foals. Ultrasonographic evaluation of the gastrointestinal tract is easier with the foal standing, as gas rises dorsad and any free fluid in the abdomen provides an acoustic win-
dow through which the sonographic evaluation can be performed.
Peritoneal cavity Ultrasonographic evaluation of the normal foal abdomen reveals little or no peritoneal fluid. Visualization of fluid with cellularity (echogenicity) is consistent with a diagnosis of hemoperitoneum or peritonitis.1, 2, 4, 6 Fibrin strands may be imaged between the serosal surface of the bowel and other intra-abdominal structures in foals with peritonitis. Maturing adhesions appear as echogenic thick bands between the serosal surface of the bowel or other intra-abdominal organs. Adhesions may also be suspected when no movement is detected between a portion of bowel and adjacent structures. A hemoperitoneum may be suspected if homogeneous swirling echogenic fluid is imaged within the peritoneal cavity. The detection of a pneumoperitoneum is indicative of a ruptured viscus, unless the foal has recently had abdominal surgery. Flocculent echogenic fluid with free gas echoes within the peritoneal cavity should also suggest a ruptured viscus (Figure 26.1) or the possibility of a bacterial peritonitis with anaerobic involvement.1, 2, 6 If a peritoneal fluid sample is desired, the ultrasonographic examination can be used to localize fluid, minimizing the risk of accidental gastrointestinal perforation from needle insertion.2, 5 Mesenteric lymphadenopathy appears as enlarged hypoechoic oval nodular structures within the gastrointestinal mesentery and can be seen with Rhodococcus equi infection and enterocolitis (Figure 26.2). Abscessation of individual mesenteric lymph nodes has been imaged in affected foals. Large intraabdominal abscesses associated with Rhodococcus equi
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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LEFT
Spleen
Figure 26.1 Sonogram from an 8-day-old Thoroughbred colt with a ruptured stomach. The spleen and small intestine can be seen floating within echoic, flocculent fluid consistent with rupture of a gastrointestinal viscus. The defect in the stomach was not visualized sonographically but was identified along the greater curvature of the stomach at postmortem.
and other organisms can often be identified ultrasonographically.2–4,7 Adhesions associated with internal abdominal abscesses (often associated with R. equi) can cause small intestinal obstructions, which can be strangulating or non-strangulating (Figure 26.3).
Surgical disorders Detection of a target or bull’s-eye appearance to the small intestine is diagnostic of a small intestinal intussusception2, 3, 8, 9 (Figure 26.4). This appearance is caused by the relative different tissue densities
LVENTRAL
Figure 26.2 Sonographic image from a 2-month-old Thoroughbred colt with mesenteric lymphadenopathy associated with Rhodococcus equi infection. Notice the enlarged heteroechoic lymph nodes in the colonic mesentery (arrows).
of the central edematous intussusceptum, the outer edematous intussuscipiens, and the interposed fluid. Distension of the more proximal portions of small intestine with fluid is imaged in foals with intussusceptions or other types of strangulating obstructions.1, 2, 6, 8 Fluid and gas distension of the stomach can be visualized from the left cranial abdomen in the 7th to 8th through the 10th to 12th intercostal spaces or from the ventral abdomen (neonates only) caudal to the liver.1–3,6, 8 In foals with a small intestinal obstruction the stomach will be distended with anechoic fluid (gastric reflux). Compromised intestine has thickened, hypoechoic, edematous walls with little or no detectable peristaltic activity. With complete volvulus of the small intestine the entire abdomen is filled with thickened, edematous, hypoechoic, distended loops of small intestine with no visible peristaltic activity.1–3 Ventral settling of the ingesta in a turgid-fluid-distended small intestine is most commonly associated with small intestinal obstruction. Intramural masses and intestinal strictures have been detected ultrasonographically in foals.1, 2 Infection, microabscesses, and hematomas of the intestinal wall are possible causes of intramural masses in young foals; neoplasms are rare. Intraluminal masses such as ascarid and meconium impactions may also be visualized ultrasonographically.1, 2 Ascarids may be visualized within the lumen of the intestinal tract, as individuals or as a mass of echogenic curvilinear parasites. Meconium impactions may be visualized as hypoechoic to echogenic masses within the lumen of the large and/or small colon (Figure 26.5).2, 3 Distension of the intestine proximal to the impaction with fluid and gas is common and may aid in the visualization of the intraluminal mass. Large bowel displacements or torsions are difficult to diagnose ultrasonographically, but may be suspected in a foal with much gaseous distension of the large bowel and persistent unrelenting pain. 2, 3 Many foals with gaseous distension of the large bowel have spasmodic colic that resolves with symptomatic therapy. Displacement of gastrointestinal viscera into another body cavity, such as into the thorax in a foal with a diaphragmatic hernia, may be diagnosed ultrasonographically (Figure 26.6). If the edge of the diaphragm is involved, direct visualization of the diaphragmatic defect may be possible ultrasonographically. Laceration of the diaphragm associated with rib fractures should be considered in neonatal foals with diaphragmatic hernia (see Figure 26.6). Gastrointestinal viscera are visualized in the thorax with dorsal displacement of lung. Obstruction of the intestine with bezoars does occur occasionally in foals, and the bezoar can be imaged as a hyperechoic shadowing, oval-to-circular mass obstructing the intestine if it is located in a portion of the intestine that is imageable ultrasonographically.2, 3 Atresia
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Figure 26.3 Sonograms of an intra-abdominal Rhodococcus equi abscess in a 4-month-old Thoroughbred foal. (a) Notice the 8.32-cm echogenic abscess (arrow) surrounded by a hypoechoic capsule axial to the fluid- and gas-distended stomach. There is a dorsal gas cap with ventral fluid in the stomach. (b) Numerous adhesions to the abscess were present causing small intestinal obstructions. Notice the distended loops of small intestine (arrowheads) adjacent to collapsed small intestinal loops (arrow).
coli may be suspected if gas and fluid distension of the more proximal portions of the intestine are visualized in the absence of edematous, hypoechoic, and distended bowel.2 Visualization of the caudal portion of the gastrointestinal tract is usually difficult in these foals, because of the large amount of distension in the more proximal intestine. If visualized, the caudal portion of the gastrointestinal tract should appear empty. Direct ultrasonographic visualization of the atretic portion of the gastrointestinal tract has not been performed successfully in foals, to the authors knowledge.
Medical disorders Medical colics can often be distinguished ultrasonographically from surgical colics based on the ultrasonographic appearance and motility of the gastrointestinal tract. Fluid-filled small or large intestines with normal to increased motility are consistent with an enteritis or enterocolitis.1–3 The wall of the affected portion of the intestine may be thicker than normal, hypoechoic, and edematous if severe inflammatory bowel disease is present. Shreds or fragments of intestinal mucosa may be seen floating within
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Figure 26.4 Transverse (a) and longitudinal (b) sonographic images from a 5-day-old Standardbred filly with a jejunojejunal intussusception. Notice the target-like appearance of the intussusception in the transverse view. In the longitudinal view the outer intussuscipiens (between black arrows) and the inner intussusceptum (between white arrows) can be clearly visualized. Their combined thickness measures 1.05 cm.
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Figure 26.5 Sonograms from a 3-day-old Standardbred filly with meconium retention within the small colon (arrows) dorsal to the bladder. Notice the hypoechoic speckled appearance of the meconium.
the gastrointestinal contents, particularly in foals with severe necrotizing enterocolitis.2, 3 Gas echoes in the wall of the gastrointestinal tract and along the mucosal surface have been seen in foals with necrotizing enteritis (Figure 26.7).2, 3 Duodenitis usually results in thickening of the duodenal wall, distension of the duodenum, and decreased duodenal motility. 2, 3 Severe thickening of the duodenal wall and subsequent stricture can occur, and these foals have a decreased duodenal lumen and gastric distension.2, 3 Thickening of the intestinal wall, particularly the small intestine, is readily imaged in foals with Lawsonia intracellularis infection.3 Foals with ileus usually have a hypomotile to amotile distended intestine, usually with a normal wall thickness.2, 3 Marked gastric distension can be readily detected from the left side of the abdomen and the luminal contents evalu-
Figure 26.7 Sonogram from a 3-day-old Thoroughbred filly with necrotizing enterocolitis. Notice the pinpoint hyperechoic echoes within the thickened wall (arrow) of the small intestine consistent with intramural gas.
ated (Figure 26.8). The nasogastric tube can be visualized within the gastric lumen as a circular hyperechoic structure (see Figure 26.8). Sand enteritis can also occur in foals. The sand grains result in the intestine laying flat against the floor of the ventral abdomen with a sparkly appearance along the mucosal surface due to the grains of sand reflecting the ultrasound beam.3 Thickening of the wall of the intestine also occurs, indicating more chronic sand ingestion.3 Clinical findings, clinical pathological data, and ultrasonographic findings can complement each other and aid the clinician in arriving at or refining a diagnosis of surgical or medial colic.
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Figure 26.6 Sonograms from a 5-day-old Thoroughbred colt with a diaphragmatic hernia and fractured ribs. (a) Sonogram of the small intestine (SI) in the right side of the thorax at the 6th intercostal space. Notice the diaphragm (arrow) ventral to the small intestine and lung. The ventral edge of the lung is irregular due to the dorsal displacement of the lung by the herniated small intestine. (b) Sonograms of the fractured 6th rib showing marked outward displacement (arrows) of the distal end of the rib.
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Lower urinary tract Ultrasonographic evaluation of the neonatal bladder may be performed from the ventral abdomen using a 15.0–7.5-MHz curved or straight linear transducer until the foal is 4 to 8 weeks old.1, 2, 4, 10 At that time, gastrointestinal viscera become superimposed between the bladder and ventral abdominal wall. Omphaloarteritis or urachitis may result in persistent visualization of the bladder in older foals. The neonatal bladder is usually round or oval and filled with relatively sonolucent fluid. This is in contrast to adult horse urine, which is often echogenic because of the large amount of mucus and calcium carbonate crystals. Distortion of the intact neonatal bladder most commonly occurs in foals with large amounts of gastrointestinal distension. Homogeneous, echogenic to hypoechoic intraluminal masses have been seen in foals associated with a blood clot in the bladder (Figure 26.9).1 Defects of the bladder wall may be visible ultrasonographically, or the appearance of the bladder may suggest uroperitoneum.1–3,11 With a bladder rupture, the bladder appears collapsed ultrasonographically. It may be folded on itself or the rent in the bladder wall may be visualized with fluid communicating between the bladder and the peritoneal cavity (Figure 26.10).2 Large amounts of usually anechoic fluid are visible within the peritoneal cavity, and the gastrointestinal viscera float in this free fluid.2, 3, 5, 11, 12 A thickened edematous hypoechoic to anechoic bladder wall has been seen in a foal with a necrotic bladder and uroperitoneum, while gas echoes in the bladder wall have been imaged in foals with an
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Figure 26.9 Sonogram of a 1-day-old Thoroughbred filly with a blood clot in the bladder. The foal was born without complications but found to be straining to urinate and passing bloodcolored urine and blood clots at 6 hours of age. Notice the heterogeneous echogenic mass (arrow) representing the blood clot surrounded by hypoechoic urine within the bladder.
emphysematous bladder and uroperitoneum.2 Rents in the urachus or ureters may also cause uroperitoneum. Tears in the ureter or urachus are difficult to visualize directly, but may be suspected because of the ultrasonographic appearance of the surrounding structures and the bladder. With a urachal tear, the bladder may be filled with anechoic fluid. Fluid may be imaged dissecting retroperitoneally along the urachus and falciform ligament, or the defect in the urachus may be visualized communicating with the peritoneal cavity.2, 12 Ureteral ruptures are more difficult to diagnose, because the ureter is not
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Figure 26.8 Sonogram of gastric hemorrhage from a 6-day-old Thoroughbred colt with neonatal encephalopathy. Notice the heteroechoic material with scattered hyperechoic gas echoes within the stomach (a). A stomach tube was passed to delineate better the wall of the stomach (arrowheads) from the lumen (b). The tube is the echoic circular structure (arrow) that casts an acoustic shadow. With the tube in place the blood clots were determined to be within the lumen and not within the wall of the stomach. The foal went on to develop significant bloody reflux and diarrhea necessitating whole blood transfusion.
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Figure 26.10 Transverse sonogram of a 1-day-old Thoroughbred colt with a ruptured bladder. Notice the collapsed bladder folded in on itself (arrow) and the umbilical arteries on either side of the bladder (arrowheads). There is a large amount of anechoic fluid free in the abdomen.
normally visible. Accumulation of anechoic fluid around the kidney in the retroperitoneal space would indicate this possibility.2 The bladder of the foal with a ureteral rupture will appear intact, but may contain less urine than that of the foal with the urachal defect. This diagnosis should be suspected in foals with a large amount of anechoic fluid visible in the peritoneal cavity, an intact bladder, and a normal urachus2, 3 Persistent patent urachus is visible ultrasonographically in the neonatal foal as a fluid communication extending from the bladder apex cranially to the external umbilical remnant.1, 2 The urachus is normally a hypoechoic area between the two umbilical arteries from the umbilicus to the bladder apex, and is a potential space. A urachal diverticulum may form at the bladder apex with closure of the patent urachus.
Upper urinary tract Evaluation of the foal kidneys can easily be performed from the flank area using a 7.5- or 5.0-MHz sector scanner or curved linear transducer, depending on the size of the foal and the kidney under investigation.1,13–15 The right kidney is normally found in the dorsal portion of the abdomen between the 14th and 16th intercostal spaces in a cranial to caudal plane, and between the dorsal and ventral aspects of the tuber coxae in a dorsal to ventral plane. Ul-
trasonographic evaluation of the right kidney can be performed with a 7.5-MHz transducer in the small foal, whereas a 5.0-MHz transducer may be needed in a larger, older individual. The left kidney is more caudal and slightly more ventral in the abdomen and is found between the 16th intercostal space and the paralumbar fossa in a cranial to caudal plane, and the dorsal aspect of the tuber coxae and the tuber ischii in a dorsal to ventral plane. This kidney is medial to the spleen and farther away from the abdominal wall. Therefore, a 5.0-MHz transducer is needed to evaluate this kidney ultrasonographically in all but the neonatal foal. Older foals (weanlings and yearlings) may require a 3.5-MHz transducer. The kidney is the least echogenic abdominal organ, with a relatively anechoic medulla, echogenic renal pelvis, and hypoechoic cortex, which is less echogenic than the adjacent hepatic parenchyma.1, 2, 16 An ultrasonographic-guided biopsy can be performed if indicated.1, 17 Congenital renal anomalies are rare in foals, but cysts may be detected as anechoic fluid-filled portions of the kidney with acoustic enhancement of the far wall (typical of a cystic structure). Hypoplastic kidneys are smaller than normal, while dysplastic kidneys are misshapen and hyperechoic with a decreased corticomedullary distinction.2, 3, 18 Renal agenesis should be suspected when a kidney cannot be identified ultrasonographically.2, 3 Acute renal failure is most commonly seen and often results in large, swollen sonolucent kidneys (more anechoic than normal).1, 2, 16, 19, 20 This ultrasonographic finding is seen in some of the foals with hypoxic renal injury, but is absent in others.2, 3 Echogenic kidneys are more commonly seen with any type of cellular infiltrate (inflammatory, granulomatous, and fibrotic) and often indicate chronicity.1, 2, 16,20–22 The administration of phenylbutazone has resulted in the development of a diffuse hyperechoic zone in the medulla near the corticomedullary junction.3 Ultrasonographic visualization of the ureter suggests ureteral obstruction and hydroureter.2 Perirenal edema is rarely seen, but may be detected in foals with a ruptured ureter or renal failure (usually acute).2 A perirenal hematoma can also occur in foals secondary to abdominal trauma and appears as an echoic homogeneous or heterogeneous mass that is often loculated in the perirenal space.
Liver The liver can be imaged from both sides of the abdomen ventral to the ventralmost lung margin as well as from the cranial ventral abdomen in the neonatal foal.1–3,14, 15, 23, 24 Ultrasonographic
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Diagnostic Ultrasonography of the Abdomen evaluation of the foal liver is performed with a 7.5MHz transducer in the neonate or a 5.0-MHz transducer in the older foal. Once the foal reaches 4 to 8 weeks of age, gastrointestinal viscera become interposed between the ventral abdominal wall and the liver, preventing its visualization from the cranial ventral abdomen. Hepatic parenchyma can normally be visualized on the right side of the foal abdomen from the 6th to the 15th intercostal spaces ventral to the ventralmost lung margins, and similarly in the left cranial abdomen from the 6th to the 9th intercostal spaces. Between 4 and 8 cm of hepatic parenchyma may normally be visualized. The entire liver is not accessible for ultrasonographic evaluation in the normal foal because of superimposed aerated lung in the thoracic cavity and gastrointestinal viscera in the cranial ventral abdomen. The caudal vena cava can often be visualized in the neonatal foal, but cannot normally be visualized in older individuals. Estimations of hepatic size are therefore based on the amount of hepatic parenchyma that can be visualized ventral to the ventralmost lung margin and are only relative estimates at best. The hepatic parenchyma is intermediate in echogenicity between the kidney and spleen.2, 14, 15, 23, 24 Visualization of the portal vein and hepatic veins is easily accomplished, whereas bile ducts are not visible in normal foals. Portosystemic shunts are rare in foals but may be detected with Doppler ultrasonography.3 Increased sonolucency and collapse of the hepatic parenchyma with a smaller than normal liver would suggest acute hepatocellular necrosis.1, 2, 24 Increased echogenicity of the hepatic parenchyma can be seen with inflammatory, granulomatous, fatty, or fibrotic infiltrates in the liver but is most commonly associated with an inflammatory cell infiltrate in the young foal.1, 2, 24, 25 Ultrasonographic-guided biopsies can be performed to assess the area of hepatic parenchyma histopathologically.1, 17 Foals with a marked inflammatory cell infiltrate in the liver will often have a liver that appears larger than normal with a relatively diffuse increase in echogenicity of the hepatic parenchyma. Tyzzer’s disease should be suspected in a foal with this ultrasonographic appearance, if the age and clinical signs are compatible.3 Miliary echogenic areas in the hepatic parenchyma have been imaged in several neonatal foals associated with microabscesses that developed secondary to neonatal septicemia.2, 3 Abscesses have also been detected in the liver of neonatal foals secondary to umbilical infections or secondary to a suppurative hepatitis.2, 3, 26 Biliary distension with sludging of bile (echogenic debris within the biliary tree) has been detected ultrasonographically in foals with proximal duodenitis.1–3 The biliary tree has thick echogenic walls and, when distended, parallel channel signs
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can be seen associated with the distended bile duct adjacent to the hepatic vein. The fluid within the biliary tree is more echogenic than blood in the hepatic vessels and lacks the rapid flow seen in the hepatic veins. The liver should also be carefully evaluated in foals with hemoabdomen since liver fractures and hematomas can occur secondary to abdominal trauma and are detectable ultrasonographically.
Spleen The spleen can be imaged from the left side of the abdomen ventral to the ventralmost lung margin as well as from the cranial ventral abdomen in the neonatal foal.1–3,14, 15, 23, 24 Ultrasonographic evaluation of the foal spleen is performed with a 7.5-MHz transducer in the neonate or a 5.0-MHz transducer in the older foal. Splenic parenchyma can normally be visualized on the left side of the foal abdomen from the 8th intercostal space ventral to the ventralmost lung margins to the paralumbar fossa. The splenic vein is a good landmark for identifying the stomach, which is in close proximity to the spleen at the level of the splenic vein. Splenic pathology is rare in foals but should be considered in foals with hemoabdomen as splenic hematomas and rupture can occur.2, 3 The spleen can be involved in foals with localized peritonitis secondary to internal umbilical remnant infection, but abscesses in the spleen are rare in foals.2
Umbilicus Umbilical remnants Umbilical remnants are visualized ultrasonographically from the ventral abdomen with a 15.0–7.5-MHz transducer with or without a hand-held standoff pad (Figure 26.11). The umbilical remnants have a characteristic size and appearance, and atrophy as the foal ages.1, 10 The umbilical vein is a small, thin-walled structure usually with a fluid lumen. It may have an echogenic core associated with clot formation. The umbilical vein normally measures 6 ± 2 mm in diameter and can be followed craniad to the liver. Umbilical arteries are thicker-walled structures and usually contain an echogenic core (clot) with little or no fluid visible in the lumen. Umbilical arteries alongside the bladder usually measure 8.5 ± 2 mm in normal foals. The urachus is a potential space between the two umbilical arteries and is imaged as a small amount of hypoechoic tissue between the two arteries. The urachus with both umbilical arteries at the bladder apex normally measures 17.5 ± 4 mm in the
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Figure 26.11 Diagram showing the umbilical vein (v), umbilical arteries (aa), and the urachus and their relationships to the umbilicus, liver, and urinary bladder. The diagram was used to locate umbilical structures ultrasonographically. (a) Anatomical region scanned to locate the umbilical vein; (b) anatomical region scanned to locate the umbilical arteries and urachus; (c) anatomical region scanned to locate the umbilical arteries and the bladder. Reproduced with permission from Reef and Collatos.10
transverse plane immediately proximal to the bladder apex. Enlargement of the umbilical remnant structures usually indicates infection1–3,26 (Figures 26.12 and 26.13). If excessive hemorrhage was noticed from the
umbilicus at parturition or if no bleeding was noticed upon rupture or cutting the umbilical cord, a large homogeneous echogenic clot may be detected within an enlarged umbilical vein or umbilical arteries. Thickening or edema of the vessel walls may
Bladder apex
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Figure 26.12 Transverse (a) and longitudinal (b) sonograms from an 11-day-old Arabian colt with a urachal abscess (arrows), arteritis (arrowheads), and tarsocrural joint sepsis. Echogenic purulent material is present within the urachus while hypoechoic fluid is present within the arteries.
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Diagnostic Ultrasonography of the Abdomen E
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Figure 26.13 Sonogram of a 20-day-old Thoroughbred colt with omphalophlebitis. Notice the thickened wall of the umbilical vein and heteroechoic material within the lumen. The heteroechoic tissue surrounding the vein is indicative of a perivasculitis. The vein was abnormal to the level of the liver prompting a complete sonographic evaluation of the liver to rule out hepatic abscesses.
also be imaged. Homogeneous echoic material may also be imaged in the retroperitoneal space along the ventral abdomen extending out from the umbilicus. These foals may also have a homogeneous echoic clot in the bladder.3 These findings should resolve rapidly if associated only with excessive hemorrhage. Hemoabdomen has also been reported secondary to rupture of an umbilical artery.3 Hypoechoic, sonolucent, or echogenic fluid in the lumen of the umbilical remnant structures is usually associated with the accumulation of purulent debris.1–3,26 If anaerobes are involved, free gas echoes are usually imaged within the lumen of the internal umbilical structures.26 Thickening of the wall of the umbilical remnant structure may also be imaged associated with an inflammatory infiltrate. In most affected foals, abnormalities of two or more umbilical remnant structures are detected ultrasonographically.26 Infections with multiple organisms are the rule, necessitating broad-spectrum antimicrobial therapy. 2, 3, 26 Metronidazole should be added to the therapeutic regimen of foals with gas within the internal umbilical remnant structures. Medical therapy alone should be considered in foals with fever of unknown origin, an elevated fibrinogen, and/or leukocytosis but who are otherwise healthy.26 Broad-spectrum antimicrobial therapy is effective in most foals with internal umbilical remnant infection when administered at the first sign of infection.27 Surgical resection of the infected umbilical remnants should be considered in foals with septicemia, septic arthritis, or other life-threatening infections.2, 26 Surgery should also be considered in foals whose umbilical vein infection extends cranially to the liver and in those with severe urachal abscessation. The physical examination find-
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ings, history, clinical pathological data, ultrasonographic findings, and previous antimicrobial history should be taken into consideration when deciding between medical or surgical therapy. If medical therapy is selected, ultrasonographic re-evaluation should be performed in 3 to 5 days, because changes can occur rapidly. Complete resolution of the infection usually takes longer and depends upon the organisms involved and the severity of the initial infection. If internal umbilical remnant structures are enlarged compared with a previous examination, a switch in antimicrobial therapy or surgical resection should be considered.
Hernias Ultrasonographic evaluation of a hernia can determine the hernial contents (intestine, omentum, and fluid), size of the hernial ring, and presence of a subcutaneous swelling or abscess1–3 (Figure 26.14). The viability of the intestine within the hernia can be evaluated by assessing wall thickness, echogenicity, distension, and motility.1–3 This can be performed successfully for umbilical, inguinal, or ventral abdominal wall hernias.
Summary In summary, ultrasonographic evaluation enables the clinician to assess non-invasively much of the abdominal viscera that was previously difficult to assess using other diagnostic techniques. The examination is well tolerated by the neonatal foal and can
Figure 26.14 Sonograms of a non-reducible umbilical hernia obtained from a 4-month-old Thoroughbred filly. These sonograms are both long-axis images of the hernia with the right image being more dorsal and the left image more ventral. Notice the loculated fluid in the hernial sac (arrowheads) in the left (ventral image), the thickened incarcerated small intestine in the right (more dorsal) image of the hernial sac, and the thickening of the wall of the small intestine as it extends ventrally into the hernia (right image). The arrows point to the serosal surface of the intestine incarcerated in the hernia.
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readily be performed with portable ultrasonographic equipment. The technique can aid in distinguishing between a medical or surgical lesion, obtaining a biopsy or aspirate, and helping to form a diagnostic plan.
References 1. Reef VB. The use of diagnostic ultrasound in the horse. Ultrasound Q 1991;9:1–34. 2. Reef VB. Equine pediatric ultrasonography. In Equine Diagnostic Ultrasound. Edited by V.B. Reef. Philadelphia, W.B. Saunders, 1998, pp. 364–403. 3. McAuliffe SB. Abdominal ultrasonography of the foal. Clin Tech Equine P 2004;3:308–16. 4. Byars TD, Halley J. Uses of ultrasound in equine internal medicine. Vet Clin N Am Equine Pract 1986;2:253–8. 5. Neal HN. Foal colic: practical imaging of the abdomen. Equine Vet Educ 2003;15:263–270. 6. Rantanen NW. Diseases of the abdomen. Vet Clin N Am Equine Pract 1986;2:67–88. 7. Hanselaer JR, Nyland TG. Chyloabdomen and ultrasonographic detection of an intra-abdominal abscess in a foal. J Am Vet Med Assoc 1983;183:1465–7. 8. Bernard WV, Reef VB, Reimer JM, Humber KA, Orsini JA. Ultrasonographic diagnosis of small-intestinal intussusception in three foals. J Am Vet Med Assoc 1989;194:395–7. 9. McGladdery AJ. Ultrasonographic diagnosis of intussusception in foals and yearlings. AAEP Proc 1996;42:239–40. 10. Reef VB, Collatos C. Ultrasonography of umbilical structures in clinically normal foals. Am J Vet Res 1988;49: 2143–6. 11. Kablack KA, Embertson RM, Bernard WV, Bramlage LR, Hance S, Reimer JM, Barton MH. Uroperitoneum in the hospitalised equine neonate: retrospective study of 31 cases, 1988–1997. Equine Vet J 2000;32:505–8. 12. Hyman SS, Wilkins PA, Palmer JE, Schaer TP, Del Piero F. Clostridium perfringens urachitis and uroperitoneum in 2 neonatal foals. J Vet Intern Med 2002;16:489–93. 13. Rantanen NW. Diseases of the kidneys. Vet Clin N Am Equine Pract 1986;2:89–103. 14. James AE. The use of compound B-mode ultrasound in abdominal disease of animals. J Am Vet Radiol Soc 1976; 17:106–12.
15. Yamaga Y, Too K. Diagnostic ultrasound imaging in domestic animals: fundamental studies on abdominal organs and fetuses. Nippon Juigaku Zasshi 1984;46:203–12. 16. Penninck DG, Eisenberg HM, Teuscher EE, Vrins A. Equine renal ultrasonography: normal and abnormal. Vet Radiol 1986; 27:81–4. 17. Modransky PD. Ultrasound-guided renal and hepatic biopsy techniques. Vet Clin N Am Equine Pract 1986;2:115– 26. 18. Ramirez S, Williams J, Seahorn TL, Blas-Machado U, Partington BP, Valdes M, McClure JR. Ultrasound-assisted diagnosis of renal dysplasia in a 3-month-old Quarter Horse colt. Vet Radiol Ultrasound 1998;39:143–6. 19. Bayly WM, Elfers RS, Liggitt HD, Brobst DF, Gavin PR, Reed SM. A reproducible means of studying acute renal failure in the horse. Cornell Vet 1986;76:287–98. 20. Kiper ML, Traub-Dargatz JL, Wrigley RH. Renal ultrasonography in horses. Compendium: continuing education for veterinarians, article #8 1990;12:993–9. 21. Ehnen SJ, Divers TJ, Gillette D, Reef VB. Obstructive nephrolithiasis and ureterolithiasis associated with chronic renal failure in horses: eight cases (1981–1987). J Am Vet Med Assoc 1990;197:249–53. 22. Hope WD, Wilson JH, Hager DA, Garry MR, CalderwoodMays MB. Chronic renal failure associated with bilateral nephroliths and ureteroliths in a two-year-old Thoroughbred colt. Equine Vet J 1989; 21:228–31. 23. Rantanen NW. Ultrasound appearance of normal lung borders and adjacent viscera in the horse. Vet Radiol 1981;22:217–19. 24. Rantanen NW. Diseases of the liver. Vet Clin N Am Equine Pract 1986;2:105–14. 25. Reef VB, Johnston JK, Divers TJ, Acland H. Ultrasonographic findings in horses with cholelithiasis: eight cases (1985–1987). J Am Vet Med Assoc 1990;196:1836–40. 26. Reef VB, Collatos C, Spencer PA, Orsini JA, Sepesy LM. Clinical, ultrasonographic, and surgical findings in foals with umbilical remnant infections. J Am Vet Med Assoc 1989;195:69–72. 27. Reimer JM. Ultrasonography of umbilical remnant infections in foals. In: Proceedings of the 39th Annual American Association of Equine Practitioners 1993;247–248.
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Chapter 27
Antibiotic Therapy K. Gary Magdesian
Antimicrobials, hemodynamic support, and nursing care form the basis of sepsis therapeutics. This section will review prophylactic and therapeutic antimicrobial use for sepsis, the leading cause of morbidity and mortality in perinatal foals. It will also review antimicrobials available for use in neonatal foals.
Neonatal pharmacology The pharmacokinetic and pharmacodynamic physiology of neonatal foals is different from that of adult horses. Foals have increased enteral drug absorption, reflecting the fact that they are not yet hindgut fermenters.1 During the first 24 h of life the neonatal gastrointestinal tract is designed to absorb intact macromolecules, especially colostral immunoglobulins. This property likely also facilitates absorption of oral medications during this time period. Examples of antimicrobials with viable oral bioavailability in foals, in contrast to adult horses, include amoxicillin, ampicillin, and first-generation cephalosporins2, 3 (Table 27.1). Neonatal foals also have lower plasma protein concentrations than adult horses, which results in relatively higher free plasma drug concentrations as compared to adult horses.4 This is particularly true for highly protein-bound drugs, such as non-steroidal anti-inflammatory drugs. Another difference between foals and adult horses is that newborns have a higher percentage of body water as compared to adults, and concomitantly a lower body fat percentage.5 This increase in body water, especially of the extracellular fluid compartment, results in an increase in the volume of distribution of water-soluble drugs such as aminoglycosides. Therefore, to obtain the same desired peak plasma concentrations of amikacin or gentamicin as that in adult horses, the administered doses often must be increased. Neonatal humans also have an increased blood–brain barrier permeability, which may be true of neonatal foals as well, which favors movement of drugs across this barrier. This works as an advantage
in the treatment of bacterial meningitis. Neonatal foal urine is acidic in contrast to that of adult horses, resulting in ion trapping and increased elimination of basic drugs, such as trimethoprim, aminoglycosides, and macrolides.6 Conversely foals have reduced elimination of acidic drugs, including beta-lactam antimicrobials and sulfonamides. Particularly during the first 2 weeks of life, especially during the first few days postpartum, foals may have slightly reduced renal and hepatic metabolic function as compared to adult horses.7 This physiological difference results in a slight delay in the elimination of water- and lipidsoluble antimicrobials. For example, the elimination of ceftiofur increases as foals age from day 1 to day 7 of life, and then again as foals continue to mature to 1 month and beyond. Based on these results, it appears that the mechanisms involved in the metabolism and elimination (primarily renal) undergo rapid maturation during the first week of life in foals.8 Similarly, the elimination of chloramphenicol increases as foals mature from day 1 to day 3 and again to day 7 of life.7 This difference in metabolism and elimination is particularly significant for lipophilic drugs with lower therapeutic indices, such as metronidazole and chloramphenicol. The recommended doses of these drugs in neonatal foals are lower than for adults, and the administration interval is longer (Table 27.1). Finally, the growing physiology of the foal puts them at unique risk for side effects of medications that are not observed in adult horses. The best example of this is the fluoroquinolone-associated arthropathy noted in foals, which precludes the use of enrofloxacin in this age group.9
Recommendations for antimicrobial use in neonatal sepsis Based on retrospective reports, the most common microbes isolated from blood cultures or cultures
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 27.1 Antimicrobial doses in neonatal foals Antimicrobial agent Aminoglycosides: Amikacin Gentamicin Beta-lactams: Ampicillin (oral pivampicillin 19.9 mg/kg) Amoxicillin Penicillin, potassium or sodium Ticarcillin-clavulanic acid Imipenem – slow infusion and diluted First-generation cephalosporins: Cefazolin Cephalothin Cefadroxil Cephalexin Second-generation cephalosporins: Cefuroxime Cefoxitin Third-generation cephalosporins: Ceftazidime Cefotaxime Ceftizoxime Cefpodoxime proxetil Ceftiofur Fourth-generation cephalosporins: Cefepime Cefquinome Macrolides: Azithromycin Clarithromycin Erythromycin Other: Chloramphenicola Doxycycline Rifampin Metronidazole for enteric clostridiosis: Trimethoprim-sulfonamide a
Dosage
25 mg/kg i.v., q 24 h 6.6–12 mg/kg i.v., q 24 h 22–30 mg/kg i.v. or p.o., q 6–8 h 13–30 mg/kg p.o., q 8 h 22,000–30,000 IU/kg i.v., q 6 h 50 mg/kg i.v., q 6 h (dose based on ticarcillin) 15 mg/kg i.v., q 6–8 h 15–20 mg/kg i.v., q 6–8 h 10–20 mg/kg i.v., q 6 h 20–40 mg/kg p.o., q 8–12 h 25–30 mg/kg p.o., q 6–8 h 16–33 mg/kg i.v., q 8 h 20 mg/kg i.v., q 6 h 50 mg/kg i.v., q 6 h, slow infusion 40–50 mg/kg i.v., q 6 h, slow infusion 20–50 mg/kg i.v., q 6 h, slow infusion 10 mg/kg p.o., q 6–8–12 h 5–10 mg/kg i.v., q 12 h 11 mg/kg i.v., q 8 h 1 mg/kg IM, i.v. q 12 h 10 mg/kg p.o., q 24 h; q 48 h after 5 d 7.5 mg/kg p.o., q 12 h 15–25 mg/kg p.o., q 8 h 40–50 mg/kg p.o., q 12 h d1–2 of life; q 8 h d3–5; q 6 h >5 d 10 mg/kg p.o., q 12 h 5 mg/kg p.o., q 12 h 10–15 mg/kg p.o.,i.v. q 8–12 h 15–30 mg/kg p.o., q 12 h
Wear gloves when handling
of tissues at necropsy of septic foals include E. coli, Actinobacillus, Klebsiella, Enterobacter, Salmonella, Enterococcus, Streptococcus, and Staphylococcus.10–14 Based on these data both Gram-negative and -positive microbes are frequently involved with equine neonatal sepsis, and they often occur together as mixed infections. In addition, both enteric organisms, such as E. coli and enterococci, as well as non-enteric agents, including Actinobacillus and streptococci, are involved. In light of these findings, broad-spectrum antimicrobials are warranted in treating sepsis, particularly when culture and susceptibility results are pending or unavailable. Antimicrobial therapy of septic neonatal foals should begin with intravenous and bactericidal drugs. The intravenous route is important because of
the hemodynamic consequences of sepsis, consisting of hypotension and hypoperfusion. Perfusion to the subcutaneous tissues, muscles, and gastrointestinal tract is reduced during sepsis and septic shock due to myocardial depression, vasodilation or vasoconstriction, and hypovolemia. Therefore, absorption of drugs from these areas is suboptimal until perfusion disturbances are corrected. Bactericidal drugs should be instituted in preference to bacteriostatic drugs, because the septic foal is in a state of relative immunocompromise as compared to adult horses and healthy neonatal foals.15, 16 Duration of treatment of sepsis should be a minimum of 14 days, with localized infections requiring longer therapy of 4–8 weeks. Antimicrobials should be instituted in any foal with suspected sepsis,
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Antibiotic Therapy as results of blood cultures are not available for > 48 hours. In addition, blood cultures are only approximately 50% sensitive and should not be relied upon to define sepsis.11 Broad-spectrum coverage with antimicrobials is indicated based on the available data for microbiological results from septic foals. Gram-negative coverage is best achieved with aminoglycosides or thirdor fourth-generation cephalosporins. Enrofloxacin, though offering an excellent Gram-negative spectrum, should not be utilized in neonatal foals due to risks of quinolone-induced arthropathies.9 Another option for Gram-negative organisms, especially Pseudomonas, is ticarcillin-clavulanic acid combination (Timentin, GlaxoSmithKline, Research Triangle Park, NC). Gram-positive coverage for neonatal sepsis is provided through beta-lactam drugs (including both penicillins and cephalosporins). A few published retrospective reports have evaluated the susceptibility patterns of isolates from cases of equine neonatal sepsis.11, 13, 17, 18 In these studies the percentage of Gram-negative enteric bacteria such as E. coli, Klebsiella, Enterobacter, and Salmonella that were susceptible to amikacin was 83–100%. In contrast, only 67% of the isolates were susceptible to gentamicin in some of the studies. Their susceptibility to ceftiofur was highly variable (30–100%). For Gramnegative non-enteric microorganisms, including Actinobacillus and Pseudomonas, the percentages that were susceptible to amikacin and gentamicin were also highly variable, ranging from 57% to 100% for both drugs. The susceptibility to ceftiofur was 85–100% for Actinobacillus, but only 29% for Pseudomonas. Approximately 83–100% of Streptococcus isolates were susceptible to penicillin or ampicillin. Staphylococcus isolates tended to be highly drug-resistant, although 94–100% of those reported were susceptible to amikacin. Enterococci were also multidrug-resistant, with ampicillin susceptibility ranging from 25% to 100% depending on the study. Chloramphenicol was effective against 80% of enterococci and 93% of staphylococci in one study.11 Based on these reports, a combination of the aminoglycoside amikacin and a beta-lactam drug (ampicillin, penicillin, or first through third-generation cephalosporin) is a highly effective first-line therapeutic in foals with sepsis. Amikacin and ampicillin together provide adequate coverage against approximately 90% of the isolates recovered from foal blood cultures at the author’s institution (University of California, Davis). The shortcomings of this combination at the author’s institution would be with certain isolates of Pseudomonas (36% of isolates resistant to amikacin), for which ticarcillin would be the drug of choice, and some isolates of enterococci (species other than E. faecalis or E. faecium, which tend to be susceptible
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to ampicillin); these enterococci are generally highly susceptible to chloramphenicol or tetracycline. The addition of ampicillin (as opposed to penicillin G or cephalosporins) to amikacin has the slight advantage of increasing the spectrum to enterococci. Either ampicillin, ceftiofur, or a firstgeneration cephalosporin (as compared to penicillin G) would also increase the spectrum of activity against Actinobacillus spp, as its susceptibility to amikacin is somewhat variable (> 90% at the author’s institution, but lower in some published reports). For Pseudomonas aeruginosa, amikacin or ticarcillin/clavulanate (or both together) should be utilized as first-line drugs, especially until susceptibility results become available. This combination of amikacin and ampicillin is also synergistic against many bacteria. Another possible advantage of the amikacin/ampicillin combination is that amikacin alone or in combination with ampicillin appears less likely than beta-lactams (ceftiofur, ampicillin or imipenem alone) to induce endotoxin and TNFα synthesis during bactericidal activity against E. coli incubated with mononuclear cells in vitro.19 Because of the nephrotoxic potential of aminoglycosides, foals with renal failure should be treated with third- or fourth-generation cephalosporins rather than aminoglycosides. The pharmacokinetics of cefepime, a fourth-generation cephalosporin, have been studied in horses and foals.20 The activity of cefquinome, labeled for use in horses in some countries, against equine bacterial pathogens have been reported.21 These drugs are best reserved for foals with renal failure, severe infections, and those infected with bacteria resistant to first-line drugs.
Clinically-oriented review of specific antimicrobials for use in neonatal foals Aminoglycosides Aminoglycosides are concentration-dependent antimicrobials, referring to the direct relationship between peak plasma concentrations and rate and extent of bacterial killing. This property is conducive to single daily dosing, which maximizes efficacy and minimizes toxicity. The mechanism of action is through inhibition of protein synthesis by binding to the 30S ribosomal subunit. They are synergistic with beta-lactam drugs, because their cellular penetration is enhanced by cell wall synthesis inhibition. Additional mechanistic actions include interference with the electron transport system and translation, as well as DNA and RNA metabolism.22
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The major determinants of aminoglycoside efficacy are the ratios of Cmax (peak plasma concentration) and area under the curve (AUC) to the minimum inhibitory concentration (MIC) of the offending bacterium. A Cmax :MIC ratio of 8–10 is recommended for optimal efficacy. In addition to providing high peaks, once daily dosing allows for plasma concentrations to decrease below targeted trough concentrations, in order to minimize renal tubular and vestibulocochlear accumulation and nephrotoxicity or vestibular/cochlear toxicity. Allowance of plasma concentrations to decrease below the MIC of the microbes with single daily dosing has its basis in the post-antibiotic effect exerted by aminoglycosides. Neonatal foals have a higher volume of distribution for aminoglycosides as compared to adult horses because of higher total body water and extracellular fluid volumes. Therefore, they may require increased doses of drug as compared to adult horses (Table 27.1). In addition, the pharmacokinetics of amikacin have been demonstrated to change as foals age from 1 to 5 and 5 to 10 days of age.23 Clearance increases, whereas half-life, AUC, and plasma concentrations decrease with age, reflecting enhanced clearance of amikacin.23 This warrants therapeutic drug monitoring every few days for foals that are on prolonged therapy to ensure optimal efficacy (see below). Because their cellular uptake is oxygendependent, aminoglycosides are ineffective against anaerobic bacteria and facultative anaerobic bacteria existing under anaerobic conditions. They are most active in alkaline environments, and therefore acidity from tissue damage may cause drug failure. Purulent material inactivates the aminoglycosides, making undrained abscesses poor targets for aminoglycosides. Another factor for consideration of aminoglycoside activity is the highly polar nature, which is an impediment to intracellular distribution. Therefore, their activity against intracellular infections, such as salmonellosis, is limited despite in vitro efficacy. The primary toxicities of aminoglycosides include renal, vestibular, and cochlear toxicity. Amikacin is regarded as less nephrotoxic than gentamicin. Risk factors for toxicity include hypovolemia, pre-existing renal disease, prolonged therapy, and increased trough concentrations.22 Calcium supplementation, high protein, and high calcium diets may decrease the risk of toxicity, as calcium and protein compete with aminoglycoside binding to the tubular epithelium.22, 24 Aminoglycosides should be administered with caution in foals with hypocalcemia, as they may cause bradycardia, reduced cardiac output, and neuromuscular blockade. Because aminoglycosides induce neuromuscular block-
Table 27.2 Therapeutic drug monitoring of aminoglycosides Aminoglycoside
Peak (µg/mL)
Trough (µg/mL)
Gentamicin (µg/mL) Amikacin (µg/mL)
≥ 20 (or 10 × MIC) ≥ 40 (or 10 × MIC)
≤1 ≤ 1–2
ade, they are contraindicated for use in foals with botulism.25 In order to minimize toxicity, therapeutic drug monitoring (TDM) should be performed and serial monitoring of creatinine should be done every 3–5 days. Therapeutic drug monitoring allows for targeting optimal peak and trough aminoglycoside concentrations (Table 27.2). If cultures are not available, these peak concentrations can be targeted empirically (Table 27.2). Peak samples should be obtained after distribution is complete. The author utilizes 1hour peaks, although 0.5–1 h time points can be used. Trough samples should be obtained prior to the next dose. The author prefers a 20-h trough, to ensure at least a 4-hour washout period. Because the pharmacokinetics of amikacin change as foals grow, TDM should be performed every 3–5 days for foals on prolonged therapy (> 7 days).23 Additional aminoglycosides include neomycin, streptomycin, tobramycin and kanamycin. Because of its high nephrotoxic potential neomycin, which is the most toxic of the aminoglycoside drugs, is limited to topical use. Acquired resistance to streptomycin, in addition to relatively high toxicity, limit its use in foals. Kanamycin has largely been replaced for parenteral administration by more active and less toxic aminoglycosides, including gentamicin and amikacin. Tobramycin has similar antibacterial activity as gentamicin, but it is more expensive. It is less active than amikacin in terms of bacterial spectrum. However, tobramycin is more active than gentamicin against Pseudomonas spp. It has slightly reduced nephrotoxic potential as compared to gentamicin, but similar ototoxicity. The pharmacokinetics of tobramycin (4 mg/kg, i.v. q 24 h) have been studied in adult horses.26 However, the drug has not been studied in foals, and data regarding clinical use are lacking except for topical use in cases of infectious keratitis, especially those associated with Pseudomonas spp. Beta-lactam antibiotics Beta-lactam antibiotics belong to a broad class that includes any antimicrobial agent with a betalactam nucleus in its molecular structure such as penicillin derivatives (penams), cephalosporins (cephems), monobactams, and carbapenems. They are the most widely used antibiotics. The
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Antibiotic Therapy mechanism of action of beta-lactam antimicrobials is inhibition of bacterial cell wall synthesis through interference with peptidoglycan synthesis. They bind penicillin-binding proteins (PBPs), which are transpeptidases and peptidoglycan-active enzymes.27 The penicillin-binding proteins catalyze cross-linking of the glycopeptide polymers that form the cell wall. The variation in activity of the different penicillin drugs is due to varying affinity of the PBPs, the ability of the drugs to penetrate the outer cell membrane, and resistance to beta-lactamase enzymes.27 The beta-lactam drugs have very low toxicity. Potential adverse effects include anaphylaxis and other forms of hypersensitivities (urticaria, angioedema), hemolytic anemia and thrombocytopenia, and antibiotic-induced colitis (such as salmonellosis or clostridiosis). Allergies to penicillins are cross-reactive with other penicillins, however crossreactions to cephalosporins occur in only 5–10% of human patients with penicillin hypersensitivity. Beta-lactam drugs are time-dependent, bactericidal drugs. This means that, unlike the aminoglycosides, the beta-lactam drugs should be maintained above the MIC of the offending bacterium for as long as possible throughout the dosing interval. Ideally, concentrations 3–4 times the MIC of the offending bacterium should be achieved and maintained at all times.
Continuous rate infusions: new administration protocol for beta-lactam antibiotics Because they exhibit time-dependent bactericidal activity, the efficacy of beta-lactam drugs is optimized through maintenance of plasma (and tissue) concentrations of drug above the MIC for the entire dosing interval. This would require frequent dosing, or more ideally continuous rate infusions (CRI).28 This dosing administration is particularly advantageous for foals in an immunocompromised state, such as neutropenia and failure of passive transfer. Foals with sepsis are conducive to CRI delivery, as they tend to be recumbent and easily instrumented with fluid lines and automated pumps. Dosage calculations for CRI administration are as follows:
r
r
The same total daily dose is used as for intermittent administration. For example, if ampicillin is administered at a dose of 22 mg/kg i.v. q 6 h, then the CRI dose rate would be 22 × 4/24 = 3.7 mg/kg/h. If the desired plasma concentration of drug and clearance (Cl) of the drug are known, the CRI dose
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can be calculated (CRI dosing rate = Cl × Desired Plasma Concentration).29 The desired drug concentration is 4 times the MIC of the microbe. For example, if clearance of drug A is known to be 0.4 L/kg/h, and the MIC of the organism is 0.5 µg/mL (or 0.5 mg/L), the CRI dosing rate is = 0.4 L/kg/h × (4 × 0.5 mg/L) = 0.8 mg/kg/h. For drugs where clearance is unknown in foals, it can be calculated from published half-life (t1/2 ) and volume of distribution (Vd) values (Cl = [0.693 × Vd]/t1/2 ). Whenever CRI dosing is used, the drug stability at room temperature must be considered, and the drug changed often enough to ensure stable and active drug. Some drugs, such as ceftiofur, are lightsensitive upon reconstitution and should be covered.
Penicillins This class of drugs includes penicillin G, ampicillin, oxacillin, methicillin, piperacillin, carbenicillin, and ticarcillin. The spectrum of activity of penicillin G is primarily Gram-positive and includes Streptococcus, Bacillus, Actinomyces, and Corynebacterium. It has moderate activity against Actinobacillus and Pasteurella. As far as anaerobes, penicillin G has activity against Clostridium, most Fusobacterium isolates, and some Bacteroides (not B. fragilis). Ampicillin is similar to penicillin G in its spectrum, but has greater activity against Actinobacillus and Pasteurella, as well as Enterococcus spp. Ampicillin is slightly less active than penicillin G against Gram-positive microbes and anaerobes, however. Ampicillin (as pivampicillin) and amoxicillin can be administered orally in neonatal foals to treat Gram-positive infections, whereas they should not be used in adult horses due to low bioavailability and high risk of diarrhea.2, 30 Methicillin, oxacillin, and nafcillin are the ‘antistaphylococcal’ penicillins in that they are resistant to penicillinases produced by non-MRSA Staphylococcus aureus. They are less active than penicillin G against other penicillin-sensitive bacteria. It should be noted that nafcillin has been associated with renal failure in dogs, and the author (KGM) has noted a high rate of thrombophlebitis in horses.31 Carbenicillin and ticarcillin are ‘anti-pseudomonal’ penicillins. They are best used with beta-lactamase inhibitors (such as clavulanic acid as in Timentin) or in conjunction with amikacin, because of the high risk of Pseudomonas isolates developing resistance to antimicrobials during treatment. Ticarcillin also has some activity against E. coli and Proteus, although Klebsiella, Citrobacter, and Enterobacter tend to be resistant. 27 Piperacillin, azlocillin, and mezlocillin also have anti-pseudomonal activity, but also have increased
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activity against Klebsiella and Bacteroides fragilis. Penicillins commonly used in foals are listed in Table 27.1.
Cephalosporins Cephalosporins are categorized into first through fourth generations. In general, the first-generation cephalosporins are primarily Gram-positive drugs. The second-generation class has relatively broadspectrum activity against selected Gram-positive and -negative microorganisms. The third-generation drugs have reduced Gram-positive activity, but increased Gram-negative activity. The fourthgeneration cephalosporins have very broad and excellent (extended spectrum) Gram-positive and -negative activity. In general, cephalosporins do not have activity against MRSA and enterococci. In general terms, only the ‘antipseudomonal’ third-generation cephalosporins (ceftazidime and cefoperazone) and fourth-generation cephalosporins have activity against Pseudomonas aeruginosa. The cephalosporin with the most potent and broadspectrum anaerobic activity, including against Bacteroides fragilis, is the second-generation drug cefoxitin. Otherwise, in general terms, the cephalosporins have limited anaerobic activity similar to ampicillin. First-generation cephalosporins In general these are Gram-positive drugs, with anaerobic activity similar to ampicillin (no activity against B. fragilis). They are active against non-methicillin resistant Staphylococcus aureus. In contrast to the situation in adult horses, some of the first-generation cephalosporins have good oral bioavailability in foals. Diarrhea associated with oral cephalosporins is far less common in foals than in adult horses. Cefadroxil and cephalexin are two examples with excellent oral bioavailability in neonates (Table 27.1). Second-generation cephalosporins Examples of this subclass include cefoxitin, cefotetan, and cefuroxime. These have moderate activity against Gram-positive organisms, but with less potency than the first-generation cephalosporins. They have increased Gram-negative activity as compared to the first-generation drugs, although they do not have efficacy against Pseudomonas aeruginosa, many enteric microbes, and enterococci. Cefoxitin has the broadest activity against anaerobes, with very potent activity against Bacteroides fragilis, Porphyromonas, and Prevotella spp.27 Third-generation cephalosporins Ceftiofur, cefotaxime, ceftizoxime, cefpodoxime, and ceftazidime are examples of third-generation
cephalosporins. These have high resistance to beta-lactamases, giving them excellent Gramnegative coverage. They are very active against both Gram-positive and -negative microbes, including Streptococcus, most enterics, Actinobacillus and Pasteurella spp, as well as Clostridium and Fusobacterium spp. Bacteroides fragilis is generally considered resistant to the third-generation cephalosporins. The third-generation class has moderate activity against Staphylococcus, although not MRSA. Cefotaxime has the best anaerobe coverage of the third-generation cephalosporins, with some activity against B. fragilis. Ceftazidime in particular has poor anaerobic activity. Pseudomonas is resistant to most third-generation drugs, with the exception of the ‘antipseudomonal cephalosporins’, ceftazidime and cefoperazone. These two are slightly less active against other bacteria relative to the other third-generation cephalosporins. Most of the third-generation cephalosporins cross the blood–brain barrier (BBB) well, particularly ceftriaxone. This property, coupled with their broad spectrum of activity, makes them excellent therapeutics for bacterial meningitis in foals. Ceftiofur does not share this property of crossing the BBB to the same extent as other third-generation drugs. Ceftiofur is the most common third-generation cephalosporin used in horses and foals. Its MICs should be interpreted with caution for Proteus and Staphylococcus, however, as its intermediate metabolite, desfuroylceftiofur, is less active than the parent drug against these two bacteria in particular.27 Ceftiofur may therefore not be appropriate for treatment of infections caused by these microbes. Otherwise, the intermediate is similar in its antibacterial spectrum to ceftiofur. Ceftiofur has a wide dose range depending on the targeted bacteria. For Streptococcus, Actinobacillus and Pasteurella, doses of 2.2 mg/kg i.m., i.v. or s.c. q 24 h can be used. Bacteria with higher MIC values, however, should be treated with higher doses (5–10 mg/kg i.m., i.v. or s.c. q 12 h for neonates). These include Gram-negative enteric microorganisms. A recent study of the pharmacokinetics of ceftiofur in foals concluded that a dose rate of 5 mg/kg every 12 h would provide sufficient concentrations for the treatment of 90% of tested isolates of E. coli, Pasteurella spp, Klebsiella spp, and beta-hemolytic streptococci, 88% of Enterobacter spp and 73% of Salmonella spp of equine origin; most Enterococcus spp were resistant to ceftiofur.32 It should be noted that CSF concentrations of ceftiofur and its active metabolite, desfuroylceftiofur, were quite low in this study. A recent study has shown that the susceptibility of all blood culture isolates from sick neonatal foals to ceftiofur was 77%.33
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Antibiotic Therapy Recently, the pharmacokinetics of cefpodoxime were reported for foals.34 This was the first oral thirdgeneration cephalosporin made available for use in foals. The recommended dose is 10 mg/kg every 6–12 h, with the more frequent dosing interval for E. coli or Salmonella infections.34 Fourth-generation cephalosporins The fourth-generation cephalosporins exhibit high activity against Gram-negative microbes, including enterics. They also have excellent efficacy against non-MRSA staphylococci and streptococci. They have only moderate activity against Pseudomonas aeruginosa. Their anaerobic coverage is only moderate, being effective only against C. perfringens and Peptostreptococcus. They are ineffective against enterococci and Bacteroides spp. The pharmacokinetics of cefepime in horses and foals have been described.20 Recently, another fourthgeneration cephalosporin, cefquinome, has been licensed for use in horses in the United Kingdom. It is approved for use with equine respiratory tract disease and foal septicemia. In an in vitro study, cefquinome showed high activity against Actinobacillus equuli, streptococci, enteric bacteria, and staphylococci.21 It should be noted that the study did not define whether the tested Staphylococcus spp isolates were methicillin-resistant. The activity of cefquinome against Rhodococcus spp and Pseudomonas spp was limited, however. Clinical use of cefquinome in horses and foals has been reported in the literature.35, 36 A field study of cefquinome (1 mg/kg q 12 h, i.v. for 3 days, then i.m. for 3–11 days) was performed in 39 foals with infections such as pneumonia, gastroenteritis, arthritis, omphalitis, or wound infections.35 Blood cultures were positive in 40.5% of cases, with E. coli, Clostridium perfringens and Staphylococcus spp being the most common isolates, and all were susceptible to cefquinome. In this study treatment was successful in terms of survival in 87.2% of cases, and average treatment duration was 7.5 days. The product was well tolerated by foals. In a second study, which was a randomized clinical trial of prophylactic and therapeutic use of cefquinome versus a penicillin/gentamicin combination in adult horses, there were no differences in terms of efficacy in the prophylactic group.36 However, in the treatment group (n = 141), there were statistically fewer complications in wound healing (2% vs 12.7%) and fewer changes in antibiotic treatment due to apparent treatment failure (0 vs 9.7%) in the cefquinome group as compared to the penicillin/gentamicin group. There were no statistical differences in the rate of diarrhea or thrombophlebitis between groups, however 2/197 horses in the study developed diarrhea and thrombophlebitis in the ce-
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fquinome group as compared to 0/190 in the penicillin/gentamicin group.36
Carbapenems Carbapenems are a class of modified penicillins, with the broadest spectrum of activity of any commercially available antimicrobial. They are highly effective against both Gram-positive and -negative microbes. These include imipenem-cilastatin, meropenem and biapenem, with imipenem-cilastatin having been administered to foals and studied in horses.37 The spectrum of imipenem covers both Grampositive and -negative bacteria, including most enterococci, Nocardia spp and anaerobes, including B. fragilis. They are highly effective against Gram-negative microbes, including enterics and Pseudomonas spp. Resistant microorganisms include MRSA, some Enterobacter spp isolates, some Pseudomonas isolates, and Enterococcus faecium. In human patients, isolates of Pseudomonas can develop resistance to imipenem rapidly; therefore, it is recommended to combine imipenem with an aminoglycoside for synergy and to minimize the emergence of resistance. Side effects of imipenem in foals include rare seizures, antibiotic-induced diarrhea, and thrombophlebitis if it is not diluted properly. The carbapenem antimicrobials should not be used as first-line therapy. Rather, they should be reserved for foals with resistant bacteria and severe infections with no other antimicrobial options available. Subacute to chronic therapy with antimicrobials in foals When oral drugs are needed for longer-term therapy in the subacute to chronic stages of disease treatment (after initial therapy with intravenous bactericidal drugs), additional antimicrobials are available for use in foals. In general, these antimicrobials should only be utilized when susceptibility data support their use as their spectra are limited. Potentiated sulfonamides (trimethoprim-sulfa; ‘TMS’ or ‘SMZ’) are moderately broad-spectrum antimicrobials, however they cannot be recommended as first-line therapies for use in foals with sepsis because of resistance by many enteric isolates and streptococci. Rather, they can be used to extend the duration of antimicrobials when MIC data become available. Foals on long-term TMS therapy should be monitored for anemia and neutropenia as markers of bone marrow suppression. Tetracyclines are broad-spectrum, but are not optimal for sepsis because they are bacteriostatic rather
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than bactericidal, and lack activity against many Gram-negative enteric microbes. They are useful for Mycoplasma infections, which can cause polyarthritis in foals. Tetracyclines can discolor teeth in young animals; they can also cause renal disease if outdated or improperly stored due to the presence of degradation products. Renal failure is also a risk when supra-antimicrobial doses (50–60 mg/kg) are used, as for contracted tendons.38 In healthy foals one large dose of oxytetracycline did not cause renal toxicity in one study.39 Foals with concurrent illnesses, which are often concomitant to flexural deformities, are at increased risk for nephrotoxicity. If foals have difficulty rising to nurse they may develop dehydration and are therefore at increased risk for renal injury. When tetracycline is used for this purpose, creatinine should be monitored closely and i.v. fluids should be administered concurrently. Doxycycline has experienced increased use in equine medicine in recent years, and has largely replaced other tetracyclines for use as an antimicrobial in foals. Doxycycline is more lipophilic and penetrates tissues well. It is safer for teeth and bones in growing animals, and does not appear to be nephrotoxic. The pharmacokinetics of oral doxycycline in foals have been studied.40 Based on this study, a dose of 10 mg/kg every 12 h orally would maintain serum, pulmonary (bronchoalveolar) cell and fluid, peritoneal and synovial fluid, and urine concentrations above the MIC of many foal pathogens, including Rhodococcus equi and beta-hemolytic streptococci. It should be noted that the clinical efficacy of doxycycline in treating R. equi infections has not been evaluated. Intravenous doxycycline should be avoided in horses and foals due to reports of cardiovascular collapse and death, and it should only be used orally.41 Doxycycline also has a number of additional effects independent of its antibiotic properties. It has anti-inflammatory, antifibrotic, anti-apoptotic, and immunosuppressive properties, which are currently the subjects of much research.42 One note of caution is that foals on long-term doxycycline therapy should be monitored for distal limb laxity, although this has not been well documented. Chloramphenicol is another broad-spectrum oral drug available for use in horses and foals; however, it is bacteriostatic and has human health risk considerations (aplastic anemia), though rare. It has excellent spectrum against enterococci and even many MRSA isolates. Liver enzymes should be monitored in foals on extended courses of chloramphenicol. Because of age-dependent pharmacokinetics, foals < 3 days of age may be administered lower doses or at more prolonged intervals, as the half-life is longer in this age group compared to those > 7 days of age.7 Due to po-
tential health hazards, handlers should wear gloves whenever handling chloramphenicol, and should not crush pills. Rifampin is very effective against many staphylococci and Gram-positive microbes, and penetrates tissues and cell membranes well. It has moderate activity against Actinobacillus and Pasteurella, and very good activity against anaerobes, including Bacteroides fragilis. Enteric microbes are resistant to rifampin. Rifampin should not be used alone, due to ready development of resistance. Rifampin can be paired with beta-lactam antimicrobials, macrolides or TMS, depending on the type of infection being treated. The risk of enterocolitis and diarrhea increases with multiple administered antimicrobials. It should be noted that rifampin may discolor urine. Macrolides and azalides have been well studied for use in foals with Rhodococcus equi infections. However, they can also be used in neonatal foals with compartmentalized or localized infections with susceptible microbes. Azithromycin, an azalide, and clarithromycin, a macrolide, have excellent Gram-positive activity; azithromycin also is effective against some Gram-negative microbes such as Pasteurella, and has moderate activity against some salmonellae isolates. However, in general most enteric microbes are resistant to azithromycin and macrolides. In general, macrolides do not exhibit anaerobic activity, however azithromycin is effective against Fusobacterium. Traditionally erythromycin has been paired with rifampin for treatment of R. equi infections. However, erythromycin does not concentrate intracellularly as well as azithromycin or clarithromycin. In one retrospective study, clarithromycin in combination with rifampin was more effective than azithromycin/rifampin or erythromycin/rifampin; however, this was not a randomized clinical trial.43 That study found a tendency toward higher incidence of diarrhea in foals treated with clarithromycin (5/18) as compared to those treated with azithromycin (1/20). A disadvantage of erythromycin is that it is associated with relatively high risks of hyperthermia, respiratory distress syndrome, and diarrhea.44 These risks appear to be less common with clarithromycin and azithromycin. The macrolides penetrate tissues very well and concentrate intracellularly. This property makes them very useful in treating walled-off infections, such as umbilical abscesses. They can be used in combination with aminoglycosides when broad-spectrum coverage is desired. Side effects include diarrhea and hyperthermia. Clarithromycin, like erythromycin, has prokinetic properties. In the author’s experience, azithromycin is less likely to cause diarrhea than clarithromycin or erythromycin. In addition, the macrolides have antibiotic-independent
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Antibiotic Therapy anti-inflammatory effects in the lungs, including inhibition of neutrophil infiltration and chemotaxis. In a study comparing the cellular disposition of azithromycin, clarithromycin and erythromycin each at a dose of 10 mg/kg, the peak drug activity in pulmonary lining fluid was significantly higher in foals treated with clarithromycin than azithromycin, and erythromycin was only detected in the pulmonary fluid of 2/6 foals.45 Peak drug activity in brochiolar lavage (BAL) cells was not different between azithromycin and clarithromycin, but both were significantly higher than erythromycin. The concentrations of azithromycin and clarithromycin were higher in BAL cells, followed by pulmonary lining fluid, as compared to serum, signifying pulmonary concentration of these drugs. In contrast, erythromycin activity in BAL cells was not higher than that of serum. Metronidazole has excellent activity against anaerobic bacteria, and is very useful for treatment of Clostridium difficile or C. perfringens enteritis in foals. Side effects of metronidazole include anorexia acutely, whereas neurological signs, peripheral neuropathies, and hepatopathies can occur with prolonged use. Because of probable increased oral bioavailability in neonatal foals, the author uses a lower dose as compared to adult horses (10 mg/kg p.o. q 12 h).
References 1. Duffee NE, Stang BE, Schaeffer DJ. The pharmacokinetics of cefadroxil over a range of oral doses and animal ages in the foals. J Vet Pharmacol Ther 1997;20(6):427–33. 2. Love DN, Rose RJ, Martin IC, Bailey M. Serum levels of amoxycillin following its oral administration to thoroughbred foals. Equine Vet J 1981;13(1):53–5. 3. Baggot JD, Love DN, Stewart J, Raus J. Bioavailability and disposition kinetics of amoxicillin in neonatal foals. Eq Vet J 1988;20(2):125–7. 4. Bauer JE, Harvey JW, Asquith RL, McNulty PK, Kivipelto J. Serum protein reference values in the foals during the first year of life: comparison of chemical and electrophoretic methods. Vet Clin Pathol 1985;14(1):14–22. 5. Spensley MS, Carlson GP, Harrold D. Plasma, red blood cell, total blood, and extracellular fluid volumes in healthy foals during growth. Am J Vet Res 1987;48(12):1703–7. 6. Edwards DJ, Brownlow MA, Hutchins DR. Indices of renal function: values in eight normal foals from birth to 56 days. Aust Vet J 1990;67(6):251–4. 7. Adamson PJ, Wilson WD, Baggot JD, Hietala SK, Mihalyi JE. Influence of age on the disposition kinetics of chloramphenicol in equine neonates. Am J Vet Res 1991;52(3):426–31. 8. Wilson WD, Mihalyi JE. Comparative pharmacokinetics of ceftiofur in neonatal foals and adult horses. Dorothy R. Havemeyer Foundation Neonatal Septicemia Workshop II. Boston, MA, 1998. 9. Vivrette SL, Bostian A, Bermingham E, Papich MG. Quinolone-induced arthropathy in neonatal foals. In: Proceedings of the 47th Annual Convention of the American Association of Equine Practitioners, 2001; pp. 376–7.
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10. Koterba AM, Brewer BD, Tarplee FA. Clinical and clinicopathological characteristics of the septicaemic neonatal foal: review of 38 cases. J Vet Int Med 1984;16: 376–82. 11. Wilson WD, Madigan JE. Comparison of bacteriologic culture of blood and necropsy specimens for determining the cause of foal septicemia: 47 cases (1978–1987). J Am Vet Med Assoc 1989;195: 1759–63. 12. Raisis AL, Hodgson JL, Hodgson DR. Equine neonatal septicaemia: 24 cases. Aust Vet J 1996;73: 137–40. 13. Marsh PS, Palmer JE. Bacterial isolates from blood and their susceptibility patterns in critically ill foals: 543 cases (1991–1998). J Am Vet Med Assoc 2001;218: 1608–10. 14. Stewart AJ, Hinchcliff KW, Saville WJ, Jose-Cunilleras E, Hardy J, Kohn CW, Reed SM, Kowalski JJ. Actinobacillus sp bacteremia in foals: clinical signs and prognosis. J Vet Int Med 2002;16: 464–71. 15. Demmers S, Johannisson A, Grondahl G, Jensen-Waern M. Neutrophil function and serum IgG in growing foals. Eq Vet J 2001;33(7):676–80. 16. McTaggart C, Penhale J, Raidala SL. Effect of plasma transfusion on neutrophil function in healthy and septic foals. Aust Vet J 2005;83(8):499–505. 17. Brewer B, Koterba A. Bacterial isolates and susceptibility patterns in foals in a neonatal intensive care unit. Comp Cont Ed for Pract Vet 1990;12: 1773–81. 18. Henson S, Barton M. Bacterial isolates and antibiotic susceptibility patterns from septicemic neonatal foals: One 15 year retrospective study (1986–2000). Dorothy Havemeyer Foundation Third Neonatal Septicemia Workshop, 2001; pp. 50–52. 19. Bentley AP, Barton MH, Lee MD, Norton NA, Moore JN. Antimicrobial-induced endotoxin and cytokine activity in an in vitro model of septicemia in foals. Am J Vet Res 2002;63(5):660–8. 20. Gardner SY, Papich MG. Comparison of cefepime pharmacokinetics in neonatal foals and adult dogs. J Vet Pharmacol Ther 2001;24(3):187–92. 21. Thomas E, Thomas V, Wilhelm C. Antibacterial activity of cefquinome against equine bacterial pathogens. Vet Microbiol 2006;115(1–3):140–47. 22. Dowling PM. Aminoglycosides. In: Giguere S, Prescott JF, Baggot JD, Walter RD, Dowling PM (eds) Antimicrobial Therapy in Veterinary Medicine, 4th edn. Ames: Blackwell Publishing, 2006; pp. 207–29. 23. Magdesian KG, Wilson WD, Mihalyi J. Pharmacokinetics of a high dose of amikacin administered at extended intervals to neonatal foals. Am J Vet Res 2004;65(4):473–9. 24. Brashier MK, Geor RJ, Ames TR, O’Leary TP. Effect of intravenous calcium administration on gentamicin-induced nephrotoxicosis in ponies. Am J Vet Res 1998;59(8):1055–62. 25. Torda, T. The nature of gentamicin-induced neuromuscular block. Br J Anaesth 1980;52: 325–9. 26. Hubenov H, Bakalov D, Krastev S, Yanev S, Haritova A, Lashev L. Pharmacokinetic studies on tobramycin in horses. J Vet Pharmacol Therap 2007;30: 353–7. 27. Prescott JF. Beta-lactam antibiotics: cephalosporins. In: Giguere S, Prescott JF, Baggot JD, Walter RD, Dowling PM (eds) Antimicrobial Therapy in Veterinary Medicine, 4th edn. Ames: Blackwell Publishing, 2006; pp. 139–57. 28. Mouton JW, Vinks AA. Continuous infusion of betalactams. Curr Opin Crit Care 2007;13: 598–606. 29. Papich MG. Improving therapy and avoiding adverse reactions in horses through application of pharmacological principles. In: Proceedings of the 25th Forum of the American College of Veterinary Internal Medicine, 2007; pp. 186–8.
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30. Ensink JM, Barneveld A, Klein WR, van Miert AS, Vulto AG. Oral bioavailability of pivampicillin in foals at different ages. Vet Q 1994;16(Suppl 2):S113–16. 31. Pascoe PJ, Ilkiw JE, Kass PH, Cowgill LD. Case-control study of the association between intraoperative administration of nafcillin and acute postoperative development of azotemia. J Am Vet Med Assoc 1996;208(7):1043–7. 32. Meyer S, Giguere S, Rodriguez R, Zielinski RJ, Grover GS, Brown SA. Pharmacokinetics of intravenous ceftiofur sodium and concentration in body fluids of foals. J Vet Pharmacol Ther 2009;32(4):309–16. 33. Sanchez LC, Giguere S, Lester GD. Factors associated with survival of neonatal foals with bacteremia and racing performance of surviving thoroughbreds: 423 cases (1982–2007). J Am Vet Med Assoc 2008;233: 1446–52. 34. Carrillo NA, Giguere S, Gronwall RR, Brown MP, Merritt KA, O’Kelley JJ. Disposition of orally administered cefpodoxime proxetil in foals and adult horses and minimum inhibitory concentration of the drug against common bacterial pathogens of horses. Am J Vet Res 2005;66(1):30–5. 35. Rohdich N, Zschiesche E, Heckeroth A, Wilhelm C, Leendertse I, Thomas E. Treatment of septicaemia and severe bacterial infections in foals with a new Cefquinome formulation: a field study. Dtsch Tierarztl Wochenschr 2009;116(9): 316–20. 36. Widmer A, Kummer M, Wehrli Eser M, Furst A. Comparison of the clinical efficacy of Cefquinome with the combination of penicillin G and gentamicin in equine patients. Eq Vet Educ 2009;21(8):430–5. 37. Orsini JAA, Moate PJ, Boston RC, Normal T, Engiles J, Benson CE, Poppenga R. Pharmacokinetics of
38.
39.
40.
41.
42.
43. 44.
45.
imipenem-cilastatin following intravenous administration in healthy adult horses. J Vet Pharmacol Ther 2005;28(4):355– 61. Vivrette S, Cowgill LD, Pascoe J, Suter C, Becker T. Hemodialysis for treatment of oxytetracycline-induced acute renal failure in a neonatal foal. J Am Vet Med Assoc 1993;203(1):105–7. Wright AK, Petne L, Papich MG, Archer J, Fretz PB. Effect of high dose oxytetracycline on renal parameters in neonatal foals. Proc Am Assoc Eq Pract 1992;38: 297–8. Womble A, Giguere S, Lee EA. Pharmacokinetics of oral doxycycline and concentrations in body fluids and bronchoalveolar cells of foals. J Vet Pharmacol Therap 2007;30: 187–93. Riond Jl, Riviere JE, Duckett WM, Atkins CE, Jernigan AD, Rikihisa Y, Spurlock SL. Cardiovascular effects and fatalities associated with intravenous administration of doxycycline to horses and ponies. Eq Vet J 1992;24(1):41– 5. Sapadin AN, Fleischmjer R. Tetracyclines: nonantibiotic properties and their clinical implications. J Am Acad Dermatol 2006;54: 258–65. Giguere S, Jacks S, Roberts GD, Hernandez J, Long MT, Ellis C. J Vet Intern Med 2004;18(4):568–73. Stratton-Phelps M, Wilson WD, Gardner IA. Risk of adverse effects in pneumonic foals treated with erythromycin versus other antibiotics: 143 cases (1986–1996). J Am Vet Med Assoc 2000;217(1):68–73. Suarez-Mier G, Giguere S, Lee EA. Pulmonary disposition of erythromycin, azithromycin, and clarithromycin in foals. J Vet Pharmacol Ther 2007;30(2):109–15.
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Chapter 28
Enteral and Parenteral Nutrition for the Neonatal Foal Wendy E. Vaala
Any feeding program for foals must address the neonate’s degree of maturity, requirements for growth, special nutritional needs created by concurrent illness, and a thorough understanding of nutritional formulas and routes of administration available. Recognition of the serious adverse effects associated with malnutrition and inadequate nutrition reinforces the importance of evaluating nutritional support in the treatment plan for any compromised neonate. Not only is the diet formulation used important, but so is route of administration. While parenteral nutrition often represents the only feasible route for many critically ill neonates with dysfunctional gastrointestinal tracts, enteral feeding remains the preferred avenue for nutrition. Recent clinical trials have demonstrated that the complete absence of enteral feeding has immunosuppressive consequences through downregulatory effects on the gastrointestinal-associated mucosal immune system.1 This chapter will review general nutritional requirements for the healthy foal, while also presenting some of the nutritional challenges posed by the orphaned and/or abnormal foal suffering from common neonatal disorders including septicemia, peripartum asphyxia, and dysmaturity. The indications, complications and guidelines for designing and implementing enteral and parenteral nutrition programs will be reviewed. A complete in-depth review of this topic is beyond the scope of this chapter. Readers are encouraged to refer to other detailed discussions of enteral and parenteral feeding strategies for healthy and sick foals.2–7
Potential consequences of malnutrition Recognition of and respect for the potentially lifethreatening effects of malnutrition and inadequate nutrition serve to emphasize the vital role nutritional support plays in the treatment of compromised foals. Malnourishment due to inadequate intake, abnormal absorption, or increased utilization results in multiple organ system dysfunction and increased morbidity and mortality. The most immediate consequence of severe underfeeding in the newborn foal is hypoglycemia, while the more devastating, long-term consequence of malnutrition is decreased resistance to infection, which may occur as a result of depressed bone marrow activity, lymphoid tissue atrophy, abnormal neutrophil function, impaired antibody production, impaired hepatic function, and depressed secretory, mucosal and cell-mediated immunity.8–10 Inadequate nutrition results in gastrointestinal atrophy and dysmotility, impaired digestion and absorption, and hepatic dysfunction.11, 12 Altered hepatic function, including decreased hepatic cytochrome P450 activity, reduces resistance to a variety of toxins and bacterial and viral pathogens. Gastrointestinal atrophy, often associated with malnutrition and lack of enteral feeding in particular, is accompanied by loss of intestinal mucosal integrity and the increased risk of bacterial translocation and secondary septicemia. Malnutrition in general has been associated with weight loss, depressed growth rates, impaired wound healing, gastrointestinal atrophy, impaired digestion and absorption of fats and carbohydrates,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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and depressed immune function.13 Nutritionally depleted patients experience compromised pulmonary function and reduced ventilatory capacity due to a loss of strength of diaphragmatic, intercostal, and abdominal muscles.14 External signs of malnourishment are fairly universal and include weight loss, generalized weakness, lethargy, glossitis, sunken eyes, and dry or scaly skin with reduced elasticity.
Broodmare nutrition Although the focus of this chapter is foal nutrition, the dam’s nutrition during late pregnancy has an important impact on fetal development. The mare’s nutritional needs increase during the last 3 months of pregnancy and reach a peak during early lactation (Table 28.1). The mare’s prepartum diet may affect placental efficiency, foal birthweight and metabolism, colostral immunoglobulin (IgG) concentration, and milk composition.16 For example, mares receiving calciumdeficient diets, prior to delivery and during lactation, raised foals that had thinner and weaker bones than foals from mares maintained on an adequate diet.17 Mares fed a diet high in soluble carbohydrates late in gestation contributed to a decrease in insulin sensitivity in their foals as weanlings.18 Maternal copper and zinc deficiencies have been implicated as a cause of developmental orthopedic disease (DOD) in growing foals (see Chapter 86 for detailed discussion). Hepatic stores become an important source of these minerals for young foals since the mineral content of mare milk is so low.19, 20 The fetal liver accumulates minerals during late gestation. Adequate supplementation of broodmares during the last trimester may help prevent certain mineral deficiencies in the neonate,21 while excessive supplementation with some minerals, such as iodine, in late-term mares may produce foals suffering from abnormal conditions such as neonatal goiter.22
Normal foal physiology During late gestation, the fetal foal has minimal glycogen stores and limited glucogenic capacity com-
pared with other domestic species.23 Foals are born with relatively few energy reserves. In the absence of enteral feeding, meager hepatic glycogen reserves will support metabolism and maintain body temperature for only a few hours, which explains why hypoglycemia develops rapidly in newborn foals that fail to nurse soon after birth. It is estimated that energy supplied by ingested colostrum lasts fewer than 20 hours. Following delivery the neonate must transition from having a constant intravenous infusion of nutrients provided via the placenta to oral ingestion, digestion, and absorption of intermittent milk meals. In order to use the relatively high carbohydrate load in mare’s milk the neonatal foal relies on appropriate endogenous production of insulin by the pancreas during the early postpartum period. Endocrine maturation, in particular maturation of pancreatic β-cell function and insulin responsiveness, develops late in gestation in the equine fetus.24 Immediately after delivery, the foal’s glucoregulatory mechanisms may not be fully functional. During the early postnatal period, pony foals demonstrated reduced glucose clearance following glucose administration compatible with mild insulin resistance.25 Due to the immaturity of the large bowel, neonatal foals rely on small intestinal digestion of starch and carbohydrates and utilize milk lactose as their primary energy source. The development of brush border disaccharidase activity in the equine small intestine follows a pattern similar to other mammals, and increases to adult levels by 7 months of age.26 Lactase, a β-galactosidase, is the predominant disaccharidase in the equine small intestine and reaches maximal levels at birth and declines steadily after 4 months of age. In contrast, the α-galasctosidases, maltase, sucrase and trehalase, are barely discernible in the equine fetus. All three enzymes increase slowly after birth to reach adult levels by 7 months of age. Glucoamylase is detectable during the first week of life and attains adult levels by 10 months. This pattern of disaccharidase development, and results of oral disaccharide absorption studies,27 suggests that newborn foals would be incapable of maintaining acceptable growth on a diet containing sucrose,
Table 28.1 Minimum nutrient requirements for broodmares: late gestation through early lactation. From National Research Council (2007)15
Stage of gestation Last 90 days of gestation Stage of lactation 0–3 months 3 months to weaning
DE (Mcal/kg)
CP (%)
Ca (%)
Phos (%)
Cu (ppm)
Zn (ppm)
Vit A (IU/kg)
Vit E (IU/kg)
2.25
10
0.43–0.45
0.32–0.35
10
40
636
80
2.6 2.45
12.5–13.2 11
0.52 0.36
0.34 0.22
10 10
40 40
523 455
80 80
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Enteral and Parenteral Nutrition for the Neonatal Foal maltose or polysaccharides as the primary carbohydrate source, a fact that should be kept in mind when selecting milk replacers. Coprophagy is normal in young foals and occurs once every few hours during the first week of life and decreases in frequency during the first 3 months. The rate of coprophagy is highest during the first 8 weeks.28 This behavior may play a role in the bacterial inoculation of the gastrointestinal tract and the acquisition of vital nutrients, vitamins and enzymes, and does not indicate dietary deficiencies. The most common source of feces is usually the mare. Some studies indicate that foals actively seek out the feces of their own dams, which suggests a response to a maternal pheromone signaling the presence of deoxycholic acid or other essential acids the foal may be deficient in and which it may require for intestinal immunocompetence and myelination of the nervous system.28, 29
Nutrient requirements of healthy neonatal foals The actual nutrient requirements of nursing foals have not been determined. Estimates of the neonate’s nutritional requirements often are based on observing foal nursing behavior, estimating the volume of milk consumed and/or calculating the amount of milk produced by various breeds of lactating mares. Average daily milk consumption is then translated into total calories, protein, and lipids consumed based on the composition of mare’s milk. Unfortunately, the composition of mare’s milk varies with stage of lactation, breed, diet, and age, as well as parity of the dam, which makes calculating a healthy foal’s nutrient requirements a daunting task. For example, mares receiving a diet rich in forage compared with mares receiving a diet rich in concentrate had higher fat, protein and linolenic acid, and lower linoleic acid concentrations.30 As lactation progresses, mare’s milk decreases in total protein, total solids, fat, and gross energy.19, 31 Healthy newborn foals nurse on average 4 to 7 times an hour during the first week of life, consume 20 to 30% of their bodyweight in mare’s milk daily, and gain an average of 1 to 3 lb/day (0.5–1.5 kg/day) or as much as 2.5% of bodyweight per day during the early neonatal period. A healthy newborn Thoroughbred foal consumes an estimated 450 mL of milk per hour. Most horse foals’ colostrum ingestion averages between 2 and 5 liters.32 Light breed mares produce approximately 3% of their bodyweight in milk/day during the first 3 months of lactation. This coincides with the healthy, term foal consuming approximately 10 to 13 L of milk/day for light breed foals and 14 to 17 L of milk/day for warmblood or draft foals. Mare’s
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milk contains approximately 480–600 kcal/L. Based on the energy, protein, and fat provided by mare’s milk, a healthy nursing foal consumes an average of 120 to 150 kcal/kg/day; 5 to 7 g protein/kg/day, and 4 to 5 g fat/kg/day. These values are generally used when designing maintenance enteral feeding programs for healthy, including orphaned, foals. The foal’s highest energy needs are reached during the first week of life. By 5 weeks of age a foal’s requirements have decreased to 98 kcal/kg/day gross energy and 3.7 g crude protein/kg/day.31 Another method of determining energy requirements uses the following equation to determine basal energy requirements: (Wt in kg).75 × 70 (species factor for mammals) = basal kcal/24 h. For a 45 kg foal this calculation is 1216 kcal/day. An adjustment must be made for activity, growth, and disease. In humans this factor is considered 4 for healthy juveniles and 5 for sick juveniles. Using these factors, a healthy 45 kg foal would require 4864 kcal daily (108 kcal/kg/ day).33 Estimating trace mineral and vitamin requirements is even more speculative. Rooney34 estimated mineral requirements for newborn foals based on the estimated daily intake of certain trace minerals from mare’s milk: iron 10 mg/day; copper 7.0 mg/day; manganese 17 mg/day; zinc 40 mg/day; selenium 0.09 mg/day. Neonatal foals have higher trace mineral requirements than adult horses, yet all trace minerals are low in mare’s milk.20, 35 Even healthy neonatal foals seem to live on the brink of copper, zinc, and iron deficiencies during early growth.36, 37 Reliance on prenatally acquired mineral reserves and increased efficiency of absorption are the likely mechanisms by which nursing foals avoid mineral deficiencies until the ingestion of solid food begins. Controversy continues regarding the efficacy of oral mineral supplementation in young foals to correct mineral deficiencies and prevent DOD.38 One commercially available product containing trace minerR (Buckeye Feeds, Dayals and vitamins is Foal Aide ton, OH). Various guidelines exist for copper and zinc supplementation and are discussed in more detail in Chapter 86. In general, foals should receive: Cu – 50 ppm of DM (dry matter); Zn – 60 ppm of DM. In addition to its role in bone development, zinc status has also been linked to immune function. Early hallmarks of zinc deficiency include lymphopenia and thymic atrophy.39 Selenium deficiency has been associated with myopathies, steatitis, and depression of the immune system with increased susceptibility to diarrhea and respiratory disease. A growing foal’s selenium requirement is estimated to be 0.16–0.2 mg/kg of dietaryDM.15 Nursing foals ingest approximately 250 to 290 mg/kg of calcium daily. Since the calcium
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retention rate in foals and the intestinal absorption coefficient of calcium in mare’s milk are not known, daily oral calcium requirements can only be estimated. One source suggests providing neonatal foals with 18 g/day of calcium and 9.0 g/day of phosphorus.34 It is recommended that the calcium and phosphorus ratio should be maintained between 1 : 1 and 3 : 1 to avoid alterations in ossification. Phosphorus excess seems to be the most common calcium/ phosphorus imbalance associated with DOD. Vitamin requirements are also extrapolated from mare’s milk composition and average milk intake of normal foals. In other species, vitamin A and E deficiencies have been associated with decreased resistance to respiratory infection. Antioxidant supplementation has been associated with improved immune function associated with increased levels of interleukin 2 (IL-2), elevated numbers of total lymphocytes and T-cells, increased killer cell activity, augmented antibody response to antigen stimulation, decreased lipid peroxidation, and decreased prostaglandin (PG) synthesis.40 Vitamin C is present in high concentrations in leukocytes and is utilized rapidly during infection. Reduced vitamin C levels are associated with reduced immune function. Ascorbate protects lipid membranes and plasma lipids from oxidative damage. In stressed horses, vitamin C supplementation has been shown to improve immune function. Vitamin E helps maintain cell membrane integrity by limiting lipid peroxidation by reactive oxygen species. Vitamin E-deficient states are associated with decreased B-cell antibody production and T-cell proliferation in response to mitogenic stimulation and an increased rate of infection. Supplementation with ascorbic acid and vitamin E has been shown to have neuroprotective effects against CNS ischemia, and is prescribed by some clinicians during the treatment of foals suffering hypoxic-ischemic encephalopathy. While supplementation with these vitamins for compromised neonates seems intuitive, there are no specific guidelines for vitamin supplementation in neonatal foals. Doses that have been used in foals are as follows: vitamin C (ascorbic acid): 100 mg/kg i.v./p.o. q 24 h; vitamin E (α-tocopherol): 5000 U p.o.; or 68 IU/kg i.v. diluted in 1 L of fluids q 24 h.41
Nutrient requirements of the sick neonate The nutrient requirements for premature and sick foals are highly variable and difficult to calculate. Critically ill foals, suffering from sepsis, prematurity, peripartum asphyxia, and a variety of gastrointestinal disorders, frequently suffer from impaired
or erratic nursing behavior, inability to digest milk efficiently, altered metabolism, as well as decreased physical activity. In addition, dysmature/premature foals and those neonates exposed to chronic placental insufficiency in utero often have very limited energy reserves at birth. Actual measurements of resting metabolic rates in one group of sick foals, suffering from varying degrees of immaturity or peripartum asphyxia syndrome, showed it was 75% of that for age-matched, healthy foals in the same environment.42 Yet some studies in people and animals provide evidence that septicemia and fever tend to increase, not decrease, the metabolic requirements.43 Severe infection may increase resting energy expenditure (REE) by 10 to 30%; fever may increase metabolic rate by 13% for each degree Celsius rise in temperature. Limited studies using indirect calorimetry to assess the nutritional requirements of two small groups of healthy and sick foals, by measuring REE and the respiratory quotient (RQ), confirmed considerable individual variation.33 The mean REE of all foals in the study was 44.36 kcal/kg, with sick and recovering foals having a slightly higher mean REE than healthy foals. The route of nutritional support adds new variables to the calculation of nutrient requirements. Many sick foals experience varying degrees of gastrointestinal dysfunction, requiring that enteral feeds be replaced by or supplemented with parenteral alimentation. It has been suggested that infants fed parenterally require only 75% of the gross energy intake of infants fed enterally to achieve the same growth rates. Little is known about a foal’s specific amino acid requirements. Glutamine has been classified as a conditionally essential amino acid in critically ill patients, including humans. Under normal conditions it is the preferred respiratory fuel for rapidly proliferating cells, including mucosal epithelial cells. Skeletal muscle is the main repository of glutamine. During sepsis and infection there is increased production and release of glutamine from the muscle, accompanied by an eight- to tenfold increase in glutamine uptake by the liver.44 The cells of the immune system (lymphocytes, macrophages) are also major glutamine consumers. Glutamine plays an important role in immunomodulation, gastrointestinal protection, and prevention of protein depletion. Glutamine supplementation in enteral and parenteral formulas administered to critically ill infants and adults has been shown to decrease hospital-acquired sepsis, improve nitrogen balance, and decrease morbidity and mortality rates.44–46 The glutamine requirements of the ill foal are not known. Premature and growth-retarded foals have poorly defined energy and protein requirements. Some foals suffering from prematurity, sepsis, and/or
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creased lipoprotein lipase activity. Lipemia develops in many septic patients, including neonates. During sepsis, the final fuel source becomes protein degradation. Uremia, production of false neurotransmitters, hepatoencephalopathy, and neurological signs occur when excessive amino acid degradation overwhelms hepatic metabolic capacity.
Enteral feeding ‘If the gut works, use it!’ Enteral alimentation is more physiological and cheaper than parenteral nutrition. Enteral nutrition seems to provide a critical stimulus for normal intestinal function and development. Oral feeds stimulate the growth of intestinal villi and the production of crypt cells, and induce hepatic and biliary secretions and brush border disaccharidase enzymes. Enterocytes rely on absorption of volatile fatty acids (VFA) such as glutamine and β-hydroxybutyrate from the intestinal lumen as their primary energy source. Therefore, even in foals that must be fed parenterally, small volumes of enteral feeds often are given just to ‘feed the gut’ to prevent gastrointestinal atrophy. During intestinal atrophy the integrity of tight junctions is destroyed, predisposing to translocation of intestinal bacteria into the bloodstream, resulting in septicemia. Oral feeds also stimulate release of enteroinsular hormones (e.g., glucagon, gastrin, secretin, and cholecystokinin) which in turn exert a tropic effect on gastrointestinal maturation. Foals that are candidates for enteral nutrition include those that have reasonable gut motility as evidenced by the presence of borborygmi on auscultation, a history of passing normal manure, and a lack of abdominal distension, gastric reflux and colic. If intestinal health is questionable, an abdominal sonogram should be performed to critically evaluate gut motility, bowel wall thickness, and any evidence of peritoneal effusion. Hypothermic and hypotensive foals should not be fed enterally until vital signs have stabilized. During low-flow perfusion states, as seen during severe septicemia and peripartum asphyxia, blood flow to the bowel is reduced and altered motility and digestion are anticipated. Feeding a poorly perfused gastrointestinal tract should be avoided since it only increases the risk of maldigestion and malabsorption which contribute to gas distended bowel, colic, and diarrhea. If a foal has not received any enteral meal for more than 48 hours, milk meals should be reintroduced slowly since brush border enzyme activity may already be reduced. If colostrum is available, it can be incorporated into the first meals due to its proven trophic effect on intestinal development and local
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mucosal immunity. Some clinicians add lactase to the early meals to reduce the risk of maldigestion associated with reduced disaccharidase activity. If intestinal integrity is still questionable, enteral feeding can be initiated using a solution containing only glucose and electrolytes, prior to reintroducing milk meals. Enteral formulas containing glutamine are attractive first meals due to the beneficial effects of glutamine on intestinal and hepatic function reported in other species. The optimum dose of glutamine for foals is unknown. Full-term and premature infants experienced beneficial effects from glutamine administered orally at a rate of 0.3 g/kg/day or intravenously at a rate of 0.4 g/kg/day.
Enteral formulas
Colostrum The epitheliochorial equine placenta prevents in utero transfer of immunoglobulins from dam to fetus prepartum. Except for trace amounts of IgM, newborn foals are born agammaglobulinemic and are dependent on absorption of IgG antibody from the colostrum. Active pinocytosis of the large proteins, including antibodies, occurs at a maximal rate during the first 6 to 10 h postpartum and declines rapidly thereafter. Gastrointestinal closure is complete by 18 to 24 h of age. Prematurity, cold stress, hypoxic gut damage, and ingestion of other medications such as ditrioctahedral smectite48 (Biosponge, Platinum Performance, Buellton, CA) may alter immunoglobulin absorption unpredictably. A recent study examined the effect of withholding macromolecules on the duration of intestinal permeability to colostral IgG in healthy neonatal foals.49 In this study, foals were fed either a glucose–electrolyte solution or commercial milk replacer for 12 h after birth before being fed a known amount of colostral IgG. Control foals were fed colostrum immediately after birth. The foals given colostrum immediately after birth transferred approximately 51% of ingested IgG into their vascular space. Delayed colostral ingestion significantly reduced IgG absorption. Withholding macromolecules for 12 hours had no effect on the subsequent efficiency of colostral IgG absorption nor did the type of fluid fed prior to colostral ingestion, suggesting that gastrointestinal closure in foals is not mediated by a finite capacity for macromolecular uptake. Colostrum has been recognized as a valuable ‘first meal’ in many species, including humans, since it contains many non-nutrient factors including hormones, growth factors, enzymes, and trophic factors in addition to calories. Equine colostrum is produced by selective secretion of humoral antibodies
into the mammary gland during the last month of gestation with peak secretion occurring during the last 2 weeks of pregnancy. Colostrum contains immunoglobulins (IgG constitutes > 80% of colostral immunoglobulins), complement and lactoferrin that aid in defense against pathogens, low molecularweight proteins that enhance efficiency of absorption of macromolecules, and epidermal growth factor that enhances gastrointestinal mucosal growth and may help prevent gastric ulcers. Colostrum is energy dense and has higher levels of protein and vitamins A and E compared to mare’s milk. Colostrum also has laxative properties. Foals that ingest and absorb sufficient amounts of colostral antibodies have serum IgG concentration > 800 mg/dL by 18–20 h of age. Colostrum can be stored frozen for at least a year without significant loss of immunoglobulin activity. However, even ‘old’ colostrum can be used for its nutrient value and laxative properties as a first meal for sick foals, regardless of their age, that are just being reintroduced to enteral feeds. Mare’s milk Mare’s milk is highly digestible and contains lactose as the primary disaccharide. When compared to cow and goat milk, mare’s milk is lower in energy, higher in water content, lower in fat and total solids, and higher in lactose (Table 28.2). During the first few weeks postpartum the concentration of total solids, protein, and lipids in mare’s milk decreases. Healthy foals digest mare’s milk very efficiently so that almost 98% of the gross energy consumed is available for metabolism and growth.50 Fresh mare’s milk fed straight from the udder is the ideal enteral formula. If a foal’s dam is deceased or uncooperative, a nurse mare represents the perfect substitute: a source for an all-milk diet available on demand and a role model, protector, and disciplinarian for the foal. If a nurse mare is not available there are several protocols that can be used on non-lactating mares to encourage them to lactate and accept a foal. One treatment protocol to induce nonpregnant mares to lactate and become foster dams for orphan foals incorporates the use of domperidone. The regimen consists of a minimum 7-day course of: domperidone (1.1 mg/kg, q 24 h, p.o.), altrenogest (44 mg q 24 h, p.o.), and estradiol – benzoate in oil (10 mg, q 24 h, i.m.).51, 52 In the author’s experience this protocol also has been used effectively to treat certain postpartum mares that are rejecting their foals and demonstrating varying degrees of agalactia. Following the use of this and similar protocols, nonparturient mares will produce milk and some may initially produce milk with elevated immunoglobulin concentrations.53
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Table 28.2 Comparison of milk from mare,a cow, goat, and selected milk replacers. From Ullrey (1966)19 and Oftedal (1983)31 Milk Nutrient Total solids DM (%) Crude protein (%) Crude fat (%) Crude fiber (%) Sugar (%) Calcium (%) Phosphorus (%) Zinc (ppm) Copper (ppm) Selenium (ppm)
Milk replacer
Mare
Cow
Goat
Mare’s Matchb
Foal-Lacc
Mare’s Milk Plusd
11.3–12.0 33–25 15–17 0 58.8 1.1 0.7 23 4 0.04
12.5 27 38 0 38.0 1.1 0.7. 40 3 0.024
13.5 25 31–34 0 31.0 1.0 0.8 30 2 –
11–13 Min 24 Min 16 Max 0.15 NA 0.65–1.15 Min 0.6 Min 40 Min 10 Min 0.3
16 Min 19.5 Min 14 Max 0.1 NA 0.9–1.2 Min 0.75 Min 50 Min 18 0.10
12.5 Min 21 Min 14 Max 0.15 NA 0.7–1.10 Min 0.65 Min 110 Min 35 Min 0.3
a
During first 4 weeks of lactation. Land O Lakes, Arden Hills, MN. c Pet-Ag Inc, Hampshire, IL. c Buckeye Nutrition, Dalton, OH. NA, not available. b
Other enteral formulas Mare’s milk remains the preferred enteral formula. Studies by Ousey et al.54 confirmed that mare’s milk is the preferred diet for a foal in terms of digestible energy and nutrients. Energy balance data indicate this diet provides enough excess energy to support growth as early as postpartum day 2. If mare’s milk is not available, other popular enteral formulas include goat’s milk and artificial milk replacers formulated for foals. Goat’s milk is higher in fat, total solids, and gross energy than mare’s milk (see Table 28.2). It is more digestible than cow’s milk due to its composition of simpler fatty acids, smaller fat globules, and better buffering capacity. Foals raised on goat’s milk occasionally exhibit constipation and metabolic acidosis. Cow’s milk can be substituted for mare’s milk if additional sugar is added and some of the fat is removed. This can be accomplished using 2% skim milk and adding 20 g dextrose/L milk (i.e., 40 mL of 50% dextrose per liter of milk). Sucrose (e.g., table sugar) should not be used since newborn foals lack the enzymes necessary to effectively digest this sugar. A variety of commercial mare’s milk replacers are available. The ideal replacer should be designed for foals (not calves), should closely match the energy density of mare’s milk (approximately 0.5 kcal/mL as fed), and should contain approximately 23% crude protein (CP), 15% crude fat, 59% sugar, and less than 0.5% fiber on a DM basis. One formula that is palatable and well tolerated by most foals is Mare’s R milk replacer (Land O Lakes, Arden Hills, Match MN). When reconstituted according to manufacturer’s directions, many formulas are more concentrated
than mare’s milk, with nearly double the DM content, and may predispose to constipation and dehydration. Free access to water reduces the risk of dehydration. A comparison of energy balance between foals allowed to nurse from their dams, bottle-fed milk replacer, or maintained on TPN (total parenteral nutrition) showed that foals fed milk replacer required several days to adapt to the formula before achieving a positive energy balance and similar growth rates as the dam-reared foals.54 Dilution of the formula was required beyond the manufacturer’s recommendations due to the excessive energy density. Based on that data, Ousey et al. advise that the final concentration of any milk replacer should be diluted until its energy density is equal to or less than that of mares’ milk (approximately 2.5 MJ/L). The foals receiving TPN at a rate of 55 kcal/kg/day experienced a negative energy balance, an increased incidence of gastric ulcers, and difficulty resuming enteral feeds due to suspected gut damage sustained as a result of temporary, but complete, deprivation of enteral feeds. Use of acidified milk replacers, such as Mare’s R (Buckeye Nutrition, Dalton, OH) allows Milk Plus milk to be left out for longer periods without the danger of it spoiling. Acidified formulas may allow foals to consume a given volume of milk over a longer time interval, thereby avoiding the foal’s tendency to quickly consume large volumes of milk over a brief time interval. Human enteral formulas also have been used in foals. These formulas contain protein, carbohydrate, and fat in complex or simple forms such as peptides, amino acids, glucose oligomers, glucose, and longand medium-chain triglycerides. These formulas are often calorie dense, which is an attractive feature for
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sick foals with reduced tolerance for large volumes of milk. The digestibility of these human products should be evaluated in light of the foal’s complement of digestive enzymes and the substrates present in the formula. The pattern of disaccharidase development in young foals and the results of oral tolerance tests performed in foals less than a week of age27 suggest that equine neonates are incapable of flourishing on any enteral diet high in sucrose, maltose, or polysaccharides as the primary carbohydrate source. Routes of administration for enteral nutrition If a newborn foal is simply physically unable to nurse from the mare, but has a functional intestinal tract, then its dam should be hand-milked at least every 2 to 3 hours, and the foal bottle-fed fresh milk. Prior to milking, the mare’s teats should be lubricated with KY jelly to reduce irritation. Administration of low doses of oxytocin (5–10 U i.v./i.m.) 5 to 10 minutes before milking will facilitate milk letdown. If the mare’s milk production remains poor, oral administration of domperidone (1.1 mg/kg orally once daily) may help improve lactation. If the foal has a productive suckle and swallow reflex, then bottle-feeding is attempted using a lamb or infant nipple. Some premature and ‘dummy’ foals may develop a suckle reflex before an effective swallow reflex. Whenever a sick or weak neonate is allowed to nurse from the bottle or the mare for the first time, frequent auscultation over the trachea is essential to be certain there is no tracheal rattle associated with dysphagia and fluid aspiration. When bottle feeding, avoid overextension of the foal’s head and neck, to reduce the risk of aspiration. The foal should be encouraged to keep its nose below the level of its eyes while nursing a bottle. Normal udder bumping and teat-seeking behavior can be stimulated by allowing the foal to approach the hand-held bottle from behind and under the handler’s armpit. When the foal demonstrates normal udder seeking behavior by nuzzling or bumping the armpit the bottle should then be offered as a reward. Bottle feeding is not a long-term option for several reasons (see discussion on feeding the orphan foal). If the foal’s swallow reflex is weak, or the foal is destined to be raised as an orphan, then bucket feeding can be tried. Drinking from a bowl or bucket with the head and neck flexed makes swallowing easier. Sometimes it is difficult to teach a foal accustomed to nurse from a mare or a bottle how to drink from a bucket. It is best to begin the introduction to bucket feeding using a shallow bowl, since some foals are reluctant to submerse their head in a deep bucket. Sometimes placing only the nipple in the milk-filled
bowl will entice the foal to use the nipple as a straw. Eventually the foal begins drinking without the nipple. Foals can also be encouraged to drink from a bucket by allowing them to suckle on a handler’s fingers and then slowly submerging your fingers in the bucket. On-demand bottle or bucket feeding is ideal but often impractical. Foals less than 7 days of age should be fed a minimum of every 2 hours. If the foal’s gastrointestinal function is normal, then at least 10% of its bodyweight in milk can be fed during a 24-h period. If mare’s milk must be reheated, a bottle warmer is preferred to the microwave to prevent overcooking, uneven heating, and the destruction of normal bacteria that would be acquired naturally by the foal while nursing from the udder. When using milk replacer it is important that it is mixed the same way each time, to avoid wide variations in concentration, and that any unused formula should be discarded. Begin offering the foal a minimum of 10% of its bodyweight in milk daily, divided into multiple feedings. A 50 kg foal would receive approximately 5 L (150 oz) of milk daily divided into 416 mL (12.5 oz) feeds every 2 hours. If the foal tolerates that diet, then the milk volume can be increased slowly (often by 1–2% of the daily intake per day) over the next 5–7 days until the foal is consuming between 15–25% of its bodyweight in milk per day. Make all dietary changes slowly. If the suckling and swallow reflexes are ineffectual, then nasogastric intubation and administration of milk via gravity flow are required. A small-bore (≥ 5 mm internal diameter), flexible feeding tube with a weighted tungsten end is preferred, to ensure a slow, gravity flow of formula and prevent overdistention of the stomach (Figure 28.1). A small-diameter tube decreases nasopharyngeal irritation and allows foals to begin to learn to nurse with the tube in place. Small indwelling tubes may be sutured to the nares or taped to half a tongue depressor, which is then taped to the side of the foal’s muzzle. The feeding tube, positioned with the end of the tube in the distal esophagus, is attached to a fleece halter (Figure 28.2). The end of an indwelling feeding tube should be positioned in the distal esophagus rather than in the stomach to reduce the risk of retrograde reflux and gastric irritation. Between feedings the indwelling stomach tube should be sealed to prevent aerophagia. Recumbent foals should be maintained in sternal recumbency during, and for at least 10–15 minutes after, tube feeding to reduce the risk of gastroesophageal reflux and aspiration of milk into the lungs. The enteral diet being used should be fed at body temperature and introduced into the tube by gravity flow using a funnel or enteral feeding bag.
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Table 28.3 Complications associated with enteral alimentation Gastrointestinal intolerance
Diarrhea Abdominal distention Gastric reflux Ileus, colic Constipation
Metabolic disturbances
Electrolyte disturbances Hypoglycemia
Nasogastric tube-related problems
Rhinitis Tube occlusion or dislodgement Nasopharyngeal trauma Gastric irritation Aerophagia, bloat, colic Aspiration pneumonia
Poor palatability and spoilage
Figure 28.1 Small-bore, flexible nasogastric feeding tube with weighted end.
Although healthy foals consume between 20–25% bodyweight in milk daily, the provision of at least 10% bodyweight is a reasonable goal when initiating enteral feeding in sick foals or those with compromised gastrointestinal function. At this rate a 50 kg foal requires 417 mL of milk every 2 h. Milk should be fed at body temperature and should not be warmed
Figure 28.2 Foal with nasogastric feeding tube taped to a tongue depressor and then taped to muzzle and foal halter. End of feeding tube is capped and closed.
in the microwave to avoid altering the composition and to prevent uneven heating to overcooking. Any changes in volume or composition of the enteral diet should be made slowly. Complications associated with enteral feeding Common complications associated with enteral feeding are listed in Table 28.3. Delayed gastric emptying and gastroduodenal dysmotility are common problems in many sick neonatal foals, and may be due to infectious and non-infectious causes. Occasionally motility modifiers are administered. These drugs should be used cautiously. If colic, ileus, abdominal distention or gastric reflux is present, an abdominal sonogram is recommended to rule out the presence of an intussusception, abnormally thickened, edematous bowel wall, or other signs suggestive of NEC, prior to initiating any prokinetic therapy. Additional bowel rest and reducing or stopping enteral feeds while increasing parenteral nutrition (PN) support are usually the preferred treatments for most cases of dysmotility. Some of the prokinetic agents that have been used include metoclopramide given as a slow intravenous infusion (0.25 mg/kg/h) that is started 10– 15 minutes prior to feeding and continued for 30– 45 minutes post-feeding. Metoclopramide can also be administered orally (0.25–0.6 mg/kg q 4–6 h). Other motility modifiers include erythromycin given orally (1.0 mg/kg q 6 h) and cisapride administered orally (10 mg/kg q 6 h). Motility modifiers should be used sparingly and only after reasonable bowel integrity and patency have been confirmed. Diarrhea may respond to decreasing the volume of milk being fed. Persistent diarrhea can be treated symptomatically with oral bismuth subsalicylate
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(1–2 mL/kg orally q 4–6 h), psyllium or ditrioctahedral smectite paste (30–60 mL orally q 12 h). Loperamide (0.1–0.2 mg/kg p.o. q 6 h) is rarely used and then only in foals with chronic diarrhea that have normal vital signs and no evidence of endotoxemia (i.e., no evidence of injected membranes and neutropenia). Lactase therapy (6000 IU orally q 3–8 h)55 may help correct osmotic diarrhea associated with disaccharidase deficiency. It is reasonable to assume that any gastrointestinal disorder in the foal that injures or destroys the brush border of the small intestinal epithelial cells has the potential to create a lactase deficiency, leading to maldigestion of milk meals and the development of an osmotic diarrhea. Transient lactase deficiency has been documented in foals with rotavirus diarrhea and clostridial enteritis.55 NEC associated with infectious agents and ischemic damage to the bowel may also affect lactase production. Affected foals often experience colic, diarrhea, and gaseous bowel distention when reintroduced to enteral milk meals. Many of these foals appear to benefit clinically from the administration of 6000 U lactase every 3 to 6 hours. If these foals are not nursing directly from the mare and are receiving their milk by bottle, bucket or tube, then lactase can be added directly to the milk before feeding. Intestinal probiotics containing Lactobacillus spp. bacteria or active culture yogurt or buttermilk often are administered frequently to foals receiving antibiotics and/or artificial diets with the premise of helping re-establish intestinal flora. Probiotics are defined as ‘living microorganisms, which upon ingestion in certain numbers exert health effects beyond inherent basic nutrition.’56 Another definition describes a probiotic as a ‘live microbial feed supplement which beneficially affects the host animal by improving its intestinal microbial balance.’57 The concept of probiotics was first reported by the Russian biologist, Eli Metchnikoff, who credited the long lives of certain Russian and Bulgarian citizens to the consumption of large amounts of fermented milk products.58 Lactobacillus acidophilus, a lactic acid producing bacteria, was later identified as a key organism in these foods.59 There is a paucity of scientific research regarding the use of probiotics in any veterinary species during health or disease. Quality control remains an issue with the currently available commercial probiotics. Most probiotics are comprised of one or more lactic acid producing bacteria and certain strains of yeast. The proposed benefits of probiotics include:
r
Competitive inhibition of enteric pathogens through competition for essential nutrients and/or adhesion receptors: beneficial bacteria compete for attachment sites within the small
r
r r r
intestine and reduce the ability of disease-causing organisms to bind to and colonize the small intestine. It is speculated that beneficial bacteria may enhance enzyme activity and thus improve nutrient absorption. Production of antimicrobial substances: such substances may include lactic acid that reduces intestinal pH, hydrogen peroxide that renders the microenvironment unfavorable for oxygenrequiring organisms, and bacteriocins that restrict the growth of other microorganisms. Stimulation of the immune system. Enhancement of growth of beneficial bacteria: lactic acid producing bacteria secrete B vitamins and other beneficial enzymes. Certain strains of yeast may enhance fiber digestion. Decreased absorption of intestinal antigens within the small intestines. This effect may be mediated by closure of large molecule transport pores. If that is true, one wonders if the use of probiotics in foals less than 24 hours of age may adversely affect immunoglobulin absorption.
In order for probiotic organisms to exert the properties mentioned, the organisms must survive transit through the acidic gastric pH, resist bile digestion, be able to adhere to intestinal epithelial cells and colonize the intestinal tract, produce antimicrobial substances, and stimulate the immune system. An excellent scientific review of these issues has been published.60 Most reports regarding the benefits of probiotic use in foals are only anecdotal. Oral administration of Saccharomyces boulardii, a subspecies of Saccharomyces cerevisiae, has been recommended as part of the treatment for and prevention of Clostridium difficile enteritis. Saccharomyces boulardii is believed to competitively inhibit the binding of C. difficile to intestinal mucosa. Nasopharyngeal irritation from repetitive tubing or chronic intubation delays the return of normal suckle and swallow reflexes and nursing vigor. The use of the smallest-diameter feeding tube helps reduce the irritation and allows the foal to swallow with the tube in place. If pharyngeal inflammation develops, insufflation of a nasopharyngeal spray containing furacin (750 mL), prednisolone (2000 mg), glycerin (1000 mL), and DMSO (250 mL) reduces mucosal inflammation. A small volume (5–7 mL) of pharyngeal spray is administered via a smalldiameter tube inserted in the nares to the level of the medial canthus, with the foal in a standing or sternal position with its head flexed and held low. Sick foals often lose weight during their illness, followed by a period of catch-up growth during the recovery phase. Growth spurts should be controlled to reduce the risk of DOD.
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Feeding the orphan foal Feeding the orphan foal is relatively straightforward since nutrient requirements and feeding regimes can be based on the nutrient intake and nursing behavior of healthy foals. The orphan usually represents a foal with a healthy gastrointestinal tract and normal nutrient requirements. Feeding decisions include what enteral formula to use, how much to feed, how often, and how to administer it. Orphan foals fare best if bonded as soon as possible to a nurse mare. Goats have also been used as surrogate dams. Bonding an orphan to a nurse mare ensures the most physiological diet and nursing regimen, allows the orphan to maintain normal social and dietary behavior patterns, and provides maternal protection and discipline. Keep rate of growth in mind when selecting a nurse mare for orphan foals. For example, a small, orphaned Quarter Horse foal is not well suited to having a Belgian mare with excessive milk production as a foster ‘mom.’ This scenario sets the stage for excessive growth and weight gain and predisposes to certain forms of DOD. If a nurse mare or goat is not available then bucket feeding is the next best option. Bucket-raised orphans that are deprived of any four-legged role model tend to exhibit delayed ingestion of solid feeds including hay, grass, and creep feed. Orphans raised in isolation do not exhibit normal coprophagy and miss the opportunity to interact with other foals. If a nurse mare is not an option, then a quiet equine chaperone should be found for the orphan. Another viable option is raising several orphans together. Orphans raised by any method, other than by nursing free choice from a mare, are more prone to gastrointestinal disturbances, ranging from constipation to diarrhea, an observation that may be related to many factors including inconsistent mixing of milk replacer formulas, regimented rather than voluntary feeding intervals, altered gut flora development, delayed intake of solid feeds, and stress. Those orphans that have been sick as neonates may have been bottle-fed initially, but bottle feeding should be considered only a temporary substitute for either a surrogate dam or a bucket. Bottle-fed foals often bond too closely with their human surrogates and are slow to develop normal social behaviors. Many bottle-fed orphans are slow to learn to eat creep feed and to graze, and without a surrogate dam they cannot be left unattended outside. Bucket feeding milk replacer is preferable to bottle feeding. Raising orphans in a group is better than rearing one by itself. Allowing orphans access to manure from healthy, recently dewormed adult horses so that they can engage in coprophagy similar to foals living
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with their dams may help reduce the incidence of intestinal disturbances. The volume of milk required by an orphan varies with the individual and is affected by the foal’s age, general health status, breed, and potential growth rate. Foals ingesting adequate amounts of milk or milk replacer should be gaining an average of 1–2.5% of their bodyweight daily, passing manure of normal consistency on a regular basis, and urinating adequate volumes of dilute urine. Fresh water should be available at all times. The foal’s daily weight gains should be monitored and growth spurts should be controlled to reduce the risk of DOD.
Parenteral nutrition Parenteral nutrition (PN) offers an important nutritional bridge during illness and is indicated whenever feeding via the alimentary tract is inadequate or contraindicated. PN is an effective way to supply complete or partial protein and energy needs for foals on a temporary or long-term basis. If a sick neonate is unable to consume at least 5% of its bodyweight in milk per day, additional parenteral nutritional support is recommended. Intravenous solutions of 5–10% dextrose can be administered at rates averaging 100–200 mL/h for short-term caloric supplementation. The caloric densities of 5% and 10% dextrose solutions are 0.17 kcal/mL and 0.34 kcal/mL, respectively. Used alone and at fluid rates considered maintenance for foals (e.g., 4–5 mL/kg/min), neither of these dextrose solutions can provide enough calories to support a foal’s energy needs. For example, administering 5% dextrose at an infusion rate of 5 mL/kg/h will provide only 20 kcal/kg/day. When using a dextrosecontaining solution at maintenance rates, a solution other than dextrose 5% in water is preferred, since this solution lacks electrolytes, is hypotonic, and is used primarily to provide free water to patients suffering from hyperosmolality. A 50% dextrose solution contains 1.7 kcal/mL and can deliver significantly more calories in a reduced volume. However, this solution is too hypertonic to be used without dilution. Parenteral nutrition involves intravenous administration of hypertonic solutions containing 50% dextrose, 8.5% amino acid solutions, and 10 or 20% lipid emulsion, in addition to varying amounts of vitamins, electrolytes, and trace minerals. These PN solutions must be administered continuously and aseptically through a large-bore (usually jugular) catheter to minimize vessel irritation and avoid fluctuations in plasma osmolarity. Jugular catheterization is the most common access route for PN therapy. Catheters should be of low reactivity and approximately 12–20 cm in length. The Arrow or MILA
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Foal: Specific Diagnostic and Management Techniques
catheters (Arrow catheter, Arrow International, Inc, Reading, PA; MILA catheter, Mila International, Florence, KY) are long, polyurethane catheters that are available with single or multiple lumens. Ideally hyperalimentation formulas should be administered through a dedicated single-lumen catheter. Another option is to use a multi-lumen catheter to allow for concurrent administration of other fluids and medications. The venipuncture site and catheterized vein should be examined daily for signs of infection or thrombosis, and all intravenous lines should be changed at least every 72 hours. The use of volumetric infusion pumps ensures accurate and continuous flow rates. Components of parenteral nutrition It has been suggested that sick foals require significantly less energy (44.36 kcal/kg/day) than healthy foals (120 kcal/kg/day).33 The non-protein energy requirements of a sick (full-term) foal have been estimated to be 30 kcal/kg/day. Energy requirements for most recumbent, sick foals are probably less than those for a healthy ambulatory foal due to inhibited growth, reduced insensible losses, and decreased activity. Therefore the goal of PN is to avoid overfeeding and to meet the basal or resting energy requirements of the sick neonate. When initiating PN with 50% dextrose (1.7 kcal/mL), to provide 3.4 kcal/g of glucose, the infusion usually begins at 10 g/kg/day. Dextrose concentration in the PN formulation can be increased to deliver up to 12–15 g/kg/day of glucose if indicated and tolerated by the patient. Concurrent insulin administration may be required to control hyperglycemia and optimize glucose metabolism in foals experiencing reduced insulin production or varying degrees of insulin resistance associated with their illness or prematurity. Lipid emulsions are the most energy-dense component of PN, contain primarily long-chain triglycerides, and deliver 9–11 kcal/g of fat. A 20% lipid emulsion delivers approximately 2.0 kcal/mL. Except for the lipid emulsion, the other two PN (i.e., 50% dextrose, 8.5% amino acids) solutions are hypertonic. Therefore, the addition of lipids to the PN formula helps reduce the overall hypertonicity of the final solution and may reduce the incidence of vein-related complications such as thrombosis and thrombophlebitis. When hyperglycemia restricts glucose administration, lipids can be used to provide up to 30–60% of non-protein calories. However, the addition of lipids to the PN formulation may not always be indicated or beneficial. Impaired lipid clearance may occur during sepsis or in association with prematurity. Lipid emulsions are prone to
contamination and may promote bacterial growth.2 Lipid emulsions are not routinely administered to infants with marked hyperbilirubinemia, since elevated concentrations of free fatty acids compete with bilirubin for albumin-binding sites and may exacerbate kernicterus. Controversy also surrounds the use of lipids in infants with severe pulmonary compromise. There is concern that lipid emulsion interferes with pulmonary microcirculation and impairs gas exchange. Hyperlipidemia can occur during lipid administration to foals, but does not appear to have adverse effects.61, 62 Resulting lipemia may respond to heparin administration. Heparin augments lipoprotein lipase activity. Lipid infusion rates often begin at 1 g/kg/day and can be increased to rates as high as 4 g/kg/day. In a recent study, the addition of lipids to the PN allowed for the provision of 40–92 kcal/kg/day to foals.62 Amino acid solutions, available as free amino acids or with electrolytes added, are hypertonic and provide approximately 4 kcal/g of protein. An 8.5% amino acid solution delivers 0.34 kcal/mL. Protein solutions are added to PN formulas to provide amino acids and nitrogen for growth and cellular repair. To prevent or reduce catabolism of protein for energy, an adequate supply of both energy and nitrogen should be provided. The recommended ratio of non-protein calories to grams of nitrogen (6.2 g protein = 1 g nitrogen) is approximately 100–200.61 In foals, the ratio of non-protein calories to grams of nitrogen was negatively correlated with weight gain.63 Parenteral nutrition formulas should be prepared in as clean an environment as possible. The use of sterile, empty 3 L or 4 L PN bags with two or three lead transfer sets (Baxter All-in-One bags, Baxter Healthcare Corp, Deerfield, IL; 3 L EVA Empty 3-in-1 Mixing Container, Abbott Labs, North Chicago, IL) facilitates sterile mixing of ingredients. When mixing PN formulas the glucose and amino acid solutions are usually added first, followed by the lipid emulsion, in order to maintain a suitable solution pH for lipid solubility. To ensure compatibility between ingredients it is advantageous to use the same manufacturer for the dextrose, amino acid, and lipid solutions. It is recommended that the concentration of dextrose in the final PN solution should not be greater than 10%. Crystalloid solutions can be added to the final formula or run concurrently in the same catheter to reduce the tonicity of the solution to 10%. Foals that are not receiving any supplemental enteral nutrition usually become hypokalemic and require potassium supplementation in the form of potassium chloride (KCl). Vitamin and trace mineral supplementation should also be considered for foals that require long-term PN. If vitamins or trace minerals are to be added, they should be introduced into the
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Enteral and Parenteral Nutrition for the Neonatal Foal formulation just before administration. PN solutions should be refrigerated before use. Parenteral nutrition formulation Reviews of two basic approaches to PN formulation for foals have been published.2, 64 The first approach involves the precise calculation of the anticipated metabolic needs and energy demands of the patient, followed by exact calculations of the volumes of dextrose, amino acid, and lipid-containing fluids required. This is a complex and time-consuming procedure and is well described elsewhere.3 The second approach is more practical and adaptable to a variety of settings, and utilizes either a dextrose–proteincontaining formulation or a dextrose–protein–lipidcontaining solution; the recipe and caloric density for these two formulas are provided in Table 28.4. Formula 1 lacks lipids and is less energy-dense and more hypertonic than formula 2, and is best used for shortterm, partial PN support. Both these formulations have been designed as suitable solutions to use when first introducing foals to PN. Both solutions provide an acceptable ratio of non-protein calories to grams of nitrogen to ensure that the protein being fed is used for growth and cellular repair rather than catabolized for energy. To initiate PN, the daily energy requirements for a given foal are estimated and the total volume of either formula 1 or 2 are calculated. The total volume is divided by 24 to obtain the targeted hourly infusion rate. Foals should be weaned onto and off PN gradually. If the foal has been receiving glucose-containing solutions prior to beginning PN, the current infusion rate of glucose can be calculated and the initial rate of PN can be calculated to deliver a similar delivery rate of glucose; otherwise the PN infusion rate can be started at approximately 50% of the calculated final target rate. Until the target rate is reached and the foal’s condition is considered stable, blood glucose concentrations should be monitored frequently to avoid both hypoglycemia (glucose < 100 mg/dL) and hyperglycemia (glucose > 180 mg/dL). The more
critical the patient, the more frequently blood glucose levels should be monitored. Urine output should be monitored frequently and urine tested for glucose intermittently. In most foals glucosuria and subsequent osmotic diuresis are observed when glucose concentrations exceed 180 mg/dL. Whenever blood samples are drawn they should be observed for gross lipemia. The PN infusion rate should be increased in slow increments every 2–4 hours until the target rate is reached. Another approach to calculating PN solutions is presented in Table 28.5. In this scenario the clinician begins with a basic, time-tested formula of 10 g/kg/day of glucose; 2 g/kg/day of protein and 1 g/kg/day of lipids. Based on the foal’s weight and the concentration of the stock solutions of dextrose, protein, and lipids being used, the final PN recipe can be formulated. Once opened, the stock solutions should be kept sealed and refrigerated. In an attempt to reduce the risk of contamination of the stock solutions, it is advisable to discard unused, open stock solutions after 72 hours. Once the foal’s condition is stabilized, the daily PN formulation can be adjusted accordingly to meet changing caloric and/or protein requirements of the patient. The rate of glucose administration can be increased slowly from 10 g/kg/ day up to 12–14 g/kg/day; protein administration can be increased from 2 g/kg/day up to 3–4 g/kg/day; lipid administration can be increased up to 3–4 gm/kg/ day. These components can be adjusted provided a few basic guidelines are remembered: total calories from lipids should not exceed 60% of the non-protein calories, and enough non-protein calories should be provided to prevent catabolism of amino acids for energy.61 This last precaution can be met by ensuring that an average of 100–200 non-protein calories are administered for every gram of nitrogen fed.61 When calculating energy needs for the sick foal, it is probably safest to err on the side of underfeeding rather than overfeeding, with the goal being to prevent undue catabolism.65 Excessive administration of glucose often precipitates hyperglycemia, which is
Table 28.4 Characteristics of two basic PN formulations for foals. From McKenzie (2009)2
Formulation
Composition
Formula 1
1500 mL 50% dextrose, 1500 mL 8.5% amino acids 1500 mL 50% dextrose, 2000 mL 8.5% amino acids, 500 mL 20% lipids
Formula 2
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Caloric density (kcal/mL)
Ratio of non-protein calories to grams of nitrogen
1.02
125
1.08
131
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Table 28.5 Plan for formulating partial parenteral nutrition (PN) for a 50 kg foal General guidelines and nutrition facts • Consider the patient’s specific nutrient requirements Healthy foal: ∼ 120 kcal/kg/day Sick foal: ∼ 40–50 kcal/kg/day • Glucose = 3.4 kcal/g; 50% dextrose solution provides 1.7 kcal/mL; 500 mg dextrose/mL • Protein = 4.0 kcal/g; 8.5% amino acid (AA) solution provides 0.34 kcal/mL; 85 mg AA/mL • Lipid = 9.0–10.0 kcal/g; 10% or 20% lipid emulsion provides 1.0 or 2.0 kcal/mL; 100 or 200 mg lipids/mL • Non-protein (NP) calories are those supplied by lipids and glucose Calories provided by lipids should represent ≤ 60% of non-protein calories To prevent catabolism of amino acids for energy, the ratio of non-protein calories to grams of nitrogen in the PN formula should average between 100 and 200 • 6.25 g protein = 1 g nitrogen Calculations for initial PN formulation Glucose: 10 g/kg/day = 500 g/day = 1000 mL of 50% dextrose solution Protein: 2 g/kg/day = 100 g/day = 1176 mL of 8.5% amino acid solution Lipids: 1 g/kg/day = 50 g/day = 500 mL of 10% lipid (or 250 mL of 20% lipid) emulsion Total fluid volume: 2676 mL Hourly infusion rate: ∼ 111.5 mL/h Nutrients provided by final formula Glucose: 500 g × 3.4 kcal/g = 1700 kcal Protein: 100 g × 4.0 kcal/g = 400 kcal Lipids: 50 g × 9.0 kcal/g = 450 kcal • Total non-protein calories provided = 2150 kcal = 43 kcal/kg/day NP calories/g of nitrogen NP calories = 2150 calories Grams of nitrogen: 100 g amino acid/6.25 = 16.0 g nitrogen NP calories/g of nitrogen = 2150/16 = 134.4 Source of NP calories in PN formula: 79% from glucose; 21% from lipids The final PN formula is hypertonic with a final dextrose concentration of ∼ 19%. Approximately 2324 mL of crystalloid solution should be added to the PN solution or run concurrently over the same 24-hour period to reduce the dextrose concentration of the overall infusion to ∼ 10%. This could be accomplished by running the PN solution at the required rate of 111.5 mL/h and running balanced crystalloid solution at 97 mL/h.
itself a proinflammatory stimulus. Excessive carbohydrate administration may lead to increased carbon dioxide production which may worsen hypercapnea in sick or premature neonates with pre-existing respiratory compromise. Hypokalemia develops in many foals that are receiving complete PN. Potassium supplementation can be achieved by calculating the potassium deficit and adding the appropriate volume of potassium
chloride to either the PN formula or the crystalloid fluids being administered concurrently. If the foal is destined to require long-term PN support, then trace mineral and vitamins are often added to the PN formula, although specific guidelines for foals have not been published. Many sick foals demonstrate varying degrees of hyperglycemia, due to insulin resistance and/or insulin deficit, which can make it difficult to increase the PN infusion to the desired target rate. If glucose concentrations exceed 180 mg/dL, and decreasing the rate of PN infusion by 30–50% for 3–6 hours does not improve the hyperglycemia, then often the only remaining remedy is exogenous administration of insulin. Regardless of whether the foal receives a continuous infusion of insulin or is given intermittent subcutaneous doses, the patient will need to be monitored closely to ensure that severe hypoglycemia does not develop. Subcutaneous administration is easier and simpler to give, but does not allow frequent changes in the dosage and has a slower onset of action. A suggested dose is 0.1–0.5 IU/kg s.c.66 Continuous rate infusions (CRI) of insulin allow a more rapid onset of effect and more control over the dosage. Cellular insulin receptors become saturated gradually and so the maximal effect of CRI insulin is not observed for as long as 90 minutes after initiating the infusion.2 For the same reason, any change in dosage or infusion rate should also be given adequate time to exert an effect on glucose concentrations. Using regular insulin, initial rates for insulin CRI vary between 0.00125 and 0.07 IU/kg/h for most foals.2, 3, 67 Blood glucose concentrations should be evaluated within 2 hours of initiating insulin therapy. For more detailed information on making and administering insulin infusions the reader is referred to more in-depth discussions of the topic.2, 4, 67, 68 Foals must be weaned onto and off PN slowly. The author does not begin to decrease the rate of PN administration until the foal is tolerating at least 5% of its bodyweight in milk per day. PN often can be discontinued completely once the foal is consistently consuming a volume of milk equivalent to 10% of its bodyweight daily. Complications associated with parenteral nutrition Some of the immediate complications associated with PN administration include fluctuations in blood osmolarity, concentrations of glucose and electrolytes, acid–base balance, and hydration status. The more common complications associated with PN are presented in Table 28.6. Therefore, in addition to closely monitoring glucose concentration, a number of other parameters should be monitored. Guidelines
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Enteral and Parenteral Nutrition for the Neonatal Foal Table 28.6 Complications associated with parenteral nutritional support
Metabolic disturbances Hyperglycemia/hypoglycemia Glucosuria/osmotic diuresis Hyperosmolar states Hyperlipemia, hypertriglyceridemia Hyperchloremic acidosis Uremia/azotemia Electrolyte imbalances Mineral/vitamin imbalances Increased carbon dioxide production secondary to glucose infusion leading to hypercapnea; may exacerbate underlying respiratory disease
Catheter-related problems Thrombosis Phlebitis Sepsis: bacterial, fungal
for monitoring foals on PN are presented in Table 28.7. A complete absence of enteral feeding encourages atrophy of the alimentary tract and associated lymphoid tissues, and may have immunosuppressive consequences for the foal, particularly if total parenteral support is continued for a prolonged period of time. Complete PN (i.e., absence of all enteral feeding) has been associated with gastrointestinal atrophy and malfunction, including alteration of intestinal cytokines and mucosal immunity.1, 69 PN has been shown to impair mucosal immunity and reduce IgA levels through depression of gut-associated lym-
phoid tissue (GALT) cytokines and GALT-specific adhesion molecules.70 Neonatal studies in rodents demonstrate that populating the intestine with immunoglobulin-producing cells and intraepithelial GALT cells depends upon some enteral nutrient stimulation.1 Another study suggests that it is the lack of enteral feeding, rather than the administration of PN itself, that is responsible for loss of mucosal immunity, changes in function of the intraepithelial lymphocytes, and resulting increase in bacterial translocation and secondary septicemia.71 Therefore, even small-volume enteral feeds delivered intermittently to foals receiving PN may help nourish gut epithelial cells and reduce gastrointestinal atrophy. PN has been linked with hepatic dysfunction, including decreased hepatic cytochrome P450 activity.72 The addition of glutamine to PN formulas has been shown to improve survival after bowel ischemia and reperfusion and to prevent many of the TPN-induced changes in intestinal and hepatic function. Recently a stable formulation of glutamine dipeptide has become commercially available; however, there are no published reports evaluating its use in sick foals. Selected studies have demonstrated that the use of total parenteral nutrition can result in negative energy balance in neonatal foals.54 In a more recent retrospective study examining complications attributed to administration of PN in 53 foals, the most common complications were hyperglycemia (glucose > 120 mg/dL) in 89% of foals, hypertriglyceridemia (TG > 200 mg/dL) in 32% of foals, and
Table 28.7 Suggested monitoring procedures for foals receiving parenteral alimentation Initial monitoring frequency
Monitoring after stabilization
Vital signs Catheter/vein inspection Fluid intake/output Weight Urine glucose Serum glucose
q4h q8h Calculate fluid balance q 8 h Daily q 6–8 h q 2–3 h initially; decrease frequency gradually as patient’s condition stabilizes
q 8–12 h q 12 h Daily Daily q 12–24 h q 24 h
Serum electrolytes Creatinine/blood urea nitrogen Triglycerides/cholesterol Visual inspection of foal’s serum for gross lipemia
q 24 h q 24 h q 24 h q 12–24 h or whenever a blood sample is drawn Baseline q 12–24 h Baseline Baseline
q 2–3 d q 2–3 d Weekly q 2–3 d
Observation
Liver enzymes and bilirubin Packed cell volume White blood cell differential Fibrinogen
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Weekly q 24–48 h Weekly Weekly
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catheter-related problems such as thrombosis or local infection in 15% of foals.73 In a separate retrospective study, the use of PN in group of 45 foals was evaluated. It was concluded that the use of lipid-containing PN solutions facilitated the delivery of energy to critically ill foals without increasing the risk of deleterious side-effects, and that severely ill foals were more prone to develop complications associated with PN and to have a poor outcome.62
Summary Nutritional support, whether it is as simple as supplemental bottle feeding or as involved as continuous parenteral nutrition, is a critical part of the treatment plan for any neonatal foal. The impact of poor nutrition on immune function should never be underestimated.
References 1. Hermsen JL, Yoshifumi S, Kudsk KA. Food Fight!: Parenteral nutrition, enteral stimulation and gut-derived mucosal immunity. Langenbecks Arch Surg 2009;394(1):17–30. 2. McKenzie III, HC, Geor RJ. Feeding management of sick neonatal foals. Vet Clin North Am Equine Pract 2009;25(1):109–19. 3. Buechner-Maxwell VA, Thatcher CD. Neonatal nutrition. In: Paradis MR (ed.) Equine Neonatal Medicine. Philadelphia: Elsevier Saunders, 2006; pp. 51–74. 4. Buechner-Maxwell VA. Nutritional support for neonatal foals. Vet Clin North Am Equine Pract 2005;21(2):487–510. 5. Settle CS, Vaala WE. Management of the critically ill foal: Initial respiratory, fluid and nutritional support. Equine Vet Edu 1991;3:49–54. 6. Vaala WE. Nutritional management of the critically ill neonate. In NE Robinson (ed). Current Therapy in Equine Medicine 3. WB Saunders CO, Philadelphia. 1992; 741–52. 7. Vaala WE, Sertich PS. Nutritional and respiratory support for the neonatal foal. In AJ Higgins, JR Snyder (eds). The Equine Manual second edition. Elsevier Saunders, Edinburgh. 2006;837–49. 8. Law DK, Dudrick SJ, Abdou NI. Immunocompetence of pateints with protein-calorie malnutrition. Ann Intern Med 1973;79:545. 9. Mullen JL, Gertner MH, Buzby GP, Goodhart GL, and Rosato EF. Implications of malnutrition in the surgical patient. Arch Surg 114(2):121–125, 1979. 10. Superina R, Meakins JL. Delayed hypersensitivity, anergy, and the surgical patient. J Surg Res 1984;37:151. 11. Viteri FE, Schneider RE. Gastrointestinal alterations in protein-calorie malnutrition. Med Clin North Am 1974;58:1487. 12. Brinson RR, Anderson WM, Singh M. Hypoalbuminemiaassociated diarrhea in critically ill patients. J Crit Ill 1987;2:72–6. 13. Haydock DA, Hill GL. Impaired wound healing in surgical patients with varying degrees of malnutrition. J Parenter Enteral Nutr 1986;10:550. 14. Rochester DF, Esau SA. Malnutrition and the respiratory system. Chest 1984;85:411.
15. National Research Council (ed.) Nutrient Requirements of Horses, 6th edn. Washington, DC: National Academies Press, 2007. 16. Thorson JF, Karren BJ, Bauer ML, Cavinder CA, Coverdale JA, Hammer CJ. Effect of selenium supplementation and plane of nutrition on mares and their foals: foaling data. J Anim Sci 2010;88(3):982–90. 17. Glade MJ. Effects of gestation, lactation, and maternal calcium intake on mechanical strength of equine bone. J Am Coll Nutr 1993;12:372. 18. George LA, Staniar WB, Treiber KH. Insulin sensitivity and glucose dynamics in foals following maternal dietary treatment. Equine Sci Soc 2007;20; 5–6. 19. Ullrey DE, Struthers RD, Hendricks DG, Brent BE. Composition of mare’s milk. J Anim Sci 1966;25:217–22. 20. Ullrey DE, Ely ET, Covert RL. Iron, zinc and copper in mare’s milk. J Anim Sci 1974;38(6):1276–7. 21. Pearce SG, Firth EC, Grace ND, Fennessy PF. Effect of copper supplementation on the evidence of developmental orthopedic disease in pasture-fed New Zealand Thoroughbreds. Equine Vet J 1998;30(3):211–18. 22. Drew B, Barber WP, Williams DG. The effect of excess dietary iodine on pregnant mares and foals. Vet Rec 1975;97:93–6. 23. Fowden AL, Mundy L, Ousey JC, McGladdery A, Silver M. Tissue glycogen and glucose-6-phosphatase levels in fetal and newborn foals. J Reprod Fertil Suppl 1991;44:537–42. 24. Fowden AL, Gardner DS, Ousey JC et al. Maturation of pancreatic beta-cell function in the fetal horse during late gestation. J Endocrinol 2005;186:467–73. 25. Holdstock NB, Allen VL, Bloomfield MR, Hales CN, Fowden AL. Development of insulin and proinsulin secretion in newborn pony foals. J Endocrinol 2004;181(3):469–76. 26. Roberts MC. The development and distribution of mucosal enzymes in the small intestine of the fetus and young foal. J Reprod Fertil Suppl 1975;23:717–18. 27. Rice L, Ott EA, Beede DK, Wilcox CJ, Johnson EL, Lieb S, Borum P. Use of oral tolerance tests to investigate disaccharide digestion in neonatal foals. J Anim Sci 1992;70: 1175–81. 28. Crowell-Davis SL, Houpt KA. Coprophagy by foals: Effect of age and possible functions. Equine Vet J 1985, 17(1): 17–19. 29. Hornicke H, Bjornhag G. Coprophagy and related strategies for digesta utilization. In: Ruckebusch Y, Thiven P (eds) Digestive Physiology and Metabolism in Ruminants. MTP Press, 1979; pp. 707–30. 30. Doreau M, Boulot S, Bauchart D, Barlet JP, Martin-Rosset W. Voluntary intake, milk production and plasma metabolites in nursing mares fed two different diets. J Nutr 1992;122:992–9. 31. Oftedal OT, Hintz HF, Schryver HF. Lactation in the horse: Milk composition and intake by foals. J Nutr 1983;113:2096–105. 32. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Colostral volume and immunoglobulin G and M determination in mares. Am J Vet Res 1989;50:466–70. 33. Paradis MR. Nutrition and indirect calorimetry in neonatal foals. Proc ACVIM, Denver, CO. 2001;19:245–7. 34. Rooney DK. Clinical nutrition. In: Equine Internal Medicine, eds. Reed and Bayly, Philadelphia: Saunders, 1998; pp. 216–50. 35. Grace ND, Pearce SG, Firth EC, Fennessy PF. Concentrations of macro- and micro-elements in the milk of pasturefed thoroughbred mares. Aust Vet J 1999;77(3):177–80.
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Enteral and Parenteral Nutrition for the Neonatal Foal 36. Okumura M, Asano M, Tagami M, Tsukiyama K, Fujinaga T. Serum copper and ceruloplasmin activity at the early growing stage in foals. Can J Vet Res 1998;62(2):122–6. 37. Brommer H, van Oldruitenborgh-Oosterbaan MM. Iron deficiency in stabled Dutch warmblood foals. J Vet Intern Med 2001;15(5):482–5. 38. Knight DA, Weisbrode SE et al. The effects of copper supplementation on the prevalence of cartilage lesions in foals. Equine Vet J 1990;22(6):426–32. 39. Fraker PJ, King LE, Laakko T, Vollmer TL. The dynamic link between the integrity of the immune system and zinc status. J Nutr 2000;130(5S Suppl):1399–406S. 40. Hall JA. Antioxidants and immune function. Proc 19th Amer Coll Vet Int Med, 2001;17–19. 41. Bain FT. Management of the foal from the mare with placentitis: A clinician’s approach. Proc Am Assoc Equine Pract 2004;50:162–4. 42. Ousey JC, Holdstock NB, Rossdale PD, McArthur AJ. How much energy do sick neonatal foals require compared with healthy foals? Pferdeheilkunde 1996;12:231–7. 43. Scrimshaw NS. Effect of infection on nutrient requirements. JPEN 1991;15:589–600. 44. Karinch AM, Pan M, Lin CM, et al. Glutamine metabolism in sepsis and infection. J Nutr 2001;131(9 Suppl):2535–50S. 45. Barbosa E, Moreira EAM, Goes JE, Faintuch J. Pilot study with a glutamine-supplemented enteral formula in critically ill infants. Rev Hosp Clin Fac Med Sao Paolo 1999;54(1):21–4. 46. Neu J. Glutamine in the fetus and critically ill low birth weight neonate: Metabolism and mechanism of action. J Nutr 2001;131:2585–89S. 47. Fowden AL, Silver M, Ellis L, Ousey J. Rossdale PD. Studies on equine prematurity 3: Insulin secretion in the foal during the perinatal period. 1984;16(4):286–91. 48. Lawler JB, Hassel DM, Magnusson RJ, Hill AE, McCue PM, Traub-Dargatz JK. Adsorptive effects of di-trioctahedral smectite on Clostridium perfringens alpha, beta, and beta-2 exotoxins and colostral antibodies. Am J Vet Res 2008;69:233–9. 49. Raidal SL, McTaggart C, Penhale J. Effect of withholding macromolecules on the duration of intestinal permeability to colostral IgG in foals. Aust Vet J 2005;83(1–2):78–81. 50. Ousey JC. Feeding the newborn foal in health and disease. Equine Vet Educ Manual 2003;6:50–4. 51. Daels PF, Duchamp G, Proter D. Induction of lactation and adoption of foals by non-parturient mares. Proc Am Assoc Equine Pract 2002;48:68–71. 52. Daels PF, Bowers-Lepore J. How to induce lactation in a mare and make her adopt an orphan foal: What 5 years of experience have taught us. Proc Am Assoc Equine Pract 2007;53:349–53. 53. Chavatte-Palmer P, Arnauld G, Duvaux-Ponter C, Brosse L, Bougel S, Daels P, Guillaume D, Clement F, Palmer E. Quantitative and qualitative assessment of milk production after pharmaceutical induction of lactation in the mare. J Vet Intern Med 2002;16(4):472–7. 54. Ousey JC, Prandi S, Zimmer J, Holdstock N, Rossdale PD. Effects of various feeding regimens on the energy balance of equine neonates. Am J Vet Res 1997;58(11):1243–51.
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55. Weese JS, Parsons DA, Staempfli HR. Association of Clostridium difficile with enterocolitis and lactose intolerance in a foal. J Am Vet Med Assoc 1999;214(2):229–32, 205. 56. Fuller R. Probiotics in human medicine. Gut 1991;32: 439–42. 57. Guarner F, Schaafsma GJ. Probiotics. Int J Food Microbiol 1998;39:237–38. 58. Metchnikoff E. The Prolongation of Life. London: Williams Heinemann, 1907. 59. Shahani KM, Ayebo AD. Role of dietary lactobacilli in gastrointestinal microecology. Am J Clin Nutr 1980;33:2448–57. 60. Weese JS. A review of probiotics: Are they really ‘functional foods’? Proc Am Assoc Equine Pract 2001;47:27–31. 61. Hansen TO. Nutritional support: parenteral feeding. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 747–62. 62. Krause JB, McKenzie III, HC. Parenteral nutrition in foals: a retrospective study of 45 cases (2000–2004). Equine Vet J 2007;39(1):74–8. 63. Spurlock SL, Donaghue S. Weight gains in foals on parenteral nutrition. International Society of Veterinary Perinatology, 2nd Scientific Conference, Cambridge, England. July, 1990; p. 27. 64. McKenzie III HC, Geor RJ. How to provide nutritional support of sick neonatal foals. Proc Am Assoc Equine Pract 2007;53; 202–8. 65. Boitano M. Hypocaloric feeding of the critically ill. Nutr Clin Pract 2006;21:617–22. 66. Koterba AM. Appendix 1: Drug doses. In: Koterba AM, Drummond WH, Kosch PC (eds) Clinical Equine Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 781–9. 67. Palmer JE. Recognition and resuscitation of the critically ill foal. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case-Based Approach. Philadelphia: Elsevier Saunders, 2006; pp. 121–34. 68. Buechner-Maxwell VA. Hyperglycemia in a neonatal foal: Management with continuous insulin infusion. Equine Pract 1994;16:13–15. 69. Ikeda S, Zarzaur BL, Johnson CD, Fukatsu K, Kudsk KA. Total parenteral nutrition supplementation with glutamine improves survival after gut ischemia/reperfusion. J Parenter Enteral Nutr 2002;26(3):169–73. 70. Kang W, Gomez FE, Lan J, Sano Y, Ueno C, Kudsk KA. Parenteral nutrition impairs gut-associated lymphoid tissue and mucosal immunity by reducing lymphotoxin beta receptor expression. Ann Surg 2006;244(3):392–9. 71. Wildhaber BE, Yang H, Spencer AU, Drongowski RA, Teitelbaum DH. Lack of enteral nutrition – effects on the intestinal immune system. J Surg Res 2005;123(1):8–16. 72. Shaw AA, Hall SD, Franklin MR, Galinsky RE. The influence of L-glutamine on the depression of hepatic cytochrome P450 activity in male rats caused by total parenteral nutrition. Drug Metab Dispos 2002;30(2):177–82. 73. Myers CJ, Magdesian KG, Kass PH, Madigan JE, Rhodes DM, Marks SL. Parenteral nutrition in neonatal foals: clinical description, complications and outcome in 53 foals (1995–2005). Vet J 2009;181(2):137–44.
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Chapter 29
Growth of Horses Clarissa Brown-Douglas, Peter Huntington and Joe Pagan
The study of growth An objective of animal study is to determine the effects of one or more variables on the growth of an animal. Growth is measured as change in weight over time, and there are several mathematical equations to explain this. The most usual way is to make a series of measurements as the body accumulates new tissue and to plot these against time. If growth is normal, the curve is a sigmoid curve with the slope of the curve being the accumulative growth rates.1 Animal growth is often studied using mathematical equations to fit growth curves. A growth model is a function describing how an animal grows and is derived from the mathematics of the physical process producing the data. Growth models include factors that affect growth, such as birthweight and gender. Growth models are usually three or four parameter non-linear exponential equations, with an inflection point coinciding with the time or age of maximum growth rate.2 Growth curves have many practical applications including:
r r r r
Measurements of the performance of an animal or group of animals with respect to time; Comparison of the growth of individuals to an expected ‘normal’ growth curve; Use in research to determine different animal responses to different nutritional trials and environmental conditions; and A predictor of future performance where no animal measurements or data may exist.2
Equine growth studies Studies of equine growth have not been extensive because the horse is not primarily a meat-producing animal. In meat-producing systems, high growth rates are considered desirable.3 This is not necessarily true in the horse, since rapid growth in the immature
horse is associated with rapid weight gain and compromised skeletal growth, predisposing the animal to developmental orthopedic disease.4–7 The regular weighing of horses is recommended,8 but is not routinely performed with most growing horses. The monitoring of an animal’s weight gain against a normal curve allows growth rates to be controlled, which is important in the management of young horses and may reduce the incidence of developmental orthopedic disease.7 Observed measurements of equine growth include bodyweight, wither and hip height, and measurement of defined body parts such as cannon bone circumference and chest width. Weight has been expressed as absolute weight in pounds (lb) or kilograms (kg)8–14 and growth rate expressed as average daily gain (ADG)8, 13, 14 or percentage of increase (%).15 Height and cannon bone circumference have also been documented,8,10–12,16, 17 as have linear measurements to assess bodyweight18 and conformation.19, 20 Condition scoring, a subjective measure of fat deposition described by Henneke et al.21 and Carroll and Huntington,22 is often used to describe body condition in growing and mature horses. Thoroughbred growth data have primarily been collected from horses raised on commercial farms.8, 9,11–14,16, 23 However, there have been some studies on horses raised at pasture.24–27 There have been few growth studies on non-Thoroughbreds, although there have been some data collected from Quarter Horses,28–30 Warmbloods,31–35 Arabians,36 Lusitanos,37 Lipizzaners,32 and ponies.38 In general, colts are heavier than fillies at equivalent ages8, 11, 12 although there was no difference in birthweights of Thoroughbred foals in one UK study.16 The mature weight of the Thoroughbred horse is assumed to be between 500 and 600 kg. Pagan et al.8 measured 472 brood mares and 25 breeding stallions and reported a mature bodyweight of 570 kg and 580 kg for mares and stallions, respectively, and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Growth of Horses concluded a mature bodyweight for a Thoroughbred in Kentucky is 575 kg. Mature bodyweights for other breeds as reported in the NRC (2007)39 are Hanoverian, 580 kg; Swedish Standardbred, 500 kg; Quarter Horse, 555 kg; Belgian draft, 863 kg; Morgan, 454 kg; Arabian, 455 kg; and pony, 195 kg.
Factors affecting equine growth The genetic make-up, or genotype, determines a horse’s mature size, but many factors influence growth rate including the environment, nutrition, and management. Gestation, birthweight, and the early growth period Growth from conception proceeds through a process of cell division and hypertrophy (enlargement), with the formation of various embryonic tissues,40 with growth in early gestation comparatively slow compared with the last trimester. At 7 months of gestation a fetal foal is a mere 20% of birthweight, less than 2% of the mare’s bodyweight;40 thus the majority of fetal growth occurs in the final months of gestation. Different tissues have varied periods of deposition both before and after birth (Figure 29.1). Birthweight of the light horse foal is on average 10% of mature bodyweight,40, 41 whereas birthweight of draft horses and Shetland ponies average 7% and 13% of mature weight, respectively.42 At birth, foals have only achieved 17% of mature bone mineral content.41 Birthweight has a significant ongoing influence on bodyweight in Thoroughbreds.27 Foals with higher birthweights are heavier throughout life, and there is one report that low-birthweight Thoroughbred foals (<40 kg) were less likely to race, and have inferior racing performance characteristics compared to heavier foals.43 As a result, breeders aim to pro-
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duce heavy foals at birth. Several management practices are used by breeders to increase the chances of a heavy foal. These include correct supplementary feeding of mares during pregnancy and selecting for large mares. Birthweight of the foal is on average 10% of mare bodyweight; therefore, larger mares commonly deliver larger foals.9, 40, 43, 44 Mare age and parity has been shown to affect foal size as mares between the ages of 7 and 11 years have foals of heavier birthweight, and foals born to maiden mares have lower birthweights than those from multiparous mares.11, 30, 45 Foal birthweight is a reflection of the balance between feto-maternal contact and placental efficiency. Both maiden and aged mares have lower microcotyledon surface density than multiparous mares of intermediate age. This may be a factor in the lower birthweights, along with lower levels of serum insulin-like growth factor-1 in foals of maiden mares, that can influence growth before and after birth.46 Wilsher and Allen47 fed maiden Thoroughbred mares a maintenance or high-energy diet (2.5–3 × maintenance) during pregnancy to compare the effect of plane of nutrition on birthweight and foal development. The experiment was compromised by a strangles outbreak in mid-gestation, but it found that a very high plane of nutrition and having mares fat before foaling increased placental area and fetal fluid volume, but not birthweight, when compared with a control group of maiden mares from the previous experiment.45 It was apparent that a strangles infection in mid-pregnancy led to substantial maternal weight loss (average 10%) and had a significant effect on reduced placental area and efficiency, along with lower birthweight and length of the foals. In humans it is thought that nutritional insults in mid-gestation can affect fetal length, and similar insults in late gestation can reduce birthweight. The
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Figure 29.1 Normal growth curve for the horse of mature weight of 500 kg. From Frape 199840 with permission
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Table 29.1 Bodyweight and height at birth, 6, 12, 18, and 24 months and their percentages of mature weight and height in Kentucky Thoroughbreds, assuming a mature weight of 575 kg and mature height of 163 cm
Age
Bodyweight (kg)
% Mature weight
Wither height (cm)
% Mature height
55 250 350 450 550
10 43 61 78 96
103 136 150 155 160
63 83 92 95 98
Birth 6 months 12 months 18 months 24 months
question of optimal weight gain during late pregnancy was studied by Powell et al.48 and Kolwaski et al.49 who found that mares can have foals of normal birthweight despite gaining little weight during the last 90 days of gestation. However, the optimal weight gain in late pregnancy has not been determined and Baker (personal communication) found that Thoroughbred mares foaling later in the season and gaining 80 kg in the last 60 days of pregnancy had heavier foals than those foaling earlier and gaining less weight. Rapid growth occurs during the first year of life. At 6 months of age, foals have achieved 43% of mature weight and 83% of mature height, and by 12 months of age foals have reached 61% of mature weight and 92% of mature height. By 24 months of age horses have reached 96% of mature weight and 98% of mature height (Kentucky Equine Research, unpublished data) (Table 29.1). Regardless of breed or geographical location, horses show a similar pattern of growth between birth and 18 months of age. The most rapid growth occurs in the first month of life, with ADG declining at a constant rate to reach a low at 10–11 months of age.8, 27, 42 ADG between birth and 6 months of age appears to be related to the degree to which their diet is composed of mares’ milk27 as growth rate in young horses is largely determined by the
amount of milk produced by their dams,40 with the peak of lactation occurring at 2 months postpartum and the greatest intakes of milk in the second and third months after birth.50 The observed nadir in growth rate around 10–11 months of age coincides with the winter months of decreased pasture quality and availability. ADG typically increases after 11 months, which has been attributed to the onset of spring pasture growth and possibly the effects of preparation for early season sales.51 Breed does influence growth patterns, with ponies maturing faster than larger horses. Smaller breeds reach a greater proportion of their mature weight or height at a given age than do larger breeds that keep growing for a longer time period (Table 29.2). Birth month and birth season have a significant effect on growth rate in horses. Foals born in the winter are generally smaller than those born later.8, 11 However, other studies have shown no seasonal differences in Quarter Horse foals,52 and early-season Thoroughbred foals were significantly larger than those born in the second half of the breeding season.53 A study of 3909 Thoroughbred mares and their foals in Kentucky has shown that winter-born foals (those born in January and February) were smaller at birth and grew more slowly during the first 2 months compared with spring-born foals.54 These winter-born foals then exhibited rapid ADG at 3 months of age, coinciding with a spring pasture flush. This compensatory growth resulted in there being no difference in bodyweights between foals born in any birth month at 5 months of age. Later-born foals (those born in May and June) exhibited the lowest ADG at 2, 3, and 4 months, which coincides with late summer pasture losing its quality, suggesting a definite seasonal effect on foal ADG. Interestingly, mare bodyweight and condition had a strong effect on foal growth, as foal bodyweight was positively correlated to mare bodyweight, foal ADG was positively associated with mare ADG, and foal fatness (indicated by body condition score or BCS) was positively related to mare BCS during the first 5 months of life. These relationships indicate that heavier mares produce
Table 29.2 Percentage of mature bodyweight and of withers height attained at different ages in various horse breeds. From Frape (1998)40 6 months Age Shetland Pony Arabian Anglo-Arab Quarter Horse Thoroughbred Percheron
12 months
18 months
Weight
Height
Weight
Height
Weight
Height
52 46 45 44 46 40
86 84 83 84 84 79
73 66 67 66 66 59
94 91 92 91 90 89
83 80 81 80 80 74
97 95 95 95 95 92
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Growth of Horses heavier foals, faster-growing foals are from mares that are gaining weight, and fatter foals (those with higher BCS) are produced from fatter mares. In addition, there was a positive relationship between foal BCS and mare daily weight change at 2 and 3 months (peak lactation), indicating that mares that are gaining weight during the first 3 moths of lactation have fatter foals. Regardless of birth month, mares that exhibited a negative energy balance postpartum (losing weight or negative daily weight change) had foals that did not gain as much weight as mares that were in a positive energy balance at any time between birth and 5 months postpartum.54 January- and Februaryfoaling mares tended to lose weight in early lactation, suggesting that their caloric intake is insufficient to meet their energy requirements for early lactation. Later in lactation, when the climate was warmer and there was access to adequate pasture, these earlyfoaling mares gained more weight and supported faster growth rates in their foals than later-foaling mares. The weanling to yearling growth period From 6 months of age, growth rate is also reported to be a function of seasonal factors. Thoroughbred weanlings in Kentucky exhibited a seasonal growth pattern, such that growth rates, regardless of birth month, were low during winter and increased during spring.8 A similar pattern was shown in New Zealand Thoroughbreds born in the spring and autumn.27 Both spring-born and autumn-born foals exhibited a similar pattern of growth during the first 6 months of age, and showed an increase in ADG during the spring when spring- and autumn-born horses were 12 months and 8 months of age, respectively.27 Weaning can have a significant influence on growth rate, and several studies report a check in ADG immediately following weaning, regardless of age.27, 55 Foals weaned at 4.5 months and 6 months both had lower ADG at 1 and 3 weeks post-weaning, but foals weaned at the younger age (4.5 months) had depressed cannon bone growth whereas those weaned at 6 months of age did not.55 Weaning can be managed to produce as little stress on the foals as possible, and anecdotal evidence suggests that staggered weaning, when the mares of the oldest foals in a group are removed one by one over time, is the least stressful on the foals. During the spring following the first winter of life, most horses will exhibit an increase in ADG. In spring-born horses this will generally occur between 13 and 16 months of age. The rapid increase in ADG has been attributed to the increase in pasture growth and quality during the spring; however, another fac-
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tor contributing to the increase in growth rates may be a peripubertal growth spurt, such as is known to occur in children.56, 57 Onset of puberty in horses occurs in the first spring of life and is related to seasonal increase in daylength.58 Pubertal growth spurts in horses have not been described; therefore it is difficult to determine whether the increased ADG observed in the spring is totally or partly related to puberty, or by the changes in nutrient supply. Interestingly, a large study observing the growth of Thoroughbred populations raised in various countries including North America, England, India, Australia, and New Zealand showed that, although the breed has a very narrow genetic base, significant differences in growth are observed depending on the location in which the horses were raised.14 In the study, which examined the growth of over 13400 Thoroughbreds, it was found that Australian and New Zealand Thoroughbreds were heavier and taller than North American Thoroughbreds, which in turn were larger (heavier and taller) than those reared in England. Thoroughbreds raised in India were significantly smaller (lighter and shorter) than all other Thoroughbred populations studied (Figures 29.2 and 29.3). These observed differences can be attributed to management of the horses during the growth period. There are several factors that may have contributed to making Australian and New Zealand Thoroughbreds larger than horses in other countries. The temperate climate in which most horses are reared allows mares and foals to remain outdoors on pasture in late winter and early spring, with access to early spring pasture plus supplementary feed to supply energy and protein requirements for milk production. This helps mares maintain or gain weight after foaling and maintain milk production at high levels. It has 500 450 400 Body weight (kg)
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Australia
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England
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India Kentucky
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New Zealand
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World Average
0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 Age (months)
Figure 29.2 Bodyweight ± 95% CI (kg) of Thoroughbreds reared in Australia, England, India, America, and New Zealand compared with the world average. From Huntington et al. (2007)59
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Figure 29.3 Wither height ± 95% CI (kg) of Thoroughbreds reared in Australia, England, India, America, and New Zealand compared with the world average. From Huntington et al. (2007)59
recently been shown that mares foaling early in the season in Kentucky tend to lose weight, and foals do not grow as fast in this period as their contemporaries born in the spring when mares are grazing spring pasture.54 In New Zealand some mares rear foals on pasture alone without supplementary feed, and in Australia, irrigation is commonly used on Thoroughbred studs to provide adequate pasture for mares and foals. As yearlings, Australasian Thoroughbreds were heavier and taller than Thoroughbreds in all other countries. Climatic and pasture quality differences in late winter and early spring contribute to this situation, and the other factor is the presence of yearling sales earlier in the year. The first yearling sales occur in January, when the oldest yearlings will be 17 months of age, whereas in the northern hemisphere the first sales are in September when the oldest yearlings could be 21 months of age. At 18 months, Australasian Thoroughbreds are significantly heavier than Thoroughbreds in all other countries. This is most likely representative of the difference in management of yearlings prepared for sale. Australasian yearlings tend to be sold at sales in a well-rounded condition with higher condition scores compared with American and English yearlings which are presented as leaner, more athletic and fit. During an 8- to 10-week yearling preparation, ADG of up to and over 1 kg per day are common as the market demands big horses carrying plenty of muscle and fat cover. This trend is beginning to change as breeders and buyers realise the potential musculoskeletal problems associated with over-conditioned yearlings. In addition, these observations may further be attributed to the management of growing horses for different racing industries under different grow-
ing conditions. In general, the North American racing industry has significantly more short-distance dirt races than the industries of England, Australia, and New Zealand and, as a result, North American Thoroughbreds are raised to be precocious sprinting horses ready to race at 2 years old with a comparatively large muscle mass and a small refined bone structure. In contrast, the English racing industry holds turf races, commonly of distances greater than 1 mile, and as a result England has traditionally produced slower-maturing Thoroughbreds with larger frames but comparatively less muscle-bulk than sprinting horses. Australia and New Zealand produce turf horses with a premium on early maturity and 2-year-old racing; however, these southern hemisphere horses are traditionally known for their size, which may be attributed to genetics, even though shuttle stallions have had a significant impact on evening out the genetic pool between hemispheres.59 In the study, Thoroughbreds in all countries except for India showed low ADG around 10 to 11 months and increases in ADG between 13 and 16 months.51 As these horses were all born in the spring and early summer, the decline in growth rate around 10 and 11 months of age coincides with the winter months of decreased pasture quality and availability. The increase in ADG after 11 months of age in these horses could be attributed to the onset of spring pasture growth. The ADG of Indian Thoroughbreds declined steadily between 1 and 18 months, did not increase between 10 and 14 months of age, and were significantly less then the rest of the world at 13–15 months of age (P < 0.05). The significant differences in growth of Indian Thoroughbreds could be attributed to differences in nutritional management as well as climatic conditions in that country. In contrast to horses in the other countries in this study, many Indian horses have restricted or no access to pasture and are reported to spend a significant amount of time indoors due to extreme heat and monsoon rains, which may hinder growth potential of these horses. Indian foals were significantly smaller on average than foals in other countries and it has been demonstrated previously that the size of the foal is a key influence on the size of the yearling. There are no reported data on the negative effect of extremely hot temperatures on the growth rates of horses; however, feed intakes were reported to be negatively related to environmental temperature in weanling horses housed in cold climates, presumably adversely affecting growth rate. For example, winter growth rates in Canadian Thoroughbreds are often half those seen in Florida.60
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Growth of Horses Effect of exercise Forced exercise during the growth period is thought to have little effect on growth rate and ultimate body size.61–63 It is understood that lack of exercise and confinement are contributing factors to growth-related skeletal disorders in young growing horses.64, 65 Weanlings that underwent forced exercise up to yearling age had increased cannon bone density when compared to non-exercised horses,63 indicating that exercise during growth and development might improve the stress-bearing characteristics of young bone.
Optimal growth rates and growth spurts Optimal growth rate results in a desirable body size at a specific age with the least amount of developmental problems. Managing the growth in horses becomes a balance between producing a desirable individual for a particular purpose without creating skeletal problems that will reduce a horse’s subsequent athletic ability (Figure 29.4). Growing a foal too slowly results in the risk of it being too small at a particular age or never obtaining optimal mature body size. Growing a foal too quickly results in the risk of developmental orthopedic disease such as epiphysitis, angular limb deformities, and osteochondritis dissecans. It is now commonly accepted that excess dietary energy intake can lead to rapid growth and increased body fat, which is thought to increase the incidence of certain types of developmental orthopedic disease in Thoroughbred foals. Thoroughbred yearlings in a Kentucky study that showed OCD of the hock and stifle were large at birth, grew rapidly from 3 to 8 months, and were heavier than the average population as weanlings.7 Vervuert et al.35 found that Warmblood foals which developed fetlock OCD were significantly lighter at 3–7 months of age and those with hock and stifle OCD had higher bodyweights in the fourth and fifth months of life. In con-
Figure 29.4 Distribution of growth-related problems during slow and fast growth. From Pagan 2005b69 With permission from Nottingham University Press
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trast, Savage et al.6 fed weanlings 130% energy requirement and induced a significantly higher incidence of dyschondroplasia joint lesions than control weanlings, without any change in growth rate. In addition to rapid growth rates, fluctuating growth rates with periods of slow or decreased growth followed by growth spurts may also be a contributing factor to developmental orthopedic disease.66 Non-uniform growth rates can occur due to dietary and environmental stress,60, 67, 68 season,8, 13, 14 and possibly puberty.15, 27 Therefore, it has been widely recommended to maintain a steady growth rate by regularly weighing and measuring horses during the growth period.39, 69 As there is no single growth rate that is desirable for all types of horse, horses should be managed differently for varying growth rates.
Is growth important to a horse’s career? Growth management of the young foal will vary depending on the commercial end point. As horses are bred and raised primarily for the enjoyment and service of humans, the purpose for which one breed of horse is raised may have no relationship to the primary purpose of another breed. There are not only differences between breeds in use or function, but also differences within breeds, and because of this there are widely different nutritional management practices to achieve the different commercial end points. For breeds that are not expected to be competitive until 4–5 years of age, such as Warmbloods, there is little incentive for rapid early growth. Instead, a slow steady growth rate that will allow the horse to reach maximal mature body size with the fewest problems as possible is desirable. For Thoroughbreds and Quarter Horses, which are expected to be successful in the sales rings as weanlings and yearlings, as well as competitive athletes as young as 18 months of age, the early growth phase is extremely important. A large, well-grown Thoroughbred yearling is desirable when offered for sale at public auction, as selling price is influenced by body size. Yearlings that sold higher than the median of the session in which they were sold were heavier and taller, but not fatter, than yearlings which sold below the session median.70, 71 Yearlings that sold were also heavier and taller than those listed as RNA (reserve not attained), and fewer lightweight yearlings (those in the lowest weight quartile) were sold compared with the heaviest yearlings.71 Early rapid growth may not be favored in Thoroughbreds that are ‘bred to race’ with no intention of being prepared for sale as weanlings or yearlings. Many breeders selling horses at public
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auction aim to produce early-born foals to maximize growing time before a July or September sale. This practice may not be that advantageous as foals born in the central Kentucky winter (January and February) were smaller at birth and grew more slowly during the first 2 months compared with spring-born foals, but compensated by growing faster later in lactation so that by 5 months of age there was no difference between bodyweights in any of the groups.54 April- and May-born yearlings at 16 months of age are also taller and heavier than winter-born yearlings.51 September sale yearlings are between 16 and 20 months of age, so buyers may mistake January-born yearlings for the largest when in fact they are only the oldest. Furthermore, recent research at Kentucky Equine Research has shown that yearling size does not necessarily predict athletic success.51 The growth and racing performance records of nearly 4000 American Thoroughbreds were analyzed to determine if certain growth characteristics affect the odds of success as a racehorse. Findings suggested some significant trends in growth and racing success. Horses that started as 2-year-olds were shorter and weighed less as foals and yearlings than those that did not, regardless of birth month. However, fewer May-born yearlings started as 2-year-olds compared with earlier-born yearlings. It is generally accepted that faster-maturing, heavier horses are more likely to be raced as 2-year-olds, but these results suggest just the opposite: smaller horses are more likely to run early. Furthermore, smaller horses had more career starts, suggesting that smaller horses may be sounder. Conversely, stakes winners, graded stakes winners, grade-1 stakes winners, and millionaires were heavier and taller than average as yearlings. Interestingly, the 21 millionaires in the study were, on average, taller than 79% of the population as yearlings. To further evaluate the relationship between size and racing performance, the yearlings were divided into four groups based on weight and height (quartiles) so that the lightest or shortest yearlings were in the lowest quartile, and the heaviest and tallest in the highest quartile. This method allowed comparison among all horses regardless of age or gender, and ranks the relative position of an individual in a population by indicating what percentage of the reference population that individual will equal or exceed for each measurement. Yearlings in the lowest weight quartile (those that weighed less than 75% of the population) had lower earnings, fewer stakes wins, and a lower sire index (sire index indicates the average racing class of foals sired by a stallion) than the rest of the population. However, yearlings below the 50th weight and height percentiles were more likely to
start as 2-year-olds and had more career starts than those above the 50th percentile. These data provide insight into managing horses for different strategies. Smaller horses are more likely to start as 2-year-olds and have more career starts; however, elite performers (graded stakes winners, grade-1 stakes winners, and millionaires) tend to be taller and heavier as yearlings.
Fueling growth Daily requirements of energy, protein, minerals, and vitamins vary depending on the growth rate and size of the individual. Because of the premium price paid for mass, young Thoroughbreds and Quarter Horses prepared for sale are grown rapidly to achieve maximal size. To fuel maximum growth, these young horses are often fed large amounts of grain. Extremely rapid growth by overfeeding energy has been implicated in developmental orthopedic disease.6 Periods of slow or decreased growth followed by growth spurts are also risky. Therefore, nutrition mistakes made during a foal’s early growth can lead to developmental problems that limit performance later in life. The best way to evaluate the success of a feeding and conditioning program of young horses is through assessment of bodyweight, height, and condition. Regular monitoring of weight allows farm managers to maintain a steady growth rate while preventing the animal from becoming too heavy.
Broodmare nutrition during pregnancy and lactation Correct nutrition of the broodmare is vital for successful growth and development of the foal. The nutritional requirements of the broodmare vary depending on her physiological state and can be divided into three stages: early pregnancy (conception until 7 months of gestation as well as barren mares and pregnant mares with no foal at foot), late pregnancy (7 months of gestation to foaling), and lactation.65 Contrary to popular belief, the fetus does not grow at a constant rate throughout the entire 11 months of gestation (Figure 29.5). Fetal growth is slow during the first 7 months of gestation, when the fetal foal equals 20% of its birthweight and weighs less than 2% of the mare’s weight. During this phase, the nutrient requirements of the growing foal are tiny compared with the maintenance requirements of the mare.65 Therefore, the mare should be provided with a nutritional regimen that meets her maintenance requirements and it is recommended that mares with
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energy requirement is approximately double that of the mare during early pregnancy, while protein needs are doubled and lysine, calcium, and phosphorus requirements are thought to be over 300% higher.39
Foal and weanling growth
Figure 29.5 Typical growth curve for a fetus expressed as a percentage of birthweight. From Frappe, 1998,40 with permission.
inadequate body condition (CS < 5) in early or midgestation should be fed additional energy to achieve a body condition score of at least 5 by the ninth month of gestation.39 After 7 months of gestation, fetal growth increases exponentially, and the nutrient requirements of the fetus become significantly greater. Digestible energy, protein, and mineral requirements for the mare all increase during the last months of gestation.39 During the last 4 months of gestation, the fetal and non-fetal tissues accumulate 77 g of protein, 7.5 g of calcium, and 4 g of phosphorus per day.65 In addition to protein and macro-minerals, trace mineral supplementation is also very important during the last trimester of pregnancy because the fetus stores iron, zinc, copper, and manganese in its liver for use during the first few months after it is born.72 This physiological process is because the trace mineral content of mare’s milk is relatively low and does not meet the requirement of the suckling foal.73 Although there is an increased nutrient requirement in late pregnancy, mares are often overfed energy in an attempt to supply adequate protein and minerals to the developing foal, which can lead to obesity.65 Obesity in broodmares, as with any type of horse, should be avoided to reduce issues such as equine metabolic syndrome, insulin resistance, and/or laminitis74 and to reduce the potential for angular limb deformities in their foals. If the pregnant mare becomes fat during late pregnancy, she should be fed a more concentrated feed that supplies the necessary protein and minerals for fetal growth, while limiting energy intake. The energy requirements of the mare increase significantly with the onset of lactation. During the first 3 months of lactation, daily milk production averages approximately 3% of the mare’s bodyweight and during months 4–6 it averages 2% of mare’s bodyweight.39, 65 During early and mid-lactation, the
During the first few months of life, foals grow rapidly, quadrupling their bodyweights by 5 months of age.8, 14, 41 During this time, foals derive the energy, protein, and minerals necessary to support rapid growth from a combination of mare’s milk, pasture, supplemental grain, and mineral stores in the foal’s body. If the broodmare has received a correctly fortified feed during late pregnancy and is producing adequate milk, in most cases it is unnecessary to supplement the foal with grain until it reaches 90 days of age. At this stage, introduction of a correctly fortified ration for growing horses should be introduced and gradually increased so that the foal’s daily grain intake is 0.5 kg feed per month of age.65 If mares are fed their ration in feed bins accessible by the foal, it is an accepted practice that the foal consumes some of the mare’s ration. Proper nutrient intake is vital during the weanling stage as the skeleton is most vulnerable to disease. Nutrition plays an important role in the pathogenesis of developmental orthopedic disease in horses as deficiencies, excesses, and imbalances of nutrients affect the incidence and severity of physitis, angular limb deformity, wobbler syndrome, and osteochondritis dissecans.6, 65,75–77 The most common feeding errors attributed to developmental orthopedic disease are excessive grain intake, feeding an inappropriate grain for the forage being fed, and inadequate fortification of grain. These three scenarios are easily rectified by feeding an appropriate grain mix fortified for the young, growing horse and feeding it at the correct intake. Young horses already suffering from developmental orthopedic disease should have their energy intakes reduced while maintaining correct levels of protein, vitamins, and minerals. In addition to excessive energy, the source of calories for young horses may also be important. Foals that experience an exaggerated and sustained increase in circulating glucose or insulin in response to a carbohydrate (grain) meal may be predisposed to osteochondritis dissecans. Research conducted by Kentucky Equine Research suggests that hyperinsulinemia may influence the incidence of osteochondritis dissecans (surgically treated) in Thoroughbred weanlings.78 In a large field trial, a high glucose and insulin response to a concentrate meal was associated with an increased incidence of osteochondritis dissecans. Plasma glucose and insulin 2 hours
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post-feeding were significantly higher in weanlings with osteochondritis dissecans than in unaffected foals (P < 0.05). Insulin/glucose ratios, however, were not significantly different. The incidence of osteochondritis dissecans was significantly higher in foals whose glucose and insulin values were greater than one standard deviation above the mean for the entire population (both osteochondritis dissecans and unaffected) in the study. Elevated insulin/glucose ratios did not appear to be correlated with an increased incidence of osteochondritis dissecans. Glycemic responses measured in the weanlings were highly correlated with each feed’s glycemic index (GI), suggesting that the GI of a farm’s feed may play a role in the pathogenesis of osteochondritis dissecans. Hyperinsulinemia may affect chondrocyte maturation, leading to altered matrix metabolism and faulty mineralisation or altered cartilage growth by influencing other hormones such as thyroxine.79 Based on the results of this study, it would be prudent to feed young horses concentrates that produce low glycemic responses such as feeds in which a proportion of the energy is supplied by fat and fermentable fiber sources (beet pulp and soy hulls). Mineral deficiency and/or excess of calcium, phosphorus, copper, and zinc may lead to developmental orthopedic disease.65, 75,80–84 The ration of a growing horse should be properly fortified because commonly fed cereal grains and forages contain insufficient quantities of several important minerals. A ration of grass hay and oats would only supply about 40% and 70% of a weanling’s calcium and phosphorus requirement, respectively, and less than 40% of its requirement for copper and zinc. It is important to provide a balanced ration as the ratio of certain minerals (calcium to phosphorus; copper to zinc) is extremely important. The ideal ratio of calcium to phosphorus in the ration of young horses is 1.5 : 1 and should never fall below 1 : 1 or exceed 2.5 : 1. Too much calcium may affect phosphorus status, particularly if the level of phosphorus in the ration is marginal; conversely, high levels of phosphorus in the ration will inhibit the absorption of calcium and will lead to a deficiency, even if the amount of calcium present was normally adequate. Forage diets with high calcium levels should be supplemented with phosphorus. The ratio of zinc to copper should be 3 : 1 to 4 : 1. The NRC39 has published daily nutrient requirements for young, growing horses.
Yearling growth The amount of supplementary grain required to maintain a desired growth rate varies, and it is important to feed each horse as an individual. ‘Easykeeping’ yearlings should be prevented from be-
coming fat by being fed a low-intake, low-calorie source of essential protein, vitamins, and minerals. On the other hand, yearlings that are large-framed with much growth potential can consume normal amounts of fortified concentrate. Most types of developmental orthopedic disease are unlikely to arise after 12 months of age. Lesions that become clinically relevant after this age have typically been formed at a younger age; nevertheless, correct nutrient balance is important in the growing yearling.65 Yearlings being prepared for sale are often fed large amounts of energy during the months before the sale to maximise size, which is favored by buyers.71 There are three different growth rates that breeders can use for their growing horses depending on when the horse is born and when or if it is sold at sale.69 A pattern of slow, early growth is more appropriate for foals that will not be sold as yearlings. Delaying rapid growth until after the ‘window of vulnerability’ for bone and joint disease (<12 months of age) significantly reduces the risk of growth-related developmental orthopedic disease.85 Foals that are early born and those that will be sold at later yearling sales could follow a moderate growth curve. This growth pattern is most commonly used for Thoroughbreds in Kentucky and results in a large, wellgrown yearling with minimal joint problems.69 A rapid growth curve, in which weight gain is concentrated over a short time period, is used for later-born foals targeted at early yearling sales. This growth pattern is most likely to result in skeletal problems, but can result in more mature yearlings earlier in the season if properly managed. Rapid growth patterns can be successfully managed by spreading the growth over several months rather than trying to add the bodyweight gain during the traditional sales preparation period of 60–90 days before a sale. Ongoing research suggests a relationship between the glycemic nature of feed and the incidence of skeletal problems in young horses. Diets high in starch and sugar (typical grain-based concentrates) produce a large blood glucose and insulin response after feeding and have been implicated in the etiology of developmental orthopedic disease.86, 87 This, in addition to the work on hyperinsulinemia in weanlings,78 indicates that it may be beneficial to replace a significant portion of the energy normally supplied by grain with fat and fermentable fiber to reduce the glycemic response to concentrate ration.65, 88
References 1. Davies AS. The animal and its growth: growth changes in the meat carcass. In: Meat Production and Processing. New Zealand Society of Animal Production, Palmerston North, New Zealand, 1989; pp. 61–72.
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Growth of Horses 2. Bridges TC, Turner LW, Gates RS, Smith EM. Relativity of growth in laboratory and farm animals. In: Representation of physiological age and growth rate time constant. Trans Am Soc Agric Eng 2000;43:1803–10. 3. Purchas RW, Butler-Hogg BW, Davies AS. Section 1: The product and the market. In: Meat Production and Processing. New Zealand Society of Animal Production, Palmerston North, New Zealand, 1989; pp. 1–11. 4. Lewis LD. Nutrition of the broodmare and growing horse and its role in epiphysitis. Proc Ann Conv Am Assoc Equine Pract 1980;25:269–96. 5. Thompson KN, Jackson SG, Rooney JR. The effect of above average weight gains on the incidence of radiographic bone aberrations and epiphysitis in growing horses. J Equine Vet Sci 1988;8:383–5. 6. Savage CJ, McCarthy RN, Jeffcott LB. Effects of dietary energy and protein on induction of dyschondroplasia in foals. Equine Vet J Suppl 1993;16:74–9. 7. Pagan JD. The incidence of developmental orthopedic disease (DOD) on a Kentucky Thoroughbred farm. In: Advances in Equine Nutrition I. Nottingham, UK: Nottingham University Press, 1998; pp. 469–75. 8. Pagan JD, Jackson SG, Caddel S. A summary of growth rates of Thoroughbreds in Kentucky. Pferdeheilkunde 1996;12:285–9. 9. McCarthy D, Mitchell J. A study of growth rate in Thoroughbred foals and yearlings. Irish J Agric Res 1974;13:111–17. 10. Hintz HF. Growth rate of horses. Proc Am Assoc Equine Pract Symp 1978;24:455–9. 11. Hintz HF, Hintz RL, van Vleck LD. Growth rate of Thoroughbreds: Effect of age of dam, year and month of birth, and sex of foal. J Anim Sci 1979;48:480–7. 12. Thompson KN. Skeletal growth rates of weanling and yearling Thoroughbred horses. J Anim Sci 1995;73:2513–17. 13. Jelan ZA, Jeffcott LB, Lundeheim N, Osborne M. Growth rates in Thoroughbred foals. Pferdeheilkunde 1996;12: 291–5. 14. Brown-Douglas CG, Pagan JD. Body weight, wither height and growth rates in Thoroughbreds raised in America, England, Australia, New Zealand and India. In: Advances in Equine Nutrition IV. Nottingham, UK: Nottingham University Press, 2009; pp. 213–20. 15. Noguiera GP, Barnabe RC, Verreschi IT. Puberty and growth rate in Thoroughbred fillies. Theriogenology 1997;48:581–8. 16. Green DA. A study of growth rate in Thoroughbred foals. Br Vet J 1969;125:539–46. 17. Green DA. Growth rate in Thoroughbred yearlings and two year olds. Equine Vet J 1976;8:133–4. 18. Staniar WB, Kronfeld DS, Hoffman RM, Wilson JA, Harris PA. Weight prediction from linear measures of growing Thoroughbreds. Equine Vet J 2004;36:149–54. 19. Mawdsley A, Kelly EP, Smith FH, Brophy PO. Linear assessment of the Thoroughbred horse: An approach to conformation evaluation. Equine Vet J 1996;28:461–7. 20. Hunt WF, Thomas VG, Stiefel W. Analysis of video recorded images to determine linear and angular dimensions in the growing horse. Equine Vet J 1999;31:402–10. 21. Henneke DR, Potter GD, Kreider JL A condition score relationship to body fat content of mares during gestation and lactation. In: Proc Equine Nutr Physiol Soc Symp 1981;7:105–10. 22. Carroll CL, Huntington PJ. Body condition scoring and weight estimation of horses. Equine Vet J 1988;20(1): 41–5.
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23. Staniar WB, Kronfeld DS, Treiber KH, Splan RK, Harris PA. Growth rate consists of baseline and systematic deviation components in Thoroughbreds. J Anim Sci 2004;82: 1007–15. 24. Grace ND, Pearce SG, Firth EC, Fennessy PF. Concentrations of macro- and micro-elements in the milk of pasturefed Thoroughbred mares. Austr Vet J 1999;77:177–80. 25. Nash DG, Avery A, Dempsey W, Nash G. Growth and development of young Thoroughbred horses on temperate Australian pastures. Proc 17 Equine Nutr Physiol Symp 2001;17:196–8. 26. Grace ND, Gee EK, Firth EC, Shaw HL. Digestible energy intake, dry matter digestibility and mineral status of grazing Thoroughbred yearlings. N Z Vet J 2002;50:63–9. 27. Brown-Douglas CG, Parkinson TJ, Firth EC, Fennessy PF. Body weights and growth rates of spring- and autumnborn Thoroughbred horses raised on pasture. N Z Vet J 2005;53(3): 326–31. 28. Ott EA, Asquith RL, Feaster JP, Martin FG. Influence of protein level and quality on the growth and development of yearling foals. J Anim Sci 1979;49:620–8. 29. Ott EA, Asquith RL. Influence of level of feeding and nutrient content of the concentrate on growth and development of yearling horses. J Anim Sci 1986;62:290–9. 30. Pool-Anderson K, Raub RH, Warren JA. Maternal influences on growth and development of full-sibling foals. J Anim Sci 1994;72:1661–6. 31. Firth EC, van Weeren PR, Pfeiffer DU, Delahunt J, Barneveld A. Effect of age, exercise and growth rate on bone mineral density (BMD) in third carpal bone and distal radius of Dutch Warmblood foals with osteochondrosis. Equine Vet J Suppl 1999;31:74–8. 32. Rastija T, Baban M, Antunovic Z, Mandic I, Opaˇcak A. A comparison of foals development of the Lipizzaner and Holstein breed. Agric Sci Prof Rev 2004;10(1): 62–7. 33. Valette JP, Robert C, Toquet MP, Denoix JM, Fortier G. Evolution of some biochemical markers of growth in relation to osteoarticular status in young horses: results of a longitudinal study in three breeds. Equine Comp Exer Physiol 2007;4:23–9. 34. Valette JP, Robert C, Denoix JM. Use of linear and nonlinear functions to describe the growth of young sport- and race-horses born in Normandy. Animal 2008;2:560–5. 35. Vervuert I, Borchers A, Granel M, Winkslett S, Christman L, Distl O, Bruns E, Hertsch B, M Coenen. Estimate of growth rates in Warmblood foals and the incidence of osteochondrosis. Pferdeheilkunde 2005;21:129–30. 36. Reed KR, Dunn NK. Growth and development of the Arabian horse. Proc Equine Nutr Physiol Soc Symp 1977; pp. 76–98. 37. Fradinho MJ, Ferreira-Dias G, Mateus L, Santos-Silva MF, Agr´ıcola R, Barbosa M, Abreu JM. The influence of mineral supplementation on skeleton formation and growth in Lusitano foals. Livestock Sci 2006;104:173–81. 38. Jordan RM, Myers V. Effect of protein levels on the growth of weanling and yearling ponies. J Anim Sci 1972;34:578–81. 39. NRC (National Research Council). Nutrient Requirements of Horses, 6th edn. Washington, DC: National Academies Press, 2007. 40. Frape D. Equine Nutrition and Feeding, 2nd edn. Oxford: Blackwell Science Ltd, 1998. 41. Lawrence LA. Principles of bone development in horses. In: Advances in Equine Nutrition III. Nottingham, UK: Nottingham University Press, 2005; pp. 289–94.
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42. Martin-Rosset W. Growth and development in the equine. In: Juliand V, Martin-Rosset W (eds) The Growing Horse: Nutrition and Prevention of Growth Disorders. EAAP publication no. 114. Wageningen: Wageningen Academic, 2005; pp. 15–50. 43. Platt H. Growth and maturity in the equine fetus. J R Soc Med 1978;71:658–61. 44. Walton A, Hammond J. The maternal effects on growth and conformation in Shire horse-Shetland Pony crosses. Proc R Soc B 1938;125:311–35. 45. Wilsher S, Allen WR. The effects of maternal age and parity on placental and fetal development in the mare. Equine Vet J 2003;35:476–83. 46. Cymbaluk N, Laarveld B. The ontogeny of serum insulinlike growth factor-1 concentrations in foals: effect of dam parity, diet, and age at weaning. Domest Anim Endocrinol 1996;13:197–209. 47. Wilsher S, Allen WR. The effects of Streptococcus equi infection-mediated nutritional insult during mid gestation in primiparous Thoroughbred fillies. Part 1: Placental and foetal development. Equine Vet J 2006;38:549–57. 48. Powell D, Lawrence L, Parrett D, DiPietro J. Body composition changes in broodmares. Proc 11th Equine Nutr Physiol Soc 1989;11:91–4. 49. Kowalski J, Williams J, Hintz HF. Weight gains of mares during the last trimester of gestation. Equine Pract 1990;12(7): 6–10. 50. Oftedal OT, Hintz HF, Schryver HF. Lactation in the horse: milk composition and intake by foals. J Nutr 1983;113:2096–106. 51. Brown-Douglas CG, Pagan, JD, Stromberg AJ. Thoroughbred growth and future racing performance. In: Advances in Equine Nutrition IV. Nottingham, UK: Nottingham University Press, 2009; pp. 231–45. 52. Huesner GL. The effect of month of birth on the size of foals. J Equine Vet Sci 1992;12(5): 297–9. 53. Thompson KN, Smith BP. Skeletal growth patterns of Thoroughbred horses. J Equine Vet Sci 1994;14(3): 148–51. 54. Pagan JD, Brown-Douglas CG, Caddel S. Body weight and condition of Kentucky Thoroughbred mares and their foals as influenced by month of foaling, season and gender. In: Advances in Equine Nutrition IV. Nottingham, UK: Nottingham University Press, 2009; pp. 137–45. 55. Warren LK, Lawrence LM, Griffin AS, Parker AL, Barnes T, Wright D. The effect of weaning age on foal growth and bone density. Proc 15th Equine Nutr Physiol Soc 1997;15: 65–70. 56. Tanner JM. Growth and maturation during adolescence. Nutr Rev 1981;39:43–55. 57. Rogol AD. Management of puberty in growth hormone deficient children. Endocrine J Suppl 1996;43:S5–11. 58. Brown-Douglas CG, Firth EC, Parkinson TP, Fennessy PF. Onset of puberty in pasture-raised Thoroughbreds born in spring and autumn. Equine Vet J 2004;36:499–504. 59. Huntington PJ, Brown-Douglas CG, Pagan JD. Growth of Australian Thoroughbreds compared with horses in New Zealand, America, England and India. Austr Equine Vet 2007;26(1): 80–92. 60. Cymbaluk NF, Christison GI. Effects of diet and climate on growing horses. J Anim Sci 1989;67:48–59. 61. Rogers CW, Firth EC, McIlwraith CW, Barneveld A, Goodship AE, Kawcak CE, Smith RK, van Weeren PR. Evaluation of a new strategy to modulate skeletal development in Thoroughbred performance horses by imposing
62.
63.
64.
65.
66.
67.
68. 69.
70.
71.
72.
73. 74.
75.
76.
77.
78.
79.
track-based exercise during growth. Equine Vet J 2008;40(2): 111–18. Orton RK, Hume ID, Leng RA. Effects of level of dietary protein and exercise on growth rates of horses. Equine Vet J 1985;17(5): 381–5. Raub RH, Jackson SG, Baker JP. The effect of exercise on bone growth and development in weanling horses. J Anim Sci 1989;67:2508–14. Owen JM. Abnormal flexion of the corono-pedal joint or ‘contracted tendons’ in unweaned foals. Equine Vet J 1975;7:40–5. Pagan JD. The role of nutrition in the management of Developmental Orthopedic Disease. In: Advances in Equine Nutrition III. Nottingham, UK: Nottingham University Press, 2005; pp. 417–31. Barneveld AR, van Weeren R. Conclusions regarding the influence of exercise on the development of the equine musculoskeletal system with special reference to osteochondrosis. Equine Vet J Suppl 1999;31:112–19. Hintz HF, Schryver HF, Lowe JE. Delayed growth response and limb conformation in young horses. In: Proceedings of the Cornell Nutrition Conference, New York, 1976; pp. 94–6. Rooney JR. Weather factors and the growth of young Thoroughbred horses. J Equine Vet Sci 1984;81:106–7. Pagan JD. Managing growth for different commercial end points. In: Advances in Equine Nutrition III. Nottingham, UK: Nottingham University Press, 2005; pp. 319–26. Pagan JD, Koch A, Caddel S, Nash, D. Size of Thoroughbred yearlings presented for auction at Keeneland sales affects selling price. Proc 19th Equine Sci Soc Symp 1998;19:234–5. Brown-Douglas CG, Pagan JD, Koch A, Caddel S. The relationship between size at yearling sale, sale price and future racing performance in Kentucky Thoroughbreds. Proc Equine Sci Soc Symp 2007;20:153–4. ¨ Meyer H, Ahlswede L. Uber das intrauterine Wachs¨ tum und die Korperzusammensetzung von Fohlen sowie ¨ den N¨ahrstoffbedarf tragender Stuten. Ubers Tierern¨ahrg 1976;4:263–92. ¨ Meyer H. Cu-Stoffwechsel und -bedarf des Pferdes. Ubers Tierern¨ahrg 1994;22:363–94. Geor RJ, Harris P. Dietary management of obesity and insulin resistance: Countering risk for laminitis. Vet Clin North Am Equine Pract 2009;25(1): 51–66. Savage CJ, McCarthy RN, Jeffcott LB. Effects of dietary phosphorus and calcium on induction of dyschondroplasia in foals. Equine Vet J Suppl 1993;16:80–3. Jeffcott LB. Developmental diseases affecting growing horses. In: Juliand V, Martin-Rosset W (eds) The Growing Horse: Nutrition and Prevention of Growth Disorders. EAAP publication no. 114. Wageningen: Wageningen Academic, 2005; pp. 243–56. McIllwraith CW. Developmental orthopedic disease in horses: a multifactorial process. Proc Equine Nutr Physiol Soc 2001;17:2–23. Pagan JD, Geor RJ, Caddel SE, Pryor PB, Hoekstra KE. The relationship between glycemic response and the incidence of OCD in Thoroughbred weanlings: A field study. Proc Am Assoc Equine Pract 2001;47:322–5. Jeffcott LB, Henson FM. Studies on growth cartilage in the horse and their application to aetiopathogenesis of dyschondroplasia (osteochondrosis). Vet J 1998;156: 177–92.
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Growth of Horses 80. Bridges CH, Moffitt PG. Influence of variable content of dietary zinc on copper metabolism in weanling foals. Am J Vet Res 1990;51:275–80. 81. Gunson DE, Kowalczyk DF, Shoop CR, Ramberg CF. Environmental zinc and cadmium pollution associated with generalized osteochondrosis, osteoporosis, and nephrocalcinosis. J Am Vet Med Assoc 1982;180:295–9. 82. Hurtig MB, Green SL, Dobson H, Mikuni-Takagaki, Choi J. Correlative study of defective cartilage and bone growth in foals fed a low copper diet. Equine Vet J Suppl 1993;16:66–73. 83. Knight DA, Weisbrode SE, Schmall LM, Reed SM, Gabel AA, Bramlage LR, Tyznik WI. The effects of copper supplementation on the prevalence of cartilage lesions in foals. Equine Vet J 1990;22:426–32. 84. Pearce SG, Firth EC, Grace ND, Fennessy PF. Effect of copper supplementation on the evidence of developmental or-
85.
86. 87.
88.
291
thopedic disease in pasture-fed New Zealand Thoroughbreds. Equine Vet J 1998;30:212–18. Hurtig MB, Pool RR. Pathogenesis of equine osteochondrosis. In: McIlwraith CW, Trotter GW (eds) Joint Disease in the Horse. Philadelphia: Saunders, 1996; pp. 335–58. Glade MJ, Belling TH. A dietary etiology for osteochondroitic cartilage. J Equine Vet Sci 1986;6:175–87. Kronfeld DS, Meacham TN, Donoghue S. Dietary aspects of developmental orthopedic disease in young horses. Vet Clin North Am Equine Pract 1990;6(2): 451–66. Harris P, Staniar WB, Ellis AD. Effect of exercise and diet on the incidence of DOD. In: Juliand V, Martin-Rosset W (eds) The Growing Horse: Nutrition and Prevention of Growth Disorders. EAAP publication no. 114. Wageningen: Wageningen Academic, 2005; pp. 275–90.
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Chapter 30
Important Gastrointestinal Parasites Eugene T. Lyons, Mariana Ionita and Sharon C. Tolliver
Introduction The horse is highlighted in this chapter but the parasites described are common in other equid types, such as ponies and donkeys. Also, similar species are found in zebras.1 Internal or endoparasites, as is implied, are located inside the horse. Albeit, some immature stages may be in the environment, in intermediate hosts, and on the exterior of the horse. Over 100 species are reported in equids.1 Approximately onehalf are in a group called small strongyles.2 The life cycle of most of the parasites is direct, but a few have intermediate hosts. A majority require part of the life cycle outside the host. The impact of parasites in horses ranges from no perceivable negative effects, to chronic problems, to actually causing death. It has been said that a socalled ‘good parasite’ does not kill its host. Parasites like bots (Gasterophilus spp.) seem well adapted to the horse and likewise the horse to them. Rarely are they the cause of death. Speculation by some people is that bots are an ‘older’ parasite compared to more pathogenic and possibly ‘younger’ ones, such as Strongylus vulgaris (a large strongyle) and ascarids (Parascaris equorum). In order for perpetuation of a parasite, a high enough level of reproduction is necessary to continue its existence. This is determined by several factors including genetic make-up of the parasites, immunology and age of the host, effectiveness of control measures such as drug treatment, environmental conditions for larval development, whether the horses are kept in a barn or dry lot versus pasture, and age groups and number of horses kept together. Viability of larval stages in the environment varies. For instance, strongyle larvae on pasture, even with ideal conditions, will die after a period of weeks
or months if not ingested by a horse. However, P. equorum has infective capability for a much longer period of time because the eggs contain larvae that are viable in the environment for years. The larvae in the ascarid eggs do not hatch while outside the host but only when ingested by it. Control of internal parasites has a two-fold objective: (i) to improve the health and well-being of the host and (ii) to lessen transmission to the host. The main difficulty in achieving these goals is resistance of some species of internal parasites, currently including ascarids and small strongyles, but potentially other species, to several of the marketed parasiticides. Also, the possibility exists for resistance of these parasites to other compounds.
Important gastrointestinal parasites Bots (Gasterophilus spp.) These are insects that spend most of their life as instars or maggots associated with the stomach.3 Adult flies, which have the appearance of honey bees (Figure 30.1), typically only live about 2 weeks. They lack, nor do they apparently need, functional mouthparts or digestive systems. The female may circle around a horse’s leg waiting to breed with the male fly. Even though they are incapable of biting or stinging, the presence of the fly and buzzing sound of its wings may alarm the horse. Female flies glue their eggs to hairs of horses typically in a species-specific pattern (Figure 30.1). Two main species, Gasterophilus intestinalis (common bot) and Gasterophilus nasalis (throat bot), and one uncommon species, Gasterophilus haemorrhoidalis (nose bot), are present in the USA. In other parts of the world, there also are other species. Eggs
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 30.1 Gasterophilus intestinalis (common bot) fly preparing to lay eggs on hairs of horse.
of the common bot are laid mostly on the hair of the inner knee. When these eggs are licked by the horse, hatching results with the first instar attaching to the tongue. At least one theory has been proposed as to why a horse would lick the knee area where bot eggs are attached. This is that some eggs may spontaneously hatch and the first instar might irritate the skin causing the horse to lick this area. First-instar throat bots hatch by themselves and crawl into the mouth. Both species, over about a 3-week period, develop from the first instar to the second instar in the mouth before being swallowed. In the mouth, the instars produce periodontal ulcerations mainly between the upper rear molars (Figure 30.2). These eroded areas heal after the bots have left, either after completing development to the second instar or after being killed by a boticide. Common bots also may burrow into the tongue. After being swallowed, the second instars, about 1 month later, have developed to the third instars. Specifically, G. intestinalis locate in the esophageal portion of the stomach and G. nasalis in the duodenum (Figure 30.3). Second and third instar bots anchor themselves by ‘mobile’ hooks to the wall of the esophageal portion of the stomach (G. intestinalis) or the duodenum
Figure 30.2 Upper molars of horse showing periodontal lesions containing first and second instars of Gasterophilus spp.
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Figure 30.3 Clusters of third instars (also a few second instars) of the common bot (Gasterophilus intestinalis) attached to the esophageal (white area) portion of stomach, and the throat bot (G. nasalis) attached to duodenal mucosa.
(G. nasalis) (Figure 30.4). Visually, the pits where the bots attach look like they would be quite damaging. However, except for rare perforation at the attachment site, other possible negative effects of bot infections, such as interfering with mastication, are difficult to assess. As with periodontal lesions, pits in the stomach or duodenum heal soon after the bots have left, either by natural passage or after a boticide is given. Without treatment for bots, the third instars detach, a few at a time, and pass down the intestinal tract and out in the feces of the horse in the spring and summer. They burrow into the ground, pupate, and adult flies emerge about 1 month later, then the cycle repeats. Even though bots inside a horse usually do not seem to cause significant problems, removal by drugs is recommended. Currently, only one class of parasiticides, the avermectins (e.g., ivermectin and moxidectin), is available as a boticide. These drugs are
Figure 30.4 Second instar Gasterophilus intestinalis anterior end portraying hooks used for attachment.
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highly effective on all bot stages including those individuals in the mouth. Stomach worms (Habronema spp., Draschia megastoma, and Trichostrongulus axei) Two types of nematodes (spirurids and trichostrongylids) infect the glandular portion of the stomach of horses.3 The spirurids include Habronema muscae, Habronema majus, and Draschia megastoma. A single species of trichostrongylid (Trichostrongylus axei) can be present. The spirurids may cause various problems. They require flies (e.g., Musca domestica and Stomoxys calcitrans) as intermediate hosts and for transmission of the larval stages. When the infected flies feed around the mouth or are swallowed by the horse, maturation of the spirurids occurs in the stomach. The flies may deposit larvae in external areas of the horse such as on the skin (called ‘summer sores’) in abrasions/wounds, causing irritation and potential development of granulomatous masses, on the outside of male genitalia (Figure 30.5), resulting in lesions and potential problems in urination, and in the eyes, causing conjunctivitis and possible granulomatous lesions. Sometimes, aberrant spirurid larvae migrate to the lungs, producing abscesses which may be associated with the bacterium Corynebacterium equi. Draschia megastoma produces nodules containing worms and necrotic and caseous debris, typically located along the margo plicatus in the glandular part of the stomach (Figure 30.6). The abscesses may have one or two openings. Usually, single nodules are < 50 mm in diameter; however, they may be larger
Figure 30.6 Draschia megastoma lesion (cut open) – stomach.
and there may be multiple nodules. Clinical signs usually are not evident unless, as occasionally occurs, abscesses break through the stomach wall directly or indirectly, involving organs like the spleen, resulting in peritonitis. Habronema muscae lives in the mucous exudate of the stomach and usually is considered benign. Trichostrongylus axei infections typically are light but, when heavy, there may be catarrhal inflammation and hypertrophic thickening of the stomach wall. This parasite is very unusual in that it is not hostspecific and it infects many species of animals including humans. Ivermectin and moxidectin are effective on the stomach stages of the spirurids and T. axei. Also, these compounds are active on larval spirurids on the exterior of the horse. Ascarids (Parascaris equorum) This roundworm is the largest nematode found in the horse, according to its combined diameter and length3, 4 (Figure 30.7). Typically, these parasites are
Figure 30.5 Cutaneous habronemiasis (‘summer sores’) causing lesions of tip of penis of a pony.
Figure 30.7 Mass of adult ascarids (Parascaris equorum) from the small intestine of a foal.
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Figure 30.8 Eggs of ascarid (round with dark outer shell) and strongyle (elongate with thin shell) found in horse feces.
Figure 30.9 Ascarid (Parascaris equorum) adults that perforated wall of foal small intestine at the mesenteric attachment.
the most prevalent and in highest numbers in adult forms in young animals, especially nursing foals and weanlings. By the yearling age, prevalence usually decreases. However, on occasion, older horses may have patent infections. The life cycle of P. equorum is as follows: Adults live in the small intestine and eggs are passed in the feces. The eggs, about 100 µm in diameter, have a heavy brown shell (Figure 30.8). In the environment, embryonation occurs but infective larvae do not hatch until ingested by the host. Due to the heavy protective eggshell and the feature of the embryo not hatching in the environment, eggs contain larvae that can remain infective for many years. Thus, once facilities like barns and pastures on horse farms are contaminated, it is very difficult to clear or clean out ascarid eggs from them completely. Infective larvae hatch from eggs ingested by the horse. The larvae then undergo hepatopulmonary migration. After development to the fourth stage (L4 ) in the lungs, they are swallowed and develop to the adult stage in the small intestine. The prepatent period (time from ingestion of larvae until egg production) is about 10–12 weeks. Females are highly prolific and, according to some sources, each individual potentially can produce 200,000 eggs per day. Various clinical problems are associated with ascarids. Migrating larvae may produce lesions in the liver. Allergic reactions may be present according to interpretations by some researchers. In experimental infections, coughing and copious mucus production in the throat have been attributed to migrating ascarid larvae. Intestinal infections produce conditions ranging from seemingly no problems to fatal situations. Motility of the small intestine may be reduced or increased, causing intussusception. A large mass of ascarids and hyper-gut-motility may cause perforation of the small intestine, usually at the mesenteric attachment. This results in peritonitis and death unless diagnosed in time for corrective surgery. Also,
if an effective ascaricide is given after maturation of ascarids, a blockage of dead worms may occur, resulting in impaction or rupture of the small intestine (Figure 30.9). As stated above, unless surgery is performed, death is predictable. Some clinical signs of ascarid infection in young horses are a ‘wormy-horse appearance’ consisting of a rough hair coat, pot belly, slow growth, and ‘poor-doing’. Drug treatment of foals is recommended to begin at about 6–8 weeks of age before ascarids mature. Formerly, several compounds were effective against ascarids, but unfortunately there are fewer effective drug classes and few drugs in the classes available now (Tables 30.1 and 30.2).5 Also, ivermectin and moxidectin are no longer effective according to reports from several countries, including the USA.6–10 This may not be too surprising because the injectible formulation of ivermectin did not always remove all of the ascarids. Currently, in small studies on pre- and posttreatment fecal examination in Thoroughbred foals in central Kentucky, effectiveness on ascarids was better for oxibendazole than fenbendazole; definite pyrantel pamoate resistance was evident on some farms.11 Presumably, piperazine is still effective but seldom is used. Table 30.1 Drugs on the market in the USA Class
Generic name
Benzimidazoles
Fenbendazole Oxibendazole Pyrantel pamoate tartrate (LLa ) Ivermectin Moxidectin Various Praziquantel
Pyrimidines
Macrocyclic lactones Piperazine Isoquinoline a
Low level.
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Table 30.2 Susceptibility of bots, ascarids, large and small strongyles, pinworms, and tapeworms to parasiticides Parasite group
Agent
Bots
Ivermectin Moxidectin BZsa Ivermectinb Moxidectinb Piperazine Pyrantel pamoateb BZs Ivermectin Moxidectin Pyrantel pamoate BZsb Ivermectinb Moxidectinb Pyrantel pamoateb BZs Ivermectin Moxidectinb Piperazine Pyrantel pamoate Praziquantel Combo (IVM)c Combo (MOX)d Pyrantel pamoate
Ascarids
Large strongyles
Small strongyles
Pinworms
Tapeworms
a
Benzimidazoles (oxibendazole and fenbendazole). Indications of lower than initial activity. c Praziquantel in combination with ivermectin. d Praziquantel in combination with moxidectin. b
Strongyles (large and small) This group of nematodes is separated into two groups, large strongyles and small strongyles.2 The distinction is not specifically by size but by certain morphologic characteristics. Taxonomically, until several years ago, large strongyles consisted of species in one genus, Strongylus spp. (S. vulgaris, S. edentatus, S. equinus, and S. assini). They have common-appearing buccal capsules and parenteral migration of larval stages. Now, four additional genera (Craterostomum, Oesophagodontus, Triodontophorus, and Bidentostomum) are classified as large strongyles because of buccal capsule similarities to Strongylus spp. However, their larval stages do not have parenteral migrations but encyst in the mucosa and submucosa of the large intestine.12 Small strongyles now generally are called cyathostomes or cyathostomins. The latter two terms are technically incorrect because of the additional four genera that recently have been added to the large strongyle group. Small strongyles are composed of about 50 species worldwide.2 In Kentucky, 33 species, including Craterostomum sp.,
Oesophagodontus sp., and Triodontophorus spp. have been recorded.12 Strongyles have a five-stage life cycle: Eggs (Figure 30.8), first-stage larvae (L1 ), second-stage larvae (L2 ), third-stage larvae (L3 ), fourth-stage larvae (L4 ), and fifth-stage larvae (L5 ) or adult stage.5, 13 Eggs are layed by females in the large intestinal tract and are voided in the feces of the horse. Outside the horse, an embryo in the egg develops to the L1 , which hatches. The L1 , now free from the egg, develops to the L2 and then to the L3 , which is enclosed in a sheath, the cuticle of the L2 . The L3 is the final stage outside the horse and is the infective stage, which the horse accidentally ingests. Inside the horse, the L3 develops to the L4 and then to the L5 . Strongylus spp. and small strongyles have different migratory routes once L3 larvae are ingested by the horse. Species in the genus Strongylus are the most pathogenic strongyles and can kill horses because of damage caused by migrating larvae in blood vessels and parenteral organs and tissues.3, 14 Strongylus vulgaris, with a prepatent period of about 6 months, is the most important parasite in horses. Its larvae cause verminous aneurysms, typically in the cranial mesenteric artery (Figure 30.10). The blood vessels balloon out and the walls thicken. Also, blood debris and larval specimens may occlude blood vessels. This is a thromboembolic process of the large arteries supplying blood to the intestines. Strongylus edentatus, with a prepatent period of about 11 months, is considered somewhat less pathogenic than S. vulgaris. Migrating S. edentatus especially can be damaging to the liver during the sojourn of the larvae there (Figure 30.11). Strongylus equinus, maturing at about 9 months, is less pathogenic than the two aforementioned Strongylus species. Mature Strongylus spp. attach to the mucosa of the cecum, ventral colon, and dorsal colon and suck blood (Figure 30.12).
Figure 30.10 Aneurysm of cranial mesenteric artery caused by migrating larvae of the large strongyle (Strongylus vulgaris).
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Figure 30.11 Liver showing migrating larvae of the large strongyle (Strongylus edentatus) and the damage they cause.
Small strongyles, except for a few species, are not as big as large strongyles. Some species produce patent infections at about 6–10 weeks. They are the most common horse parasite species and may be present in huge numbers (> 105 ). In most small strongyle species the mature parasites do not attach to the intestinal mucosa and suck blood. Typically they are plug feeders. Even though large numbers of encysted and luminal stages may be present in horses, usually little damage is evident (Figure 30.13). There may be large numbers of inhibited L3 in the lumen or encysted in the wall of the large intestine. While the effects of luminal stages are difficult to measure, larvae located in the mucosa/submucosa can be the cause of serious problems under certain circumstances. Under so-called ordinary conditions, these larvae undergo development to the fourth and sometimes fifth stage and excyst. Then they trickle into the intestinal lumen and mature. On occasion, huge numbers of larvae may emerge over a short period of time, severely damaging mucosal cells, resulting in loss of fluids and protein. This result is a condition called larval cyathostomiasis, among
Figure 30.12 Histological section of anterior end and buccal capsule of mature Strongylus vulgaris attached to the mucosa of the cecum and sucking blood.
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Figure 30.13 Encysted small strongyle larvae in mucosa of cecum.
various other names.5 The cause(s) is (are) poorly understood. Clinical larval cyathostomiasis is reported more commonly in Europe than in the USA. There, it is called winter cyathostomiasis because it typically appears in the winter and early spring. Possibly, seasonal changes of daylight length, ingestion of new vegetative growth, or other unknown factors may mobilize encysted larvae. At times, another situation seems to cause hyperactivity and release of abnormally high numbers of encysted larvae. This is after drug treatment that causes removal of luminal stages of small strongyles. Possibly, the luminal parasites have an inhibitory effect on encysted stages and, when this factor is lost, the encysted ones ‘awaken’. Larval cyathostomiasis is more prevalent in young horses (i.e., 1–3-year-olds). It even can occur in horses on regular parasite control programs. Some clinical signs of this disease are weight loss, diarrhea, pyrexia, subcutaneous edema (especially ventral), delayed shedding of hair, and colic. Diarrhea may be chronic or acute. Some diarrheas also have been associated with penetration of L3 into the mucosa. Affected animals may die at 2–3 weeks after appearance of clinical signs. Diagnosis of the disease in live animals may be difficult. Sometimes, elimination of, or at least ruling out, most other causes, is an aid. Also, lessening or cessation of signs after administration of an effective larvicidal compound helps in diagnosis. Sometimes, in feces, especially diarrheal, large numbers of specimens can be found. There may be quite obvious red worms, which are fourth-stage Cylicocyclus insigne. According to Lyons et al.,5 ‘Some specific pathologic changes caused by larval cyathostomiasis are catarrhal and fibrous inflammation of the large intestine with diffuse mucosal hemorrhagic foci, necrotic nodules in the mucosa, and mucosal disruption. On histologic examination, there is evidence of a
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pronounced cellular response characterized by the presence of mononuclear cells, eosinophils, and epitheloid macrophages. Other microscopic findings include fibrous encapsulation of larvae, mucosal and submucosal edema, widespread focal areas of submucosal hemorrhages with eosinophilic infiltrates, dilated submucosal lymphatics, and variable shedding’. Differentiation of eggs of Strongylus spp. and small strongyles passed in the feces is not possible by visual examination under a microscope (Figure 30.8). Culturing of eggs in feces allows identification of L3 Strongylus spp. from small strongyle L3 . Recent research has resulted in a molecular method of diagnosing S. vulgaris infections by detection of DNA of these parasites in feces of live horses.15 Frequent use of parasiticides for over 50 years has resulted in extensive resistance of small strongyles to most of the remaining few drugs commercially available.16 Benzimidazoles (BZs – fenbendazole and oxibendazole), pyrantel pamoate, and piperazine all have been shown to have low efficacy on these parasite species.17 Ivermectin and moxidectin have been highly efficacious, but counts of EPGs (strongyle eggs per gram of feces) currently are returning faster post treatment than they were initially.18–20 One of the ‘success’ features of persistent use of parasiticides is the virtual eradication of Strongylus spp. However, the comeback potential is always present because of possible lowered drug activity. With less and less activity of present parasiticides on small strongyles, an increase of larval cyathostomiasis is a possibility. Also, treatment of this condition may become a serious problem if there is lowered larvicidal activity of drugs like fenbendazole at the larvicidal dose rate (10 mg/kg once daily for 5 days) and moxidectin with past proven effectiveness on encysted small strongyles. Intestinal threadworms/strongyloids (Strongyloides westeri) Strongyloides westeri is a tiny nematode residing in the mucosa of the small intestine of horse foals.21 Thinwalled, embryonated eggs first appear in the feces of 2-week-old foals (Figure 30.14). These parasites are highly immunogenic. Infections decline and disappear at a few months of age. The life cycle in foals involves adult infections originating from two sources of larvae. First, and probably the most common origin, is transmission of infective larvae in milk of the mare to her foal. Larvae remain dormant in the subcutaneous tissue of the mare, especially in the ventral abdominal area, until parturition when they pass to the mammary system and then into the milk of the mare. Also, an environmental source is from L3
Figure 30.14 Egg (embryonated) of the small intestinal threadworm Strongyloides westeri.
developing from eggs passed in foal feces. These L3 may penetrate the skin or be ingested, resulting in enteric infections. Also, they can be the source of tissuelocated larval stages that pass in the milk of foaling mares. The S. westeri parasites have been incriminated as a causal agent of diarrhea in foals. Their association with this condition is usually quite difficult to differentiate from other causes such as bacterial or viral agents. Drudge and Lyons3 state that ‘Diarrheas caused by S. westeri do not have the fetid odor associated with Salmonella or other bacteria, and a febrile reaction does not occur with these nematodes. Also, feces are the normal greenish color typically found beginning a few days after birth. Fecal examinations revealing strongyloides eggs may be helpful; however, concurrent infections may be incidental. Many clinicians administer an anthelmintic (e.g. ivermectin) active against strongyloides early in the course of events. Prompt remission of the scouring indicates the problem of S. westeri in the episode’. The prevalence of S. westeri in foals in Kentucky was about 90% in the 1960s but, in current studies, is less than 10%.9, 22 Probably, highly effective drugs, such as thiabendazole and ivermectin, are the reason for the dramatic decline. It should be mentioned that some persons have believed that S. westeri was the cause of 9-day scours. There is no proof of this; in fact, all indications are that this perception is untrue. Pinworms (Oxyuris equi) This nematode species lives in the dorsal and small colon. These parasites typically live as immatures and adults in weanling and adolescent horses.3 Pinworms usually do not mature in older horses but fourth stages can be present, sometimes in large numbers. Female worms move to the anal region and lay eggs around the perineum; in the same area, ruptured females, which are quite fragile, also may be found (Figure 30.15). The females and eggs also are passed
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Figure 30.15 Pinworm (Oxyuris equi) adult females.
Figure 30.16 Egg of the tapeworm Anoplocephala perfoliata.
in horse feces. Pinworm eggs are quite sticky and are found adhered to a variety of objects in the environment. Embryonated eggs are ingested by the horse and development to the fourth stage occurs in about 3–10 days in the colon. There is a 5-month prepatent period. Pinworms mainly cause pruritus due to irritation of ruptured females and eggs around the anal region. This results in active tail-rubbing on objects such as fence posts and so forth. Hair on the base of the tail then becomes ruffled and even lost after multiple rubbings. Fourth-stage O. equi can cause damage while feeding on the mucosa of the large colon. All of the current parasiticides have label claims of activity on these parasites. One recent report indicates possible lowered activity of macrocyclic lactones.23
The most pathogenic species is A. perfoliata, possibly related to its location in the particularly vital part of the digestive system (the ileum), where it empties into the cecum. At the site of attachment at the ileocecal junction, there may be ulceration and a diphtheritic membrane. Pathologic changes may be perforation of ulcerated areas, blockage of the ileocecal opening, rupture of the cecum, and intussusception. Also, there may be hypertrophy of the terminal ileum. Presence of this tapeworm species seems to cause ‘great host reaction’ and hypermotility of the ileum. If colics, caused for instance by intussusception, are observed in time, surgery usually will correct the situation. Generally A. magna is not recognized as the cause of clinical problems, although catarrhal enteritis has been associated with these parasites. The first approved drug for removal of tapeworms in horses is the highly efficacious drug, praziquantel. It is combined with ivermectin and moxidectin in some products.24 Pyrantel pamoate at the 1X and 2X dose rates has activity on tapeworms, and now has label approval on these parasites.25–27
Tapeworms (Anoplocephala spp. and Paranoplocephala mamillana) Three species of tapeworms are found in horses in North America.3 Two of them (Anoplocephala perfoliata and A. magna) are reported more commonly than the other species (Paranoplocephala mamillana). The cecal tapeworm, A. perfoliata, has been the most prevalent species (> 50%) in surveys done in dead horses (mostly Thoroughbreds examined) in Kentucky. These studies were done before the marketing of praziquantel, which is highly efficacious on tapeworms. The eggs of tapeworms (Figure 30.16) are eaten by oribatid mites, which are intermediate hosts. The cyst stage (cysticercoid) develops in the mite, and when accidently eaten by horses these parasites have a prepatent period of about 2 months. They are found in all ages of horses. Location in the horse typically is the small intestine (SI) for A. magna (distal SI) and P. mamillana (anterior SI), and the cecum, especially the ileocecal junction, for A. perfoliata (Figure 30.17).
Figure 30.17 Anoplocephala perfoliata attached to mucosa of cecum.
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Some comments regarding parasite control There is no absolute right or wrong parasite control program. Situations vary on different farms. For instance, at present, generally there is more leeway with strongyle control than many years ago. This is because the most pathogenic parasites (Strongylus spp.) are currently rare on farms that have had good parasite control. Small strongyles can cause problems (larval cyathostomiasis) but clinical cases are more uncommon in the USA than in Europe. The larvicidal dose rate of fenbendazole (10 mg/kg once daily for 5 days) has been beneficial in treatment of larval cyathostomiasis. Whether this dose regimen is still effective on the encysted small strongyles is uncertain. As has been mentioned, ascarids are most common in young horses, especially foals. Also, it is evident that ivermectin and moxidectin are ineffective on these parasites. Therefore, for the control of ascarids, other compounds such as the BZs (oxibendazole and fenbendazole) and piperazine should be used. It is recommended that foals be treated every 6–8 weeks to remove ascarids before they mature and become the cause of more serious intestinal problems. Yearlings and older horses probably should be treated every 2–3 months, depending on the farm circumstances. Ideally, just treat those with strongyle counts of EPGs above a certain value, maybe 100 or more; however, no definite ‘cut-off’ value is absolute. It is recognized that a strongyle EPG/horse profile can be made to determine which horses may need treatment. By this, it is meant that certain horses, especially older ones, typically have consistently low EPGs. So, once this is established for specific horses, it seems unnecessary to treat them or to continue determining EPG counts. As has been discussed under the ‘Strongyle section’ of this chapter, the efficacy of all commercially available compounds on small strongyles has become less after a period of usage. Since nematode eggs typically will sink in water, a flotation solution with a specific gravity greater than 1.00 is necessary to float them. Various salt and sugar solutions are used. Sheather’s sugar solution (specific gravity 1.200–1.300) is favored by many laboratories/veterinary clinics. Veterinary clinical parasitology books and manuals are good references for use of flotation solutions and methods to detect worm eggs. These procedures for egg flotation may involve centrifugation or letting samples stand for a period of time. The BZs, piperazine, and pyrantel pamoate have very little efficacy on small strongyles now. Macrocyclic lactones (ivermectin and moxidectin) are becoming less effective; this is why they should be used
sparingly, maybe only twice a year in the spring and fall. Even though there is drug resistance of parasiticides on ascarids and small strongyles, they still seem effective on other parasites; thus, they have a place in treatment programs. The so-called ‘bottom line’ is that until newer compounds or other methods of control are found to be effective, horse owners are going to have to accept the fact that there is difficulty in reducing small strongyle burdens in their horses. Also, it should be understood that just because strongyle EPG counts may seem high, this does not necessarily mean that horses are clinically ill. Several management practices may be helpful in reducing parasite transmission in the environment. Some of them related to pastures are: rotation, clipping, chain harrowing, alternating horses and cattle, plowing and reseeding, and removal of manure. Composting manure, if done correctly, can be useful. As mentioned previously, the great dilemma in parasite control in horses and other domestic animals (e.g. ruminants) is drug resistance. Newer ideas for control of parasites need to be a high priority. These should include measures other than chemicals because, historically, parasites have developed resistance to parasiticides after a period of usage.
References 1. Krecek RC, Malan FS, Reinecke RK, de Vos V. Burchell’s zebras in South Africa. J Wildf Dis 1987;23:404–11. 2. Lichtenfels JR, Kharchenko VA, Krecek RC, Gibbons LM. An annotated checklist, by genus and species, of 93 species level names for 51 recognized species of small strongyles (Nematoda: Strongyloidea: Cyathostominea) of horses, asses and zebras of the world. Vet Parasitol 1998;79:69– 79. 3. Drudge JH, Lyons ET. Internal Parasites of Equids with Emphasis on Treatment and Control. Monograph. HoechstRoussel, 1989 (revised); 26 pp. 4. Clayton HM. Ascariasis in foals. Vet Clin N Am Equine 1986;2:313–28. 5. Lyons ET, Drudge JH, Tolliver SC. Larval cyathostomiasis. Vet Clin N Am Equine 2000;16:501–13. 6. Boersema JH, Eysker M, Nas JW. Apparent resistance of Parascaris equorum to macrocylic lactones. Vet Rec 2002;150:279–81. 7. Hearn FP, Peregrine AS. Identification of foals infected with Parascaris equorum apparently resistant to ivermectin. J Am Vet Med Assoc 2003;223:482–5. 8. Lyons ET, Tolliver SC. Prevalence of parasite eggs (Strongyloides westeri, Parascaris equorum, and strongyles) and oocysts (Eimeria leuckarti) in the feces of Thoroughbred foals on 14 farms in central Kentucky in 2003. Parasitol Res 2004;92:400–4. 9. Lyons ET, Tolliver SC, Collins SS. Field studies on endoparasites of Thoroughbred foals on seven farms in central Kentucky in 2004. Parasitol Res 2006;98:496–500. 10. Schougaard H, Nielsen MK. Apparent ivermectin resistance of Parascaris equorum in foals in Denmark. Vet Rec 2007;160:439–40.
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Important Gastrointestinal Parasites 11. Lyons ET, Tolliver SC, Ionita M, Collins, SS. Evaluation of parasiticidal activity of fenbendazole, ivermectin, oxibendazole, and pyrantel pamoate in horse foals with emphasis on ascarids (Parascaris equorum) in field studies on five farms in Central Kentucky in 2007. Parasitol Res 2008;103:287–91. 12. Tolliver SC. A Practical Method of Identification of the North American Cyathostomes (Small Strongyles) in Equids in Kenucky. University of Kentucky Agricultural Experiment Research Station Bulletin SR-2000-1, 2000; 37 pp. 13. Reinemeyer CR. Small strongyles. Recent advances. Vet Clin N Am Equine 1986;2:281–312. 14. Drudge JH, Lyons ET. Large strongyles. Recent advances. Vet Clin N Am Equine 1986;2:263–80. 15. Nielsen MK, Peterson DS, Monrad J, Thamsborg SM, Olsen SN, Kaplan RM. Detection and semi-quantification of Strongylus vulgaris DNA in equine faeces by real-time quantitative PCR. Int J Parasitol 2008;38:443–453. 16. Lyons ET, Tolliver SC, Drudge JH. Historical perspective of cyathostomes: prevalence, treatment and control programs. Vet Parasitol 1999;85:97–112. 17. Kaplan RM, Klei TR, Lyons ET, Lester G, Courtney CH, French DD, Tolliver SC, Vidyashankar AN, Zhao Y. Prevalence of anthelmintic resistant cyathostomes on horse farms. J Am Vet Med Assoc 2004;225:903–10. 18. Lyons ET, Tolliver SC, Ionita M, Lewellen A, Collins SS. Field studies indicating reduced activity of ivermectin on small strongyles in horses on a farm in Central Kentucky. Parasitol Res 2008;103:209–15. 19. Von Samson-Himmelstjerna G, Fritzen B, Demeler J, ¨ Schurmann S, Rohn K, Schnieder T, Epe C. Cases of reduced cyathostomin egg-reappearance period and failure of Parascaris equorum egg count reduction following ivermectin treatment as well as survey on pyrantel efficacy on German horse farms. Vet Parasitol 2007;144:74–8.
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20. Trawford AF, Burden F, Hodgkinson JE. Suspected moxidectin resistance in cyathostomes in two donkey herds at the Donkey Sanctuary, UK. In: Proceedings of the 20th International Conference of the World Association for the Advancement of Veterinary Parasitology, Christ Church, New Zealand, Oct 16–20, 2005. p. 196. 21. Lyons ET, Drudge JH, Tolliver SC. On the life cycle of Strongyloides westeri in the equine. J Parasitol 1973;59: 780–7. 22. Lyons ET, Tolliver SC, Drudge JH, Granstrom DE, Collins SS. Natural infections of Strongyloides westeri: prevalence in horse foals on several farms in central Kentucky in 1992. Vet Parasitol 1993;50:101–7. 23. Reinemeyer CR, Marchiondo AA, Shugart JI. Macrocyclic lactone-resistant Oxyuris equi: Anecdote or emerging problem? In: Proceedings of the American Association of Veterinary Parasitology, 51st Annual Meeting, 2006. p. 67. 24. Marley SE, Hutchens DE, Reinemeyer CR, Holste JE, Paul AJ, Rehbein S. Antiparasitic activity of an ivermectin and praziquantel combination paste in horses. Vet Ther 2004;5:105–9. 25. Lyons ET, Drudge JH, Tolliver SC, Swerczek TW, Collins SS. Determination of the efficacy of pyrantel pamoate at the therapeutic dose rate against the tapeworm Anoplocephala perfoliata in equids using a modification of the critical test method. Vet Parasitol 1989;31:13–18. 26. Marchiondo AA, White GW, Smith LL, Reinemeyer CR, Dascanio JJ, Johnson EG, Shugart JI. Clinical field efficacy and safety of pyrantel pamoate paste (19.13% w/w pyrantel base) against Anoplocephala spp. in naturally infected horses. Vet Parasitol 2006;137:94–102. 27. Reinemeyer CR, Hutchens DE, Eckblad WP, Marchiondo AA, Shugart JI. Dose-confirmation studies of the cestocidal activity of pyrantel pamoate paste in horses. Vet Parasitol 2006;138:234–9.
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Chapter 31
Vaccination of Mares, Foals and Weanlings W. David Wilson
Introduction Vaccination of horses is widely practiced and constitutes an important part of integrated infectious disease control programs based on maximizing specific and non-specific resistance to infection as well as management practices aimed at reducing the introduction and transmission of infectious agents. Fully licensed or conditionally licensed vaccines are now available in North America as aids to the prevention of tetanus, viral encephalomyelitis – Eastern equine encephalomyelitis (EEE), Western equine encephalomyelitis (WEE), Venezuelan equine encephalomyelitis (VEE) – West Nile virus (WNV) infection, influenza, equine herpesvirus type 1 (EHV1) and EHV-4 infection, strangles, rabies, equine viral arteritis (EVA), Potomac horse fever (PHF), type B botulism and, rotavirus infection. Core diseases against which all horses in North America should be vaccinated include:
r r r r
tetanus Eastern and Western equine encephalomyelitis West Nile virus (WNV) infection rabies.
The abortifacient potential of EHV-1 warrants inclusion of this disease in the core for all pregnant broodmares. The remaining diseases for which vaccines are available are considered ‘non-core’ or risk-based diseases.
Considerations for use of vaccines in broodmares The primary goals of vaccination programs for broodmares are: (i) prevention of diseases that pose
a risk to the mare or her fetus, and (ii) maximizing the level of colostral antibodies that will be passively absorbed by the neonatal foal after nursing, thus providing it with protection against diseases that pose a risk during the first few months of life. Additional considerations in selecting vaccines for use in pregnant mares include: (iii) safety to the mare and fetus, (iv) the potential for interference between multiple vaccines administered simultaneously, and (v) the influence of pregnancy on vaccine responses. Protecting the mare against diseases that pose a risk to the mare or her fetus Broodmares are at risk of exposure to the same diseases as performance and pleasure horses; therefore, they should be regularly vaccinated against all core and specific risk-based diseases according to published recommendations (Tables 31.1 and 31.2). The high horse traffic and high concentration of foals and young horses that typify many breeding farms contribute to a high risk of exposure to contagious respiratory diseases, including EHV-4, EHV-1, influenza, and strangles. Inclusion of influenza in vaccination protocols for broodmares is, therefore, routinely recommended, and addition of EHV-4 and strangles vaccines is frequently recommended when conditions of significant risk are anticipated. Vaccination of mares against EVA before breeding may be indicated when they are to be bred to a known or suspected EVA carrier stallion, either by natural cover or by artificial insemination. Whereas protection of the broodmare and fetus, or herd mates, against the abortifacient effects of EHV-1 or EVA is the primary goal underlying inclusion of these antigens in vaccination protocols for broodmares, the goal of protecting the foal features at least as
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
Inactivated vaccine: Three-dose primary series: First dose at 3–4 months of age Second dose 4–6 weeks after the first dose Third dose at 10–12 months of age, prior to the onset of the next vector season Recombinant canarypox-vectored vaccine: Three-dose primary series: First dose at 3–4 months of age Second dose 4–6 weeks after the first dose Third dose at 10–12 months of age, prior to the onset of the next vector season Flavivirus chimera vaccine: Two-dose primary series: First dose at 5–6 months of age Second dose at 10–12 months of age prior to the onset of the next vector season. Foals in the southeastern USA: The primary vaccination series should be initiated at 3 months of age due to early seasonal vector presence
Foals in the southeastern USA: The primary vaccination series should be initiated at 3 months of age due to early seasonal vector presence
Annual in spring, prior to onset of vector season
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There are no published data regarding use of the flavivirus chimera product in foals < 5 months of age. If administered to foals < 5 months of age, the recommended schedule for primary vaccination should be completed by administration of a dose of vaccine at 5 months of age or older
Month of birth and geographical location influence the risk of exposure to insect vectors at specific foal ages; therefore, scheduling of the primary immunization series may be amended by administration of WNV vaccines to foals at an earlier age if vectors are present
Month of birth and geographical location influence the risk of exposure to insect vectors at specific foal ages; therefore, scheduling of the primary immunization series may be amended by administration of EEE/WEE vaccines to foals at an earlier age if vectors are present
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West Nile virus (WNV)
Annual in spring, prior to onset of vector season
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Three-dose primary series: First dose at 3–4 months of age Second dose 4–6 weeks after the first dose Third dose 3–5 months after the second dose, prior to onset of next vector season. Foals in the southeastern USA: The primary vaccination series should be initiated at 3 months of age or earlier due to early seasonal vector presence
Annual
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Eastern and Western equine encephalomyelitis (EEE, WEE)
Three-dose primary series: First dose at 1–4 months of age Second dose 4–6 weeks after the first dose Third dose 3–5 months after the second dose
Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after the first dose Third dose at 3–5 months after the second dose (i.e., 10–12 months of age) Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after the first dose Third dose at 10–12 months of age, prior to onset of next vector season. Foals in the southeastern USA: The primary vaccination series should be initiated with an additional dose at 3 months of age due to early seasonal vector presence Inactivated vaccine: Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after the first dose Third dose at 10–12 months of age, prior to the onset of the next vector season Recombinant canarypox-vectored vaccine: Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after the first dose Third dose at 10–12 months of age, prior to the onset of the next vector season Flavivirus chimera vaccine: Two-dose primary series: First dose at 5–6 months of age Second dose at 10–12 months of age, prior to the onset of the next vector season
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Core diseasesa Tetanus (toxoid)
Foals and weanlings (< 12 months of age) of mares not vaccinated in the prepartum period
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Table 31.1 Guidelines for vaccination of foals, weanlings, and yearlings against corea and non-coreb diseases
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Equine influenza
304 Canarypox-vectored recombinant vaccine: Three-dose primary series: First dose at 5 months of age Second dose 5 weeks after the first dose Third dose at 10–12 months of age Revaccinate at 12-month intervals
Canarypox-vectored recombinant vaccine: Three-dose primary series: First dose at 5 months of age Second dose 5 weeks after the first dose Third dose at 10–12 months of age Revaccinate at 12-month intervals
(Continued)
The modified live intranasal vaccine is licensed for administration to horses 11 months of age or older with a label recommendation of one dose for primary immunization. If the vaccine is given before 11 months of age, a second dose should be administered at 11 months or older Semi-annual
Annual
An increased risk of disease may warrant vaccination of younger foals. Because potentially interfering maternal anti-influenza antibody is likely to be present, a complete primary vaccination series should be given after 6 months of age
Semi-annual
Semi-annual (6-month interval)
Annual
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Equine herpesvirus (EHV)
Three-dose primary series: First dose as early as 1–3 months of age Second dose 4 weeks after the first dose Third dose 4 weeks after the second dose Inactivated EHV-1, EHV-1/4, or modified live EHV-1 vaccine: Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after first dose Third dose 3–4 months after the second dose Revaccinate at 6-month intervals Inactivated vaccine: 3-dose primary series: First dose at 6 months of age Second dose 3–4 weeks after the first dose Third dose at 10–12 months of age Revaccinate at 6-month intervals Modified live intranasal vaccine: 2-dose primary series administered intranasally: First dose at 6–7 months of age Second dose at 11–12 months of age Revaccinate at 6-month intervals
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Three-dose primary series: First dose as early as 2–3 months of age Second dose 4 weeks after the first dose Third dose 4 weeks after the second dose Inactivated EHV-1, EHV-1/4, or modified live EHV-1 vaccine: Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after the first dose Third dose 3–4 months after the second dose Revaccinate at 6-month intervals Inactivated vaccine: Three-dose primary series: First dose at 6 months of age Second dose 3–4 weeks after the first dose Third dose at 10–12 months of age Revaccinate at 6-month intervals Modified live intranasal vaccine: Two-dose primary series administered intranasally: First dose at 6–7 months of age Second dose at 11–12 months of age Revaccinate at 6-month intervals
Anthrax vaccination is rarely indicated – only in focal endemic areas Antimicrobial drugs must not be given concurrently with this vaccine Exercise caution during storage, handling, and administration of this live bacterial product Consult a physician immediately should accidental human exposure (via mucous membranes, conjunctiva, or broken skin) occur Limited information suggests that maternal antibody does not interfere with vaccination; therefore, foals at high risk may be vaccinated as early as 2 weeks of age
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Botulism (type B toxoid)
Annual, spring
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No age-specific guidelines are available for this vaccine Manufacturer’s recommendation is for primary series of two doses administered subcutaneously (in the neck) at a 2–3 week interval
One- or two-dose primary series: First dose at 3–4 months of age Second dose 4–6 weeks after the first dose (optional if dam is seronegative) Next dose at 10–12 months of age
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Two-dose primary series: First dose at 6 months of age Second dose 4–6 weeks after the first dose Next dose at 10–12 months of age
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Rabies
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Three-dose primary series: First dose at 5–6 months of age Second dose 3–4 weeks after the first dose Third dose at 10–12 months of age Not recommended in foals Inactivated M-protein subunit vaccines: Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after the first dose Third dose 4–6 weeks after the second dose Modified live intranasal vaccine: Three-dose primary series administered intranasally: First dose at 6–9 months of age Second dose 3–4 weeks after the first dose Third dose at 11–12 months of age
Equine viral arteritis (EVA)
Potomac horse fever (PHF)
Semi-annual to annual
Three-dose primary series: First dose at 5–6 months of age Second dose 3–4 weeks after the first dose Third dose at 10–12 months of age Not recommended in foals Inactivated M-protein subunit vaccines: Three-dose primary series: First dose at 4–6 months of age Second dose 4–6 weeks after the first dose Third dose 4–6 weeks after the second dose Modified live intranasal vaccine: Three-dose primary series administered intranasally: First dose at 6–9 months of age Second dose 3–4 weeks after the first dose Third dose at 11–12 months of age
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Modified, with permission, from recommendations developed by the American Association of Equine Practitioners (AAEP) Infectious Disease Committee and posted on the AAEP website (www.aaep.org) in January 2008. ©AAEP 2008. a Core vaccines protect against diseases that are endemic to a region, are virulent or highly contagious, pose a risk of severe or fatal disease, have potential public health significance, and/or are required by law. Core vaccines have clearly demonstrable efficacy, and have a sufficiently high level of patient benefit and low level of risk to justify their use in all equids in North America. b Non-core (risk-based) vaccines are selected for use based on assessment of risk performed by, or in consultation with, a licensed veterinarian. Use of non-core vaccines will vary between individuals, populations, and/or geographical regions.
If warranted by risk, the modified live vaccine (MLV) may be safely administered to foals as young as 6 weeks of age. However, vaccine efficacy in this age group has not been determined. If the MLV product is administered to foals < 6 months of age, a third dose of vaccine should then be administered 2–4 weeks prior to weaning
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Vaccination is not routinely recommended as a strategy in outbreak mitigation; however, vaccination may be warranted on farms with endemic strangles
Prior to initial vaccination, colt (male) foals should undergo serological testing and be confirmed negative for antibodies to EAV. Testing should be performed shortly prior to, or preferably at, the time of vaccination. Maternally derived anti-EAV colostral antibodies can persist in the foal for up to 6 months; therefore, testing and vaccination should not be performed prior to 6 months of age If risk warrants, vaccine may be administered to younger foals, in which case subsequent doses should be administered at 4-week intervals until 6 months of age
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NA Semi-annual
Annual for colts intended for use as breeding stallions
Colt (male) foals: Single dose at 6–12 months of age (see comments)
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Rotavirus Strangles
Colt (male) foals: Single dose at 6–12 months of age (see comments)
Disease/vaccine
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Table 31.1 (Continued)
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Foal: Specific Diagnostic and Management Techniques
prominently in the rationale for vaccinating mares against tetanus, WNV, EEE, WEE, rabies, influenza, EHV-4, and strangles. The inclusion of rotavirus and botulism vaccines in protocols for pregnant mares is directed almost exclusively at protecting the young foal against these diseases. Maximizing maternally derived antibody (MDA) transfer Maintaining consistent broodmare vaccination protocols, which typically include administration of booster doses of vaccines during the last 2 months of gestation, will not only protect the mare but also maximize the likelihood that a uniformly high level of maternally derived antibody (MDA) transfer and passive protection will be achieved within the foal crop. This is particularly important for diseases that pose a risk to the foal during the first few weeks of life. Whereas intranasally administered vaccines may afford good protection to the mare, they are typically less effective than parenterally administered inactivated vaccines in stimulating high levels of circulating IgG, the isotype that is passively transferred to the foal in highest concentration. Parenterally administered vaccines are, therefore, preferred over intranasally administered vaccines for vaccination of mares during late gestation. Vaccine safety in broodmares Consideration of vaccine safety in broodmares must take into account the risks to maintenance of pregnancy and safety to the fetus. Potential adverse effects of vaccines on pregnancy are difficult to document, even when large numbers of mares are used, unless obvious problems occur. Because fetal organogenesis occurs early in gestation and this period is also characterized by substantial embryonic loss, even in normal mares, it is sound practice to avoid administering vaccines to mares during the first 60 days of gestation unless conditions of imminent risk prevail. Few vaccines carry specific label recommendations for use in pregnant mares and few published data exist specifically to document the safety of equine vaccines during pregnancy. Licensed vaccines that carry label recommendations for use in pregnant mares include two inactivated EHV-1 vacR +1b, Pfizer Animal Health; cines (Pneumabort-K R and Prodigy , Intervet/Schering-Plough) marketed for use in pregnant mares as an aid to prevention of R line of inactivated EHV-1 abortion; the Calvenza TM R influenza (Calvenza -03 EIV), EHV-1 (Calvenza EHV), and influenza/EHV-1 combination (CalvenzaTM -03 EIV/EHV) vaccines from Boehringer Ingelheim; two inactivated Potomac horse fever vacR , Merial; and EquovumTM cines (Equine Potomavac
PHF, Boehringer Ingelheim); and one vaccine licensed for prevention of type B botulism in foals R B, Neogen Corp.). In addition, one con(BotVax ditionally licensed vaccine (Equine Rotavirus Vaccine, Pfizer Animal Health) for prevention of rotavirus infection in foals is labeled for use in pregnant mares. While not specifically labeled for administration during pregnancy, widespread use in practice over many years has failed to document that any of the inactivated vaccines currently marketed for use in horses pose an unacceptable risk to pregnant mares. Therefore, pregnant mares are routinely vaccinated with inactivated vaccines directed against tetanus, EEE, WEE, WNV, influenza, EHV-4, strangles, and, to a lesser extent, PHF, rabies, and VEE. Similarly, adverse impacts on pregnancy have not been documented for modified live intranasally administered strangles (PinnacleTM I.N., Pfizer Animal Health) and R , Intervet/Schering-Plough) influenza (Flu Avert vaccines or the modified live parenterally adminisR tered EHV-1 vaccine (Rhinomune , Pfizer) In addition, safety of the recombinant WNV and influenza R , Merial) should not be a sigvaccines (Recombitek nificant concern because the modified live canarypox vector lacks the ability to infect mammalian cells. In addition, the equivalent canarypox-vectored influenza vaccine (ProteqFlu, Merial) marketed in the UK is labeled for use during pregnancy. Although the flavivirus chimera WNV vaccine (PreveNileTM , Intervet/Schering-Plough) is not specifically labeled for use during pregnancy, more than 300 pregnant mares were vaccinated during safety trials for licensing, without apparent adverse effects on the conceptus. In contrast, modified live virus VEE vaccines and live anthrax spore vaccines should not be used in pregnant mares. The modified live EVA vaccine should be used in pregnant mares only when conditions of imminent risk prevail. Protection of mares against the potential abortifacient effects of EVA infection is, therefore, best accomplished by completing the primary immunization series before the mare enters the broodmare band and by administering subsequent boosters during the open period before rebreeding.1 The practice of booster vaccinating mares against multiple diseases to maximize colostral transfer of antibodies to the foal, and the fact that mares in broodmare bands are generally middle aged or older, results in the typical broodmare receiving multiple doses of many vaccine antigens and adjuvants during her lifetime. In addition to stimulating high levels of antibody against a range of antigens, this practice may also predispose these mares to a higher rate of local and systemic adverse reactions, an issue that not only warrants further investigation but also may force horse owners and veterinarians carefully to consider strategies for revaccination.
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Vaccination of Mares, Foals and Weanlings Potential interference between multiple antigens administered concurrently The possibility that ‘competition’ between multiple antigens will compromise the response to some or all of the administered antigens should be considered. When administration of multiple vaccines late in gestation is indicated, it is good practice to administer no more than four antigens at one time and to allow an interval of 2 to 4 weeks between administration of different vaccines. Influence of pregnancy on vaccine responses It is widely assumed that pregnant mares are fully capable of mounting appropriate cellular and humoral immune responses to vaccines; however, this issue has received little research attention. While mares that have been primed before breeding appear to mount appropriate anamnestic responses to vaccines, we have generated preliminary data suggesting that the humoral response to primary vaccination with two inactivated vaccines (WNV and rabies) may be downregulated during gestation, resulting in failure of vaccinated mares passively to transfer specific antibodies to the foal via colostrum. We are currently researching this issue further because it has obvious ramifications for the immunoprotection of both the foal and the mare.
Considerations for use of vaccines in foals and weanlings Studies in foals to document the potential inhibitory effects of maternal antibodies are not required for licensing; therefore, it is not surprising that few vaccines carry label directions for primary immunization of foals. Of those that do, the range of recommended minimum ages is wide and includes the following: 3 months for several inactivated Potomac horse fever and rabies vaccines; 5 months for the canarypox recombinant inR Equine Influenza and WNV vaccines (Recombitek R fluenza and Recombitek WNV, Merial); 5 months for the flavirus chimera WNV vaccine (PreveNileTM , Intervet/Schering-Plough); 6 months for inactivated influenza and EHV-1 vaccines from one company (CalvenzaTM -03 EIV, EHV, and EIV/EHV, Boehringer Ingelheim); 9 months for a modified live inR I.N., Pfizer tranasal strangles vaccine (Pinnacle Animal Health); and 11 months for a modified live intranasal influenza vaccine (Flu AvertTM I.N., Intervet/Schering-Plough). Few studies have been done to determine the age at which various elements of the innate and adaptive immune system become fully competent and optimal responses to vaccines can be achieved. Maternally derived antibodies
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(MDA) and perhaps other immune effectors that are concentrated in colostrum and are passively transferred to the foal, play a crucial role in defense against pathogens encountered during the first few months of life while endogenous immune function continues to mature. Passive transfer of MDA should therefore be exploited in immunization programs for foals by consistently administering booster doses of selected vaccines to mares 4 to 8 weeks before foaling and by ensuring that foals ingest adequate amounts of highquality colostrum within 24 hours of birth. Although several immunoglobulin isotypes are present in colostrum, the sub-isotypes of IgG are absorbed into the systemic circulation of the foal in much higher concentrations than either IgM or IgA.2 Adequate passive transfer of MDA leads to titers of specific antibody in the serum of the foal that are typically very similar to the serum titer in the dam at the time of foaling. Thereafter, in the foal, maternal IgG antibodies decline with a half-life of 25 to 40 days and remain detectable for 6 months or longer in some cases.3–9 In addition to passively protecting the foal, MDAs may also exert a profound inhibitory effect on the active immune response of the foal to antigens, including those contained in vaccines. This phenomenon, known as maternal antibody interference, was brought into focus during the 1990s by several reports documenting that foals less than 6 months of age consistently failed to mount serological responses to inactivated influenza vaccines.5, 6,9–13 In addition, MDA appeared to cause misdirection of the immune response to influenza vaccines away from the more important virus-neutralizing IgGa and IgGb sub-isotypes in favor of the less effective IgG(T) sub-isotype of IgG.9 Subsequent studies in which titers of total, rather than antigen-specific, IgG sub-isotypes were determined, documented that the age-related increase in concentrations of IgGb lagged significantly behind increases in concentrations of other isotypes and remained below adult levels beyond 6 months of age.14 Maternal antibody interference has now been documented to be a significant issue for many other antigens, including tetanus, EEE, WEE, EHV-1, and EHV4, contained in vaccines administered to foals.3, 9,15–18 Even low levels of antibody, below those detectable by many routine serological tests and below those thought to be protective, can completely block the serological response to some vaccines, resulting in a potentially prolonged period of susceptibility before the foal is capable of responding appropriately to vaccines.17 These findings also indicate that it is not typically feasible to test samples from foals serologically to predict whether they will respond to particular vaccines. It is now recommended that primary vaccination with most inactivated vaccines should be delayed
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until foals are 6 months of age or older and that, with the exception of rabies, the primary immunization series for foals should comprise three doses rather than the two doses typically recommended by vaccine manufacturers. Whereas the recommended interval between the first and second doses is typically 3 to 4 weeks, a longer interval of 3 to 5 months between the second and third doses appears to be optimal. The third dose typically stimulates a serological response of greater magnitude and durability than two doses and may also contribute to a higher ‘set-point’ for the response to subsequent booster doses.9, 17, 19, 20 In contrast to the results cited above, maternal antibodies do not appear to exert a marked inhibitory effect on the immune response of foals to the inactivated (West Nile-InnovatorTM , Pfizer Animal Health), live R , Merial), live chimera recombinant (Recombitek (PreveNileTM , Intervet/Schering-Plough), or DNA R DNA, Pfizer Animal Health) (West Nile-Innovator WNV vaccines, thereby permitting antibody-positive foals as young as 3 months of age to be immunized successfully.20–22 Study results should be interpreted with caution because only humoral responses are typically assessed and infectious challenge is not typically performed to confirm protection, or lack thereof. Lack of a serological response may correlate well with lack of protection for some diseases and some vaccines, whereas for others this may not be the case. In contrast, the presence of a serological response may not correlate well with protection, as is frequently the case for respiratory tract pathogens. Because many commercially available vaccines are inactivated, adjuvanted, and administered by intramuscular injection, they have limited potential to stimulate cellular and mucosal responses; therefore, serological responses induced by these vaccines likely correlate well with their potential to induce protection. In turn, MDA interference with serological responses to inactivated vaccines likely equates to failure to induce protection. In contrast, failure to detect a serological response to a modified live, vectored DNA vaccine, or to a mucosally administered vaccine may not equate to lack of protection because vaccines of these types induce a broader array of systemic and local responses that may not be affected by MDAs. If MDA interference were not an issue for any vaccine, the approach to vaccination of foals would be greatly simplified because primary vaccination against all-important diseases could be completed before MDAs had declined to non-protective levels. In effect, the ‘window of susceptibility’ would be eliminated. In reality, an attainable goal is to maximize the beneficial effects of MDA while minimizing their negative impact on primary immunization. In order to best meet this goal it is necessary to decide which one (or both) of the following is the primary focus:
1. to protect the foal and weanling against specific high-risk infectious diseases that affect this age group and have the potential to cause significant disease, either directly or by predisposing to other secondary infections; or 2. to initiate primary immunization to protect against disease later in life. Assessing risk takes into account both the likelihood that the foal will become infected, and the risk of serious sequelae or death if the animal does become infected and develop disease. If the disease affects the foal early in life, such as is the case with rotavirus (RV) infection, there is usually insufficient time to induce a protective immune response by actively immunizing the foal. Under these circumstances, the approach should be to maximize the degree of protection passively transferred from the dam via colostrum. Other diseases, such as rabies, affect horses of all ages, but the risk of acquiring infection is generally low. Diseases of moderate to high risk to young foals but low risk to adults These include rotavirus infection (on certain breeding farms in certain years) and, in geographical areas such as Kentucky and some other eastern states, type B botulism. For these diseases, the following approach is appropriate:
r r r r
Booster-vaccinate the dam 4 to 8 weeks before foaling to maximize uniformity of passive transfer; Ensure good passive transfer of maternal antibodies; Introduce management practices to reduce exposure to the infectious agent; Vaccinate the foal if risk continues beyond the first few months of life.
Diseases of moderate to high risk for weanlings and older horses but lower risk to young foals born to vaccinated mares These include EHV-4, EHV-1, strangles, influenza, tetanus, EEE, and WNV infection. For these diseases, the following approach is appropriate:
r r r
Vaccinate the dam 4 to 8 weeks before foaling to maximize uniformity of passive transfer. Ensure good passive transfer of maternal antibodies. Start foal vaccination after the risk of maternal antibody interference is no longer present in most foals. When several vaccine types are available for a particular disease, the vaccine that is least subject to MDA interference should be used. Introduce
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management practices to reduce exposure to the infectious agent while primary vaccination is being completed. If a two-dose primary series is recommended for adult horses, use three or more doses of vaccine in the primary series to improve the chances that foals who do not respond to earlier doses will respond to additional doses administered later.
Diseases of low risk to foals In most circumstances these include rabies, PHF, WEE, and EVA. For these diseases, the following approach is appropriate:
r
r r
Vaccinate the dam before foaling if the disease is a significant risk to adult horses and a vaccine shown to be safe for use in pregnant mares is available. If the available vaccines are not considered safe for use in pregnant mares, administer boosters before breeding. Ensure good passive transfer of maternal antibodies. Start foal vaccination after the risk of maternal antibody interference is no longer present in any foal (typically 9 months to 1 year of age; except for rabies, when primary immunization can begin as early as 6 months of age).
Considerations for specific diseases (Tables 31.1 and 31.2) Tetanus All horses are at risk for developing tetanus, an often fatal disease caused by a potent neurotoxin elaborated by the anaerobic, spore-forming bacterium, Clostridium tetani. Protection against tetanus is mediated by circulating antibodies; toxin binding inhibition (ToBi) antibody titers exceeding 0.2 IU/mL are considered to be protective in the horse.19, 23 The many available vaccines are formalin-inactivated, adjuvanted toxoids that are inexpensive, safe, and potent antigens capable of inducing an excellent serological response and solid, long-lasting immunity when administered according to manufacturer’s recommendations. Annual revaccination of pregnant mares should be completed 4 to 8 weeks before foaling to enhance concentrations of specific immunoglobulins in colostrum and protect the mare if she sustains foaling-induced trauma or retained placenta. Colostrum-derived antibodies protect the foal for several months postnursing but also significantly interfere with the immune response of foals vaccinated with tetanus toxoid until they are approximately 6 months of age.9, 24
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Primary vaccination of foals that have received appropriate transfer of colostral antibodies from a vaccinated mare should include three doses of tetanus toxoid, beginning at age 6 months or older. The interval between the first two doses of vaccine should be approximately 4 weeks, and the interval between the second and third doses should be 3 to 5 months. The three-dose primary series is recommended for foals because a high proportion of foals fail to seroconvert in response to two doses of tetanus toxoid, regardless of whether or not maternal antibodies are detectable at administration of the first dose.9, 24 For foals born to non-immune mares this initial three-dose series can start at 1 to 4 months of age. Annual revaccination is recommended thereafter. Tetanus antitoxin (TAT) is produced by hyperimmunization of donor horses with tetanus toxoid. Although routine use of TAT in foals is not recommended, many practitioners continue to administer it to newborn foals born to unvaccinated mares. One vial of antitoxin (1500 IU) induces immediate passive protection that lasts up to 3 weeks.24 More prolonged protection may be accomplished with higher doses; however, the preferred approach is active immunization with toxoid. Equine encephalomyelitis (sleeping sickness) The equine encephalomyelitis viruses (EEE, WEE, and VEE) belong to the Alphavirus genus of the family Togaviridae. They are transmitted by mosquitoes, and infrequently by other bloodsucking insects, to horses from wild birds or rodents, which serve as natural reservoirs for these viruses. Correlates for protection against EEE, WEE, and VEE are not well established; however, circulating antibodies are assumed to be important because infection is acquired by vascular injection (mosquito bites) and current inactivated vaccines appear to have good efficacy.25, 26 Available vaccines are formalin-inactivated, adjuvanted, bivalent, whole-virus products containing EEE and WEE R R (Encevac with Havlogen , Intervet/Schering R InnovatorTM , Pfizer AniPlough; Encephaloid R mal Health; Cephalovac EW, Boehringer Ingelheim), or trivalent products that also contain VEE R VEW, Boehringer Ingelheim). A study (Cephalovac evaluating the serological response of horses to commercial encephalomyelitis-tetanus combination vaccines showed that the EEE neutralizing antibody responses to Encevac TTM (Intervet/ScheringPlough, carbopol adjuvant) and EquiloidTM (Pfizer Animal Health, squalene and surfactant adjuvant) were of greater magnitude and persistence than responses to Cephalovac EWTTM (Boehringer Ingelheim, saponin adjuvant).27 However, no comparative randomized challenge studies have been performed
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Table 31.2 Guidelines for vaccination of broodmares against core and non-core diseases Previously vaccinated broodmares
Broodmares not previously vaccinated or lacking a vaccination history
Annual, 4–6 weeks prepartum
Two-dose series: Second dose 4–6 weeks after the first dose Revaccinate 4–6 weeks prepartum, depending on timing of second dose
Booster at time of penetrating injury or surgery if last dose was administered more than 6 months previously
Eastern and Western equine encephalomyelitis (EEE, WEE)
Annual, 4–6 weeks prepartum
Two-dose series: Second dose 4 weeks after the first dose Revaccinate 4–6 weeks prepartum, depending on timing of second dose
Consider 6-month revaccination interval for mares residing in endemic areas with a prolonged vector season
West Nile virus (WNV)
Annual, 4–6 weeks prepartum
It is preferable to vaccinate naive mares when open. When risk is high, initiate primary series as follows: Inactivated or recombinant canarypox-vectored vaccine: Two-dose series: Second dose: 4–6 weeks after the first dose Revaccinate 4–6 weeks prepartum, depending on timing of second dose Flavivirus chimera vaccine: One dose Revaccinate 4–6 weeks prepartum, depending on timing of first dose
When using the inactivated or the recombinant product, consider a 6-month revaccination interval for mares residing in endemic areas with a prolonged vector season For naive mares being imported into an endemic area during the vector season, the preferred approach is to complete the primary vaccination series prior to importation. If this approach is not feasible, protect them from being bitten by mosquitoes if possible, and vaccinate them with one of the vaccines (flavivirus chimera or canarypox-vectored) that have the most rapid onset of immunity
Rabies
Annual, prior to breeding or 4–6 weeks prepartum
One dose; annual revaccination prior to breeding or 4–6 weeks prepartum
Because booster vaccination induces persistently elevated levels of antirabies antibody this vaccine may be given post-foaling, but prior to breeding, in order to reduce the number of vaccines given to mares prepartum
Equine herpesvirus (EHV)
Three-dose series with product labeled for prevention against EHV-1 abortion Administer during the 5th, 7th, and 9th months of gestation
Three-dose series with product labeled for prevention of EHV abortion Administer during the 5th, 7th, and 9th months of gestation
Disease/vaccine Core diseasesa Tetanus (toxoid)
Non-core (risk-based) vaccinesb Anthrax Not recommended for use during gestation Botulism Annual, 4–6 weeks prepartum
Comments
Not recommended for use during gestation Three-dose series: First dose during 8th month of gestation Second dose 4 weeks after the first dose Third dose 4 weeks after the second dose (Continued).
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Table 31.2 (Continued) Previously vaccinated broodmares
Broodmares not previously vaccinated or lacking a vaccination history
Inactivated vaccines: Semi-annual with one dose administered 4–6 weeks prepartum
Inactivated vaccine: Three-dose series: Second dose 4–6 weeks after the first dose Third dose 4–6 weeks prepartum
Canarypox-vectored vaccine: Semi-annual with one dose administered 4–6 weeks prepartum
Canarypox-vectored vaccine: Two-dose series: Second dose 4–6 weeks after first dose but no later than 4 weeks prepartum
Equine viral arteritis (EVA)
Not recommended unless risk of exposure is high
Not recommended unless risk of exposure is high
Mares potentially intended for export should undergo serological testing immediately prior to initial vaccination and be confirmed negative for antibodies to EAV
Potomac horse fever (PHF)
Semi-annual with one dose administered 4–6 weeks prepartum
Two-dose series: First dose 8–10 weeks prepartum Second dose 4–6 weeks prepartum
Strategic environmental control measures are important for effective control
Rotavirus
Three-dose series: First dose at 8 months’ gestation Second dose 4 weeks after the first dose Third dose 4 weeks after the second dose
Three-dose series: First dose at 8 months’ gestation Second dose 4 weeks after the first dose Third dose 4 weeks after the second dose
Check serum concentration of immunoglobulins in the foal to verify adequate passive transfer
Strangles
Inactivated M-protein subunit vaccines: Semi-annual with one dose given 4–6 weeks prepartum
Inactivated M-protein subunit vaccines: Three-dose series: Second dose 2–4 weeks after the first dose Third dose 4–6 weeks prepartum
The MLV intranasal strangles vaccine can be used to protect pregnant mares, but its use for the prepartum booster is not recommended because it does not reliably stimulate high levels of circulating antibody
Disease/vaccine Equine influenza
Comments The MLV intranasal influenza vaccine can be used to protect pregnant mares against influenza, but its use for the prepartum booster is not recommended because it does not reliably stimulate high levels of circulating antibody
Modified, with permission, from recommendations developed by the American Association of Equine Practitioners (AAEP) Infectious Disease Committee and posted on the AAEP website (www.aaep.org) in January 2008. ©AAEP 2008. a Core vaccines protect against diseases that are endemic to a region, are virulent or highly contagious, pose a risk of severe or fatal disease, have potential public health significance, and/or are required by law. Core vaccines have clearly demonstrable efficacy, and have a sufficiently high level of patient benefit and low level of risk to justify their use in all equids in North America. b Non-core (risk-based) vaccines are selected for use based on assessment of risk performed by, or in consultation with, a licensed veterinarian. Use of non-core vaccines will vary between individuals, populations, and/or geographical regions.
using these vaccines to document whether differences in serological responses equate to differences in efficacy. Early testing of bivalent (EEE/WEE) vaccines was performed by intracranial challenge with either EEE or WEE; the formalin-inactivated preparations demonstrated 100% protection. Veterinarians and horse owners often use combination products containing other antigens, such as tetanus, influenza, WNV, or equine herpesviruses for primary or booster immunization of horses against encephalomyelitis viruses.
Inactivated encephalomyelitis vaccines are considered to be safe for use during pregnancy; therefore, booster vaccination of pregnant mares 4 to 8 weeks before foaling is routinely recommended to enhance colostral concentrations of specific immunoglobulins. Neutralizing antibodies to WEE and EEE are transferred passively to foals through colostrum, and their concentration declines with an estimated half-life of 33 and 20 days, respectively. MDAs appear to confer protection and are detectable in the serum of many foals from vaccinated mares for at least 3 months
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and up to 7 months, depending on the post-nursing titer.3, 4, 28, 29 Several studies have shown that MDAs exert a profound inhibitory effect on the ability of foals to mount serological responses to inactivated bivalent WEE/EEE vaccines, which likely accounts for some of the reported cases of vaccine failure and resultant clinical EEE in vaccinated horses, particularly those less than 2 years of age.3, 4, 17, 28, 30 Studies have shown that 3-month-old foals born to immune mares consistently failed to mount a serological response to two doses of inactivated bivalent WEE/EEE vaccine, and the majority had not responded even after administration of a third dose.17, 20 Whereas many 6-month-old foals failed to seroconvert after administration of two doses of vaccine, most responded following administration of a third dose.17 Based on these data, inclusion of a third dose in the primary series, 3–5 months after administration of the second dose, is strongly recommended for primary immunization of foals and yearlings. Western equine encephalomyelitis has a lower mortality rate than EEE, and the prevalence of WEE in many western states is sufficiently low that the risk of foals acquiring infection during their first year of life is also low. Therefore, primary vaccination of foals of vaccinated mares in areas where mosquitoes die off in the winter and the risk of infection is low, is best completed when foals are 5–6 month of age or older in order to minimize the potential for MDA interference. Because foals born in the late spring and summer months are still less than 6 months of age by the time the mosquito season comes to an end in many regions, primary vaccination of these foals can be delayed until the spring of the yearling year. In contrast, EEE is a frequently fatal disease that poses a significant risk to foals during their first year of life, particularly in the Gulf states where competent vectors are present year-round.4, 30, 31 Therefore, most veterinarians in these regions recommend commencing primary vaccination of foals at 3 to 4 months of age using a three-dose primary series followed by a fourth dose before the onset of the next mosquito season, and semi-annual boosters thereafter, to maximize the chances of overcoming the inhibitory effects of MDA and inducing protection.4 West Nile virus West Nile virus (WNV), a member of the family Flaviviridae, is transmitted by mosquitoes, and infrequently by other bloodsucking insects, to horses, humans, and a number of other mammals from avian hosts, which serve as natural reservoirs for these viruses. West Nile virus (WNV) infection is now considered to be endemic in all mainland areas of North America and Mexico. Although cases have
been seen virtually year-round in the southeastern USA, the risk of acquiring infection is highest during those months in which mosquito activity peaks, typically July, August, September, and October in most areas of North America. The disease has been confirmed in foals as young as 3 weeks of age. Four fully licensed vaccines (West NileR , InnovatorTM , Pfizer Animal Health; Recombitek TM Merial; PreveNile , Intervet/Schering-Plough; and R West Nile-Innovator DNA, Pfizer Animal Health) are marketed for use in horses in North America. West Nile-InnovatorTM is an inactivated whole-virus vaccine that contains a metabolizable oil adjuvant.32 This vaccine is available as either a monovalent (single component) or as a multivalent vaccine containing other encephalitis virus antigens (EEE R is a carbopol-adjuvanted and WEE). Recombitek canarypox-vectored recombinant modified live virus (MLV).33–35 PreveNileTM is a non-adjuvanted chimeric yellow fever-vectored vaccine,36, 37 and West R Nile-Innovator DNA is a plasmid DNA vaccine with a metabolizable oil adjuvant.21 All four vaccines have met US Department of Agriculture (USDA) requirements for safety in tests, each involving more than 640 horses. Directions for primary immunization using West R R Nile-Innovator and Recombitek include administration of two doses of vaccine 3–6 weeks apart (consult the specific label). Optimal protection cannot be expected until 2 weeks after administration of the secR has been shown to ond dose, although Recombitek induce significant protection as early as 26 days after administration of the first dose when tested in both the mosquito challenge and intrathecal challenge models.34, 35 Primary immunization with PreveNileTM requires one dose. A challenge study in yearlings showed that 83% (five of six) were protected when challenged intrathecally 10 days after vaccination with one dose, indicating that onset of immunity is rapid.38 Rapid onset of immunity is an important feature when faced with the challenge of protecting naive horses that are being introduced into an endemic area, as is the case when horses from Europe and other non-endemic countries are imported into North America. None of the licensed vaccines currently marketed in the USA carries label recommendations for administration to pregnant mares. It is well recognized, however, that pregnant mares are at risk of acquiring infection from infected mosquitoes. Consequently, it has become accepted practice by many veterinarians to administer vaccines to pregnant mares on the reasonable assumption that the risk of adverse consequences of WNV infection far exceeds the reported adverse effects of use of vaccines in pregnant mares. Thousands of doses of West Nile-InnovatorTM
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Vaccination of Mares, Foals and Weanlings vaccine have been administered safely to pregnant mares, and a published study failed to document vaccine-associated adverse effects in a large populaR tion of pregnant mares.39 Although the Recombitek vaccine is a live vectored vaccine, the canarypox vector is incapable of replication in mammals and does not induce a viremia that could infect a fetus. In addition, a canarypox-vectored influenza vaccine available in Europe is licensed for use in horses during pregnancy; thus the vectored WNV vaccine is unlikely to be associated with an increased risk of adverse effects in pregnant mares. Similarly, data currently under review by USDA from studies involving a large number of pregnant mares suggests that PreveNileTM will likely also be shown to be safe for use in pregnant mares.40 As with other vaccines, it is sound practice to avoid administering West Nile vaccines to mares during the first 60 days of gestation unless conditions of imminent risk prevail. Booster vaccination of previously primed pregnant mares, 4 to 8 weeks before foaling, appears to induce a strong anamnestic serological response that provides their foals with passive colostral protection lasting at least 3 to 4 months.20 In contrast, a significant proportion of naive pregnant mares failed to seroconvert when the primary series of WNV-InnovatorTM vaccine was administered during the second half of gestation, perhaps reflecting pregnancy-associated downregulation of helper 2 Tcell responses.20 This observation adds further justification to the recommendation that when inactivated West Nile-InnovatorTM vaccine is used, the primary series is best completed before breeding. In a similar study, pregnancy did not appear to suppress the response of mares to primary immunization with R (W.D. Wilson et al., unpublished obserRecombitek vations, 2009). In contrast to findings with many other vaccines in the foals of immune mares, MDAs do not block the response of foals as young as 3 months of age to vaccination with the inactivated (West Nile InnovaR ), chimera (PrevetorTM ), recombinant (Recombitek TM R Nile ), or DNA (West Nile-Innovator DNA) WNV 20 vaccines. Although this finding is somewhat surprising for the inactivated vaccine, it might reasonably have been expected for the other vaccines because the vector systems accomplish transfection of cells and expression of the major E-peptide and Mpeptide antigens of WNV on the surface of antigenpresenting cells (APCs) in association with MHC class I and class II antigens. These peptide antigens are, therefore, not free in the tissues and circulation to be neutralized by MDAs. Primary vaccination of foals from properly vaccinated mares can be started by administration of the first dose of either West Nile-InnovatorTM or
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R as early as 3 months, but more typRecombitek ically at 4 to 6 months of age, followed by a second dose approximately 1 month later, then a third dose, 3 to 5 months after the second dose. A booster should be administered during the spring of the yearling year, after which the recommendations for annual revaccination of adult horses should be followed. Primary vaccination of foals from nonvaccinated, non-exposed mares should commence at 3 months of age or younger (as early as 1 month of age), depending on month of birth and seasonal level of activity of mosquito vectors in the area. The influence of MDAs on the response of foals to PreveNileTM was recently evaluated in 3-monthold and 5-month-old foals vaccinated with a single dose. Although significant titer responses were not observed post-vaccination, sensitization of cellmediated responses was accomplished, as evidenced by significant increases in expression of granzyme B, interleukin-2, perforin, and transforming growth factor-β (TGF-β), and non-significant increases in interferon-γ (IFN-γ ) expression in WNV-stimulated peripheral blood mononuclear cell(s) (PBMC) from foals vaccinated at 5 months of age.22 Similar, although non-significant, trends were seen in 3-monthold foals.22 PreveNileTM is labeled for administration of a single priming dose to foals aged 5 months or older, primarily because 5 months was the minimum age of foals used in challenge trials for licensure. Results of the above-cited study suggest that foals less than 5 months of age could be immunized successfully with PreveNileTM . Regardless, a second dose of vaccine should be administered before the onset of the next mosquito season. Preliminary data suggest R DNA vaccine that the plasmid West Nile-Innovator can also circumvent the potentially interfering effects of MDA.21
Rabies Rabies is an infrequently encountered neurological disease of equids resulting from inoculation of the rabies virus, a rhabdovirus, through the bite of infected (rabid) wildlife. It is logical to assume that the post-vaccination titer of circulating antibody correlates with protection, although this issue has not been clearly defined in horses. Available inactivated, tissue culture-derived vaccines licensed for use in horses R (RabvacTM 3, Boehringer Ingelheim; RM Imrab R TM 3, Merial; Rabguard TC , Pfizer; and EquiRab , Intervet/Schering-Plough) are potent immunogens that induce strong serological responses that peak within 28 days after intramuscular administration of a single dose to naive horses. Label directions for primary immunization include administration of one dose to horses aged 3 months or older followed by
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a second dose one year later. Thereafter, annual revaccination is recommended. Although none of the licensed vaccines carries a specific label approval for use in pregnant mares, adverse effects on the fetus have not been reported despite the fact that many veterinarians regularly administer inactivated rabies vaccines to pregnant mares. Alternatively, veterinarians may recommend that mares be vaccinated against rabies before breeding in order to reduce the number and type of vaccines given in the period before foaling. Because rabies antibodies persist in serum for a prolonged period, foals born to mares that are revaccinated before breeding or during late gestation acquire substantial titers of rabies antibody after ingesting colostrum. Studies in our laboratory have shown that the serological response of most 3-month-old foals from antibody-positive mares to vaccination is completely blocked, even when a two-dose primary vaccination series is used. Although the response to the first dose of vaccine is typically blocked in 6-month-old foals from antibody-positive mares, these foals appear to seroconvert after administration of a second dose 4 weeks later. Primary vaccination of foals from vaccinated mares should therefore be delayed until they are 6 months of age or older and should include two doses of inactivated vaccine administered approximately 4 weeks apart, followed by a third dose at 1 year of age. For foals from mares known not to be vaccinated against rabies, the primary vaccination series can be started according to manufacturers’ recommendations as early as 3 months of age and may comprise only one dose, although a two-dose series will almost certainly induce more durable immunity. For foals from mares of uncertain vaccination status, recommendations for foals from vaccinated mares can be followed. Alternately, rabies antibody titers can be determined on the mare or the foal as a prelude to determining the approach to be followed. Equine influenza Infection of the respiratory tract of horses with the orthomyxovirus, influenza A/equine/2 (H3N8), remains one of the most common causes of rapidly spreading outbreaks of respiratory disease. Immunity to the same (homologous) strain of H3N8 virus after natural infection persists for more than a year and involves both local and systemic humoral and cellular mechanisms. These include induction of large amounts of virus-specific neutralizing IgG and secretory IgA antibody in nasal secretions, high levels of circulating IgG antibodies, and genetically restricted antigen-specific cytotoxic T-lymphocytes (CTLs) that kill infected cells.41–45 Memory CTLs can
be detected in peripheral blood for at least 6 months after infection, and solid immunity persists even when circulating antibody titers have declined to low or non-detectable levels.42, 43, 46, 47 Available influenza vaccines include adjuvanted inactivated single-component vaccines (CalvenzaTM R 03 EIV, Boehringer Ingelheim; Fluvac Innovator , Pfizer Animal Health) and multicomponent vaccines (from Boehringer Ingelheim, Intervet/ScheringPlough, and Pfizer Animal Health) for intramuscular injection; one canarypox-vectored recombinant R vaccine (Recombitek Equine Influenza Virus vaccine, Merial) for intramuscular injection and one attenuated live, cold-adapted influenza vaccine R I.N., Intervet/Schering-Plough) for (Flu Avert intranasal (i.n.) administration. With the possible exception of immune stimulating complex (ISCOM) vaccines, which are not currently licensed in the USA, inactivated intramuscularly administered vaccines have limited potential to induce CTL or nasal secretory IgA responses and induce only low levels of neutralizing antibody in nasal secretions.44, 47, 48 The degree of protection induced by inactivated influenza vaccines is highly correlated with post-vaccination titers of circulating antibody, predominantly of the IgGa and IgGb sub-isotypes, as measured by hemagglutination inhibition (HI) or single radial hemolysis (SRH) tests.49–52 SRH levels ≥ 100 mm2 are considered to be at least partially protective; however, levels greater than 140 mm2 are required for successful prevention of disease.50 Protection induced by inactivated vaccines is of limited duration (up to about 7 months, depending on the vaccine) and is manifested as a reduction in clinical signs and attenuation of viral shedding in R horses exposed to infection.44, 53 Fluvac Innovator contains a KY/97 H3N8 strain in a metabolizable R oil (MetaStim ) adjuvant. CalvenzaTM EIV is adjuvanted with Carbopol (a carbopolymer) and is the only inactivated influenza vaccine currently licensed in North America that contains antigens from H3N8 viruses of both the American and European lineages (Kentucky/95 and Newmarket/2/93). The initial two doses of this vaccine are administered intramuscularly; subsequent doses may be administered intramuscularly or intranasally. It is proposed, but not proved, that administration of booster doses by the intranasal route may provide a stronger local mucosal immune response. This vaccine is licensed for use in horses aged over 6 months, including pregnant mares. Pfizer Animal Health and Boehringer Ingelheim also market several multicomponent combination vaccines that contain the same inactivated influenza antigens as in their singlecomponent products, but also contain tetanus, WEE and EEE virus, EHV, or WNV antigens. In addition,
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Vaccination of Mares, Foals and Weanlings Intervet/Schering-Plough markets multicomponent vaccines containing strains representing both North American (Kentucky/93, Kentucky/2002) and Eurasian (Newmarket/2/93) lineages. R I.N. Protection induced by intranasal Flu Avert is presumably mediated through induction of local immune responses in the respiratory tract because this vaccine does not typically induce high levels of circulating antibody.54, 55 This vaccine, which contains a Kentucky/1991 strain of North American lineage, was found to be highly efficacious in blinded, controlled challenge studies conducted 5 weeks, 6 months, and 1 year after administration of a single R dose to naive horses.55 Subsequently, Flu Avert I.N. was shown to cross-protect against European H3N8 strains, as well as against North American strains isolated during the late 1990s and early 2000s, and to induce a rapid onset of protection within 7 days of administration of a single dose to naive R horses.54, 56 Currently, Flu Avert I.N. is licensed for use in non-pregnant horses aged 11 months or older, primarily because this was the youngest age of the horses used in the challenge studies for licensing. The injectable canarypox-vectored recombinant equine R influenza vaccine (Recombitek Equine Influenza Virus vaccine, Merial) incorporates the HA gene from the Kentucky/94 and Newmarket/2/93 H3N8 strains and has been shown to induce strong protection in challenge studies.57 It contains a carbomer polymer adjuvant in the diluent and invokes a broad array of humoral and cellular immune responses.57 Challenge studies document onset of protection as soon as 2 weeks after completion of a two-dose primary series and persistence of solid protection for at least 5 months. Administration of a booster dose at 5 months induced a strong anamnestic response that provided solid protection persisting for at R Equine Influenza least 12 months.58 Recombitek Virus vaccine is licensed for vaccination of healthy horses as young as 5 months of age, and preliminary evidence suggests that this vaccine will be able to circumvent the inhibitory effect of maternal antibodies.59 Routine revaccination at an interval of 6 months appears to be appropriate for the intramuscularly administered inactivated and intranasally administered MLV influenza vaccines currently marketed in North America. A revaccination interval of 12 months is recommended for the recombinant vaccine, although this recommendation has not yet been tested in the field setting in North America. These ‘routine’ revaccination protocols should be customized, by adjusting timing of boosters or inclusion of an additional booster, to achieve maximum protection during periods when the risk of exposure is high. For example, strategic revacccination 1 month prior to being
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placed at high risk of exposure, such as at a show, sale, training barn, or another breeding farm, is justified to maximize protection. Revaccination of pregnant mares 4 to 8 weeks prior to foaling with a vaccine that stimulates a robust serological response is recommended. Although the R I.N. vaccine intranasally administered Flu Avert induces good protection and is frequently incorporated into the core influenza vaccination program for broodmares, this vaccine does not routinely stimulate high levels of circulating antibody, at least when used for primary immunization. An inactivated or canarypox-vectored recombinant injectable vaccine is therefore currently recommended for pre-foaling booster vaccination of pregnant mares.20 Primary vaccination of foals against influenza is heavily influenced by the post-nursing titer of specific antibody in the serum of the foal because MDA interference profoundly impacts the foal’s ability to respond to influenza vaccines administered during the first year of life. Foals born to seronegative, non-vaccinated mares respond appropriately to influenza vaccines; therefore, primary vaccination can commence at 3 months of age or younger if significant risk of exposure to influenza exists. In contrast, maternal antibodies have been shown to completely block the serological response of foals to a primary immunization series comprising two or more doses of inactivated influenza vaccines when the first dose is administered when the foal is younger than 6 months of age.5, 6, 9,10–13,17 Interference from MDA may persist until 9 months of age or beyond for foals with high antibody titers post-nursing; therefore, primary vaccination of foals from immune mares should be delayed as long as possible and preventive measures should focus on avoiding the introduction of infected horses.5, 6, 9,10–13,17, 20 Studies in Newmarket, UK, have shown that influenza virus infection is rare in Thoroughbred yearlings before they enter training, suggesting that the risk of influenza is low in horses less than 1 year of age born to mares in herds that are well vaccinated.60–62 Therefore, there appears to be little justification for vaccinating young foals from vaccinated mares against influenza, as was recommended in the past.29, 63, 64 R The intranasal MLV (Flu Avert I.N.) is licensed for vaccination of horses aged 11 months or older. Whereas this vaccine has been shown to be safe in foals as young as 2 months of age,65 published data regarding the potential for MDAs to interfere with the response are lacking. Unpublished observations suggest that MDAs interfere with the response of foals aged between 3 and 6 months, whereas foals vaccinated at 7 months of age were protected against virulent challenge (Holland RE and Chambers TM, 2005, personal communication). Pending publication
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of well-controlled studies, it is recommended that if R I.N. vaccine is administhe first dose of Flu Avert tered before 11 months of age, a second dose should be administered at 11 months of age or older.66 Re R combitek Equine Influenza Virus vaccine has been shown to be safe in foals as young as 4 months of age, and effective priming has been documented after administration of the first dose of this vaccine to foals age 10 to 20 weeks that had detectable MDA at the time of vaccination.59 However, the minimum age recommended on the label for vaccination of foals from immunized dams is 5 months. If the foal experiences failure of passive transfer of maternal antibodies or if the mare is seronegative for influenza, vaccination can commence at 4 months of age but should include an additional dose in the primary series. Equine herpesviruses (EHV-1 and EHV-4) The respiratory tract is the primary route of infection for both equine herpesvirus type 1 (EHV-1) and equine herpesvirus type 4 (EHV-4), both of which cause respiratory tract disease that varies in severity from subclinical to severe, and is characterized by fever, lethargy, anorexia, nasal discharge, and cough.67 EHV-1 is also an important cause of abortion, neonatal death, and myeloencephalopathy. Seroepidemiological studies indicate that the vast majority of foals become infected with EHV-1 and EHV-4 during the first few months of life but the clinical disease syndromes resulting from these infections are not always well defined, perhaps reflecting the modulating effect of maternally derived antibodies.68–70 Recurrent or recrudescent clinically apparent infections are seen in weanlings, yearlings, and young horses entering training, especially when horses from different sources are commingled.67, 71 As with herpesvirus infections in other species, horses typically fail to clear primary infections with either EHV-1 or EHV-4, the result being that most horses in the population remain latently infected with both viruses.67, 69, 72 Correlates for protection against EHV-1 and EHV4 infection are not yet clearly defined. Systemic and mucosal humoral and cellular immune responses all appear to play a role.73, 74 The finding that the presence of MHC class I-restricted CTL precursors in peripheral blood is correlated with protection against EHV-1 infection confirms the importance of cellular immunity.73 Infection with EHV-1 induces a strong humoral response but protection from reinfection is short-lived and is not achieved until the horse has experienced multiple infections with homotypic virus.67, 73 Although in one study no clear relationship between protection from EHV-1 infection and concentrations of circulating antibody induced by
vaccination or infection was found, the duration and amount of virus shedding from the nasopharynx was reduced in animals with high levels of circulating EHV-1 neutralizing antibody.73 In contrast, the efficacy of an inactivated whole-virus EHV-1/4 vaccine in reducing clinical manifestations and virus shedding was clearly correlated with vaccine-induced antibody levels in a study in which weanlings aged 5 to 8 months were challenged with virulent EHV-1 2 weeks after completion of the two-dose primary series.75 As with EHV-1, high levels of vaccineinduced circulating virus neutralizing (VN) antibody markedly reduce virus shedding and clinical signs after EHV-4 challenge infection.73,75–77 Because EHV4 replication is largely confined to epithelial cells of the upper respiratory tract, it is likely that mucosal immunity is also important in protection.73 A variety of killed vaccines are available; these include those licensed only for protection against respiratory disease, all of which currently contain R a low antigen load, and two (Pneumabort-K +1b, R , Intervet/ScheringPfizer Animal Health; Prodigy Plough) that contain a high antigen load and are licensed for protection against both abortion and respiratory disease. Performance of the killed lowantigen-load respiratory vaccines is variable, with some vaccines outperforming others. Performance of the killed high-antigen-load abortion/respiratory vaccines is superior, resulting in greater antibody responses and some evidence of cellular responses to vaccination. This factor may be a good reason to choose the high-antigen-load abortion/respiratory R EIV/EHV vaccines. Alternatively, the Calvenza (Boehringer Ingelheim) respiratory vaccine induces high titers of VN antibody comparable to those induced by the high-antigen load abortion/respiratory vaccines.27 A single manufacturer provides a liR censed modified live EHV-1 vaccine (Rhinomune , Pfizer), which to date has not been compared directly with high-antigen-load respiratory/abortion vaccines. This modified live vaccine has been shown to offer superior clinical protection and to reduce viral shedding in a comparison with a single killed lowantigen-load respiratory vaccine.78 Vaccination with either EHV-1 or EHV-4 can provide partial protection against the heterologous strain; vaccines containing EHV-1 may be superior in this regard. The principal indication for use of equine herpesvirus vaccines is prevention of EHV-1-induced abortion in pregnant mares. A secondary indication is reduction of signs and spread of respiratory tract disease (rhinopneumonitis) in foals, weanlings, yearlings, and young performance and show horses that are at high risk of exposure. Because currently available inactivated vaccines do not block infection with equine herpesviruses, the most we can hope for
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Vaccination of Mares, Foals and Weanlings when using these vaccines is reduction of severity of clinical signs and attenuation of virus shedding to help protect herd mates. Consistent vaccination appears to reduce the frequency and severity of herpesvirus-induced disease. Although convincing evidence is lacking, field experience suggests that whereas the incidence of sporadic EHV-1-induced abortion in individual mares has not changed, the incidence of abortion storms caused by EHV-1 has declined significantly since the introduction and widespread use of EHV-1 vaccines in the USA.72, 79 Outbreaks of abortion and associated perinatal foal death, however, continue to occur on occasion in herds of vaccinated mares. Of the vaccines currently licensed for use in pregnant mares in North America, only inactivated R +1b, monovalent EHV-1 vaccines (Pneumabort-K R Pfizer Animal Health; Prodigy , Intervet/ScheringPlough) containing abortifacient strains of EHV-1 carry a label claim for preventing abortion, whereas at least one bivalent EHV-1/4 vaccine is licensed for R EHVprevention of abortion in Europe (Duvaxyn 1/4, Intervet/Schering-Plough). One of the vaccines R +1b inavailable in North America, Pneumabort-K corporates both the 1p and 1b subtypes of EHV-1 to reflect the documented increase in the proportion of EHV-1 abortions caused by the 1b subtype that occurred during the 1980s as compared to earlier years.80 Pregnant mares should be vaccinated during the fifth, seventh, and ninth months of gestation. Many veterinarians also recommend a dose during the third month of gestation. Similarly, vaccination of mares with an inactivated EHV-1/EHV-4 vaccine at the time of breeding and again 4 to 6 weeks before foaling is commonly practiced to enhance concentrations of colostral immunoglobulin for transfer to the foal. However, no published reports document the effectiveness of this approach in raising titers of specific antibody in mares that have already been vaccinated against EHV-1 three times during the previous 5 months. Vaccination of barren mares and stallions with either a bivalent EHV-1/4 vaccine or a monovalent EHV-1 vaccine before the start of the breeding season, and thereafter at 6-month intervals, is recommended, with the goal of increasing herd immunity in an attempt to reduce viral shedding and challenge to pregnant mares on breeding farms.72 The modified live-virus EHV-1 vaccine R , Pfizer) has been used by some (Rhinomune practitioners for many years as an aid to prevention of EHV-1 abortion,81 even though this vaccine is not currently labeled for this use. However, several recent developments have created a renewed interest in the potential for MLV vaccines to protect horses against manifestations of EHV-1 and EHV-4 infection. Sequencing of the EHV-1 genome has made
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it possible to document the nature of the mutation encoding attenuation, mediated through truncation of the gp2 glycoprotein, of the KyA strain.82 Similar studies may soon yield information regarding the mutation underlying attenuation of the RAC-H R was derived. strain from which Rhinomune Specific antibodies against both EHV-1 and EHV-4 are passed in colostrum,7, 15, 16, 18, 83 and field studies with EHV-1 MLVs indicate that colostral antibodies exert a profound inhibitory effect on serological responses to vaccination up to at least 5 months of age.15, 84, 85 However, a cytotoxic cellular immune response to both EHV-1 and EHV-4 was induced in a substantial percentage of foals vaccinated with an EHV-1 MLV in the presence of maternal antibody, even though humoral responses were often absent.86 It is uncertain whether these responses would provide protection against natural challenge. Recent studies with two different commercially available inactivated bivalent EHV-1/4 vaccines, and one inactivated EHV-4/influenza vaccine, have shown that the majority of foals from EHV-vaccinated mares do not mount a detectable neutralizing antibody response to vaccines administered at 3 and 4 months of age, even when three doses are administered in the primary series.16–18 An increased proportion of foals responded when vaccinated with a three-dose series starting at 5 or 6 months of age but a substantial number still failed to seroconvert.17, 18 Some foals with low or undetectable levels of serum neutralizing (SN) antibody at the time of vaccination failed to mount a serological response, suggesting that low levels of antibody, below the lower limit of detection of the SN test based on EHV-1 antigen, are capable of inhibiting the serological response to inactivated EHV-1/4 vaccines.18 The failure of a large proportion of foals less than 6 months of age to mount serological responses to inactivated EHV-1/4 vaccines and the influence of antibody titer at the time of vaccination on failure to respond has been confirmed using sensitive glycoprotein D (gD) and gG enzyme-linked immunosorbent assays (ELISAs) in studies on commercial stud farms in Australia.87 In parallel studies, these researchers concluded that mares were the source of infection for foals and that intensive use of inactivated EHV-1/4 vaccines on breeding farms in Australia had minimally impacted the infection rate of young foals and weanlings with EHV-1 and EHV-4.68, 69, 88 Considering the uncertainty regarding the role of EHV-1 and EHV-4 as causes of clinically important respiratory disease, the lack of published data regarding the efficacy of available vaccines in preventing infection and establishment of latency, and results of a recent study documenting the poor serological responses of naive horses to a number of killed low-antigen-load EHV respiratory vaccines currently
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marketed in North America,27 there appears to be little rationale to support the common practice of frequent revaccination of foals, weanlings, yearlings, and young performance horses against EHV-1 and EHV-4.89 Furthermore, an obvious dilemma in designing a vaccination strategy to prevent EHV-1 and EHV-4 infection in foals and weanlings is that if primary immunization is delayed until 6 months of age or older to reduce the likelihood of MDA interference, foals are likely to encounter field infection before the three-dose primary series can be completed. It is, therefore, unreasonable to expect a high degree of efficacy for vaccination programs designed to protect foals and weanlings against EHV infection using available vaccines. Despite these uncertainties, many practitioners elect to vaccinate against both EHV-1 and EHV-4. Under these circumstances, a reasonable compromise would be to start foal vaccination at 4 to 6 months of age using two doses of an inactivated bivalent vaccine or an EHV-1 MLV administered 3 to 4 weeks apart, followed by administration of a third dose 2 to 5 months later. Revaccination at 4–6-month intervals thereafter using either an inactivated bivalent vaccine or a modified live EHV-1 vaccine appears appropriate for yearlings and young performance or show horses that experience contact with other horses. Streptococcus equi ssp. equi infection (strangles) Strangles, a highly contagious disease caused by the bacterium Streptococcus equi ssp. equi (S. equi), affects primarily weanlings and yearlings, although horses of any age can become infected if not protected by previous exposure to the organism or by vaccination. A high concentration of young susceptible horses, together with regular movement of horses on and off the facility, contributes to an increased risk of infection on breeding farms, to the extent that strangles is a persistent endemic problem on many farms. During recovery from strangles, most horses develop a solid immunity that persists in more than 75% of animals for 5 years or longer,90 indicating that induction of durable protection through vaccination is biologically feasible if the protective antigens can be identified and presented in an appropriate manner.91 Although the basis for acquired resistance to strangles is not completely understood, the finding that recovered horses rapidly clear intranasally inoculated S. equi despite not making circulating antibody to its surface proteins, indicates that to be highly effective, a strangles vaccine must stimulate local nasopharyngeal tonsillar immune clearance responses and that serum antibody is of lesser importance.92 This conclusion is further supported by the finding
that ponies with high levels of circulating antibody to multiple, unique, surface-exposed and secreted proteins after systemic vaccination remained susceptible to challenge with S. equi.92 The cell wall M-protein of S. equi (Se-M) is recognized in the acquired immune response to S. equi infection, a response that involves both production of local antibodies in the nasopharynx and circulating opsonophagocytic antibodies.93–95 The opsonophagocytic antibodies are predominantly of the IgGb sub-isotype but also include IgGa and IgA, whereas IgGb and, later, mucosal IgA predominate in nasopharyngeal secretions.2, 94 Strangles vaccines licensed for use and marketed in North America include two inactivated, adjuR vanted M-protein cell wall extracts (Strepvax R II, Boehringer Ingelheim; Strepguard with R Havlogen , Intervet/Schering-Plough; prepared by extraction with hot acid or mutanolysin plus detergent, respectively) and one attenuated live vaccine R (Pinnacle I.N., Pfizer Animal Health) derived from a non-encapsulated mutant of S. equi for intranasal administration.96 M-protein vaccines induce a good opsonophagocytic antibody response in serum but a minimal mucosal IgA response, which likely accounts for the incomplete protection observed when they are used in the field.94, 97 However, data do exist to document that vaccination using injectable SeM vaccines reduces the attack rate and severity of strangles in herds with endemic infection.97–99 The live intranasal vaccine has been shown to induce a relevant mucosal immune response and partial or complete protection, but may do so without inducing a strong serological response.100, 101 Because vaccinal organisms in the intranasal vaccine must reach the inductive sites for immunity in the pharyngeal and lingual tonsils, accurate vaccine delivery is critical to vaccine efficacy. On breeding farms, efforts should be concentrated on preventing infection of foals and weanlings by booster-vaccinating broodmares 4 to 6 weeks before foaling to maximize colostral content of antibodies. Whereas the intranasal vaccine has been shown to be safe for use in mares at all stages of pregnancy and can be used in mares in the face of an outbreak, it does not reliably stimulate high levels of circulating antibody. For this reason, intramuscularly administered inactivated SeM products are preferred for prefoaling booster immunization of mares. Antibodies of the IgG and IgA class recognizing the SeM are passively transferred to the foal through colostrum and are also present in the milk of immune mares.102 Antibodies of predominantly the IgGb isotype are absorbed from colostrum and redistribute to the nasopharyngeal mucosa.2 These IgGb antibodies and, to a lesser extent, the SeM-specific IgA antibodies that
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Vaccination of Mares, Foals and Weanlings are present in milk and passively coat the pharyngeal mucosa of nursing foals, provide protection to most foals up to the time of weaning.2, 93, 102 Serological (ELISA) responses to M-protein vaccines are poor in foals, most likely due to the inhibitory effect of maternal antibodies. When an inactivated M-protein vaccine is used for primary vaccination of foals, it is recommended that the initial series begins at 4 to 6 months of age, using three doses administered at 3- to 6-week intervals, followed by semi-annual boosters for as long as conditions of high risk prevail. Whereas the intranasal MLV may be less susceptible than the inactivated extract vaccines to MDA interference, this issue has not been investigated and the manufacturer does not recommend administration of this vaccine to horses less than 9 months of age. Considering that on farms where strangles is endemic, foals often become infected around the time of weaning, at 4 to 8-months of age, it is difficult to protect them if vaccination is delayed until 9 months of age. Therefore a reasonable compromise on breeding farms where the risk of strangles infection is high and mares are on a regular vaccination program would be to begin primary vaccination of foals using the intranasal live vaccine as early as 4 months of age. The recommended two-dose primary series administered 2 to 3 weeks apart should be followed by a third dose 3 to 4 months later and boosters at 6- to 12-month intervals thereafter, depending on risk of infection. The intranasal vaccine has been administered to foals as young as 5 or 6 weeks of age during outbreaks. If used in this manner, a third dose of the vaccine should be administered 2 to 4 weeks before the foal is weaned to optimize protection during this high-risk period. Although there are few reports of adverse effects attributable to use of the intranasal strangles vaccine in young foals, the inability of foals to mount an adequate mucosal IgA response during the first month of life and the potential for interference by maternal antibodies, suggest that foals are unlikely fully to benefit from intranasal strangles vaccine administered before 4 months of age. Injectable strangles vaccines tend to cause local reactions at the site of injection more often than do other equine vaccines. Injection in the gluteal muscles is not recommended because gravitational drainage along fascial planes can prove troublesome in the event that an abscess develops at the injection site. In addition, purpura hemorrhagica, a serious and sometimes life-threatening systemic immune complex (Arthus-type) vasculitis manifested as edema with or without petechial hemorrhages on mucosal surfaces, has been observed with low frequency in the weeks after administration of strangles vaccines. Inactivated extract vaccines are implicated more of-
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ten than the intranasal MLV, but all strangles vaccines have the potential to induce purpura. The antigen present in immune complexes is SeM, along with antibodies of the IgA class. Because a high serum IgG titer against S. equi appears to be associated with an increased risk of developing purpura, routine testing for specific IgG antibodies using a commercially available ELISA test has been recommended as a means of preventing vaccine-associated purpura.101 Horses with titers of 1:1600 or greater in the SeM ELISA and those known to have had strangles during the previous year should not be vaccinated.101 It is inadvisable to vaccinate horses that have active strangles, as well as those that are incubating infection. Potomac horse fever (equine monocytic ehrlichiosis) Potomac horse fever (PHF) is caused by Neorickettisa risticii (formerly Ehrlichia risticii). Originally described in 1979 as a sporadic disease affecting horses residing in the northeastern USA near the Potomac River, the disease has now been reported in horses in 43 states in the USA, three provinces in Canada (Nova Scotia, Ontario, and Alberta), South America (Uruguay, Brazil), Europe (The Netherlands, France), and India. PHF is seasonal, occurring between late spring and early fall in temperate areas, with most cases in July, August, and September at the onset of hot weather. Documentation of the involvement of operculate freshwater snails and aquatic insects such as caddisflies and mayflies in the life cycle of N. risticii has permitted formulation of focused control measures directed at minimizing exposure of horses to the habitats occupied by these species during the summer and fall months when disease risk is highest in endemic areas.103 Risk reduction is best accomplished by denying horses access to river banks, creek beds, and irrigation ditches, as well as pastures that have recently been flooded or flood-irrigated. Recovery after natural infection with N. risticii induces a strong antibody response and durable protection from reinfection lasting 20 months or longer. Several inactivated PHF vaccines for inR tramuscular administration (Potomavac , Merial; R PotomacGuard , Pfizer Animal Health; PHFGardTM , Pfizer; and EquovumTM PHF, Boehringer Ingelheim) are licensed for use in horses with the label claim that they aid in prevention of PHF. Two of these are also available combined with a rabies vaccine. Although challenge studies and non-controlled field studies with an aluminum hydroxide-adjuvanted R , Schering-Plough) suggested vaccine (PHF-Vax benefit in terms of reduced morbidity and mortality, large-scale epidemiological investigations in New York State have failed to document conclusively
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clinical or economic benefit from annual vaccination of adult horses with currently available vaccines.104, 105 Failure of a substantial number of individual horses to mount an immune response to inactivated PHF vaccines, heterogeneity of N. risticii isolates, the presence of only one N. risticii strain in vaccines, and much more rapid waning of immunity after vaccination than after natural infection, likely account for the observed failure of vaccines to provide protection against field infection.106, 107 Regardless, many practitioners who work in endemic areas believe that severity of disease is attenuated and mortality is reduced in vaccinated horses when vaccines are administered at 4- to 6-month intervals, with administration of one booster timed to precede the anticipated period of peak challenge. Available vaccines are licensed for use in stallions and pregnant mares and can be administered to gestating mares, 4 to 8 weeks before foaling to maximize passive transfer of specific antibodies to foals through colostrum. Whereas approximately 67% of foals from antibody-positive mares were antibody negative by 12 weeks of age, antibody was detectable in 33% of foals up to 5 months of age. On the basis of these findings, the low risk of clinical disease in young foals, and the apparent susceptibility to infection of two foals vaccinated earlier than 12 weeks of age, primary vaccination of foals from antibody-positive dams should begin with a two-dose primary series starting at 5 to 6 months of age or older, followed by administration of a booster dose 3 to 5 months later.108 However, the efficacy of this recommended regimen requires further study. If the primary series of two vaccinations is initiated before 5 months of age, additional doses should be administered at monthly intervals up to 5 months of age to maximize the likelihood that an immunological response is achieved. Vaccination of foals in endemic areas is further complicated by the distinct seasonal incidence of disease in July, August, and September, in the northern hemisphere, a time when most foals are aged between 2 and 6 months and may be subject to maternal antibody interference with vaccination. Botulism Botulism is a neuromuscular paralytic disorder caused by one of eight distinct neurotoxins (A, B, Ca, Cb, D, E, F, G) produced by Clostridium botulinum, a soil-borne, spore-forming, saprophytic, anaerobic, Gram-positive bacterium.109 Of the seven serogroups (A through G) of C. botulinum, types A, B, C, and D have been reported to cause disease in horses, with types B and C being responsible for most cases. 109 Three forms of botulism – toxicoinfectious botulism (shaker foal syndrome), forage poisoning, and
wound botulism – have been observed in horses. Shaker foal syndrome, almost all cases of which are caused by C. botulinum type B, results from toxin produced by vegetation of ingested spores in the intestinal tract. This syndrome is a significant problem in foals aged between 2 weeks and 8 months in Kentucky and in the mid-Atlantic seaboard states, and occurs sporadically in other areas.110–112 Currently toxicoinfection with C. botulinum type C is being investigated as a cause of equine grass sickness, a largely fatal, pasture-associated dysautonomia affecting horses mainly in Great Britain, continental Europe, and Australia, with reports of isolated cases in the USA. R A toxoid vaccine (BotVax B, Neogen Corp.) directed against C. botulinum type B is licensed for use in horses in the USA. Its primary indication is prevention of the shaker foal syndrome via colostral transfer of antibodies induced by vaccination of the mare. Label directions for primary vaccination of mares include administration during gestation of a series of three doses 4 weeks apart, scheduled so that the last dose will be administered 4 to 6 weeks before foaling to enhance concentrations of specific immunoglobulin in colostrum (i.e., months 8, 9, and 10 of gestation). Subsequently, mares should be revaccinated annually with a single dose administered 4 to 6 weeks prior to foaling. A similar type B toxoid is available to protect foals in endemic areas in Australia.113 Passively derived colostral antibodies appear to protect most foals for 8 to 12 weeks, although botulism has been reported in foals from properly vaccinated dams.111–113 The status of MDA transfer in each foal should, therefore, be verified. Maternal antibodies do not appear to interfere with the response of foals to primary immunization against botulism;114 therefore a primary series of three doses of vaccine, administered 4 weeks apart, can be started when foals in endemic areas are 2 to 3 months of age or older. Currently, no licensed vaccines are available for preventing botulism due to C. botulinum type C or other toxin subtypes, and cross-protection between the B and C subtypes does not occur; therefore, routine vaccination against C. botulinum type C is not currently practiced. Foals and adult horses with clinical botulism may be treated with botulinum antitoxin administered intravenously. Antitoxin is not effective against toxin that has been translocated to motor end plates; therefore, clinical sings may progress for 12 to 24 hours after administration of the antitoxin or until all internalized toxin has attached to motor end plates. The dose of botulinum type B antitoxin recommended for treating a foal is 30,000 IU, and for an adult it is 70,000 IU. Foals of unvaccinated mares born in or being moved to endemic areas may benefit from
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Vaccination of Mares, Foals and Weanlings transfusion with plasma from a vaccinated horse or from administration of C. botulinum type B antitoxin. The efficacy of these practices needs further study. Vaccination with type B toxoid as described above is a less expensive alternative to passive immunization. Equine viral arteritis Equine viral arteritis (EVA) is caused by equine arteritis virus (EAV), an RNA virus that is the prototype virus in the family Arteriviridae of the genus Arterivirus, order Nidovirales. Whereas all horse breeds appear to be equally susceptible to EAV, the prevalence of infection, as determined by seroconversion, is much higher in some breeds, notably Standardbreds and Warmbloods, than in others. Despite the high seroprevalence of infection in Standardbreds, clinical disease is rarely observed in this breed, indicating that subclinical infection is common.1, 115 Conversely, Thoroughbreds and most other breeds have a low seroprevalence of infection, but are more likely to show fulminant clinical signs when they become infected. Most primary EAV infections are subclinical or asymptomatic. Clinical signs, if they occur, typically develop 3 to 7 days post-infection and vary in severity, both within and between outbreaks. EAV is of special concern because abortion is a frequent sequel to infection in the unprotected pregnant mare. In addition, EAV can cause life-threatening pneumonia or pneumoenteritis in young foals, and infection of the post-pubertal colt or stallion frequently establishes a long-term carrier state that constitutes the most important means of persistence of infection in equine populations and poses an ever-present risk of introducing infection into new populations through breeding by natural cover or by artificial insemination.1, 116 Historically, large-scale outbreaks of EVA were relatively infrequent. However, the number of confirmed occurrences appears to be increasing, likely as a result of increased global movement of horses, increased accessibility of carrier stallions, and increased utilization of shipped cooled or frozen virus-infected semen. Outbreaks can be associated with serious economic consequences, as clearly exemplified by the 2006 multi-state outbreak in Quarter Horses that was propagated by widespread shipment of semen from the index cases, two inapparently infected carrier stallions in New Mexico. Although identification of carrier stallions is the primary means of controlling EVA, vaccination also constitutes an important approach to minimizing the spread and consequences of infection. A modified R , Pfizer Animal Health), based live vaccine (Arvac on an attenuated strain of EVA virus, has proven to be safe and effective in bringing the outbreaks un-
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der control, and for routine prevention, when used to vaccinate stallions, non-pregnant mares, and prepubertal colts.1, 117 Strategic use of the MLV has formed the cornerstone of a highly successful program to control EVA in the Kentucky Thoroughbred breeding population for many years.1 Indications for vaccination against EVA include: 1. To protect stallions against infection and subsequent development of the carrier state. 2. To immunize seronegative mares before being bred by natural cover or artificial insemination with semen from a known or suspected infected stallion. 3. To curtail outbreaks in non-breeding populations. Vaccination in the face of an EVA outbreak in concentrated populations of performance horses at racetracks has been successful in controlling horizontal disease dissemination within 7 to 10 days. Primary immunization with the modified live vaccine involves intramuscular administration of a single dose, with a booster administered annually thereafter. Virus-neutralizing antibodies are induced within 5 to 8 days after vaccination and persist for at least 2 years.1, 118 Revaccination induces high VN antibody titers that persist for several breeding seasons.118 Although the current MLV is highly attenuated and has been shown to be safe and effective in stallions and non-pregnant mares, a small proportion of first-time vaccinated horses develop mild febrile reactions and transient lymphopenia after vaccination, and vaccine virus may be isolated sporadically from the nasopharynx and buffy coat for 7 days, but occasionally for up to 32 days, after vaccination.1,118–120 Vaccinated stallions do not shed vaccine virus in either semen or urine.118 Primary vaccination provides sustained clinical protection against EVA but does not prevent reinfection and subsequent limited replication and shedding of field strains of virus.121 The frequency, duration, and amount of viral shedding via the respiratory tract are, however, significantly reduced in vaccinates. Vaccinated mares may shed field virus transiently after being bred to carrier stallions; therefore, isolation of these individuals for 21 days after breeding is recommended.1 Annual revaccination of breeding stallions, 28 days before the start of the breeding season, is highly recommended as a means of preventing establishment of the carrier state.1 Annual revaccination of mares being bred to carrier stallions should occur at least 21 days before breeding. The MLV is not recommended for use in pregnant mares, especially during the last 2 months of gestation, or in foals less than 6 weeks of age, except in emergency situations when there is a high risk of exposure. Apparent infection of
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the fetus with the modified live vaccine strain after vaccination of pregnant mares has been documented in rare instances.118, 122 Foals born to seropositive mares become seropositive after ingesting colostrum. Maternally derived antibodies decay with a mean half-life of approximately 32 days, with the result that foals become seronegative between 2 and 7 months of age.8, 123 Maternal antibodies are unlikely to interfere with the response to vaccine administered at 7 months of age or older.8 However, when foals less than 6 months of age are vaccinated during conditions of high risk, they should be revaccinated after 6 months of age. Establishment of the carrier state appears to depend on the high levels of androgens circulating in intact stallions and can be prevented by vaccinating colts, preferably before puberty and before they are used for breeding. 1 Vaccination of prepubertal colts at 6 to 12 months of age is therefore central to effective control of the spread of EAV infection and should be strongly encouraged in breeds such as Standardbreds and Warmbloods in which EVA is prevalent, and on facilities where the risk of infection is high. Persistent infection has never been documented in a stallion that was properly vaccinated with the licensed modified live vaccine prior to exposure. When planning a vaccination program against EVA, it is important to consult with state and/or federal animal health officials to ensure that any such program complies with the state’s control program for EVA, if one exists. Because it is not possible to differentiate a vaccine-induced antibody response from that due to natural infection, it is strongly recommended that, before vaccination, all first-time male vaccinates are tested and confirmed negative for antibodies to EAV by a USDA approved laboratory (http://www.aphis.usda.gov/animal health). Mares intended for export should be similarly tested. In instances where there is uncertainty or concern over whether vaccination against EVA could prevent the export of a horse to a particular country, it is advisable to consult the federal area veterinarian (http://www.aphis.usda.gov/animal health) in charge in the state to determine the specific import requirements of that country. Several countries bar entry of any equid that is serologically positive for antibodies to EAV, regardless of vaccination history. Countries that do accept EVA-vaccinated horses typically require stallions or colts to have a certified vaccination history and confirmation of prevaccination negative serological status.
Future directions A killed-virus vaccine (Artervac, Pfizer Animal Health) is licensed for use in the UK, Ireland, France,
Denmark, and Hungary, and a killed-virus vaccine is also used in Japan. As with the MLV licensed in the USA, serological responses to these inactivated vaccines cannot be distinguished from those resulting from natural infection. Development and marketing of a marker vaccine that not only affords protection but also allows vaccinated horses to be distinguished serologically from inapparently infected carriers would greatly facilitate control, and even eradication, of EAV from horse populations. Several ‘new generation’ EAV vaccines that potentially meet these criteria have been developed in recent years. These include an MLV differentiating infected from vaccinated (DIVA) vaccine with a deletion in the GP5 ectodomain,124, 125 a DNA vaccine that incorporates open reading frames (ORFs) 2b, 5, and 7,126, 127 and a subunit EAV vaccine using recombinant replicon particles derived from a vaccine strain of VEE virus that includes genes encoding both major envelope proteins (GP5 and M) of EAV.128, 129 Rotaviral diarrhea Equine rotavirus, a non-enveloped RNA virus, is one of the most important causes of infectious diarrhea in foals during the first few weeks of life and often causes outbreaks involving the majority of the foal crop on individual farms.130–132 The genus Rotavirus is one of five genera of the family Reoviridae and is divided into seven serogroups (A through G) based on differences in the inner capsid protein, VP6.132, 133 All equine rotavirus isolates to date are in group A, which is further subdivided into P (proteasesensitive, VP4-positive) and G (glycoprotein, VP7positive) serotypes, respectively.133 Although five P serotypes (P1, P6, P7, P12, and P18) and eight G serotypes (G1, G3, G5, G8, G10, G13, G14, G16) have been identified in horses,134–136 most equine rotavirus isolates from all parts of the world are of the G3 and P12 serotype and include two subtypes (A and B).137 An inactivated rotavirus A vaccine (Equine Rotavirus Vaccine, Pfizer Animal Health) containing the P12, G3 serotype (H2 strain) in a metabolizable oilin-water emulsion is conditionally licensed in the USA and is indicated for administration to pregnant mares in endemic areas as an aid to prevention of diarrhea in their foals caused by infection with rotaviruses of serogroup A. Foal vaccination is not indicated because there are no data to suggest that vaccination of the newborn foal with inactivated rotavirus A vaccine has any benefit in preventing or reducing the severity of infection. Label recommendations call for a three-dose series of the vaccine to be administered to mares during each pregnancy at 8, 9, and
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Vaccination of Mares, Foals and Weanlings 10 months of gestation. This protocol has been shown to induce significant increases in serum concentrations of neutralizing antibody in vaccinated mares and in the concentrations of antibodies of the IgG, but not IgA, subclass in the colostrum and milk of vaccinated mares.138, 139 After nursing, the concentration of passively derived rotavirus-specific antibody of the IgG subclass in the serum of foals up to 90 days of age from vaccinated mares is significantly higher than that measured in serum of foals born to non-vaccinated mares.138, 139 A field study showed circumstantial evidence of at least partial efficacy of this vaccine. An approximately two-fold higher incidence of rotaviral diarrhea was found in foals from non-vaccinated mares compared to those from vaccinated mares, although this difference did not prove to be statistically significant.138 Similarly, a controlled field study in Argentina in which an inactivated aluminum hydroxide-adjuvanted vaccine containing the SA11 (G3P2), H2 (G3P12), and Lincoln (G6P1) strains was administered to 100 mares at 60 days before foaling and again 30 days later, demonstrated a substantial reduction in the incidence and severity of rotaviral disease in foals from vaccinated mares compared with foals from nonvaccinated mares.140 As MDA titers wane at approximately 60 days of age, foals may develop rotaviral diarrhea. However, the severity of diarrhea is generally milder and of shorter duration than occurs in foals that become infected during the first 30 days of life. Challenge studies involving two inactivated rotavirus vaccines administered in a similar manner to pregnant mares in Japan showed that their foals were not completely protected against infection but had a substantial reduction in severity of clinical signs after challenge.134 The major correlate for protection against rotaviral infection appears to be mucosal immunity, predominantly mucosal IgA, in the gastrointestinal tract. Studies of the immunoglobulin isotype responses of mares after parenteral vaccination with inactivated rotavirus vaccines, and of antibodies passively transferred to their foals via colostrum, indicate that this approach is unlikely to provide foals with intestinal mucosal protection in the form of IgA.139 Consequently, it is not surprising that current protocols do not provide complete protection. In addition, because the conditionally licensed vaccine available in the USA contains only the G3 serotype of the A serogroup, it cannot be expected to protect against infection with all field strains. The potential for serological responses to primary vaccination with rotavirus vaccine to be suppressed during pregnancy should be investigated to determine whether completion of the primary series before breeding might be advantageous.
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References 1. Timoney PJ, McCollum WH. Equine viral arteritis. Vet Clin North Am Equine Pract 1993;9:295–309. 2. Sheoran AS, Timoney JF, Holmes MA, Karzenski SS, Crisman MV. Immunoglobulin isotypes in sera and nasal mucosal secretions and their neonatal transfer and distribution in horses. Am J Vet Res 2000;61:1099–105. 3. Ferguson JA, Reeves WC, Hardy JL. Studies on immunity to alphaviruses in foals. Am J Vet Res 1979;40:5–10. 4. Gibbs EPJ, Wilson JH, All BP, III. Studies on passive immunity and the vaccination of foals against eastern equine encephalitis in Florida. In: Equine Infectious Diseases V: Proceedings of the Fifth International Conference on Equine Infectious Diseases, Lexington, KY. University Press of Kentucky, 1988; pp. 201–5. 5. van Oirschot JT, Bruin G, de Boer-Luytze E, Smolders G. Maternal antibodies against equine influenza virus in foals and their interference with vaccination. Zentralbl Veterinarmed B 1991;38:391–6. 6. van Maanen C, Bruin G, de Boer-Luijtze E, Smolders G, de Boer GF. Interference of maternal antibodies with the immune response of foals after vaccination against equine influenza. Vet Q 1992;14:13–7. 7. van Maanen C, Flore PH, Minke J, Bruin G. Immune response of foals after vaccination against EHV-1/EHV-4 and persistence of maternal antibodies. In: Equine Infectious Diseases VII: Proceeedings of the Seventh International Conference on Equine Infectious Diseases, Tokyo, Japan. R & W Publications (Newmarket) Limited, 1994; pp. 351–2. 8. Hullinger PJ, Wilson WD, Rossitto PV, Patton JF, Thurmond MC, MacLachlan NJ. Passive transfer, rate of decay, and protein specificity of antibodies against equine arteritis virus in horses from a Standardbred herd with high seroprevalence. J Am Vet Med Assoc 1998;213: 839–42. 9. Wilson WD, Mihalyi JE, Hussey S, Lunn DP. Passive transfer of maternal immunoglobulin isotype antibodies against tetanus and influenza and their effect on the response of foals to vaccination. Equine Vet J 2001;33: 644–50. 10. Cullinane A, Weld J, Nelly M, McBride C. The interference of maternal antibodies with the immune response of Thoroughbred foals and yearlings to vaccination against equine influenza. In: Proceedings of the Seventh International Conference on Equine Infectious Diseases, Tokyo, Japan. 1994; p. 52. 11. Conboy HS, Berry DB, Fallon EH, Holland RE, Powell DG, Chambers TM. Failure of foal seroconversion following equine influenza vaccination. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, Phoenix, AZ, 1997; pp. 22–3. 12. Holland RE, Conboy HS, Berry DB, Fallon EH, Powell DG, Tudor LR, Chambers TM. Age dependence on foal vaccination for equine influenza: new evidence from the USA. In: Wernery U, Wade JF, Mumford JA, Kaaden OR, editors. Equine Infectious Diseases VIII: Proceedings of the Eighth International Conference, Dubai, UAE, 23–26 March 1998: R & W Publications (Newmarket) Ltd, 1999; pp. 547–8. 13. Cullinane A, Weld J, Osborne M, Nelly M, McBride C, Walsh C. Field studies on equine influenza vaccination regimes in thoroughbred foals and yearlings. Vet J 2001;161:174–85.
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14. Holznagel DL, Hussey S, Mihalyi JE, Wilson WD, Lunn DP. Onset of immunoglobulin production in foals. Equine Vet J 2003;35:620–2. 15. Burki F, Nowotny N, Rossmanith W, Pallan C, Mostl K. Training of the immune system of foals against ERP virus infections by frequent vaccination with presently available commercial vaccines. DTW Dtsch Tierarztl Wochenschr 1989;96:162–5. 16. Breathnach CC, Allen GP, Holland RE, Conboy HS, Berry DB, Jr., Chambers TM. Problems associated with vaccination of foals against Equine herpesvirus-4 and the role of anti-EHV-4 maternal antibodies. In: Wernery U, Wade JF, Mumford JA, Kaaden OR, editors. Equine Infectious Diseases VIII: Proceedings of the Eighth International Conference, Dubai, UAE, 23–26 March 1998: R & W Publications (Newmarket) Ltd, 1999; pp. 426–7. 17. Wilson WD. Vaccination programs for foals and weanlings. In: Proceedings of the 45th Annual Convention of the American Association of Equine Practitioners, 1999 pp. 254–63. 18. Wilson WD, Rossdale PD. Effect of age on the serological responses of Thoroughbred foals to vaccination with an inactivated EHV-1/EHV-4 vaccine. In: Wernery U, Wade JF, Mumford JA, Kaaden OR, editors. Equine Infectious Diseases VIII: Proceedings of the Eighth International Conference, Dubai, UAE, 23–26 March 1998: R & W Publications (Newmarket) Ltd, 1999; p. 428. 19. Heldens JG, Kersten AJ, Weststrate MW, van den Hoven R. Duration of immunity induced by an adjuvanted and inactivated equine influenza, tetanus and equine herpesvirus 1 and 4 combination vaccine. Vet Q 2001;23:210–7. 20. Wilson WD. Strategies for vaccinating mares, foals, and weanlings. Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners. Seattle, WA, 2005; pp. 421–38. 21. Chiang YW, Jennen CM, Holt TM, Waldbillig CP, Hathaway DK, Jennings NJ, Ng T, Chu HJ. Demonstration of the efficacy of a West Nile virus DNA vaccine in foals. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners, Seattle, WA, 2005; pp. 183–90. 22. Gutierrez CV, Brown SE, Long mMT, Horohov DW. Evaluation of immune responses in foals post-vaccination with a West Nile virus vaccine (live Flavivirus chimera). In: Proceedings of the 54th Annual Convention of the American Association of Equine Practitioners, San Diego, 2008; pp. 511–12. 23. Lohrer J, Radvila P. Active tetanus protection in the horses and the duration of immunity. Schwei Arch Tierheikd 1970;112:307–14. 24. Jansen BC, Knoetze PC. The immune response of horses to tetanus toxoid. Onderstepoort J Vet Res 1979;46:211–16. 25. Hays MB. Definitive efficacy and safety testing for equine encephalomyelitis vaccine. J Am Vet Med Assoc 1969;155:374–6. 26. Barber TL, Walton TE, Lewis KJ. Efficacy of trivalent inactivated encephalomyelitis virus vaccine in horses. Am J Vet Res 1978;39:621–5. 27. Holmes MA, Townsend HG, Kohler AK, Hussey S, Breathnach C, Barnett C, Holland R, Lunn DP. Immune responses to commercial equine vaccines against equine herpesvirus-1, equine influenza virus, eastern equine encephalomyelitis, and tetanus. Vet Immunol Immunopathol 2006;111:67–80.
28. Eisner RJ, Nusbaum SR. A study to determine the optimum time for vaccination of foals against eastern and western encephalitis viruses. In: Proceedings of the American Association of Veterinary Laboratory Diagnosticians, 1979; 29. Liu IKM. Duration of maternally derived antibodies in neonatal foals. Mod Vet Pract 1986;67:454–6. 30. Wilson JH, Gibbs PJ, Calisher CH, Buergelt CD, Schneider C. Investigation of vaccine-induced tolerance to Eastern equine encephalitis virus in foals. In: Proceedings of the 41st Annual Conference of the American Association of Equine Practitioners, Lexington, KY, 1995; pp. 178–80. 31. Wilson JH, Rubin HL, Lane TJ, Gibbs EPJ. A survey of eastern equine encephalomyelitis in Florida horses: prevalence, economic impact, and management practices, 1982–1983. Prev Vet Med 1986;4:261–71. 32. Ng T, Hathaway D, Jennings N, Champ D, Chiang YW, Chu HJ. Equine vaccine for West Nile virus. Dev Biol (Basel) 2003;114:221–7. 33. Minke JM, Siger L, Karaca K, Austgen L, Gordy P, Bowen R, Renshaw RW, Loosmore S, Audonnet JC, Nordgren B. Recombinant canarypoxvirus vaccine carrying the prM/E genes of West Nile virus protects horses against a West Nile virus-mosquito challenge. Arch Virol Suppl 2004;( 18): 221–30. 34. Siger L, Bowen R, Karaca K, Murray M, Jagannatha S, Echols B, Nordgren R, Minke JM. Assessment of the efficacy of a single dose of a recombinant vaccine against West Nile virus in response to natural challenge with West Nile virus-infected mosquitoes in horses. Am J Vet Res 2004;65:1459–62. 35. Siger L, Bowen RA, Karaca K, Murray MJ, Gordy PW, Loosmore SM, Audonnet JC, Nordgren RM, Minke JM. Evaluation of the efficacy provided by a Recombinant Canarypox-Vectored Equine West Nile Virus vaccine against an experimental West Nile Virus intrathecal challenge in horses. Vet Ther 2006;7:249–56. 36. Arroyo J, Miller C, Catalan J, Myers GA, Ratterree MS, Trent DW, Monath TP. ChimeriVax-West Nile virus live-attenuated vaccine: preclinical evaluation of safety, immunogenicity, and efficacy. J Virol 2004;78: 12497–507. 37. Long MT, Gibbs EP, Seino KK, Mellencamp MW, Zhang S, Beachboard SE, Humphrey PP. Safety and efficacy of a live attenuated West Nile virus chimera vaccine in horses with experimentally induced West Nile virus clinical disease. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners, Seattle, WA, 2005; pp. 177–79. 38. Long MT, Gibbs EP, Mellencamp MW, Bowen RA, Seino KK, Zhang S, Beachboard SE, Humphrey PP. Efficacy, duration, and onset of immunogenicity of a West Nile virus vaccine, live Flavivirus chimera, in horses with a clinical disease challenge model. Equine Vet J 2007;39:491–7. 39. Vest DJ, Cohen ND, Berezowski CJ, Morehead JP, Blodgett GP, Blanchard TL. Evaluation of administration of West Nile virus vaccine to pregnant broodmares. J Am Vet Med Assoc 2004;225:1894–7. 40. Long MT, Gibbs EP, Mellencamp MW, Zhang S, Barnett DC, Seino KK, Beachboard SE, Humphrey PP. Safety of an attenuated West Nile virus vaccine, live Flavivirus chimera in horses. Equine Vet J 2007;39:486–90. 41. Hannant D, Mumford JA, Jessett DM. Duration of circulating antibody and immunity following infection with equine influenza virus. Vet Rec 1988;122:125–8.
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Vaccination of Mares, Foals and Weanlings 42. Hannant D, Jessett DM, O’Neill T, Mumford JA. Antibody isotype responses in the serum and respiratory tract to primary and secondary infections with equine influenza virus (H3N8). Vet Microbiol 1989;19:293– 303. 43. Hannant D, Mumford JA. Cell mediated immune responses in ponies following infection with equine influenza virus (H3N8): the influence of induction culture conditions on the properties of cytotoxic effector cells. Vet Immunol Immunopathol 1989;21:327–37. 44. Nelson KM, Schram BR, McGregor MW, Sheoran AS, Olsen CW, Lunn DP. Local and systemic isotype-specific antibody responses to equine influenza virus infection versus conventional vaccination. Vaccine 1998;16:1306–13. 45. van Maanen C, Cullinane A. Equine influenza virus infections: an update. Vet Q 2002;24:79–94. 46. Hannant D, Jessett DM, O’-Neill T, Livesay J, Mumford JA. Cellular immune responses stimulated by inactivated virus vaccines and infection with equine influenza virus (H3N8). In: Equine Infectious Diseases VII: Proceedings of the Seventh International Conference, Tokyo, Japan. June 8–11 1994: R & W Publications (Newmarket) Ltd, 1994; pp. 169–74. 47. Daly JM, Newton JR, Mumford JA. Current perspectives on control of equine influenza. Vet Res 2004;35:411–23. 48. Crouch CF, Daly J, Hannant D, Wilkins J, Francis MJ. Immune responses and protective efficacy in ponies immunised with an equine influenza ISCOM vaccine containing an ‘American lineage’ H3N8 virus. Vaccine 2004;23:418–25. 49. Wood JM, Mumford J, Folkers C, Scott AM, Schild GC. Studies with inactivated equine influenza vaccine. 1. Serological responses of ponies to graded doses of vaccine. J Hyg (Lond) 1983;90:371–84. 50. Mumford JA, Wood J. Establishing an acceptability threshold for equine influenza vaccines. Dev Biol Stand 1992;79:137–46. 51. Townsend HGG, Morley PS, Newton JR, Wood JLN, Haines DM, Mumford JA. Measuring serum antibody as a method of predicting infection and disease in horses during outbreaks of influenza. In: Wernery U, Wade JF, Mumford JA, Kaaden OR, editors. Equine Infectious Diseases VIII: Proceedings of the Eighth International Conference, Dubai, UAE, 23–26 March 1998: R & W Publications (Newmarket Ltd), 1999; pp. 33–7. 52. Morley PS, Townsend HG, Bogdan JR, Haines DM. Risk factors for disease associated with influenza virus infections during three epidemics in horses. J Am Vet Med Assoc 2000;216:545–50. 53. Wilson WD. Equine influenza. Vet Clin N Am Equine Pract 1993;9:257–82. 54. Chambers TM, Holland RE, Tudor LR, Townsend HG, Cook A, Bogdan J, Lunn DP, Hussey S, WhitakerDowling P, Youngner JS, Sebring RW, Penner SJ, Stiegler GL. A new modified live equine influenza virus vaccine: phenotypic stability, restricted spread and efficacy against heterologous virus challenge. Equine Vet J 2001; 33:630–6. 55. Townsend HG, Penner SJ, Watts TC, Cook A, Bogdan J, Haines DM, Griffin S, Chambers T, Holland RE, Whitaker-Dowling P, Youngner JS, Sebring RW. Efficacy of a cold-adapted, intranasal, equine influenza vaccine: challenge trials. Equine Vet J 2001;33:637–43. 56. Townsend HGG. Current and new technologies for vaccines and vaccination decisions. In: Proceedings of the
57.
58.
59.
60.
61.
62.
63. 64.
65.
66.
67.
68.
69.
70.
71.
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51st Annual Convention of the American Association of Equine Practitioners, Seattle, WA, 2005; pp. 446–50. Edlund Toulemonde C, Daly J, Sindle T, Guigal PM, Audonnet JC, Minke JM. Efficacy of a recombinant equine influenza vaccine against challenge with an American lineage H3N8 influenza virus responsible for the 2003 outbreak in the United Kingdom. Vet Rec 2005;156:367–71. Minke JM, Toulemonde CE, Coupier H, Guigal PM, Dinic S, Sindle T, Jessett D, Black L, Bublot M, Pardo MC, Audonnet JC. Efficacy of a canarypox-vectored recombinant vaccine expressing the hemagglutinin gene of equine influenza H3N8 virus in the protection of ponies from viral challenge. Am J Vet Res 2007;68:213–9. Minke JM, Edlund Toulemonde C, Dinic S, Cozette V, Cullinane A, Audonnet JC. Effective priming of foals born to immune dams against influenza by a canarypoxvectored recombinant influenza H3N8 vaccine. J Comp Path 2007;137:S76–S80. Newton JR, Verheyen K, Wood JL, Yates PJ, Mumford JA. Equine influenza in the United Kingdom in 1998. Vet Rec 1999;145:449–52. Newton JR, Lakhani KH, Wood JL, Baker DJ. Risk factors for equine influenza serum antibody titres in young thoroughbred racehorses given an inactivated vaccine. Prev Vet Med 2000;46:129–41. Newton JR, Townsend HG, Wood JL, Sinclair R, Hannant D, Mumford JA. Immunity to equine influenza: relationship of vaccine-induced antibody in young Thoroughbred racehorses to protection against field infection with influenza A/equine-2 viruses (H3N8). Equine Vet J 2000;32:65–74. Smith BP. Influenza in foals. J Am Vet Med Assoc 1979;174:289–90. Liu IK, Pascoe DR, Chang LW, Zee YC. Duration of maternally derived antibodies against equine influenza in newborn foals. Am J Vet Res 1985;46:2078–80. Wilson WD, Robinson D. Field safety of a modifiedlive, cold-adapted intranasal equine influenza vaccine (HeskaTM Flu AvertTM I.N. vaccine) in horses. J Equine Vet Sci 2000;20:8–10. Wilson WD, Chambers TM, Holland RE, Lunn DP, TraubDargatz J, Townsend HG. Intranasal vaccine for equine influenza: Roundtable discussion, part 3. Equine Practice 2000;22:18–24. Slater J. Equine herpesviruses. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 134–53. Foote CE, Gilkerson JR, Whalley JM, Love DN. Seroprevalence of equine herpesvirus 1 in mares and foals on a large Hunter Valley stud farm in years pre- and postvaccination. Aust Vet J 2003;81:283–8. Foote CE, Love DN, Gilkerson JR, Whalley JM. Detection of EHV-1 and EHV-4 DNA in unweaned Thoroughbred foals from vaccinated mares on a large stud farm. Equine Vet J 2004;36:341–5. Foote CE, Love DN, Gilkerson JR, Wellington JE, Whalley JM. EHV-1 and EHV-4 infection in vaccinated mares and their foals. Vet Immunol Immunopathol 2006;111: 41–6. Pusterla N, Leutenegger CM, Wilson WD, Watson JL, Ferraro GL, Madigan JE. Equine herpesvirus-4 kinetics in peripheral blood leukocytes and nasopharyngeal secretions in foals using quantitative real-time TaqMan PCR. J Vet Diagn Invest 2005;17:578–81.
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72. Allen GP. Epidemic disease caused by equine herpesvirus-1: recommendations for prevention and control. Equine Vet Educ 2002;14:136–42. 73. Hannant D. Respiratory viral diseases of horses: overview. In: Proceedings of the World Equine Airway Symposium, Guelph, Ontario, Canada. Lifelearn Inc., 1998 on CD ROM. 74. Patel JR, Bateman H, Williams J, Didlick S. Derivation and characterisation of a live equid herpes virus-1 (EHV-1) vaccine to protect against abortion and respiratory disease due to EHV-1. Vet Microbiol 2003;91:23–39. 75. Heldens JG, Hannant D, Cullinane AA, Prendergast MJ, Mumford JA, Nelly M, Kydd JH, Weststrate MW, van den Hoven R. Clinical and virological evaluation of the efficacy of an inactivated EHV1 and EHV4 whole virus vaccine (Duvaxyn EHV1,4). Vaccination/challenge experiments in foals and pregnant mares. Vaccine 2001;19:4307–17. 76. Moore BO, Koonse HJ. Inactivated equine herpesvirus R . In: Proceedings of the 1 vaccine – Pneumabort-K 24th Annual Convention of the American Association of Equine Practitioners, St Louis, MO, 1978; pp. 75–9. 77. Burrows R, Goodridge D, Denyer MS. Trials of an inactivated equid herpesvirus 1 vaccine: challenge with a subtype 1 virus. Vet Rec 1984;114:369–74. 78. Goodman LB, Wagner B, Flaminio MJ, Sussman KH, Metzger SM, Holland R, Osterrieder N. Comparison of the efficacy of inactivated combination and modifiedlive virus vaccines against challenge infection with neuropathogenic equine herpesvirus type 1 (EHV-1). Vaccine 2006;24:3636–45. 79. Ostlund EN. The equine herpesviruses. Vet Clin N Am Equine Pract 1993;9:283–94. 80. Allen GP, Yeargan MR, Turtinen LW, Bryans JT. A new field strain of equine abortion virus (equine herpesvirus1) among Kentucky horses. Am J Vet Res 1985;46: 138–40. 81. Bass EP. Immunization with a modified live virus equine rhinopneumonitis vaccine and an aluminum hydroxide adsorbed equine influenza vaccine. In: Proceedings of the 24th Annual Convention of the American Association of Equine Practitioners, St Louis, MO, 1978; pp. 65–74. 82. Smith PM, Kahan SM, Rorex CB, von Einem J, Osterrieder N, O’Callaghan DJ. Expression of the full-length form of gp2 of equine herpesvirus 1 (EHV-1) completely restores respiratory virulence to the attenuated EHV-1 strain KyA in CBA mice. J Virol 2005;79:5105–15. 83. Kendrick JW, Stevenson W. Immunity to equine herpesvirus 1 infection in foals during the first year of life. J Reprod Fertil Suppl 1979;(27):615–8. 84. Dutta SK, Shipley WD. Immunity and the level of neutralization antibodies in foals and mares vaccinated with a modified live-virus rhinopneumonitis vaccine. Am J Vet Res 1975;36:445–8. 85. Neely DP, Hawkins DL. A two-year study of the clinical and serologic responses of horses to a modified livevirus equine rhinopneumonitis vaccine. J Equine Med Surg 1978;2:532–40 86. Ellis JA, Steeves E, Wright AK, Bogdan JR, Davis WC, Kanara EW, Haines DM. Cell-mediated cytolysis of equine herpesvirus-infected cells by leukocytes from young vaccinated horses. Vet Immunol Immunopathol 1997;57:201–14. 87. Foote CE, Love DN, Gilkerson JR, Whalley JM. Serological responses of mares and weanlings following vaccination with an inactivated whole virus equine her-
88.
89.
90.
91.
92.
93. 94.
95.
96.
97.
98.
99.
100.
101.
102.
103.
pesvirus 1 and equine herpesvirus 4 vaccine. Vet Microbiol 2002;88:13–25. Gilkerson JR, Whalley JM, Drummer HE, Studdert MJ, Love DN. Epidemiology of EHV-1 and EHV-4 in the mare and foal populations on a Hunter Valley stud farm: are mares the source of EHV-1 for unweaned foals. Vet Microbiol 1999;68:27–34. Townsend HGG. The role of vaccines and their efficacy in the control of infectious respiratory disease of the horse. In: Proceedings of the 46th Annual Convention of the American Association of Equine Practitioners, San Antonio, Texas, 2000; pp. 21–26. Hamlen HJ, Timoney JF, Bell RJ. Epidemiologic and immunologic characteristics of Streptococcus equi infection in foals. J Am Vet Med Assoc 1994;204:768–75. Sellon DC. Streptococcal infections. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 244–57. Timoney JF, Qin A, Muthupalani S, Artiushin S. Vaccine potential of novel surface exposed and secreted proteins of Streptococcus equi. Vaccine 2007;25: 5583–90. Timoney JF. Strangles. Res Vet Sci 1993;54:256–8. Sheoran AS, Sponseller BT, Holmes MA, Timoney JF. Serum and mucosal antibody isotype responses to Mlike protein (SeM) of Streptococcus equi in convalescent and vaccinated horses. Vet Immunol Immunopathol 1997;59:239–51. Flock M, Jacobsson K, Frykberg L, Hirst TR, Franklin A, Guss B, Flock JI. Recombinant Streptococcus equi proteins protect mice in challenge experiments and induce immune response in horses. Infect Immun 2004;72:3228–36. Walker JA, Timoney JF. Construction of a stable nonmucoid deletion mutant of the Streptococcus equi Pinnacle vaccine strain. Vet Microbiol 2002;89:311–21. Hoffman AM, Staempfli HR, Prescott JF, Viel L. Field evaluation of a commercial M-protein vaccine against Streptococcus equi infection in foals. Am J Vet Res 1991;52:589–92. Rief JS, George JL, Shideler RK. Recent developments in strangles research: observations on the carrier state and evaluation of a new vaccine. In: Proceedings of the 27th Annual Convention of the American Association of Equine Practitioners, 1981; pp. 33–40. Staempfli HR, Hoffman AM, Prescott JF, Viel L. Clinical evaluation of a commercial M-protein vaccine in naturally infected foals. In: Proceedings of the 37th Annual Convention of the American Association of Equine Practitioners, 1991; pp. 259–262. Li W, Fiala S, Gigson N, Timoney J, Cairns R, Acree WM, Chu H-J. Efficacy of a modified live Streptococcus equi vaccine in an equine experimental model.In: Wernery U, Wade JF, Mumford JA, Kaaden OR, editors. Equine Infectious Diseases VIII: Proceedings of the Eighth International Conference, Dubai, UAE, 23–26 March 1998: R & W Publications (Newmarket) Ltd, 1999; pp. 557–8. Sweeney CR, Timoney JF, Newton JR, Hines MT. Streptococcus equi infections in horses: guidelines for treatment, control, and prevention of strangles. J Vet Intern Med 2005;19:123–34. Galan JE, Timoney JF, Lengemann FW. Passive transfer of mucosal antibody to Streptococcus equi in the foal. Infect Immun 1986;54:202–6. Madigan JE, Pusterla N. Ehrlichial diseases. Vet Clin North Am Equine Pract 2000;16:487–99.
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Vaccination of Mares, Foals and Weanlings 104. Atwill ER, Mohammed HO. Benefit-cost analysis of vaccination of horses as a strategy to control equine monocytic ehrlichiosis. J Am Vet Med Assoc 1996;208:1295–9. 105. Atwill ER, Mohammed HO. Evaluation of vaccination of horses as a strategy to control equine monocytic ehrlichiosis. J Am Vet Med Assoc 1996;208:1290–4. 106. Palmer JE. Potomac horse fever. Vet Clin N Am Equine Pract 1993;9:399–410. 107. Dutta SK, Vemulapalli R, Biswas B. Association of deficiency in antibody response to vaccine and heterogeneity of Ehrlichia risticii strains with Potomac horse fever vaccine failure in horses. J Clin Microbiol 1998;36: 506–12. 108. Sessions J, Dawson JE. Maryland field evaluation of the Potomac horse fever vaccine. Equine Pract 1988;10:7–12. 109. Wilkins PA. Botulism. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 372–6. 110. Swerczek TW. Toxicoinfectious botulism in foals and adult horses. J Am Vet Med Assoc 1980;176:217–20. 111. Whitlock RH, Buckley C. Botulism. Vet Clin North Am Equine Pract 1997;13:107–28. 112. Wilkins PA, Palmer JE. Botulism in foals less than 6 months of age: 30 cases (1989–2002). J Vet Intern Med 2003;17:702–7. 113. Thomas RJ, Rosenthal DV, Rogers RJ. A Clostridium botulinum type B vaccine for prevention of shaker foal syndrome. Aust Vet J 1988;65:78–80. 114. Crane SA, Whitlock RH, Buckley C, Forney B, Blake Caddel L. Clostridium botulinum type-B toxoid for vaccination of adult horses, pregnant mares, and foals: a study of vaccination protocols. In: Proceedings of the 37th Annual Convention of the American Association of Equine Practitioners, San Francisco, CA, 1991; p. 611. 115. McCue PM, Hietala SK, Spensley MS, Stillian M, Mihalyi J, Hughes J. Prevalence of equine viral arteritis in California horses. Calif Vet 1991;45:24–6. 116. Timoney PJ, McCollum WH, Murphy TW, Roberts AW, Willard JG, Carswell GD. The carrier state in equine arteritis virus infection in the stallion with specific emphasis on the venereal mode of virus transmission. J Reprod Fertil Suppl 1987;35:95–102. 117. McCollum WH. Development of a modified virus strain and vaccine for equine viral arteritis. J Am Vet Med Assoc 1969;155:318–22. 118. Balasuriya UBR, MacLachlan NJ. Equine viral arteritis. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 153–64. 119. Harry TO, McCollum WH. Stability of viability and immunizing potency of lyophilized, modified equine arteritis live-virus vaccine. Am J Vet Res 1981;42:501–5. 120. McCollum WH. Pathologic features of horses given avirulent equine arteritis virus intramuscularly. Am J Vet Res 1981;42:1218–20. 121. McCollum WH, Timoney PJ, Roberts AW, Willard JE, Carswell GD. Response of vaccinated and nonvaccinated mares to artificial insemination with semen from stallions persistently infected with equine arteritis virus. In: Powell DG (editor). Equine Infectious Diseases V: Proceedings of the Fifth International Conference. Lexington, KY, USA: University Press of Kentucky, 1988; pp. 13–18. 122. Moore BD, Balasuriya UB, Nurton JP, McCollum WH, Timoney PJ, Guthrie AJ, MacLachlan NJ. Differentiation of strains of equine arteritis virus of differing virulence to
123. 124.
125.
126.
127.
128.
129.
130.
131.
132. 133.
134.
135.
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horses by growth in equine endothelial cells. Am J Vet Res 2003;64:779–84. McCollum WH. Studies of passive immunity in foals to equine viral arteritis. Vet Microbiol 1976;1:45–54. Nugent J, Sinclair R, deVries AA, Eberhardt RY, CastilloOlivares J, Davis Poynter N, Rottier PJ, Mumford JA. Development and evaluation of ELISA procedures to detect antibodies against the major envelope protein (G(L)) of equine arteritis virus. J Virol Methods 2000;90: 167–83. Castillo-Olivares J, de Vries AA, Raamsman MJ, Rottier PJ, Lakhani K, Westcott D, Tearle JP, Wood JL, Mumford JA, Hannant D, Davis-Poynter NJ. Evaluation of a prototype sub-unit vaccine against equine arteritis virus comprising the entire ectodomain of the virus large envelope glycoprotein (G(L)): induction of virus-neutralizing antibody and assessment of protection in ponies. J Gen Virol 2001;82:2425–35. Tobiasch E, Kehm R, Bahr U, Tidona CA, Jakob NJ, Handermann M, Darai G, Giese M. Large envelope glycoprotein and nucleocapsid protein of equine arteritis virus (EAV) induce an immune response in Balb/c mice by DNA vaccination; strategy for developing a DNA-vaccine against EAV-infection. Virus Genes 2001;22:187–99. Giese M, Bahr U, Jakob NJ, Kehm R, Handermann M, Muller H, Vahlenkamp TH, Spiess C, Schneider TH, Schusse G, Darai G. Stable and long-lasting immune response in horses after DNA vaccination against equine arteritis virus. Virus Genes 2002;25:159–67. Balasuriya UB, Heidner HW, Hedges JF, Williams JC, Davis NL, Johnston RE, MacLachlan NJ. Expression of the two major envelope proteins of equine arteritis virus as a heterodimer is necessary for induction of neutralizing antibodies in mice immunized with recombinant Venezuelan equine encephalitis virus replicon particles. J Virol 2000;74:10623–30. Balasuriya UB, Heidner HW, Davis NL, Wagner HM, Hullinger PJ, Hedges JF, Williams JC, Johnston RE, David Wilson W, Liu IK, James MacLachlan N. Alphavirus replicon particles expressing the two major envelope proteins of equine arteritis virus induce high level protection against challenge with virulent virus in vaccinated horses. Vaccine 2002;20:1609–17. Browning GF, Chalmers RM, Snodgrass DR, Batt RM, Hart CA, Ormarod SE, Leadon D, Stoneham SJ, Rossdale PD. The prevalence of enteric pathogens in diarrhoeic Thoroughbred foals in Britain and Ireland. Equine Vet J 1991;23:405–9. Browning GF, Sykes JE, Huntington PJ, Hollywell CA, Begg AP. Rotavirus infections in Australian foals. Aust Equine Veterinarian 1992;10:123–6. Dwyer RM. Rotaviral diarrhea. Vet Clin N Am Equine Pract 1993;9:311–19. Dwyer RM. Equine rotavirus. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 181–3. Imagawa H, Wada R, Sugita S, Fukunaga Y. Passive immunity in foals of mares immunised with inactivated equine rotavirus vaccine. In: Equine Infectious Diseases VIII: Proceedings of the Eighth International Conference on Equine Infectious Diseases, Dubai, UAE, 1999. R & W Publications (Newmarket) Ltd, 1999. Elschner M, Schrader C, Hotzel H, Prudlo J, Sachse K, Eichhorn W, Herbst W, Otto P. Isolation and molecular
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characterisation of equine rotaviruses from Germany. Vet Microbiol 2005;105:123–9. 136. Gulati BR, Deepa R, Singh BK, Rao CD. Diversity in Indian equine rotaviruses: identification of genotype G10,P6[1] and G1 strains and a new VP7 genotype (G16) strain in specimens from diarrheic foals in India. J Clin Microbiol 2007;45:972–8. 137. Browning GF, Begg AP. Prevalence of G and P serotypes among equine rotaviruses in the faeces of diarrhoeic foals. Arch Virol 1996;141:1077–89. 138. Powell DG, Dwyer RM, Traub-Dargatz JL, Fulker RH, Whalen JW, Jr., Srinivasappa J, Acree WM, Chu HJ. Field
study of the safety, immunogenicity, and efficacy of an inactivated equine rotavirus vaccine. J Am Vet Med Assoc 1997;211:193–8. 139. Sheoran AS, Karzenski SS, Whalen JW, Crisman MV, Powell DG, Timoney JF. Prepartum equine rotavirus vaccination inducing strong specific IgG in mammary secretions. Vet Rec 2000;146:672–3. 140. Barrandeguy M, Parreno V, Lagos Marmol M, Pont Lezica F, Rivas C, Valle C, Fernandez F. Prevention of rotavirus diarrhoea in foals by parenteral vaccination of the mares: field trial. Dev Biol Stand 1998;92:253–7.
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Development of the Foal Immune System M. Julia B. Felippe Flaminio and Rebecca L. Tallmadge
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Colostrum: Assessment of Quality and Artificial Supplementation Guy D. Lester
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Assessment and Modification of Passive Transfer Sally L. Vivrette
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Neonatal Isoerythrolysis Jill R. Johnson
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Immunodeficiencies in Foals M. Julia B. Felippe Flaminio
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Severe Combined Immunodeficiency Melissa T. Hines
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Thrombocytopenia and Selective IgM Deficiency Rodney L. Belgrave
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Chapter 32
Development of the Foal Immune System M. Julia B. Felippe Flaminio and Rebecca L. Tallmadge
Equine perinatal immunity Equids are under substantial selective pressure because the foal is born and immediately exposed to environmental pathogens before acquiring antibodies through colostrum. There is no transfer of maternal immunoglobulins during gestation, and nursing requires a foal that is strong and alert immediately after birth. Fortunately, the multifactorial process of adaptation from the uterine to the outside environment follows the miracle of life. More often than not, foals thrive, which is the ultimate indicator of a functional immune system. Through colostrum, passively transferred maternal immunoglobulins, opsonins, and inflammatory cytokines support the critical immune responses that are about to be developed. Foals commonly acquire infection and develop septicemia within hours to days after birth via the respiratory, oral, and umbilical routes. Failure of transfer of immunoglobulins through the colostrum increases the foal’s susceptibility to infection by environmental pathogens. The measurement of immunoglobulin (Ig) G concentration in the foal’s blood after nursing colostrum determines the need for clinical intervention with administration of immunoglobulins and other prophylactic measures. When infection occurs in the uterine environment, the foal is born with established infectious and inflammatory processes, and perhaps septicemia, which is independent of a successful, or failure of, transfer of immunoglobulins through colostrum. Immunoglobulins constitute an important fraction of the arsenal of the immune system. However, it is the combination of immunological strategies that provide the best protection against pathogens (Figure 32.1). Under physiological conditions, the immune system of the foal at birth is competent, i.e., it
controls infection and prevents disease. J.B. Solomon once wrote, ‘. . . it is the antigenic inexperience of the newborn and not its immunological capacity which is deficient’. Although there is evidence that all elements of the immune system are present in the foal at birth, some of them are not fully functional until later in life, and may not be able to manage a large burden of infection. In addition, the foal’s susceptibility to certain pathogens, including Rhodococcus equi, exclusively in the first few months of life suggests specific and physiological age-dependent development of the immune system. The foal is born with an adaptive immune system capable of generating antigen-specific immune responses, although its naive condition requires time to establish the necessary lymphocyte population expansion and memory. Therefore, the first 3 months of life pose a challenge to the foal in preparedness. In this context, environmental conditions (e.g., hygiene, pathogen load, overcrowded pastures, intense horse transit) can determine disease development in a normal foal, and it should also be the focus of attention in the prevention of diseases in early life. Some pathogens use elaborate methods of infection and replication in a still developing immune system, and, for those, strategic immunoprophylaxis of the foal preceding the period of susceptibility would be required. Alternatively, when antibody neutralization is efficient (e.g., rotavirus and botulism/tetanus toxins), maternally derived antibodies transferred through colostrum are effective and essential in protection of the neonate.
Innate immune system The innate immune system evolved to recognize and react to an array of specific pathogen structures
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Immune System
The Immune System
INNATE
phagocytes neutrophils macrophages dendritic cells opsonins complement serum amyloid A lectin lactoferrin LAK cells NK cells
antigen presenting cells
ADAPTIVE
T cells - cytokines CD4+ (Th1,Th2,Th17, Treg) CD8+ B cells - antibodies IgM IgG (IgG1-7) IgA IgE
Figure 32.1 The elements of the immune system. The innate immune system evolved to recognize pathogen-associated molecular patterns without previous encounter. Once activated, the innate immune system removes and processes pathogens or pathogen-infected cells. Importantly, it interacts with and signals the adaptive immune system to amplify the population of cells that recognize the invaders in a specific manner, including the development of memory. The combination of all elements of the immune system provides the best strategy to fight infections. LAK, lymphokine-activated killer; NK, natural killer; CD, cluster of differentiation; Ig, immunoglobulin.
(or pathogen-associated molecular patterns; PAMPs) without previous exposure or priming. The neonatal innate immune system plays an important role in protection while antigen-specific immune responses are mounting. Indeed, the innate immune system precedes, facilitates, and signals the function of the adaptive immune cells for recognition and removal of pathogens. Initially, the foal relies on its phagocytic function for removal and killing of most organisms in early life. Foals respond effectively with neutrophilia and a left shift when fighting an infection or sepsis, unless they present with prematurity or immaturity. Neutrophil function in healthy equine neonates, including phagocytosis and oxidative burst activity, is comparable to or greater than adult horses when pathogens are opsonized with pooled adult horse sera in vitro.1–3 Foal phagocytes express high levels of the CD18 receptor, which interacts with complement C3b-opsonized particles, and dimerizes with CD11 to form integrin (adhesion) molecules.4 The limiting factors in foal phagocytic function in vivo, however, are chemotactic and opsonic elements.5 This limitation may be more critical in foals with septicemia, because
they demonstrate a transitory decrease in phagocytosis and oxidative burst activity during the acute phase of disease.6 Phagocytic activity is largely facilitated by opsonins: when pathogens are bound to complement or immunoglobulins, phagocytosis is mediated by C3 or Fc receptors expressed on phagocytes, respectively. Overall opsonization capacity is much reduced in presuckle serum samples, and foals are transiently deficient in serum opsonic capacity, independent of their serum IgG concentrations.2, 3 Indeed, heat inactivation of equine serum (which preserves IgG but destroys heat-labile opsonins) has been shown to abrogate up to 90% of phagocytic function compared with normal serum. Opsonization of bacteria with foal serum yields much reduced phagocytosis compared with pooled adult horse sera. In addition, complement activity in presuckle foals has been reported to be 13% of the adult horse. Although colostrum ingestion promotes a positive effect on overall serum opsonization, colostrum is not an important source of complement to the equine neonate, and foals depend on their own production to fight infection.2, 7 Healthy foals produce complement at a rate of 0.24–0.34 mg/L serum/day in the first 10 days of life, increasing complement activity to 64% and 85% of that of the adult horse by 1 and 5 months of life, respectively.2, 6 Perhaps as a compensatory mechanism of protection, colostrum-deprived equine neonates have higher hemolytic complement activity and complement C3 concentrations within 2–5 days of life than foals receiving adequate amounts of colostrum.8 Colostrum-depleted non-sick foals also sustain a greater rate of complement and endogenous IgG production than colostrum-fed foals. Complement C3 components seem to be highly consumed and their levels are unstable in septic foals, which cannot maintain increasing age-dependent levels similarly to control foals.6 Intravenous plasma transfusion has been shown to provide and maintain opsonization capacity and phagocytic function in sick foals at levels comparable to healthy foals. In septic foals, plasma treatment does not always increase absolute IgG and complement C3 concentrations individually.9 It is possible that opsonization improvement results from the synergy of all opsonins present in plasma. Therefore, serum complement together with other heat-sensitive proteins plays a role in opsonization. When analyzing opsonization capacity against relevant pathogens to the equine neonate (Escherichia coli and Actinobacillus equuli), healthy foal serum has similar efficiency to adult horse sera. Complement activation is important for opsonization of the tested microbes, and there is no significant correlation with concentrations of IgG isotypes and serum opsonic capacity.2
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pathogen
TLR
infected cell
DC IL-12
CD4+ helper T cell
IFN
killing CD8+ cytotoxic T cell
IL-4 IL-5 IL-13
IFN
B cell
IgG1 IgG3/5 IgG4/7 IgA IgE
Figure 32.2 The interaction of dendritic cells (DCs), T-cells, and B-cells. An effective immune response requires synchrony of cell activation for appropriate cell signaling and differentiation. In the periphery, dendritic cells recognize pathogen-associated molecular patterns via toll-like receptors (TLR). Once DCs phagocytose and process the pathogens, they migrate to regional lymph nodes and present pathogen peptides to CD4+ (helper) T-cells via major histocompatibility (MHC) class II molecules. Upon recognition and binding of the T-cell receptor (TCR), co-stimulatory molecules between both cells potentiate signaling for CD4+ T-cell activation (priming). Depending on the TLR signaling in DCs, interleukin 12 (IL-12) may be secreted to promote further CD4+ T-cell activation for the generation of a Th1-type (cytotoxic) immune response. The CD4+ T-cells increase the efficiency of DC antigen presentation by secreting interferon (IFN)γ , which targets interferon regulatory factors in the DCs, and upregulates their expression of co-stimulatory molecules and cytokines. The CD8+ (cytotoxic) T-cells are activated by antigen presentation via MHC class I on DCs (cross-presentation), the co-stimulation from molecules expressed on activated CD4+ T-cells, and their secreted IFNγ. In the periphery, activated (primed) CD8+ T-cells search for infected cells that express processed pathogen peptides in the context of MHC class I molecules. CD8+ T-cells kill infected target cells using elements that alter cell permeability (e.g., perforins, granzymes). According to the DC stimulation, and in the absence of IL-12, CD4+ T-cells can generate a Th2-type (humoral) immune response. They control B-cell differentiation and antibody secretion via cytokines (IL-4, IL-5, IL-13) and co-stimulatory molecules. Depending on the cytokine milieu (including IFNγ), B-cells isotype switch into different immunoglobulins, and progress to plasma cells, which secrete immunoglobulins.
Another important opsonin is serum amyloid A (SAA), a family of hepatic acute-phase proteins that bind to high-density lipoproteins, induce chemotaxis and the upregulation of integrins in leukocytes, and promote phagocyte antimicrobial activities (phagocytosis, degranulation, and killing). Mammary-associated SAA3 has been measured in high concentrations in colostrum of mammals including the horse.10, 11 SAA3 is absorbed from colostrum, and serum levels peak in the second day of life.12 Moreover, SAA measurement has been shown to be significantly elevated in foals with sepsis compared with healthy foals, which indicates endogenous production in response to infection.6, 13 Antigen-presenting cells (APCs; e.g., dendritic cells, macrophages) (Figure 32.2) of newborn foals also show age-dependent development and response to stimulus. For instance, the expression of the major histocompatibility complex (MHC) class II molecule in foal monocyte-derived macrophages and dendritic cells at birth is 12.5 times and 11.2 times inferior, respectively, to adult horse cells.14 Those differences progressively subside by 3 months old. The MHC
class II molecule is essential for the presentation of processed foreign peptides to lymphocytes. In addition, monocyte-derived dendritic cells of foals do not seem to respond to a specific adjuvant stimulus (e.g., cytosine–phosphate–guanosine oligodeoxynucleotide (CpG-ODN)) with interleukin (IL) 12p40 and interferon (IFN) α expression at the same level as adult horse cells, although this limitation is not present when R. equi infection or IFNγ (a T-cell cytokine with positive feedback to antigen presenting cells) is the stimulus.15, 16 It is possible that foal APCs, like human neonates, require multiple stimuli, including toll-like receptor (TLR) binding to PAMPs (e.g., bacteria cell wall) present in the whole organism. When the activation threshold is achieved, APCs can then upregulate important co-stimulatory molecules (e.g., MHC class II, CD86, CD40) and cytokines (IL-12, IFNα) for lymphocyte priming and activation. Cytokines function as a front-line defense of the innate immune system of the foal. Type I interferons have been shown to play an important role at the level of the respiratory mucosa in the control of
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experimental equine herpesvirus 1 infection, before an antigen-specific response is mounted by the foal.17 They are secreted by epithelial cells and phagocytes upon infection, and have direct antiviral effect and signaling to lymphocytes. Inflammatory cytokines IL-1β, IL-6, IL-8, and tumor necrosis factor (TNF)α are secreted during the acute phase of inflammation, and promote the necessary changes in body temperature, vascular permeability, blood flow, and phagocyte recruitment and activation, for the effective removal and killing of pathogens. Paradoxically, these cytokines also promote the severe cardiovascular changes (e.g., hypertension followed by hypotension) and inflammatory damage observed during sepsis. In septic foals, the cytokine profile in the acute phase of infection is not completely understood and remains controversial. Although differences in IL-6, IL-8, IL-10, TNFα, and TGFβ mRNA expression in peripheral blood leukocytes have been suggested, their use in the prognosis for survival is not clear, and serum protein analysis does not necessarily match with leukocyte gene expression analysis due to passive transfer of cytokines through colostrum.18–21 Foal peripheral blood leukocytes stimulated with mitogen present limited but increasing IL-1 and TGF-β1 mRNA expression in the first month of life.22 When stimulated with virulent R. equi, neutrophils obtained from newborn foals or foals at 2, 4, and 8 weeks old induced greater expression of IFNγ, TNFα, IL-6, IL-8, IL-12p40, IL-12p35, and
B cells
IL-23p19 mRNA than unstimulated neutrophils, with some age-dependent impact in the magnitude of response.23
Adaptive immune system Fetal immune system The fetal immune system develops physiologically in the absence of foreign antigens. Initially, the liver and subsequently the bone marrow produce lymphoid and myeloid cells that populate the primary and secondary lymphoid organs, tissue parenchyma, and the circulation. The thymus, in which T-cell development and maturation occurs, presents a corticomedullary organization as early as 80 days of gestation.24, 25 Fetal thymic and peripheral blood T-lymphocytes respond to mitogenic and allogenic stimulation around 80 and 100 days of gestation, respectively. By that time, the spleen reveals small B-cell clusters, in addition to a large number of CD8+ T-cells, and a few CD4+ T-cells (Figure 32.3). The spleen becomes the major secondary lymphoid organ in the second third of the gestation, with lymphocyte distribution in periarteriolar sheaths and germinal centers. Although mesenteric lymph nodes and intestinal lamina propria become populated by lymphocytes by 13 weeks of gestation, most lymph node and mucosal-associated lymphoid tissue (MALT) size and
CD4+ T cells
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Figure 32.3 Immunohistochemical study of spleen sections from an equine fetus (100 days of gestation) and neonate (< 24 hours of life). Note a developing B-cell cluster in the fetal tissue, and the well-developed germinal center in the neonate. In the fetal spleen, there is a paucity of CD4+ T-cells, whereas CD8+ T-cells are abundant. In the absence of pathogens, all cell populations increase in number through gestation, which contributes to the organization of the lymphoid structure by birth. Positive cells are depicted in red. (See color plate 2).
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Development of the Foal Immune System weight increase in response to antigen exposure after birth. B-lymphocytes are functional during normal gestation despite absent or negligible exposure to organisms. By 90 days of gestation, B-cell genes including those for CD20, CD21, CD22, CD27, CD40, CD40LG, B220, CD79A, CD79B, RAG-2, and terminal deoxynucleotidyl transferase (TdT), are expressed in the bone marrow of the equine fetus, and suggest active antigen-independent B-cell development at this stage.26 Under physiological conditions, a serum IgM concentration of 10–22 mg/dL may be measured in fetuses older than 185 days of gestation, and values around 25 mg/dL in presuckle foals.25, 26 The equine fetal B-cells are capable of isotype switching, based on the measurable expression of all immunoglobulin isotype transcripts (IGHG1, IGHG2, IGHG3, IGHG4, IGHG5, IGHG6, and IGHG7) in the spleen, albeit their expression at the protein level is minor or negligible compared with the growing foal. There is minimal IgG immunoglobulin protein concentration (0.2–17 mg/dL) in the presuckle serum, and both IgG1 (5.4 ± 4.0 mg/dL) and IgG4/7 (8.0 ± 5.9 mg/dL) are present. Fetal vaccination at 200 days of gestation can elicit antigen-specific humoral response with IgG production, confirming potential antigen recognition and processing (i.e., B-cell receptor development), and isotype switching from IgM to IgG during fetal life.27, 28 In the horse, the immunoglobulin repertoire includes IgM, IgA, IgE, IgD, and seven different IgG isotypes (IgG1–7 ) defined by different amino acid sequences in the heavy chain constant region (Table 32.1).29 Immunoglobulin diversity allows the recognition of numerous foreign pathogens encountered Table 32.1 Immunoglobulin isotypes in the horse serum. IgG(T) comprises two isotypes named IgG3 and IgG5 , and IgGb includes the IGHG4 and IGHG7 , which share 96% nucleotide sequence homology. The equine immunoglobulin isotypes IgD (IGHD), IgG2 (IGHG2), and IgG6 (IGHG6) have no identified function to date Heavy Chain Constant Gene
Current Designation
Old Nomenclature
IGHA IGHD IGHE IGHG1 IGHG2 IGHG3 and IGHG5 IGHG4 and IGHG7 IGHG6 IGHM
IgA IgD IgE IgG1 IgG2 IgG3/5 IgG4/7 IgG6 IgM
IgA IgD IgE IgGa na IgG(T) IgGb IgGc IgM
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throughout life. In contrast to the heavy chain constant region, which determines the immunoglobulin isotype, the heavy and light chain variable regions bind to the antigen. The variability in the antigen-binding region is achieved through gene recombination and junctional diversity before antigen encounter, and somatic hypermutation after antigen binding. Specific enzymes account for the gene recombination during B-cell development, which include the recombination–activation gene 1 and 2 (RAG-1 and RAG-2, respectively), TdT, and the DNA-dependent protein kinase (DNA-PK). 30 These enzymes are responsible for breaking the immunoglobulin gene, filling the junctions with nontemplated nucleotides, and repairing the ends, respectively, to form the coding immunoglobulin sequence. Enzyme expression is restricted to specific stages in B-cell development (from pro-B to immature B-cell). The heavy and light chain rearrangements allow for the production of over 1 million different immunoglobulins; the diversity provided by TdT (junctional diversity) increases that number to over 100 billion unique immunoglobulins. The diversity of T-cell receptors is also a product of DNA recombination and junctional diversity. The presence of serum immunoglobulin proteins in presuckle neonatal foals indicates that recombination of immunoglobulin genes occurs during fetal life. Molecular studies confirm mRNA expression for recombination–activation genes in equine fetal liver, bone marrow, and spleen.26 Expression of lymphoidspecific TdT is detected in equine fetal liver and bone marrow, similar to several reports of fetal pig B-cells, where evidence for TdT expression is found throughout gestation in the fetal yolk sac, liver, bone marrow, and thymus.31 In contrast, it has been reported that human fetal B-cells have reduced junctional diversity at the gene segment splice sites owing to delayed TdT expression, and TdT is detected only after birth in chickens and rodents.32–36 Expression of enzymes that facilitate increased immunoglobulin diversity during fetal life may be important for equine neonatal survival. Postnatal immune system
IGH, immunoglobulin heavy chain gene; Ig, immunoglobulin; na, not applicable.
The initial development and structural organization of primary and secondary lymphoid tissues happens during fetal life in the absence of foreign antigens. Therefore, the immune system of the equine neonate is naive to environmental organisms at birth. With progressive exposure to an abundant and diverse population of pathogens in early life, the foal responds with a massive expansion of antigen-specific lymphocyte populations. This developmental response is reflected by a progressive increase in the
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number of circulating lymphocytes, and an increase in mass of secondary lymphoid tissue.37 Also, there is an increase in the circulation of cells through the gut and the appearance of gut-homing T-cells, which are likely driven by antigen. Concisely, the postnatal immune system development includes an increase in size of the total lymphocyte compartment and the establishment of primed cells.38 Peripheral blood lymphocyte counts (1000– 2000 cells/µL) in healthy equine neonates are somewhat comparable to adult horses.39 In case of infection or prematurity/immaturity, a lymphopenia may be present and persist for a few or several days. Peripheral blood lymphocyte population increases 2.5–3 times by 3 months old, and blood counts may reach 6000–8000 cells/µL by 6 months old.37 In addition, there is an age-dependent distribution of lymphocyte subpopulations in peripheral blood, with an increase in all subpopulations within the first 3 months of life, particularly CD8+ T-cells and B-cells.40 The increase in circulating lymphocytes likely reflects the intense lymphocyte priming and population expansion in the secondary lymphoid tissues, including lymph nodes and gut- and bronchus associatedlymphoid tissues (GALT and BALT). Individual variability in the distribution of the different lymphocyte subpopulations should be expected, and some foals may take longer to reach normal values than others. Similar to equine fetal lymphocytes, neonatal peripheral blood lymphocytes respond to many non-specific mitogens in vitro (e.g., phytohemagglutinin, PHA; concanavalin A, ConA).37 Moreover, mild spontaneous lymphocyte proliferation in vitro, i.e., without mitogenic stimulation, can be observed progressively in the first several months of life. This lymphocyte activation status may reflect an active response to pathogens in vivo (M.J.B.F. Flaminio, personal observation). Passive transfer of maternal leukocytes through colostrum has not been studied in the foal, but colostral leukocytes have been shown to enter the bloodstream and mesenteric lymph nodes of neonatal rats, primates, piglets, lambs, and calves.41, 42 In calves, those cells can be observed in circulation until 36 hours after colostrum ingestion, when they may reach secondary lymphoid tissues or become eliminated.43 Importantly, colostral lymphocytes are functional, and their presence in the calf’s blood enhances the in vitro response to antigens against which the dams had been vaccinated or exposed.44 Therefore, not only colostral immunoglobulins, opsonins, and cytokines but also lymphocytes can be passively transferred to the neonate for immediate protection after birth, until its own system takes over the task.
Foal peripheral blood and BALF mononuclear cells increase their efficiency of IFNγ production remarkably within the first 3–6 months of life.45 IFNγ is important for the elaboration of Th1-type responses for the killing of intracellular pathogens, such as R. equi and virus infections. Moreover, IFNγ plays an important role in the maturation of APCs for appropriate signaling and priming of lymphocytes. Although neonatal cells can produce this cytokine, it seems that the stimulus needs to overcome a much higher threshold, which could happen naturally with exposure to infection or with the use of appropriate adjuvants in vaccines. Indeed, bronchial lymph node cells from foals experimentally infected with R. equi express high levels of IFNγ, with a high IFNγ/IL-4 ratio.46 In addition, isolated bronchial lymph node lymphocytes from experimentally infected foals respond with high proliferation and IFNγ production when stimulated with R. equi virulence-associated proteins in vitro.47 Mature horse lymphocytes express the MHC class II molecules. However, foal lymphocytes show an age-dependent expression of these molecules and, by 4–6 months of age, the number of positive cells is comparable to adult horses.37, 48 The implications of the lack of MHC class II expression on lymphocytes in the young foal are unknown; yet, this molecule serves as a marker for maturation of lymphocytes. After birth, as soon as the foal is challenged by environmental organisms, the immune system reveals an antigen-dependent phase of expansion, in which B-cells go through immunoglobulin isotype switching, and develop affinity and memory for the target antigen. The capacity to produce large amounts and a diverse array of antibodies is dependent on lymphocyte population expansion and appropriate cell interactions. Therefore, endogenous antibody production to protective levels (e.g., > 400–500 mg/dL) may be reached only by 5–8 weeks of life. Meanwhile, colostral IgG concentrations are decreasing with a half-life of 28–32 days in the healthy foal.49 The onset and amount of endogenous antibody production was compared between foals fed or deprived of colostrum.50 In foals deprived of colostrum, serum IgG levels start to rise significantly at 2 weeks old, and, by 5 weeks, levels were comparable or superior to colostrum-fed foals. In foals deprived of equine colostrum but fed bovine colostrum instead, the production of endogenous antibody can be followed without the confounding element of circulating maternal antibodies: serum equine IgG levels were about 200 mg/dL by 14–19 days, 700 mg/dL by 37–50 days, and > 1000 mg/dL by 102–135 days of life.51 Therefore, endogenous antibody production in the foal is likely stimulated by the exposure to antigens.
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Relative decay of total colostral antibody is proportional to the initial amount of immunoglobulin absorbed at birth (Figure 32.4). Therefore, larger colostral IgG concentrations will maintain adequate levels for a longer period of time, perhaps beyond the time the foal’s endogenous production reaches protective levels. On the other hand, low initial colostral IgG concentrations may persist until endogenous production reaches protective levels; in this case, the foal becomes more susceptible to infection earlier in life. Each immunoglobulin isotype possesses distinct functions for neutralization, complement fixation and opsonization, and mucosal protection. The combination of isotypes provides a functional strategy for removal of pathogens, and lack of specific isotypes may facilitate certain pathogen establishment and disease. It is important to mention that immunoglobulin isotype concentrations vary according to age, breed, vaccination status, and overall management of horse populations, and such differences may have implications for the quality and duration of protection, as well the response to pathogens and vaccines. The most abundant immunoglobulin isotype in colostrum and serum of a mature horse is IgG4/7 , which accounts for approximately 60% of all isotypes.52, 53 Importantly, colostral IgG4/7 has the longest half-life in the foal (32 days), followed by IgG3/5 (21), IgG1 (17.6), IgM (5–10), and IgA (3.4). Therefore, all colostral immunoglobulin isotypes, with exception of IgG4/7 , reach their nadir concentrations in the foal by 4 weeks of life. Concurrently, production of most immunoglobulin isotypes is active, and levels comparable to mature horses are achieved by 8–12 weeks of life. In contrast, IgG4/7 nadir levels are reached by 12–16 weeks, and, because endogenous production is apparently delayed, values comparable to mature horses are reached beyond 1 year of life.53 On the other hand, the analysis of immunoglobulin isotype mRNA expression in the foal, which offers less conflicting data with colostral immunoglobulin proteins, indicates that the foal is actively expressing IGHG4 and IGHG7 genes in the secondary lymphoid tissues and peripheral blood mononuclear cells. In fact, foals less than 1 month old that were challenged with R. equi responded with increased serum IgG1 and IgG4/7 specific antibody concentrations,46 indicating their competence when the appropriate stimulus is provided (e.g., whole organism, adjuvant, etc.). Immunoglobulin E is also transferred to the foal through colostrum, and IgE is found either bound to peripheral blood leukocyte membranes or soluble in serum for a period of 2 and 4 months of life, respectively.54 In the foal, there is a long delay in endogenous IgE production; although IgE+ B-cells
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Figure 32.4 Hypothetical serum immunoglobulin (Ig)G concentrations in foals, including colostral IgG (circles), foal endogenous IgG production (squares), and the sum of both (triangles). The dashed line indicates minimal levels of protection (400–500 mg/dL). (a) Adequate transfer of immunoglobulins through the colostrum (starting at 2000 mg/dL in the foal serum) and the physiological decay based on IgG half-life. Note that the colostral antibodies and the foal endogenous IgG production sum up to protective levels at the nadir between 8 and 12 weeks. (b) Despite lower (800 mg/dL) initial serum IgG concentrations in the newborn foal, the total IgG concentration is still above protective level, but the nadir is predicted to be anticipated at 3–4 weeks. (c) With initial serum IgG concentration of 400 mg/dL, the foal is already at risk in the first week of life. A delay in endogenous antibody production in the foal creates a window of susceptibility, and the lower the initial colostral IgG concentration, the less time the foal has to compensate with endogenous production.
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can be detected in lymphoid follicles of some foals at 6 months of life, serum IgE values are detectable around 9 months of life.55 This observation creates some questions about the mechanisms of defense against parasites in the foal, and the development of allergies at a later age.
Mucosal immunity The respiratory tract is an important and common route of infection in the neonate. As described above, epithelial cells and cytokines participate in innate immunity, detecting the presence of pathogens and promoting phagocyte recruitment. While the thymus, spleen, and some systemic lymph nodes develop their structure during fetal life, foals do not present organized lymphoid tissue in the lungs at birth. Tlymphocytes and plasma cells are virtually absent in lung tissues in the first week of life, and BALT is observed only by 12 weeks old.38 The population of the lung with lymphocytes is delayed in comparison with peripheral blood and other lymphoid tissues. The concentration of leukocytes in bronchoalveolar lavage fluid (BALF) in foals less than 3 weeks old is one-half the values of adult horses.37 The percentage of CD4+ T-cells in BALF increases markedly in the first 3 weeks of life.56 B-cell counts in BALF are almost undetectable in the first 4 weeks of life, and reach values comparable to adult horses by 8 weeks of age. Around this same time, higher levels of IgA, IgG, and IgM are also detected in BALF. Therefore, the respiratory tract of the foal seems poorly protected in the first 8–12 weeks of life, and this may explain, in part, the high incidence of respiratory infections. Whereas IgA predominates in the nasal secretions of mature horses, IgG1 and IgG4/7 predominate in foals younger than 2 weeks old, and they may still be of maternal origin. Nevertheless, studies on passive transfer of antibodies against Streptococcus equi subsp equi through colostrum revealed the presence of colostral IgG and polymeric IgA on the nasopharyngeal mucosa, with peak levels at 2 days of life.57 Endogenous IgA concentrations become measurable beyond 28 days of life, reaching levels comparable to adult horses by 42 days. Likewise, the gastrointestinal tract is in close contact with environmental pathogens. The immunoglobulins present in colostrum and milk offer significant passive mucosal protection. The equine milk has a higher concentration of IgA than colostrum. Immunoglobulin A on the mucosal surface impairs bacterial binding to enterocytes, and neutralizes bacterial enterotoxins and enterotropic viruses. The development of intestinal tract lym-
phoid tissue is marked after birth, and Payer’s patches and mesenteric lymph nodes become grossly obvious beyond 3 weeks of life. The GALT protective function requires microbial stimulation provided by the initial bacterial colonization after birth and energy substrate present in the milk.58 Non-digestible milk carbohydrates create the appropriate acidic environment for the proliferation of probiotic/ commensal bacteria, which subsequently promote the development of the acquired immune response via the expression of PAMPs. Although the foal is likely born with all the elements of the mucosal immune system to detect and respond to PAMPs, it is only after exposure that secretion of mucosal IgA and the expansion of the lymphocyte population occurs. It is expected that exposure to pathogenic bacteria before this necessary priming would cause disease in the foal; in addition, elements that may affect the proliferation of resident bacteria may affect or delay protective immune response.
Foal vaccination A vaccination program has the ultimate objective of lifetime prophylaxis by inducing an immune response that provides protection against natural disease. Ideally, vaccination of foals should anticipate the drop of colostral antibodies. However, in neonatal immunology, there is a concern about maternal antibody interference with humoral response to vaccines.59, 60 The same colostral antibodies that confer protection to the foal could potentially neutralize vaccine antigens and prevent the priming of the Bcells or humoral response; alternatively, mechanisms of antibody removal or recycling may play a role. Therefore, the amount of colostral immunoglobulin absorbed by foals may delay the synthesis of the corresponding endogenous antigen-specific antibodies. However, it is possible that the foal can still respond to vaccine antigens with both humoral and cellular immunity, and memory cells, without a significant change in circulating antibodies, when given the appropriate stimulus. Whereas laboratory techniques have been more accessible to evaluate cellular immunity in the horse, humoral response in the neonate is difficult to measure because of the confounding levels of colostral-derived antibodies; yet, the presence of antigen-specific memory B-cells or resistance to post-vaccination pathogen challenges may confirm neonatal response to vaccinations despite high levels of passively transferred antibodies. Certain types of vaccines and antigen patterns may not elicit recognition and immune response in the young, and strategies should be studied and designed to overcome this limitation.
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Development of the Foal Immune System Passive humoral protection against pathogens of particular importance or danger to the foal (e.g., rotavirus, Clostridium botulinum, Clostridium tetani) can be improved by vaccinating the mare during gestation, and ensuring adequate ingestion of colostrum. Experimental vaccination of mares with Clostridium perfringens type A toxin at 8 and 4 weeks before foaling resulted in efficient transfer of antitoxin to the foal through colostrum. Colostral antitoxin titers were at least twice the serum levels of the dam within the first 3 hours after parturition.50 Foal serum antitoxin titers were approximately half of the dams’ serum levels by 12–18 hours after birth, with a half-life of 20 days. For pathogens with a lower risk of disease in early life, vaccination should commence when colostral antibody concentrations are decreasing in the foal. Pathogen-specific antibodies may be transferred to the foal through colostrum in different concentrations, and their catabolism or consumption may vary. The half-lives of maternally derived antibodies against some relevant pathogens have been described: tetanus (35 days), equine encephalomyelitis (20 days), equine influenza (27 (IgG1 ) to 39 (IgG4 ) days), and equine viral arteritis (32 days).60–62 In general, vaccination of the foal starts between 3 and 4 months of life, with at least one booster 4 weeks later. In fact, vaccination with three doses in the primary series may compensate for potential maternal antibody interference with the first dose. This practice may be effective for most vaccines that elicit immune responses in the foal, with the exception of the influenza vaccine. For the latter, primary vaccination in foals from vaccinated mares is recommended to start beyond 6 months old because earlier vaccination delays endogenous response to this vaccine even further. Unless specified by the manufacturer, modifiedlive vaccines should not be used in foals less than 6 months old. Protocols for the immunization of foals are periodically updated based on new information gained regarding the foal’s immune response, vaccine products, and applied vaccinology; therefore, in the future, innovative vaccines may be developed to overcome maternal antibody interference and/or target cellular immunity for use in the neonatal phase.
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References 1. Wichtel MG, Anderson KL, Johnson TV, Nathan U, Smith L. Influence of age on neutrophil function in foals. Equine Vet J 1991;23:466–9. ¨ 2. Grondahl G, Sternberg S, Jensen-Waern M, Johannisson A. Opsonic capacity of foal serum for the two neonatal pathogens Escherichia coli and Actinobacillus equuli. Equine Vet J 2001;33:670–5. 3. Flaminio MJBF, Rush BR, Davis EG, Shuman W, Wilkerson M. Simultaneous analysis of phagocytosis and oxidative burst activity of equine phagocytes by flow cytometry. In:
17.
18.
19.
339
Lunn DP, Wade JF (eds) Havemeyer Foundation Monograph Series, No. 4, 2001, pp. 69–70, R & W Publications, Suffolk. ¨ Grondahl G, Johannisson A, Demmers S, Jensen Waern M. Influence of age and plasma treatment on neutrophil phagocytosis and CD18 expression in foals. Vet Microbiol 1999;65:241–54. Bernoco M, Liu IK, Wuest-Ehlert CJ, Miller ME, Bowers J. Chemotactic and phagocytic function of peripheral blood polymorphonuclear leucocytes in newborn foals. J Reprod Fertil Suppl 1987;35:599–605. Gardner RB, Nydam DV, Luna JA, Bicalho ML, Matychak MB, Flaminio MJ. Serum opsonization capacity, phagocytosis, and oxidative burst activity in neonatal foals in the intensive care unit. J Vet Intern Med 2007;21:797–805. LeBlanc MM, Pritchard EL. Effects of bovine colostrum, foal serum immunoglobulin concentration and intravenous plasma transfusion on chemiluminescence response of foal neutrophils. Anim Genet 1988;19:435–45. Bernoco MM, Liu IK, Willits NH. Hemolytic complement activity and concentrations of its third component during maturation of the immune response in colostrum-deprived foals. Am J Vet Res 1994;55:928–33. McTaggart C, Penhale J, Raidal SL. Effect of plasma transfusion on neutrophil function in healthy and septic foals. Aust Vet J 2005;83:499–505. McDonald TL, Larson MA, Mack DR, Weber A. Elevated extrahepatic expression and secretion of mammaryassociated serum amyloid A 3 (M-SAA3) into colostrum. Vet Immunol Immunopathol 2001;83:203–11. Duggan VE, Holyoak GR, Macallister CG, Cooper SR, Confer AW. Amyloid A in equine colostrum and early milk. Vet Immunol Immunopathol 2008;121:150–5. Stoneham SJ, Palmer L, Cash R, Rossdale PD. Measurement of serum amyloid A in the neonatal foal using a latex agglutination immunoturbidimetric assay: determination of the normal range, variation with age and response to disease. Equine Vet J 2001;33:599–603. Paltrinieri S, Giordano A, Villani M, Manfrin M, Panzani S, Veronesi MC. Influence of age and foaling on plasma protein electrophoresis and serum amyloid A and their possible role as markers of equine neonatal septicaemia. Vet J 2008;176:393–6. Flaminio MJ, Borges AS, Nydam DV, Horohov DW, Hecker R, Matychak MB. The effect of CpG-ODN on antigen presenting cells of the foal. J Immune Based Ther Vaccines 2007; 25;5:1–17. Flaminio MJ, Nydam DV, Marquis H, Matychak MB, Gigu`ere S. Foal monocyte-derived dendritic cells become activated upon Rhodococcus equi infection. Clin Vaccine Immunol 2009;16:176–83. Sponseller BA, de Macedo MM, Clark SK, Gallup JM, Jones DE. Activation of peripheral blood monocytes results in more robust production of IL-10 in neonatal foals compared to adult horses. Vet Immunol Immunopathol 2009;127: 167–73. Chong YC, Duffus WP. Immune responses of specific pathogen free foals to EHV-1 infection. Vet Microbiol 1992;32:215–28. Pusterla N, Magdesian KG, Mapes S, Leutenegger CM. Expression of molecular markers in blood of neonatal foals with sepsis. Am J Vet Res 2006;67:1045–9. Gold JR, Perkins GA, Erb HN, Ainsworth DM. Cytokine profiles of peripheral blood mononuclear cells isolated from septic and healthy neonatal foals. J Vet Intern Med 2007;21:482–8.
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20. Secor EJ. Macrophage migration inhibitory factor (MIF) and tumor necrosis factor alpha (TNF-α ) as diagnostic markers in septic foals. A thesis presented to the Faculty of the Department of Animal Science in partial fulfillment of the requirements for a Bachelor’s Degree with Distinction in Research, Cornell University, 2008. 21. Burton AB, Wagner B, Erb HN, Ainsworth DM. Serum interleukin-6 (IL-6) and IL-10 concentrations in normal and septic neonatal foals. Vet Immunol Immunopathol 2009;132:122–8 (ePub available). 22. Boyd NK, Cohen ND, Lim WS, Martens RJ, Chaffin MK, Ball JM. Temporal changes in cytokine expression of foals during the first month of life. Vet Immunol Immunopathol 2003;92:75–85. 23. Nerren JR, Martens RJ, Payne S, Murrell J, Butler JL, Cohen ND. Age-related changes in cytokine expression by neutrophils of foals stimulated with virulent Rhodococcus equi in vitro. Vet Immunol Immunopathol 2009;127: 212–9. 24. Mackenzie CD. Histological development of the thymic and intestinal lymphoid tissue of the horse. J S Afr Vet Assoc 1975;46:47–55. 25. Perryman LE, McGuire TC, Torbeck RL. Ontogeny of lymphocyte function in the equine fetus. Am J Vet Res 1980;41:1197–200. 26. Tallmadge RL, McLaughlin K, Secor E, Ruano D, Matychak MB, Flaminio MJBF. Expression of essential B cell genes and immunoglobulin isotypes suggests active development and gene recombination during equine gestation. Dev Comp Immunol 2009;33:1027–38. 27. Martin BR, Larson KA. Immune response of equine fetus to coliphage T2. Am J Vet Res 1973;34:1363–4. 28. Morgan DO, Bryans JT, Mock RE. Immunoglobulins produced by the antigenized equine fetus. J Reprod Fertil Suppl 1975;23:735–8. 29. Wagner B. Immunoglobulins and immunoglobulin genes of the horse. Dev Comp Immunol 2006;30:155–64. 30. Kallenbach S, Doyen N, Fanton d’Andon M, Rougeon F. Three lymphoid-specific factors account for all junctional diversity characteristic of somatic assembly of T-cell receptor and immunoglobulin genes. Proc Natl Acad Sci USA 1992;89:2799–803. 31. Butler JE, Sinkora M, Wertz N, Holtmeier W, Lemke CD. Development of the neonatal B and T cell repertoire in swine: implications for comparative and veterinary immunology. Vet Res 2006;37:417–41. 32. Sugimoto M, Bollum FJ. Terminal deoxynucleotidyl transferase (TdT) in chick embryo lymphoid tissues. J Immunol 1979;122:392–7. 33. Gregoire KE, Goldschneider I, Barton RW, Bollum FJ. Ontogeny of terminal deoxynucleotidyl transferase-positive cells in lymphohemopoietic tissues of rat and mouse. J Immunol 1979;123:1347–52. 34. Bodger MP, Janossy G, Bollum FJ, Burford GD, Hoffbrand AV. The ontogeny of terminal deoxynucleotidyl transferase positive cells in the human fetus. Blood 1983;61:1125– 31. 35. Raaphorst FM, Timmers E, Kenter MJ, Van Tol MJ, Vossen JM, Schuurman RK. Restricted utilization of germ-line VH3 genes and short diverse third complementaritydetermining regions (CDR3) in human fetal B lymphocyte immunoglobulin heavy chain rearrangements. Eur J Immunol 1992;22:247–51. 36. Kincade PW, Owen JJ, Igarashi H, Kouro T, Yokota T, Rossi MI. Nature or nurture? Steady-state lymphocyte forma-
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tion in adults does not recapitulate ontogeny. Immunol Rev 2002;187:116–25. Flaminio MJ, Rush BR, Davis EG, Hennessy K, Shuman W, Wilkerson MJ. Characterization of peripheral blood and pulmonary leukocyte function in healthy foals. Vet Immunol Immunopathol 2000;73:267–85. Banks EM, Kyriakidou M, Little S, Hamblin AS. Epithelial lymphocyte and macrophage distribution in the adult and fetal equine lung. J Comp Pathol 1999;120:1–13. Becht JL, Semrad SD. Hematology, blood typing, and immunology of the neonatal foal. Vet Clin North Am Equine Pract 1985;1:91–116. Smith R 3rd, Chaffin MK, Cohen ND, Martens RJ. Agerelated changes in lymphocyte subsets of quarter horse foals. Am J Vet Res 2002;63:531–7. Parmely MJ, Beer AE. Colostral cell-mediated immunity and the concept of a common secretory immune system. J Dairy Sci 1977;60:655–65. Sheldrake RF, Husband AJ. Intestinal uptake of intact maternal lymphocytes by neonatal rats and lambs. Res Vet Sci 1985;39:10–15. Reber AJ, Hippen AR, Hurley DJ. Effects of the ingestion of whole colostrum or cell-free colostrum on the capacity of leukocytes in newborn calves to stimulate or respond in one-way mixed leukocyte cultures. Am J Vet Res 2005;66:1854–60. Donovan DC, Reber AJ, Gabbard JD, Aceves-Avila M, Galland KL, Holbert KA, Ely LO, Hurley DJ. Effect of maternal cells transferred with colostrum on cellular responses to pathogen antigens in neonatal calves. Am J Vet Res 2007;68:778–82. Breathnach CC, Sturgill-Wright T, Stiltner JL, Adams AA, Lunn DP, Horohov DW. Foals are interferon gammadeficient at birth. Vet Immunol Immunopathol 2006;112:199– 209. Jacks S, Gigu`ere S, Prescott JF. In vivo expression of and cell-mediated immune responses to the plasmid-encoded virulence-associated proteins of Rhodococcus equi in foals. Clin Vaccine Immunol 2007;14:369–74. Jacks S, Gigu`ere S, Crawford PC, Castleman WL. Experimental infection of neonatal foals with Rhodococcus equi triggers adult-like gamma interferon induction. Clin Vaccine Immunol 2007;14:669–77. Lunn DP, Holmes MA, Duffus WP. Equine T-lymphocyte MHC II expression: variation with age and subset. Vet Immunol Immunopathol 1993;35:225–38. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Absorption of bovine colostral immunoglobulins G and M in newborn foals. Am J Vet Res 1989;50:1598–603. Jeffcott LB. Studies on passive immunity in the foal. Gamma-globulin and antibody variations associated with the maternal transfer of immunity and the onset of active immunity. J Comp Pathol 1974;84:93–101. Holmes MA, Lunn DP. A study of bovine and equine immunoglobulin levels in pony foals fed bovine colostrum. Equine Vet J 1991;23:116–18. Sheoran AS, Timoney JF, Holmes MA, Karzenski SS, Crisman MV. Immunoglobulin isotypes in sera and nasal mucosal secretions and their neonatal transfer and distribution in horses. Am J Vet Res 2000;61:1099–105. Holznagel DL, Hussey S, Mihalyi JE, Wilson WD, Lunn DP. Onset of immunoglobulin production in foals. Equine Vet J 2003;35:620–2. Wagner B, Flaminio JB, Hillegas J, Leibold W, Erb HN, Antczak DF. Occurrence of IgE in foals: evidence for
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transfer of maternal IgE by the colostrum and late onset of endogenous IgE production in the horse. Vet Immunol Immunopathol 2006;110:269–78. Marti E, Ehrensperger F, Burger D, Ousey J, Day MJ, Wilson AD. Maternal transfer of IgE and subsequent development of IgE responses in the horse (Equus callabus). Vet Immunol Immunopathol 2009;127:203–11. Balson GA, Smith GD, Yager JA. Immunophenotypic analysis of foal bronchoalveolar lavage lymphocytes. Vet Microbiol 1997;56:237–46. Galan JE, Timoney JF, Lengemann FW. Passive transfer of mucosal antibody to Streptococcus equi in the foal. Infect Immun 1986;54:202–6. Forchielli ML, Walker WA. The role of gut-associated lymphoid tissues and mucosal defence. Br J Nutr 2005;93 Suppl 1:S41–8.
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59. Van Oirschot JT, Bruin G, de Boer-Luytze E, Smolders G. Maternal antibodies against equine influenza virus in foals and their interference with vaccination. Zentralbl Veterinarmed B 1991;38:391–6. 60. Wilson WD, Mihalyi JE, Hussey S, Lunn DP. Passive transfer of maternal immunoglobulin isotype antibodies against tetanus and influenza and their effect on the response of foals to vaccination. Equine Vet J 2001;33:644–50. 61. Ferguson JA, Reeves WC, Hardy JL. Studies on immunity to alphaviruses in foals. Am J Vet Res 1979;40:5–10. 62. Hullinger PJ, Wilson WD, Rossitto PV, Patton JF, Thurmond MC, MacLachlan NJ. Passive transfer, rate of decay, and protein specificity of antibodies against equine arteritis virus in horses from a Standardbred herd with high seroprevalence. J Am Vet Med Assoc 1998;213: 839–42.
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Chapter 33
Colostrum: Assessment of Quality and Artificial Supplementation Guy D. Lester
Colostrum accumulates within the mammary gland over the final weeks of pregnancy. The composition of colostrum differs from that of milk in several important areas, including higher concentrations of protein and fat, and lower concentrations of lactose. The elevated protein content largely reflects immunoglobulins (Ig), which are concentrated in the alveoli under the influence of estrogens and progesterone in late pregnancy. The principal antibody type transferred through colostrum is IgG, including subtypes IgGa, IgGb, and IgG(T), with lesser quantities of IgM, IgA, and IgE. The IgG and IgM concentrations in colostrum are approximately 166 g/L and 1.0 g/L, respectively, and the colostrum to serum ratios of each are 8:1 and 1:1, respectively.1 The foal is capable of synthesizing IgM and smaller concentrations of IgGa and IgGb during gestation. Immunoglobulin Ga, IgA, and IgG(T) are synthesized from birth and detectable by 2 months of age, but the production of IgGb, the principal IgG subtype of adult horses, is delayed. Typically, IgE is not produced by foals until 8–10 months of life. Maternal IgG not only provides immediate protection against environmental pathogens, but also has a stimulatory effect on the foal’s immune system.2 An important foal response to passively derived maternal IgG is the formation of anti-idiotypic antibodies, which are important in regulating neonatal T-cell and B-cell responses, the magnitude of immune responses, and providing additional antimicrobial protection.2 Induction of lactation in non-pregnant mares using vaginal pessaries containing altrenogest and estradiol benzoate, followed by orally administered dopamine antagonists (sulpiride and domperidone
used to increase prolactin), can produce a very small quantity of good quality colostrum in some mares.3 Colostral quality, as assessed by IgG concentration, varies widely between mares. The total IgG content in mammary secretions produced over the initial 16–20 hours after delivery varied in six mares between 199 and 855 g.1 This equated to a volume of approximately 5.1 L with a range of 3.2 to 7.0 L. Much of this volume likely included mammary secretions with little or no IgG. Massey et al. (1991)4 reported a total IgG mass of 183 ± 58 g (range 112–336 g) in 18 mares of varying breeds from milk samples that had a specific gravity ≥ 1.040. This equated to a mean volume of colostrum of 2.3 ± 0.5 L. The specific gravity declined to below 1.040 by an average of 7 hours after parturition reflecting a reduction in IgG concentration from a mean of 113 g/L to 35 g/L. Good quality colostrum is typically yellow and highly viscous, but it has been reported that these physical properties alone do not result in significant differences in foal IgG levels at 24 hours of age.5 There is a strong linear correlation between colostral specific gravity and IgG concentration with r2 values of 0.801 and 0.81.6 A modified hydrometer (Equine colostrometer, Jorgensen Laboratories, Loveland, CO) has been used to estimate the specific gravity of equine colostrum. It is important accurately to add the recommended volume of colostrum to the chamber. The temperature of the distilled water will also influence the final reading, and corrective tables are available. Hand-held refractometers designed to measure sugar or alcohol content are suitable for assessing colostral quality. The test can be performed rapidly
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Colostrum: Assessment of Quality and Artificial Supplementation by placing a single drop of mixed colostrum onto the glass prism of the refractometer. The correlation between the alcohol refractometer reading and colostral IgG content is good, with an r2 of 0.74.5 A second study reported an r-value of 0.94.7 Values greater than or equal to 16% on the alcohol scale, or 23% on the BRIX sugar scale correspond to a colostral IgG content ≥ 60 g/L. Cash (1999)7 reported the following guidelines for evaluating BRIX readings: (i) 0–15% BRIX, 0–28 g/L IgG, poor quality; (ii) 15–20% BRIX, 28–50 g/L IgG, borderline quality; (iii) 20–30% BRIX, 50–80 g/L IgG, adequate quality; and (iv) > 30% BRIX, > 80 g/L IgG, very good quality. There are several commercial kits used to estimate the IgG concentration in the foal that are based on the first-order chemical reaction between glutaraldehyde and immunoglobulins. The reaction occurs between the aldehyde groups of glutaraldehyde and free amino groups in globulins.8 The reaction time is directly proportional to the concentration of immunoglobulins. A similar reaction occurs between glutaraldehyde and fibrinogen; false positive results may theoretically occur in diseased foals with an elevated concentration of fibrinogen. Sample hemolysis is also reported to result in a false positive result. These tests are available for detection of IgG in whole blood and serum and return a result within 5 minR utes (Gamma-check E , Plasvacc). The test has also been modified by one manufacturer to estimate the IgG concentration in colostrum, with a positive test result indicating a colostral IgG content of 38 g/L or R , Plasvacc). greater (Gamma-check C A minimum dose of 60–90 g of IgG has been recommended in the first 6 hours after birth.9 This represents 1–1.5 L of good quality colostrum, as defined by an IgG content of greater than 60 g/L. Foals that receive a minimum of 1.0–1.25 g IgG/kg bodyweight within this time period should achieve a serum IgG above 8 g/L (800 mg/dL).4 There are several factors that influence colostral quality. Mares that are stressed or ill are less likely to produce good quality colostrum, as are animals with adrenal hyperplasia secondary to pituitary dysfunction or those receiving exogeneous glucocorticoids. Data derived from dairy cattle indicate that both endogenous and exogenous glucocorticoids, such as dexamethasone, can reduce the concentration of IgG1 in colostrum.10 Although not examined it would seem reasonable that a similar glucocorticoid influence on colostral quality is likely in mares. Aged mares, particularly those older than 15 years, tend to produce colostrum with a lower IgG concentration.9 In one study Arabian and Thoroughbred mares had higher colostral IgG concentrations than Standardbred mares.9 The collection of small volumes of high quality colostrum (200–250 mL) for banking is unlikely to have any significant effect on the foal’s serum IgG
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value. Colostrum should be taken within the first 2 hours after delivery from the teat opposite to that of the initial suck as foals typically stay on the same teat for the first 3–4 sucks.11 The colostrum should be tested using either a handheld refractometer or a modified hydrometer. Collection should be avoided in mares that are likely to produce colostrum of suboptimal quality. This would include older mares, those that dripped colostrum or milk prior to foaling, and those with concurrent illness.9 After collection colostrum should be filtered through gauze or cloth to remove debris and placed into a solid plastic container or heavy duty plastic bag. It can be frozen for one to two years; ideally storage is at −20◦ C (−4◦ F) although a regular household freezer is suitable for 12 months of storage. Thawing of colostrum is ideally performed using warm water (38◦ C) or placement at room temperature. Microwave use should be discouraged, as it has the potential to denature immunoglobulins. If used the power rating should be reduced to 50%. Supplementation with good quality (> 23% BRIX) banked colostrum of foals born to mares with poor quality pre-suckle colostrum (< 20% BRIX) reduces the risk of failure of passive transfer.12 The quantity of colostrum required depends on a range of factors including both mare and foal factors but may be between 500 and 1000 mL. The authors advocated the latter for mares with a pre-suckle colostral BRIX value < 15%.12 Colostral quality and quantity are not the only critical factors in determining adequacy of passive transfer of immunoglobulins to newborn foals. Timing of consumption relative to birth is also paramount. The efficiency of colostral IgG absorption into the vascular space was estimated at 51% when colostrum was consumed within the first 12 hours after birth.13 Initial consumption after 12 hours leads to a reduction in the efficiency of colostral IgG absorption to around 28%; the reduction occurred irrespective of whether the foal had received milk replacer or electrolyte solution during the 12 hours after birth. The general recommendation has been to administer colostrum within 6 hours of birth to maximize absorption, but more recent recommendations are to encourage colostrum consumption within the first 1–2 hours of postnatal life. These recommendations stem from the knowledge that many neonatal infections are likely acquired through the intestinal tract. The enhanced phagocytic activity of the newborn intestine may not readily discriminate between immunoglobulins and other large molecules. Intestinal pinocytosis of non-pathogenic E. coli was demonstrated experimentally in newborn piglets that were dosed intragastrically with the organisms between 1 and 2 hours after birth.14 Therefore, the early consumption of colostrum is recommended both to provide local
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protection against ingested bacterial pathogens and potentially to hasten gut closure. It is unlikely that early feeding of colostrum would substantially accelerate gut closure, but more detailed investigations are required. A weak but significant correlation between colostral IgG and foal serum IgG concentration has been reported, with correlation coefficients between 0.46 and 0.64.7, 15, 16 There are additional foal factors that impact the efficiency of absorption of colostral antibodies. Gestational age and the degree of maturity appear to influence colostral absorption as premature and post-mature foals commonly experience partial failure of passive transfer.9 The prophylactic administration of antidiarrheal agents, such as ditri-octahedral smectite, to foals at risk for clostridial enterocolitis could reduce the efficiency of absorption by adsorbing colostral IgG.17 It has been recommended to delay administration of oral antidiarrheal medications until 6–8 hours after colostral intake. Adequacy of passive transfer is generally accepted if the serum concentration of IgG exceeds 8 g/L (800 mg/dL). Serum levels less than 2 g/L reflect total failure of passive transfer (FPT) whereas levels between 2 and 8 mg/mL typically indicate a partial failure of passive transfer. Clabough et al.18 reported that 18% of 361 foals had an IgG concentration that was less than 4 g/L; Stoneham et al.19 reported similar findings in 15% of 1922 foals, and LeBlanc et al.9 in 13% of 253 foals. Most, but not all, studies have reported a positive correlation between failure of passive transfer and neonatal sepsis. Experimentally, deprivation of colostrum in healthy newborn foals results in a high rate of bacterial sepsis.20 Baldwin et al. (1991)21 reported a low rate of neonatal disease despite a 14% incidence of failure of passive transfer (IgG < 4 g/L) in a group of Standardbred foals, illustrating the importance of management as a critical cofactor in the development of neonatal sepsis. There are several alternatives that could be used when natural equine colostrum is unavailable and the foal is still at an age when significant intestinal antibody uptake is likely to occur, that is, less than 18–24 hours of age. Bovine colostrum can provide some important protection to certain environmental pathogens, but does not protect the foal against equine-specific pathogens, such as Actinobacillus equuli. Similar responses are reported with caprine colostrum. Bovine-derived antibodies (IgG and IgM) have a short half-life in foals but IgG concentrations were not significantly different between foals fed bovine colostrum and those consuming equine colostrum.22 Bovine IgG has a half-life of around 9 days in foals compared with maternal equine IgG, which has a half-life of approximately 26 days in normal foals. There is anecdotal evidence that feeding
bovine colostrum after gut closure may provide some additional protection against intestinal pathogens. Plasma can be administered orally to foals in lieu of colostrum although larger volumes are required due to the reduced concentration of immunoglobulins in plasma: 17–25 g IgG/L in plasma compared with 100–120 g/L in good quality colostrum. Larger volumes should be divided into smaller aliquots and administered over 2 hours. A concentrated equine IgG product collected from gelding draft horses and gamma-irradiated is available commercially in some countries (Seramune, Sera Inc., USA). The manufacturers report an IgG concentration between 100 and 120 g/L, depending on product type (oral or intravenous), delivering an IgG content of around 30 g per foal. Consequently the products are used mainly as a supplement for foals with partial failure of passive transfer. The administration of any protein-containing product intravenously should be done with caution, as anaphylactoid reactions are possible. Similarly, a commercial lyophilized equine IgG supplement has been used in foals at risk for partial failure of passive transfer (Lyphomune, Bioqual, Inc.). The coefficient of absorption was not different between foals that received pooled thawed colostrum and those that received lyophilized IgG diluted in milk replacer.23 The company also advocated use either intraveneously or orally. At the time of writing there were problems in availability of this product. Another commercial product is based on feeding lyophilized whole egg protein to newborn foals at birth and then 6–8 hours after nursing (EPIC, Bioniche Animal Health). The egg protein is derived from hens immunized with inactivated multivalent vaccines. Marketing of the product is primarily directed towards use as an adjunct to a preventative program against neonatal diarrhea. Most controlled efficacy data have been generated in newborn calves and neonatal pigs; unfortunately similar data in foals are lacking.24, 25
References 1. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Colostral volume and immunoglobulin G and M determinations in mares. Am J Vet Res 1989;50:466–70. 2. Wagner B, Flaminio JB, Hillegas J, Leibold W, Erb HN, Antczak DF. Occurrence of IgE in foals: evidence for transfer of maternal IgE by the colostrum and late onset of endogenous IgE production in the horse. Vet Immunol Immunopathol 2006;110:269–78. 3. Chavatte-Palmer P, Arnaud G, Duvaux-Ponter C, Brosse L, Bougel S, Daels P, Guillaume D, Clement F, Palmer E. Quantitative and qualitative assessment of milk production after pharmaceutical induction of lactation in the mare. J Vet Intern Med 2002;16:472–77. 4. Massey RE, LeBlanc MM, Klapstein EF. Colostrum feeding of foals and colostrum banking. In: Proceedings of the 37th
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Annual Convention of the American Association of Equine Practitioners, San Francisco, 1991, pp. 1–8. Chavatte P, Clement F, Cash R, Grongnet JF. Field determination of colostral quality by using a novel, practical method. In: Proceedings of the 44th Annual Convention of the American Association of Equine Practitioners, Baltimore, 1998, pp. 206–9. LeBlanc MM, McLaurin BI, Boswell R. Relationships among serum immunoglobulin concentration in foals, colostral specific gravity, and colostral immunoglobulin concentration. J Am Vet Med Assoc 1986;189:57–60. Cash RSG. Colostral quality determined by refractometry. Equine Vet Educ 1999;11:36–8. Brink P, Wright JC, Schumacher J. An investigation of the ability of the glutaraldehyde test to distinguish between acute and chronic inflammatory disease in horses. Acta Vet Scand 2005;46:69–78. LeBlanc MM, Tran T, Baldwin JL, Pritchard EL. Factors that influence passive transfer of immunoglobulins in foals. J Am Vet Med Assoc 1992;200:179–83. Winger K, Gay CC, Besser TE. Immunoglobulin G1 transfer into induced mammary secretions: the effect of dexamethasone. J Dairy Sci 1995;78:1306–9. LeBlanc MM. Immunologic considerations. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 275–94. Fennell LC, Anderson GA, Savage CJ, McKinnon AO. Use of colostrum banking and immunoglobulin status of foals on a commercial Thoroughbred stud farm. J Am Vet Med Assoc 2010;236:1085–90. Raidal SL, McTaggart C, Penhale J. Effect of withholding macromolecules on the duration of intestinal permeability to colostral IgG in foals. Aust Vet J 2005;83:78–81. Staley TE, Jones EW, Corley LD. Attachment and penetration of Escherichia coli into intestinal epithelium of the ileum in newborn pigs. Am J Pathol 1969;56:371–92.
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15. Morris DD, Meirs DA, Merryman GS. Passive transfer failure in horses: incidence and causative factors on a breeding farm. Am J Vet Res 1985;46:2294–99. 16. Kohn CW, Knight D, Hueston W, Jacobs R, Reed SM. Colostral and serum IgG, IgA, and IgM concentrations in Standardbred mares and their foals at parturition. J Am Vet Med Assoc 1989;195:64–8. 17. Lawler JB, Hassel DM, Magnuson RJ, Hill AE, McCue PM, Traub-Dargatz JL. Adsorptive effects of di-tri-octahedral smectite on Clostridium perfringens alpha, beta, and beta2 exotoxins and equine colostral antibodies. Am J Vet Res 2008;69:233–9. 18. Clabough DL, Levine JF, Grant GL, Conboy HS. Factors associated with failure of passive transfer of colostral antibodies in Standardbred foals. J Vet Intern Med 1991;5:335–40. 19. Stoneham SJ, Digby NJ, Ricketts SW. Failure of passive transfer of colostral immunity in the foal: incidence, and the effect of stud management and plasma transfusions. Vet Rec 1991;128:416–9. 20. Robinson JA, Allen GK, Green EM, Fales WH, Loch WE, Wilkerson CG. A prospective study of septicaemia in colostrum-deprived foals. Equine Vet J 1993;25:214–9. 21. Baldwin JL, Cooper WL, Vanderwall DK, Erb HN. Prevalence (treatment days) and severity of illness in hypogammaglobulinemic and normogammaglobulinemic foals. J Am Vet Med Assoc 1991;198:423–8. 22. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Absorption of bovine colostral immunoglobulins G and M in newborn foals. Am J Vet Res 1989;50:1598–603. 23. Burton SC, Hintz HF, Kemen MJ, Holmes DF. Lyophilized hyperimmune equine serum as a source of antibodies for neonatal foals. Am J Vet Res 1981;42:308–10. 24. Madsen FC, Stout RC. Egg protein in complexes for neonatal foal diarrhea. J Equine Vet Sci 2000;20:158–60. 25. Bennett-Steward K. Neonatal foals and hyperimmunized egg protein – Viable therapy? J Equine Vet Sci 2002;22:105–7.
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Chapter 34
Assessment and Modification of Passive Transfer Sally L. Vivrette
Introduction The immune system of the equine fetus is fairly well developed in late term but, due to the absence of antigenic stimulation, little antibody production occurs before birth.1 The diffuse epitheliochorial placenta of the horse does not allow for transfer of macromolecules, such as immunoglobulins, from the dam to the fetus. Therefore, at birth foals are basically agammaglobulinemic, although measurable quantities of IgM and trace quantities of IgG may be present in blood samples taken from the foal before colostrum ingestion. Although newborn foals are immunocompetent at birth, they require 10–14 days to mount an effective and specific response to antigens. The ability of the newborn foal’s small intestine to absorb macromolecules such as immunoglobulins is maximal early and decreases linearly over the first 24 to 36 hours of age. It is generally thought that foals should receive colostrum within 7 hours of birth to optimize absorption and health in the neonatal period.2 The level of intake, and the consequent serum IgG concentration that is protective, has not been determined for foals, and probably varies with the level of challenge (dose and pathogenicity of pathogen) the foal experiences in its environment. Foals that undergo an uncomplicated delivery and nurse within 2 hours may be tested for adequacy of passive transfer as early as 8 to 12 hours of age,3 or 6 to 8 hours after the foal first nurses. Early measurement of IgG levels allows for identification of failure of passive transfer (FPT) or partial failure of passive transfer (PFPT), and institution of appropriate treatment plans. A common recommendation is to achieve a minimum serum IgG concentration of 800 mg/dL. In foals 18–24 hours of age and older, FPT is defined as serum IgG concentrations
< 200 mg/dL (2 g/L) and PFPT as IgG concentrations between 200 and 800 mg/dL (4–8 g/L).4 Blood samples taken before 18 hours of life may show a lower IgG concentration since gastrointestinal absorption and lymphatic transport of immunoglobulin are ongoing.2 In these cases, a minimal IgG concentration of 400 mg/dL (4 g/L) is required to predict adequacy of passive transfer. Foals with suboptimal IgG levels prior to 18 hours of life should be retested at 24 hours of age to determine the need for immunoglobulin supplementation. For healthy foals housed in clean environments where presence of pathogenic bacteria is minimal, IgG concentrations in foals > 24 hours old of 400 mg/dL (4 g/L) may be sufficient.5 Normal foals have serum IgG levels of 1400 mg/dL ± 100 mg/dL.6 Serum IgG levels are used as an indicator of protection against disease, but the specificity of the immunoglobulin against individual pathogens in the foal’s environment or efficacy of protection in the gut lumen cannot be evaluated. Thus, IgG in the foal’s blood is used as an indicator only of the ability of the foal to resist disease in the neonatal period. In most cases, it is not possible to identify a foal with FPT or PFPT solely by observation or physical examination in the early neonatal period. Even when mares lose colostrum from premature lactation, it is not possible to determine by direct observation the quality and quantity of colostrum still able to be delivered via ingestion to the foal. Overt illness associated with inadequate colostrum ingestion may not be observed until the foal is 2–3 days of age, at which point treatment may be costly and the outcome possibly fatal. Owners of newborn foals are sometimes reluctant to have newborn foals examined because the foals may appear clinically normal and there is desire to keep expenses down. Known risk factors for FPT and PFPT include dystocia,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Assessment and Modification of Passive Transfer inability to stand and nurse within 6 hours of birth, or leakage of colostrum by the mare prior to foaling. However, foals that undergo uncomplicated delivery, stand and nurse in a timely fashion, and appear clinically normal at 12–24 h of age may have dangerously low immunoglobulin levels secondary to inadequate colostrum ingestion, incomplete intestinal immunoglobulin absorption, or following ingestion of colostrum with low immunoglobulin concentrations.3
Incidence of failure of passive transfer The incidence of FPT in different parts of the world is 10–18%.7 It is well established that FPT and PFPT are risk factors for the development of infections in neonatal foals.4 Inadequate colostral immunoglobulin intake is a significant risk factor for the development of bacterial infection by the foal. Foals experiencing FPT or PFPT have a 50% incidence of infectious disease in the first weeks of life.8 Such infections may result in septicemia, diarrhea, and joint and/or umbilical infections (see Chapters 16, 20, 50, 70 and 81). Treatment of these diseases may be very costly and, even with effective treatment, may result in the death of the foal.
Assessment Assessment of adequacy of passive transfer is accomplished through measurement of equine IgG in the foal’s blood, plasma or serum. Semi-quantitative test kits for determining blood or serum IgG concentrations are commercially available for use in the field, farm or veterinary clinic. The glutaraldehyde clot test (Gamma Check E, Plasvacc, Templeton, CA) is a semi-quantitative test using the foal’s whole blood or serum. The test is based on the principle that glutaraldehyde reacts with certain blood proteins (gammaglobulin and fibrinogen) to form a solid gel. The time taken for the gel to form is inversely proportional to the concentration of these proteins. When testing the foal’s whole blood, the IgG level is determined to be > 800 mg/dL when a solid clot forms in less than 5 minutes. Using the test with whole blood that is hemolyzed, or from foals that may be septic (with elevated fibrinogen levels), may result in a false-positive reading. A false-negative result may occur when testing foals that are dehydrated or have a pathologically high packed cell volume. In these situations, testing using the foal’s serum may provide more accurate results. The commercially available glutaraldehyde clot test has the advantage of low cost, ease of use, and high sensitivity.9 A disadvantage of the test is inability to clearly quantitate IgG
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levels < 800 mg/dL. Non-commercial test kits for the glutaraldehyde clot test may be prepared in the laboratory. ‘Homemade’ kits may yield inconsistent and unreliable results, possibly owing to the instability of glutaraldehyde.10 Another semi-quantitative test, SNAPTM test (IDEXX Labs, Westbrook, ME), is an enzyme immunoassay that has calibrator spots with polyclonal antibodies to equine IgG that develop color corresponding to IgG levels of 400 and 800 mg/dL. Diluted whole blood, serum or plasma from the foal can be used and results are not affected by hemolysis. The sample and supplied reagents are applied onto the surface of the device membrane. Equine IgG, if present, is captured by the immobilized anti-IgG antibodies on the sample spot. Enzyme-conjugated polyclonal antibodies are added, and bind to the captured equine IgG, forming an antibody–IgG–antibody sandwich. After washing away unbound material from the membrane, an enzyme substrate solution is added. Subsequent color development is proportional to the concentration of equine IgG captured. Color intensity on the sample spot is compared to the reference spots to determine the foal’s IgG levels. The test has high sensitivity for IgG levels below 400 or above 800 mg/dL but is less accurate at clearly distinguishing levels between these two concentrations, possibly owing to variability of sample size with the sampling loop provided in the kit.9 The test kit for the SNAP test must be warmed to 7◦ -29◦ C (45◦ -85◦ F) prior to use, and results are available in under 10 minutes. This test has a moderate cost compared to other commercially available tests on the market. A commercial test for evaluating foals for FPT using the zinc sulfate turbidity test is available (EquiZ, VMRD, Pullman, WA). The test is based on the formation of a visible precipitate when immunoblobulin combines with zinc ions in solution. The test kit needs to be warmed to room temperature and uses the foal’s serum (cannot be hemolyzed) as the test sample. One hour after addition of the sample to the test vial, the vial is compared to a photograph to determine the approximate IgG content of the serum. The sample needs to be handled carefully because the precipitate is very fragile. Light precipitates may present a problem of interpretation. In these situations the test sample can be read spectrophotometrically against a standard curve, or by using a more sensitive test such as the radial immunodiffusion (RID, discussed below). The commercially available test has the advantage of low cost, a moderate turnaround time, and good sensitivity as a screening test for FPT. Reagents for the zinc sulfate precipitate test can be prepared in the laboratory.11 Care must be taken to minimize exposure of the solution to carbon
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dioxide as this causes a pH change that may affect the results of the test. The latex agglutination test, Foalchek, is commercially available (Centaur, Overland Park, KS). The test can be run on whole blood or serum. Assessment of adequacy of passive transfer is based on visualization of the degree of agglutination between latex particles and IgG. The test reportedly can be used to distinguish between 100, 200, 400, and 800 mg/dL. Accuracy of the test is dependent on precise measurement and delivery of consistent size drops of both latex and serum/blood diluents. Studies reporting the sensitivity and specificity of this test are not available and the test kit does not provide positive controls. A turbidimetric immunoassay (TIA) can be used for measurement of plasma IgG concentrations in foals.12 The test utilizes goat anti-equine IgG antiserum, which forms a precipitate with the sample IgG. After addition of the antiserum and a 10-minute incubation period, the percent of transmission (%T) of the foal’s sample is determined via spectrophometry. The IgG concentration of the sample is determined using a conversion chart provided with the test kit. Control samples of equine IgG are supplied with the test kit. Plasma or serum samples can be evaluated and results are not affected by ambient temperatures. Sensitivity is high (97.6%), indicating that this is a very good test for screening for FPT.9 The specificity of the test ranges has been shown to be 82.8%.9 The test is commercially available (DVM Rapid Test, DVM Stat) where the spectrophotometer and test kit reagents can be purchased. The TIA has also been adapted for use on automated chemistry analyzers.13 The single radial immunodiffusion (RID) assay is generally considered to be the gold standard for immunoglobulin quantification in foals. The foal’s serum is placed in a central well on an agarose gel containing antibodies against a specific antibody, in this case equine IgG. During a 16–24 h incubation period, the immunoglobulin in the serum diffuses outward from the well and precipitates with the antibody at the zone of equivalence. The diameter of the precipitin ring is read manually and its size is a function of the immunoglobulin concentration in the serum. These measurements are plotted against a standard curve derived from measurements of the reference sera provided with the assay kit. This test is temperature dependent, and requires a degree of technical skill. The tests have excellent sensitivity and specificity for evaluation of FPT in foals. Kits for RID are commercially available (VMRD, Pullman WA; Kent Laboratories, Bellingham WA; Plasvacc USA Inc, Templeton, CA). Refractometry of neonatal foal serum may be used to evaluate adequacy of passive transfer. Orig-
inally, it was thought that this method was unreliable11 but recent research indicates measurement of serum proteins may have some clinical use in foals. It was found that neonatal serum protein concentrations < 4.5 g/dL are likely to have IgG concentrations < 400 mg/dL. Serum protein concentrations of > 5.5 g/dL indicated the foals in the study were unlikely to have an IgG concentration < 400 mg/dL (sensitivity > 96%). Additionally, it was found that serum protein concentrations > 6.0 g/dL are likely to have IgG concentrations > 800 mg/dL (negative predictive value of 96.1%). Foals with serum protein concentrations between 4.5 and 5.5 mg/dL have unpredictable IgG concentrations using refractometry.9 The test chosen to evaluate adequacy of passive transfer may depend on the facility where the test will be run. The Gamma CheckE and SNAP test are especially convenient and economical for use by veterinary practitioners working out of their practice vehicle, or by experienced horse owners foaling out a small number of mares. The DVM Rapid Test and DVM Stat are very convenient, accurate, and economical for use on horse breeding farms or at veterinary clinics and hospitals. Use of this test requires an initial investment in a spectrophotometer that costs around US$1000. The zinc sulfate turbidity test may be useful on farms or in clinics, but the turnaround time to obtain results (1 hour) and fragile nature of the precipitate make it potentially less useful than the tests listed above. The RID can be used in clinic or hospital settings to verify semi-quantitative tests or to measure IgG in foals where fast results are not needed to implement treatment.
Treatment of failure of passive transfer The decision on method of treatment of FPT or PFPT involves consideration of age of recognition, degree of immunoglobulin deficiency, health of the foal, risk of disease (pathogens in the environment), and financial resources of the foal owner. If insufficient serum IgG levels are identified in a foal less than 24 hours of age, oral treatment with equine colostrum, bovine colostrum, or a colostrum substitute is recommended. Intravenous therapy may be used; however, oral therapy is often more convenient for equine practitioners and economical for foal owners. Oral administration of immunoglobulin also confers an added benefit of local protective effects from enteric pathogens. Oral therapy involves administration of colostrum or a colostrum substitute by nasogastric tube. Large volumes (e.g., ≥ 1 L) should be divided into aliquots no greater than 250–500 mL. It is not advised to syringe-feed or bottle-feed colostrum to a
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Assessment and Modification of Passive Transfer recumbent foal as this practice may put the foal at risk to develop aspiration pneumonia. Good-quality equine colostrum is the ideal therapy for failure of passive transfer in newborn foals less than 24 hours of age. Colostrum from the foal’s dam is ideal if the quantity and quality of the colostrum is optimal. An estimate of the IgG concentration should be made by using an alcohol or sugar/Brix refractometer14 (see Chapter 33) or by measuring the specific gravity of the colostrum using a colostrometer15 (Jorgensen Laboratories). For inclusion in the colostrum bank, the Brix reading should be > 25%14 or specific gravity should be at least 1.090.15 If colostrum from the foal’s dam is not available, colostrum may be obtained from a ‘colostrum bank’ maintained by some veterinary clinics and horse breeding farms. Banked colostrum is collected from healthy mares immediately after normal foaling (term delivery, no complications), before their foals have nursed. The colostrum can be stored frozen for up to 1–2 years.7 Ideally, the freezer would be frost-free to minimize cyclic unfreezing of the colostrum which may potentiate immunoglobulin degradation. Amounts of 250 mL are usually collected, filtered, labeled with the date, mare’s name, Brix refractometry value, or specific gravity obtained from a colostrometer. The colostrum is then frozen in 250 to 500 mL aliquots. Colostrum donor mares should be EIA-negative, and the colostrum should be free of anti-erythrocyte antibodies. This may require submitting an aliquot of the colostrum or serum sample from the mare to a diagnostic laboratory for screening. Reported ranges for colostrum production in normal mares vary from 2.3 ± 4.9 L3 to 5.1 ± 0.5 L.16 The mean concentration of IgG in colostrum at foaling is 11,324 ± 4119 mg/dL.3 The minimum quantity of colostrum that a foal must receive to prevent neonatal septicemia has not been determined, and depends on the IgG concentration and quantity of the colostrum ingested (see Chapter 33). Drawing on research performed in calves, it is recommended that a newborn foal should receive colostrum or colostrum substitute with a minimum of 70 g of immunoglobulin.17 Colostrum contains a wide variety of factors that provide local and systemic protection against infection for the newborn foal. Various constituents, including complement, lactoferrin, lactoferricin, lactoperoxidase oligosaccharides, and glycoconjugates, inhibit colonization in the intestine. Colostral leukocytes also have a local protective role.18, 19 In addition, colostrum contains growth factors and hormones which promote enterocyte maturation.19 Thus, in addition to providing immunoglobulins, colostrum contains factors that provide local protection and potentiate gut closure.20
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When equine colostrum is not available, alternate sources of immunoglobulin must be sought. A conR , is marcentrated equine serum product, Seramune keted as therapy for failure of passive transfer in R foals. Currently, a 300 mL bottle of Seramune contains approximately 30 g of IgG. Thus, a foal with FPT would require the administration of 2–3 bottles R by nasogastric intubation to facilitate of Seramune reaching a serum target level IgG level > 800 mg/dL. R Use of oral Seramune in field and hospital settings is convenient and relatively inexpensive. The oral adR ministration of a single 300 mL bottle of Seramune is also marketed for prophylactic administration to newborn foals as a method of prevention of FPT. Lyophilized equine immunoglobulins can be administered orally or intravenously to newborn foals.17 In the past, the product has been sold in 10 g doses for a few hundred dollars. Thus, it is likely cost-prohibitive for administration to foals with FPT where the administration of 70 g would cost a few thousand dollars. Bovine colostrum, collected immediately after calving, has been shown to elevate serum IgG concentrations of foals to 1350–3300 mg/dL following ingestion of 4 liters of first-milking bovine colostrum. The half-life of bovine IgG in the foal was 7–9 days compared to 26 days for IgG from equine colostrum;21, 22 endogenous production of IgG was evident by 28–42 days postpartum. There were no longterm health problems with foals that received bovine colostrum. Bovine colostrum can be collected and kept frozen for up to 3 years. Administration of blood, serum or plasma orally to the foal may be attempted as treatment for FPT or PFPT; however, prohibitively large amounts of these substances would be required to raise the foal’s serum IgG levels significantly. Many features may influence the absorption of IgG from colostrum in neonatal mammals. A very important factor is the timing of colostral intake with respect to intestinal permeability. During the first 24 hours of life the selective permeability of the small intestine to macromolecules such as immunoglobulins decreases. Consumption of glucose-containing electrolyte solution or milk replacer during the first 12 hours of life of foals does not appear to hasten gut closure.23 Foals that are greater than 18–24 hours of age may have decreased immunoglobulin absorption from the small intestine. In these cases, administration of immunoglobulin intravenously as treatment for FPT or PFPT is required. The veterinary practitioner may choose to supplement the foal with equine plasma or commercially available immunoglobulin product marketed for intravenous use. Various commercial plasma products are available including those
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with increased concentrations of immunoglobulins, or specific antibodies against pathogens potentially encountered by neonatal foals including E. coli and Salmonella. The American Association of Equine Practitioners (AAEP) has recently issued a White Paper providing information on equine plasma and serum products for the equine practitioner (2009; www.aaep.org/press room.php?term=2009&id=340). It is strongly recommended that veterinarians in the United States use equine plasma and plasma products from sources licensed with the USDA or FDA (CVM). In so doing, our industry is doing its best to ensure efficacy, safety and potency of these products for our equine patients. Equine plasma may also be collected from horses in a herd owned by the mare owner or veterinarian. The plasma donors should be screened for equine infectious anemia (EIA) and other infectious diseases, and determined to be free of anti-erythrocyte antibodies (commonly and erroneously termed ‘universal donor’). Blood from the donor can be collected into sterile vacuum-loaded glass bottles containing an anticoagulant such as anticoagulant citrate dextrose solution (ACD). The bottles are then stored in a refrigerator and after a few hours the red blood cells and plasma separate, at which time the plasma can be siphoned off and administered to the foal. This method is time consuming, steps to ensure sterility must be followed, and the plasma that is obtained contains red blood cells. The long-term consequence of this has not been determined. Alternately, the blood can be collected into blood collection bags, centrifuged, and the plasma administered to the foal. This requires specialized centrifuge machinery. In certain situations, a farm or veterinary clinic may have access to a plasmaphoresis machine where plasma can be collected from screened donors. For intravenous therapy, a jugular or cephalic catheter should be placed aseptically. Plasma should be thawed in lukewarm water. Frozen plasma that is thawed too quickly can result in a significant reduction in immunoglobulin quantities. Alternatively, commercially available immunoglobulin supplements designed for intravenous administration can be used. Plasma should be administered through a blood administration set with an in-line filter, and administered at a rate of approximately 1 L per 20 minutes to a 45 kg foal. Light sedation of the foal may facilitate intravenous catheter placement and cooperation during the plasma administration. Diazepam (0.05 to 0.44 mg/kg given intravenously slowly to effect)24 and butorphanol (0.01 to 0.04 mg/kg i.v.)24 may be administered for this purpose. Xylazine may also be used (0.15 to 0.7 mg/kg i.v.).25 Doses higher than this may potentiate bradycardia and hypoten-
sion, especially in sick foals. Use of a tuberculin syringe is recommended to facilitate accurate dosage. The foal should be monitored for transfusion reaction (trembling, tachycardia, tachypnea, piloerection, hives). If these signs are observed, the administration rate of the plasma can be slowed or stopped. Administration of flunixin meglumine prior to administration of a plasma product may be recommended by the plasma or plasma substitute manufacturer. An increase in serum IgG concentrations of 200 mg/dL is anticipated after administration of 1 liter of plasma.15 The volume of plasma to be delivered depends on amount of immunoglobulin in the plasma, the degree of failure of passive transfer, health of the foal, risk of disease, and financial resources of the owner. It is often difficult to administer sufficient plasma to achieve 800 mg/dL IgG concentrations in foals with FPT. For the most part, administration of 2 liters of plasma intravenously is considered as adequate therapy for FPT in foals that are not showing signs of illness. Foals that have bacterial infections with septicemia will often be in a catabolic state and will breakdown immunoglobulins non-specifically along with other proteins. These foals may require administration of multiple liters of plasma. After treatment for FPT or PFPT, the foal’s serum IgG concentration should be rechecked to assess adequacy of immunoglobulin therapy. The precise timeframe when the foal’s immunoglobulin level can be measured after oral colostrum or intravenous plasma supplementation has not been determined. Several hours are likely needed for absorption of immunoglobulins from the intestinal lumen, passage through the lymphatics, and distribution throughout the extracellular fluid space. It has been shown that it takes 2–3 days for radioactive IgG given intravenously to neonatal foals to equilibrate with the extracellular fluid space.26 It is probably reasonable to recheck the foal 6–8 hours after immunoglobulin therapy, realizing that the value may be spuriously high. Parenteral antibiotic treatment may be included in the treatment plan for foals with FPT or PFPT. All foals that have been diagnosed with inadequate immunoglobulin levels after birth should be watched closely for signs of sepsis, including lethargy, diarrhea, and failure to nurse vigorously.
Management to prevent FPT and PFPT Management strategies for prevention of FPT and PFPT initially involves determination of risk based on factors for decreased colostrum quantity or quality by the mare. This may include production of foals
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Assessment and Modification of Passive Transfer with FPT or PFPT in the past, or illness or death of a foal associated with an undiagnosed problem. In the current pregnancy, illness or surgery of the mare, presence of placentitis or premature lactation may signal problems with colostrum production by the mare or absorption by the foal. Risk factors of the foal for development of FPT or PFPT include delivery associated with dystocia, which may be associated with fetal trauma and asphyxia. In calves, prematurity and associated immaturity of the intestinal epithelium, as well as prolonged dystocia with neonatal asphyxia may have adverse effects on colostrum absorption.27 Any foal delivery that is unobserved should be considered as possibly associated with dystocia. Foals that have reduced vigor, are premature or dysmature, have congenital defects such as contracted tendons, or appear to have poor general health have increased risk for FPT or PFPT. A foal that is slow to stand and has not nursed within a few hours of birth should be evaluated and considered at risk for having FPT or PFPT. In many areas, mares that are grazing on fescue grass pastures or fed fescue hay are at risk for having prolonged gestation, dystocia, agalactia, and the delivery of weak foals. These complications are secondary to ergot alkaloids produced by the endophytic fungus Neotyphodium coenophialum28 (formerly Acremonium coenophialum). These factors would put the mare at significant risk to have inadequate colostrum production and the foal to have inadequate colostrum consumption and absorption. Management of mares exposed to fescue grass during late gestation may include removal from fescue pastures 30 days before the expected foaling date. This is often accomplished by confining the mare on a dry lot or allowing her access to pasture free of fescue grass.29 Alternately, the mare may be adminisR tered domperidone (Equidone , Equi-Tox, Central, SC) beginning 15 days prior to the expected foaling date.30 Compounded formulations of domperidone are not recommended as the efficacy of these formulations and accuracy of drug content are variable and potentially ineffective. At foaling, the quality of the mare’s colostrum can be evaluated with a Brix refractometer or colostrometer. If the colostral quality is low (RI < 20% or specific gravity < 1.060), the foal should be supplemented with colostrum from a colostrum bank or fed an alternate immunoglobulin supplement. Colostrum supplementation may also be used if the newborn foal is observed to have poor nursing behavior, either because it is weak or because their dam is non-compliant. This management practice has been shown to reduce the incidence of FPT and PFPT on a large breeding farm.7
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The practice of ensuring colostrum ingestion prior to 7 h of age in foals has been proposed by Madigan.2 In farm situations where illness in other foals has been observed early in the neonatal period, or following a difficult delivery, it may be advisable to administer colostrum by nasogastric tube prior to the foal standing after delivery. Even though the intestines may remain permeable to immunoglobulins for many hours after birth, feeding colostrum as soon as possible will minimize transepithelial migration of bacteria, such as Escherichia coli, as has been demonstrated in the neonatal calf prior to colostrum feeding.31 Prior to foaling, mare owners should be educated by their veterinarian about risks of FPT and PFPT. With this knowledge, they can minimize exposure to fescue, monitor mares for illness and premature lactation, and make plans to observe the foaling to identify and help with dystocia. Review of the normal timeline for the newborn foal to stand and nurse, as well as normal nursing behavior, should be discussed prior to foaling. Mare owners should be encouraged to budget for a post-foaling examination. Veterinary examination of the newborn foal, in addition to evaluation of adequacy of passive transfer, also allows for identification of problems such as birth trauma (rib fractures), navel abnormalities (patent urachus), congenital abnormalities (hernias, cataracts, cardiac anomalies), and orthopedic problems such as angular limb deformity that may be improved with farrier care during the first few months of life. Examination of the mare will allow for identification of problems such as rectovaginal tears, large intestine impactions or incomplete expulsion of the placenta, with resultant risk of metritis and laminitis.
References 1. Perryman LE. Immunological management of young foals. Comp Cont Educ 3:S223–S228, 1981. 2. Madigan JE. Method for preventing neonatal septicemia, the leading cause of death in the neonatal foal. In: Proceedings of 43rd Annual Convention of the American Association of Equine Practitioners, 1997; p. 17–19. 3. Massey RE, LeBlanc MM, Klapstein EF. Colostrum feeding of foals and colostrum banking. In: Proceedings of 37th Annual Convention of the American Association of Equine Practitioners, 1991; pp. 1–8. 4. Hines MT. Immunnodeficiencies of foals. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 4th edn. Philadelphia: Saunders, 1997; pp. 581–2. 5. McClure JJ, Parish SM, Hines MT. Immunologic disorders. In: Smith BP (ed.) Large Animal Internal Medicine. St Louis, MO: Mosby, 1996; p. 1848. 6. Leblanc MM, Tran TQ. Relationships among colostral electrolytes, colostral IgG concentrations and absorption of colostral IgG by foals. J Reprod Fertil Suppl 1987;35:735– 6.
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7. Nath LC, Anderson GA, Savage CJ, McKinnon AO. Use of stored equine colostrum for the treatment of foals perceived to be at risk or failure to transfer of passive immunity. J Am Vet Med Assoc 2010;235(10):1085–1090. 8. McGuire TC, Crawford TB, Hallowell AL, Macomber LE. Failure of colostral immunoglobulin transfer as an explanation for most infections and deaths of neonatal foals. J Am Vet Med Assoc 1977;170(11):1302–4. 9. Davis R, Giguere S. Evaluation of five commercially available assays and measurement of serum total protein concentration via refractometry for the diagnosis of failure of passive transfer of immunity in foals. J Am Vet Med Assoc 2005;227(10):1640–5. 10. Sedlinska M, Krejci J, Vyskocil M. Evaluation of field methods for determining immunoglobulins in suckling foals. Acta Vet Brno 2005;74:51–8. 11. Rumbaugh GE, Ardans AA, Ginno DG, TrommershausenSmith A. Measurement of neonatal equine immunoglobulins for assessment of colostral immunoglobulin transfer: Comparison of single radial immunodiffusion with the zinc sulfate turbidity test, serum electrophoresis, refractometry for total serum protein, and the sodium sulfite precipitation test. J Am Vet Med Assoc 172:321–325, 1978. 12. McCue PM. Evaluation of a turbidimetric immunoassay for measurement of plasma IgG concentration in foals. Am J Vet Res 2007;68:1005–9. 13. Davis DG, Schaefer DMW, Hinchcliff KW, Wellman ML, Willet VE, Fletcher JM. Measurement of serum IgG in foals by radial immunodiffusion and automated turbidimetric immunoassay. J Vet Intern Med 2005;19:93–6. 14. Chavatte P, Clement F, Cash R, Grongnet JF. Field determination of colostrum quality by using a novel, practical method. In: Proceedings of the 44th Annual Convention of the American Association of Equine Practitioners, 1998; pp. 206–9. 15. LeBlanc MM. Immunologic considerations. In: Kotera AM, Drummond WA, Kosch PC (eds) Equine Clinical Neonatology. Philadephia: Lea & Febiger, 1990; p. 280. 16. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Colostral volume and immunoglobulin G and M determination in mares. Am J Vet Res 1989;50:466–70. 17. Franz LC, Landon JC, Lopes LA, Marinho LA, Sarma C, Bruemmer J, Squires EL. Oral and intravenous immunoglobulin therapy in neonatal foals. J Equine Vet Sci 1998;18(11):742–8.
18. Reiter B. Review of nonspecific antimicrobial factors in colostrum. Ann Rech Vet 1978;9:205–24. 19. Ojofeitimi EO, Elegbe IA. The effect of early initiation of colostrum feeding on proliferation of intestinal bacteria in neonates. Clin Pediatr 1982;21:39–42. 20. Heird WC, Schwarz SM, Hansen IH. Colostrum-induced enteric mucosal growth in Beagle puppies. Pediatr Res 1984;18:512–15. 21. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Absorption of bovine colostral immunoglobulins G and M in newborn foals. Am J Vet Res 1989;50:1598–603. 22. Holmes MA, Lunn DP. A study of bovine and equine immunoglobulin levels in pony foals fed bovine colostrum. Equine Vet J 1991; 23 (2): 78–80. 23. Raidal SL, McTaggart C, Yovich JV, Penhale J. Effect of withholding macromolecules on the duration of intestinal permeability to colostral IgG in foals. In: Proceedings of 46th Annual Convention of the American Association of Equine Practitioners, 2000; pp. 260–3. 24. Brock KA. Sedation and anesthesia. In: Koterba AM, Drummond WA, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 653–62. 25. Madigan JE. Sedation and anesthesia. In: Madigan JE (ed.) Manual of Equine Neonatology. Woodland, CA: Live Oak Publishing, 1997; pp. 242–7. 26. MacDougall DF. Immunoglobulin metabolism in the neonatal foal. J Reprod Fertil Suppl 1975;23:739–42. 27. Aldridge B, Garry F, Adams R. Role of colostral transfer in neonatal calf management: Failure of acquisition of passive immunity. Compend Contin Educ Pract Vet 1992;14:265–70. 28. Cross DL. Fescue toxicosis in horses. In: Bacon CW, Hill NS (eds) Neotyphodium/Grass Interactions. New York: Plenum Press, 1997; pp. 289–309. 29. Brendemuehl JP. Fescue toxicosis. In: Smith BP (ed.) Large Animal Internal Medicine, 3rd edn. St Louis, MO: Mosby, 2002; pp. 221–2. 30. Cross DL, Anas K, Bridges WC, Chappell JH. Clinical effects of domperidone on fescue toxicosis in pregnant mares. In: Proceedings of 45th Annual Convention of the American Association of Equine Practitioners, 1999; pp. 203–6. 31. Corley LD, Staley TE, Bush LJ, Jones EW. Influence of colostrum on trnsepithelial movement of Escherichia coli 055. J Dairy Sci 1977;60:1416–21.
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Chapter 35
Neonatal Isoerythrolysis Jill R. Johnson
Introduction Neonatal isoerythrolysis (NI), also known as hemolytic disease of the newborn, is an immunogenetic disease characterized by the destruction of red blood cells in a newborn foal by preformed antibodies ingested in colostrum.1, 2 The offending antibodies are produced by the dam in response to previous exposure to ‘foreign’ red cell alloantigens as a result of exposure to incompatible fetal blood during a previous pregnancy or incompatible blood transfusion. Because there is normally no transplacental passage of antibodies from a mare to her foal in utero, the problem is not manifest during gestation.
Immunology and pathogenesis The immune response to the red cell alloantigens is similar to that directed toward any foreign antigen, the difference being that the antigen is a molecule on the surface of a red cell. Over 40 of these genetically controlled red blood cell alloantigens (also known as factors) have been identified in horses (Table 35.1). Each factor is associated with just one genetic system (locus). Seven systems have been well described, each identified by an upper case letter (e.g., A, C, D, etc.). Factors (antigens) are denoted by an upper case letter to denote the system and a lower case letter for the specific factor (e.g., Aa, Ca, Dg, etc.). Alleles within a system are denoted by a series of lower case letters that represent the combination of factors that are inherited as a group (e.g., Aadg, Abc, Pacd, Qac, etc.). Red cell alloantigens are inherited as co-dominant traits, so all the factors controlled by both of an individual’s alleles will appear on the red cells.2 As an example, the A system (locus) has seven associated factors (alloantigens) and 13 recognized alleles (each specifying different combinations of factors). Each different allele controls expression of 0–4 of the seven possible A system factors (Table 35.1).
Any individual has only two of the possible 13 alleles (one inherited from each parent), which means that horses in the population are not identical for the combination of factors on red cells. The frequency of different A alleles in the population varies by breed.3 A horse cannot be induced to produce antibodies against any factor it expresses on its own cells; however, if a horse has two alleles which do not include a particular factor, for example Aa, that horse would be inclined to produce anti-Aa antibody if exposed to blood from another horse that expresses the factor. Blood group incompatibilities during pregnancy (meaning that the mare lacks a factor that is present on the red cells of the sire and the foal) of at least some blood factors are the rule rather than the exception, and yet the prevalence of sensitization (immunization) of the dam and occurrence of NI in horses is relatively low (< 2%) overall.3, 4 While, in theory, any factor could stimulate antibody production against itself in a horse that lacks the factor, two commonly occuring alloantigens (Aa, Qa)4, 5 and another six or seven rarely occuring red cell alloantigens (Ab, Db, Dc, Dg, Pa, Qc, and Ua)6–9 have been associated with clinical disease. Mules present a special case in that mares bred to donkeys can produce antibodies against ‘donkey factor,’ a red cell antigen unique to donkeys and lacking in horses.10 Virtually 100% of mule pregnancies are incompatible for this factor, with estimates as high as 10% of mule foals affected to some degree.11 For NI to occur, several things must happen. First, the dam must be negative for the alloantigen (factor) in question. Second, the mare must somehow become sensitized and produce antibody to the offending antigen (factor). Third, the foal from the current pregnancy must have inherited from its sire the antigen(s) to which the mare has been sensitized. When these conditions are met, there is a significant potential that maternal antibody directed against the foal’s
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 35.1 Blood group systems, factors, and alleles of the horse recognized by the International Society for Animal Genetics System
Factors
Recognized alleles
A C D
a,∗ , b† ,c,d,e,f,g a a,b† ,c† ,d,e,f,g† ,h,i,k,l,m,n,o,p,q,r
K P Q U
a a† ,b,c,d a∗ , b,c† a†
Aa, Aadf, Aadg, Aabdf, Aabdg, Ab, Abc, Abce, Abe, Ac, Ace, Ae, A– Ca, C– Dadl, Dadlnr, Dadlr, Dbcmq, Dcefgmq, Dcegimnq, Dcfgkm, Dcfmqr, Dcgm, Dcdmp, Dcgmq, Dcgmqr, Ddeklqr, DeIno, Ddeloq, Ddelq, Ddfkir, Ddghmp, Ddghmq, Ddghmqr, Ddki, Ddlnq, Ddlnqr, Ddlqr, Ddno, Dq, (D–) Ka, K– Pa, Pac, Pacd, Pad, Pb, Pbd, Pd, P– Qa, Qab, Qabc, Qac, Qb, Qbc, Qc, Q– Ua, U–
From G. Cothran, personal communication. ∗ Most common factors involved in neonatal isoerythrolysis (NI). † Previously reported to cause NI in at least one case.
red cell antigens will appear in the colostrum and, if subsequently ingested and absorbed by the foal, could cause destruction of red cells.
Clinical signs Foals are born healthy and usually begin to develop signs of NI by 24–36 hours old, after suckling. Progressive lethargy and weakness are early signs. Breathing becomes rapid and shallow, followed by labored breathing as the disease progresses. The foals may yawn repeatedly. Heart rate is elevated. Seizurelike activity may occur as the anemia becomes more severe. In peracute cases, death may precede the development of icterus. In acute cases, mucous membranes show initial pallor that is followed by development of icterus. In severe cases, hemoglobinemia and hemoglobinuria may be pronounced. Pigment nephropathy may occur as a result of the severe hemolysis resulting in azotemia. Severe neurological signs characterized by altered mentation and seizures may occur as a result of kernicterus or bilirubin encephalopathy when the total bilirubin is significantly increased (e.g., bilirubin ≥ 25 mg/dL).12 Affected foals are anemic. All indicators of red cell concentration (packed cell volume (PCV), hemoglobin, red cell count) show significant de-
Box 35.1
creases. PCV values often decline into the teens, and values as low as 5% have been observed. Bilirubin (mainly unconjugated) levels will be increased as a result of accelerated red cell destruction. Total bilirubin levels may be close to 20 mg/dL in severe cases. Affected foals, especially mule foals, may also be thrombocytopenic.13 Demonstration of significant amounts of antibody in the colostrum (or serum from the mare) that are directed against the red cell antigens expressed by the foal provides a definitive diagnosis of NI. These antibodies are most commonly demonstrated by lytic and agglutinating tests. Lytic tests are believed to be more reliable indicators of the presence of offending antibody.5 The presence of antibodies attached to the foal’s red cells can also be demonstrated with a direct antiglobulin test (Coombs’ test). The presence of antibodies in the mare’s serum that attach to red cells can be demonstrated with an indirect antiglobulin test.1 Blood typing laboratories maintain panels of red cells representing all known blood groups and can identify antibodies in sera to the specific factors involved. Field screening tests for antibody, such as the jaundiced foal agglutination (JFA) test (Box 35.1) indicate that antibody is present, but do not identify the specific factor toward which the antibody is directed.
Jaundiced foal agglutination test14, 20
The jaundiced foal agglutination test (JFA) is used to detect the presence of antibody in colostrum directed against red cell antigens. Colostrum from the mare is reacted with presuckle red cells from the foal. If antibody against the red cells is present in the colostrum, an agglutination reaction occurs and can be visualized. The test is most useful when performed before the foal has suckled, to assess the risk associated with colostral ingestion. The test is also useful to monitor the decline in colostral antibody over time in order to determine when it is safe to allow the foal to nurse.
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If the test is performed after suckling, colostral antibody with anti-red cell specificity may already have been absorbed and have attached to the red cells of the foal. This will give a positive reaction in all samples, including the saline control (no colostrum added) and falsely elevate the titers.
Materials needed
r r r r r r r
Centrifuge Test tubes (e.g., 13 × 100 mm disposable tubes or blood collection tubes). Eight tubes are needed per sample tested Pipette system capable of delivering approximately 1.0 mL volumes (e.g., Pasteur pipettes and rubber bulbs) 0.9% NaCl for preparing dilutions Colostrum from the mare Presuckle blood from the foal, preferably in EDTA anticoagulant Optional: blood from the mare, preferably in EDTA anticoagulant, for use as a control for effects of colostral viscosity on agglutination
Method 1. Label the first tube as SALINE CONTROL and the remaining seven with the dilution (e.g., 1:2, 1:4, 1:8, 1:16, 1:32, 1:64, 1:128) Add approximately 1 mL 0.9% NaCl to each of eight tubes (note: prepare two sets of dilutions if the mare’s cells will be tested as a negative control for the effect of colostral viscosity) 2. Prepare serial dilutions of the colostrum by adding 1 mL of colostrum to the tube labeled 1:2, then transferring 1 mL of the mixture to the tube labeled 1:4, and so on until reaching the tube labeled 1: 128. Discard 1 mL from the tube labeled 1:128 so that the total volume in each tube is approximately 1 mL 3. Add one drop of well mixed anticoagulated whole blood from the foal to each tube of one series of diluted colostrum and one drop of anticoagulated blood from the mare to the other set of diluted colostrum. Mix the samples 4. Centrifuge the tubes for 2–3 minutes at medium speed (300–500 g). The centrifugation step is important to allow cells to contact each other to promote any tendency to agglutinate. Following centrifugation, the red cells will be pelleted in the bottom of the tubes 5. One at a time, pour out the liquid contents of each tube; observe the status of the button of red cells at the bottom of the tube. When no agglutination exists the red cells easily flow down the side of the tube (Figure 35.2). Complete agglutination (4+) causes the red cells to remain tightly packed in the button; strong agglutination (3+) causes the red cells to remain in large clumps. With weaker agglutination (1 + to 2+), the red cells remain in small clumps as they run down the side of the tube. The titer is defined as the highest dilution that gives strong agglutination (3+ or 4+)
Interpretation Positive reactions at 1:16 or greater in horses and 1:64 or greater in mules are considered significant. Colostrum should be withheld until the JFA titer drops below this level. Agglutination in the saline control tube most commonly occurs when using post-suckle blood from the foal and suggests that the foal has already absorbed antibodies. In this case, the test is not a valid assessment of the amount of antibody remaining in the colostrum. The sire’s blood may be substituted as a source of red cells for testing if presuckle blood from the foal is not available. At low colostral dilutions, the physical characteristics (viscosity) of colostrum may cause clumping of the red cells. Performing the test using the mare’s red cells will serve as a control for the effect of the viscosity of colostrum. If the mare’s cells clump, the reaction at similar dilutions with the foal cells cannot be interpreted as antibody-specific agglutination.
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Treatment During the first 24 hours of life when significant amounts of immunoglobulins are present in colostrum and the foal’s gut is capable of absorption, the foal should be prevented from nursing. In most cases, by the time the problem is recognized clinically (e.g., when the foal is about 24 hours old), the bulk of colostral antibody will have been depleted from the mare’s milk and the absorptive ability of the foal’s gut will have diminished. Withholding milk at this point is of questionable benefit. Stress should be minimized and exercise restricted. Affected foals have decreased exercise tolerance and can collapse and die if forced to follow their dams. Supportive care should be administered as indicated by clinical parameters. Intravenous fluids are frequently indicated to promote diuresis to minimize the effects of the large hemoglobin load presented to the kidneys and prevent pigment nephropathy (see Chapter 21). Acid–base balance should be monitored and corrected if indicated. Intranasal oxygen supplementation may be of benefit, although oxygen saturation is often close to normal. If the anemia becomes severe (e.g., PCV 10–15%), transfusions that provide red cells should be considered. Unless the PCV drops below this level, transfusion may not be necessary if rest is enforced. The object of transfusion is to provide the affected foal with red cells that will not be destroyed by the maternal alloantibodies that were absorbed from colostrum. The foal is immunologically naive and will not have autologous alloantibody directed against any red cells that would have an immediate effect on transfused red cells. Thus, the key is to select a red cell donor whose cells will not be destroyed by the maternal antibody derived from colostrum. Cross-matching, with particular attention to the reaction of mare sera, colostrum, and foal sera (all contain the same antibody) with the donor red cells, is important for selecting a cell that will not be destroyed by the maternal antibody. Washed red cells from the dam are obviously a perfect choice in terms of cells that will not react with the alloantibodies that have been absorbed by the foal. However, to avoid administering additional antibody that would be harmful to the foal’s cells, the mare’s sera containing antibody against the foal’s cells must be removed by washing before administration.1 Up to 6–8 liters of blood can be collected from the mare (assuming body weight of approximately 450 kg) and anticoagulated with acid citrate dextrose (ACD) or sodium citrate (3.8% sodium citrate solution; one part sodium citrate/nine parts blood), although 3–4 liters usually provides sufficient red cells. The preparation of large volumes of washed cells is
aided by a large-volume centrifuge, but the procedure can be accomplished by serial sedimentations. Anticoagulated blood from the mare is allowed to settle for 1–2 hours. The plasma is aseptically drawn off, a similar or greater volume of sterile isotonic saline (0.9% NaCl) is added to the red cells and mixed, and the red cells are again allowed to settle. The saline is then drawn off and discarded. At this point, the red cells can be resuspended in an equal volume of isotonic saline for administration, or the washing procedure can be repeated. The sedimentation method is less desirable than centrifugation because it is slower and does not remove as much of the offending antibody. The aim is to dilute any harmful antibody to insignificant levels. If it is not possible to use the dam’s red cells, alternative donors can be selected. The donor should lack the antigen to which the alloantibody is directed. Because it is generally not possible to blood type donors on short notice, a previously identified horse that has been determined by blood typing to be Aa-negative, Qa-negative and free of alloantibody is a reasonable choice for donor, based on the fact that most cases of NI are associated with these two antigens. Donors that are Aa- and Qa-negative are often referred to as ‘universal donors’; however, this is not an appropriate designation because only two out of more than 40 blood factors are considered. The odds of randomly selecting a donor that lacks the common offending factors (e.g., Aa and Qa), and that would therefore be a suitable donor, varies significantly with the breed. This can be roughly calculated based on gene frequencies of alleles lacking these factors and mirrors the percentage of the population of mares ‘at risk’ (Table 35.2).14 For example, the odds of finding an Aa-negative Thoroughbred or Standardbred trotter (without considering Qa status) to serve as a red cell donor would be approximately 1:50 (2% and 3%, respectively), whereas in Quarter Horses and Standardbred pacers the odds would be approximately 1:4 (25% and 22%, respectively). Table 35.2 Estimate of the percentage of mares in selected breeds that are ‘at risk’ for producing an NI foal based on the lack of all alleles that include factors Aa or Qa14 Breed
At risk for Aa
At risk for Qa
Thoroughbred Standardbred – pacer Standardbred – trotter Saddlebred Quarter Horse Arabian
2% 22% 3% 25% 25% 3%
16% ∗ ∗ 88% 68% 72%
∗
Mares all lack factor Qa and are technically ‘at risk,’ but all stallions in that breed also lack the factor.
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Neonatal Isoerythrolysis The odds of identifying a donor negative for both Aa and Qa (e.g., the so-called ‘universal donor’) become even less likely. For example, based on gene frequency estimates, only approximately 16% of the 2% of Aa-negative Thoroughbreds would also be Qa negative; this is equivalent to approximately three in 1000 horses! Correspondingly, the odds of finding an Aa- and Qa-negative Quarter Horse would be approximately 17 in 100. Because Standardbreds lack factor Qa entirely, only the frequency of Aa needs to be considered. The sire of the foal is not a suitable donor because he shares the same target red cell antigens as the foal, and his cells will react with the maternal alloantibody present, adding more of a load to the foal’s reticuloendothelial system as they are destroyed. In the case of mules, if the dam’s cells are used, the same considerations for washing the cells would be necessary. However, because the offending antibody is generally directed against a unique donkey antigen, it appears that red cells from any horse would be compatible (e.g., would not react with maternal antibody). Horses do not appear to make naturally occurring antibodies against donkey factor; therefore, in most cases, it is not necessary to wash the cells from a horse that would not be likely to have been immunized by pregnancy against donkey factor.10 Transfusing red cells that will not react with maternal alloantibody still means introducing a likely incompatible cell into the foal (owing to other factors). These cells will probably not survive for more than a few days in the circulation; thus, such transfusions should be considered temporary measures.15, 16 Transfused cells may also sensitize the foal to future transfusions, causing reactions (perhaps not within hours or a few days, but potentially within a week). This must be considered when considering the potential good versus potential harm of such transfusions. Administration of large volumes of blood (e.g., ≥ 4 liters) has been strongly associated with non-survival and the development of liver failure, perhaps as a result of iron overload.12 Approximately 1–4 liters of washed red cells or whole blood administered at a rate that does not produce volume overload is usually adequate to produce clinical improvement. One-half of the transfused volume can be given over 30–60 minutes followed by the remainder slowly over several hours. The PCVs of foals with NI commonly increase following transfusion and then gradually decline. This decline is probably not a cause for concern if it is gradual, because the PCV of the foal generally levels off as the offending maternal antibody is metabolized. Polymerized bovine hemoglobin has been administered to provide temporary oxygen-carrying support while awaiting blood transfusion.17, 18
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Complications and outcome The prognosis varies depending on the quantity of antibody ingested, the rapidity of onset of signs, and the degree of anemia. Peracute foals may die before the problem is recognized, with no chance for administration of therapy. Foals that develop the condition more slowly generally respond to supportive care. Liver failure, kernicterus, and complications related to bacterial sepsis are the most common complications associated with death or euthanasia.12
Strategies for preventive management Neonatal isoerythrolysis is a preventable disease. When a mare is deemed to be ‘at risk’ for production of an NI foal, a suggested plan for management is in Box 35.2. Mares considered ‘at risk’ to produce an NI foal can be identified by red cell typing based on the absence of factors Aa and Qa.14 The blood type is simply an inventory of which red cell antigens are present or absent on an individual’s cells. The Internet is a good tool to locate laboratories providing equine blood typing services. To determine whether a mare is ‘at risk’ based on a blood typing report, the factors present in the A and Q systems are examined. If factor ‘a’ (Aa) is not listed under the A system, or factor ‘a’ (Qa) is not listed under the Q system on the blood typing report, the mare would be considered ‘at risk’ for producing antibodies against the missing factor(s) (Figure 35.1). Factors that only sporadically cause NI can probably best be managed through postpartum presuckle testing rather than blood typing. Lack of these or other factors in a mare does not guarantee that NI will actually occur because of the sequence of events that must take place in addition to the mare simply lacking a factor. It can, however, be evaluated prior to breeding, which suggests additional vigilance in the management of the mare. Usually, a mare is recognized to be at risk because of production of an icteric foal. These mares should be tested for the presence of anti-red cell alloantibody within a few weeks after delivery of the icteric foal. Antibodies against red cell factors may wane quickly after parturition, so delay in testing may give a false-negative result.19 If anti-red cell antibodies are identified postpartum, management of subsequent pregnancies is warranted if the mare remains in the breeding herd. If a mare is determined to be ‘at risk’ by blood typing or history, one option for management includes mating to a stallion that also lacks the offending factor(s) so that no potential for immunization during the pregnancy or inheritance of the problematic factor by the foal exists. However, blood typing
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Box 35.2
Checklist for prevention of neonatal isoerythrolysis21
1. 0 months gestation a. Record breeding date b. Calculate 330 days from last breeding date 2. Before 10 months gestation a. Identify a ‘safe’ source of colostrum (1.5–2 liters) i. Colostrum bank ii. Establish local farm equine colostrum bank iii. Bovine colostrum bank b. Identify an alternate source of immunoglobulins in case of failure of passive transfer i. Commercial plasma/serum sources ii. Local plasma bank 3. 10 months gestation a. Obtain milk calcium test kits b. Begin to check for mammary development and presence of secretions c. Once secretions are detected, begin performing daily ‘milk’ calcium measurement d. Assemble JFA test equipment including tubes, pipettes, 0.9% NaCl and centrifuge 4. When milk calcium > 400 ppm a. Check for cervical relaxation b. Check number of days gestation c. If cervix relaxed and > 330 days gestation, induce labor or observe for natural parturition (generally within 24 hours) d. Obtain EDTA-anticoagulated blood sample and colostrum sample from dam prior to induction 5. Within 10 minutes after foal is delivered a. Collect an EDTA-anticoagulated blood sample from the foal prior to suckling b. Perform JFA test using foal’s and dam’s red cells and the dam’s colostrum Note: Save the foal’s presuckle blood sample for later testing of colostrum for decline in antibodies. 6. If JFA positive with foal red cells at ≥ 1:16 in horse foals or ≥ 1:64 in mule foals a. Muzzle foal and provide an alternate source of safe colostrum per os or via nasogastric tube b. Check adequacy of passive transfer at 18 hours. If < 400 mg/dL IgG, consider plasma transfusion c. Milk out mare hourly. Test colostrum for anti-red cell antibody using JFA test. Discard colostrum with anti-red cell antibody activity d. When JFA test on colostrum (using foal’s presuckle blood) < 1:16 (horse foal) or < 1:64 (mule foal), return the foal to nurse the dam 7. If JFA < 1:16 in horse foals or < 1:64 in mule foals a. Allow foal to nurse b. Check adequacy of passive transfer at 18 hours
information is seldom readily available for stallions. Simply changing stallions without examining the blood type may not alleviate future problems since, in general, more individuals in the population will possess the offending antigens than not based on estimates from gene frequencies. Another option for managing ‘at risk’ mares is screening the mare for evidence of development of offending antibodies prior to parturition and intervening to prevent colostral ingestion by the foal if sensitization has occurred.4 Screening consists of testing of the mare’s serum for presence of anti-red cell antibodies by a blood typing laboratory during
the last month or so of gestation. If such antibodies are detected, particularly if the antibody is directed against specific factors Aa or Qa, then plans should be made to intervene to prevent ingestion of the mare’s colostrum by the foal. Horses that lack factor Ca (Ca-negative) in the C system commonly make anti-Ca antibody even without known red cell exposure, presumably because of exposure to cross-reacting antigens in the diet. However, anti-Ca antibody is generally not associated with NI; indeed, mares with anti-Ca antibody appear less likely to develop antibodies to other factors responsible for NI.19 This may occur because
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(a)
(b) Figure 35.1 Jaundiced foal agglutination test. (a) The pelleted red blood cells flow smoothly down the side of the tube in negative samples (no offending antibody) when the tube is inverted. (b) In strongly positive samples (offending antibody present), the red blood cell pellet remains in the bottom of the tube when the tube is inverted.
foreign cells with Ca are removed more rapidly because the anti-Ca antibody prevents immunization against other factors on the cells. Presuckle screening of colostrum can also be carried out immediately postpartum before allowing the foal to nurse by determining whether colostral antibodies are present which react specifically with the BLOOD GROUP MARKERS A : abcdefg Type : –bc– –
Type :
K:a –
foal’s cells. Attendance at birth is essential in order to intervene prior to ingestion by the foal of colostrum containing the offending antibody. This might involve waiting for natural birth to occur, or inducing parturition when fetal maturity is appropriate (see Chapter 232). Attendance at birth can be facilitated by measuring milk electrolytes (calcium and
System : factors C:a a P : abcd ––––
Q : abc abc
D : abcdefghiklmnopqrs –bcd–f– – k m– – U: a
a
Figure 35.2 An example of how blood group results are displayed on some blood typing reports. In this example, the system and the recognized factors are listed on the top line. On the line immediately beneath, the results of the tests for this individual horse are given. A lowercase letter indicates the factor was detected, a ‘–’ (minus) indicates the horse was tested and was negative for the factor. A blank space indicates that the horse was not tested for the factor. For the purpose of assessing risk for NI, this horse is negative for factor Aa and positive for factor Qa.
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magnesium) to predict parturition (see Chapters 232 and 286). A sharp rise in milk calcium signals readiness for birth and can prompt more vigilant watching, or indicate that fetal maturity is sufficient to allow successful induction (see Chapters 12, 232 and 286). At birth, the colostrum can be tested for antibody activity against the foal’s red blood cells and withheld if positive. An alternative source of immunoglobulins can be provided from either another source of colostrum or plasma transfusions (see Chapters 33 and 34). Testing of colostrum within the time between birth and when the foal stands and nurses can be done using the JFA test (Box 35.1). If the JFA test using colostrum and foal’s red cells is positive, an alternate source of colostrum must be provided. JFA titers of 1:16 or greater for horse foals or 1:64 or greater for mule foals are considered positive. If the JFA is negative, it is assumed that it is safe to allow the foal to nurse. In the absence of the availability of perinatal colostral testing, it is best to assume that, if a mare has produced an NI-affected foal in a previous pregnancy, she will do so again and to provide an alternative source of colostrum. Offending colostral antibodies can potentially be detected in advance of parturition by testing late term mammary secretions against the sire’s red blood cells in the JFA test if his blood is available. The presence of antibody would reinforce the need to prepare for management of possible NI; however, because the immunoglobulin content of colostrum increases dramatically near parturition, the failure to detect antibody does not eliminate the possibility of NI. If the colostrum is initially positive for antibody, the JFA test can be helpful in determining when the titer has declined to levels at which it is safe for the foal to begin nursing the mare. Colostral titers of anti-red cell antibody may drop precipitously and rather than arbitrarily withholding colostrum for 24–72 hours, which requires significant manpower to manage the feeding of the foal and limits the absorption of other beneficial things found in colostrum, testing colostrum may confirm that it is safe for the foal to nurse in as few as 6–8 hours (J.R. Johnson, unpublished data). When colostral titers in the JFA test decline to <1:16 in horses and <1:64 in mules, it is safe to allow the foal to nurse. Although the disease is more common in multiparous mares owing to pregnancy-associated exposure to fetal red cells, NI can occur with the first pregnancy, so the diagnosis should not be excluded simply because the mare is primiparous.
References 1. Scott AM, Jeffcott LB. Haemolytic disease of the newborn foal. Equine Vet J 1978;103:71–4.
2. Stormont C. Neonatal isoerythrolysis in domestic animals: a comparative review. Adv Vet Sci Comp Med 1975;19:23–45. 3. Bowling AT, Clark RS. Blood group and protein polymorphism gene frequencies for seven breeds of horses in the United States. Anim Blood Groups Biochem Genet 1985;16:93–108. 4. Bailey E. Prevalence of anti-red blood cell antibodies in the serum and colostrum of mares and its relationship to neonatal isoerythrolysis. Am J Vet Res 1982;43: 1917–21. 5. Becht JL, Page EH, Morter RL, Boon GD, Thacker HL. Evaluation of a series of testing procedures to predict neonatal isoerythrolysis in the foal. Cornell Vet 1983;73:390–402. 6. Boyle AG, Magdesian KG, Ruby RE. Neonatal isoerythrolysis in horse foals and a mule foal: 18 cases (1988–2003). J Am Vet Med Assoc 2005;227:1276–83. 7. MacLeay JM. Neonatal isoerythrolysis involving the Qc and Db antigens in a foal. J Am Vet Med Assoc 2001;219:79–81. 8. Trommershausen-Smith A, Stormont C, Suzuki Y. Alloantibodies: their role in equine neonatal isoerythrolysis. Int Symp Equine Hematol 1975;1:349–55. 9. Zaruby JF, Hearn P, Colling D. Neonatal isoerythrolysis in a foal, involving anti-Pa alloantibody. Equine Vet J 1992;24:71–3. 10. McClure JJ, Koch C, Traub-Dargatz J. Characterization of a red blood cell antigen in donkeys and mules associated with neonatal isoerythrolysis. Anim Genet 1994;25:119– 20. 11. Brumpt E. Considerations defavorables a l’hypothese de l’etiologie parasitaire de la jaunisse des muletons: jaundice in the newly born mule foals not caused by babesia infections. Ann Parasitol Hum Comp 1947;22:5–10. 12. Polkes AC, Giguere S, Lester GD, Bain FT. Factors associated with outcome in foals with neonatal isoerythrolysis (72 cases, 1988–2003). J Vet Intern Med 2008;22:1216–22. 13. Traub-Dargatz JL, McClure JJ, Koch C, Schlipf JW. Neonatal isoerythrolysis in mule foals. J Am Vet Med Assoc 1995;206:67–70. 14. Bailey E, Conboy HS, McCarthy PF. Neonatal isoerythrolysis of foals; an update on testing. In: Proceedings of the 33rd Annual Convention of the American Association of Equine Practitioners, 1987, pp. 341–53. 15. Kallfelz FA, Whitlock RH, Schultz RD. Survival of Felabeled erythrocytes in cross-transfused equine blood. Am J Vet Res 1978;39:617–20. 16. Smith JE, Dever M, Smith J, DeBowes RM. Post-transfusion survival of 50Cr-labeled erythrocytes in neonatal foals. J Vet Intern Med 1992;6:183–5. 17. Perkins GA, Divers TJ. Polymerized hemoglobin therapy in a foal with neonatal isoerythrolysis. J Vet Emerg Crit Care 2001;11:141–6. 18. Polkes AC. Neonatal isoerythrolysis: overview, management strategies, and long-term outcome. In: Proceedings of the 21st Meeting of the ACVIM Forum, 2003, pp. 248– 50. 19. Bailey E, Albright AG, Henney PJ. Equine neonatal isoerythrolysis: evidence for prevention by maternal antibodies to the Ca blood group antigen. Am J Vet Res 1988;49:1218– 22. 20. Blackmer JM, Costa LRR, Koch C. The jaundiced foal agglutination test. Vet Technician 2002;23:577–9. 21. Blackmer JM, Paccamonti D, Eilts B, Costa LRR, Koch C. Strategies for preventing neonatal isoerythrolysis. Compend Cont Educ Pract Vet 2002;24:562–9.
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Chapter 36
Immunodeficiencies in Foals M. Julia B. Felippe Flaminio
Immunodeficiency is a disorder of the immune system that results in failure to build protection against pathogens. Important clinical indicators of immunodeficiency are recurrent infections, failure to respond to treatment, and infection with opportunistic organisms. In many cases of immunodeficiency, initial clinical improvement is observed with antimicrobial therapy; however, signs of infection tend to recur. The most common sites of infection are those readily exposed to pathogens, including the lungs, intestinal tract, and skin. Immunodeficiencies are classified as primary or secondary. Primary immunodeficiencies involve a genetic defect, and are more likely to manifest at a young age, particularly when colostral antibody concentrations reach a nadir. The cause of many primary immunodeficiencies is unknown; some disorders may not have a clear pattern of inheritance, and may be found in isolated individuals or several individuals of a lineage. The prognosis for primary immunodeficiencies is poor. Secondary immunodeficiencies are acquired after immunosuppressive treatment (e.g., steroids), certain viral diseases, primary or secondary lymphoid tissue disruption (e.g., tumoral infiltration), stress or metabolic/endocrine conditions (e.g., hypercortisolemia), or malnutrition. Secondary immunodeficiencies can be transient or chronic. Immunodeficiencies can affect different elements of the immune system: (i) B-cells (humoral immunodeficiency); (ii) T-cells (cellular immunodeficiency); (iii) B- and T-cells at the same time (severe combined immunodeficiency); (iv) phagocytes; or (v) complement factors (Table 36.1).1–3 Independent of the cause, recurrent infections and fevers are common to all types of immunodeficiency. Nevertheless, the type(s) of organism(s) isolated from an immunodeficient patient may suggest which aspect of the immune system is affected because of the opportunism of disease. For example, encapsulated bacterial infections are as-
sociated with humoral deficiencies, whereas identification of intracellular pathogens suggests a cellular immune disorder. In equine neonates and weanlings, the clinical recognition of underlying primary immunodeficiency may not be obvious because of the common presentation of disease at this age, often involving failure of transfer of immunoglobulins through colostrum, and the physiological development of the immune system that predisposes to infections. Independent of the etiology, foals with immunodeficiency fail to gain weight and grow properly, present recurrent infections (pneumonia, diarrhea, osteomyelitis, meningitis) for several months, or infections with opportunistic organisms (e.g., Cryptosporidium parvum, Pneumocystis carinii, Candida spp., adenovirus).4, 5 In addition, some types of immunodeficiencies are self-limited (e.g., transient hypogammaglobulinemia of the young, transient CD4+ T-cell lymphopenia of the young), and supportive therapy, including antibiotics and intravenous plasma transfusion, may control infections during the period of immunological instability.
Humoral immunodeficiencies Impaired antibody production is the most frequently diagnosed immunodeficiency in mammals. Humoral disorders described in the foal include failure of passive transfer of immunoglobulins (see Chapter 34), transient hypogammaglobulinemia of the young, agammaglobulinemia, and selective IgM deficiency. Common variable immunodeficiency has not been described in foals to date; hence, it will be briefly discussed in this chapter, although its manifestation in young life is still possible. The Fell Pony immunodeficiency syndrome presents impaired humoral response and questionable in vivo T-cell function. Severe combined immunodeficiency, which includes
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
transient condition
< 3 weeks
1–3 months
1–3 months (only males)
total or partial failure of transfer of colostral immunoglobulins
transient hypogammaglobulinemia of the young
agammaglobulinemia
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fatal (< 2 months)
2–5 months
< 2 months
transient CD4+ T cell lymphopenia of the young
severe combined immunodeficiency
Arabian or Arabian crossbreeds
all
Fell Pony
all
values may be normal at clinical onset due to colostral-derived immunoglobulins up to 2 months of life.
fatal (< 2 months)
< 2 months
Fell Pony Syndrome
∗
poor prognosis or transient condition
2–10 months
juvenille selective IgM deficiency
Thoroughbred Standardbred Quarter Horse present functional
present functional
present functional
present functional
present functional
low to undetectable (< 25 mg/dL)
undetectable (< 25 mg/dL)
undetectable (< 25 mg/dL)
low to normal
low to normal
serum IgM
absent no response
undetectable (< 25 mg/dL)
low CD4+ T cells normal low CD4:CD8 ratio
present at birth present absent by 5 questionable weeks no function response
severe lymphopenia absent no response
normal to mild lymphopenia
moderate lymphopenia
present poor response
absent no response
normal
normal
present poor response
present functional
normal
normal
T cell distribution
serum IgA
normal
undetectable
normal∗ to undetectable
low to undetectable
normal∗ to undetectable
normal
low to normal
undetectable
low∗ to undetectable normal
low to normal
low∗
very low (<800 low mg/dL)
serum IgG
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all
all
affected breed
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Table 36.1 Immunodeficiencies described in the foal
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Immunodeficiencies in Foals a humoral component, is discussed elsewhere (see Chapter 37). In humoral immunodeficiencies, B-cells either fail to develop (B-cell lymphopenia) or differentiate into antibody-secreting plasma cells (presence of normal or subnormal numbers of B-cells). In both cases, there is hypo- or agammaglobulinemia, which affects all or certain immunoglobulin isotypes. The limitation in the production of immunoglobulins favors recurrent, severe infections caused by opportunistic or encapsulated microorganisms. Transient hypogammaglobulinemia of the young Transient hypogammaglobulinemia results from a delay in immunoglobulin (Ig) G production by the foal.6 Healthy foals produce antigen-specific antibodies immediately after birth, reaching endogenous antibody protective levels (> 400–500 mg/dL) by 5–8 weeks of life. With delayed endogenous IgG production, the foal is at risk of bacterial infection and poor development, particularly when colostral IgG concentrations decrease to levels below protection (< 400 mg/dL). Therefore, foals with transient hypogammaglobulinemia may be diagnosed with this condition by 2–4 months old, depending on the persistence of protective colostral immunoglobulin concentrations (colostral IgG half-life is 28–32 days; see Chapter 32 Development of the Foal Immune System). In general, this condition resolves by 5–6 months old, and clinical signs of infection improve or resolve. Although reported in Arabian foals, transient hypogammaglobulinemia occurs in other horse breeds (M.J.B.F. Flaminio, personal observation). This condition seems more frequent than reported owing to the lack of routine measurement of serum immunoglobulin concentrations in foals between 4 weeks and 6 months old. Therefore, in foals with recurrent respiratory infections at this age, the quantitation of serum immunoglobulin concentrations is advised. In these foals, peripheral blood B- and Tlymphocyte counts and distribution, and T-cell proliferation response to mitogen in vivo (skin) and in vitro are normal.6 Serum IgG and IgG(T) concentrations are low, and serum IgM and IgA levels may be normal or decreased. In this context, serum IgM concentration > 50 mg/dL would suggest a transient condition rather than a primary B-cell dysfunction (e.g., agammaglobulinemia, Fell Pony syndrome, common variable immunodeficiency). Vaccination may elicit a weak humoral response during the transient period. Supportive therapy with antibiotics and intravenous plasma transfusions aid the affected foal during the hyporesponsive phase. An increase
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in serum IgG concentrations with time supports the transient condition, which is likely resultant from a delayed development of the immune system. The cause of this delay is unknown, and perhaps it involves the coordinated interaction and activation of antigen-presenting cells and lymphocytes in secondary lymphoid tissues, for the proper expansion and function of the acquired immune system. Agammaglobulinemia Equine agammaglobulinemia is a rare and fatal primary disorder caused by impaired B-cell development, leading to recurrent infections in young age and death by 1.5 years old.7 Peripheral blood lymphocyte counts and T-cell function are normal. The disease manifested in young male horses presents similar features to the X-linked agammaglobulinemia (XLA) described in young male human patients. Agammaglobulinemia was first described in a Thoroughbred colt with recurrent infections (pneumonia, rhinitis, and arthritis) starting at 5.5 months old, and managed with antibiotic therapy.8 At 13 months old, the colt presented IgM, IgA, and IgG(T) agammaglobulinemia; low serum IgG concentration (16 mg/dL); failure to respond with antibody production upon antigenic stimulation; and absence of B-cells, lymphoid follicles, and plasma cells in lymphoid tissues. Functional cell-mediated immunity (T-cell) was demonstrated by positive skin response to phytolectins and in vitro lymphocyte proliferation tests. Another publication reported a 1.5-yearold Standardbred horse with persistent fever and multifocal bacterial infections since early age.9 In addition to declining serum concentrations of IgG and IgG(T) (most likely colostral in origin), serum IgM and IgA were not detectable. The colt failed to produce antibodies upon in vivo humoral response test with foreign antigen inoculation. Peripheral blood lymphocyte response to mitogens in vitro was normal, as well as a cell-mediated skin reaction to intradermal phytolectin injection. No B-cells and plasma cells were found in histopathology, and secondary lymphoid tissues lacked germinal centers. Although the immunodeficiency may only be clinically detected around the time colostral antibody concentrations start to fall, these foals present with B-cell lymphopenia and lack of plasma cells already at birth, and fail to increase serum IgM, IgG, and IgA concentrations with time. In contrast to severe combined immunodeficiency (SCID), and potentially Fell Pony syndrome, colts with agammaglobulinemia can be clinically managed until they are yearlings, likely because of their functional T-cell response. In humans, the disease is caused by a mutation of the btk gene that encodes the Bruton tyrosine
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kinase. The BTK cytoplasmic protein sustains activation signaling and transcription in response to B-cell receptor engagement, and its dysfunction prevents Bcell differentiation. The human btk gene is located in the X chromosome; hence, the manifestation of the disease in males. The involvement of the btk gene in equine agammaglobulinemia has not been confirmed to date, which would be essential for the definitive diagnosis of this disorder. An important differential diagnosis is common variable immunodeficiency (CVID). Although not diagnosed in foals yet, CVID in horses manifests in both males and females with recurrent bacterial infections, hypo- or agammaglobulinemia (particularly IgG and IgM, but also IgA), inappropriate response to tetanus toxoid vaccination, severe B-cell lymphopenia, and lack of germinal centers and plasma cells in lymphoid tissues and intestine.10 Common variable immunodeficiency seems to be a progressive disease (hence its late-onset manifestation in adult horses), in which B-cells may be present and functional in the initial phase of life or for several years after birth, but B-cell differentiation in the bone marrow and population of secondary lymphoid tissues become impaired, resulting in decreasing values of serum IgG, IgM, and IgA concentrations. To date, the etiology of common variable immunodeficiency is unknown.
cause those conditions often cause IgM hypogammaglobulinemia. Caution should be used when interpreting low serum IgM concentrations, particularly when serum IgG and IgA are within normal values; persistent serum IgM concentrations < 25 mg/dL better define this humoral dysfunction.17 Serum IgM concentrations are quite specific for B-cell function in foals, as IgM production occurs already in utero and colostral IgM has a short halflife of 5–16 days.18–20 Serum IgM and IgG concentrations should be monitored to evaluate for persistence or change in values with age. When serum IgM concentrations < 25 mg/dL are accompanied by IgG hypogammaglobulinemia and B-cell lymphopenia, other types of humoral immunodeficiency should be considered (e.g., agammaglobulinemia, common variable immunodeficiency). It is intriguing that serum IgM concentrations can be selectively low while serum IgG concentrations are normal because IgM is the initial response elicited from mature naive B-cells (which leave the bone marrow expressing IgM on the cell surface) after antigen encounter in lymph nodes; the subsequent IgG response requires robust co-stimulatory interaction with CD4+ T-cells and immunoglobulin isotype switching. Therefore, the fact that IgG is produced but not its predecessor IgM raises important questions about the mechanism of disease.
Juvenile selective IgM deficiency Selective IgM deficiency has been diagnosed in foals with chronic infections when serum IgM concentrations are more than 2 standard deviations below the normal mean, and IgG and IgA are within normal reference range for the age group.11 This condition has been reported in Arabian, Quarter Horse, and Standardbred horses with at least a 2-month history of recurrent fevers, bronchopneumonia, arthritis, enteritis, dermatitis, generalized lymph node hyperplasia (sometimes with granulomatous inflammatory response), and poor growth.12–15 Peripheral blood lymphocyte counts, B- and T-cell distribution, and response to mitogens in vitro are normal, although response to lipopolysaccharide may be decreased.16 The onset of clinical signs occurs between 2 and 10 months old, and both males and females are affected. Microorganisms isolated from the respiratory tract of affected foals include Staphylococcus aureus, Streptococcus zooepidemicus, Actinobacillus equuli, Klebsiella spp., and Escherichia coli. In foals, chronic infections leading to euthanasia have been reported, although transient conditions are possible. A genetic basis has not been identified, but two cases reported had a common sire and great-grandsire.11 Lymphoma or lymphosarcoma should be investigated using lymph node biopsy and histology be-
Fell Pony syndrome Fell Pony syndrome is a fatal primary immunodeficiency characterized by profound anemia, absolute lymphopenia, B-cell lymphopenia, and severe infections in foals less than 2 months old.21 The affected foals of the Fell pony breed are clinically normal at birth, but signs develop gradually thereafter. The foals present with lethargy, rough haircoat, severe diarrhea, pancreatitis or bronchopneumonia often caused by opportunistic organisms (e.g., C. parvum, adenovirus), frequent chewing, and ulcerative pseudomembranous coating of the tongue with Candida spp. infection.22 Although foals may be born with an adequate hematocrit, death occurs owing to rapid development of non-regenerative anemia and sepsis.23, 24 The red cells and distribution of B-cells in peripheral blood at birth in affected foals may be comparable to healthy foals.24 Nevertheless, progressive B-cell lymphopenia develops within weeks after birth. Low serum IgM concentrations (< 25 mg/dL) indicate a poor primary humoral response at an early age, and are supportive of a diagnosis of humoral immunodeficiency involving B-cell lymphopenia.25 The serum IgG concentrations may be above protective levels at the age of clinical signs, for they still
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Immunodeficiencies in Foals reflect colostral antibodies. Histopathological studies indicate paucity of B-lymphocytes in the primary and secondary lymphoid tissues, suggesting impaired B-cell development. Lymphoid follicles are absent in lymph nodes, spleen, and gut-associated lymphoid tissues, and plasma cells are not found in the intestine. The thymus demonstrates atrophy, with no distinction between cortex and medulla. The percentages of peripheral blood CD4+ and CD8+ Tlymphocytes are normal, and these cells are responsive to mitogens in vitro.26, 27 Nevertheless, T-cell in vivo function is questionable based on the absolute lymphopenia, poorly developed thymus, paucity of lymphocytes in secondary lymphoid tissues, type of pathogens that cause disease, and the fact that affected foals manifest fatal clinical signs at an early age despite normal circulating colostral IgG. In addition, the expression of major histocompatibility complex class II (MHC class II) on lymphocytes (but not monocytes) fails to increase with age in affected foals.24, 28 The phagocytic function in these foals is normal. The bone marrow reveals severe red cell hypoplasia, and abundant hemosiderophages, megakaryocytes, and myeloid precursors.24 In contrast, numerous macrophages and neutrophils infiltrate the lymphoid tissues most likely as a response to infections. Peripheral ganglionopathy has also been reported in affected foals, characterized by neuronal chromatolysis with nuclear pyknosis in the trigeminal, cranial mesenteric, and dorsal root ganglia.21 Both genders are affected, and pedigree analysis suggests inheritance through an autosomal recessive trait (M. J. Gould-Early, personal communication). The prevalence of carriers in the breed is estimated to be high, and a genetic test is being studied for ante mortem diagnosis. If breeding between carriers could be prevented, the birth of affected foals and the incidence of the mutant gene in the population would both decrease.
Cellular immunodeficiencies Primary cellular immunodeficiencies are rare and more difficult to diagnose than humoral immunodeficiencies, for they may result from the dysfunction or expression of cell surface activation molecules, or their cell signaling components. Nevertheless, conditions that affect the distribution of T-cells in peripheral blood can be recognized using immunological testing. Secondary or transient depressed cellular immunity can be observed during periods of stress, including weaning.29 Clinical signs of recurrent infections and fevers are common to cellular and humoral immunodefi-
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ciencies. However, it is the type of organism causing the disease (e.g., intracellular pathogens including viruses, fungi, protozoa, and mycobacteria) that suggests the involvement of a faulty cellular immunity. Transient CD4+ T-cell lymphopenia of the young The normal foal presents a remarkable agedependent increase in the total peripheral blood lymphocyte counts and secondary lymphoid tissue weight, which reflects the intense lymphocyte priming and population expansion upon encounter with many pathogens (see Chapter 32). Concurrently, an increase in the antigenic-primed lymphocyte subpopulation distribution is expected.30 Occasionally, foals in the initial 6 months of life may demonstrate a delay in reaching normal reference values for CD4+ lymphocytes in peripheral blood, with consequent low CD4:CD8 ratios (< 2.0).31 In addition, failure to increase the expression of MHC class II in lymphocytes (but not monocytes) is often observed in this condition (M.J.B.F. Flaminio, personal observation). Lymphocyte proliferation response in vitro upon mitogenic stimulation, serum immunoglobulin concentrations, and B-cell distribution are normal in these foals. The decrease in the peripheral blood CD4+ Tcell subpopulation is often accompanied by absolute lymphopenia. It is important to take into consideration the physiological age-dependent increase in circulating lymphocytes, and the age-specific normal reference range when assessing normal lymphocyte counts in foals (i.e., expect foals older than 3 months to have two or three times higher lymphocyte counts than adult horses). Nevertheless, standard hematology alone may not identify the problem when total lymphocyte counts are appropriate for the age; then, lymphocyte immunophenotyping becomes necessary for the identification of low circulating CD4+ T-cells. In this case, an increase in circulating natural killer (NK) cells is suspected based on the presence of CD3-negative, CD4-negative, CD8negative, B-cell-negative circulating cells.30 It is possible that transitory CD4 lymphopenia reflects secondary lymphoid tissue activity and population expansion. Clinical signs of infection often accompany this transient condition, particularly chronic pneumonia caused by opportunistic organisms that require cellular immunity (e.g., P. carinii). Therefore, in foals with recurrent respiratory infections, peripheral blood lymphocyte subpopulation distribution testing is advised. Long-term antibiotic therapy (e.g., trimethoprim–sulfadiazine) may be necessary during
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this transition period. These foals should be monitored periodically with leukograms, thoracic ultrasonography and radiographs, and immunological testing to document improvement of clinical and immunological values and determine the need for treatment.
Phagocytic immunodeficiencies There has been no description of primary phagocytic dysfunction in the horse. In the neonate foal, acquired neutrophil dysfunction has been reported during sepsis (see Chapter 16). Although healthy foals are born with competent neutrophil function, newborn foals fighting severe infection demonstrate a transient decrease in phagocytosis and oxidative burst activity that improves during the hospitalization period.32 Pelger–Hu¨et anomaly of neutrophils has been identified in the horse (M.J.B.F. Flaminio, unpublished data; M.J. Wilkerson, personal communication). Pelger–Hu¨et cell neutrophilic function is normal, i.e., neutrophils capably phagocytose and kill microorganisms, and no clinical signs of immunodeficiency are observed. Therefore, the diagnosis is incidental, during the evaluation of blood smears. Pelger–Hu¨et cells present dumbbell-shaped bi-lobed nuclei, and a reduced number of nuclear segments. In humans, Pelger–Hu¨et anomaly is a benign dominantly inherited defect of terminal neutrophil differentiation secondary to mutations in the lamin B receptor (LBR) gene. Although inherited forms of neutrophil dysfunction have not yet been described in the horse, neutrophil function testing (expression of CD18 molecule, phagocytosis, oxidative burst activity) is indicated when clinical signs include recurrent dermatitis or cutaneous abscesses caused by opportunistic or encapsulate bacteria (Serratia spp., Streptococcus spp.), or fungi (Candida spp., Aspergillus spp.). Some of the diseases associated with phagocytic disorders described in other species include leukocyte adhesion deficiency (failure to express the CD18 integrin), chronic granulomatous disease (inability to produce oxygen reactive species), cyclic neutrophil hematopoiesis (cyclic changes in blood neutrophil), and Chediak–Higashi syndrome (giant granules in neutrophils).
Complement component immunodeficiencies Primary complement dysfunction or deficiency has not been reported in the horse. However, the foal
is born with low serum complement activity, and colostrum is not a significant source of complement (see Chapter 32). Therefore, foals are transiently deficient in serum opsonic capacity, and rely on their own production of complement after birth.33, 34 In sepsis, serum complement C3 components are rapidly consumed, which delays the physiological age-dependent increase that is normally observed in healthy foals.32
Immunological testing Immunological testing for the diagnosis of immunodeficiency is indicated when clinical signs of recurrent infections and fevers are present, in the case of infections with opportunistic organisms (e.g., P. carinii, C. parvum, adenovirus), or the blood lineage history suggests inheritance of primary immune disorders. Humoral immunodeficiency can be diagnosed initially by serum immunoglobulin concentrations. In foals less than 3 months old, serum IgG concentration interpretation should take into account circulating levels of colostral antibodies, which vary individually depending on the initial amount of absorption (colostral IgG half-life 28–32 days). Therefore, serum IgM concentrations are more specific for B-cell function in foals, as IgM production already occurs in utero and colostral IgM has a short half-life of 5–16 days.18–20 Serum values < 25 mg/dL suggest an underlying humoral dysfunction. Humoral response to vaccination in foals may also be difficult to interpret because of maternal antibody interference, and the need of one or two boosters to produce a measurable response; therefore, this test may be more useful in weanlings and yearlings.35 The evaluation of peripheral blood B-cell distribution may add in the investigation of underlying humoral immunodeficiency, by suggesting normal or abnormal B-cell development. Cellular immunodeficiency is comparatively rare and/or difficult to diagnose. Diagnosis can be supported by peripheral blood T-cell distribution, T-cell proliferative response and cytokine production upon mitogenic stimulation in vitro, and cell-mediated reaction to intradermal phytolectin injection. All the immunological testing in foals should be accompanied by control samples from age-matched and, when possible, breed-matched healthy foals to account for developmental changes in the immune system. The results should also be compared with confidence intervals determined by the laboratory.30 Repeated tests should be performed to confirm a persistent versus a transient condition. A list and applications of immunological testing available for the horse are described in Table 36.2.
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Table 36.2 Immunological testing available for the horse Test
Objective
Potential Findings
Peripheral blood cytology Bone marrow cytology
count and distribution of distinct cell populations cell morphology cell morphology
neutropenia, lymphopenia, anemia, thrombocytopenia lymphoblasts, large granular neutrophils hypoplastic cell deficiencies, low myeloid-erythroid ratio
Humoral Function Serum Ig concentrations (RID or ELISA) Response to vaccination Peripheral blood lymphocyte phenotyping (FC) Peripheral blood lymphocyte proliferation (FC)
production of immunoglobulins in vivo production of antibodies distribution of circulating B cells proliferation response to B cell mitogens
IgG, IgM or IgA hypo- or agammaglobulinemia no rise in IgG titer after 3 vaccinations B cell lymphopenia or depletion poor B lymphocyte proliferative response
distribution of circulating CD4+ and CD8+ T cells proliferation response to T cell mitogens cytokine expression after mitogen stimulation MHC class II percentage positive lymphocytes in vivo T cell recruitment and proliferation
CD4+ and/or CD8+ T cell lymphopenia; low CD4/CD8 ratio poor T lymphocyte proliferative response low T cell function inadequate age-dependent T cell development poor lymphocyte response
phagocytosis capacity and production of oxygen reactive species for bacterial killing
decreased phagocytic function
identify SCID carriers and affected horses
mutation of DNA-PKc
Immune Cell Development Peripheral blood cell count
Cellular Function Peripheral blood lymphocyte phenotyping (FC) Peripheral blood lymphocyte proliferation (FC) Lymphocyte IFNγ production (FC) Lymphocyte MHC class II expression (FC) Mitogen (PHA) skin test Phagocytic Function Phagocytosis and oxidative burst activity (FC) Genetic Tests SCID VetGen, Ann Arbor, MI, USA
Interpretation of results is meaningful when compared to an age-matched control, preferably of the same breed. IG – immunoglobulin; RID – radial immunodiffusion; ELISA – enzyme-linked immunosorbent assay; FC – flow cytometry; IFNg – interferon gamma; MHC class II – major histocompatibility complex class II; PHA – phytohemagglutinin; SCID – severe combined immunodeficiency; DNA-PK – DNA-dependent protein kinase catalytic subunit
References 1. Riggs MW. Evaluation of foals with immune deficiency disorders. Vet Clin North Am Equine Pract 1987;3:515–28. 2. Perryman LE. Primary immunodeficiencies of horses. Vet Clin North Am Equine Pract 2000;16:105–16 3. Giguere S, Polkes AC. Immunologic disorders in neonatal foals. Vet Clin North Am Equine Pract 2005;21:241–72. 4. McClure JJ, Addison JD, Miller RI. Immunodeficiency manifested by oral candidiasis and bacterial septicemia in foals. J Am Vet Med Assoc 1985;186:1195–7. 5. Tanaka S, Kaji Y, Taniyama H, Matsukawa K, Ochiai K, Itakura C. Pneumocystis carinii pneumonia in a thoroughbred foal. J Vet Med Sci 1994;56:135–7. 6. McGuire TC, Poppie MJ, Banks KL. Hypogammaglobulinemia predisposing to infection in foals. J Am Vet Med Assoc 1975;166:71–5. 7. Banks KL, McGuire TC, Jerrells TR. Absence of B lymphocytes in a horse with primary agammaglobulinemia. Clin Immunol Immunopathol 1976;5:282–90. 8. McGuire TC, Banks KL, Evans DR, Poppie MJ. Agammaglobulinemia in a horse with evidence of functional T lymphocytes. Am J Vet Res 1976;37:41–6. 9. Deem DA, Traver DS, Thacker HL, Perryman LE. Agammaglobulinemia in a horse. J Am Vet Med Assoc 1979;175:469–72.
10. Flaminio MJBF, Tallmadge R, Salles-Gomes CM, Matychak MB. Common variable immunodeficiency in horses is characterized by B-cell depletion in primary and secondary lymphoid tissues. J Clin Immunol 2009;29:107–16. 11. Perryman LE, McGuire TC, Hilbert BJ. Selective immunoglobulin M deficiency in foals. J Am Vet Med Assoc 1977;170:212–15. 12. Perryman LE, McGuire TC. Evaluation for immune system failures in horses and ponies. J Am Vet Med Assoc 1980;176:1374–7. 13. Crisman MV. Selective immunoglobulin M deficiency in the horse. In: Proceedings of the 33rd American Association of Equine Practitioners, New Orleans, LA, December, 1987, pp. 313–19. 14. McGuire TC, Perryman LE, Davis WC. Analysis of serum and lymphocyte surface IgM of healthy and immunodeficient horses with monoclonal antibodies. Am J Vet Res 1983;44:1284–8. 15. Boy MG, Zhang C, Antczak DF, Hamir AN, Whitlock RH. Unusual selective immunoglobulin deficiency in an Arabian foal. J Vet Intern Med 1992;6:201–5. 16. Weldon AD, Zhang C, Antczak DF, Rebhun WC. Selective IgM deficiency and abnormal B-cell response in a foal. J Am Vet Med Assoc 1992;201:1396–8. 17. Perkins GA, Nydam DV, Flaminio MJBF, Ainsworth DM. Serum IgM concentrations in normal, fit horses and horses
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with lymphoma or other medical conditions. J Vet Internal Med 2003;17:337–42. Perryman LE, McGuire TC, Torbeck RL. Ontogeny of lymphocyte function in the equine fetus. Am J Vet Res 1980;41:1197–200. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Absorption of bovine colostral immunoglobulins G and M in newborn foals. Am J Vet Res 1989;50:1598–603. Tallmadge RL, McLaughlin K, Secor E, Ruano D, Matychak MB, Flaminio MJBF. Expression of essential B cell genes and immunoglobulin isotypes suggests active development and gene recombination during equine gestation. Dev Comp Immunol 2009;33:1027–38. Scholes SF, Holliman A, May PD, Holmes MA. A syndrome of anaemia, immunodeficiency and peripheral ganglionopathy in Fell pony foals. Vet Rec 1998;142:128– 34. Richards AJ, Kelly DF, Knottenbelt DC, Cheeseman MT, Dixon JB. Anaemia, diarrhoea and opportunistic infections in Fell ponies. Equine Vet J 2000;32:386–91. Dixon JB, Savage M, Wattret A, Taylor P, Ross G, Carter SD, Kelly DF, Haywood S, Phythian C, Macintyre AR, Bell SC, Knottenbelt DC, Green JR. Discriminant and multiple regression analysis of anemia and opportunistic infection in Fell pony foals. Vet Clin Pathol 2000;29:84–6. Tallmadge RL, Stokol T, Matychak MB, Flaminio MJBF. Prospective and molecular studies of Fell Pony Syndrome. In: Proceedings of the 8th International Veterinary Immunology Meeting, Minas Gerais, Brazil, 15–19 August, 2007. Thomas GW, Bell SC, Carter SD. Immunoglobulin and peripheral B-lymphocyte concentrations in Fell pony foal syndrome. Equine Vet J 2005;37:48–52. Bell SC, Savidge C, Taylor P, Knottenbelt DC, Carter SD. An immunodeficiency in Fell ponies: a preliminary study into cellular responses. Equine Vet J 2001;33:687–92.
27. Gardner RB, Hart KA, Stokol T, Divers TJ, Flaminio MJ. Fell Pony syndrome in a pony in North America. J Vet Intern Med 2006;20:198–203. 28. Thomas GW, Bell SC, Phythian C, Taylor P, Knottenbelt DC, Carter SD. Aid to the antemortem diagnosis of Fell pony foal syndrome by the analysis of B lymphocytes. Vet Rec 2003;152:618–21. 29. Malinowski K, Hallquist NA, Helyar L, Sherman AR, Scanes CG. Effect of different separation protocols between mares and foals on plasma cortisol and cell-mediated immune response. Equine Vet Sci 1990;10:363–8. 30. Flaminio MJ, Rush BR, Davis EG, Hennessy K, Shuman W, Wilkerson MJ. Characterization of peripheral blood and pulmonary leukocyte function in healthy foals. Vet Immunol Immunopathol 2000;73(3–4):267–85. 31. Flaminio MJ, Rush BR, Cox JH, Moore WE. CD4+ and CD8+ T-lymphocytopenia in a filly with Pneumocystis carinii pneumonia. Aust Vet J 1998;76:399–402. 32. Gardner RB, Nydam DV, Luna JA, Bicalho ML, Matychak MB, Flaminio MJ. Serum opsonization capacity, phagocytosis, and oxidative burst activity in neonatal foals in the intensive care unit. J Vet Intern Med 2007;21:797–805. ¨ 33. Grondahl G, Sternberg S, Jensen-Waern M, Johannisson A. Opsonic capacity of foal serum for the two neonatal pathogens Escherichia coli and Actinobacillus equuli. Equine Vet J 2001;33:670–5. 34. Flaminio MJBF, Rush BR, Davis EG, Shuman W and Wilkerson M. Simultaneous analysis of phagocytosis and oxidative burst activity of equine phagocytes by flow cytometry. In: Lunn DP, Wade JF (eds) Havemeyer Foundation Monograph Series, R & W Publications, Suffolk, UK, No. 4, 2001; pp. 69–70. 35. Wilson WD, Mihalyi JE, Hussey S, Lunn DP. Passive transfer of maternal immunoglobulin isotype antibodies against tetanus and influenza and their effect on the response of foals to vaccination. Equine Vet J 2001;33:644–50.
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Chapter 37
Severe Combined Immunodeficiency Melissa T. Hines
Severe combined immunodeficiency (SCID) includes a group of disorders characterized by the inability to generate antigen-specific immune responses. Due to the lack of both B- and T-cell function, affected individuals are highly susceptible to infection. SCID has been recognized in multiple species, including horses.1–4 In horses, the condition is best described in Arabians and part Arabians, although sporadic cases have been described in other breeds.5, 6
Epidemiology SCID in horses of Arabian breeding is known to be an autosomal recessive trait.7 First described in 1973, the condition is believed to have originated in a horse in Great Britain and then spread to other countries in association with the importation of horses.8 SCID has now been reported in Arabians in the United States, Canada, and Australia, as well as Great Britain.5, 9–11 The wide distribution of this disorder illustrates the importance of the founder effect in population genetics. The prevalence of SCID in Arabian foals born in the United States was initially estimated to be 2–3%, leading to the prediction that the number of SCID carriers at that time could be as high as 25% of the Arabian horse population.11 Selective breeding practices, made easier by the identification of the abnormal gene, appear to have decreased the prevalence of SCID. In 1998 a study of 250 randomly selected Arabian horses in the United States found the frequency of SCID gene carriers to be 8.4%.9 Based on this information and assuming a random breeding population, it was predicted that at this time 0.18% of Arabian foals would be affected with SCID. A second study in 2002 also found the carrier frequency to be approximately 8%.2
Pathophysiology A mutation on chromosome 9 has been identified in association with SCID in Arabians.3, 4, 12 This mutation is a 5 base-pair deletion in the gene encoding the catalytic subunit of the enzyme DNA-dependent protein kinase (DNA-PK).7,12–14 This deletion results in a frameshift and a 967 amino acid deletion from DNA-PK, yielding an unstable mutant protein. Without functional DNA-PK, lymphocyte precursors are unable to complete gene rearrangement events that lead to the expression of antigen-specific receptors on lymphocyte surfaces. Therefore, there are no mature, functional T- or B-lymphocytes.15–18 Also, interferonγ (EqIFN–γ ), which is an important cytokine produced by lymphocytes, is deficient.19 Other protective mechanisms, including neutrophils, monocytes, and natural killer cells appear to be functional in Arabian foals with SCID.20, 21 Complement levels are also normal.17 However, even with intact non-specific immune mechanisms, foals with SCID are highly susceptible to infection due to their lack of specific humoral or cellular immunity.
Clinical signs Foals that are homozygous for the defective SCID gene are immunodeficient which is manifested clinically by an increased susceptibility to infectious diseases. Heterozygotes are asymptomatic and can only be identified by genetic testing or production of an affected offspring.22 Recently, a correlation has been demonstrated between heterozygosity for the SCID allele and development of sarcoids, a virally induced tumor.2 These data suggest that DNA-PK may play a role in tumor suppression by maintaining chromosomal stability.
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Foals with SCID generally appear normal at birth, but the absence of antigen-specific immune responses renders them highly susceptible to infections once colostral protection wanes.5, 11, 23 The precise age of onset of clinical disease varies depending on the adequacy of passive transfer of immunity and the degree of environmental challenge, but affected foals typically develop infectious diseases between birth and 2 months of age and die before 5–6 months of age. A wide variety of bacterial, viral, parasitic, and fungal agents have been isolated in foals with SCID. Many of these agents are opportunists which rarely cause disease in animals that are not immunocompromised.5,24–28 While infection may occur at any site, pneumonia is particularly common. Pneumocystis carinii pneumonia and adenoviral pneumonia are found often in SCID foals and rarely in other foals.11, 28
Clinical pathology An absolute lymphopenia is a consistent finding in foals with SCID.16, 17 Lymphocyte numbers are always less than 1000 lymphocytes/µL, and are often much lower. However, the total white blood cell count is variable and may be low, normal, or elevated, depending primarily on the number of neutrophils. Therefore, when evaluating foals for SCID, it is important to determine the absolute number of lymphocytes. Persistent lymphopenia should be established for a diagnosis of SCID to be considered, as septicemic, immature and other compromised foals, as well as some normal foals, may transiently have low lymphocyte counts. Foals with SCID are unable to produce immunoglobulins due to the lack of functional B-cells.15, 17 As a result, concentrations of autologous serum immunoglobulin for age are abnormally low. As quantitative tests do not distinguish between immunoglobulin that is autologously produced and that of maternal origin, the degree of colostral transfer and the age of the foal must be considered when interpreting serum immunoglobulin concentrations. Since IgM is normally produced in utero by the foal, some IgM should be present in presuckle serum and although not pathognomonic, the absence of IgM in presuckle serum is a feature of SCID.29 Also, the absence of serum IgM after 3 to 4 weeks of age may be suggestive of SCID. This is because the concentration of maternal IgM in colostrum is lower than IgG and the half-life of IgM is shorter. As a result, most maternal IgM received from colostrum is metabolized by about 3 weeks of age, whereas the age at which maternal IgG is gone is much more variable and may be several months.
Foals affected with SCID are deficient in cellular as well as humoral immune responses. Therefore, they do not respond to intradermal phytohemagglutinin by increasing skin thickness as do normal foals, nor do they respond to stimulators of delayed-type hypersensitivity such as dinitrochlorobenzene.30 In vitro tests of cellular immunity such as blastogenesis are depressed with all mitogens.16, 17, 31 The definitive diagnosis of SCID in foals of Arabian breeding is based on demonstrating that the foal is homozygous for the defective SCID gene.14, 22 Blood or cheek swabs may be submitted to VetGen for DNA testing to determine if a horse is clear, heterozygous, or homozygous for the mutation (VetGen, Veterinary Genetic Services, Michigan, USA). Before genetic testing was available, the criteria required to confirm a diagnosis of SCID included: (i) persistent lymphopenia (<1000/µL); (ii) absence of serum IgM in presuckle samples or samples collected after 3 weeks of age; and (iii) thymic hypoplasia and characteristic histopathological changes in lymphoid tissue.32, 33
Necropsy findings Gross and histological evaluation of lymphoid tissues can be useful in assessing cases of suspected immunodeficiency. In SCID, the thymus is small and typically has a fatty appearance on gross examination.18, 32 The thymus may actually be difficult to identify in some affected foals due to its small size, and in these cases mediastinal tissue must be collected ‘blindly’ for subsequent histological evaluation and identification of thymic remnants. Histologically the thymus is largely replaced with adipose tissue, with only islands of lymphoid cells and partially formed Hassall’s corpuscles present. Occasional CD8+ cells have been identified in the thymus of SCID foals.21 While the spleen and lymph nodes may not appear grossly abnormal, a number of abnormalities have been identified histologically.32 These include an absence of germinal centers and periarteriolar lymphocytic sheaths in the spleen, as well as an absence of germinal centers with scarcity of lymphocytes in other areas of lymph nodes (Figure 37.1a). Foals with SCID often have evidence of infectious disease at necropsy (Figure 37.1b). Bronchopneumonia is a particularly common finding; however, infectious lesions in other systems are also frequently found, including colitis and hepatitis.
Treatment and prognosis There is no practical method of treatment for foals with SCID at this time. Even with intensive
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in one in four foals affected with SCID, one in four completely normal and not a carrier, and two in four asymptomatic carriers of the SCID trait. Mating of a carrier and a normal (non-carrier) does not produce any affected foals, but half of the offspring would be expected to be carriers. SCID carrier horses were previously identified by the production of an affected offspring, which identified both the sire and dam as carriers. The ability to now identify carriers by genetic testing has simplified the control of SCID by breeding practices. Mares and stallions intended for breeding should be tested to determine whether they are normal or heterozygous for the defective SCID gene. To avoid the production of an affected foal, two heterozygotes should never be mated to one another. If an owner decides to continue breeding a heterozygote, the breeding partner should be confirmed homozygous normal. All foals from such matings should be tested to determine if they are clear of or heterozygous for the defective gene (50:50 probability). Homozygous normal foals may be selected for future breeding purposes, whereas heterozygous foals should be strictly used for non-reproductive purposes only.
References
Figure 37.1 Tissue from an Arabian foal with SCID. The foal presented at 1 month of age for evaluation of persistent pneumonia. (a) 200× image of mediastinal lymph node showing extensive lymphoid atrophy. The node is poorly populated with lymphocytes which fail to form lymphoid follicles, and there is no evidence of active germinal centers. The node is comprised predominantly of loosely scattered resorbed mononuclear cells and edema. (b) 400× image of tracheal epithelium with adenovirus inclusion (black arrows), epithelial hyperplasia and suppurative tracheitis. (See color plate 3). (Photographs courtesy of Sushan Han.)
conventional therapy, such as antimicrobials, plasma, and isolation, affected foals invariably die by approximately 5 months of age.34 Although not very practical, successful bone marrow transplantation has been performed between histocompatible full siblings.35 Transplantation of tissue from unrelated horses was unsuccessful, as SCID foals developed graft versus host disease.36, 37 However, transplantation of histocompatible bone marrow cells from a healthy full sibling restored a functional immune system.35
Prevention SCID can be controlled in horses of Arabian breeding by breeding practices. As SCID is an autosomal recessive trait, mating of two carriers is expected to result
1. Bell TG, Butler KL, Sill HB, Stickle JE, Ramos-Vara JA, Dark MJ. Autosomal recessive severe combined immunodeficiency of Jack Russell terriers. J Vet Diagn Invest 2002;14(3):194–204. 2. Ding Q, Bramble L, Yuzbasiyan-Gurkan V, Bell T, Meek K. DNA-PKcs mutations in dogs and horses: allele frequency and association with neoplasia. Gene 2002;283(1–2): 263–9. 3. Leber R, Wiler R, Perryman LE, Meek K. Equine SCID: mechanistic analysis and comparison with murine SCID. Vet Immunol Immunopathol 1998;65(1):1–9. 4. Perryman LE. Molecular pathology of severe combined immunodeficiency in mice, horses, and dogs. Vet Pathol 2004;41(2):95–100. 5. McGuire TC, Poppie MJ, Banks KL. Combined (B and T lymphocyte) immunodeficiency: a fatal genetic disease in Arabian foals. J Am Vet Med Assoc 1974;164(1):70–5. 6. Perryman LE, Boreson CR, Conaway MW, Bartsch RC. Combined immunodeficiency in an Appaloosa foal. Vet Pathol 1984;21(5):547–8. 7. Perryman LE, Torbeck RL. Combined immunodeficiency of Arabian horses: confirmation of autosomal recessive mode of inheritance. J Am Vet Med Assoc 1980;176(11):1250–1. 8. McGuire TC, Poppie MJ. Hypogammaglobulinemia and thymic hypoplasia in horses: a primary combined immunodeficiency disorder. Infect Immun 1973;8(2):272–7. 9. Bernoco D, Bailey E. Frequency of the SCID gene among Arabian horses in the USA. Anim Genet 1998;29(1): 41–2. 10. Swinburne J, Lockhart L, Scott M, Binns MM. Estimation of the prevalence of severe combined immunodeficiency disease in UK Arab horses as determined by a DNA-based test. Vet Rec 1999;145(1)22–3.
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11. Studdert MJ. Primary, severe, combined immunodeficiency disease of Arabian foals. Aust Vet J 1978;54(9):411–17. 12. Shin EK, Perryman LE, Meek K. A kinase-negative mutation of DNA-PK (CS) in equine SCID results in defective coding and signal joint formation. J Immunol 1997;158(8):3565–9. 13. Shin EK, Rijkers T, Pastink A, Meek K. Analyses of TCRB rearrangements substantiate a profound deficit in recombination signal sequence joining in SCID foals: implications for the role of DNA-dependent protein kinase in V(D)J recombination. J Immunol 2000;164(3):1416–24. 14. Wiler R, Leber R, Moore BB, Van Dyk LF, Perryman LE, Meek K. Equine severe combined immunodeficiency: a defect in V(D)J recombination and DNA-dependent protein kinase activity. Proc Natl Acad Sci USA 1995;92(25): 11485–9. 15. Buening GM, Perryman LE, McGuire TC. Immunoglobulins and secretory component in the external secretions of foals with combined immunodeficiency. Infect Immun 1978;19(2):695–8. 16. Lew AM, Hosking CS, Studdert MJ. Immunologic aspects of combined immunodeficiency disease in Arabian foals. Am J Vet Res 1980;41(8):1161–6. 17. McQuire TC, Banks KL, Poppie MJ. Combined immunodeficiency in horses: characterization of the lymphocyte defect. Clin Immunol Immunopathol 1975;3(4):555–66. 18. Wyatt DR, Magnuson NS, Perryman LE. Defective thymocyte maturation in horses with severe combined immunodeficiency. J Immunol 1987;139(12):4072–6. 19. Yilma T, Perryman LE, McGuire TC. Deficiency of interferon-γ but not interferon-β in Arabian foals with severe combined immunodeficiency. J Immunol 1982;129: 931–3. 20. Banks KL, McGuire TC. Surface receptors on neutrophils and monocytes from immunodeficient and normal horses. Immunology 1975;28(3):581–8. 21. Lunn DP, McClure JT, Schobert CS, Holmes MA. Abnormal patterns of equine leucocyte differentiation antigen expression in severe combined immunodeficiency foals suggests the phenotype of normal equine natural killer cells. Immunology 1995;84(3):495–9. 22. Shin EK, Perryman LE, Meek K. Evaluation of a test for identification of Arabian horses heterozygous for the severe combined immunodeficiency trait. J Am Vet Med Assoc 1997;211(10):1268–70. 23. Don-van’t Slot HP, van der Kolk JH. Severe combined immunodeficiency disease (SCID) in the Arabian horse. Tijdschr Diergeneeskd 2000;125(19):577–81.
24. Biberstein EL, Jang SS, Hirsch DC. Norcardia asteroides infection in horses: a review. J Am Vet Med Assoc 1985;186(3):273–7. 25. Clark E.G., Turner AS, Boysen BG, Rouse BT. Listeriosis in an Arabian foal with combined immunodeficiency. J Am Vet Med Assoc 1978;172(3):363–6. 26. Gibson JA, Hill MW, Huber MJ. Cryptosporidiosis in Arabian foals with severe combined immunodeficiency. Aust Vet J 1983;60(12):378–9. 27. Mair TS, Taylor FG, Harbour DA, Pearson GR. Concurrent cryptosporidium and coronavirus infections in an Arabian foal with combined immunodeficiency syndrome. Vet Rec 1990;126(6):127–30. 28. Thompson DB, Spradborw PB, Studdert M. Isolation of an adenovirus from an Arabian foal with a combined immunodeficiency disease. Aust Vet J 1976;53(10):435–7. 29. Tizard I: Immunity in the fetus and newborn. In: Tizard I (ed.) Veterinary Immunology: An Introduction, 3rd edn. Philadelphia: Saunders, 1987. 30. Hodgin EC, McGuire TC, Perryman LE, Grant BP. Evaluation of delayed hypersensitivity responses in normal horses and immunodeficient foals. Am J Vet Res 1978;39(7): 1161–7. 31. Perryman LE, McGuire TC. Evaluation for immune system failures in horses and ponies. J Am Vet Med Assoc 1980;176(12):1374–7. 32. McGuire TC, Banks KL, Davis WC. Alterations of the thymus and other lymphoid tissue in young horses with combined immunodeficiency. Am J Pathol 1976;84(1):39–53. 33. Perryman LE. Primary and secondary immune deficiencies of domestic animals. Adv Vet Sci Comp Med 1979;23: 23–52. 34. Perryman LE, McGuire TC, Crawford TB. Maintenance of foals with combined immunodeficiency: causes and control of secondary infections. Am J Vet Res 1978;39(6): 1043–7. 35. Bue CM, Davis WC, Magnuson NS, Mottironi VD, Ochs HD, Wyatt CR, Perryman LE. Correction of equine severe combined immunodeficiency by bone marrow transplantation. Transplantation 1986;42(1):14–9. 36. Ardans AA, Trommershausen-Smith A, Osburn BI, Mayhew IG, Trees C, Park MI, Sawyer M, Stabenfeldt GH. Immunotherapy in two foals with combined immunodeficiency, resulting in graft versus host reaction. J Am Vet Med Assoc 1977;170(2):167–75. 37. Perryman LE, Liu IK. Graft versus host reactions in foals with combined immunodeficiency. Am J Vet Res 1980;41(2):187–92.
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Chapter 38
Thrombocytopenia and Selective IgM Deficiency Rodney L. Belgrave
Introduction Thrombocytopenia is a frequently encountered hematological disorder in equine neonates, generally defined by blood platelet levels below 100,000/µL.1 The thrombocytopenic state may arise as a result of reduced platelet production or shortened platelet lifespan due to either consumption or destruction. Sequestration and severe hemorrhage may also lead to significantly reduced platelet levels. Clinical conditions such as sepsis, disseminated intravascular coagulation (DIC), and immune-mediated thrombocytopenia including neonatal alloimmune disease are most commonly associated with increased platelet destruction and consumption in neonates. Myelophthisic or infiltrative diseases of the bone marrow may affect platelet production, as may potentially myelosuppressive pharmaceuticals such as phenylbutazone and chloramphenicol.2 Platelets are an integral component of the primary hemostatic process. They undergo a series of transformations in response to contact with damaged vascular endothelium, which results in adherence to the vascular defect and recruitment of additional platelets. This aggregation of platelets gives rise to the platelet plug. This plug is subsequently reinforced if needed by the production of fibrin threads during the process of blood coagulation.3 Clinical examination and diagnosis of non-specific thrombocytopenia Clinical examination findings in the severely thrombocytopenic neonate include mucous membrane and aural petechial and ecchymotic hemorrhages. Scleral hemorrhage, hyphema, and melena may also be seen, along with prolonged bleeding after venipuncture, abdominocentesis, or surgical procedures. These clin-
ical manifestations are typically seen when peripheral platelet counts decline to less than 40 000/µL.1 Pseudothrombocytopenia may occur in horses when ethylenediaminetetraacetic acid (EDTA) is used as an anticoagulant. Sodium citrate 3.8% should be used as an anticoagulant to eliminate this phenomenon for a more accurate determination of the platelet count.1 Septicemia is one of the more common maladies affecting the equine neonate. Thrombocytopenia is frequently seen in association with sepsis in neonatal foals, especially if accompanied by the phenomenon of disseminated intravascular coagulation.4 As such, efforts to rule out septicemia should be made when working up a thrombocytopenic neonate. Sepsis due to Gram-negative organisms is most frequently seen. Clinical manifestations of septicemia are nonspecific but may include lethargy, weakness, fever, injected mucous membranes, tachycardia, tachypnea, and coronary band hyperemia. Some of these clinical findings, along with hematological findings of neutropenia and low plasma immunoglobulin G (IgG) levels, can be used to determine the sepsis score, which may support a diagnosis of septicemia.4 Marked petechiation of the mucous membranes seen with cases of severe thrombocytopenia, when applied to the sepsis score, may confound the results and make it difficult to distinguish between sepsis and other causes of thrombocytopenia.5 When faced with the thrombocytopenic neonate, the presence of normal clotting profiles [prothrombin time (PT), activated partial thromboplastin time (aPTT)] and fibrin degradation products minimize the likelihood of DIC as a cause of the thrombocytopenia. Bone marrow evaluation may also aid in distinguishing between reduced platelet production and destruction or consumption as a cause of the thrombocytopenia.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Immune-mediated thrombocytopenia Immune-mediated thrombocytopenia (IMTP) is associated with antibody-mediated destruction of platelets. Such destruction may be autoimmune in nature, as seen in human infants, in which transplacental transference of maternal antibodies directed at antigens common to both maternal and fetal platelets occurs. Neonatal autoimmune thrombocytopenia has not been recognized in the equine neonate. In human fetoneonatal alloimmune thrombocytopenia, alloantibodies are produced against paternally derived fetal platelet antigens.6 This condition has been previously described in an equine neonate, where antibodies directed against the foal’s platelets were recognized in the mare’s plasma, serum, and milk.7 The syndrome has also been experimentally induced in mule foals.8 A similar but distinct syndrome of severe thrombocytopenia, ulcerative dermatitis, and neutropenia has also been described.5 The foals in this report were less than 4 days old at presentation and demonstrated oral and lingual ulceration (Figure 38.1) in addition to diffuse dermal crusting and erythema. The skin lesions and thrombocytopenia were transient, and it was postulated that colostral antibodies were responsible for the syndrome. The mares involved were multiparous, and multiple foals from the same mare were affected, suggesting alloimmune disease. The condition was prevented by offering an alternative source of colostrum to subsequent foals born to two of the affected mares. There have been reports describing the use of flow cytometric methods as a sensitive method to determine the presence of platelet-bound immunoglobulin in horses with thrombocytopenia.9–11 Platelets bound with immunoglobulin are quickly removed from the circulation via the mononuclear phagocyte system. As a result, flow cytometric assays for
Figure 38.1 Oral ulcers in a neonatal foal affected with ulcerative dermatitis, thrombocytopenia, and neutropenia.
platelet-associated antibodies should be performed early in the course of disease. Other diagnostics may include direct immunofluorescence to determine platelet surface-associated immunoglobulin or indirectly by measuring the binding of the mare’s antibodies to the foal’s platelets.5 Treatment of foals affected by this syndrome is primarily supportive and may include broad-spectrum antibiotic therapy, and whole blood or platelet-rich plasma transfusions. Use of corticosteroids, although effective in increasing platelet counts in two of the affected foals, did not appear to significantly affect the outcome of these cases.
Selective immunoglobulin M deficiency Selective immunoglobulin M (IgM) deficiency is a syndrome observed most frequently in foals of Arab and Quarter Horse lineage ranging from 2 to 8 months of age.12 It is characterized by nonexistent or markedly reduced levels of serum IgM in the face of normal concentrations of all other immunoglobulins. Normal lymphocyte numbers and immune function tests are typically documented in such cases. One report describing selective IgM deficiency also documented a concurrent abnormal B-cell response, which was suggested as a cause of the IgM deficiency.13 Foals are immunologically na¨ıve at birth, with detectably low levels of immunoglobulins. Passive immunity accounts for increased post-colostral IgM levels among other immunoglobulins, which wane at around 3–4 weeks of age. In normal foals, this is typically followed by a gradual rise in immunoglobulin levels towards those of an adult.14 This innate rise in IgM levels does not occur in affected foals, which are plagued with severe respiratory disease, arthritis, and enteritis. The condition is invariably fatal, with foals being susceptible to Gram-negative bacteria in particular. In one report, Klebsiella species were isolated from affected joints, tracheobronchial lymph nodes, and tracheal washes in three out of five affected foals.12 Some affected foals may survive temporarily, but are subjected to recurrent bacterial infections, which flare up upon discontinuation of antibiotic therapy. A small number of affected foals developed normal serum IgM concentrations over time, with concurrent resolution of their bacterial infections. The underlying mechanism for this clinical improvement remains undetermined.15 Although a hereditary basis is suspected, a pathogenesis of the disease is yet to be elucidated. Diagnosis of the condition is made by determination of serum immunoglobulin levels via various methodologies, including radial immunodiffusion
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Thrombocytopenia and Selective IgM Deficiency and enzyme-linked immunosorbent assay (ELISA). IgM levels are generally less than two standard deviations below age-matched controls.12 Immunoglobulin G levels are within normal limits, as are peripheral lymphocyte counts. Prognosis is poor, as there is no effective treatment to replenish or sustain declining IgM levels. Supportive care and broad-spectrum antibiotic therapy are usually provided.
7.
8.
9.
References 10. 1. Davis E, Wilkerson MJ. Thrombocytopenia. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. St. Louis: Saunders, 2003; p. 349. 2. Kingston J. Thrombocytopenia. In: Brown CM, Bertone JJ (eds) The Five Minute Veterinary Consult Equine. Baltimore: Lippincott, Williams & Wilkins, 2002; p. 1054. 3. Guyton AC, Hall JE. Textbook of Medical Physiology, 10th edn. Philadelphia: W.B. Saunders Co., 2000; pp. 419–29. 4. Brewer BD. Neonatal infection. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Baltimore: Lea & Febiger, 1990; pp. 295–316. 5. Perkins GA, Miller WH, Divers TJ, Clark CK, Belgrave RL, Sellon DC. Ulcerative dermatitis, thrombocytopenia, and neutropenia in neonatal foals. J Vet Intern Med 2005;19;211–16. 6. Fratellanza G, Fratellanza A, Paesano L, Scarcella A, Safoian A, Misso S, Formisano S, Scarpato N. Fetoneonatal
11.
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alloimmune thrombocytopenia (FNAIT): our experience. Transfus Apher Sci 2006;35(2):111–17. Buechner-Maxwell V, Scott MA, Godber L, Kristensen A. Neonatal alloimmune thrombocytopenia in a Quarter Horse foal. J Vet Intern Med 1997;11:304–8. Ramirez S, Gaunt SD, McClure JJ, Oliver J. Detection and effects on platelet function of anti-platelet antibody in mule foals with experimentally induced neonatal alloimmune thrombocytopenia. J Vet Intern Med 1999;13:534–9. Davis EG, Wilkerson MJ, Rush BR. Flow cytometry: clinical applications in equine medicine. J Vet Intern Med 2002;16:404–10. McGurrin MKJ, Arroyo LG, Bienzle D. Flow cytometric detection of platelet bound antibody in three horses with immune mediated thrombocytopenia. J Am Vet Med Assoc 2004;224:83–7. Nunez R, Gomes-Keller MA, Schwarzwald C, Feige K. Assessment of equine autoimmune thrombocytopenia (EAT) by flow cytometry. BMC Blood Disord 2001;1:1–13. Perryman LE, McGuire TC, Hilbert BJ. Selective immunoglobulin M deficiency in foals. J Am Vet Med Assoc 1977;170:212–15. Wheldon AD, Zhang C, Antczak DF, Rebhun WC. Selective IgM deficiency and abnormal B-cell response in a foal. J Am Vet Med Assoc 1992;201:1396–8. Thomas GW, Bell SC, Carter SD. Immunoglobulin and peripheral B-lymphocyte concentrations in Fell pony foal syndrome. Equine Vet J 2005;37:48–52. Perryman LE. Primary immunodeficiencies in horses. Vet Clin North Am Equine Pract 2000;16:105–16.
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39.
Meconium Impaction Wendy E. Vaala
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Neonatal Diarrhea K. Gary Magdesian
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Gastroduodenal Ulceration: The Neonatal Perspective L. Chris Sanchez
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Surgical Gastrointestinal Conditions Brett J. Woodie
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Other Gastrointestinal Disorders (Chyloperitoneum, Atresia, Lethal White) K. Gary Magdesian
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Hepatic Disease Thomas J. Divers and T. Douglas Byars
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Chapter 39
Meconium Impaction Wendy E.Vaala
Introduction Meconium is composed of ingested amniotic fluid, mucus, bile, and epithelial cellular debris. This first manure varies in consistency from pelleted to pasty and is usually black or dark green in color. In the absence of in utero infection, the fetal gut and its contents are sterile at birth. Meconium passage is usually complete within 12 to 24 hours of delivery and is followed by the passage of soft, yellow, pasty ‘milk feces’. Meconium can pose a problem for the newborn foal in two ways: aspiration of meconium during the birth process, and failure to void meconium postpartum. In utero passage of meconium is usually associated with prepartum or intrapartum asphyxia. In utero asphyxia or umbilical cord occlusion can result in fetal defecation of meconium into the amniotic fluid. Fetal hypoxia induces a redistribution of blood flow away from less vital organs including the gastrointestinal tract, resulting in mesenteric vasoconstriction and secondary intestinal ischemia. Transient hyperperistalsis and anal sphincter relaxation occur, thereby allowing passage of meconium. Fetal gasping before or during delivery results in meconium aspiration. Meconium aspiration can produce a variety of clinical signs including mechanical obstruction (ball-valve effect) and regional air trapping, chemical pneumonitis, alveolar edema, and displacement of surfactant by the free fatty acids in meconium, leading to decreased lung compliance and atelectasis. These events lead to increased pulmonary vascular and airway resistance and ventilation/perfusion mismatching. Meconium aspiration is discussed in detail in Chapters 64 and 65. Failure of the neonatal foal to void meconium postpartum results in varying degrees of discomfort ranging from restlessness, decreased nursing vigor and tail flagging, to overt colic with persistent rolling. Healthy newborn foals begin to pass meconium dur-
ing the first few hours after delivery and are commonly observed arching their backs and flagging their tails while defecating due to the tenacious nature of meconium. Meconium can be retained or impacted in the large colon, small or transverse colon and/or rectum, resulting in varying degrees of tenesmus. With prolonged retention the meconium plug becomes dessicated and firmly adhered to the intestinal mucosa, and prevents even the passage of gas and fluid around it. Progressive accumulation of gas and ingesta proximal to the obstruction produces progressive, tympanitic bowel distention, visible abdominal enlargement, and more severe, generalized ileus and persistent bouts of colic.
History and clinical signs Meconium retention is believed to be more common in colts due to their narrow pelvic canal.1 Newborn foals typically begin to pass meconium within the first few hours following delivery and should complete voiding meconium within the first 12 to 18 hours postpartum as a result of peristalsis. The appearance of soft, tan or yellow pasty feces associated with milk ingestion is a good indication that the foal has completed meconium passage, although occasionally scant amounts of milk feces may be passed around a meconium impaction. The tenacious nature of this first manure causes most foals to strain when defecating. If meconium is not passed, and becomes firmly impacted in the colon, the foal becomes increasingly uncomfortable due to progressive intestinal distention. Prolonged, persistent intestinal distention contributes to ileus. Early signs of meconium retention in the newborn foal may be subtle, and include reduced frequency of nursing and increased periods of recumbency. If the condition persists, the affected foal exhibits increasing restlessness, tail flagging, pacing and straining to defecate with the tail head elevated and the back
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 39.2 Meconium retention wtihin the small colon of a premature foal. Figure 39.1 A newborn foal straining to defecate with an arched back and elevated tail.
arched (Figure 39.1). Since the act of nursing stimulates defecation through a gastrocolonic reflex, signs of abdominal discomfort associated with distal meconium retention may appear shortly after each milk meal and may be mistaken for upper gastrointestinal discomfort. Likewise, tenesmus may be mistaken for dysuria, although foals that are straining to urinate usually ventroflex, rather than arch their back. Evacuation of meconium may be delayed as a result of ileus secondary to other conditions including septicemia and peripartum asphyxia syndrome (PAS). Left untreated, meconium retention leads to varying degrees of abdominal distention and colic. The foal’s abdomen becomes gas distended with tympany detected over both paralumbar fossas. Foals suffering meconium impaction can become violently colicky with pawing and sudden rolling, which frequently results in secondary self-trauma. Persistent tenesmus can result in rectal mucosa edema and eversion. Foals suffering concurrent, but unrelated, conditions such as urachal patency or inguinal/scrotal hernias may experience a worsening in those conditions as a result of persistent straining to defecate and increased abdominal pressure. Meconium impaction may cause enough discomfort and lack of nursing that a newborn foal fails to ingest and absorb adequate amounts of colostrum in a timely fashion, resulting in failure of passive transfer and an increased risk of secondary sepsis. Extensive mural damage from severe and persistent meconium impaction also can contribute to bacterial translocation and secondary septicemia.
Diagnosis of meconium retention and impaction A gentle digital rectal examination using a welllubricated, gloved finger helps identify meconium
plugs within the distal rectum as far cranial as the pelvic inlet. If the foal is relaxed or sedated enough, external palpation of the abdomen may detect firm meconium in the large colon.2 Clinical signs of colic and the lack of meconium passage confirms the diagnosis in most cases. More proximal obstructions may require other imaging modalities. Lateral abdominal radiographs obtained with the foal in a standing position can be used to identify the gas–fluid interfaces within the bowel. Both right and left lateral views should be taken since some structures are more apparent depending on which side of the abdomen is closer to the film cassette. Severe, generalized distention of the large colon is suggestive of a distal obstruction. Figure 39.2 shows meconium impaction in the small bowel of a premature foal. Contrast radiographs obtained following a barium enema help identify impactions involving the transverse colon or small colon.3 To perform this procedure, the foal should be lightly sedated and placed in lateral recumbency. A well lubricated, 24french, Foley catheter is inserted into the rectum and the cuff gradually inflated. Using gravity flow, up to 20 mL/kg bodyweight of barium is administered slowly. A lateral abdominal radiograph is obtained while the barium is retained. Transabdominal ultrasonography using a linear array or curvilinear transducer (4–7.5 MHz) can be used to evaluate the abdomen in a colicky foal. If scanning the foal in lateral recumbency, the abdomen should be viewed from both sides. It is preferable to scan the foal in a standing position since the meconium-filled colon tends to drop into the dependent portion of the abdomen, and the gas that develops proximal to the impaction may preclude a scan of deeper structures in the recumbent foal. Sonographically, meconium may appear hyperechoic, hypoechoic, or a mixture of echogenicities. The intestine is usually hypomotile and fluid or gas is visualized proximal to the obstruction. Meconium
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retained within the small colon often appears as a row of sacculated balls, while meconium retained within the large colon has a more amorphous appearance. Foals with uncomplicated meconium impactions of short duration should have relatively normal complete blood counts and serum chemistries. If the impaction persists, affected foals may develop a stress leukogram (leukocytosis with mature neutrophilia) and varying degrees of hemoconcentration, hyperproteinemia, and azotemia consistent with decreased nursing and progressive dehydration. Failure of passive transfer may occur in foals that fail to consume and absorb adequate amounts of colostrum within the early postpartum period.
Differentials for meconium retention and impaction It is important to note whether the foal has passed any meconium at all, since a complete absence of manure passage is associated with other congenital gastrointestinal defects including atresia coli, atresia recti, atresia ani, and ileocolonic aganglionosis. Most cases of atresia can be confirmed with a digital rectal examination when a total absence of meconium, coupled commonly with white or slightly discolored mucus, is evident, or with contrast radiography.3 The all-white offspring of Overo spotted Paint Horses suffer from fatal congenital ileocolonic aganglionosis. This condition is termed Overo lethal white foal syndrome (OLWFS) and results in atrophy of the distal small intestine and colon. Affected foals retain meconium and demonstrate progressive colic beginning on day 1 as a result of fatal functional intestinal obstruction. This is an untreatable condition and surgery is not an option. Lethal white foal syndrome is discussed in detail in Chapters 43 and 73. Other differentials for colic in the neonatal foal include ileus, with or without evidence of enterocolitis, gastroduodenal ulcers, intussusception, intestinal volvulus, and less common intestinal accidents such as strangulation or obstruction of the intestines by a rent in the mesentery or diaphragm. Occasionally tenesmus may be mistaken for dysuria associated with uroperitoneum.
Therapy Since most foals with meconium retention and/or impaction exhibit some degree of pain, low doses of analgesics such as dipyrone (11 mg/kg i.v.), flunixin meglumine (0.25–0.5 mg/kg i.v.), and/or butorphenol (0.01–0.05 mg/kg i.v.) may be required to prevent
Figure 39.3 Administration of an acetylcysteine retention enema using a cuffed Foley catheter inserted into the foal’s rectum. The enema solution is infused slowly.
self-trauma during colicky episodes. Xylazine can exacerbate gut stasis and causes varying degrees of respiratory depression, and should be used with caution in newborn foals. The author avoids the use of detomidine in the neonate due to unpredictable levels of sedation and cardiopulmonary depression. Therapy for simple meconium retention includes gravity-flow enemas using warm, soapy water or powdered J-lube powder dissolved in water and administered through a small-bore, well-lubricated catheter. Soft, flexible tubing, such as a stallion urinary catheter or Foley catheter, are often used (Fig. 39.3). The catheter is slowly and gently advanced until an obstruction is encountered. The enema solution (approximately 500–1000 mL) is slowly administered by gravity flow. Powdered J-lube dissolved in water seems less irritating than water and detergent solutions, especially if more than one enema is required. The foal often forces some fluid out and around the catheter. This should be allowed. Walking the foal after the enema may help to void additional meconium. Commercial over-the-counter phosphate enemas also can be used, but repeated administration may increase the risk of electrolyte disturbances including, most notably, hyperphosphatemia, hypocalcemia, and hypernatremia.4–6 The author does not
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recommend using irritating substances such as dioctyl sodium succinate (DSS) in foal enemas since this detergent can produce excessive mucosal inflammation with tenesmus. Owners should be advised not to administer more than several enemas without consulting their veterinarian. Rupture of the rectum at the site of the impaction with secondary fecal peritonitis can occur with repetitive or overly aggressive enema administration. Excessive irritation of the rectum and colon induces mucosal edema and may predispose to bacterial translocation and secondary septicemia in the debilitated foal. If meconium retention persists despite several gravity enemas, then an acetylcysteine retention enema is recommended.7–9 The supplies and procedure for a retention enema are presented in Table 39.1. The final concentration of the acetylcysteine solution described is approximately 4% with a pH between 7 and 9.
Table 39.1 Supplies and procedure for acetylcysteine retention enema Supplies
r r r r r r r r
150 mL water 6 g N-acetylcysteine powder 20 g sodium bicarbonate powder (baking soda) Cuffed 14–24 french Foley catheter and plug for the end (plunger of 3 mL or 5 mL syringe often works) Syringe and enough water to inflate the cuff of Foley catheter being used Catheter tip syringe to administer acetylcysteine solution KY lubricant; Lidocaine ointment can also be used and may further decrease tenesmus associated with the procedure White tape to attach Foley catheter to tail
Procedure
r r r r r r r
Lightly sedate the foal Mix together the first three ingredients. The bicarbonate is added to adjust solution pH to an optimum of 7 to 9. Insert a well-lubricated, cuffed Foley catheter approximately 6–10 inches into the foal’s rectum and inflate the cuff with the appropriate volume of water in order to prevent backflow of the acetylcysteine solution Slowly infuse 120–180 mL of the retention enema solution into the rectum Plug the end of the catheter and tape the end of the catheter loosely to the foal’s tail Leave the retention enema in place for a minimum of 20–40 minutes Deflate cuff and remove the catheter
The retention enema may be repeated several times. The passage of brown, meconium-stained fluid or feces is an encouraging sign.
Foals tend to tolerate and retain the acetylcysteine enema for a longer time period and with reduced straining if they are lightly sedated prior to the procedure and gently restrained in lateral recumbency during the enema administration. Lubricating the catheter with a lidocaine- or carbocainecontaining gel prior to catheter insertion also helps reduce tenesmus. In a retrospective study, 41 of 44 foals with meconium impaction responded favorably to medical therapy that included one to three acetylcysteine retention enemas.1 This study documented a higher success rate for medical therapy than an earlier study (performed prior to the routine use of acetylcysteine retention enemas) in which 8 of 24 foals with meconium impactions required surgery.10 Surgical removal of meconium should be a last resort, due to the risk of long–term complications associated with postoperative adhesion formation following abdominal surgery in neonatal foals. Acetylcysteine works by cleaving disulfide bonds and decreasing the viscosity of the meconium.9, 11 Acetylcysteine’s mucolytic effect is enhanced by prolonged contact with the meconium. In human neonates, N-acetylcysteine may be administered by orogastric tube or per rectum as a treatment for meconium obstruction and ileus.11, 12 Some clinicians advocate the use of a fecal loop to dislodge retained fecal balls.13 If a loop is used, extreme caution is warranted, to prevent trauma to rectal mucosa. The use of a forceps to grasp meconium is discouraged unless the clinician is experienced in using this technique, adequate lubrication is used, the meconium plug can be identified, grasped and removed without excessive manipulation, and the foal is properly restrained or sedated. Additional therapy for meconium impaction includes administration of intravenous polyionic fluids, oral fluids, and oral laxatives. Resolution of high impactions often requires both oral laxatives and systemic fluid therapy. If one or two enemas fail to relieve the meconium obstruction it is prudent to administer a small volume of mineral oil (60–120 mL of oil mixed with an equal volume of warm water) to the foal via nasogastric intubation. This oral laxative helps lubricate around the retained meconium and may delay dessication of the impacted fecal balls. It is the author’s experience that the longer the meconium impaction remains in place, the more likely the foal is to develop a complete intestinal obstruction resulting in serious gaseous abdominal distention accompanied by more violent bouts of colic. Another oral laxative that can be used if mineral oil alone is not sufficient is Milk of Magnesia, 30–60 mL administered orally. The use of more irritating cathartics such as castor oil and DSS is discouraged since excessive use of these agents can lead
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Meconium Impaction to severe diarrhea and may increase the risk of secondary sepsis due to inflammation of the gastrointestinal mucosal barrier. Isotonic, polyionic intravenous fluids infused at a rate that exceeds maintenance requirements are given to correct pre-existing dehydration and to increase intraluminal fluid content and help soften the consistency of retained meconium. Intravenous fluids can be given continuously or as repetitive intravenous boluses. Supplemental oral fluids can be administered by nasogastric intubation, but some foals become colicky following fluid therapy using this technique. Foals that fail to respond to initial therapy for meconium impaction may benefit from temporary withdrawal of milk, with supplemental nutritional support in the form of intravenous, dextrosecontaining fluids and/or parenteral nutrition.
Percutaneous trocarization and intestinal decompression A few foals develop severe abdominal distention as a result of complete obstruction of the small colon by firm meconium. Marked abdominal distention and tympany results in persistent and often violent colic, leading to repeated administration of systemic analgesics that can produce undesirable respiratory depression and may further reduce gastrointestinal motility. A grossly enlarged abdomen exacerbates dyspnea in some foals, especially those that are recumbent, premature or debilitated with another neonatal condition such as sepsis or hypoxic ischemic encephalopathy. Bowel overdistention antagonizes normal motility and exacerbates intestinal ileus. Severely affected foals may benefit from percutaneous trocarization and decompression of the gas-distended large bowel. To perform the procedure the foal should be sedated with a combination of diazepam (0.05–0.2 mg/kg) and xylazine (0.1 mg/kg) or butorphenol (0.01–0.04 mg/kg) and xylazine (0.1 mg/kg) given intravenously to help maintain the foal in ‘quiet’ lateral recumbency. With the foal in lateral recumbency the gas-filled large bowel distention becomes even more pronounced over the paralumbar fossa. An area over the dorsal aspect of each distended paralumbar fossa is clipped and aseptically prepared. A subcutaneous bleb of 2% carbocaine is instilled over the site of the intended troacarization. A 16-G, 3 12 -inch catheter over the stylet is used as a trocar. The catheter is attached to a sterile, 30-inch extension set. Wearing surgical gloves and using aseptic technique, the catheter and stylet are inserted briskly through the skin and sub-
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cutaneous tissues into the distended intestinal viscus in one smooth thrust. Once the bowel has been entered and gas begins escaping, submerge the free end of the extension set in a beaker or plastic specimen cup filled with sterile water or saline, to visualize gas bubbles escaping. An assistant can apply gentle pressure along the lower abdomen to facilitate decompression dorsally. When finished and gas bubbles are no longer observed, quickly withdraw the catheter while instilling 2 mL of amikacin or gentocin as the catheter passes through the subcutaneous tissues and the skin incision. Following trocarization, the foal should receive a minimum 5-day course of broad-spectrum antibiotics. Complications are rare if the foal is adequately restrained during the procedure and the catheter (with the stylet in place) is held still with minimal manipulation during the procedure. Decompression of the bowel can provide dramatic pain relief, thereby reducing the need for excessive analgesic administration. Decompression also helps promote a return of gut motility. A very small percentage of foals fail to respond to medical management and may require surgery, especially if the impaction is located in the large colon.14, 15 Specific discussion of surgical treatment is discussed elsewhere in Chapter 42.
Prophylaxis Many breeding farm managers and individual mare/foal owners administer a soapy water gravity enema or a phosphate enema to foals shortly after delivery to facilitate passage of meconium and reduce the risk of meconium retention and impaction. This is a good preventive measure that helps stimulate early meconium passage and most likely reduces the incidence of distal meconium retention.
References 1. Pusterla N, Magdesian KG, Maleski K. Retrospective evaluation of the use of acetylcysteine enemas in the treatment of meconium retention in foals: 44 cases (1987–2002). Equine Vet Educ 2004;16:133–6. 2. Ryan CA, Sanchez LC. Nondiarrheal disorders of the gastrointestinal tract in neonatal foals. Vet Clin North Am Equine Pract 2005;21(2):313–32. 3. Fischer A, Yarborough T. Retrograde contrast radiography of the distal portions of the intestinal tract in foals. J Am Vet Med Assoc 1995;207(6):734–6. 4. Hickman SA, Gill MS, Marks SL, Smith JA, Sod GA. Phosphate enema toxicosis in a pygmy goat wether. Can Vet J 2004;45:849–51. 5. Marraffa JM, Hui A, Stork CM. Severe hyperphosphatemia and hypocalcemia following the rectal administration of a phosphate-containing Fleet pediatric enema. Pediatr Emerg Care 2004;20:453–6.
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6. Biebl A, Grillenberger A, Schmitt K. Enema-induced severe hyperphosphatemia in children. Eur J Pediatr 2009;168(8):1023. 7. Madigan JE, Goetzman BW. Use of acetylcysteine solution enema for meconium retention in the neonatal foal. Proc Am Assoc Equine Pract 1990;207:734–7. 8. Madigan JE. Meconium retention. In: Madigan JE (ed.) Manual of Equine Neonatal Medicine. Woodland, CA: Live Oak Publishing, 1987; pp. 111–15. 9. Burke MS, Ragi JM, Karamanoukian HL, Kotter M, Brissear GF, Borowitz DS, Ryan ME, Irish MS, Glick PL. New strategies in nonoperative management of meconium ileus. J Pediatr Surg 2002;37(5):760–4. 10. Hughes FE, Moll HD, Sloan DE. Outcome of surgical correction of meconium impactions in 8 foals. J Equine Vet Sci 1996;16:172–5.
11. Shaw A. Safety of N-acetylcysteine in treatment of meconium obstruction of the newborn. J Pediatr Surg 1969;4:119–25. 12. Emil S, Nguyen T, Sills J. Padilla G. Meconium obstruction in extremely low-birth-weight neonates: guidelines for diagnosis and management. J Pediatr Surg 2004;39(5):731–7. 13. Wilson JH, Cudd TA. Common gastrointestinal disease. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 412–30. 14. Adams R. Exploratory celiotomy for gastrointestinal disease in neonatal foals. Equine Vet J 1988;20:9–12. 15. Adams R. Gastrointestinal surgery. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 430–42.
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Chapter 40
Neonatal Diarrhea K. Gary Magdesian
Diarrhea is a common cause of morbidity and mortality of foals, especially during the neonatal period.1 In one study evaluating the causes of disease in foals, diarrhea was second only to pneumonia as a cause of disease, with the perinatal period (≤7 days) being the highest risk for development of diarrhea.2 In another study, 48% of foals from 15 surveyed farms developed diarrhea during the neonatal period and up to 2 months of age.1 Enteritis can affect individual foals and occur as farm outbreaks, with a high morbidity. Some forms of diarrhea are associated with a high mortality rate and require intensive care, highlighting the importance of early recognition and therapeutic intervention. The etiologies, treatments, and prevention of enteritis and enterocolitis in foals are reviewed in this chapter (Table 40.1).
Risk factors associated with development of diarrhea in foals One study evaluating the development of diarrhea in 48% of 297 foals from 15 farms identified risk factors for diarrhea.1 Disinfection of foaling stalls between uses was associated with a significantly reduced risk of diarrhea, as was the practice of udder washing and tail wrapping prior to foaling. Increased risk of neonatal diarrhea was associated with use of shavings as bedding, the prophylactic administration of antimicrobials, and treatment with oral vitamins. In addition, foals born to visiting mares were at increased risk of developing diarrhea compared with those of resident mares.1 This highlights the importance of moving mares to foaling properties as soon as possible, at least 1 month prior to the anticipated due date. An epidemiological study by Cohen2 identified that the incidence of diarrhea was significantly lower among farms where foals were born on pasture than on farms where foals were born in stalls.
Infectious causes of diarrhea Bacterial causes
Salmonellosis Salmonellosis is one of the most severe forms of enterocolitis in foals. Foals are at particular risk of bacteremia with Salmonella infections, with potential for septic arthritis, osteomyelitis, and other secondary foci of infection.3 Although the clinical severity of salmonellosis varies with the serotype and host immune response, in general most foals develop moderate to severe disease. Clinical signs include diarrhea (watery to hemorrhagic), colic, fever, anorexia, and lethargy, as well as signs of systemic inflammatory response syndrome (SIRS). Group B salmonellae, such as Salmonella typhimurium or agona, are among the most virulent serotypes affecting horses. Localized and secondary infections, including septic arthritis, osteomyelitis, tenosynovitis, and peritonitis, contribute to a high morbidity and mortality rate among foals infected with Salmonella spp. Isolation of Salmonella spp. from synovial fluid has been associated with an unfavorable prognosis for survival.4 Diagnosis of salmonellosis is through fecal culture, including the use of enrichment broths. More recently, polymerase chain reaction (PCR), including quantitative (TaqMan) PCR, of feces has been used and is very sensitive, particularly when applied to overnight enrichment broth. Blood cultures should also be performed in foals with salmonellosis to evaluate for the presence of bacteremia. Hematology often shows leukopenia, immature (band) neutrophilia, and toxic cytology. Pulmonary aspergillosis and ischemic distal limb necrosis have been reported as potential complications of salmonellosis in a foal.5 Septic arthritis, osteomyelitis, tenosynovitis, and other localized infections, such as
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 40.1 Causes of neonatal foal diarrhea Type of diarrhea
Cause
Infectious
Salmonella Clostridium difficile Clostridium perfringens Rotavirus Coronavirus Cryptosporidium Strongyloides westeri
Non-infectious diarrhea
Pica Nutritional causes Lactose intolerance Perinatal asphyxia Necrotizing enterocolitis Sepsis
nephritis or peritonitis, are additional complications of infection with Salmonella spp. The cornerstone of treatment of foals with salmonellosis is a triad of antimicrobials, hemodynamic support, and nursing care. Systemic antimicrobials are to protect against bacterial translocation (Table 40.2). The use of antimicrobials does not shorten the course of enteritis, but is used for preTable 40.2 Specific therapies for infectious diarrhea Causative agent
Therapy
Salmonella
Broad spectrum antimicrobials to prevent bacteremia Amikacin (21–25 mg/kg, i.v., q. 24 hours) or third-generation cephalosporins (ceftiofur 5–10 mg/kg i.v., q. 12–24 hours; cefotaxime or ceftazidime, 50 mg/kg i.v. q. 6 hours)
Clostridium difficile
Metronidazole (10–15 mg/kg i.v. or p.o., q. 8–12 hours); smectite paste (30–60 ml p.o., q. 12 hours); lactase (6000 IU p.o., q. 3–8 hours)
Clostridium perfringens
Metronidazole; smectite; lactase as above for C. difficile
Rotavirus
Lactase (6000 IU p.o., q. 3–8 hours); NZT? (anecdotal; 25 mg/kg p.o., q. 24 hours)
Cryptosporidium
Anecdotal for severe cases not responding to supportive care: NZT? (25 mg/kg p.o. q. 24 hours); paromomycin? (100 mg/kg p.o. q. 24 hours for 5 days); azithromycin? (10 mg/kg p.o. q. 24 x 5 d, then q. 48 hours)
Strongyloides westeri
Ivermectin (200 µg/kg p.o. once); oxibendazole (15 mg/kg p.o. once)
NTZ, nitazoxanide (this product is no longer commercially available).
venting or treating bacteremia. Amikacin (21–25 mg/kg intravenously (i.v.), q. 24 hours) or thirdgeneration cephalosporins (ceftiofur 5–10 mg/kg i.v., q. 12–24 hours; cefotaxime or ceftazidime, 50 mg/kg i.v. q. 6 hours) are effective against most equine Salmonella isolates. The use of aminoglycosides for treating Salmonella infections is controversial owing to the intracellular nature of the microbe, although aminoglycosides are likely effective as prophylactic antimicrobials in preventing bacteremia. Whenever aminoglycosides are used, therapeutic drug monitoring and serial measurement of serum creatinine should be carried out. Hemodynamic support is through the administration of crystalloids and colloids, inotropes, and pressors. Initially, boluses of isotonic crystalloids, such as PlasmaLyte 148 or A (Baxter International, Deerfield, IL), Normosol R (Abbott Laboratories, North Chicago, IL), or lactate Ringer’s solution (LRS), should be administered at doses of 20 mL/kg i.v. over 30–60 minutes. Such boluses should be continued until perfusion parameters, arterial blood pressure, blood lactate, and central venous oxygen either normalize or plateau at some level, where no further improvement is observed. It is rare that neonatal foals would require more than two or three such boluses. Colloids can also be used, but at lower doses. For example, hetastarch can be administered at 3–5 mL/kg i.v. A total daily dose of 10 mL/kg/day of hetastarch should not be exceeded owing to development of coagulopathies. If no further hemodynamic improvements are observed after the fluid boluses, or if central venous pressure is at maximum (10 cmH2 O), then inotropes should be administered (dobutamine continuous rate infusion (CRI) at 3–10 µg/kg/min). Vasopressors such as norepinephrine (0.01–0.1 µg/kg/min CRI) should be administered if dobutamine does not improve perfusion parameters, blood pressure, and lactate. Plasma (hyperimmune to salmonellae or Escherichia coli endotoxins) administration is commonly used in foals with salmonellosis, although the efficacy of such products in treating diarrheic foals is unknown. In one study plasma transfusions were associated with increased survival in septic foals.6 Although administration of plasma does not alter neutrophil function in healthy foals, it does enhance the activity of those in septic foals.7 In one experimental model of endotoxemia in foals Salmonella antiserum was associated with an increase in interleukin 6 (IL-6) and tumor necrosis factor (TNF) compared with control foals.8 Autogenous Salmonella bacterin vaccines have been used on endemic farms during outbreaks, in an extralabel manner. They have been administered to pregnant mares in an attempt to reduce morbidity
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Neonatal Diarrhea and mortality associated with salmonellosis in neonatal foals (N. Slovis, personal communication). They have included Salmonella newport and S. typhimurium. Anecdotally these vaccines have been reported to be safe and felt to have reduced the incidence of diarrhea on endemic farms (N. Slovis, personal communication). A two-dose series has been used. Rigorous biosecurity protocols should be instituted on premises to prevent spread of the disease. Foals may shed salmonellae for longer periods of time than adults. Serial fecal cultures or PCR should be performed until foals are negative on several repeated tests (minimum five negatives) before they are considered non-shedders and removed from strict isolation. Salmonellae are potentially zoonotic, and affected foals should be handled with this in mind.
Clostridium difficile and Clostridium perfringens Clostridium difficile and Clostridium perfringens are among the most common causes of infectious diarrhea in newborn foals. These clostridial agents are primary pathogens in foals, and can cause outbreaks or individual cases of diarrhea.9–11 Clinical signs of enteric clostridiosis include colic and abdominal distension, variable fever, and watery to hemorrhagic diarrhea. In severe cases of C. perfringens type C infections, foals are sometimes found dead prior to development of diarrhea; this emphasizes the need for close monitoring of foals for lethargy or anorexia in the early neonatal period. C. perfringens types A, B, C, D, and E are potentially pathogenic in animals, with types A and C being the most common isolates affecting neonatal foals. Inflammation is incited by production of one or more exotoxins, including alpha, beta, epsilon, iota, and enterotoxin. Type A isolates produce alpha toxin, while type C produce alpha and beta. More recently, beta-2-producing isolates have been associated with disease in horses, and some type A equine isolates produce enterotoxin. Most severe clostridiosis occurs in young foals (<5 days old), although foals of any age can be affected. One report described a mortality rate above 54% in foals with C. perfringens, and it appears that foals with type C infections have the poorest prognosis.9 Hemorrhagic diarrhea is often seen with type C. C. difficile produces toxins A (‘enterotoxin’), B (‘cytotoxin’), and/or binary toxins; these toxins are virulence factors that result in cell necrosis and inflammation within the gut, creating varying degrees of enteritis. Diarrhea, depressed mentation, dehydration, hypovolemia, and tachypnea are clinical signs of C. difficile infection in foals. The mortality rate of foals
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with C. difficile is lower than that of C. perfringens, reported as 18% in one study.10 C. perfringens and C. difficile infections in foals are diagnosed through identification of both toxins and the microbial agent in fecal samples. Culture of both agents requires anaerobic culture and selective media. A number of commercially available fecal immunoassays are available for detections of toxins A and/or B for C. difficile. Fecal cytotoxin assays (cell culture) can be used to identify toxin B, although these are not as widely available as immunoassays. C. perfringens toxin fecal enzyme-linked immunoassay (ELISA) testing can be performed for alpha, beta, and epsilon toxins at the California Animal Health and Food Safety Laboratory (San Bernardino, CA 92408, USA). C. perfringens types A (alpha toxin), B (alpha, beta, and epsilon toxins), C (alpha and beta toxins), and D (alpha and epsilon) can be typed using this fecal ELISA. C. perfringens ‘enterotoxin’ immunoassays are available in many commercial laboratories, although only a minority of type A isolates produce this toxin, and it should therefore not be relied upon to make a diagnosis. Isolates can also be typed once cultured through the use of multiplex PCR for toxin gene sequences. PCR techniques can identify beta2-producing isolates, because immunoassays do not test for this toxin type. Gram stains of feces provide a rapid screen for clostridiosis; a large number of Gram-positive rods is suggestive of clostridial overgrowth, although it is not highly specific (Figure 40.1). C. difficile and C. perfringens (even toxin genecontaining isolates) can both be cultured from the feces of healthy foals; this fact complicates the diagnosis and requires either quantification of the agent or identification of fecal toxins in order to confirm the diagnosis. For C. perfringens, fecal concentrations >103 colony-forming units (cfu)/mL are consistent with infection. In addition, finding the toxins in feces (through immunoassays or stool cytotoxin assays) is also evidence of disease rather than subclinical colonization. Foals with clostridiosis often have hematological changes consistent with a systemic inflammatory response syndrome, including neutropenia with immature (band) neutrophilia, toxic cytological changes, hypoproteinemia, and hyperfibrinogenemia. Interestingly, most foals with clostridiosis have adequate passive transfer of colostral antibodies, and some clinicians have speculated that colostral trypsin inhibitors may protect the integrity of clostridial toxins through the stomach.9 Abdominal ultrasonography is a useful diagnostic tool in identifying suspect cases, and in monitoring the progress of cases. Foals with clostridiosis will often have distended, fluid-filled, and hypomotile small intestinal loops
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Figure 40.1 Gram stain of colon contents showing Clostridium difficile cells with subterminal spores. Photograph courtesy of Spencer Jang.
on ultrasonography, even before clinical signs of diarrhea develop. This will allow for earlier detection of cases. Finding gas shadows within the intestinal wall on ultrasound is an ominous sign and reflects invasion by the anaerobic bacteria or bowel perforation. The treatment of foals with clostridiosis is twofold: (i) supportive care, primarily focusing on maintenance of hydration and acid–base and electrolyte balance (see above; Salmonellosis); (ii) specific therapy with antimicrobials that target the agent (Table 40.2). Metronidazole (10–15 mg/kg, i.v. or per os (p.o.), q. 8–12 hours) is the first line antimicrobial against both C. difficile and C. perfringens. Because of rare metronidazole resistance among some C. difficile isolates, minimum inhibitory concentration (MIC) testing of isolates to metronidazole is recommended. Documentation of resistance on susceptibility patterns may warrant the use of oral vancomycin. However, this should be judicious and only when clear resistance has been documented. When used, foals should be kept in strict isolation using barrier protocols in a hospital setting. The use of vancomycin (4 mg/kg loading, then 2 mg/kg q. 6 hours, oral) should be extremely limited in light of its reservation in human medicine for multidrug-resistant Enterococcus or Staphylococcus isolates. Nitazoxanide (NZT), an anti-protozoal drug used in humans to
treat cryptosporidial infections, has demonstrated efficacy against C. difficile on an experimental basis and is currently an investigational drug being studied as a treatment for C. difficile in humans.12 NZT has been used to treat a small number of foals with clostridial diarrhea at a dose of 25 mg/kg p.o. once daily (N. Slovis, personal communication). This may become an accepted treatment of affected foals in the future; however, at the current time a commercial source of NZT for equine use is lacking. Nitazoxanide itself can cause severe diarrhea. Studies have demonstrated in vitro binding of both C. perfringens and C. difficile toxins by di-trioctahedral smectite (Bio-sponge, Platinum Performance Buellton, CA), suggesting that there may be a place for smectite in the treatment of equine neonatal clostridiosis.13, 14 The dose of smectite paste for a 50-kg foal is 30–60 mL p.o., q. 12 hours; however, it should not be administered within the first 12 hours of initial nursing as it can reduce the availability of colostral antibodies.14 Lactase deficiency has been reported in a foal with C. difficile infection, and is likely secondary to widespread mucosal villous damage; supplementation with exogenous lactase (6000 IU q. 3–8 hours) is therefore indicated in affected foals.15 Plasma products from horse donors hyperimmunized with C. perfringens and C. difficile antigens are available for extralabel use; however, there are no studies evaluating their efficacy. C. perfringens types C and D antitoxin has been used empirically in some foals, although safety and efficacy data are lacking.9 Slow (and dilute, if i.v.) administration of these products and possible pretreatment with diphenhydramine (0.5–1 mg/kg, slow i.v. or intramuscularly (i.m.) are warranted. The use of probiotics, especially those containing the yeast Saccharomyces boulardii, have shown some promise as an adjunctive treatment in humans with C. difficile infection. However, these should be used with caution in very young foals because of the risks of fungemia, which has been documented in human infants. Foals with enteritis, particularly clostridiosis, often demonstrate signs of abdominal pain. Analgesia can be provided through opioids or non-steroidal anti-inflammatory drugs (NSAIDs). Because of concerns over gastrointestinal (GI) and renal perfusion, NSAIDs should be used judiciously. The author uses low doses of flunixin meglumine, such as 0.25 mg/kg i.v., q. 8–12 hours, in foals that are hydrated. Butorphanol (0.01–0.02 mg/kg, i.v.) can be administered for foals with colic signs. Foals with severe enteritis may benefit from being withheld from nursing for 24–72 hours. Such gut rest can decrease the severity of diarrhea by removing an osmotic contribution to the diarrhea, as well as through allowing the intestinal mucosa a period of time to heal. Candidates include those foals with abdominal distension,
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Neonatal Diarrhea pain, or severe diarrhea. Such foals should be placed on parenteral nutrition during this period of anorexia. Because clostridial spores are very difficult to eliminate from the environment, foals should be isolated to individual stalls. These areas should then be cleaned with manual scrubbing using a detergent, followed by disinfection with bleach or peroxygen compounds (Virkon or RelyOn, Antec International, Sudbury, Suffolk, UK); the last have been shown to be sporicidal for C. difficile.
Other bacterial causes of diarrhea Other bacteria have been associated with diarrhea in foals; however, their exact role in causing enteritis is unknown. These include enterotoxigenic attaching and effacing E. coli,16 Aeromonas hydrophila, Streptococcus durans, Campylobacter jejuni, Clostridium sordelli, and Bacteroides fragilis. Viral causes
Rotavirus Rotavirus is one of the most common causes of infectious diarrhea in foals and is highly contagious. A number of group A serotypes can affect foals, including G3, G5, G10, G13, G14, P7, P12, and P18.17, 18 While most foals are between 5 and 35 days old, rotavirus has a short incubation period and can affect foals as young as 2 days old and up to 6 months old. The incubation period can be as short as 12–24 hours with heavy challenge, but is more often 2 days. Clinical signs range from mild diarrhea and anorexia to severe, watery diarrhea with fever. Hemorrhagic feces are not a feature of rotaviral diarrhea, and are more likely due to a bacterial cause. The clinical severity of rotavirus infections seems to lessen with age. The diagnosis of rotavirus diarrhea is through commercial fecal antigen tests or identification of virus particles in feces with electron microscopy. Treatment of foals with rotavirus is largely supportive, with maintenance of perfusion and hydration, as well as electrolyte and acid–base status (see above; Salmonellosis). There are currently no specific treatments for viral enteritis in foals. Systemic antimicrobials should be administered prophylactically to neonatal foals, in order to prevent secondary bacteremia across the inflamed gut. Because of villous atrophy and compensatory crypt cell proliferation, foals with rotavirus develop maldigestion and malabsorption. Lactose intolerance can develop owing to small intestinal villous atrophy, and therefore exogenous lactase (6000 IU q. 3–6 hours for a 50-kg foal) should be included in the treatment (Table 40.2).19 Otherwise, undigested lactose will enter the large colon and contribute an osmotic component to the diarrhea. Bovine colostrum administered orally has
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been used by some clinicians in the treatment of rotaviral diarrhea in foals; however, the safety and efficacy require further evaluation.20 Treatment with hyperimmune bovine colostrum from cows immunized with human rotavirus was shown to reduce the duration and severity of rotaviral diarrhea in children.21 Recently, NZT has received attention in the treatment of children with rotaviral diarrhea. NZT is a thiazolide anti-infective agent with activity against protozoa and anaerobic bacteria. In vitro NZT and its metabolites inhibit replication of rotavirus, and exhibit a profound cytoprotective effect.22 Two doubleblinded, controlled clinical trials showed that time to resolution of clinical signs was significantly reduced with a 3-day course of NZT in human patients infected with rotavirus (both adults and children), and no significant adverse events were reported.22, 23 It has been tried in a small number of foals with rotavirus; however, the efficacy and safety of NZT in foals have not been studied (N. Slovis, personal communication). It should be cautioned that NZT itself can cause severe diarrhea. Strict infection control is imperative when dealing with rotavirus, owing to its highly contagious nature, with up to 100% morbidity in some farm outbreaks. Even mildly affected foals can serve as reservoirs for younger foals, and recovering foals can shed the virus subclinically. Fomites can also transmit rotavirus. Ideally, phenolic compounds (Tek-Trol, Bio-Tek Industries, Atlanta, GA; 1 Stroke Environ, Calgon Vestal Laboratories, St. Louis, MO) should be utilized to disinfect premises contaminated with rotavirus, as the virus is resistant to many common disinfectants. Bleach can also be used, but requires 5–10 minutes of contact time and is inactivated by organic matter. Cleaning of facilities is imperative, as the virus can persist for several months. A commercial vaccine is available for mares on endemic farms. When used, a three-dose series of the vaccine is administered during months 8, 9, and 10 of gestation regardless of the mare’s previous vaccination history. Studies have demonstrated a variable reduction in morbidity, length of diarrhea, and degree of shedding of viral particles in foals resultant from vaccinated dams.17, 18 The prognosis of foals with rotavirus diarrhea is generally good with treatment.
Coronavirus and other viruses Coronavirus has been reported as a cause of diarrhea in a few individual foals; however, it does not appear to be as important as rotavirus in causing foal diarrhea.24, 25 Prior reports of coronavirus diarrhea were limited to Arabian foals with severe combined immunodeficiency (SCID), but more recent reports have described it in apparently immunocompetent
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foals. One case report describes a 5-day-old foal with severe diarrhea, dehydration, hypoalbuminemia, anemia, and thrombocytopenia, reflecting the potential severity of coronaviral diarrhea.25 The foal was euthanized due to ischemic necrosis of the distal extremities. Fecal capture ELISAs and electron microscopy can be used for diagnosis. The role of adeno-, parvo-, breda, and other viruses in foal diarrhea remains to be studied. Parasitic causes
Treatment of cryptosporidiosis in foals is primarily supportive and most foals recover; until recently, there have been no specific therapies. There is limited evidence that NZT, paromomycin, and azithromycin have efficacy in the treatment of cryptosporidiosis in immunocompetent human patients. The use of these drugs could be attempted in severely affected foals not responding to standard supportive therapy, although safety and efficacy data in this age group of horses are lacking; these drugs should therefore only be used in severe cases (Table 40.2). It should be cautioned that NZT can cause severe diarrhea.
Cryptosporidium Cryptosporidium parvum is a cause of diarrhea in individual foals as well as herd outbreaks. Most affected foals are 4–21 days old, with a 3- to 7-day incubation period. Previously thought to affect only immunocompromised foals, it is now known to cause disease in immunologically normal foals. C. parvum isolates from foals are genetically diverse, including at least three types (genotype 2, cattle genotype, and a novel horse genotype), but they are all markedly similar to human and cattle isolates.26 Cryptosporidium has zoonotic potential, and affected foals should be treated with caution and strict biosecurity protocols. Recovering foals can serve to infect other foals, highlighting the importance of isolation. Cryptosporidium is resistant to many common disinfectants, but is susceptible to ammonia, chlorine dioxide, and hydrogen peroxide. There are a number of diagnostic tests available for Cryptosporidium; oocysts can be identified in feces by means of any of the following: acid-fast staining, immunofluorescence assay, electron microscopy, or flow cytometry. The sensitivity of identification of oocysts is markedly improved with acid-fast stains, owing to their small size (Figure 40.2). PCR analysis can be used for genotyping of strains.
Figure 40.2 Acid-fast stain of Cryptosporidium. Photograph courtesy of Robin Houston. (See color plate 4).
Nematodes Strongyloides westeri (threadworm) is passed through the milk to infest neonatal foals, where it lives as an adult in the small intestine; heavy infestations may cause self-limiting diarrhea. There is an association between S. westeri and diarrhea only when >2000 eggs per gram of feces are present.27 Tissue stages of larvae in the mare are sources of milk larvae, and these are thought to be larvae that the mares accumulated as foals. The diagnosis of Strongyloides diarrhea is made based on finding embryonated eggs in the foal’s feces at 8–14 days of age (Figure 40.3). Ivermectin (200 µg/kg) administered to the mare
Figure 40.3 Strongyloides westeri ovum. Photograph courtesy of Robin Houston.
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Neonatal Diarrhea postpartum is effective in blocking or minimizing transmammary transmission. Oxibendazole (15 mg/kg; note that the dose for Strongyloides is higher than the standard dose) or ivermectin (200 µg/kg) can be used to treat S. westeri infestations in foals (Table 40.2).
Non-infectious causes of diarrhea Postpartum or ‘foal heat’ diarrhea Foal heat diarrhea commonly affects foals in the 5to 15-day age range, and is temporally related to the mare’s first estrus postpartum.28, 29 The phrase ‘foal heat’ has been used to describe this type of diarrhea because of its timing relative to the postpartum heat cycle in mares; the diarrhea was thought to occur as a result of alterations in the dam’s milk composition in response to the hormonal events of the first postpartum estrus. However, even foals not allowed to nurse their dams and fed milk replacer develop diarrhea during this same time period, disproving the relationship to estrus theory.30 Also refuting this notion, studies of mares’ milk have shown that its composition does not change significantly during the first estrous cycle.31 Others have suggested a parasitic role; however, deworming of mares with ivermectin in the postpartum period did not result in a decrease in the rate of foal heat diarrhea, suggesting that S. westeri is not a contributor.32 Similarly, the presence of yeasts in the GI tract does not appear to be a cause of foal heat diarrhea.33 The etiology of foal heat diarrhea appears to be multifactorial and associated with maturation of the GI microflora and physiology, corresponding to the initial consumption of feed and inoculation of the microbial flora from coprophagia, and leading to transient diarrhea. The resultant diarrhea is usually mild and secretory in nature, with an increase in the concentration of electrolytes (sodium, potassium, chloride) and pH of the GI contents.28, 31 Fecal osmolarity is decreased during this time, primarily due to a decrease in the concentration of volatile fatty acids.31 The immature colon appears to be unable to compensate by increasing fluid and electrolyte absorption, and therefore diarrhea ensues. Foals with foal heat diarrhea should have normal clinical and clinicopathological findings. This type of diarrhea does not require specific therapy, and signs of lethargy, anorexia, fever, or abdominal distension warrant further investigation of the cause of diarrhea. Some foals with transient postpartum diarrhea may benefit from GI protectants, such as bismuth subsalicylate or kaolin and pectin. The diarrhea is selflimiting, and usually resolves within 1 week.
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Pica Pica can result in diarrhea through mechanical irritation of the GI mucosa. Affected newborn foals inappropriately ingest foreign materials, including sand, dirt, hair, and bedding such as straw, shavings, or rice hulls. Even excessive ingestion of the dam’s hay or feed during the early postpartum period can result in diarrhea because of maldigestion and intolerance of roughage. Consumed fibrous material is not adequately digested in the large intestine of the young foal, resulting in water influx into the colon. Diagnosis of pica-induced diarrhea is through witnessing the foal ingesting dirt or bedding, or eating significant amounts of hay rather than more appropriately sampling small amounts of feed. In addition, the foal’s feces should be closely inspected for this foreign material. Feces can be submerged in water for detection of sand or dirt. A digital rectal palpation often reveals the source of the diarrhea (gritty feel or presence of large amounts of fiber). Sand and dirt are radio-opaque and accumulation can be seen on abdominal radiographs.34 With expertise, abdominal ultrasonography (ventral abdomen) may also be used to detect sand in more severe cases.35 Foals that have ingested large quantities of foreign substances may exhibit abdominal distension and colic. Resolving the diarrhea associated with pica includes removal of access to the ingested material. Preventing access to sand, muzzling the foal with intermittent elimination for nursing, and removal of bedding from stalls is necessary in severe cases. Treatment of affected foals includes hastening the evacuation of the foreign material from the GI tract, through the use of intravenous fluids for hydration of GI contents and enteral laxatives such as mineral oil (2–4 mL/kg). Mineral oil should be used cautiously in the neonate during the first day of life; the author prefers to wait until foals are >18 hours old before administering mineral oil, when most of the macromolecular absorption of immunoglobulins has already occurred. Enemas are useful in evacuation of foreign material that is present within the aboral small colon. Although controversy exists regarding the efficacy of psyllium (hydrophilic mucilloid) in eliminating sand from the colon, it has been reported to alleviate sand accumulation in one filly with sand-induced diarrhea (25–50 g of psyllium can be administered daily to a 50-kg foal).34 A combination of mineral oil and psyllium has been shown to promote evacuation of sand from adult horses compared with mineral alone.36 In severe cases where refractory abdominal pain or distension is present, foals with impactions due to pica may require surgical removal of the foreign material from the GI tract.
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Diarrhea due to nutritional causes and lactose intolerance Dietary causes of neonatal foal diarrhea include overnursing or feeding in foals not ready to handle large quantities of milk, including those with prematurity or dysmaturity, septic or SIRS shock, or severe metabolic derangements including hypotension, hypoperfusion, hypothermia, or hypoxemia. Ingestion of milk in these intolerant foals can result in an osmotic diarrhea due to excessive amounts of ingested whey or lactose and failure of complete absorption. Milk replacers can also cause an osmotic diarrhea, especially if of poor quality. Ingestion of large amounts of fibrous feedstuffs, such as grain or forage, can also lead to diarrhea in neonatal foals because of inadequate digestion and subsequent influx of water into the colon. Lactose intolerance can also contribute to an osmotic diarrhea. Transient lactase deficiency has been recognized secondary to diffuse small intestinal damage from rotaviral and clostridial enteritis in foals.15, 19 A recent report suspected primary lactose intolerance in two foals; these foals responded well to administration of exogenous lactase enzyme.37 Lactose intolerance potentially is also a problem in premature foals, as the ability to secrete lactase is a relatively late development. Although lactase is detectable in the equine fetal intestine by the third month of gestation, maximal levels are not reached until birth.38 Lactase production steadily declines after 4 months of age until weaning. Affected foals are treated with exogenous lactase (6000 IU q. 3–6 hours).
vision of calories, with no or only small amounts of milk fed for provision of local nutrition to enterocytes. Foals with mucosal injury are at risk for bacterial translocation across the GI wall; this warrants the use of prophylactic, broad-spectrum systemic antimicrobials. Foals with asphyxial injury can be monitored for progress of GI function (motility, diameter, wall thickness) through the use of serial abdominal ultrasonography. Necrotizing enterocolitis Premature foals are at risk for developing necrotizing enterocolitis (NEC). Clinical features of NEC in human infants may be non-specific and mimic generalized sepsis (SIRS, lethargy), or localize to the gastrointestinal tract with abdominal dissension, bloody stool, and postprandial gastric residuals. Radiographic evidence of gas within the bowel wall may be present in severe cases, reflecting bowel wall compromise, and is termed pneumatosis intestinalis. NEC has also been reported in foals.39 Both foals in this report exhibited ileus, gastric reflux, and pneumatosis intestinalis on radiography. Both also developed perforations of the colon. Medical therapy of NEC in human infants includes the following measures: parenteral nutrition, gastric decompression, cessation of enteral feeding, systemic antimicrobials, and general supportive measures (fluid and electrolyte management). In severe cases surgical resection of devitalized intestine may be required. Similar treatments are generally accepted for use in foals.
Perinatal asphyxia-associated diarrhea
Sepsis-associated diarrhea
GI dysfunction, including diarrhea, is a clinical feature of neonatal maladjustment syndrome or hypoxic–ischemic injury. Mucosal hypoxia or hypoperfusion and subsequent reperfusion injury can cause dysmotility, enterocyte necrosis or apoptosis, and inflammation. Clinical signs of GI injury include gastroduodenal reflux, ileus, intolerance of feeding, colic, abdominal distension, and diarrhea. The diarrhea ranges from mild to severe; foals subjected to severe or prolonged hypoxic–ischemic injury may develop hemorrhagic or necrotizing diarrhea. It should be noted that foals with peripartum asphyxial injury usually suffer varying degrees of multiple organ injury, especially neurological and renal dysfunction. Foals with asphyxia-associated intestinal dysfunction should be fed conservatively (small, frequent meals) with gradual advancement of feeding volumes. They should be closely monitored for gastric reflux prior to each feeding. Severely affected foals benefit from parenteral nutrition as the primary pro-
SIRS, especially sepsis, is commonly associated with diarrhea in foals. The pathogenesis of the diarrhea in these cases likely is altered motility induced by cytokines, as well as hemodynamic perturbations leading to underperfusion of the gut with subsequent mucosal injury. In most cases the diarrhea subsides as the SIRS diminishes. In some cases, bacterial agents, such as anaerobes or Gram-negative enteric organisms, may be associated with the diarrhea. For example, foals with sepsis due to Actinobacillus spp. are more likely to have diarrhea than those with sepsis associated with other bacterial agents.40
References 1. Traub-Dargatz JL, Gay CC, Evermann JF, Ward ACS, Zeglen ME, Gallina AM, Salman MD. Epidemiologic survey of diarrhea in foals. J Am Vet Med Assoc 1988;192:1553–6. 2. Cohen ND. Causes of and farm management factors associated with disease and death in foals. J Am Vet Med Assoc 1994;204:1644–51.
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Neonatal Diarrhea 3. Raisis AL, Hodgson JL, Hodgson DR. Equine neonatal septicemia: 24 cases. Aust Vet J 1996;73:137–40. 4. Steel CM, Hunt AR, Adams PL, Robertson ID, Chicken C, Yovich JV, Stick JA. Factors associated with prognosis for survival and athletic use in foals with septic arthritis: 93 cases (1987–1994). J Am Vet Med Assoc 1999;215:973–7. 5. Breshears MA, Holbrook TC, Haak CE, York PA. Pulmonary aspergillosis and ischemic distal limb necrosis associated with enteric salmonellosis in a foal. Vet Pathol 2007;44:215–17. 6. Peek SF, Semrad S, McGuirk SM, Riseberg A, Slack JA, Marques F, Coombs D, Lien L, Keuler N, Darien BJ. Prognostic value of clinicopathologic variables obtained at admission and effect of antiendotoxin plasma on survival in septic and critically ill foals. J Vet Intern Med 2006;20:569–74. 7. McTaggart C, Penhale J, Raidala SL. Effect of plasma transfusion on neutrophil function in healthy and septic foals. Aust Vet J 2005;83:499–505. 8. Durando MM, Mackay RJ, Linda S, Skelley LA. Effects of polymyxin B and Salmonella typhimurium antiserum on horses given endotoxin intravenously. Am J Vet Res 1994;55:921–7. 9. East LM, Savage CJ, Traub-Dargatz JL, Dickinson CE, Ellis RP. Enterocolitis associated with Clostridium perfringens infection in neonatal foals: 54 cases (1988–1997). J Am Vet Med Assoc 1998;212:1751–6. 10. Magdesian KG, Hirsh DC, Jang SS, Hansen LM, Madigan JE. Characterization of Clostridium difficile isolates from foals with diarrhea: 28 cases (1993–1997). J Am Vet Med Assoc 2002;220:67–73. 11. Magdesian KG. Neonatal diarrhea. In: Paradis MR (ed.) Equine Neonatal Medicine. Philadelphia: Elsevier Saunders, 2006; pp. 213–21. 12. Gerding DN, Muto CA, Owens RC Jr. Treatment of Clostridium difficile infection. Clin Infect Dis 2008;46 Suppl 1: S32–42. 13. Weese JS, Cote NM, deGannes RV. Evaluation of in vitro properties of di-tri-octahedral smectite on clostridial toxins and growth. Equine Vet J 2003;35:638–41. 14. Lawler JB, Hassel DM, Magnuson RJ, Hill AE, McCue PM, Traub-Dargatz JL. Adsorptive effects of di-tri-octahedral smectite on Clostridium perfringens alpha, beta, and beta2 exotoxins and equine colostral antibodies. Am J Vet Res 2008;69:233–9. 15. Weese JS, Parsons DA, Staempfli HR. Association of Clostridium difficile with enterocolitis and lactose intolerance in a foal. J Am Vet Med Assoc 1999;214:229–32. 16. Holland RE, Sriranganathan N, DuPont L. Isolation of enterotoxigenic Escherichia coli from a foal with diarrhea. J Am Vet Med Assoc 1989;194:389–91. 17. Barrandeguy M, Parreno V, Lagos Marmol M, Pont Lezica F, Rivas C, Valle C, Fernandez F. Prevention of rotavirus diarrhoea in foals by parenteral vaccination of the mares: field trail. Dev Biol Stand 1998;92: 253–7. 18. Powell DG, Dwyer RM, Traub-Dargatz JL, Fulker RH, Whalen JW Jr, Srinivasappa J, Acree WM, Chu HJ. Field study of the safety, immunogenicity, and efficacy of an inactivated equine rotavirus vaccine. J Am Vet Med Assoc 1997;211:193–8. 19. Sweeney RW. Laboratory evidence of malassimilation in horses. Vet Clin North Am Equine Pract 1987;3:507–14. 20. Watanabe T, Ohta C, Shirahata T, Goto H, Tsunoda N, Tagami M, Akita H. Preventive administration of bovine colostral immunoglobulins for foal diarrhea with rotavirus. J Vet Med Sci 1993;55:1039–40.
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21. Mitra AK, Maahalanabis D, Ashraf H, Unicomb L, Eeckels R, Tzipori S. Hyperimmune cow colostrum reduces diarrhoea due to rotavirus: a double-blind, controlled trial. Acta Paediatr 1995;84:996–1001. 22. Rossignol JF, Abu-Zekry M, Hussein A, Santoro MG. Effect of nitazoxanide for treatment of severe rotavirus diarrhoea: randomized double-blind placebo-controlled trial. Lancet 2006;368:124–9. 23. Rossignol JF, El-Gohary YM. Nitazoxanide in the treatment of viral gastroenteritis: a randomized double-blind placebo-controlled clinical trial. Aliment Pharmacol Ther 2006;24:1423–30. 24. Davis E, Rush BR, Cox J, DeBey B, Kapil S. Neonatal enterocolitis associated with coronavirus infection in a foal: a case report. J Vet Diagn Invest 2000;12:153–6. 25. Guy JS, Breslin JJ, Breuhaus B, Vivrette S, Smith LG. Characterization of a coronavirus isolated from a diarrheic foal. J Clin Microbiol 2000;38:4523–6. 26. Grinberg A, Learmonth J, Kwan E, Pomroy W, Lopez Villalobos N, Gibson I, Widmer G. Genetic diversity and zoonotic potential of Cryptosporidium parvum causing foal diarrhoea. J Clin Microbiol 2008;136:273–8. 27. Netherwood T, Wood JLN, Townsend HG, Mumford JA, Chanter N. Foal diarrhoea between 1991 and 1994 in the United Kingdom associated with Clostridium perfringens, rotavirus, Strongyloides westeri, and Cryptosporidium sp. Epidemiol Infect 1996;117:375–83. 28. Masri MD, Merritt AM, Gronwall R, Burrows CF. Faecal composition in foal-heat diarrhea. Equine Vet J 1986;18:301–6. 29. Magdesian KG. Neonatal foal diarrhea. Vet Clin North Am Equine Pract 2005;21:295–312. 30. Stowe HD. Automated orphan foal feeding. In: Proceedings of the 13th Annual Meeting of the American Association of Equine Practice, 1967, pp. 65–73. 31. Johnston RH, Kamstra LD, Kohler PH. Mares’ milk composition as related to ‘foal heat’ scours. J Anim Sci 1970;31:549–53. 32. Harris SE, Vogelsang MM, Potter GD, Bass EE. The effects of ivermectin given postpartum on the incidence and severity of foal-heat diarrhea. Prof Anim Sci 2004;20:372–5. 33. Sgorbini M, Nardoni S, Mancianti F, Rota A, Corazza M. Foal-heat diarrhea is not caused by the presence of yeasts in the gastrointestinal tract of foals. J Equine Vet Sci 2008;28:145–8. 34. Ramey DW, Reinertson EL. Sand-induced diarrhea in a foal. J Am Vet Med Assoc 1984;185: 537–8. 35. Korolainen R, Kaikkonen R, Ruohoniemi M. Ultrasonography in monitoring the resolution of intestinal sand accumulations in the horse. Equine Vet Educ 2003;5:423–32. 36. Hotwagner K, Igen C. Evacuation of sand from the equine intestine with mineral oil, with and without psyllium. J Anim Physiol Anim Nutr (Berl) 2008;92:86–91. 37. Roberts VLH, Knottenbelt DC, Williams A, McKane SA. Suspected primary lactose intolerance in neonatal foals. Equine Vet Educ 2008;20:249–51. 38. Roberts MC. The development and distribution of mucosal enzymes in the small intestine of the fetus and the young foal. Reprod Fertil Suppl 1975;23:717–23. 39. Cudd T, Pauly TH. Necrotizing enterocolitis in two equine neonates. Comp Cont Educ Pract Vet 1986;9:88–96. 40. Stewart AJ, Hinchcliff KW, Saville WJ, Jose-Cunilleras E, Hardy J, Kohn CW, Reed SM, Kowalski JJ. Actinobacillus sp. Bacteremia in foals: clinical signs and prognosis. J Vet Intern Med 2002;16:464–71.
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Chapter 41
Gastroduodenal Ulceration: The Neonatal Perspective L. Chris Sanchez
Pathophysiology In all species, gastric mucosal ulceration typically results from an imbalance between inciting and protective factors in the mucosal environment.1, 2 In most forms of the equine gastric ulcer syndrome, gastric acid is the major determining factor, but, in the sick neonate, other factors such as gastric mucosal perfusion may be more relevant. The major intrinsic factors promoting ulcer formation include hydrochloric acid (HCl), bile acids, and pepsin, with HCl the predominant factor. Intrinsic factors protecting against ulcer formation include the mucus–bicarbonate layer, adequate mucosal blood flow, mucosal prostaglandin E2 and epidermal growth factor production, and gastroduodenal motility. Foals can produce significant amounts of gastric acid by the second day of life, with consistent periods of acidity (pH < 2.0) in clinically normal animals.3, 4 In one study, foals tended to have a high gastric pH at 1 day old,3 but, in a study of critically ill foals, some foals demonstrated periods of gastric acidity on the first day of life.5 Suckling was associated with an immediate rise in gastric pH, whereas periods of rest during which foals did not suck for more than 20 minutes were associated with prolonged periods of acidity.4 Premature human infants are capable of gastric acid production at 28 weeks of gestation.6 In sheep, abomasal pH begins to drop at approximately 125 days of gestation, but does not reach values normal for adults (pH = 2.5) until after birth.7 Only one out of seven premature foals demonstrated an acidic pH recording in a study of gastric pH profiles in critically ill foals.5 While multiple factors were likely involved in those foals, the true ontogeny of gastric acid production in foals is currently unknown.
Equine squamous mucosa is very thin at birth, but becomes hyperplastic and parakeratotic within days.8 The parallel between decreasing pH and proliferation of squamous epithelium correlates with that seen in other species.9 The combination of a relatively thin gastric epithelium with a high acid output may leave neonatal foals susceptible to ulcer formation at a very young age. In addition, one must remember the difference in normal appearance of the squamous mucosa when interpreting gastric endoscopy in a neonatal population. In esophageal squamous mucosa, intercellular tight junctions and bicarbonate secretion are the major factors involved in protection against squamous acid injury in other species, although squamous bicarbonate secretion has not been documented in the horse.10–12 The principal barrier is a glycoconjugate substance secreted by cells in the stratum spinosum, with a contribution from the tight junctions in the stratum corneum.12 This barrier function is considered weak at best. In foals, the squamous mucosa adjacent to the margo plicatus likely receives exposure to acidic gastric contents on a regular basis. Recently, surface mucus demonstrating cross-reactivity to a mixture of anti-human MUC5AC (a mucin gene) antibodies has been identified in both the glandular and non-glandular regions of healthy equine gastric mucosa.13 Bile salts and pepsin have been implicated as contributing factors to ulcer disease in many species. In the adult horse, a synergistically damaging effect was found with the addition of bile salts and acid (pH 2.5) to stratified squamous mucosa in vitro in one study.14 In another study, prolonged exposure to acid alone (pH 1.5) had a damaging effect, and synergism with exposure to a combination of acid and pepsin
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Gastroduodenal Ulceration: The Neonatal Perspective or taurocholate was not found.15 In addition, several volatile fatty acids (acetic, butyric, and proprionic) decrease non-glandular mucosal barrier function,16, 17 whereas lactic acid exposure alone does not.18 Whether or not these factors play a role in neonatal squamous ulcer disease remains speculative. Several mechanisms help protect the glandular mucosa from acid injury. The mucus–bicarbonate layer serves to titrate H+ ions from the gastric lumen to CO2 and H2 O. Cellular restitution and prostaglandins of the E series, which enhance mucosal blood flow and mucus–bicarbonate secretion in the glandular mucosa, have not been documented in squamous epithelium.11, 19 Of these mechanisms, mucosal blood flow is likely the most important contributor to overall gastric mucosal health. Nitric oxide is a key regulator of mucosal blood flow and prostaglandin synthesis and thus may play a role in mucosal protection.20
Clinical syndrome Clinical signs typically associated with gastric ulceration in foals include poor appetite, diarrhea, and colic. Many foals probably never exhibit clinical signs, and some do not exhibit clinical signs until ulceration is severe or fatal perforation has occurred. Glandular ulceration is typically considered the most clinically significant type of disease in this population. The physiological stress of a concurrent illness has been associated with gastric ulceration in foals. Retrospectively, 14 (23%) of 61 foals ≤ 85 days of age with a clinical disorder were found to have lesions in the gastric glandular mucosa,21 and prospectively eight (40%) of 20 foals ≤ 30 days of age with a clinical disorder had glandular ulceration.22 By contrast, only 4–9% of clinically normal foals examined in endoscopic surveys had lesions observed in the gastric glandular mucosa.23, 24 Critically ill neonatal foals can have a markedly different pH profile from that seen in clinically normal foals, potentially because of alterations in gastric motility and acid secretion.5 Gastric ulceration was not identified in any animals at necropsy in that study; however, ulceration has been documented in a similar population.25 Thus, factors other than acid exposure, most notably mucosal perfusion, may play an important role in the ‘stress’-related ulceration seen in neonates. Gastric ulceration and rupture in the hospitalized neonatal population appears to occur less commonly now than in previous reports, despite a decline in the use of ulcer prophylaxis in one hospital.26 Advances in overall neonatal care, especially supportive care, have likely contributed to this decline.
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One recent report evaluated the use of gastric tonometry as a measurement of mucosal perfusion in neonatal foals.27 However, despite investigations into the effect of age, feeding, and omeprazole administration, the reference interval in foals appears too wide to warrant clinical application of this tool to evaluate the relationship between ulceration and mucosal perfusion in foals.
Treatment and prevention Multiple pharmacological treatments have been suggested for the treatment of ulcer disease. Because acid has been implicated as the most important pathophysiological component of squamous ulcer disease, most anti-ulcer therapy centers on suppression or neutralization of gastric acid. The principal therapeutic options for ulcer treatment include H2 -receptor antagonists (cimetidine, ranitidine, famotidine, nizatidine), proton pump inhibitors (PPIs; omeprazole, pantoprazole, rabeprazole, esomeprazole), the mucosal adherent sucralfate, and antacids. The H2 antagonists suppress hydrochloric acid secretion through competitive inhibition of the parietal cell histamine receptor that can be partially overcome with exogenous pentagastrin.28 Use of H2 antagonists has been successful in raising gastric pH and resolving gastric lesions in both foals and adult horses.4, 29, 30 Clinical and experimental evidence has demonstrated greater individual variability with lower dosages of H2 antagonists.31 Thus, dosage recommendations are based upon levels necessary to increase gastric pH and promote ulcer healing in a majority of horses. Commonly recommended dosages are 20–30 mg/kg orally every 8 hours or 6.6 mg/kg intravenously (i.v.) every 6 hours for cimetidine and 6.6 mg/kg orally every 8 hours or 1.5–2 mg/kg i.v. every 6 hours for ranitidine (Table 41.1). Although clinically normal foals respond predictably to ranitidine,4 sick neonates have shown variability in pH response to intravenous ranitidine, with a much shorter duration of action and, in some cases, no noticeable response.5 Currently, cimetidine and ranitidine are available in injectable, tablet, and liquid forms. PPIs block secretion of H+ at the parietal cell membrane by irreversibly binding to the H+ ,K+ -ATPase proton pump of the cell. These agents have a prolonged antisecretory effect, which allows for oncedaily dosing. Omeprazole (4 mg/kg orally) also has demonstrated efficacy for raising intragastric pH in clinically normal32 or critically ill33 foals and for ulcer healing in foals.34 Treatment with 1, 2 or 4 mg/kg orally every 24 hours has been shown to decrease or prevent disease or the recurrence of
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Foal: Gastrointestinal System
Table 41.1 Treatment options for gastric ulcers in foals
Drug Ranitidine Ranitidine Cimetidine Cimetidine Misoprostol Omeprazole Omeprazole Pantoprazole Sucralfate Al/Mg antacids MgOH∗
Dose (mg/kg)
Dosing interval (h)
Route of administration
6.6 1.5–2 20–25 6.6 0.005 2–4 0.5 1.5 20–40 60 AlOH/30
8 6–8 8 6–8 8 24 24 24 8 4
p.o. i.v., i.m. p.o. i.v., i.m. p.o. p.o. i.v. i.v., p.o. p.o. p.o.
R This equates to 0.5 ml/kg of Maalox TC, but the concentration of aluminum hydroxide and magnesium hydroxide can vary by product. p.o., per os; i.v., intravenously; i.m., intramuscularly.
∗
disease in adult horses maintained in training.35–38 The powder form of omeprazole is rapidly degraded in an acidic environment, thus an entericcoated capsule (as used in the human preparation) or a specially formulated paste must be used to allow delivery of the active drug to the small intestine for absorption. Compounded preparations have shown limited to no efficacy in pharmacodynamic and clinical trials,39, 40 and the proprietary formulation of omeprazole was the only product (including buffers, H2 -receptor antagonists, sucralfate, and compounded preparations) that decreased the odds of gastric ulceration in a population of racehorses.41 A compounded intravenous preparation of omeprazole (0.5 mg/kg) has been shown to raise gastric juice pH and decrease the number of non-glandular lesions in horses.42 Pantoprazole, another PPI available commercially for use in humans, given by either intravenous or intragastric routes (1.5 mg/kg) raised intragastric pH in clinically normal neonatal foals.43 Sucralfate has been used for treatment of gastric ulcers and prevention of ‘stress-induced’ ulcers in humans. The mechanism of action likely involves adherence to ulcerated mucosa, stimulation of mucus secretion, enhanced prostaglandin E synthesis, and concentration of growth factor at the site of ulceration, although the prostaglandin effects may not play an important role in ulcer healing.44 In one study, sucralfate did not promote subclinical ulcer healing in foals, relative to corn syrup.45 In another trial, simultaneous sucralfate administration significantly reduced the total area of oral ulceration and gastric epithelial necrosis experimentally induced by phenylbutazone, but did not significantly reduce the number of squamous or glandular ulcers
in foals.46 In humans, sucralfate provides protection against stress-induced ulcers with a decreased risk of pathogenic gastric colonization.47 The current recommended dose of sucralfate is 10–20 mg/kg every 6–8 hours. The efficacy of sucralfate in an alkaline pH is controversial, but appears likely.48–50 The use of antacids in the treatment of gastric ulcers has not been critically examined in the horse. Research in horses has shown that 30 g aluminum hydroxide/15 g magnesium hydroxide will result in an increase in gastric pH above 4 for approximately 2 hours.51 The use of synthetic prostaglandin E1 analogs, such as misoprostol, has been effective in the treatment of gastric and duodenal ulcers in humans, and the proposed mechanism of action involves both inhibition of gastric acid secretion and mucosal cytoprotection.52 In horses, misoprostol (5 µg/kg) has been shown to increase gastric pH53 and ameliorate deleterious effects of flunixin on mucosal recovery after ischemic injury in vitro.54 Prokinetic drugs should be considered in foals with duodenal disease, gastroesophageal reflux, and when delayed gastric emptying without a physical obstruction is suspected. Most of the cited work has been performed in adult horses; hence, recommendations are extrapolated for use in foals. Bethanechol and erythromycin have been shown to increase the rate of gastric emptying in horses.55 In cases of acute gastric atony, bethanechol 0.025–0.030 mg/kg subcutaneously (s.c.) every 3–4 hours has been effective in promoting gastric motility and emptying, followed by oral maintenance dosages of 0.35–0.45 mg/kg three or four times daily. Adverse effects can include diarrhea, inappetance, salivation, and colic, but, at the dosages stated, adverse effects have been infrequent and mild. A constant rate infusion of intravenous lidocaine provides another option with potential analgesic and prokinetic properties.56, 57 Because gastric perforation due to glandular ulcer disease has been reported in hospitalized neonates, many clinicians routinely use prophylactic anti-ulcer therapy in this population. Because some critically ill foals have a predominantly alkaline gastric pH profile, and because gastric acidity may be protective against bacterial translocation in neonates, the need for prophylactic ulcer therapy is controversial. In critically ill human neonates, while intravenous ranitidine therapy raises gastric pH and increases gastric bacterial colonization, it does not increase the risk of sepsis.58 In a retrospective study of 85 hospitalized foals < 30 days old, no difference in the frequency of gastric ulceration at necropsy was found between those foals that received prophylactic treatment for gastric ulcers and those that did not.26 Owing to the retrospective nature of that study, specific details
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Gastroduodenal Ulceration: The Neonatal Perspective regarding lesion location and severity were not available; however, none of the foals in the study died as a result of gastric ulcer disease. Thus, many clinicians, including the author, no longer recommend routine ulcer prophylaxis in all ill neonates. The major exceptions are those foals with a significant focus of pain, such as those with infectious (septic arthritis, osteomyelitis, etc.) or non-infectious (flexural deformity, etc.) orthopedic disorders or requiring abdominal surgery, as those foals typically require significant non-steroidal anti-inflammatory drug therapy. The use of prophylactic anti-acid therapy in older suckling foals with respiratory or gastrointestinal disease requires separate consideration, but is beyond the age-specific scope of the current chapter.
References 1. Nappert G, Vrins A, Larybyere M. Gastroduodenal ulceration in foals. Comp Contin Ed 1989;11:345. 2. Murray MJ, Grodinsky C. Regional gastric pH measurement in horses and foals. Equine Vet J Suppl 7, 1989;73–6. 3. Baker SJ, Gerring EL. Gastric pH monitoring in healthy, suckling pony foals. Am J Vet Res 1993;54:959–64. 4. Sanchez LC, Lester GD, Merritt AM. Effect of ranitidine on intragastric pH in clinically normal neonatal foals. J Am Vet Med Assoc 1998;212:1407–12. 5. Sanchez LC, Lester GD, Merritt AM. Intragastric pH in critically ill neonatal foals and the effect of ranitidine. J Am Vet Med Assoc 2001;218:907–11. 6. Kuusela AL. Long-term gastric pH monitoring for determining optimal dose of ranitidine for critically ill preterm and term neonates. Arch Dis Child Fetal Neonatal Ed 1998;78:F151–3. 7. Read MA, Chick P, Hardy KJ, Shulkes A. Ontogeny of gastrin, somatostatin, and the H+/K(+)-ATPase in the ovine fetus. Endocrinology 1992;130:1688–97. 8. Murray MJ, Mahaffey EA. Age-related characteristics of gastric squamous epithelial mucosa in foals. Equine Vet J 1993;25:514–17. 9. De Backer A, Haentjens P, Willems G. Hydrochloric acid. A trigger of cell proliferation in the esophagus of dogs. Dig Dis Sci 1985;30:884–90. 10. Tobey NA, Orlando RC. Mechanisms of acid injury to rabbit esophageal epithelium. Role of basolateral cell membrane acidification. Gastroenterology 1991;101:1220–28. 11. Orlando RC. Esophageal epithelial defense against acid injury. J Clin Gastroenterol 1991;13 Suppl 2:S1–5. 12. Orlando RC, Lacy ER, Tobey NA, Cowart K. Barriers to paracellular permeability in rabbit esophageal epithelium. Gastroenterology 1992;102:910–23. 13. Bullimore SR, Corfield AP, Hicks SJ, Goodall C, Carrington SD. Surface mucus in the non-glandular region of the equine stomach. Res Vet Sci 2001;70:149–55. 14. Berschneider HM, Blikslager AT, Roberts MC. Role of duodenal reflux in nonglandular gastric ulcer disease of the mature horse. Equine Vet J Suppl 29, 1999;24–9. 15. Widenhouse TV, Lester GD, Merritt AM. The effect of hydrochloric acid, pepsin, or taurocholate on the bioelectric properties of gastric squamous mucosa in horses. Am J Vet Res 2002;63(5):744–9.
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16. Nadeau JA, Andrews FM, Mathew AG, Argenzio RA, Blackford JT, Sohtell M, Saxton AM. Evaluation of diet as a cause of gastric ulcers in horses. Am J Vet Res 2000; 61:784–90. 17. Nadeau JA, Andrews FM, Patton CS, Argenzio RA, Mathew AG, Saxton AM. Effects of hydrochloric, acetic, butyric, and propionic acids on pathogenesis of ulcers in the nonglandular portion of the stomach of horses. Am J Vet Res 2003;64:404–12. 18. Andrews FM, Buchanan BR, Elliott SB, Al Jassim RA, McGowan CM, Saxton AM. In vitro effects of hydrochloric and lactic acids on bioelectric properties of equine gastric squamous mucosa. Equine Vet J 2008;40(4):301–5. 19. Argenzio RA. Comparative pathophysiology of nonglandular ulcer disease: a review of experimental studies. Equine Vet J Suppl 29, 1999;19–23. 20. Konturek PC, Brzozowski T, Sliwowski Z, Pajdo R, Stachura J, Hahn EG, Konturek SJ. Involvement of nitric oxide and prostaglandins in gastroprotection induced by bacterial lipopolysaccharide. Scand J Gastroenterol 1998;33:691–700. 21. Murray MJ. Endoscopic appearance of gastric lesions in foals: 94 cases (1987–1988). J Am Vet Med Assoc 1989;195: 1135–41. 22. Furr MO, Murray MJ, Ferguson DC. The effects of stress on gastric ulceration, T3, T4, reverse T3 and cortisol in neonatal foals. Equine Vet J 1992;24:37–40. 23. Murray MJ, Murray CM, Sweeney HJ, Weld J, Digby NJ, Stoneham SJ. Prevalence of gastric lesions in foals without signs of gastric disease: an endoscopic survey. Equine Vet J 1990;22:6–8. 24. Murray MJ, Grodinsky C, Cowles RR, Hawkins WL, Forfa RJ, Luba NK. Endoscopic evaluation of changes in gastric lesions of Thoroughbred foals. J Am Vet Med Assoc 1990;196:1623–7. 25. Wilson JH. Gastric and duodenal ulcers in foals: a retrospective study. In: Proceedings of the Second Equine Colic Research Symposium, Athens, Georgia, 1–3 October, 1985, pp. 126–8. 26. Barr BS, Wilkins PA, Del Piero F, Palmer JE. Is prophylaxis for gastric ulcers necessary in critically ill equine neonates? A retrospective study of necropsy cases 1989–1999. J Vet Intern Med 2000;14:328. 27. Sanchez LC, Giguere S, Javsicas LH, Bier J, Walrond CJ, Womble AY. Effect of age, feeding, and omeprazole administration on gastric tonometry in healthy neonatal foals. J Vet Intern Med 2008;22:406–10. 28. Campbell-Thompson ML, Merritt AM. Effect of ranitidine on gastric acid secretion in young male horses. Am J Vet Res 1987;48:1511–15. 29. Becht JL, Byars TD. Gastroduodenal ulceration in foals. Equine Vet J 1986;18:307–12. 30. Furr MO, Murray MJ. Treatment of gastric ulcers in horses with histamine type 2 receptor antagonists. Equine Vet J Suppl 7, 1989;77–9. 31. Murray MJ, Grodinsky C. The effects of famotidine, ranitidine and magnesium hydroxide/aluminium hydroxide on gastric fluid pH in adult horses. Equine Vet J Suppl 11, 1992;52–55. 32. Sanchez LC, Murray MJ, Merritt AM. Effect of omeprazole paste on intragastric pH in clinically normal neonatal foals. Am J Vet Res 2004;65:1039–41. 33. Javsicas LH, Sanchez LC. The effect of omeprazole paste on intragastric pH in clinically ill neonatal foals. Equine Vet J 2008;40:41–4.
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34. MacAllister CG, Sifferman RL, McClure SR, White GW, Vatistas NJ, Holste JE, Ericcson GF, Cox JL. Effects of omeprazole paste on healing of spontaneous gastric ulcers in horses and foals: a field trial. Equine Vet J Suppl 29, 1999;77–80. 35. Andrews FM, Sifferman RL, Bernard W, Hughes FE, Holste JE, Daurio CP, Alva R, Cox JL. Efficacy of omeprazole paste in the treatment and prevention of gastric ulcers in horses. Equine Vet J Suppl 1999;81–6. 36. McClure SR, White GW, Sifferman RL, Bernard W, Doucet MY, Vrins A, Holste JE, Fleishman C, Alva R, Cramer LG. Efficacy of omeprazole paste for prevention of gastric ulcers in horses in race training. J Am Vet Med Assoc 2005;226:1681–4. 37. McClure SR, White GW, Sifferman RL, Bernard W, Hughes FE, Holste JE, Fleishman C, Alva R, Cramer LG. Efficacy of omeprazole paste for prevention of recurrence of gastric ulcers in horses in race training. J Am Vet Med Assoc 2005;226:1685–8. 38. White G, McClure SR, Sifferman R, Holste JE, Fleishman C, Murray MJ, Cramer LG. Effects of short-term light to heavy exercise on gastric ulcer development in horses and efficacy of omeprazole paste in preventing gastric ulceration. J Am Vet Med Assoc 2007;230:1680–2. 39. Merritt AM, Sanchez LC, Burrow JA, Church M, Ludzia S. Effect of GastroGard and three compounded oral omeprazole preparations on 24 h intragastric pH in gastrically cannulated mature horses. Equine Vet J 2003;35: 691–5. 40. Nieto JE, Spier S, Pipers FS, Stanley S, Aleman MR, Smith DC, Snyder JR. Comparison of paste and suspension formulations of omeprazole in the healing of gastric ulcers in racehorses in active training. J Am Vet Med Assoc 2002;221:1139–43. 41. Orsini JA, Haddock M, Stine L, Sullivan EK, Rabuffo TS, Smith G. Odds of moderate or severe gastric ulceration in racehorses receiving antiulcer medications. J Am Vet Med Assoc 2003;223:336–9. 42. Andrews FM, Frank N, Sommardahl CS, Buchanan BR, Elliott SB, Allen VA. Effects of intravenously administrated omeprazole on gastric juice pH and gastric ulcer scores in adult horses. J Vet Intern Med 2006;20:1202–6. 43. Ryan CA, Sanchez LC, Giguere S, Vickroy T. Pharmacokinetics and pharmacodynamics of pantoprazole in clinically normal neonatal foals. Equine Vet J 2005;37:336–41. 44. Ogihara Y, Okabe S. Effect and mechanism of sucralfate on healing of acetic acid-induced gastric ulcers in rats. J Physiol Pharmacol 1993;44:109–18.
45. Borne AT, MacAllister CG. Effect of sucralfate on healing of subclinical gastric ulcers in foals. J Am Vet Med Assoc 1993;202:1465–8. 46. Geor RJ, Petrie L, Papich MG, Rousseaux C. The protective effects of sucralfate and ranitidine in foals experimentally intoxicated with phenylbutazone. Can J Vet Res 1989;53:231–8. 47. Ephgrave KS, Kleiman-Wexler R, Pfaller M, Booth BM, Reed D, Werkmeister L, Young S. Effects of sucralfate vs antacids on gastric pathogens: results of a double-blind clinical trial. Arch Surg 1998;133:251–7. 48. Danesh BJ, Duncan A, Russell RI. Is an acid pH medium required for the protective effect of sucralfate against mucosal injury? Am J Med 1987;83:11–13. 49. Konturek SJ, Brzozowski T, Mach T, Konturek WJ, Bogdal J, Stachura J. Importance of an acid milieu in the sucralfateinduced gastroprotection against ethanol damage. Scand J Gastroenterol 1989;24:807–12. 50. Danesh JZ, Duncan A, Russell RI, Mitchell G. Effect of intragastric pH on mucosal protective action of sucralfate. Gut 1988;29:1379–85. 51. Clark CK, Merritt AM, Burrow JA, Steible CK. Effect of aluminum hydroxide/magnesium hydroxide antacid and bismuth subsalicylate on gastric pH in horses. J Am Vet Med Assoc 1996;208:1687–91. 52. Leandro G, Pilotto A, Franceschi M, Bertin T, Lichino E, Di Mario F. Prevention of acute NSAID-related gastroduodenal damage: a meta- analysis of controlled clinical trials. Dig Dis Sci 2001;46:1924–36. 53. Sangiah S, MacAllister CC, Amouzadeh HR. Effects of misoprostol and omeprazole on basal gastric pH and free acid content in horses. Res Vet Sci 1989;47:350–4. 54. Tomlinson JE, Blikslager AT. Effects of cyclooxygenase inhibitors flunixin and deracoxib on permeability of ischaemic-injured equine jejunum. Equine Vet J 2005;37:75–80. 55. Ringger NC, Lester GD, Neuwirth L, Merritt AM, Vetro T, Harrison J. Effect of bethanechol or erythromycin on gastric emptying in horses. Am J Vet Res 1996;57:1771–5. 56. Robertson SA, Sanchez LC, Merritt AM, Doherty TJ. Effect of systemic lidocaine on visceral and somatic nociception in conscious horses. Equine Vet J 2005;37:122–7. 57. Malone E, Ensink J, Turner T, Wilson J, Andrews F, Keegan K, Lumsden J. Intravenous continuous infusion of lidocaine for treatment of equine ileus. Vet Surg 2006;35:60–6. 58. Cothran DS, Borowitz SM, Sutphen JL, Dudley SM, Donowitz LG. Alteration of normal gastric flora in neonates receiving ranitidine. J Perinatol 1997;17:383–8.
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Chapter 42
Surgical Gastrointestinal Conditions Brett J. Woodie
Foals showing signs of abdominal pain or colic present a challenge in which multiple diagnostic modalities are utilized to determine the proper course of treatment. In the adult horse palpation per rectum is a vital diagnostic tool that is not available to the clinician evaluating a foal showing signs of abdominal pain. However, abdominal radiography and transabdominal ultrasonography are two ancillary diagnostic modalities that can provide the clinician with valuable information. Advances in anesthesia, surgical techniques and pre- and postoperative care have contributed to the increased success rate of abdominal surgery in foals.1–6 It is important to remember that rapid intervention when a surgical lesion is present is vital to a successful outcome.
History and physical examination A complete and thorough history and physical examination can provide important information in assessing a foal with abdominal pain. In neonatal foals an abdominal disorder may be related to other abnormalities such as prematurity, septicemia, or hypoxic ischemic encephalopathy.5, 6 Crucial information to obtain when taking the history includes the age of the foal, age at onset of the problem, duration, severity, and progression of clinical signs, fecal production, urine output, medication administered, and response to treatment. Information related to parturition is also important. The level of abdominal discomfort can be variable. The severity of abdominal pain that the foal is showing does not always correlate specifically with the severity of the disease. A thorough physical examination should be performed after obtaining historical information. Initially the foal is observed for signs of abdominal
pain and abdominal distention. In order to monitor abdominal distention the clinician can measure the diameter of the foal’s abdomen with a tape. It is important to measure the abdomen at the same location each time. The site for measurement is located just caudal to the last rib and can be marked by clipping hair from the dorsal and ventral aspects of the abdomen. Progressive abdominal distention often requires surgical intervention. Physical examination parameters include but are not limited to temperature, pulse, respiratory rate, mucous membrane color, capillary refill time, auscultation for borborygmi, digital examination of the rectum, palpation of the external umbilicus, palpation of the inguinal region, and passing a nasogastric tube to determine if reflux is present. Clinicopathological findings are an important component of the diagnostic assessment. Samples should be submitted for a complete blood count, serum lactate, serum chemistry profile, and possibly immunoglobulin determination (IgG) – depending on the age of the foal. Transabdominal ultrasonography should be performed following completion of the physical examination. A thorough and complete examination of abdominal viscera, umbilical structures, bladder, and the thoracic cavity can provide valuable information. The abdominal cavity should be evaluated for the amount of fluid present. The viscera should be evaluated for location, distention, presence of motility, and thickness of the intestinal wall.6 The urachus, umbilical vein, and arteries should be evaluated for size and presence of infection. The normal size of the umbilical remnants are as follows: urachus < 2 cm in diameter, umbilical vein < 0.8 cm in diameter, and the umbilical arteries < 1 cm in diameter. Enlargement of an umbilical remnant does not always indicate a septic process
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 42.1 Standing abdominal radiograph (lateral projection) showing severe gas distention of the gastrointestinal tract.
is present. Ultrasonography of the thoracic cavity can help determine if there are other abnormalities such as pneumonia, excessive pleural fluid, or a diaphragmatic hernia. Abdominal radiographs typically provide more diagnostic information in the foal compared to adults.5 The images can be obtained with the foal standing or in lateral recumbency. The radiographs should be evaluated for distention of the intestinal tract with fluid or gas, excessive abdominal fluid, free air in the abdominal cavity, and the positioning of the intestinal tract (see Figure 42.1). Contrast radiography may be necessary to document conditions such as a delayed gastric emptying secondary to an outflow obstruction. A barium enema can be used to diagnose meconium impactions or colonic atresia. Abdominocentesis is recommended as part of the baseline information to obtain when evaluating a foal for colic. Using ultrasonographic guidance will help the clinician select a suitable site of sampling and avoid distended viscera. It is very important to remember that normal peritoneal fluid analysis does not rule out intestinal disease or ischemic compromise. When formulating a treatment plan, the clinician should consider all physical examination findings, the laboratory data, ultrasonography findings, and radiographic findings. All of this information must be used together to determine if the appropriate course of action is immediate surgical intervention or medical therapy. Differentiating surgical versus medical therapy in a foal is often a difficult task. Persistent abdominal pain despite treatment is often the deciding factor to warrant surgery. However, progressive abdominal distention, abnormal ultrasonographic/radiographic findings, and/or abnormal or
changing abdominocentesis values may indicate the need for an exploratory laparotomy. In the author’s opinion early surgical intervention is paramount. Once the decision is made to perform an abdominal exploratory laparotomy, perioperative antibiotic, non-steroidal anti-inflammatory drug, and tetanus prophylaxis should be administered at the discretion of the clinician(s) prior to induction of general anesthesia. Following anesthetic induction the foal is positioned in dorsal recumbency on a well-padded table. Hypothermia is not uncommon in foals undergoing abdominal surgery. In order to prevent hypothermia an external heat source such as a heating pad or warm water blanket should be placed under the foal. Precautions should be taken to prevent burns. Warm intravenous fluids are administered as well. The ventral abdomen is clipped and aseptically prepared for surgery. The external umbilicus may be oversewn to facilitate aseptic preparation of the abdomen. In male foals a urinary catheter can be passed and the penis directed caudally to remove it from the surgical field. Following draping of the abdomen a ventral midline incision is made beginning cranial to the external umbilicus and extending craniad. The umbilical structures need not be removed as a routine part of the exploratory laparotomy unless there is a medical reason to do so. The abdominal wall is thin in foals and entry into the abdomen should be done cautiously to avoid inadvertent injury to a distended viscus. Once the abdomen is opened a thorough and systematic evaluation should be completed. Excessive handling and unnecessary manipulation of the intestinal tract should be avoided. This will help prevent iatrogenic mesenteric tears and possibly decrease the possibility of adhesion formation. Copious lavage with warm fluids during the exploratory is beneficial. Following the completion of the abdominal exploratory the author recommends an omentectomy as another adhesion prevention technique. Closure of the abdomen is as follows: the linea alba should be closed using #1 (or #2 depending on the size of the foal) synthetic absorbable suture material such as polyglactin 910 in an appropriate pattern such as simple continuous; the subcutaneous layer is closed using 2-0 synthetic absorbable suture material in an appropriate pattern; and the skin is apposed using 2-0 or O synthetic absorbable suture material such as poliglecaprone in an appropriate pattern such as a simple continuous. The incision is protected in the immediate postoperative period by using a stent. The stent should be removed 2 days after surgery or sooner if it becomes soiled. Proper medical therapy after surgery is vital to a successful outcome. This includes but is not limited to antibiotic therapy, anti-inflammatory medication(s), intravenous fluids, maintenance of
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Surgical Gastrointestinal Conditions serum electrolyte balance, anti-ulcer medication(s), anti-adhesion therapy, and parenteral nutrition.
Gastroduodenal ulceration Foals that are treated with non-steroidal antiinflammatory drugs or that are stressed often have gastric ulceration.6 These foals are often unthrifty with a poor hair coat and may show signs of colic. Clinical signs of gastroduodenal ulceration include grinding of the teeth, salivation, gastric reflux, fever, diarrhea, and colic. Often the foal will exhibit signs of abdominal pain after nursing. Gastric outflow obstruction is diagnosed using endoscopy, ultrasonography, and contrast radiography. Endoscopic examination often shows distal esophagitis, gastric ulceration, and duodenal ulceration. Gastric distention with a large volume of reflux is common in cases with an outflow obstruction secondary to duodenal ulceration. Thickening of the duodenum may be detected on transabdominal ultrasonography. Contrast radiography can be used to determine if the stomach is emptying properly (see Figure 42.2).6 The stomach is decompressed and barium is administered followed by a series of lateral radiographs. The contrast material should enter the small intestine within 10 minutes. No contrast material should be in the stomach 35–40 minutes post-administration. Once a diagnosis of gastroduodenal ulceration is made, medical management is started. This treatment includes nasogastric decompression, intravenous anti-ulcer medication, intravenous fluids, parenteral nutrition, intravenous antibiotics, and possibly prokinetic drugs. Surgical intervention is necessary if the foal does not respond to medical management in a short period of time (within 3–5 days).
Figure 42.2 Standing abdominal radiograph (lateral projection) taken 45 minutes following the administration of barium. This is indicative of delayed gastric emptying.
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The basic principle of surgery for treatment of gastric outflow obstruction is to bypass the obstructed portion of small intestine. Prior to surgery a nasogastric tube is positioned. It is very important that the stomach is decompressed prior to performing a bypass procedure. A ventral midline approach to the abdomen is made with the incision extending to the xiphoid cartilage. The abdomen is explored and the site of the gastric outflow obstruction is determined. The most common procedure is to perform a gastrojejunostomy with a jejuojejunostomy. In order to perform this procedure access to the cranial part of the abdomen is crucial. Following completion of the abdominal exploratory the cranial abdomen is ‘packed off’ from the remainder of the abdomen using wet laparotomy sponges. A Balfour retractor is positioned at the cranial aspect of the incision and helps with exposure of the surgical site. A hand-sewn or stapled side-to-side gastrojejunostomy is performed first. Then a hand-sewn or stapled side-to-side jejunojejunostomy is performed to allow outflow of proximal small intestinal contents. Following the jejunojejunostomy procedure the small intestine mesentery is sutured to prevent internal hernia formation. The abdomen is lavaged and closure is as described earlier.
Small intestinal lesions Volvulus of the small intestine is the most common reason for intestinal surgery in the foal (see Figure 42.3).1–3, 6 Most often the ileum and distal jejunum are involved. Small intestinal volvulus is defined as rotation of the intestine around the mesenteric root. It
Figure 42.3 Small intestinal volvulus. The serosal color is abnormal secondary to vascular congestion. Note the size of the small intestine relative to that of the large colon.
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is thought to occur secondary to hyperactive peristalsis adjacent to an area of intestine that is not moving normally. Most often the foal will show signs of acute, severe abdominal pain. Foals with a small intestinal volvulus can have marked abdominal distention. Abdominal ultrasonography will show multiple loops of distended small intestine with minimal to no motility. The intestinal wall may be thickened depending on the duration of the lesion. Early surgical intervention will reduce the likelihood of needing to perform a resection and anastomosis. Once the abdomen is opened the small intestine will be distended with fluid or gas and will often have a gray/ purple color and may have an edema in the intestinal wall due to congestion. During correction of the volvulus the surgeon must be careful not to tear the mesentery. The mesentery is very friable in foals and is prone to tearing during manipulation. If a tear occurs it should be sutured using small (3-0) synthetic absorbable suture material in a simple continuous pattern. Following correction of the volvulus the small intestine should be decompressed and then assessed for viability. A resection and anastomosis will be required if a section of non-viable small intestine is present. Small intestinal intussusception is considered a common occurrence in foals. However, clinical experience and previous retrospective studies would suggest that it is not a common cause of colic in the neonatal foal.2, 3 Occasionally the small intestine can invaginate into itself or into the cecum. The intussusception begins as a simple obstruction but progresses to a strangulating lesion as the tissue becomes edematous and compromises the vascular supply to the intestine. Foals with an intussusception often present with acute colic. They may also present with signs of subacute or chronic colic. Transabdominal ultrasonography can show the classic ‘bull;s eye’ or target lesion (see Figure 42.4). This appearance is caused
by the difference between the edematous inner wall of the intussusceptum and the thinner outer wall of the intussuscipiens. In other cases ultrasonographic examination will only show distended small intestine. During surgery the intussusception is exteriorized and gentle traction applied to reduce the lesion. Once the lesion is reduced the bowel is assessed for viability and resected if necessary. If there is a long length of intestine involved in the intussusception then reduction may not be possible and a resection and anastomosis may need to be performed without reducing the intussusception.
Meconium retention Meconium consists of cellular debris, secretions from the glands, bile, and amniotic fluid. Meconium retention is the most common cause of colic in the neonate.4 Males are more commonly affected and this is thought to be related to their smaller pelvic size. Most cases respond to medical treatment with intravenous fluids, enemas, and laxatives administered via a nasogastric tube. The diagnosis is based on clinical signs, lack of meconium passage, a firm mass felt on digital rectal examination, and abdominal radiographs. Plain film abdominal radiographs show gas distention of the large intestine oral to a large volume of meconium. A barium enema can identify a high meconium impaction or small colon obstruction. Occasionally ultrasonography may be helpful with the diagnosis. Surgical intervention, although rare, may be necessary in cases that fail to respond to aggressive medical therapy. During surgery an enema can be performed in combination with gentle massage to allow passage of the meconium. A small colon enterotomy may be necessary to remove impacted meconium if the enema and massage are not successful. Gentle tissue handling is imperative to prevent adhesion formation.
Inguinal hernia
Figure 42.4 Ultrasonographic image showing the characteristic ‘bull’s eye’ lesion seen with an intussusception.
Inguinal or scrotal hernia can affect any breed but there is a higher prevalence in Standardbreds, Tennessee Walking Horses, and American Saddlebreds.5 The most common type of inguinal hernia is the indirect form. An indirect inguinal hernia has small intestine that passes through the vaginal ring into the vaginal tunic of the scrotum. Typically these hernias are noted shortly after foaling and are easily reduced. Unilateral or bilateral involvement can occur. Typically an indirect inguinal hernia will resolve by the time the foal is 3–6 months of age. Surgical intervention may be necessary if the hernia fails to resolve or
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Surgical Gastrointestinal Conditions if it is getting larger. Surgical options include closure of the external inguinal ring with and without castration, and laparoscopic closure of the internal inguinal ring without castration. A ruptured inguinal hernia occur when the common vaginal tunic ruptures and the small intestine is no longer contained within the tunic and is lying subcutaneously in the inguinal or scrotal region. A ruptured inguinal hernia is rarely reducible and causes signs of colic. Other signs include severe scrotal and preputial swelling/edema, with excoriation and splitting of the skin. Immediate surgical intervention is required for a foal with a direct inguinal hernia. The foal is positioned in dorsal recumbency, and the ventral abdomen and inguinal regions are aseptically prepared for surgery. Two approaches are utilized to resolve this type of hernia. A ventral midline incision is used to gain access to the abdomen and an inguinal incision is made to allow access to and visualization of the incarcerated intestine. The small intestine can be returned to the abdomen by a combination of gentle traction from the ‘abdominal side’ and feeding the intestine through the inguinal ring from the ‘inguinal side’. Once the intestine has been returned to the abdominal cavity it is assessed for viability. Most often a resection and anastomosis is not necessary. A unilateral castration is typically performed. The vaginal tunic and external inguinal ring should be closed. Since there is a large amount of dead space formed secondary to the intestine dissecting subcutaneously, one or two Penrose drains should be placed.
Diaphragmatic hernia A defect in the diaphragm can be congenital or acquired.6 A congenital defect or a tear in the diaphragm secondary to a rib fracture are the most likely causes in foals. Small defects in the diaphragm are more likely to strangulate intestine resulting in signs of acute abdominal pain. Large defects typically cause dyspnea from compression of the lungs by displaced abdominal viscera. Thoracic radiographs and ultrasonography can be used to make a preoperative diagnosis of a diaphragmatic hernia. However, it is not uncommon to make the diagnosis intraoperatively rather than preoperatively. At this time it is imperative to have the foal on positive pressure ventilation. The approach to the abdomen is the same except that the incision will often need to be extended more cranially to increase exposure and facilitate access to the diaphragm. It is easier to gain access to defects located in the ventral aspect of the diaphragm compared to dorsally located defects. Techniques to increase exposure include raising the front of the surgery table to displace the abdominal contents caudally, decompressing the stomach if
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it is large, and extending the cranial aspect of the incision in a paracostal fashion. Depending on the size of the defect and the status of the incarcerated portion of intestine, the hernia may need to be enlarged to free the intestine. Most defects can be repaired with large absorbable suture material using either a simple interrupted or simple continuous pattern. Mesh (absorbable or non-absorbable) can be used to facilitate repair of diaphragmatic defects. Just prior to completing the repair of the diaphragmatic defect, positive pressure ventilation can be used to restore negative pressure within the thoracic cavity. Fluid (blood, peritoneal fluid, and lavage fluid) should be suctioned from the thoracic cavity prior to completing the defect repair as well.
Umbilical hernia Umbilical hernia is a very common congenital defect in horses. Most often the hernia is reducible and repair is on an elective basis if it does not resolve as the foal matures. Hernial contents can include cecum, ileum, jejunum, or ventral colon. A Richter’s or parietal hernia occurs when just the antimesenteric portion of the intestinal wall is within the hernial ring. Incarceration of intestine within the hernial ring is not common. Typically a strangulating umbilical hernia is associated with signs of abdominal pain. The physical examination findings include a nonreducible umbilical hernia that is firm and painful. Ultrasonography of the umbilical hernia will show thickened, edematous intestinal wall. Depending on the duration of the incarceration the intestine oral to the strangulated portion may be distended with fluid or gas. A non-reducible hernia requires immediate surgical intervention. The foal is positioned and prepared as described earlier. The approach to the abdomen should be cranial to the hernia. Once the abdomen is entered the surgeon can use their finger(s) to protect the incarcerated intestine while continuing the ventral midline incision caudally to open the hernia ring. Once the ring has been opened the incarceration will be relieved and the viability of the intestine can be assessed. A resection and anastomosis may be required depending on intestinal viability. The remainder of the abdomen should be explored. Prior to closure the hernial sac and skin from over the hernia should be excised. Closure of the midline incision is as described earlier.
Large intestinal lesions Foals can show signs of colic from the same large intestinal problems that occur in adult horses. Right
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Figure 42.5 The mesodiverticular band is being elevated by the surgeon’s fingers.
and left dorsal displacements of the large colon can occur in young foals. Large colon volvulus can occur as well. However, these three large colon abnormalities are more common in adult horses than in foals. Surgically the problems are corrected via a ventral midline approach.
Congenital abnormalities During early embryonic development the vitelline artery and associated mesentery should atrophy. If this fails to occur, a mesodiverticular band is the result (see Figure 42.5).6 These bands are typically located in the distal jejunum. They are usually from one side of the mesentery to the antimesenteric surface of the small intestine, resulting in a triangular space. A problem occurs when intestine becomes strangulated in a mesenteric rent that forms in the triangular space. A volvulus of the small intestine is
often found in combination with the strangulated intestine. A mesodiverticular band can be an incidental lesion also. If a mesodiverticular band is present it should be removed. This is accomplished by using Metzembaum scissors to excise the band of mesenteric tissue. Ligatures should be applied to obtain hemostasis, if necessary. A Meckel diverticulum is a remnant of the omphalomesenteric duct that provided communication between the yolk sac and the embryonic gut.6 When present it forms a conical extension for the antimesenteric surface of the distal jejunum or ileum. The Meckel diverticulum can cause problems by strangulating intestine, creating an axis for a small intestinal volvulus, and becoming necrotic resulting in fatal peritonitis. Care must be taken when these are resected as they communicate with the lumen of the small intestine.
References 1. Bartmann CP, Freeman DE, Frauke G, von Oppen T, Lorber KJ, Bubeck K, Klug E, Deegen E. Diagnosis and surgical management of colic in the foal: literature review and a retrospective study. Clin Tech Equine Pract 2002;1(3): 125–42. 2. Cable CS, Fubini SL, Erb HN, Hakes JE. Abdominal surgery in foals: a review of 119 cases (1977–1994). Equine Vet J 1997;29(4):257–61. 3. Vatistas NJ, Snyder JR, Wilson WD, Drake C, Hildebrand S. Surgical treatment for colic in the foal (67 cases): 1980–1992. Equine Vet J 1996;28(2):139–45. 4. Bryan JE, Gaughn EM. Abdominal surgery in neonatal foals. Vet Clin Equine 2005;21:511–35. 5. Orsini JA. Abdominal surgery in foals. Vet Clin N Am Equine Pract 1997;13(2):393–413. 6. Mair T, Divers T, Ducharme N. Gastrointestinal disease in the foal. In: Mair T, Divers T, Ducharme N (eds) Manual of Equine Gastroenterology. London: Saunders, 2002; pp. 449–92.
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Chapter 43
Other Gastrointestinal Disorders (Chyloperitoneum, Atresia, Lethal White) K. Gary Magdesian
Chyloabdomen Chyloabdomen, chylous ascites, or chyloperitoneum is a rare cause of colic and abdominal distension in foals. Chyloabdomen is the presence of fluid with a high concentration of lymph and chylomicrons within the peritoneal cavity. Chylous effusions form from leaky lymphatics; causes include trauma to lymph vessels, congenital defects in lymphatic formation, lymphangiectasia, lymphatic obstruction, and idiopathic causes.1–3 In one neonatal foal, chylous effusion was attributed to segmental, midjejunal lymphangiectasia; this was suspected to represent a congenital absence of communication between afferent and efferent lymphatics or a congenital absence of efferent lymphatics from lymph nodes.2 In other reports, chyloabdomen was associated with intestinal lymphatic obstruction by an abdominal abscess in a 5-month-old foal and in association with 180◦ volvulus of the colon in a 20-year-old horse.1, 4 The author has treated one foal with transient chyloabdomen that resolved spontaneously; the cause was presumed to be traumatic in origin, as the foal had been kicked by its dam. The affected foal had clinical signs of mild colic, with effusion noted on ultrasonography. A second foal with chylous effusion had a small intestinal volvulus, with chyle leaking from a segment of strangulated jejunum. The affected jejunum was resected, and the chylous effusion resolved post-operatively. A recent report also describes spontaneous resolution of chylous peritoneal effusion in two neonatal foals with conservative management.5 Both foals had exhibited signs
of abdominal discomfort, ultrasonographic evidence of increased peritoneal effusion and small intestinal thickening, and high peritoneal triglyceride concentrations (4.49 and 16.9 mmol/L [4.0 g/L and 15.0 g/L, respectively], ref: < 1.24 mmol/L[1.1 g/L]). One foal had distended mesenteric lymphatics found upon elective exploratory celiotomy for removal fo umbilical structures a few days later.5 Other causes of chyloabdomen in humans include lymphosarcoma, mesenteric adenitis, tuberculosis, embolic occlusion of the subclavian vein, adhesive peritoneal bands, and cirrhosis. To the author’s knowledge these have not yet been reported in horses. The diagnosis of chyloabdomen is through abdominocentesis. Abdominal fluid is milky white to pinkish white in color; it is opaque, and does not clear with centrifugation. Ether or chloroform will clear the fluid of opacity, and this can be used diagnostically. The fluid is confirmed as chylous with a high triglyceride (TG) and low cholesterol concentration; these can be compared with plasma concentrations, in which the TG concentration is many times higher in the peritoneal fluid than in the plasma. In general, triglyceride concentrations exceeding 1.1 g/L and an abdominal fluid–serum triglyceride ratio greater than 12:1 are confirmatory in dogs and cats. A cholesterol–triglyceride ratio of < 1:1 is consistent with chylous effusion. In one Quarter Horse colt with congenital lymphangiectasia the cholesterol-triglyceride ratio of the fluid was 0.22:1.3 The total nucleated count and leukocyte differential are normal, provided that there is no concurrent sepsis. Free fat globules may be observed cytologically.
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Abdominal ultrasonography reveals ascites and intestinal ileus. Chylous effusions must be differentiated from pseudochylous fluid. Pseudochylous effusions do not contain chylomicrons, clear upon centrifugation, and contain cholesterol and protein–lecithin compounds, giving them a higher cholesterol concentration than serum. Treatment of chyloabdomen depends on the cause. Idiopathic or traumatic, mild chyloabdomen may resolve with time and supportive care. Foals without signs of colic and that are systemically stable warrant provision of time and monitoring of the degree of abdominal effusion through serial ultrasonography. Those with colic and marked abdominal distension may require exploratory celiotomy with resection of affected segments of bowel, or decompression of the obstructed intestine and post-operative dietary management. Lymphatic obstruction or ruptures of lymph vessels from intestinal obstruction require resolution of the primary problem. Congenital lymphatic disorders with long segments of affected intestine precluding surgical resection have a poor prognosis. Foals with moderate to marked chylous effusions may benefit from provision of parenteral nutrition. Diets low in fat and high in protein may be administered concurrently. Medium-chained triglycerides are absorbed directly into the portal vein from the intestine, and these bypass the need for intestinal absorption into lymphatics, thereby leading to decreased chyle production. Fortunately, mare’s milk contains large amounts of medium-chain fatty acids, and therefore no dietary adjustments are needed for affected foals. Chyloabdomen should be considered as a differential diagnosis in foals with signs of abdominal pain and ascites on ultrasonography. Conservative management is successful in idiopathic or traumatic cases that are not associated with severe pain, whereas surgical exploration is indicated in foals with severe colic, marked ascites, or lack of resolution in a few days’ time.
Intestinal atresia and malformations Atresia coli, recti, and ani are rare disorders of horses, but occur often enough that they should be considered differential diagnoses for neonatal colic. Most commonly the atresia occurs along the large colon, small colon, or rectum/anus.6, 7 The entire dorsal colon, segments of dorsal and ventral colon, or portions of the small colon can be absent.6 In some cases, the small colon can be normal, with the atresia involving only segments of large colon.6 The suspected pathogenesis involves an ischemic vascular event to
a segment of developing gastrointestinal tract during gestation, with subsequent necrosis. Another possible explanation includes the result of inflammation in the developing gastrointestinal tract. There are three types of atresia possible. 1. Type 1. Membrane atresia, caused by a complete membrane or diaphragm which occludes the lumen. 2. Type 2. Cord atresia, which involves a small cord of fibrous or muscular tissue joining the blind ends of bowel. 3. Type 3. Blind-end atresia, which is caused by an absence of a segment of intestine; the blind ends are disconnected and there is a concomitant gap in the mesentery. Most of the reports in foals have been type 3 atresia. Foals with atresia coli/ani develop clinical signs of colic and abdominal distension by 2–26 hours of age (mean 8.2 hours).7 The differential diagnosis list for colic in the neonatal foal is long, and includes obstructive and non-obstructive lesions. Some of the more common causes of colic include meconium impaction, ileus, uroperitoneum, intussusception, and enterocolitis. Diagnostic tests that can help in differentiation of congenital atresias from these other causes of colic include digital rectal palpation, response to enemas, barium contrast enemas, and proctoscopy. There is usually no history of meconium passage. Abdominal radiography and ultrasonography are extremely useful in ruling out other causes of colic, but they are usually non-specific as far as diagnosis of atresia. Radiographs usually reveal large colonic and cecal gas distension. Digital palpation of the rectum and enemas reveal absolutely no feces or meconium in foals with atresia; this is an important clue that should make one consider the possibility of atresia. Instead of fecal staining, the palpation and enemas reveal white mucus only. Barium enemas are very useful in the diagnosis of atresia, although they are not completely sensitive in that they can miss proximal atresias by physically not reaching the blind end.7 Proctoscopy is a very specific diagnostic tool for atretic bowel; however, like barium enemas, it can miss lesions that are relatively proximal or orad. In distal or aboral locations, proctoscopy reveals blind ends in the small colon. The only treatment available for atresia coli/ani is surgical.8 However, attempts at surgical correction in foals have met with a high rate of complications and failure, both intra- and post-operatively. Intraoperative difficulties have included mismatched diameters of bowel for anastomosis (e.g., large colon to small colon), as well as large gaps in bowel length.6, 7 Post-operative complications have
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Other Gastrointestinal Disorders (Chyloperitoneum, Atresia, Lethal White) included leakage at the anastomosis site and adhesion formation.7, 9 There has been a suspicion of a hereditary basis of atresia coli in foals; however, this remains to be proven.6 Other congenital defects have been found concurrently in affected foals. Rectourethral and rectovaginal fistulas have been reported in foals with atresia ani/recti.10, 11 These have been successfully repaired surgically.10 Anal vaginalis is the condition in which the rectum opens into the roof of the vagina and the feces exit the vulva.11 These are reparable as long as there is not a significant length of rectum that is atretic. Other concurrent defects have included ventricular septal defect and other cardiac defects, hydrocephalus, cerebellar dysplasia, and dermal hemangioma.7 Short colon syndrome, a congenital colonic malformation, was reported in one 4-month-old Standardbred colt. This was found during an exploratory laparotomy for acute colic, but was not felt to be the primary cause of the colic.12 The entire dorsal colon was missing, and the left ventral colon connected directly to the transverse colon. Post-operatively and long term, the colt had frequent bouts of mild colic, particularly associated with dietary changes or reductions in exercise, and these were felt by the authors to be due to the short colon.12 Agenesis of the mesocolon has also been reported as a cause of colic in a Standardbred foal.13 This foal also underwent exploratory laparotomy, where it was found that there was a total absence of the mesocolon ascendens. Both colonic loops formed a large ring, and the cecum and small intestine had passed through the opening and were twisted around the proximal ventral right colon. The dorsal and ventral colons were anatomically positioned and sutured in place. The foal did well long term.13 Although uncommon to rare, intestinal atresia, short colon, and mesenteric agenesis should be considered in the differential diagnosis list of colic in neonatal foals.
Intestinal aganglionosis Aganglionosis is most commonly associated with the homozygous condition of the frame overo gene. This is part of the lethal white overo syndrome, common in Paint and Pinto horses. The gene is semidominant; that is, in its heterozygous state the phenotype is a frame overo (see Figure 73.1 in Chapter 73, Inherited Diseases of Foals), whereas in its homozygous state it produces the lethal white syndrome. As such, matings of two heterozygotes (frame overos) will produce lethal white foals 25% of the time. The defect represents a mutation in the endothelin B receptor
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gene, which affects migration of neural crest-derived cells.14 Affected foals are white or nearly all white with blue eyes. Lethal white foals lack submucosal and myenteric ganglia in the distal small intestine and large and small colons. The lowest concentration of ganglia is in the small colon. This leads to a small diameter of the intestines in affected regions, with ileus and excessively folded mucosa. The result is a lack of movement of meconium from the large colon into the small colon. Foals appear clinically normal at birth, but develop clinical signs of abdominal pain and distension within a few hours to 24 hours of birth. It should be pointed out that these foals have fully patent, though severely decreased in diameter, gastrointestinal tracts and do not have atresia. Definitive diagnosis of the disease is through DNA testing for the mutation (Veterinary Genetics Laboratory, University of California, Davis). A presumptive diagnosis can be made through a combination of a nearly all or all white foal resulting from a frame overo to overo mating, which never passes meconium. These foals have blue irides and are deaf.15 In addition, digital rectal palpation and enemas produce no fecal staining and only white mucus, as with atresia coli. It should be noted that not all white foals born to two overo parents are affected. This is because of the large number of types of overos – splashed, sabino, and others. These other patterns are not associated with the lethal white gene, and can produce all white foals when crossed with frames or other pinto patterns. Therefore, a colicky white foal should not be assumed to be a lethal white without further diagnostic work-up. Also, lethal white foals can be produced from non-overo horses, especially Quarter Horses and breeds which may carry the frame lethal white overo gene. These individuals may not necessarily express the mutation phenotypically because of the presence of modifying genes.16 Additional diagnostics, other than enemas and digital rectal palpation, include barium contrast enemas, abdominal ultrasonography, and abdominal radiography. Barium contrast enemas show an abnormally small diameter of the small colon. In addition, the author has noted that affected foals do not defecate out the barium contrast normally; it is retained or has a delayed evacuation compared with non-affected foals. Abdominal radiography shows small intestinal and colonic gas distension, and ultrasonography shows ileus and small intestinal distension. There is no therapy for this condition in foals. Attempts at resection of affected segments of bowel have been unsuccessful, largely because of the extent of aganglionosis.
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The disease is completely preventable with avoidance of carrier to carrier matings. These can be avoided through genetic testing of Paint and Pinto horses, and also Quarter Horses or other breeds that may carry the allele. With revocation of the ‘white rule’ by the American Quarter Horse Association, the allele frequency may increase in this breed as frame overos with 100% Quarter Horse heritage (previously regarded as ‘crop outs’ and ineligible for registration) are now eligible for registration. Colonic hypoganglionosis has been reported in one Clydesdale foal, but this is not the same mutation as that of lethal white foal syndrome.17 This foal had chronic abdominal distension and intermittent febrile episodes, with impaction of the right dorsal colon. Intestinal aganglionosis should be suspected as a cause of colic in nearly all white to white foals of frame overo lineage. However, not all white foals have the syndrome, and they should not be assumed to be lethal without further diagnostic work-up. Genetic testing can prevent the syndrome.
References 1. Hanselaer JR, Nyland TG. Chyloabdomen and ultrasonographic detection of an intra-abdominal abscess in a foal. J Am Vet Med Assoc 1983;137:96–8. 2. Campbell-Beggs CL, Johnson PJ, Wilson DA, Miller MA. Chyloabdomen in a neonatal foal. Vet Rec 1995;137:96–8. 3. May KA, Good MJ. Congenital lymphangiectasia and chyloperitoneum in a foal. Eq Vet Educ 2007;19:16–18. 4. Mair TS, Lucke VM. Chyloperitoneum associated with torsion of the large colon in a horse. Vet Rec 1992;131:421.
5. Cesar FB, Johnson CR, Pantaleon LG. Suspected idiopathic intestinal lymphangiectasia in two foals with chylous peritoneal effusion. Eq Vet Educ 2010;22:172–178. 6. Cho DY, Taylor HW. Blind-end atresia coli in two foals. Cornell Vet 1986;76:11–15. 7. Young RL, Linford RL, Olander HJ. Atresia coli in the foal: a review of six cases. Eq Vet J 1992;24:60–2. 8. Schneider JE, Leipold HW, White SL. Repair of congenital atresia of the colon in a foal. J Eq Vet Sci 1981;1: 121–6. 9. Adams R, Koterba AM, Brown MP. Exploratory celiotomy for gastrointestinal disease in neonatal foals: a review of 20 cases. Eq Vet J 1988;20:9–12. 10. Jansson N. Anal atresia in a foal. Comp Cont Educ Pract Vet Equine 2002;24:888–9. 11. Kingston RS, Park RD. Atresia ani with an associated urogenital tract anomaly in foals. Eq Pract 1982;4:32–4. 12. Koenig JB, Rodriguez A, Colquhoun K, Stamfli H. Congenital colonic malformation (“short colon”) in a 4-month-old Standardbred foal. Can Vet J 2007;48:420–3. 13. Steenhaut M, Van Huffel X, Gasthuys F. Agenesis of the mesocolon causing colic in a foal. Vet Rec 1991;129:54–5. 14. Metallinos DL, Bowling AT, Rine J. A missense mutation in the endothelin-B receptor gene is associated with Lethal White Foal Syndrome: an equine version of Hirschsprung disease. Mamm Genome 1998;9:426–31. 15. Magdesian KG, Williams DC, Aleman M, Lecouteur RA, Madigan JE. Evaluation of deafness in American Paint Horses by phenotype, brainstem auditory-evoked responses, and endothelin receptor B genotype. J Am Vet Med Assoc 2009;235:1204–11. 16. Lightbody T. Foal with overo lethal white syndrome born to a registered quarter horse mare. Can Vet J 2002;43: 715–17. 17. Murray MJ, Parker GA, White NA. Megacolon with myenteric hypoganglionosis in a foal. J Am Vet Med Assoc 1988;192:917–19.
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Chapter 44
Hepatic Disease Thomas J. Divers and T. Douglas Byars
Hepatic disease in foals might result from infectious, parasitic, congenital, metabolic, or toxic causes. Fortunately, most cases of liver disease in foals do not progress to hepatic failure.
Clinical signs of hepatic disease The predominant signs in foals with hepatic disease are usually those related to another disease process and organ disturbance since most sick foals with hepatic disease do not have hepatic failure. If hepatic failure occurs, severe jaundice, signs of central nervous system disturbance (hepatoencephalopathy; HE), brown urine, and dermatitis may be seen. Aimless wandering, blindness with dilated but responsive to light pupils, and seizures are common signs of hepatoencephalopathy in foals (Figure 44.1). Abdominal pain (colic) may also be a clinical sign.
Diagnosis of hepatic disease and failure in foals Hepatic disease is detected by measuring liver specific enzymes such as sorbitol dehydrogenase (SDH), glutamate dehydrogenase (GDH), and gammaglutamyltransferase (GGT). SDH and GDH are both derived from hepatocytes and are frequently increased with even mild hepatocellular injury. They both have short half-lives and have similar, but not identical, sensitivity for hepatic disease. Some foals with toxic hepatic failure have had normal or only modestly elevated SDH. Gammaglutamyltransferase levels may be high in a few normal neonatal foals.1 The best laboratory aids in confirming hepatic failure are abnormally high concentrations of serum bilirubin (both conjugated and unconjugated), ammonia, and prolonged prothrombin time (Table 44.1). Plasma glucose concentrations are frequently low in neonatal hepatic failure and are commonly < 30 mg/dL (1.67 mmol). Bile acids may
be increased in foals with severe liver disease, although it must be recognized that even some healthy foals have a serum bile acid concentration well above the adult horse normal range until they are 6 weeks of age.2 Foals with portosystemic shunts will generally have normal liver enzymes and normal bilirubin concentration, but bile acids and ammonia will be increased. Ultrasonographic examination of the liver is helpful in determining fibrosis and abscessation of liver parenchyma or umbilical vein (Figure 44.2). Intrahepatic portosystemic shunts can be identified by ultrasonography in some cases. Liver biopsy can be performed on the right side of the abdomen after sonographic visualization. The liver encompasses a relatively larger area in the foal than in the adult horse and locating a site for biopsy is easy in foals. Foals can be administered a low dose of xylazine just prior to biopsy if needed. Although diazepam and midazolam are considered by some to be inappropriate for sedation in HE, they could be used to facilitate the liver biopsy. Drugs metabolized by the liver have a prolonged half-life.
Causes of liver disease and failure in neonatal foals Infectious causes
Tyzzer’s disease Tyzzer’s disease is the best documented cause of bacterial hepatitis in foals. Tyzzer’s disease affects foals 6–45 days of age, often causing rapid onset of septic shock and death. The causative organism is Clostridium piliforme, which is a soil/manure-borne Gramnegative bacteria. The disease can be reproduced in foals by feeding feces from experimentally infected adult horses. Although the disease is not considered contagious and usually is an isolated case, it may continue to occur sporadically on some farms. Foals
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 44.1 A 6-week-old Belgium foal with hyperammonemia due to portosystemic shunt. The foal had variable clinical signs, including seizure and head pressing (seen here).
Figure 44.2 A sonogram, using a 5-MHz sector scanner, of a highly echogenic liver in a 4-week-old foal with hepatic fibrosis which followed neonatal isoerythrolysis.
of non-resident mares appear to be at higher risk than foals of resident mares.3 A tentative diagnosis is based upon age of the foal, clinical signs, and laboratory findings. Affected foals are often found dead or in a coma with severe tachycardia and profound hypoglycemia, metabolic acidosis, and icterus (Figure 44.3). Ultrasonography of the abdomen usually reveals extreme hepatomegaly. Hepatocellular enzymes (AST, SDH) are markedly increased in the serum, as is total and direct bilirubin. The complete blood count (CBC) frequently shows a severe leukopenia with a left shift and thrombocytopenia. Myocarditis may also occur and cardiac troponin I would be expected to be markedly elevated in those cases. One draft mare with twin foals had one foal die with confirmed Tyzzer’s disease and the other developed liver disease within a couple of days, but was treated successfully with antibiotics (B.C. Tennant, personal communica-
tions). Another highly suspect foal was successfully treated with total parenteral nutrition along with trimethoprim–sulfamethoxazole and penicillin.4 More recently, a confirmed case was successfully
Table 44.1 Laboratory testing for hepatic disease in foals: expected approximate values in normal foalsa Biochemistry parameters
Normal range
Sorbitol dehydrogenase (SDH) Glutamate dehydrogenase (GLDH) Gamma-glutamyltransferase (GGT)b Aspartate aminotransferase (AST) Total bile acidsc Ammonia Iron saturationd Conjugated bilirubin Prothrombin time
< 18 IU/L < 22 IU/L < 42 IU/L < 380 IU/L < 30 µmol/L < 60 µg/dL < 35 µmol/L 40–60% < 0.5 mg/dL < 8.55 µmol/L < 11 seconds
a
Normal values could vary considerably between methods and testing laboratories. b May be higher for 3–4 weeks in a small percentage of normal foals. c Concentration is highly variable in normal foals, but when ‘high’ a gradual reduction towards normal adult values (< 14 µmol/L (non-fasted)) should occur over weeks to 3 months. d May approach 100% saturation for first days of life.
Figure 44.3 A 4-week-old foal with severe depression caused by Tyzzer’s disease. The foal also had tachycardia, in addition to severe icterus, and petechiation of mucous membranes. Laboratory results revealed degenerative left shift of neutrophils, thrombocytopenia, elevated liver enzymes, increased bilirubin concentration, and extremely low glucose and bicarbonate.
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Hepatic Disease treated with supportive care and ampicillin and gentamicin.5 Confirmation of the diagnosis is made at either necropsy or, less commonly, by microscopic examination of a liver biopsy.
Actinobacillus equuli and Streptococcus equi zooepidemicus bacteremia These two bacteria are common causes of acute hepatitis in young foals as part of the neonatal sepsis syndrome. Actinobacillus bacteremia and even acute death is common in 3-day-old foals. Some foals may be treated successfully with antibiotics and supportive care for septic shock. Others may be found dead or die quickly and have hundreds of multifocal microabscesses in the liver and/or other organs (kidney for Actinobacillus). Sometimes the gross appearance of the liver lesion might be difficult to distinguish from equine herpesvirus 1 (EHV-1) hepatitis.
Equine herpesvirus 1 hepatitis EHV-1 infection of a near-term fetus may result in the birth of a non-viable foal with hepatic, respiratory, and/or gastrointestinal disease.6, 7 There is generally a rapid and fatal deterioration in the clinical condition within 1–5 days after birth. Neonatal herpesvirus 1 infections are believed to be in utero infections due to either reactivation of the virus in latently infected mares or exposure to a field strain in late pregnancy. Large numbers of mares may become exposed if they are allowed contact with an EHV-1 infected and aborted fetus or the placenta and placental fluids from the aborting mare. It is well known that many mares are latently infected with EHV-1 but reasons for recrudescence in mares are poorly understood. Affected foals are severely icteric and may have severe neutropenia and lymphopenia. Many affected foals are premature and a product of precipitous delivery. Icterus is a common finding in affected foals, although icterus may be seen with other causes of sepsis in neonatal foals. Likewise, liver enzymes are high, but may not be higher than other septic foals. EHV-1-infected foals are almost always severely neutropenic and lymphopenic. Necropsy examination of an affected foal’s liver may look similar to either Actinobacillus or Streptococcus bacteremia, but severe interstitial pneumonia is characteristic of EHV-1 in neonatal foals. Diagnosis can be confirmed by virus isolation, polymerase chain reaction (PCR), and/or immunohistochemistry detection of the virus or virus proteins in the lung or liver. If the placenta is still available it can also be tested. The placenta and mare’s uterine fluid should be considered contagious. The mare may also be contagious via
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her uterine fluids or nasal secretions for several days.
Leptospirosis Leptospirosis has been reported to cause jaundice and death in a 10-day-old foal.8 Although Leptospira pomona is known to cause abortion and liver disease (giant cell hepatopathy) in the equine fetus, it is apparently a rare cause of neonatal liver disease.
Rhodococcus equi Rhodococcus equi may occasionally cause liver abscesssation in growing foals. Affected foals often also have pulmonary disease and synovitis. Some older foals have both mesenteric and hepatic abscessation due to either Streptococcus equi spp. or R. equi infection. Those foals have marked weight loss. Regardless of the site of R. equi infection, infected foals usually have marked neutrophilia and increased fibrinogen. One foal developed an R. equi abscess in the portal vein causing elevation in plasma bile acids. Long-term antibiotic treatment allowed visual (by ultrasonography) resolution of the lesion.
Systemic inflammatory diseases and hepatic disease Bacteremia and/or systemic effects of bacterial toxins in foals may initiate an exaggerated response to sepsis (systemic inflammatory response syndrome with multiple organ dysfunction), which may commonly result in multiple organ dysfunction, including hepatic disease but rarely failure. A large number of vasoactive mediators are involved in this process; the hemodynamic system becomes ineffective and certain organs, such as the liver, may fail because of hypoxia. Diffuse hepatic necrosis and hepatocellular apoptosis are the characteristic lesions. The clinical signs may be identical to Tyzzer’s disease, except the syndrome may affect a much wider age range. Diagnosis is based upon clinical findings, e.g., fever, tachycardia, and trembling, and laboratory findings indicating sepsis, e.g., leukopenia or leukocytosis with left shift and vacuolation of neutrophils. If the liver is affected, hepatocellular-specific enzymes, e.g., SDH, will be increased in the plasma. Prompt and aggressive treatment is often successful. A less common cause of liver disease associated with enteritis and systemic inflammation is acute portal vein thrombosis. Plasma bile acids and blood ammonia would be increased and diarrhea may be persistent due to portal hypertension. Loss of anticoagulant proteins, e.g., anti-thrombin III, from the inflammatory bowel disease may predispose to the
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thrombosis. It is common for umbilical vein infections to extend into the liver, but these rarely cause biochemical evidence of liver disease.
Treatment of infectious causes Treatment is mostly appropriate antimicrobial therapy and supportive care. Since most cases, except for Tyzzer’s disease, cause hepatic disease without failure, specific treatment for liver failure is not often required. Appropriate antibiotic treatments for bacterial hepatitis would be penicillin (Na or K penicillin 22 000 IU/kg q. 6 hours intravenously (i.v.) or procaine penicillin at 15 000 IU/kg intramuscularly (i.m.) q. 12 hours) or ampicillin (Na ampicillin 15 mg/kg q. 8 hours i.v. or ampicillin trihydrate 11 mg/kg q. 12 hours i.m.), combined with gentamicin (7.5 mg/kg as a loading dose followed by 6.6 mg/kg i.v. or i.m. q. 24 hours) or amikacin (25 mg/kg i.v. or i.m. as a loading dose followed by 18–20 mg/kg q. 24 hours), thereafter, for most of the bacterial causes except for R. equi, in which clarithromycin (7.5 mg/kg q. 12 hours per os (p.o.) or azithromycin (10 mg/kg q. 24 hours p.o. for 5 days and then q. 48 hours thereafter) combined with rifampicin (5–10 mg/kg q. 12 hours p.o.) is the treatment of choice. Treatment with valacyclovir (30 mg/kg q. 8 hours) may be attempted treatment for foals with neonatal herpes infection. Foals with hepatic failure due to sepsis, most commonly foals with Tyzzer’s disease, should be treated not only with specific anti-infection medication and supportive care, but should be administered therapy to combat the metabolic consequences of hepatic failure; oral metronidazole (10 mg/kg p.o. 12 hours) for 2 days, neomycin 10 mg/kg p.o. q. 12 hours for 2 days, and probiotics/prebiotics to decrease enteric ammonia production. Sodium bicarbonate can be given in an attempt to correct the often marked metabolic acidosis but only if the venous blood ph is < 7.1 since bicarbonate administration can increase the blood ammonia. Phenobarbitol can be administered (3–10 mg/kg slow i.v. infusion) if needed to control seizures. Fluids (crystalloids mixed with glucose and plasma), oxygen, judicious but early use of non-steroidal anti-inflammatory drugs (NSAIDs), and other anti-oxidants/anti-inflammatory drugs such as low-molecular-weight heparin (50 U/kg subcutaneously (s.c.) q. 24 hours), dimethylsulfoxide (1 g/kg i.v. diluted q. 12–24 hours), acetylcysteine 50 mg/kg i.v diluted for a single treatment, and/or pentoxifylline (8.5 mg/kg p.o. or i.v. q. 12 hours on day 1 followed by 5 mg/kg thereafter) are often used to help decrease inflammation and maintain organ perfusion. Foals with umbilical vein abscessation into the liver are most commonly treated by surgical removal, marsupialization, and antibiotics.
Toxic causes
Iron fumarate Iron fumarate is the best documented of the toxic causes.9, 10 Iron toxicity most often occurs when newborn foals are given iron before nursing. Colostralacquired glutathione or other protective substances in colostrum may explain the great decrease in iron hepatotoxicity when iron-containing products are administered after colostrum.9 When the iron is given at birth and before colostrum, clinical signs develop 2–5 days later. In rare cases, clinical signs may not develop until the foal is older. The initial clinical signs are associated with hepatoencephalopathy and include seizures, marked depression, ataxia, aimless wandering, head pressing, or any sign of abnormal behavior. Icterus is noted in most foals at the time neurologic signs are exhibited, although some foals may die so peracutely that icterus is not noticed.
Other toxic causes Although not documented in foals, NSAIDs, such as carprofen, are other potential toxic causes of hepatic disease. Rarely, foals treated with macrolides, trimethoprim–sulfamethoxazole, and/or H2 blockers for pneumonia, gastroduodenal ulcers, or diarrhea develop an increase in serum hepatic enzymes in spite of the foal improving clinically. On occasion, AST concentrations slowly increase to nearly 800 IU/L and SDH to 150 IU/L and GGT to 120 IU/L or more in those cases. Function tests remain normal, so there is presumed disease without failure. With changes in drug treatment, the serum enzymes return to normal, suggesting these cases may have drug-induced hepatopathy. Steroid-induced hepatic lipidosis has been observed in foals receiving both prolonged and high doses of corticosteroids, mostly for neurological diseases. Rarely, growing foals on pasture develop photosensitization and marked elevations in GGT, suggestive of a mycotoxin-induced hepatic disease, although the specific mycotoxin is rarely, if ever, discovered. Theiler’s disease (serum hepatitis) is either rare or does not occur in foals.
Liver disease/failure in foals following neonatal isoerythrolysis This is an observational disease of unknown etiology which has been seen in several foals that had neonatal isoerythrolysis and transfusion therapy (usually, but not always, more than one transfusion).11 Potential causes of the liver failure are chronic hypoxia, iron toxicity associated with multiple transfusions, immune reactions, or, more likely, a cholangiopathy
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associated with the hemolysis, bile stasis, and toxic injury to bile epithelium. Typically, affected foals have appeared to recover from the neonatal isoerythrolysis, but are noted to be lethargic and not growing normally. A chemistry panel reveals increased liver enzymes, with GGT the most abnormal, and increases in direct and indirect bilirubin and bile acids. Ultrasonographic examination of the liver often shows increased echogenicity and biopsy is reported as a hepatopathy with both regeneration and fibrosis.
Treatment of toxic hepatopathy Treatment is mostly supportive to maintain hydration and prevent further catabolism. If fulminant hepatic failure is present or seems eminent, therapy to reduce enteric ammonia production as listed under treatment of infectious causes of liver failure should be implemented. Drugs that may decrease inflammation, oxidative injury to the liver, and promote bile flow are recommended. These include pentoxifylline, 8.5 mg/kg q. 12 hours p.o., S-adenosylmethionine (SAMe) 0.5 g p.o. q. 24 hours, and supplemental vitamin E and selenium.12 Hepatic lipidosis Hepatic lipidosis from a severe hyperlipidemia frequently referred to as hyperlipemia may occur in neonatal foals, especially miniature equine foals.13, 14 The foals are usually ill with another disease which causes anorexia and marked lipolysis with visible lipidemia in the plasma leading to a rapid development of hepatic lipidosis. Plasma concentrations of hepatic enzymes are highly variable, but plasma triglyceride will be > 500 mg/dL (5.65 mmol). The clinical signs include severe depression, seizure, blindness, and ventral edema. Acute death may occur due to hepatic rupture. Treatment should be directed towards any predisposing disease and nutritional support provided to combat the hyperlipidemia/hepatic lipidosis. Parenteral nutrition with amino acids, especially L-tryptophan,15 B-vitamins, glucose, and insulin (0.1–1.0 IU regular insulin either i.m. q. 6–12 hours or i.v. infusion over time at 0.01–0.1 IU/h, depending upon glucose concentrations) can be life-saving in some cases. Enteral nutrition, with low-fat milk replacer mixed with yogurt, should also be provided if the intestinal tract is normally functioning. Crystalloids given intravenously would also be needed to maintain hydration and organ perfusion. Heparin treatment has been recommended to decrease lipemia although clinical evidence of efficiacy is lacking.
Figure 44.4 A 5-week-old foal with ptyalism and odontoporesis and icterus caused by ‘scarred’ obstruction of the duodenum and interference with the bile flow.
Obstructive causes Obstructive causes include cholangitis associated with duodenal ulcer disease, Parascaris equorum migration, and congenital anomalies such as atresia of the bile ducts. The most common of these is obstruction due to duodenal ulceration, most commonly seen in foals recovering from diarrheal diseases (Figure 44.4). Colic, bruxism, and ptyalism are usually seen with duodenal obstruction, and the stomach is markedly distended upon ultrasonographic examination. Two distinct forms of the obstruction can be seen depending upon location of the healing duodenal ulcer. If the healing (scaring ulcer) is directly over the two openings (biliary papillae) of the common bile duct, marked jaundice will occur and contrast radiographs (oral barium) will not show any barium in the bile ducts. If the obstruction is distal to the common bile duct opening, icterus is not as marked and barium may be seen entering the bile ducts.16, 17 The former is the most difficult to treat, as surgical transposition of the bile duct opening (hepaticojejunostomy) may be required, in addition to a gastroduodenojejunostomy.18 Medical therapy for duodenal ulcers and supportive care may allow some foals to improve without surgery.
Congenital or inherited causes Hyperammonemia and hepatopathy in Morgan weanlings Acute onset of signs (coma, blindness, seizure) of hyperammonemia has been seen in Morgan foals
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generally 4–7 months of age.19, 20 The onset of clinical signs often occur soon after weaning. Liver enzymes are mildly elevated, and variable degrees of portal and bridging fibrosis with bile duct hyperplasia, karyomegaly, and cytomegaly are often seen on microscopic examination, but there is not nearly enough microscopic disease to explain the hepatic failure, nor are other liver function tests abnormal in most of the foals. The cause of the disease is unknown, but appears to be mostly caused by abnormal metabolism of ammonia and possibly other amino acids. The blood ammonia is extremely high (300–600 µmol/L) in most foals, indicating they had likely developed some compensatory/somewhat protective mechanism against the persistent elevation in ammonia in order to live to be several months of age. The author has pedigree information on 12 affected foals and, interestingly, there is a common grandsire on the paternal side in 11 of the foals and, in the other foal, the stallion shows up as a grandsire on the maternal side. The disease is fatal and may end with a terminal hemolytic crisis. Some foals may survive the first crisis and live for days or even weeks before dying with a second HE-like crisis. Congenital portosystemic shunts Congenital portosystemic shunts occur infrequently in the equine. Clinical signs may not be noted until foals are 2–3 months of age and begin ingesting large amounts of grain or grass. Waxing and waning signs of encephalopathy are the most common signs in foals. An elevation in plasma bile acids and ammonia in foals with encephalopathic signs and normal concentration of hepatic-derived serum enzymes should arouse suspicion of a portosystemic shunt. Shunts may be single or multiple and may be intrahepatic or extrahepatic. Positive contrast portography is the diagnostic technique of choice. The shunt may also be detected by transrectal portoscintigraphy (in foals) or transabdominal ultrasonographic examination. Successful medical management followed by shunt ligation has been described in a foal (Figure 44.5).21 Other causes of hyperammonemia without liver disease in foals include meconium impactions and uremic encephalopathy. Other causes of icterus in foals Icterus in the neonatal ill foal is common. It may be a result of liver disease, as discussed above, or hemolysis, most commonly neonatal isoerythrolysis, or secondary to sepsis, dysmaturity, and/or neonatal encephalopathy. Foals can be surprisingly very icteric with any of the last three disorders and not have marked increases in liver enzymes, although both
Figure 44.5 Belgium foal seen in Figure 44.1 after surgical repair of a portosystemic shunt.
indirect and direct bilirubin may be increased. The cause for the increase in the direct bilirubin, sometimes as high as 3 mg/dL, with minimal or no biochemical evidence of liver disease, is not proven but could be due to either biliary stasis or, more likely, intestinal ileus and increased resorption of the conjugated (direct) bilirubin from the intestinal tract. As mentioned above, some foals with these disorders will have biochemical evidence of severe liver disease.
References 1. Patterson WH, Brown CM. Increase in serum gammaglutamyltransferase in neonatal standardbred foals. Am J Vet Res 1986;47:2461–3. 2. Barton MH, LeRoy BE. Serum bile acids concentrations in healthy and clinically ill neonatal foals. J Vet Intern Med 2007;21:508–13. 3. Fosgate GT, Hird DW, Read DH, Walker RL. Risk factors for Clostridium piliforme infection in foals. J Am Vet Med Assoc 2002;220:785–90. 4. Peek SF, Byars TD, Rueve E. Neonatal hepatic failure in a Thoroughbred foal: successful treatment of a presumptive Tyzzer’s disease. Equine Vet Educ 1994;6:307–9. 5. Borchers A, Magdesian KG, Halland S, Pusterla N, Wilson WD. Successful treatment and polymerase chain reaction (PCR) confirmation of Tyzzer’s disease in a foal and clinical and pathologic characteristics of 6 additional foals (1986–2005). J Vet Intern Med 2006;20:1212–18. 6. Hartley WJ, Dixon RJ. An outbreak of foal perinatal mortality due to equid herpesvirus type I: pathological observations. Equine Vet J 1979;11:215–18. 7. Bryans JT, Sweezle TW, Dorlington RW, Crowe MW. Neonatal foal disease associated with perinatal infection by equine herpesvirus I. Equine Med Surg 1977;1:20. 8. Hodgin EC, Miller DA, Lozano F.. Leptospira abortion in horses. J Vet Diagn Invest 1989;1:283–7. 9. Divers TJ, Warner A, Vaala WE, Whitlock RH, Acland HA, Mansmann RA, Palmer JE. Toxic hepatic failure in newborn foals. J Am Vet Med Assoc 1983;183:1407–13.
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Hepatic Disease 10. Mullaney TP, Brown CM. Iron toxicity in neonatal foals. Equine Vet J 1988;20:119–24. 11. Boyle AG, Magdesian KG, Ruby RE. Neonatal isoerythrolysis in horse foals and a mule foal: 18 cases (1988–2003). J Am Vet Med Assoc 2005;227:1276–83. 12. Medina J, Moreno-Otero R. Pathophysiological basis for antioxidant therapy in chronic liver disease. Drugs 2005;65:2445–61. 13. Tan RH, Hughes KJ, Hodgson DR. Hyperlipaemia, neonatal isoerythrolysis and hepatocellular necrosis in a 3-dayold Thoroughbred foal. Aust Vet J 2005;83:740–1. 14. Mogg TD, Palmer JE. Hyperlipidemia, hyperlipemia, and hepatic lipidosis in American Miniature Horses: 23 cases (1990–1994). J Am Vet Med Assoc 1995;207:604–7. 15. Malaguarnera M, Pistone G, Elvira R, Leotta C, Scarpello L, Liborio R. Effects of L-carnitine in patients with hepatic encephalopathy. World J Gastroenterol 2005;11:7197– 202.
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16. Buote M. Cholangiohepatitis and pancreatitis secondary to severe gastroduodenal ulceration in a foal. Can Vet J 2003;44:746–8. 17. Orsini JA, Donawick WJ. Hepaticojejunostomy for the treatment of common hepatic duct obstructions associated with duodenal stenosis in two foals. Vet Surg 1989;18:34–8. 18. Zedler ST, Embertson RM, Bernard WV, Barr BS, Boston RC. Surgical treatment of gastric outflow obstruction in 40 foals. Vet Surg 2009;38:623–30. 19. Divers TJ, Tennant BC, Murray MJ. Unusual cases of liver disease in Morgan foals. Gastroenterol Viewpoint 1994;2:6. 20. McConnico R, Duckett WM, Wood PA. Persistent hyperammonemia in two related Morgan weanlings. J Vet Intern Med 1997;11:264–6. 21. Fortier LA, Fubini SL, Flanders JA, Divers TJ. The diagnosis and surgical correction of congenital portosystemic vascular anomalies in two calves and two foals. Vet Surg 1996;25:154–60.
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45.
Anemia Jane E. Axon
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Chapter 45
Anemia Jane E. Axon
Anemia is a common clinicopathological finding in the equine neonate. The significance of the changes in the hematological values should be interpreted with reference to the normal changes that occur during the neonatal period.1 The packed cell volume (PCV) and hemoglobin concentration (Hb) are similar to those of adults at birth and then decrease over the next 2 weeks to remain in the range of low normal for an adult for the first few months of life (Table 45.1).2 The mean corpuscular volume (MCV) also decreases during the first 4 months of life, resulting in a degree of microcytosis and mild anisocytosis. Foals in the first 3 months of life have a relative iron deficiency and have a microcytosis, decreased serum ferritin concentration, and increased serum total iron-binding capacity (TIBC) (Table 45.1).3
Causes of anemia in the foal Anemia can be due to internal or external blood loss, hemolysis, or decreased erythrocyte production (Table 45.2).
Blood loss Acute hemorrhage in the foal most commonly occurs with blood loss from the umbilical remnant or laceration of major vessels by fractured ribs. Blood loss from the umbilical remnant can occur externally or internally into the internal remnant, bladder, or peritoneal cavity. Premature severance of the cord after birth results in no significant effect on the foal’s RBC parameters.4 Traumatic injury to vessels with external trauma, gastrocnemius muscle and tendon rupture, and femoral artery hemorrhage can also occur. Hemostatic disorders may result in blood loss via spontaneous hemorrhage into joints, subcutaneous bruising, and gastrointestinal hemorrhage. Heritable bleeding disorders such as von Willebrand’s disease, classic hemophilia A (factor VIII:C deficiency), and
other clotting factor deficiencies have been reported in foals.5–8 Disseminated intravascular coagulation (DIC) is the most common acquired hemostatic problem in the foal. DIC occurs secondarily to other disease processes such as sepsis and systemic inflammatory response syndrome (SIRS). Alloimmune thrombocytopenia has been reported in horses and mules and may occur in combination with neonatal isoerythrolysis.9, 10 Thrombocytopenia has been recently described with ulcerative dermatitis and mild neutropenia in foals.11 The exact cause is unknown but is suspected to be due to factors transferred in colostrum. Reduced erythrocyte lifespan Neonatal isoerythrolysis is the most common cause of hemolytic anemia in the foal. Other potential causes are rapid or concentrated administration of dimethyl sulfoxide (DMSO), immune-mediated disease, intracellular RBC parasites, and mechanical or toxin damage to the erythrocyte. Neonatal isoerythrolysis (NI) is an alloimmune hemolytic anemia of newborn foals caused by antibodies in mare’s colostrum directed against the foal’s erythrocytes (see Chapter 35). For NI to occur, the mare must be negative for the RBC factor and then be exposed to (from a previous or current pregnancy or blood transfusion), and develop antibodies against, the RBC factor. The foal from the current pregnancy inherits the RBC factor from the stallion. The foal then has to nurse and absorb the antibodies in the mare’s colostrum against the RBC factor. Although the potential is present for a higher incidence of NI in foals, the incidence of NI is 2% in Standardbreds and 1% in Thoroughbreds as only a small percentage of mares produce significant alloantibodies against the specific antigenic factors on the foals’ RBCs.12 The most common RBC factors involved in NI are Qa and Aa. Mares without a Qa or Aa factor are thus
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 45.1 Red blood cell indices, serum iron, ferritin, and total iron-binding capacity in foals2, 3 Age
RBC PCV Hb MCV MCHC Serum iron Serum ferritin TIBC
× 1012 /L L/L g/dL fL g/dL mg/L µg/L mg/L
< 1 h; before nursing
< 12 h; after nursing
24 h
7 days
1 month
3 months
6 months
9.3–12.9 0.40–0.52 13.4–19.9 37–45 33–39 3.45–5.92 34–161 3.86–6.63
9.0–12.0 0.37–0.49 12.6–17.4 36–45 32–40 2.62–4.88 – 3.39–5.35
8.2–11.0 0.32–0.46 12.0–16.6 36–46 32–40 0.78–3.48 79–263 2.08–6.20
7.4–10.6 0.28–0.43 10.7–15.8 35–44 35–40 0.30–2.73 57–173 2.22–6.19
7.9–11.1 0.29–0.41 10.9–15.3 33–40 34–40 0.49–2.88 33–140 4.37–7.77
9.2–12.0 0.32–0.42 11.7–15.3 31–38 34–40 0.61–3.06 33–223 4.10–5.96
7.9–11.6 0.29–0.41 10.8–15.4 32–39 33–40 0.85–2.64 81–331 3.41–6.35
Reference ranges determined by mean ± SD. Hb, hemoglobin; MCHC, mean corpuscular hemoglobin concentration; MCV, mean corpuscular volume; PCV, packed cell volume; RBC, red blood cell; WBC, white blood cell.
at a higher risk of producing antibodies which result in NI. An alloantibody thrombocytopenia and neutropenia may also be present. Mule foals have a higher incidence of NI owing to the RBC antigen (donkey factor) that is found on all donkey cells. Mule foals are produced from the mating of a donkey sire with a horse dam. Horse RBCs do not contain a donkey factor; thus, all mule foals are theoretically at risk of developing NI. Although many mule foals remain clinically normal after birth, the reported incidence of NI is about 10%.13 Decreased erythropoiesis Decreased erythrocyte production is most commonly seen in the foal with anemia of chronic disease. Chronic inflammatory disease causes a characteristic disturbance in iron metabolism and depressed bone marrow response; however, other factors such as inhibition of erythropoiesis by inflammatory mediators and reduced effectiveness and secretion of erythropoietin may also play a part.14 The serum iron concentration is low despite adequate total body iron stores. The total iron-binding capacity (TIBC) is normal or decreased and serum ferritin levels are increased.15 Absolute iron deficiency, which is a combination of hypoferritinemia and hypoferremia, has been described.16–18 The anemia is usually normocytic and normochromic and becomes microcytic and hypochromic in more severe cases. Bone marrow abnormalities involving erythroid precursors have been described in foals. Normal foals within 4 days of parturition have a higher myeloid–erythroid (M/E) ratio than adult horses (0.5–3.76).19, 20 Erythroid hypoplasia has been reported as part of a syndrome in Fell pony foals.21
Involvement of other cell line precursors has resulted in panleukopenia and thrombocytopenia.22 Anemia from bone marrow aplasia and hypoplasia was reported in newborn foals with concurrent congenital skin, renal, and lymphoid defects. It is suspected that administration of sulfonamides, pyramethamine with or without trimethoprim, and folic acid to pregnant mares resulted in the congenital defects.23
Clinical signs Clinical signs of anemia reflect the decreased oxygencarrying capacity of blood and include exercise intolerance, lethargy, weakness, tachycardia, tachypnea, and pale mucous membranes (Figure 45.1). The signs exhibited depend on the rate of development and severity of anemia. Acute blood loss can result in hypovolemia with poor peripheral perfusion, and shock can develop if more than one-third of the blood volume is lost acutely.16 Hemorrhage into the thoracic cavity associated with vessel laceration from a fractured rib may result in dyspnea and respiratory distress. Crepitus on rib palpation, a ‘clicking’ sound on auscultation, or edema over or ventral to the fractured rib may be seen; however, palpation of crepitus over the fracture site is not a consistent finding and rib fractures are often not detected on physical examination. The hemothorax will cause ventral dullness and lack of breath sounds on auscultation of the thorax. Sudden death may also occur when a major vessel, coronary vessel, or myocardium are lacerated. Blood loss from the umbilical remnant can result in obvious hemorrhage from the stump or formation of a hematoma around or within the external cord. Internal umbilical remnant hemorrhage can result in signs of abdominal
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Table 45.2 Causes of anemia in the foal Blood loss
Increased RBC destruction
Decreased erythropoiesis
External hemorrhage Cesarean section Bleeding umbilical remnant Trauma Gastrointestinal tract Bleeding gastric ulcers Hemorrhagic enterocolitis Urinary tract Vascular anomalies Renal hemorrhage Internal hemorrhage Ruptured femoral artery Ruptured gastrocnemius muscle/tendon Ruptured liver/spleen Internal umbilical remnant Fractured ribs Hemothorax Hemostatic disorders Thrombocytopenia Alloimmune mediated Immune mediated Disseminated intravascular coagulation Congenital factor deficiency Factor VIII:C von Willebrand’s disease Neonatal isoerythrolysis Infectious causes Clostridium spp. Piroplasmosis Equine infectious anemia Disseminated intravascular coagulation Incompatible blood transfusions Rapid administration of hypotonic fluids Immune mediated Drugs (penicillin) Rhodococcus equi Toxic causes Snake envenomation Rapid or concentrated DMSO administration Prematurity Anemia of chronic disease Iron deficiency Bone marrow defects associated with maternal administration of sulfonamides, pyramethamine, folic acid Chronic renal disease Fell pony syndrome
discomfort from a hemoabdomen, and stranguria from hematoma formation within the bladder. Hematomas within the internal remnant can be palpated through the abdominal wall or visualized on ultrasonographic examination. Hemostatic disorders may result in melena, hematuria, swollen joints associated with hemarthroses, subcutaneous swellings from hematomas, or prolonged bleeding from venipuncture sites. Foals with DIC will have numerous other clinical signs of sys-
temic disease and sepsis. Petechiae are seen with DIC and alloimmune thrombocytopenia. Icterus is characteristically present if hemolysis has occurred, but may not be apparent if the hemolysis is peracute or if the RBC destruction does not exceed the conjugating capacity of the liver. Foals with NI are typically clinically normal at birth and develop signs after ingestion of colostrum. Clinical signs can occur from 5 hours to 7 days after birth. Foals typically have pale icteric mucous membranes
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Figure 45.1 Pale mucous membranes indicative of anemia. Figure 45.2 Pale icteric mucous membranes characteristic of neonatal isoerythrolysis.
(Figure 45.2). Severe hyperbilirubinemia can result in deposition of unbound, unconjugated bilirubin in the central nervous system (CNS), resulting in kernicterus.24 The clinical signs of kernicterus need to be differentiated from those seen with severe anemia and poor tissue oxygenation or hypoxic ischemic encephalopathy. The clinical signs attributed to kernicterus tend to be permanent and the seizures are non-responsive to phenobarbital or diazepam. Anemia of chronic disease is usually seen with clinical signs of chronic infection such as ill-thrift, infectious orthopedic disease, pneumonia, or omphalophlebitis. Fell ponies with erythroid hypoplasia are usually normal at birth; however, they fail to thrive and develop signs associated with immunodeficiency, including diarrhea, nasal discharge, hypersalivation, and coughing.21
Diagnosis The diagnosis of anemia is confirmed by an absolute decrease in the RBC count. This should be supported by characteristic clinical signs described above. A cause needs to be identified for treatment and management of the foal. RBC morphology may be of some assistance. Spherocytes may be seen with immune-mediated anemia; schistocytes and kerato-
cytes with DIC; pale erythrocytes with anemia of chronic disease; and anisocytosis in regenerative anemia (Figure 45.3). Blood loss Blood loss anemia is characterized by a decreased hematocrit and protein concentration with a normal bilirubin concentration. However, immediately after acute blood loss the RBC values remain unchanged until compensatory fluid shifts occur which result in reduced RBC values 12–24 hours later. Thus, decreased plasma protein concentrations are initially the best guide to acute blood loss. Bilirubin concentrations may increase with absorption of bilirubin from RBC breakdown if internal hemorrhage occurs. Ultrasonographic examination of the thorax or abdomen may find a hemothorax or abdomen (Figure 45.4). Large hematomas may be visualized associated with internal umbilical remnant hemorrhage. Gastroscopy, occult fecal blood tests, or urinalysis should be performed if bleeding from the gastrointestinal or urinary tract is suspected. Coagulation assays should be performed if hemostatic disorders are suspected. DIC is associated with thrombocytopenia, hypofibrinogenemia, prolonged
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tor VIII:C activity in the plasma, which results in abnormal function of the intrinsic coagulation system and thus a prolonged aPTT and normal PT is seen. von Willebrand’s disease is diagnosed if there is a reduced plasma von Willebrand factor (vWF) antigen concentration. The function of the vWF may also be reduced and is measured by the ristocetin cofactor activity of vWF.7 Petechial and eccyhmotic hemorrhages of the mucous membranes, subcutaneous hematomas, hemarthrosis, and melena may be present with hemostatic disorders. Reduced erythrocyte lifespan
Figure 45.3 Alterations in erythrocyte morphology, including anisocytosis, macrocytosis, spherocytes (S), and ghost cells(G). Erythrocytes are from an 8-month-old foal with primary idiopathic immune-mediated hemolytic anemia ¨ (May–Grunwald–Giemsa, original magnification 1000×). Reprinted from Lording, PM, Erythrocytes, Veterinary Clinics North America: Equine Practice. (2008) 24(2): p. 225–237. With permission from Elsevier.
prothrombin time (PT) and activated partial thromboplastin time (aPTT), and elevated fibrin degradation products;16 however, the changes seen will depend on the balance between fibrinolysis and coagulation at the time of sampling and reflect changes associated with sepsis. Hemophilia A has low fac-
Hemolytic anemia is characterized by a decreased hematocrit with normal plasma protein concentration. Hyperbilirubinemia (due to increased unconjugated/indirect bilirubin) and icterus occur when the conjugating capacity of the liver is exceeded. This results in pale, icteric mucous membranes. Pigmenturia (hemoglobinuria) and bilirubinuria may be present. There is often an accompanying neutrophilic leukocytosis. A low platelet and neutrophil count may be present if an alloantibody thrombocytopenia and neutropenia is also present. Clinical signs of sepsis may be present if bacterial toxin-induced hemolysis, as seen with Clostridium spp., is present or if the foal has concurrent sepsis. A diagnosis of NI is based on clinical signs and demonstration of maternal alloantibodies on the foal’s RBCs. Hemolytic cross-match is the most reliable diagnostic test. The alloantibodies responsible for NI are stronger hemolysins than agglutinins; thus, tests for agglutination are not as reliable.25 A Coombs’ test can also be performed to demonstrate antibodies or complement on the foal’s RBCs; however, this is not specific for NI and can be positive with other immune-mediated causes of anemia. The jaundiced foal agglutination (JFA) assay can be performed in the field and, in experienced hands, correlates well with the hemolytic test26 (see Chapter 35). Decreased erythropoiesis
Figure 45.4 Ultrasound image of a hemothorax demonstrating the characteristic ‘smoke swirling’ appearance of blood within the thorax. Viewed through an intercostal space using a linear 7.5 MHz probe; left is dorsal.
Anemia of chronic disease is characterized by a mild anemia, normal to elevated protein concentration due to an elevated globulin fraction, an inflammatory leukogram, and hyperfibrinogenemia. Identification of a localizing infection is also supportive. Iron deficiency anemia is usually normocytic and normochromic, and in more severe cases red blood cells become microcytic and hypochromic. Low serum iron levels, high TIBC, and low saturation of TIBC are characteristic.16 A bone marrow aspiration/biopsy is useful in identifying a non-regenerative anemia and can be performed to evaluate the myeloid and erythroid
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precursors. Foals with Fell pony syndrome have a reduction in erythroid precursors reflected in an abnormal M/E ratio of 21–62:1.21 Prussian blue stain is useful for assessing iron stores.16
Treatment and management The treatment and management of foals with anemia are aimed at improving tissue perfusion and oxygenation. Foals whose vital parameters are stable and do not have findings indicative of poor tissue oxygenation should be monitored closely, have exercise and stress minimized, and given time to correct the anemia without a transfusion. Foals that are weak, not nursing, and have poor tissue perfusion and oxygenation will need a blood transfusion. Foals with a concurrent disease such as sepsis or pneumonia which impair the oxygen-carrying capacity of the blood will benefit from a transfusion sooner than the blood indices indicate.16 Foals should be placed on intranasal oxygen insufflation at 5–10 L/min to increase the free oxygen in the blood and given intravenous fluids if hypovolemia is present. Blood transfusion Determining whether a blood transfusion is necessary depends on clinical and laboratory findings. Clinical signs alone can be indicative of the necessity for a blood transfusion. In acute cases, PCV is an unreliable indicator of the need for a transfusion; thus, indicators of impaired tissue oxygenation are used (Table 45.3). In more chronic cases (2–3 days) the PCV is more useful as an indicator for a transfusion. The volume of blood required is calculated by ((PCV desired – PCV observed) × body weight (kg) × 0.15 L/kg)/PCV of donor, where 150 mL/kg is the blood volume of a 2-day-old foal.16 Blood must be given though a filtered infusion line starting slowly, monitoring for a transfusion reaction, and then inTable 45.3 Laboratory indicators for a blood transfusion Laboratory parameter
Indication for transfusion
PCV
< 18% over a 24-hour period ≤ 12% over a longer time period < 5 g/dL Pv O2 < 30 mmHg Sv O2 < 50% > 4 mmol/L > 50%
Hb concentration Venous blood gas Blood lactate concentration Oxygen exchange ratio (O2 ER) ((Sa O2 – Sv O2 ∗ /Sa O2 ) × 100)
PCV, packed cell volume; Hb, hemoglobin; P v O2 , partial pressure of venous oxygen; Sv O2 , venous oxygen saturation; Sa O2 , arterial oxygen saturation; Sv O2 , mixed or central venous oxygen saturation.
creasing to a rate of 10–20 mL/kg/h. Washed dam’s RBCs are the ideal choice for transfusion for NI foals. A Qa- and Aa-negative blood donor is recommended for transfusions; however, if cross-matching is not available a Standardbred, Morgan, or Quarter Horse gelding with no prior history of blood transfusion is also a suitable choice. Hemoglobin-based oxygen-carrying solutions (Oxyglobin; Biopure Corp. Cambridge, MA, USA) can be used to temporarily improve oxygen-carrying capacity of the blood while the whole blood transfusion is being prepared.25 Specific treatments
Blood loss anemia External hemorrhage prevents the reutilization of RBCs; however, approximately two-thirds of RBCs are reabsorbed by lymphatics in cases of internal hemorrhage.16 Thus, if there is not severe respiratory compromise with a hemothorax, it is recommended not to remove the blood. A transfixion suture can be placed around the umbilical cord to stop bleeding. Hemostatic abnormalities may benefit from whole blood or fresh plasma transfusions or specific factor supplementation. Time and supportive care may be all that is required for alloimmune abnormalities.
Reduced erythrocyte lifespan Foals with hemolytic anemia are not commonly hypovolemic unless they have stopped nursing, but they do have reduced oxygen-carrying capacity. In both instances, therapy for hypovolemia should be instituted to improve the distribution of RBCs, and thus oxygen, to the tissues. Drug-induced or autoimmune anemias are treated with corticosteroids (dexamethasone 0.05–0.1 mg/kg intramuscularly (i.m.) or intravenously (i.v.) q. 12 hours). Foals with NI that are less than 48 hours old should be muzzled and fed an alternate milk source. The mare should be milked out regularly and milk discarded. Although the antibody absorption is usually complete after 24 hours, the author prefers to wait for 48 hours prior to allowing the foal to nurse. Mares with a history of NI or with anti-RBC antibodies should have their foal muzzled at birth and an alternative source of colostrum given to the foal until the laboratory results are available. If results are positive for NI, the foal should be fed an alternative source of milk and the mare regularly milked out and milk discarded for 48 hours. The foal’s serum immunoglobulin (Ig) G concentration should be measured to ensure adequate transfer of immunoglobulins. Blood typing the mare and stallion to assess their
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Anemia compatibility can be performed, and those mares with incompatibilities can be screened for anti-RBC antibodies 30 days prior to birth. A negative result can be misleading as screening may occur prior to antibody production.
Decreased erythropoiesis Correction of the underlying localized disease will resolve the anemia of chronic disease. Documented iron deficiency in foals can be treated with oral ferric sulfate (4 mg/kg).18 Ferrous fumerate supplementation in young foals has been associated with toxic hepatopathy and death in foals.27 Normal foals less than 2 weeks old are highly susceptible to iron toxicity as they have a high serum iron concentration, high percentage saturation of transferrin, and high iron absorption, and thus should not receive iron supplementation.27
Outcome The prognosis depends on the effects of reduced tissue oxygenation and underlying cause of the anemia. If severe tissue oxygen deprivation occurs resulting in seizures, severe metabolic acidosis, and organ dysfunction, the prognosis is grave. The prognosis is good for uncomplicated cases of NI. Occasionally, foals will continue to hemolyse following the blood transfusion and require multiple blood transfusions. The prognosis for these foals is poor owing to the high rate of liver failure associated with suspected iron toxicity and high serum concentration of bilirubin resulting in kernicterus.28
References 1. Axon JE, Palmer JE. Clinical pathology of the foal. Vet Clin North Am Equine Pract 2008;24:356–86. 2. Harvey JW, Asquith RL, McNulty PK, Kivipelto J, Bauer JE. Haematology of foals up to one year old. Equine Vet J 1984;16:347–53. 3. Harvey JW, Asquith RL, Sussman WA, Kivipelto J. Serum ferritin, serum iron, and erythrocyte values in foals. Am J Vet Res 1987;48:1348–52. 4. Doarn RT, Threlfall WR, Kline R. Umbilical blood flow and the effects of premature severance in the neonatal horse. Theriogenology 1987;28:789–800. 5. Darien BJ, Feldman BF. Hemostasis in the newborn foal. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 3rd edn. Philadelphia: W.B. Saunders Co., 1992; pp. 427–31. 6. Lassen ED, Swardson CJ. Hematology and hemostasis in the horse: normal functions and common abnormalities. Vet Clin North Am 1995;11:351–89. 7. Rathgeber RA, Brooks MB, Bain FT, Byers TD. Clinical vignette: von Willebrand’s disease in a thoroughbred mare and foal. J Vet Intern Med 2001;15:63–6.
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8. Laan TTJM, Goehring LS, Sloet van OldruitenborghOosterbaan MM. Von Willebrand’s disease in an eight-dayold quarter horse foal. Vet Rec 2005;157:322–4. 9. Buechner-Maxwell V, Scott MA, Godber L, Kristensen A. Neonatal alloimmune thrombocytopenia in a quarter horse foal. J Vet Intern Med 1997;11:304–8. 10. Ramirez S, Gaunt SD, McClure JJ, Oliver J. Detection and effects on platelet function of anti-platelet antibody in mule foals with experimentally induced neonatal alloimmune thrombocytopenia. J Vet Intern Med 1999;13:534–9. 11. Perkins GA, Miller WH, Divers TJ, Clark CK, Belgrave RL, Sellon DC. Ulcerative dermatitis, thrombocytopenia, and neutropenia in neonatal foals. J Vet Intern Med 2005;19:211–16. 12. Bailey E. Prevalence of anti-red blood cell antibodies in the serum and colostrum of mares and its relationship to neonatal isoerythrolysis. Am J Vet Res 1982;43:1917–21. 13. Traub-Dargatz JL, McClure JJ, Koch CK, Schlipf JW. Neonatal isoerythrolysis in mule foals. J Am Vet Med Assoc 1995;206:67–70. 14. Means Jr RT, Krantz SB. Progress in understanding the pathogenesis of the anemia of chronic disease. Blood 1992;80:1639–47. 15. Harvey JW. Normal hematologic values. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 561–70. 16. Vaala WE. Neonatal anemia. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 571–88. 17. Brommer H, Sloet van Oldruitenborgh-Oosterbaan MM. Iron deficiency in stabled Dutch warmblood foals. J Vet Intern Med 2001;15:482–5. 18. Fleming KA, Barton MH, Latimer KS. Iron deficiency anemia in a neonatal foal. J Vet Intern Med 2006;20:1495–8. 19. Chavatte P, Brown G, Ousey JC, Silver M, Cottrill C, Fowden AL, McGladdery AJ, Rossdale PD. Studies of bone marrow and leucocyte counts in peripheral blood and fetal and newborn foals. J Reprod Fertil Suppl 1991;44:603–8. 20. Sellon DC. Disorders of the hemopoietic system. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine. St. Louis: Saunders, 2004; pp. 721–68. 21. Scholes SFE, Holliman A, May PDF, Holmes MA. A syndrome of anaemia, immunodeficiency and peripheral ganglionopathy in Fell ponies. Vet Rec 1998;142:128–34. 22. Milne EM, Pyrah ITG, Smith KC, Whitwell KE. Aplastic anemia in a Clydesdale foal: case report. J Equine Vet Sci 1995;15:129–31. 23. Toribio RE, Bain FT, Mrad DR, Messer NT, Sellers RS, Hinchcliff KW. Congenital defects in newborn foals of mares treated for equine protozoal myeloencephalitis during pregnancy. J Am Vet Med Assoc 1998;212:697–701. 24. David JB, Byars TD, Braniecki A, Chaffin MK, Storts RW. Kernicterus in a foal with neonatal isoerythrolysis. Comp Cont Educ Pract Vet 1998;20:517–22. 25. Giguere S, Polkes AC. Immunologic disorders in neonatal foals. Vet Clin North Am Equine Pract 2005;21:241–72. 26. Bailey E, Conboy HS, McCarthy PF. Neonatal isoerythrolysis of foals: an update on testing. In: Proceedings of the 33rd Annual Convention of the American Association of Equine Practice, New Orleans, 29 November-2 December 1987, pp. 341–55. 27. Mullaney TP, Brown CP. Iron toxicity in neonatal foals. Equine Vet J 1988;20:199–24. 28. Polkes AC, Giguere S, Lester GD, Bain FT. Factors associated with outcome in foals with neonatal isoerythrolysis (72 cases, 1988–2003). J Vet Intern Med 2008;22:1216–22.
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Chapter 46
Coagulation Disorders Fairfield T. Bain
Overview Overt clinical signs of coagulation disorders in foals are uncommon, and clinical attention is usually drawn to investigate clotting function after observation of excessive or recurrent hemorrhage or thrombosis. In many cases, clinical recognition of hemorrhage may require close physical examination. On occasion, clinical signs of illness are vague and the coagulation disorder may be mistaken for the similar vague signs of illness observed with birth asphyxia or early sepsis. While not a complete treatise on disorders of coagulation, the intended goal of this chapter is to describe the clinical signs and diagnostic evaluation and management of the more commonly recognized clinical disorders of coagulation affecting the newborn foal.
Clinical signs and pathogenesis Signs of hemorrhage can vary from subtle signs such as petechiae and ecchymoses to frank, persistent bleeding. Occasionally, this hemorrhage can occur into body spaces, such as hemarthrosis or hyphema, or into tissues, such as perivascular hemorrhage following venipuncture, an intramuscular hematoma following injection or trauma, or hematoma within the pelvic canal. Readers are referred to additional references for detailed discussion of the coagulation cascade (Figure 46.1).1 The coagulation system exists to maintain vascular flow, as well as to prevent blood loss, and to play a role in the healing of wounds and tissue injury. Initiation of the coagulation system occurs when there is a disruption in the vascular wall. A general view is that the platelet is the first responder in the clotting process – acting to form a temporary plug to a break in the endothelial lining of the vessel as well as a scaffold for deposition and formation of the fibrin clot. Activation of the cascade of clotting factors then
occurs to form the fibrin plug as the final product in sealing a defect in the vessel wall. The clotting process involves a ‘cascade’ of activation of several factors. Classically, this activation process has been described as being initiated by either of two ‘arms’: the intrinsic process, meaning exposure to tissue thromboplastin, or the extrinsic process, meaning contact activation by exposure to capillary endothelial basement membrane components or collagen.1 Both lead to a final common pathway that results in conversion of fibrinogen to the fibrin clot. Disorders may involve some deficiency or abnormality of any other components along this cascade to produce a failure to clot, or excessive activation and thrombosis (an abnormal tendency to form a clot). Clotting disorders observed in foals vary from hypocoagulation (tendency toward hemorrhage) to hypercoagulation (tendency toward thrombosis).2 In one prospective study of sick neonatal foals, coagulopathies were recognized in 25% of the foals in the septic shock group and in 16% of the neonates in the septic group. Hypercoagulation has generally been considered to be relatively uncommon in foals, with the exception of arterial thrombosis in the distal limb observed with sepsis, which is still an uncommon condition. More recent research indicates that microvascular thrombosis may play a role in the progression of multiple organ failure with generalized sepsis and the systemic inflammatory response syndrome (SIRS) in the foal.3 Hypocoagulation has been more commonly reported in the literature, and conditions include thrombocytopenia (alloimmune being most common) and several clotting protein deficiencies (hemophilia A, von Willebrand’s disease). In some situations, accurate diagnosis will allow for specific treatment and salvage of the foal, whereas, in others, the diagnostic process results in recognition of a genetic process; in the latter case, breeders should be advised in order to prevent recurrence of the defect in future matings.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Intrinstic Pathway Factor 12 is activated with the aid of prekallikrein and high molecular weight kininogen (12a)
Common Pathway
Factor 9
Negatively charged surfaces
Extrinsic Pathway Factor 7
Factor 10
Factor 11 is activated (11a) Activated Factor 9 (9a)
Calcium, PF3
Factor 3
Calcium
Factor 8:Ca
Activated Factor 7 complex
Activated Factor 10 (10a) Calcium, PF3, Factor 5a Factor 2
Activated Factor 2 (2a) Fibrinogen
Fibrin Factor 13a Stable fibrin clot
Figure 46.1 Classical coagulation cascade. Reprinted with permission from Baker
Diagnostic process The initial recognition of a coagulation disorder involves characterization of the pattern of hemorrhage. Small vessel hemorrhages such as petechiae or ecchymoses are most suggestive of a platelet disorder. Close examination of the mucosal membranes (oral, conjunctival, and vulvar) as well as less pigmented regions of skin (pinna of the ears) is important in detection of petechiae in neonatal foals. Larger regions of hemorrhage, whether initiated by venipuncture or trauma, or spontaneous, might indicate a coagulation cascade disorder such as hemophilia. Laboratory evaluation of a bleeding disorder should include a complete blood count (CBC) followed by screening tests for coagulation abnormalities. The CBC might reveal information about anemia from hemorrhage as well as the platelet count. EDTA-preserved blood (lavender top tubes) is used for the CBC, whereas most other coagulation testing is performed using blood preserved in sodium citrate (3.8%) with an optimal dilution of 1:9 citrate:blood. Determination of the platelet count is a first step and may be performed by some of the newer automated hematology counters or manually using a hemacytometer. Continued small vessel hemorrhage in the presence of a normal platelet count suggests the possibility of a platelet adhesion disorder, as might occur with von Willebrand’s disease. Additional testing would be necessary to document this disorder or dif-
2004.1
ferentiate it from other platelet membrane adhesion molecule defects. Screening of the coagulation cascade with the coagulation panel of prothrombin time (PT) and activated partial thromboplastin time (aPTT) is used to determine the presence of an abnormality in the clotting cascade (Table 46.1). In general, prolongation of the PT is more indicative of an abnormality of the intrinsic segment of the coagulation cascade, whereas prolongation of the aPTT is more indicative of the extrinsic arm. A bleeding time may be performed in situations where the platelet count is normal, yet there is suspicion of a platelet function abnormality.
Common disorders of coagulation in the foal Thrombocytopenia One of the more commonly recognized clotting disorders in newborn foals is petechial and ecchymotic hemorrhage associated with thrombocytopenia.4 Alloimmune thrombocytopenia is probably the most commonly recognized mechanism of neonatal thrombocytopenia in the foal. Colostral origin alloimmune antibody directed against platelet membrane antigens has been proposed as the mechanism of this disorder. The onset of clinical signs is usually within the first few days (days 2–5) of life. Petechiae and ecchymoses on the mucosal membranes are the usual
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Table 46.1 Coagulation parameters from healthy foals (N = 5). Reprinted with permission from Bentz et al.2
Age (hours)
PT (seconds)
aPTT (seconds)
AT (%)
Platelet count (×103 /mL)
Fibrinogen (mg/dL)
FDP
<7
10.4 ± 0.4a 10.5 (10.4–10.6)b
52.8 ± 8.1 50.9 (47.9–51.2)
147.6 ± 15.0 152 (144–154)
381.6 ± 58.9 286 (245–325)
226.4 ± 57.3 221 (192–249)
1.2 ± 0.3 1 (1–1)
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10.6 ± 0.9 10.7 (9.9–11.3)
51.9 ± 10.0 51.3 (43.7–60.1)
128.5 ± 22.3 128 (113–144)
229 ± 50.8 231 (205–262)
251.0 ± 64.1 234 (201–301)
1.2 ± 0.3 1 (1–1)
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11.1 ± 1.8 10.5 (10.0–12.2)
55.6 ± 10.4 57.2 (47.6–63.5)
133.0 ± 20.7 138 (119–147)
222.2 ± 60.6 200 (180–264)
316.8 ± 42.8 321 (280–353)
1 ± 0.0 1 (1–1)
Mean ± standard deviation. 50th (25th–75th) percentiles. PT, prothrombin time; aPTT, activated partial thromboplastin time; AT, antithrombin; FDP, fibrin degradation products (1, < 10 mg/mL; 2, 10–40 mg/mL; 3, > 40 mg/mL).
a
b
initial signs, but other signs including hyphema or hemorrhage into tissues, such as perirectal hematomas associated with placement of a rectal thermometer, have been observed by the author. Thrombocytopenia has also been observed as a component of a syndrome involving mucosal hemorrhage, bulla formation, ulceration, and epidermal crusting thought to be consistent with a bullous pemphigoid-like disorder.5 In both sepsis and disseminated intravascular coagulation, there is a tendency toward microvascular thrombosis and consumption of platelets. Barton et al.6 showed that, although there was evidence of prolongation of PT and aPTT and an increase in fibrin degradation products, there was no evidence of platelet consumption, indicating more of a tendency toward hypercoagulation rather than hemorrhage with endotoxemia in foals. This appears to be borne out by more recent studies showing histopathological evidence of microvascular thrombosis in tissues of foals affected by bacterial septicemia.3 Treatment of alloimmune thrombocytopenia in the neonatal foal has been debated in the literature. Corticosteroids [dexamethasone, 0.04 mg/kg, intravenously (i.v.), every 24 hr] have been recommended to decrease adherence of alloimmune antibody to the platelets as well as to decrease binding to Fc receptors on fixed macrophages. The corticosteroid therapy is usually tailored to maintain the platelet count above a level considered protective against bleeding (> 20 000–30 000/µL). In most cases, the thrombocytopenia will resolve within 3–4 weeks. Some authorities believe that affected foals may eventually recover from the thrombocytopenia without the use of corticosteroids. Additional therapy may be indicated in management of the oral ulcerations observed with this syndrome. Affected foals
may be inappetent owing to pain associated with rupture of bullae or vesicles and ulcer formation. Oral lavage with an antiseptic solution (Listerine or dilute chlorhexidine solution) may aid in resolution of the mucosal ulcerations. Systemic antimicrobial agents may also be considered as many foals affected with this syndrome are also neutropenic, and, with mucosal injury, may be at risk for secondary bacterial infection. In most cases, affected foals recover after several days to 3 weeks with supportive care. The syndrome has been observed in foals from the same mare in subsequent years even though the mare was bred to a different stallion.7 Because of this, it has been recommended that foals from mares with a history of having affected foals receive colostrum from an alternative source to avoid absorption of alloimmune antibodies of colostral origin. von Willebrand’s disease Another mucosal hemorrhage disorder that has been observed in foals is von Willebrand’s disease.8, 9 This has been documented in a family of Thoroughbreds in which clinical signs of excessive and recurrent hemorrhage have been observed. Signs included persistent small vessel bleeding from the nasal mucosa and from trauma to the skin surrounding an umbilical hernia. One adult mare in this family had prolonged bleeding from the Caslick’s procedure. Glanzmann’s thrombasthenia There are several reports of bleeding associated with platelet adhesion disorders.10, 11 Glanzmann’s thrombasthenia, a disorder of platelet membrane surface antigen glycoprotein IIb (GPIIb), has been reported in horses, and would present as a membrane bleeding
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Coagulation Disorders disorder – epistaxis with petechiae and ecchymoses. Evaluation of the bleeding disorder in these cases generally shows a normal PT, aPTT, platelet count, and fibrinogen concentration, suggesting the possibility of a platelet function disorder. Hemophilia Hemophilia is a less commonly observed disorder of coagulation in foals. Certain families might be affected in different breeds.1, 12 The inheritance pattern is a recessive trait with males being more likely to be affected because the trait is sex-linked – located on the X chromosome. The coagulation panel would show a prolonged aPTT with a normal PT and platelet count. Measurement of factor VIII:C activity is required for specific diagnosis. Documentation of this defect is important in advising the breeder regarding future matings. Other coagulopathies Hyphema was one of the clinical observations in foals affected with the mare reproductive loss syndrome seen in central Kentucky in the spring seasons of 2001 and 2002.13, 14 It is surmised that the hyphema was related to microvascular injury as occurs with a systemic inflammatory response syndrome in a manner often associated with uveitis. Treatment of intraocular (anterior chamber) hemorrhage included mydriasis with topical atropine as well as systemic and topical anti-inflammatory medication. Hemorrhagic diathesis associated with vitamin K antagonism (compounds such as warfarin, dicoumerol, and brodifacoum that inhibit the enzyme vitamin K epoxide reductase) is rarely reported in foals, but should be a consideration with effusive hemorrhages and potential for exposure. Hypercoagulation A tendency toward thrombosis may be observed in foals with sepsis and activation of the coagulation cascade. Evidence for this has been recently demonstrated by the finding of capillary microvascular thrombi in the tissues of foals having died from sepsis.3 Similar to findings in human medicine, this is considered a feature of disseminated intravascular coagulation and sepsis, and can lead to multiple organ failure and clinical deterioration in the affected foal. Organ failure might be recognized as respiratory failure, hepatic failure, or renal failure in septic foals. Similarly, elevated levels of plasma d-dimer have been found to have a significant association with sepsis in foals and are thought to indicate activation of the coagulation system.15 Antemortem detec-
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tion or prediction of hypercoagulation remains difficult, and therapeutic efforts remain more theoretical than guided by distinct alterations in the coagulation parameters. Perhaps the most common clinical observation of hypercoagulation is jugular vein thrombosis with repeated venipuncture or with the presence of an intravenous catheter. Management of jugular thrombosis generally involves removal of any offending intravenous catheter, avoidance of venipuncture, and aspirin therapy to decrease platelet adherence. Topical anti-inflammatory medications [a topical nonsteroidal medication such as 1% diclofenac sodium cream (Surpass; Idexx Pharmaceuticals, Inc.), poultice] may be beneficial in reducing perivascular inflammation. Arterial thrombosis and distal limb infarction have also been reported in foals with severe sepsis, such as enterocolitis associated with salmonellosis.16 In these patients, it was speculated that endothelial injury associated with sepsis, possibly in combination with loss of antithrombin III, might have predisposed to the arterial thrombosis. No clinical signs of generalized coagulopathy were recognized prior to the clinical detection of distal limb infarction.
References 1. Baker DC. Diagnosis of disorders of hemostasis. In: Thrall MA, Baker DC, Campbell TW, DeNicola D, Fettman MJ, Lassen ED, Rebar A, Weiser G (eds) Veterinary Hematology and Clinical Chemistry. Philadelphia: Lippincott Williams & Wilkins, 2004; pp. 179–96. 2. Bentz AI, Palmer JE, Dallap BL, Wilkins PA, Boston RC. Prospective evaluation of coagulation in critically ill neonatal foals. J Vet Intern Med 2009;23:161–7. 3. Cotovic M, Monreal L, Armengou L, Prada J, Almeida JM, Segura D. Fibrin deposits and organ failure in newborn foals with severe septicemia. J Vet Intern Med 2008;22:1403–10. 4. Buechner-Maxwell V, Scott MA, Godber L, Kristensen A. Neonatal alloimmune thrombocytopenia in a quarter horse. J Vet Intern Med 1997;11:304–8. 5. Perkins GA, Miller WH, Divers TJ, Clark CK, Belgrave RL, Sellon DC. Ulcerative dermatitis, thrombocytopenia, and neutropenia in neonatal foals. J Vet Intern Med 2005;19:211–16. 6. Barton MH, Morris DD, Norton N, Prasse KW. Hemostatic and fibrinolytic indices in neonatal foals with presumed septicemia. J Vet Int Med 1998;12:26–35. 7. Sellon DC. Foal with immune-mediated thrombocytopenia. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case-Based Approach. Philadelphia: Saunders, 2006; pp. 46–50. 8. Rathgeber RA, Brooks MB, Bain FT, Byars TD. Clinical vignette. Von Willebrand disease in a Thoroughbred mare and foal. J Vet Intern Med 2001;15:63–6. 9. Laan TT, Goerhing LS, Sloet van OldruitenborghOosterbaan MM. Von Willebrand’s disease in an eight-dayold quarter horse foal. Vet Rec 2005;157:322–4. 10. Livesey L, Christopherson P, Hammond A, Perkins J, Toivio-Kinnucan M, Insalaco T, Boudreaux MK. Platelet
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dysfunction (Glanzmann’s thrombasthenia) in horses. J Vet Intern Med 2005;19:917–19. 11. Macierira S, Rivard GE, Champagne J, Lavoie JP, Bedard C. Glanzmann thrombasthenia in an Oldenbourg filly. Vet Clin Pathol 2007;36:204–8. 12. Littlewood JD, Bevan SA, Corke MJ. Haemophilia A (classic haemophilia, factor VIII deficiency) in a Thoroughbred colt foal. Equine Vet J 1991;23:70–2. 13. Byars TD, Seahorn TL. Clinical observations of mare reproductive loss syndrome in critical mares and foals. In: Proceedings of the First Workshop on Mare Reproductive Loss
Syndrome, College of Agriculture, University of Kentucky, Lexington, KY, 2002, pp. 15–16. 14. Sebastian MM, Bernard WV, Riddle TW, Latimer CR, Fitzgerald TD, Harrison LR. Mare reproductive loss syndrome. Vet Pathol 2008;45:710–22. 15. Armengou L, Monreal L, Tarancon I, Navarro M, Rios J, Segura D. Plasma D-dimer concentration in sick newborn foals. J Vet Intern Med 2008;22:411–17. 16. Brianceau P, Divers TJ. Acute thrombosis of limb arteries in horses with sepsis: five cases (1988–1998). Equine Vet J 2001;33:105–9.
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Section (i): Musculoskeletal System
47.
Fetal Ossification and Normal Joint Development Elwyn C. Firth
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Flexural Deformities Stephen B. Adams and Timothy B. Lescun
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Fractures Commonly Seen in Foals Angus R. Adkins
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Infectious Arthritis and Osteomyelitis Fairfield T. Bain
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Muscle Disorders Erica C. McKenzie and Jennifer M. MacLeay
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Tendon and Ligament Disorders Timothy B. Lescun and Stephen B. Adams
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Chapter 47
Fetal Ossification and Normal Joint Development Elwyn C. Firth
The horse is a grazing animal and has a very highly developed musculoskeletal system at the time of birth, compared with the newborn of nest animals such as rodents, carnivores, and humans. After some weeks, development of the nervous and locomotor systems is sufficient to allow attempts at locomotion when the young of some species leave the nest, but full development in human athletes, for instance, can take many years. The young of the herbivores can stand and ambulate within hours of birth, which is an evolutionary advantage that allows escape from predators at one of the two most vulnerable stages of life. At the time of transformation from uterine to terrestrial life at foaling, the cardiovascular and respiratory systems immediately must adapt, as occurs in all mammals, but the equine neonate must be behaviorally and neurologically capable in the first hours. The neurological capability of the foal to stand and follow its dam at rapid gaits shortly after birth may be the pinnacle of the transition from prenatal to postnatal life in terrestrial mammals. This perhaps overshadows the equally crucial capability of the development of musculoskeletal tissues before and after birth, which requires normal neuromuscular development and function. Movement at speed requires near-perfect neurological capability to apply and coordinate forces and movements, and structures which can resist the forces applied. Such movement in neonates would be impossible if tissue development were not adequate. For instance, the newborn foal’s longbones must be sufficiently rigid to resist bending and torsion forces, and the cartilages must not be so thick that the forces during ambulation result in mechanical failure of the tissue. However, the degree of development at birth is not complete, and for some weeks or months after
birth the tissues retain developmental plasticity and can respond to the environment positively by adaptive change in their form and composition. This chapter deals with the closely linked process of ossification and joint development, and there are several reasons for attention to this particular area of growth and development. If chondro-osseous development is less than normal at the time of birth, there are immediate effects on the ability of the foal to ambulate, a high risk of euthanasia in premature and infected survivors of perinatal intensive care, and compromised subsequent development which often affects future performance.1 Poor cartilage development at birth means that the likelihood of infective and non-infective chondro-osseous disease is greater than in foals with normal development.2 Osteochondrosis results from abnormal cartilage development and is said to be the biggest problem for horse breeders around the world,3 and although recent study implicates ischemic necrosis of cartilage due to interruption of blood flow, particularly in intracartilaginous vessels,4, 5 the exact cause remains unknown. Lesions have been demonstrated at6 or before7 birth. Good understanding of bone and joint development is required if these common diseases are to be better recognized, managed, and prevented. Like osteochondrosis (OC), other diseases such as angular and flexural limb deformity can be present at birth or appear within some weeks or months, but have an etiology even less well understood than that of OC. The etiological factors may lie in an area not yet investigated intensively in horses, namely the environment of pregnancy. In humans, epidemiological and experimental studies have highlighted a relationship between the compromise in the early phases of life (including periconceptual, fetal, and postnatal stages) and the subsequent development of disease
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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in later life. This is the ‘developmental origins of health and disease’ model, which speculates that the fetus adapts to environmental cues in utero, with permanent adaptive readjustments in homeostatic systems to aid survival. However, if there is a mismatch between the prenatal predictions and actual postnatal environment, these adaptations may ultimately be disadvantageous in postnatal life, leading to an increased risk of disease in adulthood.8 Most research has been directed to the relationship between an adverse prenatal environment and enhanced risk of major chronic diseases of later human life, but there is little reason why the same considerations will not be valid in the context of equine fetal growth and development and the current largely unexplained prevalence of orthopedic disease in young horses.
Nomenclature The description and nomenclature of the morphological features of the mammalian longbone are not complex, but confusing nomenclature has long impeded interpretation of research9 and still does not facilitate easy study by those interested in bone growth and development. The longbone in a neonatal foal consists of diaphysis, metaphysis, and epiphysis, the last two regions consisting mostly of cancellous bone. The metaphyseal spongy bone is surrounded by a thin outer shell of cortical bone, and the epiphyseal bone by a cortical shell and by a layer of cartilage of variable thickness. The epiphysis and metaphysis are separated by the physis, or metaphyseal growth plate, synonyms of which include ‘growth plate,’ ‘discoid growth plate,’10 and ‘longitudinal growth plate.’11 By division, hypertrophy, degeneration, mineralization, and ossification of the chondrocyte columns of the physis, new bone is formed at the metaphysis by endochondral ossification, which at this site is responsible for almost all of the growth in length of the longbone. Because the physis contributes bone to the metaphysis, and plays no part in the growth of the epiphysis,12 popular terms such as ‘epiphyseal plate,’ ‘epiphyseal disc,’ ‘epiphyseal cartilage,’ and ‘epiphysis’ should not be used when referring to the physis. Until a certain period of gestation, the epiphysis consists completely of cartilage (Figure 47.1). At this time the secondary ossification center (also referred to as the ossific nucleus or bony centrum) begins to develop in the cartilaginous epiphysis (Figure 47.2). This nucleus expands radially by endochondral ossification, and the epiphysis becomes progressively ossified and enlarges in all dimensions. Epiphyseal expansion thus contributes, to a small extent, to longbone lengthening. Endochondral ossification occurs
Figure 47.1 Sagittal slice of the most lateral (left) and medial (right) aspect of the distal radius of a foal of approximately 280 days gestation. The outline of the lateral styloid process can just be discerned on the joint and section surfaces. In neither the medial nor the lateral styloid process is the secondary ossification center formed, the physis–metaphysis junction is almost flat, and the epiphyseal cartilage contains many blood vessels which course up to but not into the articular cartilage.
Figure 47.2 Cabinet radiograph of two apposing decalcified transverse thick sections of the distal radius of a foal born dead at approximately 310 days gestation. The limb was perfused with radiodense liquid through the brachial artery. The radiographs indicate the major vessels entering the epiphysis from the caudal aspect of the radius, the obvious vasculature in the epiphyseal cartilage, and the sinusoidal filling in the bone of the secondary ossification centre of the distal radial epiphysis. Bottom right (inset) is a photograph of the surface of a midsagittal section of the contralateral distal radial epiphysis, the spherical secondary ossification center of which is visible as a round focus of cancellous bone. The physeal cartilage is visibly different from the epiphyseal cartilage, and the physis–metaphysis border is flat.
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Figure 47.3 Coronal section of the third metacarpal and proximal phalangeal bone of a Thoroughbred foal born prematurely at 264 days of gestation. Proximal to the metacarpophalangeal joint space is the incompletely ossified epiphysis of the third metacarpal bone. The ossifying center of the epiphysis is expanding radially into the epiphyseal cartilage. The latter is directly apposed to the articular cartilage and to the metaphyseal growth plate (= physis), which is colored differently from the epiphyseal cartilage. Similarly structured are the proximal and distal ends of the proximal phalangeal bone, in which the secondary ossification centers are very small because they have only recently begun to form. The proximal sesamoid bones (not shown) had not begun ossification. At this stage of pregnancy, ossification is rapid, but if it is delayed or the foal is born early or small for gestational age, then the cartilage is thicker than normal.
at the junction of the bony ossification center and the surrounding cartilage of the epiphysis, at the site of a spherical growth plate referred to as the ‘epiphyseal growth cartilage,’11 ‘spherical physis,’ or ‘secondary physis.’13 It is not desirable that ‘physis’ be used in such a context, since it is defined14 as the segment of tubular bone concerned mainly with growth in length of the bone, i.e., the metaphyseal growth plate. There may be other reasons for not applying the term physis to the growth cartilage of the epiphysis (see below). More recently, the term ‘articular epiphyseal cartilage complex’ has been used in the immature animal15 which refers to both kinds of cartilage surrounding the secondary ossification center, i.e., epiphyseal cartilage and articular cartilage. Figure 47.3 shows the morphology of the bone ends of a 264-day Thoroughbred fetus. In the area where the disk-shaped metaphyseal growth plate (physis) is apposed to the epiphyseal cartilage, a bipolar growth zone, consisting of cartilage of two different origins, is present. It is clearly evident in newborn foals, even grossly (Figure 47.4). As ossification of the epiphysis proceeds, the spherical growth plate in this area becomes replaced by
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Figure 47.4 Sawn frontal section of the distal radius and lateral styloid process (L) of a stillborn term Thoroughbred foal. The cartilage visible at the point of the arrow is 3 mm thick, and consists of articular cartilage (A) (approximately 1 mm thick) and epiphyseal cartilage. The vessels in the latter cartilage are too small to be visible in this illustration. The cancellous metaphyseal (M) bone is invested in a very thin cortical shell and periosteum, the latter of which is continuous with the periosteal–perichondrial complex (PPC) which encircles the periphery of the bone at the level of the physis. The cartilage of the physis (P) is directly apposed to the epiphyseal cartilage (EC).
a subchondral bony plate, also called ‘epiphyseal bone plate,’ ‘terminal bone plate,’16 ‘epiphyseal bone seal,’17 or ‘bony end plate.’11 The growth zone in the metaphysis then consists of the metaphyseal growth plate only, which comprises resting, germinal, proliferating, and hypertrophic cell layers. Because of the changing nature of the regional blood supply,9.16, 18 emphasis has been repeatedly placed on the distinction between the temporary bipolar growth zone prior to formation of the bone plate, referred to as the ‘presumptive growth plate,’ and the metaphyseal growth plate enclosed between metaphysis and the bone plate on the epiphyseal aspect of the physis, the ‘definitive growth plate.’17 Siffert11 indicated that, regardless of their anatomical location, growth cartilages apparently function in a similar biological manner. This supports the use19 of the term ‘metaphyseal equivalent site,’ which was used to indicate the significance of the vascular arrangement in the pathogenesis of hematogenous osteomyelitis in flat, irregular bones. Others have disagreed, indicating that the two growth cartilages, i.e., the physis and the growth plate of the epiphysis, are in fact different in various aspects, the evidence for which has recently been reviewed.20 Ossification is defined as the process of bone formation which usually begins at a particular primary ossification center in the bone’s pre-existent cartilaginous matrix of the future diaphysis, and later at either
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the physis or the growth plate of the epiphysis by the process of endochondral ossification.
Ossification and joint development Bone formation is an obvious determining factor in adequate joint development by the time of birth, with implications for immediate and later life. Also, the stage of joint development is generally quantified or subjectively assessed by measures of bone development rather than that of cartilage, mainly because of the simplicity and speed of radiographic examination. For these reasons some features of ossification and bone development are dealt with together. Flatbones Ossification in the flatbones, such as the ilium and the skull, occurs largely through outward drift of the cortices such that bone size and thickness remain in proportion as growth proceeds. This is achieved through longitudinal growth of the physes in the bone, as well by bone modeling, the rate of which is highest on the periosteal surface of the outer cortex and the endosteal surface of the inner cortex, with appropriate resorption rates on the endocortical surface of the outer and the periosteal surface of the inner cortex.21 This regulates the total thickness of the flat bone, as both the inner and outer cortices move outward, enlarging the dimensions of the whole bone. Obviously the flatbones of major clinical interest in horses, the pelvis and scapula, each have a diarthrodial joint (coxofemoral and scapulohumeral, respectively), which have morphology and development that are generally similar to the more distal joints. Diaphysis The primary ossification sites become radiographically evident from approximately 60 days gestation in the case of scapular body and diaphysis of humerus, radius, femur, and tibia, with the center in the third metacarpal/metatarsal bone appearing at 70–80 days, the proximal phalangeal diaphysis at 75–85 days, middle phalangeal bone appears later (120–155 days), and distal phalangeal bone at 85–135 days. These data were taken from horses of unrecorded breed and history.22 Epiphysis The time of radiographic appearance of the secondary ossification centers at the ends of longbones apparently does not vary greatly in the different bones, all appearing at 300 ± 35 days of gestation.22
The more proximal bones begin ossification earlier than those more distal, for instance femur at 220–245 days, but the nature of the data does not allow a singular strict comparison. However, the relative amount of ossification which must occur before development is complete differs between joints, because the epiphyses of the distal bones are much smaller than those of the most proximal bones. This is consistent with cartilage in the bone ends being thicker in joints more proximally situated, which has been shown in simple linear measurement of cartilage thickness.23 Epiphyseal development can be delayed by hypothyroidism, the effects of which may not be evident for weeks or months.24 None of the potential factors which could potentially affect bone development have been studied on large numbers of foals, so the significance of each is largely unknown. Carpal and tarsal bones These bones are assumed to have generally similar morphological development as the epiphysis of longbones, although detailed study of growth cartilages of cuboidal bones is lacking. The radiographic appearance of the carpal and tarsal bones was recorded by Getty22 and Soana et al.25 In both studies, the gestational age was estimated rather than known accurately, the foals were almost certainly not of the same breed, it is unclear whether some foals were small for gestational age, in the more recent study appearance times for the distal carpal row were imputed not directly observed, and obtaining a series with sufficient animals at regularly spaced gestational ages would have been difficult. One or more of these may account for the earlier gestational ages of appearance of ossification in the more recent study. Using both sources, the calcaneus begins ossification first at 120–155 days, far earlier than the talus (from 220 days), accessory carpal bone (250 days), and others from 275 to 335 days. Soana et al.25 mention that the third, fourth, and ulnar carpal bones are the last to begin ossifying in the carpus, and they are those most affected in angular limb deformity. This is consistent with the radiographically visible (wider joint space because of thicker cartilage) less advanced ossification of the lateral styloid process compared with the medial process of the distal radius. Like epiphyseal development, severe retardation of cuboidal bone development, possibly associated with other congenital developmental defects, can occur as a result of hyperplastic goiter.26 Other causes of poor ossification have been less well defined. Foals with shorter gestation and or retarded intrauterine growth have poor cartilage development in the epiphyses and cuboidal bones, as evidenced by wide radiolucent carpal and tarsal joint spaces in
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Fetal Ossification and Normal Joint Development radiographs of foals less than 2 weeks old and of known body weight and gestational length.27 Like retained cartilage,28 poorly ossified thick cartilage most likely has a greater tendency to deform and fail under load than does thinner cartilage. This aspect of cartilage maturation may be of great importance in recognizing foals likely to sustain untoward musculoskeletal consequences owing to their being born small and/or early, because both the thick cartilage and the vessels within it may be vulnerable to forces, even the normal forces of rising, standing, and walking. Hence the recommendation to start monitoring skeletal development by radiography or ultrasonographic examination soon after birth, either radiographically as above or ultrasonographically.29 Sesamoid bones Ossification of sesamoid bones begins later than that of the epiphysis with which they articulate. In the case of proximal sesamoid bones (290–330 days) and the distal sesamoid (navicular) bone (325 days to birth, or postnatal), the ossification begins at the end of the range of gestational age at which ossification of respectively distal epiphysis of the third metacarpal bone (265–290 days) and of the epiphysis of the distal aspect of the middle phalangeal bone (310–330 days) begin. In the patella, however, ossification begins from 325 days, long after that of the distal femoral epiphysis (220–245 days). The far longer time lag between beginning of ossification of the distal femoral epiphysis (prenatal) and patella (3–12 years) in humans was related to non-conformity of the femoral and patellar cartilages.30 Bipartite sesamoid bones, presumably due to a separate ossification center, are not uncommonly evident at birth.31 The diarthrodial joint The joint is constituted of several tissues, including metaphyseal cancellous bone and the thin metaphyseal cortical shell, epiphyseal cancellous bone, and the cortical shell of the epiphysis, articular calcified cartilage, hyaline articular cartilage, the growth cartilage of the epiphysis, joint capsule, synovial membrane, collateral ligaments, intra-articular ligaments, physis, and periosteal–perichondrial complex. At least in some parts of some joints, the joint capsule may be inserted at the metaphysis, i.e., diaphysad to the physis, and thus metaphysis, periosteal–perichondrial complex and physis are partly intra-articular in some joints, and hence included in the tissues which constitute the joint as an ‘organ.’ The latter term is proposed because of the intimate association of all these tissues, which are effectively inseparable in terms of structure,
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blood supply, function, and pathology. This has long been evident in terminologies such as ‘chondroosseous,’ ‘osteochondral,’ and ‘osteoarthritis,’ and in the pathology of disease which can extend across bone, cartilage, and soft tissues, in some cases very aggressively and rapidly as in infective joint disease or over years in the case of osteoarthritis. The development of each particular tissue is outside the scope of this chapter, and the most important aspects of development have been described above. In postnatal life, tissues are known to respond to the loads placed upon them. Aside from isometric training regimens in humans, all stimuli associated with adaptive responses of musculoskeletal tissues involve movement, since it is the imposition of force by muscles which induces the responses in the tissues which are deformed by the force.32 It is presumed that these or similar mechanisms related to adaptive change in postnatal life are also operative in utero. Only recently has specific attention been paid to the vascularity of the developing joint at the particular sites affected by osteochondrosis. Vessels enter the epiphysis, usually fairly close to the physis, cross the epiphysis and pass through the developing secondary ossification center, and continue to within approximately 1 mm of the joint surface, i.e., the vessels remain in epiphyseal cartilage of the articular epiphyseal cartilage complex, and traverse to the junction of epiphyseal cartilage and articular cartilage but do not enter the latter. As the ossification front proceeds radially outward to occupy more of the cartilaginous epiphysis, more of the vessels become included within the bone tissue as it forms from the epiphyseal cartilage. Vessels which originate from the perichondrium, some of which may course roughly parallel to the advancing ossification front, also become incorporated in the tissue behind the ossification front. At the ossification front, blood flow in the vessels from either origin is affected in such a way to cause an ischemic necrosis of epiphsyeal cartilage.33 The nature of the vascular abnormality causing the ischemia is not defined, and other initiating events have been suggested for the disease.7, 34 The degree of ossification, which is affected by several factors in intrauterine life, is thus important since the lesser the ossification in a particular joint, the thicker the cartilage. Thicker cartilage contains more intracartilagenous vessels than does thin cartilage, and the vessels thus remain longer in postnatal life as epiphyseal expansion continues.
Implications of abnormal ossification and joint development The reason for the above explanation of normal ossification and joint development is that postnatal
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morbidity and mortality, reduced growth, and poor athletic performance later in life can result from compromised chondro-osseous development. Measures of adequate musculoskeletal development include stature and size at birth, amount of tissues in relation to each other (such as the relationship between bone and muscle masses), bone size (determined in utero in human fetuses), and thickness of cartilage, which can be determined from radiographic or ultrasonographic examination. The manifestations of poor development may be obviously to subtly evident soon after birth. Aside from rare conditions such as intrauterine fractures of longbones, and the multiple congenital abnormalities,26 most defects associated with abnormal ossification are the pediatric orthopedic diseases, which include angular limb deformity, tarsal collapse, osteochondrosis, joint and bone infection, flexural deformity, and digital flexor tendon hyperextension. Abnormal or delayed development results in greater than normal thickness of the articular epiphyseal cartilage complex, resulting either directly from the developmental disorder and/or secondary to the postnatal sedentarism associated with perinatal disease. Most of the clinically significant abnormalities have been shown to be associated with epiphyseal cartilage that is thicker than normal for age. There is variable change in thickness with age,23 between foals and between joints. The thick cartilage contains blood vessels which remain until the epiphyseal cartilage becomes almost fully ossified,35 and thus the thicker the epiphyseal cartilage at birth, the longer after birth the cartilage remains thicker than normalfor-age. The longer this situation pertains, the longer it is susceptible to either infective or aseptic disease. Thick cartilage thus is incriminated in the pathogenesis of the two main categories of disease, infective and non-infective. Infective chondritis and osteomyelitis, referred to as E (epiphyseal)-type bone infection,36, 37 begins at the chondro-osseous junction, particularly in predisposition sites in the epiphyses,38 and its initiation appears to be rare in foals older than 4–5 weeks of age. The reason for involvement of intracartilagenous vessels in infection settling at the predisposition site is their morphology: the vessels end in an arcade from which apparently ‘end’ vessels protrude toward but not into the articular cartilage.35 It is believed that the sluggish hypoxic conditions at the metaphyseal chondro-osseous junction, which is the main site of septic physeal–metaphyseal bone infection in other animals and in children, and known to have low oxygen pressures,39 may also occur in the epiphyseal predisposition site of bone infection. The subarticular vessel arcades are present in thicker cartilage, and
are not visible after the total cartilage (articular plus epiphyseal cartilage) thickness is less than approximately 2 mm, which probably explains the apparent immunity of normally developing foals to E-type infection after several weeks. Epiphyseal osteomyelitis in infants was first alluded to in the orthopedic literature in 1981,40 and, even after decades of use of magnetic resonance imaging, it remains a rare disease.41 Because human epiphyses are cartilaginous and contain intracartilaginous vessels for years after birth, the existence of susceptible vessels (for a few weeks) in thick equine epiphyseal cartilage might alone not explain the higher occurrence of epiphyseal than metaphyseal bone infection in foals. Perhaps the vessels are compromised as well, by the stress imposed on and in cartilage by the forces of locomotion.34 The aseptic diseases appear to be related to thicker than normal-for-age cartilage. In utero, articular cartilage appears as a relatively homogeneous, very dense mixture of cells in a highly hydrated matrix of proteoglycan and collagen. The matrix is much softer than in mature tissue, and the collagen network is organized differently. The solid content of the cartilaginous epiphysis increases slowly from 12% in the third-trimester fetus to 20% in 4- to 8-week-old calves,42 but the mechanical properties are less well studied than those of the articular cartilage itself. In foals, it appears that the locations with the thickest cartilage are associated with the common manifestations of aseptic disease. For instance, the thicker cartilage in the fourth carpal bone and lateral aspect of the third carpal bone is apparently associated with one form43 of the most common of the angular limb deformities, carpal valgus. Foals with tarsal collapse (Figure 47.5), fractures, or osteomyelitis were in many cases twins or known to have been born prematurely.2 Articular osteochondrosis has been proposed to be associated with a defect in the vascular supply which nourishes the epiphyseal cartilage of the articular–epiphyseal complex,4, 33 and in pigs the lesions are proposed to be due to an interruption of these same blood vessels at the junction of cartilage and bone.44 Such associations do not prove cause and effect, however, and detailed study of cartilage thickness, biomechanical properties of epiphyseal cartilage, and the relationship to degree of fetal development of foals is lacking. However, it seems incontrovertible that thick cartilage is the critical parameter affected and has the greatest clinical influence postnatally. The other main categories of aseptic musculoskeletal disease in foals associated with abnormal development are digital flexor tendon laxity and flexural deformities. These are often congenital and a form of benign disease, of unknown etiology, and are
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Figure 47.5 (a) Lateral radiograph of the tarsus of a 327-day Thoroughbred foal, weighing 37 kg and presented for prematurity and hypoxic ischemic syndrome (HIS). It was radiographed on admission, hospitalized for 7 days, and boxed for another 2 weeks before being allowed a restricted exercise program. (b) An example of the result of less than ‘complete for gestational age’ ossification in the tarsus from a weanling that was born at 325 days of gestation with no maturity or other health issues. The radiograph was part of a weanling pre-sale inspection series. The distal tarsal row has an obvious abnormality, and the current common presumption is that this usually is associated with inadequate ossification at birth.
not so far known to be associated with abnormalities of ossification of joint development. Some cases of the congenital flexural deformities are associated with abnormal joint development, most likely secondary to the abnormal force regimen acting on the joint after the initiation of the deformity. The etiopathogenesis remains unknown, and, although several causative associations have been recorded,45 the relatively high prevalence of the disease probably means that toxins, infection, and gene mutation are less likely to be main causative factors than are nutritional abnormality, uterine malpositioning or other causes of reduced fetal motility, and other as yet undefined maternal environmental influences which affect connective tissue metabolism.
References 1. Adams A. Non-infectious orthopedic problems. In: Koterba AM, Drummond WH, Kosch P (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; p. 333. 2. Firth EC, Goedegebuure SA, Dik KJ, Poulos PW. Tarsal osteomyelitis in foals. Vet Rec 1985;116:261–6. 3. Jeffcott LB. Osteochondrosis: an international problem for the horse industry. J Equine Vet Sci 1996;16: 32–7.
4. Carlson CS, Cullins LD, Meuten DJ. Osteochondrosis of the articular epiphyseal cartilage complex in young horses: evidence for a defect in cartilage canal blood-supply. Vet Pathol 1995;32:641–7. 5. Olstad K, Ytrehus B, Ekman S, Carlson CS, Dolvik NI. Early lesions of osteochondrosis in the distal tibia of foals. J Orthop Res 2007;25:1094–105. 6. Rejno S, Stromberg B. Osteochondrosis in the horse. II. Pathology. Acta Radiol Suppl 1978;358:153–78. 7. Lecocq M, Girard C, Girard C, Fogarty U, Beauchamp G, Richard H, Laverty S. Cartilage matrix changes in the developing epiphysis: early events on the pathway to equine osteochondrosis?’ Equine Vet J 2008;40:442–54. 8. Gluckman PD, Cutfield W, Hofman P, Hanson MA. The fetal, neonatal, and infant environments: the long-term consequences for disease risk. Early Hum Dev 2005;81:51–9. 9. Spira E, Farin I, Karplus H. The capillary conducting channels of the distal epiphyses of the radius and ulna in the rabbit. J Anat 1963;97:255–8. 10. Ogden JA. Chrondro-osseous growth and development. In: Urist MR (ed.) Fundamental and Clinical Bone Physiology. Philadelphia: Lippincott, 1980. 11. Siffert RS. Ossification of the osteochondroses. Clin Orth Related Res 1981;158:10–18. 12. Firth EC. Functional joint anatomy and its development. In: McIlwraith CW, Trotter GW (eds) Joint Disease in the Horse. Philadephia: W.B. Saunders, 1996; pp. 80–6. 13. Ogden JA. Injury to the growth mechanisms of the immature skeleton. Skeletal Radiol 1981;6:237–53.
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14. Anon. Dorland’s Illustrated Medical Dictionary. Philadelphia: Saunders, 1974. 15. Carlson CS, Hilley HD, Meuten DJ. Degeneration of cartilage canal vessels associated with lesions of osteochondrosis in swine. Vet Pathol 1989;26:47–54. 16. Trueta J. Studies of the Development and Decay of the Human Frame. London: Heinemann, 1968. 17. Moss-Salentijn AGM. The Epiphyseal Vascularisation of Growth Plates. Utrecht: Utrecht University, 1976. 18. Brookes M. The Blood Supply of Bone. New York: AppletonCentury-Crofts, 1971. 19. Nixon GW. Hematogenous osteomyelitis of metaphysealequivalent zones. Am J Roentgenol 1978;130:123–9. 20. Byers PD, Brown RA. Cell columns in articular cartilage physes questioned: a review. Osteoarthritis Cartilage 2005;14:3–12. 21. Parfitt AM. The mechanism of coupling: a role for the vasculature. Bone 2000;26:319–23. 22. Getty R. Equine osteology. In: Getty R (ed.) Sisson and Grossman’s The Anatomy of the Domestic Animals. Philadelphia: W.B. Saunders, 1975; pp. 255–317. 23. Firth EC, Greydanus Y. Cartilage thickness measurement in foals. Res Vet Sci 1987;42:35–46. 24. Irvine CHG. Hypothyroidism in the foal. Equine Vet J 1984; 16:302–6. 25. Soana S, Gnudi G, Bertoni G, Botti P. Anatomoradiographic study on the osteogenesis of carpal and tarsal bones in horse fetus. Anat Histol Embryol J Vet Med Ser C Zentralblatt Veterinarmedizin Reihe C 1998;27:301–5. 26. McLaughlin BG, Doige CE. Congenital musculoskeletal lesions and hyperplastic goitre in foals. Can Vet J 1981;22: 130–3. 27. Adams R, Poulos PW. A skeletal ossification index for neonatal foals. Vet Radiol Ultrasound 1988;29:217–22. 28. Hurtig HB, Pool RR. Pathogenesis of equine osteochondrosis. In: McIlwraith CW, Trotter GW (eds) Joint Disease in the Horse. Philadephia: W.B. Saunders, 1996; p. 342. 29. Ruohoniemi M, Hild´en L, Salo L, Tulamo R.-M. Monitoring the progression of tarsal ossification with ultrasonography and radiography in three immature foals. Vet Radiol Ultrasound 1995;36:402–10. 30. Sarin VK, Carter DR. Mechanobiology and joint conformity regulate endochondral ossification of sesamoids. J Orthop Res 2000;18:706–12.
31. Thompson KN, Rooney JR. Bipartite proximal sesamoid bones in young thoroughbred horses. Vet Radiol Ultrasound 1994;35:368–70. 32. Frost H. Bone ‘mass’ and the ‘mechanostat’: a proposal. Anat Rec 1987;219:1–9. 33. Olstad K, Ytrehus B, Ekman S, Carlson CS, Dolvik NI. Epiphyseal cartilage canal blood supply to the tarsus of foals and relationship to osteochondrosis. Equine Vet J 2008; 40:30–9. 34. Hurtig MB, Pool RR. Pathogenesis of equine osteochondrosis. McIlwraith CW, Trotter GW (eds) Joint Disease in the Horse. Philadelphia: W.B. Saunders, 1996; pp. 335–58. 35. Firth EC, Poulos PW. Vascular characteristics of the cartilage and subchondral bone of the distal radial epiphysis of the young foal. N Z Vet J 1993;41:73–7. 36. Firth EC, Dik KJ, Goedegebuure SA, Hagens FM, Verberne LRM, Merkens HW, Kersjes AW. Polyarthritis and bone infection in foals. Zbl Vet Med B 1980: 27; 102–24. 37. Firth EC. Current concepts of infectious polyarthritis in foals. Commissioned article. Equine Vet J 1983: 15; 5–9. 38. Firth EC, Goedegebuure SA. The site of focal osteomyelitis lesions in foals. Vet Q 1988;10:99–108. 39. Brighton CT. Structure and function of the growth plate. Clin Orthop Relat Res 1978;136:22–32. 40. Green NE, Beauchamp RD, Griffin PP. Primary subacute epiphyseal osteomyelitis. J Bone Joint Surg Am 1981; 63:107–14. 41. Ezra E, Cohen N, Segev E, Hayek S, Lokiec F, Keret D, Wientroub S. Primary subacute epiphyseal osteomyelitis: role of conservative treatment. J Pediatr Orthop 2002; 22:333–7. 42. Williamson AK, Chen AC, Sah RL. Compressive properties and function-composition relationships of developing bovine articular cartilage. J Orthop Res 2001;19: 1113–21. 43. McLaughlin BG, Doige CE, Fretz PB, Pharr JW. Carpal bone lesions associated with angular limb deformities in foals. J Am Vet Med Assoc 1981;178:224–30. 44. Ytrehus B, Ekman S, Lundeheim N, Mathisen L, Reinholt FP, Teige J. Focal changes in blood supply during normal epiphyseal growth are central in the pathogenesis of osteochondrosis in pigs. Bone 2004;35:1294–306. 45. Kidd JA, Barr ARS. Flexural deformities in foals. Equine Vet Educ 2002;14:311–21.
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Chapter 48
Flexural Deformities Stephen B. Adams and Timothy B. Lescun
Flexural limb deformity is a common musculoskeletal condition in foals. Deformities may be either congenital or acquired, and typically involve the lower limb joints, particularly the carpal, metacarpophalangeal, or distal interphalangeal joints. Flexural deformities are described by the affected joint, although the primary abnormality is more often related to the muscles, tendons, and ligaments that control joint position. A flexural deformity is present when a joint is positioned in a state of hyperflexion, either permanently or intermittently. The condition is not known to have a breed predisposition and has been reported to account for up to one half of all congenital defects found at necropsy in foals.1
Pathogenesis Congenital flexural deformities Congenital flexural deformity most commonly affects the carpus and the metacarpophalangeal joint (Figure 48.1). Proposed mechanisms for the occurrence of these deformities include intrauterine malpositioning, abnormal nervous and/or musculotendinous development in utero, teratogenic insults, and genetic abnormalities of the foal.2, 3 In most cases a definitive cause is not determined. Acquired flexural deformities The pathogenesis of acquired flexural deformity is likely to be multifactorial in nature. Current understanding focuses on the role of a ‘pain withdrawal response’ in the affected limb(s). This pain withdrawal response results in an increase in tone of the flexor muscles/tendons relative to the extensor muscles/tendons, resulting in hyperflexion of the joints influenced by the affected musculotendinous unit. The detection of significant numbers of myofibroblasts within the inferior check ligament and deep
digital flexor tendon of young foals,4 and clinical improvement following analgesia in acute cases adds plausibility to this theory. Underlying sources of pain may be sole bruising, physitis, or other developmental orthopedic conditions such as osteochondrosis.
Diagnosis and evaluation The diagnosis of flexural limb deformities in foals is made by observation and physical examination of the abnormal limbs. Congenital deformities may result in dystocia during parturition, or are observed during the first hours of life. Acquired deformities develop over a period of days to weeks in most cases. The clinician should determine the cause, duration, and severity of the flexural deformity. Radiographs should be taken of the affected sites in neonatal foals with severe deformities and in foals with chronic acquired deformities to determine if there are abnormalities of the skeleton. Neonatal foals should be evaluated for systemic sepsis and joint infections, particularly if initial standing and nursing were delayed. All affected joints should be extended manually to determine if they can be reduced into a normal position (Figure 48.1a). Recently acquired flexural deformities in older foals often respond quickly to treatment, whereas chronic deformities may have secondary changes to the skeleton, joint capsule, and associated ligaments due to chronic joint hyperflexion. The severity of acquired flexural deformity of the distal interphalangeal joint is evaluated by measuring the hoof–ground angle, as determined by the dorsal hoof wall and the ground surface. The hoof–ground angle is ≤ 90◦ , 90 > 115◦ , and ≥ 115◦ in foals classified with mild, moderate, and severe deformities, respectively. Recommendations for treatment are based on this classification.5 Acquired flexural deformities of the metacarpophalangeal joint in older foals should also be
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Figure 48.1 (a) Foal with moderate to severe bilateral carpal and metacarpophalangeal flexural deformity. Manual extension of the limb is being performed to assess the severity of the deformity. (b) Foal with moderate bilateral carpal flexural deformity. Note the compensation required in the rear limbs to allow ambulation. Image courtesy of Dr Danah Nuest.
evaluated for severity. The normal dorsal angle of the metacarpophalangeal joint is about 140◦ . Horses with mild flexural deformities have upright angles approaching 180◦ . These horses often stand with the joint behind the center of the hoof but intermittently knuckle forward so that the fetlock is dorsal to the hoof. This dorsal position transfers load to the extensor tendons. Horses with moderate deformities have flexion of the metacarpophalangeal joint > 180◦ while standing, with the fetlock joint positioned forward of the center of the hoof. However, when these horses walk or trot the metacarpophalangeal joint may move back to a more normal position. Severely affected horses have a metacarpophalangeal joint that is flexed past 180◦ at all times. The extensor tendons support the horse’s weight and no stress is placed on the flexor tendons or suspensory ligament. Forced extension of severely affected joints by pushing firmly back on the fetlock with the horse bearing weight will not result in a more normal posture (Figure 48.2).
Treatment Congenital flexural deformities Treatment of mild to moderate congenital flexural deformity is often successful with medical and nursing management. Severe deformities, in which the foal is unable to stand unassisted, result in an increased risk of secondary health problems, such as failure of passive transfer, pneumonia, decubital ulceration, and systemic sepsis, and attention to these factors, as well as good nursing care, are an essential part of treatment in these foals. For mild cases, where the foal is able to stand and nurse unassisted, and ambu-
Figure 48.2 Yearling with moderate to severe bilateral metacarpophalangeal flexural deformity. (a) Standing in a flexed position results in the joint positioned dorsal to the foot. (b) Forced extension of the joint results in minor improvement in the joint position.
late without additional stress on other limbs, spontaneous resolution without specific treatment is possible. However, if improvement is not observed within 2–3 days of birth, treatment should be instituted. In cases of moderate deformity, where the foal is able to stand and nurse with minimal assistance but is forced to compensate for the deformity by placing added stress on other limbs (usually rear limbs), immediate treatment is recommended (see Figure 48.1b). The application of bandages and splints or custom-fit braces to place affected joints in or close to a normal position is often an effective treatment in mild and moderate cases, combined with judicious use of analgesics. Splints should be repositioned on the limb every 6–12 hours or applied for short periods of time in an effort to avoid the development of pressure sores. Splinting may be required for 2–3 days in mild cases. Moderate and severe deformities may require splinting for several weeks. In cases with no improvement following 2–3 days of splinting, other methods of treatment should be considered. Splinting can be discontinued when the affected joint remains in a normal or near normal position without intervention. Oxytetracycline has been used successfully in neonates to relax the flexor muscles/tendons and allow improvement in the degree of flexural deformity, particularly deformity of the metacarpophalangeal joint.6, 7 The response to treatment is rapid but transient, with relaxation in normal foals shown to subside within 4 days. However, this transient period of relaxation is often enough to allow affected joints to attain a normal position and ultimately permanent correction. Dosages of 22–44 mg/kg of oxytetracycline have been recommended, repeated daily for up to three treatments. Diluting the dose is a common practice to avoid complications of rapid
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administration. Attention should be paid to the effect on other joints, as muscle/tendon relaxation is generalized and hyperextension of the metatarsophalangeal and distal interphalangeal joints may occur. The potential for nephrotoxicity exists with oxytetracycline treatment, especially in the compromised neonate.8
Surgical treatment Surgical treatment of congenital flexural deformities is rarely employed. Surgical treatment of severe cases of metacarpophalangeal joint flexural deformity may require severance of all structures on the palmar surface of the joint followed by limb casting as a salvage procedure. Even with this approach, it is our experience that few cases are able to be corrected to a normal joint position due to involvement of the entire joint capsule and supporting soft tissues. Dramatic alteration of the joint angle using this approach also risks damage to the vascular supply of the foot, with stretching and thrombosis of the digital vasculature. Alternatively, fetlock arthrodesis via osteotomy of the articulating bone ends can be performed,9 which may reduce the risk of vascular damage following correction of the deformity. Transection of the ulnaris lateralis and flexor carpi ulnaris tendons of insertion has been reported as a successful treatment for mild and moderate cases of carpal flexural deformity that do not respond to other treatments.10, 11 The procedure is straightforward, cosmetic results are excellent, and horses are not hampered athletically in the long term. In severe cases, this treatment alone is unlikely to be successful. Acquired flexural deformities
Distal interphalangeal joint Acquired flexural deformity of the distal interphalangeal joint usually occurs in foals aged 6 weeks to 8 months. The deformity causes the angle formed by the dorsal hoof wall and the ground under the sole to increase from normal. With slowly developing deformities the hoof wall grows long and the heel maintains contact with the ground. Over several months, the heel may grow as long as the toe giving the foal a club foot or box-shaped foot (Figure 48.3a). In rapidly developing flexural deformities the heels will elevate off the ground and the foal will ‘walk on its toes’, the so called ballerina foal (see Figure 48.3b). Non-surgical treatment Balanced nutrition with energy reduction should be ensured in rapidly growing foals. Exercise should be restricted to walking several times daily. Sources of
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Figure 48.3 (a) Mild acquired flexural deformity of the left front distal interphalangeal joint. Note the dorsal hoof angle is less than 90◦ to the ground surface and the long heels create a typical ‘club foot’ appearance. (b) Severe acquired bilateral flexural deformity of the distal interphalangeal joint in a foal. Note the angle of the dorsal hoof wall to the ground is greater than 115◦ and the heels do not contact the ground. This is also termed a ‘ballerina foal’. Image courtesy of Dr Kira Delfs.
pain such as bruised feet, physitis, and osteochondrosis should be identified and treated directly. Analgesics should be administered to all foals to reduce pain. To relax deep digital flexor muscles, oxytetracycline may be administered intravenously to young foals that have rapid and very recent development of the deformity. Long heels should be shortened with a rasp. In foals with mild flexural limb deformities the heels may contact the ground after being rasped. In these foals surgery may not be necessary. Worn, bruised, very short, or deformed toes should be protected with hoof acrylic or glue-on shoes. We generally recommend surgery in foals that fail to respond to these methods within 2 weeks. Surgical treatment The severity of the condition determines the surgical procedure necessary for correction of flexural deformities of the distal interphalangeal joint. Desmotomy of the accessory ligament of the deep digital flexor tendon (inferior check ligament) is the treatment of choice in foals with mild deformities in which the hoof–ground angle is 90◦ or less and that have failed to respond to non-surgical therapy.5 This surgery is relatively easy, very effective for mild deformities, and usually improves the flexural deformity immediately (Figure 48.4). Following surgery, ongoing hoof trimming or shoeing with a small toe extension is recommended to maintain a normal hoof–ground angle and distal interphalangeal joint position. The surgery may leave a small cosmetic blemish with thickening in the area of the ligament, and owners of horses
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Figure 48.4 Mild acquired bilateral flexural deformity of the distal interphalangeal joint in a weanling foal. (a) Following nonsurgical treatment efforts of hoof trimming and analgesics. (b) One day following surgical treatment with an inferior check ligament desmotomy and further lowering of the heels. Note the improvement in dorsal hoof angle. (c) Following application of toe extension shoes. Toe extensions extend to the anticipated normal position of the dorsal hoof wall to assist stretching and lengthening of the deep digital flexor musculotendinous unit.
intended for sale should be informed of this potential. Descriptions of the surgery can be found in many textbooks.12, 13 Desmotomy of the inferior check ligament may be successful in some foals with moderate flexural deformities in which marked hoof wall deformity and remodeling of the distal phalanx have not occurred. Complete correction may take several days to weeks following surgery. If correction of the deformity is not sufficient following inferior check ligament desmotomy, the deep digital flexor tendon can be transected. Foals with severe flexural deformity usually require a deep digital flexor tenotomy (see Figure 48.3b). Tenotomy of the deep digital flexor tendon can be performed at the mid-metacarpus level or the mid-pastern level.12 We prefer tenotomy at the mid-metacarpal level because it is simple to perform and does not require entering a tendon sheath. Excessive scarring at the mid-metacarpal surgical site with a cosmetic blemish has been described, but in our experience this is a minor problem if the operated limb is effectively bandaged and exercise is limited to walking for 3 weeks after surgery. With severe flexural deformities correction of the deformed hoof wall may take months. Modeling changes of the distal phalanx will also occur during this time in response to the change of position and angle of the bone relative to the ground surface and weight-bearing forces. Severe chronic flexural deformities of the distal interphalangeal joint resulting in weight bearing on the dorsal hoof wall are poor candidates for surgery. Correction of the deformity often does not occur.
Metacarpophalangeal joint Acquired flexural deformities of the metacarpophalangeal joint occur most commonly in rapidly grow-
ing horses 10 to 18 months old and are often bilateral. The deep digital flexor tendon, the superficial digital flexor tendon, and the suspensory ligament all support the fetlock joint when it is in a normal weightbearing position. Treatment of these deformities can be challenging because of the multiple structures that are involved. Non-surgical treatment A balanced diet should be fed and heavy horses should have energy restriction. Analgesic should be administered to all horses to reduce pain, which may contribute to the deformity. Manual stretching of the limb with extension of the fetlock joints can be done several times per day. Corrective shoes should be applied with a 1–2-cm toe extension and 2–3-cm heel elevation. The elevated heel reduces tension on the deep digital flexor tendon, which allows the fetlock joint to assume a more normal position. Splints that force the fetlock joint into a position behind the center of the hoof will allow loading of the flexor tendons during weight bearing. Surgical treatment Surgical treatment is indicated for mild flexural deformities that fail to respond to non-surgical therapy after 2 weeks. Surgery should be considered immediately for moderate to severe flexural deformities. Selection of the best surgical procedure or combination of procedures to correct metacarpophalangeal flexural deformities can be difficult because more than one structure supporting the fetlock may prevent normal extension of the joint.5 For mild deformities the clinician should determine which flexor tendon (and associated check ligament) is the most taut by forcing
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Flexural Deformities the fetlock caudally during weight bearing and palpating the tendons above the joint. These mild flexural deformities usually respond to inferior or superior check ligament desmotomy. When both tendons are equally taut either the superior check ligament or both superior and inferior check ligaments are transected. With moderate flexural deformities of the metacarpophalangeal joint both the superior and inferior check ligaments should be transected followed by postoperative splint application. Severe flexural deformities will require desmotomy of both inferior and superior check ligaments, or inferior check ligament desmotomy and tenotomy of the superficial flexor tendon. Postoperative splint application will be necessary. Rarely, the suspensory ligament will need to be transected to place the fetlock in a position so flexor structures are loaded during weight bearing. The goal of surgery in all horses is to get the position of the fetlock behind the center of the hoof so flexor structures are loaded during weight bearing. All horses undergoing surgery should receive postoperative analgesics and corrective shoeing. The surgical procedures for correction of metacarpophalangeal flexural deformities have been described.12–14
Prognosis The prognosis for mild and moderate congenital flexural deformities is good to excellent if response to treatment is prompt and there is no predisposing bone or joint pathology. The prognosis for severe congenital deformities is guarded, particularly if sepsis or other health conditions become a factor during treatment. The prognosis for foals with mild to moderate acquired flexural limb deformities of distal interphalangeal joints that can be corrected with medical treatment or inferior check ligament desmotomy is excellent for soundness at all activities including racing.15 Horses undergoing deep digital flexor tenotomy will be sound for light or moderate riding activities. The prognosis for horses with mild to moderate flexural limb deformities of the metacarpophalangeal joint depends on response to therapy. Horses treated with inferior or superior check ligament desmotomy or both may be sound for moderate to strenuous activities. However, the flexural deformities of some of these horses will recur when they are placed into training or if they acquire other conditions that cause lameness. Horses with severe flexural limb deformi-
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ties of the metacarpophalangeal joint can be difficult to treat, and salvage for breeding or very light use is often the outcome.
References 1. Crowe MW, Swerczek TW. Equine congenital defects. Am J Vet Res 1985;46:353–8. 2. Buoen LC, Zhang TQ, Weber AF, Turner T, Bellamy J, Ruth GR. Arthrogryposis in the foal and its possible relation to autosomal trisomy. Equine Vet J 1997;29:60–2. 3. McIlwraith CW. Adam’s lameness in horses. In: Stashak TS (ed.) Diseases of Joints, Tendons, Ligaments and Related Structures. Philadelphia: Lippincott, Williams and Wilkins, 2002; pp. 459–640. 4. Hartzel DK, Arnoczky SP, Kilfoyle SJ, Stick JA. Myofibroblasts in the accessory ligament (distal check ligament) and deep digital flexor tendon of foals. Am J Vet Res 2001;62:823–7. 5. Adams SB, Santschi EM. Management of congenital and acquired flexural limb deformities. In: Proceedings of the 46th Annual Convention of the American Association of Equine Practitioners, 2000, pp. 117–25. 6. Lokai MD. Case selection for medical management of congenital flexural deformities in foals. Equine Pract 1992;14:23–5. 7. Madison JB, Garber JL, Rice B, Stumf AJ, Zimmer AE, Ott EA. Effect of oxytetracycline on metacarpophalangeal and distal interphalangeal joint angles in newborn foals. J Am Vet Med Assoc 1994;204:246–9. 8. Vivrette S, Cowgill LD, Pascoe J, Suter C, Becker T. Hemodialysis for treatment of oxytetracycline-induced acute renal failure in a neonatal foal. J Am Vet Med Assoc 1993;203:105–7. 9. Whitehair KJ, Adams SB, Toombs JP, Parker JE, Prostredny JM, Whitehair JG, Aiken SW. Arthrodesis for congenital flexural deformity of the metacarpophalangeal and metatarsophalangeal joints. Vet Surg 1992;21:228–33. 10. Auer JA. Flexural Deformities. In: Auer JA (ed) Equine Surgery, 1st edn Philadelphia: WB Saunders, 1992, pp 957–971. 11. Charman RE, Vasey JR. Surgical treatment of carpal flexural deformity in 72 horses. Aust Vet J 2008;86:195–9. 12. Auer JA. Flexural limb deformities. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St Louis: Saunders Elsevier, 2006; pp. 1150–65. 13. Adams SB, Fessler JF. Musculoskeletal system surgery. In: Atlas of Equine Surgery. Philadelphia: WB Saunders, 2000; pp. 325–93. 14. McIlwraith CW, Nixon AJ, Wright IM, Boening KJ. Desmotomy of the accessory ligament of the superficial flexor (proximal check ligament). In: Diagnostic and Surgical Arthroscopy in the Horse, 3rd edn. Philadelphia: Mosby Elsevier, 2005; pp. 386–8. 15. Stick JA, Nickels FA, Williams MA. Long-term effects of desmotomy of the accessory ligament of the deep digital flexor muscle in standardbreds: 23 cases (1979–1989). J Am Vet Med Assoc 1992;200:1131–2.
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Chapter 49
Fractures Commonly Seen in Foals Angus R. Adkins
Introduction Fractures in foals are a relatively common occurrence and are invariably due to a single misloading or traumatic insult rather than a sequel to exercise-induced subchondral bone changes often seen with fractures in adult performance and racehorses. Prolonged stall confinement inducing weakened tendons, osteoporosis, and management procedures that allow overcrowding or overexercise can increase the risk of fractures in foals. The common fracture configurations are different to those seen in adults, reflecting the different etiopathogenesis. Physeal fractures, described by the Salter–Harris classification system, are common due to the relative weakness of this structure.1 A breed predisposition to certain fracture configurations is not commonly seen. The clinical examination of a foal with a suspected fracture is no different to that of an adult; however, extra patience is often needed to allow appropriate interpretation of clinical findings. Most foals resist handling, which will often confuse the assessment of responses to stimuli. Excessive physical or chemical restraint will also confuse the interpretation of clinical findings and should be avoided unless injury to the handlers or foal is thought likely. Radiography is usually sufficient to establish a diagnosis of a fracture, although serial radiographic examinations (every 2–5 days) are sometimes required to confirm a diagnosis. Rarely is ultrasonography, nuclear scintigraphy, computed tomography, or magnetic resonance imaging needed to establish the diagnosis of a fracture, although ultrasonography may be an excellent diagnostic tool when first evaluating suspected fractures and radiographic equipment is not readily available. These modalities may be used to better assess the accompanying soft tissue
injury or fracture configuration. The most common differential diagnosis for fractures in foals is septic arthritis/osteomyelitis which can usually be differentiated by the relevant history, clinical findings, hematology, synovial fluid cytology and culture, and radiography. Fractures in foals generally have a better prognosis than those in adults, although this is still dependent on the fracture location, configuration, and accompanying complications. Conservative treatment is undertaken more commonly in foals due to their rapid rate of healing. Relatively short periods (2– 6 weeks) of stall confinement, or external coaptation (1–3 weeks) is often sufficient to allow adequate fracture healing. Conservative treatment regimes should be monitored and modified as foals are prone to flexural and angular limb deformities. Prolonged stall confinement and a lack of exercise may also interfere with normal skeletal and tendon development and predispose to developmental orthopedic disease.2
Fractures of the distal phalanx All types of previously described distal phalanx fractures can occur in foals, with solar margin fractures of the palmar and plantar processes that typically extend further into the body of the distal phalanx than typically seen in adults (type 7) being the most common3 (Figure 49.1). These fractures usually involve the forelimbs and often occur bilaterally. Foals with distal phalanx fractures usually present with acute lameness of moderate severity (grade 2–4/5). Hoof compressors induce a painful response particularly across the affected heel for type 7 fractures. Radiography readily identifies the fracture with a dorsoproximal 45◦ lateral-palmarodistal oblique projection
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 49.1 Dorsopalmar view of the foot in an 8-week-old foal. The arrow points to a type 7 fracture of the distal phalanx.
being the most useful. Healing type 7 fractures are often found on radiographic examinations as incidental findings, and diagnostic analgesia may be necessary to establish the site of pain in a lame foal. Treatment should include stall rest (2–6 weeks) and analgesics. An improvement in the severity of the lameness is usually seen within 7 days. External coaptation with glue-on shoes or hoof casts is not required for the successful healing of pedal bone fractures in foals, and is contraindicated due to the risk of inducing or potentiating clubfoot conformation. Unlike adults, articular (i.e., type 2 and 3) distal phalanx fractures should be expected to heal with a bony union but occasionally type 7 fractures will heal with a fibrous union. Soundness should be achieved well before bony union is evident radiographically and the decision to allow increasing levels of exercise should be determined by the foal’s degree of lameness, rather than serial radiographic examinations. The differential diagnosis for distal phalanx fractures in foals includes septic osteitis of the distal phalanx, infectious arthritis of the distal interphalangeal joint or navicular bursa, or subsolar abscessation.4 Subsolar abscessation without a solar crack or penetrating injury is rare in the foal and the possibility of a septic osteitis of the distal phalanx due to hematogenous spread must always be considered.4
Fractures of the pastern The most common fracture involving the pastern in foals is a physeal fracture of the proximal first phalanx. Subluxations of the proximal interphalageal joint, although not strictly a fracture, are occasionally seen.
Figure 49.2 Dorsopalmar view of the fetlock and pastern. There is a displaced Salter–Harris type 2 fracture of the proximal first phalanx. Note the relatively large metaphyseal component to the fracture.
Physeal fractures of the proximal first phalanx are usually seen in young foals (< 2 months) and can involve the fore- or hindlimb. These foals present with moderate-severe lameness that improves rapidly with external support and stall rest. If fracture displacement has occurred then an angular limb deviation, usually valgus, will be present. Radiography is diagnostic, although care must be taken to identify non-displaced fractures. These fractures usually have a Salter–Harris type 2 configuration, although the metaphyseal component can be very small (Figure 49.2). Prolonged external coaptation should be avoided to prevent distal limb hyperextension and 7 days in a distal limb cast followed by 2–3 weeks in a support bandage is usually sufficient for healing in most foals. The prognosis for future athletic use is thought to be good. Subluxation of the proximal interphalangeal joint may occur in foals due to an avulsion injury of the palmar supporting ligaments of the proximal second phalanx occasionally with avulsion fractures of the second phalanx. Foals with this injury usually present with severe lameness and axial malalignment of the pastern while weight bearing. Radiographic examination should be undertaken with some degree of weight bearing to appreciate the severity of the subluxation. In the author’s experience the prognosis for this injury with stall rest and any form of external coaptation is poor and the most appropriate
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Figure 49.3 Dorsomedial-palmarolateral view of the fetlock. There is a displaced basilar fracture of the medial proximal sesamoid bone.
treatment is arthrodesis of the proximal interphalangeal joint. However, the relative weakness of this fixation in soft foal bone, and the reluctance to provide prolonged external coaptation for fear of inducing a severe flexural deformity, ensures the prognosis with this arthrodesis is only fair.
Fractures of the proximal sesamoid bones Fractures of the proximal sesamoid bones (PSB) are common in foals, usually occurring within the first few months of life. They are often a consequence of overexercise following turnout from stall confinement. Uniaxial or biaxial fractures are seen and they commonly occur at the apex or base of the sesamoid (Figure 49.3). Most fractures involve one front limb, although occasionally multiple limb fractures, including the hind limbs, can occur. Clinical signs include varying degrees of lameness, synovial distension of the fetlock joint, and soft tissue swelling associated with the sesamoid. Most apical fractures present with mild lameness and most basilar fractures present with moderate-severe lameness. Occasionally healing apical fractures are found as an incidental finding on radiographic examination of the fetlock joint. Careful palpation of the PSB can often identify the fracture location by the location of pain. Standard radiographic projections of the fetlock joint are usually sufficient for a diagnosis. Radiographic examination of the contralateral limb, and possibly all limbs, should be considered for all PSB fractures. Careful evaluation of the anatomical loca-
Figure 49.4 Flexed lateromedial view of the fetlock. Note one sesamoid is longer in a proximal-distal direction following healing of a displaced fracture.
tion of the body of the sesamoid should be made as disruption of the intersesamoidian ligament or suspensory apparatus can occur. This is usually associated with a poor prognosis for athletic use. Treatment of PSB fractures is usually by conservative management, including stall rest (2–8 weeks), anti-inflammatory drugs and bandaging. Prolonged bandaging should be avoided as it invariably results in hyperextension of the fetlock joint while providing little benefit to healing. Most mildly displaced PSB fractures heal with a bony union, resulting in a relatively long sesamoid (Figure 49.4). Displaced fractures, which are usually basilar, often heal with a fibrous union. The surgical stabilization of PSB fractures in foals is not usually undertaken due to the lack of bone-holding power of the sesamoid. Occasionally displaced apical or basilar fractures can be removed to prevent either a fibrous union or the appearance of a ‘long’ sesamoid, if bony healing occurs. The prognosis for future athletic soundness depends on the number and location of the fractures and the presence of bony healing. Most apical fractures heal well and their prognosis is thought to be good.5 Most basilar fractures heal poorly and their prognosis for athletic use is only fair.6
Fractures of the carpus Most fractures of the carpus in foals are crushing injuries of the cuboidal bones as a consequence of delayed endochondral ossification. Ossification of the cuboidal bones occurs from day 260 of gestation and prematurity or dysmaturity due to placental
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Figure 49.6 Lateromedial view of the elbow. There is a mildly displaced type 1 fracture of the apophysis of the olecranon. In normal foals the caudal edge of the apophysis, pointed to by the arrow, should touch the line drawn along the caudoproximal edge of the olecranon.
Figure 49.5 Dorsolateral-palmaromedial view of a carpus. There is a sagittal fracture of the intermediate carpal bone at its articulation with the ulnar carpal bone. The ulnar carpal bone is mildly displaced in a lateral direction.
insufficiency can result in the neonate ambulating on soft cartilaginous cuboidal bones.7 Excessive weight bearing on these structures causes varying degrees of crushing, lameness, and osteoarthritis. Foals with crushing injuries present with varying degrees of lameness (grade 1–4/5), synovial distension of the carpal joints and a rapid deterioration, or the development of, an angular limb deviation. Most crushing injuries primarily involve the ulnar and fourth carpal bones and a valgus deviation results (Figure 49.5). Serial radiographs over 2–4 weeks are usually needed to fully appreciate the severity of the injury. Because of the poor prognosis with crushing injuries, prevention is important and screening radiographic or ultrasonographic evaluation should occur in all foals that are suspected of being premature, dysmature, or have obvious angular limb deviations early in life. Most foals will require 4–8 weeks of stall rest once crushing has occurred and weaning of these foals should be strongly considered to reduce accelerated weight gains associated with prolonged stall rest. In the author’s experience any form of external coaptation is contraindicated due to the likelihood of it causing flexural or angular limb deviations. Once a crushing injury has occurred the prognosis for fu-
ture athletic use is poor, although many animals can achieve paddock soundness.
Fractures of the olecranon Fractures of the olecranon are the most common upper limb fracture seen in foals. A number of common fracture configurations occur and classification systems have been described.8, 9 Type 1 fractures involve a physeal separation of the apophysis and are the most common type seen in neonates (Figure 49.6). In the author’s experience this is the most commonly misdiagnosed fracture in foals as it can be mildly displaced and easily overlooked. Type 1 fractures are usually caused by an acute overload with all other configurations usually caused by direct trauma. Foals with olecranon fractures usually present with a dropped elbow and some degree of carpal flexion. The severity of lameness can vary significantly between the different fracture types, with only minor lameness seen with some non-displaced, nonarticular fractures. Extensive soft tissue swelling may occur, usually with displaced or comminuted fractures. A medial-lateral radiographic projection with the limb extended cranially is the most useful for diagnosis. A cranial to caudal radiographic projection should also be evaluated to assess for concurrent radial fractures. The treatment of olecranon fractures should include an initial assessment for any breaks in the integrity of the skin and the extent of the soft tissue injury. Immediate treatment may include wound
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lavage and debridement, antibiotics, antiinflammatory drugs, and bandaging. A splint applied to the dorsal aspect of the carpus will encourage some weight bearing but splints are difficult to maintain in place, and should only be used as a temporary form of treatment, or during transportation. In foals with simple, mildly displaced olecranon fractures conservative treatment with box rest for 4–8 weeks is usually adequate, and the prognosis for athletic soundness is good.8, 10 Anti-inflammatory drugs can be given to encourage weight bearing and avoid acquired flexural or angular limb deformities. However, this will increase the risk of fracture displacement and should be undertaken with this consideration. If conservative treatment is undertaken then the degree of lameness should be carefully assessed daily with further radiographs taken if the lameness worsens, and/or fracture displacement is suspected. The degree of lameness in foals treated successfully with conservative treatment of mildly displaced fractures should improve considerably within 3–4 weeks. Foals with displaced olecranon fractures should undergo internal fixation as the prognosis with conservative treatment is poor. The olecranon is a highly suitable bone for internal fixation as the principles of internal fixation with a tension band can be employed.11 The placement of a bone plate and screws is the most common method of internal fixation used (Figure 49.7), although Steinman pins and tension band wires have also been reported to provide good
Figure 49.7 A postoperative view of the elbow following repair of a displaced type 1 olecranon fracture using a combination of a bone plate, screws, and wires.
results, and avoid some of the complications seen with plates and screws.12, 13 Other than the commonly seen complication associated with internal fixation there is a risk that elbow subluxation may occur secondarily to internal fixation of ulnar fractures. Occasionally during internal fixation of ulnar fractures the radius must be engaged with bone screws to provide a suitable fixation. In young foals with plenty of growth potential this will cause abnormal development of the elbow joint as the proximal radial physis continues to grow. To help avoid this complication it is generally recommended that in foals less than 5 months of age implants engaging the radius should be removed once fracture healing is adequate.14 The prognosis for surgical repair of olecranon fractures in foals is generally good for all commonly seen fracture configurations other than type 1 fractures.8, 9, 15, 16 These fractures have a poor prognosis for survival due to the difficulty in achieving rigid internal fixation without post-operative failure of the fixation or elbow subluxation17 (Figure 49.8).
Fractures of the tarsus As with the carpus, most fractures of the tarsus in foals are crushing injuries of the cuboidal bones as a consequence of delayed endochondral ossification. These foals usually present with varying degrees of lameness but commonly ‘bunny hop’ when forced to move at a gait faster than a walk. Pain can often be induced with digital pressure over the dorsal aspect of the distal tarsus. The tarsus usually has a sickle or
Figure 49.8 Failure of the repair in Figure 49.7 due to a sagittal fracture of the apophysis. This is the most common method of failure following repair of type 1 fractures. The arrows point to the sections of split apophysis.
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Figure 49.9 Dorsomedial-plantarolateral view of the hock. There is ‘pinching’ of the third tarsal bone due to crushing of the distal row of tarsal bones secondarily to delayed endochondral ossification.
cow-hocked appearance, and if collapse of the proximal row of tarsal bones occurs then synovial distension of the tarsocrural joint can result. On radiographic examination a varying degree of collapse can occur, from mild wedging of the distal row of tarsal bones to displaced frontal fractures of either row of tarsal bones. Tarsal crushing injuries are treated in the same way as carpal crushing injuries, although the prognosis is better. It has been reported that foals with < 30% collapse of the tarsal bones have a good prognosis for intended use, but if > 30% collapse of affected bones with pinching or fragmentation occurs (Figure 49.9), then the prognosis for intended use is poor.18 Prevention of these injuries is important and screening radiographic or ultrasonographic examination should be undertaken on all foals suspected of having incomplete ossification. Another type of tarsal fracture commonly seen in foals is a complete, displaced fracture of the distal lateral trochlear ridge of the talus (Figure 49.10). These fractures are traumatic in origin, usually caused by a kick from another horse. A superficial skin abrasion or contusion is usually present at the lateral aspect of the tarsus. These foals usually present with only mild lameness but obvious synovial distension of the tarsocrural joint. On radiographic examination a large fragment is usually seen, involving up to 50% of the dorsal aspect of the lateral trochlear ridge. Removal of the fragment is recommended, usually via an arthrotomy, and the prognosis for future athletic use is thought to be good.
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Figure 49.10 Lateromedial view of the hock with a displaced fracture of the distal lateral trochlear ridge.
Long bone fractures Fractures of long bones usually occur due to direct trauma or misloading of a limb during exercise or getting up. Foals with long bone fractures usually present with acute, severe lameness and obvious soft tissue swelling. Instability and crepitus can often be palpated, although humeral and femoral fractures can be difficult to diagnose due to extensive soft tissue swelling and relative fracture stabilization. Radiography is necessary to establish a definitive diagnosis and determine the fracture configuration, although ultrasonography can be useful in the field to identify the presence of many long bone fractures. Many long bone fractures involve a physis with a Salter–Harris type 2 configuration being the most common (Figure 49.11). Diaphyseal fractures in foals often have a simple oblique configuration rather than being comminuted fractures that are commonly seen in adults (Figure 49.12). Stabilization of long bone fractures prior to transportation can be difficult due to the foal’s temperament and its response to any painful stimuli. However an attempt should be made to stabilize fractures distal to the elbow or stifle. In general ‘open’ fractures have a poor prognosis for survival and every attempt should be made to prevent this complication. Heavy sedation and lateral recumbency should also be considered for transportation. It can be surprising how many long bone fractures in foals can be treated successfully with the
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Figure 49.13 Caudocranial view of the tibial fracture seen in Figure 49.11 following 6 weeks of stall rest. Note the extent of bone healing achieved after this short time period. Figure 49.11 Caudocranial view of the stifle. There is a mildly displaced Salter–Harris type 2 fracture of the proximal tibia. The arrow points to the metaphyseal component of the fracture.
metacarpus, metatarsus, radius, and tibia being the most amenable to surgical repair. It is always worth consulting a surgeon before undertaking euthanasia of a foal with a long bone fracture. In general, displaced fractures require surgical fixation, although some displaced humeral and femoral
fractures can heal with prolonged stall rest to allow paddock soundness.19, 20 Furthermore, some mildly displaced physeal fractures can heal with conservative treatment (Figure 49.13). The risk of fracture displacement, and the development of angular or flexural deviations due to prolonged lameness, needs to be considered in the decision to undertake conservative treatment. The surgical treatment of long bone fractures in foals is made possible by the reduced weight of the patient, although it is sometimes made more difficult by complications associated with temperament, angular and flexural deviations due to prolonged external support or lameness, and other forms of orthopedic disease associated with prolonged stall rest. The risk of implant sepsis is high with the repair of any long bone fracture and its occurrence invariably results in failure of the fixation.21
Fractures of the pelvis
Figure 49.12 Lateromedial view of the humerus. There is a displaced, oblique, diaphyseal fracture.
Pelvic fractures in foals are usually the result of a sudden fall or collision, with the most common sites being the tuber coxae, wing of the ilium, and acetabulum. Occasionally Salter–Harris type 1 fractures of the femoral head or dislocations of the coxofemoral joint can occur. The clinical signs of lameness will depend on the fracture location and can vary from mild, shortterm lameness with some tuber coxae fractures to severe permanent lameness with displaced fractures involving the coxofemoral joint. Deep digital palpation will often elicit a painful response at the fracture site and crepitus can often be felt. Diagnostic radiographs can be taken with heavy sedation using lateral oblique and ventral oblique projections.22 Diagnostic
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Fractures Commonly Seen in Foals ultrasonography is a very useful technique for fracture identification, although a thorough understanding of the fracture configuration is difficult to achieve for all but the most experienced ultrasonographer. Rarely is scintigraphy needed to identify a pelvic fracture in foals. Treatment for pelvic fractures includes stall rest, with most non-articular fractures requiring 4–8 weeks rest and acetabular fractures up to 10 weeks rest. The judicious use of anti-inflammatory drugs should be considered to encourage some degree of weight bearing, and enable the foal to get up and down with less discomfort, while avoiding further propagation of the fracture. Most foals do not tolerate cross-tying, and this is contraindicated in most cases. Surgical repair of pelvic fractures or coxofemoral luxation is not considered a viable option for any horse other than miniatures. The prognosis for non-articular pelvic fractures in foals is generally good for athletic use; however, some degree of permanent pelvic asymmetry is common. The prognosis for articular fractures is generally thought to be poor, although some mildly or nondisplaced fractures can heal to provide paddock or even athletic soundness.23
Fractures of the vertebrae Most fractures of the vertebrae involve the cervical vertebrae, although occasionally coccygeal fractures or luxations can occur. Cervical fractures usually occur during falling or pulling back when being handled, or by colliding with solid objects. Foals with cervical fractures can present with clinical signs varying from neck pain or stiffness, a reluctance to lift their head, total tetraparesis, or even sudden death. Severe neck pain may prevent a foal nursing normally and can cause rapid dehydration. There is usually asymmetry of the neck with axial malalignment. Occasionally crepitus can be felt during movement. Foals can present with varying degrees of ataxia which should not be confused with reluctant or awkward movements due to neck pain. At the time of fracture some foals will remain recumbent and non-reactive to stimuli, and care must be taken to allow recovery before euthanasia is undertaken. Diagnosis is confirmed by plain radiography with lateral projections usually being sufficient. The interpretation of cervical radiographs in foals can be difficult due to the numerous physes and separate centres of ossification. The most common vertebral fracture seen is a physeal fracture at the dens of the axis (Figure 49.14). The dens usually remains firmly attached to the atlas with the axis becoming displaced ventrally due to the tension within the nuchal liga-
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Figure 49.14 Lateromedial view of the cranial neck. There is a displaced fracture of the dens of the axis.
ment. There is a relatively large space between the atlas and axis yet it can be surprising how little ataxia can result even with obvious fracture displacement. Treatment of cervical fractures involves prolonged stall rest (4–8 weeks) and the administration of analgesics, anti-inflammatory drugs, and supportive care. The degree of any ataxia should be expected to improve quickly if permanent resolution is to be achieved. However, as healing occurs, ataxia may redevelop due to impingement of callus on the spinal cord. Surgical treatment has been described, although it is rarely undertaken due to the potential for complications, and the associated poor prognosis.24 The prognosis for future athletic use is generally poor, although some horses can become pasture sound with varying degrees of neck stiffness and ataxia.
Fractures of the head Fractures of the head include fractures of the skull and sinuses, maxilla or mandible, and the periorbital region. Virtually all fractures are the result of direct trauma, usually from kicks, collisions, or falls. Fractures of the mandible include fractures of the incisors, interdental space, or body of the mandible. In neonates the most common fracture is a complete, displaced, compound fracture at the interdental space. These foals usually present with drooling of milk or saliva and swelling of the mandible. Surgical repair is usually advisable as these fractures are unstable with severe malocclusion, and the foals are reluctant to nurse. A number of methods have been described to repair these fractures, with simple crosspinning of the mandible being the easiest and most effective in the author’s experience (Figure 49.15). These fractures heal quickly, and 3–4 weeks of stabilisation is all that is usually required. Fractures of the
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Foal: Musculoskeletal System Fractures of the calvaria are often the result of falls or trauma during handling. The presenting clinical signs will vary considerably depending on the location and magnitude of the injury. These signs may include sudden death with severe injuries of the brain parenchyma, or a permanent head tilt with basilar skull fractures. The specific fractures, their treatment, and prognosis are beyond the scope of this chapter and are well described elsewhere.25 However, in general terms, it is the author’s experience that if clinical signs associated with calvaria fractures do not resolve quickly (i.e., within 7 days) then complete resolution is unlikely.
Fractures of the ribs
Figure 49.15 A postoperative distoproximal view of the jaw following cross-pinning with Steinman pins of a displaced fracture of the mandible at the interdental space.
body of the mandible rarely require internal fixation in foals, with the criteria for surgical repair including bilateral unstable fractures, obvious dental malocclusion, or if the foal is reluctant to nurse for prolonged periods due to pain from fracture instability. Fractures of the periorbital region in foals are often the result of being kicked by another horse. These are usually severe injuries as the thin, soft bones in this region and the protruding orbit are often severely traumatized. The extent of trauma to the orbit often dictates the specifics of treatment as rupture of the globe or severe intraocular trauma causing permanent blindness is common. There is usually extensive soft tissue swelling associated with these injuries and this should be treated before fracture stabilization is considered. A delay of 2– 4 days is not usually detrimental to surgical repair and will often make the surgical approach less invasive. Immediate surgical stabilization should be undertaken if fracture displacement has the potential to damage the globe. Surgical stabilization techniques for periorbital fractures are described elsewhere, however, the author prefers the use of monofilament absorbable suture material rather than cerclage wire for stabilization of these fractures. Fractures of the skull and sinuses occur occasionally in foals with the nasal, frontal, and maxillary bones being the most commonly involved. Surgical repair of depression fractures is usually advisable to prevent sequestrum formation, dyspnea, cosmetic blemishes, and secondary sinus diseases.
Thoracic trauma, including rib fractures or costochondral dislocation, has been reported to occur in 21% of foals aged < 3 days on a large Thoroughbred stud farm in one report26 and 69% of foals admitted to a neonatal intensive care unit in another report.27 Rib fractures can be very difficult to identify clinically with up to 75% of foals with normal thoracic symmetry having rib fractures.27 However, most foals with rib fractures causing clinical disease present with varying signs of tachypnea, dyspnea, thoracic swelling or asymmetry, or sudden death. Thoracic ultrasonography is a very useful diagnostic technique with a recent report27 identifying thoracic ultrasonography as being four times more likely to identify fractured ribs than radiography (Figure 49.16). Most fractures involve ribs 2–7 and they commonly occur
Figure 49.16 An ultrasound image of a displaced rib fracture in a neonate. Pockets of fluid can be seen between the skin and ribs.
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Fractures Commonly Seen in Foals at, or immediately proximal to, the costochondral junction. Rib fractures are thought to occur during parturition. Most ribs fractures are moderately displaced with the literature being contradictory as to the common orientation of the displacement.27, 28 Although a common finding in foals, the literature is unclear, and the author unsure, as to the best approach for treatment of this type of fracture. In most foals, conservative regimes of stall rest and supportive treatment, including analgesics, anti-inflammatory drugs, antibiotics, sedation, and intranasal oxygen, is successful. However, hemothorax, pneumothorax, diaphragmatic hernia, and sudden death can result. It has been reported that 25%28 and 28%29 of foals with rib fractures died or underwent euthanasia as result of the fractures. Pulmonary contusion and thoracic pain can also result in respiratory insufficiency and predispose to pneumonia. As such, a number of surgical techniques have been attempted to prevent these complications, including internal fixation with reconstruction plates, and a nylon strand suture repair technique.30, 31 The author prefers the use of 4-hole reconstruction plates, 3.5 mm cortical bones screws, and cerclage securing of the plate with heavy absorbable monofilament material. The criteria for when surgical intervention is required are unclear but surgery is generally reported to be advisable when intrathoracic trauma is thought to be highly likely.
References 1. Embertson RM, Bramlage LR, Herring DS, Gabel AA. Physeal fractures in the horse: classification and incidence. Vet Surg 1986;15:223–9. 2. Barneveld A, van Weeren PR. Conclusions regarding the influence of exercise on the development of the equine musculoskeletal system with special reference to osteochondrosis. Equine Vet J Suppl 1999;31:112–19. 3. Yovich JV. Fractures of the distal phalanx in the horse. Vet Clin N Am: Equine Pract 1989;5(1) 145–59. 4. Neil KM, Axon JE, Todhunter PG, Adams PL, Caron JP, Adkins AR. Septic osteitis of the distal phalanx in foals: 22 cases (1995–2002). J Am Vet Med Assoc 2007;230(11): 1683–90. 5. Schnabel LV, Bramlage LR, Mohammed HO, Emberton RM, Ruggles AJ, Hopper SA. Racing performance after arthroscopic removal of apical sesamoid fracture fragments in Thoroughbred horses age < 2 years: 151 cases (1989–2002). Equine Vet J 2007;39(1):64–8. 6. Southwood LL, McIlwraith CW. Arthroscopic removal of fracture fragments involving a portion of the base of the proximal sesamoid bone in horses: 26 case (1984–1997). J Am Vet Med Assoc 2000;217(2):236–40. 7. Auer JA. Zur intrauterinen Ossifikation der Karpal-und Tarsalknochen beim Fohlen und Behandlung von Ossificationsstorungen. Pferdeheilkunde 1986;2:35.
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8. Donecker JM, Bramlage LR, Gabel AA. Retrospective analysis of 29 fractures of the olecranon process of the equine ulnar. J Am Vet Med Assoc 1984;185(2):183–9. 9. Denny HR, Barr ARS, Waterman A. Surgical treatment of fractures of the olecranon in the horse; A comparative review of 25 cases. Equine Vet J 1987;19(4):319–25. 10. Wilson DG, Riedesel E. Nonsurgical management of ulnar fractures in the horse: A retrospective study of 43 cases. Vet Surg 1985;14(4):283–6. 11. Watkins JP. Radius and Ulner. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St Louis: Saunders, 2006;1269– 70. 12. Martin F, Richardson DW, Nunamaker DM, Ross MW, Orsini JA. Use of tension band wires in horses with fractures of the ulnar: 22 cases (1980–1992). J Am Vet Med Assoc 1995;207(8):1085–9. 13. Hanson PD, Hartwig H, Markel MD. Comparison of three methods of ulnar fixation in horses. Vet Surgery 1997;26:165–71. 14. McIlwraith CW, Robertson JT. Plate fixation of a simple fracture of the olecranon. In: McIlwraith CW, Turner AS (eds) Equine Surgery Advanced Techniques. Baltimore: Williams & Wilkins, 1998; pp. 96–102. 15. Swor TM, Watkins JP, Bahr A. Results of plate fixation of type 1b olecranon fractures in 24 horses. Equine Vet J 2003;35:670. 16. Janicek JC, Rodgerson DH, Hunt RJ, Spirito MA, Thorpe PE, Tessman RK. Racing prognosis of horses following surgically repaired olecranon fractures. Can Vet J 2006;47(3):241–5. 17. Clem MF, DeBowes RM, Douglass JP, Leipold HW, Chalman JA. The effects of fixation of the ulna to the radius in young foals. Vet Surg 1988;17(6):338–45. 18. Dutton DM, Watkins JP, Walker MA, Honnas CM. Incomplete ossification of the tarsal bone in foals: 22 cases (1988–1996). J Am Vet Med Assoc 1998;213(11):1590–4. 19. Zamos D, Parks A. Comparison of surgical and nonsurgical treatment of humeral fractures in foals: 22 cases (1980–1989). J Am Vet Med Assoc 1992;201:114. 20. McCann ME, Hunt RJ. Conservative management of femoral diaphyseal fractures in four foals. Cornell Vet 1993;83(2):125–32. 21. Hance SR, Bramlage LR, Schneider RK, Embertson RM. Retrospective study of 38 cases of femur fractures in horses less than one year of age. Equine Vet J 1992;24(5):357–63. 22. Barrett EL, Talbot AM, Diver AJ, Barr FJ, Barr AR. A technique for pelvic radiography in the standing horse. Equine Vet J 2006;38(3):266–70. 23. Rutkowski JA, Richardson DW. A retrospective study of 100 pelvic fractures in horses. Equine Vet J 1989;21(4): 256–9. 24. Nixon AJ (ed.). Equine Fracture Repair. Philadelphia: Saunders, 1996; pp. 300–1. 25. DeBowes, R.M. Fractures of the Cranium In: Nixon AJ (ed.) Equine Fracture Repair. Philadelphia: Saunders, 1996; pp. 313–22. 26. Jean D, Laverty S, Halley J, Hannigan D, Leveille R. Thoracic trauma in newborn foals. Equine Vet J 1999;31(2):149– 52. 27. Jean D, Picandet V, Macieira G, Beauregard G, D’Anjou MA, Beauchamp G. Detection of rib trauma in newborn foals in an equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 2007;39(2):158–63.
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28. Sprayberry KA, Bain FT, Seahorn TL, Slovis NM, Byars TB. 56 cases of rib fractures in newborn foals hospitalized in a referral center intensive care unit from 1997–2001. Proc Am Assoc Equine Practnrs 2001;47:395–9. 29. Schambourg KA, Laverty S, Mullim S, Fogarty UM, Halley J. Thoracic trauma in foals: post mortem findings. Equine Vet J 2003;35:78–81.
30. Bellezzo F, Hunt RJ, Provost R, Bain FT, Kirker-Head C. Surgical repair of rib fractures in 14 neonatal foals: case selection, surgical technique and results. Equine Vet J 2004;36(7):557–62. 31. Kraus, BM, Richardson DW, Sheridan G, Wilkins PA. Multiple rib fracture in a neonatal foal using a nylon strand suture repair technique. Vet Surgery 2005;34(4):399–404.
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Chapter 50
Infectious Arthritis and Osteomyelitis Fairfield T. Bain
Infectious arthritis remains a significant medical problem in the neonatal and young foal. In neonatal foals, it is thought to be at least one sequel to inadequate intake of colostrum and failure of passive transfer of immunoglobulins, but can occur in foals in the absence of failure of passive transfer. The age range of affected foals can vary from shortly after birth to a few weeks of age.
Clinical signs In younger foals with bacteremia, acute synovial distension is often the first observed clinical sign. Older foals may present with more varied clinical signs which more often include lameness with variable synovial distension. Early diagnosis requires close clinical observation for lameness or periarticular swelling. Palpation of the limbs for increased skin temperature, pain, or swelling may also be helpful, and should be part of the physical examination of any neonatal foal. Synovial distension is often easier to detect in more distal joints such as the stifle, carpus, tarsus, or fetlock joints. Deep palpation of the shoulder, hip, or elbow may elicit a pain response when these joints are affected. Infectious arthritis should remain the primary differential diagnosis for any young foal with acute lameness, until ruled out by physical examination, diagnostic imaging, and hematological evaluation. Sites of origin of bacteremia may include the umbilicus (classic ‘navel and joint ill’), the respiratory or gastrointestinal tract, or other sites such as skin wounds adjacent to the joint. Traumatic injury may also occur and may be a reason for direct infection into the joint space. In the author’s experience, localized infection of the umbilicus is not as common as it has been historically, most likely due to improved care of the umbilicus
by owners and farm personnel. With septic arthritis or osteomyelitis in foals, the disease process can follow a variable clinical course, with some presenting in a rapidly deteriorating condition with multiple joint involvement, and others demonstrating a smoldering lameness with minimal synovial or periarticular swelling. In well-managed equine facilities, close observation by management or farm personnel and early recognition can allow foals to be treated earlier, with improved outcomes.
Mechanism of infection Infectious arthritis can vary from a primary inflammatory process within the synovial space to one that involves both the synovial space and medullary space of adjacent bone. In both, the mechanism of infection is thought to be seeding of capillary vessels secondary to bacteremia. Hematogenous origin of bacteria can involve multiple areas from the synovium to the epiphyseal, physeal, or metaphyseal regions of bones. The categories described by Firth remain the most common practical means of describing the affected regions for septic arthritis in foals.1 The three general categories described include Synovial (S-type), those located solely within the synovial space; Epiphyseal (E-type), those involving the bone underlying the articular cartilage; and Physeal (P-type), those involving the metaphysis and growth plate. Figure 50.1 demonstrates severe metaphyseal and physeal necrosis subsequent to septic osteomyelitis. These categories are useful descriptors of the site of infection; multiple sites of involvement can occur simultaneously within a particular individual patient. Radiographic description of alterations allow for a more accurate means of communication between professional personnel.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 50.2 Radiograph of metacarpophalangeal joint with osteolytic change present within the physis.
Figure 50.1 Osteomyelitis – P-type. The pale tan regions in the metaphysis and physis of the distal tibia are consistent with necrosis of bone. A pathological physeal fracture is also present. Image courtesy of Dr Joe Mayhew.
Diagnostic evaluation Clinical diagnosis of septic arthritis involves observation as well as palpation of the limbs, as described earlier. Clinical laboratory data such as white blood cell count and fibrinogen concentrations are often used in the initial assessment for confirmation of an inflammatory process as well as to provide a baseline for serial monitoring of the clinical response as the treatment progresses. Beyond clinical recognition, further information is required to determine whether there is significant bone involvement. Radiographs are considered standard in evaluation of the affected joint. Radiographic alterations may lag behind the actual clinical disease process, with a delay in appearance of osteolysis, so serial evaluation is often indicated to assess the extent of bone involvement. Radiographic evidence of bony lysis could indicate a worse prognosis and may suggest the need for more invasive therapy (Figure 50.2). Ultrasonographic imaging may also be useful in evaluating the soft tissues of the joint space (synovial lining and contents of the synovial space, adjacent ligamentous or tendinous structures and their associated tendon sheaths), and may demonstrate alterations along the physis in areas of osteolysis. Normal joints are observed to have variable amounts of anechoic fluid with a thin layer of hypoechoic synonial membrane. Infected synovial spaces may have obvious distension of the synovial space with increased volume of synovial fluid of varying echogenicity, depending on the cellular content or presence of fib-
rin deposits. Because of this, ultrasonography may be useful in characterizing the content of the synovial fluid in joints where it appears to be difficult to obtain a good egress of fluid during lavage. In joints where distinct synovial distension is palpable, synoviocentesis for synovial fluid analysis and bacterial culture, followed by therapeutic joint lavage under general anesthesia is the most appropriate first step (Figure 50.3). Repeat joint lavage may be performed, based on recurrence of joint swelling and lameness. In some patients, there may be significant periarticular soft tissue swelling that may obscure definition of the synovial pouch or make it difficult to distinctly palpate the joint. Ultrasonographic imaging can occasionally be used in these cases to differentiate between periarticular soft tissue swelling and synovial distension prior to synoviocentesis. Synovial fluid analysis should include a total nucleated cell count, total protein concentration, and
Figure 50.3 Joint lavage of the tarsus. An ingress needle is attached to an extension set to administer the sterile fluid. An egress needle is present on the opposite side of the joint. Photograph courtesy of Dr Reese Hand.
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Infectious Arthritis and Osteomyelitis microscopic examination of a smear (direct or concentrated, if necessary) to search for primary evidence of microorganisms or secondary evidence by the presence of degenerate neutrophils. Total nucleated cell counts in synovial fluid is generally less than 5000 nucleated cells per µL, with septic joints commonly having total cell counts of greater than 30 000 cells per µL, but these can be quite variable depending on the severity of the inflammatory process and whether the inflammatory process is adjacent to rather than within the synovium. Total protein concentrations are generally elevated in infected joints, with normal values being less than 2.5 g/dL. Septic synovial fluid often has a total protein concentration greater than 3.5 g/dL. Collection and incubation of synovial fluid in blood culture media can enhance the potential of obtaining an organism.2 In some situations where bone lysis within the metaphysis or adjacent to the physis is apparent on radiographs, direct aspiration with a large gauge needle (14–16 G) may be useful in obtaining inflammatory exudate for bacterial culture and cytological examination. A spectrum of both Gram-positive and Gramnegative pathogens may be identified as the causative agents for septic arthritis in foals. Salmonella spp. are noted for their ability to more commonly result in osteomyelitis and destruction of bone, but this process can also occur with other organisms. Because of the variety of microorganisms, it is important to obtain a representative bacterial culture as early as possible in the disease process in order to select the best possible antimicrobial regimen in an effort to reduce potential tissue destruction and shorten the treatment time. The yield of positive culture results from synovial fluid or intraosseous exudates can vary depending on whether prior treatment with antimicrobial agents has been instituted before diagnostics are performed. Obtaining an accurate culture in cases where there is primary bone involvement on radiographs can be a challenge. In some patients, there may be a high suspicion of septic osteomyelitis, yet no primary evidence may be found on imaging. In these cases, pursuing a course of antimicrobial agents and anti-inflammatory medications with periodic reassessment may be a reasonable course. In younger foals with additional signs of systemic infection, collection of a blood culture may also be worthwhile as an additional aid in obtaining a positive culture to document the primary infectious agent. Ultrasonographic evaluation of the umbilical structures is often recommended for foals with septic arthritis in order to determine if infection of the internal umbilical structures is a nidus for bacteremia and seeding the synovial structures or bone. Should ultrasonographic evidence of infec-
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tion of the umbilical structures be found, a debate may be had as to whether surgical removal should be carried out to avoid further bacteremia. The author’s experience suggests that this may not be as common a source as it has been historically, probably due to improved management and owner education regarding umbilical care for neonatal foals in the past couple of decades. While not common in younger foals and neonates, bacterial endocarditis can occur in older foals and be a source of hematogenous bacterial emboli that result in septic arthritis in one or more joints.
Treatment The goal of therapeutic intervention in foals with septic arthritis is to control the infectious process (antimicrobial agents), remove inflammatory material from the synovial space to prevent or protect against injury to the articular cartilage (joint lavage), and provide pain relief (analgesics/anti-inflammatory medications). Another goal in cases of osteomyelitis should include minimizing destruction of bone. In many situations, broad-spectrum antimicrobial agents may be used to treat suspected sepsis or foals where a positive bacterial culture could not be obtained. Where the opportunity exists, it is recommended that diagnostic efforts be made early to obtain a culture result so that finances and time are not wasted using inappropriate antimicrobial agents. In many circumstances, a combination of both systemic and local administration of antimicrobial agents will be used for affected foals. For systemic administration of antimicrobial drugs, the intravenous route is preferred over oral or intramuscular routes for the ability to maximize the therapeutic concentration of the antimicrobial agent within the infected tissues. Local direct injection or regional perfusion for selected cases will provide for much higher local tissue levels at specific sites of infection. Selection of the agent or combination should be made based on the culture results, but in cases where no culture has been obtained, or in the interim while awaiting culture results, a broad-spectrum combination is recommended. Alterations in the regimen can be made at a later time, based on the culture results as well as the clinical response. The most commonly used initial regimen would be a beta-lactam (potassium or sodium penicillin or cefazolin) and an aminoglycoside (gentocin or amikacin). Other antimicrobial agents that might be considered include oxytetracycline, trimethoprim-sulfa combinations, and ceftiofur, based on their spectrum of antimicrobial activity. Later in the clinical course, longer-term therapy with oral antimicrobial agents (single or in
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combination) might be considered based on information obtained on the sensitivity patterns. Enrofloxacin and possibly other fluoroquinolones are considered contraindicated in neonatal foals due to concerns regarding research findings demonstrating cartilage damage in neonates.3 The use of more advanced antimicrobial agents such as third- or fourth-generation cephalosporins (e.g., ceftazidime, cefquinome) or imipenem-cilastin should be based on specific indications from the results of bacterial culture and sensitivity. Analgesic and anti-inflammatory medications may be useful, especially in the early stage of the treatment when there may be significant pain and lameness. Non-steroidal anti-inflammatory medications such as phenylbutazone (4.4–8.8 mg/kg once daily, orally or intravenously), flunixin meglumine 1.1 mg/kg once or twice daily, orally or intravenously), and ketoprofen 2.2 mg/kg two or three times daily, orally or intravenously) have been used for foals with septic arthritis in the author’s practice. Concern regarding gastric ulceration in foals when using these medications for a longer period of time has resulted in the common concurrent use of antiulcer medications such as ranitidine (6.6 mg/kg, p.o., q 8 h) or omeprazole (1–4 mg/kg, p.o., q 24 h) in an effort to reduce clinical signs of gastric ulceration. Additional pain control techniques may be necessary in some individuals, depending on the severity of disease, location of the site of septic arthritis, and degree of pain and weight-bearing. Local therapy varies depending on the degree of involvement and site of infection. Synovial type infections are commonly treated with joint lavage on an as-needed basis, with instillation of an antimicrobrial agent such as amikacin into the joint space following the lavage. In other situations where there may be infection of adjacent soft tissues or bone, additional techniques such as regional perfusion might be considered.2 Regional perfusion with concentration-dependent antimicrobial agents such as one of the aminoglycosides (gentocin, amikacin) has become a common technique for regional delivery of an antimicrobial in high concentrations within the affected tissues. High regional tissue concentrations of concentration-dependent antimicrobials such as the aminoglycosides can result in a high peak concentration : MIC ratio (Cmax : MIC ratio).4 High peak antimicrobial concentrations achieved in the synovial fluid, soft tissues, and bone may allow for a prolonged post-administration effect.5 This allows for a schedule of administration once daily or every two or three days. Antimicrobial selection should be guided by the results of bacterial culture and susceptibility testing; however, amikacin is often selected as the initial agent as it appears to have the great-
est efficacy against common equine pathogens involved with septic arthritis or osteomyelitis.5 Most commonly, regional limb perfusion with an antimicrobial agent is delivered via a small 23-gauge butterfly catheter inserted into a superficial skin vein proximal to the affected site with an occlusive tourniquet (usually elastic rubber tubing or elastic bandage over rolled gauze) placed higher on the limb. This may be performed under general anesthesia or using heavy sedation in the standing animal. An occlusive period of approximately 30 minutes is commonly used. There are no documented guidelines as to dose of drug and volume of perfusate to use in neonatal foals. Common clinical usage of one-third the total systemic dose of amikacin has been suggested, or 50 mg amikacin.5 Perfusate volumes of 10 to 30 mL or 0.1 mL/kg have also been suggested for foals.5 Direct intraosseous injection of the antimicrobial agent has also been performed for similar reasons. Occasionally, the intraosseous injection may be guided by radiographic or ultrasonographic imaging to determine the site of bone lysis where direct injection would be most appropriate. Direct intra-articular injection of an antimicrobial agent has been shown to achieve higher concentrations than with regional perfusion or intraosseous injection.6 Regional perfusion or intraosseous injection are more likely to be considered when penetration of adjacent bone or periarticular soft tissue might be necessary. Therapeutic lavage of the joint space with a balanced electrolyte solution is commonly performed in foals with septic arthritis in an effort to remove inflammatory debris and reduce injury to the articular cartilage. Large-volume lavage using large-bore cannulas is preferable in some cases, especially where fibrin or clumped debris might be present. Surgical debridement (curettage) of necrotic bone tissue along the physis has also been advocated (Figure 50.4a,b). This process would remove much of the necrotic, infected bone with the idea of speeding the resolution of the lesion and removing infectious and inflammatory debris that might prolong the clinical course. Structural integrity of the involved region (how much physeal structure could safely be removed) would have to be considered when using this procedure. Hyperbaric oxygen therapy has been used by the author and others as an adjunct to antimicrobial therapy.7 Hyperbaric oxygen has been approved for used in refractory osteomyelitis in human patients, based on the concepts that it enhances neovascularization of necrotic, poorly perfused bone, enhances delivery of antimicrobial agents into infected bone, increases neutrophil killing ability in infected bone, and raises intramedullary partial pressures of oxygen in infected bone.8–10 While no controlled trials have
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Figure 50.4 (a) Radiograph showing osteolysis along the proximal physis of the proximal phalanx (P1). Photograph courtesy of Angus McKinnon. (b) Intraoperative radiograph showing bone curette in place. Photograph courtesy of Angus McKinnon.
been performed on equine patients, anecdotal experience suggests that it is helpful in salvaging regions of necrotic bone (as determined by osteolytic change on radiographs). For most foals, this treatment requires a hyperbaric chamber custom made for a larger equine patient. In some circumstances, smaller foals might be treated in a human monoplace chamber with appropriate sedation, since they would remain recumbent. Protocols generally involve treatment of the foal at 2.0 to 2.5 atmospheres absolute (ATA) or 14.7 to 22 psi once daily for 45–60 minutes. Hyperbaric oxygen treatments are often provided for 10 days or more, dependent upon the clinical and radiographic evaluation of bone lysis. This procedure is more often reserved for foals who may fail to respond to initial medical efforts, or those with more advanced bone destruction evident on initial radiographic evaluation. It is important to adequately document radiographic alterations at the outset for proper patient selection. Those animals with significant physeal disruption, pathological fractures, or tarsal collapse are not considered appropriate patients for hyperbaric oxygen treatment.
Monitoring and prognosis Clinical improvement is generally determined by a decrease in lameness as well as a decrease in synovial distension or periarticular swelling. Resolution of fever and leukocytosis and hyperfibrinogenemia are also indicators of decreasing inflammatory process. Radiographs are considered the standard for serial monitoring of the patient with osteomyelitis. A determination of when clinical resolution is adequate to discontinue treatment is usually made from a combination of resolution of lytic bone change on the radiograph as well as resolution of clinical signs such as lameness and synovial distension. In many cases, the total duration of treatment time can be in the range of 2–4 weeks or longer. The prognosis for foals surviving septic arthritis or osteomyelitis remains unclear. At least one study has suggested that Thoroughbred foals surviving septic arthritis may be less likely to start as a racehorse than control patients.11 Permanent destruction of the articular cartilage, destruction and collapse of the physis, collapse of the joint (tarsus), or pathological fracture
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are likely outcomes of poorly controlled sepsis that result in humane destruction or poor long-term function of the animal. Salmonellosis is commonly considered a more rapidly destructive infectious agent with a poorer prognosis.
References 1. Firth EC. Current concepts of infectious polyarthritis in foals. Equine Vet J 1983;15:5. 2. Lugo J, Gaughan EM. Septic arthritis, tenosynovitis, and infections of hoof structures. Vet Clin Equine 2006;22: 363–88. 3. Egerbacher M, Edinger J, Tschulenk W. Effects of enrofloxacin and ciprofloxacin hydrochloride on canine and equine chondrocytes in culture. Am J Vet Res 2001;62: 704–8. 4. Murphey ED, Santschi EG, Papich MG. Regional intravenous perfusion of the distal limb of horses with amikacin sulfate. J Vet Pharmacol Ther 1999;22:68–71.
5. Rubio-Martinez LM, Cruz AM. Antimicrobial regional limb perfusion in horses. J Am Vet Med Assoc 2006:228(5):706–12. 6. Werner LA, Hardy J, Bertone AL. Bone gentamicin concentration after intra-articular injection or regional perfusion in the horse. Vet Surg 2003;32:559–65. 7. Slovis NM. Hyperbaric oxygen therapy in equine medicine. ACVIM Forum Proceedings, 2006; pp. 150–2. 8. Niinikoski J. Hyperbaric oxygen therapy in chronic osteomyelitis. In: Newman TS, Thom SR (eds) Physiology and Medicine of Hyperbaric Oxygen Therapy. Philadelphia: Saunders, 2008; pp. 419–26. 9. Mader JT, Adams KR, Wallace WR, Calhoun JH. Hyperbaric oxygen as adjunctive therapy for osteomyelitis. Infect Dis Clin North Am 1990;4(3):433–40. 10. Mader JT, Brown GL, Guckian JC, Wells CH, Reinarz JA. A mechanism for the amelioration by hyperbaric oxygen of experimental staphylococcal osteomyelitis in rabbits. J Infect Dis 1980;142(6):915–22. 11. Smith LJ, Marr CM, Payne RJ, Stoneham SJ, Reid SWJ. What is the likelihood that Thoroughbred foals treated for septic arthritis will race? Equine Vet J 2004;36(5):452–6.
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Chapter 51
Muscle Disorders Erica C. McKenzie and Jennifer M. MacLeay
Hereditary muscular disorders affecting foals during their first year of life include hyperkalemic periodic paralysis (HYPP), polysaccharide storage myopathy (PSSM), and glycogen branching enzyme deficiency (GBED).1–3 Congenital myotonic conditions also occur, and acquired muscular disorders of foals include trauma, nutritional myodegeneration (NMD), and myonecrosis subsequent to sepsis and hypoxia.4–8
Glycogen branching enzyme deficiency GBED is a significant cause of abortion and early neonatal mortality in Quarter horses and Paint horses.3, 9 In foals surviving to parturition, clinical signs are present at or soon after birth, and can be easily confused with sepsis and maladjustment.3 Pathophysiology Clinically affected foals are homozygous for a nonsense mutation of the Gbe1 gene resulting in an inability to produce functional glycogen branching enzyme (GBE).10 Deficiency of GBE results in accumulation of abnormal long straight chains of glycogen within multiple tissues, subsequently severely impairing glucose homeostasis.3 Horses heterozygous for the mutation are asymptomatic, consistent with the recessive nature of the condition, but have measurably reduced GBE activity compared to horses without the mutation.3 Clinical signs and laboratory findings Although abortion is a common consequence for the equine fetus homozygous for the Gbe1 mutation, foals that survive to parturition may display persistent weakness or recumbency, quadrilateral flexural limb deformities, seizures, respiratory failure, and
sudden death during mild exertion.3, 11 Characteristic laboratory abnormalities include leukopenia, mild to moderate elevations of serum CK and AST, and elevation of serum GGT. Recurrent hypoglycemia may occur in some foals, induced by even brief periods of nutritional deprivation.3 Death is inevitable in homozygous foals, usually occurring within the first 6 weeks of life. However, with intensive support some affected foals have survived until 18 weeks of age.3 Diagnosis Gross abnormalities are rarely present at necropsy and samples of cardiac and skeletal muscle, and brain and liver tissue should be obtained for histopathological examination. Hematoxylin-eosin stained tissue sections may appear unremarkable, or basophilic inclusions may be noted in skeletal and cardiac muscle sections, particularly in the cardiac purkinje fibers.3, 9, 10 Periodic acid–Schiff (PAS) staining of skeletal and cardiac muscle sections reveals decreased PAS stain intensity (consistent with low tissue glycogen concentrations) and abnormal globular or crystalline polysaccharide inclusions, which become more pronounced as affected foals age. In aborted fetuses, PAS-positive globular inclusions in the cardiac Purkinje fibers may be the only abnormal finding.3 The assay for GBE activity in blood or tissues is difficult to perform. Hence genetic testing is the best method for diagnosis of GBED or to confirm genetic carrier status in the dam, sire or siblings of affected foals or in mares that have abortions or stillbirths. A minimum of 10 mane or tail hairs, including the hair root, can be submitted to commercial laboratories offering the test (VetGenTM and the Veterinary Genetics Laboratory at the University of California, Davis). The availability of this non-invasive method of determining GBED status should facilitate responsible breeding practices to allow elimination of this
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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highly detrimental mutation. The carrier rate of the GBED mutation in Quarter horses and Paint horses is approximately 7% to 8%, however, in the same study, between 1.3% and 3.8% of aborted fetuses of Quarter horse lineage were homozygous for the mutation. This might reflect predominance in the mating of particular lines resulting in a higher incidence of affected fetuses than pure probability would suggest.9 The mutant Gbe1 allele has not been identified in other breeds such as registered Thoroughbreds.9
Hyperkalemic periodic paralysis HYPP was first identified as an autosomal dominant disorder of Quarter horses, tracing back to a single foundation stallion, ‘Impressive.’12–14 The condition has since been documented in Paints, Palominos, Appaloosas and the Pony of the Americas breed.15 Pathophysiology HYPP results from a point mutation in the gene encoding the alpha subunit of the skeletal muscle sodium channel.13 Muscle fibers containing mutant channels have a higher resting membrane potential than normal fibers, due to abnormal sodium influx at rest, making them more susceptible to excitation.16, 17 A co-dominance phenomenon of the wild-type and mutant alleles results in a more severe phenotype in homozygotes.18, 19 Clinical signs and laboratory findings Clinical signs are often spontaneous but can be induced by stress, transport, cold weather, and anesthesia.15 Affected horses display episodic fasciculations and weakness, often accompanied by third eyelid prolapse, tachypnea, and sweating. Colic, recumbency, and sudden death are less frequent.15, 20 Homozygotes may also display respiratory stridor and dysphagia related to pharyngeal and laryngeal dysfunction, frequently within the first week of life.2, 18, 19 During clinical episodes, horses are frequently hyperkalemic and mild elevations of serum creatine kinase might be documented.15
mane and tail hairs removed at the roots. Horses are designated N/N (wild-type), H/N (heterozygous), or H/H (homozygous) based on their genetic status. The American Quarter Horse Foundation (AQHA) stipulates that all foals born in 2007 onwards and derived from Impressive lineage must undergo mandatory parentage verification and HYPP testing for registration. Homozygous (H/H) foals are no longer eligible for AQHA registration.
Treatment and prevention Mild clinical signs may respond to gentle exercise and oral glucose administration. More severe attacks associated with recumbency, discomfort, dyspnea or cardiac arrhythmia should be treated aggressively.15 Tracheotomy may be required if severe dyspnea occurs. Calcium gluconate (0.2–0.4 mL/kg of a 25% solution) can be diluted in one to two liters of 5% dextrose and given intravenously to combat hyperkalemia, for cardioprotective effects, and to reduce muscle cell membrane potential by enhancing the activity of membrane Na+ /K+ ATPase.15, 20, 22 Intravenous sodium bicarbonate (0.5–1.0 mL/kg of a 1.3% solution) or dextrose (5% solution) also reduce hyperkalemia.22 Insulin and epinephrine enhance intracellular movement of potassium via stimulation of membrane Na+ /K+ ATPase, however, risks of administration include hypoglycemia and cardiac arrhythmia, respectively.15, 22 Long-term management can be achieved by feeding a low potassium ration and treatment with acetazolamide (2–4 mg/kg p.o. b.i.d.).15, 20, 23
Polysaccharide storage myopathy PSSM was first described in Quarter Horses and their crosses, and Paint and Appaloosa horses.24 The condition is now known to occur in 10% of Quarter Horses and commonly in some draft breeds such as Belgians (36% of which have PSSM) and Percherons and their crosses.25–27 Warmbloods and several other light breeds are affected by PSSM, albeit with a much lower prevalence.26, 28
Diagnosis
Pathophysiology
Electromyography is now rarely used for diagnosis, but 90% of affected horses display abnormal spontaneous activity including fibrillation potentials and myotonic or pseudomyotonic discharges.2, 20, 21 Skeletal muscle histopathology may be unremarkable or comprise non-specific mild degenerative changes and vacuolation of type II fibers.20 Genetic testing for the causative mutation can be performed on 20 to 30
Glycolytic and glycogenolytic enzyme capacities of Quarter Horses and Belgian Draft horses with PSSM are normal despite muscle glycogen concentrations between 1.5 and 4 times greater than those of normal horses.29–31 Increased insulin sensitivity and faster rates of blood glucose clearance have been documented in Quarter Horses with PSSM as early as 6 months of age.1, 32, 33 In 2008, an autosomal dominant
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Muscle Disorders mutation in the gene encoding the skeletal muscle glycogen synthase enzyme (GYS1) was identified as the primary cause of PSSM in Quarter horses, Draft horses and their crosses.34 Clinical and laboratory findings Affected foals may display persistent increases in serum CK activity as early as 4 weeks of age with no overt clinical signs, particularly if they are stall rested. However, muscle pain and cramping after short-duration exercise commonly occurs in affected foals, and muscle atrophy may also occur.1 Additionally, respiratory infections and other diseases may induce severe non-exertional rhabdomyolysis in foals with underlying PSSM; however, the mechanism for this phenomenon is currently unknown.1, 35 Clinical signs in such cases may include stiffness, fasciculations, dragging of the hindlimbs and dog sitting. Profound increases in serum CK activity and myoglobinuria may be documented in severely affected foals.35 Diagnosis Currently in adult horses, observation of characteristic amylase-resistant abnormal polysaccharide inclusions within muscle fibers on PAS-stained frozen sections of skeletal muscle is the most specific means of diagnosis, but requires a specialized laboratory.36 However, abnormal polysaccharide may not be visible in horses with PSSM until at least 7 to 18 months of age, and therefore muscle biopsies may provide a false-negative result in foals.1 Biopsy of semimembranosus muscle of the dam and sire might be considered if biopsy examination is not forthcoming in a foal suspected to have PSSM. However, genetic testing for the causative mutation is now offered through the University of Minnesota Diagnostic Laboratory and will greatly facilitate the diagnosis of PSSM in foals and young horses. Treatment and prevention Severe acute rhabdomyolysis requires prompt medical care, including intravenous fluids, correction of electrolyte abnormalities, and treatment of any underlying respiratory disease.7, 35 Simultaneous provision of a low-starch, high-fat diet and regular graduated exercise successfully reduces episodes of rhabdomyolysis in adult horses with PSSM.37 Although these measures have not been specifically studied in young foals, daily turnout onto pasture and provision of a balanced, low-starch, fat-supplemented diet that meets caloric demands and protein, vitamin, and mineral requirements as dictated by the foal’s age is likely to be beneficial.1 This diet should also be fed to
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the dam until the foal is weaned to prevent the foal from accessing high starch in the mare’s feed. Although early studies suggested that PSSM was an autosomal recessive trait, more recent findings indicate PSSM is autosomal dominant, in line with the substantial prevalence observed in some breeds.1, 26 The recent availability of genetic testing should facilitate sound reproductive management of breeds in which PSSM has a high prevalence, with the aim of reducing prevalence.26
Myotonia Myotonia is characterized by delayed relaxation of muscle following mechanical stimulation or voluntary contraction. Both myotonia congenita and myotonia dystrophica have been reported in foals.5, 8, 38 Pathophysiology Although abnormalities of sarcolemmal chloride and sodium conduction have been identified in congenital myotonias of other species, a similar phenomenon has not yet been identified in the horse.39 Additionally, dystrophic forms of myotonia possibly represent different sarcolemmal defects to those occurring with myotonia congenita, despite similarities in clinical presentation.39 It is currently unknown whether equine myotonia is heritable; however, Thoroughbred, Quarter horse and Anglo-Arabian foals have reportedly been affected.40 Clinical and laboratory findings Affected animals classically display a stiff gait, particularly after rising, and prolonged muscle contraction after local percussion. Both atrophy and hypertrophy of affected musculature has been described, and hindlimb musculature may be particularly prominent.8, 41 Additional congenital abnormalities occasionally occur in affected foals and elevation of serum CK and AST activity may also occur in some affected foals.8, 41 Myotonia is frequently progressive, with signs present at birth or developing during the first 6 months of life.5, 8, 38 Although clinical signs may plateau in severity, affected foals are usually euthanized prior to adulthood. Diagnosis Myotonia is diagnosed on the basis of clinical signs, electromyography, and histological examination of skeletal muscle tissue. Electromyography of affected foals reveals classic waning (‘dive-bomber’) or
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waxing and waning (‘revving motorcycle’) sounds of the EMG trains of high-frequency myotonic discharges.8 Biopsies of affected limb muscles can be submitted for analysis by a laboratory accustomed to examining equine muscle biopsies. Histological findings may be unremarkable or fiber size variation, fiber type grouping, increased numbers of central nuclei, necrosis, degeneration and increased connective tissue may be observed. Histological abnormalities may be more profound with dystrophic myotonia.5, 41
Treatment and prevention A variety of pharmaceutical agents have been used to treat horses with myotonia, with little success. Affected horses are usually euthanized due to the poor prognosis for normal function, and breeding of affected horses should be strongly discouraged due to the unknown heritability of myotonia syndromes in horses.
Nutritional myodegeneration Nutritional myodegeneration (NMD) is an acquired muscular disorder arising from dietary deficiency of vitamin E and/or selenium. The condition is also known as ‘white muscle disease’ due to the characteristic appearance of affected musculature.
Clinical and laboratory findings Clinical signs of NMD can vary substantially depending on the distribution and extent of muscular lesions. Signs most commonly occur in animals less than 2 months of age and often progress rapidly. In severely affected foals, the myocardium, diaphragm, and respiratory muscles are often affected, leading to signs of heart failure and respiratory compromise, including tachycardia, arrhythmias, cardiac murmurs, respiratory distress, and sudden death. Less severely affected foals may display weakness, stiffness, lethargy, recumbency, and discomfort on palpation of affected musculature. Dysphagia is often noted due to involvement of the tongue, pharyngeal, and sometimes masticatory muscles, and may result in aspiration pneumonia. Regurgitation, starvation, and ptyalism can also occur. Recovery can occur within days or weeks with appropriate treatment.4 Elevation of serum AST and CK activity is observed in acute disease but values decline with chronicity and recovery. In one study, muscle enzyme activity was elevated in approximately 25% of mares with affected foals and in 20% of clinically normal foals in the same stable, suggesting subclinical disease may occur.42 Severe rhabdomyolysis may re-
sult in hyperkalemia, hyponatremia, hypocalcemia, hypochloremia, and myoglobinuria in affected foals.7 Diagnosis The combination of appropriate signalment, clinical signs, and elevations in serum CK and AST activities, in addition to response to appropriate treatment, is strongly suggestive of NMD in foals. Selenium and vitamin E status can be determined in affected foals prior to supplementation. Selenium concentrations can be measured in whole blood or a liver biopsy, or the activity of selenium-dependent glutathione peroxidase (GSH-Px) in red cells can be determined. In non-deficient horses whole blood selenium concentrations should exceed 0.07 ppm, liver selenium concentrations range from 1.05 to 3.5 µg/g, and adequate GSH-Px activity ranges from 20 to 50 U/mg of hemoglobin per minute although the specific reference range for the laboratory where measurements are made should be applied.43 Plasma vitamin E concentrations should normally exceed 1.1 ppm, however, samples should be protected from heat and light prior to analysis due to the labile nature of plasma vitamin E.43 Testing of the dam may help support a need for supplementation or a diagnosis of NMD in foals if marginal or low concentrations of vitamin E or selenium are observed in the mare. Necropsy reveals bilaterally symmetrical myodegeneration particularly in limb muscles, with paleness and streaking of affected musculature. The diaphragm, tongue, pharynx, cervical musculature, and muscles of mastication are commonly affected and myocardium may also be involved. Macroscopic findings in peracute cases may be minimal. Histopathological examination of skeletal muscle may reveal myofiber degeneration or phagocytosis, and evidence of regeneration depending on the chronicity of the lesions.43 Treatment and prevention Affected foals should be confined to reduce physical activity, and promptly supplemented with vitamin E and selenium. Selenium can be administered as a sinR , 1 mL gle intramuscular injection (0.06 mg/kg, E-Se per 45 lb bodyweight) and vitamin E (2–10 IU/kg) may be given orally or by intramuscular injection although commercial injectable preparations with a high concentration of alpha-tocopherol (300 IU/mL) are preferable.43 Dysphagic, anorectic or recumbent foals may require additional supportive care including nasogastric feeding and appropriate treatment for concurrent disorders including pneumonia and sepsis. The prognosis is guarded in severely or chronically affected foals.
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Muscle Disorders Foals require 2–6 IU/kg bodyweight of vitamin E per day. Natural forms of vitamin E (RRRα-tocopherol, formerly known as d-α-tocopherol) should be prioritized since they have superior bioavailability to synthetic vitamin E (all-rac-αtocopherol, formerly known as dl-α-tocopherol). In deficient areas, mares should be supplemented with 1 mg of selenium orally per day and 600–2000 IU of vitamin E during gestation.44 Supplementation of selenium in mares and foals should be performed judiciously due to the risk of toxicity. However, placental transfer of selenium is limited, and selenium does not concentrate in milk, which may explain why NMD may occur in the offspring of supplemented mares.45 Therefore effective prevention of NMD in seleniumdeficient areas may require supplementation of foals at birth, repeating at periodic intervals every 2 to 3 months during the first 6 months of life.4, 45
Gastrocnemius rupture Rupture of the gastrocnemius muscle has been described in neonatal foals. The condition usually presents during the first 4 days after a complicated birth.6 Pathophysiology Affected foals usually have a history of dystocia and assisted delivery. Inappropriate positioning of the hindlimbs during delivery using traction likely results in traumatic damage to the gastrocnemius muscle of one or both limbs. Dystocia resulting from hip lock in particular may predispose to the condition.6 Clinical and laboratory findings Affected foals may be unable to bear weight on the affected limb and swelling is often appreciable in the caudal thigh. Partial tears may result in a ‘dropped hock’ appearance with failure of the reciprocal apparatus. Additionally, laxity or effusion of the stifle joint may also occur. In severe cases, significant hemorrhage into the muscle belly may result in clinical signs of hypovolemic shock.6 Diagnosis The diagnosis can be made from the clinical appearance when disruption of the reciprocal apparatus is appreciated in the standing patient. Ultrasonography of the gastrocnemius muscles may reveal muscle tissue disruption, fibrosis or mineralization depending on the chronicity and severity of the injury. Radiographs of the stifle and tarsal joints should also be
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considered to detect avulsion fractures and other abnormalities.6 Treatment and prevention Mild gastrocnemius injuries are expected to respond well to restricted exercise and anti-inflammatory treatment. Severe injury may require intensive supportive care with forced recumbency and application of a splint to limit flexion of the tarsal joint. The prognosis for survival of foals with severe gastrocnemius injury is guarded; however, foals with mild unilateral injuries likely have a good prognosis for survival and athletic function, as long as concurrent abnormalities are not present and significant fibrosis of the damaged musculature does not occur.6
References 1. DeLaCorte FD, Valberg SJ. Developmental onset of polysaccharide storage myopathy. J Vet Intern Med 2002;16(5): 581–7. 2. Traub-Dargatz JL, Ingram JT, Stashak TS, Kiper ML, Tarr S, Child G, MacAllister CG. Respiratory stridor associated with polymyopathy suspected to be hyperkalemic periodic paralysis in four quarter horse foals. J Am Vet Med Assoc 1992;201(1):85–9. 3. Valberg SJ, Ward TL, Rush B, Kinde H, Hiraragi H, Nahey D, Fyfe J, Mickelson JR. Glycogen branching enzyme deficiency in quarter horse foals. J Vet Intern Med 2001;15(6):572–80. 4. Dill S, Rebhun W. White muscle disease in foals. Comp Contin Educ Pract Vet 1985;7: S627–36. 5. Hegreberg GA, Reed SM. Skeletal muscle changes associated with equine myotonic dystrophy. Acta Neuropathol 1990;80(4):426–31. 6. Jesty SA, Palmer JE, Parente EJ, Schaer TP, Wilkins PA. Rupture of the gastrocnemius muscle in six foals. J Am Vet Med Assoc 2005;227(12):1965–8, 1929. 7. Perkins G, Valberg SJ, Madigan JM, Carlson GP, Jones SL. Electrolyte disturbances in foals with severe rhabdomyolysis. J Vet Intern Med 1998;12(3):173–7. 8. Reed SM, Hegreberg GA, Bayly WM, Brown CM, Paradis MR, Clemmons RM. Progressive myotonia in foals resembling human dystrophia myotonica. Muscle Nerve 1988;11(4):291–6. 9. Wagner ML, Valberg SJ, Ames E.G., Bauer MM, Wiseman JA, Penedo MC, Kinde H, Abbitt B, Mickelson JR. Allele frequency and likely impact of the glycogen branching enzyme deficiency gene in Quarter Horse and Paint Horse populations. J Vet Intern Med 2006;20(5):1207–11. 10. Ward TL, Valberg SJ, Adelson DL, Abbey CA, Binns MM, Mickelson JR. Glycogen branching enzyme (GBE1) mutation causing equine glycogen storage disease IV. Mamm Genome 2004;15(7):570–7. 11. Sponseller BT, Valberg SJ, Ward TL, Fales-Williams AJ, Mickelson JR. Muscular weakness and recumbency in a Quarter horse colt due to glycogen branching enzyme deficiency. Equine Vet Educ 2003;15(4):182–8. 12. Bowling AT, Byrns G, Spier S. Evidence for a single pedigree source of the hyperkalemic periodic paralysis susceptibility gene in quarter horses. Anim Genet 1996;27(4):279–81.
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13. Rudolph JA, Spier SJ, Byrns G, Rojas CV, Bernoco D, Hoffman EP. Periodic paralysis in Quarter horses: a sodium channel mutation disseminated by selective breeding. Nat Genet 1992;2(2):144–7. 14. Spier SJ, Carlson GP, Harrold D, Bowling A, Byrns G, Bernoco D. Genetic study of hyperkalemic periodic paralysis in horses. J Am Vet Med Assoc 1993;202(6):933–7. 15. Naylor JM. Hyperkalemic periodic paralysis. Vet Clin North Am Equine Pract 1997;13(1):129–44. 16. Hanna WJ, Tsushima RG, Sah R, McCutcheon LJ, Marban E, Backx PH. The equine periodic paralysis Na+ channel mutation alters molecular transitions between the open and inactivated states. J Physiol 1996;497(pt 2):349–64. 17. Pickar JG, Spier SJ, Snyder JR, Carlsen RC. Altered ionic permeability in skeletal muscle from horses with hyperkalemic periodic paralysis. Am J Physiol 1991;260(5 Pt 1):C926–33. 18. Carr EA, Spier SJ, Kortz GD, Hoffman EP. Laryngeal and pharyngeal dysfunction in horses homozygous for hyperkalemic periodic paralysis. J Am Vet Med Assoc 1996;209(4):798–803. 19. Naylor JM, Nickel DD, Trimino G, Card C, Lightfoot K, Adams G. Hyperkalaemic periodic paralysis in homozygous and heterozygous horses: a co-dominant genetic condition. Equine Vet J 1999;31(2):153–9. 20. Spier SJ, Carlson GP, Holliday TA, Cardinet GH 3rd, Pickar JG. Hyperkalemic periodic paralysis in horses. J Am Vet Med Assoc 1990;197(8):1009–17. 21. Robinson JA, Naylor JM, Crichlow EC. Use of electromyography for the diagnosis of equine hyperkalemic periodic paresis. Can J Vet Res 1990;54(4):495–500. 22. Meyer TS, Fedde MR, Cox JH, Erickson HH. Hyperkalemic periodic paralysis in horses: a review. Equine Vet J 1999;31(5):362–7. 23. Duren S. Feeding management of horses with hyperkalemic periodic paralysis (HYPP). World Equine Vet Rev 1998;3: 5–8. 24. Valberg SJ, Cardinet GH 3rd, Carlson GP, DiMauro S. Polysaccharide storage myopathy associated with recurrent exertional rhabdomyolysis in horses. Neuromusc Disord 1992;2(5–6):351–9. 25. Firshman AM, Baird JD, Valberg SJ. Prevalences and clinical signs of polysaccharide storage myopathy and shivers in Belgian Draft Horses. J Am Vet Med Assoc 2005;227(12):1958–64. 26. McCue ME, Ribeiro WP, Valberg SJ. Prevalence of polysaccharide storage myopathy in horses with neuromuscular disorders. Equine Vet J Suppl 2006;36: 340–4. 27. Valentine BA, Credille KM, Lavoie JP, Fatone S, Guard C, Cummings JF, Cooper BJ. Severe polysaccharide storage myopathy in Belgian and Percheron draught horses. Equine Vet J 1997;29(3):220–5. 28. Valentine BA, McDonough SP, Chang YF, Vonderchek AJ. Polysaccharide storage myopathy in Morgan, Arabian and Standardbred related horses and Welsh-cross ponies. Vet Pathol 2000;37(2):193–6.
29. Firshman AM, Valberg SJ, Baird JD, Hunt L, DiMauro S. Insulin sensitivity in Belgian horses with polysaccharide storage myopathy. Am J Vet Res 2008;69(6):818–23. 30. Valberg SJ, Townsend D, Mickelson JR. Skeletal muscle glycolytic capacity and phosphofructokinase regulation in horses with polysaccharide storage myopathy. Am J Vet Res 1998;59(6):782–5. 31. Valberg SJ, Mickelson JR, Gallant EM, MacLeay JM, Lentz L, de la Corte F. Exertional rhabdomyolysis in Quarter horses and Thoroughbreds: one syndrome, multiple aetiologies. Equine Vet J Suppl 1999;30:533–8. 32. Annandale EJ, Valberg SJ, Mickelson JR, Seaquist ER. Insulin sensitivity and skeletal muscle glucose transport in horses with equine polysaccharide storage myopathy. Neuromusc Disord 2004;14(10):666–74. 33. De La Corte FD, Valberg SJ, MacLeay JM, Williamson SE, Mickelson JR. Glucose uptake in horses with polysaccharide storage myopathy. Am J Vet Res 1999;60(4):458–62. 34. McCue ME, Valberg SJ, Miller MB, Wade C, DiMauro S, Akman HO, Mickelson JR. Glycogen synthase (GYS1) mutation causes a novel skeletal muscle glycogenosis. Genomics 2008;91(5):458–66. 35. Byrne E, Jones SL, Valberg SJ, Zimmel DN, Cohen N. 2000. Rhabdomyolysis in two foals with polysaccharide storage myopathy and concurrent pneumonia. Comp Contin Educ Pract Vet 2000;22: 503–7. 36. Firshman AM, Valberg SJ, Bender JB, Annandale EJ, Hayden DW. Comparison of histopathologic criteria and skeletal muscle fixation techniques for the diagnosis of polysaccharide storage myopathy in horses. Vet Pathol 2006;43(3):257–69. 37. Firshman AM, Valberg SJ, Bender JB, Finno CJ. Epidemiologic characteristics and management of polysaccharide storage myopathy in Quarter Horses. Am J Vet Res 2003;64(10):1319–27. 38. Steinberg S, Botelho S. Myotonia in a horse. Science 1962;137: 979–80. 39. McKerrell RE. Myotonia in man and animals: Confusing comparisons. Equine Vet J 1987;19(4):266–7. 40. Aleman M. A review of equine muscle disorders. Neuromusc Disord 2008;18(4):277–87. 41. Valberg SJ, Hodgson DR. Disorders of muscle tone. In: Smith BP (ed.) Large Animal Internal Medicine, 3rd edn. St Louis: Mosby, 2002; pp. 1271–2. 42. Higuchi T, Ichijo S, Osame S, Ohishi H. Studies on serum selenium and tocopherol in white muscle disease of foal. Nippon Juigaku Zasshi 1989;51(1):52–9. 43. Maas J, Parish S, Hodgson D, Valberg SJ. Nutritional myodegeneration. In: Smith BP (ed.) Large Animal Internal Medicine, 3rd edn. St Louis: Mosby, 2002; pp. 1279–82. ¨ onen ¨ 44. Ron´eus BO, Hakkarainen RV, Lindholm CA, Tyopp JT. Vitamin E requirements of adult Standardbred horses evaluated by tissue depletion and repletion. Equine Vet J 1986;18(1):50–8. 45. Maylin GA, Rubin DS, Lein DH. Selenium and vitamin E in horses. Cornell Vet 1980;70(3):272–89.
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Chapter 52
Tendon and Ligament Disorders Timothy B. Lescun and Stephen B. Adams
Tendon disorders of the foal can be classified into three general areas; tendon laxity, tendon ruptures, and flexural deformities (see Chapter 48). Tendon laxity and tendon ruptures will be discussed in this chapter.
ossified cuboidal carpal and tarsal bones, which can be assessed with radiographs. Radiographs will also detect fractures of the sesamoid bones.
Tendon laxity
Treatment of digital hyperextension will vary depending on the severity of the condition. Foals with mild laxity in which the fetlock is mildly hyperextended and the hooves rock back and forth from weight bearing on the sole to the coronary band often improve spontaneously over 5–10 days as the foals gain muscle tone.3 These foals benefit from restricted activity in a large stall or small paddock, light bandages to protect the caudal aspect of the coronary band, and hoof trimming. Strenuous exercise places the foal at risk for skin abrasions, sesamoid bone injury, crushing of cuboidal bones, or worsening of the condition. Some clinicians advocate swimming older foals to encourage muscle strengthening while minimizing the risk of injury.4 In many foals the weight-bearing surface of the hoof is not flat because the heels are crushed. The hooves should be trimmed with the heel low to maximize a flat weight-bearing surface giving the hoof a larger imprint (Figure 52.2). Foals with moderate laxity have toes permanently elevated off the ground, constant weight bearing on the caudal aspect of the coronary band, and a very low fetlock that does not touch the ground. These foals will benefit from glue-on shoes with heel extensions in addition to the medical treatment described for mild laxity (Figure 52.3). The shoes keep the sole of the hoof flat on the ground, which indirectly supports the fetlock joint. Glue-on shoes can constrict the growing hoof and cause contraction of the hoof capsule. They should be replaced every 10–14 days until no longer needed.
Pathogenesis Tendon laxity is common in newborn foals and may be particularly prevalent in Shire foals.1 However, digital hyperextension or flexural laxity are more descriptive terms since flaccidity from loss of normal flexor muscle tone to the digits is the primary problem. The condition occurs more frequently in premature or dysmature foals than in normal foals.2 The most common presentation is that of digital hyperextension, in which the foal presents with dropped fetlocks, usually with the toes of affected hooves being elevated off the ground. Weight bearing often occurs on the coronary band of the heels or the lower pastern, or in severe cases directly on the palmar or plantar aspect of the fetlock joint (Figure 52.1). Hindlimbs are more often affected than forelimbs, and all limbs in the same foal may exhibit laxity. Diagnosis and evaluation Diagnosis is made by history and clinical examination of the affected limbs. The foal should be carefully examined for other problems associated with dysmaturity or prematurity. Radiography and ultrasonography rarely contribute to the diagnosis or understanding of the cause, since structural damage to the tendons and muscles is not evident. However, affected foals often have incompletely
Treatment
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 52.2 Photograph of a foal with mild flexor tendon laxity. The dashed white line shows where trimming of the crushed heels should be performed to create a larger weight-bearing surface for the foot. Trimming alone in this foal was adequate to manage the tendon laxity and allow strengthening of flexor muscles over time.
This is a salvage procedure only. The authors have only performed this procedure on the forelimbs. Figure 52.1 Photograph of a neonate with moderate rear limb flexor tendon laxity. Note the hyperextended position of the fetlock joints, the toe position elevated off the ground, and the weight bearing on the plantar pastern and coronary band of the heels.
Many of the foals with moderate laxity will respond to treatment within several weeks. Severe laxity, with the foal walking on the caudal aspect of the fetlock and pastern area, presents a treatment challenge. Foals may respond to corrective shoes, bandaging of the pastern and coronet, and restricted exercise. Some clinicians will apply splints to support the fetlock in a more normal position while muscle tone is being developed. However, splints rapidly and easily cause pressure sores in foals due to their thin, delicate skin. The splints should not incorporate the hoof and may be counterproductive by further relaxing the muscle-tendon units, particularly if immobilization is rigid. This treatment should be reserved for the worst cases only as they are frequently associated with complications.
Surgical treatment Foals that do not respond to medical management may require surgical arthrodesis of the fetlock joints.
Prognosis The prognosis is good for all but the most severely affected foals. Most mildly affected foals recover spontaneously. Complications such as severe pressure sores on the palmar or plantar aspect of the limbs and
(a)
(b)
Figure 52.3 (a) Photograph of a foal with moderate flexor tendon laxity prior to shoeing. (b) Photograph of the same foal as in (a) following application of glue-on heel extension shoes. Notice the improved weight-bearing position of the distal limb and that the angle of the fetlocks, although improved, remains in hyperextension. The fetlock position will improve as the underlying tendon laxity resolves. The distal interphalangeal joint is in flexion following placement of the shoes.
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damage to cuboidal bones or sesamoid bones will make the prognosis less favorable.
Ruptured common digital extensor tendon The most frequent tendon rupture encountered in foals involves the common digital extensor tendon, which is typically noticed around the time of birth associated most commonly with carpal flexural deformities. It may occur unilaterally, but in the authors’ experience is more commonly bilateral. Ruptured common digital extensor tendon may be an isolated finding, it may occur secondary to flexural limb deformity, or it may result in the development of secondary flexural limb deformities.5
Figure 52.4 Photograph of a foal with bilaterally ruptured common digital extensor tendons. Characteristic dorsolateral swellings are present at the musculotendinous junction (upper arrows) and where effusion is present in the tendon sheath at the distal region of the carpus and proximal metacarpus (lower arrows).
Pathogenesis The pathogenesis of ruptured common digital extensor tendon in foals and young horses is largely unknown; however, in some cases an underlying pathology or weakness has been suggested as a contributing factor. A syndrome of congenital deformities has been reported in foals in western Canada with hyperplastic goiter.6 These reports include foals with ruptured common digital extensor tendon in approximately half the cases, and cases of other ruptured tendons, including the extensor carpii radialis tendon.6, 7 Although the etiology of the goiter and its relationship with the congenital abnormalities (which include ruptured tendons, mandibular prognathism, incomplete ossification of cuboidal bones, and flexural deformities) have not been fully elucidated, several reports implicate an underlying pathology linking these conditions.6–9 A potential link between winter greenfeed intake (typically cereal crops such as oats, high in nitrates) or a lack of mineral supplementation of the dam during pregnancy (a diet potentially low in iodine) and an increased risk of foals being born with this syndrome has been reported.9 Other reports suggest an association between congenital flexural deformities and ruptured common digital extensor tendon.5, 10 In some cases common digital extensor tendon ruptures are observed to occur in foals with pre-existing flexural deformity of the carpus and/or fetlock, presumably as a result of excessive strain on the tendon while the foal is compensating to stand and ambulate with the flexural deformity. The common digital extensor tendon consistently ruptures within its tendon sheath, suggesting an area of biomechanical weakness relative to the other regions of this musculotendinous unit. Tendon ruptures seen in neonatal foals may be associated with prematurity or dysmaturity or may
be a result of mechanical overload of immature tendinous tissues.5, 10 Diagnosis and evaluation Diagnosis of common digital extensor tendon rupture in a foal is usually straightforward, with characteristic postural and gait deficits accompanied by swelling in the affected carpal region. Following common digital extensor tendon rupture, foals typically stand with a slightly varus or bow-legged posture, and when walking have an exaggerated and slightly prolonged swing phase of the stride in the forelimbs to enable placement of the digit without knuckling at the fetlock. Initially, many foals will knuckle at the fetlock and/or carpus while walking until gait compensations are made. Some foals, however, will continue to knuckle over, and severe damage or even erosion into the fetlock joint can occur within a few days. Swelling of the common digital extensor tendon sheath on the dorsolateral aspect of the carpus is the most consistent physical finding, and the ends of the tendon are usually palpable within the swelling at the level of the distal carpal joints (Figure 52.4). Ultrasonography may be used to confirm and characterize the ruptured tendon ends but is usually unnecessary for making the diagnosis of common digital extensor tendon rupture (Figure 52.5). Evaluation of foals with ruptured common digital extensor tendons should include determining the presence or absence of concurrent flexural deformity, radial nerve dysfunction, and other musculoskeletal or congenital deformities that may reduce the prognosis or alter case management. Recognition of concurrent abnormalities such as mandibular
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RF CARPUS
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Figure 52.5 (a) Longitudinally oriented ultrasound image of the distal extent of the right common digital extensor tendon sheath in a foal with rupture of the tendon. Notice the enlarged appearance of the tendon end surrounded by fibrin within a distended tendon sheath. (b) Transverse ultrasound image of the mid-portion of the common digital extensor tendon sheath from the same foal as in (a). Notice the remaining but separated tendon fibers within the sheath. (c) Transverse ultrasound image of the proximal common digital extensor tendon sheath from the same foal as in (a) and (b) showing the proximal tendon stump and fibrinous adhesions forming within the sheath.
prognathism, poorly ossified cuboidal bones, flexural limb deformity, or thyroid hyperplasia should alert the veterinarian to the possibility of an exposurerelated dietary cause during gestation, particularly if more than one foal is affected on the same farm.7, 9 Treatment Rupture of the common digital extensor tendon that is not complicated by the presence of other musculoskeletal conditions is treated conservatively with stall confinement for up to 6 weeks. Support bandaging and splinting may be necessary to allow the foal to ambulate without knuckling over and to protect the limbs from secondary wounds, particularly on the dorsum of the fetlock. The presence of incompletely ossified cuboidal bones, flexural deformities, or other factors may result in the need for an adjusted treatment and management plan, depending on the severity of concurrent conditions. Restricting activity to a stall or small paddock area is recommended to allow fibrosis and healing of the ruptured tendon ends. Stall confinement is recommended for a total of 4–6 weeks, followed by a further 6–8 weeks in a small paddock. Support bandaging and splinting serve two purposes: preventing the postural and gait deficits associated with the rupture (allowing the foal to ambulate more easily and stand to nurse); and providing counter-pressure on the distended tendon sheath to reduce swelling and effusion. Bandaging beyond the period of stall confinement is not recommended, and splinting should be used for as short a period as possible in uncomplicated cases. Many foals learn to adjust their gait within the first week or two to compensate for the ruptured tendon,
making splinting unnecessary beyond this time. Systemic analgesic administration in the form of nonsteroidal anti-inflammatory drugs may be used in uncomplicated cases to reduce initial swelling; however, treatment beyond 48–72 hours is typically unnecessary unless concurrent problems (such as flexural deformity) require ongoing treatment. Surgical drainage of the tendon sheath and placement of a drain has been suggested to provide excellent cosmetic results,3 and drainage of the sheath with tendon debridement is also reported to provide good results.10, 11 However, excellent functional and cosmetic results are typically seen with conservative management of these cases, and currently surgical intervention is not recommended for uncomplicated cases. Prognosis The prognosis for ruptured common digital extensor tendon is good to excellent. Other concurrent factors such as the presence of incompletely ossified cuboidal bones or flexural deformity should be considered when deciding upon the prognosis for an individual foal.
Ruptured peroneus tertius Pathogenesis Rupture of the origin of the peroneus tertius tendon has been reported in two foals. In both cases an avulsion fracture of the distal lateral femur was also present.12, 13 In one of these cases there was a history of trauma associated with the affected limb being caught in a wire fence.
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Diagnosis and evaluation
Diagnosis and evaluation
Rupture of the origin of the peroneus tertius in a foal is accompanied by the postural abnormality described classically in adults, with flexion of the stifle while the tarsus is able to be extended.12 These injuries can involve the lateral femorotibial joint, and associated effusion may be apparent on the lateral aspect of the stifle. Radiographic and ultrasonographic evaluation is indicated to define the extent of the tendon injury, degree of articular involvement, and any associated or concurrent injuries.
Rupture of the gastrocnemius tendon, or more commonly the muscle, presents as a postural deficit with the hock dropped or positioned in hyperflexion during standing, and with swelling and pain of the caudal thigh and gastrocnemius region. Some foals will be unable to stand unassisted, and a definitive diagnosis is made using ultrasonographic examination. Evaluation of the foal’s packed cell volume is always indicated in cases of muscle rupture or tearing as several reports document sufficient bleeding in these cases to be life threatening.16–18
Treatment Rupture of the peroneus tertius can be treated conservatively with stall confinement initially followed by a graduated increase in activity after 12–16 weeks.13 Appropriate management of any joint involvement with the injury is indicated. Surgical treatment, with removal of the avulsion fracture off the distal lateral femur from the lateral femorotibial joint, was performed in one foal.12 Limb stabilization is typically unnecessary.
Prognosis Rupture of the peroneus tertius origin resulted in a persistent lameness 1 year following treatment in one foal12 and no residual lameness in another foal.13 These injuries involved the lateral femorotibial joint. Simple rupture of the peroneus tertius in older animals carries a good prognosis.13
Treatment Conservative management of gastrocnemius muscle rupture in neonates involves primarily nursing care with confinement and limb stabilization. Foals that have other abnormalities may require intensive care.17 Limb stabilization following disruption of the reciprocal apparatus can be effectively achieved using a modified Thomas splint-cast combination.15 This method of immobilization effectively transfers weight from the pelvis to the ground, allowing the gastrocnemius muscle to be immobilized, and can reduce ongoing muscle damage during weight bearing. Pressure sores are inevitable with this and other methods of limb immobilization for disruption of the reciprocal apparatus. Surgical repair of a gastrocnemius tendon avulsion has been performed successfully.14 Prognosis
Ruptured gastrocnemius muscle or tendon Pathogenesis Rupture of the caudal components of the reciprocal apparatus in foals and young horses has been reported. Rupture of the gastrocnemius tendon in a 3-month-old foal was presumed to be a result of injury,14 and rupture of both the gastrocnemius and superficial digital flexor muscles in a 6-month-old foal was also presumed to be a result of overexertion.15 However, partial tearing or rupture of the gastrocnemius muscle appears to be a more common clinical finding in neonates, with most cases recognized following dystocia or soon after birth.16, 17 A history of ‘hip lock’ as the cause of dystocia has been proposed to result in flexion of the tarsus with extension of the stifle during traction if a hind foot (or both hind feet) were positioned over the pelvic brim, preventing normal reciprocal extension of the tarsus.17
Rupture of the gastrocnemius tendon/muscle in a foal carries a fair prognosis for survival with a guarded to poor prognosis for athletic use.14, 15, 17
Ruptured superficial digital flexor tendon Pathogenesis Flexor tendon laxity was suggested to contribute to increased strain on the suspensory apparatus during an episode of overexertion in a 3-month-old foal that subsequently ruptured the straight sesamoidean ligament and the insertion of the superficial digital flexor tendon bilaterally resulting in subluxation of the forelimb proximal interphalangeal joints.19 The authors have diagnosed two foals with unilateral rupture of the straight sesamoidean ligament and superficial digital flexor tendon in the hindlimb resulting in pastern joint subluxation. Flexor tendon laxity was not present in these foals prior to injury.
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Diagnosis and evaluation Rupture of the superficial digital flexor tendon insertion associated with the straight sesamoidean ligament results in pastern joint subluxation due to the loss of palmar or plantar support of the joint. Radiographic and ultrasonographic evaluation is indicated to thoroughly assess the injury. Treatment The preferred treatment for rupture of the straight sesamoidean ligament and the superficial digital flexor tendon insertion is arthrodesis of the pastern joint to counteract the loss of palmar joint support and stabilize the digit. Splinting the limb with a lightly padded bandage in an upright position, keeping the fetlock angle at 180◦ , with the splint contacting the dorsal hoof wall distally and the cannon region proximally, should be undertaken for initial stabilization and transport. Long-term management with a splint is unlikely to return the pastern joint to a normal position, and development of osteoarthritis of the pastern joint following this injury makes conservative management less than ideal. Prognosis The authors have treated two foals with this injury involving a single hindlimb in each case. Both were treated by arthrodesis of the pastern joint and both were sound as yearlings when owners were contacted at follow up.
References 1. Knottenbelt DC, Holdstock N, Madigan JE. Flexural laxity (digital hyperextension deformity). In: Knottenbelt DC, Holdstock N, Madigan JE (eds), Equine Neonatology Medicine and Surgery. Saunders, 2004; pp. 297–9. 2. Trumble TN. 2005. Orthopedic disorders in neonatal foals. Vet Clin North Am Equine Pract 21(2):357–385. 3. Auer JA. Flexural limb deformities. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St Louis: Saunders Elsevier, 2006; pp. 1150–65.
4. Fackelman GE. Deformities of the appendicular skeleton. In: Jennings PB (ed.) The Practice of Large Animal Surgery. 1984; pp. 980–1. WB Saunders Co. 5. Yovich JV, Stashak TS, McIlwraith CW. Rupture of the common digital extensor tendon in foals. Comp Cont Educ Pract Vet 1984;6: S373–S378. 6. McLaughlin BG, Doige CE. Congenital hyperplastic goitre and musculoskeletal lesions in foals. Can Vet J 1981;22:130–3. 7. Allen AL, Doige CE, Fretz PB, Townsend HGG. Hyperplasia of the thyroid gland and concurrent musculoskeletal deformities in western Canadian foals: Reexamination of a previously described syndrome. Can Vet J 1994;35: 31–8. 8. McLaughlin BG, Doige CE, McLaughlin PS. Thyroid hormone levels in foals with musculoskeletal lesions. Can Vet J 1986;27:264–7. 9. Allen AL, Townsend HGG, Doige CE, Fretz PB. A casecontrol study of the congenital hypothyroidism and dysmaturity syndrome of foals. Can Vet J 1996;37:349–58. 10. Myers VS, Gordon GW. Ruptured common digital extensor tendons associated with contracted flexor tendons in foals. In: Proceedings of the 21st Annual Convention of the American Association of Equine Practitioners 1976, 21, pp. 67–74. 11. Stevenson WL, Stevenson WG. Rupture of the common digital extensor in foals. Can J Comp Med 1944;6:197–203. 12. Blikslager AT, Bristol DG. Avulsion of the origin of the peroneus tertius tendon in a foal. J Am Vet Med Assoc 1994;204:1483–5. 13. Koenig J, Cruz A, Genovese R, Fretz P, Trostle S. Rupture of the peroneus tertius tendon in 27 horses. Can Vet J 2005;46:503–6. 14. Valdez H, Coy CH, Swanson T. Flexible carbon fiber for repair of gastrocnemius and superficial digital flexor tendons in a heifer and gastrocnemius tendon in a foal. J Am Vet Med Assoc 1982;181:154–7. 15. Lescun TB, Hawkins JF, Seims JJ. Management of rupture of the gastrocnemius and superficial digital flexor muscles with a modified Thomas splint-cast combination in a horse. J Am Vet Med Assoc 1998;213:1457–9. 16. Sprinkle FP, Swerczek TW, Crowe MW. Gastrocnemius muscle rupture and hemorrhage in foals. Equine Pract 1985;7:10–17. 17. Jesty SA, Palmer JE, Parente EJ, Schaer TP, Wilkins PA. Rupture of the gastrocnemius muscle in six foals. J Am Vet Med Assoc 2005;227:1965–8. 18. Pascoe RR. Death due to rupture of the origin of the gastrocnemius muscle in a filly. Aust Vet J 1975; 51:107. 19. Harrison LJ, May SA. Bilateral subluxation of the pastern joint in the forelimbs of a foal. Vet Rec 1992;131:68–70.
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Neurological Evaluation Robert J. MacKay
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Neurologic Disorders in the Neonatal Foal Robert J. MacKay
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Chapter 53
Neurological Evaluation Robert J. MacKay
Neurological evaluation Normal neurological development Abnormal neurological function of foals can only be appreciated if the examiner has a thorough understanding of the milestones of normal neurological development, especially during the first few days of life.1–4 Foals are minimally responsive during transit through the birth canal but immediately after expulsion acquire righting reflexes and muscle tone.5 Most foals are in sternal recumbency with the head up even before the umbilical cord separates and certainly should have become sternal within 15 minutes after birth. The head adopts a noticeably more flexed angle at the atlanto-occipital joint in newborn foals than it does in adults. Within 5 minutes of birth, the foal should look bright and alert and be responsive to tactile, visual, and auditory stimuli. Both the basic6 and modified2 APGAR scoring systems for foals include evaluations of responses to stimulation by the examiner. The five criteria (Appearance, Pulse, Grimace, Activity, Respiration) used as a mnemonic learning aid were devised in the 1952 by Dr Virginia Apgar as an aid in assessing the health of babies after obstetrical anesthesia. By 30 minutes, the foal should have vocalized, usually in response to the dam’s nickering. Periods of limb extension and restlessness in sternal recumbency alternate with periods of sleep in lateral recumbency. Usually beginning within 30 minutes of birth, the foal makes the first of several lurching attempts to stand.1 Initially there is a base-wide swaying posture and the foal rocks from side to side or back and forth. The foal may pitch into the ground several times before getting solidly to its feet. By the time it stands, the foal’s head should follow movements of the mare. Once it stands solidly (approximately 1 hour after birth on average), the foal begins a series of increas-
ingly accurate approaches to the mare’s udder. This period often includes investigations of the chest between the mare’s front legs and attempts to get to the udder by going between the mare’s back legs. Coarse bobbing of the head and rocking of the body during teat seeking at this stage is considered normal. By 3 hours after birth (average of 2 hours), the foal should begin suckling for periods of 1–5 minutes. Suckling is typically followed by a period of obvious drowsiness. The head drops repeatedly and the foal often collapses awkwardly to the ground, then may sleep in lateral recumbency before standing to suckle again a few minutes later. Over the first 3 days of life, the foal becomes increasingly confident and coordinated and fully bonded with its dam. Although it usually stays close to the mare, the foal develops growing interest in the environment away from the dam. At the walk, the gait is noticeably bouncy and hypermetric but, usually within a day, the foal is capable of trotting and galloping. Foals spend considerably more time in REM sleep than adults do. Because REM sleep is associated with relaxation of skeletal muscles and inhibition of reflexes, foals usually appear to sleep more deeply than adults. However, normal sleeping foals also may actively move in a manner reminiscent of humans responding to dreams during REM sleep. These movements, which usually continue for several minutes, may include vocalization, extension of the neck and torso, and thrashing or galloping actions of the limbs. In some cases, the movements are so pronounced and unexpected that they are mistaken for seizures.
Neurological examination Detailed history-taking and a complete physical examination should precede any neurological evaluation. Previous medical and reproductive conditions of the mare, vaccination history, and environmental
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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conditions all could point to risk factors for the development of neurological disorders of the newborn foal.4, 5 In order to perform the passive part of the examination, have an assistant restrain the foal in standing position by placing one arm around the front of the chest and another around the hindquarters. If the foal is recumbent, position it sternally in order to examine the head. The examiner must be flexible in interpretation of the responses of the newborn foal. In comparison to the older foal, the responses of any neonate can seem random, demonstrably abnormal, unpredictable, and variable over time. For this reason it is very important that results of neurological examinations be recorded on a standardized form and repeated often in order to evaluate progress and solidify interpretations. The following is an outline of a minimal examination protocol for the foal less than 3 days of age.1–3 1. Observe the foal while it is free to move around. Note particularly any interactions with the dam and the examiner. This is the first opportunity to evaluate the foal’s behavior and level of alertness. Evaluation of alertness and behavior should continue throughout the examination. 2. Assess the foal’s overall conformation and muscle tone. The head should be above the horizontal plane or the neck in standing foals and the long axis (i.e., from poll to nasal philtrum) should be in the median plane when viewed from the front. Note the direction of any head tilt or rotation. The skull may have a domed appearance in some foals with hydrocephalus or the nose may be deviated laterally in cases of ‘wrynose’ (also known as ‘squiffy face’). Abnormal lateral, ventral, or dorsal curvatures of the spine are scoliosis, lordosis, or kyphosis, respectively. 3. Gently touch the inside of the ear canal then touch the side of the nasal septum. These noxious stimuli should elicit avoidance responses wherein the foal moves its head away from the touched side. Compare the responses on either side. 4. Stand in front of the foal and evaluate facial expression, tone, and symmetry. Expression usually changes from ‘blank’ to alert over the first few hours of life in normal foals. Unilateral facial paralysis is evident as drooping of the ear, eyelid, and lip on the affected side and deviation of the muzzle away from the affected side. Examine the flick reflexes on each side of the face including the lip, palpebral, and ear reflexes. These are best elicited by gently touching the tip of a hemostat to the commisure of the lip, the medial and lateral canthi of the eye, and the pinna of the ear. A normal response is brisk lip retraction, eyelid clo-
sure, and ear flick, respectively. Remember that flick reflexes involve only the hindbrain and do not require any conscious reaction by the foal. 5. Examine the eyes. Use a light to check pupillary size, orientation, and diameter. There normally is slight ventromedial rotation of the eyeballs in neonatal foals compared to older foals. A strong light directed into the eye should elicit both a blink in the ipsilateral eye (dazzle reflex) and pupillary contraction in both eyes (direct and consensual pupillary light reflexes). These reflexes should be brisk and symmetric. As with older horses, the eyeballs should be oriented centrally within the orbits and should maintain their absolute positions when the chin is lifted (i.e., they should appear to rotate ventrally relative to the long axis of the head). Look for rhythmic movements of the eyes when the head is held stationary in normal position. Such spontaneous nystagmus is always abnormal. Next, move the head from side to side in a horizontal plane and look for normal physiological nystagmus with the fast phase in the direction of head movement. Finally, in alert foals more than 24 hours old, try to elicit a visual avoidance response. Tap the face firmly to gain the foal’s attention then make a threatening gesture toward the eye. Although neonatal foals less than 1–2 weeks have not yet acquired a menace response, many move the whole head away in reaction to such a gesture. 6. Insert a clean (preferably gloved) finger into the mouth and assess the suckle reflex and jaw tone. The suckle reflex should be present within a few minutes of birth and should be strong within an hour. Allow the foal to suckle the mare and assess head and neck coordination as the foal seeks the teat, then look for swallowing movements during suckling. Check after suckling for regurgitation of milk from the nose. (Auscultation over the trachea immediately post suckle may detect a tracheal rattle suggestive of dysphagia.) 7. Check long spinal reflexes. First, test the cervicofacial reflex by prodding the skin over the brachiocephalicus muscle at intervals along the neck. An appropriate response is simultaneous shrugging of the shoulder (brachiocephalicus contraction) and twitching of the face. Evaluate the cutaneous trunci reflex (also known as panniculus). Prod the skin over each intercostal space and look for twitching of the cutaneous muscle. The reflex is present continuously from the back part of the triceps to the last intercostal space. If any deficits are found in the cervicofacial or cutaneous trunci reflexes, systematically evaluate cutaneous sensation by squeezing pinched skin with hemostats and looking for behavioral reaction by the foal.
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Next, run the tip of the hemostat briskly in a caudal direction along the skin over the longissimus muscles to test postural responses. The foal should extend then relax its lumbosacral spine without buckling or wobbling the pelvic limbs. Finally, run the hemostat caudally along the gluteal region. The normal response is brisk flexion then relaxation of the lumbosacral spine and pelvis. The ‘slap’ test or laryngeal adductor reflex, which is commonly performed in adult horses, is too inconsistent in neonates to be of value. Test limb strength with one hand by pushing downwards over the withers then over the pelvis. Pull sideways steadily on the tail. The normal response by the foal to these tests is to give slightly to the pressure then firmly resist. Check caudal structures. Lift the tail and note its tone. Test the anal/tail clamp reflex by tapping the anus with the tip of a hemostat. The expected response is contraction of the anal sphincter and flexion of the tail. If this reflex is abnormal carefully check perineal cutaneous sensation. Evaluate gait. This is of necessity a very limited examination in neonates compared to older foals or adults. Watch the gait during spontaneous unconstrained movement, then lead the dam in lines and circles and watch the foal as it follows. In most cases, the walk is precise and symmetric and has a four-beat cadence. The foal typically alternates between walking and trotting to keep pace with the mare. The trunk and limbs should move smoothly around turns without toe-dragging or circumduction. In some breeds a two-beat lateral gait is considered normal for neonates. Compared with adults, the trunk ‘bounces’ when a foot contacts the ground and the limbs are a little more stiff and hypermetric during the swing phases. Place the foal in lateral recumbency. Test the tone of each limb during passive flexion and extension. With the hand against the sole of the hoof, keep the phalangeal joints extended while pushing the rest of the leg into a flexed position and watch for the limb to extend in response. This extensor thrust reflex is normal. Pinch the skin over the distal part of the limb in several areas and watch for reflex flexion of the ipsilateral limb (flexor or withdrawal reflex) and extension of the opposite limb (crossed extensor reflex). A brisk flexor reflex is expected in normal foals and some normal foals also exhibit a crossed extensor reflex. Use a reversed hemostat or plexor to test additional reflexes in the upper limbs. The patellar, sciatic, and gastrocnemius reflexes can reliably be elicited in the pelvic limb. The triceps and biceps
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reflexes can be obtained in the thoracic limbs. If results of reflex testing of the limbs are abnormal, evaluate sensation over the limbs by pinch testing. ‘Autonomous zones’, wherein sensation to particular areas of skin are supplied by single nerves, have been described for the horse.7 Neuroanatomical localization Use the results of neurological examination in combination with anatomical information in Table 53.1 to localize the lesion(s) and generate a list of possible differential diagnoses.
Subarachnoid puncture and cerebrospinal fluid collection Cerebrospinal fluid (CSF) is collected for cytological analysis or other testing or in preparation for injection of contrast material for myelography. CSF can be obtained from atlanto-occipital (AO) or lumbosacral (LS) sites. The advantages of AO versus LS collection are: (i) the technique is simpler because the target space is larger and more superficially located; (ii) there is less risk of contaminating the CSF sample with blood. The advantages of LS versus AO collection are: (i) it can be performed in conscious foals; (ii) there is less risk of ‘coning’ (herniation of the cerebellum into the foramen magnum) during CSF collection from a foal with increased intracranial pressure; (iii) the needle can safely be advanced through the subarachnoid space. (If this were done at the AO site, the needle would penetrate the spinal cord.) Atlanto-occipital collection 1. Unless the foal is comatose, it should be briefly anesthetized for the procedure. A combination of i.v. diazepam (0.05–0.2 mg/kg) with i.v. ketamine (2.2 mg/kg) is adequate for most foals. If necessary, a single half-dose can be repeated for each of these drugs to prolong anesthesia. In hemodynamically unstable foals, sedation combined with local anesthesia or gas anesthesia combined with controlled ventilation via endotracheal tube may be used as alternatives to i.v. anesthesia. 2. Place the foal in lateral recumbency and palpate the puncture site as the intersection of the dorsal midline of the neck with an imaginary line joining the cranial edge of the wings of the atlas (Figure 53.1b,c). This landmark is a distinct corner found by following the wing of the atlas cranially and medially until it turns sharply medially toward the midline. Clip a 10-cm square centered on this site, then clip an additional strip which includes the
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Table 53.1 Neuroanatomical localization based on results of neurological examination Evaluation
Pathways
Major signs of disorder
Behavior
Forebrain (mostly cerebrum)
Reduced affinity for dam, restlessness, head-pressing, compulsive walking
Alertness Avoidance – nasal Avoidance – visual Head position
All of brain V,a pons, thalamus, cerebrum II, thalamus, thalamus, cerebrum VIII, medulla
Lethargy, stupor, semi-coma, coma Facial hypalgesia Blindness Head tilt, turn, body lean, walking in circles, ataxia, nystagmus
Eye position and movement Flick reflexes
III, IV, VI, VIII, midbrain, hindbrain V, VII, pons, medulla
Strabismus, nystagmus Facial paralysis, facial hypalgesia, absent flick reflexes
Dazzle reflex Pupillary light reflex Suckle Swallow Cervicofacial reflex
II, subcortex, midbrain, VII II, midbrain, III V, VII, XII, medulla, cerebrum IX, X, medulla Cervical spinal nerves and spinal cord, VII, (medulla)
Absent dazzle reflex Absent pupillary light reflex Weak or absent suckle Flow of milk from the nose Diminished cervicofacial reflex
Cutaneous trunci reflex
Thoracic spinal nerves and spinal cord, brachial plexus, lateral thoracic nerve
Diminished reflex caudal to spinal cord lesion
Cutaneous sensation
Peripheral nerves, spinal cord, cerebrum
Cutaneous hypalgesia/anesthesia
Gait
VIII, cerebellum, medulla, spinal cord, peripheral nerves
Ataxia
Limb strength
Spinal cord, peripheral nerves
Limb weakness at or caudal to the level of the lesion
Flexor reflex (pelvic), patella, sciatic reflexes
Spinal cord, peripheral nerves (L3–S2)
Pelvic limb weakness, diminished or absent reflexes
Flexor reflex (thoracic), biceps, triceps reflexes
Spinal cord, peripheral nerves (C6–T2)
Weakness of thoracic ± pelvic limbs, diminished or absent reflexes
Anal/tail clamp reflex
Spinal cord, cauda equina, peripheral nerves (S2–Co)
Reduced or absent anal/tail clamp reflex
a
Roman numerals refer to cranial nerves; Co = coccygeal spinal cord segments.
cranial edge of the wing of the atlas. Scrub and surgically prepare the clipped areas. 3. Flex the foal’s AO joint so that the long axes of the cervical vertebrae and skull are at about 90◦ (Figure 53.1a), then use a gloved hand to palpate the cranial corner of the wing of the atlas on the top side (Figure 53.1b). Use this landmark to identify the skin puncture site on the dorsal midline. 4. Introduce a needle (18–20 ga 3.8 cm needle in foals < 60 kg; 6.4–8.9 cm styletted spinal needle in larger foals) through the skin and, staying in the median plane, aim the needle toward the symphysis of the mandible. When the needle pierces the dorsal atlanto-occipital ligament and dura mater (approximately 2.5–4.5 cm from the skin surface in 40to 100-kg foals), a distinct ‘pop’ is usually felt. At this stage, stop advancing the needle, remove the stylet (if a styletted needle is used) and check for
CSF at the hub of the needle. If CSF does not appear when a pop is felt, rotate the needle several times in each direction. If CSF still does not appear, carefully advance the needle until another pop is felt and check again for CSF. If blood (undiluted with CSF) appears at the hub or if the needle strikes bone, replace the stylet, withdraw the needle point to a subcutaneous position, and redirect. Atlanto-occipital cisternal puncture can be performed either blind or under ultrasound guidance, as has been described for adult horses.8 An advantage of the latter technique is the opportunity to image the spinal cord and surrounding structures, including CSF. 5. Collect the CSF from the hub of the needle (Figure 53.1f), replace the sytlet, then slowly withdraw the needle and allow the foal’s head to return to a normal or extended position.
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(c)
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(b)
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Figure 53.1 Atlanto-occipital CSF collection. (a) Location of the cranial margin of the wing of the atlas (black line) and the cranial corner of the wing of the atlas (landmark for locating skin puncture site). (b and c) Use of the cranial corner(s) of the wing(s) of the atlas to locate skin puncture site. (d and e) Needle correctly placed in atlanto-occipital subarachnoid space. (f) Aspiration of CSF from the hub of the needle.
Lumbosacral collection 1. The procedure can be performed with the foal standing, in sternal recumbency, or in lateral recumbency. If standing lumbosacral puncture is attempted in foals less than 4 weeks old, there is a tendency for the foal to collapse in the pelvic
limbs during the procedure. It is helpful to angle the pelvic limbs forward in laterally recumbent foals to help open up the lumbosacral space. Lumbosacral puncture can be performed under ultrasound guidance, as described.9 2. Sedation should be provided as necessary (e.g., diazepam 0.05–0.2 mg/kg i.v. in combination with
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butorphanol 0.05–0.1 mg/kg i.v.) to prevent movement during the procedure. 3. Locate the approximate site for skin puncture as the intersection of an imaginary line between the caudal aspects of the tuber coxae and the dorsal midline. Palpate and identify the dorsal edges of the tuber sacrale as they form a V-shape with the vertex pointing caudally. The skin puncture site usually lies within the V. Beginning several centimeters behind the tuber sacrale, use a thumb (usually the thumb of the dominant hand) to palpate the dorsal spinous processes of the sacrum. Slide the thumb cranially along the midline until a depression is felt, usually at a site between the tuber sacrale. This is the space between the dorsal spinous processes of the sixth lumbar and first or second sacral vertebrae and is the correct site for skin puncture. Clip and surgically prepare a 6 × 6 cm square of skin centered on the skin puncture site. 4. Lumboscaral puncture usually is performed with an 8.9 cm styletted needle. The lumbosacral subarachnoid space is located approximately 4 to 8 cm beneath the skin surface. After infiltration of 0.5–1.0 mL of local anesthetic along the intended needle path, make a stab incision through the skin with a no. 15 scalpel blade. 5. Using gloved hands, introduce the needle at right angles to the skin surface and advance it until there is an involuntary reaction by the foal or the point strikes bone. Often there will be a slight pop as the needle penetrates the dorsal interarcuate ligament and meninges. Withdraw the stylet, attach a 3 mL syringe to the needle and apply gentle suction pressure to aspirate CSF. To maneuver the needle tip into a subarachnoid space, it may
be necessary to rotate the needle in either direction while gradually withdrawing it. If CSF does not flow into the syringe, replace the stylet and withdraw the needle tip to a subcutaneous position before redirecting. It is advisable to collect several aliquots of CSF to clear any blood contamination associated with the initial meningeal puncture.
References 1. Knottenbelt D, Holdstock N, Madigan JE. Equine Neonatology: Medicine and Surgery. Edinburgh: Saunders, 2004. 2. Vaala WE, House JK, Madigan JE. Initial management and physical examination of the neonate. In: Smith BP (ed.) Large Animal Internal Medicine. St Louis: Mosby, 2002; pp. 277–93. 3. Green SL, Mayhew IG. 1990. Neurologic disorders. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 496–530. 4. Vaala WE, House JK. Perinatal adaptation, asphyxia, and resuscitation. In: Smith BP (ed.) Large Animal Internal Medicine. St Louis: Mosby, 2002; pp. 266–76. 5. Wilkins PA. Disorders of foals. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine. St Louis: Saunders, 2004; pp. 1381–431. 6. Martens RJ. Pediatrics. In: Mansmann RA, McAllister ES, Pratt PW (eds) Equine Medicine and Surgery. Santa Barbara: American Veterinary Publications, 1982. 7. Mayhew IG. Large Animal Neurology: A Handbook for Veterinary Clinicians. Philadelphia: Lea & Febiger, 1989. 8. Audigi´e F, Didierlaurnt J-M, Denoix J-M. Ultrasoundguided atlanto-occipital puncture for myelography in the horse. Vet Radiol Ultrasound 2004;45:340–4. 9. Aleman M, Borchers A, Kass PH, Puchalski SM. Ultrasound-assisted collection of cerebrospinal fluid from the lumbosacral space in equids. J Am Vet Med Assoc 2007;230:378–84.
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Chapter 54
Neurological Disorders in the Neonatal Foal Robert J. MacKay
A detailed list of neurological disorders of the neonatal period (defined here as the first month of life), classified by presenting clinical signs, is provided in Table 54.1. Below is a discussion of the more important of those disorders.
Hydrocephalus Etiology Foals are born with hydrocephalus because of genetic defects in the cerebral ventricular system.1 The condition also may be acquired because of inflammatory changes associated with meningitis. In all cases with clinical signs, there is attenuation of cerebral structures associated with abnormally distended ventricles.
Clinical signs Most foals appear physically normal at birth although a few may have obvious doming of the skull. Signs vary greatly in severity but are referable to cerebral dysfunction. There may be dullness, poor bonding with the mare, head-pressing, compulsive walking, seizures, and variable development of menace responses. Involvement of the brainstem is indicated by strabismus or limb ataxia and weakness. Diagnosis The diagnosis is presumptive unless confirmed by cross-sectional imaging techniques – computed tomography (CT) and magnetic resonance imaging (MRI) – which have replaced the older technique of ventriculography.
Treatment and prognosis There is no practical treatment in foals with substantial attenuation of the forebrain. For mildly affected foals, a surgical procedure has been described for shunting cerebrospinal fluid (CSF) from the ventricular system to the peritoneal cavity.2 Foals born with hydrocephalus are unlikely to ingest adequate colostrum and are therefore unusually susceptible to neonatal sepsis. Even if they are intensively supported, most foals remain ill-thrifty.
Hypoxic-ischemic encephalopathy (HIE) Hypoxic-ischemic encephalopathy refers to noninfectious syndromes of foals aged less than 3 days and characterized by signs of central nervous system (CNS) disease; it excludes diseases caused directly by congenital/genetic defects or trauma involving the CNS. Synonyms for HIE include hypoxic-ischemic encephalomyelopathy, neonatal maladjustment syndrome, neonatal encephalopathy, dummy, wanderer, and perinatal asphyxia syndrome.3 Etiology Hypoxemia, anemia, hypoglycemia, ischemia, hemorrhage, and/or reperfusion during the perinatal period likely initiate hypoxic damage to CNS neurons.7, 55 Cascades of injurious mediators including excitatory amino acids and oxygen free radicals are released and cause neuronal necrosis, hemorrhage, and tissue edema.5, 6 Placental disease, premature placental separation, maternal illness, dystocia, cesarean section, birth ‘trauma’, pulmonary hypertension, airway obstruction, or other causes of neonatal asphyxia are risk factors for HIE.7, 55
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 54.1 Neurological disease of foals during the first month of life Non-neurological causes (breed)
Clinical presentation
Neurological causes (breed)
1
Abnormal body positions Aimless wandering Compulsive walking Disinterest in dam Head-pressing Obsessive licking Walking into objects
Brain abscess Brain malformationa Electrolyte derangement Encephalitis, incl. West Nile/EEE H. gingivalis36 Fe toxicityb 4, 37 Head trauma HHH (Morgan38 ) HIE Meningitis Moxidectin toxicity39 Porto-systemic shuntb 40 Post-seizure
Blindness associated with disorders of the eyes
2
Convulsions Muscle spasms Stiffness Sudden collapse
As for presentation 1 Inherited myoclonus (Paso41 ) Kernicterus42 Lavender foal (Arabian43 ) Narcolepsy/cataplexy Tetanus
GSD IV (QH44 ) Nettle reaction45 Nutritional myodegeneration46 OLWS (Overo Paint47 ) Other colic
3
Obtundation Lethargy Stupor Semi-coma Coma
As for presentation 1 Hypoglycemia Kernicterus Tyzzer’s diseaseb 48
EHV-1 infection Heart defect Hypothermia NI Prematurity Sepsis Starvation
4
Facial paralysis Head tilt
Brain abscess Encephalitis Head trauma HIE Meningitis Otitis media/interna
OAAM Skull malformation Wry neck (non-neurological) Vertebral malformation Wry nose
5
Ataxic, not weak Head bob Shaking Tremoring
Cerebellar abiotrophy Cerebellar dysplasia/hypoplasia49 HIE Prematurity Head trauma Fungal tremorgense
Hypothermia
6
Weak all four limbs Often recumbent
As for presentation 3 Botulism EDM Focal spinal cord lesion (C1–T2) OAAM Other vertebral malformation/stenosis Spinal abscess Vertebral trauma Inherited myoclonus (Paso) Lavender foal (Arabian) Snake bited Spinal cord malformationc Tick paralysis50
As for presentation 3 Arthrogryposis Nutritional myodegeneration GSD IV (QH) Tendon laxity Septic arthritis ALD FLD Musculoskeletal injury
(Continued).
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Table 54.1 Neurological disease of foals during the first month of life (continued) Non-neurological causes (breed)
Clinical presentation
Neurological causes (breed)
7
Weak in thoracic limb(s) only
Brachial plexus injury HIE (spinal cord) Nerve root injury (C6–T2) Peripheral nerve injury
ALD Arterial thrombosis51 FLD Musculoskeletal injury Septic arthritis Tendon laxity
8
Weak in pelvic limb(s) only
Focal spinal cord lesion (T3–S2) Spinal abscess Vertebral malformation/stenosis Vertebral trauma Lumbosacral plexus injury Meningitis Nerve root injury – L3–S2 Peripheral nerve injury Spinal cord malformation (see Figure 54.8)c >
Tendon laxity Gastrocnemius/SDF rupture Septic arthritis FLD ALD Arterial thrombosis52 Musculoskeletal injury
9
CES Perineal hypalgesia Rectal/anal paralysis Tail paralysis Urinary incontinence
Meningitis Spinal cord/cauda equina (S2–caudal) Spinal abscess Vertebral malformation/stenosis Vertebral trauma Spinal cord malformationc53, 54
Ectopic ureter Musculoskeletal injury
a
Mainly hydrocephalus. Signs due to hepatoencephalopathy. c Mainly myelodysplasia. d Only reported in Australia. e Mainly perennial ryegrass staggers. ALD, angular limb deformity; CES, cauda equina syndrome; EDM, equine degenerative myeloencephalopathy; EEE, Eastern equine encephalitis; FLD, flexural limb deformity; GSD IV, glycogen storage disease IV; HHH, hyperammonemia-homocitrullinuria-hyperornithinemia, a urea cycle disorder; HIE, hypoxicischemic encephalopathy; NI, neonatal isoerythrolysis; OAAM, occipitoatlantoaxial malformation; OLWS, Overo lethal white syndrome; QH, Quarter Horse; SDF, superficial digital flexor. b
Clinical signs Within the continuum of clinical signs of HIE, two major categories have been discerned.8, 9 Category 1 foals are born normally then develop signs within the first 48 hours. Category 2 foals usually have risk factors for HIE and are abnormal from birth. Clinical signs are mostly referable to cerebral dysfunction (Figure 54.1) but some foals (usually category 2) show additional signs indicating involvement of the brainstem and/or spinal cord (Figure 54.2).10 There are behavioral abnormalities, including lack of affinity for or interest in the dam, restlessness, hyperresponsiveness to handling, abnormal posturing, tongue protrusion, abnormal jaw and facial movements, ‘star-gazing’, head-pressing, obsessive licking, and abnormal vocalization. Recurrent seizures are common; signs range from mild abnormal movements of the face and jaw to generalized seizures with recumbency and paddling of the limbs. In addition to behavioral abnormalities, there may be lethargy or stupor, head-tilt, facial paralysis, abnormal breathing
patterns, and ataxia and weakness of the limbs. Signs of multiple organ dysfunction involving pulmonary, renal, hemostatic, gastrointestinal, or hepatic systems may complicate the clinical picture of HIE.7, 55 Diagnosis The diagnosis of HIE relies on compatible history and clinical findings and exclusion of competing diagnoses.9, 10 Hematology and serum chemistry results usually are normal unless the presentation is complicated by sepsis or organ dysfunction. Cerebrospinal fluid may be normal or xanthochromic with increased concentration of red blood cells (RBC) and protein.5, 10 Necrosis, edema and hemorrhage of the CNS are characteristic post-mortem findings. Treatment and prognosis The aims of therapy are (i) general support; (ii) control of seizures; (iii) prevention of sepsis; and
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(a)
(b)
(c)
(d)
Figure 54.1 Abnormal neurological signs typical of foals with hypoxic-ischemic encephalopathy (category 1). (a) Involuntary retraction of the lips and twitching of the muzzle, often seen during mild localized seizures or at the onset of generalized seizures (image from video). (b) Excessive tongue wringing or protrusion. (c) Generalized seizures. (d) Obsessive licking of the wall or other objects. Image in panel (c) courtesy of I.G. Mayhew, Massey University, Palmerston North, New Zealand.
(iv) treatment of CNS edema and hemorrhage. Broad-spectrum antimicrobial coverage and general support, according to the principles of critical care treatment discussed elsewhere (see Chapter 19), are the cornerstones of management. Seizures should be controlled with one or two doses of diazepam (0.1–0.2 mg/kg i.v.). If seizures persist, sodium phenobarbital should be started (10–20 mg/kg i.v. over 15 min, then 5 mg/kg i.v. or p.o. twice daily) and continued for at least 7 days. In foals with intractable seizures, a continuous infusion of midazolam (0.1–0.5 mg/kg/h) has been effective. Total
intravenous anesthesia (e.g., infusion of propofol) for several hours may be necessary if status epilepticus persists. The most uncertain part of therapy is the use of anti-inflammatory/anti-edema drugs designed to treat or prevent CNS injury. The following therapies are used empirically in foals with HIE but such uses have not yet been validated by the results of clinical trials (in humans or horses):
r r
flunixin meglumine (1.1 mg/kg i.v. every 12 h) MgSO4 (0.05 g/kg/h loading dose, then 0.025 g/kg/h)
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positive and Gram-negative bacteria have been implicated in published reports. Clinical signs Hyperesthesia and rigidity of the neck in combination with signs of CNS disease are highly suggestive of bacterial meningitis. There may be degrees of obtundation ranging to coma, opisthotonus, seizure activity, cranial nerve abnormalities, and ataxia and weakness of the limbs. An associated abscess may cause additional localizing neurological signs. Signs of extraneural sepsis such as fever, joint distension, pneumonia, or umbilical abscess are expected. Diagnosis
Figure 54.2 Brainstem signs in hypoxic-ischemic encephalopathy (HIE) (category 2). This 3-week-old Peruvian Paso foal presented at 1 day of age with signs as described for Figure 54.1 plus obtundation, head tilt, head and neck turn, and tight circling to the right. The foal’s condition did not improve and it was euthanized. Necrosis of the brainstem involving the right vestibular nuclei was found at necropsy.
r r r r r r r
thiamine (10–20 mg/kg i.v. slowly or s.c. once daily) dimethyl sulfoxide (DMSO; 1 g/kg as a 10% solution i.v. every 12 or 24 h) vitamin E (20 IU/kg s.c. or p.o. once daily) vitamin C (100 mg/kg i.v. or p.o. once daily) mannitol (0.5–2.0 g/kg i.v. every 6 h) hypertonic saline (2 mL/kg of 7.2% NaCl every 4 h for 5 treatments) furosemide (1 mg/kg i.v. every 12–24 h).
In the author’s opinion, the risk of inducing immunosuppression-associated sepsis outweighs any potential advantage of corticosteroid therapy. Uncomplicated (category 1) HIE generally resolves completely within several days if treated appropriately. Survival rate for these foals is estimated to be at least 75%.10, 11 The spinal cord, brainstem, or multiple organ dysfunction typical of category 2 HIE dramatically worsens the prognosis.
Hematology and serum chemistry results reflect systemic sepsis and organ failure. The serum immunoglobulin G (IgG) concentration typically is low (< 8 g/L) if there is failure of passive transfer. Definitive diagnosis is by CSF analysis. There is a neutrophilic pleocytosis (usually > 5 × 108 cells/L) with cellular degeneration and bacteria are often seen on microscopic examination of CSF sediment. Treatment and prognosis Effective broad-spectrum coverage can be provided by the third-generation cephalosporins cefotaxime (40 mg/kg given i.v. every 6 or 8 h) or ceftriaxone (25 mg/kg i.v. every 12 h). Although expensive, these drugs likely have excellent penetration into the CNS. Alternative antimicrobial regimens include ceftiofur (5 mg/kg i.v. every 12 h), amikacin sulfate (25 mg/kg i.v. every 24 h) plus ampicillin sodium (20 mg/kg i.v. every 6–8 h), trimethoprim–sulfamethoxazole (30 mg/kg i.v. or p.o. every 12 h), or chloramphenicol sodium succinate (50 mg/kg i.v. every 12 h in foals less than 7 days old, then every 6 h). Dexamethasone sodium phosphate (0.4 mg/kg i.v.) or dexamethasone (0.3 mg/kg i.v.) every 12 hours for 2 days may improve survival.12 The steroid should begin with or before the first dose of antimicrobial. With neck stiffness and hyperesthesia only (meningitis only), the prognosis is fair (> 50% recover); in foals that are also obtunded or weak (meningoencephalomyelitis) the prognosis is guarded to poor.
Central nervous system trauma Bacterial meningitis Etiology Etiology Bacterial meningitis is a rare complication of sepsis or cranial trauma. Multiple different species of Gram-
Brain trauma most commonly results from a blow to the head over the frontal or parietal bones (Figure 54.3).13 Such injuries often occur as a result of a
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Figure 54.3 Head trauma with signs of asymmetric forebrain injury (image from video). This 2-week-old Quarter Horse filly ran into a post 12 h before the image was recorded. There were contusions and a small laceration above the left eye, but no skull fracture. The foal was obtunded, walked compulsively in large circles to the right, turned its head and neck to the right without tilting its head (as in this image), and had no interest in its dam. With supportive therapy, the foal completely recovered within 24 h.
kick from the dam or other horse. The brain also may be injured when a foal flips over backward (usually while being handled) and strikes the poll or temporal area. Spinal trauma usually is seen in foals over 1 week of age. Vertebrae are fractured or dislocated when galloping foals pitch headfirst into the ground or collide with an immobile object (Figure 54.4). Often these accidents occur at night. In all cases of trauma to the CNS, neurological signs result from both the immediate injury and the subsequent tissue reaction to the injury.13
Figure 54.4 Vertebral fracture in a 4-week-old Thoroughbred colt (image from video). The foal struggled frequently into a dog-sitting position. Thoracic limb reflexes were normal. The withdrawal reflex was intact in the pelvic limbs but the patella reflex could not be elicited. The cutaneous trunci reflex was absent from T16 caudally. Radiographs revealed dislocation of the joints between T15 and T16 vertebrae, and a fracture through the body of L2.
weakness of the limbs. Vertebral fracture/dislocation causes paralysis or weakness and ataxia of the limbs at or caudal to the site of injury. With upper cervical involvement, reluctance to move the head may be the only sign. Diagnosis Radiographic, CT, or MRI imaging is required to document fractures and dislocations. Skull fractures often are difficult to identify on radiographs because of the complex anatomy of the skull; in neonates, interpretation is further complicated by the numerous suture lines and synchondroses evident normally. Contrast myelography is still commonly used to determine spinal cord compression, although MRI and CT eventually will obviate the need for this procedure.
Clinical signs
Treatment and prognosis
There may be skin abrasions, soft-tissue swellings, sensitivity, resistance to palpation of affected areas, bleeding from the nostrils and external ears, or crepitus associated with skull or vertebral fractures. With any head injury, there often is a brief period of unconsciousness (concussion) following the trauma. After a blow to the front of the head, subsequent signs are suggestive of cerebral dysfunction with disorientation, disinterest in the dam, head-pressing, lethargy or stupor, and a tendency to walk compulsively, often in circles. After impact to the poll or temporal area, signs predominantly reflect injury to the brainstem and cranial nerves. There is often profound obtundation, head tilt or facial paralysis, and ataxia and
In addition to general supportive therapy and attention to any skin wounds (including the use of broad-spectrum antimicrobials), the therapies outlined above for HIE may be used in an attempt to suppress the secondary phase of CNS injury. Evidence from a large prospective study in adult humans suggests that corticosteroids are contraindicated in the treatment of traumatic brain injury.14 By contrast, a slight benefit has been demonstrated for treatment of spinal cord injury with high-dose methylprednisolone sodium succinate (30 mg/kg i.v. over 30 min, then 5.4 mg/kg/h for the next 48 h).15 Surgery is indicated to elevate and stabilize bone fragments in depressed fractures of frontal or parietal
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Peripheral nerve injury Etiology
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anti-inflammatory effect can be provided locally by the use of skin-permeant drugs such as DMSO or diclofenac. The signs associated with type 1 nerve injuries and partial nerve tears should resolve in days to several weeks. Regrowth by axonal sprouting from the proximal stump of a type 2-injured nerve should occur at the rate of about 2.5 cm (1 inch)/month.
Equine degenerative myeloencephalopathy (EDM)
Peripheral nerve and plexus injury can occur as a result of forced extraction of the fetus, secondary to injection, or because of soft tissue trauma or fractures. Type 1 injuries cause loss of function (neurapraxia) but axonal continuity is not affected. Type 2 injuries result in disruption of the axon (axonotmesis) or the entire nerve (neurotmesis).
Equine degenerative myeloencephalopathy has been seen in horses of many different breeds.21–23 It is speculated that clinical expression of the disease involves an interaction between a vitamin E-deficient diet and an inherited predisposition to the disease.
Clinical signs
Clinical signs
There is weakness and hyporeflexia of the affected limb, and there may be areas of cutaneous anesthesia. 20 Examples include signs of brachial plexus injury or femoral nerve paralysis after dystocia, and sciatic or tibial nerve paralysis after injections into the muscles of the rump or back of the leg, respectively. After about 2 weeks there may be obvious muscle atrophy.
There is symmetric weakness and ataxia of all limbs that begins any time from birth to 2 years of age. Signs usually are progressive for several months then level out. Cutaneous trunci and cervicofacial reflexes often are poor or absent.
Diagnosis The diagnosis is presumptive based on history and clinical signs. Electrodiagnostic studies, including nerve conduction and needle electromyography (after 10–14 days), can be performed to support the diagnosis and define the nerves involved.
Etiology
Diagnosis The disease may be suspected because of characteristic clinical signs and exclusion of other possible diagnoses such as vertebral malformation or myelodysplasia (Figure 54.5). A supportive finding is demonstration of a serum vitamin E concentration
Treatment and prognosis Wounds associated with nerve injury must be cleaned and debrided, and fractured bones stabilized. If the nerve is severed, the ends should be identified. Anastomosis after epineurial alignment is technically possible although rarely performed in horses. A support wrap or splint should be kept on the affected limb to prevent contracture and protect against further injury. Joints on the affected limb can be kept flexible by passive flexion and extension routines several times daily. The use of faradic or galvanic electrical stimulation of affected muscles may prevent denervation atrophy; however, this therapy is controversial. Acutely, anti-inflammatory treatment should be provided with a cyclooxygenase inhibitor such as flunixin meglumine (1.1 mg/kg i.v. once or twice daily) and DMSO (1 g/kg as a 10% solution i.v. or by nasogastric [NG] tube). Additional
Figure 54.5 Myelodysplasia in a 3-week-old American Paint filly (image from video). Since birth this foal had tail, anal, rectal, and bladder paralysis (cauda equina syndrome) and analgia of the perineal region. The pelvic limbs were incoordinated and moved in a synchronous ‘bunny-hopping’ manner even at walking speed. At necropsy there were signs of incomplete closure of the neural tube caudal to the 7th thoracic spinal cord segment.
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Figure 54.6 Narcolepsy-cataplexy in a 9-day-old Morgan foal (images from video). The sequence of images was recorded during a 5-minute period, beginning in the upper left panel and progressing clockwise. The foal collapsed 4–6 times daily, often while suckling, and lay for several minutes in a completely flaccid state with occasional rapid eye movements. Muscle tone and reflexes returned gradually over several minutes and the foal was clinically normal by the time the lower left image was recorded.
< 1 mg/L.24 Characteristic post-mortem findings are demyelination and axonal degeneration within the white matter of the spinal cord. Treatment and prognosis The disease is incurable, although it is thought that large doses of vitamin E (e.g., 6000 IU p.o. daily) can prevent progression of clinical signs or even effect some slight improvement.23 Supplemental vitamin E likely can prevent EDM in genetically susceptible horses.
deficient production by the hypothalamus of arousal peptides (orexins/hypocretins). Clinical signs Foals may collapse suddenly, usually during restraint or while suckling, then remain laterally recumbent for several minutes in a hypotonic, hyporeflexic state (Figure 54.6). Rapid eye movements may be obvious after collapse. Diagnosis
Narcolepsy/cataplexy Etiology ‘Fainting’ attacks occur as transient events in all breeds of foals and as syndromes involving persistent and repeated episodes in some lines of Shetlands, Suffolks, Morgan Horses, and American Miniature Horses. The cause is not known, but some narcoleptic syndromes in human beings have been traced to
The clinical signs are definitive. A physostigmine test has been used to provoke the signs in American Miniature Horse foals. Treatment and prognosis Most foals have only a few episodes and then grow out of it. In other foals signs persist for months or even for life. There is no treatment.
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Cerebellar abiotrophy Etiology Cerebellar abiotrophy is an inherited disorder of certain lines of Arabian horses and Gotland ponies.25–27 The cerebellum undergoes full development, then degenerates before or shortly after birth.
491
may demonstrate that the cerebellum is small. It is reported that CSF protein concentration is increased in foals during the progressive stage of abiotrophy.26 Post-mortem examination reveals degeneration of Purkinje cells and thinning of the molecular and granular layers of the cerebellar cortex.27 Treatment and prognosis
Clinical signs Signs usually are present at birth or begin within the first few months of life and progress for weeks or months. Affected foals may stand base-wide and sway back and forth at rest. Movements are jerky and dysmetric and foals are hyperresponsive to handling (Figure 54.7). Such foals are extremely difficult to train to accept a halter and be led. They are particularly likely to fall when handled or when changing direction at gaits above the walk. There usually is a coarse head bob that is exaggerated during teat-seeking activity. Although foals with cerebellar abiotrophy can see, they never develop a competent menace response. Diagnosis The breed association and signs of symmetric ataxia without weakness are highly suggestive of the diagnosis. Cross-sectional CT or MR scanning of the brain
Most foals are euthanized for humane reasons. Some mildly affected horses have had a normal lifespan.
Occipitoatlantoaxial malformation (OAAM) Etiology This syndrome occurs sporadically in all breeds but is an inherited disorder of Arabians.28 Signs result from instability and stenosis of the vertebral canal because of anomalies of the occiput, atlas, and axis. Clinical signs Foals may be born dead or comatose. Typically, however, there is tetraparesis and ataxia of all four limbs. Signs range in severity from inapparent to quadriplegia. There may be a head tilt or the head may be held extended or deviated to the side. Movements of the head and neck may elicit obvious clicking sounds. Diagnosis The diagnosis can be made on plain radiographs. Typically, there is fusion of the atlas and occiput and hypoplasia of the dens (Figure 54.8). Treatment and prognosis Mildly affected foals may survive but most are euthanized for humane reasons.
Botulism Etiology
Figure 54.7 Cerebellar abiotrophy. This 4-day-old Arabian colt had been severely ataxic since birth and had a coarse vertical head bob that was exaggerated during approaches to the dam’s udder. As shown in this image, the colt was hyperresponsive to handling, often rearing and falling over backward when attempts were made to handle him.
Botulism is caused by the actions of botulinum toxin, an exotoxin produced by the Gram-positive sporeforming anaerobic bacillus Clostridium botulinum. Most equine cases are caused by C. botulinum types B and C.29, 30 The toxin prevents acetylcholine release at neuromuscular junctions and results in signs of generalized flaccid paralysis. The disease occurs endemically in areas of the eastern USA and sporadically elsewhere. Endemic cases in foals are known as
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Foal: Nervous System breathing is labored. Death occurs within 72 h of the onset and is due to respiratory paralysis. Diagnosis The diagnosis is presumptive based on clinical signs. Hypercapnia is found on blood gas analysis. Definitive diagnosis additionally requires the demonstration of toxin in blood or feces by mouse inoculation or immunoassay.31 Detection of toxin or spores in feed sources or culture of the organism from feces is strongly supportive of the diagnosis of botulism. (a)
Treatment and prognosis
(b) Figure 54.8 Occipitoatlantoaxial malformation (OAAM). Radiographs of the craniocervical region of a normal neonatal foal (a) and a 3-week-old Arabian foal with OAAM (b). Note slightly shortened dens, ventral luxation of the dens, cranioventral movement of the body of the atlas to encroach the guttural pouch, and fusion of the atlas to the occiput with occipitalization of its caudal surface, in the foal with OAAM. Images courtesy of Dr I.G. Mayhew.
shaker foal syndrome and are thought to be due to ingestion of the organism and subsequent colonization of the gastrointestinal tract by C. botulinum type B. Botulism also may occur because of ingestion of preformed toxin or proliferation of the organism in a wound or abscess. Clinical signs Foals may be found dead. Usually, there is a sudden onset of weakness and stilted gait in previously healthy foals.31, 32 An early sign is dribbling of milk from the nose and mouth after suckling. The facial expression is blank, and pupillary dilatation and ptosis may be noted. Within hours of the first signs, there is trembling of triceps and quadriceps muscles, and progression to increasingly prolonged periods of recumbency. Terminally, the neck is extended and
A polyvalent antitoxin made in horses is commercially available and is effective, especially if given early in the course. It is recommended that 200 mL be given intravenously to foals. Foals that become recumbent because of botulism are unlikely to survive without mechanical ventilation.52 Ventilation must be continued for several days up to 2 weeks before foals are able to breathe on their own. Full recovery may take months. Without treatment, more than 90% of affected foals die within 72 hours. Conversely, over 80% of affected foals survive if given antitoxin within several hours of the onset of signs.31 The incidence of endemic botulism has been reduced by vaccination of mares during pregnancy with a type B toxoid.
Tetanus Etiology Tetanus usually occurs in foals older than 7 days and is caused by the actions of exotoxins produced by the Gram-positive, spore-forming anaerobic bacillus Clostridium tetani. Tetanospasmin is released from proliferating C. tetani, circulates to peripheral nerve terminals, then is transported centrally in axons to presynaptic inhibitory neurons. The toxin acts by blocking the release of the neurotransmitter gammaamniobutyric acid.33 Loss of motor neuron inhibition leads to the signs of muscle rigidity, muscle spasm, and autonomic overactivity. The disease has become rare because of the widespread use of vaccines containing tetanus toxoid. Clinical signs The disease is characterized by rapidly progressive muscular rigidity and hyperresponsiveness to stimuli of any kind (Figure 54.9).34, 35 Affected foals may at first appear to have trouble suckling because of
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or failure of passive transfer of immunity to the foal.
Treatment and prognosis
(a)
(b) Figure 54.9 Tetanus in a 10-day-old Quarter Horse foal (images from video). The foal was in lateral recumbency in an extended posture caused by rigidity (tonus) of skeletal muscles (a). Stimulation of the foal (in this case by touching) caused generalized muscle spasms (b).
spasms of the masticatory muscles (trismus) and neck that occur during teat-seeking. Even if the foal can suckle, reflex pharyngeal spasms prevent swallowing and milk is returned from the mouth and nose. Limb and back extension result in a ‘sawhorse’ posture with elevation of the tail and rigid extension of the head and ears. Attempts to move or change direction or gait may provoke muscle spasms and cause the foal to fall. The face often has a wooden anxious expression (risus sardonicus) and there may be intermittent prolapse of the nictitans in response to external stimuli. Sympathetic overactivity may result in tachycardia, fluctuant heart rate, and profuse sweating. Eventually the foal becomes laterally recumbent in a rigidly extended position and is subject to rounds of severe myoclonic contractions. Death is due to respiratory failure or, possibly, the cardiac effects of sympathetic overactivity. Diagnosis Diagnosis is based on clinical signs in the context of a history of poor vaccination of the dam against tetanus
Any wound or abscess should be cleaned, flushed, and debrided to remove the source of toxin. Circulating toxin can be neutralized by intravenously administered antitoxin (arbitrary dose of 5000 units i.v.). Metronidazole (10–25 mg/kg i.v. or p.o. every 12 h) should be given to kill residual organisms. Penicillin is no longer recommended because of potential GABA antagonism.33 The basis of therapy in horses is drug-induced sedation and muscle relaxation. Diazepam and midazolam are effective for shortterm control. Sodium pentobarbital (3–10 mg/kg i.v. every 8 h or by continuous infusion), a GABA agonist, and chlorpromazine (0.04–0.08 mg/kg i.v. or i.m.), an alpha-adrenergic antagonist, are used in combination. Intensive supportive therapy is essential. It is theoretically possible to paralyze and mechanically ventilate an affected foal for the several weeks required for toxin to be removed. Recently, the use of magnesium as sole therapy has been adopted for treatment of human neonatal tetanus in underdeveloped countries.33 Magnesium has the advantage of blocking both neuromuscular transmission and sympathetic overactivity. Although not yet tested for this purpose in horses, the use of continuous infusion of MgSO4 seems logical. Try 0.05 g/kg/h loading dose, then 0.025 g/kg/h. The dose should be titrated to control clinical signs without causing respiratory depression. The fatality rate reported for horses of all ages was 75%.35 Thus, the prognosis is at best guarded for foals that are treated before they become recumbent, and poor for foals that are unable to stand.
References 1. Ojala M, Ala-Huikku J. Inheritance of hydrocephalus in horses. Equine Vet J 1992;24:140–3. 2. Bentz BG, Moll HD. Treatment of congenital hydrocephalus in a foal using a ventriculoperitoneal shunt. J Vet Emerg Crit Care 2008;18:170–6. 3. Vaala WE, House JK. Manifestations of disease in the neonate. In: Smith BP (ed.) Large Animal Internal Medicine. St Louis: Mosby, 2002; pp. 319–86. 4. Divers TJ, Warner A, Vaala WE, Whitlock RH, Acland HA, Mansmann RA, Palmer JE. Toxic hepatic failure in newborn foals. J Am Vet Med Assoc 1983;183:1407–13. 5. Mayhew IG. Observations on vascular accidents in the central nervous system of neonatal foals. J Reprod Fertil Suppl 1982;32:569–75. 6. Palmer AC, Leadon DP, Rossdale PD, Jeffcott LB. Intracranial haemorrhage in pre-viable, premature and full term foals. Equine Vet J 1984;16:383–9.
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7. Wilkins PA. Disorders of foals. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine. St Louis: Saunders, 2004; pp. 1381–431. 8. Hess-Dudan F, Rossdale PD. Neonatal maladjustment syndrome and other neurologic signs in the newborn foal. Equine Vet Educ 1996;8:24–32. 9. Knottenbelt D, Holdstock N, Madigan JE. Neonatal Syndromes In: Equine Neonatology: Medicine and Surgery. Edinburgh: Saunders, 2004; pp. 155–364. 10. Green SL, Mayhew IG. Neurologic disorders. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 496–530. 11. Baker SM, Drummond WH, Lane TJ, Koterba AM. Followup evaluation of horses after neonatal intensive care. J Am Vet Med Assoc 1986;189:1454–7. 12. Chaudhuri A. Adjunctive dexamethasone treatment in acute bacterial meningitis. Lancet Neurol 2004;3:54–62. 13. MacKay RJ Brain injury after head trauma: pathophysiology, diagnosis, and treatment. Vet Clin North Am Equine Pract 2004;20:199–216. 14. Roberts I, Yates D, Sandercock P, Farrell B, Wasserberg J, Lomas G, Cottingham R, Svoboda P, Brayley N, Mazairac ˜ G, Lalo¨e V, Munoz-S´ anchez A, Arango M, Hartzenberg B, Khamis H, Yutthakasemsunt S, Komolafe E, Olldashi F, Yadav Y, Murillo-Cabezas F, Shakur H, Edwards P, CRASH trial collaborators. Effect of intravenous corticosteroids on death within 14 days in 10008 adults with clinically significant head injury (MRC CRASH trial): randomised placebocontrolled trial. Lancet 2004;364:1321–8. 15. Bracken MB, Shephard MJ, Holford TR, Leo-Summers L, Aldrich EF, Fazl M. Administration of methylprednisolone for 24 or 48 hours or tirilazad mesylate for 48 hours in the treatment of acute spinal cord injury. Results of the Third National Acute Spinal Cord Injury Randomized Controlled Trial. National Acute Spinal Cord Injury Study. JAMA 1997;277:1597–604. 16. McIlwraith CW, Robertson JT. Surgical repair of depression fractures of the skull. In: McIlwraith & Turner’s Equine Surgery: Advanced Techniques. Philadelphia: Lippincott, Williams & Wilkins, 1998; pp. 276–7. 17. McCoy DJ, Shires PK, Beadle R. Ventral approach for stabilization of atlantoaxial subluxation secondary to odontoid fracture in a foal. J Am Vet Med Assoc 1984;185:545–9. 18. Nixon AJ, Stashak TS. Laminectomy for relief of atlantoaxial subluxation in four horses. J Am Vet Med Assoc 1988;193:677–82. 19. Owen R, Maxie LL. Repair of fractured dens of the axis in a foal. J Am Vet Med Assoc 1978;173:854–6. 20. Mayhew IG. Physical and chemical causes. In: Large Animal Neurology. Ed 2. Oxford: Wiley-Blackwell, 2009; pp. 294–320. 21. Adams R, Mayhew IG. Neurologic diseases. Vet Clin North Am Equine Pract 1985;1:209–34. 22. Gandini G, Fatzer R, Mariscoli M, Spadari A, Cipone M, Jaggy A. Equine degenerative myeloencephalopathy in five Quarter Horses: clinical and neuropathological findings. Equine Vet J 2004;36:83–5. 23. Mayhew IG, Brown CM, Stowe HD, Trapp AL, Derksen FJ, Clement SF. Equine degenerative myeloencephalopathy: a vitamin E deficiency that may be familial. J Vet Intern Med 1987;1:45–50. 24. Blythe LL, et al. Serially determined plasma alphatocopherol concentrations and results of the oral vitamin E absorption test in clinically normal horses and in
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29.
30.
31. 32.
33. 34. 35.
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38.
39.
40.
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horses with degenerative myeloencephalopathy. Am J Vet Res 1991;52:908–11. Bjorck G, Everz KE, Hansen HJ, Henricson B. Congenital cerebellar ataxia in the gotland pony breed. Zentralbl Veterinarmed A 1973;20:341–54. DeBowes RM, Leipold HW, Turner-Beatty M. Cerebellar abiotrophy. Vet Clin North Am Equine Pract 1987;3: 345–52. Palmer AC, Blakemore WF, Cook WR, Platt H, Whitwell KE. Cerebellar hypoplasia and degeneration in the young Arab horse: clinical and neuropathological features. Vet Rec 1973;93:62–6. Mayhew IG, Watson AG, Heissan JA. Congenital occipitoatlantoaxial malformations in the horse. Equine Vet J 1978;10:103–13. Haagsma J, Haesebrouck F, Devriese L, Bertels G. An outbreak of botulism type B in horses. Vet Rec 1990; 127:206. Kinde H, Bettey RL, Ardans A, Galey FD, Daft BM, Walker RL. Clostridium botulinum type-C intoxication associated with consumption of processed alfalfa hay cubes in horses. J Am Vet Med Assoc 1991;199:742–6. Whitlock RH, Buckley C. Botulism. Vet Clin North Am Equine Pract 1997;13:107–28. Wilkins PA, Palmer JE. Botulism in foals less than 6 months of age: 30 cases ( 1989-2002). J Vet Intern Med 2003;17: 702–7. Attygalle D, Rodrigo N. New trends in the management of tetanus. Expert Rev Anti Infect Ther 2004;2:73–84. Beroza, GA. Tetanus in the horse. J Am Vet Med Assoc 1980;177:1152–4. Blythe LL, Craig AM, Lassen ED, Rowe KE, Appell LH. Tetanus in the horse: a review of 20 cases (1970 to 1990). J Vet Intern Med 1994;8:128–32. Spalding MG, Greiner EC, Green SL. Halicephalobus (Micronema) deletrix infection in two half-sibling foals. J Am Vet Med Assoc 1990;196:1127–9. Acland HM, Mann PC, Robertson JL, Divers TJ, Lichtensteiger CA, Whitlock RH. Toxic hepatopathy in neonatal foals. Vet Pathol 1984;21:3–9. McCornico RS, Duckett WM, Wood PA. Persistent hyperammonemia in two related Morgan weanlings. J Vet Intern Med 1997;11:264–6. Johnson PJ, Mrad DR, Schwartz AJ, Kellam L. Presumed moxidectin toxicosis in three foals. J Am Vet Med Assoc 1999;214:678–80. Fortier LA, Fubini SL, Flanders JA, Divers TJ. The diagnosis and surgical correction of congenital portosystemic vascular anomalies in two calves and two foals. Vet Surg 1996;25:154–60. Gundlach AL, Kortz G, Burazin TC, Madigan J, Higgins RJ. Deficit of inhibitory glycine receptors in spinal cord from Peruvian Pasos: evidence for an equine form of inherited myoclonus. Brain Res 1993;628: 263–70. Loynachan AT, Williams NM, Freestone JF. Kernicterus in a neonatal foal. J Vet Diagn Invest 2007;19:209–12. Page P, Parker R, Harper C, Guthrie A, Neser J. Clinical, clinicopathologic, postmortem examination findings and familial history of 3 Arabians with lavender foal syndrome. J Vet Intern Med 2006;20:1491–4. Ward TL, Valberg SJ, Adelson DL, Abbey CA, Binns MM, Mickelson JR. Glycogen branching enzyme (GBE1) mutation causing equine glycogen storage disease IV. Mamm Genome 2004;15:570–7.
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Neurological Disorders in the Neonatal Foal 45. Leaman TR. Nettle reaction in a foal. Vet Rec 2008;162:164. 46. Lofstedt J. White muscle disease of foals. Vet Clin North Am Equine Pract 1997;13:169–85. 47. McCabe L, Griffin LD, Kinzer A, Chandler M, Beckwith JB, McCabe ER. Overo lethal white foal syndrome: equine model of aganglionic megacolon (Hirschsprung disease). Am J Med Genet 1990;36:336–40. 48. Borchers A, Magdesian KG, Halland S, Pusterala N, Wilson WD. Successful treatment and polymerase chain reaction (PCR) confirmation of Tyzzer’s disease in a foal and clinical and pathologic characteristics of 6 additional foals (19862005). J Vet Intern Med 2006;20:1212–18. 49. Fox J, Duncan R, Friday P, Klein B, Scarratt W. Cerebelloolivary and lateral (accessory) cuneate degeneration in a juvenile American Miniature horse. Vet Pathol 2000;37:271–4. 50. Loomis EC, Bushnell RB. Tick paralysis in California livestock. Am J Vet Res 1968;29:1089–93.
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51. Triplett EA, O’Brien RT, Wilson DG, Steinberg H, Darien BJ. Thrombosis of the brachial artery in a foal. J Vet Intern Med 1996;10:330–2. 52. Moore LA, Johnson PJ, Bailey KL. Aorto-iliac thrombosis in a foal. Vet Rec 1998;142:459–62. 53. Cho DY, Leipold HW. Syringomyelia in a thoroughbred foal. Equine Vet J 1977;9:195–7. 54. Jacobsen B, Venner M, Gerdwilker A, Wohlsein P. Ventral meningomyelocele in a German warmblood foal. Dtsch Tierarztl Wochenschr 2007;114:470–72. 55. Vaala WE, House JK. Perinatal adaptation, asphyxia, and resuscitation, In: Smith BP (ed.) Large Animal Internal Medicine. St Louis: Mosby, 2002, pp. 266–76. 56. Wilkins PA, Palmer JE. Mechanical ventilation in foals with botulism: 9 cases (1989–2002). J Vet Intern Med 2003;17:708–12.
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55.
Techniques in Diagnosis and Monitoring of Cardiovascular Disease Anne Desrochers
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Congenital Abnormalities of the Cardiovascular System Celia M. Marr
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Acquired Abnormalities of the Cardiovascular System C.J. (Kate) Savage
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Chapter 55
Techniques in Diagnosis and Monitoring of Cardiovascular Disease Anne Desrochers
Introduction The primary function of the cardiovascular system is the rapid delivery of nutrients and oxygen to tissues and organs. In addition, the cardiovascular system plays a crucial role in waste product elimination, hormone distribution, immune function, coagulation, and body temperature regulation. The importance of this vital system cannot be overstated and patients with suspect cardiovascular dysfunction should be promptly evaluated. The purpose of this chapter is to review the current available information regarding the techniques for diagnosing and monitoring cardiovascular disease in foals.
Cardiovascular evaluation Background information The signalment of the patient and a detailed history regarding management practices, diet, vaccination status, deworming program, previous medical problems, and the nature and duration of the current clinical signs can all provide valuable information for the initial assessment of foals with suspected cardiovascular disease. Physical examination A complete investigation of the cardiovascular system must include evaluation of mucous membrane color and capillary refill time, arterial pulse quality, jugular vein refill and pulsation, and presence of peripheral venous distention, and dependent edema. Careful assessment of the respiratory system should
also be performed to detect concurrent pulmonary involvement or disease. Normal mucous membranes should be pink when evaluated in good lighting and capillary refill time should be less than 2 seconds. Pale mucous membranes and prolongation of the capillary refill time are consistent with decreased peripheral tissue perfusion. Cyanosis is more often seen with severe respiratory disease resulting in hypoxia but can also develop with congenital cardiac conditions causing right-toleft shunting of blood, thereby bypassing pulmonary oxygenation. Normal jugular vein filling and pulses should not extend above one-fourth to one-third of the distal portion of the neck, provided the neck is in normal upright position. Jugular vein distension and higher pulses reflect increased right atrial pressure and suggest presence of congestive heart failure, tricuspid regurgitation, tetralogy of Fallot, or atrial septal defect. Normal peripheral arterial pulses should be strong and regular on palpation. Irregular arterial pulses often indicate the presence of a cardiac arrhythmia such as atrial fibrillation. Bounding pulses, reflecting large difference in pressure between systole and diastole, are palpable in patients suffering from left-to-right shunt such as patent ductus arteriosus and ventricular septal defect or from severe aortic insufficiency, although rare in foals. Weak peripheral pulses are detectable in congestive heart failure or conditions causing cardiovascular collapse such as septic or hypovolemic shock and severe hemorrhage. Poor peripheral perfusion secondary to low cardiac output causes cold extremities, particularly legs and ears, as blood is shunted centrally. Pectoral, ventral abdominal or limb edema develops secondary
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Cardiovascular System
to poor cardiac output in patients suffering from congestive heart failure. Tachypnea, increased bronchovesicular sounds, and nostril flair can be present when interstitial pulmonary edema develops as a consequence of marked elevation in left atrial pressure secondary to left-sided heart failure. If the pulmonary edema involves the alveoli, respiratory distress, fluidy lung and tracheal sounds as well as white frothy fluid visible from the nares or expelled from the mouth can develop. In foals, non-specific clinical signs of cardiac disease include exercise intolerance, slow growth rate, lethargy, weakness, collapse, and sudden death. These signs usually develop as consequences of congestive heart failure or complex congenital defects. Cardiac auscultation Cardiac auscultation allows for the determination of heart rate, cardiac rhythm, and presence of murmur(s). Both sides of the thorax and all valve areas should be carefully auscultated in a quiet environment.
Heart rate and rhythm Because significant change in heart rate can occur following variation in the autonomic nervous system input and level of physical activity, it is important to measure heart rate under resting state when muscular activity and environmental disturbances are minimal. Normal resting heart rates in foals of different ages and breeds have been published.1–5 Variations from 91 ± 15 beats per minute (bpm) at 1 day old, 88 ± 27 bpm at 1 week old, 77 ± 12 bpm at 1 month to 69 ± 13 bpm at 3 months old were recorded in a study evaluating full-term healthy pony and Thoroughbred foals.3 The normal cardiac rhythm of foals is regular on auscultation. A suitable period of time should be spent during the examination for the detection of sustained and intermittent arrhythmias.
Cardiac murmurs Murmurs are manifestations of normal or abnormal blood flow in the heart or blood vessels which develop during a normally silent period of the cardiac cycle. Murmurs may be associated with increased velocity of blood flow or decreased blood viscosity. When present, cardiac murmurs should be classified according to their grade or intensity, timing, duration, shape, character, point of maximal intensity, radiation, and palpation of a precordial thrill. This characterization can help to differentiate physiological from pathological murmurs and identify foals requiring additional diagnostic investigations. Physiological murmurs are extremely common in foals early in life and are generally caused by the nor-
mal transition from fetal to pediatric circulation and by the vibrations produced by the ejection of blood during systole or during the rapid filling of the ventricles in early diastole. Typical physiological murmurs are soft (grade 1–3/6), localized, short in duration, and vary in intensity with heart rate and body position. In foals, certain pathological conditions may lead to the development of physiological murmurs and include fever, sepsis, moderate to severe anemia, and conditions causing peripheral vasodilation or high sympathetic activity such as colic and other causes of pain. The continuous machinery murmur of patent ductus arteriosus audible over the left base of the heart is considered a normal finding in fullterm foals. At birth, the ductus arteriosus normally closes in response to increased local oxygen tension and inhibition of prostaglandins.6 The murmur typically disappears within the first 4 days of life.2, 6 However, the systolic component occasionally persists for a longer period of time after birth.1, 7 Pathological murmurs are less common and are usually associated with the presence of patent remnants of the fetal circulation, congenital cardiac malformations, and acquired pathological conditions. The presence of a grade 3/6 or higher holosystolic or pansystolic murmur, multiple systolic murmurs, holodiastolic murmur, bilateral murmurs, persistent continuous machinery murmur, widely radiating murmur associated with a precordial thrill, or pericardial friction rubs and muffled heart sounds can all be indicative of cardiac pathology.
Ancillary tests Several auxiliary tests can provide indirect but useful additional information about the cardiovascular disease processes and their effects. Laboratory studies Complete blood count, fibrinogen level, and serum amyloid A concentration can help to document the presence of inflammation, infection, and anemia. Biochemistry profile and blood gas analysis allow assessment of electrolyte levels, acid–base status, as well as pulmonary, hepatic, and renal function. Blood abnormalities may reflect the presence of metabolic disorders and can lead to the development of cardiovascular diseases such as cardiac arrhythmias secondary to electrolyte imbalances. Conversely cardiovascular disorders may affect blood parameters. For instance, azotemia may result from poor renal perfusion due to low cardiac output and persistent PaO2 lower than 100 mmHg despite nasal insufflation of 100% oxygen suggests the presence of a right-to-left cardiac shunt. Blood cultures are indicated when an
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Techniques in Diagnosis and Monitoring of Cardiovascular Disease infective etiology is suspected, such as in cases of bacterial endocarditis. Urine output monitoring, urinalysis, and urine chemistry can provide valuable information about renal function and perfusion. Biomarkers of myocardial cell injury such as cardiac troponins (I and T) and isoenzymes of creatine kinase (CKMB) can be released in the circulation and blood levels can increase in cases of myocardial inflammation, toxicosis, and necrosis.8, 9 Reported values from healthy neonates using mouse monoclonal antibodies immunoassays varied from 0.01 to 0.51 ng/mL and 0.009 to 0.041 ng/mL for cTnl and cTnT respectively.8 Normal CKMB values varied between 0.40 to 9.3 ng/mL.8 Another reference indicates that cTnI values greater or equal to 1 ng/mL are highly suggestive of myocardial disease.10 Rarely, vitamin E and selenium deficiency can lead to myocardial degeneration in foals. Vitamin E (alpha-tocopherol) and selenium blood concentrations as well as glutathione peroxidase activity can help to reach a definitive antemortem diagnosis.11 Adequate glutathione peroxidase activity in horses should be between 20 and 50 U/mg of hemoglobin/minute.12 Whole blood selenium concentrations ranging from 0.07 to greater than 0.1 ppm (µg/g)13, 14 are considered normal in horses and normal alpha-tocopherol level should be greater than 1.1–2.0 ppm (µg/g).12, 14 Other specific toxicological screening tests are recommended when exposure to cardiac toxins such as oleander and ionophores is suspected.15, 16 Radiography Thoracic radiographs can be helpful in the estimation of heart size and evaluation of the pleural space, lower airways, pulmonary parenchyma, and lung vasculature. Standing and recumbent lateral views of the thorax can be obtained with portable radiographic equipment. With appropriate sedation, a ventrodorsal view can also be performed in neonates. The heart occupies a greater proportion of the thoracic cavity in foals compared to adults. In neonates, normal maximal craniocaudal and apicobasilar cardiac dimensions should not exceed 5.6 to 6.3 times and 6.7 to 7.8 times the length of a mid-thoracic vertebral body respectively.17 Marked enlargement of the cardiac silhouette may cause dorsal displacement of the trachea or rounded appearance of the heart and can be associated with primary cardiac or pericardial disease (Figure 55.1). Increased number and size of the pulmonary vessels are suggestive of pulmonary hyperperfusion due to congestive cardiac disease or presence of a left-to-right shunt such as ventricular septal defect or patent ductus arteriosus. Pulmonary hyperperfusion may lead to the development of interstitial to alveolar edema characterized
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Figure 55.1 Right lateral thoracic radiograph showing cardiac silhouette enlargement, dorsal displacement of the trachea, and evidence of pulmonary edema.
by increased lung opacity with or without presence of air bronchograms (Figure 55.1). Decreased lung vascular markings can result from poor pulmonary perfusion. Tetralogy of Fallot, pulmonary stenosis, pulmonary valve hypoplasia or atresia, and neonatal pulmonary hypertension should be considered in those cases.
Specific diagnostic aids in equine cardiology In-depth evaluation of foals suffering from cardiovascular disease typically requires more advanced and accurate diagnostic aids to determine the underlying dysfunction and to recommend appropriate monitoring and treatment. Electrocardiography Electrocardiography (ECG) is a valuable tool for the determination of heart rate, cardiac rhythm, and conduction times. Unlike in small animals, ECGs can not be reliably used to detect chamber enlargement in equine medicine. Several lead systems have been found to be useful in equine medicine and lead placements for these systems are described in previous literature.18, 19 The base apex lead system is commonly used to record ECG in foals. To apply base apex leads, the (+) left arm electrode is positioned over the left cardiac apex on the left hemithorax at the level of and just caudal to the olecranon. The (−) right arm electrode is placed at the top of the right scapular spine or two thirds of the way down the jugular groove on the right. The left leg electrode (ground) is placed in a remote position away from the heart. The ECG is recorded from lead 1 on the ECG
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Foal: Cardiovascular System electrocardiography and may be the result of underlying myocardial disease or secondary to systemic illness. Potential causes of rhythm disturbance in foals include hypoxic-ischemic syndrome, sepsis, trauma (fractured ribs), and electrolyte imbalances such as hyperkalemia. Echocardiography
Figure 55.2 24-hour Holter monitor.
machine. A paper speed of 25 mm/s and amplitude of 1 cm/mV are typically used for ECG recordings in horses.9 The speed and calibration can be adjusted to optimize complex size and to compensate for normally higher heart rates observed in foals. Recumbency during ECG recordings is often preferable to a standing position in foals in order to limit interference from muscle tremors and movement. The electrodes can be attached to the skin using contact or adhesive electrodes. The use of alcohol or ultrasound coupling gel can help to improve electrical contact between electrode and skin. Additional modalities, including 24-hour Holter monitoring (Figure 55.2) and radiotelemetry, allow for the acquisition of continuous electrocardiographic recordings. These devices are used in hospital settings for identification and monitoring of infrequent rhythm disturbances, quantifying the severity of an arrhythmia, or evaluation of the efficacy of drug therapy. In newborn foals, transient cardiac arrhythmias are common after birth and usually disappear without treatment within the first 15 to 20 minutes of life.4, 5 Most of these arrhythmias are considered normal during the adaptive period to extrauterine life and are attributed to high vagal tone and transient physiological hypoxia at birth.4, 5 Persistent abnormal cardiac rhythm should be characterized by
Echocardiography can provide important information in the diagnosis of congenital heart defects, cardiac murmurs, myocardial and pericardial disease, heart failure, and cardiac arrhythmias. Echocardiography can also help to determine the severity of a condition, estimate prognosis for life and athletic performance, and monitor response to therapy. It is a non-invasive, highly accurate technique which has no known risks or side effects. Two-dimensional and M-mode echocardiography techniques produce images of the heart in realtime motion. These modalities are used to assess valvular and myocardial structure and function, to evaluate the anatomy, dimensions, and shape of the cardiac chambers and great vessels, and to identify pericardial disease and congenital abnormalities. In foals, Doppler ultrasonography techniques are mainly used to detect intra- or extracardiac shunts and to characterize valvular insufficiency. Various forms of Doppler ultrasonography exist. Pulsed-wave echocardiography can detect the precise location of low velocity blood flow. Colorflow Doppler provides information about the location, direction, and velocity of abnormal blood flow. Continuous-wave Doppler echocardiography helps to determine the peak velocity of blood flow and estimate pressure gradients between cardiac chambers. Intravenous contrast-enhanced ultrasonography can be performed by injecting gas-filled microbubbles into the venous system to improve tissue and blood delineation. This technique can aid in confirming the presence of congenital abnormalities and valve dysfunction. Although transthoracic echocardiography can be performed in foals in lateral or sternal recumbency, it is easier to execute and interpret in a standing position (Figure 55.3). Sedation should be avoided since its use can alter some cardiac dimensions and indices of function. If a sedative agent is required, acepromazine maleate would be the most appropriate drug to use since it has no significant effect on cardiac function and echocardiographic measurements of heart dimensions.20 Many commercially available ultrasonographic machines, preferably equipped with ECG leads and ECG display, can be used. Low-frequency (3.5 or 5.0 MHz) sector, phased array, or small footprint curvilinear transducers provide adequate penetration and image resolution.
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for unanesthetized foals of different ages, sizes, and breeds have been published1, 22, 23 (Table 55.2). The values of arterial pressure are not static, but undergo natural variations between heartbeats and throughout the day in a circadian rhythm. Blood pressures also change in response to stress, nutritional factors, drugs, or disease. Arterial blood pressure is a basic hemodynamic index representing perfusion and oxygen delivery to tissues. In foals, primary cardiovascular disease and systemic illness such as sepsis and hypoxic-ischemic encephalopathy may result in marked hemodynamic alterations. Arterial blood pressure measurement is utilized in critical care situations to help guide treatment and monitor the effects of intravenous fluids, vasopressors, and inotropic agents on the systemic circulation. In-depth discussions of blood pressure monitoring in critically ill foals has been published.24–27
Figure 55.3 Standing transthoracic echocardiography.
In foals, a depth setting of 12 to 20 cm should provide adequate visualization of the heart and great vessels. The examination should be performed from the right (intercostal space 4) and left (intercostal spaces 3 to 5) parasternal windows. All standard longand short-axis views of the heart and great vessels should be obtained. A more detailed descriptions of the echocardiographic recording techniques have been previously given.1, 3, 21 In addition, the reference values of cardiac dimensions and functional indices from foals of different ages and breeds have been published1, 3 (Table 55.1).
Arterial blood pressure monitoring Blood pressures are lower in neonates and rapidly rise during the first month of life to reach the normal adult ranges.1, 22 Normal blood pressure values
Direct blood pressure measurement Peripheral arterial blood pressure is most accurately measured invasively by placing an indwelling intra-arterial catheter. The facial, radial, and dorsal metatarsal arteries are among the most common sites for cannulation in foals. The catheter is attached to a sterile, fluid-filled tubing system, which is connected to a zeroed and calibrated electronic pressure transducer. The transducer converts pressure energy into electrical signals which are used to generate a pressure waveform on an oscilloscope. The advantage of this system is that it allows real-time and continuous monitoring of a patient’s arterial blood pressure. Arterial catheterization is infrequently associated with complications such as thrombosis, infection, embolization, hemorrhage, and hematoma formation. To avoid complications and dislodgement
Table 55.1 Normal ranges of selected cardiac dimensions in 16 healthy pony and thoroughbred foals from 1 day to 3 months of age3 Age
1 day
1 week
1 month
3 months
Weight (kg) LVEDD (cm) LVESD (cm) D(%) RVEDD (cm) AoRD (cm) LAD (cm)
46.96 ± 8.98 5.81 ± 0.53 4.67 ± 0.56 21.33 ± 2.93 2.37 ± 0.40 3.70 ± 0.25 3.01 ± 0.70
56.89 ± 10.05 6.50 ± 0.53 5.50 ± 0.41 20.08 ± 5.01 2.61 ± 0.27 3.61 ± 0.29 3.23 ± 0.50
80.66 ± 13.67 7.40 ± 0.65 6.52 ± 0.11 16.25 ± 1.77 2.64 ± 0.34 4.19 ± 0.21 3.52 ± 0.92
111.83 ± 16.80 7.76 ± 0.65 6.92 ± 0.88 21.06 ± 1.37 2.74 ± 0.23 4.34 ± 0.32 3.82 ± 0.60
LVEDD, left ventricular end-diastolic dimension. LVESD, left ventricular end-systolic dimension. D, fractional shortening. RVEDD, right ventricular end-diastolic dimension. AoRD, aortic root dimension. LAD, left atrial dimension.
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Table 55.2 Normal arterial blood pressure values from healthy foals using direct and indirect methods of measurement No. of foals
Age (days)
Breed
Technique
Systolic (mmHg)
Diastolic (mmHg)
Mean (mmHg)
12 23 12 3 21 6 12 12 12 12
1 1 1 1 3 6 7 14 30 90
Pony TB Pony TB/QH TB TB/QH Pony Pony Pony Pony
Doppler Osc Osc Direct Osc Direct Doppler Doppler Doppler Doppler
81 ± 10 144 ± 15 128 ± 17 NA 155 ± 12 NA 104 ± 21 100 ± 16 114 ± 21 111 ± 15
35 ± 7 74 ± 9 64 ± 10 NA 80 ± 11 NA 40 ± 14 41 ± 7 54 ± 13 57 ± 19
50 ± 81 95 ± 1322 85 ± 1022 85 ± 2.523 101 ± 922 96 ± 623 61 ± 151 61 ± 61 74 ± 111 75 ± 161
TB, Thoroughbred. QH, Quarterhorse. NA, not available. Doppler, Doppler technique. Osc, oscillometric technique. Direct, direct arterial blood pressure measurement.
during normal motion and manipulation of recumbent foals, the arterial catheter must be carefully secured and protected. Due to its invasiveness and difficulty in maintaining catheter placement in unanesthetized foals, this method is not routinely used. In intensive care units, continuous direct monitoring is generally reserved for patients where rapid variations in arterial pressure are anticipated and for foals receiving therapy for cardiovascular support capable of inducing sudden hemodynamic changes. Direct measurements are also indicated in cases where severe hypotension and vasoplegia result in inaccurate indirect blood pressure monitoring.28, 29
Indirect blood pressure measurement Alternative non-invasive blood pressure values are simpler and faster to obtain than invasive measurements, require less expertise, and have virtually no complications. The non-invasive measurements can be obtained from indirect techniques including manual or automatic auscultatory methods and via ultrasonic Doppler techniques. Because of the frequent need for prolonged monitoring of blood pressure among critically ill patients, automated oscillometric measurements are commonly used in intensive care units. Oscillometric methods use an electronic pressure sensor able to detect flow-induced oscillations in the arterial wall. The transducer is located in a cuff encircling the patient’s appendage (Figure 55.4). In foals, the most accessible locations for cuff placement are over the median, coccygeal, or dorsal metatarsal arteries. The cuff is electronically inflated and then slowly released. When the pressure decreases in the expanded cuff, the artery starts to emit pulsations. The systolic blood pressure is measured near the ini-
tial increase in oscillations, the mean arterial pressure at the point of maximal amplitude of the oscillations, and the diastolic pressure at the lowest point, just before oscillations cease.29 The blood pressure values are recorded and are digitally or graphically displayed on the monitor of the machine. Inaccurate indirect measurements often result from inappropriate cuff selection and placement. Adequate bladder width-to-girth ratio is essential to minimize errors. To improve accuracy, the cuff should encircle at least 80% of the circumference of the leg or tail and the cuff width should be at least 40% of the appendage’s circumference.26 Undersized cuffs yield too high values, whereas oversized cuffs result in too low readings.27 Blood pressures should be measured in lateral recumbency during a quiet or sleep state and be obtained with minimal physical restraint. Consistent cuff size and location should be used for each consecutive measurement. Serial measurements should be performed with sufficient time for total cuff deflation between readings until consistent results are obtained.
Figure 55.4 Indirect blood pressure measurement. Photograph courtesy of Dr Jonathan Palmer.
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Techniques in Diagnosis and Monitoring of Cardiovascular Disease Blood flow to tissues and organ perfusion are crucially dependent on mean arterial blood pressure.30 The mean arterial pressure is sensed by baroreceptors located in the carotid sinuses and the arch of the aorta. The role of these receptors is to control the arterial pressure by adjusting heart rate and arteriolar vessel radius. The autoregulation of blood supply in vital organ systems such as the kidneys, heart, and brain is dependent upon maintenance of an adequate mean arterial pressure. The mechanism of autoregulation consists of the automatic adaptation of the radius of the arterioles to maintain constant blood flow over a wide range of mean pressures in order to protect organ function.31 Maintaining a mean arterial pressure above 60–65 mmHg is critical.30, 32 Below this value autoregulatory control ceases and organ perfusion becomes pressure dependent. However, therapeutic decisions should not be made based solely on low mean arterial pressure results but in association with the presence of other signs of hypoperfusion such as cold extremities, high heart rate, weak arterial pulses, pale mucous membranes, prolonged capillary refill time, abnormal mentation, decreased urine production, metabolic acidosis, high lactate concentration, or other signs of poor organ perfusion.
Advanced hemodynamic diagnostic techniques Cardiac catheterization, angiocardiographic studies, and advanced methods of cardiac output measurements may be of additional benefit in obtaining a diagnosis, establishing a prognosis, and guiding therapy in cases suffering from cardiovascular disease. These tests are usually limited to specialty practices and referral institutions which have sophisticated equipment and experienced personnel capable of interpreting results. However, the need for invasive methods of investigation has decreased, as the majority of the required information can now be obtained non-invasively, safely, and more readily with two-dimensional and Doppler echocardiography.
Conclusion The information gained from thorough clinical assessment combined with judicious selection of adjunctive diagnostic aids can lead to prompt and accurate diagnosis of cardiovascular system dysfunctions. Once a specific diagnosis has been made, formulation of appropriate management strategies including the monitoring of disease progression and response to treatment can have a significant positive impact on patient outcome.
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References 1. Lombard CW, Evans M, Martin L, Tehrani J. Blood pressure, electrocardiogram and echocardiogram measurements in the growing pony foals. Equine Vet J 1984;16(4):342–7. 2. Rossdale PD. Clinical studies on the newborn Thoroughbred foal: II. Heart rate, auscultation and electrocardiogram. British Vet J 1967;123:521–31. 3. Stewart JH, Rose RJ, Barko AM. Echocardiography in foals from birth to three months old. Equine Vet J 1984;16(4): 332–41. 4. Yamamoto K, Yasuda J, Too K. Arrhythmias in newborn Thoroughbred foals. Equine Vet J 1992;23(3):169–73. 5. Yamamoto K, Yasuda J, Too K. Electrocardiographic findings during parturition and blood gas tensions immediately after birth in Thoroughbred foals. Jpn J Vet Res 1991;39:143–57. 6. Machida N, Yasuda J, Too K, Kuno N. A morphological study on the obliteration processes of the ductus arteriosus in the horse. Equine Vet J 1988;20(4):249–54. 7. Livesey LC, Marr CM, Boswood A, Freeman S, Bowen IM, Corley KTT. Auscultatory and two-dimensional, M mode, spectral and colour flow Doppler echocardiographic findings in pony foals from birth to seven weeks of age. J Vet Intern Med 1998;12:255–67. 8. Slack JA, McGuirk SM, Erb HN, Lien L, Coombs D, Semrad SD, Riseberg A, Marques F, Darien B, Fallon L, Burns P, Murakami MA, Apple FS, Peek SF. Biochemical markers of cardiac injury in normal, surviving septic, or nonsurviving septic neonatal foals. J Vet Intern Med 2005;19(4):577–80. 9. Durango, MM, Young LE. 2003. Cardiovascular examination and diagnostic techniques. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. Philadelphia:Saunders, 2003; pp. 572–85. 10. Voss E. The cardiovascular system. In: McAuliffe SB, Slovis NM (eds) Color Atlas of Diseases and Disorders of the Foal. Philadelphia:Elsevier Saunders,2008; pp. 189–211. ¨ 11. Lofstedt J. White muscle disease of foals. Vet Clin North Am: Equine Pract 1997;13(1):169–85. 12. Mass J, Parish SM, Hodgson DR, Valberg SJ. Diseases of muscle. In: Smith BP (ed.) Large Animal Internal Medicine, 3rd edn. St Louis:Mosby, 2002; pp. 1279–82. 13. Moore RM, Kohn CW. Nutritional muscular dystrophy in foals. Comp Cont Educ Pract Vet 1991;13:476–86. 14. McAuliffe SB. The muscular system. In: McAuliffe SB, Slovis NM (eds) Color Atlas of Diseases and Disorders of the Foal. Philadelphia:Elsevier Saunders, 2008; pp. 212–23. 15. Hall JO. Toxic feed constituents in the horse. Vet Clin North Am Equine Pract 2001;17(3):479–89. 16. Hughes KJ, Dart AJ, Hodgson DR. Suspected Nerium oleander (Oleander) poisoning in a horse. Aust Vet J 2002;80(7):412–15. 17. Butler JA, Colles CM, Dyson SJ, Kold SE, Poulos PW. The thorax. In: Butle JA, Colles CM, Dyson SJ, Kold SE, Poulos PW (eds) Clinical Radiology of the Horse, 2nd edn. Iowa:Blackwell Science, 2000; pp. 483–528. 18. Patteson MW. Diagnostic aids in equine cardiology. In: Patteson MW (ed.) Equine Cardiology. Cambridge:Blackwell Science, 1996; pp. 70–115. 19. Marr CM. Electrophysiology and arrhythmogenesis. In: Marr CM (ed.) Cardiology of the Horse. Philadelphia: Saunders, 1999; pp. 51–69. 20. Buhl R, Ersbøll AK, Larsen NH, Eriksen L, Koch J. The effects of detomidine, romifidine or acepromazine on
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echocardiographic measurements and cardiac function in normal horses. Vet Anaesth Analg 2007;34(1):1–8. Bonagura JD, Blissitt KJ. Echocardiography. Equine Vet J 1995; Suppl.19;5–17. Franco RM, Ousey JC, Cash RSG. Study of arterial blood pressure in newborn foals using an electronic sphygmomanometer. Equine Vet J 1986;18(6):475–8. Madigan JE. 1986. Cardiopulmonary function in normal newborn foals from 1–14 days of age. In: Proceedings of the ACVIM, 1986; pp. 101–16. Vaala WE, House JK. Supportive care of the abnormal newborn. In: Smith BP (ed.) Large Animal Internal Medicine, 3rd edn. St. Louis:Mosby, 2002; pp. 294–302. Vaala WE, Webb AI. Cardiovascular monitoring of the critically ill foal. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia:Lea & Febiger, 1990; pp. 262–272. Palmer J. Recognition and resuscitation of the critically ill foal. In: Paradis MR (ed.) Equine Neonatal Medicine. Philadelphia:Elsevier Saunders, 2006; pp. 121–34.
27. Magdesian GK. Monitoring the critically ill equine patient. Vet Clin North Am: Equine Pract 2004; 20:11–39. 28. Henneman EA, Henneman PL. Intricacies of blood pressure measurement: Reexamining the rituals. Heart Lung 1989;19:263–73. 29. Bridges MEJ, Middleton R. Direct arterial vs oscillometric monitoring of blood pressure: stop comparing and pick one (a decision-making algorithm). Crit Care Nurse 1997;17(3):58–72. 30. Hollenberg SM, Ahrens TS, Astiz MD, Chalfin DB, Dasta JF, Heard SO, Martin C, Susla GM, Vincent JL. Practice parameters for hemodynamic support of sepsis in adult patients. Crit Care Med 1999;27:639– 60. 31. McGhee BH, Bridges MEJ. Monitoring arterial blood pressure: what you may not know. Crit Care Nurse 2002;22(2):60–79. 32. Gigu`ere S, Knowles HA, Valverde A, Bucki E, Young L. Accuracy of indirect measurement of blood pressure in neonatal foals. J Vet Intern Med 2005;19:517–76.
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Chapter 56
Congenital Abnormalities of the Cardiovascular System Celia M. Marr
Introduction Congenital abnormalities of the equine cardiovascular system are uncommon, both in surveys of pathological findings of foals1–4 and in surveys of cases with clinical evidence of cardiovascular disease.5 Congenital cardiac disease can be classified as simple or complex anomalies based on the number of specific anatomical derangements that are present, with simple defects being the more prevalent. Simple defects such as ventricular or atrial septal defects may not produce clinical signs for many months or years after birth, whereas complex congenital cardiac disease will often produce significant hemodynamic compromise, and consequently overt clinical signs of cardiac disease early in life. The diagnosis of congenital cardiac disease relies on the identification of anatomical derangements and, thus, echocardiography is the main tool used in its diagnosis.5–10 Contrast echocardiography,11 survey and contrast radiography,12–14 and cardiac catheterization5 may provide further characterization of structural abnormalities and their hemodynamic consequences. Surgical correction of cardiac anomalies is rarely, if ever, undertaken in equine patients; therefore, the main emphasis in case management in equine congenital cardiac disease is in providing the most accurate prognosis possible by careful evaluation of the full range of anatomical derangements present in individual cases.
Ventricular septal defect Ventricular septal defect (VSD) is the most common form of congenital cardiac abnormality that occurs in the horse. Defects can be single or multiple, and the most common location is in the membranous or
non-muscular portion of the septum immediately below the aortic valve (Figure 56.1). VSDs can also be located in the right ventricular outflow tract and the muscular septum.15, 16 In the latter location, VSDs may take the form of multiple channels or defects. Foals with small VSDs in the typical location will often show no overt clinical signs until well into adulthood. The hemodynamic impact of a VSD is dependent on its size and this, in turn, influences the severity of the presenting and clinical signs and the prognosis. It is important to recognize that a VSD is often a component of complex cardiac disease and the presence of additional anatomical derangements will increase significantly the likelihood of severe cardiac compromise. Consequently, clinical evaluation of horses with VSDs should include a careful search for additional lesions such as pulmonic valve pathology,17, 18 over-riding aorta, and atrial septal defects19 as well as the more complex anomalies such as tetralogy of Fallot.5 The VSD is typically associated with at least two murmurs. The first, due to the intracardiac shunt through the VSD from left to right, is loudest over the right sternal border and is usually panasytolic and coarse in nature. The second, the murmur of relative pulmonic stenosis, is due to the excessive quantity of blood leaving the right side of the heart as a result of the left to right intracardiac shunt. This murmur is loudest over the pulmonic valve, in the left third intercostal space, and is typically one grade less loud than the VSD murmur, holosystolic, and crescendo–decrescendo in nature. When a different configuration of murmurs is present, clinicians should suspect that the VSD may be in an atypical location or that there may be additional anomalies present. With large subaortic defects, the aortic valve may become diseased; this can be associated with a
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 56.1 A pathological specimen demonstrating the most common location for a ventricular septal defect (VSD) in the membranous septum immediately beneath the aortic valve.
murmur of aortic regurgitation, occurring in diastole, and usually decrescendo in nature. It may be loudest on either side of the thorax, although this is most often the left. The size of a VSD can be determined most accurately with echocardiography (Figure 56.2). Doppler echocardiography can be used to estimate the pressure gradient between the left and right ventricle. With smaller VSDs, flow across the defect is restricted, a large pressure gradient is maintained between the left and right ventricles, and the speed of the intracardiac shunt will usually exceed 4 m/s.20 In Thoroughbreds and Standardbreds, membranous defects of less than 2.5 cm are generally well tolerated; these breeds with such a defect may be successful at low levels of competition, although they are unlikely to be elite athletes.20 In other breeds, it is more helpful to compare the size of the defect with that of the aorta: restrictive defects will typically be less than one-third the diameter of the aorta.5, 21 These guidelines apply specifically to horses with a VSD in the subaortic location and guidelines for assessment of a VSD in other locations are not available currently.
Atrial septal defect Atrial septal defects (ASDs) are uncommon: they occur singly22 or in combination with other lesions.19, 23, 24 They are associated with a systolic murmur at the left and right heart base and frequently carry a poor prognosis. Diagnosis is generally by echocardiographic visualization.
ported with pulmonic stenosis being the most prevalent.17,25–30 Foals with severe pulmonic stenosis may show clinical signs from birth; however, the majority of cases with aortic, tricuspid, or mitral dysplasia are aged from 3 to 18 months at diagnosis. The specific cardiac murmur(s) found will be determined by the specific valve(s) affected. Echocardiography may demonstrate cardiomegaly, valvular regurgitation, and thickening of the affected heart valve. The main differential diagnosis for valvular pathology in young animals is infective endocarditis, in which generally vegetative lesions are present rather than more diffuse nodular pathology that is typical of congenital valvular dysplasia. In infective endocarditis there is often concurrent laboratory evidence of infection, such as hyperfibrinogenemia and leukocytosis. The prognosis for horses with congenital valvular dysplasia is poor.
Patent ductus arteriosus The ductus arteriosus is a normal communication between the left and right sides of the circulation in the fetus, serving to allow part of the right heart output to bypass uninflated lung. It closes within the first few days of life and it is common to auscult systolic or continuous murmurs over the left heart base in newborn foals prior to complete closure of the ductus arteriosus. Persistence of a patent ductus arteriosus (PDA) is an uncommon congenital cardiac abnormality which, like the VSD and ASD, can occur either singly31–33 or in conjunction with other anomalies in complex cardiac disease.23,34–37 When it is an isolated and small lesion, affected horses may have a relatively normal lifespan, although there are very limited reports on which to base this assessment.31 Larger defects may lead to heart failure within the first 18 months of life, and affected horses may present with a continuous cardiac murmur loudest over the left heart base due to the shunt within the PDA; alternatively, the onset of biventricular dilation can lead to atrioventricular valvular insufficiency with bilateral systolic murmurs.5 Echocardiographic visualization of the PDA is difficult as it lies between the pulmonary artery and the descending aorta which is not visible from the right or left cardiac windows beyond the first day or two of life. Retrograde flow may be detected within the pulmonary artery and echocardiography will allow the identification of chamber dilation and heart failure if not the specific cause.
Valvular dysplasia
Complex congenital cardiac disease
Valvular dysplasia appears to be less common in horses than it is in dogs; however, congenital abnormalities of all of the heart valves have now been re-
A wide array of complex congenital cardiac diseases have now been described in horses. Broadly, these can be divided into defects of cardiac chamber
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(a)
509
(b)
(c)
Figure 56.2 Right parasternal color flow Doppler echocardiograms of the left ventricular outflow tract from two cases with ventricular septal defect (VSD). (a) A long-axis image of a large intracardiac jet within the VSD. (b) In this long-axis image, the defect is smaller and, consequently, it is not visible in this plane, but the intracardiac shunt is readily identified. (c) In a short-axis image from the same case, a small VSD is visible. RV, right ventricle; IVS, interventricular septum; LV, left ventricle; AO, aorta; LVOT, left ventricular outflow tract. (See color plate 5).
anatomy,23, 36, 38 those affecting the great vessels,39, 40 and combinations of both.41 All are extremely rare; however, within that context, tetralogy of Fallot,42 hypolastic left heart,43 and atresia of the right atrioventricular valve44, 45 appear to be the most prevalent forms of abnormalities of chamber anatomy while there are a number of anomalies of the great vessels including truncus arteriosus,32, 46 transposition of the great vessels, and anomalous origins of various branches of these vessels.37 In all complex congenital cardiac diseases, the onset of clinical signs can be expected at a young age, often within the first 3 months, whereas signs of severe cardiac compromise include poor growth rates, increased respiratory rate and effort, and poor exercise tolerance. Loud
cardiac murmurs are almost always present and echocardiography is used to demonstrate the specific structural abnormalities. Each vessel and chamber must be identified and its size and relationship to other cardiac structures determined.9 In complex congenital cardiac disease, the prognosis is invariably hopeless.
References 1. Crowe MW, Swerczek TW. Equine congenital defects. Am J Vet Res 1984;46:353–8. 2. Collobert-Laugier C, Tariel G. Congenital abnormalities in foals: results of a seven year postmortem survey. Prat Vet Equine 1993;25:105–10.
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3. Puyalton-Moussu C, Collobert C, Tariel G, Foucher N. Study of congenital abnormalities in horses in Lower Normandy: incidence (1994–1998) and risk factors. Epidemiol Sante Anim 1999;35:87–96. 4. Buergelt CD. Equine cardiovascular pathology: an overview. Anim Health Res Rev 2003;4(2):109–29. 5. Marr CM. Cardiac murmurs: congenital heart disease. In: Marr CM (ed.) Cardiology of the Horse. London: WB Saunders, 1999; pp. 210–32. 6. Lombard C, Evans M, Martin L, Tehrani J. Blood pressure, electrocardiogram and echocardiogram measurements in the growing pony foal. Equine Vet J 1984;16:342–7. 7. Reef VB. Cardiovascular disease in the equine neonate. Vet Clin North Am Equine Pract 1985;1:117–29. 8. Bonagura JD, Herring DS, Welker F. Echocardiography. Vet Clin North Am Equine Pract 1985;1(2):311–33. 9. Reef VB. Echocardiographic findings in horses with congenital cardiac disease. Comp Contin Educ 1991;13:109–17. 10. Stewart J, Rose RJ, Barko, A. Echocardiography in foals from birth to three months old. Equine Vet J 1984;16:332– 41. 11. Kvart C, Carlsten J, Jeffcott LB, Nilsfors L. Diagnostic value of contrast echocardiography in the horse. Equine Vet J 1985;17(5):357–60. 12. Steyn PF, Holland P, Hoffman JS. The angiographic diagnosis of a persistent truncus arteriosus in a foal. J S Afr Vet Assoc 1989;60:106–8. 13. Lamb C, O’Callaghan MW, Paradis MR. Thoracic radiography in the neonatal foal: a preliminary report. Vet Radiol Ultrasound 1990;31:11–16. 14. Hinchcliff KW, Adams WM. Critical pulmonic stenosis in a newborn foal. Equine Vet J 1991;23:318–20. 15. Ueno Y, Tomioka Y, Kaneko M. Muscular ventricular septal defect in a horse. Bull Equine Res Inst 1992;29:15–19. 16. Deniau V, Delecroix A. A case of interventricular communication in a racehorse. Prat Vet Equine 2004;36:25–31. 17. Critchley KL. An interventricular septal defect, pulmonary stenosis and bicuspid pulmonary valve in a Welsh pony foal. Equine Vet J 1976;8(4):176–8. 18. Young LE, Blunden AS, Bartram DH, Edgar A. Pulmonary atresia with an intact ventricular septum in a thoroughbred foal. Equine Vet Educ 1997;9:123–7. 19. Reppas GP, Canfield PJ, Hartley WJ, Hutchins DR, Hoffman KL. Multiple congenital cardiac anomalies and idiopathic thoracic aortitis in a horse. Vet Rec 1996;138:14– 16. 20. Reef VB. Evaluation of ventricular septal defect in horses using two-dimensional and Doppler echocardiography. Equine Vet J 1995; Suppl 19: 86–95. 21. Bonagura JD, Blissitt KJ. Echocardiography. Equine Vet J 1995; Suppl 19: 5–17. 22. Taylor FG, Wotton PR, Hillyer MH, Barr FJ, Luce VM. Atrial septal defect and atrial fibrillation in a foal. Vet Rec 1991;128:80–1. 23. Chaffin MK, Miller MW, Morris EL. Double outlet right ventricle and other associated congenital cardiac anomalies in an American miniature horse foal. Equine Vet J 1992;24(5):402–6. 24. Physick-Sheard PW, Maxie MG, Palmer NC, Gaul C. Atrial septal defect of the persistent ostium primum type with hypoplastic right ventricle in a Welsh pony foal. Can J Comp Med 1985;49(4):429–33. 25. Clark ES, Reef VB, Sweeny CR, Lichtensteiger C. Aortic valvular insufficiency in a one-year-old colt. J Am Vet Med Assoc 1987;191:841–4.
26. King JM, Flint TJ, Anderson WI. Incomplete subaortic stenotic rings in domestic animals: a newly described congenital anomaly. Cornell Vet 1988;78:262–71. 27. Davis JL, Gardner SY, Schwabenton B, Breuhaus BA. Congestive heart failure in horses: 14 cases (1984–2001). J Am Vet Med Assoc 2002;220(10):1512–15. 28. McGurrin MK, Physick-Sheard PW, Southorn E. Parachute left atrioventricular valve causing stenosis and regurgitation in a Thoroughbred foal. J Vet Intern Med 2003;17(4):579–82. 29. Taylor SE, Else RW, Keen JA. Congenital aortic valve dysplasia in a Clydesdale foal. Equine Vet Educ 2007;19:463–8. 30. Reimer JM. Congenital defects and reversions to foetal circulation. In: Reimer JM (ed.) Atlas of Equine Ultrasonography. St. Louis: Mosby, 1998; pp. 138–47. 31. Carmichael JA, Buergelt CD, Lord PF, Tachjian RJ. Diagnosis of patent ductus arteriosus in a horse. J Am Vet Med Assoc 1971;158:767–75. 32. Steyn PF, Holland P, Hoffman J. The angiographic diagnosis of a persistent truncus arteriosus in a foal. J S Afr Vet Assoc 1989;60:106–8. 33. Guarda HI, Schifferlis RCA, Alvarez MLH, Probst A, Konig HE. A patent ductus arteriosus in a foal. Wien Tierarztl Monatsschr 2005;92:233–7. 34. Hinchcliff KW, Adams WM. Critical pulmonic stenosis in a newborn foal. Equine Vet J 1991;23:318–20. 35. van der Linde-Sipman JS, Goedegebuure SA, Kroneman J. Persistent right aortic arch associated with a persistent left ductus arteriosus and an interventricular septal defect in a horse. Tijdschr Diergeneeskd 1979;104(20): 189–94. 36. Bayly WM, Reed SM, Leathers CW, Brown CM, Traub JL, Paradis MR, Palmer GH. Multiple congenital heart anomalies in five Arabian foals. J Am Vet Med Assoc 1982;181(7):684–9. 37. Reimer JM, Marr CM, Reef VB, Saik JE. Aortic origin of the right pulmonary artery and patent ductus arteriosus in a pony foal with pulmonary hypertension and right sided heart failure. Equine Vet J 1993;25:466–70. 38. Kraus MS, Pariaut R, Alcaraz A, Gelzer ARM, Malik N, Renaud-Farrell S, Charter ME, Fox PR. Complete atrioventricular canal defect in a foal: clinical and pathological features. J Vet Cardiol 2005;7:59–64. 39. Jesty SA, Wilkins PA, Palmer JE, Reef VB. Persistent truncus arteriosus in two Standardbred foals. Equine Vet Educ 2007;19:307–11. 40. Valdes-Martinez A, Easdes SC, Strickland KN, Roberts ED. Echocardiographic evidence of an aortic-pulmonary septal defect in a 4-day-old thoroughbred foal. Vet Radiol Ultrasound 2006;47:87–9. 41. Zamora CS, Vitums A, Nyrop KA, Sande RD. Atresia of the right atrioventricular orifice with complete transposition of the great arteries in a horse. Anat Histol Embryol 1989;18(2):177–82. 42. Gessell S, Brandes K. Tetralogy of Fallot in a 7-year-old gelding. Pferdeheikunde 2006;22:427–30. 43. Musselman EE, LoGuidice RJ. Hypoplastic left ventricular syndrome in a foal. J Am Vet Med Assoc 1984;185(5):542–3. 44. Wilson RB, Haffner JC. Right atrioventricular atresia and ventricular septal defect in a foal. Cornell Vet 1987;77(2):187–91. 45. Meurs KM, Miller MW, Hanson C, Honnas C. Tricuspid valve atresia with main pulmonary artery atresia in an Arabian foal. Equine Vet J 1997;29(2):160–2. 46. Sojka JE. Persistent truncus arteriosus in a foal. Equine Pract 1987;9:24–6.
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Chapter 57
Acquired Abnormalities of the Cardiovascular System C. J. (Kate) Savage
Cardiac abnormalities in foals are of particular concern, as the manifestations of disease may have serious consequences for longevity, performance, and safety. Congenital abnormalities of the cardiovascular system are covered in Chapter 56. Many acquired abnormalities will not present until the foal is older; however, this is true also of a number of congenital cardiac abnormalities, so the differential diagnostic list can be extensive as diagnostic investigation is commenced. It is prudent to remember that murmurs are simply audible vibrations caused by turbulent blood flow with transmission to the body surface. Turbulence is most easily generated when blood viscosity is low, so both large luminal size (e.g., arteries, ventricles, and atria) and anemia are important in murmur generation. Consequently, when a murmur is detected in a foal, it is important to know whether the patient is anemic; a packed cell volume (PCV, hematocrit) should be measured, and important history should be obtained, for example, sepsis, deworming.
Recognition of an acquired cardiovascular abnormality in a foal History A comprehensive history is mandatory if one is effectively to evaluate the foal’s cardiovascular system. Even if the foal is beyond the neonatal period, it is imperative to know certain facts pertaining to the birth, including:
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prematurity/dysmaturity; presence of placentitis; presence of dystocia; presence of known or suspected systemic disease in the dam or foal.
The links provided by a pertinent history are invaluable when trying to choose diagnostic tests practically for both patient welfare and client economics. Other useful historical data include:
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Timing of onset of clinical signs – i.e., pre- or postweaning, or after illness of another body system (e.g., septicemia, pneumonia, enterocolitis, presence of fever of unknown origin in the last 2 months). History or presence of an intravenous catheter and/or thrombophlebitis – this is of interest in itself as a vascular disease in foals, but also may be associated with secondary endocarditis. Vaccination history. Deworming history – for example, if the foal has a murmur, the deworming history is important in trying to assess the likelihood of anemia, etc. It is helpful to know the deworming history in cases of foal/weanling diarrhea, as there is a link between enterocolitis and septicemia, and that in turn may be linked with endocarditis. Soil selenium – likelihood of being a candidate for white muscle disease.1 Exercise tolerance (although this may be misleading in cases of both acquired and congenital cardiovascular disease).
Physical examination of the foal Clinical evaluation and knowledge of normal values are essential for prompt placement of cardiovascular disease on the differential list in a sick foal. Mucous membranes (both oral and vulval/preputial) should be pink and have a refill time of less than 2 seconds. It is important to examine mucous membranes before contaminating your hands (or gloves) by touching the mare or foal (especially by taking the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 C. J. (Kate) Savage.
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rectal temperature), as the veterinarian must not act as a fomite (e.g., Salmonella sp.). Auscultation of the heart on both left and right sides should be performed whilst simultaneously palpating the arterial pulse (e.g., facial artery or digital artery) to determine whether there are any pulse deficits. It is also important to establish the pulse amplitude and contour where possible. This may be helpful in determining prognosis in some cases; for example, if the aortic valve is affected and the pulse is hyperkinetic, then prognosis is worsened. Heart rate (HR) should be taken immediately to decrease the chance of erroneous measurements, due to stress. The HR varies with the age of the foal, even during the neonatal period. Normal equine juvenile HR data include:
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newly born neonate HR = 40–80 beats per minute (bpm); neonate (1–5 minutes of age) HR > 60 bpm; neonate (at approximately 1 hour or when trying to stand2 HR = 120–150 bpm; neonate > 24 hours of age (resting) HR = 80– 100 bpm; neonate > 24 hours of age (active) HR = 100– 150 bpm; neonate by 7 days of age HR = 80–100 bpm; neonate by 14 days of age HR = 80–90 bpm;2 weanling HR = 40–66 bpm; yearling HR = 32–56 bpm.
When tachycardia is diagnosed, one has to judge the severity and whether it is related to sympathetic stimulation (stress, pain), non-cardiac disease, or primary cardiac disease. The apex beat is usually loudest in the left hemithorax in the 5th intercostal space. The veterinarian should still use the ‘pA M ’ system in the left hemithorax, where the pulmonic valve (‘P’) is auscultated most cranially (3rd intercostal space ventral to the shoulder), and the aortic valve (‘A’) is auscultated dorsal and more caudal (the 4th intercostal space) and the mitral valve (‘M’) is in the 5th intercostal space (i.e., most caudal), ventral to the aortic valve and pulmonic valve positions. However, in the foal subtle movements of the stethoscope head are needed as the foal heart is much smaller than that of an adult. Murmurs can be auscultated and, indeed, palpated if there is a thrill (grade V/VI or VI/VI), in which case they are invariably pathological.3 Grades4, 5 can be defined as follows: Grade I: Murmur barely audible. Discernible only after careful auscultation over a focal area. Grade II: Low-intensity murmur. Heard immediately to a few seconds after the stethoscope is placed
over the point of maximal impulse of the murmur. Softer than the first heart sound (S1). Grade III: Loud murmur that is immediately audible. May be heard over a wide area of the chest. Equally as loud as S1. Grade IV: A prominent, widespread murmur that is louder than S1 and may be heard over a wide area of the chest. Grade V: A prominent murmur (the loudest that becomes inaudible when the stethoscope is removed) with a palpable thrill. Grade VI: A loud murmur with accompanying precordial thrill that can be auscultated with the stethoscope off the chest wall. In the early neonatal period, before taking steps to diagnose other congenital causes or acquired causes of cardiac disease, it is important to remember that the common congenital cardiac murmur in the neonate is a patent ductus arteriosus (PDA). Systolic murmurs were auscultated in all 10 pony foals soon after birth (i.e. first 3 days) in a study by Livesey et al., 6 but continuous murmurs were only detected in 5 of these 10 pony foals.6 It was interesting and clinically relevant that six pony foals still had systolic murmurs on day 35, and two pony foals had systolic murmurs on day 49. Interestingly, there was still a large amount of turbulent flow detected echocardiographically in the ductus arteriosus on days 35 and 49. On day 1 of this study, auscultation of a continuous murmur indicating flow in the ductus arteriosus only had a sensitivity of 40%.6 Prior to echocardiographic findings by Lombard et al.7 and Livesey et al.,6 PDA had usually been assumed significant after 4 days; however, more recently clinicians have used 7 days as the threshold of clinical significance using auscultation alone. These data are particularly important as a veterinarian may only see a foal at 10, 20, or 30 days of age and without a prior auscultation history, making it much harder to make purely clinical decisions without further diagnostic testing (e.g., echocardiography). The latter is easily accomplished in foals in referral hospitals, but now many general equine veterinarians have access to excellent ultrasonographic equipment with transducers (e.g., 4–7 MHz) and even Doppler capabilities, so that quick and practical evaluation of the heart valves’ appearance and chamber sizes may be performed on the farm. The rectal temperature must be recorded – it is prudent to advise the owner or stud manager to record the temperature twice daily. This gives the veterinarian more information regarding inflammation and infection, despite usage of non-steroidal antiinflammatory drugs. The jugular vein in the jugular furrow should be non-distended, but should readily fill when occluded. Jugular pulsations should be seen
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Acquired Abnormalities of the Cardiovascular System in the recumbent and standing foal. When standing with the head in normal carriage (i.e., not held up or down) the pulsation should only progress to a third or half of the way up the jugular furrow. This jugular pulsation reflects normal pressure changes in the thorax and right atrium.8
Diagnostic testing
Clinical pathology In foals with a significant cardiovascular inflammatory process such as valvular bacterial endocarditis, pericarditis, and septic thrombophlebitis (which is important in itself and also as a causal factor for endocarditis), the complete blood count (CBC) can be extremely useful in acting as a prompt to perform further diagnostics (especially echocardiography and ultrasonography of veins during and after intravenous catheterization). It is imperative to perform echocardiography in foals that have a leukocytosis characterized by a neutrophilia (especially with a significant left shift) and a hyperfibrinogenemia if the foal is fevered or has a history of fever and another system has not been identified. Frequently, tachycardia, a murmur, and even tachypnea will be obvious, but echocardiography in foals is easy and quick to perform and should not be delayed. The biochemical profile can be useful in these patients too, as there may be a hyperglobulinemia as well, and subsequently the total protein may be mildly to moderately elevated. Anemia may be present due to chronic inflammation in these patients, so the PCV should be evaluated critically in the light of the white cell count (WCC) and fibrinogen and globulin levels. However, even when a low-grade murmur is identified in a foal the PCV should be measured, as anemia can allow decreased viscosity and increased turbulence. However, if the murmur is greater than or equal to grade III/VI, then echocardiography should be performed, even in the face of anemia without other characteristics of inflammation. Biochemical profile measurements of specific use in foals with arrhythmias include the potassium concentration and the acid–base status of the foal; however, provision of a full biochemical profile may elucidate a number of abnormalities. Biomarkers used in human medicine have been used as indicators of cardiac injury in adult horses and foals. The cardiac troponins I (cTnI), T (cTnT), and C (cTnC), as well as the myocardial bound isoform of creatine kinase (CK-MB) and brain (B-type) natriuretic peptide (BNP) have been used to assess cardiac injury in humans. It appears that BNP is predominantly synthesized in the ventricular myocardium in humans,9, 10 but this has not been veri-
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fied in horses. Cardiac troponins I and T are highly conserved across species and have been used in humans, cats, dogs, and adult horses and foals to detect myocardial cell damage.11–21 Slack et al.22 established a reference range for cTnI, cTnT, and CK-MB in clinically normal neonatal foals, as well as evaluating septic foals in a prospective study. Cardiac troponin I inhibits the structural interaction of myosin heads with actin binding sites, while cTnT binds the troponin complex to tropomyosin. Release of cTnI and cTnT occurs when there is cardiac ischemia and necrosis, and may occur in dogs when the primary clinical signs are due to pericardial effusion secondary to hemangiosarcoma.15 Measurement of circulating cTnI has been thought to be more sensitive than measurement of either cTnT or the cardiac isoenzyme of creatine kinase (CK-MB). In the prospective foal study, cTnI, cTnT, and CK-MB were compared in 52 clinically normal foals (aged between 12 and 48 hours) and septic neonates.22 There were significant differences between clinically normal neonates and septic foals, suggesting myocardial damage during sepsis. However, there was no difference in cTnT and CK-MB in surviving and nonsurviving foals, so the usefulness of cTnT to predict morbidity appears minimal. Muurlink et al.23 demonstrated large elevations in cTnI after collapse and resuscitation of a foal with splenic rupture and hemoabdomen. The foal was in ventricular tachycardia, which was uniform at the time of collapse and then multiform/multicentric after surgery. Echocardiographic evaluation revealed regions of myocardial hyperechogenicity. The arrhythmia was abolished using lidocaine pre-surgery and was monitored post-surgery. The cTnI level decreased slowly (monitored for months) as the foal’s clinical parameters and echocardiogram improved.23 B natriuretic peptide is also of interest as a myocardial biomarker in horses, and work on this is progressing (C.M. Fennell and C.J. Savage, University of Melbourne, personal communication).
Electrocardiography (ECG) Foals with myocardial damage, electrolyte or acid–base abnormalities may have arrhythmias. Foals with ruptured bladder frequently have electrolyte abnormalities (especially hyperkalemia) and may be arrhythmic. Some neonatal foals have been diagnosed with atrial fibrillation (AF). This appears to be acquired/ adaptive and of varying, but usually low, clinical significance.24, 25 In most foals, the AF has been transitory (lasting between 5 and 170 minutes postpartum). No treatment was administered in most cases;24 however, Machida et al.25 described treatment with
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300 mg procainamide intravenously after 200 minutes of AF postpartum associated with collapse. Sinus rhythm was immediately established. However, AF is rarely observed clinically in foals that are presented for cardiac disease in referral hospitals. Ventricular tachycardia has been documented in a 3month-old foal with splenic rupture and cardiovascular collapse.23 Electrocardiography is easy to perform on the neonatal foal; however, weanlings and yearlings may behave poorly if clip-on electrodes are used. Adhesive electrodes are easy to use and readily available with many systems of ECG, especially newer telemetric systems. The latter have the added advantage of allowing monitoring when the machine is outside the stall or yard, so that ECG associated with less stressed behavior and exercise may both be achieved.
Echocardiography The echocardiogram is easy to perform in foals whether they are recumbent or standing. As foals and weanlings have relatively low bodyweight, even ultrasound machines with less sophisticated capabilities may be used successfully to give the equine practitioner some idea of valvular, myocardial, and pericardial structure. The ratio of left ventricular free wall thickness to right ventricular free wall thickness in neonates is usually 1:1; however, this alters over time, until the adult ratio of 3:1 is attained. The endocardium, myocardium, epicardium, and pericardium as well as the valvular structures should be evaluated echocardiographically. Echocardiography has been well described in foals and adult horses.7,26–29 A limited description is given here for ‘on-farm’ echocardiography of a foal. Use a machine and transducer with a depth availability of at least 14 cm for neonates, 20 cm for weanlings, and 24 cm for yearlings. Machines with depth capabilities greater than 24 cm are advantageous, but not widely available. Clip the right axillary region or wet it with alcohol and/or coupling gel. The hair from the right elbow will often be pulled down towards your imaging area when you apply the transducer with pressure; however, this is much more evident on adults. If it is a problem, simply clip a strip of hair from the elbow. Try to point the transducer head (rotating the reference point to 1 o’clock)27 cranially so a picture of the right ventricular outflow is seen, and the tricuspid and pulmonic valves can be evaluated. Then point the transducer head slightly caudally so that you bring the aorta and aortic valve into view. Then bring the transducer head back slightly for a fourchambered view, so the mitral and tricuspid valves can be observed. The best review of equine echocar-
diographic technique is given by Reef (1998)27 – these techniques can be applied to foals with ease.
Acquired valvular heart disease The veterinarian must recognize that when a murmur or arrhythmia is detected, congenital cardiovascular disease will initially need to be considered as well as acquired conditions. Congenital abnormalities may often predispose to degeneration of other valves and endocardium. Foals with ventricular septal defect (VSD)30 may have additional murmurs associated with aortic, tricuspid, and mitral insufficiencies, and a murmur of relative pulmonic stenosis (due to increased flow through the pulmonic artery). The following is a differential diagnostic list: 1. Endocarditis. Any valve can be affected, but lesions of mitral, aortic, and tricuspid valves appear to be most prevalent. 2. Myocardial diseases: a myocardial infarcts and hypoxia/ischemia;23 b myocardial and pericardial trauma – fractured ribs may cause perforation of the pericardium (and hemopericardium) and myocardial damage; c myocardial abscessation – Streptococcus zooepidemicus;31 d myocarditis: bacteria, virus, parasites – with fibrosis and/or calcification;27 e white muscle disease – vitamin E and selenium deficiency; f toxins – ionophore, heavy metal toxicities, Cassia spp. – unlikely but possible. Vitamin E (alpha-tocopherol) has been suggested to have some protective antioxidant effects in other species with myocardial disease. This is a very safe oral supplement and can be used in equine cardiac patients; however, its effects in the horse are unknown. 3. Pericardial disease 4. Neoplasia 5. Arrhythmias. Endocarditis Endocarditis occurs rarely in foals. In adult horses it may transpire due to chronic insult to the heart valves, thereby ensuring that induction of endocarditis is easier. If the foal has a history of septicemia, pneumonia, enterocolitis, and/or fever with leukocytosis characterized by neutrophilia and hyperfibrinogenemia (even if a murmur cannot be detected), then the foal should undergo a cardiac examination and
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monary artery due to cranial mediastinal abscessation is possible, but unlikely. Mediastinal tumors in foals are unlikely in comparison to young cattle, in which they are regularly reported.32, 33 Acquired myocardial disease
Myocardial infarcts and hypoxia/ischemia These can occur for a variety of reasons, including endotoxemia and lack of perfusion in foals with colic,23 pneumonia, or septicemia.
Figure 57.1 Echocardiogram from a 14-month-old Thoroughbred filly with a 2-month history of fever and leukocytosis of unknown origin. Aortic valve endocarditis was diagnosed using auscultation and echocardiography. The markedly thickened septal leaflet of the aortic valve is seen in long axis at the level of AV. A small regurgitant jet (green) is seen using colorflow Doppler here. (See color plate 6).
echocardiographic evaluation, which will include concurrent ECG. In a foal or weanling, the clarity of image should be excellent as the ultrasound machine is not challenged with penetration issues as it is in adult horses. If the foal requires referral for these tests it should be done immediately, as delays to intensive care can affect prognosis. If a valvular lesion is detected in a febrile foal, weanling, or yearling (Figure 57.1) with an inflammatory clinical pathological profile, then it is imperative to test (e.g., multiple blood cultures) and treat aggressively. Antimicrobial treatment may be based on blood cultures, but initially should be broadspectrum. Antimicrobials usually should be used for 4–6 weeks, but each patient’s therapy requires individual attention. Non-steroidal anti-inflammatory drugs (NSAIDs) may be used to decrease platelet function. Aspirin is often the drug of choice and can be used at 10 mg/kg/q 24 hrs p.o. It is rare that adult horses on chronic administration of aspirin suffer from gastric ulceration or right dorsal colitis, but as these foals are compromised, it may be prudent to treat prophylactically with omeprazole and misoprostol. Although other NSAIDs may depress platelet function for more than 24 hours, aspirin affects adhesion of platelets irreversibly throughout their circulating life. Depending on the degree of insufficiency (characterized by the murmur and more importantly the color-flow Doppler echocardiographic findings) the foal may require treatment with an ACE (angiotensin-converting enzyme) inhibitor (e.g., enalapril) or management of congestive heart failure (e.g., digoxin). Pulmonary stenosis is rare in horses. A murmur secondary to supravalvular stenosis of the pul-
Myocardial and pericardial trauma Fractured ribs may cause perforation of the pericardium (and thus hemopericardium) and myocardial damage. This can lead to potentially fatal arrhythmias. This may occur to foals during parturition; but may also be inflicted by the dam, or by other foals and horses.34–36 Jean et al.36 found that on one studfarm only 43.4% of foals with thoracic trauma were colts, which is contrary to most anecdotal reports in which colt foals have been thought to be over-represented due to their size. In 2007, Jean et al.37 reported that ultrasonography was more accurate in detecting rib fractures. Interestingly the percentage of fillies (68%; 13/19) with rib fracture was greater than that of similarly affected colts.37
Myocardial abscessation Jackson et al.31 described a 1-month-old male Thoroughbred foal with a large (3 × 4-cm) abscess of the right ventricular free wall. Streptococcus equi var. zooepidemicus was cultured.31 The foal was found dead with no signs of trauma or illness. Unfortunately details of the foal’s birth and maturity, and the mare’s parity and placenta were not given. It is likely that a foal like this may have had episodes of life-threatening ventricular arrhythmias.
Myocarditis Etiological agents in cases of myocarditis can include bacteria, virus, and parasites, and the ramifications may be fibrosis and/or calcification.27 It is possible that foals, weanlings, and yearlings with myocarditis and myocardial abscessation could have severe and possibly terminal arrhythmias.
White muscle disease This is a nutritional muscular dystrophy caused by alpha-tocopherol (vitamin E) and selenium deficiency.38 Anecdotally, Dill and Rebhun39 and Valberg40 have described putative myocardial involvement with sudden death after arrhythmia
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and weakness. Perkins et al.41 described myocardial necrosis in one of four foals diagnosed with electrolyte disturbances, including low ionized calcium, and rhabdomyolysis associated with selenium and possibly vitamin E deficiency. Only two of the four foals had necropsies, although three underwent euthanasia. Although hyperkalemia, hyponatremia, hypochloremia, and hyperphosphatemia are more commonly associated with ruptured bladder or renal disease in foals, white muscle disease in foals has been associated with these electrolyte abnormalities.42
Ionophore toxicity This scenario is possible, but rare.43 Ionophore antibiotic contamination could theoretically occur in a batch of feed to which foals have access. The usual way proposed for ionophore toxicity is when there is contamination of horse feed after using the same facilities to manufacture poultry, pig, and cattle feeds, in which the ionophore is used as a coccidiostat or growth promoter, respectively.44 As horses are very sensitive to ionophore antibiotics, accidental toxicity is usually disastrous. The author has not seen an affected foal, but adults can have clinical signs of cardiac failure (myocardial degeneration) and neuropathy,44 and foals and weanlings, if accidentally exposed, could have severe signs because of their smaller bodyweight (depending on the amount of feed ingested). Echocardiographic signs include lack of interventricular septum (IVS) and free wall motility leading to extremely reduced fractional shortening measurements. Depending on the degree of myocardial degeneration, there may also be paradoxical movement of the IVS. It is advisable to perform echocardiography and measure cTnI and other biomarkers as soon as ionophore toxicity is suspected. Information on other cardiac toxins, predominantly affecting older horses, can be found in equine medicine texts.
Figure 57.2 Echocardiogram from a 6-month-old Thoroughbred colt with fibrinous pericarditis. The tricuspid valve is seen at the level of TV. The roughened surface can be seen clearly (arrows) and there is excessive pericardial fluid (hypoechoic portion around arrows).
the control group horses was 7 years) and exposure to Eastern tent caterpillars on the farm.46 Bivariate analyses also identified some other factors including lack of direct contact with cattle, and farm being affected with MRLS. Whether caterpillars are directly toxic, harbor a toxin, or are a marker for exposure of another sort is not known. Interestingly, at necropsy, the predominant bacterial species identified was Actinobacillus sp., although Streptococcus zooepidemicus and Enterococcus faecalis were identified in some. Actinobacillus sp. may be pericardiotrophic, but it is unlikely to have been the primary agent in this epidemic of fibrinous pericarditis in Kentucky.47 Hemopericardium may occur in some foals with trauma, including fractured ribs, and veterinarians should be aware of this complication in these foals. In cases with hemothorax, the author frequently manages these foals medically; however, if sharp fracture fragments appear to be in close proximity to the cardiac silhouette (clinically and ultrasonographically), then it may be prudent to perform surgical fracture repair (usually with plating).
Acquired pericardial abnormalities Pericarditis may be effusive, fibrinous (Figure 57.2) or constrictive (which may be a further stage of the fibrinous type). Although this is a well-known cause of cardiovascaular disease, most publications refer to adults45 and young horses,46 rather than foals. In the spring and summer of 2001, there was an epidemic of equine fibrinous pericarditis in Kentucky, USA, in association with mare reproductive loss syndrome (MRLS)47 and exposure to Eastern tent caterpillars.46 Risk factors for development of fibrinous pericarditis identified included younger age (median age of case horses was 1 year whereas the median age for
Neoplasia This is rare in foals. Other than lymphoma, few tumors have been diagnosed in young horses. In adult horses, hemangiosarcoma and mesothelioma have been diagnosed in pericardium;48, 49 and an infiltrating lipoma was found in myocardium.50 Metastatic lymphoma and hemangiosarcoma have been reported in adult horses.51, 52 Hepatocellular carcinoma has been diagnosed definitively in foals,53 but to the author’s knowledge no myocardial metastases have been found, including in a 3-month-old foal and a yearling that the author has seen.
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Acquired Abnormalities of the Cardiovascular System Arrhythmias When an arrhythmia is detected there may be an underlying cardiac problem, an electrolyte or acid–base problem, or, in the first 15 minutes of life, vagal tone and hypoxemia may cause a variety of arrhythmias.24
Atrial fibrillation This was noted in 15/50 foals in the immediate postpartum period, and these cases were considered physiological.24 Machida et al.25 also reported on three foals with AF, and one required treatment at 200 minutes postpartum. There was no correlation between fetal and postpartum ECG in foals described by either study.24, 25
Bladder rupture This occurs in neonates usually aged less than 7 days and predominantly in colt foals. Numerous electrolyte abnormalities may occur including hyponatremia, hypochloremia, and hyperkalemia. It is recommended that anesthesia and surgical repair be delayed until electrolyte abnormalities are corrected. Otherwise a number of ventricular arrhythmias and third-degree atrioventricular block may occur. Twenty-five years ago 37.5% of foals in one retrospective study had cardiac arrest or third-degree atrioventricular block.54
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there is bacteremia and/or sepsis. When foals have indwelling intravenous catheters, the vein should be monitored at a minimum of twice daily by distending the vein and ensuring that it is still able to ‘rise’ and then collapse as soon as the vein is released. However, care must be taken not to compromise the cleanliness or sterility of the area. If the vein appears to be raised already then palpation of the vein is prudent. If the vein will not collapse or feels hard, then the catheter should be removed and the tip cultured (aerobically and anaerobically). If the temperature becomes and/or remains elevated, blood culture is beneficial. During daily examination, the catheter entrance to the vein should also be assessed to ensure there is no purulent discharge present. After catheter removal, the area should be evaluated ultrasonographically with attention paid to any ‘gas sparkling’ indicative of anaerobic activity with gas production, as that will affect treatment and prognosis. Topical anti-inflammatory ointments and hot-packs may be used, but the benefit of these is not known. The temperature, white cell count and distribution, and fibrinogen concentration should be measured and monitored. The heart should be monitored closely for murmurs, as endocarditis can be a sequela to thrombophlebitis. Occasionally septic veins with thrombophlebitis, that are unresponsive to medical treatment, must be removed surgically. Catheter loss
Other arrhythmias Yamamoto et al.24 reported that 48 of 50 foals studied in the first 15 minutes postpartum had one or multiple arrhythmias. The type of arrhythmia could not be predicted by fetal ECG. Most commonly observed were sinus arrhythmias, including wandering pacemaker (32/50) and atrial premature contraction (10/50). Also seen were AF (15/50), ventricular premature contraction (10/50), partial atrioventricular block (7/50), ventricular tachycardia (4/50), atrial tachycardia (3/50), and idioventricular rhythm (1/50). The authors suggested that these were due to high vagal tone and hypoxemia; it may be possible that electrolyte changes also played a role, but all arrhythmias had disappeared by 15 minutes of age.24
Acquired vascular abnormalities Venous thrombosis and thrombophlebitis Foals, like adults, can suffer from venous thrombosis and thrombophlebitis after catheter placement, especially if polyurethane catheters are not used and if
If a portion of a catheter is lost, then the foal should be sedated and ultrasonography of the local region and then circulation including the heart should be undertaken. This is important as retrieval, especially from the jugular vein just distal to the site of loss, can be managed at referral hospitals with immediate surgery.
References 1. Ferrans VJ, Van Vleet JF. Cardiac lesions of seleniumvitamin E deficiency in animals. Heart Vessels 1985;1:294– 7. 2. Machida N, Yasuda J, Too K. Auscultatory and phonocardiographic studies on the cardiovascular system of the newborn thoroughbred foal. Jpn J Vet Res 1987;35:235– 50. 3. Fregin GF. The purchase examination: the cardiovascular system. In: Proceedings of the 24th Annual Convention of the AAEP, St Louis, MO, 1978, pp. 583–9. 4. Levine SA, Harvey WP. Clinical Auscultation of the Heart. Philadelphia: W.B. Saunders, 1949. 5. Blissitt KJ. Auscultation. In: Marr CM (ed.) Cardiology of the Horse. Philadelphia: W.B. Saunders, 1999. 6. Livesey LC, Marr CM, Boswood A, Freeman S, Bowen IM, Corley KTT. Auscultatory and two-dimensional M mode,
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spectral and colour flow doppler echocardiographic findings in pony foals from birth to seven weeks of age. J Vet Int Med 1998;12: 255. Lombard CW, Evans M, Martin L, Tehrani J. Blood pressure, electrocardiogram and echocardiogram measurements in the growing pony foal. Equine Vet J 1984;16:342–7. Reef VB. Cardiovascular disease in the equine neonate. Vet Clin N Am Equine Pract 1985;1:117–29. de Lemos JA, Morrow DA, Bentley JH, Omland T, Sabatine MS, McCabe BS, Hall C, Cannon CP, Braunwald E. The prognostic value of B-type natriuretic peptide in patients with acute coronary syndromes. N Engl J Med 2001;345:1014–21. Mukoyama M, Nakao K, Hosoda K, Suga S, Saito Y, Ogawa Y, Shiakami G, Jougasaki M, Obata K, Yasue H. Brain natriuretic peptide as a novel cardiac hormone in humans. Evidence for an exquisite dual natriuretic peptide system, atrial natriuretic peptide and brain natriuretic peptide. J Clin Invest 1991;87:1402–12. Adams JE, Bodor GS, Davila-Roman D, et al. A marker with high specificity for cardiac injury. Circulation 1993;88:101–6. Peek SF, Slack JA, Darien BJ, Semrad SD, Marques F, Risberg A, Erb HN, Apple FS, McGuirk SM. Biochemical markers of cardiac injury in normal and surviving versus non-surviving septicemic neonatal foals. J Vet Emerg Crit Care 2004;14:S15. Herndon WE, Kittleson MD, Sanderson K, Drobatz KJ, Clifford CA, Gelzer A, Summerfield NJ, Linde A, Sleeper MM. Cardiac troponin I in feline hypertrophic cardiomyopathy. J Vet Intern Med 2002;16:558–64. Cornelisse CJ, Schott HC, Olivier NB, Mullaney TP, Koller A, Wilson DV, Derksen FJ. Concentration of cardiac troponin I in a horse with a ruptured aortic regurgitation jet lesion and ventricular tachycardia. J Am Vet Med Assoc 2000;217:231–5. Shaw SP, Rozanski EA, Rush JE. Cardiac troponins I and T in dogs with pericardial effusion. J Vet Intern Med 2004;18:322–4. Oyama MA, Sisson DD. Cardiac troponin-I concentration in dogs with cardiac disease. J Vet Intern Med 2004;18: 831–9. Phillips W, Giguere S, Franklin RP, Hernandez J, Adin D, Peloso JG. Cardiac troponin I in pastured and race-trained Thoroughbred horses. J Vet Intern Med 2003;17:597–9. Davis JL, Gardner SY, Schwabenton B, Breuhaus BA. Congestive heart failure in horses: 14 cases ( 1984–2001). J Am Vet Med Assoc 2002;220:1512–15. Schwarzwald CC, Hardy J, Buccellato M. High cardiac troponin I serum concentration in a horse with multiform ventricular tachycardia and myocardial necrosis. J Vet Intern Med 2003;17:364–8. Begg LM, Hoffman KL, Begg RA. Serum and plasma cardiac troponin I concentrations in clinically normal Thoroughbreds in training in Australia. Aust Vet J 2006;84:336– 7. Durando MM, Reef VB, Kline K, Birks EK. Acute effects of short duration, maximal exercise on cardiac troponin I in healthy horses. Equine Comp Exercise Physiol 2006;3:217– 23. Slack JA, McGuirk SM, Erb HN, Lien L, Coombs D, Semrad SM, Riseberg A, Marques F, Darien BJ, Fallon L, Burns P, Murakami MA, Apple FS, Peek SF. Biochemical markers of cardiac injury in normal, surviving septic, or nonsurviving septic neonatal foals. J Vet Intern Med 2005;19:577–80.
23. Muurlink A, Walmsley J, Savage CJ, Whitton RC. Splenectomy in a foal to control intra-abdominal hemorrhage caused by splenic rupture. Equine Vet Educ 2008;20: 362–366. 24. Yamamoto K, Yasuda J, Too K. Arrhythmias in newborn thoroughbred foals. Equine Vet J 1992;24:169–73. 25. Machida N, Yasuda J, Too K. Three cases of paroxysmal atrial fibrillation in the Thoroughbred newborn foal. Equine Vet J 1989;21:66–8. 26. Slater JD, Herrtage ME. Echocardiographic measurements of cardiac dimensions in normal ponies and horses. Equine Vet J (Suppl.) 1996;19:28–32. 27. Reef VB. Equine Diagnostic Ultrasound, 1st edn. Philadelphia: W.B. Saunders, 1998. 28. Long KL, Bonagura JD, Darke PGG. Standardised imaging technique for guided M-mode and Doppler echocardiography in the horse. Equine Vet J 1992;24:226–35. 29. Stewart JH, Rose RJ, Barko AM. Echocardiography in foals from birth to three months old. Equine Vet J 1984;16:332–41. 30. Hughes KJ. Diagnostic challenge: Lethargy and weakness in an Arabian foal with cardiac murmurs. Aust Vet J 2006; 84:209–12. 31. Jackson C, Newman D, Locke S. Streptococcus zooepidemicus myocardial abscessation in a 1-month-old Thoroughbred foal. J Equine Vet Sci 2008;28:103–4. 32. Nasir KS. Sporadic juvenile thymic lymphoma in a 6month-old Holstein heifer. Can Vet J 2005: 46; 831–3. 33. Radostits OM, Gay CC, Blood DC, Hinchcliff KW. Veterinary Medicine, 9th edn. London: W.B. Saunders, 2000. 34. Kraus BM, Richardson DW, Sheridan G, Wilkins PA. Multiple rib fracture in a neonatal foal using a nylon strand suture repair technique. Vet Surg 2005;34:399–404. 35. Reimers J. Cardiovascular ultrasonography. In: Equine diagnostic ultrasonography (Eds) Rantanen NW, McKinnon AO Williams and Wilkins, 1988, pp 583–4. 36. Jean D, Laverty S, Halley J, Hannigan DRL Leveille R. Thoracic trauma in newborn foals. Equine Vet J 1999;31: 149–52. 37. Jean D, Picandet V, Macieira S, Beauregard G, D’Anjou MA, Beauchamp G. Detection of rib trauma in newborn foals in an equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 2007;39:158–63. 38. Higuchi T, Ichijo S, Osame S, Ohishi H. Studies on serum selenium and tocopherol in white muscle disease of foal. Nippon-Juigaku-Zasshi 1989;51:52–9. 39. Dill SG, Rebhum WG. White muscle disease in foals. Compend Cont Edu Practicing Vet 1985; 7:627–635. 40. Valberg SJ. A review of the diagnosis and treatment of rhabdomyolysis in foals. In: Proc AAEP 2002; 48:117–121. 41. Perkins G, Valberg SJ, Madigan JM, Carlson GP., Jones SL. Electrolyte disturbances in foals with severe rhabdomyolysis. J Vet Intern Med 1998;12:173–7. 42. Gunn HM. A comparison of the proportion of muscle, bone and fat and the distribution of muscle between thorough breds and other horses as adults and during growth. In: Equine exercise physiology 2 (eds Gillespie JR and Robinson NE). ICEEP Publications, Davis, CA. pp 253–64. 43. Stoker JW. Monensin sodium in horses (coccidiostat toxicity). Vet Rec 1975;97:137–8. 44. Kart A. Ionophore antibiotics: toxicity, mode of action and neurotoxic aspect of carboxylic ionophores. J Anim Vet Adv 2008;7:748–51. 45. Perkins S, Magdesian G, Thomas WP, Spier SJ. Pericarditis
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Acquired Abnormalities of the Cardiovascular System and pleuritis caused by Corynebacterium pseudotuberculosis in a horse. J Am Vet Med Assoc 2004;224:1133–8. 46. Seahorn JL, Slovis NM, Reimer JM, Carey VJ, Donaghue JG, Cohen ND. Case-control study of factors associated with fibrinous pericarditis among horses in central Kentucky during spring 2001. J Am Vet Med Assoc 2003;223:832–8. 47. Bolin DC, Donahue JM, Vickers ML, Harrison L, Sells S, Giles RC, Hong CB, Poonacha KB, Roberts J, Sebastian MM, Swerczek TW, Tramontin R, Williams NM. Microbiologic and pathologic findings in an epidemic of equine pericarditis. J Vet Diagn Invest 2005;17:38–44. 48. Birks EK, Hultgren BD. Pericardial haemangiosarcoma in a horse. J Comp Path 1998;99:105–7.
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49. Carrime BL, Schneider G, Cook JE, Leipold HW. Pericardial mesothelioma in a horse. Vet Pathol 1977;14:513–15. 50. Baker JR, Kreeger J. Infiltrative lipoma in the heart of a horse. Cornell Vet 1987;77:258–62. 51. Frye FL, Knight HD, Brown SI. Hemangiosarcoma in a horse. J Am Vet Med Assoc 1983;182:287–9. 52. Weldon AD, Step DL, Moise NS. Lymphosarcoma with myocardial infiltration in a mare. Vet Med 1992;87:595– 8. 53. Jeffcott LB. Primary liver-cell carcinoma in a young Thoroughbred horse. J Pathol 1969;97:394–7. 54. Richardson DW, Kohn CW. Uroperitoneum in the foal. J Am Vet Med Assoc 1983;182:267–71.
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58.
Endocrine Abnormalities Melissa T. Hines
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Chapter 58
Endocrine Abnormalities Melissa T. Hines
The hypothalamic-pituitary-adrenal axis in foals Activation of the hypothalamic-pituitary-adrenal (HPA) axis is one of the major adaptive responses to stress. The HPA axis responds to stress largely by regulating the secretion of cortisol, the primary stress hormone. Essentially, physiological stressors, such as pain, hypotension, metabolic abnormalities, or tissue injury, result in activation of the nervous system and the release of cytokines. These signals of stress are integrated in the hypothalamus, resulting in the release of corticotropin-releasing hormone (CRH). The CRH then stimulates the release of adrenocorticotropic hormone (ACTH or corticotropin) by the anterior pituitary gland, which in turn stimulates the release of cortisol from the adrenal cortices. While the secretion of cortisol is primarily regulated via this pathway, other factors play a role as well. For example, in horses, arginine vasopressin (AVP) produced by the hypothalamus acts as an ACTH secretagogue in addition to its osmoregulatory and pressor functions.1, 2 Approximately 90–95% of the cortisol released from the adrenal gland into the circulation becomes protein-bound. The majority is bound to corticosteroid-binding globulin (CBG or transcortin), a high-affinity plasma protein transporter, while a small percentage is bound to albumin. It is the unbound, free fraction of cortisol that is biologically active. Cortisol has a number of systemic effects that are essential to the adaptation to physiological stressors. While the functions of cortisol are complex, some of the major effects involved in restoring homeostasis include the maintenance of blood pressure, provision of energy to tissues, and regulation of the inflammatory response. The HPA axis in neonatal foals is dynamic in nature, and its function varies with age as well as
between individuals. Evidence suggests that maturation of the HPA axis in foals occurs late in development, beginning during the last 4 to 5 days prior to parturition and continuing during the first 1 to 2 weeks of life.3–6 Plasma concentrations of ACTH peak approximately 5–10 minutes after parturition and decline to adult concentrations by 6–12 hours of age (Table 58.1). As a result of the late rise in ACTH, serum concentrations of cortisol remain low until immediately prior to parturition, which is later in gestation than in some other species, such as lambs. Cortisol concentrations tend to peak at approximately 30–60 minutes postpartum and decline within 6–12 hours. The concentrations during the first year of life generally remain lower than in adult horses. ACTH stimulation tests, which are widely accepted as a means to evaluate adrenal function, have been performed in foals, utilizing both high, supraphysiological doses and low, physiological doses of ACTH (Table 58.2).7–12 Recently, a paired lowdose/high-dose ACTH stimulation test was evaluated in healthy neonatal foals of various ages.8 As found in other studies, normal neonatal foals responded to stimulation with both low and high doses of exogenous ACTH, but their response was blunted in comparison to both neonates of other species and mature horses (Table 58.1).The highest response occurred during the first few days of life, and as with baseline ACTH and cortisol concentrations, the ACTH responsiveness declined in the days after birth. In addition to this decreased response to ACTH, the release of cortisol in response to either insulin-induced hypoglycemia or acute hypotension appears to be diminished in foals as compared to adult horses.1, 3 Data in foals related to the percentage of circulating cortisol that is free rather than protein-bound are limited. However, as compared to adults and infants, foals may have a lower protein-binding
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Endocrinology System Table 58.1 Hypothalamic–pituitary–adrenal axis assessment in normal neonatal foals: baseline plasma cortisol and ACTH concentrations and response to a paired low-dose (LD)/high-dose (HD) ACTH stimulation test8 Age
Birth (n = 11)
12–24 h (n = 10)
36–48 h (n = 10)
5–7 days (n = 11)
Basal cortisol (µg/dL) Basal ACTH (pg/mL) ACTH : cortisol ratio LD delta cortisol (µg/dL) HD delta cortisol (µg/dL)
10.2 ± 2.3 285.5 ± 284.8 27.4 ± 23.8 3.6 ± 2.0 6.4 ± 4.1
3.6 ± 1.6 19.6 ± 5.3 6.2 ± 2.0 5.5 ± 2.1 9.4 ± 3.5
2.6 ± 1.0 32.7 ± 26.4 13.9 ± 7.4 3.5 ± 3.5 6.8 ± 3.4
2.0 ± 0.8 33.8 ± 23.1 15.9 ± 5.2 1.3 ± 0.6 3.5 ± 1.3
capacity for cortisol and thus a greater free cortisol fraction.13 In adult horses, the free cortisol fraction is approximately 10%, while in foals it is 53% at birth, declining to 40% by 5–7 days of age. In human infants, the percentage of free cortisol is approximately 30% at birth, gradually decreasing to 19% by 3 months of age.67 The relative immaturity of the HPA axis in neonatal foals may impact the ability of foals to respond to stress and disease. Even normal neonatal foals may have a diminished response to stressors as compared to adults. Recently, there has been interest in the possible role of the HPA axis in abnormal foals, such as those with prematurity or sepsis. Function of the HPA axis in premature foals The HPA axis plays an important role in fetal maturation. The intrafetal administration of ACTH has been shown to cause precocious maturation of the equine fetus and a significant reduction in gestational length.3, 14, 15 This is most likely mediated via the adrenal regulation of fetal maturation and production of maternal progestagens. Limited studies in premature foals suggest that there is dysfunction of the HPA axis and cortisol insufficiency, which is consistent with both the observation that maturation of the HPA axis occurs late in development and with findings in premature human infants.4, 5, 10 As compared to term foals, premature foals (gestational age < 320 days) have been shown to have lower concentrations of serum cortisol (18 ng/mL vs 80 mg/mL) and higher concentrations of endogenous ACTH (400 pg/mL vs 250 pg/mL).4 These findings suggest that the low
concentration of cortisol results from a failure at the level of the adrenal gland rather than a central problem, and this is supported by the diminished response of premature foals to ACTH stimulation tests. On the first day of life, premature foals were found to have only a 28% increase in plasma cortisol following the administration of exogenous ACTH as compared to a 208% increase in term foals.5 Premature foals have also been shown to have other abnormalities consistent with cortisol insufficiency, including low neutrophil to lymphocyte ratios, hypoglycemia, and hypoinsulinemia. Relative adrenal insufficiency in sick foals Relative adrenal insufficiency (RAI) has been defined as a cortisol response that is inadequate for the degree of illness.16, 17 This dysfunction, also referred to as critical illness-related corticosteroid insufficiency (CIRCI), occurs secondary to a primary severe illness or stress, and while the exact mechanisms are poorly understood, it is likely to be a multifactorial problem influencing several levels of control of cortisol secretion and action.16–19 Due to the important immunomodulatory, anti-inflammatory, and pressor effects of cortisol, RAI can lead to hypotension and overwhelming inflammation, with the end result in some cases being severe systemic inflammatory response syndrome (SIRS) or multiple organ dysfunction syndrome (MODS).
Diagnosis of RAI The accurate diagnosis of RAI can be difficult due to the many host, pathogen, and environmental factors
Table 58.2 Examples of protocols used for assessment of the hypothalamic–pituitary–adrenal axis in foals: paired low-dose/high-dose ACTH stimulation test and low-dose ACTH stimulation test8, 12 Paired low-dose/high-dose ACTH stimulation test
Time 0: measure baseline ACTH and cortisol: administer 10 µg cosyntropin IV; 30 min: measure cortisol; 90 min: measure cortisol; administer 100 µg cosyntropin IV; 120 min: measure cortisol; 180 min: measure cortisol
Low-dose ACTH stimulation test
Time 0: measure baseline ACTH and cortisol administer 0.1 pg/kg cosyntropin IV; 30 min: measure cortisol; 60 min: measure cortisol
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Endocrine Abnormalities that affect the physiological response to stress and infection. In human medicine, some current guidelines for establishing a diagnosis of RAI include basal serum cortisol levels below those commonly induced by stress in healthly people and/or a blunted cortisol response to either a high-dose or low-dose ACTH stimulation test.8, 16, 20 It has been suggested that the low-dose ACTH stimulation test is more physiological and may more accurately diagnose RAI in human patients than the high-dose test; however, this may not be the case in neonatal foals.21–23 Other diagnostic tests, such as measurement of free rather than total cortisol concentrations or an insulin tolerance test, may also be useful in assessing RAI.
Significance of RAI The significance of RAI has been investigated in multiple species. In human patients, including pediatric patients, HPA dysfunction has been shown to be associated with an increased disease severity and mortality.16, 20, 24 For example, in one study of 189 human patients with septic shock, patients with RAI had a mortality rate of 82% as compared to a mortality rate of 26% in septic patients who did not meet the criteria of RAI.20 Limited data in critically ill dogs also suggest that RAI is associated with an increased incidence of hypotension and a decreased survival rate.25 Importantly, in some human patients with RAI, treatment with low, physiological doses of corticosteroids has been shown to be beneficial.26, 27
RAI in critically ill foals Neonatal foals may have a high risk of RAI due to the relative immaturity of the HPA axis and the prevalence of sepsis, which is commonly associated with RAI in human critical care. In 1998, Couteil et al. reported a case of transient adrenal insufficiency in a foal with septicemia and hypovolemic shock.28 The foal had a low basal serum cortisol concentration and a poor cortisol response to a high-dose ACTH stimulation test. The foal was treated with supportive care, antimicrobial therapy, and a tapering course of prednisolone supplementation. The foal survived and had a normal ACTH stimulation test at 2 months of age. Gold et al. demonstrated in septic foals that the ACTH : cortisol ratio was elevated in non-survivors (59.00 ± 79.01) as compared to survivors (27.26 ± 48.01), providing additional evidence for HPA axis dysfunction in severely ill septic foals, possibly at the level of the adrenal gland.29 While most septic foals in this study had relatively high ACTH and cortisol concentrations as compared to healthy foals, some septic foals had normal or low ACTH concentrations in the face of low cortisol con-
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centrations, suggesting a poor pituitary response to stress. Recently, function of the HPA axis was assessed in 72 hospitalized neonatal foals less than 7 days of age and correlated with disease severity and outcome.22, 30 Seventy six percent of the foals in the study met the criteria for sepsis, which was defined by a sepsis score ≥ 11 and/or a positive blood culture. Basal ACTH and cortisol concentrations were measured in all 72 foals, and a paired low-dose/highdose ACTH stimulation test was performed in 58 foals. Based on low basal concentrations of cortisol, HPA dysfunction was diagnosed in 49% of the entire foal population and 37% of septic foals, while based on inadequate responsiveness to ACTH, HPA dysfunction was diagnosed in 52% of the entire foal population and 41% of septic foals. When all foals were considered, increased baseline cortisol concentrations were correlated with disease severity and non-survival. In the septic foals, diminished responsiveness to the high-dose ACTH stimulation test was correlated with non-survival. Also, 90% of the septic foals with both an increased basal serum cortisol concentration and a low ACTH responsiveness did not survive, as compared to a non-survival rate of only 9% of foals with a high cortisol concentration and an appropriate ACTH response. In a separate study of 52 ill foals, there were no differences in baseline cortisol or ACTH concentrations or in the ACTH to cortisol ratio either between ill foals and healthy controls or between ill foals with and without septicemia.23 Wide individual variation in values was noted. The majority of ill foals responded to a low-dose stimulation test; however, the cortisol response was significantly higher in foals that survived as compared to those that died or had euthanasia performed. Septic foals have also been shown to frequently have elevated concentrations of AVP, which acts to stimulate ACTH along with its pressor functions.19, 22, 31 Foals with the highest AVP concentrations had the highest ACTH concentrations and were more likely to die. Other endocrine abnormalities A number of other endocrine abnormalities may exist in foals, either associated with HPA dysregulation or as separate entities. Abnormalities of glucose homeostasis are common in neonates of all species, especially in cases of prematurity and critical illness. In foals, insulin can be detected in fetal plasma as early as 150 days of gestation, and pancreatic beta-cells are responsive to changes in glucose concentration during the last trimester.32–34 At birth, the concentrations of insulin (∼ 15 uU/mL) and glucose (∼ 75 mg/dL) are lower than in adult horses (∼ 24 uU/ml and 104 mg/dL, respectively). These concentrations
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increase in the first 24–48 hours of life. In foals of all ages, there is a positive correlation between the concentrations of insulin and glucose and the beta-cells response to changes in glucose concentration. However, both the release of insulin and the subsequent glucose clearance appear to be slightly diminished during the first 24 hours of life. Abnormalities of glucose regulation, resulting in either hypo- or hyperglycemia, occur frequently in premature or sick neonatal foals. In premature foals, the concentrations of insulin (∼ 9 uU/mL) and glucose (∼ 42 mg/dL) tend to be lower than in term foals.33 In a study of 515 critically ill foals, only 29.1% had blood glucose concentrations within the reference range at presentation (76–131 mg/dL), while 36.5% were hyperglycemic and 34.4% were hypoglycemic.35 Foals with glucose concentrations that were either < 10 mg/dL or > 180 mg/dL at admission were less likely to survive. In addition, as compared to surviving foals, non-surviving foals had lower mean blood glucose concentrations at admission, higher maximum and lower minimum concentrations during the first 24 hours of hospitalization, and higher concentrations at 24 and 36 hours. Hypoglycemia at admission was associated with sepsis, a positive blood culture, and systemic inflammatory response syndrome (SIRS). Given the importance of glucose in metabolism, regulating blood glucose concentrations via either glucose infusions or insulin therapy may be beneficial in managing sick foals. Dosages of insulin that have been suggested for glucose control in foals are 0.15 U/kg given intramuscularly or subcutaneously every 12 hours and 0.00125–0.2 U/kg/h given intravenously as a continuous rate infusion.36 However, the value of neonatal insulin replacement therapy is still under investigation in infants, and more data are needed in foals.37 Abnormalities of calcium regulation also occur in foals. A syndrome of idiopathic hypocalcemia has been described in five Thoroughbred foals ranging in age from 4 days to 5 weeks.38 Signs included tachycardia, hyperhidrosis, diarrhea, and muscle rigidity or a stiff gait. In four of the foals, recumbency, seizure-like activity, or pronounced extensor muscle rigidity were also seen. The foals did not respond to oral calcium supplementation and all foals died or had euthanasia performed. Critical illness in human patients has been associated with disorders of calcium regulation and the same may be true in foals. In a prospective study of foals < 7 days of age, septic foals were more likely to have disorders of calcium regulation than healthy foals.39 Concentrations of calcium, magnesium, phosphorus, parathyroid hormone (PTH), calcitonin, and parathyroidrelated peptide were measured in 82 septic (blood culture positive or sepsis score ≥14), 40 sick non-
septic, and 24 healthy foals. Septic foals had significantly decreased concentrations of ionized calcium (5.6 vs 6.1 mg/dL), with increased concentrations of PTH (16.2 vs 3.2 pmol/L) and phosphorus (7.1 vs 6.3 mg/dL). Among septic foals, non-survivors had higher PTH concentrations than survivors (41.1 vs 10.7 pmol/L).
The hypothalamic–pituitary–thyroid axis in foals Basic physiology of the thyroid gland The production of thyroid hormones is regulated by thyroid-releasing hormone (TRH) from the hypothalamus which stimulates the anterior pituitary to release thyroid-stimulating hormone (TSH, thyrotropin). TSH then regulates the synthesis and release of the thyroid hormones from the thyroid gland. Circulating thyroid hormones feed back on both the hypothalamus and anterior pituitary to regulate thyroid hormone concentrations. A wide variety of factors influence thyroid hormone production, such as age, dietary factors, climate, activity, pharmaceuticals, and systemic illness. The thyroid hormones are stored within the follicles of the thyroid gland as part of thyroglobulin. When thyroid hormones are needed, thyroxine (T4 ) and triiodothyronine (T3 ) are cleaved from thyroglobulin and enter the circulation. Once in the circulation, most thyroid hormones become proteinbound, primarily to thyroid-binding globulin, but also to T4 binding prealbumin and albumin. The thyroid hormones are slowly released from the protein, crossing the capillary endothelium primarily in the unbound or free state. The major thyroid hormones found in the circulation include the following.
r
r r
T4 (thyroxine): Approximately 90% of the thyroid hormone released from the thyroid gland is T4 . T4 is less biologically active than T3 , and ultimately most is converted to T3 by peripheral deiodination. T3 (triiodothyronine): Accounting for about 10% of the hormone secreted by the thyroid gland, most T3 is formed by peripheral deiodination. It is about 3–5 times as active as T4 . rT3 (reverse T3): The major product of peripheral deiodination is T3 ; however, some rT3 is formed by deiodination of the carboxyl end. The precise function of rT3 is unclear. In humans, concentrations are high in the fetus and patients with systemic illness.
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Table 58.3 Thyroid hormone concentrations in foals of various ages43–48 Age
Total T4
Total T3
Birth 1 day 7 days 14 days 1 month 6 months 1–10 hours 4 days 4 days 28 days
233 ng/mL 207 ng/mL 92 ng/mL 49 ng/mL 26 ng/mL 35 ng/mL 28.86µg/dL 11.2µg/dL 231.7 ± 61.8 nmol/L 30.6 ± 17.4 nmol/L
7.9 ng/mL 6.7 ng/mL 4.2 ng/mL 2.4 ng/mL 1.6 ng/mL 0.9 ng/mL 991 ng/dL 935 ng/dL 7.8 ± 4.2 nmol/L 3.1 ± 0.4 nmol/L
fT4
fT3
12.12 pg/mL
2.99 pg/mL
1.2 ± 0.4 ng/dL
3.4 ± 1.1 pg/mL
T4 thyroxine; T3 triiodothyronine; fT4 free thyroxine; fT3 free triiodothyronine
r
r
fT3 and fT4 (free T3 and free T4 ): Only about 1% of the total concentration of thyroid hormone in the circulation is unbound to carrier proteins. It is this unbound hormone that is biologically active and acts in the feedback loop to regulate the synthesis and release of thyroid hormones. Other derivatives of thyroid hormones include 3-iodothyronamine (T1 a) and thyronamine (T0 a), which result from further decarboxylation and deiodinataion of thyroid hormone.
The thyroid hormones are essential throughout the body. They primarily act by increasing the nuclear transcription of multiple genes, leading to a general increase in protein synthesis and metabolic activity. Some specific functions of the thyroid hormones include maintenance of the basal metabolic rate, neural transmission, and thermogenesis, as well as stimulation of the heart rate, cardiac output, and blood flow. In the fetus and neonate, the thyroid hormones promote differentiation and play an important role in overall growth and development, especially of skeletal and neuronal tissues. The accurate assessment of thyroid gland function can be challenging. Single measurements of resting concentrations of thyroid hormones are often misleading due to both variation in normals and the influence of extrathyroidal factors. Both TSH and TRH stimulation tests have been advocated for the diagnosis of thyroid disease, but unfortunately approved TRH and TSH products are often unavailable or too costly to be practical. Measurement of serum TSH concentration has proven to be a reliable means of diagnosing hypothyroidism in people, but has not been as consistent in dogs. TSH concentrations in neonatal foals have been evaluated, but data remain limited.40, 41 Aspiration or biopsy of the thyroid gland may help establish a pathological process, but the high vascularity of the thyroid should be considered. Other means of assessing the thyroid include scinti-
graphic imaging, ultrasonography, and measurement of the basal metabolic rate.
Thyroid function in neonatal foals Thyroid hormones are clearly important in fetal development in foals as in other species. In a study by Allen et al., four foals were partially thyroidectomized at approximately 215 days of gestation.42 These foals had both abnormal behavior and locomotor skills as well as numerous abnormalities in their skeletal development. Postnatal changes in thyroid hormone concentrations have been studied in neonatal foals (Table 58.3).43–48 While the specifics vary somewhat depending on which thyroid hormone is assessed, thyroid hormone concentrations are significantly higher in neonatal foals than adult horses, with concentrations gradually declining over several months. Also, the concentrations of thyroid hormones in foals tend to be higher than in neonates of other species. Baseline serum TSH concentrations in normal foals are not significantly different from adult horses.40, 41 Consistent with the high thyroid hormone production, neonatal foals are fully capable of responding to TSH.49 The high concentrations of thyroid hormones in foals may be responsible for their high thermogenic capacity and rapid pre- and postnatal rates of growth, especially of the musculoskeletal and nervous systems.
Hypothyroidism in foals Several syndromes of thyroid dysfunction have been identified in neonatal foals. An immature hypothalamic–pituitary–thyroid axis may contribute to decreased thyroid function in premature foals. Also, as in all species, thyroid function in foals is affected by numerous extrathyroidal factors, especially concurrent illness. In addition, several specific
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syndromes of congenital hypothyroidism have been identified in neonatal foals.
Thyroid function in premature and sick foals Premature neonatal foals may be transiently hypothyroid since thyroid function peaks late in gestation. In one study, serum concentrations of total and free thyroid hormones were decreased in premature foals as compared to normal term foals.41 While baseline serum concentrations of TSH were not different between the groups, the TSH responses to TRH were exaggerated in the premature foals. Based on these findings, it was hypothesized that thyroid hormone supplementation in premature foals might accelerate organ system maturation and thus improve survival. A study in 23 newborn foals and 5 fetuses catheterized in late gestation demonstrated a relationship between circulating cortisol and T3 and that prematurity is associated with low concentrations of both hormones.50 Systemic illness has been shown in multiple species to cause decreases in thyroid hormones, especially T3 . This syndrome, known as euthyroid sick syndrome or non-thyroidal illness syndrome, has been investigated in foals. In one study, concentrations of T3 were shown to be lower in sick foals than in healthy foals, approximating the diminished concentrations found in premature foals.41 Concentrations of T4 in sick foals were intermediate between those concentrations found in premature and normal
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foals. In another study of 16 neonatal foals stressed by disease, 7 had low concentrations of T3 , 12 had low concentrations of T4 , and 10 had low concentrations of rT3 .51
Congenital hypothyroidism and dysmaturity A syndrome of hypothyroidism in foals known as congenital hypothyroidism and dysmaturity (CHD) or thyroid hyperplasia and musculoskeletal deformities has been described in association with a number of abnormalities, primarily in the musculoskeletal system.52–55 While the syndrome is primarily identified in foals born at term, it has also been identified as a cause of abortion.53 First reported in the early 1980s in foals from the western provinces of Canada, it was originally thought that the syndrome might be limited to this region.54 However, while it may be more common in certain geographic locations, cases have been reported from several areas in the United States and in Finland.53, 56 Due to the often guarded prognosis, especially for soundness, and the fact that multiple foals may be affected in a given year, the syndrome can be devastating. The predominant clinical characteristics of CHD include prolonged gestation (average 360 days), mandibular prognathism, hypo-ossification, and contracted tendons, often with accompanying rupture of the common digital extensor tendon (Figures 58.1 and 58.2).42, 53, 54,56–58 In addition, despite the gestation length, some affected foals may have signs of
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Figure 58.1 Foals with congenital hypothyroidism and dysmaturity (CHD). (a) This foal has severe tendon contracture, which is one of the classical clinical signs of the syndrome. (b) Another classical sign of CHD is mandibular prognathism, which is mild in this case. While prolonged gestation is another common feature of the syndrome, affected foals often display signs of immaturity.
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Figure 58.2 Radiographs of (a) carpi and (b) hock of a foal with severe hypo-ossification of the cuboidal bones associated with congenital hypothyroidism and dysmaturity (CHD). The severity of signs varies widely between cases.
immaturity such as a soft, silky hair coat and pliable ears. Also, incomplete closure of the abdominal wall and poor muscle development, particularly evident over the sternum, have been described. While some foals are weak and non-responsive at birth, many are bright and alert with a strong suckle reflex. Importantly, there is generally no palpable enlargement of the thyroid glands, although there is histological evidence of thyroid hyperplasia. The severity of signs varies widely among cases. Complete blood counts, serum chemistries, and electrolytes are normal if the complications of failure of passive transfer and septicemia are avoided.52–54,56, 57 Baseline concentrations of T3 and T4 may be either low or within normal ranges for neonatal foals, but there is a diminished response to TSH.56, 57 Dams have been found to have normal thyroid function at the time of parturition. Biopsy of the thyroid gland reveals marked thyroid hyperplasia and can be used to confirm the diagnosis (Figure 58.3). Treatment of CHD has been largely supportive: prevention of failure of passive transfer, nutritional supplementation, splints, physical therapy, and restricted exercise.52–54,56 It is unknown if supplementation of thyroid hormones is of any benefit in cases of CHD. Limited data suggest that surviving affected foals become responsive to TSH over time even without thyroid supplementation, and the thyroid tissue becomes histologically normal.56 The prognosis for foals with CHD varies with the severity of the condition and whether concur-
rent septicemia develops. A variety of orthopedic problems have been identified in surviving foals, including tarsal collapse, angular limb deformities, physitis, and lesions associated with osteochondrosis. The prognosis for soundness is often poor in foals with CHD unless the musculoskeletal deformities are mild. The underlying defect in CHD appears to be hypothyroidism, which is supported by the observation that thyroidectomy in foals and in other species results in a similar clinical syndrome.42, 59 However, in most cases, relatively little is known about the specific cause of the impaired thyroid function. In general, the occurrence appears sporadic, and as there is no sex or breed predisposition, a genetic cause is considered unlikely. Since many factors potentially influence thyroid function, the insult to the fetal thyroid might not be the same in all cases. In the Pacific Northwest, foals that are at risk tend to be born in the latter half of the foaling season.56 Currently, an exposure-related cause is suspected for CHD and theories based primarily on epidemiological studies include exposure to high concentrations of nitrate or exposure to mustard plants.52, 53, 56 The active pump that transfers iodide ions into the thyroid cells can also pump nitrate ions, and thus high concentrations of nitrate can cause competitive inhibition of iodide transport into the cell. Plants in the mustard family contain varying amounts of glucosinolate, a known thyrotoxic compound.60 Thus exposure to high concentrations of either of these compounds could impair thyroid function and contribute to CHD.
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Figure 58.3 (a) The thyroid gland of a normal neonatal foal. Note the colloid-containing follicles. (b) The thyroid gland of a foal affected with congenital hypothyroidism and dysmaturity (CHD). There is a lack of normal colloid with diffuse, severe hyperplasia of thyroid follicular epithelial cells.
Epidemiological studies have also suggested that inadequate mineral supplementation may be a risk factor for the syndrome.52, 56 Deficiencies of minerals such as iodine and selenium may adversely affect thyroid function.
Other causes of hypothyroidism in neonatal foals Syndromes of hypothyroidism in foals other than CHD have been identified, including both a deficiency or excess of dietary iodine and exposure to feed contaminated with fungi (Acremonium coenophialum, Claviceps purpurea).56, 61, 62 The thyroid glands in these cases are typically grossly enlarged and histological lesions vary slightly from those seen in foals affected with CHD in Canada and the Pacific Northwest. A dietary deficiency of iodine has been associated with goiter.44 In addition, high concentrations of dietary iodine interfere with several aspects of thyroid synthesis and release. There have been several case reports of foals with goiter due to excess dietary iodine, which has been associated in some cases with feeding seaweed as a supplement.63–65 Foals born to mares grazing endophyteinfected fescue have been weak with clinical signs of hypothyroidism, including musculoskeletal abnormalities.61, 62 In experimental studies, foals of mares grazing on Acremonium coenophialum-infected fescue pasture have had increased gestation duration, decreased serum T3 concentrations, and reduced sur-
vivability.61, 62 The concentrations of T4 and rT3 were not significantly different from control foals.61 Histologically, the thyroid glands of foals born to mares exposed to endophyte had large distended thyroid follicles lined by flat cuboidal epithelial cells.
Therapy for decreased thyroid function The effects of thyroid supplementation in foals with the various syndromes of thyroid dysfunction are unknown. In premature infants with decreased thyroid function, supplementation of thyroid hormones has yielded variable results, but has been beneficial in some cases. Anecdotal dosage recommendations for foals are 20–50 µg/kg/day for T4 (levothyroxine) and 1 mg/kg/day for T3 .66 During treatment, periodic monitoring of serum thyroid hormone concentrations is recommended. It has been theorized that prolonged supplementation could suppress endogenous hormone production. Thus, it has been suggested that if foals are treated with exogenous thyroid hormone, they be gradually weaned from supplementation.
Summary The endocrine system is critical in perinatal development and the maintenance of homeostasis. The role of the endocrine system is complex, and even in human medicine where large-scale clinical studies are more feasible, the diagnosis and appropriate management of endocrine abnormalities remain
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Endocrine Abnormalities controversial. More information relative to RAI and other endocrine disorders in neonatal foals is needed. In particular, the potential benefit of therapeutic intervention, such as corticosteroid, insulin or thyroid replacement therapy, needs to be investigated.
References 1. Alexander SI, Roud HK, Irvine CH. Effect of insulininduced hypoglycemia on secretion patterns and rates of corticotrophin-releasing hormone, arginine vasopressin and adrenocorticotrophin in horses. J Endocrinol 1997;153(3):401–9. 2. Livesey JH, Evans MJ, Mulligan R, Donald RA. Interactions of CRH, AVP and cortisol in the secretion of ACTH from perifused equine anterior pituitary cells: ‘permissive’ roles for cortisol and CRH. Endocr Res 2000;26(3):445–63. 3. O’Connor SJ, Gardner DS, Ousey JC, Holdstock N, Rossdale P, Edwards CM, Fowden AL, Giussani DA. Development of baroreflex and endocrine responses to hypotensive stress in newborn foals and lambs. Pflugers Arch 2005;450(5):298–306. 4. Rossdale PD, Ousey JC. Studies on equine prematurity 6: Guidelines for assessment of foal maturity. Equine Vet J 1984;16: 300. 5. Silver M, Ousey JC, Dudan FE, Fowden AL, Knox J, Cash RS, Rossdale PD. Studies on equine prematurity 2: Postnatal adrenocortical activity in relation to plasma adrenocorticotrophic hormone and catecholamine levels in term and premature foals. Equine Vet J 1984;16(4):278–86. 6. Webb PD, Steven DH. Development of the adrenal cortex in the fetal foal: an ultrastructural study. J Dev Physiol 1981;3(1):59–73. 7. Hart KA, Ferguson DC, Heusner GL, Barton MH. Synthetic adrenocorticotropic hormone stimulation tests in healthy neonatal foals. J Vet Intern Med 2007;21(2):314–21. 8. Hart KA, Heusner GL, Norton NA, Barton MH. Hypothalamic-pituitary-adrenal axis assessment in healthy term neonatal foals utilizing a paired low dose/high dose ACTH stimulation test. J Vet Intern Med 2009;23: 344–351. 9. Ousey JC, Fowden AL, Wilsher S, Allen WR. The effects of maternal health and body condition on the endocrine responses of neonatal foals. Equine Vet J 2008;40(7): 673–9. 10. Rossdale PD, Silver M, Ellis L, Frauenfelder H. Response of the adrenal cortex to tetracosactrin (ACTH1–24) in the premature and full-term foal. J Reprod Fertil Suppl 1982;32: 545–53. 11. Silver M, Fowden AL. Prepartum adrenocortical maturation in the fetal foal: responses to ACTH. J Endocrinol 1994;142(3):417–25. 12. Wong DM, Vo DT, Alcott CJ, Stewart AJ, Peterson AD, Sponseller BA, Hsu WH. Adrenocorticotropic hormone stimulation tests in healthy foals from birth to 12 weeks of age. Can J Vet Res 2009;73(1):65–72. 13. Barton MH, Hart K. Hypothalamic-pituitary-adrenal axis in healthy neonatal foals. In: Proceedings of the 26th American College of Veterinary Internal Medicine Forum, 2008; pp. 198–200. 14. Ousey JC, Rossdale PD, Dudan FE, Fowden AL. The effects of intrafetal ACTH administration on the outcome of pregnancy in the mare. Reprod Fertil Dev 1998;10(4):359–67.
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15. Rossdale PD, McGladdery AJ, Ousey JC, Holdstock N, Grainger L, Houghton E. Increase in plasma progesterone concentrations in the mare after foetal injection with CRH, ACTH or betamethasone in late gestation. Equine Vet J 1992;24(5):347–50. 16. Marik PE, Pastores SM, Annane D, Meduri GU, Sprung DL, Arit W, Keh D, Briegel J, Beishuizen A, Dimopoulou I, Tsagarakis S, Singer M, Shrousos GP, Zaloga G, Bokhari F, Vogeser M, American College of Critical Care Medicine. Recommendations for the diagnosis and management of corticosteroid insufficiency in critically ill adult patients: consensus statements from an international task force by the American College of Critical Care Medicine. Crit Care Med 2008;36(6):1987–9. 17. Venkataraman S, Munoz R, Candido C, Witchel SF. The hypothalamic-pituitary-adrenal axis in critical illness. Rev Endocr Metab Disord 2007;8(4):365–73. 18. Maxime V, Siami S, Annane D. Metabolism modulators in sepsis: the abnormal pituitary response. Crit Care Med 2007;35(9 Suppl):S596–601. 19. Toribio RE. Endocrine dysregulation in septic foals. In: Proceedings of the 26th American College of Veterinary Internal Medicine Forum, 2008; pp. 204–6. 20. Annane D, Maxime V, Ibrahim F, Alvarez JC, Abe E, Boudou P. Diagnosis of adrenal insufficiency in severe sepsis and septic shock. Am J Resp Crit Care Med 2006;174(12):1282–4. 21. Dicstein G, Saiegh L. Low-dose and high-dose adrenocorticotropin testing: indications and shortcomings. Curr Opin Endocrinol Diabetes Obes 2008;15(3):244–9. 22. Hart KA, Slovis NM, Barton MH. Hypothalamic-pituitaryadrenal axis dysfunction in hospitalized neonatal foals. J Vet Intern Med 2009;23: 901–13. 23. Wong DM, Vo DT, Alcott CJ, Peterson AD, Sponseller BA, Hsu WH. Baseline plasma cortisol and ACTH concentrations and response to low-dose ACTH stimulation testing in ill foals. J Am Vet Med Assoc 2009;234(1):126–32. 24. Clark L, Preissig C, Rigby MR, Bowyer F. Endocrine issues in the pediatric intensive care unit. Pediatr Clin North Am 2008;55(3):805–33. 25. Burkitt JM, Haskins SC, Nelson RW, Kass PH. Relative adrenal insufficiency in dogs with sepsis. J Vet Intern Med 2007;21(2):226–31. 26. Keh D, Sprung CL. Use of corticosteroid therapy in patients with sepsis and septic shock: an evidence-based review. Crit Care Med 2004;32(11 Suppl):S527–33. 27. Minneci PC, Deans KJ, Banks SM, Eichacker PQ, Natanson C. 2004. Corticosteroids for septic shock. Ann Intern Med 2004;141(9):742–3. 28. Couetil LL, Hoffman AM. Adrenal insufficiency in a neonatal foal. Equine Vet J 1995;27(2):140–6. 29. Gold JR, Divers TJ, Barton MH, Lamb SV, Place NJ, Mohammed HO, Bain FT. Plasma adrenocorticotropin, cortisol, and adrenocorticotropin/cortisol ratios in septic and normal term foals. J Vet Intern Med 2007;21(4): 791–6. 30. Hart KA, Barton MH. Stressed out? Relative adrenal insufficiency in critically ill foals. In: Proceedings of the 26th American College of Veterinary Internal Medicine Forum, 2008; pp. 201–3. 31. Hurcombe SD, Toribio RE, Slovis N, Kohn CW, Refsal K, Saville W, Mudge MC. Blood arginine vasopressin, adrenocorticotropin hormone, and cortisol concentrations at admission in septic and critically ill foals and their association with survival. J Vet Intern Med 2008;22(3):639–47.
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32. Fowden AL, Ellis L, Rossdale PD. Pancreatic beta cell function in the neonatal foal. J Reprod Fertil Suppl 1982;32: 529–35. 33. Fowden AL, Silver M, Ellis L, Ousey J, Rossdale PD. Studies on equine prematurity 3: Insulin secretion in the foal during the perinatal period. Equine Vet J 1984;16:286–91. 34. Holdstock NB, Allen VL, Bloomfield MR, Hales CN, Fowden AL. Development of insulin and proinsulin secretion in newborn pony foals. J Endocrinol 2004;181: 469–476. 35. Hollis AR, Furr MO, Meagdesian KG, Axon JE, Axon JE, Luslow V, Boston RC, Corley KT. Blood glucose concentrations in critically ill foals. J Vet Intern Med 2008;22: 1223–7. 36. Wilkins PA. Disorders of foals. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 3rd edn. St Louis: Saunders, in press. 37. Sinclair JC, Bottino M, Cowett RM. Interventions for prevention of neonatal hyperglycemia in very low birth weight infants. Cochrane Database Syst Rev 2009; (3):CD007615. 38. Beyer MJ, Freestone JF, Reimer JM, Bernard WV, Rueve ER. Idiopathic hypocalcemia in foals. J Vet Intern Med 1997;11: 356–60. 39. Hurcombe SD, Toribio RE, Slovis NM, Saville WJ, Mudge MC, Macgillivray K, Frazer ML. Calcium regulating hormones and serum calcium and magnesium concentrations in septic and critically ill foals and their association with survival. J Vet Intern Med 2009;23:335–43. 40. Berg EL, McNamara DL, Keisler DH. Endocrine profiles of periparturient mares and their foals. J Anim Sci 2007;85: 1660–8. 41. Breuhaus B. Thyroid function in normal, sick and premature foals. J Vet Intern Med 2005;19(3):445. 42. Allen AL, Fretz PB, Card CE, Doige CE. The effects of partial thyroidectomy on the development of the equine fetus. Equine Vet J 1998;30(1):53–9. 43. Chen DL, Riley AM. Serum thyroxine and triiodothyronine concentrations in neonatal foals and mature horses. Am J Vet Res 1981;6:1415–17. 44. Irvine CH. Hypothyroidism in the foal. Equine Vet J 1984;16(4):302–6. 45. Irvine CH, Evans MJ. Postnatal changes in total and free thyroxine and triiodothyronine in foal serum. J Reprod Fertil Suppl 1975;23: 709–15. 46. Murray MJ, Luba NK. Plasma gastrin and somatostatin, and serum thyroxine (T4), triiodothyronine (T3), reverse triiodothyronine (rT3) and cortisol concentrations in foals from birth to 28 days of age. Equine Vet J 1993;25:237–9. 47. Shaftoe S. Peripartum endocrinologic adaptation. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 40–54. 48. Toribio RE, Duckett WM. Thyroid gland. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis, MO: Saunders, 2004; pp. 1340–56. 49. Shaftoe S, Schick MP, Chen CL. Thyroid-stimulating hormone response tests in one-day-old foals. J Equine Vet Sci 1988;8:310–12. 50. Silver M, Fowden AL, Knox J, Ousey J, Cash R, Rossdale PD. Relationship between circulating tri-iodothyronine and cortisol in the perinatal period in the foal. In: Proceedings of the 5th International Symposium on Equine Reproduction, 1991; pp. 619–26.
51. Furr MO, Murray MJ, Ferguson DC. The effects of stress on gastric ulceration, T3, T4, reverse T3 and cortisol in neonatal foals. Equine Vet J 1992;24(1):37–40. 52. Allen AL, Townsend HG, Doige CE, Fretz PB. A case–control study of the congenital hypothyroidism and dysmaturity syndrome of foals. Can Vet J 1996;37(6):349–51, 354–8. 53. Allen AL, Doige CE, Fretz PB, Townsend HG. Hyperplasia of the thyroid gland and concurrent musculoskeletal deformities in western Canadian foals: reexamination of a previously described syndrome. Can Vet J 1994;35(1):31–8. 54. Doige CE, McLaughlin BG. Hyperplastic goiter in newborn foals in Western Canada. Can Vet J 1981;22(2):42–5. 55. Gawrylash SK. Thyroid hyperplasia and musculoskeletal deformity in a standardbred filly in Ontario. Can Vet J 2004;45(5):424–6. 56. Hines MT, Doles J, Gay C, Schott HC, Cudaback L. Thyroid hyperplasia and musculoskeletal deformity in neonatal foals. In: Proceedings of the 14th American College of Veterinary Internal Medicine Forum, 1996; pp. 544–5. 57. McLaughlin BG, Doige CE, McLauglin PS. Thyroid hormone levels in foals with congenital musculoskeletal lesions. Can Vet J 1986;27(7):264–7. 58. McLaughlin BG, Doige CE. A study of ossification of carpal and tarsal bones in normal and hypothyroid foals. Can Vet J 1982;23(5):164–8. 59. D’Amours GH, Taylor SM, Olfert ED, Simko E, Allen AL. Evaluation of experimental methods to induce congenital hypothyroidism in guinea pigs for use in the study of congenital hypothyroidism in horses. Am J Vet Res 2004;65(9):1251–8. 60. Vermorel M, Davicco MJ, Evvard J. Valorization of rapeseed meal. 3. Effects of glucosinolate content on food intake, weight gain, liver weight and plasma thyroid hormone levels in growing rats. Reprod Nutri Dev 1987;27(1A):57–66. 61. Boosinger TR, Brendemuehl JP, Bransby DL, Wright JC, Kemppained RJ, Kee DD. Prolonged gestation, decreased triiodothyronine concentration and thyroid gland histomorphologic features in newborn foals of mares grazing Acremonium coenophialum-infected fescue. Am J Vet Res 1995;56(1):66–9. 62. Putnam MR, Bransby DI, Schumacher J, Boosinger TR, Bush L, Shelby Ra, Vaughan JT, Ball D, Brendemuehl JP. Effects of the fungal endophyte Acremonium coenophialum in fescue on pregnant mares and foal viability. Am J Vet Res 1991;52(12):2071–4. 63. Drew B, Barber WP, Williams DG. The effect of excess dietary iodine on pregnant mares and foals. Vet Rec 1975;97(05):93–5. 64. Eroksuz H, Eroksuz y, Ozer H, Ceribasi AO, Yaman I, Ilhan N. Equine goiter associated with excess dietary iodine. Vet Hum Toxicol 2004;46(3):147–9. 65. Miyazawa K, Motoyoshi S, Usui K. Nodular goiters of three mares and their foals, induced by feeding excessive amount of seaweed. Nippon Juigaku Zasshi 1978;40(6):749–53. 66. Breuhaus G. Thyroid function and dysfunction in equine neonates. In: Proceedings of the 24th Annual Meeting of the American College of Veterinary Internal Medicine, 2006; pp. 162–4. 67. Rokicki W. Forest MG, Loras B, Bonnet H, Bertrand J. Free cortisol of human plasma in the first three years of life. Biol Neonate 1990;57(1);21–9.
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Ophthalmic Disorders of the Foal Caryn E. Plummer and Dennis E. Brooks
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Chapter 59
Ophthalmic Disorders of the Foal Caryn E. Plummer and Dennis E. Brooks
The eye examination The horse is a precocious animal and, as such, is born with fully developed eyes and open eyelids.1 Good functional vision is present in the normal animal since individuals in the wild must be able to flee predators practically immediately. Vibrissae are present above and below the eyes in the normal foal, as are long upper lid cilia, or eyelashes. These cilia should be oriented outward in the normal, comfortable eye. The nictitating membrane should be present and mobile and may or may not be pigmented. In front of the nictitans lies a well-developed caruncle at the medial canthus. The globes should be symmetrically mobile and the cornea should be the prominent structure visible through the open palpebral fissure. An ocular examination in a foal should proceed anterior to posterior in the same manner as an examination of an adult horse, and should include evaluation of all adnexal structures and the various parts of the orbit and globe. However, there are some important differences between young animals and adults that must be recognized in order to avoid misdiagnosis of a normal condition. The youngest foals are often easily examined with manual restraint alone. Sedation is usually necessary in the older individuals in order to perform a thorough examination of the eyes. The eyes of small individuals can often be examined without the aid of an auriculopalpebral block, which blocks the branch of the facial nerve that innervates the upper eyelid. The examination of older, stronger individuals will be greatly facilitated by this motor block. The sensory innervation of the face of foals is well-developed early on and may even be hypersensitive in newborns.1, 2 A thorough ocular examination begins with an evaluation of the pupillary light reflexes (PLRs),
the menace response, and the dazzle reflex.1,3–6 The PLRs at birth are present, but sluggish, and may not acquire the normal rapid response typical of an adult horse until 5 days of age. It is important to use a very bright light source to assess the PLRs. A poor PLR may result from insufficient photic stimulation from a poor light source, or severe retinal disease. The pupils of neonatal foals are round to slightly ovoid in shape initially, but will assume the typical horizontal oval shape by 3 to 5 days after birth. The menace response is a learned response and is, therefore, not present at birth. It usually develops within 5 to 9 days, but may not be present in some individuals until 2 weeks after birth. Rapid hand motion should elicit a blink response if the menace reflex is present. The menace response is a crude assessment of vision. Shining a very bright light into the pupil will result in a blink response in testing the dazzle reflex. The dazzle reflex is a test of light perception and is a sensitive indicator of retinal function. Often, visual deficits in foals are not apparent until the foal begins to assert its independence or has been weaned, since the mare acts as a very efficient guide.3 Occasional neonates will have an obvious blindness that is noted soon after birth (< 48 hours old, bumping into objects when not in direct contact with the dam). It resolves in time (3–7 days). The cornea should be smooth and clear, free of opacities or defects (Figure 59.1). In foals, the average corneal diameters are 26.1 mm horizontally and 19.8 mm vertically. This compares to 34 mm across the horizontal axis and 26.5 mm across the vertical axis of the cornea in the average adult horse. Corneal sensitivity in foals is increased when compared to normal adults; however, sick foals have diminished corneal sensitivity which may predispose them to acquired corneal disease.2 Corneal cytological
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 59.1 Heterochromic iris with blood vessels present in the iridocorneal angle of a normal foal eye. The dorsal corpora nigra are small. (See color plate 7).
specimens can be obtained with topical anesthesia and a metallic instrument such as the handle end of a scalpel blade. Cotton swabs generally do not provide enough material for examination. The iridocorneal angle, with its porous meshwork appearance, should normally be visible along the temporal and nasal aspects of the corneoscleral junction (Figures 59.1 and 59.2). The intraocular pressure is measured with topical anesthesia and a handheld tonometer. The irides of most horses are moderate brown in color; however, blue irides, or partially blue irides are not uncommon and are normal.1 Color-dilute animals, such as Palominos, may have light brown irides as a manifestation of their color variation. Corpora nigra are cystic structures usually present along the dorsal margin of the pupil and
Figure 59.2 The iridocorneal angle is gray and open in this normal foal. (See color plate 8).
Figure 59.3 Y-suture opacity is present in this eye. (See color plate 9).
may be present, albeit small, ventrally as well. The lens should be clear and its anterior surface smooth. The lens sutures may be visible in the normal eye as stellate or ‘Y’-shaped linear shapes within the lens. The sutures at the anterior surface of the lens, if ‘Y’-shaped, tend to be upright, while the suture lines along the posterior surface are inverted. Lens sutures may be misdiagnosed as cataract if they are prominent1 (Figure 59.3). A remnant of the hyaloid system, the vascular system that feeds the intraocular structures during fetal development, is often present along the axial posterior surface of the lens in newborn foals (Figure 59.4). This remnant is easily seen in the first day of life and may appear as fine, wavy gray lines oriented between the posterior aspect of the lens and the optic nerve head. Some hyaloid remnants, however, may be black or appear as obviously blood-filled vessels traversing the vitreal
Figure 59.4 A vascular remnant of the hyaloid system is present posterior to the lens in this foal. (See color plate 10).
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Ophthalmic Disorders of the Foal
Figure 59.5 A normal optic nerve head and retina are present in this foal. (See color plate 11).
cavity. These hyaloid structures will regress rapidly over the first few days to weeks after birth. Aside from the hyaloid structures, the vitreous should be free of any opacities. It is a clear, gel-like medium through which light travels after having been bent by the focusing abilities of the lens. The ocular fundus of the foal is similar to that of the adult horse (Figure 59.5). The optic disc or optic nerve head is located in the non-tapetal fundus, just ventral to the junction of the tapetal and non-tapetal portions of the fundus. The main difference between the foal and the adult fundus is the color of the optic nerve head, which in foals tends to appear more red than the salmon color of the adult disc. The optic disc is typically ovoid in shape, but may be slightly rounded. The retinal vascular pattern is paurangiotic, in which 40–60 small retinal vessels radiate from the periphery of the optic nerve head and extend one to two disc diameters from the nerve head. There is a segment at the 6 o’clock position in the normal horse optic nerve head that is avascular and appears as a white notch. There is vast variation in the tapetal color and size in horses regardless of age. Most of this variation is related to the genetics dictating coat color and the overall level of pigmentation of the individual. Most horses have a yellow-green tapetum, but the range varies from yellow to blue. End-on choroidal vessels, known as stars of Winslow, are visible in the normal tapetal fundus as regularly spaced brown-black dots.1, 3, 5
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in order to provide the most accurate prognosis and make the most appropriate treatment and breeding decisions for that individual. In particularly young foals, acquired lesions may often be misinterpreted as being congenital. Congenital defects are usually the result of some intrauterine insult, hereditary factors, or an idiopathic insult of ocular development. Knowledge of embryology often allows identification of the developmental defect that leads to the ocular deformity; however, identification of a specific etiological agent in such cases is rarely possible. In utero causes may include maternal infections (bacterial or viral) or inflammatory processes, trauma, toxins, drugs, nutritional deficiencies or excesses, radiation, and idiopathic forces. Regardless of the mechanism responsible, the most important determinant of the severity of the abnormality is the stage of development during which the insult occurs, with the more severe ocular anomalies resulting from earlier insults. Inherited ocular defects can either be present at birth or develop some time later. Many ocular anomalies may be the result of a genetic predisposition encouraged by an environmental factor or set of factors.1, 3, 5 Fortunately, the overall incidence of congenital ocular anomalies is relatively low in foals. The remainder of this chapter will discuss disorders of the orbit, eyelid, nasolacrimal system, cornea, uvea, lens, and posterior segment that have been documented in foals.
Diseases and disorders of the globe and orbit Congenital abnormalities
Microphthalmia One of the more common congenital ocular defects noted in horses is microphthalmia1, 3, 5 (Figure 59.6) wherein the globe is reduced in size to at least two standard deviations below the mean. It represents
Ocular abnormalities: overview Ocular abnormalities in the foal are classified as congenital, acquired, or inherited.1, 3, 5 It is important to determine if a lesion is either congenital or acquired
Figure 59.6 Microphthalmos in this foal resulted in a flattening of the skull on the ipsilateral side. (See color plate 12).
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between 7% and 14.7% of all congenital ocular defects in the horse. Microphthalmia or microphthalmos is the result of failure of the globe to develop appropriately during gestation and may be either unilateral or bilateral. Most cases are idiopathic and occur sporadically. However, there have been a few case reports of maternal ingestion of potential teratogens, such as sulfadimethoxine, resulting in abnormal ocular development in foals. There is no known genetic defect or predisposition identified in the horse for microphthalmia, although Thoroughbreds seem to have a higher incidence of this condition than other breeds. The severity of this anomaly varies tremendously and may range from a slightly small, but comfortable and visual eye (pure microphthalmos) to the apparent lack of an eyeball at all (anophthalmos). Complete anophthalmia, or absence of the globe, is very rare. Even when no recognizable ocular tissue is visible, there is usually some histological evidence of rudimentary ocular tissue when the orbits are processed and examined. Most cases fall somewhere in between these two extremes. The severity of the microphthalmia depends upon the time of gestation at which the insult occurs with the more extreme ocular abnormalities resulting at the earlier stages of development. There is often a small palpebral fissure and an elevated or prominent third eyelid since the position of this structure relies upon a normally sized and positioned globe filling the orbit. Microphthalmia may be accompanied by other ocular anomalies, such as cataracts or retinal dysplasias and detachments, many of which are the cause of the visual deficits. Microphthalmic globes are predisposed to ulcerative keratitis due to the reduced volume of the orbital contents allowing the eyelids to secondarily roll inwards causing entropion. Ocular discharge is common due to ocular irritation from the abnormal ocular and orbital anatomy or external irritants or from a poorly functioning lacrimal drainage system. Surgical correction of any problematic entropion is often necessary and enucleation of excessively small, nonvisual eyes is in the best interest of the comfort of the foal. It must be noted, however, that continued growth and development of the structure of the orbit and the face rely on the presence of a globe within the orbit. The orbits and face of enucleated or microphthalmic globes may fail to grow appropriately, resulting in asymmetry of the periocular region. There is no treatment available to correct the size of the small globes themselves.
Figure 59.7 Lateral strabismus is present associated with a congenital cataract in a Paint filly. It resolved following cataract surgery. (See color plate 13).
1 month of age1, 3, 5 (Figures 59.7 and 59.8). The pupil of the normal adult horse is oriented at a very slight dorsomedial angle to the palpebral fissure. In foals, the axis of the pupil may be slightly ventromedial relative to the horizontal palpebral fissure. If this deviation remains in the foal past its first post-parturient month or if the globe is deviated in another position, this suggests there may be another structural or visual defect that is preventing proper globe orientation. Inappropriate development of one or more extraocular muscles will result in globe deviation in the direction of the abnormal muscle. Cataracts or other abnormalities that prevent light stimulus from interacting appropriately with the visual pathways may cause strabismus as well. Any significant or persistent strabismus warrants a complete ophthalmic examination for other underlying or confounding conditions. Strabismus may occur in any breed of horse, but has been reported with greater frequencies in Appaloosas and may be associated with congenital stationary night blindness (CSNB) in this breed. The
Strabismus Strabismus, or the deviation of the globe from its normal position, is a normal finding in foals less than
Figure 59.8 Ventral strabismus is present in this foal. (See color plate 14).
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encroachment upon the adjacent cornea. They have also been reported in both bulbar and palpebral conjunctiva, the eyelids, and the third eyelids. An orbital dermoid cyst causing exophthalmos was reported in one horse. Dermoids may limit vision if they are large and obscure the visual axis of the cornea, and may cause irritation, particularly if they are haired. They may result in eyelid or nictitans deformity if they are located upon or within theses structures. The only treatment possible for these lesions is surgical excision of the entire lesion, which is usually curative. Fortunately, most corneal and scleral dermoids in the equine eye are superficial, although the rare lesion will involve the full thickness of the cornea. If the entire lesion is not removed, there is potential for regrowth. Small lesions may not require any treatment if they are not causing tissue distortion or irritation. Figure 59.9 A pigmented dermoid and gray area of cornea are present near the dorsal limbus in this Standardbred foal. (See color plate 15).
strabismus that accompanies CSNB is usually convergent and dorsomedial.1, 3, 5 Strabismus also occurs in mules at a much higher incidence than in horses, and is usually convergent but asymmetrical.1, 3, 5
Dermoids Dermoids, also known as choristomas, are masses of histologically normal tissue occurring in an abnormal location, and are rare in the equine eye (Figures 59.9 and 59.10). They are usually some form of dark brown skin-like tissue with focal pink glandular areas that may or may not have hair follicle development.1, 3, 5, 7 They can occur in almost any position on the globe or about the orbit, but are most commonly found on the temporal or dorsal limbus with
Figure 59.10 The pale spots on the dermoid are glandular tissue. Hairs are not present. (See color plate 16).
Congenital glaucoma The glaucomas are a group of diseases resulting from alterations of aqueous humor dynamics that cause an intraocular pressure (IOP) increase above that which is compatible with normal function of the retinal ganglion cells and optic nerve1, 3, 5, 8 (Figure 59.11). In most cases, glaucoma in the horse is an acquired condition secondary to chronic intraocular inflammation. Aged horses, those afflicted with equine recurrent or persistent uveitis, and Appaloosas are predisposed.1, 3, 5 Congenital glaucoma is reported in foals and is usually associated with developmental anomalies of the iridocorneal angle or drainage angle through which aqueous humor exits the globe. The IOP is easily measured with a handheld tonometer and topical anesthesia. There normal IOP ranges from 17 to 30 mmHg in the horse. Any animal with
Figure 59.11 Corneal edema is present in this foal with congenital glaucoma. (See color plate 17).
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Table 59.1 Topically applied medications for equine glaucoma Drug
Type
Dose
Timolol maleate 0.5% Dorzolamide 2% Latanoprost 0.005% Atropine 1% Prednisolone acetate 1%
Beta blocker CAI Prostaglandin Anticholinergic Corticosteroid
0.2 mL b.i.d. topically∗ 0.2 mL t.i.d. topically∗ 0.2 mL s.i.d. topically∗∗ 0.2 mL bid topically∗∗∗ 0.2 mL t.i.d. topically
CAI, carbonic anhydrase inhibitors. ∗ CosoptTM (combination of timolol and dorzolamide), Merck & Co., Whitehouse Station, NJ. ∗∗ XalatanTM , Pharmacia and Upjohn, Kalamazoo, MI. Not recommended unless close tonometric monitoring is available. May cause phthisis bulbi. ∗∗∗ Selected cases with strict IOP tonometric monitoring.
pressures consistently above the upper values in this range and with the accompanying clinical signs of corneal edema, buphthalmia, and corneal striae can be diagnosed as having glaucoma. Topical and systemic medications to control inflammation and lower pressure may be attempted, however, most congenital cases are blind at birth and the IOP is not easily controlled. However, if the foal is sighted, medical management should by all means be attempted in order to maintain vision and keep the animal comfortable. Medical therapy is included in Table 59.1. If medical therapy is not adequate to control IOP, cyclophotocoagulation or cyclocryotherapy are surgical options to decrease the production of aqueous humor in sighted animals. Salvage procedures such as enucleation, evisceration with implantation of an intraocular prosthesis, or chemical ciliary ablation are means to lower IOP in irreversibly blind eyes. Often there are other abnormalities present in eyes with congenital glaucoma, such as aniridia, microphakia (Figure 59.12), and any other findings associated with dysgenesis of the anterior segment.1, 3, 5 Acquired ocular abnormalities Most acquired orbital disease in the neonate or older foal are attributable to trauma. There are sporadic reports of neoplastic diseases occurring in the eye or periocular regions in foals, but they are rare and most often derangements of primitive or nondifferentiated tissues. Deviation of globe position or strabismus may occur secondary to a traumatic insult, particularly one which results in fractures of the bones of the orbit that displace the globe or result in impingement of the extraocular muscles. Strabismus may also result from space occupying lesions of the orbit, including orbital inflammatory disease, or because of a muscle avulsion from a traumatic event, although this is unusual in the horse.1, 3, 5
Figure 59.12 Microphakia and a cataract are noted in this foal eye. (See color plate 18).
Exophthalmos refers to the anterior displacement of the globe within the orbit.1, 3, 5 In its purest description, exophthalmos affects a normal-sized globe, rather than an enlarged globe that appears to protrude from the orbit, as is the case when an eye is afflicted with chronic glaucoma and subsequently becomes buphthalmic. Trauma severe enough to cause immediate exophthalmos usually also results in significant damage to the orbital and periorbital structures. Due to the protective complete bony orbit of the horse, proptosis, or the protrusion of the globe from the orbit with entrapment of the eyelid margins posterior to the equator of the globe, is very rare and when it occurs usually leaves the globe severely injured, usually necessitating its removal. Often there are extensive fractures of the orbital bones as well, especially the dorsal orbital rim. If a fracture results in bony fragments that threaten the globe or entrap an extraocular muscle and limit globe mobility, it must be repaired promptly as fibrous union will begin within days of the injury. Inflammatory orbital disease may also result in exophthalmos, and is usually a diffuse process such as cellulitis rather than a discrete abscess.1, 3, 5 Blunt trauma, introduction of foreign material and infectious agents, and hemorrhage are common precursors to orbital cellulitis in the horse. Treatment should be initiated promptly and aggressively as progressive disease can damage the globe and lead to vision loss or even threaten the life of the animal if the process is septic. Systemic broad-spectrum antibiotics and nonsteroidal anti-inflammatory agents are generally indicated. Sampling for cytology and culture will permit directed therapy (see Table 59.2). Enophthalmos, or the retraction of the globe into the orbit, can be an active event wherein the animal engages the retractor bulbi muscle to pull the globe
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Table 59.2 Medical therapy for stromal abscesses Drug
Dose and frequency
Topical natamycin (5%), miconazole (%), or voricohazoic (1%) Topical antibiotic: bacitracin, neomycin, polymyxin B Topical atropine (1%) Systemic flunixin meglumine Systemic omeprazole Systemic antibiotics: Doxycycline (D); Trimethoprim Sulfa (TMS) Systemic antifungals: itraconazole (Itra) or fluconazole (Flu)
0.2 ml q.i.d 0.2 ml q.i.d 0.2 ml q.i.d, reduce after dilation 1 mg/kg i.v., i.m., p.o. b.i.d. 4 mg/kg p.o. s.i.d. for 28 days D: 5 mg/kg b.i.d. p.o. TMS: 15 mg/kg p.o. b.i.d. Itra: 3 mg/kg p.o. b.i.d. Fluc: 5 mg/kg p.o. b.i.d. × 7 days, then 5 mg/kg p.o. s.i.d.
inward in either as protective mechanism or a sign of pain or a passive process wherein the globe slips backward as a result of the loss of orbital contents. Dehydration is one of the most common causes of enophthalmos in the young, sick foal. Phthisis bulbi, or shrinkage of the globe, may simulate globe retraction. A careful ophthalmic and physical examination will distinguish between etiologies.
Diseases and disorders of the eyelid, conjunctiva, and nasolacrimal system Congenital abnormalities
Coloboma The term coloboma refers to an absence of tissue resulting from the failure of complete development.1, 3, 5, 9 Any ocular structure may be involved including the eyelid, iris (Figure 59.13), lens, choroid, retina, or optic nerve. Most cases occur due to failure
of closure of the optic fissure during embryological development. Fortunately, all forms of coloboma in the horse are rare. Eyelid colobomas, also referred to as eyelid agenesis, may be the most common form, and may or may not occur from embryological fissure anomalies. Ischemic damage to a rapidly developing embryonic eyelid may result in incomplete development and, thus, a defect. Short full-thickness colobomas in the foal are most common in the lower lid and have been observed in cases accompanying multiple ocular anomalies. Treatment depends upon the extent of the defect and the degree of secondary ocular exposure or keratitis present. Mild cases appear as a focal notch in the eyelid with no clinical signs; more extensive lesions may lead to corneal or conjunctival disease if the eyelashes or facial hair contact the globe or if the eyelids are unable to cover the globe or appropriately distribute the tear fluid. Surgical blepharoplasty is necessary in these cases to protect the globe and establish adequate eyelid function. Cryoepilation of misdirected eyelashes or trichiasis may relieve discomfort if the focal defect is small and the hairs are the main irritant.
Entropion
Figure 59.13 An iris coloboma in the pigmented portion of the iris exposes the peripheral lens. (See color plate 19).
Entropion is the inward displacement, or inversion, of the eyelid margin (Figure 59.14). It may occur as either a congenital defect or an early acquired condition in the equine neonate. More often the lower eyelid is affected, but severe cases may have involvement of both eyelids. The infolded eyelid margin permits contact of the external skin and hair with the cornea and conjunctiva and thus leads to mechanical irritation and ulcerative corneal lesions.1, 3, 5, 10, 11 Painful ocular conditions, such as the ulcerative keratitis and conjunctivitis that are associated with entropion, can lead to or exacerbate entropion already present. Spastic entropion is a secondary event. The problem may result from a weakness in the tarsal plate of the eyelid and is reportedly inherited
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(a)
(b)
(c)
Figure 59.14 Entropion of the lower lid is found in this foal. (b) Entropion repaired with sutures. (c) Entropion repaired with staples. (See color plate 20).
in the Thoroughbred, although the mechanism of inheritance is unknown. Treatment involves surgical repositioning of the eyelid margin, typically via placement of horizontal or vertical mattress sutures of 4-0 non-absorbable suture material, or surgical staples within the skin adjacent to the inversion (Figure 59.14c). These sutures are temporary. Many cases will reverse themselves in time and not require permanent surgical correction. Permanent correction of an anatomic entropion should be delayed until the animal has attained adult or near adult size.
length of the nasolacrimal duct. Most cases are not diagnosed in foals until the animals have reached several months of age, often because the clear epiphora that is the first clinical sign is unnoticed until the
Nasolacrimal atresia and aplasia Congenital malformations of the nasolacrimal apparatus have been reported in foals both unilaterally and bilaterally12–15 (Figures 59.15–59.17). The most common and easily addressed manifestation of this defect is atresia of the distal punctum in the nasal meatus, otherwise known as an imperforate punctum. More severe developmental anomalies including duct aplasia are possible anywhere along the
Figure 59.15 Severe, purulent dacryocystitis is present in this foal with atresia of the nasolacrimal duct. (See color plate 21).
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Figure 59.16 Dacryocystorhinography of the horse in Figure 59.15 reveals a distal blockage of the duct.
character of the discharge changes to mucopurulent and a secondary dacryocystitis and conjunctivitis occurs. Diagnosis is also delayed because neonatal foals have a reduced tear volume compared to older foals, and the portion of the nasolacrimal duct that has formed has the ability to distend and accommodate higher volumes. Tear fluid can be absorbed transmucosally across the duct as well. Once present, however, the amount of mucopurulent discharge is often copious and of strong odor. Diagnosis is accomplished by physically observing the lack of puncta, observing or palpating distension of nasal mucosa above the duct, and the inability of fluid flushed through the apparatus to exit the system. If the termination of the duct is not appreciable via direct observation of a dilated spot or palpation of same, contrast dacryocystorhinography is necessary to determine the extent of the malformation. Surgery is necessary to correct the defect and establish patency of the nasolacrimal excretory system.
Figure 59.17 The duct was reformed surgically and a stent placed in the duct for 1 month to resolve the condition of the horse in Figures 59.15 and 59.16. (See color plate 22).
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Fortunately, most cases are mild and involve only the nasal punctum or the distal third of the duct. If the nasal punctum is visible but covered with a membrane, excision of the membrane, followed by flushing of the duct and treatment with topical antibiotics and anti-inflammatory medications, is all that is necessary. If the distal portion of the duct has not formed and a discrete punctum is not visible, a #5 French catheter should be inserted into the proximal duct as far distal as is possible. Often the catheter will be palpable from the nasal opening. If so, excision of the overlying tissue above the catheter with a no.11 blade is necessary to establish drainage. A pliable catheter should then be placed along the length of the patent duct, sutured to the skin of the face distally at the nostril and proximally near the medial canthus, and left in place for 4–6 weeks to allow for epithelialization of the newly formed portion of the duct. Appropriate topical therapy for the secondary dacryocystitis should follow. If the condition is severe with minimal to no formation of the nasolacrimal apparatus or if correction of the atresia or agensis is not possible with the techniques described above, a more elaborate surgical approach is necessary. Conjunctivorhinostomy, canaliculorhinostomy, or conjunctivosinotomy will create an ostium into the rostral maxillary sinus or nasal cavity allowing drainage of the tear fluid from the lacrimal lake and resolution of the epiphora from tear overflow. Acquired eyelid abnormalities Due to the nature of the horse, trauma to the eye is exceedingly common. Younger animals tend to be quick and excitable and therefore quite prone to traumatic insults. Eyelids are easily lacerated, particularly upon exposed nails or bucket hooks, and should be immediately repaired.1, 3, 5 Upper eyelids especially are critical for the protection and lubrication of the globe. Profound corneal injury and disease can occur if the eyelids are not properly functional. Hanging pedicles of full-thickness eyelid should be sutured back in place with particular attention paid to the repair of the eyelid margin, and never amputated. Two-layer closures are stronger; however, care must be taken to ensure that no suture contacts the cornea from exposure through the palpebral conjunctival surface. Auriculopalpebral nerve blocks, sedation, topical anesthesia, and ocular lubricants aid repair of lid lacerations. If a repaired laceration dehisces or the wound contracts excessively during the healing process, more sophisticated blepharoplastic procedures may be necessary. Acquired entropion is usually secondary either to systemic disease or ocular pain in the foal. Dehydration and emaciation may result in loss of orbital
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Figure 59.19 Subconjunctival hemorrhage in a foal was caused by a difficult birth. (See color plate 24). Figure 59.18 Conjunctivitis is pronounced in a foal exposed to air pollution from a forest fire. (See color plate 23).
tissue and therefore enophthalmia. The eyelid margins roll inward passively and can irritate the globe. Treatment is aimed at the systemic disease and support of the ocular condition. Spastic entropion can occur when ocular pain is present. Finding the cause of the pain from an ulcer or uveitis and treatment with systemic non-steroidal drugs can elimination the spastic entropion. In these cases, there is no anatomic anomaly, but squinting from ocular pain can mimic it. Most cases of enopthalmic or spastic entropion require temporary tacking to keep the eyelid margins everted while the systemic disease or ocular condition is being corrected. Eyelid trauma or surgery can result in cicatricial entropion from scarring. Reconstructive blepharoplasty and scar revision is necessary to correct these cases.1, 3, 5 Conjunctivitis is common in foals and is most often a secondary disease process (Figure 59.18). Hyperemia, chemosis, and ocular discharge are the most common clinical signs of conjunctivitis. Chronic conjunctival inflammation may lead to the development of raised lymphoid follicles within the affected tissue. Systemic bacterial or viral infections are often accompanied by a serous or mucopurulent conjunctivitis. Etiological agents include adenovirus, equine herpesvirus (EHV)-1, equine viral arteritis, influenza virus, Streptococcus equi ssp equi (strangles), Rhodococcus, and Actinobacillus sp.1, 3, 5 Proliferation of opportunistic conjunctival bacterial flora may exacerbate conjunctival inflammation and cause a serous discharge to transform into a mucopurulent one. Bacterial conjunctivitis should be treated with a broadspectrum topical antibiotic ointment such as bacitracin neomycin and polymixin, or chloramphenicol.
Conjunctivitis may also result from airborne irritants, insects or trauma; however, primary conjunctivitis is rare and the presence of conjunctival inflammation should prompt a thorough physical and ophthalmic examination to exclude other primary disorders. If intraocular inflammation is present, anti-inflammatory therapy with dexamethasone or prednisolone acetate is indicated. If other signs of systemic disease are discovered, further diagnostics may be indicated as well and systemic therapy for the inciting disorder. Subconjunctival hemorrhages are relatively common in neonatal foals and are often due to birthing trauma1, 3, 5 (Figure 59.19). Most hemorrhages are located dorsally or dorsonasally and are not associated with any other ocular abnormality. Most subconjunctival hemorrhages resolve within 1–2 weeks with no untoward effects on the eye. If foci of hemorrhage are discovered elsewhere in the body and other corresponding clinical signs are observed, evaluation for the presence of a bleeding disorder may be warranted. Dacryocystitis is the most common acquired condition of the nasolacrimal apparatus.1, 3, 5 It is an almost invariable secondary consequence of nasolacrimal punctal, or ductal occlusion or atresia. Conjunctivitis often occurs secondary to nasolacrimal obstructions. While all young animals should be evaluated for congenital anomalies that may result in dacryocystitis, other causes of acquired disease should be considered, including trauma that might have resulted in a facial fracture or swelling that impinges upon the outflow of the nasolacrimal apparatus or the presence of a parasite within the medial canthus or NLD that is impairing tear drainage. Topical antibioticcorticosteroid combination preparations are usually necessary for treatment.
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Diseases and disorders of the cornea Congenital abnormalities Fortunately, congenital defects of the cornea are rare. Corneal opacities from dermoids or iris-to-cornea persistent pupillary membranes are the most common anomalies. The cornea is the most common location of dermoids (Figures 59.9 and 59.10). Local irritation is common and obstruction of the visual axis is possible with large lesions. Surgical resection via superficial keratectomy is the only viable treatment option and is curative if complete removal is achieved. Other congenital corneal opacities have not been well described or characterized. A group of Thoroughbred foals born with irregular superficial punctuate opacities was reported. The etiology of the opacities was unknown, but suspected to be an infectious or environmental insult suffered by the mare during late pregnancy. The opacities resolved in time without treatment.1, 3, 5 A few reports have cited the appearance of focal corneal melanosis or perilimbal superficial corneal vascularization in newborn foals. The vessels resolved without treatment within a week postpartum. Cyclosporine-responsive corneal pigmentation has been noted in foals. Anomalies of corneal size are possible in newborn foals. Microcornea has occasionally been reported and usually occurs in conjunction with microphthalmia or multiple ocular anomalies. Megalocornea, wherein the corneal diameter is larger than normal while the biometry of other ocular structure remains normal, has been seen in Rocky Mountain Horses and Miniature Horses as a manifestation of anterior segment dysgenesis.1, 16
Figure 59.20 Severe, central melting ulcer with corneal edema, hypopyon, and corneal vascularization are present in this foal. (See color plate 25).
tissue turnover in rapidly growing foals, it is relatively easy for the normal housekeeping balance dictating the destruction of old corneal collagen and the production of new corneal collagen to be tipped in favor of the destructive enzymes which are naturally present in the corneal tissue with the end result a melting corneal ulcer. More often than not, melting ulcers in foals are sterile; however, they are susceptible to secondary bacterial and fungal infection. All melting ulcers should be cultured for the presence of microbes and their specific drug sensitivities. Aggressive topical medical therapy with antimicrobials, antiproteases, and antiuveitic drugs is imperative for ulcer resolution (see Table 59.4).
Acquired corneal abnormalities The most ocular common disorder affecting both foals and adult horses is ulcerative keratitis1, 3, 5 (Figures 59.20 and 59.21). Corneal ulcers are usually secondary to some sort of trauma, even if microscopic, and can be exacerbated by low tear production in foals, decreased corneal sensation in sick foals, and intraocular inflammation affecting the overall health and nutrition of the cornea. Corneal ulcers require prompt evaluation and treatment in horses since they can progress rapidly to perforating insults in many instances. Cytology of the ulcer and culturing is recommended. Treatment may be medical or surgical, depending upon the depth of the lesion, its etiology, and response to therapy. In all cases, topical antimicrobial and antiprotease medications (see Table 59.3) should be employed and any secondary uveitis should be addressed with topical cycloplegics and systemic anti-inflammatory medications. Due to fast
Figure 59.21 Fluorescein staining reveals an ulcer of the cornea in this foal. (See color plate 26).
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Foal: Ocular System Table 59.3 Antiproteinase drugs for topical use Proteinase-types inhibited and mechanism of action
Antiproteinase drugs
Dose
Serum: (α-2-macroglobulin, α-1-proteinase inhibitor, and platelet-derived growth factors)
Undiluted; q1-2h
Plasma: (α-2-macroglobulin, and α-1-proteinase inhibitor)
Undiluted; q1-2h
MMP and serine proteinase inhibitor; chelates Ca and Zn
EDTA 0.2%
q1-2h
MMP inhibitor
Acetylcysteine 10%
q2-4h
MMP inhibitor
Doxycycline 0.1%
q8h
MMP inhibitor
α-1-proteinase inhibitor 0.1%
q1-2h
Serine proteinase inhibitor; protease entrapment
Diseases of the uvea Congenital abnormalities Defects of the anterior uvea can range from very mild focal tissue thinning or hypoplasia, to near complete absence of the iris or aniridia1, 3, 5,17–21 (Figures 59.22 and 59.23). Iris hypoplasia is not an uncommon finding, particularly in blue-eyed animals, and the most mild form occurs as a bulging of iris toward the cornea at the 6 or 12 o’clock positions. These lesions must not be confused with neoplastic mass lesions. In hypoplastic irides, the affected area will change shape with changes in pupillary dilation, may appear thin, and often the area will transilluminate with light reflected from the ocular fundus. As the spectrum advances, full-thickness iris colobomas may be present and appear as holes within the iris. The most common location for these lesions is at the 6 o’clock position in the ventral iris that corresponds to the closure site of the optic fissure. They may occur elsewhere, Table 59.4 Melting ulcer therapy Drugs
Initial dose frequency
Topical antibiotics: gentamicin; tobramycin; bactitracin-neomycinpolymyxin; ciprofloxacin; amikacin; doxycyclin, or cefazolin
q1-2h
Topical antifungal if cytology is positive for hyphae: Natamycin (5%)
q4h
Topical autogenous serum
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Topical EDTA (0.2%)
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Topical Ilomostat
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Topical atropine (1%): reduce dose when pupil is dilated
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Systemic flunixin meglumine
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Systemic doxycycline
5 mg/kg p.o. b.i.d.
Figure 59.22 Iris hypoplasia with absence of the corpora nigra are present in this White foal. (See color plate 27).
however, and should not be confused with iris atrophy which occurs in aged horses. True aniridia is rare in horses, but has been reported as an autosomal dominant inherited condition in Quarter Horses and Belgians. Affected
Figure 59.23 Heterochromia iridis is present in this eye. (See color plate 28).
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Figure 59.24 A persistent pupillary membrane is present but causing no harm in this foal. (See color plate 29).
animals present for evaluation of an abnormally large pupil, blepharospasm, and photophobia bilaterally. The equator of the lens is visible as are the ciliary processes to which the lens zonules attach. Several theories of abnormal development have been proposed to explain this phenomenon, but failure of mesodermal ingrowth to form the iris is most widely accepted. Other ocular anomlies may be present as well, including limbal dermoids and cataracts. No specific treatment is available for this condition and affected animals should be supported for their clinical signs, protected from excessively bright conditions with either differential turnout or masks that provide shade, and eliminated from breeding programs. Another form of uveal maldevelopment is the lack of regression of the papillary membrane that forms during embryogenesis. Persistent pupillary membranes
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(PPMs) (Figure 59.24) may occur as small strands connecting the iris collarette to itself or as bands connecting the iris to the lens posteriorly or the cornea anteriorly.1, 3, 5 Posterior adhesions result in focal cataracts and anterior adhesions cause focal corneal opacities. When large regions of the cornea are affected, focal endothelial dysfunction may result in corneal edema. A syndrome of ocular maldevelopment known as anterior segment dysgenesis has been reported in several horse breeds, but the most spectacular examples have been found in the Rocky Mountain Horse.1, 16 There is a range of affliction that varies from cysts of the ciliary body alone, to megalocornea with congenital miosis, abnormal iridocorneal angle structure, cataracts, retinal dysplasia, and retinal detachments. Severe cases with aqueous outflow abnormalities may develop glaucoma. There is a heritable basis for this anomaly and is believed to be related to pigmentation and coat color, with chocolate-colored animals with flaxen manes and tails more likely to be affected. Acquired uveal abnormalities Uveitis is a non-specific inflammatory reaction in the iris, ciliary body, or choroid to any number of insults, either local or systemic1, 3, 5 (Figure 59.25a,b). In most cases, uveitis in the foal is either the result of a traumatic insult, or a systemic bacterial or viral infection. Sepsis or focal infections elsewhere in the body will often produce uveitis as a manifestation of disease, either by direct translocation of the infectious agent into the eye or via a systemic inflammatory response. Clinical signs may include miosis, corneal edema and vascularization, aqueous flare, fibrin within the anterior chamber, hypopyon or hyphema, ocular hypotony, change in iris color, iridal neovascularization,
(b)
Figure 59.25 (a) Fibrin obstructs the pupil, and the iris vessels are very congested in this eye with uveitis. (b) Uveitis from Rhodococcus is manifested by severe fibrin deposition of the anterior chamber in both eyes. (See color plate 30).
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scleral injection, and evidence of ocular pain. Rhodoccocus equi, Actinobacillus equili, Escherichia coli, Streptococcus equi and the Salmonella spp are notorious for causing uveitis in addition to other signs of systemic disease. The uveitis in these cases, especially in foals with Rhodococcus pneumonia, is often fibrinous with a large amount of fibrin accumulating within the anterior chamber, often obscuring observation of the interior structures of the globe. Treatment is directed at the primary disorder and symptomatic therapy for the intraocular inflammation. Topical corticosteroids such as dexamethasone or prednisolone acetate and mydriatic/cycloplegic agents such as 1% atropine as well as systemic anti-inflammatory medications are usually indicated. The goal of therapy for the eye is to relieve discomfort and to minimize damage to the interior structures of the eye, such as the development of synechiae and cataracts, in order to maintain adequate visual function. Antifibrinolytic agents such as tissue plasminogen activator are often needed to remove excessive amounts of anterior chamber fibrin.
Diseases of the lens Congenital abnormalities
Cataracts As in other species, equine cataracts may be variously classified according to their anatomic location within the lens, their physical appearance, their presumptive etiogenesis, or stage of progression or maturity.1 No single classification wholly or satisfactorily encompasses the very wide range of cataracts occurring in the horse. A pragmatic compromise in this species is a classification based upon a combination of anatomic location, appearance, and etiogenesis.1, 3, 5 The majority of equine cataracts are nonprogressive, and classification of equine cataract according to stage of progression, i.e., incipient (implying progression), immature, mature, and hypermature, is appropriate only in a very few cases, e.g., some post-uveitic cataracts. Morgagnian cataract, a hypermature cataract where the nucleus ‘sinks’ within the liquefied cortex, has been described in the horse, but is probably extremely rare.1 Early equine clinical texts1 describe the basic anatomical division of equine cataracts into capsulolenticular cataracts (involving the capsule and subjacent cortex) and lenticular cataracts (involving the lens cortex and nucleus). This may be further refined by more specific localization of the cataract (e.g., anterior capsular, sutural, perinuclear, equatorial).1 A fundamental and practical division of equine cataracts by etiogenesis can be made into acquired
Figure 59.26 An immature cataract is present in the right eye of a Thoroughbred foal. (See color plate 31).
or secondary cataracts, and developmental cataracts. Acquired or secondary cataracts occur as a result of influences arising from outside the lens, and in the horse these most commonly arise as a consequence of intraocular disease. Developmental cataracts result from disrupted evolution and accretion of the crystalline lens both in utero and postnatally, and have been elegantly described as aberrations rather than arrests.1 They include all congenital cataracts, with the exception of very rare congenital uveitic cataracts in foals. Their origins lie in interrupted or abnormal differentiation and growth of embryonic ectodermal derivatives, or in disruption of the normal maturation and laying down of secondary fibers in the fetal and postnatal lens. Most developmental cataracts can be morphologically related to a specific stage or event in lens embryogenesis and growth. More than one form of developmental cataract may be present in any one eye.1 Cataracts are by far the most common congenital abnormality of the horse eye, accounting for 35% of all ocular anomalies in foals1, 3, 5 (Figures 59.7, 59.26, 59.27). Excluding traumatic injuries, they are also the most common cause of blindness or vision impairment in foals. They may occur as the sole ocular abnormality, which is the case in most situations, or may accompany other findings such as microphthamia, aniridia, PPMs, persistent hyaloid structures, or syndromes of multiple ocular anomalies. They may occur in one or both eyes, but when both eyes are affected, the degree of cataract development is often asymmetrical, and they may be either focal or diffuse. Most cases are idiopathic and probably the result of an intrauterine insult, and either inflammatory, nutritional, toxic, or traumatic in nature. However, heritable cataracts have been described in several breeds including Thoroughbreds, Belgians, Quarter Horses and Morgans.1, 3, 5, 10, 22 . Examination
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Figure 59.27 Light aimed from various angles can aid diagnosis of a cataract. This is the same eye as in Figure 59.26. (See color plate 32).
of both sire and dam for the presence of cataract and a thorough medical history, including the ophthalmic status of previous offspring or other close relatives and the health, and the diet and environmental condition of the dam during gestation may help with the determination of etiology. Several types of congenital cataract have been reported in foals, including focal nuclear cataracts that are unlikely to progress, cataracts strictly associated with the Y-sutures of the lens, posterior cataracts associated with persistent hyaloid structures, and complete mature cataracts. The location of the opacity within the lens and the amount of the lens involved affects the cataract’s implications for vision. Small focal cataracts may not affect vision to any appreciable degree while complete, or nearly complete, cataracts will have a dramatic deleterious effect on vision. Axial cataracts will affect performance to a much greater degree than peripheral lesions.1, 3, 5 The only treatment available to eliminate cataract is surgical removal. Eyes with cataracts in association with other ocular anomalies or multiple ocular anomalies may not be reasonable candidates for surgery.1, 3, 5 Even if cataract is the sole ocular abnormality, preoperative testing to ensure suitability for surgery is recommended. Ocular ultrasonography will determine if a retinal detachment is present and electroretinography will determine if the retina is functional and likely to allow vision once the cataract is removed. The preferred method is via phacoemulsfication in order to minimize surgical time and intraocular surgical trauma, and to remove the lenticular material through a small incision. Cataract extraction is relatively successful in young horses as long as their temperaments permit frequent administration of postoperative medications and the stall confinement necessary during convalescence.
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Medications to control postoperative uveitis are critical to the health of the eye and the outcome of the procedure. Also, successful outcomes are influenced by the experience of the handlers and the diligence with which they handle their charges after surgery. The highest success rates for cataract surgery in foals is in the youngest animals since they tend to be easier to medicate, and their cataracts are generally less mature, softer, and easier to extract. Recently prosthetic intraocular lenses have become available for implantation at the time of cataract removal. The goal of these lenses is to improve visual acuity in the aphakic animal in order to more closely approximate vision of the normal phakic eye.1, 3, 5 Deprivation amblyopia is a possibility in foals with congenital cataracts and results from insufficient visual stimulation of the central nervous system. These foals have permanent functional and structural abnormalities of the lateral geniculate nucleus and visual cortex and cataract extracton will not restore vision. Animals that display a searching or wandering nystagmus may be suspected of having this condition. Abnormalities in lens shape, position, and size may occur as a congenital lesions in foals, although their incidence is rare. Lenticonus or lentiglobus, wherein the lens assumes a conical shape most often at posterior pole, has been sporadically reported as a unilateral affliction in horses. Microphakia (Figure 59.12. See also colour plate) or a congenitally small lens is usually noted associated with other ocular abnormalities such as microphthalmia or lens luxations. Congenitally luxated lenses are often cataractous and are usually the result of defects in zonular number or forms.1 Acquired lens abnormalities Cataracts can be an acquired affliction as well as a congenital one, particularly if the foal has suffered from a particularly severe or long-standing uveitic process. Treatment will include therapy for uveitis. Surgical removal of uveitic cataracts may or may not be possible depending upon the extent of the intraocular damage from the inflammation, the presence of synechiae, the status of the retina, and the potential for vision.1, 3, 5 Lens luxations or displacements are rare occurrences in the foal and when present are either a consequence of inappropriate development of the lens itself and its associated lens zonules, or because of trauma to the globe. Luxations secondary to chronic intraocular inflammation occur in aged animals that have suffered for months to years from their affliction rather than in foals. Treatment will be aimed at controlling intraocular inflammation and
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maintaining normotension of the globe. Occasionally surgical removal of the lens is possible, but carries with it many potential surgical complications. Anterior luxations are far easier to address surgically than posterior luxations.1, 3, 5
Diseases of the posterior segment Congenital abnormalities The embryonic hyaloid vascular system normally regresses shortly after birth, but in rare cases will persist into adulthood. The implications on vision and the health of the eye depend upon the extent of the residual structure. The most serious complication is the development of cataract at the posterior pole of the lens.1, 3, 5, 19, 23 Congenital lesions of the equine fundus are uncommon. Retinal dysplasia (Figure 59.28) or abnormal retinal differentiation is the result of either retinal non-attachment, defective retinal pigment epithelial tissue, or necrosis of developing retinal. In many species it has been associated with either in utero exposure to viral pathogens or with hereditary factors, but the etiology in horses remains undetermined. It can occur alone but has most often been associated with multiple ocular defects. Visual function of an affected animal will depend upon the severity and extent of the maldevelopment and the presence of other ocular abnormalities. Congenital retinal detachments (Figures 59.29 and 59.30) are usually complete and bilateral and have been reported most frequently in the Thoroughbred and Standardbred.1, 3, 5, 24 Ocular ultrasound can aid
the diagnosis of a retinal detachment. They may occur alone or in conjunction with multiple ocular anomalies. This abnormality is usually due to retinal non-attachment, rather than normal attachment followed by detachment. Successful treatment of a congenital detachment or nonattachment has not been achieved. Colobomas of the retina and optic nerve have been observed in horses.1, 3, 5, 9, 25 Retinal colobomas usually occur as sharply demarcated zones of depigmentation in the peripapillary fundus. Unless they are large or encompass a significant portion of the area
Figure 59.28 Retinal dysplasia is found in the tapetum of this foal. (See color plate 33).
Figure 59.30 A retinal detachment is present in this ultrasonographic image.
Figure 59.29 A retinal detachment appears as a gray veil obstructing a view of the optic nerve head. (See color plate 34).
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Ophthalmic Disorders of the Foal centralis; they are believed to have little impact on functional vision. This contrasts with colobomas of the optic nerve, which are often associated with significant, if not complete, visual deficits. Optic nerve colobomas are most often typical, or located at the 6 o’clock position on the disc and represent failure of closure of the optic fissure. Bilateral optic disc colobomas have been reported in Quarter Horses. Optic nerve hypoplasia, wherein the optic nerve is smaller than normal due to failure of retinal ganglion cells to develop, may be either unilateral or bilateral. It may occur alone, or with other ocular lesions. Visual impairment may be seen in severe cases, but does not occur in all cases. On ophthalmoscopic examination, the nerve head will often appear either small, pale, or with reduced retinal vasculature.1, 3, 5 Congenital stationary night blindness is a disease state in which visual disturbances occur in dimlighting conditions and at night. Affected animals often have a history of self-inflicted injury or apprehension at night. Most perform normally in daylight conditions, but a few suffer visual impairment during the day as well. Most affected animals are Appaloosas, in which a recessive or sex-linked mode of inheritance has been proposed. Up to 25% of Appaloosas may be affected. However, the disease has been reported sporadically in other breeds. The fundic examination of these animals is normal and diagnosis is dependent upon history and abnormal electroretinography. The defect is thought to be one of neural transmission and appears not to be progressive. Many affected individuals will demonstrate bilateral dorsomedial strabismus and stargazing behaviors and may actively seek illuminated conditions. No treatment is available for the condition and it is advisable to remove affected animals from breeding programs due to the hereditary nature of the disorder.1, 3, 5 Acquired posterior segment abnormalities Retinal hemorrhages are noted with some frequency in neonatal foals26 (Figure 59.31. See also colour plate). Most resolve without treatment and appear to have little or no long-term untoward effects on the animals’ vision or ocular health. There is a higher incidence of retinal hemorrhages in foals that underwent prolonged parturition. Some suggest that retinal hemorrhages are also a common finding in foals with neonatal maladjustment syndrome; however, recent surveys of equine neonates do not support this correlation. Acquired abnormalities of the posterior segment in foals are usually either traumatic or inflammatory in nature. Chorioretinitis and optic neuritis may occur with or without corresponding anterior segment
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Figure 59.31 Retinal hemorrhages are present in the eye of this foal immediately after birth. (See color plate 35).
inflammation. Both are most often the consequence of a systemic infection or inflammatory process and their presence should prompt a thorough physical examination. Systemic anti-inflammatory medications such as flunixin meglumine are warranted if the disease process is active, since topical medications will not affect the posterior segment of the eye. Acquired retinal detachments may occur secondary to intraocular inflammation, external trauma, or the hypotony that occurs with intraocular surgery. Prognosis is dependent upon the extent of the detachment, presence of concurrent retinal tears, damage to other ocular structures, and response to therapy. Ultrasounds of injured globes and orbits can assist clinicians towards making a prognosis for vision1, 3, 5 (see Figure 59.30).
References 1. Brooks DE, Matthews AG. Equine ophthalmology. In: Gelatt KN (ed.) Veterinary Ophthalmology, 4th edn. Ames, IA: Blackwell, 2007; pp. 1165–274. 2. Brooks DE, Clark CK, Lester GD. Cochet-Bonnet aesthesiometer-determined corneal sensitivity in neonatal foals and adult horses. Vet Ophthalmol 2000;3(2–3):133–7. 3. Brooks DE. Ophthalmology for the Equine Practitioner, 2nd edn. Jackson WY: Teton New Media, 2008. 4. Cook CS. Ocular embryology and congenital malformations. In: Gelatt KN (ed.) Veterinary Ophthalmology, 4th edn. Ames, IA: Blackwell, 2007; pp. 3–36 5. Gilger BC. Equine Ophthalmology. Philadelphia: Elsevier, 2005. 6. Enzerik E. The menace and papillary light reflex in neonatal foals. Equine Vet J 1998;30(6):546–8. ˜ ˜ T. Retrobulbar der7. Munoz E, Leiva M, Naranjo C, Pena moid cyst in a horse: a case report. Vet Ophthalmol 2007; 10(6):394–7. 8. Halenda RM, Grahn BH, Sorden SD, Collier LL. Congenital equine glaucoma: clinical and light microscopic findings in two cases. Vet Comp Ophthalmol 1997;7(2):105–9.
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9. Bildfell R, Watrous B, Schlipf J, Reynolds M. Bilateral optic disc colobomas in a Quarter Horse filly. Equine Vet J 2003;35(3):325–7. 10. Crowe MW, Swerczek TW. Equine congenital defects. Am J Vet Res 1985;46:353–8. 11. Gelatt KN. Congenital and acquired ophthalmic disease in the foal. Anim Eye Res 1993;1–2:15–27. 12. Latimer CA, Wyman M. Neonatal ophthalmology. Vet Clin North Am Eq Pract 1985;1(1):235–57. 13. Latimer CA, Wyman M, Hamilton J. An ophthalmic survey of the neonatal horse. Equine Vet J (Suppl) 1983;2:9–14. 14. Latimer CA, Wyman M, Diesem CD, Burt JK. Radiographic and gross anatomy of the nasolacrimal duct of the horse. Am J Vet Res 1984;45:451–8. 15. Latimer CA, Wyman M. Atresia of the nasolacrimal duct in three horses. J Am Vet Med Assoc 1984;184(8):989–92. 16. Ramsey DT, Ewart SL, Render JA, Cook CS, Latimer CA. Congenital ocular abnormalities of Rocky Mountain Horses. Vet Ophthalmol 1999;2:47–59. 17. Joyce JR, Martin JE, Storts RW, Skow L. Iridal hypoplasis (aniridia) accompanied by limbic dermoids and cataracts in a group of related Quarterhorses. Equine Vet J Suppl 1990;10:26–8.
18. Mosier DA, Engelman RW, Confer AW, McCarroll GD. Bilateral multiple congenital ocular defects in Quarterhorse foals. Equine Vet J Suppl 1983;2:18–20. 19. Munroe GA, Barnett KC. Congenital ocular disease in the foal. Vet Clin North Am Large Anim Pract 1984;6:519– 37. 20. Roberts SM. Congenital ocular anomalies. Vet Clin North Am Eq Pract 1992;8:459–78. 21. Turner AG. Ocular conditions of neonatal foals. Vet Clin North Am Eq Pract 2004;20(2):429–40. 22. Beech J, Irby N. Inherited nuclear cataracts in the Morgan horse. J Heredit 1985;76:371–2. 23. Matthews AG, Crispin SM, Parker J. The equine fundus. II: normal anatomical variants and colobomata. Equine Vet J Suppl 1990;10:50–4. 24. Cutler TJ, Brooks DE, Andrew SE, Denis HM, Biros DJ, Gelatt KN. Disease of the equine posterior segment. Vet Ophthalmol 2000;3:73–82. 25. Wheeler CA, Collier LL. Bilateral colobomas involving the optic discs in a Quarterhorse. Equine Vet J Suppl 1990;10: 39–41. 26. Munroe G. Survey of retinal hemorrhages in neonatal Thoroughbred foals. Vet Rec 2000;146:95–101.
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60.
Diseases of the Skin Derek C. Knottenbelt
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Chapter 60
Diseases of the Skin Derek C. Knottenbelt
Introduction The skin of a normal full-term newborn foal is soft and pliable and the hair coat is short, uniform, and silky. The mane and tail are usually short and slightly curly. Foals that are born premature (and sometimes the dysmature ones also) have sparse coats and may even be bald. Skin disorders are a relatively uncommon complaint in newborn and young foals, but congenital disorders (i.e., those present at birth) are encountered along with problems that are acquired at or after birth as a result of direct injury or infection. Congenital skin diseases are, by definition, those that are present (though not necessarily visible) at birth; some are only manifest later in life. For example, atopy and dermoid cysts usually manifest after some years and may be undetectable in the young or neonatal foal. Hereditary equine regional dermal asthenia (HERDA syndrome) may not be detected until the animal is subjected to skin pressure from a saddle and girth. Other conditions such as mane and tail dystrophy in horses of the Appaloosa breed may manifest much later than that. It is also possible that the breeders of such horses will be more than aware of the risks and may even be blas´e. The fact that these conditions occur congenitally does not mean that they are hereditary. Although some conditions are hereditary, little is known about most of them. Specific breed-related disorders and abnormalities are usually well recognized. Conditions such as insect bite hypersensitivity (sweet itch) and atopy have genetic origins and so there is a breeding implication. There are also a few non-genetic congenital conditions (e.g., congenital viral papilloma) and sporadic disorders such as nevus and cavernous hemangioma. Breeding implications should always be considered for the recognized hereditary skin disorders. However, some breeders simply accept a low rate of affected foals. For example, the lethal white syn-
drome in Overo Paint horses is an accepted, if rare, congenital condition with a dermal component. Most of the genetic congenital disorders are, by their nature, untreatable. Some do not materially affect the welfare of the horse and so can be either tolerated or managed, but others are incompatible with normal life either because they do not resolve or because they are associated with other concurrent potentially or actually fatal systemic abnormalities. The skin abnormality may be of no or little material significance, but it may act as a marker for the diagnosis of other more serious abnormalities, e.g., lethal white syndrome and Fell Pony immunodeficiency syndrome. There are dangers in making this association, however, and an albino (white) foal presented for neonatal colic may simply have an impacted small colon and may not be a lethal white syndrome foal. Furthermore, it may be difficult to convince an owner that a foal warrants euthanasia when the extent of cutaneous defects in a case of epidermolysis bullosa or cutaneous agenesis appears to be trivial. The acquired conditions are those that occur as result of injury or infection or are secondary to underlying disease and can be grouped broadly into infectious and non-infectious categories. Although the skin of neonatal foals is strong and resistant to damage, trauma and inflammation induced by body fluids such as diarrhea, urine, saliva, and wound exudate do occur with some frequency. Examples of these include traumatic injuries as a result of savaging by the dam, predation, or trauma, and decubitus ulcers over the bony prominences in foals that are recumbent, struggling to stand, or are seizuring. Systemic infections may also manifest with skin diseases. Some of the conditions with skin defects such as epidermolysis bullosa and cutaneous agenesis provide a portal of entry for serious secondary systemic infections. The attending clinician should establish quickly whether the skin lesions are in fact secondary to some deeper problem. For example, owners may
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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blame a skin deficit on savaging or predation, but if that is not plausible it may be a primary congenital condition. Clinical objectives for a clinician include the establishment of a definitive diagnosis so that a realistic prognosis and rational treatment can be suggested. Diagnosis can be assisted in many cases by collection of samples such as biopsy, microbiological swabs, and aspirates as well as hair plucks and brushings from representative skin lesions. These will either help to establish a positive diagnosis or rule out some of the differentials. The clinician should, however, consider whether the diagnosis is in fact an imperative for treatment; if not, then the precise diagnosis may become academic. Some conditions will respond to empirical or symptomatic treatment and so the diagnosis becomes less important. However, for client and professional satisfaction it is important to pursue a definitive conclusion if possible.l
Primary skin disorders occurring in newborn foals These conditions are primary cutaneous problems, although they may have systemic implications such as providing a portal of entry for bacteria. Some are hereditary and others are accidental genetic mutational or acquired disorders. Although some are of little significance, others are life-threatening. Epitheliogenesis imperfecta (aplasia cutis) This is a rare congenital and inherited cutaneous defect of foals reported to be associated with a single autosomal recessive gene.1 The American Saddlebred is over-represented but other breeds are occasionally involved. Many of the previously published reports of this condition can probably be attributed to epidermolysis bullosa rather than cutaneous agenesis.2 There is a complete or partial absence of one or more of the dermal components, including the epidermis, dermis, subcutis, and fat.3 Absence of varying areas of the skin is the most common syndrome, but sometimes it is complicated by concurrent defects in the subcutis resulting in loss of normal adhesion between the skin and the subcutaneous tissues. Defects on the limbs are probably more common, but the body trunk and hooves, and even the tongue and proximal esophagus, may also be affected. Areas of presumed ‘trauma’ are detected at or shortly after birth, but close examination will confirm the lack of inflammatory components and the paucity of healing. The defects may be significant portals for entry of bacteria, resulting in septicemia. Repeated reconstructive surgical procedures can be attempted but are seldom effective. Recurrent
wound breakdown and local and systemic infections are common. The prognosis is very poor but foals with limited defects can survive provided that the affected region can be managed effectively. Epidermolysis bullosa This is a rare, complex heritable junctional mechanobullous disease occurring principally in the Belgian and Saddlebred breed in both sexes. Other breeds and donkeys are sporadically affected. It is characterized by fragile ‘blister’ formation on the skin and oral mucosa, following mild trauma. Although the human condition carries the same name and there are some pathological similarities, there is limited information on how this relates to the equine condition.4 The junctional form of the condition is the most common and probably results from a lamina lucida defect.5, 6 Variations in location of the clefting within the epidermis are theoretically interesting but have no material significance for the management of the condition. It has been suggested that some of the previous reports of cutaneous agenesis in foals may in fact have been epidermolysis bullosa. Minor trauma results in bulla (blister) formation and, when these rupture, they leave an open ulcerated area that bleeds and is easily traumatized and infected. The limbs are more often affected than the body trunk and head. The mouth and coronary band may be affected severely. Treatment is not possible and euthanasia is the only effective treatment for most cases. Attempts to reconstruct the deficits are usually fruitless. Breeding from both parents should probably be stopped. The disorder can initially resemble the recently described, self-resolving bullous/ulcerative dermatosis associated with ingestion of colostrally derived alloantibodies.7, 8 Differentiation is important because of the profound difference in prognosis; the latter is self-resolving whereas the former leads inevitably to euthanasia. Generalized primary seborrhea This is a rare congenital condition most noticeable in younger horses (< 2 years of age), characterized by epidermal cell hyperproliferation with inappropriate sebum production.9 Generalized greasy scale and flake occur over the whole body from a young age. Secondary bacterial and Malassezia sp. infection is common. Diagnosis is made only when the condition has been present from birth or shortly afterwards, and affects the whole body, and when secondary causes of seborrhea have been eliminated. Biopsy is largely
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Figure 60.1 (a) A typical ‘curly coated’ yearling. The coat is often less curly in summer. Note the prominent curling of the mane and tail hairs. This is regarded as normal in some breeds. Natural (genetic) patterning (b) and curling of the hair coat at different anatomic sites (c) is an incidental finding of no medical significance. Some of these patterning syndromes are hereditary.
unrewarding with non-specific epidermal hyperplasia and hypertrophy and excessive scale. Treatment is palliative and symptomatic with degreasing and descaling shampoos. The condition is a persistent but clinically insignificant problem. Congenital curly coat syndrome A congenital ‘curly coat’ syndrome is recognized in Paint, Quarterhorse, Percheron, Arabian, Appaloosa, Missouri Fox Trotter, Tennessee Walking Horses, Morgan, and Paso Fino breeds.10, 11 Defined genetic lines are possibly more liable to the trait. This is sometimes viewed as a desirable feature and so active steps are taken to maintain the condition in some breeds and lines within breeds. Curly coat patterns are also found in other horses from time to time although most such cases are only manifest later in life. A prominent curly, dense coat is present from birth and develops into a dense wavy coat quality (Figure 60.1). The condition may be restricted to the mane and tail (wavy mane syndrome) but can affect the whole body. Some hair loss, including the tail and mane, is also often present. The condition has no apparent deleterious effect. Treatment is not usually expected and in any case there is none. The condition should not be confused with either the hirsutism characteristic of very old horses arising as a result of pituitary pars intermedia dysfunction or the natural patterning of the hair coat that is sometimes encountered (Figure 60.1b).
(pink or light blue) irides (and usually have no pigment in the retina) (Figure 60.2). Albinism is an autosomal dominant trait and a foal is only viable in the heterozygous state (Ww). The homozygous state (WW) is lethal and affected embryos/fetuses are resorbed or aborted early in gestation. Albinism does not, in itself, have any disease implication for the horse although the lack of pigment may make the skin more prone to actinic dermatitis and squamous cell carcinoma. Lethal white foal syndrome is an inherited autosomal recessive trait occurring with the mating of Overo and Paint horses of the Pinto breed (see Chapter 73). Cases have been encountered in other breeds also.12 Affected foals are white (albino) but sometimes have a few brown flecks on the ears or face (Figure 60.3). Affected foals develop severe non-responsive colic within hours of birth associated with an atretic (incomplete) gastrointestinal tract in which there is a complete closure or absence of a section usually of the large intestine or ileus resulting from ileocolonic aganglionosis,6, 8 or both.
Albinism/lethal white foal syndrome Albinism is a condition caused by a biochemical defect preventing normal melanin synthesis in melanocytes. True albinos have a congenital lack of pigment (melanin) in the skin, hair, and iris and so have white hair, pink skins, and non-pigmented
Figure 60.2 This mare had a similarly colored albino foal. Both had completely non-pigmented retinas and blue/white irises. Neither animal had any problems over many years apart from a tendency to sunburn on the muzzle in summer months.
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Figure 60.3 This Overo Paint foal showed typical non-responsive colic after birth (a) and had an incomplete intestinal tract in addition to no fecal material detectable at the anus (b).
Treatment for either condition is not possible. Albinotic horses probably need to be given extra sun protection (mask for face and eyes and light rugs) but most seem to cope well anyway. Lethal white foals must be destroyed immediately; any attempts to remove lengths of affected intestine or restore patency of the gut will fail.
Parents of the affected foal should probably not be used for breeding, but little is known about the genetics of the syndrome as yet.
Lavender foal syndrome Lavender foal syndrome is a rare poorly characterized, (supposedly) hereditary disorder of Arabian foals derived from a particular Egyptian line.13 The name derives from the unusual hair coat color, which can be tan to brown but which has a slightly purplish, ‘lavender’ appearance. Cases have been reported in the USA and South Africa (Figure 60.4). The syndrome is invariably fatal and has a highly complex pathogenesis involving critical neurological deficits. Prominent neurological symptoms are present from birth and affected foals usually fail to stand. Convulsions, blindness, and opisthotonus are present and these will inevitably require differentiation from acquired (and possibly treatable) conditions such as trauma, cerebral hypoxia syndromes, tetanus, and septicemic states with systemic infectious response syndrome (SIRS). It may be coupled with other congenital/genetic defects such as cardiac deformities, wry nose, or cleft palate and possible intestinal aganglionosis. There is no treatment; the condition is fatal.
Figure 60.4 This ‘lavender foal’ shows the typical color associated with the condition but the more serious signs are associated with the nervous system – the coat color is simply a visible component of the wider deformity. (Picture courtesy Dr. Alan Nesser.)
Fell Pony immunocompromise disorder This heritable condition resulting in severe immunocompromise is associated with a single recessive gene in pure bred Fell Pony foals under 7–12 weeks of age (see Chapter 36).14–16 There appears to be a high prevalence of the carrier status in the Fell Pony
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Diseases of the Skin population and cases have been reported in several countries including the USA17 and The Netherlands.16 Specific breeding lines carry a major risk of the disease and it is likely that the disease also exists in the closely related Dales pony breed. The condition is not similar to any of the other recognized immunocompromising disorders of foals since the disease is associated with a progressive profound anemia characterized by an aplastic bone marrow (see Figure 60.5) and early onset failure of the cellular immune processes. The white cell counts are usually either within reference or marginally low with a marginal lymphopenia. Colostrally derived immunoglobulin G (IgG) is absorbed normally, but even high concentrations are not usually enough to prevent the early development of opportunistic infections. Cutaneous signs are comparatively minor but provide a good (if non-specific) early warning indicator (Figure 60.5). Symptoms include opportunistic skin (and systemic) infections and a ‘halo’ of russet, long, coarse hairs. This latter sign is sometimes more obvious in particular cases and develops over the first 3 weeks of life, so that, when the foal is viewed from a moderate distance, a faint brown halo, created by the longer brown hairs, is visible. Close-up examination reveals longer russet hairs within the normal black color of the Fell Pony foal (Figure 60.5) and a profound progressive non-regenerative anemia (Figure 60.5). Opportunistic infections, including bacterial dermatitis (Figure 60.5), are frequently encountered and commonly a fungal glossitis with a pseudomembrane develops on the tongue when Candida sp. infection is present (Figure 60.5). Other signs include diarrhea, poor growth and weight gain, and oral and intestinal infections (Figure 60.5). Affected foals are often described as ‘friendly and gentle’ but this masks a lethargy and depression (Figure 60.5) and is singularly unusual in Fell foals. The diagnosis is easily established at necropsy when a profound aplastic bone marrow (Figure 60.5) and severe generalized lymphoid hypoplasia is found. Treatment attempts are fruitless – the condition is 100% fatal. Neither parent of an affected foal should be used for breeding. A specific genetic testing method to identify carriers has recently been developed in the UK. Congenital vascular hamartoma/cavernous hemangioma This is an uncommon congenital cutaneous tumor of vascular endothelial cells that is usually visible on the skin of newborn foals as a papillomatous or sessile exophytic mass (Figure 60.6). It is best viewed
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as a hamartoma; a hamartoma is defined as a local malformation that resembles a neoplasm but is actually a faulty development of an organ or tissue type. In horses they most often affect muscle (muscular hamartoma), blood vessels (vascular hamartoma), and dermal melanocytes. The tumors progress slowly or occasionally more rapidly so that some may be very small at birth and become visible at the weanling stage. Some have an aggressive and invasive nature, expanding to involve adjacent tissues (Figure 60.6d). The commonest sites include the distal limbs (Figure 60.6a–c) and the umbilical area (Figure 60.6d), but the tumors can be found on the face, head, and neck. Solitary, well-circumscribed lesions are possibly more common than the extensive and more aggressive forms with extensive tissue invasion. In the latter case, hemangioma is a better descriptive name. Diagnosis is fairly straightforward as there are few alternative diagnoses. Congenital papilloma is the only major differential. Biopsy is helpful but it does, however, result in significant bleeding. There is no evidence-based information on the possibility of exacerbation following biopsy. High-frequency ultrasonography with color flow is a useful method of defining the extent of the mass and therefore the prospects for success with treatment. Treatment is limited to surgical excision (assuming this is anatomically feasible) with or without radiation. Large surgical defects may require grafting, but in the author’s experience with these cases this has a high rate of failure. Incomplete excision results in recurrence and topical anti-tumor compounds are largely ineffective. Cryosurgery may seem to be an attractive option but again the results are poor. The prognosis is very guarded as some cases are highly invasive and many require repeated interventions before resolution takes place.
Melanotic nevus/melanocytoma/congenital melanoma Occasionally, isolated heavily pigmented, nonencapsulated cutaneous nodules varying in size from 0.5 to 2–3 cm in diameter and number are identified in the skin of gray or non-gray foals at birth or within the first few months of life. The neck, face, and body trunk are more often involved. Individual nodules are often raised and commonly ulcerated to reveal the black/purple tumor tissue (Figure 60.7a,b). The tumors are regarded as benign and, although they do expand with time in some cases, many remain static. Diagnosis is simply by clinical examination and confirmation through submission of surgical excisional biopsy.
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Figure 60.5 The Fell Pony Immunodeficiency Syndrome is characterised by lethargy (a) and often dissociation from the mare (b). There is usually a russet colour to the coat with an apparent halo of longer finer hair (c, d). Opportunistic infections affect the tongue where fungal glossitis (e) is common, the skin, where bacterial (dermatophilosis – Figure f) or fungal infections occur. The most characteristic features of the disease are a progressive non-regenerative anaemia with server mucous membrane pallor (g) and enteric and respiratory tract infections with opportunistic viruses and bacteria. Diarrhoea is common (g). Post mortem findings include a profound paucity of lymphoid tissue and an almost startling lack of active bone marrow (i).
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Figure 60.5 (Continued)
Treatment is limited to surgical excision, which should be successful. Congenital equine cutaneous mastocytosis This is a rare congenital form of mast cell tumor that is present at birth or becomes obvious shortly afterwards.18 Multiple small raised circular/ovoid nodules (3–25 mm diameter) develop along the body trunk and lateral aspects of the hind limbs, in particular. The lesions may ulcerate and become fluctuant. The diagnosis can be confirmed through excisional biopsy. Treatment is not required; in most cases, the lesions regress spontaneously over a variable period. Individual lesions can be surgically removed. Congenital papillomatosis (neonatal wart/epidermal nevus) The equine papillomavirus apparently has the ability to pass across the placenta to infect the skin of the developing fetus. Viral papillomata are not an uncommon incidental condition encountered in newborn foals and they may also be identified on the allantoic surface of the placenta. This may be regarded as a nevus (birthmark or mole) and some cases have no detectable viral involvement. Clinically, the isolated papillomatous lesions of variable size and number are found on any part of the body. Single lesions are most common and some can be quite extensive with ulcerated surfaces (Figure 60.8). Although most are visibly ‘warty’ in appearance, some can be fragile and easily traumatized, causing bleeding and secondary infection (Figure 60.8). Diagnosis will rely on the typical clinical appearance and biopsy.
In contrast to the very common juvenile viral papilloma (milk warts/grass warts) spontaneous resolution does not usually occur. Surgical excision or cryosurgery is, however, curative; recurrence does not occur. The presence of a congenital papilloma does not mean that the foal will not get viral papilloma (‘grass warts’) or sarcoid later in life.
Secondary skin disorders occurring in newborn foals Disorders of other body systems that have cutaneous signs are an important group of conditions in foals. The skin is a major organ and as such is affected by some of systemic, metabolic, nutritional, and toxic conditions that affect foals. The importance of establishing the primary disease is obvious; failure to recognize the underlying primary condition will invariably mean disappointing responses to treatment.
Congenital hypothyroidism Thyroid hormone deficiencies will affect almost every developing organ, but the main visible effects are related to cutaneous and musculoskeletal deformities.19 A paradoxical (hyperiodic) hypothyroidism is recognized in foals born to mares ingesting abnormally high dietary iodine (including nutritional supplement seaweed) during the early/mid-stages of gestation.20, 21 It is common that such affected foals have prominent, bilaterally symmetrical goiter, and the common mistake is to assume that such foals and their dams are hypoiodic. This leads to further supplementation and exacerbation of the problem in the next pregnancy. Multiple skeletal abnormalities (including fused joints, superior or inferior brachygnathism, flexural deformities, incomplete cuboidal bone ossification),22
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(d3) Figure 60.6 Congenital vascular/cavernous hemangioma is a relatively common birth abnormality. Usually, these lesions remain local but become progressively more aggressive (a–c). In some cases they expand widely to become really serious life-threatening tumors (d1–3). The three examples shown here were treated by combinations of surgery and radiation and finally resolved with extensive scarring. In (d) the condition was present at birth (d1) and expanded progressively over 2 months (d2) before becoming ulcerated and hemorrhagic at 4 months (d3). The foal was destroyed.
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Figure 60.7 Solid black nodules or plaques such as are seen in these two cases may be present at birth. Often they are overlooked until the foal is much older. Cream and albino foals seem more likely to be affected. The nodular forms are usually regarded as melanoma while the flat type are probably best termed nevus. (a) a non-progressive melanotic naevus in the lower lid that was present from birth. (b) congenital melanomas on the back of a yearling that started as small solid nodules and gradually progressed. Histologically these had evidence of early malignancy and were removed without recurrence.
poor growth (dysmaturity), skin abnormalities, and prominent goiter are presented. The skin signs are non-specific and of relatively minor consequence but include epidermal atrophy and abnormal keratinization and hair formation. Some cases are almost alopecic and some severe localized or generalized scaling and flaking may be present. Biochemical evidence of hypothyroidism is seldom present if the tests are performed after 24 hours of age, and so static and dynamic thyroid tests such as tri-iodothyronine (T3), thyroxine (T4), thyroid stimulating hormone (TSH) response tests are not significantly helpful in reaching a diagnosis. Much more relevance is paid to the management of the mare and an accurate history and analysis of nutritional sources during pregnancy and the presence of a goiter with a variety of seemingly unrelated developmental abnormalities. Treatment is impossible. Once the physical abnormalities are present they will persist. Thyroid supplementation has no beneficial effect. The prognosis for affected foals is very poor – few survive usefully to become effective athletes.
Ulcerative dermatitis, thrombocytopenia, and neutropenia syndrome This is a recently described transient, but alarming, syndrome affecting neonatal foals less than 4 days old.23 There is no breed, sex, or color predisposition. The etiopathogenesis is uncertain but an alloantibody-related subepidermal clefting is consistent with the signs. Cases have been reported in foals
delivered to mares with pemphigus foliaceus.24 The source of the antibody is likely to be colostral ingestion.25 The panel of signs includes neutropenia and thrombocytopenia, but, as yet, it is unclear what the primary target for the alloantibodies is. Skin crusting, decreased nursing, and hemoptyalism (‘slobbering with blood’) are the primary presenting signs. Extensive cutaneous and mucosal petechial and ecchymotic hemorrhages occur (with nodular hemorrhages in the oral mucosa in particular) (Figure 60.9). Less obvious bulla formation occurs on the head, body, and limb skin. Usually, the foals remain reasonably alert. The condition is transient with spontaneous recovery over 3–7 days. Supportive treatment with corticosteroids, antibiotics, and anti-ulcer drugs helps hasten recovery. Affected foals do not warrant euthanasia! The prognosis for affected foals is excellent in spite of the apparent severity of the early signs. Most cases are significantly improved within 2–5 days. The condition must be differentiated from the incurable epidermolysis bullosa (see above).
Primary congenital/hereditary conditions that become significant/noticeable in older foals and adults One of the serious implications for breeders is the late detection of significant congenital, hereditary diseases. Several significant skin diseases fall into this
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Figure 60.8 Viral papillomas are encountered with some frequency in newborn foals. They may remain static and have no significance clinically but they do not usually resolve spontaneously. Although milder, smaller, localized lesions are commonest (a, b) some can be very large (c). Photograph courtesy of Marianne Sloet.
category. In some cases the disease is hereditary and, by the time it becomes obvious, the animal may have even been used for breeding. Many are more of cosmetic significance and the breeding implications may not be as serious. This is, however, a major reason for breeders to continue to take an interest in the animals they breed for sale.
Linear keratosis/linear alopecia Linear keratosis is an idiopathic, possibly inherited dermatosis characterized by unilateral, linear, vertical bands of hyperkeratosis on the neck, thorax, and upper hind leg. The disorder is probably more common than is appreciated, with most owners ignoring it or simply accepting it. Its pathogenesis and
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Figure 60.9 Ulcerative dermatitis, thrombocytopenia, and neutropenia syndrome. (I) Oral mucosal bullae (I), hemoptyalism, and inappetance are the main signs of this syndrome. Concurrent coronary band (II) and skin bullae and ulceration (III, IV) are also encountered. (Pictures courtesy of Alex Dugdale.)
indeed its pathology are unclear. There may be some heritable tendency but the condition is seen in many breeds to varying extents.26 Most cases occur in Quarter Horses between 1 and 5 years of age. It passes unnoticed in most affected horses. Linear keratosis is otherwise very similar to the condition known as linear epidermal nevus. A particular form also occurs on the palmar aspect of the hind cannons from the fetlock upwards, where it can be classified as an epidermal nevus (Figure 60.10). Linear alopecia is a similar disorder in which alopecia is the main sign. It is seen most often in Quarter Horses and, again, an inherited aspect has been proposed.27 Although both disorders are probably present from an early age, they are usually noted only when the horse is clipped out or is being groomed during summer months. Young foals
may be presented with prominent linear alopecic, hyperkeratotic ‘lines’ on the neck and buttocks – this is sometimes coupled with a brindling of the hair coat. Clinical signs are obvious in most cases and they usually remain constant for many years (if not for life). Few or several, non-painful, non-pruritic linear strips or bands of alopecia and/or hyperkeratinized skin are visible. The lesions are non-inflammatory and may progress slowly or remain static. Treatment is neither indicated nor possible. Keratolytic shampoos and synthetic retinoids may be helpful but have little benefit. They may simply widen the affected area and make it painful! The prognosis is excellent since the condition has no effect on the horse at all and it is best regarded as a cosmetic blemish even when the condition is extensive.
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Figure 60.10 (a) This 2-year-old Haflinger had a prominent hyperkeratotic linear condition from within days of birth. Over the life of the mare it showed no tendency to improvement or exacerbation. Histology confirmed that it was a benign epidermal nevus. (b) Cannon keratosis is a similar condition that occurs on the dorsal metatarsal region almost exclusively.
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Figure 60.11 This heavy horse had prominent scaling and crusting for many years on the palmer carpus (‘mallenders’) (a) and dorsal tarsus (‘sallenders’) (b). Biopsy revealed only a marked hyperkeratosis, but repeated attempts to resolve the problem with keratolytic medications resulted in gradual deterioration.
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Diseases of the Skin Epidermal nevus An epidermal nevus is a form of hamartoma and is a well-defined developmental hyperplasia of the epidermis usually restricted to the plantar metatarsal region (Figure 60.10). A hamartoma is defined as an abnormal accumulation of normal tissue at a normal site. Although it is often tumorous it is not a neoplasm. Many different nevus types can occur, and some of the conditions are given special status with their own name and description. There is no obvious breed, color, or sex predisposition but feathered breeds are possibly more often affected, especially with a hyperkeratotic skin on the dorsal tarsal and the palmar carpal regions; historically, these lesions were called mallenders and sallenders and have been recognized for hundreds of years (Figure 60.11). Often, they remain static for life, but, as they are frequently subjected to unnecessary treatments, gradual exacerbation and extension occur in many cases. They also appear to be an attractive site for surface feeding mites such as Chorioptes spp. An area of partial alopecia and thickened skin with prominent scaling and flaking is usually the main sign. The condition is non-inflammatory and there is no pain or pruritus unless the area is secondarily infested with mites and is then subjected to self-trauma. Treatment is not indicated but keratolytic shampoos and synthetic retinoids may be used if the condition becomes severe. However, repeated treatments will usually result in progressive deterioration and in any case the treatment may be painful.
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Hereditary equine regional dermal asthenia (HERDA) (hyperelastosis cutis/cutaneous asthenia/Ehlers–Danlos syndrome) This condition includes a spectrum of autosomal recessive, inherited, connective tissue diseases, occurring principally in young Quarter Horses, which appear at birth or later in life.28, 29 Thoroughbreds and Arabian horses have also been diagnosed with this condition.30 At least nine different subtypes occur in humans and it has been classified on clinical, genetic, and biochemical differences. The precise nature of the genetic problem in horses is not yet known. Alterations and disorientation of dermal collagen fibrils are thought to be the main pathological basis for the condition.31 A second recognizable syndrome may occur in newborn foals in which an unexplained wound draws attention to a region of abnormal skin attachment (Figure 60.12). The chest wall, medial thigh, and buttock skin are more often affected in the author’s experience. This particular syndrome is not yet widely recognized and it is uncertain whether it falls into this category rather than epitheliogenesis imperfecta (congenital cutaneous aplasia) (see above). Clinical signs are usually delayed until the skin is subjected to significant trauma such as saddle placement. Large areas of skin necrosis and eschar formation occur and either slough and heal slowly or do not heal at all Treatment is impossible. Attempts to graft the defects or remove the affected skin are fruitless. The prognosis is hopeless, although a few cases may be managed provided they are not ridden. Wound care is a long-term challenge.
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Figure 60.12 This extensive skin defect was visible in this newborn foal. There was an apparent complete lack of any connection between the skin and the body wall. Attempts to restore the connection surgically are fruitless, but some cases that involve smaller areas can survive usefully. (Pictures courtesy Jacintha Wilmink.)
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Hypotrichosis/mane and tail dystrophy (follicular dysplasia) Hypotrichosis is a rare hereditary disease found most often in some lines of Arabian horses in which there is less than the normal amount/density of hair. It can affect other breeds on an individual basis and it does not appear to be linked to hair color (Figure 60.13). Lesser forms are sometimes recognized in which there is simply a reduction in hair density over variable areas. The latter are possibly more common than is appreciated. In some cases, the changes can result in severe hair loss. As part of this group of hair disorders, bizarre hair growth patterns are sometimes encountered. These are of no material consequence, but the disorder may result in loss of value for some uses. A more common manifestation of hair dystrophy/thinning is well recognized in the Appaloosa breed, in which the long tail and mane hairs in particular are sparse and brittle. This is regarded as a normal trait in some lines of the Appaloosa horse (Figure 60.14). Mane and tail dystrophy occurring simultaneously have been recorded in a variety of breeds.
Clinical signs are variable depending on the syndrome, but generalized hair loss in a very young foal or mane and tail loss in older Appaloosa horses are typical. Biopsy can help to confirm the scarcity of functioning hair follicles, and characteristic follicular pathology in the mane and tail form may be recognized. No treatments are possible. The conditions do not affect the horse and so it is best viewed as a cosmetic nuisance.
Appaloosa parentage syndrome Appaloosa cross-bred foals often do not have their full color change until 3–4 years of age and then a fading of the spots is sometimes seen so that the horse has a progressive diminishing of the hair color without any associated changes in hair density (Figure 60.15). Treatment is not possible. The trait is sometimes regarded as desirable and is viewed as normal for the breed.
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Figure 60.13 This 5-year-old Thoroughbred cross mare had prominent hypotrichosis of the face (a, b) and tail (c) from birth and by 4 years of age the remaining sparse tail hairs had fallen out. The mane was unaffected.
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Figure 60.14 This 7-year-old Appaloosa shows the typical genetic mane and tail dystrophy syndrome that is prevalent in the breed. Over the following 8 years, the condition showed little progression.
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Figure 60.15 These two pictures show the same pure bred Appaloosa 4 years apart. The definite patterning at the weanling stage is shown in (a) whereas (b) shows the gelding at 5 years old. The gelding continued to lose color definition and at 12 was a uniform very pale roan color. Note also the poor mane and tail density characteristic of mane and tail dystrophy.
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(Arabian) fading syndrome (pinky Arab syndrome) This occurs spontaneously in Arabian horses at any age (2–23 years) but has also been observed in Welsh mountain ponies and Clydesdale horses. The condition is largely symptom free apart from the loss of pigmentation and hair in some cases and may be similar or identical to vitiligo. The latter condition is usually reserved for the more localized periocular condition that occurs in a larger number of breeds and is most often only facial. The face and perineum are more often affected (Figure 60.16). Gray horses are over-represented. There is no known link to copper deficiency or any other pathology but a relationship to endocrine changes in pregnancy has been suggested.32 This does not correspond with the occurrence of the condition in an almost equal number of males (geldings and stallions). Treatment is impossible. Tattooing and skin protection with masks or rugs/sheets are the only options. The main implications lie in the cosmetic aspects and the increased tendency to actinic dermatitis and squamous cell carcinoma in the affected skin. (a)
Dermoid cyst Dermoid cysts are congenital or hereditary cutaneous cysts in young horses, and are thought to be due to embryonic displacement of ectoderm into the subcutis. They are usually single (or few) and most commonly occur along the dorsal midline or the girth region although individual cysts can occur at any cutaneous site. No particular breed predilection has been established. The clinical features are usually distinctive – a few (or single), isolated, slow-growing (or static), noninflammatory nodules in the dorsal midline of young horses are characteristic (Figure 60.17).
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Figure 60.16 This pure bred Arabian stallion developed progressive facial and periocular depigmention which increased his sensitivity to sunburn. He also developed extensive progressive depigmentation in the perineal region.
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Figure 60.17 This horse developed a dermoid cyst in the dorsal midline at 6 months old. It gradually progressed until it was removed surgically when the horse was 3 years old. The lesion was typically in the dorsal midline, solitary, and contained typical sebaceous and hair materials.
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Diseases of the Skin In some cases inflammatory flares do occur, making the site hot and painful. This may be a result of local immune or inflammatory responses to secretions or break down of the products of the cyst. When this occurs, the clinician is faced with a different set of diagnostic differentials but simple historical questions will establish their likely nature. Surgical excision is the only alternative to leaving them alone. They do not generally resolve spontaneously.
References 1. Lieto LD, Cothran EG. The epitheliogenesis imperfecta locus maps to equine chromosome 8 in American Saddlebred horses. Cytogenet Genome Res 2003;102:207–10. 2. Scott DW, Miller WM. Congenital and hereditary skin diseases. In: Equine Dermatology. Philadelphia: W.B. Saunders, 2003; p. 628. 3. Scott DW. Structure and function of the skin. In: Large Animal Dermatology. Philadelphia: W.B. Saunders, 1988; pp. 1–4. 4. Linder KE. Mechanobullous disease of Belgian foals resembles lethal (Herlitz) junctional epidermolysis bullosa of humans and is associated with failure of laminin-5 assembly. Vet Dermatol 2000;11:24–30. 5. Berthelsen H, Eriksson K. Epitheliogenesis imperfecta neonatorum in a foal possibly of hereditary nature. J Comp Pathol Ther 1935;48:285–6. 6. Lieto LD, Swerczek TW, Cothran EG. Equine epitheliogenesis imperfecta in two American Saddlebred foals is a lamina lucida defect. Vet Pathol 2002;39:576–80. 7. Perkins GA, Miller WH, Divers TJ, Clark CK, Belgrave R, Sellon DC. Ulcerative dermatitis, thrombocytopenia, and neutropenia in neonatal foals. J Vet Intern Med 2008;19:211–16. 8. Ginn PE, Hillier A, Lester GD. Self-limiting subepidermal bullous disease in a neonatal foal. Vet Dermatol 1998;9:249–56. 9. Scott DW, Miller WH. Equine seborrhea. In: Equine Dermatology. Philadelphia: W.B. Saunders, 2003; pp. 575–81. 10. Sponenberg DP. Peculiarities of hair growth. In: Equine Color Genetics 3rd edition. Ames, Wiley Blackwell, 2009; pp. 129–30. 11. Knottenbelt DC. Pascoe’s Principles and Practice of Equine Dermatology. Oxford: Saunders, 2009; pp. 233–4. 12. Lightbody T. Foal with overo lethal white syndrome born to registered quarter horse mare. Can Vet J 2002;43:715– 17. 13. Page P, Parker R, Harper C, Guthrie A, Neser J. Clinical, clinicopathologic, postmortem examination findings and familial history of 3 Arabians with lavender foal syndrome. J Vet Intern Med 2006;20:1491–4.
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14. Scholes SF, Holliman A, May PD, Holmes MA. A syndrome of anaemia, immunodeficiency, and peripheral ganglionopathy in Fell Pony foals. Vet Rec 1998;142:1283–9. 15. Richards AF, Kelly DF, Knottenbelt DC, Cheeseman MT, Dixon JB. Anaemia, diarrhoea and opportunistic infection in Fell ponies. Equine Vet J 2000;32:386–91. 16. Butler CM, Westermann CM, Koeman JP, Sloet van Oldruitenborgh-Oosterbaan MM. The Fell Pony immunodeficiency syndrome also occurs in the Netherlands: a review and six cases. Tijdschr Diergeneeskd 2006;131:114–18. 17. Gardner RB, Hart KA, Stokol T, Divers TJ, Flaminio MJ. Fell Pony syndrome in a pony in North America. J Vet Intern Med 2006;20:198–203. 18. Johnson PJ. Dermatologic tumours (excluding sarcoids). Vet Clin North Am Equine Pract 1998;14:633–4. 19. Mair TS. The endocrine system: In: Higgins A, Snyder JR (eds) The Equine Manual, 2nd edn. Oxford: Saunders, 2006; pp. 651–3. 20. McLaughlin BG, Doige CE, McLaughlin PS. Thyroid hormone levels in foals with congenital musculoskeletal lesions. Can Vet J 1986;27:264–6. 21. McLaughlin BG, Doige CE. Congenital musculoskeletal lesions and hyperplastic goitre in foals. Can Vet J 1981;22:130–3. 22. McLaughlin BG, Doige CE. A study of ossification of carpal and tarsal bones in normal and hypothyroid foals. Can Vet J 1982;23:164–8. 23. Perkins GA, Miller WH, Divers TJ. Ulcerative dermatitis, thrombocytopenia and neutropenia in neonatal foals. J Vet Intern Med 2005;19:211–16. 24. Schott HC II, Petersen AD. Cutaneous markers of disorders affecting young horses. Clin Tech Equine Pract 2005;4: 314–23. 25. Perkins GA, Miller WH, Divers TJ, Clark CK, Belgrave RL, Sellon D. Ulcerative dermatitis, thrombocytopenia and neutropaenia in neonatal foals. J Vet Intern Med 2005;19:211–16. 26. Moriello KA. Diseases of the skin. In: Reed SM, Bayley WM (eds) Equine Internal Medicine. Philadelphia: W.B. Saunders, 1998; pp. 531–2. 27. Fadok VA. Update on four unusual equine dermatoses. Vet Clin North Am Equine Pract 1995;11:105–15. 28. Bridges CH, McMullen WC. Dermatosparaxis in quarter horses. Proc Am Coll Vet Pathol 1984;35:12–14. 29. Brounts SH, Rashmir-Raven AM, Black SS. Zonal dermal separation: a distinctive histopathological lesion associated with hyperelastosis cutis in a quarter horse. Vet Dermatol 2001;12:219–24. 30. Rashmir Raven AM, Winand NJ, Read RW. Equine hyperelastosis cutis update. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2004, p. 1409. 31. Finno CJ, Spier SJ, Valberg SJ. Equine diseases caused by known genetic mutations. Vet J, 2009;179:336–47. 32. Thomsett LR. Pigmentation and pigmentary disorders of the equine skin. Equine Vet Educ 1991;3:131–4.
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Diagnostic Approach to Respiratory Disorders Guy D. Lester
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Acute Respiratory Distress Syndrome (ARDS) Pamela A. Wilkins
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Respiratory Diseases Specific to the Neonate Guy D. Lester
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Aspiration Pneumonia Mary Rose Paradis
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Other Disorders of Breathing Pamela A. Wilkins
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Bacterial and Viral Respiratory Disease of the Neonate ´ Renaud Leguillette
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Congenital Abnormalities of the Respiratory System Eric J. Parente
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Genetic Basis of Respiratory Disorders Eliane Marti, Caroline Tessier and Vincent Gerber
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Chapter 61
Diagnostic Approach to Respiratory Disorders Guy D. Lester
In adult animals a thorough physical examination can be highly effective at ruling respiratory disease in or out. In contrast, physical assessment of the respiratory tract of newborn foals can be misleading. It is important to understand which aspects of the examination can provide reliable information and what additional tests may improve the accuracy of a diagnosis of respiratory disease.
Respiratory rate The respiratory rate of the neonatal foal can be elevated for numerous reasons unrelated to pulmonary disease. These include exercise, excitement, pain, hyperthermia, fever, or as a compensatory response to metabolic acidosis. Immediately after delivery the normal foal will increase minute ventilation by raising both the respiratory rate and tidal volume. This can produce rates of 60–80 breaths per minute, which reduce to 30–40 breaths per minute by 1 hour postpartum.1 Respiratory rate and depth can be a misleading indicator of respiratory function. The initial response to hypoxemia is hyperventilation, but both rate and depth of breathing will decline to levels at or below normal when hypoxemia becomes chronic. The response of the neonatal foal to hypercapnia can also be attenuated, particularly if the hypercapnia is persistent or if the foal has experienced an excessive period of asphyxia. In these situations the expected ventilatory response to hypercapnia is lost, with the primary respiratory drive most likely controlled by peripheral chemoreceptors in response to hypoxemia. Treatment of these foals with oxygen (O2 ) therapy will potentially further exacerbate hypercapnic acidosis by removing hypoxemic drive.
Respiratory rhythm Healthy awake neonatal foals usually have a constant respiratory rhythm but may show a variety of ventilation patterns during sleep. The most exaggerated patterns are observed during sleep periods in premature or dysmature foals, and in foals that have experienced prolonged or excessive asphyxia around the time of birth. Patterns include several forms of periodic breathing, including Cheyne–Stokes respiration and cluster breathing. In the former the phase of breathing that follows a period of apnea contains breaths that wax and wane in amplitude before the next period of apnea. In cluster breathing the apneic period ends with a large breath that is greater in amplitude than the breaths that follow. In this type of periodic breathing the apnea is typically longer than the breathing phase; in Cheyne–Stokes breathing the opposite is true. Many post-asphyxia foals will have ataxic breathing patterns, where there is no clear pattern to ventilation. Apneic periods can be prolonged, occasionally more than 30 seconds. During episodes of periodic or ataxic breathing both respiratory frequency and tidal volume are affected,2 and it is possible that these breathing patterns could further contribute to the pathology associated with peripartum asphyxia. Lengthy apneic periods can acutely lead to retention of CO2 and a pronounced respiratory acidosis. These foals may benefit from a respiratory stimulant, such as doxapram hydrochloride (0.02–0.05 mg/kg/h as i.v. constant rate infusion (CRI)) or caffeine (10 mg/kg loading dose orally, then 2.5–5.0 mg/kg p.o. once a day), or assisted ventilation. Chronically, the primary respiratory acidosis is often well compensated metabolically, such that the arterial pH may approach normal. Treatment of these foals will often produce a pronounced alkalosis.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Thoracic and abdominal wall movement A flexible, and therefore highly compliant thoracic wall is critical for safe delivery through the maternal pelvic canal. Unfortunately, this can have a negative impact on postnatal ventilation in compromised foals.3 When the diaphragm contracts to initiate inspiration there is a reduction in pleural pressure that not only causes the lungs to inflate but also exerts an inward pull on the thoracic wall. The highly compliant wall of the newborn is prone to distortion. These inward forces are offset, in part, by contraction of the external intercostal muscles in healthy animals. In foals with sustained high ventilation rates these muscles may fatigue leading to paradoxical thoracic wall motion, i.e., inward thoracic wall movement during inspiration. This is a sign of impending respiratory failure. Premature or dysmature foals may also show thoracic distortion due to structural immaturity and incomplete proprioceptive control of intercostal muscles. There is an interaction between thoracic wall compliance and positioning, such that lateral recumbency leads to reduced minute ventilation and poor gas exchange. This is further complicated by dependent atelectasis of the down lung.4 Placement of foals into sternal recumbency not only improves ventilation but also may reverse paradoxical thoracic wall motion. Thoracic trauma and rib fracture can occur as a complication of second-stage labor. Costochondral dislocation or fractures of the rib shaft can cause significant clinical disease and death. This can include hemothorax, lung laceration and pneumothorax, and pericardial/myocardial puncture. Diagnosis is by observation of thoracic wall symmetry and synchrony, palpation, and ultrasonography. Management is typically conservative unless displacement of shaft fragments is identified over key underlying structures. Normal expiration in adult horses begins with passive recoil of lung tissue and is facilitated by active abdominal muscle contraction at the end of expiration. A similar pattern of expiration is seen in foals, irrespective of positioning.3, 4 An exaggerated abdominal movement is abnormal and would be consistent with pulmonary disease. A prolonged abdominal component during expiration may be associated with audible grunt at the end of the respiratory cycle. Foals with failing lung function will close their glottis during expiration to maintain pressure in the respiratory tree, thereby maximizing gas exchange. The grunt is caused by the sudden release of expired air associated with opening of the glottis. A pronounced abdominal movement during inspiration could be associated with an extrathoracic airway obstruction or restrictive lung disease.
Examination of the upper airway Both nares should be examined closely for evidence of abnormal or absent airflow and discharge. Absence of airflow from a nostril could be consistent with choanal atresia. Mucoid or purulent nasal discharge in response to lower airway disease is uncommon during the early neonatal period, but will often develop as the disease becomes chronic. The presence of milk should prompt further observation after sucking, and examination of the oral cavity to assess integrity of the soft and hard palate. A cleft in the palate will characteristically result in nasal regurgitation of milk immediately after sucking; however, if the defect is small the regurgitation may be inconsistent and of small volume making diagnosis difficult. The palate can be visually assessed and palpated using an upturned and clean middle finger. Most defects occur in soft palate. Small clefts may be difficult to diagnose and may require endoscopy of the nasal passages and/or oral cavity under anesthesia for confirmation. The presence of milk at the nares after feeding is not pathognomonic for a cleft palate. On the contrary, many foals with this complaint do not have a palate defect but rather delayed or altered coordination of the swallowing reflex. Clinical signs may be noted as early as a few hours of age, and persist variably from hours through weeks postpartum. The basis of the condition is not known, but generalized weakness, hypoxic-ischemic brain disease, or nutritional myodegeneration (white muscle disease) have all been suggested as possible causes. Chronically affected foals will develop signs consist with aspiration pneumonia, including cough, ill-thrift and failure to grow. The diagnosis is confirmed through endoscopy of the nasopharynx, where persistent dorsal displacement of a flaccid soft palate and dorsal collapse of the nasopharynx are typical findings. The palate will not replace during frequent attempts to swallow. The procedure is important to rule out congenital defects of the palate and epiglottis, although endoscopy of the oral cavity under short-acting anesthesia may be necessary to rule out more obscure problems. Soft palate displacement may also occur in newborn foals due to a persistent frenulum between the ventral aspect of the epiglottis and base of the tongue or with hypoplasia of the epiglottis.5 Radiography and ultrasonography are useful diagnostic tests to confirm the presence and extent of a ventral consolidating pneumonia that develops secondary to aspiration. Inspiratory stridor or dyspnea can be due to several developmental abnormalities including choanal atresia, guttural pouch tympany, pharyngeal or subepiglottic cysts, and tracheal collapse. A case of progressive ethmoidal hematoma was reported in a 4-week-old foal resulting in respiratory stridor,
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Diagnostic Approach to Respiratory Disorders epistaxis, and facial deformity.6 Airway stridor has also been reported in neonatal Quarter Horse foals experiencing episodic attacks of hyperkalemic periodic paralysis.7
Cough Cough is uncommon in foals with respiratory disease during the early neonatal period. This is most likely attributable to a delayed maturation of irritant receptors within the airway. The laryngo-pharyngeal cough reflex,which is induced through vocal fold stimulation, also appears delayed in newborn foals. As a consequence milk aspiration may occur without any overt signs. This is particularly relevant when inexperienced carers attempt to force-feed sick foals. Foals with lower airway disease will usually develop cough after several days of age.
Rectal temperature The rectal temperature of the neonatal foal is typically between 37.2 and 38.3◦ C (99.0–101.0◦ F) although temperatures up to 38.8◦ C (102.0◦ F) may be considered normal. Body temperature is influenced by a range of factors including environmental conditions (temperature, humidity, wind), exercise, maturity (premature and dysmature foals often have difficulty in regulating body temperature), seizure activity, and drug therapy (e.g., phenobarbitone). Fever is an inconsistent finding in neonatal foals with earlyonset pulmonary disease; as the disease becomes chronic, fever along with cough and nasal discharge become more consistent clinical findings.
Auscultation Auscultation of the respiratory tract is an essential diagnostic technique but can be misleading. Moist rales are normally present throughout the respiratory cycle after birth due to residual fluid within the airways. Asymmetry of air movement and endinspiratory crackles are also a normal finding due to simple atelectasis of the dependent lung during lateral recumbency. These typically resolve within minutes of standing. Conversely, foals can have significant lung disease yet few abnormal findings on thoracic auscultation.
Activity Foals with mild-to-moderate respiratory insufficiency may still appear to be relatively bright and active. This may be due, in part, to a small but lingering adaptation to the relative hypoxic environment in which the fetus develops. Reported umbil-
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ical vein and artery oxygen tensions in late gestational fetal foals are around 48 and 31 mmHg, respectively.8 These values compare with uterine artery and vein partial pressures of 101 and 51 mmHg. The mean difference in Po2 between uterine and umbilical veins is approximately 2–5 mmHg, well below that of pregnant cows and sows, reflecting the highly efficient placental exchange that occurs between mare and foal.8 In contrast to most other species studied, equine fetal hemoglobin is structurally identical to that of the adult animal.9 Lower concentrations of 2,3-diphosphoglycerate (2,3-DPG) in fetal erythrocytes result in shifting of the oxyhemoglobin dissociation curve to the left, promoting an increase in O2 affinity.10 Enhanced O2 affinity of fetal blood facilitates transplacental O2 exchange in the developing fetus and enhanced pulmonary exchange in the immediate postpartum period. Red cell 2,3-DPG concentrations slowly rise to adult levels over the first 5 days of postpartum life.10 Increased O2 affinity of hemoglobin will also occur with hypothermia and alkalosis, but conversely will be decreased in hyperthermia and acidosis. A decrease in affinity is advantageous under certain conditions, as it promotes unloading of O2 in tissues with metabolic needs.
Mucous membrane color Mild to moderate hypoxemia will often fail to result in distinct membrane color change. It is only when the hypoxemia is severe that overt membrane cyanosis becomes apparent. This usually coincides with markedly reduced hemoglobin saturation and partial pressures of O2 , often less than 40 mmHg.
Advanced diagnostic techiques Identification of infectious agents A high index of suspicion of infectious respiratory disease is based on history, physical examination findings, and laboratory data. Hematology is commonly abnormal in bacterial and viral respiratory disease, but changes are not consistent, and vary with agents involved and chronicity. Acutely infected foals tend to be leukopenic, characterized by neutropenia and lymphopenia. Neutrophil toxicity is common. As bacterial infections become chronic then a leukocytosis with mature neutrophilia is expected. The acutephase protein response to infection, as measured by fibrinogen or serum amyloid A concentration, is also delayed. There are a number of techniques that can aid in the isolation of causative infectious agents. Blood culture can provide good indirect evidence of offending respiratory pathogens as bacteremia occurs commonly. Using aseptic technique, 10 mL of whole
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blood is collected and placed into a blood culture medium, which is incubated at 37◦ C before plating onto culture media. Ideally blood is collected before antibiotic treatment is started. Sampling directly from the large airway increases the chances of recovering offending bacteria but should be done with great caution in foals with overt respiratory signs. Tracheal aspiration is best performed transcutaneously using aseptic technique. Commercial kits are available that include a Teflon introduction catheter with a 70-cm 16-gauge flushing catheter (Mila International, Erlanger, KY). Culturing the tip of a nasotracheal tube after replacement is indicated if the practice has facilities for mechanical ventilation. Submission of nasal swabs for bacterial culture is not recommended as recovery of mixed normal upper airway flora is expected. The identification of viral infections can be difficult. Most viral infections of the newborn or young foal that involve the lower respiratory tract are fatal, the best described of which are equine herpesvirus 1 (EHV-1) and equine influenza. An antemortem diagnosis of both of these diseases could be achieved through PCR testing of nasal swabs. Arterial blood gas (ABG) analysis The mainstay of assessment of the respiratory system is measurement of arterial partial pressures of oxygen (Pa o2 ), carbon dioxide (Pa co2 ), oxygen saturation (Sa o2 ), and pH. The equipment to process the ABG has traditionally been limited to institutional or large referral practices. The introduction of less R , Abbott expensive, point-of-care units (e.g., i-Stat R Point of Care Inc., NJ; IRMA TruPoint , ITC, NJ; R R pHOx , Nova Biomedical, MA; and Stat Profile R ABL80 FLEX , Radiometer, OH) enables greater use of this technology. Direct measurement of Sa o2 using a co-oximeter is the preferred method; however, most blood gas analyzers do not have inbuilt cooximeters and calculate Sa o2 from Pa o2 , using the oxygen-dissociation curve and assume that no abnormal forms of hemoglobin are present. Pulse oximetry can also be used to estimate hemoglobin saturation with Pa o2 again calculated using a standard oxygen-dissociation curve. The technique compares the absorption of red and infrared light wavelengths. Oxyhemoglobin absorbs infrared light (910 nm) and allows red light (660 nm) to pass, while reduced hemoglobin absorbs red light and permits infrared light to pass. There are some issues relating to reliability in horses, including movement artifact and inaccuracy associated with poor peripheral perfusion.11 The ABG sample is usually collected from the dorsal metatarsal artery, easily palpated in most foals on the lateral aspect of the third metatarsal bone. Alternative sites include the brachial artery, located at
the level of the medial collateral ligament of the elbow joint, or the carotid artery. Hematoma formation is a common sequel to sampling from the carotid artery. The sample needs to be handled appropriately, paying strict attention to avoidance of air contamination, which will artificially increase Pa o2 and decrease Pa co2 . The sample should be processed within 15 min of collection. Oxygen will leak through plastic syringes and if a delay is anticipated then collection into a less permeable glass syringe and placement on ice is preferred. This should preserve the oxygen content for up to 90 min.11 Plastic syringes are not suitable for preservation of sample Po2 for more than 15 min, irrespective of whether they are stored on ice or at ambient temperature. In contrast both sample Pco2 and pH are relatively stable in plastic syringes at room temperature for up to 90 min.12 The interpretation of the ABG sample involves consideration of the amount of struggling and position of the foal during sample collection. Lateral recumbency can reduce the Pa o2 by as much as 30 mmHg.4 The two most common respiratoryderived ABG derangements include hypoxemia with normo- or hypo-capnia, and hypoxemia with hypercapnia. It is important to distinguish acute from chronic hypercapnia. Acute hypercapnia is associated with a more substantial drop in blood pH and may lead to circulatory collapse and coma, particularly if accompanied by acute hypoxemia. Chronic exposure to elevated CO2 permits adaptation and more subtle clinical effects. The change in pH is less dramatic primarily because of enhanced bicarbonate reabsorption in the proximal tubules of the kidney. This effect begins within 6–12 hours of exposure to increased concentrations of CO2 and is maximal by 3 to 4 days. Hypercapnia can be exacerbated by fever or the administration of carbohydrates or bicarbonate. The latter is often clinically relevant and highlights the danger of giving sodium bicarbonate to foals with pulmonary disease. The four main causes of hypoxemia in any animal are ventilation/perfusion mismatch (V/Q mismatch), right-to-left shunting, alveolar hypoventilation, and diffusion impairment. Except for right-toleft shunting, these problems typically respond favorably to O2 therapy. Oxygen therapy is indicated in neonatal foals when the Pa o2 is less than 60 mmHg or the Sa o2 is less than 90%, with the goal of therapy being a Pa o2 between 75 and 110 mmHg and an Sa o2 above 92%.13
Thoracic radiography Radiographs can add important information as to the nature, extent, and location of disease, as well as providing evidence of disease progression and response
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Diagnostic Approach to Respiratory Disorders to treatment. The diagnostic technique is commonly limited by equipment constraints. The most common artifact is produced by motion, often due to a combination of a fast respiratory rate, foal movement, and inadequate equipment.14 Longer exposure times may be required to compensate for the reduced amperage of some portable machines. Ideally large cassettes (35 × 43 cm) are available and a grid (8:1 to 10:1 grid ratio) is used to reduce image-degrading scatter radiation. The views required are dictated by the foal’s age and disease severity. Most commonly, right and left lateral projections are obtained with the foal standing or in lateral recumbency. The portion of the lung cranial to the heart is partly obscured by the triceps muscle mass in the standing foal. In the nonrecumbent animal the side of the thorax closest to the cassette is preferentially visualized. Lesions on the opposite side of the thorax, i.e., furthest away from the cassette, will be magnified with less distinct margins. In recumbent views the down lung undergoes a degree of dependent atelectasis; the uppermost lung is better aerated, providing greater contrast for detecting abnormal lung patterns. This highlights the importance of taking both left and right projections, particularly in recumbent foals. Ventrodorsal projections can provide some additional lateralizing information, but are usually avoided due to stress on both the foal and its handlers, as well as producing a negative effect on ventilation. The normal newborn foal will have an increased lung density and blurring of the small vascular structures; however, these changes, which are due to a combination of residual small airway fluid, uptake of fluid into the lung interstitium, and incomplete lung inflation, should clear by 12–24 hours of age. The thymus is a significant soft tissue opacity that lies cranial to the heart in the foal, peaking in size at around 2 months of age before regressing.
Figure 61.1 thorax.
Lateral thoracic radiograph of a normal foal
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Figure 61.2 Lateral thoracic radiographic demonstrating a moderate diffuse interstitial pattern.
Figures 61.1 to 61.4 show normal and examples of abnormal radiographic patterns.14 Slight blurring of the pulmonary vessels represents the mildest form of the interstitial lung pattern. In a moderate interstitial pattern some vessels remain visible, bronchial walls are more evident, and there may be reticulated soft tissue opacities. When the pattern becomes severe vessels become indistinct, there is progression of the bronchial markings, and coalescence of reticulated soft opacities. Extension to this pattern is the alveolar pattern, where air bronchograms become visible within the soft opacities. This pattern is caused by the contrast of a large air-filled bronchus with adjacent fluid-filled alveoli. The term ‘interstitial pattern’ is a radiographic description and does not strictly imply a specific anatomical location. Processes that result in this pattern include viral and bacterial infections and non-infectious causes, such as recumbency (dependent atelectasis), abdominal distension, meconium
Figure 61.3 Severe alveolar pattern in a newborn foal with neonatal respiratory distress syndrome.
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Figure 61.4 An example of a severe diffuse reticulated interstitial pattern.
sponsible for the accumulation of pleural fluid in foals with uroperitoneum is not known but could be due to re-expansion pulmonary edema, acute lung injury, and/or altered pleural lymphatic drainage.16 Pleural effusion as an extension of viral or bacterial pneumonia is rare in neonates. Pleuropneumonia due to Streptococcus equi ssp. zooepidemicus has been seen in foals as young as 3 months of age. The dorsocaudal lung region is the most commonly affected area radiographically in neonates, but as foals age and ambulate any infiltrates tend to settle caudoventrally, such that this region tends to worsen over time and is the last to clear.14 In the majority of cases improvement in respiratory function precedes the clearing of radiographic infiltrates.
aspiration, and lung immaturity. Differentiation between infectious and non-infectious causes is often not possible using radiographic interpretation alone. Alveolar patterns are most commonly associated with severe bacterial and viral infections, or neonatal respiratory distress syndrome. Other patterns include miliary nodular, often attributable to Actinobacillus equuli infection, and nodular. Rhodococcus equi infection in older foals classically produces a nodular pattern, although mixed interstitial and alveolar patterns are possible. From a practical perspective ultrasound is the preferred image modality for detection of preclinical and clinical cases.15 A single bulla or multiple bullae can occur congenitally or in response to ventilation trauma; bullae may or may not be associated with clinical signs. Ruptured bullae have been associated with pneumomediastinum, pneumothorax, and subcutaneous emphysema (Figure 61.5).15 Accumulation of pleural fluid is uncommon, but may occur with thoracic wall trauma or rib fracture (hemothorax) or uroperitoneum. The mechanism re-
Thoracic ultrasonography
Figure 61.5 Widespread subcutaneous emphysema secondary to a ruptured lung bullae.
Figure 61.6 Ultrasonographic appearance of a pyogranulomatous lesion caused by Rhodococcus equi in a 6-week-old foal.
Ultrasonography is the preferred modality to identify rib pathology. Linear higher-frequency probes (7.5 MHz or higher) are preferred. Ultrasound is reportedly more sensitive than physical examination or radiography in identifying rib disease.17 Most rib fractures reportedly occur within 3 cm of the costochondral junction, with the distal rib fragment displaced laterally in almost all cases. Fractures can result in a range of clinical scenarios, from none through fatal hemorrhage. Signs can include overt distortion of the thoracic wall, subcutaneous emphysema, respiratory distress, hypovolemic shock, and cardiac arrest. Ultrasonography is the preferred imaging tool for assessment of the pleural space, and can be a useful aid in identifying and monitoring the progression of some lung parenchymal diseases. The technique is ideally suited to identify foals with multinodular lesions caused by infection with Rhodococcus equi (Figure 61.6).
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Diagnostic Approach to Respiratory Disorders
References 1. Rossdale PD. Measurements of pulmonary ventilation in normal newborn thoroughbred foals during the first three days of life. Br Vet J 1969;125:157–61. 2. Lorenzi-Filho G, Genta PR, Figueiredo AC, Inoue D. Cheyne-Stokes respiration in patients with congestive heart failure: causes and consequences. Clinics 2005;60:333–44. 3. Koterba AM, Wozniak JA, Kosch PC. Respiratory mechanics of the horse during the first year of life. Respir Physiol 1994;95:21–41. 4. Kosch PC, Koterba AM, Coons TJ, Webb AI. Developments in management of the newborn foal in respiratory distress 1: Evaluation. Equine Vet J 1984;16:312–18. 5. Yarbrough TB, Voss E, Herrgesell EJ, Shaw M. Persistent frenulum of the epiglottis in four foals. Vet Surg 1999;28:287–91. 6. Colbourne CM, Rosenstein DS, Steficek BA, Yovich JV, Stick JA. Surgical treatment of progressive ethmoidal hematoma aided by computed tomography in a foal. J Am Vet Med Assoc 1997;211:335–8. 7. Traub-Dargatz JL, Ingram JT, Stashak TS, Kiper ML, Tarr S, Child G, MacAllister CG. Respiratory stridor associated with polymyopathy suspected to be hyperkalemic periodic paralysis in four quarter horse foals. J Am Vet Med Assoc 1992;201:85–9. 8. Comline RS, Silver M. A comparative study of blood gas tensions, oxygen affinity and red cell 2,3-DPG concentrations in foetal and maternal blood in the mare, cow and sow. J Physiol 1974;242:805–26.
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9. Stockell A, Perutz MF, Muirhead H, Glauser SC. A comparison of adult and foetal horse hemoglobins. J Molec Biol 1961;3:112–16. 10. Bunn HF, Kitchen H. Hemoglobin function in the horse: the role of 2,3-diphosphoglycerate in modifying the oxygen affinity of maternal and fetal blood. Blood 1973;42: 471–9. 11. Wilson, DV. Monitoring respiratory function. In: Doherty T, Valverde A (eds) Manual of Equine Anesthesia and Analgesia. Ames: Blackwell, 2006; pp. 191–9. 12. Picandet V, Jeanneret S, Lavoie JP. Effects of syringe type and storage temperature on results of blood gas analysis in arterial blood of horses. J Vet Intern Med 2007;21: 476–81. 13. Palmer JE. Ventilatory support of the critically ill foal. Vet Clin North Am Equine Pract 2005;21:457–86. 14. Lester GD, Lester NV. Abdominal and thoracic radiography in the neonate. Vet Clin North Am Equine Pract 2001;17:19–46. 15. Marble SL, Edens LM, Shiroma JT, Savage CJ. Subcutaneous emphysema in a neonatal foal. J Am Vet Med Assoc 1996;208:97–9. 16. Wilkins PA. Respiratory distress in foals with uroperitoneum: possible mechanisms. Equine Vet Educ 2004;16:293–5. 17. Jean D, Picandet V, Macieira S, Beauregard G, D’Anjou MA, Beauchamp G. Detection of rib trauma in newborn foals in an equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 2007;39:158–63.
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Chapter 62
Acute Respiratory Distress Syndrome (ARDS) Pamela A. Wilkins
Introduction Acute lung injury (ALI) and acute respiratory distress syndrome (ARDS) are syndromes of severe pulmonary dysfunction and respiratory failure caused by physical or chemical injury or an exaggerated pulmonary immune response.1–4 In human medicine ALI/ARDS is defined by an acute onset of respiratory dysfunction, bilateral pulmonary infiltrates, a ratio of pulmonary arterial oxygen pressure (Pa o2 ) to fraction of inspired oxygen (Fi o2 ) of < 300 mmHg for the less severe form, ALI, and < 200 mmHg for ARDS and no clinical evidence of left atrial hypertension or a pulmonary arterial occlusion pressure of < 18 mmHg.1 The definition acknowledges ALI/ARDS as a clinically recognized condition, regardless of the inciting cause of the respiratory failure.5 In ALI/ARDS an imbalance of pro- and antiinflammatory factors combined with activation of the pulmonary endothelium and derangement of the coagulation cascade favors the development of a fulminate, uncontrolled pulmonary inflammatory response and a widespread procoagulant environment in alveoli and pulmonary microcirculation.5 The triggering event for ALI/ARDS in humans, or any of the veterinary forms, may be an intrapulmonary (viral or bacterial pneumonia, smoke inhalation, near drowning, or food aspiration) or extrapulmonary (trauma, multiple transfusions, systemic inflammatory response, or sepsis) insult.5, 6 The effect on the patient is severe hypoxemia with hypo- or hypercapnia caused by ventilation/perfusion (VA /Q) mismatch and decreased pulmonary compliance. Resolution of ALI/ARDS depends on functional distal lung epithelium. If damage to the pulmonary epithelium is minor, pulmonary edema can be cleared swiftly, pulmonary function improves
rapidly, and chances of survival increase. If large areas of alveolar epithelium are destroyed, repair must occur before the edema can be cleared.5 Severe damage to the epithelium corresponds with prolonged respiratory failure, slow recovery by gradual regeneration of the respiratory epithelium, and a high mortality.7 Less than 5% of human patients die of refractory hypoxemia. The majority of patients succumb to their primary disease process and multiorgan failure.8 An ALI/ARDS-like syndrome has been described in 1- to 7-month-old foals and occasionally in neonatal foals.2,9–12 In earlier reports the condition was named acute interstitial or bronchointerstitial pneumonia. Multiple infectious and non-infectious agents are able to initiate the inflammatory response. In foals, the most common underlying condition appears to be bacterial pneumonia and, in rare cases, viral pneumonia.
Clinical signs and diagnosis Clinically, it is almost impossible to differentiate foals with ALI/ARDS from foals with severe bacterial or viral pneumonia. ALI/ARDS is likely triggered in many cases by an underlying bacterial infection. Clinical signs include profound depression, fever, and respiratory distress with tachycardia, tachypnea, nostril flare, and cyanotic mucous membranes. Auscultation of the lungs can reveal a variety of abnormal sounds or even complete silence over non-ventilated areas. Laboratory findings include severe hypoxemia with hypo- or hypercapnia, and, depending on the underlying disease process, leukocytosis or leukopenia and hyperfibrinogenemia. Transtracheal lavage fluid demonstrates neutrophilic inflammation with
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Acute Respiratory Distress Syndrome (ARDS)
Figure 62.1 Radiographic appearance of the thorax of a foal with severe ARDS. Note the diffuse interstitial and alveolar patterns throughout the lung that obscure the normal vascular pattern, the caudal vena cava, and the caudal cardiac silhouette.
or without signs of infection, depending on the underlying etiology. Thoracic radiographs demonstrate a diffuse, dense bronchointerstitial pattern often coalescing to a focal or diffuse alveolar pattern with prominent air bronchograms (Figure 62.1). Computed tomography (CT) reveals bilateral dependent density gradients. The initial radiographic appearance of the lungs may worsen dramatically within hours due to rapid progression of pulmonary edema. On the other hand, absorption of the alveolar fluid may result in impressive improvement of the radiographic appearance within days.2, 11 On gross pathological examination, the affected lungs are wet, firm, fail to collapse, and airways contain variable amounts of pink, foamy liquid. Histologically, necrosis of alveolar and terminal bronchial epithelium is observed with extensive filling of the alveolar spaces with neutrophils, macrophages, hem-
orrhage, protein-rich edema fluid, or hyaline membranes. Occasionally, microthrombi can be detected in interstitial capillaries. In more chronic cases, diffuse proliferation of cuboidal pneumocytes and fibroblasts and beginning fibrosis may be present. Diffuse alveolar damage (DAD) is the common histopathological injury. Recently, diagnostic criteria for equine neonatal ALI/ARDS (EqNALI/NARDS), and also for primary surfactant deficiency ALI/ARDS, known now as neonatal equine respiratory distress syndrome (NERDS), were described in a veterinary ALI/ARDS consensus statement (Table 62.1 and Boxes 62–3).3 Presence of a primary disease process such as bacterial or viral pneumonia does not rule out the concurrent presence of ALI/ARDS. It is likely that the provided Pa o2 :Fi o2 ratios will be altered with further investigation, but are presented as rational initial evaluation cut-points. Therapy In human medicine, mechanical ventilation using low tidal volumes (6.0 mL/kg) and some form of positive end-expiratory pressure (PEEP) is the treatment of choice.13 Owing to a lack of availability for mechanical ventilation in many areas, the most important and practical treatment remains intranasal insufflation of humidified oxygen via uni- or bilateral intranasal cannulas at high flow rates (10–30 L/min). Alternatively, intratracheal O2 insufflation rates have been described, achieving results similar to high bilateral intranasal flow rates.14 Anti-inflammatory treatment is also essential; although the use of corticosteroids in human ALI/ ARDS is controversial, a recent small study reported reduced mortality and decreased need for ventilation in the group treated with low doses of corticosteroids (2.0 mg/kg methylprednisolone intravenous (i.v.) loading dose followed by 2.0 mg/kg/day i.v.
Table 62.1 Diagnostic criteria for equine neonatal acute lung injury/respiratory distress syndrome (Eq NALI/NARDS): same as for Veterinary acute lung injury/Veterinary acute respiratory distress syndrome (VetALI/VetARDS) but adhering to the following age-dependent adaptation of the Pa o2 /FI o2 ratio in term foals in lateral recumbency breathing room air (FI o2 = 0.21) based on age-dependent normal values for Pa o2 under similar conditions. Reprinted with permission from Wilkins et al.3 Postnatal age 60 min 12 hours 24 hours 48 hours 4 days 7 days
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Normal P a o2 (mmHg)
Normal P a o2 /F I o2 ratio
NALI
NARDS
60.9 ± 2.7 73.5 ± 3.0 67.6 ± 4.4 74.9 ± 3.3 81.2 ± 3.1 90.0 ± 3.1
> 300 mmHg > 350 mmHg > 350 mmHg > 350 mmHg > 400 mmHg > 430 mmHg
< 175 mmHg < 200 mmHg < 200 mmHg < 200 mmHg < 250 mmHg < 280 mmHg
< 115 mmHg < 140 mmHg < 140 mmHg < 140 mmHg < 160 mmHg < 190 mmHg
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Box 62.1 Neonatal equine respiratory distress syndrome (NERDS) Etiology: Primary surfactant deficiency due to failure of final fetal pulmonary surfactant metabolism maturation. Diagnosis requires meeting all criteria listed below: 1. Persistent hypoxemia, progressive hypercapnia a Hypoxemia defined as Pa o2 < 60 mmHg taken with foal in lateral recumbency while breathing room air 2. Appropriate risk factors present: (1 or more of 3) a Very early gestational age: less than 290 days of gestation OR less than 88% of average of dam’s previous gestations b Induction of parturition c Cesarean section 3. Failure to develop normal immediate postpartum respiratory patterns: development/persistence of paradoxical breathing over first several hours of life, persistent tachypnea 4. At 24 hours of age or less ‘ground glass’ appearance of lateral radiograph(s) of lungs taken either standing or in lateral recumbency 5. Absence of evidence of fetal inflammatory response syndrome (FIRS) at birth a Normal white blood cell count, differential, and fibrinogen concentration for gestational age 6. Congenital cardiac disease ruled out as a cause of hypoxemia Reprinted with permission from Wilkins et al.3
Box 62.2 Veterinary acute lung injury and acute respiratory distress syndrome (VetALI/VetARDS): must meet at least one each of the first four criteria; criterion 5 is a recommended but optional measure: 1. Acute onset (< 72 hours) of tachypnea and labored breathing at rest Known risk factors (see Box 62.3) 2. Evidence of pulmonary capillary leak without increased pulmonary capillary pressure∗ – (any one or more of the following): a Bilateral/diffuse infiltrates on thoracic radiographs (more than 1 quadrant/lobe) b Bilateral dependent density gradient on CT c Proteinaceous fluid within the conducting airways d Increased extravascular lung water
3. Evidence of inefficient gas exchange (any one or more of the following): a Hypoxemia without PEEP or CPAP and known FI o2 i. Pa o2 /FI o2 ratio ii. ≤ 300 mmHg VetALI iii. ≤ 200 mmHg VetARDS iv. Increased A–a gradient v. Venous admixture (non-cardiac shunt) b Increased ‘dead-space’ ventilation 4. Evidence of diffuse pulmonary inflammation a Transtracheal wash/bronchoalveolar lavage cytological neutrophilia b Transtracheal wash/bronchoalveolar lavage biomarkers of inflammation c Molecular imaging (PET) ∗ No evidence of cardiogenic edema (one or more of the following): PAOP <18 mmHg (adult horse) No clinical or diagnostic evidence supporting left heart failure, including echocardiography. Reprinted with permission. Wilkins et al.3 CT, computed tomography; PAOP, pulmonary artery occlusion pressure; PEEP, positive end expiratory pressure; CPAP, continuous positive airway pressure; Fi o2 , fractional inspired oxygen concentration; PET, positron emission tomography. divided in four treatments).15 Considering that ventilation is rarely an option and that in two reports all but two foals surviving ALI/ARDS received corticosteroids, anecdotal evidence is in favor of steroid
Box 62.3 Risk Factors for veterinary acute lung injury and acute respiratory distress syndrome (VetALI/VetARDS) 1. 2. 3. 4. 5.
6. 7. 8. 9. 10.
Inflammation Infection Sepsis Systemic inflammatory response syndrome (SIRS) Severe trauma a Long bone fracture b Head injury c Pulmonary contusion Multiple transfusions Smoke inhalation Near-drowning Aspiration of stomach contents Drugs and toxins
Reprinted with permission. Wilkins et al.3
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Acute Respiratory Distress Syndrome (ARDS) administration.2, 11 Intravenous prednisolone sodium succinate or methylprednisolone at 1–2 mg/kg/day divided in 2–4 doses may be useful. Antimicrobial treatment, if required, should be directed against an identified underlying disease process. Bronchoconstriction is not a prominent feature of ALI/ARDS and bronchodilators are therefore of limited use, but may be beneficial in an individual case. As described earlier, clearance of edema fluid from the lungs depends on active transepithelial ion transport, a process that can be stimulated by β2 -agonists such as salmeterol, terbutaline, and epinephrine. There is some evidence in experimental models of ALI that β2 -agonists delivered into the airways reduce pulmonary edema by attenuating pulmonary vascular injury and upregulating fluid clearance from the air spaces.16 Care should be taken when administering bronchodilators as they can induce sudden worsening of V/Q mismatch and lead to acute decompensation of the severely hypoxic patient. Judicious intravenous fluid therapy may be necessary in dehydrated patients. Measurement of central venous pressure (CVP) may aid in determination of the patient’s hydration status, but applied intrathoracic pressure changes should be considered carefully in ventilated foals as CVP may be spuriously altered. If foals are too depressed to nurse or eat, nutritional support is essential. Several other treatments have been suggested in human medicine including vasodilators such as inhaled nitric oxide or prostacycline, surfactant to reduce surface tension, and scavengers of reactive oxygen species (tocopherol, ascorbic acid); however, none have been proven to increase survival despite the fact that individual treatments improved pulmonary function.8 Considering that, in contrast to human medicine, most equine patients die as a direct result of respiratory failure and severe hypoxemia, some of those therapeutic options may benefit the equine patient. Based on the limited information available, the prognosis for survival and future athletic performance is guarded with survival rates ranging between 60% and 69% in two clinical reports.2, 11 Nevertheless, survival and successful careers as racehorses have been reported even in severely affected animals.2
References 1. Bernard GR, Artigas A, Brigham KL, Carlet J, Falke K, Hudson L, Lamy M, LeGall JR, Morris A, Spragg R.
2.
3.
4.
5. 6.
7.
8.
9.
10.
11.
12.
13. 14.
15.
16.
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The American-European Consensus Conference on ARDS. Definitions, mechanisms, relevant outcomes, and clinical trial coordination. Am J Respir Crit Care Med 1994;149: 818–24. Dunkel B, Dolente B, Boston RC. Acute lung injury/acute respiratory distress syndrome in 15 foals. Equine Vet J 2005;37:435–40. Wilkins PA, Otto CM, Dunkel B, Bedenice D, Paradis MR, Staffieri F, Baumgardner JE, Syring RS, Slack J, Grasso S, Pranzo G. Acute lung injury (ALI) and acute respiratory distress syndromes (ARDS) in veterinary medicine: consensus definitions. J Vet Emerg Crit Care 2007;17:333–9. Dunkel B, Wilkins PA. Acute respiratory distress syndrome. In: Smith BP (ed.) Large Animal Internal Medicine. Philadelphia: Mosby Inc., 2008; pp. 536–8. Piantadosi CA, Schwartz DA. The acute respiratory distress syndrome. Ann Intern Med 2004;141:460–70. Giannoudis PV. Current concepts of the inflammatory response after major trauma: an update. Injury 2003; 34:397–404. Matthay MA, Robriquet L, Fang X. Alveolar epithelium: role in lung fluid balance and acute lung injury. Proc Am Thorac Soc 2005;2:206–13. Jain R, DalNogare A. Pharmacological therapy for acute respiratory distress syndrome. Mayo Clin Proc 2006; 81:205–12. Buergelt CD, Hines SA, Cantor G, Cantor G, Stirk A, Wilson JH. A retrospective study of proliferative interstitial lung disease of horses in Florida. Vet Pathol 1986;23: 750–6. Prescott JF, Wilcock BP, Carman PS, Hoffman AM. Sporadic, severe bronchointerstitial pneumonia of foals. Can Vet J 1991;32:421–425. Lakritz J, Wilson WD, Berry CR, Schrenzel MD, Carlson GP, Madigan JE. Bronchointerstitial pneumonia and respiratory distress in young horses: clinical, clinicopathologic, radiographic, and pathological findings in 23 cases (1984–1989). J Vet Intern Med 1993;7:277–88. Peek SF, Landolt G, Karasin AI, Slack JA, Steinberg H, Semrad SD, Olsen CW. Acute respiratory distress syndrome and fatal interstitial pneumonia associated with equine influenza in a neonatal foal. J Vet Intern Med 2004;18: 132–4. Mols G, Priebe HJ, Guttmann J. Alveolar recruitment in acute lung injury. Br J Anaesthesiol 2006;96:156–66. Hoffman AM, Viel L. A percutaneous transtracheal catheter system for improved oxygenation in foals with respiratory distress. Equine Vet J 1992;24:239–41. Lee HS, Lee JM, Kim MS, Kim HY, Hwangbo B, Zo JI. Low-dose steroid therapy at an early phase of postoperative acute respiratory distress syndrome. Ann Thorac Surg 2005;79:405–10. McAuley DF, Frank JA, Fang X, Matthay MA. Clinically relevant concentrations of beta2-adrenergic agonists stimulate maximal cyclic adenosine monophosphate-dependent airspace fluid clearance and decrease pulmonary edema in experimental acid-induced lung injury. Crit Care Med 2004;32:1470–6.
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Chapter 63
Respiratory Diseases Specific to the Neonate Guy D. Lester
Bacterial pneumonia Bacterial infection of the newborn lung occurs most commonly during the early neonatal period from either direct aspiration or inhalation of bacteria or from the hematogenous spread of organisms in foals that are bacteremic. Infection can also become established before parturition, through aspiration of infected amniotic fluid or hematogenous spread via the umbilical cord. This may take place in mares with bacterial placentitis or amnionitis. The most common bacterial organisms that have been associated with pulmonary disease in foals are identical to those that cause systemic sepsis and include Escherichia coli, Klebsiella pneumoniae, Pasturella spp, Actinobacillus spp and Streptococci spp.1 Less common isolates include, but are not limited to, Salmonella spp, Enterobacter spp, Pseudomonas spp, Serratia marcesans, Staphylococcus spp, and Yersinia pseudotuberculosis. Streptococcal and Actinobacillus species appear to be particularly common in foals infected in utero. Diagnosis The definitive diagnosis of bacteria pneumonia can be difficult, and treatment is usually based on clinical suspicion rather than direct evidence. Signs of lower respiratory tract infection can be vague and features common to adults, such as nasal discharge and cough, may be absent in the early stages of infection, particularly during the early neonatal period. Signs could be limited to restlessness or tachypnea. The combination of any signs of respiratory tract disease together with laboratory evidence of sepsis (e.g., leukopenia, leukocytosis, neutrophil toxicity, an increase in band neutrophils, or hyperfibrinogenemia) suggests that bacterial bronchopneumonia is possible. Ultrasonography can be useful in identifying
lung pathology but, as with radiography, the differentiation of pneumonia from simple atelectasis can be difficult. The radiographic distribution of lung disease can provide some insight into cause and progression. Foals that aspirate contaminated amniotic fluid in utero often develop a focal interstitial pattern in the dorsocaudal lung fields, presumably because the aspiration occurred when the foal was in dorsal recumbency.2 Pneumonia acquired postnatally may be diffuse or focal, but over time most will tend to ‘settle’ caudoventrally. There is also anecdotal evidence that radiographic progression, for better or worse, will lag clinical change. The identification of causative organisms should be an important component of the diagnostic workup. Isolation of bacteria can be attempted from blood culture, or culture of amniotic fluid or placental tissue if in utero infection is suspected. Lower airway culture can be difficult, as a transtracheal aspirate can be dangerous in a compromised neonate. An alternative method involves passage of a guarded swab through a nasotracheal tube into the lower airway. The tip of the nasotracheal tube can also be cultured after removal. Therapy The treatment of bacterial lung disease involves a combination of respiratory support techniques and antimicrobial therapy. The neonatal foal readily develops dependent atelectasis in lateral recumbency. Consequently, positioning in sternal rather than lateral recumbency results in improved ventilatory capacity and higher arterial oxygen tension. Sternal recumbency is an unnatural position for the foal and maintenance is aided by use of a V-pad (Dandy Products Inc, Goshen, OH; Shank’s Veterinary Equipment, Milledgeville, IL).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Respiratory Diseases Specific to the Neonate
Figure 63.1 Delivery of humidified oxygen through nasal insufflation tube taped to a non-sterile tongue depressor which is taped to the foal’s muzzle.
Accurate assessment of pulmonary function is through arterial blood gas analysis. The most common blood gas derangement associated with bacterial pneumonia is mild to moderate hypoxemia with normal or reduced Pa co2 . In this situation, humidified oxygen is the treatment of choice and is best delivered through a nasal insufflation tube. The aim is to place the tube tip into the nasal passage to a point around the level of the medial canthus. A good method to ensure consistent placement is to form a Ushape with the insufflation tube and tape it to a nonsterile tongue depressor, leaving the end 10 cm free. The free end is inserted into the nasal passage and the tongue depressor is taped to the muzzle (Figure 63.1). A good starting point for most foals is to deliver humidified oxygen at a flow rate of 5 liters per minute. Rates of up to 10 liters per minute may be required and probably translate into a fraction of inspired oxygen (Fi o2 ) of around 1.0. The response to therapy needs to be carefully evaluated. Ideally, arterial blood gas (ABG) analysis is obtained to confirm an appropriate rise in Pa o2 . The aim should be to maintain the arterial oxygen concentration at around 70 to 105 mmHg. If an ABG analysis is not available then clinical improvement can be confirmed by noting a reduction in rate and or depth of ventilation, improvement in mucous membrane color, reduced agitation, etc. It is possible to maintain oxygen therapy in ambulatory foals by attaching the gas line to a current or old coiled fluid line. The nasal cannula should be cleaned twice daily and the oxygen lines and humidifier should be replaced every 1 to 2 days. In contrast to neonatal foals with mild respiratory disease, where arterial CO2 concentrations are typically normal or low, animals with moderate to severe
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bacterial pneumonia often have ABG changes consistent with hypercapnic respiratory failure, ie., reduced oxygen and elevated CO2 concentrations. This produces a primary respiratory acidosis. If the respiratory acidosis is sustained, the effect of CO2 as a respiratory stimulant may be lost, with respiratory drive controlled by peripheral chemoreceptors in response to hypoxemia. Consequently, oxygen therapy may result in an acute elevation in Pa co2 and a further reduction in pH due to a loss of hypoxic drive and hypoventilation. The management of hypercapnic respiratory failure is through assisted ventilation. Mechanical or assisted ventilation is indicated during resuscitation after birth or if the arterial concentration of CO2 exceeds 70 mmHg. The resultant acidemia is potentially life-threatening, and prolonged high levels of CO2 can result in generalized central nervous system depression and circulatory failure. Adult human resuscitation bags are effective in providing temporary intermittent positive airway pressure. Bags with adjustable positive end-expiratory pressure (PEEP) valves are recommended, and can be used with room air or can be attached to a humidifed oxygen source. An oxygen demand valve (Hudson, Temecula, CA) can assist spontaneous breathing in intubated foals. Negative airway pressure created by the foal during spontaneous ventilation will cause the valve to open and supplement the breath with oxygen-enriched gas at an increased tidal volume. The ideal method of reversing hypercapnic respiratory failure is through mechanical ventilation3 (see Chapter 22). The most important aspect of treatment for bacterial pneumonia is the correct selection of an antimicrobial agent. In an ideal clinical scenario, drug selection is based on recovery of a specific pathogen with a comprehensive antimicrobial sensitivity screen. Broad-spectrum therapy is indicated initially and, although the combination of a penicillin and aminoglycoside provide such a spectrum, third- or fourthgeneration cephalosporins have better penetration and activity in many cases of pneumonia. Examples of third-generation cephalosporins used in foals include ceftiofur, ceftazidime, cefotaxime, and cefpodoxime, the latter of which can be administered orally to neonatal foals (see Chapter 27).
Viral pneumonia Several equine viruses have been identified as causes of pneumonia in the neonatal foal. These include equine herpesvirus type 1 (EHV-1) and type 4 (EHV4), equine influenza, equine viral arteritis virus, and adenovirus. Of these, EHV-1 appears to be the most common.
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Herpesviral pneumonia is frequently fatal, even in the face of aggressive supportive therapies such as mechanical ventilation. One difficulty is developing a high index of suspicion of viral disease early in the disease course. Several factors appear common to EHV-1 infected foals, but none should be considered pathognomonic. Affected foals are typically delivered early without warning. The placenta is usually grossly normal. Clinically foals show signs of cardiovascular insufficiency along with respiratory difficulty. Mucous membranes are icteric and moderately to severely congested. Laboratory findings include leukopenia (often severe) with neutropenia, lymphopenia, and neutrophil toxicity. There is depletion of the myeloid cell lines on cytological examination of bone marrow aspirates. Dilation of retinal vessels and a red discoloration to the optic disc on fundic examination may also be present. An antemortem diagnosis of EHV-1 pneumonia could be achieved through submission of a nasal swab or whole blood sample for PCR detection of virus. The anti-viral drug, acyclovir, has been used, but efficacy data are lacking in foals with significant pneumonia.4 A greater bioavailability of the acyclovir pro-drug, valacyclovir, in adult horses would suggest that this drug may have enhanced efficacy in EHV-1 infected foals.5 Infection with adenovirus can be a problem in any immunocompromised foal, especially Arabian foals with combined immunodeficiency syndrome. Equine influenza virus (EIV) was associated with significant neonatal mortality in a recent outbreak in a naive population of horses in Australia.6 Foals commonly presented in acute respiratory distress or developed fulminant bronchointerstitial pneumonia while recovering from other ailments. Death occurred between 2 and 12 days of age. Histopathological findings were similar to those reported in older suckling foals or weanlings with bronchointerstitial lung disease.7 In those foals a clear etiology was not determined but a viral cause was postulated. An antemortem diagnosis of EIV is through PCR analysis of a nasal swab. Infection of foals with EIV is rare if there has been adequate colostral transfer from recently vaccinated or previously exposed mares. Equine viral arteritis is another cause of abortion and perinatal mortality. The virus causes a systemic fibrinoid vasculitis and progressive severe respiratory failure.8
Fungal pneumonia Fungal pneumonia is very rare in neonatal foals. Several different fungi have been associated with neonatal pneumonia. In utero infection with Histo-
plasma capsulatum can result in placentitis, abortion, or birth of an infected foal with multiple organ disease, including granulamotous pneumonia.9 An antemortem diagnosis can be difficult to establish, but is aided by tracheal aspiration or bronchoalveolar lavage where characteristic yeast-like organisms (3–5 µm in diameter) can be seen within macrophages. In addition, neonatal and maternal serum should be positive for anti-Histoplasma antibodies using an agar gel immunodiffusion test. The disease has been successfully treated in adults using amphotericin B, but reports of neonatal survival are lacking. Infection with Candida species (especially Candida albicans) is an infrequent complication in foals with chronic bacterial infection. Lengthy antimicrobial use is a risk factor for infection, and many cases begin with oral candidiasis. The diagnosis is based on a history that often includes persistent low-grade fever, worsening respiratory disease or the development of synovitis, and isolation of the organism through blood culture. Successful treatment of neonatal candidiasis has been achieved with ketoconazole or fluconazole.10 Newer antifungal agents are also likely effective, but data are lacking.
Meconium aspiration Any pre- or intrapartum stressful event to the foal can cause defecation of meconium into the amniotic fluid. Subsequent fetal breathing or gasping movements can lead to aspiration of meconium into the airways (see Chapters 64 and 65). The condition is recognized by the presence of yellow-orange staining of the amniotic fluid, the foal, or more accurately, by the observation of contaminated fluid within the nasal passages. Meconium contamination predisposes the foal not only to respiratory problems but also to other problems as a consequence of presumed asphyxia. Such an observation should be a warning of possible complications, including neurological dysfunction (perinatal apshyxia) and respiratory disease. Meconium is a sterile concretion of sloughed intestinal cells and mucus, and when aspirated may obstruct airways, interfere with gas exchange, or act as a nidus for bacterial infection. The diagnosis of meconium aspiration is based on direct observation of meconium staining of amniotic fluid, of the foal itself, or the presence of a brown fluid within the nasal passages. Radiographically, meconium aspiration may produce a focal increase in interstitial density. The site of the infiltrate can be variable, and can be located dorsally if aspiration occurred with the foal in a dorsopubic position within the mare. The degree of respiratory compromise or distress is clearly related to the volume of meconium
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Respiratory Diseases Specific to the Neonate aspirated, and whether or not secondary bacterial infection has occurred. The treatment of meconium aspiration should include clearance of the nasal passages and suctioning of the trachea shortly after birth. This can be achieved through passage of a soft, sterile catheter into the airway and gentle aspiration with a 60 mL dose syringe or connection to a portable or fixed suction device. Nasotracheal intubation will usually be necessary for effective suctioning of the lower airway. Care must be taken to avoid excessive or prolonged negative pressure during this procedure. Many foals with meconium aspiration require minimal treatment or support, but it is recommended to treat all foals with known or suspected meconium aspiration with a course of broad-spectrum antibiotics. If tachypnea or respiratory distress occur, then additional therapy with intranasal oxygen or assisted ventilation may be required. The prognosis for uncomplicated meconium aspiration is generally good, but close monitoring is indicated for affected foals.
Milk aspiration Aspiration of milk into the lower airway may occur as a complication of a wide range of conditions (see Chapter 64). Most foals that aspirate milk will also demonstrate nasal regurgitation of milk. Unfortunately, the decreased sensitivity of the upper and lower airway to foreign material may make diagnosis of milk aspiration difficult. Aspiration can occur in foals with cleft palate, persistent dorsal displacement of the soft palate (DDSP), epiglottic cysts, white muscle disease, botulism, perinatal asphyxia, or generalized weakness due to sepsis or prematurity. Iatrogenic contamination of the airway can occur when bottle-feeding is forced or if the foal is too weak or sleepy to receive feeding. Substantial and often fatal pneumonia can result from inappropriate placement of a nasogastric tube. The diagnosis of milk aspiration is supported by historical data (nasal regurgitation of milk), physical examination findings (abnormal lung and tracheal sounds), and laboratory data (inflammatory leukogram, elevated fibrinogen, hypoxemia). Radiographic examination commonly reveals a heavy, perihilar and/or ventrally located interstitial density, with or without air bronchograms. Treatment for the pneumonia involves long-term, broad-spectrum antimicrobial therapy and prevention of further contamination of the airway. The underlying cause should be pursued diagnostically and treated. This may necessitate the use of further diagnostics tests, including endoscopy and plain and contrast radiography. Enteral feeding through a naso-
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gastric or esophagostomy tube is indicated until the underlying problem has been resolved. Persistent or intermittent DDSP in the neonate will frequently resolve over time, but may take weeks to months.11 A staphylectomy has coincided with successful resolution of DDSP in young foals.12 A lack of improvement after resolution of the underlying problem should prompt re-consideration of the antibiotic choices, noting that anaerobic coverage may be needed in some cases.
The immature lung Most premature or dysmature foals will experience a degree of respiratory insufficiency, the severity of which is influenced by maturation of the surfactant system and structural development of the lung. Lung surfactant is produced from type II alveolar epithelial cells and is composed of phospholipids, inert lipids, and protein. It prevents spontaneous collapse of the alveoli by reducing alveolar surface tension. Surfactant activity can be detected in the developing fetus by 100 days but is not fully developed until 300 days, or in some foals, not until after birth.13 Deficiency of surfactant, superimposed on a structurally immature lung, has been implicated as the major contributing factor to the development of neonatal acute respiratory distress syndrome. A highly proteinaceous fluid is leaked into the interstitium and alveoli, producing a characteristic radiographic alveolar pattern. Although relatively uncommon in premature foals, affected animals develop severe respiratory acidosis with secondary, hypoxia-induced pulmonary arterial hypertension, reduced cardiac output, and tissue hypoxia. The prognosis for ARDS-affected foals remains poor, irrespective of the level of support available. The administration of bovine surfactant to premature foals at risk for ARDS has produced inconsistent results. Most premature and dysmature foals develop a clinically less severe form of lung dysfunction than ARDS, characterized by reduced ventilation capacity, tachypnea, hypoxemia, and progressive hypercapnia. Radiographically, there is a diffuse increase in interstitial density without alveolar changes. Atelectasis, with or without inflammatory cell infiltrates, is observed histologically, but hyaline membrane formation is rare. Treatment is based on severity of changes, but frequently involves intranasal administration of humidified oxygen and occasionally a respiratory stimulant, such as doxapram or caffeine. Foals should receive broad-spectrum antibiotic therapy. Fetal lung maturation is accelerated in foals exposed to chronic in utero stress, as is seen with bacterial placentitis.
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Persistent pulmonary hypertension Pulmonary hypertension in the newborn foal is most commonly associated with pneumonia or ARDS. There are other circumstances where pulmonary hypertension may exist in the absence of pulmonary disease. Persistent pulmonary hypertension (PPH), or persistent fetal circulation, has been suspected in foals with systemic sepsis and after perinatal asphyxia. The condition may develop in response to hypoxemia or acidosis.14 PPH should be suspected in foals with hypoxemia without evidence of significant pulmonary or cardiac disease. In some situations it is possible to invoke right-to-left shunting across the ductus arteriosus, with resultant caudal hypoxemia. Supplementation with intranasal oxygen will often produce suboptimal increases in Pa o2 . Treatment is difficult, but should be aimed at treating underlying problems such as acidosis, and providing ventilatory support, often utilizing 100% oxygen. Inhaled nitric oxide is the therapeutic agent most likely to reverse PPH, and limited clinical data supports its efficacy. Unfortunately, the optimal delivery of nitric oxide requires connecting the patient to a ventilator circuit (see Chapter 65).
Idiopathic or transient tachypnea Idiopathic or transient tachynpea is a syndrome observed most frequently in Clydesdale foals, but also reported in other breeds, including Thoroughbreds and Arabians. The condition is usually seen in association with warm, humid conditions and is probably related to either immaturity or dysfunction of thermoregulatory mechanisms. Clinically affected animals usually have an elevated respiratory rate and rectal temperature. The signs develop within a few days of birth and will usually spontaneously resolve within days to weeks. A complete blood count, fibrinogen estimation, auscultation of the thorax, arterial blood gas analysis, and thoracic radiography should be close to or within normal limits. Treatment with antipyretics such as flunixin are usually of little help, but substantial improvement can occur with body clipping, alcohol baths, and placement into a cooler environment. Given the difficulty in differentiating idiopathic tachypnea from infectious causes, it is usually recommended to place these foals on a course of broad-spectrum antibiotics.
Fractured ribs Thoracic trauma occurs commonly during the birthing process, and can result in rib shaft fracture or, more commonly, costochondral dislocation. Potential clinical problems include local pain and hematoma formation, hemothorax, lung lacera-
tion and pneumothorax, and pericardial/myocardial puncture. Diagnosis is by observation of thoracic wall symmetry and synchrony, palpation, ultrasonography findings, and/or radiographic examination. Ultrasonographic examination is superior to radiography in detecting fractures or dislocation.15 Treatment involves recognition and treatment of potential complications and avoidance of further thoracic trauma.
Pneumothorax Pneumothorax is rare in the young foal, but can develop after extrathoracic trauma, as a complication to over-exuberant assisted ventilation, or rarely, secondary to necrotizing bronchopneumonia. The condition should be suspected in any foal with sudden deterioration in lung function during assisted ventilation. Conservative treatment is indicated in mild cases and involves placement on a high inspired oxygen concentration for several hours. Moderate –to severe cases require placement of thoracic drains. Pneumothorax, pneumomediastinum, and subcutaneous emphysema are seen in neonates either spontaneously or in foals with a single lung bulla or multiple bullae.16 Some foals will present for body emphysema and have little or no respiratory insufficiency.
Pleural effusion Accumulation of large volumes of fluid in the peritoneal cavity, as is seen in advanced cases of uroperitoneum, will compromise respiratory function. In addition to atelectasis, there is frequently an accumulation of pleural fluid. Resolution of the primary abdominal problem has an immediate positive effect on lung function. Primary infectious pleuritis or pleural effusion secondary to lung disease is rare in the neonate.
References 1. Sanchez LC, Giguere S, Lester GD. Factors associated with survival of neonatal foals with bacteremia and racing performance of surviving Thoroughbreds: 423 cases (1982–2007). J Am Vet Med Assoc 2008;233(9):1446–52. 2. Lester GD, Lester NV. Abdominal and thoracic radiography in the neonate. Vet Clin North Am Equine Pract 2001;17(1):19–46. 3. Palmer JE. Ventilatory support of the critically ill foal. Vet Clin North Am Equine Pract 2005;21(2):457–86. 4. Murray MJ, del Piero F, Jeffrey SC, Davis MS, Furr MO, Dubovi EJ, Mayo JA. 1998. Neonatal equine herpesvirus type 1 infection on a thoroughbred breeding farm. J Vet Intern Med 1998;12(1):36–41. 5. Maxwell LK, Bentz BG, Bourne DW, Erkert RS. Pharmacokinetics of valacyclovir in the adult horse. J Vet Pharmacol Ther 2008;31(4):312–20.
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Respiratory Diseases Specific to the Neonate 6. Patterson-Kane JC, Carrick JB, Axon JE, Wilkie I, Begg AP. The pathology of bronchointerstitial pneumonia in young foals associated with the first outbreak of equine influenza in Australia. Equine Vet J 2008;40(3):199–203. 7. Lakritz J, Wilson WD, Berry CR, Schrenzel MD, Carlson GP, Madigan JE. Bronchointerstitial pneumonia and respiratory distress in young horses: clinical, clinicopathologic, radiographic, and pathological findings in 23 cases (1984–1989). J Vet Intern Med 1993;7(5):277–88. 8. Szeredi L, Hornyak A, Denes B, Rusvai M. Equine viral arteritis in a newborn foal: parallel detection of the virus by immunohistochemistry, polymerase chain reaction and virus isolation. J Vet Med B Infect Dis Vet Public Health 2003;50(6):270–4. 9. Rezabek GB, Donahue JM, Giles RC, Petrites-Murphy MB, Poonacha KB, Rooney JR, Smith BJ, Swerczek TW, Tramontin RR. Histoplasmosis in horses. J Comp Pathol 1993;109(1):47–55. 10. Reilly LK, Palmer JE. Systemic candidiasis in four foals. J Am Vet Med Assoc 1994;205(3):464–6.
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11. Altmaier K, Morris EA. Dorsal displacement of the soft palate in neonatal foals. Equine Vet J 1993;25(4):329– 32. 12. Shappell KK, Caron JP, Stick JA, Parks AJ. Staphylectomy for treatment of dorsal displacement of the soft palate in two foals. J Am Vet Med Assoc 1989;195(10):1395– 8. 13. Pattle RE, Rossdale PD, Schock C, Creasey JM. The development of the lung and its surfactant in the foal and in other species. J Reprod Fertil Suppl 1975;23: 6517. 14. Wilkins PA. Lower respiratory problems of the neonate. Vet Clin North Am Equine Pract 2003;19(1):19–33. 15. Jean D, Picandet V, Macieira S, Beauregard G, D’Anjou MA, Beauchamp G. Detection of rib trauma in newborn foals in an equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 2007;39(2):158–63. 16. Marble SL, Edens LM, Shiroma JT, Savage CJ. Subcutaneous emphysema in a neonatal foal. J Am Vet Med Assoc 1996;208(1):97–9.
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Chapter 64
Aspiration Pneumonia Mary Rose Paradis
Introduction Aspiration pneumonia in the neonatal foal can occur perinatally. Prepartum or intrapartum aspiration generally occurs when the foal experiences in utero stress such as a dystocia. This stress may result in hypoxia and meconium release into the amniotic fluid. If the foal is not delivered quickly then it may inhale contaminated amniotic fluid into its lungs as it gasps for breath. Meconium itself is a sterile material but large particles will cause airway obstruction, regional lung atelectasis, and chemical pneumonitis in neonates.1–3 The most common substance involved in postnatal aspiration in the foal is milk. Milk aspiration is usually associated with dysphagia, or difficulty in swallowing. Nursing in the foal normally begins with the foal wrapping its tongue around the mare’s teat, compressing it against the hard palate creating a negative pressure to draw milk into the mouth. Swallowing begins with the milk bolus being transported through the oropharynx to the nasopharynx to the esophagus by a series of coordinated events.4 Disruption of these events leads to milk regurgitation through the nares and aspiration of milk down the trachea.
Clinical signs The clinical signs of meconium aspiration include meconium staining of the amniotic fluid and foal. This staining is generally brown to yellow in appearance, and particulate matter in the fluid may be visualized. The contaminated fluid can be seen coming from the oral and nasal cavities when the foal is placed in a head-down, elevated chest position and coupaged (gentle tapping of the chest with a cupped hand) to achieve drainage of amniotic fluid from the lungs. Signs of respiratory impairment may occur immediately after birth. These signs will include an increased respiratory rate and effort. It is sometimes
difficult to distinguish from the initial breathing pattern of a normal newborn, but with meconium aspiration the increased effort persists beyond the initial few minutes after birth and usually becomes progressively worse. The clinical signs of milk aspiration may or may not be directly apparent at the first nursing bout. This depends on the etiology of the dysphagia and the amount of milk that is aspirated. Dysphagia in the newborn foal manifests outwardly as milk regurgitation from the foal’s nares after nursing (Figure 64.1). A benign cause of milk in a foal’s nares post-nursing would be the result of milk streaming from the mare’s udder as the foal nurses. External splashing of milk onto the nares needs to be differentiated from actual regurgitation of milk from the throat to the nares. Milk regurgitation should not be ignored because it is one of the signs that the foal is having trouble swallowing and that aspiration of milk into the lung may be occurring at the same time. Milk regurgitation must also be differentiated from milk reflux. The timing of the appearance of milk at the nose may be helpful in making this distinction. Regurgitation is a disruption of the swallowing mechanism and the milk appears immediately after nursing. Milk reflux is generally thought to be associated with an esophageal dysfunction such as megaesophagus. This may not be evident at nursing but is apparent when the foal lowers its head. The actual aspiration of milk secondary to dysphagia results in pneumonia. The severity of the pneumonia is dependent on the amount of milk that is aspirated, the period of time over which it has been aspirated, and the age and immune status of the foal. In the beginning no changes will be noted in auscultation or respiratory rate or effort. As the problem progresses, wheezes and crackles will be heard on auscultation of the foal’s lungs. A distinct rattle in the trachea from the aspirated milk can be felt after the foal suckles. Respiratory rate and
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Figure 64.1 Milk reflux from the nares of a foal with dysphagia.
effort will increase as the condition worsens. Because milk is not sterile, foals may develop an elevated temperature due to bacterial growth in the lungs. Depending on the cause of the dysphagia, foals may present with stridor as well. If upper airway obstruction occurs as part of the dysphagia, then there may be an increase in upper airway noise and inspiratory effort. Severe stridor may warrant an emergency tracheotomy.
Differential diagnosis for dysphagia Dorsal displacement of the soft palate (DDSP) is the most reported condition that results in dysphagia and aspiration pneumonia in the newborn foal.4–9 Persistent displacement of the soft palate results in failure of the soft palate to form a seal with the dorsal pharyngeal wall preventing milk regurgitation, and in failure of the epiglottis to prevent the bolus from entering the larynx. Dorsal displacement of the soft palate can be seen alone or in combination with other pharyngeal/laryngeal abnormalities such as rostral displacement of the palatopharyngeal arch, redundancy of the soft palate, or persistent epiglottal frenulum4–9 (see also Chapter 67). Generalized conditions such as prematurity, perinatal asphyxia, and nutritional myodegeneration (vitamin E/selenium deficiency) may also play a role in the development of DDSP. Approximately half the foals with nutritional myodegeneration have dysphagia as one of their clinical signs.10, 11 Poor pharyngeal muscle tone may lead to pharyngeal collapse, upper airway obstruction, stridor, and respiratory distress. Other causes for impaired nursing or swallowing in foals may include cleft palate, the presence of subepiglottic or dorsal pharyngeal cysts, bilateral laryngeal paralysis, and arytenoid chondritis.4, 9,12–14 Cleft palates in the foal are a fairly rare occurrence.
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Figure 64.2 Endoscopic view of a complete cleft hard and soft palate. Note base of tongue is visible from nasal cavity.
The heritability of this defect in horses is unknown. Defects can occur in both the hard and soft palate (Figure 64.2). They prevent the development of negative pressure in the oral cavity during nursing and allow milk to flow directly from the mouth to the nasal passage. Subepiglottic and dorsal pharyngeal cysts are also not a common problem in foals. Respiratory distress may be the first clinical sign that is noted due to upper airway obstruction.13, 14 The presence of a cyst in the pharynx disrupts proper swallowing and results in both milk regurgitation and aspiration pneumonia.
Diagnostic tests Diagnostic tests should be directed at examining both the cause of the dysphagia and the result of the dysfunction, which is aspiration pneumonia. The upper airway abnormalities are best visualized with endoscopy. A 1-meter, 1–1.5-cm diameter endoscope can be passed in most foals weighing over 40 kg. It is inadvisable to use sedation when examining the upper airway of the affected foal because some sedatives may relax the pharyngeal walls giving a false perception of the degree of pharyngeal collapse or soft palate displacement. In addition, tranquilization may lead to a sudden narrowing of the pharynx or larynx, precipitating acute asphyxia. Most foals will tolerate upper airway endoscopy with manual restraint.9 Dorsal displacement of the soft palate is readily apparent on endoscopic examination if there are no other abnormalities. In the normal foal the triangularly shaped epiglottis should be easily seen. In DDSP the epiglottis is hidden and you can only visualize the caudal edge of the soft palate as it encircles the arytenoid cartilages (Figure 64.3). If rostral displacement of the palatopharyngeal arch
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Figure 64.3 Endoscopic examination of the upper airway of a foal with DDSP. Note the absence of the epiglottis.
Figure 64.5 Dorsal pharyngeal cyst in a foal with milk aspiration. Right guttural pouch opening on left.
and collapse of the pharyngeal walls is present then it may be difficult to find the larynx. In very small foals, the presence of the endoscope in the nares may cause enough obstruction and negative pressure to again falsely elevate the soft palate. In these cases or in foals that will not stand unsedated for an endoscopic procedure, high detailed radiography of the lateral pharyngeal area may allow you to visualize a displaced soft palate9 (Figure 64.4). Endoscopy will help to illustrate congenital abnormalities of the upper airway.9 Cleft palates will be seen as defects in either the hard or soft palate. Small clefts in the caudal soft palate may be missed on endoscopy if obscured by the epiglottis. These are best found orally using a long-bladed laryngoscope inserted over the tongue. Large clefts in the hard palate may be visualized by a simple oral examination and can be confusing to the inexperienced examiner when viewed by an endoscope via the nasal passage.
Dorsal pharyngeal cysts are fluid-filled masses viewed endoscopically that may fill the field of view and obscure your other landmarks (Figure 64.5).9 Subepiglottic cysts arise from the ventral pharynx and may obscure or elevate the epiglottis. After examination of the pharyngeal region, the endoscope can be passed into the trachea. In foals with meconium aspiration, a brown-tinged fluid may be seen in the trachea. The presence of milk in the trachea is a positive indication of milk aspiration. Lateral radiographs of the foal’s chest are the best diagnostic for determining the extent of the aspiration pneumonia. A caudal ventral and dorsal granular infiltrate can be seen on chest radiographs of a foal with meconium aspiration. Foals with milk aspiration typically have a caudal ventral distribution of consolidation. The pattern can vary from a mild interstitial to heavy consolidation (Figure 64.6). A baseline complete blood count (CBC) and biochemical profile may be helpful in determining the degree of systemic involvement. An elevated or decreased white blood cell (WBC) count may
Figure 64.4 Lateral radiographic appearance of a dorsally displaced soft palate.Note the epiglottis is ventral to the edge of the soft palate. Reprinted with permission from Paradis.9 Copyright Elsevier 2006. Used with permission.
Figure 64.6 Lateral thoracic radiograph of a foal with aspiration pneumonia. Note area of consolidation caudal to heart.
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Aspiration Pneumonia indicate the presence of an infection and possibly sepsis. Significantly elevated creatinine kinase values are suggestive of nutritional myodegeneration.
Treatment and outcome Rapid recognition that there is meconium staining of a foal is the first step in treatment of potential meconium aspiration. If the foal is stained with meconium, the foal’s head should be lower than it body and the chest coupaged to expel as much meconium-contaminated fluid from the lungs and trachea as possible. The mouth, nares, and trachea should be suctioned to remove contaminated fluid as well. Supportive care and oxygen administration are the primary treatments if the foal has aspirated meconium. If the aspiration is severe the treatment is very unrewarding. For foals that have milk aspiration, the first treatment involves cessation of nursing. The foal should be muzzled and fed through an indwelling nasogastric tube until the cause of the dysphagia is identified and resolved. The normal foal drinks between 25 and 35% of its bodyweight in milk on a daily basis. This is approximately 11–15 liters of milk per day for a 45-kg foal.15 Because we are unable practically to simulate the feeding schedule of the normal foal, which nurses five to seven times per hour, bolus feeding every 2 hours is recommended. One may not be able to deliver by bolus the volume of milk needed to make up 25% of its bodyweight. It is best to start bolus feeding at a lower percentage bodyweight, perhaps 10%, and build up to the higher amounts as the foal adjusts to this type of feeding. For example, a 45-kg foal may start with 4.5–5 L/day divided into 12 feedings of 375 mL each. The mare’s milk is the best nutrition for the foal and can be obtained by milking the mare at frequent intervals. Unfortunately mares will often decrease their milk production if the foal is not actually nursing. Mare’s milk can usually be supplemented with a commercial mare’s milk replacer. Because milk is not sterile, aspiration of milk results in a mixed bacterial pneumonia. A broadspectrum antibiotic coverage is recommended. Generally an aminoglycoside for Gram-negative coverage plus a beta-lactam or a cephalosporin for Gram-positive coverage is adequate. Specific therapy for the cause of the dysphagia needs to be addressed. In cases where the foal is experiencing severe upper airway obstruction, an emergency tracheotomy may be necessary. This would most often be seen in foals with pharyngeal cysts, arytenoiditis, bilateral paralysis of the arytenoids, and severe pharyngeal collapse.
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A temporary tracheotomy is performed in the middle third of the neck. A vertical incision is made in the skin over the trachea. Blunt dissection through the soft tissue will expose the trachea. A horizontal incision, approximately one-third the diameter of the trachea, is made between the tracheal rings. An appropriately sized human tracheotomy tube can then be inserted through the incision. The foal’s stridor should be immediately relieved if the problem is an upper airway obstruction. Care of the tracheotomy tube involves daily cleaning.9 Removal of pharyngeal cysts can be accomplished by a laryngotomy or by transendoscopic laser surgery. Once the tissue is removed the foals generally do well. Surgical repair of cleft palates has been attempted with limited success in foals. Bilateral paralyses of the larynx or bilateral arytenoiditis are uncommon in the foal. Surgical options are possible but have not been evaluated in the foal. The prognosis for these problems is poor. Specific treatment for DDSP depends on its etiology. If it is secondary to a persistent frenulum then surgical correction is warranted.8 If DDSP is secondary to nutritional myodegeneration, then administration of a vitamin E/selenium supplement is recommended.10, 11, 16 Many times no etiology for DDSP can be found. If the foal is restrained from nursing and placed on an effective antibiotic the clinical signs of pneumonia resolve fairly rapidly, usually within 5–7 days. Re-examination of the pharyngeal region of the foal, either endoscopically or radiographically, should be performed every 2–3 days to assess the resolution of the DDSP. Many cases will resolve spontaneously within a week. A few may take longer and may require weaning from the mare. These foals can often drink from a bucket or be switched to solid milk pellets without aspirating.
References 1. Tyler DC, Murphy J, Cheney FW. Mechanical and chemical damage to lung tissue caused by meconium aspiration. Pediatrics 1978;62:454. 2. Lopez A, Bildfell R. Pulmonary inflammation associated with aspirated meconium and epithelial cells in calves. Vet Pathol 1992;29:104–11. 3. Korhonen K, Soukka H, Halkola L, Peuravuori H, Aho H, Pulkki K, Kero P, Kaapa PO. Meconium induces only localized inflammatory lung injury in piglets. Pediatr Res 2003;54:192–7. 4. Barton MH. Nasal regurgitation of milk in foals. Compend Cont Educ Pract 1993;15:81–91. 5. Shappell KK, Caron JP, Stick JA, Parks AJ. Staphylectomy for treatment of dorsal displacement of the soft palate in two foals. J Am Med Vet Assoc 1989;195:1395–8.
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6. Koterba AM, Paradis MR. Specific respiratory conditions. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1991; pp. 177–199. 7. Altmaier K, Morris EA. Dorsal displacement of the soft palate in neonatal foals. Equine Vet J 1993;25:329–32. 8. Yarbtough TB, Voss E, Herrgesell EJ, Shaw M. Persistent frenulum of the epiglottis in four foals. Vet Surg 1999;28:287–91. 9. Paradis MR. In: Paradis MR (ed.) Aspiration pneumonia secondary to dysphagia. Equine Neonatal Medicine: A CaseBased Approach. Philadelphia: Elsevier, 2006; pp. 148–56. 10. Dill SG, Rehbun WC. White muscle disease in foals. Compend Cont Educ Pract 1985;7:s627–s635.
11. Moore RM. Nutritional muscular dystrophy in foals. Compend Cont Educ Pract 1991;13:476–90. 12. Balstone JHF. Cleft palate in the horse. Br J Plast Surg 1966;19:327. 13. Stick JA, Boles C. Subepiglottic cyst in three foals. J Am Vet Med Assoc 1980;180:62–4. 14. Hardy J. Upper airway obstruction in foals, weanlings, and yearlings. Vet Clin North Am (Equine Pract) 1991;7: 105–22. 15. Paradis MR. Nutritional support: enteral and parenteral. Clin Tech Equine Pract 2003;2:87–95. 16. Valberg SJ. A review of the diagnosis and treatment of rhabdomyolysis in foals with severe rhabdomyolysis. AAEP Proc 2002;48:117–21.
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Chapter 65
Other Disorders of Breathing Pamela A. Wilkins
Persistent pulmonary hypertension of the newborn Persistent pulmonary hypertension (PPH) is also known as reversion to fetal circulation or persistent fetal circulation and its genesis lies in the failure of the fetus to successfully make the respiratory and cardiac transition to extrauterine life, or reversion of the newborn to fetal circulatory patterns in response to hypoxia and/or acidosis.1–4 Differentiating this problem from other causes of hypoxemia in the newborn, such as sepsis or bacterial or viral pneumonia, or congenital cardiac malformation, requires some investigation and multiple serial arterial blood gas analyses are necessary to confirm suspicion of this problem. However, the condition should be suspected in any neonate with hypercapnic hypoxemia that persists and worsens; these foals are in hypoxemic respiratory failure. Hypoxemia is generally severe (Pa o2 < 50 mmHg) while hypercapnia is more variable, but generally > 50 mmHg. The fetal circulatory pattern, with pulmonary hypertension and right-to-left shunting of blood through the patent foramen ovale and ductus arteriosus, is maintained in these cases. Treatment of PPH is two-pronged: abolishment of hypoxemia and correction of the acidosis, as both abnormalities only encourage the fetal circulatory pattern. Initial therapy is provision of intranasal oxygen (INO2 ) at 8–10 L/min. Some foals will respond to this therapy and establish neonatal circulatory patterns within a few hours. Failure to improve, or worsening of, hypoxemic respiratory failure following INO2 should prompt intubation and mechanical ventilation with 100% oxygen (Figure 65.1). This serves two purposes, one diagnostic and one therapeutic. Ventilation with 100% O2 may resolve PPH and, if intrapulmonary shunt and altered ventilation perfusion relationships are causing the hypoxemic respiratory
failure, arterial oxygen tension (Pa o2 ) should exceed 100 mmHg under these conditions. Failure to improve Pa o2 suggests PPH or large right-to-left extrapulmonary shunt due to congenital cardiac anomaly. Based on evidence presently available, it appears reasonable to use inhaled nitric oxide in an initial concentration of ∼ 20 ppm in the ventilatory gas for term and near-term foals with hypoxemic respiratory failure and PPH that fails to respond to mechanical ventilation using 100% oxygen alone.5, 6 Experience to date suggests clinical cases will require treatment for less than 4–6 hours, minimizing the potential for deleterious consequences, such as methemoglobinemia observed in human infants receiving prolonged (more than several days) NO treatment. Foals requiring more prolonged treatment should be re-evaluated for the presence of other contributing problems to persistent unresponsive hypoxemia. Further information may be found in Chapter 63.
Meconium aspiration syndrome Meconium aspiration is thought to occur secondary to acute pre- or intrapartum asphyxiation associated with severe fetal distress, subsequent passage of meconium by the fetus, and aspiration of meconium during fetal gasping efforts. Neonates with meconium aspiration syndrome (MAS) have marked surfactant dysfunction. Airways and alveoli of affected neonates contain meconium, inflammatory cells, inflammatory mediators, edema fluid, protein, and other debris. MAS can present clinically with different degrees of severity, ranging from a mild form of respiratory compromise to severe forms that may result in perinatal death despite respiratory management up to and including mechanical ventilation and extracorporeal membrane oxygenation (ECMO). Current treatments being investigated in the management of human infants with MAS include surfactant
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Figure 65.1 Foal on ventilator receiving 100% oxygen and nitric oxide as treatment for persistent pulmonary hypertension of the neonate (PPHN). Foal responded to treatment within a few hours and fully recovered.
administration, surfactant administration combined with partial liquid ventilation, and surfactant lavage.7–9 Refer to Chapters 63 and 64.
Rib fractures/thoracic trauma Rib fractures have been recognized in 3–5% of all neonatal foals and can be associated with respiratory distress. The incidence of rib fractures in foals presenting to a neonatal intensive care unit is much higher, with reports ranging up to 30%. Potential complications of rib fractures include fatal myocardial puncture, hemothorax, pneumothorax, and fatal hemorrhagic shock.10–12 Rib fractures are frequently found during physical examination by palpation of the ribs, palpation of crepitus associated with the ribs, or by auscultation over the fracture sites. Audible ‘clicking’ or grating sounds may be heard over fracture sites of affected ribs. Affected foals may exhibit painful responses when affected ribs are palpated or may have short shallow respirations consistent with thoracic ‘guarding’. Displacement of several ribs will result in deformation of the normal thoracic contour. Affected foals may exhibit painful respiration characterized by groaning or grunting. Diagnosis is most reliably confirmed by ultrasonographic evaluation.10, 13 Usually multiple ribs are affected on one side of the chest and occur in close proximity to the costochondral junction, usually 1–3 cm dorsal (Figure 65.2). Specific treatment is generally unnecessary for non-displaced or minimally displaced rib fractures, beyond stall confinement for a period of 2–3 weeks and care in handling, but direct pressure on the thorax should be avoided in all cases. Foals with thoracic trauma and/or rib fractures should be monitored carefully and frequently as acute displacement of fractured ribs may result in po-
Figure 65.2 Ultrasonographic image of rib fracture. Note fluid surrounding fracture and medial displacement of ventral rib fracture. This foal underwent surgical repair of multiple rib fractures and survived.
tentially fatal intrathoracic injury at any time. Treatment may need to be more aggressive in foals presenting to intensive care units with rib fractures as a part of their clinical picture and some specific patients may benefit from surgical stabilization of fractures, particularly those displaced fractures overlying the heart or resulting in intrathoracic injury14, 15 (Figure 65.3). More information may be found in Chapters 49 and 57.
Pneumothorax Pneumothorax can occur spontaneously or secondary to thoracic trauma or excessive positive pressure ventilation16 (Figure 65.4). It can also occur
Figure 65.3 Intraoperative image of rib fracture repair using the monofilament technique described by Kraus et al.15 This image shows monofilament being passed through needle inserted into the ventral fracture segment in preparation for stabilization. This foal underwent surgical repair of multiple rib fractures and survived. Image courtesy of Dr Joanne Hardy.
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(c) Figure 65.4 Thoracic radiograph obtained in standing foal with multiple rib fractures. Note cranial pneumomediastinum (heart pushed caudally) (a) and pneumothorax (b) outlining one dorsal lung lobe. Rib fractures are not readily apparent. (c) Thoracic radiograph obtained just prior to discharge. Note resolution of pneumomediastinum and pneumothorax and the presence of radiodense skin staple and clips securing monofilament repair of fracture site(s).
secondary to tracheostomy, surgery or trauma. Any foal being mechanically ventilated that suddenly presents with respiratory distress and hypoxemia should be evaluated for pneumothorax. Pneumothorax is suspected by auscultation and percussion of the thorax, but is best confirmed by radiographic evaluation of the thorax. Treatment is required in cases where clinical signs are moderate to severe and/or progressive and involves closed suction of the pleural space. Subcutaneous emphysema can complicate both diagnosis and treatment.
Idiopathic or transient tachypnea Idiopathic or transient tachypnea has been observed in Clydesdale, Thoroughbred and Arabian breed foals primarily. In human infants, transient tachypnea can be related to delayed absorption of fluid
from the lung, perhaps due to immature sodium channels.17 In foals, it is generally seen when environmental conditions are warm and humid and is thought to result from immature or dysfunctional thermoregulatory mechanisms. Clinical signs are primarily increased respiratory rate and rectal temperature. Signs develop within a few days of birth and may persist for several weeks. Treatment involves moving the foal to a cooler environment, body clipping, and provision of cool water or alcohol baths. These foals are frequently treated with broad-spectrum antimicrobial drugs until infectious pneumonia can be ruled out.
References 1. Murphy JD, Rabinovitch M, Goldstein JD, Reid LM. The structural basis of persistent pulmonary hypertension of the newborn infant. J Pediatr 1981;98(6):962–7.
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2. Wilkins PA. Lower respiratory problems of neonates. In: Parente EJ (ed.) Equine Respiratory Disease, Veterinary Clinics of North America, Equine Practice, 2003;19: 19–34. 3. Wilkins PA. Disorders of foals. In: Reed SM, Bayley WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis: Saunders, 2004; pp. 1381–1431. 4. Magdesian KG, Wilkins PA. Neonatal diseases. In: Orsini JA, Divers TJ (eds) Manual of Equine Emergencies, 3nd edn. St Louis: Elsevier/Saunders, 2008; pp. 523–31. 5. Lester GD, DeMarco VG, Norman WM. Effect of inhaled nitric oxide on experimentally induced pulmonary hypertension in neonatal foals. Am J Vet Res 1999;60(10): 1207–12. 6. Finer NN, Barrington KJ. Nitric oxide for respiratory failure in infants born at or near term (Cochrane Review). Cochrane Database Syst Rev 2001;4:CD000399. 7. Ghidini A, Spong CY. Severe meconium aspiration syndrome is not caused by aspiration of meconium. Am J Obstet Gynecol 2001;185(4):931–8. 8. Chappell SE, Wolfson MR, Shaffer TH. A comparison of surfactant delivery with conventional mechanical ventilation and partial liquid ventilation in meconium aspiration injury. Respir Med 2001;95(7):612–7. 9. Wiswell TE, Knight GR, Finer NN, Donn SM, Desai H, Walsh WF, Sekar KC, Bernstein G, Keszler M, Visser VE, Merritt TA, Mannino FL, Mastrioianni L, Marcy B, Revak SD, Tsai H, Cochrane CG. A multicenter, randomized, controlled trial comparing Surfaxin (Lucinactant) lavage with standard care for treatment of meconium aspiration syndrome. Pediatrics 2002;109(6):1081–7.
10. Jean D, Laverty S, Halley J, Hannigan D, L´eveill´e R. Thoracic trauma in newborn foals. Equine Vet J 1999;31(2):149– 52. 11. Sprayberry KA, Bain FT, Seahorn TL, Slovis NM, Byars TD. 56 cases of rib fractures in neonatal foals hospitalized in a referral center intensive care unit from 1997–2001. In: Proceedings of the American Association of Equine Practitioners, San Diego, 2001; pp. 395–9. 12. Schambourg MA, Laverty S, Mullim S, Fogarty UM, Halley J. Thoracic trauma in foals: post mortem findings. Equine Vet J 2003;35:78–81. 13. Jean D, Picandet V, Macieira S, Beauregard G, D’Anjou MA, Beauchamp G. Detection of rib trauma in newborn foals in an equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 2007;39(2):158–63. 14. Bellezzo F, Hunt RJ, Provost R, Bain FT, Kirker-Head C. Surgical repair of rib fractures in 14 neonatal foals: case selection, surgical technique and results. Equine Vet J 2004;36(7):557–62. 15. Kraus BM, Richardson DW, Sheridan G, Wilkins PA. Multiple rib fracture repair in a neonatal foal using nylon strand suture repair technique. Vet Surg 2005;34(4):399–404. 16. Marble SL, Edens LM, Shiroma JT, Savage CJ. Subcutaneous emphysema in a neonatal foal. J Am Vet Med Assoc 1996;208(1):97–9. 17. O’Brodovich HM. Immature epithelial Na+ channel expression is one of the pathogenetic mechanisms leading to human neonatal respiratory distress syndrome. In: Proceedings of the Association of American Physicians, 1996;108(5):345–55.
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Chapter 66
Bacterial and Viral Respiratory Disease of the Neonate ´ Renaud Leguillette
The lungs of newborn foals must go through a series of changes to ensure a successful transition from fetus to neonate during the birth process. Healthy foals are born hypoxemic and hypercapneic1–3 and adapt to extrauterine life during the first 4 days following delivery.4 Bacterial and viral respiratory diseases of the neonate can quickly compromise the gas exchange in the lungs.
Neonatal respiratory physiology Establishment of normal respiratory function in the neonatal foal can be affected by four critical events. First, the pulmonary hypertension in the fetus that limits blood flow to the lungs in utero (only 15% of the blood flow)5 must be abolished to permit pulmonary blood flow to increase to 100%. Lung inflation caused by the first gasping breaths (expiring against a closed larynx) as well as pulmonary artery vasodilation caused by the increasing Pa o2 at birth are the main factors responsible for a decrease in pulmonary hypertension and an increase in blood flow to the lungs. Any disease process that produces a decrease in Pa o2 , often accompanied by acidosis, can increase pulmonary hypertension and prompt a return to fetal circulation. Second, neonatal lung function is marginal at birth due to immature lung mechanics.6 Neonatal lungs are more elastic (i.e., decreased compliance) due in part to fluid and edema in the parenchyma, while the chest wall is very compliant.7 These dynamics result in increased work of breathing and decreased tidal volume.8 Third, surfactant production and maturation appears late in the gestation,9 which may further compromise respiratory function, especially in premature foals. Fourth, the ventilation–perfusion ratio is not optimum in foals as a result of right-to-left shunting, either intracardiac
or intrapulmonary, and may not respond to oxygen therapy.10, 11 Consequently, bacterial or viral respiratory diseases can easily overwhelm an already fragile cardiopulmonary system.
Epidemiology Bacterial pneumonia is often associated with septicemia in newborn foals. Pneumonia and septicemia are leading causes of morbidity and mortality in the equine neonate.12 A prospective study on 2468 foals found that 37% of the fatalities occurred during the first 7 days of age and 57% occurred during the first month. Although septicemia represented the leading cause of mortality, the morbidity for pneumonia was high for foals 0–31 days old.12 The prevalence of pneumonia in foals younger than 1 month old is probably much higher than reported because pneumonia may be underdiagnosed in practice, especially when foals are suffering from septicemia. The incidence of pneumonia in older foals can be as high as 82% on Thoroughbreds farms.13
Pathogenesis Bacterial and viral respiratory disease can be acquired in utero, during parturition, or postpartum. Pneumonia can be respiratory or hematogenous in origin. A retrospective study found that foals with Actinobacillus sp. septicemia were three times more likely to have pneumonia than foals with positive hemocultures due to other pathogens.14 Pneumonia in newborns is often secondary to hematogenous infections, probably because of the intense vascularization of the lung combined with slow blood flow in small-diameter capillaries. Another common cause of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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pneumonia in newborn foals is milk aspiration, often associated with bottle feeding. Since many pneumonic neonates are also septicemic, signs of respiratory disease may be overshadowed by systemic signs of sepsis. Inflammation and infection of the lung alter the gas exchange properties. Inflammation and exudates in the small airways can impede airflow to the alveoli and alter the function of the surfactant. Surfactant is a lipoprotein complex that covers the alveoli to reduce alveolar surface tension and to facilitate alveoli opening and lung expansion. Alterations in surfactant quality or quantity will increase respiratory work, alter gas exchange, and increase ventilation–perfusion mismatch.
Pathogens The bacterial pathogens most commonly associated with newborn foal pneumonia are usually the same isolates associated with septicemia. This assumption means that the causative agents for pneumonia in newborn foals are very different from those found in older foals and adult horses (see Chapters 76 and 78). Although neonatal septicemia is often associated with Gram-negative bacteria, retrospective studies found an increase in the prevalence of Gram-positive bacteria in newborn foals; up to 34% of bacteremic cases were found to have a single Gram-positive isolate.15, 16 . The incidence of viral infections in newborn foals is thought to be low compared to bacterial sepsis. The most common viral pathogens involved are equine herpesviruses (EHV-1 and EHV-4)17–19 and equine arteritis virus (EAV), all of which can cause respiratory disease as well as abortion.20, 21 Influenza has also been described in a few cases.22 Viral infections usually induce interstitial pneumonia, but bronchopneumonia is also possible. The role of EHV-2 is uncertain in foals’ respiratory diseases. Adenoviral infections in foals can occur in Arabians with combined immunodeficiency and are usually fatal.23
Physical examination Prior to physical examination of the sick newborn, a history of respiratory disease outbreaks on the farm and the medical history of the pregnancy and parturition should be obtained. The mare and her placenta should be examined for signs of pathology. If either the placenta or the foal shows meconium staining, the foal should be closely monitored for depression, decreased nursing, and septicemia. Foals should be assessed right after birth and re-evaluated once the effects of the postpartum catecholamine rush
have vanished. Catecholamines have a protective effect and may mask clinical signs associated with infection. Examination of the respiratory system includes the respiratory rate and its pattern when evaluated in a resting foal, the presence of abdominal effort, lung auscultation, and the palpation of the chest for the detection of broken ribs. Signs of pneumonia can be vague and difficult to identify;24 indeed, a retrospective study on bacteremic foals found that 30% of neonates were considered bright, and 51% were able to stand. Pneumonia was diagnosed in 28% of the foals with bacteremia.6 Foals affected with viral infection affecting the lower respiratory tract function show even more severe clinical signs and are often depressed and not able to nurse. The respiratory rate is normally elevated at birth and decreases quickly within the first few days of age. The respiratory rate of newborn foals varies between 53.9 ± 9.9 bpm at day 2 of age and 40.9 ± 5.2 bpm by 7 days of age.8 The breathing pattern may be irregular with short periods of apnea in premature foals and foals suffering from the maladjustment syndrome even in the absence of pneumonia. Mucous membrane color can be misleading and is an unreliable indicator of hypoxia in newborn foals. Very low blood oxygen content may be reached before the membranes are cyanotic, in part because of the presence of fetal hemoglobin. Chest palpation should be performed with the foal maintained in dorsal recumbency for the detection of rib fractures.25 In the newborn foal there is a poor correlation between lung auscultation and the presence and severity of pneumonia. Positioning of the foal just before and during the lung auscultation can affect the findings. It is not uncommon to detect crackles (rales) in the lower lung of a foal that has been placed in sternal recumbency after resting in lateral recumbency for a period of time. The presence of wheezes (rhonchi) and crackles may also be noted in a foal during resolution of pneumonia. Neonates with pneumonia do not necessarily have any fever, cough or nasal discharge. In one study only 5% of foals with pneumonia had pyrexia, and only 17% had tachypnea (> 40/min), although most of them were probably older than a month in this study.13 If the foal’s condition allows, lung auscultation should be repeated after brief exercise or the use of a rebreathing bag. The cranioventral lung field may be the first area to exhibit an absence of lung sounds. Importantly, the presence of tachypnea, abnormal respiratory sounds, fever, weakness, and milk reflux from the nares observed at the clinical examination are not correlated with the survival and outcome of the foal.26 Clinical signs of respiratory dysfunction do not permit differentiation between infectious and
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non-infectious etiologies nor between viral and bacterial pathogens. Viral pneumonia may be acquired in utero or postnatally, which causes abortion or interstitial pneumonia with lymphoid depletion. Premature delivery is usually found in mares that have been infected with EHV-1. Typical clinical signs observed in newborn foals affected by EHV-1 are usually obvious respiratory distress, icterus, and potentially petechiae and ecchymotic hemorrhages. Dilated retinal vessels and retinal hemorrhages are also often noted. These foals rarely survive and die shortly after birth. Foals infected with EAV present with severe respiratory distress of acute onset and almost always systematically die soon after the occurrence of the clinical signs.
Diagnosis Imaging:thoracic radiography Diagnostic imaging can be performed in the field in most cases. The advantage of radiography over chest ultrasonography is the ability to visualize the whole depth of the thorax while the ultrasonographic examination is more superficial.27, 28 Chest radiography is well suited for air/tissue contrast and the entire thorax of newborn foals can usually fit on one big cassette. Digital radiographs (DR) of the chest may be easier to take and to read but some DR cassettes, being smaller than the conventional or computed radiographs (CR) cassettes, it may sometimes be easier to use a bigger cassette in a conventional system if the clinician is familiar with the technique. It may be beneficial to perform both lateral and dorsoventral views if the foal’s condition will allow. Radiographic lesions are described by pattern and localization. Patterns can be normal, interstitial, alveolar (see Figure 66.1) or nodular, in the caudodorsal, caudoventral, hilar, or cranioventral anatomical regions of the chest. Severe lesions in the caudodorsal region are associated with a poorer prognosis for survival than in the other areas.24, 26 Radiographs are useful for re-evaluation of pneumonia; the cranioventral region is usually the last area to improve on the x-rays.24 Radiographs should be interpreted in the clinical context; for example, a study found no correlation between the presence of radiographic lesions in young foals and the presence of dyspnea; in addition, the severity of the radiographic changes did not correlate with abnormal respiratory sounds.29 It is important to note that the severity of the pneumonia in newborn foals, as observed on the radiographs, has little impact on long-term performance. A retrospective study found that the location and severity of the lesions observed on the lung radiographs of newborn
Figure 66.1 Lateral thoracic radiograph from a 10-day-old foal with pneumonia. The caudoventral and caudodorsal lung fields show a pronounced alveolar pattern with air bronchograms. The caudal silhouette of the heart and of the vena cava are completely obscured. Courtesy of Dr S. Giguere, University of Georgia.
foals with pneumonia (mean age at admission 3.7 days) had no correlation with the race performance.24 Imaging: thoracic ultrasonography Although ultrasounds do not penetrate the layer of air found in normal lungs, ultrasound imaging can be a useful diagnostic tool for pneumonia and for the detection of fractured ribs.25 In newborn foals pneumonia is usually diffuse, thus making it easier to detect with ultrasonography. Because of the shape of the chest, contact is better if a smaller convex probe is used. When pneumonia is present, the images show a rough and irregular lung parenchyma surface. Because lung atelectasis (when the parenchyma is not filled with air) is frequent in the ventral parts of the lungs, the ultrasound is able to penetrate deeper lung tissues. One can sometimes see areas with fluid containing hyperechogenic particles and fibrin in the most severe and chronic cases. The most severe lesions are usually observed in the ventral half of the chest. It is the author’s experience that newborn foals rarely present with pleuropneumonia and thus significant pleural fluid accumulation is rarely seen on the ultrasound. Hemothorax may occur secondary to fractured ribs.
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Isolation of pathogens
Blood culture Since pneumonia in newborn foals is often secondary to a systemic infection, blood culture(s) should be performed to identify the bacteria involved. The sampling technique should be performed in aseptic conditions and better results are usually obtained when using the blood culture vials recommended by the testing laboratory. Ideally, aerobic and anaerobic condition cultures should be performed. Repeated sampling and sampling during a fever peak will increase the chances of isolating the pathogen. With the exception of aspiration pneumonia, it can be assumed that the bacteria isolated from the blood culture are cause of the pneumonia. However, further studies are necessary to determine the correlation between the results from blood cultures and transtracheal wash in newborn foals with pneumonia.
Transtracheal wash In newborn foals with septicemia and systemic signs of disease, a transtracheal wash culture may not be more informative than a blood culture. However, in cases where the suspected etiology is milk aspiration or dysphagia, a transtracheal wash is probably more useful than a blood culture to identify the responsible pathogens. Although more often carried out in older foals and adult horses, a transtracheal wash can still be performed safely in newborn foals by taking a few precautions. The diameter of the trachea is small in young foals and the addition of a significant amount of fluid before sampling can increase tremendously the resistance to airflow in the trachea. Therefore, the smallest possible amount of saline (or no saline at all) should be used for the wash. We usually use ≤ 10 mL of saline in young foals, depending on the volume and viscosity of the secretions in the trachea as well as the size of the foal. The procedure should be postponed or cancelled if the foal is showing signs of respiratory distress. The procedure should be done in aseptic conditions (sterile gloves, skin shaved and aseptically prepared, sterile catheters, and sterile saline). The skin is clipped and shaved on an area approximately 5 × 5 cm at the junction between the upper and middle third of the trachea. Subcutaneous local anesthesia is then applied. If necessary and if the clinical condition of the foal allows, short-term sedation with i.v. diazepam can be used. While stabilizing the trachea with one hand, a vertical 2 cm skin incision is performed. The skin incision is not mandatory but does reduce the risk of subcutaneous infections and emphysema by allowing a better drainage of the transtracheal wound (which may become in-
Figure 66.2 Transtracheal wash in a 6-day-old foal using a long intravenous polypropylene catheter inserted percutaneously at a 45◦ angle down into the trachea. 5 mL of sterile saline will be injected in the trachea and aspirated back.
fected by the bacteria responsible for the pneumonia). A transtracheal wash can be performed using a double or triple sheath catheter (Mila International inc.), a long polypropylene catheter passed through an i.v. catheter (14G, 31/2 -in, Angiocath, Becton Dickinson, New Jersey) (Figure 66.2), or a long intravenous catheter (16G 24-in Intracath, Becton Dickinson, New Jersey). During insertion the catheter should be oriented down at a 45◦ angle (Figure 66.2). Once the catheter reaches the horizontal part of the trachea, saline can be slowly injected and immediately aspirated back (or secretions can be aspirated directly if no saline is used). For shipping, place the sample in Port-a-Cul culture media (Becton Dickinson, New Jersey Inc.) or any culture milieu recommended by the testing laboratory. The sample can also be Gramstained immediately on site to provide information on the bacteria involved.
Viruses Viral infections in foals are severe and often lead to a rapid demise. Isolation and culture of viruses is more difficult and time-consuming than for bacteria. However, PCR testing permits quick identification of many equine viruses. Conventional PCR provides a non-quantitative, positive or negative answer, which is sufficient for diagnostic purpose. Absolute or relative PCR quantification indicates the number or load of viruses in the sample but is difficult to do properly on a large scale. Clinicians should also be aware that while PCR is very sensitive, contamination of samples during collection or shipping can decrease the test’s specificity. PCR results should be interpreted with the prevalence of the disease in mind. Pharyngeal or nasal swabs are usually submitted for PCR
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Table 66.1 Normal values for blood gases in newborn foals (modified from Stewart et al. 1984)3 Age
Pa o2 (mmHg)
Pa co2 (mmHg)
pH
Standard bicarbonate (mmol/L)
60 min 12 h 48 h 7 days
60.9 ± 2.7 73.5 ± 3.0 74.9 ± 3.3 86.9 ± 2.2
47.3 ± 2.2 44.3 ± 1.2 46.1 ± 1.1 46.7 ± 1.1
7.362 ± 0.013 7.357 ± 0.024 7.396 ± 0.008 7.374 ± 0.014
25.3 ± 1.0 23.2 ± 1.3 25.7 ± 0.6 25.6 ± 0.8
testing, although a transtracheal wash can also be sent. The ideal swab should have a plastic and not a wooden shaft, and a Dacron or synthetic tip, not cotton. Samples should be kept in a dry tube on ice and should not be put into a bacterial growth milieu. Contact the diagnostic laboratory for specific sampling and shipping guidelines. Arterial blood gases Blood gases analysis is helpful to evaluate the degree of respiratory dysfunction and the response to therapy. Portable blood gas analyzers which have been validated for equine medicine30–32 are now available for quick and reliable bedside analysis of the samples. Sampling should be done as quickly as possible once the foal is put in lateral recumbency since there is a significant decrease in Pa o2 values (average 14 mmHg) after shifting the foal from sternal to lateral recumbency.3 The great metatarsal arteries are usually the most accessible sampling sites. Normal values for blood gases are indicated in Table 66.1.3 A retrospective study on foals with pneumonia and radiographic lung lesions found that 73% of these foals had respiratory acidosis (decreased pH with increased Pa co2 ). Lower pH values (< 7.3) were significantly correlated with non-survival, while the most important criterion for non-survival was found to be the anion gap in several foal studies.29, 33 Pa o2 appears to have a prognostic value for survival in newborn foals with pneumonia.24 Pa o2 < 60 mmHg is indicative of hypoxemia but may also be affected by body position and respiratory rate. Young foals with pneumonia can be severely hypoxic due to ventilation–perfusion mismatch. The presence and severity of shunts can be evaluated by administrating 100% O2 at a high rate (10 L/min.). If the Pa o2 does not increase above 100 mmHg then the foal suffers from shunts that allow unoxygenated blood to bypass ventilated alveoli. Most foals with pneumonia have more difficulties maintaining adequate perfusion (e.g., blood oxygenation) versus adequate ventilation; thus hypoxemia often develops well before hypercapnia. Sick foals with pneumonia are usually hyperventilating, thus eliminating properly CO2 which is much more soluble than O2 .
Treatment Antibiotic therapy Due to the close association between neonatal pneumonia and septicemia, antimicrobial therapy should be initiated as soon as possible without waiting for the results of the blood and transtracheal wash cultures. Therefore, the choice of antibiotic(s) often is an educated guess about the possible pathogens involved in the infection. A recent retrospective study on bacterial isolates from blood cultures in foals in intensive care units found approximately a third of them had a single Gram-positive organism growth and most of the other pathogens were Gram-negative bacteria.16 Although there are no retrospective studies examining bacterial isolates from transtracheal washes in foals less than a month old, one study reported that greater than 50% of cases with a positive blood culture had the same bacteria isolated from other body fluid cultures including transtracheal wash, synovial fluid, umbilical exudate, or cerebrospinal fluid. It is likely that foals with septicemia that develop pneumonia are affected by the same pathogens in their lungs.34 Broad-spectrum antibiotic therapy is recommended after the adequate samples have been sent for culture and sensitivity, although the blood culture does not seem to be affected even if antibiotic therapy was started before the sample collection.34 In one study16 the pathogens isolated from blood cultures were (in descending order of frequency) for the Gram-negative: E. coli, Enterobacter spp, Actinobacillus spp, Klebsiella, Salmonella, and Pseudomonas, and for the Gram-positive: Streptococcus spp, Enterococcus spp, and Staphylococcus spp. Amikacin has better efficacy than gentamicin against all Gram-negative bacteria, except for some Actinobacillus spp.16, 35 Amikacin has a concentration dependent activity and a post-antibiotic effect on bacteria and should be administered at a high dose once daily (21 mg/kg s.i.d. i.v.) to maximize its efficacy while reducing nephrotoxicity.36 Ceftiofur is an interesting alternative but should probably be used after confirmation that the offending pathogens are susceptible. The recommended dosage of ceftiofur when
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administered by the i.v. route in septic foals is much higher (> 5 mg/kg b.i.d., up to 10 mg/kg t.i.d.) than in adult horses.37 In severe cases of septicemia associated with resistant bacteria a combination of ticarcillin and clavulanic acid may be used (50 to 100 mg ticarcillin i.v., q.i.d.).35, 38 Management of foals with viral pneumonia Specific therapy against viral infections has not been evaluated in foals and measures for supportive therapy as well as treatment of secondary infections should be put in place first. The status of the foal’s immune system should be evaluated by measuring and rechecking the plasma levels of IgG. Combined immunodeficiency should be considered in Arabian and Thoroughbred foals with a confirmed diagnostic of EAV pneumonia. Frequent administration of hyperimmune plasma may be performed in these foals as a temporary treatment. The pharmacokinetics of the antiviral drug acyclovir, whose activity is restricted against herpes viruses, has not been studied in foals. However, it has been used in neonates during an EHV-1 outbreak in a herd with partial success in mildly affected foals.39 When a viral infection is suspected, good hygiene and biosecurity measures are important to avoid contaminating the environment with infective virions. Isolation of the foal and mare is recommended for a suspected viral infection. Disinfection of the boxstall and contaminated materials with viricidal products should be performed. Oxygen insufflation and ventilation Oxygen supplementation is indicated when foals are hypoxemic (Pa o2 < 65 mmHg) while in sternal recumbency. Oxygen insufflation is best performed by inserting an intranasal catheter to the level of the medial canthus of the eye. In cases of severe hypoxemia, oxygen supplementation can also be delivered using a percutaneous transtracheal catheter.40 It is important to use a bubbling system with sterile water to increase the moisture content of the gas. Flow rates between 5 and 10 L/min are often used when initiating the therapy. Excessive Pa o2 should be avoided as it may result in excessive free oxygen radical production. If a foal does not respond to O2 insufflation, then mechanical ventilation using positive end expiratory pressure should be considered. Mechanical ventilation may not be sufficient to improve the ventilation/perfusion (V/P) mismatch and is usually more effective in cases suffering from hypoventilation and hypercapnia. Cases necessitating mechanical ventilation are challenging and better managed in intensive care units. Assisted ventilation can induce
lung trauma, decrease cardiac output, and spread the lung infection throughout the airways. Maintaining adequate perfusion is very important to optimize V/P ratio in septic foals. Administration of i.v. fluids (colloids and crystalloids) may help improve cardiac output and pulmonary perfusion. Caution should be used to avoid administrating an excessive amount of i.v. fluids which may in turn impair the gas exchange in the lungs. In addition to the usual physical examination parameters, urinary output and precise weighing of the foal may help estimate the volume of i.v. fluids needed. In addition, pressor support using continuous rate infusion of dobutamine or/and dopamine may be needed in hypotensive, septic foals. The use of such medications requires frequent or constant monitoring of the arterial pressure. Bronchodilators and respiratory stimulants Bronchodilators are not used as a first-line treatment in young foals with pneumonia, but may be necessary when hypoxia is persistent and the response to oxygen therapy is not satisfactory. The purpose of bronchodilator therapy is to increase the volume of lung tissues ventilated to improve the V/P ratio. There are three classes of bronchodilators available that act on different receptors (anticholinergic, β2-adrenergic agonists, methylxanthines) and are available for different routes of administration. Oral β2-adrenergic agonists like clenbuterol may be used in foals at a dose of 0.8–3.2 µg/kg b.i.d.. The higher doses may be necessary to obtain an effect in many cases but they are more likely to come with side effects like tachycardia, excitation, and sweating. Caution and very close monitoring of the cardiopulmonary functions should be used when administering clenbuterol to very young neonates with critical cardiopulmonary parameters. Theophylline, a methylxanthine, is also available for oral administration (1 mg/kg b.i.d. to t.i.d.), but its therapeutic index is narrow and side effects like excitation can be observed before the bronchodilatory effect is obtained. Ipratropium, an inhaled anticholinergic, can be used with a special device like the AeroHippus or the small Aeromask (both from Trudell Medical International). Salmeterol is also available as an inhaled β2-adrenergic agonist that may have a longer activity. However, the doses for foals have not been validated yet. Caffeine, a central stimulant, has been used in foals suffering from the hypoxic-ischemic encephalopathy, but should not be used in young foals with pneumonia that already have a high ventilatory rate in response to hypoxia or hypercapnia. A recent study found that doxapram (continuous
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Bacterial and Viral Respiratory Disease of the Neonate rate infusion 0.02–0.05 mg/kg/h), a molecule that improves ventilation by direct stimulation of the medullary respiratory centers, is more effective than caffeine (p.o. initial dose 7.5–12 mg/kg followed by 2.5–5 mg/kg daily) to decrease the Pa co2 and HCO3 − levels. Nebulization therapy Although nebulization therapy has only been reported in a limited number of sick foals,41 high levels of medications can be achieved in the lower respiratory tract of normal lungs using a facemask coupled to a nebulizer. Jet nebulizers that use compressed air seem to be more effective at delivering medications into the lungs than ultrasonic nebulizers.42 The author recommends jet nebulizers that are effective even when tilted at an angle. High therapeutic concentrations of gentamicin and ceftiofur can be achieved in the respiratory secretions of horses by nebulization, without side effects.43, 44 To increase the diffusion of the medication in parts of the lung that are not optimally ventilated (which is frequent in foals with pneumonia) it is usually recommended to precede the antibiotic nebulization by the administration of an inhaled bronchodilator. Nebulization of sterile water, saline or N-acetylcysteine have been shown to decrease the viscosity of airway mucus,45 which facilitates their expectoration thus increasing the airflow through the lungs. Hygienic and management precautions It is important to encourage newborn foals with compromised respiratory function to spend as much time as possible in sternal or standing positions to optimize ventilation. Sick newborn foals should be kept comfortable and dry, in a warm, well-ventilated environment. Handling and stressful procedures should be kept to a minimum. Providing adequate nutrition and fluid intake are also extremely important to avoid catabolism and dehydration.
Prognosis Pa o2 and Pa co2 have a prognostic value for premature foals2 but further studies are necessary for young foals with pneumonia. In a population of foals in intensive care units, independently of the diagnosis, the anion gap as well as venous Pa o2 had a prognostic value that has been put into a model for calculating the probability of non-survival in these cases;33 a majority of these foals were affected by septicemia. Similarly another study on septic foals found that foals presented with a respiratory rate of 60 or more per
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minute, being able to stand and with a blood pH higher than 7.35 and a positive base excess, had an increased chance of survival.15 Septic arthritis with multisystemic disease is associated with an unfavorable prognosis for survival and racing.46 Lastly, EHV1 infections in neonates have a very poor prognosis and are usually associated with severe lesions in the lungs, liver, and lymphoreticular tissues that induce death shortly after birth. EAV infections usually present with severe clinical signs of respiratory insufficiency in foals with severe immunodeficiency and have a very poor outcome.
References 1. Rossdale PD. Clinical studies on 4 newborn thoroughbred foals suffering from convulsions with special reference to blood gas chemistry and pulmonary ventilation. Res Vet Sci 1969;10(3):279–91. 2. Rose RJ, Rossdale PD, Leadon DP. Blood gas and acid–base status in spontaneously delivered, term-induced and induced premature foals. J Reprod Fertil Suppl 1982;32:521–8. 3. Stewart JH, Rose RJ, Barko AM. Respiratory studies in foals from birth to seven days old. Equine Vet J 1984;16(4):323–8. 4. Rossdale PD. The adaptive processes of the newborn foal. Vet Rec 1970;87(2):37–8. 5. Rose RJ. Cardiorespiratory adaptations in neonatal foals. Equine Vet J Suppl 1988;(5):11–13. 6. Gillespie JR. Postnatal lung growth and function in the foal. J Reprod Fertil Suppl 1975;( 23): 667–71. 7. Koterba AM, Wozniak JA, Kosch PC. Respiratory mechanics of the horse during the first year of life. Respir Physiol 1994;95(1):21–41. 8. Koterba AM, Kosch PC. Respiratory mechanics and breathing pattern in the neonatal foal. J Reprod Fertil Suppl 1987;35:575–85. 9. Pattle RE, Rossdale PD, Schock C, Creasey JM. The development of the lung and its surfactant in the foal and in other species. J Reprod Fertil Suppl 1975;(23):651–7. 10. Stewart JH, Young IH, Rose RJ, Costas L, Barko AM. The distribution of ventilation-perfusion ratios in the lungs of newborn foals. J Dev Physiol 1987;9(4):309–24. 11. Stewart JH, Rose RJ, Young IH, Costas L. The distribution of ventilation-perfusion ratios in the lungs of a dysmature foal. Equine Vet J 1990;22(6):442–6. 12. Cohen ND. Causes of and farm management factors associated with disease and death in foals. J Am Vet Med Assoc 1994;204(10):1644–51. 13. Hoffman AM, Viel L, Juniper E, Prescott JF. Clinical and endoscopic study to estimate the incidence of distal respiratory tract infection in thoroughbred foals on Ontario breeding farms. Am J Vet Res 1993;54(10):1602–7. 14. Stewart AJ, Hinchcliff KW, Saville WJ, Jose-Cunilleras E, Hardy J, Kohn CW, Reed SM, Kowalski JJ. Actinobacillus sp. bacteremia in foals: clinical signs and prognosis. J Vet Intern Med 2002;16(4):464–71. 15. Gayle JM, Cohen ND, Chaffin MK. Factors associated with survival in septicemic foals: 65 cases (1988–1995). J Vet Intern Med 1998;12(3):140–6. 16. Marsh PS, Palmer JE. Bacterial isolates from blood and their susceptibility patterns in critically ill foals: 543 cases (1991–1998). J Am Vet Med Assoc 2001;218(10):1608–10.
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17. Bryans J, Swerczek T, Darlington R. Neonatal foal disease associated with perinatal infection by equine herpes virus 1. J Eq Med Surg 1977;1:20. 18. Blunden A, Smith K, Binns M, Zhang L, Gower S, Mumford J. Replication of equid herpesvirus 4 in endothelial cells and synovia of a field case of viral pneumonia and synovitis in a foal. J Comp Pathol 1995;112(2):133–40. 19. McCartan C, Russell M, Wood JL, Mumford J. Clinical, serological and virological characteristics of an outbreak of paresis and neonatal foal disease due to equine herpesvirus-1 on a stud farm. Vet Rec 1995;136(1):7–12. 20. Buergelt C, Hines S, Cantor G. A retrospective study of proliferative interstitial lung disease of horses in Florida. Vet Pathol 1986;23:750. 21. Carman S, Rae C, DuBovi E. Equine arteritis virus isolated from a Standardbred foal with pneumonia. Can Vet J 1988;29(11):937. 22. Peek SL, Landolt G, Karasin AI, Slack JA, Steinberg H, Semrad SD, Olsen CW. Acute respiratory distress syndrome and fatal interstitial pneumonia associated with equine influenza in a neonatal foal. J Vet Intern Med 2004;18(1):132– 4. 23. Henry J, Gagnon A. Adenovirus pneumonia in an Arabian foal. Can Vet J 1976;17(8):220–1. 24. Lester GD, Ackerman N, Steible-Hartless C. Respiratory disease in the newborn foal: radiographic and clinical findings. Dorothy Havemeyer Foundation Neonatal Septicemia Workshop. Boston, MA, 1998. 25. Jean D, Picandet V, Macieira S, Beauregard G, D’Anjou MA, Beauchamp G. Detection of rib trauma in newborn foals in an equine critical care unit: a comparison of ultrasonography, radiography and physical examination. Equine Vet J 2007;39(2):158–63. 26. Bedenice D, Heuwieser W, Brawer R, Solano M, Rand W, Paradis MR. Clinical and prognostic significance of radiographic pattern, distribution, and severity of thoracic radiographic changes in neonatal foals. J Vet Intern Med 2003;17(6):876–86. 27. Lester GD, Lester NV. Abdominal and thoracic radiography in the neonate. Vet Clin N Am Equine 2001;17(1):19– 46, v. 28. Ramirez S, Lester GD, Roberts GR. Diagnostic contribution of thoracic ultrasonography in 17 foals with Rhodococcus equi pneumonia. Vet Radiol Ultrasound 2004;45(2):172–6. 29. Bedenice D, Heuwieser W, Solano M, Rand W, Paradis MR. Risk factors and prognostic variables for survival of foals with radiographic evidence of pulmonary disease. J Vet Intern Med 2003;17(6):868–75. 30. Grosenbaugh DA, Gadawski JE, Muir WW. Evaluation of a portable clinical analyzer in a veterinary hospital setting. J Am Vet Med Assoc 1998;213(5):691–4. 31. Looney AL, Ludders J, Erb HN, Gleed R, Moon P. Use of a handheld device for analysis of blood electrolyte concentrations and blood gas partial pressures in dogs and horses. J Am Vet Med Assoc 1998;213(4):526–30.
32. Klein LV, Soma LR, Nann LE. Accuracy and precision of the portable StatPal II and the laboratory-based NOVA stat profile 1 for measurement of pH, P(CO2 ), and P(O2 ) in equine blood. Vet Surg 1999;28(1):67–76. 33. Hoffman AM, Staempfli HR, Willan A. Prognostic variables for survival of neonatal foals under intensive care. J Vet Intern Med 1992;6(2):89–95. 34. Henson S, Barton M. Bacterial isolates and antibiotic sensitivity patterns from septicemic neonatal foals: a 15year retrospective study (1986–2000). Dorothy R. Havemeyer Foundation Neonatal Septicemia Workshop. Talloires, France, 2001. 35. Giguere S. Antimicrobial drug use in horses. In: Giguere, S, Prescott JF, Baggot JD, Walker RD, Dowling PM (eds) Antimicrobial Therapy in Veterinary Medicine. Ames, IA: Blackwell, 2006; p. 449. 36. Magdesian KG, Wilson WD, Mihalyi J. Pharmacokinetics of a high dose of amikacin administered at extended intervals to neonatal foals. Am J Vet Res 2004;65(4):473–9. 37. Wilson WD, Mihalyi JE. Comparative pharmacokinetics of ceftiofur in neonatal foals and adult horses. Dorothy R. Havemeyer Foundation Neonatal Septicemia Workshop, Boston, MA, 1998. 38. Palmer JE. Recognition and resuscitation of the critically ill foal. In Paradis MR (ed.) Equine Neonatal Medicine: A Case-Based Approach. Philadelphia: Elsevier Saunders, 2006; p. 121. 39. Murray MJ, del Piero F, Jeffrey SC, Davis MS, Furr MO, Dubovi EJ, Mayo JA. Neonatal equine herpesvirus type 1 infection on a thoroughbred breeding farm. J Vet Intern Med 1998;12(1):36–41. 40. Hoffman AM, Viel L. A percutaneous transtracheal catheter system for improved oxygenation in foals with respiratory distress. Equine Vet J 1992;24(3):239–41. 41. Morresey PR. How to deliver respiratory treatments to neonates by nebulization. Proceedings of the 54th Annual Convention of the American Association of Equine Practitioners, San Diego, 2008. 42. Votion D, Ghafir Y, Munsters K, Duvivier DH, Art T, Lekeux P. Aerosol deposition in equine lungs following ultrasonic nebulisation versus jet aerosol delivery system. Equine Vet J 1997;29(5):388–93. 43. McKenzie HC. Characterization of antimicrobial aerosols for administration to horses. Vet Ther 2003;4(2):110–19. 44. McKenzie HC 3rd, Murray MJ. Concentrations of gentamicin in serum and bronchial lavage fluid after once-daily aerosol administration to horses for seven days. Am J Vet Res 2004;65(2):173–8. 45. Dixon P. Therapeutics of the respiratory tract. In: Robinson NE (ed.) Current Therapy in Equine Medicine. Philadelphia: Saunders, 1992;303–10. 46. Steel CM, Hunt AR, Adams PL, Robertson ID, Chicken C, Yovich JV, Stick JA. Factors associated with prognosis for survival and athletic use in foals with septic arthritis: 93 cases (1987–1994). J Am Vet Med Assoc 1999;215(7):973–7.
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Chapter 67
Congenital Abnormalities of the Respiratory System Eric J. Parente
Introduction Most upper respiratory abnormalities in neonatal foals are not life threatening but can lead to lifethreatening consequences such as aspiration pneumonia. Thus recognition in the early stages is critical. Often treatment of the primary problem can be or even should be postponed until the animal is more mature, but management changes or treatment of the secondary problems may be recommended more immediately. This is particularly true if the abnormality results in some degree of dysphagia. Evaluation of a suspected abnormality should start with a physical examination. It is frequently difficult to do a good endoscopic examination on a young foal because of the limitations of the nasal passage relative to the diameter of most endoscopes. Looking for asymmetry of the head, evaluating symmetry of airflow through the nostrils, looking for signs of dysphagia (discharge at the nostrils or coughing), and palpation of the larynx all provide valuable information in determining the diagnosis. Endoscopic examination of the foal can be very difficult. If the foal has a large enough nasal passage to accommodate an endoscope, significant restraint is required to minimize the risk of any bleeding that would otherwise significantly compromise the ability to see anything of the upper respiratory tract. Sedation, such as small doses of xylazine, can be employed if necessary but it must be kept in mind that sedation will alter pharyngeal and laryngeal tone. Alternatively topical application of carbocaine at the opening of the nares will minimize the foal’s resistance to passing the endoscope. Functional assessments of the larynx or pharynx should not be made in the sedated animal. Even in the unsedated foal, the pharynx will appear much more flaccid than the adult horse. It is very typical to note the dorsal roof
of the pharynx rolling over the corniculates of the arytenoids and this should not be considered abnormal. Displacement of the soft palate is much more easily induced in the foal versus the adult, and this is not abnormal. While displacement may be easily induced, the palate should be below the epiglottis at all times unless provoked to displace, and displacements should be corrected with swallowing (Figure 67.1).
Nasal/sinus abnormalities Choanal atresia Choanal atresia is the only true upper respiratory emergency in foals. The choanal membrane separates the oral and nasal cavity and should undergo resorption during embryonic development under normal circumstances. Its persistence results in a complete obstruction at the junction of the caudal nasal passage and the nasopharynx (Figure 67.2). The obstruction can be unilateral or bilateral. As an obligate nasal breather, the foal will die if both nasal passages are obstructed at birth and it does not receive immediate attention. Passing an endotracheal tube or performing a tracheostomy will establish an airway, but severe organ damage from a period of hypoxia may have already occurred. Therefore, even with restoration of the airway, and reconstruction of the nasal passage, treatment for peripartum asphyxia will likely be required. If unilateral, choanal atresia often goes unnoticed until the foal is older. Since the nasal passage is the major contributor to airway resistance, increased resistance from unilateral choanal atresia could have a significant impact on the strength of the pharynx and larynx, but most animals can grow normally with
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Respiratory System hemorrhage at the first surgery and the likelihood of stricture formation and the need for further surgery. An owner committed to a successful outcome should be prepared for multiple revision surgeries until the animal is almost fully grown. Dilation after radial cuts of any stricture appears to be more effective than stenting to maximize the size of the lumen.1 An alternative approach to laser treatment would be a direct surgical resection via a nasal flap surgery under general anesthesia. Regardless of which approach is used, the prognosis for performance is still somewhat guarded. The subjectively narrow width of the nasal passage in foals with this abnormality may impact the long-term prognosis Wry nose
Figure 67.1 Dorsal displacement of the soft palate. The epiglottis is not visible on endoscopic view of the nasopharynx.
unilateral nasal obstruction. Generally it is prudent to wait until the animal with unilateral choanal atresia is more mature and larger before attempting surgical correction. Invariably the obstruction in horses is completely membranous, unlike other species that may have a bony component. This means laser fenestration under videoendoscopic guidance is feasible. The complications associated with laser fenestration are S
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These deviations involve both a lateral deviation as well as some rotational deviation along the axis of the nasal passage. Mild deviations may not cause significant respiratory obstruction, but moderate to severe deviations will cause respiratory obstructions from accompanying septal and turbinate deviations, usually on the convex side of the curvature. Furthermore, nares development is adversely affected and there is a limited ability to dilate the nostril on the convex side (Figure 67.3). While surgery can be performed to correct the bony deviation, it is extremely difficult to correct the septal, turbinate, or nares deviations without major resection of these tissues. The prognosis for obtaining an athletic animal even after surgery is still poor except in relatively mild cases.
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Figure 67.2 Endoscopic view of the caudal right nasal passage of a foal with choanal atresia. Note the obstructing membrane centrally which should be an opening into the nasal passage (A), the wall of the ventral turbinate (B), and the septum (C).
Figure 67.3 Wry nose in a yearling that was allowed to mature with this severe deviation. Note the collapse of the right nares.
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Congenital Abnormalities of the Respiratory System Paranasal sinus cysts It is very uncommon, but infrequently a paranasal sinus cyst can develop in utero. The foal will be born with obvious unilateral facial swelling/deviation and likely obstruction of the adjacent nasal passage. Radiography and endoscopic examination are required to confirm the diagnosis as would be performed in the adult. Surgical extirpation through a sinusotomy is required and the prognosis is generally excellent. As with any animal undergoing sinus surgery, considerations should be made prior to surgery for whole blood transfusion if necessary.
Pharyngeal abnormalities Persistent dorsal displacement of the soft palate The clinical evidence of a persistent displacement of the soft palate is usually not manifested by respiratory noise or dyspnea but more typically by dysphagia (milk at the nares) or a secondary aspiration pneumonia. Abnormal expiratory noise may be associated with structural abnormalities of the epiglottis. These might include a large subepiglottic cyst or a persistent frenulum of the epiglottis.2 Primary functional soft palate displacements are more common. While we do not know what causes a primary displacement of the soft palate, it is suspected to be associated with neuromuscular weakness. Any disorder which causes profound whole body neuromuscular weakness in the foal (botulism, white muscle disease) will cause pharyngeal dysfunction and dysphagia. Focal pharyngeal dysfunction could be a result of a transient episode of hypoxia or toxin exposure during gestation, but the latter is very unlikely given how quickly the problem resolves under most conditions. Both surgical and medical treatments have been employed3, 4 and both have reported success. While the surgical approach of staphylectomy (shortening the palate) may allow the foal to resolve the displacement in the short term, there should be concern about the long-term upper respiratory function during exercise, and some long-term dysphagia has been reported.3 Since most of these foals will have pneumonia and will be undernourished and weak, aggressive medical management is required prior to any surgical intervention. Antimicrobials, anti-inflammatory therapy, and feeding via a nasogastric tube or parenteral nutrition are necessary. Rarely is tracheostomy required. Typically as foals gain increased strength and resolve concurrent respiratory infection, they will correct their pharyngeal dysfunction.4 A slow transition to bucket feeding is necessary to main-
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tain adequate nutrition. Most foals respond to medical management within days and those that take longer may have some residual pharyngeal weakness. Their long-term ability to exercise successfully or race has not been well established. Those individuals that do not resolve their displacement within days of medical management should be re-evaluated for an underlying structural abnormality as described above. Cleft palate Dysphagia manifested by milk at the nose can also be caused by a cleft palate. Clefts are the result of a failure of the lateral palatine processes to fuse on midline. They can be secondary to genetic or environmental (toxic) factors. Exposure to pesticides and ingestion of plants of certain Lupinus spp has been associated with cleft palates and other teratogenic abnormalities. The cause is rarely defined in the individual equine patient. In most foals, the cleft spans almost the entire soft palate and rarely involves any of the hard palate. Diagnosis may be made during an oral examination more easily than a videoendoscopic evaluation through the nares. A cleft palate undiagnosed for any significant length of time will typically result in pneumonia. The pneumonia should be treated aggressively with medical management prior to any surgical intervention. Surgical repair of a cleft palate is difficult, requires a mandibular symphysiotomy, and possibly a laryngotomy to access the palate. In many reports there is some partial dehiscence of the repair. Even if the repair is completely successful, it is unlikely to be functionally normal to the point which the animal can become a racehorse.5, 6 Nasopharyngeal cysts Most pharyngeal cysts do not cause any clinical abnormalities. Those on the wall of the pharynx should not be considered the source of any clinical abnormalities unless they are so large that they impede visualization of the larynx. Surgical resection is usually unnecessary. Cysts within the palate are very different and do cause significant problems. Their location will impede the normal positioning of the epiglottis above the palate and likely create some degree of dysphagia and possibly respiratory compromise. A lateral radiograph can be helpful in determining the presence of any structural abnormalities that result in palate displacement if endoscopy is not available (Figure 67.4). Resection of these cysts can result in defects in the palate and therefore it is better to ablate the cyst with laser energy via laryngotomy and not resect any tissue. Medical management of the foal as described
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Foal: Respiratory System with displacement of the soft palate should take place concurrently. Guttural pouch tympany This is not a congenital abnormality but is most often observed in the young foal. For an unknown reason, the opening to the guttural pouch allows air to enter but not escape, resulting in an increased accumulation of air and distension of the guttural pouch. The diagnosis can usually be made on physical examination by the appearance of a soft, easily compressible bulge behind the vertical ramus of the mandible. It is most commonly unilateral but often appears bilateral since there is often swelling bilaterally. The side with the greater bulge is the affected side (Figure 67.5a-c) Endoscopic examination is typically unremarkable. In the more severe cases there
Figure 67.4 Lateral radiograph of a foal with a cyst at the free edge of the palate (arrow) resulting in persistent displacement of the soft palate. An esophageal feeding tube (arrowhead) is in place to prevent aspiration from nursing.
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(c) Figure 67.5 (a) Foal with guttural pouch tympany and asymmetric left-sided swelling. (b) The right side of the same foal has some right-sided swelling despite it being a unilateral left-sided problem. (c) The appearance of the foal immediately after a left-sided salpingopharyngeal fistula was created.
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Congenital Abnormalities of the Respiratory System will also be some history of respiratory stridor associated with concurrent pharyngeal collapse over the larynx that can be observed on endoscopic examination. There are multiple treatment options all with the goal of creating a way to allow air to escape from the guttural pouch. The preferred method by the author is to create a salpingopharyngeal fistula, which is a direct opening from the pharynx into the guttural pouch. This can be performed with transendoscopic laser surgery in the standing, sedated patient with just local anesthetic. It provides immediate resolution of the tympany and appears less likely to stricture than a previously described method of creating a fistula in the median septum between the pouches.
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Laryngeal abnormalities Laryngeal dysfunction Laryngeal paresis or paralysis in the neonate is similar to pharyngeal dysfunction. It appears to be a neuromuscular dysfunction that should resolve over time and the primary objective is to manage the patient during this time. The dysfunction is most frequently bilateral (unlike the idiopathic form of recurrent laryngeal neuropathy in the adult) and typically not complete paralysis. Respiratory distress or stridor at birth or shortly thereafter is the primary clinical sign and tracheostomy or endotracheal intubation can be life saving. While there is no information on foals, our experience is that most foals regain adequate function within days if provided with an opportunity to breathe easily through a tracheostomy and not stressed. In humans, the majority of babies regain good function quickly, but the duration to complete recovery may be several months.7
Figure 67.6 Oropharyngeal view of a foal with persistent displacement of the soft palate secondary to a large subepiglottic cyst. (A) Tip of the epiglottis within the oropharynx, (B) ventral surface of the soft palate, (C) large subepiglottic cyst.
tic cyst and the endoscopic examination appears normal. Subepiglottic cysts have been resected many ways. They can be grasped through a laryngotomy, orally, or nasally. Transendoscopic laser surgery can be used to resect them through the mouth or transnasally.8 If this technology is not available, a snare can be created from obstetric wire to resect the cyst orally with the foal under anesthesia. The prognosis is generally excellent.
Laryngeal malformations Subepiglottic cysts Subepiglottic cysts can present similarly to palatal cysts. Their location may impede the position of the epiglottis and result in persistent palate displacement and severe dysphagia (Figure 67.6). If they are smaller, the palate could remain in normal position at rest but the foal can present with intermittent dysphagia. Initial endoscopic evaluation may assist in making the diagnosis if the cyst is above the palate. If the epiglottis is displaced ventrally, or if the epiglottis is dorsal but the cyst is ventral during the examination, the cyst will not be observed. A lateral radiograph of the larynx is recommended to see what is below the palate if there is clinical suspicion of a subepiglot-
Fourth branchial arch defects are congenital abnormalities that do not present as a clinical disease in the foal, but will likely present as exercise intolerance in adults.9 They are commonly misdiagnosed as right-sided laryngeal hemiplegia or ‘roarers’. The muscles and cartilages of the larynx form from the fourth and sixth branchial arches. Developmental defects in the fourth branchial arch will lead to malformation of the cartilages and their relative position to each other on the right side of the larynx. This has been most commonly observed in Thoroughbreds. The diagnosis is made by a combination of laryngeal palpation and endoscopic examination. There will often be significant asymmetry in the space between the ventral thyroid cartilages and the cricoid. Endoscopically there may be deformity of the right
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Figure 67.7 Endoscopic view of a young horse with a fourth branchial arch abnormality. Palatopharyngeal arch displacement and an irregular right corniculate process are evident.
corniculate, displacement of the palatopharyngeal arch, and typically inability to fully abduct the right arytenoid (Figure 67.7). Depending on the degree of deformation, laryngoplasty may not be an effective surgical technique to prevent respiratory compromise during exercise. While rare, laryngeal cysts will cause a more severe deviation of laryngeal structure than fourth branchial arch abnormalities and typically result in a more acute respiratory problem in the neonate or young foal (Figure 67.8). Depending on their specific position and size, laryngeal cysts can result in severe dyspnea. A cursory endoscopic examination may give the impression of typical hemiplegia, but closer inspection should give the impression that the arytenoid is pushed in rather than hanging in and palatopharyngeal arch displacement is likely. Their origin is debatable. A branchial cleft cyst is most likely given that it occurs within the laryngeal structure and is off midline.10 Surgical resection may be performed successfully,11 but cysts which have already caused permanent malformations of the laryngeal cartilages will preclude return of normal function. If such an abnormality is suspected, imaging modalities like magnetic resonance or at least radiographs should be pursued prior to surgery.
Figure 67.8 Endoscopic view of a foal with a branchial cyst. Medial deviation of the right arytenoid (arrow) and displacement of the palatopharyngeal arch (arrowhead) suggests a mass or swelling lateral to the arytenoid.
References 1. James FM. Management of bilateral choanal atresia in a foal. J Am Vet Med Assoc 2006;229(11):1784–9. 2. Yarbrough TB. Persistent frenulum of the epiglottis in four foals. Vet Surg 1999;28(4):287–91. 3. Shappell KK. Staphylectomy for treatment of dorsal displacement of the soft palate in two foals. J Am Vet Med Assoc 1989;195(10):1395–8. 4. Altmaier K. Dorsal displacement of the soft palate in neonatal foals. Equ Vet J 1993;25(4):329–32. 5. Kirkham LEMcN, Vasey JR. Surgical cleft soft palate repair in a foal. Austr Vet J 2002;80(3):143–6. 6. Keeling NG. Use of mucosal graft to augment cleft palate repair in a foal. Equ Prac 1995;17(10):34–9. 7. De Gaudemar I. Outcome of laryngeal paralysis in neonates: A long-term retrospective study of 113 cases. Int J Pediatr Otorhinolaryngol 1996;34(1–2):101–10. 8. Parente EJ. Laser surgery of the upper respiratory tract. In: McGorum BC et al. (eds) Equine Respiratory Medicine and Surgery. Philadelphia: Saunders, 2007; pp. 533–41. 9. Lane JG. Fourth branchial arch defects. In: McGorum BC et al. (eds) Equine Respiratory Medicine and Surgery. Philadelphia: Saunders, 2007; pp. 467–72. 10. Wetmore RF. Differential diagnosis of neck masses. In: Cummings CC et al. (eds) Cummings Otolaryngology Head & Neck Surgery, 4th edn. Philadelphia: Elsevier Mosby, 2005; pp. 4210–22. 11. Baxter GM. Paralaryngeal accessory bronchial cyst as a cause of laryngeal hemiplegia in a horse. Equ Vet J 1992;24(1):67–9.
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Chapter 68
Genetic Basis of Respiratory Disorders Eliane Marti, Caroline Tessier and Vincent Gerber
Introduction In the horse, congenital abnormalities such as fourth branchial arch defects and diseases caused by a mutation in a single gene, such as hyperkalemic periodic paralysis, are rare. Furthermore, respiratory diseases due to a single gene mutation are so far unknown. However, for a number of important equine respiratory diseases such as pulmonary infections and obstructive diseases of the upper and lower airways, it has long been recognized that, within groups of horses living in the same environment, some individuals become affected while others remain healthy. Clinical expression of these respiratory diseases results from complex interactions between the environment, including infectious agents, allergens, and airway irritants, and the genetic make-up of each individual horse. These differences in genetic susceptibility to respiratory diseases are likely due to variations in several genes, i.e., they are polygenic. The majority of airway diseases of foals and younger horses have an infectious origin. However, while ‘mechanical’ problems of the upper airways such as idiopathic hemiplegia laryngis are usually identified when horses start to be trained and are expected to compete, they can sometimes already be identified in foals.1 In contrast, chronic obstructive conditions of the lower airways are more frequent in mature and older horses, and their prevalence increases with age.2,3 Nevertheless, as there is a genetic basis for recurrent airway obstruction (RAO), some horses are already born with an increased susceptibility for this disease. Therefore, breeders can reduce the number of RAO-susceptible horses by selecting appropriate parents, i.e., by avoiding matings of two RAO-affected parents. Furthermore, optimal management of genetically RAO-susceptible foals and young horses may prevent the clinical ex-
pression of this condition. Thus, knowledge about genetics of equine respiratory diseases can influence decisions in reproduction.
Infectious diseases of the respiratory tract The notion that susceptibility to infectious diseases may be inherited is widely accepted. In the horse, susceptibility/resistance to clinical infectious respiratory disease probably also has a genetic component, but only a few studies have been performed in this field. Two studies have shown that transferrin alleles are associated with severity of respiratory diseases. The first study showed that Thoroughbred foals that succumbed to Rhodococcus equi infection showed an excess of the transferrin F allele and a deficiency of the D allele.4 In the second study, Newton et al.5 investigated which factors and infections are associated with naturally occurring respiratory disease in recently weaned ponies. They found that the predominant infection associated with clinical disease was S. zooepidemicus and that foals with the transferrin D allele demonstrated significantly less clinical disease, while ponies carrying the F2 transferrin allele had more severe clinical signs. Thus, although different respiratory pathogens were studied in these two investigations, in both cases transferrin D alleles were associated with less severe clinical signs of respiratory disease. Transferrin is an iron-binding protein that has bacteriostatic properties in the blood. The specific mechanism by which transferrin alleles are related to differences in severity of infectious respiratory diseases is not known. Further studies are necessary to substantiate these findings. Furthermore, many other genes will probably also contribute to resistance/susceptibility to infectious respiratory diseases in the horse. A preliminary study suggests that
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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the solute carrier family 11 member 1 (SLC11A1) gene is associated with susceptibility to R. equi pneumonia in Arabian and Thoroughbred foals from two farms.6 Various other genes are likely to be involved. For example, in humans, alleles of the major histocompatibility complex and of the mannose binding complex are associated with susceptibility to infectious diseases of the respiratory tract.7,8
Non-infectious diseases of the respiratory tract Upper airway disease
Guttural pouch tympany Guttural pouch tympany (GPT) is a condition that affects foals and only rarely affects older horses. The condition requires surgical treatment. The exact causes of GPT are not known. GPT develops when air cannot leave the guttural pouch through the pharyngeal orifice, suggesting that either an abnormally large mucosal flap at the pharyngeal orifice or a functional failure of the pharyngeal orifice can lead to air entrapment in the guttural pouch and subsequently to GPT. Fillies are more frequently affected than colts.9 Guttural pouch tympany has been described in various breeds, including Thoroughbreds, Warmbloods, and Arabian horses. Blazyczek et al.10 observed that, of 51 cases treated between 1994 and 2001 at the clinic for horses at the School of Veterinary Medicine in Hannover, 24 were Arabian foals, and 22 of these foals belonged to only four different families. Additionally, some were full- or half-siblings, suggesting a genetic basis for GPT. A segregation analysis performed with 276 animals of these four families showed that, among all models tested, a polygenic and a mixed monogenic-polygenic model best explained the segregation of GPT in Arabian foals,10 demonstrating that a genetic component significantly contributes to the development of GPT in Arabian foals. Moreover, heritability for GPT was estimated at 0.49 ± 0.28 in Arabian foals9 and 0.81 ± 0.16 in German Warmblood foals,11 suggesting that GPT is also strongly influenced by genetic factors in other breeds. A recent study performed by the same investigators has enabled identification of quantitative trait loci for GPT on equine chromosomes 2 and 15.12 This is a first step towards identifying the genes responsible for GPT.
Laryngeal hemiplegia Laryngeal hemiplegia has long been recognized as the most important upper airway disease in horses. It causes inspiratory stridor and exercise intolerance at higher speeds and workloads. This disease occurs
worldwide and is of particular relevance in sport horses with poor performance. Characteristics of the disease include atrophy of the laryngeal muscles innervated by the recurrent laryngeal nerve. There are numerous possible causes of this disease. In a limited number of cases, a specific etiology can be identified: perivascular/perineural injections of irritant products, guttural pouch mycosis, trauma to the neck, organophosphate intoxication, fourth branchial arch defects, and lead and plant poisoning.1 However, these specific causes can explain only a small proportion of the observed cases. In a recent study of 375 horses of various breeds with laryngeal hemiplegia, 94% were idiopathic in origin.13 Idiopathic laryngeal hemiplegia (ILH) is also referred to as recurrent laryngeal neuropathy. ILH is thought to be due to nerve damage, either as a result of stretching of the recurrent laryngeal nerve or following nerve compression in the region of the aortic arch.1,14 Thiamin deficiency has also been incriminated in the development of ILH,15 as well as bacterial and viral infections, and unknown plant and chemical toxicities. Pathological changes Lesions in the laryngeal muscles are characteristic of neurogenic disease. Denervation of the adductor muscles precedes abductor involvement and typical changes include scattered angular fibers and groups of atrophied fibers adjacent to hypertrophied fibers with central nuclei.16,17 These lesions have been referred to as distal axonopathy, with the much more severe pathology in the left recurrent laryngeal nerve being explained by its greater length. Myelinopathies and distal axonopathies typically affect multiple nerves and would be expressed in other species as polyneuropathy and progressive. ILH has been reported in young animals18–20 and was shown to be progressive in some horses.21 However, there has been no report of diseases progressing to produce right-sided hemiplegia. Furthermore, affected horses do not display other lesions of polyneuropathy such as mega-esophagus, limb paresis, or muscle atrophy. Anecdotal reports have shown involvement of other long nerves in horses affected by ILH but they are rare.22,23 Epidemiological studies Although a genetic predisposition for laryngeal hemiplegia has been proposed for a long time, obtaining evidence has been complicated by the confounding effects of other factors such as breed, sex, age, size, and conformation. Many authors have reported that ILH can occur in foals and yearlings.18–20 In 15% of cases, ILH has been
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Genetic Basis of Respiratory Disorders shown to be a progressive disease, and muscular atrophy seems to progress in some cases as the horse ages.21 In a study evaluating the prevalence of ILH in competing draft horses, 35% of the animals were affected by laryngeal hemiplegia, which is a higher proportion than that reported in other breeds.24 ILH prevalence differed significantly among draft breeds: 42% of Belgians, 31% of Percherons, and 17% of Clydesdales examined were affected, suggesting that genetic factors contribute to ILH in draft horses as well. Several studies have evaluated families of horses and the effect of affected parents on the laryngeal function of their offspring:
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Galizzi and Galanti25 have shown that members of eight affected Thoroughbred families suffered in larger numbers from ILH (29%) than did the control population (3.4%). Poncet et al.26 examined 47 offspring of one affected Warmblood stallion and found that the prevalence was 47% versus 10% in the control population. However, a significant difference was observed in the average height at the withers of the affected stallion’s affected offspring compared to that of the unaffected offspring. Ohnesorge et al.1,27 evaluated 24 affected stallions and their offspring (240 foals and 474 adults), as well as the dams (308 mothers of 35 foals and 216 adults). They showed that the incidence and degree of ILH depended on the age of the offspring and the occurrence of the same dysfunction in the parents. Foals were affected significantly less frequently (24.7%) than adults (49.7%). Furthermore, the progeny from affected parents was significantly more often affected (41% of the foals, 60.9% of the adults) than comparable progeny of healthy parents (8.9% and 39.6%, respectively). Miesner,28 using the same populations as mentioned above, and using an index to grade ILH, found a heritability of 0.61 (standard error = 0.17). Length of the neck and narrow intermandibular space were positively correlated to the grade of ILH. There was no correlation between the occurrence of ILH and sex. A recent study performed in draft horses confirms that conformation of the horse is, at least in some breeds, associated with IHL: in Belgian and Percheron horses, but not in Clydesdales, horse height was significantly associated with the risk of ILH.24 As conformational traits usually have a high heritability, it is thus difficult to determine if there is a true genetic influence on ILH which is separate from size.
These findings show that genetic factors probably play an important role in the etiology of ILH.
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However, to date, no studies have demonstrated a simple recessive or dominant mode of inheritance, which suggests that ILH is likely a polygenic trait. The genes contributing to the genetic predisposition for ILH have not yet been identified. Investigations focusing on specific loci have so far yielded negative results. McClure et al.29 showed that in Thoroughbred and Quarter horses there was no association between ILH and the equine lymphocyte antigens. These results were confirmed by Poncet et al.26 in a Swiss Warmblood family.
Fourth branchial arch defect Fourth branchial arch defect is a syndrome of irreparable congenital defects resulting from a failure of development of some or all of the derivatives of the fourth branchial arch. The structures involved are the wings of the thyroid cartilage, the cricothyroid articulation, the cricothyroideus muscles, and the cricopharyngeal sphincter. Permutation of aplasia or hypoplasia of these structures may arise uni- or bilaterally. Fourth branchial arch defect is one of the causes of rare right-sided laryngeal hemiplegia. Since fourth branchial arch defect is congenital, breeders are questioning if the condition may be inherited. To date, there is no evidence that this disease is genetically transmitted. More studies of this uncommon and so far poorly understood condition are needed. Chronic lower airway diseases Non-infectious chronic lower airway disease is a major health problem of stabled mature horses. In conventional indoor stable environments with hay feeding and straw bedding, the air in the breathing zone of horses contains very high levels of organic dust with allergens and non-specific irritants such as endotoxin.30 Consequently, horses in conventional stable environments show a high incidence of mild to severe non-infectious lower airway disease.3 Horses with less severe forms of chronic airway disease are said to have inflammatory airway disease (IAD), but the definition of this phenotype is difficult and presently requires cytological documentation of lower airway inflammation or demonstration of lowgrade changes in lung function.31 Little is known about the pathogenesis of this condition and both bacterial and allergic etiologies have been discussed. To our knowledge, a genetic basis for IAD has not been investigated, although an association between transferrin alleles (see above, under infectious diseases) and IAD has been reported.32 RAO is a severe, debilitating disease characterized by coughing and increased breathing effort due to cholinergic bronchospasm and airway hyperreactivity as well as neutrophil and mucus
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accumulation in the airways as a result of hypersensitivity reactions to various inhaled allergens.33 Removal of all exposure to hay and straw often leads to remission of the disease. The immunological basis of RAO is still poorly understood. There is some evidence for IgE-mediated type I hyperreactivity,34,35 and some studies show a Th2-type bias in equine RAO with increased numbers of interleukin (IL)-4 and IL-5 positive cells36 and increased mRNA levels of IL-4 and IL-13 in bronchoalveolar lavage fluid (BALF) of RAO horses.37,38 However, Th1-type cytokines, such as interferon (IFN)-γ , can also be increased in RAO.39 Furthermore, non-specific responses to endotoxin, alone and through potentiation of reactions to mold allergens, also play a role.40 Endotoxin increases Toll-like receptor (TLR)-4 and TLR2 mRNA in horse lungs.41 However, the relative importance of IgE-mediated reactions, Th1- and Th2-type profiles, innate immunity and individual cytokines in the pathogenesis of RAO is still unclear. We suspect that some contradictory results are in part due to investigations of different groups of RAO-affected horses displaying the same clinical phenotype, but with differing immunogenetic make-ups.
Equine recurrent airway obstruction Familial aggregation, epidemiological and population genetic analyses A genetic basis of equine RAO, based on observations of family histories and familial aggregation, has been postulated for over 70 years.42 Two different studies now clearly document the presence of a genetic predisposition for RAO. In the first study,2 horses from two studs (German Warmbloods and Lipizzans) were grouped according to their own lung status and the lung status of their parents. In both studs the percentage of offspring suffering from RAO increased with increasing number of affected parents, and thus the relative risk (RR, Mantel-Haenzel statistics) of developing RAO was significantly increased in offspring with one (RR = 3.2; P < 0.05) and, even more so, with two affected parents (RR = 4.6; P < 0.05). Interestingly, these findings were quite similar to the results from a study on the inheritance of atopy in humans.43 Additionally, offspring of three RAO-affected and three healthy sires were investigated; 39 (49%) of the 79 offspring of the RAO-affected sires had a history of RAO versus only 12% of the 74 offspring of healthy sires. An analysis of variance demonstrated that the sire, the age, and the stable environment had a significant influence on the clinical condition of the offspring. In this study the diagnosis of RAO was based primarily on the history of a chronic cough in hay-fed and stabled horses which improved when horses were removed from hay. In the second investigation,
more objective criteria were used both to define the phenotype of the horse as well as to evaluate the effect of the environment. In the second study3 over 200 offspring from two RAO-affected sires were collected and compared with approximately 100 controls which were half-siblings (same dam) or unrelated horses. A standardized questionnaire was used to gather the phenotypic information on the horses’ history of chronic coughing, increased breathing effort, and nasal discharge, and detailed questions on management and feeding, deworming strategies, past and concurrent diseases, and the use of the horse. This information was used in a ‘horse owner assessed respiratory signs index’ (HOARSI) ranging from 1 to 4, with 1 being the clinically healthy horses and 3 and 4 the RAO-affected group. HOARSI-3/4 animals in clinical exacerbation showed signs consistent with RAO: coughing, nasal discharge, abnormal lung sounds and breathing pattern, as well as increased numbers of neutrophils in tracheobronchial secretions and broncheolar lavage fluid, excessive mucus accumulation, and airway hyperresponsiveness to methacholine. HOARSI-3/4 horses in remission only have increased amounts of tracheal mucus and neutrophil percentages in tracheobronchial secretions. Clinical phenotypes were not significantly different between the two families. HOARSI thus reliably identified RAO-affected horses in this population.44 Significantly more offspring of the affected sires (29.6% and 27.3%) showed moderate to severe clinical signs compared to random age-matched and halfsibling age-matched control groups (between 5 and 10%). Frequent coughing, increased nasal discharge, poor performance, and abnormal breathing pattern were all significantly increased in the offspring of the RAO-affected sires. Multivariate regression analysis showed that sire, hay feeding, and age (in decreasing order of strength) were associated with more severe clinical signs, more frequent coughing and nasal discharge. The risk of sire 1 and 2 offspring for developing RAO was increased 4.1 and 5.5 fold, respectively. Coughing, which is known to be associated with airway inflammation and obstruction in equine RAO,45 most markedly distinguished genetically predisposed RAO-affected offspring.3 Genetic models tested by segregation analysis showed the highest likelihood for a mixed inheritance model with rather high heritabilities. Segregation analyses clearly revealed the presence of a major gene playing a role in RAO. In one family, the mode of inheritance was autosomal dominant, whereas in the other family it was autosomal recessive. Although the expression of RAO is influenced by exposure to hay, these findings suggest a strong, complex genetic background for RAO.46
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Genetic Basis of Respiratory Disorders Candidate gene and genome scanning approaches After having identified a genetic basis for RAO the next step is to identify which genes could be responsible for susceptibility to this disease. For this purpose, two different approaches can be chosen: a genome scan or a candidate gene approach. In a genome scan, a set of markers which are evenly distributed over the whole genome are tested for linkage with the phenotype of interest. A genome scan using microsatellite markers was performed for RAO with the horses included in the study by Ramseyer et al.3 Analysis using a regression method for half-sibling family structures indicated two chromosome regions (on ECA13 and ECA15) with a genome-wide significant association with RAO. An additional 11 chromosome regions showed a more modest association.47 Several candidate genes are located in these regions. In the candidate gene approach, genes which could be involved in the pathogenesis of a disease are tested, or genes that have been shown to be involved in a similar condition in another species, such as asthma in humans, for example. A limited number of candidate genes have been investigated so far for RAO. No association was found between RAO and the equine leukocyte antigens (ELA)2 nor with markers for the key epithelial signaling factors CLCA1, EGFR, and BCL2 genes (Gerber, unpublished results). Of the microsatellites located near candidate genes for atopy such as IL-10, IL-9, CMA1, and NOS1, only the ECA 13q13 region containing the IL4RA gene was linked to HOARSI and coughing frequency in the offspring of one RAO-affected sire (sire 1) but not in other (sire 2).48 Interestingly, association of polymorphisms in IL4RA and equine RAO in American Quarter Horses was reported in a conference abstract.49 The IL4R gene is a strong candidate gene for RAO because, together with IL-4 and IL-13, it drives the production of IgE in B-cells and stimulates the differentiation of naive T-cells into allergypromoting T-helper-2 cells. Furthermore, the IL4R gene was also associated with asthma or other atopic phenotypes in humans.50 In the horse, IL4RA was further explored as a candidate gene by single nucleotide polymorphism (SNP) association and linkage studies, and gene expression experiments. Gene expression data indicate that there may be differences in the immunological background between the two families: offspring of sire 2 showed a Th2-type bias (high IL13/IFNγ ratio) compared to offspring of sire 1. On the other hand, expression of IL4RA was increased in RAO-affected offspring of sire 1 in exacerbation compared to the same group of sire 2 and the clinically healthy horses (Gerber, unpublished results). These findings support a role of the IL-4RA gene in RAO susceptibility in the family of sire 1.
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Interestingly, IL4RA also plays in important role in the defense against parasites and RAO was shown to be associated with resistance against strongylid parasites in the offspring of sire 1.51
Relationship of RAO and genetics of the humoral immune response Although the pathogenesis of RAO is still not fully understood, a number of studies suggest that immunoglobulin E (IgE) against molds or other allergens may be involved in the pathogenesis of this disease.34,52 Measurement of IgE levels against molds in sera of 450 Lipizzan horses from six different studs showed that, as is the case in humans, the environment as well as genetic factors influence the specific IgE response.53 Heritability estimates of 0.33 were found for IgE levels against molds. This means that 33% of the variation of the IgE levels was due to genetic factors. However, no data on the lung status of these Lipizzans was available. Further studies are thus needed to elucidate if genes influencing the IgE response account for the genetic predisposition to RAO, and to identify the genes responsible for the heritability of IgE levels. Genes within the major histocompatibility gene region have been demonstrated to influence the IgE response against some specific mold allergens.54 However, various other genes will probably also influence specific and total IgE levels in the horse. Interestingly, a recent study indicates that offspring of sire 13 have significantly higher IgE levels against a recombinant mold allergen than the offspring of sire 255 , confirming that the ability to mount a high IgE response is also influenced by genetic factors in the horse.
Summary and conclusion In this chapter we have given an overview of what is presently known on the genetics of respiratory diseases of the horse. There is now clear evidence that genetic factors influence the susceptibility to recurrent airway obstruction, guttural pouch tympany, and ILH. There is also increasing evidence to suggest that the degree of severity of infectious diseases of the respiratory tract is influenced by genetic factors not only of the pathogen but also of the host. However, more studies on this aspect are needed to understand the impact of the genetic make-up of the horse on both susceptibility/resistance to infectious disease as well as the ability to mount a protective immune response after vaccination. Furthermore, epidemiological/genetic studies on other respiratory diseases may show that other respiratory diseases, such as exercise-induced pulmonary hemorrhage, for example, have a genetic basis.
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Knowing which diseases have a genetic basis and, ideally, identification of the genes or genetic markers for these diseases could allow:
tools should enable a better understanding of the genetic background of these diseases in the future.
References
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A reduction of the prevalence of some disease through selective breeding. However, we have to keep in mind that some of these susceptible horses may have other positive traits which breeders aim to select for, and that the breeders/owners will always have to weigh the risk of having a horse predisposed to RAO, for instance, against other valuable traits of this individual. The identification of foals with a higher risk for particular diseases – even for conditions which will only become clinically apparent later in life, such as RAO. This can allow owners to prevent the clinical manifestation of these conditions through management of the horse’s environment, avoiding dust and allergens in the case of RAO-susceptible individuals. A better understanding of the pathogenesis of various diseases which will subsequently allow veterinarians to choose the most efficient treatment for these conditions. Pharmacogenetics56 is a research area in which the genetic basis of a disease is used to improve current treatments and to develop novel approaches. The understanding of genetic factors which contribute to disease status is important when trying to identify new therapeutic targets and optimizing currently available therapy for individual cases.
Horse genetics is expected to progress rapidly in the near future; in 2007 the National Human Genome Research Institute (USA) completed the sequencing of the whole horse genome (www.ncbi.nlm.nih. gov/entrez/query.fcgi?db=genomeprj).57 The initial sequencing assembly is based on 6.8-fold coverage of the horse genome, which means, on average, each base pair has been sequenced almost seven times over. In addition, researchers have produced a map of horse genetic variation using DNA samples from a variety of modern and ancestral breeds, including the Akal Teke, Andalusian, Arabian, Icelandic, Quarter Horse, Standardbred, and Thoroughbred.57 This map, comprised of 1 million signposts of variation called single nucleotide polymorphisms, or SNPs (www.broad.mit.edu/mammals/horse/snp), will provide scientists with a genome-wide view of genetic variability in horses and help them identify the genetic contributions to physical and behavioral differences, as well as to disease susceptibility. Provided that samples of groups or families of horses are phenotypically well characterized for particular traits are available, then these new genetic
1. Ohnesorge B, Deegen E, Miesner K, Geldermann H. [Laryngeal hemiplegia in warmblood horses – a study of stallions, mares and their offspring.] Zentralbl Veterinarmed A 1993;40(2):134–54. 2. Marti E, Gerber H, Essich G, Ouhlela J, Lazary S. On the genetic basis of equine allergic diseases. I. Chronic hypersensitivity bronchitis. Equine Vet J 1991;23:457–60. 3. Ramseyer A, Gaillard C, Burger D, Straub R, Jost U, Boog C, Marti E, Gerber V. Effects of genetic and environmental factors on chronic lower airway disease in horses. J Vet Intern Med 2007;21:149–56. 4. Mousel MR, Harrison L, Donahue JM, Bailey E. Rhodococcus equi and genetic susceptibility: assessing transferrin genotypes from paraffin-embedded tissues. J Vet Diagn Invest 2003;15(5):470–2. 5. Newton JR, Woodt JL, Chanter N. Evidence for transferrin allele as a host-level risk factor in naturally occurring equine respiratory disease: a preliminary study. Equine Vet J 2007;39(2):164–71. 6. Halbert ND, Cohen ND, Slovis NM, Faircloth J, Martens RJ. Variations in equid SLC11A1 (NRAMP1) genes and associations with Rhodococcus equi pneumonia in horses. J Vet Intern Med 2006;20(4):974–9. 7. Ng MH, Lau KM, Li L, Cheng SH, Chan WY, Hui PK, Zee B, Leung CB, Sung JJ. Association of human-leukocyteantigen class I (B∗ 0703) and class II (DRB1∗ 0301) genotypes with susceptibility and resistance to the development of severe acute respiratory syndrome. J Infect Dis 2004;190(3):515–18. 8. Gong MN, Zhou W, Williams PL, Thompson BT, Pothier L, Christiani DC. Polymorphisms in the mannose binding lectin-2 gene and acute respiratory distress syndrome. Crit Care Med 2007;35(1):48–56. 9. Blazyczek I, Hamann H, Ohnesorge B, Deegen E, Distl O. [Population genetic analysis of the heritability of gutteral pouch tympany in Arabian purebred foals.] Dtsch Tierarztl Wochenschr 2003;110(10):417–19. 10. Blazyczek I, Hamann H, Ohnesorge B, Deegen E, Distl O. Inheritance of guttural pouch tympany in the Arabian horse. J Hered 2004;95(3):195–9. 11. Blazyczek I, Hamann H, Ohnesorge B, Deegen E, Distl O. [Gutteral pouch tympany in German warmblood foals: influence of sex, inbreeding and blood proportions of founding breeds as well as estimation of heritability.] Berl Munch Tierarztl Wochenschr 2003;116(7–8):346–51. ¨ 12. Zeitz A, Spotter A, Blazyczek I, Diesterbeck U, Ohnesorge B, Deegen E, Distl O. Whole-genome scan for guttural pouch tympany in Arabian and German warmblood horses. Anim Genet 2009;40(6):917–24. 13. Dixon PM, McGorum BC, Railton DI, Hawe C, Tremaine WH, Pickles K, McCann J. Laryngeal paralysis: a study of 375 cases in a mixed-breed population of horses. Equine Vet J 2001;33(5):452–8. 14. Rooney JR, Delaney FM. An hypothesis on the causation of laryngeal hemiplegia in horses. Equine Vet J 1970;2:34–6. 15. Loew FM. Thiamin and equine laryngeal hemiplegia. Vet Rec 1973;92:372–3. 16. Duncan ID, Baker GJ, Heffron CJ, Griffiths IR. A correlation of the endoscopic and pathological changes in subclinical pathology of the horse’s larynx. Equine Vet J 1977;9:220–5.
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Genetic Basis of Respiratory Disorders 17. Duncan ID, Amundson J, Cuddon PA, Sufit R, Jackson KF, Lindsay WA. Preferential denervation of the adductor muscles of the equine larynx I: muscle pathology. Equine Vet J 1991;23:94–8. 18. Hillidge CJ. Prevalence of laryngeal hemiplegia on a Thoroughbred horse farm. J Equine Vet Sci 1985;5:252–4. 19. Lane JG, Ellis DR, Greet TRC. Observations on the examination of Thoroughbred yearlings for idiopathic laryngeal hemiplegia. Equine Vet J 1987;19:531–6. 20. Sloet-Oosterbaan MM, Gruys E, Barneveld A. Laryngeal hemiplegia in foals in relation to histopathological lesions of the laryngeal musculature and nervous tissues, and to vitamin E and selenium concentrations. In: World Equine Airways Symposium, Canada, 3–6 August 1998. 21. Dixon PM, McGorum BC, Railton DI, Hawe C, Tremaine WH, Pickles K, McCann J. Clinical and endoscopic evidence of progression in 152 cases of equine recurrent laryngeal neuropathy (RLN). Equine Vet J 2002;34(1):29–34. 22. Cahill JI, Goulden BE. Equine laryngeal hemiplegia I. A light microscopic study of peripheral nerves. N Z Vet J 1986;34:161–9. 23. Cahill JI, Goulden BE. Equine laryngeal hemiplegia IV. Muscle pathology. N Z Vet J 1986;34:186–90. 24. Brakenhoff JE, Holcombe SJ, Hauptman JG, Smith HK, Nickels FA, Caron JP. The prevalence of laryngeal disease in a large population of competition draft horses. Vet Surg 2006;35(6):579–83. 25. Galizzi VG, Galanti R. Contributo allo studio degli aspetti genetici del corneggio del cavallo. VII Congresso Naz Soc Ital d’Ippologia, Vieste 8: 1985; 108–12. 26. Poncet PA, Montavon S, Gaillard C, Barrelet F, Straub R, Gerber H. A preliminary report on the possible genetic basis of laryngeal hemiplegia. Equine Vet J 1989;21(2):137–8. 27. Ohnesorge, B. Hemiplegia larynges bei Warmblutpferden – eine Untersuchung an Hengsten, Stuten und deren Nachkommen. Vet Med Thesis, Veterinary School of Hannover, Germany, 1990. ¨ 28. Miesner K. Untersuchung uber die abduktorische Funk¨ tionsstorung des Kehlkopfes beim Pferd: Genetische und ¨ ¨ umweltbedingte Einflussfaktoren sowie mogliche zuchterische Massnahmen. Thesis of the Agricultural Faculty of the Friedrich-Wilhelms University, Bonn, Germany, 1996. 29. McClure JJ, Koch C, Powell M, McClure JR. Association of arytenoid chondritis with equine lymphocyte antigens but no association with laryngeal hemiplegia, umbilical hernias and cryptorchidism. Anim Genet 1988;19(4):427–33. 30. Woods PS, Robinson NE, Swanson MC, Reed CE, Broadstone RV, Derksen FJ. Airborne dust and aeroallergen concentration in a horse stable under two different management systems. Equine Vet J 1993;25(3):208–13. 31. Ewart SL, Robinson NE. Genes and respiratory disease: a first step on a long journey. Equine Vet J 2007;39(3):270–4. 32. Newton JR. Investigations of Respiratory Diseases of the Thoroughbred Racehorse, Diploma of Fellowship Thesis, Royal College of Veterinary Surgeons, London, 2003. 33. Robinson NE. Chairperson’s introduction: International Workshop on Equine Chronic Airway Disease, Michigan State University, 16–18 June 2000. Equine Vet J 2001;33:5–19. 34. Halliwell REW, McGorum PC, Irving P, Dixon PM. Local and systemic antibody production in horses affected with chronic pulmonary disease. Vet Immunol Immunopathol 1993;38:201–15. 35. Hare JE, Viel L, Conlon PD, Marshall JS. In vitro allergeninduced degranulation of pulmonary mast cells from
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42.
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44.
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horses with recurrent airway obstruction (heaves). Am J Vet Res 1999;60:841–6. Lavoie JP, Maghni K, Desnoyers M, Taha R, Martin JG, Hamid QA. Neutrophilic airway inflammation in horses with heaves is characterized by a Th2-type cytokine profile. Am J Respir Crit Care Med 2001;164:1410–13. Cordeau ME, Joubert P, Dewachi O, Hamid Q, Lavoie JP. IL-4, IL-5 and IFN-gamma mRNA expression in pulmonary lymphocytes in equine heaves. Vet Immunol Immunopathol 2003;97:87–96. Horohov DW, Beadle RE, Mouch S, Pourciau SS. Temporal regulation of cytokine mRNA expression in equine recurrent airway obstruction. Vet Immunol Immunopathol 2005;108:237–45. Ainsworth CG, Grunig G, Matychak MB, Young J, Wagner B, Herb AN, Antczak DF. Recurrent airway obstruction (RAO) in horses characterized by IFN-gamma and IL-8 production in bronchoalveolar lavage cells. Vet Immunol Immunopathol 2003;96:83–91. Pirie RS, Dixon PM, McGorum BC. Endotoxin contamination contributes to the pulmonary inflammatory and functional response to Aspergillus fumigatus extract inhalation in heaves horses. Clin Exp Allergy 2003;33:1289–96. Singh Suri S, Janardhan KS, Parbhakar O, Caldwell S, Appleyard G, Singh B. Expression of Toll-like receptor 4 and 2 in horse lungs. Vet Res 2006;37:541–51. Schaeper W. Untersuchungen ueber die Erblichkeit und das Wesen des Lungendampfes beim Pferd. Tieraerztliche Rundschau 1939;31:595–9. Cookson WO, Hopkin JM. Dominant inheritance of atopic immunoglobulin-E responsiveness. Lancet 1988;1(8577):86–8. Laumen E, Doherr MG, Gerber V. Relationship of horse owner assessed respiratory signs index to characteristics of recurrent airway obstruction in two Warmblood families. Equine Vet J 2010;42(2):142–8. Robinson NE, Berney C, Eberhart S, deFeijter-Rupp HL, Jefcoat AM, Cornelisse, CJ, Gerber VM, Derksen FJ. Coughing, mucus accumulation, airway obstruction, and airway inflammation in control horses and horses affected with recurrent airway obstruction. Am J Vet Res 2003;64(5): 550–7. ¨ Gerber V, Baleri D, Klukowska-Rotzler J, Swinburne JE, Dolf G. Mixed inheritance of equine recurrent airway obstruction. J Vet Intern Med 2009;23(3):626–30. ¨ ¨ ¨ Swinburne JE, Bogle H, Klukowska-Rotzler J, Drogem uller M, Leeb T, Temperton E, Dolf G, Gerber V. A wholegenome scan for recurrent airway obstruction in Warmblood sport horses indicates two positional candidate regions. Mamm Genome 2009;20(8):504–15. Jost U, Klukowska-Rotzler J, Dolf G, Swinburne JE, Ramseyer A, Bugno M, Burger,D, Blott S, Gerber V. A region on equine chromosome 13 is linked to recurrent airway obstruction in horses. Equine Vet J 2007;39(3):236–41. Watson JL. IL4R in the horse: SNPa of the tale. 5th International Equine Gene Mapping Workshop conducted by the Dorothy Russell Havemeyer Foundation at Kruger National Park, Pretoria- South Africa, 11–14 August 2003. Ober C, Leavitt SA, Tsalenko A, Howard TD, Hoki DM, Daniel R, Newman DL, Wu X, Parry R, Lester LA, Solway J, Blumenthal M, King RA, Xu J, Meyers DA, Bleecker ER, Cox NJ. Variation in the interleukin 4-receptor alpha gene confers susceptibility to asthma and atopy in ethnically diverse populations. Am J Hum Genet 2000;66:517–26.
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51. Neuhaus S, Bruendler P, Frey CF, Gottstein B, Doherr MG, Gerber V. Increased Parasite Resistance and Recurrent Airway Obstruction in Horses of a High-Prevalence Family. J Vet Intern Med 2010 Jan 22. [Epub ahead of print] 52. Eder C, Crameri R, Mayer C, Eicher R, Straub R, Gerber H, Lazary S, Marti E. Allergen-specific IgE levels against crude mould and storage mite extracts and recombinant mould allergens in sera from horses affected with chronic bronchitis. Vet Immunol Immunopathol 2000;73:241–53. 53. Eder C, Curik I, Brem G, Crameri R, Bodo I, Habe F, ¨ Lazary S, Solkner J, Marti E. Influence of environmental and genetic factors on allergen-specific immunoglobulin E levels in sera from Lipizzan horses. Equine Vet J 2001;33: 714–20. 54. Curik I, Fraser D, Eder C, Achmann R, Swinburne J, Binns ¨ M, Crameri R, Brem G, Solkner J, Marti E. Association between the MHC gene region and variation of serum IgE levels against specific mould allergens in the horse. Genetics, Selection, Evolution 2003;35:117–90. 55. Scharrengberg A, Gerber V, Swinburne JE, Wilson AD, ¨ Klukowska-Rotzler J, Laumen E, Marti E. IgE, IgGa, IgGb
and IgGT serum antibody levels in offspring of two sires affected with equine recurrent airway obstruction. Animal Genetics 2010 (accepted). 56. Palmer LJ, Silverman ES, Weiss ST, Drazen J.F. Pharmacogenetics of asthma. Am J Crit Care Med 2002;165:861–6. 57. Wade CM, Giulotto E, Sigurdsson S, Zoli M, Gnerre S, Imsland F, Lear TL, Adelson DL, Bailey E, Bellone RR, ¨ Blocker H, Distl O, Edgar RC, Garber M, Leeb T, Mauceli E, MacLeod JN, Penedo MC, Raison JM, Sharpe T, Vogel J, Andersson L, Antczak DF, Biagi T, Binns MM, Chowdhary BP, Coleman SJ, Della Valle G, Fryc S, Gu´erin G, Hasegawa T, Hill EW, Jurka J, Kiialainen A, Lindgren G, Liu J, Magnani E, Mickelson JR, Murray J, Nergadze SG, Onofrio R, Pedroni S, Piras MF, Raudsepp T, Rocchi M, Røed KH, Ryder OA, Searle S, Skow L, Swinburne JE, Syv¨anen AC, Tozaki T, Valberg SJ, Vaudin M, White JR, Zody MC; Broad Institute Genome Sequencing Platform; Broad Institute Whole Genome Assembly Team, Lander ES, Lindblad-Toh K. Genome sequence, comparative analysis, and population genetics of the domestic horse. Science 2009;326(5954):865–7.
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Disorders of the Bladder, Ureters and Urethra Thomas J. Divers
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Disorders of the Umbilicus and Urachus Kirsten M. Neil
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Chapter 69
Disorders of the Bladder, Ureters and Urethra Thomas J. Divers
Disorders of the bladder Ruptured bladder Ruptured bladder causing uroperitoneum is the most common disorder of the bladder in foals.1 The cause of the rupture in most cases is presumed to be a result of maternal pelvic pressure on the foal’s urine-filled bladder during parturition. The ‘tear’ in the bladder is most commonly in the dorsal wall, suggesting that this is the weakest area in the bladder muscle. Ruptured bladder at delivery may occur in both sexes but is more common in colts presumably because of their longer urethra and smaller internal pelvic diameter. Rupture of the bladder is also common in sick and recumbent neonatal foals because in some recumbent foals the bladder becomes overly distended when urinary catheters are not in place.2 In these cases, frequent lifting or moving of the foal with a distended bladder is thought to cause the rupture. Foals with neonatal encephalopathy may rarely have disruption of neurological micturition pathways resulting in bladder distension, stranguria, and eventually rupture (dummy bladders) unless they are catheterized. Urachal infection or necrosis at the apex of the bladder may also result in rupture and uroperitoneum3 and may be seen in foals as old as 3 months. Interestingly, some of these older foals with bladder rupture may not have the expected electrolyte abnormalities seen in neonatal foals with uroperitoneum, although they will all be azotemic. Likewise, foals that are being treated for encephalopathy, prematurity, or sepsis prior to developing a ruptured bladder may also have unremarkable electrolyte findings.4 Non-parturient trauma (i.e., a kick, or mare falling on her foal) may be a rare cause of bladder rupture in foals.
Clinical signs Progressive abdominal distension (ascites) and stranguria are the most common clinical signs, often first noted on day 2 or 3 of life in foals with presumed parturition-associated rupture. The initial tear to the bladder does not appear to cause noticeable discomfort to the foal unless the tear is associated with a urachal infection or a bladder hematoma is present. If the rupture has been present for 3 or more days, seizures may occur as a result of the severe hyponatremia or respiratory distress and colic may be found associated with severe ascites. Laboratory findings Foals with uroperitoneum characteristically have hyponatremia, hypochloremia, hyperkalemia, and azotemia with an increase in both blood urea nitrogen (BUN) and creatinine.1 Blood lactate may also be increased as a result of hypotension and poor perfusion of abdominal viscera. Differential diagnoses The most common differential diagnosis for the neonatal foal would be meconium impaction. Theoretically, foals with meconium impaction (also more common in colts) and tenesmus would have their hind legs more rostral than would foals with stranguria from a ruptured bladder, but this can be difficult to determine clinically especially in cases that have had multiple enemas (Figure 69.1a,b). Other differentials for stranguria in neonatal foals include subcutaneous rupture of the urachus (Figure 69.2)5 or bladder distension associated with neonatal encephalopathic syndrome (dummy bladder)
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 69.1 (a) A 3-day-old Thoroughbred colt demonstrating stranguria associated with a ruptured bladder. (b) A 2-day-old foal with tenesmus caused by meconium impaction. Notice the difference in hind limb placement in this foal compared with the foal in (a).
(Figure 69.3). On rare occasions, a neonatal foal may have a blood clot in the bladder or an emphysematous bladder causing dysuria. In older foals, dysuria can occur following a urachal abscess with adhesions keeping the bladder more ventral in the abdomen than normal. Rupture of one or both ureters can cause uroabdomen and electrolyte abnormalities similar to a ruptured bladder. Diagnosis The diagnosis of uroabdomen is made by clinical signs, laboratory findings, and sonographic evidence of large amounts of anechoic fluid in the abdomen (Figures 69.4 and 69.5). If there is a large amount of fluid in the abdomen, the small intestine may appear as numerous collapsed circles on the abdom-
inal sonogram. In some cases, the bladder will be inverted (Figure 69.6). Since the sonographic findings of uroabdomen and serum chemistries can be nearly identical with either ruptured bladder or ruptured urachus or ruptured ureter(s), the entire abdomen should be scanned to help rule out a ureter(s) or urachal rupture. Treatment The treatment of a ruptured bladder is surgical repair, although, rarely, this may heal on its own.6 Before anesthesia and surgery are performed, medical treatment to correct electrolyte abnormalities, especially severe hyponatremia and/or hyperkalemia, should be accomplished. Hyperkalemia and hyponatremia can be corrected by treating the foal with
0TCE/10/0MHZ
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P 100% G 100% D SCM FR 127 AUSONICS
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RUA
Figure 69.2 Prepucial and ventral body wall edema caused by subcutaneous rupture of the urachus. The foal responded well to urinary catheterization, tranquilization to decrease straining, and antibiotics. Some cases require surgical removal of the umbilicus and placement of subcutaneous drains.
Figure 69.3 Sonogram of a distended bladder in a foal with severe stranguria and neonatal encephalopathic syndrome (URA, urachus; RUA, right umbilical artery; LUA, left umbilical artery).
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Disorders of the Bladder, Ureters and Urethra
Figure 69.4 Depiction of ultrasound examination of a foal with uroperitoneum. Drawing by Kip Carter, MS, CMI and previously published in Urinary and Hepatic Disorders of Neonatal Foals, T.J. Divers, Clin. Techniques in Eq. Pract 2003, 2(1), p72.
dextrose and sodium bicarbonate. If the hyperkalemia is severe (> 8.0 mEq/L) and/or there are cardiac changes on an ECG (prolonged PR interval, spiked T wave, flat or absent P wave) calcium borogluconate (30 mL) should be administered with the first 0.5 L of fluids (e.g., 0.9% sodium chloride). If hyperkalemia and/or bradycardia are severe, care should be taken in placing a jugular catheter so as not to irritate the vagus nerve. Foals with severe abdominal distension and hyperkalemia should not be over-restrained or restrained in lateral recumbency for a prolonged time as this could further exacerbate cardiopulmonary dysfunction. If the hyponatremia is severe (< 115 mEq/L), the sodium should ideally be corrected slowly (2–3 mEq/L/h), but this is not always feasible in equine practice and I am not
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sure how important it is. In humans, too rapid correction of hyponatremia can cause irreversible central pontine myelosis. To my knowledge, this lesion has not been confirmed in foals and I believe most of the seizures in hyponatremic foals are due to the hyponatremia itself as opposed to rapid correction of the hyponatremia. Regardless, some consideration should be given to prevent rapid correction of the hyponatremia beyond 125 mEq/L. If the abdomen is tight, the urine should be drained with a teat cannula or drain prior to anesthesia (Figure 69.7). Surgery should be performed soon after draining the fluid, or subcutaneous fluid (urine) may accumulate at the tap site. Prolapse of the omentum through the puncture site is also common, and any prolapsing omentum should be cut and the site bandaged to help prevent further prolapse. There is no uniform agreement among surgeons as to whether to leave a Foley catheter in the colts (not fillies) for 2–3 days to keep the bladder from filling. I would recommend this only if a leak in the bladder occurs following the first surgery, necessitating a second repair (not common).
Prognosis The prognosis is generally excellent if surgery is performed within the first couple of days of life, the electrolyte abnormalities and abdominal distension are only modest, and the foal has no other illnesses (e.g., sepsis).
Prevention of bladder rupture There is nothing that can be done to prevent most ruptures associated with parturition. Critically ill
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Figure 69.5 (a and b) Sonograms of the abdomen of two foals with uroperitoneum showing the large amount of free anechoic fluid.
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Figure 69.6 ‘Inverted’ bladder observed on abdominal sonogram of a foal with uroperitoneum.
and/or recumbent foals (both colts and fillies) should have their bladder catheterized (Figure 69.8) to prevent distension of the bladder. If a recumbent foal is already noted to be urinating, it might not be necessary to catheterize the bladder. If this is the case, the bladder size should be frequently monitored by ultrasound. Foals with neonatal encephalopathy and dysuria due to interruption of normal micturition pathways should be catheterized and treated for hypoxia/ischemic encephalopathy until normal urination can occur. In some cases, this may take several days. Inflammatory diseases of the bladder (cystitis)
Causes and clinical signs Cystitis is rare in foals but may occur from either bacterial sepsis or secondary yeast infection following antibiotic therapy. Stranguria and pollakiuria would be the most common clinical signs although these are relatively mild with yeast cystitis. Viscous urine, often caused by a yeast infection (Candida spp.),
Figure 69.8 Cook urinary catheter used in critically ill foals. The catheter can be attached to a used fluid bag to both maintain a closed system and allow monitoring of urine value.
can cause obstruction of urine flow through a urinary catheter in critical care foals. Bacterial infections of the bladder may be the result of ascending infection associated with catheterization or from bacteremia. Dysuria and stranguria may be pronounced in those cases, even causing bladder prolapse in fillies. Bacteremia that lodges in the bladder may cause necrosis.
Diagnosis Diagnosis is made from the clinical history, clinical signs, urinalysis, and culture. Urinalysis results in foals with cystitis include pyuria and often hematuria for bacterial cystitis. Candida cystitis would cause pyuria and large numbers of yeast organisms on microscopic examination. Culture and antimicrobial sensitivity of a catheterized or midstream free flow urine sample for bacterial cystitis is recommended.
Treatments
Figure 69.7 Drainage of urine from the abdomen of a 4-day-old foal with severe abdominal distension and a serum potassium of 7.6 mEq/L.
Treatments for cystitis in foals should revolve around correcting any predisposing risk factors, e.g., improper catheterization, and appropriate antimicrobial treatments based upon culture and sensitivity of the offending organism in addition to consideration of the pharmacodynamics and safety of selected antimicrobials. Yeast cystitis usually resolves when antibiotic therapy is discontinued. Clotrimazole can be infused into the bladder if the yeast infection is causing obstruction of urine flow though a urinary catheter that must be maintained. Phenazopyridine (4 mg/kg p.o. q. 8–12 h) can be used to diminish
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Disorders of the Bladder, Ureters and Urethra discomfort caused by cystitis and/or indwelling urinary catheters. This drug will cause some discoloration of the urine. Catheterization of the foal bladder should be performed as a ‘sterile’ procedure and continuous drainage should be maintained in a closed system.
Disorders of the ureters There are three clinical syndromes involving ureteral dysfunction in neonatal foals. These are ectopic ureter, ureter rupture, and hydroureter.7
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tal to this. This same procedure could be attempted in foals with hydroureter(s). With either method, the prognosis is good for recovery from clinical signs and use as an athletic and/or breeding animal. If a nephrectomy is performed, the serum chemistry and ultrasonographic findings should be supportive of normal renal function. Following a nephrectomy, the serum creatinine often increases by approximately 0.5–0.7 mg/dL, but, owing to nephron hypertrophy (presumed) of the opposite kidney, the serum creatinine returns to nearly presurgical levels within 4–6 months.7 Ureteral rupture
Ectopic ureter Ectopic ureters are rare in the equine and often go unnoticed until the foal is several weeks old when urine scalding is noted on the perineum in fillies. It seems to be much more commonly recognized in fillies. Signs may be subtle if the ectopic ureter enters at the urethral sphincter. Endoscopy is the best method to determine whether an ectopic ureter is the cause of the incontinence and which side is affected (Figure 69.9). Endoscopy (cystostomy) of the bladder is easier to perform in foals that are several months of age. Surgical removal of the affected side (ureter and kidney) is the easiest method of repair as opposed to the more technically challenging ureteral transposition. Another option would be to create a hole in the ureter where it is seen coursing through the dorsal bladder wall and ligating the ureter just dis-
Rupture of the ureter(s) is diagnosed infrequently in 3- to 7-day-old foals. Both sexes are affected and in some cases the rupture is bilateral. The ‘tear’ is most commonly found in the proximal one-third of the ureter. The cause is unknown but may, similar to bladder rupture, be associated with parturition. Clinical findings and serum chemistries are similar to those found with a ruptured bladder or urachus except stranguria may be less pronounced and uroperitoneum may not be present in all cases if the peritoneal membrane remains intact. If the accumulation of urine is strictly retroperitoneal, the vulvar membranes of fillies may bulge externally as a result of the large amount of retroperitoneal urine.8 In most cases, the urine ruptures into the peritoneal cavity and may look identical to a ruptured bladder. In those cases, the diagnosis may not be discovered until the surgical approach for presumed ruptured bladder is performed. Treatment of a ruptured ureter requires surgical repair of the defect and ideally placement of a long ureteral catheter extending from the kidney and exiting the urethra. The catheters will frequently fall out in 1–3 days, but, during that time, may be helpful in maintaining consistent urine flow from the affected side(s). The prognosis is good for recovery if further leakage does not occur. In one case, leakage occurred postoperatively, but an abdominal drain was inserted and the defect apparently healed on its own in 2–3 days (Figure 69.10). Follow-up ultrasound of the kidneys should be performed weeks and months after surgery; one case of bilateral ureteral rupture that was surgically repaired developed hydronephrosis in one kidney as a 2 year old. Hydroureter syndrome in neonatal foals
Figure 69.9 Cystoscopic examination of a filly with ectopic ureter. Only the left ureteral opening could be seen in the bladder trigone area (arrow).
Although there are currently no scientific publications on this syndrome, I have seen and/or am aware of 15–20 foals that have hydroureters and clinical disease caused by the hydroureters. The cause of the syndrome is unknown but could be related to either
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Figure 69.11 Sonogram of distended ureter(s) in a 4-day-old foal with hyponatremia and encephalopathy (∗, kidney; arrows, dilated ureter; +, bladder). Figure 69.10 A 7-day-old filly with bilateral ureteral rupture causing uroperitoneum. The surgical repair has been completed and ureteral and urethral catheters can be seen exiting the vulva.
ureteral motility dysfunction or partial obstruction of the ureter at the bladder junction. I cannot find identical examples of this in literature reports on hydroureters in other species. The most common case history would be a 3- to 7-day-old foal that develops neurological signs suggestive of cerebral cortical disease (e.g., seizure, obtunded). Affected foals were generally healthy on days 1–3, and in most cases there was no history of recent illness in the mare or any problems during delivery.7 Hydroureters have been seen in both fillies and colts. This clinical syndrome with severe electrolyte abnormalities and neurological signs should not be confused on ultrasonographic examination with the mild to moderate ureteral and renal pelvic distension often seen in neonatal foals receiving large volumes of intravenous fluids. Clinical laboratory findings Severe hyponatremia and hypochloremia are consistently observed in the syndrome. The hyponatremia, which is frequently < 114 mEq/L, is presumed to be the cause for the neurological signs. In most all cases, the neurological signs are the first signs and are most likely the result of cerebral edema caused by the severe hyponatremia. Urinary fractional excretion of sodium and chloride can be very high at the time of clinical illness and may be caused by hydronephrosis and/or an abnormal aldosterone response. Hyperkalemia and azotemia are usually present, but often not as severe as seen in foals with ruptured bladders. Ultrasonographic examination reveals distorted ureters (usually bilateral) (Figure 69.11) and hydronephrosis. The distension of the ureters can be
in the proximal one-third (near the kidney) in some cases and throughout the ureter in other cases. Affected foals still void urine normally and the bladder appears normal. Differentials for the combination of severe hyponatremia and neurological signs include ruptured bladder or ureter(s), infectious diarrheal diseases, or rarely white muscle disease. There is a case report of transient pseudohypoaldosteronismin a 10-day-old foal.9 The salt-losing condition is due to renal medullary dysfunction (dilation and incomplete response to aldosterone; hence, pseudohypoaldosteronism). Most ill foals with various disorders have been found to have very high levels of aldosterone.10 Treatment and prognosis Supportive treatment that includes slow correction of the hyponatremia usually resolves the neurological signs. In a small number of cases, there may be spontaneous resolution of the hydroureter. Resolution may also be accomplished by an indwelling catheter placed in the ureters of fillies (this would be more difficult to do in a colt). Unfortunately, the ureteral distension may recur following removal of the catheter in most foals. The problem will resolve in a small number of affected foals, which suggests that the distension may be a developmental motility disorder. Until aldosterone has been measured in affected foals, it is not possible to know what role (primary or secondary) aldosterone abnormalities might play in the disorder. A brief treatment with a corticosteroid with mineralocoid properties could be attempted. Affected foals should be treated with antibiotics since hydroureters may predispose to urinary tract infection. Another unproven option would be to make a new opening of the ureter into the bladder at
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Disorders of the Bladder, Ureters and Urethra the site where the ureter first enters the dorsal bladder wall.
References 1. Richardson DW, Kohn CW. Uroperitoneum in the foal. J Am Vet Med Assoc 1983;182:267–71. 2. Kablack KA, Embertson RM, Bernard WV, Bramlage LR, Hance S, Reimer JM, Barton MH. Uroperitoneum in the hospitalised equine neonate: retrospective study of 31 cases, 1988–1997. Equine Vet J 2000;32:505–8. 3. Hyman SS, Wilkins PA, Palmer JE, Schaer TP, Del Piero F. Clostridium perfringens urachitis and uroperitoneum in 2 neonatal foals. J Vet Intern Med 2002;16:489–93. 4. Dunkel B, Palmer JE, Olson KN, Boston RC, Wilkins PA. Uroperitoneum in 32 foals: influence of intravenous fluid therapy, infection, and sepsis. J Vet Intern Med 2005;19:889–93.
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5. Lee MJ, Easley KJ, Sutherland RJ, Yovich JV, Bolton JR. Subcutaneous rupture of the urachus, its diagnosis and surgical management in three foals. Equine Vet J 1989;21(6): 462– 464. 6. Butters A. Medical and surgical management of uroperitoneum in a foal. Can Vet J 2008;49:401–3. 7. Divers TJ, Perkins G. Urinary and hepatic disorders in neonatal foals. Clin Tech Equine Pract 2003;2:67–78. 8. Divers TJ, Byars TD, Spirito M. Correction of bilateral ureteral defects in a foal. J Am Vet Med Assoc 1988;192: 384–6. 9. Arroyo LG, Vengust M, Dobson H, Viel L. Suspected transient pseudohypoaldosteronism in a 10-day-old quarter horse foal. Can Vet J 2008;49:494–8. 10. Hollis AR, Boston RC, Corley KT. Plasma aldosterone, vasopressin and atrial natriuretic peptide in hypovolaemia: a preliminary comparative stud of neonatal and mature horses. Equine Vet J 2008;40: 64–9.
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Chapter 70
Disorders of the Umbilicus and Urachus Kirsten M. Neil
Abnormalities of the umbilicus and urachus are common in foals, particularly neonates, with infection, herniation, and urachal patency among the most frequently encountered disorders. The external appearance of the umbilicus can be misleading; internal abnormalities cannot be excluded even if the external umbilical remnant appears normal. Likewise, enlargement or swelling of the umbilicus may be due to herniation, infection, hemorrhage, and edema and a definitive diagnosis may be difficult on physical examination alone. Ultrasonography is an extremely useful modality that enhances the diagnosis and management of affected foals in both hospital and field situations.
Normal anatomy and development The umbilicus comprises an external remnant and four internal remnants: the urachus, two umbilical arteries, and the umbilical vein. The internal remnants are contained within the umbilical or amnionic sheath.1 Early in embryonic development, the allantoic duct forms as a cranial continuation of the bladder,2 passing through the umbilical foramen and expanding external to the fetus to form the allantoic sac. This duct becomes the urachus, connecting the fetal bladder to the allantois.1, 2 When the umbilical cord ruptures after parturition, the urachus retracts within the amnionic sheath into the abdomen, and atrophies to become a disorganized mass of scar tissue (cicatrix) on the vortex of the bladder and the median ligament of bladder.1 The two umbilical arteries provide efferent blood flow from the internal iliac arteries of the fetus to the placenta. After parturition, the arteries retract up to 6 cm within the abdomen, and ultimately become the round ligaments of the bladder on its lateral
aspects.1 In the neonate, the round ligaments course within paired vesicular folds and retain narrow lumens through which some blood flow reaches the cranial portion of the bladder.2 A third vesicular fold supports the urachus in the fetus but is empty in the adult.2 The umbilical vein is the afferent blood supply from the placenta to the fetal liver and porta cava. After parturition, the vein regresses to become the round ligament of the liver along the edge of the falciform ligament.1
Ultrasonographic evaluation of the umbilicus Ultrasonographic visualization of the umbilicus from the ventral abdomen is simple and practical due to the close proximity of the internal umbilical structures to the skin surface. The normal ultrasonographic appearance of the umbilicus has been described.3–6 The examination may be performed with the foal standing or in lateral recumbency, and is usually well tolerated with minimal restraint and/or sedation. A 7.5–10.0 MHz sector scanner or a ≥5.0 MHz linear array transducer may be used, with or without a stand-off. A standard transrectal probe provides satisfactory images in the field. Clipping the abdomen and the use of coupling gel improves detail. While short-axis (transverse) views are the most useful, a longitudinal view is also helpful for assessing the attachment of the urachus to the apex of the bladder. To completely assess the umbilicus, both the external and the internal remnants need to be evaluated; each has a characteristic size and appearance. The normal external umbilical remnant contains hyperechoic structures, with the umbilical arteries and vein distinguishable based on wall thickness, and the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 70.1 (a) Ultrasonogram of a normal umbilical vein (arrow) in a 1-day-old foal. The wall of the vein is hyperechoic with an hypoechoic center. (b) Ultrasonogram of a normal umbilicus in a 4-day-old foal. Note the hyperechoic center (arrow) due to the presence of a blood clot.
urachus occupying the potential space between the arteries. These structures are visible for several days after birth but normally atrophy rapidly with the external remnant detaching from the underlying skin and by around 1–2 months of age are inconsistently visualized unless abnormal.3 The umbilical vein is a small, oval or elliptical structure with a thin echogenic wall and small anechoic fluid center that may vary in echogenicity3 (Figure 70.1). The vein should be followed from the external umbilical remnant cranially into the liver. The diameter of the vein varies along its length and is usually slightly larger at the external stump and just adjacent or caudal to the liver.3 Measurements should be made at at least two levels: just cranial to the external umbilical remnant and just caudal to its entrance into the liver3 (Table 70.1). Measurements can also be taken midway between the external remnant and the liver where the vein is typically slightly smaller,3 or at any other location along its length.
The umbilical arteries and urachus are examined together in the transverse plane from just behind the external remnant and followed caudally. Measurement of the combined cross-sectional area of the urachus and both arteries is made just cranial to the apex of the bladder3 (Figure 70.2). The arteries are small, thick-walled vessels and may contain sludging blood with pulsations detectable for up to 24 hours after birth.4 The center of the vein and arteries can vary in appearance from hypoechoic to echogenic (clot), and is typically hyperechoic by around 7 days of age.3, 4 The arteries should be followed individually along the lateral aspects of the bladder, and measured as caudal as possible.3 Additional measurements may also be taken in other locations between the external remnant and the caudal edge of the bladder.5 The urachus occupies the potential space between the arteries, does not normally contain fluid,3, 6, 7 and should be followed from the umbilical stump to the apex of the bladder. A longitudinal view of the urachus is useful to identify urachal patency
Table 70.1 Ultrasound measurements of normal and abnormal umbilical structures Umbilical vein
Source
Reef and Collatos (1988)3 Reimer (1998)7 Reimer (1998)7 Reef et al. (1989)8
External stump Normal 0.61 ±0.2 cm Abnormal >1 cm Mean 1.97 ± 0.3 cm
Umbilical artery
Mid-abdomen
Liver
Single artery
Urachus and both umbilical arteries
0.52±0.19 cm <0.8 cm
0.6±0.19 cm
0.85±0.21 cm <1 cm
1.75±0.37 cm <2.1 cm
>1.3 cm Mean 1.6±0.36 cm
>2.5 cm Mean 3.35±0.81 cm
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Foal: Urogenital System between 7% and 50% of foals had omphalitis.10–13 In an early study, 13% of septicemic foals had omphalitis,14 while more recently, only 5% of bacteremic foals had external signs of umbilical disease.15 However, when ultrasonographic examination of the umbilicus was specifically performed in foals with positive sepsis scores, bacteremia, or other foci of infection, abnormalities were detected in 57%.16 Signalment and clinical presentation
Figure 70.2 Ultrasonogram of a normal transverse view of the urachus and left and right umbilical arteries just cranial to the apex of the bladder. The left and right umbilical arteries are visible, and the urachus occupies the potential space between them (arrow).
and diverticulae. Slight dilation of the lumen of the urachus was reported in 6.2% of 145 clinically normal foals in one study.1 Little alteration in size or ultrasonographic appearance of the umbilical vessels was reported in a group of 13 clinically normal foals followed from birth to 4 weeks of age.3 In a larger group of foals, a significant reduction in mean umbilical vessel diameter was found over the first 7 days, except for the combined urachus/left and right umbilical artery diameter which remained unchanged.5 Repeated ultrasonographic examination of the umbilicus is useful, with alterations in size and appearance often indicating abnormalities such as infection.
Umbilical remnant infection Infection of the umbilical remnants (omphalitis) is a common problem in foals. Ultrasonography is an essential diagnostic modality as external abnormalities are not always apparent. Sonographic evaluation is recommended to determine whether one or more remnant is affected and the extent of involvement, with more specific descriptive terminology applied if the urachus alone (urachitis), umbilical vein (omphalophlebitis), or one or both umbilical arteries (omphaloarteritis) are infected. Incidence In an early study investigating joint ill and infections in nearly 2800 foals, the incidence of omphalitis was 0.21%.9 Omphalitis is a common concurrent disease process in foals with septicemia or other foci of infection, especially septic arthritis. In retrospective studies of foals with infectious orthopedic disease,
Omphalitis can affect foals of any age with the majority of foals afflicted less than 2 months old.8, 9 Rarely, older foals are affected, with omphalophlebitis reported in a 16-month-old colt.17 In two retrospective studies, the mean age of affected foals was 12.3 and 17.7 days.8, 18 Neonates comprised 24–31% of affected foals, with the majority between 8–30 days of age. External signs are variable and may include heat, pain, swelling, thickening, discharge, perinaval edema and/or urachal patency.18, 19 External signs are not always apparent. Fifteen of 16 foals had external abnormalities or urachal patency in one study; however, in another study less than 50% had external abnormalities visible.8, 18 Affected foals often present for other complaints.19 The umbilicus may be the primary source of infection or become infected secondary to dissemination of bacteria. Failure of passive transfer, septicemia, pyrexia, or other foci of infection, in particular septic arthritis or osteomyelitis, are common. Pneumonia, diarrhea, meningitis, hypopyon, and uveitis are other potential sequelae. Septic arthritis, septicemia, failure of passive transfer, vertebral body abscess and/or pneumonia were documented in 18 of 33 foals in one study.8 Although 30% of foals had joint ill and 27% were septic, there was no significant association between the presence of umbilical infection and septic arthritis or septicemia. However, in another study, 81% of foals had additional foci of infection, with septic arthritis documented in 43%.18 Omphalitis may also occur as a complication of other umbilical abnormalities such as trauma, hematomas, and urachal patency. Foals with a patent urachus present from birth are susceptible to secondary infection; conversely, omphalitis is the primary cause of acquired patency. Local extension of infection from the umbilicus can result in cystitis, peritonitis, adhesions, and hepatic abscessation.8, 20 An infected urachus can rupture causing uroperitoneum, and rarely urachal abscesses may rupture into the bladder. Pathophysiology Historically, the umbilicus was considered the main potential portal for entry of pathogens in foals with
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Disorders of the Umbilicus and Urachus septicemia18, 21 and septic arthritis,10 which led to the term ‘navel ill’ being used.9 Poor umbilical hygiene and environmental contamination are implicated as risk factors for the development of omphalitis. The time delay between separation of the cord at birth and complete atrophy of the remnants provides an avenue for bacteria to enter from the environment. A variety of postnatal treatments have been recommended to chemically cauterize the external umbilical remnant, including silver nitrate, concentrated phenol, 2% or 7% iodine, 1% povidone iodine, and 0.5% or 4% chlorhexidine.22, 23 Strong cauterants may irritate and damage surrounding tissue, predisposing to infection,24 abscess formation,18, 25 and increased risk of urachal rupture if omphalitis is present.26 Postnatal treatment has been extensively examined as a potential risk factor in human infants. Umbilical sepsis is relatively uncommon in the developed world, but endemic in less developed countries where it remains a leading cause of neonatal morbidity and mortality.27 Under conditions of good hygiene, topical antimicrobials or antiseptics have little benefit over daily cleaning and drying of the cord. However, in developing countries where the baseline risk of omphalitis is high, topical treatment (4% chlorhexidine) has been associated with reduced bacterial colonization rates,28 infection, and neonatal mortality.29 Chlorhexidine (0.5%) in 70% ethanol has also been used which has been shown to be effective in reducing bacterial colonization and bacterial dissemination in human infants.30 Less scrutiny of postnatal umbilical treatments has been made in foals. One study compared the effect of disinfectants on the normal bacterial flora of the umbilicus in 139 neonatal foals.31 The umbilicus was dipped from birth with either 2% or 7% iodine, 0.5% chlorhexidine diacetate, 1% povidone iodine, or no treatment applied. Treatment with 7% iodine and 0.5% chlorhexidine were the most effective in reducing the number of bacteria isolated; however, 40% of foals treated with 7% iodine later developed urachal patency, attributed to rapid desiccation of the umbilical stump and tissue necrosis of surrounding skin. Manipulation of the umbilical cord has been anecdotally implicated as a predisposing factor for the development of omphalitis.19, 25 Premature severance and ligation of the cord after birth may prevent retraction of umbilical vessels, increase the urachal lumen diameter, and delay time to closure. Causative organisms The most common bacteria isolated from the normal equine umbilicus at birth are coagulase-negative staphylococci (59%) and diptheroids (40%).31 A
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variety of organisms have been associated with omphalitis in foals, most of which are also associated with sepsis.8 The most common organisms isolated are Escherichia coli and beta-hemolytic Streptococcus spp.8, 9, 18 Mixed infections are common (50–68%).8, 18, 32 Other organisms commonly isolated include Klebsiella spp., Enterococcus spp., Actinobacillus spp., Salmonella spp., and Enterobacter spp.8, 18, 33, 34 Gram-positive organisms are isolated less frequently, as are anaerobes.8, 18 The predominant anaerobes isolated are Clostridium spp.8, 35, 36 While neonatal septicemia due to Listeria monocytogenes has been reported, isolation of this organism from the umbilicus has been rare.37 Diagnosis
Ultrasonography Ultrasonography is the most useful modality available to diagnose omphalitis, and is recommended when the external umbilical remnant is abnormal, and also in foals with septicemia, pyrexia, failure of passive transfer, leukocytosis, hyperfibrinogenemia, or other foci of infection. Abnormalities consistent with infection include enlargement of the affected structure due to perivascular thickening or enlargement with anechoic fluid, purulent echogenic material, and/or gas hyperechoic echoes, the latter suggestive of anaerobic infection8, 38 (Figure 70.3 and Table 70.1). Purulent material can vary in echogenicity, and fibrous tissue may be visualized associated with resolving infections. Edema or cellulitis may be seen in the surrounding tissue. Infection and abscessation of the umbilical vein tends to involve the vein in its entirety, and purulent material can often be seen extending into the liver (Figure 70.4). Encapsulation of a structure suggests abscessation (Figure 70.5), and may involve one or more structure.8 Visualization of an urachal lumen is often due to purulent material.38 More than one internal umbilical structure is usually infected8, 18 as they lie in close proximity and are bound within the amnionic sheath.1 In early retrospective studies of small numbers of foals with omphalitis, the urachus was most commonly infected with or without involvement of other structures.8, 18 Infection of one or both umbilical arteries was more common than infection of the vein.
Additional diagnostic tests Culture of infected remnants is recommended if there is discharge present or at surgery. The majority of umbilical cultures taken from foals with omphalitis are positive.8, 18, 32 Blood cultures should be performed in neonates, and when other foci of infection are present, appropriate fluid samples such as
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Figure 70.3 (a) Ultrasonogram of an infected umbilical vein in a 6-day-old foal. The vein is enlarged due to perivascular thickening and contains a small anechoic fluid center of purulent material. (b) Ultrasonogram of the umbilicus of a 5-day-old foal with omphaloarteritis and urachal patency. The right umbilical artery is enlarged due to infection, the left umbilical artery is of normal size, and the combined urachal/left and right umbilical artery diameter is enlarged. The lumen of the urachus (arrow) is visible due to the presence of purulent material and acquired urachal patency.
synovial fluid or peritoneal fluid should also be obtained for culture and cytology. As failure of passive transfer has been associated with omphalitis,8 the foal’s IgG level should be checked. A hematological profile should be performed, not only as a diagnostic aid but also to provide a baseline for monitoring response to treatment. Leukocytosis and neutrophilia are the predominant abnormalities seen, and hyperfibrinogenemia is common.32 A biochemical profile is recommended due to the propensity for other disease processes in affected
Figure 70.4 Ultrasonogram of an infected umbilical vein in a 6week-old foal. The vein was enlarged with an anechoic lumen cranial to the external umbilical remnant and was followed cranially into the liver. The vein was considered enlarged for a foal of this age and was of mixed echogenicity.
foals. Hepatic abscessation may result in elevations in liver enzymes, although this appears to be rare.22 Other acute phase inflammatory proteins such as serum amyloid A may also be useful. Treatment Antimicrobials are mandatory whether affected foals are treated medically or surgically. The overall health of the foal, the presence of septicemia or other foci of infection, intraoperative and postoperative risks, risk of bacterial shedding from the umbilicus, remnants involved and the extent of infection, and complicating factors such as hepatic abscessation, urachal patency or peritonitis should be considered. Omphalitis often resolves with medical treatment alone,21 and may be preferred if the lesion is small and well localized or when the foal is too debilitated for general anesthesia.39 Ultrasonography and hematology should be repeated to monitor the response to treatment. Failure to respond medically as determined by lack of reduction in size of infected remnants, increasing size or extension into other structures, persistent fever, persistent leukocytosis or hyperfibrinogenemia, and lack of clinical improvement are also criteria for surgical intervention. In some cases, a change in antimicrobial regime may result in improvement. Pending culture and sensitivity results the use of broad-spectrum antimicrobials (e.g., beta-lactam and aminoglycoside) is appropriate initially. If anaerobic infection is suspected, use of metronidazole is warranted. Most foals require antimicrobial treatment for
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Figure 70.5 (a) Ultrasonogram of the external umbilical remnant in a 2-week-old foal with an urachal abscess (arrow). The external umbilical remnant was palpably enlarged and thickened on palpation. The urachus is enlarged and the lumen of the urachus is visible with anechoic fluid representing purulent material. (b) Ultrasonogram of urachal abscess from the same foal as in Figure 70.5a. The image was obtained just caudal to the external remnant.
a minimum of 2–3 weeks.39 When long-term treatment is required, oral cephalosporin, chloramphenicol or oral trimethoprim sulfadiazine alone or with rifampin are useful.38 Topical disinfection is usually performed if the urachus is patent. Postnatal treatments include topical dilute povidone iodine, 2% iodine, 4% chlorhexidine, and 0.5% chlorhexidine in 70% ethanol.18, 19, 40 Surgery has been recommended based on ultrasonographic findings, with enlargement, involvement of multiple umbilical structures, and abscessation recommended criteria for surgical intervention.8 Surgery has also been recommended in foals with other foci of infection such as septic arthritis, with umbilical resection often performed at the same time as joint lavage. Surgical resection of the urachus and umbilicus has been described.18, 41 En bloc resection of the umbilicus is performed via laparotomy with the urachus removed at its junction with the apex of the bladder. Marsupialization of the umbilical vein is recommended when infection extends into the hepatic parenchyma, precluding removal of the vein in its entirety.18 A two-stage procedure has been described in calves42 and has been used in a foal.18 Using this technique, the vein is secured cranial to the celiotomy incision; however, a second surgery is required to obliterate the stalk. An alternative technique that does not require follow-up surgery involves mobilizing the vein so that it can be secured lateral to the celiotomy incision.43 Potential complications of both techniques include ascending infection, dehiscence, herniation, and cellulitis. Laparoscopic resection has also been described, with incision size, improved intraoperative visibility,
and reduced swelling and edema reported benefits.32 Eleven foals aged between 8 and 42 days (mean 17.5 days) were treated using this technique, including one foal which had had umbilical vein marsupialization performed. Ten of the 11 foals survived with few complications reported. Prognosis With early diagnosis and appropriate treatment, omphalitis is a manageable condition. The prognosis for affected foals depends not only on the extent of involvement of umbilical structures, but also on which concurrent disease processes or complications are present. Complications such as hepatic abscessation or the development of adhesions negatively influence survival. While only 50% of foals with hepatic abscessation survived in one study,8 other clinicians suggest that the prognosis is usually good.22 While there is no reported association between the structure affected and outcome, extensive abscessation of the umbilical vein requires more extensive surgical intervention and increases the likelihood of complications. Medical treatment avoids both intra- and postoperative risks, but the risk of dissemination of bacteria to other sites remains. While 66.6% of foals treated surgically survived compared to only 42.9% that received medical treatment alone in one study,18 similar studies have found no association.8, 16 Concurrent disease processes, notably septicemia and septic arthritis/osteomyelitis, greatly influence outcome; only 28.6% of foals with concurrent joint ill survived.18
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Patent urachus Urachal patency is a common disorder in neonatal foals.22 A patent or persistent urachus occurs when the urachus fails to close at or shortly after birth (congenital or primary) or when a previously closed urachus reopens, usually secondary to infection of the umbilical remnants (acquired, secondary). Pathophysiology Congenital patency has been proposed to occur due to partial obstruction of the umbilical cord from torsion in utero, resulting in dilation of the urachus and a delay in closure postpartum.1 Excessively long umbilical cords may be predisposed to excessive torsion during parturition and increase tension at the attachment to the abdominal wall,1, 24 which may compromise the urachus and umbilical vessels. Torsion may also lead to pooling of urine proximal to the site where the lumen of the urachus is constricted or obstructed.44 Overdistension of the urachus may prevent normal closure between the apex of the bladder and the urachus,1 or lead to rupture of the urachus or bladder or overdistension of the bladder in utero.1, 39, 44 Traumatic rupture of the umbilical cord postpartum may also prevent closure of the urachus,44 with early severance or ligation of the umbilical cord24 thought to predispose to opening of the urachus, especially after a ligature is removed.44 Acquired patency of the urachus is more common44 and may occur secondary to recumbency in critically ill foals or due to infection or inflammation of the urachus or umbilicus.18, 24 Urachitis or omphalitis should be suspected in all foals with acquired urachal patency. In one study 81% of foals with omphalitis had urachal patency.18 Acquired patency is not uncommon in septic foals, with the condition documented in 7% of bacteremic foals15 and 21% of foals with septicemia.14 Although omphalitis is the leading cause, increased recumbency, reduced movement, and lack of normal stance when urinating are contributing factors in debilitated foals.39, 45 A previously closed urachus may also reopen under conditions which increase the intra-abdominal pressure, such as in foals with tenesmus from a meconium impaction or dysuria from cystitis or uroperitoneum.44 Bladder catheterization may also cause inflammation or infection of the urachus and predispose to urachal patency 24–48 hours after the catheter has been removed. Diagnosis The diagnosis is usually based on physical examination, with characteristic clinical signs of urine drib-
bling from the urachus and umbilical stump. Urine may dribble continuously from the urachus or as a separate stream during micturition. The external remnant is usually persistently moist. Some foals strain to urinate due to urachal irritation and inflammation. Chronic cases may have urine scalding around the umbilicus. Some foals with concurrent omphalitis will have a grossly enlarged umbilicus, and there may be evidence of purulent discharge and urine from the external umbilical remnant associated with urachal fistulas.18 Ultrasonography should be performed in all foals with a patent urachus to determine if omphalitis is a complicating factor. Normally the urachus appears as a poorly defined echogenic structure between the two umbilical arteries, but may be fluid filled when patent3, 7 (Figure 70.3b). Identification of vesiculourachal patency confirms the diagnosis,46 and is best seen on a longitudinal view as a communication between the urachus and the bladder apex. Occasionally, a urachal diverticulum may be visible. Other modalities are usually not required to establish the diagnosis. Positive contrast radiographs can be used to identify a patent urachus as a triangular appearance to the apex of the bladder that may extend to the ventral body wall, and contrast may also be evident dripping from the umbilicus.
Treatment Treatment depends on whether the condition is congenital or acquired, and if there is evidence of omphalitis. Given time, most uncomplicated cases resolve without surgical treatment; congenital patency often resolves spontaneously. When there is no evidence of omphalitis, topical treatment and antimicrobials usually resolve the condition without the need for surgery.21, 45 Recumbent foals should be kept on clean dry bedding and have broad-spectrum antimicrobial coverage. Once these foals are more active, the urachal patency usually resolves associated with improved abdominal wall tone and more regular micturition.44 Straining associated with irritation of the urachus may be treated with phenazopyridine hydrochloride (200 mg p.o. q 8 h), a urinary tract analgesic that anesthetizes urinary tract epithelial surfaces.39, 44 Concurrent antimicrobial coverage is also recommended.39 Surgery may be required if there is concurrent omphalitis or if the urachal patency fails to respond medically within 5–7 days. Surgical management is essentially the same as described for omphalitis, with the urachus removed along with the umbilical vessels via a cystoplasty and modified midline celiotomy.41, 46
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Urachal rupture Although the bladder is the most common site of urine leakage in the neonate, rupture of the urachus has also been reported,26,47–50 and was the second most common site in retrospective studies of foals with uroperitoneum with between 22% and 39% of foals affected.47–50 Nearly half of the foals with urachal tears in one study had concurrent tears in the bladder wall,49 and more than one tear in the urachus has been detected at the time of surgery in some cases.47 Traditionally, uroperitoneum was associated with 1- to 2-day-old foals with bladder involvement, and 4- to 5-day-old foals if the urachus was involved, perhaps due to the thought that urachal rupture occurred secondary to infection and necrosis of the urachus.51 However, recent studies document a wider age range with foals presenting at an average age of 4.8 days49 and 6.2 days,50 and uroperitoneum documented in foals up to 42 days of age.50 Only three of the five foals with urachal tears in one study were less than 1 week and one foal with an urachal tear secondary to urachitis was 2 months old.47 Although foals with urachal tears were predominately male,47, 48 gender predilection has not been predominant in more recent retrospective studies of foals with uroperitoneum, although whether this applies specifically for foals with urachal tears could not be established.49, 50
Pathophysiology Urachal rupture has commonly been attributed to trauma or necrosis of the urachus secondary to infection. Hypoxic damage has also been speculated as a predisposing factor.52 Rupture of the urachus may occur during foaling if the umbilical cord is avulsed, which may be more likely if the mare foals while standing.51 Traumatic rupture of the umbilical cord has also been implicated.50 Tenesmus associated with gastrointestinal disease such as meconium impaction has been speculated as a contributing factor in the development of urachal tears.48 Urachal rupture due to congenital defects has not been reported in foals. Infection is being increasingly recognized as a cause of urachal rupture35,47–51 (Figure 70.6). Infection can lead to necrosis of the urachus and has been documented histologically from samples taken at surgery.49, 50 Urachal rupture secondary to infection may result in subcutaneous accumulation of urine rather than uroperitoneum18 and/or peritonitis.35, 47, 48
Figure 70.6 Postmortem specimen from a 10-day-old septic foal treated for omphalitis and septic arthritis. The foal developed uroperitoneum and septic peritonitis secondary to rupture of the infected urachus. The broken arrow depicts purulent discharge through the urachal tear. Cranial is to the right.
Diagnosis Clinical signs associated with uroperitoneum have been well described52 and include straining to urinate, decreased urination frequency and volume, weakness, tachycardia, tachypnea, excessive weight gain, and abdominal distension. Urachal tears that occur near or external to the abdominal wall may present with subcutaneous accumulation of urine with or without uroperitoneum51, 52 and may be confused with ventral edema.52, 53 Peritoneal tears can also result in fluid leaking into the subcutaneous space.6 Foals may also have signs of concurrent sepsis, omphalitis, urachitis, or patent urachus.47 Laboratory findings associated with uroperitoneum have been well described. Hyponatremia, hypochloremia, hyperkalemia, and increased serum creatinine concentration are common. A peritoneal:serum creatinine ratio >2 : 1 is considered diagnostic. Foals with urachal lesions had significantly higher peritoneal : serum creatinine ratios than those with bladder lesions in one study.49 Accumulation of subcutaneous fluid would also be expected to have a similar creatinine ratio as peritoneal fluid.51 Calcium carbonate crystals may also be visualized. Uroperitoneum is usually sterile unless infection preceded the urachal tear. Cytological evaluation and culture of peritoneal fluid is recommended when there is a suspicion of infection from the history or physical examination. Ultrasonography is the preferred diagnostic modality for uroperitoneum and can detect increased peritoneal fluid volume or on occasion a tear in the urachus.52 Observation of an intact fluid-filled
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bladder with fluid dissecting out along the urachus and into the retroperitoneal or subcutaneous space along the ventral abdomen is consistent with the diagnosis.4, 53 Urachal tears are not always visible sonographically.49 A ‘bubblegram’ is a useful extension of the ultrasonographic examination.46 Sterile saline is shaken and instilled into the bladder via a urinary catheter, and gas echoes can be visualized escaping from the bladder into the abdominal cavity. Other diagnostic modalities are used less frequently. Installation of new methylene blue dye or fluorescein into the bladder followed by abdominocentesis can aid in the diagnosis of uroperitoneum, but not in differentiating between the bladder and the urachus as the primary lesion.46, 52 Retrograde contrast cystography may be useful to diagnose uroperitoneum and establish the site of urine leakage.51 Treatment It is essential that foals with uroperitoneum are stabilized medically prior to surgical correction. Correction of electrolyte abnormalities, abdominal drainage, bladder catheterization and antimicrobials are the mainstay of medical treatment. Intravenous saline (0.9% or 0.45%) should be used initially, and intravenous glucose, sodium bicarbonate, calcium gluconate, and insulin can be used to correct life-threatening hyperkalemia. For foals with subcutaneous urine accumulation, conservative management with a Foley catheter may be useful for the short term; however, surgery is usually required if the tear is too large to heal spontaneously.51 Surgical correction via a cystoplasty has been well described.41, 46
risk,57 and Thoroughbreds at greater risk than Standardbreds.58 Umbilical hernias occur when the umbilicus fails to develop normally or fails to close during fetal development. The abdominal viscera are normally herniated through the umbilicus and contained in the extra-embryonic coelomic cavity, but return to the abdominal cavity later in gestation,2 leaving only a small space containing the umbilical cord.59 The linea alba forms as an aponeurosis of the abdominal musculature and fuses as a fibrous ring at the umbilicus.23 An umbilical hernia occurs when the peritoneal ring closes but the overlying abdominal musculature fails to close.2 A hereditary component has been proposed, with a significant difference in the prevalence of herniation in foals from different Warmblood sires in one study.60 In this study of 44 Warmblood foals monitored from birth, 43% had a defect palpable at birth.60 After 4 days, the defect was no longer palpable except in one foal. However, 12 foals later developed a hernia between 5 and 8 weeks of age, only 58% of which had a defect palpable at birth. The fibrous tissue at the umbilicus may soften or incompletely granulate after birth and possibly predispose to later herniation. Increased intra-abdominal pressure may also be a factor.60 Other predisposing factors for the development of umbilical hernias include omphalitis,56, 61, 62 trauma to the umbilical cord during birth, and excessive straining.23, 63, 64 In calves, umbilical infection greatly increases the risk of herniation.65 In equine studies, only 8%56 and 25%60 of foals treated for mild omphalitis developed herniation. Manually breaking the cord and cord ligation have also been implicated.63, 64
Prognosis Foals with uroperitoneum have a good prognosis, although a poorer prognosis has been documented when the lesion was in a location other than the bladder.47 Only 20% and 29% of foals with urachal lesions survived in two retrospective studies.47, 48 Nonsurvival was attributed to peritonitis, atresia recti or anesthetic complications. Concurrent sepsis has been associated with a poor prognosis in foals with uroperitoneum.49
Umbilical hernias Umbilical hernias are the most common type of hernia54 and the second most common congenital defect in the horse.55 Between 0.5% and 2% of foals have been estimated to have umbilical hernias.56 Females and Quarterhorses were considered at greater
Diagnosis Umbilical hernias are often asymptomatic and an incidental finding on physical examination.59 Digital palpation of the hernial ring is recommended to determine the size and shape of the defect, contents of the hernial sac, and reducibility of the hernia.54 Most are small, non-painful, and reducible.56 Small hernias are often considered as a cosmetic defect but are a potential site for incarceration. Hernial sacs usually contain subperitoneal fat, omentum or intestine,54 but as it is not always possible to differentiate between these contents on physical examination, ultrasonography is recommended. Ultrasonography is useful to exclude other causes of umbilical swelling such as omphalitis and umbilical abscessation, and is recommended prior to surgery or when complications are suspected.64
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Disorders of the Umbilicus and Urachus Foals, and especially 6- to 12-month-old horses with umbilical hernias present since birth, should always have the hernia carefully evaluated when there has been a history of colic or increase in size or swelling around the hernia. Firmness, warmth, pain on palpation, and a previously reducible hernia that becomes non-reducible are signs consistent with both complicated hernias (incarceration or strangulation), and abscessation.64 Differentiation between these conditions can be difficult. Leukocytosis and hyperfibrinogenemia would be expected to be diagnostic of abscessation; however, these changes can also occur in some foals with incarcerated and strangulated hernias.56 Horses with intestinal incarceration usually present for colic, with swelling of the hernia sac and pain on palpation. Incarceration may also occur in parietal hernias, and can progress to strangulation or enterocutaneous fistulas. Foals with strangulated umbilical hernias are usually pyrexic, tachycardic, and tachypneic.56 Signs of colic are not apparent in all cases or may be intermittent, especially if the lumen of the affected viscera is not completely occluded. An elevated peritoneal fluid white cell count and protein level can occur with strangulating hernias.56 Complications Complications of umbilical hernias occur less frequently in horses compared to cattle,63 and can include strangulation, incarceration, enterocutaneous fistulas, parietal (Richter) hernias, and abscessation. Complications have been reported in 2–10% of foals with umbilical hernias,20 and occur more often in foals 6 months or older with large defects.56 Incarceration is the most commonly reported complication. 20 The likelihood of intestinal incarceration increases the longer a hernia is present.66 Strangulation occurs when the blood supply to the incarcerated tissue is compromised. Strangulating umbilical hernias have been reported in foals ranging in age from 1 day to 30 months.56 The most consistent signs were increased size and firmness of the hernial sac with localized edema; signs of colic were evident in only 31%. Small intestine (jejunum or ileum) was most commonly strangulated, with ventral colon, cecum, and omentum less frequently affected. Parietal (Richter) hernias are defined as incarceration of only a portion of the intestinal wall; the antimesenteric surface of the ileum is most commonly incarcerated.66 Enterocutaneous fistulas occur infrequently associated with intestinal incarceration and parietal hernias.56, 67, 68 They can occur spontaneously or secondary to umbilical abscessation, with umbilical clamps and subcutaneous injection of irritant solutions around the hernial ring also implicated.20 Such
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fistulas develop after a period of incarceration leads to compromised vasculature to the entrapped intestinal wall and ischemic necrosis. Necrosis of the adjacent hernial sac leads to leakage of intestinal contents into the subcutaneous tissue, necrosis of the skin, and leakage of intestinal contents through the skin defect.66 Most affected horses have a history of increased swelling of the umbilical hernia in the 1–3 weeks prior to evidence of intestinal leakage and purulent discharge.67 Most do not recover spontaneously,67 with en bloc resection the preferred surgical technique.66 Occasionally, trauma to the umbilical cord during birth can cause evisceration immediately postpartum, necessitating immediate surgery or destruction.54 Treatment A variety of non-surgical and surgical treatment options have been described. Most umbilical hernias are small and resolve spontaneously by 12 months of age as the foal matures.60 Usual recommendations for reducible uncomplicated hernias with a diameter of <3 cm are for daily manual reduction of the hernia and close monitoring. If the hernia has not resolved by 4–6 months of age, surgical repair is recommended,54 although up to 12 months has also been recommended.62, 64 Larger defects carry an increased risk for complications such as incarceration, strangulation, and abscessation.54, 69 Such defects rarely close spontaneously and usually require surgery.64 Most umbilical hernias are repaired for cosmetic reasons.63 Non-surgical treatment options include hernial clamps and abdominal bandages.63, 64 Hernial clamps were used in 40% of all foals treated for umbilical hernias over a 3-year period at a university hospital.63 The aim of the clamp is to obliterate the hernia sac and stimulate healing of the abdominal wall defect. Infection and incarceration are contraindications for the use of clamps, which have been recommended in small (<5 cm), reducible hernias without evidence of infection.63 However, many surgeons do not recommend their use due to the possibility of complications such as accidental entrapment of tissue (e.g., omentum or intestine) in the clamp,54, 64 and rarely evisceration may occur.54 Abdominal bandages are usually not effective and are prone to slipping. Surgical repair of umbilical hernias has been described in detail.54, 59, 63, 64 The hernia sac may be inverted and the hernia repaired without removal of the hernia sac and entering the abdominal cavity (closed), or more often the hernial sac is incised along the edge of the hernial ring (open). Strangulating or incarcerated hernias and enterocutaneous fistulas should be corrected using an open technique to
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allow for evaluation of intestinal integrity, adhesions, and the need for resection and anastomosis.64 Enterocutaneous fistulas are repaired via an en bloc resection of the affected intestinal segment within the hernial ring. The preferred method of closure of the hernial ring is with a simple appositional closure for both open and closed herniorrhaphies.63, 64 Postoperative complications of herniorrhaphy may include incisional hernias, incisional infection, peritonitis, adhesions, and wound dehiscence with complications documented in up to 10% of cases.56, 63 Repair of large umbilical hernias in older foals and adults may require the use of prosthetic mesh to reduce tension,69 but this is contraindicated if there is evidence of infection. Potential complications of mesh herniorrhaphy include infection, adhesions, peritonitis and incisional drainage, with failure more likely if used in an infected site. Laparoscopic repair is usually not indicated.54 The prognosis for foals with umbilical hernias is good,69 even when complications are present, provided they are diagnosed early and treated aggressively.20, 56
Premature severance of the umbilical cord has long been viewed as a predisposing factor to the development of umbilical problems or to anemia in the neonate by preventing transfer of blood from the placenta to the foal immediately postpartum. However when the umbilicus was manually separated immediately postpartum in a group of neonates, there was no significant difference in hematological parameters or physical examination findings compared to a control group, suggesting that separation of the cord was not harmful.70 Umbilical hematomas usually present immediately after birth, and are a differential for sudden enlargement of the umbilicus, which is not usually warm or painful (Figure 70.7a). Ultrasonography is used to establish the diagnosis (Figure 70.7b).
Miscellaneous conditions of the umbilicus and urachus Urachal diverticula and cysts Other conditions associated with incomplete regression of the urachus include urachal diverticula that occur when the proximal portion of the urachal communication with the bladder persists. Diverticula can be diagnosed on a long-axis ultrasound view as a cranial outpouch of the apex and wall of the bladder.7, 38 Urachal diverticula may be an incidental finding but are usually associated with stranguria and occasionally predispose to the development of urachal patency.38 Urachal cysts may be associated with dysuria and occur when the proximal and distal ends of the urachus close but the mid portion remains open. Urachal cysts appear as sacculated structures on ultrasonographic examination. When only the umbilical opening of the urachus persists, an urachal sinus may form although this is infrequent.
(a)
Hemorrhage, hematomas, and seromas Trauma to the umbilical cord and vessels at birth or excessive stretching of the cord can lead to hemorrhage or formation of hematomas or seromas, overt hemorrhage from the umbilicus, and hemoperitoneum if the intra-abdominal portion of the umbilical artery or vein is affected. Foals with clotting disorders would be expected to display other signs consistent with abnormal bleeding.
(b) Figure 70.7 (a) Swollen umbilicus in a 12-hour-old foal that presented as a possible umbilical hernia. The foal had evidence of acute swelling shortly after birth, which continued to increase in size. (b) Ultrasonogram of the umbilicus in Figure 70.7a. An umbilical hematoma was diagnosed and confirmed by cytological analysis of an aspirate of the fluid. The foal was treated medically and the hematomas reduced in size over the next 7 days.
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Disorders of the Umbilicus and Urachus Hemorrhage may manifest as enlargement of the internal vessels, which may contain fibrin clots, and/or loculated fluid in the surrounding perivascular tissues.4 Ultrasonography can detect the occasional rupture of the umbilical vessel and hemoperitoneum, although abdominocentesis may be required to confirm the diagnosis. Damage to the umbilical vessels as well as the urachus can lead to retrograde bleeding into the proximal urachus and bladder. Cystic hematomas have been reported as a cause of hematuria, stranguria, and colic in foals.71 Affected foals had a history of excessive hemorrhage from the umbilicus at birth and ligation of the cord. Urachal and cystic hematomas were diagnosed sonographically, with blood clots within the bladder visible as echogenic masses. Trauma can lead to seroma formation, which appear as loculated anechoic fluid sonographically.7 It may be difficult to distinguish between abscessation, hematomas, and seromas since both blood clots and purulent material can range in echogenicity.38 Seromas usually contain sterile fluid, although this can become infected. Foals with excessive hemorrhage, hematomas, and seromas are usually placed on broad-spectrum antimicrobials as infection is a common sequelae.4 Hematomas and seromas are usually treated conservatively, and decrease in size within a few weeks.
Omphalocele An omphalocele is an uncommon congenital defect in the abdominal wall at the umbilicus through which viscera protrude into the base of the umbilical cord.72 While umbilical hernias have a hernial sac covered by overlying skin, an omphalocele is only covered by epithelium of the umbilical cord, i.e., amnion. These occur when the intestine fails to return to the abdominal cavity during fetal development. An omphalocele was diagnosed in an Arabian colt foal with a large abdominal mass noted at birth and surgically repaired.72
Other conditions Evagination of the bladder through the umbilicus has been described in a neonatal filly that presented with an umbilical mass and stranguria.73 The apex of the bladder had prolapsed through a proximal tear in the urachus. Absence of the urachus has been reported in a foal, resulting in anomalous fusion of the bladder to the umbilical foramen and the development of an enlarged bladder.74 Neoplastic conditions are extremely rare, with one case of an urachal cystadenoma in an older (2-year-old) filly.75
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Persistent attachments of viscera to the umbilicus or urachus are rare. Persistent attachment of an urachal remnant to the bladder has been described in a 15-month-old filly.76 Persistent vitelline circulation has also been reported. This circulation exists early in embryonic development to provide nourishment for the embryo from the yolk sac, and later regresses as the placental circulation predominates. The vitelline artery usually persists as the mesodiverticular band (Meckel diverticulum) and is usually associated with small intestinal volvulus, although an umbilical hernia containing the diverticulum has been reported in a 6-month-old filly with acute colic and swelling of the umbilicus.77 Persistence of the vitelline vein between the umbilicus and portal vein has been reported as a cause of intestinal strangulation in a 3-day-old foal.78
References 1. Whitwell KE. Morphology and pathology of the equine umbilical cord. J Reprod Fert Suppl 1975;23:599–603. 2. Dyce KM, Sack WO, Wensing CJG. The urogenital apparatus. In: Dyce KM, Sack WO, Wensing CJG (eds) Textbook of Veterinary Anatomy. Philadelphia: Saunders, 1987; pp. 162–204. 3. Reef VB, Collatos C. Ultrasonography of umbilical structures in clinically normal foals. Am J Vet Res 1988;49(12):2143–6. 4. Reef VB. Pediatric abdominal ultrasonography. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia: WB Saunders, 1998; pp. 364–403. 5. Lavan RP, Craychee T, Madigan JE. Practical method of umbilical ultrasonographic examination of one-week-old foals: the procedure and the interpretation of age correlated size ranges of umbilical structures. J Eq Vet Sci 1997;17(2):96–101. 6. Reimer JM. Neonatal sonography. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore, MD: Williams & Wilkins, 1998; pp. 73–8. 7. Reimer JM. The abdomen. In: Reimer JM (ed.) Atlas of Equine Ultrasonography. St Louis, MO: Mosby, 1998; pp. 180–99. 8. Reef VB, Collatos C, Spencer PA, Orsini JA, Sepesy LM. Clinical, ultrasonographic and surgical findings in foals with umbilical remnant infections. J Am Vet Med Assoc 1989;195(1):69–72. 9. Platt H. Joint-ill and other bacterial infections on Thoroughbred studs. Equine Vet J 1977;9(3):141–5. 10. Firth EC, Dik KJ, Goedegebuure SA, Hagens FM, Verberne LR, Merkens HW, Kersjes AW. Polyarthritis and bone infections in foals. Zentrlabl Veterinarmed B 1980;27:102– 24. 11. Firth EC, Goedegebuure SA. The site of focal osteomyelitis lesions in foals. Vet Q 1988;10:99–108. 12. Paradis MR. Septic arthritis in the equine neonate: a retrospective study. In: Proceedings of 10th American College Veterinary Internal Medicine Forum, 1992; pp. 458–9. 13. Smith LJ, Marr CM, Payne RJ, Stoneham SJ, Reid SW. What is the likelihood that Thoroughbred foals treated for septic arthritis will race? Equine Vet J 2004;36:452–6.
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14. Gayle JM, Cohen ND, Chaffin MK. Factors associated with survival in septicemic foals: 65 cases (1988–1995). J Vet Int Med 1998;12:140–6. 15. Corley KTT, Pearce G, Magdesian KG, Wilson WD. Bacteraemia in neonatal foals: clinicopathological differences between Gram-positive and Gram-negative infections, and single organism and mixed infections. Equine Vet J 2007;39(1):84–9. 16. White SL, Huff T. Retrospective study of surgical vs medical management of umbilical remnant infections in neonates. Paper read at 2nd Dorothy Havemeyer Foundation Neonatal Septicemia Workshop, November 1998 at Boston, MA. 17. Collatos C, Reef VB, Richardson DW. Umbilical cord remnant abscess in a yearling colt. J Am Vet Med Assoc 1989;195(9):1252. 18. Adams SB, Fessler JF. Umbilical cord remnant infections in foals: 16 cases (1975–1985). J Am Vet Med Assoc 197;190(3):316–18. 19. Nolen-Watson R. 2006. Umbilical and urinary disorders: umbilical infection/patent urachus. In: Paradis MR (ed.) Equine Neonatal Medicine: a case-based approach. Philadelphia: Elsevier Saunders, 2006; pp. 231–7. 20. Freeman DE, Orsini JA, Harrison IW, Muller NS, Leitch M. Complications of umbilical hernias in horses: 13 cases (1972–1986). J Am Vet Med Assoc 1988;192(6):804–7. 21. Sanchez LC. Equine neonatal sepsis. In: Sanchez LC (ed.) Neonatal Medicine and Surgery: Vet Clin North Am Equine. Philadelphia: Elsevier Saunders, 2005; pp. 273–93. 22. Divers TJ, Perkins G. Urinary and hepatic disorders in neonatal foals. Clin Tech Equine Pract 2003;2(1):67–78. 23. Smith M. Management of umbilical disorders in the foal. In Practice 2006;28:280–7. 24. Turner TA, Fessler JF, Evert KM. Patent urachus in foals. Equine Pract 1982;4:24–31. 25. Firth EC. Current concepts of infectious polyarthritis in foals. Equine Vet J 1983;15:5–9. 26. Ford J, Lokai MD. Ruptured urachus in a foal. Vet Med Small Anim Clin 1982;77(1):94–5. 27. Zupan J, Garner P. Topical umbilical cord care at birth. Cochrane Database Syst Rev 2004; (3):CD001057. 28. Meberg A, Schoyen R. Bacterial colonization and neonatal infections: effects of skin and umbilical disinfection in the nursery. Acta Paediatr Scand 1985;74(3):366–71. 29. Mullany LC, Darmstadt GL, Khatry SK, Katz J, LeClerq SC, Shrestha S, Adhikai R, Tielsch JM. Topical applications of chlorhexidine to the umbilical cord for prevention of omphalitis and neonatal mortality in southern Nepal: a community-based, cluster-randomised trial. Lancet 2006;367(9514):910–18. 30. Bygdeman S, Hambraeus A, Henningsson A, Nystrom B, Skoglund C, Tunell R. Influence of ethanol with and without chlorhexidine on the bacterial colonization of the umbilicus of newborn infants. Infect Control 1984;5(6): 275–8. 31. Lavan R, Madigan J, Walker R, Muller N. Effect of disinfectant treatments on the bacterial flora of the umbilicus of neonatal foals. Proceedings of the 40th Annual American Assoc Equine Practitioners conference, 1994; pp. 63–5. 32. Fischer AT Jr. Laparascopically assisted resection of umbilical structures in foals. J Am Vet Med Assoc 1998;214:1813–16. 33. Vaala WE, Clark ES, Orsini JA. Omphaloarteritis and osteomyelitis associated with Klebsiella septicemia in a premature foal. J Am Vet Med Assoc 1988;193:1273.
34. Patterson-Kane JC, Bain FT, Donahue JM, Harrison LR. Meningoencephalitis in a foal due to Salmonella agona infection. N Z Vet J 2001;49(4):159–61. 35. Hyman SS, Wilkins PA, Palmer JE, Schaer TP, Del Piero F. Clostridium perfringens urachitis and uroperitoneum in 2 neonatal foals. J Vet Int Med 2002;16:489–93. 36. Ortega J, Daft B, Assis RA, Kinde H, Anthenill L, Odani J, Uzal FA. Infection of internal umbilical remnant in foals by Clostridium sordelli. Vet Pathol 2007;44:269–75. 37. Monteiro F, Wong D, Scarratt WK, Buechner-Maxwell V, Grisman M. Listeria monocytogenes septicaemia in 2 neonatal foals. Eq Vet Educ 2006;18(1):27–31. 38. Reimer JM, Bernard WV. Abdominal sonography of the foal. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore, MD: Williams & Wilkins, 1998; pp. 627–36. 39. Wilkins PA. 2004. Disorders of foals. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis MO: Saunders, 2004; pp. 1381–440. 40. Elce YA. Infections in the equine abdomen and pelvis: perirectal abscesses, umbilical infections, and peritonitis. In: Southwood LL (ed.) Advances in Diagnosis and management of infection: Vet Clin N Am Equine. Philadelphia: Elsevier Saunders, 2006; pp. 419–36. 41. McIlwraith CW, Robertson JT. Surgery of the urogenital system. In: McIlwraith CW, Robertson JT (eds) McIlwraith and Turner’s Equine Surgery Advanced Techniques. Baltimore, MD: Williams & Wilkins, 1998; pp. 371–414. 42. Steiner A, Lischer CJ, Oertle C. Marsupialization of umbilical vein abscesses with involvement of the liver in 13 calves. Vet Surg 1993;22(3):184–9. 43. Edwards R, Fubini S. A one-stage marsupialization procedure for the management of infected umbilical vein remnants in calves and foals. Vet Surg 1995;24:32–5. 44. Vaala WE. The urinary system: patent urachus. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Williams & Wilkins, 1993; pp. 1032–5. 45. Russell CM, Wilkins PA. Evaluation of the recumbent neonate. Clin Tech Equine Pract 2006;5:161–71. 46. Lillich JD, Fischer AT Jr, DeBowes RM. Bladder. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St Louis, MO: Saunders Elsevier, 2006; pp. 877–87. 47. Richardson DW, Kohn CW. Uroperitoneum in the foal. J Am Vet Med Assoc 1983;182(3):267–70. 48. Adams R, Koterba AM, Cudd TM, Baker WA. Exploratory celiotomy for suspected urinary tract disruption in neonatal foals: a review of 18 cases. Equine Vet J 1988;20(1): 13–17. 49. Kablack KA, Embertson RM, Bernard WV, Bramlage LR, Hance S, Reimer JM, Barton MH. Uroperitoneum in the hospitalized equine neonate: retrospective study of 31 cases, 1988–1997. Equine Vet J 2000;32:505–8. 50. Dunkel B, Palmer JE, Olson KN, Boston RC, Wilkins PA. Uroperitoneum in 32 foals: influence of intravenous fluid therapy, infection and sepsis. J Vet Int Med 2005;19:889–93. 51. Hardy J. Uroabdomen in foals. Eq Vet Educ 1998;10(1): 21–5. 52. Wilkins PA, Dunkel B. Umbilical and urinary disorders: rupture of the urinary bladder. In: Paradis MR (ed.) Equine Neonatal Medicine: a case-based approach. Philadelphia: Elsevier Saunders, 2006; pp. 237–45. 53. Lees MJ, Easley KJ, Sutherland RJ, Yovich JV, Klein KT, Bolton JR. Subcutaneuous rupture of the urachus, its diagnosis and surgical management in three foals. Equine Vet J 1989;21:462.
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Disorders of the Umbilicus and Urachus 54. Stick JA. Abdominal hernias. In: Auder JA, Stick JA (eds) Equine Surgery, 3rd edn. St Louis, MO: Saunders Elsevier, 2006; pp. 491–9. 55. Preister WA, Glass WG, Waggoner NS. Congenital defects in domesticated animals: general considerations. Am J Vet Res 1970;31:1871–9. 56. Markel MD, Pascoe JR, Sames AE. Strangulated umbilical hernias in horses: 13 cases (1974–1985). J Am Vet Med Assoc 1987;190(6):692–6. 57. Hayes HM. Congenital umbilical and inguinal hernias in cattle, horses, swine, dogs and cats: risk by breed and sex among hospital patients. Am J Vet Res 1974;35:839–42. 58. Freeman DE, Spencer PA. Evaluation of age, breed, and gender as risk factors for umbilical hernias in horses of a hospital population. Am J Vet Res 1991;52(4):637–9. 59. Peyton LC. Surgical correction of equine umbilical hernias. Vet Med Small Anim Clin 1981;76(8):1212–15. 60. Enzerink E, Van Weeren PR, Van Der Velden MA. Closure of the abdominal wall at the umbilicus and the development of umbilical hernias in a group of foals from birth to 11 months. Vet Rec 2000;147:37–9. 61. Holt PE. 1986. Hernias and ruptures in the horse. Eq Prac 1986;8:13–16. 62. Hance SR, Debowes RM, Clem MF, Welch RD. Umbilical, inguinal and ventral hernias in horses. Comp Cont Educ Pract Vet 1990;12:862–71. 63. Fretz PB, Hamilton GF, Barber SM, Ferguson JG. Management of umbilical hernias in cattle and horses. J Am Vet Med Assoc 1983;183(5):550–2. 64. Orsini JA. Management of umbilical hernias in the horse: treatment options and potential complications. Eq Vet Educ 1997;9(1):7–10. 65. Steenholdt C, Hernandez J. Risk factors for umbilical hernia in Holstein heifers during the first two months after birth. J Am Vet Med Assoc 2004;224(9):1487–90.
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66. Baxter GM. Umbilical enterocutaneous fistula. Comp Cont Educ Equine 2007;2(2):96–9. 67. Bristol DG. Enterocutaneous fistulae in horses: 18 cases (1964–1992). Vet Surg 1994;23:167–71. 68. Rijkenhuizen ABM, Van Der Velden MA, Back W. Incarcerated umbilical hernia with enterocutaneous fistulae in two foals. Equine Vet Educ 1997;9(1):3–6. 69. Kawcak CE, Stachak MS. Predisposing factors, diagnosis, and management of large abdominal defects in horses and cattle. J Am Vet Med Assoc 1995;206:607–11. 70. Doarn RT, Threlfall WR, Kline R. Umbilical blood flow and the effects of premature severance in the neonatal horse. Theriogenology 1987;28(6):789–800. 71. Arnold CE, Chaffin MK, Rush BR. Hematuria associated with cystic hematomas in three neonatal foals. J Am Vet Med Assoc 2005;227(5):778–80. 72. Steinman A, Kelmer G, Avni G, Johnston DE. Omphalocele in a foal. Vet Rec 2000;146:341–3. 73. Textor JA, Goodrich L, Wion L. Umbilical evagination of the urinary bladder in a neonatal filly. J Am Vet Med Assoc 2001;219:953–6. 74. Dubs B. Megavesica due to the absence of an urachus in a newborn foal. Schweiz Arch Tierheilkd 1976;118(9): 393–5. 75. Gibson KT, Cantley CE, Donald JJ, Alley M. Urachal cystadenoma in a filly. Aust Vet J 1999;77(10):638– 40. 76. Dean PW, Robertson JT. Urachal remnant as a cause of pollakiuria and dysuria in a filly. J Am Vet Med Assoc 1988;192(3):375–6. 77. Hilbert BJ, Jacobs CV, Cullen LK. Umbilical hernia of a diverticulum of the vitelline duct in a horse. Aust Vet J 1981;57(4):190–2. 78. De Bosschere H, Simoens P, Ducatelle R. Persistent vitelline vein in a foal. Vet Rec 1999;145:77–9.
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Chapter 71
Renal Problems Jonathan E. Palmer
Mild renal dysfunction is common in neonatal foals who have suffered prenatal, intranatal or postnatal distress. Placentitis leading to fetal inflammatory insults and late-term hypoxic ischemic insults can induce FIRS (fetal inflammatory response syndrome). Intranatal hypoxia or asphyxia, neonatal sepsis or hypoperfusion, and anemia may all result in renal insults leading to acute kidney injury and decreased renal function. These insults can also lead to significant renal damage resulting in acute renal failure (ARF), but this is a much more uncommon outcome. Abnormalities of fluid and electrolyte balance are common in ill neonatal foals. Renal problems may be the underlying cause in some cases and may significantly complicate other cases. Aggressive fluid therapy required for emergency cardiovascular resuscitation and fluid therapy necessary for nutritional support are often complicated by inadequate renal function, leading to water and electrolyte imbalances. It is important to recognize renal abnormalities and understand the underlying pathophysiology so that a rational approach to therapy can be followed. This chapter will discuss important renal problems commonly encountered in equine neonatology, how to recognize them, and suggested approaches to therapy.
Birth transition It is important to understand some aspects of the birth transition undergone by the kidneys to understand neonatal renal function. The birth transition of all the organ systems begins days to weeks before birth and continues through the neonatal period (about 4 weeks). In fact the neonatal period is marked by this transition. Most of the information known about this transition is from studies in lambs and infants. Although there may be species differences, there are likely many more similarities which make this information useful in understanding neonatal foal renal function.
Fetal kidneys receive about 3% of the cardiac output, which translates into a low filtration rate.1 Just before birth, urine output decreases as fetal blood pressure increases.1 At birth the renal share of cardiac output rises quickly to about 15%, followed by a slower increase to 20%. Although some of this increase may be secondary to the rise in systemic blood pressure, most is from a decrease in renal vascular resistance.1, 2 There is also a redistribution of renal blood flow from the inner cortex to outer superficial cortex.1 The timing of the transition to the adult pattern varies with species, with no information specifically in the foal.2 Renal hemodynamics Renal hemodynamics during the transition is regulated by the interaction of angiotensin II, the renal sympathetic nervous system, circulating adrenergics, prostaglandins, the kallikrein-kinin system, nitric oxide, vasopressin, adenosine, atrial natriuretic factor, and endothelin.1 All of these factors have a somewhat different function and activity in the fetus and neonate than in the adult.2 In general, vasodilators increase renal blood flow and glomerular filtration, but because of the interaction of afferent and efferent glomerular arterioles and glomerular filtration, the relationship is not always straightforward. Angiotensin II, a major vasoconstrictor with its primary site of action the efferent arterials, will simultaneously decrease perfusion to the tubules as it increases glomerular filtration rate (GFR).2 The renal sympathetic nervous system has a dominant role in regulating renal blood flow during the transition. Circulating adrenergic agents and sympathetic tone increase vascular resistance and thus decrease renal blood flow more than in the adult. Angiotensin II is also a major reason for high vascular resistance during the neonatal period.2, 3 Intrinsic prostaglandins (PG) also have a more important
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Renal Problems role in neonatal renal hemodynamic regulation and function. PG is the most important vasodilator which counteracts the highly activated vasoconstricted state of neonatal microcirculation. Neonates treated with antiprostaglandin drugs will experience significant decrease in renal blood flow, increase in renal vascular resistance, and decrease in urine production.1, 3 Nitric oxide (NO) is also a major renal vasodilator during the neonatal period. The decrease in renal vascular resistance resulting in increased renal blood flow (increase in percent of cardiac output) is most likely from decline of vasoconstrictor forces in the face of maintained vasodilatory forces.2 The fetus and neonate are able to maintain a degree of autoregulation despite low blood pressure (mean arterial pressure 40–60 mmHg).2, 3 Autoregulation of GFR and renal blood flow appears to be a myogenic mechanism resulting in an intrinsic ability of arterials to constrict with high blood pressures and vasodilate with low blood pressures.3 In neonates the range of autoregulation is set to a lower perfusion pressure than adults, and renal pressure–flow relationship changes with renal maturation.4, 5 This reduction in autoregulation pressure range appears to be mediated through PG-dependent renin release causing vasoconstriction at lower levels of perfusion pressure.4, 5 Tubuloglomerular feedback occurs when macula densa cells sense changes in NaCl delivered to the distal tubules. With a drop in perfusion pressure there is less filtration, resulting in less NaCl delivery to the distal tubes. When this is sensed by the macula densa, angiotensin II is released from juxtaglomerular cells, resulting in constriction of efferent arterioles, thus increasing GFR.3 Maturation of tubuloglomerular feedback also occurs with growth and is maximally sensitive to a tubular flow that corresponds to the normal operating range. As GFR increases, maximum response and flow range also increases, so that relative sensitivity of tubuloglomerular feedback is unaltered during growth.2
Normal neonatal renal function Despite being born with an increased plasma creatinine level, foals appear to have a GFR nearly the same as adult horses by the time they are a day old. Many species have a slow increase in GFR to adult levels but, like the calf, the foal reaches the adult level soon after birth.6–9 Birth creatinine level in normal foals may range from 133 to 354 µmol/L (1.5 to 4.0 mg/dL). The creatinine level at birth is not a reflection of renal function but most likely originates from fetal fluid stores of creatinine transferred secondary to fluid shifts just before and during birth.10 At term in normal fetal
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foals the creatinine in the amniotic fluid averages 972 ± 442 µmol/L (11 ± 5 mg/dL) and allantoic fluid 13 170 ± 2475 µmol/L (149 ± 28 mg/dL).10 Foals suffering intrauterine distress are often born with much higher plasma creatinine levels. They usually range from 530 to 1800 µmol/L (6 to 20 mg/dL) but can be up to 3500 µmol/L (40 mg/dL) or higher. Again, these levels do not reflect renal function but may result from entrainment of creatinine secondary to shifting large amounts of fetal fluids into the interstitial reserves as a result of stress response of the fetal cardiovascular system.10 No matter what the cause of the high birth creatinine, the rapidity of drop to normal levels which usually occurs within 48 to 72 hours of birth, even when the creatinine value begins at a very high level, reflect efficacy of renal function and GFR level. Normal neonatal creatinine after this initial drop is usually < 100 µmol/L (1.1 mg/dL) and may be as low as 35 µmol/L (0.4 mg/dL). The final level is a balance between creatinine production from normal muscle metabolism and GFR. The blood creatinine will vary inversely with the muscle mass. Thus the cachectic, intrauterine growth-restricted foal or proportionally small premature foal will have lower levels and the well-muscled, well-grown foal higher levels. In normal Thoroughbred foals the creatinine level usually stays between 80 and 100 µmol/L (0.9 and 1.1 mg/dL) until they begin to have an increase in their muscle development. Unlike in other species, the first urine passed by the foal is usually delayed, not occurring until about 12 hours after birth. This first urine is very concentrated, having a specific gravity greater than 1.040. Urine in the urachus at birth is similarly concentrated, with mean creatinine level of 24 800 µmol/L (280 mg/dL). The reason for the delayed urination is low production of concentrated urine. After passing the first concentrated urine, the normal foal begins urinating every 1 to 2 hours, with each urination resulting in more dilute urine, and within a few hours the urine specific gravity falls below 1.005 and remains dilute. The underlying physiological mechanism leading to the concentrated urine during the perinatal period in the foal has not been studied. It may be secondary to the high vasopressin tone which is part of the normal cardiovascular birth transition in all neonates, but in other species the vasopressin does not lead to concentrated urine probably, because of a downregulation of renal V2 receptors. It is more likely that a more complex interaction of vasogenic mediators leads to a modification of afferent and efferent renal arterial tone, resulting in a transient decrease in GFR which is rapidly followed by a decrease in renal vascular resistance, leading to a rise in GFR to adult levels by the end of the first day of life.
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Table 71.1 Causes of acute renal problems in neonatal foals Classification
Renal disease
Examples of initiating factors
Glomerular dysfunction
Prerenal azotemia Neonatal Vasogenic Nephropathy
Septic shock Failure of birth transition Flunixin or Phenylbutazone Other nonselective and COX 2 selective NSAID Immune mediated diseases Sepsis – systemic or local
Glomerulonephritis Tubular dysfunction
Acute Tubular Necrosis Osmotic Nephrosis Renal Tubular Acidosis
Hypoxic ischemic insults Sepsis – systemic or local Synthetic colloids, mannitol Genetic defects Drug reactions: aminoglycoside, trimethoprim-potentiated sulfa drugs, NSAID, tetracycline Idiosyncratic Drug Reactions: Flunixin, other NSAID, antimicrobials Sepsis – local or systemic Immune mediated interstitial disease
Interstitial dysfunction
Interstitial Nephritis
Renal infectious insults
Infectious Nephropathies
Systemic sepsis Interstitial abscesses Pyelonephritis
Toxic nephropathy
Antimicrobial Nephropathies
Aminoglycosides Beta-lactams: cephalosporins, penicillins, carbapenems Trimethoprim-sulfa drugs Macrolides Fluoroquinolones Flunixin Phenylbutazone COX 2 specific inhibitors Furosemide
Drug Nephropathies
NSAID, nonsteroidal anti-inflammatory drugs; COX, cyclooxygenase.
Neonates need a positive sodium balance for growth, but fresh milk is sodium poor. Yet sodium absorption by the proximal tubule is not mature, resulting in a higher percentage of filtered sodium escaping proximal tubule reabsorption. The neonate makes up for this deficit by having enhanced distal tubule absorption, resulting in very efficient conservation of sodium as reflected by a very low fractional excretion of sodium.11 In normal neonatal foals the fractional excretion of sodium on day 1 of life ranges from 0.3% down to as low as 0.001% or lower. This enhanced ability to reabsorb sodium in the distal tubule results in blunted sodium excretion in the face of a sodium load. This partially explains why even moderate sodium administration can result in extracellular volume expansion and widespread edema12 in the neonatal foal.
ter and mature the distal sodium carrier mediated absorption responsible for sodium conservation by the neonatal kidney.13 However for the most part, fetal distress, intranatal distress and neonatal distress lead to failure of the birth transition and abnormal renal function which may progress to renal failure. Renal failure is characterized by a decrease in the GFR, resulting in an increase in the blood concentration of creatinine and nitrogenous waste products and by the inability of the kidney to appropriately regulate fluid, electrolyte and acid-base homeostasis. Acute renal failure (ARF) is an acute reduction in GFR with failure to remove solutes and water leading to concurrent net solute and water retention resulting in oligoanuric renal failure.14 Table 71.1 presents the causes of acute renal problems in foals. Glomerular disease
Renal problems Fetal stress and preterm birth may accelerate the renal transition.1 The accompanying cortisol surge may increase GFR and accelerate development of tubular reabsorption capacity for sodium, potassium, wa-
Decreased GFR may result from a loss of the number of filtering nephrons units or a decrease in rate of filtration in individual nephrons. Short of trauma or significant renal vessel thrombosis, loss of numbers usually does not lead to ARF although it may
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Renal Problems be important in chronic renal disease.15 Decrease in GFR in the acute situation is secondary to decreased GFR in each nephron, referred to as decreased single nephron GFR (SNGFR).16 Decreased SNGFR usually occurs because of decreased glomerular plasma flow, decreased glomerular transcapillary hydraulic pressure, increased plasma colloid osmotic pressure, or changes in permeability properties of the glomerular capillary.15
Neonatal vasogenic nephropathy Neonatal vasogenic nephropathy (NVN) is the most common cause of renal disease in the neonatal foal as marked by a slow decrease in birth creatinine, low volumes of concentrated urine, and water overload. This clinical term is used to describe the intrinsic renal vascular cause of decreased GFR.17 The most important factors in acute decrease in GFR in neonatal foals are decreased plasma flow rate and decreased transcapillary hydrostatic pressure. Decreased glomerular plasma flow rate may be caused by prerenal or intrarenal blood flow deficits. Even when the total renal blood flow is normal, there may be a vascular basis of decreased GFR. Autoregulation controls both afferent and efferent vascular tone, allowing for consistent GFR. With decreased renal perfusion the usual response is afferent dilation and efferent constriction, resulting in an unchanged GFR. Thus, in the face of decreased perfusion, autoregulation maintains GFR by maintaining transcapillary hydrostatic pressure with decreased renal vascular resistance on the afferent side but at times sacrificing tubular blood flow with increased efferent vascular resistance. Autoregulation is maintained effectively in neonates with physiological low blood pressure.18, 19 But when the low blood pressure is caused by pathological mechanisms such as sepsis or hypoxic ischemic insults, autoregulation may fail. Autoregulation frequently fails in the face of ARF.15 Angiotensin II causes vasoconstriction of efferent arterioles, an important part of autoregulation. But neonates, especially those with renal compromise, may be less responsive to vasoconstriction so sometimes, even though renal blood flow increases because of afferent vasodilatation, GFR may fail to increase as the efferent arteriole does not vasoconstrict. Neonates appear to have an abundance of angiotensin II receptors and other renin–angiotensin system components but also appear less responsive. This lack of responsiveness may be from a postreceptor insensitivity or different physiological actions mediated by the receptors, but also may be because of changes in vascular distribution caused by sepsis negating the expected response to receptor stimulation. In the normal fetus and neonate, angiotensin II may cause a vasodilatory response of the
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efferent arterials. This results in a disassociation of afferent dilation and GFR as, despite afferent vasodilatation, GFR may not increase as efferent vasodilatation will negate its effect.15, 17 These variations in the vasogenic response of sick neonates are the pathophysiological basis for NVN. The most common reason for decreased GFR in the neonatal foal is hypoperfusion of the glomerulus. This may be extrarenal, due to global hypoperfusion secondary to generalized cardiovascular insufficiency, or intrarenal due to local vasogenic issues. The former has been traditionally called prerenal azotemia and considered benign. The latter, due to intrarenal vasogenic responses, has been considered indicative of renal damage. However, clinical experience shows that there is no clear separation of these two pathogenic mechanisms in the patient, both often occurring simultaneously. Also, the usual stated reason for using clinical judgment to separate the two (e.g., the presence or absence of renal damage) does not follow uniformly. Fortunately it is rare for NVN to decrease tubular perfusion to the degree that tubular damage occurs, unless systemic factors such as septic shock result in global hypoperfusion. Many foals with intrinsic vasogenic decrease in GFR have no lasting damage once the vasogenic mishap has been corrected, whereas some foals with primary extrarenal hypoperfusion may have significant renal damage. With pathological stresses and increased adrenergic tone (increased renal sympathetic tone and circulating adrenergics) as may occur with sepsis or hypoxic ischemic insults, the resulting vasoconstriction will overwhelm the finely tuned regulatory balance of vasodilators (local prostaglandins, local NO) and vasoconstrictors (local adrenergics and angiotensin II), resulting in a decreased GFR and tubular blood flow. Any decrease in local prostaglandin production, as may occur intrinsically with failure of the birth transition or iatrogenically with the use of NSAIDs, may compound the problem. Neonates with renal injury or volume depletion have higher renal vascular resistance, resulting in lower GFR, and potentially may suffer more injury from lack of prostaglandins. In this instance even minimal decreases in perfusion could cause additional renal ischemia.
Glomerulonephritis Glomerulonephritis is a disease process that arises from damage to the glomerular filtration unit, resulting in reduce renal function. The initiator may be drugs, sepsis or immune-mediated diseases. The resulting inflammatory processes result in swelling of glomerular epithelial cells and a reduction in the intracellular gaps necessary for the filtration of plasma. Immune deposits along the basement
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membrane can form barriers to filtration. Antibodies may form complexes with circulating extrarenal antigens that deposit on the basement membrane, or antibody–antigen complexes may form at the level of the basement membrane. Antibodies may react with renal antigens or extrarenal antigens which collect in the glomeruli. The resulting damage to the glomeruli results in decreased GFR and glomerular leak of protein. Glomerulonephritis is an uncommonly recognized renal disease of neonatal foals. Tubular dysfunction Foals with NVN often have profound oliguria, infrequently producing small volumes of very concentrated urine with low sodium concentration. In fact oliguria is the most reliable early indication of NVN. Foals with oliguria have intact tubular function.16 Hypotension and stress-mediated vasopressin release results in transforming the distal tubule and collecting duct to their fully ‘water-permeable’ state. In general, the low flow in the tubules as a result of decreased GFR for any reason helps concentration mechanisms. Thus urine volume is minimized and concentration maximized. When tubules are injured, concentration is impaired and results in isosthenuric urine with a more normal volume. The foal with highly concentrated urine (urine specific gravity 1.020–1.040) and low urine sodium (fractional excretion Na < 0.3%) has normal tubular function but not necessarily normal renal function.16
Acute tubular necrosis Acute tubular necrosis (ATN) is the clinical term used to describe tubular dysfunction.16 It excludes renal disease caused by intrinsic vascular, glomerular or interstitial lesions.15 Interestingly, true necrosis of tubular cells is rare in this syndrome.16 Experimentally ATN can be induced after ischemia for at least 1 hour followed by reperfusion. After this severe insult, necrosis of tubular cells occurs in the outer medulla and in proximal convoluted tubules, areas with the highest oxygen demand and simultaneously limited perfusion reserve. The distal nephron is usually spared and only segments of the nephron are affected rather than an entire nephron. But clinically the term does not really describe a morphological change but rather a functional change. There may be minimal parenchymal compromise despite severe organ dysfunction. In sepsis, which is a common predisposing cause of ATN, endothelial dysfunction, coagulation abnormalities, and systemic inflammation all result in the renal dysfunction.16 Normal tubular epithelial cell function depends on defined apical and basolateral membranes. Integrins, which are membrane-spanning glycoproteins
on the basolateral cell surface, mediate tubular epithelial cell adhesion, defining the cellular orientation by anchoring the cell cytoskeleton. Ischemia resulting in ATP depletion causes integrins to loosen and relocate to the apical membrane, changing actin cytoskeleton orientation. This results in cellular rounding and detachment from the basement membrane. Some cells may be lost to the tubular lumen and adhere to each other by the apically displaced integrins, causing tubular obstruction, resulting in increased intratubular pressure and thus decreased GFR. Some of the luminal cells remain viable and may reattach. Ischemia also causes reorientation of Na+ ,K+ -ATPase from the basolateral membrane to the apical membrane, reversing sodium absorption, resulting in active secretion of sodium and increased sodium wasting, accounting for the very high urine sodium concentrations in some cases where urine sodium concentrations can be well in excess of plasma sodium concentrations. The appearance of higher concentrations of sodium in the distal tubule stimulates a vasoactive decrease in renal blood flow through the normal tubuloglomerular feedback mechanism.15 Also tubular injury will interrupt the structural integrity enough to disrupt the tight junctions, allowing backleak of creatinine from the tubular lumen to the blood. So some of the failure to clear creatinine is from damaged tubular integrity and not decreased GRF.20
Osmotic nephrosis Synthetic colloids, mannitol, and other hyperoncotic/hyperosmotic solutions have been associated with renal failure. Although the mechanism is not clearly defined and decreased glomerular filtration may be secondary to increased plasma oncotic pressure, osmotically induced tubular damage also occurs. Uptake of non-metabolizable molecules such as synthetic colloids by pinocytosis into proximal tubular cells generates an oncotic gradient, with swelling and vacuolization of tubular cells and tubular obstruction. Stimulation of the tubuloglomerular feedback due to the high distal solute delivery may also contribute to decreased GFR.21, 22 Neonatal foals suffering from NVN who have decreased filtration and thus slow movement of fluid through the tubules may be more susceptible to osmotic nephrosis. In such cases synthetic colloids and mannitol therapy should be avoided.
Renal tubular acidosis Renal tubular acidosis (RTA) is a collection of renal tubular disorders which characteristically cause a hyperchloremic (non-anion gap) acidosis without impaired glomerular filtration.23, 24
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Renal Problems Both genetic and acquired defects in proton and bicarbonate transporters with additional involvement of chloride and sodium transporters may be involved.24, 25 There are three types of RTA which have been described in humans: proximal RTA (type 2), distal RTA (type 1), and hyperkalemic distal RTA (type 4). A fourth type (heterogeneous RTA – type 3) had been reported but now is thought to be a combination of type 1 and type 2 RTA in the same patient and not a unique syndrome.26 Only proximal (type 2) and distal (type 1) RTA have been described in horses.27 In clinical equine neonatology RTA is a rare, sporadic disease occurring too infrequently to clearly characterize. Proximal RTA (type 2) is characterized by impaired proximal recovery of bicarbonate. This may be combined with other proximal tubular defects, such as the Fanconi syndrome (defective reabsorption of glucose, amino acids, electrolytes, and organic acids).23–26 Proximal RTA is characterized by bicarbonaturia, with a fractional bicarbonate excretion of greater than 15% when on bicarbonate replacement therapy.24, 25 In untreated cases the urine is usually acidic as plasma bicarbonate drops low enough for reabsorption to keep pace with the low filter load. Treatment may be difficult because administered base is often excreted at such a high rate that the desired normalization is not achieved. The acidosis in proximal RTA can be viewed in the conventional manner as loss of bicarbonate, or from the physicochemical approach to acid–base balance as the retention of chloride resulting in a hyperchloremic acidosis (decreased strong ion difference).24 Distal RTA (type 1) is characterized by impaired ability to acidify the urine in the distal tubules.23 With distal RTA, ammonium (NH4 + ) ions are not excreted in amounts adequate to keep pace with a normal rate of acid production.24, 25 Distal RTA is recognized by the inability to decrease urine pH below 5.5 in spite of metabolic acidosis and a low urine Pco2 after bicarbonate loading, indicating a lack of distal hydrogen ion secretion.24, 25 In humans, nephrocalcinosis and nephrolithiasis frequently occur secondary to this condition because of hypercalciuria. Hyperkalemic RTA (type 4) is a heterogeneous group of disorders that is characterized by failure to excrete acid and hyperkalemia usually associated with aldosterone deficiency or defective aldosterone signaling.23 RTA in humans occurs as a primary (persistent or transient) or secondary disorder. Secondary RTA occurs as a result of a great number of other diseases, exposure to certain drugs and toxins, a variety of genetic defects of carrier systems in the renal tubular cells, or structural disruptions of renal tubules caused by trauma or other primary re-
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nal diseases.24 Several drugs which may be used in foals have been reported to cause RTA including aminoglycosides, trimethoprim-potentiated sulfa drugs, carbonic anhydrase inhibitors, amphotericin B, non-steroidal anti-inflammatory drugs, tetracyclines (outdated or degraded tetracycline products), and lithium carbonate.26–28 The timeframe for development and recovery from drug-induced RTA is variable between individuals, but may begin within a week of exposure to the drug and can resolve as quickly as 3–4 days after discontinuing the drug.26 Foals with RTA usually present with lethargy, failure to thrive, growth retardation, generalized weakness, ataxia, anorexia, colic, constipation, tachycardia, tachypnea, polyuria, and polydipsia.25, 27 The signs may be quite vague. Any foal found to have a hyperchloremic acidosis (decreased strong ion difference, normal anion gap) without possible gastrointestinal or other origin such as treatment with large volumes of saline, should be suspected of RTA. To help support this diagnosis the blood creatinine should be normal and the urine strong ion difference (urine Na + urine K – urine Cl) should be positive (normal near 80). If the origin of the hyperchloremic acidosis is extrarenal and renal function is normal, then large amounts of chloride would be expected to be excreted in the urine with NH4 + to excrete the acid load. In the latter situation, because of the high concentration of chloride in the urine, the urine strong ion difference will be well below 80 and is in fact usually negative. Failure to excrete chloride in the urine in the face of acidosis is a central defect in RTA. If RTA appears to be present, a urine pH can help determine if it is proximal or distal disease. The pH should be measured on fresh urine using a pH meter, as dipstick readings are not reliable enough.25 In patients who have not been treated and have systemic acidosis (plasma HCO3 − < 15 mEq/L), a urine pH < 5.5 supports the diagnosis of proximal RTA. With distal RTA the urine would not be expected to be acid. Once placed on therapy (plasma HCO3 − > 22 mEq/L), a fractional excretion of HCO3 − > 15% supports the diagnosis of proximal RTA.28 The most important symptomatic treatment of RTA is correcting the acidosis. In distal RTA this is usually easily accomplished using 2–4 mEq/kg per day of bicarbonate. Proximal RTA is much more refractory, requiring up to 20 mEq/kg per day of bicarbonate or more.
Interstitial nephritis Clinical interstitial nephritis usually has a sudden onset with a rapid decline in renal function caused by acute renal inflammation. The disease is
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characterized by inflammation of the renal interstitium, interstitial edema, and damage to tubular epithelial cells. The reaction may be initiated by drugs (especially NSAIDs), sepsis, or immune-mediated disease. Idiosyncratic drug reactions may be responsible. In the author’s experience flunixin is the most common initiating drug in foals although antimicrobials may also be initiators. Usually only one or two doses of the drug are sufficient to induce the reaction. Withdrawing the drug is vital to successful recovery. Sepsis is also a common cause of interstitial nephritis in neonatal foals. Foals with interstitial nephritis may present with non-specific signs of systemic inflammatory response syndrome (SIRS) but will also have a rising creatinine, an elevated fractional excretion of sodium, hematuria, pyuria, mild proteinuria, and occasionally casts. Definitive diagnosis requires a biopsy.
Infections: systemic sepsis, interstitial abscesses, pyelonephritis Sepsis and severe septic shock promote renal failure through a variety of mechanisms. Hypotension and low cardiac output states result in decreased renal blood flow. Central venous pressure may rise as a result of cardiac dysfunction, or more commonly aggressive volume resuscitation, increasing central venous pressure, further reducing organ perfusion pressure. The vascular endothelium can be injured resulting in capillary dysfunction, the loss of NO-mediated autoregulatory mechanisms, and development of coagulopathy causing a diffuse microangiopathic thrombosis. Catecholamine levels are increased by both endogenous release and therapeutic infusions, resulting in decreased renal perfusion. The influx of neutrophils may result in free radical injury and release of cytokines. In addition, antimicrobials administered during sepsis may have nephrotoxic effects. Sepsis-induced ARF is multifactorial. As the individual response to sepsis is quite variable it is often difficult to predict if sepsis will lead to ARF or if there will be no renal compromise. Occasionally foals with bacteremia will develop miliary cortical abscess, especially those with Actinobacillus equuli infections. These infections can be manifested as an interstitial nephritis but rarely causes ARF. Pyelonephritis, especially ascending disease, is very uncommon in neonatal foals.
Toxic nephropathies Nephrotoxins can cause a variety of renal lesions such as acute tubular necrosis, acute allergic interstitial nephritis, thrombotic microangiopa-
thy, immunologically mediated damage, glomerular dysfunction leading to a nephrotic syndrome, altered intraglomerular renal perfusion, crystal accumulation, and osmotic nephrosis.29 A wide variety of antimicrobials and other drugs have been associated with renal toxicity including aminoglycosides, cephalosporins, penicillins, carbapenems, trimethoprim-sulfa drugs, tetracyclines, macrolides, fluoroquinolones, furosemide, and NSAIDs.29 The toxicity may be an idiosyncratic reaction independent of dose or may be associated with high doses. Although there are many nephrotoxins, the most common affecting neonatal foals are aminoglycoside antimicrobials and NSAIDs. Toxic effects of nephrotoxins (as well as ischemic effects) are not uniform throughout the kidneys. Nephrons and nephron segments, including the straight segment of the proximal tubule and the ascending limb located in the outer medulla, are especially susceptible to injury. This is likely related to the low oxygen tension in this region relative to the high workload and the lack of anaerobic adaptability of these segments. Nephrotoxic renal insults are more likely to result in acute renal failure with normal urine output as apposed to oliguria.15, 30 Non-steroidal anti-inflammatory drug toxicity may be a result of an idiosyncratic interstitial reaction or prostaglandin suppression resulting in decreased perfusion. Normally when renal perfusion is decreased, the afferent arteriole relaxes its vascular tone to decrease renal vascular resistance and maintain renal blood flow. Decreased renal perfusion results in increased catecholamine secretion, activation of the renin–angiotensin system and the generation of prostaglandins. During renal hypoperfusion, the intrarenal generation of vasodilatory prostaglandins, including prostacyclin, mediates vasodilatation of the renal microvasculature to maintain renal perfusion. Administration of NSAIDs can inhibit this compensatory mechanism and precipitate acute renal insufficiency with decreased GFR during renal hypoperfusion. The effect is usually transient and reverses once the NSAID clears. Selective COX-2 inhibitors are not renal sparing and can adversely affect renal hemodynamics similar to the effects of nonselective COX inhibitors.30, 31 Neonates are especially sensitive to the effects of NSAID as normal renal perfusion patterns are highly dependent on the presence of prostaglandins, even when global hemodynamics are normal. So even when normal foals are treated with NSAIDs there can be a decrease in glomerular perfusion as well as possible disruption of autoregulation and tubular-glomerular feedback. Because of this, NSAID use should be avoided whenever possible in neonates.
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Renal Problems Chronic failure Chronic renal failure is an uncommon sequela to acute renal failure in neonatal foals. A small percentage of foals who have acute renal insults may appear to regain renal function, as indicated by a creatinine which decreases almost into the normal range, only to have it increase again at 7 to 10 days of age. The creatinine may remain high for up to a year. Some affected foals will not grow well and have small fibrotic kidneys whereas others appear to recover; however, they probably have compromised renal function and even minor insults may lead to clinical renal failure.
Diagnosis Although renal failure has been defined as azotemia and oligoanuria, these clinical changes are also the hallmark of the normal renal response to hypovolemia.16 So, oliguria (defined as < 1 mL/kg/h)32 or an increase in creatinine or blood urea nitrogen (BUN) is not always diagnostic of renal disease.16 When diagnosing renal problems, laboratory findings are used to support historic and clinical findings and not as the final arbiter of the diagnosis. In fact, using laboratory findings without knowledge of the clinical condition may be misleading. Creatinine and blood urea nitrogen The most common measure of renal function is blood creatinine level. Creatinine is a metabolic product of creatine and phosphocreatine, both found almost exclusively in muscle. Thus, creatinine production is proportional to muscle mass. As a result, young foals, after clearing the increased birth creatinine, have normal creatinine levels between 80 and 100 µmol/L (0.9–1.1 mg/dL). Premature foals and foals with intrauterine growth restriction with a small muscle mass often have creatinine levels as low as 50 µmol/L (0.6 mg/dL) or even 35 µmol/L (0.4 mg/dL). As the foal matures and muscle mass increases, so will the creatinine level. Thus neonatal foals have a lower normal creatinine level than older foals. Creatinine is small, not bound to plasma proteins, and freely filtered by the renal glomerulus. It is also secreted by the renal tubule. Secretion is a saturable process that probably occurs via the organic cation pathway and is blocked by some medications including cimetidine, trimethoprim, pyrimethamine, and dapsone.33, 34 Normal foals placed on trimethoprimpotentiated sulfa drugs may have a 0–20% increase in blood creatinine levels without changes in GFR.33 The amount of increase is variable between individuals and may change with time. The percent of creatinine secreted appears to vary in the same individual over
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time, between individuals, and between laboratories depending on the analysis methodology.The proportion of creatinine secreted by the tubules increases as creatinine approaches normal, thus blood creatinine will not be sensitive to early decreases of GFR.35 The GFR may be reduced by 50% before serum creatinine increases above the normal range.3, 33, 35 Until recently most routine serum creatinine assays in current use had evolved from the reaction first described by Jaffe in 1886.36 The Jaffe reaction is a colorimetric method which detects creatinine and non-creatinine chromogens. Ever since its original description it has been constantly evolving through modification to decrease and limit the interference by many substances such as protein, bilirubin, glucose, pyruvate, acetoacetate, and cephalosporins.37 Although some current methods are not significantly affected by interfering substances, it is important that the practitioner is well aware of the accuracy limits to the technique used in reporting the creatinine level in their patients.37 Creatinine is distributed uniformly in body water and diffuses readily into the gastrointestinal tract. At normal creatinine levels this may not be clinically important but, as GFR decreases leading to volume overload, the creatinine will distribute into the retained fluid, dampening the effect of the decreased GFR on blood creatinine levels. Also if the foal has diarrhea some creatinine may be excreted through the gastrointestinal tract, again dampening the rise of blood creatinine in the face of decreased GFR. Changes in BUN levels are not a reliable measure of renal function in the neonatal foal. BUN originates from normal intestinal flora and liver catabolism of proteins. So it is not surprising that BUN levels are very low in fetuses and remain low in normal neonatal foals. Increases of BUN levels in the neonatal foal seemed to be closely correlated with the onset and degree of catabolism in critical neonates rather than a reflection of renal function, until renal dysfunction is marked. Clearance tests and fractional excretion indices Although blood creatinine is most commonly used to estimate GFR, clearance tests are more sensitive. Both endogenous creatinine clearance and inulin clearance have been described in neonatal foals.6, 7, 9 Neither is perfect. Creatinine clearance can be influenced by the tubular secretion of creatinine, resulting in an overestimate of true GFR. Inulin clearance can be inaccurate if sampling is taken before distribution is complete, perhaps explaining the inconsistent results between studies.6, 9 In addition, inulin may be redirected away from glomeruli in the neonate, making it
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Foal: Urogenital System Table 71.2 Renal diagnostic indices in neonatal foals Test
Normal value
Blood creatinine level
Birth Cr ≤ 350 µmol/L (4 mg/dL) Birth to 72 h – slow decrease from birth levels > 72 h to 2 wk ≤ 95 µmol/L (1.1 mg/dL) ≤ 2000 µmol/L (12 mg/dL) 1st urination > 1.030 Ad lib nursing < 1.005 Normal volume dependent on intake Oliguria < 1 mL/kg/h
BUN Urine specific gravity Urine output Creatinine clearance
1.46–2.15 mL/min/kg9 1.97–2.10 mL/min/kg6 2.8±0.55 mL/min/kg7
FxNa
≤ 0.3%
RFI (Renal failure index)
≤ 0.4
Urine Cr to plasma Cr ratio Excretion of Na
Mean 32, range 10–95 0.48 ± 0.24 mg/kg/day7
Formula
ClCr =
UCr × UVol /time/wt PCr
UNa P F xNa = Na × 100 UCr PCr UNa RF I = UCr PCr UCr PCr ENa = UNa × UVol /wt/time
Cr, creatinine; BUN, blood urea nitrogen; Clcr , creatinine clearance; Ucr , urine creatinine concentration; Uvol , urine volume; Pcr , plasma creatinine concentration; wt, bodyweight in kg; FxNa , fractional excretion of sodium; UNa , urine sodium concentration; PNa , plasma sodium; RFI, renal failure index; ENa , sodium excretion.
underestimate GFR.3 Clinically, creatinine clearance is convenient, as the only requirement beyond usual clinical practice is collection of all urine produced during the test period. However, as this usually requires an indwelling urinary catheter, it is invasive. Although it is better to avoid catheterization of the bladder in order to prevent nosocomial infection, it may be necessary in sick neonates. In cases with mild renal disease, creatinine clearance is accurate enough for clinical practice and can be quite helpful in following changes in renal function. Whereas estimates of GFR are used to monitor glomerular perfusion and function, monitoring sodium excretion is frequently used to monitor tubular function. Despite some limitations, the most useful clinical measurement of tubular sodium handling is fractional excretion of sodium (FxNa ). FxNa is the sodium clearance divided by the creatinine clearance which results in a measurement of tubular sodium transport normalized for GFR. The calculation of FxNa only requires simultaneous measurement of plasma and urine sodium and creatinine levels (see Table 71.2). In normal neonatal foals the FxNa should be less than 0.3%. Values greater than 1% suggest some degree of tubular dysfunction. The consistent fluid intake in the form of regular milk meals in the case of the ambulatory foal, or constant-rate infusion fluids in the case of the critically ill foal, make spot samples used for FxNa calculations more repre-
sentative than those in adults, where intake is more episodic. But there are a number of confounders. If the test urine is collected after rapid administration of sodium-containing fluids, as during fluid resuscitation in hypovolemia, after furosemide administration or in the presence of glucosuria, the resulting diuresis will result in a high FxNa despite normal tubular function. Foals placed on a high sodium diet (e.g., milk replacer) or those given large amounts of sodium in intravenous fluids or plasma will initially conserve sodium and maintain a low FxNa , but eventually will appropriately excrete excess sodium, resulting in a high FxNa . The transition from the low FxNa appears to be delayed in many critically ill foals. Some foals with extraordinary high FxNa values have disruption of the normal tubular cell orientation, resulting in active sodium excretion. These foals will have severe tubular disease but the measured fractional excretion value will not reflect the true FxNa . Despite its limitations, FxNa is a valuable clinical tool. The renal failure index (RFI) (see Table 71.2) is closely related to FxNa but removes the undue influence of plasma sodium in the situations when that value is well outside the normal range. Although not a percentage, the numeric value of the RFI is just slightly higher that the FxNa value. Another useful value which can be easily calculated using the information collected to measure FxNa is the ratio of urine creatinine to plasma creatinine. This value is a
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Renal Problems more accurate measure of urine concentration than estimates of urine specific gravity from refractometery. The most accurate way of determining sodium handling of the tubules is by measuring sodium excretion and calculating the sodium balance. In the critically ill neonatal foal which has dysmotility and thus is not passing feces, all of the sodium excretion occurs in the urine. By measuring the total urine output and concentration of sodium in the urine, sodium excretion can be derived. If the foal’s intake is controlled then the amount of sodium in the fluids, parenteral and enteral nutrition, and medications can be compared to the sodium excretion. A negative sodium balance in the absence of earlier sodium overload is strong evidence of a sodiumwasting nephropathy. Urinalysis Careful attention to the results of serial urinalysis examinations can be helpful in characterizing renal abnormalities in foals. Urine concentration is a useful clinical reflection of renal function. Urine concentration is best estimated by measuring urine osmolality, which depends on the concentration of all particles in the urine.38 However, in routine clinical practice urine specific gravity, which correlates with concentration, is estimated by urine refractive index.38, 39 Unlike osmolality, specific gravity and refractive index are dependent on the size (refractive index) and weight (specific gravity) of the particles in the sample as well as their concentration. As refractive index is more heavily influenced by size of the particles than their total weight or numbers, it should be noted that the presence of large molecules such as glucose or protein will increase the refractive index disproportionately to the urine concentration, whereas small molecules such as electrolytes will have almost no influence on refractive index, resulting in a low estimation from refractive index of the true urine concentration. Despite these limitations, refractive index-derived specific gravity is a useful clinical estimate of urine concentration reflecting renal function.38–40 The first urine produced by normal foals, which is usually passed at about 12 hours after birth, normally has a specific gravity > 1.030. Each subsequent urination during the first day, which usually occurs at about 2-hour intervals after the initial urination, will have a lower urine specific gravity until it is < 1.005. Normal foals consuming 10–25% of their bodyweight in mare’s milk per day will have urine specific gravity values 1.005–1.000. Foals receiving milk replacer will have slightly higher urine specific gravity values as they excrete extra solutes consumed in the replacer. The urine specific gravity of foals which are
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ill will depend on their underlying disease(s) and the therapy they are receiving. For example, critically ill foals with normal renal function receiving the author’s formulated maintenance fluid therapy41 will have a urine specific gravity ranging between 1.005 and 1.012. Foals on high levels of medications which are excreted in the urine, such as antimicrobials, may have slightly higher values (1.008–1.018). Foals with normal renal tubular function and appropriate antidiuresis (as from dehydration) or inappropriate antidiuresis (as from NVN or SIADH [syndrome of inappropriate antidiuretic hormone secretion]) may have values ranging from 1.020 to > 1.040. Foals with abnormal renal tubular function may have values in the isosthenuric range of 1.009–1.012. Taking into account the clinical presentation, urine specific gravity is a simple and inexpensive test can be very useful in investigating renal function in the neonatal foal. Urine pH is primarily helpful when investigating abnormal renal tubular function as occurs in renal tubular acidosis. In general, normal foals are born with aciduria which quickly changes to alkaline urine as they become anabolic. Foals which are catabolic secondary to inflammatory disease or lack of caloric intake have acid urine. But once they become anabolic their urine usually becomes alkaline associated with their recovery. During the first 48 hours of life, normal foals often have blood or protein in their urine, as indicated by a positive dipstick reaction. Persistence beyond that period may indicate either lower or upper urinary tract disease. Glucosuria associated with hyperglycemia is common in treated neonatal foals but glucosuria in the face of euglycemia can indicate tubular dysfunction such as acute tubular necrosis or Fanconi syndrome. Foals with normal tubular function and decreased GFR, such as occurs in NVN, may not spill glucose despite extreme hyperglycemia since there is ample opportunity for complete glucose absorption, as the small amount of filtered urine moves slowly through the tubules. The ketone reaction on the urine dipstick is seldom useful in neonatal foals but it is well to note that very high levels of ceftiofur administered intravenously can result in a false-positive ketone reaction. Examination of urine sediment can be very useful in detecting and characterizing renal disease. Sediment from 10 mL of urine should be examined, taking into account the urine concentration in its interpretation. The differential diagnosis of hematuria can be categorized as originating in the upper or lower urinary tract. Hematuria that is accompanied by red blood cell casts, dysmorphic red cells or marked proteinuria is most likely to be glomerular in origin. In the absence of these findings, distinguishing glomerular from post-glomerular bleeding can be
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difficult.40 White blood cells can enter the urine from anywhere along the excretory system. The presence of proteinuria and casts suggests a glomerular source. In the absence of other formed elements, the clinician must look beyond the urine sediment for additional clues to find the origin of urine leukocytes. There are no effective methods to identify the origin of white blood cells found in the urine. Most leukocytes in the urine are polymorphonuclear, but they are not all neutrophils as eosinophils may be present, especially associated with drug-induced interstitial nephritis. When examining the urine for casts it is ideal to examine the sediment within 10 minutes of collection, and the urine specific gravity and pH should be taken into account. As formed elements are more stable in concentrated and acidic urine than in dilute and alkaline urine, the finding of a few casts in the latter type of sample is as significant as finding many in the former. Casts are cylindrical bodies several-fold larger than leukocytes and red blood cells. They form in distal tubules and collecting ducts where TammHorsfall glycoprotein precipitates and entraps cells present in the urinary space. Dehydration and the resulting increased tubular fluid concentration favor the formation of casts. An acid urine is also conducive to cast formation. Observing casts in the urine sediment often provides helpful diagnostic information. Hyaline or finely granular casts can be seen in normal individuals but also may be present in the face of disease. Cellular casts are generally more helpful. Red blood cell casts, for example, are distinctive and most often indicate glomerular disease. White blood cell casts are most commonly associated with interstitial nephritis but can also be seen in glomerular nephritis. Casts made up of renal tubular epithelial cells are always indicative of tubular damage. Coarsely granular casts often result from the degeneration of cellular casts. They also contain protein aggregates. Thus, the presence of granular casts implies pathology but not the specific location of the problem. Waxy casts are also non-specific. They are believed to result from the degeneration of cellular casts and, thus, can be seen in a variety of kidney diseases. The presence of cellular casts confirms renal injury but, since neonatal foals usually have very dilute, alkaline urine, the absence of casts or other formed elements does not rule out renal disease.40
Therapy Unfortunately there is no therapy that has been found to reverse acute renal injury.42 Management of the neonatal foal with acute renal injury is largely supportive, removing all adverse conditions and minimizing the demands on the kidneys while allowing time for recovery to occur.42 Although the presence of oliguria may be an indication of serious renal disease,
conversion of oliguria to a diuresis will not decrease the renal injury or speed recovery.42 Even mild renal compromise may be associated with the potential for complications, such as volume overload due to impaired water excretion (decreased GFR), disrupted acid–base homeostasis due to impaired strong ion regulation, electrolyte disorders (hyponatremia, hyperkalemia), retention of uremic solutes, and impaired elimination of a variety of other potential toxins such as drugs and their metabolites. Volume overload may be further aggravated by the use of high fluid infusion rates in the face of oliguria. In any neonatal foal which shows indications of renal compromise, such as the slow drop of birth creatinine to normal levels indicating reduced GFR, nephrotoxic drugs should be avoided. It is especially important to avoid the use of NSAIDs in these cases as often the underlying problem is NVN, which will be exacerbated by decreasing renal prostaglandin levels. Fluid balance/Fluid therapy The most crucial aspect of therapy for foals with renal compromise is the careful management of fluid balance. If there is any clinical indication that poor perfusion may be playing a role in decreased renal function, as indicated by oliguria, a test fluid bolus of 5–10 mL/kg over 30 minutes should be given. If there is a positive response then careful attention to optimizing global perfusion is in order. If there is no response to the fluid challenge or if it is clear that global perfusion is normal, making it evident that there is an intrinsic renal problem causing the decreased GFR, it is not rational to continue high fluid infusion rates.42 In fact, if there is evidence of significant fluid overload such as substantial weight gain or edema, serious consideration should be given to fluid restriction. As long as normovolemia and perfusion are maintained, decreasing fluid infusion rate to half or less of the calculated maintenance rate is usually well tolerated and helpful in controlling the fluid balance. Following the bodyweight by measured weights once or twice a day is much more valuable in assessing fluid balance than traditional methods of estimating hydration state by physical findings. In fact physical findings in neonates are often misleading when trying to estimate hydration status. When possible, tracking intake and output can be equally valuable. The common practice of delivering increased fluid volumes in the face of decreased GFR should be avoided, unless the underlying problem is decreased global perfusion (prerenal azotemia) or the renal failure is manifested as a high output problem.42 If the working diagnosis is NVN in an oliguric foal with significant fluid overload, other approaches to therapy with the aim of increasing glomerular perfusion include an inotrope trial, dopamine trial, and
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Renal Problems furosemide trial. The conversion of oliguric to nonoliguric acute renal failure has not been shown to alter the course of acute renal failure.42, 43 But inducing a diuresis in an oliguric foal will help manage the fluid overload while waiting for the renal function to improve with the spontaneous resolution of the renal disease. Before inducing a diuresis, cystomegaly should be ruled out as the cause of decreased urine output, by abdominal palpation or abdominal ultrasonography. Even if the clinical examination of the foal suggests adequate perfusion, an inotrope trial with dobutamine may be rewarding. If a diuresis does not occur with the dobutamine trial and there is no other clinical indication for inotropic support, the dobutamine infusion should not be continued as its alpha-adrenergic properties could potentially cause afferent renal arterial vasoconstriction, exacerbating the vasogenic nephropathy. Dopamine is another rational therapeutic choice. It may cause renal vasodilatation counteracting the vasogenic basis of the nephropathy as well as act as an inotrope, and could induce a diuresis. It also has tubular effects which may contribute to a diuresis. Like dobutamine, if a diuresis is not induced, therapy should be discontinued as the alphaadrenergic effects could potentiate the NVN. The use of a furosemide trial (0.5–2 mg/kg i.v.) is the most rational therapy for NVN as it will induce the production of vasodilatory prostaglandins, which directly addresses the imbalance of vasodilators and vasoconstrictors which is the underlying cause of NVN.17,44–46 When furosemide therapy is successful in NVN, a diuresis will be induced with one or two doses and will be sustained without the need for further furosemide therapy. Repeated furosemide therapy will complicate fluid management by inducing electrolyte imbalances. In such cases fluid restriction is more effective and less metabolically disruptive than continued furosemide therapy. As stated earlier, inducing a diuresis is only important in helping fluid management and will not change the outcome of the renal disease. Continued decreased glomerular perfusion will not result in enough compromise of tubular perfusion to result in secondary damage in these neonates.17, 42 If a diuresis can not be achieved in foals with NVN, patience is the key to successful therapy. Fluid restriction should be instituted and oliguria tolerated until it spontaneously resolves, which may take as long as 72 hours.
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be placed on decreasing the water overload and not with supplementing with sodium. On the other hand, in a sodium-wasting nephropathy it is important to replace the sodium losses. Fractional excretion of sodium can help to differentiate the two conditions. Without measuring the sodium excretion, the amount of sodium replacement must be empirically judged based on the degree of hyponatremia, the fractional excretion of sodium, and degree of edema. Although placing an indwelling urinary catheter, even with closed collection, can increase the risk of serious nosocomial infection, when it is indicated based on concurrent problems such as cystomegaly, the information gained from total urine collection can be utilized to help sodium and fluid management as well as characterize and follow the renal dysfunction. As most critical neonates with oliguria during the first few days of life pass little or no feces, sodium loss in the urine will account for almost all the sodium loss from the body, as long as the clinical course is not complicated by reflux or hemorrhage. If the urine output is accurately measured (as with a closed urinary collection device) and if a sodium level is measured from a mixed aliquot of urine saved each time the collection bag is changed, the sodium excretion can easily be calculated and sodium replacement (total sodium delivered in fluids, drugs, PPN formula, oral supplementation, etc.) matched to sodium losses, maintaining sodium balance. As long as the patient is not sodium overloaded from excessive amounts of sodium from initial resuscitation fluids, this approach can be very helpful. Similarly, in the fluidoverloaded patient, adjusting fluid delivery every 6 hours to match urine output in the past 6 hours will result in gradual decrease in fluid overload as insensible losses will not be replaced. Losses can be followed by changes in bodyweight but can be confounded by catabolism or growth. So management of sodium balance and fluid balance are simplified by using total urine collection, but secondary nosocomial infection from the indwelling urinary catheter may preclude this technique, although the risk varies between care settings. Other considerations when managing renal disease in neonatal foals includes careful management of potassium balance along with other electrolytes, special attention to acid–base balance, and consideration of adjusting the dose of all medications excreted through the kidneys. It is important not to let such details go overlooked.
Sodium management Sodium management in neonatal foals with renal disease can be difficult. Hyponatremia is common. It may be secondary to water overload in the face of normal total body sodium or even sodium overload. In such cases the therapeutic emphasis should
Outcome and prognosis Most neonatal foals with signs of mild renal compromise recover fully and show no indications of further problems after the neonatal period. The best
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indication that serious chronic renal disease is present and may evolve into chronic renal failure is a failure of the blood creatinine level to decrease to normal values within the first week of life. An even more ominous sign is a rising blood creatinine. It should be remembered that, even when an insult is brief and the damage finite, it may take several days for the creatinine to reach a new steady state; so continued rise does not equate to continued insult. In any foal which has had signs of renal compromise during the first week of life, serial creatinine values should be taken even after the creatinine level is normal. A percentage of foals will have a secondary rise in blood creatinine between 2 and 4 weeks of age, probably reflecting the kidneys’ marginal function and difficulty dealing with the increased creatinine produced as the foal’s muscle mass increases. Some such foals will have an increased creatinine level of between 180 and 350 µmol/L (2.0–4.0 mg/dL) for a year or more, reflecting the chronic loss of function and lower GFR. In any case, even in foals with mild indications of renal compromise during the neonatal period, renal precautions such as avoiding nephrotoxic drugs or situations which could lead to renal compromise is a prudent approach.
References 1. Brophy PD, Robillard JE. Functional development of the kidney in utero. In: Polin R, Fox W, Abman S (eds) Fetal and Neonatal Physiology, 3rd edn. Philadelphia: Elsevier, 2004; pp. 1229–41. 2. Solhaug MJ, Jose PA. Postnatal maturation of renal blood flow. In: Polin R, Fox W, Abman S (eds) Fetal and Neonatal Physiology, 3rd edn. Philadelphia: Elsevier, 2004; pp 1242–8. 3. Guignard JP. Postnatal development of glomerular filtration rate in neonates. In: Polin R, Fox W, Abman S (eds) Fetal and Neonatal Physiology, 3rd edn. Philadelphia: Elsevier, 2004; pp 1256–66. 4. Buckley NM, Brazeau P, Frasier ID. Renal blood flow autoregulation in young swine. Am J Physiol 1983;245(1):H1–6. 5. Chevalier RL, Carey RM, Kaiser DL. Endogenous prostaglandins modulate autoregulation of renal blood flow in young rats. Am J Physiol 1987;253(1):F66–75. 6. Brewer BD, Clement SF, Lotz WS, Gronwall R. A comparison of inulin, para-aminohippuric acid, and endogenous creatinine clearances as measures of renal function in neonatal foals. J Vet Intern Med 1990;4:301–5. 7. Brewer BD, Clement SF, Lotz WS, Gronwall R. Renal clearance, urinary excretion of endogenous substances, and urinary diagnostic indices in healthy neonatal foals. J Vet Intern Med 1991;5(1):28–33. 8. Dalton R. Renal function in neonatal calves: Inulin, thiosulphate and para-aminohippuric acid clearance. Br Vet J 1987;125:498–502. 9. Holdstock NB, Ousey JC, Rossdale PD. Glomerular filtration rate, effective renal plasma flow, blood pressure and heart rate in the equine neonate during the first 10 days post partum. Equine Vet J 1998;30(4):335–43. 10. Palmer JE. Recognition and resuscitation of the critically ill foal. In: Paradis MR (ed.) Equine Neonatal Medicine: A Case-
11.
12.
13.
14. 15.
16. 17.
18.
19.
20.
21. 22. 23. 24.
25. 26. 27. 28.
29.
30. 31.
32.
33.
Based Approach. Philadelphia: Elsevier Saunders, 2006; pp. 121–34. Banks RO. Segmental nephron sodium and potassium reabsorption in newborn and adult dogs during saline expansion. Proc Soc Exp Biol Med 1983;173(2):231–7. Feld LG, Corey HE. Renal transport of sodium during early development. In: Polin R, Fox W, Abman S (eds) Fetal and Neonatal Physiology, 3rd edn. Philadelphia: Elsevier, 2004; pp. 1267–78. Slotkin TA, Seidler FJ, Kavlock RJ, Gray JA. Fetal dexamethasone exposure accelerates development of renal function: Relationship to dose, cell differentiation and growth Inhibition. J Dev Physiol 1992;17(2):55–61. Gouyon JB, Guignard JP. Management of acute renal failure in newborns. Pediatr Nephrol 2000;14(10–11):1037–44. Hunley TE, Kon K. Pathophysiology of acute renal failure in the neonatal period. In: Polin R, Fox W, Abman S (eds) Fetal and Neonatal Physiology, 3rd edn. Philadelphia: Elsevier, 2004; pp. 1335–40. Kellum JA. Acute kidney injury. Crit Care Med 2008;36(4 Suppl):S141–S145. Toth-Heyn P, Drukker A, Gllinard JP. The stressed neonatal kidney: from pathophysiology to clinical management of neonatal vasomotor nephropathy. Pediatr Nephrol 2000;14(3):227–39. Baer PG, Navar LG. Renal vasodilation and uncoupling of blood flow and filtration rate autoregulation. Kidney Int 1973;4(1):12–21. Robillard JE, Nakamura KT, Matherne GP, Jose PA. Renal hemodynamics and functional adjustments to postnatal life. Semin Perinatol 1988;12(2):142–50. Fanning AS, Mitic LL, Anderson JM. Transmembrane protein in the tight junction barrier. J Am Soc Nephrol 1999;10(6):1337–45. Schetz M, Dasta J, Goldstein S, Golper T. Drug-induced acute kidney injury. Curr Opin Crit Care 2005;11(6):555–65. Szeto CC, Chow KM. Nephrotoxicity related to new therapeutic compounds. Ren Fail 2005;27(3):329–33. Rocher LL, Tannen RL. The clinical spectrum of renal tubular acidosis. Ann Rev Med 1986;37:319–31. Ring T, Frische S, Nielsen S. Clinical review: Renal tubular acidosis – a physicochemical approach. Crit Care 2005;9(6):573–80. Bagga A, Sinha A. Evaluation of renal tubular acidosis. Indian J Pediatr 2007;74(7):679–86. Hemstreet BA. Antimicrobial-associated renal tubular acidosis. Ann Pharmacother 2004;38(6):1031–8. Arroyo LG, Stampfli HR. Equine renal tubular disorders. Vet Clin Equine 2007;23(3):631–9. Firmin CJ, Kruger TF, Davids R. Proximal renal tubular acidosis in pregnancy. Gynecol Obstet Invest 2007;63(1):39– 44. Fanos V, Cuzzolin L. Causes and manifestation of nephrotoxicity. In: Geary DF, Schaefer F (eds) Comprehensive Pediatric Nephrology. Philadelphia: Mosby Elsevier, 2008; pp. 1003–16. Andreoli SP. Acute renal failure in the newborn. Semin Perinatol 2004;28(2):112–23. Guignard JP. The adverse renal effects of prostaglandinsynthesis inhibitors in the newborn rabbit. Semin Perinatol 2002;26(6):398–405. Subramanian S, Agarwal R, Deorari AK, Paul VK, Bagga1 A. Acute renal failure in neonates. Indian J Pediatr 2008;75(4):385–91. Naderer O, Nafziger AN, Bertino JS. Effects of moderatedose versus high-dose trimethoprim on serum creatinine
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34.
35.
36.
37.
38.
39.
and creatinine clearance and adverse reactions. Antimicrob Agents Chemother 1997;41(11):2466–70. Silkensen JR, Kasiske BL. Laboratory assessment of renal disease: Clearance, urinalysis, and renal biopsy. In: Brenner BM (ed.) Brenner and Rector’s The Kidney, 7th edn. Philadelphia: Saunders, 2004; pp. 1107–50. Levey AS, Berg RL, Gassman JJ, Hall PM, Walker WG. Creatinine filtration, secretion and excretion during progressive renal disease. Kidney Int 1989;36(suppl 27):S73–80. Jaffe M. Uber den niederschlag, welchen pikrinsaure in normalen hrn erzeugt und uber eine neue reaction des kreatinins. Z Physiol Chem 1886;10:391–400. Peake M, Whiting M. Measurement of serum creatinine – current status and future goals. Clin Biochem Rev 2006;27(4):173–84. Dossin O, Germain C, Braun JP. Comparison of the techniques of evaluation of urine dilution/concentration in the dog. J Vet Med A Physiol Pathol Clin Med 2003;50(6):322–5. Reine NJ, Langston CE. Urinalysis interpretation: how to squeeze out the maximum information from a small sample. Clin Tech Small Anim Pract 2005;20(1):2–10.
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40. Patel HP. The abnormal urinalysis. Pediatr Clin N Am 2006;53(3):325–37. 41. Palmer JE. Fluid therapy in the neonate: not your mother’s fluid space. Vet Clin North Am Equine Pract 2004;20(1):63– 75. 42. Bagshaw SM, Bellomo R, Kellum JA. Oliguria, volume overload, and loop diuretics. Crit Care Med 2008;36(4 Suppl):S172–8. 43. Kellum JA. Use of diuretics in the acute care setting. Kidney Int 1998;53(suppl 66):67–70. 44. Antonucci R, Cuzzolin L, Arceri A, Fanos V. Urinary prostaglandin E2 in the newborn and infant. Prostagland Lipid Mediat 2007;84(1):1–13. 45. Mann B, Hartner A, Jensen BL, Kammerl M, Kramer BK, Kurtz A. Furosemide stimulates macula densa cyclooxygenase-2 expression in rats. Kidney Int 2001;59:62–8. 46. Nies AS, Gal J, Fadul S, Gerber JG. Indomethacin– furosemide interaction: the importance of renal blood flow. J Pharmacol Exper Therapeut 1983;226(1):27–32.
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Inherited Problems of Foals Sharon J. Spier, Carrie J. Finno and Stephanie J. Valberg
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Chapter 72
Congenital Problems of Foals Antonio M. Cruz
Introduction Congenital defects or abnormalities in foals are poorly described from an epidemiological perspective. By definition a congenital abnormality is one that is present at birth, which may be the result of genetic abnormalities, intrauterine environment, errors of morphogenesis, or a chromosomal abnormality. The outcome of the disorder will further depend on complex interactions between the prenatal deficit and the postnatal environment.1 In addition environmental toxins, known as teratogens, could cause birth defects. In equine medicine, most of the descriptions of birth defects arise from case reports, retrospective studies or small case series as there is no centralized reporting mechanism to capture their real incidence or population characteristics. Such a database would aid in the investigation of potential causes and serve as a surrogate indicator of environmental health, in an effect similar to the ‘canary in the coal mine’. For instance, natural exposure to zinc has been associated with the development of osteochondrosis.2 Since no mandatory reporting of congenital abnormalities exists, many abnormalities go undocumented and unreported. Many congenital anomalies may not be diagnosed until later in life and diagnosed rather as developmental. Genetic and congenital diseases are a very important consideration for the equine industry, particularly in breeds where the genetic pool may be narrow and where genetic etiologies have been postulated. Such diseases affect the value of the breeding stock and the animal itself and lessen genetic gain. Control measures in breeding programs may require costly changes such as exclusion of valuable animals from breeding, to eradicate genetic defects. Some examples might be the eradication of the carrier status for hyperkalemic periodic paralysis (HYPP) and combined immunodeficiency (CID). Genetic conditions are perhaps one of the most important problems affecting equine health today.
The technical advances in cytogenetics have enabled the identification of chromosome specific aberrations. Advances in genomic tools have been used to more precisely define chromosome abnormalities.3 The day when chromosomal aberrations are linked to specific defects and diagnostic methods implemented to eliminate them has already appeared. It is beyond the scope of this chapter to produce a treaty of all the congenital abnormalities reported in horses. A search of the PubMed database under the terms ‘congenital’ and ‘equine’ yielded 517 responses. In a retrospective study spanning 13 years in Kentucky and including 608 foals, the following congenital anomalies were observed: contracted foal syndrome (33.2%), miscellaneous limb contraction (20%), multiple defects (5.3%), microphthalmia (4.6%), craniofacial malformations (4.3%), cleft palate (4.0%), heart defects (3.5%), umbilical defects (3.5%), and hydrocephalus (3.0%).4 In another study involving 3520 foals, causes of perinatal death were investigated and 10% of foals were reported to have died due to congenital abnormalities.5 Among a total of 137 717 cases in clinics of veterinary colleges in the United States and Canada, 6455 patients with congenital defects were reported of which 17.5% were horses.6 This data confirms congenital abnormalities as a significant clinical problem with a potentially large impact in the overall equine industry. The urogenital and musculoskeletal systems are the ones most commonly affected by congenital anomalies in the horse.
Respiratory system The most commonly reported congenital problems of the respiratory system include cleft palate, guttural pouch tympany, dorsal displacement of the palatopharyngeal arch, and wry nose. Cleft palate Cleft palate is an uncommon congenital defect manifested by nasal discharge of milk during or after
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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nursing as well as coughing. No specific breeds appear to be affected preferentially although in the author’s experience Fell ponies may be overrepresented. Most of these foals also develop aspiration pneumonia, as they are unable to entirely protect the airway during nursing or eating. The owner usually seeks veterinary advice, alarmed by the nasal discharge present in an otherwise healthy foal. Alternatively, they may seek veterinary advice once the foal becomes clinically affected by aspiration pneumonia, which may take several days to weeks to develop. A cleft palate can affect the hard or soft palate or both and the amount of palate missing is variable. A thorough approach to the diagnosis of dysphagia and nasal discharge includes endoscopic and radiographic examination of the nasal cavity, pharynx, and larynx as well as an examination of the oral cavity. Similar clinical signs can arise from pharyngeal dysfunction or other deformities affecting the palatopharyngeal locking mechanism. Foals affected with cleft palate should be treated surgically or euthanized. There are no reports regarding athletic prognosis of affected horses surgically treated. Surgical repair is associated with almost 100% morbidity and 50% mortality.7 Affected horses probably should not be bred. Benign neglect is not an acceptable treatment. Depending on what part of the palate is affected, different surgical approaches may be selected and the reader is directed to a surgical textbook where the technique is described.8 More recently, a transoral technique using laparoscopic instrumentation has been used successfully, avoiding the traditional surgical approach which is fraught with complications. (L. Boure, personal communication). Guttural pouch tympany This is a poorly explained condition, which preferentially affects Arabian fillies.9 Although it can be seen in older foals, this condition is mostly seen shortly after birth. Although clinical signs are not present at birth, it is thought that the functional defect of the plica salpingopharyngea is already present. Occasionally some older foals develop the condition due to scarring of the plica salpingopharyngea. This condition can be unilateral (most common) or bilateral. The latter may represent an emergency if the airway is severely obstructed. The plica salpingopharyngea acts as a one-way valve allowing air into the diverticulum but preventing it from exiting. As a result the guttural pouch fills with air resulting in the characteristic tympanic appearance from the outside. The retropharyngeal swelling is soft, indentable, and non-painful. An endoscopic and radiographic evaluation of the guttural pouch is mandatory in these cases. Once the endoscope is passed into the guttural
pouch, the air exits and the visible swelling diminishes dramatically but normally reappears shortly after the examination. During radiographic examination, a large, air-filled radiolucent area is present, extending from the pharyngeal region to the level of the second or third cervical vertebrae. Some affected foals may present with a secondary empyema and mild nasal discharge or they can be completely normal otherwise. Fortunately most cases are unilateral and respond well to surgical treatment. Temporary relief can be obtained by maintaining an indwelling Foley catheter in the affected pouch. Occasionally the condition resolves after the tube is maintained for a period of 4–6 weeks. A catheter as large as possible should be placed in the affected pouch. The author prefers the use of a 20–24 Fr Foley, as the inflated balloon may prevent premature dislodgement. In addition the Foley catheter seems to be less traumatic than a plastic material, although its placement can be a bit more challenging due to its flexible nature. A blunt thorax stylet into the Foley assists with placement and sutures at the end of the catheter to the alar fold in two different spots approximately 180◦ apart is recommended. The balloon is filled to its maximum volume with saline. The surgical correction of unilateral pouch tympany involves the fenestration of the mesial septum either by an open technique or an endoscopic transnasal approach using electrocautery or laser. In cases of bilateral tympany, the plica salpingopharyngea can be resected or, ideally, a bilateral fistula can be created between the guttural pouch and nasopharynx at the most dorsal aspect of the plica. Although there appears to be no long-term sequelae of this technique, the surgeon may accidentally damage the maxillary vessels during the procedure; therefore a thorough knowledge of the region’s anatomy and optimal visualization are mandatory prior to performing transnasal endoscopic electrosurgery or laser surgery. It is important once any fenestration has been performed to maintain a Foley catheter through it for a period of 4–6 weeks to prevent scarring of the fenestration and subsequent closure. The placement of a catheter facilitates formation of a permanent fenestration, resolving the problem effectively. Rostral displacement of the palatopharyngeal arch This condition is frequently a symptom of a larger problem described as a defect of the fourth branchial arch during embryogenesis. The extrinsic structures affected include the thyroid wing, the cricothyroid articulation, the cricothyroid muscle and the upper esophageal sphincter10 which lead to malformation (aplasia/hypoplasia) of one or more cartilaginous or
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Congenital Problems of Foals muscular structures. Typically these horses may be presented later on in life due to abnormal respiratory noise and/or dysphagia associated with performance, but they are often not identified as abnormal during their early life. The clinician should be aware that the endoscopic diagnosis may misrepresent the severity of the defect and other tests such as radiography and ultrasonography may be pursued, although palpation is usually diagnostic of an anatomic abnormality. To diagnose these cases purely by identification of a piece of tissue lying over the corniculate processes of the arytenoid cartilage can be a mistake. Fourth branchial arch defect cases should be thoroughly investigated prior to recommending any treatments. Unfortunately there are few treatment options available for this condition, although resection of the displaced portion of the palatopharyngeal arch in horses not intended for any athletic pursuits may ease some of the associated clinical signs. Choanal atresia The persistence of the buccopharyngeal septum (choanal atresia) prevents communication between the nose and the nasopharynx, leading to partial (unilateral) or complete (bilateral) occlusion of the respiratory tract. In cases of bilateral atresia, becausethe horse is an obligate nasal breather an astute clinician may recognize the inability of the newborn to breathe and perform a quick life-saving tracheostomy. However, since many deliveries may be unattended, the affected foal often dies shortly after being born and the cause misdiagnosed as simply a stillbirth unless a detailed postmortem investigation is performed. In cases of unilateral atresia, successful surgical management has been described through different approaches depending on its nature (bony vs membranous) although the prognosis for high-level athleticism remains guarded.11 Wry nose Wry nose is a congenital deviation of the maxillae, premaxillae, nasal bones, and vomer bone. Its incidence appears to be increased in Arabian horses and there is no predisposition to sex or direction of the deviation.12 Its identification is relatively simple due to its characteristic appearance. Additional signs may include dyspnea, regurgitation, dysphagia, and ill-thrift. Recently successful surgical correction by using external fixators and the principle of distraction osteogenesis has been reported.13 Alternatively a much more involved one- or two-stage surgery can also be attempted, with a poor to guarded prognosis.14 Surgical reconstruction should not be attempted prior to 6 months of age.
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Urogenital system In the urogenital system the most common congenital conditions include inguinal hernia, cryptorchidism, intersex, and ureteral ectopia. Inguinal hernia The early recognition of herniated abdominal contents into the scrotum is important to reduce the chances of colic or the need for an emergency laparotomy or herniorraphy. Many newborn foals may have an enlarged scrotum due to the presence of fluid and/or intestines. Initially scrotal hernias do not require surgical repair unless they cannot be reduced easily or are associated with intestinal strangulation and/or rupture of the peritoneum. These conditions are associated with scrotal pain, swelling, and vascular compromise to the scrotal skin leading to edema15 (Figure 72.1) and, in advanced cases, skin necrosis. A foal with a strangulated hernia will display signs of colic and cardiovascular response compatible with those seen associated with an intestinal vascular obstruction. The clinician should be able to recognize these clinical signs early and recommend a surgical herniorraphy and/or laparotomy. If the hernia has not ruptured the peritoneum and is reducible, it can be dealt with by daily reduction since most uncomplicated hernias resolve spontaneously as the foal grows older. In the author’s experience inguinal bandages are not very practical and are cumbersome to use for many owners. Inguinal herniorraphy of uncomplicated cases can also be done at the owner’s request. A bilateral procedure and castration should be performed if the procedure is performed through an inguinal approach, since leaving the testicles intact may increase the chances of reccurrence. Castration at an early age does not appear to have long-lasting effects on development and performance. Recently a successful laparoscopic testicle-sparing technique has also been described and should be available at most referral hospitals.16 The prognosis for both techniques is excellent as long as intestinal vascular compromise has not occurred. Since there is a strong indication that this condition is inherited, there should be serious reservations about testicle-sparing techniques, which may allow subsequent breeding and transmission of the defect. Cryptorchidism The reader is referred to Chapters 106 and 158. Intersex The reader is referred to Chapter 106.
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(a)
(b)
Figure 72.1 (a) Ruptured inguinal hernia in a foal presenting with skin edema. (b) Hernia as seen at surgery.
Ureteral ectopia This is the most common congenital anomaly of the equine urinary tract and is more common in fillies. This condition is caused by an abnormal development of the metanephric ducts leading to the relocation of the ureters draining into the uterus, vagina, or urethra.17 Foals affected with ectopic ureters commonly present with urinary incontinence and resulting skin scalding and a persistently wet perineum and hind legs. Other congenital defects of the urinary system may be concurrently present and the clinician should investigate those possibilities prior to recommending any treatment or discussing prognosis with the owner.18 Surgical repair and conservative treatments have been reported.19
Musculoskeletal system Flexural and angular limb deformities All neonatal foals have some degree of flexural and angular deformity due to weakness of the supporting connective tissues (tendons and ligaments). Most correct over the first few days of life as the foal exercises and gains muscle strength. Occasionally the defect may be so severe that it prevents the mare from delivering normally per vagina (carpal contracture) or requires treatment. Weak flexor tendons leading to hyperextension of the distal extremity are best treated with controlled exercise and corrective shoeing (heel extensions). Occasionally the affected limbs require light bandaging of the legs to protect them, but ideally the tendons should experience adequate loading and weight-bearing.
‘Contracted’ tendons produce hyperflexion of the distal extremity and require prompt treatment. Administration of 3 g of intravenous oxytetracycline, splinting and/or surgical treatment can be used by themselves or in combination to treat these problems. Ideally the invasiveness of treatment should match the severity of the condition. Congenital medial/lateral angular deformities (Figure 72.2) are usually due to laxity of the periarticular structures and usually resolve with time and adequate support in the form of bandaging or splinting. 20 Additional corrective trimming and shoeing may be necessary. As a general rule the extension should be placed on the part of the foot that corresponds to the convex part of the deformity: medially in cases of valgus (lateral deviation) and laterally in cases of varus (medial deviation), and the hoof should be rasped on the side opposite to where the extension is placed. The prognosis for these deformities is usually favorable. Deformities that occur as a result of diaphyseal deviation within the third metacarpus or metatarsus should be addressed with a corrective osteotomy if conservative treatment fails. Patellar luxation Patellar luxation is seen mostly in Miniature foals and in Shetland ponies. This condition prevents extension of the stifle and the foal adopts a characteristic squatting stance. This condition should be differentiated from femoral nerve paralysis. Most congenital patellar luxations are lateral as a result of hypoplasia of the lateral trochlear ridge of the femur.21 Lateral patellar luxation has been linked to a homozygosity of a single autosomal recessive gene.22 Successful surgical treatment has been reported.23
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the longer bone (either mandible or maxillae) by the use of intraoral braces.25 Arthrogryposis This condition is defined as a permanent joint contracture at birth and may be neurogenic, myogenic, or caused by lesions in joint surfaces. It may involve the front or hind legs and may be accompanied by other defects such as cleft palate, maxillary brachygnathia, and torticollis. In Norwegian Fjord horses, the syndrome of bilateral hind legs arthrogryposis associated with cleft palate, maxillary brachygnathia and polydactylism of the hind leg is lethal and considered to be genetically transmitted.26 No current treatment is available for this condition and most foals are born dead due to prolonged dystocia. Contracted foals
Figure 72.2 Severe congenital angular limb deformity (carpus valgus).
This syndrome includes asymmetric skull development, torticollis, scoliosis of thoracolumbar area, bilateral flexion contracture of distal limbs more commonly involving front limbs, and defects of the ventral abdominal wall. The syndrome varies from mild, reversible, flexure contractures to severe lesions of skull and vertebral column combined with joint contractures.27, 28
Upward fixation of the patella
Joint dysplasia
This is a fairly common condition in yearling, 2-yearold, and adult horses. It rarely occurs in foals. The foal shows a partial or complete inability to flex the stifle and therefore the hip and hock, resulting in temporary or permanent leg extension. The patella is ‘locked’ in an abnormal location due to the medial patellar ligament being ‘hooked’ over the large medial trochlear ridge of the femur. This condition can be treated successfully in many instances.24
Shoulder and hip dysplasia have been described in the horse.25 Shoulder dysplasia affects Shetland and Miniature ponies preferentially.29,30, 31 It is characterized by a disparity of size between the glenoid cavity and the head of the humerus leading to severe juvenile osteoarthritis. Scapulohumeral arthrodesis has been successfully used to treat this condition and salvage affected horses.32 Hip dysplasia leads to severe osteoarthritis and its successful treatment has not been reported.
Brachygnathia (parrot mouth and monkey mouth)
Vertebral malformations
Mandibular brachygnathia (parrot mouth) is more common than its maxillary counterpart (monkey mouth). This congenital defect may lead to important abnormalities in dental wearing and associated problems. This abnormality is considered to be linked to a single autosomal recessive gene. Surgical correction should be accompanied by adequate counseling to the owner and an inability to register the affected animal in its corresponding breed registry. The condition can be surgically fixed through growth retardation of
These malformations include kyphosis (dorsal deviation), lordosis (ventral deviation), scoliosis (lateral deviation),33 torticollis (twisted neck), and all of their combinations. The most commonly reported vertebral malformation in horses is atlanto-occipital junction fusion with hypoplasia of the dens. This condition affects Arabian horses exclusively and may go undetected unless there is spinal cord damage and associated defects.34 Horses may also develop ‘wobbler’ syndrome as a result of cervical vertebral malformations.35
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Digestive system Ileocolonic aganglionosis (Overo lethal white syndrome) This syndrome is caused by the congenital absence of myenteric ganglia in the terminal portion of the ileum, cecum, large colon, and small colon. It is seen in foals born to parents of Overo lineage which themselves do not have to be of Overo coloration. The gene responsible for this defect causes a prominent Overo color pattern. The lethal mutation in the responsible gene produces an abnormality in the endothelin signaling pathway during the development of the neural crest cells. Foals born with this condition are born white with ‘glass eyes’ appearance and invariably develop signs of colic due to a functional obstruction. Affected foals should be euthanized.36 Breeding stock can be tested and the disease avoided by not breeding heterozygotes.37 Intestinal atresia Atresia is the lack of development of a part of the intestinal tract. Typically it can affect the colon (coli), rectum (recti), or anus (ani). Foals affected with atresia suffer from intestinal obstruction and colic signs. Some forms of atresia can be successfully treated surgically, particularly atresia ani. The prognosis for atresia coli and recti remains guarded to poor. Differential diagnosis include meconium impaction and other causes of intestinal obstruction in foals. Contrast radiographs with barium and proctoscopy can be used to confirm a diagnosis and determine the location of the obstruction. The cause of atresia remains unknown but it is speculated that congenital loss of vascular supply to the affected intestine is implicated.38 Umbilical hernia This is the second most common congenital defect of horses occurring in 0.5–2% of foals.39 The diagnosis is relatively simple as a defect can be palpated in the body wall with abdominal contents filling a hernia sac. Surgical correction (herniorrhaphy) is indicated in cases where the hernia is non-reducible or if the hernia is large (> 4 fingers wide). Smaller hernias may resolve spontaneously with age. Different techniques have been used to successfully treat hernias including use of hernia clamps,40 surgical herniorraphy, and abdominal bandaging.41
Ophthalmic abnormalities Many ocular congenital abnormalities have been described in foals. Entropion is occasionally seen in the
foal. It is the inversion of one or both of the eyelids, with the lower one preferentially affected. Cataracts are the most common cause of blindness in newborn foals and represent about a third of congenital ocular defects. Congenital cataracts can be inherited but they can also be the result of toxic or traumatic insult. Cataracts described in foals include complete mature, nuclear, associated with the Y sutures of the lens, and those linked to hyaloids remnants.42 Bilateral absence of the iris, referred as aniridia, has been reported most commonly in the Belgian breed and is considered a genetic defect most likely transmitted as a simple autosomal recessive.43 Microphthalmia has been reported.4 Additionally congenital corneal opacities can be found in the neonate and are attributed to a thin and abnormal Descemet membrane.44 Retinal defects such as detachment and colobomas are rare but have been described in isolation or in combination with other defects.45 Finally, congenital stationary night blindness has been described in Appaloosa horses and is believed to be inherited as a recessive gene.45
Metabolic and endocrine abnormalities Hyperkalemic periodic paralysis Hyperkalemic periodic paralysis (HYPP) affects Quarter Horses, Paint horses, Appaloosas and any other horses with Impressive lineage. HYPP results from an altered electrochemical gradient across the sarcolemma. Affected horses are well muscled and exhibit episodes of sustained muscle contraction and generalized muscle tension. Recumbency may occur in some horses and these episodes last for 30–60 minutes. Serum potassium concentration is increased during the episodes and returns to normal between episodes. Diagnosis is done by a genetic test using DNA. In mild cases the disease is manageable and horses compete successfully. Dietary modifications such as feeding the horse a readily absorbable source of carbohydrates (oats, timothy hay) and avoiding alfalfa hay as well as the use of acetazolamide are beneficial. In severe episodes horses should be treated with 23% calcium gluconate (0.2–0.4 mL/kg) or bicarbonate (1–2 mEq/kg).46
Polysaccharide storage myopathy Polysaccharide storage myopathy (PSSM) affects Quarter Horses and related breeds. It is characterized by repeated episodes of tying-up due to abnormal accumulation of polysaccharide within the myoplasm.47 A different disease, or perhaps
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Congenital Problems of Foals a variant of PSSM, termed equine polysaccharide storage myopathy has been described in draft breed horses.48 Signs may vary and can be similar to a tying episode or manifested by progressive weakness, poor performance, shivers, muscle wasting, and potentially death. Diagnosis is by muscle biopsy. Dietary and exercise control are usually adequate in mild or early cases. The diet is modified to provide enough calories, limiting the intake of carbohydrates, but increasing dietary fat.
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Hydranencephaly The complete or almost complete absence of cerebral hemispheres is called hydranencephaly. The cranium has normal conformation and the resulting space is filled with cerebrospinal fluid surrounded by a thin membranous cerebral tissue. The defect is uncommon in horses, and its cause is unknown. It may be associated with hydrops allantois.51
Internal hydrocephalus Congenital hypothyroidism This condition is part of a syndrome characterized by multiple congenital musculoskeletal deformities (prognathia, delayed ossification of cuboidal bones, angular limb deformities, and rupture of common digital extensor tendons) and thyroid gland hyperplasia. In addition to the musculoskeletal deformities, these foals have signs of dysmaturity, including a silky, short hair coat, floppy ears, and tendon laxity. Supportive therapy is indicated. Long-term outcomes seem to be poor.49
Central nervous system Congenital defects of brain and spinal cord may involve the central nervous system only or may be combined with skeletal lesions. These defects are best classified by combining anatomical and functional factors: cerebral defects and malformation involving primarily the cerebrum, defects involving primarily the cerebellum and brainstem, and spinal cord defects.50–52 Equine central nervous system defects were common in one study and ranged from cyclopia to hydrocephalus.4 Unfortunately etiological studies of central nervous system defects in horses appear to be rare. The frequency of these defects is not a fixed proportion of all foal births, but varies with genetic and environmental factors. Neurological abnormalities are common in the newborn foal and may be the result of sepsis, electrolyte disturbances, hypoglycemia, hypoxic or ischemic injuries, and congenital defects.53 Morphological information obtained by computed tomography scanning will greatly improve the diagnosis of neonatal neurological disorders.50
Excessive fluid accumulating in the cranial cavity within the ventricular system (internal hydrocephalus) may cause dystocia, because of the large size of the head (Figure 72.3). Hydrocephalus is a commonly reported defect of unknown origin in horses.52
Microencephaly An abnormally small but grossly normal brain, called microencephaly, has been described in foals.52, 54 Defects of the cerebellum and brainstem
Dandy-Walker syndrome This syndrome consists of hydrocephalus, aplasia or hypoplasia of cerebellar vermis, and cystic enlargement of the fourth ventricle, covered by a markedly thinned medullary velum. It is rare in foals.50, 55
Cerebellar hypoplasia The clinical signs are present at birth and include recumbency with extended limbs, intermittent opisthotonus, and ataxia. Whereas earlier investigators invariably ascribed cerebellar disease in calves to
Cerebral defects
Agenesis of the corpus callosum Agenesis of the corpus callosum is commonly associated with other defects such as hydranencephaly, agenesis of the septum pellucidum, internal hydrocephalus, porencephaly or microencephaly, or Dandy-Walker syndrome.50
Figure 72.3 Hydrocephalus present in a foal delivered by cesarean section due to dystocia.
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hereditary causes, researchers have documented that intrauterine fetal infection with bovine viral diarrhea virus causes cerebellar hypoplasia.52 No prenatal viral infection is reported to be teratogenic in foals.
Cerebellar disease in Arabian foals Signs of cerebellar disease are present at birth or appear at 3 to 6 months of age and are characterized by intention tremor. Histological lesions include degeneration of the Purkinje cells and mineralized neurons of the thalamus.56 Spinal cord Spina bifida, defective closure of the dorsal vertebral laminae, has been described in foals.54 Other rare spinal cord defects reported include atlanto-occipital fusion and spinal dysraphism.54, 57
Skin Congenital defects involving skin and adnexa, with a few exceptions, are rare in horses and may be localized or generalized. Epitheliogenesis imperfecta Epitheliogenesis imperfecta has been reported in most species including the horse and is characterized at birth by lack of development of the epithelium in patches, usually just above the carpal and tarsal area. The tongue may have similar lesions, and one or more hooves may be partially or completely missing. The disease is fatal and is transmitted by a single autosomal recessive trait.58 Epidermolysis bullosa Epidermolysis bullosa refers to a group of mechanobullous diseases in which lesions are caused by a defect in the dermal-epidermal junction at the level of the basement membrane. The condition is the result of a defect in one of the genes encoding laminin, a glycoprotein present in the basement membrane. These diseases have been described primarily in American Saddlebred and Belgian horses.59–62 Hyperelastosis cutis This group of inherited connective tissue disorders has also been referred to as cutaneous asthenia, Ehlers-Danlos syndrome, and dermatosparaxis. 63 This disease has only been seen in Quarter Horses. In the United States it is primarily seen in cutting horse lines. A similar condition has been described
in an Arabian cross-bred. The condition is apparent at birth or shortly thereafter. Males and females are equally affected. The disease in Quarter Horses is inherited as an autosomal recessive trait. The skin is hyperfragile, tears easily, and exhibits impaired healing. Many of the defects have a depressed surface best appreciated with proper lighting. Pain may be elicited when traction is applied to the edge of the defect.63
Curly coat Curly coat is reported in Percherons and is considered to be caused by homozygosity of a simple autosomal recessive trait. The trait does not affect the health and well-being of the foals.64
Immune system Combined immunodeficiency The reader is referred to Chapter 37.
Cardiovascular system Congenital heart and vascular defects may come to the attention of owners and veterinarians at birth. Affected horses may do poorly or have impaired athletic capabilities. Most cardiovascular defects are case reports and do not contain data about the frequency of these defects. In addition nothing is known about their cause. Interventricular septal defect is the most common heart defect in young horses.4 Affected foals may be ill-thrifty and may have intolerance to exercise, although clinical signs are highly variable, and likely related to the size of the defect. The defect is variable in size and located in the upper part of the membranous septum.65 Tetralogy and pentalogy of Fallot are rare defects in horses and, in addition to an interventricular defect, include pulmonic stenosis, dextroposition of aorta, and right ventricular hypertrophy.66, 67 Pentalogy of Fallot also presents additionally a patent ductus arteriosus. Multiple heart defects in foals ranging in age from newborn to 3 weeks are recognized and characterized by ventricular septum defects, stenosis of the pulmonary artery, and persistence of a common truncus arteriosus.68 Another congenital malformation, persistence of foramen ovale, has also been blamed for neonatal death.69 Various defects of the large vessels such as patent ductus arteriosus also can occur in foals. Persistence of the right aortic arch and dextroposition of aorta are rare defects in horses.70, 71 In summary, all of these cases require careful clinical
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Congenital Problems of Foals examination using ultrasonography and confirmation at necropsy if indicated. 20.
References 1. Birth Defects Research at http://www.cdc.gov/ncbddd/bd/ research.htm, accessed October 17, 2008). 2. Kowalczyk DF, Gunson DE, Shoop CR, Ramberg CF Jr. The effects of natural exposure to high levels of zinc and cadmium in the immature pony as a function of age. Environ Res 1986;40(2):285–300. 3. Lear TL, Bailey E. Equine clinical cytogenetics: the past and future. Cytogenet Genome Res 2008;120(1–2):42–9. 4. Crowe MW, Swerczek TW. Equine congenital defects. Am J Vet Res 1985;46(2):353–8. 5. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203(8):1170–5. 6. Priester WA, Glass AG, Waggoner NS. Congenital defects in domesticated animals: general considerations. Am J Vet Res 1970;31(10):1871–9. 7. Semevolos SA, Ducharme N. Surgical repair of congenital cleft palate in horses: eight cases (1979–1997). Proc Am Ass Eq Pract 1998;44:267–8. 8. Ducharme NG. Pharynx. In: Auer J, Stick J (eds) Equine Surgery, 3rd edn. St Louis, MO: Saunders, 2006; pp. 560–4. 9. Blazyczek I, Hamann H, Deegen E, Distl O, Ohnesorge B. Retrospective analysis of 50 cases of guttural pouch tympany in foals. Vet Rec 2004;154(9):261–4. 10. Lane JG. Fourth Branchial arch defects. In: McGorum BC, Robinson NE, Schumacher J, Dixon PM (eds) Equine Respiratory Medicine and Surgery. Chicago: Saunders, 2006; pp. 467–72. 11. James FM, Parente EJ, Palmer JE. Management of bilateral choanal atresia in a foal. J Am Vet Med Assoc 2006;229(11):1784–9. 12. Vandeplassche M, Simoens P, Bouters R, De Vos N, Verschooten F. Aetiology and pathogenesis of congenital torticollis and head scoliosis in the equine foetus. Equine Vet J 1984;16(5):419–24. ´ 13. Puchol JL, Herr´an R, Durall I, Lopez J, D´ıaz-Bertrana C. Use of distraction osteogenesis for the correction of deviated nasal septum and premaxilla in a horse. J Am Vet Med Assoc 2004;224(7):1147–50. 14. Schumacher J, Brink P, Easley J, Pollock P. Surgical correction of wry nose in four horses. Vet Surg 2008;37(2): 142–8. 15. Orsini J. Small intestinal diseases associated with colic in the foal. In: Mair T, Divers T, Ducharme N (eds) Manual of Equine Gastroenterology. St. Louis, MO: Saunders, 2002; p. 477. 16. Caron JP, Brakenhoff J. Intracorporeal suture closure of the internal inguinal and vaginal rings in foals and horses. Vet Surg 2008;37(2):126–31. 17. Schott HC II. Disorders of the urinary system. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis, MO: Saunders, 2004; pp. 1169–83. 18. Houlton JEF, Wright IM, Matic S., Herrtage ME. Urinary incontinence in a Shire foal due to ureteral ectopia. Equine Vet J 1987;19(3):244–7. 19. Gettman LM, Ross MW, Elce YA. Bilateral ureterocystostomy to correct left ureteral atresia and right ureteral
21.
22.
23.
24.
25.
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32.
33. 34.
35. 36.
37.
38. 39.
40.
41.
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ectopia in an 8-month-old standardbred filly. Vet Surg 2005;34:657–61. Embertson RM. Congenital abnormalities of tendons and ligaments. Vet Clin N Am Equine Pract 1994;10(2):351–64. Rooney JR, Rake CW, Harmany KJ. Congenital lateral luxation of the patella in the horse. Cornell Vet 1971;61: 670–3. Hermans WA, Kersjes AW, van der Mey GJ, Dik KJ. Investigation into the heredity of congenital lateral patellar (sub)luxation in the Shetland pony. Vet Q 1987;9:1–8. Kobluk CN. Correction of patellar luxation by recession sulcoplasty in three foals. Vet Surg 1993;22(4): 298–300. Dumoulin M, Pille F, Desmet P, Dewulf J, Steenhaut M, Gasthuys F, Martens A. Upward fixation of the patella in the horse: a retrospective study. Vet Comp Orthop Traumatol 2007;20(2):119–25. Gift LJ, DeBowes RM, Clem MF, Rashmir-Raven A, Nyrop KA. Brachygnathia in horses: 20 cases (1979–1989). J Am Vet Med Assoc 1992;200(5):715–19. Nes N, Lømo OM, Bjerk˚as I. Hereditary lethal arthrogryposis (muscle contracture) in horses. Nord Vet Med 1982;31:425–30. Rooney JR. Contracted foals. Cornell Vet 1966;56:172–87. Boyd JS. Congenital deformities in two Clydesdale foals. Equine Vet J 1976;8:161–4. Speirs VC, Wrigley R. A case of bilateral hip dysplasia in a foal. Equine Vet J 1979;11(3):202–4. Clegg PD, Dyson SJ, Summerhays GE, Schramme MC. Scapulohumeral osteoarthritis in 20 Shetland ponies, miniature horses and falabella ponies. Vet Rec 2001;148(6):175–9. Boswell JC, Schramme MC, Wilson AM, May SA. Radiological study to evaluate suspected scapulohumeral joint dysplasia in Shetland ponies. Equine Vet J 1999;31(6):510–14. Semevolos SA, Watkins JP, Auer JA. Scapulohumeral arthrodesis in miniature horses. Vet Surg 2003;32(5): 416–20. Wong D, Miles K, Sponseller B. Congenital scoliosis in a quarter horse filly. Vet Radiol Ultrasound 2006;47(3):279–82. Watson AG, Mayhew IG. Familial congenital occipitoatlantoaxial malformation (OAAM) in the Arabian horse. Spin. 1986;11(4):334–9. Wagner PC, Grant BD, Reed SM. Cervical vertebral malformations. Vet Clin N Am Equine Pract 1987;3(2):385–96. Santschi EM. Ileocolonic aganglionosis. In: Mair T, Divers T, Ducharme N (eds) Manual of Equine Gastroenterology. St Louis, MO: Saunders, 2002; pp. 488–9. Santschi EM, Purdy AK, Valberg SJ, Vrotsos PD, Kaese H, Mickelson JR. Endothelin receptor B polymorphism associated with lethal white foal syndrome in horses. Mamm Genome 1998;9(4):306–9. Young RL, Linford RL, Olander HJ. Atresia coli in the foal: a review of six cases. Equine Vet J 1992;24(1):60–2. Orsini J. Small intestinal diseases associated with colic in the foal. In: Mair T, Divers T, Ducharme N (eds) Manual of Equine Gastroenterology. St Louis, MO: Saunders, 2002; p. 478. Riley CB, Cruz AM, Bailey JV, Barber SM, Fretz PB. Comparison of herniorrhaphy versus clamping of umbilical hernias in horses: a retrospective study of 93 cases (1982–1994). Can Vet J 1996;37(5):295–8. Fretz PB, Hamilton GF, Barber SM, Ferguson JG. Management of umbilical hernias in cattle and horses. J Am Vet Med Assoc 1983;183(5):550–2.
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42. AG Turner. Ocular conditions of neonatal foals. Vet Clin N Am Eq Pract 2004;20(2):429–40. 43. Eriksson K. Hereditary aniridia with secondary cataract in horses. Nord Vet Med 1955;7:773–93. 44. Latimer CA, Wyman M. Neonatal ophthalmology. Vet Clin N Am Eq Pract 1985;1(1):235–60. 45. Roberts SM. Congenital ocular anomalies. Vet Clin N Am Eq Pract 8 (3): 459–478, 1992. 46. Naylor JM, Jones V, Berry SL. Clinical syndrome and diagnosis of hyperkalemic periodic paralysis in Quarter horses. Eq Vet J 1993;25:227–30. 47. Valberg SJ, Mickelson JR, Gallant EM, MacLeay JM, Lentz L, de la Corte F. Exertional rhabdomyolysis in quarter horses and thoroughbreds: one syndrome, multiple aetiologies. Eq Vet J Suppl 1999;30:533. 48. Valentine B. Polysaccharide storage myopathy in draft and draft-related horses and ponies. Eq Pract 1999;21:16. 49. Allen AL, Doige CE, Fretz PB, Townsend HG. Hyperplasia of the thyroid gland and concurrent musculoskeletal deformities in western Canadian foals: reexamination of a previously described syndrome. Can Vet J 1994;35:31–8. 50. Cudd TA, Mayhew IG, Cottrill CM. Agenesis of the corpus callosum with cerebellar wermian hypoplasia in a foal resembling the Dandy-Walker syndrome: pre-mortem diagnosis by clinical evaluation and CT scanning. Eq Vet J 1989;21:378–81. 51. Waelchli RO, Ehrensberger F. Two related cases of cerebellar abnormality in equine fetuses associated with hydrops of fetal membranes. Vet Rec 1988;123:513–14. 52. Leipold HW, Troyer DL. Congenital neurologic abnormalities. Proc Am Coll Vet Int Med 1990;8:611–15. 53. Adams A, Mayhew IG. Neurologic disease. Vet Clin N Am Eq Pract 1985;1:209–34. 54. Rivas LJ, Hinchcliff KW, Robertson JT. Cervical meningomyelocele associated with spina bifida in a hydrocephalic miniature colt. J Am Vet Med Assoc 1996;209(5):950–3. 55. Wong D, Winter M, Haynes J, Sponseller B, Schleining J. Dandy-Walker-like syndrome in a quarter horse colt. J Vet Intern Med 2007;21(5):1130–4. 56. Turner-Beatty M et al. Cerebellar disease in Arabian horses. Proc Am Assoc Eq Pract 1986; pp. 241–55.
57. Cho DY, Leipold HW. Myelodysplasia in a foal. Eq Vet J 1977;9:195–7. 58. Yager JA, Scott DW The skin and appendages. In: Jubb KVF, Kennedy PC (eds) The Pathology of Domestic Animals, 4th edn. San Diego, CA: Academic Press, 1993; pp. 531–738. 59. Lieto LD, Swerczek TW, Cothran EG. Equine epitheliogenesis imperfecta in two American Saddlebred foals is a lamina lucida defect. Vet Pathol 2002;39(5):576–80. 60. Lieto LD, Cothran EG. The epitheliogenesis imperfecta locus maps to equine chromosome 8 in American Saddlebred horses. Cytogenet Genome Res 2003;102(1–4): 207–10. 61. Graves KT, Henney PJ, Ennis RB. Partial deletion of the LAMA3 gene is responsible for hereditary junctional epidermolysis bullosa in the American Saddlebred Horse. Anim Genet 2009;40(1):35–41. 62. Spirito F, Charlesworth A, Linder K, Ortonne JP, Baird J, Meneguzzi G. Animal models for skin blistering conditions: absence of laminin 5 causes hereditary junctional mechanobullous disease in the Belgian horse. J Invest Dermatol 2002;119(3):684–91. 63. Stannard AA. Congenital diseases. Vet Dermatol 2000;11:211–15. 64. Blakeslee LH, Hudson RS, Hunt HR. Curly coat in horses. J Hered 1943;34:115–18. 65. Reef VB. Cardiovascular disease in the equine neonate. Vet Clin N Am Eq Pract 1985;1:117–25. 66. Greene HJ, Wray DD, Greenway JA. Two equine congenital cardiac anomalies. Irish Vet J 1975;29:115–17. 67. Prickett ME, Reeves JT, Zent WW. Tetralogy of Fallot in a thoroughbred mare. J Am Vet Med Assoc 1973;162: 552–5. 68. Rooney JR, Franks WC. Congenital cardiac anomalies in horses. Pathol Vet 1964;1:454–64. 69. Wilson AP. Persistent foramen ovale in a foal. Vet Med 1943;38:491–2. 70. Bartels JE, Vaughan JT. Persistent right aortic arch in the horse. J Am Vet Med Assoc 1969;154:4069. 71. Vitums A., Grant BD, Stone EC, Spencer GR. Transposition of the aorta and atresia of the pulmonary trunk in a horse. Cornell Vet 1973;63:41–57.
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Chapter 73
Inherited Problems of Foals Sharon J. Spier, Carrie J. Finno and Stephanie J. Valberg
Introduction A genetic disease is suspected when a specific breed is affected, when several related offspring are affected, or when developmental, congenital, or lethal traits are involved. The horse’s long gestation, single births, dispersion of horses after weaning, and existence of many diseases with delayed onset of expression or variable penetrance have hampered the identification of genetic diseases. The pace of genetic discovery of mutations responsible for inherited diseases has accelerated in recent years. The development of equine genome maps by the equine genome community1 and the complete sequencing of the horse genome performed at the Broad Institute under the auspices of the National Human Genome Research Institute have accelerated the pace of genetic discovery. Advances in genetic technologies will provide the means to identify polygenic traits in the near future. This chapter will focus on a review of the genetic diseases in the horse for which a mutation is currently known, including hyperkalemic periodic paralysis, overo lethal white syndrome, junctional epidermolysis bullosa, glycogen branching enzyme deficiency, and hereditary equine regional dermal asthenia. Two inherited diseases with known mutations, severe combined immunodeficiency (SCID) and polysaccharide storage myopathy (PSSM), are covered in other sections in this text (see Chapters 37 and 51). For each disease, the prevalence, clinical signs, etiology, diagnosis, treatment, and prognosis are mentioned.
Hyperkalemic periodic paralysis (HYPP) Hyperkalemic periodic paralysis (HYPP) is an autosomal dominant trait affecting Quarter Horses, American Paint Horses, Appaloosas, and Quarter Horse crossbred animals worldwide. The genetic
disease has been linked to the Quarter Horse sire named Impressive, and current estimates indicate that 4% of the 5 000 000 registered Quarter Horses are affected.2, 3 Affected horses may have been preferentially selected as breeding stock due to their phenotypic expression of well-developed musculature, and there is evidence of selection of HYPP-affected horses as superior halter horses by show judges.4 In 1996, the American Quarter Horse Association (AQHA) officially recognized HYPP as a genetic defect or undesirable trait, and mandatory testing for HYPP with results designated on the Registration Certificate began for foals descending from Impressive born after 1 January 1998. Foals born in 2007 or later that test homozygous affected for HYPP (H/H) are not eligible for registration by the AQHA. Clinical signs Clinical signs among horses carrying the same mutation range from asymptomatic to daily muscle fasciculations and weakness resulting in recumbency. Foals homozygous for HYPP usually show clinical signs of disease in the first few days of life. Clinical signs in H/H foals include respiratory stridor and periodic obstruction of the upper respiratory tract. Foals may present for dysphagia or respiratory distress, and endoscopic findings include pharyngeal collapse and edema, laryngopalatal dislocation, and laryngeal paralysis.5 Affected homozygous horses also exhibit dysphonia (high-pitched whinny) even between episodes. Foals that are heterozygous (N/H) are less severely affected, and typically do not demonstrate clinical signs of disease until they are weaned. By the time training is initiated, typically by 2 to 3 years of age, most N/H horses have shown intermittent clinical signs with no apparent abnormalities between episodes.6–8 In most cases, clinical episodes begin with a period of brief myotonia
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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(twitching or delayed relaxation of muscles), with some horses showing prolapse of the third eyelid. Sweating and muscle fasiculations are observed commonly in the flanks, neck, and shoulders. Stimulation and attempts to move may exacerbate muscle tremors, and some horses develop severe muscle cramping. During mild attacks, horses remain standing. In more severe attacks, clinical signs may progress to swaying, staggering, dogsitting, or recumbency within a few minutes. Horses may be tachycardic and tachypneic during episodes, yet remain relatively bright and alert. Episodes last for variable periods, usually 15–60 minutes. Several horses have died during acute episodes.9 Respiratory distress, due to paralysis of upper respiratory muscles, occurs in some animals and may require a tracheostomy. After an episode of HYPP subsides, horses appear normal. Electromyographic examination of asymptomatic HYPP horses reveals abnormal fibrillation potentials, complex repetitive discharges with occasional myotonic potentials and trains of doublets between episodes.6, 8 Episodes of HYPP may commonly be triggered by diets containing more than 1.1% of total daily intake as-fed dry weight as potassium. Feeds high in potassium include alfalfa hay, molasses, electrolyte supplements, and kelp-based supplements.10 Other precipitating factors include fasting, anesthesia or heavy sedation, trailer rides, and stress.3 Exercise does not appear to stimulate clinical signs, and serum creatine kinase shows no change or a very modest increase during episodic fasciculations and weakness. Etiology Hyperkalemic periodic paralysis is due to a missense mutation (C to G substitution) resulting in a phenylalanine → leucine substitution in the voltagedependent skeletal muscle sodium channel alpha subunit (SCN4A).7 The SCN4A gene was mapped to chromosome 11.11 In horses with HYPP, the resting membrane potential is closer to firing than in normal horses.12 Sodium channels are normally briefly activated during the initial phase of the muscle action potential. The HYPP mutation results in a failure of a subpopulation of sodium channels to inactivate when serum potassium concentrations are increased. As a result, an excessive inward flux of sodium and outward flux of potassium ensues, resulting in persistent depolarization of muscle cells followed by temporary weakness. Diagnosis Descent from the stallion Impressive on the sire’s or dam’s side in a horse with episodic muscle tremors,
weakness, or collapse is strongly suggestive of HYPP. However, as other diseases can produce similar signs (e.g., myotonia, exertional rhabdomyolysis, neurological disorders, electrolyte derangements) and veterinarians may not be present during acute episodes, the definitive test for identifying HYPP is the demonstration of the base-pair sequence substitution in SCN4A.7 Submission of mane or tail hair with roots should be made to a licensed laboratory such as the Veterinary Genetics Laboratory at the University of California at Davis (www.vgl.ucdavis.edu). In most cases, hyperkalemia (6–9 mEq/L), hemoconcentration, and mild hyponatremia occur during clinical manifestations of the disease with normal acid–base balance.8 Serum potassium concentration returns to normal after the cessation of clinical signs. Some affected horses may have normal serum potassium concentrations during minor episodes of muscle fasciculations.6 Treatment If horses are just beginning to exhibit clinical signs, light exercise can sometimes abort an episode in mild cases. Feeding grain or corn syrup to stimulate insulin-mediated movement of potassium across cell membranes may be of benefit. Other treatment options that may abort an episode include administration of epinephrine (adrenaline) (3 mL of 1:1000/500 kg i.m.) and administration of acetazolamide (3 mg/kg every 8–12 h orally). Many horses spontaneously recover from episodes of paralysis and may appear normal by the time a veterinarian arrives. In severe cases, administration of calcium gluconate (0.2–0.4 mL/kg of a 23% solution diluted in 1 L of 5% dextrose) will often provide immediate improvement. An increase in extracellular calcium concentration raises the muscle membrane threshold potential, which attenuates membrane hyperexcitability. To reduce serum potassium, intravenous dextrose (6 mL/kg of a 5% solution) alone or combined with sodium bicarbonate (1–2 mEq/kg) can be used to enhance intracellular movement of potassium. With severe dyspnea due to laryngeal or pharyngeal obstruction, a tracheostomy may be necessary. Control Decreasing dietary potassium and increasing renal losses of potassium are the primary steps taken to prevent HYPP episodes. Regular exercise and frequent turnout are beneficial. Horses with HYPP can graze pastures because the high water content of the pasture grass makes it unlikely that horses will consume large amounts of potassium in a short
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Inherited Problems of Foals time. Ideally, horses with recurrent episodes of HYPP should be fed a balanced diet containing between 0.6 and 1.1–1.5% total potassium concentration, and meals containing less than 33 g of potassium.10 For horses with recurrent episodes of muscle fasciculations even after dietary alterations are instituted, acetazolamide (2–3 mg/kg every 8–12 h, orally) or hydrochlorothiazide (0.5–1 mg/kg every 12 h, orally) may be helpful. These agents exert their effects through different mechanisms; however, both cause increased renal potassium ATPase activity.9 In addition, acetazolamide stabilizes blood glucose and potassium by stimulating insulin secretion. Breed registries and other associations have restrictions on the use of these drugs during competitions.
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Figure 73.1 Frame overo coat color pattern associated with overo lethal white syndrome. Courtesy of Dr Gary Magdesian.
Prognosis In most cases, HYPP is a manageable disorder; however, recurrent bouts may occur and severe episodes can be fatal. Owners of affected horses should be strongly discouraged from breeding these animals, for the long-term health of the Quarter Horse and other breeds. Since HYPP is a dominant trait, breeding an affected horse to a normal horse results in a 50% chance of producing a foal with HYPP. All affected horses share the same mutation, regardless of whether or not owners have witnessed clinical signs in their horses.13 Owners of affected horses should advise veterinarians of HYPP status before anesthesia or procedures requiring heavy sedation as anesthesia has been associated with cardiac or respiratory arrest in affected horses. Horses of Impressive lineage should be tested for HYPP during pre-purchase examination. Registered horses will have the results of HYPP status printed on the registration certificate.
Ileocolonic aganglionosis (overo lethal white foal syndrome) Ileocolonic aganglionosis, or overo lethal white foal syndrome (OLWS), is an autosomal recessive trait affecting foals of American Paint Horse, Quarter Horse, and rarely Thoroughbred breeds. Breeding of carriers of OLWS may produce foals that are all white or nearly all white and die shortly after birth of colic due to functional intestinal obstruction. Some affected foals may have flecks of black hair in the mane or tail or a small black body spot. A very high incidence (> 94%) of OLWS heterozygotes is found in frame overo (Figure 73.1), highly white calico overo, and frame blend overo coat patterns.14 White coat-colored patterns with a lower incidence of heterozygous OLWS genotypes (< 21%) include tobiano (Figure 73.2), sabino, minimally blend overo, and
breeding-stock solid.14 Descriptions of coat color patterns are available at the American Paint Horse website (http://www.apha.com). It is important to note that the many solid-colored (non-white coat color pattern) horses with Paint Horse bloodlines are heterozygous, therefore the genotype cannot necessarily be inferred from coat color patterns.15 Clinical signs OLWS is characterized by a white coat and intestinal tract abnormalities that result in colic and related signs within 12 hours of birth.16 The colic is not responsive to analgesics and progressive abdominal distension will be seen with no fecal material passed. The intestinal abnormalities are due to the complete absence of intrinsic myenteric plexus in the terminal small intestine, cecum, and entire colon, with the ileum most severely affected.16–18 Embryologically, both melanocytes and myenteric ganglia cells are of neural crest origin and their failure to migrate from
Figure 73.2 Tobiano coat color pattern. Courtesy of Dr Gary Magdesian.
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the neural crest results in the absence of melanocytes in the skin and aganglionosis of the intestine.17 Some foals may be deaf and have blue eyes in addition to the pigment defects and aganglionosis.16, 18 Foals that have all white coat color patterns do not always have OLWS. Dominant white color patterns, albinos, and some foals heterozygous for OLWS may have allwhite coats without aganglionosis. Etiology The genetic defect responsible for OLWS is a single base-pair change (missense mutation) resulting in an isoleucine → lysine substitution at codon 118 of the endothelin receptor B (EDNRB) gene located on chromosome 17.19 Affected foals are homozygous for the Lys gene (Lys118/Lys118) and carriers are heterozygous (Ile118/Lys118).19 The equine Ile to Lys EDNRB substitution is in transmembrane domain 1 of a seven-transmembrane-domain G protein-coupled receptor for the endothelins. Both endothelin B receptor and endothelin 3 are essential for normal development of the enteric ganglia and melanocytes within the neural crest.20, 21 Diagnosis A white foal with signs of colic that does not pass any meconium is almost pathognomonic for the disease; however, radiographs, contrast studies, and an ultrasound examination may be indicated to document complete bowel obstruction. White Paint foals without evidence of colic may not be homozygous for the OLWS mutation and should be genetically tested. The genetic test is also useful to determine carrier status, especially for non-frame overos, tobianos, and out-cropped Quarter Horses (horses with white color markings). Testing is available through the Veterinary Genetics Laboratory at the University of California at Davis (www.vgl.ucdavis.edu). Treatment/prognosis There is no treatment available and the prognosis is invariably grave.
Junctional epidermolysis bullosa (JEB) Junctional epidermolysis bullosa is an autosomal recessive trait affecting Belgians, other draft breeds, and American Saddlebred horses.22–25 A mutation causing JEB has been identified in Belgian horses. In North America, 17% of Belgian horses were found to be carriers of the mutation, and in Europe, 8% to 27% of horses of the Breton, Comtois, Vlaams Paard, and Belgische Koudbloed Flander draft horse breeds were
Figure 73.3 A 1-day-old foal with junctional epidermolysis bullosa (JEB) showing blisters, ulceration of gingiva and abnormal, serrated incisor teeth. Courtesy of Dr John Baird.
carriers.26 The draft horse JEB mutation was not identified in a screening of 107 American Saddlebreds. Clinical signs Foals are typically born alive, but irregular, reddened erosions and ulcerations develop in the skin and mouth over pressure points or after mild trauma.27 Since the skin is so fragile, intact bullae are rarely seen and blisters are commonly noted instead.28 Extensive erosions may be present at mucocutaneous junctions of the mouth, rectum, and vulva, and along the coronary bands (Figure 73.3). Granulation tissue along the coronary bands may result in separation of the coronary bands from the hoof wall and sloughing of the hooves.23 Corneal ulcers and dystrophic teeth are also described.27 The foal may have temporary incisor teeth visible at birth that are white with irregular serrated edges and pitted enamel.26 Secondary bacterial infections, scarring, impaired alimentation due to oral lesions, and death usually follow.29 Etiology In the Belgian and European draft breeds, the genetic defect responsible for JEB is a cytosine insertion
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Inherited Problems of Foals (1368insC) creating a premature stop codon in the Lamc2 gene, which encodes for the laminin γ 2 chain.30 The Lamc2 gene is located on chromosome 5.31 The truncated γ 2 subunit chain lacks the Cterminal domain so it cannot interact with the other two subunits thereby preventing the formation of laminin 5.30 Laminin 5 is widely distributed in the basement membrane of epithelial tissues. The absence of laminin 5 results in a cleft between the basement membrane zone of the dermal-epidermal junction, which is evident on microscopic evaluation of skin biopsies from affected foals.29 The mutation for JEB in American Saddlebreds has not been identified but is suspected to involve the LAM α3 gene32 located on chromosome 8.31 In Saddlebreds the clinical presentation has been compared to epitheliogenesis imperfecta.24
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Etiology Glycogen branching enyzme deficiency is due to a C→A point mutation at base 102 that results in a stop codon in exon 1 of the GBE1 gene encoding glycogen branching enzyme.37 The GBE1 gene was mapped to equine chromosome 26.38 Glycogen is a required energy source in the rapidly growing fetus and neonate and is synthesized by glycogen synthase, which creates straight chains of glucose with α-1,4 glycosidic linkages, and by glycogen branching enzyme, which creates a branched structure through α-1,6 linkages. Tissues from GBED foals have no measurable GBE activity or immunodetectable GBE and cannot form normally branched glycogen.35 As a result, cardiac and skeletal muscle, liver, and the brain cannot store and mobilize glycogen to maintain normal glucose homeostasis.
Diagnosis
Diagnosis
A high index of suspicion is raised by typical clinical signs combined with skin biopsy findings of separation of the epidermis from the dermis by subepidermal clefts that are relatively free of inflammatory cells and debris.29 Definitive diagnosis in draft horses requires DNA testing for JEB, which is available through the University of California Veterinary Genetics Laboratory (www.vgl.ucdavis.edu).
Common hematological findings include a leukopenia and moderate elevations in serum creatine kinase, aspartate transaminase, and gamma-glutamyl transferase.35 Skeletal muscle, purkinje cells, or cardiac myocytes may contain basophilic globules and eosinophilic crystalline material in hematoxylin and eosin (H&E) stains.33, 35 Periodic acid–Schiff (PAS) stains are required for a histopathological diagnosis as they clearly show PAS-positive globular inclusions with decreased normal background staining for glycogen in cardiac and skeletal muscle (Figure 73.4).35 Definitive diagnosis requires identification of the mutation in GBE1 by the University of California Veterinary Genetics lab (www.vgl.ucdavis.edu) or Vet Gen (www.vetgen.com), which are licensed by the University of Minnesota to perform the test. Mane or tail hairs with roots intact or fetal liver tissue can
Treatment/prognosis There is no treatment for affected foals and they will eventually succumb to secondary infections or complete sloughing of the hooves.
Glycogen branching enzyme deficiency (GBED) Glycogen branching enyzme deficiency (GBED) is an autosomal recessive disease affecting Quarter Horse and American Paint Horse breeds. Carrier frequency estimates of 7.1% and 8.3% in the Paint and Quarter Horse breeds, respectively, have been reported.33
Clinical signs Many affected foals may be aborted or stillborn.33–35 If the foal survives to term, it may appear weak and hypothermic at birth and may progress to sudden death from hypoglycemic seizures, cardiac arrest, or respiratory failure.35, 36 Affected foals may have flexural limb deformities. All affected foals studied to date have died or been euthanized by 18 weeks of age due to the severity of muscle weakness.36
Figure 73.4 Semimembranosus muscle biopsy from a foal affected with GBED stained with periodic acid–Schiff (PAS) stain. Note the scant normal background staining for glycogen and accumulation of large globular as well as smaller crystalline inclusions of polysaccharide. (See color plate 36).
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be submitted to determine if horses are affected or are carriers of GBED.
Treatment/prognosis There is no treatment for GBED and the prognosis is grave. It is important to test aborted foals and stillborns for this disease or to test the dams for carrier status.
Hereditary equine regional dermal asthenia (hyperelastosis cutis) Hereditary equine regional dermal asthenia (HERDA), also known as hyperelastosis cutis, is an autosomal recessive trait affecting Quarter Horses and horses with Quarter Horse lineage. Carrier frequency in Quarter Horses has been recently estimated at 3.5%,39 with previous estimates ranging from 1.8% to 6.5%.40 Males and females are affected equally.41 The prevalence of HERDA is higher in horses bred for certain western performance events such as cutting and reining.
Clinical signs Heterozygotes appear clinically normal and are highly successful in athletic events. Clinical signs of HERDA are restricted to the homozygous affected horse. These are not visible at birth, but typically appear at 1.5 to 2 years of age, most often associated with initial saddling or trauma.39, 40, 42, 43 Horses homozygous for the mutation causing HERDA may present with loose, easily tented skin that does not return to its original position (Figure 73.5), seromas or hematomas, open wounds or sloughing skin (Figure 73.6), scars, and white hairs at areas of hair regrowth.41 Affected areas are most striking along the dorsum, although lesions can be found in other locations associated with trauma. Owners report that lesions heal slowly and poorly. There has been no evidence of collagen-associated abnormalities in any internal organs of affected animals at postmortem examination.41, 43
Figure 73.5 Phenotype of homozygous HERDA-affected horses. Note the extensible skin in affected tissue that can be easily separated from underlying fascia. Courtesy of Dr Stephen White.
discovered mutation causes disease is unknown and is under investigation. Diagnosis Histological examination of skin biopsies was used as a means to diagnose HERDA prior to the development of the genetic test; however, samples from grossly normal skin are not helpful in distinguishing HERDA-affected horses from those not affected.41–43 Commonly observed histological changes on skin biopsies include thinning of the dermis, as well as thinning, fragmentation, and disorientation of collagen fibers in mid to deep dermis.41,46–48 A distinctive horizontal linear zone in which separation of the collagen bundles resulted in the formation of a large empty cleft between the upper and lower
Etiology The genetic defect responsible for HERDA was identified through homozygosity mapping. The defect was initially localized to equine chromosome 1 and subsequently a G→A substitution at codon 115 was identified in equine cyclophilin B (PPIB).39 This gene likely plays a role in the protein folding of collagens, 44, 45 although the exact means by which the recently
Figure 73.6 Phenotype of homozygous HERDA-affected horses. A large hematoma developed at approximately 1.5 years of age along the dorsum of this affected horse. Courtesy of Dr Stephen White.
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Inherited Problems of Foals regions of the deep dermis has been described in two HERDA cases.41, 49 A genetic test to screen for the mutation in equine PPIB is available through the University of California Davis Lab. Treatment/prognosis At this time, there is no effective therapy for HERDA. Horses appear less likely to develop lesions during the winter and it has been suggested to keep horses indoors (protected from light) and away from other horses to prevent the development or progression of lesions.42 Affected horses are often euthanized due to the severity of lesions and associated discomfort.
References 1. Spencer G, Davis N. Horse Genome Assembled. National Institutes of Health, August 2007 (http://www. genome.gov/20519480). 2. Bowling AT, Byrns G, Spier S. Evidence for a single pedigree source of the hyperkalemic periodic paralysis susceptibility gene in quarter horses. Anim Genet 1996;27:279–81. 3. Spier SJ. Hyperkalemic periodic paralysis: 14 years later. In: Proceedings of the 52nd Annual American Association of Equine Practitioners Convention, 2006, pp. 347–50. 4. Naylor JM. Selection of Quarter Horses affected with hyperkalemic periodic paralysis by show judges. J Am Vet Med Assoc 1994;204:926–8. 5. Carr EA, Spier SJ, Kortz GD, Hoffman EP. Laryngeal and pharyngeal dysfunction in horses homozygous for hyperkalemic periodic paralysis. J Am Vet Med Assoc 1996;209:798–803. 6. Naylor JM. Equine hyperkalemic periodic paralysis: review and implications. Can Vet J 1994;35:279–85. 7. Rudolph JA, Spier SJ, Byrns G, Rojas CV, Bernoco D, Hoffman EP. Periodic paralysis in Quarter Horses: a sodium channel mutation disseminated by selective breeding. Nat Genet 1992;2:144–7. 8. Spier SJ, Carlson GP, Holliday TA., Cardinet GH III, Pickar JG. Hyperkalemic periodic paralysis in horses. J Am Vet Med Assoc 1990;197:1009–17. 9. Cox JH. An episodic weakness in four horses associated with intermittent serum hyperkalemia and the similarity of the disease to hyperkalemic periodic paralysis is man. In: Proceedings of the 31st Annual American Association of Equine Practitioners Convention, 1985, pp. 383–91. 10. Reynolds JA, Potter GD, Green LW, Wu G, Carter GK, Martin MT, Peterson TV, Murray Gerzik M, Moss G, Erkert RS. Genetic-diet interactions in the hyperkalemic periodic paralysis syndrome in Quarter horses fed varying amounts of potassium: III. The relationship between plasma potassium concentration and HYPP symptoms. J Equine Vet Sci 1998;18:731–5. 11. Chowdhary BP, Raudsepp T, Kata SR, Goh G, Millon LV, Allan V, Piumi F, Guerin G, Swinburne J, Binns M, Lear TL, Mickelson J, Murray J, Antczak DF, Womack JE, Skow LC. The first-generation whole-genome radiation hybrid map in the horse identifies conserved segments in human and mouse genomes. Genome Res 2003;13:742–51. 12. Pickar JG, Spier SJ, Snyder JR, Carlsen RC. Altered ionic permeability in skeletal muscle from horses
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16. 17. 18.
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with hyperkalemic periodic paralysis. Am J Physiol 1991;260:C926–C933. Zhou J, Spier SJ, Beech J, Hoffman EP. Pathophysiology of sodium channelopathies: correlation of normal/mutant mRNA ratios with clinical phenotype in dominantly inherited periodic paralysis. Hum Mol Genet 1994;3:1599–603. Santschi EM, Vrotsos PD, Purdy AK, Mickelson JR. Incidence of the endothelin receptor B mutation that causes lethal white foal syndrome in white-patterned horses. Am J Vet Res 2001;62:97–103. Metallinos DL, Bowling AT, Rine J. A missense mutation in the endothelin-B receptor gene is associated with Lethal White Foal Syndrome: an equine version of Hirschsprung disease. Mamm Genome 1998;9:426–31. Vonderfecht SL, Bowling AT, Cohen M. Congenital intestinal aganglionosis in white foals. Vet Pathol 1983;20:65–70. Hultgren BD. Ileocolonic aganglionosis in white progeny of overo spotted horses. J Am Vet Med Assoc 1982;180:289–92. McCabe L, Griffin LD, Kinzer A, Chandler M, Beckwith JB, McCabe ER. Overo lethal white foal syndrome: equine model of aganglionic megacolon (Hirschsprung disease). Am J Vet Res 1990;36:336–40. Santschi EM, Purdy AK, Valberg SJ, Vrotsos PD, Kaese H, Mickelson JR. Endothelin receptor B polymorphism associated with lethal white foal syndrome in horses. Mamm Genome 1998;9:306–9. Baynash AG, Hosoda K, Giaid A, Richardson JA, Emoto N, Hammer RE, Yanagisawa M. Interaction of endothelin3 with endothelin-B receptor is essential for development of epidermal melanocytes and enteric neurons. Cell 1994;79:1277–85. Hosoda K, Hammer RE, Richardson JA, Baynash AG, Cheung JC, Giaid A, Yanagisawa M. Targeted and natural (piebald-lethal) mutations of endothelin-B receptor gene produce megacolon associated with spotted coat color in mice. Cell 1994;79:1267–76. Frame SR, Harrington DD, Fessler J, Frame PF. Hereditary junctional mechanobullous disease in a foal. J Am Vet Med Assoc 1988;193:1420–4. Kohn CW, Johnson GC, Garry F, Johnson CW, Martin S, Scott DW. Mechanobullous disease in two Belgian foals. Equine Vet J 1989;21:297–301. Lieto LD, Swerczek TW, Cothran EG. Equine epitheliogenesis imperfecta in two American saddlebred foals is a lamina lucida defect. Vet Pathol 2002;39:576–80. Milenkovic D, Chaffaux S, Taourit S, Guerin G. A mutation in the LAMC2 gene causes the Herlitz junctional epidermolysis bullosa (H-JEB) in two French draft horse breeds. Genet Sel Evol 2003;35:249–56. Baird JD, Millon LV, Dileanis S, Penedo MC, Charlesworth A, Spirito F, Meneguzzi G. Junctional epidermolysis bullosa in Belgian draft horses. In: Proceedings of the 49th Annual American Association of Equine Practitioners Convention, New Orleans, LA, 2003, pp. 122–5. Shapiro J, McEwen B. Mechanobullous disease in a Belgian foal in eastern Ontario. Can Vet J 1995;36:572. Knottenbelt DC, Holdstock N, Madigan JE. Congenital abnormalities and inherited disorders. In: Knottenbelt DC, Holdstock N, Madigan JE (eds) Equine Neonatal Medicine and Surgery, vol. 1. New York: Saunders, 2004; pp. 96–153. Johnson GC, Kohn CW, Johnson CW, Garry F, Scott D, Martin S. Ultrastructure of junctional epidermolysis bullosa in Belgian foals. J Comp Pathol 1988;99:329–36. Spirito F, Charlesworth A, Linder K, Ortonne JP, Baird J, Meneguzzi G. Animal models for skin blistering
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conditions: absence of laminin 5 causes hereditary junctional mechanobullous disease in the Belgian horse. J Invest Dermatol 2002;119:684–91. Milenkovic D, Oustry-Vaiman A, Lear TL, Billault A, Mariat D, Piumi F, Schibler L, Cribiu E, Guerin G. Cytogenetic localization of 136 genes in the horse: comparative mapping with the human genome. Mamm Genome 2002;13:524–34. Lieto LD, Cothran EG. The epitheliogenesis imperfecta locus maps to equine chromosome 8 in American Saddlebred horses. Cytogenet Genome Res 2003;102:207–210. Wagner ML, Valberg SJ, Ames EG, Bauer MM, Wiseman JA, Penedo MC, Kinde H, Abbitt B, Mickelson JR. Allele frequency and likely impact of the glycogen branching enzyme deficiency gene in Quarter Horse and Paint Horse populations. J Vet Int Med 2006;20:1207–11. Render JA, Common RS, Kennedy FA, Jones MZ, Fyfe JC. Amylopectinosis in fetal and neonatal Quarter Horses. Vet Pathol 1999;36:157–60. Valberg SJ, Ward TL, Rush B, Kinde H, Hiraragi H, Nahey D, Fyfe J, Mickelson JR. Glycogen branching enyzme deficiency in quarter horse foals. J Vet Intern Med 2001;15:572–80. Valberg SJ, Mickelson J. Glycogen branching enzyme deficiency. In: Proceedings of the 52nd Annual American Association of Equine Practitioners Convention, 2006, pp. 351–3. Ward TL, Valberg SJ, Adelson DL, Abbey CA, Binns MM, Mickelson JR. Glycogen branching enzyme (GBE1) mutation causing equine glycogen storage disease IV. Mamm Genome 2004;15:570–7. Ward TL, Valberg SJ, Lear TL, Guerin G, Milenkovic D, Swinburne JE, Binns MM, Raudsepp T, Skow L, Chowdhary BP, Mickelson J R. Genetic mapping of GBE1 and its association with glycogen storage disease IV in American Quarter Horses. Cytogenet Genome Res 2003;102:201–6. Tryon RC, White SD, Bannasch DL. Homozygosity mapping approach identifies a missense mutation in equine cyclophilin B (PPIB) associated with HERDA in the American Quarter Horse. Genomics 2007;90:93–102.
40. Tryon RC, White SD, Famula TR, Schultheiss PC, Hamar DW, Bannasch DL. Inheritance of hereditary equine regional dermal asthenia in Quarter Horses. Am J Vet Res 2005;66:437–42. 41. White SD, Affolter VK, Bannasch DL, Schultheiss PC, Hamar DW, Chapman PL, Naydan D, Spier SJ, Rosychuk RA, Rees C, Veneklasen GO, Martin A, Bevier D, Jackson HA, Bettenay S, Matousek J, Campbell KL, Ihrke P J. Hereditary equine regional dermal asthenia (“hyperelastosis cutis”) in 50 horses: clinical, histological, immunohistological and ultrastructural findings. Vet Dermatol 2004;15:207–17. 42. Rashmir-Raven AM, Winand NJ, Read RW, Hopper RM, PL Ryan, Poole MH, Erb HN. Equine hyperelastosis eutis update. In: Proceedings of the 50th Annual American Association of Equine Practitioners Convention, 2004, pp. 47– 50. 43. White SD, Affolter VK, Schultheiss PC, Ball BA, Wessel MT, Kass P, Molinaro AM, Bannasch DL, Ihrke PJ. Clinical and pathological findings in a HERDA-affected foal for 1. 5 years of life. Vet Dermatol 2007;18:36–40. 44. Bachinger HP. The influence of peptidyl-prolyl cis-trans isomerase on the in vitro folding of type III collagen. J Biol Chem 1987;262:17144–8. 45. Steinmann B, Bruckner P, Superti-Furga A. Cyclosporin A slows collagen triple-helix formation in vivo: indirect evidence for a physiologic role of peptidyl-prolyl cis-transisomerase. J Biol Chem 1991;266:1299–303. 46. Hardy MH, Fisher KR, Vrablic OE, Yager JA, NimmoWilkie JS, Parker W, Keeley FW. An inherited connective tissue disease in the horse. Lab Invest 1988;59:253– 62. 47. Lerner DJ, McCracken MD. Hyperelastosis in 2 horses. J Equine Med Surg 1978;2:350–2. 48. Stannard AA. Stannard’s illustrated equine dermatology notes. Vet Dermatol 2000;11:211–15. 49. Brounts SH, Rashmir-Raven AM, Black SS. Zonal dermal separation: a distinctive histopathological lesion associated with hyperelastosis cutis in a Quarter Horse. Vet Dermatol 2001;12:219–24.
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Gastroduodenal Ulcer Complex of the Older Foal Bonnie Barr
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75. Rhodococcus Pneumonia: Epidemiology and Prophylaxis Noah D. Cohen
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76. Rhodococcus Pneumonia: Pathogenesis, Diagnosis and Therapy ` Steeve Giguere
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Strangles John F. Timoney
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Other Bacterial Respiratory Diseases of the Older Foal ` Steeve Giguere
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Viral Respiratory Diseases of the Older Foal Lutz S. Goehring and D. Paul Lunn
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Neurological Conditions (EPM, EMND Wobblers, Trauma, etc) Guy D. Lester
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Diarrhea Kirsten M. Neil
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Colic Joanne Hardy
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Weight Loss Kirsten M. Neil
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Angular Limb Deformities Earl M. Gaughan and Sarah E.T. Hanna
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Developmental Orthopedic Disease C. Wayne McIlwraith
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Feeding the Growing Horse to Avoid Developmental Orthopedic Disease (DOD) Paul D. Sicilano
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Presale Endoscopy Rolf M. Embertson
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Presale Radiographs James P. Morehead and Brent N. Cassady
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Presale Radiographic Interpretation and Significance C. Wayne McIlwraith
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Chapter 74
Gastroduodenal Ulcer Complex of the Older Foal Bonnie Barr
Pathophysiology The exact cause of gastroduodenal ulcers is unknown. Initially the pathophysiological mechanism of gastric ulcer development was thought to be due to increased gastric acid production and the use of nonsteroidal drugs. It is now known that ulcer development is more complex and is described as an imbalance between protective and aggressive factors.1, 2 Hydrochloric acid and stomach pH have been incriminated as aggressive factors causing gastric ulcers, along with organic acids and pepsin. Protective factors for the gastric mucosa include maintenance of adequate blood flow, adequate mucus and bicarbonate production, prostaglandins, and gastric innervation. The glandular mucus has extensive protective mechanisms, including a bicarbonate-rich mucus layer, an extensive capillary network, and rapid restitution of epithelium when injured.3 Factors predisposing foals to gastroduodenal ulcers include use of non-steroidal anti-inflammatory drugs, stress, or a disease process that results in an imbalance between the protective and aggressive factors of the gastric mucosa. Non-steroidal anti-inflammatory drugs have been documented to cause gastroduodenal ulceration owing to the inhibition of prostaglandins which results in vasoconstriction, ischemic injury to the mucosa, and hence the development of ulcers.1 Stress due to the disease itself, transportation, herd pressures, and weaning result in changes in acid production/secretion, delay in gastric emptying, and mucosal blood flow (decrease in splanchnic blood flow). There have been a few reports of several foals on one farm being affected, leading to the belief that an infectious agent may be involved, although one has not definitively been identified. Infectious agents that have been suspected as an etiological agent include rotavirus, Clostridium species, Campylobacter species,
and Escherichia coli. In addition, many foals will have a concurrent illness or history of a prior illness.
Clinical syndromes Ulcerative disorders involving the esophagus, stomach, and duodenum have been recognized as an important condition in foals for many years. In 1986, Becht and Byars4 described four clinical syndromes of gastric ulceration in foals: silent ulcers, active ulcers, perforating ulcers, and pyloric or duodenal obstruction from healing ulcers. Silent ulcers are usually incidental findings at necropsy and most heal without treatment or apparent clinical problems. The location of these ulcers is typically in the non-glandular portion of the stomach along the margo plicatus. These lesions are most commonly seen in the younger foal. Clinically active gastric ulcers can occur as a primary problem or secondary to an underlying problem and can be seen in foals of any age. These ulcers occur in both the non-glandular and glandular portion of the stomach. Classical clinical signs of active gastric ulcers include bruxism, ptyalism, and abdominal discomfort. Other signs include depression, poor nursing, fever, or chronic poor condition. Perforation of an ulcer is a dramatic sequel to gastroduodenal ulceration. In many cases, the typical signs of gastroduodenal ulceration do not precede a perforation, but instead the foal is found acutely toxic, depressed, colicky, or dead.5 The final syndrome is associated with lesions in the gastric mucosa extending from pylorus into proximal duodenum. These lesions are usually seen in older foals and severe lesions can result in gastric outflow obstruction. Gastric emptying normally results from coordinated myoelectric activity that originates in the gastric antrum and propagates toward the pylorus and into the proximal duodenum.3 This
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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myoelectric activity is regulated by the autonomic nervous system. Stimulation of the vagus nerve increases motility and stimulation of the sympathetic nervous system inhibits motility. The composition of ingesta also determines the rate of gastric emptying.3 Inflammation, erosion, or ulceration can cause impairment of gastric emptying by affecting the myoelectric activity of the gastrointestinal tract. Ulceration that has progressed to fibrosis will result in stricture of the pylorus or proximal duodenum. Thus, severe gastroduodenal ulcers can lead to delayed gastric emptying due to a functional gastric outflow obstruction from inflammation and ulceration, a mechanical gastric outflow obstruction from fibrosis and stricture formation, or a combination of both.6–8 Clinical signs of gastric outflow obstruction are difficult to differentiate from gastric ulceration and include bruxism, pytalism, and abdominal discomfort.1, 2 Some foals may be dull and lethargic with no signs of ulcer pain; others will occasionally paw or remain recumbent. Oftentimes the foal is inappetant, but some will begin to nurse and then stop abruptly probably due to gastric discomfort. If the gastric distension is severe, the foal may present with gastric contents leaking from the nostrils. Affected foals are often unthrifty in appearance and smaller than those foals of the same age. The foal may have diarrhea or hairless areas on the rump indicating recent diarrhea. Many foals have a history of a previous episode of enteritis or enterocolitis.
Diagnostics The gold standard of diagnosis of gastroduodenal ulcers is gastroscopy. This procedure permits direct examination of the stomach and allows for proper assessment as to the presence and severity of gastroduodenal lesions. The ulcers may range from mild to severe and may be located in the glandular region of the stomach, the non-glandular region of the stomach, or both areas and extend into the proximal duodenum (Figures 74.1 and 74.2). A thorough evaluation should include evaluation of the esophagus, pylorus, and proximal duodenum. Often, foals with severe gastroduodenal ulcers have evidence of reflux esophagitis. These lesions may appear as diffuse inflammation and erosions or linear ulcerative lesions in the distal esophagus (Figure 74.3). If endoscopy is not available, but the suspected diagnosis is gastroduodenal ulcers, ulcer therapy can be started. A response to therapy confirms the diagnosis. Foals with continual abdominal discomfort, continuous gastric reflux, or minimal response to anti-ulcer medication warrant further diagnostics. Ultrasonography of the abdomen with a 7.5-MHz
Figure 74.1 Severe, diffuse gastric ulceration of the nonglandular mucosa. (See color plate 37).
transducer may reveal distended loops of small intestine, peritoneal effusion, or gastric dilation. The stomach can be imaged on the left side between rib spaces 8 and 14.9 If the gastric distension is severe, the stomach may take up the entire left side and extend ventrally. A fluid–gas interface in a severely distended stomach may be imaged on a foal with delayed gastric emptying. The proximal small intestine can be imaged in the right paralumbar fossa cranial to the right kidney.9 Abnormalities of the small intestine that may be observed include mural edema, increased wall thickness, and poor motility. Ultrasonography of the abdomen may also show findings suggestive of enteritis or enterocolitis such as fluid content within the small intestine and large intestine. Ultrasonographic evidence of a distended stomach and thick-walled proximal small intestine are strongly suggestive of delayed gastric emptying (Figure 74.4).
Figure 74.2 Ulcerations on the mucosa of the proximal duodenum in a foal. (See color plate 38).
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non-specific and are most often associated with inflammation, dehydration, and inappetence. Possible abnormalities include electrolyte abnormalities, leukopenia, neutropenia, acidosis, azotemia, and hemoconcentration. Foals that have had problems for a longer period of time may have changes in the blood profile including anemia, leukocytosis, neutrophilia, hypoproteinemia, and hyperfibrinogenemia. Anemia may be due to blood loss from ulcers or due to the chronicity of the problem. Changes in the blood profile may be helpful in pointing to the presence of concurrent disease. Fecal occult blood test is usually negative because of colonic bacterial digestion of hemoglobin. Figure 74.3 Diffuse erosions and ulcers of the distal esophagus due to reflux esophagitis. (See color plate 39).
Therapy Radiography without contrast detects only intestinal atony. Radiography with contrast agent is an additional diagnostic test that can provide useful information to determine whether the stomach is able to empty in an appropriate amount of time, which is important if there is delayed gastric emptying. The procedure is performed as follows: the stomach is completely emptied, barium contrast (10 mL/kg) is administered through a nasogastric tube, and serial standing lateral radiographs of the abdomen are taken.10 One radiograph is taken prior to barium administration to determine the correct placement of the radiograph plate, then the rest are taken immediately after barium administration and at 5, 10, 15, 20, 30, and 60 minutes after barium administration. A normal foal should have evidence of barium in the duodenum by 2–5 minutes after barium administration.10 Hematology and serum biochemistry in foals affected by gastroduodenal ulcer complex are
Figure 74.4 Ultrasound of duodenal outflow underneath the liver on the right-hand side.
The primary goal in the treatment of gastroduodenal ulcers is to suppress acid and increase stomach pH. Anti-ulcer medications include antacids, histamine type 2 receptor antagonists, proton pump inhibitors, and mucosal protectants. Antacids decrease gastric acid pH by neutralization of luminal acid. Liquid antacids include magnesium hydroxide, aluminum hydroxide, and calcium carbonate. The efficacy of these products in foals has been debated. The mechanism of action of liquid antacids is exchange of cations for anions, thus removing hydrogen ion activity from the gastric milieu.11 Studies have shown that, to be effective, these drugs need to be administered every 2 hours.11 Adverse effects include constipation with the products containing aluminum and calcium; loose stool with magnesium-containing products, and prolonged use may result in interference with absorption of electrolytes. Histamine type 2 receptor antagonists include cimetidine and ranitidine. These agents act at the histamine type 2 receptors of the parietal cells by competitively inhibiting histamine, thereby decreasing gastric acid output both during basal conditions and when stimulated by food, pentagastrin, histamine, or insulin.12–14 Ranitidine is more potent than cimetidine. Omeprazole is a proton pump inhibitor that exerts its action by selectively and irreversibly binding to parietal cell enzyme H+ ,K+ -adenosine triphosphatase (ATPase). This enzyme catalyzes the exchange of H+ for K+ ions in the final step of HCl production by parietal cells. The binding of omeprazole to this enzyme effectively decreases acid secretion regardless of the stimulus to the cell. Omeprazole irreversibly binds to H+ ,K+ -ATPase for the life of the parietal cell.13, 15, 16 Therefore, it is suitable for once a day administration.
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Table 74.1 Therapeutic agents for treating gastric ulcers Drug
Dose
Route
Frequency
Cimetidine Cimetidine Ranitidine Ranitidine Ranitidine Omeprazole Omeprazole (prevention)
12–20 mg/kg 6.6 mg/kg 6.6–7.5 mg/kg 11.3 mg/kg 1.3–1.5 mg/kg 4 mg/kg 1–2 mg/kg
p.o. i.v. p.o. p.o. i.v. p.o. p.o.
q8 hours q4–6 hours q8 hours q12 hours q8–12 hours q24 hours q24 hours
i.v., intravenous; p.o., per os.
Sucralfate is a mucosal protectant that acts locally in the treatment of gastric ulcers. Sucralfate, a complex polysaccharide that is composed of sucrose octasulfate and aluminum hydrozide, binds to negatively charged particles in the ulcerated mucosa forming an acid-resistant layer and protects against acid and pepsin. Sucralfate has been shown to enhance normal gastric defense mechanisms by stimulating mucus and bicarbonate production, deactivating pepsin, and promoting mucosal blood flow through stimulation of endogenous prostaglandins.14 Sucralfate is believed to enhance epithelial healing and stimulate cell proliferation (Table 74.1). Treatment of a foal with delayed gastric emptying due to gastroduodenal ulcers is more complex. In addition to anti-ulcer medication the foal may require intravenous fluids, frequent decompression of the stomach, prokinetic medications, intravenous antibiotics, and nutritional support. Often, these foals have electrolyte abnormalities, thus balanced polyionic fluids with appropriate supplementation is recommended. A constant-rate infusion is best, but, if it is not practical, fluids can be administered every 4–6 hours. Foals with severe gastric distension and abdominal discomfort require frequent decompression of the stomach (every 2–4 hours), thus an indwelling nasogastric tube may cause less trauma and stress to the foal than continuous passing of the nasogastric tube. Foals with hypoproteinemia due to concurrent enteric disease may require administration of plasma or hetastarch to restore oncotic pressure. Motility-enhancing drugs, such as bethanecol and metoclopramide, may be helpful at promoting gastric emptying in foals without a physical obstruction. Bethanecol is a cholinergic agonist that promotes gastric emptying and gastric motility through stimulation of the muscarinic receptors of the gastrointestinal smooth muscle.17 Bethanecol can be given by the oral route at dosages of 0.3–0.4 mg/kg every 6–8 hours or by the subcutaneous route at dosages of 0.025–0.1 mg/kg every 6–8 hours. Potential side-effects include diarrhea, inappetance, sali-
vation, and abdominal discomfort. Metoclopramide is a benzamide with the main mechanism of action through activation of 5-hydroxytryptamine receptors, which result in activation of gastrointestinal smooth muscle.18 Metoclopramide can be given at a dosage of 0.02–0.1 mg/kg by slow intravenous infusion. Adverse reactions include sedation, excitement, involuntary muscle spasms, and behavioral changes. These reactions are due to the ability of metoclopramide to cross the blood–brain barrier and antagonize dopamine receptors. Nutritional support can range from intravenous dextrose (5% solution) as an additive to fluids to partial or total parenteral nutrition. The need for intravenous nutritional support is based on the debilitation or body condition of the foal. An adequate total parenteral nutrition formula is dextrose 10 g/kg/day, amino acids 2 g/kg/day, and lipids 1 g/kg/day. The rate of administration varies according to the foal’s caloric needs, weight, and ability to tolerate the total parenteral nutrition. Usually, it is best to start at one-fourth the foal’s caloric needs and then increase it over a 12- to 24-hour period depending on the foal’s ability to tolerate the total parenteral nutrition. A foal that is on any intravenous glucose supplementation should have its blood glucose monitored every 2–6 hours. Signs of improvement include a decrease in or resolution of reflux, decrease in or resolution of the bruxism and ptyalism, improvement in attitude, resolution of the signs of abdominal discomfort, and the foal’s ability to nurse without any signs of abdominal discomfort. If medical management is not successful after 2–3 days, surgical intervention should be considered. The goal of surgical intervention is to bypass the inflamed or strictured portion of the pylorus or duodenum. Gastrojejunostomy is performed in foals with obstruction of the duodenum distal to the pylorus, whereas a gastroduodenostomy is performed in foals with obstruction of the pylorus.19, 20 A jejunojejunostomy may also be performed in addition to the procedures described above depending on the severity of the lesion. Postsurgical therapy includes allowing nothing by mouth for 2 days, appropriate fluid therapy, anti-ulcer medication, antibiotics, and nutritional support if needed. Initial reports of prognosis for survival following gastroenterostomy were 46%, but recently it was noted to be 71%.19, 20
Prevention Because the exact cause of gastroduodenal ulcers is unknown, effective preventative measures can be hard to define. Minimal usage of non-steroidal antiinflammatory drugs (NSAIDs) is recommended. If
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Gastroduodenal Ulcer Complex of the Older Foal NSAIDs are used, anti-ulcer medication can be administered at the same time. In addition, good management practices should include avoiding overcrowding, reducing stress, and limiting exposure to possible infectious diseases.
References 1. Andrews FM, Nadeau JA.. Clinical syndromes of gastric ulceration in foals and mature horses. Equine Vet J Suppl 1999;29:30–3. 2. Murray MJ. Stomach diseases of the foal. In: Mair T, Divers T, Ducharme N (eds) Equine Gastroenterology: Gastrointestinal Disease in the Foal. London: W.B. Saunders, 2002; pp. 469–76. 3. Merritt AM. Normal equine gastroduodenal secretion and motility. Equine Vet J Suppl 1999;29:7–13. 4. Becht JL, Byars TD. Gastroduodenal ulceration in foals. Equine Vet J 1986;18:307–12. 5. Borrow HA. Duodenal perforations and gastric ulcers in foals. Vet Rec 1993;132:297–9. 6. Sprayberry KA. Gastric outflow obstruction in young horses. In: Robinson NE (ed.) Current Therapy in Equine Medicine. Philadelphia: Saunders, 2003; pp 101–3. 7. Murray MJ. Gastroduodenal ulceration in foals. Equine Vet Educ 1999;11:199–207. 8. Barr BS. Duodenal stricture in a foal. In: MacLeay JM (ed.) The Veterinary Clinics of North America Equine Practice. Philadelphia: W.B. Saunders, 2006; pp. 37–42. 9. Reimer JM. The abdomen. In: Reimer JM (ed.) Atlas of Equine Ultrasonography. St. Louis: Mosby Year Book, 1998; pp. 172–242 10. Campbell M, Ackerman N, Peyton LC. Radiographic gastrointestinal anatomy of the foal. Vet Radiol 1984;25:194–204.
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11. Clark CK, Merritt AM, Burrow JA, Steible CK. Effect of aluminum hydroxide/magnesium hydroxide antacid and bismuth subsalicylate on gastric pH in horses. J Am Vet Med Assoc 1996;208:1687–91. 12. Sangiah S, MacAllister CC, Amouzadeh HR. Effects of cimetidine and ranitidine on basal gastric pH, free and total acid contents in horses. Res Vet Sci 1988;45:291– 5. 13. Murray MJ. Suppression of gastric acidity in horses. J Am Vet Med Assoc 1997;211:37–40. 14. MacAllister CG. A review of medical treatment for peptic ulcer disease. Equine Vet J Suppl 1999;29:45–9. 15. Nieto JE, Spier SJ, VanHoogmoed L, Pipers F, Timmerman B, Snyder JR. Comparison of omeprazole and cimetidine in healing of gastric ulcers and prevention of recurrence in horses. Equine Vet Educ 2001;18:260–4. 16. Buchanan B, Andrews F. Treatment and prevention of equine gastric ulcers syndrome. In: Jones S (ed.) The Veterinary Clinics of North America Equine Practice. Philadelphia: W.B. Saunders, 2003; pp. 575–97. 17. Ringger NC, Lester GD, Neuwirth L, Merritt AM, Vetro T, Harrison J. Effect of bethanechol or erythromycin on gastric emptying in horses. Am J Vet Res 1996;57:1771– 5. 18. Nieto JE, Rakestraw PC, Synder JR, Vatistas NJ. In vitro effects of erythromycin, lidocaine, and metoclopramide on smooth muscle from the pyloric antrum, proximal portion of the duodenum, and middle portion of the jejunum of horses. Am J Vet Res 2000;61:413–19. 19. Orsini JA, Donawick WJ. Surgical treatment of gastroduodenal obstructions in foals. Vet Surg 1986;15:205– 13. 20. Zedler ST, Embertson RM. Surgical resection of gastric outflow obstruction in the horse. Paper presented at the 8th International Equine Colic Research Symposium. August 3–5, 2005, Quebec City, Quebec, Canada.
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Chapter 75
Rhodococcus Pneumonia: Epidemiology and Prophylaxis Noah D. Cohen
Introduction Pneumonia caused by the bacterium Rhodococcus equi is an important cause of disease and death among foals. The objective of this chapter is to review current knowledge of the epidemiology and prevention of this disease. While the case for curative medicine tends to be more compelling than that for prevention, prophylaxis is likely to be the most effective approach for reducing the burden of R. equi pneumonia at horse breeding farms.
Epidemiology Occurrence of R. equi pneumonia among foals Pneumonia or other clinical disorders caused by infection with R. equi are rare among horses greater than 6 months of age: disease caused by R. equi occurs primarily among foals from 3 to 12 weeks of age.1, 2 Serological and ecological evidence, however, indicates that exposure of mature horses and foals to R. equi is widespread.3–5 Few reports describe the cumulative incidence or prevalence of R. equi pneumonia. The disease appears most often as a problem of foals at breeding farms. Characterizing the frequency of the disease is complicated because the distribution of disease varies by farm: some farms experience recurrent problems with the disease (so-called endemic farms), some farms only sporadically have foals affected by the disease, and some farms remain unburdened by the disease.1, 5 Thus, the occurrence of the disease is best reflected by considering farm-level
data. Another problem with estimating the occurrence of the disease is that many veterinarians eschew performing transcutaneous or transendoscopic tracheobronchial aspiration of foals 3 weeks or older with pneumonia to obtain samples for definitive diagnosis of R. equi pneumonia at farms with a previous history of R. equi pneumonia. The rationale for this approach is two-fold. First, the positive predictive value of results of non-specific diagnostic or screening testing (e.g., clinical signs of pneumonia and multifocal areas of pulmonary consolidation seen using thoracic ultrasonography) is high among foals at endemic farms. Second, other common causes of pneumonia among foals of this age (e.g., Streptococcus equi subspecies zooepidemicus) will be responsive to treatments used for presumed R. equi pneumonia. Bearing these caveats in mind, there have been a few studies involving multiple farms in Australia and the USA from which estimates of disease occurrence can be estimated.6–9 Cumulative incidences of foals with clinical signs of disease tend to be around 10%, but cumulative incidences greater than 20% are not unusual. Use of ultrasonographic screening is increasingly common to identify foals with subclinical or preclinical pulmonary consolidation,10 and the cumulative incidences of such pulmonary lesions attributed to R. equi are considerably higher, sometimes exceeding 66% of foals in a season. Reported case fatality rates derived from studies of multiple farms have generally ranged from 10% to 30%, but higher rates have been observed. Case fatality rates should be expected to be considerably lower when diagnosis is based on results of testing to detect pre- or
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Rhodococcus Pneumonia: Epidemiology and Prophylaxis subclinical disease, such as ultrasonographic findings of pulmonary consolidation in the absence of clinical signs of disease. Determinants of R. equi pneumonia among foals A common framework for considering determinants of infectious diseases is the triad of agent, environment, and host factors. This framework will be used for examining determinants of the occurrence of R. equi pneumonia in foals.
Agent factors Our understanding of agent-related factors has been enhanced by application of a variety of molecular methods, including genotyping. Host-related associations and virulence of R. equi isolates have been characterized on the basis of analysis of plasmids. Isolates from horses are primarily those bearing plasmids which encode the traA (a conjugal transferassociated gene) and vapA (a gene that encodes a surface-expressed protein associated with virulence known as the virulence-associated protein A) genes, but not the vapB gene (a gene related to vapA that also encodes a surface-expressed protein). In contrast, isolates from pigs are most often those with plasmids bearing traA and vapB (but not vapA) and bovine isolates tend to have traA but not vapA or vapB.11 Isolates obtained from foals with disease almost invariably bear a plasmid with vapA, whereas isolates from soil and the environment may carry a plasmid encoding vapA or may lack a plasmid.5, 12 Thus, equine isolates have been characterized as being of a virulent biotype (namely, those with an 85- to 95kb plasmid that includes the vapA gene) or an avirulent biotype (namely, those lacking this plasmid). Although there is some degree of variation in plasmid genotypes among regions, molecular epidemiological studies conducted to date have revealed considerable heterogeneity in chromosomal DNA of isolates of R. equi, even among isolates from foals at the same farm.9,13–16 The most practical implication of these findings is that there is no evidence that the occurrence of disease is explained by a farm-specific strain or spread of a particular isolate at the farm. Moreover, any isolate of R. equi bearing the plasmid with the pathogenicity island including the gene for vapA should be considered as capable of causing disease in susceptible foals. As will be described in the ensuing section, occurrence of vapA-positive isolates is widespread in the environment of foals. Thus, while virulent isolates of R. equi appear to be a necessary cause of R. equi, they are not sufficient alone to cause disease in foals.
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Environmental factors Data regarding the role of the environment are conflicting. A number of lines of evidence indicate that environmental factors are not of critical importance. First, serological studies indicate that most horses and foals have been exposed to R. equi.3, 4, 17 Numerous reports have documented R. equi in the feces from mares and foals, and in soil, feed, and air samples from horse breeding farms. Moreover, recent studies from our laboratory indicate that exposure to virulent isolates of R. equi at horse breeding farms is widespread: nearly all periparturient fecal samples from mares and soil samples from breeding farms contain virulent isolates of R. equi, irrespective of the R. equi disease status of either the foal or the farm, and concentrations of R. equi or virulent R. equi in feces or soil do not appear to be associated with cumulative incidence of disease.18, 19 The finding that exposure is widespread at farms, but that the disease is not, indicates that exposure to environmental isolates is not sufficient to cause disease. The concentration of virulent isolates in feces of mares or in soil samples also does not appear to be correlated with the incidence of R. equi pneumonia.9, 18, 19 There also is evidence from Texas that soil geochemistry does not differ significantly between farms affected by R. equi and farms without current or historical problems with the disease from the same region;20 the extent to which this finding is true for other geographic regions needs further exploration. Despite the aforementioned findings, the fact that the disease tends to be clustered by farm (i.e., occurs at some farms whereas others remain unaffected) suggests that there are environmental determinants of disease. Of note, the incidence of disease appears to be higher at farms that use practices generally deemed to be desirable for preventing disease (such as testing foals for passive transfer of maternal antibody, having someone attend foalings, etc.).7, 8 In this respect, R. equi pneumonia appears to be a disease of affluence. Evidence exists that the cumulative incidence of disease is higher at farms with greater density of mares and foals.6, 19 The proportions of virulent isolates in soil and foal (but not mare) fecal samples were higher at a farm in Japan that had foals affected by R. equi pneumonia than at another farm in Japan that was unaffected.21 In a study from Brazil, the cumulative incidence of foal pneumonia was lower among foals born and maintained at pasture than foals that were born in stalls and maintained in stalls at least twice daily for feeding.22 The most intriguing new environmental data have come from a group of investigators in Australia. This group has shown that the incidence of R. equi pneumonia at Thoroughbred breeding farms was
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correlated with the airborne concentration of virulent R. equi at farms, but not with soil concentrations.9 They also observed that certain environmental characteristics were associated with increased airborne concentrations of virulent R. equi, such as location (stalls, holding pens, or lanes had higher concentrations than did paddocks), warmer ambient temperature, reduced soil moisture, reduced grass height, and later date in the foaling season. Work from our laboratory has demonstrated that an increase in the proportion of virulent isolates from soil samples was positively and significantly associated with incidence of R. equi pneumonia at farms in central Kentucky.19 A challenge for the interpretation of these environmental epidemiological studies is that the temporality of infection with respect to exposure remains unclear. For associations to be causal, the exposure should precede the event. It remains to be elucidated whether the observed correlation of incidence of pneumonia with increased airborne concentrations (or proportion of virulent isolates in soil) causes foals to develop R. equi pneumonia, or whether it results from greater contamination of the environment by clinically affected foals, which amplify the organism in their gastrointestinal tract and exhale greater concentrations of the organism. The group from Australia has also demonstrated that the concentration of virulent R. equi in air exhaled by foals (i.e., air sampled near the nares of foals) that are maintained together in groups increases prior to the time of diagnosis of clinical signs of pneumonia among some members of the herd.23 These data were interpreted by the authors as suggesting the possibility that R. equi might be transmitted directly from foal to foal by aerosolization. Although data from other studies indicating direct transmission are lacking, these intriguing results merit further evaluation.
Host factors The findings that any isolate of R. equi harboring the virulence-associated plasmid appears capable of resulting in disease and that exposure to such isolates is commonplace in the environment of foals indicates that interindividual variation in susceptibility to infection is likely to play a role in the occurrence of R. equi pneumonia. This concept is supported by recent evidence that intrabronchial infection with a mutant strain of R. equi of reduced virulence, generated as a potential vaccine strain, resulted in severe pneumonia in two of five infected pony foals, while the remaining foals were unaffected clinically by the mutant strain and were protected from subsequent intrabronchial challenge with a virulent strain of R. equi.24
To date, studies have failed to identify significant associations of risk of R. equi pneumonia and a variety of foal-level factors, such as month of birth, level of maternal antibody transfer via colostrum, and parity or age of the dam.25 Evidence exists that host immunity plays a role in R. equi pneumonia. Although immunophenotypic differences do not exist at the time of disease between affected and unaffected agematched foals, the ratio of CD4+ to CD8+ lymphocytes was significantly lower at 2 weeks of age among foals that subsequently developed R. equi pneumonia than among age-matched foals from the same farms that remained free of the disease.26 In this same study, the peripheral blood neutrophil concentrations were significantly lower at 2 weeks of age in foals that subsequently developed pneumonia than among agematched foals from the same farm. Although other immunological correlates with disease are lacking, it seems likely that they exist. The disease occurs almost exclusively among foals, whereas horses ≥ 1 year old rarely have clinical signs in the absence of immunodeficiency, and R. equi is recognized most commonly in people infected with human immunodeficiency virus or who receive immunosuppressive therapy for cancer or organ transplantation.1, 12 Adaptive immune responses involving lymphocytes have been demonstrated to be critical for pulmonary clearance of experimental infection with R. equi in mice and horses.27, 28 The observation that newborn foals produce relatively less interferon-γ29, 30 indicates that younger foals might be particularly susceptible to infection.
Prophylaxis As for any infectious disease, preventing new cases is preferable to treating cases as they develop. This is particularly true for an insidious disease like R. equi pneumonia, in which clinical signs may not be apparent until the horse is severely ill. An effective vaccine has not yet been developed to prevent this disease. To date, only two interventions have proven effective for reducing the incidence of disease at farms: (i) transfusion of hyperimmune plasma and (ii) chemoprophylaxis with azithromycin. Martens et al.31 first demonstrated that transfusion of hyperimmune plasma to pony foals could prevent development of pneumonia following experimental challenge. Subsequent studies describing the impact of transfusion of hyperimmune plasma to foals to reduce the cumulative incidence in the field have yielded conflicting results (Table 75.1). Considering current knowledge, it appears that transfusion of plasma may reduce the incidence (and perhaps the severity) of disease, but it is only partially effective and may be ineffective for preventing disease
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Table 75.1 Summary of data from observational (field) studies of the efficacy of hyperimmune plasma for reducing the incidence of R. equi pneumonia (REP) among foals
Study Madigan et al.43 Mueller et al.44 Becu et al.45 Malschitzky et al.22 Hurley and Begg46 Higuchi et al.47 Giguere ` et al.48 Chaffin et al.7
Incidence of REP with plasma (a)
Incidence of REP without plasma (b)
Protective proportion 1 – (a/b)
Difference statistically significant?
3% 8% 0.2% 17% (stalled) 4% (not stalled) 27% 6% 19% 6%
43% 50% 5.8% 30% (stalled)
93% 84% 96% 44% 87% −28% 77% 37% 60%
Yes Yes Yes (historical controls) Yes, for both comparisons
21% 26% 30% 15%
in some circumstances. In the author’s opinion, it is important that plasma be administered very early in life for optimal results. Because the age at which foals become infected is unknown and because of decay of passively transferred antibodies, some veterinarians advocate administration of a second transfusion (typically 1 liter) 3–4 weeks after the initial transfusion. To date, data regarding chemoprophylaxis are limited and conflicting. In a report from Germany, administration of azithromycin (AZT) daily for the first 28 days of life did not significantly reduce the cumulative incidence of pulmonary abscessation, clinical severity score, the number of pulmonary abscesses detected sonographically per foal, or the ultrasonographically determined abscess severity score relative to untreated controls from the same farm.32 The age at diagnosis of clinical signs attributed to pulmonary abscessation, however, was significantly older among treated foals (N = 25) than the untreated controls (N = 45); none of the AZT-treated foals developed abscessation during the period of treatment. In contrast, Chaffin et al.33 have recently reported that administration of AZT during the first 2 weeks of life to foals at farms in the USA with a history of R. equi pneumonia significantly reduced the cumulative incidence of pneumonia attributed to R. equi. In this randomized, controlled trial, a total of 338 foals at 10 farms were followed from birth through weaning. Foals were randomly assigned to receive either AZT or no treatment. Although there was no formal blinding of those administering AZT to foals, the veterinarians responsible for ascertaining disease status of foals did not administer chemoprophylactic AZT and thus were unaware of the AZT status of foals. The proportion of foals that developed clinical signs attributed to R. equi pneumonia was significantly lower among the AZT group (5%) than among controls (21%). After accounting for farm effects, the estimated percent efficacy (a quantity often used to characterize vaccines) for reducing R. equi pneumo-
No No No No
nia was about 80–85%. There was no significant difference in the age at onset of clinical signs of pneumonia among affected foals in the two groups (i.e., AZT did not appear to delay the onset of clinical signs of disease). Adverse reactions attributed to AZT were not reported, and neither the proportion of foals that developed diarrhea during the first 2 or 4 weeks of life nor the reported number of days of diarrhea during the first 2 or 4 weeks of life differed significantly between groups. The reason for the discrepancy between the 2 studies is unclear, but may relate to differences in study design (randomized treatment assignment for the US study versus non-randomized, time periodbased treatment assignment in the German study, differences in the extent of blinding, etc.), case definitions, environmental/ecological differences between Europe and the USA, etc. Further work is needed to determine the efficacy of chemoprophylaxis. Resolving the apparent discrepancy will be important not only for establishing the value of chemoprophylaxis for prevention, but also for adding further evidence to support or refute the notion that most affected foals become infected early in life, perhaps at a time of enhanced susceptibility to infection. Widespread use of an antimicrobial such as AZT that is used to treat humans raises important medical and ethical concerns regarding development of antimicrobial resistance. Thus, our laboratory is exploring the efficacy of gallium maltolate, a semimetal compound that we have demonstrated to have in vitro efficacy against R. equi, even when the R. equi are within macrophages.34–36 Gallium accumulates in areas of inflammation and infection, and within macrophages, neutrophils, and many bacteria.37, 38 Trivalent gallium competes with trivalent iron for binding sites on iron-binding proteins. Unlike trivalent iron, trivalent gallium cannot be reduced to the divalent state under physiological conditions. Consequently, gallium does not enter
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ferrous-bearing molecules such as heme and thus does not interfere with oxygen transport or other vital bodily functions. Bacteria acquire protein-bound trivalent gallium instead of protein-bound trivalent iron. Gallium inhibits bacterial replication because bacterial DNA polymerases require divalent iron as a cofactor, and acquisition of gallium that cannot be reduced leads to failure of bacterial DNA polymerases. Clearly, there is great need for a vaccine, an alternative immunoprophylactic approach, or chemoprophylaxis to prevent this important disease in foals. In the absence of effective methods for prophylaxis, control of the disease is most likely to result from using screening tests to detect foals with disease that is preor subclinical (hereafter termed subclinical). The rationale for using screening tests is that detecting the disease early in its course is expected to shorten the duration and improve the clinical outcome of treatment. A variety of screening methods exist, including visual inspection, physical examination, hematological or serum assays, and thoracic imaging. Visual inspection to detect changes in attitude, nasal discharge, coughing, increased effort of breathing, or extrapulmonary disorders such as polyarthritis can be useful to detect foals suspected to have R. equi pneumonia. Detecting elevated rectal temperatures or abnormal lung sounds may be useful screening tests, although the latter is often insensitive early in the course of disease. A white blood cell concentration > 15 000 cells/µL was observed to have a sensitivity of nearly 80% and specificity of 91% as a screening test for R. equi among foals at an endemic farm; in the same study, specificity of fibrinogen concentration > 400 mg/dL was only 51%, indicating that it lacks accuracy as a screening test. 39 Although sensitivity of fibrinogen concentration > 400 mg/dL was 91%, the corresponding specificity was only 51% in that same study. Serological tests have been developed to detect antibodies against R. equi in serum, including enzymelinked immunsorbent assay (ELISA), agar-gel immunodiffusion (AGID), and serum hemolysis inhibition (SHI) methods. These tests lack accuracy for screening for,40 and for diagnosis of,40, 41 R. equi pneumonia. Serum amyloid A testing also does not appear to be useful as a screening test for R. equi.42 Thoracic imaging can be used to screen foals for evidence of R. equi pneumonia. Imaging results are likely to be more specific than those from visual inspection, physical examination, or hematology. Radiography can identify both axial and peripheral pulmonary lesions in foals that may be subclinical. Applying radiography repetitively to multiple foals can become too laborious, time-consuming, and costly to be practical. Moreover, early lesions can be
subtle and easily missed, and the radiation exposure to personnel is undesirable. Ultrasonography is increasingly used to detect foals with subclinical pneumonia attributed to R. equi infection.10 The technique obviates the need for radiation exposure, and can be efficiently and effectively used at large breeding farms. Disadvantages of sonographic screening include that R. equi is not the only agent that can cause ultrasonographically apparent pulmonary abscessation or consolidation in foals, that only peripheral pulmonary lesions can be visualized sonographically, that costs associated with repetitive examinations can be high, and that there is an increase in the apparent cumulative incidence of disease due to screening. The apparent increase in incidence results because of the identification of foals with subclinical disease that will progress to cause pneumonia (i.e., preclinical disease) and of foals with subclinical disease that might regress spontaneously without causing overt clinical signs. The proportion of foals with positive results of sonographic screening that would progress to develop clinical signs is unknown, but is presumed to be small. Thus, many foals with pulmonary lesions that would have remained subclinical are being identified and treated at breeding farms which use sonographic screening. Although it is likely that the outcome for foals with subclinical infection that would progress to clinical disease would be improved by earlier detection and treatment, there are limitations to the approach resulting from increasing the number of foals being treated for R. equi pneumonia. These limitations include increasing the risk of adverse events from treatment (such as antimicrobial-associated diarrhea) and increasing the pressure for development of antimicrobial resistance by increasing the number of treated foals. Screening tests for R. equi pneumonia need to be applied iteratively and should begin when foals are approximately 3 weeks of age. The approach to screening must be tailored to the preferences of the farm and the veterinarian(s) performing screening, and the time interval between tests should include consideration of the test to be used (e.g., visual inspection versus thoracic radiography), cost of the test, ease of the test, and resources available at the farm. There is no single screening program that is optimal for all farms, and screening can employ only a single test (e.g., ultrasonography) or multiple tests. When multiple screening tests are used, they may be used either in parallel or in series. It is imperative to discuss with all interested parties the limitations and advantages of screening for R. equi pneumonia, including the risks and benefits of treating false-positive screening test results. It bears repeating that, in the absence of effective methods for chemo- or
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Rhodococcus Pneumonia: Epidemiology and Prophylaxis immunoprophylaxis, control of R. equi pneumonia will best be achieved by the diligent application of screening tests to detect foals with subclinical disease. Further evaluation of the relative merits of screening programs is warranted.
16.
References
17.
1. Prescott JF. Rhodococcus equi: an animal and human pathogen. Clin Microbiol Rev 1991;4:20–34. 2. Gigu`ere S, Prescott JF. Clinical manifestations, diagnosis, treatment, and prevention of Rhodococcus equi infections in foals. Vet Microbiol 1997;56:313–34. 3. Hietala SK, Ardans AA, Sansome A. Detection of Corynebacterium equi-specific antibody in horses by enzyme-linked immunosorbent assay. Am J Vet Res 1985;46:13–15. 4. Takai S, Kawazu S, Tsubaki S. Enzyme-linked immunosorbent assay for diagnosis of Corynebacterium (Rhodococcus) equi infection in foals. Am J Vet Res 1985;46:2166–70. 5. Takai S. Epidemiology of Rhodococcus equi infections: a review. Vet Microbiol 1997;56:167–76. 6. Chaffin MK, Cohen ND, Martens RJ. Evaluation of equine breeding farm characteristics as risk factors for development of Rhodococcus equi pneumonia in foals. J Am Vet Med Assoc 2003;222:467–75. 7. Chaffin MK, Cohen ND, Martens RJ. Evaluation of equine breeding farm management and preventative health practices as risk factors for development of Rhodococcus equi pneumonia in foals. J Am Vet Med Assoc 2003;222:476–85. 8. Cohen, ND, O’Conor MS, Chaffin MK, Martens RJ. Farm characteristics and management practices associated with development of Rhodococcus equi pneumonia in foals. J Am Vet Med Assoc 2005;226:404–13. 9. Muscatello G, Anderson GA, Gilkerson JR, Browning GF. Associations between the ecology of virulent Rhodococcus equi and the epidemiology of R. equi pneumonia on Australian thoroughbred farms. Appl Environ Microbiol 2006;72:6152–60. 10. . Slovis, NM, McCracken JL, Mundy G. How to use thoracic ultrasound to screen foals for Rhodococcus equi at affected farms. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practice, 2005, pp. 274–8. 11. O’Campo-Sosa AA, Lewis DA, Navas J, Quigley F, Callejo R, Scortti M, Leadon DP, Fogarty U, Vazquez-Boland JA. Molecular epidemiology of Rhodococcus equi based on traA, vapA, and vapB virulence plasmid markers. J Infect Dis 2007;196:763–9. 12. Takai S, Sasaki Y, Tsubaki S. Rhodococcus equi infections in foals – current concepts and implication for future research J Equine Sci 1995;6:105–19. 13. Morton AC, Baseggio N, Peters MA, Browning GF. Diversity of isolates of Rhodococcus equi from Australian thoroughbred horse farms. Antonie van Leeuwenhoek 1998;74:21–5. 14. Morton AC, Begg AP, Anderson GA, Takai S, L¨ammler C, Browning GF. Epidemiology of Rhodococcus equi strains on Thoroughbred horse farms. Appl Environ Microbiol 2001;67:2167–75. 15. Cohen ND, Smith KE, Ficht TA, Takai S, Libal MC, West BR, DelRosario LS, Becu T, Leadon DP, Buckley T, Chaffin MK, Martens RJ. Epidemiologic study of pulsed-field gel electrophoresis of isolates of Rhodococcus equi obtained
18.
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20.
21.
22.
23.
24.
25.
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27. 28.
29.
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from horses and horse farms. Am J Vet Res 2003;64: 153–61. Venner M, Meyer Hamme B, Verspohl J, Hatori F, Shimizu N, Sasaki Y, Kakuda T, Tsubaki S, Takai S. Genotypic characterization of VapA positive Rhodococcus equi in foals with pulmonary affection and their soil environment on a warmblood horse breeding farm in Germany. Res Vet Sci 2007;83:311–17. Martens RJ, Cohen ND, Chaffin MK, Takai S, Doherty CL, Angulo AB, Edwards RE. Evaluation of 5 serologic assays to detect Rhodococcus equi pneumonia in foals. J Am Vet Med Assoc 2002;221:825–33. Grimm MB, Cohen ND, Slovis NM, Mundy GD, Harrington JR, Libal MC, Takai S, Martens RJ. Evaluation of fecal samples from mares as a source of Rhodococcus equi for their foals by use of quantitative bacteriologic culture and colony immunoblot analysis. Am J Vet Res 2007;68: 63–71. Cohen ND, Carter CN, Scott HM, Chaffin MK, Smith JL, Grimm MB, Kuskie KR, Takai S, Martens RJ. Association of soil concentrations of Rhodococcus equi and incidence of pneumonia attributable to Rhodococcus equi in foals on farms in central Kentucky. Am J Vet Res 2008;69:385–95. Martens RJ, Cohen ND, Chaffin MK, Waskom JS. Association of Rhodococcus equi foal pneumonia with farm soil geochemistry. Am J Vet Res 2002;63:95–8. Takai S, Ohbushi S, Koike K, Tsubaki S, Oishi H, Kamada M. Prevalence of virulent Rhodococcus equi in isolates from soil and feces of horses from horse-breeding farms with and without endemic infections. J Clin Microbiol 1991;29:2887–9. Malschitzky E, Neves AP, Gregory RM, Mattos RC. Reduction of stall use reduces the incidence of Rhodococcus equi infections in foals. A Hora Veterinaria 2005;24:35–8. . Muscatello G, Lowe JM, Flash ML, McBride KL, Browning GF, Gilkerson JR. Review of the epidemiology and ecology of Rhodococcus equi. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practice, 2007, pp. 214–17. Pei Y, Nicholson V, Woods K, Prescott JF. Immunization by intrabronchial administration to 1-week-old foals of an unmarked double gene disruption strain of Rhodococcus equi strain 103+. Vet Microbiol 2007;125:100–10 Chaffin MK, Cohen ND, Martens RJ, Edwards RF, Nevill M. Foal-related risk factors associated with development of Rhodococcus equi pneumonia on farms with endemic infection. J Am Vet Med Assoc 2003;223:171–9. Chaffin MK, Cohen ND, Martens RJ, Edwards RF, Nevill M, Smith 3rd R. Hematologic and immunophenotypic factors associated with development of Rhodococcus equi pneumonia of foals at equine breeding farms with endemic infection. Vet Immunol Immunopathol 2004;100:33–48. Hines SA, Kanaly ST, Byrne BA, Palmer GH. Immunity to Rhodococcus equi. Vet Microbiol 1997;56:177–85. Hines SA, Stone DM, Hines MT, Alperin DC, Knowles DP, Norton LK, Hamilton MJ, Davis WC, McGuire TC. Clearance of virulent but not avirulent Rhodococcus equi from the lungs of adult horses is associated with intracytoplasmic gamma interferon production by CD4 +and CD8+ T lymphocytes. Clin Diagn Lab Immunol 2003;10:208–15. Boyd NK, Cohen ND, Lim W-S, Martens RJ, Chaffin MK, Ball JM. Temporal changes in cytokine expression of foals during the first month of life. Vet Immunol Immunopathol 2003;92:75–85. Breathnach CC, Sturgill-Wright T, Stiltner JL, Adams AA, Lunn DP, Horohov DW. Foals are interferon
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gamma-deficient at birth. Vet Immunol Immunopathol 2006;112:199–209. Martens RJ, Martens JG, Fiske RA, Hietala SK. Rhodococcus equi foal pneumonia: protective effects of immune plasma in experimentally infected foals. Equine Vet J 1989;21:249–55. Venner M, Reinhold B, Beyerbach M, Feige K. Efficacy of azithromycin in preventing pulmonary abscesses in foals. Vet J 2009;179:301–3. Chaffin MK, Cohen ND, Martens RJ. Chemoprophylactic effects of azithromycin against Rhodococcus equi pneumonia among foals at endemic equine breeding farms. J Am Vet Med Assoc 2008;232:1035–47. Harrington JR, Martens RJ, Cohen ND, Bernstein LR. Antimicrobial activity of gallium against virulent Rhodococcus equi in vitro and in vivo. J Vet Pharmacol Ther 2006;29: 121–7. Martens RJ, Miller NA, Cohen ND, Harrington JR, Bernstein LR. Chemoprophylactic antimicrobial activity of gallium maltolate against intracellular Rhodococcus equi. J Equine Vet Sci 2007;27:341–4. Martens RJ, Mealey K, Cohen ND, Harrington JR, Chaffin MK, Taylor RJ, Bernstein LR. Pharmacokinetics of gallium maltolate after intragastric administration in neonatal foals. Am J Vet Res 2007;68:1041–4. Bernstein LR. Mechanisms of therapeutic activity for gallium. Pharmacol Rev 1998;50:665–82. Bernstein LR, Tanner T, Godfrey C, Noll B. Chemistry and pharmacokinetics of gallium maltolate, a compound with high oral gallium bioavailability. Metal Based Drugs 2000;7:33–47. Gigu`ere S, Hernandez J, Gaskin J, Miller C, Bowman JL. Evaluation of white blood cell concentration, plasma fibrinogen concentration, and an agar gel immunodiffusion test for early identification of foals with Rhodococcus equi pneumonia. J Am Vet Med Assoc 2003;222:775–81.
40. Martens RJ, Cohen ND, Chaffin MK, Takai S, Doherty CL, Angulo AB, Edwards RF. Evaluation of 5 serologic assays to detect Rhodococcus equi pneumonia in foals. J Am Vet Med Assoc 2002;221:825–33. 41. Gigu`ere S, Hernandez J, Gaskin J, Prescott JF, Takai S, Miller C. Performance of five serological assays for diagnosis of Rhodococcus equi pneumonia in foals. Clin Diagn Lab Immunol 2003;10:241–5. 42. Cohen ND, Chaffin MK, Vandenplas ML, Edwards RF, Nevill M, Moore JN, Martens RJ. Study of serum amyloid A concentrations as a means of achieving early diagnosis of Rhodococcus equi pneumonia. Equine Vet J 2005;37:212–16. 43. Madigan, JE, Hietala S, Muller N. Protection against naturally acquired Rhodococcus equi pneumonia in foals by administration of hyperimmune plasma. J Reprod Fertil Suppl 1991;44:571–8. 44. Muller N, Madigan J. Methods of implementation of an immunoprophylaxis program for the prevention of Rhodococcus equi pneumonia: results of 5 years study. In: Proceedings of the 38th Annual Convention of the American Association of Equine Practice, 1992, pp. 274–8. 45. Becu T, Polledo G, Gaskin JM. Immunoprophylaxis of Rhodococcus equi pneumonia in foals. Vet Microbiol 1997;56:193–204. 46. Hurley J, Begg A. Failure of hyperimmune plasma to prevent pneumonia caused by Rhodococcus equi in foals. Aust Vet J 1995;72:418–20. 47. Higuchi T, Arakaw A, Hashikura S, Inui T, Senba H, Takai S. Effect of prophylactic administration of hyperimmune plasma to prevent Rhodococcus equi infection on foals from endemically affected farms. J Vet Med B 1999;46:641–8. 48. Gigu`ere S, Gaskin JM, Miller C, Bowman JL. Evaluation of a commercially available hyperimmune plasma product for prevention of naturally acquired pneumonia caused by Rhodococcus equi in foals. J Am Vet Med Assoc 2002;220:59–63.
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Rhodococcus Pneumonia: Pathogenesis, Diagnosis and Therapy ` Steeve Giguere
Introduction Rhodococcus equi, a Gram-positive facultative intracellular pathogen, is one of the most common causes of pneumonia in foals between 3 weeks and 5 months of age. Although R. equi can be cultured from the environment of virtually all horse farms, the clinical disease in foals is endemic and devastating on some farms, sporadic on others, and unrecognized on most. On farms where the disease is endemic, costs associated with veterinary care, long-term therapy, and mortality of some foals may be very high. This text reviews the clinical manifestations, pathogenesis, diagnosis, and treatment of R. equi infections in foals.
Clinical manifestations The most common manifestation of R. equi infections in foals is a chronic suppurative bronchopneumonia with extensive abscessation. The slow spread of the lung infection combined with the remarkable ability of foals to compensate for the progressive loss of functional lung, make early clinical diagnosis difficult. Early clinical signs often consist only of a mild fever or a slight increase in respiratory rate that may not be apparent unless foals are exercised or stressed by handling. As the disease progresses, clinical signs may include decreased appetite, lethargy, fever, tachypnea, and labored breathing. Cough and bilateral nasal discharge are inconsistent findings. However, coughing associated with exercise (moving mares and foals to the veterinary examination area) can occasionally be useful in identifying some suspected cases.
Extrapulmonary manifestations of rhodococcal infections may also occur. Clinical signs associated with the abdominal form of the disease may include fever, depression, anorexia, weight loss, colic, and diarrhea. In one study, intestinal lesions were present in approximately 50% of foals with R. equi pneumonia presented for necropsy.1 However, the majority of foals with R. equi pneumonia do not show clinical signs of intestinal disease. Abdominal lesions may include ulcerative enterocolitis and typhlitis over the area of the Peyer’s patches, granulomatous or suppurative inflammation of the mesenteric and/or colonic lymph nodes, or in some cases a single large abdominal abscess may be the only lesion.1 Polysynovitis is present in approximately one-third of cases with chronic R. equi infections.2 However, polysynovitis is rare in subclinical cases or in cases diagnosed early in the disease process. The tibiotarsal and stifle joints are most commonly affected but, occasionally, all joints are affected. The degree of joint effusion is variable and, in most cases, lameness is mild or absent. Cytological examination of the synovial fluid usually reveals a non-septic mononuclear pleocytosis, and bacteriological culture of the synovial fluid is negative.2 Histological examination of the synovial membrane of a few affected foals revealed lymphoplasmacytic synovitis suggesting an immune-mediated process.3, 4 Immune-mediated processes may also contribute to the development of uveitis, anemia, and thrombocytopenia in some foals infected with R. equi.5 Bacteremic spread of the organism from the lungs or gastrointestinal tract may occasionally result in septic arthritis and, more commonly, osteomyelitis.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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However, foals can occasionally develop R. equi septic arthritis or osteomyelitis without apparent lung or other source of infection. R. equi vertebral osteomyelitis or diskospondylitis resulting in spinal cord compression has also been reported.6–8 Other rare extrapulmonary manifestations of R. equi infections in foals include panophthalmitis, guttural pouch empyema, sinusitis, pericarditis, nephritis, and hepatic, renal, inguinal, and intracranial abscessation.5, 9 Survival rate was significantly higher among foals without extrapulmonary diseases (EPDs) (80%) than among foals with EPDs (46%), but many EPDs were recognized only at postmortem.10
Virulence and pathogenesis The ability of R. equi to induce disease in foals likely depends on both host and microbial factors. R. equi is a facultative intracellular pathogen and its ability to persist in, and eventually destroy, alveolar macrophages seems to be the basis of its pathogenicity. Knowledge of the virulence mechanisms of R. equi were largely speculative until the discovery of a virulence plasmid.11, 12 Unlike most environmental R. equi, isolates from pneumonic foals typically contain an 80- to 90-kb plasmid. Plasmid-cured derivatives of virulent R. equi strains lose their ability to replicate and survive in macrophages.13 Plasmid-cured derivatives also fail to induce pneumonia and are completely cleared from the lungs of foals within 2 weeks following heavy intrabronchial challenge, confirming the absolute necessity of the large plasmid for the virulence of R. equi.13, 14 Nucleotide sequencing of the large plasmid of isolates from two foals revealed the presence of 69 open reading frames (ORFs).15 Comparisons of the plasmid sequence with genes previously identified in other microorganisms has identified three functional regions. Two of these regions contain genes encoding proteins involved in conjugation and in plasmid replication stability, and segregation.15 The third region of 27.5 kb bears the hallmark of a pathogenicity island and contains the genes for a family of eight closely related virulence-associated proteins designated VapA and VapC to VapI. VapA is expressed on the bacterial surface and its expression is temperature regulated, occurring between 34◦ C and 41◦ C.16 VapC, -D, and -E are secreted proteins concomitantly regulated by temperature with VapA.17 In a recent study, a R. equi mutant lacking a 7.9-kb DNA region spanning six vap genes (vapA, -C, -D, -E, -F, -I) was attenuated for virulence in mice and failed to replicate in macrophages.18 Complementation with vapA alone could restore full virulence, whereas complementation with vapC, vapD, or vapE could not. Conversely, a recombinant plasmid-cured derivative expressing wild-type levels of VapA failed to survive
and replicate in macrophages and remained avirulent for foals, showing that expression of VapA alone is not sufficient to restore the virulence phenotype.13 These findings show that, although VapA is essential for virulence, other plasmid-encoded products also contribute to the ability of R. equi to cause disease. Consistent with these findings, two R. equi mutants lacking expression of pathogenicity island genes (ORF4 and ORF8) are fully attenuated.19 All vap genes as well as five other ORFs within the pathogenicity island are upregulated when R. equi is grown in macrophage monolayers.20 Of all the vap genes, vapA, vapD, and vapG are preferentially expressed in the lungs of infected foals, and expression of these genes in the lungs is significantly higher than that achieved during in vitro growth.21 Regulation of expression of the genes of the pathogenicity island is complex and depends on at least five environmental signals, including temperature, pH, oxidative stress, magnesium, and iron.16, 20, 22 The precise role of each of these genes in the pathogenesis of R. equi infections remains to be determined. Although it has been firmly established that the virulence plasmid is essential for infection of foals it is also clear that chromosomally encoded factors, such as regulatory genes and metabolic pathways, are also important in allowing the pathogen to thrive within the host.23–25 R. equi is closely related to Mycobacterium tuberculosis, as evidenced by a partial genome sequence of R. equi which showed that the majority of R. equi genes have most homology with M. tuberculosis.26 To date, only one chromosomal gene, aceA, has been conclusively identified as a virulence factor.25 This gene encodes isocitrate lyase, a key enzyme in fatty acid metabolism. Inhalation of virulent R. equi is the major route of pneumonic infection. The incubation period following experimental intrabronchial challenge varies from approximately 9 days after administration of a heavy inoculum to approximately 2–4 weeks when a lower inoculum is administered.13, 27 Lung consolidation can be detected as early as 3 days following heavy intrabronchial challenge.13 The incubation period under field conditions is unknown but likely depends on the number of virulent bacteria inhaled as well as various host-related factors. Ingestion of the organism is a significant route of exposure, and likely also of immunization, but rarely leads to hematogenously acquired pneumonia unless the foal has multiple exposures to large numbers of bacteria.28 As opposed to foals, adult horses are typically resistant to R. equi infections. Clearance of R. equi in adult horses is associated with lymphoproliferative responses to R. equi antigens, development of R. equispecific cytotoxic T lymphocytes, and interferon-γ (IFN-γ ) induction.29–32 The recent finding that young foals are deficient in their ability to produce IFN-γ in
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response to mitogens has led to the hypothesis that an IFN-γ deficiency may be at the basis of their peculiar susceptibility to R. equi infections.33 However, the fact that oral administration of live virulent R. equi to newborn foals confers complete protection against subsequent heavy intrabronchial challenge34, 35 and the fact that most foals on farms where the disease is endemic do not develop disease or develop subclinical disease and eventually clear the infection unequivocally, demonstrates that most foals have the ability to mount protective immune responses to R. equi. Consistent with these findings, experimental infection of young foals with R. equi results in IFN-γ induction and antibody responses similar to or greater than that of adult horses undergoing the same experimental challenge.36
Diagnosis The distinction between lower respiratory tract infections caused by R. equi and that caused by other pathogens is problematic especially on farms with no previous history of R. equi infections. Many diagnostic tests, including complete blood count (CBC), measurement of fibrinogen concentrations, ultrasonography, and radiography, may help distinguish pneumonia caused by R. equi from that caused by other pathogens. However, bacteriological culture and/or polymerase chain reaction (PCR) amplification of the vapA gene combined with cytological examination of a tracheobronchial aspirate (TBA) are necessary to make a definitive diagnosis of pneumonia caused by R. equi. Hyperfibrinogenemia is the most consistent laboratory finding in foals with R. equi pneumonia, although rare cases may have normal fibrinogen concentrations. Neutrophilic leukocytosis with or without monocytosis is also common. In a review of 40 cases of lung abscesses there was a trend toward higher fibrinogen concentrations and white blood cell (WBC) counts for the foals from which R. equi was isolated from a TBA as opposed to foals from which another pathogen was isolated from a TBA.37 However, there was considerable overlap in ranges, which precludes the use of fibrinogen concentrations and WBC counts as diagnostic tests or prognostic indicators for an individual animal. Thoracic radiography is useful in evaluating the severity of pneumonia and in assessing response to therapy. A prominent alveolar pattern characterized by ill-defined regional consolidation is the most common radiographic abnormality (Figure 76.1). The consolidated lesions are often seen as more discrete nodular and cavitary lesions consistent with pulmonary abscessation (Figure 76.1). Ultrasonography is a helpful diagnostic tool when lung involvement includes peripheral areas, but may not be as useful as
Figure 76.1 Thoracic radiograph from a 3-month-old foal with pneumonia caused by Rhodococcus equi. The study shows a combination of alveolar and nodular interstitial patterns. Additionally, cavitary lesions consistent with pulmonary abscessation can be seen dorsally.
radiography to evaluate the full extent of lung lesions since abscesses with overlying aerated lung will not be detected. However, in most horses and foals with pulmonary abscessation, the periphery of the lung is affected, enabling the ultrasonographer to successfully image some of the abscesses. Early ultrasonographic lesions are non-specific and may only include irregularities of the pleural surface. These lesions may progress to form focal areas of consolidation of various sizes (Figure 76.2). In more chronic cases, well-circumscribed, encapsulated abscesses can be detected (Figure 76.3). Ultrasonography is very useful in evaluating the severity of pneumonia and in
Figure 76.2 Sonogram of the left thorax in a foal with mild pneumonia caused by Rhodococcus equi. There is an approximately 1 cm2 area of focal consolidation.
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Figure 76.3 Sonogram of the left thorax in a foal with severe pneumonia caused by Rhodococcus equi. There is consolidation of the ventral aspect of the lung surrounding a large encapsulated abscess.
assessing response to therapy, especially for equine practitioners who do not have access to thoracic radiography. Ultrasonography is also a useful tool for detection of some abdominal abscesses (Figure 76.4) and for screening for R. equi-infected foals on farms where the disease is endemic (see Control). PCR amplification of the vapA gene is more sensitive than bacterial culture.38–40 However, culture offers the advantage of detecting other bacterial pathogens present and permits in vitro susceptibility testing of the recovered pathogens. As a result, PCR amplification of the vapA gene may be done in association with, but should not replace, bacterial culture. On endemic farms, many foals without clinical disease have R. equi in their trachea as a result of inhalation of contaminated dust or as a result of a subclinical infection.41 For this reason, culture or PCR amplification of R. equi from a TBA should be interpreted in the context of cytological evalua-
Figure 76.5 Wright–Giemsa stain of tracheobronchial fluid collected from a 2-month-old foal with clinical signs of respiratory disease showing septic suppurative inflammation with pleomorphic cocobacilli (arrow) suggestive of Rhodococcus equi.
tion and clinical examination (Figure 76.5). A light growth of R. equi from a foal with no clinical signs of respiratory disease, no cytological evidence of airway inflammation, and no ultrasonographic or radiographic evidence of pulmonary lesions is likely an incidental finding. Several independent studies have evaluated the performance of available serological tests for diagnosis of infection caused by R. equi on endemic farms. The serological tests evaluated were found to have either low sensitivity or low specificity, or both.42, 43 Improving either sensitivity or specificity of enzymelinked immunosorbent assays (ELISAs) by changing the cut-off value of the tests could be done only at the detriment of the other. The performance of each assay was not improved by sequential sampling of each foal at 2-week intervals over time.43 Reliance on serology as the sole diagnostic test for R. equi infections results in overdiagnosis of the disease and in missing infections in the early stages. Serological tests may be more useful at the farm level to detect overall exposure, but they have little value in establishing or excluding a diagnosis of R. equi pneumonia.
Treatment
Figure 76.4 Sonogram of the abdomen obtained from a 3month-old Thoroughbred foal with severe pneumonia caused by Rhodococcus equi and multiple abdominal abscesses. The sonogram shows an abdominal abscess.
A wide variety of antimicrobial agents are active against R. equi in vitro. However, because R. equi is a facultative intracellular pathogen surviving and replicating in macrophages and therefore causes granulomatous lesions with thick caseous material, many of these drugs are ineffective in vivo. For example, in one study all 17 foals with R. equi pneumonia treated with the combination of penicillin and gentamicin died despite the fact that all isolates were sensitive to gentamicin.2 The combination of
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Table 76.1 Recommended dosages, minimum inhibitory concentrations, as well as serum and pulmonary pharmacokinetics of macrolides commonly used for the treatment of Rhodococcus equi infections in foals.
Recommended oral dosage (mg/kg) MIC90 of R. equi isolates (µg/mL) Serum pharmacokinetics Oral bioavailability (%) T 1/2 (h) C max (µg/mL) C min (µg/mL) Pulmonary pharmacokineticsb BAL cells T 1/2 (h) BAL cells C max (µg/mL) PELF T 1/2 (h) PELF C max (µg/mL) a b
Erythromycin
Clarithromycin
Azithromycin
25 q. 6–8 h 0.25
7.5 q. 12 h 0.12
10 q. 24–48 ha 1.00
18 1 1.10 0.13
57 5.4 0.88 0.20
56 16 0.68 0.23
ND 1.02 ND ND
10.7 74.2 8.6 49.9
54.4 49.9 34.8 10.0
Administration q. 24 hours for 5 days and q. 48 hours thereafter. Pulmonary pharmacokinetics following administration of a single dose of 10 mg/kg by nasogastric tube.
MIC90 , minimum inhibitory concentration that inhibit growth of at least 90% of isolates; T 1/2 , half-life; C max , maximum serum concentration; C min , minimum serum concentration at the end of the dosing interval; PELF, pulmonary epithelial lining fluid; BAL, bronchoalveolar lavage; ND, not determined because quantifiable erythromycin activity was not detected in a sufficient number of foals or time points.
rifampin and erythromycin became the treatment of choice in the 1980s and has dramatically reduced foal mortality, at least compared with historical data, since its introduction.2, 44 In recent years, clarithromycin or azithromycin, two newer generation macrolides, often replace erythromycin in the combination with rifampin.45 Macrolides and rifampin are highly active against R. equi in vitro but exert only bacteriostatic activity.46 As a result, macrolides exert time-dependent activity against R. equi in vitro. Of the three macrolides listed above, clarithromycin is the most active against R. equi in vitro. The minimum inhibitory concentration at which 90% of R. equi isolates are inhibited (MIC90 ) is 0.12, 0.25, and 1.0 µg/mL, for clarithromycin, erythromycin, and azithromycin, respectively.47 The combination of a macrolide and rifampin is synergistic both in vitro and in vivo, and the use of the two classes of drugs in combination reduces the likelihood of R. equi resistance to either drug.46, 48 Rifampin and macrolides are lipid soluble, allowing them to penetrate cell membranes and caseous material. The recommended dosages are listed in Table 76.1. Several formulations of erythromycin are commercially available. Although they all show slight differences in bioavailability and elimination, they all result in therapeutic concentrations at recommended dosages. Advantages of azithromycin and clarithromycin over erythromycin in foals include enhanced oral bioavailability, prolonged half-lives, and much higher concentrations in bronchoalveolar cells and pulmonary epithelial lining fluid.49–52 These properties of the newer generation macrolides contribute to their lower dosages and longer dosing
intervals. Concentrations of clarithromycin in pulmonary epithelial lining fluid and bronchoalveolar cells of foals at steady state are considerably higher than concentrations reported following daily administration of azithromycin to foals.50, 51 However, clarithromycin concentrations at these sites decrease rapidly, whereas the release of azithromycin from cells is much slower, resulting in sustained concentrations of azithromycin in tissues for days following discontinuation of therapy.49–52 In a retrospective study, the combination clarithromycin–rifampin was significantly more effective than erythromycin– rifampin or azithromycin–rifampin, especially in foals with severe radiographic lesions.45 Tulathromycin, a semisynthetic macrolide recently approved for use in swine and cattle, was recently compared with azithromycin–rifampin for the treatment of foals with subclinical pneumonia as identified by ultrasonographic screening on a farm with a high incidence of R. equi infections. Pulmonary lesions 1 week after initiation of treatment were significantly smaller and duration of therapy was significantly shorter in foals treated with azithromycin–rifampin, indicating tulathromycin is not as effective as standard therapy.53 Tulathromycin is currently not recommended for use in foals at least until the activity of the drug against R. equi is confirmed and the drug is proven effective in foals with clinical pneumonia caused by R. equi. Tilmicosin, another macrolide approved for use in swine and cattle, is not active against R. equi.51 Resolution of clinical signs, normalization of plasma fibrinogen concentrations, and radiographic or ultrasonographic resolution of lung lesions are
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commonly used to guide the duration of therapy, which generally ranges between 2 and 12 weeks depending on the severity of the initial lesions. Although well tolerated by most foals, macrolides commonly cause diarrhea. Most of the time the diarrhea is self-limiting and does not necessitate cessation of therapy, but affected foals should be monitored carefully because some may develop severe diarrhea, leading to dehydration and electrolyte loss that necessitate intensive fluid therapy and cessation of oral macrolides. The incidence of diarrhea in foals treated with erythromycin–rifampin has ranged between 17% and 36%.45, 54 In most cases, diarrhea is mild and self-limiting. During surges of very hot weather an idiosyncratic reaction characterized by severe hyperthermia and tachypnea has been described in foals treated with erythromycin.54 Anecdotal reports suggest that these reactions may occasionally occur with newer macrolides as well. Treated foals should not be allowed outside to exercise on hot days. Administration of antipyretic drugs and placing the foal in a cool environment or cold hosing may help treat observed hyperthermia. Severe enterocolitis has also been reported in mares whose foals are being treated with erythromycin, presumably due to disruption of the mare’s normal colonic microflora following ingestion of small amounts of active drug during coprophagia or from contamination of feeders or water buckets with drug present on the foal’s muzzle.55 Even though the vast majority of R. equi isolates from foals are highly susceptible to macrolides and rifampin, resistant strains to either class of drug have been encountered. The percentage of R. equi isolates resistant to either macrolides or rifampin has ranged between 0% and 4% depending on the studies.47, 56 Rifampin should not be used in monotherapy because this increases the chance of resistance developing.46, 57 Rifampin resistance is conferred by mutations in the RNA polymerase β subunit encoded by the rpoB gene.58, 59 Progressive development of resistance to both erythromycin and rifampin during treatment is extremely rare, but it has been reported.3 The molecular mechanisms of macrolide resistance in R. equi isolates have not been determined but R. equi isolates resistant to one of the three macrolides listed above are almost invariably resistant to the other two as well. Therapy of foals developing severe diarrhea during macrolide therapy or therapy of foals infected with resistant isolates is problematic because of the limited range of alternative effective drugs. Oral doxycycline in combination with rifampin has been used successfully for the treatment of foals with pneumonia caused by R. equi. Recommended dosage for doxycycline in foals is 10 mg/kg every 12 hours orally.60 This dosage results in serum, pulmonary epithelial lining fluid, and bronchoalveolar
cell concentrations above the MIC90 of R. equi isolates (1.0 µg/mL) for the entire dosing interval. Doxycyline is bacteriostatic against R. equi but the drug is highly synergistic with rifampin and with macrolides in vitro. Therefore, doxycycline could also be combined with a macrolide for the treatment of rifampinresistant isolates. Chloramphenicol can be administered orally and achieves high concentrations within phagocytic cells in other species. The recommended dosage regimen is 50 mg/kg every 6 hours orally. However, the fact that only 70% of R. equi isolates are susceptible to this drug and the potential human health risk make this drug a less attractive alternative. High doses of a trimethoprim–sulfonamide (TMS) combination (30 mg/kg of combination every 8 or 12 hours orally) have been used alone or in combination with rifampin in foals with mild or early R. equi pneumonia or for continued therapy in foals responding well to other antimicrobials.2 However, TMS is not subjectively as effective as the combination macrolide–rifampin. In an experimental model of R. equi infection in immunosuppressed mice the most effective drugs in monotherapy were found to be, in order of effectiveness, vancomycin, imipenem, and rifampin.61 Amikacin, erythromycin, ciprofloxacin, or minocycline in monotherapy did not lead to a significant decrease in bacterial counts. The most active drugs in combination were those including vancomycin or rifampin, but these combinations were not significantly different from vancomycin monotherapy.61 Vancomycin and imipenem in foals should be used only for the treatment of life-threatening R. equi infections caused by isolates confirmed to be resistant to all other possible alternatives. Nursing care, provision of adequate nutrition and hydration, and maintaining the foal in a cool and well-ventilated environment are important. Oxygen therapy using humidified oxygen by nasopharyngeal insufflation in moderately hypoxemic foals, or by percutaneous transtracheal oxygenation in severely hypoxemic animals, is indicated.62 Judicious use of non-steroidal anti-inflammatory drugs is of value in reducing fever and improving attitude and appetite in febrile depressed anorectic foals. Nebulization may be helpful in selected cases with tenacious secretions and non-productive cough. However, most cases of R. equi pneumonia do not benefit from nebulization and the procedure is stressful to some foals. Similarly, bronchodilators are rarely helpful clinically in foals with pneumonia caused by R. equi. In addition to appropriate systemic antimicrobial therapy, foals with R. equi septic arthritis or osteomyelitis often require aggressive local therapy such as joint lavage, surgical debridement, and intravenous or intraosseous regional limb perfusion with
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Rhodococcus Pneumonia: Pathogenesis, Diagnosis and Therapy antimicrobial agents. The prognosis of foals with abdominal abscesses is poor, although rare cases will respond to long-term antimicrobial therapy. Surgical removal or marsupialization has been attempted in some foals, but abdominal adhesions usually result in the inability to resect the lesion and in chronic intermittent episodes of colic.
Prognosis Prior to the introduction of the combination of erythromycin and rifampin as the treatment of choice in the early 1980s, the prognosis of R. equi-infected foals was poor, with reported survival rates as low as 20%.63 Using erythromycin and rifampin, Hillidge44 reported a successful outcome (as assessed by survival) in 50 (88%) out of 57 foals with confirmed R. equi pneumonia. Studies in referral centers, more likely to see the most severely affected cases, have revealed survival rates ranging between 59% and 72%.5, 45, 64 Of the survivors 54% had at least one racing start as opposed to 65% for the control population, suggesting that horses contracting R. equi pneumonia as foals may be slightly less likely to race as adults. However, the racing performance of foals that raced was not significantly different from that of the US racing population.64
References 1. Zink MC, Yager JA, Smart NL. Corynebacterium equi infections in horses, 1958–1984: a Review of 131 Cases. Can J Vet Res 1986;27:213–17. 2. Sweeney CR, Sweeney RW, Divers TJ. Rhodococcus equi pneumonia in 48 foals: response to antimicrobial therapy. Vet Microbiol 1987;14:329–36. 3. Kenney DG, Robbins SC, Prescott JF, Kaushik A, Baird JD. Development of reactive arthritis and resistance to erythromycin and rifampin in a foal during treatment for Rhodococcus equi pneumonia. Equine Vet J 1994;26:246–8. 4. Madison JB, Scarratt WK. Immune-mediated polysynovitis in four foals. J Am Vet Med Assoc 1988;192:1581–4. 5. Chaffin MK, Martens RJ. Extrapulmonary disorders associated with Rhodococcus equi pneumonia in foals: retrospective study of 61 cases (1988–1996). In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 79–80. 6. Chaffin MK, Honnas CM, Crabill MR, Schneiter HL, Brumbaugh GW, Briner RP. Cauda equina syndrome, diskospondylitis, and a paravertebral abscess caused by Rhodococcus equi in a Foal. J Am Vet Med Assoc 1995;206:215– 20. 7. Gigu`ere S, Lavoie JP. Rhodococcus equi vertebral osteomyelitis in 3 Quarter Horse colts. Equine Vet J 1994;26:74–7. 8. Olchowy TW. Vertebral body osteomyelitis due to Rhodococcus equi in two Arabian foals. Equine Vet J 1994;26:79–82. 9. Janicek JC, Kramer J, Coates JR, Lattimer JC, Lacarrubba AM and Messer NT. Intracranial abscess caused by Rhodococcus equi infection in a foal. J Am Vet Med Assoc 2006;228:251–3.
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10. Reuss SM, Chaffin MK, Cohen ND. Extrapulmonary disorders associated with Rhodococcus equi infection in foals: a retrospective study of 150 Cases (1987–2007). In: Proceedings of the 54th Annual Convention of the American Association of Equine Practitioners, 2008, pp. 528. 11. Takai S, Sekizaki T, Ozawa T, Sugawara T, Watanabe Y, Tsubaki S. Association between a large plasmid and 15to 17-kilodalton antigens in virulent Rhodococcus equi. Infect Immun 1991;59:4056–60. 12. Tkachuk-Saad O, Prescott J. Rhodococcus equi plasmids: isolation and partial characterization. J Clin Microbiol 1991;29:2696–700. 13. Gigu`ere S, Hondalus MK, Yager JA, Darrah P, Mosser DM, Prescott JF. Role of the 85-kilobase plasmid and plasmid-encoded virulence-associated protein a in intracellular survival and virulence of Rhodococcus equi. Infect Immun 1999;67:3548–57. 14. Wada R, Kamada M, Anzai T, Nakanishi A, Kanemaru T, Takai S, Tsubaki S. Pathogenicity and virulence of Rhodococcus equi in foals following intratracheal challenge. Vet Microbiol 1997;56:301–12. 15. Takai S, Hines SA, Sekizaki T, Nicholson VM, Alperin DA, Osaki M, Takamatsu D, Nakamura M, Suzuki K, Ogino N, Kakuda T, Dan H, Prescott JF. DNA sequence and comparison of virulence plasmids from Rhodococcus equi ATCC 33701 and 103. Infect Immun 2000;68:6840–7. 16. Takai S, Fukunaga N, Kamisawa K, Imai Y, Sasaki Y, Tsubaki S. Expression of virulence-associated antigens of Rhodococcus equi is regulated by temperature and pH. Microbiol Immunol 1996;40:591–4. 17. Byrne BA, Prescott JF, Palmer GH, Takai S, Nicholson VM, Alperin DC, Hines SA. Virulence plasmid of Rhodococcus equi contains inducible gene family encoding secreted proteins. Infect Immun 2001;69:650–6. 18. Jain S, Bloom BR, Hondalus MK. Deletion of vapA encoding virulence associated protein a attenuates the intracellular actinomycete Rhodococcus equi. Mol Microbiol 2003;50:115– 28. 19. Ren J, Prescott JF. The effect of mutation on Rhodococcus equi virulence plasmid gene expression and mouse virulence. Vet Microbiol 2004;103:219–30. 20. Ren J, Prescott JF. Analysis of virulence plasmid gene expression of intra-macrophage and in vitro grown Rhodococcus equi ATCC 33701. Vet Microbiol 2003;94:167–82. 21. Jacks S, Gigu`ere S, Prescott JF. In vivo expression of and cell-mediated immune responses to the plasmid-encoded virulence-associated proteins of Rhodococcus equi in foals. Clin Vaccine Immunol 2007;14:369–74. 22. Benoit S, Benachour A, Taouji S, Auffray Y, Hartke A. H(2)O(2), which causes macrophage-related stress, triggers induction of expression of virulence-associated plasmid determinants in Rhodococcus equi. Infect Immun 2002;70:3768– 76. 23. Boland CA, Meijer WG. The iron dependent regulatory protein ider (dtxr) of Rhodococcus equi. FEMS Microbiol Lett 2000;191:1–5. 24. Navas J, Gonzalez-Zorn B, Ladron N, Garrido P, VazquezBoland JA. Identification and mutagenesis by allelic exchange of choE, encoding a cholesterol oxidase from the intracellular pathogen Rhodococcus equi. J Bacteriol 2001;183: 4796–805. 25. Wall DM, Duffy PS, Dupont C, Prescott JF, Meijer WG. Isocitrate lyase activity is required for virulence of the intracellular pathogen Rhodococcus equi. Infect Immun 2005;73:6736– 41.
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26. Rahman MT, Herron LL, Kapur V, Meijer WG, Byrne BA, Ren J, Nicholson VM, Prescott JF. Partial genome sequencing of Rhodococcus equi ATCC 33701. Vet Microbiol 2003;94:143–58. 27. Barton MD, Embury DH. Studies of the pathogenesis of Rhodococcus equi infection in foals. Aust Vet J 1987;64:332–9. 28. Johnson JA, Prescott JF, Markham RJ. The pathology of experimental Corynebacterium equi infection in foals following intragastric challenge. Vet Pathol 1983;20:450–9. 29. Hines MT, Paasch KM, Alperin DC, Palmer GH, Westhoff NC, Hines SA. Immunity to Rhodococcus equi: antigenspecific recall responses in the lungs of adult horses. Vet Immunol Immunopathol 2001;79:101–14. 30. Hines SA, Stone DM, Hines MT, Alperin DC, Knowles DP, Norton LK, Hamilton MJ, Davis WC, McGuire TC. Clearance of virulent but not avirulent Rhodococcus equi from the lungs of adult horses is associated with intracytoplasmic gamma interferon production by CD4(+) and CD8(+) T lymphocytes. Clin Diagn Lab Immunol 2003;10:208–15. 31. Lopez AM, Hines MT, Palmer GH, Alperin DC, Hines SA. Identification of pulmonary T-lymphocyte and serum antibody isotype responses associated with protection against Rhodococcus equi. Clin Diagn Lab Immunol 2002;9:1270–6. 32. Patton KM, McGuire TC, Fraser DG, Hines SA. Rhodococcus equi-infected macrophages are recognized and killed by CD8+ T lymphocytes in a major histocompatibility complex class I-unrestricted fashion. Infect Immun 2004; 72:7073–83. 33. Breathnach CC, Sturgill-Wright T, Stiltner JL, Adams AA, Lunn DP, Horohov DW. Foals are interferon gammadeficient at birth. Vet Immunol Immunopathol 2006;112: 199–209. 34. Chirino-Trejo JM, Prescott JF, Yager JA. Protection of foals against experimental Rhodococcus equi pneumonia by oral immunization. Can J Vet Res 1987;51:444–7. 35. Hooper-McGrevy KE, Wilkie BN, Prescott JF. Virulenceassociated protein-specific serum immunoglobulin Gisotype expression in young foals protected against Rhodococcus equi pneumonia by oral immunization with virulent R. equi. Vaccine 2005;23:5760–7. 36. Jacks S, Gigu`ere S, Crawford PC, Castleman WL. Experimental infection of neonatal foals with Rhodococcus equi triggers adult-like gamma interferon induction. Clin Vaccine Immunol 2007;14:669–77. 37. Lavoie JP, Fiset L, Laverty S. Review of 40 cases of lung abscesses in foals and adult horses. Equine Vet J 1994; 26:348–52. 38. Pusterla N, Wilson WD, Mapes S, Leutenegger CM. Diagnostic evaluation of real-time PCR in the detection of Rhodococcus equi in faeces and nasopharyngeal swabs from foals with pneumonia. Vet Rec 2007;161:272–5. 39. Rodriguez-Lazaro D, Lewis DA, Ocampo-Sosa AA, Fogarty U, Makrai L, Navas J, Scortti M, Hernandez M, Vazquez-Boland JA. Internally controlled real-time PCR method for quantitative species-specific detection and vapA genotyping of Rhodococcus equi. Appl Environ Microbiol 2006;72:4256–63. 40. Sellon DC, Besser TE, Vivrette SL, McConnico RS. Comparison of nucleic acid amplification, serology, and microbiologic culture for diagnosis of Rhodococcus equi pneumonia in foals. J Clin Microbiol 2001;39:1289–93. 41. Ardans AA, Hietala SK, Spensley MS, Sansome A. Studies of naturally occurring and experimental Rhodococcus equi. In: Proceedings of the 32nd Annual Convention of the American Association of Equine Practitioners, 1986, pp. 129–44.
42. Gigu`ere S, Hernandez J, Gaskin J, Prescott JF, Takai S, Miller C. Performance of five serological assays for diagnosis of Rhodococcus equi pneumonia in foals. Clin Diagn Lab Immunol 2003;10:241–5. 43. Martens RJ, Cohen ND, Chaffin MK, Takai S, Doherty CL, Angulo AB, Edwards RE. Evaluation of 5 serologic assays to detect Rhodococcus equi pneumonia in foals. J Am Vet Med Assoc 2002;221:825–33. 44. Hillidge CJ. Use of erythromycin-rifampin combination in treatment of Rhodococcus equi pneumonia. Vet Microbiol 1987;14:337–42. 45. Gigu`ere S, Jacks S, Roberts GD, Hernandez J, Long MT, Ellis C. Retrospective comparison of azithromycin, clarithromycin, and erythromycin for the treatment of foals with Rhodococcus equi pneumonia. J Vet Intern Med 2004; 18:568–73. 46. Nordmann P, Ronco E. In-vitro antimicrobial susceptibility of Rhodococcus equi. J Antimicrob Chemother 1992;29:383– 93. 47. Jacks S, Gigu`ere S, Nguyen A. In vitro susceptibilities of Rhodococcus equi and other common equine pathogens to azithromycin, clarithromycin and 20 other antimicrobials. Antimicrob Agents Chemother 2003;47:1742–5. 48. Prescott JF, Nicholson VM. The effects of combinations of selected antibiotics on the growth of Corynebacterium equi. J Vet Pharmacol Ther 1984;7:61–4. 49. Davis JL, Gardner SY, Jones SL, Schwabenton BA, Papich MG. Pharmacokinetics of azithromycin in foals after i.v. and oral dose and disposition into phagocytes. J Vet Pharmacol Ther 2002;25:99–104. 50. Jacks S, Gigu`ere S, Gronwall PR, Brown MP, Merritt KA. Pharmacokinetics of azithromycin and concentration in body fluids and bronchoalveolar cells in foals. Am J Vet Res 2001;62:1870–5. 51. Womble A, Gigu`ere S, Murthy YV, Cox C, Obare E. Pulmonary disposition of tilmicosin in foals and in vitro activity against Rhodococcus equi and other common equine bacterial pathogens. J Vet Pharmacol Ther 2006;29:561–8. 52. Suarez-Mier G, Gigu`ere S, Lee EA. Pulmonary disposition of erythromycin, azithromycin, and clarithromycin in foals. J Vet Pharmacol Ther 2007;30:109–15. 53. Venner M, Kerth R, Klug E. Evaluation of tulathromycin in the treatment of pulmonary abscesses in foals. Vet J 2007;174:418–21. 54. Stratton-Phelps M, Wilson WD, Gardner IA. Risk of adverse effects in pneumonic foals treated with erythromycin versus other antibiotics: 143 cases (1986–1996). J Am Vet Med Assoc 2000;217:68–73. 55. Baverud V, Franklin A, Gunnarsson A, Gustafsson A, Hellander-Edman A. Clostridium difficile associated with acute colitis in mares when their foals are treated with erythromycin and rifampicin for Rhodococcus equi pneumonia. Equine Vet J 1998;30:482–8. 56. McNeil MM, Brown JM. Distribution and antimicrobial susceptibility of Rhodococcus equi from clinical specimens. Eur J Epidemiol 1992;8:437–43. 57. Takai S, Takeda K, Nakano Y, Karasawa T, Furugoori J, Sasaki Y, Tsubaki S, Higuchi T, Anzai T, Wada R, Kamada M. Emergence of rifampin-resistant Rhodococcus equi in an infected foal. J Clin Microbiol 1997;35:1904–8. 58. Fines M, Pronost S, Maillard K, Taouji S, Leclercq R. Characterization of mutations in the RpoB gene associated with rifampin resistance in Rhodococcus equi isolated from foals. J Clin Microbiol 2001;39:2784–7. 59. Asoh N, Watanabe H, Fines-Guyon M, Watanabe K, Oishi K, Kositsakulchai W, Sanchai T, Kunsuikmengrai K,
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Rhodococcus Pneumonia: Pathogenesis, Diagnosis and Therapy Kahintapong S, Khantawa B, Tharavichitkul P, Sirisanthana T, Nagatake T. Emergence of rifampin-resistant Rhodococcus equi with several types of mutations in the RpoB gene among AIDS patients in northern Thailand. J Clin Microbiol 2003;41:2337–40. 60. Womble A, Gigu`ere S, Lee EA. Pharmacokinetics of oral doxycycline and concentrations in body fluids and bronchoalveolar cells of foals. J Vet Pharmacol Ther 2007;30: 187–93. 61. Nordmann P, Kerestedjian JJ, Ronco E. Therapy of Rhodococcus equi disseminated infections in nude mice. Antimicrob Agents Chemother 1992;36:1244–8.
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62. Hoffman AM, Viel L. A percutaneous transtracheal catheter system for improved oxygenation in foals with respiratory distress. Equine Vet J 1992;24:239–41. 63. Elissalde GS, Renshaw HW, Walberg JA. Corynebacterium equi: an interhost review with emphasis on the foal. Comp Immunol Microbiol Infect Dis 1980;3:433–45. 64. Ainsworth DM, Eicker SW, Yeagar AE, Sweeney CR, Viel L, Tesarowski D, Lavoie JP, Hoffman A, Paradis MR, Reed SM, Erb HN, Davidow E, Nalevanko M. Associations between physical examination, laboratory, and radiographic findings and outcome and subsequent racing performance of foals with Rhodococcus equi infection: 115 cases (1984–1992). J Am Vet Med Assoc 1998;213:510–15.
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Chapter 77
Strangles John F. Timoney
Introduction Equine strangles is a purulent rhinopharyngitis/tonsillitis and lymphadenitis caused by Streptococcus equi (S. equi ssp. equi) of Lancefield group C. S. equi appears to be a clonal descendant or biovar of an ancestral S. zooepidemicus (S. equi ssp. zooepidemicus) with which it shares extensive DNA homology and expresses most of the same proteins. However, host immunity engendered by the two subspecies is not cross-protective.1 Comparison of the genomic sequences of equi and zooepidemicus has revealed only a few genes unique to the highly virulent S. equi. These include pyrogenic exotoxins, iron-binding equibactin, and a few other phageencoded proteins.2, 3 Two potent antiphagocytic proteins, SeM and Se18.9, are expressed by S. equi but not by S. zooepidemicus.4, 5 S. equi but not S. zooepidemicus constitutively expresses a well-developed hyaluronic acid capsule. Consistent with its clonal status, the highly hostadapted S. equi shows very little variation in the amino acid sequences of its immunogenic proteins and so is antigenically uniform. In contrast, S. zooepidemicus exists in a large number of different serotypes and causes disease in many different animal hosts. Recent studies on phage sequences have indicated that the clonal S. equi probably emerged during domestication of the horse more than 6000 years ago. SeP9, a temperate bacteriophage that infects S. equi, but not contemporary S. zooepidemicus, shares extensive DNA sequence homology with phage 315.6 of S. pyogenes and with prophage sequences in the genome of S. equi 4047 but not of S. zooepidemicus.6 This suggests a scenario in which an ancestral S. zooepidemicus in a horse was lysogenized by a phage from S. pyogenes of human origin, a process that involved transduction of genes for the pyrogenic exotoxins from S. pyogenes. The resulting lysogeny may also have generated genetic instability resulting in a great increase and dispersion of inser-
tion elements. This in turn could have led to gene inactivation or loss of gene regulation in the ancestral clone of S. zooepidemicus.
Virulence factors Virulence factors involved in infection and the pathogenesis of strangles include the hyaluronic acid capsule, the antiphagocytic proteins SeM, Se18.9, IdeE, streptokinase, streptolysin S, pyrogenic exotoxins SePE-I, H, L, and M, tonsil-binding SzPSe, and fibronectin-binding protein FNE. The constitutively expressed hyaluronic acid capsule is optimally expressed in logarithmic-phase cultures (Figure 77.1), and functions by blocking access of surface-bound C3b and antibody to the CR1 and Fc receptors on the phagocyte. It is also important in the display of hydrophobic surface proteins, which in its absence are poorly functional. Resistance to phagocytosis is a crucial feature of S. equi infection of tonsillar and lymphatic tissue of susceptible equids. SeM functions by binding fibrinogen and limiting deposition of C3b. Se18.9 binds factor H, the complement control factor,5 and IdeE reduces the bactericidal activity of neutrophils for S. equi.
Clinical signs Immunologically naive foals are highly susceptible to S. equi infection and may experience attack rates of 90% or more. Clinical signs include sudden onset of high fever, depression, progressive swelling of mandibular and retropharyngeal lymph nodes, abscessation, nasal discharge (serous to mucoid to mucopurulent), accelerated pulse, slight cough, and, in some cases, pharyngitis and conjunctivitis (Figure 77.2). Abscesses in the retropharyngeal lymph node(s) may cause narrowing of the airway, dyspnea, painful swallowing, and an extended neck
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 77.1 Mucoid colonies of Streptococcus equi on blood agar. The colonies are asymmetric and have a flowing, honeylike texture.
posture (Figure 77.3). Local lesions in the throat may be extensive involving the tonsils, and breath odor is foul. Pneumonia is sometimes a complication in foals with a poor prognosis. In these cases, the lungs show congestion, edema, and abscesses. Mortality is also associated with extensive metastatic abscessation involving the liver, spleen, kidneys, musculature, and central nervous system (CNS). The mammary gland may become infected during nursing resulting in mastitis. Maturation of abscesses may take several days to weeks. Rupture brings clinical relief. Retropharyngeal lymph node abscesses that drain into the pharynx or guttural pouch will be associated with a profuse nasal discharge. Accumulation of pus may result in subcutaneous swelling posterior to the vertical mandible in the throat latch area. Affected foals may lose weight. Myocarditis has been reported as an aftermath of strangles in young horses but has not been well characterized electrocardiographically. Purpura hemorrhagica, a vasculitis associated with circulating immune complexes of antibody and streptococcal proteins, is rarely seen in young foals as
Figure 77.2 Mandibular lymphadenitis caused by Streptococcus equi.
Figure 77.3 Weanling pony with acute strangles showing extended neck posture and swelling in the throat latch region associated with abscessation of retropharyngeal lymph nodes.
a sequela to strangles or vaccination. Older animals that make very strong serum antibody responses to microbial proteins may develop purpura hemorrhagica with dependant edema and mucosal hemorrhages on the gums (Figure 77.4). The prognosis is poor in severe cases. Metastatic spread to the brain, thoracic and abdominal organs, or associated lymph nodes (bastard strangles) occurs in a small percentage of cases. This percentage increases with intensity and duration of exposure and so it is important to segregate acutely affected horses that are possibly shedding large numbers of S. equi.
Clinical pathology Hematological evidence of an acute phase response in the earlier stages of strangles includes elevations in white cell count (20 750 ± 1583 cells/µL),
Figure 77.4 Ecchymotic hemorrhages on the gum of a horse with purpura hemorrhagica.
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segmented neutrophils (15 059 ± 1604 cells/µL), and monocytes (6200 ± 1600 cells/µL). Plasma protein (7.08 ± 0.17 g/dL) and fibrinogen (560 ± 48 mg/dL) are also increased. Hemoglobin and packed cell volumes (PCV) may also be slightly or moderately reduced during recovery. These decreases have not been explained but are unlikely to be related to iron deficiency or chronic inflammation since the lifespan of the equine erythrocyte is 140–155 days.
Laboratory diagnosis Culture of nasal swabs, nasal washes, or pus from abscesses on Columbia CNA (colistin, nalidixic acid) agar with 5% sheep or horse blood added is essential for confirming the presence of S. equi. This may also be established by polymerase chain reaction (PCR). Nasal washes are more sensitive for the detection of small numbers of organisms because a greater surface area within the internal nares is sampled. The technique involves instilling approximately 50 mL of warm normal saline via a 15-cm length of soft rubber tubing (5–6 mm in diameter) about 12 cm into the internal nares and collecting the washings. These are centrifuged, and the pellet is cultured.7 Culture may, however, be unsuccessful during the incubation and early clinical phases. S. equi is normally not detectable on the mucosa until 24–48 hours after the onset of fever. Culture of nasal swabs may fail to detect guttural pouch carriage in apparently normal older horses for several months following recovery from strangles. Shedding of S. equi may resume later and continue sporadically. The only reliable means of identifying these carriers is endoscopic examination to confirm empyema and/or chondroids and to sample the pouch content.8 However, this is often not practical as a routine screening measure. Polymerase chain reaction combined with culture greatly increases the carrier detection rate. Some of these carriers may remain PCR-positive for months after viable organisms are cultured, suggesting persistence of DNA following death of S. equi.9 Persistence of DNA in the guttural pouch of long-term carriers contrasts with rapid disappearance of DNA from nasal washes and swabs from animals following recovery from clinical strangles.10 This is explained by efficient mucociliary clearance. PCR is at least three times as sensitive as culture in detection of S. equi on rayon- or Dacron-tipped swabs of the internal nares, or in pellets from nasal washes, and can be completed in the laboratory in a few hours.10 Its potential and use for specimens from horses about to be shipped or introduced to another farm, racetrack, show-ring, etc. are clearly evident. Gene sequences
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Figure 77.5 Serum antibody responses of six yearling horses to the antiphagocytic SeM protein of Streptococcus equi following occurrence of strangles induced by commingling exposure that began on Day 0. ELISA OD, enzyme-linked immunosorbent assay optical density.
used for design of primers include SeM, SePE-I, sod A, and 16SrRNA. PCR may be applied to samples of drinking water from communal tanks to monitor shedding by a group of animals. Recent technological advances based on thermophilic helicase-dependent amplification (tHDA) and use of a rapid detection cartridge indicate the likelihood that S. equi DNA detection will eventually become available as a test performed in the veterinarian’s office.
Serology Measurement of SeM specific antibody in sera is useful in the diagnosis of metastatic abscesses, recent infection, and purpura hemorrhagica associated with S. equi infection or vaccination. Titers of paired sera are often helpful in determining exposure and infection status. Antibody levels peak about 5 weeks after exposure and persist for 6 months (Figure 77.5).7, 11 Extract-based vaccines elicit responses that peak in 2 weeks and persist for the same period. These responses differ from those induced by infection in that they do not include IgGa (IgG1, 2) and are primarily IgGb (IgG4) and IgG(T) (IgG3, 5, 6). Responses of individual horses vary considerably; hyperresponders are at risk of immune complexmediated vasculitis should S. equi proteins enter the bloodstream. Titers in the SeM-specific enzymelinked immunosorbent assay (ELISA) range from zero to weak positive (1:200 to 1:400) to very high positive > 1:12 800 in cases of metastatic abscess or purpura. Foals that receive colostrum from high-titer mares will have correspondingly high serum antibody levels for 2 to 3 months. The half-life of the principal isotype, IgGb, in foal sera is about 32 days.
Management of outbreaks Foals born of mares that have had strangles during the previous 2 to 4 years are often resistant to
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Strangles infection for 2 to 3 months. However, this immunity is easily overwhelmed by continuing heavy challenge, and so mares and their foals should be separated in small groups during the first 2 to 3 months. Ideally, mare and foal should be accommodated in a separate paddock during this time to minimize risk of transmission of S. equi in herds with active infection. The severity of clinical disease in foals, weanlings, and yearlings is in part determined by level and duration of exposure to S. equi. Thus, it has long been known that strangles mortality in large groups of horses increases as an outbreak progresses. Also, it should be remembered that older mares with fading protective immunity may experience the catarrhal form of strangles with no clinical disease other than a purulent nasal discharge; S. equi from such a case is nevertheless highly virulent for younger, more susceptible animals. Reduction of transmission and exposure during an outbreak in a breeding population can be achieved by following these guidelines:
r r r r r r r
daily measurement of rectal temperatures of exposed animals and immediate isolation of those with increases of 1.5◦ F or more; isolation and culture in separate stalls or in small groups of clinically affected and recently febrile cases; isolation of newly arrived animals including nurse mares for 3 weeks before admission to the general population; daily disinfection of water tanks; use of separate labeled feed and water buckets for stalled horses; pasture rotation/resting for 1 month before repopulation; segregation of recently recovered cases for 3 weeks combined with culture/PCR to verify cessation of nasal shedding of S. equi before release.
Persisting infection in a breeding herd suggests the presence of one or more clinically silent carrier mares that intermittently shed S. equi from the guttural pouch. Detection of these carriers may be accomplished by separating the herd into small groups of 10 horses or less and screening by culture/PCR of nasopharyngeal swabs, or communal drinking water. Three sequential negative samples collected a few days apart are required to establish a high level of certainty that shedding has ceased. Shedder horses with guttural pouch empyema must be treated by guttural pouch lavage and debridement followed by infusion of an antimicrobial solution or gel via a Foley catheter.12
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Treatment Penicillin therapy at onset of fever is often curative. Cases with high fever, airway obstruction, and severe depression should be treated with antiinflammatory medications (flunixin meglumine, 1 mg/kg every 12–24 h; phenylbutazone, 4.4 mg/kg q12 h reducing to 2.2 mg/kg q12 h after 24 h). Emergency tracheostomy may be required in cases of airway obstruction caused by enlarging abscesses in retropharyngeal lymph nodes. Prompt administration of procaine penicillin (22,000 IU/kg i.m.), benzylpenicillin G (10 000 µg/kg) i.m. every 12–24 h, or sodium/potassium penicillin (22 000 U/kg i.v.) every 6–8 h will usually reduce swelling and bring clinical improvement within a few hours. Although producing clinical improvement, penicillin administration may not result in complete killing of S. equi in abscesses and so clinical relapses are common. In general, uncomplicated cases of strangles are best left untreated since most cases recover uneventfully and develop a high level of immunity to future exposure. Resistance to penicillin and other antibiotics including cephalosporins and macrolides with known therapeutic efficacy against S. equi has not been recorded. However, trimethoprim–sulfonamide combinations have low bactericidal activity for S. equi and should be avoided.
Immunity Acquired immunity derived from exposure to S. equi during the course of strangles stimulates a solid immunity to infection in most horses.11 Immunity persists in over 70% of animals but may be overwhelmed by intense exposure to S. equi in herds where the disease is endemic, and by stress of overcrowding and bad weather. Mares that experience a second attack of strangles are usually less severely affected and develop an afebrile purulent rhinitis (catarrhal form) with occasional moderate abscessation of the mandibular lymph nodes. Subclinical infections in nurse mares, if not recognized at time of introduction, may be the source of a strangles outbreak on a previously uninfected farm. The immunological basis of protective immunity to S. equi has not been fully characterized. Immune, resistant horses usually have high levels of serum antibody to the antiphagocytic SeM and Se18.9, to the pyrogenic exotoxins SePE-I, H, L, and M, and to a number of other surface exposed proteins found also on S. zooepidemicus.11 Se18.9, Se46.8, and SeM are the proteins most consistently and strongly reactive with IgA in nasopharyngeal washings of recently recovered horses. However, it is not known which of these
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Table 77.1 Strangles vaccines
Vaccine
Route of administration
Schedule
Adverse reactions
Live attenuated Intranasal non-encapsulated mutant
Two doses 2–3 weeks apart; annual booster
Lymphadenitis, purpura, ‘shot’ abscesses
Equilis Strep Eb
Live attenuated AroA deletion mutant
Submucosal (upper lip)
Two doses 4 weeks apart; booster every 6 months
Lymphadenitis, local inflammation, ‘shot’ abscesses
Strepguardc
Enzymic extract, adjuvanted (Havlogen)
Intramuscular
Two doses 2–3 weeks apart; annual booster
Local systemic reactions, purpura
Strepvax IId Equivax Se
Acid extract adjuvanted (aluminum hydroxide)
Intramuscular
Three doses at 3-week intervals; annual booster
Local systemic reactions, purpura
R Pinnacle I.N.a
Type
a
Fort Dodge Animal Health, Fort Dodge, Iowa. Intervet UK Ltd, Walton, Milton Keynes, UK. c Intervet, Millsboro, Delaware. d Boehringer Ingelheim, St Joseph, Missouri. e Pfizer Animal Health NZ, Auckland, New Zealand. b
IgAs, if any, participates in protective immunity. It is known that protective immunity derived from infection by S. equi induces a blocking immunity at the tonsillar level. Re-exposed horses do not exhibit anamnestic serum antibody responses to the major immunoreactive proteins SeM, SzPSe, and Se46.8.11
Vaccination Vaccines currently used to control/prevent strangles include adjuvanted hot acid and mutanolysinextracted protein-rich preparations (North America, Australia), an unencapsulated attenuated mutant (North America), and an aroA deletion mutant (UK)16 (Table 77.1). Extract vaccines are injected intramuscularly and are highly potent. They do not elicit mucosal antibodies.7 Injection-site reactions are frequent and efficacy is moderate. Foals are about 50% less likely to develop severe symptoms in the months immediately after vaccination but remain susceptible to infection and subsequent shedding.13 Mares vaccinated prepartum with extract vaccines 2 to 3 months before foaling have increased colostral levels of S. equi antibody that is subsequently passed to their foals. R A live attenuated mutant of S. equi, Pinnacle , is widely used as an intranasal vaccine in North America to induce protective immune responses in the tonsillar complex. Successfully vaccinated horses may make mucosal and serum antibody responses similar to those elicited by infection with virulent S. equi and are resistant to exposure to virulent S. equi.14 Disadvantages of the attenuated intranasal vaccine include its ability to produce abscesses in tissue when inadvertently inoculated with another injectable medica-
tion or vaccine. A low frequency of submandibular or pharyngeal lymphadenopathy, and bastard strangles abscesses have also been reported. Lack of efficacy can result from failure of a vaccine ‘take’ in the tonsillar complex or from inadequate application. There is also a slight risk of purpura with this and other forms of strangles vaccine in foals/adult horses. R Finally, an injectable, live S. equi, Equilis Strep E , in which the aroA gene has been deleted, has had limited use in the UK. The vaccine, in a dose volume of 0.2 mL, is injected into the inside of the upper lip. It has been shown to be relatively safe in most animals aged over 4 months. Deletion of aroA renders the vaccine strain unable to persist in tissue because of the inability to synthesize aromatic amino acids and p-aminobenzoic acid. Immunity generated is effective for about 3 months and so is much inferior to that accompanying recovery from naturally occurring strangles. Safety issues include painful lesions in the lip and occasional abscesses in cranial lymph nodes. The immunological basis of immunity stimulated by Equilis Strep E is not known. In general, available evidence suggests that successful vaccination against strangles, as for many other infectious diseases, is most effectively achieved by stimulation with live organisms of reduced virulence. It is, however, possible that failure of SeM, by itself or in combination with other S. equi proteins, to elicit an effective level of protective immunity following parenteral injection, is explained by inappropriate presentation. Tonsillar immunity may involve cell-mediated responses that must be stimulated by presenting the antigen(s) in an adjuvant that drives a T-helper 1 lymphocyte (Th1) and not a T-helper 2 lymphocyte (Th2) response.
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Strangles
Additional information The American Association of Equine Practitioners (AAEP) guidelines for vaccination of horses against strangles are available on the AAEP website (www.aaep.org/vaccination guidelines.htm). In addition, the American College of Veterinary Internal Medicine (ACVIM) consensus statement provides information on treatment, control, and prevention of strangles.15
8.
9.
10.
References 1. Bazely PL. Studies with streptococci. 4. Cross-immunity to Streptococcus equi. Aust Vet J 1942;18:189–94. 2. Artiushin SC, Timoney JF, Sheoran AS, Muthupalani SK. Characterization and immunogenicity of pyrogenic mitogens SePE-H and SePE-I of Streptococcus equi. Microb Pathogenesis 2002;32:71–85. 3. Heather Z, Holden MTG, Steward KF, Parkhill J, Song L, Challis GL, Robinson C, Davis-Poynter N, Waller AS. A novel streptococcal integrative conjugative element involved in iron acquisition. Mol Microbiol 2008;70:1274–92. 4. Timoney JF, Artiushin SC, Boschwitz JS. Comparison of the sequences and functions of Streptococcus equi M-like proteins SeM and SzPSe. Infect Immun 1997;65:3600–5. 5. Tiwari R, Qin A, Artiushin S, Timoney JF. Se18.9, an antiphagocytic factor H binding protein of Streptococcus equi. Vet Microbiol 2007;121:105–15. 6. Tiwari R. Bacteriophage SeP9 of Streptococcus equi and the antibacterial activity of its lysin. PhD dissertation, University of Kentucky, 2007. 7. Sheoran AS, Sponseller BT, Holmes MA, Timoney JF. Serum and mucosal antibody isotype responses to
11.
12.
13.
14.
15.
16.
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M-like protein (SeM) of Streptococcus equi in convalescent and vaccinated horses. Vet Immunol Immunopathol 1997;59: 239–51. Newton JR, Verhezen K, Talbot NC, Timoney JF, Wood JFN, Lakhani JH, Chanter N. Control of strangles outbreaks by isolation of guttural pouch carriers using PCR and culture of Streptococcus equi. Equine Vet J 2000;32:515–26. Gronbaek M, Angen O, Vigre H, Olsen SN. Evaluation of a nested PCR test and bacterial culture of nasal swabs and swabs from abscesses in relation to diagnosis of strangles. Equine Vet J 2006;38:59–63. Timoney JF, Artiushin SC. Detection of Streptococcus equi in equine nasal swabs and washes by DNA amplification. Vet Rec 1997;141:446–7. Timoney JF, Qin A, Muthupalani S, Artiushin S. Vaccine potential of novel surface exposed and secreted proteins of Streptococcus equi. Vaccine 2007;25:5583–90. Verheyen K, Newton JR, Talbot NC, DeBrauwere MN, Chanter N. Elimination of guttural pouch infection and inflammation in asymptomatic carriers of Streptococcus equi. Equine Vet J 2000;32:527–32. Hoffman AM, Staempfli HR, Prescott JF. Field evaluation of a commercial M protein vaccine against Streptococcus equi infection in foals. Am J Vet Res 1991;52:589–95. Timoney JF, Gal´an JE. The protective responses of the horse to an avirulent strain of Streptococcus equi. In: Kimura Y, Kotami S, Shiokawa Y. (eds) Recent Advances in Streptococci and Streptococcal Diseases. Reedbooks Ltd, 1985; pp. 294–5. Sweeney CR, Timoney JF, Newton JR, Hines MT. Streptococcus equi infections in horses: Guidelines for treatment control and prevention of strangles. J Vet Intern Med 2005; 19: 123–34. Jacobs, A.A.C., Goovaerts, D., Nuijten, P.J.M., Theelen, R.P.H., Hartford, O.M., and Foster, T.J. 2000. Investigation towards an efficacious and safe strangles vaccine: submucosal vaccination with a live attenuated Streptococcus equi. Vet. Rec. 147:563-567.
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Chapter 78
Other Bacterial Respiratory Diseases of the Older Foal ` Steeve Giguere
Introduction Bronchopneumonia is one of the leading causes of both morbidity and mortality in foals between 1 and 6 months of age.1 Foal pneumonia has a major financial impact on the horse industry. Not only does pneumonia account for significant mortality on some farms but therapy is expensive and the effect on subsequent performance of the equine athlete may be substantial if the disease is not recognized and treated early. The morbidity rate has been estimated between 6% and 9% across the USA.1, 2 It is likely, however, that the true incidence of distal respiratory tract infection is much higher and that many cases go unrecognized and resolve spontaneously. Indeed, careful weekly physical examination of more than 200 Thoroughbred foals on 10 farms demonstrated an average morbidity from distal respiratory tract infection of 82%.3 In foals, lower respiratory tract infections may be localized to the bronchi (bronchitis) or also involve the pulmonary parenchyma (pneumonia). In equine medicine, the term bronchopneumonia is often used to refer to lower respiratory tract infection regardless of whether the infection is localized to the bronchi or also involves the lung parenchyma. The spectrum of clinical signs shown by affected foals is broad and reflects the severity of the disease process. Early identification of affected foals and immediate initiation of appropriate therapy are essential to prevent mortality and functional impairment of the respiratory system.
Etiological agents Bacteria Although a wide variety of microorganisms have been associated with bronchopneumonia in foals, the vast majority of cases are bacterial in origin. Bacterial bronchopneumonia in foals is generally caused by
opportunistic pathogens that are normal inhabitants of the equine upper respiratory tract or gastrointestinal tract, or saprophytic environmental microorganisms. Streptococcus equi subspecies zooepidemicus (S. zooepidemicus) is by far the most common bacterial pathogen isolated from foals with bronchopneumonia. In a prospective study, S. zooepidemicus was isolated from 88% of foals with lower respiratory tract disease and was positively correlated with the percentage of neutrophils in pulmonary airway secretions.4 Streptococcus equi subspecies equi (strangles) infections are common in foals and young horses, but this organism is rarely isolated from the lungs of foals with bronchopneumonia. Rhodococcus equi pneumonia may occur sporadically or be enzootic on some farms. R. equi is considered elsewhere (see Chapters 75 and 76). Non-enteric Gram-negative bacteria such as Pasteurella spp., Actinobacillus spp., and Bordetella bronchiseptica are also frequently isolated, either alone or in combination with S. zooepidemicus. Enteric Gram-negative bacteria such as Klebsiella spp., Escherichia coli, and Salmonella spp. may also be isolated. Other aerobic bacteria such as Pseudomonas aeruginosa and Staphylococcus spp. are occasionally isolated. Pseudomonas spp. are rarely a primary cause of pneumonia in horses and their presence often reflects contamination of equipment used for taking airway samples (such as endoscopes). Streptococcus pneumoniae, a common pathogen of humans, has been positively correlated with lower airway inflammation in young Thoroughbred racehorses in the UK.5 The microorganism can also induce pneumonia in ponies following heavy intrabronchial challenge.6 However, S. pneumoniae is rarely isolated from pneumonic foals in the USA. Anaerobic bacteria are isolated much less frequently from foals with bronchopneumonia than from adult horses.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Other Bacterial Respiratory Diseases of the Older Foal Viruses Primary viral infections of the respiratory tract are considered elsewhere (see Chapter 79). Viral agents may predispose to secondary bacterial pneumonia by inducing ulceration of the pulmonary epithelium, reducing mucociliary clearance, and impairing alveolar macrophage function. However, with the possible exception of equine herpesvirus 2 (EHV-2), it is rarely possible to isolate a viral agent or demonstrate seroconversion by the time foals present with signs of bronchopneumonia.4, 7 In one study, the rate of isolation of EHV-2 in tracheobronchial aspirates (TBAs) of foals with bacterial bronchopneumonia (20/30, 66%) was significantly greater than from clinically normal foals (1/20, 5%).7 In contrast, in another study, EHV-2 or other viral agents were not isolated from TBAs obtained from 101 foals with bacterial lower respiratory tract infections.4 In the same study, seroconversion to viral agents was demonstrated in only two foals (EHV-1).4 EHV-2 remains latent in B-lymphocytes and can be isolated from the blood of approximately 80–90% of clinically healthy foals.7, 8 Experimental infection only induces mild upper respiratory signs.9 The role of EHV-2 in development of bacterial infections of the lower respiratory tract infections in foals remains to be determined. Mycoplasma Several Mycoplasma species have been isolated from the respiratory tract of both diseased and healthy horses with Mycoplasma felis and Mycoplasma equirhinis being the most common isolates. M. felis has been isolated from adult horses with pleuropneumonia, and experimental infection with this organism induced pleuritis.10, 11 M. felis has also been associated with outbreaks of lower respiratory tract disease in horses.12 In one study, there was an association between the presence of Mycoplasma spp. in the nasopharynx and respiratory disease in foals.13 In another study, Mycoplasma spp. were cultured from TBAs obtained from only four of 101 foals with lower respiratory tract infection.4 Fungi Molds and fungi isolated from tracheobronchial aspirates of foals are usually environmental contaminants and rarely contribute to the disease process. Infections with pathogenic fungi such as Cryptococcus neoformans, Histoplasma capsulatum, and Coccidioides immitis are rare in foals.14–16 Opportunistic fungi such as Aspergillus spp., Phycomycetes, Mucor, Rizopus, and Candida spp. are rare causes of severe pulmonary disease in immunocompromised patients, in foals
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with severe systemic diseases (most commonly enterocolitis), or in foals treated with prolonged broadspectrum antibacterial agents.17–19 Pneumocystis jirovecii (formerly P. carinii) is a unicellular eukaryote that has been classified as a fungus based on phylogenic analysis of rRNA.20 P. jirovecii lung infection develops in approximately 33% of Arabian foals with combined immunodeficiency disease (CID).21 It has also been reported in a variety of other breeds although, in most cases, no attempt was made to assess immune function.22–24 In one case, P. jirovecii pneumonia was associated with CD4+ and CD8+ lymphopenia.25 In another report, corticosteroid therapy may have induced immunosuppression and contributed to the development of P. jirovecii pneumonia.26 The onset of clinical signs in affected foals range from a 3-week history of weakness, weight loss, and nasal discharge to acute respiratory distress. P. jirovecii may be isolated in conjunction with R. equi, S. zooepidemicus, or other bacterial pathogens.27 Foals with P. jirovecii infections should be treated with trimethoprim–sulfonamide combinations (30 mg/kg orally q. 12 hours). Parasitic Parasitic pneumonia in foals is caused by either migrating larvae of Parascaris equorum or adults and larvae of Dictyocaulus arnfieldi. Eggs shed by P. equoruminfected horses mature to infective stages in 1–2 weeks. After infection, larvae appear in the liver in 2 days and in the lungs by 7–14 days.28 The larvae then travel up the bronchial tree and are swallowed. The prepatent period is 10–12 weeks. Larval migration of P. equorum results in eosinophilic interstitial pneumonia. Clinical signs of respiratory disease are first noted 10–13 days following experimental infection and typically last 5–10 days. Lung lesions may persist for more than 80 days post-infection.28 D. arnfieldi may cause respiratory disease in foals kept on pasture with donkeys or mules. Donkeys and mules may be asymptomatic carriers and contaminate the environment. After ingestion, infective thirdstage larvae penetrate the intestinal wall and travel to the lungs via the mesenteric lymphatics. Larvae reach the lungs approximately 1 week after infection where they mature in peripheral bronchi.29 The parasite may be seen during bronchoscopy. In adult horses, infections are usually non-patent, whereas in foals patent infection may develop. In a patent infection, eggs are laid in the bronchi and hatch quickly so that first-stage larvae ascend by the mucociliary apparatus, are swallowed, and passed in the feces. The prepatent period is 2–3 months. Fenbendazole (10 mg/kg/day for 5 consecutive days) is effective against both adult and migrating
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larvae of P. equorum and D. arnfieldi. Although widely reported as effective in older literature, P. equorum resistance to ivermectin (200 µg/kg) is now widely documented.30, 31
Epidemiology The highest frequency of distal respiratory tract infections occurs in foals between 1 and 6 months old with a peak incidence around 4 months of age.3 The management and environmental factors associated with development of bronchopneumonia in foals have never been critically investigated. In a prospective study, shipment of foal with dam to stud, foal dystocia, immunoglobulin (Ig)G status as neonate, foaling location (resident farm vs other farm) and housing type (outdoors with access to shed vs daily turnout from stable) were not significantly associated with development of lower respiratory tract infection.3 Infection was also as likely to be detected before as after weaning.3
study, abnormal lung sounds were detected in 30 of 86 (35%) foals with pneumonia, whereas every affected foal had abnormal lung sounds when using a rebreathing bag.3 The bronchovesicular lung sounds of foals are much louder than those of adult horses. The lung sounds of foals with distal respiratory tract infection vary considerably and may be confused by sounds referred from the upper airway. Foals with a large amount of secretions in the trachea often have an audible and palpable tracheal rattle. Most foals with bronchopneumonia cough when the rebreathing bag is applied whereas normal foals do not. Occasional inspiratory and expiratory crackles and/or wheezes may be heard over affected areas, which are more commonly located cranioventrally. Because consolidated lung parenchyma is a good acoustic medium, mild consolidation sometimes results in only increased bronchial sounds. In contrast, the lung sounds may be diminished in areas of severe consolidation, extensive abscess formation, or pleural effusion. Pleural effusion is a rare finding in foals with bronchopneumonia
Clinical signs and physical examination
Pathogenesis
The spectrum of clinical signs ranges from an otherwise normal appearing foal with mild bilateral nasal discharge or intermittent cough to one with severe cough, profuse purulent nasal discharge, fever, anorexia, depression, and respiratory distress. Mucoid to mucopurulent bilateral nasal discharge, sometimes evidenced only by crusting around the nostrils, is the most common clinical finding in foals with bronchopneumonia.3 However, the absence of nasal discharge does not rule out lower respiratory tract disease as exudate from the lungs can be swallowed and not appear as nasal discharge. Coughing is detected in approximately 50% of foals with lower respiratory tract infections. Mild tachypnea and altered respiratory character are also common features. Resting respiratory rates greater than 40 breaths per minute at rest in foals older than 3 weeks are usually considered abnormal and deserve further evaluation. Increased intercostal effort, often characterized by asynchronous rib excursion, is a subtle but common early sign.2 Most affected foals are well grown and in good body condition, but weight loss may become apparent with chronicity in severely affected cases. Although septicemia is not as common in older foals as it is in neonates, the examiner should be alert for other disease processes such as septic arthritis and diarrhea. Careful auscultation after application of a rebreathing bag (if not precluded by respiratory distress) is extremely valuable in defining the presence and sometimes extent of lung involvement. In one
As opposed to neonates that most commonly acquire pneumonia secondary to septicemia and hematogenous spread of bacteria, older foals most commonly acquire pneumonia by aspiration of organisms that transiently colonize the nasopharynx (most bacterial agents) or inhalation of aerosolized (viral agents) or environmental pathogens (R. equi). Aspiration pneumonia secondary to dysphagia or improper bottle-feeding techniques in orphan foals is also encountered occasionally. Colonization of the lungs by opportunistic bacteria occurs when pulmonary defense mechanisms are overwhelmed by massive challenge or when defenses are compromised by factors such as viral infection, transport, stress, dust, or noxious gases. Other factors that have been shown to significantly increase bacterial contamination of the lower respiratory tract include confinement with the head elevated, transportation, and high-intensity exercise.32–34 In one study, confinement of horses with the head elevated resulted in a significant increase in bacterial numbers as well as neutrophilic inflammation in the lower respiratory tract as early as 6 hours after initiating confinement.33 Actinobacillus spp., Pasteurella spp. and β-hemolytic streptococci were the predominant bacterial isolates. These findings indicate that periods of lowered head posture are absolutely essential for normal mucociliary clearance in horses.35 The inflammatory response induced by bacterial invasion will result in infiltration with neutrophils and other inflammatory cells into the airways and
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Other Bacterial Respiratory Diseases of the Older Foal pulmonary parenchyma. Inflammatory cells and their mediators cause damage to the airway epithelium and capillary endothelium, leading to flooding of the terminal airways with inflammatory cells, serum cellular debris, and fibrin. This process is generally more severe in the cranioventral portions of the lung. These lesions interfere with gas exchange and, if severe enough, the resulting ventilation–perfusion mismatch leads to hypoxemia and clinical signs of respiratory disease.2
Diagnosis Diagnosis of lower respiratory tract infection is generally based on clinical signs and careful auscultation of the lungs with a rebreathing bag. The need for additional diagnostic procedures is determined by: the herd history, number and the value of the foals, severity and duration of the clinical signs, treatment used, and response to therapy. On large farms, the disease process is often well advanced in the first animal(s) diagnosed, but additional foals are often affected but not yet showing clinical signs. Diagnostic evaluation should therefore be directed at the entire herd because it is unlikely that a single foal will be affected. In foals suspected of having pneumonia, the goal of diagnostic evaluation is to rule out diseases of the upper respiratory tract, and to determine the etiology and severity of lung involvement. A complete blood count (CBC) including plasma protein and fibrinogen concentrations may reveal neutrophilic leukocytosis and/or increased fibrinogen concentrations in some affected foals. There does not seem to be a good correlation between the severity of clinical signs and the presence or magnitude of hematological changes. In a prospective study, CBC and fibrinogen concentrations were not significantly different between foals with confirmed lower respiratory tract infection and healthy controls.3 When elevated, sequential measurement of plasma fibrinogen concentrations provides a useful means of monitoring response to treatment and is a useful guide in the decision to discontinue treatment. Persistent lymphopenia (<1000/µL) indicates the need for further evaluation of immune function, especially in Arabian foals at risk for combined immunodeficiency. Parasitic pneumonia should be considered in foals with clinical signs of lower respiratory tract disease and peripheral eosinophilia. A TBA for cytological examination and bacterial culture is the most definitive diagnostic procedure available. Whenever possible, antimicrobial therapy should be discontinued at least 24 hours before performing a tracheal wash. TBA can easily be obtained by percutaneous transtracheal aspiration or, alternatively, with a sterile polyethylene tube passed
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through the biopsy channel of a flexible fiberoptic endoscope. Endoscopy has the advantage of allowing selective aspiration of exudate when it is present (which may, therefore, enhance recovery of bacteria). However, bacterial contamination of nasal or pharyngeal origin commonly occurs with endoscopy. The use of a sterile cellulose sheath over the endoscope,36 or a guarded endoscopy swab or aspiration catheter (Mila International Inc.) passed through the biopsy channel of the endoscope37 significantly reduces, but may not completely eliminate, upper airway contamination. Under field conditions, it is not always practical or even indicated to do a TBA on every pneumonic foal. A reasonable approach is to perform TBA on the first few cases during an outbreak or in cases that do not respond to appropriate antimicrobial therapy. The trachea is not a perfectly sterile site and potentially pathogenic bacteria can be isolated from the trachea of clinically normal foals.38 Therefore, culture results should always be interpreted in the context of clinical signs and cytological examination. Pulmonary alveolar macrophages and columnar ciliated epithelial cells predominate in healthy individuals. The percentage of neutrophils is quite variable in a population of apparently healthy horses but less than 40% well-preserved neutrophils is usually considered normal.38, 39 Occasional plant spores and fungal hyphae may be present, either free or in large mononuclear cells. Their presence does not necessarily indicate fungal infection and probably reflect the horse’s environment. If bacteria are seen in the absence of cytological evidence of sepsis, it is unlikely that they are the cause of the foal’s respiratory problem. Presence of squamous epithelial cells indicates contamination with the upper respiratory tract or pharynx and is more common in samples obtained via the endoscope. In cases of lower respiratory tract infection, the neutrophil is the primary cell type seen and mucus is abundant. The neutrophils are degenerate (karyolysis, hypersegmented or pyknotic nuclei) and intraor extracellular bacteria are often present. Presence of larvae or eosinophilic inflammation in foals is suggestive of parasitic pneumonia. Special staining techniques may be necessary to identify unusual pathogens such as P. jirovecii, which is best demonstrated by a silver stain. Bronchoalveolar lavage (BAL) fluid is more sensitive than a tracheobronchial aspirate for the detection of P. jirovecii. Thoracic radiography and/or ultrasonography are indicated in more severe cases in which consolidation or lung abscessation is suspected. In addition to being useful for determining the severity of pneumonia, these two techniques are also useful in assessing response to therapy. Thoracic ultrasonography
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Figure 78.2 Lateral thoracic radiograph of a 5-month-old foal with bacterial bronchopneumonia showing a pronounced diffuse bronchointerstitial pattern.
Figure 78.1 Sonogram of the left thorax obtained from a 2month-old Thoroughbred foal with bacterial bronchopneumonia. There is mild focal consolidation of the lung (open arrow). Notice the liver-like appearance of the consolidated the lung. Normal lung can be seen dorsally and ventrally (solid arrows).
can be performed using a range of transducers and machine types. The normal aerated lung parenchyma is not penetrated by the ultrasound beam, rendering only the pleural space and superficial lung surface available for study. Ultrasonography is a helpful diagnostic tool when lung involvement includes peripheral areas but may not be as useful as radiography to evaluate the full extent of lung lesions since lesions with overlying aerated lung will not be detected. However, in most foals with bronchopneumonia the periphery of the lung is affected, enabling the clinician to successfully image some of the lesions. Early ultrasonographic lesions are non-specific and may include only irregularities of the pleural surface. These lesions may progress to form focal areas of consolidation of various sizes. Consolidated lung varies in appearance from dimples of the pleural surface to large wedge-shaped areas of sonolucent lungs (Figure 78.1). In horses with mild septic inflammatory airway disease thoracic radiographs may be normal or reveal only mild bronchial or bronchointerstitial patterns (Figure 78.2). In more severely affected animals, radiographs demonstrate irregular opacities in the ventral thorax that may obscure the normal vasculature and cardiac silhouette. Air bronchograms are sometimes visible (Figure 78.3). Abscesses are often present as circular soft-tissue opacities of varying sizes. In some cases they may be cavitated with thick walls and a distinct horizontal line representing fluid–gas interface. Pleural effusion is rare in
foals with bronchopneumonia but, when present, it appears as a horizontal line demarcating a ventral soft-tissue opacity that obscures the heart and ventral lung fields. Although ultrasonography is more convenient than radiography, the latter should still be performed when possible to detect and monitor resolution of deep pulmonary abscesses that may be missed during ultrasonography. Upper airway endoscopy is useful to rule out upper airway infection when physical examination and auscultation of the lungs is not conclusive. Presence of mucopurulent secretions in the trachea and bronchi in foals is suggestive of a bacterial infection of the lower respiratory tract. Other procedures that may be useful in the diagnostic evaluation of foals with lower respiratory tract infection include arterial blood gas analysis to evaluate oxygenation and
Figure 78.3 Lateral thoracic radiograph of a foal with bacterial bronchopneumonia showing a pronounced alveolar pattern with air bronchograms (some indicated by arrows). The caudal silhouette of the heart is completely obscured.
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Other Bacterial Respiratory Diseases of the Older Foal ventilation in severely affected foals, blood cultures, virus isolation, polymerase chain reaction testing for respiratory viruses, serology (paired samples) for respiratory viruses, fecal flotation, and Baermann test on feces when parasitic pneumonia is suspected.
Therapy The goal of therapy is to kill or inhibit bacterial pathogens, improve respiratory function, minimize stress, and maximize patient comfort and environmental quality. Restricting exercise is important in severely affected cases to limit ventilatory demand. In milder or resolving cases, limited exercise may facilitate expectoration. Affected foals should be kept in a cool, clean, and well-ventilated environment. Antimicrobial therapy Administration of antimicrobial agents is the most important part of the therapeutic plan. The choice of the antimicrobial agent depends on the severity of the clinical signs, cost, ease of administration, experience within the herd, and, when available, results of bacteriological culture and susceptibility testing of a TBA. Since a high percentage of pneumonia in foals older than 1 month is caused by bacteria with a predictable susceptibility profile such as S. zooepidemicus and Pasteurella spp. (this is not true for neonatal foals), ceftiofur (2.2–4.4 mg/kg q. 12–24 hours intramuscularly (i.m.)) is often used for initial therapy on farms where R. equi is not enzootic. Ceftiofur, a third-generation cephalosporin, has a broad spectrum of activity that includes most of the etiologic agents of foal pneumonia, except R. equi. A single daily dose at 2.2 mg/kg i.m. is usually sufficient to maintain serum concen-
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trations above the minimum inhibitory concentration (MIC) of S. zooepidemicus. However, when enteric Gram-negative bacteria are suspected or confirmed, a higher dose of 4.4 mg/kg q. 12 hours is necessary because these microorganisms have a much higher MIC than S. zooepidemicus.40 Procaine penicillin G or ampicillin is also highly effective against S. zooepidemicus. If resistant Gram-negative organisms are present, aminoglycosides (gentamicin or amikacin) are often combined with penicillin. Trimethoprimsulfonamide (TMS) combinations are attractive because they can be administered orally and for long periods of time. Unfortunately, the usefulness of TMS combinations for the treatment of bacterial respiratory tract infections in foals is limited by its lack of in vivo activity against S. zooepidemicus. In contrast to penicillin, TMS is ineffective in eradicating S. zooepidemicus in a tissue chamber model of infection in horses.41, 42 This failure of in vivo response was observed despite in vitro susceptibility of the isolate and high concentrations of TMS in the tissue chamber fluid.41, 42 The therapeutic regimen should be reevaluated if the foal does not show clinical improvement within 3–5 days following initiation of therapy. Chronic cases generally respond more slowly than do acute cases. Table 78.1 provides recommendations for selection of an antimicrobial agent for the most common respiratory pathogens of foals. Table 78.2 provides dosage recommendations for commonly used antimicrobial agents. Treatment should be continued for a minimum of 10 days, and much longer in severe cases. Clinical signs, lung auscultation, measurement of fibrinogen concentrations, and imaging techniques (ultrasonography, radiography) can be used to assess efficacy
Table 78.1 Suggested choices of antimicrobial agents for common bacterial pathogens of horses Microorganisms Gram positive β-hemolytic streptococcia Staphylococcus spp. Rhodococcus equi Gram negative Pasteurella–Actinobacillus spp. Actinobacillus equuli Bordetella bronchiseptica Escherichia coli Klebsiella spp. Enterobacter spp. Pseudomonas spp. a
Antimicrobial of choice
Alternative antimicrobials
Penicillin Cefazolin Macrolideb and rifampin
Ceftiofur; ampicillin; macrolide; rifampin; chloramphenicol TMS; amikacin; rifampin; chloramphenicol Doxycycline–rifampin; chloramphenicol–rifampin
TMS Ceftiofur Gentamicin; amikacin Amikacin Amikacin Amikacin Amikacin
Ceftiofur; doxycycline; gentamicin Gentamicin; amikacin TMS; doxycycline; oxytetracycline Gentamicin; ceftiofur Gentamicin; ceftiofur; cefotaxime; cefepime Cefepime Cefepime; ticarcillin
Streptococcus equi subspecies equi, S. zooepidemicus, and S. dysgalactiea subspecies equisimilis. Erythromycin, clarithromycin, or azithromycin. TMS, trimethoprim–sulfonamide. b
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Table 78.2 Antimicrobial agents commonly used to treat bacterial bronchopneumonia in older foals Antimicrobiala
Dose
Frequency (h)
Route
25 000 IU/kg 25 000 IU/kg
6 12
i.v. i.m.
20 mg/kg 20 mg/kg
8 12–24
i.v. i.m.
10–20 mg/kg 2.2–4.4 mg/kg
6 12–24
i.v. i.v. or i.m.
25 mg/kg 7.5 mg/kg 10 mg/kg
6–8 12 24–48b
p.o. p.o. p.o.
10 mg/kg (> 6 weeks) 25 mg/kg (< 6 weeks) 6.6 mg/kg (> 6 weeks) 12 mg/kg (< 6 weeks)
24 24 24 24
i.v. or i.m. i.v. or i.m. i.v. or i.m. i.v. or i.m.
Tetracycline Doxycyclinec
10 mg/kg
12
p.o.c
Others Rifampin TMS
5 mg/kg 30 mg/kg (combined)
12 12
p.o. p.o.
β-lactams Benzyl penicillins Penicillin G (Na, K) Penicillin G (procaine) Aminobenzyl penicillins: Ampicillin sodium Ampicillin trihydrate Cephalosporins Cefazolin Ceftiofur
Macrolides Erythromycin Clarithromycin Azithromycin Aminoglycosides Amikacin Gentamicin
a Dosages and agents listed may be extralabel and based on pharmacokinetic studies performed on small numbers of horses. As a result, efficacy and safety studies are often not available. Many of the drugs listed have the potential to cause adverse effects. Veterinarians using this table are encouraged to consult current literature and product label for information regarding efficacy, safety, and potential adverse reactions. b Once a day for 5 days and q. 48 hours thereafter. c Administer orally only. Intravenous doxycycline has resulted in severe cardiovascular effects including collapse and death in some horses .
and determine duration of therapy. Significant reduction in the frequency and number of S. zooepidemicus isolated during therapy supported a causal role for this microorganism in development of lower respiratory tract infection. In contrast, the frequency of β-hemolytic streptococci, Staphylococcus epidermidis and Streptomyces spp. increased during treatment suggesting a commensal or competitive role for these microorganisms.43 Most cases make a complete recovery if diagnosed early and treated appropriately. In a prospective study involving 178 foals with lower respiratory tract infection, complete resolution of the clinical signs was observed during the treatment period in all but seven (4%) cases.43 In the same study, the relapse rate of lower respiratory infections was approximately 30%. Relapses were identified from 7 to 35 days after discontinuation of therapy. Relapse was not associated with development of resistant organisms. In a preliminary study looking at the influence of foal pneumonia on future racing performance, a negative influence of β-hemolytic Streptococcus pneumonia was identified.44 These findings will need to be confirmed in a larger popu-
lation of foals before the true effect of foal pneumonia on future racing performance can be determined. Aerosolized antimicrobial agents may be a useful adjunct to oral or systemic administration particularly in foals with chronic septic inflammatory airway disease and no or minimal involvement of the lung parenchyma. Aerosol administration of antimicrobial agents can result in high drug concentrations in the respiratory tract while minimizing systemic concentrations and their resulting toxicity.45 Antimicrobial delivery by inhalation is greatly influenced by the product formulation and type of nebulizer. In one study, the particle size distribution and particle density of gentamicin sulfate and ceftiofur sodium aerosols were affected by the antimicrobial concentration of the solution.46 Gentamicin solutions at a concentration of 50 mg/mL or ceftiofur solutions at a concentration of 25 mg/mL produced the optimal combinations of particle size and aerosol density when using a medical ultrasonic nebulizer.46 The major limitation to the use of aerosolized gentamicin in horses is its lack of activity against S. zooepidemicus, the most common bacterial
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Other Bacterial Respiratory Diseases of the Older Foal pathogen of the equine respiratory tract. In contrast, ceftiofur is active against virtually all S. zooepidemicus isolates. Aerosolized antimicrobial agents should be considered in cases that do not respond to systemic antimicrobial agents or cases that relapse following appropriate therapy. Other therapies Nursing care, provision of adequate nutrition and hydration, and maintaining foals in a cool and well-ventilated environment are also important. Oxygen therapy using humidified oxygen by pharyngeal insufflation in moderately hypoxemic foals (Pa o2 < 65 mmHg), or by percutaneous transtracheal oxygenation in severely hypoxemic animals (Pa o2 < 50 mmHg)47 is indicated in foals with persistent hypoxemia or cyanosis. Non-steroidal anti-inflammatory drugs (NSAIDs) such as flunixin meglumine (1.1 mg/kg intravenously (i.v.) or per os (p.o.) q. 12 hours) may minimize the damaging effects of the cyclooxygenase-derived inflammatory mediators and are of value in reducing fever and improving attitude and appetite in febrile depressed anorectic foals. The use of bronchodilators in treating foal pneumonia is usually not rewarding since bronchoconstriction represents a relatively small part of the overall lung pathology. Nevertheless, rare foals with severe bronchopneumonia and a positive bronchodilator response test with inhaled albuterol may benefit from inhaled bronchodilators. Immunomodulators Immunomodulators have been used for the prevention and therapy of infectious respiratory diseases in horses. These agents function predominantly via non-specific macrophage activation and release of cytokines such as interleukin (IL) 1, IL-6, interferon, and tumor necrosis factor (TNF)-α.48 The assumption is that cytokines released by activated macrophages will enhance immune responses to a variety of antigens, thereby increasing resistance to infectious or neoplastic diseases. Mild fever, anorexia, and lethargy are common side-effects of immunomodulators.48 Purified mycobacterial cell wall extracts R (Equimune I.V. Vetrepharm Inc), inactivated ProR pionibacterium acnes (EqStim Neogen Corporation), R , Pfizer and inactivated parapoxvirus ovis (Zylexis Animal health) are available in North America. Data in adult horses suggest that these agents may be useful in the prevention or therapy of some infectious respiratory diseases.49–51 In healthy yearling horses, P. acnes administration increases the number of CD4+ T-lymphocytes in the blood, enhances the lymphokine-activated killing activity of BAL and
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blood lymphocyte, and enhances non-opsonized phagocytic activity peripheral blood leukocytes.52 In another study, administration of inactivated parapoxvirus ovis on days 0, 2, and 9 post-weaning and transport significantly reduced clinical signs of respiratory disease in foals compared with placebotreated animals.53 The use of immunomodulators in the treatment of foal bacterial bronchopneumonia has, to the author’s knowledge, never been addressed in a controlled study. The use of immunomodulators, in association with appropriate antimicrobial therapy, may be considered in pneumonic foals that do not respond to, or relapse following, conventional therapy.
Control strategies Good herd management is essential for successful prevention of foal pneumonia. Efforts should be made to maintain good hygiene and environmental quality, avoid overcrowding, reduce dust, maintain fixed herd groups, and isolate newly arrived and ill horses. Detection and treatment of cases of passive transfer of immunity, strict parasite control (Chapter 30) and adequate vaccination programs (Chapter 31) are also important. Because the success and cost of therapy depend on early diagnosis, foals should be monitored carefully for clinical signs such as nasal discharge, cough, increased respiratory rate, abnormal breathing pattern, exercise intolerance, anorexia, and fever. Abnormal foals should be immediately subjected to appropriate diagnostic evaluation and treated appropriately.
References 1. Cohen ND. Causes of and farm management factors associated with disease and death in foals. J Am Vet Med Assoc 1994;204:1644–51. 2. Wilson WD. Foal pneumonia: an overview. Proc Am Assoc Equine Pract 1992;38:203–29. 3. Hoffman AM, Viel L, Juniper E, Prescott JF. Clinical and endoscopic study to estimate the incidence of distal respiratory tract infection in thoroughbred foals on Ontario breeding farms. Am J Vet Res 1993;54:1602–7. 4. Hoffman AM, Mazan MR, Ellenberg S. Association between bronchoalveolar lavage cytologic features and airway reactivity in horses with a history of exercise intolerance. Am J Vet Res 1998;59:176–81. 5. Wood JL, Newton JR, Chanter N, Mumford JA. Association between respiratory disease and bacterial and viral infections in British racehorses. J Clin Microbiol 2005;43:120–6. 6. Blunden AS, Hannant D, Livesay G, Mumford JA. Susceptibility of ponies to infection with Streptococcus pneumoniae (Capsular Type 3). Equine Vet J 1994;26:22–8. 7. Murray MJ, Eichorn ES, Dubovi EJ, Ley WB, Cavey DM. Equine herpesvirus type 2: Prevalence and seroepidemiology in foals. Equine Vet J 1996;28:432–6.
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8. Kemeny L, Pearson JE. Isolation of herpesvirus from equine leukocytes: Comparison with equine rhinopneumonitis virus. Can J Comp Med 1970;34:59–65. 9. Borchers K, Wolfinger U, Ludwig H, Thein P, Baxi S, Field HJ, Slater JD. Virological and molecular biological investigations into equine herpes virus type 2 (EHV-2) Experimental Infections. Virus Res 1998;55:101–6. 10. Hoffman AM, Baird JD, Kloeze HJ, Rosendal S, Bell M. Mycoplasma felis pleuritis in two show-jumper horses. Cornell Vet 1992;82:155–62. 11. Ogilvie TH, Rosendal S, Blackwell TE, Rostkowski CM, Julian RJ, Ruhnke L. Mycoplasma felis as a cause of pleuritis in horses. J Am Vet Med Assoc 1983;182:1374–6. 12. Wood JL, Chanter N, Newton JR, Burrell MH, Dugdale D, Windsor HM, Windsor GD, Rosendal S, Townsend HG. An outbreak of respiratory disease in horses associated with Mycoplasma felis infection. Vet Rec 1997;140:388–91. 13. Antal A, Szabo I, Vajda G, Antal VD, Polner A, Totth B, Szollar I, Stipkovits L. Immunoglobulin concentration in the blood serum of foals suffering from pneumonia associated with Mycoplasma infection. Arch Exp Veterinarmed 1989;43:747–50. 14. Blanchard PC, Filkins M. Cryptococcal pneumonia and abortion in an equine fetus. J Am Vet Med Assoc 1992;201:1591–2. 15. Petrites-Murphy MB, Robbins LA, Donahue JM, Smith B. Equine cryptococcal endometritis and placentitis with neonatal cryptococcal pneumonia. J Vet Diagn Invest 1996;8:383–6. 16. Rezabek GB, Donahue JM, Giles RC, Petrites-Murphy MB, Poonacha KB, Rooney JR, Smith BJ, Swerczek TW, Tramontin RR. Histoplasmosis in horses. J Comp Pathol 1993;109:47–55. 17. Reilly LK, Palmer JE. Systemic candidiasis in four foals. J Am Vet Med Assoc 1994;205:464–6. 18. Sweeney CR, Habecker PL. Pulmonary aspergillosis in horses: 29 cases (1974–1997). J Am Vet Med Assoc 1999;214:808–11. 19. Breshears MA, Holbrook TC, Haak CE, York PA. Pulmonary aspergillosis and ischemic distal limb necrosis associated with enteric salmonellosis in a foal. Vet Pathol 2007;44:215–17. 20. Edman JC, Kovacs JA, Masur H, Santi DV, Elwood HJ, Sogin ML. Ribosomal RNA sequence shows Pneumocystis carinii to be a member of the fungi. Nature 1988;334:519– 22. 21. Perryman LE, McGuire TC, Crawford TB. Maintenance of foals with combined immunodeficiency: Causes and control of secondary infections. Am J Vet Res 1978;39:1043–7. 22. Perron Lepage MF, Gerber V, Suter MM. A case of interstitial pneumonia associated With Pneumocystis carinii in a foal. Vet Pathol 1999;36:621–4. 23. Peters SE, Wakefield AE, Whitwell KE, Hopkin JM. Pneumocystis carinii pneumonia in thoroughbred foals: identification of a genetically distinct organism by DNA amplification. J Clin Microbiol 1994;32:213–16. 24. Tanaka S, Kaji Y, Taniyama H, Matsukawa K, Ochiai K, Itakura C. Pneumocystis carinii pneumonia in a thoroughbred foal. J Vet Med Sci 1994;56:135–7. 25. Flaminio MJ, Rush BR, Cox JH, Moore WE. CD4+ and CD8+ T-lymphocytopenia in a filly with Pneumocystis carinii pneumonia. Aust Vet J 1998;76:399–402. 26. Ewing PJ, Cowell RL, Tyler RD, MacAllister CG, Meinkoth JH. Pneumocystis carinii pneumonia in foals. J Am Vet Med Assoc 1994;204:929–33.
27. Ainsworth DM, Weldon AD, Beck KA, Rowland PH. Recognition of Pneumocystis carinii in foals with respiratory distress. Equine Vet J 1993;25:103–8. 28. Srihakim S, Swerczek TW. Pathologic changes and pathogenesis of Parascaris equorum infection in parasite-free pony foals. Am J Vet Res 1978;39:1155–60. 29. Burks BS. Parasitic pneumonitis in horses. Compend Contin Educ Pract Vet 1998;20:378–83. 30. Slocombe JO, deGannes RV, Lake MC. Macrocyclic lactoneresistant Parascaris equorum on stud farms in Canada and effectiveness of fenbendazole and pyrantel pamoate. Vet Parasitol 2007;145:371–6. 31. Lyons ET, Tolliver SC, Ionita M, Collins SS. Evaluation of parasiticidal activity of fenbendazole, ivermectin, oxibendazole and pyrantel pamoate in horse foals with emphasis on ascarids (Parascaris equorum) in field studies on five farms in Central Kentucky in 2007. Parasitol Res 2008;103: 287–91. 32. Raidal SL, Love DN, Bailey GD. Effect of a single bout of high intensity exercise on lower respiratory tract contamination in the horse. Aust Vet J 1997;75:293–5. 33. Raidal SL, Love DN, Bailey GD. Inflammation and increased numbers of bacteria in the lower respiratory tract of horses within 6 to 12 hours of confinement with the head elevated. Aust Vet J 1995;72:45–50. 34. Racklyeft DJ, Love DN. Influence of head posture on the respiratory tract of healthy horses. Aust Vet J 1990;67:402– 5. 35. Raidal SL, Love DN, Bailey GD. Effects of posture and accumulated airway secretions on tracheal mucociliary transport in the horse. Aust Vet J 1996;73:45–9. 36. Hoffman AM, Viel L, Muckle CA, Tesarowski DB. Evaluation of a guarded bronchoscopic method for microbial sampling of the lower airways in foals. Can J Vet Res 1991;55:325–31. 37. Darien BJ, Brown CM, Walker RD, Williams MA, Derksen FJ. A tracheoscopic technique for obtaining uncontaminated lower airway secretions for bacterial culture in the horse. Equine Vet J 1990;22:170–3. 38. Crane SA, Ziemer EL, Sweeney CR. Cytologic and bacteriologic evaluation of tracheobronchial aspirates from clinically normal foals. Am J Vet Res 1989;50:2042–8. 39. Beech J. Tracheobronchial aspirates. In: Beech J (ed.) Equine Respiratory Disorders. Philadelphia: Lea & Febiger, 1991; pp. 41–53. 40. Meyer S, Gigu`ere S, Rodriguez R, Zielinski RJ, Grover GS, Brown SA. Pharmacokinetics of intravenous ceftiofur sodium and concentration in body fluids of foals. J Vet Pharmacol Ther 2009;32:309–16. 41. Ensink JM, Smit JA, van Duijkeren E. Clinical efficacy of trimethoprim/sulfadiazine and procaine penicillin G in a Streptococcus equi subsp. zooepidemicus infection model in ponies. J Vet Pharmacol Ther 2003;26:247– 52. 42. Ensink JM, Bosch G, van Duijkeren E. Clinical efficacy of prophylactic administration of trimethoprim/sulfadiazine in a Streptococcus equi subsp. zooepidemicus infection model in ponies. J Vet Pharmacol Ther 2005;28:45–9. 43. Hoffman AM, Viel L, Prescott JF. Microbiologic changes during antimicrobial treatment and rate of relapse of distal respiratory tract infections in foals. Am J Vet Res 1993;54:1608–14. 44. Bernard B, Dugan J, Pierce S, Gardiner I. The influence of foal pneumonia on future racing performance. Proc Am Assoc Equine Pract 1991;37:17–18.
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Other Bacterial Respiratory Diseases of the Older Foal 45. McKenzie III HC, Murray MJ. Concentrations of gentamicin in serum and bronchial lavage fluid after intravenous and aerosol administration of gentamicin to horses. Am J Vet Res 2000;61:1185–90. 46. McKenzie HC. Characterization of antimicrobial aerosols for administration to horses. Vet Ther 2003;4:110–19. 47. Hoffman AM, Viel L. A percutaneous transtracheal catheter system for improved oxygenation in foals with respiratory distress. Equine Vet J 1992;24:239–41. 48. Rush BR, Flaminio MJ. Immunomodulation in horses. Vet Clin North Am Equine Pract 2000;16:183–97. 49. Cormack S, Alkemade S, Rogan D. Clinical study evaluating a purified mycobacterial cell wall extract for the treatment of equine respiratory disease. Equine Pract 1991;13:18.
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50. Nestved A. Evaluation of an immunostimulant in preventing shipping related respiratory disease. J Equine Vet Sci 1996;16:78. 51. Vail CD, Nestved A, Martens JB. Adjunct treatment of equine respiratory disease complex (ERDC) with the Propionibacterium acnes, immunostimulant, EqStim. J Equine Vet Sci 1990;10:399. 52. Flaminio MJ, Rush BR, Shuman W. Immunologic function in horses after non-specific immunostimulant administration. Vet Immunol Immunopathol 1998;63:303–15. 53. Ziebell KL, Steinmann H, Kretzdorn D, Schlapp T, Failing K, Schmeer N. The use of Baypamun N in crowding associated infectious respiratory disease: efficacy of Baypamun N (freeze dried product) in 4–10 month old horses. Zentralbl Veterinarmed [B] 1997;44:529–36.
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Chapter 79
Viral Respiratory Diseases of the Older Foal Lutz S. Goehring and D. Paul Lunn
General introduction Viral respiratory tract infection in foals is a common cause of clinical or subclinical disease during the foal’s first 12 months of life.1 Multiple intrinsic and extrinsic factors determine the outcome after exposure to an infectious agent. One critical factor is passive, colostrum-derived maternal immunity. Colostral antibodies can provide protection against a variety of infectious agents in foals for their first 2–5 months of life. Factors that can limit the effect of passive transfer include failure to produce goodquality colostrum by the dam and failure of ingestion and absorption by the foal. Maximizing colostral protective immunity against specific pathogens requires that the dam has been exposed to the relevant pathogens prior to the initiation of colostrum production, either through natural infection or vaccination.2 Some of the pathogens to be discussed in the following section exhibit repetitive, subclinical cycles of replication in horses, and the mare, typically asymptomatic, may serve as a natural reservoir for viral infection of the foal. These silent cycles of viral replication may produce antigen-specific antibodies, which are transferred into colostrum; however, the antibody titers can be greatly enhanced by strategic vaccinations towards the end of a pregnancy (see also Chapter 31). Key extrinsic factors determining the occurrence of disease in foals include the dose of infectious agent to which the foal is exposed, stress, transportation, and changes in environment that may facilitate the viral colonization of the respiratory epithelium of the foal. Even subclinical infections can still allow production of vast numbers of pathogens in the respiratory tract of the foal, which are expelled into the environment and can infect other foals. Typical farm man-
agement favors turnout of mares and foals in groups, and crowding can significantly enhance the likelihood of spread of respiratory pathogens from one foal to another. Transportation of the mare and foal for breeding increases the risk for infection. Weaning followed by regrouping foals with subsequent social ranking changes are further stress factors. There are only a handful of viruses known to affect the foal’s respiratory tract: equine influenza virus (EI), equine arteritis virus (EAV), alphaherpesviruses (EHV-1 and -4) and gammaherpesviruses (EHV-2 and -5), adenovirus, and rhinitis viruses (A and B).3 All respiratory viruses are capable of causing some degree of damage to the respiratory epithelium. They can impair the respiratory tract’s mucociliary clearance mechanisms, thereby allowing common environmental bacteria to colonize affected areas. As a consequence of a secondary bacterial infection the clinical signs of coughing and mucopurulent nasal discharge will worsen.
Diagnostics utilized in viral respiratory disease Clinical signs may be very similar regardless of what virus caused the infection. Therefore laboratory diagnostics are necessary to determine the causative agent. Generally utilized laboratory diagnostics for equine respiratory tract viruses are detection of antigen or specific antibody.37 Antigen detection may use conventional virus isolation in tissue culture; direct antigen immunoassay membrane tests, such R Flu A (BD-Diagnostics, Franklin as the Directigen Lakes, NJ) for influenza virus; other immunoassays; electron microscopy; or polymerase chain reaction (PCR). Antigen-specific antibodies in serum
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Viral Respiratory Diseases of the Older Foal can be measured or quantified by enzyme-linked immunosorbent assay (ELISA), immunofluorescence, immunoperoxidase, or, rarely, by radioimmunoassays.3 A few multiplex real-time PCR assays are available that screen for a variety of specific antigens and often offer analysis for a panel of respiratory agents. More typically, the clinician needs to choose one or more tests based on the presenting clinical signs. Currently, routine PCR diagnostics for equine respiratory viruses are only available for EHV-1, EHV-4, equine influenza virus and EAV. Virus culture requires an experienced and dedicated laboratory, and a sufficiently high viral load in the submitted sample; however, it may provide a diagnosis in otherwise PCR-negative specimens. Electron microscopy (EM) may be helpful if all tests return with negative results, and it may detect an agent in affected lung tissue with a potential diagnosis made on viral characteristics such as cell type infected, viral size, and presence or absence of an envelope. Antigen-specific antibodies in a foal’s serum can be due to specific subclinical or clinical disease, passive antibody transfer after colostrum ingestion (which in foals may last until 5 or 6 months of age), or it may be due to vaccination. Passively acquired antibodies in foal plasma may also originate from plasma transfusions. Serology for a specific disease can be a false negative in the acute phase of infection, and should, therefore, always be assessed by seroconversion (e.g., observing a four fold titer increase) over a 3–4-week period. Routine laboratory diagnostics are typically only offered for influenza virus, EHV1 and -4, and EAV. Nasal swabs, immersed in virus transportation medium, tracheal washes, or guttural pouch lavage fluids should be submitted as appropriate. These samples, with the exception of nasal swabs are typically also submitted for bacterial culture. Chronic-active signs of respiratory disease are most likely due to a secondary bacterial infection of the respiratory tract, and can be most accurately diagnosed by means of culture and cytology of a tracheal lavage fluid, while viral antigen detection at that stage of the disease may be negative. At this stage seroconversion will provide the best evidence of viral infection.4 Most viral respiratory infections in foals and yearlings are cured with rest, very limited exercise, and antibiotics if secondary bacterial infection develops. Although not proven in experimental settings, inadequate care and time for recuperation from viral infections, and in particular from equine influenza, may predispose horses to chronic, immune-mediated conditions such as inflammatory airway disease (IAD) and recurrent airway obstruction (RAO).3 Viral causes of respiratory disease in foals include:
r r r r r
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equine influenza (EIV) equine arteritis virus (EAV) equine herpesviruses: r alphaherpesviruses (EHV-1 and EHV-4) r gammaherpesviruses (EHV-2 and EHV-5) equine rhinitis virus (type A and B) adenoviruses.
For each disease-causing agent the epidemiology, pathogenesis, diagnosis, treatment, and quarantine recommendations will be discussed under the corresponding subheadings. ‘Immunoprophylaxis and vaccination’ will be discussed only briefly, and these authors refer the reader to Wilson’s excellent chapter 31 on Vaccination of Mares, Foals and Weanlings. Additional and continuously updated information on vaccination protocols and guidelines can be found on the website of the American Association of Equine Practitioners (AAEP) at www.aaep.org/vaccination guidelines.htm under ‘Guidelines For The Vaccination of Horses’.
Influenza virus The severity of clinical disease caused by equine influenza virus (EIV) will depend on three factors: strain characteristics, quantity of virus in the inoculum, and host immunity. Maintenance of the circulation of EIV in the equine population depends on the virus’s ability to change its antigenicity through ‘antigenic drift’ and ‘antigenic shift’. Antigen-specific immunity from previous infection or vaccination may not recognize new viral antigens, and the immune system will be less efficacious in providing protection against new viral strains. The immunologically naive animal will have the most profound clinical signs of disease. Historically, horses have been affected by A/equine-1 viruses with an H7N7 strain type, and by A/equine-2 viruses with an H3N8 strain type. Currently, H7N7 viruses appear to have disappeared from equine populations, and only the H3N8 virus type needs to be included in vaccines. Within the H3N8 strain, two principal lineages are recognized, Eurasian and American. The latter appears to be more pathogenic, and is increasingly the most common lineage isolated worldwide in outbreaks. Vaccination recommendations typically suggest inclusion of EIV antigens from both H3N8 lineages.
Epidemiology Equine influenza virus can cause significant damage to a horse’s respiratory tract, particularly in the immunologically naive foal or yearling. Spread of EIV among a group of horses is very rapid because of
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the speed of viral replication, massive nasal shedding of virus, and aerosolization due to coughing. Inhalation of aerosolized virus is the most common way of transmission, and infectivity of virus may continue for 72 hours in a favorable environment.
Pathogenesis There is rapid replication of EIV in the upper respiratory tract, including the trachea and bronchi. Clinical signs include a frequent and hacking cough, a high fever, which typically peaks 72–96 hours postinfection, mucopurulent nasal discharge, and lymphadenopathy. Because of the intense damage to the airway’s epithelium there is decreased mucociliary clearance with subsequent trapping of cellular debris, inhaled bacteria, and mucus. Viable bacteria will colonize these otherwise sterile parts of the respiratory tract leading to secondary bacterial infection. While EIV does not typically cause a viremia, additional clinical signs are seen including limb edema and occasionally myositis. Myocarditis has been associated with EIV historically, but this has not been well documented. In isolated and severe cases, a severe interstitial viral pneumonia can develop, and this has been described in neonates and weanlings.6
search and potentially for vaccine development and updating.5
Treatment and quarantine recommendations Although there is some experience with virustatics such as amantadine, rimantadine, and the R neuraminidase inhibitor oseltamivir (Tamiflu ) in 7–9 an EIV infection is typically treated with horses, rest in a well-ventilated environment. Depending on secondary complications and clinical presentation, symptomatic therapy may include bronchodilators, mucolytics, antibiotics, and judicious use of non-steroidal anti-inflammatory drugs. Rest for at least 4 weeks, and ideally for 2 months, remains the key to a full recovery as the mucociliary escalator of the respiratory tract is impaired throughout this period. Quarantine can be lifted for EIV-infected horse premises in the absence of new cases. Recording and documentation of daily rectal temperatures from all horses on a quarantined horse operation is very helpful in this decision-making process; for more information see the AAEP website (www.aaep.org/pdfs/control guidelines). Vaccination
Diagnosis Equine influenza virus is the virus most likely to cause extensive coughing in the infected animal. Coughing as a clinical sign is less common with the other viral agents, as long as a secondary bacterial infection does not complicate the clinical picture. Clinically, EIV also appears to spread more quickly among susceptible horses of any age group, and fevers may be higher when compared to the other viral respiratory causes. However, for an accurate diagnosis laboratory testing is essential. During the acute febrile stage of EIV infection, for 7–10 days after onset of infection, nasal secretions may be positive for influenza virus antigen using stall-side tests such as the DirectigenTM Flu-A assay. Detection sensitivity is higher with either real-time quantitative, or with conventional gel-based PCR. Seroconversion should be demonstrated in an acute and converted serum sample with 3 weeks between sampling. The hemagglutination inhibition test is the most commonly available test and is offered by many laboratories. Nasal swabs in virus transportation medium should be sent to diagnostic laboratories whenever possible to confirm diagnosis and determine the strain type using culture techniques. This will make the strain available for future re-
A detailed immunization plan for horses in this age group is presented in Wilson’s Chapter 31 on vaccination, and current recommendations can also be found on the AAEP website (www.aaep.org). Four important points should be considered when vaccinating this age group:
1. Only contemporary vaccines should be used, i.e. ones that incorporate recently isolated strains, or vaccines that have been documented as effective against currently circulating EIV strains. 2. Passive immunity interferes with EIV vaccination, and foals should not be vaccinated against EIV before maternal immunity has waned. This occurs at 6 months of age if mares were boostered during late pregnancy. 3. Weaning, regrouping, or long-distance transportation should be planned appropriately to minimize immunosuppressive effects on vaccination and immunity development. Vaccination should also be timed to protect horses during these periods of increased risk. 4. A third vaccination 3 months after V2 can significanlty boost vaccine-induced antibody titers.38
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Viral Respiratory Diseases of the Older Foal
Equine arteritis virus (EAV)10 Equine viral arteritis is a highly contagious disease caused by the equine arteritis virus (EAV).10–12 The epidemiology and pathogenesis of this disease is described in greater detail in the sections focusing on abortion and stallion management, while the immunoprophylaxis for EAV is described in detail in Wilson’s Chapter 31 on Vaccination of Mares, Foals and Weanlings. Epidemiology Equine arteritis virus (EAV), causing equine viral arteritis (EVA), is a highly contagious disease that can spread through both respiratory and venereal routes. Seroprevalence differs between breeds and countries. A chronically infected intact male plays a central role in maintaining the virus in a horse population. Seronegative equids, including horses, ponies, mules, and donkeys of any age, are susceptible to infection, which can cause abortion or early embryonic loss in the pregnant mare (see also Chapter 246), and (sub)clinical disease with edema, conjunctivitis, excessive lacrimation, respiratory signs such as nasal discharge and coughing, and fever. An EAV infection of seronegative post-pubertal intact males will induce a carrier state in 30–50% of cases, where virus is shed with semen (see Chapters 121 and 246 for more details). Venereal infection of a seronegative mare will lead to disseminated disease involving the respiratory tract. A respiratory tract infection in seronegative pregnant mares will lead to abortion and seroconversion. Neonatal foals infected with EAV may show severe respiratory signs and a high mortality. It is sometimes unclear whether they were infected in utero or post partum; however, it shows their vulnerability to this infection.10–12 Clinical signs other than pregnancy losses during a recent outbreak of EVA in the western USA were described as mild, or even inapparent.11 Others describe outbreaks of EVA with high morbidity but low mortality, where high fevers are common as well as signs of a vasculitis (conjunctivitis, limb edema, petechial hemorrhage). EVA can cause significant morbidity in older, antibody-negative foals; however, mortality rates are low when compared to those for the neonatal foal.13,14 Pathogenesis Equine arteritis virus infection in the 2-week- to 12month-old group is strictly by aerosol inhalation. The respiratory tract and its associated lymphoid tissues are the locations of primary replication, and virus can be isolated from the respiratory tract between 2 and 14 days post-infection. Viremia follows pri-
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mary infection, and virus can be isolated from serum, plasma, or buffy coat up to 19 days post-infection. EAV replicates in endothelial cells causing cell damage and the clinical signs of a disseminated vasculitis. Immediately following the viremic phase virus can be isolated from a variety of tissues and body fluids. EAV is generally not isolated beyond 28 days after infection except from the semen of carrier stallions.10 Diagnosis Infection results in a high fever, which may last 2–9 days, a generalized vasculitis causing petechial hemorrhage, conjunctivitis, increased nasal secretions, interstitial pneumonia, and limb edema. However, during a recent outbreak of EVA in the western USA clinical signs were mild or absent. Diagnostics in the acute phase focus on virus isolation via culture, or PCR from nasal secretions, buffy coat, or plasma/serum. A detectable antibody titer in a foal from a seronegative mare indicates recent infection, which needs to be confirmed by seroconversion (four-fold titer increase over a period of 3–4 weeks). Treatment and quarantine recommendations A solid immunity develops post-infection, and with the appearance of antibody the excretion of virus through the respiratory tract ceases approximately 14 days post-infection. Virus may then be excreted through the semen of carrier stallions. Therapy for EAV-infected horses with clinical signs is palliative, and may include non-steroidal anti-inflammatory drugs, diuretics, hydrotherapy, and support wraps to reduce edema in the more severely affected cases. Quarantine is discontinued when no more clinical cases or serological evidence of infection are observed for three consecutive weeks; see the AAEP website for more information (www.aaep.org/pdfs/control guidelines).10, 11 Vaccination The most important steps in EVA control are the identification of carrier stallions (see Chapters 121, 246 and 321), and the prevention of the development of the carrier status of future breeding stallions. This can be achieved by vaccination of all prepubertal colts with breeding potential followed by yearly boosters. Details about the timing of vaccination can be obtained from Chapter 31 on Vaccination of Mares, Foals and Weanlings. Maternal immunity usually persists until age 2–5 months and may interfere with vaccine efficacy. Documentation of EVA vaccination and identification of the vaccinated colt is extremely important.15
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Equine herpesviruses16 To date all known herpesviruses of the horse fall into two subfamilies: alphaherpesviruses (EHV-1, EHV-3, EHV-4) and gammaherpesviruses (EHV-2, EHV-5).17 All members, with the exception of EHV-3, are associated with respiratory disease in the horse. In addition to respiratory disease, EHV-1, and to a much lesser degree EHV-4, are important causes of abortion and myelopathy. EHV-2 and EHV-5 are also associated with respiratory disease in horses of various ages; however, their role in disease has not been clearly established. EHV-1 and EHV-2 may also play a significant role in keratoconjunctivitis in horses. A survival strategy of herpesviruses, both alphaand gammaherpesviruses, is to establish lifelong latent infections that evade the host’s immune response. EHV-1 and -4 latency establishes in lymphoid tissue and in the trigeminal ganglia, while EHV-2 and -5 latency is restricted to lymphoid tissue.16 Equine herpesvirus research focuses primarily on EHV-1 and its associated syndromes of abortion/neonatal death and myelopathy. Key findings and developments over the past 10 years include the introduction of rapid and sensitive PCR diagnostics;18 the association of a point mutation in the polymerase gene of EHV-1 associated with neuropathology;19 the importance of antigen-specific cytotoxic lymphocytes in counteracting cell-associated viremia in abortion20 and myelopathy;21 and the detection of risk factors associated with myelopathy.22 Epidemiology Equine alpha- and gammaherpesviruses have a worldwide distribution. EHV-1 and -4 can cause respiratory disease in horses. Both are able to cause abortion/neonatal death; however, EHV-4 is only sporadically isolated from aborted fetuses (see also Chapters 240 and 245). While EHV-1 is most commonly associated with myelopathy outbreaks there are only anecdotal cases of EHV-4-associated myelopathy. Investigation of natural outbreaks of EHV-1 has shown that horses under 3 years of age are at a lower risk of developing myelopathy compared to older age groups.22, 23 The role of EHV-2 and -5 in disease is less clear. Both have an association with respiratory disease; however, there is usually an association with a secondary bacterial infection whenever clinical signs are present. Bell et al. isolated EHV-2 from nasal swabs in a group of foals on several occasions, but without clinical respiratory disease.24 Clinical or subclinical herpesvirus infections occur early in the life of a horse. Usually by 1 year of age serum antibodies against EHV-1, -2, -4, and -5 are
present, and it is believed that silent cycles of replication in the dams will infect their foals, followed by additional horizontal foal-to-foal spread. An aborted fetus, placenta, and vaginal fluids post-abortion are other sources of environmental spread for EHV-1. It is common practice on breeding operations to protect against EHV abortion by vaccinating pregnant mares at the beginning of the fifth, seventh, and ninth months of pregnancy (see also Wilson’s Chapter 31 on Vaccination of Mares, Foals and Weanlings). This will also lead to colostral antibody transfer to the foal. However, an Australian study has shown that these antibodies cannot prevent EHV-1 infections in foals, which typically occur in the first weeks or months of life.25 Pathogenesis Replication in the epithelium of the upper respiratory tract is common to EHV-1, -2, -4, and -5. With EHV-1 and -4 this can lead to mild to moderate clinical signs of respiratory disease with fevers for 1–7 days, mild lymphadenopathy, and serous nasal discharge. Virus is then transported to regional lymph nodes,16 where further replication leads to development of viremia. Viral replication causes damage to the respiratory epithelial cell layer, which can lead to secondary bacterial infections. A rare presentation of an EHV-1 infection in yearling horses has been described by Del Piero et al., where an EHV-1 infection apparently resulted in interstitial pneumonia and death.26 The contribution of gammaherpesvirus to respiratory disease is not clear. Murray et al. showed that respiratory disease in foals was associated with EHV-2 virus isolation from tracheal aspirates, but not from age-matched, asymptomatic foals.27 Some authors have speculated that gammaherpesvirus infections may predispose to infection with other agents. One report suggests a role for EHV-2 infections as a prerequisite for Rhodococcus equi infections, and the study claimed that vaccination against EHV-2 decreased the number of Rhodococcus equi infections among foals on the involved premises.28, 29 EHV-5 has been associated with a syndrome of multinodular pulmonary fibrosis in horses aged 4 years or older, but not in the age group discussed here.30 Diagnosis A diagnosis cannot be made on the basis of clinical signs alone. The most typical respiratory tract signs of EHV-1 and EHV-4 infections in this age group are serous nasal discharge, pyrexia, and enlarged mandibular lymph nodes. Coughing is uncommon with herpesvirus infections unless secondary
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Viral Respiratory Diseases of the Older Foal bacterial infections occur. Based on the findings of Bell et al., primary gammaherpesvirus infections were typically subclinical.24 Many diagnostic laboratories nowadays provide PCR diagnosis for EHV-1 and EHV-4 from nasal swabs collected from affected horses. Few laboratories offer virus isolation. Nasal swabs submitted for this purpose have to be transported in the appropriate virus isolation medium. Serology for EHV-1 and -4 using a serum neutralization (SN) assay is routinely offered by many diagnostic laboratories. A four-fold titer increase between an acute sample and a convalescent sample collected 3 weeks later is key to a proper diagnosis. However, it is important to understand that an SN assay does not distinguish between antibodies produced against EHV-1 or EHV-4. Few laboratories offer a complement fixation (CF) assay, which more readily shows recent infection; however, similar to the SN assay, the CF assay does not distinguish between antibodies produced against EHV-1 or EHV-4. A glycoprotein G (gG)-specific ELISA is currently the only test to confirm an antibody titer specifically caused by EHV1 infection (or maternal EHV-1 antibodies, or EHV1 vaccination). This ELISA is currently commercially available in Europe and Australia.31 Routine diagnostics are infrequently available for EHV-2 and EHV-5 detection. Virus culture and PCR are most frequently used to demonstrate this viral infection, but tests for gammaherpesviruses are typically only available from research laboratories. Treatment and quarantine Clinical signs of respiratory disease due to EHV-1 or -4 infections are mild and may not require therapy other than rest. EHV-2 and -5 infections are typically subclinical. However, foals infected in utero with EHV-1 and not aborted often present icteric and usually with severe respiratory distress due to interstitial pneumonia within days of birth (for more information see Chapters 63, 66 and 245). These foals shed EHV-1 and are an important risk of spread of infection. Infected foals and weanlings may form an important link in transmission of virus from one horse to the other, because of the potential of increased virus replication in their respiratory tracts when compared to adult horses with previous exposure. This is one reason not to commingle pregnant mares with foals, weanlings, or yearlings. Strategies for quarantine in the face of an EHV-1 outbreak are described in detail in the AAEP Outbreak Guidelines (www.aaep.org/pdfs/control guidelines). The key elements are disinfection of contaminated areas (post-abortion), isolation of affected horses, sub-
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mission of diagnostic samples, and implementation of hygiene procedures to spread further infection. Lifting of quarantine should ideally be delayed until 28 days after the last new clinical case, although shorter quarantine periods are used in conjunction with monitoring for fever daily, and then testing all horses for EHV-1 shedding after 14 days free of fever. Even in these circumstances, horses transported after outbreaks should be quarantined again at their new destinations.
Vaccination Several killed-virus vaccines and one modified-live virus vaccine against EHV-1 are available in the USA. Some killed-virus vaccines against EHV-1 also contain EHV-4 antigen. Two Food and Drug Administration (FDA)-registered (killed-virus) vaccines have an anti-abortion claim. For vaccination guidelines see Chapter 31 and the vaccination guidelines on the AAEP website (www.aaep.org/vaccination guidelines.htm).
Equine rhinitis A and B viruses (ERAV and ERBV)32, 33 Equine rhinitis A and B viruses are picornaviruses. The seroprevalence of rhinitis virus antibodies ranges worldwide from 20% to 80%. Foals apparently receive colostral antibodies against ERV and the titer decreases during the first 5 months of a foal’s life. A four-fold titer increase is indicative of ERV infection, and is more commonly observed in horses older than 24 months of age.3 Both viruses spread through respiratory aerosol infection; however, ERAV is also excreted through urine. This route of excretion actually is more prolonged than excretion through the respiratory tract. Clinical signs of respiratory disease are considered mild: serous nasal discharge, a fever, pharyngeal follicular hyperplasia, and mild lymphadenopathy. ERV infections are associated with coughing, which is probably the result of secondary bacterial infections. Although the diagnosis can be made through antigen detection via PCR or seroconversion on paired serum samples, there are only a limited number of laboratories that routinely run samples for ERV detection. There is currently no vaccine available.
Adenovirus34–36 Equine adenovirus type 1 (EAdV1) is able to cause respiratory disease in horses younger than 1 year of age, although signs are mild and infections typically
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go undiagnosed. This virus replicates readily in respiratory epithelium causing upper respiratory disease, conjunctivitis, bronchopneumonia, and, in addition, mild enteritis as a result of gastrointestinal viral replication. Virus is transmitted in respiratory or ocular secretions. Fecal shedding is suspected in foals with gastrointestinal infection. The virus is found worldwide. Seroprevalence varies between 2% and 100%. Antibody titers to EAdV1 persist throughout life due to chronic or recurrent infection. A persistent and disseminated infection with EAdV1 in foals may indicate a compromised immune system and is a frequent finding in Arabian foals affected with the severe combined immunodeficiency syndrome (SCID), where adenoviral pneumonia and pancreatitis are a frequent cause of death.3, 8 Currently no vaccine against EAdV1 is available.34–36
References 1. Barr BS. Pneumonia in weanlings. Vet Clin North Am Equine Pract 2003;19:35–49. 2. Tizard IR. Immunity of the Fetus and the Newborn. In: Veterinary Immunology 8th ed.; Saunders Elsevier; chapter 18, pp 223–238. 3. Landolt GA, Lunn DP. Equine respiratory viruses. In: Smith BP (ed.) Large Animal Internal Medicine. St Louis: Mosby Elsevier, 2008; pp. 542–9. 4. Davis GE, Freeman DE, Hardy J. Respiratory infections. In: Sellon DC, Long, MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007, pp. 1–20. 5. Landolt GA, Townsend HGG, Lunn DP. Equine influenza infection. In: Sellon DC, Long, MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007, pp. 124–34. 6. Patterson-Kane JC, Carrick JB, Axon JE. The pathology of bronchointerstitial pneumonia in young foals associated with the first outbreak of equine influenza in Australia. Equine Vet J 2008;40:199–203. 7. Rees WA, Harkins JD, Lu M. Pharmacokinetics and therapeutic efficacy of rimantadine in horses experimentally infected with influenza virus A2. Am J Vet Res 1999;60: 888–94. 8. Rees WA, Harkins JD, Woods WE. Amantadine and equine influenza: pharmacology, pharmacokinetics and neurological effects in the horse. Equine Vet J 1997;29:104–10. 9. Yamazaki Y, Ishii T, Honda A. The influence of oseltamivir carboxylate and oseltamivir on hemagglutinin inhibition and microneutralization test. Antiviral Res 2008;80:354–9. 10. Balasuriya UBR, MacLachlan NJ. Equine viral arteritis. In: Sellon DC, Long, MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 153–64. 11. Holyoak GR, Balasuriya UB, Broaddus CC. Equine viral arteritis: current status and prevention. Theriogenology 2008;70:403–14. 12. Timoney PJ, McCollum WH. Equine viral arteritis. Vet Clin North Am Equine Pract 1993;9:295–309. 13. Szeredi L, Homy´ak A, D´enes B. Equine viral arteritis in a newborn foal: parallel detection of the virus by immunohistochemistry, polymerase chain reaction and virus isolation. J Vet Med B Infect Dis Vet Public Health 2003;50:270–4.
14. Vaala WE, Hamir AN, Dubovi EJ. Fatal, congenitally acquired infection with equine arteritis virus in a neonatal thoroughbred. Equine Vet J 1992;24:155–8. 15. Hullinger PJ, Wilson WD, Rossitto PV. Passive transfer, rate of decay, and protein specificity of antibodies against equine arteritis virus in horses from a Standardbred herd with high seroprevalence. J Am Vet Med Assoc 1998;213:839–42. 16. Slater J.D. Equine herpesviruses. In: Sellon DC, Long, MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007, pp. 134–53. 17. Roizman B, Pellett PE. The family Herpesviridae: A brief introduction. In: Knipe, DM, Howley PM (eds) Field’s Virology. Philadelphia: Lippincott Williams & Wilkins, 2001; pp. 2381–445. 18. Hussey SB, Clark R, Lunn KF. Detection and quantification of equine herpesvirus-1 viremia and nasal shedding by real-time polymerase chain reaction. J Vet Diagn Invest 2006; 18:335–42. 19. Nugent J, Birch-Machin I, Smith KC. Analysis of equid herpesvirus 1 strain variation reveals a point mutation of the DNA polymerase strongly associated with neuropathogenic versus nonneuropathogenic disease outbreaks. J Virol 2006;80:4047–60. 20. Kydd JH, Wattrang E, Hannant D. Pre-infection frequencies of equine herpesvirus-1 specific, cytotoxic T lymphocytes correlate with protection against abortion following experimental infection of pregnant mares. Vet Immunol Immunopathol 2003;96:207–17. 21. Allen GP. Risk factors for development of neurologic disease after experimental exposure to equine herpesvirus-1 in horses. Am J Vet Res 2008;69:1595–600. 22. Goehring LS, van Winden SC, van Maanen C. Equine herpesvirus type 1-associated myeloencephalopathy in The Netherlands: a four-year retrospective study (1999–2003). J Vet Intern Med 2006;20:601–7. 23. Henninger RW, Reed SM, Saville WJ. Outbreak of neurologic disease caused by equine herpesvirus-1 at a university equestrian center. J Vet Intern Med 2007;21:157–65. 24. Bell SA, Balasuriya UB, Gardner IA. Temporal detection of equine herpesvirus infections of a cohort of mares and their foals. Vet Microbiol 2006;116:249–57. 25. Foote CE, Love DN, Gilkerson JR. EHV-1 and EHV-4 infection in vaccinated mares and their foals. Vet Immunol Immunopathol 2006;111:41–6. 26. Del Piero F, Wilkins PA, Timoney PJ. Fatal nonneurological EHV-1 infection in a yearling filly. Vet Pathol 2000;37:672–6. 27. Murray MJ, Eichom ES, Dubovi EJ. Equine herpesvirus type 2: prevalence and seroepidemiology in foals. Equine Vet J 1996;28:432–6. 28. Nordengrahn A, Rusvai M, Merza M. Equine herpesvirus type 2 (EHV-2) as a predisposing factor for Rhodococcus equi pneumonia in foals: prevention of the bifactorial disease with EHV-2 immunostimulating complexes. Vet Microbiol 1996;51:55–68. 29. Varga J, Fodor L, Rusvai M. Prevention of Rhodococcus equi pneumonia of foals using two different inactivated vaccines. Vet Microbiol 1997;56:205–12. 30. Williams KJ, Maes R, Del Piero F. Equine multinodular pulmonary fibrosis: a newly recognized herpesvirusassociated fibrotic lung disease. Vet Pathol 2007;44: 849–62. 31. Gilkerson JR, Love DN, Drummer HE. Seroprevalence of equine herpesvirus 1 in thoroughbred foals before and after weaning. Aust Vet J 1998;76:677–82.
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Viral Respiratory Diseases of the Older Foal 32. Carman S, Rosendal S, Huber L. Infectious agents in acute respiratory disease in horses in Ontario. J Vet Diagn Invest 1997;9:17–23. 33. Klaey M, Sanchez-Higgins M, Leadon DP. Field case study of equine rhinovirus 1 infection: clinical signs and clinicopathology. Equine Vet J 1998;30:267–9. 34. McChesney AE, England JJ. Adenoviral infection in foals. J Am Vet Med Assoc 1975;166:83–5. 35. Thompson DB, Spradborw PB, Studdert M. Isolation of an adenovirus from an Arab foal with a combined immunodeficiency disease. Aust Vet J 1976;52:435–7.
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36. Thomson GR. Adenovirus infections of horses. In: Coetzer JAW, Tustin RC (eds) Infectious Diseases of Livestock. Cape Town: Oxford University Press, 2004; pp. 823–6. 37. Tizard I. Immunodiagnostic techniques. In: Veterinary Immunology. St Louis, MO: Elsevier, 2009; Chapter 38, pp. 509–27. 38. Holmes, MA, Townsend HG, Kohler AK. Immune responses to commercial equine vaccines against equine herpesvirus-1, equine influenza virus, eastern equine encephalomyelitis, and tetanus. Vet Immunol Immunopathol, 2006. 111(1-2): p. 67–80.
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Chapter 80
Neurological Conditions (EPM, EMND, Wobblers, Trauma, etc) Guy D. Lester
There are several neurological conditions that could affect the older foal, the most important of which is cervical vertebral compressive myelopathy. Trauma to the central nervous system is also very common. The principles of diagnosis and therapy in response to trauma are discussed in Chapter 54.
Cervical vertebral compressive myelopathy Cervical vertebral compressive myelopathy (CVCM) is one of the most common neurological diseases of young horses. There are a number of synonyms for CVCM used in the veterinary literature, including cervical stenotic myelopathy, cervical vertebral stenotic myelopathy, cervical compressive myelopathy, cervical spondylotic myelopathy, and cervical vertebral malformation. The term ‘wobbler’ or ‘wobbler’s syndrome’ is also used colloquially, but is not recommended as the term is broadly descriptive of a range of spinal cord diseases. The syndrome is part of a larger group of developmental orthopedic diseases (DOD) that are expressed most commonly in young, rapidly growing horses. Other manifestations of DOD include osteochondrosis dissecans, epiphysitis, and subchondral cystic lesions. The neurological signs are caused by bony malformation or exostoses that result in extradural compression of the spinal cord. The distribution, severity, and symmetry of limb signs is determined by the nature and site of the bony deformities. Typically the earliest signs of CVCM are not seen until the late suckling period. These are usually attributable to dynamic ventrodorsal compression of
the spinal cord between the C2 and C5 vertebrae. Neurological abnormalities may include weakness, ataxia, and dysmetria. These neurological deficits are typically symmetrical (left versus right) and more severe in the pelvic limbs. Extensor weakness is noted by muscular trembling, collapse of the limb when turning, and a decreased resistance to tail pull when moving forward. Flexor weakness is manifest clinically through frequent toe dragging and altered limb height when moving forward or turning. Proprioceptive deficits are detected through walking the horse in small circles and by tight pivoting. The small circles should be of sufficient size such that the horse is always moving forward but constantly changing direction. The abnormal horse will succumb to inertia and move the outside pelvic limb widely in a circular motion before placing (circumduction). They may also have a tendency to ‘fall’ towards the handler with the inside thoracic limb. When a horse is pivoting, the handler should try and maintain a central position within a tight circle. The normal horse will rotate its inside pelvic limb approximately 15–20 degrees before deliberately and precisely placing it in front of the outside limb. The abnormal horse will pivot excessively on the inside pelvic limb, with severely affected animals rotating more than 360 degrees on the inside limb. The outside front foot should be placed across and in front of the inside thoracic limb; the abnormal horse will often throw the leg in a stiff and exaggerated manner and may at times step on the inside foot. Pivoting can be difficult to assess in belligerent horses or in animals with neck pain.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Neurological Conditions (EPM, EMND, Wobblers, Trauma, etc)
Figure 80.1 A young Thoroughbred demonstrating pronounced hypometria of the right thoracic limb moving down a slope with the head elevated.
Spasticity or hypometria is often a prominent feature of thoracic limb movement in horses with cervical spinal cord disease, and is exacerbated by walking an affected animal on a downward slope with the head elevated. This gait is often described as akin to that of a ‘tin –soldier’, where the forelimbs are rapidly thrust forward with limited limb flexion and the animal appears to place the foot several centimeters above the ground (Figure 80.1). Animals with CVCM will also frequently knuckle excessively in their rear fetlocks when moving down a slope with head elevated. A similar distribution of signs in newborn or younger foals should also warrant consideration of occipitoatlantoaxial malformation or equine degenerative myelopathy (see Chapter 54). A second subtype of CVCM is more commonly seen in older horses (2 years +) and is due to dorsolateral compression of the spinal cord, typically between the C5 and T1 veterbrae.1 The compression is constant or static, and is due to osteoarthritis of the dorsal articular facets. Variability in facet size and shape can lead to asymmetry in neurological signs, and the difference in signs between thoracic and pelvic is often less than that associated with the dynamic form of compression. The diagnosis of CVCM is based on neurological assessment and imaging. Standing plain lateral radiographs are very useful in increasing the accuracy of a diagnosis. Radiographic findings consistent with CVCM include flexion-fixation of adjacent vertebrae resulting in the appearance of joint subluxation (as illustrated in Figure 80.2), overt narrowing or ‘wedging’ of the vertebral canal, elongation of the dorsal laminae such that the caudal aspect may project behind the cranial physis of the trailing vertebrae, and epiphyseal flaring. An adjusted minimal sagittal diameter (MSD) ratio has been used to predict spinal cord compression
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Figure 80.2 A plain standing radiograph, illustrating narrowing of the cranial aspect of the vertebral canal at C4 with the appearance of joint subluxation at C3–4.
in horses with CVCM.2 The ratio is calculated by dividing the minimum sagittal diameter of the vertebral canal by the maximal sagittal height of the cranial half of the body of the vertebrae (Figure 80.3). The values have been determined from C3–4 through C6–7 and a corrected MSD less than or equal to 0.51 predicts cord compression with more than 95% sensitivity and specificity. Values less than 0.50 were associated with likelihood ratios of having CVCM of 28.6, 26.1, and 41.5 for C4, C5 and C6, respectively. A ratio of less than 0.52 was associated with likelihood ratio of 39.0 at C7.2 It is important to note that these values have not been validated in horses less than 12 months of age and should be used with caution. Furthermore,
Figure 80.3 The adjusted MSD ratio is calculated by dividing the minimum sagittal diameter of the vertebral canal (C) by the maximal sagittal height of the cranial half of the body of the vertebra (D). The intervertebral ratios described by Hahn et al.4 use the smaller of the values A or B as the numerator; A is measured from the dorsocaudal aspect of the vertebral body to the cranial aspect of the dorsal lamina of the more caudal vertebra. B is measured from the caudal aspect of the dorsal lamina of the arch to the dorsocranial aspect of the body of the caudal vertebra. The denominator, as described in their paper, was the maximal sagittal height of the cranial half of the body of the more cranial vertebra.
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the technique does not always accurately identify the precise site of compression, prompting the need for myelography, should surgery be considered. A semi-quantitative scoring system was developed to predict compressive spinal cord lesions in Thoroughbred foals.3 The system used common radiographic features including, but not limited to, canal stenosis (as assessed using MSD corrected by vertebral body length), angular fixation of joints, enlargement of physeal plates, and degenerative changes in the dorsal articular facets. Foals with clinical evidence of CVCM had a mean score of 17.0 while those without neurological signs had a score of 5.7. An alternative method of evaluating plain standing cervical films has been described.4 Data were collected from 26 Warmblood and Thoroughbred horses with an age range of 6 months to 18 years. The authors compared sagittal ratios generated using intraand intervertebral measurements. The intravertebral measurements were obtained in a similar manner to those published previously, except that the minimum measurement could be taken at any point along each vertebra. The intervertebral distance used in the calculation was the smallest of the following: a line drawn from the caudal aspect of the dorsal lamina of the cranial vertebra to a point on the dorsocranial aspect of the body of the more caudal vertebra or a line from the dorsocaudal aspect of the floor of the vertebral canal to a point of the cranial dorsal lamina of the vertebral arch of the more caudal vertebra (Figure 80.3). The maximum height of the cranial vertebral physis was used as the denominator to establish each ratio. Greater accuracy of radiographic detection of CVCM was achieved using intervertebral sagittal diameter ratios rather than intravertebral sagittal diameter measurements. An inter- or intravertebral sagittal ratio of less than or equal to 0.485 accurately predicted cord compression in this study and, importantly, the technique accurately predicted the site of cord compression. Myelography is often described as the most accurate method to detect cord compression, and should be performed when considering surgical management of CVCM. The criteria used to evaluate contrast myelography are, however, not well described and care must be taken when interpreting any reduction in the height of contrast columns.5 A reduction in the dorsal dye column to less than 2 mm has been used to predict compression in flexed views, based on published normal minimal heights.6 A recent review of the ‘2 mm rule’ concluded that the method was indeed sensitive, but lacked specificity.5 The most commonly used criterion is a 50% reduction in the dorsal dye column when compared to the maximal column height at the vertebral body cranial to the reduction. Some have described a 50% reduction in
both the dorsal and ventral columns at diametrically opposed sites as diagnostic of compression. The 50% reduction criterion has also been evaluated for accuracy and the authors concluded that this definition resulted in a relatively low sensitivity (56%) and moderate specificity (83%), and did not perform as well in predicting compression as did plain films evaluated with a scoring system.7 Van Bierliet and colleagues5 concluded that at C6–7 a 20% or greater reduction in dural diameter predicted cord compression with high sensitivity and specificity. Furthermore, the criterion was accurate independent of neck position, i.e., flexion or extension. Unfortunately, the use of fixed criteria were problematic at more proximal sites where a 20% reduction in dural diameter was highly specific but lacked sensitivity; flexed views increased the sensitivity but led to a decline in specificity, i.e., an increase in the number of false-positives. A reasonable conclusion from all of these studies is that a combination of sound neurological assessment and plain standing films are likely to predict compression with a relatively high degree of accuracy, and that interpretation of myelographic studies can be problematic. A diagnosis of CVCM carries a guarded to poor outlook. The options for correction are limited, although some success has been reported for foals less than 12 months of age, placed on a strict regimen of calorie and exercise restriction.8 The technique involves stall confinement and feeding a formulated diet that provides essential nutrients while delivering a reduction in energy content to delay growth. The technique is only suitable for young animals less than 12 months of age, with early and relatively mild signs of cord compression. Consideration may be given to surgical intervention in the management of CVCM. The decision for surgery is based on wide range of criteria, including the number and location of sites of compression, the severity of neurological signs, a realistic expectation of the degree of improvement, the intended use of the horse, appreciation of the time involved in convalescence, the cost of the surgery and after-care, and the availability of surgical expertise to perform the procedure.
Equine protozoal myeloencephalitis Equine protozoal myeloencephalitis (EPM) is a disease of the nervous system of horses caused by Sarcocystis neurona. A similar disease has also been attributed to Neospora hughesi and Neospora caninum. The disease occurs in the Americas and is widespread throughout North America. The definitive host of S. neurona is the opossum, with the raccoon, armadillo, skunk, and domestic cat all identified as viable intermediate hosts. The presence of cats residing on horse properties was identified as one of several
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Neurological Conditions (EPM, EMND, Wobblers, Trauma, etc) risk factors in a multicenter epidemiological study of EPM.9 Although the mean age of affected horses is between 3 and 6 years, a diagnosis of EPM in an animal less than 2 years of age would be unusual. Furthermore, in regions where there is a high rate of seropositivity to these parasites in the adult horse population, the exposure to S. neurona or N. hughesi between birth and 2.5 years of age is low.10 Disease caused by S. neurona is typically described as multifocal and asymmetric, although these features are often inconsistent from case to case. In an epidemiological study of 183 horses with EPM, 48% had neurological evidence of multifocal disease and 66% had asymmetry of signs.9 Most cases have lesions that localize to the cervical spinal cord, causing signs that may mimic those of CVCM (see above). Asymmetry of limb signs can be a feature of static compression due to uneven changes in dorsal articular facets. Establishing an antemortem diagnosis of EPM can be very difficult. A complete neurological examination is a critical component in reaching a provisional diagnosis of EPM. The disease should be strongly suspected in cases where there is evidence of multifocal asymmetric neurological disease, assuming the animal is in, or has been in, an endemic area. Focal muscle atrophy is another feature of disease, with gluteal muscle involvement the most commonly reported. Involvement of cranial nerves may be seen in around 10% of cases.11 The accuracy of a provisional diagnosis can be increased through laboratory testing. The subject of testing is controversial, and readers are referred to other texts that address this in greater detail.11 Essentially, most tests are designed to detect the presence of antibodies to parasite surface proteins. Methodologies include Western blot, immunofluorescence, and SAG-1 ELISA. Sensitivity and specificity vary between methods and laboratories. Antibodies are commonly found in the sera of normal horses, and more recently it has become clear that antibody may also be present in the cerebrospinal fluid (CSF) of neurologically normal animals.11 Blood contamination of CSF during the act of sampling can also account for a false-positive CSF antibody determination. At postmortem the lesions predominate in gray matter. The gross findings can vary in size and location. Histologically there is mononuclear cell cuffing of small vessels, parenchymal necrosis, and prominent astrocyte proliferation in more chronic cases. Parasites are seen in less than 50% of cases.12 Treatment strategies may include ponazuril R , Bayer Animal Heath) for a minimum (Marquis of 28 days and/or a combination of sulfadiazine and pyrimethamine. A pelleted form of diclazuril R (Protazil , Intervet Schering-Plough Animal Health)
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has been licensed but not marketed. Nitazoxanide has been used but is not currently available as an approved medication. Complete recovery is unusual, but most horses will improve with treatment. It has been suggested that prompt diagnosis and introduction of treatment can significantly improve the chances of complete recovery. Relapse can occur in around 10% of cases, and response rates to repeat treatment is generally not good.
Viral diseases Equine herpes type-1 myeloencephalopathy Most of the viral encephalitidies of horses are rarely seen in young animals. In a retrospective study of nine outbreaks of equine herpesvirus type-1 (EHV1) myeloencephalopathy in the Netherlands, none of the clinically affected animals was less than 3 years of age.13 The disease has been described in younger horses; weanlings were affected as part of a disease outbreak on a Thoroughbred breeding farm in the United Kingdom,14 and subtle signs were induced experimentally in two of nine young foals.15 EHV-1 myeloencephalopathy is primarily a disease of spinal cord, with the brain less commonly involved. Clinical signs can include limb ataxia, weakness, and dysmetria. The limb signs are often restricted to the pelvic limbs but can involve all limbs. Bladder dysfunction is also common. Signs are typically sudden in onset and may worsen over the initial 72 hours, with many making slow but steady improvement thereafter. The prognosis for animals that become recumbent or those with tetraplegia is guarded. The diagnosis is by PCR testing of a nasopharyngeal swab or peripheral blood leukocytes. The underlying lesion is a vasculopathy involving thrombosis and vascular leakage. The CSF is consequently discolored yellow (xanthochromic) with a normal leukocyte concentration and a normal or elevated total protein concentration. Antiviral drugs have been used (valacyclovir 40 mg/kg p.o. t.i.d.) although experimental data are not compelling.16 Aspirin may also be of some value in the early stages of the disease. Strict property quarantine for at least 3 weeks from the final day of fever in the last affected animal is recommended.17 West Nile virus West Nile virus (WNV) is a flavivirus that continues to have an enormous economic and emotional impact on the horse industries of many regions of the world. The recent spread across North America has raised the importance of this viral disease. Although uncommon, foals can develop clinical WNV
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disease. In a survey of 569 horse cases from North Dakota, the median age of affected animals was 8 years with a range from 3 months to 30 years.18 Similarly, a study of cases from Colorado and Nebraska reported a mean age of cases of 9.5 years with a range from 3 months to 35 years.19 Another epidemiological study in Indiana reported the youngest affected animal as 5 months, with 6% of cases less than 12 months of age.20 Clinical signs frequently include low-grade fever, ataxia and paresis which may progress to recumbency, and muscle fasciculations and tremor. The diagnosis of WNV infection is based on location, typical clinical findings, and laboratory testing. These include an IgM capture ELISA, a plaque reduction neutralization test, or a hemagglutination inhibition assay.13 The prognosis for survival appears to be around 70%. Vaccination is available in several endemic regions (see Chapter 31). Eastern/Western/Venezuelan equine encephalitis Clinical signs include high fever and evidence of diffuse forebrain disease, including obtundation, compulsive walking, head pressing, ataxia, recumbency, and seizure. Cortical blindness may also be present. Mortality rates are high, in the range 75–95% for EEE, 19–50% for WEE, and 40–90% for VEE. Surviving horses gradually recover over weeks to months, but many retain signs of CNS damage. The diagnosis is based on location, time of the year (due to presence of mosquito vector), clinical signs, and laboratory data. CSF in the early stages of infection reveals a neutrophilic pleocytosis which tends towards a mononuclear response in surviving animals. Antigen-specific IgM and IgG antibody detection in serum is helpful in confirming the diagnosis. Protection of foals against Eastern/Western/Venezuelan Equine Encephalitis viruses is through maternal vaccination and ensuring adequate passive transfer of immunoglobulin.
Hyperammonia syndromes Differentials for hyperammonemia include hepatitis/hepatopathy or a portosystemic shunt. There is evidence to indicate that an inherited condition of Morgans exists that is similar to hyperornithinemiahyperammonemia-homocitrullinuria (HHH) syndrome in humans.21 This syndrome is attributed to an inherited defect in a mitochondrial ornithine transporter protein, such that the synthesis of urea is impaired, leading to increased serum concentrations of ammonia and ornithine. Signs often develop around the time of weaning and may include ob-
tundation, dementia, and reduced feed intake. Foals can aimlessly wander, circle, and head press. Laboratory values include a persistently elevated blood ammonia concentration. Management could include feeding a low-protein diet and lactulose (0.2 g/kg p.o. t.i.d.). Acute signs can also be managed with an intravenous infusion of dextrose/glucose. The long-term outcome for affected foals is poor. Most foals with congenital portosystemic shunts become clinically apparent within the first 12 months of life.22–24 Signs can be variable and intermittent, and have included obtundation, lethargy, ataxia, and blindness. Most foals are poorly grown. Serum levels of ammonia and bile acids are elevated, with little change in liver-derived enzymes. Shunts can be diagnosed using mesenteric portovenography or nuclear scintigraphy. In the latter the hepatic uptake of 99m Technetium-labeled sulfur colloid after intravenous administration is measured using a gamma camera. Equine motor neuron disease Equine motor neuron disease (EMND) is a neurodegenerative disease of motor neurons in the brain and spinal cord. The disease is highly unlikely to occur in juvenile horses, with the lower end of the reported age of onset at 2 years and the median age of onset around 10 years of age.25 The disease is characterized clinically by progressive weakness, muscle fasciculations, and loss of muscle mass. EMND has been induced by feeding a diet low in vitamin E and high in copper and iron, along with grass hay that had been in storage for 12 months. Horses were maintained in dirt lots and four of eight horses developed signs consistent with EMND after 21–28 months.26 A follow-up investigation by the same group documented EMND in 10 horses maintained on a vitamin E deficient diet with disease evident at a median time of 38.5 months after enrolment, with the earliest case at 18 months post-enrolment.27 Vitamin E deficiency is also associated with retinal degeneration. Not only is there a long period of deficiency required to induce EMND, the histopathological changes differ from those seen in other neurological diseases attributed, in part, to vitamin E deficiency, such as equine degenerative myeloencephaloapthy (EDM). EDM is an axonal degenerative disorder that produces changes in the distal axons of spinal proprioceptive tracts without involvement of motor neurons.
References 1. Levine JM, Adam E, MacKay RJ, Walker MA, Frederick JD, Cohen ND. Confirmed and presumptive cervical vertebral
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compressive myelopathy in older horses: a retrospective study (1992–2004). J Vet Intern Med 2007;21(4):812–19. Moore BR, Reed SM, Biller DS, Kohn CW, Weisbrode SE. Assessment of vertebral canal diameter and bony malformations of the cervical part of the spine in horses with cervical stenotic myelopathy. Am J Vet Res 1994;55(1):5–13. Mayhew IG, Donawick WJ, Green SL, Galligan DT, Stanley EK, Osborne J. Diagnosis and prediction of cervical vertebral malformation in thoroughbred foals based on semi-quantitative radiographic indicators. Equine Vet J 1993;25(5):435–40. Hahn CN, Handel I, Green SL, Bronsvoort MB, Mayhew IG. Assessment of the utility of using intra- and intervertebral minimum sagittal diameter ratios in the diagnosis of cervical vertebral malformation in horses. Vet Radiol Ultrasound 2008;49(1):1–6. van Biervliet J, Scrivani PV, Divers TJ, Erb HN, de Lahunta A, Nixon A. Evaluation of decision criteria for detection of spinal cord compression based on cervical myelography in horses: 38 cases (1981–2001). Equine Vet J 2004;36(1):14– 20. Mayhew IG, deLahunta A, Whitlock RH, Krook L, Tasker JB. Spinal cord disease in the horse. Cornell Vet 1978; 68 Suppl 6: 1–207. Hudson NPH, Mayhew IG. Radiographic and myelographic assessment of the equine cervical vertebral column and spinal cord. Equine Vet Educ 2005;17(1):34–8. Donawick WJ. Mayhew IG, Galligan DT, Osborne J, Green S, Stanley EK. Early diagnosis of cervical vertebral malformation in young Thoroughbred horses and successful treatment with restricted, paced diet and confinement. Proceedings of the 35th annual convention of the American Association of Equine Practitioners, 1989; p. 525–8. Cohen ND, Mackay RJ, Toby E, Andrews FM, Barr BS, Beech J, Bernard WV, Clark CK, Divers TJ, Furr MO, Kohn CW, Levy M, Reed SM, Seahorn TL, Slovis NM. A multicenter case–control study of risk factors for equine protozoal myeloencephalitis. J Am Vet Med Assoc 2007;231(12):1857–63. Duarte PC, Conrad PA, Wilson WD, Ferraro GL, Packham AE, Bowers-Lepore J, Carpenter TE, Gardner IA. Risk of postnatal exposure to Sarcocystis neurona and Neospora hughesi in horses. Am J Vet Res 2004;65(8):1047–52. Furr MO. Equine protozoal myeloencephalitis. In: Furr MO, Reed S (eds) Equine Neurology. Ames, IA: Blackwell Publishing, 2008; pp. 197–212. MacKay RJ, Granstrom DE, Saville WJ, Reed SM. Equine protozoal myeloencephalitis. Vet Clin North Am Equine Pract 2000;16(3):405–25. Goehring LS, van Winden SC, van Maanen C, Sloet van Oldruitenborgh-Oosterbaan MM. Equine herpesvirus type 1-associated myeloencephalopathy in The Netherlands: a
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four-year retrospective study (1999–2003). J Vet Intern Med 2006;20(3):601–7. Greenwood RE, Simson AR. Clinical report of a paralytic syndrome affecting stallions, mares and foals on a thoroughbred studfarm. Equine Vet J 1980;12(3):113–17. Patel JR, Edington N, Mumford JA. Variation in cellular tropism between isolates of equine herpesvirus-1 in foals. Arch Virol 1982;74(1):41–51. Garre B, Gryspeerdt A, Croubels S, De Backer P, Nauwynck H. Evaluation of orally administered valacyclovir in experimentally EHV1-infected ponies. Vet Microbiol 2009;135(3–4):214–21. Goehring L. Viral diseases of the nervous system. In: Furr MO, Reed S (eds) Equine Neurology. Ames, IA: Blackwell Publishing, 2008; pp. 169–96. Ndiva Mongoh M, Hearne R, Dyer NW, Khaitsa ML. The economic impact of West Nile virus infection in horses in the North Dakota equine industry in 2002. Trop Anim Health Prod 2008;40(1):69–76. Salazar P, Traub-Dargatz JL, Morley PS, Wilmot DD, Steffen DJ, Cunningham WE, Salman MD. Outcome of equids with clinical signs of West Nile virus infection and factors associated with death. J Am Vet Med Assoc 2004;225(2): 267–74. Ward MP, Levy M, Thacker HL, Ash M, Norman SK, Moore GE, Webb PW. Investigation of an outbreak of encephalomyelitis caused by West Nile virus in 136 horses. J Am Vet Med Assoc 2004;225(1):84–9. McConnico RS, Duckett WM, Wood PA. Persistent hyperammonemia in two related Morgan weanlings. J Vet Intern Med 1997;11(4):264–6. Buonanno AM, Carlson GP, Kantrowitz B. Clinical and diagnostic features of portosystemic shunt in a foal. J Am Vet Med Assoc 1988;192(3):387–9. Hillyer MH, Holt PE, Barr FJ, Weaver BM, Brown PJ, Henderson JP. Clinical signs and radiographic diagnosis of a portosystemic shunt in a foal. Vet Rec 1993;132(18): 457–60. Fortier LA, Fubini SL, Flanders JA, Divers TJ. The diagnosis and surgical correction of congenital portosystemic vascular anomalies in two calves and two foals. Vet Surg 1996;25(2):154–60. MacKay RJ. Neurodegenerative disorders. In: Furr MO, Reed S (eds) Equine Neurology. Ames, IA: Blackwell Publishing, 2008; pp. 238–46. Divers TJ, Cummings JE, de Lahunta A, Hintz HF, Mohammed HO. Evaluation of the risk of motor neuron disease in horses fed a diet low in vitamin E and high in copper and iron. Am J Vet Res 2006;67(1):120–6. Mohammed HO, Divers TJ, Summers BA, de Lahunta A. Vitamin E deficiency and risk of equine motor neuron disease. Acta Vet Scand 2007;49: 17.
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Chapter 81
Diarrhea Kirsten M. Neil
Introduction
Infectious diseases
Diarrhea is a common problem of the older foal, weanling, and yearling, with up to 80% of foals reported to have an episode of diarrhea in the first 6 months.1 Single foals may be affected, or the clinician may be presented with an outbreak amongst foals of similar ages. As both infectious and non-infectious disorders can cause diarrhea, identification of the underlying cause is vital so that appropriate treatment, control, and preventative measures can be instigated. However, similar to adult horses with enterocolitis, the specific etiological agent may not be identifiable in all cases.2 Infectious causes of diarrhea are common. Some pathogens, such as Clostridia spp and Salmonella spp, can cause diarrhea in all age groups but infection tends to be most severe in neonates.3 Other infectious organisms which commonly cause colitis in the suckling foal (1 week–2 months) include rotavirus, Cryptosporidium, and Strongyloides westeri. The same organisms can also cause diarrhea in the 2- to 6-month-old foal, although other parasites, Rhodococcus equi, and equine proliferative enteropathy are the main differentials in this age group. For 6- to 12-month-old foals, parasites are commonly implicated. Common non-infectious causes of diarrhea include gastroduodenal ulceration and sand enteropathy. Other less frequent causes of colitis in this age group are common to adults and include toxicological causes (e.g., antimicrobial and non-steroidal antiinflammatory drug induced), feed-related causes (grain overload, dietary change), and rarely infiltrative bowel disease. Diarrhea is most commonly a sign of gastrointestinal disease; however it may be associated with disease of other organ systems such as peritonitis.
Diagnosis of the cause of diarrhea can be problematic. Clinical signs are usually similar regardless of the cause, with acute profuse diarrhea, dehydration and electrolyte imbalances, endotoxemia, hypoproteinemia, and pyrexia common. Chronic diarrhea, ill-thrift, and edema are also common to many etiological agents. Isolation of the organism from feces of affected foals is often the basis of the diagnosis; however some organisms such as Rhodococcus equi, Clostridium perfringens, and rotavirus can be isolated from normal foals.4 Newer molecular tests are now available which provide rapid results and are a valuable aid in differentiating between causes of infectious diarrhea. Intestinal clostridiosis
Clostridium difficile Clostridium difficile is a Gram-positive spore-forming obligate anaerobe associated with enterocolitis in both foals and adult horses. A number of clinical manifestations have been identified. C. difficile is the most common cause of antimicrobial-associated diarrhea, and is also a primary pathogen infecting foals of any age group, with the most severe disease recognized in septic neonates. Diarrhea has been associated with the use of erythromycin, potentiated sulfonamides, and beta-lactam antimicrobials in older foals and yearlings.5 Acute colitis in mares whose foals are being treated with erythromycin has been well documented.6 The role of C. difficile in the development of antimicrobial-associated diarrhea in foals is less clear. Colonization by the organism may occur following antimicrobial treatment without evidence of clinical disease. C. difficile was cultured from feces of 44% of 1- to 3-month-old foals without
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Diarrhea diarrhea that were treated with erythromycin or gentamicin with rifampin, and from foals without diarrhea treated with penicillin and/or trimethoprim sulphonamides but only if they were less than 2 weeks of age.7 C. difficile is infrequently isolated from normal healthy foals, suggesting that healthy foals may be a potential reservoir of infection. However isolation appears to be age dependent with the organism cultured from feces of foals less than 2 weeks of age, but not from 1- to 6-month-old foals.6, 7 C. difficile has also been recognized as a primary cause of acute diarrhea in older foals, albeit with variable frequency.7–9 In a retrospective study of 28 foals with C. difficileassociated diarrhea, the mean age of affected foals was 21.5 days (range 1–120 days), with 29% of foals between 3 weeks and 4 months of age.9 In a recent study, 32% of horses with diarrhea associated with the organism were less than 6 months of age.10 Although C. difficile should be strongly suspected in septic neonates with acute, severe hemorrhagic diarrhea, clinical signs associated with the organism in older foals are typically indistinguishable from disease caused by other infectious organisms. Diarrhea may be mild, but acute profuse watery diarrhea is more typical. Signs attributable to endotoxemia are common including pyrexia, tachycardia, and tachypnea.9, 10 Colic due to ileus has also been reported.9, 10 Common clinicopathological abnormalities include hemoconcentration, elevated band neutrophil counts, hyperfibrinogenemia, and electrolyte abnormalities such as hypocalcemia and hyponatremia.9, 10 While other modalities such as ultrasonography are helpful to establish a diagnosis of enterocolitis, findings of thickened hypomotile intestine are not specific for clostridial disease, although the presence of gas-filled loops of small intestine or the appearance of hyperechoic areas which cast an acoustic shadow suggestive of air in the wall of the intestine should raise suspicion. Confirmation of the diagnosis can be difficult. Isolates of C. difficile are considered virulent if they produce one or more exotoxin (toxins A and B) which are responsible for causing intestinal inflammation.9 Isolation of the organism from feces requires anaerobic culture using specific culture media. A positive culture is usually considered significant, however definitive diagnosis requires detection of toxins A (enterotoxin) and/or B (cytotoxin) in feces using cytotoxin assays. PCR and enzyme immunoassay tests are also available.4, 9 Although toxin A ELISA has been considered less sensitive than PCR,9 sensitivity and specificity of a toxin A ELISA was good compared to microbial culture.10 Although not definitive, the organism can be suspected as a potential etiological agent before other test results become
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available, based on the detection of large numbers of Gram-positive rods on cytology of feces from foals with diarrhea. Multiple daily samples may need to be submitted as the organism and its toxins may not be detected until late in the course of the disease.11, 12 Young foals should have blood cultures submitted for anaerobic culture, due to the likelihood of concurrent sepsis and bacteremia. The prognosis for foals with C. difficile diarrhea is fair to good;9, 10 however, the survival of neonates with severe hemorrhagic diarrhea is generally poor. In a recent retrospective study, horses with toxin A detected in feces had more severe disease and a higher mortality rate than those negative for toxin A.10 Treatment and infection control are discussed in a subsequent section.
Clostridium perfringens In a retrospective study of infectious diarrhea, C. perfringens was significantly associated with diarrhea in foals less than 6 months of age.1 C. perfringens has also been isolated from the feces of normal foals.1, 2, 13 In one study, the organism was isolated from 57% of foals with diarrhea compared to only 27% of healthy foals,1 while in a another study the prevalence was similar between healthy foals and foals with diarrhea.2 Isolation of the organism from healthy foals was significantly greater in foals that were in contact with foals with diarrhea compared to foals not in contact with diarrhea. Isolation of the organism from feces is common in healthy neonatal foals up to 3 days of age, but a lower percentage of 1- to 2-month-old foals and adults shed C. perfringens type A.13 There are five major biotypes of Clostridium perfringens, categorized based on the production of several pathogenic exotoxins which cause necrosis of the intestinal wall. Biotypes A and C are most commonly associated with fatal hemorrhagic enterocolitis in neonatal foals.14 The organism has been significantly associated with fatal illness with most deaths occurring in neonates,1 and the mortality rate is higher with biotype C than biotype A.15 C. perfringens enterotoxin is most commonly associated with type A strains;16 however, the role of this toxin in foal diarrhea is unclear. Enterotoxin was detected in the feces of 29% of foals with diarrhea but not in healthy foals,8 while in another study enterotoxin was detected in 16% of fecal samples from horses with diarrhea.16 Clinical signs associated with C. perfringensinduced diarrhea are most severe in neonates less than 10 days of age, with sudden death, colic, hemorrhagic diarrhea, and sepsis common.3, 14 In older foals, diarrhea may be milder but clinical signs and clinicopathological findings are similar to other
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causes of acute enterocolitis such as C. difficile. Culture of high numbers of the organism from feces is suggestive of infection (> 1 × 106 CFU/g, normal horses shed < 100 CFU/g);14 however, definitive diagnosis relies on identification of toxin in feces. An ELISA is commercially available for the detection of enterotoxin in feces.16 However, as enterotoxin is not produced by all pathogenic strains of C. perfringens, the reliability of enterotoxin immunoassays has been questioned, with PCR biotyping suggested as a preferred method of diagnosis.17 As for C. difficile, cytology of fecal samples may be a useful rapid screening test but is not definitive for disease due to this organism.
Treatment Principles of treatment of diarrhea discussed in this section are common to the management of intestinal clostridiosis and other cause of enterocolitis. Treatment of diarrhea is generally supportive regardless of the initiating cause, and should be tailored to the individual foal depending on the severity of diarrhea and associated conditions such as endotoxemia. Hydration status, glucose and electrolyte concentrations, acid–base balance, and, if available, oncotic pressure should be evaluated and monitored regularly. Intravenous fluid therapy, colloids, plasma, and correction of electrolyte abnormalities are the mainstay of most treatment regimes. Fluid therapy and correction of electrolyte and acid–base balance is discussed in detail in Chapters 19 and 21. In young foals initial fluid replacement, based on level of dehydration, is with isotonic crystalloids (Lactated R ) Ringer’s Solution, Plasmalyte 148, Nomosoi administered as i.v. boluses of 10–12 mL/kg over 20–30 minutes.18 Maintenance fluids are usually administered at 2–3 times normal maintenance rates (3 mL/kg/h older foal, 2 mL/kg/h yearling) to compensate for estimated fluid losses in diarrhea. A constant rate infusion is ideal, however fluids can be bolused every 4–6 hours in less severe cases. Additional electrolyte supplementation with potassium, bicarbonate, magnesium, and calcium is often required. Hypertonic saline is an effective resuscitative therapy (7–7.5% solution, 2–4 mL/kg) as it provides immediate improvement in perfusion, but should be administered with care in foals with unknown electrolyte levels, as rapid correction of hyponatremia can predispose to central pontine myelinolysis. Synthetic colloids (e.g., Hetastarch 5–10 mL/kg) and plasma (20–40 mL/kg) are often needed to increase oncotic pressure and maintain circulating volume in foals with hypoproteinemia and hypoalbuminemia. Concurrent administration of
colloids with resuscitative fluids may be indicated if circulating fluid volume is not restored with the initial crystalloid fluid boluses. Crystalloid fluids rapidly leave the vascular space to equilibrate with extracellular fluid. Administration of colloids during volume resuscitation preserves oncotic pressure and results in more effective volume expansion.18 Plasma not only improves oncotic pressure, but also supplies fibronectin, complement, and other factors, while potentially reducing the effects of endotoxemia. Withholding or restricting milk intake is beneficial for nursing foals, especially if there is evidence of colic, ileus, or abdominal distension. Nutritional support in the form of partial or total parenteral nutrition (see Chapter 28) may be required. Mild cases may simply require the addition of intravenous glucose (5%). Debilitated foals will often require parenteral nutrition with dextrose, amino acid and lipid solutions. The use of antimicrobials in the treatment of horses with enterocolitis is controversial. However, broad-spectrum antimicrobial administration is indicated in young foals or those with evidence of sepsis or severe neutropenia because of the risk of translocation of bacteria across the intestinal tract. Suggested antimicrobials are potassium penicillin (20–40 × 103 IU/kg i.v. q 6 h) or ceftiofur sodium (5–10 mg/kg i.v. q 6–12 h) with an aminoglycoside (amikacin 25 mg/kg i.v. q 24 h young foal or 10–15 mg/kg older foal, gentamicin 15 mg/kg i.v. q 24 h young foal or 6.6–8.8 mg/kg older foal). For foals with intestinal clostridiosis, specific antimicrobial treatment is indicated; however, if diarrhea developed associated with antimicrobial administration, antimicrobials should be discontinued. Metronidazole (neonatal foals: 10–15 mg/kg p.o/i.v. q8–12hr; older foals and adults: 15–20 mg/kg p.o. q8hr) is recommended for treatment of intestinal clostridiosis; however, susceptibility testing of isolates should be performed as resistance to metronidazole has been infrequently reported for C. difficile.9 Bacitracin and vancomycin may also be effective, but should not be considered as first-line drugs of choice.12 The use of vancomycin should be limited due to concerns about developing resistance. Administration of flunixin meglumine (0.25–0.5 mg/kg i.v. q 8 h) is a mainstay of anti-endotoxin therapy but may reduce mucosal perfusion. Polymixin B (5000–6000 IU/kg i.v. q 8–12 h) acts as a chelating agent binding to lipopolysaccharide,19 and in a study in foals administered endotoxin, ameliorated clinical signs of fever and tachycardia, and decreased TNFα concentrations.20 In healthy horses, no adverse effects were seen,21 but caution is recommended in dehydrated horses and neonates.22 Pentoxifylline (8 mg/kg p.o. q 8–12 h) may decrease
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Diarrhea endothelial permeability, quicken mucosal healing, and reduce leukocyte activation and migration,23 and is often used in adult horses for treatment of endotoxemia. The efficacy of pentoxifylline has been questioned and its safety in foals has not been determined.24 Its efficacy may be enhanced with concurrent use of flunixin meglumine.25 Non-steroidal anti-inflammatories should be used with care in young foals, and low doses or alternative analgesics used in the colicky foal. Effective pain management may be achieved through the use of butorphanol (0.04–0.1 mg/kg i.v.). Lidocaine (1.3 mg/kg slow i.v. bolus then 0.05 mg/kg/min constant rate infusion i.v.) is used as a prokinetic in foals with ileus, and may have a place in the treatment of entercolitis due to its potential analgesic, antiinflammatory, and ischemia-reperfusion effects. Hyperimmune plasma specific for C. perfringens and C. difficile is available in some countries and may be beneficial.26 Proton pump inhibitors and H2 antagonists (omeprazole 4 mg/kg p.o. q 24 h, ranitidine 5–10 mg/kg p.o. q 6–12 h) are often administered concurrently, due to the propensity for foals with other clinical disorders to develop gastric ulcers. In addition to gastroprotection, sucralfate (10–20 mg/kg p.o. q 6 h) may have other beneficial effects such as increasing local prostaglandin production and blood flow.22 Intestinal protectants and absorbents such as kaolin/pectin or bismuth salicylate are often administered with the aim of limiting absorption of endotoxins and exotoxins and coating mucosa. Kaolin/pectin products (0.5–4 mL/kg p.o. q 6–24 h) bind to mucosa and have a coating effect. Possible beneficial effects of bismuth subsalicylate (0.5–4 mL/kg p.o. q 4–6 h) include stimulation of fluid and electrolyte absorption, inhibition of prostaglandin synthesis, and binding of bacterial toxins. Additionally, salicylate may have some antiprostaglandin activity but should be used with care in foals with impaired renal function. Use of ditrioctahedral (DTO) smectite (BioR sponge ) has gained popularity in recent years. This product is a natural hydrated clay aluminomagnesium silicate which has been used in human medicine to treat infectious colitis. In vitro, ditrioctahedral smectite was effective in absorbing C. difficile toxins A and B and C. perfringens exotoxins and enterotoxin.27, 28 Clinical trials demonstrating its efficacy in the treatment of enterocolitis in horses are lacking, however postoperative diarrhea was significantly reduced when DTO smectite was administered (0.5 kg by NGT q 24 h for 3 days) in a clinical trial of mature horses with large intestinal surgical diseases.29 Recommended dose rates are foals 23.5 g p.o. q 12h and adults 97.5 g p.o. q12h. Lactose intolerance may
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develop secondary to loss of the small intestinal brush border. Affected foals typically develop diarrhea again once nursing has been reintroduced. As lactose intolerance has been reported secondary to C. difficile enteritis,11 lactase supplementation (6000 units/50 kg p.o. q 3–4 h) may be beneficial. Probiotics are commonly administered to horses with diarrhea in an attempt to re-establish normal intestinal flora, although their efficacy is questionable. Most commercial preparations contain Lactobacillus, with administration of 1 × 109 –1 × 1010 CFU per 50 kg bodyweight suggested.30 Efficacy trials in older foals are lacking. In a clinical trial evaluating the efficacy of probiotic administration (Lactobacillus pentosus) for prevention of diarrhea in normal 24to 48-hour-old foals, administration was associated with the development of diarrhea, anorexia, and colic.31 The non-pathogenic yeast Saccharomyces boulardii (1–2 g per 50 kg bodyweight per day or 5 × 109 CFU p.o. q 12 h) may compete with C. difficile toxin A for binding sites on intestinal epithelium,15 and has been suggested for the prevention and treatment of intestinal clostridiosis. In a prospective study of horses with acute enterocolitis, the severity and duration of gastrointestinal disease was significantly reduced in horses administered Saccharomyces boulardii compared to placebo.32 Prebiotics such as psyllium (1 g/kg divided daily) may be beneficial. Provision of a source of fermentable fiber such as psyllium may modify fecal consistency through increasing fecal bulk and stimulating the production of short-chain fatty acids such as butyrate which stimulate enterocyte and colonocyte proliferation and mucosal healing.30, 33 Administration of C. perfingens type C antitoxin has been suggested,15, 22 but requires off-label use. Efficacy of this treatment in foals is unknown.
Prevention, infection control and management strategies Infection control strategies should be employed for foals with acute diarrhea to prevent spread of infection and minimize environmental contamination. Affected foals should be isolated, and disposable protective clothing, gloves, and boots worn by caregivers. Good hygiene is important, not only to prevent spread of infection, but also because clostridial organisms are human pathogens. Hand washing with antimicrobial soap and the use of alcohol-based hand sanitizers is recommended. Clostridial spores are highly resistant and can survive in the environment for years. Most disinfectants are not effective against clostridial spores. Sodium hypochlorite (bleach, 1:10 dilution) can be effective as long as there is no organic debris present.34 Newer
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R peroxygen products (e.g., Virkon S ) are also likely to have sporicidal activity and are not inactivated by organic matter. Administration of prophylactic antimicrobials (metronidazole) to newborn foals on farms with outbreaks of clostridial enterocolitis has been suggested,22 although this practice may predispose to the development of metronidazole resistance, and is unlikely to prevent clostridial infections in older foals. There are no vaccines currently available with proven efficacy and safety in horses.
Salmonellosis Salmonellosis is an important cause of acute enterocolitis and outbreaks of diarrhea in all age groups of horses, and has significant zoonotic and nosocomial potential. In two epidemiological surveys of foal diarrhea, the prevalence of Salmonella was low,1, 2 but when present was significantly associated with a fatal outcome.1 Diarrhea is both secretory and malabsorptive in nature, the latter secondary to intestinal inflammation. A number of virulence factors have been identified, including exotoxin, cytotoxin, and enterotoxin.35 Infection with Salmonella group B isolates is common.17 The most commonly reported serotype is Salmonella typhimurium, however a number of other serotypes have been associated with colitis in horses, including S. agona, S. newport, S. dublin, S. agona, and S. infantis35 The presence of carriers which actively shed Salmonella in feces is reportedly high (10–20%), but their prevalence in the general population may be less than 1%.36, 37 When risk factors associated with fecal shedding of Salmonella in a hospitalized population were investigated, the prevalence of shedding was significantly higher in foals up to 1 year of age compared to adult horses.38 Similarly, foals with gastrointestinal disease were significantly more likely to be shedding than adults with gastrointestinal disease. The age range of foals in this study was 1–250 days, with foals older than 38 days more likely to shed Salmonella in feces compared to younger foals. Foals with diarrhea have been the index case in a number of outbreaks in veterinary hospitals.39, 40 Serotype virulence, host susceptibility, environmental factors, and immune status are important determinants of disease severity. S. typhimurium is the most pathogenic serotype and has been associated with a high mortality rate, particularly in outbreaks. Multidrug-resistant S. newport has also become an important pathogen in recent years. Transportation, administration of antimicrobials, underlying gastrointestinal disease, gastrointestinal surgery, and season (peak incidence in late summer and fall) have been associated with an increased risk of infection.17, 41
Salmonella causes severe fibrinonecrotic typhlocolitis and is associated with clinical signs of endotoxemia, fever, anorexia, dehydration, acute profuse watery diarrhea, and occasionally colic.17 Depression, fever, anorexia, and neutropenia may occur without diarrhea. Foals less than 1 month of age are frequently bacteremic, and extraintestinal signs of disease such as septic arthritis, osteomyelitis, pneumonia, uveitis, and meningitis are usually restricted to foals less than 2–3 months of age.4 Chronic diarrhea may occur. Common clinicopathological abnormalities include hemoconcentration, neutropenia, increased band neutrophils, toxic changes, hypoproteinemia, azotemia, electrolyte abnormalities (hyponatremia, hypochloremia, hypokalemia), acidosis, and hyperlactatemia; however, these changes are not specific for Salmonella enterocolitis. Coagulopathies, thrombocytopenia, and disseminated intravascular coagulation (DIC) may also be evident. Diagnosis is based on repeated fecal culture, with five daily cultures recommended as the organism is shed intermittently. Large fecal samples are preferred (10–30 g). The sensitivity of fecal culture is low (55%) with single samples,42 but improved with multiple fecal cultures (> 90%).3, 43 The concurrent use of fecal cultures and rectal mucosal biopsy cultures increases the sensitivity to 60–75%.44 PCR of feces is rapid, and is the most sensitive and specific test currently available,43, 45 with a reported sensitivity of 100% and specificity of 98%;45 however, false-negative results are possible. Serotyping and antimicrobial sensitivity testing cannot be achieved with PCR, so concurrent fecal cultures should also be submitted. Blood cultures should be performed in young foals due to the high incidence of septicemia. Adult horses are rarely bacteremic;17 however, blood cultures should be considered as concurrent sepsis with Pseudomonas aeruginosa has been reported in an adult horse with enteric salmonellosis,46 and positive blood cultures in adult horses with diarrhea have recently been associated with a poorer prognosis.47 Affected horses usually require intensive intravenous fluid therapy, colloids, and plasma as well as other supportive care measures. The reader is referred to the previous section on treatment of intestinal clostridiosis for further information. The use of antimicrobials in the treatment of enterocolitis in adults is controversial and is not thought to alter the course of the disease17 or reduce fecal shedding.42 In contrast, antimicrobials are strongly indicated when treating affected foals, especially those that are immunocompromised, bacteremic, septicemic, or neutropenic. Broad-spectrum antimicrobials are usually recommended until antibiogram results are available. Specific antimicrobials which are
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Diarrhea usually effective against Salmonella are aminoglycosides (amikacin, gentamicin) and third-generation cephalosporins such as ceftiofur sodium. Systemic fluoroquinolones (enrofloxacin 5 mg/kg i.v. q 24 h) are currently effective against most Salmonella isolates;48 however, their use should be restricted to avoid the development of resistance.26 The use of enrofloxacin in young horses is discouraged due to potential detrimental effects on cartilage. Administration of hyperimmune plasma may be beneficial, however no positive effect was seen in a study in 3- to 5-month-old foals administered endotoxin, and clinical signs were in fact exacerbated by the administration of hyperimmune plasma.20 Administration of probiotics has had no effect on Salmonella shedding or prevalence of diarrhea in clinical trials of postoperative colic patients.49, 50 . Catheter sites should be closely monitored as endotoxemia and salmonellosis are documented risk factors for the development of catheter-associated thrombophlebitis.51 Despite aggressive treatment, the mortality rate associated with salmonellosis is often high due to complications such as DIC, multi-organ dysfunction, laminitis, acute renal failure, and gastrointestinal infarction.17, 52 The prognosis for horses with chronic diarrhea is poor, as is the prognosis for foals with extraintestinal signs of disease such as septic arthritis and osteomyelitis.53 Biosecurity and management practices to limit spread of infection should be implemented as described previously for clostridial disease. In particular, an infectious etiology such as Salmonella should be assumed in horses with acute profuse diarrhea until proven otherwise. Foals should be isolated, and strict attention paid to personal hygiene and sanitation as the zoonotic potential of this organism is high. Most disinfectants are effective against Salmonella including bleach, quaternary ammonium disinfectants, peroxygen compounds, and phenolic disinfectants.54 There is no licensed vaccine currently available in the United States for use in horses. An inactivated Salmonella typhimurium vaccine is available in Australia for use in pregnant mares and foals. The vaccine is thought to be effective;52 however, clinical trials demonstrating its efficacy in reducing fecal shedding and clinical disease are lacking. Lawsonia intracellularis Lawsonia intracellularis is the causative organism of equine proliferative enteropathy (EPE), an enteric disease most commonly documented in 2- to 8-month-old foals, especially after stress such as weaning.55–57 The condition has been reported in
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Figure 81.1 Ultrasonogram of increased small intestinal mural thickness (broken line) in a weanling with diarrhea, weight loss, edema, and colic.
foals up to 13 months of age. Diarrhea is a common presenting sign, and affected foals often have concurrent signs of weight loss, anorexia, lethargy, depression, and ventral edema due to hypoproteinemia. Diarrhea is typically acute, but may be intermittent in some cases.56, 58 Manure consistency varies from soft ‘cow pat’ feces to profuse watery diarrhea;17, 55 however, some foals may have normal feces. In a recent retrospective study of 57 foals with EPE, ventral edema was the most common clinical sign, and only 26% of foals had diarrhea of varying severity.57 Diarrhea occurs due to mucosal thickening and proliferation of the small intestine, resulting in a secretory diarrhea. Increased small intestinal mural thickness is often detected sonographically (Figure 81.1). The most consistent laboratory finding is hypoproteinemia due to hypoalbuminemia.55, 57, 59 Although infrequently tested, glucose and xylose absorption tests have been normal in most foals;55, 60, 61 however, complete or partial small intestinal malabsorption has been documented in recent cases.62, 63 Currently available diagnostic tests include fecal PCR and determination of serum antibody titer using immunoperoxidase monolayer assay (IPMA). Both tests should be performed in suspect cases, as only 50% of affected foals tested positive on both fecal PCR and IPMA in a recent study.57 Diarrhea typically resolves quickly once appropriate antimicrobial treatment has been instigated. Effective antimicrobials include a macrolide (erythromycin estolate 25 mg/kg p.o. q 8 h; azithromycin 10 mg/kg p.o. q 24 h; clarithromycin 7.5 mg/kg p.o. q 12 h) alone or in combination with rifampin (5–10 mg/kg p.o. q 12 h), oxytetracycline (6.6 mg/kg i.v. q 12 h), doxycycline (10 mg/kg p.o. q 12 h) or chloramphenicol (44 mg/kg p.o. q 6–8 h).55–57,59 Additional supportive therapy in the form of intravenous fluid therapy and gastroprotectants is recommended. Administration of colloids (Hetastarch and/or plasma) is indicated in most foals as hypoproteinemia is a common finding. Readers are referred to
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Chapter 83 where this topic is addressed in more detail. Rhodococcus equi Rhodococcus equi is a common cause of pneumonia in foals 1–6 months of age, with most foals displaying clinical signs between 1 and 3 months of age. Extrapulmonary manifestations of infection may also occur, with or without signs of pneumonia. Intestinal involvement was evident on postmortem examination in around 50% of pneumonic foals in one study.64 In a recent study of 150 foals presenting to a university hospital, extrapulmonary disorders (EPD) were identified in 77% of foals.65 The most common EPD were diarrhea, immune mediated synovitis, ulcerative enterotyphlocolitis, intraabdominal abscessation, abdominal lymphadenitis, and pyogranulomatous hepatitis.65 The intestinal form of infection is characterized by multifocal ulcerative enterocolitis and typhlitis with suppurative or granulomatous inflammation of mesenteric lymph nodes.66 Diarrhea, colic, weight loss, and poor growth are common. Diagnosis can be difficult, as Rhodococcus equi may be cultured from feces of normal foals without evidence of infection.4 In foals with concurrent pneumonia, diagnosis of R. equi as the cause of diarrhea is often assumed when the organism is established as the cause of the pneumonia; however, other causes of diarrhea should be excluded. In a recent study, the survival of foals with EPD was only 46% compared with 80% of foals without EPD; however, many of the EPD were only identified on postmortem examination.65 Leukocytosis, neutrophilia, and azotemia were commonly reported, suggesting that foals with profound leukocytosis and azotemia should be thoroughly assessed for possible EPD. Diarrhea may also occur in affected foals as a side effect of treatment with erythromycin67 associated with disruption of gastrointestinal flora and stimulation of motilin receptors. Softening of feces is generally self-limiting; however, profuse diarrhea can occur in some foals. Diarrhea occurred more commonly in pneumonic foals treated with erythromycin compared to clarithromycin, with the lowest incidence of diarrhea documented as a side effect of azithromycin treatment.68 Treatment of rhodococcal diarrhea is generally supportive. Affected foals should be treated with a macrolide in combination with rifampin. Although there are no reports of the efficacy of different macrolides in treating the intestinal form of disease, azithromycin or clarithromycin may be superior over erythromycin due to the lower incidence of adverse effects and higher success rates associated with their
use in foals with pneumonia.68 Foals which develop diarrhea as a result of macrolide therapy often respond to cessation of treatment for 24–48 hours, although in mild cases treatment may be continued along with supportive therapy for the diarrhea. Rotavirus Rotavirus is a significant cause of diarrhea in foals. In epidemiological studies of foals with infectious diarrhea, rotavirus was isolated from up to 53% of foals,1, 2, 69 with more than 90% of outbreaks attributable to rotavirus in one study.69 Rotavirus group A is a common cause of diarrhea in foals from 1 week to 3 months of age, but has been reported in up to 5-month-old foals.3 The peak incidence of infection occurred in foals 1–2 months of age in one study,2 while in another, the average age of affected foals was 77 days.69 Shedding has been documented in 2–8% of normal foals.1, 2 The virus has also been isolated from healthy foals in contact with diarrheic foals.1 Most adult horses have positive titers indicating exposure, but do not shed the virus in feces. Rotavirus is highly contagious and has a short incubation period, with several foals typically affected. Transmission is via the feco-oral route. The virus affects microvilli of the jejunum and ileum, causing secretory and osmotic diarrhea and malabsorption. As intestinal crypt cells are unaffected, they continue to replicate and replace the damaged villous epithelial cells so most cases are generally self-limiting. Foals stop nursing, are depressed, and develop diarrhea within 12–24 hours post-infection. Diarrhea may be watery or pasty in nature but is typically non-hemorrhagic with a characteristic odor. Fever is an inconsistent finding. Disease severity is dependent on immune status, inoculation dose, and age of the foal.4 Younger foals are more severely affected, and are more likely to be severely dehydrated with marked electrolyte derangements. Foals older than 3 months may be protected by active immunity and maturation of intestinal function.2 Diarrhea usually lasts for 2–3 days but may be more protracted in some cases (10–12 days).69 Recurrent infection and chronic diarrhea are uncommon. Foals may continue shedding rotavirus for an average of 3 days after feces have normalized. The disease is associated with a high morbidity but low mortality rate with appropriate treatment. A number of diagnostic tests are available such as virus isolation, latex particle agglutination, enzyme immunoassay, and polyacrylamide electrophoresis.3 Fecal antigen tests are sensitive and specific and provide rapid results. Treatment is generally supportive with antibiotics generally not indicated except in foals less than 2
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Diarrhea weeks.4 Fluid and electrolyte deficits need to be corrected, and administration of gastroprotectants may be beneficial as an association between rotavirus and ulceration has been suggested.70 Lactose intolerance has been reported, so affected foals may benefit from lactase supplementation during treatment and the recovery phase. Vaccination of pregnant mares (see Chapter 31) is suggested to be efficacious as a preventative measure.71 Mares are vaccinated during the eighth, nine, and tenth month of gestation, with a single booster given a month prior to foaling in subsequent years. Vaccination has been shown to reduce the incidence and severity of disease. Rotavirus is very stable and can survive in the environment for up to 9 months, including freezing up to 3 months.72 Effective disinfectants are phenolic compounds, 10% povidone iodine, quaternary ammonium disinfectants, and peroxygen compounds. Bleach and chlorhexidine are not effective. As for other infectious causes of diarrhea, isolation of foals with diarrhea is important, and all in contact foals should be considered exposed and potentially incubating the disease.
Protozoal diarrhea
Cryptosporidium The role of Cryptosporidium parvum in foal diarrhea has been controversial. These coccidial parasites invade the distal small intestine, especially the ileum, causing enteritis characterized by villous atrophy and inflammation.73 Oocysts have been isolated from the feces of both normal foals and foals with diarrhea in the same frequency.2 In another epidemiological survey of foals with diarrhea, Cryptosporidium was again isolated from both normal and diarrheic foals but was significantly associated with diarrhea in this population.1 Fecal shedding of oocysts is intermittent and age dependent, with infection rates of 15–31% reported in suckling foals but in less than 3% of weanlings.74 Age less than 6 months and a history of diarrhea are established risk factors for fecal oocyst shedding.75 Immunosuppression is also important, with cryptosporidiosis often reported in immunodeficient foals or foals compromised by concurrent diseases. Arabian foals with severe combined immunodeficiency typically present at 1–3 months of age with pneumonia, weight loss, and cryptosporidial diarrhea.76, 77 Less frequently, cryptosporidiosis has been reported as a cause of mild diarrhea in immunocompetent foals up to 8 weeks of age,78, 79 and has been implicated in an outbreak of diarrhea in 4- to 21-day-old foals.73 Chronic intermittent diarrhea has been reported in older foals.79
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The incubation period is 3–7 days. Cryptosporidium oocysts are sporulated and infective when excreted. Oocysts are difficult to identify on routine fecal flotation due to their small size. Diagnosis is based on detection of oocysts in feces via acid-fast staining (modified Ziehl–Neelsen), immunofluorescence assays, and flow cytometry.4 Most cases are self-limiting; however cryptosporidiosis has been associated with fatal severe watery diarrhea.1 Treatment is generally supportive, as there is no known specific therapy. Human patients with cryptosporidium have been treated with azithromycin, clarithromycin, nitazoxanide, and the amioglycoside paromomycin. These drugs have been suggested for treatment of affected foals but efficacy data are lacking.4 Biosecurity is essential due to the nosocomial and zoonotic potential of Cryptosporidium. Oocysts are highly resistant to most disinfectants, so disposal of fecal material and contaminated bedding is important. Steam, undiluted bleach, 5% ammonia, and formalin may be effective but require prolonged exposure. Peroxygen compounds may also be effective. Giardia and Eimeria leukarti The role of Giardia in foal diarrhea is unclear. The organism has been isolated from feces of normal foals and foals with diarrhea of all age groups, with an infection rate of 17–35% reported in foals.74 Excretion of cysts was documented in foals between 2 and 22 weeks of age, and in weanlings. Eimeria leukarti has been isolated from feces of normal foals and foals with diarrhea, with oocysts detected in up to 41% of asymptomatic foals between 29 and 191 days old.80 Intestinal coccidiosis has been infrequently reported in foals 45 days–6 months of age.81 However, its role as a primary enteric pathogen is unclear, with other causative organisms such as Salmonella spp81 and Lawsonia intracellularis82 often isolated concurrently. Parasitism (see also Chapter 30) Parasites should be considered as a cause of diarrhea in foals older than 2 weeks, with larval cyathostominosis an important differential for diarrhea in older foals (>2 months of age), and yearlings. Young horses (6 months-2 years) are most likely to have high tapeworm burdens.83 Colic is the most common clinical sign.84, 85 Ascarids are common in foals less than 10 months; however illthrift or colic is the most common clinical sign.
Strongyloidosis Strongyloides westeri infrequently produces clinical disease. S. westeri was associated with foal diarrhea
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in an epidemiological study but only when present in large numbers (> 2000 eggs/g).1 Infection is common in younger foals (1 week–2 months), although regular deworming of the postpartum mare with ivermectin (or oxibendazole) has reduced the incidence of disease. Fecal flotation is used to diagnose infection.
Strongyles and cyathostominosis Recently weaned 3- to 6-month-old foals grazing pasture contaminated with large and small strongyles have been reported to develop diarrhea.86 Diarrhea is more commonly seen with chronic disease than acute strongylosis in naive horses.17 Clinical disease associated with large strongyles occurs infrequently, in comparison cyathostominosis is the most common cause of chronic diarrhea.87 Other clinical signs of chronic disease include weight loss, and intermittent diarrhea. The acute form of the disease, larval cyathostomiasis, is associated with synchronous emergence of larvae from the intestine. Diarrhea is the most common clinical sign;88 however, weight loss and edema have been reported in horses without diarrhea.89 The disease may affect horses of all ages, with young horses (1–3 years) affected more often.90 Risk factors associated with cyathostomiasis include age, season (late winter, spring in temperate climates) and time since last anthelmintic administration, with anthelmintic treatment within the previous 2 weeks a risk factor for the development of clinical signs.91 A negative fecal egg count does not rule out parasites as a cause of diarrhea as many parasites cause clinical disease during the prepatent period.3 Diagnosis is usually presumptive based on history and clinical signs, with larvae, adults, or eggs seen infrequently on fecal examination during clinical disease. Definitive diagnosis of cyathostominosis requires postmortem examination or examination of large intestinal biopsies. Rectal mucosal biopsies may be useful in some cases. Fecal excretion of larvae and adults has been associated with diarrhea, hypoproteinemia, and hypoalbuminemia.92 Moxidectin (400 µg/kg p.o.) and fenbendazole (10 mg/kg p.o. s.i.d. 5 days) are effective treatments. Supportive care including intravenous fluid therapy, colloids, and anti-inflammatories (oral prednisolone) is often indicated.93 Other infectious causes Potomac horse fever (equine monocytic ehrlichiosis) is a disorder characterized by acute enterocolitis, due to infection with Neorickettsia risticii. Clinical signs include anorexia, depression, and fever. Diarrhea is common but may not be evident in all horses. Potomac horse fever can affect any age group of horses, although infection is rare in horses less than 1 year of
age. A vaccine is available which does not completely prevent the disease but may lessen its severity (see Chapter 31). Aeromonas hydrophila is more frequently recovered from the feces of foals with diarrhea than from normal foals, although its role as a primary pathogen is not clear.2 Bacteroides fragilis has been associated with diarrhea in foals, but is also isolated from the feces of normal foals.4 Coronavirus has been isolated at a low prevalence from foals with diarrhea,2 and has been associated with colitis in immunocompromised foals,77 but only rarely as a primary pathogen in immunocompetent neonates.4 Adenovirus has been isolated from foals with concurrent rotavirus infection.94 Recently, the Gram-negative spirochaete Brachyspira pilosicoli has been isolated from weanlings 8–10 months of age with chronic diarrhea.95
Non-infectious causes Gastroduodenal ulceration (see also Chapters 41 and 74) Equine gastric ulcer syndrome (EGUS) may affect horses of any age, with distinct clinical syndromes identified in neonatal foals, suckling/weanling foals, and yearlings/adult horses.96 The pathophysiology of EGUS has been described in detail,96, 97 and is similar in all age groups; however, inciting causes and clinical signs are often different.98 Typical clinical signs in yearlings include anorexia, colic, and loss of condition while, diarrhea is a common sign in suckling foals and weanlings. The prevalence of EGUS in foals, including neonates, has been reported as 25–57%.99, 100 In an early endoscopic survey of asymptomatic foals between 2 and 85 days, ulcers were more prevalent in foals less than 10 days and least prevalent in foals older than 70 days.100 In another study of foals that underwent gastroscopy at a referral hospital, 56% of foals less than 85 days had lesions compared to 41% of foals between 90 and 310 days.99 Desquamation of squamous epithelium at the margo plicatus without ulceration is common in foals less than 30 days, but is not typically associated with clinical signs.99–101 Gastric ulcers in foals less than 50 days of age are most frequently located in the squamous mucosa adjacent to the margo plicatus along the greater curvature, whereas lesions are more prevalent along the lesser curvature in older foals.100, 101 Diarrhea is the most common clinical sign in symptomatic foals with squamous mucosal lesions.96 Other signs include poor appetite, ill-thrift, poor growth rates, pot-bellied appearance, bruxism, and colic.
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Diarrhea Concurrent clinical disease, such as diarrhea is a common predisposing factor for the development of gastric ulcers in foals.100 Squamous ulcers have been associated with pre-existing clinical disorders, especially in foals more than 90 days of age.99 Ulcers involving the glandular mucosa are reported less frequently,99 and may also be associated with administration of non-steroidal anti-inflammatory agents (NSAIDs).98, 102 Duodenal ulcers are less common than gastric ulcers, and can affect foals of any age. Gastroduodenal ulcer disease is a syndrome affecting suckling foals and weanlings in particular, with pyloric and duodenal ulcers most common in 3- to 5-month-old foals.103 The exact cause of duodenal ulceration is unknown, although a similar pathophysiology as gastric ulcers is proposed. Gastroduodenal ulceration has also been associated with NSAID use.102 Outbreaks have been reported, although an association with an infectious agent has not been established.104, 105 Signs are usually identical to those associated with gastric ulcers, except delayed gastric emptying and gastroesophageal reflux may occur. Complications include gastric or duodenal rupture, peritonitis, duodenal stricture, gastric outflow obstruction, ascending cholangitis, and aspiration pneumonia.102,105–107 The diagnosis of gastric ulceration may be suspected on the basis of clinical signs and response to treatment; however, gastroscopy is required to definitively diagnose the condition (Figure 81.2). In foals with duodenal ulceration, contrast radiography may be used to document delayed gastric emptying. A number of agents are available for treatment of gastric ulcers, and their mechanism of action has been described in detail elsewhere96, 108 (see Chapters 41 and 74). The main components of most treatment regimes are the use of H2 antagonists, proton pump blockers, and sucralfate. The H2 antagonists suppress hydrochloric acid secretion, and have been shown to be efficacious.109 Recommended dosages are 20–30 mg/kg p.o. q 8 h for cimetidine and 6.6 mg/kg p.o. q 8 h for ranitidine, with a minimum of 2–3 weeks duration of treatment.96 Proton pump inhibitors such as omeprazole (4 mg/kg p.o. q 24 h for 28 days) block hydrogen ion secretion, and have documented efficacy in foals and adults,110 although currently available products are not registered for use in foals less than 4 weeks of age. Sucralfate (20–40 mg/kg p.o. q 8 h) adheres to ulcerated mucosa, stimulates mucus secretion, and enhances prostaglandin synthesis, and is likely to be more effective in the treatment of glandular ulcers.96 The efficacy of sucralfate has been questioned, with one study showing no significant effect compared to treatment with corn syrup. 111 Antacids have a short-lived effect in horses, ne-
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Figure 81.2 Gastric ulceration in a 4-month-old foal with diarrhea, inappetance, and ill-thrift. Ulceration of the squamous mucosa (arrow) was evident above the margo plicatus (broken arrow).
cessitating administration at 2- to 4-hour intervals, and their efficacy has not been determined.108 Intravenous fluid therapy and nutritional support in the form of partial or total parenteral nutrition may be required. In severe cases of duodenal ulceration or stricture unresponsive to medical treatment, surgery is indicated.104, 105, 107 Foals with severe gastroduodenal ulceration have a guarded prognosis even with surgical therapy.96 Early diagnosis, early surgical intervention and postoperative management are important determinants of surgical success. Sand Sand accumulation in the gastrointestinal tract has been reported as a cause of chronic diarrhea, colic, and weight loss in horses of any age.112 The shorter transit time and smaller diameter of the large intestine in foals was thought to reduce the likelihood of sand accumulation in younger horses;113 however, the condition has been reported in 4-week-old,114 6week-old,115 and 3- to 4-month-old foals112, 116 and yearlings.117 Fifteen percent of horses in a recent retrospective study of sand colic impactions were less than a year of age.114 Sand settles ventrally in the colon causing chronic mucosal irritation and inflammation, malabsorption, and disruption of intestinal motility. Diarrhea, when it occurs, is usually intermittent and not severe.
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Figure 81.3 Lateral abdominal radiograph of a weanling with ill-thrift and chronic diarrhea. A large accumulation of sand is evident as a radiodense mass in the cranioventral abdomen (cranial is to the right).
Physical examination findings suggestive of sand enteropathy are characteristic sand sounds on auscultation of the cranioventral abdomen, and finding abnormal amounts of sand in feces. Blood tests are usually unremarkable,112, 116 although endotoxemia and peritonitis have been reported.112, 115 Abdominal radiography is a reliable diagnostic modality (Figure 81.3), and is useful for monitoring resolution and response to treatment.115 Ultrasonography of the ventral abdomen using a 5 MHz rectal transducer demonstrated similar findings when compared with radiographic findings in one study.118 Typical ultrasonographic findings include the colon lying flat against the ventral abdominal wall with non-moving sacculations and reduced motility.117 Suggested treatment regimes include the use of psyllium mucilloid, mineral oil, magnesium sulphate, and/or dioctyl sodium sulphosuccinate (DSS). Psyllium is a soluble dietary fiber administered via nasogastric tube or in feed (1 g/kg divided daily). Proposed mechanisms of action include dislodgement of sand, hydrating and lubricating effects, stimulation of intestinal motility,112 and promotion of colonocyte proliferation and facilitation of mucosal healing.33 Although the efficacy of psyllium has been debated, resolution of diarrhea within a few days and radiographic clearance of sand has been reported.112, 115, 116 However, there was no apparent effect in another study.119 Other non-infectious disorders Other disorders are common to horses of all age groups and are not specific to foals and yearlings. Abrupt dietary alterations and carbohydrate overload can alter enteric bacterial flora, and prolonged anorexia (> 72 hours) may lead to villous atrophy
and diarrhea.35 Infiltrative bowel disease is uncommon in young horses and is discussed in more detail in Chapter 83 on weight loss. NSAIDs should be used with caution in foals and young horses, as this age group is particularly sensitive to the development of toxic side effects.120 Adverse effects on the gastrointestinal tract occur secondary to inhibition of protective prostaglandins, resulting in decreased mucosal blood flow, disruption of the mucus–bicarbonate barrier and mucosal ulceration. Gastric ulceration and right dorsal colitis have been reported in horses overdosed with NSAIDs, as well as in horses which have received appropriate doses (particularly phenylbutazone) for short periods.121, 122 Anorexia, lethargy, colic, and diarrhea are signs of right dorsal colitis. Weight loss and soft feces may be a feature in chronic cases. Endotoxemia and petechial hemorrhages may also be detected. Clinicopathological abnormalities include hypoproteinemia, hypoalbuminemia, neutropenia and changes in volatile fatty acid production.123 Gastroscopy is recommended in suspect cases. Abdominal ultrasonography has demonstrated increased mural thickness and alterations in the appearance of the right dorsal colon in horses with right dorsal colitis.124 Flunixin meglumine appears to be less toxic than phenylbutazone, but both have been associated with glandular gastric ulceration, hypoproteinemia, and right dorsal colitis.100, 125 Chronic flunixin meglumine administration in foals was associated with oral and glandular gastric ulceration.126 Ketoprofen has been shown to have less gastrointestinal side effects than phenylbutazone.125 If NSAID toxicity is suspected, NSAID use should be discontinued and gastroprotectants administered. Pelleted feeds, psyllium, corn oil, sucralfate, and misoprostol have been used in the treatment of right dorsal colitis. Pelleted feeds and eliminating roughage from the diet are thought to minimize colonic load.33 The efficacy of sucralfate in the treatment of right dorsal colitis has not been determined. Misoprostol (2 µg/kg p.o. q 8 h) is a synthetic prostaglandin analog often used by practitioners in the treatment of right dorsal colitis due to its potential to enhance mucosal healing. Clinical trials demonstrating its efficacy are lacking. Recently, selective COX-2 inhibitors have been investigated for use in horses. Gastric ulcers were not detected when firocoxib was administered for 30 days; however, the safety of this product in horses less than a year old has not been determined. Pharmacokinetics of meloxicam, another specific COX-2 inhibitor, has recently been investigated in healthy foals 2–23 days of age.127 Single dose and shortterm administration (0.6 mg/kg p.o. q 12 h for 2 weeks) was not associated with the development of
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Diarrhea significant gastric ulceration. Further studies are warranted to investigate the safety of these products in clinically ill foals.
References 1. Netherwood T, Wood WL, Townsend HG, Mumford JA, Chanter N. Foal diarrhea between 1991 and 1994 in the United Kingdom associated with Clostridium perfringens, rotavirus, Strongyloides westeri and Cryptosporidium spp. Epidemiol Infect 1996;117(2):375–83. 2. Browning GF, Chalmers RM, Snodgrass DR, Batt RM, Hart CA, Ormarod SE, Leadon D, Stoneham SJ, Rossdale PD. The prevalence of enteric pathogens in diarrhoeic thoroughbred foals in Britain and Ireland. Equine Vet J 1991;23(6):405–9. 3. Dunkel B, Wilkins PA. Infectious foal diarrhea: pathophysiology, prevalence and diagnosis. Equine Vet Educ 2004;16(2):94–101. 4. Lester GD. Infectious diarrhea in foals. In Proceedings 47th annual meeting American Association Equine Practitioners, 2001; pp. 468–71. 5. B˚averud V, Gustafsson A, Franklin A, Lindholm A, Gunnarsson A. Clostridium difficile associated with acute colitis in mature horses treated with antibiotics. Equine Vet J 1997;29(4):279–84. 6. B˚averud V, Franklin A, Gunnarsson A, Gustafsson A, Hellander-Edman A. Clostridium difficile associated with acute colitis in mares when their foals are treated with erythromycin and rifampin for Rhodococcus equi pneumonia. Equine Vet J 1998;30:482–8. 7. B˚averud V, Gustafsson A, Franklin A, Aspan A, Gunnarsson A. Clostridium difficile: prevalence in horses and environment and antimicrobial susceptibility. Equine Vet J 2003;35(5):465–71. 8. Weese JS, Staempfli HR, Prescott JF. A prospective study of the roles of Clostridium difficile and enterotoxigenic Clostridium perfringens in equine diarrhea. Equine Vet J 2001;33(4):403–9. 9. Magdesian KG, Hirsh DC, Jang SS, Hansen LM, Madigan JE. Characterization of Clostridium difficile isolates from foals with diarrhea: 28 cases (1993–1997). J Am Vet Med Assoc 2002;220(1):67–73. 10. Ruby R, Magdesian KG, Hass PH. Comparison of clinical, microbiologic and clinicopathologic findings in horses positive and negative for Clostridium difficile infection. J Am Vet Med Assoc 2009;234:777–84. 11. Weese JS, Parsons DA, Staempfli HR. Association of Clostridium difficile with enterocolitis and lactose intolerance in a foal. J Am Vet Med Assoc 1999;214:229–32. 12. Jones RL. Clostridial enteritis. In: Timoney PJ (ed.) Emerging infectious diseases: Vet Clin N Am Equine 2000;16(3): 471–86. 13. Tillotson K, Traub-Dargatz JL, Dickinson CE, Ellis RP, Morley PS, Hyatt DR, Magnuson RJ, Riddle WT, Bolte D, Salman MD. Population-based study of fecal shedding of Clostridium perfringens in broodmares and foals. J Am Vet Med Assoc 2002;220:342–7. 14. East LM, Savage CJ, Traub-Dargatz JL, Dickinson CE, Ellis RP. Enterocolitis associated with Clostridium perfringens infection in neonatal foals: 54 cases (1988–1997). J Am Vet Med Assoc 1998;212:1751–6. 15. MacKay RJ. Equine neonatal clostridiosis: treatment and prevention. Compendium Cont Educ 2001;23:280–5.
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16. Donaldson MT, Palmer JE. Prevalence of Clostridium perfringens enterotoxin and Clostridium difficile toxin A in feces of horses with diarrhea and colic. J Am Vet Med Assoc 1999;215:358–61. 17. Jones SL. Inflammatory diseases of the gastrointestinal tract causing diarrhea. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis, MO: Saunders, 2004; pp. 884–912. 18. Seahorn JL, Seahorn TL. Fluid therapy in horses with gastrointestinal disease. In: Jones SL (ed.) Therapeutics for gastrointestinal disease: Vet Clin N Am Equine 2003;19(3):665–79. 19. Kelmer G. Update on treatments for endotoxemia. In: Andrews FM (ed.) New perspectives in equine colic: Vet Clin N Am Equine 2009;25:259–70. 20. Durando MM, MacKay RJ, Linda S, Skelley LA. Effects of polymixin B and Salmonella typhimurium antiserum on horses given endotoxin intravenously. Am J Vet Res 1994;55:921–7. 21. Barton MH, Parviainen A, Norton N. Polymyxin B protects horses against induced endotoxemia in vivo. Equine Vet J 2004;36:396–401. 22. Beard L. Managing foal diarrhea. Compend Equine 2009;4:16–26. 23. Jones SL. Treatment of acute and chronic gastrointestinal inflammation. In: Jones SL (ed.) Therapeutics for gastrointestinal disease: Vet Clin North Am Equine 2003; 19(3):697–714. 24. Barton MH, Moore JN, Norton N. Effects of pentoxifylline infusion on response of horses to in vivo challenge exposure with endotoxin. Am J Vet Res 1997;58:1300–7. 25. Baskett A, Barton MH, Norton N, Anders B, Moore JN. Effect of pentoxifylline, flunixin meglumine and their combination on a model of endotoxemia in the horse. Am J Vet Res 1997;58:1291–9. 26. Feary DJ, Hassel DM. Enteritis and colitis in horses. In: Southwood LL (ed.) Advances in diagnosis and management of infection: Vet Clin N Am Equine 2006;22:437–79. 27. Weese JS, Cole NW, deGannes RV. Evaluation of in vitro properties of di-tri-octahedral smectite on clostridial toxins and growth. Equine Vet J 2003;35(7):638–41. 28. Lawler JB, Hassel DM, Magnuson RJ, Hill AE, McCue PM, Traub-Dargatz JL. Adsorptive effects of di-tri-octahedral smectite on Clostridium perfringens alpha, beta and beta2 exotoxins and equine colostral antibodies. Am J Vet Res 2008;69:233–9. 29. Hassel DM, Smith PA, Nieto JE, Beldomenico P, Spier SJ. Di-tri-octahedral smectite for the prevention of postoperative diarrhea in equids with surgical disease of the large intestine: results of a randomized clinical trial. Vet J 2009;182:210–14. 30. Tillotson K, Traub-Dargatz JL. Gastrointestinal protectants and cathartics. In: Jones SL (ed.) Therapeutics for gastrointestinal disease: Vet Clin N Am Equine 19(3):599– 615. 31. Weese JS, Rousseau J. Evaluation of Lactobacillus pentosus WE7 for prevention of diarrhea in neonatal foals. J Am Vet Med Assoc 2005;226:2031–4. 32. Desrochers Am, Dolenta BA, Roy MF, Boston R, Carlisle S. Efficacy of Saccharomyces boulardii for treatment of horses with acute enterocolitis. J Am Vet Med Assoc 2005;227:954–9. 33. Bueno AC, Seahorn TL, Moore RM. Diagnosis and treatment of right dorsal colitis in horses. Comp Cont Educ Pract Vet 2000;22(2):173–81.
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34. B˚averud V. Clostridium difficile diarrhea: infection control in horses. In: Baom FT. Weese JS (eds) Infection Control: Vet Clin N Am Equine 2004;20:615–30. 35. Chapman AM. Acute diarrhea in hospitalized horses. In: Andrews FM (ed.) New perspectives in equine colic: Vet Clin N Am Equine 2009;25:363–80. 36. Murray MJ. Salmonellosis in horses. J Am Vet Med Assoc 1996;209:558–60. 37. Traub-Dargatz JL, Garber LP, Fedorka-Cray PJ, Ladely S, Ferris KE. Fecal shedding of Salmonella spp by horses in the United States in 1998 and 1999 and detection of Salmonella spp in grain and concentrate sources on equine operations. J Am Vet Med Assoc 2000;217:226–30. 38. Ernst NE, Hernandez JA, MacKay RJ, Brown MP, Gaskin JM, Nguyen AD, Giguere S, Colahan PT, Troedsson MR, Haines GR, Addison IR, Miller BJ. Risk factors associated with fecal Salmonella shedding among hospitalized horses with signs of gastrointestinal tract disease. J Am Vet Med Assoc 2004;225(2):275–81. 39. Schott HC 2nd , Ewart SL, Walker RD, Dwyer RM, Dietrich S, Eberhart SW, Kusey J, Stick JA, Derksen FJ. An outbreak of Salmonellosis among horses at a veterinary teaching hospital. J Am Vet Med Assoc 2001;218: 1152–9. 40. Ward MP, Brady TH, Couetil LL, Liljebjelke K, Maurer JJ, Wu CC. Investigation and control of an outbreak of salmonellosis caused by multidrug-resistant Salmonella typhimurium in a population of hospitalized horses. Vet Microbiol 2005;107:233–40. 41. Carter JD, Hird DW, Farver TB, Hjerpe CJ. Salmonellosis in hospitalized horses: seasonality and case fatality rates. J Am Vet Med Assoc 1986;188:163–7. 42. van Duijkeren E, Flemming C, Sloet van OldruitenborghOosterbann M, Kalsbeek HC, van der Geissen JW. Diagnosing Salmonellosis in horses: culturing of multiple versus single fecal samples. Vet Q 1995;17:63–6. 43. Cohen ND, Martin LJ, Simpson RB, Wallis DE, Neibergs HL. Comparison of polymerase chain reaction and microbiological culture for detection of Salmonellae in equine feces and environmental samples. Am J Vet Res 1996;57:780–6. 44. Palmer JE, Whitlock RH, Benson CE, Becht JL, Morriss DD, Acland HM. Comparison of rectal mucosal cultures and fecal cultures in detecting Salmonella infection in horses and cattle. Am J Vet Res 1988;46:697–8. 45. Kurowski PB, Traub-Dargatz JL, Morley PS, GentryWeeks CR. Detection of Salmonella spp in fecal specimens by use of real-time polymerase chain reaction assay. J Am Vet Med Assoc 2002;63(9):1265–8. 46. Johns IC, Jesty SA, James FM. Pseudomonas aeruginosa sepsis in an adult horse with enteric Salmonellosis. J Vet Emerg Crit Care 2006;16:219–23. 47. Johns I, Tennent-Brown B, Dallap Schaer B, Southwood L, Boston R, Wilkins P. Blood culture status in mature horses with diarrhea: a possible association with survival. Equine Vet J 2009;41:160–4. 48. Papich MG. Antimicrobial therapy for gastrointestinal diseases. In: Jones SL (ed.) Therapeutics for gastrointestinal disease: Vet Clin North Am Equine 2003;19(3):645–63. 49. Parraga ME, Spier SJ, Thurmond M, Hirh D. A clinical trial of probiotic administration for prevention of Salmonella shedding in the postoperative period in horses with colic. J Vet Intern Med 1997;11:36–41. 50. Kim LM, Morley PS, Traub-Dargatz JL, Salman MD, Gentry-Weeks C. Factors associated with Salmonella
51.
52.
53.
54.
55.
56.
57. 58.
59.
60. 61.
62.
63.
64.
65.
66.
67.
68.
shedding among equine colic patients at a veterinary teaching hospital. J Am Vet Med Assoc 2001;218:740–8. Dolente BA, Beech J, Lindborg S, Smith G. Evaluation of rsk factors for development of catheter-associated jugular thrombophlebitis in horses: 50 cases (1993–1998). J Am Vet Med Assoc 2005;227:1134–41. Begg AP, Johnston KG, Hutchins DR, Edwards DJ. Some aspects of the epidemiology of equine salmonellosis. Aust Vet J 1988;65:221–3. Steel CM, Hunt AR, Adams PL, Robertson ID, Chicken C, Yovich JV, Stick JA. Factors associated with prognosis for survival and athletic use in foals with septic arthritis: 93 cases (1987–1994). J Am Vet Med Assoc 1999;215:937–9. Dwyer RM. Environmental disinfection to control equine infectious diseases. In: Bain FT, Weese JS (eds) Infection Control: Vet Clin North Am Equine 2004;20:531–42. Lavoie JP, Drolet R, Parsons D, Leguillette R, Sauvageau R, Shapiro J, Houle L, Halle G, Gebhart CJ. Equine proliferative enteropathy: a cause of weight loss, colic, diarrhoea and hypoproteinemia in foals on three breeding farms in Canada. Equine Vet J 2000;32(5):418–25. Feary DJ, Gebhart CJ, Pusterla N. Lawsonia intracellularis proliferative enteropathy in a foal. Schweiz Arch Tierheilk 2007;149(3):129–33. Frazer ML. Lawsonia intracellularis infection in horses: 2005–2007. J Vet Intern Med 2008;22(5):1243–48. McClintock SA, Collins AM. Lawsonia intracellularis proliferative enteropathy in a weanling foal in Australia. Aust Vet J 2004;82(12):750–2. Sampieri F, Hinchcliff KW, Toribio RE. Tetracycline therapy of Lawsonia intracellularis enteropathy in foals. Equine Vet J 2006;38(1):89–92. Bihr TP. Protein-losing enteropathy caused by Lawsonia intracellularis in a weanling foal. Can Vet J 2003;44:65–6. Dauvillier J, Picandet V, Harel J, Gottschalk M, Desrosiers R, Jean D, Lavoie JP. Diagnostic and epidemiological features of Lawsonia intracellularis enteropathy in 2 foals. Can Vet J 2006;47:689–91. Allen KJ, Pearson GR, Fews D, McOrist S, Brazil TJ. Lawsonia intracellularis proliferative enteropathy in a weanling foal, with a tentative histological diagnosis of lymphocytic plasmacytic enteritis. Eq Vet Educ 2009;21(8):411–14. Wong DM, Alcott CJ, Sponseller BA, Young JL, Sponseller BT. Impaired intestinal absorption of glucose in 4 foals with Lawsonia intracellularis infection. J Vet Intern Med 2009;23(4):940–4. Giguere S, Prescott JF. Clinical manifestations, diagnosis, treatment, and prevention of Rhodococcus equi infections in foals. Vet Microbiol 1997;56:313–34. Reuss SM, Chaffin MK, Cohen ND. Extrapulmonary disorders associated with Rhodococcus equi infection in foals: a retrospective study of 150 cases (1987–2007). Proc Am Assoc Eq Pract 2008;54:528. Giguere S. Rhodococcus equi infections. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. St Louis, MO: Saunders, 2003; pp. 60–3. Stratton-Phelps M, Wilson WD, Gardner IA. Risk of adverse effects in pneumonic foals treated with erythromycin versus other antibiotics: 143 cases (1986–1996). J Am Vet Med Assoc 2000;217(1):68–72. Giguere S, Jacks S, Roberts GD, Hernandez J, Long MT, Ellis C. Retrospective comparison of azithromycin, clarithromycin and erythromycin for the treatment of foals with Rhodococcus equi pneumonia. J Vet Intern Med 2004;18:568–73.
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Diarrhea 69. Dwyer RM, Powell DG, Roberts W, Donahue M, Lyons ET, Osborne M, Woode G. A study of the etiology and control of infectious diarrhea among foals in central Kentucky. Proc Am Assoc Equine Pract 1990;36:337–55. 70. Lester GD. Foal diarrhea. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. St Louis, MO: Saunders, 2003; pp. 677–9. 71. Powell DG, Dwyer RM, Traub-Dargatz JL, Fulker RH, Whalen JW Jr, Srinivasappa J, Acree WM, Chu HJ. Field study of the safety, immunogenicity, and efficacy of an inactivated equine rotavirus vaccine. J Am Vet Med Assoc 1997;211(12):193–8. 72. Dwyer RM. Rotaviral diarrhea. Vet Clin N Am Equine Pract 1993;9:311–19. 73. Grinberg A, Oliver L, Learmonth JJ, Leyland M, Roe W, Pomroy WE. Identification of Cryptosporidium parvum ‘cattle’ genotype from a severe outbreak of neonatal foal diarrhea. Vet Rec 2003;153:628–31. 74. Xiao L, Herd RP. Epidemiology of equine Cryptosporidium and Giardia infections. Equine Vet J 1994;26:14–17. 75. Cole DJ, Cohen ND, Snowden K, Smith R. Prevalence of and risk factors for fecal shedding of Cryptosporidium parvum oocysts in horses. J Am Vet Med Assoc 1998;213(9):1296–302. 76. Gibson JA, Hill MW, Huber MJ. Cryptosporidiosis in Arabian foals with severe combined immunodeficiency. Aust Vet J 1983;60(12):378–9. 77. Mair TS, Taylor FG, Harbour DA, Pearson GR. Concurrent cryptosporidium and coronavirus infections in an Arabian foal with combined immunodeficiency syndrome. Vet Rec 1990;126(6):127–30. 78. Gajadhar AA, Caron JP, Allen JR. Cryptosporidiosis in two foals. Can Vet J 1985;26(4):132–4. 79. Xiao L, Herd RP. Review of equine Cryptosporidium infection. Equine Vet J 1994;26:9–13. 80. Lyons ET, Tolliver SC. Prevalence of parasite eggs (Strongyloides westeri, Parascaris equorum, and strongyles) and oocysts (Eimeria leuckarti) in the feces of Thoroughbred foals on 14 farms in central Kentucky in 2003. Parasitol Res 2004;92:400–4. 81. Reppas GP, Collins GH. Eimeria leuckarti infections in three foals. Aust Vet J 1995;72(2):63–4. 82. Schumacher J, Schumacher J, Rolsma M, Brock KV, Gebhart CJ. Surgical and medical treatment of an Arabian filly with proliferative enteropathy caused by Lawsonia intracellularis. J Vet Intern Med 2000;14:630–2. 83. Proudman CJ, Holmes MA, Sheoran AS, Edwards SE. Trees AJ. Immunoepidemiology of the equine tapeworm Anoplocephala perfoliata: age-intensity profile and agedependency of antibody subtype responses. Parasitology 1997;114:89–94. 84. Proudman CJ, Trees AJ. Tapeworms as a cause of intestinal disease in horses. Parasitol Today 1999;15:156–9. 85. Reinemeyer CR, Nielsen MK. Parasitism and colic. In: Andrews FM (ed.) New perspectives in equine colic: Vet Clin N Am Equine 2009;25:233–45. 86. Thamsborg SM, Leifsson PS, Grondahl C, Larsen M, Nansen P. Impact of mixed strongyle infection in foals after one month on pasture. Equine Vet J 1998;30:240–5. 87. Love S, Mair TS, Hillyer MH. Chronic diarrhea in adult horses: a review of 51 referred cases. Vet Rec 1992;130:217–19. 88. Peregrine AS, McEwen B, Bienzie D, Koch TG, Weese JS. Larval cyathostominosis in horses in Ontario: an emerging disease? Can Vet J 2006;47:80–2.
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89. Mair TS. Outbreak of larval cyathostomiasis among a group of yearlings and two year old horses. Vet Rec 1994;135:598–600. 90. Lyons ET, Drudge JH, Tolliver SC. Larval cyathostomiasis. In: Timoney PJ (ed.) Emerging Infectious Diseases: Vet Clin N Am Equine 2000;16(3):501–13. 91. Reid SW, Mair TS, Hillyer MH, Love S. Epidemiological risk factors associated with a diagnosis of clinical cyathostomiasis in the horse. Equine Vet J 1995;27:127–30. 92. Smets K, Saw DJ, Deprez P, Vercruysse J. Diagnosis of larval cyathostomiasis in horses in Belgium. Vet Rec 1999;144:665–8. 93. Love S. Treatment and prevention of intestinal parasite-associated disease. In: Jones SL (ed.) Therapeutics for gastrointestinal disease: Vet Clin N Am Equine 2003;19(3):791–806. 94. Corrier DE, Montgomery D, Scuthfield WL. Adenovirus in the intestinal epithelium of a foal with prolonged diarrhea. Vet Pathol 1982;19(5):564–7. 95. Hampson DJ, Lester GD, Phillips ND, La T. Isolation of Brachyspira pilosicoli from weanling horses with chronic diarrhea. Vet Rec 2006;158:661–2. 96. Sanchez LC. Diseases of the stomach. In: Reed SM, Bayly WM, Sello DC (eds) Equine Internal Medicine, 2nd edn, St Louis, MO: Saunders, 2004; pp. 863–73. 97. Murray MJ. Pathophysiology of peptic disorders in foals and horses: a review. Equine Vet J Suppl 1999;29:14–19. 98. Andrews FM, Nadeau JA. Clinical syndromes of gastric ulceration in foals and mature horses. Equine Vet J Suppl 1999;29:30–3. 99. Murray MJ. Endoscopic appearance of gastric lesions in foals: 94 cases (1987–1988). J Am Vet Med Assoc 1989;195(8):1135–41. 100. Murray MJ, Murray CM, Sweeney HJ, Weld J, Digby NJ, Stoneham SJ. Prevalence of gastric lesions in foals without signs of gastric disease: an endoscopic survey. Equine Vet J 1990;22(1):6–8. 101. Murray MJ, Grodinsky C, Cowles RR, Hawkins WL, Forfa RJ, Luba NK. Endoscopic evaluation of changes in gastric lesions of Thoroughbred foals. J Am Vet Med Assoc 1990;196(10):1623–7. 102. Becht JL, Byars TD. Gastroduodenal ulceration in foals. Equine Vet J 1986;18(4):307–12. 103. Nappert G, Vrins A, Larybyere M. Gastroduodenal ulceration in foals. Comp Cont Educ Pract Vet 1989;11: 338–45. 104. Campbell-Thompson ML, Brown MP, Slone DE, Merritt AM, Moll HD, Levy M. Gastroenterostomy for treatment of gastroduodenal ulcer disease in 14 foals. J Am Vet Med Assoc 1986;188(8):840–4. 105. Orsini JA, Donawick WJ. Surgical treatment of gastroduodenal ulceration in foals. Vet Surg 1986;15:205–8. 106. Acland HM, Gunson DE, Gilette DM. Ulcerative duodenitis in foals. Vet Pathol 1983;20(6):653–61. 107. Barr BS. Duodenal stricture in a foal. In: MacLeay JM (ed.) Medical case management: Vet Clin N Am Equine 2006;22:37–42. 108. MacAllister CG. A review of medical treatment for peptic ulcer disease. Equine Vet J Supp 1999;29:45–9. 109. Sanchez LC, Lester GD, Merritt AM. Effect of ranitidine on intragastric pH in clinically normal neonatal foals. J Am Vet Med Assoc 1998;212:1407–12. 110. Murray MJ, Eichorn ES, Holste JE, Cox JL, Stanier WB, Cooper WL, Cooper VA. Safety, acceptability and endoscopic findings in foals and yearling horses treated with a
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Foal: Problems of Older Foals, Weanlings and Yearlings paste formulation of omeprazole for 28 days. Equine Vet J Suppl 1999;29:67–70. Borne AT, MacAllister CG. Effect of sucralfate on healing of subclinical gastric ulcers in foals. J Am Vet Med Assoc 1993;202(9):1465–8. Bertone JJ, Traub-Dargatz JL, Wrigley RW, Bennett DG, Williams RJ. Diarrhea associated with sand in the gastrointestinal tract of horses. J Am Vet Med Assoc 1988;193(11):1409–12. Specht TE, Colahan PT. Surgical treatment of sand colic in equids: 48 cases (1978–1985). J Am Vet Med Assoc 1988;193:1560–4. Granot N, Milgram J, Bdolah-Abram T, Shemesh I, Steinman A. Surgical management of sand colic impactions in horses: a retrospective study of 41 cases. Aust Vet J 2008;86:404–7. Ruohoniemi M, Kaikkonen R, Raekallio, Luukkanen L. Abdominal radiography in monitoring the resolution of sand accumulations from the large colon of horses treated medically. Equine Vet J 2001;33(1):59–64. Ramey DW, Reinertson EL. Sand induced diarrhea in a foal. J Am Vet Med Assoc 1984;185(5):537–8. Korolainen R, Kaikkonen R, Ruohoniemi M. Ultrasonography in monitoring the resolution of intestinal sand accumulations in the horse. Equine Vet Educ 2003;15(6):331–6. Korolainen R, Ruohoniemi M. Reliability of ultrasonography compared to radiography in revealing intestinal sand accumulations in horses. Equine Vet J 2002;34(5): 499–504.
119. Hammock PD, Freeman DE, Baker GJ. Failure of psyllium mucilloid to hasten evacuation of sand from the equine large intestine. Vet Surg 1998;27:547–54. 120. Traub JL, Gallina AM, Grant BD, Reed SM, Gavin PR, Paulsen LM. Phenylbutazone toxicosis in a foal. Am J Vet Res 1983;44:1410–18. 121. Cohen ND, Carter GK, Mealey RH, Taylor TS. Medical management of right dorsal colitis in 5 horses: a retrospective study (1987–1993). J Vet Intern Med 1995;9:272–6. 122. Hough ME, Steel CM, Bolton JR, Yovich JV. Ulceration and stricture of the right dorsal colon after phenylbutazone administration in four horses. Aust Vet J 1999; 77:785–8. 123. McConnico RS, Morgan TW, Williams CC, Hubert JD, Moore RM. Pathophysiologic effects of phenylbutazone on the right dorsal colon in horses. Am J Vet Res 2008; 69:1496–505. 124. Jones Sl, Davis J, Rowlingson K. Ultrasonographic findings in horses with right dorsal colitis: five cases (2000–2001). J Am Vet Med Assoc 2003;222:1248–51. 125. MacAllister CG, Morgan SJ, Borne AT, Pollet RA. Comparison of adverse effects of phenylbutazone, flunixin meglumine and ketoprofen in horses. J Am Vet Med Assoc 1993;202:71–7. 126. Traub-Dargatz JL, Bertone JJ, Gould DH, Wrigley RH, Weiser MG, Forney SD. Chronic flunixin meglumine therapy in foals. Am J Vet Res 1988;49:7–12. 127. Raidal SL, Pippia J, Noble G. Pharmacokinetics of single and multiple oral doses of meloxicam in foals less than 6 weeks of age. Aust Equine Vet 2008;27:52.
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Chapter 82
Colic Joanne Hardy
Introduction Older foals and weanlings presented for abdominal pain will often have conditions that are different than those seen in the adult, and the pain level is not necessarily reflective of the severity of disease, as young horses are often more demonstrative than adults. Conversely, with advancing intestinal devitalization, signs of abdominal pain may subside and foals may become lethargic. Conditions that cause colic in older foals and weanlings include severe enteritis, umbilical and inguinal hernias, torsions, large colon or small colon impactions and fecaliths, and small intestinal problems such as ascarid impaction, small intestinal intussusception, and gastric outflow obstruction. Evaluation of a foal with a painful abdomen can be a diagnostic challenge for the clinician as some of the diagnostic modalities applied in the adult, such as rectal palpation, cannot be applied to the foal. However, other techniques, such as abdominal radiographs and abdominal ultrasonography, are useful in the foal, but require a good understanding of normal anatomy and the ability to identify problems. The elements of diagnosis of colic in the older foal and weanling are similar to those in the adult and include history, physical examination, nasogastric intubation, and abdominocentesis. Additional diagnostic procedures include ultrasonography, abdominal radiographs, and clinicopathological data.
Initial assessment of the colicky foal History Age, signalment, and sex can help identify certain causes of colic in foals. A summary of the different causes of colic in foals of different age groups is listed in Table 82.1. Certain breeds are predisposed to certain types of colic. For example, female Quarter Horses are at
risk for umbilical hernia.1 Standardbreds, Tennessee Walkers, and Belgian draft breeds are at increased risk for inguinal hernias.2 Miniature foals are more commonly affected with small colon fecalith obstruction.3 Sex may also increase the risk for certain types of colic. For example, male foals are at risk for inguinal hernia and females are more commonly affected with umbilical hernia. A complete history should include information regarding the foal’s neonatal period, and whether any neonatal problems such as sepsis were encountered. Prior or current medication and any surgical procedure such as umbilical hernia repair, ruptured bladder repair, or exploratory celiotomy are important information. A history of previous sepsis may suggest abdominal or umbilical abscess, and abdominal surgery may have led to adhesion formation. Feeding and deworming practices should be obtained, and any concurrent or prior disease should be identified. For example, recent deworming in an older foal may suggest an ascarid impaction. Diarrhea before the colic episode may suggest an intussusception. Presence of pneumonia or farm disease prevalence may suggest abdominal abscesses from Rhodococcus equi. Physical examination
Evaluation of pain The severity of pain is important. Foals with a simple impaction or ulcers usually have mild intermittent pain that requires minimal medication. In contrast, foals with intussusceptions are often uncontrollably painful. It is important to remember that a strangulating obstruction will be extremely painful initially, but foals can become mostly depressed and in shock when the bowel is completely devitalized.4 The manifestation of pain is important. Foals with ulcer disease will often roll up on their backs, in a position of relief. Foals with small colon impaction often strain
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 82.1 Causes of colic in foals of different age groups Newborn foal
2–5 days old
Older foals
Meconium impaction
Ruptured bladder
Ascarid obstruction
Ileus/enteritis
Ileus/enteritis
Small intestinal volvulus
Ulcers
Ulcers
Small colon obstruction (fecalith)
Inguinal hernia with ruptured tunic
Atresia coli
Intussusception Enteritis Ulcers with/without gastric outflow obstruction
to defecate. Foals with small colon obstruction may stretch, strain, and flag their tail. The timing of pain is important. Pain immediately after nursing is consistent with proximal disease, as in gastric ulceration. Pain associated with a simple large colon obstruction will be more severe as gas distension progresses. Pain can be controlled with an α 2 -agonist such as xylazine (0.2–1 mg/kg intravenously (i.v.)), and may be combined with butorphanol (0.01–0.1 mg/kg i.v., intramuscularly (i.m.)). However, the need to repeat such medication is an indication of a severe and unresolved problem. Non-steroidal anti-inflammatory drugs (NSAIDs) such as flunixin meglumine are useful, but should be used with caution because of the risk of gastrointestinal ulceration. When using flunixin meglumine, a maximum dose of 1.1 mg/kg twice a day can be used. However, it is better to use the minimal effective dose, starting with 0.25 mg/kg two or three times a day.5 As an alternative ketoprofen (1.1–2.2 mg/kg i.v.) may be considered for its decreased toxicity.6 Other techniques can be performed to relieve the pain such as nasogastric intubation, or trocarization. It can sometimes be difficult to obtain reflux on a foal, and the use of a tube with lateral fenestrations may help. Trocarization should be performed only in cases of large colon distension, and if extreme abdominal distension is present with evidence of abdominal compartment syndrome. Trocarization should be performed in experienced hands because of the risk of peritonitis or bowel laceration. It is essential to determine whether the distension is caused by large colon distension (by radiographs or preferably ultrasonography) before trocarizing the foal. Sedation is recommended prior to this procedure to decrease the likelihood of bowel trauma.
Evaluation of the cardiovascular status Temperature, pulse, and respiration should be obtained. Normal temperature in foals is higher than
in the adult, and up to 102◦ F (38.8◦ C) is considered normal. The presence of a fever may suggest the presence of enteritis, however. Older foals and weanlings are considered to have the same temperature as adults. A febrile response may indicate an impending enterocolitis, although with advanced intestinal devitalization a fever may also be present; hypothermia may indicate advanced shock. Cardiovascular parameters are important and reflect the degree of cardiovascular compromise. Although these evaluations do not provide information as to the etiology of the colic, they are needed to address the urgency of the problem and to provide an assessment of prognosis. For this purpose, heart rate, mucous membrane color, and capillary refill time should be assessed. If evidence of compromise or shock is noted, immediate intravenous fluid replacement should be initiated. Examination of the gastrointestinal tract
Signs of gastrointestinal disease There are multiple signs of gastrointestinal disease in the foal. An overall examination should first be performed to rule out obvious causes of colic, such as inguinal herniation (non-reducible hernia or ruptured vaginal tunic), or strangulating umbilical hernia. Bruxism and rolling on their back is often seen with gastric ulceration. Severe hypersalivation and even spontaneous reflux is more often seen with duodenal ulcers. Regurgitation of feed material is possible in the foal. The presence of feed at the nostril indicates the need for nasogastric decompression. Abdominal distension can be caused by fluid or gas accumulation. Palpation and succussion can help differentiate the two. Typically, gas distension will manifest as tympany in the paralumbar fossa, whereas abdominal fluid will manifest as a pendulous abdomen (Figure 82.1). In foals, severe abdominal distension can be caused by small intestinal or large colon distension.
Figure 82.1 Illustration of a pendulous abdomen in a foal with uroperitoneum. This foal also has subcutaneous edema associated with subcutaneous accumulation of urine as a result of a ruptured urachus.
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Auscultation for the presence of borborygmi should be performed. Hypermotility is suggestive of enterocolitis, whereas hypomotility is more indicative of obstructive disease. Digital palpation is useful, particularly if sand impaction is suspected; sedimentation of collected fecal material within a rectal sleeve may reveal accumulation of sand. Severe abdominal distension, the absence of feces, and the palpation of mucus only are signs suggestive of complete small colon obstruction, such as in a fecalith obstruction.
Abdominocentesis Abdominocentesis should be performed with caution and possibly with sedation in young horses. The risk of bowel laceration is greater because of greater fractiousness during the procedure. For this reason, the author prefers to use a blunt cannula rather than a needle (although this increases the risk of omentum prolapse). Abdominocentesis should not be performed if it will not alter the decision-making process. Normal values in foals are reportedly lower than those of the adults, with total protein (TP) < 1.6 g/dL and white blood cell (WBC) count < 1500 cells/µL.7 In the face of a strangulating obstruction with progression of bowel necrosis, the expected changes are: increasing total protein within the first hour; increased TP and red blood cell (RBC) count within the first 3–4 hours, and increased WBC count starting at 5–6 hours. For unknown reasons, simple small colon obstructions are often associated with an inflammatory response manifested by an increase in both protein and WBC count. Comparison of peritoneal to plasma lactate can help in assessing the presence of intestinal compromise. An increased peritoneal fluid lactate compared with plasma concentration indicates intestinal compromise, and may be an earlier indicator of intestinal compromise than peritoneal fluid WBC concentration.
Figure 82.2 Ultrasonographic image of a weanling with small intestinal strangulation showing distended, amotile, thickened small intestine.
overall evaluation of the abdomen and intestinal tract. Examination of umbilical structures is best performed using a 7.5-MHz scan head. Ultrasonography can be used to determine the location of abdominal distension. Foals with enteritis will usually have a distended but motile loop of small intestine, whereas in small intestinal obstructions thickened, amotile loops are imaged (Figure 82.2). There is some overlap, however. In cases where intussusception is suspected, ultrasonography is useful to identify the intussuscepted loop (Figure 82.3). Ultrasonography is also very useful to identify abdominal effusion or accumulation of fluid in association with uroabdomen. Ultrasonography can be used to identify a location for abdominocentesis. In cases of uroabdomen, the tear in the bladder can often be imaged.
Abdominal radiographs Although ultrasonography has almost replaced radiographs in the examination of small bowel disease, radiographs remain more helpful for diagnosis of large colon obstructive disease, such as impaction
Nasogastric intubation Nasogastric intubation should be performed in all colicky foals, particularly if there is regurgitation of feed material, or gastric or small intestinal distension is seen on radiographs or ultrasonography. If there is reflux, oral fluids or oral medication should not be given, and appropriate parenteral rehydration, to account for the fluid loss in the reflux, should be instituted.
Abdominal ultrasonography Abdominal ultrasonography is now a routine diagnostic procedure in the evaluation of colic in young horses. A 3.5- or 5-MHz scan head can be used for
Figure 82.3 Transabdominal ultrasonographic image of a small intestinal intussusception in a foal (arrow).
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150min.
Figure 82.4 Contrast radiographic study of the proximal gastrointestinal tract in a foal presented for colic showing markedly delayed gastric emptying, indicating a gastric outflow obstruction. There is a nasogastric tube inserted in the stomach for administration of contrast material.
or sand colic. Contrast radiographs are useful for diagnosis of gastric outflow obstruction (Figure 82.4), or small colon obstruction. An upper gastrointestinal study can be performed after withholding food for 4–12 hours by administration of a barium sulfate suspension (5 mL/kg per os (p.o.)) by nasogastric tube.8 A lateral projection of the stomach is obtained at 1, 15, and 30 minutes, and then every hour until the barium has passed the area of interest. In normal foals the barium should start to leave the stomach within 30 minutes and should have left the stomach within 2 hours. Delayed gastric emptying can indicate pyloric outflow obstruction although some delayed gastric emptying can be present in sick foals owing to decreased gastrointestinal motility. A lower gastrointestinal study can be performed under sedation by administration of a 30% w/v barium sulfate enema by gravity flow (up to 20 mL/kg), using a 24-Fr Foley catheter with its bulb inflated, until the barium overflows around the catheter. Lateral and ventrodorsal radiographs are obtained immediately after barium administration. Clinicopathological evaluation A minimum laboratory database in the evaluation of the young horse with colic should include a complete blood count (CBC), evaluation of fibrinogen and lactate concentration, as well as a chemistry profile. The presence of anemia suggests chronicity whereas an increased fibrinogen would alert towards an inflammatory response such as abscessation. The WBC count and differential are useful to detect possible enteritis (leukopenia, high number of
bands); an increased WBC count may be present in response to stress or corticosteroid administration; however, a marked and persistent increase indicates an inflammatory, probably septic, response. The chemistry profile is useful for evaluation of hydration (creatinine), electrolyte and acid–base status, and metabolic status. The normal creatinine concentration of young foals is lower than that of the adult; adult values can be used in young horses older than 3 months of age. Lactate concentration can help determine and monitor perfusion. Electrolyte abnormalities can be significant with certain disease processes. Foals with chronic diarrhea can have hyponatremia, hypochloremia, and hypokalemia. Seizures can be observed with severe hyponatremia (Na < 110 mEq/L) or hypoglycemia. Increased liver enzymes can indicate primary liver disease, or indicate biliary outflow obstruction such as seen with duodenal stricture.
Common causes of colic in older foals and weanlings Enteritis Foals with severe enteritis can occasionally present for signs of colic, and abdominal distension as a result of severe ileus. In these foals, severe small and/or large intestinal distension is present. The challenge is to differentiate ileus from an acute abdominal obstruction. The presence of fever, leukopenia, and neutropenia can help differentiate these cases from a surgical abdomen. Occasionally, surgery is recommended in the absence of a clear-cut diagnosis. Clostridial diseases are the most common organism isolated from these cases, and management includes supportive care, antimicrobials, and intestinal rest by providing parenteral nutrition.
Ulcers Gastric/duodenal ulcer disease in foals is a controversial subject, as there is evidence that the mechanism of disease, at least in neonates, may be different than that of adults. In neonates, impaired gastric blood flow, as part of the shock/systemic inflammatory response syndrome (SIRS) present in this category of patients, may be the principal culprit whereas, in older foals, ulcers secondary to another illness may be more prevalent. In sick neonates, the pH of gastric juices has been shown to be alkaline, and HCl secretion is more intermittent and erratic. In addition, there is concern that alkalinization of the gastric juices removes a protective mechanism necessary in the defense of gastrointestinal (GI)
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Colic pathogens. Therefore, in neonates ulcer prevention and treatment is not universal. In older sick foals or foals that are receiving NSAIDs for pain management prevention and treatment of ulcers may be more indicated. Ketoprofen may be less ulcerogenic than flunixin meglumine or phenylbutazone, but its efficacy for pain management is also decreased. Signs of ulcer disease include colic after nursing, poor appetite, ill-thriftiness, bruxism, and hypersalivation. Foals with duodenal ulcer disease particularly show bruxism, hypersalivation, and poor doing. The treatment and prevention of ulcers is medical, using omeprazole, ranitidine, famotidine, or cimetidine. In contrast to adults, there is currently no evidence of efficacy for any of these drugs. Foals with duodenal ulcers may develop gastric outflow obstruction with or without bile duct obstruction. In these foals, surgical intervention with gastric bypass is indicated.9, 10 It is important to realize that foals with significant esophageal reflux from partial gastric outflow obstruction can develop significant esophagitis and subsequent esophageal stricture, which will worsen an already guarded prognosis. Congenital problems: umbilical and inguinal hernias
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hernial sac results in herniation of multiple loops of small intestine (Figure 82.6a,b). The treatment of complicated umbilical hernia is surgical with reduction of the hernia and, if indicated, resection of the segment of bowel. In the case of enterocutaneous fistulation, en bloc resection of the bowel and skin is indicated
Inguinal hernia Congenital inguinal hernias can present as an incidental finding, in which case they are reducible and do not cause signs of colic. Daily reduction of these hernias will lead to self-correction in approximately 80% of foals by 6 months of age. If self-correction does not occur, surgical management by castration, tunic reduction, or laparoscopy is indicated. Inguinal hernias can become strangulating, in which case they are irreducible, and cause signs of colic (Figure 82.7a,b). In certain instances, rupture of the tunic can occur, with migration of intestinal contents in the subcutaneous space (Figure 82.8a,b). These types of hernias are a surgical emergency.
Small colon obstruction
Umbilical hernias can be a cause of colic in the foal or weanling by strangulation of a segment of small intestine (Richter’s hernia) (Figure 82.5a,b) or cecum or large colon. In the last case, obstruction to intestinal flow is not impaired, and the strangulated segment of bowel can undergo necrosis without overt signs of abdominal discomfort, leading to an enterocutaneous fistula. Occasionally, rupture of the
Small colon obstruction is particularly prevalent in foals that are 1–3 months of age, particularly when they start to explore their environment. They are caused by dietary indiscretions, and can be made of hair, forage, shavings, or any other foreign material. Foals cannot chew roughage well so any highly fibrous forage can result in an obstruction. These foals will not be significantly systemically ill, but they can show significant signs of abdominal pain, distension, and straining to defecate. Miniature
(a)
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Complicated umbilical hernias
Figure 82.5 (a) External appearance of a small intestinal strangulation through an umbilical hernia in a foal. (b) Intraoperative illustration of small intestinal herniation through an umbilical hernia in the same foal.
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Figure 82.6 (a) Strangulating umbilical hernia with rupture of the hernial sac in a weanling. (b) Intraoperative illustration in the same foal showing herniation of multiple loops of small intestine and small colon through the umbilical defect.
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(b)
Figure 82.7 (a) External appearance of a horse with a left-sided inguinal hernia (arrow). (b) Intraoperative illustration of an inguinal hernia. Note that the testicle (arrow) is located within the vaginal tunic and that there is small intestine adjacent to it (arrowhead).
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Figure 82.8 (a) External appearance of a foal with an inguinal hernia with a ruptured tunic. Notice the large swelling extending beyond the end of the prepuce and migrating down the medial aspect of the leg. (b) Intraoperative illustration of the same foal showing the small intestine next to the testicle. In contrast to (b), the small intestine and testicle are not contained within the vaginal tunic.
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Colic
Figure 82.9 Intraoperative illustration of a small colon fecalith in a miniature foal (arrow).
foals are particularly predisposed with this type of condition.3 These foals usually can be managed medically, although the dramatic level of pain manifested may lead to surgical intervention. Fecaliths are usually removed by surgical intervention (Figure 82.9).
Small intestinal obstruction Small intestinal obstruction is more prevalent in the older foal (2–6 months of age). Conditions that can lead to small intestinal obstruction include small intestinal volvulus (Figure 82.10), intussusception (Figure 82.11), ascarid impaction (Figure 82.12), Meckel’s diverticulum (Figure 82.13), and mesodiverticular bands (Figure 82.14) to name a few. Signs include abdominal pain, small intestinal distension on radiographs and ultrasonography, and reflux. It is important to remember that in early strangulation foals demonstrate marked abdominal
Figure 82.10 Intraoperative illustration of the small intestine of a foal affected with a small intestinal volvulus (volvulus nodosus). This particular form of volvulus is found in foals 2–4 months of age and is localized at the junction of the jejunum and ileum.
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Figure 82.11 Foal presented for diarrhea that suffered an acute episode of colic. A jejunojejunal intussusception was identified at surgery (arrow).
pain, but, as intestinal necrosis progresses, foals become systemically compromised and the level of pain subsides. Abdominocentesis should be performed with caution because of the risk of enterocentesis, particularly if there is marked small intestinal distension. In the case of strangulation, abdominocentesis values will be consistent with intestinal necrosis, with increasing protein, lactate, and WBC count depending on duration of intestinal compromise. Ultrasonography of the abdomen can help identify the cause of the obstruction, such as ascarid impaction (Figure 82.15), intussusception, or strangulation. The treatment of small intestinal obstruction is surgical. An exception is ascarid impaction, in which medical therapy may be attempted. Ascarid impactions in foals are associated with a high parasite load, and, although recent deworming is often present in the history, it is not necessary for this condition to occur. The presence of adult ascarids in the intestinal tract is thought to result in hypomotility and contribute to the obstruction.11 Initial
Figure 82.12 Intraoperative illustration of a foal with an ascarid impaction and subsequent rupture of the distal jejunum.
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Figure 82.13 Intraoperative illustration of a weanling with Meckel’s diverticulum. Meckel’s diverticulum is a remnant of the embryonic vitelline duct.
presentation includes signs of simple small intestinal obstruction, including small intestinal distension visible on ultrasonography and normal abdominocentesis. With time, however, pressure necrosis at the site of impaction may occur; in addition, ascarid impaction may be associated with other conditions such as volvulus or intussusception, resulting in strangulating obstructions.10 Because the obstruction is most commonly located in the ileum, reflux may be delayed for several hours after the onset of signs of abdominal pain. Occasionally, the reflux will contain ascarids. Medical management can be attempted with intravenous fluid administration, gastric decompression, and use of parenteral NSAIDs such as flunixin meglumine or ketoprofen. However, if severe ileus, toxemia, or changes in the abdominal fluid are observed, surgical intervention is recommended. Surgical resolution is achieved by intraluminal
Figure 82.15 Transabdominal ultrasonography of a foal with colic showing obstruction of the small intestine by ascarids (arrow).
injection of a lubricant (the author prefers carboxymethylcellulose, as extravasation into the peritoneal cavity does not cause peritonitis) and manual massage, with or without an enterotomy. In foals with a suspected or known high ascarid load, deworming should be performed with caution and at a reduced dose to avoid massive killing. Low-dose fenbendazole (5 mg/kg) or pyrantel pamoate at one-third of the dose for 3 days can help prevent ascarid impaction in susceptible animals. The use of benzimidazoles in foals with heavy parasite burdens may be preferred as this class of drug has a slower mode of action.12 A full deworming regimen should follow within 1 week. Ascarid infestation can be prevented by careful management of pastures, decreasing overcrowding of susceptible animals, and effective deworming practices. There is rising concern of resistance to ivermectin and moxidectin; therefore, fenbendazole (10 mg/kg) can used if resistance is suspected.13 Foals with abdominal pain accompanied by weight loss, diarrhea, and hypoproteinemia should be suspected of proliferative enteropathy caused by Lawsonia intracellularis.14 Several foals may be affected on farms where the problem is endemic. Transabdominal ultrasonography will reveal increased small intestinal wall thickness. The diagnosis is based on polymerase chain reaction (PCR) analysis of serum or feces, and this condition is responsive to antibiotic therapy such as tetracycline (doxycycline) or macrolides. Miscellaneous conditions
Figure 82.14 Intraoperative illustration of a weanling with a mesodiverticular band (arrows). Mesodiverticular bands are remnant of the embryonic vitelline arteries. These can cause colic by chronic partial obstruction, or by entrapment of small intestine within the mesentery and subsequent strangulation.
Older foals and weanlings presented for colic can present with various conditions, such as abdominal abscessation, ovarian abscessation, ovarian torsion, and cecocolic intussusception.15, 16 In the case of abscessation, these conditions will usually cause
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from systemic illness such as sepsis, which worsens the prognosis. The reported survival in foals following abdominal surgery is 10% for foals < 14 days old and 45% in foals 15–150 days old, with a short-term survival of 63–65%.17
References
Figure 82.16 Intraoperative illustration of an abdominal abscess in a weanling. Intestinal adhesions are present. Aspiration of the abscess (arrow) resulted in a pure culture of Streptoccocus equi var. zooepidemicus.
chronic colic because of partial intestinal obstruction (Figure 82.16). Occasionally, an acute abdominal crisis will occur owing to acute strangulation. Cecocecal and cecocolic intussusception can present for acute or chronic colic depending on the degree of cecal invagination (Figure 82.17). These conditions are often associated with tapeworm infestation. Prognosis following abdominal surgery in foals and young horses Foals and young horses are at increased risk of developing adhesions following abdominal surgery for colic for several reasons: surgery is often delayed; lesions are often in the small intestine, with an associated worsened prognosis; the neonate’s bowel is much more fragile than the adult, therefore manipulations lead to more surgical trauma, mesenteric rent, and subsequent problems; and foals often suffer
Figure 82.17 Intraoperative illustration showing a cecocolic intussusception in a yearling. A right ventral colon enterotomy has been performed (black arrow) in order to exteriorize the invaginated ceum (arrowhead). Tapeworms were present within the cecum (white arrow).
1. Freeman DE, Spencer PA. Evaluation of age, breed, and gender as risk factors for umbilical hernia in horses of a hospital population. Am J Vet Res 1991;52:637–9. 2. Gaughan EM. Inguinal hernias in horses. Comp Cont Educ Pract Vet 1998;20:1057–9. 3. Haupt JL, McAndrews AG, Chaney KP, Labbe KA. Surgical treatment of colic in the miniature horse: a retrospective study of 57 cases (1993–2006). Equine Vet J 2008;40:364–7. 4. Bartmann CP, Freeman DE, Glitz F, von Oppen T, Lorber KJ, Bubeck K, Klug E. Diagnosis and surgical management of colic in the foal: Literature review and a retrospective study. Clin Tech Equine Pract 2002;1:125–42. 5. Semrad SD, Hardee GE, Hardee MM, Moore JN. Flunixin meglumine given in small doses: pharmacokinetics and prostaglandin inhibition in healthy horses. Am J Vet Res 1985;46:2474–9. 6. MacAllister CG, Morgan SJ, Borne AT, Pollet RA. Comparison of adverse effects of phenylbutazone, flunixin meglumine, and ketoprofen in horses. J Am Vet Med Assoc 1993;202:71–7. 7. Grindem CB, Fairley NM, Uhlinger CA, Crane SA. Peritoneal fluid values from healthy foals. Equine Vet J 1990;22:359–61. 8. Fisher A, Kerr L, O’Brien T. Radiographic diagnosis of gastrointestinal disorders in the foal. Vet Radiol 1987;28:42–8. 9. Orsini JA. Abdominal surgery in foals. Vet Clin North Am Equine Pract 1997;13:393–413. 10. Zedler ST, Embertson RM, Bernard WV, Bass BS, Boston RC. Surgical treatment of gastric outflow obstruction in 40 foals. Vet Surg 2009;38:623–30. 11. Clayton HM. Ascarids: recent advances. Vet Clin North Am Equine Pract 1986;2:313–28. 12. Austin S, DiPietro J, Foreman J. Parascaris equorum infections in horses. Compend Contin Educ Pract Vet 1990;12:1110–19. 13. Lyons ET, Tolliver SC, Ionita M, Collins SS. Evaluation of parasiticidal activity of fenbendazole, ivermectin, oxibendazole, and pyrantel pamoate in horse foals with emphasis on ascarids (Parascaris equorum) in field studies on five farms in Central Kentucky in 2007. Parasitol Res 2008;103:287–91. 14. Lavoie JP, Drolet R, Parsons D, Leguillette R, Sauvageau R, Shapiro J, Houle L, Hall´e G, Gebhart CJ. Equine proliferative enteropathy: a cause of weight loss, colic, diarrhoea and hypoproteinaemia in foals on three breeding farms in Canada. Equine Vet J 2000;32:418–25. 15. Santschi EM, Juzwiak JS, Moll HD, Slone DE. Diaphragmatic hernia repair in three young horses. Vet Surg 1997;26:242–5. 16. Serata V. Cecocolic intussusception in a foal. Acta Unio Int Contra Cancrum 2002;13:27–30. 17. Cable CS, Fubini SL, Erb HN, Hakes JE. Abdominal surgery in foals: a review of 119 cases (1977–1994). Equine Vet J 1997;29:257–61.
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Chapter 83
Weight Loss Kirsten M. Neil
Introduction Although weight loss, ill-thrift, and a failure to thrive are a common clinical presentation in the older foal, weanling and yearling, they are non-specific signs of disease. Loss of weight is usually insidious and chronic in nature; however, acute rapid weight loss may occur with some disorders. The main mechanisms involved are reduced dietary intake, increased nutritional demands, increased metabolic activity, and catabolism. A thorough history is important, in particular in relation to management practices, nutrition, and anthelmintic regimes. One or more horses in a group may be affected. Malnutrition may be a contributing factor and needs to be excluded as a cause of weight loss in this age group. A thorough physical examination is vital. Dentition should be examined, although dental problems are reported less frequently compared to in older horses. A pot-bellied appearance, poor body condition, and rough hair coat are common (Figure 83.1). Blood tests may reveal leukocytosis, hyperfibrinogenemia, and/or hyperglobulinemia suggesting an ongoing infectious process. Biochemical abnormalities vary depending on the underlying disorder, and may aid in identifying the organ systems involved and hence which other diagnostic tests are required. Ultrasonography is a powerful tool for examining affected horses, especially when the primary disease process is not obvious from the clinical examination or blood tests.
Gastrointestinal disease Weight loss and diarrhea are common clinical manifestations of gastrointestinal disease, frequently attributable to malabsorption, inflammation, and protein-losing enteropathy. Additional diagnostic tests such as gastroscopy, ultrasonography, abdominocentesis, radiography, glucose or xylose
absorption tests, fecal flotation and egg counts, or rectal mucosal or intestinal biopsies are often required to establish the diagnosis. Although weight loss and diarrhea can occur acutely, chronic diarrhea is a common manifestation of a variety of conditions affecting this age group. Parasitism Parasitism is a leading cause of weight loss in horses. Weight loss occurs due to malabsorption and proteinlosing enteropathy due to intestinal inflammation, increased nutritional requirements, organ and vessel damage from migrating larvae, and anorexia in advanced cases. The prevalence of parasite eggs in feces of more than 700 foals up to 8 months of age was 27.6% strongyles, 22.4% Parascaris equorum and 1.5% Strongyloides westeri.1 The prevalence of each parasite varied depending on the age group, with S. westeri common in younger foals. This parasite is infrequently associated with clinical disease;2 however, weight loss and edema were presenting signs in a 6-month-old foal with overwhelming strongyloidosis.3 The most important parasite of young horses is Parascaris equorum. Clinical disease is typically seen in foals less than 10–12 months of age, particularly suckling and weanling foals, with ill-thrift the most common presenting complaint.4 Weight loss, dull hair coat, pot-bellied appearance, and poor growth rates are attributed to both hepato-pulmonary larval migration, as well as the presence of mature ascarids in the small intestine which compete for nutrients and cause malabsorption. Other clinical manifestations include respiratory signs secondary to larval migration, diarrhea, and colic. Colic is typically seen in 5- to 6-month-old foals with heavy parasite burdens and a poor deworming history. Affected foals present with acute small intestinal obstruction, usually within 24 hours after treatment with anthelmintics (e.g.,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 83.1 Weight loss, ill-thrift and pot-bellied appearance in a weanling with an intra-abdominal abscess due to Rhodococcus equi infection.
piperazine, pyrantel pamoate, ivermectin) which cause sudden ascarid death.5 Ascaridiasis should be suspected as a cause of illthrift in foals with a poor deworming history, evidence of farm management practices which predispose to high parasite burdens, or when anthelmintic resistance is suspected. Overcrowding and repeated use of the same foaling and weanling paddocks can expose foals to high levels of environmental contamination, as ascarid eggs are extremely resistant to dessication and can survive in the environment for up to 5 years. Anthelmintic regimes of pregnant mares should also be considered as foals are usually infected shortly after birth. No serological tests are available to detect larval migration, and eggs do not appear in feces of affected foals until 10–12 weeks of age. Fecal flotation should be performed for suspect cases, although fecal egg counts are not a good indicator of the mature worm burden as female ascarids are prolific egg layers. Affected foals may pass adult worms within a few days of anthelmintic treatment, and passage of ascarids in both feces and nasogastric reflux has been reported in foals with colic.5 Worms may also be detected in the small intestinal lumen with abdominal ultrasonography. Hematological and biochemical abnormalities are non-specific, with hypoproteinemia, hypoalbuminemia, and occasionally anemia reported. Anthelmintic treatment is recommended for confirmed or suspect cases. Anthelmintics effective against luminal stages of Parascaris equorum include fenbendazole (10 mg/kg), oxibendazole (10 mg/kg), ivermectin (200 µg/kg), moxidectin (400 µg/kg; not registered for use in foals less than 4 months of age), piperazine (88 mg/kg), pyrantel pamoate
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(6.6 mg/kg), and pyrantel tartrate (2.64 mg/kg/ day).6 Recent studies have shown oxibendazole and fenbendazole to be the most effective anthelmintics.7, 8 Foals with ill-thrift and a poor deworming history are prime candidates for post-treatment impactions and colic, so initial treatment with a reduced dose of a dewormer that does not kill via paralysis (e.g., fenbendazole 5 mg/kg is recommended). Ascarids are the most important parasite to target in anthelmintic control programs in young horses. Routine anthelmintic treatment of foals should begin at 8 weeks of age, followed by 6- to 8-week dosing intervals. More frequent dosing intervals, in particular monthly treatment with ivermectin, have been associated with resistance.9 Anthelmintic resistance has been reported to pyrantel pamoate, even at twice the recommended dosage,8 and resistance to macrocyclic lactones (ivermectin and moxidectin) is an emerging problem. Macrocyclic lactone resistance appears to be widespread, with reports in America,7 Canada,10, 11 and Europe. 12, 13 The exact mechanism of resistance is unclear; however, it may relate to reduced susceptibility of migrating larvae compared to intestinal stages at recommended dosages.14 Fecal egg count reduction tests should be performed at least annually to monitor for anthelmintic resistance (see Chapter 30). Strongyle parasites are less commonly associated with weight loss in young horses. Foals exposed to heavy infestations and young horses with a poor deworming history are at risk; however, clinical disease may still occur despite a regular deworming program.15 Recently weaned 3- to 6-month-old foals grazing pasture contaminated with large and small strongyles developed weight loss or reduced weight gain as well as diarrhea.16 Heavy contamination was associated with debilitation, inappetance, and severe weight loss. Both acute and chronic cyathostominosis may be associated with weight loss, diarrhea, and edema. The acute form of the disease, larval cyathostominosis, is more common in young horses (1–3 years old).15 Diarrhea is the most common clinical sign;17 however, weight loss and edema without diarrhea has also been reported.18 Diagnosis can be problematic as clinical signs are non-specific. Supportive hematological and biochemical abnormalities include hypoalbuminemia and microcytic anemia. Adult cyathostomes or fourth-stage larvae may be detected in feces, but fecal egg counts may be negative.17 Strongyle eggs detected in feces of foals less than 6 months of age are assumed to be small strongyle eggs, although large strongyle eggs may be ingested by coprophagia and pass through the foal’s intestinal tract.1 Treatment with anthelmintics with larvicidal efficacy is indicated: fenbendazole (10 mg/kg once daily for 5 consecutive days) and
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moxidectin (400 µg/kg) are most effective. Supportive care in the form of intravenous fluid therapy including colloids may be indicated in foals with concurrent diarrhea, and affected horses may also benefit from treatment with corticosteroids (dexamethasone 0.02–0.05 mg/kg q 24 h or prednisolone 0.5 mg/kg p.o. q 12–24 h). Gastroduodenal ulceration (see also Chapter 74) Ill-thrift may be a presenting sign of gastroduodenal ulceration. In symptomatic suckling and weanling age foals with gastric ulcers, diarrhea and poor appetite are the most common clinical signs;19 however, ill-thrift, poor growth, rough hair coat, pot-bellied appearance, bruxism, excess salivation, and colic may also occur.20 Gastroduodenal ulceration should be considered as a cause of ill-thrift in 2- to 6-monthold foals, especially with a history of or evidence of concurrent clinical disease.21 Yearlings with gastric ulcers typically present with anorexia and chronic or intermittent colic.19 Vague signs such as poor condition, poor quality hair coat, reduced appetite, and failure to thrive are also reported. Although rare, pyloric stenosis has been documented as a cause of weight loss, inappetance, and bruxism in a yearling.22
cases, pigs have been reported in close proximity24, 27 or on the premises prior to affected foals.28–30 In one study, a dog tested positive on fecal PCR.30 Between 10–75% of in-contact horses on farms with clinical cases have been seropositive,30–33 while in other reports positive titers from in-contact foals and adults were uncommon.24, 27, 34 In-contact horses have either been negative on fecal PCR,30 or positive results were obtained in only 6% of age-matched clinically normal herd mates from farms with confirmed cases,31 despite positive serological titers in 75%30 and 33%31 of in-contact horses, respectively. Wildlife have been postulated to be the source of contamination.27 In a recent epidemiological study, PCRpositive fecal samples were detected from striped skunks, Virginian opossums, jackrabbits, and coyotes at equine farms with confirmed cases of EPE, suggesting these species may be a potential source of infection for equids.35 Recent equine epidemiological studies have shown that positive serum antibody titers were detected at the time of foaling in 55% of Thoroughbred broodmares on a farm endemic for EPE.33 Passive transfer of colostral antibodies was documented in 37 of 68 foals on the farm. Colostral antibodies were detectable in the serum of most foals for up to a month (range 1–3 months), suggesting that positive titers obtained in foals ≥ 2 months of age are likely to reflect postnatal exposure.36
Equine proliferative enteropathy Proliferative enteropathy is a transmissible enteric disease that can affect a number of species, but has been best described in pigs. Equine proliferative enteropathy (EPE) is an important emerging disease of young horses. The first equine report was a postmortem diagnosis of intestinal adenomatosis, attributed to Campylobacter-like organisms based on the histopathological similarity to porcine intestinal adenomatosis.23 The causative organism has since been identified as Lawsonia intracellularis, an obligate intracellular Gram-negative bacterium.
Epidemiology Equine proliferative enteropathy has a worldwide distribution, with reports in foals in North America, Canada, Australia, and Europe. The incidence in horses is unknown and while most reports have been of sporadic cases, endemic outbreaks have also been reported.24, 25 Transmission occurs via the fecooral route. Although the incubation period has not been determined in horses, the incubation period is 2–3 weeks in pigs, and the organism may survive in the environment for up to 2 weeks.26 The source of contamination is usually not known. In some equine
Signalment Affected foals are usually 3–6 months old,24 although EPE has been reported in foals as young as 2 months and horses up to 13 months of age.24, 27, 29, 31,37–39 A variety of breeds may be affected and no sex predilection has been evident. Infection in pigs usually occurs within 2 weeks of a stressful event such as weaning, with a similar association documented in equine cases.24, 27, 30, 34, 38 A seasonal component was suggested in a retrospective study of weanlings with EPE in Kentucky.31 All foals presented in fall to early winter (August to January), with 53% of cases in November and December, although this may also reflect the normal age distribution of foals in this area during that time of year.
Clinical signs Affected foals typically present with rapid weight loss, anorexia, diarrhea, colic, lethargy/depression, and/or ventral edema due to hypoproteinemia.24, 38 Affected foals may have a rough hair coat and potbellied appearance.24 Weight loss and ill-thrift are attributed to malabsorption, intestinal inflammation,
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Weight Loss protein-losing enteropathy, and catabolism. Pyrexia may be a feature in some foals.25, 37, 40, 41 Concurrent disease processes such as parasitism, respiratory infections, dermatitis, and gastric ulcers are not uncommon.24, 27 The most common presenting signs in sporadic cases treated at a university hospital were depression or lethargy, anorexia, weight loss, and ventral edema.38 In a recent outbreak on a Standardbred farm, affected foals presented with depression, colic, pyrexia, and ventral edema but without diarrhea.25 Conversely, in an outbreak on three breeding farms, depression, diarrhea, and colic predominated, and although rapid and severe weight loss was also a feature, most foals maintained a normal appetite.24 In a recent retrospective study of 57 foals with EPE, ventral edema was the most common clinical sign (81%) followed by lethargy (32%), diarrhea (26%), pyrexia (19%), and colic (7%).31
Diagnosis Ultrasonographic findings consistent with EPE are markedly increased small intestinal mural thickness (Figure 83.2). In one study, 89% of foals had 4–8 mm thickening,38 and up to 10 mm has been reported.30 However, in another study, 7 of 25 horses had small intestinal thickness within reference range, indicating that a normal sonogram should not rule out EPE.31 The most common clinicopathological abnormality is hypoalbuminemia.24, 25, 31, 38 Panhypoproteinemia is common with hypoglobulinemia, low albumin : globulin ratio24, 29 and decreased colloid oncotic pressure.38 Leukocytosis is detected
Figure 83.2 Ultrasonogram of markedly increased small intestinal mural thickness (A. . .A) in a weanling with equine proliferative enteropathy.
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more consistently than hyperfibrinogenemia.24, 31, 38 Other abnormalities are non-specific and include anemia or hemoconcentration, prerenal azotemia, and electrolyte imbalances such as hypochloremia, hyponatremia, hypocalcemia, and metabolic acidosis.24, 29–31, 38, 40 Mild elevations in creatine kinase are also common.24, 38 Oral glucose absorption tests (OGAT) have been adequate in some foals,24, 27, 40 but were abnormal in two recent studies in which foals had evidence of complete or partial small intestinal malabsorption.42, 43 In early studies, the diagnosis of EPE was established on postmortem examination.23, 28, 29, 44 Lawsonia intracellularis targets enterocytes, disrupting the normal cell cycle resulting in enlarged, elongated crypt cells and mucosal hyperplasia.44 Gross lesions of EPE include mucosal thickening of the distal small intestine, the surface of which may appear corrugated. Lesions may be segmental, irregular, multifocal, or diffuse and are most commonly found in the distal jejunum and ileum. Focal erosions and submucosal edema may also be present.45 Infrequently, lesions extend to the duodenum.29, 39, 44 Distal extension into the large intestine is also possible39 but appears uncommon, with concurrent mucosal necrosis and edema documented in the large intestine in only one foal.37 Histological changes include crypt hyperplasia and proliferation, villous atrophy, inflammatory infiltrates, and occasionally abscessation within the crypts.29, 39 The detection of curved rod-shaped intracellular bacteria within the apical cytoplasm of proliferating crypt epithelial cells with Warthin-Starry silver stain confirms the diagnosis.26 Immunohistochemistry of small intestinal samples44 and PCR of small intestinal biopsies or postmortem samples have also yielded positive results.28, 37, 44 Rectal mucosal and duodenal biopsies are unlikely to be useful diagnostic modalities as lesions are typically confined to the distal jejunum and ileum, although concurrent lymphocytic plasmacytic enteritis has been diagnosed by duodenal biopsy in a foal with EPE.42 Until recently, antemortem diagnosis of EPE was challenging with typical clinical findings (weight loss, diarrhea, ventral edema), clinical pathology (hypoproteinemia), ultrasonographic findings (thickened small intestine), and exclusion of other causes of enteric disease considered supportive of the diagnosis. A rapid clinical response to appropriate antimicrobial treatment is also considered suggestive of EPE.24 Due to its intracellular location, isolation of the organism requires cell culture media and is not routinely performed. Extrapolating from diagnostics used in pigs, fecal PCR and serological tests have become the definitive means by which to establish the diagnosis. PCR offers the advantage of fast results in the early stages of infection before antibody
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titers have risen.46 Serological tests use a Lawsonia intracellularis-specific immunoperoxidase monolayer assay (IPMA) to determine antibody titer in serum. IPMA is considered a more reliable indicator of active or recent infection compared to PCR, with an IPMA titer ≥ 60 considered positive.31 In pigs, PCR has a sensitivity of 39–67% with 100% specificity, and IPMA has a sensitivity of 100% and a specificity of 90%.47 In horses, fecal PCR tests are often positive at admission but IPMA titers may be negative, with time required for seroconversion.27, 30 Fecal PCR is specific but has a low sensitivity, with false-negative results possible.26, 31 A negative fecal PCR has been documented after 4–7 days of antimicrobial treatment;27, 41 thus fecal PCR may not be appropriate to establish the diagnosis after antimicrobial treatment has been initiated. Affected foals can remain seropositive for up to 6 months post-infection.27 Although fecal PCR indicates the presence of L. intracellularis, unaffected in-contact foals have also tested positive on PCR without clinical disease.31 Both fecal PCR and serum IPMA samples should be tested in suspect cases, as only 50% of foals with EPE tested positive on both fecal PCR and IPMA in one study.31
bial treatment, the prognosis for survival of affected foals is good. Nine of 11 foals treated at a university hospital survived38 and 82% of foals in an outbreak of EPE recovered.24 Fifty-three of 57 foals survived in a retrospective study of foals with EPE in Kentucky.31 Although only 14 of these foals were subsequently sold as yearlings at auction, affected foals sold at an average of 68% of the average price of other yearlings by the same sire, suggestive of growth retardation in affected foals.31 Clinical improvement, including resolution of diarrhea, is usually seen within 2–3 days of treatment with appropriate antimicrobials. Hypoproteinemia and hypoalbuminemia usually take longer to resolve; however, some improvement is usually seen within 1–2 weeks. Serum protein and albumin concentrations returned to within normal range 50–90 days after presentation24, 34, 41 but in other foals hypoproteinemia has been more prolonged.43 In the latter study, continued partial malabsorption of glucose was also documented 58 days post-infection despite resolution of abdominal ultrasonographic abnormalities.43
Prevention Treatment Erythromycin estolate (25 mg/kg p.o. q 8 h) alone or in combination with rifampin (5–10 mg/kg p.o. q 12 h) for a minimum of 3 weeks are the most commonly used antimicrobials and have been shown to be effective.24 Other macrolides such as azithromycin (10 mg/kg p.o. q 24 h) and clarithromycin (7.5 mg/kg p.o. q 12 h) have also been successfully used.30, 31 Chloramphenicol (44 mg/kg p.o. q 6–8 h) treatment has also been effective.24, 31 Extrapolating from pig isolates, penicillin and ampicillin may also be effective; however, a response to treatment is often not seen in vivo with these antimicrobials.30, 34, 41 Foals have also been successfully treated with oxytetracycline (6.6 mg/kg i.v. q 12 h for 7 days) and doxycycline (10 mg/kg p.o. q 12 h for 8–17 days).25, 31, 38 Antimicrobial choice was not associated with survival in a recent retrospective study.31 Other supportive care in the form of intravenous fluids, colloids (plasma, hetastarch), electrolytes, and gastroprotectants is usually required. Surgical treatment (jejunocecostomy) has been described in one filly which presented with colic.34
Prognosis In initial reports of foals with EPE, death occurred before the diagnosis was made and appropriate therapy could be initiated.23, 28, 44 With appropriate antimicro-
Strategies to prevent EPE are currently being investigated. Affected foals should be isolated and herd mates tested and monitored closely. Stresses such as overcrowding should be avoided. Disinfectants such as povidone-iodine and quaternary ammonium compounds may be effective. Two recent studies have involved administration of an avirulent live vaccine of L. intracellularis.48, 49 No adverse reactions were detected. Foals were vaccinated twice, 3 weeks apart, with the vaccine administered either orally, orally following premedication with omeprazole (to determine the effect of gastric acidity on vaccine efficacy), or intra-rectally.48 The humoral immune response and onset and duration of fecal shedding were documented. Fecal shedding lasted 1–12 days, and was predominately evident in foals that received the intra-rectal vaccine. The strongest humoral response was seen in foals vaccinated intrarectally; all foals in this group seroconverted after the first vaccine. In comparison, none of the foals vaccinated intra-orally without premedication seroconverted, and only 50% of foals vaccinated orally after pre-medication with omeprazole seroconverted. In a subsequent study, intra-rectal vaccine was administered to broodmares 3–5 weeks before expected foaling dates.49 The pattern of fecal shedding was similar to that described in the weanling foal vaccination trial,48 but of a lower magnitude. Mares seroconverted 14–28 days post-vaccination (mean 20.1 days), with antibody detectable for 42–70 days (mean
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Weight Loss 59.5 days). Colostral antibodies were detectable in some but not all mares, perhaps related to a delay in the production of serum antibody relative to the time of foaling. Passive antibody transfer was documented in most foals from vaccinated mares, with serum antibody titers in foals detectable for 11–56 days (mean 44.5 days). Although serum antibody titers in the dams were not predictive of colostral and antibody titers in their foals, all mares with undetectable serum antibody titers at time of foaling also had undetectable colostral antibody levels. Although production of humoral antibody was demonstrated in both studies, cell-mediated immunity and local production of mucosal IgA are also likely to be important.49 The usefulness of vaccination in prevention of infection on endemic farms remains to be determined.
Infiltrative and inflammatory bowel disease Infiltrative and inflammatory bowel disease are infrequently reported in horses less than 1 year old50 and rarely manifest clinically in foals less than 6 months of age.51 Chronic wasting, poor condition, chronic diarrhea, edema, and colic are common clinical signs. The most common proliferative inflammatory bowel disease in weanling foals is EPE. Other conditions which infrequently affect yearlings include granulomatous enteritis and multisystemic eosinophilic epitheliotrophic disease.52 Idiopathic eosinophilic enteritis has been reported in a 10-week-old Quarter Horse, although presenting signs related to colic and diarrhea rather than weight loss.53 Glucose or xylose absorption tests and rectal mucosal biopsy are often required to establish the diagnosis in young horses.54
Other gastrointestinal disorders Weight loss and diarrhea may be associated with chronic salmonellosis and/or ulcerative enterocolitis due to Rhodococcus equi infection. Sand accumulation in the gastrointestinal tract is associated with weight loss and difficulty maintaining weight, as well as diarrhea.45, 55, 56
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Pneumonia is a common condition in weanlings and yearlings, associated with both bacterial and viral infections. However, weight loss is not usually a feature unless the disease is chronic or complications such as pleuropneumonia develop. Ultrasonography and radiography are valuable to characterize the pneumonia, and culture of transtracheal aspirates is recommended. Peritonitis has been reported in foals 2 months of age and older57 and may occur associated with a number of disorders such as gastroduodenal ulcer perforation, septicemia, enteritis, parasites (verminous arteritis, ascarid impactions), umbilical infections, and intra-abdominal abscessation. Peritonitis associated with Actinobacillus equuli has been reported in horses as young as 9 months old.58 Clinical signs usually depend on the primary disorder; however colic, weight loss, fever, depression, and anorexia are common. Although bacterial endocarditis occurs infrequently, the disorder is more common in young horses, with cases documented in foals from 2 to 3 months of age.59, 60 The clinical presentation can vary, although intermittent fever, weight loss, and depression are common.61 Murmurs are frequently audible, and signs of congestive heart failure may be evident in advanced cases. Organisms frequently isolated include Streptococcus sp., Actinobacillus sp., and Pasteurella sp.60, 61 Endocarditis due to Rhodococcus equi infection has also been reported.62 Echocardiography should be performed in any suspect cases, with vegetative lesions most common on the mitral and aortic valves.60 Treatment requires long-term antimicrobials, and the prognosis is generally considered guarded to poor. Weight loss is an uncommon clinical manifestation of pericarditis. In a recent outbreak of fibrinous pericarditis, foals from 3 weeks of age were affected, and 48% of cases were less than 2 years old.63 More common clinical signs are fever, lethargy, and edema.61 The diagnosis is confirmed by echocardiography, and antimicrobial treatment and pericardial drainage are usually required. Streptococcus sp. and Actinobacillus sp. have been most frequently isolated.63, 64 Internal abscessation
Chronic infection Chronic infection in any organ system can result in weight loss or ill-thrift. Anorexia, lethargy, and pyrexia are usually evident. Affected foals often present with weight loss associated with fever of unknown origin, with ancillary diagnostic tests required to establish the origin of the infection. Common hematological abnormalities include leukocytosis, hyperfibrinogenemia, and hyperglobulinemia.
Hematogenous spread of bacteria or local extension of infection may lead to the formation of internal abscesses. Internal abscessation is a common cause of chronic infection, fever of unknown origin, weight loss, and ill-thrift in older foals, weanlings, and yearlings. Extrapulmonary abscessation such as in the cranial mediastinum may occur as a complication of conditions such as pleuropneumonia. Intraabdominal abscessation is common in this age group,
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Figure 83.3 Ultrasonogram of intra-abdominal abscesses. (a) Large consolidated abscess in the caudoventral abdomen from foal depicted in Figure 83.1. (b) Abscess (arrows) in cranioventral abdomen of yearling with weight loss and fever of unknown origin.
either as a primary infectious disease process or as a complication of other disorders such as gastroduodenal ulceration, parasites,65 or compromised intestinal viability. Intra-abdominal abscesses may involve organs such as the liver, kidney, spleen or mesenteric lymph nodes, and adhesions and peritonitis are frequent sequelae. Diagnosis may be difficult as laboratory findings simply indicate chronic infection, and abnormalities are not always detected on abdominocentesis. Abdominal ultrasonography is useful (Figure 83.3a,b) and other helpful modalities include radiography and radiolabeled autologous leukocytes, although this is not commonly available.64 Exploratory celiotomy is required to establish the diagnosis in some cases. A number of organisms have been implicated; however, Rhodococcus equi, Streptococcus sp., and Corynebacterium pseudotuberculosis are commonly isolated. Treatment commonly requires long-term antimicrobial administration, and although surgical resection may be attempted in some cases, adhesions to intestinal viscera are common.
bination with erythromycin estolate (25 mg/kg p.o. q 8 h), azithromycin (10 mg/kg p.o. q 24 h), or clarithromycin (7.5 mg/kg p.o. q 12 h) (see Chapter 76).
Streptococcus equi Streptococcus equi subsp. equi infection is common in young horses, and metastatic abscessation (‘bastard strangles’) is an infrequent complication. Spread may occur hematogenously or via lymphatics, with mesenteric lymph nodes, liver, spleen, and kidneys most commonly affected.66 Diagnosis can be difficult and may rely on a history of exposure to the organism and elevated serum antibody titers. Occasionally peritoneal fluid antibody titers are also elevated.66 Treatment requires long-term antimicrobial therapy; however, metastatic abscesses may be refractory to treatment due to inadequate penetration
Rhodococcus equi The most common manifestation of Rhodococcus equi infection is bronchopneumonia; however, intestinal manifestations are also documented. Intraabdominal abscessation occurs infrequently and should be suspected in ill-thrifty weanlings in particular, with or without a history of respiratory disease (Figure 83.4). Prognosis for foals with this form of Rhodococcus equi infection is generally poor. Treatment requires long-term antimicrobial administration with rifampin (5–10 mg/kg p.o. q 12 h) in com-
Figure 83.4 Postmortem sectioned view of intra-abdominal abscess depicted in Figure 83.3a. Multiple loops of jejunum and omentum were adhered to the abscess.
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Weight Loss or inactivation in pus. The antimicrobial of choice is penicillin (22 × 103 IU/kg i.m. q 12 h penicillin G). Cephalosporins (ceftiofur sodium 2.2 mg/kg i.m. q 12–24 h) or macrolides may also be effective, and most isolates are also sensitive to trimethoprimsulfadiazine (30 mg/kg p.o. q 12 h). Streptococcus equi subsp. zooepidemicus may also cause internal abscessation, as a consequence of hematogenous spread from a respiratory infection.
Corynebacterium pseudotuberculosis Although infrequent in foals less than 6 months old, 1- to 2-year-old horses are at increased risk of Corynebacterium pseudotuberculosis infection.67 External abscessation (‘pigeon fever’) is the most common form of the disease.68 Clinical signs are predominantly fever, edema, and non-healing wounds; however, weight loss and anorexia may also be a feature. Weight loss is not usually associated with ulcerative lymphangitis unless the disease becomes chronic. Internal infections occur in approximately 8% of affected horses68 and has a high case fatality rate. This form of the disease has been reported in foals as young as 10 months.69 The most common clinical signs are concurrent external abscesses, anorexia, lethargy, weight loss, and signs of respiratory disease or abdominal pain.69 The liver, lungs, and kidney are most commonly involved. The organism may be cultured from accessible abscesses (usually external), and from peritoneal and pleural fluid. Serology is useful for diagnosis of internal infection, with synergistic hemolysis inhibition (SHI) titers ≥ 512 considered diagnostic.68 Antimicrobial treatment of internal infections is vital, with a 100% fatality rate reported in horses not receiving antimicrobials.68 Most antimicrobials are effective and are often used in combination with rifampin (5–10 mg/kg p.o. q 12 h).69
Other disorders Other disease processes are infrequently reported to cause weight loss or failure to thrive in older foals, weanlings, or yearlings. Weight loss is a non-specific sign and is often not the primary presenting complaint. Weight loss has been reported along with limb edema, anorexia, and fever in foals as young as 6 months with purpura hemorrhagica.70 Congestive heart failure (CHF) is uncommon in young horses, but may be associated with signs of weight loss, edema, and exercise intolerance. In addition to endocarditis, congenital abnormalities such as ventricular septal defects are possible causes of CHF even in yearlings. Weight loss due to chronic renal failure is infrequent in young horses. Renal hypoplasia and renal
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dysplasia have been reported as causes of weight loss, lethargy, poor growth, anorexia, and renal azotemia in 3- to 12-month-old horses.71, 72 Hepatic disorders are uncommon, although chronic hepatic insufficiency can lead to significant weight loss and failure to thrive.73 Possible causes include hepatic abscessation, ascending cholangitis associated with duodenal ulcers, plant toxicities (e.g. pyrrolizidine alkaloid), congenital abnormalities, and neoplasia. Stunted growth has been reported in 2- to 6-monthold foals with hepatic encephalopathy due to portosystemic shunts.73 Hepatocellular carcinoma has been reported as a cause of poor growth, erythrocytosis, and hypoglycemia in a yearling filly.74
References 1. Lyons ET, Tolliver SC. Prevalence of parasite eggs (Strongyloides westeri, Parascaris equorum, and strongyles) and oocysts (Eimeria leuckarti) in the feces of Thoroughbred foals on 14 farms in central Kentucky in 2003. Parasitol Res 2004;92:400–4. 2. Netherwood T, Wood WL, Townsend HG, Mumford JA, Chanter N. Foal diarrhea between 1991 and 1994 in the United Kingdom associated with Clostridium perfringens, rotavirus, Strongyloides westeri and Cryptosporidium spp. Epidemiol Infect 1996;117(2):375–83. 3. Brown CA, MacKay RJ, Chandra S, Davenport D, Lyons ET. Overwhelming strongyloidosis in a foal. J Am Vet Med Assoc 1997;211(3):333–4. 4. Love S. Treatment and prevention of intestinal parasiteassociated disease. In: Jones SL (ed.) Therapeutics for Gastrointestinal Disease: Vet Clin North Am Equine, 2003;19(3): 791–806. Philadelphia: Elsevier Saunders. 5. Cribb NC, Cote NM, Boure LP, Peregrine AS. Acute small intestinal obstruction associated with Parascaris equorum infection in young horses: 25 cases (1985–2004). N Z Vet J 2006;54:338–43. 6. Reinemeyer CR, Nielsen MK. Parasitism and colic. In: Andrews FM (ed.) New Perspectives in Equine Colic: Vet Clin N Am Equine 2009;25:233–45. 7. Lyons ET, Tolliver SC, Collins SS. Field studies on endoparasites of Thoroughbred foals on seven farms in central Kentucky in 2004. Parasitol Res 2006;98:496–500. 8. Lyons ET, Tolliver SC, Ionata M, Collins SS. Evaluation of parasiticidal activity of fenbendazole, ivermectin, oxibendazole, and pyrantel pamoate in horse foals with emphasis on ascarids (Parascaris equorum) in field studies on five farms in central Kentucky in 2007. Parasitol Res 2008;103:287–91. 9. Craig TM, Diamond PL, Ferwerda NS, Thompson JA. Evidence of ivermectin resistance by Parascaris equorum on a Texas horse farm. J Eq Vet Sci 2007;27:67– 71. 10. Hearn FP, Peregrine AS. Identification of foals infected with Parascaris equorum apparently resistant to ivermectin. J Am Vet Med Assoc 2003;223:482–5. 11. Slocombe JO, de Gannes RV, Lake MC. Macrocyclic lactoneresistant Parascaris equorum on stud farms in Canada and effectiveness of fendendazole and pyrantel pamoate. Vet Parasitol 2007;145:371–6. 12. Boersema JH, Eysker M. Apparent resistance of Parascaris equorum to macrocyclic lactones. Vet Rec 2002;150:279–81.
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13. Schougaard H, Nielsen MK. Apparent ivermectin resistance of Parascaris equorum in foals in Denmark. Vet Rec 2007;160:439–40. 14. DiPietro JA, Lock TF, Todd KS, Reuter VE. Evaluation of ivermectin paste in the treatment of ponies for Parascaris equorum infections. J Am Vet Med Assoc 1987;190:1181–83. 15. Lyons ET, Drudge JH, Tolliver SC. Larval cyathostomiasis. In: Timoney PJ (ed.) Emerging Infectious Diseases: Vet Clin N Am Equine 2000;16(3):501–13. 16. Thamsborg SM, Leifsson PS, Grondahl C, Larsen M, Nansen P. Impact of mixed strongyle infection in foals after one month on pasture. Equine Vet J 1998;30:240–5. 17. Peregrine AS, McEwen B, Bienzie D, Koch TG, Weese JS. Larval cyathostominosis in horses in Ontario: an emerging disease? Can Vet J 2006;47:80–2. 18. Mair TS. Outbreak of larval cyathostomiasis among a group of yearlings and two year old horses. Vet Rec 1994; 135:598–600. 19. Sanchez LC. Diseases of the stomach. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis: Sanders, 2004; pp. 863–73. 20. Andrews FM, Nadeau JA. Clinical syndromes of gastric ulceration in foals and mature horses. Equine Vet J Supp 1999;29:30–3. 21. Murray MJ. Endoscopic appearance of gastric lesions in foals: 94 cases (1987–1988). J Am Vet Med Assoc 1989;195(8): 1135–41. 22. Heidmann P, Saulez MN, Cebra CK. Pyloric stenosis with reflux oesophagitis in a Thoroughbred filly. Eq Vet Educ 2004;16(4):172–6. 23. Duhamel GE, Wheeldon EB. Intestinal adenomatosis in a foal. Vet Pathol 1982;19:447–50. 24. Lavoie JP, Drolet R, Parsons D, Leguillette R, Sauvageau R, Shapiro J, Houle L, Halle G, Gebhart CJ. Equine proliferative enteropathy: a cause of weight loss, colic, diarrhoea and hypoproteinemia in foals on three breeding farms in Canada. Equine Vet J 2000;32(5):418–25. 25. Merlo JL, Sheats MK, Elce Y, Hunter S, Brehaus BA. Outbreak of Lawsonia intracellularis on a standardbred breeding farm in North Carolina. Equine Vet Educ 2009;21(4):179– 82. 26. Lavoie JP, Drolet R. Lawsonia intracellularis proliferative enteropathy. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. St Louis: Saunders, 2003; pp. 164–6. 27. Dauvillier J, Picandet V, Harel J, Gottschalk M, Desrosiers R, Jean D, Lavoie JP. Diagnostic and epidemiological features of Lawsonia intracellularis enteropathy in 2 foals. Can Vet J 2006;47:689–91. 28. Williams NM, Harrison LR, Gebhart CJ. Proliferative enteropathy in a foal caused by Lawsonia intracellularis like bacterium. J Vet Diagn Invest 1996;8:254–6. 29. Brees DJ, Sondhoff AH, Kluge JP, Andreasen CB, Brown CM. Lawsonia intracellularis-like organism infection in a miniature foal. J Am Vet Med Assoc 1999; 215(4):511–14. 30. Feary DJ, Gebhart CJ, Pusterla N. Lawsonia intracellularis proliferative enteropathy in a foal. Schweiz Arch Tierheilk 2007;149(3):129–33. 31. Frazer ML. Lawsonia intracellularis infection in horses: 2005–2007. J Vet Int Med 2008;22(5):1243–48. 32. Pusterla N, Higgins JC, Smith P, Mapes S, Gebhart C. Epidemiological survey on farms with documented occurrence of equine proliferative enteropathy due to Lawsonia intracellularis infection. Vet Rec 2008;163(5):156–8. 33. Pusterla N, Jackson R, Wilson R, Collier J, Mapes S, Gebhart C. Temporal detection of Lawsonia intracellularis using
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36.
37.
38.
39.
40. 41.
42.
43.
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45.
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serology and real-time PCR in Thoroughbred horses residing on a farm endemic for equine proliferative enteropathy. Vet Microbiol 2009;136(1):173–6. Schumacher J, Schumacher J, Rolsma M, Brock KV, Gebhart CJ. Surgical and medical treatment of an Arabian filly with proliferative enteropathy caused by Lawsonia intracellularis. J Vet Int Med 2000;14:630–2. Pusterla N, Mapes S, Rejmanek D, Gebhart C. Detection of Lawsonia intracellularis by real-time PCR in the feces of free-living animals from equine farms with a documented occurrence of equine proliferative enteropathy. J Wildl Dis 2008;44(4):992–8. Lavoie JP, Drolet R. Equine proliferative enteropathy: an emerging disease of foals. Equine Vet Educ 2009;21(4):183– 5. Deprez P, Chiers K, Gebhart CJ, Ducatelle R, Lef`ere L, Vanschandevijl K, Van Loon G. Lawsonia intracellularis infection in a 12-month-old colt in Belgium. Vet Record 2005;157:774–6. Sampieri F, Hinchcliff KW, Toribio. Tetracycline therapy of Lawsonia intracellularis enteropathy in foals. Equine Vet J 2006;38(1):89–92. Wuersch K, Huessy D, Koch C, Oevermann A. Lawsonia intracellularis proliferative enteropathy in a filly. J Vet Med Assoc 2006;53:17–21. Bihr TP. Protein-losing enteropathy caused by Lawsonia intracellularis in a weanling foal. Can Vet J 2003;44:65–6. McClintock SA, Collins AM. Lawsonia intracellularis proliferative enteropathy in a weanling foal in Australia. Aust Vet J 2004;82(12):750–2. Allen KJ, Pearson GR, Fews D, McOrist S, Brazil TJ. Lawsonia intracellularis proliferative enteropathy in a weanling foal, with a tentative histological diagnosis of lymphocytic plasmacytic enteritis. Eq Vet Educ 2009;21(8):411–14. Wong DM, Alcott CJ, Sponseller BA, Young JL, Sponseller BT. Impaired intestinal absorption of glucose in 4 foals with Lawsonia intracellularis infection. J Vet Int Med 2009;23(4):940–4. Frank N, Fishman CE, Gebhart CJ, Levy M. Lawsonia intracellularis proliferative enteropathy in a weanling foal. Equine Vet J 1998;30(6):549–52. Jones SL. Inflammatory diseases of the gastrointestinal tract causing diarrhea. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis: Saunders, 2004; pp. 884–912. Pusterla N, Madigan JE, Leutenegger CM. Real-time polymerase chain reaction: a novel molecular diagnostic toll for equine infectious diseases. J Vet Int Med 2006;20:3–12. Guedes RM, Gebhart CJ, Deen J, Winkelman NL. Validation of an immunoperoxidase monolayer assay as a serologic test for porcine proliferative enteropathy. J Vet Diagn Invest 2002;14:528–30. Pusterla N, Higgins H, Wattanaphansak S, Collier JR, Mapes SM, Stenbom RM, Gebhart C. Evaluation of the humoral immune response and fecal shedding in weanling foals following oral and intra-rectal administration of an avirulent live vaccine of Laswonia intracellularis. Proc Am Assoc Equine Pract 2008;54:314–15. Pusterla N, Collier J, Mapes SM, Wattanaphasak S, Gebhart C. Effects of administration of an avirulent live vaccine of Lawsonia intracellularis on mares and foals. Vet Rec 2009;164:783–5. Scott EA, Heidel JR, Snyder SP, Ramirez S, Whitler WA. Inflammatory bowel disease in horses: 11 cases (1988–1998). J Am Vet Med Assoc 1999;214(10):1527–30.
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Weight Loss 51. Merritt AM. Idiopathic eosinophilic enteritis: etiology and pathophysiology. Compend Cont Educ Equine 2002;24(4): 344–7. 52. Roberts MC. Proliferative and inflammatory intestinal diseases associated with malabsorption and maldigestion. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis: Saunders, 2004; pp. 878– 84. 53. Stanar LS, Little D, Redding WR, Jones SL. Idiopathic eosinophilic enteritis in a 10-week-old colt. Comp Cont Educ Equine 2002; 24(4):342–4. 54. Lindberg R, Nygren A, Persson SGB. Rectal biopsy diagnosis in horses with clinical signs of intestinal disorders: a retrospective study of 116 cases. Equine Vet J 1996;28(4):275–84. 55. Ramey DW, Reinertson EL. Sand induced diarrhea in a foal. J Am Vet Med Assoc 1984;185(5):537–8. 56. Bertone JJ, Traub-Dargatz JL, Wrigley RW, Bennett DG, Williams RJ. Diarrhea associated with sand in the gastrointestinal tract of horses. J Am Vet Med Assoc 1988;193(11): 1409–12. 57. Dyson S. Review of 30 cases of peritonitis in the horse. Equine Vet J 1983;15(1):25–30. 58. Matthews S, Dart AJ, Dowling BA, Hodgson JL, Hodgson DR. Peritonitis associated with Actinobacillus equuli in horses: 51 cases. Aust Vet J 2001;79(8):536–9. 59. Buergelt CD, Cooley AJ, Hines SA, Pipers FS. Endocarditis in six horses. Vet Pathol 1985;22(4):333–7. 60. Maxson AD, Reef VB. Bacterial endocarditis in horses: ten cases (1984–1995). Equine Vet J 1997;29(5):394–9. 61. Bonagura JD, Reef VB. Disorders of the cardiovascular system. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis: Saunders, 2004; pp. 355– 459. 62. Collatos C, Clark S, Reef VB, Morris DD. Septicemia, atrial fibrillation, cardiomegaly, left atrial mass and Rhodococcus equi septic osteoarthritis in a foal. J Am Vet Med Assoc 1990;197:1039–42. 63. Bolin DC, Donahue JM, Vickers ML, Harrison L, Sells S, Giles RC, Hong CB, Poonacha KB, Roberts J, Sebastian MM, Swerczek TW, Tramontin R, Williams NM. Microbiologic
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and pathologic findings in an epidemic of equine pericarditis. J Vet Diagn Invest 2005;17:38–44. Jesty SA, Reef VB. Septicemia and cardiovascular infections in horses. In: Southwood LL (ed.) Advances in diagnosis and management of infection: Vet Clin N Am Equine 2006;22:481–95. DiPietro JA, Boero M, Ely RW. Abdominal abscess associated with Parascaris equorum infection in a foal. J Am Vet Med Assoc 1983;182(9):991–2. Sweeney CR, Timoney JF, Newton JR, Hines MT. Streptococcus equi infections in horses: guidelines for treatment, control and prevention of strangles. J Vet Int Med 2005;19:123–34. Doherr MG, Carpenter TE, Hanson KM, Wilson WD, Gardner IA. Risk factors associated with Corynebacterium pseudotuberculosis infection in California horses. Prev Vet Med 1998;35(4):229–39. Aleman M, Spier SJ, Wilson WD, Doherr M. Corynebacterium pseudotuberculosis infection in horses: 538 cases (1982–1993). J Am Vet Med Assoc 1996;209(4):804–9. Pratt SM, Spier SJ, Carroll SP, Vaughan B, Whitcomb MB, Wilson WD. Evaluation of clinical characteristics, diagnostic test results, and outcome in horses with internal infection caused by Corynebacterium pseudotuberculosis: 30 cases (1995–2003). J Am Vet Med Assoc 2005;227(3):441–8. Pusterla N, Watson JL, Affolter VK, Magdesian KG, Wilson WD, Carlson GP. Purpura haemorrhagica in 53 horses. Vet Rec 2003;153(4):118–21. Andrew FM, Rosol TJ, Kohn CW, Reed SM, DiBartola SP. Bilateral renal hypoplasia in four young horses. J Am Vet Med Assoc 1986;189(2):209–12. Plummer PJ. Congenital renal dysplasia in a 7-month-old quarterhorse colt. In: MacLeay JM (ed.) Medical Case Management: Vet Clin N Am Equine 2006;22:e63–9. Barton MH. Disorders of the liver. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis: Saunders, 2004; pp. 951–94. Roby KA, Beech J, Bloom JC, Black M. Hepatocellular carcinoma associated with erythrocytosis and hypoglycemia in a yearling filly. J Am Vet Med Assoc 1990;196(3):465–7.
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Chapter 84
Angular Limb Deformities Earl M. Gaughan and Sarah E.T. Hanna
Angular limb deformities (ALDs) are distortions of the limb typically considered to have a lateral (outward) or medial (inward) deviation of the long axis of an affected limb.1, 2 This deviation is often associated with a joint, or with a physeal region. Angular limb deformities do not appear to be associated with any particular breed or gender. The two classifications of angular limb deformities are valgus and varus – deviations laterally and medially, respectively. Valgus deviations often appear as knock-kneed and varus as bowlegged. These classifications can be further described by their location on the limb and what joints are involved, for example, carpal valgus.
Etiology Angular limb deformities can be present at the time of foaling, as congenital deformities, or can be acquired with time and growth after birth.3, 4 Congenital deformities are often associated with laxity of tendons and ligaments, which in turn fail to support a joint, causing a deformation of limb conformation. Deformed or unformed bones can also play a role in congenital angular deformities.3 For example, the structure of the cuboidal bones in the carpus is instrumental to joint architecture. If these bones are not completely ossified, the result can be an alteration in the normal angulation of the limb due to the nonrigid structure of the joint. Incomplete ossification can result from perinatal events that affect the uterine environment (Figure 84.1). Events that alter the intrauterine environment, such as systemic illness or parasitic infection, can result in incomplete ossification of the cuboidal bones, just as placentitis, premature foaling, and twinning can and may be related to intrauterine growth retardation. If the angulation of a limb is normal at birth, but an ALD occurs later, this deformity is considered acquired and can result from changes in the physes of long bones. Physeal abnormalities can substantially
influence acquired angular limb deformities and can be associated with physitis, or inflammation of the growth plate. Physitis can result from excessive, asynchronous, or abnormal growth and from overt or micro-trauma.5 Physeal development can be retarded or accelerated by unbalanced weight bearing secondary to an existing lameness or pain. If weight bearing is not symmetrical, uneven strains initiated at the hoof can be distributed proximally through the long bones. The differential of this strain within the limb can result in uneven growth at the physes, leading to abnormal growth of the long bone, and an alteration in the angulation of the limb. Although a common cause for ALD, physeal abnormalities are not the sole consideration in the development of acquired ALDs. Trauma-related changes can also cause change in angles of a limb. Fractures of the carpal or tarsal bones, hypoplasia, or displacement of the carpal, tarsal, metacarpal, or metatarsal bones can also cause abnormal angles of the limbs.
Examination and diagnosis When examining a foal with ALD, a complete physical examination is essential. It is important to determine if the defect is likely of soft tissue origin or skeletal, as this will influence the treatment options that need to be employed. If the defect can be manually reduced, the angular deformity is most likely due to incomplete ossification of the cuboidal bones or to a soft tissue abnormality, such as periarticular ligamentous/tendinous laxity. If the defect cannot be manually reduced, the deformity most likely involves osseous structures, and radiographs are typically indicated. Observation of an affected horse’s gait is important as well. If an apparent angulation results from rotation of the entire limb (i.e., due to a thin chest or other conformation faults) all of the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 84.1 A dorsal to caudal radiographic projection of a foal with premature cuboidal bones in the carpus.
joints in the limb will move together in the same way. No rotation or angulation of an individual joint will be demonstrated if the entire limb is rotated around the long axis. Radiography can be very helpful in the assessment of any ALD. Radiographs can help to determine the angle of deviation in an angular limb deformity and in determining complete ossification of the cuboidal bones. A dorsopalmar projection of an affected site is typically obtained and evaluated using either the pivot point or individual joint angle technique. The pivot point technique involves drawing a line bisecting the long axis of each long bone until the lines cross each other (Figure 84.2). The point where the lines cross identifies the location of the abnormal deviation. Individual joints can also be assessed. Using this technique can provide localization of an affected region.
Treatment Overall, an angle of 3◦ or less can be considered within normal limits and no therapy is indicated. It is important to identify incomplete ossification of the cuboidal bones. Due to the soft precursor cartilage, the shape of the cuboidal bones can easily be altered due to uneven weight bearing. This can result in lifelong debilitating, angulation that cannot be rectified by surgical intervention.
Figure 84.2 A dorsal to caudal radiographic projection of a foal with carpal valgus demonstrating bisecting lines at a pivot point, which indicates the most likely site of angulation.
It is important to determine how quickly, if at all, the angle is changing, and also to recognize the time period in which one has to alter the conformation of the limb. In order to ascertain the time frame in which alteration of the physis will result in adequate correction of the angulation, it is important to know when each physis ceases bone development. There are differences between the distal and proximal limb. A rule of thumb is that more distal long bone physes ‘close’ earlier than more proximal long bone physes (i.e., a fetlock will require interventional therapy in the first weeks of life whereas the distal radial physes can be manipulated through the first year of life). Treatment options vary with the severity of the deviation. For mild angulation deformities, rest to prevent further injury or angulation may be all that is required. Diet should be appropriately monitored, specifically the phosphorus:calcium ratio and total caloric intake. A high concentration of phosphorus in the diet has been associated with physitis. Medical therapy can include non-steroidal anti-inflammatory drugs to reduce inflammation of affected physes. Occasionally mild cases of angular limb deformity may be assisted toward correction with hoof trimming.6 For example, the most common deviation, carpal valgus, can be treated by placing medial hoof supports
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to extend the medial wall. This type of hoof manipulation has been recommended every 2 to 3 weeks in order to maintain the correct therapeutic hoof shape. However, cases of substantial ALD should not be expected to respond to hoof trimming alone. Congenital abnormalities may not require aggressive therapy and are classified under two different categories. Congenital abnormalities with tendon laxity can often be attributed to prematurity of the foal. These foals can be treated with exercise restriction, often stall rest with controlled exercise such as hand walking, if possible. Rigid external support can provide support to lax tendons and ligaments; however, constant support of the limb can stress protect and exacerbate tendon laxity. Congenital abnormalities without tendon laxity usually only require rest. Young horses with uncorrecting ALDs are perhaps best treated with rest until 2–3 months of age if the ALD is unchanging. If the ALD corrects, or the angle of deviation decreases, it is probably best to continue rest and observe changes that may occur. If an ALD worsens with time, aggressive intervention is likely best for the athletic future of the horse. Surgical treatment is indicated for young horses with deformities that do not correct or become worse. Also, foals greater than 4 months of age will most likely require surgery for angular limb deformity correction. Surgical options include periosteal stripping, considered to be a growth-accelerating procedure, and transphyseal bridging, a growth-retarding procedure. Different abnormalities require different surgical techniques. Periosteal stripping involves a hemicircumferential transection of the periosteum on the side of the bone where growth stimulation is desired (the concave or growth-deficient aspect of the affected long bone). Periosteal stripping has been thought to stimulate the osteocytes to replicate more readily, therefore causing long bone growth to increase.7, 8 Recently, some doubt has been cast on the efficacy of periosteal stripping, as work has suggested the procedure has no effect on long bone growth when compared to the control, non-stripped limb of the same horse.9 Another option for surgical correction of an angular limb deformity is transphyseal bridging.10 This can be performed with transphyseal staples, screws, and wires, or a single transphyseal screw. The physis on the growth-rich side of an affected long bone is bridged with a rigid material to retard growth (Figure 84.3). This restriction of the physis on the longer (convex) side allows the slower side to catch up. Careful observation and follow-up radiographs are required to avoid the risk of overcorrection of the original ALD. If the bridging materials are left in the limb for too long, the limb can overcorrect, causing the opposite ALD to develop. In fact, the
Figure 84.3 A post-surgical dorsal to caudal radiographic projection of a foal with carpal valgus treated with a single transphyseal screw. The screw will need to be removed as angulation correction is achieved.
optimal time to remove the bridging device is just before the limb straightens completely. Surgical success has been documented with angulations up to 29◦ .7 Dependent on timing, physeal manipulation can achieve partial or complete correction with even more severe defects.
Conclusions In conclusion, if ALDs are identified early and classified correctly after a thorough physical examination and imaging, a favorable outcome for athleticism can be achieved for most affected young horses. The prognosis for a successful race career is potentially affected by the presence of ALD in early life and the need for treatment. One study indicated that affected Thoroughbred foals having periosteal stripping surgery did not reach similar numbers of starts or earnings per start as unaffected siblings.11 Similar studies are not available for young horses treated with other techniques or destined for other sporting activity.
References 1. Caron JP. Angular deformities in foals. Equine Vet J 1988;20:225–8.
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Angular Limb Deformities 2. Mitten LA, Bertone AL. Angular limb deformities in foals. J Am Vet Med Assoc 1994;204:717–20. 3. Orsini JA, Kreuder C. Musculoskeletal disorders of the neonate. Vet Clin North Am Equine Pract 1994;10:137– 66. 4. O’Donohue DD, Smith FH, Strickland KL. The incidence of abnormal limb development in the Irish thoroughbred from birth to 18 months. Equine Vet J 1992;24:305–9. 5. Bramlage LR. The science and art of angular limb deformity correction. Equine Vet J 1999;31:182–3. 6. Greet TR, Curtis SJ. Foot management in the foal and weanling. Vet Clin North Am Equine Pract 2003;19:501–17. 7. Bertone AL, Park RD, Turner AS. Periosteal transaction and stripping for treatment of angular limb deformities in foals: radiographic observations. J Am Vet Med Assoc 1985;187:153–6.
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8. Bertone AL, Turner AS, Park RD. Periosteal transaction and stripping for treatment of angular limb deformities in foals: clinical observations. J Am Vet Med Assoc 1985;187:145–52. 9. Read EK, Read MR, Townsend HG, Clark CR, Pharr JW, Wilson DG. Effect of hemi-circumferential periosteal transaction and elevation in foals with experimentally induced angular limb deformities. J Am Vet Med Assoc 2002; 221:536–40. 10. Brauer TS, Booth TS, Riedesel E. Physeal growth retardation leads to correction of intracarpal angular limb deviations as well as physeal valgus deformity. Equine Vet J 1999;31:193–6. 11. Mitten LA, Bramlage LR, Embertson RM. Racing performance after hemicircumferential periosteal transaction for angular limb deformities in thoroughbreds: 199 cases (1987–1989). J Am Vet Med Assoc 1995;207:746–50.
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Chapter 85
Developmental Orthopedic Disease C. Wayne McIlwraith
Introduction The term developmental orthopedic disease (DOD) was coined in 1986 to encompass all orthopedic problems seen in the growing foal1 and has become a generally accepted term. It is a term that encompasses all general growth disturbances of horses and is therefore non-specific. It is felt that one has to be non-specific when talking about the various limb abnormalities of young horses because previous terms such as metabolic bone disease and osteochondrosis implied that they all had a common cause and pathogenesis. While there are similarities in pathogenesis for some entities of DOD, the term should not be used synonymously with osteochondrosis, nor should subchondral cystic lesions, physitis, angular limb deformities, and cervical vertebral malformations to be presumed be manifestations of osteochondrosis (a condition defined below). The spectrum of conditions currently classified as DOD was previously designated ‘metabolic bone disease’.2 However, it is felt that this term is misleading because it refers specifically to bone, whereas many of these problems are seen as problems associated with the articular-epiphyseal cartilage complex. Severe forms of DOD are seen sometimes when there is little or no aberration in bone histomorphometry, implying questionable change in bone metabolism.3 While use of the term ‘metabolic bone disease’ has been defended,4 ‘developmental orthopedic disease’ has become the generally accepted term.
The different entities of DOD When the term DOD was first coined, it was categorized to include the following: (i) osteochondritis dissecans; (ii) subchondral cystic lesions; (iii) angular limb deformities; (iv) physitis; (v) flexural
deformities (these may have no defined cause, or may be secondary to osteochondrosis or physitis); (vi) cuboidal bone abnormalities; and (vii) juvenile osteoarthritis.
Osteochondritis dissecans This condition involves a dissecting lesion of the articular surface of a joint with the formation of a cartilage or cartilage and bone flap.5 Flaps may become detached and form ‘joint mice’. Blood vessels from the periphery of the joint frequently remain in communication with cartilage flaps or detached flakes, leading to calcification or ossification of the separated cartilage. It is commonly considered that release of debris from under the flap causes synovitis (inflammation of the synovial membrane) and pain, but we do not think it is that simple now because we find instances where the cartilage is intact but the horse still has lameness and synovial effusion.6, 7 Osteochondritis dissecans (OCD), as well as subchondral bone cysts, are classically considered to be a manifestation of osteochondrosis5 (Fig. 85.1). The normal epiphyseal ossification center advances out until ossification ceases, leaving a layer of cartilage (this layer of cartilage becomes the articular cartilage). If there is a disturbance in endochondral ossification, an area of retained cartilage can be formed with a consequent defect in the bone (Fig. 85.2). Subsequent cracking or separation in this retained cartilage can lead to the development a flap or fragment of cartilage that may contain bone. These flaps and fragments on the surface of the joint result in the clinical entity of OCD. Although the concept of osteochondrosis (a defect in endochondral ossification) seems to be the pathogenesis in many clinical cases (based on detection
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 85.1 Classical pathway by which osteochondrosis can lead to osteochondritis dissecans or subchondral bone cysts.
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Figure 85.3 Section of medial trochlear ridge of femur demonstrating separation of cartilage of normal thickness from bone that does not have a defect in it (and presumably no obvious defect in endochondral ossification).
of a subchondral bone defect with radiographs and bone disease at arthroscopic surgery), clinical OCD can also develop unassociated with a defect in the bone (and therefore, presumably, no defect of endochondral ossification) (Figures 85.3 and 85.4). It is becoming increasingly common to find elevated articular cartilage at arthroscopic surgery that is of normal thickness and unassociated with a clear defect in the subchondral bone. In addition to being a diagnostic challenge (because of the absence of a de-
fect in the subchondral bone) there is implication of a different pathway because there seems to be a specific structural breakdown between the articular cartilage and bone. Analysis of serum with biomarkers provides evidence for a collagen defect in the pathogenesis of OCD8 and changes in type II collagen have been found in equine osteochondrosis cartilage samples.9 Such a defect could feasibly cause separation of
Figure 85.2 Section through lateral trochlear ridge of femur affected with OCD. This demonstrates the classic role whereby osteochondrosis (defective endochondral ossification) leads to OCD.
Figure 85.4 Arthroscopic view of elevated cartilage on medial trochlear ridge of femur in femoropatellar joint. This cartilage was of normal thickness but has separated from the bone.
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cartilage of normal thickness from the bone because of loss of integrity of the collagen framework.
Subchondral cystic lesions Subchondral cystic lesions occur in various locations, but the most common place is in the medial condyle of the femur in the stifle, followed by the distal metacarpus in the fetlock joint. Subchondral cystic lesions have also been classically represented as a manifestation of osteochondrosis by a number of authors and there is pathological evidence to support this.5, 10 However, research in the author’s laboratory demonstrated that subchondral cystic lesions could develop from defects in the joint surface and in mature animals.11 Also, in other research from our laboratory examination of the contents of these bone cysts after their removal has shown that the linings have high levels of degradative enzymes and can actively resorb bone.12 The same findings probably explain how cysts expand after endochondral ossification has ceased.
Angular limb deformities Angular limb deformities involve deviations from 180◦ associated with the carpus, fetlock or tarsus. The most common one, carpal valgus, is a normal conformation for a foal and retention of a mild degree of carpal valgus in adults is desirable13 (Figure 85.5) On the other hand, if the deformity is in the other direction (carpal varus), this is a severe problem and needs immediate attention. The pathogenesis of angular limb deformities will be discussed in Chapter 84. In a minor number of cases osteochondrosis occurs in the physis and in these instances it is presumed that pathogenetic factors are the same as for osteochondrosis in a joint (discussed below).
Physitis This condition presents as swelling around the growth plates of the distal radius and distal metacarpus. It is usually a clinical diagnosis, but sometimes radiographic lesions with spurring and some lysis at the edge of the growth plate will be seen. In other instances it may be an area of retained cartilage (osteochondrosis) or pathological change more consistent with trauma (Figure 85.6). In some recent work in New Zealand, it has been noted that swelling of the distal metacarpus and distal metatarsus previously considered physitis (of distal MCIII or MTIII) is unassociated with any problems in the physis.14 With such a variation of change in this syndrome, any etiological associations are difficult.
Figure 85.5 Carpal valgus in a foal of varying degrees in each limb, but both considered normal at this stage of development. Correction of the left leg would only be indicated if it still had this angle at 6–9 months of age.
Cuboidal bone abnormalities This problem is generally one of immaturity and is most commonly seen in premature foals or foals with other causes of intrauterine growth retardation. The normal small bones of the carpus (knee) and tarsus (hock) ossify initially as a circle and later form a bony cube. If a foal is born prematurely with angulations of the same joint, wedging of the bone can potentially occur. The occurrence of the condition will appear clinically as an angular limb deformity in the carpus or, in the case of tarsal bone collapse, as a sickle hock conformation. Radiographs confirm the diagnosis. These problems should always be considered in obvious angular limb deformities that are not improving. This condition has also been reported in association with hypothyroidism.15 Flexural deformities Flexural deformities are manifested as increased angulations around a joint as seen from the side. They can be congenital or acquired. Congenital flexural deformities may involve the carpus, fetlock, or distal interphalangeal joint. The pathogenesis is uncertain, but response to oxytetracycline implies excessive
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Figure 85.6 A clinical physitis of carpus (a) and lesions on both sides of the metaphyseal growth plate (physis) at postmortem (b).
muscular contraction and perhaps activity of myofibroblasts within the tendon is involved.16 Acquired flexural deformities occur in the first 18 months of life and involve either a flexural deformity of the distal interphalangeal joint (also called contracture of the deep digital flexor tendon or ‘clubfoot’) that tends to occur at 6 to 12 months16 or a flexural deformity of the fetlock joint (usually metacarpophalangeal joint) which occurs slightly later (Figure 85.7). This has previously been called contracted superficial flexor tendons, but the actual muscle-tendon involvement is more complicated than that. Acquired flexural deformities can occur in association with osteochondrosis and there have been various etiological associations made including high energy diets, mineral imbalances (similar to osteochondrosis), and plant toxicities.16, 17
surveys of horses not necessarily showing clinical signs. This has led to some questionable conclusions, not only regarding the causative factors of DOD but also the effect of treatment or the significance of
Juvenile arthritis Inclusion of juvenile arthritis or juvenile OA in DOD is appropriate.2 The joints most commonly affected are the proximal interphalangeal and distal tarsal joints and there can be overlap with osteochondrosis. Bony fragments of the palmar/plantar surface of the proximal phalanx could also be considered in this category, but their pathogenesis is considered to be traumatic rather than osteochondrosis and the hereditability is very low.10
Defining the incidence of DOD There has been a problem or confusion between reports of clinical instances of DOD and radiographic
Figure 85.7 A flexural deformity of the metacarpophalangeal (fetlock) joint.
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various problems. There are many instances where radiographs on a pre-purchase examination will show OCD, for instance, but that OCD may not be causing clinical problems because the fragment has not separated. It has also been demonstrated that lesions detected radiographically in the hock and stifle can heal spontaneously (based on a 1-year study with monthly radiographs).18 In one radiographic study in Quebec, each horse’s racing performance at 2 years of age was related to the radiographic lesions diagnosed at approximately 17 months of age and before training.19 Although the radiographic lesions did not seem to be associated with poor racing performance, the authors of the study noted the lack of clinical data and the relatively small numbers. What are needed are improved studies in which clinical signs are correlated with radiographs and, possibly more important, all horses are radiographed and then followed so we know how many of those horses with x-ray changes develop clinical problems. This would solve many of the current issues at the yearling sales and one study with 1200 yearlings has been reported.20 Mild to moderate physitis and flexural deformities (concurrent with physitis in most cases) was reported in 88% of 42 weanlings between weeks 6 to 8 of a study looking at the effect of dietary energy and phosphorus on blood chemistry and development of growing horses.21 In these instances, the clinical signs largely resolved on their own by 5 weeks. Dietary treatment did not influence the incidence, nor was it related to daily weight gain. There has been another study defining incidence of DOD in Thoroughbred horses in Ireland over a period of 18 months.22 It was found that angular limb deformities and physitis together constituted 72.9% of the cases treated. It was also noted that 67.7% of the animals showed some evidence of DOD, but only 11.3% were deemed to need treatment. This study points out the need to have definition of what disease process we have. I think it can be seen from these studies that when ‘developmental orthopedic disease’ occurs, it commonly involves angular limb deformities and physitis problems that spontaneously self-correct. It is the ones that often do not self-correct, such as OCD or subchondral bone cysts, that are more critical to us as clinicians. In other words we have DODs of little concern and DODs of major concern.
Osteochondrosis: important clinical causative factors As mentioned previously, most cases of OCD, many cases of subchondral cystic lesions, and some cases of physitis and angular limb deformities are associ-
ated with the pathological process of osteochondrosis. It is probably useful to consider osteochondrosis involving a problem of the articular-epiphyseal cartilage complex and, therefore, susceptibility of the joint surface to injury during development, at least in the cases of OCD and subchondral cystic lesions.23, 24 It has been conventional to use the term ‘osteochondrosis’ to describe the process of abnormal bone and cartilage formation and to use the term ‘osteochondritis dissecans’ to describe the subsequent lesions that penetrate the joint surface creating inflammation and effusion.6, 24 Although there have been arguments that dyschondroplasia may be a more descriptive term because it emphasizes a primary defect in cartilage formation rather than bone, it can be confused with other generalized metabolic conditions, and it has yet to be proven that the clinical entities we see start in cartilage per se. It could well be a critical primary lesion. It is known that endochondral ossification is controlled by a complex interaction of systemic factors, including growth and thyroid hormones, as well as growth factors such as transforming growth factor-beta (TGF-β), insulin-like growth factors 1 and 2 (IGF-1 and -2), fibroblast growth factor (FGF), and bone morphogenic proteins (BMPs) and that parathyroid hormone-related protein (PTHrP) and its receptor has a major coordinating role. Expression of PTHrP is regulated by Indian hedgehog (Ihh) signals through the PTH-PTHnrP receptor.25 Differential and site-specific roles for the cysteine proteinases cathepsin B and L have been demonstrated.26 Various pathogenic processes in dyschondroplasia have been reviewed.25 Other workers have also shown changes in molecular expression of aggrecan and collagen types I, II, and X, IGF-1, TGF-β, and BMPs in articular cartilage obtained from horses with natural acquired osteochondrosis.27, 28 Examination of expression of BMP-2, BMP-6, and a BMP antagonist (Noggin) in equine OCD have shown that BMP-6 induces expression of Ihh, and that BMP-2 stimulates chondrocyte proliferation and hypertrophy and induces apoptosis. Noggin prevents binding of BMPs to receptors, hence a negative feedback on BMP and cartilage differentiation.28 Elevated PTH-rP protein and mRNA expression were identified in the deeper layers of equine osteochondrosis cartilage27 and, more recently, increased expression of Ihh in osteochondrosis suggests a role of Ihh in diseased cartilage and the author suggested that a different primary prescription factor Ihh or the presence of a elevated Ihh inhibitor in these osteochondrotic cartilage samples could be a factor.29 These factors could all be involved in the primary lesion of OCD. Certain etiological factors have been associated with the development of lesions of osteochondrosis.
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Developmental Orthopedic Disease These factors have varied, but the idea that there is a multifactorial etiology has generally been accepted. The information presented here comes from a combination of clinical and pathology reports, as well as experimental studies in the horse. Genetic predisposition Radiographic studies in Swedish Trotters and Warmbloods found thatfor one stallion of each breed there was a significantly high frequency of OCD amongst his progeny, compared with the progeny of the other stallions (P < 0.001).24 In another study in Denmark, radiographic evidence was seen of a significantly high proportion of osteochondrosis in the progeny of one of eight stallions, even though the stallion itself did not show radiographic signs of osteochondrosis.30 A study by Philipsson et al. examined 793 progeny of 24 Swedish Standardbred trotter stallions, and was based on radiographs of the tarsocrural joint for OCD and the fetlock joint for palmar/plantar osteochondral fragments. The study horses were a random sample of about 35 progeny for each of 24 stallions.31, 32 The occurrence of OCD was nearly three times as high among the progeny of stallions affected with OCD in the hock as amongst those of the nonaffected stallions but the difference was much smaller with plantar osteochondral fragments (not considered OCD). The results in all progeny groups showed differences in the incidence of hock OCD in different progeny stallions from 0 to more than 40%, and the overall heritability estimates were 0.27 for tarsocrural OCD and 0.17 for proximal palmar/plantar lesions. The authors also felt that the higher estimate of heritability reported in a Norwegian study33 was due to the high incidence in one particular progeny group (9/13). Even though the defects have proven to be partly under genetic control, breeding policies must also consider the importance of these defects in future soundness and the ability to perform. There has been little work done outside Scandinavia with regard to heredity and we certainly have not been able to develop any type of screening program for osteochondrosis in stallions and mares that will ensure freedom from that condition. However, it would appear likely that there are genetic components to this disease in all breeds. Individual instances of certain stallions and mares producing these individuals have been seen. Growth and body size A high growth rate was first recognized to be associated with a high incidence of osteochondrosis in dogs and pigs.34, 35 Stromberg reported a predominance of osteochondrosis in large-framed and rapidly grow-
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ing horses5 and many others have noted this anecdotally. In a more recent work, the incidence of OCD lesions occurring in either the fetlock, hock, shoulder, or stifle on a commercial thoroughbred farm were studied over a 4-year period.36 All lesions were clinically significant (needed surgery) and OCD was diagnosed in 10% of the 271 foals monitored. Fetlock OCDs tended to occur before 180 days of age, while hock, shoulder, and stifle OCDs occurred around 300–350 days of age. Foals that developed hock OCDs averaged 5 kg heavier than the Kentucky average at 25 days of age and by 240 days these foals were 14 kg heavier than the population average. Foals that developed stifle or shoulder lesions averaged 5.5 kg heavier than the Kentucky average at 25 days of age and 17 kg heavier at 120 days of age. The authors postulated that these types of lesion could have been the result of excessive biomechanical forces exerted on otherwise normal cartilage. Early fetlock OCDs could have resulted from inadequate subchondral bone formation due to restricted activity in foals born early in the year and housed indoors at night. Hock and stifle lesions may have occurred in heavy foals that grew rapidly after weaning.36 One caveat is that this study has yet to be published in a refereed journal.
Mechanical stress and trauma It has long been recognized clinically that mechanical stresses often precipitate the onset of clinical signs with OCD and it is presumed that this is by avulsing the OCD flap or fragment.25 Whether trauma or physical stress is involved in the primary induction of an OCD lesion is more controversial, but it has been suggested that shear forces may disrupt capillaries in the subchondral bone (bone under the cartilage) and give rise to chondrocyte or cartilage cell damage.37 A recent experiment tested this hypothesis. Using an osteotome arthroscopically, a portion of the distal intermediate ridge of tibia and lateral trochlear ridge of femur was fractured off and these lesions examined some months later. The histological appearance of these fragments was quite distinct from that of naturally occurring OCD, arguing that these lesions are not the result of a primary traumatic injury.38
Defects in vascularization OCD was initially attributed to vascular and ischemic necrosis of the subchondral bone.39, 40 Some histological studies of various lesions in the horse have revealed little evidence of primary bone or cartilage and necrosis and subchondral avascularity was not recognized,3, 41 despite work in the pig suggesting a defect in blood supply to the pathogenesis of
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osteochondrosis.42 Hurtig and Pool feel that the idea of direct mechanical shearing of cartilage canals is more likely to account for distribution of lesions in equine osteochondrosis than simple ischemic necrosis of epiphyseal bone.24 They feel that the idea is consistent with some sites in the horse where biomechanical forces are high and the cartilage is normally quite thick, making it more prone to shearing forces. The lateral trochlear ridge of the femur is one such site, where running at speed can create direct compression of the lateral ridge.43
Nutrition Overnutrition as a predisposing cause of osteochondrosis was originally extrapolated from work in dogs44 and pigs.35, 45 A series of controlled experiments showed that high-energy diets of 129% NRC requirements consistently produced lesions of osteochondrosis in weanling foals, compared with a controlled diet based on 100% NRC requirements.46 Clinical and radiographic signs of osteochondrosis were seen in 6/12 foals fed the high-DE (129% NRC levels) diet. None of the foals fed the high-CP or controlled diet showed any clinical or radiographic signs of osteochondrosis. Multiple lesions of osteochondrosis were detected in 11 foals fed the high-DE diet, and in one foal fed the high-CP diet, and in one foal fed the control diet. The number of histological lesions was significantly greater in the high-DE foals than the control group (P < 0.0001). The incidence of lesions in the foals fed high-CP did not differ significantly than that in the control foals (P = 0.11).46 Glade and Belling found that high-energy/protein diets fed to young horses resulted in earlier glucose and insulin peaks and proposed that the intense early insulin secretion and clearance stimulated conversion of thyroxine (T4) to triiodinethyronine (T3).47 Higher T3 concentrations inhibit thyroid-stimulating hormone via negative feedback, so T4 secretion decreases and a periodic postprandial transient thyrotoxemia may be induced. Because final stages of chondrocytic differentiation and collagen and proteoglycan synthesis appear to depend on thyroid hormones, there is the potential for a problem here.
Mineral imbalances Various mineral imbalances have been implicated as causative factors with OCD including excessive dietary phosphorus46 while excessive levels of dietary calcium did not have an effect on OCD incidence.48–51 Hurtig et al. conducted a blind randomized study in Quarter Horse foals with high (25 ppm) and low (7 ppm) copper diets.50 Foals in the low copper group
had declining liver copper values and developed OCD lesions, physitis, and limb deformities over a 6-month period. Erosion and separation of cartilage from the subchondral bone in the dorsal facets of the cervical spine was the most consistent postmortem lesion (it was not possible to diagnose these lesions clinically) and foals fed the high copper diet had no significant musculoskeletal lesions. Biochemical assays showed significantly reduced collagen, pyridinoline cross-linkages in articular cartilage, growth plate cartilage, and bone in the OCD-affected foals and the results suggested that the concept of cartilage thickening was a myth, at least in the growth plate, and that the defect was in the transition between calcified cartilage and primary spongiosa.52 They interpreted the lesion as one of reduced structural strength rather than arrested or abnormal endochondral ossification, that there is biomechanically inferior connective tissue rather than failure of chondrocyte differentiation and provisional calcification, and that this defect could be associated with defective collagen cross-linking and altered matrix remodeling. In another study in New Zealand, the authors reported a low incidence and severity of lesions in their foals on low copper diets and suggested that there is a greater dietary copper requirement in America compared with that of pasture-fed foals in New Zealand.53–56 Copper supplementation of the mare, however, improved radiographic indices of physitis in foals at 150 days and it was suggested that copper supplementation of mares could provide a convenient treatment regime on farms experiencing an increased incidence of DOD.54 Cymbaluk and Smart57 have cautioned that equine veterinarians and owners could be overemphasizing copper intake to the exclusion of adequate energy, protein, and macromineral intakes. Although there is evidence that copper may control some aspects of osteochondrosis, attention needs to be paid to generally sound nutrition or management and not presume that copper supplementation will cure this multifactorial problem.57, 58 Because zinc appears to be the main copper antagonist for horses, dietary copper supplementation must always consider zinc intake. Zinc to copper ratios of 4:1 or 5:1 for all breeds of horse have been recommended. Excessive zinc intake has also been related directly to equine osteochondrosis. Environmental exposure to zinc and cadmium in pregnant pony mares have been associated with observations of lameness, swollen joints, and ill-thrift, particularly in foals.44 Endocrine factors Endocrine causes have been implicated in the pathogenesis and it has been postulated by one investigator that the production of osteochondrosis
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Developmental Orthopedic Disease lesions in association with overfeeding is mediated by the endocrine system.59 Certainly the longterm administration of dexamethasone has produced osteochondrosis-type lesions and it is considered that glucocorticoids induced a parathyroid hormone resistance at the level of the osteocyte causing an inhibition of normal remodeling. Glucocorticoids also induced decreased glucosaminoglycan (GAG) levels and this decrease in turn inhibits capillary penetration of the cartilage, which is a very important step in forming bone from cartilage. The failure of ossification could also be mediated through induced defects in vitamin D metabolism. In addition to 1,25OH2 (D3) promoting the availability of both calcium and phosphate, it has been shown that the other active vitamin D metabolite [24,25-OH2 (D3)] promotes the synthesis of proteoglycan by chondrocytes and is necessary for the normal differentiation of epiphyseal cartilage.60 It has also been pointed out that corticosteroids inhibit lysyl oxidase and lysyl oxidase is involved in cartilage cross-linking. As noted previously Glade and Belling suggested that the episodic insulin secretion and thyroid hormone abnormalities produced by high-soluble-carbohydrate diets could cause osteochondrosis.47 Site vulnerability Because the lesions of equine osteochondrosis occur at specific anatomic sites, this obviously suggests site vulnerability. This predilection could be related to an ossification defect or trauma caused by excessive stress in that region. In nearly all instances, the sites of occurrence of OCD are very close to the limits of articulation. Differences in GAG content have been demonstrated between articular and non-articular lesions in canine shoulders61 and, also, GAG contents can be modulated by changing a non-articular into an articular surface by making the dog step up onto a race surface. OCD lesions are frequently bilateral in the stifle and hock and quadrilateral in the fetlock joint, while they frequently involve different joints in the same animal. It is felt this may suggest a ‘window of vulnerability’ in the endochondral ossification of that specific joint when an environmental insult may have occurred.37 If the causative factor were present intermittently or for a transient period during the foal’s growth period, this would explain the development of the disease in only one pair of joints. It is not possible from this data to ascertain different periods of onset of the disease process in different joints. Exercise (or lack thereof) Adequate exercise in foals would logically be important to maintain cartilage and bone quality. There
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was a report in 1994 claiming a reduction of 20% to 6% in the prevalence of osteochondrosis in the hock due to an exercise regimen.62 This study prompted Jeffcott63 to suggest that exercise should be investigated. A study on the effect of exercise on distribution and manifestation of osteochondrosis in the Warmblood foal was published in 1999.64 Forty-three foals, all from sires having osteochondrosis in either the femoropatellar or tibiotarsal joint, were reared to weaning at 5 months, under similar conditions except for the type and amount of exercise. Fifteen foals remained at pasture, 14 were kept in box stalls, and 14 foals were kept in the same box stalls but daily given an increased number of gallop sprints. After weaning, 8 foals from each group were euthanized and the remaining 19 animals were given an identical light exercise regime for an additional 6 months to euthanasia at 11 months when all major diarthrodial joints were inspected and histology performed. Exercise did not significantly influence the number of lesions, but at 5 months there was a tendency for more severe lesions in the box-rested foals. In the femoropatellar/femorotibial joint, lesions were found mainly in the femoral condyles of the boxrested foals and the lateral trochlear ridge of the femur in the trained foals. The authors also pointed out how many lesions regress and do not become clinically evident, making the number of horses with lesions at mature age into an underestimation of the total amount animals that have suffered from the conditions. The period during which a lesion develops and possibly regresses is limited and variable per site and, while exercise at the level given in this study may have had some influence on the appearance and distribution of the lesions, it did not appear to have an etiological role.
References 1. McIlwraith CW. (ed.) What is developmental orthopedic disease, osteochondrosis, osteochondritis, and metabolic bone disease? In: Proceedings of meeting of the American Association of Equine Practitioners, 1993;39:35–44. 2. Gabel AA, Knight OA, Reed SM, Pultz JA, Powers JD, Bramlage LR, Tyznik WJ. Comparison of incidence and severity of developmental orthopedic disease on 17 farms before and after adjustment of ration. In: Proceedings of the 33rd Annual Convention of the American Association of Equine Practitioners, New Orleans, LA, 1987; pp. 163– 70. 3. Reino S, Stromberg B. Osteochondrosis in the horse. II. Pathology. Acta Radiol Suppl 1978;358:153–78. 4. Gabel AA. Metabolic bone disease: Problems in terminology. Equine Vet J 1988;20:4–6. 5. Stromberg J. A review of the salient features of osteochondrosis in the horse. Equine Vet J 1979;11:211–14. 6. McIlwraith CW. Inferences from referred clinical cases of osteochondrosis dissecans. Equine Vet J 1993;S16:27–30.
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Foal: Problems of Older Foals, Weanlings and Yearlings
7. McIntosh SC, McIlwraith CW. Natural history of femoropatellar osteochondrosis in three crops of Thoroughbreds. Equine Vet J 1993;S16: 54–61. 8. Billinghurst RC, Brama PHA, van Weeran R, McIlwraith CW. Evaluation of serum concentrations of biomarkers of skeletal metabolism and results of radiography as indicators of severity of osteochondrosis in foals. Am J Vet Res 2004;65:143–50. 9. Laverty S, Ionescu M, Marcoux M, Bour´e L, Doize B, Poole AR. Alterations in cartilage type-II procollagen and aggrecan content in synovial fluid in equine osteochondrosis. J Orthop Res 2000;18:399–405. 10. Sandgren B. Bony fragments in the tarsocrural and metacarpophalangeal or metatarsophalangeal joints in the Standardbred horse: A radiographic survey. Equine Vet J 1988;S6:67–70. 11. Ray CS, Baxter GM, McIlwraith CW, Trotter GW, Powers BE, Park RD, Steyn PF. Development of subchondral cystic lesions after articular cartilage and subchondral bone damage in young horses. Equine Vet J 1996;28:225–32. 12. von Rechenberg B, Guenther H, McIlwraith CW, Leutenegger RC, Frisbie DD, Akens MK, Auer JA. Fibrous tissue of subchondral cystic lesions in horses produce local mediators and neutral metalloproteinases and cause bone resorption in vitro. Vet Surg 2000;29:420–9. 13. Anderson TA, McIlwraith CW, Douay P. The role of conformation in musculoskeletal problems in the racing Thoroughbred. Equine Vet J 2004;36:571–5. 14. Gee E, PhD Thesis, Massey University, New Zealand, 2004. 15. McLaughlin PG, Doige CE. Study of ossification of carpal and tarsal bones in normal and hypothyroid foals. Can Vet J 1982;23:164–8. 16. McIlwraith CW. Diseases of joints, tendons, ligaments, and related structures. In: Stashak TS (ed.) Adams Lameness in Horses, 5th edn. Lippincott, Williams & Wilkins, 2002; pp. 459–644. 17. McIlwraith CW, James LF. Limb deformities in foals associated with ingestion of loco weed by mares. J Am Vet Med Assoc 1982;181:255–8. 18. Dik KJ, Enzerink E, van Weeran. Radiologic development of osteochondral abnormalities in the hock and stifle of Dutch Warmblood foals, from age 1 to 11 months. Equine Vet J Suppl 1999;131:9–15. 19. Alvarado AF, Marcoux M, Berton L. The incidence of osteochondrosis on a Standardbred breeding farm in Quebec. In: Annual Convention of the American Association of Equine Practitioners, 1989; 35: 293–307. 20. Kane AJ, McIlwraith CW, Park RD, Rantanan NW, Morehead JP, Bramlage LR. Radiographic changes in Thoroughbred yearlings: Part II: Association with racing performance. Equine Vet J 2003;35:354–65. 21. Cymbaluk NF, Christison GI. Effects of dietary energy and phosphorus content on blood chemistry and development of growing horses. J Anim Sci 1989,67;951–8. 22. O’Donaghue DD, Smith EH, Strickland KL. The incidence of abnormal limb development in the Irish Thoroughbred from birth to 18 months. Equine Vet J 1992;24: 305–9. 23. Bramlage LR. Investigation of farm-wide incidence of bone formation problems in the horse. In: 39th Annual Convention of the American Association of Equine Practitioners, 1993; pp. 45–8. 24. Hurtig MB, Pool RR. Pathogenesis of equine osteochondrosis. In: McIlwraith CW, Trotter GW (eds) Joint Disease in the Horse. Philadelphia: Saunders, 1996; pp. 335–7.
25. McIlwraith CW. Osteochondrosis and more ideas on an enzymatic pathogenesis. Equine Vet J 2003;35:7–8. 26. Glaser KE, Davies ME, Jeffcott LB. Differential distribution of cathepsins B and L in articular cartilage during skeletal development in the horse. Equine Vet J 2003;35:42–7. 27. Semevolos SA, Nixon Brower-Toland. Changes in molecular expression of aggrecan and collagen types I, II and X, insulin-like growth factor I, and transforming growth factor-beta 1 in articular cartilage obtained from horses with naturally acquired osteochondrosis. Am J Vet Res 2001;62:985–1180. 28. Semevolos SA, Nixon AJ, Strausheim ML. Expression of bone morphogenetic protein-6 and -2 and a bone morphogenetic protein antagonist in horses with naturally acquired osteochondrosis. AJVR 2004;65:110–15. 29. Semevolos SA, Strassheim ML, Haupt JL, Nixon AJ. Expression patterns of hedgehog signaling peptides in naturally acquired equine osteochondrosis. J Orthop Res 2005;23:1152–9. 30. Schoougaard H, Falk-Ronne J, Phillipson J. A radiographic survey of tibiotarsal osteochondrosis in a selected population of trotting horses in Denmark and its possible genetic significance. Equine Vet J 1990;22:288–9. 31. Philipsson J. Pathogenesis of osteochondrosis: Genetic implications. In: McIlwraith CW, Trotter GW (eds) Joint Disease in the Horse. Philadelphia: Saunders, 1996; pp. 359–62. 32. Philipsson J, Andreasson E, Sandgren B, Dalin G, Carlsten J. Osteochondrosis in the tarsocrural joint and osteochondral fragments in the fetlock joints in Standardbred trotters. II. Heritability. Equine Vet J 1993;S16:38–41. 33. Grondahl AM, Dolvik NI. Heritability estimations of osteochondrosis in the tibiotarsal joint and of bony fragments in the palmar/plantar portion of the metacarpophalangeal and metatarsophalangeal joints of horses. J Am Vet Med Assoc 1993;203:101–4. 34. Hedhammar A, Wu F, Krook L, Schryver HF, De Lahunta A, Whalen JP, Kallfez FA, Nunez EA, Hintz HF, Sheffy BE, Ryan GD. Overnutrition and skeletal disease: An experimental study in growing Great Dane dogs. Cornell Vet 1972;(Suppl)5:1. 35. Reiland S, Ordell N, Lundeheim N, Olsson SE. Heredity of osteochondrosis, body constitution and leg weakness in pigs. Acta Radiol Suppl 1978;358:123–37. 36. Pagan JD, Jackson SG. The incidence of developmental orthopedic disease on a Kentucky Thoroughbred farm. World Equi Vet Rev 1996;20–26. 37. Pool RR. Pathologic manifestations of osteochondrosis. In: McIlwraith CW (ed.) Proceedings of Developmental Orthopedic Disease Symposium. Amarillo, TX: American Quarter Horse Association, 1986; pp. 3–7. 38. Bertone AL, Bramlage LR, McIlwraith CW, Malemud CL. Comparison of articular cartilage proteoglycon and collagen in equine dyschondroplasia and healing experimentally. Induced osteochondral fragments. Am J. Vet Res 2005;66:1881–1890. 39. Adams OR. Lameness in Horses, 3rd edn. Philadelphia: Lea & Febiger, 1974. 40. Schebitz H. Degenerative arthritis of the shoulder joint following aseptic necrosis of the humeral head in foals. In: 11th Annual Convention of the American Association of Equine Practitioners, 1965; pp. 359–70. 41. Meagher DM, Pool RR, O’Brien TR. Osteochondritis of the shoulder joint in the horse. In: Proceedings of 19th Annual Meeting of the American Association of Equine Practitioners, Atlanta, 1973;247–56.
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Developmental Orthopedic Disease 42. Carlson CS, Meuten DJ, Richardson DC. Ischemic necrosis of cartilage in spontaneous and experimental lesions of osteochondrosis. J Orthop Res 1991;9:317–29. 43. Maquet P. Biomechanique du genou et gonarthrose. Rev Med Liege 1969;24:170. 44. Gunson DE, Kowalczyk OF, Shoop CR, Ramberg CF. Environmental zinc and cadmium pollution associated with generalized osteochondrosis, osteoporosis and nephrocalcinosis in horses. J Am Vet Med Assoc 1982;180:295–9. 45. Reiland S. Morphology of osteochondrosis and sequelae in the pig. Acta Radiol Suppl 1978;358:45–90. 46. Savage CJ, McCarthy RN, Jeffcott LB. Effects of dietary energy and protein on induction of dyschondroplasia in foals. Equine Vet J 1993;516:74–9. 47. Glade MJ, Belling TH. A dietary etiology for osteochondrotic cartilage. J Equine Vet Sci 1986;6:151–5. 48. Bridges CH, Harris ED. Experimentally induced cartilaginous fractures (osteochondritis dissecans) in foals fed lowcopper diets. J Am Vet Med Assoc 1988;193:215–21. 49. Bridges CH, Womack JE, Harris ED, Scrutchfield WL. Considerations of copper metabolism in osteochondrosis of suckling foals. J Am Vet Med Assoc 1984;185:173–8. 50. Hurtig MB, Green SL, Dobson H, Burton J. Defective bone and cartilage in foals fed a low copper diet. In: Proceedings of the 35th Annual Convention of the American Association of Equine Practitioners, Lexington, KY, 1990; pp. 637–43. 51. Knight DA. Gabel AA, Reed SM, Embertson RM, Bramlage LR, Tyznik WI. Correlation of dietary minerals to incidence and severity of metabolic bone disease in Ohio and Kentucky. In: Proceedings of 31st Annual Meeting of the American Association of Equine Practitioners, Toronto, Canada, 1985; pp. 445–561. 52. Hurtig MB, Green SL, Dobson H, Burten J. Correlation of the clinical signs, diagnostic imaging and gross pathology of defective cartilage and bone growth in copper-deficient foals [abstract]. In: Proceedings of Equine Osteochondrosis in the 90’s, Cambridge University, 1992; pp. 27–8.
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53. Pearce SG, Firth EC, Grace ND, Fennessy PF. Effect of copper supplementation on the evidence of developmental orthopaedic disease in pasture-fed New Zealand Thoroughbreds. Equine Vet J 1998;30:211–18. 54. Pearce SG, Grace ND, Firth EC, Wichtel JJ, Holle SA, Fennessy PF. Effect of copper supplementation on the copper status of pasture-fed young Thoroughbreds. Equine Vet J 1998;30:204–10. 55. Pearce SG, Grace ND, Wichtel JJ, Firth EC, Fennessy PF. Effect of copper supplementation on copper status of pregnant mares and foals. Equine Vet J 1998;30:200–3. 56. Petersson H, Sevelius F. Subchondral bone cysts in the horse: a clinical study. Equine Vet J 1986;1:75. 57. Cymbaluk NF, Smart ME. A review of possible metabolic relationships of copper to equine bone disease. Equine Vet J 1993;S16:19–26. 58. Jeffcott LB, Davies ME. Copper status and skeletal development in horses: Still a long way to go. Equine Vet J 1998;30:183–5. 59. Hintz HF. Factors which influence developmental orthopedic disease. In: Proceedings of the 33rd Annual Convention of the American Association of Equine Practitioners, New Orleans LA, 1987; pp. 159–62. 60. Kanis JA. Vitamin D metabolism and its clinical application. J Bone Joint Surg [Br] 1982;64:542. 61. Kincaid SA, Van Sickle DC. Regional histochemical and thickness variations of adult articular cartilage. J Am Vet Med Assoc 1981;42:428. 62. Bruin G, Creemers JJHM, Smolders EEA. Effect of exercise on osteochondrosis in the horse [abstract]. In: Proceedings of Equine Osteochondrosis in the 90’s, Cambridge University, 1992; pp. 41–2. 63. Jeffcott LB. Problems and pointers in equine osteochondrosis. Equine Vet J 1993;S16:1–3. 64. van Weeran PR, Barneveld A. The effect of exercise on the distribution and manifestation of osteochondral lesions in the Warmblood foal. Equine Vet J Suppl 1999;31:16–25.
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Chapter 86
Feeding the Growing Horse to Avoid Developmental Orthopedic Disease (DOD) Paul D. Siciliano
Introduction Diet is one of several factors potentially contributing to developmental orthopedic disease (DOD). Effective prevention strategies must consider all contributing factors (e.g., biomechanical, genetic, diet, etc.). The relationship between diet and DOD is not always clear, and many unanswered questions surrounding the relationship exist. The complexities of these relationships make it a challenging area of study. Nonetheless there are several dietary components for which at least some relationship with DOD has been demonstrated. The purpose of this chapter is to provide current knowledge regarding selected nutrient requirements of growing horses, to discuss various relationships between diet and DOD, and also feeding strategies for its prevention. For a comprehensive discussion of growing horse nutrient requirements the reader is referred to the National Research Council’s (NRC’s) Nutrient Requirements of Horses (6th edition).1
Effect of bodyweight gain on DOD It has been suggested that a relative high overall nutritional plane facilitating accelerated bodyweight gain impacts negatively on bone health in dogs2–4 and pigs.3 Evidence in dogs suggests that rapid growth, initiated by increased nutritional plane, is associated with enlarged bone organs formed of cancellous and compact bone having relatively low density and consequently lowered resistance to biomechanical loading.4 These findings in other species led to hypotheses that high nutritional plane
may negatively impact bone growth and development and potentially contribute to DOD in horses. Relationships between accelerated bodyweight gain and DOD have been reported in horses;5–7 however, accelerated bodyweight gain is not always related to the incidence and development of DOD.8 The results from studies evaluating the direct effect of nutritional plane on bone growth and development suggest that accelerated bodyweight gain alone is not sufficient to cause DOD, but that other factors such as compensatory growth and absolute bodyweight may play a role. Hintz et al.9 reported that four of six Standardbred weanlings fed diets aimed at restricting average daily gain to 0.3 kg/d or less for 4 months followed by ad libitum feeding of the same diet for an additional 4 months led to compensatory bodyweight gain and fetlock flexure deformities. Interestingly the rate of bodyweight gain following the switch from restricted to ad libitum was not different from a control group of weanlings fed the ad libitum diet for the entire 8-month experiment. However, the rate of bodyweight gain for the second 4-month period when weanlings were fed the ad libitum diet was double that of the first 4-month period when they were fed the restricted diet. The fetlock flexure deformities self-corrected toward the end of the experiment, coinciding with a time of increased daily exercise. This finding suggests that an abrupt change in daily bodyweight gain may be a more important factor in the precipitation of flexure deformities than the absolute rate of daily bodyweight gain. Average daily bodyweight gain (ADG) did not appear negatively to affect bone mineral density and bone mineral content of selected regions of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Feeding the Growing Horse to Avoid Developmental Orthopedic Disease (DOD) the appendicular skeleton in weanlings fed two different balanced rations formulated to achieve 0.91 or 0.34 kg bodyweight gain/d.10 These average daily bodyweight gains were considered slightly above rapid ADG (0.85 kg/d) and below moderate ADG (0.65 kg/d) as defined by the NRC’s Nutrient Requirements of Horses (5th edition).11 Although ADG did not appear negatively to impact bone growth and development, it was noted that three of the six horses fed to gain 0.91 kg/d developed physitis and mild flexure deformities of the carpus and metacarpophalangeal joints following approximately 50 to 56 days of dietary treatment, but were resolved by day 75 of dietary treatment. Three horses showing clinical signs of DOD were approximately 20 kg heavier than those without clinical signs, but average daily gain was similar between the two groups. This may suggest that absolute bodyweight can be a factor in the development of DOD. No clinical signs of DOD were reported in the weanlings fed balanced rations formulated to achieve 0.34 kg daily bodyweight gain.
Digestible energy Metabolic roles, dietary sources, and requirements Energy is required for body temperature maintenance, biomolecule synthesis and degradation, ion gradient maintenance, and muscle contraction. Adenosine triphosphate (ATP) is the common energy currency used by cells and is synthesized during the metabolism of carbohydrates, lipids, and amino acids. The primary carbohydrate metabolized for ATP is glucose. Glucose originates from dietary starch contained in cereal grains, through metabolism of the volatile fatty acid propionate produced by microbial fermentation of dietary carbohydrates in the gut, and by metabolism of some amino acids (i.e., glucogenic amino acids). The primary lipids metabolized for energy include fatty acids originating from dietary triglyceride or that stored in the body; and volatile fatty acids (propionate, acetate, and butyrate) originating from microbial fermentation of carbohydrate in the horse’s gut. Amino acids, originating from dietary protein, in excess of those required for protein synthesis, can be metabolized for energy as well. Energy requirements for horses are expressed in units of megacalories (Mcal; 106 calories) or megajoules (MJ; 4.184 joules = 1 calorie) of digestible energy (DE). Digestible energy is the total caloric content of the feed (i.e., intake energy or gross energy) minus energy lost in feces. The digestible energy fraction contains energy that is useful to the horse (i.e.,
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net energy for maintenance and production), energy ultimately lost in gas and urine, and energy lost from the body as heat in association with ingestion, digestion, absorption, transport, and storage of energycontaining compounds (i.e., heat increment). Therefore, only a fraction of the total DE is used for ATP synthesis. In addition to the DE system a net energy (NE) system for expressing requirements and feed energy density has been developed and employed by the French.12 The NE system attempts to estimate more accurately the ‘useful’ energy present in feeds and may have an advantage over the DE system in this regard. NE values for a broad range of feedstuffs throughout the world have not been defined. In contrast, several equations have been derived to predict the DE value of a range of different feeds based on their chemical composition.1 Therefore the DE system is used more extensively throughout the world and will be emphasized in this chapter. Digestible energy requirements for growing horses consist of two main components: maintenance and tissue growth. The maintenance and tissue growth components are estimated using Equations 1.1 and 1.2: DEMaintenance (Mcal/kg BW/d) = 0.0565x −0.145 DETissue growth (Mcal/kg BW gain/d) = 1.99 + 1.21x − 0.021x 2 where x is age in months. These two equations illustrate that total DE required per day for growing horses is a function of age, bodyweight, and bodyweight gain. The total DE requirement for growing horses (maintenance + tissue growth) expressed as Mcal/d increases with age due to the net effect of changes in bodyweight, age, and bodyweight gain. Therefore a horse’s daily DE intake must change periodically to facilitate growth to maturity. Digestible energy intake is increased by increasing dry matter (DM) intake, increasing dietary DE concentration, or a combination of both. Dry matter intake can be increased to a maximum of approximately 2.5% of bodyweight in growing horses. Increasing dietary DE concentration is achieved by increasing the ratio of energy concentrates (e.g., cereal grains and fat) to roughage (e.g., forage) or by selecting energy concentrates and forage having a high caloric density. Therefore, the maximum increase in daily DE intake is constrained by DM intake and dietary DE concentration. Much of the age-related increase in daily DE intake can be achieved by increasing DM intake because DM intake increases with increasing
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Table 86.1 Changes in digestible energy (DE) requirement, dry matter (DM) intake, and ration energy density with age
Age (months) 4 6 8 10 12 14 16 18 24 a
Bodyweight (kg)
Bodyweight gain (kg/day)
DE requirement (Mcal/day)
DM intake (kg/day)a
DE requirement (Mcal/kg DM)
168 216 257 291 321 347 369 387 429
0.84 0.72 0.62 0.53 0.45 0.39 0.33 0.29 0.18
13.3 15.5 17.1 18.1 18.9 19.1 19.3 19.3 18.7
4.2 5.4 6.4 7.3 8.0 8.7 9.2 9.7 10.7
3.2 2.9 2.7 2.5 2.4 2.2 2.1 2.0 1.7
Assuming dry matter intake = 2.5% of bodyweight.
bodyweight, and thus age. Although daily DE requirement also increases with age, the magnitude of DM intake increase is greater than the daily DE requirement increase and has a net effect of decreasing required dietary DE concentration (i.e., Mcal DE/kg DM). This is illustrated in Table 86.1, showing the change in DE requirements and DM intake with age, of a weanling having a 500-kg mature bodyweight. Growing horses consume approximately 2.5% of their bodyweight per day in dry matter, which means that the required DE concentration ranges from 1.7 to 3.2 Mcal/kg DM (see Table 86.1). Forage DE content varies considerably with degree of maturity and forage type (1.8 to 2.9 Mcal/kg DM). Therefore, a forage-only ration may not meet the daily DE requirement within DM intake constraints, especially for weanlings (i.e., less than 12 months of age). Concentrated forms of DE are usually incorporated into weanling rations to complement forage during this phase of growth. The same is true for yearlings when an accelerated rate of growth is desired (e.g., sales yearlings). The DE content of most commercially formulated grain-mix-concentrates for growing horses ranges from 3.2 to 3.6 Mcal/kg DM. Grain-mix-concentrates containing added fat sources (e.g., vegetable oils, rice bran) have a slightly higher DE depending upon the amount of fat added. Environment and exercise are two additional factors that can influence daily DE requirements of growing horses. Energy required for maintenance increases as the environmental temperature decreases below the lower critical temperature of the thermoneutral zone.13, 14 At environmental temperatures below the lower critical temperature more energy is lost from the body as heat and energy metabolism must increase to replace that heat and maintain body temperature. Failure to provide additional calories during cold weather can lead to repartitioning of energy away from tissue growth toward maintenance needs. The net result is an attenuated growth rate.
The increased caloric demand associated with cold temperature appears to be short lived, potentially due to physiological changes associated with cold adaptation (e.g., hair growth).1 Exercise increases the daily DE requirement by 20–40% of the maintenance requirements depending upon the level of exercise.1 Similar to environmental temperature, a failure to provide calories adequate for supporting work will lead to a partitioning of calories toward work and away from tissue growth resulting in attenuated growth rate. Digestible energy intake and DOD Caloric intake above and below recommended daily DE intakes can influence skeletal growth and development and may contribute to DOD. Dietary calorie source (e.g., starch, fat, fiber) can influence skeletal growth and development; although its relationship with DOD is less clear. Growth plate morphology was altered in weanling Thoroughbred horses fed 20.75 versus 15.96 Mcal DE/250 kg BW (130% and 100% of NRC requirements,15 respectively), and the morphological alterations were similar to those associated with osteochondrosis in horses affected by equine hypothyroidism.16 Chondrocyte maturation during endochondrial ossification is dependent upon thyroid hormone. Serum thyroxine (T4 ) concentrations were transiently decreased in association with increased serum insulin concentration as a result of increased starch intake, which was proposed as the potential mechanism by which increased caloric intake might contribute to altered growth plate morphology.17 In a follow-up study a similar effect of starch intake on serum T4 concentration was reported following a single meal, but when the same protocol was repeated 6 months later there was no effect of starch intake on serum T4 concentration.18 Associations between osteochondrosis and post-prandial
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Feeding the Growing Horse to Avoid Developmental Orthopedic Disease (DOD) hyperinsulinemia/hyperglycemia have been reported in horses, but definitive cause and effect relationships have not been established.19–21 Glade and Belling16 also reported that growth plate morphology in a third group of horses fed 11.17 Mcal DE/250 kg BW (70% of requirements15 ) was considered normal, but matrix production, indicators of growth plate metabolism, and foreleg growth rate were decreased indicating impaired endochondrial ossification due to inadequate caloric intake. This finding emphasizes the point that both nutritional deficiency and excess can alter bone growth and development and potentially contribute to DOD. Dyschondroplasia incidence (assessed by clinical signs, radiographs, postmortem examination, and histological assessment) was greater in weanlings of light horse breeding fed 19.7 versus 15.3 Mcal DE/d (approximate average bodyweight = 216 kg).8 The dietary DE content was increased by adding corn oil to a rice-based pellet, which does not support the insulin-mediated mechanism proposed by Glade et al.17 as corn oil would not contribute to a post-prandial serum insulin concentration increase. Additional research by others also failed to support the insulin-mediated mechanism proposed by Glade et al.17 Radiographic osteochondritic lesion incidence and bone mineral deposition were not different between weanlings fed diets containing 11% to 24% starch and providing a total caloric intake of approximately 17 Mcal/d (average bodyweight = 239 ± 8 kg), and radiographic osteochondritis dissecans (OCD) lesion incidence decreased by approximately 29% from start to finish of the 112-day study.22 Hoffman et al.23 also studied the effect of carbohydrate source on bone growth and development and reported that weanlings fed approximately 2 to 3 kg/d of a high-fat high-fiber (26.5% non-structural carbohydrates) supplement had significantly lower (∼ 7%) third metacarpal bone mineral content (radiographic estimate) as compared to those fed similar amounts of a high-starch high-sugar (62.4% nonstructural carbohydrates) supplement. The total diet of both groups of weanlings was intended to meet or exceed 140% for all dietary requirements.11 Although third metacarpal bone mineral content was lower in weanlings fed the fat and fiber supplement, the incidence and severity of physitis, joint effusion, and flexural and angular limb deformities was relatively low and was unaffected by dietary carbohydrate source (fat and fiber vs starch and sugar). Carbohydrate source (approximately 75:25 vs 33:67 forage to concentrate ratio) did not influence weanling physitis or flexural deformity incidence.24 Mild to moderate physitis was reported in 88% of the weanlings, but occurred transiently between weeks 6 and 8 of the study when the horses were ap-
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proximately 7 months of age. Daily caloric intake (∼ 21 Mcal/d) in both weanling groups was 36% above required daily intake.15 Radiographic and growth characteristics of the third metacarpal bone have been shown to be influenced by the level of caloric intake. Thompson et al.25 reported that 4-month-old ponies consuming diets giving them digestible energy and crude protein intakes of 150% and 275%, respectively, above requirements,15 independent of other nutrients, led to increased skeletal growth rate, greater values for third metacarpal bone breaking strength (56% greater than controls), but lower values for percentage of calcium (4% less than controls) and total bone mineral content (4% less than controls). Based on these findings, the authors suggested during accelerated growth, facilitated by increased DE and crude protein intake, bone mineral deposition may be reduced. However, in follow-up studies using horses (suckling foals and weanlings) and a similar experimental protocol the authors reported that the increased DE and/or CP did not significantly affect bone quality as measured by third metacarpal radiographic bone mineral density, cortical width, and cortical area.26, 27 However, based on numerically, but not statistically significant, lower values for third metacarpal bone percent cortical area in weanlings fed diets containing a low protein:calorie ratio (i.e., DE was increased independent of CP) the authors suggested that protein intake needs to be increased proportionally to energy intake to maintain skeletal quality during rapid periods of growth.27 When osteochondrosis incidence and severity was used as an end point in exercised (35 min circular trotting, 5 d/week) weanlings fed diets providing either 110% or 135% of digestible energy and crude protein requirements,11 there was no difference between the two groups.28 The severity, but not the incidence, of osteochondrosis in these two groups of exercised weanlings tended to be less, 47% and 59% less, than that of a third group receiving no forced exercise and fed a diet supplying 110% of requirements,11 suggesting that exercise may have a greater impact on the severity of DOD than dietary imbalance of digestible energy and crude protein intake. In conclusion, there is evidence to suggest that increasing DE intake independent of other nutrients can contribute to dyschondroplasia in weanling horses, yet the mechanism by which this occurs has not been defined. Increasing DE intake, independent of other nutrients, appears to have a minimal effect on radiographic parameters intended to reflect third metacarpal bone quality. Alternatively, decreasing DE (in conjunction with crude protein), relative to other nutrients, may negatively affect endochondrial ossification and subsequent long bone growth. The effect of calorie source (starch vs fat and fiber) on
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bone growth and development is unclear. Given these findings it would seem prudent to insure that DE intake is balanced relative to other nutrients, which can be accomplished by following the current DE recommendations,1 which coincidentally are similar to the caloric intake of control horses showing no indication of dyschondroplasia in the study conducted by Savage et al.8
Crude protein Metabolic roles, dietary sources, and requirements Proteins play several important roles in the body. Protein is a primary structural component of the body; in addition proteins play regulatory (e.g., enzymes) and physiological (e.g., some hormones) roles. Protein in excess of that required for structural, regulatory, and physiological roles can be degraded to amino acids, which are further metabolized for ATP synthesis. Proteins in the diet vary in concentration and in their amino acid profile. For example, some cereal grains such as corn contain 9% crude protein while soybean meal contains 50%. Forage crude protein concentration is inversely related to forage maturity. As the plant matures, crude protein concentration decreases. For a given level of maturity, grasses have lower crude protein content than legumes (e.g., alfalfa and clover). As a result forage crude protein can range from as low as 5% to over 20% crude protein depending upon plant type and level of maturity. The amino acid profile of dietary protein also varies. Lysine is the first limiting amino acid for growth. Even though two protein sources may be similar in crude protein they can differ in lysine content. The comparison of soybean meal (50% crude protein, 3.1% lysine) to cottonseed meal (45% crude protein, 1.85% lysine) illustrates this point. Failure to meet the lysine requirement, even if the crude protein requirement is met, results in a reduced growth rate.1 Protein requirements are expressed using the crude protein system. Additionally, growing horses have a daily lysine requirement. Dietary crude protein requirements consist of a maintenance requirement plus that for growth, and are estimated as follows: g crude protein/dmaintenance requirement = BW × 1.44 g crude protein/kg g crude protein/dgrowth requirement = [(ADG × 1000 × 0.20)/E]/0.79 where BW is bodyweight in kg, ADG is average daily gain in kg/d, and E is efficiency of dietary protein
use, which is estimated to be 0.50, 0.45, 0.40, 0.35, and 0.30 for horses 4 to 6, 7 to 8, 9 to 10, 11, and 12 and older months of age, respectively.1 Lysine requirements are estimated as a percentage of the daily crude protein requirement as follows: g lysine/d = 0.043 × g crude protein required/d Given a dry matter intake of 2.5% of bodyweight, the daily crude protein and lysine requirement expressed as a percentage of the diet ranges from 16% and 0.7%, respectively, at 4 months of age, to 7% and 0.3%, respectively, at 24 months of age. Therefore, rations for weanlings require high-quality forage and/or grain-mix-concentrations containing supplemental protein (e.g., soybean meal). As the growing horse approaches maturity the concentration of crude protein and lysine is easily met by forage. Crude protein and DOD Excessive dietary crude protein is often implicated as a cause of DOD. Glade et al.29 suggested that dietary crude protein in excess of requirement may lead to an increase in renal calcium and phosphorus excretion, which may in turn have a negative impact on bone health. Schryver et al.30 investigated this hypothesis by feeding foals diets containing deficient (9%), adequate (14%), or excess (20%) crude protein and found lower growth rates and decreased bone turnover when deficient levels of crude protein were fed as compared to adequate and excess levels of crude protein. There were no negative effects of feeding excess crude protein on bone growth and development in these horses. Savage et al.8 also reported that weanlings fed excess crude protein (26% above a control ration formulated for moderate growth according to the NRC11 ) did not have a greater incidence of dyschondroplasia than horses fed the control ration. Thompson et al.25 suggested that the protein to calorie ratio could influence skeletal growth in that when energy levels of rations are increased, protein should also be increased to optimize bone growth and development. In summary, the results of these studies suggest that a deficiency in crude protein is more likely to create a problem in bone growth and development as compared to excess. There is no scientific evidence to suggest that excess crude protein has a negative effect on bone growth and development. It should also be noted that there does not appear to be any benefit in feeding excess crude protein in terms of growth rate or bone growth and development.30, 31
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Calcium and phosphorus Metabolic roles, dietary sources, and requirements The best-recognized role of calcium and phosphorus is as structural components of bone mineral. However, both calcium and phosphorus play several other roles. Calcium influences a number of physiological processes including muscle contraction, hormone release, neurotransmitter release, vision, glycogen metabolism, and cellular differentiation and proliferation.32 Phosphorus is a structural component of cell membrane phospholipids, DNA, and RNA, and plays a key role in metabolic regulation (e.g., various kinase enzymes) and energy metabolism as a component of ATP.33 The calcium and phosphorus contents of equine diets vary with ingredients. Grains are low in calcium (e.g., 0.01–0.1%) while forages are higher (0.5–1.5%) when compared to calcium requirements for growing horses (e.g., a 5-month-old weanling requires 0.8% Ca in the diet, assuming a DM intake of 2.5% of bodyweight).1 Among forages, legumes contain two to three times the calcium concentration as compared to grasses. Grains are relatively high in phosphorus (0.3–0.4%) while forages can have slightly lower (0.25%) to similar (0.33%) concentrations when compared to requirements for growing horses (e.g., a 5-month-old weanling requires 0.45% phosphorus in the diet, assuming a DM intake of 2.5% of bodyweight). Calcium availability from forage ranges from 50% to 70% (apparent absorption), with availability from alfalfa being higher than that from grasses.1 The phosphorus present in cereal grains is complexed with phytate. The phytate phosphorus is thought to be poorly absorbed by horses, as is the case in other monogastrics. van Doorn et al.34 reported 11.1 ± 1.5% apparent phosphorus digestibility in mature horses fed a diet containing a high proportion of phytate-bound phosphorus (i.e., 55%). Addition of microbial phytase (300 phytase units/kg diet) to the diet did not affect phosphorus apparent digestibility (11.1 vs 13.2 ± 1.5%), but did improve calcium apparent digestibility from 26.4 to 31.5 ± 1.4%. Equine diets are often supplemented with both calcium and phosphorus depending upon the base diet ingredients used. Supplementation is achieved through several inorganic sources (e.g., dicalcium phosphate, monocalcium phosphate), which have a relatively high availability. Published calcium and phosphorus requirements are estimated using the factorial approach, which takes into consideration the daily endogenous losses of these minerals associated with maintenance func-
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tions plus the daily amount of calcium and phosphorus retained in growing tissues, and divides them by the apparent absorption.1 The prediction equations for each are as follows:
calcium g/d = (0.072 × kg BW) + (32 g × kg BW gain/d) phosphorus g/d = (0.04 g × kg BW) + (17.8 g × kg BW gain/d)
The first quantity in the above equations reflects daily endogenous losses, whereas the second reflects tissue mineral accretion associated with growth. The above calcium and phosphorus requirements assume 50% and 45% apparent absorption, respectively. These absorption coefficients are average values. In the event of the diet being fed having a higher or lower absorption coefficient, the requirement is effectively decreased or increased, respectively. Rations for the growing horse can be formulated using forage to concentrate ratios ranging from 50:50 to 70:30 and are generally supplemented with inorganic calcium and phosphorus sources. Therefore the above estimates of apparent absorption are realistic. However, a ration containing a lower forage to concentrate ratio and unsupplemented with appropriate calcium and phosphorus may result in inadequate calcium and phosphorus intakes due to the low calcium concentration in grains and low phosphorus availability of grains.
Calcium, phosphorus, and DOD Both a deficiency of calcium and phosphorus can negatively impact bone growth and development.1 Additionally, phosphorus consumption in excess of calcium (i.e., inverted Ca:P) has been shown to cause dyschondroplasia in weanling horses. Five of six weanlings consuming 388% (1.71% P; Ca:P = 0.33:1) of required phosphorus had histologically confirmed lesions of dyschondroplasia as compared to 2 of 12 weanlings consuming the control diet (0.44% P; Ca:P = 1.3:1).35 However, excess calcium (1.95% Ca; Ca:P = 4.43:1) did not appear to induce dyschondroplasia lesions when all other requirements were in balance. Inverted Ca:P ratios occur practically when rations containing a relatively high proportion of an unfortified grain (e.g., > 60%) are fed in conjunction with grass hay.
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Copper Metabolic roles, dietary sources, and requirements The ability of Cu to cycle between oxidized and reduced states allows it to function as a catalytic cofactor for selective oxidoreductases, which are involved in a variety of different biological processes including ATP synthesis, antioxidant defense, iron metabolism, maturation of extracellular matrix and neuropeptides, and neuroendrocrine signaling.36 Maturation of extracellular matrix is of particular importance for bone growth and development. Lysyl-oxidase, a copper-dependent oxidoreductase, is necessary for pyridinoline cross-link formation of newly formed collagen. The pyridinoline cross-links facilitate formation of nucleation sites for bone mineral deposition. Impaired lysyl-oxidase activity due to copper deficiency is thought to impair bone mineral apposition and increase subchondral bone mineral fragility contributing to osteochondrosis dissecans in growing horses.37 The Cu concentration of typical feedstuffs used in rations for growing horses tends to be low (e.g., < 10 mg Cu/kg DM) relative to daily requirements (10 mg Cu/kg DM assuming a 2.5% BW DM intake). The availability of Cu in unsupplemented hay-grain diets is thought to range between 14% and 50% of dietary Cu.38 As a result Cu is commonly supplemented to base feeds. Copper can be supplemented using either inorganic or organic forms, or a combination of the two. Inorganic sources consist of Cu plus an inorganic ligand (e.g., copper sulfate; CuSO4 ), whereas organic forms consist of Cu and an organic ligand (e.g., copper lysine; Cu is centrally bound to the carboxyl and amino groups of two lysine molecules). Other commonly used inorganic and organic forms of copper include copper oxide and copper proteinates, respectively. Estimates of copper absorption from diets supplemented with organic and inorganic copper sources are limited and vary from 29% to 49% of total dietary copper.39, 40 Copper sulfate, copper-lysine, and copper-proteinates appear similar in bioavailability (i.e., the ability of the mineral to be absorbed and utilized in metabolism).1 Copper oxide is relatively low in bioavailability (e.g., relative bioavailability is 30% cf. copper sulfate).41 Copper absorption can be influenced by the composition of other minerals in the diet. Some have reported that high zinc levels result in decreased copper status (1200 mg Zn/kg diet42 ); whereas other reported no effect of dietary zinc level on copper status.43, 44 Based on current zinc and copper requirements, the zinc to copper ratio is 3.2:1. Iron has also been suggested as a
factor that can decrease copper absorption in horses, based on reports in other species,38 but the effect of iron on copper absorption in horses has not been determined. The current Cu requirement for growing horses is 0.25 mg Cu/kg BW.1 Assuming a total daily DM intake of 2.5% BW, 10 mg Cu/kg DM is required in the total diet for growing horses. The current requirement is in line with values reported by others based on a factorial model approach (i.e., estimate of endogenous loss + tissue accretion divided by a digestion coefficient).38, 45 Copper and DOD Copper deficiency has been implicated in osteochondrosis, and a mechanism has been proposed.37 However, the clinical significance of cartilage lesions developing during Cu deficiency has been questioned by some.37, 46 Also the dietary Cu threshold necessary to produce a cartilage lesion is unclear.47 The relationship between Cu and osteochondrosis has been a topic of research in the horse for more than 50 years. The hypothesis that Cu deficiency causes cartilage lesions in horses originated from an experiment conducted with calves in which Cu supplementation (15 mg Cu/kg DM) for nurse cows prevented cartilage lesions in calves.48 Cupps and Howell49 tested this hypothesis in growing foals and determined that their requirement was less than 8 mg Cu/kg DM and that the relationship between Cu and joint lesions was not clear. Secondary Cu deficiency due to excessive dietary Zn intake consumed by horses grazing pasture contaminated by industrial pollution was suggested as a cause of lameness, enlarged epiphyses, and stiffness of gait in growing horses.50, 51 Bridges et al.52 reported an association between low serum Cu concentrations and osteochondrosis. A subsequent report supported the hypothesis that Cu deficiency was associated with dissecting cartilage lesions in foals weaned from their dams at 1 day of age and fed liquid milk replacer for 5–7 months; however, the experimental design involved a small number of animals (n = 4) and the dietary Cu concentration was extremely low (1.7 mg Cu/kg DM) relative to typical feedstuffs.53 In a field survey study of 19 farms representing Thoroughbred, Standardbred, Quarter Horse, and Arabian breeds, Knight et al.54 reported a negative correlation between a ‘farm score’ reflecting the incidence and severity of DOD (high score = greater incidence and severity) and dietary Cu concentration. However, when two outliers in the analysis were removed the relationship no longer existed. A subsequent controlled clinical trial was conducted to determine the effect of Cu
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Feeding the Growing Horse to Avoid Developmental Orthopedic Disease (DOD) supplementation for mares (13 vs 32 mg Cu/kg DM; last 3–6 months of gestation) and foals (15 vs 55 mg Cu/kg DM; for 90 or 180 days) and the incidence of DOD.55 Although the average number of cartilage lesions per horse was numerically lower in the Cusupplemented group (1.6 and 2.8 lesions/horse on days 90 and 180, respectively) as compared to unsupplemented controls (4.4 and 12 lesions/horse on days 90 and 180, respectively), no inferential statistics were included in the report, making a final conclusion regarding the effect of Cu supplementation difficult. Furthermore, a single horse in the control group accounted for 59% and 35% of the lesions reported on days 90 and 180, respectively. Results from another clinical trial, in which 3-month-old weanlings were fed either 8 or 25 mg Cu/kg DM for 180 days, suggest that weanlings fed the supplemented Cu diet (i.e., 25 mg Cu/kg DM) had a lower incidence of OCD, and that the mechanism by which low Cu diets contribute to OCD may be due to reduced collagen quality resulting from impaired lysyl oxidase activity, which leads to poorly mineralized subchondral bone and subsequent subchondral bone fragility.37 However, the authors conclude that although the experiment created lesions in multiple sites throughout the body, the lesions were not typical of naturally occurring OCD in horses. Subsequent experiments support this idea that low Cu diets (< 8 mg/kg DM) may contribute to abnormalities in growth plate cartilage, but the clinical significance of these lesions is questionable.46, 56, 57 van Weeren et al.57 provided evidence that Cu deficiency is likely not a significant factor in the etiopathogenisis of osteochondrosis, but Cu status may play an important role in the natural process of early repair of lesions. This conclusion was based on the finding that foals having a higher liver Cu status had a reduction in the number and severity of radiographic osteochondrotic lesion scores in the stifle and hock from 5 to 11 months of age.
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transcription and gene expression. As a result Zn is important for growing and developing tissue such as that occurring in the skeleton of growing horses. Zinc deficiency impairs endochondral ossification in rodents58 and chicks.59 Excess dietary Zn may interfere with Cu, as discussed in the section on Cu (see above). The current dietary requirement for Zn is 40 mg/kg DM. Common forages used in feeding horses often contain less than this amount.
Vitamin A Metabolic roles, dietary sources, and requirements In addition to its role in vision, vitamin A functions in cell differentiation by the regulation of gene expression via nuclear retinoic acid receptors, and as a result plays crucial roles in reproduction and embryogenesis.60 Raw feed ingredients such as forage and cereal grains contain not vitamin A but its precursor betacarotene. Beta-carotene is metabolized into vitamin A in the small intestine. It is assumed that beta-carotene in equine diets is converted to vitamin A at a rate not exceeding 1 mg beta-carotene to 400 IU vitamin A. This estimate is likely high because the calculated vitamin A concentration using this conversion rate is well above the upper safe limit of 16 000 IU vitamin A/kg DM when applied to some forages containing high beta-carotene concentrations (e.g., alfalfa), yet no vitamin A toxicity due to high beta-carotene intake has been reported in horses.1 The current vitamin A requirement for growing horses is 45 IU/kg bodyweight. This requirement is based on intakes associated with clinically normal horses and is not a true minimum dietary requirement. Vitamin A and DOD
Other trace minerals Manganese and zinc are two trace minerals involved in skeletal growth and development. However, little is known about their role in DOD. Manganese is necessary for activation of glycosyl transferases involved in the synthesis of mucopolysaccharides, components of cartilage. Abnormal cartilage development due to failed chondroitin sulfate synthesis results from Mn deficiency in other species, and presumably in the horse. However, manganese deficiency has not been described in the horse. The current dietary Mn requirement is 40 mg/kg DM. Zinc is a cofactor for over 300 different enzymes and plays roles in the initiation and regulation of
Kronfeld et al.61 reported that excessive vitamin A consumption (supplied by retinyl palmitate) by two of three growing horses was associated with clinical and histological signs of physitis. The vitamin A intakes associated with these findings were 100 and 1000 times recommended intakes.1 This finding emphasizes the need to avoid haphazard vitamin A supplementation of growing horses.
Other vitamins Vitamin D plays a role in both Ca and P homeostasis and also influences cell growth and differentiation. Equine vitamin D status assessed by blood values for calcidiol and calcitriol is low relative to other
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species;62 however, the classical deficiency symptom of rickets has not been reported.1 Vitamin D deficiency produced by absence of sunlight and a dietary deficiency resulted in impaired bone growth and development in ponies and could be prevented with 1000 IU/day.63 Low vitamin D status has been implicated in osteochondrosis,64 but the vitamin’s exact role is unclear. The true minimum dietary vitamin D requirement for horses is not known. It is assumed that most of the metabolic requirement for horses is met by exposure to sunlight. In situations where sunlight is limited the following estimates have been suggested for growing horses: 22.2, 17.4, 15.9, and 13.7 IU/kg BW for 0–6, 7–12, 13–18, and 19–24 months of age, respectively. The recommended requirement for adult horses aged over 24 months is 6.6 IU/kg BW. Vitamin K functions as a cofactor for vitamin Kdependent carboxylase, which is necessary for the synthesis of proteins containing γ-carboxyglutamic acid (Gla-proteins). The Gla-proteins’ most widely known role is in blood clotting; however, they also play roles in bone metabolism.65 At the time of writing there are no published vitamin K minimum dietary requirements for horses. Horses’ vitamin K requirements are thought to be met by consumption of plant (phylloquinones present in forage) and microbial sources (menaquinones synthesized in the equine intestine).1 Only limited information regarding vitamin K status in growing horses exists, and indicates that vitamin K status is lowest in foals prior to weaning and increases at the time of weaning. These changes in vitamin K status were not correlated with changes in radiographic bone density of the third metacarpal bone.66 The effect of lower vitamin K status on bone metabolism remains to be determined.
Feeding growing horses Growing horses consume up to 2.5% of their bodyweight in dry matter daily. The total required nutrient amounts must be contained within this dry matter intake constraint. Because the nutrient requirements, expressed per unit of DM intake, are relatively high during the initial phases of growth and then decrease with age the diet must be relatively concentrated for young horses, but can be less nutrient dense for older growing horses. Diets for the growing horse should be forage (e.g., hay, pasture) based. Forage-based nutrients are well utilized by horses, even growing horses as young as weanlings.10 Horses are periodic grazers. They prefer to graze (i.e., many small frequent meals) as opposed to consuming one or two large meals.67 Additionally, small frequent forage-based meals are beneficial to the health of the gastrointestinal tract.68, 69 Forage
used in rations for the growing horse should be of high quality. Forage quality is influenced by several factors, including stage of maturity at harvest, plant variety, storage conditions, and weather conditions at harvest. Among these factors, stage of maturity at harvest has the greatest impact on forage quality. As a plant becomes more mature its nutrient concentration and digestibility decrease. Forage quality can be assessed visually or by analytical chemistry. Visual analysis is based primarily on indicators of maturity. As a plant matures the ratio of leaves to stems decreases and the presence of seed heads or flowers increases. Therefore forage for young horses should be relatively ‘leafy’ and contain minimal seed heads and/or flowers. Several analytical chemistry methods are available to assess forage quality;1 among them, crude protein (CP) and acid detergent fiber (ADF) are important indicators of quality. CP and ADF are related to forage quality directly and inversely, respectively. Forage fed to growing horses should contain at least 13–14% CP and have less than 35% ADF. Although forage is an important source of nutrients for growing horses, forage alone is often insufficient to meet the nutrient requirements of growing horses less than 1 year of age due to restricted DM intake capacity and relatively high requirements. Forage is typically supplemented with a complementary grain-mix-concentrate. Grain-mixconcentrate consists of energy concentrates (e.g., cereal grains, fat), protein supplements, mineral supplements, and vitamin supplements and is a concentrated source of calories, protein, minerals, and vitamins. Grain-mix-concentrates fed to growing horses should be formulated specifically for growing horses, and should not be intended for use in other feeding classes (e.g., performance horses). An example of appropriate grain-mix-concentrate nutrient concentrations is shown in Table 86.2. Table 86.2 Example of appropriate grain-mix-concentrate nutrient concentrations for selected nutrientsa Item Digestible energy (Mcal/kg) Crude protein (%) Lysine (%) Ca (%) P (%) Cu (mg/kg or ppm) Zn (mg/kg or ppm) Se (mg/kg or ppm) Vitamin A (IU/kg) Vitamin E (IU/kg) a
Nutrient concentration 3 to 3.2 14 to 16 0.85 0.75 to 1 0.6 30 to 50 150 0.3 to 0.5 1,000 80
All nutrient concentrations are expressed on an as-fed basis.
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Feeding the Growing Horse to Avoid Developmental Orthopedic Disease (DOD) Table 86.3 Suggested forage:concentrate ratios for growing horses assuming a daily dry matter intake of 2.5% of bodyweight
Age (months) 4 to 8 9 to 11 12 to 17 18 to 24: not in traininga 18 to 24: light to moderate traininga 19 to 24: heavy traininga
Forage (%)
Grain-mixconcentrate (%)
50 60 80 90 80 50
50 40 20 10 20 50
a Light = mean heart rate of 80 beats per minute (bpm), 1–3 h/week. Moderate = mean heart rate 90 bpm, 3–5 h/week. Heavy = mean heart rate 110 bpm, 4–5 h/week. Light and moderate training correspond to early stages of breaking and training, whereas heavy training corresponds to middle stages of race training.
Rations for the growing horse can be categorized into four different phases: early weaning (4–8 months), late weaning (9–11 months), early yearling (12–17 months), and long yearling (18–24 months). The ratio of forage:grain-mix-concentrate varies from 50:50 to almost 100:0 during these phases, as shown in Table 86.3. A bodyweight estimate is necessary to determine total dry matter intake using the values above. The bodyweight of a growing horse can be estimated using prediction equations based on heart girth circumference70 or age and mature bodyweight. 1, 71 For example, if a 5-month-old weanling weighs 200 kg it will consume 5 kg of DM/day (i.e., 200 kg BW × 0.025). Given the forage:concentrate ratio of 50:50 in Table 86.3, the weanling should be fed 2.5 kg DM of forage and 2.5 kg DM of grain-mix-concentrate daily. These DM intakes must be converted to asfed intake by dividing the amounts by the percent dry matter. Most air-dried feeds such as hay and grain-mix-concentrates contain approximately 90% dry matter. Therefore, dry matter amounts of hay and grain-mix-concentrate are divided by 0.9 to obtain the as-fed intake actually fed to the horse (e.g., 2.5/ 0.9 = 2.78 kg as-fed). The total as-fed amounts of forage and grain-mix-concentrate are typically divided into two or three feeds per day. Horses should have access to water free of choice. Rations should be fed in at least two equal feedings, but more feedings may be beneficial to the health of the gastrointestinal tract and relieve boredom in stalled horses. Care should be taken in groupfeeding settings to insure that all horses consume their intended rations. Rations should be updated at least every other month to account for changing bodyweight, dry matter intake, and nutrient requirements. Bodyweight should be monitored monthly to
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determine the effectiveness of the feeding program. Average daily gain can be calculated by subtracting the beginning and ending weight for a defined period (usually monthly) and dividing by the total number of days between beginning and ending weight. The actual average daily gain can be compared to predicted values for horses of similar mature bodyweight and age using the NRC’s Nutrient Requirements of Horses Model Program,1 and used to monitor the ration’s effectiveness. Body condition, an estimate of body fat covering, can also be monitored using the equine body condition scoring system.1 The equine body condition scoring system assigns a score from 1 to 9 based on fat covering over the neck, withers, top-line, and ribs. A body condition score of 5 is generally desired; this can be defined as a horse whose ribs are easily felt upon palpation but not seen.
Conclusions Nutrition is one of many factors contributing to DOD. Several nutrients have been shown to play a role in DOD, either by deficiency, excess, or imbalance. Aside from feeding digestible energy in excess of requirements, there still does not appear to be a dominant nutrient deficiency, excess, or imbalance accounting for the majority of DOD occurring in practical settings. Additionally, the nutrient intake thresholds that precipitate DOD are not well defined. The current body of evidence suggests that rations for the growing horse should be evaluated and formulated against current requirements1 to insure that all nutrient requirements are met, and that no imbalances or excesses exist. The growing horse’s nutrient requirements are constantly changing with increasing age and bodyweight, and therefore the ration must also change. Therefore, the most effective nutritional/dietary strategy for prevention of DOD is accurate ration formulation followed by frequent (e.g., at least monthly) ration evaluation by a knowledgeable equine nutritionist.
References 1. National Research Council. Nutrient Requirements of Horses, 6th edn. Washington, DC: National Academy Press, 2007. 2. De Lahunta A, Hedhammar A, Wu FM, Krook L. Overnutrition and skeletal disease. An experimental study in growing Great Dane dogs. VII. Cervical vertebrae and spinal cord. Cornell Vet 1974;64(2): Suppl. 5:58–71. 3. Olsson SE. Osteochondrosis in domestic animals. Introduction. Acta Radiol 1978;358(Suppl.):9–14. 4. Dammrich K. Relationship between nutrition and bone growth in large and giant dogs. J Nutr 1991;121(11 Suppl.):S114–S121. 5. Ruff SJ, Wood CH, Aaron DK, Lawrence LM. A comparison of growth rates of normal Thoroughbred foals and foals
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diagnosed with cervical vertebral malformation. J Equine Vet Sci 1993;13:596–9. Sandgren B, Dalin G, Carlsten J, Lundeheim N. Development of osteochondrosis in the tarsocrural joint and osteochondral fragments in the fetlock joints of Standardbred trotters. II. Body measurements and clinical findings. Equine Vet J 1993;16(Suppl.):48–53. Pagan JD, Jackson SG. The incidence of developmental orthopedic disease on a Kentucky Thoroughbred farm. Pferdeheilkunde 1996;12:351–4. Savage C, McCarthy RN, Jeffcott LB. Effects of dietary energy and protein on induction of dyschondroplasia in foals. Equine Vet J 1993;16(Suppl.):7–79. Hintz HF, Schryver HF, Lowe JE. Delayed growth response and limb conformation in young horses. In: Proceedings of the Cornell Nutrition Conference, Ithaca, NY, 1976, pp. 94–6. Petersen ED, Siciliano PD, Turner AS, Kawcak CE, McIlwraith CW. Effect of growth rate on bone mineral-content and -density of selected regions of the appendicular skeleton in growing horses. In: Proceedings of the 17th Equine Nutrition and Physiology Symposium, Lexington, KY, 2001, pp. 123–4. National Research Council. Nutrient Requirements of Horses, 5th edn. Washington, DC: National Academy Press, 1989. Vermorel M, Martin-Rosset W. Concepts, scientific bases, structure and validation of the French horse net energy system (UFC). Livest Prod Sci 1997;47:261–75. Cymbaluk NF, Christison GI. Effects of diet and climate on growing horses. J Anim Sci 1989;67:48–59. Cymbaluk NF. Cold housing effects on growth and nutrient demand of young horses. J Anim Sci 1990;68:3152–62. National Research Council. Nutrient Requirements of Horses, 4th edn. Washington, DC: National Academy Press, 1978. Glade MJ, Belling TH Jr. Growth plate cartilage metabolism, morphology and biochemical composition in over- and underfed horses. Growth 1984;48:473–82. Glade MJ, Gupta S, Reimers TJ. Hormonal responses to high and low planes of nutrition in weanling Thoroughbreds. J Anim Sci 1984;59:658–65. Glade MJ, Reimers TJ. Effects of dietary energy supply on serum thyroxine, tri-iodothyronine and insulin concentrations in young horses. J Endocrinol 1985;104:93–8. Ralston SL. Hyperglycemia/hyperinsulinemia after feeding a meal of grain to young horses with osteochondritis dissecans (OCD) lesions. Pferdeheilkunde 1996;3:320–2. Henson FM, Davenport C, Butler L, Moran I, Shingleton WD, Jeffcott LB, Schofield PN. Effects of insulin and insulin-like growth factors I and II on the growth of equine fetal and neonatal chondrocytes 1. Equine Vet J 1997;29:441–7. Pagan JD. The relationship between glycemic response and the incidence of OCD in Thoroughbred weanlings: a field study. In: Proceedings of the Kentucky Equine Research Nutrition Conference, Lexington, KY, 2003, pp. 119–24. Ott EA, Brown MP, Roberts GD, Kivipelto J. Influence of starch intake on growth and skeletal development of weanling horses. J Anim Sci 2005;83:1033–43. Hoffman RM, Lawrence LA, Kronfeld DS, Cooper WL, Sklan DJ, Dascanio JJ, Harris PA. Dietary carbohydrates and fat influence radiographic bone mineral content of growing foals. J Anim Sci 1999;77:3330–8. Cymbaluk NF, Christison GI, Leach DH. Energy uptake and utilization by limit- and ad libitum-fed growing horses. J Anim Sci 1989;67:403–13.
25. Thompson KN, Baker JP, Jackson SG. The influence of high dietary intakes of energy and protein on third metacarpal characteristics of weanling ponies. J Equine Vet Sci 1988;8:391–4. 26. Thompson KN, Baker JP, Jackson SG. The influence of supplemental feed on growth and bone development of nursing foals. J Anim Sci 1988;66:1692–6. 27. Thompson KN, Jackson SG, Baker JP. The influence of high planes of nutrition on skeletal growth and development of weanling horses. J Anim Sci 1988;66:2459–67. 28. Anderson KP, Raub R, Leedle R, Warren J. Influence of an imbalanced nutrient:calorie ratio and exercise on the incidence of developmental orthopedic disease in weanling horses. In: Proceedings of the Equine Nutrition and Physiology Symposium, Calgary, Alberta, 1991, pp. 15–16. 29. Glade MJ, Beller D, Bergen J, Berry D, Blonder E, Bradley J, Cupelo M, Dallas J. Dietary protein in excess of requirements inhibits renal calcium and phosphorus reabsorption in young horses. Nutr Rep Int 1985;31:659. 30. Schryver HF, Meakim DW, Lowe JE, Williams J, Soderholm LV, Hintz HF. Growth and calcium metabolism in horses fed varying levels of protein. Equine Vet J 1987;19:280–7. 31. Ott EA, Asquith RL. Influence of level of feeding and nutrient content of the concentrate on growth and development of yearling horses. J Anim Sci 1986;62:290–9. 32. Weaver CM. Calcium. In: Bowman BA, Russell RM (eds) Present Knowledge in Nutrition, 8th edn. Washington, DC: ILSI, 2001; pp. 273–80. 33. Anderson JJB, Sell ML, Garner SC, Calvo MS. Phosphorus. In: Bowman BA, Russell RM (eds) Present Knowledge in Nutrition, 8th edn. Washington, DC: ILSI, 2001; pp. 281– 91. 34. van Doorn DA, Everts H, Wouterse H, Beynen AC. The apparent digestibility of phytate phosphorus and the influence of supplemental phytase in horses. J Anim Sci 2004;82:1756–63. 35. Savage C, McCarthy RN, Jeffcott LB. Effects of dietary phosphorus and calcium on induction of dyschondroplasia in foals. Equine Vet J 1993;16(Suppl.):80–3. 36. Failla ML, Johnson MA, Prohaska JR. Copper. In: Bowman BA, Russell RM (eds) Present Knowledge in Nutrition, 8th edn. Washington, DC: ILSI, 2001; pp. 373–383. 37. Hurtig M, Green SL, Dobson H, Mikuni-Takagak Y, Choi J. Correlative study of defective cartilage and bone growth in foals fed a low-copper diet. Equine Vet J 1993;16(Suppl.):66–73. 38. Cymbaluk NF, Smart ME. A review of possible metabolic relationships of copper to equine bone disease. Equine Vet J 1993;16(Suppl.):19–26. 39. Miller ED, Baker LA, Pipkin JL, Bachman RC, Haliburton JT, Veneklasen GO. The effect of supplemental inorganic and organic forms of copper and zinc on digestibility in yearling geldings in training. In: Proceedings of the Eighteenth Equine Nutrition and Physiology Symposium, East Lansing, MI, 2003, pp. 107–12. 40. Baker JP, Wrigley M, Pipkin JL, Haliburton JT, Bachman RC. Digestibility and retention of inorganic and organic sources of copper and zinc in mature horses. In: Proceedings of the Nineteenth Equine Science Society Symposium, Tucson, AZ, 2005, pp. 162–7. 41. McDowell LR. Minerals in Animal and Human Nutrition, 2nd edn. Amsterdam, The Netherlands: Elsevier, 2003. 42. Coger LS, Hintz HF, Schryver HF, Lowe JE. The effect of high zinc intake on copper metabolism and bone development in growing horses. In: Proceedings of the Tenth
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Equine Nutrition and Physiology Symposium, Fort Collin, CO, 1987, pp. 173–7. Young JK, Potter GD, Greene LW, Web SP, Evans JW, Web GW. Copper balance in miniature horses fed varying amounts of zinc. In: Proceedings of the Tenth Equine Nutrition and Physiology Symposium, Fort Collins, CO, 1987, pp. 153–7. Bridges CH, Moffitt PG. Influence of variable content of dietary zinc on copper metabolism of weanling foals. Am J Vet Res 1990;51:275–80. Grace ND, Pearce SG, Firth EC, Fennessy PF. Content and distribution of macro- and micro-elements in the body of pasture-fed young horses. Aust Vet J 1999;77:172–6. Gee EK, Firth EC, Morel PC, Fennessy PF, Grace ND, Mogg TD. Enlargements of the distal third metacarpus and metatarsus in Thoroughbred foals at pasture from birth to 160 days of age. NZ Vet J 2005;53:438–47. Pearce SG, Firth EC, Grace ND, Fennessy PF. Effect of copper supplementation on the evidence of developmental orthopaedic disease in pasture-fed New Zealand Thoroughbreds. Equine Vet J 1998;30:211–18. Washburn LE. skeletal abnormality of probable nutritionalendocrine origin observed in cattle receiving antirachitogenic rations. J Anim Sci 1946;5:395. Cupps PT, Howell CE. The effects of feeding supplemental copper to growing foals. J Anim Sci 1949;8:286–9. Gunson DR, Kowalczyk DF, Shoop CR, Ramberg CF. Environmental zinc and cadmium pollution associated with generalized osteochondrosis, osteoporosis, and nephrocalcinosis in horses. J Am Vet Med Assoc 1982;180:295–9. Eamens GJ, Macadam JF, Laing EA. Skeletal abnormalities in young horses associated with zinc toxicity and hypocuprosis. Aust Vet J 1984;61:205–7. Bridges CH, Womack JE, Harris ED, Scrutchfield WL. Considerations of copper metabolism in osteochondrosis of suckling foals. J Am Vet Med Assoc 1984;185:173–8. Bridges CH, Harris ED. Experimentally induced cartilaginous fractures (osteochondritis dissecans) in foals fed lowcopper diets. J Am Vet Med Assoc 1988;193:215–21. Knight DA, Gabel AA, Reed SM, Embertson RM, Bramlage LR, Tyznik WJ. Correlation of dietary minerals to incidence and severity of metabolic bone disease in Ohio and Kentucky. In: Annual Symposium of the American Association of Equine Practitioners, 1985, pp. 445–61. Knight DA, Weisbrode SE, Schmall LM, Reed SM, Gabel AA, Bramlage LR, Tyznik WI. The effects of copper supplementation on the prevalence of cartilage lesions in foals. Equine Vet J 1990;22:426–32. Pearce SG, Firth EC, Grace ND, Fennessy PF. Effect of copper supplementation on the evidence of developmental
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62.
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66. 67. 68.
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orthopaedic disease in pasture-fed New Zealand Thoroughbreds. Equine Vet J 1998;30:211–18. van Weeren PR, Knaap J, Firth EC. Influence of liver copper status of mare and newborn foal on the development of osteochondrotic lesions. Equine Vet J 2003;35:67–71. Rossi L, Migliaccio S, Corsi A, Marzia M, Bianco P, Teti A, Gambelli L, Cianfarani S, Paoletti F, Branca F. Reduced growth and skeletal changes in zinc-deficient growing rats are due to impaired growth plate activity and inanition 1. J Nutr 2001;131:1142–6. Wang X, Fosmire GJ, Gay CV, Leach RM Jr. Short-term zinc deficiency inhibits chondrocyte proliferation and induces cell apoptosis in the epiphyseal growth plate of young chickens 1. J Nutr 2002;132:665–73. Solomons NW. Vitamin A and carotenoids. In: Bowman BA, Russell RM (eds) Present Knowledge in Nutrition, 8th edn. Washington, DC: ILSI, 2001; pp. 127–45. Kronfeld DS, Meacham TN, Donoghue S. Dietary aspects of developmental orthopedic disease in young horses. Vet Clin N Am Equine Pract 1990;6:451–66. Maenpaa PH, Koskinen T, Koskinen E. Serum profiles of vitamins A, E and D in mares and foals during different seasons. J Anim Sci 1988;66:1418–23. Elshorafa WM, Feaster JP, Ott EA, Asquith RL. Effect of Vitamin-D and sunlight on growth and bone-development of young ponies. J Anim Sci 1979;48:882–6. Sloet van Oldruitenborgh-Ooste, Mol JA, Barneveld A. Hormones, growth factors and other plasma variables in relation to osteochondrosis. Equine Vet J Suppl 1999;31(Nov):45–54. Ferland G. Vitamin K. In: Bowman BA, Russell RM (eds) Present Knowledge in Nutrition, 8th edn. Washington, DC: ILSI, 2001; pp. 164–72. Siciliano PD, Warren LK, Lawrence LM. Changes in vitamin K status of growing horses. J Equine Vet Sci 2000;20:726–9. Houpt KA. Ingestive behavior. Vet Clin N Am Equine Pract 1990;6:319–37. Clarke LL, Roberts MC, Argenzio RA. Feeding and digestive problems in horses. Physiologic responses to a concentrated meal. Vet Clin N Am Equine Pract 1990;6: 433–50. Hudson JM, Cohen ND, Gibbs PG, Thompson JA. Feeding practices associated with colic in horses. J Am Vet Med Assoc 2001;219:1419–25. Caroll CL, Huntington PJ. Body condition scoring and weight estimation of horses. Equine Vet J 1988;20:41–5. Ringler JE, Lawrence LM. Comparison of Thoroughbred growth data to body weights predicted by the NRC. In: Proceedings of the Twentienth Symposium of the Equine Science Society, Hunt Valley, MD, 2007, pp. 40–1.
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Chapter 87
Presale Endoscopy Rolf M. Embertson
Pre-sale evaluation of the upper airway has become commonplace at 2-year-old, yearling, and weanling Thoroughbred sales. The examination is performed to identify abnormalities that may eventually adversely affect the ability of that horse to train and race. Much of the information obtained regarding normal and abnormal findings in the upper airway of the immature horse has come from experience with the Thoroughbred intended for sale. This information can most likely be extrapolated to other breeds. A thorough upper airway examination should include visualization and palpation of the head and neck and endoscopy of the nasal passages, guttural pouches, pharynx, larynx, and trachea. However, the examination usually consists of brief endoscopy of the pharynx and larynx, with the endoscope passed up one nasal passage. Generally, this is adequate to assess the upper airway for the large majority of abnormalities that may occur. However, some findings are of debatable significance and some abnormalities may be overlooked. Thus, prior to making purchase recommendations, a thorough understanding of the normal and abnormal findings in the upper airway of the immature horse is important.
be an explosive event and unsafe for all involved. Excessive restraint, especially with the twitch, may also induce dorsal displacement of the soft palate. Removal of the twitch with the endoscope still in place may enable normal positioning of the epiglottis. The author prefers to pass the endoscope through the right nasal passage to gain access to the pharynx and larynx. Regardless of the nasal passage used, the operator should be consistent in the method of examination of each horse. After observing the upper airway at rest, maximal movement of the arytenoids should be evaluated by induction of swallowing and/or deep breaths. Introduction of water through the endoscope water channel or deflecting the endoscope tip to touch the side of the pharynx will induce swallowing. Bumping the epiglottis or arytenoid cartilages with the endoscope to induce swallowing should be discouraged. This can create visible signs of irritation that, although small, may affect the opinion of another veterinarian during a subsequent examination. Transient obstruction of the nasal passages will force the horse to breathe deeply and usually maximally abduct the arytenoid cartilages.
Examination of the upper airway
Maturation of the upper airway
The mechanics of performing an upper airway endoscopic examination varies between veterinarians and depends somewhat on the situation encountered. To obtain the most accurate picture of the upper airway, the horse should be examined in a stall, restrained by a competent handler, and not sedated. In the author’s opinion, by using the twitch to twist the muzzle away from the nasal passage of intended use and then placing a hand on the halter, the handler provides the most reliable means of stabilizing the head and restraining the horse for the examination. A lip chain is also commonly used for restraint. Excessive restraint may lead to the horse rearing and striking, which can
The ideal upper airway structure and function of a normal mature horse is used as the standard for the immature horse. However, structural and functional differences, although often subtle, are found when comparing the mature and immature horse. The immature horse generally has a higher grade of pharyngeal lymphoid hyperplasia (PLH), a shorter, narrower, and more flaccid epiglottis, and an increased incidence of dorsal displacement of the soft palate (DDSP) when the upper airway is stressed by nasal occlusion. The young horse will often have a row of small, raised nodules on the dorsal midline of the epiglottis. (Figure 87.1). In addition, arytenoid
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 87.1 Epiglottic midline ridge. This is occasionally seen in the immature horse and dissipates with age.
movement in the immature horse is less consistent and repeatable. These differences gradually disappear as the horse matures. The PLH nodules decrease in size and number. Changes in the structural appearance of the epiglottis are most evident between the yearling and 2-year-old, when the epiglottis becomes larger and more rigid and the small nodules disappear. Epiglottic appearance compared to average often remains similar as it matures, e.g., if shorter and more flaccid than average as a yearling, it usually remains shorter and more flaccid than average as an adult. As the horse nears adulthood, it becomes increasingly difficult to induce DDSP during an upper airway examination. In addition, arytenoid movement becomes more consistent between examinations. Based on observations in the Thoroughbred, the maturation process is usually complete late in their 2-year-old year.
Variation in upper airway findings The structure and function of some of the upper airway tissues change as they mature. These tissues can also vary in appearance over shorter periods, unrelated to maturation. This observation has been based on experiences at or near the time of sales, when the horses are most critically evaluated. The epiglottis can appear to vary mildly in size and stiffness over a few days or even a few hours. This is probably a result of the immediate level of excitement, the degree of induced respiratory stress (nasal occlusion), and effect of the endoscope in the pharynx. These factors may induce more contraction of the hyoepiglotticus muscle, which may transiently deform the epiglottis, making it appear more flaccid and possibly creating dorsal displacement of the palate.1 The incidence and the relative ease of induction of DDSP in the young horse may vary. This can occur over a matter of hours or days and seems to be related to several factors. Dorsal displacement of the
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soft palate more often occurs in horses with a smaller and more flaccid than normal epiglottis. It also occurs more often with increasing grades of PLH, which may be related to the level of local inflammation. Horses that are more excited and uncooperative tend to be more prone to DDSP. The method of restraint during the examination may play a role, for example, use of a twitch may make the young horse more prone to DDSP than a lip chain. Arytenoid movement is the most studied area during upper airway evaluation in the young horse. The concern regards the ability to predict future function of the arytenoids during the horse’s athletic career. Unless easily identified as normal, this can be difficult, as the degree of arytenoid movement can vary mildly with time. Symmetrical, full abduction of both arytenoids may occur during the entire examination in an excited yearling. In some yearlings, both the left and right arytenoids cannot be induced to fully abduct while evaluated at rest, using nasal occlusion to stress the airway. In some young horses, one arytenoid (usually the left) abducts to 94–96% of maximum, yet can maintain that degree of abduction. The above findings are considered normal by the author. Infrequently, the grade of arytenoid movement (described below) appears different over a period of a few hours to days. Arytenoid movement in yearlings has been observed during the time on the sales grounds to change from a grade II to a grade III and vice versa.2 As noted above, the variations in findings during subsequent endoscopic examinations of the upper airway in the young horse may be related to their level of excitement, fatigue, cooperation, and the amount of inflammation in the upper airway. Examination of the horse when minimally stressed generally provides the most accurate assessment of the upper airway. This may require re-evaluation at a different time. Variation in arytenoid movement in immature horses over a longer duration has been examined in longitudinal studies. Although different arytenoid movement grading systems were used, two reports examining changes from weanling to yearling yielded similar findings. In one study of 187 Thoroughbred foals (probable weanlings), 60% remained the same grade, 20% improved, and 20% worsened.3 In the other study of 243 Thoroughbred weanlings, 61% remained the same grade, 21% improved, and 18% worsened.4 In another study examining primarily Thoroughbred and some Standardbred horses from 6 months to 2 years of age, then again at least 16 months later, somewhat similar findings were noted. Overall, 43% remained the same grade, 29% improved, and 28% worsened.5 In these studies, changes in the arytenoid movement grades were usually not remarkable. However, they do
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support the belief that predictions of future arytenoid function based on an examination as an immature horse can be less than accurate. The results of the latter study5 may have been mildly affected by some horses developing recurrent laryngeal neuropathy (RLN). This abnormality usually starts to become evident on endoscopic evaluation of the upper airway in the Thoroughbred as a late yearling and early 2-year-old, and progresses at a variable rate.
Grading upper airway findings Grading systems for upper airway findings are useful for communication between veterinarians and to quickly assess one’s own notes regarding previous findings. The grading scale for PLH (I–IV) has been well accepted and is frequently used.6 Many grading systems for arytenoid movement have been used. In 2003, the Havemeyer Foundation supported a workshop on RLN where a consensus grading system was devised, built upon the structure of the previous Cornell system.7, 8 It seems that the Havemeyer grading system has become widely accepted. It is useful for any age of horse and is reasonably accurate in predicting arytenoid function, if that horse was maximally exercised. This grading system for arytenoid function performed in the standing, unsedated horse is as follows:
I All arytenoid cartilage movements are synchronous and symmetrical and full arytenoid cartilage abduction can be achieved and maintained. II Arytenoid cartilage movements are asynchronous and/or larynx asymmetric at times but full arytenoid cartilage abduction can be achieved and maintained. II.1 Transient asynchrony, flutter or delayed movements are seen. II.2 There is asymmetry of the rima glottidis much of the time due to reduced mobility of the affected arytenoid and vocal fold but there are occasions, typically after swallowing or nasal occlusion, when full symmetrical abduction is achieved and maintained. III Arytenoid cartilage movements are asynchronous and/or asymmetric. Full arytenoid cartilage abduction cannot be achieved and maintained. III.1 There is asymmetry of the rima glottidis much of the time due to reduced mobility of the arytenoid and vocal fold but there are occasions, typically after swallowing or nasal occlusion, when full symmetrical abduction is achieved but not maintained.
III.2 Obvious arytenoid abductor deficit and arytenoid asymmetry. Full abduction is never achieved. III.3 Marked but not total arytenoid abductor deficit and asymmetry with little arytenoid movement. Full abduction is never achieved. IV Complete immobility of the arytenoid cartilage and vocal cord. Grading epiglottic structure is not commonly done. The epiglottis in the young horse varies widely in size and stiffness and most veterinarians record a description of its physical appearance. An epiglottic structure grading system has been created to help quantify epiglottic characteristics in the yearling and examine any association with racing performance.9, 10 Epiglottic structure was graded as follows: 0 Adequate thickness length, and definition with serrated edges; I Slightly thin, of adequate length and texture, slightly flaccid, without serrated edges; II Mildly thin, of adequate length, mildly flaccid with curled edges, no obvious vasculature on the dorsal surface (Figure 87.2); III Very thin, of adequate length, moderately flaccid and easily bent; IV Extremely thin, markedly short, severely flaccid and easily bent.
Figure 87.2 Immature epiglottis. Note the epiglottis is mild to moderately narrow and flaccid, with curled-up edges. The dorsal vasculature is difficult to visualize. The epiglottic structure would be considered a grade II or III based on the system used here.
Approach to the upper airway regarding the purchase examination The Thoroughbred sales companies in North America, United Kingdom, Australia, New Zealand, and probably others have conditions of sale regarding the upper airway described in their catalogues. If
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Presale Endoscopy certain abnormalities are found during a veterinary examination immediately post-sale, the horse is returnable to the seller. This has compelled the consignors to have the upper airway of their sales horses examined pre-sale. Generally, those individuals that do not pass the conditions of sale are not brought to the sale. These conditions of sale have decreased the incidence of the upper airway abnormalities found in weanlings, yearlings, and 2-year-olds offered at these sales to a level where post-sale disputes are now rare. Although this author only has direct knowledge of the effect of these conditions on the sales in North America, the results appear the same in the United Kingdom.11 The upper airway abnormalities listed as returnable in the Keeneland sales catalog are as follows: laryngeal hemiplegia (consistent immobility or inability to fully abduct the arytenoid cartilage – Figure 87.3), rostral displacement of the palatopharyngeal arch (Figure 87.4), epiglottic entrapment (Figure 87.5), permanent dorsal displacement of the soft palate (Figure 87.6), chondroma (Figure 87.7) or severe arytenoid chondritis (Figure 87.8), subepiglottic cyst(s) (Figure 87.9), or cleft palate (Figure 87.10).12 Other sales companies have fairly similar conditions regarding the upper airway in their sales catalogs. At the Tattersalls sale, regarding arytenoid function, the horse is returnable if it makes a characteristic abnormal inspiratory sound and has laryngeal hemiplegia (RLN).11 It is also stated in the Keeneland catalog that surgical intervention of the upper respiratory tract must be disclosed by placing a veterinary certificate in the repository or be announced at the time of sale. Acupuncture and/or electro-stimulation with the intent of altering laryngeal function are prohibited once the horse enters the Keeneland sales ground.
Figure 87.3 Left arytenoid paralysis, grade IV. Although the grading system is based on movement, there appears to be no abduction of the left arytenoid when the right arytenoid has been stimulated to fully abduct.
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Figure 87.4 Rostral displacement of the palatopharyngeal arch. This is the most commonly seen manifestation of a fourth branchial arch defect congenital abnormality during endoscopic examination of the upper airway.
The upper airway of most horses at the sale intended for racing are examined pre-sale by the buyer’s veterinarian. This is done to screen for the above listed abnormalities, but also for abnormalities not listed as returnable,for example, cysts in the nasal septum or maxillary region, or findings that may be within normal limits, but close enough to abnormal to be of concern. Examples of the latter are grade II.2 arytenoid movement, a short, flaccid, and thin epiglottis, or frequent DDSP. It is much easier to avoid purchase than to attempt to return a borderline passable horse. The incidence of upper airway abnormalities in the young horse is low. A few studies of large numbers of Thoroughbred yearlings provide some information regarding incidence.10,13–15 However, these studies primarily examined yearlings at sales, which would not reflect the true incidence in the general population of Thoroughbred
Figure 87.5 Epiglottic entrapment. In a young horse, the tissue entrapping the epiglottis is usually smooth and not ulcerated. This is in contrast to an adult race horse, where the tissue tends to be swollen and ulcerated.
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Figure 87.6 Dorsal displacement of the soft palate. This occurs commonly in the immature horse as a transient event. Permanent DDSP is rare and usually associated with neurological disease.
Figure 87.8 Arytenoid chondritis. The arytenoid cartilage, in this case the right, becomes inflamed and thickened. Once deformed, the cartilage rarely returns to normal.
yearlings. The ranges of the percentage of yearlings in these studies found with the following abnormalities are: grade IV arytenoid movement (0–0.48%), fourth branchial arch defects (0.05–0.2%), entrapped epiglottis (0.06–0.12%), subepiglottic cyst (0–0.14%), and arytenoid chondritis or chondroma (0–0.63%). The incidence of arytenoid chondritis/chondroma/mucosal lesion is higher in Australia than in other countries with large Thoroughbred yearling sales.
Relationship between upper airway findings and racing performance The relationship between upper airway findings in the Thoroughbred yearling and subsequent racing performance has been examined.10, 16 In one study, 427 sales yearlings were examined by four
Figure 87.9 Subepiglottic cyst. The cyst can sometimes be easily seen during upper airway endoscopy, or positioned in the oropharynx and hidden from view by the soft palate. It is not uncommon to see cysts associated with an entrapped epiglottis in the immature horse.
Figure 87.7 Arytenoid chondroma. This generally refers to a mass that protrudes from the arytenoid cartilage into the laryngeal lumen.
Figure 87.10 Cleft palate. This is generally found shortly after birth when the reason for milk dribbling from the nose following nursing is investigated. Some of these horses are able to reach adulthood without developing aspiration pneumonia.
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Presale Endoscopy veterinarians.16 The 4-point Cornell grading system was used to grade arytenoid movement and a subjective assessment of normal or abnormal was used to categorize epiglottic appearance. The percentage of yearlings in each of the arytenoid function grades were: I – 35%, II – 40%, III – 22%, IV – 3%. Race records were examined at 2, 3, and 4 years of age. Results of this study revealed that visual epiglottic abnormalities such as thin, flaccid and hypoplastic epiglotti (8.7% of these yearlings) were not associated with racing performance. Horses with grades I and II arytenoids raced significantly better than those with grades III and IV arytenoids. In a later study, 2954 sales yearlings were examined by one veterinarian.10 An expanded Cornell grading system,2 later retrofit into the Havemeyer system, was used to grade arytenoid function. A previously devised system (see above) was used to grade epiglottic structure. The percentage of yearlings in each of the arytenoid function grades were: I – 19%, II.1–70%, II.2–9%, III (all) – 2%, IV – 0%. The horses with grades III.1, III.2, and III.3 were combined for comparison in this study. Epiglottic structure grades 0–II (99%) were combined and compared to grades III–IV (1%). Race records were examined at 2, 3, and 4 years of age. Results of this study showed that horses with arytenoid function grades I, II.1, and II.2 had significantly more starts, and more earnings per year at 3 and 4 years of age than grade III horses, with a similar trend at 2 years of age. In addition, horses with grades I and II.1 were significantly better in most parameters examined than grade II.2 horses. Horses with epiglottic structure grades 0–II performed significantly better in most parameters studied than grades III– IV horses. The difference between these studies, regarding the percentage of yearlings in each of the arytenoid function grades, is likely related to the evaluator’s assessment. It is possible that many yearlings that were graded II.2 in the second study may have been graded III in the first study. In addition, a single evaluator (second study) may have been more consistent than four evaluators (first study). The results regarding prediction of racing performance were similar, i.e., grade I and II yearlings perform better as racehorses than grade III and IV yearlings. The above findings should also be considered when evaluating for purchase a yearling with grade II.2 arytenoid function. The results regarding visual epiglottic structure and racing performance differ between these studies. This is probably explained by the more critical grading of epiglottic structure performed in the second study. In reality, yearlings with markedly small and flaccid epiglotti are discriminated against at the sale.
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The second study provides evidence to support this practice. In summary, critical evaluation of the upper airway of the Throughbred sales yearling is important to avoid purchase of a horse that will be unable to perform as desired. These studies help further support opinions based on experience.
References 1. Holcombe SJ, Cornelisse CJ, Berney C, Robinson NE. Electromyographic activity of the hyoepiglotticus muscle and control of epiglottic position in horses. Am J Vet Res 2002;63:1617–21. 2. Embertson RM. Evaluation of the upper respiratory tract in weanlings and yearlings. In: White NA, Moore JN (eds) Current Techniques in Equine Surgery and Lameness. Philadelphia: Saunders, 1998; pp. 122–7. 3. Lane JG. Long-term longitudinal study of laryngeal function in 187 foals. In: Dixon PM, Robinson E, Wade JF (eds) Havemeyer Foundation Monograph Series No. 11. Proceedings of a workshop on equine recurrent laryngeal neuropathy. R&W Publications (Newmarket) Ltd, 2004; pp. 31–2. 4. Pierce SW, Embertson RM. Changes in the upper respiratory tracts from weanling to yearling in 371 Thoroughbreds. Proc Am Assoc Equine Pract 2005;51:309–11. 5. Anderson BH, Kannegieter NJ, Goulden BE. Endoscopic observations on laryngeal symmetry and movements in young race horses. N Z Vet J 1997;45:188–92. 6. Raker CW, Boles CL. Pharyngeal lymphoid hyperplasia in the horse. J Equine Med Surg 1978;2:202. 7. Rakestraw PC, Hackett RP, Ducharme NG, Nielan GJ, Erb HN. Arytenoid cartilage movement in resting and exercising horses. Vet Surg 1991;20:122–7. 8. Workshop summary: Clinical grading of RLN. In: Dixon PM, Robinson E, Wade JF (eds) Havemeyer Foundation Monograph Series No. 11. Proceedings of a workshop on equine recurrent laryngeal neuropathy. R & W Publications (Newmarket) Ltd, 2004; pp. 95–6. 9. Pierce SW, Embertson RM. Correlation of racing performance to yearling endoscopic evaluation. Proc Am Assoc Equine Pract 2001;47:113–14. 10. Garrett KS, Pierce SW, Embertson RM, Stromberg AJ. Endoscopic evaluation of arytenoid function and epiglottic structure in Thoroughbred yearlings and association with racing performance at two to four years of age: 2,954 cases (1998–2001). J Am Vet Med Assoc 2010;236:669–73. 11. Ellis DR, Greet TRC, Lane JG. Sales: Problems in Diagnosis of RLN – UK Perspective. In: Dixon PM, Robinson E, Wade JF (eds) Havemeyer Foundation Monograph Series No. 11. Proceedings of a workshop on equine recurrent laryngeal neuropathy. R&W Publications (Newmarket) Ltd, 2004; pp. 39–41. 12. Conditions of this sale. January horses of all ages sale. Keeneland Association, Inc. 2010; pp. 38–60. 13. Anderson BH. Non-RLN URT disorders identified during post sale endoscopic examination of 5,559 TB yearlings (1997–2002) in New Zealand. In: Dixon PM, Robinson E, Wade JF (eds) Havemeyer Foundation Monograph Series No. 11. Proceedings of a workshop on equine recurrent laryngeal neuropathy. R&W Publications (Newmarket) Limited. 2004; pp. 51–4.
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14. Kelly G, Lumsden JM, Dunkerly G, Williams T, Hutchins DR. Idiopathic mucosal lesions of the arytenoid cartilages of 21 Thoroughbred yearlings: 1997–2001. Equine Vet J 2003;35:276–81. 15. Lane JG. Non-RLN upper respiratory tract disorders found in a survey of 3,497 Thoroughbred yearlings. In: Dixon PM, Robinson E, Wade JF (eds) Havemeyer Foundation Monograph Series No. 11. Proceedings of a workshop on equine recur-
rent laryngeal neuropathy. R&W Publications (Newmarket) Ltd. 2004; pp. 49–50. 16. Stick JA, Peloso JG, Morehead JP, Lloyd J, Eberhart S, Padundtod P, Derksen FJ. Endoscopic assessment of airway function as a predictor of racing performance in Thoroughbred yearlings: 427 cases (1997–2000). J Am Vet Med Assoc 2001;219:962–6.
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Chapter 88
Presale Radiographs James P. Morehead and Brent N. Cassady
Radiographic assessment of the sale horse does not rule out all pathology; instead it is an attempt to economically identify known issues affecting the use or proposed use of the individual. One must remember that the ability to race does not require a clean radiographic assessment. Many young horses have had issues or trauma that have resulted in radiographic changes from normal. The correlation of radiographic findings and their effect on selling, training, or racing is a topic for another chapter. All are dependent on a consistent quality of radiographic evaluation, and this is the topic of this chapter. The ability to reproduce a standard for each exam performed on sales horses facilitates your evaluation of those individuals. Variations in techniques, marker orientation, or artifacts from faulty equipment or dirt on the limb add only to the frustration level of those evaluating the films. Consistency is the requirement and each veterinarian who generates radiographic images is in charge of their own quality control. Quality of films generated is a direct reflection of the professionalism of the veterinarian who radiographed the horse. Repository films for use in auctions are meant to reduce the radiographic exposures needed and to facilitate the inspection of sale horses. It is very important to examine the entire radiograph and not just focus on the perceived area of interest. If there is a radiographic change anywhere on the radiograph then that change should be noted and described on the accompanying radiographic report. Generating an accurate radiograph report allows the veterinarian to suggest a knowledgeable prognosis or therapy for the horse whereas a radiographic report without sufficient detail may lead everyone – buyer, seller, and their veterinarians – down the wrong path. The title of this chapter refers to the radiographic evaluation of the distal limbs. Skull, cervical, elbow, shoulder, pelvic, or even foot radiographs are commonly performed but are not included routinely in the evaluation of sale horses.
Radiation safety The radiographic evaluation of any horse must include adequate radiation safety. The veterinarian, who performs the procedure many times, is the sole educated party concerning the risks, and he or she must be diligent in keeping all personnel involved out of harm’s way, by both proper restraint of the horse and minimizing any radiation hazard; they must continually monitor the techniques without endangering the assistants through undue exposure to radiation. Ensuring the use of protective gear along with keeping laypersons out of the primary beam are the responsibilities of the veterinarian. No doubt some countries regulate radiation safety more than others, and veterinarians are advised to educate themselves not only about the laws of their own area but also about radiation safety in general. Monitoring badges may or may not be required but are most certainly recommended so as to protect all who assist in this procedure, and also to protect the veterinarian as an employer of those same assistants. The monitoring of exposure levels is a priority of responsible employers. There are many cassette holders available that keep the assistant’s hand out of the primary beam. Remember that the lead protective gear does not block the primary beam. Lead is intended to block scatter radiation. There are also devices designed to hold the radiograph machine itself during exposures, but the levels of sedation will need to be adjusted to the level of additional equipment used in the procedure.
Radiographic labeling Proper labeling of radiographs is a well-established protocol. The marker system should inform the viewer of the individual horse being examined, by either sale hip number, name, or pedigree. The marker should also tell who generated the film, on which
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date, and which limb is being examined. With the use of plain films and proper marker placement those viewing the radiographs can easily ascertain which view is being examined. The simple rule of the marker placement on the cassette is always lateral if possible, with the exception being on the lateralmedial projections and then the marker is placed on the dorsal aspect of the joint. The only views with the markers on the dorsal aspect of the limb are the standing lateral, or the flexed lateral views. In all other views the marker should be placed laterally. The digital radiography units currently available may allow for the veterinarian to customize the marker placement, but most label the view with standard nomenclature identifying the view.
Sedation In the field of human radiology the technician can request the patient to lie still once positioned or even to assist with the positioning. This highlights the need for adequate restraint including the use of sedatives or tranquilizers in equine radiology. Familiarity with the different agents and the expected results with the horse when used alone or in combination will come with experience. The veterinarian should use caution during this period so as not to compromise him or herself, the assistants, the horse, or the equipment. The ability to ‘take control’ of the situation and accomplish the task is a learned behavior. Saving a couple of dollars on tranquilizers may land the veterinarian in the emergency room or the radiograph machine in the shop. Either one will reduce the efficiency of accomplishing the task, at least for a while. We currently prefer detomidine alone, or in combination with butorphanol, for sedation while radiographing for sales.
Examination procedures Adequate sedation, protected, educated, and sufficient staff, along with quality radiographic equipment all add to the efficient production of sales radiographs. Do not lose focus on the primary goal, which is the generation of standardized quality films to facilitate the examination of sales horses. Speed comes with any procedure after repetition. The primary focus cannot be speed when learning the required views. Motion is a common problem and it can be motion of the patient, the cassette, or the radiograph machine. All motion needs to be eliminated to achieve quality radiographs. The views required adequately to evaluate a horse pre-sale depend on the type of horse, its age, and the budget predetermined by the potential buyer. The problematic lesion may only be visible on one view,
which makes that view required if you are to achieve an accurate evaluation. That said, you are in the position of acquiring the views with the highest probability of showing pathology as determined by your experience. The required views can be different on a weanling or yearling Thoroughbred intended for racing compared with a 2-year-old or adult Warmblood or Quarter Horse. The required list of views on any category of horse may also change based on the examination. The yearling Thoroughbred market may not require a radiographic assessment of the foot, but some veterinarians may advise it or some buyers may request it. Sufficient radiographic views of pertinent anatomical areas are desired and those are based on a thorough examination, knowledge of the intended use of the horse, and the buyer’s expectations. The views described are the current views recommended by most sales companies with repositories (Table 88.1). Fetlock (metacarpophalangeal) joint A routine examination of the front fetlock joint requires five views and the hind fetlock four views: dorsopalmar/dorsoplantar (DP), dorsal lateral palmaro/plantarodistal medial oblique (DLPMO), dorsal medial palmaro/plantarodistal lateral oblique (DMPLO), both a flexed lateral-medial (FLM) and a standing lateral-medial (SLM) of the front fetlock and a standing lateral-medial (SLM) of the hind fetlock. The discussion of the fetlock joint could include additional views but remember we are taking high probability radiographs. If lameness, local pain, or swelling is present then the required views may change to fit the clinical presentation. The 20-degree elevated proximal DP view is the standard DP projection of the fetlock. The elevation of the generator lifts the sesamoids above the distal end of the cannon bone (third metacarpal/metatarsal). This separation of the sesamoids above the fetlock joint allows for evaluation of the distal aspect of the sesamoids and the distal end of the cannon bone. A true DP view (without elevation = parallel to the ground) results in a view with overlap of the sesamoids and the distal end of the cannon. Subchondral bone cysts of either the distal cannon bone or proximal first phalanx (P1) are not infrequent in youngsters, as are some basilar changes in the sesamoids, and these are best evaluated on this view. Previously fractured sesamoids that have since healed may result in an elongated sesamoid, and this view allows comparison of the medial and lateral sesamoids. Radiographic cassette placement routinely is perpendicular to the ground surface in all other views but not on this view. Placing the plate parallel to the
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Table 88.1 Generally recommended views for radiographic repository for sale of horses that have not raced and the most common lesions revealed by that projection. For horses that have raced or trained, a minimum of three additional views of the carpus are recommended (DP, standing lateral, and a skyline projection of the distal row of carpal bones). Table reproduced by courtesy of Elizabeth M. Santschi, DVM, Diplomate ACVS Joint
Views
Common traumatic lesions
Developmental lesions
Fetlock
DP
DJD, physitis, angular limb deformities, sesamoid fractures, proximal plantar P1 fractures Dorsal proximal P1 fractures
Subchondral bone cysts (also pastern), OCD median sagittal ridge
Flexed lateral (forelimb) Standing lateral (hindlimb and forelimb) DLPMO DMPLO
Carpus
Flexed lateral DLPMO DMPLO
Hock
DP Lateral DMPLO
Stifle
Lateral Lateral 20-degree caudal-mediocranial oblique (CaLCrMO) Posterior/Anterior (PA)
OCD median sagittal ridge, supracondylar lysis of distal MC3
Sesamoid fractures, dorsal proximal P1 fractures Sesamoid fractures, dorsal proximal P1 fractures
Palmar/plantar P1 fragments
Cuboidal bone fractures or remodeling, accy. carpal bone fracture Cuboidal bone fractures or remodeling Cuboidal bone fractures or remodeling
Subchondral bone cysts (rare), osteochondroma (rare) Subchondral bone cysts Subchondral bone cysts
DJD of DIT and TMT joints, suspensory ligament avulsion (rare) DJD of DIT and TMT joints DJD of DIT and TMT joints
Medial malleolar OCD
Distal patellar fractures Intercondylar area fractures, insertional desmitis of meniscal ligaments
Trochlear ridge and patellar OCD Subchondral bone cyst medial femoral condyle
Intercondylar area fractures, insertional desmitis of meniscal ligaments and DJD
Subchondral bone cyst medial femoral condyle
Palmar/plantar P1 fragments
Trochlear ridge OCD Distal intermediate ridge of tibia OCD
DIT, distal intertarsal; DJD, degenerative joint disease; DLPMO, dorsal lateral palmaro/plantarodistal medial oblique; DMPLO, dorsal medial palmaro/plantarodistal oblique; OCD, osteochondritis dissecans; P1, first phalanx; TMT, tarsometatarsal
pastern and touching the heels and the back of the fetlock joint will usually allow the examiner to visualize the pastern (proximal interphalangeal) joint also (Figure 88.1). The dorsal 10-degree proximal 30-degree lateral palmaro/plantarodistal medial oblique (DLPMO) (Figure 88.2) and the dorsal 10-degree proximal 30-degree medial palmaro/plantarodistal lateral oblique (DMPLO) (Figure 88.3) views are views routinely used as the standard obliques in the evaluation of the front or hind fetlocks. The elevation allows the separation of the distal aspect of the sesamoid from the proximal palmar, or plantar, eminences of P1. The radiographic plate is perpendicular to the ground surface. Excessive elevation leads to distortion of the dorsal proximal aspect of P1 and may cause sufficient overlap to disguise chip fractures. The standing lateral view of hind fetlocks is easily obtained if the examiner remembers that hindlimbs are pronated. True lateral is actually 10–15◦ caudal to lateral due to the outward rotation of the hindlimbs.
The beam is parallel to the ground surface. The flexed lateral view of the fetlock increases the likelihood of motion, as with all non-weight-bearing radiographs. The assistant holding the limb and cassette should avoid abducting the limb. The fetlock should be moderately flexed. Practicing this view repeatedly will educate the examiner on beam direction. The advent of direct radiology is a highly prized asset in learning these techniques. The ability to expose the plate and view the image instantly allows the examiner to modify the beam direction to achieve the true flexed lateral view (Figure 88.4). Both the standing and flexed lateral views should allow visualization of the median sagittal ridge, the dorsoproximal aspect of the P1, and the true overlap of the sesamoids. The standing lateral view of the hind fetlock allows another view of the proximal plantar aspect of P1 and the distal margin of the sesamoids. As we refine our radiographic techniques it will also be possible for us to evaluate additional parameters. A true flexed or standing lateral view
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Figure 88.1 Dorsopalmar/dorsoplantar (DP) view of fetlock.
Figure 88.3 Fetlock: dorsal medial palmaro/plantarodistal lateral oblique (DMPLO) view.
allows for accurate interpretation of the distal MC3 or MT3 for supracondylar lysis, which is an important prognostic indicator (Figure 88.5). Carpus (knee) Yearling and weanling pre-sale radiographs of the carpus are commonly the flexed lateral, the dorsal 65degree medial-palmarolateral oblique, and the dorsal 65degree lateral-palmaromedial oblique. Horses in training
Figure 88.2 Fetlock: dorsal lateral palmaro/plantarodistal medial oblique (DLPMO).
Figure 88.4 Fetlock: flexed L-M view.
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Presale Radiographs
Figure 88.5 Fetlock: standing lateral view.
will also have a skyline (flexed dorsoproximaldorsodistal oblique [flexed D30Pr-DDiO]) view of the third carpal bone. It must be remembered, as we stated with regard to the fetlocks, that more views may be required if the examiner finds clinical issues that warrant further investigation. Due to being nonweight bearing, the flexed lateral view of the carpus has a higher incidence of motion. This view allows for assessing both the proximal and distal borders of the intermediate and radiocarpal bones, the accessory carpal bone, and the distal radius. Avoiding total overlap of the proximal row of carpal bones is important or the evaluation will be hampered. The medial oblique (dorsal 65-degree medial-palmarolateral oblique) view is facilitated by moving the limb slightly forward. The beam is parallel to the ground, and the common problem is being too dorsally positioned. Without moving the limb slightly forward it will be more difficult to achieve this view without getting the other carpus in the field. This view will highlight the intermediate carpal bone. The lateral oblique (dorsal 65-degree lateral-palmaromedial oblique) view is best achieved with the limb perpendicular to the ground and the beam parallel to the ground surface. This view highlights the dorsal border of the radiocarpal and third carpal bone (Figures 88.6–88.8).
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Figure 88.6 Carpus: dorsal lateral palmaro/plantarodistal medial oblique (DLPMO) view.
the lateral, and the DMPLO (dorsal 25-degree medial plantarolateral oblique). The dorsal 10-degree lateral-plantaromedial oblique view highlights the medial malleolus, the distal rows of tarsal bones, and the proximal third metatarsal
Tarsus (hock) The common views obtained for sales horses are the DP (dorsal 10-degree lateral-plantaromedial oblique),
Figure 88.7 Carpus: dorsal medial palmaro/plantarodistal lateral oblique (DMPLO) view.
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Figure 88.8 Carpus: flexed L-M view.
bone. The beam is parallel to the ground on all three standard radiographs of the tarsus. The x-ray beam for the lateral-medial view (L-M), due to the pronated limb, appears to be directed slightly dorsally. This view shows the distal intermediate ridge of the tibia, the trochlear ridge, the distal rows of the tarsal bones, and the proximal third metatarsal bone. The dorsal 25-degree medial- plantarolateral oblique view of the hock and the dorsal 10-degree proximal 30-degree medial plantarolateral oblique view of the hind fetlock are shot from the opposite side of the horse and underneath its abdomen. The beam on the hock is parallel to the ground. This view of the hock highlights the distal tibia and the lateral trochlear ridge of the talus (Figures 88.9–88.11).
Figure 88.9 Tarsus: dorsal 10-degree lateral-plantaromedial oblique (D10L-PLMO) .
joint. The use of a lip twitch or shoulder roll will facilitate the acquisition of the needed views. Merely rubbing the medial aspect of the stifle prior to cassette placement is also helpful. The limb being radiographed should be a half step behind the off limb.
Stifle The stifle was not routinely included in radiographic evaluations during the 1990s but has since become standard. Although we have stressed the highprobability radiographs to detect common pathology and developmental issues, no other joint commonly evaluated has such a large surface area. This surface area might suggest the need for additional views, but currently the standard evaluation and the requirement of the sales companies is for three views. The joint also requires attentive staff and careful horse handling as the subject will feel threatened by cassette placement. Sedatives have been discussed, but frequently manual restraint will be added for this
Figure 88.10 Tarsus: dorsal medial palmaro/plantarodistal lateral oblique (DMPLO) view.
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Figure 88.11 Tarsus: L-M view. Figure 88.12 Stifle: lateral-medial (L-M) view.
Once the patient has been adequately restrained and is aware of the massage to the medial aspect of the stifle the cassette is inserted. Assessing both the medial and the lateral trochlear ridges of the distal femur is important and is best achieved by the lateral view (L-M) of the stifle (Figure 88.12). Adequate evaluation of both trochlear ridges is facilitated by two lateral views, one highlighting the medial and then the lateral 20-degree caudal-mediocranial oblique view to highlight the lateral trochlear ridge (Figure 88.13). Hindlimbs are pronated (or rotated outward) so a lateral view will actually have the beam parallel to the ground and directed slightly forward. When using a digital radiology system leave the cassette on the medial aspect of the stifle for the lateral and then move only the radiograph machine to generate the next view. Changing cassettes is required with plain films between exposures but with a digital system you should hold the plate still so as not to stimulate the horse. The lateral 20-degree caudal-mediocranial oblique view, required by sales companies, has the same cassette placement as the true lateral to highlight the medial trochlear ridge, but the radiograph machine moves caudally and with the cassette inserted sufficiently deep the examiner can visualize the lateral trochlear ridge and the distal medial femoral condyle. This view entails a high risk of damage to the radiograph machine, so all participants are encouraged to be calm, yet alert. The beam is parallel to the ground.
The posterior-anterior (PA) view of the stifle requires a quick trigger finger and a willing patient but is routinely acquired in sales horses. The limb being evaluated remains a half step behind the off limb. The beam is directed slightly downward at the
Figure 88.13 Stifle: lateral 20-degree caudal-mediocranial oblique (CaLCrMO) view.
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Foal: Problems of Older Foals, Weanlings and Yearlings ciate ligaments. The radiograph cassette is moved to the dorsal surface of the stifle for this view (Figure 88.14).
Summary Adequate pre-sale radiographs of the horse require that the examining veterinarian has knowledge of the intended use, the ability to correlate the radiograph with the clinical picture, and the ability to obtain a quality set of images. If we critically evaluate the radiographs that we obtain and refuse to settle for images with motion or artifacts while striving to generate repeatable, quality, well-positioned radiographs then there will be less imagination required to accurately access our radiographs (see Table 88.1).
Further reading
Figure 88.14 Stifle: caudal-cranial (Cd-Cr) or posterior-anterior (PA) view.
stifle to separate the distal femur from the proximal tibia. Overlapping of these bones will not allow accurate interpretation of the radiographs as subchondral bone cysts are not uncommon on the distal medial femoral condyle. The PA view also allows visualization of the areas of origin and the insertion of the cru-
The Prevalence of Radiographic Changes in Thoroughbred Yearlings – Kane, AJ, DVM, MPVM, PhD; McIlwraith, CW, BVSc, PhD; Park, RD, DVM, PhD; Rantanen, NW, DVM, MS; Morehead, JP, DVM; and Bramlage, LR, DVM, MS How to Properly Position Thoroughbred Repository Radiographs – Garrett, KS, DVM; and Berk, JT, VMD Radiographing Thoroughbred Yearlings for the Repository – Hance, SR, DVM; Morehead, JP, DVM Optimal Radiographic Views for Evaluating Thoroughbred Yearlings – Quality Control of the Radiographic Image – Park, RD, DVM, PhD
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Chapter 89
Presale Radiographic Interpretation and Significance C. Wayne McIlwraith
Introduction The pre-purchase examination is one of the most important professional services offered to the horse industry be equine practitioners.1 While pre-purchase examinations are an important part of the equine veterinarian’s work, it has also been pointed out recently that they receive relatively little attention in modern textbooks.2 Pre-purchase examination of Thoroughbred yearlings at public auction is a special form of pre-purchase examination, in that the degree of thoroughness appropriate in other situations is not possible. In the USA, at least, yearlings are purchased on a visual physical and physical examination including: walking in hand, limb palpation, presale radiographs, and an upper tract endoscopic examination.3 Evaluation of conditions that could possibly lead to problems in later training is limited to outwardly visible or palpable changes and the radiographic examination. On the other hand, radiographic changes are often detected in young horses prior to training or athletic competition and, in many instances, these changes represent normal variations in the appearance or development of bone structures or are the result of acquired, congenital, or developmental abnormalities. It has been emphasized that the clinical objective should be the opinion of identifying an acceptable horse as much as the return of an unacceptable one.4 The prevalence of selected radiographic changes in young horses has been described for several breeds. Fifteen percent of 99 clinically normal Swedish Warmblood horses less than 3 years of age had a radiographic diagnosis of osteochondritis dis-
secans (OCD) of the intermediate ridge of the distal tibia in the tarsus.5 The reported prevalence of tarsal OCD in clinically normal young Standardbreds (including intermediate ridge of the distal tibia as well as lesions of the lateral trochlear ridge of the talus) range from 10% to 26% and these lesions were bilateral in about half of the horses examined.5–9 The prevalence of OCD has been reported in 14–29% of young Standardbreds. The prevalence of OCD in 80 yearling feral horses has been reported at about 5% for the fetlock and 1% for the tarsal joints.10 In Thoroughbred yearlings, Howard11 described the results of survey radiographs of 582 Thoroughbred sale yearlings. OCD lesions of the distal intermediate ridge of the tibia or distal lateral trochlear ridge of the talus were noted in 19 of 710 (3%) tarsi examined. A diagnosis of OCD or fragmentation of the proximal first phalanx was made for 89 of 1018 (9%) of the fore fetlocks and 46 of 700 (7%) of hind fetlocks examined. The same study also reported approximately 10% of fore and hind fetlocks affected by sesamoiditis (defined as large linear defects > 1 mm in width with an ill-defined margin). McIntosh and McIlwraith described the occurrence of femoropatellar OCD in three crops of yearlings between 11 and 24%: however, this included cases identified by clinical signs.12 Spike-Pierce and Bramlage have correlated racing performance with radiographic changes in the proximal sesamoid bones in 487 Thoroughbred yearlings.3 Overinterpretation is another hazard when examining yearling radiographs. Becht and Park reported on normal radiographic variations of the equine fetlock, carpus, tarsus, and the stifle. Selected normal
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radiographic variations in the fetlock included: flattening of the sagittal ridge of the third metacarpal bone with no subchondral lysis and/or fragment, flattening of distal palmar/plantar articular surface of third metacarpal/metatarsal bone with no subchondral lysis, variation in size and visibility of transverse ridge on the distal articular surface of the third metacarpal bone, false fracture line produced by superimposition of the base of the proximal sesamoid bones over the proximal palmar eminence of proximal phalanx, variation in location of vascular foramena through the dorsal cortex of the proximal phalanx, circular lucency representing the medullary cavity of the proximal and middle phalanges, medial proximal sesamoid bones being more cuboidal than lateral and fore proximal sesamoid bones being more elongated than hind, separate centers of ossification in the proximal aspect of proximal sesamoid bones, and distal border of hind proximal sesamoid bones being flatter than distal border of fore proximal sesamoid bones.13 Selected normal radiographic variations in the equine carpus included variation in number (0–3) and location of carpal fat pads, residual ulna on distal palmar aspect of radius, presence of first carpal bone, presence of fifth carpal bone, fossa for first carpal bone on palmar surface of second carpal bone without presence of first carpal bone, fossa for fifth carpal bone on palmar surface of fourth carpal bone without presence of fifth carpal bone, proximal-distal orientation of accessory carpal bone, variation in shape and size of transverse ridge of third carpal bone, circular lucency in mid-distal radius representing the radial fossa, and incomplete fusion between the radius and ulna on the lateropalmar aspect of the distal radius.13 Selected normal radiographic variation in equine tarsus included: flattening of the dorsal aspect of the medial trochlear ridge of the talus, visualization of talocalcaneal joint space, variation in the shape of the bone eminence at the distal aspect of the medial trochlear ridge of the talus, variation in visualization of the medial and lateral aspect of the intertarsal joint spaces, variation in shape of the distal intermediate ridge of the tibia, superimposition of proximal medial tuberosity of the talus over the distal intermediate ridge of the tibia, and visualization and clarity of the tarsal canal.13 The selected normal radiographic variations in the equine stifle included irregularity of proximal femoral trochlear ridges at patella in young horses, fusiform lucency in proximal aspect of the lateral femoral trochlear ridge, variation in visibility of extensor fossa in distal lateral femoral condyle for attachment of the long digital extensor tendon, varying degrees of irregularity of apophysis of the tibial crest, variation in the degree of roundness of the medial and lateral femoral condyles, and
variation in linear radiolucency in the intercondyloid fossa of the tibia at the area of attachment of the caudal cruciate ligament.13
Keeneland yearling study In the interest of defining what radiographic changes could be acceptable and did not affect racing performance, a larger study was carried out and most of the author’s statements in this chapter are based on these findings.14, 15 The goal of this study was to identify problems that were likely to limit racing performance, without unnecessarily condemning a yearling with minor radiographic changes. To do this, the radiographs of the front and hind fetlocks, carpi, tarsi, stifles, and fore feet in the radiographs of 1162 Thoroughbred yearlings were examined. Changes associated with racing performance as well as clinical problems or surgery through the 3-year-old year were obtained and correlated with the various changes identified. Fetlock joints Changes recorded in the fore fetlocks included proximal dorsal (Figure 89.1) and proximal palmar fragments of the first phalanx. Palmar fragments were
Figure 89.1 A dorsolateral-palmaromedial oblique view of a front fetlock joint with an OCD fragment off the proximal dorsal aspect of the first phalanx (arrow). Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
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Presale Radiographic Interpretation and Significance
(a)
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811
(c)
Figure 89.2 Flexed lateromedial views of the front fetlock showing a notch (a), a lucency (b), and fragment (c) at the dorsal aspect of the distal third metacarpus. Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
classified as non-articular or articular, with articular fragments evidenced by a visible defect in the joint margin or surface. Cystic lesions in the distal third metacarpus or proximal first phalanx were also recorded (defined as areas of increased lucency extending through subchondral bone). On the distal dorsal aspect of the third metacarpus, changes categorized included the presence of a well-defined semicircular notch at the proximal aspect of the sagittal ridge, irregular-shaped lucencies (type I OCD) and fragments (type II OCD) (Figure 89.2), as well as an evaluation of the distal sagittal ridge of the third metacarpus on the dorsopalmar and flexed lateromedial views for flattening or lucency (Figure 89.3). Medial and lateral proximal sesamoid bones of the forelimb were categorized as elongated if they were
(a)
> 2 mm longer than the other proximal sesamoid bone in the same limb and abnormal shape was defined as proximal, distal or abaxial enlargement of proximal sesamoid bones. Fractures of the proximal sesamoid bone were recorded as apical, abaxial, basal, mid-body, comminuted, or other. Periarticular bone production at the proximal or distal articulation of the fetlock joint was recorded as an osteophyte and bone production within the suspensory or distal sesamoidean ligaments where they attach to the proximal or distal palmar respectively of the sesamoid bone were recorded as an enthesophyte (Figure 89.4). Well-defined lucencies in the proximal sesamoid bones were recorded and the number of linear lucencies (regular and irregular vascular channels) was recorded for each proximal sesamoid
(b)
Figure 89.3 Dorsopalmar (a) and flexed lateromedial (b) views of a front fetlock with lucency on the distal aspect of the distal metacarpus. Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
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Figure 89.4 A dorsolateral-palmaromedial oblique view of the front fetlock with a large enthesophyte on the palmar distal aspect of the lateral sesamoid bone. Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
bone (Figure 89.5). Regular vascular channels were defined as linear lucencies that had parallel sides for the entire length and were ≤ 2 mm in width, whereas irregular vascular channels were defined as linear lucencies that had non-parallel sides for any portion of their length or were > 2 mm in width. Hind fetlock and sesamoid changes were classified in the same manner as the fore fetlock. Hind proximal sesamoids with moderately pointed proximal or distal ends seen on the oblique views were regarded as normal and were not categorized as having osteophytes or enthesophytes. The fetlock joints were the most common joints in which radiographic changes were observed. With 1127 fore and 1102 hind fetlock radiographic series examined, flattened areas on the distal palmar aspect of the distal articular surface of the third metacarpus were found on 461 (41%) yearlings and had no effect on racing performance, joint problems, or surgery. Flattening (110 yearlings, 10%) or lucency (196 yearlings, 17%) of the dorsal aspect of the distal sagittal ridge was also quite common and had no effect on racing performance, joint problems, or surgery.
All other fore fetlock changes observed affected less than 3% of the yearlings examined. Compared with the forelimb, proximal P1 fragments of the hind limb were more common. Lucencies and fragments on the dorsal aspect of the distal metatarsus were more common. Sagittal ridge and condylar changes were much less common and supracondylar lysis was not found in any hind limb. Supracondylar lysis was not seen in hind fetlock joints. The finding of palmar supracondylar lysis of the distal third metacarpus and its significance was a surprise and had not been noted as a finding among yearlings in the author’s experience (Figure 89.6). Only 14 of the 24 moderately or severely affected horses (58%) started a race compared with 82% of unaffected horses that started, making the odds of these horses starting a race one-third of those for unaffected horses (P < 0.01). Affected horses also placed in only 25% of their starts compared with 40% of starts placed for unaffected horses (P < 0.09). Two of the four horses with supracondylar lysis for which clinical follow-up was available, reported a fetlock problem. Only 8 of 14 (57%) affected horses started in a race making the odds of starting a race for yearlings with enthesophyte formation on fore proximal sesamoid bones one-third of those for unaffected horses (P = 0.04). For yearlings with enthesophyte formation on the hind proximal sesamoid bone there was also a lower tendency (P = 0.13) for starting a race with 9 of 14 (64%) affected horses starting a race. The percentage of starts placed (16%) was also significantly (P = 0.01) lower for these yearlings compared to horses without this change (40%) and earnings per starts were significantly reduced in the group with enthesophytes. A notch occurred on the sagittal ridge of the distal third metacarpus in 349 (31%) fetlocks examined. These had no effect on performance. Lucency occurred in 22 (2%) of yearlings and also did not have any effect. Because fragments and loose bodies only occurred in 9 (1%) of yearlings, it was difficult to say that we correctly found that they had no influence even though after statistical analysis there was no effect on racing performance nor on joint problems or surgery. The actual figures are interesting. Of the ones with a notch, 84% started (290 of 347 affected), of the 22 with lucency 15 (68%) started, and of those with a loose body or fragment (OCD), 7 of 9 or 78% started. Fragments of the proximal dorsal aspect of the first phalanx occurred in the front limb with 18 (2%) yearlings affected. With these fragments, 78% of those affected started a race but with only 27% of starts placed compared with 40% in the non-affected population (P = 0.07). Half of these horses (one out of two with follow-up) had surgery. With proximal palmar
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Plate 1 (Figure 4.5 – Volume 1) B cells
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Plate 13 (Figure 59.7 – Volume 1)
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Plate 21 (Figure 59.15 – Volume 1)
Plate 24 (Figure 59.19 – Volume 1)
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Plate 25 (Figure 59.20 – Volume 1)
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Plate 32 (Figure 59.27 – Volume 1) Plate 33 (Figure 59.28 – Volume 1) Plate 34 (Figure 59.29 – Volume 1)
Plate 35 (Figure 59.31 – Volume 1)
Plate 36 (Figure 73.4 – Volume 1)
Plate 37 (Figure 74.1 – Volume 1)
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Satge V
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Plate 49 (Figure 152.4 – Volume 1) Plate 50 (Figure 153.3 – Volume 1)
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Plate 53 (Figure 153.7 – Volume 1)
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Plate 52 (Figure 153.6a-c – Volume 1)
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Plate 55 (Figure 155.2 – Volume 1)
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Plate 83 (Figure 208.6 – Volume 2)
Plate 84 (Figure 208.8 – Volume 2)
Plate 85 (Figure 208.9 – Volume 2)
Plate 86 (Figure 208.12 – Volume 2)
Plate 88 (Figure 209.1 – Volume 2)
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(d)
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Maximum
Blood Flow
Minimum
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Apex Ovulation
)
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Maximum Minimum
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Serration Apex ? Septated Evacuation, HAF, Atresia
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Hours before expected evacuation
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Plate 101 (Figure 215.10 – Volume 2)
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9 12
6 3
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100 1
Apical area
10
2
Preovulatory Follicle
9
3
Basal area
8
% of mares (n=21)
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0 Serrated granulosa Vascularized theca
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7 6
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10 11 12
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Septated Evacuation
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F
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F
apex
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Normal Evacuation
Septated Evacuation
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Plate 104 (Figure 215.14a-f – Volume 2)
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Septated evacuation
Normal evacuation
apex apex
base
base
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(b) Blood flow
100
Septated evacuations 80 % of circumference
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40
20 -11
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Plate 105 (Figure 215.15a-b – Volume 2)
Normal evacuation
Septated evacuation
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2
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80 Pregnant 70 60 Nonpregnant
50
Prominence (score)
Color-Doppler signals
3.0 2.8 2.6 2.4
(a) Vascular indices 1.7 0.7
1.5 PI
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0 30 Hours after hCG treatment
Plate 108 (Figure 215.18a-b – Volume 2)
30 (b)
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Luteal Gland Development a)
B-Mode Ultrasonography
b)
B-Mode/Color-Doppler Ultrasonography
Early diestrus
Mid-diestrus
Late diestrus
Plate 109 (Figure 216.2a-b – Volume 2)
Days after ovulation
Sonogram of uterus and conceptus
Days after Sonogram of uterus ovulation and conceptus
Embryonic vesicle Blastoceole
Capsule
Day 10
Day 18 Inner cell mass
Embryonic vesicle
Ectoderm (trophoblast)
Bilamina (ectoderm & endoderm) Trilamina (ectoderm, mesoderm & endoderm)
Yolk sac
Amniotic sac
Day 12 Embryonic disc
Endoderm
Day 22 Embryo proper
Day 14 Mesoderm
Day 24
Chorion (SomatoPleure)
Day 16 Day 26
Plate 110 (Figure 217.2 – Volume 2)
Chorionic girdle
Allantoic sac
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Embryonic vesicle
Day 29 Allantochorionic placenta
Day 36
Fetus
Day 40
Umbilical chord
Day 45
Plate 110 (Figure 217.2 – Volume 2, continued) Orientation vs Dysorientation of Embryo Proper Ventral position of embryonic pole
Dorsal position of embryonic pole Ma
Ma
Ep Ep
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Ys
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Day 19
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Ma
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As
Ep As
Ep
Day 29
Ys
Day 28
Ma
Ma
Uc
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As
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Fetus Fetus Day 45
Plate 111 (Figure 217.3 – Volume 2)
Day 44
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Plate 113 (Figure 219.7 – Volume 2)
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Figure 89.5 Regular vascular channels defined as having parallel sides and ≤ 2 mm width (a) and irregular (b) vascular channels defined as having non-parallel sides or > 2 mm width were recorded from dorsolateral-palmaromedial and dorsomedialpalmarolateral oblique views of the fetlock. Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
Figure 89.6 Flexed lateromedial views of the front fetlock showing dorsopalmar narrowing due to severe palmar supracondylar lysis (double arrows). Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
P1 fragments, all of the five affected horses started with 15% of the starts placed compared with 40% in the unaffected population (P = 0.16). In the author’s opinion cystic lesions of the proximal aspect of the proximal phalanx are never a problem and cystic lesions of the distal metacarpus are rarely a problem. In the study, there were six cases of cysts in the distal metacarpus and two cases in the proximal first phalanx. Five of the eight (62%) started a race and it was concluded that there was no substantial effect on outcome (P = 0.22). Elongation and enlargement of the sesamoid bones had no substantial affect on outcome. Three of five horses affected with apical or abaxial sesamoid fragments started (60%) and 6 yearlings with other types of fractures showed no substantial affect on outcome examined. The author had usually considered chip fragments to be an indication for arthroscopic surgery prior to the study. Of horses with fragments off the proximal dorsal aspect of the first phalanx in the hind limb, 25 of 36 (69%) started and the odds of starting were half of those for unaffected horses. Two out of nine of these horses with follow-up (22%) had a problem compared to 5% that had a problem without a fragment. Four of the nine (56%) had surgery compared to 5% that had surgery without a fragment so the data implies that surgery would be a benefit. Plantar fragment of the hind limb had no detectable effect on racing performance, but there were clinical effects. With no lesion, 4% had a problem and 4% had surgery. Of the 20 yearlings with non-articular fragments (2%
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of those examined) 25% had a problem, 3 of 12 with follow-up (P = 0.0002) and 50% had surgery (6 of 12) (P < 0.0001). There were 45 yearlings with articular plantar fragments (4% of those examined) and 1 of 6 (17%) had a problem (P = 0.002) and 2 of 6 (33%) had surgery (P < 0.0001). Circular lucencies in the fore sesamoid bones were found in 164 yearlings (15%) and 133 (81%) started. However, two of five yearlings with two or more circular lucencies and clinical follow-up (40%) had a sesamoid problem compared with 7% that had a problem without a lucency (P = 0.02). Vascular channels in fore and hind proximal sesamoids were common with 1101 of 1127 (98%) of yearlings examined having at least one regular or irregular lucency, and most having two or three. These changes were not associated with any of the outcomes examined. In contrast, Spike-Pierce and Bramlage found a significant decrease in the mean number of starts and mean total earning at 2 and 3 years for Thoroughbred yearlings with more than two abnormally shaped linear defects in a proximal sesamoid bone compared with yearlings that had no linear defect.3 Although we attempted to follow similar definitions in our study, it is probable that there was some difference in the criteria to identify irregular vascular channels. Only 20 of 487 yearlings (2%) were described as having more than two abnormal linear defects in the study of Spike-Pierce and Bramlage but 366 of 1127 yearlings (32%) were described as having more than two irregular vascular channels in our study. It has been recently pointed out by Bramlage (personal communication, 2009) that in our study a ‘cone’-shaped entrance to the bone by the vasculature on the abaxial surface of the sesamoid bone was considered irregular; however, in their study these were regarded as normal. The author now uses more restrictive definition of irregular vascular channels.
no detectable effects on other outcomes examined. The most common finding on carpal radiographs was circular lucencies in the ulnar carpal bone (177 unilateral of 227 affected) seen on the dorsolateralpalmaromedial oblique view. Among affected yearlings, 183 of 227 (81%) started (P = 0.68) and there were no associations with any other outcomes, confirming that these changes on radiographs are insignificant. Seven of nine horses (78%) with carpal chip fragments started and no association was identified with any of the outcomes examined. Osteophytes were seen most commonly on the radial carpal (11 yearlings) and intermediate carpal bones (6 yearlings), with one each seen on the distal radius and third carpal bone. Of these 19 horses, 13 (68%) started (P = 0.17). Of four with follow-up, three had a problem (75%) compared with only 10% that had a problem when no fragment was present (P = 0.004). In addition, two of four with follow-up had surgery (50%, P = 0.01) compared with 3% that had surgery without a fragment. Because carpal chip fragments were so rare, below a 1% prevalence, as were the three carpal
Carpus The author has always been concerned with changes in the carpus of a yearling, as he has been particularly unimpressed with the results of arthroscopic surgery in cases of OCD. The term dorsal medial intercarpal joint disease was used in the Keeneland study if the radial carpal bone had a rounded appearance and/or thickened dorsal cortex or there were proliferative changes, enthesophytes, or fragments involving the radial carpal or third carpal bones (Figure 89.7).14 Thirty of 1130 yearlings with complete carpal examinations had signs of dorsal medial intercarpal disease (22 unilateral in these 30). Of the 30 affected, 19 started (63%) (P = 0.02). Analysis revealed that the odds of starting a race were one-third of those for unaffected horses (P < 0.01), but there were
Figure 89.7 Dorsolateral-palmaromedial oblique view of the carpus showing a thickened dorsal cortex of the radial carpal bone with proliferative changes and a distal fragment at the articular margin. These changes, regarded as signs of dorsal medial intercarpal joint disease, were shown to affect racing performance. Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
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Presale Radiographic Interpretation and Significance cysts recorded in the distal radius, ulnar carpal, and second carpal bone respectively (different yearlings), our ability to demonstrate significant effects on the outcomes examined had very low power. Tarsus Osteochondritis dissecans (OCD) has always been the most common issue when examining tarsal radiographs of sale yearlings. The author has never been very concerned about them. Arthroscopic surgery has always been recommended when there is synovial effusion present in a horse of any age. When looking at OCD lesions without effusion in a sale yearling, the advice has been that we do not know how often clinical signs will develop in training (we certainly have instances where they do not), but if clinical signs develop, the problem can be treated successfully. For this reason, some buyers elect to purchase and immediately have the problem treated surgically to obviate problems developing later in training. There were 1101 complete tarsal series. Thirtythree of 48 yearlings with radiographic changes in the intermediate ridge of the distal tibia had fragments visible; five of these were bilateral. Only one of the 15 yearlings with a defect or concavity alone was affected bilaterally. One yearling had a flattened area on the lateral trochlear ridge of the talus, four had defects or lucencies alone, and 10 additional year-
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lings had fragments with or without visible defects. All were affected unilaterally. Except for the most distal aspects, no fragments were observed from the medial trochlear ridge, but 12 yearlings had flattened areas and 7 had defects (2 bilateral) without a visible fragment. Seventy-two yearlings (6.5%) had some lucency or fragmentation of the medial malleolus, intermediate ridge of the distal tibia or lateral or medial trochlear ridge of the talus. At the most distal aspect of the medial trochlear ridge of the talus, 5 of 39 yearlings with a protuberance and 2 of 8 yearlings with fragments had these changes bilaterally. Of the 48 horses with concavity or fragmentation of the distal intermediate ridge of the tibia, there were no detectable effects on racing performance. However, 3 of 10 with follow-up (30%) had a clinical problem compared with 7% that had a problem without this change (P = 0.03). In addition, 3 of 10 (30%) had surgery compared with 1% that had surgery without this change (P < 0.0001). The changes most commonly observed in the distal tarsal joints were the presence of osteophytes or enthesophytes involving the distal intertarsal or tarsometatarsal joints (Figure 89.8). Both of these findings were usually unrelated. In the study reviewed here, osteophytes or enthesophytes involving the distal intertarsal or tarsometatarsal joints were the only changes significantly (P = 0.03) associated with starting a race; however, the magnitude of this effect was small (147 or 193 yearlings or 75% started) and no
(b)
Figure 89.8 Osteophytes and enthesophytes at the tarsometatarsal (a) and distal intertarsal (b) joints were the most common findings in the tarsus of yearlings but only had a small effect on the likelihood of starting a race during the 2- or 3-year-old years. Reproduced with permission from Kane AJ, Park RD, McIlwraith CW et al. Radiographic changes in Thoroughbred yearlings. Part I: Prevalence at the time of yearling sales. Equine Vet J 2003;5:354–65.
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association with the odds of starting was identified in the final multivariate model. Seven of 36 horses with clinical follow-up (19%) had a clinical problem compared with 6% that had a problem without osteophytes (P = 0.04). When tarsal bone wedging or collapse was observed it involved the third tarsal bone in 10 of 13 yearlings. Two of these were affected bilaterally. Of the 13 yearlings with wedging or collapse of tarsal bones, there was no significant association with starting a race, the percent of starts placed, or the clinical outcomes. Money earned with these changes was somewhat (P = 0.11) decreased ($1937 compared to $9335) and earnings per start were $440 compared with $1386 for horses without wedging (P = 0.15). Some previously published work in Quarter Horses (non-racing) divided tarsal bone collapse into two grades. In grade II (fragmentation of wedged bone), there was a negative effect on prognosis. Less than 2% of yearlings were affected with lucencies in the medial malleolus or changes along the curvature of the trochlear ridges of the talus and therefore there was no significant effects could be fully evaluated for any outcomes examined. Stifles The use of stifle radiographs has become more frequent in recent years and is now standard as part of the Thoroughbred yearling evaluation. The two main areas of concern are OCD changes on the femoral trochleas in the femoropatellar joint and cystic lesions in the medial femoral condyle in the femorotibial joint. It has been previously demonstrated by McIntosh and McIlwraith12 (1993) that defects on the trochlear ridges can heal even when clinical signs are present. The clinical detection of synovial effusion gives a strong indicator as to the degree of compromise in these cases. The presence of cystic lesions in the medial femoral condyles is more difficult. Because of numbers, we did not really answer this question well in the Keeneland study. There were 600 yearlings with a stifle series included in this study; however, the medial femoral condyle and proximal tibia could be clearly visualized in only 170 yearlings. Eighteen yearlings had fragments (four bilateral) and 16 had defects without a visible fragment (three bilateral) in the lateral trochlear ridge. Two yearlings had defects with fragments in the medial trochlear ridge. There were no cysts detected in the femoral condyles or proximal tibia, but a limited number radiographic series depicted this area. Four of 660 yearlings had flattened areas and there was no significant association with any outcome. Thirty-four of 660 yearlings (5%) had defects with or without fragments. Of five where
follow-up was obtained, two (40%) had surgery compared with a 1% chance of having surgery without a defect (P = 0.0008). There was no significant association with any other outcome. Fore feet Fore feet series were available for 300 of the yearlings. Sixteen of 33 yearlings identified with signs of pedal osteitis had bilateral changes. Other changes recorded in the feet included synovial invaginations (nutrient foramena) in the distal aspect of the navicular bone (18 yearlings, 5 bilateral), palmar fragments of the third phalanx (15 yearlings, 4 bilateral), focal lucencies or cysts in the third phalanx (7 yearlings), osteophytes or enthesophytes in the extensor process of the third phalanx (2 yearlings, 1 bilateral), extensor process fragmentation, dished appearance of the third phalanx, and rotation of the third phalanx. There were no detectable effects on any outcomes examined.
Summary The priority of the study reviewed here was to examine each radiographic change in detail without combining different types of changes in the same location (e.g., flattening, lucency, and fragmentation in the fetlock) or similar changes in different locations of the same joint (e.g., fragmentation of the distal intermediate ridge, lateral or medial trochlear ridges of the talus) into broad categories. This limited the power of the analysis to detect significant effects for those changes that only affect a small number of yearlings. However, it preserved the detail with which radiographs are routinely examined at yearling sales and should be more useful for veterinarians trying to predict the effect of a specific radiographic change during sale examination. Many of the changes that can be of concern clinically were rare in this study and so along with difficulty in getting follow-up in all cases made it difficult statistically to demonstrate an effect on performance or clinical outcomes. Although return of questionnaires limited the analysis for clinical outcomes and prevented adjustment of racing performance analysis for surgical intervention, it is important to recognize that further studies focused on some of these conditions will no doubt demonstrate associations with performance. This study confirmed that some lesions established in the literature as incidental findings (e.g., ulnar carpal bone lucencies) are indeed so. Other changes clearly not associated with lower performance are flattened areas on the sagittal ridge or condyles of the distal third metacarpus, circular lucencies, and vascular channels in sesamoid bones.
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Presale Radiographic Interpretation and Significance New areas of concern were palmar supracondylar lysis of the distal third metacarpus, enthesophyte formation on proximal sesamoids, and fragmentation of proximal dorsal P1 in the hind limb and dorsomedial intercarpal joint disease. It also needs to be recognized that with the current situation where foals are radiographed when weanlings and many are treated surgically, the population of horses presented for sale now could be very different than those included in our study with data collected from 1993 to 1996. Radiographic examination of sale yearlings is best conducted with using results of this study in parallel with personal clinical experience. On the other hand, the veterinarian new to the evaluation of sale yearlings now has some figures to draw from regarding certain radiographic changes as incidental findings.
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References 1. Moyer WA, Werner H. Risk reduction in the reporting of the purchase examination. Proc Am Ass Equine Practnrs 1999;54:24. 2. Bladon BM, Maine JPM. Clinical evidence in the evaluation of presale radiography: Are we in a desert on a horse with no name? Equine Vet J 2003;35:341–342. 3. Spike-Pierce DL, Bramlage LR. Correlation of racing performance with radiographic changes in the proximal sesamoid bone of 487 Thoroughbred yearlings. Equine Vet J 2003;35:350–3. 4. Bramlage LR. Osteochondrosis and the sale horse. Proc Am Ass Equine Practnrs 1993;39:87–9. 5. Hoppe F. Osteochondrosis in Swedish Horses, A Radiological and Epidemiological Study with Special Reference to
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Frequency and Heredity. PhD Thesis, Swedish University of Agricultural Sciences, Uppsala. Alvarado AF, Marcous M, Breton L. The incidence of osteochondrosis in a Standardbred breeding farm in Quebec. Proc Am Ass Equine Practnrs 1989;35:293–307. Grondahl AM. The incidence of osteochondrosis in the tibiotarsal joint of Norwegian Standardbred trotters – a radiographic study. Equine Vet Sci 1991;11:272–4. Jorgensen HS, Proschowsky H, Falk-Ronne J, Willeberg P, Hesselholt M. The significance of routine radiographic findings with respect to subsequent racing performance and longevity in Standardbred trotter. Equine Vet J 1997;29: 55–9. Sandgren B. Bony fragments in the tarsocrural and metacarpo- or metatarsophalangeal joints in the Standardbred horse – a radiographic survey. Equine Vet J Suppl 1988;6:66–70. Valentino LW, Lilich D, Gaughan EM, Biller DR, Raub RH. Radiographic prevalence of osteochondrosis in yearling feral horses. Vet Comp Orthop Traumatol 1999;1: 151–5. Howard BA, Embertson R, Rantanen NW, Bramlage LR. Survey radiographic findings in thoroughbred sales yearlings. Proc Am Ass Equine Practnrs 1992;38:397–402. McIntosh SC, McIlwraith CW. Natural history of femoropatellar osteochondrosis in three crops of Thoroughbreds. Equine Vet J Suppl 1993;16:54–61. Becht JL, Park, RD. A review of selected normal radiographic variations of the equine fetlock, carpus, tarsus and stifle. Proc Am Assoc Equine Pract 2000;46:362–64. Kane AJ, Park RD, McIlwraith CW, Rantanen NW, Morehead JP, Bramlage LR. Radiographic changes in Thoroughbred yearlings. Part 1: Prevalence at the time of the yearling sales. Equine Vet J 2003;5:354–65. Kane AJ, McIlwraith CW, Park RD, Rantanen NW, Morehead JP, Bramlage LR. Radiographic changes in Thoroughbred yearlings. Part 2: Associations with racing performance. Equine Vet J 2003;35:366–74.
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Section (s): Appendix
90.
Development of a NICU Nathan M. Slovis and Lynne M. Hewlett
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Diagnostic Laboratory Resources and Critical Care Supplies Nathan M. Slovis and Wendy E. Vaala
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Formulary Robert P. Franklin
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Hematology and Biochemical Reference Values Robert P. Franklin and Joan Palmero
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SI Conversion Table
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Chapter 90
Development of a NICU Nathan M. Slovis and Lynne M. Hewlett
Today the public demands and insists in more specialized care for their animals. This specialized care has resulted in the growth in the size and number of referral equine hospitals around the world. It is within these hospitals that advanced patient standards of care are being practiced every day. However, many of these hospitals are utilizing their stalls as both a temporary neonatal intensive care unit (NICU) during the breeding season and regular hospital stall facility during the off-season. Utilization of a stall for dual purposes can result in an increased number of nosocomial infections among our neonatal patients. Neonates by definition in the authors’ hospital are foals < 7 days of age. Neonatal foals at birth have a naive gastrointestinal tract and immune system. Neonates are therefore inherently more susceptible to acquire nosocomial infections. That is why it is imperative that, if a hospital is planning on developing an NICU, then a section of the hospital floor plan (or individual stalls) must be dedicated to the treatment and housing of the sick neonatal patient. This designated section cannot be utilized for any other purpose but for the treatment of critically ill neonatal foals. In the authors’ hospital setting the NICU is utilized for 7 months of the year. The remainder of the year the NICU is shut down and disinfected appropriately in anticipation for the following foaling season. No animal or human traffic is permitted in and out of the NICU during the off-season. Many equine hospitals cannot afford to keep a section of their hospital dedicated as an NICU. If this is the case, then during the breeding season it is strongly recommended that a group of stalls or area be designated for the use of sick neonatal patients only. No other hospitalized patient should utilize these stalls during the specified time that is agreed upon. NICUs were developed in the 1950s and 1960s by human pediatricians to provide better temperature, respiratory and nutritional support, isolation
from infectious diseases, and access to specialized equipment and resources. Some NICU patients such as low birth-weight infants require respiratory support ranging from supplemental oxygen to continuous positive airway pressure or mechanical ventilation. Public access to human and veterinary NICUs is limited, and staff and visitors are required to take precautions to reduce transmission of infection. An NICU in a veterinary hospital is typically directed by one or more board certified internal medicine specialists and staffed by registered veterinary technologists and veterinary technicians. Nearly all veterinary schools in the United States have NICUs. Many equine practitioners have recognized the importance of the NICU and would like to organize and maintain such units within their hospital setting. This chapter provides details of how to design, staff, and provide instruments in an equine NICU whether the caseload is 10 or 400 neonates per season.
Facility The average caseload per year will dictate the size of the NICU. Cinder block construction for stalls provides the best surface for disinfection between patients. Wood stalls are very difficult to clean although painting wood surfaces with two coats of marine varnish or SPAR polyurethane can provide an effective seal and greatly enhance the ability to properly disinfect surfaces. The authors’ NICU consists of 13 stalls measuring 12 ft × 14 ft each. An additional 12 ft × 14 ft room is stationed at the end of the barn which is used for storage of stall cleaning equipment, straw and hay. Stall cleaning utensils are hung on the wall in an orderly fashion with color-coded labels for each individual stall (Figure 90.1). Equipment includes pitchfork, rake, broom, and scrubbing brush, all of which are dipped into a disinfection bath situated below the racks after every stall cleaning. Bales of hay and straw are placed on the opposite side of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 90.1 All stalls in the NICU have their own designated stall cleaning instruments which are hung in an orderly fashion in the storage room. Having individual stall cleaning instruments may help decrease the incidence of nosocomial infection within the NICU.
the room, away from the stall cleaning equipment to prevent contamination. Sixteen-foot wide aisles provide easy access for staff as well as mares with foals. Each stall is equipped with in-wall oxygen outlets which eliminates the need for portable tanks (Figure 90.2). In addition, multiple electrical outlets are situated conveniently on the outside of each stall. Stall flooring consists of a seamless textured rubber flooring which helps with the biosecurity and cleaning of the stalls. Having a seamless flooring system reduces the chances of colonizing pathogenic bacteria compared to porous or non-seamless flooring systems. The seamless flooring should possess bacterial- and mildew-resistant properties. Every stall has a window for optimal natural ventilation. An adjustable window that can be partially or completely opened is ideal. In addition, cross-ventilation is improved when the main doors on either side of the barn are open. The NICU should have a separate heating and cooling unit installed to save on expenses during the off-peak season. In the authors’ NICU, hardware for an optional containment system is permanently installed in each stall to separate the mare from the foal should this become necessary. The divider panels which complement this system measure 5 ft 9 in. in length and 3 ft 9 in. in height (Figure 90.3). They are made of steel and are extremely durable, portable, and easily cleaned. Small sinks are located on each side of the barn for ease of hand washing. All sink stations in the NICU have permanently installed antibacterial soap dispensers as well as disposable paper towel dispensers within reach. An alternative to traditional hand-operated faucets with individual hot/cold water knobs is the use of knee-operated or large lever
Figure 90.2 Oxygen is supplied to every stall in the NICU without the use of stall-side portable tanks. All patients receiving oxygen must have a humidifier attached to help decrease the incidence of mucosal injury associated with dry air. Most foals receive between 4 and 6 L/min of oxygen via an indwelling nasal oxygen line.
Figure 90.3 A containment system that can be easily removed from the stall if not needed is essential. This system is composed of steel and is used to separate the mare and foal for ease of fluid and oxygen administration.
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Technical staff
Figure 90.4 Plenty of organized cabinet space is a necessity for the NICU.
arm faucets. The utilization of the non-traditional faucets may help decrease the incidence of recontamination of the veterinary staff hands. The aisle and stalls should provide for adequate drainage, giving careful consideration to the direction of the drainage flow, and the distance it covers. Liquid waste and water flowing out of a contaminated stall into the barn aisle can be a source of cross-contamination. A garden hose on both ends of the NICU should be installed, with provision of both hot and cold water. Our NICU design includes a large workstation situated in the center of the barn for easy access to either side. Workstations should include a refrigerator, microwave, sufficient cabinet space for oral and injectable medications with syringes, additional cabinets for supplies, a double sink, and large cabinet for fluid storage (Figure 90.4). The authors’ facility has two additional rooms in the NICU. The first is a large laundry room with a high-efficiency washer and dryer, and additional shelving units for miscellaneous items such as fleeces and foal beds. Another large sink is located in this room for items considered soiled or dirty. The importance of having a large storage room to keep the NICU looking tidy and clutter-free cannot be over-emphasized. A second room contains additional cabinets and large shelving units with a double sink. This room may be used to prepare for procedures such as a soapy enema or uterine lavage. It is large enough to store extra equipment including two ventilators, an EKG machine, and a crash cart. Hooks secured to the wall keep items such as muzzles and extra halters organized. Barns should have good access lighting outside for optimal visibility. Furthermore, a loading ramp located nearby is invaluable for staff trying to unload and load patients in a time-efficient manner.
The technical staff in an organized NICU includes technicians and barn cleaning personnel in addition to the veterinarians. This group might include certified or licensed veterinary technicians and other technicians with extensive ‘hands on’ experience. Training new technicians to give the precise nursing care required in an NICU is essential to the success of the clinic. At the authors’ institution, technicians receive 2 weeks of intensive training with a senior technician. Training topics covered include neonatal nursing care techniques and principles, guidelines for how to perform a physical examination, basic laboratory procedures, a review of general medicine topics, pertinent safety instruction for staff, patient handling and restraint, infectious disease protocols, as well as a review of how to care for special needs patients such as those suffering neurological impairment or those requiring intubation and mechanical ventilation. In addition, emphasis is placed on patient assessment, intravenous catheter care, fluid therapy monitoring (including care and maintenance of fluid lines), nutrition guidelines and delivery techniques, nasogastric tube feeding, principles of sterile technique, and organization of the emergency crash carts and kits. Other topics of importance include a discussion of commonly used medications and the routes of administration, bandage application including abdominal support band application, oxygen set-up and delivery, care and maintenance of urinary and thoracic catheters, and care of the post-foaling mare. All technicians are trained on whole blood collection and delivery and basic physical therapy. Basic knowledge of NICU equipment is required, including proper use of the suction machine, capnograph, glucometer, intravenous fluid pumps, EKG/blood pressure monitor, and ultrasound machine. Every technician receives a training manual which explains expectations and includes guidelines, charts, and diagrams as well as color pictures which can be used as an easy reference. Furthermore, technicians are taught the importance of correct and accurate documentation in patient charts. The average caseload ratio in the authors’ NICU is three or four foals per technician, of which at least one foal is recumbent. New technicians must successfully pass a written self-assessment test (passing score is 80% or higher) as well as a didactic test by the end of their 2week training session before permission is granted to work in the NICU. A senior technician as well as an intern is present at all times on every shift to ensure quality care for all patients. Technicians are required to wear designated clothing in this facility and must change into clean attire should they become soiled.
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Strict adherence to infectious disease protocol is of paramount importance and, therefore, no technician may work in another area of the clinic during their shift while assigned to the NICU.
Equipment The intensive nursing care provided in the neonatal ward requires a list of items necessary to promote the well-being and comfort of each foal. Recumbent foals require additional equipment and supplies to prevent decubital ulcers and head trauma should they be prone to seizures. The authors’ facility places R a pressure relieving mattress (Tempurpedic ) with a custom-made washable cover under the recumbent foal. The washable cover is a necessity for keeping the mattresses from becoming soiled. Additional padding is placed on the sides of the bed to minimize head trauma. These foam pads covered in vinyl are durable, washable and measure 26 in. wide by 43 in. long by 3 in. thick (Figure 90.5). Keeping the sick recumbent neonate in sternal recumbency is of the utmost importance in maintaining proper ventilation; therefore an additional V-shaped pad is placed under the foal, with the front legs extended forward to prevent the foal from lying lateral. The NICU must be well-stocked with pillows, plastic and cotton pillowcases, fleeces, blankets, and underpads to keep the patient comfortable and dry. Heat lamps, and heating pads with soft covers, may be needed to aid in temperature regulation. Although thick fleece
pads are convenient, they may not be ideal material to have in an NICU because of the straw and debris that become entangled and are difficult to remove during washing. Heat lamps should be secured to a fixture which is solid and secured to the stall in such a fashion that there is no risk of it dislodging onto the bedding or the patient. The authors’ NICU uses a heavy metal bracket with two heat lamps attached that is secured to the gate with permanent hooks welded on to the bracket. Each stall in this department has a stainless steel stall-side cart on wheels (Figure 90.6a,b). The cart has two shelves in addition to a drawer. Items needed for the patient are placed on the cart with additional supplies such as extra needles, tape, pens, and documentation papers placed in the drawer. Only a box of examination gloves and the patient chart are placed on the top shelf. A bottle of disinfectant hand wipes and a sharps container are secured to the sides of the cart. These carts are invaluable to keep technicians from running back and forth to the main barn workstation for supplies, thereby reducing foot traffic in and out of the stall, while maximizing efficiency. Additional time-saving NICU equipment includes fluid pumps and fluid harnesses. Fluid pumps promote accuracy when delivering exact time-and-volume sensitive fluids such as total parenteral nutrition (TPN), dopamine, dobutamine, or lidocaine infusions. Fluid harnesses, equipped with a ring to adhere to the fluid line, greatly aid in the delivery of fluids to the active foal (Figure 90.7). The harness and ring prevent the fluid line from pulling on the intravenous catheter.
Infectious disease control strategies
R Figure 90.5 Padding: foals are placed on a Tempurpedic mattress inserted into a waterproof cover, topped with fleece. Side pads are placed between the bed and the wall to prevent head trauma. These pads measure 26 in. × 43 in. × 3 in. Side pads and V-pads are purchased from Dandy Products; Ohio, USA. Website: www.dandyproductsinc.com
The vast majority of patients who have access to veterinary services today are healed. There are some, however, who suffer unintended consequences of care, such as healthcare-associated infections (HAI). To ensure that such life-saving care does not result in HAI, modern veterinary care has developed an extensive system for infection prevention. Regardless of the approaches taken, veterinary care facilities must strive for 100% adherence to the institution’s infection control strategies. To achieve this caliber of adherence, proper education of the staff will be needed to succeed. The transmission of infectious agents requires three elements: a source (or reservoir) of the infectious agents, a susceptible host with a portal of entry receptive to the agent, and a mode of transmission for the agent. Identification of areas or processes where transmission of pathogens is likely to occur (control points) and implementation of measures aimed at
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Figure 90.7 The use of fluid harnesses help prevent foals from breaking the fluid lines and decreases the incidence of foals from pulling out their intravenous catheters. Fluid harness sets may be obtained from Mila International, Florence KY, US (www.milaint.com).
(a)
(b) Figure 90.6 a,b A stainless steel cart is placed in front of every NICU stall. All carts are equipped with disinfecting hand wipes, containers for sharps, examination gloves, lubricant, thermometer, patient’s medical records, and other necessary equipment. The carts are very useful in reducing foot traffic both into and out of the stall.
minimizing the possibility of such transmission, while allowing for reasonable flow and function within the veterinary hospital or animal facility, is an important component of biosecurity.
To help identify a potential reservoir for an infectious disease all foals arriving to the authors’ ICU > 24 hours of age are tested for salmonella and rotavirus in their feces. Foals are then routinely tested daily for a total of three salmonella cultures and then tested weekly until time of discharge. Before implementing disease prevention and control strategies, several issues need to be considered which include the risk, feasibility, cost, and effectiveness. Risk can be defined by the potential for exposure. To help better understand the risks of the animal to disseminate a disease there is a need to know the past disease status and exposure status in order to best implement targeted control measures for contagious disease. Admittance-based alert systems to notify infection control and clinical personnel about readmitted or transferred patients with a history of infection or colonization, and routine assessment of education and training among personnel, can improve infection prevention at the facility level. One such mechanism would be self-certification by the owner or representative of the animal. The owner/authorized caretaker/transporter would sign a form and state the past history of the animal in relation of exposure to a specific disorder. The reason for such self-certification is the inability to establish risk level without this type of information, as those receiving the animal would have no mechanism to gather this information other than through self-certification for animals that have no overt signs of disease at the time of presentation to the facility, yet the animal could have had exposure to contagious disease agent that would pose a substantial risk to other animals in the facility if not managed accordingly. In the authors’ ICU before a foal can be admitted the farm’s personnel or owner must indicate if
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the foal has been housed in a facility that has noticed an increased number of foals with diarrhea. If yes, then specific personal protective equipment (disposable booties and booties) must be worn when treating this foal. Foot baths with a phenol disinfectant would also be placed outside the foal’s stall. All animals arriving to a veterinary facility should be classified by its relative risk level. The risk level will help determine where the foal is to be housed, stall set-up, and what personal protective wear must be worn when handling the foal. Risk levels can be adjusted for a patient as warranted (i.e., test results from fecal cultures, etc.). Table 90.1 represents the patient classification system used at the authors’ hospital. Table 90.2 lists the guidelines to be strictly adhered to when caring for each caution level. Hand hygiene in the authors’ ICU consists of regular hand washing with alcohol antiseptics. Alcohol solutions containing 60% to 95% alcohol are most effective, and higher concentrations are less potent because proteins are not easily denatured in the absence of water. Alcohol antiseptics have been documented in the prevention of healthcare-associated pathogens. These products have also been noted to be more effective for standard hand washing or hand antisepsis by healthcare workers than soap or antimicrobial soaps. What volume of alcohol-based hand hygiene product is needed can be extrapolated from several studies. The ideal volume to be applied to the hands is not exactly known and is affected by the type of alcohol used and its concentration. However, it can be speculated that if hands feel dry after rubbing hands together after 10–15 seconds, an insufficient volume of product was likely applied. Interestingly, because alcohol-impregnated towelettes contain limited amounts of alcohol, their effectiveness is comparable to soap and water. It therefore may be recommended to use more towelettes when utilized for hand antisepsis. In a field trial of alcoholbased hand rubs (low-viscosity rinses, gels, and foams) it was noted that ethanol gel was slightly more effective than a comparable ethanol solution at reducing bacterial counts on the hands of healthcare workers. Once an animal is discharged from the authors’ ICU the stall will be cleaned and disinfected with a phenolic compound. The stalls are then cultured for salmonella. Once a negative result is obtained the stall will be reopened to admit patients.
Cost capturing In the authors’ ICU there are several fixed costs associated with (i) the cost for hospitalization and
Table 90.1 Example of patient classification system Caution Level Yellow The following patients will be considered medium risk and will be identified with a yellow sticker on the stall, medicine box and chart:
r r r r r r r r
Colic patients Inpatients presenting to medicine from surgery Surgical patients requiring inhalation anesthesia (excluding orthopedic cases) Immunocompromised patients Newborn foals Foals Nurse mares Recipient mares that are under quarantine, unknown disease status. These mares will be identified with a ‘yellow’ tag.
Caution Level Orange The following patients will be considered medium-high risk and will be identified with an orange sticker on the stall, medicine box and chart:
r r r r r
Newborn foals less than 24 hours old, presenting with diarrhea with no farm history of problems. Joint ill foals less than 7 months of age Patients with bone infections Patients that have one of the following, of unknown origin: fevers greater than 102◦ F low WBC (less than 4800) loose manure Recipient mares that have been exposed to a horse that tested positive for an infectious disease but are testing negative. These mares will be identified with an ‘orange’ tag.
Code Red: must be moved to an isolation facility The following patients will be considered high risk and will be identified with a red sticker on the stall, medicine box and chart:
r r r r
Patients positive for infectious disease Suspect neurological patients for infectious disease Patients greater than 24 hours of age, presenting with diarrhea Patients that have two of the following, of unknown origin: fevers greater than 102◦ F low WBC (less than 4800) loose manure
All inpatients shall be handled with gloves. Gloves shall be easily accessible and shall be changed as necessary during examination, procedures, treatments, and between each patient. Staff must wash hands or use available hand sanitizers between each patient. Doctors may examine a patient without gloves, only if he/she feels that it is vital to the proper examination of the patient. The doctor must wash hands before examining another patient.
(ii) the cost for the daily doctor’s progress reports (i.e., physical examination and patient management). In ordered to capture the costs associated with technical staff the patients will be charged according to the patient level of care. At the authors’ hospital patients are classified according to one of six levels of care. For example, patient care level 1 is associated with an animal that requires only b.i.d. TPR and oral
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Table 90.2 Guidelines to be adhered to in caring for patients according to caution level
Table 90.2 Guidelines to be adhered to in caring for patients according to caution level
Caution Level Yellow
Code Red
r r r r r
r r r r r r r r r r r r
Wash hands. Prepare all oral and IV medications in new, properly labeled syringes. Put on a single pair of gloves. Put on booties. Enter stall; administer all treatments in the following order: 1 Pulse and respiration 2 IV treatments 3 Oral treatments 4 Any other treatments required 5 Temperature shall be taken last. Exit stall and close stall door. Discard all oral syringes. Place all IV syringes on stall ledge. Clean thermometer with alcohol swab. Remove booties and clip to garbage can, if in good condition. Remove gloves and discard. Disinfect stethoscope. Dip feet in dip pan. Take syringes and thermometer to treatment room. Discard of all other used IV syringes in sharps container. Store heparin flush syringe and thermometer in medicine box. Wash hands.
All heparin flush syringes shall be changed each 8-hour shift. A new syringe shall be used if the current syringe is dropped in a stall. A new needle shall be used for all treatments. Caution Level Orange
r r r r r
r r r r r r r r r r r r r
Wash hands. Prepare all oral and IV medications in new, properly labeled syringes. Put on a single pair of gloves. Put on disposable gown, and then put on booties. Enter stall; administer all treatments in the following order: 1 Pulse and respiration 2 IV treatments 3 Oral treatments. 4 Any other treatments required 5 Temperature shall be taken last. Temperature shall be taken. Exit stall and close stall door. Discard oral syringes into garbage can. Discard IV syringes into sharps container. Heparin flush syringe may be stored in the box on the stall ledge. If used, clean thermometer with alcohol wipes and store in box on stall ledge. Remove booties and clip to garbage can, if in good condition. Remove disposable gown and discard. Remove gloves and discard. Disinfect stethoscope. Use hand sanitizer to sanitize hands. Dip feet in dip pan. Wash hands.
All heparin flush syringes shall be changed each 8-hour shift. A new syringe shall be used if the current syringe is dropped in a stall. A new needle shall be used for all treatments.
r r r r r r r
r r r r r r r r r r r r r r r
Wash hands. Prepare all oral and IV medications in new, properly labeled syringes. Enter red box or area in front of stall door. Put on first pair of gloves. Put on disposable gown and then put on booties. Put on second pair of gloves over the first pair of gloves. Enter stall; administer all treatments in the following order: 1 Pulse and respiration 2 IV treatments 3 Oral treatments. 4 Any other treatments required 5 Temperature shall be taken last. After treatments have been completed, remove outer layer of gloves. Hold discarded gloves, inside out, in one hand. Open stall door, exit and close stall door. Discard outer layer of gloves and oral syringes in garbage can. Discard IV syringes into sharps container. Heparin flush syringe may be stored in box on the stall ledge. If used, clean thermometer with alcohol wipes and store in box on stall ledge. Dip feet, then remove booties and clip to garbage can, if in good condition. Remove disposable gown and discard. Remove inner layer of gloves and discard. Disinfect stethoscope. Use hand sanitizer to sanitize hands. Submerse feet in dip pan. Exit red box or area in front of stall door. Wash hands.
Figure 90.8 Large stretcher to aid in moving the recumbent foal from the horse trailer into the stall.
medications that do not exceed b.i.d. administration. Care level 6 would be assigned to a recumbent ICU foal that requires continuous rate infusion of fluids and treatments that may be required every 1 to 2 hours. These patient level of care costs are calculated
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(a)
(b)
Figure 90.9 a,b Example of a well-stocked resuscitation cart and fishing tackle box containing emergency drugs and syringes. Table 90.3 Resuscitation kit contents
r r r r r r r
r r r r
r r r r r
Full oxygen tank (not needed if oxygen is supplied stall side) Ambu resuscitator with a 3 ft corrugated tubing tail piece (Jorgenson www.jorvet.com, 1-800-525-5614 USA) Nasotracheal tubes used for intubation (7, 8, and 9 mm for foals) (Surgivet, www.surgivet.com, 1-888-745-6562 USA) and 10 mL syringe to inflate the cuff Fluids: Hetastarch, Hypertonic saline, and Lactated Ringer solution or Normosol Fluid administration sets, including a pressure bulb for rapid administration of fluids Fluid additives: 50% Dextrose and 50% Magnesium Sulfate Oxygen supplies Tubing: rubber tubing 3/16 in. diameter which connects the nasal cannula to the humidifier (Grogans Health Care, www.grogans.com, 1-800-365-1020 USA) R Levin tube (item number 0044180 www.crbard.com, Nasal cannulas: used for intranasal oxygen administration 18 Fr Bard 1-800-367-2273 USA) and 24 Fr Kendall Harris Flush Tube (item number 155733, www.kendallhq.com, 1-800-962-9888 USA) Humidifier and connectors Intravenous catheters 16 g × 3 in. MilaTM over-the-needle polyurethane (Mila International, www.milaint.com, 1-888-645-2468 USA) 16 g × 5 14 in. MilaTM over-the-needle polyurethane 16 g × 16 cm ArrowTM or MilaTM antimicrobial over-the-wire polyurethane (Arrow International, www.arrowintl.com, 1-800-523-8446 USA) Feeding tubes 24 Fr, 60 in. long (Kendall Harris Flush Tube (item number 155733, www.kendallhq.com, 1-800-962-9888 USA) or stallion urinary catheter Mila International has a variety of feeding tubes Sedatives for the mare: Xylazine, Detomidine and Romifidine Tackle box with emergency medications already drawn up and labeled 1. Epinephrine 1 mg/mL (asystole or bradycardia). Dose: 0.01 to 0.02 mg/kg i.v., have three syringes with 1 mL of the medication drawn up 2. Doxapram 20 mg/mL (respiratory stimulant), Dose: 0.5 mg/kg i.v., have two syringes with 1 mL of the medication drawn up 3. Glycopyrollate (anticholinergic agent used to treat bradyarrhythmias). Dose: 0.0022 mg/kg i.v., have two syringes with 1 mL of the medication drawn up 4. Yohimbine (alpha-2-adrenergic antagonist). Used for dystocias or C-sections in which the mare is heavily sedated with an alpha-2-adrenergic. Dose 0.75 mg/kg i.v., have bottle available, give 1.5 mL i.v. per 100 lb (45 kg) 5. Naloxone (narcotic antagonist). Used in C-section foals to help reduce the endogenous opioids that cause mental depression. Dose 5 mg i.v., have two 4 mg bottles available 6. Prednisolone sodium succinate. Dose 2 mg/kg i.v., have three 100 mg bottles available 7. Lidocaine 2% (used for ventricular tachycardia); Dose 1 mg/kg i.v. 8. Furosemide (diuretic) Used for pulmonary edema): Dose 1 mg/kg i.v., have bottle available Umbilical clamp and Nolvasan umbilical dip (0.5% concentration) Gloves (sterile and non-sterile) Eye lubricant Suture kit (suture holders, Mayo scissors, 2 small hemostats, scapel handle, Metzenbaum scissors) and scalpel blades Tracheostomy tubes (plastic preferable) used if upper respiratory lesion that does not permit proper airflow from nares: 5 mm ID and 7.3 mm OD, 7 mm I.D and 10 mm OD
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Development of a NICU into an equation that is associated with an average technician hourly salary (including health benefits, etc.) and a designated profit margin.
Resuscitation kits No NICU is complete without ready access to a wellstocked resuscitation kit and stretcher (Figure 90.8).
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These kits must be organized and contain all the medications and supplies needed to provide emergency resuscitation (Figure 90.9a,b). All drugs and supplies should be clearly labeled. Table 90.3 lists the essential items for the resuscitation kit.
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Chapter 91
Diagnostic Laboratory Resources and Critical Care Supplies Nathan M. Slovis and Wendy E. Vaala
This chapter serves as a brief summary of practical laboratory assays and critical care supplies used frequently to monitor and care for late-term mares and their foals.
Selected laboratory assays There are several tests, readily performed by most clinical laboratories, that aid the veterinarian in making important diagnostic decisions for the health and well-being of the foal. These tests include milk electrolyte assays used to predict parturition and assess fetal maturity, neonatal isoerythrolysis (NI) screening tests to identify pregnancies and/or foals at risk for NI, and a variety of tests designed to measure concentrations of immunoglobulin G (IgG) in foal blood or mare colostrum. Several commonly used point-ofcare (POC) monitors used to measure glucose and lactate are also discussed. To ensure consistent and reliable results for any diagnostic assay, the laboratory should routinely perform internal quality controls on a regular basis. This test validation procedure usually requires the use of high and low control samples or positive and negative controls (depending on type of test) to ensure accurate testing of patient sampless, especially when sample values fall near the upper or lower limits of the assay being used. Milk electrolyte concentrations Pre-foaling mammary secretion electrolytes are related to fetal readiness for birth in normal pregnancies. The three electrolytes that can be assessed readily in the mare’s colostrum are sodium, potassium,
and calcium. The concentrations and proportions of these electrolytes can be used to predict impending parturition and to help assess fetal maturity in the late-term mare. Evaluation of fetal maturation becomes critical when managing pregnancies that may require emergency or elective induction or cesarean section. Changing concentrations in mammary secretions have also been used to help predict delivery. Calcium (Ca) concentrations in mammary secretions rise sharply 24–48 hours prior to parturition and should attain Ca concentration greater than 40 mg/dL, which is equivalent to 400 ppm, 10 mmol/L, and 20 mEq/L of calcium. If a serum chemistry analyzer is used to determine milk electrolyte concentrations it is important to note that calcium, and not calcium carbonate (CaCO3 ), is the analyte being measured. Water hardness test kits, which are often used stall-side to predict parturition, measure only CaCO3 . If a water hardness test kit is used, calcium carbonate levels of 300–500 ppm are indicative of impending parturition within 12–18 hours. To convert ppm calcium carbonate to ppm calcium, results should be divided by 2.5 (since the molecular weight of calcium carbonate is 100 and the molecular weight of calcium is 40). However, adjustments often need to be made for dilution factors as well, so be wary when relying on water hardness test kits. The most accurate method is to use a laboratory analyzer that measures calcium (not CaCO3 ) and use a cut-off value >40 mg/kg (10 mmol/L) as the criterion compatible with a term pregnancy and mature fetus. Sodium (Na) concentration is typically greater than potassium (K) concentration until 3–5 days before delivery, at which time the Na/K ratio inverts
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Diagnostic Laboratory Resources and Critical Care Supplies and K concentration increases above 35 mEq/mL (35 mmol/mL) and Na concentration decreases below 30 mEq/mL (30 mmol/mL). This milk electrolyte profile is usually indicative of a mature or term fetus in the normal, uncomplicated pregnancy. Precise measurements of these electrolytes are best obtained using a laboratory chemistry analyzer. In summary, as parturition approaches, calcium concentrations increase, accompanied by an inversion of sodium and potassium concentrations. Mature foals are usually produced from deliveries associated with the following milk electrolyte profile: calcium ≥ 40 mg/dL, sodium ≤ 30 mEq/L, and potassium ≥ 35 mEq/L.1,2 Chapter 286 discusses the evaluation, interpretation, and clinical application of milk electrolyte profiles in greater detail. Neonatal isoerythrolysis screens When a mare is bred to a stallion that carries red cell alloantigens that are not present on the mare’s red blood cells, the mare has the potential to produce antibodies against these antigens. During the last 2 weeks of pregnancy, these antibodies will be produced in high concentrations in the serum and there will be correspondingly high concentrations in the colostrum as well. The neonatal isoerythrolysis (NI) screen detects any antibodies in the dam’s colostrum or blood that are known to cause a hemolytic reaction in the foal. If the colostrum and/or serum are positive for any of these antibodies, the foal should be restricted from nursing colostrum from its dam immediately postpartum to avoid a potentially fatal hemolytic reaction caused by the antibodies present in the colostrum. An alternate source of colostrum (tested negative for offending antibodies) should be administered to the foal. The mare is then handmilked for an average of 24–36 hours following delivery or until samples of her milk are no longer positive for the offending antibodies. An in-depth discussion of NI and NI screening procedures is presented in Chapter 35.
Neonatal isoerythrolysis screens for the dam A blood sample must be collected from the pregnant mare within 2 weeks of parturition to ensure the presence of a high enough level of antibody for detection in the serum. A panel of cells with red cell antigens that are most commonly associated with NI is washed with saline to eliminate any proteins in the serum or plasma that may cause non-specific agglutination. The washed cells are mixed with serial saline dilutions of the dam’s serum in a 96-well plate. Rabbit complement is added to enhance the hemolytic reac-
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tion. The plate is mixed on a plate shaker and the cells are allowed to settle out for a few hours. The plate is then read, looking for hemolysis. If no hemolysis is observed in any of the wells, the patient is considered negative for hemolytic antibodies. If hemolysis is observed, the pattern of hemolysis will determine which antibody is present. If the mare does not foal within 2 weeks of when the NI screen was performed, it is suggested that another blood sample be collected and the test repeated. This follow-up screen helps prevent false-negative results that may occur if the mare was not producing antibody at a high enough level to be detected during earlier NI screens.
Neonatal isoerythrolysis screens for colostrum NI screens can also be performed on colostrum to determine if any antibodies are present that may cause a hemolytic reaction. This is useful if the mare foals early and an NI screen was not performed on the dam’s blood prior to parturition. This test can be run as a prophylactic measure to ensure that no hemolytic reaction will occur. The test can also be used as an early diagnostic test to help confirm the likelihood of hemolysis in a foal that is suspected of having NI; a positive NI screen on the colostrum supports a tentative diagnosis of NI in the foal. This test is also useful to screen samples of good-quality colostrum (IgG concentration > 4000 mg/dL) that a farm intends to collect and add to its colostrum bank to be used for future foals that are born to mares with either positive NI screens or poor-quality colostrum. Jaundice foal agglutination test A jaudice foal agglutination (JFA) test uses the mare’s colostrum or milk and the foal’s red cells to determine if a hemolytic reaction may occur. The test will be ordered if a mare had a positive NI screen or if a NI screen was not performed on the mare prior to delivery. To perform the test, a milk sample is obtained from the mare and a pre-suckle blood sample is obtained from the foal. Using saline, serial dilutions are made of the mare colostrum. Varying dilutions of the mare’s colostrum are mixed with her foal’s washed red cells and rabbit complement in a test tube and incubated at 37◦ C. After incubation, the test tube is centrifuged and then shaken lightly while watching the cell button in the bottom of the tube and checking for macroscopic agglutination. After checking for macroscopic agglutination, microscopic agglutination is evaluated by placing a drop on a slide and covering the specimen with a coverslip and observing for agglutination under the microscope
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Foal: Appendix Assessment of colostral immunoglobulin G Evaluation of colostral IgG concentration immediately post-foaling can help predict if FPT is likely to occur, since ingestion of poor-quality colostrum is one of the risk factors for hypogammaglobulinemia in foals. Colostral IgG concentrations above 4000 mg/dL are desirable to ensure the foal’s serum IgG concentration will reach acceptable post-suckle levels (i.e., > 800 mg/dL).
r Figure 91.1 Jaundiced foal agglutination test. Slide showing several clumps of agglutinating red blood cells. Positive microagglutination indicates the dam’s colostrum contains enough anti-red cell antibodies to produce a hemolytic reaction in the foal. Reprinted with permission from McAuliffe SB and Slovis NM, Color Atlas of Diseases and Disorders of the Foal, copyright Elsevier, 2008.
(Figure 91.1). Agglutination in any of the dilutions represents a positive result at that dilution. The farm is advised to milk the mare out for a minimum of 24 hours, at which time another sample of milk is retested to ensure the antibody level is low enough not cause a hemolytic reaction in the foal. In author’s (NS) clinical experience, if after 24 hours the colostral antibody level is still greater than 1 : 16 there have been some foals that develop hemolytic reactions resulting in neonatal isoerythrolysis if they are allowed to return to the mare prematurely. It is not unusual for a hemolytic crisis to occur in foals > 24 hours when they ingest colostrum with elevated antibody levels above 1 : 16. Therefore we recommend not allowing the foal to nurse until the colostral antibody level is < 1 : 16 or the foal is at least 48 hours of age.
r
r
Tests to measure immunoglobulin G Due to the lack of in utero transplacental transfer of antibodies from dam to fetus, the foal is usually born without detectable levels of IgG. The neonatal foal’s immunity to infectious agents relies on the ingestion and absorption of maternal IgG from colostrum. Healthy newborn foals that have consumed adequate amounts of good-quality colostrum typically have serum IgG concentrations above 800 mg/dL by 12–24 hours of age. Foals with IgG levels less than 400 mg/dL are considered to have failure of passive transfer (FPT) and are at increased risk for infection during the first few weeks postpartum. Foals identified with FPT are candidates for IgG supplementation using colostrum, other oral IgG products, or a plasma transfusion.
r
Colostrum can be subjectively assessed on the basis of appearance. A thick, yellow, sticky secretion is most likely of good quality, whereas a dilute, white or translucent secretion is most likely inadequate. However, physical characteristics alone are often rather subjective. Quantitative measurement of IgG levels in colostrum can be obtained using a serial radial immunodiffusion (SRID) assay. Unfortunately, the SRID technique cannot be performed easily on a farm and requires approximately 24 hours to obtain results. It is most suitable for testing colostrum that is to be banked, or to confirm questionable IgG results obtained using other assays. Qualitative assessment of colostral IgG concentrations can be performed more rapidly using R commercially available kits (Gamma-Check C , Plasvacc, Templeton, CA) that are based on a precipitation reaction between glutaraldehyde and immunoglobulins. The time required for precipitation (clot formation) to occur is directly proportional to the amount of IgG in the sample. Such kits are easy to use in the clinic or on the farm. To perform the test, a sample of mare’s colostrum is mixed with a glutaraldehyde reagent and, if sufficient IgG is present, a clot is formed within the recommended time period (< 5 minutes). A positive test result indicates colostral IgG concentration is greater than 38 g/L. Semi-quantitative assessment of colostrum can also be performed using one of several techniques to measure specific gravity (SG). Specific gravity can be estimated using a modified hydrometer (Equine Colostrometer, Jorgensen Laboratories, Loveland, CO). A measured volume of filtered colostrum is placed into a chamber that is then suspended in a column of distilled water. Goodquality colostrum should have an SG >1.065. To improve the accuracy of this method it is important to add precisely the correct volume (15 mL) of colostrum to the chamber and to ensure that the distilled water is at the appropriate temperature. Alternatively, a hand-held refractometer, designed
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to measure sugar or alcohol content, is also suitable for assessing colostrum quality. There is a direct correlation between the alcohol refractometer reading and IgG content. Values ≥ 16% on the alcohol scale or ≥ 23% on the BRIX sugar scale correspond to colostral IgG concentration ≥ 60 g/L. The refractometer is easy to use and simply involves placing a drop of colostrum on the refractometer and obtaining a refractive index reading. Chapters 33 and 34 discuss these procedures in greater detail.
Assessment of foal serum imunoglobulin G Ideally newborn foals should have a serum IgG concentration greater than 400–800 mg/dL within the first 12–24 hours following delivery. A variety of easy-to-use, rapid tests are available to measure IgG concentration in the foal’s serum. The details and accuracy of different assay technologies used to measure IgG as well as the recommended treatment strategies for foals suffering from FPT are discussed in Chapter 34. Zinc sulfate turbidity test The basis of the zinc sulfate turbidity test is the formation of a precipitate when immunoglobulins in the patient’s serum react with zinc ions in the reagent. The degree of turbidity produced by the reaction is related to the IgG concentration in the sample. The degree of precipitation can be subjectively evaluated by direct visualization, or the sample can be read spectrophotometrically against a standard curve. This test is inexpensive and easy to perform. It is best used as a screening test since accuracy decreases when serum samples contain lower levels of IgG (< 400 mg/dL). Presence of hemolysis in the foal’s serum sample will produce falsely elevated results. Serum should be used instead of plasma since elevated plasma fibrinogen concentrations may result in inaccurate results. Since it take time for serum to be obtained, as opposed to whole blood or plasma, this test may be unsuitable if rapid testing is required. R , VMRD, A commercial test is available (EquiZ Pullman, WA) or the reagents for the zinc sulfate test can be prepared in the clinic. The recipe for preparing the zinc sulfate solution is as follows: 1. 250 mg of hydrated zinc sulfate added to 1 L of freshly boiled water. 2. 6 mL aliquots of the warm solution are placed into sealed, 7–10 mL plain vacuum collection tubes. These tubes remain useful for several months. When used, the solution in the tube should be clear. Any cloudiness before the test commences indicates that carbon dioxide has been absorbed
Figure 91.2 Zinc sulfate turbidity test. The zinc sulfate solution is clear prior to the addition of serum (left) and becomes cloudy after the addition of serum (right), indicating the presence of immunoglobulin. Reprinted with permission from McAuliffe SB and Slovis NM, Colour Atlas of Diseases and Disorders of the Foal, copyright Elsevier, 2008.
into the solution, resulting in a pH change. The solution should be discarded. 3. Add 0.1 mL of the patient’s serum to the solution. Mix the tube gently. 4. Wait 10 minutes for a qualitative result. Turbidity indicates a positive result, meaning that adequate levels of IgG are present (Figure 91.2). 5. Wait 60 minutes for a quantitative result (measure against calibrated barium sulfate standards).
Latex agglutination test This test involves specific equine IgG antiserum which is coated onto latex reagent particles and mixed with the patient’s sample. Assessment of IgG concentration in the sample is based on the visualization of the degree of agglutination that occurs between latex particles and patient immunoglobulin. Positive agglutination occurs within 15 minutes and results can be quantified. Commercial kits are available. Whole blood, plasma or serum can be used depending on the kit. The test is simple and rapid and can be performed stable-side, although kits may be expensive. The test is not affected by hemolysis but can be adversely affected by temperature. There is a high accuracy at or below 4 g/L; however, above this level, accuracy may be poor. Precise measurement of all components used in the test is critical to ensure test accuracy. Concentration immunoassay technology test R test) (CITE A variety of commercial kits are available. They all require accurate pipetting, and the instructions
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Foal: Appendix 4. Clot formation in less than 10 minutes = [IgG] > 8 g/L. Clot formation in 60 minutes = [IgG] between 4–8 g/L.
Figure 91.3 Glutaraldehyde precipitation test. Note the firm clot that has formed in the bottom of the tube after the addition of serum to the glutaraldehyde reagent.Reprinted with permission from McAuliffe SB and Slovis NM, Colour Atlas of Diseases and Disorders of the Foal, copyright Elsevier, 2008.
provided with the kit should be followed exactly. Whole blood, serum or plasma can be used. The test uses a color spot with calibration standards that correlate to concentrations of 2, 4, and 8 g/L of IgG. The test is rapid (10–15 minutes) and results correlate well with SRID results. Glutaraldehyde coagulation test Chemical grade 25% glutaraldehyde is used in the test. This reagent forms an insoluble precipitate when mixed with basic proteins, including immunoglobulins (see Figure 91.3). R Commercial kits are available (Gamma-Check E, Plasvacc Inc, Templeton, CA) or the glutaraldehyde solution can be prepared in-house. It is inexpensive and reliable, but glutaraldehyde is a hazardous substance. Hemolysis in the patient sample will result in overestimation of IgG, producing a potentially false-positive result. This test should be periodically checked against the SRID test to maintain quality control. The test procedure is as follows: 1. Prepare a 10% solution of glutaraldehyde using deionized water. 2. Add 50 µL glutaraldehyde to 5 mL of serum and start the clock. 3. Measure the time for clot formation to develop.
R Foal IgG IDEXX SNAP R The SNAP Foal IgG Test (IDEXX Labs, Westbrook, ME) is an enzyme-linked immunoassay for semi-quantitative detection of IgG) in equine serum, plasma or whole blood. Diluted whole blood, serum or plasma and conjugate reagent are added to the R SNAP device. The device is then activated, releasing reagents stored within the device. Color development on the sample spot of the device is proportional to IgG concentration in the test sample. Calibrator spots on the device develop color corresponding to sample IgG levels of 400 mg/dL and 800 mg/dL. Comparison of the intensities of color development on the sample spot and the calibrator spots allows an assessment of IgG level in the test sample. Color development on the calibrator spots indicates that the assay reagents are active. Recent studies have found that the best results were noted with low (< 400 mg/dL) and high (> 800 mg/dL) concentrations of IgG, with an accuracy of 80% and 89%, respectively, when compared to single radial immunodiffusion.
DVM Rapid TestTM for equine IgG The DVM-StatTM analyzer (Immunosystems Inc, Spring Valley, WI) is a microprocessor-controlled, LED-derived filter photometer suitable for colorimetric testing in the laboratory or the field. A highintensity light-emitting diode (LED) transmits a narrow bandwidth light beam through the reagenttreated sample to a light-sensitive photovoltaic cell, assuring high test sensitivity and very long light source and battery life. Reagent chemistries have been selected that provide a color intensity related to the concentration of the substance under test (e.g., IgG). This test is a quantitative immunoassay with high specificity and sensitivity for equine IgG concentrations. It is quickly becoming an industry standard. Point-of-care (POC) analyzers
Lactate analyzer Lactate is the normal end product of anaerobic metabolism and is used as a biochemical marker of oxygen delivery to tissues. Elevated lactate levels in sick foals are associated with tissue hypoxia which in turn may be due to hypovolemia, hypotension, cardiac failure, sepsis/endotoxemia or anemia. Lactate levels are used to determine whether foals have had a hypoxic/ischemic event. Lactate values can also be used to monitor the neonate’s response to fluid
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Table 91.1 Critical care supplies for foals Category
General stall supplies
Bedding supplies
Foal bed options
r r r
Potential source
Twin mattress or equivalent with removable, vinyl protective cover R Tempurpedic mattress with protective cover Custom water bed
Support devices to encourage sternal recumbency
r
Dandy ‘V’ support pads
r
Pillows with removable protective covers
Dandy Products, Goshen, OH; www.dandyproductsinc.com
Protective pads and mats
r r r
Heating supplies
Fluid supplies
Synthetic sheepskin mattress pads Wrestling mats for floors Soft floor and wall covering
r r r
Infrared radiant brooder lamps with thermostat R Bair Hugger warming blanket Thermacare Convective warming blankets; R Circulating hot water pads, blankets and T/Pump Long-term polyurethane i.v. catheters
r
14 or 16 gauge 20 cm catheters with extension set attached; single or double lumen
Coiled primary and extension i.v. sets with swivel Gyro Hanger with supreme swivel (fluid hanging system for stalls)
SoftStallTM System; SoftWallTM System: www.softstall.com Kalglo Electronics Co., Bethlehem, PA DRE Veterinary, Louisvile, KY; www.dreveterinary.com Gaymar Industries, NY
Arrow International Inc, Reading, PA MILA International Inc, Florence, KY [email protected] International WIN Ltd, PA MILA International Inc, KY International WIN Ltd
Devices to control fluid flow rates
r r r r r
Enteral and parenteral nutrition supplies
Respiratory support supplies
r
Dial-A-Flow extension set Buretrol fluid regulator with 150 mL reservoir chamber Soluset with 150 mL reservoir chamber IV fluid/enteral feeding backpack and surcingle with pressure infusor Infusion pumps
r r
Small diameter (12, 14 Fr) flexible, self-capping, enteral feeding tubes 3-in-1 3 L TPN mixing bags 8.5% amino acid solution (Aminosyn), 10 or 20% lipid emulsion (Liposyn) 50% dextrose Intranasal oxygen cannulas and oxygen tubing
r
Oxygen humidifiers and flow meters
r r
Abbott Labs, Inc, North Chicago, IL Travenol Labs, Deerfield, IL Abbott Labs, IL MILA International Inc., KY IMED Corp, San Diego, CA Baxter Healthcare Corp, Marion, NC Mila International, Inc Abbott Labs, IL Hospira, Inc
Jorgensen Labs, Loveland, CO Allegiance Medical Supply, Shreveport, LA Allegiance Medical Supply, LA Puritan Bennett, Lenexa, KS
Oxygen source
r r r r r r
Portable oxygen cylinders Portable oxygen concentrators with flow meter; 1–10 L/min flow rates; purchase new or used Cuffed nasotracheal tubes, 50–55 cm long; selection of sizes: 7, 8, 9, 10, 11, 12 cm internal diameter Self-inflating resuscitation bag: Ambu SPUR adult disposable resuscitator McCullough Constant Delivery Foal Resuscitator, aspirator, and oxygen mask Ventilator with selection of modes including SIMV, CPAP
Home healthcare pharmacies Jorgensen Labs Home care pharmacies; various medical supply distributors Bivona Inc, Gary, IN Cook Veterinary Products Ambu Inc, Glen Burnie, MD McCullough Products Ltd, Auckland, NZ; available through SurgiVet, Inc, Waukesha, WI; www.surgivet.com
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Foal: Appendix
Table 91.1 (Continued) Category
General stall supplies
Supplies to diagnose/treat hypogammaglobulinemia
Tests to measure IgG in colostrum
r r r
Potential source
Jorgensen Labs Plasvacc, Inc, Templeton, CA
Equine Colostrometer R Gamma-Check C BRIX refractometer
Tests to measure IgG in foal serum
r r r r
R SNAP Foal IgG test R Gamma-Check E Midland Quick Test kit DVM Rapid TestTM and DVM StatTM spectrophotometer ARS Foal IgG Test kit and densimeter R FoalChek Equine IgG Latex agglutination assay IgG-containing products
r r r
Monitoring/diagnostic equipment
Point-of-care diagnostic assays
r r r r r r r r r r r
Miscellaneous supplies
Hi-Gamm Plasma; Plasvacc Plasma; Visit website: [email protected] to see list of licensed products R Seramune (oral IgG supplement) ECG machine Capnograph: end-tidal CO2 monitors Indirect blood pressure monitor Ultrasound machine, preferably sector scanner Radiograph unit
Lake Immunogenics, Ontario, NY Plasvacc Inc, Templeton, CA Sera Inc, Shawnee Mission, KS Datascope Corp, Paramus, NJ Datascope Corp
R i-Stat System: analyzes blood chemistry/electrolytes, blood gases IRMA Blood Analysis System Series: analyzes electrolytes, blood gases STAT PROFILEpHOx: analyzes blood gases Glucometers: Accu-chek Aviva; One Touch Ultra; Optimum Xceed R Lactate analyzers: Lactate Accutrend , Lactate Scout, Lactate Plus
Foley urinary catheters with balloon cuff and wire stylets Blood culture media Selection of splinting material/braces
r
IDEXX Laboratories, Westbrook, ME Plasvacc Inc Midland BioProducts Corp, Boone, IA Value Diagnostics, a Division of Immunosystems Inc, Spring Valley, WI Animal Reproduction Sytems, Chino, CA Centaur Inc, Overland, KS
Abbott Point of Care Inc, Princeton, NJ Diametrics Medical, St Paul, MN Nova Biomedical, Waltham, MA www.glucosemeters4u.com www.Lactate.com MILA International Inc Bectin Dickinson, Sparks, MD DynaSplint Systems, Inc, Severna Park, MD
R DynaSplint Systems Microhematocrit centrifuge and refractometer
CPAP, continuous positive airway pressure; ECG, electrocardiogram; i.v., intravenous; SIMV, synchronous intermittent mandatory ventilation; TPN, total parenteral nutrition.
therapy. Persistent hyperlactatemia and decreased removal of excess lactate may both be manifestations of mitochondrial derangement and energy failure, even in the presence of normoxia and good tissue perfusion, secondary to the initial hypoxic event. Lactate concentration can be measured using handheld lactate analyzers or an open-ended chemistry analyzer. The handheld analyzers are easy to operate and are very similar to using a glucometer. Lactate levels should be < 2.0 mmol/L. Measuring lactate during initial evaluation of the sick foal and sequentially over time may be particularly useful to veterinarians managing critically ill neonates, especially in light of the increasing validation of stall-side
analyzers. In one study, for each 1.0 mmol/L increase in blood lactate, the odds of survival at 96 hours postbirthing decreased significantly by 18% at admission, 39% at 24 hours, and by 53% at 48 hours.
Glucometer Glucose reserves in the newborn foal are low. Hypoglycemia can be observed with lack of nursing, increased catabolism as seen with sepsis or reduced gluconeogenesis associated with prematurity and sepsis. Hyperglycemia can result from overzealous glucose administration or insulin resistance
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Diagnostic Laboratory Resources and Critical Care Supplies associated with septicemia, hypoxic-ischemia syndrome or prematurity. Glucose levels can be measured using glucometers or an open-ended analyzer. With the use of stall-side glucometers the veterinarian can easily assess a sick foal’s glucose levels and treat accordingly. If the glucose concentration is below 70 mg/dL then dextrose supplementation is indicated, either by adding dextrose to intravenous fluids and/or by instituting enteral feeding if the foal is stable and has a functioning gastrointestinal tract. Blood cultures Blood cultures are performed if a foal is suspected of being septic. Blood cultures are collected aseptically, preferably by direct venipuncture, following sterile preparation of the skin overlying the vein. Gloves should be worn for the procedure. Cultures also can be obtained by withdrawing a sample of blood through a sterile catheter immediately after its insertion. Once collected the blood sample is placed into a broth culture media and incubated overnight. The culture bottle is examined grossly for visible growth and then subcultured onto culture plates. If the culture broth appears turbid a sample should be obtained aseptically and a Gram stain performed to identify bacteria. The culture plates and blood culture vial are then re-incubated overnight and the plates and bottle re-examined for evidence of bacterial growth. If there is growth on the plates, Gram stain identification and antibiotic sensitivity testing of the organism will be performed. The blood culture will be subcultured again onto more culture plates, and the plates and culture bottle will be re-incubated and re-examined for growth for the following two days. Often foals have received antibiotics prior to having a blood culture performed. Even one dose of antibiotics administered 24–48 hours earlier may make
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it difficult to recover pathogenic organisms from a blood culture. If a foal was administered antibiotics before blood cultures are collected, a blood culture vial with some type of antimicrobial/antibiotic inhibitor should be used to increase optimal recovery of pathogenic organisms.
Critical care supplies Equipment needs are dictated by the level of intensive care to be provided by the farm personnel or in the clinic. A detailed list of supplies, basic monitoring equipment and diagnostic POC analyzers is presented in Table 91.1. Potential sources for specialized equipment are also presented. This list is by no means exhaustive. Additional equipment and suggested supplies are described throughout this entire section on foal care. The reader is referred to Chapters 13, 14, 19, 21, 24, 27 and 92 for a review of fluid and drug supplies used frequently in the treatment of the young foal. The most critical component in any foal intensive care unit or foaling barn is the nursing staff. Without their patient care and diligent monitoring skills, foal morbidity and mortality rates will remain unacceptably high regardless of the inventory of critical care equipment available. Careful selection and training of nursing personnel is essential to the success of any foal unit.
References 1. Ousey JC, Dudan F, Rossdale PD. Preliminary studies of mammary secretions in the mare to assess foetal readiness for birth. Equine Vet J 1984;16:259–63. 2. Leblanc MM. Induction of parturition in the mare: Assessment of readiness for birth. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 34–9.
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Chapter 92
Formulary Robert P. Franklin
Table 92.1 Medical use formulary. Drug
Amount/route/frequency
Indication
Comments
Acepromazine
0.02–0.066 i.v., i.m. q.6 h or p.r.n.
Sedative, vasodilator, treatment of rhabdomyolysis
May cause paraphimosis in stallions
Acetazolamide
2.5 mg/kg p.o. q.12 h
Potassium wasting diuretic for prevention of HYPP attacks
Acetylcysteine
Enema: Administer via Foley catheter; retain for 30 min G.P. lavage: 20–60 mL of 20% into guttural pouches Neonatal encephalopathy: 30 mg/kg bolus followed by 20 mg/kg/h i.v. as 10% solution Ophthalmic: 0.2 mL topically as 5% q.1–4 h
Meconium impaction Mucolytic for treating empyema
ACTH: cosyntropin
0.1 mg/foal i.v.
High-dose ACTH stimulation test. Pull post-cortisol samples at 30/90 min
Sepsis, MODS, prematurity and a negative prognosis may be indicated by a poor response
Acyclovir
20 mg/kg p.o. q.8 h 10 mg/kg i.v. over 1 h q.12 h
Antiviral, prevention and treatment of EHV
Poor oral bioavailability, see Valaciclovir
Albuterol
1–2 µg/kg or 1–2 puffs of 90 µg/100 kg via AeromaskTM or EquinehalerTM . Use as needed but not to exceed 4× per week unless combined with corticosteroids. With fluticasone: administer prior to fluticasone
Short-acting (rescue) bronchodilator for treatment of respiratory distress associated with bronchoconstriction
β 2 -agonist receptors are downregulated very quickly unless combined with corticosteroids
Allopurinol
44 mg/kg p.o. within 4 h of birth
Prevention of hypoxic, ischemic damage in at-risk foals, by limiting free radical formation
Altrongenist
Mare 0.088 mg/kg p.o. q24 hrs
Used as tocolytic agent to treat placentitis; also used with domperidone and estradiol to create surrogate dams
Extra-label uses
Amikacin
Systemic Foals: 25 mg/kg i.v. or i.m. q.24 h Local Intra-articular: 250 mg per joint q.24 h IVRP: 250 mg to 1 g q.s. to 20–60 mL with saline q.24–48 h
Aminoglycoside antibiotic. Gram-negative or Staphylococcus infections Local: septic arthritis, septic physitis, osteomyelitis, joint lacerations
Monitor creatinine levels. For broad-spectrum activity add with penicillin. Trough ≤ 1–2 µg/mL IVRP: tourniquet proximal to area of infection. Leave tourniquet on for 30 min post injection
Aminocaproic acid
70 mg/kg i.v. over 20 min, then CRI of 15 mg/kg/h
Antifibrinolytic for horses at risk of severe hemorrhage
Enema preparation: 40 mL 20% acetylcysteine + 160 mL warm water + 20 g bicarbonate
Prevention of NE Antiproteinase for melting ulcers
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Robert P. Franklin.
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Formulary
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Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Amphotericin B
0.3 mg/kg day 1, 0.45 mg/kg day 2, 0.6 mg/kg day 3, then 0.6 mg/kg q.48 h for 28 days, i.v. in 1 L 5% dextrose over 1 h
Antifungal therapy
Monitor creatinine levels
Ampicillin sodium
Systemic: 22 mg/kg i.m. q.12 h or i.v. q.6–8 h
Beta-lactam antibiotic. Gram-positive bacterial infections and Escherichia coli
Combine with aminoglycoside for broad-spectrum activity
Arginine vasopressin (ADH)
0.25–0.5 mU/kg/min i.v. CRI
Treatment of refractory hypotension in cases of septic shock
Aspirin
12 mg/kg p.o. q.48 h
Antithrombotic, phlebitis, horses at risk of thromboemobolic disease
Atracurium besylate
0.2–0.28 mg/kg i.v.
Paralyzing agent
Atropine
Bradycardia: 0.02–0.10 mg/kg i.v., i.m., s.c. Ophthalmic: 0.2 mL topically q.2–48 h p.r.n.
Parasympatholytic Anterior uveitis, synechia
Aurothioglucose (gold salts)
50 mg test dose i.m., 1 mg/kg i.m. q.7 days, decrease to q.14–28 days
Pemphigus foliaceus
Azathioprine
1–3 mg/kg p.o. q.24 h
Immunosuppressant, in lieu of corticosteroids
Azithromycin
10 mg/kg p.o. q.24 h for 5 days then q.48 h
Azalide antibiotic for treatment of Rhodococcus equi or Salmonella in foals; usually combine with rifampin
Azosulfamide
2 mg/kg i.v.
Urine dye for detecting urinary leaks
Bacillus Calmette–Guerin ´ (BCG)
Intralesional injection of 0.3–0.5 mL/cm3
Sarcoids
Benztropine mesylate
500 kg: 8 mg i.v. given slowly
Parasympatholytic and antihistamine drug used to treat acute priapism or acute fluphenazine toxicity
Colic
Bethanachol
0.025 mg/kg s.c. q.6 h
Parasympathomimetic, prokinetic
May cause salivation
Bicarbonate
mEq=25: bicarb value × 0.6 (foal) or 0.3 (adult) × bodyweight in kg. 1 L of 5% has 595 mEq (500 mL bottle has 297.5 mEq)
Treatment of normovolemic metabolic acidosis or hyperkalemia
Biosponge : smectite clay
Foals: 30–60 mL p.o. q.12 h Adult: 1.5 kg initial dose via NGT followed by 500 g via NGT q.12 h
Smectite clay shown to bind Clostridium difficile and Clostridium perfringens toxins in vitro
Do not administer to foals prior to screening for colostrum absorption
Bismuth subsalicylate
Foal: 1 mL/kg p.o. q.6–12 h
Local anti-inflammatory and binding agent for diarrhea
May combine with Biosponge
Botulinum antitoxin
Foal: 200 mL i.v.
Treatment of botulism
Bretylium tosylate
3–10 mg/kg i.v. bolus
Chemical defibrillation for ventricular fibrillation
Low success, but worth a try if defibrillator is not available
Butorphanol
Foal: 0.05 mg/kg i.v. or i.m. CRI: bolus 0.02 mg/kg i.v., then 10–15 µg/kg/h
Analgesic/sedative, antitussive. CRI useful for visceral pain
May be combined with romifidine, xylazine or detomidine. CRI may also be combined with low-dose ketamine
Butylscopolamine bromide (Buscopan R Compositum )
0.3 mg/kg i.v. once
Spasmolytic, analgesic for treatment of colic
R
If used frequently or for prolonged time, monitor for ileus
May cause diarrhea or hyperthermia esp. in foals > 3 months of age Keep foals in cool environment
R
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Foal: Appendix
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Cabergoline
500 kg: 2–3 mg p.o. q.12–24 h
Dopamine receptor agonist for suppressing lactation
Caco–copper–iron
1 mL/100 kg i.v. diluted into 55 mL saline
Trace mineral supplement.
Caffeine
10 mg/kg loading dose p.o., then 2.5 mg/kg p.o. q.12 h
Respiratory stimulant for hypoventilation
Calcium gluconate 23%
Up to 1 mL/kg in fluids
Hypocalcemia, cardioprotectant during hyperkalemia
Carprofen
1 mg/kg p.o. q.12–24 h
NSAID, Cox-2 selective
Cefazolin
Systemic: 20 mg/kg i.v. q.6 h Ophthalmic: 0.2 mL 5% topically q.1–12 h
First-generation cephalosporin with activity against several Gram-negative organisms, Staphylococcus and most Gram-positive organisms
Ophthalmic: reconstitute 1 g vial with 5 mL sterile water and 15 mL artificial tears
Cefepime
Foals: 11 mg/kg i.v. or i.m. q.8 h
Fourth-generation cephalosporin. Broad-spectrum activity
Should be reserved for multidrug-resistant infections
Cefotaxime
Systemic Foals: 40 mg/kg i.v. q.6 h Local IVRP: 2 g in 30–60 mL saline IA: 500 mg
Third-generation cephalosporin. Broad-spectrum activity. Treatment of choice for bacterial meningitis
Should be reserved for multidrug-resistant infections. Pretreat meningitis cases with dexamethasone sodium phosphate
Cefpodoxime proxetil
Foals: 10 mg/kg p.o. q.6–12 h
Third-generation cephalosporin. Broad-spectrum activity
Oral bioavailability documented in foals
Cefquinome
1 mg/kg i.m., i.v., q 12h
Fourth-generation cephalosporin
Broad spectrum activity
Ceftazidime
Foals: 20 mg/kg i.v. or i.m. q.8 h
Third-generation cephalosporin. Broad-spectrum activity
Should be reserved for multidrug-resistant infections
Ceftiofur sodium
Systemic Foals: 5 mg/kg i.v., i.m., s.c. q.12 h Adults: 2.2 mg/kg i.v., i.m., s.c. q.12–24 h Excede (ceftiofur crystalline free acid) Adults: 6.6 mg/kg i.m. q.4 days (divide dose into ≤ 10 mL/site) Foals: 6.6–13.2 mg/kg s.c. q.3–5 days Local IA: 150 mg
Third-generation cephalosporin. Broad-spectrum activity
i.v. pharmacokinetics in foals recently established. Ceftiofur crystalline free acid: do not administer in the neck, when treating foals for Gram-negative bacteria use higher dose and frequency. Excede may cause injection site reactions and diarrhea.
Ceftriaxone
25–50 mg/kg i.v. or i.m. q.12 h
Third-generation cephalosporin. Broad-spectrum activity
Chloramphenicol
Systemic: 50 mg/kg p.o. q.6 h Ophthalmic: 0.2 mL q.1–24 h
Broad-spectrum antibiotic. Bacteriostatic
Cimetidine
6.6 mg/kg i.v. q.8 h 15 mg/kg p.o. q.8 h
H2 antagonist for gastric ulcers
Ciprofloxacin
Ophthalmic: 0.2 mL topically q.2–12 h
Fluoroquinolone for Gram-negative or Staphylococcus keratitis. Good corneal penetration
Useful in neonatal encephalopathy cases with marginally elevated P a co2
Human health hazard
May slow corneal epithelialization
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Formulary
841
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Cisapride
0.5 mg/kg p.o. q.8 h
Prokinetic for cecal motility disorders
Clarithromycin
7.5 mg/kg p.o. q.12 h
Macrolide antibiotic effective in treating Rhodococcus equi infections. Combine with rifampin
Most effective treatment for severe Rhodococcus equi infections. May cause diarrhea at same incidence as erythromycin especially in foals older than 3 months. May also cause hyperthermia in older foals. Keep foals in cool environment.
Clenbuterol
0.5–2 mL/50 kg bodyweight p.o. q.12 h
Bronchodilator and tocolytic
β 2 -agonist receptors are downregulated very quickly unless combined with corticosteroids
Colchicine
0.06 mg/kg p.o. q.24 h for 5 days/week
Inhibit liver cirrhosis/fibrosis
Dantrolene
10 mg/kg p.o., then 2.5 mg/kg p.o. q.2 h or 15–25 mg/kg i.v. slow q.6 h
Prevention/treatment of rhabdomyolysis
Denosyl (S-adenosylmethionine ademetionine)
20 mg/kg p.o. q.24 h
Antioxidant for liver dysfunction
Detomidine
0.005–0.02 mg/kg i.v. or i.m.
Sedation/analgesia
May add butorphanol to enhance analgesic properties
Dexamethasone
Systemic: 0.02–0.2 mg/kg p.o., i.v., i.m. q.24 Meningitis: 0.4 mg/kg i.v. q.12 h × 2 days Ophthalmic: Topically 0.1% q.4–24 h
Corticosteroid Non-septic anterior uveitis, conjunctivitis
Oral bioavailability is 50% of i.v./i.m.
Dextrose 50%
5% solution: 5–10 mL/kg/h 10% solution: 2.5–5 mL/kg/h
Hypoglycemia Hyperkalemia
100 mL/L LRS = 5% dextrose solution 200 mL/L LRS = 10% dextrose solution
Diazepam
Seizure control: 0.1 – 0.2 mg/kg i.v. Anesthesia induction:- 50 µg/kg i.v. with ketamine
Antiepileptic, induction of anesthesia
R
Diclofenac
R ): 0.2 mL topically Ophthalmic (Voltran q.4–6 h R ): apply locally q.12–24 h Topical (Surpass
Comments
Topical NSAID
Digoxin
0.002 mg/kg i.v. q.12 h 0.01 mg/kg p.o. q.12–24 h
Cardiac inotrope, negative chronotrope for congestive heart failure
Monitor serum levels to keep < 2.0 µg/mL
Dimethyl sulfoxide
Systemic 0.25–1.0 g/kg i.v., NGT as 10% solution q.12–24 h
ROS scavenger. Treatment of a number of equine maladies
Concentration is 999 mg/dL. Give as 10% solution. May cause hemolysis if not diluted
Diphenhydramine
1 mg/kg i.m. or slow i.v.
Adjunctive treatment for anaphylaxis. Reversal agent for fluphenazine reaction
May cause initial excitement phase
Dipyrone
22 mg/kg i.v. or i.m.
Analgesic, antipyretic, antispasmodic
Dobutamine
2–15 µg/kg/min
Cardiac inotrope for hypotension
Correct volume deficit first. May combine with norepinephrine
Domperidone
1.1 mg/kg p.o. q.12–24 h for 10 days prior to anticipated foaling date or if known to be grazing fescue pasture
Agalactia
Used with altrenogest and estradiol to create foster dams from non-lactating mares
P1: SFK/UKS c92
P2: SFK
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842
13:31
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Printer Name: Yet to Come
Foal: Appendix
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Dopamine
1–10 µg/kg/min
Afferent renal arteriole vasodilator and at higher doses increases cardiac output
2–3 µg/kg/min dose for ARF, although human meta-analysis has failed to show significant effect
Doxapram
0.02–0.05 mg/kg/min i.v. CRI 1 mg/kg i.v. q.5 min
Respiratory stimulant
CRI for hypoventilating patients. i.v. push for postpartum apnea; during secondary apnea respiratory center no longer responsive to doxapram
Doxycycline
10 mg/kg p.o. q.12 h
Broad-spectrum antibiotic
Enrofloxacin
5 mg/kg i.v. q.24 h 7.5 mg/kg p.o. q.24 h
Fluoroquinolone antibiotic. Gram-negative or Staphylococcus infections
Caution in foals as may cause cartilage damage. Irritating to veins when used in IVRP
Epinephrine
Low dose: 0.01–0.02 mg/kg High dose: 0.1–0.2 mg/kg 25 mg/kg p.o. q.6–8 h
Anaphylaxis, cardiac standstill
Use high dose if unresponsive to 3 rounds of low dose Monitor for colitis and hyperthermia. Keep foals in a cool environment
Erythromycin injectable
1 mg/kg i.v. q.6 h
Prokinetic
Erythromycin phosphate or stearate
37.5 mg/kg p.o. q.12 h
Rhodococcus equi pneumonia combined with rifampin
Monitor for colitis and hyperthermia. Keep foals in a cool environment
Ethylenediamine dihydroiodide, EDDI
20–40 mg/kg p.o. q.24 h
Fungal infections
Organic iodide
Etodolac
23 mg/kg p.o. q.24 h
NSAID for musculoskeletal pain
Famotidine
3 mg/kg p.o. q.8 h
H2 antagonist for treatment or prevention of gastric ulcers
Fentanyl (Duragesic R patches )
Foal: 100 µg/h patch per 50 kg
Analgesic for visceral pain, especially when NSAIDs are contraindicated. Postoperative pain control in colic, umbilical, bladder surgeries
Filigrastim
50-kg foal: 300 µg s.c.
Granulocyte colony-stimulating factor for severe neutropenia
Fluconazole
14 mg/kg loading, 5 mg/kg q.24 h
Antifungal
Fludrocortisone acetate
0.1 mg/70 kg p.o. q.24 h
Mineralocorticoid for treatment of adrenal insufficiency
Fluids, i.v. maintenance crystalloids
Juvenile: 7.5% bodyweight per day Neonate: 10% bodyweight per day Add ongoing losses and residual deficits to calculation
Maintenance of hydration in NPO patients. 20–40 mEq KCl is often added per liter in normokalemic patients
Maintenance fluids: Plasmalyte 56, 0.45% saline with 2.5% dextrose. Replacement fluids can substitute if potassium added
Fluids, i.v. replacement crystalloids
L = fluid deficit (BW × % dehydration) × 0.5 rapid i.v. infusion
Volume replacement. 25% of isotonic crystalloids remain in the intravascular space. Short-lived effects
Replacement fluids include: LRS, Normosol-R, 0.9% saline, Plasmalyte 148, Hartmann’s
Flunixin meglumine
1.1 mg/kg i.v., p.o. q.12 h or 1.5 mg/kg i.v. q.24 h ‘Antiendotoxic dose’: 0.25 mg/kg i.v. q.6 h
NSAID with a wide range of usages
Foals may have delayed clearance and SID dosing may be appropriate
Flurbiprofen
0.2 mL topically q.8–12 h
NSAID for anterior uveitis
Erythromycin estolate
Rhodococcus equi pneumonia combined with rifampin
Clip hair, do not scrub, secure by gluing edges to skin and wrapping with Elastikon. Sites: neck, forearm
P1: SFK/UKS c92
P2: SFK
Color: 1C
BLBK232-McKinnon
February 10, 2011
13:31
Trim: 279mm X 216mm
Printer Name: Yet to Come
Formulary
843
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Fluticasone
4.4 µg/kg or 2 puffs of 220 µg/100 kg via AeromaskTM or EquinehalerTM . Begin at q.12 h × 10 days, then q.24 h × 10 days, then q.48 h × 10 days
Inhaled corticosteroid for interstitial pneumonia
BAL results should indicate increased levels of non-degenerative neutrophils
Furosemide
0.5–1 mg/kg i.v. or i.m. q.6–12 h
Diuretic for pulmonary edema
Gabapentin
2.5 mg/kg p.o. q.12 h
Neuropathic pain
Taper dose at end of treatment
Gentamicin
Systemic Foal < 14 days: 12 mg/kg i.v. q.24 h Foal > 14 days; Adult: 4.4–6.6 mg/kg i.v., i.m. q.24 h Local IA: 250 mg per joint q.24 h IVRP: 500 mg q.s. to 60 mL with saline q.24 h Ophthalmic: 0.2 mL q.2–12 h
Aminoglycoside. Gram-negative and Staphylococcus bacterial infections Foals: septic arthritis, physitis, osteomyelitis
Monitor creatinine levels. Target trough ≤ 1 µg/mL
Glycopyrrolate
0.005–0.01 mg/kg i.v., i.m., s.c.
Anticholinergic
Griseofulvin
5–10 mg/kg p.o. q.24 h × 7–28 days
Dermatophytosis
Combine with miconazole shampoo and lime sulfur dip
Guaifenesin
Anesthesia: 5% solution For induction: To effect i.v. Maintenance: ∼ 1 drop/s for adult horse as triple drip
Muscle relaxant, expectorant. Induction and maintenance of anesthesia
Triple drip: 500 mg xylazine, 1000–2000 mg ketamine in 1 L guaifenesin 5%
Heparin
25 iu/kg s.c. q.6, or 100 iu/kg/day i.v. as CRI
Prevention of adhesions and thromboembolic disease
Taper dose if PCV decreasing or clinical bleeding occurs
Heparin: low molecular weight R (Enoxaparin )
1 mg/kg s.c. q.24 h
Prevention of thromboembolic disease in septic patients
Low risk of clinical bleeding. Costly
Heparinized saline
1000 iu heparin per liter saline
i.v. catheter flush
Hetastarch
10 mL/kg i.v. q.24 h
Synthetic colloid
Hyaluronate sodium
1 vial IA
Treatment for synovitis osteoarthritis. Postoperative arthroscopy
Hydrocortisone R (Solu Cortef )
1 mg/kg i.v. q.6 h
Physiological dose: may assist in prevention of sepsis-induced hypotension or ARDS
Hydroxyzine
1 mg/kg p.o. q.12–24 h
Antihistamine for pruritus/urticaria
Hypertonic saline
2–4 mL/kg i.v. bolus
Low-volume plasma expander
Ibuprofen
25 mg/kg p.o. q.8–12 h
NSAID
Imidocarb diproprionate
2 mg/kg i.m. q.24 h for 2 days (Babesia caballi) 4 mg/kg i.m. q.3 days for 4 treatments (Babesia equi)
Babesiosis
Unlikely to cause remission in Babesia equi cases. Monitor liver/kidney function
Imipenem/cilastatin
Foal: 5–10 mg/kg i.v. over 20 min. q.6–8 h
Carbapenem antibiotic. Broad-spectrum. Very expensive
Reserve for multidrug-resistant bacterial infections
Insulin regular
0.1–0.5 iu/kg s.c. 0.00125–0.05 iu/kg/h i.v. as CRI
Hyperglycemia
Monitor glucose and potassium levels closely
Iohexol
0.1 mL/kg over 3 min intrathecal
Myelography
Pretreat horses with phenylbutazone to reduce risk of chemical meningitis
Maintains oncotic pressure for at least 24 h. Clotting abnormalities at dosages > 15 mL/kg
Monitor sodium as may result in hypernatremia. May combine with fludrocortisone
Follow with isotonic fluids. Use is controversial in neonatal foals
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Foal: Appendix
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Itraconazole
5 mg/kg p.o. q.24 h
Systemic antifungal
Itraconazole 1%/DMSO 30%
0.2 mL topically q.4–12 h
Fungal keratitis, stromal abscess
Ketamine
Analgesia 0.25 mg/kg i.m. q.6 h 2–10 µg/kg/min i.v. CRI Anesthesia Induction: 2 mg/kg i.v. Maintenance: 5–20 µg/kg/min
NMDA receptor antagonist. Prevention of spinal ‘wind up’ phenomenon. Maintenance dose used to lower inhalant anesthesia
Preserves activity of morphine/detomidine epidural, minimizes NSAID use. May be used to block NMDA in encephalopathic foals
Ketoconazole
10 mg/kg p.o. q.24 h
Antifungal
Very expensive
Ketoprofen
2.2 mg/kg i.v. q.24 h
NSAID
Lactase supplement R (Lactaid )
1–2 tablets p.o. q.2 h
Lactose-intolerant foals or foals recovering from diffuse enteritis where villous tips are damaged and unable to produce lactase
Lactobacillus rhamnosus GG R (Culturelle )
Foals: 1 tablet/50 kg p.o. q.12 h
Well-studied probiotic. For rotaviral and other enteritis/colitis cases
Levamisole
2 mg/kg p.o. q.48 h or 1–2 mg/kg p.o. q.24 h for 3 days per week
Immunomodulator
Lidocaine
Analgesic/prokinetic 1.3 mg/kg i.v. bolus over 5 min, then 0.05 mg/kg/min i.v. CRI Epidural Mares: 50 mg combine with 75 mg xylazine q.s. to 6 mL with sterile saline
Analgesic, prevention of neutrophil margination/activation. Questionable prokinetic
Loperamide R (Immodium AD )
0.1–0.2 mg/kg p.o. q.6 h
Antidiarrheal
Lime sulfur R (LymDyp )
Apply dip to entire body after shampooing with miconazole shampoo daily for 1 week
Ringworm
May also combine with griseofulvin
Magnesium hydroxide
0.5 mL/kg p.o. q.4 h
Antacid
Short-lived effects
Magnesium sulfate
Hypomagnesaemia: 5–15 mg/kg i.v. CRI in i.v. fluids Neonates: 50 mg/kg/ h i.v. over 1 h immediately post dystocia, then 10 mg/kg/ h CRI over 24 h Tetanus: i.v. CRI to diminish all neurological activity yet preserving the patellar reflex
NMDA receptor antagonist. May prevent neural injury from hypoxia/ischemia. Treatment of tetanus may require ventilator support
Treatment for neonatal encephalopathy is purely theoretical use
Magnesium sulfate (Epsom salts)
1 g/kg via NGT with 4–8 L water q.12–24
Increases intraluminal water content for impaction colics
Works synergistically with i.v. fluid therapy
Mannitol
0.25–1.0 g/kg i.v. as a 10% solution, give over 0.5 h
Osmotic diuretic for treatment of cerebral edema. ROS scavenger
Past concerns on impact of drug on cerebral hemorrhage are likely unwarranted
Methionine
500 kg: 30–60 g p.o. q.24 h
Urinary acidifier
Methocarbamol
5–25 mg/kg i.v. slow q.12 h, 50 mg/kg p.o. q.12 h
Muscle relaxant
Methylprednisolone sodium succinate (Solu Medrol)
30 mg/kg i.v. bolus then 5 mg/kg/h i.v. CRI for 23 h
Antioxidant/anti-inflammatory for spinal/head trauma
Recent data in dogs and humans suggest it may not affect outcome
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BLBK232-McKinnon
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Formulary
845
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Metoclopramide
0.1–0.2 mg/kg i.v. or i.m. q.6–12 h 0.04 mg/kg/h i.v. CRI
Prokinetic for gastric hypomotility
Often used in treatment of GDUD
Metronidazole
10 mg/kg i.v. q.12 h 10–25 mg/kg p.o. q.12 h 20–50 mg/kg per rectum q.12 h
Anaerobic infections (esp. Bacteroides fragilis, Clostridium difficile, perfringens) antidiarrheal
i.v. route is useful in foals with necrotizing enteritis and associated ileus
Miconazole 1%
0.2 mL topically q.4–12 h
Fungal keratitis
Miconazole shampoo R (Malaseb )
Shampoo daily and follow with lime sulfur dip
Dermatophytosis
Midazolam
0.1–0.2 mg/kg i.v. bolus 0.1–0.4 mg/kg/h i.v. CRI
Intractable neonatal seizure activity
Milk replacer
Start at 10% of bodyweight per day. Increase by 2% per day to a maximum of 25%
Orphaned or rejected foals
Mineral oil Misoprostol
7.5 mL/kg via NGT p.r.n. 0.004 mg/kg p.o. q.12–24 h
Laxative Prostaglandin E for treatment of right dorsal colitis
Morphine
0.1 mg/kg i.v. or i.m. q.4–12 h 0.16 mg/kg loading dose then 0.1 mg/kg/ h i.v. CRI
Narcotic analgesic
Morphine/detomidine epidural
Morphine 0.1–0.2 mg/kg Detomidine 0.03 mg/kg q.s. to a volume of 20 mL with sterile saline q.6–24 h
Analgesic for use with epidural catheter. Combine with ketamine to preserve effects. May see initial sedation due to lipid solubility of detomidine
Use at a frequency to control pain. Use caution with morphine containing preservative
Naloxone
NE or PPH: 0.01–0.02 mg/kg i.v. once Prokinetic: 0.05 mg/kg i.v.
Opioid antagonist. NE, PPH, prokinetic
Theoretical use
Naproxen
10 mg/kg p.o. q.12–24 h
NSAID
Natamycin
0.2 mL topically q.4–12 h
Fungal keratitis
Neostigmine
0.005–0.02 mg/kg s.c. q.1 h
Prokinesis for SI and LC
Nitroglycerin
Apply topically wearing gloves q.24 h
Treatment of thrombophlebitis
Norepinephrine
0.1–1.5 µg/kg/min in 5% dextrose i.v. CRI
Treatment of vasoresponsive hypotension
Ofloxacin
0.2 mL topically q.2–12 h
Fluoroquinolone for Gram-negative or Staphylococcus keratitis. Good corneal penetration
Omeprazole
Therapy: 4 mg/kg p.o. q.24 h, 0.5 mg/kg i.v. q.24 h Prophylaxis: 1 mg/kg p.o. q.24
Proton pump inhibitor for gastric ulcers
Oxacillin
25 mg/kg i.v. q.6 h
Penicillinase-stable penicillin. Indicated for treating many non-MRSA Staphylococcus infections
MRSA resistance is a growing concern. Reserve for resistant infections
Oxyglobin
30 mL/kg i.v., 3–5 mL/kg i.v. if combined with hetastarch
Hemoglobin replacement fluid for acute blood loss
Also has strong colloidal properties
May also combine with griseofulvin
Buckeye Mare’s Milk PlusTM , Progressive Foal’s FirstTM , Land O’Lakes Mare’s MatchTM are recommended Will cause abortion
May dilute to 3.3% with sterile water for SPL use
Should be combined with positive inotrope (dobutamine) to maintain splanchnic perfusion
P1: SFK/UKS c92
P2: SFK
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BLBK232-McKinnon
February 10, 2011
846
13:31
Trim: 279mm X 216mm
Printer Name: Yet to Come
Foal: Appendix
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Oxytetracycline
Contracted carpi/fetlocks 44 mg/kg i.v. q.24 h in 1 L physiological saline Antimicrobial: 7.5 mg/kg i.v. q.12–24 h
Tendon relaxation. Broad-spectrum antibiotic. Anti-MMP-9 activity
Monitor creatinine in foals with FLD, especially after multiple doses
Oxytocin
Choke: 0.11–0.22 iu/kg i.v. Reproduction: 10–60 iu i.m. or 30 iu i.v. q.30 min to 24 h Induction of labor: 2–10 iu slow i.v., parturition in 30–90 min; or 40–100 iu i.v., i.m. RFM: 100 iu in 1 L saline over 15–45 mins to effect
Treatment of choke, uterine fluid expulsion, retained fetal membranes. Low dose is preferred for induction of parturition
Pantoprazole
1.5 mg/kg i.v. q.24 h
Proton pump inhibitor for foals not tolerating oral medications
Partial parenteral nutrition
2–4 mL/kg/h
Useful for short-term parenteral nutrition (<24 h)
Penicillin, potassium
20 000–50 000 iu/kg i.v. q.6 h
Beta-lactam antibiotic. Gram-positive, not Staphylococcus, and anaerobic bacteria
Penicillin, procaine
22 000 IU/kg i.m. q.12 h
Beta-lactam antibiotic. Gram-positive, not Staphylococcus, and anaerobic bacteria
Combine with aminoglycoside to cover Gram-negative and Staphylococcus infections
Pentafusion pain management
0.15 mL/kg/h i.v. CRI and reduce as pain diminishes Loading doses at the beginning of treatment: Lidocaine: 1.3 mg/kg i.v. slow Morphine: 0.1 mg/kg i.v.
Combination of analgesics and sedatives for the control of severe pain
Add the following to a 1-L empty i.v. bag: 1000 mL 2% lidocaine 4000 mg ketamine 170 mg morphine 30 mg detomidine 15 mg acepromazine
Pentobarbital sodium
Seizures: 2–10 mg/kg i.v. Euthanasia: 0.1–0.2 mL/kg i.v.
Antiepileptic, euthanasia
Cardiorespiratory depressant
Pentoxyfilline
8.5 mg/kg p.o. q.8 h or 10 mg/kg p.o. q.12 h
Anti-TNF-α and MMP-9 for endotoxemia, vasodilator for placentitis. Also allows RBC membrane deformability but this effect takes days to occur
Also used with antibiotics and altrenogest to treat placentitis
Pharyngeal flush
10 mL via nasal applicator q.12–24 h
Treatment of lymphoid hyperplasia, soft palate ulcerations and other inflammatory conditions of the pharynx
Ingredients: 200 mL glycerin, 100 mL dexamethasone 0.2%, 100 mL DMSO, 150 mL furacin solution
Phenazopyridine
4 mg/kg p.o. q.6–8 h
Provides urinary tract anesthesia in foals with stranguria
Will cause urine discoloration
Phenobarbital
5–10 mg/kg p.o. q.12 h 5 mg/kg i.v. as loading dose (can use up to 20 mg/kg)
Barbiturate for seizure control
Therapeutic drug monitoring is encouraged after 4 days: therapeutic levels = 15–45 µg/mL
Phenylpropanolamine
1–2 mg/kg p.o. q.12 h
Increase urethral tone in incontinent horses
Phenytoin extended release
2.8–16.5 mg/kg p.o. q.8 h
Anticonvulsant in foals
1 L amino acids 8.5% or 10% with or without electrolyte 1 L 50% dextrose to 5 L LRS Combine with aminoglycoside to cover Gram-negative and Staphylococcus infections
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BLBK232-McKinnon
February 10, 2011
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Trim: 279mm X 216mm
Printer Name: Yet to Come
Formulary
847
Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Plasma, hyperimmune
FPT: 20–40 mL/kg Endotoxemia: 2–4 mL/kg Hypoproteinemia: 10 mL/kg
Treatment of failure of passive transfer, endotoxemia, hypoproteinemia
Caution when giving more than 40 mL/kg as volume overload may occur leading to pulmonary edema
Plasma, Rhodococcus
1 L i.v. on day 1, 2, or 3 and again on day 21
Prevention of/reduction of risk of infection
May reduce incidence by 33%
Polymixin B
6000 IU/kg i.v. q.12 h in 1 L LRS over 30 min
Subtherapeutic dose that binds endotoxin
Continue as long as insult persists
Ponazuril
5 mg/kg p.o. q.24 h for minimum of 28 days
EPM therapy
Potassium bromide
25–35 mg/kg p.o. q.24 h
Anticonvulsant
Potassium chloride
Spike fluids with 20–40 mEq/L, not to exceed 0.5 mEq/kg/h
Fluid supplement for anorexic horses or treatment of hypokalemia
Potassium iodide
2 mg/kg p.o. q.24 h
Systemic antifungal treatment
Povidone iodine
Ophthalmic: 0.2 mL 2% topically q.24 h
Fungal keratitis
Prednisolone
Anti-inflammatory/immunosuppressive 0.5–4 mg/kg p.o. q.24 to 48 h Ophthalmic (Prednisolone acetate 1%) 0.2 mL topically q.4–12 h
Prednisolone sodium succinate (Solu Delta R ) Cortef
1–5 mg/kg i.v. q.12–24 h
Preferred treatment for anaphylaxis
Procainamide
1 mg/kg/min i.v. to a total of 20 mg/kg
Ventricular tachycardia
Propantheline bromide
500 kg: 30 mg i.v.
Antispasmodic
Propofol
Induction: 2 mg/kg i.v. to effect Maintenance: 0.3 mg/kg/min
i.v. anesthetic for foals
Premedicate with xylazine with or without butorphanol
Psyllium powder
Therapy: 1 g/kg via NGT q.12–24 h.
Sand impaction
Mix with mineral oil when giving via NGT to prevent tube clot
Pyrilamine maleate
1 mg/kg i.m., s.c. or slow i.v.
Antihistamine
Pyrimethamine/ sulfadiazine
PYR: 1 mg/kg p.o. q.24 h Sulfadiazine: 30 mg/kg p.o. q.24 h
EPM therapy. Maintain on therapy as long as clinical signs are improving and there are no signs of bone marrow suppression
Quinapril
500 kg: 120 mg p.o. q.24 h
ACE inhibitor for managing CHF
Quinidine sulfate
Acute atrial fibrillation: 1.1–2.2 mg/kg i.v. q.10 min until conversion or total dose of 12 mg/kg reached Chronic: 22 mg/kg via NGT q.2 h until conversion, max. of 5 doses, QRS prolongs, or other complications occur. Next try q.6 h with digoxin at 11 µg/kg via NGT
Atrial fibrillation
Therapeutic range is 1–2 mg/mL with phenobarbital and 3 mg/mL as monotherapy
Monitor for signs of iodism such as lacrimation
Prednisone is poorly absorbed from GIT
20 mg/kg dose into 250 mL saline over 30 min or to effect
May cause non-regenerative anemia that responds once medication is terminated. Treatment in pregnant mares may be associated with the birth of weak foals, including birth defects, bone marrow aplasia and renal agenesis
Side-effects may include urticaria, airway obstruction, supraventricular tachycardia, paraphimosis, colitis
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Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Ranitidine
1.4 mg/kg i.v. q.8 h 6.6 mg/kg p.o. q.8 h
H2 antagonist for treatment/ prevention of gastric ulcers
Reserpine
2.5 mg/450 kg animal i.m. q.30 days 2.5 mg/450 kg animal p.o. q.24 h
Long-acting tranquilizer. Orally useful for agalactia
Mild diarrhea that may last for 2 days. May cause neurological signs at higher doses
Rifampin
5–10 mg/kg p.o. q.12 h
Gram-positive bacteria, especially Rhodococcus equi. Good bone/abscess penetration
Do not use as a monotherapy as resistance develops quickly. For treatment of Rhodococcus equi always use in combination with macrolide or azalide
Rimantadine
5 mg/kg i.v. q.4 h 30 mg/kg p.o. q.24 h
Antiviral treatment aimed at preventing influenza
Romifidine
Sedation: 0.04–1 mg/kg i.v. Preanesthetic: 0.075 mg/kg i.v.
α2 -agonist for sedation or preanesthetic
Saccharomyces boulardii (SynerGI-EQ)
5 billion cells/50 kg p.o. q.12 h
Well-studied yeast. Treatment/prevention of antibiotic-induced diarrhea or Clostridium enteritis/colitis
Salmeterol
0.4 µg/kg or 2 puffs of 25 µg/100 kg via AeromaskTM or EquinehalerTM q.12–24 h
Long-acting bronchodilator for use in patients with severe bronchospasm
Serum (autologous) Silver sulfadiazine
0.2 mL topically q.1–4 h 0.2 mL topically q.4–12 h
Antiproteinase for melting ulcers Fungal keratitis
Sodium iodide
30 mg/kg i.v. q.24 h
Systemic antifungal therapy
Sodium polystyrene sulfonate
50 kg foal: 15 g p.o. or per rectum q.12 h
Decreases blood potassium levels
Stanozolol
0.5–1.0 mg/kg i.m. q.7–14 days for up to 4 doses
Anabolic steroid
Sucralfate
20 mg/kg p.o. q.6 h
Sucrose aluminum hydroxide compound that binds to GI ulcers
Sulfamethoxazole/ trimethoprim
30 mg/kg p.o. or i.v. q.12 h
Broad-spectrum antibiotic
Telithromycin
15 mg/kg p.o. q.24 h
Macrolide-resistant infections in foals (esp. Rhodococcus equi)
Tetanus antitoxin
10,000 iu i.v., i.m.
Treatment of tetanus
Thiamine (vitamin B1 )
1–5 mg/kg i.m., s.c., i.v., p.o. q.24 h, or 10 mg/kg in i.v. fluids q.24 h
Treatment of CNS disease
Ticarcillin with clavulanic acid R (Timentin )
Systemic: Foals: 50 mg/kg i.v. q.6 Local: IA: 400–800 mg
Generally effective against common Gram-positive and Gram-negative bacteria including Staphylococcus
Tissue plasminogen activator (TPA)
50–150 µg intracameral with 27-gauge needle at limbus
Treatment of anterior chamber fibrin. Accelerates fibrinolysis
Tobramycin
0.2 mL topically q.2–12 h
Gram-negative bacterial keratitis
Tolazoline
Prokinesis: 0.5 mg/kg i.v. Alpha 2 antagonist: 4 mg/kg i.v., give slowly at 1 mL/s to effect
Prokinesis, α2 -antagonist
Comments
Combine with butorphanol for analgesia
β 2 -agonist receptors are downregulated very quickly unless combined with corticosteroids Mix 1 mL of cream with 8 mL artificial tears for use with subpalpebral lavage system
Potentially useful in foals with uroperitoneum
Use caution in older foals: may cause diarrhea, hyperthermia
Avoid with recent hemorrhage < 48 h
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Table 92.1 (Continued). Drug
Amount/route/frequency
Indication
Comments
Total parenteral Nutrition
Foal: 100 kcal/kg/day Adult: 30 kcal/kg/day
i.v. nutrition
Add 1L of 50% dextrose, 1L of 8.5% amino acids, 250 mL 20% lipids to 2-L sterile bag to make 1 kcal/mL TPN. May also give at fractional rate to provide partial parenteral nutrition
Tramadol R (Ultram )
Neonates: 5 mg/kg p.o. q.6 h Weanlings: 10 mg/kg p.o. q.8 h
Synthetic opioid analgesic for chronic pain
Trifluridine solution
1% topically q.2–6 h
Viral keratoconjunctivitis or keratitis
Tripelennamine
1 mg/kg i.m. q.6–8 h
Antihistamine for treatment of urticaria, anaphylactoid reactions
Triple antibiotic solution
Ophthalmic: 0.2 mL topically q.2–12 h
Broad-spectrum bacterial keratitis
Valaciclovir
20 mg/kg p.o. q.8 h
Antiviral. Herpes prophylaxis or therapeutic
Vancomycin
7.5 mg/kg i.v. q.8 h in 1 L physiologic saline 4 mg/kg p.o. loading dose, then 2 mg/kg p.o. q.6 h
Gram-positive infections, esp. MRSA, resistant Rhodococcus equi. Oral for resistant Clostridium difficile
Use only with pandrug-resistant infections
Vitamin B complex
1 mL/50 kg i.m., i.v. or in fluids
Appetite stimulant and supplement
Dilute to 60 mL with sterile saline
Vitamin B12
1 mL/50 kg i.m., i.v. or in fluids
Appetite stimulant and supplement
Dilute to 60 mL with sterile saline
Vitamin C
10–100 mg/kg i.v. (in fluids) or p.o. q.12–24
ROS scavenger
Vitamin E
20 iu/kg s.c., p.o. q.24 h, first dose use Vital R E s.c
ROS scavenger for neonatal foals with NE
Vitamin E/selenium
1 mL /50 kg i.m. q.10 days
Treatment of white muscle disease
Vitamin K1
1 iu/kg s.c. q.24 h
Adjunct to hemostasis, antidote to warfarin toxicity
Voriconazole
Systemic: 3 mg/kg p.o. q.12 h or 4 mg/kg p.o. q.24 h Ophthalmic: 0.2 mL topically q.2–24 h
Antifungal for Aspergillus
Whole blood
L = bodyweight × 0.08 × (desired PCV – actual PCV/donor PCV), or 15 mL/kg. Maximum harvest is 1.6% of donor’s bodyweight
Xylazine
Sedation: 0.2–1 mg/kg i.v. Preanesthetic: 1 mg/kg i.v. Epidural: 75 mg combine with 50 mg lidocaine q.s. to 6 mL with sterile saline
Sedative, analgesic
Yohimbine
0.075 mg/kg i.v.
α 2 -antagonist for reversal of xyalzine and prokinetic
Vitamin E concentration is very low in this preparation
Combine with butorphanol 0.02 mg/kg for more analgesia
ACE, Angiotensin converting enzyme; ACTH, Adrenocorticotropic hormone; ADH, Antidiuretic hormone; ARDS, Acute respiratory distress syndrome; ARF, Acute renal failure; BAL, Bronchoalveolar lavage; CHF, Congestive Heart Failure; CNS, Central nervous system; Cox-2, Cyclooxygenase 2; CRI, Constant Rate Infusion; EHV, Equine Herpes Virus; EPM, Equine Protozoal myeloencephalitis; FLD, Flexural limb deformity; GDUD, Gastroduodenal ulcer disease; GITgastrointestinal tract; HYPP, hyperkalemic periodic paralysis; IM, intramuscular; IV, intravenous; IA, intraarticular; IVRP, intravenous regional limb perfusion; LC, large colon; LRS, lactated ringers solution; MODS, multiple organ dysfunction syndrome; MRSA, methicillin resistant Stayphylococcus aureus; NE, neonatal encephalopathy; NGT, nasogastric tube; NMDA, N-methyl-D-aspartic acid; NPO, nil per os; NSAID, nonsteroidal antiinflammatory drug; PRN, Pro re nata, “as the circumstance arises”; PPH, post partum hemorrhage; PYR, pyrimethamine; RBC, red blood cell; RFM, retained fetal membranes; ROS, reactive oxygen species; SI, small intestine; SID, sem’el in di’e “once per day”; SPL, subpalpebral lavage system; TNF, tumor necrosis factor; Tpn, total parenteral nutrition.
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Table 92.2 Conversion of electrolytes from molar equivalents (mEq/L) to a measurement of mass concentration (mg/dL). To make the conversion from molar equivalents multiply by the conversion factor to obtain mass concentration units. Mass concentration units are converted to molar equivalents by dividing by the conversion factor
Electrolyte
Molar equivalent
Conversion factor
Mass concentration
Calcium Chloride Bicarbonate Magnesium Potassium Sodium
mEq/L mEq/L mEq/L mEq/L mEq/L mEq/L
2.005 3.55 6.1 1.215 3.91 2.30
mg/dL mg/dL mg/dL mg/dL mg/dL mg/dL
Table 92.3 Conversion of conventional units to a measurement in Syst`eme International (SI) units. To make the conversion from conventional units, multiply by the conversion factor to obtain SI units. SI units are converted to conventional units by dividing by the conversion factor
Component Albumin Ammonia (NH3 ) Bicarbonate Bilirubin Calcium Calcium Carbon dioxide Chloride Coriticotropin (ACTH) Cortisol Creatinine Diazepam Digoxin Epinephrine Estradiol Estrone Fibrinogen Glucose Lactate Magnesium Magnesium Phenobarbital Potassium Progesterone Protein Quinidine Sodium Triglycerides
Conventional unit g/dL µg/dL
mEq/L mg/dL mg/dL mEq/L mEq/L mEq/L pg/mL µg/dL mg/dL µg/mL ng/mL pg/mL pg/mL ng/dL mg/dL mg/dL mg/dL mg/dL mEq/L mg/L mEq/L ng/mL g/dL µg/mL mEq/L mg/dL
Conversion factor
SI unit
10 0.587 1.0 17.1 0.25 0.50 1.0 1.0 0.22 27.59 88.4 3.512 1.281 5.46 3.671 37 0.0294 0.0555 0.111 0.411 0.50 4.31 1.0 3.18 10.0 3.08 1.0 0.0113
g/L µmol/L mmol//L µmol/L mmol/L mmol/L mmol/L mmol/L pmol/L nmol/L µmol/L µmol/L nmol/L pmol/L pmol/L pmol/L µmol/L mmol/L mmol/L mmol/L mmol/L µmol/L mmol/L nmol/L g/L µmol/L mmol/L mmol/L
Table 92.4 Measurement and conversion of salts Salts
Conversion
CaCO3 45 g (18 g Ca) KCl 1 g KCl 30 g KCl 40 g KHCO3 1 g MgO 30 g NaCl 1 g NaCl 6.33 g NaCl 33 g NaCl 43 g NaHCO3 1 g NaHCO3 18 g NaHCO3 23.4 g
12-mL syringe case 14 mEq 6-mL syringe case 12-mL syringe case 10 mEq 12-mL syringe case 17 mEq 1 teaspoon 6-mL syringe case 12-mL syringe case 12 mEq 1 tablespoon 12-mL syringe case
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Chapter 93
Hematology and Biochemical Reference Values Robert P. Franklin and Joan Palmero
Table 93.1 Normal complete blood count parameters1, 2
Age
Packed cell Red blood volume Hemoglobin cell (%) (g/dL) (×106 /µ L)
Mean cell volume (fL)
Mean cell hemoglobin Total concentration leukocytes Neutrophils Lymphocytes (×103 /µ L) (g/dL) (×103 /µ L) (×103 /µ L)
Monocytes (×103 /µ L)
Eosinophils (×103 /µ L)
Basophils (×103 /µ L)
< 12 hours
37–49
12.6–17.4
9.0–12.0
36–45
32–40
6.9–14.4
5.55–12.38
0.46–2.54
0.04–0.43
0
0–0.02
1 day
32–46
12.0–16.6
8.2–11.0
36–46
32–40
4.9–11.7
3.36–9.57
0.67–2.12
0.07–0.39
0–0.02
0–0.03
1 week
28–43
10.7–15.8
7.4–10.6
35–44
35–40
6.3–13.6
4.35–10.55
1.43–2.28
0.03–0.54
0–0.09
0–0.18
1 month
29–41
10.9–15.3
7.9–11.1
33–40
34–40
5.3–12.2
2.76–9.27
1.73–4.85
0.05–0.63
0–0.12
0–0.08
3 months
32–42
11.7–15.3
9.2–12.0
31–38
34–40
6.7–16.8
3.92–10.35
2.88–7.15
0.12–0.76
0–0.55
0–0.07
6 months
29–41
10.8–15.4
7.9–11.6
32–39
33–40
7.8–11.6
2.89–5.56
3.2–6.01
0.04–0.45
0–0.55
0–0.06
Table 93.2 Normal ranges for complete blood count continued
Age < 12 hours 1 day 1 week 1 month 3 months 6 months
Total plasma protein (g/dL)
Fibrinogen (mg/dL)
Haptoglobin (mg/dL)
Serum amyloid A (µg/mL)
Icterus index (units)
Platelets (×103 /µL)
5.1–7.6 5.2–8.0 5.2–7.5 5.1–7.1 5.5–7.1 5.9–7.1
100–350 100–400 150–450 200–700 100–800 200–550
8–120 0–136 0–143 0–214 23–187 0–125
7.8 58.9 – – – –
15–50 10–75 5–25 5–20 5–25 5–20
105–446 129–409 111–387 136–468 200–376 128–368
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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13–39 18–43 14–164 17–99 0–27 0–26
0.2–4.8 0.6–4.6 0.8–8.2 1.2–5.9 1.1–3.9 0.3–3.3
97–315 146–340 237–620 252–440 282–480 300–620
0–47 0–49 4–50 5–47 8–65 7–20
65–380 40–909 52–143 81–585 57–204 97–396
CK 108–190 121–233 121–192 130–216 88–179 110–210
Glucose 12–27 9–40 4–20 6–21 7–20 15–30
Blood urea nitrogen 1.7–4.2 1.2–4.3 1.0–1.7 1.1–1.8 0.7–2.2 1.2–2.1
0.9–2.8 1.3–4.5 0.8–3.0 0.5–1.7 0.4–2.0 0.3–1.3
Total Creatinine bilirubin 0.3–0.6 0.3–0.7 0.3–0.7 0.1–0.6 0.1–0.7 0.1–0.7
Conjugated bilirubin 0.8–2.5 1.0–3.8 0.5–2.3 0.4–1.2 0.4–1.4 0.1–0.6
Unconjugated bilirubin
ALP, alkaline phosphatase; ALT, alkaline transaminase; AST, aspartate aminotransferase; CK, creatine kinase; GGT, gamma-glutamyl transferase; SDH, sorbitol dehydrogenase.
152–2835 861–2671 137–1169 210–866 206–458 155–226
ALT
111–432 110–562 139–445 83–233 110–226 83–173
Cholesterol
24–88 30–193 30–239 45–155 28–151 35–76
Triglyceride
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Table 93.3 Normal serum biochemistry values (range, IU/L)3
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Table 93.4 Normal serum electrolyte concentrations3
Age
Na+ (mEq/L)
K+ (mEq/L)
Cl− (mEq/L)
CO2 (mEq/L)
HPO4 (mg/dL)
Ca2+ (mg/dL)
Mg2+ (mg/dL)
Anion gap (mEq/L)
< 12 hours 1 day 1 week 1 month 3 months 6 months
133–163 123–159 130–154 136–154 140–156 133–153
3.4–5.4 3.6–5.6 3.8–5.8 3.6–5.4 3.4–5.8 2.8–5.6
93–117 90–114 94–110 97–109 102–110 98–112
20–30 21–33 24–32 22–32 24–30 22–30
3.1–6.3 3.6–7.2 5.4–9.4 6.9–8.9 6.3–8.3 4.8–7.6
10.8–14.8 9.7–13.7 11.3–13.7 11.0–13.4 11.2–13.2 10.2–13.4
0.7–2.3 0.6–4.2 1.4–2.6 1.0–2.0 1.6–2.8 1.7–3.1
9–23 8–24 11–25 13–25 12–28 11–25
Table 93.5 Normal clotting profile tests4–7 1 day
1 week
> 1 week
6.8–12.8 124–333 11.6–14.6 32.6–55.8 102 000–298 000 – 5.5–10.9
1.7–8.1 130–288 10.5–14.9 25.8–56.3 127 000–273 000 152–495 2.2–9.0
2.5–5.8 158–281 9.4–15.2 10.7–59.7 151 000–249 000 – 4.5–3.1
28.7 66.1–148.1 51.6–75.4 89.9–135.9 66.5–98.1 0.9–3.5
– 129.4–199.4 – – 83.1–115.3 –
– 130.9–211.8 82.8–104.0 72.6–86.6 83.7–112.5 0.8–10.0
13.2–64.8
–
10.7–32.3
104–389
160–418
278–495
Test BT (min) ACT (s) PT (s) APPT (s) Platelet/µL D-Dimer (ng/mL) Fibrin (ogen) degradation products (µg/mL + 1)1/2 Antithrombin III (mg/dL) Antithrombin III activity (%) Protein C antigen (%) Protein C activity (%) Plasminogen (%) Tissue plasminogen activator (u/mL) Plasminogen activator inhibitor (u/mL) Fibrinogen (mg/dL)
ACT, activated clotting time, APPT, activated partial thromboplastin time; BT, bleeding time; PT, prothrombin time.
Table 93.6 Normal ranges for serum iron, iron-binding capacities and ferritin concentrations in foals8
Age < 12 h 1 day 1 week 1 month 3 months 6 months
Iron (µg/dL)
Unsaturated iron-binding capacity (µg/dL)
Total iron-binding capacity (µg/dL)
Iron saturation (%)
Ferritin (ng/mL)
262–488 78–348 30–273 49–288 61–306 85–264
10–133 28–416 35–503 250–668 163–478 159–467
339–535 208–620 222–619 437–777 410–596 341–635
69–98 22–90 10–72 9–50 12–63 16–58
Not done 79–263 57–173 33–140 33–223 81–331
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Table 93.7 Normal lactate values in foals9
Table 93.10 Mean cortisol values in foals11
Age
Age
Mean lactate value (mmol/L)
At birth 24 h 48 h
2.38 1.24 1.08
At birth 1h 6h 2–3 days
Full term
Premature
80 ng/mL 120–140 ng/mL 60 ng/mL 52.5 ng/mL
18–19 ng/mL 30 ng/mL 30 ng/mL
Table 93.8 Cerebrospinal fluid analysis10 Table 93.11 Adrenocorticotropic hormone11
Parameter
Value
Color Total protein
Clear 100 mg/dL ± 50 (Biuret method) 1.3347–1.3350 (refractive index) 80% of blood glucose value 7.34–7.40 < 5 cells/µL 0 to 500 cells/µL 15.2 ± 9.2 142.6 ± 2.8 3.6 ± 2.1 109 ± 3.4
Glucose pH WBC count RBC count Creatine kinase IU/L Sodium (mmol/L) Potassium (mmol/L) Chloride (mmol/L)
RBC, red blood cell; WBC, white blood cell .
Age
Full term
Premature
At birth 2h
250 pg/mL 50 pg/mL
400 pg/mL 100 pg/mL
Table 93.12 Mean maximum increase in cortisol following 0.125 mg exogenous adrenocorticotropic hormone intramuscularly within 2–29 h after birth11 Age
Full term
Premature
2–29 h Increase by 85.8 ± 10.1 ng/mL Increase by 9.3 ± 2.2 ng/mL 31–48 h Increase by 46.5 ± 5.7 ng/mL
Table 93.9 Mean total triiodothyronine (T3 ) and thyroxine (T4 ) concentrations11, 12
Age 1–10 h 1–3 days 5–11 days 12–16 days 28 days 22–90 days
Total T3 (ng/dL)
Total T4 (µg/dL)
Free T3 (pg/mL)
Free T4 (pg/mL)
991 940 631
28.86 28.1 7.4
2.99
12.12
292
2.7
192
2.6
Table 93.13 Response to adrenocorticotropic hormone stimulation using cosyntropin13 Age 3–4 days
3.4
10 µg
100 µg
Peak at 30 min
Peak at 90 min
12.0
Table 93.14 Normal arterial blood gas values8 Age 1–12 h 12–48 h 48 h to 1 week
pH
P a co2 (mmHg)
P a o2 (mmHg)
HCO3 (mEq/L)
Position
7.378 ± 0.015 7.374 ± 0.004 7.384 ± 0.014
42.2 ± 1.8 44.5 ± 1.2 42.4 ± 1.0
77.3 ± 3.1 83.2 ± 3.1 88.2 ± 5.9
26.3 ± 1.1 25.3 ± 1.0 26.2 ± 1.1
Lateral Lateral Lateral
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Table 93.15 Normal values for standard urinalysis14 Specific gravity
pH
1.001–1.027
Protein
5.5–8.0 Neg to +30
Glucose
Crystals
Neg
Usually none; rare calcium oxalate or amorphous urate
Casts Hemoprotein Neg
Neg to +2
Bacteria
Epithelial cells
Neg except Squamous free catch or caudate
RBC
WBC
Mucus
Neg
+3/HPF
Neg to abundant
HPF, high powered field; RBC, red blood cell; WBC, white blood cell.
Table 93.16 Urinary diagnostic indices14
Age 1 day 4 days
Serum urea nitrogen (mg/dL)
Serum creatinine (SCr ) (mg/dL)
Urine creatinine (Ucr ) (mg/dL)
26.5 ± 13.7 5.9 ± 3.3
1.61 ± 0.79 1.0 ± .10
26.5 ± 13.7
UCr /SCr
Serum osmolarity (SOsm ) (mOsm/kg)
Urine osmolarity (UOsm ) (mOsm/kg)
UOsm /SOsm
25.47 ± 14.98
244.5 ± 18.7
101.7 ± 24
0.41 ± .09
Table 93.17 Urinary diagnostic indices continued14
Age 1 day 4 days
Urine alkaline phosphatase (IU/L) 2.3 ± 1.8
Urine alkaline phosphatase/UCr
Serum alkaline phosphatase (IU/L)
Urine GGT (IU/L)
38.5
1500–3000
2.4 ± 2
UGGT /UCr 8.2 ± 5.7 0–16.6
Serum GGT (IU/L) 17–58
GGT, gamma-glutamyl transferase.
Table 93.18 Fractional excretion of endogenous substances14 Age 4 days Adult
Na
K
Cl
P
Ca
0.31 ± 0.18 0.02–1
13.26 ± 4.49 15–65
0.42 ± 0.32 0.04–1.6
3.11 ± 3.81 0–0.5
2.85 ± 3.26 ≥ 1.49–2.5
Table 93.19 Peritoneal fluid analysis15
RBC count
Total nucleated cell count
Neutrophilsa
100–19 900 × 106 /L
< 1400/µL
2–94%
Lymphocytes
Large mononuclear cellsa
Eosinophils
Basophils
Total protein
0–7%
5–98%
0–4
0
< 2.0 g/dL
a Neutrophils and large mononuclear cells should be near equal proportion. RBC, red blood cell.
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Table 93.20 Synovial fluid analysis15 Factor
Description
Appearance
Pale yellow, clear or slightly turbid, high viscosity
Total RBC
Very few, < 1400/µL
Total nucleated cell count
< 500/µL
Total protein
< 1.2 g/dL
Cytology
< 10% neutrophils, > 90% mononuclear cells
RBC, red blood cells.
References 1. Duggan VE, Holyoak GR, MaCallister CG, Confer AW. Influence of induction of parturition on the neonatal acute phase response in foals. Theriogenology 2007;67:372– 81. 2. Harvey JW. Normal hematologic values. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Lea & Febiger, Philadelphia & London, 1990; pp. 561–70. 3. Bauer JE. Normal blood chemistry. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Lea & Febiger, Philadelphia & London, 1990; pp. 602–14. 4. Armengou L, Monreal L, Taracon I, Navarro M, Rios J, Segura D. Plasma d-dimer concentration in sick newborn foals. J Vet Intern Med 2008;22:411–17. 5. Henry Barton M, Deem Morris D, Crowe N, Collatos C, Prasse KW. Hemostatic indices in healthy foals from birth to one year of age. J Vet Diagn Invest 1995;7:380–5.
6. Darien BJ, Carleton C, Kurdowska A, Stickle J, Estry DW, Williams MA, Brown CM, Travis J. Haemostasis and antithrombin III in the full-term newborn foal. Comp Haematol Int 1991;1:161–5. 7. Clemmons RM. Approach to hemostatic problems. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Lea & Febiger, Philadelphia & London, 1990; p. 617. 8. Koterba AM. Respiratory disease: approach to diagnosis. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Lea & Febiger, Philadelphia & London, 1990; pp. 168–9. 9. Magdesian KG. Blood lactate levels in neonatal foals: normal values and temporal effects in the post-partum period. J Vet Emerg Crit Care 2003;13:174. 10. Green SL, Mayhew IG. Neurologic disorders. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Lea & Febiger, Philadelphia & London, 1990; p. 501. 11. Shaftoe S. Peripartum endocrinologic adaptation. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Lea & Febiger, Philadelphia & London, 1990; pp. 45–53. 12. Toribio RE, Duckett WM. Thyroid gland. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine. Saunders, St Louis, 2004; p. 1347. 13. Hart KA, Ferguson DC, Heusner GL, Barton MH. Synthetic adrenocorticotropic hormone stimulation tests in healthy neonatal foals. J Vet Intern Med 2007;21:314–21. 14. Brewer BD. Renal disease. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Lea & Febiger, Philadelphia & London, 1990; pp. 448–9. 15. Parry BW. Normal clinical pathology data. In: Robinson NE (ed.) Current Therapy in Equine Medicine 5. Saunders, St Louis, 2003; pp. 870–84.
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Chapter 94
SI Conversion Table The source of the SI conversion table is the JAMA instructions for authors and is reprinted with permission from http://jama.amaassn.org/content/ vol295/issue1/images/data/103/DC6/JAMA auinst si.dtl.
Conventional units of measure are preferred, with Syst`eme International (SI) units expressed secondarily (in parentheses). To convert values from conventional units to SI units, multiply by the conversion factor.
Component
Conventional unit
Conversion factor
SI unit
Acetaminophen Acetoacetic acid Acetone Acid phosphatase Alanine Alanine aminotransferase (ALT) Albumin Alcohol dehydrogenase Aldolase Aldosterone Alkaline phosphatase Aluminum Aminobutyric acid Amitriptyline Ammonia (as NH3 ) (dual report) Amylase Androstenedione Angiotensin I Angiotensin II Anion gap Antidiuretic hormone Antithrombin III α 1 -Antitrypsin (dual report) Apolipoprotein A Apolipoprotein B Arginine Asparagine Aspartate aminotransferase (AST) Bicarbonate Bilirubin (dual report) Blood gases (arterial) P a co2
µg/mL mg/dL mg/dL units/L mg/dL units/L g/dL units/L units/L ng/dL units/L ng/mL mg/dL ng/mL µg/dL units/L ng/dL pg/mL pg/mL mEq/L pg/mL mg/dL mg/dL mg/dL mg/dL mg/dL mg/dL units/L mEq/L mg/dL
6.62 0.098 0.172 1.0 112.2 1.0 10 1.0 1.0 0.0277 1.0 0.0371 97 3.61 0.587 1.0 0.0349 0.772 0.957 1.0 0.923 10 0.184 0.01 0.01 57.4 75.7 1.0 1.0 17.1
µmol/L mmol/L mmol/L U/L µmol/L U/L g/L U/L U/L nmol/L U/L µmol/L µmol/L nmol/L µmol/L U/L nmol/L pmol/L pmol/L mmol/L pmol/L mg/L µmol/L g/L g/L µmol/L µmol/L U/L mmol/L µmol/L
mmHg
1.0
mmHg
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Foal: Appendix
Component pH P a o2 Bromide C-peptide C1 esterase inhibitor C3 complement C4 complement Calcitonin Calcium (dual report) Carbon dioxide Carboxyhemoglobin Carotene Ceruloplasmin Chloride Cholesterol (dual report) Citrate Copper Coproporphyrins (urine) Corticotropin (ACTH) Cortisol Cotinine Creatine Creatine kinase (CK) Creatinine (dual report) Creatinine clearance (dual report) Cyanide Dehydroepiandrosterone (DHEA) Desipramine Diazepam Digoxin (dual report) Epinephrine Erythrocyte sedimentation rate Estradiol (dual report) Estriol Estrone Ethanol (ethyl alcohol) Ethylene glycol Ferritin α-Fetoprotein Fibrinogen Fluoride Folate Follicle-stimulating hormone Fructose Galactose Glucagon Glucose (dual report) Glutamine
Conventional unit
Conversion factor
pH units mmHg mg/dL ng/mL mg/dL mg/mL mg/mL pg/mL mg/dL mEq/L mEq/L % of hemoglobin saturation µg/dL mg/dL mEq/L mg/dL mg/dL µg/dL µg/24 hr pg/mL µg/dL ng/mL mg/dL units/L mg/dL mL/min
1.0 1.0 0.125 0.333 10 0.01 0.01 1.0 0.25 0.50 1.0 0.01 0.0186 10 1.0 0.0259 52.05 0.157 1.527 0.22 27.59 5.68 76.26 1.0 88.4 0.0167
pH units mmHg mmol/L nmol/L mg/L g/L g/L ng/L mmol/L mmol/L mmol/L Proportion of hemoglobin saturation µmol/L mg/L mmol/L mmol/L µmol/L µmol/L nmol/d pmol/L nmol/L nmol/L µmol/L U/L µmol/L mL/s
mg/L ng/mL
23.24 3.47
µmol/L nmol/L
ng/mL µg/mL ng/mL pg/mL mm/h
3.75 3.512 1.281 5.46 1.0
nmol/L µmol/L nmol/L pmol/L mm/h
pg/mL ng/mL ng/dL mg/dL mg/L ng/mL ng/mL mg/dL µg/mL ng/mL mIU/mL
3.671 3.467 37 0.217 16.11 2.247 1.0 0.0294 52.6 2.266 1.0
pmol/L nmol/L pmol/L mmol/L µmol/L pmol/L µg/L µmol/L µmol/L nmol/L IU/L
mg/dL mg/dL pg/mL mg/dL mg/dL
55.5 55.506 1.0 0.0555 68.42
µmol/L
SI unit
µmol/L ng/L mmol/L µmol/L
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SI Conversion Table
Component γ -Glutamyltransferase (GGT) Glycated hemoglobin (glycosylated) hemoglobin A1 , A1C ) Glycerol (free) Glycine Haptoglobin Hematocrit Hemoglobin (whole blood)Mass concentration High-density lipoprotein cholesterol (HDL-C) (dual report) Histidine Homocysteine (total) Human chorionic gonadotropin (HCG) Hydroxybutyric acid Hydroxyproline Immunoglobulin A (IgA) Immunoglobulin D (IgD) Immunoglobulin E (IgE) Immunoglobulin G (IgG) Immunoglobulin M (IgM) Insulin Iron, total (dual report) Iron-binding capacity, total (dual report) Isoleucine Isopropanol Lactate (lactic acid) Lactate dehydrogenase Lactate dehydrogenase isoenzymes (LD 1 –LD5 ) Lead (dual report) Leucine Lipase Lipids (total) Lipoprotein (a) Lithium Low-density lipoprotein cholesterol (LDL-C) (dual report) Luteinizing hormone (LH, leutropin) Lysine Magnesium (dual report) Manganese Methanol Methemoglobin
Conventional unit
Conversion factor
SI unit
units/L
1.0
U/L
% of total hemoglobin
0.01
Proportion of total hemoglobin
mg/dL mg/dL mg/dL % g/dL
108.59 133.3 0.10 0.01 10.0
µmol/L µmol/L Proportion of 1.0 g/L
mg/dL
0.0259
mmol/L
mg/dL mg/L mlU/mL
64.45 7.397 1.0
µmol/L
mg/dL mg/dL mg/dL mg/dL mg/dL mg/dL mg/dL µIU/mL µg/dL µg/dL
96.05 76.3 0.01 10 10 0.01 0.01 6.945 0.179 0.179
µmol/L g/L mg/L mg/L g/L g/L pmol/L µmol/L µmol/L
mg/dL mg/L mg/dL units/L %
76.24 0.0166 0.111 1 0.01
µmol/L mmol/L mmol/L U/L Proportion of 1.0
µg/dL mg/dL units/L mg/dL mg/dL mEq/L mg/dL
0.0483 76.237 1.0 0.01 0.0357 1.0 0.0259
µmol/L µmol/L U/L g/L µmol/L mmol/L mmol/L
IU/L
1.0
IU/L
mg/dL mg/dL mEq/L ng/mL mg/L % of total hemoglobin
68.5 0.411 0.50 18.2 0.0312 0.01
µmol/L mmol/L mmol/L nmol/L mmol/L Proportion of total hemoglobin
µmol/L
µmol/L IU/L µmol/L
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Foal: Appendix
Component Methionine Myoglobin Nicotine Nitrogen, non-protein Norepinephrine Ornithine Osmolality Osteocalcin Oxalate Parathyroid hormone Phenobarbital Phenylalanine Phenytoin Phosphorus (dual report) Plasminogen Plasminogen activator inhibitor Platelets (thrombocytes) Potassium Pregnanediol (urine) Pregnanetriol (urine) Progesterone Prolactin Proline Prostate-specific antigen Protein, total Prothrombin Prothrombin time (protime, PT) Protoporphyrin, erythrocyte Pyruvate Quinidine Red blood cell count Renin Reticulocyte count Salicylate Serine Serotonin (5-hydroxytryptamine) Sodium Somatomedin-C (insulin-like growth factor) Somatostatin Taurine Testosterone (dual report) Theophylline Thiocyanate Threonine Thyroglobulin Thyrotropin (thyroid-stimulating hormone, TSH)
Conventional unit
Conversion factor
mg/dL mg/L mg/dL pg/mL mg/dL mOsm/kg µg/L mg/L pg/mL mg/L mg/dL µg/mL mg/dL mg/dL % mIU/mL
67.02 0.0571 6.164 0.714 0.00591 75.67 1.0 0.171 11.1 1.0 4.31 60.54 3.96 0.323 0.113 0.01 1.0
mmol/L nmol/L µmol/L mmol/kg nmol/L µmol/L ng/L µmol/L µmol/L µmol/L mmol/L µmol/L Proportion of 1.0 IU/L
×103 /µL mEq/L mg/24 h mg/24 h ng/mL µg/L mg/dL ng/mL g/dL g/L s
1.0 1.0 3.12 2.97 3.18 43.478 86.86 1.0 10.0 13.889 1.0
×109 /L mmol/L µmol/d µmol/d nmol/L pmol µmol/L µg/L g/L µmol/L s
µg/dL
0.01777 113.6 3.08 1.0 0.0237 0.01 0.00724 95.2 0.00568
µmol/L
mg/dL µg/mL ×106 /µL pg/mL % of RBCs mg/L mg/dL ng/mL mEq/L ng/mL
1.0 0.131
pg/mL mg/dL ng/dL µg/mL mg/L mg/dL ng/mL mIU/L
0.611 79.91 0.0347 5.55 17.2 83.95 1.0 1.0
µg/L
SI unit µmol/L
nmol/L µmol/L
µmol/L µmol/L ×1012 /L pmol/L Proportion of 1.0 mmol/L µmol/L µmol/L
mmol/L nmol/L (coagulation factor II) pmol/L µmol/L nmol/L µmol/L µmol/L µmol/L µg/L mIU/L
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SI Conversion Table
Component Thyroxine, free (T4 ) (dual report) Thyroxine, total (T4 ) (dual report) Transferrin Triglycerides (dual report) Triiodothyronine Free (T3 ) (dual report) Resin uptake Total (T3 ) (dual report) Troponin I (cardiac) Troponin T (cardiac) Tryptophan Tyrosine Urea nitrogen (dual report) Uric acid Valine Vasoactive intestinal polypeptide Vitamin A (retinol) Vitamin B6 (pyridoxine) Vitamin B12 (cyanocobalamin) Vitamin C (ascorbic acid) Vitamin D 1,25-Dihydroxyvitamin D 25-Hydroxyvitamin D Vitamin E Vitamin K Warfarin White blood cell count White blood cell differential count (number fraction) Zinc
Conventional unit
Conversion factor
SI unit
ng/dL
12.87
pmol/L
µg/dL
12.87
nmol/L
mg/dL mg/dL
0.01 0.0113
g/L mmol/L
pg/dL % ng/dL ng/mL ng/mL mg/dL mg/dL mg/dL mg/dL mg/dL pg/mL
0.0154 0.01 0.0154 1.0 1.0 48.97 55.19 0.357 59.48 85.5 1.0
pmol/L Proportion of 1.0 nmol/L µg/L µg/L µmol/L µmol/L mmol/L µmol/L µmol/L ng/L
µg/dL
0.0349 4.046 0.738
µmol/L
ng/mL pg/mL mg/dL
56.78
µmol/L
pg/mL ng/mL mg/dL ng/mL µg/mL ×103 /µL %
2.6 2.496 23.22 2.22 3.247 1.0 0.01
pmol/L nmol/L µmol/L nmol/L µmol/L ×109 /L Proportion of 1.0
µg/dL
0.153
µmol/L
861
nmol/L pmol/L
Dual report, because of the continued use of conventional units in many laboratories in the USA, some of the more common analytes currently are dual reported, with the SI value and unit listed first, followed by the conventional value and unit. Analytes with a 1-to-1 conversion between SI and conventional units or with a conversion factor that is a multiple of 10 usually are not dual reported. Information on how to convert from SI units to conventional units may be given in the text, in a table footnote, or in a figure legend (e.g., to convert ethanol from millimoles per liter to milligrams per deciliter, divide millimoles per liter by 0.217). From Iverson et al.,1 Tietz,2 Jacobs et al.,3 Young and Huth,4 and Kratz and Lewandrowski.5
References 1. Iverson C, Flanagin A, Fontanarosa PB, Glass RM, Geitman P, Lantz JC, Meyer HS, Smith JM, Winker MA, Young RK. American Medical Association Manual of Style: A Guide for Authors and Editors, 9th edn. Baltimore, MD: Williams & Wilkins, 1998. 2. Tietz NW (ed.). Clinical Guide to Laboratory Tests, 3rd edn. Philadelphia: W.B. Saunders Co, 1995.
3. Jacobs DS, Demott WR, Grady HJ, Horvat RT, Huestis DW, Kasten Jr BL (eds). Laboratory Test Handbook, 4th edn. Hudson, OH: Lexi-Comp Inc., 1996. 4. Young DS, Huth EJ. SI Units for Clinical Measurement. Philadelphia: American College of Physicians, 1998. 5. Kratz A, Lewandrowski KB. Normal reference laboratory values. N Engl J Med 1998; 339: 1036–42.
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PART II
The Stallion
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Section (a): Anatomy, Physiology and Endocrinology
95.
Functional Anatomy of the Adult Male Rupert P. Amann
867
96.
Physiology and Endocrinology Rupert P. Amann
881
97.
From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell Dickson D. Varner and Larry Johnson
909
98.
Oxidative Stress in Sperm Barry A. Ball
991
99.
Endocrine–Paracrine–Autocrine Regulation of Reproductive Function in the Stallion Janet F. Roser
996
100.
Puberty Noah L. Heninger
1015
101.
Spermatogenesis Larry Johnson, Cleet E. Griffin and Michael T. Martin
1026
102.
Spermatozoa Function Rupert P. Amann and James K. Graham
1053
103.
Sperm-Oviduct Interactions Barry A. Ball
1085
104.
Sperm–Uterine Interactions Terttu Katila
1092
105.
Testicular Descent Mimi Arighi
1099
865
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Chapter 95
Functional Anatomy of the Adult Male Rupert P. Amann
To evaluate properly a stallion suspected of having reproductive problems, the normal location, shape, and size of each organ must be known. The male reproductive organs consist of two testes, each suspended by a spermatic cord and external cremaster muscle; two epididymides; two deferent ducts each with an ampulla; paired vesicular glands; a prostate gland; paired bulbourethral glands; the penis; and the associated urethralis, ischiocavernosus, bulbospongiosus, and retractor penis muscles∗ (Figure 95.1). The vesicular glands, prostate gland, and bulbourethral glands often are termed the accessory sex glands. The reproductive tract is supported within the pelvic cavity by the hammock-like genital fold and externally by the scrotum and prepuce.
Scrotum The scrotum is an outpouching of skin composed of two scrotal sacs, one for each testis, separated by the septum scroti. The sacs lie on either side of the penis (Figure 95.2). If one testis is larger than the other, the scrotum may appear asymmetric or lopsided. The scrotum consists of four layers. The outermost layer is the skin, which contains an unusually large number of sweat glands. Underlying the skin and associated connective tissue is the tunica dartos. The tunica dartos is a layer of smooth muscle fibers intermingled with connective tissue, rather than a discrete muscle. This layer forms the outermost component of each scrotal sac and, by raising or lowering the testis, the smooth muscle fibers aid in control of testicular temperature. The third layer is loose connective tissue, or scrotal fascia, which allows the testis great mobility for ver-
tical or horizontal movement within the scrotal sac. Normally, the loose connective tissue prevents 180◦ rotation of the testis within the scrotal sac, although in certain stallions such rotation does occur.7–9 The impact of rotation of a testis on reproductive capacity is discussed in Chapter 110. The innermost layer of the scrotum is the parietal vaginal tunic (some anatomists consider this covering as part of the testis). The parietal vaginal tunic (lamina parietalis of tunica vaginalis, also called the common vaginal tunic) is a membranous sac that extends from the abdominal cavity through the inguinal canal (cannalis inguinalis, an opening in the abdominal wall through which the spermatic cord passes) to the bottom of the scrotum. Part of this membrane covers the testis and epididymis and also contributes to the spermatic cord (funiculus spermaticus). The space within the vaginal cavity (cavum vaginale) – between the parietal vaginal tunic and visceral vaginal tunic (lamina visceralis of tunica vaginalis), which is the outermost covering around the testis and epididymis (Figure 95.2) – contains a watery, serous fluid, which serves as a lubricant and facilitates movement of the testis within the sac formed by the parietal vaginal tunic. In older stallions, adhesions sometimes develop between the parietal vaginal tunic and the visceral vaginal tunic, which covers the testis. Possibly, adhesions arise following slight hemorrhage from capillaries within the tunics, induced by slight trauma to the scrotum and testes as part of the normal actions of a stallion. Such adhesions impede the mobility of the testis and may reduce effectiveness of temperature control mechanisms.
∗
The general anatomic description is based on the author’s observations1 and standard texts.2−6 References for newer anatomic observations and for specific physiological principles are supplied in the text.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Stallion: Anatomy, Physiology and Endocrinology Ductus deferens Ampulla
Spine
Vesicular gland Rectum Prostate gland
Genital fold Pelvic urethra Urinary bladder
Bulbourethral gland
Testicular artery and vein
Floor of pelvis
Retractor penis muscle
Internal inguinal ring Penile urethra Spermatic cord within inguinal canal Bulbospongiosus muscle
Corpus spongiosum penis Glans penis Corpus cavernosum penis Prepuce
Testis
Scrotum Epididymis
Figure 95.1 The reproductive tract of the stallion as seen in a left lateral dissection. (Modified from Pickett et al.1 )
Descent of the testes into the scrotum10 is described in Chapter 105. The truly cryptorchid testis, one retained within the abdominal cavity, apparently occurs when the testis fails to enter the inguinal canal before closure of the internal inguinal ring (annulus inguinalis profundus) during the first 2 weeks after birth. The high incidence of failure of the left testis to descend9 might result from the relatively slow rate of descent of the left epididymis and testis. Obviously, this could result in a higher incidence of retention of the left testis compared with the right.
Diagnosis of cryptorchidism should include careful external palpation of the scrotum and external inguinal ring, as well as palpation per rectum of the internal inguinal ring and pelvic area. Alternatively, cryptorchidism can be diagnosed by ultrasonography.11 During the first several weeks after birth, the gubernaculum may be quite large and should not be confused with a testis. At birth, the weight of each testis is 5–20 g, and testicular size increases very little through 10 months of age. More rapid development of the testes generally starts around 12–18 months of
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Functional Anatomy of the Adult Male
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Corpus cavernosum penis Penis Lymphatic vessel Urethra Vaginal cavity
Fibroblast
Corpus epididymidis Ductus deferens Central vein
Blood vessel
Skin Tunica dartos Tunica albuginea Testicular parenchyma
Parietal vaginal tunic
Leydig cells
Visceral vaginal tunic
Figure 95.2 The penis, scrotum and testes of the stallion as seen in a vertical cross-section. (Modified from Pickett et al.1 )
age, but the age at which rapid growth of the testes commences varies considerably.
Testis The testis is the male gonad and the site of production for both spermatozoa and the predominant male sex hormone, testosterone. The testes are ovoid, slightly compressed from side to side with their long axis almost horizontal (Figure 95.1). When a testis is retracted, the long axis becomes more vertical, so the cauda epididymidis becomes ventrally rather than caudally located. Testes of postpubertal stallions range considerably in size. An average testis might measure 80–140 mm in length by 50–80 mm in width and weigh about 225 g. However, age and season greatly affect testis weight; no definitive data show that draft stallions have larger (or smaller) testes than light horse breeds, but, for cattle, significant breed differences have been reported.12 If the testis is exposed, as during open castration, the thick tunica albuginea is seen (Figure 95.2). Fused to the outer surface of this connective tissue capsule is the thin visceral vaginal tunic. Supporting strands of connective tissue extend from the tunica albuginea to divide the testis into lobules (lobuli testis). The parenchyma, or non-capsular part of the testis, is lightly pigmented in a young postpubertal stallion, moderately pigmented in a 4- to 5-year-old stallion, and darkly pigmented in an aged stallion.13 The testicular parenchyma consists of seminiferous
Seminiferous tubules
Figure 95.3 A stallion testis showing the relationship among the blood vessels, lymphatic vessels, and Leydig cells of the interstitial tissue and seminiferous tubules. (From Pickett et al.1 )
tubules (tubuli seminiferi) and interstitial tissue (Figure 95.3). The seminiferous tubules (tubuli seminiferi contorti) (Figure 95.4) are lined by a seminiferous epithelium (epithelium spermatogenicum) that consists of different types of germinal cells (cellulae spermatogenicae) (see Chapters 97 and 101 and the Sertoli cells (sustentacular cells). The seminiferous tubule is limited by a lamina propria consisting of fibroblasts, myoid cells (specialized smooth muscle cells), and laminin. Rhythmic contractions of the myoid cells presumably help to move spermatozoa and fluids from the seminiferous tubules. The germinal cells are the most conspicuous feature of the seminiferous epithelium, but the Sertoli cells have a pivotal role in coordinating germinal cell differentiation and isolating most germinal cells from the host stallion’s immune system. Junctional complexes between adjacent Sertoli cells (Figure 95.4) form the major component of the blood–testis barrier and divide the seminiferous epithelium into two functional compartments, or regions: basal (peripheral) and adluminal (inner). The blood–testis barrier isolates the more differentiated germinal cells from the immune system of the host stallion. Because the developing immune system is not exposed to differentiated spermatocytes or spermatids, it considers them to be foreign cells. Therefore, except for this isolation provided by the blood–testis barrier, germinal cells would be
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Adluminal compartment
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Spermatids
Primary spermatocyte
Basal compartment
Junctional complex
Sertoli cell
Spermatogonium Lamina propria Interstitial compartment
c95
Leydig cell Blood vessel
Figure 95.4 A section of a stallion seminiferous tubule showing the relationship of germinal cells and adjacent Sertoli cells in seminiferous epithelium. Spermatogonia, primary spermatocytes, secondary spermatocytes, and spherical spermatids all develop in the space between two or more Sertoli cells and are in contact with them. Primary spermatocytes are moved, by the Sertoli cells, from the basal compartment through the junctional complexes and into the adluminal compartment. During elongation of spermatids, they are repositioned by the Sertoli cells to become embedded within long pockets in cytoplasm of individual Sertoli cells. Note the intercellular bridges between adjacent germinal cells in the same cohort or generation. (From Pickett et al.1 )
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Functional Anatomy of the Adult Male destroyed. Damage to the blood–testis barrier is rare in stallions, but can cause damage to the testis, reduce spermatozoal production, or induce sterility. Because Sertoli cells largely surround developing germinal cells, the immediate environment around all germinal cells, except spermatogonia, is controlled by Sertoli cells.14–17 Sertoli cells produce several proteins needed to carry vitamin A, iron, and copper to developing germ cells and also a number of unique proteins involved in regulating formation of spermatozoa. Many compounds adversely affecting spermatogenesis probably act on Sertoli cells rather than directly on germinal cells. Furthermore, the general number of Sertoli cells per testis and maximum number of germinal cells per Sertoli cell are characteristics of a species,18 but the number of Sertoli cells per testis is a trait with high heritability.19 Any treatment that decreases the number of Sertoli cells formed before puberty or alters Sertoli cell function will probably adversely affect spermatozoal production. The interstitial tissue (interstitium testis) includes blood vessels, lymphatic channels, nerves, connective tissue, and Leydig cells (endocrinocytus interstitialis). Leydig cells, which are responsible for production of steroid hormones including testosterone, are the major component of the interstitial tissue in adults.13 As a stallion grows older, postpubertal Leydig cells are gradually replaced by adult Leydig cells, which contain abundant pigmented lipids and are interconnected by numerous interdigitations, 20 and they may produce more testosterone per cell than Leydig cells of younger stallions. The percentage composition of the testis, on a volume basis, changes with age. The ratio of Leydig cells to seminiferous tubules increases from 1:12 in stallions 2–3 Table 95.1 Age-related differences in testicular compositiona Age (years)
Testicular weight (g) Parenchyma Tunica albuginea Parenchymal composition (%) Leydig cells Other interstitial tissue Seminiferous tubules Seminiferous tubule Diameter (m) Length/testis (m) Daily spermatozoal production (109 /testis)
2–3
4–5
13–20
105a 12a
146b 15b
184c 29c
6a 22a 72
12b 16b 72
18c 10c 72
212a 2040b 1.3a
230b 2390a,b 2.7b
242c 2790b 3.2c
a,b,c Means in the same row with different superscripts differ (P < 0.05). Adapted from Johnson and Neaves.13
871
200
40
150
30
20
100
10
50
0
2–3 4–5 13–20 2–3 4–5 13–20 Age (years)
µL Leydig cells / g testis
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Figure 95.5 Effect of age on number of Sertoli cells and volume of Leydig cells per gram of testicular parenchyma. Data for Sertoli cells are for testes collected in June and July and data for Leydig cells are for testes obtained between February and May. (From Pickett et al.1 )
years old to 1:4 for those 13–20 years old (Table 95.1). Although certain of the changes presented in Table 95.1 are a consequence of testicular growth, composition of the testis on a per gram basis also changes with age. This is especially evident for the number of Sertoli cells per gram of testicular parenchyma,21 and the decline with advancing age (Figure 95.5) may be partially responsible for the decline in daily spermatozoal production which occurs in some older stallions. Although percentage of the testis occupied by seminiferous tubules does not change with age (Table 95.1), the total volume of Leydig cells per gram of testicular parenchyma increases more than threefold with age (Figure 95.5). The total length of seminiferous tubules in the testis also increases by about onethird.13 A seminiferous tubule is arch shaped and consists of three zones. The major portion of a seminiferous tubule, the convoluted seminiferous tubule, is highly coiled and is the site of spermatozoal production. A convoluted seminiferous tubule is lined by a seminiferous epithelium, which contains Sertoli cells and several types of germinal cells.1, 2,13–15,17,21–27 Sertoli cells and spermatogonia line the basement membrane of the tubules, and Sertoli cells extend toward the lumen in a radial pattern (Figure 95.4) to occupy 15–25% of the seminiferous epithelium in an adult stallion. Although it is conventionally accepted that Sertoli cells do not divide after puberty,8, 14, 16, 27 the results of recent research demonstrate that Sertoli cells in the stallion proliferate as the breeding season approaches.21, 24, 28, 29 The number of Sertoli cells in a testis is an important determinator of the number of spermatozoa that a testis can produce.22
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Figure 95.6 A scanning electron micrograph (×50) of the interconnecting tubules within the rete testis near the central vein (CV). Rete tubules (R) and smaller blood vessels (B) are comingled in that area. (From Amann et al.30 )
Both ends of a convoluted seminiferous tubule continue as tapered transitional zones leading to the straight portions of the seminiferous tubule. Straight tubules (tubuli seminiferi recti) are the first component of the passageway through which spermatozoa pass from the seminiferous epithelium to the epididymis. Straight tubules converge in the cranial twothirds of the testis30 in an area termed the rete testis (Figures 95.6 and 95.7). Tubules of the intratesticular rete testis penetrate the tunic albuginea and continue as an extratesticular rete.30 Eventually, each rete tubule fuses with one of the 13–15 efferent ducts (ductuli efferentes testis) that lead to the epididymal duct (ductus epididymidis).31 Horses are seasonal breeders and characteristics of the testes (Table 95.2) change markedly throughout the year. However, unlike some seasonal breeders, stallions continue to produce spermatozoa throughout the year. Maximum testicular development and function occur in May, June, and July.23, 29,32–34 From September through February, the testes are regressed; they are probably minimal in size and function from November to January. At that time of year, the testes of typical stallions are 25% lighter; contain about 35% fewer Leydig cells and consequently less smooth endoplasmic reticulum (SER),
the part of the cell involved in production of testosterone; contain 31% fewer Sertoli cells; and produce 40–50% fewer spermatozoa (Table 95.2). Concentrations in the blood of hormones involved in control of reproductive function also tend to be lower in the non-breeding season.
Spermatic cord The spermatic cord extends from the abdominal inguinal ring to its attachment on the testis. It suspends the testis in the scrotum and acts as a passageway for the deferent duct (ductus deferens), nerves, and blood vessels associated with the testis (Figure 95.8). The external cremaster muscle is conspicuous on the lateral aspect of the spermatic cord, although it is not part of the cord. This striated muscle helps to support the testis and aids in control of testicular temperature. The spermatic cord includes the highly coiled testicular artery (Figure 95.7). The veins draining the testis form an intimate network of small veins around the highly coiled artery.35 This network of veins is called the pampiniform plexus and serves as a heat exchange system.
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Ductus deferens Testicular artery Testicular veins Reflected parietal vaginal tunic
Pampiniform plexus Cut edge of visceral vaginal tunic
Corpus epididymidis Convoluted seminiferous tubule
Rete tubule Caput epididymidis
Straight tubule
Ductus epididymidis Efferent duct
Tunica albugines
Cauda epididymidis
Figure 95.7 The location of straight tubules and rete testis in the stallion testis. The straight tubules converge in a group of interconnecting rete tubules that penetrate the tunica albuginea and fuse with the efferent ducts. The efferent ducts merge in the epididymal duct. The testicular artery becomes highly coiled in the pampiniform plexus. After emerging from the pampiniform plexus, the testicular artery passes along the dorsal aspect of the testis to the caudal pole where it starts to branch to vascularize the parenchyma. Venous drainage of the parenchyma is via the central vein and superficial testicular veins. After leaving the testis, the veins form an anastomosing plexus of veins, termed the pampiniform plexus, which is in intimate contact with the testicular artery. About 7–10 cm above the testis, the veins converge into the testicular vein. (Modified from Pickett et al.1 ) Table 95.2 Effect of season on characteristics of testes of stallions 4–20 years olda
Parenchyma weight, paired testes (g) Leydig cells (106 /g testis) (109 /2 testes) Leydig cell SERc (mL/2 testes) Testosterone (µg/g testis) (mg/2 testes) Sertoli cells (106 /g testis) (109 /2 testes) Elongated spermatids per Sertoli cell Daily spermatozoal production (106 /g testis) (109 /2 testes) Serum FSHd (ng/mL) LHe (ng/mL) Testosterone (ng/mL)
Breeding
Non-breeding
Difference (%)b
154
116
−25
22 7.26 34.0
20 4.70 22.2
−9 −35 −35
700 224
580 154
−17 −31
24.2 3.69 9.36
23.6 2.54 7.54
−2 −31 −19
19.1 5.96
14.6 3.53
−23 −41
108 36 0.36
81 22 0.27
−21 −39 −25
Not all data are for the same stallions. For most values n ≥ 48. Not all differences between the breeding and non-breeding seasons were statistically significant in the original studies. Because data were compiled across studies, statistical significance was not determined. c Smooth endoplasmic reticulum. d Follicle-stimulating hormone. e Luteinizing hormone. Based on Johnson L, Thompson Jr DL. Seasonal variation in the total volume of Leydig cells in stallions is explained by variation in cell number rather than cell size. Biol Reprod 1986;35:971–9; and Johnson L, Thompson Jr DL. Effect of seasonal changes in Leydig cells and intratesticular testosterone content in stallions. J Reprod Fertil 1987;81:227–32. From Pickett et al.1 a
b
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Stallion: Anatomy, Physiology and Endocrinology External cremaster muscle
Mesorchium
Parietal vaginal tunic
Mesoductus deferens
Ductus deferens Visceral vaginal tunic
Vaginal cavity
Testicular artery, veins, and nerves
Figure 95.8 Cross-section through the right spermatic cord and external cremaster muscle above the pampiniform plexus. (From Pickett et al.1 )
Excurrent duct system
(a)
After penetrating the tunica albuginea, each rete tubule fuses with an efferent duct. These, in turn, lead to the epididymal duct (ductus epididymidis), which continues as the deferent duct, and terminates in the colliculus seminalis. Epididymis The epididymis is divided anatomically into three parts: caput, corpus, and cauda. The caput curves around the testis and lateral aspect of the spermatic cord (Figures 95.1 and 95.7) and continues as the corpus of the epididymis. The caput is rather flat, has a J shape and is closely attached to the testis. The corpus epididymidis is a cylindric structure loosely attached to the dorsal surface of the testis (Figure 95.2). The cauda epididymidis is large, bulbous, and loosely attached to the caudal pole of the testis. The proximal caput epididymidis actually contains the distal ends of 13–15 highly coiled efferent ducts that lead from the tubules of the extratesticular rete testis.30, 31 Within the caput epididymidis, efferent ducts fuse into a single duct termed the epididymal duct.31 This single duct, possibly 45 m long, is folded in pleats and continues in a tortuous pattern (Figure 95.7) through the caput, corpus, and cauda epididymidis and is continuous with the deferent duct. Based on cellular structure, six to eight regions can be distinguished in the stallion epididymis, and the function of each region is probably different.36 From a functional point of view (see Chapter 97), the epididymis has three segments.37–40 Epithelia of the efferent ducts plus the initial segment of the caput (Figures 95.9 and 255.10) are involved in resorption of most of the fluid and solutes entering from the testis
(b) Figure 95.9 (a) Section through an efferent duct showing ciliated and non-ciliated cells in this epithelium (×725). (b) The luminal surface of an efferent duct and a spermatozoon with a proximal cytoplasmic droplet (normal for a spermatozoon from this region) also are shown (×3075). (From Johnson et al.40 )
and also secrete some compounds. The middle segment includes major portions of the caput and corpus epididymidis and is involved in spermatozoal maturation, a process that depends on specific secretions of the epithelium. The terminal segment is composed of the cauda epididymidis (Figure 95.11) and proximal deferent duct and is involved in storage of fertile spermatozoa.
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crypts and glands, although luminal diameter does increase slightly.
Accessory sex glands
(a)
The accessory sex glands (glandulae genitales accessoriae) are sometimes considered to include the ampulla of the deferent duct, but herein are considered to include only the glandula vesicularis, prostata, and glandula bulbourethralis. Functions of these glands, and their contributions to semen are considered in Chapters 97 and 107. The two vesicular glands (previously termed seminal vesicles) are elongated, hollow pouches about 15–20 cm long and 5 cm in diameter (Figure 95.12). In spite of the common use of the term seminal vesicles to describe these glands, the vesicular glands do not serve as a storage organ for spermatozoa. The prostate gland is a single, firm, nodular gland (Figure 95.12) with two narrow lobes (each 7 × 4 × 1 cm) connected by a thin transverse isthmus (about 3 cm long). The two bulbourethral glands (previously termed Cowper’s glands) are positioned on either side of the pelvic urethra near the ischial arch (Figures 95.1 and 95.12).
Urethra
(b) Figure 95.10 Sections through the caput epididymidis showing (a) the epithelium and thin layer of smooth muscle (×60) and (b) the surface of the epithelium and spermatozoa within the duct (×300). (From Johnson et al.40 )
The urethra is a long mucous-secreting tube that extends from the bladder to the free end of the penis. The pelvic portion of the urethra is overlaid by a thick, striated muscle termed the urethralis muscle (Figure 95.12), which contracts vigorously during ejaculation. The urethra terminates in a free extension called the urethral process (processus urethrae) (Figure 95.13). The penile urethra is surrounded by the corpus spongiosum penis (Figure 95.14), which is an area of cavernous, erectile tissue. The urethra serves as the joint excretory canal for urine and semen.
Deferent duct The deferent duct (ductus deferens) is a continuation of the epididymal duct and extends from the cauda epididymidis through the spermatic cord to the pelvic urethra (Figures 95.1 and 95.7). The deferent duct of the stallion has an extremely thick wall of smooth muscle. Thus the proximal deferent duct can be palpated readily through the scrotal skin. As the deferent duct approaches the pelvic urethra, it widens into a structure termed the ampulla of the deferent duct (Figures 95.1 and 95.12). The ampulla is about 18 mm in diameter compared with 4–5 mm for the deferent duct. The increased diameter in the ampullar region is primarily caused by a thickening of the wall associated with the presence of
Penis The penis is the male organ of copulation and consists of three regions (Figure 95.13): the root, which is the site of attachment to the skeletal system; the body or shaft, which is the main portion of the penis; and the glans penis, which is the enlarged, free end of the penis. The primary functional components of the penis are the corpus cavernosum penis; corpus spongiosum penis, which is continuous with the corona glandis of the glans penis; urethra; bulbospongiosus muscle; and associated blood vessels and nerves (Figure 95.14). The stallion penis is a musculocavernosus type and undergoes considerable enlargement in length and diameter during erection, which is in
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Figure 95.11 Sections through the cauda epididymidis. (a) In the proximal cauda, projections of epithelium radiate from the wall of the duct and the smooth muscle layer is of moderate thickness (×53). (b) Details of the projections and smooth muscle (SM) (×225). (c) In the distal cauda, projections are much longer and the smooth muscle layer is thick (×35). (From Johnson et al.40 )
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Urinary bladder Ductus deferens
Ampulla of ductus deferens
Genital fold Ureter
Vesicular gland
Lobe of prostate gland
Ischium
Isthmus of prostate gland
Urethralis muscle Bulbourethral gland Retractor penis muscle
Bulbospongiosus muscle Ischiocavernosus muscle
Figure 95.12 A dorsal view of the pelvic portion of the reproductive tract. The connective tissue and fat have been dissected away to expose the genital fold that supports the pelvic portion of the tract. (From Pickett et al.1 )
Obturator artery
Rectal part of retractor penis muscle
Tuber ischii
Ischiocavernosus muscle
External pudendal artery
Cranial artery of penis
Middle artery of penis
Suspensory ligament of penis
Dorsal process of glans
Dorsal artery of penis Retractor penis muscle
Corona glandis
Bulbospongiosus muscle Corpus cavernosum penis
Urethral process
Neck of glans
Figure 95.13 A left lateral view of the penis and its attachment to the ischium. (From Pickett et al.1 )
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Dorsal artery of penis
Dorsal veins of penis
Dorsal nerve of penis
Tunica albuginea
Corpus cavernosum penis
Trabeculae
Urethra Bulbospongiosus muscle
Corpus spongiosum penis
Retractor penis muscle
nective tissue capsule, the tunica albuginea (Figure 95.14). The root of the penis is tightly attached to the tuber ischii by the paired crua, overlain by the paired ischiocavernosus muscles and stabilized by two suspensory ligaments (Figure 95.13). When viewed in cross-section (Figure 95.14), the corpus cavernosum penis forms the major component of the body of the penis. The corpus cavernosum penis is spongy, erectile tissue, which is continuous with veins draining the penis. The corpus spongiosum penis is a small area of spongy, erectile tissue that immediately surrounds the urethra but is not enclosed by the tunica albuginea. The corpus spongiosum penis is continuous into the glans penis (Figure 95.15), which is richly endowed with nerve endings. The corpus spongiosum penis and corona glandis become engorged with blood and erect during sexual excitement. When not erect, the penis is about 50 cm long by 2.5–5.0 cm in diameter. About 15–20 cm are free in the prepuce (Figure 95.15). Erection increases the length and diameter of the penis about 50%, whereas the glans penis increases 300–400% in diameter through engorgement.
Figure 95.14 A cross-section through the body of the penis. (Adapted from Pickett et al.1 )
contrast to the fibroelastic penis of the bull, which becomes straight and stiff during erection, but does not increase in size. The stallion penis contains a large amount of erectile tissue, some of which surrounds the urethra, but most of which is enclosed in a con-
Hypothalamus and adenohypophysis The hypothalamus and adenohypophysis are described in Chapters 96 and 99. Their secretions are pivotal in the regulation of male reproductive function as considered in Chapters 96 and 99.
Dorsal venous plexus
Abdominal wall Corpus cavernosum penis
Corona glandis Urethra
Urethral sinus
Fossa glandis
Corpus spongiosum penis
Urethral process Preputial fold Preputial orifice Figure 95.15 A left lateral view of the glans penis and prepuce. (Adapted from Pickett et al.1 )
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Acknowledgement Some individuals, and some manuscripts have wonderful insight and historical value. Such is the case with Dr. Rupert P. Amann and four chapters that he so ably produced for the first edition of Equine Reproduction. While Dr. Amann is now retired and has turned the baton over to others, these pieces of work have endured the test of time and continue to be an invaluable resource for the equine practitioner and academician. As such, we decided to reprint these chapters for you.
References 1. Pickett BW, Amann RP, McKinnon AO, Squires EL, Voss JL. Management of the Stallion for Maximum Reproductive Efficiency, II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins: Colorado State University, 1989. 2. Banks WJ. Applied Veterinary Histology, 2nd edn. Baltimore: Williams & Wilkins, 1986. 3. Dyce KM, Sack WO, Wensing CJG. Textbook of Veterinary Anatomy. Philadelphia: W.B. Saunders, 1987. 4. Krolling O, Grau H. Lehrbuch der Histologie und Vergleichenden Microskopischen Anatomie der Haustiere. Berlin: Paul Parey, 1960. 5. Montane M, Bourdelle E, Bressou C. Anatomic Regionale des Animaux Domestiques. I. Equides, 9th edn. Paris: J.B. Bailliere, 1949. 6. Nickel R, Schummer A, Seiferle E, Sack WO. The Viscera of the Domestic Mammals. New York: Springer-Verlag, 1973. 7. Pickett BW, et al. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. In Proceedings of the 34th Annual Convention of the American Association of Equine Practitioners, 1988, pp. 487–518. 8. Setchell BP, Brooks DE. Anatomy, vasculature, innervation and fluids of the male reproductive tract. In: Knobil E, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 753–836. 9. Stickle RL, Fessler JF. Retrospective study of 350 cases of equine cryptorchidism. S Am Vet Med Assoc 1978;172:343–6. 10. Bergin WC, Gier HT, Marion GB, Coffman JR. A developmental concept of equine cryptorchism. Biol Reprod 1970;3:82–92. 11. Varner DD, Schumacher J, Blanchard TL, Johnson L. Diseases and Management of Breeding Stallions. Galeta, CA: American Veterinary Publications, 1991. 12. Lunstra DD, Ford JJ, Echternkamp SE. Puberty in beef bulls: hormone concentrations, growth, testicular development, sperm production and sexual aggressiveness in bulls of different breeds. J Anim Sci 1978;46:1054–62. 13. Johnson L, Neaves WB. Age-related changes in the Leydig cell population, seminiferous tubules, and sperm production in stallions. Biol Reprod 1981;24:703–12. 14. De Kretser DM, Kerr JB. The cytology of the testis. In: Knobil E, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 837–932. 15. Fawcett DW. Ultrastructure and function of the Sertoli cell. In: Hamilton DW, Greep RO (eds) Handbook of Physiology, vol. 5. Washington DC: American Physiology Society, 1975; pp. 21–55. 16. Setchell BP. The Mammalian Testis. Ithaca, NY: Cornell University Press, 1978.
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17. Hochereau-de Reviers M-T, Courtens J, Courot M, De Reviers M. Spermatogenesis in mammals and birds. In: Lamming GE (ed.) Marshall’s Physiology of Reproduction, vol. 2. Reproduction in the Male, 4th edn. London: Churchill Livingstone, 1990; pp. 106–82. 18. Lok D, Weenk D, De Rooij DG. Morphology, proliferation, and differentiation of undifferentiated spermatogonia in the Chinese hamster and the ram. Anat Rec 1982;203: 83–99. 19. Hochereau-de Reviers M-T, Monet-Kuntz C, Courot M. Spermatogenesis and Sertoli cell numbers and function in rams and bulls. J Reprod Fertil Suppl 1987;34:101–14. 20. Almahbobi G, Papadopoulos V, Carreau S, Silberzahn P. Age-related morphological and functional changes in the Leydig cells of the horse. Biol Reprod 1988;38:653–65. 21. Johnson L, Thompson Jr DL. Age-related and seasonal variation in the Sertoli cell population, daily sperm production and serum concentrations of follicle stimulating hormone, luteinizing hormone and testosterone in stallions. Biol Reprod 1983;29:777–89. 22. Berndtson WE, Igboeli G, Parker WG. The numbers of Sertoli cells in mature Holstein bulls and their relationship to quantitative aspects of spermatogenesis. Biol Reprod 1987;37:60–7. 23. Berndtson WE, Squires EL, Thompson Jr DL. Spermatogenesis, testicular composition and the concentration of testosterone in the equine testis as influenced by season. Theriogeriology 1983;20:449–57. 24. Johnson L. A new approach to quantification of Sertoli cells that avoids problems associated with the irregular nuclear surface. Anat Rec 1986;214:231–7. 25. Johnson L, Amann RP, Pickett BW. Scanning electron and light microscopy of the equine seminiferous tubule. Fertil Steril 1978;29:208–15. 26. Swierstra EE, Gebauer MR, Pickett BW. Reproductive physiology of the stallion. I. Spermatogenesis and testis composition. J Reprod Fertil 1974;40:113–23. 27. Fawcett DW. Observations on the organization of the interstitial tissue of the testis and on the occluding cell junctions in the seminiferous epithelium. Adv Biosci 1973;10:83–99. 28. Johnson L, Nguyen HB. Annual cycle of the Sertoli cell population in adult stallions. J Reprod Fertil 1986;76:311–16. 29. Johnson L, Tatum ME. Sequence of seasonal changes in numbers of Sertoli, Leydig, and germ cells in adult stallions. In: Proceedings of the 11th International Congress on Animal Reproduction and Artificial Insemination, 1988, p. 373. 30. Amann RP, Johnson L, Pickett, BW. Connection between the seminiferous tubules and the efferent ducts in the stallion. Am J Vet Res 1977;38:1571–9. 31. Hemeida NA, Sack WO, McEntee K. Ductuli efferentes in the epididymis of boar, goat, ram, bull, and stallion. Am J Vet Res 1978;39:1892–1900. 32. Clay CM, Squires EL, Amann RP, Pickett BW. Influences of season and artificial photoperiod on stallions: Testicular size, seminal characteristics and sexual behavior. J Anim Sci 1987;64:517–25. 33. Clay CM, Squires EL, Amann RP, Nett TM. Influences of season and artificial photoperiod on stallions: Luteinizing hormone, follicle stimulating hormone and testosterone. J Anim Sci 1988;66:1246–55. 34. Clay CM, Squires EL, Amann RP, Nett TM. Influences of season and artificial photoperiod on stallions: pituitary and testicular responses to exogenous GnRH. J Anim Sci 1988;67:763–70.
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35. Ippensen E, Kiug-Simon C, Klug E. Der Verlauf der Blutgefasse vom loden des Pferdes im Hinblick auf eine Biopsiemoglichkeit. Zuchthygiene 1972;7:35–45. 36. Nicander L. Studies on the regional histology and cytochemistry of the ductus epididymidis in stallions, rams and bulls. Acta Morphol Neerl Scand 1958;1:337–62. 37. Amann RP. Function of the epididymis in bulls and rams. J Reprod Fertil Suppl 1987;34:115–31.
38. Amann RP. Maturation of spermatozoa. In: Proceedings of the 11th International Congress on Animal Reproduction and Artificial Insemination, 1988, pp. 320–8. 39. Glover TD, Nicander L. Some aspects of structure and function in the mammalian epididymis. J Reprod Fertil Suppl 1971;13:39–50. 40. Johnson L, Amann RP, Pickett BW. Scanning electron microscopy of the epithelium and spermatozoa in the equine excurrent duct system. Am J Vet Res 1978;39:1428–34.
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Chapter 96
Physiology and Endocrinology Rupert P. Amann
The production and ejaculation of normal spermatozoa in numbers adequate for conception is imperative for reproduction. Unfortunately, one or more of the reproductive organs may not function normally, and the probability of conception and development of a normal embryo is reduced. Development of a prognosis and effective treatment for such stallions, or effective management of a normal stallion, require an understanding of normal reproductive function. The testes have a pivotal dual role: (i) production of spermatozoa and (ii) secretion of hormones essential for normal function of the epididymis and accessory sex glands as well as expression of sexual behavior. Consequently, knowledge of testicular function is crucial for people working with stallions, so they can avoid actions which interfere with normal testicular function or recognize possible origins of detected alterations in spermatozoal production. In addition, some knowledge of the role of the epididymis and accessory sex glands is desirable because their functions influence spermatozoal quality. Because of the predilection of horse owners and some veterinarians to administer hormones to normal stallions, endocrine regulation of reproductive function in adult stallions is considered in detail in this chapter to provide a basis for consideration of changes involved in puberty and to serve as a setting for Chapter 114. Finally, the process of ejaculation is briefly considered.
The testes The gross anatomy and microanatomy of the testis are considered in Chapters 95, 97 and 101. To learn about testis function, examine Figures 95.3 and 95.4 and read the associated text. These figures show how the testis is organized, and familiarity with the figures is assumed herein.
Thermoregulation of testicular temperature The scrotum serves to cover and protect the testes (see Figure 95.2), but its primary function is regulation of temperature in the testis and cauda epididymidis. This is accomplished by two basic mechanisms: vascular transfer of heat and changes in position relative to the abdominal wall. The first is a cooling mechanism whereas the latter can facilitate or minimize heat loss. A stallion’s testes must be below body temperature (Figure 96.1) for normal spermatogenesis. Based on research with other species,1–5 if intratesticular temperature is elevated to body temperature for a long interval, or to 40.5◦ C for as little as 2 hours, certain germinal cells developing within seminiferous tubules will be affected and die. Especially sensitive to heat are certain pachytene primary spermatocytes, but B-spermatogonia and spherical spermatids can also be affected. Consequently, a transitory decrease in number of spermatozoa ejaculated would be expected starting about 40 days later (Figure 96.2). With an exposure of only a few hours, the maximum depression of number of spermatozoa ejaculated should occur at about 50 days and improvement should be detected within 2 weeks thereafter. However, it would take ≥ 70 days after cessation of elevated testicular temperature before seminal characteristics should approach normal. The influence of elevated temperature on spermatozoa within the epididymis has not been delineated for stallions. In other species, however, prolonged elevation of intrascrotal temperature affects the quality and functionality of spermatozoa within the epididymis,4, 6 although conflicting data exist. A stallion might have normal fertility for 3–4 days after a severe traumatic injury to the testes, scrotal swelling or a marked increase in body temperature.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 96.1 Temperatures at different points within the scrotum and adjacent tissue as measured by thermography. The globular left testis is the major tissue mass in the lower portion of the figure and the arrows designate the approximate level of the abdominal wall. The central core of cool tissue, leading from the testis, is the pampiniform plexus. Note that the area near the cauda epididymidis was the coolest tissue (white). The scale shows the temperature of designated areas. The recording was made when air temperature was 28.5◦ C. (From Pickett et al.7 )
However, the increased temperature within the cauda epididymidis, induced by such an event, might cause a decline in seminal quality as soon as 4 days later, as evidenced by spermatozoa with de-
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tached tails.7 A severe decline in spermatozoal quantity and quality should be anticipated in 1–2 months (Figure 96.2), because of impaired spermatogenesis resulting from the temporary rise in testicular temperature. Thermal regulation of the testis and epididymis depends on the combined action of the scrotum, pampiniform plexus (see Figure 95.7), tunica dartos muscle, and external cremaster muscle. Based on data for sheep (temperature values given are for rams), blood within surface veins of the testis is cooled through evaporation of moisture from the scrotal skin.2, 3 Thus, the temperature of testicular venous blood (33◦ C) is similar to that beneath the scrotal skin. The cooler venous blood enters the pampiniform plexus, which serves as a countercurrent heat exchange area and functions similarly to the radiator of an automobile. Heat is transferred from warm arterial blood (39◦ C) to cooler venous blood so that arterial blood leaving the pampiniform plexus (34◦ C) is cooler than that in the testicular artery just above the pampiniform plexus. Thus, arterial and capillary blood perfusing the testis is several degrees cooler than the blood in visceral organs, and tissue temperature remains low (34◦ C). The extent of cooling of arterial blood passing through the pampiniform plexus is governed by temperature of the venous blood. Position of the testis relative to the abdominal wall is determined by two muscles. In cold weather, the tunica dartos muscle fibers, within the scrotal wall, contract to raise the testes close to the warm abdominal wall and heat loss is minimized by decreasing the area of the scrotal surface. In warm weather, the tunica dartos relaxes to make the scrotum more pendulous and maximize cooling of testicular venous blood by evaporation from scrotal skin. The external cremaster muscle can contract to raise the testis for a short interval.
Spermatogenesis
0 0
20 40 60 Days after onset of elevated testicular temperature
80
Figure 96.2 The pattern of changes in spermatozoal output and morphology after 48 hours of elevated testicular temperature (intrascrotal temperature averaged 37.3◦ C). Based on data for 12 stallions. (Data kindly provided by S.E. Heath.)
Spermatogenesis is the sum of cell divisions and cellular changes that result in formation of spermatozoa from spermatogonia. A more complete definition is that spermatogenesis is the lengthy, chronological process whereby a few stem spermatogonia divide by mitosis to maintain their own number and to cyclically produce differentiated spermatogonia that divide by mitosis to produce primary spermatocytes that, in turn, undergo meiosis to produce spermatids which differentiate into spermatozoa. This chapter explains spermatogenesis using available data for stallions,7–22 but some details of the process must be extrapolated from data for other species.2,23–32
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Physiology and Endocrinology Spermatozoa are produced within the seminiferous epithelium (epithelium spermatogenicum) of the convoluted seminiferous tubules (tubuli seminifer convolutus). The seminiferous epithelium of a mature stallion is composed of somatic cells, termed Sertoli cells (epitheliocytus sustentans), and different types of germinal cells (cellulae spermatogenicae), namely spermatogonia, primary spermatocytes, secondary spermatocytes, and spermatids (Figure 96.3). In a cross-section of a normal seminiferous tubule, four or five generations of developing germinal cells
EMBRYO
Primordial germ cell
BIRTH
Gonocytes
COLT
spermatogonia start to proliferate near puberty
POSTPUBERAL STALLION
Spermatocytogenesis (multiplication phase)
itted Uncommatogonia rm e p s A1 ted Commitrm gonia spe ato A1 - & A 1.4 A1.B - spermatogonia
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are arranged in well-defined cellular associations. 12, 13, 20, 21 In the stallion, each succeeding layer or generation of germinal cells within a cellular association is 12.2 days more advanced toward spermatozoa.23 The time required to produce a spermatozoon from a committed spermatogonium, or duration of spermatogenesis, is not influenced by season. The duration of spermatogenesis is about 57 days in stallions,7, 12, 23, 24 rather than 55 days as previously reported.21 Consequently, the interval between an event detrimental to spermatogenesis and a decline in seminal quality might be as long as 2 months. An interval of at least 2 months may be required for restoration of normal spermatogenesis after damage from an increase in testicular temperature or administration of drugs such as steroids. The duration of spermatogenesis (57 days) represents three phases: spermatocytogenesis (19.4 days), characterized by mitosis and differentiation of spermatogonia; meiosis (19.4 days), characterized by exchange of genetic material between homologous chromosomes in primary spermatocytes, followed by the two divisions of meiosis, producing haploid spermatids; and spermiogenesis (18.6 days), characterized by differentiation and specialization of function to result in fully differentiated spermatids, which are termed spermatozoa after release from the seminiferous epithelium.
A2 - spermatogonia A3 - spermatogonia B1 - spermatogonia
Sertoli cells
B2 - spermatogonia
As noted in Chapter 95, Sertoli cells rest on the lamina propria of the seminiferous tubule and their extensive cytoplasm extends around the germinal cells to the tubular lumen (see Figure 95.4). All details of the role of Sertoli cells in spermatogenesis are unknown, but they play a pivotal role in the process. In addition, the more Sertoli cells a testis contains, the more spermatozoa that testis can produce.12, 16, 17, 22, 26, 30 The probable functions of Sertoli cells include (i) formation of the blood–testis barrier, (ii) structural support for and nutrition of germinal cells, (iii) movement of developing germinal cells within the seminiferous epithelium, (iv) discharge of mature spermatids by a process termed spermiation, (v) phagocytosis of degenerating germinal cells and residual bodies of cytoplasm left behind as mature spermatids undergo spermiation, (vi) secretion of fluids and proteins to bathe the developing germinal cells and convey spermatozoa through the seminiferous tubules to the rete testis, and (vii) cell-to-cell communication with developing germinal cells, the underlying myoid cell layer (part of the lamina propria), and Leydig cells.
Primary spermatocytes Meiosis Secondary spermatocytes Spermatids
Spermiogenesis (differentiation phase)
Spermiation
Spermatozoa after spermiation
Figure 96.3 The pattern of spermatogenesis in stallions during development of spermatogonia to form primary spermatocytes, secondary spermatocytes, spermatids, and spermatozoa. The horizontal connections between adjacent germ cells in a cohort depict intercellular bridges that connect germ cells. A cohort is a group of two or more germ cells that are the progeny of a less developed type of germ cell. The vertical lines show the pattern of cell division and dashed lines designate that some potential germinal cells degenerate. The pattern of spermatogonial division and approximate life spans of the several types of spermatogonia are detailed in Figure 96.13. (From Pickett et al.7 )
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Figure 96.4 A scanning electron micrograph (×4500) of stallion spermatozoa just as they complete spermatogenesis and are about to be released from the seminiferous epithelium into the lumen of a convoluted seminiferous tubule. After their release by breakage of their cytoplasmic stalks (arrow), spermatozoa are washed into the straight seminiferous tubules by flowing fluid. Proximal cytoplasmic droplets (CD) are normal components of such spermatozoa. (From Johnson et al.13 )
Junctional complexes between adjacent Sertoli cells form a blood–testis barrier functionally dividing the seminiferous epithelium into basal and adluminal compartments. Spermatocytogenesis occurs in the basal compartment, where spermatogonia and preleptotene primary spermatocytes are found. In the early leptotene phase of meiosis, primary spermatocytes migrate through the blood–testis barrier, by a process analogous to moving a ship through a pair of locks, into the adluminal compartment where meiosis continues and spermiogenesis occurs. Integrity of the blood–testis barrier is maintained by forming new junctional complexes below the leptotene primary spermatocytes before dissolution of junctional complexes above the cells. The blood–testis barrier isolates spermatocytes and spermatids from the immune system of the host stallion. Because the developing immune system is not exposed to differentiated spermatocytes or spermatids, it considers them to be foreign cells and would react to antigens expressed on their surface. Therefore, except for this isolation provided by the blood–testis barrier, germinal cells would be destroyed and sterility would ensue. Damage to the blood–testis barrier can cause damage to the testis, re-
duce spermatozoal production, or induce sterility. Although rare in stallions, it is common in black mink.33 The blood–testis barrier also restricts the flux of macromolecules and other components of interstitial fluid to spermatocytes and spermatids. Sertoli cells provide structural support for developing germinal cells, as evident from their intimate contact, interdigitation of plasma membranes, and cellular specializations allowing communication. During spermiogenesis, spherical spermatids are moved, by actions of microtubular and microfibrillar components of Sertoli cells, toward the basal membrane and then back toward the luminal face; these changes occur concomitantly with changes in shape of spermatids, which are at least partially induced by Sertoli cells. As spermiation approaches, mature spermatids are pushed out into the tubular lumen, but remain attached by a thin stalk (Figure 96.4) to the major portion of the cytoplasm, contained in a residual body. The cytoplasmic residual body remains surrounded by Sertoli cells, and after rupture of the stalk, resulting in spermiation and discharge of spermatozoa, the residual bodies are phagocytized by Sertoli cells. Many germinal cells die during the process of spermatogenesis and these dead cells are rapidly phagocytized by Sertoli cells. Although degenerating cells can be seen within the seminiferous epithelium, because of their rapid phagocytosis by Sertoli cells, the number of degenerating cells seen does not accurately reflect the effect of cell death on the potential production of spermatozoa. Sertoli cells secrete a number of products into the fluid surrounding the germinal cells and hence into the luminal fluid of the seminiferous tubule. Although certain of the products are unique and not produced elsewhere in the body, others are similar to secretions of the epididymis or other organs. Lactate is secreted as an energy source for developing germinal cells. Published reviews provide an excellent overview of the biochemical nature of more complex secretions.29, 34, 35 Some serve as transport molecules to move essential metals, vitamins, or hormones from Sertoli cells to developing germinal cells, whereas others may help in the regulation of epididymal function. Some secretions of Sertoli cells, such as the protein hormone inhibin, are secreted both from the luminal face of the seminiferous tubule to enter the epididymis in rete testis fluid and also from the basal aspect of Sertoli cells to pass into the interstitial fluid of the testis and ultimately enter by the lymphatic and venous drainage. The function of Sertoli cells depends on two hormones: follicle-stimulating hormone (FSH) and testosterone. In addition, in the last several years it
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Table 96.1 A comparison between spermatogenesis in stallions and other mammals Daily sperm production
Paired testes weight (g) Human Bull Hereford Dairy Dog Stallionc Ram, Ile de France Boar Rat, Wistar Rabbit
34 650 725 34 340 500 720 3.7 6.4
Per gram parenchyma (106 /g) 4.4 10 12 12 16 21 23 24 25
Duration of cycle(days)a
Duration spermatogenesis (days)b
0.125
16.0
72
5.9 7.5 0.37 5.37 9.5 16.2 0.086 0.160
13.5 13.5 13.6 12.2 10.4 8.6 13.3 10.7
61 61 61 57 47 39 60 48
By both testes (109 )
a
Duration of one cycle of the seminiferous epithelium. Duration of spermatogenesis, assuming that it requires 4.5 cycles of the seminiferous epithelium, except for stallions for which a value of 4.7 cycles was used. c Averaged across ages and seasons, see also Figures 96.12, 96.13, and 96.15. From Pickett et al.7 b
has become evident that Sertoli cells participate in two-way communication with both germinal cells and Leydig cells. For example, both Sertoli cells and Leydig cells apparently secrete or are responsive to a variety of regulatory factors such as β-endorphin, produced by Leydig cells apparently to cause cessation of mitosis of indifferent supporting cells before puberty.36 Insulin-like growth factor (IGF), epidermal growth factor (EGF), and transforming growth factor-β (TGF-β) also may have a role in regulating testis function. Mitogenic polypeptides produced by Sertoli cells may stimulate or coordinate mitosis and meiosis of germinal cells.36, 37 Many compounds adversely affecting spermatogenesis probably act on Sertoli cells rather than directly on germinal cells. Furthermore, the number of Sertoli cells per testis and the maximum number of germinal cells per Sertoli cell are characteristic of a species,32 but the number of Sertoli cells per testis is a trait with high heritability.26, 30 In bulls, a direct relationship is found between the number of Sertoli cells in the testis and daily spermatozoal production by that testis. A treatment that decreases the number of Sertoli cells formed before puberty or that alters Sertoli cell function will probably adversely affect spermatozoal production.
Spermatogenesis: a species comparison In the adult stallion, billions of spermatozoa are produced daily in the convoluted seminiferous tub-
ules.7–9,12,15–20,22,38–40 A comparison of productivity of the stallion testis with data for other species is presented in Table 96.1. Although the two testes of an adult stallion produce about 70,000 spermatozoa each second during the breeding season, production of each individual spermatozoon requires about 57 days. When spermatozoa are liberated from seminiferous epithelium (Figure 96.4), fluid carries them from the convoluted seminiferous tubules into straight seminiferous tubules and the tubules of the rete testis where additional fluid may be added. The suspension of spermatozoa is moved rapidly through ductuli efferentes testis into the proximal epididymis. The cycle of the seminiferous epithelium From histological examination of stallion testes, researchers have determined that adjacent crosssections through the seminiferous tubules are usually different. Detailed analyses have enabled recognition of eight different cellular associations, or stages, based on four or five specific types of germinal cells grouped together.12, 13, 20, 21 The exact number of cellular associations depends on the criteria used for identification of each grouping of germinal cells.2, 12, 25, 27–29,41–43 In each stage or cellular association, the four or five types of germinal cell are associated in a specific layered pattern (Figures 96.5 and 95.4). Each layer is one generation of germinal cells, which is 12.2 days more developed than the layer below. The
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Figure 96.5 The cycle of the seminiferous epithelium in the stallion. The lamina propria is at the bottom, and the lumen of the seminiferous tubule is at the top. Each of the five upper rows represents one generation of germinal cells that are increasingly (from bottom to top) more mature. The columns represent the eight cellular associations as explained in the text. Germinal cells present in each cellular association can be discerned by reading up in each column. The complete series of cellular associations is termed the cycle of the seminiferous epithelium. In the stallion, duration of one cycle of the seminiferous epithelium is 12.2 days. Duration of each cellular association is also shown. Spermatogenesis starts when a single A1 spermatogonium divides to form a pair of A1 -spermatogonia (designated A1–2 ) this is the commitment of these germinal cells to divide further and differentiate into A2 -spermatogonia and ultimately give rise to spermatids and spermatozoa. Other germinal cells are designated: B1 -, B1 -spermatogonia; B2 -, B2 spermatogonia; Pl, preleptotene; L, leptotene; Z, zygotene; P, pachytene; and D, diplotene primary spermatocyte; 2, secondary spermatocyte; and Ss, Se, Sc, and Sm, progressively more mature spermatids. Approximately 4.7 cycles of the seminiferous epithelium pass between division of a single A1 spermatogonium into a pair of committed A1–2 -spermatogonia and release of resulting spermatozoa from the seminiferous epithelium. Thus duration of spermatogenesis is 57 days in the stallion. (From Pickett et al.7 )
youngest generation is located along the wall or lamina propria of the seminiferous tubule. Older generations are found closer to the tubular lumen. In a normal testis, germinal cells are always found in these specific cellular associations or stages. The cells present in each cellular association can be determined by reading upward in a column (Figure 96.5) from the lamina propria toward the tubular lumen (generations five to one). The width of each column in Figure 96.5 depicts the relative duration of each cellular association.
If a fixed point within a seminiferous tubule were viewed over time, germinal cells developing at that point would sequentially acquire the appearance of each of the eight cellular associations (Figures 96.6–96.9) characteristic of stallions.7, 12, 13, 20, 21 This progression through the series of cellular associations occurs repeatedly in a predictable manner. The complete series of cellular associations (Figure 96.5) is termed the cycle of the seminiferous epithelium. The interval required for one complete series of cellular associations to occur at one point within the tubule is termed the duration of the cycle of the seminiferous epithelium. Swierstra et al.21 used 12 stallions to study spermatogenesis and was able to identify eight stages in the cycle of the seminiferous epithelium. The cycle was divided into eight stages, based on the structure of the germinal cells and their relative position within the seminiferous epithelium (Figures 96.6–96.8). A current interpretation of descriptions of each stage or cellular association follows.12, 20, 21 Stage I From complete disappearance of mature spermatids lining the tubular lumen to onset of elongation of spermatid nuclei. Stage II From onset of elongation to end of elongation of spermatid nuclei. Stage III From end of elongation of spermatid nuclei to start of the first meiotic division. Stage IV From start of the first to end of the second meiotic division. Stage V From end of the second meiotic division to initial appearance of type B2 -spermatogonia. Stage VI From initial appearance of type B2 spermatogonia to when all bundles of elongated spermatids begin to migrate toward the lumen of the seminiferous tubule. Stage VII From the time all bundles of elongated spermatids have begun to migrate toward the tubular lumen until they reach the lumen and B2 spermatogonia are no longer present. Stage VIII From appearance of preleptotene primary spermatocytes and when elongated spermatids line the tubular lumen until complete disappearance of mature spermatids lining the tubular lumen. The relative frequency of occurrence of the eight stages was expressed as a percentage of the total number of tubular cross-sections evaluated for each testis (Table 96.2). Two generations of primary spermatocytes are found in stages I–III (Figure 96.5), and thus about 35% of cross-sections through seminiferous tubules should contain two generations of primary spermatocytes in a normal testis. Similarly, two generations of spermatids are found in stages V–VIII
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Figure 96.6 Photomicrograph (a) and scanning electron micrograph (b) of the same cross-section of a seminiferous tubule showing stages I and II. Both cellular associations contain two generations of primary spermatocytes (P) and one generation of spermatids. The spermatids are spherical (SS) in stage I and elongating (ES) in stage II (×510). (Modified from Johnson et al.13 )
so that about 50% of cross-sections in a normal testis should contain two generations of spermatids, and 16% should have spermatids lining the tubular lumen (stage VIII). Although most cross-sections through seminiferous tubules of a stallion testis contain a single cellular association, some cross-sections contain two or three stages of cellular associations.13, 20 These
cross-sections presumably represent locations within a seminiferous tubule where different cellular associations adjoin.23, 29 Knowledge of time required to produce a spermatozoon is essential for understanding the time course of events after drug injection or trauma to the testis. Swierstra et al.20, 21 measured duration of the cycle of the seminiferous epithelium using six stallions.
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Figure 96.7 Photomicrograph (a) and scanning electron micrograph (b) of the same cross-section of a seminiferous tubule showing stages Ill, IV, and V. Stage Ill is characterized by two generations of primary spermatocytes (P) and deeply embedded bundles of elongated spermatid nuclei (ES). In stage IV, metaphase figures (M) and secondary spermatocytes (2S) are seen. Stage V contains newly formed spermatids with spherical nuclei (SS) and older spermatids with elongated nuclei (×510). (Modified from Johnson et al.13 )
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Figure 96.8 Photomicrograph (a) and scanning electron micrograph (b) of the same cross-section of a seminiferous tubule showing stages VI and VII. Both stages are characterized by elongated spermatid nuclei (ES), spherical spermatid nuclei (SS) and one generation of primary spermatocytes (P). In stage VI, bundles of elongated spermatid nuclei are deeply embedded, while in stage VII the elongated spermatids have begun to migrate toward the lumen (×510). (Modified from Johnson et al.13 )
This was accomplished by injecting a radioisotope, [3 H]thymidine, into both testicular arteries of each stallion. The compound [3 H]thymidine was taken up by germinal cells that were synthesizing DNA at the time of injection, and such cells or their progeny were identified on the basis of their radioactivity. The 12 testes from these stallions were removed 4.5 hours to 35 days after injection of [3 H]thymidine. Radioactive germinal cells were detected by coating the histological sections with a photographic emulsion. When the film was developed, black ‘grains’ were evident over each radioactive germinal cell in a histological section.
Table 96.2 Relative frequency and mean duration of stages of the cycle of the seminiferous epitheliuma Stage
Figure 96.9 Scanning electron micrograph of the seminiferous epithelium in late stage VIII, showing the entire length of the elongated spermatids extending into the tubule lumen. A Sertoli cell (S) and primary spermatocyte (P) are seen. The basement membrane is visible at the bottom of the picture (×970). (From Johnson et al.13 )
I II III IV V VI VII VIII a
Relative frequency (%)
Duration (days)
16.9 14.9 3.2 15.8 7.4 13.5 12.6 15.7
2.1 1.8 0.4 1.9 0.9 1.6 1.5 1.9
Mean for 18 stallions. Adapted from Swierstra EE et al.20
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Lumen
Sperm
2
3
Sm
Adluminal compartment
1
4
Sc
Se
1
Ss
Z
L
B1
A3
2
2°
2°
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D
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Blood-testis barrier Basal
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A I
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VI VII VIII
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I
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VI VII VIII
Cycle 2
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I
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IV
Cycle 3
VI VII VIII
32
I
II
IV
Cycle 4
VI VII VIII
45
I
II
IV
VI VII VIII
Cycle 5
57
Figure 96.10 Spermatogenesis in the stallion. Four or five generations (cohorts) of germinal cells develop simultaneously and synchronously at any point in a seminiferous tubule. Spermatogenesis is a continuous process and involves an infinite number of subtle changes at a given point in the tubule. These changes in cellular association have been classified as eight stages; some of them are depicted and demarcated by light vertical lines. The complete sequence of stages occurs at a given point in a tubule over an interval (12.2 days) termed the duration of the cycle of the seminiferous epithelium. Because development of generations is synchronized, the same stages recur every 12.2 days in stallions. The figure depicts (from left to right) changes over time, at a given point, in a seminiferous tubule during 57 days, or 4.7 cycles of the seminiferous epithelium, required for the complete process of spermatogenesis. Cellular associations are designated as stages I–VIII along the x-axis and their width is proportional to their duration. The four or five generations of germinal cells in any stage are designated along the y-axis, and designated: A1 -, A2 -, A3 -, B1 -, and B2 -spermatogonia; Pl, preleptotene; L, leptotene; Z, zygotene; P, pachytene; and D, diplotene primary spermatocyte; 2◦ , secondary spermatocyte; and Ss, Se, Sc, and Sm, progressively more mature spermatids. Development of one A1 -spermatogonium through spermatogenesis, to form a spermatozoon can be followed by the dark path. Near the beginning of the leptotene phase of development, new junctional complexes between adjacent Sertoli cells form below these germinal cells and the old junctions disappear. In this way, integrity of the blood–testis barrier (stippled band) is maintained. In stallions, one, but occasionally two or three, stages (usually sequential ones) are typically found in one cross-section through a seminiferous tubule. Details of spermatogonial renewal have been simplified (see Figure 96.14). (From Pickett et al.7 )
At 4.5 hours after injection of [3 H]thymidine, young (leptotene) primary spermatocytes in stage I were the most mature radioactive germinal cells. For horses castrated at longer intervals after injection of the radioisotope, progressively more advanced germinal cells were labeled. By analyzing these data, Swierstra et al.20, 21 concluded that duration of one cycle of seminiferous epithelium was 12.2 days. Thus, at any particular point within a seminiferous tubule the same cellular association or stage is repeated every 12.2 days. Cells in stage I, which were young (leptotene) primary spermatocytes in generation 3 (Figure 96.5), will be old (pachytene) primary spermatocytes 12.2 days later (in generation 2) and spherical spermatids after another 12.2 days in generation 1 (Figure 96.10). Using information on relative frequency for each stage and data for duration of the cycle of the seminiferous epithelium, absolute duration of each stage was calculated (Table 96.2). The life spans of primary spermatocytes, secondary sperma-
tocytes, and spermatids are 18.7, 0.7, and 18.6 days, respectively.
Duration of spermatogenesis At a given point within a seminiferous tubule, A1 spermatogonia periodically produce committed A1–2 spermatogonia (Figures 96.3 and 96.5); this occurs only at an interval equal to the duration of one cycle of the seminiferous epithelium.2, 7, 12, 23, 28, 29, 43 Similarly, at some point in the cycle, groups of mature spermatids, originating from spermatogonia committed to differentiate approximately 4.7 cycles earlier, are released from the seminiferous epithelium (Figure 96.10). Thus, because of the division of A1 spermatogonia every 12.2 days,20 a new cohort (family or chain) of germinal cells begins to develop every 12.2 days, and all cells in this cohort develop synchronously (Figure 96.10). Because total duration of spermatogenesis apparently is 4.7 cycles of the
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seminiferous epithelium (12.2 days), spermatogenesis requires about 57 days in the stallion. Because A1 -spermatogonia at different sites in the testis develop asynchronously, hundreds of spermatogonia committed to form spermatozoa are produced every second. Therefore, spermatozoa are released continuously from seminiferous tubules. Daily spermatozoal production Daily spermatozoal production (DSP) is the number of spermatozoa produced per day by a testis or the two testes of a stallion.23 Efficiency of spermatozoal production is the number of spermatozoa produced per gram of testicular parenchyma. During the breeding season, efficiency of spermatozoal production is reasonably similar for normal stallions, although testicular size may differ greatly among stallions.8, 9, 11, 12, 16, 17, 22 Consequently, >spermatozoal production can be estimated with fair accuracy by measuring testis size. Because testis size increases as a stallion grows from 2 or 3 year old to a sexually mature stallion, DSP also increases12, 17 (Figure 96.11). Daily sperm production also is affected by season (Chapter 133), as is especially evident for 6- to 20-year-old stallions (Figure 96.11). The magnitude of this decline during the non-breeding season averages 50% in stallions 6–20 years old (6.40 vs. 3.19 billion spermatozoa per day). Data for groups of similar adult stallions slaughtered in each month of the year are shown in Figure 96.12. Daily sperm production was low from September through February but increased by March.12, 16 Maximum DSP occurred
400 6
Breeding Non-breeding
300
5 4
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Figure 96.12 Daily spermatozoal production (DSP) of stallions in each month of the year. Data for 13–17 stallions, 4–20 years old, slaughtered in each month. Daily sperm production was estimated by morphometric analysis of spermatids with a spherical nucleus (similar to columns designated Ss in Figure 96.10). (From Pickett et al.7 )
in May and June, whereas values for July and August were markedly lower. However, DSP is not influenced by frequency of use for breeding. Efficiency of spermatozoal production in a stallion during the breeding season averages 19 million spermatozoa per day per gram of testicular parenchyma,8, 9, 11, 12,15–17,26, 40 and declines to about 15 million per day per gram in the non-breeding season. It is evident (Figure 96.11) that for stallions 6–12 and especially 13–20 years old, the decline in DSP during the non-breeding season appears to be greater than that for testicular weight (49 vs. 29% for 13to 20-year-old stallions).17 Differences in efficiency of spermatozoal production between breeding and nonbreeding season for stallions 2–3, 4–5, 6–12, and 13–20 years old were –21%, –12%, –19%, and –34%, respectively.17 Moreover, for stallions 13–20 years old, the tunica albuginea represents a larger proportion of testicular weight than in stallions 4–8 or 2–3 years old (14%, 9%, and 10%, respectively).15 The efficiency of spermatozoal production in stallions is greater than in bulls, but less than in rams or boars (Table 96.1). However, because of difference in testicular size, DSP is similar for bulls and stallions.
0
Age (years)
Figure 96.11 Effects of age and season on testes weight and daily spermatozoal production (DSP) of stallions. Stallions were slaughtered during June and July (breeding season) and December and January (non-breeding season). Means for 7–15 stallions in each group. For testicular weight or DSP, means (pooled across season) were significantly different for stallions <2, 2–3, and 4–20 years old. Effects of season also were significant as were interactions of age and season. (From Pickett et al.7 )
Germinal cell degeneration and renewal of spermatogonia In stallions, spermatozoa are produced continuously after puberty. This is accomplished by continuous production of differentiated spermatogonia that produce primary spermatocytes, which ultimately give rise to spermatids (Figures 96.3 and 96.10). At the same time, uncommitted spermatogonia must be
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Stage 1
A3 A3
Uncommited Undiffer. A-paired A-aligned
2 3
B1 B1
A1.2
4 5
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7 8 1
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Differ.
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A3
2 3
A1
Degen. in NBS
B1
B1
4 5
B2 B2
B2 B2
6 Degen. in BS
7 8
Pl
Pl
Figure 96.13 A possible pattern for the process of spermatogonial renewal (formation of two new isolated and uncommitted A1 -spermatogonia) and commitment of A1 -spermatogonia toward differentiation by forming a pair of undifferentiated A1–2 -spermatogonia that divide to form a chain of committed, but undifferentiated, A1–4 - and then A1–8 -spermatogonia. Formation of a pair of committed A1–2 -spermatogonia is considered to be the onset of spermatogenesis (see Figure 96.3). Commitment of another A1 -spermatogonium to form a pair of A1–2 -spermatogonia occurs only in stage III of the cycle of the seminiferous epithelium, once every 12.2 days at a given point in a seminiferous tubule. In theory, each differentiated A2 -spermatogonium could give rise to 64 spermatozoa. Much less than this number is produced because of degeneration of germinal cells. Although degeneration of potential spermatozoa can occur at any point in spermatogenesis, degeneration during the breeding season involves mostly B2 -spermatogonia, whereas in the non-breeding season it involves A1–8 - and A3 spermatogonia. Pl, Preleptotene; NBS; BS. (From Pickett et al.7 )
replaced so that spermatogenesis will not stop because the seminiferous epithelium becomes depleted of spermatogonia.2, 8, 12, 25,27–29,31, 43 Details of the process in stallions are now being unraveled10–12,16 (L. Johnson, personal communication) (Figure 96.13). Present interpretations of data from ongoing research with stallions, based on personal communications with L. Johnson and published data,18 and data for other species,29, 31 are as follows:
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1. An isolated, uncommitted A1 -spermatogonium divides to form either two new uncommitted A1 spermatogonia, which then are moved apart and serve as new ‘stem cells’, or two cells that remain together joined by an intercellular bridge as a pair of A1 -spermatogonia (designated A1–2 spermatogonia in Figure 96.13) that continue to divide by mitosis to form a chain of eight aligned A1 spermatogonia (designated A1–8 -spermatogonia). 2. Some of the A1 -spermatogonia degenerate, especially in the non-breeding season, but in the breeding season most A1–8 -spermatogonia divide to form A2 -spermatogonia that are termed ‘differentiated’ cells because they are functionally different from A1 -spermatogonia. 3. A2 -spermatogonia divide by mitosis to form further differentiated A3 -spermatogonia, which in turn divide into morphologically different B1 spermatogonia, although in the non-breeding season some of A3 -spermatogonia degenerate. 4. B1 -spermatogonia divide by mitosis to form B2 spermatogonia, which in turn divide into preleptotene primary spermatocytes, although in the breeding season some B2 -spermatogonia degenerate. 5. Soon after their formation, preleptotene primary spermatocytes enter the long prophase of the first division of meiosis, ultimately to give rise to secondary spermatocytes, then spermatids. Another population of A-spermatogonia, few in number, termed reserve spermatogonia, exist, but are not shown in Figure 96.13. Their lifespan is unknown, but is probably > 60 days. Normally, they do not participate in spermatogenesis but serve as a source of germinal cells to repopulate the testis in seasonal breeders in which the testes ‘shut down’ completely except in the breeding season (such as mule deer) or after an event that causes loss of most or all of the regular germinal cells (exposure to a toxin). Not all spermatozoa, which potentially should be produced by the seminiferous epithelium actually are formed and released from seminiferous epithelium (Figures 96.11 and 96.13). Considerable degeneration of germinal cells occurs in normal stallions even during the breeding season. Because testicular weight decreases in the non-breeding season (Figure 96.11), researchers find it most informative to consider degeneration of germinal cells in terms of number of spermatozoa that theoretically should be produced per gram of testis. During the breeding season an excessive number of B2 -spermatogonia are produced and 32% of the potential number of ‘young’ primary spermatocytes are not formed (Figure 96.14). Presumably, this is because each Sertoli cell can accommodate only about 9 or 10 spermatids
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30
years old (Figure 96.11). In addition to environmental factors such as day length or temperature, drugs or unknown factors can lead to increased degeneration of germ cells.2, 12, 29, 44, 45
*a –32%
Potential DSP/g testis
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* b
20
b
Leydig cells
* b * b
a
a
a
–23% b b
10
B 0
Y
O S
Breeding
E
B
Y
O S
E
Nonbreeding
Figure 96.14 Effects of season on degeneration of potential spermatozoa. Potential daily spermatozoal production (DSP) per gram of testicular parenchyma was calculated assuming all cells of a given type resulted in the theoretical number of spermatozoa without degeneration. Values for potential DSP/g testis were calculated from morphometric determinations of numbers of B2 -spermatogonia (B); preleptotene, leptotene, plus zygotene spermatocytes (Y); pachytene plus diplotene spermatocytes (O); spherical spermatids (S); and elongated spermatids (E) in testes of 28 stallions slaughtered in the breeding season (June and July) and 28 stallions slaughtered in the non-breeding season (December and January). (From Pickett et al.7 )
of a given generation and proportionately fewer primary spermatocytes (about 2.5 of a given generation).10, 17, 22 Virtually all primary spermatocytes formed ultimately give rise to four spermatozoa.11 However, during the non-breeding season, 35% fewer A-spermatogonia per gram of testis are found and 40% fewer B2 -spermatogonia.11, 16, 18 Nearly all B2 -spermatogonia give rise to two ‘young’ primary spermatocytes, so that the number of these cells per gram of testis is not different in non-breeding and breeding seasons (Figure 96.14). However, in the nonbreeding season, about 23% of the potential number of spherical spermatids are not formed; germinal cells apparently degenerate during aberrant meiotic divisions or as newly formed spermatids. Presumably, the environment provided by Sertoli cells is deficient in some way, as evidenced by the reduced number of germinal cells per Sertoli cell.11, 12, 30 As was true for the breeding season, in the non-breeding season virtually all spherical spermatids that develop for 2 or 3 days are transformed into spermatozoa. The result is that in the non-breeding season, only 75% as many spermatozoa are formed per gram of testis as in the breeding season. However, because of differences in testis weight, DSP per stallion is reduced by 20% for stallions 4–5 years old and by 50% for stallions 6–20
Within the interstitial tissue, Leydig cells (endocrinocytus interstitialis) are in close proximity to blood vessels and lymphatic channels, as well as the basal lamina of the seminiferous tubules. The primary role of Leydig cells is secretion of steroid hormones, which help to regulate function of the seminiferous epithelium, the hypothalamic–hypophyseal axis, and the accessory sex glands. Trivial names used for steroid hormones are testosterone (17β-hydroxy-androst-4-en-3-one), androstenedione (3,17-diketo4 -androstene), androstenediol (3β 2 17β-dihydroxy5 -androstene), dihydrotestosterone (17β-hydroxy-5α-androstan-3-one), 3α-androstanediol (5α-androstan-3α,17β-diol), 3βandrostanediol (5α-androstane-3β,17β-diol), progesterone (pregn-4-ene3,20-dione), estrone [3βhydroxyestra-1,3,5(10)-triene-17-one], and estradiol [estra-1,3,5(10)-triene-3β,17β-diol]. Because Leydig cells are the site of production for most of these hormones, interstitial fluid contains a much higher concentration of testosterone and other secretory products of Leydig cells than does peripheral blood (e.g., serum from blood drawn from the jugular vein). Concentration of testosterone averaged 416 ng/g and 640 ng/g parenchyma in two studies,8, 46 although these values might be artifactually high because of continued production of testosterone by the tissue after cessation of blood flow from the testis. Leydig cells constantly secrete basal amounts of testosterone and several other hormones. However, periodically Leydig cells are stimulated to increase their production of testosterone. Thereafter, twofold to fourfold elevations of testosterone concentration, each lasting 2–4 hours, are found in peripheral blood.47 Three to eight episodic bursts of testosterone production occur each day in most, but not all, stallions. Consequently, if a single blood sample from a stallion is analyzed for concentration of testosterone, an unusually high value may be obtained that is not typical of peripheral concentrations in blood of that horse. During episodic production of testosterone, concentrations of testosterone within the testis are probably elevated to > 10 times basal concentration. Consequently, seminiferous tubules are continuously exposed to a high concentration of testosterone. This high concentration of testosterone around seminiferous tubules is probably essential for normal spermatogenesis, as has been shown with other
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Physiology and Endocrinology Table 96.3 Hormonal concentrations in testicular vein and jugular vein blood
17β-hydroxy-androgen (ng/mL) Total non-conjugated estrogen (ng/mL)
61.5 25.4
CH3 C O
CH3 C O O
HO
Testis vein
Jugular vein
∆5-Pregnenolone
Progestrone
CH3 C O OH
1.3 0.058
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CH3 C O OH
HO
O
17α-Hydroxypregnenolone
17α-Hydroxyprogesterone
From Amann and Ganjam.49
species.2, 3, 29, 35, 41, 48 However, minimum concentration of testosterone necessary within the seminiferous epithelium for normal spermatogenesis in stallions is unknown. Researchers do not know whether normal intratesticular concentrations of testosterone could be maintained by injecting stallions with massive doses of testosterone. This is possible with rats48 but not humans (E. Steinberger, personal communication). The concentration of total 17β-hydroxy-androgen in blood leaving the testis through the testicular vein is about 45 times the concentration in blood taken concurrently from the jugular vein49 (Table 96.3). During intervals when Leydig cells are under maximal stimulation of endogenous luteinizing hormone (LH), concentrations of testosterone in testicular vein blood can exceed 500 ng/mL, or approximately 100fold typical concentrations of testosterone in jugular vein blood.49, 50 In stallions, Leydig cells secrete greater amounts of estrogens than testosterone, although testosterone is the steroid of greatest physiological importance. In addition to estrone and estradiol, the stallion testis also secretes estriol and two unusual compounds, equilin and equilenin51–56 (Figure 96.15). Although testicular vein blood contains extraordinarily high concentrations of free estrone and estradiol, most estrogen secreted by the stallion testis is conjugated with a sulfate (or glucuronide) side chain. This makes the compound more water soluble and restricts bioactivity. The biological role of the conjugated estrogens produced by the equine testis remains obscure. Enzymes involved in production of steroid hormones in Leydig cells are localized on the smooth endoplasmic reticulum and mitochondria.57, 58 No evidence exists to support the notion that other elements of interstitial tissue, or seminiferous tubules, produce steroid hormones. The first step in steroidogenesis is formation of cholesterol, either by de novo synthesis or breakdown from low-density lipoproteins of blood origin. Based on data for rodent and human testes,59 researchers logically assume that in a stallion testis secreting testosterone at a minimal level most cholesterol is derived from intra-Leydig cell sources (cholesterol droplets). However, during stim-
HO
HO
O
Dehydroepiandrosterone
∆4-Androstenedione
HO 5-Androstenediol
H
Estrone
OH
OH
OH
∆
O
O
O
HO
O
Testostrone
H
Estradiol
OH
O
O
HO H
Dihydrotestosterone
Equilin
OH O
HO
H
5α-Androstane- 3 α / β, 17 β–Diol
HO
Equilenin
Figure 96.15 The pathway for steroid biosynthesis in the stallion testis. Pregnenolone, formed from cholesterol, is converted by one of several possible pathways to androstenediol or androstenedione, which are the two immediate precursors of testosterone; the exact pathway is unknown for stallions. A portion of the androstenedione produced by Leydig cells is converted to estrone, rather than testosterone, and secreted as estrone sulfate. Testosterone in turn can be converted to dihydrotestosterone or 3α-androstanediol (androgens active in many tissues) or to estradiol (generally considered as a female sex hormone although it also may be important in the male) and secreted as estradiol sulfate. Note the structural similarity of many of these hormones. Synthetic compounds having a similar structure may affect a stallion by blocking or mimicking an action of a natural hormone. Two estrogens unique to horses, equilin and equilenin, also are depicted. (Modified from Pickett et al.7 )
ulation by LH, or exogenous human chorionic gonadotropin (hCG), most of the cholesterol is probably derived from blood-borne low-density lipoprotein. The steroidogenic pathway begins with conversion of cholesterol to pregnenolone, which occurs within the mitochondria. Pregnenolone is transported to the smooth endoplasmic reticulum where other enzymes involved in the pathway (Figure 96.15) are located. Detailed consideration of the enzymes and thermodynamics involved in steroidogenesis are outside the scope of this chapter, but excellent reviews are available.57, 58
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Figure 96.15 shows that two general pathways exist from pregnenolone to testosterone: (i) the socalled 5 pathway through dehydroepiandrosterone and 5 -androstenediol and (ii) the 4 pathway via progesterone and 4 -androstenedione. Which pathway, or combination of pathways, predominates in equine Leydig cells is unknown. However, in rat Leydig cells the 4 pathway predominates, whereas in rabbit Leydig cells the 5 pathway is used almost exclusively.60 Because steroidogenic enzymes are localized on the smooth endoplasmic reticulum, a direct and linear relationship exists between total surface area of smooth endoplasmic reticulum associated with Leydig cells of a testis and its ability to secrete testosterone. The relationship even holds true across species.60 Luteinizing hormone or hCG stimulates steroidogenesis by Leydig cells. This stimulatory effect probably is a consequence of LH stimulating transport of cholesterol from intracellular stores to the outer mitochondrial membrane, and also within the mitochondria, to provide cholesterol to the side-chain cleavage enzyme (a mitochondrial cytochrome P450 enzyme), which converts cholesterol to pregnenolone in a three-step sequence of reactions.58 Thus conversion of cholesterol to pregnenolone is the ratelimiting step in production of testosterone. With gonadotropin stimulation, production of pregnenolone increases and this enables increased production of testosterone and estrogens. Leydig cells contain specific receptors for LH integral to their plasma membrane. Binding of LH to the membrane receptor activates a guanosine triphosphate (GTP)-binding protein which, in turn, stimulates adenylate cyclase and increases local concentration of cyclic adenosine monophosphate (cAMP).58 Cyclic AMP, in turn, phosphorylates specific proteins by a post-translational modification; in this case a cholesterol-ester hydrolase becomes more active and releases increased amounts of free cholesterol for transport by microfilaments to mitochondria. Because the process does not involve synthesis of a new protein and enzymes involved in conversion of pregnenolone to testosterone are working far below their maximum capacity, Leydig cells rapidly increase production of testosterone in response to LH stimulation. Other proteins probably exist for which function is modified by LH stimulation to increase the transport flux of cholesterol to the inner mitochondrial membrane where the side-chain cleavage enzyme is located.
Epididymis Most aspects of epididymal function have not been studied in the stallion. However, from available
data,14, 61 and those from other species,62–69 the function of the excurrent duct can be deduced. After leaving the testis, spermatozoa enter the ductuli efferentes testis and then are transported, successively, through several zones of the epididymis. These include the initial segment of the epididymal duct (ductus epididymidis), two or three spermatozoal maturation zones in the caput and corpus epididymidis, and a zone where fertile spermatozoa are stored in the cauda epididymidis until ejaculated (Figure 96.16). Most of the fluid, protein, and other material entering from the testis is resorbed in the ductuli efferentes and proximal caput epididymidis, to be replaced by secretions of the epididymal epithelium. However, spermatozoa are not normally removed in the epididymis. The composition of luminal fluid surrounding spermatozoa is different in successive functional zones and regional differences in mechanisms regulating epithelial function exist. Spermatozoa leaving the testis are infertile, whereas spermatozoa recovered from the cauda epididymidis are fertile.62, 63, 65, 66 The process by which spermatozoa develop capacity for fertilization is termed spermatozoal maturation, but this is only one aspect of a continuous spectrum of changes in these cells from when they are formed until they fertilize an egg63 (see Chapter 97). Spermatozoal maturation depends on sequential exposure of spermatozoa to epididymal fluids of differing composition. Enzymes and other proteins in the fluid modify the plasma membrane, and certain other components, of spermatozoa (Figure 96.16). Availability of testosterone to the epithelium, especially in the caput and corpus epididymidis,62, 65, 66 is essential for secretion of certain proteins by the epididymal epithelium, although other secretions are produced without androgenic stimulation. Evidence for spermatozoal maturation includes acquisition of fertilizing capacity and progressive motility as well as changes in spermatozoal structure, characteristics of the plasma membrane of spermatozoa, and spermatozoal metabolism.14, 61, 63,65–68 However, simple retention of spermatozoa within a given segment of the ductus epididymidis is insufficient to induce spermatozoal maturation.65, 66, 68 Equine spermatozoa from the caput or proximal corpus epididymidis are immotile when released into a physiological salt solution (Table 96.4). However, the percentage of motile spermatozoa in samples removed from the cauda epididymidis and diluted in buffer is similar to that in ejaculates collected from the same stallions. Thus, as measured by progressive motility, maturation of stallion spermatozoa is completed before spermatozoa enter the cauda epididymidis.14 As accessed by ability of their plasma membrane to exclude the pink dye eosin,
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Physiology and Endocrinology Testis
Rete testis
Efferent ducts
Initial segment
Proteins Inositol
Proteins Growth factors Steroids
High Na:K ratio Absorb much of protein & fluid; Unique protein secrete protein; alter ionic balance spectrum; low Na:K ratio in lumen
Proteins
Cauda
Proteins
?
Proteins
?
?
ABP ASF
FMP Oxytocin Inhibin
? ?
?
CORPUS
Corpus epididymidis
Caput epididymidis
895
LIPID GPC CARNITINE
?
CAPUT CAUDA ED EPIDIDYMAL REGIONS
Immotile, infertile sperm
Concentrate suspension and initiate maturation of sperm Functions independent of androgens
Functions dependent of DHT 1 1 2 3 4 5
1. Changes in exposed CHOs 2. Changes in membrane lipids 3. Decrease in intracellular pH 4. Translocation of cytoplasmic droplet 5. Increased disulfide bonding 6. Increased permeability of plasma membrane 7. Decreased resistance to cold shock 8. Secretion of forward motility protein 9. Express motility 10. Secretion of acrosome stabilizing factor 11. Capable of binding to oocytes 12. Express fertilizing ability 13. Improved pregnancy ensuance
Major maturational changes of sperm
Store fertile sperm
Functions dependent on unknown regulators
6 7 8 9 10 11 12 13
STALLION EVENTS IN EPIDIDYMIS AND CHANGES IN SPERM
14 Motile, fertile sperm
Figure 96.16 A diagrammatic representation of the regional compartmentalization of epididymal function and changes in spermatozoa, associated with maturation, as they are transported through the epididymis. In the inset, the efferent ducts are designated as ED. Molecules present in the luminal fluid include: ABP, androgen binding protein; ASF, acrosome stabilizing factor; DHT, dihydrotestosterone; FMP, forward motility protein; and GPC, glycerylphosphorylcholine. Deduced from the literature, primarily for bulls, rams, and rats.
spermatozoa from all regions of the epididymis are resistant to ‘cold shock’, induced by rapid cooling to 0◦ C. Ejaculated spermatozoa, however, are altered by this treatment (Table 96.4). Based on these and other observations, Johnson et al.14 concluded that maturation of spermatozoa in the stallion was not completed until spermatozoa left the corpus epididymidis. This is consistent with data for other species,62,65–67 although changes in spermatozoa that result in a greater percentage of surviving embryos occur in the cauda epididymidis, at least in rams.62 Available laboratory data and limited data on fertility of stallion spermatozoa from the cauda epididymidis lead to the conclusion that spermatozoa from the cauda epididymidis of a valuable stallion could be used to inseminate mares, if such spermatozoa were recovered within a few hours after death or castration.70 Although spermatozoa are found throughout the epididymis,21, 23, 38, 69, 71 the cauda epididymidis and deferent duct (including the ampulla) are the major spermatozoal storage areas (Table 96.5). The two caudae epididymidum of a typical adult stallion
(5–16 years old) contain about 54 billion spermatozoa (Table 96.5), or 61% of the total number found within the excurrent duct system.38 In stallions, the caudae epididymidum contain more spermatozoa than the Table 96.4 Change in spermatozoa during passage through the epididymis Unstained spermatozoa (%) Spermatozoa from
Progressive motility (%)
Not cooled
Cooled to 0◦ C in ≤2 min
Central caput Proximal corpus Distal corpus Central cauda Ejaculated
0 0 11a 56b 64
96 97 93 90 68
96 95 93 87 27c
a
Spermatozoa from two of five stallions were progressively motile. The percentage of motile spermatozoa was similar (P > 0.05) to that in ejaculated semen. c The percentage of unstained spermatozoa was reduced (P < 0.05) by cold shock. Adapted from Johnson et al.13 b
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Table 96.5 Spermatozoal reserves in the adult stalliona Segment of excurrent duct
Spermatozoa in both sides (109 )
Relative distribution (%)
12 17 54 4 2
13 19 61 2 2
Caput epididymidis Corpus epididymidis Cauda epididymidis Deferent duct Ampulla of deferent duct a
Data for stallions 5–16 years old. Adapted from Amann et al.38
caudae from a bull, but a much lower number than the caudae from a ram or boar. Sufficient numbers of spermatozoa are present within the caudae epididymidum in the horse for several ejaculates.23, 38 Testis size, daily spermatozoal production, and number of spermatozoa stored within the epididymis are influenced by age. Spermatozoa do not swim through the epididymis and deferent duct. Movement of spermatozoa through the epididymal duct is primarily by continuous peristaltic contractions of smooth muscle in the wall of the duct within the caput and corpus epididymidis. In the cauda, the epididymal duct normally is inactive, except when smooth muscle is stimulated to contract. Consequently, time required for movement of spermatozoa through the caput and corpus epididymidis is not altered by ejaculation and averages about 4.1 days in stallions.21, 23, 38, 71 Fertility of spermatozoa is usually not depressed in males ejaculating frequently, because rate of spermatozoal transport through the caput and corpus epididymidis is not influenced by ejaculation; decreased fertility could result if number of spermatozoa ejaculated were less than the number required for maximum reproductive efficiency. Extensive data for bulls ejaculating daily, or at a similar high frequency, show that spermatozoal fertility is equivalent, if not slightly superior, to that for bulls ejaculating once a week72–74 (J.O. Almquist, personal communication). When seven successive ejaculates were collected from beef bulls and the spermatozoa was used to artificially inseminate cattle, fertility did not differ among the seven successive ejaculates.75 Fertility of stallion spermatozoa used for artificial insemination should be similar whether stallions are collected daily, every other day, or every 4 days. The interval that spermatozoa spend in the cauda epididymidis is influenced by ejaculation.23, 38 The number of spermatozoa in the cauda epididymidis is maximal in sexually rested stallions and is reduced in males ejaculating daily or every other day.38 Because fewer spermatozoa are present in the cauda
epididymidis of a stallion ejaculating regularly than in an inactive male, transit time for spermatozoa through the cauda epididymidis of a sexually active stallion is reduced by 2 or 3 days from the 10 days characteristic of a sexually rested stallion.21, 38, 71 Although the time required for movement of spermatozoa through the caput and corpus epididymidis is reasonably similar among species,23 the interval that spermatozoa spend in the cauda epididymidis differs greatly, and ranges from < 4 days in humans and some beef bulls to > 12 days in rams. Spermatozoa are produced continuously, regardless of ejaculation frequency. Because spermatozoa enter the epididymis at a constant rate, they must also leave the excurrent duct system at a relatively constant rate, although this rate is altered by ejaculation. Based on research with several species,76, 77 all spermatozoa that enter the excurrent duct system of a stallion likely leave through the urethra. Resorption of spermatozoa probably does not occur within the excurrent duct system.23, 76, 77 In bulls and rams, spermatozoa that are not ejaculated at copulation or voided by masturbation are eliminated periodically during urination.77 Probably in a normal, sexually inactive stallion, spermatozoa intermittently pass from the deferent duct into the pelvic urethra (pars pelvina) and are voided during urination. The direct cause for or interval between such emissions is unknown. Certain stallions accumulate an abnormally large number of spermatozoa in the epididymis and perhaps to some extent in the deferent duct, including the ampulla. In such a horse, spontaneous emissions probably do not occur and spermatozoa accumulate in the epididymis until the limit of distensibility of the epididymal duct is reached. As a consequence of this accumulation, storage interval of spermatozoa in the cauda epididymidis of such a stallion is much longer than 7–10 days, and spermatozoa may undergo marked alterations. Much lower percentages of spermatozoa are structurally normal or motile in the first several ejaculates collected from a stallion accumulating spermatozoa than in a normal horse because of the prolonged storage interval.
Accessory sex glands Collectively, the prostate gland, bulbourethral glands, and vesicular glands are termed the accessory sex glands (glandulae genitales accessoriae) (see Figures 95.1 and 95.12). These glands contribute most of the fluid to the ejaculate. Spermatozoa from the cauda epididymidis and deferent duct are immotile until mixed with accessory sex gland fluids at ejaculation (or mixed with a buffer by human intervention). Exact factors causing initiation of motility are unknown, but intracellular pH may be
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Physiology and Endocrinology involved. Although certain proteins in accessory sex gland fluids are bound by spermatozoa, exposure to these fluids is not essential for normal fertility of the spermatozoa; cells from the cauda epididymidis are of normal fertility. Indeed, increasing evidence is accumulating that for some stallions a component of seminal plasma may be responsible for reduced survival of sperm during storage or low fertility. Normal function of all accessory sex glands depends on availability of testosterone in peripheral blood.78 The secretion of the prostate gland is thin and watery.79 This secretion probably helps cleanse the urethra during ejaculation and also constitutes a major portion of seminal plasma, especially if a second ejaculation occurs 1–3 hours after an earlier ejaculation. Depending on season and individual stallion, fluid secreted by the vesicular glands may (or may not) contribute a major portion of seminal plasma in an ejaculate. The gelatinous material often found in seminal plasma, especially in April to July is secreted by these glands.79 This seasonal change in secretion of gel by the vesicular glands, as well as the stallion-to-stallion difference, may reflect differences in concentration of testosterone in blood. Two bulbourethral glands are positioned on either side of the pelvic urethra near the ischial arch. Their secretion contributes to the seminal plasma but probably only a minor portion in terms of volume.
Endocrine hormones Function of reproductive organs is controlled by the neuroendocrine system. The neuroendocrine system includes specialized groups of nerve cell bodies and endocrine tissues that secrete chemical messengers termed hormones (Chapter 99), which are carried through the blood from one organ to control the function of another organ. The autonomic nervous system has a role in controlling function of the reproductive organs; transport of spermatozoa from the testes through the epididymides and deferent ducts; and the processes of erection, emission, and ejaculation. However, maintenance of normal function of reproductive organs depends more on the neuroendocrine system.
Hypothalamic–hypophyseal axis The hypothalamus is part of the diencephalon of the brain, but its exact boundaries have not been defined critically in the horse. From work with other species, the hypothalamus seems to be involved in regulation of appetite and thirst, body temperature, vasomotor activity, emotion, use of body nutrient reserves, activity of the intestinal tract and bladder, states of
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sleep and wakefulness, sexual behavior, and release of tropic hormones. This latter role is essential for controlling reproductive function. The hypophysis is connected to, and extends downward from, the hypothalamus. The adenohypophysis synthesizes and discharges a number of hormones that control reproductive processes in the stallion and mare.80, 81 The neurohypophysis, on the other hand, does not produce hormones but simply serves as a storage reservoir for hormones produced by neural tissue within the brain. The hypothalamus and adenohypophysis are linked by portal vessels, which extend from the hypothalamus through the infundibulum to the pars distalis. Within this portal system, blood normally flows directly from the hypothalamus to the pars distalis. The portal vessels are the only direct link between the hypothalamus and the adenohypophysis. Under appropriate neural stimulation, the hypothalamus synthesizes and discharges a number of ‘releasing hormones’. The releasing hormone directly involved in controlling reproductive function is gonadotropin-releasing hormone (GnRH) (Chapter 99). GnRH is discharged by the hypothalamus in short, pulsatile bursts. This fact, coupled with rapid removal of GnRH from the blood, results in a pulsatile stimulation of the gonadotropin-secreting cells in the adenohypophysis. A large number of compounds similar to GnRH, called structural analogues, have been synthesized and used in research to study and control reproductive function. Two types of analogues of GnRH are available: those that block the action of the natural hormone and those that are more effective in evoking a response than natural GnRH. The adenohypophysis produces at least six tropic hormones, but only two or three have a direct role in male reproduction. The adenohypophysis produces LH and FSH in direct response to stimulation by GnRH. Luteinizing hormone and FSH (Chapter 99) are gonadotropic hormones because they act on the gonads (testes in males) and stimulate their function, including production of steroid hormones and spermatozoa. Although details of the secretory pattern of GnRH are not available for stallions, GnRH must be secreted in a pulsatile manner. Only pulsatile secretion of GnRH could account for secretion of LH as a series of distinct pulses.82–85 However, in some stallions, pulsatile secretion of LH is not evident from analyses of jugular blood, especially during the breeding season.47, 85 Secretion of FSH also is pulsatile, although more short-term variability exists,47, 85 possibly reflecting intermittent discharges of small amounts of hormone. Pulsatile secretion of FSH is generally associated with an episode of LH secretion. Although several studies of concentrations
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of hormones in blood of stallions have been made, 17, 44, 47, 56, 78,82–91 concepts for endocrine control of reproductive function are based primarily on data from other species.34, 58, 80, 92, 93 The concepts outlined herein are true regardless of season or age. However, great seasonal differences occur in secretory rates and concentrations of all hormones in blood. Testicular hormones Steroid hormones produced by Leydig cells (Figure 96.15) are the primary endocrine product of the testes. However, in the last decade the importance of other hormones secreted by the testes has become evident. Sertoli cells secrete inhibin and activin, a pair of related glycoprotein hormones, which share a common subunit.94 The inhibin/activin family include three subunits: α, β A , and β B . Inhibin is the combination α/β A or α/β B , and activin is the combinations β A /β B, β A /β A , or β B /β B . The relative amounts of different inhibins, or activins, secreted differ as a function of cell type (Sertoli cells of male or granulosa cells of female, species, and stage of development). Although stallion Sertoli cells secrete inhibin and activin, the exact molecular form(s) of each remains to be established. Numerous reports have been made of secretion of GnRH-like peptides by the rat testis. This appears to be unique to rats, because attempts to detect secreted GnRH or responses of testicular tissue to applied GnRH in other species all have been unsuccessful. No such study with stallion tissue has been reported. Based on data for rats and rams, Leydig cells likely secrete oxytocin into the interstitial fluid.95, 96 Apparently, oxytocin facilitates rhythmic contraction of seminiferous tubules to help evacuate sperm. Oxytocin seems to be transported by Sertoli cells into the luminal fluid or secreted by the rete testis, because in rams the rete testis fluid draining the testis contains about 550 pg/mL as compared with 124 pg/mL in serum from testicular venous blood.96 Oxytocin may be important in facilitating contractions of the efferent ducts and proximal epididymal duct. Veeramachaneni and Amann97 speculated that a deficiency of testicular oxytocin in fluid entering the excurrent ducts or its uptake by the epithelium therein could be a cause of sperm stasis and formation of granulomatous lesions (common in cattle, sheep, and goats, but the frequency in horses is unknown). Research in the coming decade certainly will reveal an abundance of regulatory peptides, important for signaling between Leydig cells, Sertoli cells, and germinal cells. The testes may secrete unidentified hormones that act on other reproductive organs or
the hypothalamic–hypophyseal axis. Certainly progesterone and conjugated estrogen may have some systemic role. Regulation of hormone secretion In the adult male, production of testosterone is controlled by episodic bursts of LH secretion, which periodically elevate the concentration of LH in blood reaching the testis far above the basal level. Consequently, basal production of testosterone is augmented by episodic bursts of production of testosterone49, 54, 82, 85, 87, 90 (Figure 96.17). Testosterone produced by Leydig cells enters venous blood draining the testis, passes into the general circulation, and then moves to the hypothalamus and adenohypophysis (Figure 96.18). In this manner, testosterone feeds back via a long loop and modulates discharge of GnRH and LH.78, 83 If the concentration of testosterone reaching the hypothalamus and adenohypophysis is relatively high, discharge of GnRH by the hypothalamus is suppressed and response of the adenohypophysis to available GnRH also is suppressed. This negative effect of gonadal steroids has been attributed to testosterone per se, but it could result from a combined effect of testicular testosterone, estradiol, and possibly progesterone. Furthermore, testosterone might be converted to estradiol in the target cells of the hypothalamus or adenohypophysis. That negative effects of steroids are expressed on both the hypothalamus and adenohypophysis is likely, but unproven. In any case, because of this negative feedback, concentration of LH would be low in blood entering the testis so Leydig cells would be exposed to a low concentration
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Figure 96.17 Concentrations of LH and testosterone (T) in serum from jugular blood taken every 20 minutes from a stallion. Although pulsatile secretion of both LH and testosterone is not always as evident as in this example, pulsatile secretion of LH was evident for 96% of 231 such sequences of blood samples (72 samples/sequence) taken from 21 stallions in all seasons of the year. (From Pickett et al.7 )
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Acessory sex glands + +
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Leyding cells in interstitial tissue
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Testis Sertoli cell in seminiferous tubule
Figure 96.18 Interrelationship of hypophyseal hormones acting on Leydig cells and Sertoli cells of the seminiferous tubules and feedback control of gonadal hormones on the hypothalamus and adenohypophysis. An increased level of testosterone in peripheral blood, either as a result of increased production by the testes or after injection of hormone, feeds back on the hypothalamus and adenohypophysis to suppress discharge of GnRH and LH, respectively. Therefore, Leydig cells produce less testosterone and concentration of testosterone around the seminiferous tubules drops. Circulating FSH acts directly on Sertoli cells, which secrete two protein hormones: inhibin, which acts on the adenohypophysis to suppress selectively the amount of FSH secreted in response to GnRH, and activin, which stimulates FSH secretion. Adequate concentrations of testosterone and FSH must be present to stimulate Sertoli cells to produce an environment appropriate for normal spermatogenesis. A, activin; ABP, androgen-binding protein; E, estradiol or other estrogens; GnRH, gonadotropin-releasing hormone; I, inhibin; LH, luteinizing hormone; FSH, follicle-stimulating hormone; PRL, prolactin; T, testosterone. (Modified from Pickett et al.7 )
of LH, and consequently, Leydig cells would only secrete testosterone at a basal rate. As concentration of testosterone in peripheral blood declines, the negative block is removed and episodic discharge of GnRH from the hypothalamus can occur. This is followed by a discharge of LH from the adenohypophysis, an elevated concentration of LH in blood flowing to the testis, and rapid stimulation of Leydig cells to produce and discharge testosterone (compare data for LH and testosterone in Figure 96.17). Thus, Leydig cells, the hypothalamus, and the adenohypoph-
ysis are involved in a circular feedback loop that regulates concentrations of LH and testosterone in peripheral blood (Figure 96.18). Although secretion of FSH also is stimulated by GnRH, secretion of FSH and LH must be controlled by different mechanisms. Bursts of secretion of LH are sometimes not accompanied by a discharge of FSH and vice versa.47, 85 Furthermore, a differential effect of season on secretion of LH and FSH occurs. Synthesis and secretion of FSH may be much less dependent on GnRH than secretion of LH.98 Based
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on research with non-equine species, FSH seems to act exclusively on Sertoli cells within the seminiferous tubules.3, 34 Among the products of Sertoli cells are several protein hormones, including closely related inhibin and activin. Among other functions, these hormones act on the adenohypophysis to suppress (inhibin) or stimulate (activin) secretion of FSH, with little or no effect on secretion of LH. Thus, the inhibin/activin system and probably the ratio of testosterone to estradiol impinging on the adenohypophysis control the relative amounts of LH and FSH secreted in response to GnRH from the hypothalamus. Although the stallion testis produces uniquely high concentrations of estrogens, researchers are not certain if estrogens are produced in Sertoli cells, Leydig cells, or some other component of the testis. Figure 96.18 incorporates the speculation that a portion of testosterone produced by Leydig cells is transformed within Leydig cells to estradiol or other estrogens. In any case, blood draining from the stallion testis contains high concentrations of estrogens, and high concentrations of estrogens in blood flowing to the hypothalamus and adenohypophysis may suppress discharge of GnRH or LH and FSH in a negative feedback loop.78, 98 Because of feedback loops involving the hypothalamus, adenohypophysis, and testis, injecting hormones will alter the delicate hormonal balance in a stallion and may profoundly disturb reproductive function. Sequelae of such manipulations, often with undesirable results, are detailed in Chapter 114.
Hormonal control of spermatogenesis The hormonal requirements for normal spermatogenesis in a stallion are unknown. Based on research with rats, duration of the cycle of seminiferous epithelium and total duration of spermatogenesis are probably not modified by hormonal balance within the testis.2, 27, 29, 43 However, degree of germ cell degeneration is partially under hormonal control.2, 27, 41, 43, 99 Seminiferous tubules are surrounded by Leydig cells producing testosterone, so the seminiferous epithelium is exposed to a higher concentration of testosterone than found in peripheral blood. This high level of testosterone is essential for normal spermatogenesis.41, 99 In rams, FSH is necessary for spermatogenesis30, 41, 99 and LH may have a direct role in regulating division of spermatogonia in addition to its role at stimulating production of testosterone.99 The role of FSH in bringing about normal development of spermatids apparently is mediated via Sertoli cells.30 Definitive conclusions on hormonal control of spermatogenesis in stallions must await
data obtained with stallions, because some differences among species exist.99
Descent of the testes A description of the embryological development of the equine reproductive system is beyond the scope of this chapter. However, understanding the process by which the testes normally descend from the abdominal cavity into the scrotum is essential. In the normal colt, both testes should descend into the scrotum between 30 days before and 10 days after birth.100, 101 Failure of normal testicular descent is common in horses102 and is termed cryptorchidism. By day 40 of gestation, the testis is suspended from the abdominal wall (Figure 96.19) and the mesonephric duct, which later gives rise to the epididymis and ductus deferens, leads into the pelvic area.100 A narrow evagination, termed the vaginal process (process vaginalis peritonei), starts to form about day 43 of gestation and progressively develops to form the inguinal canal (canalis inguinalis) (Figure 96.19). Around day 150, the developing cauda epididymidis is drawn to or just within the internal inguinal ring (annulus inguinalis profundus) (Figure 96.20), but the testis is large and cannot enter the inguinal canal.100 Entrance of the testis into the inguinal canal typically begins between 270 and 300 days of gestation.100 This occurs only after the vaginal process and internal inguinal ring have been stretched sufficiently by the enlarging cauda epididymidis (Figure 96.21) to allow entrance of the testis that has diminished in size from 50 to 30 g. Pressure from fluid in the abdominal cavity, and possibly from the intestines, forces the testis down through the inguinal canal. This descent places the lamina visceralis or the tunica vaginalis – the outer covering of the testis – into apposition with the lamina parietalis of the tunica vaginalis – the former vaginal process – separated by the cavum vaginale (Figure 96.21). Bergin et al.100 reported that the earliest complete descent of both testes was at 315 days of gestation, about 25 days before parturition. In 32 fetuses between 9 months of gestation and birth, descent of the right testis was more advanced than the left in 78% of the fetuses, while the left testis was more advanced in only 3%. Of 12 fetuses collected at term, 42% had completely descended testes, 25% had both testes within the inguinal canals, 17% had one testis in the scrotum and one in the inguinal canal, and 17% had both testes within the abdominal cavity. A total of five of nine colts less than 1 week old had complete bilateral descent of the testes into the scrotum. As reviewed by Bergin et al.,100 failure of the testes to descend has been attributed to abnormalities of the testis, development of adhesions between the testis
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Cephalic ligament of testis
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Dorsal body wall Mesorchium Mesonephric duct Peritoneal covering over testis A
A Testis
Tunica albuginea of testis
Caudal gonadal ligament
B
Invaginated peritoneal lining Gubernacular cord
Vaginal processes
B Vaginal processes Gubernacular bulb
Figure 96.19 A horse fetus at 75 days of gestation. The testis is suspended within the abdominal cavity by a thin double-layered band of fused tissue termed the mesorchium. The caudal gonadal ligament, extending from the testis to its point of fusion with the mesonephric duct (future epididymis and ductus deferens), is continuous with the gubernacular cord that extends to the gubernacular bulb. The vaginal process (future lamina parietalis of the tunica vaginalis) encloses about one third of the gubernacular bulb. (Modified from Pickett et al.7 )
and adjacent structures, or an abnormal outpouching of the vaginal process. Bergin et al. discounted these factors as causes of cryptorchidism and suggested that the most obvious reasons for the testis to remain in the abdominal cavity included the following:
Ductus epididymidis Ductus deferens
Mesorchium
1. Insufficient abdominal pressure to properly expand the vaginal process. 2. Stretching of the gubernacular cord (chorda gubernaculi).
Cephalic ligament of testis Dorsal body wall Mesorchium Ductus epididymidis
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A testis Caudal gonadal ligament
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Peritoneal covering of testis (future visceral layer of vaginal tunic)
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Invaginated peritoneal lining (inner = future visceral layer of vaginal tunic outer = future parietal layer of vaginal tunic)
Gubernacular bulb Vaginal processes
Figure 96.20 A horse fetus at 175 days of gestation. The vaginal process extends almost to the scrotum and has virtually enclosed the gubernacular bulb. The mesorchium is continuous from the gubernaculum and the vaginal process to the testis. The epididymis and ductus deferens have formed from the mesonephric duct and the future cauda epididymidis will form where the duct is sharply reflected. (Modified from Pickett et al.7 )
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Corpus epididymids Ductus deferens Tunica albuginea Testis
Ductus deferens
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Corpus epididymidis
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Mesorchium Ductus deferens surrounded by mesoductus deferens Vainal cavity
A Mesorchium B Cauda epidiymidis Gubernacular bulb
Testis
Caudal gonadal ligament
Blood vessels + nerve surrounded by visceral laeyr of vaginal tunic Parietal layer of vaginal tunic
Figure 96.21 A horse fetus near term. The testis has passed through the inguinal canal, but is not fully within the scrotum. The gubernacular bulb, gubernacular cord, cauda epididymidis, and caudal gonadal ligament all have regressed. The testis now is connected by the mesorchium to the dorsal wall of the lamina parietalis of the tunica vaginalis (formally the vaginal process). The deferent duct, blood vessels, and lymphatics are connected to the lamina parietalis of the tunica vaginalis by folds of the mesorchium. (Modified from Pickett et al.7 )
3. Insufficient growth of the gubernaculum and cauda epididymidis so that they are unable to expand the inguinal ring sufficiently to allow entrance of the testis. 4. Displacement of the testis to a position where intestinal pressure prevents tension from the gubernaculum, via the gubernacular cord, pulling the testis into the vaginal process.
Puberty At birth, the testis contains few (if any) functional Leydig cells and only indifferent supporting cells and gonocytes (progenitors of Sertoli cells and spermatogonia). The stallion soon enters the infantile stage of its life. Based on data for other species, this infantile stage is characterized by a declining number of gonocytes, virtual absence of gonadotropin secretion by the adenohypophysis, and limited steroidogenesis (at least in terms of testosterone) by Leydig cells.41,103–107 Spermatozoa are not produced. This infantile stage continues through ≥ 6 months in stallions, when changes are initiated which continue through a prepubertal stage and culminate in puberty. Timing of the prepubertal changes differs among stallions and might be influenced by breed and season of birth. Puberty (meaning ‘to beget’) is the time when a stallion is first capable of successfully participating in reproduction. In literature on domestic animals, puberty is considered to be a definitive end point – being capable of reproduction – and,
by definition, implies achievement of spermarche (production of first spermatozoa) and completion of prepubertal events. In literature pertaining to humans, however, the term puberty has no specific end point, but refers rather to the series of events termed the prepubertal stage herein. With some stallions, a few spermatozoa are available for ejaculation by 14 months of age. Following puberty, development of reproductive capacity continues (postpubertal stage), and 2–4 years after puberty, a stallion achieves sexual maturity (maximum reproductive capacity). Years later, reproductive senescence may occur. For most stallions, no change in DSP between 4 and 20 years of age occurs.22 Little is known about testicular function or changes in the neuroendocrine system of stallions during the infantile stage of development or exactly when the prepubertal stage is initiated. Timing of specific events likely ranges considerably, especially comparing light horse and draft breeds. Starting at about 9 months, concentrations of FSH and especially LB in blood increase (Figure 96.22), and, around 12 months, the testes start to grow and develop rapidly.88, 108, 109 Several months later, they gradually begin to produce spermatozoa.88, 110 Total scrotal width increases in a linear manner from 42 mm at 42 weeks to 85 mm at 96 weeks of age (about 1 mm/ week), based on data for 15 colts born in July and early August.88 Increased production of testosterone by Leydig cells after 80 weeks of age is evident from concentrations of testosterone in jugular vein blood
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Figure 96.22 Concentrations of LH, FSH, and testosterone in serum prepared from jugular blood. Age at puberty averaged 83 weeks, but ranged from 56 to 97 weeks (based on seminal characteristics). Based on data for eight colts born in July or early August. Blood samples were taken from each colt at 8, 16, and 24 weeks of age and every 4 weeks thereafter. (From Pickett et al.7 )
(Figure 96.22).88 These prepubertal developments are culminated in puberty when a stallion produces spermatozoa and would be fertile if allowed to breed a mare. For 15 colts born in July and early August, and carefully monitored for reproductive development, age at puberty averaged 83 weeks (puberty defined as first ejaculate containing 50 million spermatozoa of which ≥ 10% are motile).88 A total of 11 of the 15 colts attained puberty by 90 weeks of age. After puberty, the quantity and quality of spermatozoa produced slowly increased (Table 96.6). At 2 years old, ejaculates contained 3.3 billion spermatozoa, although percentages of motile and structurally normal spermatozoa still were low. With bulls, fertility of spermatozoa increases for about 6 months after puberty (J.O. Almquist, personal communication). In the absence of complete information for stallions, changes in the hypothalamic– hypophyseal–testicular axis must be deduced based on information for bulls, rams, and other species.92, 93, 103, 107 Typically, little secretion of LH, FSH, or testosterone occurs in the infantile stage, but transition from the infantile to prepubertal stage is evidenced by a marked increase in frequency and amplitude of discharges of LH.92, 93, 103, 105 The relatively low secretion rate of LH in stallions before about 36 weeks of age (Figure 96.22) probably reflects a low concentration of GnRH receptors in the adenohypophysis (based on data for bulls)105 and little secretion of GnRH from the hypothalamus. In bulls, high secretion of LH, evidenced by frequent discharges of high amplitude, starts at 10–12 weeks of age and continues for about 6 weeks.103 A similar
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event apparently occurs in stallions, as evidenced by the high concentrations of LH in jugular vein blood between 36 and 45 weeks of age (Figure 96.22). In stallions, however, the surge of LH secretion may reflect simply a change in amplitude of LH pulses rather than a combined effect of increased frequency of discharges of LH with each discharge containing more hormone; in adult stallions, the seasonal rise of serum concentration of LH was solely caused by an increase in LH pulse amplitude.47, 57, 82 The stallion hypophysis has the capability to secrete considerably more LH at 32 weeks of age than actually is secreted,89 although the amount of LH released after GnRH administration increased from 32 to 48 weeks of age. For the eight stallions studied by Naden et al.,88 Leydig cells were unresponsive to the increased LH stimulation which occurred from 36 to 45 weeks of age because no change in testosterone concentration occurred in peripheral blood88 (Figure 96.22). This also is true for bulls, for which more extensive data are available.92, 93, 103, 105 Prolonged stimulation of Leydig cells by LH likely is necessary to complete their differentiation so that they can secrete testosterone. In stallions, an interval of almost 12 months is seen between initial increase of LH concentration in the blood and increase of testosterone concentration88 (Figure 96.22), whereas in bulls the interval is about 3 months.103 Perhaps this difference is a consequence of the more pronounced seasonal effect on reproductive function in stallions than in bulls, coupled with the fact that in Naden’s study Leydig cells were exposed to high concentrations of LH in May to July, and about 3 months later the effect of season may have suppressed secretion of testosterone that otherwise might have occurred.88 In adult stallions, secretion of testosterone is reduced and blood concentrations are low during September through January.88 Table 96.6 Characteristics of light horse stallions and their semen at pubertya and 96 weeks of age
Characteristic Total scrotal width (mm) Gel-free seminal volume (mL) Sperm concentration (106 /mL) Total sperm per ejaculate (109 ) Motile spermatozoa (%) Morphologically normal spermatozoa (%)
Puberty 77 12 99 1.1 26 33
96 weeksb 88 21 162 3.3 26 44
Puberty was defined as the first ejaculate that contained 50 × 106 spermatozoa of which ≥ 10% were motile. Age at puberty averaged 83 weeks, but ranged from 56 to 97 weeks for the 15 colts. Seminal data are averages for nine ejaculates per colt, collected every third day. From Naden et al.88
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With onset of the next breeding season, secretion of LH, FSH, and testosterone all increased in the young colts born in July and early August (Figure 96.22). What causes inactivity of the neuroendocrine system during the infantile stage? Extrapolating from data for bulls and rams,99,103–105 I speculate that in stallions an estradiol-mediated block suppresses secretion of GnRH by the hypothalamus before about 16 weeks of age. Then pulsatile discharge of GnRH begins or increases, provided neurochemical stimuli are present. Concomitantly, an increased concentration of estradiol receptors in the adenohypophysis might allow initiation of positive feedback of estradiol on this organ, with a resultant increase in concentration of GnRB receptors in the adenohypophysis and increased responsiveness of the adenohypophysis to discharges of GnRH initiated concurrently. This course of events allows secretion of limited amounts of LH by 16 weeks of age (Figure 96.22), although the gland is not operating at maximum capacity. Continued GnRH stimulation of the adenohypophysis presumably induces greatly increased synthesis of mRNA for the β-subunit of LH between 20 and 32 weeks of age, which, coupled with increased stimulation from GnRH, enables copious production of LH present at 32–45 weeks of age. Naden et al.88 speculated that the testicular secretion involved in maturation of the hypothalamic–hypophyseal axis might be estradiol or androstenedione, rather than testosterone.88 Secretion of a gonadal steroid, other than testosterone, which acts on the hypothalamic–hypophyseal axis, could be a cause of the seasonal suppression in LH secretion from 55 to 85 weeks of age in the colts studied88 (Figure 96.22). The interplay between hypophyseal stimulation of Leydig cells and the seminiferous epithelium and between Leydig cells and the seminiferous epithelium are unknown for stallions. For other species, however, indifferent supporting cells apparently must be exposed to FSH and receive androgenic stimulation to complete their differentiation into Sertoli cells capable of supporting spermatogenesis. Given the data depicted in Figure 96.22, sufficient androgenic stimulation appears to be provided by the intratesticular milieu at 70–75 weeks of age to enable onset of spermatogenesis and ejaculation of spermatozoa at 83 weeks of age (puberty), although secretion of testosterone continued to increase for at least several months after the initial rise around 72 weeks of age. Based on data for bulls and rats,23, 106, 107 the efficiency of spermatozoal production in stallions likely increases rapidly through the interval from puberty to > 100 weeks of age, although no data exist for stallions.
Ejaculation The process commonly considered to be ejaculation actually involves three sequential processes: erection, emission, and ejaculation. Erection is the lengthening and stiffening of the penis, resulting from engorgement with blood of the corpus cavernosum penis and, later, the corpus spongiosum penis. Emission is the movement and deposition of spermatozoa and fluid from the ductus deferens and cauda epididymidis, as well as fluids from the accessory sex glands, into the pelvic urethra (pars pelvina). Ejaculation is the actual expulsion of semen through the urethra. Erection of the penis is initiated by sensory stimuli of the glans penis or psychic stimulation of the cerebral cortex. Erection is initiated during teasing because visual stimuli presented by location or teaser animal (or dummy) initiates responses in the cerebrum. Parasympathetic impulses pass from the second, third, and fourth sacral segments of the spinal cord, via splanchnic nerves, to the penis. This overrides sympathetic stimulus normally keeping the arterioles in the penis partly constricted and causes dilation of penile arterioles. More blood enters the corpus cavernosus penis and corpus spongiosum penis. Concurrently, extrinsic muscles of the penis (i.e., ischiocavernosus, bulbospongiosus, and urethralis) contract and compress the deep and dorsal veins of the penis against the ischial arch, impeding venous return from the cavernous bodies of the penis. Consequently, a shunting of blood to fill and distend the corpus cavernosum penis and corpus spongiosum penis occurs, and enlargement of the penis results. Cardiac output may also increase. The penis returns to its flaccid state when the arteries constrict to their normal state as a result of sympathetic impulses, and pressure on the veins is relieved by relaxation of the ischiocavernosus and bulbospongiosus muscles. In the stallion, emission (caused by sympathetic impulses) and ejaculation (caused by parasympathetic stimulation) occur as a series of strong, pulsatile contractions of the urethralis and bulbospongiosus muscles so that several successive jets of semen are ejaculated.111 The general sequence is probably that the prostate gland secretes a watery fluid into the pelvic urethra and some prostatic fluid is probably ejaculated as a prespermatozoal fraction. Next, emission and ejaculation of the spermatozoal-rich fraction occurs. This fraction consists of spermatozoa and epididymal secretions and probably prostatic fluids and watery bulbourethral gland fluid. Typically, three to six sequential discharges of sperm-rich fluid occur. Although not always present, the gel or post-spermatozoal fraction
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Physiology and Endocrinology is derived from the vesicular glands. The function of the gel is unknown, but it is not necessary for fertility. Some ejaculates, especially the second ejaculate in a 2-hour period, contain little or no gel. Little is known about the secretions of the glands in the ampulla of the ductus deferens or urethra in terms of chemical composition or contribution to an ejaculate. The characteristics of semen from a given stallion vary greatly as a function of season, age, testicular size (a highly heritable trait), interval of abstinence since the previous ejaculation(s), and the extent of sexual arousal or courtship before ejaculation. Composition of semen and details of spermatozoal structure and function are presented in the chapter on spermatozoal function (Chapter 102).
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Acknowledgement Some individuals, and some manuscripts have wonderful insight and historical value. Such is the case with Dr. Rupert P. Amann and four chapters that he so ably produced for the first edition of Equine Reproduction. While Dr. Amann is now retired and has turned the baton over to others, these pieces of work have endured the test of time and continue to be an invaluable resource for the equine practitioner and academician. As such, we decided to reprint these chapters for you.
References 1. Harrison RG. Effect of temperature on the mammalian testis. In: Hamilton DW, Greep RO (eds) Handbook of Physiology, vol. 5, section 7. Washington DC: American Physiology Society, 1975; pp. 219–23. 2. Setchell BP. The Mammalian Testis. Ithaca: Cornell University Press, 1978. 3. Setchell BP, Brooks DE. Anatomy, vasculature, innervation and fluids of the male reproductive tract. In: Knobil E, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 753–836. 4. Waites GMH. Temperature and fertility in mammals. In: Proceedings of the Sixth International Congress on Animal Reproduction and Artificial Insemination, vol. 1, 1968, pp. 235–56. 5. Waites GMH, Ortavant R. Effets precoces d’une breve elevation de la temperature testiculaire sur la spermatogenese du Belier. Ann Bio Anim Biochim Biophys 1968;8:323–31. 6. Austin JW, Hupp EW, Murphree RL. Effect of scrotal insulation on semen of Hereford bulls. J Anim Sci 1961;20:307–10. 7. Pickett BW, Amann RP, McKinnon AO, Squires EL, Voss JL. Management of the Stallion for Maximum Reproductive Efficiency, II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins: Colorado State University. 8. Berndtson WE, Squires EL, Thompson Jr DL. Spermatogenesis, testicular composition and the concentration of
15.
16.
17.
18. 19.
20.
21.
22.
23.
24.
25. 26.
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testosterone in the equine testis as influenced by season. Theriogenology 1983;20:449–57. Johnson L. Increased daily sperm production in the breeding season of stallions is explained by an elevated population of spermatogonia. Biol Reprod 1985;32: 1181–90. Johnson L. A new approach to quantification of Sertoli cells that avoids problems associated with the irregular nuclear surface. Anat Rec 1986;214:231–7. Johnson L. Effect of season on spermatocytogenesis in stallions. In: Proceedings of the Annual Meetings of the American Association of Veterinary Anatomists, 1987, p. 28. Johnson L. Spermatogenesis. In: Cupps PT (ed.) Reproduction in Domestic Animals, 4th edn. New York: Academic Press, 1990; pp. 173–219. Johnson L, Amann RP, Pickett BW. Scanning electron and light microscopy of the equine seminiferous tubule. Fertil Steril 1978;29:208–15. Johnson, L, Amann RP, and Pickett BW. Scanning electron microscopy of the epithelium and spermatozoa in the equine excurrent duct system. Am J Vet Res 39:1428-1434, 1978. Johnson L, Neaves WB. Age-related changes in the Leydig cell population, seminiferous tubules, and sperm production in stallions. Biol Reprod 1981;24:703–12. Johnson L, Tatum ME. Sequence of seasonal changes in numbers of Sertoli, Leydig and germ cells in adult stallions. In: Proceedings of the Eleventh International Congress on Animal Reproduction and Artificial Insemination, vol. 3, 1988, pp. 373–4. Johnson L, Thompson Jr DL. Age-related and seasonal variation in the Sertoli cell population, daily sperm production and serum concentrations of follicle-stimulating hormone, luteinizing hormone and testosterone in stallions. Biol Reprod 1983;29:777–89. Johnson L. Seasonal differences in equine spermatocytogenesis. Biol Reprod 1991;44:284–91. Johnson L, Varner DD, Thompson Jr DL. Effect of age and season on the establishment of spermatogenesis in the horse. J Reprod Fertil Suppl 1991;44:87–97. Swierstra EE, Gebauer MR, Pickett BW. Reproductive physiology of the stallion. I. Spermatogenesis and testis composition. J Reprod Fertil 1974;40:113–23. Swierstra EE, Pirkett BW, Gebauer MR. Spermatogenesis and duration of transit of spermatozoa through the excurrent ducts of stallions. J Reprod Fertil Suppl 1975;23:53– 7. Johnson L, Varner DD, Tatum ME, Scrutchfield WL. Season but not age affects Sertoli cell number in adult stallions. Biol Reprod 1991;45:404–10. Amann RP. A critical review of methods for evaluation of spermatogenesis from seminal characteristics. J Androl 1981;2:37–58. Amann RP, Johnson L, Thompson Jr DL, Pickett BW. Daily spermatozoal production, epididymal spermatozoal reserves and transit time of spermatozoa through the epididymis of the Rhesus monkey. Biol Reprod 1976;15:586–92. Berndtson WE. Methods for quantifying mammalian spermatogenesis: a review. J Anim Sci 1977;44:818–33. Berndtson WE, Igboeli G, Parker WG. The numbers of Sertoli cells in mature Holstein bulls and their relationship to quantitative aspects of spermatogenesis. Biol Reprod 1987;37:60–7.
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27. Clermont Y. Kinetics of spermatogenesis in mammals: seminiferous epithelium cycle and spermatogonial renewal. Physiol Rev 1972;52:198–236. 28. Courot M, Hochereau-de Reviers M-T, Ortavant R. Spermatogenesis. In: Johnson AD, Gomes WR, VanDemark NL (eds) The Testis, vol. 1. New York: Academic Press, 1970; pp. 339–42. 29. de Kresta DM, Kerr JB. The cytology of the testis. In: Knobil E, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 837–932. 30. Hochereau-de Reviers M-T, Monet-Kuntz C, Courot M. Spermatogenesis and Sertoli cell numbers and function in rams and bulls. J Reprod Fertil Suppl 1987;34:101–14. 31. Lok D, Weenk D, De Rooij DG. Morphology, proliferation, and differentiation of undifferentiated spermatogonia in the Chinese hamster and the ram. Anat Rec 1982;203:83–99. 32. Russell LD, Peterson RN. Determination of the elongate spermatid-Sertoli cell ratio in various mammals. J Reprod Fertil 1984;70:635–41. 33. Tung KSK, Teuscher C, Meng AL. Autoimmunity to spermatozoa and testis. Immunol Rev 1981;55:217–55. 34. Bardin CW, Chang CY, Musto NA, Gunsalus GL. The Sertoli cell. In: Knobil E, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 933–74. 35. Purvis K, Hansson V. Hormonal regulation of spermatogenesis. Int J Androl Suppl 1981;3:81–143. 36. Wright WW. Intragonadal control of testis function. In: Medically Assisted Contraception. Washington, DC: National Academy Press, 1989; pp. 191–210. 37. Belv´e AR. The molecular biology of mammalian spermatogenesis. Oxf Rev Reprod Biol 1979;1:159–261. 38. Amann RP, Thompson Jr DL, Squires EL, Pickett BW. Effects of age and frequency of ejaculation on sperm production and extragonadal sperm reserves in stallions. J Reprod Fertil Suppl 1979;27:1–6. 39. Berndtson WE, Hoyer JH, Squires EL, Pickett BW. Influence of exogenous testosterone on sperm production, seminal quality and libido of stallions. J Reprod Fertil Suppl 1979;27:19–23. 40. Gebauer MR, Pickett BW, Swierstra EE. Reproductive physiology of the stallion. II. Daily production and output of sperm. J Anim Sci 1974;39:732–6. 41. Courot M, Hochereau de Reviers MT, Manet-Kuntz C, Locartelli A, Pisselet C, Blanc MR, Dacheux JL. Endocrinology of spermatogenesis in the hypophysectomized ram. J Reprod Fertil Suppl 1979;26:165–73. 42. Ellery JC. Spermatogenesis, accessory sex gland histology and the effects of seasonal change in the stallion. PhD thesis, St. Paul, University of Minnesota, 1971. 43. Steinberger E, Steinberger A. Spermatogenic function of the testis. In: Hamilton DW, Greep RO (eds) Handbook of Physiology, vol. 5, section 7. Washington DC: American Physiology Society, 1975; pp. 1–19. 44. Hoyer J. The effect of testosterone on reproductive function in stallions. MS thesis, Fort Collins, Colorado State University, 1978. 45. Squires EL, Todter GE, Berndtson WE, Pickett BW. Effect of anabolic steroids on reproductive function in young stallions. J Anim Sci 1982;54:576–82. 46. Johnson L, Thompson Jr DL. Effect of seasonal changes in Leydig cell number on the volume of smooth endoplasmic reticulum in Leydig cells and intratesticular testosterone content in stallions. J Reprod Fertil 1987;81:227– 32.
47. Clay CM. Influences of season and artificial photoperiod on reproduction in stallions. PhD thesis, Fort Collins, Colorado State University, 1988. 48. Berndtson WE, Desjardins C, Ewing LL. Inhibition and maintenance of spermatogenesis in rats implanted with polydimethylsiloxane capsules containing various androgens. J Endocrinol 1974;62:125–35. 49. Amann RP, Ganjam VK. Effects of hemicastration or hCGtreatment on steroids in testicular vein and jugular vein blood of stallions. J Androl 1981;2:132–9. 50. Seamans MC, Roser JF, Linford RL, Liu IKM, Hughes JP. Gonadotrophin and steroid concentrations in jugular and testicular venous plasma in stallions before and after GnRH injection. J Reprod Fertil Suppl 1991;44:57–67. 51. Silberzahn P, Zwain I, Guerin P, Benoit E, Jouany JM, Bannaire Y. Testosterone response to human chorionic gonadotropin injection in the stallion. Equine Vet J 1988;20:61–3. 52. Bedrak E, Samuels LT. Steroid biosynthesis by the equine testis. Endocrinology 1969;85:1186–95. 53. Gaillard J-L, Silberzahn P. Aromatization of 19norandrogens by equine testicular microsomes. J Biol Chem 1987;262:5717–22. 54. Ganjam VK. Episodic nature of the 4 -ene and 5 ene steroidogenic pathways and their relationship to the adrenogonadal axis in stallions. J Reprod Fertil Suppl 1979;27:67–71. 55. Lindner HR. Androgens and related compounds in the spermatic vein blood of domestic animals. IV. Testicular androgens in the ram, boar and stallion. J Endocrinol 1961;23:171–8. 56. Raeside JI. Seasonal changes in the concentration of estrogens and testosterone in the plasma of the stallion. Anim Reprod Sci 1979;1:205–12. 57. Ewing L, Brown BL. Testicular steroidogenesis. In: Johnson AD, Gomes WR (eds) The Testis, vol. 4. New York: Academic Press, 1977; pp. 239–87. 58. Hall PF. Testicular steroid synthesis: organization and regulation. In: Knobil E, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 975–98. 59. Zirkin BR, Samfulli R, Awoniyi CA, Ewing EL. Maintenance of advanced spermatogenic cells in the adult rat testis: quantitative relationship to testosterone concentrations within the testis. Endocrinology 1989;124:3043–9. 60. Zirkin BR, Ewing LL, Kormann N, Cochran RC. Testosterone secretion by rat, rabbit, guineapig, dog, and hamster testes perfused in vitro: Correlation with Leydig cell ultrastructure. Endocrinology 1980;107:1867–74. 61. Lopez ML, de Souza W, Bustos-Obregon E. Cytochemical analysis of the anionic sites on the membrane of the stallion spermatozoa during the epididymal transit. Gamete Res 1987;18:319–32. 62. Amann RP. Function of the epididymis in bulls and rams. J Reprod Fertil Suppl 1987;34:115–31. 63. Amann RP. Maturation of spermatozoa. In: Proceedings of the International Congress of Animal Reproduction and Artificial Insemination, vol. 5, 1988, pp. 320–8. 64. Hamilton DW. Structure and function of the epithelium lining the ductus efferentes, ductus epididymidis, and ductus deferens in the rat. In: Hamilton DW, Greep RO (eds) Handbook of Physiology, vol. 5, section 7. Washington DC: American Physiology Society, 1975; pp. 259–301. 65. Orgebin-Crist M-C, Danzo BJ, Davies J. Endocrine control of the development and maintenance of sperm fertilizing ability in the epididymis. In: Hamilton DW, Greep RO
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66.
67.
68. 69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79. 80. 81.
82.
83.
(eds) Handbook of Physiology, vol. 5, section 7. Washington DC: American Physiology Society, 1975; pp. 319–38. Robaire B, Hermo L. Efferent ducts, epididymis, and vas deferens: structure, functions, and their regulation. In: Knobil E, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 999–1080. Glover TD, Nicander L. Some aspects of structure and function in the mammalian epididymis. J Reprod Fertil Suppl 1971;13:39–50. Cooper TG. The Epididymis, Sperm Maturation and Fertilization. Berlin: Springer-Verlag, 1986. Nicander L. Studies on the regional histology and cytochemistry of the ductus epididymis in stallions, rams and bulls. Acta Morphol Neerl Scand 1957;1:337–62. Barker CAV, Gaudier JCC. Pregnancy in a mare resulting from frozen epididymal spermatozoa. Can J Comp Med Vet Sci 1957;21:47–51. Gebauer MR, Pickett BW, Swierstra EE. Reproductive physiology of the stallion. III. Extra-gonadal transit time and sperm reserves. J Anim Sci 1974;39:737–42. Almquist JO, Hale EB, Amann RP. Sperm production and fertility of dairy bulls at high collection frequencies with varying degrees of sexual preparation. J Dairy Sci 1958;41:733. Hafs HD, Hoyt RS, Bratton RW. Libido, sperm characteristics, sperm output, and fertility of mature dairy bulls ejaculated daily or weekly for thirty-two weeks. J Dairy Sci 1959;42:626–36. Martig RC, Almquist JO. Reproductive capacity of beef bulls. III. Postpubertal changes in fertility and sperm morphology at different ejaculation frequencies. J Anim Sci 1969;28:375–8. Martig RC, Almquist JO, Foster J. Reproductive capacity of beef bulls. V. Fertility and freezability of successive ejaculates collected by different methods. J Anim Sci 1978;30:60–2. Amann RP, Kavanaugh JF, Griel Jr LC, Voglmayr JK. Sperm production of Holstein bulls determined from testicular spermatid reserves, after cannulation of rete testis or vas deferens, and by daily ejaculation. J Dairy Sci 1974;57:93–9. Lino BF, Braden AWH. The output of spermatozoa in rams. I. Relationship with testicular output of spermatozoa and the effect of ejaculations. Aust J Biol Sci 1972;25:351–8. Thompson Jr DL, Pickett BW, Squires EL, Nett TM. Effect of testosterone and estradiol-17β alone and in combination on LH and FSH concentrations in blood serum and pituitary of geldings and in serum after administration of GnRH. Biol Reprod 1979;21:1231–7. Mann T, Lutwak-Mann C. Male Reproductive Function and Semen. Berlin: Springer-Verlag, 1981. Cupps PT. Reproduction in Domestic Animals, 4th edn. New York: Academic Press, 1990. Waites GMH, Setchell BP. Physiology of the testis. In: Lamming GE (ed.) Marshall’s Physiology of Reproduction, 4th edn. London: Churchill Livingstone, 1981; pp. 1–105. Clay CM, Squires EL, Amann RP, Nett TM. Influences of season and artificial photoperiod on stallions: Luteinizing hormone, follicle stimulating hormone and testosterone. J Anim Sci 1988;66:1246–55. Irvine CHG, Alexander SL, Turner JE. Seasonal variation in the feedback of sex steroid hormones on serum LH concentrations in the male horse. J Reprod Fertil 1986;76:221–30.
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84. Thompson Jr DL, Johnson L, St. George RL, Garza Jr F. Concentrations of prolactin, luteinizing hormone and follicle stimulating hormone in pituitary and serum of horses: effect of sex, season and reproductive state. J Anim Sci 1986;63:854–60. 85. Thompson Jr DL, St. George RL, Jones LS, Garza Jr F. Patterns of secretion of luteinizing hormone, follicle stimulating hormone and testosterone in stallions during the summer and winter. J Anim Sci 1985;60:741–8. 86. Clay CM, Squires EL, Amann RP, Nett TM. Influences of season and artificial photoperiod on stallions: Pituitary and testicular responses to exogenous GnRH. J Anim Sci 1988;67:763–70. 87. Irvine CHG, Alexander SL, Hughes JP. Sexual behavior and serum concentrations of reproductive hormones in normal stallions. Theriogenology 1985;23:607–17. 88. Naden J, Amann RP, Squires EL. Testicular growth, hormone concentrations, seminal characteristics and sexual behavior in stallions. J Reprod Fertil 1990;88:167–76. 89. Naden J, Squires EL, Nett TM, Amann RP. Effect of maternal treatment with altrenogest on pituitary response to exogenous GnRH in pubertal stallions. J Reprod Fertil 1990;88:177–83. 90. Squires EL, Todter GE, Berndtson WE, Pickett BW. Effect of ejaculation on systemic levels of testosterone and LH in stallions. In: Proceedings of the Annual Meetings of the American Society of Animal Science, 1977, p. 210. 91. Thompson Jr DL, Pickett BW, Nett TM. Effect of season and artificial photoperiod on levels of estradiol17β and estrone in blood serum of stallions. J Anim Sci 1978;47:184–7. 92. Lacroix A, Gamier D-H, Pelletier J. Temporal fluctuations of plasma LH and testosterone in Charolais bull calves during the first year of life. Ann Biol Anim Biochim Biophys 1977;17:1013–19. 93. McCarthy MS, Convey FM, Hafs HD. Serum hormonal changes and testicular response to LH during puberty in bulls. Biol Reprod 1979;20:1221–7. 94. Bardin CW. Inhibin structure and function in the male. Ann N Y Acad Sci 1989;564:102–23. 95. Pickering BT, Birkett SD, Buldenaar SE, Nicholson HD, Worley RT, Yavachev L. Oxytocin in the testis: what, where and why? Ann N Y Acad Sci 1989;564:198. 96. Knickerbocker JJ, Sawyer HR, Amann RP, Tekpetey FR, Niswender GD. Evidence for the presence of oxytocin in the ovine epididymis. Biol Reprod 1988;39:391–7. 97. Veeramachaneni DNR, Amann RP. Oxytocin in the ovine ductuli efferentes and caput epididymides: immunolocalization and evidence for receptor-mediated endocytosis. Endocrinology 1990;126:1156–64. 98. Garcia Jr F, et al. Active immunization of intact mares against gonadotropin releasing hormone: differential effects on secretion of luteinizing hormone and follicle stimulating hormone. Biol Reprod 1986;35:347–52. 99. Courot M. The effects of gonadotrophins on testicular function (spermatogenesis). In: Proceedings of the International Congress on Animal Reproduction and Artificial Insemination, vol. 5, 1988, pp. 311–19. 100. Bergin WC, Gier HT, Marion GB, Coffman JR. A developmental concept of equine cryptorchism. Biol Reprod 1970;3:82–92. 101. Gier HT, Marion GB. Development of the mammalian testis. In: Johnson AD, Gomes WR, VanDemark NL (eds) The Testis, vol. 1. New York: Academic Press, 1970; pp. 1–45.
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102. Stickle RL, Fessler JF. Retrospective study of 350 cases of equine cryptorchidism. J Am Vet Med Assoc 1978;172:343–6. 103. Amann RP. Endocrine changes associated with the onset of spermatogenesis in Holstein bulls. J Dairy Sci 1983;66:2606–22. 104. Amann RP, Schanbacher BD. Physiology of male reproduction. J Anim Sci 1983;57 (Suppl. 2):380–403. 105. Amann RP, Wise ME, Glass JD, Nett TM. Prepubertal changes in the hypothalamic-pituitary axis of Holstein bulls. Biol Reprod 1986;34:71–80. 106. Robb GW, Amann RP, Killian GJ. Daily sperm production and epididymal sperm reserves of pubertal and adult rats. J Reprod Fertil 1978;54:103–7.
107. Curtis SK, Amann RP. Testicular development and establishment of spermatogenesis in Holstein bulls. J Anim Sci 1981;53:1645–57. 108. Glassneck HW. Histometrische Untersuchungen uber die Entwicklung des Pferdehodens zwischen 1. und 18. Lebensmonat. DMV dissertation, Tierarztliche Hochschule Hannover, 1978. 109. Nishikawa Y. Studies on Reproduction in Horses. Tokyo: Japan Racing Association, 1959. 110. Cornwell JC, Hauer EP, Spillman TE, Vincent CK. Puberty in the Quarter Horse colt. J Anim Sci 1973;36:1215. 111. Tischner M, Kosiniak K, Bielanski W. Analysis of the pattern of ejaculation in stallions. J Reprod Fertil 1974;41:329–35.
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Chapter 97
From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell∗ Dickson D. Varner and Larry Johnson
Introduction The stallion represents one-half of the breeding equation; yet, apportioned time and funding has been considerably less for efforts to advance the discipline of stallion reproduction compared with that of mare reproduction. Modern-day fathers of stallion reproduction, such as Robert M. Kenney, Bill W. Pickett, and Marion Tischner, made significant strides in our understanding of stallion reproductive physiology and clinical disease, but this fascinating area of study begs for individuals with the aptitude and tenacity required to take the discipline to new heights. In recent years, the level of interest in research directed toward the breeding stallion has increased noticeably, as evidenced by the 2006 Congress of the International Society of Equine Reproduction (ISER). The stallion was represented in the largest section of papers presented and published at this meeting, a first since the inception of the ISER in 1975.
This chapter will be directed primarily at the male gamete, the spermatozoon, and all the fascination that it bestows upon those of us who have dedicated a large portion of our professional lives eavesdropping on its microcosm. We will begin with a detailed overview of spermatozoal structure and function and the events that accompany a spermatozoon’s sojourn through the female reproductive tract, followed by a discussion of some potential clinical ramifications. Not all published work could be represented in this chapter, and we preface it with an apology for unintentional exclusion of pertinent data. While we direct readers to some relevant studies conducted in equids, most of the information provided in this chapter stems from investigations conducted in other species, where the process of discovery has been most pronounced. Although this comparative approach can be quite enlightening, a drawback is that the relevance to stallions of findings unveiled in other species will require documentation.
∗ After lengthy pondering about both the subject area and the title of this manuscript, I (Varner) arrived at that shown above. Much to my chagrin, while conducting a literature search during the preparation of this paper, I found that one of my mentors from the University of Pennsylvania, Dr. Bayard T. Storey, authored a paper over a decade ago with a similar title.1 Rather than change the title, I elected to retain it as a tribute to Dr. Storey. Now a Professor Emeritus of Reproductive Biology and Physiology at the University of Pennsylvania, Dr. Storey established himself as an international leader in the area of spermatozoal physiology. With hundreds of masterful manuscripts to his credit and an exhaustive list of trainees that have emerged to become authorities in their own right, Dr. Storey is a scientist and educator of the highest order. As one trainee that spent only a short time in his laboratory, I would like to honor Dr. Storey’s professional accomplishments through this writing. Likewise, many of my colleagues and I received our residency training through the watchful eye and nurturing environment offered by Dr. Robert M. Kenney, University of Pennsylvania. Dr. Kenney has not only touched our lives in such meaningful fashion, but opened his laboratory and offered his wisdom to virtually anyone, in any part of the world, who sought to gain a more thorough understanding of equine reproduction. His personable nature, combined with an encyclopedic knowledge base, is praised by all the individuals that he has touched. We thank you both for your diligence in the field, your tenacity for intellectual growth, and your amiable approach to the education of others! Reprinted with permission of the American Association of Equine Practitioners (Proceedings, Annual Convention of the American Association of Equine Practitioners, Orlando, Florida, 1997, pp. 104–177).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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A chapter directed exclusively at the spermatozoon would appear to represent a minuscule part of the total reproduction picture. Upon close scrutiny, however, one becomes enlightened about the level of scientific interest in this cell. As an example, approximately 50 000 entries are logged in response to a query for the keywords ‘spermatozoa’ or ‘spermatozoon’ in PubMed, a database of indexed citations offered by the National Institutes of Health. An uninformed individual might describe a spermatozoon as a highly specialized, but simple, cell with only one role to fulfill – that of fertilization. While fertilization is the end point of spermatozoal function, this cell must be extremely sophisticated and adaptable to achieve this task, and the process involves a series of highly coordinated cellular and molecular events. The following is a list of some requirements ascribed to a mammalian spermatozoon.
1. Loss of most organelles and cytoplasm during formation in the testis and maturation in the epididymis (requires a host of intracellular and intercellular signaling events). 2. Remodeling of spermatozoal chromatin within the epididymis as a protective mechanism against environmental injury (requires repackaging of nuclear DNA into a highly condensed form through the aid of specialized proteins termed protamines). 3. Plasma membrane alterations within the epididymis to yield proteins important to fertilization (requires various enzyme-linked alterations of existing proteins, as well as uptake of proteins from epididymal fluid or from the epididymal epithelium). 4. Passage through the uterus and utero-tubal junction (UTJ) of the female at the time of insemination (requires activated flagellar movements and protection against immunological attack). 5. Binding to oviductal epithelial cells to form a spermatozoal reservoir (requires specific cell–cell attachment, possibly mediated through spermatozoal surface carbohydrate-binding proteins, termed lectins). 6. Acquisition of additional maturational changes, collectively termed capacitation, that permit a spermatozoon to fertilize an oocyte (requires an assortment of signal transduction cascades). 7. Release from oviductal epithelial cells and passage to the vicinity of the oocyte at the isthmic– ampullar junction of the oviduct (requires a coordinated spermatozoa-release mechanism, hyperactivated motility, and probably chemotaxis). 8. Penetration through the extracellular matrix of the oocyte cumulus (possibly mediated by hyperactivated motility and redistribution/unmasking
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of surface-associated hyaluronidase, as the cumulus matrix is rich in hyaluronic acid). Binding to the zona pellucida, a highly glycosylated protein matrix surrounding the oocyte (appears to involve specific affinity between spermatozoal surface molecules and the zona pellucida components). Acquisition of the acrosome reaction, a regulated form of exocytosis (requires reorganization of the outer acrosomal and overlying plasma membranes necessary for fusion and vesiculation). Penetration of the zona pellucida (release of acrosomal proteins with enzymatic activity is required for this event to occur). Binding and fusion with the oolemma (requires specific region-dependent molecular interactions). Dispersion of nuclear contents (requires specific fusogenic alterations of the lipid membranes of the spermatozoon and oocyte). Oocyte activation (a spermatozoon-derived factor is required for activation of the oocyte and embryonic development). Pronucleus formation (requires decondensation of the highly compact spermatozoal nucleus). Organization of the mitotic spindle after pronuclear formation (requires contribution of a proximal centriole from the spermatozoon).
After viewing this lengthy list of functions, one becomes quite appreciative of the highly complex and specialized features of a spermatozoon. In fact, the biochemical and biophysical features are so sophisticated that many of the cellular and molecular mechanisms remain unresolved to this day. Furthermore, spermatozoa (and oocytes) represent some of the most highly differentiated cells in the mammalian body; yet, when sperm and oocyte are combined, they retain their potential for totipotency (ability to divide and produce all cell types of the body) through creation of a zygote.
Origin of the spermatozoon The life of a spermatozoon begins within the testes, unless, of course, one wishes to consider the embryonic origin of the primordial germ cells. The testis, an elaborately designed organ, is classically considered to possess two functions: (i) exocrine – spermatogenesis and (ii) endocrine – production of hormones important to spermatogenesis, sexual differentiation, development of secondary sex characteristics, and libido. While this simplistic description provides one with the general concept of testicular function, it does not portray the extremely complex nature and elegant interplay of these two processes. Conventional descriptions convey the role of hypothalamic- and
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell pituitary-derived hormones on regulation of testicular function, as well as feedback mechanisms required for homeostasis. While such pathways are undoubtedly the key orchestrators of testicular function, emerging information is revealing a multitude of subcellular, molecular-mediated events that ‘cloud’ our understanding of the events that actually occur within the testes. As with any area of study, the more learned we become about a topic, the more queries surface that require additional clarification. Such is the case with testicular function. Without question, a thorough understanding of testicular function will require a keen appreciation of the mechanisms by which genes and gene products are expressed and repressed.2–8 As an example of the genetic complexity surrounding control of testicular function, new information has revealed that single nucleotide polymorphisms (SNPs) have been identified in the follicle-stimulating hormone (FSH) receptor gene of humans, resulting in a mutation of the FSH receptor (as is found on the surface of the Sertoli cell) that can influence its activity.9 As another example, use of transgenic mice deficient in estrogen receptor genes has demonstrated that estrogens are likely to play a more important role in testicular function than was once thought to be the case.10 Similarly, use of mice with a selective androgen receptor knockout in Sertoli cells revealed that the androgen receptor in the Sertoli cell is an absolute requirement for normal spermatogenesis.11, 12 Furthermore, experimentation with germ cell-specific androgen receptor knockout mice revealed normal spermatogenesis, suggesting that germ cell androgen receptors may play different roles as the germ cells progress through spermatogenesis.13 As seen here, to more fully understand the processes that take place in the testes, we must capitalize on the powerful molecular tools that have been developed in this capacity. Most of these types of studies are conducted in human and laboratory specimens, so it will be important to test the relevance to the horse. Organization of the testis The testes are composed of testicular parenchyma encapsulated by the thick fibrous tunica albuginea. Extensions of the tunica albuginea penetrate the underlying testicular parenchyma, dividing it into numerous lobules (Figures 97.1 and 97.2).14 The tunica albuginea is composed of collagen and elastic fibers, myoid cells, and a network of blood vessels (Figure 97.3).15–17 Testicular parenchyma occupies nearly 90% of the total testicular mass in adult horses,16 and over 70% of the testicular parenchyma is occupied by seminiferous tubules.18 The interstitial component of the testis is located between seminiferous tubules and consists primarily of the Leydig cells intermin-
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(b) Figure 97.1 Drawings of the left equine testis with attached epididymis and spermatic cord in lateral view (a) and medial view (b). The vascular pattern over the surface of the testis is demonstrated. The testis is normally oriented in the scrotum with the long axis directed horizontally. The epididymis courses over the dorsolateral aspect of the testis. a, Cranial pole of testis (extremitas capitata); b, caudal pole of testis (extremitas caudata); c, appendix testis [vestigial remnant of the embryological mullerian (paramesonephric) duct]; d, epididymal border of the testis, showing the entrance to the testicular bursa; e, proper ligament of the testis; i, ligament of the tail of the epididymis; g, reflected parietal tunic (tunica vaginalis parietalis); h, ductus deferens.14
gled with fibroblasts, lymphocytes, mast cells, blood vessels, lymphatic vessels, and extracellular matrix (Figures 97.4–97.10).17–19 Nerves are rarely seen in the testicular interstitium,17 but they are well established in the area of the spermatic cord leading to the testis. Spermatogenesis Spermatozoal production, i.e., spermatogenesis, occurs within the seminiferous tubules. Both ends of these highly coiled tubules open directly into the rete
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Figure 97.2 Schematic representation of the testis and associated epididymis. The thick tunica albuginea (a) covers the surface of the testis and also extends into the testicular parenchyma, thereby separating the parenchyma into lobules that contain both seminiferous tubules (b) and interstitial tissue. The luminae of the seminiferous tubules empty into the interconnecting tubules of the rete testis (f). The tubules of the extratesticular portion of the rete testis connect to the epididymis via several efferent ducts (c) on the dorsocranial aspect of the testis. The epididymis forms a single duct that spans up to 45 meters in length, and consists of an initial segment (d), a caput (or head; e), a corpus (or body; g), and a cauda (or tail; h), which, in turn, connects with the ductus deferens (i).
testis, such that the products (both cellular and noncellular) of the seminiferous tubules are excreted into the rete testis and delivered to the excurrent duct system (e.g., the epididymis and ductus deferens). The numerous and tortuous seminiferous tubules, with a combined average length of 2419 meters per testis in the stallion,20 are composed of an epithelial wall, termed the seminiferous or germinal epithelium, and a lumen. The seminiferous epithelium consists of germ cells in various steps of development intermingled with Sertoli cells that serve to provide structural support and a nurturing source to the germ cells.17 The Sertoli cells are anchored to the basement membrane, and extend to the lumen, of the seminiferous tubules. The seminiferous tubules are bordered by peritubular myoid cells (myofibroblasts) that, through peristaltic contractions, may aid in evacuation of luminal contents into the rete testis. These myoid cells are also considered to be involved in paracrine signaling events.21–23 A critical feature of the seminiferous epithelium is the formation of tight junctional complexes that develop between Sertoli cells, thereby physically dividing the seminiferous epithelium into basal and adlu-
(c) Figure 97.3 The equine testis viewed in a gross cross-section (a), in a low-magnification histological section (b), and a higher magnification histological section (c). (a) The testis is composed of the tunica albuginea (capsule; TA) and testicular parenchyma (TP). The central vein (CV) has associated connective tissue. (b) The testicular parenchyma contains numerous tightly coiled seminiferous tubules (ST) which are composed of Sertoli cells (SC), and a variety of germ cells, including spermatogonia (Sg), primary spermatocytes (P), round spermatids (Sa), elongated spermatids (Sd2) and residual bodies (RB). (c) The interstitium between seminiferous tubules (ST) contains numerous Leydig cells (LC), blood vessels (BV), myoid cells (MC) and lymphatic vessels (LV). Scale bars, 10 mm in (a), 1 mm in (b), and 30 µm in (c).15–17
minal compartments (Figure 97.11).17 This structural complex is termed the blood–testis barrier because it restricts direct access of bloodborne substances into the adluminal compartment. The described actions of the blood–testis barrier are (i) to segregate meiotic (except the earliest preleptotene spermatocytes) and post-meiotic germ cells from immunological attack, as these germ cells are considered to be in an immunologically privileged site, and (ii) to provide
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Figure 97.4 Micrograph of cross-sections of equine seminiferous tubules and interstitial tissue, after toluidine blue staining of 0.5-µm Epon section. Stages I, III, VI, and VII of the seminiferous epithelial cycle are depicted, and illustrate respective groupings of spermatogonia (A, B), primary spermatocytes (L, Z, P), and spermatids (Sa, Sb1 , Sc, Sd1 , Sd2 ) that are intimately associated with Sertoli cells (SC) and contained by myoid cells (MC) marking the outer limits of the seminiferous tubules. The interstitium between tubules is characterized by robust Leydig cells (LC), lymphatic ducts (LD), and blood capillaries (BC). Scale bar, 10 µm.
a unique microenvironment for the final stages of germ cell development within the adluminal compartment.24 Although several blood barriers exist in the general body, there is no counterpart to the blood–testis barrier in the female. In the female, development of primary oocytes occurs prior to the immune system recognizing self, and the female does develop the counterpart to spermatids (i.e., ootids). In the male, germ cell development to spermatocytes and spermatids does not occur
Figure 97.6 Transmission electron micrograph of equine Leydig cells found in testicular interstitium. Equine Leydig cells have an abundance of cytoplasm containing smooth endoplasmic reticulum (SER) and a large number of mitochondria (M). Gap junctions (GJ) for intercellular communication are located in the plasma membranes of two adjacent cells. The nucleus (N) is spherical and largely euchromatic, and has a distinct nucleolus (No). Scale bar, 3 µm.18
until puberty. This is well after the immune system recognizes self and considers these cells as foreign. Especially intriguing is the molecular control over the transient disassembly of the blood–testis barrier to facilitate migration of germ cells from the basal into the adluminal compartment.24, 25 Spermatogenesis is an extremely complex process that involves germ cell proliferation, germ cell differentiation, and, paradoxically, programmed germ cell death (termed apoptosis). This lengthy process, 57 days in the stallion,20, 26 is controlled by a vast array of messengers acting through endocrine, paracrine, and autocrine pathways.27–31 Spermatogenesis not only involves transformation of undifferentiated diploid germ cells into highly differentiated and specialized haploid spermatozoa (Figures 97.12 and 97.13),17, 26 but it also involves profound transcriptional modifications within the cells.3, 32 As a partial list, the finished product of spermatogenesis has (i) a 1× chromosomal complement that is profoundly repackaged, (ii) a newly formed and intricately designed flagellum, (iii) the
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Figure 97.5 Scanning electron micrograph of the equine testis revealing seminiferous tubules (ST) and Leydig cells (LC). Tails of developing spermatids (T) can be seen projecting into the lumen of seminiferous tubules. Scale bar, 50 µm.19
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Figure 97.7 The effect of stallion age on pigmentation of unfixed equine testicular parenchyma, as observed grossly. (a) Light parenchyma is from a post-pubertal stallion (2–3 years old). (b) Moderately dark parenchyma is characteristic of adult (4–5 years old) stallions. (c) Dark parenchyma characterizes aged stallions (13–20 years old).18
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Figure 97.8 Composite of the interstitium between seminiferous tubules (ST) and Leydig cells (LC) in stallions at different ages in the middle (a, c, e) or onset (b, d, f) of the breeding season. (a, b) The post-pubertal (2–3 years old) stallion has large Leydig cell clusters in the interstitium. (c, d) The young adult stallion (4–5 years old) has larger clusters of Leydig cells that increase the density. (e, f) The aged stallion testis (13–20 years old) has mostly Leydig cells in its interstitium; however, some Leydig cells may contain a large accumulation of lipofuscin (Lf). No difference in Leydig cell structure is noted between the onset and middle of the breeding season. Scale bars, 75 µm in (a, c, e) and 25 µm in (b, d, f).18
biogenesis of a highly complex secretory vesicle, the acrosome,33 and (iv) retention of some mRNA (in a largely depleted cytoplasmic package) that likely has implications in fertilization and post-fertilization events.32, 34, 35 Spermatogenesis is initiated by the differentiation of spermatogonia from a stem cell pool that is continually replenished for most of a stallion’s adult life. Under normal circumstances, this is a highly productive process, yielding of the order of 5–6 billion spermatozoa per day for an adult stallion. This translates into 30–40 trillion spermatozoa produced over the course of a stallion’s life. From an equally impressive view, an average healthy stallion produces 60 000–70 000 spermatozoa per second. These newly formed spermatogonia enter a proliferative phase, whereby continuous mitotic amplifications yield a
dramatic increase in spermatogonial numbers. Interestingly, the cytoplasmic component of mitotic divisions is often incomplete, resulting in daughter cells that remain connected by intracytoplasmic bridges (Figure 97.14).36 Such an arrangement permits direct communication within this syncytium of developing germ cells, thereby assisting in their synchronous development. Following completion of spermatocytogenesis, the spermatozoa enter a meiotic phase, characterized by duplication and exchange of genetic information (i.e., genetic recombination) and two meiotic divisions which reduce the chromosome complement to form haploid round spermatids. It is during the meiotic stage that germ cells pass through the blood–testis barrier to enter the adluminal compartment. During the final phase of development, spermiogenesis, the round spermatids undergo
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(b) Figure 97.9 Transmission electron micrograph of Leydig cells in young adult (4–5 years old; a) and aged stallions (13–20 years old; b). In both cases, Leydig cells contain an abundance of smooth endoplasmic reticulum (SER) and mitochondria (M). In aged stallions (b), Leydig cells may have an accumulation of lipofuscin (Lf). Scale bar, 1 µm.18
Figure 97.11 Transmission electron micrograph of Sertoli cells and germ cells and Sertoli cell–Sertoli cell junctional complexes (inset). Junctional complexes (JC) between adjacent Sertoli cells (SC) separate the basal compartment that contains spermatogonia (Sg) from the adluminal compartment that contains primary spermatocytes (PS) and spermatids (St). Junctional complexes between adjacent Sertoli cells in the stallion are similar to those of other mammals. These include fusion (F) of outer leaflets of the plasma membranes of two Sertoli cells and the presence of juxtaposed endoplasmic reticulum (ER) and bundles of fine filaments (BF). Scale bars, 1 µm and 2 µm (inset).17
a dramatic transformation that includes nuclear reshaping through chromatin compaction, creation of a flagellum, development of the acrosome, and considerable loss of cytoplasm (Figures 97.13 and 97.14). The fully developed spermatids are released as spermatozoa into the lumen of the seminiferous tubules by a process termed spermiation (Figure 97.15).19, 37 It is during spermiogenesis that the germ cells may be most vulnerable to both structural and genetic defects.34 Cellular associations during spermatogenesis
Figure 97.10 Scanning electron micrograph of the equine testicular interstitium and seminiferous tubule. Clusters of Leydig cells (LC) are seen in the interstitium between seminiferous tubules (ST). One seminiferous tubule contains spermatids (Spermatids) with spherical nuclei and spermatids whose tails already project into the lumen. Spermatozoa (Sperm) in transit are seen in the lumen. The open arrows mark the height of the seminiferous epithelium. Scale bar, 20 µm.17
The spermatogenic cycle, or cycle of the seminiferous epithelium, represents the series of changes in a given region of a seminiferous tubule between two appearances of the same developmental stages.38–41 For instance, if spermiation were used as a reference point, the cycle would consist of all the cellular associations occurring within a given cross-section of a seminiferous tubule between two consecutive spermiations (Figure 97.16). These cross-sectional cellular associations can be divided into eight distinct stages (Figures 97.16–97.19).19, 42, 43 The spermatogenic cycle length is constant at 12.2 days in the stallion.20, 26
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Figure 97.12 Equine spermatogonia and spermatocytes viewed by bright-field microscopy of 5-µm methacrylate sections stained with toluidine blue. Small arrows designate cells of interest. The most primitive spermatogonia have small, oval, or flattened nuclei (A1 ). Other subtypes include those with a light center (A2 ), a large single nucleolus (A3 ), two or three nucleoli (A3 ), or large nucleoli plus fragmented nucleoli (B1 ), or with a small spherical nucleus (B2 ). Preleptotene (Pl), leptotene (L), zygotene (Z), early pachytene (EP), late pachytene (LP), diplotene primary spermatocytes (D; open arrow), and secondary spermatocytes (SS; closed arrow) are indicated. Scale bar, 10 µm.17, 26
The spermatogenic wave, on the other hand, refers to the spatial, sequential order of stages along the length of a seminiferous tubule at a given point in time (Figure 97.20).42 These two characteristics of the seminiferous epithelium can be defined because of the synchronous nature of development for cohorts of differentiating germ cells along the length of the seminiferous tubules. Histological evaluation of the stages of the seminiferous epithelial cycle permits one to assess the efficiency of spermatogenesis.39, 44 Germ cell degeneration Germ cell degeneration can be amplified in stallions with abnormal testicular function, but it is also a normal phenomenon in spermatogenesis.45 In fact, ‘normal’ spermatogenesis is a relatively inefficient process, and has been reported to result in an estimated loss of 25–75% of the potential number of spermatozoa produced by spermiation in the rat.46, 47 In the stallion, germ cell degeneration is more profound during the physiological breeding season than during the non-breeding season.41 Developing spermatogonia are the most vulnerable to apoptotic degeneration (Figure 97.21).43 It is possible that the ‘physiological’ form of germ cell degeneration is a homeostatic mechanism that prevents
overloading the Sertoli cells, i.e., to maintain a fine balance in the germ cell–Sertoli cell ratio. Apoptosis is a well-defined physiological process of cell elimination,48 and the apoptotic process is required for normal spermatogenesis in mammals.49, 50 A protective role of luteinizing hormone (LH) against germ cell apoptosis has been reported in rats, based on expression of various apoptotic genes following immunoneutralization of LH in germ cells.51 Apoptosis of spermatogonia and spermatocytes has been reported in normal stallion testes, a finding that is consistent with the expected time that germ cells may be susceptible to removal from the system.43 Of interest, formation of the developing seminiferous tubules and the Sertoli cell (i.e., blood–testis) barrier coincides with increased germ cell apoptotic rates in stallions, providing evidence that apoptosis may play an intricate role in initiation of spermatogenesis.52 As expected, these changes are coincident with gene expression patterns.53 Studies involving rats indicate that FSH is an important regulator of this event.54, 55
Epididymal transit and maturation Mammalian spermatozoa are incapable of in vivo fertilization upon exiting the testis. The cells must undergo considerable post-testicular remodeling
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Figure 97.13 Transmission electron micrographs of equine spermatids representing different steps of development during spermiogenesis. (a–e) The Sa spermatids (Sa) are characterized by a large Golgi apparatus (GA), which produced vesicles (V) that fuse to form larger vesicles and ultimately to form the acrosomic vesicle (AV). The acrosomic vesicle indents the nuclear envelope and then flattens over the nucleus. (b) Developing flagella (F) from neighboring Sa spermatids have extended away from the spermatid cell body into the extracellular space between the Sertoli cell and spermatid. (c) Cytoplasmic bridges connect adjacent spermatids. (f) In the Sb1 spermatids (Sb1 ), the acrosome had formed a head cap over the nucleus. (g, h) The Sc spermatid (Sc) has a distinct manchette (Mn), elongating and condensing nucleus, attached flagellum with distinct annulus (An), and well-defined acrosome (A) over the anterior portion of the nucleus. (i, j) The Sd spermatids (Sd1 and Sd2 ) have lost their manchette and have further condensation of the nucleus. (b) The formation of the flagellum begins in the early Sa spermatid, (g, h) appears as a growing axoneme in Sc spermatids, but (i) develops outer dense fibers (DF), the fibrous sheath (FS), and has mitochondrial migration around the flagellum in early Sd1 spermatids. (j) Late Sd2 spermatids are mostly extended into the lumen (TL), and have a completely formed flagellum, with mitochondria (M) surrounding the midpiece, and a distinct fibrous sheath (FS). Excess cytoplasm of the spermatids remains in the proximal region of the midpiece as a cytoplasmic droplet (CD). Scale bar, 2 µm.17, 26
within the epididymis to acquire this ability. Each epididymis is an unbranched tortuous duct that spans 70–80 meters in the stallion.56 The epididymis is connected to the rete testis by several efferent ducts (Figure 97.2). The anatomical features of the rete testis and efferent ductules have been well characterized in the stallion (Figures 97.22–97.24).15, 57 For descriptive purposes, each epididymis is typically divided anatomically into a caput (or head), a corpus
(or body), and a cauda (or tail) (Figure 97.2). The caput is closely applied to the cranial pole of the testis, the corpus courses over the dorsolateral surface of the testis, and the bulbous cauda is attached loosely to the caudal pole of the testis via the proper ligament of the testis (Figure 97.1). An initial segment and an intermediate zone have also been ascribed to the beginning portion of the epididymal duct within the caput.58
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Figure 97.14 High-voltage electron micrographs of equine Sertoli and germ cells embedded in Epon 812 and sectioned at 1.0 µm. The basal (a) and apical (b) regions of the Sertoli cells are characterized. (a) The close association between Sertoli cells and germ cells representing different developmental steps is evident. Laminar processes (LP) extend from the main body of a Sertoli cell and can be seen between germ cells. One forms a rim of Sertoli cell cytoplasm from the main body of the Sertoli cell to the site of an intercellular bridge (IB) between two primary spermatocytes (PS). Long slender mitochondria (arrows) oriented with the long axis of the Sertoli cell characterize the main body cytoplasm, while numerous dense lipid and lipofuscin granules are found in the basal cytoplasm. Mitochondria (M) of germ cells are contrasted with those of Sertoli cells. Rough endoplasmic reticulum (RER) can be seen in the cytoplasm of germ cells. Portions of Sertoli cell nuclei (N) depict the highly indented nature of the nuclear surface. An A spermatogonium shares the intercellular space (IS) with a Sertoli cell. While Golgi-phase spermatids (Sa) have not yet developed an acrosomic vesicle, the developing flagellum (F) from the centriole (Ct) can be seen within the cytoplasm of one such spermatid. (b) Apical region of a Sertoli cell containing long slender mitochondria (arrows) and four embedded Sd1 spermatids with elongated nuclei are shown in this profile. A portion of the nucleus and three mitochondria of a Sa spermatid (Sa) are present. By following Sertoli cell cytoplasm marked by intercellular space (IS) between Sd1 spermatids and the Sertoli cell, it is evident that Sertoli cell cytoplasm is located between each spermatid. These Sd1 spermatids are deeply embedded in the apex of the Sertoli cell via tubular indentations in the Sertoli cell plasma membrane. The annulus (An), manchette (Mn), and mitochondria (M) of the Sd1 spermatids are indicated. The flagellar canal (FC) is located between the developing flagellum and surrounding manchette. The manchette is situated to provide structural support to the flagellar canal. This canal is lined by the plasma membrane enfolded from the surface of the spermatid at the lumen into which the flagellum extends. Scale bars, 2 µm (a) and 1 µm (b).36
On average, 90 billion spermatozoa reside in the excurrent ducts of reproductively mature stallions when sexually rested, with a lower number (60 billion) reported for 2- to 4-year-old stallions.59 On average, 62% of the spermatozoa within the excurrent duct system are in the caudae epididymides, and 7% are in the ductus deferens.59 Based on radiolabeled studies, the average time required for spermatozoal transit through the epididymis of the stallion is reported to be 8–11 days.60 Spermatozoal transit time through the caput and corpus segments is constant at 4 days and is unaffected by frequency of ejaculation, age, or season,60–62 whereas transit time through the cauda epididymis can be accelerated by increased ejaculation frequency.61, 63 Spermatozoal number in the caudae epididymides and ductus deferens (i.e., spermatozoa available for ejaculation) may be reduced by approximately 30% when semen
is collected once every 2 days, as compared with sexually rested stallions.59 The normal spermatozoal transit time through the cauda epididymis of the stallion ranges from 4 to 7 days. Spermatozoa enter the epididymis at a constant rate in reproductively normal stallions (average of over 5 billion spermatozoa per day). Spermatozoal phagocytosis within the epididymis is considered to be rare,58, 63 so spermatozoa must also exit the epididymis at a similar rate to prevent epididymal overload. The length of time that spermatozoa can remain in the cauda epididymis without suffering storage-related injury has not been critically studied. Under normal conditions in sexually rested stallions, rhythmic contractions of the smooth musculature in the caudae epididymides and deferent ducts are thought to result in spontaneous emission of spermatozoa into the urethra. Some stallions are known to accumulate excessive numbers of
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Figure 97.15 Scanning electron micrograph of stage VII (a), early stage VIII (b) and late stage VIII tubules (c,d). Spermatid tails project into the lumen in stages VII and VIII, but the entire spermatid projects into the lumen in late stage VIII, attached only by cytoplasmic stalks (CS). (c,d) An enlarged luminal view of (c) is seen in (d). Maturation-phase spermatids are located in the lumen; however, tails (T) of Sa spermatids with round nuclei already project into the lumen. One luminal spermatozoon, recently released, has its characteristic cytoplasmic droplet (CD) in the proximal position where the midpiece attaches to the head. Scale bars, 10 µm (a,c), 5 µm (b), and 2.6 µm (d).19
spermatozoa in the caudae epididymides, deferent ducts, and ampullar luminae (the ampullae surround the terminal segment of the ductus deferens). This condition, termed ‘spermiostasis’64 or ‘plugged ampullae’,65 is thought to be associated with an intrinsic dysfunction in contractility of the smooth musculature surrounding the cauda epididymis and ductus deferens. It leads to a marked reduction in spermatozoal quality, or azoospermia if the ductal luminae become obstructed completely.14 A separate
subset of stallions is known to experience reduced semen quality and fertility if sexually rested for more than 12–24 hours (Varner D and Love C, unpublished observations, 2006). In this instance, the microenvironment within the cauda epididymis may impart factors that impede spermatozoal survival rather than favoring maximal survival time. As epididymal spermatozoa are known to be susceptible to oxidative injury,66, 67 it is possible that reduced elimination of potentially harmful agents or reduced synthesis
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12.2 DAYS Figure 97.16 The kinetics of equine spermatogenesis and the stages of the cycle of the seminiferous epithelium. Three major divisions of spermatogenesis (spermatocytogenesis, meiosis, and spermiogenesis) combine to constitute the eight stages of the cycle of the equine seminiferous epithelium (I–VIII). During the 19 days of spermatocytogenesis, A1 spermatogonia enter cyclic activity during stage III and undergo mitotic division to produce B1 and B2 spermatogonia, followed by production of preleptotene primary spermatocytes (Pl). During the 20 days of meiosis, preleptotene primary spermatocytes differentiate through zygotene meiotic division to produce Sa spermatids (Sa). During the 18 days of spermiogenesis, Sa spermatids differentiate through Sb1 , Sb2 , Sc, Sd1 , and Sd2 steps of development before spermiation (i.e., release from the seminiferous epithelium) as spermatozoa. While the letters indicate the development step, the numbers associated with each germ cell step indicate the developmental age (in days) of each cell type in the middle of each spermatogenic stage. The cycle length is 12.2 days, and the duration of spermatogenesis is 57 days in stallions.
and secretion of antioxidants contributes to this disorder.58 Alternatively, spermatozoal membranes which are overly abundant in polyunsaturated fatty acids may be at increased risk of oxidative injury.67 As is the case with many other species studied, stallion spermatozoa are immotile upon entering the epididymis and do not gain motility until reaching the distal corpus. A motility pattern consistent with ejaculated spermatozoa is not acquired until spermatozoa reach the cauda epididymidis.68 As such, movement of spermatozoa through the epididymis is primarily attributed to rhythmic contractions of the smooth musculature that surrounds the epididymal duct (Figures 97.25–97.27).57, 58 In men, both the thickness of the smooth musculature and its adrenergic innervation increase from the proximal to the distal end of the epididymis, and into the ductus deferens.69 Autonomic drugs (both cholinergic and adrenergic) increase contractility of all segments of the epididymis.70 Prostaglandins (PGF2α and PGE) have been isolated from the testes and excurrent ducts of male rats, with a concentration
noted to be lowest in the testes, intermediate in the epididymides, and highest in the ductus deferens.71 These same workers demonstrated that aspirin, which suppresses production of PGF2α , strongly inhibited contractility of the caput epididymidis in vitro. Oxytocin receptors are also present in the epididymal epithelium,72, 73 and the frequency of caput epididymal contractions increases, and tubule diameter decreases, in a dose-responsive manner to oxytocin exposure.74 While the epididymis in considered to be an androgen-dependent tissue,75, 76 estrogens appear to play a more important role than androgens in promotion of epididymal motility. This is achieved through estrogen-mediated upregulation of both the oxytocin receptor gene and the receptor protein.77 In addition to serving as a conduit, the epididymis bestows upon spermatozoa specific structural and physiological alterations, the result of which yields acquisition of fertilizing potential. Maturational changes in spermatozoa occur predominantly within the caput and corpus, as spermatozoa recovered from the cauda have attained a fertilizing
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Figure 97.17 Bright-field micrographs depicting the eight stages of the cycle of the seminiferous epithelium in paraffin sections stained with periodic acid–Schiff and toluidine blue. Stage I, characterized by two generations of leptotene (L) and pachytene (P) primary spermatocytes. Sb1 round spermatids can be observed with a well-developed and easily recognizable acrosomal cap. Developing tails are sometimes visible, although these structures can be difficult to distinguish in paraffin sections. Stage II, two generations of L and P primary spermatocytes. Sb2 spermatids become irregularly shaped as elongation and orientation begins. Acrosomal caps are plainly evident. Stage III, two generations of zygotene (Z) and pachytene spermatocytes are evident, as are characteristic Sc spermatids which are oriented with acrosomal caps toward the basement membrane. No primary spermatocytes with meiotic figures are evident. Stage IV, contains the most germ cell types of any state. B1 spermatogonia are sometimes evident (not shown) along the basement membrane next to zygotene primary spermatocytes. In the adluminal compartment, large pachytene spermatocytes begin their divisions and are seen with numerous meiotic figures (MF; metaphases I and II). Secondary spermatocytes (SS), newly formed Sa round spermatids, and Sc elongated spermatids are also evident. Stage V, B1 spermatogonia are still evident. Only one generation of pachytene primary spermatocytes may be found. The newly formed Sa round spermatids have not yet begun formation of the acrosomal cap over the nucleus. Sd1 elongated spermatids with densely packed chromatin may be found stacked in clusters, deeply embedded within the seminiferous epithelium. Stage VI, small, round B2 spermatogonia with small clumps of heterochromatin have formed and one generation of pachytene spermatocytes may be found near the basement membrane. Sa round spermatids may be observed with an acrosomic vesicle not yet touching the nucleus. Sd1 elongated spermatids begin their migration toward the lumen. Stage VII, B2 spermatogonia and one generation of pachytene primary spermatocytes may be found. Acrosomal caps of Sa round spermatids begin to flatten and cover the surface of the nucleus. Densely packed Sd2 elongated spermatids are migrating closer to the lumen. Stage VIII, B2 spermatogonia have divided to form small round preleptotene (Pl) primary spermatocytes along the basement membrane. One generation of pachytene primary spermatocytes and late Sa round spermatids with developing acrosomal caps can be found. Residual bodies (Rb) accumulate as mature spermatozoa are being released into the lumen as spermiation occurs. Scale bar, 30 µm.43
capacity similar to that of ejaculated spermatozoa for most species studied.58, 78, 79 The cauda epididymis is considered to be a storage reservoir for spermatozoa, but this capacity extends into the ductus deferens and ampullae. In one study, sufficient spermatozoa were
present in the ductus deferens and ampullae of stallions to account for average spermatozoal numbers in an ejaculate.62 A paucity of information exists with regard to the specific mechanisms that control spermatozoal
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Figure 97.18 Bright-field light (a, c, e) and scanning electron (b, d, f) micrographs of different stages of the equine spermatogenic cycle. (a, b) Stages I and II are illustrated. (c, d) Stages III, IV and V are represented. (e, f) Stages VI and VII reveal changes in preparation of the release of spermatozoa in stage VIII. Scale bar, 20 µm.19
maturation. Studies to date indicate that a salient feature of the epididymis is a continual change in the microenvironment to which spermatozoa are exposed during epididymal transit. This is portrayed by the variation in histomorphological characteristics of the numerous epididymal cell types that line the various regions of the duct (Figures 97.24–97.27),57, 58, 80 as well as the distinct regional and cell-specific patterns of gene expression and product secretion.81–84 Although a variety of such localization studies have
been conducted, these must be followed by mechanistic and functional approaches before we can fully appreciate the molecular interactions that impart fertilizing power to a spermatozoon.85–87 The unique composition of epididymal plasma can be attributed, in large part, to the formation of a blood–epididymis barrier.88, 89 This barrier is formed by tight junctional complexes near the luminal border between adjacent principal cells of the epididymal epithelium.90 This mechanism restricts entry of
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Figure 97.19 A clockwise arrangement of eight stages of the cycle of the equine seminiferous epithelium observed by Nomarski optics in unstained, 20-µm Epon histological sections. Both nuclear and cytoplasmic details are revealed in Leydig cells (LC), Sertoli cells (SC), various types of germ cells, and myoid cells (MC). A spermatogonia (A) of different types and Sertoli cells are found in all stages of the cycle. Classification of spermatid development was based on that described for humans and used in the horse. Briefly, the Sa spermatid is the earliest form of spermatid, as it contains a spherical nucleus and a large Golgi with either no acrosomic vesicle, a developing acrosomic vesicle, or an acrosomal cap. The Sb1 spermatid has a spherical nucleus but also has an attached flagellum and a distinct acrosome covering half of the nuclear surface. The Sb2 spermatid has begun nuclear elongation with the appearance of the manchette. The Sc spermatid has a distinct manchette and a more elongated nucleus than the Sb1 spermatid. The Sd1 spermatid is undergoing final maturation with the disappearance of the manchette and migration of mitochondria around the tail. The Sd2 spermatid is the final form with a large portion of the cell, including the head, projecting into the tubular lumen in stage VIII. Stage I (I) is characterized by preleptotene or leptotene (L) and pachytene (P) primary spermatocytes as well as Sb1 spermatids (Sb1 ). These spermatids are characterized by spherical nuclei, attached acrosomal caps (AC), and developing flagellum (F). Stage II (II) is characterized by leptotene primary spermatocytes (L), pachytene primary spermatocytes (P), and Sb2 spermatids (Sb2 ). The labeled pachytene primary spermatocyte is displaying its large spherical Golgi apparatus. The Sb2 spermatid has an elongating nucleus, manchette (Mn), and annulus (An). Stage III (III) has zygotene (Z) and pachytene (P) primary spermatocytes, and bundles of Sc spermatids (Sc). The Sc spermatid has an elongating and condensing nucleus, distinct manchette (Mn), and annulus (An). Stage IV (IV) is characterized by the presence of zygotene primary spermatocytes (Z), diplotene primary spermatocytes or secondary spermatocytes (SS), meiotic figures (MF), and Sc spermatids (Sc). Stage V (V) is composed of pachytene primary spermatocytes (P), newly formed Sa spermatids (Sa), and Sd1 spermatids (Sd1 ). The Sa spermatid has a distinct Golgi apparatus (GA), developing acrosomic vesicle (AV), and chromatoid body (CB); and Sd1 spermatids form bundles which are deeply embedded in the seminiferous epithelium. Stage VI (VI) has type B spermatogonia (B), pachytene primary spermatocytes (P), Sa spermatids (Sa) with their grouped mitochondria (GM) and Sd1 spermatids (Sd1 ) with their annulus (An) and less distinct manchette. Stage VII (VII) is characterized by type B spermatogonia (B), pachytene primary spermatocytes (P), Sa spermatids (Sa), and Sd2 spermatids (Sd2 ). Sd2 spermatids, the most advanced form, are migrating toward the tubular lumen in this stage. Stage VIII (VIII) has newly formed preleptotene (Pl) and pachytene (P) primary spermatocytes, late Sa spermatids (Sa) with their acrosomic granule (AG) and newly attached developing flagellum, and Sd2 spermatids lining the lumen with their migrated annulus, enlarged midpieces (MP), and attached cytoplasmic droplet (CD). Residual bodies (RB) left behind by spermiation of spermatids can be seen near the luminal surface and in transit toward the base within Sertoli cell (SC) cytoplasm. Scale bar, 10 µm.42
various bloodborne molecules into the epididymal lumen, thereby allowing spermatozoa to be bathed in a milieu that is controlled locally, and affording spermatozoal protection against immunological attack.58 Disruption of the blood–testis barrier91 and the blood–epididymis barrier92 may be a contribu-
tor to reduced semen quality in older males. The corpus may be the most vulnerable segment of the epididymis to age-related barrier dysfunction.92 Barrier-mediated isolation of the epididymal lumen from the blood plasma probably necessitates more reliance on alternative signaling pathways.
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Figure 97.21 Light microscopic view of a cross-section of an equine seminiferous tubule, showing the nuclei of some spermatogonia that label positive (brown–black) with an immunohistochemical apoptotic detection assay (termed the TUNEL assay). The background tissue stain is periodic acid–Schiff/ toluidine blue. The germ cell associations in the cross-sectional view are consistent with stage V of the seminiferous epithelial cycle.43 (See color plate 40).
Figure 97.20 The spermatogenic wave revealed by an enzymatically isolated equine seminiferous tubule that was fixed in glutaraldehyde, followed by osmium tetroxide, then infiltrated with Epon, mounted in toto in Epon, and observed by brightfield microscopy. The corresponding drawing reveals the consecutive stages along the length of the tubule as determined by observation with Nomarski optics. While modulations (reversal of order) occur, adjacent stages are in consecutive order. Latter stages (more developed; higher numbers) are observed in the same direction of two sets of all stages I–VIII. Scale bar, 200 µm.42
Paracrine pathways (where target cells in the epididymis are issued signals from secretions of neighboring cells) and juxtacrine pathways (where target cell membrane receptors are activated by molecules on the plasma membrane of neighboring cells) are likely to exist.93 Lumicrine pathways (where
signaling molecules originate in the testes and are transported via the rete testis into the epididymal duct, or originate in one region of the epididymis and have an effect in another region of the excurrent duct system) are also key to epididymal function.58, 93, 94 Testosterone (synthesized by the Leydig cells) and androgen-binding protein (synthesized by the Sertoli cells) represent prime examples of a lumicrinederived mechanism of action in the epididymis. Of interest, androgen-binding protein can also be synthesized and secreted by principal cells along the entire length of the epididymis, possibly exerting a paracrine- or lumicrine-related action.58, 95 An assortment of growth factors derived from the testis have been proposed to have an action on the epididymis in a lumicrine fashion,93 and many growth factors are also produced locally within the epididymis.93 Close interactions between spermatozoa and the epididymal epithelium are crucial for the maturation process to occur, so it is not surprising that spermatozoa may also serve a lumicrine role within the epididymis.96 In summary, the epididymis serves to concentrate, mature, transport, and store spermatozoa. Massive resorption of luminal water within the efferent ductules and proximal caput and, to a much lesser extent,
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shedding of the cytoplasmic droplet. The cytoplasmic droplet of caput spermatozoa is consistently located in a proximal position near the neck of the middle piece. A few spermatozoa have bent middle pieces associated with translocation of the cytoplasmic droplet and spermatozoal maturation. When spermatozoa reach the proximal cauda epididymidis, most have completed translocation of the cytoplasmic droplet, as noted by droplets located near the junction of the middle piece and the principal piece. Under normal circumstances, few ejaculated spermatozoa have cytoplasmic droplets.68 Other more subtle, but necessary, changes acquired by spermatozoa during epididymal maturation include remodeling of the surface lipids, proteins, and glycocalyx;97–104 reorganization of acrosomal molecules;105 further changes in the structural stability of the nuclear chromatin (Figure 97.28);68, 106 development of signaling pathways;107 and alterations in spermatozoal metabolism.108 The collective biochemical changes, both intracellular and extracellular, incurred by spermatozoa during epididymal maturation remain a mystery, but experimentation is gradually revealing new findings.
Spermatozoa structure: form to function
(b) Figure 97.22 Scanning electron microscopic view of the equine testicular parenchyma, illustrating (a) the rete testis complex (RTC) housed in the connective tissue of the mediastinum around the central vein (CV) of the equine testis, and (b) a cross-support within the complex. (a) Seminiferous tubules (ST) are adjacent to the connective tissue around the rete testis complex. Cross-supports (open arrow) are found in the rete testis complex and in rete testis tubules (RTT). (b) The cross-supports are composed of a simple cuboidal epithelium overlying a connective tissue core. Testicular spermatozoa may be attached to this epithelium. Scale bars, 500 µm (a) and 4.6 µm (b).15
in the remainder of the epididymal duct leads to a 105 -fold increase in spermatozoal concentration in the cauda epididymidis, as compared with the rete testis.58 Observable changes in spermatozoa during epididymal maturation, based on light microscopic analysis, include (i) acquisition of flagellar movement within the corpus epididymis, followed by a progressive pattern of spermatozoal motility within the cauda epididymis, and (ii) translocation and
The general form of the spermatozoon was first described by van Leeuwenhoek over 330 years ago (1677). General acceptance of his proposal that this tadpole-shaped form contributed to fertilization required approximately 200 years. Thereafter, the structure of the spermatozoon received mounting attention. Resolving power of early microscopes hampered the ability to provide an exquisite description of spermatozoal anatomy, but introduction of electron microscopes in the middle of the twentieth century paved the way for intricate viewing of the spermatozoon and its substructure. Excellent descriptions of spermatozoal ultrastructure were provided by Saacke and Almquist in 1964,109, 110 and by Fawcett in 1975.111 More recently, Eddy112 provided a exceptional review of the topic, and included molecular-related insights that have been acquired in recent years. The spermatozoon is typically divided anatomically into a head and a flagellum (or tail). The head contains the nucleus, overlying acrosome, and a reduced complement of cytosolic elements. The head can be subdivided into an acrosomal region, equatorial segment, post-acrosomal region, and posterior ring, which demarcates the junction between the head and flagellum. The posterior ring is the site of plasma membrane anchoring to the nuclear envelope, and is thought to produce a tight seal that separates
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Figure 97.23 Transmission (a, b) and scanning (c, d) electron microscope views of the efferent ducts. (a, b) Cells lining the efferent ducts are composed of pseudostratified columnar epithelial cells and basal cells (BC). Ciliated cells (CC) provide a mechanism to move the spermatozoa through the duct, while non-ciliated cells (NcC) contain microvilli that enhance absorption of fluid by endocytosis and pinocytosis. Vesicles (V) located at the apex of the cells near the lumen are evidence of their absorptive functions. (c, d) Spermatozoa in the efferent ducts still have proximal droplets characteristic of spermatozoa in the testis and efferent ducts. Both ciliated and non-ciliated epithelial cells can be seen. Scale bars, 4.7 µm (a), 1.33 µm (b), 5 µm (c), and 2.2 µm (d).57
cytosolic components of the head and flagellum. The flagellum can be subdivided into a connecting piece, middle piece (or midpiece), principal piece, and end piece (Figures 97.29 and 97.30).19, 26 These various parts of the spermatozoon are surrounded by a common plasma membrane, although the composition of the plasma membrane can be subdivided into regional domains that impact its multiple functions, such as sperm–oviductal adhesion; penetration of the cumulus–oophorus matrix; sperm–zona adhesion; the acrosome reaction; acquisition of activated motility and hyperactivated motility; and sperm–oocyte adhesion and fusion.112 The equine spermatozoon has similar general structures to that described for the bull, ram, boar,
ram, dog, and human. The average length of an equine spermatozoon is reported to be 61–86 µm, with the average lengths of the head, midpiece, principal piece, and end piece reported as 5.8–7.0, 9.8–10.5, 43.8–67, and 2.79 µm, respectively.113–115 The maximum width of the head is reported to be 2.9–3.9 µm.113–115 The size of an equine spermatozoon, relative it body size, is remarkably smaller than that of some other species, such as the mouse, rat, hamster, and honey possum, where spermatozoal lengths are 123, 190, 189, and 356 µm, respectively.113 For further comparison, the average length of a human spermatozoon is 57 µm, with an average head length of approximately 4.5 µm.113
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Figure 97.24 Low-magnification scanning electron micrographs of an efferent duct (a) and the various profiles of the same single duct in the caput epididymidis (b). (b) A cluster of spermatozoa (arrow) is seen filling the lumen of one cross-section of this duct. Sparse smooth muscle surrounds the efferent ducts (a) and caput epididymidis (b). Scale bars, 10 µm (a) and 112 µm (b).57
The equine spermatozoon head is defined as splatulate-shaped, in contrast to the falciformshaped sperm heads characteristic of some species (e.g., the mouse, rat, and hamster). Of the laboratory animal species, the equine spermatozoon most closely resembles that of the rabbit. The spermatozoal head is somewhat elliptical in shape, is flattened in one plane (dorsoventrally), and is thicker in the posterior portion of the head than in the apical portion of the head (Figures 97.30 and 97.31).14, 19, 26 The nucleus, which occupies the majority of space within the spermatozoon head, contains the paternal genetic material. Specifically, the male genome is composed of the X or Y chromosome and a haploid number of somatic chromosomes. This genetic material is
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packaged for delivery to the oocyte for fertilization, where two haploid (male and female) genomes are combined to produce a diploid offspring. The chromatin (i.e., the DNA and associated proteins) within the nucleus of the mature spermatozoon is highly condensed, resulting in a volume that may approach only 5–10% of that of a somatic cell. This packing is a result of a marked alteration in the composition of nucleoproteins which occurs during epididymal transit. The haploid genome of the round spermatid encodes for unique spermatozoal proteins, termed protamines, which predominate as nucleoproteins during spermatozoal maturation in the epididymis. The cysteine residues of this protein establish intramolecular and intermolecular disulfide linkages
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Figure 97.25 Scanning (a) and transmission (b) electron micrographs of the epithelium of the caput epididymidis. Tall columnar cells are lined with stereo cilia (extremely long microvilli; Sc). (a) The cytoplasm (C) and luminal surface of these cells are revealed. (b) Section through the apex of these cells reveals forming vesicles (FV) in the process of endocytosis. Scale bars, 10 µm (a) and 2 µm (b).57
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Figure 97.26 Scanning electron micrographs of spermatozoa in the caput epididymidis with their proximally located cytoplasmic droplets (CD; a), epithelium of the corpus epididymidis with its tall columnar cells with shorter stereo cilia than found in the caput (b), corpus epididymidis spermatozoa undergoing translocation of the cytoplasmic droplet (c), and epithelium and thick smooth muscle (SM) wall of the proximal cauda epididymidis (d). (c) The corpus epididymidis is the site of droplet translocation from the proximal to the distal position. The translocation process appears to occur by bending (arrow) of the tail around the droplet followed by re-straightening of the tail. Scale bars, 10 µm (a), 45 µm (b), 10 µm (c), and 240 µm (d).57
that result in compaction and stabilization of the associated DNA. This design is thought to provide protection to the chromosomes during their perilous journey within the female reproductive tract, and to streamline the spermatozoon so as to enhance its mobility upon activation at the time of ejaculation. The nuclear envelope, consisting of a double membrane (each with a lipid bilayer), separates the contents of the nucleus from the surrounding cytoplasm. While the nuclear envelope is regularly perforated by nuclear pore complexes in somatic cells, such pores are absent over most of the spermatozoal nuclear envelope, i.e., in the area under the acrosome and in the post-acrosomal region. The exception is the region of the redundant nuclear envelope, a portion of the nuclear envelope posterior to the chromatin that folds and ex-
tends back into the neck region. This portion of the nuclear envelope contains abundant hexagonally arranged pores. The caudal portion of the nuclear envelope forms a concavity, termed the implantation fossa, which is the site of attachment with the flagellum. This region of the nuclear envelope is overlain with a thick sheet of material, termed the basal plate. The acrosome is a membrane-bound exocytotic organelle that overlies the rostral two-thirds of the nucleus, with a fit resembling that of a bathing cap. Anatomically, the membrane is subdivided into an inner acrosomal membrane which is continuous with an outer acrosomal membrane. These connecting membranes enclose a narrow heterogeneous compartment, termed the acrosomal matrix. The inner acrosomal membrane is closely apposed to the
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Figure 97.27 Scanning electron micrographs of epithelium and part of the smooth muscle (SM) wall of the proximal cauda epididymidis (a), spermatozoa of the cauda epididymidis (b), cross-section of the distal cauda epididymidis revealing the epithelium, supportive connective tissue, and thick smooth muscle (SM) wall (c), and ejaculated spermatozoa, most of which have lost their cytoplasmic droplets (d). Scale bars, 50 µm (a), 10.8 µm (b), 556 µm (c), and 4.5 µm (d).57
nuclear envelope whereas the outer acrosomal membrane underlies the plasma membrane. These two membranes converge at the level of the equatorial segment. The acrosome originates from the Golgi apparatus in round spermatids during spermiogenesis. Light microscopic changes of the developing acrosome can be visualized in cross-sections of the seminiferous tubules (Figures 97.4 and 97.13). Refinements in acrosomal morphology and biochemical composition continue as spermatozoa traverse the epididymis.116 In the mature spermatozoon, the acrosome contains a bountiful supply of active molecules, both within the acrosomal matrix and as components of the inner acrosomal membrane. These molecules, which include an assortment of protein receptors and hydrolytic enzymes, are thought to be important for adhesion to, and penetration of, the zona pellucida (ZP) and sperm–oolemma interactions.116–119
During the course of the acrosome reaction, the outer acrosomal membrane fuses with the overlying plasma membrane, thereby creating hybrid vesicles and pores that lead to release and exposure of acrosomal contents (Figure 97.32).116 Although the spermatozoal acrosome contains several enzymes characteristic of a typical cellular lysosome (in addition to those enzymes specific to the spermatozoon), the enzymes are not used within the cell as in autophagy for cellular organelle remodeling and renewing, or in heterophagy as occurs in phagocytic cells. The cytosolic compartment of the mature spermatozoal head is virtually free of organelles other than the acrosome, and the mature spermatozoon is considered to be transcriptionally quiescent. Nonetheless, the cytoplasm of the head region contains cytoskeletal proteins that are important to spermatozoal function. Structural changes occur
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Figure 97.28 Transmission electron micrographs of spermatozoa treated with the ionic detergent, sodium codicil sulfate, from the proximal corpus epididymidis (a) and distal corpus epididymidis (b). (a) Spermatozoa from the testis to the proximal corpus epididymidis have little resistance to detergent treatment, as noted by loss of all tail components and membranes and the decondensation of the nucleus (DN). (b) Spermatozoa from the distal corpus epididymidis, through the cauda epididymidis, and in the ejaculate have acquired some resistance to this detergent treatment during maturation in the epididymidis as noted by the presence of the outer membrane of mitochondria (OM), the outer dense fibers (DF) of the tail, and nucleus (N) that have not decondensed. Scale bars, 0.6 µm (a) and 1 µm (b).68
in the cytoskeleton (both microfilaments and microtubules) during spermiogenesis that lead to the reshaping of the spermatid head to an elongated form. This appears to be primarily mediated by the acrosome–acroplaxome–manchette complex and the perinuclear theca.120–122 The acroplaxome is formed at the leading edge of the developing acrosome. This structure contains both filamentous actin and keratin, and anchors the developing acrosome to the underlying nuclear envelope. The manchette consists of a circumferential bundle of microtubules that assemble posterior to the developing acrosome at the perinuclear ring, and extend along the developing flagellum to a point where the fibrous sheath begins (Figures 97.13 and 97.14).123 The manchette may aid nuclear elongation, as well as directional migration of molecules necessary for both nuclear condensation and tail formation.120, 121 The perinuclear theca is located between the inner acrosomal membrane and the nuclear envelope (i.e., the subacrosomal region), as well as between the nuclear envelope and the plasma membrane
posterior to the developing acrosome (i.e., the post-acrosomal region). This theca is composed of a complex of highly condensed cytotoskeletal proteins that have actin-binding properties, and appear to be involved in reshaping of the spermatozoal head.121 The connecting piece of the flagellum consists primarily of a capitulum, segmented columns of fibers, and the proximal and distal centrioles. The connecting piece serves to attach the flagellum to the head, and also functions to produce and stabilize some structural components of the flagellum. The capitulum articulates with the head at the level of the implantation fossa, attaching by fine filaments that connect the capitulum to the basal plate. The segmented columns anchor the dense fibers of the flagellum (Figure 97.31). The centrioles, oriented at right angles to each other, are involved in development of the connecting piece and the axoneme. The proximal centriole remains attached at the implantation fossa, but the distal centriole gives rise to the axoneme during tail development.
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The primary structural elements of the flagellum include the axoneme, the outer dense fibers, the fibrous sheath, and the mitochondria (Figures 97.29–97.31 and 97.33). The midpiece is characterized by the presence of an axoneme, an array of outer dense fibers, and an overlying sheath of helically arranged mitochondria. The midpiece joins the principal piece at the annulus, a point where the mitochondrial sheath is replaced with a fibrous sheath. The principal piece thereby consists of a centralized axoneme, outer dense fibers of variable length, and a fibrous sheath. The fibrous sheath terminates at the junction of the principal piece and end piece, with the end piece consisting of
a small extension of the axoneme (or individually arranged acrosomal microtubules) past the termination site of the fibrous sheath. The entire flagellum is enveloped by a plasma membrane. The axonome is a cylindrical array of nine doublet microtubules that surround two singlet microtubules (termed the central pair) connected by regularly spaced bridges, thus forming the ‘9 + 2’ configuration that is characteristic of both cilia and flagella throughout the plant and animal kingdoms. The microtubules are composed primarily of the tubulin family of proteins. Each of the nine outer microtubule doublets consists of an ‘A’ subunit, which is
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(j) Figure 97.30 Phase contrast (j), scanning electron (b), and transmission electron (a, c–i) micrographs of equine spermatozoa. (a) The plasma membrane (PM) encloses the entire head and tail. The head is composed of the nucleus (N), the overlying acrosome (Ac), and post-acrosomal region (PR). The acrosome can be divided into the apical segment (ASA), principal segment (PSA), and equatorial segment (ESA). The inner acrosomal membrane (IAM) is in juxtaposition with the nuclear membrane (NM). The outer acrosomal membrane (OAM) fuses with the plasma membrane during the acrosome reaction to discharge the acrosomal contents and expose molecules attached to the inner acrosomal membrane. (j) The tail is composed of the midpiece (MP), which houses the mitochondria (M), the principal piece (PP), which contains the fibrous sheath (FS), and the end piece (EP). (c) The tail attaches at the implantation fossa (IF) (d–i). The distal centriole gives rise to the ‘9 + 2’-arranged microtubular complex of the axoneme, with the nine outer doublets (DI) surrounding the central pair (CP) of microtubules. The nine dense fibers (DF) parallel the axoneme and extend to different lengths within the principal piece. In the end piece, the axonemal doublets become disorganized and ultimately separate into 20 single microtubules (SM), but still are enclosed in the plasma membrane. The cytoplasmic droplet (CD), located at the proximal end of the midpiece in spermatozoa recovered from the equine efferent ducts (b, c), is characteristic of spermatozoa that have not yet matured in the epididymis. These cytoplasmic droplets contain remnants of spermatid cytoplasm that are not lost in the residual body upon spermiation. Contents may include endoplasmic reticulum, mitochondria, and microtubules, including remnants of manchettes. Scale bars, 0.5 µm (a, b), 0.75 µm (c), 0.24 µm (d–i), and 1.28 µm (j).19, 26
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Figure 97.31 Further magnified illustrations of Figure 97.29, revealing midsagittal and partially resected views of equine spermatozoa. (a) Plasma membrane; (b) acrosome; (c) outer acrosomal membrane; (d) inner acrosomal membrane; (e) nuclear envelope; (f) post-acrosomal lamina; (g) nucleus; (h) basal lamina and underlying capitulum; (i) proximal centriole; (j) segmented column; (k) outer dense fibers; (l) outer doublets of axoneme; (m) center pair of microtubules within the axoneme; (n) mitochondria.14
completely cylindrical and composed of 13 protofilaments, and a ‘B’ subunit, which is C-shaped and composed of 10 or 11 protofilaments (Figure 97.33). The A subunits serve as an anchor for the outer dynein arms and inner dynein arms which possess
ATPase activity and generate the force required for axonemal and, thereby, flagellar motion through an attachment–detachment cycle between A and B subunits of adjacent doublets. Filamentous nexin links or nexin arms connect adjacent doublets, and radial
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Stallion: Anatomy, Physiology and Endocrinology The outer dense fibers course from the connecting piece through the midpiece and into the principal piece. These fibrous structures are thought to provide structural support as well as passive elasticity during flagellar bending. Nine irregularly shaped outer dense fibers occupy sites overlying the nine axonemal doublets along the entire length of the midpiece. The outer dense fibers taper at varying locations within the principal piece. Two of the outer dense fibers terminate in the proximal portion of the principal piece, and the structural support of these two fibers is replaced by inward extensions of the fibrous sheath. The mitochondrial sheath forms near the end of spermiogenesis, i.e., after the axoneme and outer dense fibers are fully formed. The axoneme of the flagellum in the developing spermatid originates from the centriolar apparatus, and extends
Figure 97.32 Transmission electron micrograph revealing sagittal views of adjacent equine spermatozoal heads. One spermatozoon has an intact acrosome (black arrow) and one has undergone the acrosome reaction induced by the calcium ionophore A23187 (white arrow).
spokes connect the axonemal doublets to a helical sheath surrounding the central pair of microtubules. The mechanism of axonemal action is discussed below under spermatozoal motility.
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Figure 97.33 Magnified illustrations of cross-sections through the midpiece and principal piece regions of equine spermatozoa. (a) Plasma membrane; (b) mitochondria; (c) outer dense fiber; (d) axonemal doublet; (e) central pair of axonemal microtubules; (f) sheath surrounding central pair of axonemal microtubules; (g) radial arm; (h) fibrous sheath; (i) outer dynein arm; (j) inner dynein arm; (k) longitudinal column of fibrous sheath; (l) connecting bridge between central pair of axonemal microtubules; (m) nexin links; (n) annulus.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell outwardly in the flagellar canal (Figure 97.14). This canal is created by movement of the annulus and attached plasma membrane to the posterior aspect of the nucleus, resulting in an infolding of the plasma membrane toward the posterior aspect of the nucleus. The flagellar canal prevents attachment of the mitochondria to the midpiece of the spermatid until the annulus migrates distally to the junction of the midpiece and principal piece.26, 124 Coincidently, the mitochondria assume their final position in an end-to-end spiral arrangement around the outer dense fibers of the midpiece. The length of the midpiece, and the number of mitochondrial gyres, varies greatly among mammalian species, but is relatively constant within a given species. The equine spermatozoon contains 40–50 mitochondrial gyres. The order and symmetry of the mitochondria along the midpiece may convey a functional effect, as disruptions in this organization have been associated with reduced fertility of stallions.125 The fibrous sheath extends along the entire length of the principal piece, i.e., from the annulus to the end piece. It is composed of two longitudinally arranged fibrous columns that are bridged by a series of interconnecting, circumferentially arranged fibrous ribs. The fibrous sheath is thought to provide rigid structural support and elasticity to the flagellum, but it also serves an important role as a scaffold for a variety of cell-signaling and metabolic events.
Physiological considerations Spermatozoal motility Under natural conditions, regulation of spermatozoal motility occurs at three critical points: epididymal reservoir (cauda epididymis and ductus deferens) – suppression of motility; ejaculation – activation of motility; and oviductal reservoir – hyperactivation of motility. Spermatozoa in the cauda epididymis are intrinsically capable of motility,68, 126 but do not exhibit motility until released from the epididymis. A threshold level of cyclic adenosine monophosphate (cAMP) is present in spermatozoa within the cauda epididymis.127 Specific motility-inhibiting proteins have been identified in rat cauda epididymal fluid that, when removed, allow initiation of motility.128, 129 A pH-dependent inhibitory factor has been reported in bulls.130, 131 Although such inhibitory factors may exist, it is possible that sperm motility may simply be suppressed by the acidic pH of the epididymal environment. The pH of bull cauda epididymal fluid is reported to be 5.5, and the cytosolic pH of bull epi-
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didymal spermatozoa is reported to be 6.5–6.6.130, 132 Caudal epididymal fluid of bulls, rams, boars, and stallions does not contain measurable quantities of bicarbonate (HCO3 − ),133 and HCO3 − is known to be a key effector of spermatozoal motility.134, 135 Bicarbonate is present at fairly high concentrations in seminal plasma, and may be higher in seminal plasma of stallions than some other mammals studied.136 Simple exposure of normal spermatozoa to seminal plasma or physiological fluids will activate spermatozoal motility that is characterized by a moderate amplitude and symmetrical flagellar beat leading to a forward propulsive trajectory.137, 138 This form of spermatozoal movement is defined as activated motility and is to be differentiated from hyperactivated motility discussed below. Activated motility is considered necessary for propelling sperm into the oviductal reservoir. Environmental cues activate spermatozoal motility through a signal transduction mechanism. Although the signaling pathway(s) of activated spermatozoal motility are not resolved completely, an ever-increasing body of information indicates that HCO3 − , calcium ions (Ca2+ ), and cAMP are key signaling components.112, 139 Transmembrane movement of HCO3 − into the spermatozoal cytosol is associated with an increase in intracellular pH ([pH]i ), leading to regulation of cAMP.140, 141 Activity of Na,K-ATPase and Na+ /K+ exchangers are also of central importance in regulating [pH]i and sperm motility.142–146 Spermatozoal Na,K-ATPase is important for generation of the Na+ gradient required for ion transport. This Na+ gradient permits the Na+ /K+ exchangers to catalyze the coupled exchange of extracellular Na+ for intracellular H+ .143 A sperm-specific Na+ /K+ exchanger has been localized to the principal piece and likely plays an important role in regulating [pH]i .142, 145, 146 Activation of motility requires that this exchanger is functional.145 It also likely that a Na+ /HCO3 − cotransporter plays a role in controlling cytosolic pH (Figure 97.34).140 Although increased alkalinization of the spermatozoal cytosol is known to activate membrane Ca2+ channels, this may be of primary importance in hyperactivation of spermatozoal motility where increased cytosolic Ca2+ is required.141,147–152 Spermatozoal motility can be activated and maintained for a short time in Ca2+ -free media for many species,126, 138, 153 but presence of extracellular Ca2+ maximizes spermatozoal motility.154–156 The flagellum is known to contain a variety of Ca2+ membrane transport channels, including voltage-gated, cyclic nucleotide-gated, transient receptor potential, Ca2+ -release, and CatSper channels. While the roles of some of these channels remain unknown, their
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Figure 97.34 A schematic model of some documented and hypothesized molecular events associated with activated and hyperactivated forms of spermatozoal motility. Please refer to the text for an explanation of these signaling pathways and for figure abbreviations.
mere presence suggests that they probably contribute in some manner. Sperm [Ca2+ ]i is also known be regulated by Ca2+ -ATPases, Na+ /H+ exchangers, and Ca2+ /H+ exchangers.157 The signaling pathway for activated spermatozoal motility involves the cAMP-dependent protein kinase A (PKA) pathway.156,158–165 Intracellular production of cAMP, as it relates to motility, is catalyzed by soluble adenylyl cyclase (sAC). sAC has been identified in the flagellum of spermatozoa, and is required for activated motility166 and capacitation (see below). Interestingly, the cAMP–sAC–PKA does not appear to be required for hyperactivated motility150 or the acrosome reaction.166 sAC is stimulated directly by HCO3 −167, 168 or Ca2+ ,169, 170 thereby catalyzing phosphorylation of PKA, which then leads to phosphorylation of flagellar proteins that are central to activated motility.138,171–174 One such protein is A kinase anchoring protein (AKAP). AKAP has been detected at the level of the axonemal central pair, the fibrous sheath (in the principal piece), and the outer dense fibers (in the middle piece).112, 138, 171, 175, 176 PKA is known to bind to AKAP, thus suggesting that the action of PKA can be regionalized, and its substrate specificity aided, through this anchoring
mechanism.112, 138, 171, 177, 178 Although PKA is known to be located in the vicinity of the outer dynein arms,175 the need for a localized pattern of PKA distribution remains unresolved.179 In addition to regulating the cAMP–sAC–PKA pathway, Ca2+ affects dynein activity through direct control of the central pair of microtubules within the axoneme.180 As Ca2+ can inhibit the activity of the axonemal central pair to regulate dynein arm function,180 excessive Ca2+ can lead to cessation of spermatozoal motility.181 Calmodulin (CaM), a Ca2+ -binding and sensor protein, is a key component of spermatozoal signaling mechanisms.181–186 Calmodulin associates with the outer dynein arms and the radial spokes, and may play a role in Ca2+ -dependent propagation of flagellar bending.172, 175 Calmodulin kinase II (CaMK)151 and calmodulin-dependent protein phosphatase172, 187, 188 are central to dynein function. One study indicates that CaM can also interact with Ttype voltage-gated Ca2+ channels by a mechanism independent of PKA, CaMK, or calmodulin-dependent protein phosphatase.189 In addition, calmodulin may be involved in regulation of adenylyl cyclase.182
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell A substantial and continuous supply of energy, in the form of ATP, is required for activated spermatozoal motility, and this requirement is heightened for hyperactivated motility.150 The mechanisms by which ATP is generated and transferred in the flagellum remain unsolved. Certainly, oxidative respiration within the mitochondria yields a plentiful supply of ATP. Some investigators propose that the ATP produced in this manner is capable of diffusing along the entire length of the flagellum in a manner suitable for initiation and maintenance of spermatozoal motility.138,190–192 Possibly, an adenylate kinase shuttle may assist in the rapid distribution of mitochondrial-produced ATP.192 Recently, adenylate kinases were regionalized to the principal piece of mice spermatozoa.193 Others contend that local glycolysis within the principal piece is critical to the timely generation and distribution of ATP required for spermatozoal motility.171, 194, 195 Certainly, a variety of glycolytic enzymes have been identified in association with the fibrous sheath and/or outer dense fibers,196–202 and it is well known that the glycolytic pathway to energy production is essential under anaerobic conditions. Arguments in favor of oxidative respiration and of glycolysis exist for generation and supply of ATP for spermatozoal motility. A sperm-specific glycolytic enzyme, glyceraldehyde3-phosphate dehydrogenase (GAPDH), has been localized to the principal piece, and targeted deletion of this enzyme is reported to lead to a severe reduction in motility of mouse spermatozoa.194 In contrast, another study revealed that chemical inhibition of GAPDH did not affect spermatozoal motility or ATP concentration.192, 203 At present, it appears possible that either oxidative respiration or glycolysis, or both, can support the ATP generation and availability needed to drive spermatozoal motility. The length of the principal piece in stallions is similar to that in bulls, boars, rams, and dogs. It is also considerably shorter than the principal piece of mice, rats, hamsters, and guinea pigs, in which many experimental studies are conducted.113, 192 This may explain some of the inconsistencies among laboratories. The axoneme, dynein arms, outer dense fibers, mitochondrial sheath, and fibrous sheath are often portrayed as the fundamental elements of the flagellum. While each is vital to flagellar function, one must also be aware that these elements are embedded in a network of other molecules that are equally important. Bending of the flagellum is caused by reciprocal sliding between doublet microtubules within the axoneme. The dynein arms are permanently attached to one doublet microtubule with arms extending to intermittently engage an adjacent doublet microtubule. The dynein arms are situated at 24-nm (outer dynein arms) to 96-nm (inner dynein
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arms) intervals along the entire length of the axoneme.204, 205 When the crossbridges created by the dynein arms are complete between neighboring doublets, the corresponding microtubules are prevented from sliding. Dynein is a high-molecular-weight ATPase, so the dynein arms have the capacity to transform chemical energy into unidirectional mechanical force. Each dynein arm has three ATP-sensitive binding sites and all sites must bind with ATP for the dynein arm to detach from the microtubule,206 i.e., the crossbridge attachment–detachment cycle is dependent on binding and hydrolysis of ATP. The conformational change in the dynein arms results in the force generation required for microtubule sliding. The microtubules of the axoneme enhance the rate of ADP release following the dephosphorylation to aid the reactivation step.207 The nexin arms also undergo periods of displacement to allow sliding of microtubules.137 The basic mechanism is similar to that of muscle myosin; however, the loss and rebinding of ATP products occurs at two to three orders of magnitude faster for dynein than for myosin.208 Bending of the tail occurs during axonemal sliding because the microtubules are fixed at the base of the flagellum. 209 The passive elastic nature of the overlying outer dense fibers and fibrous sheath permits the flagellum to bend, and also provides elastic recoil.210, 211 Bending of the flagellum requires a patterned activation of microtubule sliding around the circumference, and along the length, of the cylindrical array of microtubules that compose the axoneme. This dynein-generated activity is regulated by the central pair of microtubules and their associated structures, collectively called the central apparatus. The asymmetry of components in the central apparatus forms the basis of a ‘timing device’ that creates the spatially timed sliding of various microtubules.180, 212 As such, the central apparatus both constrains and activates the dynein arms via communication through the radial spokes.137, 213 Spermatozoal metabolism Spermatozoa require a constant supply of energy for maintenance of cellular order and functions needed for survival. This energy requirement increases significantly with the onset of activated motility,214, 215 and becomes even more pronounced when hyperactivated motility is initiated.150 Approximately 500 common metabolic reactions are known to occur in most somatic cells, many of which require energy.216 A spermatozoon is a ‘stripped down’ cell that is designed expressly for propagation of genes and it is one of the smallest cells in the body, but it accomplishes its mission at a distance far from its origin in the epididymis. While spermatozoa require a
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relatively small amount of energy for ‘housekeeping’ purposes,197, 214 a plentiful supply of substrate is required to provide a spermatozoon with the stamina required for this arduous journey. Spermatozoa are also capable of prolonged survival both within and outside the body. Exogenously derived nutrients are needed to fulfill the energy demands of these very metabolically active cells. These nutrients (or substrates) are metabolized intracellularly, resulting in the release of chemical bond energy. Usable energy is made available for cellular processes primarily in the form of the activated carrier molecule, ATP, with other macromolecules such as NADH, NADPH, FADH2 , and acetyl CoA also providing vital high-energy intracellular linkages.216, 217 Spermatozoa possess the metabolic machinery required for glycolysis, the citric acid cycle, and oxidative phosphorylation.218 Glycolysis occurs in the cytosol and has both an anaerobic and aerobic mode,219 whereas the citric acid cycle and oxidative phosphorylation are strictly aerobic and proceed within the mitochondria. A variety of substrates can be garnered for energy extraction, including monosaccharides, pyruvic acid, lactic acid, and even fatty acids and amino acids, the last four of which require an aerobic environment. Sorbitol, a sugar alcohol obtained by reduction of glucose through the polyol pathway, is thought to serve as a substrate for metabolism.218 Although the seminal plasma of stallions does contain this product,136 stallion spermatozoa appear to be incapable of metabolizing it to produce ATP.218, 220 Generally speaking, glycolysis is considered the ‘backbone’ of the energy-procurement pathways because it can proceed in situations of low oxygen tension and its product, pyruvate, can be further catabolized to CO2 and H2 O by oxidative respiration. Efficiency of energy production is lowest in the glycolysis pathway, where one molecule of a monosaccharide can generate a net increase of two molecules of ATP. Conversely, complete oxidation of glucose through the citric acid cycle and oxidative phosphorylation generates an additional 34 molecules of ATP per monosaccharide molecule. Despite the increased efficiency of oxidative metabolism, a monosaccharide cannot directly enter the citric acid cycle, but must first be metabolized to pyruvate; hence, the need for glycolysis as the first stage of energy production where monosaccharides are concerned.216 Glycolysis involves a sequence of 10 separate reactions, the first of which is catalyzed by hexokinase in an ATP-consuming manner. This enzyme can phosphorylate several types of monosaccharides, including glucose, fructose, mannose, and galactose. The affinity of hexokinase for glucose is reported to be 20 times higher than its affinity for fructose.221
Another account indicated that, in mammalian spermatozoa, glucose is used in preference to fructose, but that mannose has the highest affinity of the three monosaccharides to hexokinase.218 Of interest, a study involving astroglial cells revealed that mannose-phosphorylating ability is only 40% of glucose-phosphorylating ability.222 Fructose was identified as the ‘seminal sugar’ in 1946.223 Subsequent studies have revealed that the concentration of this sugar is high in seminal plasma of bulls, rams, and man (range 40–1000 mg/100 mL), but extremely low (< 1 mg/100 mL) in seminal plasma of stallions.218 Interestingly, another report indicated that the fructose concentration in stallion semen ranged from 7 to 11 mg/100 mL.224 Seminal plasma concentrations of some other metabolizable substrates, such as citric acid, lactic acid, and pyruvic acid (utilized exclusively by oxidative respiration), are also much lower in stallion seminal plasma than that of bulls, rams, or man.218 Bull and ram spermatozoa utilize fructose at a rate of 2 mg/109 spermatozoa/h under anaerobic conditions at 37◦ C, whereas boar spermatozoa utilize fructose at a rate 10 times lower.218 One report suggests that stallion spermatozoa have a limited capacity to utilize fructose.220 Interestingly, fructose is not found in seminal plasma of dogs and glucose is used preferentially to fructose by dog spermatozoa; yet, fructose maintains higher spermatozoal motility than does glucose under in vitro storage conditions.225 Heterogeneity may also exist among motile dog spermatozoa, with subpopulations experiencing increased responsiveness to both fructose and glucose.226 Either glucose or fructose can increase motility and ATP concentration of human spermatozoa.227 One study with human semen revealed that spermatozoal motility was maximized in medium containing 1 mM glucose or 15 mM fructose; however, motility was further enhanced when both monosaccharides were provided.228 Conversely, in vitro fertilization in mice and rats is not attainable when fructose is the only available substrate, but high when glucose is included in the media.229–232 Other studies certainly support the importance of glucose in gamete interaction.233–235 Transport of monosaccharides across the plasma membrane requires the presence of carrier proteins and is an ATP-dependent process. One report involving bull spermatozoa indicated that the glucose transport protein has only a limited ability to transport fructose, and that another carrier protein may be the primary means by which fructose gains access into the cytosol.214, 236 This carrier protein may be absent in stallion spermatozoa, thus explaining why fructose may not be a primary, or even a suitable, energy source in this species.214
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell The extent to which spermatozoa utilize glycolysis or oxidative respiration for generation of energy varies considerably among mammalian species, and probably with the profile of spermatozoal motility, and environmental conditions.214, 218 Few studies have been directed toward stallion spermatozoa, so much of the information must be extrapolated from information garnered from other species. This approach may be precarious, however, because of the known species variation in spermatozoal metabolic preferences. One report suggests that bull and ram spermatozoa utilize the glycolytic pathway at a much higher rate than stallion spermatozoa, and that stallion spermatozoa rely more heavily on aerobic metabolism for energy production.214, 220 Others report that ram and bull spermatozoa oxidize acetic acid in preference to fructose and glucose.218 Human, boar, and mouse spermatozoa appear to obtain an high proportion of their ATP by glycolysis.237–239 Although more intensive study with stallion spermatozoa is required to more fully elucidate metabolic pathway preferences, glycolytic breakdown of glucose is currently considered to be a major source of ATP production.214 The minimal oxygen tension required to maintain a linear rate of O2 uptake by rabbit spermatozoa has been reported to be approximately 10 mmHg.218 As such, the luminal environments of the uterus and oviduct are thought to be sufficiently high in oxygen content to allow aerobic respiration to proceed.218, 240, 241 Storage of spermatozoa outside the body cavity, however, can affect availability of oxygen and metabolic processes. If raw or extended semen is left undisturbed in a laboratory setting, utilization of dissolved O2 by aerobic respiration leads to depletion of O2 and the need to resort to glycolysis for meeting energy demands.214 For oxidative metabolism, bull and ram spermatozoa utilize O2 at a rate of approximately 10–20 µL/hour/108 spermatozoa at 37◦ C.218 A potential drawback of oxidative respiration is the production of reactive oxygen species (ROS), which can lead to disruption of ATP production, lipid peroxidation, or other cellular mechanisms that result in spermatozoal dysfunction.242–246 Metabolism of endogenous substrates by spermatozoa for the production of energy is thought to be minimal, possibly representing 10% or less of total energy production, based on calorimetric and carbon balance techniques.247 For bull spermatozoa, endogenous metabolism occurs at a constant rate at storage temperatures between 20◦ C and 35◦ C.248 Spermatozoal capability for glycogen synthesis and storage was once thought to be non-existent. Recent reports, however, indicate that spermatozoa of dogs, boars, rams, and horses do contain glycogen,
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as well as the enzymatic machinery required for its production.192, 249, 250 The pentose phosphate pathway does not appear be directly involved with spermatozoal motility,218 but may be central to sperm–oocyte interactions,251 possibly through production of NADPH and the resultant generation of radical oxygen species required for sperm–oocyte fusion.252 Furthermore, the pentose phosphate pathway may aid in protection against oxidative injury through generation of NADPH, which is required to return expended glutathione peroxidase to its reduced (protective) form (see Reactive oxygen species).253, 254 Reactive oxygen species A discussion directed at ROS at this point in the chapter seems appropriate because of the pathological consequences that ROS can have on spermatozoal motility and spermatozoal metabolism. ROS are products derived from the reduction of (i.e., addition of electrons to) diatomic oxygen (O2 ), and include radicals and other reactive products. Radicals are atomic or molecular species that have unpaired electrons in their orbits, thus making them quite unstable, i.e., highly reactive, and likely to participate in a variety of chemical reactions in order to displace, receive, or share electrons so that they may once again become stable.255 Other O2 derivatives are not radicals but are highly reactive; hence, these products are collectively termed reactive oxygen species. Examples of ROS (where ‘·’ implies the presence of an unpaired electron) include superoxide radical (O2 − ·), hydroxyl radical (OH·), hydroperoxyl radical (HO2 ·), peroxyl radical (RO2 ·), alkoxyl radical (RO·), hydrogen peroxide (H2 O2 ), and subclasses of ROS that contain reactive nitrogen or chlorine species, such as nitric oxide (NO·), peroxynitrite (ONOO− ; where NO· + O2 − · → ONOO− ), or hypochlorous acid (HOCl).255 ROS may assume physiological roles (see Capacitation); however, their presence can also lead to spermatozoal demise.66,256–260 Spermatozoa are exposed to ROS that are derived both intracellularly and extracellularly. Production of superoxide occurs continuously within the mitochondrial electron transport chain where O2 acts as an electron carrier during oxidative respiration. A molecule of O2 must pick up a total of four electrons in order to form water (i.e., O2 + 4H+ + 4e− → 2H2 O). During this process, the reactive oxygen forms are generally caged within the respiratory enzyme complexes. A small percentage of the superoxide produced does not stay within the mitochondrial respiratory chain, but escapes to exert actions on other cellular components, including lipids, sugars, proteins, and nucleic acids.255, 258,261–264
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Recently, presumed non-mitochondrial production of O2 − · has been shown to occur in spermatozoa of both men265 and stallions.266 Evidence is mounting that spermatozoa contain NADPH oxidase, which catalyzes the formation of superoxide, presumably for physiological reasons,267–270 although some workers conclude that spermatozoal derived NADPH oxidase activity is insignificant.254, 271, 272 An aromatic amino acid oxidase, released from dead spermatozoa or present in seminal plasma, has also been reported to generate production of H2 O2 in semen.273–276 Generation of ROS is known to be more pronounced in morphologically abnormal spermatozoa, a feature that can be associated with retention of excessive amounts of residual cytoplasm.258,277–281 Damaged or morphologically abnormal equine spermatozoa are known to generate more ROS than do morphologically normal sperm.282 Neutrophils are known to be a primary exogenous source of ROS in semen of men, where a relatively high concentration of leukocytes is commonplace.283, 284 While spiking equine semen with activated neutrophils can lead to reduced motility,285 contamination of equine semen with sufficient neutrophils to affect spermatozoal motility is quite uncommon. Another source of ROS is the uterine environment, such as the ROS generated from estrogen-induced uterine NADPH oxidase of the endometrial epithelium.276, 286 Cycle-regulated synthesis of uterine nitric oxide has also been described.287, 288 Leukocytes of mares produce H2 O2 in response to spermatozoa or bacteria, and this activity is heightened by leukocyte exposure to seminal plasma,289 suggesting a means for uterine clearance of bacteria and spermatozoa. Others, however, report a protective effect of seminal plasma against ROS.280, 290, 291 Seminal plasma of some subfertile men is known to possess reduced antioxidant capacity.292, 293 Unsaturated lipids are quite susceptible to peroxidative injury when exposed to various ROS, and mammalian spermatozoa are especially susceptible because their plasma membranes are rich in polyunsaturated fatty acids.242, 245, 257,294–299 One report300 indicated that the spermatozoa of individual men vary in their membranous unsaturated fatty acid content, and that a superabundance of polyunsaturated fatty acids predisposes spermatozoa to oxidative injury. Peroxidative injury has been thought to adversely affect membrane fluidity,258 although some work suggests that the oxidative action may not occur through lipid peroxidation.301 Oxidation may directly affect proteins and membrane permeabilization (possibly through oxidation of sulfhydryl groups on enzymes and membrane proteins), as opposed to disturbing lipid fluidity.302 Others have described an effect of
ROS on the sperm axoneme and ATP generation that can lead to an irreversible loss of motility.303, 304 In stallion spermatozoa, a ROS-associated reduction in motility was not associated with a detectable increase in lipid peroxidation, or a decrease in viability or mitochondrial membrane potential.260 This finding suggests that ROS-related effects on equine spermatozoa may occur through a mechanism unrelated to lipid peroxidation, and also indicates that motility may be a more sensitive indicator of ROSrelated injury than the other experimental end points, i.e., viability, mitochondrial membrane potential, and lipid peroxidation. Others report that lipid peroxidation is a good predictor of sperm motility and consider a lipid peroxidation assay to be a potential clinical test.305 The mechanisms of ROS-induced damage to DNA have been evaluated intently.306 Sperm DNA is known to be susceptible to oxidative injury, resulting in reduced fertility and perhaps even pregnancy loss or a variety of pathological entities in offspring.307 Aberrant DNA repair by spermatozoa is thought to increase the mutagenic load of any resulting conceptus, thereby leading to post-fertilization crises.281, 308 Mitochondrial DNA is more susceptible to H2 O2 induced damage than is nuclear DNA, possibly making it a good marker of oxidative injury.309 Spermatozoa of stallions are susceptible to ROS-induced DNA fragmentation.310 Protection against the effects of ROS in spermatozoa is afforded by an assortment of scavenging molecules, including three enzyme systems: (i) superoxide dismutase (SOD), (ii) catalase (CAT), and the glutathione peroxidase system (GPx), as indicated by the following reactions: O2 −· + O2 −· + 2H+ → H2 O2 +O2
(97.01)
2H2 O2 → 2H2 O +O2
(97.02)
2GSH + H2 O2 → GSSG + 2H2 O
(97.03)
SOD
CAT
where GSH is monomeric glutathione and GSSG is oxidized glutathione disulfide. Glutathione reductase (GRD) then reduces GSSG to GSH to complete the cycle as follows: GSSG + NADPH + H+ → 2GSH +NADP+ GRD
Superoxide dismutase is considered to be prevalent in spermatozoa of many species,261,311–313 and rapidly reduces O2 − · to H2 O2 . The direct cellular action of O2 − · is thought to be minimal, with its major effects produced indirectly, through conversion to H2 O2 243, 256, 260, 274, 294, 301, 307, 310, 314 or other radicals.255
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell Spermatozoa of stallions are thought to have limited intrinsic SOD, with SOD activity derived primarily from the seminal plasma through adsorption to the plasma membrane.315 Glutathione peroxidase has also been identified in spermatozoa or seminal plasma,311, 313, 316 and is considered to impart protection against oxidative injury incurred by H2 O2 ;316, 317 however, a primary action of glutathione may be protection against peroxyl radicals of polyunsaturated fatty acids (RO2 ·) through conversion to relatively inert hydroxyl fatty acids.313, 318 Glutathione also plays a structural role, as demonstrated by sulfhydryl oxidation and crosslinking associated with nuclear condensation and assembly of the midpiece.317, 319 Glutathione is present in stallion semen,315, 320 but the source in semen appears to be primarily of seminal plasma origin.315 Spermatozoa from most species contain little to no catalase;243, 245,321–324 however, seminal plasma is generally high in catalase activity.67 The amount of catalase in rabbit semen is genetically influenced (estimated heritability value of 0.48).325 Stallion semen contains catalase activity311, 326 that is primarily derived from prostatic secretions.326 Several studies have demonstrated that addition of catalase to semen of various species will help neutralize the detrimental effects of H2 O2 on spermatozoa following in vitro storage.260, 285, 290, 301, 310,327–330 Similarly, oviductal fluids may contain catalase in sufficient quantity to protect sperm from oxidative injury.330, 331 Surprisingly, stallion semen with higher initial activity of CAT, SOD, and GSH did not lead to improved retention of spermatozoal motility or membrane integrity following cooled storage.311 Addition of CAT to extended equine semen did not improve maintenance of motility following cooled storage.332 Similarly, addition of CAT, SOD, or GSH, ascorbic acid, or α-tocopherol to semen extender did not improve post-thaw quality of stallion spermatozoa.333 Studies with ram spermatozoa suggest that CAT, SOD, and GSH are ineffective at preventing acute peroxidative injury to lipids,302, 334 and incorporation of CAT in extenders for sex-sorted or non-sorted semen does not improve post-thaw quality of ram spermatozoa.335 Overall, spermatozoa have limited endogenous CAT, SOD, and GPx activity; therefore, spermatozoa depend on extracellular availability of these enzymes to counter oxidative stress, namely from seminal plasma. Seminal plasma contains these enzymes, in addition to a host of other free radical scavengers.276 Molecules with antioxidant activity include, but are not limited to, ascorbic acid,336, 337 α-tocopherol,246, 334, 336,338–340 taurine,341, 342 hypotaurine,342 butylated hydroxyanisole (BHA; an antioxidant used for long-term preservation of food, cosmetics, and pharmaceuticals),246, 334, 339 al-
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bumin,261, 342, 343 cysteine,344, 345 lipoic acid,346–348 xanthurenic acid,349–351 carnitine,352, 353 ergothioneine,354 and pyruvate.349, 355 Many of these have been used as dietary supplements, or as direct semen treatments, with mixed results.267, 299, 332, 340,356–358 As the end product of glycolysis, pyruvate serves as an important substrate for oxidative phosphorylation, but when present at relatively high concentrations, pyruvate can also exhibit antioxidant properties.355, 359 Addition of pyruvate to milk-based extender at a concentration of 2 mM is reported to improve equine spermatozoal motility following cooled storage.349 Presently, there is much controversy over types, doses, and delivery methods for antioxidants as they relate to semen processing. Further study is required to determine whether antioxidant therapy will maximize spermatozoal function, especially following cooled or frozen storage. Spermatozoal transport Spermatozoa are virtually immotile in the epididymis, but develop motility (or, more precisely, activated motility) upon ejaculation. Deposition of spermatozoa is intrauterine when artificial insemination is used, and a large portion of an ejaculate is deposited directly into the uterine body at the time of natural coitus if the mare’s cervix is dilated at the time of breeding and if the stallion does not dismount prematurely during the ejaculatory process. Following intrauterine deposition of semen, the spermatozoa are rapidly transported to the oviduct where a spermatozoal reservoir forms and the spermatozoa gain fertilizing potential prior to interaction with the vestments of the oocyte near the ampullar–isthmic junction. Spermatozoal migration to the oviducts is dependent, to a large part, on uterine contractions.360 The effect of insemination volume on the frequency of these uterine contractions appears variable,361, 362 but, within the range tested (5–50 × 106 spermatozoa/mL), spermatozoal concentration had no apparent effect on spermatozoal number recovered from mare oviducts at 4 hours following insemination.363 The time required for spermatozoa to gain access into the oviduct has not been studied extensively. Spermatozoa have been detected in the oviducts as early as 2 hours post-insemination, based on recovery of spermatozoa in abattoir specimens.364 Sufficient spermatozoa to establish pregnancy may be transported into the oviduct as early as 30 minutes following insemination, based on extensive lavage of the uterus post-insemination with an iodine-based solution to immobilize intrauterine spermatozoa. Pregnancy rates are maximized by delaying the uterine lavage until 4 hours post-insemination.365 Location
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of intrauterine insemination appears to have a significant impact on the spermatozoal number that gain access into the oviduct. In one study, a higher number of spermatozoa were recovered from the ipsilateral oviduct following deep uterine horn insemination than resulted from uterine body insemination. In the same study, the number of oviductal spermatozoa recovered from reproductively normal mares and mares susceptible to post-mating endometritis were similar following artificial insemination of 500 million total spermatozoa.366 Others have found more spermatozoa, and a higher percentage of motile spermatozoa, in the isthmic oviductal region of reproductively normal mares than in mares susceptible to chronic uterine infection.367 The same investigators reported more spermatozoa in the oviducts when mares received semen from a fertile stallion, as opposed to a subfertile stallion.367 Another group reported an effect of stallion on spermatozoal numbers in recovered oviducts of mares following artificial insemination, although the fertility or semen quality of these stallions was not provided.368 Administration of oxytocin to mares immediately after insemination did not improve pregnancy rates in mares bred by either fertile or subfertile stallions.369 This may be due to a directional pattern of luminal flow toward the cervix following oxytocin administration, as occurs naturally during the expulsive stage of labor (i.e., uterine evacuation).370, 371 It is interesting to note, however, that dynamic scintigraphic analysis of radiolabeled spermatozoa in the mare uterus revealed radioactivity in the tips of the uterine horns as early as 8 minutes following uterine body insemination (5 mL volume).360 In this study, uterine contractions did not propagate spermatozoa in one direction only; rather, movement continued in both tubal and cervical directions. Similarly, ultrasonographic evaluation of the uterus following natural mating of mares revealed similar patterns of uterine contractions in cervico-tubular and tubo-cervical directions.362 The peristaltic contractions were increased in frequency, amplitude, and duration in comparison with the precoital findings. It appears that bidirectional myogenic activity in the early period following insemination assists spermatozoal transport to the UTJ, and it is possible that this mechanism is impeded by administration of supraphysiological doses of oxytocin or other drugs with oxytocic properties. Studies in pigs support this line of thought, as intrauterine infusion of cloprostenol prior to insemination reduced oviductal recovery rates of spermatozoa and increased cervical reflux of inseminates.372 In another report involving pigs, spiking a semen dose with 10 IU oxytocin reduced fertilization rate, whereas intravenous administration of the same dose at 5 minutes following
insemination improved fertilization rate.373 The addition of another uterotonic agent, PGF2α , to extended semen (final concentration of 125 µg/mL) did not improve the pregnancy rate in a small group of mares.374 The involvement of other factors that may affect spermatozoal transport in horses requires further examination. As an example, studies involving pigs revealed that the mere presence of a boar (i.e., nontactile presence) could increase peripheral plasma oxytocin concentration in the sow,375 and uterine activity following boar exposure was most enhanced in sows that initially had a reduced frequency of uterine contractions.376, 377 Similar studies involving horses are contradictory, with some reports of increased myometrial activity and plasma oxytocin concentrations following simple stallion exposure,378–382 and another report indicating no increase in uterine contractions until mechanical stimulation of the vagina and cervix occurs.383 It seems logical that seminal plasma contributes to uterine transport of spermatozoa following natural cover, as it provides a medium to aid peristaltic movement of spermatozoa toward the oviducts. Mann et al.384 demonstrated the presence of two chemical markers of seminal plasma in the uterine horns at 50 minutes following mating, with concentrations similar to those detected in fresh ejaculates. These data are consistent with the concept that most of the post-coital fluid in the uterus is derived from the seminal plasma. Using the same chemical markers, these scientists also demonstrated that seminal plasma may gain access into the oviducts. The physiological significance of seminal plasma in spermatozoal transport extends beyond that of a simple vehicle. Seminal plasma contains a rich assortment of both organic and inorganic constituents.136 Prostaglandins have been identified in relatively high concentration in the seminal plasma of humans, monkeys, and the great apes, while the concentration in seminal plasma of stallions is minuscule.136 Of interest, the seminal plasma of boars contains an appreciable concentration of estrogens (up to 12 µg per ejaculate), and this steroid is thought to stimulate myometrial contractions by inducing the release of PGF2α from the endometrium.376, 385 Spiking boar semen with estradiol-17β, however, did not improve pregnancy rate or fetal number following artificial insemination of gilts.386 We are unaware of reports relating to estrogens in seminal plasma of stallions. Seminal plasma is a well-known modulator of spermatozoon-induced uterine inflammation, a feature that likely plays an integral role in removal of spermatozoa from the uterus.387, 388 Troedsson et al.388 reported that some proteins in equine seminal plasma can protect viable, but not damaged,
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell spermatozoa from binding to neutrophils. While this finding may not affect spermatozoal transport at the first breeding of an estrous cycle, it could affect spermatozoal transport mechanisms if a second insemination were placed in an inflamed uterus induced by the first insemination. Furthermore, it could potentially affect semen transport in mares bred for the first time in an estrous cycle if the mare had a pre-existing acute form of endometritis. To that effect, pregnancy rates in mares were dramatically increased on the second insemination of an estrous cycle (i.e., 12 hours following insemination of 1 × 109 killed spermatozoa in semen extender or infusion of semen extender only to induce an endometritis) when the inseminate contained washed spermatozoa in seminal plasma (17/22; 77%), as compared with washed spermatozoa in semen extender only (1/22; 5%).389 Clement et al.390 reported embryo-recovery statistics for mares inseminated two or three times in an estrous cycle (at 2-day intervals) until ovulation occurred, using different stallions for each insemination. For mares inseminated twice in an estrous cycle, 14 of 17 embryos recovered resulted from the first insemination (82%), whereas when mares were inseminated thrice in an estrous cycle, only one of six embryos recovered resulted from the first insemination (17%). Therefore, the first insemination was not uniformly the most fertile. This study was not designed to evaluate the effect of seminal plasma on pregnancy rates, and separation of insemination times by 48 hours may have resulted in a uterine environment that was not as hostile as that reported by Troedsson et al.389 Seminal plasma concentration of inseminates was not reported in this study, but is presumed to be > 5%. In a study reported by Metcalf,391 mares were inseminated with frozen–thawed semen from two different stallions during an estrous cycle at a 4- to 10-hour interval, followed by parentage determination of the resulting foals.391 Of nine foals born, four were the result of the first insemination (44%) and five were the result of the second insemination (56%). While the second inseminate likely contained some seminal plasma, the amount would have been relatively low, if the semen were processed for freezing in a standardized manner. The seminal plasma concentration of the frozen semen was not provided in the report. Based on the combined data from these three reports, it would appear that seminal plasma does provide some protective effect for spermatozoa entering an inflamed uterine environment, but the seminal plasma concentration in the inseminate can be quite low and still achieve this effect. Regardless of the method(s) used to promote uterine transport of equine spermatozoa, in the end only a small fraction of inseminated sperm reach the oviductal luminae. In one report, only
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0.0006–0.0007% of inseminated sperm were recovered from oviductal flushings of mares at 18 hours following intrauterine insemination.366 Barring oviductal occlusion, deposition of spermatozoa in the uterine body of mares leads to similar spermatozoal numbers entering either oviduct.366, 368 This finding suggests that chemotactic effects are not present to attract uterine spermatozoa to the oviduct ipsilateral to a periovulatory follicle, nor is a signal elicited to induce differential control of the oviductal papillae. The process of spermatozoal passage into the oviduct, however, does not appear to be a passive one, i.e., controlled only through the reproductive tract of the female. Studies to date suggest that a dynamic interaction occurs among spermatozoa, reproductive fluids, and the epithelial surface of the UTJ and caudal isthmus of the oviduct.392 Based on studies with rats, it seems that only motile spermatozoa can negotiate passage through the UTJ.393 Spermatozoa tend to associate closely with the epithelium while traversing through the oviductal papilla of the mare,392, 394 a phenomenon also observed with other species.395 Cross-talk between spermatozoa and the epithelium is further manifested by studies with transgenic mice lacking a gene for two different spermatozoal surface proteins. In these males, all functional characteristics of the spermatozoa appear normal, except that spermatozoal migration into the oviducts is hampered.396, 397 A preferential selection process occurs for the few spermatozoa that successfully pass through the UTJ into the protective confines of the caudal isthmus. Scott et al.394 reported that over 90% of spermatozoa visualized by scanning electron microscopy at the UTJ were morphologically normal, even when inseminates contained a high percentage of spermatozoa with morphological abnormalities. The most common morphological abnormality noted in the bound spermatozoa was a proximal cytoplasmic droplet. All of the spermatozoa were noted to have normal head morphology. A high incidence of normal morphological features in oviductal spermatozoa has also been reported in cattle,398, 399 even when the inseminate contained a high proportion of spermatozoa with abnormal heads.399 A recent report conveyed that cytoplasmic droplets in boar spermatozoa may impede binding to the oviductal epithelium under in vitro conditions.400 Thomas et al.401 ably demonstrated that co-culture of equine spermatozoa with oviductal epithelial cell monolayers yielded higher percentages of bound morphologically normal and motile spermatozoa than were present in the neat semen. When using oviductal explants to assess sperm–oviductal interactions, this team of investigators also found that more equine spermatozoa were bound to isthmic explants than to ampullar implants,
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and that more spermatozoa bound to explants procured during the periovulatory period, as compared with the luteal phase of the estrous cycle.402 The caudal isthmus is generally considered to be the reservoir site for oviductal spermatozoa,367 although the UTJ also appears to harbor a considerable number of spermatozoa.403 The effect of the harsh post-inseminate conditions within the uterus on these spermatozoa requires further study. The existence of a spermatozoal reservoir in the mare is further supported by the documentation in the literature that spermatozoa of some fertile stallions can persist in the mare for up to 6 days prior to ovulation, yet result in establishment of pregnancy.404, 405 The mechanism of spermatozoal attachment to the oviductal epithelial cells has received considerable study and appears to consist of a specific sperm–ligand interaction involving glycoconjugates (i.e., lectin-like molecules on the surface of spermatozoa interacting with carbohydrate-containing moieties on the surface of oviductal epithelial cells).406–412 Recently, galactose-binding proteins were characterized on equine spermatozoa that may prove to be involved in the sperm–oviductal binding mechanism.413 This intimate oviductal cell contact with spermatozoa in the oviductal reservoir seems to play two divergent roles: assisting with the final maturational events of spermatozoa that must occur prior to fertilization of an oocyte,414–416 while also maintaining spermatozoa in a viable quiescent state to allow for an extended storage period.417–420 Binding of equine spermatozoa to oviductal epithelial cells under in vitro culture conditions results in both a quantitative and a qualitative change in protein synthesis and secretion by the epithelial cells.421 Purified oviductal glycoprotein and polypeptides secreted by oviductal epithelial cells have been shown to have a positive effect on spermatozoal capacitation, including sperm–oocyte interactions.422, 423 Peripheral proteins of the oviductal membrane appear to contribute to maintenance of boar spermatozoal viability.424 Stallion spermatozoa that are bound to oviductal epithelial cells in culture exhibit flagellar motion for up to 4 days, with gradual release of spermatozoa during that time frame.425 The oviductal support system and gradual spermatozoal release pattern are consistent with the concept of ensuring that appropriately prepared spermatozoa are available to ‘greet’ an ovulated oocyte soon after its arrival in the oviduct. Spermatozoa attached to the isthmic epithelium will subsequently detach and traverse the oviduct to the vicinity of an oocyte. The mechanism for detachment is speculative, but likely involves acquisition of hyperactivated spermatozoal motility to break the connection with the oviductal epithelial cells.426 The lectin-like molecules on the spermatozoal surface
responsible for specific binding with the oviductal cells may also be released during the capacitation process, thereby assisting with spermatozoal detachment.427 Detachment of spermatozoa probably yields cells that are both hypermotile and primed for spermatozoon–oocyte interaction. Spermatozoal capacitation In mammals, freshly ejaculated spermatozoa are not immediately capable of fertilizing an oocyte. Early studies demonstrated that spermatozoa require residence time in the female reproductive tract to gain this capability,428, 429 later termed capacitation.430 Subsequent studies have revealed that similar changes are required by spermatozoa subjected to fertilization by in vitro methods [i.e., conventional in vitro fertilization (IVF)], as reviewed by Yanagimachi.431 As such, capacitation is considered to be an absolute requirement of mammalian spermatozoa destined to fertilize an oocyte without mechanical assistance [i.e., by intracytoplasmic sperm injection (ICSI)]. The process involves a series of preparative biochemical and biophysical changes in the spermatozoon that prime it to respond to signals originating from the oocyte and its surrounding cumulus. If capacitation is incomplete, spermatozoa are unable to penetrate the cumulus matrix,432, 433 and have greatly reduced ability to penetrate the ZP, undergo a zona-induced acrosome reaction, and fuse with the oolema.434, 435 Two early laboratory hallmarks for verifying completion of capacitation were acquisition of a hyperactivated pattern of motility and inducibility of the acrosome reaction by various stimulants (Figure 97.35). While capacitation has been defined by some as the changes that render a spermatozoon capable of undergoing the acrosome reaction,436–439 spermatozoa are also known to exhibit a hyperactivated state of motility when exposed to capacitating conditions.440–442 The term was originally defined as the physiological changes of the spermatozoa in the female genital tract before they are capable of penetrating and fertilizing eggs;443 thus, hyperactivated motility and the acrosome reaction would both be considered components of capacitation, based on its original meaning. Even though the acrosome reaction116, 444 and hyperactivated motility441,445–448 are important for fertilization, the kinetic and temporal relationships of these two features are not well understood.449, 450 The signaling pathways for hyperactivated motility and the preparative changes required for the acrosome reaction appear to be divergent.150, 171, 441, 451, 452 Interestingly, human spermatozoa that undergo hyperactivated motility are predominantly cells with normal morphology.453 In recent years, a variety of molecular markers have
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Figure 97.35 Examples of traditional and alternative biomarkers for assessing spermatozoal capacitation.
been identified that allow more critical evaluation of the molecular events that emerge during capacitation (Figure 97.35).454–477 Unfortunately, many of these assays can be technically challenging or have not become feasible for clinical use.478 Neither the precise signaling mechanisms that govern the attainment of capacitation nor the exact cellular characteristics of a capacitated spermatozoon have been fully elucidated despite considerable investigation in this area over the past few decades. Nonetheless, some key signaling pathways and cellular events have been identified, and include surface (membrane) alterations, cytoskeletal modifications, influx and efflux of cytosolic constituents, and a myriad of enzymatic processes.479 The reader is directed to several reviews on this topic.431, 451, 452,479–490 Interestingly, mature spermatozoa (i.e., those in the caudae epididymis or in ejaculated semen) appear to be pre-programmed to undergo capacitation and the process can be induced by a variety of signals, as opposed to a specific molecular event. This is exemplified by the fact that in vitro culture of spermatozoa in many different media types can evoke capacitation. It appears that nature has provided spermatozoa with a broad array of capacitation induction alternatives to ensure that they may retain fertilizing potential even if some evokers are rendered non-functional. Figure 97.36 provides the reader with some of the proposed signal transduction pathways
for spermatozoal capacitation. Although the literature is replete with potential activation mechanisms, in our view those presented in Figure 97.36 represent an accurate reflection of the most widely accepted signaling pathways. Species differences are known to exist in these regulatory pathways, even among eutherian mammals. Few in-depth studies have been conducted with stallion spermatozoa, so much of our predictions must be extrapolated from studies performed with other species. Conclusions drawn from work in non-equids may be misleading, and thereby require verification through experiments involving stallion spermatozoa. It is also important to point out that ejaculates contain a heterogeneous population of spermatozoa and the cells do not undergo capacitation in a highly synchronous fashion. An advantage gained by such an arrangement is the continuous replenishment of capacitating spermatozoa available in the oviduct to await exposure to an ovulated oocyte, because spermatozoal lifespan is shortened appreciably by the capacitation process. Spermatozoa are bathed in epididymal and ejaculatory fluids that contain ‘decapacitation factors’491 that become adsorbed to the surface of the plasma membrane.450,492–496 Although these factors and their regulatory roles have not been fully elucidated, they aid in suppressing premature initiation of the capacitation process during spermatozoal passage to the isthmic portion of the oviduct.479 Desorption of these
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extracellular factors may unmask some key signaling mechanisms for capacitation.497, 498 A cysteine-rich protein family, termed CRISP, associate with spermatozoa in the epididymis as well as through the seminal plasma.104,499–504 These proteins have been shown to inhibit tyrosine phosphorylation and capacitation of spermatozoa,505 possibly through their ability to block ion channels within the plasma membrane.104 Alterations in membrane-associated cholesterol, and cytosolic concentrations of bicarbonate and calcium ions ([HCO3 − ]i and [Ca2+ ]i , respectively) appear to be central evokers of capacitation in spermatozoa studied under in vitro conditions,141, 183, 454, 458, 470, 488,506–523 and the same likely applies in vivo. Efflux of cholesterol from the plasma membrane requires the presence of sterol acceptors in the milieu, such as albumin or lipoproteins.483, 497, 508, 514, 523 Under laboratory conditions, bovine serum albumin483, 514, 523 or β-methyl cyclodextrin524–526 are generally used to achieve this effect. The net result of cholesterol efflux appears to be destabilization and increased fluidity of the plasma membrane, combined with externalization of key surface receptors.478, 478, 508, 515, 527, 528 The cholesterol efflux is hypothesized to result in an increased permeability of the membrane to HCO3 − and Ca2+ .520 Both of these ions are thought to enter the cytosol from the extracellular space through ion-specific channels.529, 530 A variety of channels, transporters, and stores exist in spermatozoa that aid in regulation of [Ca2+ ]i .480, 529 Voltage-dependent Ca2+ channels (VDCCs) exist in the plasma mem-
brane of spermatozoa, and membrane depolarization mediates Ca2+ entry through these channels.529, 531 Additionally, an increase in cytosolic pH ([pH]i ) markedly increases the activity of VDCCs.532 Internalization of HCO3 − has been shown to further enhance Ca2+ entry through a PKA-dependent mechanism.533, 534 Interestingly, Ca2+ -Pi symporters535, 536 and Na+ /HCO3 − cotransporters140 are also known to exist in mammalian spermatozoal membranes and are thought to be involved in the capacitation process. Plasma membrane Ca2+ -ATPase (PMCA), an enzyme pump that extrudes Ca2+ , is activated by the presence of decapacitation factors. In their absence, PMCA activity is decreased, resulting in a net increase in [Ca2+ ]i .537–539 A family of transmembrane Ca2+ -selective ion channels, called CatSpers, has been identified in recent years. These channels are thought to play an important role in both activated (non-capacitated) and hyperactivated (capacitated) motility.540, 541 CatSper2 channels have been localized to the flagellum,542 and CatSper1 channels have been detected in the principal piece of the flagellum.543 Receptors for adenosine, calcitonin, and fertilization-promoting peptide (seminal plasma constituents) have been identified in the plasma membrane of spermatozoa, and appear to play a modulating role in capacitation through regulation of membrane-associated adenylyl cyclase (mAC). These molecules activate mAC in uncapacitated spermatozoa but downregulate mAC in capacitated cells, possibly to prevent ‘overcapacitation’ of cells.544 Another method of preventing overcapacitation
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell of spermatozoa is the polymerization of actin.545 These molecules have been identified in many areas of the spermatozoon, including the periacrosomal space, the equatorial and post-acrosomal regions, and the flagellum.546–548 Actin polymerization is a phospholipase D (PLD)-dependent event, and this enzyme is regulated by both PKA and protein kinase C (PKC).549, 550 The formation of a filamentous form of actin in the periacrosomal space may prevent premature fusion of outer acrosomal and overlying plasma membranes when the properties of these two membranes become more fusogenic during the capacitation process. The actin must depolymerize for the acrosome reaction to occur.545 An increase in [HCO3 − ]i and [Ca2+ ]i activate a soluble cytosolic form of adenylyl cyclase (sAC), leading to elevated intracellular concentration of cyclic adenosine 5 -monophosphate (cAMP).167, 463, 551 This nucleotide subsequently activates cAMPdependent PKA activity, eventually leading to an upregulation in protein tyrosine phosphorylation.520 A large number of proteins are known to be tyrosine phosphorylated during capacitation, but the roles of many remain unclear.552 Indeed, tyrosine phosphorylation of spermatozoal proteins appears to be crucial to acquisition of a capacitated state.489 Tyrosine phosphorylation of head proteins occurs during capacitation, but this activity is even more pronounced in the flagellum,477, 553 thereby suggesting an importance of these proteins in hyperactivation.554 Two members of a tyrosine-containing family of proteins, termed ‘A kinase anchoring proteins’ (AKAPs), are localized in the fibrous sheath and appear to tether PKA and various other signaling enzymes of the flagellum. These AKAP members are phosphorylated during capacitation and are thought to provide regional control of signal transduction.187, 523, 554, 555 Similarly, a calcium-binding protein, CABYR, has been localized to the fibrous sheath and is known to be tyrosine phosphorylated during capacitation.556 Phospholipid hydroperoxide glutathione peroxidase (PHGPx) has been identified in the mitochondrial capsule of the spermatozoal midpiece and tyrosine phosphorylation of this enzyme during capacitation may play a role in hyperactivated motility.452 The phosphotyrosine protein valosin-containing protein is known to redistribute from the neck region to the anterior head region and may be involved with the acrosome reaction.552 Other phosphotyrosine proteins may be involved with recognition of the ZP and sperm–zona binding.557 Another spermatozoonspecific enzyme, scramblase, may be activated through the PKA–tyrosine phosphorylation pathway, leading to a collapse of the asymmetric bilayer distribution of phospholipids, lipid packing disor-
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ders, and phospholipase activation.481, 513,557–559 Some workers consider the HCO3 − –cAMP–PKA–tyrosine phosphorylation (scramblase) pathway to precede an efflux of cholesterol from the plasma membrane.472, 481, 513,557–560 Flippases, floppases, and an amnophospholipid transporter localized to the acrosomal region of mouse spermatozoa are thought to be important in maintaining the lipid asymmetry of the lipid bilayers prior to capacitation by establishing higher concentrations of phosphatidylserine and phosphatidylethanolamine in the inner leaflet and higher concentrations of sphingomyelin and phosphatidylcholine in the outer leaflet.561 Another probable pathway to tyrosine phosphorylation involves formation of ROS that trigger downstream events that lead to capacitation. The effects of superoxide anion (O2 ·) and H2 O2 on spermatozoal capacitation were first described 15 years ago.562, 563 More recently, nitric oxide (NO·) has been shown to regulate capacitation.564, 565 These findings have stimulated intensive investigations regarding the physiological roles of ROS in this process.266, 268, 272, 485,566–576 Current evidence supports the concept that ROS induce the cAMP-mediated tyrosine-phosphorylation cascade,577 but additional routes of action are also quite likely.485, 578 As indicated above, the plasma membrane undergoes considerable remodeling during capacitation, with an appreciable efflux of sterols and transverse asymmetry of phospholipids. These events increase the fluidity of the membrane, thereby allowing horizontal redistribution of membrane molecules. Recent studies reveal that lipid rafts participate in this redistribution process.579 These cholesterol-rich microdomains are laden with signaling molecules that are known to be involved with capacitation and zona binding.580, 581 The capacitation process can increase the number of membrane rafts, as well as their affinity for zona binding.582 Actin polymerization may play a role in association/dissociation of the membrane rafts.583 This has become an intense area of study, as these signal-carrying lipid rafts appear to play an increasingly important role in capacitation and spermatozoon–oocyte interaction.584, 585 The list of other potential endogenous signaling mechanisms is extensive, including involvement of mitogen-activated protein kinase (MAPK),566 extracellular signal-regulated kinases (ERK),586 calmodulin-dependent kinases,182, 587 protein tyrosine K (PTK),485, 588 PKC,549 Na+ /Ca2+ exchangers,589–591 Na+ /K+ ase,482 KATP channels,592 and the renin–angiotensin system.593–596 This list is not meant to be all inclusive, but to demonstrate that capacitation involves an extremely complex series of molecular events. Future investigations will more
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Figure 97.37 Some documented and hypothesized molecular events associated with the spermatozoal acrosome reaction. Please refer to the text for an explanation of these signaling pathways and for figure abbreviations.
clearly define the precise molecules and mechanisms that control capacitation, including spatial, temporal, and kinetic patterns. Acrosome reaction Regulated acrosomal exocytosis is a fundamental element of fertilization, and might be a more appropriate defining term than ‘acrosome reaction’.116, 597, 598 It must be preceded by a complex series of events, including spermatozoal capacitation, followed by spermatozoon–oocyte recognition, spermatozoon–zona binding, and elicitation of signaling pathways.431,599–601 An excellent review of the acrosome reaction was recently provided by Bailey,554 and readers are referred to this source for an in-depth account of the biochemical mechanisms involved in the process. Other resources are also available for those who desire a thorough grasp of the subject.116, 444, 451, 481, 484, 602 Capacitation primes a spermatozoon for the acrosome reaction through alterations in the membrane lipids; membrane hyperpolarization; relocation, aggregation, and externalization of membrane-signaling molecules; and changes in cytosolic and cytoskeletal components. A trigger, however, is required for induction of the acrosome reaction, and most experimental evidence suggests that the ZP provides the inciting factor.602–604 Spermatozoal interaction with the zona appears to involve a precise species-specific receptor–ligand interaction, although the exact identity of the spermatozoal receptor remains elusive. Evidence abounds, however, that designated glycoproteins of the ZP, termed ZP3, are responsible for initiating the signaling cascade(s) that lead(s)
to the acrosome reaction.1, 444, 601, 603,605–611 The exact signal transduction pathway(s) for acquisition of the acrosome reaction is (are) not fully understood, but decades of intensive study have yielded much enlightening information on the subject. Figure 97.37 conceptualizes possible pathways involved. As with capacitation, it is quite possible that more than one reaction cascade can elicit the acrosome reaction. The ZP is an extracellular glycoprotein matrix surrounding the oocyte. In most mammalian species studied, excluding humans,612, 613 the matrix is composed of only three glycoproteins, typically termed ZP1, ZP2, and ZP3.609, 611, 614 The peptide component of these glycoproteins appears to be well conserved across species, but post-translational modifications (i.e., glycosylation) lead to species heterogeneity and, hence, species-specific binding affinity.615–619 The interactions appear to involve recognition of specific carbohydrate moieties present on the polypeptide backbone of ZP3 by complementary zona-binding proteins of the plasma membrane of the spermatozoon.601, 615, 620, 621 In the mouse, β1,4galactosyltransferase I (GalT I) is the putative receptor on the spermatozoal surface for a glycoside ligand of ZP3, the binding of which leads to induction of the acrosome reaction.621–627 A complex of spermatozoal surface molecules is likely involved in spermatozoon–zona interaction.628 Whereas ZP3 can be considered the most likely signaling molecule of the ZP for induction of the acrosome reaction, various experiments also suggest that ZP2 may be involved in the anchoring of acrosome-reacted spermatozoa to the zona.605, 620 The ZP2 and ZP3 exist as heterodimers in filamentous arrangement cross-linked by ZP1, so it seems a fitting motif for integrated
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell roles of ZP2 and ZP3 in spermatozoon–oocyte interaction.444, 606,629–631 As evidence that the intricacies of spermatozoon–zona interactions are not yet resolved, a recent report revealed that, in bovine spermatozoa, glycosylation of ZP3 may not be an absolute requirement for induction of the acrosome reaction.632 Extensive literature indicates that elevated intracellular Ca2+ is a steadfast requirement of the acrosome reaction. Spermatozoa utilize a range of intracellular messengers for various functions. Cytosolic Ca2+ is certainly a key regulator in these processes and is subject to complex and sophisticated spatiotemporal control so that Ca2+ -sensitive responses can be activated discretely within various compartments of the cell.529, 633 A capacitated spermatozoon has an increased [Ca2+ ]i ,484 created primarily by influx of transients through voltageoperated Ca2+ channels in the plasma membrane. Exposure of spermatozoa to solubilized ZP has been shown to result in a further increase in [Ca2+ ]i transients. Although intracellular Ca2+ stores are thought to be minimal, the acrosome may serve as an intracellular Ca2+ depot that is subsequently mobilized during the acrosomal reaction cascade.484, 634 The efflux of such intracellular stores may then activate store-operated channels (SOCs) in the plasma membrane,529 probably transient receptor potential (TRP) channels,635 thus creating the sustained rise in intracellular Ca2+ that is required to drive the acrosome reaction.636–638 The mechanism of TRP channel activation is unresolved, but both depletion of Ca2+ from internal stores and direct receptor activation have been proposed as methods of activation.636, 639 The TRP channels have been expressed in both the head and flagellar regions of spermatozoa, suggesting that they may play a role in both the acrosome reaction and in hyperactivated spermatozoal motility.640 Receptor-activated Gi proteins appear to play a central role in the acrosome reaction, as evidenced by their activation by ZP3–GalT I binding,641 and their inactivation by pertussis toxin (which inactivates Gi proteins).642, 643 Activation of Gi proteins leads to a rise in [Ca2+ ]i .642 Spermatozoon–ZP3 adhesion directly evokes Ca2+ influx via low-voltage-activated calcium channel (T-type channel) activation in membranes. Hyperpolarization of the membrane potential, as occurs during capacitation, is thought to release the channels from inactivation in order that they may respond effectively to a stimulus derived from the ZP,462, 480, 592, 644 i.e., membrane potential changes are translated into intracellular Ca2+ signals.531 A ZPmediated increase in mAC has also been reported to occur via Gi protein activation,645 presumably resulting in the increase in cAMP that is known to occur following spermatozoal exposure to the ZP.645, 646 Evidence for a cAMP/PKA-mediated acrosome re-
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action exists following Ca2+ entry into the cell,647, 648 but little is known about the role of this pathway in the acrosome reaction. Breitbart and co-workers suggested that PKA may activate a voltage-dependent channel in the outer acrosomal membrane that leads to release of Ca2+ stores from the acrosome.634, 649, 650 Zona-induced acrosomal exocytosis can be blocked by an inhibitor of PKA.651 The phospholipase C (PLC) family appears to be central to the events leading to the acrosome reaction, and these enzymes have been localized to the spermatozoal head region.652 Activation of PLC requires Ca2+ ,604, 653, 654 but at relatively low concentrations,451 as might be produced by the pathways in the preceding paragraph. Zona-mediated coactivation of membrane-bound PLCβ1 (through a Gi-protein-coupled receptor) and membrane-bound PLCγ [through a tyrosine kinase (TK) receptor] has been proposed.634, 652, 655 Activation of PLC is thought to promote the acrosome reaction through a couple of routes. Firstly, PLC activates hydrolysis of phosphatidylinositol 4,5-biphosphate (PIP2 ) within the membrane, resulting in production of two active intracellular messengers, diacylglycerol (DAG) and inositol 1,4,5-triphosphate (IP3 ). DAG mediates the activation of PKC, and IP3 releases Ca2+ from intracellular stores, i.e., the acrosome.451, 655 Activation of PLC may also lead to disassembly of F-actin filaments that are formed between the outer acrosomal membrane and overlying plasma membrane during capacitation, presumably to separate these increasingly fusogenic membranes.484, 634, 656 PIP2 is purported to inhibit actin-severing proteins. Another member of the PLC family, PLCδ4, plays a central role in the acrosome reaction. This enzyme appears to be important for mobilizing Ca2+ stores in the acrosome and also activating SOCs in the plasma membrane, thereby creating a sustained increase in cytosolic Ca2+ concentration.484, 634,655–659 The IP3 receptor and PLCδ4 have been localized to the acrosome.655, 657 and both probably play a role in TRP channel regulation of Ca2+ entry into the cell.635,660–664 The role of the PKC, as a signaling pathway in the acrosome reaction, remains somewhat unclear. The PKC activity of spermatozoa is dramatically lower than that detected in some other tissues,665–667 but, nonetheless, has been identified in various regions of the spermatozoa.666, 668 Certainly, DAG, produced through the activity of PLC, is known to be a potent activator of PKC,665, 669 possibly leading to opening of a Ca2+ channel in the plasma membrane.634, 649, 650 DAG is also a membrane-destabilizing molecule that increases membrane fusibility.451, 670, 671 Experimentation has revealed that phosphorylation of spermatozoal proteins occurs upon spermatozoal stimulation by progesterone, a known inducer of the acrosome
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reaction, and the activity can be mimicked with various activators of PKC and inhibited by various inhibitors of PKC.672 Synaptotagmins are a family of calcium-binding transmembrane proteins purported to be calcium sensors responsible for triggering exocytosis in various cells.673, 674 Synaptotagmin VI has been localized to the outer acrosomal membrane of human spermatozoa.675 Recently, PKC-mediated phosphorylation of spermatozoal synaptotagmin VI has been demonstrated.672 These workers proposed a model by which PKC has a regulatory role in synaptotagmin function through phosphorylation of synaptotagmin C2 domains;672, 673 Hence, PKC may play a vital role in controlling the molecular machinery that mediates membrane fusion (discussed below). Phospholipase A2 (PLA2 ) is recognized as an important enzyme for the acrosome reaction. Certainly, indirect evidence suggests that treatment of spermatozoa with PLA2 inhibitors will impede this event.676–679 Both Ca2+ and DAG (in the presence of Ca2+ ) are capable of activating PLA2 .654,680–683 The resulting enzymatic action is deacylation of fatty acids from phospholipids, producing fatty acids (such as arachidonic acid and lysophospholipids). A key target is phosphatidylcholine, yielding lysophosphatidylcholine. This ‘metabolite’ is a likely contributor to the acrosome reaction, as it is known to trigger the reaction in capacitated spermatozoa.684–687 Exposure of capacitated spermatozoa to solubilized zona results in increased metabolism of phosphatidylcholine to arachidonic acid and lysophosphatidylcholine, resulting in a high proportion of acrosome-reacted cells. The PLA activation is regulated by a signal transduction pathway involving Gi protein and DAG.682 Lysophosphatidylcholine, or similar metabolites of PLA2 activity, appear to be important to the final stages of cAMP-mediated acrosomal exostosis.688 Regulated exocytosis, as it relates to membrane fusion at neural synapses, has been studied extensively.689 Expanding these investigations to spermatozoa has revealed that a similar mechanism of action occurs between the outer acrosomal and plasma membranes,672,690–692 culminating in release of acrosomal contents and exposure of previously hidden domains on the inner acrosomal membrane. Studies to date indicate that the machinery required for this exocytotic event (which requires the finely regulated merger and fusion of two membranes) includes several core components: (i) soluble Nethylmaleimide-sensitive factor attachment protein receptor (SNARE) proteins; (ii) N-ethylmaleimidesensitive factor (NSF); (iii) soluble NSF attachment protein (α-SNAP); (iv) Rab-GTPase; (v) synaptotagmin; and (vi) calcium ions. This basic machinery is considered obligate for acrosomal exocytosis.690, 693
Of interest, the SNARE proteins, NSF, and synaptotagmin have recently been identified immunocytochemically in stallion spermatozoa, with predominant staining in the regions of the acrosomal cap and equatorial segment.694 The SNARE proteins are functionally classified as v-SNAREs and t-SNAREs, and membrane fusion is executed when v-SNAREs on the acrosomal (i.e., vesicle) membrane pair the complementary t-SNARES on the plasma (i.e., target) membrane to form trans-SNARE complexes or SNAREpins. Both vand t-SNARE proteins are thought to be secured to their respective membranes by hydrophobic anchors that span both leaflets of the lipid bilayers.689, 695, 696 NSF and its cofactor, α-SNAP, are cytosolic proteins that act cooperatively to regulate SNARE protein function. The α-SNAPs bind to the SNARE protein so as to correctly position NSF, and are responsible for stimulation of the ATPase activity of NSF which dissociates trans-SNARE complexes.697–701 This is a mechanism for recycling of components of the fusion machinery in other cell types, but would not appear necessary in the acrosome, where the reaction is a one-time event. Recent work with spermatozoa indicate that they also serve essential prefusion roles,691, 692 probably by inducing conformational changes in the SNARE protein complexes, i.e., catalyzing disassembly of an inactive cis-SNARE complexes, to allow re-association as active trans complexes.691 The NSF has also been reported to act in a pre-fusion step in non-spermatozoal cells by catalyzing SNARE protein-induced membrane rearrangement that is more suitable for Ca2+ -mediated fusion.702 Rabs are a family of proteins that are included in the Ras superfamily of G proteins. They are anchored to the acrosomal membrane, a process augmented by cholesterol efflux,703 and, upon activation [via guanosine triphosphate (GTP) binding], can interact with Rab effector molecules on the plasma membrane to form an initial physical link, allowing the v-SNARE and t-SNARE proteins to interact to form a trans-SNARE complex.689, 704, 705 The assembly of the trans-SNARE complex is thought to generate sufficient energy to overcome the repulsive forces generated by the polar head groups of the two membranes.689 In essence, the acrosome becomes tethered to the plasma membrane. It is possible that that the filamentous F-actin interspersed between the outer acrosomal and plasma membranes could dampen the repulsive forces.451 Epac, a known guanine nucleotide exchange factor (i.e., catalyzes replacement of GDP with GTP on Rab proteins), is activated by cAMP, resulting in induction of the acrosome reaction;706 hence, another signaling cascade enters the mix of possible regulatory mechanisms for the acrosome reaction.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell Ca2+ -binding synaptotagmin, also a major Ca2+ sensor, is central to both the spatial and the kinetic elements of the fusion process. Synaptotagmin is known to have direct interaction with t-SNARE and SNAP proteins, a feature augmented by Ca2+ , and may actually assist in SNARE complex formation.673, 707 This protein is a large transmembrane protein whose cytosolic component is primarily C2 domains, similar to that of PKC isoforms.673 In other cell types, the protein is thought to facilitate docking of secretory vesicles close to Ca2+ channels where they might be exposed to high local concentrations of this ion.673, 708, 709 As mentioned above, studies with spermatozoa indicate that synaptotagmin may also inhibit spontaneous fusion, through regulation by PKC. This control is likely to be driven by [Ca2+]i . Without doubt, Ca2+ is the driving force for the final stages of acrosomal preparedness for reaction, from Rab3 activation through final pore formation.691 It appears that Ca2+ is maintained in locally low concentrations during SNARE complex formation, possibly through synaptotagmin control. When transSNARE complexes are fully assembled for the final fusion event, synaptotagmin likely triggers release of Ca2+ .673, 691 Studies involving non-spermatozoal cells reveal that the docking mechanism pulls the pre˚ 691, 710 fusion membranes to within a distance of 2–3 A. Subsequent bridging of adjacent phosphate head groups by Ca2+ allows regional intermembranous displacement of water (i.e., by calcium hydration) so that the resulting anhydrous complex is amenable to lipid mixing.691,710–712 The regions of fusion are restricted by the circular array of SNARE complexes.691, 711 Similar mechanisms probably exist for spermatozoa since the basic machinery is similar to that of somatic secretory cells. As with capacitation, the precise pathways involved in the acrosome reaction are not fully elucidated despite considerable study in this area, and species differences are likely. The potential for involvement of signaling pathways and endogenous molecules, other than those listed above, is exemplified by the potential role of angiotensin II in the acrosome reaction of stallion spermatozoa.593, 594
Hyperactivated motility A hyperactivated form of motility is required to free the spermatozoa from the oviductal reservoir (through spermatozoal release from the oviductal epithelium and subsequent passage through the mucinous environment of the oviduct) and to penetrate the cumulus matrix and ZP surrounding the oocyte.149, 152, 441, 445 It is of interest that hyperactivated spermatozoa tend to have normal morphological characteristics.453 When observed in aqueous media,
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the motility pattern of a hyperactivated spermatozoon is characterized by a high-amplitude, often asymmetric, whiplash flagellar beat pattern, leading to a circular or non-progressive trajectory.149, 152, 441 Studies have revealed that this flagellar wave form actually improves the progressivity of spermatozoa over that of activated motility when spermatozoa are exposed to viscoelastic conditions such as those that exist in the oviduct.441, 713 Activated motility and hyperactivated motility may require different environmental signals. Presumably, a signal for hyperactivated motility is elicited within the oviduct under natural conditions in order to initiate the event at a time that is conducive to fertilization.441 Although the precise signal(s) for initiation of hyperactivation remain(s) unsolved, it is possible that chemotactic and/or thermotactic factors serve in this capacity.714 It is also possible that spermatozoal exposure to alkalinizing conditions, as exists in the oviduct and above that which initiates activated motility, is the primary initiator of hyperactivated motility.150, 152, 715, 716 The signaling pathway is also dissimilar between activated and hyperactivated forms of motility. Activated motility primarily utilizes the cAMP–sAC–PKA pathway and requires a relatively low concentration of Ca2+ for phosphorylation of flagellar proteins (Figure 97.34). Conversely, hyperactivated motility appears to require an elevated concentration of Ca2+ ,148, 164,717–719 and no cAMP149, 150 or sAC.166 The precise mechanisms controlling hyperactivated motility are subject to continued investigation, but an increasing body of literature suggests that an increase in extracellular pH leads to an increase in [pH]i , thereby potentiating the action of Ca2+ channels.147, 467, 716, 718, 720 The most likely mechanism for an increase in cytosolic Ca2+ is through CatSper channels, and through release from internal stores. Four CatSper channels (CatSper1–4) have been localized to the principal piece and they appear to have a physical interaction.716 One report suggests that all four CatSper channels must be operational in order for hyperactivation to occur;716 however, others provide evidence that this may not be the case.721 CatSper1 may serve as an internal pH sensor, as evidenced by the composition of its N terminus.147, 543 The redundant nuclear envelope (RNE) at the base of the flagellum has been identified as a possible internal store for Ca2+ . This structure represents the remnants of the nuclear membrane following spermiogenesis. The RNE is reported to contain IP3 receptor-gated Ca2+ channels which are thought to release Ca2+ stores and regulate hyperactivated motility independent of similar channels in the acrosome.719, 722 As the RNE is located at the base of the flagellum, where flagellar bending is propagated, it seems a
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logical location for an internal Ca2+ store.138, 721 A specific activator of PKC has been shown to induce hyperactivated motility in human spermatozoa;723 thus, a PKC signal transduction pathway may also be important to stimulation of hyperactivated motility. Although Ca2+ is known to affect the bending pattern of the flagellum, and added Ca2+ is required for hyperactivated motility, the mechanism by which Ca2+ controls this event remains unknown. The signaling pathways and cellular events leading to hyperactivated motility and to the acrosome reaction are different, and the events of each can occur independently.138, 148 Nonetheless, internal alkalinization appears to be a common inciter for the two events. The pathways, although not the same, achieve the critical spermatozoal priming required for interaction with the oocyte. As capacitation was originally defined with this end point in mind, it seems appropriate to consider hyperactivated motility as a component of the capacitation process. It has been quite difficult, however, to understand the interplay of the various components of capacitation. Indeed, the various events of the capacitation process are not tightly coupled. Penetration of zona-free hamster eggs by human spermatozoa is maximal at 18 hours under in vitro conditions, whereas changes in tyrosine phosphorylation occur after 1–2 hours, and hyperactivation is maximal after 3 hours of incubation.187, 449, 598, 724, 725 Furthermore, different groups have demonstrated variability among men in spermatozoal capacitation time.598, 726, 727 An uncoupling of the acrosome reaction and hyperactivated spermatozoal motility has also been demonstrated in hamsters and bulls.506, 719, 728 Indeed, we now know a great deal about the molecular control of various facets of the capacitation process, but many more discoveries are needed to uncover the secrets of the spermatozoon. Sperm–oocyte interaction While spermatozoal capacitation is not a site-specific phenomenon and can be induced in a variety of artificial media, the caudal segment of the oviduct appears to be the principal location for spermatozoal capacitation and storage under in vivo conditions.436 Interactions with an ovulated oocyte, however, require spermatozoal migration to the ampullar region of the oviduct, and only a small percentage of spermatozoa that gain access into the oviduct will eventually arrive at this fertilization site.436 Such spermatozoa are thought to have achieved full fertilizing potential. The precise mechanisms by which spermatozoa migrate to the ampullar region of the oviduct remain speculative, but contractile movements of the oviduct and hyperactivated sper-
matozoal motility are thought to play key roles in this migratory phase. Evidence is mounting that chemotactic factors are also important to directional migration of oviductal spermatozoa.729–741 Some investigators assert that chemotactic responsiveness is also a part of the capacitation process.729, 730 Spermatozoa possess specific chemotactic receptors.478 Olfactory receptors are considered to represent the largest family of genes in the human genome, with up to 1000 members,742, 743 and odorant receptors have been identified, and genes encoding for olfactory receptors have been expressed, in the testes of mammals.744–750 Olfactory receptor genes have even been identified in the primordial germ cells of fetuses.751 Odorant receptors have been localized to the flagellar base and proximal midpiece of spermatozoa,752, 753 suggesting that odorant-like molecules emanating from the female reproductive tract might induce hyperactivated spermatozoal motility, possibly by release of Ca2+ from internal stores,152, 747 or another signal-transduction pathway.754 Spermatozoa do not appear to acquire a capacitated state, complete with chemotactic competence, in a synchronous manner; rather, a small percentage of a spermatozoal population is thought to achieve this potential at any given time. This is thought to serve as a safeguard to ensure that capacitated spermatozoa, which are short-lived, are available over a protracted period for prompt interaction with an ovulated oocyte.729, 741 Oriented migration of spermatozoa to follicular factors has been demonstrated under in vitro conditions,714, 741, 755 and progesterone is thought to be an important mediator of this event.714, 739 Follicular fluid is not thought to pass down the oviduct to the caudal isthmic region, however, and migrating capacitated sperm can be detected in the more proximal regions of the oviduct prior to ovulation.738 Studies involving oocyte transfer in horses have also demonstrated that fertilization can occur when oocytes are transferred to the oviduct contralateral to an ovary containing a pre-ovulatory follicle or when transferred to oviducts of non-cyclic mares supplemented with exogenous steroids.756 One working hypothesis is that spermatozoa are transported to the general site of fertilization by thermotaxis and contractions of the oviduct and, when in the direct vicinity of the oocyte, are directed by chemoattractants secreted by the cumulus cells.714, 757, 758 Certainly, studies regarding spermatozoal chemotaxis remain scanty at this juncture, but continued investigation may divulge a specific role of chemoattractants in spermatozoal migration patterns. Before a spermatozoon can interact directly with the oocyte, it must first negotiate passage through the two sizable extracellular matrices which surround
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell the oocyte, i.e., the cumulus complex and the ZP. An excellent description of the three-dimensional structure of the ZP was recently provided.759 Acquisition of hyperactivated motility and translocation/exposure of glycosylphosphatidylinositolanchored surface hyaluronidase (termed PH-20) likely permit penetration of the cumulus matrix.760–762 Binding of acrosome-intact spermatozoa to the ZP via species-specific carbohydrate moieties of the ZP and corresponding receptors on the spermatozoal surface lead to induction of the acrosome reaction.605, 609, 610,763–766 An assortment of spermatozoal surface lectins and glycoenzymes have been shown to have zona-binding ability.767–774 Previous studies involving mouse spermatozoa suggest that the binding of the spermatozoal surface receptor, GalT I, to specific oligosaccharide moieties of ZP3 on the ZP is important to induction of the acrosome reaction.608, 775 More recent work, however, indicates that other mechanisms may be responsible for initial adhesion of spermatozoa to the ZP.775–777 A newly voiced theory is that the ZP acts as a spermatozoal scaffold, with ZP2 acting to orchestrate zona permissibility to zona penetration by a spermatozoon, and initial zona penetration by the spermatozoon activating the acrosome reaction through ‘mechanosensory’ signals.778 One report involving horses suggests that spermatozoal surface GalT and zona glycoprotein ZPC (probably similar to ZP3 in the mouse) are not required for spermatozoal binding to the ZP.779 Acquisition of the acrosome reaction was not an experimental end point in this study, so it remains possible that a GalT–ZPC-independent mechanism is responsible for initial adhesion, but that a GalT–ZPC-dependent interaction may be required for the acrosome reaction. Passage through the cumulus may also displace the surface-associated sperm glycoproteins, glycodelin-A and glycodelin-F, to promote spermatozoal–zona binding.780 Not surprisingly, the ZP of ovulated oocytes is also known to contain products secreted by the oviducts that may play a role in initial sperm adhesion.781, 782 Further research may lead to identification of a multitude of receptors that are involved with spermatozoal–zona binding. The cutting thrust issued by hyperactivated spermatozoa,783 possibly combined with externalization/ release of proteases and glycosidases within the acrosome,598, 760, 784, 785 allow spermatozoal penetration of the ZP. Studies involving hamster spermatozoa indicate that the process of spermatozoal adhesion to, and penetration of, the ZP requires only 10–15 minutes.598, 786, 787 Once situated within the perivitelline space, the spermatozoon binds to, and fuses with, the egg plasma membrane (or oolemma), leading to internalization of the sperm haploid chromosomal
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complement by the oocyte. Only spermatozoa that have undergone an acrosome reaction are thought to be capable of fusion with the plasma membrane of the oocyte,788 and this binding/fusion process involves specific domains of the post-equatorial region of the sperm head and the microvillar region of the oolemma (Figure 97.38).760, 789 The specific sperm proteins that are involved with spermatozoal–oocyte binding remain an enigma, although several candidate proteins have been identified.504, 760, 789, 790 A family of sperm-associated cysteine-rich secretory proteins (CRISPs) have been characterized in many mammalian species. These proteins are synthesized and secreted by the epididymis and associate with the spermatozoal membrane, most prominently in the distal corpus and cauda epididymis.504, 791 CRISPs have been localized to the equatorial region of capacitated and acrosome-reacted spermatozoa.792 Interestingly, stallion spermatozoa possess an abundance of CRISPs, with largest amounts detected in ejaculated spermatozoa.503 Three members of the CRISP family have been identified in the testis, epididymis, ampullae, or seminal vesicles of stallions.499, 503 A portion of CRISP is tightly associated with the spermatozoa, and is localized to the post-acrosomal and equatorial regions of the sperm head, as well as the midpiece.503 The regionalized distribution of the protein hints to a potential role in spermatozoon–oocyte interaction; however, complementary molecules have not been characterized on the oocyte membrane of any species studied.504 In addition, the lack of hydrophobic domains in this molecule suggests that it may not be directly involved in membrane fusion events.598, 790 The spermatozoal ADAM (a disintegrin and metalloprotease) family of proteins [including fertilin α (ADAM1), fertilin β (ADAM2), and cyritestin (ADAM3)] have been implicated in spermatozoon–oocyte binding,504, 760 and are known to possess fusogenic potential.793 Nonetheless, gene disruption data do not corroborate an essential role of these proteins in fertilization.760, 794 The protein Izumo has withstood the spatiotemporal, immunological, and gene-knockout tests required to consider it as a probable spermatozoal candidate in gamete fusion.789, 795 Treatment of spermatozoa with antibodies to integrins or osteopontin also reduce spermatozoon–oocyte binding and fertilization in vitro.796 Recently, proteomic analysis of the spermatozoal membrane regions involved in spermatozoon–oocyte interactions has uncovered an abundance of proteins that could be participants, including some previously uncharacterized proteins.797 A tetraspanin protein, CD9, has been implicated as a very viable candidate oocyte protein,608, 760, 788, 789, 798 for spermatozoal adhesion and fusion, although additional proteins might also be involved.798 The
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Figure 97.38 Conceptualization of spermatozoon–oocyte interactions. (a) Spermatozoon negotiates passage through the intercellular matrix of the cumulus oophorus. (b) Initial adhesion of the equatorial region of the spermatozoon with the zona pellucida. (c) Induction of the acrosome reaction. (d) Spermatozoal penetration of the zona pellucida at an oblique angle, with entry into the perivitelline space. (e) Fusion of the post-acrosomal spermatozoal membrane with the microvillar region of the oolemma. (f) Establishment of a gateway for spermatozoal nucleus entry into the cytoplasm of the oocyte, as well as entry of spermatozoon cytosolic molecules (presumably spermatozoon-specific phospholipase C) that serve as a signaling mechanism for activation of the oocyte and exocytosis of oocyte cortical granules. Modified from Primakoff and Myles760 and Rubinstein et al.789
microvilli of the oolemma appear to be key to spermatozoon–oocyte fusion, and CD9 is enriched in the microvillar portion of the oolemma.799 In addition, CD9 may be involved in coordination of microvillar shape and distribution along the oolemma.799 Membrane lipid molecules are also of fundamental importance to spermatozoon–oocyte fusion.800 In addition, polymerization of actin filaments may be critical to spermatozoal incorporation into the oocyte cytoplasm, decondensation of the nucleus, and activation of the oocyte block to polyspermy.545,801–803 Entry of the sperm into the ooplasm elicits an initial increase in intracellular Ca2+ concentration, followed by repetitive Ca2+ oscillations. This signaling mechanism induces exocytosis of oocyte cortical granules (to prevent polyspermy); release of the oocyte from meiotic arrest; pronuclear formation; mediation of genomic union; progression into mitosis
with reorganization of both nuclear and cytoskeletal components; and stimulation of oocyte mitochondrial respiration.804–810 The factor responsible for this activation is derived from the spermatozoal cytosol, and a spermatozoon-specific PLC is considered to be the most likely candidate molecule.811–813 In a manner similar to that of the acrosome reaction, PLC might activate hydrolysis of PIP2 within the membrane, resulting in production of IP3 which, in turn, could mediate release of Ca2+ from intracellular (i.e., endoplasmic reticulum) stores.814 Internal energy generation is required for spermatozoon–oocyte interactions. In the mouse, spermatozoon–oocyte fusion is inhibited in the absence of glucose.233 In this species, glucose appears to mediate tyrosine phosphorylation of proteins in midpiece-specific sites.234 Sperm entry into the mouse oocyte is characterized by pentose phosphate pathway activity and redox regulation, in addition to
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell glycolysis.251, 252 Interestingly, oocytes are incapable of metabolizing glucose; thus pyruvate or cumulus cells (which convert glucose or lactate to pyruvate) must be present in IVF medium.598, 815
Clinical considerations: semen evaluation Given the lengthy and detailed nature of the above text, one may wonder about its relevance to the clinical arena. Assisted stallion reproduction spans hundreds of years, with references to the topic dating back to Arabic texts (c. 1300) and scientific reports issued from the late 1700s to the early 1900s by noted authorities such as Spallanzani, Repiquet, Hoffman, and Ivanoff.816 Advancements were furthered by the works of investigators such ´ as Bielanski, Day, Mann, Merkt, and Nishikawa, and the next generation of dedicated scientists, including Amann, Foote, Hughes, Kenney, Palmer, Pickett, and Tischner. These individuals took the discipline of stallion reproduction to new heights. As seen by the information provided in the previous sections, we continue to garner a deeper understanding of spermatozoal structure and function. Although progress in equine reproduction is curbed by funding limitations, our discipline has gained considerable insights from work conducted in humans, laboratory animals, and food-producing animals. An appreciation of the molecular basis of spermatozoal function, spermatozoal–oviductal interactions, and spermatozoon–oocyte engagement will undoubtedly lead to many practical applications in the clinical front, such as assemblage of a battery of in-depth laboratory tests to assess spermatozoal function, expanded treatment options for subfertile stallions, improved methods for preservation of semen, and heightened applications for assisted reproductive technologies such as conventional IVF and ICSI techniques. The bottom line is that the more we learn, the more informed we can be in decision-making regarding diagnostic and therapeutic strategies as they relate to spermatozoal function and reproductive health in stallions. It becomes incumbent upon us, and future clinicians and academicians, to convert these opportunities into practical applications. Conventional laboratory tests for assessment of spermatozoal quality have included light microscopic evaluation of spermatozoal morphological characteristics and estimation of spermatozoal motility (including percentages of motile and progressively motile sperm; velocity of spermatozoal movement; and longevity of spermatozoal motility following in vitro storage).17,817–820 While other features of semen quality are also considered, including
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spermatozoal concentration, semen volume, and presence of blood, urine, or potentially pathogenic bacteria, the basic features of this examination process date back several decades. The value of the examination is predicated, to a large extent, on reliable equipment and personnel with good observational power. Even when these stipulations are in effect, the predictive value of the examination can be limited.114, 115, 125,821–833 The same holds true for spermatozoa of other mammalian species.834–837 After viewing the previous sections of this paper, one can appreciate that expanded analysis of equine spermatozoal populations might improve the predictive value of the testing process. In fact, additional tests shown to be of diagnostic value are presently incorporated into the spermatozoal examination process within some reference laboratories today. One of these tests is the sperm chromatin structure assay (SCSA). This assay, introduced by Evenson et al. in 1980,838 has been applied to spermatozoa from a number of species, including horses.839–841 The SCSA tests a compartment of spermatozoa that is not monitored by conventional methods, i.e., nuclear chromatin. The SCSA is a flow cytometric procedure that utilizes the metachromatic fluorochrome acridine orange, and tests the denaturability of spermatozoal chromatin challenged with acid treatment. The literature provides variable results regarding the relationship of stallion spermatozoal chromatin denaturation to the extent of disulfide bonding within and between protamine molecules;842, 843 however, chromatin susceptibility to denaturation is correlated with the level of actual DNA strand breaks.841 The DNA strand breaks can be associated with a myriad of factors, including idiopathic apoptosis, oxidative stress, heat stress, radiation injury, or protamine deficiency,843–850 and may involve double-stranded or single-stranded DNA fragmentation or oxidized nucleosides.844, 851 Such lesions could create genetically defective spermatozoa, leading to germline mutations. Interestingly, spermatozoa affected by such damage may appear to be normal, based on laboratory parameters such as spermatozoal motility and membrane integrity, but may induce post-fertilization embryonic failure.852 Owing to the highly condensed nature of the spermatozoal chromatin, mature spermatozoa are known to be transcriptionally inactive,35 so it is logical that DNA damage might not be expressed until mitosis occurs at the time of spermatozoon–oocyte fusion. This becomes quite important clinically as it represents a potential non-compensible defect, i.e., affected spermatozoa in an ejaculate may not be impaired for fertilization, so increasing the insemination number will not increase pregnancy rate. Ejaculated spermatozoa are known to retain a cohort of cytoplasmic mRNAs as well as translational
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ability,32, 35 so it is also possible that fragmentation of mRNA could have a negative impact on some other spermatozoal functions leading to reduced fertilization potential. Assays other than the SCSA are available to measure spermatozoal DNA fragmentation/chromatin disruption, including a TdTmediated-dUTP nick end labeling (TUNEL) assay, an in situ nick translation (NT) assay, a sperm chromatin dispersion (SCD) assay, and an electrophoresisbased comet assay.310, 844, 845,853–857 While these assays have not been used to the same extent as the SCSA in the equine arena, they are commonly applied in the human field. An immunofluorescence assay has also been developed for evaluation of human spermatozoal protamine levels,846 and a similar assay for equine spermatozoa could have diagnostic value. Another assay that we now commonly employ in our battery of tests is the acrosomal responsiveness assay (ARA). This assay is directed at testing the functionality of the spermatozoal acrosome, i.e., its ability to acrosome react when challenged with a potent inducer of the event, the Ca2+ ionophore A23187. Conventional tests are unable to predict the fertility of a subset of stallions with acrosomal dysfunction, because these stallions present with normal spermatozoal morphology and motility, as well as normal chromatin quality (based on SCSA testing) and normal acrosomal structure (based on fluorescent light microscopy and transmission electron microscopy). Meyers and co-workers858 first reported that the acrosomes of subfertile stallions with poor spermatozoal motility did not react readily in response to progesterone exposure. Fertile stallions averaged a 17% acrosomal reaction rate following 5 hours of incubation in capacitating conditions followed by exposure to progesterone, while subfertile stallions averaged only a 6% response rate. This study indicated that progesterone was capable of stimulating the acrosome reaction in equine spermatozoa exposed to capacitating conditions, and the response in subfertile stallions was reduced.858 Others have reported that the plasma membrane of stallion spermatozoa contains progesterone receptors, and indicate that this may be a pathway for induction of the acrosome reaction.859 In this regard, the percentage of spermatozoa with exposed progesterone receptors has been shown to be highly correlated with fertility of stallions.860 Other workers identified a subset of subfertile stallions whose only distinguishing spermatozoal characteristic was the drastically reduced ability to acrosome react when exposed to A23187 for up to 3 hours (i.e., mean reaction rate of 84% in fertile stallions versus mean reaction rate of 6% in subfertile stallions),861 thereby suggesting the spermatozoal defect may be isolated to the acrosome. Measurement of the cholesterol–phospholipid ratio in semen of these stal-
lions revealed that the ratio was significantly greater in both seminal plasma and whole sperm from the subfertile stallions when compared with fertile stallions.862 Further studies are required to dissect the underlying etiology of this condition and to determine appropriate treatment strategies. A keen understanding of the molecular pathways of capacitation and the acrosome reaction, as discussed above, provides us with logical investigative approaches. While transmission electron microscopy remains a decisive means for detecting the acrosome reaction in stallion spermatozoa,863 acrosomal status can be assessed by bright-field microscopy using Coomassie blue stain.864, 865 Fluorescence-tagged lectins or acidotrophic probes can be used with light microscopy or flow cytometric methods to assess the acrosome reaction or acrosomal damage.858,866–873 Although the more popular fluoresceinated lectin markers have generally replaced chlortetracycline-based assays for detecting the acrosome reaction, the latter assay remains useful because of its ability to gage capacitation, as well as the acrosome reaction.863, 874 Another marker demonstrated to be useful for detection of capacitation in stallion spermatozoa is merocyanine 540, an impermeant lipophilic probe which permits evaluation of the architecture and disorder of lipids in the outer leaflet of the plasma membrane bilayer.455, 481, 513, 560 Other probes, such as fluorochrome-conjugated annexin V or Ro-09-0198, are being used increasingly to monitor membrane asymmetry. These two probes bind with phosphatidylserine and phosphatidylethanolamine, respectively. Surface exposure of these phospholipids is considered to represent spermatozoal capacitation because they generally reside in the cytoplasmic leaflet of the lipid bilayer in uncapacitated spermatozoa.472, 875, 876 We know from the information provided in the previous sections that a spermatozoon is laden with special molecular domains that are designed, and modulated, for a multitude of functions within the female reproductive tract. In some respects, however, the spermatozoon might be considered a relatively simple structure, as it is only composed of a head and tail, with a dramatic reduction in cellular organelles in comparison with somatic cells or its homolog, the ovum. As such, a spermatozoon might be subdivided into: (i) a nucleus, (ii) an overlying acrosome, (iii) the fibrous and microtubular components of the flagellum, (iv) a mitochondrial sheath, and (v) an enveloping plasma membrane. With this in mind, one can appreciate that a battery of tests devised to evaluate these various compartments might aid in improved localization of cellular dysfunction and might also improve the predictive value of a laboratory-based semen evaluation.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell Assays developed for the nucleus and the acrosome were discussed above. In addition, techniques specific for mitochondrial function, flagellar substructure, and plasma membrane integrity are available. While the importance of mitochondrial-based oxidative metabolism for ATP production in spermatozoa has been challenged,237, 877 a key role of the mitochondria in sustenance of spermatozoal functions remains quite likely.192 Mitochondria-selective fluorescent markers abound, and include rhodamine-123; an R probes (which differ in assortment of MitoTracker spectral characteristics, adaptability to permeabilization and fixation, and reliance on respiration state of the mitochondria); and 5,5 ,6,6 -tetrachloro-1,1 ,3,3 tetraethylbenzimidazolylcarbocyanine (JC-1). These reporter molecules passively diffuse across the plasma membrane and accumulate in the mitochondria. Rhodamine-123 was one of the original mitochondrial probes, but has largely been replaced by other markers because of its reduced photostability and its tendency to lose retention in the mitochondria following a loss in membrane potential. Some R probes do not fluoresce until of the MitoTracker oxidized, so these markers permit easy assessment of the respiration state of the mitochondria. The JC1 may be optimally suited for evaluation of the energetic state of mitochondria because it produces a membrane potential-sensitive shift in color pattern, i.e., the color changes from green to red–orange as membrane polarization increases. This is due to the reversible formation of red–orange J-aggregates in a state of high membrane potential, as compared with green monomeric JC-1 aggregates in a state of low to medium membrane potential.873,878–881 The semipermeable plasma membrane, which is composed of a wide assortment of lipids, proteins, and carbohydrates, envelops the entire spermatozoon. This structure is vital to regulation of spermatozoal functions by establishing ion gradients, facilitating cytosolic entry of larger molecules, and orchestrating various cell-signaling events, to name a few. Thus, assessment of its integrity would seem an important component of a semen evaluation. To this end, a variety of laboratory procedures have been used to evaluate the integrity of the plasma membrane. One method is to evaluate the ability of spermatozoa to exclude extracellular dyes, such as eosin Y, which are non-permeable when the membrane is intact. Another approach is to expose spermatozoa to hypotonic media (50–100 mOsm range) to test their osmoregulatory function [termed the hypo-osmotic swelling test (HOST)].882, 883 With this assay, membrane-intact spermatozoa theoretically permit excessive water entry into the cytosol, resulting in a variety of morphological changes in the flagellum associated with the cytosolic swelling.
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Conversely, osmoregulation-incompetent spermatozoa will not experience noticeable changes in flagellar shape. While these tests can serve an adjunctive role in semen evaluation, they are not widely applied in the clinical setting because of the potential for misinterpretation. For instance, humidity affects the cellular permeability of eosin Y.884 The incidence of spermatozoa with a bent flagellum prior to HOST will also confound the interpretation of results following exposure of a spermatozoal population to hypo-osmotic media. A broad array of membrane-impermeable fluorescent dyes is available commercially and can be used to test membrane intactness. Examples include the DNA dyes propidium iodide, bis-benzimide R R -1, TOTO -1, and ethid(Hoechst 33258), YO-PRO ium homodimer-1. As an alternative, spermatozoa can be bathed in cell-permeable probes that become hydrolyzed to form membrane-impermeant fluorescent products in the cytosol. Examples include carboxyfluorescein diacetate (hydrolyzed by nonspecific cytosolic esterases to form carboxyfluorescein); calcein AM or dihydrocalcein AM (hydrolyzed by esterases to form calcein, which, in turn, forms fluorescent complexes with Ca2+ , and other metals); R -14 (deacylated in the cytosol, with the reand SYBR sulting product expressing strong fluorescence when complexed with nucleic acids). Of interest, staining R of porcine spermatozoa with SYBR -14, which complexes with DNA, does not affect their ability to fertilize oocytes.885 Certain membrane-impermeable and membrane-permeable dyes can be combined in solution prior to spermatozoal exposure in an effort to provide a more accurate reflection of membrane R -14 integrity. For instance, a combination of SYBR and propidium iodide yields three populations of R stained spermatozoa: (i) membrane-intact, SYBR 14-stained cells (green); (ii) membrane-damaged, propidium-iodide-stained cells (red), and (iii) moribund cells (double stained).886, 887 In one study, the percentage of motile stallion spermatozoa (based on computerized motility analysis) was highly correlated (r = 0.98) with the percentage of spermatozoa with intact plasma membranes, based on staining R with SYBR -14 and propidium iodide.888 Some fluorescent plasma membrane dyes can be also be combined with certain mitochondrial dyes888, 889 or acrosomal dyes890–894 to provide more thorough compartmental coverage in the assay. The literature reports triple-stain fluorescent techniques for use with R stallion spermatozoa: propidium iodide/SYBR 888 14/JC1 and propidium iodide/FITC-PNA (an acrosomal lectin probe)/carboxy-SNARF-1 (an intracellular pH indicator).890, 895 Although images can be ascertained with the various fluorophores described above by using fluorescence microscopy,
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Table 97.1 Biochemical markers for assessing spermatozoa function Biochemical marker
Potential value of test
Acrosin/amidase activity898–900 SNARE proteins694, 901 Caspases902–905 Heparin-binding proteins499, 906 Protein phosphotyrosine activity907–909 Superoxide anion266 Chromomycin A3 910–913 Acetylcarnitine/carnitine Phospholipase Cζ 914, 915 C11 -BODIPY(581/591)340,916–920 Malondialdehyde921–928 Glutathione peroxidase320,929–932 Reactive oxygen species257, 282, 835, 928, 931,933–938 Lactate dehydrogenase939–942 Creatine kinase/heat shock protein A2943–946 CRISPs104, 499, 503, 790,947–949 SP20/hyaluronidase456, 761, 950 Spermadhesion proteins499,951–953 P34H protein954–957 Zonadhesin958–964 SP22 protein965–968
Acrosome integrity Acrosome reaction Apoptosis Capacitation Capacitation Capacitation Chromatin packing Motility/morphology Oocyte activation Oxidative injury Oxidative injury Oxidative injury Oxidative injury Spermatozoal motility Spermatozoal maturity Sperm–oocyte binding Sperm–oocyte interaction Sperm–zona binding Sperm–zona binding Sperm–zona binding Spermatozoal fertility
BODIPY, boron-dipyrromethene; CRISPs, cysteine-rich secretory proteins; SNARE, soluble N-ethylmaleimide-sensitive factor attachment protein receptor; SP, sperm protein.
flow cytometry is typically applied because of the high throughput and objectivity associated with this approach.870, 871, 881, 894, 896 Utilization of a fluorescence microplate reader assay is also reported for use with JC-1.897 Considerable effort has been directed toward identification of biochemical markers of spermatozoal function that might aid in laboratory-based detection of fertility by targeting specific subcellular compartments or domains. Many of these methods have been devised for use with non-equine subjects, but several have been proposed for potential use with stallion spermatozoa. Although the value of such tests requires further scrutiny and standardization, Table 97.1 provides examples of assays that may prove valuable as diagnostic tools. Incorporation of in-depth tests, such as those above, for semen evaluation does not replace, or diminish the value of, the classical measurements of spermatozoal motility or morphology, and these two methods are likely to remain the hallmarks for semen evaluation. Evaluation of spermatozoal motility in both raw and extended forms is considered to be a fundamental laboratory test for assessing the fertilizing capacity of spermatozoa in an ejaculate. Evaluation of raw (undiluted or neat) semen gives one an idea of how well spermatozoa perform in their natural fluid milieu. Determining motility in the raw
form can be hampered by higher sperm concentrations and spermatozoal agglutination to the coverslip, making it difficult for the evaluator to discern individual motility patterns. To overcome this limitation, an aliquot of semen can also be appropriately diluted (e.g., to 25 × 106 sperm/mL) in a good quality semen extender that is free of debris when visualized microscopically. The extender may slightly alter the motility pattern, usually by increasing the velocity measures. After initial extension, a high percentage of spermatozoa may exhibit a circular motility pattern; however, this behavior usually resolves following 5–10 minutes of exposure time in the extender. Spermatozoal motility is exceptionally susceptible to environmental conditions (such as excessive heat or cold, lubricants, light, disinfectants, and osmolarity/pH of semen extender), so it is necessary to protect the semen from injurious agents or conditions prior to analysis. To enhance the reliability of motility estimation, the procedure should be performed by an experienced person using a properly equipped microscope, i.e., one with a built-in stage warmer and properly adjusted phase-contrast optics. Estimations of the percentages of motile and progressively motile spermatozoa are generally determined, in addition to an estimation of spermatozoal velocity [based on an arbitrary scale of 0 (stationary) to 4 (fast)]. Subjective assessment of motility is generally quite
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Table 97.2 Effect of equine spermatozoa centrifugation through a silanized silica-particle solution on spermatozoal motion characteristics Stallion
Semen treatment
Fertile Fertile Subfertile Subfertile
Before After Before After
MOT (%)
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MOT, percentage motility; PMOT, percentage progressive motility; VCL, mean curvilinear velocity.
acceptable, provided personnel are experienced in analysis of spermatozoal motility. Several different techniques and instruments have been developed in an effort to secure an objective (i.e., unbiased) evaluation of spermatozoal motility; however, these methods (e.g., time-lapse photomicrography, frame-by-frame playback videomicrography, spectophotometry, or computerized analysis) are generally considered to be too tedious or expensive for routine use. Computerized systems are currently in place in many reference laboratories with the intent to objectively assess motion characteristics of spermatozoa. Despite the commercial availability of various generations of computer-assisted spermatozoal analysis (CASA) systems for over 20 years, their presence has not provided the definitive assay for measuring spermatozoal fertilizing potential.969 Such an expectation, however, is unrealistic given the numerous independent spermatozoal attributes that are required for a spermatozoon to possess fertilization competence. What these systems do provide is the prospect of objective measurement and protocol standardization. These instruments permit customized selection of various features, including frequency and length of frame capture; threshold demarcations for presence of spermatozoal motion, progressivity of motion, and velocity measures; and gating freedom for both size and luminosity representative of spermatozoal heads, as a means to maximize capture of spermatozoa while minimizing capture of non-spermatozoal material in the sample of interest. Such manipulations are important for improving measurement accuracy and repeatability within a given laboratory, but make it virtually impossible for reliable comparisons among laboratories or among different CASA brands.970, 971 Computerized analysis of spermatozoa is primarily reserved for the research setting, where standardization, accuracy, and precision are a prerequisite to measurement of experimental end points. A distinct value of CASA instruments in the commercial environment (at a veterinary hospital or an equine breeding operation) is the ability to garner objective results for a variety of motility variables. Confusion arises, however, regarding the
relationship of the myriad of obtainable CASA variables to fertility of the sample. As an example, we recently conducted a fertility trial with a subfertile stallion whose semen was subjected to density-gradient centrifugation in an effort to improve semen quality prior to insemination. Values for percentage motility (MOT), percentage progressive motility (PMOT), and mean curvilinear velocity (VCL; µm/s) before and after semen processing for the subfertile stallion and a fertile control stallion are shown in Table 97.2. Based on these results, it would appear that semen treatment for the subfertile stallion yielded a spermatozoal population with quality similar to, or exceeding (based on velocity values), that of the fertile control stallion. Nonetheless, when fertile mares were inseminated hysteroscopically with 20 × 106 progressively motile spermatozoa (100 µL volume), the resulting pregnancy rates were 15/20 (75%) for the fertile stallion, compared with 7/20 (35%) for the subfertile stallion (P = 0.01). This demonstrates that spermatozoal motility does not provide absolute discrimination power, again emphasizing that spermatozoal attributes other than motility play critical roles in spermatozoal fertilizing ability. Some CASA systems can be outfitted with fluorescence optics, and the predictive value of CASA might be improved by incorporation of fluorescent dyes in the media such that motility variables can be segregated by the presence or absence of membrane intactness.972 Further studies are required to determine whether the predictive value of spermatozoal motility can be improved by this approach. A study involving boars suggests that CASA analysis might have an improved relationship with fertility if the spermatozoa are also incubated under capacitating conditions to observe changes in spermatozoal velocity over time.973 The morphology or structure of spermatozoa is typically examined with a light microscope at 1000× magnification. Standard bright-field microscope optics can be used to examine air-dried semen smears, provided appropriate stains are used in slide preparation. Specific sperm stains include those developed by Williams974 and Casarett.975 General purpose cellular stains (e.g., Wright’s, Giemsa,
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hematoxylin–eosin) have also been used to accent both germinal and somatic cells in semen smears. Background stains (e.g., eosin–nigrosin, India ink) probably are the most widely used stains because of their ease of application. Visualization of the structural detail of spermatozoa can be greatly enhanced by fixing the cells in buffered formol saline or a similar fixative, then viewing the unstained cells as a wet mount with either phase-contrast or, preferably, differential interference contrast (DIC) microscopy.884 In addition, the incidence of artifactual changes is reduced in comparison with stained smears. At least 100 spermatozoa should be evaluated for evidence of morphological defects. The type and incidence of each defect should be recorded. Abnormalities in spermatozoal morphology traditionally have been classified as primary, secondary, or tertiary. Primary abnormalities are considered to be associated with a defect in spermatogenesis and, therefore, are of testicular origin. Secondary abnormalities are created in the excurrent duct system. Tertiary abnormalities, as opposed to the previous two types, develop in vitro as a result of improper semen collection or handling procedures. The current trend is to record the numbers of specific morphological defects, such as knobbed acrosomes, proximal protoplasmic droplets, swollen midpieces, and coiled tails. This method of classification is considered superior to the traditional system because it reveals more specific information regarding a population of sperm while avoiding erroneous assumptions about the origin of these defects. The origin of some spermatozoal morphological defects is unknown. Additionally, some morphological abnormalities such as detached heads can be primary, secondary, or tertiary in nature, thereby introducing the possibility of error when using the traditional classification system exclusively. Readers are directed to Figure 97.39 for further information on normal and abnormal sperm morphological features, as might be viewed when using DIC microscopy for analysis. The percentage of morphologically normal spermatozoa is positively correlated with spermatozoal motility. Moreover, close morphological inspection provides additional information about characteristics of individual spermatozoa. This information is important because semen can possess good spermatozoal motility, yet have a relatively high incidence of spermatozoal morphological abnormalities. Furthermore, stallions can have many spermatozoal abnormalities, yet display normal fertility. Some morphological defects (e.g., cytoplasmic droplets, bent tails) appear to have a minor effect on fertility of stallions bred by natural cover, whereas other defects (e.g., detached heads, abnormally shaped heads, abnormally shaped midpieces, coiled tails, and premature germ
cells) have a deleterious effect on fertility.125 In general, morphologically abnormal spermatozoa do not exert a direct negative influence on normal spermatozoa. Therefore, the total number of morphologically normal sperm in ejaculates may provide more information regarding the fertility of a stallion than the percentage or absolute number of morphologically abnormal spermatozoa. Generally speaking, the percentage of morphologically normal sperm in a semen sample is similar to the percentage of progressively motile spermatozoa. If spermatozoal motility is low and the percentage of morphologically normal sperm is high, it suggests that laboratory errors occurred which led to a lowering of spermatozoal motility. One cannot discount, however, a potentially negative effect of seminal plasma on spermatozoal motion characteristics. As seen in the previous paragraphs, a variety of techniques and protocols are available for evaluation of the spermatozoon. Application of newly developed assays can be cumbersome and expensive. Nonetheless, incorporation of some of these diagnostic tools into a standard semen analysis may yield improved discrimination ability regarding the competence of these complex, yet intriguing, cells. Similar applications would be valuable for critical evaluation of laboratory techniques applied to liquid preservation or cryopreservation of spermatozoa.
Concluding remarks Although a plethora of scientific information surrounds spermatozoal structure and function, many unresolved issues remain, even with human and rodent spermatozoa in which the largest body of information has been assimilated. Only when we know precisely the specific molecular interactions required for attaining full spermatozoal fertilizing potential, including the spatial and temporal changes, energetics, and gaseous environment involved, will we be able to reliably manipulate spermatozoa to meet the growing needs within the equine breeding industry. Goals might include devising methods for longterm cooled semen preservation, improving the fertility of cryopreserved semen, and incorporation of IVF (both conventional IVF and ICSI) into commercial programs. While these might appear to be lofty goals, a more absolute understanding of spermatozoal structure and function would certainly take a lot of the ‘guess work’ out of current approaches to analysis and manipulation of equine spermatozoa. Attempts to gain a better understanding of equine spermatozoa solely from extrapolation of data acquired from other mammalian species are likely destined to failure because of the well-known species
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Figure 97.39 Drawings of spermatozoa to resemble images of normal and abnormal equine spermatozoal morphology, as viewed by differential interference contrast (DIC) microscopy of a fixed and unstained wet-mount semen specimen. The DIC optics provide a three dimensional image of spermatozoa. (A) Normal spermatozoal morphology, in dorsoventral (a1, a2) and side (a3) views. Abaxial midpieces (a1) are considered to be morphologically normal. (b) Variations in abnormal head morphology, including macrocephalic (large head) morphology (b1), microcephalic (small head) morphology (b2), nuclear vacuoles (or crater defects; considered by our laboratory to be normal if low in number and size, and located randomly over acrosomal region; (b3), tapered head (b4), pyriform head (b5), constricted or hour-glass head (b6), and degenerate head (b7). (c) Acrosomal defect in dorsoventral (c1, c2) and side (c3) views. This morphological defect is termed ‘knobbed acrosome’. (d) Proximal (d1) and distal (d2, d3) cytoplasmic droplets. (e) Abnormalities of the midpiece, including segmental aplasia of the mitochondrial sheath (e1), roughed midpiece from uneven distribution of mitochondria (e2), enlarged mitochondrial sheath (e3), bent midpiece (e4, e5, e6), and double midpiece/double head (e7). (f) Bent tail (or hairpin tail), involving the mid-region of the principal piece (f1–f4) or with a singular bend (f5) or proximal bend (f6) involving the midpiece–principal piece junction. (g) Coiled tail, with the tail tightly encircling the head (g1) or the tail coil not encircling the head (g2, g3). (h) Fragmented sperm, including head detachment (termed detached heads or tailless heads; (h1) or fragmentation at the level of the annulus (h2). Fragmentation can also occur at other sites, as demonstrated in f4. (i) Premature germ cells with a single nucleus (i1) or multiple nuclei (i2).
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differences in spermatozoal attributes and physiology. Nonetheless, much information derived from other species may have relevance to equids, and should be investigated for applicability. Examples include identification of candidate genes for specific spermatozoal traits, targeted mutation of genes for specific proteins to study the resulting effect on reproductive function, and use of gene-silencing agents for regulatable ablation of gene function. Incorporation of molecular techniques would appear to be the key to elucidation of mechanisms that control spermatozoal development and function in stallions. Indeed, the ‘omics’ era (e.g., genomics, toxicogenomics, transcriptomics, metabonomics, and proteomics) is upon us. Technological progress in the field of molecular genetics has been remarkable, and has opened the door to new paradigms for the study of spermatozoal function and dysfunction. Significant inroads have been made in horse genome mapping in recent years, including availability of synteny, linkage, radiation-hybrid, cytogenetic, and human–horse comparative maps.976–981 The National Institutes of Health recently reported (www.nih.gov/news/pr/feb2007/nhgri-07.htm) that the 2.7 billion DNA base-pair horse genome has been sequenced and partially assembled, and the first draft of the sequence is now accessible through public databases; hence, the importance of bioinformatics to biological research. Continued progress in genomic research will lead to isolation and identification of genes responsible for spermatozoal development and function, as well as genetic factors that underlie various types of reproductive failure. High-throughput microarray technology (cDNA and oligonucleotide) is being used with increased intensity to study differential gene expression.982 Reverse transcription and amplification steps are being applied to quantify gene activity in targeted cells or processes, and to unravel the complexity of genes.983, 984 The large-scale information obtained from these emergent technologies is being applied to male fertility,985 including that of horses.53, 986, 987 Interpretation of such expression profiling and its biological significance, however, can be a quagmire and continued effort is required to overcome this obstacle.988 Given the intensity of investigation in this area, the basic mechanisms that control gene expression will emerge in the years ahead, and genetic markers for stallion fertility may one day abound.989 Robust mass spectrometry proteomics methodologies allow inventory of proteins in cells, organelles, and body fluids,990–992 and offer the potential for identification of biomarkers for diagnostic tests.993–995 As with humans and other animal species, the impacts that environmental chemicals have on stal-
lion fertility deserve attention. An apparent decline in spermatozoal quality and quantity in men has been described in recent years,996–998 Increased rates of other reproductive tract problems, such as cryptorchidism and testicular cancer, have also been reported.999, 1000 Environmental reproductive toxicants are considered to be a major contributor to these patterns.1001–1003 Both complimentary DNA (cDNA) microarray and real-time reverse-transcriptase polymerase chain reaction (qRT-PCR)approaches have revealed differential expression profiles in some spermatogenesis-related genes (i.e., heat shock protein 70-2, insulin growth factor binding protein 3, and glutathione S transferase pi) following toxicant exposure, so genes such as these may serve as useful biomarkers when screening for testicular toxicity.1004 A better understanding of the molecular mechanisms regulating spermatozoal development and function will likely lead to new diagnostic techniques and therapeutic strategies for reduced fertility in stallions and may possibly translate to methodologies designed to curb gonadal aging. Such progress can be curtailed only by funding deficits or diverted interests of investigators in the years ahead. Undoubtedly, future generations will look beyond descriptive morphology and motility in characterizing structural and functional features and deficiencies of spermatozoa. In addition, they will likely use the spermatozoon as a vehicle for production of transgenic horses,1005, 1006 as well as spermatogonial stem cell transplantation or testicular grafting techniques to study testicular function or to propagate a genetic line.1007–1014 It seems befitting to conclude this manuscript with a quote from Professor Storey, in his article entitled ‘Interactions between gametes leading to fertilization: the sperm’s eye view’:1 ‘the reactions that lead to fertilization of the mammalian egg by its conspecific sperm are many, complex, and exquisitely coordinated. One might add that they are also vital.’
Acknowledgements Financial assistance was provided by the Patsy Link Endowment Fund, Texas A&M University; the Shining Spark-Carol Rose Stallion Reproduction Fund (through the American Quarter Horse Association); the Azoom, Corona Cartel, and Teller Cartel syndicates (through Lazy E Ranch); Black Rock Ranch; and Coolmore Stud. Special thanks go to Mr. Ricardo Levario, Ms. Nicole Hilborn, Ms. Victoria Star Varner, Mr. Vince Hardy, and Dr. Noah Heninger for assistance with manuscript figures, and to Ms. Sarah Chellappan for reference management.
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References 1. Storey BT. Interactions between gametes leading to fertilization: the sperm’s eye view. Reprod Fertil Dev 1995; 7:927–42. 2. MacLean II JA, Wilkinson MF. Gene regulation in spermatogenesis. Curr Top Dev Biol 2005;71:131–97. 3. Tanaka H, Baba T. Gene expression in spermiogenesis. Cell Mol Life Sci 2005;52:344–54. 4. Rossi P, Dolci S, Sette C, Capolunghi F, Pellegrini M, Loiarro M, Agostino SD, Paronetto MP, Grimaldi P, Merico D, Martegani E, Geremia R. Analysis of the gene expression profile of mouse male meiotic germ cells. Gene Expr Patterns 2004;4:267–81. 5. Elliott D. Pathways of post-transcriptional gene regulation in mammalian germ cell development. Cytogenet Genome Res 2003;103:210–16. 6. Ronfani L, Bianchi ME. Molecular mechanisms in male determination and germ cell differentiation. Cell Mol Life Sci 2004;61:1907–25. 7. Grimes SR. Testis-specific transcriptional control. Gene 2004;343:11–22. 8. van der Weyden L, Arends MJ, Chausiaux OE, Ellis PJ, Lange UC, Surani MA, Affara N, Murakami Y, Adams DJ, Bradley A. Loss of TSLC1 causes male infertility due to a defect at the spermatid stage of spermatogenesis. Mol Cell Biol 2006;26:3595–609. 9. Gromoll J, Simoni M. Genetic complexity of FSH receptor function. Trends Endocrinol Metab 2005;16:368–73. 10. Akingbemi BT. Estrogen regulation of testicular function. Reprod Biol Endocrinol 2005;3:51. 11. De Gendt K, Swinnen JV, Saunders PTK, Schoonjans L, Dewerchin M, Devos A, Tan K, Atanassova N, Claessens F, Lecureuil C, Heyns W, Carmeliet P, Guillou F, Sharpe RM, Verhoeven G. A sertoli cell-selective knockout of the androgen receptor causes spermatogenic arrest in meiosis. Proc Natl Acad Sci USA 2004;101:1327–32. 12. Verhoeven G. A sertoli cell-specific knock-out of the androgen receptor. Andrologia 2005;37:207–8. 13. Tsai M, Yeh SD, Wang RS, Yeh S, Zhang C, Lin HY, Tzeng CR, Chang C. Differential effects of spermatogenesis and fertility in mice lacking androgen receptor in individual testis cell. Proc Natl Acad Sci USA 2006;103:18975– 80. 14. Varner DD, Blanchard TL, Brinsko SP, Love CC, Taylor TS, Johnson L. Techniques for evaluating selected reproductive disorders of stallions. Anim Reprod Sci 2000;60–61:493–509. 15. Amann RP, Johnson L, Pickett BW. Connection between the seminiferous tubules and the efferent ducts in the stallion. Am J Vet Res 1977;38:1571–9. 16. Johnson L, Thompson Jr DL. Age-related and seasonal variation in the Sertoli cell population daily sperm production and serum concentrations of follicle-stimulating hormone, luteinizing hormone and testosterone in stallions. Biol Reprod 1983;29:777–89. 17. Varner DD, Schumacher J, Blanchard TL, Johnson L. Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991. 18. Johnson L, Neaves WB. Age-related changes in the Leydig cell population, seminiferous tubules, and sperm production in the stallion. Biol Reprod 1981;24:703–12. 19. Johnson L, Amann RP, Pickett BW. Scanning electron and light microscopy of the equine seminiferous tubule. Fertil Steril 1978;29:208–15.
963
20. Swierstra EE, Gebauer MR, Pickett BW. Reproductive physiology of the stallion. I. Spermatogenesis and testis composition. J Reprod Fertil 1974;40:113–23. 21. Maekawa M, Kamimura K, Nagano T. Peritubular myoid cells in the testis: their structure and function. Arch Histol Cytol 1996;59:1–13. 22. Skinner MK, Fritz IB. Identification of a non-mitogenic paracrine factor involved in mesenchymal-epithelial cell interactions between testicular peritubular cells and Sertoli cells. Mol Cell Endocrinol 1986;44:85–97. 23. Hettle JA, Balekjian E, Tung PS, Fritz IB. Rat testicular peritubular cells in culture secrete an inhibitor of plasminogen activator activity. Biol Reprod 1988;38:359– 71. 24. Wong C, Cheng CY. The blood-testis barrier: its biology, regulation, and physiological role in spermatogenesis. Curr Topics Dev Biol 2005;71:263–96. 25. Li MWM, Xia W, Mruk DD, Wang CQF, Yan HHN, Siu MKY, Lui W, Lee WM, Cheng CY. Tumor necrosis factor (alpha) reversibly disrupts the blood-testis barrier and impairs Sertoli-germ cell adhesion in the seminiferous epithelium of adult rat testes. J Endocrinol 2006;190: 313–29. 26. Johnson L. Spermatogenesis. In: Cupps PT (ed.) Reproduction in Domestic Animals. San Diego: Academic Press, 1991; pp. 173–219. 27. Huleihel M, Lunenfeld E. Regulation of spermatogenesis by paracrine/autocrine testicular factors. Asian J Androl 2004;6:259–68. 28. O’Donnell L, Meachem SJ, Stanton PG, McLachlan RI. Endocrine regulation of spermatogenesis. In: Neill JD (ed.) Knobil and Neill’s Physiology of Reproduction. Boston, MA: Elsevier, 2006; pp. 1017–69. 29. Holdcraft RW, Braun RE. Hormonal regulation of spermatogenesis. Int J Androl 2004;27:335–42. 30. Perrard M-H, Vigier M, Damestoy A, Chapat C, Silandre D, Rudkin BB, Durand P. Beta-nerve growth factor participates in an autocrine/paracrine pathway of regulation of the meiotic differentiation of rat spermatocytes. J Cell Physiol 2007;210:51–62. 31. Abo-Elmaksoud A, Sinowatz F. Expression and localization of growth factors and their receptors in the mammalian testis. Part I. Fibroblast growth factors and insulin-like growth factors. Anat Histol Embryol 2005;34: 319–34. 32. Miller D, Ostermeier GC. Towards a better understanding of RNA carriage by ejaculate spermatozoa. Hum Reprod Update 2006;12:757–67. 33. Moreno RD, Alvarado CP. The mammalian acrosome as a secretory lysosome: new and old evidence. Mol Reprod Dev 2006;73:1430–4. 34. Dadoune JP, Siffrol JP, Alfonsi MF. Transcription in haploid male germ cells. Int Rev Cytol 2004;237:1–56. 35. Miller D, Ostermeier GC. Spermatozoal RNA: why is it there and what does it do? Gynecol Obstet Fertil 2006;34: 840–6. 36. Johnson L. A new approach to quantification of Sertoli cells that avoids problems associated with the irregular nuclear surface. Anat Rec 1986;214:231–7. 37. Holstein AF, Schulze W, Davidoff M. Understanding spermatogenesis is a prerequisite for treatment. Reprod Biol Endocrinol 2003;1:107. 38. Lebland CP, Clermont Y. Definition of the stages of the cycle of the seminiferous epithelium in the rat. Ann N Y Acad Sci 1952;55:548–73.
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39. Johnson L. Efficiency of spermatogenesis. Microsc Res Tech 1995;32:385–422. 40. Clermont Y. The cycle of the seminiferous epithelium in man. Am J Anat 1963;112:35–51. 41. Johnson L. Increased daily sperm production in the breeding season of stallions is explained by an elevated population of spermatogonia. Biol Reprod 1985;32:1190. 42. Johnson L, Hardy VB, Martin MT. Staging equine seminiferous tubules by Normarski optics in unstained histologic sections and in tubules mounted in toto to reveal the spermatogenic wave. Anat Rec 1990;227:167– 74. 43. Heninger NL, Staub C, Blanchard TL, Johnson L, Varner DD, Forrest DW. Germ cell apoptosis in the testes of normal stallions. Theriogenology 2004;62:297. 44. Johnson L, Blanchard TL, Varner DD, Scrutchfield WL. Factors affecting spermatogenesis in the stallion. Theriogenology 1997;48:1199–216. 45. Blanchard TL, Johnson L. Increased germ cell degeneration and reduced germ cell:Sertoli cell ratio in stallion with low sperm production. Theriogenology 1997;47: 665–77. 46. Clermont Y. Quantitative analysis of spermatogenesis of the rat: a revised model for the renewal of spermatogonia. Am J Anat 1962;111:111–29. 47. Huckins C. The morphology and kinetics of spermatogonial degeneration in normal adult rats: an analysis using a simplified classification of the germinal epithelium. Anat Rec 1978;190:905–26. 48. Kerr JF, Wyllie AH, Currie AR. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer 1972;26:239–57. 49. Cisternas P, Moreno RD. Comparative analysis of apoptotic pathways in rat, mouse, and hamster spermatozoa. Mol Reprod Dev 2006;73:1318–25. 50. Allan DJ, Harmon BV, Roberts SA. Spermatogonial apoptosis has three morphologically recognizable phases and shows no circadian rhythm during normal spermatogenesis in the rat. Cell Prolif 1992;25:241–50. 51. Joshi AR, Dighe RR. Expression of apoptotic genes in testicular germ cells and their modulation by luteinizing hormone. J Steroid Biochem Mol Biol 2006;101:22–30. 52. Heninger NL, Staub C, Johnson L, Blanchard TL, Varner D, Ing NH, Forrest DW. Testicular germ cell apoptosis and formation of the Sertoli cell barrier during the initiation of spermatogenesis in pubertal stallions. Anim Reprod Sci 2006;94:127–31. 53. Ing NH, Laughlin AM, Varner DD, Welsh Jr TW, Forrest DW, Blanchard TL, Johnson L. Gene expression in the spermatogenically inactive ‘dark’ and maturing ‘light’ testicular tissues of the prepubertal colt. J Androl 2004; 25:535–44. 54. Meachem SJ, Ruwanpura SM, Siolkowski J, Ague JM, Skiner MK, Loveland KL. Developmentally distinct in vivo effects of FSH on proliferation and apoptosis during maturation. J Endrocrinol 2005;186:429–46. 55. Sluka P, O’Donnell L, Bartles JR, Stanton PG. FSH regulates the formation of adherens junctions and ectoplasmic specialisations between rat Sertoli cells in vitro and in vivo. J Endocrinol 2006;189:381–95. 56. Nickel R, Schummer A, Seiferle E, Sack WO. The Viscera of Domestic Animals. New York: Springer-Verlag, 1973; pp. 304–50. 57. Johnson L, Amann RP, Pickett BW. Scanning electron microscopy of the epithelium and spermatozoa in the
58.
59.
60.
61. 62.
63.
64.
65.
66.
67. 68. 69.
70.
71.
72.
73.
74.
75.
equine excurrent duct system. Am J Vet Res 1978;39: 1428–34. Robaire B, Hinton BT, Orgebin-Crist M-C. The epididymis. In: Neill JD (ed.) Knobil and Neill’s Physiology of Reproduction. New York: Elsevier, 2006; pp. 1071–148. Amann RP, Thompson Jr DL, Squires EL, Pickett BW. Effects of age and frequency of ejaculation on sperm production and extragonadal sperm reserves in stallions. J Reprod Fertil Suppl 1979;27:1–6. Swierstra EE, Pickett BW, Gebauer MR. Spermatogenesis and duration of transit of spermatozoa through the excurrent ducts of stallions. J Reprod Fertil Suppl 1975; 23:53–7. Amann RP. A review of anatomy and physiology of the stallion. J Equine Vet Sci 1981;1:83–105. Gebauer MR, Pickett BW, Swierstra EE. Reproductive physiology of the stallion. III. Extra-gonadal transit time and sperm reserves. J Anim Sci 1974;39:737–42. Amann RP. A critical review of methods for evaluation of spermatogenesis from seminal characteristics. J Androl 1981;2:37–58. Varner DD, Schumacher J, Blanchard TL, Johnson L. Diseases of the epididymis. In: Varner DD, Schumacher J, Blanchard TL, Johnson L (eds) Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991; pp. 233–40. Love CC. Ultrasonographic evaluation of the testis, epididymis, and spermatic cord of the stallion. In: Blanchard TL, Varner DD (eds) Veterinary Clinics of North America: Equine Practice (Stallion Management). Philadelphia: W.B. Saunders, 1992; pp. 167–82. Alvarez JG, Storey BT. Spontaneous lipid peroxidation in rabbit epididymal spermatozoa: its effects on sperm motility. Biol Reprod 1982;27:1102–8. Vernet P, Aitken RJ, Drevet JR. Antioxidant strategies in the epididymis. Mol Cell Endocrinol 2004;216:31–9. Johnson L, Amann RP, Pickett BW. Maturation of equine epididymal spermatozoa. Am J Vet Res 1980;41:1190–6. Baumgarten HG, Holstein AF, Rosengren E. Arrangement, ultrastructure and adrenergic innervation of smooth musculature of the ductuli efferentes, ductus epididymis and ductus deferens of man. Z Zellforsch Mikrosk Anat 1971;120:37–79. Pholpramool D, Triphrom N. Effects of cholinergic and adrenergic drugs on intraluminal pressures and contractility of the rat testis and epididymis in vivo. J Reprod Fertil 1984;71:181–8. Cosentino MJ, Takihara H, Burhop JW, Crockett AT. Regulation of rat caput epididymides contractility by prostaglandins. J Androl 1984;5:216–22. Filippi S, Vannelli GB, Granchi S, Luconi M, Crescioli C, Mancina R, Ratali R, Brocchi S, Vignozzi L, Bencini E, Noci I, Ledda F, Forti G, Maggi M. Identification, localization and functional activity of oxytocin receptors in epididymis. Mol Cell Endocrinol 2002;193:89–100. Whittington K, Assinder SJ, Parkinson T, Lapwood KR. Function and localization of oxytocin receptors in the reproductive tissue of rams. Reproduction 2001;122:317–25. Studdard PW, Stein JL, Cosentino MJ. The effects of oxytocin and arginine vasopressin in vitro on epididymal contractility in the rat. Int J Androl 2002;25:65–71. Ganjam VK, Amann RP. Testosterone and dihydrotestosterone concentrations in the fluid milieu of spermatozoa in the reproductive tract of the bull. Acta Endocrinol (Copenh) 1973;74:186–200.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 76. Turner TT, Jones CE, Howards SS, Ewing LL, Zegeye B, Gunsalus GL. On the androgen microenvironment of maturing spermatozoa. Endocrinology 1984;115:1925–32. 77. Filippi S, Luconi M, Granchi S, Vignozzi L, Bettuzzi S, Tozzi P, Ledda F, Forti G, Maggi M. Estrogens, but not androgens, regulate expression and functional activity of oxytocin receptor in rabbit epididymis. Endocrinology 2002;143:4271–80. 78. Amann RP, Hammerstedt RH, Veeramachaneni DNR. The epididymis and sperm maturation: a perspective. Reprod Fertil Dev 1993;5:361–81. 79. Bedford JM. Report of a workshop: maturation of the fertilizing ability of mammalian spermatozoa in the male and female reproductive tract. Biol Reprod 1974;11:346– 62. 80. Nicander L. Studies on the regional histology and cytochemistry of the ductus epididymidis in stallions, rams and bulls. Acta Morphol Neerl Scand 1958;1:337–62. 81. Robaire B, Viger RS. Regulation of epididymal epithelial cell functions. Biol Reprod 1995;52:226–36. 82. Gatti J-L, Castella S, Dacheux F, Ecroyd H, Metayer S, Thimon V, Dacheus J-L. Post-testicular sperm environment and fertility. Anim Reprod Sci 2004; 82–83:321–39. 83. Dacheux J-L, Gatti JL, Dacheux F. Contribution of epididymal secretory proteins for spermatozoa maturation. Microsc Res Tech 2003;61:7–17. 84. Dacheux JL, Belghazi M, Lanson Y, Dacheux F. Human epididymal secretome and proteome. Mol Cell Endocrinol 2006;250:36–42. 85. Lye RJ, Hinton BT. Technologies for the study of epididymal-specific genes. Mol Cell Endocrinol 2004;216: 23–30. 86. Suzuki K, Drevet J, Hinton BT, Huhtaniemi I, Lareyre J-J, Matusik RJ, Pons E, Poutanen M, Sipila P, OrgebinCrist M-C. Epididymis-specific promoter-driven targeting: a new approach to control epididymal function. Mol Cell Endocrinol 2004;216:15–22. 87. Turner TT, Johnston DS, Jelinsky SA. Epididymal genomics and the search for a male contraceptive. Mol Cell Endocrinol 2006;250:178–83. 88. Cyr DG, Finnson K, Dufresne J, Gregory M. Cellular interactions and the blood-epididymis barrier. In: Robaire B, Hinton BT (eds) The Epididymis: From Molecules to Clinical Practice. New York: Kluwer Academic/Plenum, 2002; pp. 103–18. 89. Agarwal A, Hoffer AP. Ultrastructural studies on the development of the blood-epididymis barrier in immature rats. J Androl 1989;10:425–31. 90. Cyr DG, Robaire B, Hermo L. Structure and turnover of junctional complexes between principal cells of the rat epididymis. Microsc Res Tech 1995;30:54–66. 91. Levy S, Serre V, Hermo L, Robaire B. The effects of aging on the seminiferous epithelium and the blood-testis barrier of the Brown Norway rat. J Androl 1999;20:356–65. 92. Levy S, Robaire B. Segment-specific changes with age in the expression of junctional proteins and the permeability of the blood-epididymis barrier in rat. Biol Reprod 1999;60:1392–401. 93. Tomsig JL, Turner TT. Growth factors and the epididymis. J Androl 2006;27:348–57. 94. Hinton BT, Lan ZJ, Rudolph DB, Labus JC, Lye RJ. Testicular regulation of epididymal gene expression. J Reprod Fertil Suppl 1998;53:47–57. 95. Hermo L, Barin K, Oko R. Androgen binding protein secretion and endocytosis by principal cells in the adult rat
96.
97.
98.
99. 100.
101.
102.
103.
104.
105.
106.
107.
108.
109.
110.
111. 112.
113. 114.
115.
965
epididymis and during postnatal development. J Androl 1998;19:527–41. Garrett JE, Garrett SH, Douglass JA. A spermatozoaassociated factor regulates proenkephalin gene expression in the rat epididymis. Mol Endocrinol 1990;4:108– 18. Jones R. Plasma membrane structure and remodeling during sperm maturation in the epididymis. J Reprod Fertil Suppl 1998;53:73–84. Belmonte SA, Romano PS, Sosa MA. Mannose-6phosphate receptors as a molecular indicator of maturation of epididymal sperm. Arch Androl 2002;48:53–63. Martin-DeLeon PA. Epididymal SPAM1 and its impact on sperm function. Mol Cell Endocrinol 2006;250:114–21. Zhu G-Z, Myles DG, Primakoff P. Testase 1 (ADAM 24) a plasma membrane-anchored sperm protease implicated in sperm function during epididymal maturation or fertilization. J Cell Sci 2001;114:1787–94. Schroter S, Osterhoff C, McArdle W, Ivell R. The glycocalyx of the sperm surface. Hum Reprod Update 1999;5: 302–13. Tulsiani DRP. Glycan modifying enzymes in luminal fluid of rat epididymis: are they involved in altering sperm surface glycoproteins during maturation? Microsc Res Tech 2003;61:18–27. Tulsiani DRP. Glycan-modifying enzymes in luminal fluid of the mammalian epididymis: an overview of their potential role in sperm maturation. Mol Cell Endocrinol 2006;250:58–65. Roberts KP, Ensrud KM, Wooters JL, Nolan MA, Johnston DS, Hamilton DW. Epididymal secreted protein CRISP-1 and sperm function. Mol Cell Endocrinol 2006; 250:122–7. Yoshinaga K, Toshimori K. Organization and modifications of sperm acrosomal molecules during spermatogenesis and epididymal maturation. Microsc Res Tech 2003;61:39–45. Rodriguez CM, Kirby JL, Hinton BT. Regulation of gene transcription in the epididymis. Reproduction 2001; 122:41–8. Lin M, Lee YH, Xu W, Baker MA, Aitken RJ. Ontogeny of tyrosine phosphorylation-signaling pathways during spermatogenesis and epididymal maturation in the mouse. Biol Reprod 2006;75:588–97. Yeung C-H, Cooper TG. Developmental changes in signalling transduction factors in maturing sperm during epididymal transit. Cell Mol Biol 2003;49:341–9. Saacke RG, Almquist JO. Ultrastructure of bovine spermatozoa. I. The head of normal, ejaculated sperm. Am J Anat 1964;115:143–61. Saacke RG, Almquist JO. Ultrastructure of bovine spermatozoa. II. The neck and tail of normal, ejaculates sperm. Am J Anat 1964;115:163–83. Fawcett DW. The mammalian spermatozoon. Dev Biol 1975;44:394–436. Eddy EM. The Spermatozoon. In: Neill JD (ed.) Knobil and Neill’s Physiology of Reproduction. San Diego: Elsevier Academic Press, 2006; pp. 3–54. Cummins JM, Woodall PF. On mammalian sperm dimensions. J Reprod Fertil 1985;75:153–75. Bielanski W, Kaczmarski F. Morphology of spermatozoa in semen from stallions of normal fertility. J Reprod Fertil Suppl 1979;27:39–45. Dott HM. Morphology of stallion spermatozoa. J Reprod Fertil Suppl 1975;23:41–6.
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116. Gerton GL. Function of the sperm acrosome. In: Hardy DM (ed.) Fertilization. San Diego: Academic Press, 2002; pp. 265–302. 117. Allison AC, Hartree EF. Lysosomal enzymes in the acrosome and their possible role in fertilization. J Reprod Fertil 1970;21:501–15. 118. Abou-Haila A, Tulsiani DRP. Mammalian sperm acrosome: formation, contents, and function. Arch Biochem Biophys 2000;379:173–82. 119. Herr JC. The acrosome. In: Proceedings of the Havemeyer Foundation Workshop on Stallion Reproduction, Bandera, Texas, 2007. 120. Kierszenbaum AL, Tres LL. The acrosome-acroplaxomemanchette complex and the shaping of the spermatid head. Arch Histol Cytol 2004;67:271–84. 121. Toshimori K, Ito C. Formation and organization of the mammalian sperm head. Arch Histol Cytol 2003;66: 383–96. 122. Tachibana M, Terada Y, Murakawa H, Murakami T, Yaegashi N, Okamura K. Dynamic changes in the cytoskeleton during human spermiogenesis. Fertil Steril 2005; 84:1241–7. 123. Holstein AF, Roosen-Runge EC. Atlas of Human Spermatogenesis. Berlin: Grosse Verlag, 1981. 124. Phillips DM. Spermiogenesis. New York: Academic Press, 1974. 125. Love CC, Varner DD, Thompson JA. Intra- and interstallion variation in sperm morphology and their relationship with fertility. J Reprod Fertil Suppl 2000;56:93–100. 126. Storey BT. Energy metabolism of spermatozoa. IV. Effect of calcium on respiration of mature epididymal sperm of the rabbit. Biol Reprod 1975;13:1–9. 127. Hoskins DD, Casillas ER. Function of cyclic nucleotides in mammalian spermatozoa. In: Hamilton DW, Greep RO (eds) Handbook of Physiology. Bethesda, MD: American Physiological Society, 1975; pp. 453–60. 128. Turner TT, Giles RD. A sperm motility-inhibiting factor in the rat epididymis. Am J Physiol 1982;242:R199–203. 129. Usselman MC, Cone RA. Rat sperm are mechanically immobilized in the cauda epididymis by ‘immobilin’, a high molecular weight glycoprotein. Biol Reprod 1983;29: 1241–53. 130. Carr DW, Acott TS. Inhibition of bovine spermatozoa by caudal epididymal fluid. I. Studies of a sperm motility quiescence factor. Biol Reprod 1984;30:913–25. 131. Acott TS, Carr DW. Inhibition of bovine spermatozoa by caudal epididymal fluid. II. Interaction of pH and a quiescence factor. Biol Reprod 1984;30:926–35. 132. Babcock DF. Examination of the intracellular ionic environment and of ionophore action by null point measurements employing the fluorescein chromophore. J Biol Chem 1983;258:6380–9. 133. Zaneveld LJD. The epididymis. In: Zaneveld LJD, Chatterton RT (eds) Biochemistry of Mammalian Reproduction. New York: John Wiley & Sons, 1982; pp. 37–63. 134. Holt WV, Harrison RA. Bicarbonate stimulation of boar sperm motility via a protein kinase a-dependent pathway: between-cell and between ejaculate differences are not due to deficiencies in protein kinase A activation. J Androl 2002;23:557–65. 135. Okamura N, Tajima Y, Soejima A, Masuda H, Sugita Y. Sodium bicarbonate in seminal plasma stimulates the motility of mammalian spermatozoa through direct activation of adenylate cyclase. J Biol Chem 1985;260: 9699–705.
136. Mann T, Lutwak-Mann C. Biochemistry of seminal plasma and male accessory fluids: application to andrological problems. In: Mann T, Lutwak-Mann C (eds) Male Reproductive Function and Semen. New York: SpringerVerlag, 1981; pp. 269–336. 137. Gagnon C. Regulation of sperm motility at the axonemal level. Reprod Fertil Dev 1995;7:847–55. 138. Luconi M, Baldi E. How do sperm swim? Molecular mechanisms underlying sperm motility. Cell Mol Biol 2003;49:357–69. 139. Tash JS, Means AR. Regulation of protein phosphorylation and motility of sperm by cyclic adenosine monophosphate and calcium. Biol Reprod 1982;26:745– 63. 140. Demarco IA, Espinosa F, Edwards J, Sosnik J, DeLaVegaBeltran JL, Hockensmith JW, Kopf GS, Darszon A, Visconti PE. Involvement of a Na+/HCO-3 cotransporter in mouse sperm capacitation. J Biol Chem 2003;278:7001–9. 141. Garbers DL, Tubb TJ, Hyne RV. A requirement of bicarbonate for Ca2+ -induced elevations of cyclic AMP in guinea pig spermatozoa. J Biol Chem 1982;257:8980–4. 142. Quill TA, Wang D, Garbers DL. Insights into sperm cell motility signaling through sNHE and the CatSpers. Mol Cell Endocrinol 2006;250:84–92. 143. Woo AL, James PF, Lingrel JB. Roses of the Na,K-ATPase alpha 4 isoform and the Na+/H+ exchanger in sperm motility. Mol Reprod Dev 2002;62:348–56. 144. Woo AL, James PF, Lingrel JB. Sperm motility is dependent on a unique isoform of the Na,K-ATPase. J Biol Chem 2000;275:20693–9. 145. Wang D, King SM, Quill TA, Doolittle LK, Garbers DL. A new sperm-specific Na+/H+ exchanger required for sperm motility and fertility. Nat Cell Biol 2003;5:1117–22. 146. Malo ME, Fliegel L. Physiological role and regulation of the Na+ /H+ exchanger. Can J Physiol Pharmacol 2006; 84:1081–95. 147. Kirichok Y, Navarro B, Clapham DE. Whole-cell patchclamp measurements of spermatozoa reveal an alkalineactivated Ca2+ channel. Nature 2006;439:737–40. 148. Marquez B, Suarez SS. Different signaling pathways in bovine sperm regulate capacitation and hyperactivation. Biol Reprod 2004;70:1626–33. 149. Suarez SS, Ho HC. Hyperactivated motility in sperm. Reprod Domest Anim 2003;38:119–24. 150. Ho HC, Granish KA, Suarez SS. Hyperactivated motility of bull sperm is triggered at the axoneme by Ca2+ and not cAMP. Dev Biol 2002;250:208–17. 151. Ignotz GG, Suarez SS. Calcium/calmodulin and calmodulin kinase II stimulate hyperactivation in demembranated bovine sperm. Biol Reprod 2005;73:519–26. 152. Ho HC, Suarez SS. Hyperactivation of mammalian spermatozoa: function and regulation. Reproduction 2001; 122:519–26. 153. Morita Z, Chang MC. The motility and aerobic metabolism of spermatozoa in laboratory animals with special reference to the effects of cold shock and the importance of calcium for the motility of hamster spermatozoa. Biol Reprod 1970;3:169–79. 154. Morton BE, Sagadraca R, Fraser C. Sperm motility within the mammalian epididymis: species variation and correlation with free calcium levels in epididymal plasma. Fertil Steril 1978;29:695–8. 155. Chinoy JJ, Verma RJ, Patel KG. Effect of calcium on sperm motility of cauda epididymis in vitro. Acta Eur Fertil 1983;14:421–3.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 156. Aoki F, Sakai S, Kohmoto K. Regulation of flagellar bending by cAMP and Ca2+ in hamster sperm. Mol Reprod Dev 1999;53:77–83. 157. Fraser LR. Cellular biology of capacitation and the acrosome reaction. Hum Reprod 1995;10:22–30. 158. Tash JS. Protein phosphorylation: the second messenger signal transducer of flagellar motility. Cell Motil Cytoskeleton 1989;14:332–9. 159. Vijayaraghavan S, Hoskins DD. Regulation of bovine sperm motility and cyclic adenosine 3’,5’monophosphate by adenosine and its analogues. Biol Reprod 1986;34:468–77. 160. Skalhegg BS, Huang Y, Idzerda RL, McKnight GS, Burton KA. Mutation of the Calpha subunit of PKA leads to growth retardation and sperm dysfunction. Mol Endocrinol 2002;16:630–9. 161. Brandt H, Hoskins DD. A cAMP-dependent phosphorylated motility protein in bovine epididymal sperm. J Biol Chem 1980;255:982–7. 162. Garbers DL, Kopf GS. The regulation of spermatozoa by calcium and cyclic nucleotides. In: Greengard P, Robison GA (eds) Advances in cyclic nucleotide research. New York: Raven Press, 1980; pp. 251–306. 163. Tash JS, Means AR. Cyclic adenosine 3’,5’ monophosphate, calcium and protein phosphorylation in flagellar motility. Biol Reprod 1983;28:75–104. 164. Lindermann CB, Kanous KS. Regulation of mammalian sperm motility. Arch Androl 1989;23:1–22. 165. Luconi M, Proazz I, Ferruzzi P, Marchiani S, Forti G, Baldi E. Tyrosine phosphorylation of the a kinase anchoring protein 3 (AKAP3) and soluble adenylate cyclase are involved in the increase of human sperm motility by bicarbonate. Biol Reprod 2005;72:22–32. 166. Hess KC, Jones BH, Marquez B, Chen Y, Ord TS, Kamenetsky M, Miyamoto C, Zippin JH, Kopf GS, Suarez SS, Levin LR, Williams CJ, Buck J, Moss SB. The ‘soluble’ adenylyl cyclase in sperm mediates multiple signaling events required for fertilization. Dev Cell 2005;9:249– 59. 167. Chen Y, Cann MJ, Litvin TN, Iourgenko V, Sinclair ML, Levin LR, Buck J. Soluble adenylyl cyclase as an evolutionarily conserved bicarbonate sensor. Science 2000;289: 625–8. 168. Wuttke MS, Buck J, Levin LR. Bicarbonate-regulated soluble adenylyl cyclase. JOP 2001;2:154–8. 169. Litvin TN, Kamenetsky M, Zarifyan A, Buck J, Levin LR. Kinetic properties of ‘soluble’ adenylyl cyclase; synergism between calcium and bicarbonate. J Biol Chem 2003; 278:15922–6. 170. Jaiswal BS, Conti M. Calcium regulation of the soluble adenylyl cyclase expressed in mammalian spermatozoa. Proc Natl Acad Sci USA 2003;100:10676–81. 171. Turner RM. Moving to the beat: a review of mammalian sperm motility regulation. Reprod Fertil Dev 2006;18: 25–38. 172. Tash JS. Protein phosphorylation: the second messenger signal transducer of flagellar motility. Cell Motil Cytoskeleton 1989;14:332–9. 173. Leclerc P, deLamirande E, Gagnon C. Cyclic adenosine 3’,5’ monophosphate-dependent regulation of protein tyrosine phosphorylation in relation to human sperm capacitation and motility. Biol Reprod 1996;55:684–92. 174. Tash JS, Bracho GE. Regulation of sperm motility: emerging evidence for a major role for protein phosphatases. J Androl 1994;15:505–9.
967
175. Inaba K. Molecular architecture of the sperm flagella molecules for motility and signaling. Zoolog Sci 2003;20: 1043–56. 176. Visconti PE, Johnson LR, Oyaski M, Fornes M, Moss SB, Gerton GL, Kopf GS. Regulation, localization, and anchoring of protein kinase A subunits during mouse sperm capacitation. Dev Biol 1997;192:351–63. 177. Hall DD, Davare MA, Shi M, Allen ML, Weisenhaus M, McKnight SG, Hell JW. Critical role of cAMP-dependent protein kinase anchoring to the L-type calcium channel Cav 1. 2 via A-kinase anchor protein 150 in neurons. Biochemistry 2007;46:1635–46. 178. Turner RMO, Casas-Dolz R, Schlingmann KL. Characterization of an A-kinase anchor protein in equine spermatozoa and examination of the effect of semen cooling and cryopreservation on the binding of that protein to the regulatory subunit of protein kinase-A. Am J Vet Res 2005;66:1056–64. 179. Burton KA, Treash-Osio B, Muller CH, Dunphy EL, McKnight GS. Deletion of type II alpha regulatory subunit delocalizes protein kinase A in mouse sperm without affecting motility or fertilization. J Biol Chem 1999;274:24131– 6. 180. Nakano I, Kobayashi T, Yoshimura M, Shingyoji C. Central-pair-linked regulation of microtubule sliding by calcium in flagellar axonemes. J Cell Sci 2003;116:1627– 36. 181. Bannai H, Yoshimura M, Takahashi K, Shingyoji C. Calcium regulation of microtubule sliding in reactivated sea urchin sperm flagella. J Cell Sci 2000;113:831–9. 182. Gross MK, Toscano DG, Toscano Jr WA. Calmodulinmediated adenylate cyclase from mammalian sperm. J Biol Chem 1987;262:8672–6. 183. White DR, Aitken RJ. Relationship between calcium, cyclic AMP, ATP, and intracellular pH and the capacity of hamster spermatozoa to express hyperactivated motility. Gamete Res 1989;22:163–77. 184. Si Y, Olds-Clark P. Evidence for the involvement of calmodulin in mouse sperm capacitation. J Biol Chem 2000;262:8672–6. 185. Ahmad K, Bracho GE, Wolf DP, Tash JS. Regulation of human sperm motility and hyperactivation components by calcium, calmodulin, and protein phosphatases. Arch Androl 1995;35:187–208. 186. Carafoli E, Santella L, Branca D, Brini M. Generation, control, and processing of cellular calcium signals. Crit Rev Biochem Mol Biol 2001;36:107–260. 187. Carrera A, Moos J, Ning XP, Gerton FL, Texarik J, Kopf FS, Moss SB. Regulation of protein tyrosine phosphorylation in human sperm by a calcium/calmodulindependent mechanism: identification of A kinase anchor proteins as major substrates for tyrosine phosphorylation. Dev Biol 1996;180:284–96. 188. Tash JS, Bracho GE. Identification of phosphoproteins coupled to initiation of motility in live epididymal mouse sperm. Biochem Biophys Res Commun 1998;251:557–63. 189. Lopez-Gonzalez I, DeLaVega-Beltran JL, Santi CM, Florman HM, Felix R, Darszon A. Calmodulin antagonists inhibit T-type Ca2+ currents in mouse spermatogenic cells and the zona pellucida-induced sperm acrosome reaction. Dev Biol 2001;236:210–19. 190. Nevo AC, Rikmenspoel R. Diffusion of ATP in sperm flagella. J Theor Biol 1970;26:11–18. 191. Adam DE, Wei J. Mass-transport of ATP within motile sperm. J Theor Biol 1975;49:125–45.
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192. Ford WCL. Glycolysis and sperm motility: does a spoonful of sugar help the flagellum go round? Hum Reprod Update 2006;12:269–74. 193. Cao W, Haig-Ladewig L, Gerton GL, Moss SB. Adenylate kinase 1 and 2 are part of the accessory structures in the mouse sperm flagellum. Biol Reprod 2006;75:492–500. 194. Miki K, Qu W, Goulding EH, Willis WD, Bunch DO, Strader LF, Perreault SD, Eddy EM, O’Brian DA. Glyceraldehyde 3-phosphate dehydrogenase-S, a spermspecific glycolytic enzyme, is required for sperm motility and male fertility. Proc Natl Acad Sci USA 2004;101: 16501–6. 195. Muka C, Okuno M. Glycolysis plays a major role for adenosine triphosphate supplementation in mouse sperm flagellar movement. Biol Reprod 2004;71:540–7. 196. Eddy EM, Toshimori K, O’Brien DA. Fibrous sheath of mammalian spermatozoa. Microsc Res Tech 2003;61: 103–15. 197. Storey BT, Kayne FJ. Properties of pyruvate kinase and flagellar ATPase in rabbit spermatozoa: relation to metabolic strategy of the sperm cell. J Exp Zool 1980;211: 361–7. 198. Travis AJ, Foster JA, Rosenbaum NA, Visconti PE, Gerton GL, Kopf GS, Moss SB. Targeting of a germ cell-specific type 1 hexokinase lacking a porin-binding domain to the mitochondria as well as to the head and fibrous sheath of murine spermatozoa. Mol Biol Cell 1998;9:263–76. 199. Westhoff D, Kamp G. Glyceraldehyde 3-phosphate dehydrogenase is bound to the fibrous sheath of mammalian spermatozoa. J Cell Sci 1997;110:1821–9. 200. Mori C, Nakamura N, Welch JE, Gotoh H, Goulding EH, Fujioka M, Eddy EM. Mouse spermatogenic cell-specific type 1 hexokinase (mHk1-s) transcripts are expressed by alternative splicing form the mHk1 gene and the HK1-S protein is localized mainly in the sperm tail. Mol Reprod Dev 1998;49:374–85. 201. Bunch DO, Welch JE, Magyar PL, Eddy EM, O’Brian DA. Glyceraldehyde 3-phosphate dehydrogenase-S protein distribution during mouse spermatogenesis. Biol Reprod 1998;58:834–41. 202. Bradley MP, Geelan A, Leitch V, Goldberg E. Cloning, sequencing, and characterization of LDH-C4 from a fox testis cDNA library. Mol Reprod Dev 1996;44:452–9. 203. Ford WCL, Harrison A. The effect of 6-chloro-6deoxysugars on adenine nucleotide concentrations in and motility of rat spermatozoa. J Reprod Fertil 1981;63: 75–9. 204. Nicastro D, Schwartz C, Pierson J, Gaudette R, Porter ME, Mcintosh JR. The molecular architecture of axonemes revealed by cryoelecron tomography. Science 2006; 313:944–8. 205. Lupetti P, Lanzavecchia S, Mercati D, Cantele F, Dallai R, Mencarelli C. Three-dimensional reconstruction of axonemal outer dynein arms in situ by electron tomography. Cell Motil Cytoskeleton 2005;62:69–83. 206. Shimizu T, Johnson KA. Kinetic evidence for multiple dynein ATPase sites. J Biol Chem 1983;258:13841–6. 207. Holzbaur EL, Johnson KA. Microtubules accelerate ADP release by dynein. Biochemistry 1989;28:7010–16. 208. Johnson KA. Pathway of the microtubule-dynein ATPase and the structure of dynein: a comparison with actomyosin. Annu Rev Biophys Biophys Chem 1985;14:161–88. 209. Fujimura M, Okuno M. Requirement of the fixed end for spontaneous beating in flagella. J Exp Biol 2006;209: 1336–43.
210. Baltz JM, Willams O, Cone RA. Dense fibers protect mammalian sperm against damage. Biol Reprod 1990;43: 485–91. 211. Hinsch KD, DePinto V, Aires VA, Schneider X, Messina A, Hinsch E. Voltage-dependent anion-selective channels VDAC2 and VDAC3 are abundant proteins in bovine outer dense fibers, a cytoskeletal component of the sperm flagellum. J Biol Chem 2004;279:15281–8. 212. Wargo MJ, Smith EF. Asymmetry of the central apparatus defines the location of active microtubule sliding in Chlamydomonas flagella. Proc Natl Acad Sci USA 2003; 100:137–42. 213. Porter ME, Johnson KA. Dynein structure and function. Annu Rev Cell Biol 1989;5:119–51. 214. Amann RP, Graham JK. Spermatozoal function. In: DiRienzi D (ed.) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 715–45. 215. Hammerstedt RH, Volonte C, Racker E. Motility, heat, and lactate production in ejaculated bovine sperm. Arch Biochem Biophys 1988;266:111–23. 216. Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P. Cell chemistry and biosynthesis. In: Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P (eds) Molecular Biology of the Cell. New York: Garland Science, 2002; pp. 47–127. 217. Brooks DE, Mann T. Relation between the oxidation state of nicotinamide-adenine dinucleotide and the metabolism of spermatozoa. Biochem J 1972;129:1023– 34. 218. Mann T, Lutwak-Mann C. Biochemistry of spermatozoa: chemical and functional correlations in ejaculated semen, andrological aspects. In: Mann T, Lutwak-Mann C (eds) Male Reproductive Function and Semen. New York: Springer-Verlag, 1981; pp. 195–268. 219. Storey BT, Kayne FJ. Energy metabolism of spermatozoa. V. The Embden-Myerhof pathway of glycolysis: activities of pathway enzymes in hypotonically treated rabbit epididymal spermatozoa. Fertil Steril 1975;26:1257–65. 220. Mann T. Metabolism of semen: fructolysis, respiration and sperm energetics. In: Mann T (ed.) The Biochemistry of Semen and of the Male Reproductive Tract. New York: Barnes and Noble, 1964; pp. 265–307. 221. Stryer L. Glycolysis. In: Stryer L (ed.) Biochemistry. New York: W.H. Freeman, 1988; pp. 349–72. 222. Dringen R, Bergbauer K, Wiesinger H, Hamprecht B. Utilization of mannose by astroglial cells. Neurochem Res 1994;19:23–30. 223. Mann T. Studies on the metabolism of semen. 3. Fructose as a normal constituent of seminal plasma. Site of formation and function of fructose in semen. Biochem J 1946;40:481. 224. Kosiniak K. Characteristics of the successive jets of ejaculated semen of stallions. J Reprod Fertil Suppl 1975;23: 59–61. 225. Ponglowhapan S, Essen-Gustavsson B, Forsberg CL. Influence of glucose and fructose in the extender during long-term storage of chilled canine semen. Theriogenology 2004;62:1498–517. 226. Rigau T, Farre M, Ballester J, Mogas T, Pena A, Rodriquez-Gil JE. Effects of glucose and fructose on motility patterns of dog spermatozoa from fresh ejaculates. Theriogenology 2001;56:801–15. 227. Williams AC, Ford WC. The role of glucose in supporting motility and capacitation in human spermatozoa. J Androl 2001;22:680–95.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 228. Nagai T, Yamaguchi K, Moriwaki C. Studies on the effects of sugars on washed human sperm motility. J Pharmacobiodyn 1982;5:564–7. 229. Hoppe PC. Glucose requirement for mouse sperm capacitation in vitro. Biol Reprod 1976;15:39–45. 230. Niwa K, Iritani A. Effect of various hexoses on sperm capacitation and penetration of rat eggs in vitro. J Reprod Fertil 1978;53:267–71. 231. Fraser LR, Quinn PJ. A glycolytic product is obligatory for initiation of the sperm acrosome reaction and whiplash motility required for fertilization in the mouse. J Reprod Fertil 1981;61:25–35. 232. Sakkas D, Urner F, Menezo Y, Leppens G. Effects of glucose and fructose on fertilization, cleavage, and viability of mouse embryos in vitro. Biol Reprod 1993;49:1288– 92. 233. Urner F, Sakkas D. Glucose participates in sperm-oocyte fusion in the mouse. Biol Reprod 1996;55:917–22. 234. Urner F, Leppens-Luisier G, Sakkas D. Protein tyrosine phosphorylation in sperm during gamete interaction in the mouse: the influence of glucose. Biol Reprod 2001; 64:1350–7. 235. Urner F, Sakkas D. Protein phosphorylation in mammalian spermatozoa. Reproduction 2003;125:17–26. 236. Hiipakka RA, Hammerstedt RH. 2-deoxyglucose transport and phosphorylation by bovine sperm. Biol Reprod 1978;19:368–79. 237. Mukai C, Okuno M. Glycolysis plays a major role for adenosine triphosphate supplementation in mouse sperm flagellar movement. Biol Reprod 2004;71:540–7. 238. Ford WCL, Rees JM. The bioenergetics of mammalian sperm motility. In: Gagnon C (ed.) Controls of Sperm Motility: Biological and Clinical Aspects. Boca Raton, FL: CRC Press, 1990; pp. 175–202. 239. Marin S, Chiang K, Bassilian S, Lee W-NP, Boros LG, Fernandez-Novell JM, Centelles JJ, Medrano A, Rodriguez-Gil JE, Cascante M. Metabolic strategy of boar spermatozoa revealed by a metabolomic characterization. FEBS Lett 2003;20:342–6. 240. Bishop DW. Oxygen concentration in the rabbit genital tract. In: Proceedings of the 3rd International Congress on Animal Reproduction, 1956, pp. 53. 241. Mastroianni L, Jones R. Oxygen tension within the fallopian tube. J Reprod Fertil 1956;9:99. 242. Jones R, Mann T, Sherins R. Peroxidative breakdown of phospholipids in human spermatozoa, spermicidal properties of fatty acid peroxides, and protective action of seminal plasma. Fertil Steril 1979;31:531–7. 243. Holland MK, Storey BT. Oxygen metabolism of mammalian spermatozoa. Generation of hydrogen peroxide by rabbit epididymal spermatozoa. Biochem J 1981;198: 273–80. 244. Alvarez JG, Storey BT. Assessment of cell damage caused by spontaneous lipid peroxidation in rabbit spermatozoa. Biol Reprod 1984;30:323–31. 245. Alvarez JG, Storey BT. Lipid peroxidation and the reactions of superoxide and hydrogen peroxide in mouse spermatozoa. Biol Reprod 1984;30:833–41. 246. Aitken RJ, Clarkson JS. Significance of reactive oxygen species and antioxidants in defining the efficacy of sperm preparation techniques. J Androl 1987;9:367–76. 247. Hammerstedt RH, Lovrien RE. Calorimetric techniques for metabolic studies of cells and organisms under normal conditions and stress. J Exp Zool 1983;228:459– 69.
969
248. Inskeep PB, Hammerstedt RH. Endogenous metabolism by sperm in response to altered cellular ATP requirements. J Cell Physiol 1985;123:180–90. 249. Ballester J, Fernandez-Novell JM, Rutlant J, Garcia-Rocha M, Palomo MJ, Mogas T, Pena A, Rigau T, Guinovart JJ, Rodriguez-Gil JE. Evidence for a functional glycogen metabolism in mature mammalian spermatozoa. Mol Reprod Dev 2000;56:207–19. 250. Palomo MJ, Fernandez-Novell JM, Pena A, Guinovart JJ, Rigau T, Rodriguez-Gil JE. Glucose- and fructoseinduced dog-sperm glycogen synthesis shows specific changes in the location of the sperm glycogen deposition. Mol Reprod Dev 2003;64:349–59. 251. Urner F, Sakkas D. Characterization of glycolysis and pentose phosphate pathway activity during sperm entry into the mouse oocyte. Biol Reprod 1999;60:973–8. 252. Urner F, Sakkas D. Involvement of the pentose phosphate pathway and redox regulation in fertilization in the mouse. Mol Reprod Dev 2005;70:494–503. 253. Ford WCL, Whittington K, Williams AC. Reactive oxygen species in human sperm suspensions: production by leukocytes and the generation of NADPH to protect sperm against their effects. Int J Androl 1997;20 (Suppl 3): 44–9. 254. Williams AC, Ford WCL. Functional significance of the pentose phosphate pathway and glutathione reductase in the antioxidant defenses of human sperm. Biol Reprod 2004;71:1309–16. 255. Halliwell B, Gutteridge JMC. The chemistry of free radicals and related ‘reactive species’. In: Halliwell B, Gutteridge JMC (eds) Free Radicals in Biology and Medicine. Oxford: Oxford University Press, 1999; pp. 36–104. 256. Alvarez JG, Storey BT. Role of glutathione peroxidase in protecting mammalian spermatozoa from loss of motility caused by spontaneous lipid peroxidation. Gamete Res 1989;23:77–90. 257. Aitken RJ, Clarkson JS, Fishel S. Generation of reactive oxygen species lipid peroxidation and human sperm function. Biol Reprod 1989;40:183–97. 258. Aitken RJ. Free radicals, lipid peroxidation and sperm function. Reprod Fertil Dev 1995;7:659–68. 259. Aitken RJ. Oxidative considerations relating to capacitation and the acrosome reaction: from studs to sterility. In: Proceedings of the Havemeyer Foundation Workshop on Stallion Reproduction, Bandera, Texas, 2007. 260. Baumber J, Ball BA, Gravance CG, Medina V, DaviesMorel MCG. The effect of reactive oxygen species on equine sperm motility, viability, acrosomal integrity, mitochondrial membrane potential, and membrane lipid peroxidation. J Androl 2000;21:895–902. 261. Ford WCL. Reactive oxygen species and sperm. Hum Fertil 2001;4:77–8. 262. Agarwal A, Prabakaran S, Allamaneni S. What an andrologist/urologist should know about free radicals and why. Urology 2006;67:2–8. 263. Chandel NS, Budinger GRS. The cellular basis for diverse responses to oxygen. Free Radic Biol Med 2007;42:165– 74. 264. Sikka SC, Rajasekaran M, Hellstrom WJG. Role of oxidative stress and antioxidants in male infertility. J Androl 1995;16:464–8. 265. De Iuliis GN, Wingate JK, Kopers AJ, McLaughlin EA, Aitken RJ. Definitive evidence for the nonmitochondrial production of superoxide anion by human spermatozoa. J Clin Endrocrinol Metab 2006;91:1968–75.
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266. Burnaugh L, Sabeur K, Ball BA. Generation of superoxide anion by equine spermatozoa as detected by dihydroethidium. Theriogenology 2007;67:580–9. 267. Ball BA, Baumber J, Sabeur K. Role of reactive oxygen species on normal and abnormal function of equine spermatozoa. Theriogenology 2002;58:299–300. 268. Banfi B, Molnar G, Maturana A, Steger K, Hegedus B, Demaurex N, Krause K-H. A Ca2+-activated NADPH oxidase in testis, spleen, and lymph nodes. J Biol Chem 2001;276:37594–601. 269. Armstrong JS, Bivalacqua TJ, Chamulitrat W, Sikka S, Hellstrom WJG. A comparison of the NADPH oxidase in human sperm and white blood cells. Int J Androl 2002; 25:223–9. 270. Sabeur K, Ball BA. Detection of superoxide anion generation by equine spermatozoa. Am J Vet Res 2006;67:701–6. 271. Richer SC, Ford WC. A critical investigation of NADPH oxidase activity in human spermatozoa. Mol Hum Reprod 2001;7:237–44. 272. Ford WCL. Regulation of sperm function by reactive oxygen species. Hum Reprod Update 2004;10:387–99. 273. Upreti GC, Jensen K, Munday R, Vishwanath R, Smith JG. Studies on ram spermatozoal aromatic amino acid oxidase. In: Proceedings of the 26th Annual Conference of the Australian Society for Reproductive Biology, 1994, p. 115. 274. Tosic J, Walton A. Metabolism of spermatozoa. The formation and elimination of hydrogen peroxide by spermatozoa and effects on motility and survival. Biochem J 1950;47:199–212. 275. Shannon P, Curson B. Kinetics of the aromatic L-amino acid oxidase from dead bovine spermatozoa and the effect of catalase on fertility of diluted bovine semen stored at 5C and ambient temperatures. J Reprod Fertil 1982;64:463–7. 276. Wai-sum O, Chen H, Chow PH. Male genital tract antioxidant enzymes: their ability to preserve sperm DNA integrity. Mol Cell Endocrinol 2006;250:80–3. 277. Gomez E, Buckingham DW, Brindle J, Lanzafame F, Irvine DS, Aitken RJ. Development of an image analysis system to monitor the retention of residual cytoplasm by human spermatozoa: correlation with biochemical markers of the cytoplasmic space, oxidative stress, and sperm function. J Androl 1996;17:276–87. 278. Aitken RJ, Krausz C, Buckingham DW. Relationship between the biochemical marker for human sperm quality, creatine kinase, and the mechanisms responsible for the generation of reactive oxygen species. Mol Reprod Dev 1994;39:268–79. 279. Rao B, Soufir JC, Martin M, David G. Lipid peroxidation in human spermatozoa as related to midpiece abnormalities and motility. Gamete Res 1989;24:127–34. 280. Iwasaki A, Gagnon C. Formation of reactive oxygen species in spermatozoa of infertile patients. Fertil Steril 1992;57:409–16. 281. Aitken RJ. Founders’ Lecture. Human spermatozoa: fruits of creation, seeds of doubt. Reprod Fertil Dev 2004; 16:655–64. 282. Ball BA, Vo AT, Baumber J. Generation of reactive oxygen species by equine spermatozoa. Am J Vet Res 2001; 62:508–15. 283. Krausz C, West K, Buckingham D, Aitken RJ. Development of a technique for monitoring the contamination of human semen samples with leukocytes. Fertil Steril 1992;57:1317–25.
284. Aitken RJ, West K, Buckingham D. Leukocytic infiltration into the human ejaculate and its association with semen quality, oxidative stress, and sperm function. J Androl 1994;15:343–52. 285. Baumber J, Vo A, Sabeur K, Ball BA. Generation of reactive oxygen species by equine neutrophils and their effect on motility of equine spermatozoa. Theriogenology 2002;57:1025–33. 286. Jain S, Saxen D, Kumar PG, Laloraya M. NADPH dependent superoxide generation in the ovary and uterus of mice during estrous cycle and early pregnancy. Life Sci 2000;66:1139–46. 287. Norman JE, Cameron IT. Nitric oxide in the human uterus. Rev Reprod 1996;1:61–8. 288. Huang Y, Roby KF, Pace JL, Russell SW, Hung JS. Cellular localization and hormonal regulation of inducible nitric oxide synthase in cycling mouse uterus. J Leukoc Biol 1995;57:27–35. 289. Hansen PJ, Hoggard MP, Rathwell AC. Effects of stallion seminal plasma on hydrogen peroxide release by leukocytes exposed to spermatozoa and bacteria. J Reprod Immunol 1987;10:157–66. 290. Kovalski NN, de Lamirande E, Gagnon C. Reactive oxygen species generated by human neutrophils inhibit sperm motility: protective effect of seminal plasma and scavengers. Fertil Steril 1992;58:809–16. 291. Smith R, Vantman D, Ponce J, Escobar J, Lissi E. Total antioxidant capacity of human seminal plasma. Hum Reprod 1996;11:1655–60. 292. Lewis SEM, Boyle PM, McKinney KA, Young IS, Thompson W. Total antioxidant capacity of seminal plasma is different in fertile and infertile men. Fertil Steril 1995;64:868–70. 293. Lewis SE, Steriling ESL, Young IS, Thompson W. Comparison of individual antioxidants of sperm and seminal plasma in fertile and infertile men. Fertil Steril 1997;67: 142–7. 294. Aitken RJ, Clarkson JS. Cellular basis of defective sperm function and its association with the genesis of reactive oxygen species by human spermatozoa. J Reprod Fertil 1987;81:459–69. 295. Jones R, Mann T. Lipid peroxides in spermatozoa: formation, role of plasmalogen and physiological significance. Proc R Soc Lond B 1976;193:235–43. 296. Jones R, Mann T. Toxicity of exogenous fatty acid peroxides towards spermatozoa. J Reprod Fertil 1977;50:255–60. 297. Jones R, Mann T, Sherins R. Peroxidative breakdown of phospholipids in human spermatozoa, spermicidal properties of fatty acid peroxides and protective action of seminal plasma. Fertil Steril 1979;31:531–7. 298. Lenzi A, Picardo M, Gandini L, Dondero F. Lipids of the sperm plasma membrane: from polyunsaturated fatty acids considered as markers of sperm function to possible scavenger therapy. Hum Reprod Update 1996;2:246–56. 299. Ball BA, Vo A. Detection of lipid peroxidation in equine spermatozoa based upon the lipophilic fluorescent dye C11-BODIPY581/591. J Androl 2002;23:259–69. 300. Aitken RJ, Wingate JK, De Iuliis GN, Koppers AJ, McLaughlin EA. Cis-Unsaturated fatty acids stimulate reactive oxygen species generation and lipid peroxidation in human spermatozoa. J Clin Endocrinol Metab 2006; 91:4154–63. 301. Armstrong JS, Rajasekaran M, Chamulitrat W, Gatti P, Hellstrom WJ, Sikka SC. Characterization of reactive oxygen species induced effects on human spermatozoa
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movement and energy metabolism. Free Radic Biol Med 1999;26:869–80. Yonka C, James PS, Jones R. Lipid diffusion in sperm plasma membranes exposed to peroxidative injury from oxygen free radicals. Mol Reprod Dev 2004;68:365–72. de Lamirande E, Gagnon C. Reactive oxygen species and human spermatozoa. I. Effects on the motility of intact spermatozoa and on sperm axonemes. J Androl 1992;13:368–78. de Lamirande E, Gagnon C. Reactive oxygen species and human spermatozoa. II. Depletion of adenosine triphosphate plays an important role in the inhibition of sperm motility. J Androl 1992;13:379–86. Williams AC, Ford WCL. Relationship between reactive oxygen species production and lipid peroxidation in human sperm suspensions and their association with sperm function. Fertil Steril 2005;83:929–36. Dizdaroglu M, Jaruga P, Birincioglu M, Rodriguez H. Free radical-induced damage to DNA: mechanisms and measurement. Free Radic Biol Med 2002;32:1102–15. Aitken RJ, Baker MA. Oxidative stress, sperm survival and fertility control. Mol Cell Endocrinol 2006;250:66–9. Aitken RJ, Koopman P, Lewis SE. Seeds of concern. Nature 2004;432:48–52. Sawyer DE, Mercer BG, Wiklendt AM, Aitken RJ. Quantitative analysis of gene-specific DNA damage in human spermatozoa. Mutat Res 2003;529:21–34. Baumber J, Ball BA, Linfor JJ, Meyers SA. Reactive oxygen species and cryopreservation promote deoxyribonucleic acid (DNA) damage in equine sperm. Theriogenology 2003;58:301–2. Pagl R, Aurich C, Kankifer M. Anti-oxidative status and semen quality during cooled storage in stallion. J Vet Med A Physiol Pathol Clin Med 2006;53:486–9. Holland JK, Alverez JG, Storey BT. Production of superoxide and activity of superoxide dismutase in rabbit epididymal spermatozoa. Biol Reprod 1982;27:1109–18. Storey BT. Biochemistry of the induction and prevention of lipoperoxidative damage in human spermatozoa. Mol Hum Reprod 1997;3:203–13. McLeod J. The role of oxygen in the metabolism and motility of human spermatozoa. Am J Physiol 1943;138: 512–18. Baumber J, Ball BA. Determination of glutathione peroxidase and superoxide dismutase-like activities in equine spermatozoa, seminal plasma, and reproductive tissues. Am J Vet Res 2005;66:1415–19. Li T. The glutathione and thiol content of mammalian spermatozoa and seminal plasma. Biol Reprod 1975;12: 641–6. Drevet JR. The antioxidant glutathione peroxidase family and spermatozoa: a complex story. Mol Cell Endocrinol 2006;250:70–9. Chance B, Sies H, Boveris A. Hydrogen peroxide metabolism in mammalian organs. Physiol Rev 1979;59: 527–605. Maiorino M, Ursini F. Oxidative stress, spermatogenesis and fertility. Biol Chem 2002;383:591–7. Stradaioli G, Rubei M, Zamparini M, Tubaro F, Valentini S, Degl’Innocenti S, Monaci M. Enzymatic evaluation of phospholipid hydroperoxide glutathione peroxidase (PHGPx) in equine spermatozoa. Anim Reprod Sci 2006;94:29–31. Foote RH. Catalase content of rabbit, ram, bull and boar semen. J Anim Sci 1962;21:966–8.
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322. White IG. Studies on semen. Aust J Biol Sci 1959;21: 208–14. 323. Alvarez JG, Touchstone JC, Blasco L, Storey BT. Spontaneous lipid peroxidation and production of hydrogen peroxide and superoxide in human spermatozoa. J Androl 1987;8:338–48. 324. Bilodeau JF, Chatterjee S, Sirard MA, Gagnon C. Levels of antioxidant defenses are decreased in bovine spermatozoa after a cycle of freezing and thawing. Mol Reprod Dev 2000;55:282–8. 325. Foote RH, Hare E. High catalase content of rabbit semen appears to be inherited. J Androl 2000;21:664–8. 326. Ball BA, Gravance CG, Medina V, Baumber J, Liu IKM. Catalase activity in equine semen. Am J Vet Res 2000;61:1026–30. 327. Aitken RJ, Buckingham D, Harkiss D. Use of a xanthine oxidase oxidant generating system to investigate the cytotoxic effects of reactive oxygen species on human spermatozoa. J Reprod Fertil 1993;97:441–50. 328. Griveau JF, Dumont E, Renard P, Callegari JP, Le Lannou D. Reactive oxygen species, lipid peroxidation and enzymatic defence systems in human spermatozoa. J Reprod Fertil 1995;103:17–26. 329. Bilodeau JF, Blanchette S, Cormier N, Sirard M-A. Reactive oxygen species-mediated loss of bovine sperm motility in egg yolk Tris extender: protection by pyruvate, metal chelators and bovine liver or oviductal fluid catalase. Theriogenology 2002;57:1105–22. 330. Lapointe S, Sirard M-A. Catalase and oviductal fluid reverse the decreased motility of bovine sperm in culture medium containing specific amino acids. J Androl 1998;19:31–6. 331. Lapointe S, Sullivan R, Sirard M-A. Binding of a bovine oviductal fluid catalase to mammalian spermatozoa. Biol Reprod 1998;58:747–53. 332. Ball BA, Medina V, Gravance CG, Baumber J. Effect of antioxidants on preservation of motility, viability and acrosomal integrity of equine spermatozoa during storage at 5C. Theriogenology 2001;56:577–89. 333. Baumber J, Ball BA, Linfor JJ. Assessment of the cryopreservation of equine spermatozoa in the presence of enzyme scavengers and antioxidants. Am J Vet Res 2005;66:772–9. 334. Christova Y, James PS, Jones R. Lipid diffusion in sperm plasma membranes exposed to peroxidative injury from oxygen free radicals. Mol Reprod Dev 2004;68:365–72. 335. de Graaf SP, Evans G, Gillan L, Guerra MMP, Maxwell WMC, O’Brien JK. The influence of antioxidant, cholesterol and seminal plasma on the in vitro quality of sorted and non-sorted ram spermatozoa. Theriogenology 2007;67:217–27. 336. Krishnamoorthy G, Venkataraman P, Arunkumar A, Viqnesh RC, Aruldhas MM, Arunakaran J. Ameliorative effect of vitamins (alpha-tocopherol and ascorbic acid) on PCB (Aroclor 1254) induced oxidative stress in rat epididymal sperm. Reprod Toxicol 2006;23:239–45. 337. Yazama F, Furuta K, Fujimoto M, Sonoda T, Shigetomi H, Horiuchi T, Yamada M, Nagao N, Maeda N. Abnormal spermatogenesis in mice unable to synthesize ascorbic acid. Anat Sci Int 2006;81:115–25. 338. Hatamoto LK, Baptista Sobrinho CA, Nichi M, Barnabe VH, Barnabe RC, Cortada CNM. Effects of dexamethasone treatment (to mimic stress) and vitamin E oral supplementation on the spermiogram and on seminal plasma spontaneous lipid peroxidation and antiox-
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Stallion: Anatomy, Physiology and Endocrinology idant enzyme activities in dogs. Theriogenology 2006;66: 1610–14. Upreti GC, Jensen K, Oliver JE, Dugarizich DM, Munday R, Smith JF. Motility of ram spermatozoa during storage in a chemically-defined diluent containing antioxidants. Anim Reprod Sci 1997;48:269–78. Almeida J, Ball BA. Effect of alpha-tocopherol and tocopherol succinate on lipid peroxidation in equine spermatozoa. Anim Reprod Sci 2005;87:321–37. Hansen SH, Andersen MD, Birkedal H, Cornett C, Wibrand F. The important role of taurine in oxidative metabolism. Adv Exp Med Biol 2006;583:129–35. Alverez JG, Storey BT. Taurine, hypotaurine, epinephrine and albumin inhibit lipid peroxidation in rabbit spermatozoa and protect against loss of motility. Biol Reprod 1983;29:548–55. Twigg J, Fulton N, Gomez E, Irvine DS, Aitken RJ. Analysis of the impact of intracellular reactive oxygen species generation on the structural and functional integrity of human spermatozoa: lipid peroxidation, DNA fragmentation and effectiveness of antioxidants. Hum Reprod 1998;13:1429–36. Funahashi H, Sano T. Select antioxidants improve the function of extended boar semen stored at 10 degrees C. Theriogenology 2005;63:1605–16. Bilodeau JF, Blanchette S, Gagnon C, Sirard MA. Thiols prevent hydrogen peroxide-mediated loss of sperm motility in cryopreserved bull semen. Theriogenology 2001;56:275–86. Yang R, Le G, Li A, Zheng J, Shi Y. Effect of antioxidant capacity on blood lipid metabolism and lipoprotein lipase activity of rats fed a high-fat diet. Nutrition 2006;22:1185–91. Prahalathan C, Selvakumar E, Varalakshmi P. Modulatory role of lipoic acid on adriamycin-induced testicular injury. Chem Biol Interact 2006;160:108–14. Selvakumar E, Prahalathan C, Sudharsan PT, Varalakshmi P. Chemoprotective effect of lipoic acid against cyclophosphamide-induced changes in the rat sperm. Toxicology 2006;217:71–8. Bruemmer JE, Coy RC, Squires EL, Graham JK. Effect of pyruvate on the function of stallion spermatozoa stored for up to 48 hours. J Anim Sci 2002;80:12–18. Denniston DJ, Squires EL, Bruemmer JE, Brinsko SP, McCue PM, Graham JK. The effect of antioxidants on the motility and viability of cooled stallion spermatozoa. J Reprod Fertil 2000;56 (Suppl):121–6. Lopez-Burillo S, Tan DX, Mayo JC, Saijnz RM, Manchester LC, Reiter RJ. Melatonin, xanthurenic acid, resveratrol, EGCG, vitamin C and alpha-lipoic acid differentially reduce oxidative DNA damage induced by Fenton reagents: a study of their individual and synergistic actions. J Pineal Res 2003;34:269–77. Dokmeci D. Oxidative stress, male infertility and the role of carnitines. Folia Med (Plovdiv) 2005;47:26–30. Stradaioli G, Sylla L, Zelli R, Chiodi P, Monaci M. Effect of L-carnitine administration on the seminal characteristics of oligoasthenospermic stallions. Theriogenology 2004;62:761–77. Mann T. Biochemistry of stallion semen. J Reprod Fertil Suppl 1975;23:47–52. Upreti GC, Jensen K, Munday R, Dugarizich DM, Vishwanath R, Smith JF. Studies on aromatic amino acid oxidase activity in ram spermatozoa: role of pyruvate as an antioxidant. Anim Reprod Sci 1998;29:275–87.
356. Agarwal A, Sushil A, Prabakaran A, Said TM. Prevention of oxidative stress injury to sperm. J Androl 2005;26: 654–60. 357. Ford WCL, Whittington K. Antioxidant treatment for male subfertility: a promise that remains unfulfilled. Hum Reprod 1998;13:1416–19. 358. Donnelly ET, McClure N, Lewis SE. The effect of ascorbate and alpha-tocopherol supplementation in vitro on DNA integrity and hydrogen peroxide-induced DNA damage in human spermatozoa. Mutagenesis 1999; 14:505–12. 359. Milzer E, Schmidt HL. Carbon isotope effects on the decarboxylation of carboxylic acids: comparison of the lactate oxidase reaction and the degradation of pyruvate by hydrogen peroxide. Biochem J 1988;252:913–15. 360. Katila T, Sankari S, M¨akela O. Transport of spermatozoa in the reproductive tracts of mares. J Reprod Fertil Suppl 2000;56:571–8. 361. Sinnemma L, Jarvimaa T, Lehmonen N, Makela O, Reilas T, Sankari S, Katila T. Effect of insemination volume on uterine contractions and inflammatory response and on elimination of semen in the mare uterus- scintigraphic and ultrasonographic studies. J Vet Med 2005;52:466–71. 362. Campbell MLH, England GCW. Effects of coitus and the artificial insemination of different volumes of fresh semen on uterine contractions in mares. Vet Rec 2006; 159:843–9. 363. Fiala SM, Pimentel CA, Mattos ALG, Gregory RM, Mattos RC. Effect of sperm numbers and concentration on sperm transport and uterine inflammatory response in the mare. Theriogenology 2007;67:556–62. 364. Bader H. An investigation of sperm migration into the oviducts of the mare. J Reprod Fertil Suppl 1982;32:59–64. 365. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post insemination on pregnancy rate in mares. Theriogenology 1991;35:1111–17. 366. Rigby S, Derczo S, Brinsko SP, Blanchard TL, Taylor T, Forrest DW, Varner DD. Oviductal sperm numbers following proximal uterine horn or uterine body insemination. In: Proceedings of the 46th Annual Convention of the American Association of Equine Practitioners, 2000, pp. 332–4. 367. Scott MA, Liu IKM, Overstreet JW. Sperm transport to oviducts: abnormalities and their clinical implications. In: Proceedings of the 41st Annual Convention of the American Association of Equine Practitioners, 1995, pp. 1–2. 368. Parker WG, Sullivan JJ, First NL. Sperm transport and distribution in the mare. J Reprod Fertil Suppl 1975;23: 63–6. 369. Rigby S, Hill J, Miller C, Thompson J, Varner DD, Blanchard TL. Administration of oxytocin immediately after insemination does not improve pregnancy rates in mares bred by fertile or subfertile stallions. Theriogenology 1998;51:1143–50. 370. Haluska GJ, Currie WB. Variation in plasma concentrations of oestradiol-17 beta and their relationship to those of progesterone, 13,14-dihydro-15-keto-prostaglandin F2 alpha and oxytocin across pregnancy and at parturition in pony mares. J Reprod Fertil 1988;84:635–46. 371. Allen WE, Chard T, Forsling ML. Peripheral plasma levels of oxytocin and vasopressin in the mare during parturition. J Endocrinol 1973;57:175–6. 372. Langendijk P, Bouwman EG, Kidson A, Kirkwood RN, Soede NM, Kemp B. Role of myometrial activity in sperm
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transport through the genital tract and in fertilization in sows. Reproduction 2002;123:683–90. Stratman FW, Self HL, Smith VR. The effect of oxytocin on the fertility in gilts artificially inseminated with a low sperm concentration and semen volume. J Anim Sci 1959;18:634–40. Penrod L, Adams K, Arns MJ. The influence of exogenous prostaglandin F2-alpha on spermatozoa motility and fertility. Anim Reprod Sci 2006;94:36–8. Claus R, Schams D. Influence of mating and intra-uterine oestradiol infusion on peripheral oxytocin concentration in the sow. J Endocrinol 1990;126:361–5. Langendijk P, Bouwman EG, Soede NM, Taverne MA, Kemp B. Myometrial activity around estrus in sows: spontaneous activity and effects of estrogens, cloprostenol, seminal plasma and clenbuterol. Theriogenology 2002;57:1563–77. Langendijk P, Soede NM, Kemp B. Uterine activity, sperm transport, and the role of boar stimuli around insemination in sows. Theriogenology 2005;63:500–13. Stecco R, Paccamonti D, Gutjahr S, Pinto C, Eilts B. Day of cycle affects changes in equine intrauterine pressure in response to teasing. Theriogenology 2003;60:727–33. Nikolakopoulos E, Kindahl H, Gilbert CL, Goode J, Watson ED. Release of oxytocin and prostaglandin F2 alpha around teasing, natural service and associated events in the mare. Anim Reprod Sci 2000;63:89–99. Madill S, Troedsson MHT, Alexander SL, Shand N, Santschi EM, Irvine CHG. Simultaneous recording of pituitary oxytocin secretion and myometrial activity in oestrous mares exposed to carious breeding stimuli. J Reprod Fertil Suppl 2000;56:351–61. Alexander SL, Irvine CHG, Shand N, Evans MJ. Is luteinizing hormone secretion modulated by endogenous oxytocin in the mare? Studies on the role of oxytocin and factors affecting its secretion in estrous mares. Biol Reprod Monogr 1995;1:361–71. Taverne MA, Van der Weyden GC, Rontijne P, Dieleman SJ, Pashen RL, Allen WR. In vivo myometrial electrical activity in the cyclic mare. J Reprod Fertil 1979;56:521–32. Campbell MLH, England GCW. M-mode ultrasound imaging of the contractions of the equine uterus. Vet Rec 2002;150:575–7. Mann T, Polge C, Rowson LEA. Participation of seminal plasma during the passage of spermatozoa in the female reproductive tract of the pig and horse. J Endocrinol 1956;13:133–40. Claus R, Hoang-Vu C, Ellendorf F, Meyer HD. Seminal oestrogens in the boar: original and functions in the sow. J Steroid Biochem 1987;27:331–5. Willenburg KL, Miller GM, Rodriguez-Zas SL, Knox RV. Influence of hormone supplementation to extended semen on artificial insemination, uterine contractions, establishment of a sperm reservoir, and fertility in swine. J Anim Sci 2003;81:821–9. Troedsson MHT, Liu IKM, Crabo BG. Sperm transport and survival in the mare: a review. Theriogenology 1998; 50:807–18. Troedsson MHT, Desvousges A, Alghamdi AS, Dahms B, Dow CA, Hayna J, Valesco R, Collahan PT, Macpherson ML, Pozor M, Buhi WC. Components in seminal plasma regulating sperm transport and elimination. Anim Reprod Sci 2005;89:171–86. Troedsson MHT, Alghamdi AS, Mattisen J. Equine seminal plasma protects the fertility of spermatozoa in an
390.
391.
392.
393.
394.
395.
396.
397.
398.
399.
400.
401.
402.
403.
404.
405. 406.
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inflamed uterine environment. Theriogenology 2002;58: 453–6. Clement F, Vincent P, Mahla R, Meriaux JC, Palmer E. Which insemination results in fertilization when several are performed before ovulation? J Reprod Fertil Suppl 2000;56:579–85. Metcalf ES. The effect of postinsemination endometritis on fertility of frozen stallion semen. In: Proceedings of the 46th Annual Convention of the American Association of Equine Practitioners, 2000, pp. 330–1. Scott MA. A glimpse at sperm function in vivo: sperm transport and epithelial interaction in the female reproductive tract. Anim Reprod Sci 2000; 60–61:337–48. Gaddum-Rosse P. Some observations on sperm transport through the uterotubal junction of the rat. Am J Anat 1981;160:333–41. Scott MA, Liu IKM, Overstreet JW, Enders AC. The structural morphology and epithelial association of spermatozoa at the uterotubal junction: a descriptive study of equine spermatozoa in situ using scanning electron microscopy. J Reprod Fertil Suppl 2000;56:415–21. Nilsson O, Reinius S. Light and electron microscope structure of the oviduct. In: Hafez ESE, Blandau RJ (eds) The Mammalian Oviduct: Comparative Biology and Methodology. Chicago: University of Chicago Press, 1969; pp. 57–83. Cho C, Bunch DO, Faure JE, Goulding EH, Eddy EM, Primakoff P, Myles DG. Fertilization defects in sperm from mice lacking fertilin b. Science 1998;281:1857–9. Hagaman JR, Moyer JS, Bachman ES, Sibony M, Magyar PL, Welch JE, Smithies O, Krege JH, O’Brien DA. Angiotensin-converting enzyme and male fertility. Proc Natl Acad Sci USA 1998;95:2552–7. Hunter RHF, Flechon B, Flechon JE. Distribution, morphology and epithelial interactions of bovine spermatozoa in the oviduct before and after ovulation: a scanning electron microscope study. Tissue Cell 1991;23:641–56. Larsson B. Distribution of spermatozoa in the genital tract of heifers inseminated with large numbers of abnormal spermatozoa. J Vet Med Ser A 1988;35:721–8. Waberski D, Magnus F, Ardon F, Petrunkina AM, Weitze KF, Topfer-Petersen E. Binding of boar spermatozoa to oviductal epithelium in vitro in relation to sperm morphology and storage time. Reproduction 2006;131:311–18. Thomas PGA, Ball BA, Miller PG, Brinsko SP, Southwood L. A subpopulation of morphologically normal, motile spermatozoa attach to equine oviductal epithelial cell monolayers. Biol Reprod 1994;51:303–9. Thomas PGA, Ball BA, Brinsko SP. Interaction of equine spermatozoa with oviduct epithelial cell explants is affected by estrous cycle and anatomic origin of explant. Biol Reprod 1994;51:222–8. Scott MA, Varner DD, Liu IKM, Enders AC. Presumptive evidence of a preovulatory sperm reservoir in the mare: morphological investigations using scanning electron microscopy. Theriogenology 2002;58:639–42. Wood J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic-loss rate in mares. Equine Vet J 1990;22:410–15. Bain AM. Thoroughbred mare in Australasia. J Am Vet Med Assoc 1957;131:179–85. Dobrinski I, Ignotz GG, Thomas PGA, Ball BA. Role of carbohydrates in the attachment of equine spermatozoa to uterine tubal (oviductal) epithelial cells in vitro. Am J Vet Res 1996;57:1635–9.
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407. Ekhasi-Hundrieser M, Gohr K, Wagner A, Tsolova M, Petrunkina AM, Topfer-Petersen E. Spermadhesin AQN1 is a candidate receptor molecule involved in the formation of the oviductal sperm reservoir in pig. Biol Reprod 2005;73:536–45. 408. Gwathmey TYM, Ignotz GG, Suarez SS. PDC-109 (BSPA1/A2) promotes bull sperm binding to oviductal sperm release. Biol Reprod 2003;69:809–15. 409. Marini PE, Cabada MO. One step purification and biochemical characterization of a spermatozoa-binding protein from porcine oviductal epithelial cells. Mol Reprod Dev 2003;66:383–90. 410. Perez FA, Roma SM, Cabada MO, Marini PE. Sperm binding glycoprotein is differentially present surrounding the lumen of isthmus and ampulla of the pig’s oviduct. Anat Embryol (Berl) 2006;211:619–24. 411. Ignotz GG, Lo MC, Perez CL, Gwathmey TM, Suarez SS. Characterization of a fucose-binding protein from bull sperm and seminal plasma that may be responsible for formation of the oviductal sperm reservoir. Biol Reprod 2006;64:1806–11. 412. Gwathmey TYM, Ignotz GG, Mueller JL, Manjunath P, Suarez SS. Bovine seminal plasma proteins PDC-109, BSP-A3, and BSP-30-kDa share functional roles in storing sperm in the oviduct. Biol Reprod 2006;75:501–7. 413. Sabeur K, Ball BA. Characterization of galactose-binding proteins in equine testis and spermatozoa. Anim Reprod Sci 2007;101:74–84. 414. Ellington JE, Ignotz GG, Miller PG, Currie WB, Meyers SA. Oviduct epithelial cell co-culture modifies stallion and bull sperm cell surface proteins. Biol Reprod 1993;48(suppl 1):107. 415. Ellington JE, Ball BA, Blue BJ, Wilker CE. Capacitationlike membrane changes and prolonged viability in vitro of equine spermatozoa cultured with uterine tube epithelial cells. Am J Vet Res 1993;54:1505–10. 416. Ellington JE, Ball BA, Yang X. Binding of stallion spermatozoa to the equine zona pellucida after coculture with oviductal epithelial cells. J Reprod Fertil 1993;98:203–8. 417. Smith TT, Yanagimachi R. Attachment and release of spermatozoa from the caudal isthmus of the hamster oviduct. J Reprod Fertil 1991;91:567–73. 418. Pollard JW, Plante C, King WA, Hansen PJ, Betteridge KJ, Suarez SS. Fertilizing capacity of bovine sperm may be maintained by binding of oviductal epithelial cells. Biol Reprod 1991;44:102–7. 419. Dobrinski I, Smith TT, Suarez SS, Ball BA. Membrane contact with oviductal epithelium modulates the intracellular calcium concentration of equine spermatozoa in vitro. Biol Reprod 1997;56:861–9. 420. Smith TT, Nothnick WB. Role of direct contact between spermatozoa and oviductal epithelial cells in maintaining rabbit sperm viability. Biol Reprod 1997;56:83–9. 421. Thomas PG, Ignotz GG, Ball BA, Brinsko SP, Currie WB. Effect of coculture with stallion spermatozoa on de novo protein synthesis and secretion by equine oviduct epithelial cells. Am J Vet Res 1995;56:1657–62. 422. Killian GJ. Evidence for the role of oviduct secretions in sperm function, fertilization and embryo development. Anim Reprod Sci 2004; 82–83:82–3. 423. Ellington JE, Ignotz GG, Ball BA, Meyers-Wallen VN, Currie WB. De novo protein synthesis by bovine uterine tube (oviduct) epithelial cells changes during coculture with bull spermatozoa. Biol Reprod 1993;48: 851–6.
424. Fazeli A, Elliott RMA, Duncan AE, Moore A, Watson PF, Holt WV. In vitro maintenance of boar sperm viability by a soluble fraction obtained from oviductal apical plasma membrane preparations. Reproduction 2003;125: 509–17. 425. Ellington JE, Ignotz GG, Varner DD, Marcucio RS, Mathison P, Ball BA. In vitro interaction between oviduct epithelia and equine sperm. Arch Androl 1993;31:79–86. 426. Pacey AA, Davies N, Warren MA, Barratt CL, Cooke ID. Hyperactivation may assist human spermatozoa to detach from intimate association with the endosalpinx. Hum Reprod 1995;10:2603–9. 427. Topfer-Petersen E. Molecules on the sperm route to fertilization. J Exp Zool 1999;285:259–66. 428. Austin CR. Observations on the penetration of the sperm into the mammalian egg. Aust J Biol Sci 1951;4:581–96. 429. Chang MC. Fertilizing capacity of spermatozoa deposited into fallopian tubes. Nature 1951;168:697–8. 430. Austin CR. Capacitation and the release of hyaluronidase from spermatozoa. J Reprod Fertil 1960;3:310–11. 431. Yanagimachi R. Mammalian fertilization. In: Knobil E, Neill JD (eds) The Physiology of Reproduction. New York: Raven Press, 1994; pp. 189–317. 432. Talbot P. Sperm penetration through oocyte investments in mammals. Am J Anat 1985;174:331–46. 433. Corselli J, Talbot P. In vitro penetration of hamster oocyte-cumulus complexes using physiological numbers of sperm. Dev Biol 1987;122:227–42. 434. Cardullo RA, Thaler CD. Function of the egg’s extracellular matrix. In: Hardy DH (ed.) Fertilization. San Diego: Academic Press, 2002; pp. 119–52. 435. Visconti PE, Bailey JL, Moore GD, Pan D, Olds-Clarke P, Kopf GS. Capacitation of mouse spermatozoa. I. Correlation between the capacitation state and protein tyrosine phosphorylation. Development 1995;121:1129–37. 436. Yanagimachi R. Sperm capacitation and gamete interaction. J Reprod Fertil 1989;38:27–33. 437. Austin CR. Capacitation of spermatozoa. Int J Fertil 1967;12:25–31. 438. Bedford JM. Sperm capacitation and fertilization in mammals. Biol Reprod Suppl 1970;2:128–58. 439. Yanagimachi R. Mechanisms of fertilization in mammals. In: Mastroianni L, Biggers JD (eds) Fertilization and Embryonic Development In Vitro. New York: Plenum Press, 1981; pp. 81–182. 440. Yanagimachi R. In vitro capacitation of hamster spermatozoa by follicular fluid. J Reprod Fertil 1969;18:275–86. 441. Suarez SS, Ho HC. Hyperactivation of mammalian sperm. Cell Mol Biol 2003;49:351–6. 442. Mortimer ST, Maxwell WM. Kinematic definition of ram sperm hyperactivation. Reprod Fertil Dev 1999;11:25–30. 443. Chang MC. The meaning of sperm capacitation: a historical perspective. J Androl 1984;5:45–50. 444. Kopf GS. Signal transduction mechanisms regulating sperm acrosomal exocytosis. In: Hardy DM (ed.) Fertilization. San Diego: Academic Press, 2002; pp. 181–223. 445. DeMott RP, Suarez SS. Hyperactivated sperm progress in the mouse oviduct. Biol Reprod 1992;46:779–85. 446. Stauss CR, Suarez SS. Sperm motility hyperactivation facilitates penetration of the hamster zona pellucida. Biol Reprod 1995;53:1280–5. 447. Suarez SS, Dai X. Intracellular calcium reaches different levels of elevation in hyperactivated and acrosome reacted hamster sperm. Mol Reprod Dev 1995;42:325–33.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 448. Suarez SS, Katz DF, Owen DH, Andrew JB, Powell RL. Evidence for the function of hyperactivated motility in sperm. Biol Reprod 1991;44:375–81. 449. Robertson L, Wolf DP, Tash JS. Temporal changes in motility parameters related to acrosomal status: identification and characterization of populations of hyperactivated human sperm. Biol Reprod 1988;39:797–805. 450. Mortimer ST, Swan MA, Mortimer D. Effect of seminal plasma on capacitation and hyperactivation in human spermatozoa. Hum Reprod 1998;13:2139–46. 451. Breitbart H. Signaling pathways in sperm capacitation and acrosome reaction. Cell Mol Biol 2003;49:321–7. 452. Nagdas SK, Winfrey VP, Olson GE. Tyrosine phosphorylation generates multiple isoforms of the mitochondrial capsule protein, phospholipid hydroperoxide glutathione peroxidase (PHGPx), during hamster sperm capacitation. Biol Reprod 2004;72:164–71. 453. Green S, Fishel S. Morphology comparison of individually selected hyperactivated and non-hyperactivated human spermatozoa. Hum Reprod 1999;14:123–30. 454. Harrison RA, Ashworth PJ, Miller NG. Bicarbonate/CO2, an effector of capacitation, induces a rapid and reversible change in the lipid architecture of boar sperm plasma membranes. Mol Reprod Dev 1996;45:378–91. 455. Rath R, Colenbrander B, Bevers MM, Gadella BM. Evaluation on in vitro capacitation of stallion spermatozoa. Biol Reprod 2001;65:462–70. 456. Meyers SA, Rosenberger AE. A plasma membraneassociated hyaluronidase is localized to the posterior acrosomal region of stallion sperm and is associated with spermatozoal function. Biol Reprod 1999;61:444–51. 457. Jimenez I, Gonzalez-Marquez H, Ortiz R, Herrera JA, Garcia A, Betancourt M, Fierro R. Changes in the distribution of lectin receptors during capacitation and acrosome reaction in boar spermatozoa. Theriogenology 2003; 59:1171–80. 458. Cross NL. Decrease in order of human sperm lipids during capacitation. Biol Reprod 2003;69:529–34. 459. Smith TT, McKinnon-Thompson CA, Wolf DE. Changes in lipid diffusibility in the hamster sperm head plasma membrane during capacitation in vivo and in vitro. Mol Reprod Dev 1998;50:86–92. 460. Benoff S. Carbohydrates and fertilization: an overview. Mol Hum Reprod 1997;3:599–637. 461. Cross NL, Overstreet JW. Glycoconjugates of the human sperm surface: distribution and alterations that accompany capacitation in vitro. Gamete Res 1987;16:23–35. 462. Zeng Y, Clark EN, Florman HM. Sperm membrane potential: hyperpolarization during capacitation regulates zona pellucida-dependent acrosomal secretion. Dev Biol 1995;171:554–63. 463. Visconti PE, Moore GD, Bailey JL, Leclerc P, Connors SA, Pan D, Olds-Clark P, Kopf GS. Capacitation of mouse spermatozoa. II. Protein tyrosine phosphorylation and capacitation are regulated by a cAMP-dependent pathway. Development 1995;121:1139–50. 464. Piehler E, Petrunkina AM, Ekhlasi-Hundrieser M, Topfer-Petersen E. Dynamic quantification of the tyrosine phosphorylation of the sperm surface proteins during capacitation. Cytometry A 2006;69:1062–70. 465. Linares-Hernandez L, Guzman-Grenfell AM, HicksGomez JJ, Gonzalez-Martinez MT. Voltage-dependent calcium influx in human sperm assessed by simultaneous optical detection of intracellular calcium and membrane potential. Biochim Biophys Acta 1998;1372:1–12.
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466. Bray C, Son JH, Meizel S. Acetylcholine causes an increase of intracellular calcium in human sperm. Mol Hum Reprod 2005;11:881–9. 467. Fraire-Zamora JJ, Gonzalez-Martinez MT. Effect of intracellular pH on depolarization-evoked calcium influx in human sperm. Am J Physiol Cell Physiol 2004;287:1688–96. 468. Neri-Vidaurri PD, Torres-Flores V, Gonzalez-Martinez MT. A remarkable increase in the pHi sensitivity of voltage-dependent calcium channels occurs in human sperm incubated in capacitating conditions. Biochem Biophys Res Commun 2006;343:105–9. 469. Garcia MA, Meizel S. Regulation of intracellular pH in capacitated human spermatozoa by a Na+ /H+ exchanger. Mol Reprod Dev 1999;52:189–95. 470. Harrison RA, Mairet B, Miller NG. Flow cytometric studies of bicarbonate-mediated Ca2+ influx in boar sperm populations. Mol Reprod Dev 1993;35:197–208. 471. Magistrini M, Guitton E, Levern Y, Nicolle JC, Vidament M, Kerboeuf D, Palmer E. New staining methods for sperm evaluation estimated by microscopy and flow cytometry. Theriogenology 1997;48:1229–35. 472. Gadella BM, Harrison RAP. Capacitation induces cyclic adenosine 3’,5’-monophosphate-dependent, but apoptosis-unrelated, exposure of aminophospholipids at the apical head plasma membrane of boar sperm cells. Biol Reprod 2002;67:340–50. 473. Colenbrander B, Brouwers JFHM, Neild DM, Stout TAE, daSilva P, Gadella BM. Capacitation dependent lipid rearrangements in the plasma membrane of equine sperm. Theriogenology 2002;58:341–5. 474. Flesch FM, Brouwers JFHM, Nievelstein PFEM, Verkleij AJ, vanGolde LMG, Colenbrander B, Gadella BM. Bicarbonate stimulated phospholipid scrambling induces cholesterol redistribution and enables cholesterol depletion in the sperm plasma membrane. J Cell Sci 2001;114:3543–55. 475. Flesch FM, Colenbrander B, vanGolde LMG, Gadella BM. Capacitation induces tyrosine phosphorylation of proteins in the boar sperm plasma membrane. Biochem Biophys Res Commun 1999;262:787–92. 476. Gadella BM, Lopes-Cardozo M, vanGolde LMG, Colenbrander B, Gadella Jr TWJ. Glycolipid migration from the apical to the equatorial subdomains of the sperm head plasma membrane precedes the acrosome reaction. Evidence for a primary capacitation event in boar spermatozoa. J Cell Sci 1995;108:935–46. 477. Pommer AC, Rutlant J, Meyers SA. Phosphorylation of protein tyrosine residues in fresh and cryopreserved stallion spermatozoa under capacitating conditions. Biol Reprod 2003;68:1208–14. 478. Jaiswal BS, Eisenbach M. Capacitation. In: Hardy DM (ed.) Fertilization. San Diego:, Academic Press, 2002; pp. 57–117. 479. Florman HM, Ducibella T. Fertilization in mammals. In: Neill JD (ed.) Knobel and Neill’s Physiology of Reproduction. Boston: Elsevier, 2006; pp. 55–112. 480. Felix R. Molecular physiology and pathology of Ca2+ conducting channels in the plasma membrane of mammalian sperm. Reproduction 2005;129:251–62. 481. Gadella BM, Rathi R, Brouwers JFHM, Stout TAE, Colenbrander B. Capacitation and the acrosome reaction in equine sperm. Anim Reprod Sci 2001;68:249–65. 482. Thundathil JC, Anzar M, Buhr MM. Na+ /K+ ATPase as a signaling molecule during bovine sperm capacitation. Biol Reprod 2006;75:308–17.
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483. Langlais J, Roberts KD. A molecular membrane model of sperm capacitation and the acrosome reaction of mammalian spermatozoa. Gamete Res 1985;12: 183–224. 484. Aitken RJ. Molecular mechanisms regulating human sperm function. Mol Hum Reprod 1997;3:169–73. 485. O’Flaherty C, deLamirande E, Gagnon C. Positive role of reactive oxygen species in mammalian sperm capacitation: triggering and modulation of phosphorylation events. Free Radic Biol Med 2006;41:528–40. 486. deLamirande E, Leclerc P, Gagnon C. Capacitation as a regulatory event that primes spermatozoa for the acrosome reaction and fertilization. Mol Hum Reprod 1997;3: 175–94. 487. Darszon A, Acevedo JJ, Galindo BE, HernandezGonzalez EO, Nishigaki T, Trevino CL, Wood C, Beltran C. Sperm channel diversity and functional multiplicity. Reproduction 2006;131:977–88. 488. Jha KN, Kameshwari DB, Shivaji S. Role of signaling pathways in regulating the capacitation of mammalian spermatozoa. Cell Mol Biol 2003;49:329–40. 489. Visconti PE, Kopf GS. Regulation of protein phosphorylation during sperm capacitation. Biol Reprod 1998;59:1–6. 490. Mitra K, Shivaji S. Proteins implicated in sperm capacitation. Indian J Exp Biol 2005;43:1001–15. 491. Chang MC. A detrimental effect of seminal plasma on the fertilizing capacity of sperm. Nature 1957;175:258–9. 492. Fraser LR. Interactions between a decapacitation factor and mouse spermatozoa appear to involve fucose residues and GPI-anchored receptor. Mol Reprod Dev 1998;51:193–202. 493. Lopes CH, Mazzini MN, Tortorella H, Konrath RA, Brandelli A. Isolation, partial characterization and biological activity of mannosyl glycopeptides from seminal plasma. Glycoconj J 1998;15:477–81. 494. Villemure M, Lazure C, Manjunath P. Isolation and characterization of gelatin-binding proteins from goat seminal plasma. Reprod Biol Endocrinol 2003;1:39. 495. Parks JE, Hough SR. Platelet-activating factor acetylhydrolase activity in bovine seminal plasma. J Androl 1993;14:335–9. 496. Huang YH, Chen YH, Lin CM, Ciou YY, Kuo SP, Chen CT, Shih CMS, Chang EE. Suppression effect of seminal vesicle autoantigen on platelet-activating factor-induced mouse sperm capacitation. J Cell Biochem 2007;100:941–51. 497. Therien I, Moreau R, Manjunath P. Major proteins of bovine seminal plasma and high-density lipoprotein induce cholesterol efflux from epididymal sperm. Biol Reprod 1998;59:768–76. 498. Lusignan M-F, Bergeron A, Crete M-H, Lazure C, Manjunath P. Induction of epididymal boar sperm capacitation by pB1 and BSP-A1/-A2 proteins, members of the BSP protein family. Biol Reprod 2006;76:424–32. 499. Topfer-Petersen E, Ekhasi-Hundrieser M, Kirchhoff C, Leeb T, Sieme H. The role of stallion seminal proteins in fertilisation. Anim Reprod Sci 2005;89:159–70. 500. Nolan MA, Wu L, Bang HJ, Jelinsky SA, Roberts KP, Turner TT, Kopf GS, Johnston DS. Identification of rat cysteine-rich secretory protein 4 (Crisp4) as the ortholog to human CRISP1 and mouse CRISP4. Biol Reprod 2006; 74:984–91. 501. Jalkane J, Huhtaniemi I, Poutanen M. Mouse cysteinerich secretory protein 4 (CRISP4): a member of the CRISP family exclusively expressed in the epididymis
502.
503.
504.
505.
506.
507.
508. 509.
510.
511.
512.
513.
514.
515.
516.
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518.
in an androgen-dependent manner. Biol Reprod 2005;72: 1268–74. Udby L, Bjartell A, Malm J, Egesten A, Lundwall A, Cowland JB, Borregaard N, Kjeldsen L. Characterization and localization of cysteine-rich secretory protein 3 (CRISP-3) in the human male reproductive tract. J Androl 2005;26:333. Schambony A, Hess O, Gentzel M, Topfer-Petersen E. Expression of CRISP proteins in the male equine genital tract. J Reprod Fertil Suppl 1998;53:67–72. Evans JP. The molecular basis of sperm-oocyte membrane interactions during mammalian fertilization. Hum Reprod Update 2002;8:297–311. Roberts KP, Wamstad JA, Ensrud KM, Hamilton DW. Inhibition of capacitation-associated tyrosine phosphorylation signaling in rat sperm by epididymal protein CRISP1. Biol Reprod 2003;69:572–81. Boatman EE, Robbins RS. Bicarbonate: carbon-dioxide regulation of sperm capacitation, hyperactivated motility, and acrosome reactions. Biol Reprod 1991;44:806–13. Cross NL. Effect of cholesterol and other sterols on human sperm acrosomal responsiveness. Mol Reprod Dev 1996;45:212–17. Cross NL. Role of cholesterol in sperm capacitation. Biol Reprod 1998;59:7–11. DasGupta S, Mills EL, Fraser LR. Ca2+ -related changes in the capacitation state of human spermatozoa assessed by a chlortetracycline fluorescence assay. J Reprod Fertil 1993;99:135–43. Davis BK, Byrne R, Hungund B. Studies on the mechanism of capacitation. II. Evidence for lipid transfer between plasma membrane of rate sperm and serum albumin during capacitation in vitro. Biochem Biophys Acta 1979;558:257–66. Davis BK, Byrne R, Bedigian K. Studies on the mechanism of capacitation: albumin-mediated changes in plasma membrane lipids during in vitro incubation of rat sperm cells. Proc Natl Acad Sci USA 1980;77: 1546–50. Davis BK. Timing of fertilization in mammals: Sperm cholesterol/phospholipid ratio as a determinant of the capacitation interval. Proc Natl Acad Sci USA 1981;78: 7560–4. Gadella BM, Harrison RAP. The capacitating agent bicarbonate induces protein kinase A-dependent changes in phospholipid transbilayer behavior in the sperm plasma membrane. Development 2000;127:2407–20. Go KJ, Wolf DP. Albumin-mediated changes in sperm sterol content during capacitation. Biol Reprod 1985; 32:145–53. Harrison RAP. Capacitation mechanisms, and the role of capacitation as seen in eutherian mammals. Reprod Fertil Dev 1996;8:581–94. Lee MA, Storey BT. Bicarbonate is essential for fertilization of mouse eggs: mouse sperm require it to undergo the acrosome reaction. Biol Reprod 1986;34:349–56. Okamura N, Tajima Y, Soejima A, Masuda H, Sugita Y. Sodium bicarbonate in seminal plasma stimulates the motility of mammalian spermatozoa through the direct activation of adenylate cyclase. J Biol Chem 1985;260: 9699–705. Shi Q-X, Roldan ERS. Bicarbonate/CO2 is not required for zona pellucida- or progesterone-induced acrosomal exocytosis of mouse spermatozoa but is essential for capacitation. Biol Reprod 1995;52:540–6.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 519. Suzuke F, Yanagimachi R. Changes in the distribution of intramembranous particles and filipin-reactive membrane sterols during in vitro capacitation of golden hamster spermatozoa. Gamete Res 1989;23:335–47. 520. Visconti PE, Galantino-Homer H, Ning X, Moore GD, Valenzuela JP, Jorgez CJ, Alvarez JG, Kopf GS. Cholesterol efflux-mediated signal transduction in mammalian sperm. Beta-cyclodextrins initiate transmembrane signaling leading to an increase in protein tyrosine phosphorylation and capacitation. J Biol Chem 1999;274:3235–42. 521. Visconti PE, Stewart-Savage J, Blasco A, Battaglia L, Miranda P, Kopf GS, Tezon JG. Roles of bicarbonate, cAMP, and protein tyrosine phosphorylation on capacitation and the spontaneous acrosome reaction of hamster sperm. Biol Reprod 1999;61:76–84. 522. Visconti PE, Ning X, Fornes MW, Alvarez JG, Stein P, Connors SA, Kopf GS. Cholesterol efflux-mediated signal transduction in mammalian sperm: cholesterol release signals an increase in protein tyrosine phosphorylation during mouse sperm capacitation. Dev Biol 1999;214: 429–43. 523. Visconti PE, Westbrook VA, Chertihin O, Demarco I, Sleight S, Diekman AB. Novel signaling pathways involved in sperm acquisition of fertilizing capacity. J Reprod Immunol 2002;53:133–50. 524. Choi YH, Toyoda Y. Cyclodextrin removes cholesterol from mouse sperm and induces capacitation in a protein free medium. Biol Reprod 1998;59:1328–33. 525. Cross NL. Effect of methyl-beta-cyclodextrin on the acrosomal responsiveness of human sperm. Mol Reprod Dev 1999;53:92–8. 526. Osherroff JE, Visconti PE, Valenzuela JP, Travis AJ, Alverez J, Kopf GS. Regulation of human sperm capacitation by a cholesterol efflux-stimulated signal transduction pathway leading to protein kinase A-mediated upregulation of protein tyrosine phosphorylation. Mol Hum Reprod 1999;5:1017–26. 527. Nolan JP, Hammerstedt RH. Regulation of membrane stability and the acrosome reaction in mammalian sperm. FASEB J 1997;11:670–82. 528. Shinitzky M. Membrane fluidity and cellular functions. In: Shinitzky M (ed.) Physiology of Membrane Fluidity. Boca Raton, FL: CRC Press, 1984; pp. 1–51. 529. Jimenez-Gonzalez C, Michelangeli F, Harper CV, Barratt CLR, Publicover SJ. Calcium signalling in human spermatozoa: a specialized ‘toolkit’ of channels, transporters and stores. Hum Reprod Update 2006;12:253–67. 530. Spira B, Breitbart H. The role of anion channels in the mechanism of acrosome reaction in bull spermatozoa. Biochim Biophys Acta 1992;1109:65–73. 531. Darszon A, Lopez-Martinez P, Acevedo JJ, HernandezCruz A, Trevino CL. T-type Ca2+ channels in sperm function. Cell Calcium 2006;40:241–52. 532. Neri-Vidaurri Pdel C, Torres-Flores V, GonzalezMartinez MT. A remarkable increase in the pHi sensitivity of voltage-dependent calcium channels occurs in human sperm incubated in capacitating conditions. Biochem Biophys Res Commun 2006;343:105–9. 533. Wennermuth G, Westenbroek RE, Xu T, Hille B, Babcock DF. Cav2.2 and Cav2.3 (N- and R-type) Ca2+ channels in depolarization-evoked entry of Ca2+ into mouse sperm. J Biol Chem 2000;275:21210–17. 534. Wennermuth G, Babcock DF, Hille B. Calcium clearance mechanisms of mouse sperm. J Gen Physiol 2003;122: 115–28.
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535. Breitbart H, Webbie R, Lardy H. Regulation of calcium transport in bovine spermatozoa. Biochim Biophys Acta 1990;1027:72–8. 536. Zarca A, Rubinstein D, Breitbart H. Transport mechanism for calcium and phosphate in ram spermatozoa. Biochim Biophys Acta 1988;994:351–8. 537. DasGupta S, Mills CL, Fraser LR. A possible role for Ca2+ -ATPase in human sperm capacitation. J Reprod Fertil 1994;102:107–16. 538. Roldan ERS, Flemming AD. Is a Ca2+ -ATPase involved in Ca2+ regulation during capacitation and the acrosome reaction of guinea-pig spermatozoa. J Reprod Fertil 1989;85:297–308. 539. Fraser LR, Abeydeera LR, Niwa K. Ca2+ -regulating mechanisms that modulate bull sperm capacitation and acrosomal exocytosis as determined by chlortetracycline analysis. Mol Reprod Dev 1995;40:233–41. 540. Quill TA, Sugden SA, Rossi KL, Doolittle LK, Hammer RE, Garbers DL. Hyperactivated sperm motility driven by CatSper2 is required for fertilization. Proc Natl Acad Sci USA 2003;100:14869–74. 541. Nikpoor P, Mowla SJ, Movahedin M, Ziaee SZ, Tiraihi T. CatSper gene expression in postnatal development of mouse testis and in subfertile men with deficient sperm motility. Hum Reprod 2004;19:124–8. 542. Quill TA, Ren D, Clapham DE, Garbers KL. A voltagegated ion channel expressed specifically in spermatozoa. Proc Natl Acad Sci USA 2001;98:12527–31. 543. Ren D, Navarro B, Perez G, Jackson AC, Hsu S, Shi Q, Tilly JL, Clapham CE. A sperm ion channel required for sperm motility and male fertility. Nature 2001;413: 603–9. 544. Fraser LR, Adeoya-Osiguwa S, Baxendale RW, Mededovic S, Osiguwa OO. First messenger regulation of mammalian sperm function via adenylyl cyclase/cAMP. J Reprod Dev 2005;51:37–46. 545. Breitbart H, Cohen G, Rubinstein S. Role of actin cytoskeleton in mammalian sperm capacitation and the acrosome reaction. Reproduction 2005;129:263–8. 546. Clarke GN, Clarke FM, Wilson S. Actin in human spermatozoa. Biol Reprod 1982;26:319–27. 547. Ochs D, Wolf DP. Actin in ejaculated human sperm cells. Biol Reprod 1985;33:1223–26. 548. Virtanen I, Badley RA, Paasivuo R, Lehto VP. Distinct cytoskeletal domains revealed in sperm cells. J Cell Biol 1984;99:1083–91. 549. Breitbart H, Rubinstein S, Etkovitz N. Sperm capacitation is regulated by the crosstalk between protein kinase A and C. Mol Cell Endocrinol 2006;252:247–9. 550. Cohen G, Rubinstein S, Gur Y, Breitbart H. Crosstalk between protein kinase a and c regulates phospholipase d and f-actin formation during sperm capacitation. Dev Biol 2004;267:230–41. 551. Cross NL, Overstreet JW. Glycoconjugates of the human sperm surface; distribution and alterations that accompany capacitation in vitro. Gamete Res 1987;16:23–35. 552. Ficarro S, Chertihin O, Westbrook VA, White F, Jayes F, Kalab P, Marto JA, Shabanowitz J, Herr JC, Hunt DF, Visconti PE. Phosphoproteome analysis of capacitated human sperm. Evidence of tyrosine phosphorylation of a kinase-anchoring protein 3 and valosincontaining protein/p97 during capacitation. J Biol Chem 2003;278:11579–89. 553. Urner F, Sakkas D. Protein phosphorylation in mammalian spermatozoa. Reproduction 2003;125:17–26.
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Stallion: Anatomy, Physiology and Endocrinology
554. Bailey JL. Cell signalling during capacitation and the acrosome reaction. In: Proceedings of the Havemeyer Foundation Workshop on Stallion Reproduction, Bandera, Texas, 2007. 555. Vijayaraghavan S, Liberty GA, Mohan J, Winfrey VP, Olson GE, Carr DW. Isolation and molecular characterization of AKAP110, a novel, sperm-specific protein kinase A-anchoring protein. Mol Endocrinol 1999;13:705–17. 556. Naaby-Hansen S, Mandal A, Wolkowicz MJ, Sen B, Westbrook VA, Shetty J, Coonrod SA, Klotz KL, Kim YH, Bush LA, Flickinger CJ, Herr JC. CABYR, a novel calciumbinding tyrosine phosphorylation- regulation of tyrosine phosphorylation in human spermatozoa and its role in the control of human sperm function. Dev Biol 2002; 242:236–54. 557. Aisquith KL, Baleato RM, McLaughlin EA, Nixon B, Aitken RJ. Tyrosine phosphorylation activates surface chaperones facilitating sperm-zona recognition. J Cell Sci 2004;117:3657. 558. Brewis IA, Moore HD, Fraser LR, Holt WV, Baldi E, Luconi M, Gadella BM, Ford WC, Harrison RAP. Molecular mechanisms during sperm capacitation. Hum Fertil 2005;8:253–61. 559. Harrison RAP, Miller NG. cAMP-dependent protein kinase control of plasma membrane lipid architecture in boar sperm. Mol Reprod Dev 2000;55:220–8. 560. Harrison RA, Gadella BM. Bicarbonate-induced membrane processing in sperm capacitation. Theriogenology 2005;63:342–51. 561. Wang L, Beserra C, Garbers DL. A novel aminophospholipid transporter exclusively expressed in spermatozoa is required for membrane lipid asymmetry and normal fertilization. Dev Biol 2004;267:203–15. 562. deLamirande E, Gagnon C. A positive role for the superoxide anion in triggering hyperactivation and capacitation of human spermatozoa. Int J Androl 1993;16: 21–5. 563. Bize I, Santander G, Cabello P, Driscoll D, Sharpe C. Hydrogen peroxide is involved in hamster sperm capacitation in vitro. Biol Reprod 1991;44:398–403. 564. Kameshwari DB, Siva AB, Shivaji S. Inhibition of in vitro capacitation of hamster spermatozoa by nitric oxide synthase inhibitors. Cell Mol Biol 2003;49:421–8. 565. Herrero MB, de Lamirande E, Gagnon C. Nitric oxide regulates human sperm capacitation and protein tyrosine phosphorylation in vitro. Biol Reprod 1999;61:575–81. 566. O’Flaherty C, deLamirande E, Gagnon C. Reactive oxygen species modulate independent protein phosphorylation pathways during human sperm capacitation. Free Radic Biol Med 2006;40:1045–55. 567. Aitken RJ, Paterso M, Fisher H, Buckingham DW, Duin M. Redox regulation of tyrosine phosphorylation in human spermatozoa and its role in the control of human sperm function. J Cell Sci 1995;108:2017–25. 568. Aitken RJ, Fisher HM, Fulton N, Gomez E, Knox W, Lewis B, Irvine S. Reactive oxygen species generation by human spermatozoa is induced by exogenous NADPH and inhibited by the flavoprotein inhibitors diphenylene iodonium and quinacrine. Mol Reprod Dev 1997; 47:482. 569. Aitken RJ, Harkiss D, Knox W, Paterson M, Irvine DS. A novel signal transduction cascade in capacitating human spermatozoa characterised by a redoxregulated cAMPmediated induction of tyrosine phosphorylation. J Cell Sci 1998;111:645–56.
570. Natarajan V, Forman HJ. Introduction to serial reviews on redox regulation of phospholipases and sphingomyelinase in cell signaling. Free Radic Biol Med 2006;40: 363. 571. Thundathil J, de Lamirande E, Gagnon C. Nitric oxide regulates the phosphorylation of the threonineglutamine-tyrosine motif in proteins of human spermatozoa during capacitation. Biol Reprod 2003;68: 1291–8. 572. Rodriguez PC, O’Flaherty CM, Beconi MT, Beorlegui NB. Nitric oxide-induced capacitation of cryopreserved bull spermatozoa and assessment of participating regulatory pathways. Anim Reprod Sci 2005;85:231–42. 573. O’Flaherty C, de Lamirande E, Gagnon C. Reactive oxygen species and protein kinases modulate the level of phospho-MEK-like proteins during human sperm capacitation. Biol Reprod 2005;73:94–105. 574. Baumber J, Sabeur K, Vo A, Ball BA. Reactive oxygen species promote capacitation and tyrosine phosphorylation in equine sperm. Anim Reprod Sci 2001;68: 337–8. 575. Baker MA, Atiken RJ. The importance of redox regulated pathways in sperm cell biology. Mol Cell Endocrinol 2004;216:47–54. 576. Baumber J, Sabeur K, Vo A, Ball BA. Reactive oxygen species promote tyrosine phosphorylation and capacitation in equine spermatozoa. Theriogenology 2003;60: 1239–47. 577. Lewis B, Aitken RJ. A redox-regulated tyrosine phosphorylation cascade in rat spermatozoa. J Androl 2001; 22:611–22. 578. Aitken RJ, Harkiss D, Knox W, Paterson M, Irvine DS. A novel signal transduction cascade in capacitating human spermatozoa characterized by a redox-regulated, cAMPmediated induction of tyrosine phosphorylation. J Cell Sci 1998;111:645–56. 579. Cross NL. Reorganization of lipid rafts during capacitation of human sperm. Biol Reprod 2004;71:1367–73. 580. vanGestel RA, Brewis IA, Ashton PR, Helms JB, Brouwers JFHM, Gadella BM. Capacitation-dependent concentration of lipid rafts in the apical ridge head area of porcine sperm cells. Mol Hum Reprod 2005;11:583–90. 581. Weerachatyanukul W, Probodh I, Kongmanas K, Tanphaichitr N, Johnston LJ. Visualizing the localization of sulfoglycolipids in lipid raft domains in model membranes and sperm membrane extracts. Biochim Biophys Acta 2006;1768:299–310. 582. Khalil MB, Chakrabandhu K, Xu H, Weerachatyanukul W, Buhr MM, Berger T, Carmona E, Vuong N, Kumarathasan P, Wong PTT, Carrier D, Tanphaichitr N. Sperm capacitation induces an increase in lipid rafts having zona pellucida binding ability and containing sulfogalactosylglycerolipid. Dev Biol 2006;290:220–35. 583. Thaler CD, Thomas M, Ramalie JR. Reorganization of mouse sperm lipid rafts by capacitation. Mol Reprod Dev 2006;73:1541–9. 584. Sleight SB, Miranda PV, Plaskett NW, Maier B, Lysiak J, Scrable H, Herr JC, Visconti PE. Isolation and proteomic analysis of mouse sperm detergent-resistant membrane fractions: evidence for dissociation of lipid rafts during capacitation. Biol Reprod 2005;73:721–9. 585. Tanphaichitr N, Carmona E, Bou Khalil M, Xu H, Berger T, Gerton GL. New insights into sperm-zona pellucida interaction: involvement of sperm lipid rafts. Front Biosci 2007;12:1748–66.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 586. deLamirande E, Gagnon C. The extracellular signalregulated kinase (ERK) pathway is involved in human sperm function and modulated by the superoxide anion. Mol Hum Reprod 2002;8:124–35. 587. Wasco WM, Orr GA. Function of calmodulin in mammalian sperm: presence of a calmodulin-dependent cyclic nucleotide phosphodiesterase associated with demembranated rat caudal epididymal sperm. Biochem Biophys Res Commun 1984;118:636–42. 588. Luconi M, Krausz C, Forti G, Baldi E. Extracellular calcium negatively modulates tyrosine phosphorylation and tyrosine kinase activity during capacitation of human spermatozoa. Biol Reprod 1996;55:207–16. 589. Ashraf M, Peterson RN, Russell LD. Evidence for Ca2+ /Na+ antiport in boar sperm plasma membrane vesicles. Biol Reprod 1982;26:37A. 590. Bradley MP, Forrester IT. A sodium-calcium exchange mechanism in plasma membrane vesicles isolated from ram sperm flagella. FEBS Lett 1980;121:15–18. 591. Rufo BA, Schoff PK, Lardy HA. Regulation of calcium content in bovine spermatozoa. J Biol Chem 1984;259: 2547–52. 592. Acevedo JJ, Mendoza-Lujambio I, Vega-Beltran JL, Felix R, Darszon A. KATP channels in mouse spermatogenic cells and sperm, and their role in capacitation. Dev Biol 2006;289:395–405. 593. Ball BA, Gravance CG, Wessel MT, Sabeur K. Activity of angiotensin-converting enzyme (ACE) in reproductive tissues of the stallion and effects of angiotensin II on sperm motility. Theriogenology 2003;59:901–14. 594. Sabeur K, Vo A, Ball BA. Affects of angiotensin II on the acrosome reaction in equine spermatozoa. J Reprod Fertil 2000;120:135–42. 595. Fraser LR, Adeoya-Osiquwa SA, Baxendale RW, Gibbons R. Regulation of mammalian sperm capacitation by endogenous molecules. Front Biosci 2006;11:1636–45. 596. Mededovic S, Fraser LR. Mechanisms of action of angiotensin II on mammalian sperm function. Reproduction 2005;129:211–18. 597. Hutt DM, Cardullo RA, Baltz JM, Ngsee JK. Synaptotagmin VIII is localized to the mouse sperm head and may function in acrosomal exocytosis. Biol Reprod 2002;66:50–6. 598. Olds-Clark P. Unresolved issues in mammalian fertilization. Int Rev Cytol 2003;232:129–84. 599. Miller DJ, Shur BD. Molecular basis of fertilization in the mouse. Dev Biol 1994;5:255–64. 600. Snell WJ, White JM. The molecules of mammalian fertilization. Cell 1996;85:629–37. 601. Wassarman PM. Mammalian fertilization: molecular aspects of gamete adhesion, exocytosis and fusion. Cell 1999;96:175–83. 602. Kopf GS, Ning XP, Visconti PE, Purdon M, GalantinoHomer H, Fornes M. Signaling mechanisms controlling mammalian sperm fertilization competence and activation. In: Gagnon C (ed.) The Male Gamete: From Basic Science to Clinical Applications. Vienna, IL: Cache River Press, 1999; pp. 105–18. 603. Florman HM, Storey BT. Mouse gamete interactions: the zona pellucida is the site of the acrosome reaction leading to fertilization in vitro. Dev Biol 1982;91:121–30. 604. Roldan ERS, Murase T, Qi-Xian S. Exocytosis in spermatozoa in response to progesterone and zona pellucida. Science 1994;266:1578–81. 605. Wassarman PM. Contribution of mouse egg zona pellu-
606.
607. 608. 609.
610.
611.
612.
613.
614. 615.
616.
617.
618.
619.
620.
621.
622.
623.
624.
979
cida glycoproteins to gamete recognition during fertilization. J Cell Physiol 2005;204:388–91. Wassarman P, Chen J, Cohen N, Litscher E, Liu C, Qi H, Williams Z. Structure and function of the mammalian egg zona pellucida. J Exp Zool 1999;285:251–8. Wassarman PM. Sperm receptors and fertilization in mammals. Mt Sinai J Med 2002;69:148–55. Wasserman PM, Jovine L, Litscher E. A profile of fertilization in mammals. Nat Cell Biol 2001;3:E59–64. Bleil JD, Wassarman PM. Structure and function of the zona pellucida: identification and characterization of the proteins of the mouse oocyte’s zona pellucida. Dev Biol 1980;76:185–202. Bleil JD, Wassarman PM. Mammalian sperm-egg interaction: identification of a glycoprotein in mouse egg zonae pellucidae possessing receptor activity for sperm. Cell 1980;20:873–82. Bleil JD, Wassarman PM. Sperm-egg interactions in the mouse: sequence of events and induction of the acrosome reaction by a zona pellucida glycoprotein. Dev Biol 1983;95:317–24. Caballero-Campo P, Chirinos M, Fan XJ, GonzalezGonzalez ME, Galicia-Chavarria M, Larrea F, Gerton GL. Biological effects of recombinant human zona pellucida proteins on sperm function. Biol Reprod 2006;74:760–8. Conner SJ, Lefievre L, Hughes DC, Barratt CL. Cracking the egg: increased complexity in the zona pellucida. Hum Reprod 2005;20:1148–52. Wassarman PM. Fertilization in mammals. Sci Am 1988;259:78–84. Florman HM, Wassarman P. O-linked oligosaccharides of mouse egg ZP3 account for its sperm receptor activity. Cell 1985;41:313–24. Epifano O, Liang LF, Familiari M, Moos ML, Dean J. Coordinate expression of the three zona pellucida genes during mouse oogenesis. Development 1995;121:1947–56. McLeskey SB, Dowds C, Carballada R, White RR, Saling PM. Molecules involved in mammalian sperm-egg interaction. Int Rev Cytol 1998;177:57–113. Florman HM, Bechtol KB, Wassarman PM. Enzymatic digestion of the functions of the mouse egg’s receptor for sperm. Dev Biol 1984;106:243–55. Sinowatz F, Topfer-Petersen E, Kolle S, Palma G. Functional morphology of the zona pellucida. Anat Histol Embryol 2001;30:257–63. Bleil JD, Wassarman PM. Galactose at the nonreducing terminus of O-linked oligosaccharides of mouse egg zona pellucida glycoprotein ZP3 is essential for the glycoprotein’s sperm receptor activity. Proc Natl Acad Sci USA 1988;85:6778–82. Miller DJ, Macek MB, Shur BD. Complementarity between sperm surface beta-1,4-galactosyltransferase and egg-coat ZP3 mediates sperm-egg binding. Nature 1992;357:589–93. Shi Q-X, Amindari S, Paruchuru K, Skalla D, Burkink H, Shur BD, Miller DJ. Cell surface beta-1,4galactosyltransferase-I activates G protein-dependent exocytotic signaling. Development 2001;128:645–54. Rodeheffer C, Shur BD. Sperm from B1,4galactosyltransferase I-null mice exhibit precocious capacitation. Development 2004;131:491–501. Lopez LC, Bayna EM, Litoff D, Shaper NL, Shaper JH, Shur BD. Receptor function of mouse sperm surface galactosyltransferase during fertilization. J Cell Biol 1985;101:1501–10.
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625. Scully NF, Shaper JH, Shur BD. Spatial and temporal expression of cell surface galactosyltransferase during mouse spermatogenesis and epididymal maturation. Dev Biol 1987;124:111–24. 626. Shur BD, Hall NG. A role for mouse sperm surface galactosyltransferase in sperm binding to the egg zona pellucida. J Cell Biol 1982;95:574–9. 627. Shur BD, Neely CA. Plasma membrane association, purification, and partial characterization of mouse sperm Beta 1,4-galactosyltransferase. J Biol Chem 1988;263: 17706–14. 628. Rodeheffer C, Shur BD. Characterization of a novel ZP3independent sperm-binding ligand that facilitates sperm adhesion to the egg coat. Development 2004;131:503–12. 629. Jovine L, Darie CC, Litscher E, Wassarman PM. Zona pellucida domain proteins. Annu Rev Biochem 2005;74: 83–114. 630. Wassarman PM. Fertilization. Mol Biol 2003;16:117–204. 631. Greve JM, Wassarman PM. Mouse egg extracellular coat is a matrix of interconnected filaments possessing a structural repeat. J Mol Biol 1985;181:253–64. 632. Hinsch E, Aires VA, Hedrich F, Oehninger S, Hinsch KD. A synthetic decapeptide from a conserved ZP3 protein domain induces the G protein-regulated acrosome reaction in bovine spermatozoa. Theriogenology 2005;63:1682–94. 633. Berridge MJ, Lipp P, Bootman MD. The versatility and universality of calcium signaling. Nat Rev Mol Cell Biol 2000;1:11–21. 634. Breitbart H, Spungin B. The biochemistry of the acrosome reaction. Mol Hum Reprod 1997;3:195–202. 635. Jungnickel MK, Marrero H, Birnbaumer L, Lemos JR, Florman HM. Trp2 regulates entry of Ca2+ into mouse sperm triggered by egg ZP3. Nat Cell Biol 2001;3: 499–502. 636. O’Toole CM, Arnoult C, Darszon A, Steinhardt RA, Florman HM. Ca2+ entry through store-operated channels in mouse sperm is initiated by egg ZP3 and drives the acrosome reaction. Mol Biol Cell 2000;11:1571–84. 637. Gonzalez-Martinez MT, Galindo BE, De La Torre L, Zapata O, Rodriguez E, Florman HM, Darszon A. A sustained increase in intracellular Ca2+ is required for the acrosome reaction in sea urchin sperm. Dev Biol 2001; 236:220–9. 638. Hirohashi N, Vacquier D. Store-operated calcium channels trigger exocytosis of the sea urchin sperm acrosomal vesicle. Biochem Biophys Res Commun 2003;304:285–92. 639. Hartneck C, Plant TD, Schultz G. From worm to man: three subfamilies of TRP. Trends Neurosci 2000; 159–66. 640. Castellano LE, Trevino CL, Rodriguez D, Serrano CJ, Pacheco J, Tsutsumi V, Felix R, Darszon A. Transient receptor potential (TRPC) channels in human sperm: expression, cellular localization and involvement in the regulation of flagellar motility. FEBS Lett 2003;541:69–74. 641. Gong X, Dubois DH, Miller DJ, Shur BD. Activation of a G protein complex by aggregation of beta-1,4galactosyltransferase on the surface of sperm. Science 1995;269:1718–21. 642. Bailey JL, Storey BT. Calcium influx into mouse spermatozoa activated by solubilized mouse zona pellucida, monitored with the calcium fluorescent indicator, fluo-3. Inhibition of the influx by three inhibitors of the zona pellucida induced acrosome reaction: tyrphostin A48, pertussis toxin, and 3-quinuclidinyl benzilate. Mol Reprod Dev 1994;39:297–308.
643. Lee MA, Check JH, Kopf GS. A guanine nucleotidebinding regulatory protein in human sperm mediates acrosomal exocytosis induced by the human zona pellucida. Mol Reprod Dev 1992;31:78–86. 644. Arnoult C, Kazam IG, Visconti PE, Kopf GS, Villaz M, Florman HM. Control of the low voltage-activated calcium channel of mouse sperm by egg ZP3 and by membrane hyperpolarization during capacitation. Proc Natl Acad Sci USA 1999;96:6757–62. 645. Leclerc P, Kopf GS. Mouse sperm adenylyl cyclase: general properties and regulation by the zona pellucida. Biol Reprod 1995;52:1227–33. 646. Noland TD, Garbers DL, Kopf GS. An elevation in cyclic AMP concentration precedes the zona pellucida-induced acrosome reaction of mouse spermatozoa. Biol Reprod 1998;38 (Suppl. 1):138. 647. Garde J, Roldan ER. Rab 3-peptide stimulates exocytosis of the ram sperm acrosome via interaction with cyclic AMP and phospholipase A2 metabolites. FEBS Lett 1996;39:263–8. 648. De Jonge CJ, Han J-L, Lawrie H, Mack SR, Zaneveld LJD. Modulation of human sperm acrosome reaction by effectors of the adenylate cyclase/cAMP second messenger pathway. J Exp Zool 1991;258:113–25. 649. Spungin B, Breitbart H. Calcium mobilization and influx during sperm exocytosis. J Cell Sci 1996;109:1947–55. 650. Breitbart H, Naor Z. Protein kinases in mammalian sperm capacitation and the acrosome reaction. Rev Reprod 1999;4:151–9. 651. Bielfeld P, Faridi A, Zaneveld LJD, DeJonge CJ. The zona pellucida-induced acrosome reaction of human spermatozoa is mediated by protein kinases. Fertil Steril 1994;61:536–54. 652. Tomes CN, McMaster CR, Saling PM. Activation of mouse sperm phosphatidylinositol-4,5 bisphosphatephospholipase C by zona pellucida is modulated by tyrosine phosphorylation. Mol Reprod Dev 1996;43: 196–204. 653. Roldan ER, Fragio C. Phospholipase A2 activity and exocytosis of the ram sperm acrosome: regulation by bivalent cations. Biochem Biophys Acta 1993;1168:108–14. 654. Roldan ER. Role of phospholipases during sperm acrosomal exocytosis. Front Biosci 1998;3:1109–19. 655. Walensky LD, Snyder SH. Inositol 1,4,5-triphosphate receptors selectively localized to the acrosomes of mammalian sperm. J Cell Biol 1995;130:857–69. 656. Spungin B, Margalit I, Breitbart H. Sperm exocytosis reconstructed in a cell-free system: evidence for the involvement of phospholipase C and actin filaments in membrane fusion. J Cell Sci 1995;108:2525–35. 657. Fukami K, Nakao K, Inoue T, Kataoka Y, Kurokawa M, Fissore RA, Nakamura K, Katsuki M, Mikoshiba K, Yoshida N, Takenawa T. Requirement of phospholipase Cdelta4 for the zona pellucida-induced acrosome reaction. Science 2001;292:920–3. 658. Fukami K, Yoshida M, Inoue T, Kurokawa M, Fissore RA, Yoshida N, Mikoshiba K, Takenawa T. Phospholipase C-4 is required for Ca2+ mobilization essential for acrosome reaction in sperm. J Cell Biol 2003;161:79–88. 659. Putney JWJ. Capacitative calcium entry revisited. Cell Calcium 1990;11:611–24. 660. Hikida M, Kurosaki T. Regulation of phospholipase C-γ 2 networks in B lymphocytes. Adv Immunol 2005;88:73–96. 661. Montell CL, Birnbaumer L, Flockerzi V. The TRP channels, a remarkably functional family. Cell 2002;108:595–8.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 662. Kiselyov K, Mignery GA, Zhu MX, Muallem S. The Nterminal domain of the IP3 receptor gates store-operated hTrp3 channels. Mol Cell 1999;4:423–9. 663. Boulay G, Brown DM, Quin N, Jiang M, Dietrich A, Zhu MX, Chen Z, Birnbaumer M, Mikoshiba K, Birnbaumer L. Modulation of Ca2+ entry by polypeptides of the inositol 1,4,5-triphosphate receptor (IP3 R) that bind transient receptor potential (TRP): evidence for roles of TRP and IP3 R in store depletion-activated Ca2+ entry. Proc Natl Acad Sci USA 1999;96:14955–60. 664. Xu XZ, Sternberg PW. A C. elegans sperm TRP protein required for sperm-egg interactions during fertilization. Cell 2003;114:285–97. 665. O’Toole CM, Roldan ER, Fraser LR. Protein kinase C activation during progesterone-stimulated acrosomal exocytosis in human spermatozoa. Mol Hum Reprod 1996;2:921–7. 666. Breitbart H, Lax J, Rotem R, Naor Z. Role of protein kinase C in the acrosome reaction of mammalian spermatozoa. Biochem J 1992;281:473–6. 667. Rotem R, Paz GF, Homonnai ZT. Further studies on the involvement of protein kinase C in human flagellar motility. Endocrinology 1990;127:2571–7. 668. Rotem R, Paz GF, Homonnai ZT, Kalina M, Lax Y, Breitbart H, Naor Z. Protein kinase C is present in human sperm: possible role in flageller motility. Proc Natl Acad Sci USA 1990;87:7305–8. 669. Naor Z, Breitbart H. Protein kinase C and mammalian spermatozoa acrosome reaction. Trends Endocrinol Metab 1997;8:337–42. 670. Luk AS, Kaler EW, Lee SP. Phospholipase C-induced aggregation and fusion of cholesterol-lecithin small unilamellar vesicles. Biochemistry 1993;32:6965–73. 671. Siegel DP, Banschbach J, Alford D, Ellens H, Lis L-J, Quinn PJ, Yeagle PL, Bentz J. Physiological levels of diacylglycerols in phospholipid membranes induce membrane fusion and stabilize inverted phases. Biochemistry 1989;28:3703–9. 672. Roggero CM, Tomes C, deBlas GA, Castillo J, Michaut MA, Fukuda M, Mayorga LS. Protein kinase C-mediated phosphorylation of the two polybasic regions of synaptotagmin VI regulates their function in acrosomal exocytosis. Dev Biol 2005;285:422–35. 673. Tucker WC, Chapman ER. Role of synaptotagmin in Ca2+ -triggered exocytosis. Biochem J 2002;366:1–13. 674. Sudhof TC. Synaptotagmins: why so many? J Biol Chem 2002;277:7629–32. 675. Michaut M, deBlas GA, Tomes CN, Yunes R, Fukuda M, Mayorga LS. Synaptotagmin VI participates in the acrosome reaction of human spermatozoa. Dev Biol 2001;235:521–9. 676. Ono K, Yanagimachi R, Huang TTF. Phospholipase A of guinea pig spermatozoa: its preliminary characterization and possible involvement in the acrosome reaction. Dev Growth Diff 1982;24:305–10. 677. Lui CW, Meizel S. Further evidence in support of a role for hamster sperm hydrolytic enzymes in the acrosome reaction. J Exp Zool 1979;207:173–86. 678. Singleton CL, Killian GJ. A study of phospholipase in albumin and its role in inducing the acrosome reaction of guinea pig spermatozoa in vitro. J Androl 1983;4:150–6. 679. Kyono K, Hoshi K, Saito A, Tsuiki A, Hoshiai H, Suzuki M. Effects of phospholipase A, lysophosphatidylcholine, and fatty acid on the acrosome reaction of human spermatozoa. Tohoku J Exp Med 1984;144:257–63.
981
680. Roldan ER, Vazquez JM. Bicarbonate/CO2 induces rapid activation of phospholipase A2 and renders boar spermatozoa capable of undergoing acrosomal exocytosis in response to progesterone. FEBS Lett 1996;396:227–32. 681. Roldan ER, Fragio C. Phospholipase A2 activation and subsequent exocytosis in the Ca2+ /ionophore-induced acrosome reaction of ram spermatozoa. J Biol Chem 1993;268:13962–70. 682. Yuan YY, Che WY, Shi QX, Mao LZ, Yu SQ, Fang X, Roldan ER. Zona pellucida induces activation of phospholipase A2 during acrosomal exocytosis in guinea pig spermatozoa. Biol Reprod 2003;68:904–13. 683. Roldan ER, Fragio C. Diradylglycerols stimulate phospholipase A2 and subsequent exocytosis in ram spermatozoa. Biochem J 1994;297:225–32. 684. Flemming AD, Yanagimachi R. Effects of various lipids on the acrosome reaction and fertilizing capacity of guinea pig spermatozoa with special reference to the possible involvement of lysophospholipids in the acrosome reaction. Gamete Res 1981;4:253–73. 685. Ohzu E, Yanagimachi R. Acceleration of acrosome reaction in hamster spermatozoa by lysolecithin. J Exp Zool 1982;224:259–63. 686. Llanos MN, Morales P, Riffo MS. Studies of lysophospholipids related to the hamster sperm acrosome reaction. J Exp Zool 1993;267:209–16. 687. Parrish JJ, Susko-Parish JL, Winer MA, First NL. Capacitation of bovine sperm by heparin. Biol Reprod 1988;38: 1171–80. 688. Garde J, Roldan ER. Stimulation of Ca2+ -dependent exocytosis of the sperm acrosome by cAMP acting downstream of phospholipase A2. J Reprod Fertil 2000;118: 57–68. 689. Sollner TH. Regulated exocytosis and SNARE function (Review). Mol Membr Biol 2003;20:209–20. 690. Ramalho-Santos J, Moreno RD, Sutovsky P, Chan AW, Hewitson L, Wessel GM, Simerly CR, Schatten G. SNAREs in mammalian sperm: possible implications for fertilization. Dev Biol 2000;223:54–69. 691. deBlas GA, Roggero CM, Tomes C, Mayorga LS. Dynamics of SNARE assembly and disassembly during sperm acrosomal exocytosis. PLoS Biol 2005;3:1801–12. 692. Tomes CN, deBlas GA, Michaut MA, Farre EV, Cherhitin O, Visconti PE, Mayorga LS. Alpha-SNAP and NSF are required in a priming step during the human sperm acrosome reaction. Mol Hum Reprod 2004;11:43–51. 693. Katafuchi K, Mori T, Toshimori K, Iida H. Localization of a syntaxin isoform, syntaxin 2, to the acrosomal region of rodent spermatozoa. Mol Reprod Dev 2000;57:375– 83. 694. Gamboa S, Ramalho-Santos J. SNARE proteins and caveolin-1 in stallion spermatozoa: possible implications for fertility. Theriogenology 2005;64:275–91. 695. McNew JA, Weber T, Parlati F, Johnson RJ, Melia TJ, Sollner TH, Rothman JE. Close is not enough: SNAREdependent membrane fusion requires an active mechanism that transduces force to membrane anchors. J Cell Biol 2000;150:105–17. 696. Grote E, Baba M, Ohsumi Y, Novick PJ. Geranylgeranylated SNAREs are dominant inhibitors of membrane fusion. J Cell Biol 2000;151:453–66. 697. He P, Southard C, Chen D, Whiteheart SW, Copper RL. Role of alpha-SNAP in promoting efficient neurotransmission at the crayfish neuromuscular junction. J Neurophysiol 1999;82:3406–16.
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698. Barnard RJ, Morgan A, Burgoyne RD. Stimulation of NSF ATPase activity by alpha-SNAP is required for SNARE complex disassembly and exocytosis. J Cell Biol 1997;139:875–83. 699. Clary DO, Griff IC, Rothman JE. SNAPs, a family of NSF attachment proteins involved in intracellular membrane fusion in animal and yeast. Cell 1990;61:709–21. 700. Nagiec EE, Bernstein A, Whiteheart SW. Each domain of the N-ethylmaleimide-sensitive fusion protein contributes to its transport activity. J Biol Chem 1995;270: 29182–8. 701. Weidman PJ, Melanncon P, Block MR, Rothman JE. Binding of an N-ethylmaleimide-sensitive fusion protein to Golgi membranes requires both a soluble protein(s) and an integral membrane receptor. J Cell Biol 1989; 108:1589–96. 702. Banerjee A, Barry VA, DasGupta BR, Martin TFJ. NEthylmaleimide-sensitive factor acts at a prefusion ATPdependent step in Ca2+ -activated exocytosis. J Biol Chem 1996;271:20223–6. 703. Belmonte SA, Lopez CI, Roggero CM, deBlas GA, Tomes CN, Mayorga LS. Cholesterol content regulates acrosomal exocytosis by enhancing Rab3A plasma membrane association. Dev Biol 2005;285:393–408. 704. Pfeffer SR. Transport-vesicle targeting: tethers before SNAREs. Nat Cell Biol 2001;1:E17–22. 705. Waters MG, Hughson FM. Membrane tethering and fusion in the secretory and indocytic pathways. Traffic 2000; 1:588–97. 706. Branham MT, Mayorga LS, Tomes CN. Calcium-induced acrosomal exocytosis requires cAMP acting through a protein kinase A-independent, Epac-mediated pathway. J Biol Chem 2006;281:8656–66. 707. Littleton JT, Bai J, Vyas B, Desai R, Baltus AE, Garment MB, Carlson SD, Ganetzky B, Chapman ER. Synaptotagmin mutants reveal essential functions for the C2B domain in Ca2+ -triggered fusion and recycling of SVs in vivo. J Neurosci 2001;21:1421–33. 708. Augustine GJ, Neher E. Neuronal Ca2+ signaling takes the local route. Curr Opin Neurobiol 1992;2:302–7. 709. Catterall WA. Interactions of presynaptic Ca2+ channels and snare proteins in neurotransmitter release. Ann N Y Acad Sci 1999;868:144–59. 710. Leabu M. Membrane fusion in cells: molecular machinery and mechanisms. J Cell Mol Med 2006;10:423–7. 711. Jeremic A, Kelly M, Cho JA, Cho SJ, Horber JK, Jena BP. Calcium drives fusion of SNARE-apposed bilayers. Cell Biol Int 2004;28:19–31. 712. Portis A, Newton C, Pangborn W, Papahadjopoulos D. Studies on the mechanism of membrane fusion: evidence for an intermembrane Ca2+ -phospholipid complex, synergism with Mg2+ , and inhibition by spectrin. Biochemistry 1979;18:780–90. 713. Suarez SS, Dai X. Hyperactivation enhances mouse sperm capacity for penetrating viscoelastic media. Biol Reprod 1992;46:686–91. 714. Eisenbach M, Giojalas LC. Sperm guidance in mammals: an unpaved road to the egg. Nat Rev Mol Cell Biol 2006;7:276–85. 715. Suarez SS, Vincenti L, Ceglia MW. Hyperactivated motility induced in mouse sperm by calcium ionophore A23187 is reversible. J Exp Zool 1987;244:331–6. 716. Qi H, Moran MM, Navarro B, Chong JA, Krapivinsky G, Krapivinsky L, Kirichok Y, Ramsey IS, Quill TA, Clapham DE. All four CatSper ion channel proteins are
717.
718.
719.
720.
721.
722.
723.
724.
725. 726.
727. 728.
729.
730.
731.
732.
733. 734.
required for male fertility and sperm cell hyperactivated motility. Proc Natl Acad Sci 2007;104:1219–23. Lindemann CB, Goltz JS. Calcium regulation of flagellar curvature and swimming pattern in triton X-100: extracted rat sperm. Cell Motil Cytoskeleton 1988;10:420– 31. Marquez B, Suarez SS. Bovine sperm hyperactivation is promoted by alkaline-stimulated Ca2+ influx. Biol Reprod 2007;76:660–5. Ho HC, Suarez SS. An inositol 1,4,5-triphosphate receptor-gated intracellular Ca2+ store is involved in regulating sperm hyperactivated motility. Biol Reprod 2001; 65:1606–15. Santi CM, Santos T, Hernandez-Cruz A, Darszon A. Properties of a novel pH-dependent Ca2+ permeation pathway present in male germ cells with possible roles in spermatogenesis and mature sperm function. J Gen Physiol 1998;112:33–53. Marquez B, Ignotz GG, Suarez SS. Contributions of extracellular and intracellular Ca2+ to regulation of sperm motility: release of intracellular stores can hyperactivate CatSper1 and CatSper2 null sperm. Dev Biol 2007; 303:214–21. Ho HC, Suarez SS. Characterization of the intracellular calcium store at the base of the sperm flagellum that regulates hyperactivated motility. Biol Reprod 2003;68: 1590–696. Zhang Z. The effect of 12-myristate 13-acetate phorbol ester on human sperm hyperactivation. Fertil Steril 1997;67:1140–5. Burkman LJ. Discrimination between nonhyperactivated and classical hyperactivated motility patterns in human spermatozoa using computerized analysis. Fertil Steril 1991;55:363–71. Rogers BJ. The sperm penetration assay: Its usefulness reevaluated. Fertil Steril 1985;43:821–40. Wramsby H, Liedholm P. Importance of the use of short and long sperm pre-incubation periods in assessing the fertilizing capacity of human spermatozoa by hamster test. Human Reprod 1986;1:171–3. Muller CH. Rationale, interpretation, validation and use of sperm function tests. J Androl 2000;21:10–30. DeMott RP, Lefebvre R, Suarez SS. Carbohydrates mediate the adherence of hamster sperm to oviductal epithelium. Biol Reprod 1995;52:1395–403. Eisenbach M, Ralt D. Precontact mammalian sperm-egg communication and role in fertilization. Am J Physiol 1992;262:C1095–101. Cohen-Dayag A, Tur-Kaspa I, Dor J, Mashiach S, Eisenbach M. Sperm capacitation in humans is transient and correlates with chemotactic responsiveness to follicular factors. Proc Natl Acad Sci USA 1995;92:11039–43. Thomas MB, Haines SL, Akeson RA. Chemoreceptors expressed in taste, olfactory and male reproductive tissues. Gene 1996;178:1–5. Fabro G, Rovasio RA, Civalero S, Frenkel A, Caplan SR, Eisenbach M, Giojalas LC. Chemotaxis of capacitated rabbit spermatozoa to follicular fluid revealed by a novel directionality-based assay. Biol Reprod 2002;67: 1565–71. Kirkman-Brown JC, Sutton KA, Florman HM. How to attract a sperm. Nat Cell Biol 2003;5:93–6. Riffell JA, Krug PJ, Zimmer RK. The ecological and evolutionary consequences of sperm chemoattraction. Proc Natl Acad Sci USA 2004;101:4501–6.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 735. Eisenbach M. Towards understanding the molecular mechanism of sperm chemotaxis. J Gen Physiol 2004;124: 105–8. 736. Ralt D, Manor M, Cohen-Dayag A, Tur-Kaspa I, BenShlomo I, Makler A, Yuli I, Dor J, Blumberg S, Mashiach S, Eisenbach M. Chemotaxis and chemokinesis of human spermatozoa to follicular factors. Biol Reprod 1994;50:774–85. 737. Cohen-Dayag A, Ralt D, Tur-Kaspa I, Manor M, Makler A, Dor J, Mashiach S, Eisenbach M. Sequential acquisition of chemotactic responsiveness by human spermatozoa. Biol Reprod 1994;50:786–90. 738. Hunter RHF, Petersen HH, Greve T. Ovarian follicular fluid, progesterone and Ca2+ ion influences on sperm release from the fallopian tube reservoir. Mol Reprod Dev 1999;54:283–91. 739. Villanueva-Diaz C, Arias-Martinez J, Bermejo-Martinez L, Vadillo-Ortega F. Progesterone induces human sperm chemotaxis. Fertil Steril 1995;64:1183–8. 740. Sliwa L. Chemotaction of mouse spermatozoa induced by certain hormones. Arch Androl 1995;35:105–10. 741. Eisenbach M, Tur-Kaspa I. Do human eggs attract spermatozoa? Bioessays 1999;21:203–10. 742. Feldmesser E, Olender T, Khen M, Yanai I, Ophir R, Lancet D. Widespread ectopic expression of olfactory receptor genes. BMC Genomics 2006;7:121. 743. Vanderhaeghen P, Schurmans S, Vassart G, Parmentier M. Specific repertoire of olfactory receptor genes in the male germ cells of several mammalian species. Genomics 1997;39:239–46. 744. Spehr M, Schwane K, Riffell JA, Zimmer RK, Hatt H. Odorant receptors and olfactory-like signaling mechanisms in mammalian sperm. Mol Cell Endocrinol 2006; 250:128–36. 745. Fukuda N, Touhara K. Developmental expression patterns of testicular olfactory receptor genes during mouse spermatogenesis. Genes Cells 2006;11:71–81. 746. Walensky LD, Ruat M, Bakin RE, Blackshaw S, Ronnett GV, Snyder SH. Two novel odorant receptor families expressed in spermatids undergo 5’-splicing. J Biol Chem 1998;273:9378–87. 747. Spehr M. Identification of a testicular odorant receptor mediating human sperm chemotaxis. Science 2003;299: 2054–8. 748. Fukuda M, Yomogida K, Okabe M, Touhara K. Functional characterization of a mouse testicular olfactory receptor and its role in chemosensing and in regulation of sperm motility. J Cell Sci 2004;117:5835–45. 749. Vosshall LB. Olfaction: attracting both sperm and the nose. Curr Biol 2004;14:R918–20. 750. Tatsura H, Nagao H, Tamada A, Sasaki S, Kohri K, Mori K. Developing germ cells in mouse testis express pheromone receptors. FEBS Lett 2001;488:139– 44. 751. Goto T, Salpekar A, Monk M. Expression of a testisspecific member of the olfactory receptor gene family in human primordial germ cells. Mol Hum Reprod 2001;7:553–8. 752. Walensky LD, Roskams AJ, Lefkowitz RJ, Snyder SH, Ronnett GV. Odorant receptors and desensitization proteins colocalize in mammalian sperm. Mol Med 1995;1:130–41. 753. Vanderhaeghen P, Schurmans S, Vassart G, Parmentier M. Olfactory receptors are displayed on dog mature sperm cells. J Cell Biol 1993;123:1441–52.
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754. Spehr M, Schwane K, Riffell JA, Barbour J, Zimmer RK, Neuhaus EM, Hatt H. Particulate adenylate cyclase plays a key role in human sperm olfactory receptor-mediated chemotaxis. J Biol Chem 2004;279:40194–203. 755. Giojalas LC, Rovasio RA. Mouse spermatozoa modify their motility parameters and chemotactic response to factors from the oocyte microenvironment. Int J Androl 1998;21:201–6. 756. Carnevale EM, Maclellan LJ, Coutinho da Silva MA, Checura CM, Scoggin CF, Squires EL. Equine spermoocyte interaction: results after intraoviductal and intrauterine inseminations of recipients for oocyte transfer. Anim Reprod Sci 2001;68:305–14. 757. Sun F, Bahat A, Gakamsky A, Girsh E, Katz N, Giojalas LC, Tur-Kaspa I, Eisenbach M. Human sperm chemotaxis: both the oocyte and its surrounding cumulus cells secrete sperm chemoattractants. Hum Reprod 2005;20:761–7. 758. Suarez SS, Pacey AA. Sperm transport in the female reproductive tract. Hum Reprod Update 2006;12:23–37. 759. Familiari G, Relucenti M, Heyn R, Micara G, Correr S. Three-dimensional structure of the zona pellucida at ovulation. Microsc Res Tech 2006;69:415–26. 760. Primakoff P, Myles DG. Penetration, adhesion, and fusion in mammalian sperm-egg interaction. Science 2002;296:2183–5. 761. Meyers SA. Equine sperm-oocyte interaction: the role of sperm surface hyaluronidase. Anim Reprod Sci 2001;68: 291–303. 762. Olds-Clarke P. How does poor motility alter sperm fertilizing ability. J Androl 1996;17:183–6. 763. Wassarman PM. Role of carbohydrates in receptormediated fertilization in mammals. Ciba Found Symp 1989;145:135–49. 764. Vierira A, Miller DJ. Gamete interaction: is it speciesspecific? Mol Reprod Dev 2006;73:1422–9. 765. Velasquez JG, Canovas S, Barajas P, Marcos J, JimenezMovilla M, Gallego RG, Ballesta J, Aviles M, Coy P. Role of sialic acid in bovine sperm-zona pellucida binding. Mol Reprod Dev 2007;74:617–28. 766. Skutelsky E, Ranen E, Shalgi R. Variations in the distribution of sugar residues in the zona pellucida as possible species-specific determinants of mammalian oocytes. J Reprod Fertil 1994;100:35–41. 767. Tantibhedhyangkul J, Weerachatyanukul W, Carmona E, Xu H, Anupriwan A, Michaud D, Tanphaichitr N. Role of sperm surface arylsulfatase A in mouse sperm-zona pellucida binding. Biol Reprod 2002;67:212–19. 768. Cheng A, Le T, Placios M, Bookbinder LH, Wassarman PM. Sperm-egg recognition in the mouse: characterization of sp56, a sperm protein having specific affinity for ZP3. J Cell Biol 1994;125:867–78. 769. Mori E, Mori T, Takasaki S. Binding of mouse sperm to beta-galactose residues on egg zona pellucida and asialofetuin-coupled beads. Biochem Biophys Res Commun 1997;238:95–9. 770. Nixon B, Lu Q, Wassler MJ, Foote CI, Ensslin MA, Shur BD. Galactosyltransferase function during mammalian fertilization. Cells Tissues Organs 2001;168:46– 57. 771. Thaler CD, Cardullo RA. Defining oligosaccharide specificity for initial sperm-zona pellucida adhesion in the mouse. Mol Reprod Dev 1996;45:535–46. 772. Cornwall GA, Tulsiani DRP, Orgebin-Crist MC. Inhibition of the mouse sperm surface alpha-d-mannosidase
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Stallion: Anatomy, Physiology and Endocrinology inhibits sperm-egg binding in vitro. Biol Reprod 1991; 44:913–21. Tulsiani DRP, Abou-Haila A. Mammalian sperm molecules that are potentially important in interaction with female genital tract and egg vestments. Zygote 2001; 9:51–69. Rattanachaiyanont M, Weerachatyanukul W, Leveille M-C, Taylor T, D’Amours D, Leader A, Tanphaichitr N. Anti-SLIP1-reactive proteins exist on human sperm and are involved in zona pellucida binding. Mol Hum Reprod 2001;7:633–40. Rodeheffer C, Shur BD. Characterization of a novel ZP3independent sperm-binding ligand that facilitates sperm adhesion to the egg coat. Development 2004;131:503–12. Shur BD, Rodeheffer C, Ensslin MA, Lyng R, Raymond A. Identification of novel gamete receptors that mediate sperm adhesion to the egg coat. Mol Cell Endocrinol 2006;250:137–48. Dean J. Reassessing the molecular biology of sperm-egg recognition with mouse genetics. Bioessay 2004;26:29–38. Baibakov B, Gauthier L, Talbot P, Rankin TL, Dean J. Sperm binding to the zona pellucida is not sufficient to induce acrosome exocytosis. Development 2007;134: 933–43. Mugnier S, Lahuec C, Yvon JM, Magistrini M, Goudet G. Equine sperm surface galactosyltransferase and zona glycoprotein ZPC are not necessary for sperm binding to the zona pellucida. Anim Reprod Sci 2006;94:331–2. Chiu PCN, Chung M-K, Koistinen R, Koistinen H, Seppala M, Ho P-C, Ng EHY, Lee K-F, Yeung WSB. Cumulus oophorus-associated glycodelin-C displaces sperm bound glycodelin-A and -F and stimulates spermatozoazona pellucida binding. J Biol Chem 2006;282: 5378–88. O’Day-Bowman MB, Mavrogianis PA, Minshall RD, Verhage HG. In vivo versus in vitro oviductal glycoprotein (OGP) association with the zona pellucida (ZP) in the hamster and baboon. Mol Reprod Dev 2002;62:248–56. Verhage HG, Mavrogianis PA, O’Day-Bowman MB, Schmidt A, Arias EB, Donnelly KM, Boomsma RA, Thibodeaux JK, Fazleabas AT, Jaffe RC. Characteristics of an oviductal glycoprotein and its potential role in the fertilization process. Biol Reprod 1998;58:1098–101. Bedford JM. Mammalian fertilization misread? Sperm penetration of the eutherian zona pellucida is unlikely to be a lytic event. Biol Reprod 1998;59:1275–87. Ohmura K, Kohno N, Kobayashi Y, Yamagata K, Sato S, Kashiwabara S, Baba T. A homologue of pancreatic trypsin is localized in the acrosome of mammalian sperm and is released during acrosome reaction. J Biol Chem 1999;274:29426–32. Tulsiani DRP, Abou-Haila A, Loeser CR, Pereira BMJ. The biological and functional significance of the sperm acrosome and acrosomal enzymes in mammalian fertilization. Exp Cell Res 1998;240:151–64. Drobnis EZ, Yudin AI, Cherr GN, Katz DF. Hamster sperm penetration of the zona pellucida: kinematic analysis and mechanical implications. Dev Biol 1988;130: 311–23. Katz DF, Cherr GN, Lambert H. The evolution of hamster sperm motility during capacitation and interaction with the ovum vestments in vitro. Gamete Res 1986;14:333–46. Wassarman PM, Jovine L, Qi H, Williams Z, Darie C, Litscher ES. Recent aspects of mammalian fertilization research. Mol Cell Endocrinol 2005;234:95–103.
789. Rubinstein E, Ziyyat A, Wolf JP, Le Naour F, Boucheix C. The molecular players of sperm-egg fusion in mammals. Semin Cell Dev Biol 2006;17:254–63. 790. Cuasnicu PS, Ellerman DA, Cohen DJ, Busso D, Morgenfeld MM, Da Ros VG. Molecular mechanisms involved in mammalian gamete fusion. Arch Med Res 2001;32:614–18. 791. Cameo MS, Blaquier JA. Androgen-controlled specific proteins in rat epididymis. J Endrocrinol 1976;69:47–55. 792. Rochwerger L, Cuasnicu PS. Redistribution of a rat sperm epididymal glycoprotein after in vitro and in vivo capacitation. Mol Reprod Dev 1992;31:34–41. 793. Gan SW, Xin L, Torres J. The transmembrane homotrimer of ADAM 1 in model lipid bilayers. Protein Sci 2007; 16:285–92. 794. Kim E, Yamashita M, Nakanishi T, Park K-E, Kimura M, Kashiwabara S-I, Baba T. Mouse sperm lacking ADAM1b/ADAM2 fertilin can fuse with the egg plasma membrane and effect fertilization. J Biol Chem 2006; 281:5634–9. 795. Inoue N, Ikawa M, Isotani A, Okabe M. The immunoglobulin superfamily protein Izumo is required for sperm to fuse with eggs. Nature 2005;434:234–8. 796. Goncalves RF, Wolinetz CD, Killian GJ. Influence of arginine-glycine-aspartic acid (RGD), integrins (alphav and alpha-5) and osteopontin on bovine sperm-egg binding, and fertilization in vitro. Theriogenology 2007; 67:468–74. 797. Stein KK, Go JC, Lane WS, Primakoff P, Myles DG. Proteomic analysis of sperm regions that mediate sperm-egg interactions. Proteomics 2006;6:3533–43. 798. Stein KK, Primakoff P, Myles D. Sperm-egg fusion: events at the plasma membrane. J Cell Sci 2004;177: 6269–74. 799. Runge KE, Evans JE, He Z-Y, Gupta S, McDonald KL, Stahlberg H, Primakoff P, Myles DG. Oocyte CD9 is enriched on the microvillar membrane and required for normal microvillar shape and distribution. Dev Biol 2007;304:317–25. 800. Atif SM, Hasan I, Ahmad N, Khan U, Owais M. Fusogenic potential of sperm membrane lipids: Nature’s wisdom to accomplish targeted gene delivery. FEBS Lett 2006;580:2183–90. 801. Tsaadon A, Eliyahu E, Shtraizent N, Shalgi R. When a sperm meets an egg: Block to polyspermy. Mol Cell Endocrinol 2006;252:107–14. 802. Kumakiri J, Oda S, Kinoshita K, Miyazaki S. Involvement of Rho family G protein in the cell signaling for sperm incorporation during fertilization of mouse eggs: inhibition by Clostridium difficile toxin B. Dev Biol 2003;260:522–35. 803. Sanchez-Gutierrez M, Contreras RG, Mujica A. Cytochalasin-D retards sperm incorporation deep into the egg cytoplasm but not membrane fusion with the egg plasma membrane. Mol Reprod Dev 2002;63:518–28. 804. Tremoleda JL, Colenbrander B, Stout TAE, Bevers MM. Reorganization of the cytoskeleton and chromatin in horse oocytes following intracytoplasmic sperm injection. Theriogenology 2002;58:697–700. 805. Bedford SJ, Kurokawa M, Hinrichs K, Fissore RA. Patterns of [Ca2+ ]i oscillations after intracytoplasmic sperm injection of in vitro and in vivo matured horse oocytes. Theriogenology 2002;58:701–4. 806. Comizzoli P, Urner F, Sakkas D, Renard JP. Upregulation of glucose metabolism during male pronucleus formation determines the early onset of the s phase in bovine zygotes. Biol Reprod 2003;68:1934–40.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 807. Gardner AJ, Evans JP. Mammalian membrane block to polyspermy: new insights into how mammalian eggs prevent fertilisation by multiple sperm. Reprod Fertil Dev 2006;18:53–61. 808. Yanagimachi R. Male gamete contributions to the embryo. Ann N Y Acad Sci 2005;1061:203–7. 809. Dumollard R, Duchen M, Sardet C. Calcium signals and mitochondria at fertilisation. Semin Cell Dev Biol 2006;17:314–23. 810. Kline D, Kline JT. Repetitive calcium transients and the role of calcium in exocytosis and cell cycle activation in the mouse egg. Dev Biol 1992;149:80–9. 811. Kurokawa M, Sato K, Wu H, He C, Malcuit C, Black SJ, Fukami K, Fissore RA. Functional, biochemical, and chromatographic characterization of the complete [Ca2+ ]i oscillation-inducing activity of porcine sperm. Dev Biol 2005;285:376–92. 812. Swann K, Saunders CM, Lai FA. The cytosolic sperm factor that triggers Ca2+ oscillations and egg activation in mammals is a novel phospholipase C: PLCzeta. Reproduction 2004;127:431–9. 813. Saunders CM, Swann K, Lai FA. PLCzeta, a spermspecific PLC and its potential role in fertilization. Biochem Soc Symp 2007;74:23–36. 814. Evans JP, Florman HM. The state of the union: the cell biology of fertilization. Nat Cell Biol 2002;4 Suppl.:57–63. 815. Leese HJ, Barton AM. Production of pyruvate by isolated mouse cumulus cells. J Exp Zool 1985;234:231–6. 816. Brinsko SP, Varner DD. Artificial insemination and preservation of semen. In: Blanchard TL, Varner DD (eds) The Veterinary Clinics of North America: Equine Practice (Stallion Management). Philadelphia: W.B. Saunders Co., 1992; pp. 205–18. 817. Kenney RM, Hurtgen J, Pierson R, Witherspoon D, Simons J. Theriogenology and the Equine. Part II. The Stallion (Society For Theriogenology Manual For Clinical Fertility Evaluation of the Stallion). Hastings, NE: Society For Theriogenology, 1983. 818. Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL. Examination of the stallion for breeding soundness. In: Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL (eds) Manual of Equine Reproduction. St. Louis: Mosby, 2003; pp. 143–64. 819. Jasko DJ. Evaluation of stallion semen. In: Blanchard TL, Varner DD (eds) The Veterinary Clinics of North America: Equine Practice (Stallion Management). Philadelphia: W.B. Saunders Co., 1992; pp. 129–48. 820. Pickett BW. Reproductive evaluation of the stallion. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 755–68. 821. Jasko DJ, Little TV, Lein DH, Foote RH. Determination of stallion semen quality and its relationship with fertility. J Reprod Fertil Suppl 1991;44:649–50. 822. Jasko DJ, Little TV, Lein DH, Foote RH. Comparison of spermatozoal movement and semen characteristics with fertility in stallions: 64 cases (1987–1988). J Am Vet Med Assoc 1992;200:979–85. 823. Kenney RM, Kingston RS, Rajamannon AH, Ramberg Jr CF. Stallion semen characteristics for predicting fertility. In: Proceedings of the 17th Annual Convention of the American Association of Equine Practitioners, 1971, pp. 53–67. 824. Colenbrander B, Puyk H, Zandee AR, Parlevliet J. Evaluation of the stallion for breeding. Acta Vet Scand Suppl 1992;88:29–37.
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825. Amann RP. Can the fertility potential of a seminal sample be predicted accurately? J Androl 1989;10:89–98. 826. Bielanski W, Dudek E, Bittmar A, Kosiniak K. Some characteristics of common abnormal forms of spermatozoa in highly fertile stallions. J Reprod Fertil Suppl 1982;32:21–6. 827. David JSE. Semen examination for fertility potential in stallions. J Reprod Fertil Suppl 1982;32:635. 828. Voss JL, Pickett BW, Loomis PR. The relationship between seminal characteristics and fertility in thoroughbred stallions. J Reprod Fertil Suppl 1982;32:635–6. 829. Rousset H, Chanteloube P, Magistrini M, Palmer E. Assessment of fertility and semen evaluations of stallions. J Reprod Fertil Suppl 1987;35:25–31. 830. Dowsett KF, Pattie WA. Characteristics and fertility of stallion semen. J Reprod Fertil Suppl 1982;32:1–8. 831. Voss JL, Pickett BW, Squires EL. Stallion spermatozoal morphology and motility and their relationships to fertility. J Am Vet Med Assoc 1981;178:287–9. 832. Clement F, Ladonnet Y, Magistrini M. Sperm morphology and fertility. Anim Reprod Sci 2001;68:362–3. 833. Kenney RM, Evenson DP, Garcia MC, Love C. Relationships between sperm chromatin structure, motility, and morphology of ejaculated sperm, and seasonal pregnancy rate. Biol Reprod Monogr 1995;6:647–53. 834. Wang C, Chan SYW, Ng M, So WWK, Tsoi W-L, Lo T, Leung A. Diagnostic value of sperm function tests and routine semen analyses in fertile and infertile men. J Androl 1988;9:384–9. 835. Aitken RJ. Sperm function tests and fertility. Int J Androl 2006;29:69–75. 836. Rodriguez-Martinez H. Can we increase the estimative value of semen assessment? Reprod Domest Anim 2006; 41:2–10. 837. Graham JK, Moce E. Fertility evaluation of frozen/ thawed semen. Theriogenology 2005;64:492–504. 838. Evenson DP, Darzynkiewica Z, Melamed MR. Relation of mammalian sperm chromatin heterogeneity to fertility. Science 1980;210:1131–3. 839. Love CC. The sperm chromatin structure assay: a review of clinical applications. Anim Reprod Sci 2005;89:39–45. 840. Love CC, Kenney RM. The relationship of increased susceptibility of sperm DNA to denaturation and fertility in the stallion. Theriogenology 1998;57:955–72. 841. Evenson DP, Sailer BL, Jost LK. Relationship between stallion sperm deoxyribonucleic acid (DNA) susceptibility to denaturation in situ and presence of DNA strand breaks: implications for fertility and embryo viability. Biol Reprod Monogr 1995;6:655–9. 842. Evenson DP, Jost LK, Varner DD. Relationship between sperm nuclear protamine free -SH status and susceptibility to DNA denaturation. J Reprod Fertil Suppl 2000;56:401–6. 843. Love CC, Kenney RM. Scrotal heat stress induces altered sperm chromatin structure associated with decrease in protamine disulfide bonding in the stallion. Biol Reprod 1999;60:615–20. 844. Makhlouf AA, Niederberger C. DNA integrity tests in clinical practice: it is not a simple matter of black and white (or red and green). J Androl 2006;27:316–23. 845. O’Brien J, Zini A. Sperm DNA integrity and male infertility. Urology 2005;65:16–22. 846. Aoki VW, Emery BR, Liu L, Carrell DT. Protamine levels vary between individual sperm cells of infertile human males and correlate with viability and DNA integrity. J Androl 2006;27:890–8.
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Stallion: Anatomy, Physiology and Endocrinology
847. Lewis SEM, Aitken RJ. DNA damage to spermatozoa has impacts on fertilization and pregnancy. Cell Tissue Res 2005;322:33–41. 848. Aoki VW, Moskovtsev SI, Willis J, Liu L, Mullen JB, Carrell DT. DNA integrity is compromised in protaminedeficient human sperm. J Androl 2005;26:741–8. 849. Sailer BL, Sarkar LJ, Bjordahl JA, Jost LK, Evenson DP. Effects of heat stress on mouse testicular cells and sperm chromatin structure. J Androl 1997;18:294–301. 850. Oliva R. Protamines and male infertility. Hum Reprod Update 2006;12:417–35. 851. Fraga CG, Motchnik PA, Shigenaga MK, Hellbock HJ, Jacob RA, Ames BN. Ascorbic acid protects against endogenous oxidative DNA damage in human sperm. Proc Natl Acad Sci USA 1991;88:11003–6. 852. Fatehi AN, Bevers MM, Schoevers E, Roelen BAJ, Colenbrander B, Gadella BM. DNA damage in bovine sperm does not block fertilization and early embryonic development but induces apoptosis after the first cleavages. J Androl 2006;27:176–88. 853. Agarwal A, Said TM. Role of sperm chromatin abnormalities and DNA damage in male infertility. Hum Reprod Update 2003;9:331–45. 854. Chohan KR, Griffin JT, Lafromboise M, De Jonge CJ, Carrell DT. Comparison of chromatin assays for DNA fragmentation evaluation in human sperm. J Androl 2006;27:53–9. 855. Erenpreiss J, Spano M, Erenpreisa J, Bungum M, Giwercman A. Sperm chromatin structure and male fertility: biological and clinical aspects. Asian J Androl 2006;8:11– 29. 856. Evenson DP, Wixon R. Comparison of the Halosperm test kit with the sperm chromatin structure assay (SCSA) infertility test in relation to patient diagnosis and prognosis. Fertil Steril 2005;84:846–9. 857. Linfor JJ, Meyers SA. Detection of DNA damage in response to cooling injury in equine spermatozoa using single-cell gel electrophoresis. J Androl 2002;23:107–13. 858. Meyers SA, Overstreet JW, Liu IKM, Drobnis EZ. Capacitation in vitro of stallion spermatozoa: comparison of progesterone-induced acrosome reactions in fertile and subfertile males. J Androl 1995;16:47–54. 859. Cheng FP, Gadella BM, Voorhout WF, Fazeli A, Bevers A, Colenbrander B. Progesterone-induced acrosome reaction in stallion spermatozoa is mediated by a plasma membrane progesterone receptor. Biol Reprod 1998;59:733–42. 860. Rathi R, Nielen M, Cheng FP, Van Buiten A, Colenbrander B. Exposure of progesterone receptors on the plasma membranes of stallion spermatozoa as a parameter for prediction of fertility. J Reprod Fertil Suppl 2000;56:87– 91. 861. Varner DD, Brinsko SP, Blanchard TL, Love CC, Macpherson ML, Heck RS, Johnson L. Subfertility in stallions associated with acrosome dysfunction. In: Proceedings of the 47th Annual Convention of the American Association of Equine Practitioners, 2001, pp. 227–8. 862. Brinsko SP, Love CC, Bauer JE, Macpherson ML, Varner DD. Cholesterol-to-phospholipid ratio in whole sperm and seminal plasma from fertile stallions and stallions with unexplained subfertility. Anim Reprod Sci 2007;99:65–71. 863. Varner DD, Ward CR, Storey BA, Kenney RM. Induction and characterization of the acrosome reaction in equine spermatozoa. Am J Vet Res 1987;48:1383–9.
864. Brum AM, Thomas AD, Sabeur K, Ball BA. Evaluation of coomassie blue staining of the acrosome of equine and canine spermatozoa. Am J Vet Res 2006;67:358–62. 865. Cardullo RA, Florman HM. Strategies and methods for evaluating acrosome reaction. Methods Enzymol 1993; 225:136–53. 866. Bosard T, Love C, Brinsko SP, Blanchard T, Thompson J, Varner D. Evaluation and diagnosis of acrosome function/dysfunction in the stallion. Anim Reprod Sci 2005;89:215–17. 867. Varner DD, Thompson JA, Blanchard TL, Heck R, Love CC, Brinsko SP, Johnson L. Induction of the acrosome reaction in stallion spermatozoa: effects of incubation temperature, incubation time, and ionophore concentration. Theriogenology 2002;58:303–6. 868. Bosard TS. Flow cytometric evaluation of acrosome function/dysfunction in the stallion. Master’s Thesis, Texas A&M University, 2006. 869. Carrell BP. Validation of pisum sativum agglutinin fluorescent marker for stallion spermatozoal acrosomes with transmission electron microscopy. Master’s Thesis, Texas A&M University, 2001. 870. Graham JK. Assessment of sperm quality: a flow cytometric approach. Anim Reprod Sci 2001;68:239–47. 871. Silva PFN. Detection of damage in mammalian sperm cells. Theriogenology 2006;65:958–79. 872. Farlin ME, Jasko DJ, Graham JK, Squires EL. Assessment of Pisum sativum agglutinin in identifying acrosomal damage in stallion spermatozoa. Mol Reprod Dev 1992;32:23–7. 873. Thomas CA, Garner DL, DeJarnette JM, Marshall CE. Effect of cryopreservation on bovine sperm organelle function and viability as determined by flow cytometry. Biol Reprod 1998;58:786–93. 874. Ward CR, Storey BT. Determination of the time course of capacitation in mouse spermatozoa using a chlortetracycline fluorescence assay. Dev Biol 1984;104:287–96. 875. de Vries KJ, Wiedmer T, Sims PJ, Gadella BM. Caspaseindependent exposure of aminophospholipids and tyrosine phosphorylation in bicarbonate responsive human sperm cells. Biol Reprod 2003;68:2122–34. 876. Gadella BM, Miller NGA, Colenbrander B, van Golde LMG, Harrison RAP. Flow cytometric detection of transbilayer movement of fluorescent phospholipid analogues across the boar sperm plasma membrane: elimination of labelling artefacts. Mol Reprod Dev 1999;53:108–25. 877. Miki K, Qu W, Goulding EH, Willis WD, Bunch DO, Strader LF, Perreault SD, Eddy EM, O’Brian DA. Glyceraldehyde 3-phosphate dehydrogenase-S, a spermspecific glycolytic enzyme, is required for sperm motility and male fertility. Proc Natl Acad Sci USA 2004;101: 16506. 878. Smiley ST, Reers M, Mottola-Hartshorn C, Lin M, Chen A, Smith TW, Steele Jr GD, Chen LB. Intracellular heterogeneity in mitochondrial membrane potentials revealed by a J-aggregate-forming lipophilic cation JC-1. Proc Natl Acad Sci USA 1991;88:3671–5. 879. Cossarizza A, Ceccarelli D, Masini A. Functional heterogeneity of an isolated mitochondrial population revealed by cytofluorometric analysis at the single organelle level. Exp Cell Res 1996;222:84–94. 880. Garner DL, Thomas CA, Joerg HW, DeJarnette JM, Marshall CE. Fluorometric assessments of mitochondrial function and viability in cryopreserved bovine spermatozoa. Biol Reprod 1997;57:1401–6.
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From a Sperm’s Eye View: Revisiting Our Perception of this Intriguing Cell 881. Gravance CG, Garner DL, Miller MG, Berger T. Fluorescent probes and flow cytometry to assess rat sperm integrity and mitochondrial function. Reprod Toxicol 2001;15:5–10. 882. Neild D, Chaves G, Flores M, Mora N, Beconi M, Aguero A. Hypoosmotic test in equine spermatozoa. Theriogenology 1999;51:721–7. 883. Jeyendran RS, Van der Ven HH, Zaneveld LJ. The hypoosmotic swelling test: an update. Arch Androl 1992;29: 105–16. 884. Kenney RM. Manual for Clinical Fertility Evaluation of the Stallion. Hastings, NE: Society for Theriogenology, 1983. 885. Garner DL, Dobrinsky JR, Welch GR, Johnson LA. Porcine sperm viability, oocyte fertilization and embryo development after staining spermatozoa with SYBR-14. Theriogenology 1996;45:1103–13. 886. Garner DL, Johnson LA. Viability assessment of mammalian sperm using SYBR-14 and propidium iodide. Biol Reprod 1995;53:276–84. 887. Garner DL, Thomas CA, Allen CH. Effect of semen dilution on bovine sperm viability as determined by dual-DNA staining and flow cytometry. J Androl 1997; 18:324–31. 888. Love CC, Thompson JA, Brinsko SP, Rigby SL, Blanchard TL, Lowry VK, Varner DD. Relationship between stallion sperm motility and viability as detected by two fluorescence staining techniques using flow cytometry. Theriogenology 2003;60:1127–38. 889. Garner DL, Thomas CA. Organelle-specific probe JC1 identifies membrane potential differences in the mitochondrial function of bovine sperm. Mol Reprod Dev 1999;53:222–9. 890. Kavak A, Johannisson A, Lundeheim N, RodriguezMartinez H, Aidnik M, Einarsson S. Evaluation of cryopreserved stallion semen from Tori and Estonian breeds using CASA and flow cytometry. Anim Reprod Sci 2003;76:205–16. 891. Kirk ES, Graham JK, Bruemmer JE, Squires EL. Evaluating frozen equine semen by flow cytometry. Anim Reprod Sci 2001;68:348–9. 892. Cross NL, Morales P, Overstreet JW, Hanseon FW. Two simple methods for detecting acrosome-reacted human sperm. Gamete Res 1986;15:213–26. 893. Cross NL, Meizel S. Methods for evaluating the acrosomal status of mammalian sperm. Biol Reprod 1989;41: 635–41. 894. Graham JK, Kunze E, Hammerstedt RH. Analysis of sperm cell viability, acrosomal integrity, and mitochondrial function using flow cytometry. Biol Reprod 1990;43: 55–64. 895. Kavak A, Johannisson A, Malmgren L, RodriguezMartinez H, Aidnik M, Einarsson S. Post-thaw evaluation of stallion spermatozoa using triple fluorescent staining and flow cytometry. Anim Reprod Sci 2001;68: 349–50. 896. Cai K, Yang J, Guan M, Ji W, Li Y, Rens W. Single UV excitation of Hoechst 33342 and propidium iodide for viability assessment of rhesus monkey spermatozoa using flow cytometry. Arch Androl 2005;51:371–83. 897. Gravance CG, Garner DL, Baumber J, Ball BA. Assessment of equine sperm mitochondrial function using JC-1. Theriogenology 2000;53:1691–703. 898. Ball BA, Fagnan MS, Dobrinski I. Determination of acrosin amidase activity in equine spermatozoa. Theriogenology 1997;48:1191–8.
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899. Kennedy WP, Kaminski JM, Van der Ven HH, Jeyendran RS, Reid DS, Blackwell J, Bielfeld P, Zaneveld LJD. A simple, clinical assay to evaluate the acrosin activity of human spermatozoa. J Androl 1989;10:221–31. 900. Cross NL. A modified and improved assay for sperm amidase activity. J Androl 1990;11:409–13. 901. Sousa APM, Gomes-Santos CSS, Ramalho-Santos J. Localization of snares, NSF and Caveolin 1 in human spermatozoa: relationship with seminal parameters. Arch Androl 2006;52:347–53. 902. Desvousges AL, Dow CA, Hayna JT, Miller L, Jousan D, Hansen PJ, Buhi WC, Troedsson MHT. Heat shock induces apoptosis in equine spermatozoa. Anim Reprod Sci 2006;94:125–6. 903. Brum A, Sabeur K, Ball BA. Apoptotic-like changes in equine spermatozoa after separation by density-gradient centrifugation or after cryopreservation. Anim Reprod Sci 2006;94:138–9. 904. Wang X, Sharma RK, Sikka SC, Thomas Jr AJ, Falcone T, Agarwal A. Oxidative stress is associated with increased apoptosis leading to spermatozoa DNA damage in patients with male factor infertility. Fertil Steril 2003;80:531–5. 905. Said TM, Paasch U, Glander H-J, Agarwal A. Role of caspases in male infertility. Hum Reprod Update 2004;10: 39–51. 906. Bellin ME, Oyarzo JN, Hawkins HE, Zhang H, Smith RG, Forrest DW, Sprott LR, Ax RL. Fertility-associated antigen on bull sperm indicates fertility potential. J Anim Sci 1998;76:2032–9. 907. Buffone MG, Calamera JC, Verstraeten SV, Doncel GF. Capacitation-associated protein tyrosine phosphorylation and membrane fluidity changes are impaired in the spermatozoa of asthenozoospermic patients. Reproduction 2005;129:697–705. 908. Yunes R, Doncel GF, Acosta AA. Incidence of sperm-tail tyrosine phosphorylation and hyperactivated motility in normozoospermic and asthenozoospermic human sperm samples. Biocell 2003;27:29–36. 909. Tardif S, Dube C, Chavalier S, Bailey JL. Capacitation is associated with tyrosine phosphorylation and tyrosine kinase-like activity in pig sperm proteins. Biol Reprod 2001;65:784–92. 910. Bianchi PG, Manicardi GC, Urner F, Campana A, Sakkas D. Chromatin packaging and morphology in ejaculated human spermatozoa: evidence of hidden anomalies in normal spermatozoa. Mol Hum Reprod 1996;2:139–44. 911. Nasr-Esfahani MH, Razavi S, Mardani M. Relation between different human sperm nuclear maturity tests and in vitro fertilization. J Assist Reprod Genet 2001;18: 219–25. 912. Iranpour FG, Nasr-Esfahani MH, Yalojerdi MR, al-Taraihi TM. Chromomycin A3 staining as a useful tool for evaluation of male fertility. J Assist Reprod Genet 2000;17:60–6. 913. Franken DR, Franken CJ, de la Guerre H, de Villiers A. Normal sperm morphology and chromatin packaging: comparison between aniline blue and chromomycin A3 staining. Andrologia 1999;31:361–6. 914. Gradil C, Yoon S-Y, Brown J, He C, Visconti P, Fissore RA. PLC-Zeta: a marker of fertility for stallions. Anim Reprod Sci 2006;94:23–5. 915. Swann K, Saunders CM, Rogers NT, Lai FA. PLCzeta: a sperm protein that triggers calcium ion oscillations and egg activation in mammals. Semin Cell Dev Biol 2006; 17:264–73.
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916. Neild DM, Gadella BM, Colenbrander B, Aguero A, Brouwers JFHM. Lipid peroxidation in stallion spermatozoa. Theriogenology 2002;58:295–8. 917. Neild DM, Brouwers JFHM, Colenbrander B, Aguero A, Gadella BM. Lipid peroxide formation in relation to membrane stability of fresh and frozen thawed stallion spermatozoa. Mol Reprod Dev 2005;72:230–8. 918. Guthrie HD, Welch GR. Use of fluorescence-activated flow cytometry to determine membrane lipid peroxidation during hypothermic liquid storage and freezethawing of viable boar sperm loaded with C11-BODIPY 581/591. J Anim Sci 2007;85:1402–11. 919. Brouwers JFHM, Gadella BM. In situ detection and localization of lipid peroxidation in individual bovine sperm cells. Free Radic Biol Med 2003;35:1382–91. 920. Brouwers JF, Silva PFN, Gadella BM. New assays for detection and localization of endogenous lipid peroxidation products in living boar sperm after BTS dilution or after freeze-thawing. Theriogenology 2005;63:458–69. 921. Roca J, Gil MA, Hernandez M, Parrilla I, Vazquez JM, Martinez EA. Survival and fertility of boar spermatozoa after freeze-thawing in extender supplemented with butylated hydroxytoluene. J Androl 2004;25:397–405. 922. Stradaioli G, Gennevieve A, Magistrini M. Evaluation of lipid peroxidation by thiobarbituric acid test in equine spermatozoa. Anim Reprod Sci 2001;68:320–2. 923. Stradaioli G, Magistrini M. A comparative evaluation of thiobarbituric acid methods for determination of malondialdehyde in equine spermatozoa. Theriogenology 2002;58:347–50. 924. Aitken RJ, Harkiss D, Buckingham DW. Analysis of lipid peroxidation mechanisms in human spermatozoa. Mol Reprod Dev 1993;35:302–15. 925. Martinez P, Proverbio F, Camejo MI. Sperm lipid peroxidation and pro-inflammatory cytokines. Asian J Androl 2007;9:102–7. 926. Agarwal A, Prabakaran SA. Mechanism, measurement, and prevention of oxidative stress in male reproductive physiology. Indian J Exp Biol 2005;43:963–74. 927. Peris SI, Bilodeau JF, Dufour M, Bailey JL. Impact of cryopreservation and reactive oxygen species on DNA integrity, lipid peroxidation, and functional parameters in ram sperm. Mol Reprod Dev 2007;74:878–92. 928. Kumar TR, Muralidhara. Induction of oxidative stress by organic hydroperoxides in testis and epididymal sperm of rats in vivo. J Androl 2007;28:77–85. 929. Foresta C, Flohe L, Garolla A, Roveri A, Ursini F, Maiorino M. Male fertility is linked to the selenoprotein phospholipid hydroperoxide glutathione peroxidase. Biol Reprod 2002;67:967–71. 930. Stradaioli G, Parillo F, Sylla L, Tubaro F, Gargiulo AM, Monaci M. Enzymatic and immunohistochemical evaluation of phospholipid hydroperoxide glutathione peroxidase (PHGPx) in bovine spermatozoa. In: Proceedings of the International Congress on Animal Reproduction, 2004, p. 192. 931. Weir CP. Spermatozoa have decreased antioxidant enzymatic capacity and increased reactive oxygen species production during aging in the brown Norway rat. J Androl 2007;28:229–40. 932. Mesequer M, de los Santos MJ, Simon C, Pellicer A, Remohi J, Garrido N. Effect of sperm glutathione peroxidases 1 and 4 on embryo asymmetry and blastocyst quality in ooctye donation cycles. Fertil Steril 2006;86:1376– 85.
933. Aitken RJ, Clarkson JS, Hargreave TB, Irvine DS, Wu FCW. Analysis of the relationship between defective sperm function and the generation of reactive oxygen species in cases of oligozoospermia. J Androl 1989;10: 214–20. 934. Ball BA, Baumber J, Vo A, Sabeur K. Effect of oxidative stress on equine spermatozoa. Anim Reprod Sci 2001; 68:364. 935. Guthrie HD, Welch GR. Determination of intracellular reactive oxygen species and high mitochondrial membrane potential in Percoll-treated viable boar sperm using fluorescence-activated flow cytometry. J Anim Sci 2006;84:2089–100. 936. Said TM, Agarwal A, Sharma RK, Mascha E, Sikka SC, Thomas Jr AJ. Human sperm superoxide anion generation and correlation with semen quality in patients with male infertility. Fertil Steril 2004;82:871–7. 937. Aitken RJ, Irvine DS, Wu FC. Prospective analysis of sperm-oocyte fusion and reactive oxygen species generation as criteria for the diagnosis of infertility. Am J Obstet Gynecol 1991;164:542–51. 938. Whittington K, Harrison SC, Williams KM, Day JL, McLaughlin EA, Hull MGR, Ford WCL. Reactive oxygen species (ROS) production and the outcome of diagnostic tests of sperm function. Int J Androl 1999;22:236–42. 939. Pesch S, Bergmann M, Bostedt H. Determination of some enzymes and macro- and microelements in stallion seminal plasma and their correlations to semen quality. Theriogenology 2006;66:307–13. 940. Mollova M, Atanassov B, Nedkova R, Kyrukchiev S. Biochemical and immunochemical characterization of boar sperm flagellar protein with role in hyperactivation/capacitation process. Reprod Biol 2006;6:79–94. 941. Yousef MI, Esmail AM, Baghdadi HH. Effect of isoflavones on reproductive performance, testosterone levels, lipid peroxidation, and seminal plasma biochemistry of male rabbits. J Environ Sci Health B 2004;39:819–33. 942. Noquera Velasco JA, Tovar Zapata I, Martinez Hernandez P, Perez Albacete M, Tortosa Oltra J, Parrilla Paricio JJ. Lactic dehydrogenase-C4 activity in seminal plasma and male infertility. Fertil Steril 1993;60:331–5. 943. Huszar G, Corrales M, Vigue L. Correlation between sperm creatine phosphokinase activity and sperm concentration in normospermic and oligospermic men. Gamete Res 1988;19:67–75. 944. Huszar G, Vigue L. Incomplete development of human spermatozoa is associated with increased creatine phosphokinase concentration and abnormal head morphology. Mol Reprod Dev 1993;34:292–8. 945. Huszar G, Stone K, Dix D, Vigue L. Putative creatine kinase M-isoform in human sperm is identified as the 70-kilodalton heat shock protein HspA2. Biol Reprod 2000;63:925–32. 946. Huszar G, Ozkavukcu S, Jakab A, Celik-Ozenci C, Sati GL, Cayli S. Hyaluronic acid binding ability of human sperm reflects cellular maturity and fertilizing potential: selection of sperm for intracytoplasmic sperm injection. Curr Opin Obstet Gynecol 2006;18:260–7. 947. Ellerman DA, Cohen DJ, Da Ros VG, Morgenfeld MM, Busso D, Cuasnicu PS. Sperm protein ‘DE’ mediates gamete fusion through an evolutionarily conserved site of the CRISP family. Dev Biol 2006;297:228–37. 948. Reineke A, Hess O, Schambony A, Petrunkina AM, Bader H, Sieme H, Topfer-Petersen E. Sperm-associated seminal plasma proteins: a novel approach for the evalua-
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955.
956.
957. 958.
959.
960.
961.
962.
963.
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tion of sperm fertilizing ability of stallions. Pferdcheikunde 1999;6:531–7. Jude R, Giese A, Piumi F, Guerin G, Sieme H, TopferPetersen E, Distl O, Leeb T. Molecular characterization of the CRISP 1 gene-a candidate gene for stallion fertility. Theriogenology 2002;58:417–20. Meyers SA, Rosenberger A, Orpneck K. Localization and cellular distribution of a unique hyaluronidase in stallion spermatozoa during epididymidal transit. J Reprod Fertil Suppl 2000;56:79–86. Jonakova V, Manaskova P, Kraus M, Liberda J, Ticha M. Sperm surface proteins in mammalian fertilization. Mol Reprod Dev 2000;56 (Suppl. 2):277. Rodriguez-Martinez H, Iborra A, Martinez P, Calvete JJ. Immunoelectronmicroscopic imaging of spermadhesin AWN epitopes on boar spermatozoa bound in vivo to the zona pellucida. Reprod Fertil Dev 1998;10:491–7. Jonakova V, Kraus M, Veselsky L, Cechova D, Bezouska K, Ticha M. Spermadhesins of the AQN and AWN families, DQH sperm surface protein and HNK protein in the heparin-binding fraction of boar seminal plasma. J Reprod Fertil 1998;114:25–34. Boue F, Sullivan R. Cases of human infertility are associated with the absence of P34H an epididymal sperm antigen. Biol Reprod 1996;54:1018–24. Sullivan R, Legare C, Villeneuve M, Foliguet B, Bissonnette F. Levels of P34H, a sperm protein of epididymal origin, as a predictor of conventional in vitro fertilization outcome. Fertil Steril 2006;85:1557–9. Xia WY, Huang YF, Xu XF. Epididymal sperm protein P34H and male reproduction. Zhonghua Nan Ke Xue 2002;8:356–8. Sullivan R. Male fertility markers, myth or reality. Anim Reprod Sci 2004; 82–83:341–7. Bailey LB, Brady HB, Tardif S, Thompson LD, Hardy DM. Zonadhesin expression and localization in stallion testes. Anim Reprod Sci 2006;94:56–9. Breazeale KR, Brady HA, Bi M, Uppuluri S, Thompson LD, Bruemmer JE, Hardy DM. Biochemical properties and localization of zonadhesin in equine spermatozoa. Theriogenology 2002;58:359–62. Hardy DM, Garbers DL. A sperm membrane protein that binds in a species-specific manner to the egg extracellular matrix is homologous to von Willebrand factor. J Biol Chem 1995;270:26025–8. Bi M, Hickox JR, Winfrey VP, Olson GE, Hardy DM. Processing, localization and binding activity of zonadhesin suggest a function in sperm adhesion to the zona pellucida during exocytosis. Biochem J 2003;375:477–88. Lea IA, Sivashanmuqam P, O’Rand MG. Zonadhesin: characterization, localization, and zona pellucida binding. Biol Reprod 2001;65:1691–700. Gao A, Garbers DL. Species diversity in the structure of zonadhesin, a sperm-specific membrane protein containing multiple cell adhesion molecule-like domains. J Biol Chem 1998;273:3415–21. Eager FE, Brady HA, Thompson LD, Bruemmer JE, Hardy DM. Biochemical properties of zonadhesin in spermatozoa of fertile and subfertile stallions. In: Proceedings of the 19th Annual Convention of the American Association of Equine Practitioners, 2005, pp. 95–100. Klinefelter GR, Welch JE, Perreault SD, Moore HD, Zucker RM, Suarez JD, Roberts NL, Bobseine K, Jeffay S. Localization of the sperm protein SP22 and inhibition of fertility in vivo and in vitro. J Androl 2002;23:48–63.
989
966. Miller LMJ, Greene ES, Roberts KP, Troedsson MHT. Expression of equine sperm protein 22kDa (SP22) in testicular and epididymal tissue. Anim Reprod Sci 2006;94:54–5. 967. Miller LT, Troedsson MH, Duoos LA, Klinefelter GR, Roberts KP. Immunocytochemical detection and localization of sperm protein 22 (SP22) in fresh and cryopreserved equine semen. Biol Reprod 2003;68 (Suppl. 1):166–7. 968. Wrench N, Pinto CRF, Klinefelter GR, Farin CE. Seasonal expression and pattern of SP22 immunolocalization in stallion semen. Anim Reprod Sci 2006;94:32–5. 969. Amann RP, Katz DF. Reflections on CASA after 25 years. J Androl 2004;25:317–25. 970. Jasko DJ, Lein DH, Foote RH. A comparison of two computer-automated semen analysis instruments for the evaluation of sperm motion characteristics in the stallion. J Androl 1990;11:453–9. 971. Varner DD, Vaughan SD, Johnson L. Use of a computerized system for evaluation of equine spermatozoal motility. Am J Vet Res 1991;52:224–30. 972. Wessel MT, Althouse GC. Validation of an objective approach for simultaneous assessment of viability and motility of fresh and cooled equine spermatozoa. Anim Reprod Sci 2006;94:21–2. 973. Holt C, Holt WV, Moore HDM, Reed HCB, Curnock RM. Objectively measured boar sperm motility parameters correlate with the outcomes of on-farm inseminations: results of two fertility trials. J Androl 1997;18:312–23. 974. Williams WW. Cytology of the human spermatozoon. Fertil Steril 1950;1:199–215. 975. Casarett GW. A one-solution stain for spermatozoa. Stain Technol 1953;28:125–7. 976. Penedo MC, Millon LV, Bernoco D, Bailey E, Binns M, Cholewinsko G, Ellis N, Flynn J, Gralak B, Guthrie A, Hasegawa T, Lindgren G, Lyons LA, Roed KH, Swinburne JE, Tozaki T. International equine gene mapping workshop report: a comprehensive linkage map constructed with data from new markers and by merging four mapping resources. Cytogenet Genome Res 2005; 111:5–15. 977. Chowdhary BP, Taudsepp T, Kata SR, Goh B, Millon LV, Allan V, Piumi F, Guerin G, Swinburne J, Binns M, Lear TL, Mickelson J, Murray J, Antczak DF, Womack JE, Skow LC. The first-generation whole-genome radiation hybrid map in the horse identifies conserved segments in human and mouse genomes. Genome Res 2003;13:742–51. 978. Chowdhary BP, Bailey E. Equine genomics: galloping to new frontiers. Cytogenet Genome Res 2003;102:184–8. 979. Tozaki T, Hirota KI, Hasegawa T, Ishida N, Tobe T. Whole-genome linkage disequilibrium screening for complex traits in horses. Mol Genet Genomics 2007;277: 663–72. 980. Tozaki T, Swinburne J, Hirota KI, Hasegawa T, Ishida N, Tove T. Improved resolution of the comparative horsehuman map: investigating markers with in silico and linkage mapping approaches. Gene 2007;392:181–6. 981. Shiue Y-L, Bickel LA, Caetano AR, Millon LV, Clark RS, Eggleston ML, Michelmore R, Bailey E, Guerin G, Godard S, Mickelson JR, Valberg SJ, Murray SJ, Murray JD, Bowling AT. A synteny map of the horse genome comprised of 240 microsatellite and RAPD markers. Anim Genet 1999;30:1–9. 982. Wrobel G, Priming M. Mammalian male germ cells are fertile ground for expression profiling of sexual reproduction. Reproduction 2005;129:1–7.
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983. Chan W-Y, Wu SM, Ruszczyk L, Law E, Lee TL, Baxendale V, Pang ALY, Rennert OM. The complexity of antisense transcription revealed by the study of developing male germ cells. Genomics 2006;87:681–92. 984. Martins RP, Krawetz SA. RNA in human sperm. Asian J Androl 2005;7:115–20. 985. He Z, Chan WYC, Dym M. Microarray technology offers a novel tool for the diagnosis and identification of therapeutic targets for male infertility. Reproduction 2006;132:11–19. 986. Laughlin AM, Forrest DW, Varner DD, Blanchard TL, Love CC, Welsh Jr TW, Johnson L, Ing NH. DNA microarray analysis of gene expression in testicular tissue of stallions. Theriogenology 2002;58:413–15. 987. Turner RM, Casas-Dolz R. Differential gene expression in stallions with idiopathic testicular degeneration. Theriogenology 2002;58:421–4. 988. Smith GW, Rosa GJM. Interpretation of microarray data: trudging out of the abyss towards elucidation of biological significance. J Anim Sci 2007;85 (E.Suppl.):E20–3. 989. Leeb T, Sieme H, Topfer-Petersen E. Genetic markers for stallion fertility-lessons from humans and mice. Anim Reprod Sci 2005;89:21–9. 990. Andersen JS, Mann M. Organellar proteomics: turning inventories into insights. EMBO Rep 2006;7:874–9. 991. Pilch B, Mann M. Large-scale and high-confidence proteomic analysis of human seminal plasma. Genome Biol 2006;7:R40. 992. Martinez-Heredia J, Estanyol JM, Ballesca JL, Oliva R. Proteomic identification of human sperm proteins. Proteomics 2006;6:4356–69. 993. Chu DS, Liu H, Nix P, Wu TF, Ralston EJ, Yates III JR, Meyer BJ. Sperm chromatin proteomics identifies evolutionarily conserved fertility factors. Nature 2006;443:101–5. 994. Bailey JL, Tardif S, Dube C, Beaulieu M, Reyes-Moreno C, Lefievre L, Leclerc P. Use of phosphoproteomics to study tyrosine kinase activity in capacitating boar sperm. Kinase activity and capacitation. Theriogenology 2005; 63:599–614. 995. Lalancette C, Faure RL, Leclerc P. Identification of the proteins present in the bull sperm cytosolic fraction enriched in tyrosine kinase activity: a proteomic approach. Proteomics 2006;6:4523–40. 996. Swan SH, Elkin EP, Fenster L. The question of declining sperm density revisited: an analysis of 101 studies published 1934–1996. Environ Health Perspect 2000;108:961–6. 997. Carlsen E, Giwercman A, Keiding N, Skakkebaek NE. Evidence for decreasing quality of semen during past 50 years. Br Med J 1992;305:609–13. 998. Swan SH, Elkin EP, Fenster L. Have sperm densities declined? A reanalysis of global trend data. Environ Health Perspect 1997;105:1228–32.
999. Huyghe E, Matsuda T, Thonneau P. Increasing incidence of testicular cancer worldwide: a review. J Urol 2003;170:5–11. 1000. Paulozzi L. International trends in rates of hypospadias and cryptorchidism. Environ Health Perspect 1999; 107:297–302. 1001. Swan SH. Does our environment affect our fertility? Some examples to help reframe the question. Semin Reprod Med 2006;24:142–6. 1002. Hauser R. The environment and male fertility: recent research on emerging chemicals and semen quality. Semin Reprod Med 2006;24:156–67. 1003. Lahousse SA, Wallace DG, Liu D, Gaido KW, Johnson KJ. Testicular gene expression profiling following prepubertal rat mono-(2-ethylhexyl) phthalate exposure suggests a common initial genetic response at fetal and prepubertal ages. Toxicol Sci 2006;93:369–81. 1004. Fukushima T, Yamamoto T, Kikkawa R, Hamada Y, Komiyama M, Mori C, Horii I. Effects of male reproductive toxicants on gene expression in rat testes. J Toxicol Sci 2005;30:195–206. 1005. Ball BA, Sabeur K, Allen WR. Uptake of exogenous DNA by equine spermatozoa and applications in spermmediated gene transfer. Anim Reprod Sci 2006;94:115–16. 1006. Robl JM, Wang Z, Kasinathan P, Kuroiwa Y. Transgenic animal production and animal biotechnology. Theriogenology 2007;67:127–33. 1007. Dobrinski I. Germ cell transplantation and testis tissue xenografting in domestic animals. Anim Reprod Sci 2005;89:137–45. 1008. Hamra FK, Chapman KM, Nguyen DM, WilliamsStephens AA, Hammer RE, Garbers DL. Self renewal, expansion, and transfection of rat spermatogonial stem cells in culture. Proc Natl Acad Sci USA 2005;102:17430–5. 1009. Honaramooz A, Li MW, Penedo MC, Meyers S, Dobrinski I. Accelerated maturation of primate testis by xenografting into mice. Biol Reprod 2004;70:1500–3. 1010. McLean DJ. Spermatogonial stem cell transplantation and testicular function. Cell Tissue Res 2005;322:21– 31. 1011. Rathi R, Honaramooz A, Zeng W, Turner R, Dobrinski I. Germ cell development in equine testis tissue xenografted into mice. Reproduction 2006;131:1091–8. 1012. Turner RM, Rathi R, Zeng W, Dobrinski I. Xenografting of degenerate stallion testis onto a mouse host does not rescue the testicular degeneration phenotype. Anim Reprod Sci 2005;89:253–5. 1013. Turner RM, Rathia R, Zeng W, Honaramooz A, Dobrinski I. Xenografting to study testis function in stallions. Anim Reprod Sci 2006;94:161–4. 1014. Zeng W, Avelar GF, Rathi R, Franca LR, Dobrinski I. The length of the spermatogenic cycle is conserved in porcine and ovine testis xenografts. J Androl 2006;27:527–33.
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Chapter 98
Oxidative Stress in Sperm Barry A. Ball
Introduction Oxidative stress is a component of many disease processes, and spermatozoa are particularly sensitive to the effects of oxidation. The production of reactive oxygen species (ROS) may occur during oxidative metabolism or may result from specific molecular mechanisms designed to produce ROS, such as the oxidative burst of leukocytes. It is now clear that ROS play an important role in normal sperm function; however, an imbalance in either the production or degradation of ROS may have serious deleterious effects on sperm. During sperm storage by either cooling or cryopreservation, the deleterious effect of oxidative stress may be further increased in situations in which much of seminal plasma is removed from a semen sample because much of the antioxidant capacity in semen resides within seminal plasma.
Reactive oxygen species scavengers in equine semen The enzymatic scavengers for ROS in semen include catalase, superoxide dismutase (SOD), and glutathione peroxidase (GPx) (Figure 98.1). There are wide species variations in the relative importance of these scavengers in seminal plasma, although catalase, SOD, and GPx appear to be particularly abundant in equine seminal plasma.1, 2 Much of the specific activity of catalase appears to originate in the prostate gland whereas SOD and GPx appear to derive from the accessory sex glands as well as epididymal fluid and testis. Although these enzyme scavengers are abundant in seminal plasma, sperm, with their small amount of cytoplasm, have very limited ability to scavenge ROS. Therefore, most of these scavengers are provided by seminal plasma, which apparently functions to protect ejaculated equine sperm from the adverse effects of ROS. Thus, the removal of seminal plasma during semen processing
may increase the level of susceptibility of spermatozoa to oxidative stress. In addition to the enzyme scavengers, a number of other components of seminal plasma may act to remove ROS. In particular, low-molecular-weight components such as albumin, urate, taurine, hypotaurine, pyruvate, lactate, ascorbic acid, tocopherol, and ergothioniene may act as antioxidants.3–6 Although little research has been conducted concerning the role of low-molecular-weight antioxidants in equine seminal plasma, assessment of the total antioxidant capacity of seminal plasma in other species suggests that these components may constitute an important part of the antioxidant capacity of semen.7
Production of reactive oxygen species by equine sperm The superoxide anion (O2 − ) appears to be the primary ROS generated by equine sperm, and this short-lived free radical rapidly dismutates to form hydrogen peroxide (H2 O2 ).8, 9 Production of the superoxide anion by equine sperm is derived from either a sperm-specific NADPH oxidase (NOX5) present in the plasma membrane of the sperm head or from sperm mitochondria (Figure 98.2).10, 11 Because the superoxide anion has poor membrane permeability, it is likely that H2 O2 accounts for the major cytotoxic effect in sperm.12 Cryodamaged, non-viable, or morphologically abnormal sperm, particularly sperm with proximal cytoplasmic droplets or abnormalities of the midpiece, have an increased production of ROS.8 This increased generation of ROS is principally driven by electron leakage from the mitochondrial electron transport chain with subsequent reduction of molecular oxygen to form the superoxide anion.10 During cryopreservation, sperm mitochondria frequently demonstrate moderate to marked swelling of the midpiece, representing distension of mitochondria (Figure 98.3). In turn,
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1) O2 + e- → O2-. SOD 2) O2-. + O2-. + 2H + → H2O2 CAT 3) 2H2O2 → 2H2O + O2 GPx 4) 2GSH + H2O2 → GSSG + 2H2O Figure 98.1 Enzyme scavengers and removal of reactive oxygen species. Equation 1, oxygen is reduced in the presence of a free electron (e− ) to the superoxide anion (O2 − ·). Equation 2, the superoxide anion undergoes dismutation either spontaneously or in the presence of superoxide dismutase (SOD) to form hydrogen peroxide (H2 O2 ). Equation 3, hydrogen peroxide is catabolized by catalase (CAT) to form water and oxygen. Equation 4, In the presence of reduced glutathione (GSH), hydrogen peroxide is used to oxidize GSH to form the oxidized glutathione (GSSG) and water.
damage to sperm mitochondria may result in uncoupling of normal oxidative metabolism and generation of increased levels of ROS.
Effects of reactive oxygen species on equine sperm capacitation Although ROS clearly have detrimental effects on sperm function when present in excess, low-level generation of ROS has important functional effects in the mature sperm cell. Under physiological conditions, the membrane-associated NADPH oxidase,
Figure 98.3 Differential interference contrast micrograph of equine sperm after cryopreservation. Sperm midpiece demonstrates marked thickening (arrow) due to mitochondrial swelling.
NOX5, appears to be the likely mechanism for this generation of ROS.11 Low-level ROS generation by equine sperm is important in sperm capacitation, which is accompanied by a concomitant increase in tyrosine phosphorylation.13 During sperm capacitation in vitro there is also a detectable increase in production of the superoxide anion, which further confirms the role of ROS on sperm function.9 The ability of ROS to induce capacitation in equine sperm may also affect sperm function following cryopreservation. Subsequent to cryopreservation, sperm have an increased intracellular calcium concentration, an increased generation of ROS, and a reduced antioxidant capacity because of removal of seminal plasma, which could lead to premature capacitation of sperm. 14 This ‘cryocapacitation’ of sperm results in similar but not identical changes in sperm membrane, as detected during capacitation in vitro, and may be responsible for the reduced longevity of sperm typically noted after freezing and thawing.15
Adverse effects of reactive oxygen species on sperm function
Figure 98.2 Nitroblue tetrazolium (NBT) cytochemical analysis of equine sperm. Sperm were stained with NBT in the presence of 1 mM NADPH. The generation of reactive oxygen species (ROS) is demonstrated by the intense black staining over the sperm midpiece (white arrow) as well as the fainter staining over the rostral portion of the sperm head (black arrow) .
As noted above, H2 O2 is more stable and more membrane permeable than the superoxide anion and appears to be the most important ROS resulting in damage to equine sperm.12 Treatment of equine sperm with the xanthine/xanthine oxidase-generating system (which produces the superoxide anion) increases H2 O2 production and decreases sperm motility.12 The addition of the enzyme scavenger catalase (which catabolizes H2 O2 ), but not SOD (which catabolizes the superoxide anion), maintained normal motility secondary to this induced oxidative stress. The decrease in sperm motility associated with ROS occurs in the absence of any detectable decrease in plasma
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Figure 98.4 Photomicrograph showing a fluorescently labeled sperm nucleus demonstrating DNA fragmentation in the COMET assay. During electrophoresis, fragmented sperm DNA moves out of the sperm nucleus forming the diffuse comet ‘tail’. The greater the fragmentation, the larger the comet tail.
membrane integrity, acrosomal integrity, or mitochondrial membrane potential. Therefore, sperm motility appears to be a sensitive indicator of oxidative stress and may be one of the first parameters affected during oxidative stress.
Effects of reactive oxygen species and sperm storage on DNA integrity of equine sperm Oxidative stress is well characterized as a cause of DNA damage in a variety of cell types, including sperm. In equine sperm, DNA damage increases in a dose-dependent fashion with increasing concentrations of ROS.16 DNA fragmentation (Figure 98.4) was blocked in the presence of catalase or reduced glutathione (GSH) but not SOD, which suggested that H2 O2 and not superoxide anion was responsible for DNA damage in equine sperm.16 Interestingly, damage to sperm DNA was initiated at levels of oxidative stress comparable to those previously shown to adversely affect sperm motility. DNA fragmentation in equine sperm increases with both cooled storage17 and frozen storage.16 Unfortunately, the addition of selected antioxidants (vitamin E, reduced glutathione, vitamin C) or enzyme scavengers (catalase, SOD) to freezing extenders did not reduce DNA fragmentation during cryopreservation of equine sperm.18 In fact, the addition of SOD to freezing extenders significantly increased DNA fragmentation, suggesting again that H2 O2 is the primary ROS resulting in DNA damage in equine sperm. It is unclear why the addition of these antioxidants to equine sperm failed to reduce DNA fragmentation
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during cryopreservation, and future research should address this important topic. In human sperm, DNA damage secondary to oxidative stress increased before there were any measurable changes in sperm–oocyte fusion or alterations in sperm motility.19 Because low levels of oxidative stress may increase the capacitation state of sperm and also initiate DNA fragmentation, oxidative stress may allow sperm with DNA damage to fertilize the oocyte. Studies from other species indicate that, although fertilization may occur, the rate of subsequent embryonic development is reduced and the rate of early embryonic death is increased in situations in which fertilization is initiated by DNAdamaged sperm.20, 21 Similar data are not available for the horse; however, it appears likely that DNA damage subsequent to oxidative damage might also lead to reduced fertility and increased rates of early embryonic loss.
Effect of reactive oxygen species on membrane lipid peroxidation Mammalian sperm membranes are characterized by a relatively high concentration of polyunsaturated fatty acids,22 which are susceptible to peroxidative damage.23 Neither H2 O2 nor O2 − · are energetic enough to initiate lipid peroxidation, and a transition metal catalyst (such as Fe2+ ) is typically required to cause lipid peroxidation.24 Once initiated, lipid peroxidation can proceed in a chain reaction with the formation of lipid peroxides and cytotoxic aldehydes such as malondialdehyde and 4-hydroxynonenol.23 Once initiated, membrane lipid peroxidation alters membrane fluidity, which can affect the ability of sperm membranes to fuse, an important function during acrosomal exocytosis. The presence of ironbinding proteins, such as lactoferrin, in equine seminal plasma25 may play a role in reducing available iron to help prevent membrane lipid peroxidation. Equine spermatozoa appear to be more resistant to membrane lipid peroxidation than sperm of other domestic animals.12, 26 However, cryopreservation of equine sperm increased their susceptibility to lipid peroxidation, which was more pronounced over the region of the sperm midpiece.26 Likewise, storage of equine semen at 5◦ C resulted in a detectable increase in lipid peroxidation.24 The addition of vitamin E significantly reduced lipid peroxidation in equine sperm,24 and the vitamin E analog tocopherol succinate was superior to vitamin E in preventing lipid peroxidation.27 Unfortunately, the succinate ester of tocopherol suppressed motility of equine sperm to a greater extent than vitamin E alone, which precludes its practical application.
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Effects of antioxidants on storage of equine sperm To further evaluate the effects of antioxidants on sperm storage, catalase as well as lipid- and watersoluble antioxidants were added to skim-milk-based extenders during storage of equine sperm at 5◦ C.28 The addition of catalase did not improve maintenance of motility during 72-hour storage at 5◦ C. Likewise, lipid-soluble antioxidants butylated hydroxytoluene (BHT), vitamin E, or the nitroxide compound tempo were evaluated by addition to skim-milk extenders. The addition of BHT significantly reduced progressive motility during storage, and there were no positive treatment effects of either vitamin E or tempo on maintenance of sperm motility. In a third experiment, semen was diluted in skim-milk extender containing trolox, vitamin C, bovine serum albumin, or combinations of these antioxidants, and again no positive effects of treatment on the maintenance of sperm motility were detected. Aurich et al.29 observed a positive effect of addition of vitamin C but not catalase on preservation of membrane integrity of cooled equine sperm; however, there was no effect of addition of catalase under similar conditions.29 Likewise, the addition of N-acetylcysteine had no effect on maintenance of equine sperm motility or membrane integrity during cooled storage.30 Together, these studies indicate that the addition of catalase or a variety of lipid- or water-soluble antioxidants does not improve the maintenance of motility during short-term cooled storage of equine sperm in the presence of skim-milk-based extenders. Disappointingly, most studies that have examined the addition of antioxidants to cryopreserved equine sperm have also not shown an improvement in postthaw parameters or fertility. The addition of the enzyme scavengers catalase or SOD, or low-molecularweight antioxidants such as reduced glutathione, vitamin C, or vitamin E, did not improve DNA fragmentation, mitochondrial membrane potential, viability, or motility of frozen equine sperm after thawing.18 However, Aguero et al.31 reported a positive effect of addition of vitamin E to cryopreserved equine sperm. In cattle, Foote et al.32 examined a variety of antioxidants and combinations of antioxidants added to either liquid or frozen bovine sperm with limited, if any, positive effect on fertility. Interestingly, there were interactions between extender type (skim-milk-based vs egg yolk-based extenders) and the addition of antioxidants. Foote and co-workers32 suggested that the addition of casein and other milk proteins in extenders may provide abundant ROS scavenging capability and may, therefore, obviate the need for addition of lipid- or water-soluble antioxidants to semen extenders containing milk
products. Similarly, in studies with equine semen, the addition of skim-milk-based extenders increased the activity of GPx, SOD, and catalase compared with raw semen.33
Dietary supplementation with antioxidants Although the literature concerning the addition of vitamin E to semen extenders appears to demonstrate a variable response on maintenance of sperm function and fertility, several studies suggest that dietary supplementation of vitamin E may positively affect semen quality and maintenance of sperm during in vitro storage. Dietary supplementation of the male with vitamin E appears to be effective in limiting lipid peroxidation of sperm membranes in both turkeys and chickens.34 Furthermore, the addition of organic selenium to the diet also increases activity of selenium-dependent GPx in seminal plasma.34 Supplementation of the female with dietary vitamin E and organic selenium also appeared to improve fertility, perhaps through effects on sperm storage in the oviduct.34 In boars, dietary levels of vitamin E and selenium affected the percentage of motile, morphologically normal sperm present in the ejaculate as well as the fertilization rate in gilts mated.35 At present, two reports are available regarding dietary addition of antioxidants in the stallion and subsequent effects on semen quality or storage.36, 37 The addition of 3000 IU D-alpha-tocopherol daily for 14 weeks to the diet of stallions with low (< 35% progressively motile sperm) post-thaw motility did not improve motility parameters of sperm after freezing and thawing compared with untreated control stallions.37 However, there were significant improvements in the maintenance of sperm motility in cooled semen stored for 48 hours compared with controls. In a separate study, the dietary addition of antioxidants (tocopherol 300 mg/day; ascorbic acid 300 mg/day; l-carnitine 4000 mg/day; folic acid 12 mg/day) did not improve motility parameters of sperm stored for 24 hours. There was, however, a small, but significant, improvement in sperm morphological parameters in stallions fed the antioxidant diet.36 These studies suggest that there may be some benefit to the dietary addition of antioxidants in the stallion, although the effects are not clear cut.
References 1. Ball BA, Gravance CG, Medina V, Baumber J, Liu IKM. Catalase activity in equine semen. Am J Vet Res 2000;61: 1026–30. 2. Baumber J, Ball BA. Determination of glutathione peroxidase and superoxide dismutase-like activities in equine
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3.
4.
5.
6. 7.
8.
9.
10. 11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
spermatozoa, seminal plasma, and reproductive tissues. Am J Vet Res 2005;66:1415–19. Mann T, Minotakis CS, Polge C. Semen composition and metabolism in the stallion and jackass. J Reprod Fertil 1963; 5:109–22. Alvarez J G, Storey BT. Taurine, hypotaurine, epinephrine and albumin inhibit lipid peroxidation in rabbit spermatozoa and protect against loss of motility. Biol Reprod 1983; 29:548–55. De Lamirande E, Gagnon C. Reactive oxygen species and human spermatozoa. I. Effects on the motility of intact spermatozoa and on sperm axonemes. J Androl 1992;13:368–78. Halliwell B, Gutteridge JMC. Free Radicals in Biology and Medicine. New York, NY: Oxford University Press, 1999. Thiele JJ, Friesleben HJ, Fuchs J, Ochsendorf FR. Ascorbic acid and urate in human seminal plasma: determination and interrelationships with chemiluminescence in washed semen. Hum Reprod 1995;10:110–15. Ball BA, Vo AT, Baumber J. Generation of reactive oxygen species by equine spermatozoa. Am J Vet Res 2001;62: 508–15. Burnaugh L, Sabeur K, Ball BA. Generation of superoxide anion by equine spermatozoa as detected by dihydroethidium. Theriogenology 2007;67:580–9. Sabeur K, Ball BA. Detection of superoxide anion generation by equine spermatozoa. Am J Vet Res 2006;67:701–6. Sabeur K, Ball BA. Characterization of NADPH oxidase 5 in equine testis and spermatozoa. Reproduction 2007;134: 263–70. Baumber J, Ball BA, Gravance CG, Medina V, DaviesMorel MCG. The effect of reactive oxygen species on equine sperm motility, viability, acrosomal integrity, mitochondrial membrane potential and membrane lipid peroxidation. J Androl 2000;21:895–902. Baumber J, Sabeur K, Vo A, Ball BA. Reactive oxygen species promote tyrosine phosphorylation and capacitation in equine spermatozoa. Theriogenology 2003;60:1239–47. Neild DM, Gadella BM, Chaves MG, Miragaya MH, Colenbrander B, Aguero A. Membrane changes during different stages of a freeze-thaw protocol for equine semen cryopreservation. Theriogenology 2003;59:1693–705. Thomas AD, Meyers SA, Ball BA. Capacitation-like changes in equine spermatozoa following cryopreservation. Theriogenology 2006;65:1531–50. Baumber J, Ball BA, Linfor JJ, Meyers SA. Reactive oxygen species and cryopreservation promote DNA fragmentation in equine spermatozoa. J Androl 2003;24:621–8. Linfor JJ, Meyers SA. Detection of DNA damage in response to cooling injury in equine spermatozoa using single-cell gel electrophoresis. J Androl 2002;23:107–13. Baumber J, Ball BA, Linfor JJ. Assessment of the cryopreservation of equine spermatozoa in the presence of enzyme scavengers and antioxidants. Am J Vet Res 2005;66:772–9. Aitken RJ, Clarkson JS, Fishel S. Generation of reactive oxygen species, lipid peroxidation, and human sperm function. Biol Reprod 1989;41:183–97. Ahmadi Ali, Ng SC. Fertilizing ability of DNA-damaged spermatozoa. J Exp Zool 1999;284:696–704.
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21. Morris ID, Ilott S, Dixon L, Brison DR. The spectrum of DNA damage in human sperm assessed by single cell gel electrophoresis (Comet assay) and its relationship to fertilization and embryo development. Hum Reprod 2002;17: 990–8. 22. Parks JE, Lynch DV. Lipid composition and thermotropic phase behavior of boar, bull, stallion, and rooster sperm membranes. Cryobiology 1992;29:255–66. 23. Aitken RJ. Free radicals, lipid peroxidation and sperm function. Reprod Fertil Dev 1995;7:659–68. 24. Ball BA, Vo A. Detection of lipid peroxidation in equine spermatozoa based upon the lipophilic fluorescent dye C11-BODIPY581/591. J Androl 2002;23:259–69. 25. Inagaki M, Kikuchi M, Orino K, Ohnami Y, Watanabe K. Purification and quantification of lactoferrin in equine seminal plasma. J Vet Med Sci 2002;64:75–7. 26. Neild DM, Brouwers JFHM, Colenbrander B, Aguero A, Gadella BM. Lipid peroxide formation in relation to membrane stability of fresh and frozen thawed stallion spermatozoa. Mol Reprod Dev 2005;72:230–8. 27. Almeida J, Ball BA. Effect of [alpha]-tocopherol and tocopherol succinate on lipid peroxidation in equine spermatozoa. Anim Reprod Sci 2005;87:321–37. 28. Ball BA, Medina V, Gravance CG, Baumber J . Effect of antioxidants on preservation of motility, viability and acrosomal integrity of equine spermatozoa during storage at 5◦ C. Theriogenology 2001;56:577–89. 29. Aurich JE, Schonherr U, Hoppe H, Aurich C. Effects of antioxidants on motility and membrane integrity of chilledstored stallion semen. Theriogenology 1997;48:185–92. 30. Pagl R, Aurich C, Kankofer M. Anti-oxidative status and semen quality during cooled storage in stallions. J Vet Med Ser A Physiol Pathol Clin Med 2006;53:486–9. 31. Aguero A, Miragaya MH, Mora NG, Chaves MG, Neild DM, Beconi MT. Effect of vitamin E addition on equine sperm preservation. Comun Biol 1995;13:343–56. 32. Foote RH, Brockett CC, Kaproth MT. Motility and fertility of bull sperm in whole milk extender containing antioxidants. Anim Reprod Sci 2002;71:13–23. 33. Kankofer M, Kolm G, Aurich J, Aurich C. Activity of glutathione peroxidase, superoxide dismutase and catalase and lipid peroxidation intensity in stallion semen during storage at 5 [deg]C. Theriogenology 2005;63:1354–65. 34. Breque C, Surai P, Brillard JP. Roles of antioxidants on prolonged storage of avian spermatozoa in vivo and in vitro. Mol Reprod Dev 2003;66:314–23. 35. Marin-Guzman J, Mahan DC, Chung YK, Pate JL, Pope WF. Effects of dietary selenium and vitamin E on boar performance and tissue responses, semen quality, and subsequent fertilization rates in mature gilts. J Anim Sci 1997; 75:2994–3003. 36. Deichsel K, Palm F, Koblischke P, Budik S, Aurich C. Effect of a dietary antioxidant supplementation on semen quality in pony stallions. Theriogenology 2008;69:940–5. 37. Gee EK, Bruemmer JE, Siciliano PD, McCue PM, Squires EL. Effects of dietary vitamin E supplementation on spermatozoal quality in stallions with suboptimal post-thaw motility. Anim Reprod Sci 2008;107:324–5.
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Chapter 99
Endocrine–Paracrine– Autocrine Regulation of Reproductive Function in the Stallion Janet F. Roser
Introduction In most mammalian male species, including the stallion, normal reproductive function is dependent upon a dynamic and integrative hypothalamic– pituitary–testicular (HPT) axis involving classic endocrine hormones. Melatonin regulates the release of gonadotropin-releasing hormone (GnRH) from the hypothalamus; GnRH regulates the synthesis and release of luteinizing hormone (LH) and folliclestimulating hormone (FSH) from the anterior pituitary gland and LH and FSH regulate the synthesis and release of steroids, proteins and other local factors from the testis. Feedback effects on the hypothalamus and anterior pituitary are controlled by steroids such as testosterone and estrogen and proteins such as inhibin and activin.1–6 In the last decade it has become evident that other hormones such as prolactin (Prl), growth hormone (GH), thyroid hormones (T4 and T3 ), and opioids regulate testicular function by way of the HPT axis in the stallion.7–9 Local testicular factors called paracrine–autocrine factors such as growth factors, oxytocin, transferrin, and insulin-like peptide 3 (INSL3), appear to modulate testicular and epididymal cell function.8–12 The presence of these local factors and their dynamic interactions with the endocrine system are responsible for initiating and maintaining testicular and epididymal function. The integrative roles of the local paracrine–autocrine factors and the endocrine system have received close scientific scrutiny in other mam-
malian species, but remain largely undefined in the stallion. This review summarizes current information regarding the nature of the multiple endocrine– paracrine–autocrine systems that regulate reproductive function in the stallion including seasonal effects, puberty, and epididymal regulation.
Dynamics of the endocrine system (Figure 99.1) Melatonin, GnRH, LH, and FSH A regulator of reproductive function is photoinducable melatonin, which when secreted from the pineal gland during the non-breeding season suppresses hypothalamic GnRH production.13–15 During the breeding season when melatonin production is low, GnRH output is at its highest levels to promote optimum testicular function and sperm output by way of pituitary LH and FSH. The hypothalamus of the stallion releases GnRH in a pulsatile fashion, which, in turn, stimulates pulsatile release of the gonadotropins.16, 17 The frequency and amplitude of the pulses and the ratio of LH to FSH may contribute to the effectiveness of LH and FSH to stimulate testicular function in the stallion. LH binds to the membrane receptors on the equine testicular Leydig cells and stimulates the production and release of both testosterone (T) and estrogens.18 In contrast, and as demonstrated in the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Endocrine–Paracrine–Autocrine Regulation of Reproductive Function Brain
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–
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Hypothalmus: GnRH, GHRH, dopamine, TRH
Pituitary
T– ?? ?
E ±?
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E– ?
INH–
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Thyroid
T, E, INH
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Autocrine Paracrine
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Figure 99.1 Hypothalamic–pituitary–testicular axis of the stallion (HPT). Melatonin from the pineal gland and opioids from the brain regulate gonadotropin-releasing hormone (GnRH) in a negative fashion. GnRH regulates the release of luteinizing hormone (LH) and follicle-stimulating hormone (FSH) from the anterior pituitary. The major regulator of Leydig cells is LH. The major endocrine products of the Leydig cells are testosterone (T) and estrogens (E) in the adult stallion. The major regulator of the Sertoli cells is FSH. Estrogen, inhibin (INH), and probably activin (ACT) in the stallion are the major endocrine products of the Sertoli cells. Growth hormone-releasing hormone (GHRH) regulates production and secretion of growth hormone (GH) from the anterior pituitary. Dopamine negatively regulates the release of prolactin (PRL) from the pituitary. Thyroid-releasing hormone (TRH) produced in the hypothalamus stimulates thyroid stimulating hormone (TSH) from the anterior pituitary which thereafter regulates the release of thyroid hormones (T3 and T4) from the thyroid gland. The role of growth hormone, prolactin, and thyroid hormone on testicular function in the stallion are unclear. Testosterone inhibits GnRH but has no action on LH release at the level of the pituitary. Its role in regulating pituitary FSH is unclear. Estrogen modulates GnRH-induced LH release, but its action on FSH release at the pituitary and GnRH at the hypothalamus is unclear. Inhibin feeds back on the pituitary to inhibit the release of FSH. Activin has been shown to positively modulate the release of pituitary FSH in other species, but its action in the stallion is unknown.9 With permission from Elsevier.
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linked oligosaccharide and one O-linked carbohydrate moiety.25 Equine LH is the only LH that possesses an O-linked glycosylated C-terminal peptide (CTP) consisting of an extra 28 residues previously believed to be restricted to chorionic gonadotropins. 25 Although the alpha subunit of equine FSH (eFSH) is similar to the LH alpha, the beta subunit is composed of 111 amino acid residues that are different than the beta subunit of eLH. FSH beta has two Nlinked oligosaccharide and no O-linked CTP.25 The beta subunit of LH and FSH defines the biological specificity of the whole molecule.26 The horse pituitaries produce a number of gonadotropin isoforms with varying degrees of sialic acid, which increases the half-life of these hormones.27 Testosterone Testicular steroid hormones feed back on the hypothalamus and pituitary via the peripheral circulation to modulate the synthesis and secretion of GnRH, LH, and FSH.1, 2, 9 Muyan and co-workers28 clearly demonstrated that estrogens, particularly estradiol-17β (E2 ), and not the androgens, modulate LH release from cultured stallion pituitary cells. The results demonstrated that estradiol significantly enhanced the GnRH-induced LH release from stallion pituitary cells, whereas testosterone and dihydrotestosterone (DHT) had no effect. The major role of testosterone in other male species appears to be a negative feedback effect on the release of GnRH,29 which may be the case in the stallion. Thompson and co-workers30 reported a negative effect of testosterone on serum LH levels and pituitary LH content but no effect on the GnRH-induced release of LH in geldings, suggesting that the feedback effects of testosterone in the stallion occurs at the hypothalamus and the pituitary.30 Estrogen
stallion and other species, FSH binds to Sertoli cells, which stimulate production of estrogens, inhibin, activin, androgen-binding protein (ABP), transferrin, insulin-like growth factor-1 (IGF-1), and other factors needed for spermatogenesis.2,19–21 Melatonin is produced in the pinealocytes of the pineal gland by enzymatic conversion of serotonin to N-acetyl-5-methoxytryptamine.22 GnRH is a neuro-decapeptide synthesized in the hypothalamus. 23 Pituitary LH and FSH are glycoproteins that have a heterodimeric structure in which an alpha subunit, common to the LH and FSH, is non-covalently joined to the beta subunit by disulfide bonds.24 The equine alpha subunit is composed of 96 amino acid residues and contains two N-linked oligosaccharides. The beta subunit contains 114 residues with one N-
The stallion testes produce a large amount of estrogen compared with most other mammalian male species. 31 Although it has been shown to have both negative and positive feedback effects on the hypothalamus in other species,29, 32 its mechanism of action at the hypothalamus in the stallion is yet to be revealed. Thompson and co-workers30 demonstrated that treatment with estradiol decreased circulating FSH levels, increased circulating LH levels, and decreased GnRH-induced FSH secretion in long-term geldings. Thompson and Honey33 further demonstrated a role of estradiol on FSH secretion when they reported that immunization of colts against estradiol increased circulating FSH levels without changing LH levels. These studies do not identify the site of action of estradiol, however. Estradiol could be
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acting at the hypothalamus to change the amplitude and frequency of GnRH and thus the ratio of LH and FSH secreted from the pituitary as observed in the rat and primate,34, 35 or estradiol could be acting at the level of the pituitary to modulate GnRH-induced LH and FSH release as observed in other species, including the horse.28, 32, 36 Inhibin Inhibin is a member of the transforming growth factor-beta super family.37 It consists of glycosylated polypeptide dimers involving subunits linked by a disulfide bond. In the case of inhibin, the subunits consist of an alpha (α) subunit linked to a beta (β) subunit; either β A or β B . The subunits have been localized in both the Leydig and Sertoli cells by immunocytochemistry in several species, including the horse.37–39 Because inhibin inhibits synthesis of FSH at the level of the pituitary in several mammalian species, including the horse,38–40 inhibin may act in conjunction with estrogen or testosterone in the stallion to control FSH secretion. The FSH suppression hypothesis is supported by the fact that in the mare immunization against inhibin is followed by an increase in FSH levels.40 Although it has been demonstrated that circulating FSH levels increase after immunization against inhibin,41 there have been no such studies carried out in the stallion. The supposition that inhibin is important for testicular recrudescence in the stallion is supported by the fact that both FSH and inhibin exhibit similar seasonal changes; peripheral FSH and inhibin increase in the spring and decrease in the fall.39, 42 The role of inhibin as a paracrine–autocrine factor in the testes of other mammalian male species also suggests that it may be important in spermatogenesis (see Testicular inhibin and activin). Activin Activin is also a member of the transforming growth factor-beta super family.37 Activin consists of two beta subunits linked by a disulfide bond: β A β A, β B β B or β A β B . Not much is known about the role of activin in the stallion. In other species, activin appears to be stimulated by FSH from Sertoli cells and to act at the level of the pituitary to release FSH.38 Expression of inhibin/activin β A subunit mRNA activity has been demonstrated in the endometrial glands of the mare during pregnancy.43 One could surmise that activin is also localized in the testicular somatic cells of the stallion given the fact that activins, like inhibins, consist of beta subunits that have been localized in equine Leydig and Sertoli cells.39 Studies investigating activin activity at the level of the hypothalamus, pituitary, and testes in the stallion have not been done.
Prolactin, thyroid hormone, and growth hormone
Prolactin Prolactin is a single-chain protein hormone synthesized by the lactotrophs in the anterior pituitary and plays a role in reproductive function in the male.44, 45 It modulates testicular function in the boar,46 ram,47 rodent,48, 49 and human.49 In other species, prolactin plays a role in inducing transcription of the testicular estrogen receptor.50 It has been shown to affect androgen-sensitive tissues and, together with LH and growth hormone, controls synthesis of LH receptors in the testes, activates androgen synthesis, and affects spermatogenesis.46–48 The role of prolactin on reproductive function in the stallion is yet to be determined. Prolactin fluctuated with the season in the stallion and mare.51, 52 Moreover, prolactin secretion tended to increase with age in the stallion.51 Prolactin did not stimulate testosterone or estrogen production in equine Leydig cells in culture.18
Thyroid hormone The thyroid hormones [thyroxine (T4 ) and triiodothyronine (T3 )] are tyrosine-based hormones produced by the thyroid gland.53 These hormones are crucial to the onset of adult Leydig cell and Sertoli cell differentiation.54, 55 Hypothyroidism during testicular development leads to proliferation of the testicular somatic cells and bigger size testes.54 One critical molecular target of thyroid hormone in the Leydig cell may be the steroidogenic acute regulatory (StAR) protein, which is responsible for cholesterol transport across the outer mitochondrial membrane in Leydig cells.55 A lack of thyroid hormone leads to a decrease in StAR mRNA and protein, causing a decrease in testosterone production.55 In Sertoli cells, thyroid hormone stimulates lactate secretion as well as mRNA expression of mullerian inhibiting substance, aromatase, estradiol receptor, and androgen binding protein.56 Thyroid hormone receptors have been identified in sperm, developing germ cells, and Sertoli, Leydig, and peritubular myoid cells.56 The seasonal changes and effects of thyroid hormone have not been investigated in the stallion. In the mare, thyroid hormone fluctuates with season,52 but thyroid function was not associated with level of fertility.57
Growth hormone Growth hormone is a protein hormone of about 190 amino acids that is synthesized and secreted by cells called somatotrophs in the anterior pituitary.58
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Endocrine–Paracrine–Autocrine Regulation of Reproductive Function Growth hormone affects testicular function by modulating gonadal steroid synthesis and gametogenesis.59 It has been demonstrated that GH mediates the release of pituitary LH as well as the synthesis and release of testicular IGF-1.60 GH deficiency is associated with abnormally small testes61 and reduced or absent sperm motility.62 Treatment with GH can increase plasma IGF-1 concentrations and restore sperm motility.62 Growth hormone resistance is associated with reduced fertility in men63 and may alter gametogenesis by affecting testosterone synthesis. Testosterone is necessary for sperm production and mRNA encoding GH receptors in rat Leydig cells.64 GH increases basal or hCG-stimulated testosterone production in GH-deficient men.65 Growth hormone does not appear to fluctuate with season in the stallion.66 However, treatment with a somatostatin analog (GH inhibitor) caused a transient decrease in semen motility and hCGinduced testosterone release, suggesting a role for GH in the regulation of testicular function in stallions.66 A direct effect of growth hormone on Leydig cell steroidogenesis in culture was not observed.67 Opioids In the horse as well as other species, it appears that seasonal changes in GnRH/LH release are partly regulated by opioids.7 Opioids act as neurotransmitters that modulate hypothalamic function by suppressing GnRH release during the winter months.7 Aurich and co-workers68 showed that treatment with the opioid antagonist naloxone caused an acute LH release in stallions outside of, but not during, the breeding sea-
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son. The opioidergic regulation of LH release appears to require the presence of the gonads.68
Dynamics of the testicular paracrine–autocrine system (Figure 99.2) In addition to the endocrine control on testicular function and sperm production, local testicular steroids, proteins, and peptides called paracrine– autocrine factors coordinate the various functions of the different testicular cell types and/or modulate the testicular actions of endocrine factors according to local conditions and requirements (Figure 99.2). Paracrine factors are produced by one cell and act on a different cell type, whereas autocrine factors are produced by one cell and act back on the same cell type. Potential paracrine–autocrine regulators of testicular function and spermatogenesis of proven physiological significance include such factors as estrogen, testosterone, inhibin, activin, GnRH-like peptides, growth factors like IGF-1, and transforming growth factor alpha and beta (TGFα and -β), oxytocin, vasopressin, proenkephalins, enkephalins, propiomelanocortin (POMC), and POMC-derived peptides such as β-endorphins. In addition, cytokines, transferrin, sulfated glycoprotein-l (sGP-l) and sGP2, insulin-like peptide 3 (INSL3) and Pmods, a paracrine factor secreted from peritubular myoid cells that modulates Sertoli cell function, may also play a paracrine–autocrine role in the testis.3, 10, 11,69–81 It is beyond the scope of this chapter to discuss the roles of each of these local factors, so only those that have been investigated in the stallion will be
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Figure 99.2 Paracrine–autocrine regulation of testicular function. Many of the local interactions in the equine testis are unknown. The figure presents a hypothetical paracrine–autocrine system based on what has been observed in other species: gonadotropinreleasing hormone (GnRH), insulin-like growth factor 1 (IGF-1), insulin-like peptide 3 (INSL3), inhibin (INH), activin (ACT), betaendorphin (β-EP), transferrin (Trans), basic fibroblast growth factor (bFGF), transforming growth factor (TGF), transforming growth factor beta (TGF-β), transforming growth factor alpha (TGF-α), interleukin-1 (IL-1), sulfated glycoprotein (SGP), nerve growth factor (NGF), peritubular modifying substance (PmodS), oxytocin (OT), vasopressin (VP), testosterone (T), estradiol (E2 ). Those factors highlighted have been identified in the stallion testis. (Modified from Roser.9 ) With permission from Elsevier.
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presented: IGF-1, transferrin, inhibin, activin, estrogen, testosterone, INSL3, POMC, and β-endorphins and oxytocin.82 Testicular IGF-1 A publication by Gnessi and co-workers73 provides an excellent overview of the expression, localization, production, and testicular function of IGF-1 in mammalian species. IGF-1, a single-chain polypeptide, is present in peritubular myoid cells, germ cells, Sertoli cells, and Leydig cells. Receptors for IGF-1 have been found on the cell membrane of germ cells, Leydig and Sertoli cells, and myoid cells. IGF-1 receptors can be upregulated by LH, FSH, and GH, with LH being the most important factor. The actions of IGF-1 can be modified by its binding protein (IGFBP), which may enhance or inhibit the effects of IGF-1. IGF-1 stimulates the proliferation of pre-pubertal Sertoli cells and has a small mitogenic effect on immature Leydig cells. IGF-1 stimulates testosterone production and potentiates hCG-induced testosterone formation. Furthermore, IGF-1 is believed to be involved in spermatogenesis. It stimulates spermatogonial DNA synthesis and has a maintaining effect on premeiotic DNA synthesis in the rat under in vitro conditions, and it induces the differentiation of mouse type A spermatogonia. More recently, IGF-1 has been shown to have anti-apoptotic effects on Leydig cells and germ cells.83, 84 Testicular IGF-1 in the stallion testes IGF-1 has been identified in testicular tissue and in seminal fluid in the stallion.85, 86 Hess and Roser85 reported that in colts less than 2 years old, IGF-1 concentrations in plasma and testicular extracts were higher than in the other age groups and were higher in the breeding season than in the non-breeding season. Seasonal changes in IGF-1 were not observed in stallions older than 2 years. The results of this study demonstrate that plasma and testicular IGF-1 levels are higher in developing colts than in older stallions, suggesting that IGF-1 may be involved in testicular development. IGF-1 alone does not appear to stimulate a steroidogenic response from equine Leydig cells in culture.67 Circulating IGF-I and IGF-1 in testicular extracts did not appear to correlate with declining fertility in stallions tested, suggesting that IGF-1 may not be a reliable biomarker for the diagnosis of subfertility and infertility using plasma or testicular extracts.85 Preliminary studies using a radioimmunoassay (RIA) indicate that both equine Leydig cells and Sertoli cells in culture produce IGF-1.5 Basal secretion of IGF-1 levels in the Sertoli cell culture medium was highest in pre-pubertal colts, whereas basal IGF-1
concentrations in the Leydig cell culture medium were highest in peri-pubertal animals. The basal levels of IGF-1 in Sertoli cell culture medium were dramatically lower than in the culture medium of Leydig cells.5 Although LH did not regulate IGF-1 in Leydig cells, FSH stimulated IGF-1 from Sertoli cells but only prior to puberty.5 Recently, using immunocytochemistry (ICC), there is evidence in our laboratory that IGF-1 is localized in Leydig cells and germ cells but not in Sertoli cells. The dichotomy of these two studies regarding Sertoli production of IGF-1 suggests that IGF-1 may not have been localized using ICC in the Sertoli cells owing to the very low basal concentrations produced. In addition, two different primary antibodies were used: one in an RIA to measure IGF-I in culture medium and another one for ICC. The antibody in the RIA could have been more sensitive than the antibody in the ICC procedure. It would be interesting to preculture the Sertoli cells with eFSH to increase production of IGF-1 and then run ICC on the cells after treatment. There is some evidence to suggest that measurement of IGF-1 in seminal plasma from stallions may be a biomarker for fertility.86 A positive but variable relationship between IGF-1 and sperm morphology and motility was noted. Stallions with high levels of IGF-1 in seminal plasma achieved higher pregnancy rates in mares, suggesting that IGF-1 may play a role in sperm function. Preliminary studies in our laboratory indicate that IGF-1 may have anti-apoptotic effects on equine Leydig and Sertoli cells and stimulate steroidogenesis from Leydig cells when incubated with equine LH. Testicular transferrin An excellent review by Sylvester and Griswold87 discusses the physiology of transferrin in the mammalian testes. Transferrin is an iron-binding protein that delivers iron as a nutritional component to germ cells. Sertoli cells synthesize transferrin and, when bound to iron, secrete the complex into the adluminal space. The adluminal testicular iron-bound transferrin is then available to receptors on spermatocytes. Histological examination of testes from mutant mice lacking normal transferrin revealed a decreased number of germ cells and a greatly reduced level of spermiation.87 Transferrin has been identified in human seminal plasma and evidence indicates that the measurement of seminal fluid transferrin may be an indicator of the function of Sertoli cells in intact human males.88 In bulls there was a strong positive correlation between sperm output and transferrin concentrations in seminal fluid, suggesting that transferrin production should be a good indicator of Sertoli function and spermatogenic potential in bulls.89
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Endocrine–Paracrine–Autocrine Regulation of Reproductive Function Testicular transferrin in the stallion testes In the stallion, preliminary studies indicate that transferrin is produced by Sertoli cells, with the highest production rate occurring at the time of puberty.21, 90 A preliminary study suggested that seminal plasma transferrin concentrations are not correlated with various degrees of fertility in the stallion, but more work in this area is warranted.91 Testicular androgens and estrogens In other species, it has been demonstrated that androgens and estrogens regulate the function of Sertoli cells, Leydig cells, peritubular myoid cells, and germ cells by binding to their respective receptors and eliciting a cellular response.74, 79,92–95
Testicular androgens Testicular androgens induce their effect by way of their receptors in Sertoli and Leydig cells.93, 96, 97 By acting through its receptors in Sertoli cells, testosterone appears to be responsible for maintaining an adequate blood–testis barrier function98 and inducing meiosis and post-meiotic development of germ cells.74, 94 Testosterone has been shown to inhibit germ cell apoptosis.99 There also appears to be an androgen action on Sertoli cells that modulates their gene expression, proliferation, and differentiation.74, 94 Studies using a tissue-specific knockout mouse with the androgen receptor gene deleted demonstrated an alteration in the expression of several key steroidogenic enzymes in Leydig cells, suggesting that testosterone is an autocrine factor regulating its own production.100 The androgen receptor knockout mouse also demonstrated an arrest of spermatogenesis predominately at the round spermatid stage.100
Testicular estrogen Increasing evidence over the last decade has clearly established the importance of estrogen as a paracrine–autocrine factor in regulating testicular function.78, 79, 101 The production of estrogen comes from the enzymatic conversion of androgens to estrogens by the aromatase enzyme. According to the literature reviewed by O’Donnell and co-workers79 and Hess and Carnes101 the production of estrogen from the different testicular cell types changes with age and species. For example, during the early stages of testicular development in the rat, basal aromatase activity is found in both the immature Leydig and Sertoli cells, with the Sertoli cells being more active in producing estrogen than the Leydig cells. However,
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aromatase appears to have an age-dependent pattern of expression in the rat. As the animal matures, the Leydig cell basal aromatase activity increases threeto fourfold, whereas the aromatase activity in the Sertoli cell remains or declines with age. In other species, it appears that the amount of aromatase activity is equally present in both the adult Sertoli and Leydig cells. Of interest is that germ cells express aromatase activity during different stages of development and into maturity, which suggests that sperm themselves could control the levels of estrogen present in the luminal fluid and epididymis. The localization of estrogen receptors in testicular cells varies depending on the species, developmental stage of the cell, and type of receptor.78, 79, 101 Although two different receptors have been identified, estrogen receptor alpha (ERα) and estrogen receptor beta (ERβ), the differences in their mechanisms of action are still not clear except that certain cell types are regulated by one receptor or the other. In general, according to O’Donnell and co-workers,79 it appears that both ERα and ERβ are localized in Leydig cells, whereas ERβ is mostly confined to Sertoli cells and developing germ cells. Testicular testosterone and estrogen in the stallion testes Changes in intratesticular testosterone and estrogen concentrations during testicular development and stages of fertility have been investigated in the stallion.102 Intratesticular concentrations of estradiol and estrogen conjugates increased with testicular maturation whereas there was no age effect on testosterone concentrations. There was no difference detected in intratesticular concentrations of estradiol, estrogen conjugates, and testosterone among the fertile, subfertile, and infertile groups of stallions although plasma concentrations of the estrogens but not testosterone were significantly lower in the infertile group. By using in vitro equine testicular cell culture techniques, studies have demonstrated that increasing doses of equine LH stimulate the production of testosterone and estrogens from Leydig cells and increasing doses of FSH influence the production of estrogen from Sertoli cells.18, 20 By using ICC techniques it has been demonstrated that aromatase expression changes with age in the stallion such that both Leydig cells and Sertoli cells stain positive for aromatase prior to and during puberty, but in the adult stallion positive staining remains in Leydig cells and to a lesser degree in the Sertoli cells.103, 104 In the stallion testis, estrogen and androgen receptors have been localized in equine Leydig, Sertoli, myoid, or germ cells depending on the age of the animal.93, 105 The stallion makes more estrogens than androgens in the testis, as demonstrated by using blood
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samples obtained by catheterization of the testicular vein.31 Taken together, the studies above demonstrate an important regulatory role of estrogens and testosterone in testicular cell function and spermatogenesis in the stallion. Testicular inhibin and activin de Kretser and co-authors106 have written an extensive review of inhibin and activin. Inhibin and activin are glycoproteins of the transforming growth factor superfamily produced mainly by the Sertoli cells, but subunits have been localized in Leydig and germ cells depending on the stage of testicular development and mammalian species. Both proteins have endocrine, paracrine, and autocrine functions. As discussed above, inhibin and activin are released into the peripheral circulation and regulate pituitary FSH. In the testis, inhibin and activin receptors, coreceptors, and intracellular signaling molecules have been identified in the various somatic cell types and the germ cells.81 Evidence indicates that inhibin and activin act as paracrine factors influencing Leydig cell steroidogenesis107–109 as well as spermatogenesis.110 The paracrine role of activin appears to be an inhibitory one on steroidogenesis in Leydig cells.109 Activin also appears to have an autocrine role in stimulating follistatin and inhibin from Sertoli cells.106 In rat testes activin has age-dependent effects on Sertoli cell division and affects the development of germ cells in the male.111, 112 In human males, measurement of inhibin B levels in the peripheral circulation appears to be useful as an indicator of functional spermatogenesis and as a marker for male fertility.113 Testicular inhibin and activin in the stallion testes A paracrine–autocrine role of inhibin and activin, or the presence of their receptors, has not been established in the stallion testes. It has been established that inhibin is present in equine testicular tissue, and its subunits are predominantly located in both the Leydig and Sertoli cells.39, 102 The observation that intratesticular inhibin concentrations were significantly lower in colts less than 1 year of age than in pubertal and adult stallions102 suggests that inhibin and possibly activin play a local role in testicular development. Changes in hormone levels based on fertility status were different between plasma concentrations and intratesticular concentrations in stallions. In plasma, inhibin, estradiol and estrogen conjugates were significantly lower and testosterone higher in idiopathic infertile stallions but no change was observed in idiopathic subfertile stallions compared with fertile stallions. In contrast, intratesticular steroid concentra-
tions did not change among fertility groups. However, intratesticular levels of inhibin tended to be lower in subfertile stallions and were significantly lower in infertile stallions, suggesting that intratesticular inhibin may be an early marker for declining fertility in stallions. This phenomenon is similar to what has been observed in the peripheral circulation of human males. As previously mentioned, plasma concentrations of inhibin B were significantly lower in men with fertility problems, suggesting that inhibin may be a good biomarker for declining fertility.113 Testicular insulin-like peptide 3 (INSL3) The relaxin-like peptide hormone INSL3 (previously known as relaxin-like factor, RLF) is a circulating hormone in the male that has recently been identified.114 It is produced almost exclusively by the Leydig cells of the testis, with anorchid men having undetectable circulating levels of the hormone.115 INSL3 is made by both fetal and adult-type populations of Leydig cells, but only when these have attained their mature phenotype.116 Various studies have indicated that the Insl3 gene and protein (INSL3) are expressed constitutively once Leydig cells are mature, thus making INSL3 an excellent marker of Leydig cell differentiation status.116, 117 It has been used in this context to study changes in testicular function in seasonally breeding animals 118, 119 and during aging120 as well as to identify unequivocally Leydig cells within testis sections and in primary culture.121 Although INSL3 appears to be constitutively expressed in adult species, a recent finding using male patients with hypogonadotropic hypogonadism clearly showed that INSL3 production and secretion was dependent on LH.115 Currently, little is known about the function of INSL3 in the testis. INSL3 produced by prenatal Leydig cells is essential for the transabdominal part of testis descent via stimulation of outgrowth and differentiation of the gubernaculum, as shown in rodents.122, 123 Disruption of the Insl3 gene causes bilateral cryptorchidism.122, 123 The receptor for INSL3 has been recently identified as the novel G-proteincoupled receptor (GPCR) LGR8, also known as Great. 124, 125 Studies have demonstrated that the observed expression of LGR8 transcripts in the gubernaculum and the INSL3 stimulation of cAMP production by these cells were consistent with the common cryptorchid phenotypes of this ligand–receptor pair in earlier transgenic mouse studies.124, 125 Recently it was demonstrated that LGR8 is expressed in meiotic and particularly post-meiotic germ cells and in Leydig cells, though not in Sertoli or peritubular myoid cells.114 The recent finding of LGR8 expression in germ cells and the suppression of apoptosis through
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binding of INSL3 to LGR8 suggest a paracrine role of INSL3 in germ cell survival.126 Human male patients with testicular disorders often have reduced or absent circulating levels of INSL3.115
peptides. Studies indicate that POMC-derived peptides have differential effects on the function of the Sertoli cells. ACTH and MSH-like peptides are stimulatory while β-endorphin is inhibitory.
Testicular INSL3 in the stallion testes
Testicular POMC and β-endorphin in the stallion testes
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Recent studies by Klonisch and co-workers have identified INSL3 and its receptor LGR8 in the testis of the horse. Equine INSL3 (eINSL3) was identified as a marker of Leydig cells in pre- and post-pubertal equine testis. The localization of eInsl3 gene transcripts in Leydig cells was independent of the location of the testis in the animal, whether it is found in the abdomen or scrotum. By contrast, eInsl3 gene activity in testicular Leydig cells appeared to be reduced in cryptorchid testes, despite the presence of multiple Leydig cells. No INSL3 or LGR8 mutations have so far been identified in cryptorchid stallions. Interestingly, however, it was demonstrated that, in unilateral cryptorchid stallions, INSL3 expression is downregulated and LGR8 expression upregulated in the cryptorchid versus the descended testis.127 It is still a matter of debate whether unilateral cryptorchidism significantly affects stallion fertility. However, in artificial insemination (AI) stallions, the 50% reduction in sperm production may become relevant if the stallion is used frequently or is a marginal breeder. In the equine testis, Klonisch and coworkers127 also detected eINSL3 expression despite marked differences in the functional state of the seminiferous epithelial cell cycle. However, the quantitative differences they observed in post-pubertal normal compared with cryptorchid and pre-pubertal testes suggest regulatory mechanisms that affect INSL3 production in testicular Leydig cells. Klonisch and co-workers127 also demonstrated the expression of eLgr8 gene transcripts in the equine testis indicating that the testes were a target tissue for the actions of equine INSL3. Preliminary observations in our laboratory demonstrate an increase in basal production of INSL3 in equine Leydig cells cultured for 24 hours and after treatment with eLH. Further studies are ongoing. Testicular propiomelanocortin (POMC) and β-endorphin POMC is a precursor protein that contains the sequence for several bioactive peptides including adrenocorticotropin (ACTH), β-endorphin, and melanocyte-stimulating hormone (MSH). A review by Bardin and co-workers128 indicates that the Leydig cells may be the only testicular cell that produces POMC-derived peptides and that there is evidence that LH may be involved in the production of these
Soverchia and co-workers129 investigated the presence of POMC and β-endorphin in equine testicular and epididymal tissue. POMC gene expression was not found in either testicular or epididymal tissue. However, by using ICC techniques, they demonstrated the presence of β-endorphin in those tissues. Testicular oxytocin Oxytocin is recognized as having endocrine and paracrine roles in male reproduction. Oxytocin and its receptor have been localized in the testes of several mammalian species.130 Oxytocin appears to be produced and secreted by Leydig cells and its role is by way of receptors on Sertoli cells. Evidence supports the role of oxytocin on the contractility of the peritubular myoid cells to eject sperm into the lumen of the tubule and on spermiation.130 Oxytocin may have an autocrine role in modulating Leydig cell steroidogenesis.130 Testicular oxytocin in the stallion testes The role of oxytocin in the stallion has been investigated by Watson and co-workers131 using RIAs and they detected oxytocin in seminal fractions, washed lysed sperm, and in extracts from the testis and epididymis. By using ICC, immunostaining for oxytocin was present in occasional interstitial cells in the testis and in the epididymal epithelium and smooth muscle. The authors concluded that the staining was due to binding of oxytocin to its receptor in these tissues, suggesting that oxytocin is not locally produced in the testis or epididymis of the stallion.
Seasonal effects on the HPT axis in the stallion (Figure 99.3) Historically, the stallion has been classified as a ‘longday breeder’ because maximum reproductive capacity is attained during periods of increasing day length. During the spring and summer, there is an increase in testicular size and weight, sperm production and output, libido, and plasma concentrations of LH, FSH, testosterone, E2 , inhibin, and prolactin.14, 39, 52,132–136 (Figure 99.3). The endocrine events involved in initiating testicular recrudescence in the
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Figure 99.3 Seasonal effects on circulating luteinizing hormone (LH), follicle-stimulating hormone (FSH), testosterone (T), estradiol (E2 ), and inhibin (INH) concentrations. Monthly blood collections. Hormone profiles demonstrate a rise in all hormones during the breeding season and fall during the non-breeding season. (Data from Nagata et al.39 )
stallion have not been adequately characterized but appear to start with a photoinducible decrease in melatonin from the pineal gland, which, in turn, allows for an increase in hypothalamic GnRH and a subsequent increase in pituitary LH and FSH release into the peripheral circulation, albeit reports on seasonal or photoinducible changes in FSH release are conflicting.13, 14,135–139 The increase in plasma concentrations of gonadotropins and perhaps prolactin and thyroid hormone act on the testicular Leydig and Sertoli cells to increase production of T, E2 , and inhibin and spermatogenesis in the stallion. In the stallion as well as other species, it appears that seasonal changes in GnRH/LH release are partly regulated by opioids.7 Although it is yet to be demonstrated that prolactin plays a role in testicular function in the stallion, dopaminergic inhibition of prolactin secretion undergoes seasonal changes.140 Circulating inhibin concentrations are lower in the non-breeding season and increase during testicular recrudescence,39 which is consistent with reports that the number of Sertoli cells and spermatogonia increase with recrudescence.138 There are very few published studies on the seasonal effects on testicular paracrine–autocrine factors in male species, including the stallion. Since testosterone, estrogen, and inhibin are both endocrine and paracrine–autocrine factors, it would not be surprising to find that these hormones induce local changes in testicular function and spermatogenesis during testicular recrudescence. Further involvement
of other local testicular factors or changes in localization of testicular factors between the breeding and non-breeding season most likely occur in the stallion testis. In the roe deer testis, local growth factors such as IGF-I, IGF-II, and basic fibroblast growth factor (bFGF) were maximally transcribed during testis recrudescence in spring.141 The cellular localization of inhibin/activin subunits in the male raccoon dog and the male ground squirrel showed seasonrelated changes in the breeding and non-breeding seasons.142, 143 Although the hypothalamic–pituitary–testicular axis of the stallion is stimulated by photoperiod in a similar fashion as the hypothalamic– pituitary–ovarian (HPO) axis of the mare,139 it is unclear why ∼75% of mares shut down completely while stallions have the capacity to continue to breed throughout the non-breeding season, albeit with lower sperm numbers per ejaculate.144 Photoperiod does affect gonadotropin secretion in the male horse, and these effects appear to be gonadal dependent given the patterns of gonadal steroids and gonadotropins. However, distinct seasonal changes in LH and FSH were detected in long-term geldings, suggesting that, in part, seasonal changes are gonadal-independent.145 Perhaps from an evolutionary standpoint, the stallion’s HPT axis is probably less sensitive to changes in day length in order to ensure maximum breeding efficiency throughout the year. More research in this area is warranted.
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Endocrine–Paracrine–Autocrine Regulation of Reproductive Function A comprehensive 2-year study by Clay and Clay136 on the effects of season and photoperiod on endocrine and testicular function in stallions has revealed that the stallion, like other mammalian species, has an endogenous circannual cycle associated with testicular function that is entrained by an external environmental rhythm or zeitgeber (timegiver) identified as photoperiod.136 In this study, stallions exposed to 8 hours of light and 16 hours of dark (8:16) for 20 months starting in July continued to display a significant seasonal cycle in testicular size, sperm output, and peripheral hormone levels similar to the control group exposed to naturally occurring changes in day length. Thus, stallions maintained on continuous short days have the capacity to recrudesce despite the lack of exposure to a stimulatory photoperiod. On the other hand, testicular recrudescence was accelerated earlier in the season compared with control subjects when stallions were exposed to an 8:16 photoperiod for 20 weeks beginning in July and then 16:8 for 15 months thereafter, indicating that seasonal sexual recrudescence can be photoinducible. In addition, the study demonstrated that continued exposure to a stimulatory photoperiod does not maintain a heightened reproductive capacity indefinitely and testicular regression does ensue. Cox and co-workers146 found that increasing day length using artificial lights at the end of the breeding season in the northern hemisphere significantly decreased testosterone concentrations, suggesting a negative impact on the fertility of stallions being transported to the southern hemisphere. Therefore, shorter day length in autumn may provide a
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critical trigger in resensitizing the stallion to the photoinducible events of long days. This concept may be important to keep in mind when shipping breeding stallions from the northern hemisphere to the southern hemisphere, especially those with marginal fertility. To maintain peak reproductive performance in these stallions throughout the year, it may be important to put them on a short day lighting regime for 1–3 months before shipping them from one hemisphere to another. Further research in this area is warranted.
Endocrine–paracrine–autocrine factors during puberty in the stallion (Figures 99.4 and 99.5a–h) FSH, LH, and testosterone The role of FSH, LH, and LH-induced testosterone production in the development and maintenance of testicular function and spermatogenesis in the stallion are not entirely clear. Testosterone is known to act as an endocrine factor to modulate the release of LH but the fact that it is produced by the Leydig cells, and it acts at the level of the testes by way of androgen receptors, clearly indicates that it has a role as a paracrine–autocrine factor. Endocrine and testicular profiles of colts prior to and during puberty and post-puberty suggest a differential role of FSH, LH, and LH-induced testosterone, estradiol, and inhibin production on testicular function and spermatogenesis (Figures 99.4 and 99.5a–h).102, 147 Prior to puberty, serum FSH concentrations in the colt appeared to
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Figure 99.5 (a–h) Changes in plasma and testicular extract hormones at different ages in the breeding (B) season. (a,b) Plasma luteinizing hormone (LH) and follicle-stimulating hormone (FSH); (c,d) plasma and testicular testosterone (T); (e,f) plasma and testicular estradiol (E2 ); (g,h) plasma and testicular inhibin. In general, plasma and testicular hormone concentrations were lower in the < 1 year age group and increased thereafter except plasma FSH and testicular T. (Modified from Stewart and Roser.102 )
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Endocrine–Paracrine–Autocrine Regulation of Reproductive Function peak at about 40 weeks of age, declined, and then increased again at the time of puberty (∼83 weeks) (Figure 99.4).147 Serum concentrations of LH also peaked at around 40 weeks of age, declined to baseline, and then rose again at the time of puberty (Figure 99.4). Testosterone levels prior to puberty appeared to remain low and did not increase with the surge of LH and FSH at 40 weeks but then rapidly peaked in conjunction with LH at the time of puberty (Figure 99.4). In another study, FSH remained relatively high, fluctuated, and then started to decline at about 3.5 years while LH and testosterone steadily rose.148 Based on data presented by Stewart and Roser102 (Figure 99.5a), circulating LH was significantly higher in colts < 1 year old than in adult stallions. Circulating FSH levels were high in colts < 1 year old but did not change with age (Figure 99.5b). Plasma testosterone concentrations were significantly higher in colts less than 1 year old (Figure 99.5c), while, in contrast, testicular testosterone did not change with age (Figure 99.5d). The dichotomy between the profiles of plasma and testicular testosterone is yet to be explained. The above studies on the dynamics of FSH, LH, and LH-dependent testosterone release suggest that FSH may be critical for Sertoli cell development and proliferation and early spermatogenesis prior to puberty, whereas LH is important for pre-pubertal Leydig cell development which includes production of testosterone. The LH-induced testosterone release at puberty may trigger Sertoli cell differentiation and maturation necessary for gene expression of paracrine–autocrine factors important for germ cell maturation.94 In post-pubertal animals androgens maintain spermatogenesis,94 whereas the role of FSH declines. It has been reported that FSH receptor numbers per milligram of protein are lower in the mature testes of the rat and bull than in the developing testes.149–151 Although this has not been investigated in the stallion, it does support the concept that the actions of LH-induced testosterone and FSH mediate early spermatogenesis by way of the Sertoli cells.152 In post-pubertal animals, testosterone is the major regulator of spermatogenesis while the role of FSH declines.153
Estrogen, inhibin, IGF-1, and transferrin as testicular paracrine–autocrine factors during puberty in the stallion
Testicular estrogen during puberty Estrogen may not only have an endocrine role in modulating the release of LH and FSH but it appears to play a paracrine–autocrine role in Sertoli and germ cell development. Pre-pubertal colts at 6 months old were immunized against estrogen. At castration at
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27 months old, estrogen-immunized colts had greater testicular and parenchymal weights and produced more spermatozoa per horse than did control colts.33 In boars, reducing endogenous estrogen by treatment with an aromatase inhibitor was consistent with a delay in testicular maturation/puberty that allowed for a longer window of proliferation of Sertoli cells and maturation of Leydig cells, resulting in larger testes and higher spermatid production.154 The increase in estradiol (Figure 99.5e,f) suggests that this steroid may be important for maintenance of spermatogenesis. These data suggest that estrogen prior to puberty may be involved in hastening Sertoli cell differentiation and Leydig cell maturation. In addition, estradiol may be important in maintaining spermatogenesis.
Testicular inhibin during puberty There are data to suggest that inhibin and IGF-1 are important for events during puberty. Inhibin can inhibit spermatogonial proliferation in mice and hamsters.155 The observation that inhibin is significantly lower in equine plasma and testicular extracts from colts less than 1 year old and higher in colts 2–5 years old102 (Figure 99.5g,h) supports the concept that low inhibin prior to puberty allows for spermatogonial proliferation and that perhaps higher inhibin is important for differentiation of Sertoli and germ cells during maturation and spermatogenesis.
Testicular IGF-1 during puberty As mentioned above under the section on testicular IGF-1, IGF-1 stimulates the proliferation of prepubertal Sertoli cells and has a small mitogenic effect on immature Leydig cells.73 It stimulates spermatogonial DNA synthesis and has a maintaining effect on premeiotic DNA synthesis in the rat under in vitro conditions, and it induces the differentiation of mouse type A spermatogonia.73 It has been demonstrated in the colt that the highest amount of IGF-1 in testicular tissue is prior to puberty compared with post-pubertal and mature stallions,85 suggesting that IGF-1 is important for equine testicular development and early spermatogenesis.
Testicular transferrin during puberty Transferrin delivers iron as a nutritional component to germ cells.87 It is known to be a good indicator of Sertoli function and spermatogenic potential in bulls.89 It has been observed in the stallion that transferrin production in cultured Sertoli cells is significantly higher in the 2-year-old age group than in post-pubertal or mature stallions,21, 90 suggesting that
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in the equine testes transferrin is important in differentiation of Sertoli cells and germ cell development. More work needs to be done to determine other paracrine–autocrine factors in the equine testis that are important during puberty.
Epididymis Structure and function The epididymis is a highly convoluted duct which receives spermatozoa from the ductuli efferentes of the testis and conducts them to the more distal vas deferens.157 The epididymis is generally divided into the caput epididymis (head), corpus epididymis (body), and cauda epididymis (tail). However, it can be subdivided into many regions possessing their own distinct epithelial cell types.158, 159 The epididymis has four basic functions: (i) it serves as a sperm reservoir; (ii) it disposes of old and superfluous sperm; (iii) it is the site of spermatozoal maturation, which includes motility and fertilizing capability; and (iv) it is responsible for the composition of the fluid surrounding epididymal spermatozoa.157 Fluid reabsorption takes place in the efferent ductules160 and caput, sperm maturation occurs in the corpus and proximal end of the cauda, and sperm storage occurs in the cauda.12
Epididymal sperm transit time influenced by prostaglandins, oxytocin, vasopressin, and sympathetic innervation Although the length of the epididymis varies between species,161 the mean duration of epididymal sperm transit is surprisingly constant at around 9– 11 days. In stallions, epididymal transit time has been reported to vary between 5 and 14 days.162 Movement of the spermatozoa through the epididymis is probably due to the presence of prostaglandin F2α (PGF2α ), which induces continuous peristaltic contractions of smooth muscles in the wall of the caput and corpus epididymis.157 The contraction of the cauda epididymis has been found to be under the influence of sympathetic innervation, vasopressin, oxytocin, and both PGF2α and prostaglandin E (PGE). PGE is known to inhibit contractility, which may be responsible for the smooth muscles in the cauda being quiescent until stimulated to contract.157
Epididymal androgens The epididymis has long been known to be an androgen-dependent tissue in other species.163, 164 Under androgen regulation [testosterone and/or di-
hydrotestosterone (DHT)], cells of the epididymal epithelium produce proteins, glycoproteins, glycolipids, and phospholipids, all components of epididymal fluid which are necessary for maturation and survival of spermatozoa.93 The profile of proteins secreted by the epididymis was dramatically altered following castration,165 but showed improvement following testosterone replacement.166 However, testosterone replacement alone was insufficient to completely restore levels and regionalization of protein secretion to the pre-castration pattern, suggesting that other testicular products such as estrogen may be involved in regulating protein secretion in the epididymis. Epididymal estrogens Recent studies have revealed that estrogen has a much greater role in male reproduction than previously believed, particularly at the level of the epididymis. It has clearly been demonstrated that estrogen regulates the reabsorption of luminal fluid in the efferent ducts of mice and, without this function, these animals were infertile.101 Male mice without a functional estrogen receptor (estrogen receptor knockout mice) were infertile.167 Sperm in the epididymis of these mice appeared abnormal.168 Young adult mice given ICI 182,780, an estrogen receptor antagonist, were initially fertile but became infertile after treatment, indicating that the infertility was related to a loss of functional estrogen receptors and not just developmental abnormalities.169 Estrogen and androgen receptors Steroid receptors, specifically AR, ERα, and ERβ, have been identified in the epididymis of several mammalian species, including the stallion.170–174 AR and ERβ localize to all regions of the epididymis.175, 176 ERα was generally found in the efferent ductules across species, but its localization in the epididymis was much more variable between species and even within a species, as seen in the rat.160 Estrogen and androgen and their receptors in the stallion epididymis Parlevleit and co-workers174 demonstrated that estrogens and androgens are found in the luminal fluid of the epididymis and their receptors (ERα, ERβ, and AR) are located throughout the epididymis during development in the stallion. It appears that ERα distribution is both age and tissue/segment dependent whereas ARs and ERβ are present in all segments in all age groups. ERα was not present in the epididymis epithelium of the cauda of pre-pubertal animals or in
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Endocrine–Paracrine–Autocrine Regulation of Reproductive Function the corpus of post-pubertal animals. It was present in all regions of peri-pubertal colts and the same for adult animals. ERβ was present in the epididymis epithelium of all three epididymal regions throughout development and in adulthood in stallions. ARs were also present in the epididymis epithelium of all three segments in all age groups. Although epididymal function is predominantly regulated by testicular steroids, it has recently been suggested that the sperm themselves may influence epididymal function.177
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didymis compared with other regions of the epididymis and the testis. It was also demonstrated that the amount of transferrin in a particular region was age dependent. The caput had the highest amount of transferrin in a pre-pubertal group of colts compared with pubertal, post-pubertal, and older stallions. Parlevliet and co-workers91 demonstrated that transferrin was present in the seminal fluid. The amount of transferrin was not correlated with fertility, but more studies are warranted to definitively prove this observation.
Epididymal secretions in the stallion Proteins produced by the equine epididymal epithelium and secreted into the lumen have been characterized using two-dimensional electrophoresis.156 Using this proteomic approach, 201 proteins were found in the lumen and 117 were found secreted by the epididymal epithelium. The major proteins secreted by the epithelium were lactoferrin and clusterin along with other components such as procathepsin D, cholesterol transfer protein (HE1/CTP), glutathione peroxidase (GPX), beta-Nacetyl-hexosaminidase, and prostaglandin D2 synthase (PDGS). The major proteins and compounds found in the luminal fluid were albumin, lactoferrin, PGDS, GPX, HE1/CTP, and hexosaminidase.156 Lactoferrin and transferrin in the equine epididymis Two proteins that have recently been investigated in the stallion epididymis are lactoferrin178 and transferrin.90 Preliminary results indicate that lactoferrin, a defense protein and antioxidant, was localized in the principal cells in the corpus and cauda but not in the caput throughout epididymal development in the stallion. Lactoferrin was also localized to the sperm membrane of the midpiece and tail regions. It appears that, when testosterone was incubated with equine epididymal tissue sections, it did not increase the secretion of lactoferrin above baseline in the corpus but did increase secretion in the cauda by approximately 20%. Estradiol increased lactoferrin secretion above baseline approximately 85%, 16%, and 18% in the caput, corpus, and cauda, respectively.178 Together, these results suggest that lactoferrin secretion from the equine epididymal epithelium can be influenced by steroids and that it binds to sperm as a defense protein to protect sperm from damage due to oxidative stress. Preliminary studies by Parlevliet and co-workers90 demonstrated that transferrin, an iron-binding protein, was present in all regions of the epididymis in all age groups. However it was highest in the tissue extracts of the cauda region of the equine epi-
Summary The reproductive tract of the stallion is regulated by an interactive system of endocrine–paracrine– autocrine factors. These factors and their membrane receptors play an integral role in regulating normal reproduction during pre-pubertal, pubertal, and post-pubertal maturation and maintenance of the testis and epididymis. Seasonal effects can change the dynamics of the interplay between these factors and their membrane receptors in testicular and epididymal tissue. This review presents factors and systems that may be regulating stallion reproduction by drawing on information from other species. It also presents an up-to-date account of the endocrine–paracrine–autocrine regulation of reproductive function in the stallion.
References 1. Matsumoto A. Hormonal control of spermatogenesis. In: Burger H, Kretser D (eds) The Testis, 2nd edn. New York: Raven Press, 1989; pp. 181–96. 2. Amann RP. Physiology and endocrinology. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 658–85. 3. Weinbauer GF, Nieschlag E. Hormonal control of spermatogenesis. In: de Kretser D (ed.) Molecular Biology of Male Reproductive System. New York: Academic Press, 1993; pp. 99–142. 4. McLachlan RI. The endocrine control of spermatogenesis. Baillieres Best Pract Res Clin Endocrinol Metab 2000;14:345–62. 5. Roser JF. Endocrine and paracrine control of sperm production in stallions. Anim Reprod Sci 2001;68:139–51. 6. Schanbacher BD. Hormonal interrelationships between hypothalamus, pituitary, and testis of rams and bulls. J Anim Sci 1982;55 (Suppl. 2):56–67. 7. Gerlach T, Aurich JE. Regulation of seasonal reproductive activity in the stallion, ram and hamster. Anim Reprod Sci 2000;58:197–213. 8. Roser JF. Regulation of testicular function in the stallion: an intricate network of endocrine, paracrine and autocrine systems. Anim Reprod Sci 2008;107:179–96. 9. Roser JF. Reproductive endocrinology of the stallion. In: Equine Breeding Management and Artificial Insemination. St Louis: Saunders Elsevier, 2009; pp. 17–31.
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10. Huhtaniemi I, Toppari J. Endocrine, paracrine and autocrine regulation of testicular steroidogenesis. Adv Exp Med Biol 1995;377:33–54. 11. Huleihel M, Lunenfeld E, Samper JC. Regulation of spermatogenesis by paracrine/autocrine testicular factors. Asian J Androl 2004;6:259–68. 12. Sostaric E, Aalberts M, Gadella BM, Stout TAE. The roles of the epididymis and proteasomes in the attainment of fertilizing capacity by stallion sperm. Anim Reprod Sci 2008;107:237–48. 13. Sharp DC, Cleaver BD. Melatonin. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 100–8. 14. Sharp DC, Cleaver BD, Davis SD. Photoperiod. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 179–85. 15. Malpaux B, Migaud M, Tricoire H, Chemineau P. Biology of mammalian photoperiodism and the critical role of the pineal gland and melatonin. J Biol Rhythms 2001;16:336–47. 16. Irvine CHG, Alexander SL. A novel technique for measuring hypothalamic and pituitary hormone secretion rates from collection of pituitary venous effluent in the normal horse. J Endocrinol 1987;113:183–92. 17. Irvine CH, Alexander SL. Secretory patterns and rates of gonadotropin-releasing hormone, follicle stimulating hormone, and luteinizing hormone revealed by intensive sampling of pituitary venous blood in the luteal phase mare. Endocrinology 1993;132:212–18. 18. Eisenhauer KM, Roser JF. Effects of lipoproteins, eLH, eFSH and ePRL on equine testicular steroidogenesis in vitro. J Androl 1995;16:18–27. 19. Griswold MD. Protein secretion by Sertoli cells: General considerations. In: Russell LD, Griswold MD (eds) The Sertoli Cell. Clearwater, FL: Cache River Press, 1993; pp. 195–200. 20. Dean KJ, Roser JF. Inhibin and estradiol production by Cultured equine Sertoli cells. In: Proceedings of the 16th Equine Nutrition and Physiology Society, 1999, pp. 190–1. 21. Bidstrup LA, Dean DJ, Pommer AC, Roser JF. Transferrin production in cultured Sertoli cells during testicular maturation in the stallion. Biol Reprod Suppl 2002;66:493. 22. Karasek M, Winczyk K. Melatonin in humans. J Physiol Pharmacol 2006;57(Suppl 5):19–39. 23. Okubo K, Nagahama Y. Structural and functional evolution of gonadotropin-releasing hormone in vertebrates. Acta Physiol (Oxf) 2008;193:3–15. 24. Lambert A, Talbot TA, Anobile CJ, Robertson WR. Gonadotropin heterogeneity and biopotency: implications for assisted reproduction. Mol Hum Reprod 1998;4:619–29. 25. Bousfield GR, Butney VY, Gotshcall RR, Baker VL, Mooreb WT. Structural features of mammalian gonadotropins. Mol Cell Endocrinol 1996;125:3–19. 26. Pierce JG, Parsons TF. Glycoprotein hormones: structure and function. Annu Rev Biochem 1981;50:465–5. 27. Irvine CHG. Kinetics of gonadotrophins in the mare. J Reprod Fertil Suppl 1979;27:131–41. 28. Muyan M, Roser JF, Dybdal N, Baldwin DM. Modulation of gonadotropin-releasing hormone-stimulated luteinizing hormone release in cultured male equine anterior pituitary cells by gonadal steroids. Biol Reprod 1993;49:340–5. 29. Tilbrook AJ, Clarke IJ. Negative feedback regulation of the secretion and actions of gonadotropin-releasing hormone in males. Biol Reprod 2001;64:735–42.
30. Thompson Jr DL, Pickett BW, Squires EL, Nett TM. Effect of testosterone and estradiol-17β alone and in combination on LH and FSH concentrations in blood serum and pituitary of geldings and in serum after administration of GnRH. Biol Reprod 1979;21:1231–7. 31. Seamans MC, Roser JF, Linford RL, Liu IK, Hughes JP. Gonadotrophin and steroid concentrations in jugular and testicular venous plasma in stallions before and after GnRH injection. J Reprod Fertil Suppl 1991;44:57–67. 32. Fink G. The G.W. Harris Lecture. Steroid control of brain and pituitary function. J Exp Physiol 1988;73:257–93. 33. Thompson Jr DL, Honey PG. Active immunization of prepubertal colt against estrogens: Hormonal and testicular responses after puberty. J Anim Sci 1984;59:189–96. 34. Barraclough CA, Wise PM, Turgeon J, Shander D, Depaulo L, Range N Recent studies on the regulation of pituitary LH and FSH secretion. Biol Reprod 1979;20:86–97. 35. Knobil E. The neuroendocrine control of the menstrual cycle. Prog Hormone Res 1980;36:53–88. 36. Baldwin DM, Roser JF, Muyan M, Lasley B, Dybdal N. Direct effects of free and conjugated steroids on GnRHstimulated LH release in cultured equine pituitary cells. J Reprod Fertil Suppl 1991;44:327–32. 37. Loveland KL, Dias V, Meachem S, Raipert-De Meyts E. The transforming growth factor-beta super family in early spermatogenesis: potential relevance to testicular dysgenesis. Int J Androl 2007;30:377–84. 38. Vale W, Bilezikjian LM, Rivier C. Reproductive and other roles of inhibins and activins. In: Knobil E, Neill JD (eds) The Physiology of Reproduction, 2nd edn. New York: Raven Press, 1994; pp. 1861–71. 39. Nagata S, Tsunoda N, Nagamine N, Tanaka Y, Taniyama H, Nambo Y, Watanabe G, Taya K. Testicular inhibin in the stallion: cellular source and seasonal changes in its secretion. Biol Reprod 1998;59:62–8. 40. McKinnon AO, Brown RW, Pashen RL, Greenwood PE, Vasey JR. Increased ovulation rates in mares after immunization against recombinant bovine inhibin alphasubunit. Equine Vet J 1992;24:144–6. 41. Nambo Y, Kaneko H, Nagata S, Oikawa M, Yoshihara T, Nagamine N, Watanabe G, Taya K. Effect of passive immunization against inhibin on FSH secretion, folliculogenesis and ovulation rate during the follicular phase of the estrous cycle in mares. Theriogenology 1998;50: 545–57. 42. Roser JF, McCue P, Hoye E. Inhibin activity in the mare and stallion. Domest Anim Endocrinol 1994;11:87–100. 43. Yamanouchi K, Sugawara Y, Tojo H, Takahashi M. Expression of inhibin/ activin subunit genes in reproductive organs of pregnant mare and equine fetus. In: Kawashima S, Kikuyama S (eds) Advances in Comparative Endocrinology I. Bologna, Italy: Monduzzi Editor, 1997; p. 487. 44. Ben-Jonathan N, LaPensee CR, LaPensee EW. What can we learn from rodents about prolactin in humans? Endocr Rev 2008;29:1–41. 45. Bachelot A, Binart N. Reproductive role of prolactin. Reproduction 2007;133:361–9. 46. Jedlinska M, Rozewiecka I, Ziecik AJ. Effect of hypoprolactinaemia and hyperprolactinaemia on LH secretion, endocrine function of testes and structure of seminiferous tubules in boars. J Reprod Fertil 1995;103:265–72. 47. Regisford EGC, Katz LS. Effects of bromocriptineinduced hyperprolactinaemia on gonadotrophin secretion and testicular function in rams (Ovis aries) during two seasons. J Reprod Fertil 1993;99:529–37.
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Endocrine–Paracrine–Autocrine Regulation of Reproductive Function 48. Hondo E, Kuromaru M, Sakai S, Ogawa K, Hayashi Y. Prolactin receptor expression in rat spermatogenic cells. Biol Reprod 1995;52:1284–90. 49. Bartke A. Hyperprolactinemia and male reproduction. In: Paulson J, Negro-Vilar A, Lucena E, Martini L (eds) Andrology: Male Fertility and Sterility. New York: Academic Press, 1986; pp. 101–23. 50. Frasor J, Gibori G. Prolactin regulation of estrogen receptor expression. Trends Endocrinol Metab 2003;14:118–23. 51. Thompson Jr DL, Johnson L, Wiest JJ. Effects of month and age on prolactin concentrations in stallion serum. J Reprod Fertil Suppl 1987;35:67–70. 52. Johnson AL. Serum concentrations of prolactin, thyroxine and triiodothyronine relative to season and the estrous cycle in the nonpregnant mare. J Anim Sci 1986;62:1012–20. 53. Zoeller RT, Tan SW, Tyl RW. General background on the hypothalamic-pituitary-thyroid (HPT) axis. Crit Rev Toxicol 2007;37:11–53. 54. Mendis-Handagama C, Ariyaratne S. Prolonged and transient neonatal hypothyroidism on Leydig cell differentiation in the postnatal rat testis. Arch Androl 2004;50:347–57. 55. Cooke PS, Holsberger DR, Witorsch RJ, Sylvester PW, Meredith JM, Treinen KA, Chapin RE. Thyroid hormone, glucocorticoids, and prolactin at the nexus of physiology, reproduction, and toxicology. Toxicol Appl Pharmacol 2004;194:309–35. 56. Maran RRM. Thyroid hormones: their role in testicular steroidogenesis. Arch Androl 2003;49:375–88. 57. Meredith TB, Dobrinski I. Thyroid function and pregnancy status in broodmares. J Am Vet Med Assoc 2004;224:892–4. 58. Higham CE, Trainer PJ. Growth hormone excess and the development of growth hormone receptor antagonists. Exp Physiol 2008;93:1157–69. 59. Zachman M. Interrelations between growth hormone and sex hormones-physiology and therapeutics consequences. Horm Res Suppl 1992;38:1–8. 60. Chandrashekar V, Bartke A. The role of growth hormone in pituitary and testicular function in adult mice Endocrinology 1998;139:1067–74. 61. Spiteri-Grech J, Nieschlag E. The role of growth hormone and insulin-like growth factor I in the regulation of male reproductive function. Horm Res Suppl 19921:22–7. 62. Breier BH, Vickers MH, Gravance CG, Casey PJ. Therapy with growth hormone: major prospects for the treatment of male subfertility? Endocrinol J Suppl 1998;45:53–60. 63. Laron Z, Klinger B. Effect of insulin-like growth factor-I treatment on serum androgens and testicular penile size in males with Laron syndrome (primary growth hormone resistance). Eur J Endocrinol 1998;138:176–80. 64. Kanzaki M, Morris PL. Growth hormone regulates steroidogenic acute regulatory protein expression and steroidogenesis in Leydig cell progenitors. Endocrinology 1999;140:1681–6. 65. Shoham Z, Conway GS, Ostergaard H, Lahlou N, Bouchard P, Jacobs HS. Cotreatment with growth hormone for induction of spermatogenesis in patients with hypogonadotropic hypogonadism. Fertil Steril 1992;57:1044–51. 66. Aurich JE, Kranski S, Parvizi N. Somatostatin treatment affects testicular function in stallions. Theriogenology 2003;60:163–74. 67. Hess MF, Roser JF. A comparison of the effects of equine luteinizing hormone (eLH), equine growth hor-
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mone (eGH) and human recombinant insulin-like growth factor (hrIGF-I) on steroid production in cultured equine Leydig cells during sexual maturation. Anim Reprod Sci 2005;89:7–19. Aurich C, Sieme H, Hoppe H, Schlote S. Involvement of endogenous opioids in the regulation of LH and testosterone release in the male horse. J Reprod Fertil 1994;102:327–36. Heindel JJ, Treinen KA. Physiology of the male reproductive system: endocrine, paracrine and autocrine regulation. Toxicol Pathol 1989;17:411–45. Jegou B, Sharpe RM. Paracrine mechanisms in testicular control. In: de Kretser D (ed.) Molecular Biology of Male Reproductive System. New York: Academic Press, 1993; pp. 271–310. Skinner MK. Secretion of growth factors and other regulatory factors. In: Russell LD, Griswold MD (eds) The Sertoli Cell. Clearwater, FL: Cache River Press, 1993; pp. 237–47. Jegou B, Pineau C. Current aspects of autocrine and paracrine regulation of spermatogenesis. In: Mukhopadhyay AK, Raizada MK (eds) Tissue Renin–Angiotensin Systems. New York: Plenum Press, 1995; pp. 67–86. Gnessi L, Fabbri A, Spera G. Gonadal peptides as mediators of development and functional control of the testis: an integrated system with hormones and local environment. Endocr Rev 1997;18:541–609. Holdcraft RW, Braun RE. Hormonal regulation of spermatogenesis. Int J Androl 2004;27:335–42. Spiteri-Grech J, Nieschlag E. Paracrine factors relevant to the regulation of spermatogenesis-a review. J Reprod Fertil 1993;98:1–14. Le Roy C, Lejeune HA, Chuzel F, Saez JM, Langlois D. Autocrine regulation of Leydig cell differentiated functions by insulin-like growth factor I and transforming growth factor beta. J Steroid Biochem Mol Biol 1999;69:379–84. Hull KL, Harvey S. Growth hormone: a reproductive endocrine-paracrine regulator? Rev Reprod 2000;5:175–82. Abney TO. The potential role of estrogens in regulating Leydig cell development and function: a review. Steroids 1999;64:610–17. O’Donnell L, Robertson KM, Jones ME, Simpson ER. Estrogen and spermatogenesis. Endocr Rev 2001;22:289–318. Petersen C, Soder O. The Sertoli cell: a hormonal target and ‘super’ nurse for germ cells that determines testicular size. Horm Res 2006;66:153–61. Weldt C, Sidis Y, Keutmann H, Schneyer A. Activins, inhibins and follistatins: From endocrinology to signaling. A paradigm for the new millennium. Exp Biol Med 2002;227:724–52. Roser JF. Endocrine diagnostics and therapeutics for the stallion with declining fertility. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 244–51. Colon E, Zaman F, Axelson M, Larsson O, CarlssonSkwirut C, Svechnikov KV, Soder O. Insulin-like growth factor-I is an important antiapoptotic factor for rat Leydig cells during postnatal development. Endocrinology 2007;148:128–39. Ozkurkcugil C, Yardimoglu M, Dalcik H, Erdogan S, Gokalp A. Effect of insulin-like growth factor-1 on apoptosis or rat testicular germ cells induced by testicular torsion. Br J Urol Int 2004;93:1094–7.
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85. Hess ME, Roser JF. The effects of age, season and fertility status on plasma and intratesticular insulin-like growth factor I concentration in stallions. Theriogenology 2001;56:723–33. 86. Macpherson ML, Simmen RC, Simmen FA, Hernandez J, Sheerin BR, Varner DD, Loomis P, Cadario ME, Miller CD, Brinsko SP, Rigby S, Blanchard TL. Insulinlike growth factor I and insulin-like growth factor binding protein-2 and -5 in equine seminal plasma: association with sperm characteristics and fertility. Biol Reprod 2002;67:648–54. 87. Sylvester SR, Griswold MD. The testicular iron shuttle: a ‘nurse’ function. J Androl 1994;15:381–5. 88. Zalata A, Hafez T, Schoonians F, Comhaire F. The possible meaning of transferrin and its soluble receptors in seminal plasma as markers of the seminiferous epithelium. Hum Reprod 1996;11:761–4. 89. Gilmont RR, Senger PL, Sylvester SR, Griswold MD. Seminal transferrin and spermatogenic capability in the bull. Biol Reprod 1990;43:151–7. 90. Parlevliet JM, Bidstrup LA, Famula T, Roser JF. Transferrin concentrations in the different regions of the epididymis and in the testes in horses of different ages. In: Proceedings 15th ICAR, 2004, p. 160. 91. Parlevliet JM, Bidstrup LA, Roser JF. Transferrin concentrations in seminal fluid in horses: a marker for fertility. Anim Reprod Sci 2008;107:337–8. 92. Sierens JE, Sneddon SF. Collins F, Millar MR, Saunders PT. Estrogens in testis biology. Ann N Y Acad Sci 2005;1061:65–76. 93. Bilinska B, Wiszniewska B, Kosiniak-Kamysz K, KotulaBalak M, Gancarczyk M, Hejmej A, Sadowska J, Marchlewicz M, Kolasa A, Wenda-Rozewicka L. Hormonal status of male reproductive system: androgens and estrogens in the testis and epididymis. In vivo and in vitro approaches. Reprod Biol Suppl 2006;1:43–58. 94. Dohle GR, Smit M, Weber RF. Androgens and male fertility. World J Urol 2003;21:341. 95. Kotula M, Tuz R, Frczek B, Wojtusiak A, Bilinska B. Immunolocalization of androgen receptors in testicular cells of prepubertal and pubertal pigs. Folia Histochem Cytobiol 2000;38:157–62. 96. Collins LL, Lee HJ, Chen YT, Chang M, Hsu HY, Yeh S, Chang C. The androgen receptor in spermatogenesis. Cytogenet Genome Res 2003;103:299–301. 97. Ramesh R, Pearl CA, At-Taras E, Roser, JF, Berger T. Ontogeny of androgen and estrogen receptor expression in porcine testis: effect of reducing testicular estrogen synthesis. Anim Reprod Sci 2007;102:286–99. 98. Meng J, Holdcraft RW, Shima JE, Griswold MD, Braun RE. Androgens regulate the permeability of the bloodtestis barrier. Proc Natl Acad Sci USA 2005;102:16696–700. 99. Singh J, O’Neill C, Handelsman DJ. Induction of spermatogenesis by androgens in gonadotropin-deficient (hpg) mice. Endocrinology 1995;136:5311–21. 100. Xu Q, Lin HY, Yeh SD, Yu IC, Wang RS, Chen YT, Zhang C, Altuwaijri S, Chen LM, Chuang KH, Chiang HS, Yeh S, Chang C. Infertility with defective spermatogenesis and steroidogenesis in male mice lacking androgen receptor in Leydig cells. Endocrine 2007;32:96–106. 101. Hess RA, Carnes K. The role of estrogen in testis and the male reproductive tract: a review and species comparison. Anim Reprod 2004;1:5–30. 102. Stewart BL, Roser JF. Effects of age, season and fertility status on plasma and intratesticular immunoreactive
103.
104.
105.
106.
107.
108.
109.
110.
111.
112.
113.
114. 115.
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(Ir) inhibin concentrations in stallions. Domest Anim Endocrinol 1998;15:129–39. Sipahutar H, Sourdaine P, Moslemi S, Plainfosse B, Seralini GE. Immunolocalization of aromatase in stallion Leydig cells and seminiferous tubules. J Histochem Cytochem 2003;51:311–18. Hess MF, Roser JF. Immunocytochemical localization of cytochrome P450 aromatase in the testis of prepubertal, pubertal and postpubertal horses. Theriogenology 2004;61:293–9. Clement H, Conley A, Mapes S, Roser J. Immunolocalization of estrogen receptor α, estrogen receptor β, and androgen receptors in the equine testis. Biol Reprod Suppl 2002;66:114. de Kretser DM, Buzzard JJ, Okuma Y, O’Connor AE, Hayashi T, Lin SY, Morrison JR, Loveland KL, Hedger MP. The role of activin, follistatin and inhibin in testicular physiology. Mol Cell Endocrinol 2004;225:57–64. Hsueh AJ, Dahl KD, Vaughan J, Tucker E, Rivier J, Bardin CW, Vale W. Heterodimers and monodimers of inhibin subunits have different paracrine actions in the modulation of luteinizing hormone-stimulated androgen biosynthesis. Proc Natl Acad Sci USA 1987;84:5082–6. Lin T, Calkins JK, Morris PL, Vale W, Bardin CW. Regulation of Leydig cell function in primary culture by inhibin and activin. Endocrinology 1989;125:2134–40. Lejeune H, Chuzel F, Sanchez P, Durand P, Mather JP, Saez JM. Stimulating effect of both human recombinant inhibin A and activin A on immature porcine Leydig cell functions in vitro. Endocrinology 1997;138:4783–91. Hakovirta H, Kaipia A, Soder O, Parvinen M. Effects of activin-A, inhibin-A and transforming growth factorbeta 1 on stage specific deoxyribonucleic acid synthesis during rat seminiferous epithelial cycle. Endocrinology 1993;133:1664–8. Boitani C, Stefanini M, Fragale A, Morena AR. Activin stimulates Sertoli cell proliferation in a defined period of rat testis development. Endocrinology 1995;136: 5438–44. Meehan T, Schlatt S, O’Bryan MK, de Kretser DM, Loveland KL. Regulation of germ cell and Sertoli cell development by activin, follistatin, and FSH. Dev Biol 2000;220:225–37. Kumanov P, Nandipati K, Tomova A, Agarwal A. Inhibin B is a better marker of spermatogenesis than other hormones in the evaluation of male factor infertility. Fertil Steril 2006;86:332–8. Ivell R, Hartung S, Anaud-Ivell R. Insulin-like factor 3: where are we now? Ann N Y Acad Sci 2005;1041:486–96. Bay K, Hartung S, Ivell R, Schumacher M, Jurgensen D, Jorgensen N, Holm M, Skakkebaek NE, Andersson AM. Insulin-like factor 3 serum levels in 135 normal men and 85 men with testicular disorders: relationship to the luteinizing hormone-testosterone axis. J Clin Endocrinol Metab 2005;90:3410–18. Balvers M, Spiess AN, Domagalski R, Hunt N, Kilic E, Mukhopadhyay AK, Hanks E, Charlton HM, Ivell R. Relaxin-like factor expression as a marker of differentiation in the mouse testis and ovary. Endocrinology 1998;139:2960–70. Sadeghian H, Anand-Ivell R, Balvers M, Relan V, Ivell R. Constitutive regulation of the Insl3 gene in rat Leydig cells. Mol Cell Endocrinol 2005;124:10–20. Ivell R, Balvers M, Anand RJK, Paust HJ, McKinnell C, Sharpe R. Differentiation-dependent expression of
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17beta-HSD type 10 in the rodent testis: effect of aging in Leydig cells. Endocrinology 2003;144:3130–7. Hombach-Klonisch S, Schon J, Kehlen A, Blottner S, Klonisch T. Seasonal expression of INSL3 and Lgr8/Insl3 receptor transcripts indicates variable differentiation of Leydig cells in the roe deer testis. Biol Reprod 2004;71:1079–87. Paust HJ, Wessels J, Ivell R, Mukhopadhyay AK. The expression of the RLF/INSL3 gene is reduced in Leydig cells of the aging rat testis. Exp Gerontol 2002;37: 1461–7. Caprio M, Fabbrini E, Ricci G, Basciani S, Gnessi L, Arizzi M, Carta AR, De martino MU, Isidori AM, Frajese GV, Fabbri A. Ontogenesis of leptin receptor in rat Leydig cells. Biol Reprod 2003;68:1199–207. Nef S, Parada LF. Cryptorchidism in mice mutant for Insl3. Nat Genet 1999;22:295–9. Zimmermann S, Steding G, Emmen JM, Brinkmann AO, Nayernia K, Holstein AF, Engel W, Adham IM. Targeted disruption of the Insl3 gene causes bilateral cryptorchidism. Mol Endocrinol 1999;13:681–91. Overbeek PA, Gorlov IP, Sutherland RW, Houston JB, Harrison WR, Boettger-Tong HL, Bishop CE, Agoulnik A. A transgenic insertion causing cryptorchidism in mice. Genesis 2001;30:26–35. Kumagai J, Hsu SY, Matsumi H, Roe J-S, Fu P, Wade JD, Bathgate RAD, Hsueh AJW. INSL3/Leydig Insulin-like peptide activates the LGR8 receptor important in testis descent. J Biol Chem 2002;277:31283–6. Kawamura K, Kumagai J, Sudo S, Chun SY, Pisarska M, Morita H, Toppari J, Fu P, Wade JD, Bathgate RA, Hsueh AJ. Paracrine regulation of mammalian oocyte maturation and male germ cell survival. Proc Natl Acad Sci USA 2004;101:7323–8. Klonisch T, Steger K, Kehlen A, Allen WR, Froehlich C, Kauffold J, Bergmann M, Hombach-Klonisch S. INSL3 ligand-receptor system in the equine testis. Biol Reprod 2003;68:1975–81. Bardin CW, Shaha C, Mather J, Salomon Y, Margioris AN, Liotta AS, Gerendai I, Chen CL, Krieger DT. Identification and possible function of pro-opiomelanocortin-derived peptides in the testis. Ann N Y Acad Sci 1984;438:346–64. Soverchia L, Mosconi G, Ruggeri B, Ballarini P, Catone G, Degl’Innocenti S, Nabissi M, Polzonetti-Magni AM. Proopiomelanocortin gene expression and beta-endorphin localization in the pituitary, testis, and epididymis of stallion. Mol Reprod Dev 2006;73:1–8. Thackare H, Nicholson HD, Whittington K. Oxytocin: its role in male reproduction and new potential therapeutic uses. Hum Reprod Update 2006;12:437–48. Watson ED, Nikolakopoulos E, Gilbert C, Goode J. Oxytocin in the semen and gonads of the stallion. Theriogenology 1999;51:855–65. Pickett BW, Amann RP, McKinnon AO, Squires EL, Voss JL. Management of the Stallion for Maximum Reproductive Efficiency, II. Animal Reproduction Laboratory General Series Bulletin No. 05. Fort Collins, CO: Colorado State University, 1989. Harris JM, Irvine CHG, Evans MJ. Seasonal changes in serum levels of FSH, LH, testosterone and semen parameters in stallions. Theriogenology 1983;19:311–22. Thompson Jr DL, Pickett BW, Nett TM. Effect of season and artificial photoperiod on levels of estradiol17β and estrone in blood serum of stallions. J Anim Sci 1978;47:184–7.
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135. Thompson Jr DL, Johnson L, St George RL, Garza Jr F. Concentrations of prolactin, luteinizing hormone and follicle stimulating hormone in pituitary and serum of horses: effects of sex, season and reproductive state. J Anim Sci 1986;63:854–60. 136. Clay CM, Clay JN. Endocrine and testicular changes associated with season, artificial photoperiod, and the peripubertal period in stallions. Vet Clin North Am Equine Pract 1992;8:31–56. 137. Burns PJ, Jawad MJ, Edmundson A, Cahill C, Boucher JK, Wilson EA, Douglas RH. Effect of increased photoperiod on hormone concentrations in thoroughbred stallions. J Reprod Fertil Suppl 1982;32:103–11. 138. Johnson L, Thompson Jr DL. Age-related and seasonal variation in the Sertoli cell population, daily sperm production and serum concentrations of follicle-stimulating hormone, luteinizing hormone and testosterone in stallions. Biol Reprod 1983;29:777–89. 139. Sharp DC. Environmental influences on reproduction in horses. Vet Clin North Am Equine Pract 1980;2:207–23. 140. Aurich C, Gerlach T, Aurich JE, Hoppen HO, Lange J, Parvizi N. Dopaminergic and opioidergic regulation of gonadotropin and prolactin release in stallions. Reprod Domest Anim 2002;37:335–40. 141. Wagener A, Blottner S, Goritz F, Streich WJ, Fickel J. Differential changes of a and b FGF, IGF-1 and -2, and TGFalpha during seasonal growth and involution of roe dear testis. Growth Factors 2003;21:95–102. 142. Weng Q, Medan MS, Xu M, Tsubota T, Watanabe G, Taya K. Seasonal changes in immunolocalization of inhibin/activin subunits and testicular activity in wild male raccoon dogs (Nyctereutes procyonoides). J Reprod Dev 2006;52:503–10. 143. Sheng X, Zhang H, Zhang W, Song M, Zhang M, Li B, Weng Q, Watanabe G, Tava K. Seasonal changes in spermatogenesis and immunolocalization of inhibin/activin subunits in the wild male ground squirrel (Citellus dauricus Brandt). J Reprod Dev 2008;54:460–4. 144. Pickett BW. Factors affecting sperm production and output. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 689–704. 145. Hoffman LS, Adams TE, Evans JW. Circadian, circhoral and seasonal variation in patterns of gonadotrophin secretion in geldings. J Reprod Fertil Suppl 1987;35:51–8. 146. Cox JE, Redhead PH, Jawad NMA. The effect of artificial photoperiod at the end of the breeding season on plasma testosterone concentrations in stallions. Austr Vet J 1988;65:239–41. 147. Naden J, Amann RP, Squires EL. Testicular growth, hormone concentrations, seminal characteristics and sexual behaviour in stallions. J Reprod Fertil 1990;88:167–76. 148. Johnson AL, Varner DD, Thompson DL. Effect of age and season on the establishment of spermatogenesis in the horse. J Reprod Fertil Suppl 1991;44:87–97. 149. Bortolussi M, Zanchetta R, Belvedere P, Colombo L. Sertoli and Leydig cell numbers and gonadotropin receptors in rat testis from birth to puberty. Cell Tissue Res 1990;260:185–91. 150. Dias JA, Reeves JJ. Testicular FSH receptor numbers and affinity in bulls of various ages. J Reprod Fertil 1982;66:39–45. 151. Sundby A, Andersen D, Purvis K, Hansson V. Testicular gonadotropin receptors, testicular testosterone, dihydrotestosterone and androstenedione in the developing bull. Arch Androl 1984;12:59–64.
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152. Sairam MR, Krishnamurthy H. The role of follicle stimulating hormone in spermatogenesis: lessons from knockout animal models. Arch Med Res 2001;32:601–8. 153. O’Shaughnessy PJ Morris ID, Huhtaniemi I, Baker PJ Abel MH. Role of androgen and gonadotrophins in the development and function of the Sertoli cells and Leydig cells: data from mutant and genetically modified mice. Mol Cell Endocrinol 2009;306:2–8. 154. At-Taras EE, Berger T, McCarthy MJ, Conley AJ, NittaOda BJ, Roser JF. Reducing estrogen synthesis in developing boars increases testis size and total sperm production. J Androl 2006;27:552–9. 155. van Dissel-Emiliani FM, Grootenhuis AJ, de Jong FH, de Rooij DG. Inhibin reduces spermatogonial numbers in testes of adult mice and Chinese hamsters. Endocrinology 1989;125:1899–903. 156. Fouchecourt S, Metayer S, Locatelli A, Dacheux R, Dacheux JL. Stallion epididymal fluid proteome: Qualitative and quantitative characterization; secretion and dynamic changes of major proteins. Biol Reprod 2000;62; 1790–803. 157. Cosentino MJ, Cockett ATK. Review Article: Structure and function of the epididymis. Urol Res 1986;14:229–40. 158. Hoskins DD, Brandt H, Acott TS. Initiation of sperm motility in the mammalian epididymis. Fed Proc 1978;37:2534–6. 159. Reid BL, Cleand KW. The structure and function of the epididymis. I. The histology of the rat epididymis. Aust J Zool 1957;5:233–46. 160. Hess RA. Estrogen in the adult male reproductive tract: a review. Reprod Biol Endocrinol 2003;1:52–66. 161. Jones R, Glover TD. Interrelationship between spermatozoa, the epididymis and epididymal plasma. In: Dukett JG, Racey PA (eds) The Biology of the Male Gamete. Biol J Linnean Soc 1975;7 Suppl 1:367–88. 162. Franca L, Avelar G, Almeida F. Spermatogenesis and sperm transit through the epididymis in mammals with emphasis on pigs. Theriogenology 2005;63:300–18. 163. Robaire B, Viger RS. Regulation of epididymal epithelial cell functions. Biol Reprod 1995;52:226–36. 164. Cooper TG. Role of the epididymis in mediating changes in the male gamete during maturation. Adv Exp Med Biol 1995;377:87–101. 165. Ezer N, Robaire B. Gene expression is differentially regulated in the epididymis after orchidectomy. Endocrinology 2003;144:975–88. 166. Syntin P, Dacheux JL Dacheux F. Postnatal development and regulation of proteins secreted in the boar epididymis. Biol Reprod 1999;61:1622–35.
167. Hess RA, Bunick D, Lee KH, Bahr J, Taylor JA, Korach KS, Lubahn DB. A role for estrogens in the male reproductive system. Nature 1997;390:509–12. 168. Eddy EM, Washburn TF, Bunch DO, Goulding EH, Gladen BC, Lubahn DB, Korach KS. Targeted disruption of the estrogen receptor gene in male mice causes alteration of spermatogenesis and infertility, Endocrinology 1996;137:4796–805. 169. Lee KH, Hess RA, Bahr JM, Lubahn DB, Taylor J, Bunick D. Estrogen receptor alpha has a functional role in the mouse rete testis and efferent ductules. Biol Reprod 2000;63:1873–80. 170. Zhou Q, Nie R, Prins GS, Saunders PT, Katzenellenbogen BS, Hess RA. Localization of androgen and estrogen receptors in adult male mouse reproductive tract. J Androl 2002;23:870–81. 171. Goyal HO, Bartol FF, Wiley AA, Neff CW. Immunolocalization of receptors for androgen and estrogen in male caprine reproductive tract tissues: unique distribution of estrogen receptors in efferent ductule epithelium. Biol Reprod 1997;56:90–101. 172. Hejmej A, Gorazd M, Kosiniak-Kamysz K, Wiszniewska B, Sadowska J, Bilinska B. Expression of aromatase and estrogen receptors in reproductive tissues of the stallion and a single cryptorchid visualized by means of immunohistochemistry. Domest Anim Endocrinol 2005;29: 534–47. 173. Bilinska B, Hejmeh A, Gancarczyk M, Sadowska J. Immunoexpression of androgen receptors in the reproductive tract of the stallion. Ann N Y Acad Sci 2005;1040: 227–9. 174. Parlevliet JM, Pearl CA, Hess MF, Famula TR, Roser JF. Immunolocalization of estrogen and androgen receptors and steroid concentrations in the stallion epididymis. Theriogenology 2006;66:755–6. 175. Yamashita S. Localization of estrogen and androgen receptors in male reproductive tissues of mice and rats. Anat Rec 2004;279A:768–78. 176. Nie R, Zhou Q, Jassim E, Saunders PT, Hess RA. Differential expression of estrogen receptors alpha and beta in the reproductive tracts of adult male dogs and cats. Biol Reprod 2002;66:1161–8. 177. Reyes-Moreno C, Laflamme J, Frenette G, Sirard MA, Sullivan R. Spermatozoa modulate epididymal cell proliferation and protein secretion in vitro. Mol Reprod Dev 2008;75:512–20. 178. Pearl CA, Roser JF. Lactoferrin in the stallion epididymis: localization and steroid regulation. In: Proceedings of the 39th Annual SSR Meeting, 2006, p. 153.
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Chapter 100
Puberty Noah L. Heninger
Introduction Puberty for most species, including the stallion, has traditionally been difficult to define. The universally accepted definition of puberty (meaning to ‘beget’) is first achievement of the capability to successfully sexually reproduce. This is generally characterized by the maturation of genital organs, development of secondary sex characteristics, and psychosocial aspects of adolescent development. Purists will argue that puberty in domestic animals should be strictly defined as the point in time when an individual has the capability to successfully reproduce. Others define puberty as the age at first ejaculate containing 50 million sperm with ≥ 10% motility. Previous research in Quarter Horse-type stallions, based on seminal characteristics, showed that puberty occurs at 1–2.2 years1 or 1.1–1.9 years.2 If testicular development is taken into consideration, puberty has started to occur between 1.4 and 1.8 years of age in AngloNorman stallions. These definitions have a definitive end point suitable for comparisons in research. Some stallions achieve the capability to breed, but are not allowed to demonstrate this maturity due to herd hierarchy, libido, environmental restrictions, and/or interactions with man. Anatomical abnormalities (i.e., cryptorchidism, penile deviations, etc.) may also interfere with an individual’s capability to fulfill the current defining criteria and thereby contribute to difficulties in defining the term ‘puberty’. Stallions, as described above, do not fit these research models for puberty. Puberty in itself is not a single event, but a collection of dynamic events and interactions over time that ultimately lead to the ability to produce offspring. To fully understand puberty we should attempt to define various events or stages of growth leading to a stallion’s ability to reproduce. Four rather indistinct stages/phases of development appear to exist. The first phase consists of embryonic development of the male sex organs. This should probably
also include important perinatal events (i.e., testicular descent) which are critical for proper anatomical and functional development. The second ‘quiescent phase’ consists of growth and development of the young foal, i.e., this stage is associated with a strict lack of any sexual maturation. The third phase consists of subtle developmental changes leading to normal reproductive function including achievement of spermarche (production of the first sperm), normal endocrine function, and development of libido. If the definition of puberty is ‘achievement’ of these functions, then all changes leading to this development should be considered ‘pre-pubertal’ or ‘peripubertal’. Finally, post-pubertal development leads to sexual maturation, and achievement of reproductive efficiency. Puberty should not be confused with sexual maturity. This is especially true in the stallion since sexual maturity, based on sperm numbers in ejaculates, does not occur until 4–5 years of age (long after puberty has occurred).
Embryonic development Prior to birth, primordial germ cells migrate and colonize the genital ridge of the undifferentiated fetal gonads.3 Differentiation of these gonads requires differentiation of fetal Sertoli cells and Leydig cells. Primordial germ cells divide to produce gonocytes in the male sex cords. Hormone production begins and mullerian inhibiting substance (MIS) is secreted by the Sertoli cells to regress the paramesonephric ducts by the induction of apoptosis. At birth, the stallion testis contains only indifferent supporting cells (early Sertoli cells), gonocytes (early spermatogonia),4 and interstitial cells (i.e., undifferentiated Leydig cells, fibroblasts and myoid cells). Leydig cells, present in high numbers, decline rapidly before birth, when numbers remain relatively stable until pre-pubertal changes occur. Descent of the testes generally occurs between 30 days before and 10 days after birth.5 The
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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gubernaculum testis is a mesenchymal cord that contains fibroblasts, collage, and mucopolysaccharides.6 Proximally, the gubernaculum attaches to the caudal pole of the testis and its distal attachment runs through the inguinal canal and fixes to the scrotum. Failure of the testis to descend into the appropriate location in the scrotum is termed cryptorchidism. This condition can be unilateral or bilateral. A testis that has descended through the inguinal ring, but remains in the inguinal canal, is termed a ‘high flanker’. Spermatogenesis for testes in the inguinal canal is impaired due to inappropriate thermoregulation, but may depend on relative location within the canal. During the first few months following birth, testicular function remains relatively quiescent. This extended period (encompassing approximately the first 9 months of a young stallion’s life) appears to be characterized by limited gonadotropin secretion from the adenohypophysis, and limited steroidogenesis. A lack of secretion of gonadotropin-releasing hormone (GnRH) and low numbers of GnRH receptors have been suggested as the basis for this relatively quiescent gonadotropin state, based on data from bulls. 7 This is postulated to occur by negative feedback mechanisms of estrogen on the hypothalamus and adenohypophysis existing before 16 weeks.4 Further studies are necessary to elucidate these mechanisms in the young stallion. Gonocytes and Sertoli cells persist in the male sex cords (rudimentary seminiferous tubules) until peripubertal events begin to occur. At this time, gonocytes divide to produce the first spermatogonia. In most species, Sertoli cells proliferate extensively before puberty. The number of Sertoli cells at puberty are thought to remain for the breeding life of the animal. Stallions appear quite unique in this respect and research suggests that Sertoli cells in this species maintain the capability to divide prior to each breeding season.8 The number of Sertoli cells within the testis is important because it is ultimately a major determining factor in level of sperm production.
Endocrine events One of the first significant changes in stallion endocrine development occurs about 9 months of age (Figure 100.1). Increased frequency and amplitude of pulses of luteinizing hormone (LH) and folliclestimulating hormone (FSH) in response to GnRH occur at 9–10 months.2 For this study, early pulses did not result in measurable increases of testosterone in peripheral circulation. Leydig cells appeared to be unresponsive to circulating gonadotropins since testosterone remained at baseline levels. Significant testosterone production for this particular study did not occur until almost 80 weeks of age, during the
Figure 100.1 Concentrations of luteinizing hormone (LH), follicle-stimulating hormone (FSH), and testosterone in serum prepared from jugular blood. Age at puberty averaged 83 weeks, but ranged from 56 to 97 weeks (based on seminal characteristics). Based on data for eight colts born in July or early August. Blood samples were taken from each colt at 8, 16, and 24 weeks of age and every 4 weeks thereafter. From Pickett et al.36 with permission
spring of the second year of age. Seasonal effects likely influence the secretion of testosterone at this time. However, the 12-month interval between initial surges of gonadotropins from the adenohypophysis and the production of testosterone may indicate other roles for gonadotropins in early development prior to the increase in testosterone. It is possible that the rise in LH without a concomitant surge in testosterone may imply a function to mature the Leydig cells, and that the rise in FSH may trigger/facilitate the initiation of spermatogenesis. This important point should be noted and will be discussed later in the chapter.
Histological development of spermatogenesis As spermatogenic development progresses, spermatogonia (AS or A0 ) differentiate into the various morphological forms (A1–A4 and B1–B2) of spermatogonia, giving rise to more advanced generations of germ cells (primary spermatocytes and spermatids) as pubertal development progresses. Seminiferous tubules begin to form a tubule lumen by unidirectional fluid secretion forming small vacuoles in the area of the developing tubule lumen,9, 10 forming what will be ducts to transport mature spermatozoa from their site of production in the seminiferous tubule to the epididymis. Spermatogonia differentiate to produce spermatocytes by 12 months and round spermatids by 16 months of age. Complete spermatogenesis, with a full complement of germ cells, is generally observed in most tubules by 24–36 months of age.8 However, the timing of the appearance of specific germ cells during puberty
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Puberty depends heavily on the region of the testis sampled, season, and stallion-to-stallion variation. These histological observations roughly correspond to some stallions having spermatozoa available for ejaculation by 14 months and significant sperm production by 83 ± 2.9 weeks (20 months).4 Another study11 supporting this research demonstrated that daily sperm production per gram of testicular parenchyma for stallions castrated during the long summer months was 6 × 106 at 2–3 years, and 18 × 106 (i.e., a 300% increase) at 4–5 years of age.
Initiation of spermatogenesis Initiation of spermatogenesis for most mammalian species, except the stallion, begins with the conversion of gonocytes into spermatogonia in random, scattered locations throughout the mammalian testis.12 This results in uniformity of parenchymal coloration or shading13 across the testis. In the stallion, development of spermatogenesis begins centrally and evolves progressively toward the periphery.12, 14, 15 This pattern of development may suggest that some local control mechanisms exist for the initiation of spermatogenesis.16 A grossly observed section through a developing testis from a 1- to 2-year-old stallion reveals both ‘light’ testicular parenchyma, with expanding seminiferous tubules in the center, and surrounding ‘dark’ testicular parenchyma in the periphery where tubules are less developed.17 The spermatogenically inactive dark tissue gives way to spermatogenically active light tissue as development progresses.13 The observable difference in color is attributed to an abundance of fetal Leydig cells and numerous large macrophages (pigment cells) in the interstitium of dark immature parenchyma.16 Advancing pubertal development leads to increased volume density of germ cells and tubular lumen formation, as well as a decreased density of macrophages and fetal Leydig cells in the light tissue.13 To date, the stallion is the only known mammal that develops regionally in this manner. This provides an excellent research model to study multiple phases of testicular development within the same testis from the same animal. Seminiferous tubule cross-sections were classified according to a modification of the lumen scoring system previously reported by Clemmons et al.13 Testes from 67 Quarter Horse-type yearling stallions (aged 8–13 months) were used to establish a normal pattern of development (N.L. Heninger, unpublished observations). Testicular weights for these stallions ranged from 8.5 to 68.7 g. Seminiferous tubules from light, dark, and intermediate areas were histologically examined based on patterns of gross development. Samples
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Figure 100.2 Classification of the equine seminiferous epithelium during the initiation of spermatogenesis based on lumen score (LS; 1–7). Scale bar represents 40 µm. Lumen score 1, seminiferous tubule contains only gonocytes/spermatogonia and Sertoli cells which are randomly dispersed throughout the tubule. There is no evidence of vacuole formation. Tubule appears quiescent. Lumen score 2, tubule contains centrally located gonocytes/spermatogonia and Sertoli cells which have migrated to the basal compartment. A single formed vacuole is present. Lumen score 3, tubule contains gonocytes/spermatogonia and occasional early (leptotene and zygotene) spermatocytes. Several independent, randomly located vacuoles are present within the seminiferous epithelium. Lumen score 4, tubule contains several connecting/aggregating vacuoles. The most advanced germ cell type is the pachytene spermatocyte. Lumen score 5, tubule is characterized by an open lumen and a thin (one cell thick) seminiferous epithelium which contains only spermatogonia and Sertoli cells. Lumen score 6, tubule is characterized by a complete lumen and an incomplete complement of germ cell types. Lumen score 7, tubule is characterized by a full complement of germ cells and is capable of classification by the stage of seminiferous epithelium (similar to adult tubules) according to previously reported methods.33
were classified in terms of events leading to the formation of a seminiferous tubule lumen (lumen score) and the presence of advanced germ cell types (Figure 100.2). The progression of seminiferous epithelium development was observed under bright-field microscopy. Lumen scores (LSs) were assigned for each tubule based on the number of germ cell types present (spermatogonia, spermatocytes, round spermatids, and elongated spermatids), the number or aggregation of vacuoles, and the formation of a complete lumen. Tubules were scored on a scale of 1 (quiescent tubule with no lumen) to 7 (complete lumen formation with a full complement of germ cell types) based on the naturally occurring progression of development observed in these young stallions (for a full description, see the legend to Figure 100.2). Although a somewhat subjective observation, LS tubules 1–4 were most often found in the dark parenchyma, and LS 5–7 were found exclusively in light parenchyma. A simple schematic map based on observable developmental events demonstrates the full range of seminiferous tubule development during the initiation of spermatogenesis in the stallion (see Figure 100.7).
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Figure 100.3 Initiation of spermatogenesis in a single, representative pubertal stallion testis. (a) Tubule development and initial lumen formation in light testicular parenchyma (LT with white arrows) (lumen score 2–4) in central regions of the testis surrounding the mediastinum (M) and central vein (CV). (b) Tubule development and lumen formation (within the same testis) beginning separately, and concurrently, in the testicular parenchyma proximal to the epididymis. The vaginal tunic (TN) was partially dissected from the periphery of the testis, leaving only a small segment of the tunic to indicate the histological orientation of each testis to indicate the dorsal pole. Spermatogenically inactive dark tubules (DT with black arrows) (lumen score 1) surround the region of developing tubules in both panels. Scale bar, 230 µm at 10× magnification.
Little is known about the specific signals along the neuroendocrine pathway that initiate testicular development. Testicular development has been shown to vary by breed, season, and the individual. At birth, testis weights may be 8–13 g but vary by breed. Testicular weights are known to remain relatively constant until 9 months of age. At the earliest initiation of spermatogenesis (testis weights roughly 8–10 g) lumen development within tubules begins centrally around the mediastinum and separately, but concurrently, in the testicular parenchyma located closest to the epididymis at the middle dorsal pole (Figures 100.3 and 100.4). Development progresses from both the mediastinum and the dorsal pole of the testis until all tubules in between the two locations have initiated
vacuole/lumen formation. Development then progresses outwards, peripherally creating a ‘horseshoeshaped’ ring of less developed parenchyma along the outer distal boundaries. Initiation of tubule development proceeds until no dark parenchyma can be found (i.e., when testes weight exceeds 50 g). Initiation in both areas occurred in all observed cases adjacent to vascular areas. This new finding may challenge current views that spermatogenesis initiates locally. If spermatogenesis begins separately and concurrently in two locations it is possible that specific endocrine signals (i.e., the gonadotropins FSH and LH) may play a more specific undefined role for the initiation of testicular development than previously thought. The timing of this early
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Figure 100.4 Schematic representation depicting testicular development in medial cross-sections. TN, tissue marker representing location of the epididymis and dorsal pole; CV, the central vein of the testis. A testis with dark parenchyma and no active tubules (a). Initiation begins centrally around the mediastinum and separately, but concurrently, in the testicular parenchyma located closest to the epididymis at the middle dorsal pole (b). As development progresses from dark parenchyma to light a horseshoe-shaped ring of dark parenchyma surrounds the lighter, more spermatogenically active, areas (c and d). A testis with active tubules containing no dark tissue (e). Testis size (not represented) increased from 8 g to 50 g, respectively (a–e).
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Puberty development observed corresponds to previous studies noting the initial surge of gonadotropins at 36–45 weeks (9–12 months). Speculation as to the nature of additional mechanisms suggest localized changes in vasculature,18, 19 paracrine/autocrine signaling, and/or patterns of gene expression locally. Our research supports the availability of a small number of sperm cells for ejaculation by 14 months of age in some stallions. Stallions ejaculating sperm at 14 months likely have proportional amounts of dark parenchyma and light parenchyma capable of producing mature sperm. The most important factor for relative sperm production appears to be testicular size (L. Johnson, personal communication). In these Quarter Horse-type stallions, as testis weights increased from baseline (8 g) to 10–12 g, active tubules were always observed to some degree in the center and periphery of the testis.
Blood–testis barrier formation Spermatogenesis cannot occur without the presence of a barrier to prevent blood (and its components) from interfering with normal development of germ cells. Endothelial cells, peritubular myoid cells, and Sertoli cells are all normal components of a functional blood–testis barrier. The most commonly studied component of the blood–testis barrier are the tight junctions (protein bonds) between Sertoli cells.20 Logically, these apical junctions serve to isolate advanced germ cells during development. These bonds are postulated to isolate non-self cells to prevent immunological destruction by major histocompatibility complexes (MHCs). Type A spermatogonia along the basement membrane are not isolated by these bonds between adjacent Sertoli cells. Seminiferous tubule lumen formation has been used as an indirect marker for establishment of a functional blood–testis barrier.9 It has also been reported that the appearance of a tubule lumen within the seminiferous tubules coincides with the development of tight junctions between Sertoli cells of many species.10, 19, 21, 22 In pubertal animals, the formation of the small lumina (vacuoles) has been shown to be due to an increase in fluid secretion by the Sertoli cells which occurs in concert with the formation of the Sertoli cell junctional complexes.9, 10 For the stallion, formation and aggregation of vacuoles (the initial formation of a tubule lumen) may coincide with the formation of a Sertoli cell barrier.23 To assess the presence of a Sertoli cell barrier according to the various LSs (as characterized by lumen scores; see Figure 100.2), testes from six yearling stallions (aged 9–14 months) were obtained by surgical castration (N.L. Heninger, unpublished ob-
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servations). Using previously established methods,24 testes were perfused via the testicular artery for 30–60 minutes (or until firm) with either 2% glutaraldehyde (450 mOsm) or 2% glutaraldehyde with 10% dextrose (1214 mOsm). The presence/absence of the Sertoli cell barrier was examined in tubules which contain advanced germ cells (i.e., primary pachytene spermatocytes or round spermatids, because these cell types are expected to reside in the adluminal compartment). A lack of shrinkage artifact for cells in the adluminal compartment in the presence of a vascularly perfused hypertonic (i.e., high dextrose concentration) fixation fluid suggests the presence of an impermeable barrier (i.e., a Sertoli cell barrier). Early spermatocytes (i.e., leptotene and zygotene spermatocytes) could not be used to evaluate the presence of a barrier due to the uneven distribution of chromatin in the nuclei of these cells and their basal location within the tubule. Less developed tubules (LSs 1 and 2) were examined without advanced germ cells, since these tubules only rarely contained advanced germ cell types. Interestingly, the hypertonic fixation test revealed profound shrinkage of all seminiferous tubules and interstitial cells (Leydig cells and macrophages). Within crosssections of seminiferous tubules, shrinkage of every cell type (germ cells and Sertoli cells) was evident for LSs 1–4 (Figure 100.5a,b,c). In LS 4 tubules (the first LS with pachytene spermatocytes), shrinkage artifact was detected in pachytene spermatocytes, indicating that an impermeable barrier to high concentrations of dextrose had not been formed. LS 5 tubules demonstrated shrinkage artifact for both spermatogonia and Sertoli cells, but assessment of the Sertoli cell barrier was not possible since advanced germ cell types are not present in this LS. In LS 6 tubules (the first LS with round spermatids) (Figure 100.5d), advanced germ cells (pachytene spermatocytes and round spermatids) in the adluminal compartment showed no signs of shrinkage artifact. The most advanced tubules (LS 7 tubules) in pubertal stallions with a full complement of germ cells demonstrated pronounced shrinkage artifact for early germ cells such as spermatogonia as well as preleptotene, leptotene (Figure 100.5e), and zygotene (Figure 100.5f) spermatocytes in the basal compartment only. Advanced germ cells in the adluminal compartment showed no signs of shrinkage. In short, these findings support the notion that an intact Sertoli cell barrier is formed in LS 5 and that a functional barrier exists for LSs 6 and 7. One notable point is that differentiating germ cells were found in increasing numbers during early seminiferous tubule development (LSs 1–4). This development occurred prior to the formation of a Sertoli cell barrier (as judged by the hypertonic fixation
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Figure 100.5 Hypertonic fixation test according to lumen score (LS). Marked shrinkage of all seminiferous tubules occurred due to the hypertonic fixative. (a1 and a2) The effects of either 2% glutaraldehyde or 2% glutaraldehyde with 10% dextrose (hypertonic fixative) on LS 1 tubules. All other control tubules were similar to tubules in Figure 100.1. Exaggerated shrinkage artifact (arrows) due to the hypertonic fixative can be observed for all germ cells and Sertoli cells within the seminiferous epithelium of LS 1–5 as shown in (a2, b, and c), indicating that an impermeable barrier (Sertoli cell barrier) to high concentration of dextrose had yet to be formed. (b) A LS 4 tubule with shrinkage artifact surrounding advanced germ cells (white arrows) and the lack of a Sertoli cell barrier. (d) The effects of the same hypertonic fixative on LS 6 tubules. Marked shrinkage artifact (arrows) occurred in the basal compartment of the seminiferous tubule, while no shrinkage artifact was observed for advanced germ cells (arrowheads; pachytene spermatocytes) in the adluminal compartment indicating the presence of an intact Sertoli cell barrier. (e and f) LS 7 tubules with a full complement of germ cells (stages IV and I, respectively) and an intact Sertoli cell barrier indicated by shrinkage artifact in the basal compartment including leptotene (L) and zygotene (Z) spermatocytes. Scale bar, 40 µm.
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Puberty test). Previous studies seem to support these results. One previous study found that spermatogenesis can proceed in the absence of a Sertoli cell barrier up to, and including, the DNA synthesis phase of spermatocytes without triggering an immune response.25 Collectively, these results suggest that initial differentiation of germ cells occurs prior to, or perhaps stimulates the formation of, a Sertoli cell barrier. Development to the round spermatid stage, however, was never observed before LS 6. This was determined by the region within the testis that the tubule was located. The lumen formation phase begins with the development of a single vacuole (LS 2). Small vacuoles increase in number (LS 3) and then aggregate (LS 4) to initiate the development of a tubule lumen. Fluid secretion alone, even without the presence of a barrier, has been suggested to be responsible for the formation of very small lumina.10 Testicular perfusion with a hypertonic fixative caused cytoplasmic and nuclear shrinkage artifact in conjunction with exaggerated extracellular space throughout the seminiferous epithelium of all early tubules (LSs 1–5), indicating that a permeability barrier (Sertoli cell barrier) to high concentrations of dextrose had yet to be formed. Therefore, in the stallion, formation of small vacuoles in early LS tubules is likely due to secretion of fluids by Sertoli cells prior to the formation of a Sertoli cell barrier. The final process in lumen formation results in a dramatic increase in tubule lumen diameter (LS 5) coupled with a reduction in seminiferous epithelium height (reduced/spread to a single cell layer). One previous study in the mouse noted that a gradual increase in lumen diameter occurred prior to formation of the Sertoli cell barrier, but that more than 90% of the increase took place after the Sertoli cell barrier was formed, reflecting an increased tightness of the Sertoli cell barrier, increased fluid secretion by Sertoli cells, and/or remodeling of structural elements with increased tubule size.10 Applying this information to the developmental pattern observed in the pubertal stallion would imply that the Sertoli cell barrier is formed early in LS 5 when the contact between Sertoli cells is maximized. This is observed with the dramatic difference in lumen size between LS 4 and LS 5 (Figure 100.2). The hypertonic fixation test appears to be an effective method for determining the initial formation of a permeability barrier but it is not a sensitive method for determining the degree of tightness between the junctions.10 This test detects only the presence or absence of a Sertoli cell barrier, and future studies should consider the use of exclusion properties of molecules to compare with current results. Advanced germ cells in the adluminal compartment of LS 4 demonstrated shrinkage artifact, but this was
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not observed for advanced germ cells in the adluminal compartment of LS 6. It should then be considered that formation of the Sertoli cell barrier occurs in LS 5, coinciding with the increase in lumen diameter. Results of this study also demonstrate that LS 5 tubules can be found throughout the range of testicular weights used in this study. Therefore, we conclude that, for the stallion, the Sertoli cell barrier appears to develop regionally, over time (many months), rather than asynchronously along the length of a tubule,26 or by stage of seminiferous epithelium27–29 as suggested in other species. Additional studies are required to elucidate the dynamics of the Sertoli cell barrier and tubule lumen formation in the stallion.
Apoptosis during the initiation of spermatogenesis Apoptosis or ‘programmed cell death’ is a process of orderly cell destruction minimizing inflammation, and recycling components for further use. Apoptosis has been a large focus of testicular research in many species, including the stallion. In other species, an early physiological apoptotic wave occurs among testicular germ cells during the first division and differentiation. This has been shown to be necessary for the development of normal, mature spermatogenesis.29–31 The significance of this apoptotic wave during the first round of spermatogenesis has not been elucidated in detail. Studies in mice that inhibit apoptosis by knocking out key apoptotic genes have resulted in accumulation of spermatogonia and spermatocytes and subsequent infertility presumably due to inadequate formation of Sertoli–Sertoli cell junctions.31, 32 Apoptosis of testicular germ cells has been shown to be a normal occurrence during spermatogenesis of mature stallions.33 Rates according to stage of seminiferous epithelium have been established. Research involving the stallion has also shown that apoptosis is a vital part of early events during testicular development. Large numbers of degenerating apoptotic germ cells can be detected during the initiation of spermatogenesis. It is thought that germ cell apoptosis serves to remove germ cells from the seminiferous epithelium at a critical time during seminiferous epithelial development to facilitate the formation of the Sertoli cell barrier. Ten testes from different horses were chosen for the evaluation of LS per unit area, and the number of apoptotic germ cells per unit area within the predominant LS (described below). Testis samples were selected (based on testicular weight, development, and completeness of the histological section) to represent the complete range of parenchymal development
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[all dark (n = 2), mostly dark with some light (n = 3), mostly light with some dark (n = 3), and all light (n = 2)] observed in these young stallions. Tissue samples from three adult Quarter Horse stallions (≥ 5 years) with normal testes size and semen quality (as determined by a previous study in our laboratory33 ) were used for comparison with the pubertal stallions. Immunohistochemical apoptotic detection of testicular germ cells was performed on 5-µm sections, usR ing the Apoptag (Intergen Co., Purchase, NY) TdTmediated dUTP nick end labeling (TUNEL) assay, previously validated for use in the stallion.33 The predominant LS per unit area (squares containing at least 75% of a representative LS in a gridbased counting system) and the number of TUNELpositive germ cells within seminiferous tubules representing the predominant LS were recorded. Using the data generated by this analysis, representative schematic maps were created to facilitate an understanding of the developmental process for the stallion (see Figure 100.7). The predominant LS for each respective square was color-coded using various shades of blue as follows: the darkest blue represents immature, undeveloped parenchyma (LS 1), with progressively lighter shades of blue reflecting progressive seminiferous tubule development and higher LSs (LS 2–7). Squares with three or more (empirically determined to account for the low level of apoptosis observed in LS 1–3 tubules) apoptotic germ cells per square were identified and shaded red. Numbers within blue or red squares represent the predominant LS for that square. Squares containing less than an estimated 25% seminiferous tubules (interstitial space, vasculature, visceral tunic, and mediastinum) are unnumbered and color-coded gray. The number of apoptotic germ cells was not recorded for these squares. An estimated 20% of each testis was not observed due to shading from the grid lines between each square. Results of this study are shown in Figure 100.6. A small, primary wave of apoptosis occurred prior to the development of a complete lumen. LS 4 tubules had a higher mean number of apoptotic cells within the predominant LS per unit area (P < 0.05) than LS 1–3 or LS 5 tubules (Figure 100.6a). A second, larger apoptotic wave occurred after the formation of a complete lumen. LS 6 tubules had a higher number of apoptotic cells per unit area than all other LS tubules. The rate of germ cell apoptosis in LS 7 tubules was similar to the level found in adult seminiferous tubules. Schematic, color-coded maps were generated as a result of recording the predominant LS for each square, and the number of TUNEL-positive germ cells within the predominant LS for each square (Figure 100.7). Waves of regional development cor-
responding to LS in shades of blue are evident. A similar relationship was seen for the percentage of unit area with at least three apoptotic cells and LS when compared with the mean number of apoptotic germ cells (Figure 100.6b). Squares containing three or more apoptotic germ cells per unit area (flagged red) completely surround the mediastinal area and indicate a regionally higher apoptotic rate for LSs 6 and 7 when compared with less mature parenchyma in the periphery. The magnitude of the first increase in the number of apoptotic cells in LS 4 tubules was minimal, though statistically significant. Previous estimates of the number of germ cells removed by this primary wave of apoptosis are > 80%.30 For our study, the increased apoptotic rate in the early wave arises from the degeneration of a small number of germ cells engaged in spermatogenesis prior to the formation of a tubule lumen and a Sertoli cell barrier. Germ cells that enter spermatogenesis during LSs 1–4 must be removed from the developing seminiferous epithelium, since no differentiated germ cells are present in LS 5 tubules. Therefore, it can be suggested from our study that, during the initial differentiation of germ cells, apoptosis serves as an important mechanism to remove these germ cells from the seminiferous epithelium. Once germ cells have been removed, Sertoli cell contact is maximized so that attachment/organization of junctional proteins34 and the establishment of a functional barrier can ensue. The second increase in number of apoptotic cells in LS 6 tubules (Figure 100.6) is threefold larger than the first and corresponds to removal of the first germ cells that could potentially complete spermatogenesis. This ‘second initiation’ of spermatogenesis after LS 5 consists of several consecutive cycles of the seminiferous epithelium, each developing further than the previous cycle. The extraordinarily high losses of germ cells (above that observed for normal, mature spermatogenesis) may be attributed to a gaining of ‘competence’ perhaps associated with the complexity of producing haploid sperm cells. During pubertal development in the stallion, the number of germ cells have been shown to dramatically increase, while the number of Sertoli cells per gram decreases.13 Apoptosis would be a logical mechanism for establishing a critical germ cell–Sertoli cell ratio beyond the capacity of a single Sertoli cell.31 Another explanation for the dramatic increase in apoptosis might be the absence of survival signals secreted by round spermatids.35 Although round spermatids were found in LS 6, their numbers may be too few for effective signal propagation. In summary, recent research provides new insight into pubertal development for the stallion. Much
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Average number of apoptotic cells per unit area
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(b) Figure 100.6 Mean number of apoptotic germ cells per unit area (a) and the percentage of unit area with at least three apoptotic germ cells (b) according to the predominant lumen score within a square for yearling (n = 10) and adult (n = 3) stallions. Means with different superscripts (A, B, C, D) are different (P < 0.05).
information has been extrapolated from data in other species, but, as more data become available, a better understanding of the stallion’s unique method for testicular development may contribute to a better understanding of the initiation of spermatogenesis across all species. Quite literally, a cross-section through a developing testis reveals the entire range of spermatogenic development from start to finish within a single testis! This pattern of testicular development can allow future studies to more closely examine treatment effects (i.e., toxins, glucocorticoid therapy) on spermatogenic development. Initial germ cell differentiation precedes the formation of a
Sertoli cell barrier. However, formation of the barrier coincides with the development of a well-defined tubule lumen. Critical roles for both local and endocrine control clearly exist during the initiation of equine spermatogenesis. Higher rates of germ cell apoptosis that occur during specific steps of seminiferous tubule development can be viewed grossly on schematic maps. Apoptosis is clearly involved in initiation of normal spermatogenesis, although its role is still ambiguous. Future studies should exploit the unequivocal pattern of testicular development in the stallion for a better understanding of the events leading to the initiation of spermatogenesis and puberty.
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(c) Figure 100.7 Schematic color-coded (see color picture on enclosed DVD) maps representing pubertal development in stallion testis. The predominant lumen score for each respective square (600 µm2 ) is represented by various shades of blue. The darkest blue represents immature, spermatogenically inactive parenchyma (dark tubules), with lighter shades of blue representing progressive seminiferous tubule development (light tubules) and a higher lumen score (identified by the number in the square). Squares with three or more apoptotic germ cells per square (empirically determined) were shaded red. Squares containing less than an estimated 25% seminiferous tubules [interstitial space, vasculature, visceral tunic indicating histological orientation at the dorsal pole of the testis∗ , and mediastinum (M)] are unnumbered, and color-coded gray. The number of apoptotic germ cells was not recorded for these squares. (a; 9.2-g testis) The initiation of spermatogenesis centrally near the mediastinum, and separately, but concurrently, in the parenchyma near the dorsal pole of the testis (arrow). (b; 18.4-g testis) Regional, progressive tubule development starting from the center, toward the periphery with a horseshoe-shaped ring of less developed tubules (lumen scores 1 and 2) in the outer margins. (c; 52.2-g testis) Progressive, regional seminiferous tubule development and regionally higher apoptotic rates in the center of the testis (apoptotic wave) (lumen scores 6 and 7) surrounding the mediastinum.
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Puberty
References 1. Cornwell JC. MS thesis. Seasonal variation in stallion semen and puberty in the Quarter Horse colt. Louisiana State University, Baton Rouge, LA, 1972. 2. Naden J, Amann RP, Squires EL. Testicular growth, hormone concentrations, seminal characteristics and sexual behavior in stallions. J Reprod Fertil 1990;88:167–76. 3. Mintz B. Continuity of the female germ cell line from embryo to adult. Arch Anat Microsc Morphol Exp 1959;48(Suppl):155–72. 4. Amann RP. Physiology and endocrinology. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp 658–85. 5. Bergin WC, Gier HT, Marion GB, Coffman JR. A developmental concept of equine cryptorchism. Biol Reprod 1970;3:82–92. 6. Johnston SD, Kustritz MVR, Olson PNS. Canine and Feline Theriogenology. Philidelphia: Saunders, 2001; pp. 275–86. 7. Amann RP, Wise ME, Glass JD, Nett TM. Prepubertal changes in the hypothalamic-pituitary axis of Holstein bulls. Biol Reprod 1986;34:71–80. 8. Johnson L, Varner DD, Thompson Jr DL. Effect of age and season on the establishment of spermatogenesis in the horse. J Reprod Fertil Suppl 1991;44:87–97. 9. Curtis SK, Amann RP. Testicular development and establishment of spermatogenesis in Holstein bulls. J Anim Sci 1981;53:1645–57. 10. Russell LD, Bartke A, Goh JC. Postnatal development of the Sertoli cell barrier, tubular lumen, and cytoskeleton of Sertoli and myoid cells in the rat, and their relationship to tubular fluid secretion and flow. Am J Anat 1989;184: 179–89. 11. Johnson L, Thompson Jr DL. Age-related and seasonal variation in the Sertoli cell population, daily sperm production and serum concentrations of follicle-stimulation hormone, luteinizing hormone and testosterone in stallions. Biol Reprod 1983;29:777–89. 12. Courot M, Hochereau-de Reviers M-T, Orvant R. Spermatogenesis. In: Johnson AD, Gomes WR, VanDemark NL (eds) The Testis, vol. 1.New York: Academic Press, 1970; pp. 339–442. 13. Clemmons AJ, Thompson Jr DL, Johnson L. Local initiation of spermatogenesis in the horse. Biol Reprod 1995;52:1258–67. 14. Bouin P, Ancel P. La glande interstitelle du testicule chez le cheval. Arch Zool Exp Gen 1905;3:391–437. 15. Nishikawa Y, Horie T. Studies on the development of the testes and epididymides of the horse. 1. Studies on the development of the testes of the horse, with special reference to the singularity and the age of sexual maturity. Nat Inst Agric Sci Jpn Bull Ser G Anim Husb 1955;10: 229–349. 16. Johnson L, Varner DD, Tatum ME, Schrutchfield WL. Season but age affects Sertoli cell number in adult stallions. Biol Reprod 1991;45:404–10. 17. Cai L, Hales BF, Robaire B. Induction of apoptosis in the germ cells of adult male rats after exposure to cyclophosphamide. Biol Reprod 1997;56:1490–7. 18. Kormano M. An angiographic study of the testicular vasculature in the postnatal rat. Z Anat Entwicklungsgesch 1967;126:138–53. 19. Setchell BP, Pollanen P, Zupp JL. Development of the blood-testis barrier and changes in vascular permeability at puberty in rats. Int J Androl 1988;11:225–33.
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20. Wong C, Cheng CY. The blood-testis barrier: its biology, regulation, and physiological role in spermatogenesis. Curr Top Dev Biol 2005;71:263–96. 21. Gondos B. Development and differentiation of the testis and male reproductive tract. In: Steinberger A, Steinberger E (eds) Testicular Development, Structure and Function. New York: Raven Press, 1980; pp. 3–20. 22. Vitale R, Fawcett DW, Dym M. The normal development of the blood-testis barrier and the effects of clomiphene and estrogen treatment. Anat Rec 1973;176:331–44. 23. Heninger NL, Staub C, Johnson L, Blanchard TL, Varner D, Ing NH, Forrext DW. Testicular germ cell apoptosis and formation of the Sertoli cell barrier during the initiation of spermatogenesis in pubertal stallions. Anim Reprod Sci 2006;94:127–31. 24. Johnson L, Neaves WB. Age-related changes in the Leydig cell population, seminiferous tubules, and sperm production in stallions. Biol Reprod 1981;24:703–12. 25. Pelletier RM. Cyclic formation and decay of the blood-testis barrier in the mink (Mustela vison), a seasonal breeder. Am J Anat 1986;175:91–117. 26. Vignon X, Terquem A, Dadoune JP. The postnatal development of the junctional complexes of hamster Sertoli cells as revealed by HRP and freeze-fracture. J Submicrosc Cytol 1987;19:303–9. 27. Bergmann M, Dierichs R. Postnatal formation of the bloodtestis barrier in the rat with special reference to the initiation of meiosis. Anat Embryol (Berl) 1983;168:269–75. 28. Cavicchia JC, Sacerdote FL. Correlation between bloodtestis barrier development and onset of the first spermatogenic wave in normal and in busulfan-treated rats: a lanthanum and freeze-fracture study. Anat Rec 1991;230: 361–8. 29. Jahnukainen K, Chrysis D, Hou M, Parvinen M, Eksborg S, Soder O. Increased apoptosis occurring during the first wave of spermatogenesis is stage-specific and primarily affects midpachytene spermatocytes in the rat testis. Biol Reprod 2004;70:290–6. 30. Rodriguez I, Ody C, Araki K, Garcia I, Vassalli P. An early and massive wave of germinal cell apoptosis is required for the development of functional spermatogenesis. EMBO J 1997;16:2262–70. 31. Russell LD, Chiarini-Garcia H, Korsmeyer SJ, Knudson CM. Bax-dependent spermatogonia apoptosis is required for testicular development and spermatogenesis. Biol Reprod 2002;66:950–8. 32. Knudson CM, Tung KS, Tourtellotte WG, Brown GA, Korsmeyer SJ. Bax-deficient mice with lymphoid hyperplasia and male germ cell death. Science 1995;270:96–9. 33. Heninger NL, Staub C, Blanchard TL, Johnson L, Varner DD, Forrest DW. Germ cell apoptosis in the testes of normal stallions. Theriogenology 2004;62:283–97. 34. Knudson CM, Tung KS, Korsmeyer SJ. Molecular regulation of testicular cell death. In: Zirkin BR (ed.) Proceedings of the XIVth Testis Workshop in Germ Cell Development, Division, Disruption and Death. New York: Springer-Verlag, 1997; pp. 140–9. 35. Zhao GQ, Deng K, Labosky PA, Liaw L, Hogan BL. The gene encoding bone morphogenetic protein 8B is required for the initiation and maintenance of spermatogenesis in the mouse. Genes Dev 1996;10:1657–69. 36. Pickett BW, Amann RP, McKinnon AO, Squires EL, Voss JL. Management of the stallion for maximum reproductive efficiency, II. Animal Reproduction Laboratory Bulletin no. 05. Fort Collins: Colorado State University, 1989.
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Chapter 101
Spermatogenesis Larry Johnson, Cleet E. Griffin and Michael T. Martin
Introduction Spermatogenesis represents the processes of cellular division and differentiation by which spermatozoa are produced within seminiferous tubules of the testis. It is a lengthy process that incorporates in sequence spermatogonia, spermatocytes, and spermatids. Spermatogonia, serving as stem-type cells lining the base of seminiferous tubules, divide by mitosis to maintain their own numbers and produce primary spermatocytes in a cyclic pattern. Primary spermatocytes then undergo meiosis, which consists of one duplication of chromosomes and two divisions, to produce haploid spermatids that differentiate into spermatozoa without division; the spermatozoa are then released into the lumen of the seminiferous tubules (Figures 101.1 and 101.2). Seminiferous tubules constitute the majority of the testis (Figure 101.1). Seminiferous tubules contain two types of parenchymal somatic cells – myoid cells and Sertoli cells – and three types of germ cells – spermatogonia, spermatocytes, and spermatids (Figures 101.2 and 101.3). Leydig cells, another somatic parenchymal cell of the testis, are predominant in the interstitium between seminiferous tubules (Figures 101.1 and 101.2). The lifespan of the three types of germ cells sequentially divides spermatogenesis into three almost equal phases: spermatocytogenesis (mitosis) of spermatogonia, meiosis of spermatocytes, and spermiogenesis of spermatids (Figure 101.4). In spermatocytogenesis, stem cell (i.e., primitive/precursor capable of self-replication) spermatogonia divide by mitosis to produce other stem cells. To continue the lineage of stem cells throughout the adult life of the male progeny of these stem cells, spermatocytes divide cyclically by mitosis to produce committed spermatogonia, which immediately divide in sequence to produce primary spermatocytes. Primary spermatocytes are in the second phase of spermato-
genesis, i.e., meiosis. During meiosis, exchanges in genetic material between homologous chromosomes are followed by the first of two linked divisions. The first division separates duplicate paternal from duplicate maternal chromosomes, thereby sexing the secondary spermatocyte (e.g., each spermatid contains the duplicated paternal or duplicated maternal chromosomes but not both paternal and maternal). The second meiotic division produces haploid spermatids which mark the beginning of spermiogenesis. Spermiogenesis is the differentiation without division of spherical spermatids into spermatozoa. Spermatozoa are released (spermiated) in the luminal free surface of seminiferous tubules as spermatozoa (Figures 101.1, 101.2, and 101.4). Spermatozoa are removed hydrostatically as fluid flows through the lumen of seminiferous tubules (Figure 101.5) into the rete testis tubules (Figure 101.6). The rete testis tubules provide an outlet for the spermatozoa from the testis into the extragondal ducts (efferent ducts and epididymis) for maturation and storage.
The spermatozoon A description of the spermatozoon may be useful to understand the series of events that are necessary during spermatogenesis to create the remarkable cells which, with the oocyte, are responsible for future generations of man and many animals. A spermatozoon is composed of a head and tail (Figure 101.7), and the general structures for domestic species are similar to those described for the bull1 and the stallion.2 Horse spermatozoa are unusual in that, for 40–60% of cells, the tail is abaxial, i.e., the tail is not attached to the center of the base of the head (Figure 101.7b). This poses no known difficulty for the horse but is considered abnormal in other species. The nucleus occupies the major portion
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 101.1 Photomicrographs of the equine testis. Presented are a complete cross-section (a), a low-magnification histological section (b), and a higher magnification histological section (c). Labeled structures include: the tunica albuginea (TA), the central vein (CV), testicular parenchyma (TP), seminiferous tubules (ST), Leydig cells (LC), lymph vessels (LV), and blood vessels (BV), residual bodies (RB), spermatogonia (Sg), spermatocytes (Sa) elongated spermatids (Sd2), and Sertoli cells (SC). Bar lengths equal 10 mm for (a), 1 mm for (b), and 30 µm for (c). Reprinted with permission.86
Figure 101.2 Micrographs of the equine testis viewed by scanning electron microscopy (a and d), high-voltage transmission electron microscopy (b), and light microscopy with Nomarski optics (c). Labeled structures include: a seminiferous tubule (ST), interstitial tissue (IT), Leydig cells (LC), myoid cells (MC), Sertoli cells (SC), primary spermatocytes (PS), Sd1 spermatids (Sd1 ), Sd2 spermatids (Sd2 ), and released spermatozoa (Sp). Bar lengths equals 100 µm for (a), 3 µm for (b), and 10 µm for (c) and (d). Reprinted with permission.7, 87
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Figure 101.3 High-voltage electron micrographs of equine Sertoli and germ cells. These were embedded in Epon 812 and sectioned at 1.0 µm. The basal (a) and apical (b) regions of the Sertoli cells are shown. Labeled structures include: laminar processes (LP), an intercellular bridge (IB), nucleus of primary spermatocyte (PS), mitochondria of Sertoli cells (white arrows), mitochondria of germ cells (M), rough endoplasmic reticulum (RER), Sertoli cell nucleus (N), intercellular space (IS), Golgiphase spermatid (Sa), developing flagellum (F), centriole (Ct), portion of the nucleus and three mitochondria of an Sa spermatid (Sa), annulus (An), manchette (Mn), and flagellar canal (FC). Bar length equals 2 µm for (a) and 1 µm for (b). Reprinted with permission.41
of the head, and contains the genetic material (male genome) to be delivered to the egg. An acrosome, over the anterior surface of the nucleus, is a bag of hydrolytic enzymes necessary for penetration of the egg vestments during fertilization. The acrosomal membrane encloses an apical ridge (more distinct in bull, boar, and ram), a principal segment, and equatorial segments (Figure 101.7). The tail includes a neck region, where it is attached to the head. The middle piece contains mitochondria for energy production. The principal piece contains the fibrous sheath for flexibility and is attached to the end piece. The tail is motile due to the presence of an axoneme which runs its length and terminates in the end piece. In the horse, as in many species, there are nine dense fibers
that run to a varying degree down the length of the middle and principal piece. The plasma membrane envelops the entire cell, although it is regionally specialized. The neck region of the tail is attached to the base of the head. Both proximal and distal centrioles remain in the neck, but the distal centriole gives rise to the axoneme (nine microtubular doublets and a central pair of microtubules) during development of the tail. An axoneme (composed of a doublet and central pair of microtubules) continues through the middle piece, principal piece, and end piece. The axoneme is surrounded by nine dense fibers. The role of the outer dense fibers remains uncertain.3 Fawcett4 proposed that they function as passive elastic structures. A ring-like structure, the annulus, marks the end of the middle piece and the beginning of the principal piece. This latter area is the longest portion of the tail, and contains a fibrous sheath with rib-like structures to facilitate bending during movement and dense fibers that disappear near the distal end of the principal piece. An axoneme (composed of a doublet of microtubules and a central pair) continues through the middle piece, principal piece, and end piece. The end piece is composed of only a decreasing number of microtubules and the plasma membrane (Figure 101.7). There are species differences in size and shape of spermatozoa.5 Bull spermatozoa, like boar and ram, have flattened heads.1 A stallion spermatozoon2 is more similar in size and shape to those of humans6 and other primates. In most species, spermatozoa are not mature when released from the seminiferous tubules but require morphological and physiological changes in the epididymis to become motile and have fertilizing compatibility. Just prior to spermiation of spermatids as spermatozoa, the excess cytoplasm that is not lost by the sperm at the time of spermiation accumulates at the neck end of the developing tail. This cytoplasm becomes the cytoplasmic droplet (located in the proximal position near the head) of released spermatozoa (Figures 101.2d, 101.6b).7, 8
Germ cells in the three major events of spermatogenesis Spermatogonia (spermatocytogenesis) Spermatocytogenesis is the first major event of spermatogenesis and involves the mitotic divisions of spermatogonia of different types, resulting from a sequential set of mitotic divisions (Figures 101.4 and 101.8). Spermatogonia arise from gonocytes positioned in precursor seminiferous tubules during the second month of gestation. Spermatogonia are
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12.2 days Figure 101.4 Three major phases of spermatogenesis. Spermatocytogenesis, meiosis, and spermiogenesis are overlaid by the eight stages of the cycle of equine seminiferous epithelium. During the 19 days required for spermatocytogenesis, A-spermatogonia (A) enter cyclic activity during stage III and undergo division to produce B1 spermatogonia (B1), B2 spermatogonia (B2), and preleptotene primary spermatocytes (Pl). During the 20 days of meiosis, newly formed preleptotene primary spermatocytes (Pl) differentiate through leptotene (L), zygotene (Z), pachytene (P), and diplotene (D) before the first meiotic division to produce secondary spermatocytes (SS), and the second meiotic division to produce Sa spermatids (Sa). During the 18 days of spermiogenesis, Sa spermatids differentiate through Sb1 (Sb1 ), Sb2 (Sb2 ), Sc (Sc), Sd1 (Sd1 ), and Sd2 (Sd2 ) steps of development before their spermiation as spermatozoa. The letters indicate the cell type, while the numbers associated with the developmental step of each germ cell indicate the total time to formation of that cell type, calculated to the middle of that spermatogenic stage. In the horse, the cycle length is 12.2 days, and the duration of spermatogenesis is 57 days. Reprinted with permission.87
located at the peripheral region of the seminiferous tubules in adults. There are two types of spermatogonia. Stem cell spermatogonia divide by mitosis to produce other stem cells to continue the lineage of stem cells throughout the male’s adult life. These stem cells repopulate the testis after damage of the more advanced germ cells, or seasonal regression in some species that intermittently discontinue spermatogenesis seasonally. Committed spermatogonia resulting from stem cell divisions periodically enter a series of proliferative and/or differentiative mitotic
divisions culminating in production of primary spermatocytes. Hence, spermatocytogenesis functions to (i) continue the lineage of stem cells and (ii) produce committed spermatogonia, which result in the production of primary spermatocytes. In the horse (Figure 101.8), there are five different subtypes of spermatogonia characterized by (i) a small flattened nucleus (A1 ), (ii) a large nucleus with apparently empty centers composed of euchromatin (A2 ), (iii) the largest nucleus with either one large nucleolus or large fragments of nucleoli (A3 ), (iv) a
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Figure 101.5 An equine seminiferous tubule and some interstitium viewed by scanning electron microscopy. Clusters of Leydig cells (LC) are in the interstitium around the seminiferous tubule (ST). This seminiferous tubule contains spermatids with spherical nuclei whose tails already project into the lumen. Spermatozoa (sperm) are in transit through the lumen. The open arrows designate the height of seminiferous epithelium. Bar length equals 20 µm. Reprinted with permission.86
large oval to spherical nucleus with large chromatin flakes (intermediate or B1 ), or (v) a small spherical nucleus with small chromatin flakes (B2 ), whose division yields preleptotene primary spermatocytes.9 The numbers of each subtype of equine spermatogonia in a given cross-section of the seminiferous tubules vary with the stages of spermatogenic cycle (Figure 101.9).9 These cyclic differences in cell numbers reflect mitotic activity and degeneration of the specific subtypes of spermatogonia in different stages of the spermatogenic cycle. Likewise, the number of subtypes of spermatogonia varies with the season of the year (Figure 101.10). In the natural breeding season, the total number of spermatogonia per testis and the ‘yield’ of B2 spermatogonia per A1 spermatogonium is greater than in the non-breeding season (Figure 101.10). However, as the end of spermatocytogenesis is reached, the overabundance of B2 spermatogonia in the breeding season is so great that many degenerate rather than produce two viable primary spermatocytes. Although the yield of this division (B2 to primary spermatocytes) is less in the breeding season than in the non-breeding season, division of the overwhelmingly larger number of A1 spermatogonia in the breeding seasons produces significantly more primary spermatocytes in the breeding season than in the non-breeding season. In the stallion, and in both seasons, spermatocytogenesis produces an overabundance of spermatogonia, and germ cell degeneration is an important mechanism to reduce its yield to a manageable number of spermatocytes and spermatids.10 Hence, seasonal
Figure 101.6 Scanning electron micrographs of equine testicular parenchyma. (a) The rete testis complex (RTC) housed in connective tissue around the central vein (CV) of the equine testis. (b) A cross-support within the complex. In (a), seminiferous tubules (ST) are adjacent to the rete testis complex, which contains cross-supports (open arrow) with a simple cuboidal epithelium. At higher magnification (b) is a support (open arrow) with spermatozoa passing through. These spermatozoa have the typical proximal cytoplasmic droplet-located neck region. Bar lengths equal (a) 500 µm and (b) 4.6 µm. Reprinted with permission.88
differences in spermatogonial number largely control seasonal differences in daily spermatozoan production, even though the yield of B2 spermatogonia to primary spermatocytes is greater in the non-breeding season.10 Based on data quantifying A-spermatogonia and their mitotic activity during different stages of the spermatogenic cycle,10 a pattern for the process of spermatogonial renewal in the stallion has been proposed (Figure 101.11). Increased yield early in spermatocytogenesis (A1 spermatogonia) explains
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Figure 101.7 Photomicrographs of equine spermatozoa. Views include those produced by transmission electron (a, c–i), scanning electron (b), and phase contrast (j) microscopy. Attaching lines between (a) and (b) or (b) and (c) and between (d–i) and (j) are corresponding regions of the spermatozoon. Labeled structures include (a) the plasma membrane (PM), nucleus (N), acrosome (Ac), post-acrosomal region (PR), nuclear membrane (NM); the apical (ASA), principal (PSA), and equatorial (ESA) segments of the acrosome; the inner (IAM) and outer (OAM) acrosomal membranes; (b) and (c) the implantation fossa (IF) of the head, mitochondria (M) of the middle piece, the fibrous sheath of the tail (FS), and cytoplasmic droplets (CD) of this immature spermatozoon; (d) the central pair (CP) of microtubules and the outer nine doublet microtubules (Dl) of the axoneme, surrounded by the dense fibers (DF); (i) the single microtubules (SM) in the end piece of the tail, rather than doublets as in the middle or principal piece; (j) the middle piece (MP), principal piece (PP), and end piece (EP) of the tail. Reprinted with permission.2, 8
increased daily sperm production in the breeding season of the horse.9 The mechanism by which the number of Aspermatogonia is seasonally modulated in horses is unclear. Spermatogonial number is dependent upon (i) the number of stem cells per testis, (ii) the scheme of stem cell renewal, (iii) the number of cell divisions from the stem cell to the primary spermatocyte,11 and (iv) the amount of degeneration of spe-
cific subtypes of spermatogonia.12 Only the interval at which newly committed spermatogonia enter the spermatogenic cycle (i.e., 12-day cycle length) has been determined for the horse.13, 14 The number of spermatogonial subtypes has not been confirmed for the horse, but is considered to be 4–6 subtypes for a given species.11 This number of mitotic divisions is not likely to be changed seasonally or to account for seasonal changes in daily sperm production (DSP).
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Figure 101.8 Micrographs of spermatogonia, and primary and secondary spermatocytes viewed by bright-field microscopy. Presented are 5-µm methacrylate sections of a stem spermatogonium (As); different subtypes of spermatogonia (A1 , A2 , A3 , B1 , or B2 ); preleptotene (Pl), leptotene (L), zygotene (Z), early (EP) or late (LP) pachytene, and diplotene (D) primary spermatocytes; and secondary spermatocytes (SS). Small arrows designate the cell of interest when other cells are in view. The open arrow distinguishes the diplotene primary spermatocyte from the secondary spermatocyte designated by the solid arrow. Bar length equals 10 µm. Reprinted with permission.59
All germ cells, including spermatogonia, spermatocytes, spermatids, or residual bodies, are attached to one another by intercellular bridges (Figure 101.3). These bridges, between cells in the same developmental step (e.g., spermatocytes), may function to facilitate (i) the synchronous development or degeneration of similar germ cells, (ii) the production of committed spermatogonia,12 (iii) the differentiation of haploid spermatids, which now have only one sex chromosome, and/or (iv) the phagocytosis and digestion of residual bodies left after spermiation.
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Figure 101.9 Number of each subtype (A1 , A2 , A3 , B1 , and B2 ) of spermatogonia per 100 Sertoli cells in each of the eight stages of the spermatogenic cycle in equine testes obtained in the breeding season. Reprinted with permission.9
Spermatocytogenesis plays a pivotal role in the regulation of spermatogenesis. Meiosis is the process by which genetic material is exchanged between homologous chromosomes and haploid (half of the number of chromosomes to be joined by half of the egg’s chromosomes to produce a diploid number of chromosomes at fertilization) spermatids are produced. Meiosis occurs only in germ cells in gonads (testes and ovaries). In males, these cells are spermatocytes (Figure 101.8). After the mitotic division of B2 spermatogonia, preleptotene primary spermatocytes result (Figures 101.4, 101.8, and 101.11). Without dividing, spermatocytes differentiate (change in function and appearance); this is most obvious from changes in nuclear size and chromatin appearance (Figure 101.8). Chromatin clumps located inside and near the nuclear envelope in B2 spermatogonia and preleptotene primary spermatocytes disperse to produce the fine chromatin filaments of leptotene primary spermatocytes as these primary spermatocytes are undergoing development (Figure 101.4). The preleptotene primary spermatocytes immediately begin meiosis. During the preleptotene phase, homologous chromosomes replicate themselves15 by actively synthesizing DNA for approximately 18 hours.15 Incorporation of [3 H]-thymidine into DNA during this synthesis facilitates the study of the timing of the kinetics of spermatogenesis, as labeled cells can be followed through this process.16 This procedure allows determination of the spermatogenic cycle length (12.2 days in the horse).14 Given these data, the duration of spermatogenesis (57 days in the horse), the duration of each spermatogenic stage, and the duration of the major phases of spermatogenesis can be determined (Figure 101.4). Type B spermatogonia divide by mitosis to produce preleptotene primary spermatocytes. These develop into leptotene spermatocytes as different phases of meiosis are completed. Diplotene primary spermatocytes, which develop from the pachytene phase, complete a second meiotic division. The zygotene phase marks the beginning of the exchange of genetic material through the pairing of homologous chromosomes, which continues in the pachytene phase. The sex chromosomes are separated into the sex vesicles,17 but some exchange does occur between sex chromosomes. During the first meiotic division, primary spermatocytes rapidly undergo metaphase, anaphase (separation), and telophase (complete separation). Secondary spermatocytes result from this division (Figure 101.8) and contain the haploid number of chromosomes, but each has two conjoined copies of the DNA sequence. They have spherical nuclei
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Figure 101.10 Effect of season on numbers of germ cells. (a) The total number of A1 , A2 , A3 , B1 , and B2 spermatogonia (A1 , A2 , A3 B1 , and B2 , respectively) or primary spermatocytes (PS) per testis. (b) The yields of specific spermatogonial subtypes per B2 spermatogonium, and the yield of B2 -spermatogonia per pachytene primary spermatocytes. The conversion of A1 to B2 is greater in the breeding season. Reprinted with permission.9
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Figure 101.11 Model of the postulated pattern for renewal of equine stem cells. The formation of two new isolated and uncommitted A1 -spermatogonia and commitment of A1 -spermatogonia toward differentiation by forming a pair of undifferentiated A1.2 spermatogonia that divide to form a chain of committed but undifferentiated A1.4 - and then A1.8 -spermatogonia is presented. Spermatogenesis begins with formation of a pair of committed A1.2 spermatogonia. Commitment of another A1 -spermatogonium to form a pair of A1.2 -spermatogonia occurs only in stage III of the cycle of the seminiferous epithelium. Although degeneration of spermatogonia can occur at any point in spermatocytogenesis, degeneration during the breeding season involves mostly B2 spermatogonia, whereas in the non-breeding season it involves A1.8 - and A3 -spermatogonia. Degeneration of the B2 -spermatogonia during their mitosis in the breeding season results from an overpopulation of A-spermatogonia whose progeny are too numerous to be supported. Reprinted with permission.59, 74
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Figure 101.12 Transmission electron micrographs of different stages of development of horse spermatids. These are represented by (a–d) Sa spermatids, (e) Sb1 spermatid, (f) Sb2 spermatid, (g,h) Sc spermatids, (i) Sd1 spermatid, and (j) Sd2 spermatid at different stages of development during spermatogenesis. Labeled structures include: a Golgi apparatus (GA), vesicles (V) that fuse to form the spherical acrosomic vesicle (AV), an intracellular bridge (IB), developing flagellum (F), developing manchette (Mn), annulus (An), mitochondria (M), dense fibers (DF), and fibrous sheath (FS) of the tail, future cytoplasmic droplet (CD), and a portion of the tubular lumen (TL). Bar length equals 2 µm. Reprinted with permission.59
with chromatin flakes of varying size. Within a few hours, and without DNA synthesis, the second meiotic division occurs; this involves splitting the conjoined copies of the chromosome and results in spermatids with a haploid number of chromosomes. Hence, one duplication of chromosomes followed by two divisions results in the production of haploid numbers of chromosomes in the spermatids. Spermatids (spermiogenesis) Spermiogenesis is the morphological differentiation of spherical spermatids into spermatozoa with their unique shape (Figures 101.12 and 101.13). Spermatids, produced by the second meiotic division, differentiate from spherical cells with spherical nuclei to cells that have (i) a streamlined head with a condensed nucleus, containing the male genome; (ii) a bag containing penetrative enzymes (termed
the acrosome); and (iii) a tail that provides cellular motility. Since spermiogenesis does not involve cell division, DNA synthesis is minimal during spermiogenesis and probably reflects DNA repair. Chromatin proteins (histone) change in arginine and cystine content, and the number of disulfide bonds changes during and after elongation. Spermiogenesis is divided into Golgi, cap, elongation, and maturation phases, depending on maturation changes within the spermatid noted by shape and predominant organelles.
Golgi phase Golgi phase spermatids have a prominent Golgi apparatus that produces membrane-bound vesicles on their mature face (Figure 101.12). These small vesicles fuse to form the acrosomic vesicle adjacent to the nucleus, and this differentiates into the
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Figure 101.13 Phase contrast micrographs of equine spermatids. The micrographs are of cells in testicular homogenates, and illustrate different (Sa through Sd2 ) steps of development during spermiogenesis. Arrows indicate an acrosomic granule in (c), the first distinct appearance of a flagellum in (f), the acrosomic cap in (h), a developing flagellum in (i), the first appearance of the manchette in (j), mitochondria around the middle piece in (s), and a cytoplasmic droplet in (u). Bar length equals 5 µm. Reprinted with permission.46
acrosome, a membrane-bound, enzyme-containing vesicle over the nucleus of the spermatozoon (Figure 101.7). The centrioles migrate to a region near the nuclear envelope, where the distal centriole gives rise to the developing axoneme; initially, this is entirely within the spherically shaped cell (Figure 101.12), but later its growth is projected away from the nucleus toward the lumen (Figure 101.2). The nuclear envelope is indented, ‘nuclear indentation,’ by the developing articular fossa of the attaching flagellum (Figure 101.3). The flagellar canal is a tubular infolding of the plasma membrane of the spermatid, from its surface to the annulus located just below the attachment of the flagellum to the nucleus, through which the flagellum extends toward or into the lumen. The flagellar canal provides a mechanism by which new growth of the flagellum can be extended from the spermatid
area of the articulation fossa to allow subsequent flagellar growth. The annulus also is moved distally. Flagellation (vibrating) of maturation-phase spermatids can be observed on slides made from freshly diced seminiferous tubules. The flagellar canal also may facilitate minute flagellation (i) to assure nuclear membrane contact during flagellar attachment to the nucleus and (ii) to direct the need for ATP (energy) to the middle piece to draw mitochondria into this region where they will be attached in the spermatozoon. It is known that cells with high energy use or regions within cells with high energy, such as in the apical cytoplasm intestinal absorptive cells, have a large number of mitochondria. Alternatively, the flagellar canal (Figure 101.3) may function to prevent mitochondria from attaching to the axoneme or outer dense fibers during development of these structures.
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Cap phase When the acrosomic vesicle flattens and begins to spread, it forms a cap over the nucleus (Figures 101.12 and 101.13). The Golgi apparatus becomes less conspicuous as it moves to the opposite side (away from the acrosomic vesicle or acrosome) of the cell. Flagellar development becomes more obvious as it projects further from the cell’s surface. The nucleus is still spherical.
ovulation in females. The developing spermatid is attached to its residual body by cytoplasmic stalks in the neck region (Figure 101.2). A small amount remains in the neck region as a cytoplasmic droplet. Upon spermiation, a large portion of cytoplasm found in the spermatid is left behind, creating the residual body (Figures 101.1 and 101.2), which is phagocytized and digested by Sertoli cells. However, it appears that lysomal action within the residual body has already begun self-degradation.
Elongation phase As the cap extends further over the nucleus, the nucleus begins to elongate (Figures 101.12 and 101.13). The manchette, a transient organelle found only in spermatids, is composed of microtubules attached by linking arms and arranged in a sheath that forms around the lower half (caudal region) of the elongating nucleus and extends down and outside of the developing flagellum (Figures 101.3, 101.12, and 101.13), all within the cytoplasm, which is elongating. As spermatids develop further, they are embedded in deep recesses within the Sertoli cell cytoplasm (Figure 101.3). In the horse, components of manchette remain in the residual body that is left behind at spermiation, where it degenerates. In the horse, as in other species, the spermatozoon is not considered normal if it retains its manchette.
Maturation phase The final phase of spermatid development is characterized by migration of the manchette caudally, where it may provide a shaft that supports the flagellar canal and freedom of movement during flagellar development (Figures 101.2, 101.3, and 101.12). Indeed, the dissolution of the manchette corresponds to the migration of the annulus to its permanent location at the junction of the middle and principal pieces of the spermatozoon and the shortening and disappearance of the flagellar canal. Mitochondria move in around the flagellum in the region of the middle piece after annulus migration (Figure 101.12). This process appears to occur quickly as it is difficult to find a spermatid whose annulus has migrated but mitochondria have not surrounded its middle piece. The mechanism of production of the nine dense fibers and fibrous sheath of the spermatozoon (Figure 101.7) is unclear. The fibrous sheath is two columns attached by ribs that extend down the principal piece between the plasma membrane and outer dense fibers. The release of spermatids from the seminiferous epithelium as individual and free-floating spermatozoa is known as spermiation, the counterpart to
Kinetics of spermatogenesis Stages of the spermatogenic cycle The three phases of spermatogenesis (spermatocytogenesis, meiosis, and spermiogenesis) can be superimposed as the spermatogenic cycle (Figure 101.4). The spermatogenic cycle (cycle of the seminiferous tubular epithelium)18 is a series of changes in a given region of seminiferous epithelium between two appearances of the same developmental stages. If we use spermiation as a reference point, the cycle would be all the events that occur between two consecutive occasions of spermatozoan released from a given region of the seminiferous epithelium. Once the process of spermatogenesis starts, it continues at a defined rate, such that cells (Golgi phase spermatids) at a given developmental step are almost always associated with other cells (maturation phase spermatids) at some respective developmental step. Indeed, any combination of germ cells or classes that differ in age of development by a multiple of the cycle length will constitute a stage of the cycle. A stage in the cycle of spermatogenesis is defined as an association of four or five generations of germ cells in specific, chronological developmental steps, whose developmental age differs by a multiple of the cycle length.19 Cell division magnifies the yield in spermatogenesis. Each committed spermatogonium entering spermatogenesis has a potential yield of 64 spermatozoa. In spermatogenesis, the frequency of product release is greater than in our college model. In spermatogenesis, the production of newly committed spermatogonia is not synchronized among tubules and is not simultaneous along the length of the same tubule. Somewhere in the testis newly committed spermatogonia enter the process each second. Hence, each second, thousands of spermatozoa are being released into the lumen of the seminiferous tubules. While this continual release seems wasteful, this assures that at least some male gametes are always available, even after exhaustive sexual activity, whenever female gametes are available for fertilization.
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Figure 101.14 The eight stages (I–VIII) of the cycle of the equine seminiferous epithelium. These micrographs are presented in a clockwise arrangement, and observed by Nomarski optics in unstained, 20-µm Epon histological sections. Designated are Leydig cells (LC), Sertoli cells (SC), myoid cells (MC), and germ cells including: A-spermatogonia (AS), B-spermatogonia (B), preleptotene (Pl), leptotene (L), zygotene (Z), and pachytene (P) primary spermatocytes, secondary spermatocytes (SS), Sa spermatids (Sa), Sb1 spermatids (Sb1 ), Sb2 spermatids (Sb2 ), Sc spermatids (Sc), Sd1 spermatids (Sd1 ), and Sd2 spermatids (Sd2 ). Labeled structures include: flagella (F), acrosomal cap (AC), manchette (Mn), annulus (An), meiotic figures (MF), Golgi apparatus (GA), developing acrosomic vesicle (AV), chromatoid bodies (CB), grouped mitochondria (GM), acrosomic granule (AG), enlarged middle pieces (MP), cytoplasmic droplet (CD), and residual bodies (RB). Bar length equals 10 µm. Reprinted with permission.30
The spermatogenic cycle has been divided into stages, which are man-made divisions of the naturally occurring and continuously changing cellular associations in a given region of the seminiferous tubule (Figure 101.14). Eight stages were first described in the horse by Swierstra et al.14 Classification of stages of the spermatogenic cycle has been based on the presence and location of specific germ cells (i.e., B-spermatogonia, one or two generations of primary spermatocytes or spermatids, or secondary spermatocytes), and on changes associated with nuclear elongation, location, and release of spermatids.20 Although this has been studied and illustrated for the bull, ram, boar, and horse using conventional histology, it is presented here for the horse using differential interference contrast images of tissue embedded in Epon (Figure 101.14). This advantageous approach reveals more detail of the equine spermatogenic cycle. Details regarding differences among stages have been included in the legend to Figure 101.14.
Cycle length and duration of spermatogenesis As mentioned earlier, the length of a cycle is the amount of time between consecutive releases of spermatozoa (spermiations) and is dictated by the frequency of newly committed spermatogonia entering the process. Hence, the cycle length is neither arbitrary nor man-made. Cycle lengths, in days, for some species are 7.2 for the prairie vole, 8.6 for the boar, 8.7 for the hamster, 8.9 for the mouse, 10.4 for the ram, 9.5 for the rhesus monkey, 12.2 for the horse, 12.2 for the rabbit, 12.9 for the rat, 13.5 for the bull, 13.6 for the beagle dog, and 16 for man.13,21–23 The duration of spermatogenesis can only be approximated for any species studied, owing to the difficulty of determining the point at which committed A-spermatogonia enter the process. Usually, it has been considered to be approximately 4.5 times the cycle length for a given species.21 This corresponds to four or five generations of germ cells
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Table 101.1 Relative frequency and mean duration of stages of the cycle of the seminiferous epithelium Stage
Relative frequency (%)
Duration (days)
I II III IV V VI VII VIII
16.9 14.9 3.2 15.8 7.4 13.5 12.6 15.7
2.1 1.8 0.4 1.9 0.9 1.6 1.5 1.9
Modified from Swiestra et al.14
found in each stage of the spermatogenic cycle. The duration of spermatogenesis can be approximated at 39 days in boars, 47 days in Ile de France rams, 57 days in stallions, 60 days in rats, 61 days in bulls and dogs, and 74 days in humans.21
Frequencies of stages The frequency of each stage of the spermatogenic cycle is similar for adult members of the same species, but differs among species. The frequency of a spermatogenic stage is expressed as a percentage and is determined histologically by scoring many crosssections of the seminiferous tubule in histological sections, and dividing the number of tubular crosssections in a given stage by the total number of all tubular cross-sections scored. This frequency multiplied by the cycle length represents the duration of a given stage. From these data, and those for the presence of certain cells in certain stages, the lifespan of a given cell type can be estimated. The relative frequencies and mean durations of the eight spermatogenic stages in the horse, as defined by Swierstra et al.,14 are summarized in Table 101.1.
Figure 101.15 Spermatogenic wave in the equine testis is evident in an enzymatically isolated equine seminiferous tubule observed by bright-field microscopy. The tubule was fixed in glutaraldehyde then in osmium tetroxide, infiltrated with Epon, photographed, and studied. The corresponding drawing shows stages are consecutive along the length of the tubule, based on observations with Nomarski optics. Bar length equals 200 µm. Reprinted with permission.30
Spermatogenic wave The spermatogenic wave is the spatial order of stages along the length of seminiferous tubules at any given time (Figure 101.15). The order, inevitably, is sequential. The origin of the wave is unknown, but it results from synchronized but not simultaneous division of stem cell spermatogonia in adjacent tubular segments. The spermatogenic wave may function as a mechanism to (i) assure a constant release of spermatozoa, (ii) reduce competition for hormones and metabolites used in a given stage, (iii) reduce tubular congestion that would result if spermiation occurred simultaneously along the entire length of a tubule, and (iv) assure a constant
flow of seminiferous tubular fluid to maintain the vehicle for spermatozoan transport. The spermatogenic wave refers to the distribution of consecutive stages along the length of the tubules at any given time (Figure 101.15). The spermatogenic cycle, however, is the series of changes seen over time in a given region of the tubule. In 1901, Regaud24 expanded Von Ebner’s25 concept of the wave of the seminiferous epithelium by stating that ‘the wave is in space what the cycle is in time.’ The spermatogenic wave has been exploited to determine stage-specific changes in the seminiferous
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Spermatogenesis epithelium.26 Both follicle-stimulating hormone (FSH) and testosterone, essential hormones for spermatogenesis, seem to have different preferential stages during which they act on Sertoli cells. Also, there are stage-dependent differences in Sertoli cell function, as measured by the production of androgen-binding protein, plasminogen activator, and meiosis-inducing substance.26 These kinds of studies were made possible by staging live, dispersed rat or mouse seminiferous tubules by transillumination.27, 28 Seminiferous tubules in domestic species are tightly bound in the testicular parenchyma and are not easily isolated for these types of studies. Transillumination has been successfully employed to stage enzymatically dispersed horse seminiferous tubules.29, 30 This procedure should help to improve our understanding of spermatogenesis in domestic animals.
Evaluation of spermatogenesis The circumference of the scrotum in bulls and rams is highly correlated with testicular weight and spermatozoan production.31–33 This measurement is easily made and is valuable in examination for breeding soundness.34 In stallions, similar information can be obtained by measuring testis length plus width plus height. Spermatogenesis can be evaluated qualitatively by the general appearance of tubules viewed in histological sections and quantitatively by counting different germ cell types in tubule cross-sections of a specific stage.35 When using the general appearance, it must be considered that species differ in duration and relative frequency of stages that contain maturation phase spermatids or spermiation. The percentage of the horse testis occupied by seminiferous epithelium (72%) is higher than for rats (60%) and for humans (40%).35 Likewise, seminiferous tubules occupy 76% of the testis in bulls, 77% in boar, and 84% in dogs.14 Given the lifespan and theoretical yield of a specific germ cell, an estimate of the DSP can be obtained from the number of germ cells of that type in the testis (Figure 101.14). The duration of spermatogenic stages in which a cell type occurs equals that cell’s life span. The theoretical yield is equal to 2n , where n is the number of cell divisions between that cell type and spermatids.19 Daily sperm production per gram of decapsulated testis is useful for species comparisons as it is a measure of spermatogenic efficiency. This efficiency is lower in humans (4–6 × 106 /g = 4–6 million/gram of testicular tissue) than in all other species evaluated.35 The values (× 106 /g) for other species are estimated at 12 in bulls, 16 in dogs, 16–19 in stallions, 21 in rams,
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20–24 in rats, 23 in boars and rhesus monkeys, and 24 in hamsters.19, 21, 22,36–38
Germ cell degeneration during spermatogenesis Germ cells that degenerate Germ cell degeneration occurs throughout spermatogenesis, as evidenced by comparison of spermatozoan production rates from numbers of various types of germ cells. However, the greatest amount and impact occurs during spermatocytogenesis and meiosis. Degeneration rate during spermatocytogenesis is unknown for the horse, but has been determined to be 25% in mice,39 11% in Sherman rats, and 75% in adult Sprague–Dawley rats.12 In rams, long-day illumination of the non-breeding season causes increased spermatogonial degeneration.40 Likewise, in the stallion, the seasonally increased population of A-spermatogonia during the breeding season is offset by increased degeneration of B-spermatogonia (Figure 101.16).12 It has been reported9 that a twofold increase in the number of A-spermatogonia in the breeding season results from an increased yield of early spermatogonia (yield of A2 -spermatogonia or less degeneration of A3 -spermatogonia).9 Possibly 25% of germ cell loss in horses during the breeding season is during the meiotic divisions, but this is < 5% during the non-breeding season.19
Etiology of germ cell degeneration The number of cells that can be sustained by the available Sertoli cell population probably determines the amount of degeneration.12, 35 In stallions, if a large number and high percentage of B-spermatogonia did not degenerate during the breeding season when the number of A-spermatogonia is doubled, each Sertoli cell would have to accommodate 44 instead of the typical 28 germ cells (Table 101.2)9, 41 or far more than accommodated by Sertoli cells of any species. The seasonal differences in the number of germ cells that are supported by available Sertoli cells are shown in Table 101.2. Germ cell degeneration may be the result of a mechanism to eliminate cells containing abnormal chromosomes.39, 42 However, simple selection to eliminate chromosomal abnormalities cannot explain the fact that only certain types of spermatogonia usually are lost and the magnitude is relatively constant. 12 While germ cell degeneration plays a pivotal role in quantitative and qualitative spermatogenesis, its mechanism and approaches to its prevention remain to be discovered.19
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Figure 101.16 Effect of month (January = 1) on numbers of germ cells in the testis. Cells designated as type A-spermatogonia (A), type B-spermatogonia (B), preleptotene and leptotene plus zygotene primary spermatocytes (YP), Leydig cells (LC), and Sertoli cells (SC) per testis in adult (4- to 20-year-old) stallions (13–17 each month). Numbers of A- and B-spermatogonia in the breeding season were twice their values in the non-breeding season. However, monthly differences in numbers of cells were seen in each cell type. Reprinted with permission.50
Sertoli cells Role in spermatogenesis Sertoli cells, the somatic component of the seminiferous epithelium (Figure 101.3), are critical to germ cell development during spermatogenesis. They provide physical support and direct nutrients and regulating factors to the enveloped, partially enclosed germ cells. The numbers of Sertoli cells
are important as they probably determine the maximum numbers of the spermatogonia and spermatids and, hence, spermatozoan production rates (Figure 101.17).41 The entire role of Sertoli cells in spermatogenesis is unknown, but many functions are established. Sertoli cells provide the immediate microenvironment for each developing germ cell, based on their intimate contact (Figure 101.3), communication via gap
Table 101.2 Seasonal variation in number of Sertoli cells per testis and number of germ cells accommodated by a single Sertoli cell in stage VIII tubules based on 28 horses per season Season Item Number of Sertoli cells per testis (109 ) Germ-cell type A-spermatogonia Preleptotene and leptotene plus zygotene primary spermatocytes Pachytene plus diplotene primary spermatocytes Spermatids with spherical nucleus Spermatids with elongated nucleus All germ cell types combined NS, not significant. From Johnson41 and Johnson and Thompson.46
Non-breeding
Breeding
Significance
2.6 ± 0.2
3.6 ± 0.2
P < 0.01
1.1 ± 0.1 2.9 ± 0.4
1.5 ± 0.1 2.0 ± 0.2
P < 0.01 NS
2.6 ± 0.2 8.1 ± 0.8 8.0 ± 0.8 22.8 ± 2.1
3.0 ± 0.2 10.9 ± 0.8 10.2 ± 0.7 28.5 ± 1.7
NS P < 0.05 P < 0.05 P < 0.05
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Number of Sertoli cells per horse (109)
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Figure 101.17 Relationships between number of Sertoli cells or weight of testicular parenchyma and spermatogenesis. These data are from 184 adult horses (a–d), or from various germ cell populations in a subset of 19 horses (e–f) whose testes were obtained throughout the year. All correlation coefficients ((a) r = 0.827, (b) r = 0.887, (c) r = 0.806, (d) r = 0.802, (e) r = 0.744, and (f) r = 0.845) are significant (P < 0.01). Reprinted with permission.89
junctions,43 and exchange of nutrients and paracrine signals. Tight junctional complexes attach adjacent Sertoli cell processes and constitute the blood–testis barrier, which blocks the flux of serum components to spermatocytes and spermatids,44 thereby providing immunological isolation for these germ cells. The unique environment in which spermatocytes proliferate and spermatids differentiate into spermatozoa
is created by Sertoli cells and their junctional complexes.45 Sertoli cell junctional complexes divide the seminiferous epithelium into basal and adluminal compartments.4 Spermatocytogenesis occurs in the basal compartment where preleptotene primary spermatocytes are formed (Figure 101.3). These preleptotene primary spermatocytes pass through the blood–testis
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barrier to the adluminal compartment where meiosis and spermiogenesis occur. Sertoli cells are nurse cells as they provide material to stimulate growth, but also must function in removing waste. Sertoli cells phagocytize degenerating germ cells and residual cytoplasm left behind in the residual bodies at the end of spermiation. The transit of the residual bodies from the face of the seminiferous epithelium to the base of Sertoli cells, easily seen in rats, also can be seen in domestic species with the use of differential interface contrast optics and 20-µm-thick histological sections (Figure 101.14) (horse stage VIII). Pubertal development and effect of age on Sertoli cell numbers
Effect of season on Sertoli cell numbers in adults
Fetal, prepubertal, and pubertal horse testes are compared in Figure 101.18. The effects of developmental age on DSP, number of Sertoli cells, development of seminiferous tubules, and hormonal concentrations are summarized in Figure 101.19. In stallion testes obtained during the breeding season (summer), the
(a)
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Sertoli cells generally are assumed to have a stable population in adults. However, based on a large number of adult stallions, the number of Sertoli cells was found to be greater in the breeding season
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(c)
(f)
Figure 101.18 Phases in establishment of spermatogenesis in the horse. These phases extend from (a) 270 days of gestation through (b, c) 10–12 months, and (d, e) 16 months to (f) 36 months of age. Labeled structures include: a high density of robust interstitial cells (IC), pre-Sertoli cells (Ps), gonocytes (G), Sertoli cells (SC), spermatogonia (Sg), primary spermatocytes (PS), Sa spermatids (Sa), and Sd2 spermatids (Sd2 ). Bar length equals 10 µm. Reprinted with permission.59
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Figure 101.19 Development of the equine testis. Measurements include testicular weight, daily sperm production (DSP), number of Sertoli cells, elongated spermatids per Sertoli cell and serum hormonal concentrations in a group of young and adult horses whose numbers (n) in each age group are indicated. In (a) are right (RT-Wt) or left (LT-Wt) testicular weights and paired parenchymal weight (PPar-Wt), which increased (P < 0.01) with age and sexual development. Also presented in (b) are DSP/g of parenchyma in the breeding season (B DSP/g) and non-breeding season (NB DSP/g) as well as DSP per horse (DSP/H), which increased (P < 0.01) with age; units are — and —, respectively. The number of Sertoli cells per horse (c) in the breeding (B SC/H) and non-breeding (NB SC/H) season increased with age. Over all age groups, B SC/H was greater (P < 0.01) than NB SC/H. Despite incomplete spermatogenesis in some of the youngest stallions (d), DSP/horse was directly related (P < 0.01) to the number of Sertoli cells per horse (SC/H). Numbers (e) of elongated spermatids accommodated by a single Sertoli cell (ES/SC) and elongated spermatids per horse (ES/H) increased with age. Serum concentrations (f) of follicle-stimulating hormone (FSH), luteinizing hormone (LH), and testosterone (T) increased with age. FSH and LH levels are in ng/mL of serum and testosterone in 10 pg/mL serum. Values for percentages of elongated spermatids (% ES) and residual bodies (% RB) in adult stallions (g), are established by 2–4 years old, as noted by a second group of horses whose numbers for each age group are designated by n. Levels of paired parenchymal weight (Ppar-wt) and DSP/adult horse are reached by age 4 years. Reprinted with permission.47
(summer) than in the winter.46 In a subsequent experiment involving 186 other stallions,48 it was found that adult stallions had more Sertoli cells in the breeding season than in the non-breeding season, and intermediate absolute values were found between seasons (Figure 101.16). Photoperiod drives seasonal changes in serum concentrations of sex hormones in the stallion,49 and probably is responsible for seasonal changes in the number of Sertoli cells and other testicular cells.50
Growth along the length compared with the ends of the tubule, possibly by mitosis, appears to be a logical means by which Sertoli cell augmentation occurs in the breeding season of stallions.48 While the length of the tubules increases significantly from 1.9 to 2.9 km during the breeding season, there is no seasonal difference in tubular diameter, size of individual Sertoli cell nuclei, or number of Sertoli cells with a distinct nucleolus per cross-section of tubule.46
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The number of Sertoli cells per horse and paired parenchymal weight are directly related to the DSP and the number of spermatogonia (Figure 101.17). The number of Sertoli cells per horse was directly related not only to the number of young (preleptotene, leptotene, and zygotene) primary spermatocytes but also to the number of A1 -spermatogonia (Figure 101.18). This relationship between Sertoli cell numbers in the horse and spermatogenesis evident in the most primitive cell type, A1 -spermatogonia (Figure 101.17), is consistent with regulatory mechanisms working between Sertoli cells and A1 -spermatogonia.
Leydig cells Leydig cells are found all around the seminiferous tubules (Figures 101.1 and 101.2) and produce high concentrations of testosterone, which supports spermatogenesis. The role of testosterone will be discussed below. Equine Leydig cells (Figure 101.20, Table 101.3) have an abundant quantity of smooth endoplasmic reticulum (SER), where choles-
Figure 101.20 Photomicrograph of Leydig cells. Transmission electron micrograph of a Leydig cell in the stallion during the breeding season, which was also characteristic of the nonbreeding season. Labeled structures include: profiles of smooth (SER) and rough endoplasmic reticulum (RER), mitochondria (M), secondary lysosomes (L), nucleus (N) with numerous nuclear pores (NP), and the plasma membrane (PM) where gap junctions (GJ) and desmosomes (Ds) are found. At the free surface of the cells, the plasma membrane forms microvilli (Mv). Bar length equals 1 µm. Reprinted with permission.52
Table 101.3 Volume density (%) of various components in Leydig cell cytoplasm Season Component
Non-breeding
Breeding
SER RER Mitochondria Lysosome Lipid Swirled membrane Other components
78.1 ± 3.8 1.24 ± 0.26 11.2 ± 0.7 1.33 ± 0.30 0.05 ± 0.02 1.11 ± 0.22 7.0 ± 3.6
81.3 ± 1.6 1.01 ± 0.22 9.9 ± 0.22 1.18 ± 0.20 0.16 ± 0.05 0.88 ± 0.09 5.6 ± 1.8
RER, rough endoplasmic reticulum; SER, smooth endoplasmic reticulum. From Johnson and Thompson.52
terol is synthesized and pregnenolone is rapidly metabolized to testosterone. Leydig cells have many mitochondria that house the rate-limiting step of testosterone production, namely cholesterol sidechain cleavage and formation of pregnenolone. Species differences in the volume density of Leydig cell SER almost totally explains the species differences in in vitro production of testosterone.51 In addition, significant correlations between SER volume per testis and serum concentrations of testosterone or intratesticular testosterone content were found for the horse (Table 101.4).52 Leydig cells are functional in the embryo during the second month of gestation (Figure 101.18), regress in late embryonic and early neonatal periods, and become functional (Figure 101.20) again at puberty.53 Leydig cell development corresponds to changes in the darkness of the testicular parenchyma, and tubular development, in the horse.54 Fetal horse testes are laden with plump Leydig cells surrounding the sex cords, which later become the seminiferous tubules (Figure 101.20) and contain the darkpigmented cells that determine the dark appearance of the parenchyma when viewed grossly. The parenchyma becomes lighter as debris from fetal Leydig cells is removed, and the seminiferous tubules develop before puberty. In 15- to 20-g testes from 1- to 1.5-year-old horses, the parenchyma is light in color centrally, but still dark peripherally (Figure 101.21). Dark regions are characterized by an interstitium in which degenerating fetal interstitial cells are removed, and are associated with small undeveloped tubules with mostly Sertoli cells and gonocytes. In the light regions, seminiferous tubules are larger, have formed a lumen, and contain spermatogonia and spermatocytes (Figure 101.21). Further, there is an abundance of large, pigmented cells (macrophages) and unrecognizable Leydig cells.30, 55, 56 Hence, local development within a testis is obvious.
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Table 101.4 Effect of season on the endocrine status, number of Leydig cells, volume of Leydig cell cytoplasm, and volume of smooth endoplasmic reticulum (SER) in stallions Season
Number of stallions Age Testicular parenchymal weight (g) Serum FSH (ng/mL) Serum prolactin (ng/mL) Serum LH (ng/mL) Serum testosterone (pg/mL) Intratesticular testosterone (mg) Per g tissue Per testis Leydig cell number Per g parenchyma (× 10−6 ) Per testis (× 10−9 ) Volume of Leydig cell cytoplasm (mL) Volume of SER Per g (µL) Per testis (mL)
Non-breeding
Breeding
Significance
47 9.0 ± 0.8 126 ± 6 104 ± 10 2.1 ± 0.3 24 ± 5 300 ± 36
41 9.8 ± 0.9 163 ± 7 134 ± 14 6.0 ± 0.4 37 ± 6 402 ± 40
NS P < 0.01 NS P < 0.01 NS NS
0.6 ± 0.08 77 ± 12
0.7 ± 0.08 112 ± 12
NS P < 0.05
21 ± 1 2.6 ± 0.2 14.2 ± 0.9
24 ± 1 4.0 ± 0.2 20.9 ± 1.7
P < 0.05 P < 0.01 P < 0.01
89 ± 4 11.1 ± 0.7
102 ± 7 17.0 ± 1.4
NS P < 0.01
NS, not significant; FSH, follicle-stimulating hormone; LH, luteinizing hormone. From Johnson and Thompson.52
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(c)
Figure 101.21 Testicular parenchyma from a 1.5-year-old horse showing local initiation of spermatogenesis. Gross view of the parenchyma (a) showing the dark region (D) encircling the light (L) region, designated by the open arrows. Dark parenchyma (b) is characterized by undifferentiated seminiferous tubules (ST), as in Figure 101.19b, and large macrophages (open arrows) in the interstitium. Seminiferous tubules (ST) (c) in the light regions are developed to the extent that some mature spermatids are present (open arrows), as in Figure 101.19e or 101.19f. Bar length equals (a) 1 cm and (b, c) 100 µm. Reprinted with permission.65
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Leydig cell development appears first in the left testis. The left testis was heavier than the right 80% of the time in a large series of horses aged 1–54 months. 56 The left testis preceding the right in development, and the center preceding the peripheral region in development, is consistent with a local control of Leydig cell development (or removal of fetal interstitial cells) and local control in the establishment of spermatogenesis,47 provided the impinging peripheral hormonal influence is appropriate. Establishment of the Leydig cell population corresponds temporarily with changes in testicular size and parenchymal darkness. Leydig cell numbers increase with age in horses from 1.4 × 109 at 2–3 years old to 4.7 × 109 at 13–20 years old.54 This corresponds to an increase in Leydig cell volume per testis from 6 mL at 2–3 years old to 32 mL in 13- to 20-year-olds, an increase in testicular size from 117 to 213 g and a redarkening of the parenchyma. Indeed, a parenchymal pigmentation score reflecting the increased number and concentration of Leydig cells and the corresponding redarkening of the testis was shown to be indicative of a horse’s age from 2 to 20 years old.54 The Leydig cell population, like those of germ cells or Sertoli cells, cycles yearly (Figure 101.16).50, 57 Seasonal changes in Leydig cell number are responsible for seasonal changes in the volume of SER in Leydig cells, and in the intratesticular testosterone content in stallions (Table 101.3).52 Daily sperm production is positively correlated with the number of Leydig cells (r = 0.76) in the horse57 and the amount of Leydig cell SER in humans.58
Establishment of spermatogenesis Establishment of spermatogenesis takes a long time in domestic animals. The process starts before birth when primordial germ cells of the early developing embryo migrate from the yolk sac into the undifferentiated fetal gonads.59 These primordial germ cells undergo several divisions in the gonads to produce gonocytes within the fetal male sex cords (Figure 101.20). Sex cords are composed of the lightly staining gonocytes and the darkly staining supporting cells that differentiate into Sertoli cells (Figure 101.18) after birth. Gonocytes persist in the male sex cords until before puberty, at which time they differentiate and divide to produce spermatogonia. In the lamb, spermatogonia proliferate to produce spermatocytes by 100 days and spermatids by 120–125 days, with complete spermatogenesis by 140–150 days.60 In the calf, spermatogonial proliferation occurs during 16–20 weeks after birth, primary spermatocytes are present by 24 weeks, spermatids by 28 weeks, and completion of spermatogenesis by 32 weeks.61
Figures 101.18 and 101.19 show the establishment of spermatogenesis in the stallion. Gonocytes are present in the sex cords throughout most of gestation. By 12 months postnatal age, spermatogonia and primary spermatocytes are present, followed by spherical spermatids at 16 months and mature spermatids at 36 months. In the horse, as in other species,61–63 many degenerating germ cells can be found during the establishment of spermatogenesis. Once spermatogenesis is complete, the efficiency of yield increases over several months and then is stabilized at the adult level (Figure 101.19). Testicular size increases for a longer interval. At 28 weeks, 50% of bulls have a DSP ≥ 0.5 × 106 /g, for a mean 0.60 × 106 /g parenchyma. At 32 weeks, the efficiency of spermatogenesis measured by DSP/g was 4.0 × 106 /g,61 and increased through the first year when it reached a mature level of 12 × 106 .64 In stallion testes obtained in the summer, DSP/g of parenchyma was 6 × 106 at 1–2 years old, 15 × 106 at 2–3 years old, 18 × 106 at 4–5 years old, and 20 × 106 at 6–20 years old.46 Establishment of spermatogenesis apparently occurs at random throughout the testis in most species.15 The horse is an exception in that development of seminiferous tubules starts in the central portion of the testis and moves to peripheral regions.55, 65 As the result of this peculiar regional development, a gross section through a developing testis from a 1-year-old horse has light testicular parenchyma with expansion of seminiferous tubules in the center and dark on the peripheral regions where the degenerating interstitium cells predominate. Regional development of spermatogenesis within the testis as well as differences in the timing of development between the paired testes (the left testis usually develops before the right testis) implies a local control mechanism for the initiation of spermatogenesis. In stallions, as in other domestic animals, the maximum efficiency of spermatogenesis is obtained prior to maximum spermatozoan production per testis. Based on seminal characteristics, puberty occurs at 1–2.2 years old66 or 1.1–1.9 years old67 in Quarter Horse colts. Based on testicular development,68 puberty occurs between 1.4 and 1.8 years in Anglo-Norman stallions. This difference between breeds may reflect different approaches and different studies. By 2.5 years of age (Figure 101.19), equine testes from lightweight breeds reach adult levels for the percentage of seminiferous tubules with elongated spermatids (Sd1 and Sd2 spermatids) (Figure 101.19) or residual bodies left by released spermatozoa.47 Adult levels of DSP/g of parenchyma are obtained by 3 years old in the stallion, but testicular parenchymal weight and DSP increased to year 4 (Figure 101.19).
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Spermatogenesis The percentage of the testis occupied by seminiferous tubules was similar at about 72% for horses aged 2–3, 4–5, and 13–20 years old when testicular weight increased from 117 to 213 g/testis, and the volume of seminiferous tubules/testis increased from 72 to 127 mL.54 The increase in seminiferous tubular volume was partly due to a slight increase in the diameter, but was primarily due to an increased length from 2.0 km to 2.8 km over the same time period.
Control of spermatogenesis Hormonal influences The time over which a spermatozoon develops (duration of spermatogenesis) and the spermatogenic cycle
length are not influenced by hormones in normal or hypophysectomized males.26 However, the amount of germ cell degeneration and DSP are influenced by hormones.23,69–73 The initiation of spermatogenesis is thought to depend largely upon concentrations of luteinizing hormone (LH), FSH, and testosterone. High concentrations of LH stimulate production of testosterone by Leydig cells in juxtaposition with seminiferous tubules. Hormonal control of spermatogenesis differs among species.70 In the ram, FSH is necessary71, 72 and LH may also have a direct role in regulation of spermatogonial proliferation.70 In contrast, spermatogenesis can be maintained by testosterone alone in the rat. In the horse, seasonal photoperiod-driven49
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Figure 101.22 Scanning electron micrographs of the epididymis and spermatozoa within the epididymal duct. (a) Several profiles of the epithelium and lumen of the head of the epididymis. (b) A cross-section through the body of the epididymis, which has a thicker wall than that of the head. (c) A profile of spermatozoa in the proximal body of the epididymis, where translocation of the cytoplasmic droplet occurs. (d) A cross-section of the tail of epididymis, which has a large lumen and a thick muscular wall that expels spermatozoa via the ductus deferens. Bar lengths equal (a) 130 µm, (b) 126 µm, (c) 5.3 µm, and (d) 263 µm. Reprinted with permission.8
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serum concentrations of FSH, LH, testosterone, and prolactin were highest in the summer when testicular weight, intratesticular testosterone levels, numbers of Sertoli cells and Leydig cells, numbers of spermatogonia, and DSP are highest (Figure 101.19).9, 41, 46, 48, 50, 52, 57,74–76
Other influences Elevated temperature or adversely cold conditions are detrimental to spermatogenesis in many species. Artificially increasing scrotal temperature to 41◦ C for 3 hours in rams and boars causes rapid destruction of pachytene spermatocytes.77, 78 A transient effect on spermatogonial divisions occurs with increased numbers of cells in prophase and metaphase.78 While elevated scrotal temperature also caused loss of spherical spermatids in the boar,77 pachytene spermatocytes in meiotic prophase appeared to be the most thermosensitive.78 High ambient temperature reduced spermatozoan production in boars.79 Other factors, such as radiation, drugs, vitamin A deficiency, limited diet, and toxic chemicals, are detrimental to spermatogenesis. Advanced age reduces the numbers of germ cells, thus DSP, in rabbits,80 mice,81 and rats.82 In horses, DSP reaches a high plateau at about 4–5 years old, and remains at this level until 20 years old.46, 54 What happens beyond 20 years old remains unknown because of lack of data. Approximately 46% of Holstein bulls selected for both genetic and reproductive desirability at artificial insemination centers are removed because of reproductive problems83 and replaced by genetically superior bulls. Few bulls of any breed are retained past 20 years old for genetic reasons. The fact that some males retain their fertility, even with advancing age, is indicated by fertility in 19-year-old bulls,84 and a successful paternity in a 94-year-old man.85
Timing between developmental steps in spermatogenesis and the appearance of spermatozoa in the ejaculate The interval required for sperm to be transported through the head and body is approximately 4 days, but transport through the tail of the epididymis is affected by ejaculation frequency. In sexually inactive stallions, spermatozoa often remain in the tail for approximately 10 days, but in sexually active stallions this interval is reduced to 6–8 days (Figure 101.22). Hence, in a sexually active stallion (high frequency of ejaculation), sperm transit time through the entire epididymis is 10–11 days. Since spermatogenesis in
Figure 101.23 Scanning electron micrograph of ejaculated equine spermatozoa. Most of the spermatozoa in the ejaculate are normal. However, distal cytoplasmic droplets (CD) on some spermatozoa are still attached in the annulus area of the spermatozoan tail. Reprinted with permission.8
the stallion is 57 days, the number of days between a given developmental step of a germ cell and the time when resulting spermatozoa will appear in the ejaculate (Figure 101.23) can be estimated. Figure 101.24 shows the number of days from each developmental step in the eight stages of the spermatogenic cycle to the time the progeny of that developmental step will appear in the ejaculate. This information is useful to note which developmental step in spermatogenesis is susceptible to a given testicular insult. Hence, the number of days between the testicular insult and the appearance of poor seminal quality can be used to calculate the developmental age of germ cells and identify the germ cell type that is sensitive to that specific testicular insult.
Summary and conclusions Spermatogenesis is the production of spermatozoa via the division of spermatogonia by mitosis (spermatocytogenesis) to form spermatocytes, which undergo meiosis to form spermatids, which then differentiate (spermiogenesis) from spherical spermatids to mature spermatids, and are released from the seminiferous epithelium as spermatozoa.
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25.2 Sb1 23.2
Sb2 22.1
Sc 21.0
Sc 19.6
Sd1 18.3
Sd1 16.7
Sd2 15.0
Sd2
37.4
P 35.4
P 34.3
D 33.2
SS 31.8
Sa 30.5
Sa 28.9
Sa 27.2
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P 44.0
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B1 57.6
B1 56.2
B1 54.9
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12.2 Days Figure 101.24 Time until appearance in the ejaculate, based on a 14-day epididymal transit time. The eight stages of the spermatogenic cycle in the horse. This figure is similar to Figure 101.4, with timing of cell types to correspond to when spermatozoa are produced from these cells and appear in the ejaculate. This timing is based on a 10-day transit time of spermatozoa to travel through the equine epididymis. If a testicular insult influences both B1 -spermatogonia and Sb2 -spermatids, the effect will be seen as poor semen quantity or quality 23 days later and again 56 days later from the same insult. Reprinted with permission.87
The duration of spermatogenesis from committed spermatogonia to release of spermatozoa is 57 days in the horse. The length of the spermatogenic cycle is 12.2 days in the horse. This is the interval between two consecutive spermiations (releases of spermatozoa) from a given region of the seminiferous tubule. The cycle of the seminiferous epithelium in the horse has been divided (man-made divisions) into eight stages of the spermatogenic cycle. The horse testis is composed of 72% seminiferous tubules. The percentage of tubules with two generations (two cohorts with 12.2 days different in developmental ages) of primary spermatocytes for the horse is 35%, or the duration of stages I–III. The percentage of tubules with two generations of spermatids is 50%, or stages V–VIII. The efficiency of spermatogenesis (DSP/g of testicular parenchyma) averages 19 × 106 for the summer (natural breeding season) and 15 × 106 for
the winter. The efficiency is maximal prior to the maximal spermatozoan production per horse as the total testicular weight continues to increase after the efficiency of spermatozoa has been reached, but the efficiency will be influenced seasonally in adult horses. There is a latency in the time before a testicular insult and the resulting decline in seminal quality, and the decline can persist for > 57 days (duration of spermatogenesis) plus 14 days (epididymal transit time). Even longer will be required for the seminal quality to recover from a major or protracted testicular insult.
Acknowledgements Special thanks are extended to Vince B. Hardy for expert technical assistance and Penny Churchill for secretarial assistance with this manuscript.
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References 1. Saacke RG, Almquist JO. Ultrastructure of bovine spermatozoa. II. The neck and tail of normal, ejaculated sperm. Am J Anat 1964;115:163–84. 2. Johnson L, Amann RP, Pickett BW. Maturation of equine epididymal spermatozoa. Am J Vet Res 1980;41:1190–6. 3. Olson GE, Sammons DW. Structural chemistry of outer dense fibers of rat sperm. Biol Reprod 1980;22:319–32. 4. Fawcett DW. Ultrastructure and function of the Sertoli cell. In: Hamilton DW, Greep RO (eds) Handbook of Physiology, vol. 5, section 7. Washington, DC: American Physiology Society, 1975; pp. 21–55. 5. Garner DL, Hafez ESE. Spermatozoa. In: Hafez ESE (ed.) Reproduction in Farm Animals, 4th edn. Philadelphia: Lea & Febiger, 1980; pp. 167–88. 6. Johnson L. A re-evaluation of daily sperm output of men. Fertil Steril 1982;37:811–16. 7. Johnson L, Amann RP, Pickett BW. Scanning electron and light microscopy of the equine seminiferous tubule. Fertil Steril 1978;29:208–15. 8. Johnson L, Amann RP, Pickett BW. Scanning electron microscopy of the epithelium and spermatozoa in the equine excurrent duct system. Am J Vet Res 1978;39:1428–34. 9. Johnson L. Seasonal differences in equine spermatocytogenesis. Biol Reprod 1991;44:284–91. 10. Johnson L. Increased daily sperm production in the breeding season of stallions is explained by an elevated population of spermatogonia. Biol Reprod 1985;32:1181–90. 11. Hochereau-de Reviers MT. Control of spermatogonial multiplication. In: McKerns KW (ed.) Reproductive Processes and Contraception. New York: Plenum Press, 1981; pp. 307–31. 12. Huckins C. The morphology and kinetics of spermatogonial degeneration in normal adult rats: an analysis using a simplified classification of the germinal epithelium. Anat Rec 1978;190:905–26. 13. Amann RP. A critical review of methods for evaluation of spermatogenesis from seminal characteristics. J Androl 1981;2:37–58. 14. Swierstra EE, Gebauer MR, Pickett BW. Reproductive physiology of the stallion. I. Spermatogenesis and testis composition. J Reprod Fertil 1974;40:113–23. 15. Courot M, Hochereau-de Reviers MT, Ortavant R. Spermatogenesis. In: Johnson AD, Gomes WR, VanDemark NL (eds) The Testis, vol. I. New York: Academic Press, 1970; pp. 339–442. 16. Amann RP. Sperm production rates. In: Johnson AD, Gomes WR, VanDemark NL (eds) The Testis, vol. I. New York: Academic Press, 1970; pp. 433–82. 17. Burgos MH, Vitale-Calpe R, Aoki A. Fine structure of the testis and its functional significance. In: Johnson AD, Gomes WR, VanDemark NL (eds) The Testis, vol. I. New York: Academic Press, 1970; pp. 551–649. 18. Leblond CP, Clermont Y. Spermiogenesis of rat, mouse, hamster and guinea pig as revealed by the periodic acid-fuchsin sulfurous acid technique. Am J Anat 1952;90:167–215. 19. Johnson L. Spermatogenesis (animal species and humans). In: Proceedings of the International Symposium on Gamete Physiology, Newport Beach, CA, 1988, Serono Symposia. 20. Roosen-Runge EC, Giesel LO. Quantitative studies on spermatogenesis in the albino rat. Am J Anat 1950;87:1–30. 21. Amann RP. Detection of alterations in testicular and epididymal function in laboratory animals. Environ Health Perspect 1986;70:149–58.
22. Amann RP, Johnson L, Thompson DL Jr, Pickett BW. Daily spermatozoal production, epididymal spermatozoal reserves and transit time of spermatozoa through the epididymis of the Rhesus monkey. Biol Reprod 1976;15:586–92. 23. Clermont Y. Kinetics of spermatogenesis in mammals: seminiferous epithelium cycle and spermatogonial renewal. Physiol Rev 1972;52:198–236. 24. Regaud C. Etude sur la structure des tubes seminiferes et sur la spermatogenese chez les mammiferes. Arch Anat Microscop Morphol Expt 1901;4:101–56,231–80. 25. Von Ebner V. Untersuchungen uber den Bau der Samenkanalchen und die Entwicklung der spermatozoiden bei den Saugentieren und beim Menschen. In: Rollet’s Untersuchungen aus dem Institut fur ¨ Physiologie und Histologie in Graz. Leipzig, 1871; pp. 200–36. 26. Parvinen M. Regulation of the seminiferous epithelium. Endocrine Rev 1982;3:404–17. 27. Parvinen M, Hecht NB. Identification of living spermatogenic cells of the mouse by transillumination-phase contrast microscopic technique for in situ analyses of DNA polymerase activities. Histochemistry 1981;71:567–79. 28. Parvinen M, Vanha-Perttula T. Identification and enzyme quantification of the stages of the seminiferous epithelial wave in the rat. Anat Rec 1972;174:435–50. 29. Johnson L, Kattan-Said AF, Hardy VB, Scrutchfield WL. Isolation and staging of horse seminiferous tubules by transillumination. J Reprod Fertil 1990;89:689–96. 30. Johnson L, Hardy VB, Martin MT. Staging equine seminiferous tubules by Nomarski optics in unstained histologic sections and in tubules mounted in toto to reveal the spermatogenic wave. Anat Rec 1990;227:167–74. 31. Coulter GH. Testicular development: its management and significance in young beef bulls. Proc. 8th National Assoc Anim Breeders Tech. Conf. AI Reprod, 1980; pp. 106–11. 32. Foote RH. Factors influencing the quantity and quality of semen harvested from bulls, rams, boars and stallions. J Anim Sci 1978;47(Suppl. 2):1–11. 33. Willett EL, Ohms JI. Measurement of testicular size and its relation to production of spermatozoa by bulls. J Dairy Sci 1957;40:1559–69. 34. Amann RP, Schanbacher BD. Physiology of male reproduction. J Anim Sci 1983;63(Suppl. 2):380–403. 35. Johnson L. Review article: spermatogenesis and aging in the human. J Androl 1986;7:331–54. 36. Johnson L, Lebovitz RM, Samson WK. Germ cell degeneration in normal and microwave-irradiated rats: potential sperm production rates at different developmental steps in spermatogenesis. Anat Rec 1984;209:501–7. 37. Johnson L, Petty CS, Neaves WB. Age-related variation in seminiferous tubules in men: a stereologic evaluation. J Androl 1986;7:316–22. 38. Lebovitz RM, Johnson L. Testicular function of rats following exposure to microwave radiation. Bioelectromagnetics 1983;4:107–14. 39. Clermont Y. Quantitative analysis of spermatogenesis of the rat: a revised model for the renewal of spermatogonia. Am J Anat 1962;111:111–29. 40. Ortavant R. Le cycle spermatogenetique chez le Belier. Thesis. University of Paris, France, 1958. 41. Johnson L. A new approach to quantification of Sertoli cells that avoids problems associated with the irregular nuclear surface. Anat Rec 1986;214:231–7. 42. Oakberg EF. A description of spermiogenesis in the mouse and its use in analysis of the cycle of the seminiferous epithelium and germ cell renewal. Am J Anat 1956;99:391–413.
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Spermatogenesis 43. Russell L. Desmosome-like junctions between Sertoli and germ cells in the rat testis. Am J Anat 1977;148:301–12. 44. Setchell BP, Waites GMH. The blood-testis barrier. In: Hamilton DW, Greep RO (eds) Handbook of Physiology, vol. 5, section 7. Washington, DC: American Physiology Society, 1975; pp. 143–72. 45. Waites GMH. Fluid secretion. In: Johnson AD, Gomes WR (eds) The Testis, vol. IV. New York: Academic Press, 1977; pp. 91–123. 46. Johnson L, Thompson DL Jr. Age-related and seasonal variation in the Sertoli cell population, daily sperm production and serum concentrations of follicle-stimulating hormone, luteinizing hormone and testosterone in stallions. Biol Reprod 1983;29:777–89. 47. Johnson L, Varner DD, Thompson DL Jr. Effect of age and season on the establishment of spermatogenesis in the horse. J Reprod Fertil Suppl 1991;44:87–97. 48. Johnson L, Nguyen HB. Annual cycle of the Sertoli cell population in adult stallions. J Reprod Fertil 1986;76: 311–16. 49. Clay CM, Squires EL, Amann RP, Nett TM. Influences of season and artificial photoperiod on stallions: luteinizing hormone follicle-stimulating hormone and testosterone. J Anim Sci 1988;66:1246–55. 50. Johnson L, Tatum ME. Temporal appearance of seasonal changes in numbers of Sertoli cells, Leydig cells, and germ cells in stallions. Biol Reprod 1989;40:994–9. 51. Zirkin BR, Ewing LL, Kromann N, Cochran RC. Testosterone secretion by rat, rabbit, guinea pig, dog, and hamster testes perfused in vitro: correlation with Leydig cell ultrastructure. Endocrinology 1980;107:1867–74. 52. Johnson L, Thompson DL Jr. Effect of seasonal changes in Leydig cell number on the volume of smooth endoplasmic reticulum in Leydig cells and intratesticular testosterone content in stallions. J Reprod Fertil 1987;81:227–32. 53. Hooker CW. The intertubular tissue of the testis. In: Johnson AD, Gomes WR, VanDemark NL (eds) The Testis, vol. I. New York: Academic Press, 1970; pp. 483–550. 54. Johnson L, Neaves WB. Age-related changes in the Leydig cell population, seminiferous tubules, and sperm production in stallions. Biol Reprod 1981;24:703–12. 55. Bouin P, Ancel P. La glande interstitelle du testicule chez le cheval. Arch Zool Exp Genet 1905;3:391–437. 56. Nishikawa Y, Horie T. Studies on the development of the testes and epididymides of the horse. I. Studies on the development of the testes of the horse, with special reference to singularity and the age of sexual maturity. Natl Inst Ag Sci JPN Bull Ser G Anim Husb 1955;10:229–349. 57. Johnson L, Thompson DL Jr. Seasonal variation in the total volume of Leydig cells in stallions is explained by variation in cell number rather than cell size. Biol Reprod 1986;35:971–9. 58. Johnson L, Grumbles JS, Chastain S, Goss HFJ, Petty CS. Leydig cell cytoplasmic content is related to daily sperm production in men. J Androl 1990;11:155–60. 59. Johnson L. Spermatogenesis. In: Cupps PT (ed.) Reproduction in Domestic Animals, 4th edn. New York: Academic Press, 1991; pp. 173–219. 60. Ortavant R, Courot M, Hochereau-de Reviers MT. Spermatogenesis in domestic animals. In: Cole HH, Cupps PT (eds) Reproduction in Domestic Animals, 3rd edn. New York: Academic Press, 1977; pp. 203–27. 61. Curtis SK, Amann RP. Testicular development and establishment of spermatogenesis in Holstein bulls. J Anim Sci 1981;53:1645–57.
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62. Attal J, Courot M. Testicular development and onset of spermatogenesis in bulls. Ann Biol Anim Biochim Biophys 1963;3:219–41. 63. Clermont Y, Perey B. Quantitative study of the cell population of the seminiferous tubules in immature rats. Am J Anat 1957;100:241–67. 64. Amann RP, Almquist JO. Bull management to maximize sperm output. In: Proceedings of the 6th Tech. Conf. AI Reprod, 1976, pp. 1–10. 65. Clemmons AJ, Thompson DL Jr, Johnson L. Local initiation of spermatogenesis in the horse. Biol Reprod 1995;52:1258–67. 66. Cornwell JC. Seasonal variation in stallion semen and puberty in the Quarter Horse colt. Master’s Thesis, Louisiana State University, 1972. 67. Naden J, Amann RP, Squires EL. Testicular growth, hormone concentrations, seminal characteristics and sexual behaviour in stallions. J Reprod Fertil 1990;88:167–76. 68. Nishikawa Y. Studies on Reproduction in Horses. Tokyo: Japan Racing Association, 1959; p. 27. 69. Amann RP. Structure and function of the normal testis and epididymis. J Am Coll Toxicol 1989;8:457–71. 70. Courot M. The effects of gonadotropins on testicular function (spermatogenesis). In: Proc. 11th Internat. Congr. Anim. Reprod. AI, 5:311–319, 1988. 71. Courot M, Hochereau-de-Reviers MT, Monet-Kuntz C, Locatelli A, Pisselet C, Blanc MR, Dacheux JL. Endocrinology of spermatogenesis in the hypophysectomized ram. J Reprod Fertil Suppl 1979;26:165–73. 72. Hochereau-de Reviers MT, Monet-Kuntz C, Courot M. Spermatogenesis and Sertoli cell numbers and function in rams and bulls. J Reprod Fertil Suppl 1987;34:101–14. 73. Steinberger E, Steinberger A. Spermatogenic function of the testis. In: Hamilton DW, Greep RO (eds) Handbook of Physiology, vol. 5, section 7. Washington, DC: American Physiology Society, 1975; pp. 1–19. 74. Pickett BW, Amann RP, McKinnon AO, Squires EL, Voss JL. Management of the Stallion for Maximum Reproductive Efficiency, II. Colorado State University Animal Reproduction Laboratory Bulletin No. 05, 1989. 75. Thompson DL Jr, Johnson L. Effects of age, season and active immunization against estrogen on serum prolactin concentrations in stallions. Dom Anim Endocrinol 1987;4:17–22. 76. Thompson DL Jr, Johnson L, St. George RL, Garza F Jr. Concentrations of prolactin, luteinizing hormone and follicle stimulating hormone in pituitary and serum of horses: effect of sex, season and reproductive state. J Anim Sci 1986;63:854–60. 77. Mazzarri G, du Mesnil du Buisson F, Ortavant R. Action de la temperature et de la lumiere sur la spermatogenese, la production et le pouvoir fecondant du sperme chez le verrat. Proc. 6th Internat. Congr. Anim. Reprod. AI, 1:305–308, 1968. 78. Waites GMH, Ortavant R. Early cytological changes observed after the testis was subjected to elevated temperature. Ann Biol Anim Biochim Biophys 1968;8:323–31. 79. Wettemann RP, Wells ME, Omtvedt IT, Pope CE, Turman EJ. Influence of elevated ambient temperature on reproductive performance of boars. J Anim Sci 1976;42:664–9. 80. Ewing LL, Johnson BH, Desjardins C, Clegg RF. Effect of age upon the spermatogenic and steroidogenic elements of rabbit testes. Proc Soc Exp Biol Med 1972;140:907–10. 81. Gosden RG, Richardson DW, Brown N, Davidson DW. Structure and gametogenic potential of seminiferous tubules in ageing mice. J Reprod Fertil 1982;64:127–33.
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82. Johnson L, Neaves WB. Enhanced daily sperm production in the remaining testis of aged rats following hemicastration. J Androl 1983;4:162–6. 83. Kratz JL, Wilcox CJ, Martin FG, Becker RB. Vital statistics and reasons for disposal of United States and Canadian artificial insemination dairy sires. J Dairy Sci 1983;66: 646–6. 84. Bishop MWH. Ageing and reproduction in the male. J Reprod Fertil Suppl 1970;12:65–87. 85. Seymour FI, Duffy C, Koerner A. A case of authenticated fertility in a man, aged 94. JAMA 1935;105:1423–4. 86. Varner DD, Schumacher J, Blanchard TL, Johnson L. Reproductive anatomy and physiology. In Pratt PW (ed.) Diseases
and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991; pp. 1–59. 87. Johnson L. Efficiency of Spermatogenesis in Humans and Animals. College Station, TX: College of Veterinary Medicine Texas A&M University, 1997. 88. Amann RP, Johnson L, Pickett BW. Connection between the seminiferous tubules and the efferent ducts in the stallion. Am J Vet Res 1977;38:1571–9. 89. Johnson L, Carter GK, Varner DD, Taylor TS, Blanchard TL, Rembert MS. The relationship of daily sperm production with number of Sertoli cells and testicular size in adult horses: role of primitive spermatogonia. J Reprod Fertil 1994;100:315–21.
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Chapter 102
Spermatozoal Function Rupert P. Amann and James K. Graham
Spermatozoa are unique among cells produced by a stallion, not only by virtue of their haploid number of chromosomes but especially because of their highly specialized function and limited capacity for repair. Although spermatozoa are released from the seminiferous epithelium at the time of spermiation, designating the end of spermatogenesis, this represents but one point in a continuous process of cell modification (Chapters 96, 97 and 101). The saga of the spermatid/spermatozoon is initiated with formation of spermatids, their development in the seminiferous epithelium during spermatogenesis, and their release as spermatozoa; continues with modifications, termed spermatozoal maturation, resulting from changes induced by components of luminal fluid within the epididymis; further continues with admixture of spermatozoa and seminal fluids at emission and ejaculation; progresses with exposure to the environment of the female reproductive tract and entrance into the microenvironment of the oocyte; and is culminated following penetration into the investments around the oocyte and ultimately the oocyte proper (Figure 102.1).1 Unfortunately, veterinarians and reproductive biologists tend to consider spermatozoa from the perspective of their own narrow interests, such as characteristics of ejaculated spermatozoa, preservation of ejaculated spermatozoa, or in vitro fertilization. This approach ignores the fact that an event modifying a spermatid/spermatozoon at any time during its normal 5-week life span can reduce the ability of that spermatozoon to fertilize an oocyte. During spermatogenesis, organelles of the spermatid are modified in shape and function from those characteristic of a somatic cell into those appropriate for a cell designed to fertilize an oocyte.2–5 Nuclear shape is modified from that of a sphere, and the chromatin is condensed and compacted to minimize
its volume within the nucleus. A propulsion mechanism is provided as the axoneme, dense fibers, and the fibrous sheath; mitochondria are arrayed in close proximity to the proximal portion of the propulsion system to facilitate transport of adenosine triphosphate (ATP) from the mitochondria to the contracting elements of the axoneme; and a highly specialized structure, the acrosome, which contains hydrolytic enzymes, is placed over the rostral portion of the nucleus to facilitate penetration by the spermatozoon of the investments around the oocyte. Finally, the plasma membrane is anchored to underlying structures at several points over the tail of the spermatozoon but is relatively unattached to underlying structures in the head region. The major portion of the cytoplasm, including organelles characteristic of a somatic cell, is left behind as a residual body retained by the seminiferous epithelium at the time of spermiation. Consequently, although spermatozoa have the capacity for both aerobic and anaerobic metabolism of glucose, lactate, or pyruvate, and oxidation of certain amino acids and lipids, they lack the conventional machinery for biosynthesis and cell repair. For a spermatozoon to fertilize an oocyte, it must develop and retain at least five general attributes:6,7 1. metabolism for production of energy 2. progressive motility 3. enzymes, located within the acrosome, that are essential for penetration of the spermatozoon through structures surrounding the oocyte 4. proper distributions of lipids in the plasma and acrosomal membranes to stabilize these structures until fertilization and ultimately allow membrane fusion at the proper time 5. proteins in the plasma membrane that are essential for survival of a spermatozoon within the female reproductive tract, by locally suppressing the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Spermatid
Embryo Fertilization
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Acrosome reaction
Ejac.
Storage
Testis
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In vitro Figure 102.1 The saga of the stallion spermatid/ spermatozoon. An abnormal event, or human intervention, at any point on the lifeline can alter subsequent events. An alteration may be apparent immediately, but it often is not apparent until later in the life line or even by failure of a fouror eight-cell embryo to develop into a foal. (Adapted from Amann.1 )
immune system and also providing needed interactions with epithelial cells at selected sites and for attachment of the spermatozoon to the plasma membrane of the oocyte at the time of fertilization. Although certain of these attributes are provided during spermatogenesis, their final development is not completed until maturation of spermatozoa within the epididymis and admixture of spermatozoa with seminal plasma1,8 (Chapter 96). Although the exact nature and sequence of the multiple changes involved in developing a fertile spermatozoon are unknown for the stallion, researchers have no doubt that each of the attributes listed above is crucial for accomplishment of its task. Furthermore, although spermatozoa from the distal cauda epididymidis are equally as effective as ejaculated spermatozoa for fertilizing oocytes, with resulting embryos developing into newborns, seminal plasma alters spermatozoa in a manner presumed to be beneficial. Although seminal plasma probably is important in modifying spermatozoa,9 as well as providing a large volume to facilitate their distribution within the uterus of a mare, it is not an ideal medium for storage of spermatozoa10 and sometimes contains deleterious components. Even what appear to be normal spermatozoa may not be competitive. Spermatozoa from some males are able to ‘beat out’ spermatozoa from another male, or several males, when deposited simultaneously in the female reproductive tract by heterospermic insemination11 or sequentially, as might happen in the wild when several males copulate with an estrous female. This probably is an evolutionary feature to help perpetuate a species.12 Similarly, strains of mammals or poultry can be selected for fertility. A horse breeder who goes to extreme efforts to obtain progeny of a subfertile male by application of
techniques of modern animal biotechnology is imprudent, because that might only perpetuate undesirable reproductive traits.
Structure of equine spermatozoa A spermatozoon usually is considered as having a head, neck, middle piece, principal piece, and end piece2,13–17 (Figure 102.2). Each of these five structural regions will be considered in detail, together with functions of the components of each region. However, from the perspective of storage of spermatozoa for use in artificial insemination, either at 5◦ C or especially –196◦ C, it is more relevant to consider spermatozoa as consisting of a highly condensed nucleus and microtubular, fibrous, and membranous structures, because of the differential response of these types of structural components to cold shock, reduced temperature, or cryopreservation.18–20 Microtubular and fibrous structures are important for spermatozoal motility, because they constitute the doublets of the axoneme and the dense fibers and fibrous sheath of the middle piece and principal piece. However, membranous structures are more important from the perspective of cold shock. Cold shock is a type of damage which could be inadvertently caused during handling of spermatozoa associated with seminal collection or artificial insemination, even when semen is not intentionally cooled in preparation for storage.20 Because of the importance of the changes in spermatozoa invoked by thermal stress, the topic is considered in detail in a following section.
Plasma membrane The plasma membrane encompasses the entire spermatozoon and is its outermost component. Although it is continuous over the surface of the spermatozoa except after the acrosome reaction, which is a prelude to fertilization, or with senescence and death, the nature and function of the plasma membrane differs regionally. Not all regional differences in function of the plasma membrane are known, but portions over the rostral and caudal surfaces of the head, middle piece, and principal piece have different roles in spermatozoal function and survival. Regardless of location in a spermatozoon, the plasma membrane consists of three zones: lipid bilayer, phospholipid–water interface, and glycocalyx (Figure 102.3). In addition, proteins can be adsorbed to the surface of the membrane. The lipid bilayer is composed of polar phospholipids oriented with their hydrophobic fatty acyl
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Spermatozoal Function
Plasma membrane
Acrosome Head
Mitochondria
Neck Middle piece
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Dense fibers Nucleus
Central pair Doublet
Plasma membrane
Equatorial segment Principal piece
Plasma membrane Mitochondria Annulus Longitudinal column Dense fibers Doublet Plasma membrane Longitudinal column
Post acrosomal lamina
Fibrous sheath
Plasma membrane Mitochondria End piece
Plasma membrane Doublets
Doublets
Dense fibers
Figure 102.2 Diagrams of a stallion spermatozoon. The entire spermatozoon is overlaid by the plasma membrane, which usually is in apposition with the underlying structures and is anchored to the caudal margin of the head, at the annulus, and along the longitudinal columns of the principal piece. The head includes the nucleus (containing the genetic information in highly condensed DNA), the bag-like acrosome (containing enzymes necessary for fertilization), a specialized portion of the acrosome termed the equatorial segment, and the postacrosomal lamina. The neck is the point of attachment of the tail to the head, by a ball-and-socket arrangement. The central pair and nine doublets of microtubules, which constitute the axoneme, are surrounded by nine dense fibers. All extend from the neck region, through the middle piece and principal piece, into the end piece where they terminate at slightly different sites. Because these dense fibers are tapered, the tail becomes progressively thinner. The doublets are the contractile elements which contract differentially to induce a sliding motion and flex the tail in a helical pattern. This propels the spermatozoon. Mitochondria are membranous structures where most of the energy necessary for spermatozoal motion is produced. The longitudinal columns and fibrous sheath of the principal piece and the dense fibers provide the rigidity necessary for normal motion of the tail. Dimensions of stallion spermatozoa are approximately as follows: head length, 7 µm; middle piece length, 10 µm; middle piece diameter, 0.9 µm; principal piece length, 40 µm; principal piece diameter, 0.6 to <0.5 µm; and end piece length, 4 µm. (Modified from Amann et al.7 )
chains directed internally and hydrophilic, charged polar headgroups directed externally toward the polar solvent, water7,18,19 (Figure 102.4). The predominant lipids are phospholipids and cholesterol. Proteins are intermingled with the lipids and represent about 50% of the weight of the membrane. Proteins within these lipids are considered either integral (essential for structure of the membrane) or peripheral (associated with the membrane, but easily removed). Some integral proteins serve as pores or channels through the membrane or are surface receptors for other molecules, whereas others are found between the two bilayers of the membrane. Many proteins on the external surface contain carbohydrate side chains, which tend to have a net negative charge and attract, and loosely bind, other proteins in the medium
around the spermatozoon. Because of this adsorbed material, the outer aspect of the glycocalyx region of a spermatozoon can change, depending on its history and the medium it is in. Only limited data exist on the composition of the plasma membranes of stallion spermatozoa.21 The cholesterol to phospholipid ratio in the plasma membrane is 0.36, which is midway between values for boar and bull spermatozoa. As in spermatozoa from other domestic species, choline, ethanolamine, and sphingomyelin are the major phospholipid classes. In spermatozoa from common mammals, composition and localized distribution of phospholipids, and the nature of their fatty acyl side chains, differ within certain domains or regions of the plasma membrane. This probably also is true of stallion spermatozoa.
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Plasma membrane phospholipid bilayer
Glycocalyx region
Adsorbed protein
Integral protein
Peripheral protein
Carbohydrates Extracytoplasmic surface
0.7nm
5nm
Phospholipid Cholesterol
5nm
15nm
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Figure 102.3 Functional view of the plasma membrane of a spermatozoon, showing spatial relationships and distances. Both small (molecular weight 20 000) and large (molecular weight 60 000) solutes that might approach the membrane are depicted. Their approach to the lipid bilayer is influenced by the nature and amount of proteins adsorbed to the glycocalyx and the nature of the glycocalyx per se. (From Hammerstedt et al.18 )
Normally, different phospholipids are randomly arranged in a lamellar fashion and are free to move about laterad within one leaflet of the membrane bilayer.7,18 This is because membranes are fluid at body temperature. Furthermore, the lipid molecules are in a liquid rather than a gel or crystalline state. The ratio of cholesterol to phospholipids with polyunsaturated acyl side chains, as well as the nature of the phospholipid, determines fluidity of the membrane. In general, the more cholesterol present, the less fluid or flexible is that portion of the membrane. Cholesterol aids in keeping the phospholipids in a random, lamellar arrangement. Normally, an adequate amount of cholesterol and the random distribution of integral membrane proteins impose a lamellar configuration on phospholipid and a normal bilayer is maintained. Under certain conditions, however, the lipids may change into crystalline arrays (because of a phase transition) and the proteins become aggregated.18 This induces instability in the membrane and the membrane may be damaged irreversibly. Cooling is one factor inducing such changes.
Peripheral protein Channel or pore
Cytoplasmic surface
Figure 102.4 A diagrammatic representation of the plasma membrane with the outer face up. This figure is an oversimplification, but it illustrates the complexity of membranes. Other membranes in a spermatozoon are similar in nature, but they may lack the extensive array of exposed carbohydrates constituting the glycocalyx. The membrane is composed of lipids (mainly phospholipids and cholesterol) and proteins (peripheral and integral). The structure of integral proteins allows formation of both hydrophilic (dark gray) and hydrophobic (striped) regions. The hydrophobic regions allow intercalation of proteins into the hydrophobic interior of the lipid bilayer. Carbohydrate groups (depicted on the upper surface) probably are found only on the extracellular side of plasma membranes. The channel, or pore, facilitates transport of small molecules through the membrane. The head groups of phospholipids are depicted as spheres, although in actuality they differ in size and shape, as does the nature of fatty acyl side chains. (From Zafian PT. Plasma membrane alterations induced in bovine spermatozoa by cryopreservation. MS thesis, Colorado State University, 1984.)
At least three areas of specialization of the plasma membrane serve to anchor it to underlying structures.4,5 These are over the caudal ring of the head, the annulus, and the principal piece (Figure 102.5). The plasma membrane overlying the caudal ring, around the caudal end of the nucleus, is characterized by a striated band of intramembrane particles and fusion of the plasma membrane and the posterior ring. Over the annulus, the plasma membrane contains a densely packed band of particles which probably are involved in attachment of these structures. Finally, the principal piece has a longitudinal row of particles in or on the plasma membrane called the zipper. This is presumed to provide a firm attachment of the membrane and underlying structures in the principal piece. The plasma membrane overlaying the postacrosomal lamina may also be stabilized to underlying structures, because it is much less susceptible to vesiculation such as occurs during the acrosome reaction or senescent degeneration; alternatively it may have a unique lipid composition.
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Plasma membrane Perforatorium Acrosome Nuclear envelope Nucleus
Equatorial segment
Post-acrosomal lamina
Capitulum Segmented column Centriole Dense fiber Mitochondria Axoneme
Figure 102.5 Scale drawings of the head of a stallion spermatozoon. The head and neck are shown in longitudinal section and as viewed from the broad aspect. Major components include plasma membrane, acrosome, equatorial segment region of acrosome, postacrosomal lamina, and proximal centriole. Not shown is the close apposition of the plasma membrane and the underlying structures or its attachment to the underlying nuclear envelope at the caudal ring.
Head The head of a spermatozoon includes the nucleus with its nuclear envelope, the acrosome, postacrosomal lamma, and the plasma membrane (Figure 102.5). The shape of the head is determined primarily by shape of its nucleus. For stallion spermatozoa, the head and nucleus are broad and relatively flat. The nucleus is gradually tapered from a narrow rostral end to a thicker caudal end, with maximum width of the broad aspect in the center (Figure 102.5). The nucleus contains the highly condensed chromatin, DNA complexed with protamine, which typically is homogenous in appearance when viewed by an electron microscope. The nucleus is enclosed by the double-layered nuclear envelope, which contains few pores. The rostral portion of the nucleus is overlain by the acrosome, which is a specialized vesicle formed from a double-layered membrane (Figure 102.6). The acrosome in turn is overlain by the plasma membrane.2,4,5,16,17,22 The acrosome contains glycolipids and enzymes, at least some of which are bound
Figure 102.6 The head, neck, and proximal middle piece of a stallion spermatozoon. In the head, perforatorium (P), acrosome (A), plasma membrane (PM), nucleus (N) with nuclear vacuoles, postacrosomal lamina (PAL), caudal ring (CR), and basal plate (BP) are evident. In the neck and middle piece, the mitochondria (M), plasma membrane (PM), two dense fibers (DF), two doublets (D), and the central pair (CP) are designated. A tear or vesiculation of the plasma membrane, evident on the right side (arrow), is a common artifact seen in transmission electron micrographs of spermatozoa. Bar is 0.5 µm; magnification ×23 000.
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to the inner face of the inner acrosomal membrane rather than being contained in the contents (which appears amorphous under the electron microscope). Hyaluronidase, proacrosin/acrosin, and lipases are the primary enzymes. In stallion spermatozoa, the acrosome is slightly thickened but sharply tapered at the rostral end.16,17 No conspicuous thickening is found on one side (apical segment or ridge) as is characteristic of bull, boar, and ram spermatozoa. Occasionally, evidence for a stabilizing structure, the perforatorium, can be seen under the rostral aspect of the acrosome. Caudad, the acrosome is thinner and this area is termed the equatorial segment. The outer acrosomal membrane rostral to the equatorial segment is uniquely fusogenic, and fuses with the plasma membrane during the acrosome reaction, which precedes fertilization (Chapter 97). The equatorial segment of the acrosome does not contain enzymes and is not involved in the acrosome reaction, but the plasma membrane in this area fuses with that of the oocyte. The postacrosomal lamina covers the caudal portion of the nucleus, equatorial segment of the acrosome, and posterior ring (see Figures 102.2 and 102.5). This ill-defined structure may have a secondary role in attachment of a spermatozoon to the plasma membrane of the oocyte at fertilization. The caudal ring is a point of fusion between the plasma membrane and the nuclear envelope, at the caudal end of the head. The implantation fossa, at the base of the head, is a ball-and-socket articulation which serves to attach the neck (and the rest of the spermatozoon) to the head. The outer layer of the double-layered nuclear envelope lining the implantation fossa is thickened into a distinct basal plate. The basal plate provides the actual attachment with the neck. This attachment is fragile. In stallion spermatozoa, in contrast to those from other common mammals, the implantation fossa often is acentric in position, with respect to the breadth of the cell, and thus about 50% of stallion spermatozoa have an ‘abaxial tail’23 (Figure 102.7). In the rare cases when a spermatozoon actually has two tails, two implantation fossae exist. Neck The neck is the connection between the middle piece and the head (Figure 102.8). It contains a complex structure termed the connecting piece, the proximal centriole, several small mitochondria, and redundant nuclear envelope.2,4,5,17 The neck region is fragile, and the connecting piece contains several specialized elements, namely the segmented columns and capitulum. Malconnection of the head and neck may oc-
(a)
N BP CAP C
SC
SC
M
(b)
N BP SC C
(c) Figure 102.7 A, Scanning electron microscopic view of spermatozoa with axial or abaxial (arrows) attachment of the tail. Bar is 5.0 µm, magnification ×34 000. The implantation fossa and neck region of spermatozoa with a central attachment (B) or an abaxial attachment (C) of the tail, as viewed by transmission electron microscopy. The nucleus (N), basal plate (BP), capitulum (CAP), centriole (C), segmented columns (SC), mitochondria (M), and dense fibers plus axoneme of the middle piece are evident. Bar is 0.5 µm; magnification ×26 600.
cur during spermatogenesis as a hereditary defect, or separation can be induced because of hereditary epididymal malfunction or other reasons.13 The nine segmented columns are formed from a series of about 15 overlapping plate-like structures, formed from a fibrous protein. Each segmented column is fused, in the neck region, to the rostral origin of one of the nine dense fibers (Figure 102.8), which are components of the middle and principal pieces.
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Spermatozoal Function
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Nucleus Nuclear envelope Postacrosomal lamina Plasma membrane Basal plate Posterior ring Capitulum Proximal centriole
Segmented column Dense fiber
Dense fiber
Doublet
Mitochondrion
Radial spoke Central pair Axoneme Figure 102.8 Sketch of the neck region of a spermatozoon. The implantation fossa, at the base of the head, includes the basal plate which is attached to the capitulum, the most proximal structure of the principal piece. The capitulum consists of the cranial segments of the segmented columns which are coupled to the dense fibers. When the tail is detached from the head, usually it is because of separation of the basal plate and capitulum.
The segmented columns are not continuous with the dense fibers. Two pairs of the nine segmented columns are fused together over most of their length to form two major segmented columns; the remaining five are termed minor segmented columns. The two major segmented columns differ in shape (Figures 102.7 and 102.8). The primary one bends sharply and gives rise to the major portion of the capitulum, which is an enlarged head or ball which serves as the attachment with the basal plate of the head. The two stacks of plates in the other major segmented column are not fused together until they are rostral to the centriole, and then extend as a single fused stack of plates fused to the capitulum. Together, the two major segmented columns and capitulum form a continuous, elongated, articulation which has its long axis in the same plane as the broad axis of the spermatozoal head. Thus, the major segmented columns might be thought of as running up the ‘sides’ of the neck. The five minor segmented columns bend over and merge with the capitulum on the dorsal and ventral aspects of the neck area. The proximal centriole is located between the major segmented columns, within the anterior end of the connecting piece. This centriole has the characteristic nine-fibril structure and is positioned at a 45–60◦ angle to the tail axis. In stallion spermatozoa, the proximal centriole is relatively large in comparison with diameter of the neck, and typically one end is near the plasma membrane (see Figure 102.7). The proximal centriole and connecting piece likely constitute the site where beat of the tail is initiated in mature
spermatozoa. The distal centriole, seen in the neck region of a spermatid during development, disappears during formation of the connecting piece. A membranous outpocketing of redundant nuclear envelope lies lateral to the connecting piece. These loops or scrolls represent excess nuclear envelope remaining as the spermatid nucleus underwent condensation and a reduction in volume during spermiogenesis. Middle piece The middle piece extends from the caudal end of the neck, distad through the annulus.2,4,5,17 It is characterized by the presence of numerous mitochondria arranged circumferentially, end to end, in a continuous double spiral (Figures 102.9 and 102.10). The terminations of two mitochondria in one winding of the coil are bordered by the middle of individual mitochondria in the turns above and below. A typical stallion spermatozoon has about 50 gyri, or helical turns, of mitochondria. Cristae are clearly visible by transmission electron microscopy within individual mitochondria. The mitochondria contain the enzymes and cofactors necessary for production of ATP (see next section). The exact mechanism by which energy stored in ATP is converted to contractile activity of the tail is unknown. The outer shell or membrane of a mitochondrion is rich in thiol bonds, and resistant to dissolution. Mitochondrial DNA is present, but its role in cellular function or reproduction is obscure. Central to the gyri of mitochondria are the nine dense fibers. These have a tough, keratin-like fibrous
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Mitochondrion
Cross-bridge
Dense fibers Plasma membrane
1
Nexin link
2
9 1
Doublet 2
9
3
8 Radial spoke
3
8
4
7 4
7 Inner dynein arm
6
6
5
Longitudinal column
5
Fibrous rib
Outer dynein arm
Central sheath
Central pair of tubules Figure 102.9 Sketch of the middle piece (left) and principal piece (right) showing the plasma membrane, mitochondria or fibrous ribs, and nine dense fibers arranged around the axoneme, consisting of nine doublets and the central pair. Dense fibers and doublets are numbered as designated. Passing distally through the principal piece, the dense fibers become thinner and terminate; dense fibers 1, 5, and 6 are the longest.
structure. They extend from their origin in apposition to segmented columns, in the neck of the spermatozoon, through the length of the middle piece and most of the principal piece. They gradually taper away in the caudal principal piece and are not present in the end piece. Among the nine outer dense fibers (see Figure 102.9), numbers 1, 5, and 6 are larger than the rest; number 9 is intermediate in size; and the rest are small. The dense fibers do not contract, but probably dampen the arc of flagellar beat by providing rigidity concurrently with flexibility. A convention exists for numbering the dense fibers and doublets. A line tangential to the midpoints of the microtubules in the central pair divides the axoneme and dense fibers into halves (see Figure 102.9). Four dense fibers or doublets are in either half, and one (pair) is bisected. The bisected dense fiber or doublet is called doublet 1, and doublet 2 is that on the side of doublet 1 to which its arms point. Dense fibers are numbered the same as the corresponding doublet. The axoneme of a spermatozoon, central to the dense fibers, is identical in organization with the contractile structures of motile cilia of the trachea or certain microorganisms. The axoneme is the propulsive element of a spermatozoon, and the method by which spermatozoa move is considered in a subsequent section. The axoneme consists of a central pair of micro-
tubules surrounded by a ring of nine doublets (Figure 102.11). The role of the central pair is unclear. Each doublet is composed of a small cylindrical A microtubule, with an attached C-shaped B microtubule5 (Figures 102.9 and 102.11). The cylindrical A microtubule has two longitudinal rows of arms pointing toward the next doublet. Both microtubules of a doublet contain tubulin molecules arranged to form 13 protofilaments in the A microtubule and 9 or 10 in the B microtubule. The arms contain dynein, which is rich in ATPase and transduces chemical energy to mechanical motion. The entire doublet is about 70% tubulin and 15% dynein. The doublets are interconnected by nexin links, and a series of nine radial spokes extend from the central pair to the doublets (see Figure 102.10). The nexin links probably are elastic and maintain symmetry while regulating displacement of the doublets during their sliding and contraction. The spokes probably provide structural support. The annulus is an electron-dense structure which lies between the caudal-most gyrus of mitochondria and the rostral end of the fibrous sheath of the principal piece (see Figure 102.10). Thus it demarcates the end of the middle piece. In cross-section, the annulus appears as an isosceles triangle with its base against the mitochondria. The annulus is a point where the plasma membrane is firmly attached.
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Spermatozoal Function A B
MP
Protofilament
Nexin link 6
5
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Nexin link
4 Radial spoke 3
PP Outer inner Contral sheath
(a)
dynein arm
Radial spoke
Figure 102.11 Sketches of the axoneme showing the relationship between the A and B microtubules, dynein arms, nexin links, radial spokes, and central pair of microtubules. (Adapted from Satir82 and Linck.84 )
A
MP
Principal piece
PP
(b)
PM
M
1
6
3
5
(c)
3
The dense fibers and axoneme of the middle piece continue through the principal piece, although the dense fibers become narrower and terminate in the caudal principal piece.2,4,5,17 First dense fibers 3 and 8 taper away, then dense fibers 4 and 7, then dense fiber 9, and finally dense fibers 1, 5, and 6. The unique feature of the middle piece is the fibrous sheath with its fibrous ribs and longitudinal columns, both of proteinaceous material. In a sagittal section through the principal piece (see Figures 102.9 and 102.10), the fibrous ribs are cut in cross-section, and appear as dense strands, which pass halfway around the fibrous sheath, from one longitudinal column to the other. Sometimes the fibrous ribs branch. Longitudinal columns run the length of the principal piece, and overlay doublets 3, 4, and 8. Thus longitudinal columns are found on the dorsal and ventral aspects of the principal piece. Progressing caudad, the fibrous sheath lies closer to the axoneme, as the dense fibers taper away. The fibrous sheath probably provides the structural support plus flexibility essential for effective translation of sliding motion and contraction of the doublets into flagellar beats of defined flexure and amplitude.
3 End piece
Figure 102.10 (a) Surface views by scanning electron microscopy of a middle piece (MP) and a principal piece (PP) of spermatozoa. Bar is 0.2 µm; magnification ×41 000. (b) Longitudinal section through the middle piece and principal piece, showing the annulus (A). Bar is 0.2 µm; magnification ×45 000. (c) Cross-sections through the middle and principal piece. Compare with Figure 102.9. The vertical line bisects dense fiber 1 and the central pair and passes between dense fibers 5 and 6. Dense fibers 1, 5, and 6 and doublet 3 are designated. Bars are 0.2 µm, and magnifications are ×60 000.
The doublets and central pair of the axoneme continue intact about halfway through the end piece, and then taper out over a short distance of 1–2 µm. Thus the 9 + 2 pattern is lost in the caudal end of the end piece.
Biochemistry and metabolism of spermatozoa Semen The chemical compositions of stallion semen, seminal plasma, and spermatozoa have been
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tabulated.24,25 Although there have been few comprehensive studies of either seminal plasma or spermatozoa, biochemical characteristics of stallion semen are similar to those for semen from other domestic mammals.26 However, distinctive features are noted. Stallion semen contains considerable glucose and very little fructose, 0.82 and 0.02 mg/mL, respectively, unlike bull or ram semen in which fructose is the main glycolyzable sugar.25 Stallion semen also contains a high concentration of sorbitol and a substantial amount of lactic acid.26 The pH of normal stallion semen is 6.2–7.8, and the osmolality is between 300 and 334 mOsm/kg.24,25 Differences in pH between first and second ejaculates (7.47 vs. 7.59) have been reported.27 Season also affects seminal pH and osmolality. Stallion semen contains glyceryl phosphoryl choline (GPC), ergothioneine, and citric acid which are produced by the epididymis, ampullary glands, and vesicular glands, respectively. Analysis of semen for these compounds can provide information concerning the secretory contribution by each of these glands to a given ejaculate.24 When several ejaculates from a single stallion were analyzed, GPC, ergothioneine, and citric acid were present at 3.8, 0.8, and 2.6 mg/mL of semen.24 The sequence in which fluids from these glands are expelled during ejaculation has been determined by analyzing fractions collected at 5- to 10-second intervals during ejaculation.24,26,28 Ejaculation begins with a presperm fraction, which is watery in appearance and contains no spermatozoa, GPC, ergothioneine, or citric acid. It probably is from the bulbourethral glands and prostate gland. The second fraction is delivered a few seconds later. This fraction, which may encompass several jets of semen, has a milky appearance, is rich in spermatozoa, GPC, and ergothioneine but contains little citric acid or gel. This fraction, therefore, contains secretions from the epididymis (spermatozoa and GPC) and ampulla of the ductus deferens, but not the vesicular glands. The third fraction, contains few spermatozoa or ergothioneine, but large quantities of gel and citric acid. This fluid originates primarily from the vesicular glands and washes out the few spermatozoa remaining in the urethra. The final fraction, postcoital fluid which drips from the stallion’s penis at dismount, has a watery appearance and contains few spermatozoa and little ergothioneine. It is not characteristic of the entire ejaculate.24 Occasionally, infertility might result from defective function of the epididymis, ductus deferens, ampulla, or vesicular glands or from disturbances in sequence of the ejaculatory process (vesicular gland secretions discharged before or simultaneously with the sperm-rich fraction).26
Composition of spermatozoa Only limited information exists on lipid composition of stallion spermatozoa.21,29 Total lipid content of stallion spermatozoa is 14.2% (w/w), and the ratio of cholesterol to phospholipid is 0.23.29 By comparison, cholesterol to phospholipid ratios are 0.22 for ejaculated bull spermatozoa30 and 0.44 for ram cauda epididymal spermatozoa.31 Parks et al.30 reported that the plasma and outer acrosomal membranes contain different lipid and protein compositions. Presumably, regional differences in lipid composition give the different membrane compartments their specific functions (see previous section). Analysis of the lipid composition of ram epididymal spermatozoa revealed that the major phospholipids were phosphatidylcholine (63%), phosphatidylethanolamine (15%), and lysophosphatidylcholine (10%); diphosphatidylglycerol, phosphatidylinositol, and phosphatidylserine were present in minor quantities (< 10%).30 The fatty acid docosahexanoic acid (22 carbons and 6 double bonds) is unique to spermatozoa and makes up about 50% of the total fatty acids in spermatozoa. Palmitic acid, a 16-carbon saturated fatty acid, makes up about 30% of the total spermatozoal fatty acids, while myristic acid (14 carbons, saturated) and stearic acid (18 carbons, saturated) each constitute about 10% of the total fatty acids in ram spermatozoa.30 Proteins constitute > 50% of the total weight of spermatozoa.24 Although many of these are histone proteins involved in DNA packaging,24 others are involved as enzymes in the acrosome, receptors on the plasma membrane, elements of membrane function (ion and carbohydrate transport), and cytoskeletal structures (giving shape to head or providing filaments of the tail). Nothing is known about specific proteins in stallion spermatozoa, and a detailed discussion of these proteins would be beyond the scope of this book. Several surface proteins have been visualized in spermatozoa from other species using fluorescently labeled lectins32,33 and antibodies.34–36 The distribution patterns (quantity and location) of some of these proteins change during epididymal maturation and during capacitation. Immunosuppressive proteins in seminal plasma apparently bind to spermatozoa.37 Other proteins originating from epididymal or seminal plasma are removed from spermatozoa during capacitation to facilitate membrane fusion and exposure of spermatozoa–oocyte receptors.9,37 However, functions of most proteins on the plasma membrane remain unknown. Metabolism of spermatozoa Spermatozoa are unique in that they lack the capability to divide and have only limited biosynthetic
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Spermatozoal Function Glucose-6-phosphate
Glucose
Phosphohexoisomerase Fructose-6-phosphate
Fructose Anaerobic metabolism (glycolysis)
Lactate + 2 ATP + 2 H2O
Tricarboxylic acid cycle
36 ATP/Glucose 36 ATP/Fructose
Aerobic metabolism
Figure 102.12 Overview of carbohydrate metabolism in stallion spermatozoa. Not all metabolites are shown. Glucose is readily metabolized to fructose-6-phosphate and then to lactate and energy. Alternatively, fructose can enter the glycolytic pathway one reaction step below glucose. Both substrates yield identical end products and identical amounts of energy (2 ATP during anaerobic metabolism and 36 ATP during aerobic metabolism).
capacity.38 Mature spermatozoa lack systems to repair cell damage or synthesize new enzymes. Therefore, most constituents necessary for spermatozoal function or metabolism are synthesized during spermatogenesis, and spermatozoa only perform maintenance functions and produce needed energy. Energy is used by spermatozoa to (i) initiate catabolic processes such as glycolysis, (ii) maintain motility, and (iii) maintain ion balance and miscellaneous cell functions.38,39 Spermatozoa rely primarily (about 90%) on extracellular substrates to meet their energy requirements. This energy is principally derived from carbohydrates (Figure 102.12). Spermatozoa readily metabolize monosaccharides such as glucose and fructose but do not metabolize all sugars or more complex carbohydrates.24,39 Stallion spermatozoa, however, possess limited capacity to use fructose when compared with spermatozoa from other species and cannot metabolize sorbitol, a major component of stallion seminal plasma.24 Other exogenous substrates used by spermatozoa include lactic acid, glycerol, fatty acids, and amino acids.24 Extensive use of exogenous glucose and fructose by ram24 and bull40 spermatozoa has been demonstrated. Ram spermatozoa in neat (undiluted) semen will metabolize all available monosaccharides in seminal plasma within 15–20 minutes at 37◦ C and stop swimming,24 but this is not a problem with stallion spermatozoa.
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Metabolizable sugars are transported across the plasma membrane by specific transport proteins, a process which has not been studied in stallion spermatozoa. In bull spermatozoa, however, the transport of glucose is the rate-limiting step in metabolism of exogenous sugars.39 Glucose transport in spermatozoa is an ATP-dependent process which can be blocked by inhibitors such as cytochalasin B,41 and is not coupled to Na+ or K+ transport39 as is the case for many other cell types. At least in bull spermatozoa, the glucose transport protein has only limited capability to transport fructose.39 A separate fructose transport protein may be present in bull and ram spermatozoa, which enables them to readily metabolize fructose. A fructose transport protein may be absent in stallion spermatozoa, which would explain the limited capability of stallion spermatozoa to metabolize fructose. Metabolism of endogenous compounds contributes about 10% of the cell’s energy and ATP production in a physiological state.42 Spermatozoa can metabolize endogenous substrates, such as by oxidation of phospholipids in the mitochondria.24,42 Spermatozoa have no glycogen stores and lack the enzymes necessary to use glycogen as an energy source.38 In bull spermatozoa, ATP production from endogenous substrates remains constant regardless of the presence or absence of exogenous substrates or the temperature of the cell.38 However, production of ATP from exogenous substrates, and hence total production of ATP, is temperature dependent. Use of ATP also has been studied, but not for stallion spermatozoa. In bull spermatozoa, about 60% of ATP produced is used to maintain motility, but nearly 40% of ATP produced by spermatozoa is wasted by substrate cycling. Substrate cycling is the repeated phosphorylation and dephosphorylation of glycolytic intermediates,43,44 so that no net production of ATP occurs. Spermatozoa are different from other cell types, in that they spend very little energy on transmembrane ion movement.44 Only negligible amounts of ATP are used for maintenance of ion pumps and all other cell processes. Several external ions affect spermatozoal motility and respiration. Compounds which stimulate spermatozoa include PO4 – and Na2 HCO3 2– and low concentrations of K+ or Mg2+ .24,45–48 Ions which inhibit spermatozoal motility and metabolism include H+ , Mn2+ , Ca2+ , and high concentrations of Mg2+ .24 Species differences exist in the capability of spermatozoa to use aerobic respiration. Bull and ram spermatozoa can use anaerobic respiration at a much higher rate than stallion spermatozoa, which rely heavily on aerobic metabolism.24 Glycolytic breakdown of certain monosaccharides, especially glucose
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for stallion spermatozoa, provides the major source of ATP in spermatozoa. Human spermatozoa maintain a high rate of glycolysis even during aerobic respiration and maintain motility in the absence of oxygen and/or in the presence of mitochondrial inhibitors; thus, they have only a limited need for aerobic respiration.49,50 One consequence of aerobic metabolism is production of hydrogen peroxide in the mitochondria, which induces peroxidation of lipids in the plasma membrane.51–55 Lipid peroxidation causes dysfunction of certain metabolic enzymes; increases permeability of the plasma membrane;53 and is deleterious to structural integrity, motility, and viability of spermatozoa.51 Lipid peroxidation is depressed by seminal plasma and antioxidants such as butylated hydroxytoluene (BHT).51,55 When neat or extended semen remains undisturbed, aerobic metabolism exhausts the oxygen dissolved in the extender or seminal plasma and the spermatozoa are forced to rely on anaerobic metabolism. This occurs, for example, when semen is cooled slowly from body temperature to 5◦ C. After oxygen is depleted, spermatozoa use only glycolytic metabolic pathways and the end product is lactic acid.24 Accumulation of lactic acid reduces the pH of the medium, which may cause a decrease in metabolism and ATP production with a resulting loss in spermatozoal motility. Rabbit spermatozoa rapidly die when exposed to a pH of less than 5.8.24 Maintaining a proper intracellular pH for stallion spermatozoa (accomplished by maintaining the extracellular medium at a pH between 6.2 and 7.8) is an important factor in sustained spermatozoal motility and metabolism. Maintenance of proper intracellular pH is complicated during storage of spermatozoa at low temperatures, because pH of the extender is affected by temperature, buffer(s) in the extender, buffer capacity, organic co-solvents used as cryoprotectants, and increased ionic strength of the medium during freezing when water leaves the solution by crystallizing into ice. Changes in pH caused by reduced temperature arise in part from changes in the pH scale, which is temperature dependent, and changes in the acid dissociation constant (pKa) of water and many biological buffers; both are markedly affected by temperature. Taylor illustrated this pH dependence on temperature by the pH of water at 25◦ C (pH 7.0), 0◦ C (pH 7.47), and –35◦ C (pH 8.4).56 Changes in pH induced by organic co-solvents are typified by addition of dimethyl sulfoxide to aqueous solutions, which can increase the pH by 0.1 to 0.2 pH units.57 Increased ionic strength, incurred during freezing, lowers hydrogen activity coefficients and therefore alters pH.58
Compartmentalization of spermatozoal function Spermatozoa contain only those cellular components necessary for them to traverse the female reproductive tract and fertilize an oocyte. Specialized compartments include the plasma membrane, nucleus, acrosome, neck, mitochondria, and axoneme. Each has a specific function and consists of several classes of molecules (i.e., lipids and proteins), which because of the molecules and/or their stoichiometry, make each compartment different from all others. Some compartment differences are easily recognizable. For example, the spermatozoal nucleus contains the DNA, the middle piece contains all mitochondria, and the axoneme contains primarily microtubules and contractile microfilaments. Other, more subtle, differences can also be detected within many compartments. Most intracellular compartments are bound by a membrane which has a distinct composition of lipids and proteins, different from all other compartmental membranes, which impart a designed characteristic to that compartment. Mixing of membrane proteins or lipids from one compartment to another rarely occurs. However, transfer of molecules between the acrosomal and plasma membranes does occur during the acrosome reaction when vesiculation of those membranes occurs. Uniqueness of the lipid and protein constituents of each compartment membrane is obvious from several types of evidence. The plasma and acrosomal membranes contain different relative amounts of phospholipid and cholesterol. Molar cholesterol to phospholipid ratios of 0.38 for plasma membrane and 0.26 for acrosomal membrane are reported.30 Differences exist in abundance and location of individual peripheral proteins as detected by antibodies.34–36 Differences in carbohydrate residues on membrane proteins, detected using lectins, also exist.33,59,60 These techniques reveal that although proteins are specific to certain membrane compartments, they can migrate to other membrane compartments. Differences in membrane potential, specific protein content, and macromolecular permeability between compartments have made it possible to analyze the integrity of each compartment in a spermatozoon. Specific fluorescent stains specific for DNA have been used to assess the status of this cell compartment.61–64 Integrity of the plasma membrane covering the head can be assessed using specific stains,13,65–67 whereas the integrity of the plasma membrane covering the tail can be assessed using a hypo-osmotic swelling assay68 or column filtration using Sephadex or glass wool.69,70 The acrosome contains hydrolytic enzymes (including acrosin, acid phosphatases, β-glucuronidase,
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Spermatozoal Function neuraminidase, N-acetylglucosaminidase, and hyaluronidase), at least some of which are required for fertilization.5,71 The acrosome includes a large amount of carbohydrate which enables staining of the acrosome using the periodic acid–Schiff reaction.72 The acrosome also can be visualized by other stains73 and the acrosomal contents of spermatozoa from several species, including the stallion, specifically bind Pisum sativum agglutinin (PSA) lectin.73–76 A specific monoclonal antibody to acrosomal protein(s) in stallion spermatozoa is available.77 All of these probes can be used to assess acrosomal integrity in stallion spermatozoa with a reasonable degree of accuracy. Fluorescent probes also have been developed for assessment of mitochondrial function.75,78,79 The ability to use multiple assays for individual spermatozoal compartments illustrates the uniqueness of each compartment and the functional isolation of one compartment from all others. The underlying principle is that any spermatozoon capable of fertilizing an oocyte must have each of at least four attributes or functions operational at 100% of normal at the correct time, when it approaches the oocyte. The proportion of completely functional spermatozoa in a sample can be determined only by evaluating individual spermatozoa for each of a battery of important traits. Population averages are less sensitive; for a given characteristic, a value of 0.2 might result because 20% of all spermatozoa gave a 100% response (a potentially desirable situation) or because 100% of the cells gave a 20% response (a potentially undesirable situation). The percentage of motile cells is useful information because it distinguishes between these two cases. However, oxygen consumption or acrosin content of a spermatozoal population (rather than for individual cells) is less informative. Multiparameter tests of individual spermatozoa75,78,80,81 circumvent this problem.
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surrounding medium also alters beat frequency and bending pattern of the flagellum.85 Bending of the tail results from shear forces generated between neighboring doublets which cause one doublet to slide, temporarily, a short distance past the other.5,82,83 Dynein arms are the pivotal component of this sliding system, and the vigor of sliding depends on ATP and a cyclic adenosine monophosphate (cAMP) dependent protein kinase. The sliding or shear force is not developed simultaneously over the entire length of a doublet, but probably is localized in a portion of a doublet(s). Doublets 1–4 alternate contractions with doublets 6–9, and the site of localized contractions may pass around the nine doublets in a circular manner. How do the doublets slide to induce contraction? In the neutral or immotile state, dynein arms extending from microtubule A are long and associated with microtubule B of the adjacent doublet5,82,83 (Figure 102.13). Although the dynein arm is permanently attached to microtubule A, attachment with microtubule B is transitory and broken by availability of ATP. Thus local availability of ATP causes the dynein arm to detach from microtubule B and, concurrently, to shorten. This shortening is similar to ATP-driven contraction of a muscle fiber. The free arm then tilts at an angle of about 40◦ , elongates, and reattaches to microtubule B at a new site. Contraction and return of the attached arm to a 90◦ (horizontal) angle with the doublet creates a shear or sliding force sufficient to move the second doublet past the first doublet, toward the tip of the axoneme. By this slight movement of about 16 nm, the dynein arms attached to microtubule A ‘walk’ along microtubule B of the neighboring doublet. The initial detachment and shortening of the dynein arm occur only with availability of ATP, tilting to a 40◦ angle and elongation of the detached dynein arm depend on hydrolysis of ATP to
No ATP Rigor
+ ATP Release
ATP ADP + energy Re-extension Re-binding
Elevation Sliding
Mechanics of doublet sliding Forward motion of a spermatozoon results from bending of the flagellum, a consequence of coordinated bending waves propagated from the neck. The bending movement probably results from the interplay of two systems: (i) a sliding filament system in the doublets, which generates the force for the bending wave, and (ii) a control system, which coordinates the sliding and regulates flagella beat.5,82–84 The ultimate pattern of bending of the flagellum results from active forces within the tail, working against constraints imposed by the nexin links, radial spokes, dense fibers, and fibrous sheath. The viscosity of the
Dynein arm elongated and attached
Dynein arm detaches and shortens
Dynein arm tilts, elongates and attaches at lower site
Dynein arm shortens and returns to horizontal ‘‘rigor’’-producing sliding
Figure 102.13 Mechanism of doublet sliding within the axoneme resulting from detachment, shortening, elongation, and reattachment of the dynein arms in a localized region (only one is depicted), followed by return to the horizontal and rigor position. The return to horizontal slides one doublet relative to the other. (Adapted from Satir.83 )
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provide needed energy, and final shortening of the arm is independent of ATP because it is the ‘resting state’ (Figure 102.13). Although both the inner and outer dynein arms probably contribute an equivalent sliding force, contraction can occur in the absence of the outer dynein arms. Sliding per se cannot create the bending motion of the flagellum. Presumably, bending is induced by anchorage of the doublets and dense fibers in the neck region and resistance offered along the entire length of the tail by the nexin links and other structures. Bends appear to form passively, some distance from the point of active formation and breaking of dynein attachments, as a consequence of resistance to displacement of the microtubules. As noted above, the doublets on one side of the axoneme contract more or less in synchrony. Thus a planar tail beat results from active shear forces and sliding on one side of the axoneme, induced by the dynein arms, concurrent with passive sliding and relaxation on the opposite side. In the case of a helical tail beat, frequently evident as spermatozoa rotate or roll along their long axis, a slight twisting may be generated as the bending motion progresses along the principal piece, because of a staggered or rotational activation of dynein arms in doublets 1–9, rather than synchronous contractions of doublets 1–4 or 6–9. The control mechanism activating opposing groups of doublets to contract possibly is modulated by Ca2+ , acting through calmodulin. Initiation or maintenance of spermatozoal motility depends on maintenance of a necessary intracellular concentration of cAMP, synthesized from ATP by adenylate cyclase, and the calcium/calmodulin system is involved.5,38,44 In addition, adenosine, bicar-
bonate, and the protein carboxymethylation enzyme system also are likely involved. The interactions of these regulatory systems are unclear for mammalian spermatozoa in general and have not been studied for stallion sperm. The pattern of spermatozoal motion To a casual observer viewing spermatozoa through a microscope, not all stallion spermatozoa swim in a similar manner. Detailed analysis of the motion characteristics of motile spermatozoa can be achieved by computer-assisted image analysis; comprehensive discussions of these instruments have been published.86–90 Systems for computer-assisted analysis of spermatozoal motion are accurate if properly used.87,91,92 Basically, the microscopic image of a sample of spermatozoa in an optically clear medium is detected by a video camera and sent, either directly or via a videotape interface, to a computer which captures information in 15–60 frames (typically 0.5–1.0 at 30 frames/s), locates each spermatozoon in each frame, and reconstructs their paths. The conventional terminology used to describe spermatozoal motion is summarized in Table 102.1 and Figure 102.14. Several instruments are available from commercial vendors. Although all use similar principles, actual algorithms and strategies for considering spermatozoa which collide or leave the field of view differ. Thus slightly different data will be obtained with each brand of instrument even if the same samples are analyzed. Also, values selected for user-defined settings can influence results. If properly used, these instruments can provide valuable objective data on spermatozoal motion, but data for other attributes
Table 102.1 Nomenclature used to describe spermatozoal motiona Term
Description
Curvilinear velocity (VCL) Average path velocity (VAP)
The centroid-to-centroid path, and velocity along that path The smoothed centroid-to-centroid path, as calculated by one of several smoothing algorithms, and velocity along that path The linear path between the first and last centroid in a sequence, and velocity along that path VSL/VCL VSL/VAP VAP/VCL Spermatozoa with a VAP or VCL > v µm/s Spermatozoa with a VAP or VCL > v µm/s and a LIN or STR > 1 Spermatozoa with a VAP or VCL > v µm/s and radius of the average path < r µm/s Mean distance of the centroids from the average path The number of times the curvilinear path crosses the average path, expressed as intersections per second
Straight-line velocity (VSL) Linearity (LIN) Straightness (STR) Wobble (WOB) Motile spermatozoa (MOT) Progressively motile spermatozoa (PMOT) Circularly motile spermatozoa (CIR) Lateral head displacement (LHD) Beat cross frequency (BCF)
a Distance is expressed as µm and velocity as µm/s. Note that VCL > VAP > VSL for any given spermatozoon (see Figure 102.14). In determining if a spermatozoon is motile, a minimum cut-off value can be selected, but in some instruments it is for VCL and for other instruments it is for VAP. In most cases, an equine spermatozoon with VCL <15 or 20 µm/s or VAP <10 or 15 µm/s should be considered as immotile.
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1 Straight line path
3 2 Actual path
Curvilinear path
4
12 11
8
5
14
10 13
6 9 7 = Location of head centroid in each frame
1 3 2
Actual path Average path 4
12 8
11
5 6
14
10
9
13
Lateral head displacement
7
Figure 102.14 Path of the centroid (midpoint) of a spermatozoal head and some parameters measured or typically reported. The curvilinear path joins the centroids, and the average path is calculated by a smoothing algorithm. The straight line path connects the first and last centroid. Lateral head displacement is the distance between a centroid and the average path, and beat cross frequency is the number of times the curvilinear path crosses the average path. (From Amann.89 )
also should be considered when predicting potential fertility of a stallion. Although some spermatozoa are immotile, and may actually be dead, other spermatozoa pause momentarily and appear immotile only to resume swimming after a short interval. Watching motile spermatozoa, a researcher might note that they swim with different velocities, in a rather progressive manner with or without rotation along the long axis of the cell or sometimes swim in a curvilinear manner. This is evident in computer-based analysis of spermatozoal motion (Figure 102.15). Other spermatozoa might actually be swimming backward, because of a 180◦ reflection of the tail near the juncture of the principal piece and middle piece. In addition, the pattern of flagellar beat and the nature of head motion can be influenced by the viscosity and chemical composition of the medium in which the spermatozoa are suspended, temperature, and depth of the preparation.85,88,93 Given the size of a stallion spermatozoon, the amplitude of flexations of the tail around the theoretical longitudinal axis of a swimming cell, and the wobble of the head as a spermatozoon rotates around its longitudinal axis, unfettered motion of a spermatozoon requires a sample depth of at least 50 µm and possibly 100 µm. Unfortunately, microscopic examination of spermatozoa in a preparation with a depth ≥ 50 µm is impossible with conventional instruments, because of the mutual exclusivity of the requirements for adequate magnification and retention of a motile
cell within the depth of field of the microscope objective viewing the preparation.93 Thus, to enable critical observation of spermatozoal motility, we must restrict the freedom of spermatozoal movement, in the belief that this does not negate validity of the observations. This assumption probably is correct if the depth of the preparation is ≥ 10–12 µm, but if depth of the preparation is ≤ 10 µm spermatozoa find it increasingly difficult to move.93,94 Erroneous conclusions are likely if depth of the preparation is < 10 µm. Preparations typically used by a theriogenologist are 15–30 µm in depth, assuming an appropriatesized drop on a microscope slide is overlain with a cover glass. Restriction of the sample to a depth of 15–20 µm facilitates viewing and is obligatory with most computer-assisted systems for automated analysis of spermatozoal motion.86,87 This can be achieved by placing 7 µL under an 18 × 18-mm cover glass,95 10 µL under a 22 × 22-mm cover glass, or use of special chambers (µ-Cell, Fertility Technologies, Inc., Natick, MA) made for computer-assisted analysis of spermatozoal motion. For visual observation, with continuous focusing on different levels in the preparation, volumes of 12–18 µL under a 22 × 22-mm cover glass are better. The problem of sample thickness must be solved by compromise; use of a preparation ≥ 12 µm deep, but not so deep as to make use of the microscope difficult. Using an HTM-2000 motility analyzer (HamiltonThorne Research, Danvers, MA), with dark field
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S7 S8 Average path Centroid Curvilinear path S9
Velocity average path (µm/sec)
c102
110
9.0
90
7.0
70
5.0
50 3.0 30 1
5
7 8 9 10 15 0.5 - Sec segment of track
Characteristics
Slideb
Capillaryb
Total motile spermatozoa (% ≥ 5 µm/s) Motile spermatozoa(% ≥ 20 µm/s) Progressively motile spermatozoa (%)c Average path velocity (µm/s) Straight-line velocity (µm/s) Lateral head displacement (/s) Linearity
58 55 31 91 46 13 0.51
44∗ 42∗ 28 98 57† 11∗ 056†
a
LHD (µm)
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Figure 102.15 Motion of a typical stallion spermatozoon over 10 seconds. The centroid location, curvilinear path, and average path during three 0.5-second segments (segments 7, 8, and 9) are shown in the upper portion of the figure. Mean path velocity (velocity over the average path, open squares) and lateral head displacement (LHD, open circles) are shown for all 20 segments in the lower portion. For all 20 segments, curvilinear velocity (centroid-to-centroid), path velocity, and LHD averaged 65 µm/s, 94 µm/s, and 5.1 µm/s. For segments 7, 8, and 9, values were 48, 81, and 102 µm/s for curvilinear velocity, 32, 39, and 81 µm/s for path velocity, and 3.1, 7.5, and 6.4 µm for LHD. Although curvilinear velocity was 68% greater during segment 8 than during segment 7, path velocity was only 22% greater; this was because LHD was large. (Modified from Amann.86 )
illumination and infrared light, Amann et al.88 studied the motility of spermatozoa in a fiat capillary tube 200 µm deep and in a preparation about 16 µm deep on a glass slide. Semen was diluted to 25 × 106 spermatozoa/mL in an optically clear medium (10 g sucrose plus 3 g ovine serum albumin per 100 mL water). They found that sample thickness affected percentage of motile spermatozoa, straight-line velocity, lateral head displacement, and linearity (Table 102.2). In other words, spermatozoa swam differently in the 16-µm preparation than in the deep capillary tube, where proximity to a glass surface did not impede their motion. Surprisingly, a lower percentage of spermatozoa were motile in capillary tubes than on slides, but they had higher straight-line velocities and linearities. The authors gave no explanation for the differences in velocity,88 and none is evident for the difference in percentage of motile spermatozoa. However, a reasonable conclusion is that restrictions imposed by upper and lower surfaces of the 16-µn preparation precluded many spermatozoa from rotating smoothly about their long axis and caused them to
Means for spermatozoa from 10 ejaculates extended in 10% (w/v) sucrose plus 3% (w/v) bovine serum albumin and evaluated 5–20 minutes and 95–110 minutes after collection, using five fields per evaluation; variance associated with time was not significant. Values designated by ∗ or † differ (P <0.05 or <0.01). b The slide sample had a depth of about 16 µm and each field was scanned twice. The capillary had a depth of 200 µm and each field was scanned once. c Spermatozoa with a path velocity > 5 µm/s and a straightness > 0.5. Adapted from Amann.86
thrash their heads and flagella more to complete each 180◦ rotation. If this assumption is correct, it could explain the increased lateral head displacement (a measure of how much the head is wobbling around the general path of movement) and decreased linearity and straight-line velocity (measures of how effectively a spermatozoon is moving from point A to point B in a linear manner). Changes detected probably should be interpreted as evidence that the 16µm preparation somewhat restricts movement of a spermatozoon from point A to point B within such a preparation (Table 102.2). Nevertheless, the evaluation of percentage of motile spermatozoa was not compromised by use of preparations only 16 µn deep. Effects of temperature on spermatozoal motion often are ignored by clinicians, because they are considered to be unimportant or because adequate temperature control of equipment used to prepare the slide, and of the slide during viewing, requires extra effort or expense. None of these reasons for inadequate control of temperature while viewing a sample of semen is justified and, depending on environmental conditions, can lead to erroneous results. Under most environmental conditions, failure to control temperature adequately while performing a seminal evaluation will provide incorrect information to the client; because the clinician should have known better, this is malpractice. Effects of temperature on motion of stallion spermatozoa88 when suspended in a 200-µm-deep flat capillary tube are summarized in Table 102.3. Temperature affected all parameters of spermatozoal motion (P < 0.05), although spermatozoa in some ejaculates were affected more than those in others; the ejaculate by temperature interaction was significant for percentage of motile spermatozoa, velocity
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Table 102.3 Effects of temperature on motion characteristics of stallion spermatozoaa Characteristic
22◦ C
27◦ C
32◦ C
37◦ C
42◦ C
Total motile spermatozoa (% ≥ 5 µm/s) Motile spermatozoa (% ≥ 20 µm/s) Progressively motile spermatozoa (%) Curvilinear velocity (µm/s) Average path velocity (µm/s) Straight-line velocity (µm/s) Linearity Lateral head displacement (µm)
53b 49b 33b 87b 51b 28b 34b 7.0b
66c 63c 46c 100c 57c 35c 36c 8.1c
66c 64c 53d 107d 62d 42d 42d 8.4c
68c 67c 58e 114e 69e 51e 47e 8.4c
66c 63c 56d,e 117e 75f 56f 50f 86c
a
Based on analyses of spermatozoa in each of five ejaculates; semen was diluted in an aqueous solution of 10% (w/v) sucrose and 3% (w/v) bovine serum albumin and held at about 30◦ C until analysis. All analyses using an HTM-2000 instrument were completed within 3 hours of seminal collection, and each sample was evaluated in order of ascending temperature; time probably had a minimal effect in confounding effects of temperature. Means in a row with a different superscript differ (P <0.05). Adapted from Amann.86
parameters, and lateral head displacement. The important point is that at 22◦ C, essentially room temperature, the percentages of motile spermatozoa or progressively motile spermatozoa were significantly lower than at higher temperatures. Maximum percentage of motile spermatozoa was obtained only at 37◦ C or 42◦ C.86,88 Spermatozoa swam with greater velocity and linearity at higher temperatures, and more spermatozoa had a velocity of > 60 µm/s for observations at 37◦ C or 42◦ C than at cooler temperatures (Figure 102.16). However, the effect of temperature on velocity characteristics is at least partially independent of its effect on percentage of motile spermatozoa. What does this mean for a clinician? It is unlikely that a clinician will consider a spermatozoon with a linear velocity < 20 µm/s as motile. Assuming this to be correct, and based on the data summarized in Table 102.3, Amann et al.86,88 calculated that for a sample in which 72% of spermatozoa were motile at 37◦ C, only 56% would be judged to be motile at 22◦ C. Thus, failure to have a controlled temperature introduced a 22% reduction, or error, in the percentage of motile spermatozoa. A different human observer might require a spermatozoon to be moving at 30 or even 40 µm/s to be considered motile. In these cases, the errors would be 33% and 51%, respectively. However, if the observers requiring motile spermatozoa to be moving 30 or 40 µm/s had been evaluating samples at 37◦ C, the errors would have been only % and 14%, respectively. Thus, adequate temperature control at 37◦ C is essential to allow spermatozoa to swim with their potential characteristics. More important, observation at 37◦ C is essential to minimize differences from sample to sample associated with velocity of spermatozoa within that sample or among observers. The take-home message is obvious: failure to evaluate samples at 37◦ C might preclude 30% of potentially motile spermatozoa from swimming with suf-
ficient velocity to be considered motile by a human observer. Therefore, evaluations of seminal quality at room temperature often will lead to invalid estimates of spermatozoal quality. No doubt the nature of the medium in which spermatozoa are suspended10,85,93 and the physiological
22 °C 27 °C 32 °C
80 60
37 °C 42 °C
40 Percentage of sperm (v > 0)
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20
60
100
150
200
22 °C
80
37 °C 60 40 20
0
20
60
100
150
200 > 200
Velocity average path (µm/sec)
Figure 102.16 Effect of temperature on average path velocity of stallion spermatozoa. Expressed as the percentage of distribution for about 800 spermatozoa with v < 0 in each of five ejaculates, as determined using an HTM-2000 instrument. Although means for average path velocity differed (P < 0.05) at 22◦ C, 27◦ C, 32◦ C, 27◦ C, and 42◦ C, distributions at 22◦ C and 27◦ C were similar, as were those at 37◦ C and 42◦ C (upper; ignores values for v < 200). The distribution was different (P <0.01) at 22◦ and 37◦ C (lower). (Modified from Amann.86 )
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status of the cells92,96 influence how spermatozoa swim. However, the important question is not what are the motion characteristics of spermatozoa in vitro when in medium A or B or even how spermatozoa swim at sites U, I, or F within the female reproductive tract. The important questions are whether evaluations of spermatozoal motility, either subjectively by a human observer or by a computer-assisted system, accurately portray the quality of spermatozoa in a given sample and whether they indicate the potential for those cells to achieve fertilization.6 The answer to both questions probably is a qualified yes,6,70,86,87,89 but the important proviso is that spermatozoal motility is not the only attribute required for survival within the female reproductive tract or achievement of fertilization. No data for motion characteristics of spermatozoa in vivo within the uterus or uterine tube exist, but spermatozoa do not necessarily swim in the same manner as they do in vitro. Indeed, based on studies focused on in vitro fertilization, clinicians have concluded that spermatozoa which have undergone the acrosome reaction, a prelude to fertilization, swim differently from spermatozoa which have not undergone this modification of the plasma membrane and underlying structures.97 As outlined previously in this chapter, a successful spermatozoon must have attributes in addition to those of ‘a good dancer’ to fertilize an ovum. The successful spermatozoon must elude the natural defense mechanisms of the female, receive nurturing at receptive sites, retain the ability to respond to chemoattractants produced by oocytes (if they exist in mammals), and ultimately penetrate the investments of the oocyte. Motility certainly is a prerequisite for normal fertilization in terms of penetration through the zona pellucida and attachment to the plasma membrane of the oocyte. However, this does not mean that all spermatozoa which are motile when examined in vitro have the necessary attributes to fertilize an oocyte. Indeed, recent research strongly supports the conclusion that a subpopulation of motile spermatozoa, shortly after ejaculation or after preservation at 5◦ C or –196◦ C, have altered plasma membranes which probably preclude their successful participation in fertilization.70 Thus, not all motile spermatozoa are capable of fertilizing an ovum, although most motile spermatozoa likely have the potential to achieve this task.
Responses of spermatozoa to thermal stress Cooling stallion spermatozoa to 5◦ C is necessary for storing spermatozoa to be used within a few days either at the location of collection or after ship-
Table 102.4 Damage to specific membrane compartments of stallion spermatozoa after cold shock (%) Membrane compartment status Intact plasma membrane; intact acrosome Damaged plasma membrane; intact acrosome Damaged plasma membrane; damaged acrosome
Control 30◦ C
Cold shockeda 0–4◦ C
81
36
9
23
11
42
Cells were cold shocked by immersion into water at 0–4◦ C. Data from Watson et al.112
a
ment to another location or as a prerequisite for cryopreservation.23 Cooling stallion spermatozoa, however, stresses the cell and can cause cellular injury.20,22,98 Understanding the causes of cooling damage and how to minimize it is essential if spermatozoa stored at low temperatures are to retain maximal fertility. Cellular injury can be caused directly by affecting cellular structures (such as by rupturing membranes) or indirectly by altering cellular functions (such as slowing down metabolic processes).20 Rapid cooling of stallion spermatozoa from 20◦ C (room temperature) to 5◦ C induces partially irreversible damage characterized by an abnormal pattern of swimming, rapid loss of motility, damage to acrosomal and plasma membranes, reduced metabolism, and loss of intracellular components. Collectively, this damage is termed cold shock.7,20,99 Stallion spermatozoa are susceptible to cold shock, and cooling damages several spermatozoal membrane compartments (Table 102.4).Indirect cooling damage is more difficult to discern and may not be evident until sometime after the spermatozoa have reached 0–5◦ C.7,99 Responses of membranes to cooling To understand how cooling and/or cold shock damages spermatozoa, the clinician must consider both spermatozoal structure (see Figure 102.2) and function. The original concepts of the fluid-mosaic membrane model100 provided a membrane model of proteins floating like icebergs in a sea of lipids composed primarily of phospholipids and cholesterol. The membrane consists of a bilayer having the hydrophilic portions of both lipids and proteins oriented externally and their hydrophobic portions oriented toward the bilayer center.100–102 Modifications of this model include asymmetry of membrane lipids and proteins and complex interactions between lipids and proteins that regulate membrane receptors, ion pores or enzymes, and ultimately membrane
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(a)
(b) Figure 102.17 Schematic representation of the two polymorphic phase forms for lipid in biological membranes. Normally, the bilayer (a) has a planar configuration with phospholipid heads (spheres) oriented toward the bulk water and fatty acyl chains inward. The bilayer forms a continuous permeability barrier excluding polar molecules. (b) The relatively rare hexagonal-II phase has a cylindric form with phospholipid head groups toward the center and hydrocarbon tails oriented outward. This structure weakens the permeability barrier but may play an important role during physiological fusion of membranes. (Adapted from Hammerstedt et al.18 )
function.18,101,102 In addition, alternative forms of lipid aggregates exist within a membrane.101–103 A minor lipid aggregate form, the hexagonal-II phase (Figure 102.17), provides a point defect capable of inducing membrane fusion rather than maintaining a permeability barrier.104 However, a similar aggregation to a hexagonal-II phase may be necessary for the acrosome reaction and fertilization. Under stress, such as induced by cooling, membranes may undergo rearrangement into the hexagonal-II phase, and the formed point defects then may induce excessive membrane permeability or even membrane disruption.18 Each spermatozoal membrane (e.g., plasma, acrosomal, and mitochondrial) is a unique aggregate of lipids and proteins arranged in a bilayer. The lipid and protein compositions of each membrane are unique, and little or no exchange of lipid or protein among them occurs. This compartmentalization arises during spermiogenesis and is maintained during epididymal modifications; it permits each membrane to maintain its unique function.4,102,105 Few data are available on the lipid composition of mem-
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branes from stallion spermatozoa. However, cholesterol is the major sterol in stallion spermatozoa, and the cholesterol to phospholipid ratio for whole stallion spermatozoa is 0.36.21 Glycolipids make up < 10% of the polar lipids in stallion spermatozoa.21 More complete biochemical analyses are available for bull spermatozoa.30 In addition to lipid analysis of whole spermatozoa, the isolation of membranes from individual compartments of bull spermatozoa revealed compositional differences. The plasma membrane of a bull spermatozoon has a higher cholesterol to phospholipid ratio (0.38) than the outer acrosomal membrane (0.26). Differences in amounts of individual phospholipid species also were detected between membrane compartments.30 Lipid composition is important in membrane function, because each lipid species contributes its unique characteristics to the membrane. Lipids undergo a phase transition, from a fluid or liquidcrystalline state, in which the fatty acyl chains are relatively disordered, to the gel state, in which the fatty acyl chains are increasingly rigid and parallel to each other as temperature is reduced. Proper membrane function requires a fluid membrane. Each lipid species undergoes this phase transition at a temperature specific for that lipid. The phase transition temperature of a lipid is determined by the nature of the head group, length of fatty acyl chains (short chains have a lower transition temperature), and degree of unsaturation of fatty acyl chains (chains with more unsaturated bonds have lower transition temperatures). When the temperature of a membrane is decreased below the transition temperature of an individual lipid species, that lipid (but not other lipids) will undergo a phase transition and aggregate in microdomains of lipid gel in an otherwise liquid membrane. This has two effects. First, lipids which were randomly dispersed throughout the membrane interacting with specific lipid species and proteins aggregate into microdomains and no longer interact with their specific membrane partners. Second, borders between gel microdomains and fluid portions of the membrane create ion-permeable gaps which are unstable and can lead to membrane fusion or rupture. Lipid asymmetry is an additional level of molecular complexity beyond the uniqueness of each membrane compartment. Membranes of cells consist of a stabile bilayer having the total phospholipid pool equally allocated to the inner and outer leaflets of the bilayer. Each phospholipid species, however, has a preference for one of these two leaflets. Phosphatidylcholine and sphingomyelin prefer the outward facing leaflet, whereas phosphatidylethanolamine, phosphatidylserine, and phosphatidylinositol prefer the inner, cytosolic leaflet. Membrane stressors (i.e., cooling) that alter orientations of lipids could affect
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membrane stability, particularly as each lipid species has an individual preference for the bilayer or hexagonal-II phase states when under stress.106 Lipid–protein interactions provide another level of membrane molecular complexity. Not all lipids in a membrane prefer the bilayer orientation. In a stable bilayer membrane, these lipid species, which tend to impede retention of a bilayer, are found in close association with integral membrane proteins.106 These lipid–protein interactions mediate smooth melding of the protein into the bilayer, eliminating holes or pores where proteins contact the lipids of the bilayer, and may be requisite to proper function of the protein as an enzyme, receptor, or ion channel by inducing proper protein conformation.102 For each membrane compartment in a stallion spermatozoon, the proper lipid and protein constituents have evolved to permit proper function when the membrane is in the fluid state at 39◦ C. As spermatozoa are cooled, the bulk membrane undergoes a phase transition into the gel state. This occurs at 20.7◦ C for stallion spermatozoa.21 In addition, individual lipid species undergo a phase transition and aggregate into microdomains, the proteins associated with these lipids may cease to function as designed because lipid–protein interactions are disrupted (Figure 102.18). Upon rewarming, the nonbilayer-preferring lipids within any microdomain may not return to apposition with the proper protein but aggregate in a hexagonal-II phase.106 Aggregation of lipids into gel-phase microdomains forces membrane proteins of the glycocalyx to aggregate in the remaining fluid lipid domains.101 Function of such glycocalyx components can be altered as stoichiometry of proteins in the membrane is altered. This can be caused by alterations in attachment of integral proteins with cytoskeletal assemblies or by protein coupling.101,107 Responses of proteins to cooling Intracellular proteins are affected by cooling. For example, as temperature is reduced, enzyme function is reduced. The relative rate of activity of adenylate cyclase was reduced by 90%, from 1630 pmol cAMP formed per minute per milligram protein when bull spermatozoa were cooled from 37◦ C to 10◦ C.108 Phosphodiesterase activity decreased similarly.108 Proteins involved in transport of substances across the membrane also have reduced activity at low temperature. Proteins which form the Na+ /K+ and Ca2+ pumps are decreased to 0.25% of their activity at 37◦ C when at ≤ 6◦ C. This decrease in pumping ability appears to be associated with the protein itself and not
Native membrane
Cooled to 4°C
Warmed to 37°C
Bilayer at 37°C with hexII phase
= Lipid favoring bilayer configuration = Lipid favoring nonbilayer configuration Figure 102.18 Schematic representation of probable effects of cooling and rewarming on distribution of lipids around integral membrane proteins. The native membrane represents the spermatozoal plasma membrane at seminal collection; lipids tend to impede retention of bilayer associated with integral membrane proteins (blocks). Cooling to 4◦ C causes lipids tending to impede bilayer formation to undergo a phase transition and cluster together in the gel state; this forces membrane proteins to cluster together and associate with lipids tending to favor retention of the bilayer. However, clustering of membrane proteins alters membrane function. Rewarming to 37◦ C produces a membrane that has lipid–protein associations different from those of the native membrane, and lipid–lipid associations may form hexagonal-II configurations. A complete return to the native membrane configuration will occur only if lipid mobility is sufficient to allow all membrane components to find their ‘original’ partners. (Adapted from et al.18 )
the surrounding lipids.109 Glucose transport in the red blood cell is reduced at 0◦ C to < 0.1% of its transport at 37◦ C.109 Reductions in both enzyme and transport activities can be caused by combination of (i) decreased kinetics of molecules at lower temperatures; (ii) altered pH of aqueous portions of the cell and surrounding medium at low temperature, inducing a change in pK of the protein, its electrical charge, and its function;109,110 (ii) altered lipid–protein interactions, because of removal of preferred lipids (those tending to prefer the hexagonal-II phase) from the protein and reduced lateral movement of proteins in membranes in which lipids are in the gel phase;101 and (iv) irreversible denaturation of proteins.107,110
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With the preceding background on effects of cooling on cell components, one can begin to understand effects of cooling on spermatozoa. Rapid cooling of spermatozoa induces cold shock, but slow cooling does not entirely circumvent the problem.20 Spermatozoa stored at 5◦ C undergo changes observed in cells undergoing senescence (see later), but at a reduced rate.99 Cooling appears to dissociate decreased fertility and decreased motility in contrast to senescence. Cooled ram spermatozoa maintain motility (upon warming), but have reduced fertility, whereas aged spermatozoa lose both attributes simultaneously.99,107 This dissociation probably arises from differences in the composition of the membrane compartments. Cooling a membrane induces gel state microdomains with leaky borders. Normally, ions which leak into the cell (Na+ and Ca2+ ) are pumped back out by ion transporters. However, at 5◦ C permeability of spermatozoal membranes to Ca2+ is increased107 and the effectiveness of the Ca2+ transporter is decreased;107,109 thus Ca2+ accumulates in the cell. Probably the acrosomal membrane becomes more permeable to Ca2+ at 5◦ C than mitochondrial membranes associated with motility, and the high Ca2+ concentration in the acrosome induces a premature vesiculation of these membranes.107 Differences in the biochemical composition of these membrane compartments (acrosome vs. mitochondria) may explain why fertility declines more rapidly than motility in cooled spermatozoa. Cold-shocked spermatozoa exhibit acute and extensive damage. Changes in spermatozoal motion are easily recognized (Figure 102.19). The percentage of motile cells is decreased7,20,98,107 and motion characteristics of the motile spermatozoa are altered. In spermatozoa from many species, a non-reversible bending of the middle piece and coiling of the tail occurs which causes the tail to fold back on itself and the spermatozoa to swim backward.20,107 Investigations on metabolism of cold-shocked ram and bull spermatozoa revealed decreased anaerobic glycolysis111 and respiration.107,111 This was caused by losses of ATP, glycolytic enzymes, and cytochrome c as a result of membrane damage. However, uptake of a fluorescent dye, rhodamine 123, by mitochondria was evidence that the mitochondria still were functioning.107 Microscopic examinations of cold-shocked ram spermatozoa revealed increased permeability of the plasma membrane to vital stains, localized swellings of the flagellum, and damage to the acrosome (swollen, broken, or missing) in a high proportion of spermatozoa.107 Electron microscopic evaluation
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Figure 102.19 Effects of slow cooling to 4◦ C and cold shock, by direct immersion of vial containing extended semen in 4◦ C water, on velocity of spermatozoa. Velocity for most spermatozoa in semen extended in E-Z Mixin shortly after ejaculation was < 30 µm/s (I) and averaged 84 µm/s. After cooling extended semen slowly to 4◦ C, the velocity distribution for motile spermatozoa was not greatly altered (SC) and still averaged 84 µm/s; fewer spermatozoa were motile (73 vs. 84%, initially). For spermatozoa subjected to cold shock (CS), however, few spermatozoa had a velocity < 30 µm/s. Means are for one ejaculate from each of three stallions. Semen are extended to 25 × 106 /mL in EZ Mixin. Samples were evaluated with a Stromberg-Mika Cell Motion Analysis System (SM-CMA, 4.3 Bad Fellienbach, Germany).
revealed that the plasma membrane overlying the acrosome was particularly vulnerable to damage and damage was accompanied by loss of the acrosomal matrix.107 Stallion spermatozoa subjected to cold shock exhibit similar cellular damage as ram and bull spermatozoa.112 Extensive disruption of the plasma membrane covering the head of stallion spermatozoa was noted. In contrast to ram spermatozoa, mitochondria in stallion spermatozoa also were damaged, indicating that cold stress directly affected the mitochondrial cristae.112 Subtle mitochondrial damage probably is the cause of loss of motility, because little structural damage was observed in the spermatozoal middle piece or principal piece. Although cooling spermatozoa to 4◦ C or 5◦ C has some deleterious effects on spermatozoal function, when properly done it reduces metabolism and the
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rate of senescence so that a high proportion of spermatozoa are motile and fertilizing capacity is retained after 24 hours at 5–80◦ C.98,113–116 Minimizing thermal stress The maturity of spermatozoa affects their susceptibility to thermal stress. Epididymal spermatozoa are more resistant to cold shock than are ejaculated spermatozoa.1 Although no data for stallion spermatozoa exist, incubation for ≥ 20 minutes in seminal plasma possibly will reduce sensitivity to cold. Indeed, boar spermatozoa are sensitive to cold shock immediately after ejaculation, but acquire resistance to cold-shock damage after incubation in seminal plasma.117,118 A similar but less pronounced change is observed with ram spermatozoa.107 A variety of additives reduce cold-shock injury in spermatozoa.99,107 Compounds such as ethylenediaminetetraacetic acid (EDTA), which bind Ca2+ and Mg2+ , are likely to reduce leakage of Ca2+ into the cell. Most other protective compounds contain lipids or lipophilic molecules and likely affect the plasma membrane, although the exact mechanism by which they affect the cell is unknown. Phillips and Lardy119 demonstrated that egg yolk protected bull spermatozoa from cold shock. Egg yolk also protects ram spermatozoa120 and stallion spermatozoa121–123 but fails to protect boar spermatozoa from cold shock.124 The protective action is provided by the low-density lipoproteins of the egg yolk,125–129 especially phospholipids.125,128,130–132 Phospholipids are believed to stabilize the membrane, but their mode of interaction is not understood. Lipid composition of the membrane apparently is not altered after addition of egg yolk,125,130 and washing spermatozoa free of egg yolk removes its protective capabilities.128,130,131 Extenders containing skim milk or cream have been used for preservation of stallion spermatozoa.23,98,113–115,133 Little research has been done investigating the mechanism of milk’s protective action against cold-shock damage to spermatozoa. However, milk lipoproteins probably act similarly to egg yolk lipoproteins in protecting spermatozoal membranes. Milk proteins also may act by directly stabilizing protein elements in the spermatozoal membrane.99 Other additives, such as butylated hydroxytoluene (BHT)134–137 and phosphatidylserine,138,139 have been used with bull and boar spermatozoa. These compounds directly alter the composition of the plasma membrane and affect the membrane’s fluidity and susceptibility to cold-shock damage.99,134 Whatever their mechanism for minimizing coldshock injury to spermatozoa, these compounds per-
mit the cooling of spermatozoa for storage as liquid semen or in conjunction with cryopreservation. Spermatozoa cooled to or stored at 5◦ C maintain fertilizing capacity, although cooled spermatozoa possess slightly altered function (motility and acrosomal integrity) upon rewarming and insemination. To maximize retention of fertilizing capacity, stallion spermatozoa should be (i) mixed with at least three volumes of an appropriate extender, (ii) cooled no faster than 0.05◦ C/min between 18◦ C and 8◦ C,107 and (iii) incubated at low temperature (3–6◦ C) for as short an interval as possible and preferably < 36 hours.
Changes in spermatozoa occurring before fertilization or during aging and senescence The biochemistry of capacitation and the acrosome reaction Freshly ejaculated stallion spermatozoa, like spermatozoa from all mammals, are unable to fertilize oocytes without undergoing further modification (see Figure 102.1). One major modification involves changes in the spermatozoal plasma membrane which facilitate fusion of the plasma membrane with the outer acrosomal membrane during the acrosome reaction. In addition, spermatozoal motion is modified and spermatozoa change from moving progressively forward to a non-progressive motion in which a spermatozoon exhibits whiplash-like tail movements causing the head of the spermatozoon to move in a star-like or figure-of-eight pattern. This modified motility is referred to as hyperactivated motility or hyperactivation.5,140 The processes which induce these spermatozoal modifications are referred to as spermatozoal capacitation. Some of these processes are enhanced by the female reproductive tract and its secretions, but most will proceed to some extent in vitro. Spermatozoa undergo aging and senescence after ejaculation as well, which include spontaneous changes in the spermatozoal plasma membrane and metabolism that render spermatozoa infertile. These changes are similar to changes seen during spermatozoal capacitation. The requirement for spermatozoal modification before fertilization first was observed in 1949 by Noyes, Finkle, and Rock, although never published,141 and was reported by Austin142 and Chang.143 Austin coined the term capacitation to describe these changes.144 Spermatozoal capacitation is defined as changes in a spermatozoon, particularly the plasma membrane, that allow the cell to
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Spermatozoal Function undergo the acrosome reaction, which is a reaction to physiological concentrations of free Ca2+ which induces vesiculation of the plasma membrane and outer acrosomal membrane and is accompanied by an altered beat pattern of the flagellum resulting in hyperactivated motility.4,5,145 These changes require 18–20 hours for stallion spermatozoa in vitro.146 However, the time required for capacitation to occur in utero is not known. Comprehensive studies of this process have been difficult because no obvious morphological change occurs in spermatozoa during capacitation.147 Reviews of molecular mechanisms underlying capacitation and the acrosome reaction are available.140,145,147–152 However, clinicians should be aware of the reversible changes in spermatozoal metabolism, cell surface components, and membrane alterations that occur during capacitation as a prelude to the acrosome reaction and hyperactivated motility. After ejaculation, major changes occur in surface components of spermatozoa during capacitation. Although the female tract supplies spermatozoa with energy substrates, this environment is not necessarily optimal for spermatozoa. Therefore, spermatozoa acquire a glycoprotein coat during epididymal transit, secreted by epididymal epithelium, to which is added proteins from the seminal plasma upon ejaculation. These proteins help maintain membrane integrity during their sojourn in the female reproductive tract. One important part of capacitation is the gradual removal or alteration of the adsorbed protein coat to alter transmembrane ion flux, to expose spermatozoal membrane receptor sites, and to remove compounds covering the tail which may restrict flagellar hyperactive motion.141 Similar changes occur over the caudal and rostral portion of the spermatozoal head, the middle piece, and principal piece. Changes in the plasma membrane over the head facilitate binding of the spermatozoon to the zona pellucida and occurrence of the acrosome reaction. Changes in the plasma membrane over the tail facilitate hyperactive motility, and those over the mitochondria facilitate increased energy output.140 In addition to changes in the glycoprotein coat covering the spermatozoa! plasma membrane, the plasma membrane itself undergoes specific changes during spermatozoal capacitation. One major change is efflux of cholesterol from the plasma membrane.150,153 Cholesterol is transferred nonspecifically to albumins in the female tract and is transferred to a high-density lipoprotein of follicular origin (apolipoprotein A-I) in the uterine tube by a specific transfer protein.153 Cholesterol plays a significant role in stabilizing membranes. Thus, loss of cholesterol tends to destabilize the sperma-
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tozoal plasma membrane, particularly that portion over the acrosome, because this portion of the plasma membrane has a high concentration of phospholipids containing docosahexanoic acid, a fatty acid which enhances membrane fluidity. Destabilization of the membrane by removal of cholesterol permits reorganization of components in the bilayer, including redistribution of integral proteins. Clustering of these proteins facilitates receptor–ligand interactions, exposure of membrane-bound enzymes, and alters binding characteristics of surface receptors.153 Destabilization affects the lipid portion of the membrane altering ion permeability, particularly Ca2+ and membrane fusogenic properties, both of which are important for the acrosome reaction.140 During capacitation, spermatozoal metabolism increases before onset of hyperactivated motion. Respiration of rabbit spermatozoa increases fourfold during that time and appears to be directly linked to an increase in intracellular cAMP.148 Increased ATP production is required as spermatozoal velocity and the amplitude of flagellar movement increase during capacitation.148 Hyperactivated motility of spermatozoa at the site of fertilization may be necessary for spermatozoa to escape binding to epithelial cells of the uterine tube and to penetrate through the zona pellucida.154 The time required to induce these changes, via capacitation, ensures induction of the acrosome reaction at the surface of the zona pellucida and not earlier during a spermatozoon’s passage up the female tract. The acrosome reaction must occur in the vicinity of the egg, because fusion of the outer acrosomal membrane with the overlying plasma membrane releases the acrosomal enzymes needed for spermatozoal penetration through the zona pellucida;154 premature loss of those enzymes would preclude fertilization. In addition, loss of the outer acrosomal/plasma membrane vesicles after the acrosome reaction exposes the inner acrosomal membrane, which contains receptors for sperm–egg binding.140 Spermatozoa with an intact outer acrosomal membrane cannot bind to the egg plasma membrane and initiate fertilization. The biochemistry of spermatozoal senescence induced by aging The natural saga for ejaculated stallion spermatozoa includes (i) deposition into the reproductive tract of the mare, (ii) transport through the female tract (i.e., through cervical mucus, uterus, and uterine tube155 ), (iii) capacitation, (iv) exhibition of the acrosome reaction and hyperactived motility, (v) binding to and passing through the zona pellucida, (vi) penetration of the plasma membrane of the oocyte, and (vii) nuclear and chromosome decondensation and union of
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Table 102.5 Effect of aging of spermatozoa in vitro on fertility (%)a Age of semen at time of insemination (days) Species
1
2
3
4
5
Reference
Horseb Cattleb Cattleb Rabbitc
71 64 55 73
85 65 66 52
50 61 63 95
67 54 54 63
– 49 39 27
160 161 162 163
a
Based on use of aliquots of the same semen sample each day. Fertility reported is 180-day non-return rate. c Fertility based on number of 6-day embryos and corpora lutea. b
male and female pronuclei to produce a one-cell fertilized egg.152,156 In horses, this sequence usually progresses normally if the stallion breeds the mare or artificial insemination occurs within 0–36 hours before ovulation. If deposition of spermatozoa into the mare occurs 0–6 hours after ovulation, fertility will not be greatly reduced, but deposition of semen in the mare 12 to ≥ 24 hours after ovulation will result in reduced fertility and foaling rate (Chapter 123). The stallion, type of sperm (fresh cooled or frozen–thawed), and mare all probably affect the interval after ovulation when a successful insemination can occur. If spermatozoa are deposited in the mare early or after spermatozoa have been stored in vitro (i.e., when spermatozoa are stored at 5◦ C or frozen–thawed), the timing of this sequence of events may be altered. Changes from this scenario arise because of spermatozoal aging. Although no research reports the effects of aging on stallion spermatozoa, stallion spermatozoa likely respond in a manner similar to spermatozoa from other species. Roche et al.157 demonstrated that when freshly collected rabbit spermatozoa were mixed in equal numbers with rabbit spermatozoa aged for 24 hours at 5◦ C, the freshly collected spermatozoa were responsible for 95% of the resulting offspring.157 Thus spermatozoal senescence can occur before insemination and during normal handling of spermatozoa between the time of semen collection and insemination. Aging also can occur in the female tract if spermatozoa are inseminated too early before ovulation and must reside in the female tract longer than is optimal.158 It is difficult to distinguish aging phenomena from capacitation because some results of both processes are similar.159 However, aging results in decreased fertilizing capacity, decreased spermatozoal motility, and increased embryonic mortality. Spermatozoal capacitation leads to a spermatozoon becoming competent to fertilize an egg. Inseminating females with spermatozoa that had been stored for 1–5 days before insemination revealed gradually reduced pregnancy rates in mares,
cattle, and rabbits (Table 102.5). However, fertility often is greater on day 2 than on day 1. Storage of spermatozoa for 4–5 days resulted in lower pregnancy rates. Assessing spermatozoal fertility can be difficult because fertility and pregnancy rates are not the same because of embryonic loss. This discrepancy arises because defective as well as physiologically fit spermatozoa can initiate embryonic development.164 Perhaps for this reason, fertility of semen stored for short periods of time resulted in higher fertilization rates than semen used immediately. An ejaculate may contain defective spermatozoa which can compete with normal spermatozoa immediately after ejaculation, but these spermatozoa may not survive storage and are unable to compete with normal spermatozoa when insemination occurs after 2 or 3 days of storage.159,162 The loss of fertility resulting from aged spermatozoa probably is caused by the inability of aged spermatozoa to initiate fertilization because of defective motility, capacitation, acrosome reaction, acrosomal enzymes, binding capability, or genomic abnormality. Rabbit or hamster spermatozoa aged in the uterus gave a decreased fertilization rate and development of resulting embryos was slower than normal. However, no increase in percentage of embryos with chromosomal abnormalities was noted.163,165 Rabbit spermatozoa aged in vitro at 5◦ C for 5 hours or 5 days resulted in kindling rates of 87% and 44% of ovulated oocytes, respectively.165 Part of the decrease in live young was caused by abnormal embryonic development resulting from aged spermatozoa. Size of 6day-old embryos was reduced for embryos resulting from spermatozoa aged 5 days (9 µL volume) compared with embryos resulting from fresh spermatozoa (26 µL volume). Embryonic death accounted for about one-third of the difference in kindling rate after insemination of aged rather than fresh spermatozoa. Maurer et al.164 concluded that when aged spermatozoa fertilize an oocyte, embryonic development is delayed which, in some cases, results in embryonic
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Spermatozoal Function death. However, most of these embryos result in live young. Mammalian (cow) and amphibian (frog) embryos resulting from aged spermatozoa lack several proteins found in normal embryos, probably because of defective genome translation, and this arrests embryonic development.162,166 However, the major loss in offspring born, accounting for about two-thirds of the differences in kindling rates, was not attributed to increased embryonic mortality. Rather, reduced fertilization rate by aged spermatozoa was the major cause of reduced kindling rate.164 No critical study using aged stallion spermatozoa has been reported. Use of transported semen is increasing and some reduction in fertility by aged spermatozoa might be expected. Therefore, age-related changes in spermatozoal function will be considered in detail. The physiochemical changes that occur during storage in vitro and in utero are largely irreversible.167 Spermatozoal defects induced by aging can be classified as (i) nuclear instability, (ii) membrane defects, (iii) loss of intracellular components (enzymes, ions, metabolites, etc.), and (iv) lipid peroxidation. Nuclei of spermatozoa are altered during long incubation at 37◦ C or 5◦ C.159 The actual DNA content of the spermatozoon remains unchanged during storage,159,167 but DNA–protein associations change.167 Samples of bull semen in which a high proportion of the spermatozoa had altered DNA packaging had low fertility.61,62 This also may be true for stallion spermatozoa,168 but additional studies are necessary. Therefore, alterations in DNA–protein associations, induced by aging, are likely to affect fertility of spermatozoa adversely. As spermatozoa age, plasma membrane defects arise. The acrosomal membrane swells, and all membranes become increasingly permeable.159 Permeability changes can be detected with live–dead or differential stains.73 Increased membrane permeability results in an inability to maintain proper intracellular ionic concentrations, because Ca2+ leaks into the cell faster than it can be pumped out. In addition, aged spermatozoa lose ATP and are incapable of resynthesizing it.159 Acrosomal enzymes, glycolytic enzymes, and transaminase enzymes also are lost. The loss of glutamic-oxaloacetic transaminase (GOT) has been correlated with spermatozoal damage.169–171 Lipid peroxidation is detrimental to spermatozoa. Lipid peroxidation is the interaction of oxygen with unsaturated fatty acids to produce hydrogen peroxide. This reaction occurs in spermatozoa from all species examined,54,172,173 with docosahexanoic acid, a primary fatty acid of spermatozoa, being especially vulnerable to peroxidation. The resulting lipid peroxides affect membranes, altering their permeability and inducing breakage.172 Because docosahexanoic
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acid comprises up to 67% of the lipids in the acrosomal membranes,172 acrosomal swelling and disintegration are commonly seen as a result of spermatozoal aging.159 The percentage of motile mouse spermatozoa decreases in a linear fashion as extent of peroxidation increases.54 Lipid peroxidation is a major cause of the membrane defects seen in aged spermatozoa. Permeability of the plasma membrane increases, so that vital stains, such as eosin, enter the cell. Similarly, intracellular constituents, such as glycolytic enzymes, leak out.172 Lipid peroxides also directly destroy and/or inactivate certain proteins and enzymes. Enzymes of the acrosome are susceptible to inactivation by lipid peroxidation.172 Loss of enzymes through inactivation, as well as membrane leakage, radically alter spermatozoal metabolism, resulting in a rapid and irreversible loss of motility and fertilizing capacity.54,172 Seminal plasma contains several substances that help minimize lipid peroxidation. Naturally occurring antioxidants containing thiols, such as ergothioneine, glutathione, and cysteine, are effective in eliminating peroxides.54 Stallion semen contains relatively high concentrations of ergothioneine (76 µg/mL semen) when compared with bull semen (trace amounts).24 Bull spermatozoa receive protection from their seminal plasma, which contains proteins which virtually eliminate lipid peroxidation, but seminal plasma of the stallion, ram, and human lack such proteins.172 Lipid peroxidation also is affected by diluent and handling procedures. Diluents containing a high concentration of K+ increase the rate of peroxidation,54 while addition of egg yolk decreases peroxidation,172 as do added antioxidants.174 In addition, rapid cooling, cold shock, and centrifugation increase peroxidation rates, whereas slow cooling and passing carbon dioxide gas over semen reversibly arrests motility, metabolism, and peroxidation.167 In summary, in early stages of senescence, lipid peroxidation triggers degenerative changes in spermatozoal membranes. These lead to acrosomal swelling and increased permeability of the acrosomal and plasma membranes. Increased membrane permeability results in loss of essential substances (both ions and enzymes), decreased metabolism and motility, and decreased fertility. Inclusion of certain additives in a seminal extender and avoidance of deleterious handling procedures can minimize this damage to spermatozoa.
Potential for separating X- and Y-chromosome-bearing spermatozoa Interest in determining the sex of offspring was expressed as early as 460–370 bc but has only recently
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been achieved. Unfortunately, even methods proven effective with mammalian spermatozoa in controlled small-scale studies, and carefully documented in the literature,171–178 are not appropriate for use in commercial artificial insemination, for which numerous doses of > 300 × 106 spermatozoa are needed. Two basic approaches exist for altering sex ratio at birth by manipulating spermatozoa before artificial insemination.179,180 The first is a physical separation to provide at least one population, if not two populations, enriched for either spermatozoa bearing an X chromosome or spermatozoa bearing a Y chromosome (hereafter termed X-spermatozoa and Y-spermatozoa). Obviously, enrichment to 80–85% for either type of spermatozoa would be of great economic importance, provided that fertility was not suppressed and that losses of spermatozoa in the separation procedure were minimal. This is the concept usually thought of for sexing sperm. However, a second and equally valid approach would be to selectively alter the function of either X-spermatozoa or Y-spermatozoa by a procedure that caused death of one type or enhanced the probability that spermatozoa of one type would fertilize the oocyte. With this strategy, a functional advantage is established, rather than a physical separation. Less than 30 years ago the critical role of the Y chromosome in determining the sex of mammals was established. Based on data for several species of mammals, evidence shows that if the primitive gonads develop as fetal testes, during the second month of gestation for colts, the previously bipotential reproductive system will develop into one characteristic of a male; following birth and puberty, the testes will produce spermatozoa. This is because the fetal testes secrete hormones, including testosterone and ¨ mullerian-inhibiting substance.181 In a normal fetus, the presence of a Y chromosome induces development of the gonads as testes. After a furious research effort during the past decade, researchers established that a single gene on the Y chromosome is responsible for determining how the primitive gonad develops. Based on data for mice and humans, the Sry gene is located on the Y chromosome; it encodes for a factor which. is obligatory for making a male gonad.182–185 This gene apparently is conserved across all common mammals and is expressed in somatic cells of the genital ridge only for a short interval just before overt testis differentiation. Animals with a Y chromosome lacking this gene develop as phenotypic females. Animals with two X chromosomes, one of which contains a translocated Sry gene, develop as sex-reversed phenotypic males rather than females. However, certain genes present on the X chromosome and autosomes, in addition to the Sry gene, are necessary for complete expression of maleness in the adult.
Occasionally, the Sry gene and other nearby genes are translocated from the Y chromosome to the X chromosome, as an aberration occurring in the primary spermatocyte that gave rise to the fertilizing spermatozoon. Such an XX embryo would develop as a male, because of presence of the translocated portion of the Y chromosome.184 Nevertheless, in most cases an XX embryo will develop as a mare and an XY embryo will develop as a stallion. An X-spermatozoon contains more DNA than a Yspermatozoon. For common domesticated mammals, this difference is about 3.5% of the total amount of DNA.63 However, because of variations in components of spermatozoa other than DNA content, scientists have been unable to separate X-spermatozoa from Y-spermatozoa consistently by techniques relying on total mass, size, or sedimentation velocity.179,180 A recent brief report, however, provided evidence that serial passage of bull spermatozoa through two 10-layer Percoll gradients gave a separation of X- and Y-spermatozoa.186 Confirmation of this report is awaited. With a technique termed flow sorting, scientists theoretically can distinguish an X-spermatozoon from a Y-spermatozoon on the basis of DNA content and collect spermatozoa of the desired type.63,64,175,178 Until recently, the problem had been that the treatments imposed on spermatozoa to enable precise quantitation of their DNA content, and thus separation, killed the spermatozoa. This problem has been overcome by modifying the treatment imposed on the spermatozoa, so that viable spermatozoa emerge from the flow sorting instrument.175–178 US Department of Agriculture (USDA) scientists developing this procedure have applied for a patent, and the USDA has licensed the procedure. If a patent is granted, the exclusive license for use of this process will influence attempts to use the technology. In any case, access to an instrument costing at least $300 000 would be necessary. Publications document birth of cattle, rabbits, and swine sired with spermatozoa isolated by flow sorting.175,177,178 Although fertility rates were low, the sex ratios of animals born were skewed in the expected directions (Table 102.6). Improvements in both the efficacy of separation and fertility of spermatozoa subjected to this procedure are likely to occur in the coming years. Unfortunately, use of this procedure probably will be restricted to sorting spermatozoa in a few samples from truly superior sires, to enhance the probability of obtaining male or female offspring from such individuals. This technique is likely to be used with in vitro fertilization of oocytes obtained from very valuable females. Appropriate methods are in place for cattle and could be developed for horses, if the techniques were not precluded
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Table 102.6 Success in predetermining sex of offspring by artificial insemination of flow-sorted spermatozoa Sex of offspring Species Cattle Rabbit Rabbit
Type of species sperm
Number of females
Fertility (%)
Male (no.)
Female (no.)
Correct sex (%)
X Y X Y X Y X+Y
75 75 60 60 14 16 17
15 27 32 35 21 31 29
3 12 4 10 1 17 6
8 8 15 11 15 4 8
73 60 79 48 94 81 –
by breed registry associations. At least for cattle, sufficient spermatozoa can be obtained by flow sorting to enable success using conventional artificial insemination, although with low fertility. Success with separating X-spermatozoa from Yspermatozoa by flow sorting175–178 or possibly Percoll gradients186 has not eliminated the need for development of a technique for isolating large populations of cells highly enriched for either X-spermatozoa or Y-spermatozoa of normal fertility or for differentially blocking fertilizing capacity of one type of spermatozoa. No information exists to convince a knowledgeable individual that a given procedure works consistently with spermatozoa from stallions. To conveniently obtain numerous, large samples of sperm treated to predetermine sex of offspring remains an unsolved intellectual challenge. Availability of samples highly enriched for either X-spermatozoa or Yspermatozoa with intact plasma membranes should facilitate development of large-scale methods. For example, several groups have attempted to detect proteins in the plasma membrane which are specific to either X-spermatozoa or Y-spermatozoa and use antibodies against sex-specific plasma membrane proteins to isolate, inactivate, or kill one type of spermatozoa.187 Although this approach has not proven successful, access to populations of viable spermatozoa isolated by flow sorting should facilitate development of this approach, if sex-specific plasma membrane proteins actually exist. Both the scientific literature and popular magazines are filled with articles and anecdotal data purporting to prove or disprove that a given method for treating semen alters sex ratio at birth. Virtually without exception, these reports should be given little credence. Reasons for discounting most literature on this popular topic have been discussed elsewhere.177,178 Most publications purporting to prove or disprove that a given procedure results (or does not result) in an alteration of sex ratio of embryos or offspring are meaningless, because the authors have failed to
Reference 175 175 177
consider limitations imposed by binomial variation (male vs. female). If the true sex ratio was 70:30 rather than 50:50, then 125 observations of embryos or offspring from treated spermatozoa and 125 observations for control spermatozoa would be needed to have a 95% chance of detecting a difference for that sample of processed semen. However, if the true sex ratio was 80:20, then only 50 observations would be needed. Most publications, however, contain far fewer than 100 observations on treated semen and 100 observations on contemporaneous control semen from the same males. Editorial Note This chapter was written in 1993, and provides much historical information regarding the use of sexed semen. Nonetheless, considerable progress has been made in this area over the past several years, especially with regard to commercial possibilities in the horse. The reader is encouraged to view Chapter 324 for a current overview of sexed semen technologies as they relate to equine breeding programs.
Acknowledgement Some individuals, and some manuscripts have wonderful insight and historical value. Such is the case with Dr. Rupert P. Amann and four chapters that he so ably produced for the first edition of Equine Reproduction. While Dr. Amann is now retired and has turned the baton over to others, these pieces of work have endured the test of time and continue to be an invaluable resource for the equine practitioner and academician. As such, we decided to reprint these chapters for you.
References 1. Amann RP. Maturation of spermatozoa. In: Proceedings of the 11th International Congress on Animal Reproduction and Artificial Insemination, vol. 5, 1988, pp. 320–8.
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2. Johnson L. Spermatogenesis. In: Cupps PT (ed.) Reproduction in Domestic Animals, 4th edn. New York: Academic Press, 1991; pp. 173–219. 3. Hochereau-de Reviers M-T, Courtens JL, Courot M, de Reviers M. Spermatogenesis in mammals and birds. In: Lamming GE (ed.) Marshall’s Physiology of Reproduction, vol. 2, 4th edn. London: Churchill Livingstone, 1990; pp. 106–82. 4. Eddy EM. The spermatozoon. In: Knobil F, Neill J (eds) The Physiology of Reproduction. New York: Raven Press, 1988; pp. 27–68. 5. Bedford JM, Hoskins DD. The mammalian spermatozoon: morphology, biochemistry and physiology. In: Lamming GE (ed.) Marshall’s Physiology of Reproduction, vol. 2, 4th edn. London: Churchill Livingstone, 1990; pp. 379–568. 6. Amann RP. Can the fertility potential of a semen sample be predicted accurately? J Androl 1989;10:89–98. 7. Amann RP, Pickett BW. Principals of cryopreservation and a review of cryopreservation of stallion spermatozoa. J Equine Vet Sci 1987;7:145–73. 8. Amann RP. Function of the epididymis in bulls and rams. J Reprod Fertil Suppl 1987;34:115–31. 9. Metz KW, Berger T, Clegg ED. Adsorption of seminal plasma proteins by boar spermatozoa. Theriogenology 1990;34:691–700. 10. Jasko DJ, Moran DM, Farlin ME, Squires EL. Effect of seminal plasma dilution or removal on spermatozoal motion characteristics of cooled stallion semen. Theriogenology 1991;35:1059–67. 11. Saacke RG, Marshall CE, Vinson WE, O’Connor ML, Chandler JE, Mullins KJ, Amann RP, Wallace RA, Vincel WN, Kellgren HC. Semen quality and heterospermic insemination in cattle. In: Proceedings of the 9th International Congress on Animal Reproduction and Artificial Insemination, vol. 5, 1980, pp. 75–8. 12. Small MF. Sperm wars. Discover 1991; 48–53. 13. Barth AD, Oko RJ. Abnormal Morphology of Bovine Spermatozoa. Ames: Iowa State University Press, 1989. 14. Garner DL. Artificial insemination. In: Cupps PT (ed.) Reproduction in Domestic Animals, 4th edn. New York: Academic Press, 1991; pp. 251–78. 15. Setchell BP. Male reproductive organs and semen. In: Cupps PT (ed.) Reproduction in Domestic Animals, 4th edn. New York: Academic Press, 1991; pp. 221–49. 16. Johnson L, Amann RP, Pickett BW. Maturation of equine epididymal spermatozoa. Am J Vet Res 1980;41:1190–6. 17. Baumgartl C. Licht- und elektronenmikroskopische Untersuchungen Uber Veranderungen der Plasmamembran und Akrosomstruktur von Pferdespermien. DMV dissertation, Tier`arztliche Hochschule Hannover, 1980. 18. Hammerstedt RH, Graham JK, Nolan JP. Cryopreservation of mammalian sperm: what we ask them to survive. J Androl 1990;11:73–88. 19. Hammerstedt RH, Graham JK. Cryopreservation of poultry sperm: the enigma of glycerol. Cryobiology 1992;29:26–38. 20. Watson PF. Artificial insemination and the preservation of semen. In: Lamming GE (ed.) Marshall’s Physiology of Reproduction, vol. 2, 4th edn. London: Churchill Livingstone, 1990; pp. 747–869. 21. Parks JE, Lynch DV. Lipid composition and thermotropic phase behavior of boar, bull stallion and rooster sperm membranes. Cryobiology 1992;29:255–66. 22. Baumgartl C, Bader H, Drommer W, Liming I. Ultrastructural alterations of stallion spermatozoa due to se-
23.
24.
25.
26. 27.
28.
29.
30.
31.
32. 33.
34.
35.
36.
37.
38.
39.
40.
41.
men conservation. In: Proceedings of the 9th International Congress on Animal Reproduction and Artificial Insemination, vol. 5, 1980, pp. 134–7. Pickett BW, Squires EL, McKinnon AO. Procedures for Collection, Evaluation and Utilization of Stallion Semen for Artificial Insemination. Animal Reproduction Laboratory Bulletin No. 03. Fort Collins, Colorado State University, 1987. Mann T. Metabolism of semen: fructolysis, respiration and sperm energetics. In: Mann T (ed.) The Biochemistry of Semen and of the Male Reproductive Tract. New York: Barnes and Noble, 1964, pp. 265–307. Polakoski KL, Kopta M. Seminal plasma. In: Zaneveld LJD, Chatterton RT (eds) Biochemistry of Mammalian Reproduction. New York: John Wiley & Sons, 1982, pp. 89–118. Mann, T. Biochemistry of stallion semen. J Reprod Fertil Suppl 1975;23:47–52. Pickett BW, Faulkner LC, Voss JL. Effect of season on some characteristics of stallion semen. J Reprod Fertil Suppl 1975;23:25–8. Tischner M, Kosiniak K, Bielanski W. Analysis of the pattern of ejaculation in stallions. J Reprod Fertil 1974;41:329–35. Komarek RJ, Pickett BW, Gibson EW, Lanz RN. Composition of lipids in stallion semen. J Reprod Fertil 1965;10:337–42. Parks JE, Anon JW, Foote RH. Lipids of plasma membrane and outer acrosomal membrane from bovine spermatozoa. Biol Reprod 1987;37:1249–58. Parks JE, Hammerstedt RH. Developmental changes occurring in the lipids of ram epididymal spermatozoa plasma membrane. Biol Reprod 1985;32:653–68. Koehler JK. Lectins as probes of the spermatozoon surface. Arch Androl 1981;6:197–217. Magargee SF, Kunze E, Hammerstedt RH. Changes in lectin-binding features of ram sperm surfaces associated with epididymal maturation and ejaculation. Biol Reprod 1988;38:667–85. Eddy EM, Vernon RB, Muller CH, Hahnel AC, Fenderson BA. Immunodissection of sperm surface modifications during epididymal maturation. Am J Anat 1985;174:225–37. Feuchter FA, Vernon RB, Eddy EM. Analysis of the sperm surface with monoclonal antibodies: topographically restricted antigens appearing in the epididymis. Biol Reprod 1981;24:1099–110. Chakraborty J, Constatinou A, McCorquodale M. Monoclonal antibodies to bull sperm surface antigens. Anim Reprod Sci 1985;9:101–9. Hunter AG, Nornes HO. Characterization and isolation of a sperm-coating antigen from rabbit seminal plasma with capacity to block fertilization. J Reprod Fertil 1969;20:419–27. Inskeep PB, Hammerstedt RH. Endogenous metabolism by sperm in response to altered cellular ATP requirements. J Cell Physiol 1985;123:180–90. Hiipakka RA, Hammerstedt RH. 2-deoxyglucose transport and phospho4ylation by bovine sperm. Biol Reprod 1978;19:368–79. Flipse RJ. Metabolism of bovine semen. XI. Factors affecting the transport of fructose in bovine spermatozoa. J Dairy Sci 1962;45:917–20. Peterson RN, Bundman D, Freund M. Binding of cytochalasin B to hexose transport sites in human spermatozoa and inhibition of binding by purines. Biol Reprod 1977;17:198–206.
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Spermatozoal Function 42. Hammerstedt RH, Lovrien RE. Calorimetric techniques for metabolic studies of cells and organisms under normal conditions and stress. J Exp Zool 1983;228:459–69. 43. Hammerstedt RH. Use of sperm cells as a model for the study of metabolism. In: Lennon DLF, Stratman FW, Zahlten RN (eds) Biochemistry of Metabolic Processes. New York: Elsevier, 1983, pp. 29–38. 44. Hammerstedt RH, Volonte C, Racker E. Motility, heat, and lactate production in ejaculated bovine sperm. Arch Biochem Biophys 1988;266:11–123. 45. Hoskins DD, Brandt H, Acott TS. Initiation of sperm motility in the mammalian epididymis. Fed Proc 1978;37:2534–42. 46. Vijayaraghavan S, Critchlow LM, Hoskins DD. Evidence for a role for cellular alkalization in the cyclic adenosine 3’,5’-monophosphate-mediated initiation of motility in bovine caput spermatozoa. Biol Reprod 1985;32: 489–500. 47. Wales RG, Murdoch RN. Factors influencing the response of ram spermatozoa to bicarbonate and carbon dioxide. Aust J Biol Sci 1971;24:345–54. 48. Peterson RN, Freund M. Effects of [H+ ], [N+ ], [K+ ] and certain membrane-active drugs on glycolysis, motility, and ATP synthesis by human spermatozoa. Biol Reprod 1973;8:350–7. 49. Peterson RN. The sperm tail and midpiece. In: Zaneveld LJD, Chatterton RT (eds) Biochemistry of Mammalian Reproduction. New York: John Wiley & Sons, 1982, pp.153– 73. 50. Setchell BP. Spermatogenesis and spermatozoa. In: Austin CR, Short RV (eds) Reproduction in Mammals. I. Germ Cells and Fertilization, 2nd edn. New York: Cambridge University Press, 1982, pp. 63–101. 51. Jones R, Mann T, Sherins R. Peroxidative breakdown of phospholipids in human spermatozoa, spermicidal properties of fatty acid peroxides, and protective action of seminal plasma. Fertil Steril 1979;31:531–7. 52. Holland MK, Storey BT. Oxygen metabolism of mammalian spermatozoa. Generation of hydrogen peroxide by rabbit epididymal spermatozoa. Biochem J 1981;198: 273–80. 53. Alvarez JG, Storey BT. Assessment of cell damage caused by spontaneous lipid peroxidation in rabbit spermatozoa. Biol Reprod 1984;30:323–31. 54. Alvarez JG, Storey BT. Lipid peroxidation and the reactions of superoxide and hydrogen peroxide in mouse spermatozoa. Biol Reprod 1984;30:833–41. 55. Aitken RJ, Clarkson JS. Significance of reactive oxygen species and antioxidants in defining the efficacy of sperm preparation techniques. J Androl 1987;9:367–76. 56. Taylor MJ. The meaning of pH at low temperatures. Cryoletters 1981;2:231–9. 57. Taylor MJ. Acid dissociation constants for some biological buffers in aqueous solutions containing dimethylsulfoxide at 25◦ C and –12◦ C. Cryoletters 1980;1:449–60. 58. Taylor MJ. Physico-chemical principles in low temperature. In: Grout BWW, Morris GJ (eds) The Effects of Low Temperatures on Biological Systems. Baltimore: Edward Arnold, 1987; pp. 3–71. 59. Sinowatz F, Friess AE. Localization of lectin receptors on bovine epididymal spermatozoa using a colloidal gold technique. Histochemistry 1983;79:335–44. 60. Friess AE, Sinowatz F. Con A- and WGA-binding sites on bovine spermatozoa: TEM of specimens in toto. Biol Cell 1984;50:279–84.
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61. Evenson DP, Darzynkiewicz Z, Melamed MR. Relation of mammalian sperm chromatin heterogeneity to fertility. Science 1980;210:1131–3. 62. Ballachey BE, Evenson DP, Saacke RG. The sperm chromatin structure assay relationship with alternate tests of semen quality and heterospermic performance of bulls. J Androl 1988;9:109–15. 63. Garner DL, Gledhill BL, Pinkel D, Lake S, Stephenson D, Van Dilla MA, Johnson LA. Quantification of the X- and Y-chromosome-bearing spermatozoa of domestic animals by flow cytometry. Biol Reprod 1983;28:312–21. 64. Pinkel D, Garner DL, Gledhill BL, Lake S, Stephenson D, Johnson LA. Flow cytometric determination of the proportions of X- and Y-chromosome-bearing sperm in samples of purportedly separated bull sperm. J Anim Sci 1985;60:1303–7. 65. Mayer DT, Squires CD, Bogart R, Oloufa MM. The technique for characterizing mammalian spermatozoa as dead or living by differential staining. J Anim Sci 1951;10: 226–35. 66. Swanson EW, Bearden HJ. An eosin-nigrosin stain for differentiating live and dead bovine spermatozoa. J Anim Sci 1951;10:981–7. 67. Hancock JL. A staining method for the study of temperature shock in semen. Nature 1951;167:323–4. 68. Jeyendran RS, Van der Ven HH, Perez-Pelaez M, Crabo BG, Zaneveld LJ. Development of an assay to assess the functional integrity of the human sperm membrane and its relationship to other semen characteristics. J Reprod Fertil 1984;33:113–18. 69. Graham EF, Schmehl MKL, Evensen BK. An overview of column separation of spermatozoa. In: Proceedings of the 7th National Association of Animal Breeders Technical Conference on Artificial Insemination and Reproduction, 1978, pp. 69–73. 70. Samper JC, Hellander JC, Crabo BG. Relation between the fertility of fresh and frozen stallion semen and semen quality. J Reprod Fertil Suppl 1991;44:107–14. 71. Bhattacharyya AK, Zaneveld LJD. The sperm head. In: Zaneveld LJD, Chatterton RT (eds) Biochemistry of Mammalian Reproduction. New York: John Wiley & Sons, 1982, pp. 119–52. 72. Leblond CP, Clermont Y. Definition of the stages of the cycle of the seminiferous epithelium in the rat. Ann N Y Acad Sci 1952;55:548–73. 73. Cross NL, Meizel S. Methods for evaluating the acrosomal status of mammalian sperm. Biol Reprod 1989;41: 635–41. 74. Cross NL, Overstreet JW. Glycoconjugates of the human sperm surface: Distribution and alterations that accompany capacitation in vitro. Gamete Res 1987;16:23–35. 75. Graham JK, Kunze E, Hammerstedt RH. Analysis of sperm cell viability, acrosomal integrity, and mitochondrial function using flow cytometry. Biol Reprod 1990;43:55–64. 76. Farlin M, Jasko DK, Graham JK, Squires EL. Assessment of Pisum sativum agglutinin in identifying acrosomal damage in stallion spermatozoa. Mol Reprod 1992;32:23–7. 77. Blach EL, et al. Use of a monoclonal antibody to evaluate integrity of the plasma membrane of stallion sperm. Gamete Res 1988;21:233–41. 78. Evenson DP, Darzynkiewicz Z, Melamed MR. Simultaneous measurement by flow cytometry of sperm cell viability and mitochondrial membrane potential related to cell motility. J Histochem Cytochem 1982;30:279–80.
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79. Auger J, Ronot X, Dadoune JP. Human sperm mitochondrial function related to motility: a flow image cytometric assessment. J Androl 1989;10:439–48. 80. Evenson DP, Ballachey BE. Flow cytometric evaluation of bull sperm chromatin structure, mitochondrial activity, viability and concentration. In: Proceedings of the 11th National Association of Animal Breeders Technical Conference on Artificial Insemination and Reproduction, 1986, p. 109. 81. Didion BA, Dobrinsky JR, Giles JR, Graves CN. Staining procedure to detect viability and the true acrosome reaction in spermatozoa of various species. Gamete Res 1989;22:51–7. 82. Satir P. How cilia move. Sci Am 1974;231:44–52. 83. Satir P. The generation of ciliary motion. J Protozool 1984;31:8–12. 84. Linck RW. Advances in the ultrastructural analysis of the sperm flagellar axoneme. In: Fawcett DW, Bedford JM (eds) The Spermatozoon. Baltimore: Urban and Schwarzenberg, 1979, pp. 99–115. 85. Rikmenspoel R. Movements and active moments of bull sperm flagella as a function of temperature and viscosity. J Exp Biol 1984;108:205–30. 86. Amann RP. Computerized evaluation of stallion spermatozoa. In: Proceedings of the 34th Annual Convention of the American Association of Equine Practitioners, 1988, 453–73. 87. Budworth PR, Amann RP, Chapman PL. Relationships between computerized measurements of motion of frozenthawed bull sperm and fertility. J Androl 1988;9:41–54. 88. Amann RP, Squires EL, Pickett BW. Effects of sample thickness and temperature on spermatozoal motion. In: Proceedings of the 11th International Congress on Animal Reproduction and Artificial Insemination, vol. 3, 1988, pp. 221a–221c. 89. Amann RP. Relationship between computerized evaluations of spermatozoal motion and competitive fertility index. In: Proceedings of the 12th National Association of Animal Breeders Technical Conference on Animal Reproduction and Artificial Insemination, 1988, pp. 38– 44. 90. Boyers SP, Davis RO, Katz DF. Automated semen analysis. Curr Probl Obstet Gynecol Fertil 1989;12:173–99. 91. Varner DD, Vaughan SD, Johnson L. Use of a computerized system for evaluation of equine spermatozoal motility. Am J Vet Res 1991;52:224–30. 92. Jasko DK, Lein DH, Foote RH. The repeatability and effect of season on seminal characteristics and computeraided sperm analysis in the stallion. Theriogenology 1991;35:317–27. 93. Amann RP, Hammerstedt RH. Validation of a system for computerized measurements of spermatozoal velocity and percentage of motile sperm. Biol Reprod 1980;23:647–56. 94. Makler A. The thickness of microscopically examined seminal sample and its relationship to sperm motility estimation. Int J Androl 1978;1:213–19. 95. Jasko DJ, Lein DH, Foote RH. A comparison of two computer-automated semen analysis instruments for the evaluation of sperm motion characteristics in the stallion. J Androl 1990;11:453–9. 96. Blach EL, Amann RP, Bowen RA, Frantz D. Changes in quality of stallion spermatozoa during cryopreservation: plasma membrane integrity and motion characteristics. Theriogenology 1989;31:283–98.
97. Robertson L, Wolf DP, Tash JS. Temporal changes in motility parameters related to acrosomal status: identification and characterization of populations of hyperactivated human sperm. Biol Reprod 1988;39:797–805. 98. Kayser J-P, Amann RP, Shideler RK, Squires EL, Jasko DJ, Pickett BW. Effects of linear cooling rate on motion characteristics of stallion spermatozoa. Theriogenology. 1992;38:601–14. 99. Watson PF. The effects of cold shock on sperm cell membranes. In: Morris GJ, Clarke A (eds) Effects of Low Temperatures on Biological Membranes. London: Academic Press, 1981, pp. 189–218. 100. Singer SJ, Nicholson GL. The fluid mosaic model of the structure of cell membranes. Science 1972;175:720–31. 101. Houslay MD, Stanley KK (eds). Dynamics of Biological Membranes. New York: John Wiley & Sons, 1982. 102. Aloia RC, Curtin CC, Gordon LM (eds). Lipid Domains and the Relationship to Membrane Function. New York: Liss, 1988. 103. Cullis PR, Hope MJ. Physical properties and functional roles of lipids in membranes. In: Vance DE, Vance JE (eds) Biochemistry of Lipids and Membranes. Menlo Park: Benjamin/Cummings, 1985, pp. 25–72. 104. Ohki S, et al. (eds). Molecular Mechanisms of Membrane Fusion. New York: Plenum, 1988. 105. Hammerstedt RH, Parks JE, Doyle D, Flanagan TD, Hui SW, Mayhew E. Changes in sperm surfaces associated with epididymal transit. J Reprod Fertil Suppl 1987;34:133–49. 106. Quinn PJ. Principles of membrane stability and phase behavior under extreme conditions. J Bioenerg Biomembr 1989;21:3–19. 107. Watson PF, Morris GJ. Cold shock injury in animal cells. In: Bowler K, Fuller BJ (eds) Temperature and Animal Cells. Cambridge: Company of Biologists, 1987, pp.311–40. 108. Hammerstedt RH, Hay SR. Effect of incubation temperature on motility and cAMP content of bovine sperm. Arch Biochem Biophys 1980;199:427–37. 109. Ellory JC, Hall AC. Temperature effects on red cell membrane transport processes. In: Bowler K, Fuller BJ (eds) Temperature and Animal Cells. Cambridge: Company of Biologists, 1987, pp. 53–66. 110. Morris GJ, Clarke A. Cells at low temperatures. In: Grout BWW, Morris GJ (eds) The Effects of Low Temperatures on Biological Systems. Baltimore: Edward Arnold, 1987; pp. 72–119. 111. Morris GJ. Direct chilling injury. In: Grout BWW, Morris GJ (eds) The Effects of Low Temperatures on Biological Systems. Baltimore: Edward Arnold, 1987; pp. 120–46. 112. Watson PF, Plummer JM, Allen WE. Quantitative assessment of membrane damage in cold-shocked spermatozoa of stallions. J Reprod Fertil Suppl 1987;35:651–3. 113. Douglas-Hamilton DH, Osol R, Osol G, Driscoll D, Noble H. A field study of the fertility of transported equine semen. Theriogenology 1984;22:291–303. 114. Province CA, Squires EL, Pickett BW, Amann RP. Cooling rates, storage temperatures and fertility of extended equine spermatozoa. Theriogenology 1985;23:925–33. 115. Francl AT, Amarm RP, Squires EL, Pickett BW. Motility and fertility of equine spermatozoa in a milk extender after 12 or 24 hours at 20◦ C. Theriogenology 1987;27:517–25. 116. Varner DD, Blanchard TL, Meyers PJ, Meyers SA. Fertilizing capacity of equine spermatozoa stored for 24 hours at 5◦ or 20◦ C. Theriogenology 1989;32:515–25.
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Spermatozoal Function 117. Pursel VG, Johnson LA, Schulman LL. Effect of dilution, seminal plasma and incubation period on cold sock susceptibility of boar spermatozoa. J Anim Sci 1973;37: 528–31. 118. Berger T, Clegg ED. Effect of male accessory gland secretions on sensitivity of porcine sperm acrosomes to cold shock, initiation of motility and loss of cytoplasmic droplets. J Anim Sci 1985;60:1295–302. 119. Phillips PH, Lardy HA. A yolk-buffer pablum for the preservation of bull sperm. J Dairy Sci 1940;23: 399–404. 120. Blackshaw AW. The prevention of temperature shock of bull and ram semen. Aust J Biol Sci 1954;7:573–82. 121. Nishikawa Y, Waide Y, Shinomiya S. Studies on deep freezing of horse spermatozoa. In: Proceedings of the 6th International Congress on Animal Reproduction and Artificial Insemination, vol. 2, 1968, pp. 1589–96. 122. Oshida H, et al. Fertility of frozen stallion semen and some factors affecting to it. In: Proceedings of the 6th International Congress on Animal Reproduction and Artificial Insemination, vol. 2, 1968, pp. 1597–9. 123. Pickett BW, Burwash LD, Voss JL, Back DG. Effect of seminal extenders on equine fertility. J Anim Sci 1975;40:1136–43. 124. Pursel VG, Johnson LA, Schulman LL. Interactions of extender composition and incubation period on cold shock susceptibility of boar spermatozoa. J Anim Sci 1972;35:580–4. 125. Kampschmidt RF, Mayer DT, Herman HA. Lipid and lipoprotein constituents of egg yolk in the resistance and storage of bull spermatozoa. J Dairy Sci 1953;36:733– 42. 126. Masuda H, Nishikawa Y. Studies on the substances in egg yolk effective on viability and metabolism of spermatozoa. III. Substances in the non-dialyzable portion of egg yolk effective on the survival of spermatozoa of goats, bull and horses. Jpn J Zootech Sci 1972;43:355–9. 127. Pace MM, Graham EF. Components of egg yolk which protect bovine spermatozoa during freezing. J Anim Sci 1974;39:1144–9. 128. Watson PF. The protection of ram and bull spermatozoa by the low density lipoprotein fraction of egg yolk during storage at 5◦ C and deep-freezing. J Thermal Biol 1976;1:137–41. 129. Foulkes JA. The separation of lipoproteins from egg yolk and their effect on the motility and integrity of bovine spermatozoa. J Reprod Fertil 1977;49:277–84. 130. Quinn PJ, Chow PYW, White IG. Evidence that phospholipid protects spermatozoa from cold shock at the plasma membrane site. J Reprod Fertil 1980;60:403–7. 131. Parks JE, Meacham TN, Saacke RG. Cholesterol and phospholipids of bovine spermatozoa. II. Effect of liposomes prepared from egg phosphatidylcholine and cholesterol on sperm cholesterol, phospholipids and viability at 4◦ C and 37◦ C. Biol Reprod 1981;24:399–404. 132. Evans RW, Setchell BP. Association of exogenous phospholipids with spermatozoa. J Reprod Fertil 1978;53: 357–62. 133. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: techniques and preliminary findings. In: Proceedings of the 21st Annual Convention of the American Association of Equine Practitioners, 1975, pp. 327–36. 134. Hammerstedt RH, Amann RP, Rucinsky T, Morse PD Jr, Lepcock J, Snipes W, Keith AD. Use of spin labels
135.
136.
137.
138.
139.
140.
141. 142.
143. 144. 145.
146.
147.
148.
149.
150.
151.
152.
153.
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and electron spin resonance spectroscopy to characterize membranes of bovine sperm: effect of butylated hydroxytoluene and cold shock. Biol Reprod 1976; 14:381–97. Pursel VG. Effect of cold shock on boar sperm treated with butylated hydroxytoluene. Biol Reprod 1979;21: 319–24. Watson PF, Anderson WJ. Influence of butylated hydroxytoluene (BHT) on the viability of ram spermatozoa undergoing cold shock. J Reprod Fertil 1983;68:229–35. Graham JK, Hammerstedt RH. Differential effects of butylated hydroxytoluene (BHT) analogs on bull sperm subjected to cold-induced membrane stress. Cryobiology 1992;29:106–7. Butler WJ, Roberts TK. Effects of some phosphatidyl compounds on boar spermatozoa following cold shock or slow cooling. J Reprod Fertil 1975;43:183–7. Graham JK, Foote RH. Effect of several lipids, fatty acyl chain length, and degree of unsaturation on the motility of bull spermatozoa after cold shock and freezing. Cryobiology 1987;24:42–52. Yanagimachi R. Capacitation and the acrosome reaction. In: Asch RH, Balmaceda JP, Johnston I (eds) Gamete Physiology. Norwell, MA: Serono Symposia, 1990; pp. 31–42. Noyes WR. The fertilizing capacity of spermatozoa. West J Surg Obstet Gynecol 1953;61:342–9. Austin CR. Observations on the penetration of the sperm into the mammalian egg. Aust J Sci Res Ser B 1951;4: 581–96. Chang MC. Fertilizing capacity of spermatozoa deposited into the fallopian tubes. Nature 1951;168:697–8. Austin CR. The ‘capacitation’ of the mammalian sperm. Nature 1952;170:326. Bedford JM. Significance of the need for sperm capacitation before fertilization in eutherian mammals. Biol Reprod 1983;28:108–20. Brackett BG, Cofone MA, Boice ML, Bousquet D. Use of zona-free hamster ova to assess sperm fertilizing ability of bull and stallion. Gamete Res 1982;5:217–27. Fraser LR. Sperm capacitation and its modulation. In: Bavister BD, Cummins J, Roldan ERS (eds) Fertilization in Mammals. Norwell, MA: Serono Symposia, 1990; pp. 141–54. Rogers BJ, Brentwood BJ. Capacitation, acrosome reaction and fertilization. In: Zaneveld LJD, Chatterton RT (eds) Biochemistry of Mammalian Reproduction. New York: John Wiley & Sons, 1982, pp. 203–30. Clegg ED. Mechanisms of mammalian sperm capacitation. In: Hartmann JF (ed.) Mechanism and Control of Animal Fertilization. New York: Academic Press, 1983; pp. 177–212. Langlais J, Roberts KD. A molecular membrane model of sperm capacitation and the acrosome reaction of mammalian spermatozoa. Gamete Res 1985;12:183–224. Oliphant G, Reynolds AB, Thomas TS. Sperm surface components involved in the control of the acrosome reaction. Am J Anat 1985;174:269–83. Yanagimachi R. Mechanisms of fertilization in mammals. In: Mastroiarmi L, Biggers JD (eds) Fertilization and Embryonic Development In Vitro. New York: Plenum Press, 1981; pp. 81–182. Parks JE, Ehrenwald E. Cholesterol efflux from mammalian sperm and its potential role in capacitation. In: Bavister BD, Cummins J, Roldan ERS (eds) Fertilization in Mammals. Norwell, MA: Serono Symposia, 1990; pp. 155–64.
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154. Suarez SS, Drost M, Redfern K, Gottlieb W. Sperm motility in the oviduct. In: Bavister BD, Cummins J, Roldan ERS (eds) Fertilization in Mammals. Norwell, MA: Serono Symposia, 1990; pp. 111–24. 155. Overstreet JW, VandeVoort CA. Sperm transport in the female genital tract. In: Asch RH, Balmaceda JP, Johnston I (eds) Gamete Physiology. Norwell, MA: Serono Symposia, 1990; pp. 43–52. 156. Fraser LR, Ahuja KK. Metabolic and surface events in fertilization. Gamete Res 1988;20:491–519. 157. Roche JF, Dziuk PJ, Lodge JR. Competition between fresh and aged spermatozoa in fertilizing rabbit eggs. J Reprod Fertil 1968;16:155–7. 158. Tesh JM. Effects of ageing of rabbit spermatozoa in utero on fertilization and prenatal development. J Reprod Fertil 1969;20:299–306. 159. Mann T, Lutwak-Mann C. Biochemical aspects of aging in spermatozoa in relation to motility and fertilizing ability. In: Blandau RJ (ed.) Aging Gametes: Their Biology and Pathology. New York: S. Karger, 1975, pp. 122–50. 160. Hughes JP, Loy RG. Artificial insemination in the equine: a comparison of natural breeding and artificial insemination from six stallions. Cornell Vet 1970;60:463–75. 161. Salisbury GW, Flerchinger FH. In vitro aging of spermatozoa and evidence for embryonic or early fetal mortality in cattle. In: Proceedings of the 4th International Congress on Animal Reproduction and Artificial Insemination, vol. 2, 1961, pp. 601–6. 162. Salisbury GW. Fertilizing ability and biological aspects of sperm storage in vitro. In: Proceedings of the 6th International Congress on Animal Reproduction and Artificial Insemination, vol. 2, 1968, pp. 1189–204. 163. Nicolai P, Shaver EL. The chromosome complement of rabbit blastocysts resulting from spermatozoa stored at 5◦ C. Biol Reprod 1977;17:640–4. 164. Maurer RR, Stranzinger GF, Paufier SK. Embryonic development in rabbits after insemination with spermatozoa stored at 37, 5 or –196◦ C for various periods. J Reprod Fertil 1976;48:43–9. 165. Bell CL, Shaver EL. Analysis of preimplantation golden hamster conceptuses resulting from spermatozoa aged in utero. Gamete Res 1982;6:199–207. 166. Hart RG, Salisbury GW. The effect of sperm age on embryonic mortality in the frog. Fed Proc 1967;26: 645. 167. Rowson LEA. Prolonged storage of gametes in relation to fertility and progeny characteristics in farm animals. In: Blandau RJ (ed.) Aging Gametes: Their Biology and Pathology. New York: S. Karger, 1975, pp. 249–64. 168. Kenney RM, Kent MG, Garcia MC, Hurtgen JP. The use of DNA index and karyotype analyses as adjuncts to the estimation of fertility in stallions. J Reprod Fertil Suppl 1991;44:69–75. 169. Flipse R.J. Metabolism of bovine semen. IX. Glutamicoxaloacetic and glutamic-pyruvic transaminase activities. J Dairy Sci 1960;43:773–6. 170. Graham EF, Pace MM. Some biochemical changes in spermatozoa due to freezing. Cryobiology 1967;4:75–84.
171. Pace MM, Graham EF. The release of glutamic oxaloacetic transaminase from bovine spermatozoa as a test method of assessing semen quality and fertility. Biol Reprod 1970;3:140–6. 172. Jones R, Mann T. Damage to ram spermatozoa by peroxidation of endogenous phospholipids. J Reprod Fertil 1977;50:261–8. 173. Calamera JC, Giovenco P, Quiros MC, Brugo S, Dondero F, Nicholson RF. Effect of lipid peroxidation upon human spermatic adenosine triphosphate (ATP). Relationship with motility, velocity and linearity of the spermatozoa. Andrologia 1989;1:48–54. 174. Killian G, Honadel T, McNutt T, Henault M, Wegner C, Dunlap D. Evaluation of butylated hydroxytoluene as a cryopreservative added to whole or skim milk diluent for bull semen. J Dairy Sci 1989;72:1291–5. 175. Morrell JM, Keeler KD, Noakes DE, Mackenzie NM, Dresser DW. Sexing of sperm by flow cytometry. Vet Rec 1988;122:322–4. 176. Johnson LA, Clarke RN. Flow sorting of X and Y chromosome-bearing mammalian sperm: activation and pronuclear development of sorted bull, boar, and ram sperm microinjected into hamster oocytes. Gamete Res 1988;21:335–43. 177. Johnson LA, Flook JP, Hawk HW. Sex pre-selection in rabbits: Live births from X and Y sperm separated by DNA and cell sorting. Biol Reprod 1989;41:199–203. 178. Johnson LA. A flow cytometric/sorting method for sexing mammalian sperm validated by DNA analysis and live births. Cytometry 1990;4(Suppl.):42. 179. Amann RP. Treatment of sperm to predetermine sex. In: Proceedings of the National Association of Animal Breeders Technical Conference, 1988, pp. 127–35. 180. Amann RP. Treatment of sperm to predetermine sex. Theriogenology 1989;31:49–60. 181. Jost A, Magre S. Testicular development phases and dual hormonal control of sexual organogenesis. In: Serb M Motta M, Zarisi M, Martini L (eds) Sexual Differentiation: Basic and Clinical Aspects. New York: Raven Press, 1984, pp. 11–15. 182. Berta P, Hawkins JR, Sinclair AH, Taylor A, Griffiths BL, Goodfellow PN, Fellous M. Genetic evidence equating SRY and the testis-determining factor. Nature 1990;348:448–51. 183. Jager RJ, Anvret M, Hall K, Scherer G. A human XY female with a frame shift mutation in the candidate testisdetermining gene SRY. Nature 1990;348:452–4. 184. McLaren A. The making of male mice. Nature 1991;351:96. 185. Koopman P, Gubbay J, Vivian N, Goodfellow P, LovellBadge R. Male development of chromosomally female mice transgenic for Sry. Nature 1991;241:117–21. ¨ 186. Blottner S, Bottcher M, Schwerin M, Rommel P. Preconceptional sex determination by in vitro fertilization in cattle. In: Proceedings of the 7th Annual Meeting European Embryo Transfer Association, 1991, p. 124. 187. Bradley MP. Immunological sexing of mammalian semen: Current status and future options. J Dairy Sci 1989;72:3372–80.
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Chapter 103
Sperm-Oviduct Interactions Barry A. Ball
Introduction Although billions of spermatozoa are deposited into the mare’s uterus at the time of natural mating, only relatively few sperm reach the site of fertilization at the ampullar–isthmic junction of the oviduct.1 This marked attrition of sperm during adovarian movement through the mare’s reproductive tract serves to reduce the number of sperm present at the site of fertilization and thereby may reduce the possibility of polyspermic fertilization. In addition, there appears to be a strong selection of motile, morphologically normal sperm with progression through the reproductive tract which also enhances the prospects of normal fertilization and embryonic development.2, 3 A possible adaptation to the long estrus of mares is the ability of equine sperm to survive for a relatively long period of time in the mare’s reproductive tract with intervals between mating and ovulation that are routinely 2–3 days and upwards of 6–7 days in some reports.4 The oviduct of the mare appears to be critical in the reduction of sperm numbers reaching the site of fertilization, the enrichment of the fertilizing population of sperm, as well as the relatively long survival of fertile sperm in the mare’s reproductive tract. This chapter will focus upon the formation of this ‘oviductal sperm reservoir’ and its impact on fertility in the horse, as well as its relationship to assisted reproductive technologies in the horse.
Characterization of the oviductal sperm reservoir With natural mating, stallion semen is generally deposited directly through the cervix into the uterus. Although studies have not addressed the immediate transit of sperm to the uterotubal junction in the horse, data from other species suggest that there is a rapid (within minutes) transit of sperm to the
oviduct, which is due primarily to uterine contractility, followed by a more sustained (hours) transit of sperm, which is probably responsible for establishing the oviductal sperm reservoir.5 In the mare, approximately 4 hours is required from the time of insemination until an adequate sperm reservoir is established in the oviduct to maintain normal fertilization.6 Beyond this time, in the normal mare uterine lavage to remove sperm does not adversely affect subsequent pregnancy rates, implying that the sperm reservoir is safely established in the oviduct by this point. The uterotubal junction (UTJ) in the mare is characterized by an extremely narrow lumen lined with multiple longitudinal folds of mucosa which jut into the uterine lumen (Figure 103.1). Changes in the amount of edema present at the UTJ, along with the presence of mucus and the narrow lumen at this site, are thought to restrict the passage of sperm into the oviduct, thereby helping to reduce the number of sperm reaching the site of fertilization.3, 7 Although not definitively established for the mare, it appears that the oviductal isthmus (region closest to the UTJ) and possibly the UTJ are the anatomical sites for sperm storage in the mare.3, 7 The isthmus of the mare’s oviduct is lined by both ciliated and non-ciliated epithelial cells (Figure 103.2) which are thrown into many longitudinal folds.8 Formation of the oviductal sperm reservoir involves adhesion of sperm to both ciliated and non-ciliated epithelial cells (Figure 103.3) lining the oviductal isthmus with the sperm attachment occurring through the rostral portion of the sperm head.3, 7 Sperm binding to the oviductal epithelium appears to be a critical event in the formation of the sperm reservoir, and sperm remain viable and fertile in this protected location for an extended period of time prior to ovulation and release of the oocyte. Current evidence suggests that the cell–cell interaction between equine spermatozoa and oviductal epithelium modifies both sperm and oviductal epithelial
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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(c) Figure 103.1 The uterotubal junction in the mare. (a) Hysteroscopic view of the uterotubal papilla (arrow) which juts into the uterine lumen. (b) Photomicrograph of the uterotubal junction showing complex folds (courtesy of Dr. A.C. Enders, University of California, Davis). (c) Photomicrograph showing a cross-section of the oviductal isthmus. The narrow lumen is surrounded by a thick muscular coat.
cell (OEC) function. The interaction with OECs preferentially selects morphologically normal sperm with intact plasma and acrosomal membranes – a process that may be important for enriching the population of sperm that will ultimately participate in fertilization.2, 3 Several lines of evidence suggest that adhesion of spermatozoa prevents capacitation and prolongs sperm viability until near the time of ovulation.9–11 Adhesion of equine sperm to OECs acts to prevent a premature rise in intracellular calcium concentration [Ca2+ ]i in sperm, thereby preventing capacitation. Release of sperm from oviductal epithelium is associated with a rise in [Ca2+ ]i , capacitation, and the ability to bind the zona pellucida and initiate acrosomal exocytosis.12 The effect of adhesion on maintenance of low [Ca2+ ]i and prevention of capacitation appears to be tissue specific and does not depend upon secreted factors from the oviductal ep-
ithelium. Although the mechanism for maintenance of low [Ca2+ ]i as a function of cell adhesion is unknown in this system, it is likely related to activation of a calcium-ATPase in spermatozoa or a suppression of tyrosine phosphorylation.13, 14 Adhesion of sperm to oviductal epithelium also alters oviductal cell function. Sperm adhesion to oviductal cells increased [Ca2+ ]i in oviductal cells,15 and altered the synthesis and secretion of a number of oviductal proteins.16 Although the identity of these proteins has not been determined, some of these proteins associate with the sperm membrane and likely alter sperm function.17 Although an estrus-associated oviductal glycoprotein has not been identified in the horse,18 work in other mammalian species suggests that oviductal glycoproteins associate with spermatozoa and alter sperm adhesion to the extracellular matrix known as the zona pellucida.19 Together, these studies indicate that there is a dynamic interaction
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Sperm-Oviduct Interactions
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Figure 103.2 Scanning electron micrographs of the isthmic oviductal mucosa. The oviductal mucosa is thrown into a series of folds (a) which are covered by a ciliated epithelium (b). The oviductal epithelium includes both ciliated (black arrow) epithelial cells and epithelial cells covered by microvilli (white arrow).
between spermatozoa and OECs that alters the function of both cell types as part of events preceding fertilization.
Molecular basis for sperm–oviductal interaction
Figure 103.3 Scanning electron micrograph of equine sperm adherent to the cilia of oviductal epithelial cells.
The molecular basis for adhesion of sperm to oviductal epithelium appears to be mediated, at least in part, by a calcium-dependent lectin. The carbohydrate moiety recognized by the sperm lectin varies with species and has been characterized as galactosyl or fucosyl residues for equine or bovine sperm, respectively.20, 21 Galactosyl residues appear to be among the most abundant present in the isthmic oviduct of the mare, based upon lectin histochemistry (Figure 103.4), and may represent the ligand for sperm adhesion in the equine oviduct.22 The expression of these galactosyl residues varied with cycle stage in the mare, and expression of galactosyl glycoconjugates was greater in the isthmus than in the ampulla of the equine oviduct.22 Therefore, putative sperm binding sites were more readily available in the isthmic oviduct during estrus, a finding that is consistent with enhanced sperm adhesion to isthmic oviductal explants during estrus.23 Consistent with
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Figure 103.4 Fluorescent micrograph showing the isthmic oviduct of a mare stained with the lectin Dolichos biflorus (DBA), which binds galactosyl residues. Intense staining for galactosyl residues are present on the luminal epithelium and represent putative ligands for equine sperm adhesion in the oviductal isthmus.
this observation, both galactose and the glycoprotein asialofetuin, which expresses terminal galactose residues, inhibit adhesion of equine sperm to OECs.20 Galactose-binding proteins are present on equine spermatozoa and represent possible candidates for sperm adhesion to OECs.24 This sperm-specific, 25kDa, galactose-binding protein is localized over the rostral sperm head and is redistributed to the equatorial region during sperm capacitation. Further work is required to determine whether this galactosebinding protein is involved in equine sperm adhesion to the oviductal epithelium. Bovine sperm adhesion to oviductal epithelium is at least partially mediated via seminal plasma proteins, including bovine seminal plasma protein (BSPA1/-A2; also known as PDC-109), which are secretory proteins of the vesicular gland,25 as well as BSP-A3 and BSP-30-kDa.26 These heparin-binding proteins coat sperm at the time of ejaculation and anchor to choline phospholipids of the sperm head. In turn, these BSPs appear to recognize fucosyl residues expressed on annexins present on oviductal epithelial cells during the process of sperm adhesion.27 Epididymal sperm lack these BSPs and, as a consequence, have a significantly reduced adhesion to OECs; however, the addition of purified BSPs enhances the ability of epididymal sperm to bind OECs.26 During heparin-induced capacitation and release of bovine sperm from OECs, PDC-109 is lost from the sperm head.26 Once capacitated, bovine sperm lose the ability to bind to OECs; however, the reintroduction of PDC-109 to capacitated sperm facilitates their adhesion to OECs. In pigs, it appears that a different family of seminal plasma proteins, the spermadhesins, are responsible for adhesion of boar sperm to OECs.28 The spermadhesin AQN1 is a carbohydrate binding protein which
appears to bind to OECs via a mannose-binding site. Therefore, two different classes of seminal plasma proteins appear to be responsible for sperm adhesion to OECs in ruminants and swine. In the horse, seminal plasma proteins structurally related to both BSP and spermadhesins have been characterized.29 The horse seminal plasma proteins (SP1 and SP2) are heparin- and phosphorylcholinebinding proteins that constitute the major proteins of equine seminal plasma and are homologous to the BSPs.30 Horse SP1 and SP2 are produced primarily in the ampulla of the vas deferens rather than in the vesicular glands, as in cattle. Likewise, one member of the spermadhesin family is present in equine seminal plasma (HSP7) and may play a role in sperm–zona pellucida interaction in the horse.30, 31 A third major group of seminal plasma proteins in the horse are the cysteine-rich secretory proteins (CRISPs).29 Although no research has examined the specific role of any of these proteins in sperm–OEC adhesion in the horse, it seems likely that one or more of these seminal plasma proteins may play a role in sperm adhesion to OECs as part of establishing the oviductal sperm reservoir in mares based upon information available in cattle and swine.
Sperm release from the oviductal sperm reservoir Release of sperm from the oviductal sperm reservoir appears to be closely tied to maturation and ovulation of the oocyte in most species, and sperm which are released appear to undergo capacitation in preparation for fertilization of the oocyte.32 This carefully orchestrated process of ovulation of the metaphase II oocyte along with the release of capacitated sperm from the oviductal sperm reservoir helps to ensure that fertile gametes are available at the site of fertilization. This timing is particularly important because the fertile lifespan of both the ovulated ovum and the capacitated spermatozoon is limited. A number of mechanisms have been proposed for the release of sperm from the oviductal reservoir and subsequent capacitation. In cattle, heparinbinding sites on PDC-109 are thought to play a role in the loss of the adsorbed seminal plasma protein from the sperm with subsequent release from the oviductal epithelium.26, 32 Glycosaminoglycans are well-characterized components of equine follicular fluid,33 and are able to induce capacitation of equine sperm in vitro.34 In the horse, a combination of heparin and estradiol increase release of sperm from OECs in vitro, suggesting that follicular fluid entering the oviduct might affect sperm release. However, the role of oviductal fluid components in
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Sperm-Oviduct Interactions eliciting sperm release from the OECs in vivo has been debated in other species.32
Clinical implications Establishment of the oviductal sperm reservoir appears to be an important process relative to normal fertilization in horses and thereby has important implications in fertility. Subfertile stallions have a reduced ability to establish a reservoir of sperm in the oviduct based upon the reduced numbers of sperm found in this site after insemination compared with fertile stallions.35 Under in vitro conditions, sperm from different stallions demonstrated differences in ability to bind OECs,36 and, in cattle and swine, the ability of sperm to bind OECs in vitro has been related to fertility.37–41 Although studies comparing the ability of sperm to bind OECs in vitro and fertility have not been published for the stallion, existing information suggests that this might be a useful assay to consider for the horse. Cryopreservation of equine sperm also reduced their ability to bind OECs compared with fresh sperm despite adjustment for equal numbers of motile or membrane-intact sperm between treatments.36 In swine, fewer frozen–thawed than fresh sperm were recovered from the oviduct at 8 hours after insemination of comparable numbers of live sperm cells.40, 41 These studies suggest that frozen–thawed sperm may be less capable of establishing a sperm reservoir, and may account, at least in part, for the reduction in fertility observed with frozen semen as well as the need to inseminate mares closer to ovulation with frozen–thawed sperm. As noted earlier, the seminal plasma protein CRISP3 has been postulated to play a role in the interaction of sperm with the female reproductive tract. Interestingly, CRISP3 expression on equine sperm is correlated with stallion fertility,42, 43 and a nonsynonymous single nucleotide polymorphism (SNP) detected in the CRISP3 gene was associated with variable fertility in stallions.42 Stallions that were heterozygous for the CRISP3 SNP (E208K) had a per cycle pregnancy rate that was approximately 7% less than homozygotes.42 Although the function of CRISP3 in male fertility and its putative role in interaction with the female reproductive tract have not been defined, further research is warranted to understand the importance of this seminal plasma protein. In addition to the effects of the stallion on establishing the oviductal sperm reservoir, Scott et al.35 reported that mares deemed susceptible to endometritis had fewer sperm in the caudal isthmus of the oviduct than did normal mares following insemi-
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nation with comparable numbers of sperm. These workers also described alterations in the luminal epithelium that were detected with scanning electron microscopy in susceptible mares. In contrast, Rigby et al.44 found no difference in the number of oviductal sperm recovered from reproductively normal mares compared with mares that exhibited delayed uterine clearance. Based upon these studies, it is unclear whether differences in sperm numbers in the oviduct of susceptible mares translate into a reduction in fertilization rates in these mares. Several strategies may be considered in situations where a reduction in stallion fertility could potentially be associated with a reduced longevity of sperm in the mare’s reproductive tract with an implied reduction in formation or function of the oviductal sperm reservoir. Practically speaking, reducing the interval between mating or insemination and ovulation by the careful monitoring of follicular development and the use of ovulatory induction agents has been used in attempts to improve fertility of some subfertile stallions. Likewise, some clinicians have advocated multiple matings or inseminations of mares bred to subfertile stallions in attempts to enhance the formation of the oviductal sperm reservoir. Some studies suggest that increasing the number of matings or inseminations prior to ovulation may increase conception rates per cycle in mares. In a field study of Thoroughbred mares, there was a significant increase in conception rates per cycle when mares were bred twice rather than once per cycle.45 Likewise, a large field study based upon insemination of mares with frozen–thawed equine sperm demonstrated a significant increase in pregnancy rates when mares were bred twice before ovulation rather than once,46 although another study reported no difference in fertility in mares bred once versus twice with cryopreserved sperm.47 Clearly a number of factors influence these studies, including the type of mating or insemination, individual stallion effects, timing of mating or insemination relative to ovulation, and mare fertility. Nonetheless, techniques such as shortening the interval from mating to ovulation or using multiple inseminations per estrus may be useful in managing some subfertile stallions.
References 1. Parker WG, Sullivan JJ, First NL. Sperm transport and distribution in the mare. J Reprod Fertil Suppl 1975;23: 63–6. 2. Thomas PGA, Ball BA, Miller PG, Brinsko SP, Southwood L. A subpopulation of morphologically normal, motile spermatozoa attach to equine oviduct epithelial cells in vitro. Biol Reprod 1994;51:303–9.
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3. Scott MA, Liu IKM, Overstreet JW, Enders AC. The structural morphology and epithelial association of spermatozoa at the uterotubal junction: A descriptive study of equine spermatozoa in situ using scanning electron microscopy. J Reprod Fertil Suppl 2003;56:415–21. 4. Day FT. Survival of spermatozoa in the genital tract of the mare. J Agric Sci 1942;32:108–11. 5. Scott MA. A glimpse at sperm function in vivo: sperm transport and epithelial interaction in the female reproductive tract. Anim Reprod Sci 2000;60–61:337–48. 6. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post insemination on pregnancy rate in mares. Theriogenology 1991;35:1111–19. 7. Boyle MS, Cran DG, Allen WR, Hunter RHF. Distribution of spermatozoa in the mare’s oviduct. J Reprod Fertil Suppl 1987;35:79–86. 8. Ball BA. Scanning electron microscopy of the equine oviduct and observations on ciliary currents in vitro at day 2 after ovulation. Theriogenology 1996;46:1305–11. 9. Ellington JE, Ball BA, Blue BJ, Wilker CE. Capacitationlike membrane changes and prolonged viability in vitro of equine spermatozoa cultured with uterine tube epithelial cells. Am J Vet Res 1993;54:1505–10. 10. Dobrinski I, Suarez SS, Ball BA. Intracellular calcium concentration in equine spermatozoa attached to oviductal epithelial cells in vitro. Biol Reprod 1996;54:783–8. 11. Dobrinski I, Smith TT, Suarez SS, Ball BA. Membrane contact with oviductal epithelium modulates the intracellular calcium concentration of equine spermatozoa in vitro. Biol Reprod 1997;56:861–9. 12. Ellington JE, Ball BA, Yang X. Binding of stallion spermatozoa to the equine zona pellucida after coculture with oviductal epithelial cells. J Reprod Fertil 1993;98:203–8. 13. Petrunkina AM, Friedrich J, Drommer W, Bicker G, Waberski D, Topfer-Petersen E. Kinetic characterization of the changes in protein tyrosine phosphorylation of membranes, cytosolic Ca2+ concentration and viability in boar sperm populations selected by binding to oviductal epithelial cells. Reproduction 2001;122:469–80. 14. Gualtieri R, Boni R, Tosti E, Zagami M, Talevi R. Intracellular calcium and protein tyrosine phosphorylation during the release of bovine sperm adhering to the fallopian tube epithelium in vitro. Reproduction 2005;129:51–60. 15. Ellington JE, Varner DD, Burghardt RC, Meyers-Wallen VN, Barhoumi R, Brinsko SP, Ball BA . Cell-to-cell communication of equine uterine tube (oviduct) cells as determined by anchored cell analysis and sorting in culture. Anim Reprod Sci 1993;30:313–24. 16. Thomas PGA, Ignotz GG, Ball BA, Currie WB. Effect of coculture with stallion spermatozoa on de novo protein synthesis and secretion by equine oviduct epithelial cells. Am J Vet Res 1995;56:1657–62. 17. Ellington JE, Ignotz GG, Varner DD, Marcucio RS, Mathison P, Ball BA. In vitro interaction between oviduct epithelia and equine sperm. Arch Androl 1993;31:79–86. 18. Battut I, Palmer E, Driancourt MA. Proteins synthesized by equine oviducts: Characterization, variations, and interactions with spermatozoa. Biol Reprod Monogr 1995;1:131–40. 19. Verhage HG, Mavrogianis PA, O’Day-Bowman MB, Schmidt A, Arias EB, Donnelly KM, Boomsma RA, Thibodeaux JK, Fazleabas AT, Jaffe RC. Characteristics of an oviductal glycoprotein and its potential role in the fertilization process. Biol Reprod 1998;58:1098–101. 20. Dobrinski I, Ignotz GG, Thomas PG, Ball BA. Role of carbohydrates in the attachment of equine spermatozoa to uter-
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
ine tubal (oviductal) epithelial cells in vitro. Am J Vet Res 1996;57:1635–9. Suarez SS, Revah I, Lo M, Kolle S. Bull Sperm binding to oviductal epithelium is mediated by a Ca2+-dependent lectin on sperm that recognizes Lewis-a trisaccharide. Biol Reprod 1998;59:39–44. Ball BA, Dobrinski I, Fagnan MS, Thomas PG. Distribution of glycoconjugates in the uterine tube (oviduct) of horses. Am J Vet Res 1997;58:816–22. Thomas PGA, Ball BA, Brinsko SP . Interaction of equine spermatozoa with oviduct epithelial cell explants is affected by estrous cycle and anatomic origin of explant. Biol Reprod 1994;51:222–8. Sabeur K, Ball BA. Characterization of galactose-binding proteins in equine testis and spermatozoa. Anim Reprod Sci 2007;101:74–84. Ignotz GG, Lo MC, Perez CL, Gwathmey TYM, Suarez SS. Characterization of a fucose-binding protein from bull sperm and seminal plasma that may be responsible for formation of the oviductal sperm reservoir. Biol Reprod 2001;64:1806–11. Gwathmey TM, Ignotz GG, Mueller JL, Manjunath P, Suarez SS. Bovine seminal plasma proteins PDC-109, BSPA3, and BSP-30-kDa share functional roles in storing sperm in the oviduct. Biol Reprod 2006;75:501–7. Ignotz GG, Cho MY, Suarez SS. Annexins are candidate oviductal receptors for bovine sperm surface proteins and thus may serve to hold bovine sperm in the oviductal reservoir. Biol Reprod 2007;77:906–13. Ekhlasi-Hundrieser M, Gohr K, Wagner A, Tsolova M, Petrunkina A, Topfer-Petersen E. Spermadhesin AQN1 is a candidate receptor molecule involved in the formation of the oviductal sperm reservoir in the pig. Biol Reprod 2005;73:536–45. Topfer-Petersen E, Ekhlasi-Hundrieser M, Kirchhoff C, Leeb T, Sieme H. The role of stallion seminal proteins in fertilisation. Anim Reprod Sci 2005;89:159–70. Ekhlasi-Hundrieser M, Schafer B, Kirchhoff C, Hess O, Bellair S, Muller P, Topfer-Petersen E. Structural and molecular characterization of equine sperm-binding fibronectin-II module proteins. Mol Reprod Dev 2005;70:45– 57. Reinert M, Calvete JJ, Sanz L, Topfer-Petersen E. Immunohistochemical localization in the stallion genital tract, and topography on spermatozoa of seminal plasma protein SSP-7, a member of the spermadhesin protein family. Andrologia 1997;29:179–86. Hunter RH. Sperm release from oviduct epithelial binding is controlled hormonally by peri-ovulatory graafian follicles. Mol Reprod Dev 2008;75:167–74. Varner DD, Forrest DW, Fuentes F, Taylor TS, Hooper RN, Brinsko SP, Blanchard TL. Measurements of glycosaminoglycans in follicular, oviductal and uterine fluids of mares. J Reprod Fertil Suppl 1991;44:297–306. Varner DD, Bowen JA, Johnson L. Effect of heparin on capacitation/acrosome reaction of equine sperm. Arch Androl 1993;31:199–207. Scott MA, Liu IKM, Overstreet JW. Sperm transport to the oviducts: Abnormalities and their clinical implications. In: Proceedings of the 41st Annual Convention of the American Association of Equine Practitioners, 1995, pp. 1–2. Dobrinski I, Thomas PGA, Ball BA. Cryopreservation reduces the ability of equine spermatozoa to attach to oviductal epithelial cells and zonae pellucdiae in vitro. J Androl 1995;16:536–42.
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Sperm-Oviduct Interactions 37. De Pauw IMC, Van Soom A, Laevens H, Verberckmoes S, De Kruif A. Sperm binding to epithelial oviduct explants in bulls with different nonreturn rates investigated with a new in vitro model. Biol Reprod 2002;67:1073–9. 38. Waberski D, Magnus F, Ferreira FM, Petrunkina AM, Weitze KF, Topfer-Petersen E. Importance of spermbinding assays for fertility prognosis of porcine spermatozoa. Theriogenology 2005;63:470–84. 39. Waberski D, Magnus F, Ardon F, Petrunkina AM, Weitze KF, Topfer-Petersen E. Binding of boar spermatozoa to oviductal epithelium in vitro in relation to sperm morphology and storage time. Reproduction 2006;131:311–18. 40. Petrunkina AM, Waberski D, Gunzel-Apel AR, TopferPetersen E. Determinants of sperm quality and fertility in domestic species. Reproduction 2007;134:3–17. 41. Abad M, Sprecher DJ, Ross P, Friendship RM, Kirkwood RN. Effect of sperm cryopreservation and supplementing semen doses with seminal plasma on the establishment of a sperm reservoir in gilts. Reprod Domest Anim 2007;42:149–52. 42. Hamann H, Jude R, Sieme H, Mertens U, Topfer-Petersen E, Distl O, Leeb T. A polymorphism within the equine
43.
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CRISP3 gene is associated with stallion fertility in Hanoverian warmblood horses. Anim Genet 2007;38:259–64. Reineke A, Hess O, Schambony A, Petrounkina AM, Bader H, Sieme H, Topfer-Petersen E. Sperm-associated seminal plasma proteins – a novel approach for the evaluation of sperm fertilizing ability of stallions? Pferdeheilkunde 1999;15:531–7. Rigby S, Derczo S, Brinsko S, Blanchard T, Taylor T, Forrest DW, Varner D. Oviductal sperm numbers following proximal uterine horn or uterine body insemination. In: Proceedings of the 46th Annual Convention of the American Association of Equine Practitioners, 2000, pp. 332–4. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of Flatrace and National Hunt Thoroughbred mares and stallions in England. Equine Vet J 2007;39:438– 45. Vidament M, Dupere AM, Julienne P, Evain A, Noue P, Palmer E. Equine frozen semen: freezability and fertility field results. Theriogenology 1997;48:907–17. Sieme H, Schafer T, Stout TAE, Klug E, Waberski D. The effects of different insemination regimes on fertility in mares. Theriogenology 2003;60:1153–64.
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Chapter 104
Sperm–Uterine Interactions Terttu Katila
Large numbers of spermatozoa mixed in seminal plasma are ejaculated into the uterus, but only very small numbers gain access into the oviducts. Spermatozoa are rapidly transported through the uterus to the utero-tubal junction (UTJ) by frequent myometrial contractions. Excessive spermatozoa in the uterus are eliminated in due course. Spermatozoa induce a uterine inflammatory reaction by cleaving complement, resulting in powerful chemoattractants for polymorphonuclear leukocytes (PMNs), which phagocytize sperm and bacteria. Finally, the destroyed spermatozoa, PMNs, and inflammatory byproducts have to be removed from the uterus. For this, uterine contractions are again needed. They expel uterine contents through the cervix or drain them via lymphatic vessels. The uterus has a dual role: it ascertains that spermatozoa are safely transported into the UTJ, but it becomes a hostile environment for excessive sperm, leading to their rapid destruction. Contact of sperm with female genital fluids can induce sperm capacitation, an event which precedes the acrosome reaction. While it has been described that primarily progressively motile, morphologically normal sperm gain access into the oviduct, the mechanism by which this occurs in not well understood. It is also unclear whether sperm reservoirs in the uterus exist or whether the spermatozoa are safely stored only in the UTJ and oviductal isthmus. The handling of semen for artificial insemination (AI), e.g., dilution, centrifugation, and freezing of semen, may influence the fate of spermatozoa in the uterus.
Ejaculate During ejaculation, the greatly enlarged glans of the fully erect stallion penis is pressed against the external cervical os of the mare. The penile thrusts create a vacuum in the uterus that aids movement of ejaculated semen through the relaxed estrous cervix
into the uterus.1 Although the stallion’s penis is in the vagina, semen is generally deposited directly into the uterus. Unlike in many other species, stallion semen typically has minimal contact with the hostile acidic vaginal environment. The contact with the structurally very simple cervix of the mare lasts only seconds and does not involve a selection mechanism for sperm. During ejaculation, spermatozoa are mixed with seminal plasma. Seminal plasma consists of secretions from the testes and epididymides, but the majority of the seminal plasma is derived from the accessory sex glands (bulbourethral glands, prostate, vesicular glands, and ampullae). Stallions ejaculate in five to eight jets, the compositions of which vary due to the different contribution of the accessory sex glands. The first two or three jets have high sperm concentration and contain testicular and epididymal secretions. In the last portions of the ejaculate, sperm concentration is low and vesicular gland secretions dominate.2, 3 The composition of seminal plasma differs in the successive jets of the ejaculation, but it is unknown whether the seminal plasma fractions exert different physiological effects in the mare. Seminal plasma contains many different substances, e.g., several proteins. Proteins attach to the spermatozoa rapidly during ejaculation. Some of the seminal plasma components are considered to be beneficial for sperm transport and survival in the female genital tract,4 but some other components decrease sperm survival during storage. Therefore, the proportion of seminal plasma is reduced either by dilution or by centrifugation to optimize sperm quality following cooled or frozen storage. Mammalian spermatozoa can fertilize an oocyte only after they have undergone capacitation and then acrosome reaction. Capacitation is probably initiated by sperm contact with the genital tract fluids of the female. Conversely, decapacitation factors in seminal plasma may prevent premature onset of capacitation.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Sperm–Uterine Interactions This is thought to be achieved through spermatozoal membrane stabilization through maintenance of a proper cholesterol–phospholipid ratio. During capacitation, spermatozoa release decapacitation factor(s).5
Sperm transport By 10–20 minutes after AI, radioactively labeled stallion sperm have been imaged by gamma camera in the tips of the uterine horns.6 The swimming speed of human sperm in aqueous medium is about 5 mm/min.7 It is clear that even highly motile sperm cannot traverse the length of a mare’s uterus rapidly enough without the aid of myometrial contractility. For at least 30 minutes after AI, frequent uterine contractions move semen back and forth between the uterine body and the horns. This ensures efficient distribution of sperm in the uterus, but simultaneously initiates the elimination process by backflow of semen through the cervix.6 In a study by Fiala et al.,8 mares were slaughtered 0.5, 1, 1.5, and 2 hours after AI. Thirty minutes after AI, 67% of the mares showed spermatozoa in oviductal flushes. The percentages of mares with sperm in the oviducts or the numbers of sperm were not different between 0.5 and 2 hours or between the oviducts contralateral or ipsilateral to the periovulatory follicle.8 It is unclear whether semen – spermatozoa and/or seminal plasma – in the uterus induces uterine contractions. Stallion seminal plasma contains high amounts of estrogens, prostaglandin F2α (PGF2α ),9 and oxytocin (OT).10 However, the highest OT concentrations are found in the gel,10 which is removed for AI. Despite the high concentrations of estrogens and PGF2α , seminal plasma does not increase uterine contractions in sows.11 In mares, the numbers of uterine contractions were lower when washed sperm were mixed with seminal plasma instead of skimmilk extender.12 It is possible that uterine contents, spermatozoa, or seminal plasma are not key players in myometrial contractility after mating or AI; rather, the cervix and vagina may play an important role in these contractions. Cervical manipulations in connection with AI or vaginal pressure by the stallion’s penis activate the autonomic nervous system. The contact stimuli interact with adrenergic receptors concurrently with OT release.13
Selection and reservoirs of sperm The stallion ejaculates billions of sperm, but only some hundreds or thousands of spermatozoa will enter the oviduct8, 14 and only a single sperm cell fertilizes the oocyte. Sperm are rapidly transported to the tips of the horns in a current of fluid propelled by strong uterine contractions. The longitudinal fur-
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rows between endometrial folds of the mare may act as channels for rapid migration of sperm. The oviductal papilla is bathed with sperm to ensure that sperm are in close proximity to the UTJ. Sperm induce a physiological uterine inflammation, which is characterized by a rapid influx of PMNs.15 This is the first line of non-specific defense of the uterus and eliminates bacteria and excessive sperm. It is unclear whether the interaction between sperm cells and PMNs is random attachment or whether it is mediated by sperm surface molecules. 16 PMNs did not bind to membrane-damaged boar sperm cells in an in vitro study, whereas membraneintact motile sperm did bind with PMNs. This suggests that the spermatozoal membrane and possible molecules or residues on its surface mediate the interaction between sperm and PMNs. None of the lectins tested were able to block PMN–sperm binding, thereby suggesting that it is not lectin dependent.16 It seems odd that only viable sperm can interact with PMNs. This may represent negative selection, preventing viable but otherwise unfit sperm from entering the UTJ.16 The entrance requirements to the UTJ are strict: normal motility, normal morphology, and, presumably, the presence of certain surface proteins.17 Because damaged sperm are not allowed into the UTJ anyway, their elimination by phagocytosis is not so important, as they will be removed by backflow. The PMNs and sperm numbers in uterine fluid 5 hours after AI were the same in mares inseminated with live compared with killed spermatozoa.18 Troedsson et al.19 found out that two separate seminal plasma proteins were involved in sperm elimination by PMNs in horses. These proteins protected viable, but not damaged, spermatozoa from PMN binding. Other proteins promoted binding and phagocytosis of dead sperm. In mice, acrosomes of ejaculated spermatozoa become coated by immunoglobulin G (IgG), which masks the species-specific antigenic sites over the acrosome. The IgG originates from seminal plasma or from uterine fluids. Since oviductal spermatozoa do not have an IgG coating, it is proposed that labeling of sperm with IgG marks the sperm for phagocytosis.20 The UTJ is presumably the major selector and reservoir of sperm in the mare.14, 21 Sperm numbers in the UTJ are higher than in the more cranial parts of the oviduct.14 The intimate contact and orientation with the epithelium of the UTJ suggest sequestration (Figure 104.1).21 Spermatozoa residing in the UTJ are hidden from PMNs, whereas uterine sperm are readily exposed to PMN attacks (Figure 104.2).14 The elimination of sperm from the uterus is rapid: it starts 0.5 hours after AI and by 4 hours the majority of sperm have been eliminated.6, 22
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Figure 104.1 Scanning electron micrograph of equine spermatozoa fixed in situ at the utero-tubal junction of a mare, 18 hours after insemination. These spermatozoa are located deep in an epithelial fold of the uterine papilla and show a characteristic side-by-side arrangement and orientation that is suggestive of a specific interaction with the epithelium of this region (bar, 2 µm). Reproduced from Scott21 with permission from Elsevier.
In the cat and dog, uterine glands have been suggested as sperm reservoirs.23, 24 Almost all endometrial glands of bitches contained spermatozoa 24 hours after pre-ovulatory AI, albeit in 60% of glands only one spermatozoon was detected.24 In 70% of mares, spermatozoa were located in uterine glands 1 hour after pre-ovulatory AI.8 Also in mares, only very few sperm were detected in a gland. It is unclear whether uterine glands serve as sperm reservoirs in the mare or whether sperm indiscriminately gain access into the glands and are eliminated later. In many investigations about sperm–uterine interactions, results on sperm numbers have been based on flushings of the genital tract. However, histological methods for enumeration of sperm may be more reliable. The attachment between sperm and epithelial cells in the uterus, uterine glands, and oviducts is strong, and flushing could easily underestimate the number of sperm.16, 23, 24
Uterine inflammation Kotilainen et al.15 showed that sperm induce leukocytosis in the equine uterus and that the intensity of PMN influx was positively correlated to sperm concentration (Figure 104.3). The chemotaxis for PMNs is associated with the ability of sperm to activate complement, resulting in cleavage products C5a, C5b, and C3b. C3b is an opsonin and C5a is chemotactic for spermatozoa.19 In mice, the inflammatory response is initiated when seminal plasma proteins interact with estrogen-primed cervical and uterine epithelial cells and activate synthesis of proinflammatory cytokines, e.g. granulocyte–macrophage colony-stimulating factor (GM-CSF), and interleukins IL-6, IL-8, and IL10.25 In seminal plasma of mice, transforming growth
factor β (TGF-β) is thought to be the most important mediator of inflammation and immunity. Its concentration is much higher in seminal plasma than in any other fluids of the body. Plasmin present in the genital tract fluids of the female converts the latent TGF-β to an active form. TGF-β stimulates the release of other cytokines.26 The cytokines attract inflammatory leukocytes including, among others, macrophages, dendritic cells, and granulocytes.25 Similarly, in mares, not only PMNs are recruited in the early inflammatory phase, but also helper T cells (CD4+ ) are significantly increased in numbers 6 hours after AI.27 In pigs, AI with washed sperm induced a greater PMN influx than seminal plasma or whole semen.28 In horses, sperm prewashed to remove seminal plasma induce PMN influx into the uterine lumen.12, 15 Semen extender alone will also induce PMN influx, although not as intensive as that induced by sperm or seminal plasma (Figure 104.3).15 In vitro studies have demonstrated the downregulatory role of stallion seminal plasma in inflammation through decreased complement activation, PMN chemotaxis, sperm opsonization, binding, and phagocytosis.29, 30 Under in vivo conditions, however, seminal plasma itself causes PMN influx,15 and sperm mixed with seminal plasma induce a stronger inflammation than sperm mixed with skim-milk extender.12 The presence of seminal plasma fluid is not required for pregnancy when mares with a normal uterine environment are inseminated. Portus et al.12 washed sperm populations three times and mixed them with either seminal plasma or skim-milk extender; in both groups, 62% of mares conceived following insemination of only 50 × 106 total sperm. On the other hand, when mares are inseminated into an inflamed uterine environment, seminal plasma may be needed for sperm protection. Troedsson et al.30 inseminated mares with and without seminal plasma (500 × 106 progressively motile sperm) after having induced uterine inflammation 12 hours earlier; Only 0.5% (1/22) of mares conceived without seminal plasma and 77% (17/22) conceived with seminal plasma. Later, Alghamdi and Foster31 discovered that spermactivated PMNs extrude their DNA, which traps sperm cells and prevents their movement. Seminal plasma contains DNase that digests the extruded DNA and may free entangled spermatozoa. In practice, mares are often inseminated every 24 hours with frozen semen and achieve good pregnancy rates despite the presence of PMNs in the uterus at that time.22 During semen freezing, the majority of seminal plasma is removed by supernatant aspiration after centrifugation, but some seminal plasma (about 5%) is generally present in the final sample subjected to cryopreservation. Presumably, this amount of seminal plasma is sufficient to
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Figure 104.2 (a) The appearance of freshly ejaculated stallion spermatozoa free of surface of membrane disruptions in the acrosomal region (SEM). (b) Spermatozoa and leukocytes in the uterine horn 4 hours after insemination (SEM). (c) Phagocytosis of a spermatozoon in the uterine lumen (SEM). (d) Spermatozoa in phagocytosis (TEM); magnification 14 800×. (e) Spermatozoon recovered from the uterine lumen 4 hours after insemination showing disruptions of the plasma membrane (SEM). (f) Loss of the acrosome 4 hours after insemination (SEM). SEM, scanned electron microscopy; TEM, transmission electron microscopy. Reproduced from Bader14 with permission from Elsevier.
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Stallion: Anatomy, Physiology and Endocrinology Neutrophils in uterine washings 6 h post infusion No treatment PBS
Skim milk extender Egg yolk extender Raw semen Extended semen Centrifuged semen Frozen semen Frozen semen+skim m. Frozen seminalplasma Frozen semen+sem.pl. 0
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Millions/ml (mean ± s.e.m. Figure 104.3 Neutrophil numbers in uterine lavage fluid of mares 6 hours after intrauterine infusion of semen or extenders. Dashed line indicates typical neutrophil numbers after intrauterine bacterial inoculation. PBS, phosphate-buffered saline. (Reproduced from Kotilainen et al.15 )
provide sperm protection. It has been hypothesized that the low pregnancy rates after frozen semen AI could be explained by the lack of seminal plasma or by the strong inflammatory reaction induced by frozen semen. Addition of seminal plasma to frozen semen, however, does not improve pregnancy rates or decrease inflammation.15 Initially, the inflammation caused by frozen semen was thought to be related to components of freezing extenders (e.g., egg yolk), but the inflammatory response provoked by freezing extenders containing egg yolk is very mild (Figure 104.3).15, 32 Freezing of semen does not appear to influence the degree of uterine inflammation, but concentration of semen – whether frozen or fresh – increases PMN response (Figure 104.3).15 It is common knowledge that reproductively abnormal mares have very low pregnancy rates with frozen semen,33 but it is not explained by a more pronounced inflammatory response to frozen semen.34 Although inflammatory response is an important part of uterine defense, excessive inflammation causes tissue damage by release of destructive enzymes. The control of inflammation is therefore important. In swine, spermadhesin PSPI/PSP-II induces migration of PMNs by stimulating macrophages to release tumor necrosis factor α (TNFα), but this activity is modulated by IL-4 liberated by
mast cells. Dexamethasone, but not indometacin, inhibits chemotactic activity.35 Mast cells have not been studied in mares, but improved pregnancy rates have been reported in barren mares and in mares with intrauterine fluid accumulations by treatment with prednisolone before AI. However, leukocyte numbers did not decrease in the treated mares.36 In a ¨ report by Konig et al.37 involving postpartum cows, dexamethasone-treated cows had higher PMN numbers 6 hours after intrauterine infusion of recombinant human IL-8 (rhIL-8) than the control cows. Dexamethasone decreased plasma concentrations of cortisol and estrogen and reduced the generation of reactive oxygen species (ROS) by uterine PMNs. Excessive extracellular ROS are known to cause tissue damage; thus, the beneficial effect of prednisolone in the study by Dell’Aqua et al.36 may have been due to the prevention of ROS generation. The inflammatory reaction is intensive and rapid, but transient. The PMN numbers in the uterine lumen peak between 6 and 12 hours and only very few are left at 48 hours. Elimination of sperm is also fast: the majority have been eliminated within 4 hours and practically all by 48 hours post insemination.22 This is the timeframe in which reproductively normal mares display efficient uterine contractility. Uterine contractions are needed in the uterine clearance both via lymphatics and by backflow through the cervix. For the latter, a dilated cervix is also a prerequisite. Delayed uterine clearance is diagnosed by ultrasonically visible intrauterine fluid > 2 cm in height after breeding.38 Such fluid represents transudate, increased glandular secretions and inflammatory byproducts.
Additional semen effects Semen has functions beyond fertilization. Seminal plasma deposited in the tip of a ligated uterine horn ipsilateral to the pre-ovulatory follicle advanced ovulation in gilts.39 Later, it was demonstrated that semen-induced cytokines in the uterine lymphatic fluid are delivered by countercurrent transfer to the ovary to accelerate the maturation of pre-ovulatory follicles.40 Similar studies have not been conducted with mares. LeBlanc et al.41 showed that the lymphatic vessels drain uterine contents of mares, and India ink infused into the uterine lumen was visible in lymph nodes after AI. It is possible also that dead sperm or their components could be transported in the lymphatic system. The inflammatory cascade after mating is likely to be essential for induction of immune tolerance to seminal antigens. It may prevent activation of
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Sperm–Uterine Interactions aberrant immunity to sperm and may prime an appropriate female immune response to embryo implantation, since seminal antigens are shared by the conceptus.26 This has been demonstrated in species with a deciduate-type placenta, such as mice and humans. The CD4+ CD25+ T-regulatory (Treg) cells are mediators of immune tolerance in pregnant mice and women. Reduced Treg function has been associated with infertility in women.42 Inadequate expression of IL-6 and IL-1α mRNAs in endometrial tissue has been discovered in women with recurrent miscarriage. The following mechanisms of action have been suggested: perturbed maternal immune response, decidual tissue remodeling and angiogenesis, and dysregulated trophoblast differentiation and invasion.43 Endometrial cups have been suggested as a means of inducing immune tolerance to paternal antigens expressed in the equine embryo.44 The pig has a placenta similar to the horse, but the embryo implants much earlier than the horse embryo. In swine, cytokines induced by seminal plasma proteins have been associated with immunostimulation, modulation of uterine immunity, and regulation of preimplantation embryo development. Seminal plasma treatment has increased the number of embryos and their viability, suggesting that appropriate modulation of uterine immune activity is necessary for reproductive success.45, 46 In addition, sperm DNA, and particularly protein kinase, are probably involved in the modulation of the environment which embryos will face.47 The functions of seminal plasma and spermatozoa show similarities and differences across species. Mice and rabbits are the experimental animals most intensively studied. Pigs and horses have many things in common, e.g., the large volume of seminal plasma. Perhaps we can learn lessons from studies involving pigs; knowledge about semen–uterine interactions is scarce in horses at this juncture.
References 1. Millar R. Forces observed during coitus in thoroughbreds. Aust Vet J 1952;28:127–8. 2. Kosiniak K. Characteristics of the successive jets of ejaculated semen of stallions. J Reprod Fertil Suppl 1975;23: 59–61. 3. Magistrini M, Lindeberg H, Koskinen E, Beau P, Seguin F. Biophysical and H magnetic resonance spectroscopy characteristics of fractionated stallion ejaculates. J Reprod Fertil Suppl 2000;56:101–10. ¨ 4. Topfer-Petersen E, Waberski D, Hess O, Bellair S, Schambony A, Ekhlasi-Hundrieser M, Gentzel M, Reineke A. ¨ die Befruchtung – ein Bedeutung des Seminalplasma fur ¨ kurzer Uberblick [The role of seminal plasma in fertilization]. Tier¨arztl Umsch 1998;53: 447–54. 5. Begley AJ, Quinn P. Decapacitation factors in semen. Clin Reprod Fertil 1982;1:167–75.
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6. Katila T, Sankari S, M¨akel¨a O. Transport of spermatozoa in the reproductive tracts of mares. J Reprod Fertil Suppl 2000;56:571–8. 7. Mortimer ST, Swan MA. Variable kinematics of capacitating human spermatozoa. Hum Reprod 1995;10: 3178–82. 8. Fiala SM, Jobim MIM, Katila T, Gregory RM, Mattos RC. Sperm distribution in the oviduct and uterus of mares within two hours after artificial insemination. Pferdeheilkunde 2008;24: 96–8. 9. Claus R, Dimmick MA. Estrogens and prostaglandin F2α in the semen and blood plasma of stallions. Theriogenology 1992;38:687–93. 10. Watson ED, Nikolakopoulos E, Gilbert C, Goode J. Oxytocin in the semen and gonads of the stallion. Theriogenology 1999;51:855. 11. Langendijk P, Bouwman EG, Soelde NM, Taverne MA, Kemp B. Myometrial activity around estrus in sows: spontaneous activity and effects of estrogens, cloprostenol, seminal plasma and clenbuterol. Theriogenology 2002; 57:1563–77. 12. Portus BJ, Reilas T, Katila T. Effect of seminal plasma on uterine inflammation, contractility and pregnancy rates in mares. Equine Vet J 2005;37:515–19. 13. Madill S, Troedsson MHT, Alexander SL, Shand N, Santschi EM, Irvine CHG. Simultaneous recording of pituitary oxytocin secretion and myometrial activity in oestrous mares exposed to various breeding stimuli. J Reprod Fertil Suppl 2000;56:351–61. 14. Bader H. An investigation of sperm migration into the oviducts of the mare. J Reprod Fertil Suppl 1982;32:59–64. 15. Kotilainen T, Huhtinen M, Katila T. Sperm induced leukocytosis in the equine uterus. Theriogenology 1994;41:629– 36. 16. Taylor U, Rath D, Zerbe H, Schuberth HJ. Interaction of intact porcine spermatozoa with epithelial cells and neutrophilic granulocytes during uterine passage. Reprod Domest Anim 2008;43:166–75. 17. Suarez SS, Pacey AA. Sperm transport in the female reproductive tract. Hum Reprod Update 2006;12:23–37. 18. Katila T. Neutrophils in uterine fluid after insemination with fresh live spermatozoa or with killed spermatozoa. Pferdeheilkunde 1997;13:540. 19. Troedsson MHT, Desvousges A, Alghamdi AS, Dahms B, Cow CA, Hayna J, Valesco R, Callahan PT, Macpherson ML, Pozor M, Buhi WC. Components in seminal plasma regulating sperm transport and elimination. Anim Reprod Sci 2005;89:171–86. 20. Cohen J, Werrett DJ. Antibodies and sperm survival in the female tract of the mouse and rabbit. J Reprod Fertil 1975;42:301–10. 21. Scott MA. A glimpse at sperm function in vivo: sperm transport and epithelial interaction in the female reproductive tract. Anim Reprod Sci 2000; 60–61: 337–48. 22. Katila T. Onset and duration of uterine inflammatory response of mares after insemination with fresh semen. Biol Reprod Monogr 1995;1:515–17. 23. Chatdarong K, Lohachit C, Linde-Forsberg C. Distribution of spermatozoa in the female reproductive tract of the domestic cat in relation to ovulation induced by natural mating. Theriogenology 2004;62:1027–41. 24. Rijsselaere T, Van Soom A, Van Cruchten S, Coryn M, Gortz K, Maes D, deKruif A. Sperm distribution in the genital tract of the bitch following artificial insemination in relation to the time of ovulation. Reproduction 2004;128:801–11.
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25. Robertson SA. Seminal fluid signaling in the female reproductive tract: lessons from rodents and pigs. J Anim Sci 2007; 85:E36–44. 26. Robertson SA, Ingman WV, O’Leary S, Sharkey DJ, Tremellen KP. Transforming growth factor β – a mediator of immune deviation in seminal plasma. J Reprod Immunol 2002; 57:109–28. ´ A-M, Katila T, Magnusson U, Nummij¨arvi A, 27. Tunon Rodriguez-Martinez H. T-cell distribution in two different segments of the equine endometrium 6 and 48 hours after insemination. Theriogenology 2000;54:835–41. 28. Rozeboom KJ, Troedsson MH, Molitor TW, Crabo BG. The effect of spermatozoa and seminal plasma on leukocyte migration into the uterus of gilts. J Anim Sci 1999;77: 2201–6. 29. Troedsson MHT, Lee C-S, Franklin RD, Crabo BG. The role of seminal plasma in post-breeding uterine inflammation. J Reprod Fertil Suppl 2000;56:341–9. 30. Troedsson MHT, Alghamdi AS, Mattisen J. Equine seminal plasma protects the fertility of spermatozoa in an inflamed uterine environment. Theriogenology 2002;58: 453–6. 31. Alghamdi SA, Foster DN. Seminal DNase frees spermatozoa entangled in neutrophil extracellular traps. Biol Reprod 2005;73:1174–81. 32. Palm F, Walter I, Budik S, Aurich C. Influence of different semen extenders and seminal plasma on the inflammatory response of the endometrium in oestrous mares. Anim Reprod Sci 2006;94:286–9. 33. Sieme H, Bonk A, Hamann H, Klug E, Katila T. Effects of artificial insemination techniques and sperm doses on fertility of normal mares and mares with abnormal reproductive history. Theriogenology 2004;62:915–28. ¨ 34. Guvenc K, Reilas T, Katila T. Effect of frozen semen on the uterus of mares with pathological uterine changes. Reprod Nutr Dev 2004;44:243–50. 35. Assreuy AMS, Alencar NMN, Cavada BS, Rocha-Filho DR, Feitosa RF, Cunha FQ, Calvete JJ, Ribeiro RA. Porcine spermadhesin PSP-I/PSP-II stimulates macrophages to release a neutrophil chemotactic substance: modulation on by mast cells. Biol Reprod 2003;68:1836–41. 36. Dell’Aqua Jr JA, Papa FO, Lopes MD, Alvarenga MA, Macedo L, Melo CM. Modulation of acute uterine inflam-
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matory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94:270–3. ¨ Konig T, Schuberth HJ, Leibold W, Zerbe H. Dexamethasone depresses the expression of L-selectin but not the in vivo migration of bovine neutrophils into the uterus. Theriogenology 2006;65:1227–41. Brinsko SP, Rigby SL, Varner DD, Blanchard TL. A practical method for recognizing mares susceptible to post-breeding endometritis. In: 49th Annual Convention of the American Association of Equine Practitioners, 2003. www.ivis.org Waberski D, Kremer H, Borchardt Neto G, Jungblut PW, Kallweit E, Weitze KF. Studies on local effect of boar seminal plasma on ovulation time in gilts. Zentralbl Veterin¨armed A 1999;46:431–8. ¨ Waberski D, Dohring A, Ardon F, Ritter N, Zerbe H, Schuberth HJ, Hewicker-Trautwein M, Weitze KF, Hunter RH. Physiological routes from intra-uterine seminal contents to advancement of ovulation. Acta Vet Scand 2006;48: 13. LeBlanc MM, Johnson RD, Calderwood Mays MB, Valderrama C. Lymphatic clearance of India ink in reproductively normal mares and mares susceptible to endometritis. Biol Reprod Monogr 1995;1:501–6. Jasper MJ, Tremellen KP, Robertson SA. Primary unexplained infertility is associated with reduced expression of the T-regulatory cell transcription factor Foxp3 in endometrial tissue. Mol Hum Reprod 2006;12:301–8. Jasper MJ, Tremellen KP, Robertson SA. Reduced expression of IL-6 and IL-1alpha mRNAs in secretory phase endometrium of women with recurrent miscarriage. J Reprod Immunol 2007;73:74–84. Allen WR. Fetomaternal interactions and influences during equine pregnancy. Reproduction 2001;121:513–27. Leshin LS, Raj SM, Smith CK, Kwok SC, Kraeling RR, Li WI. Immunostimulatory effects of pig seminal plasma proteins on pig lymphocytes. J Reprod Fertil 1998;114:77–84. O’Leary S, Jasper MJ, Warnes GM, Armstrong DT, Robertson SA. Seminal plasma regulates endometrial cytokine expression, leukocyte recruitment and embryo development in the pig. Reproduction 2004;128:237–47. Chaykin S, Watson G. Reproduction in mice: spermatozoa as factors in the development and implantation of embryos. Gamete Res 1983;7:63–73.
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Chapter 105
Testicular Descent Mimi Arighi
Normal testes In the normal equine male fetus, both testes descend into the scrotum between 30 days before and 10 days after birth.1 Three anatomic structures are involved in normal descent: the vaginal process, the gubernaculum, and the inguinal canal. The vaginal process is a flask-shaped evagination of the peritoneum in the inguinal region. This evagination exits the abdominal wall at the inguinal canal and passes into the scrotum. In the adult, this structure is divided into two parts: (i) a long, narrow neck containing the spermatic cord consisting of blood vessels, nerves, and lymphatics of the testis, the epididymis, the vas deferens, and the internal cremaster muscle and (ii) a dilated body containing the testis, epididymis, and the proximal portion of the vas deferens.2, 3 The opening of the vaginal process, through which the gubernaculum proprium passes, forms the vaginal cavity.1–4 The vaginal ring is at the level of the internal inguinal ring.2, 5 The testes are encased in the tunica albuginea and lie within an outpocket of abdominal peritoneum known as the tunica vaginalis. The tunica vaginalis consists of visceral and parietal tunics. The visceral tunic is tightly adhered to the testis, ducts, and vessels, and the parietal tunic is continuous with the parietal peritoneum. The fetal gubernaculum is divided into three main parts: (i) the portion connecting the testis to the tail of the epididymis – the caudal gonadal ligament or the proper ligament of the testis, (ii) a portion between the tail of the epididymis and vaginal process – the caudal ligament of the epididymis, and (iii) a remaining portion extending from the vaginal process to the scrotal fold – the scrotal ligament.2, 3, 6, 7 The gubernacular ligament passes from the end of the vaginal process to the scrotum but does not form a firm anchor to the process. The gubernaculum helps to keep the passageway open by expanding in size and stretching the vaginal ring.1–4,6,8–12
The inguinal canal is a passageway between the internal inguinal ring and the external inguinal ring. The internal inguinal ring is formed from the internal abdominal oblique muscle, and the external inguinal ring is formed from both the internal and external abdominal oblique muscles.2, 3 The external cremaster muscle, which lies on the outside of the vaginal tunic, is derived from the internal abdominal oblique muscle and is continuous with the parietal layer of the tunica vaginalis inserting at the caudal pole of the testis. When it contracts, the cremaster muscle elevates the associated testis toward the external inguinal ring.3, 13 During embryonic development, the equine gonad becomes differentiated into a testis at 51/2 weeks of gestation and lies ventral to the mesonephric kidney.2–4,8,14–16 At 4 months of gestation, the cranial pole of the testis overlies the caudal pole of the mesonephric kidney, whereas its caudal pole is still some distance from the vaginal ring. The equine gonad begins to hypertrophy at 6 weeks of gestation through an increase in the number of interstitial cells; by 5 months of gestation, this hypertrophy results in the caudal pole of the testis lying close to the vaginal ring.1, 4, 8, 14,17–19 The high growth rate of the fetal testis corresponds to a period of high blood estrogen in the pregnant mare.17, 19 The equine fetal testis weighs about 20 g at 5 months of gestation, 50 g at 8 months, and 30 g at 10 months.1, 8, 9, 14 After about 71/2 months of gestation, the testis begins to decrease in size owing to degeneration of the interstitial cells; by 10 months, there is a clear separation between the testis and the kidney.1, 4, 8, 14, 16, 19 Between 5 months and 10 months of gestation, the inguinal canal is too small for the passage of the testis. The developing gubernaculum and vaginal process hold the testis at the vaginal ring. Other abdominal viscera also aid in retaining the testes in the pelvic region. After 5 months of gestation, the enlarged distal end of the gubernaculum proprium becomes the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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main force preventing inversion of the vaginal process. Firm fibrous connections between the process and the inguinal canal also exist. After 8 months of gestation, the gubernaculum proprium begins to decrease in size and only the fibrous connections in the inguinal canal hold the vaginal process outside the vaginal ring.1, 2, 4,8–10,14, 16, 17, 19 The vaginal process and gubernaculum continue to lengthen at about the same rate up to 81/2 months of gestation. During this time, the caudal gonadal ligament, between the testis and cauda epididymis, increases in length faster than the gubernaculum, resulting in the cauda epididymis becoming progressively displaced from the caudal pole of the testis. The cauda epididymis does, in fact, descend into the inguinal canal at about 5 months of gestation, well in advance of the testis.1–4,11, 14, 15, 17, 18 After 81/2 months of gestation, there is a gradual decrease in the length of the gubernaculum while the vaginal process remains static, so that the caudal pole of the regressing testis is drawn into the opening of the vaginal ring, which has been dilated by the increasing diameter of the caudal gonadal ligament and the body of the epididymis.2, 4, 14 Owing to the length of the caudal gonadal ligament, hypertrophy of the fetal testis, the small diameter of the vaginal ring, and the lack of tension on the gubernacula during midgestation, descent of the testes through the inguinal canal must occur late in gestation or in the early postnatal period.18 The actual passage of the testis through the vaginal ring is a rapid process, which typically occurs during the last month of gestation. At this stage of gestation, the gubernacular length continues to decrease while there is a rapid increase in the length of the vaginal process, which increases tension of the gubernaculum on the small flaccid testis. The testis does not pass immediately into the scrotum because the gubernaculum mass remains below it. At birth, most testes lie in the inguinal canal whereas the gubernaculum lies in the scrotal region and can easily be mistaken for a testis. Further passage of the testis into the scrotum after birth is affected by a progressive reduction in length and mass of the gubernaculum, which finally becomes a small dense fibrous band.1, 2, 4, 6, 17, 19 During the first 2 weeks of neonatal life, the internal vaginal ring typically constricts to about 1 cm in diameter and fibroses; thus, the testis can generally not be forced in either direction after this time.4, 6, 8, 9, 16, 17 At birth, the weight of each testis ranges from 5 g to 20 g and testicular size does not change much throughout the first 10 months of life. There is slight growth between 11 and 16 months of age, and then rapid development of the testes starting around 18 months of age.1 Puberty in pony stallions was reported to occur between 11 and 15 months of age
based on the appearance of any sperm in the ejaculate.20–22 Puberty in Quarter Horse stallions was found to occur between 13 months and 23 months of age, and this was based on the first ejaculate containing 50 × 106 spermatozoa with ≥ 10% motile.23 Although stallions produce spermatozoa throughout the year, their testes show seasonality, with maximum production of sperm occurring in the summer. In one study, testes of 17 stallions (which ranged from 2 to 20 years of age) were found to weigh an average of 167 g, with an average length of 8.5 cm and an average width of 5.0 cm.24 In another study, 48 stallions (age range from 2 to 16 years) were evaluated and their testes had an average weight of 164 g, with an average length of 10.4 cm and an average width of 5.6 cm.25 In the second study, the testes of stallions 7 years or older were shown to be significantly larger than those of the younger horses.25 In the first study, the age factor was not evaluated. Normal testicular descent involves a process in which the essential physical components are: a testis with sufficient freedom of attachment to allow it to be moved from its original dorsal abdominal location to its final scrotal position; patent inguinal rings leading to a properly developed tunica vaginalis; the proper size relationship between the testis and the inguinal ring; and sufficient force to move the testis to the inguinal ring.
Cryptorchid testes Cryptorchidism, or the failure of normal testicular descent, is most frequently found in humans, swine, and horses, although it does occur in many other mammalian species.8, 26 In horses, cryptorchidism, with one or both testes retained, is a relatively common occurrence. The percentage of foals having cryptorchidism is estimated to be 5–8% of male foals, with most being unilateral cryptorchids. It is not known how many diagnosed retained testes would descend at a later date, nor how many would remain undescended.19, 27 The highest incidence of cryptorchidism has been reported in Quarter Horses, followed by Percheron, American Saddle Horses, and ponies.6, 27, 28 When both testes are retained in the abdomen, the animal is sterile owing to the detrimental effect of higher temperatures on spermatogenesis, and such testes have been shown to be more prone to development of secondary tumors.5, 19, 29 Unilateral cryptorchids are fertile if spermatogenesis occurs in the descended testis.19 Cryptorchid testes are usually smaller than normal, are soft and flaccid, and weigh between 25 and 131 g, which is considerably less than normal scrotal testes (Figure 105.1).11,30–32 Histologically, cryptorchid testes have arrested spermatogenesis with changes seen mainly in the germ cell layers. Their
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(a)
(b)
Figure 105.1 Comparison of size between a scrotal testis (a) and an abdominal testis (b).
Leydig cells appear normal.11, 19, 30, 33, 34 Spermatogenesis does not seem to proceed beyond A- or Bspermatogonia in abdominal testes, or primary spermatocytes in the case of inguinal testes.3, 17, 18, 31 Mean tubular diameters and spermatogenic cell layers were significantly less in inguinal and abdominal testes than in normally descended scrotal testes (Figures 105.2–105.4).31 In horses, development of seminiferous tubules in abdominal testes appeared to proceed to the extent normally found in foals of 3–4 months of age, whereas development in the inguinal testes roughly corresponded to that of a 9- to 12month-old normal foal.3, 6, 11, 17, 18
Figure 105.2 Histological section of a portion of a scrotal testis from a stallion at medium power. Numerous seminiferous tubules are depicted with multiple layers of germinal epithelium. Tubules are separated by scanty interstitial cells. From Arighi M, Bosu WTK, Raeside JL. Hormonal Diagnosis of Equine Cryptorchidism and Histology of the Retained Testes. Proc. Am. Ass. Equine Practitioners 1985; 31:591–602.
Figure 105.3 Histological section of a portion of an inguinal testis from a stallion at medium power that shows a decrease in the layers of germinal epithelium that line the tubules in comparison with a scrotal testis.
The interstitial (Leydig) cells, which produce testosterone and other hormones, are not as heat sensitive as are the cells of the seminiferous epithelium. Thus, although bilateral abdominal cryptorchid horses may not produce viable sperm, they will usually still exhibit normal secondary sex characteristics, including libido.29 This is especially true in horses, since Leydig cells are very abundant and increase in numbers with age.35 A cryptorchid horse may have either one or both testes retained in an abdominal and/or inguinal location and diagnosis can be complicated if a single normal descended testis has been removed.1, 2, 6,8–10,15, 36 The retained abdominal testis can be complete or incomplete. When all parts of the testis and epididymis are located in the abdominal cavity, the cryptorchidism is complete. If either the epididymis or part of the testis is located in the inguinal canal the cryptorchidism is incomplete.8, 19, 28 In the horse, the epididymis descends first, thus an abdominally retained testis may have an attached
Figure 105.4 Histological section of a portion of an abdominal testis from a stallion at medium power that shows a decrease in the layers of germinal epithelium that line the tubules in comparison with either an inguinal or a scrotal testis, and an increase in the number of interstitial cells.
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epididymis within the inguinal canal. If the change in the gubernaculum does not occur at the proper time (decrease in size), or if sufficient gonadal shrinkage does not take place, abdominal testes may never descend because, following birth, gonadal size begins to increase, and the vaginal ring constricts.1, 6, 8, 17 If the gonad does not escape the abdomen at about the time of parturition, the horse will likely become an abdominal cryptorchid.6, 8, 17 Causes of maldescent remain unsolved and may be the result of several factors. Several explanations for maldescent have been proposed in humans: 1. defective hypothalamic–pituitary axis and deficiency of luteinizing hormone (LH);8, 15, 18 2. anatomic defects such as gubernacular abnormalities;8, 15, 18 3. genetic factors, based on familial and inherited tendencies;15, 18, 19 4. defects in the testis itself, resulting in deficiency of testicular production of androgens which influence ductus deferens, epididymis, and gubernaculum during descent;8, 15, 18,37–39 5. testicular dysgenesis based on abnormal ‘testicular’ chromosomes.18, 33, 40, 41 A deficiency of gonadotropic hormonal stimulation appears to be a popular theory to explain bilateral cryptorchidism in humans, and perhaps horses. Based on experiments with rats, it is now known that premature descent can be induced by treatment with gonadotropin. In addition, since normal testicular descent occurs at a time when endogenous gonadotropin levels are rising, it is assumed that descent of the testes into the scrotum in rats is under regulation of the pituitary gonadotropins. Testicular descent can also be inhibited in the rat by daily injections of estradiol, and can be reversed by simultaneous treatment with human chorionic gonadotropin (hCG). It is thought that estrogen inhibits testicular descent by suppressing pituitary gonadotropin release, at least in the rat. Only dihydrotestosterone, not testosterone or any other androgen, can induce premature testicular descent in the rat.15 Although the findings in rats may not be directly applicable to humans or horses, the finding of bilateral cryptorchidism supports a need for an intact hypothalamic–pituitary axis and adequate levels of LH for appropriate testicular descent to occur. However attractive the hormonal theory is, it is difficult to apply to unilateral cryptorchidism, which is far more common in both humans and horses (three to five times) than bilateral cryptorchidism.15 Other causes of, or complicating factors in, failure of testicular descent are thought to be undue stretching of the gubernacular cord; abdominal pres-
sure inadequate for proper expansion of the vaginal process; inadequate growth of the gubernaculum and epididymis resulting in failure of the testis to expand the inguinal ring; or displacement of the testis into the pelvic cavity where it is held by organ pressure resulting in ineffective gubernacular tension.1, 3, 8, 9, 14 In addition, some authors have suggested that cryptorchidism in the horse can be an inherited abnormality with dominant inheritance,17, 18, 40, 42 whereas others have concluded that transmission is a simple autosomal recessive trait.19 At least two genetic factors have been reported to be involved, one of which is located on the sex chromosome.19 The descent of the right testis in equine fetuses is more advanced than the left in the majority of the fetuses between 9 months of gestation and birth.1, 6, 16 The right testis is also found to be significantly smaller in size than the left. The difference in size seems to persist after birth. According to some authors, the smaller size of the right testis hinders its passage from the abdomen to the inguinal area, and retention in the inguinal area.4, 16, 32 Overall, inguinal retention is reportedly more common than abdominal retention.28 Some authors have reported more left abdominally retained testes,4, 6, 16, 33,43–48 whereas others have reported equal incidence of side.28 Reports of location of inguinal testes are more variable, with some authors reporting a higher incidence of right testis involvement,4, 11, 16, 17, 44, 47, 49 while others have found an equal incidence on both sides.6, 33, 45 Bilateral cryptorchid horses are much less commonly seen (14% of cryptorchid horses) but, when encountered, both testes may be in the abdomen, in the inguinal areas, or one in each site.27 Some studies report predominance of both retained testes being inguinal,11 whereas others have reported a greater incidence of both being abdominal.6, 17, 44, 45 Inguinal testes are more commonly found in yearlings and 2-year-old horses than in older horses, and the ratio of inguinal to abdominal cryptorchid diagnoses decreases with advancing age.16, 28, 33 These findings could be explained by the fact that some of the inguinal testes may advance into the scrotal area as the animal becomes older.16, 28 Cryptorchid horses have a variety of undesirable behavioral characteristics, as do colts with both testes normally descended. They can be irritable and nervous, have abnormal sexual excitement, or vicious behavior. Owners might prefer to castrate these animals as soon as possible because their behavioral traits make them poor candidates as riding horses, and their retained testis(es) might make them not ideal breeding prospects.19 Surprisingly, however, many equine breed registries condone the use of unilateral cryptorchid stallions for breeding purposes.
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Testicular Descent
Diagnostic tests for cryptorchidism At 1–2 years of age a stallion should receive an examination of the inguinal and scrotal areas to determine that both testes have descended into the scrotal sac. If either one or both testes cannot be palpated in the scrotal and/or inguinal areas, then other tests must be done to determine the location of the testis(es) so the best surgical approach can be chosen to locate and remove that testis(es). Rectal examination Some practitioners carry out rectal examinations on stallions to determine testis location; however, the usefulness of rectal palpation to determine the location of a retained testis is controversial. Some authors have found this test reliable whereas others have not. In a study carried out by Stickle and Fessler,6 preoperative diagnosis by rectal palpation of the vaginal rings was found to be 87% accurate. If an incorrect diagnosis was made, it was usually because the ductus deferens was felt to enter the vaginal ring from the abdomen, and so it was assumed incorrectly that the testis had also left the abdomen and would be in the inguinal canal. However, instead, it was later found that the testis was still in the abdomen (partial abdominal cryptorchid). Rectal palpation can also be dangerous, especially in a young stallion, since a rectal tear could occur if the stallion moves suddenly, or the veterinarian could be injured by a stallion’s kick. During rectal palpation, a normal vaginal ring is about 1.2 cm in diameter with the ductus deferens located at the caudomedial aspect of the ring if the testicular structures have passed normally into the scrotum. Rectal palpation findings may include either the absence or presence of the ductus deferens passing through the vaginal ring towards the scrotum, or the presence of the testis in the abdomen. If the ductus deferens leaving the abdomen through the vaginal ring is not felt, the testis is most likely in the abdomen. If it is felt, then it has traditionally indicated that either the testis is in the inguinal canal/scrotum or the ductus deferens is in the inguinal canal with the testis still in the abdomen.6, 50 Ultrasonography This can be done inguinally, rectally, or both. Animals need sedation in most cases before ultrasonography can be safely done, and xylazine and butorphanol have successfully been used. A 5-MHz transrectal transducer is needed for a successful ultrasonographic examination of this area. For inguinal scans, the operator first palpates the inguinal area and then places the transducer longitudinally over
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the external inguinal ring on the side of the retained testis and scans the inguinal ring thoroughly. Rectal scans are started at the pelvic brim and moved forward in an axial to abaxial pattern over the side of the retained testis. Most abdominal testes are located over or within 6 cm of the deep inguinal ring. In 11 horses reported by Jann and Rains,50 only two of the 11 retained testes could be palpated rectally or inguinally whereas 100% of the testes were found via ultrasonography. Also, testicular size determined presurgically via ultrasound in all cases closely approximated the actual size obtained by gross measurement of the excised testis.50, 51 Basal testosterone levels Hormone concentrations of animals in which no scrotal/inguinal testes are palpable have been used to determine whether testicular tissue is present. Basal hormonal concentrations can differentiate geldings from animals with testicular tissue. Although low testosterone concentrations in some stallions and high concentrations in some geldings may overlap,52, 53 it has been found that, in all ages of horses, resting testosterone was the most reliable method for differentiating geldings from animals with testicular tissue. Geldings had a mean testosterone value of 0.12 ng/mL (range 0.03–0.15 ng/mL), whereas animals with testicular tissue had mean values ranging from 0.72 to 0.98 ng/mL (range 0.38–1.2 ng/mL).30, 34 Only 5% of the animals (5/101) were incorrectly diagnosed using this method. Eleven percent (11/101) would have needed further testing.30, 34 In one study, it was concluded that resting testosterone concentrations of ≤ 0.24 ng/mL were diagnostic of absence of testicular tissue (gelding), whereas resting concentrations of ≥ 0.44 ng/mL were indicative of the presence of testicular tissue.30, 34 Others have confirmed this finding.41 Values between 0.24 and 0.44 ng/mL were not diagnostic, and in these cases another type of hormonal testing, such as hCG stimulation or basal levels of estrone sulfate, should be run concurrently.30, 34 hCG stimulation test hCG has also been utilized to identify the presence or absence of testicular tissue in a horse. The administration of hCG increases synthesis and secretion of testosterone by Leydig cells in the stallion.18, 54 Testosterone secretion in response to hCG is longer than secretion following a natural, episodic discharge of LH.1, 55, 56 The injection of hCG is followed within 1–2 hours by an elevation of testosterone to a level much higher than the normal episodic changes. The testing procedure compares a base plasma testosterone level with a second one taken at
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1 hour, 3 hours, 24 hours, or even 3 days after the injection of 6000–12,000 international units (IU) of hCG.7, 9, 30, 34, 47, 52,56–59 In known geldings, basal testosterone concentrations averaged 0.015 ng/mL (0.011–0.02 ng/mL). Stallions, on the other hand, had basal concentrations ranging from 0.065 ng/mL to 1.6 ng/mL, whereas cryptorchids had a mean testosterone concentration of 0.42 ng/mL.52 In intact stallions, testosterone concentrations increased four- to 30-fold between 0 and 120 minutes following intravenous (i.v.) injection of hCG and the testosterone concentration in these animals was always > 0.1 ng/mL. In geldings, the concentration increased only zero- to twofold with the testosterone concentration still at < 0.04 ng/mL; and in cryptorchids, a threefold increase was seen with the testosterone concentration rising to > 0.1 ng/mL.59, 60 In one study, it was reported that a plasma sample taken before stimulation, then stimulation with 10,000 IU hCG given intravenously, and then two more samples taken at 1 hour and 24 hours after stimulation captured two testosterone peaks if testicular tissue was present.30, 34 Location of the testis, age of the animal, and time of year affect the response of the testis to hCG. Abdominal testes were much less responsive to gonadotropin stimulation than scrotal testes.30, 34, 59 The response is more pronounced in the summer than in the winter, possibly because of the seasonal variation in testosterone production. Colts less than 18 months of age did not, in many instances, produce enough testosterone in response to hCG to differentiate them from geldings.59 This was also seen with bilateral abdominal cryptorchid stallions.30 Basal estrone sulfate levels Stallion testes produce uniquely high amounts of estrogens.54, 61, 62 The stallion’s daily estrogen output in urine exceeds that of a non-pregnant mare by a factor of 100–200.57, 63 The urine or blood of geldings contains very little estrogen.63 Urinary estrogen levels in immature stallions are only 0.2–0.5% of a mature stallion, and, as sexual maturity approaches, a rise in urinary estrogen secretion occurs.63 The highest level of estrogen production and secretion is attained at the age of 4–5 years.1 Estrone sulfate is by far the most abundant plasma estrogen in the stallion.64, 65 Findings of large quantities of estrone sulfate and only trace amounts of estrogens as free compounds in stallion testes may partly answer the question of how males retain their masculinity in spite of enormous estrogen secretion.65 Different groups of researchers have found significantly higher total estrogen levels in the peripheral plasma of bilaterally cryptorchid horses and boars,
which suggests that the cryptorchid testis synthesizes more estrogens than does the scrotal testis.48, 66 One of these groups66 also saw a higher concentration of total estrogens in the cryptorchid testis than in the scrotal testis in unilaterally cryptorchid animals. The germinal epithelium and the Sertoli cells are destroyed in cryptorchid testes, and, since the Sertoli cells in the testis normally secrete a follicle-stimulating hormone (FSH)-inhibiting substance (inhibin), a lack of FSH inhibition could perhaps explain the marked increase in estrogen levels observed in bilateral cryptorchids. Since the total estrogen levels in unilateral cryptorchids are only slightly higher than those of normal stallions, the scrotal testis in the unilateral cryptorchid may be producing sufficient inhibin (a FSH inhibitor) to control the overall production and secretion of estrogen by the cryptorchid tissues.1, 66 These results also suggest that the measurement of total estrogen in mature horses may be useful in determining whether testicular tissue is still present in a particular suspect animal.66 Since the concentration of estrone sulfate in the plasma of mature stallions (> 3 years of age) is nearly 100-fold greater than that of testosterone, and geldings’ plasma contains very little estrone sulfate, the measurement of estrone sulfate in the plasma of a horse > 3 years old has been shown to be a good way to determine whether testicular tissue is still present in that horse.30, 41, 60, 67 This has been shown, however, to not be true for donkeys,41 which have an absence of conjugated estrogen. Thus, this test would not be good for determining whether testicular tissue was still present in a supposedly castrated donkey. In a few studies, it was shown that geldings had resting estrone sulfate concentrations of ≤ 0.12 ng/mL whereas animals with testicular tissue had concentrations of ≥ 1.0 ng/mL. There was a lower likelihood of false positive results if the animal being tested was > 3 years of age.30, 34, 41 If the hormone concentration falls in the area between these two values, the test should be re-run or an assay for basal level testosterone or hCG stimulation should be carried out simultaneously.30, 34, 41, 68 In conclusion, there are three possible hormonal tests that can be run: resting testosterone concentration, resting estrone sulfate concentration, or hCG stimulation. In horses with no palpable testes, many times one will need to run more than one of these tests to confirm for sure whether testicular tissue is present in the horse. Most times, resting testosterone is the first test that is run since it is more specific than resting estrone sulfate concentration in young horses and is an easier test to run than hCG stimulation. If the results of that test are non-diagnostic, then the second test that will be run will depend on the age of the horse. If the horse is > 3 years of age, then an
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Testicular Descent estrone sulfate test would be a good second test to run; if the horse is younger than that, then the hCG stimulation test is more reliable.
References 1. Amann RP. A review of anatomy and physiology of the stallion. J Equine Vet Sci 1981;1:83–106. 2. Ashdown RR. Symposium: the inguinal canal and its relationship to health and disease. I. The anatomy of the inguinal canal in the domestic mammals. Vet Rec 1963;75:1345–51. 3. Nickel R, Schummer A, S´eiferle E, Sack WO. The Viscera of the Domestic Mammal. New York: Verlag Paul Parey, 1973; pp. 351–92. 4. Smith JA. The development and descent of the testis in the horse. Vet Annu 1975;15:156–61. 5. Stick JA. Teratoma and cyst formation of the equine cryptorchid testicle. J Am Vet Med Assoc 1980;176:211–14. 6. Stickle RL, Fessler JF. Retrospective study of 350 cases of equine cryptorchidism. J Am Vet Med Assoc 1978;172: 343–6. 7. Trotter GW, Aanes WA. A complication of cryptorchid castration in three horses. J Am Vet Med Assoc 1981;178: 246–8. 8. Bergin WC, Gier HT, Marion GB, Coffman JR. A developmental concept of equine cryptorchidism. Biol Reprod 1970;3:82–92. 9. Collier MA. Equine cryptorchidectomy: surgical considerations and approaches. Mod Vet Pract 1980;61:511–14. 10. Sisson S. Male genitalia. In: Sisson S, Grossman JD (eds) The Anatomy of the Domestic Animal, vol. 1, 5th edn. Toronto: W.B. Saunders Co., 1975; pp. 531–41. 11. Bishop MWH, David JSF, Messervy A. Some observations on cryptorchidism in the horses. Vet Rec 1964;76:1041–8. 12. Frank ER. Veterinary Surgery, 7th edn. Minneapolis: Burgess Publishing, 1964; pp. 267–74. 13. Searle D, Dart AJ, Dart CM, Hodgson DR. Equine castration: review of anatomy, approaches, techniques, and complications in normal, cryptorchid and monorchid horses. Aust Vet J 1999;77:428–34. 14. Gier HT, Marion GB. Development of the mammalian testes. In: Johnson AD, Gomes WR, Vandemark NL (eds) The Testes, vol. 1. New York: Academic Press, 1970; pp. 1–45. 15. Shapiro SR, Bodai BI. Current concepts of the undescended testis. Surg Gynecol Obstet 1978;147:617–25. 16. Genetzky RM, Shira MJ, Schneider EJ, Easley JK. Equine cryptorchidism: pathogenesis, diagnosis and treatment. Comp Cont Educ 1984;6:S577–81. 17. Arthur GH. The surgery of the equine cryptorchid. Vet Rec 1961;73:385–9. 18. Vaughan JT. Surgery of the male equine reproductive system. In: Jennings PB (ed.) The Practice of Large Animal Surgery. Toronto: W.B. Saunders Co., 1984; pp. 1083–105. 19. Leipold HW, DeBowes RM, Bennett S, Cox JH, Clem MF. Cryptorchidism in the horse: genetic implications. In: Proceedings of 31st Annual Convention of the American Association of Equine Practitioners, Toronto, Canada, 1986; pp. 579–90. 20. Skinner JD, Bowen J. Puberty in the welsh stallion. J Reprod Fertil 1968;16:133–5.
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21. Wesson JA, Ginther OJ. Plasma gonadotrophin concentrations in intact female and intact and castrated male prepubertal ponies. Biol Reprod 1980;22:541–9. 22. Wesson JA, Ginther OJ. Puberty in the male pony: plasma gonadotropin concentrations and the effects of castration. Anim Reprod Sci 1981;4:165–75. 23. Naden J, Amann RP, Squires EL. Testicular growth, hormone concentrations, seminal characteristics, and sexual behavior in stallions. J Reprod Fertil 1990;88:167–76. 24. Love CC, Garcia MC, Riera FR, Kenney RM. Use of testicular volume to predict daily sperm output in the stallion. In: Proceedings of 36th Annual Convention of the American Association of Equine Practitioners, 1990; pp. 15–21. 25. Thompson DL, Pickett BW, Squires EL, Amann RP. Testicular measurements and reproductive characteristics in stallions. J Reprod Fertil Suppl 1979;27:13–17. 26. Williams WL. Diseases of the Genital Organs of Domestic Animals, 3rd edn. Massachusetts: Ethel Williams Plimpton Publishing Co., 1943; p. 23. 27. Hayes HM. Epidemiological features of 5009 cases of equine cryptorchidism. Equine Vet J 1986;18:467–70. 28. Cox JE, Edwards GB, Neal PA. An analysis of 500 cases of equine cryptorchidism. Equine Vet J 1979;11:113–16. 29. Eik-Nes KB. Secretion of testosterone by the eutopic and the cryptorchid testes in the same dog. Can J Physiol Pharmacol 1966;44:629–33. 30. Arighi M, Bosu WTK. Comparison of hormonal methods for diagnosis of cryptorchidism in horses. Equine Vet Sci 1989;9:20–6. 31. Arighi M, Singh A, Bosu WTK, Horney FD. Histology of the normal and retained equine testis. Acta Anat 1987;129:127–30. 32. Cox JE. Factors affecting testis’ weight in normal and cryptorchid horses. J Reprod Fertil Suppl 1982;32:129–34. 33. Coryn M, DeMoor A, Bouters R, Vandeplassche M. Clinical, morphological and endocrinological aspects of cryptorchidism in the horse. Theriogenology 1981;16: 489–96. 34. Arighi M, Bosu WTK, Raeside JI. Hormonal diagnosis if equine cryptorchidism and histology of the retained testes. In: Proceedings of the 31st Annual Convention of the American Association of Equine Practitioners, 1985; pp. 591–602. 35. Johnson L, Neaves WB. Age-related changes in the Leydig cell population, seminiferous tubules and sperm production in stallions. Biol Reprod 1981;24:703–12. 36. Valdez H, Taylor TS, McLaughlin SA, Martin MT. Abdominal cryptorchidectomy in the horse using inguinal extension of the gubernaculum testis. J Am Vet Med Assoc 1979;174:1110–12. 37. Nalbandov AV. Reproductive Physiology, 2nd edn. San Francisco: Freeman, 1964; pp. 44–5. 38. Cour-Palais IJ. Spontaneous descent of testicle. Lancet 1966;I:1403–5. 39. Federman DD. Disorders of sexual development. N Engl J Med 1967;277:351–60. 40. Roberts SJ. Infertility in male animals. In: Roberts SJ (ed.) Veterinary Obstetrics and Genital Diseases, 2nd edn. Ann Arbor: Edwards Brothers Inc., 1971; pp. 752–826. 41. Cox JE, Redhead PH, Dawson FE. Comparison of the measurement of plasma testosterone and plasma estrogens for the diagnosis of cryptorchidism in the horse. Equine Vet J 1986;18:179–82. 42. Jones WE, Bogart R. Genetics of the Horse. East Lansing: Caballus, 1971; pp. 333–9.
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43. Wright JG. Laparo-orchidectomy in the horse with abdominal cryptorchidism. Vet Rec 1960;72:57–60. 44. Wright JG. The surgery of the inguinal canal in animals. Vet Rec 1963;75:1352–63. 45. Lowe JE, Higginbotham R. Castration of abdominal cryptorchid horses by a paramedian laparotomy approach. Cornell Vet 1969;59:121–6. 46. O’Connor JP. Rectal examination of the cryptorchid horse. Ir Vet J 1971;25:129–31. 47. Moore JN, Johnson JH, Tritschler LG, Garner HE. Equine cryptorchidism: presurgical considerations and surgical management. Vet Surg 1978;7:43–7. 48. Ryan PL, Friendship RM, Raeside JI. Impaired estrogen production by Leydig cells of the naturally retained testis in unilaterally cryptorchid boars and stallions. J Androl 1986;7:100–4. 49. Arthur GH. The surgery of the inguinal canal in animals. Vet Rec 1963;75:1365–6. 50. Jann HW, Rains JR. Diagnostic ultrasonography for evaluation of cryptorchidism in horses. J Am Vet Med Assoc 1990;196:297–300. 51. Schaumbourg MA, Farley JA, Marcoux M, Laverty S. Use of transabdominal ultrasonography to determine the location of cryptorchid testes in the horse. Equine Vet J 2006;38: 242–5. 52. Cox JE, Williams JW, Rowe PH, Smith JA. Testosterone in normal, cryptorchid and castrated male horses. Equine Vet J 1973;5:85–90. 53. Crowe CW, Gardner RE, Humbug JM, Nachreiner RF, Purohit PC. Plasma testosterone concentrations and behavior characteristics in geldings with intact epididymides. J Equine Med Surg 1977;1:387–90. 54. Amann RP, Ganjam VK. Effects of hemicastration or hCG: treatment on steroids in testicular vein and jugular vein blood of stallions. J Androl 1981;3:132–9. 55. Cox JE, Redhead PH. Prolonged effect of a single injection of human chorionic gonadotrophin on plasma
56.
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59. 60.
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62. 63. 64.
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67. 68.
testosterone and estrone sulphate concentrations in mature stallions. Equine Vet J 1990;22:36–8. Silberzahn P, Pouret EJM, Zwain I. Androgen and estrogen response to a single injection of hCG in cryptorchid horses. Equine Vet J 1989;21:126–9. Lindner HR. Androgens and related compounds in the spermatic vein blood of domestic animals. J Endocrinol 1961;23:171–8. Cox JE, Williams JH. Some aspects of the reproductive endocrinology of the stallion and cryptorchid. J Reprod Fertil Suppl 1975;23:75–9. Cox JE. Experiences with a diagnostic test for equine cryptorchidism. Equine Vet J 1975;7:179–83. Cox JE. Testosterone or estrone sulphate assay for the diagnosis of cryptorchidism. Equine reproduction III. Proc. 3rd Int. Symp. Eq. Reprod., Sydney. J Reprod Fertil Suppl 1982;32:625. Nyman MA, Geiger JA, Goldzieher JW. Biosynthesis of estrogen by the perfused stallion testis. J Biol Chem 1959;234:16. Bedrak E, Samuels LT. Steroid biosynthesis by the equine testis. Endocrinology 1969;85:1186. Pigon H, Lunaas T, Velle W. Urinary estrogens in the stallion. Acta Endocrinol 1961;36:131–40. Raeside JI. Seasonal changes in the concentration of estrogens and testosterone in the plasma of the stallion. Anim Reprod Sci 1978/79;1:205–12. Raeside JI. The isolation of estrone sulphate and estradiol17β sulphate from stallion testis. Can J Biochem Physiol 1969;47:811–15. Ganjam VK, Kenney RM. Androgens and estrogens in normal and cryptorchid stallions, J Reprod Fertil Suppl 1975;23:67–73. Cox JE. Blood test for equine cryptorchidism. Vet Rec 1982;110:211. Burba DJ. Theriogenology question of the month. J Am Vet Med Assoc 1996;15:1705–6.
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Developmental Abnormalities of the Male Reproductive Tract Mimi Arighi
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107.
Abnormalities of the Accessory Sex Glands Dickson D. Varner and James Schumacher
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108.
Abnormalities of the Ejaculate Regina M.O. Turner
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Abnormalities of the Penis and Prepuce James Schumacher and Dickson D. Varner
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Abnormalities of the Spermatic Cord James Schumacher and Dickson D. Varner
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Epididymal Abnormalities Aime K. Johnson and Charles C. Love
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Abnormalities of the Testicles Warren Beard
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Neoplasia of the Reproductive Tract John F. Edwards
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Chapter 106
Developmental Abnormalities of the Male Reproductive Tract Mimi Arighi
Congenital and developmental defects are defined as abnormalities of either structure or function present at birth. They are the result of disruptive events in embryogenesis or fetal development and can be either genetic or environmental in origin.
Anorchidism
Monorchidism
Ectopic testes
Monorchidism is defined as the complete absence of one testis, which may result from agenesis of the testis and mesonephric duct or a unilateral testicular vascular accident. It is reported to be rare in horses. The differentiation between agenesis and a cryptorchid with total testicular degeneration is difficult; however, agenesis is considered to be very rare. In most cases of congenital monorchidism, a vaginal process is present on the side where the testis is missing, leading one to believe that the lack of testis on that side was due to a severe degeneration of a previously existing testis while in utero as opposed to a complete agenesis of that testis. The other testis can be scrotal, inguinal, or abdominal in location. The diagnostic approach to monorchid and cryptorchid horses should be similar. A definitive diagnosis of monorchidism is based on history, behavior, physical examination, hormonal testing, and surgical exploration.1–8
Horses have been diagnosed with ectopic testes, but it is uncommon. An ectopic testis is one that has deviated from the normal pathway of descent and may lie alongside the penis cranial to the scrotum. This is much more common in cattle than in horses.6
Polyorchidism This abnormality has been reported in horses but is rare. It is described as more than two testes present in an animal. It is possible that true polyorchidism has not really been identified, but instead a cyst on the spermatic cord or an epididymis has been mistaken for another testis.6, 8
This condition has been described in horses but is very rare. It is the absence of all testicular tissue, and it can be differentiated from degeneration of both retained testes only by histopathology.7, 8
Testicular hypoplasia Hypoplastic testes are those that have failed to grow to a normal size. It occurs in cryptorchidism, and some intersex conditions. It can be unilateral or bilateral and can affect both scrotal and abdominal testes. The small size is due to a reduction in the amount of seminiferous epithelium. Otherwise, these testes are the same as a normal sized testis.9, 10 Suspected causes of this abnormality include transplacental infections and intoxications, zinc deficiency, hormonal insufficiency, impaired testicular descent, abnormal karyotype, or vascular disturbances. When the condition is bilateral, these animals are usually subfertile to sterile, owing to azoospermia, severe oligospermia, or dysfunctional spermatozoa. This condition is considered to be congenital and may be hereditary. This diagnosis should not be made before the horse is 2 years old; otherwise, this condition could be misdiagnosed in stallions that have delayed testicular growth.9, 10
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Fused testes There is a report of a miniature horse that was found to have two abdominal testes fused together at their caudal poles. The fused testes had separate epididymides and spermatic cords. On histological examination, there was no evidence of spermatogenesis, so this stallion was sterile.11
Congenital testicular tumors Testicular neoplasms in the horse can arise from germinal or non-germinal cells and are usually benign. Germinal neoplasms are the most common type and include teratomas, seminomas, and teratocarcinomas. Non-germinal tumors arise from stromal cells and include Sertoli and Leydig cell tumors.8 Teratoma This tumor is considered to be an embryonic tumor. Embryonic tumors are those that arise during embryonic, fetal, or early postnatal development from a particular organ rudiment or tissue while it is still immature. They are simultaneously neoplasms and malformations. Teratomas are the most common type of testicular tumor type detected in horses and are most commonly identified in retained testes; nonetheless, they are also uncommonly seen in scrotal testes (Figure 106.1). A teratoma contains tissue derived from cells from more than one of the three primary germ layers: ectoderm, mesoderm, and endoderm. They can be cystic-lined or can be solid masses composed of fibrous, adipose, nervous, cartilaginous and/or bony tissue. They can also contain hair and/or teeth (Figure 106.2). They have been called dermoid cysts
Figure 106.2 Hair and bone identified in a cut section of a teratoma in an abdominal testis.
by some people owing to their appearance. This type of tumor is rarely malignant in horses and an affected horse will generally show no clinical signs.6, 7, 12 Embryonic testicular carcinoma This tumor type is very rare in equine testes. They are composed of poorly differentiated epithelial tissue that produces α-fetoprotein. They have been known to metastasize to other organs in the body when they do occur. A teratocarcinoma, a neoplasm with features of both teratoma and embryonal carcinoma, has also been reported in horses.7, 12 Hamartomas Hamartomas have also been described in horses. They are benign vascular tumor-like nodules composed of an overgrowth of mature cells that normally occur in that area but one element predominates.12
Hydrocele A hydrocele is an excess of fluid within the lumen of the vaginal process, i.e., the vaginal cavity. If blood is mixed with the fluid, it is called a hematocele. This condition is sometimes congenital and this type of hydrocele is most commonly seen in Shire horses. It can be bilateral or unilateral, and the presence of the increased fluid around the testis on the affected side may cause atrophy of that testis and thus reduction in spermatogenesis. This condition must be differentiated from an inguinal hernia and a variocele.6
Inguinal/scrotal hernias Figure 106.1 A teratoma in a retained abdominal testis.
An inguinal/scrotal hernia is a protrusion of an organ, part of an organ, or other structure through the
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Developmental Abnormalities of the Male Reproductive Tract wall of the cavity normally containing it. Hernias can be direct (hernial contents outside the tunic) or indirect (hernial contents within the tunic). A hernia is composed of three parts: the ring (opening in the cavity wall through which the hernia occurs), the sac (which encloses the contents of the hernia), and the contents of the hernia. Hernias can be reducible or irreducible.6, 9,13–16 In the horse, congenital inguinal hernias are either unilateral or bilateral, are normally indirect, and normally form through the vaginal ring, which is the hernial ring. It is normally seen on the left side if unilateral and usually first appears at birth to 30 days of age. It is more common in Standardbreds and Draught horses and it is recommended that affected horses are castrated. The sac of the hernia is the vaginal tunic, and the contents are normally either intestine or mesentery.6, 9,13–16 Congenital inguinal hernias in the horse may be inherited. Most of the time, if a foal has a congenital inguinal hernia, spontaneous resolution can occur by 3–12 months of age if the hernial contents are manually reduced on a regular basis. If the inguinal hernia is secondary to trauma and is a direct hernia, then surgery is necessary to reduce the hernia and close the rent in the vaginal tunic.6, 9,13–16
Congenital abnormalities of the epididymis Epididymal abnormalities are rare in the stallion, but bilateral congenital hypoplasia of the distal corpus epididymis has been reported. This condition results in azoospermia despite normal libido and ejaculatory behavior. Congenital failure of the gubernaculum to regress to form the proper ligament of the testis or the ligament of the tail of the epididymis can result in cryptorchidism.17 Segmental aplasia of the mesonephric duct is an important, but rare, condition that has been reported in horses. It may be inherited and is usually unilateral. The part normally missing is the tail of the epididymis. Production of sperm continues and so spermatostasis occurs, which causes tubular dilation, sperm granulomas, pressure within the testis, and finally testicular atrophy. Hypertrophy of the opposite testis usually occurs secondary to this condition and it is normally recognized at the time of puberty.7
Penile/preputial abnormalities Congenital abnormalities of the penis in horses are rare, but include a short penis, retroverted penis, aplasia or dysgenesis of the corpus spongiosum glandis, aplasia of the prepuce or penis, hypospadias (in which the urethra fails to close on the ventral surface
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of the penis or in the perineal region), and epispadias (in which the urethra fails to close on the dorsal surface of the penis).6, 14, 18 Constriction or stenosis of the preputial ring is a narrowing of the preputial opening that inhibits the extrusion of the penis. It is normally a congenital disorder and clinical signs can show up at birth or may not be evident until months later. Clinical signs are usually consistent with phimosis.19 Congenital aplasia or hypoplasia of the prepuce has also been described in the horse. The animals appear to have paraphimosis but, in actuality, it is due to a lack of sufficient preputial tissue to retain the penis within the prepuce.19 Congenital abnormalities of the penis and prepuce are rare in the horse but, if seen, it is most likely to be in a male pseudohermaphrodite.20
Intersex Several manifestations of abnormal male external genitalia have been described. The male pseudohermaphrodite is the most common type of intersex in the horse. Penile-like tissues are seen as an exaggerated clitoris in association with normal- or abnormal-appearing vaginal structures. These horses often have variations of phenotypically female external anatomy but are genetically male and they have testicles which are almost always intra-abdominal. Male pseudohermaphrodites can demonstrate either stallion-like or nymphomanic mare behavior. Castration is normally suggested as the treatment of choice. True hermaphrodism has been reported in foals, but is rare. The clinical signs include a poorly defined penis and absence of a scrotum. Definitive diagnosis is based on the presence of ovotestes. Even rarer is the female pseudohermaphrodite, and these animals have ovaries and a chromosome make-up that is female but show male external genitalia.6, 7, 13, 18,19–22
References 1. Arighi M, Bosu WTK. Comparison of hormonal methods for diagnosis of cryptorchidism in horses. Equine Vet Sci 1989; 9: 20–6. 2. Cox JE, Redhead PH, Dawson FE. Comparison of the measurement of plasma testosterone and plasma estrogens for the diagnosis of cryptorchidism in the horse. Equine Vet J 1986; 18: 179–82. 3. Parks AH, Scott EA, Cox JE, Stick JA. Monorchidism in the horse. Equine Vet J 1989; 21: 215–17. 4. Strong M, Dart AJ, Malikides N, Hodgson DR. Monorchidism in two horses. Aust Vet J 1997; 75: 333–5. 5. Petrizzi L, Varasano V, Robbe D, Valbonetti L. Monorchidism in an appaloosa stallion. Vet Rec 2004; 155: 424–5. 6. Cox JE. Developmental abnormalities of the male reproductive tract. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Malvern: Lea & Febiger, 1993; pp. 895–904.
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7. Foster RA, Ladds PW. Male genital system. In: Maxie MG (ed.) Jubb, Kennedy, and Palmer’s Pathology of Domestic Animals, 5th edn. Philadelphia: Elsevier Saunders, 2007; pp. 565–619. 8. Turner RMO. Testicular abnormalities. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders/Elsevier, 2007; pp. 195–204. 9. Robertson JT, Embertson RM. Surgical management of congenital and perinatal abnormalities of the urogenital tract. Vet Clin North Am Equine Pract 1988; 4: 359–79. 10. Blanchard TL, Varner DD, Bretzlaff KN, Morris DL, Elmore RG. Male reproductive disorders (testicular aplasia and hypoplasia). In: Smith BP (ed.) Large Animal Internal Medicine, 2nd edn. St. Louis: Mosby, 1996; p. 1571. 11. Sherman K, Myhre G, Wells R. Abdominal fused testicles in a miniature horse: a case report. J Equine Vet Sci 1989; 9: 191–2. 12. Misdorp W. Congenital tumors and tumor-like lesions in domestic animals. 3. Horses: a review. Vet Q 2003; 25: 61–71. 13. Huston R, Saperstein G, Leipold HW. Congenital defects in foals. J Equine Med Surg 1977; 1: 146–61. 14. Adams R. The genital system. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 490–5
15. Spurlock GH, Robertson JT. Congenital inguinal hernias associated with a rent in the common vaginal tunic in five foals. J Am Vet Med Assoc 1988; 193: 1087–8. 16. Walker DF, Vaughan JT. Surgery of the testes, In: Bovine and Equine Urogenital Surgery. Philadelphia: Lea & Febiger, 1980; pp. 163–5. 17. Burns T. The epididymis. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders/Elsevier, 2007; pp. 171–3. 18. Walker DF, Vaughan JT. Surgery of the penis and prepuce. In: Bovine and Equine Urogenital Surgery. Philadelphia: Lea & Febiger, 1980; pp. 140–3. 19. Gaughan EM, Van Harreveld PD. Penile abnormalities. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders/Elsevier, 2007; pp. 220–1. 20. Vaughan JT. Penis and prepuce. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Malvern: Lea & Febiger, 1993; pp. 885–94. 21. Mittwoch U. Sex reversal in the horse: 2 sides of a common coin. Equine Vet J 1997; 29: 333–4. 22. Schumacher J, Vaughan JT. Surgery of the penis and prepuce. Vet Clin North Am Equine Pract 1988; 4: 473–91.
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Chapter 107
Abnormalities of the Accessory Sex Glands Dickson D. Varner and James Schumacher
Introduction The accessory genital glands of the stallion include the paired ampullae, the bi-lobed prostate gland, the paired seminal vesicles (also called vesicular glands), the indistinct disseminate urethral glands, and the paired bulbourethral glands (Figure 107.1). Each ampulla is evident as a prominent, 1- to 2-cm diameter, circumferential enlargement in the distal 10- to 15-cm (terminal) segment of the deferent duct. The paired seminal vesicles are thin-walled, elongated, slightly pyriform sacs that lie lateral to the ampullae. The base, or fundus, of each seminal vesicle lies within the abdominal cavity and is covered by the genital fold (peritoneum). Each seminal vesicle expands greatly during sexual stimulation owing to secretory products emitted into its lumen. These glands can reach 15–20 cm in length and 5 cm in diameter when fully distended. The duct of each seminal vesicle typically converges with the ipsilateral deferent duct to form a common ejaculatory duct that enters the lumen of the pelvic urethra through the seminal colliculus (Figure 107.2).1 The prostate gland of the stallion is a bi-lobed structure with a connecting isthmus. The lobes are situated lateral to the pelvic urethra and caudal to the corresponding vesicular glands. Each lobe is 5–8 cm long, 2–3 cm wide, and 1–2 cm thick. The isthmus is a thin, traverse band that courses over the dorsum of the pelvic urethra at the level of the seminal colliculus. The prostate gland has several ducts that enter the urethral lumen lateral to the seminal colliculus (Figure 107.2). The paired bulbourethral glands lie 6–8 cm caudal to the prostate gland and are situated on the dorsolateral surface of the pelvic urethra near the ischial arch. These ovoid glands are 3–4 cm long and 2–2.5 cm wide. Each bulbourethral gland has several
excretory ducts that enter the urethral lumen distal to the seminal colliculus and near the dorsal midline. The disseminate urethral glands of the stallion are poorly characterized but appear to be located along the distal aspect of the pelvic portion of the urethra and the proximal aspect of the penile portion of the urethra, as evidenced by small papillae of the excretory ducts entering the urethral lumen (Figure 107.2). Uncommonly, the accessory sex glands of the stallion can become diseased. Abnormalities of the urethral and bulbourethral glands have not been reported, and the authors have documented only one case of prostatic neoplasia, which was an extension of an ampullary adenocarcinoma. The most common abnormality of the accessory sex glands involves excessive accumulation of spermatozoa within the ampullar region of the ductus deferens. Seminal vesiculitis is also identified infrequently in stallions.
Plugged ampullae The size and glandular nature of the ampullae of the stallion provide a potential for excessive accumulation of spermatozoa, which occasionally become inspissated and block the luminae. This disorder is often termed ‘plugged ampullae’, although the terms ‘ampullar spermiostasis’, ‘ductal spermiostasis’, ‘ductal sperm accumulation’, and ‘rusty load’ have been used to describe this condition. With complete bilateral blockage, the stallion is azoospermic. Blockage can be sporadic, causing affected stallions to have poor spermatozoal morphology and motility owing to a high percentage of tailless heads. Densely packed spermatozoal casts may also be present in the semen (Figure 107.3). The condition probably originates in the terminal segment of the ductus deferens because the lumen of the ductus deferens narrows at the level of its communication with the seminal colliculus.1–3
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 107.1 The accessory genital glands of the stallion (dorsal view). a, genital fold; b, bladder; c, ampulla; d, seminal vesicle (vesicular gland); e, lobe of prostate gland; e´ , isthmus of prostate gland; f, pelvic urethra (enclosed by urethralis muscle); g, bulbourethral gland; h, retractor penis muscle. (From Varner et al.2 ). With permission from Elsevier.
Stallions with large testes appear to be most commonly affected by this disorder, probably owing to their increased rates of sperm production. Nonetheless, the condition has also been identified occasionally in stallions with average- to small-sized testes. Stallions with only one scrotal testis (owing to unilateral cryptorchidism or surgical removal of one testis) often undergo compensatory testicular hypertrophy, leading to an increased rate of sperm production in the single, scrotal testis and an increased probability of overloading the ipsilateral extragonadal duct with spermatozoa. Diagnosis is based on palpation and ultrasonographic examination of the ampullae per rectum, and characteristics of semen quality. Palpation may reveal enlarged and rigid ampullae, and ultrasonographic examination may reveal distension of the ampullar lumen with fluid that is either hypoechoic or of mixed echogenicity. The condition can be unilateral, but it usually involves both ampullae. Although the presence of tailless spermatozoal heads in high proportion is a classic feature of this disorder, an increased frequency of distal protoplasmic droplets or bent tails is also an indicator of prolonged storage of spermato-
Figure 107.2 Pelvic urethra, revealing ductal openings, as viewed by endoscopic approach. a, bladder opening; b, seminal colliculus; b , common opening of seminal vesicles and ampullae; c, opening of prostatic ducts; d, opening of urethral ducts; e, opening of bulbourethral ducts. (From Varner et al.2 ). With permission from Elsevier.
zoa within the extragonadal ducts. The incidence of this problem among stallions, as presented in varying degrees of severity, can be as high as 30–40% when the stallions are allowed extended periods of sexual abstinence. Occasionally, the condition may be severe enough to result in complete blockage of the ductal
Figure 107.3 Differential interference contrast image of a sperm clump in an ejaculate containing numerous tailless heads from a stallion with plugged ampullae.
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Abnormalities of the Accessory Sex Glands lumina and azoospermia. In such instances, measurement of the alkaline phosphate concentration in semen can be used to help differentiate between complete obstruction of the ampullar lumina (negligible or low alkaline phosphatase concentration) and azoospermia resulting from profound testicular dysfunction (high alkaline phosphatase concentration.4, 5 Treatment consists of repeated attempts at semen collection. This procedure can be preceded by extensive massage of the ampullae per rectum to aid in dislodging inspissated ampullar contents. Intravenous injection of oxytocin (10–20 IU) 2–5 minutes prior to an attempt at semen collection may also aid in evacuation of ampullar contents through contraction of the musculature enveloping the ductus deferens. Horses refractory to this treatment have responded favorably to intramuscular injection of cloprostenol (25–125 µg) 5–10 minutes before an attempt at semen collection. Horses with intractable plugging of the ampullae may require antegrade catheterization of the ductus deferens near the tail of the epididymis to relieve the obstruction using fluid pressure.6 Affected stallions are generally at increased risk for recurrence of the condition when they are allowed extended periods of sexual rest.
Seminal vesiculitis Stallions develop seminal vesiculitis, or vesicular adenitis, uncommonly, but seminal vesiculitis is a noteworthy disorder because of its persistent nature and interference with fertility. Reported bacterial etiological organisms include Pseudomonas aeruginosa, Klebsiella pneumoniae, Streptococcus spp., Staphylococcus spp., Brucella abortus, and Acinetobacter calcoaceticus. Although mycoplasmata, ureaplasmata, chlamydia, protozoa, and viruses have been associated with seminal vesiculitis in other species of animals, these microorganisms have not been isolated from inflamed seminal vesicles of horses. Although noninfectious seminal vesiculitis of stallions has not been described, we have identified air-filled seminal vesicles in a young Quarter Horse stallion in performance training. The precise pathogenesis of seminal vesiculitis remains an enigma. Potential routes for infection of the seminal vesicles include: (i) ascent through the urethra, (ii) descent from the upper portion of the genital or urinary tract, (iii) lymphogenous or hematogenous spread, or (iv) direct invasion from periglandular tissues. Seminal vesiculitis can be bilateral or unilateral. Although the infection is usually limited to the seminal vesicles, spread to the ampullae is possible. Urethritis is generally not identified in horses with seminal vesiculitis.1
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Characteristically, stallions with seminal vesiculitis do not manifest overt physical signs of impairment, although colic associated with the disorder has been reported.7 In general, affected stallions tend to be afebrile and do not exhibit signs of pain at the time of urination, defecation, or copulation. Presumptive diagnosis of seminal vesiculitis is usually made by gross and microscopic analysis of semen. During cytological examination, variable numbers of neutrophils may be seen. Grossly, the semen sample is generally discolored, and it may be blood, brown, or beige tinged. It may also be flocculent, with clumps of purulent material. Large numbers of bacteria may be detectable microscopically. The quality of spermatozoa may appear unaffected when semen is examined immediately after ejaculation, but longevity of spermatozoa is usually reduced. The appearance of discolored semen toward the end of an ejaculate suggests that the source of the disorder is the seminal vesicles.2 Determining the site of inflammation can be difficult when neutrophils are detected in semen, because neutrophils can arise from lesions involving the penile integument, urethra, upper portion of the urinary tract, deferent ducts, epididymides, testes, or any of the accessory genital glands. An internal source of infection should be suspected if no lesions are observed on the penile integument or urethral process. Bacterial types and numbers recovered from the exterior of the penis, urethra, and fractionated semen can be compared to help localize an infectious process. Culture swabs of the unwashed penis and preputial cavity should be obtained first to determine the resident bacterial flora in these areas. This step is important because many horses suspected of having seminal vesiculitis simply have potentially pathogenic bacteria on the integument of the penis and prepuce. Next, the penis should be washed with a bactericidal soap and thoroughly rinsed and dried in preparation for more bacterial culturing. The distal portion of the urethral lumen is then swabbed before and after the horse ejaculates. Semen is also swabbed for bacterial culture. Semen is collected using an artificial vagina that is free of bacteria prior to use to prevent possible contamination. A heavy, pure growth of bacteria in the semen and in the urethra after ejaculation is consistent with infection of the internal genitalia. Gross, microscopic, and microbial evaluation of semen fractionated into sperm-rich (first two or three seminal jets) and sperm-poor (later jets) portions can be used diagnostically to determine the source of infection.1 Both bacterial culture and cytological evaluation of expressed secretions of the vesicular glands aid in diagnosis of seminal vesiculitis. To cause the luminae
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of the vesicular gland to fill with secretions, the stallion is first stimulated sexually (teased) for approximately 15 minutes by exposing it to a mare in estrus. The distal portion of the penis is washed, rinsed, and dried. A sterile, 100-cm-long rubber catheter with an inflatable cuff is then passed into the urethra until its tip is immediately caudal to the seminal colliculus, where the ducts of the vesicular glands enter the urethra.8 Proper placement of the catheter tip is aided by palpation of the urethra and catheter per rectum. After the cuff has been inflated, each vesicular gland is identified by palpation per rectum, and the contents are removed by manually compressing the gland. Fluid can be collected for bacterial culture and cytological evaluation, using a sterile container attached to the opening of the catheter at the urethral orifice. Palpation per rectum of the seminal vesicles of stallions with seminal vesiculitis is often not rewarding diagnostically. Typically, no changes are evident in the size, shape, or consistency of the affected glands, although on occasion they may feel enlarged, firm, and lobulated. Pain may be elicited during palpation if the horse is acutely affected with seminal vesiculitis. Formation of abscesses in a vesicular gland of stallions has been reported, but occurs rarely. Such lesions may be detectable by palpation per rectum. The accessory genital glands, including the seminal vesicles, of normal stallions have been evaluated using transrectal ultrasonography.9 This procedure has been reported to be capable of detecting seminal vesiculitis through assessment of the echotexture of the vesicular luminae.10 We have used a flexible endoscope for definitive diagnosis of seminal vesiculitis in four stallions. A small-diameter (e.g., 7–9 mm) fiberoptic or video endoscope can be passed directly through the opening of the duct of the seminal vesicle at the seminal colliculus into the vesicle’s lumen. Care should be taken to prevent undue trauma to the opening of the ducts of the seminal colliculus when the endoscope is inserted into each vesicular gland. The luminae can be visualized directly to facilitate sampling for bacterial culture and cytology analysis. Culture instruments or catheters designed for endoscopes can be passed into position via the biopsy channel for acquisition of samples. Using this endoscopic approach through the urethra, the character of fluid expressed manually per rectum from the seminal vesicles, prostate gland, and bulbourethral glands can also be visualized. Treatment of stallions for seminal vesiculitis has generally consisted of systemically administered antimicrobial therapy, the selection of which is based on the results of sensitivity testing of bacterial cultures. Parenterally administered antimicrobial therapy tends to be unrewarding because
most antimicrobial drugs are incapable of reaching therapeutic concentrations in the luminae of the accessory genital glands. The characteristics of an antimicrobial drug that is capable of passing across the epithelial membranes of the accessory genital glands include: (i) high lipid solubility, (ii) low molecular size, (iii) low binding affinity with plasma proteins, and (iv) high pKa to increase the fraction of un-ionized antimicrobial drug. Unfortunately, few antimicrobial drugs possess these characteristics. Studies involving other species have revealed that trimethoprim–sulfonamide and enrofloxacin pass readily into the prostate gland; therefore, these antimicrobial drugs may be the antibiotics of choice for parenteral therapy of stallions with seminal vesiculitis caused by bacteria susceptible to these drugs. A stallion with seminal vesiculitis caused by enrofloxacin-susceptible A. calcoaceticus, however, was refractory to parenteral treatment for 15 days.11 An antimicrobial drug can sometimes be deposited directly into the seminal vesicles of affected stallions by passing a catheter via the urethra to the level of the seminal colliculus. By manipulating the catheter tip per rectum, the catheter can be directed blindly into each vesicular duct. If attempts at cannulating the ducts are unsuccessful, the orifice of the bladder can be occluded by compressing it with a hand in the rectum, and antimicrobial drug ejected from the pre-placed catheter in the proximal portion of the urethra. Transrectal ultrasonography is used to verify filling of the vesicular glands with the antimicrobial solution. We prefer to use an endoscopic approach for instillation of antimicrobial drugs into the seminal vesicles and have used a flexible endoscope for treatment of several stallions with seminal vesiculitis. Using this approach, each seminal vesicle is cannulated via the opening of its duct at the seminal colliculus by passing a catheter through the biopsy channel of an endoscope placed in the pelvic urethra so that the seminal colliculus can be easily viewed (Figures 107.4 and 107.5). The affected glands are first lavaged with a large quantity of normal saline solution or balanced salt solution, and then the appropriate antimicrobial drug, some of which must be buffered to prevent localized tissue reaction, is instilled directly into the luminae of the glands. This protocol is generally repeated daily for 5 consecutive days. This regimen of therapy, in most instances, resolves seminal vesiculitis. Because other accessory genital glands may be infected concomitantly and are not amenable to this therapeutic approach, successful treatment may be limited to horses with infection of only the seminal vesicles.12 Infected seminal vesicles have been surgically removed from bulls refractory to medical therapy. The
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Figure 107.4 Videoendoscopic view of the seminal colliculus demonstrating a catheter placed into a seminal vesicle for lavage of the gland’s lumen and instillation of antimicrobial drugs.
surgical procedure is rather difficult and can result in postoperative complications, including hemorrhage, infection, and damage to the intrapelvic nerves or muscles, preventing ejaculation. Using a surgical technique developed for use in bulls, an infected seminal vesicle was reportedly removed successfully from a stallion.13 Seminal
Figure 107.5 Interior of seminal vesicle, as viewed after placement of the insertion tube of a videoendoscope into the seminal colliculus and advancement into lumen of the gland. A catheter directed through the biopsy channel of the videoendoscope is being used to lavage the gland’s lumen with a balanced salt solution.
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vesiculectomy, when performed experimentally on reproductively normal stallions, does not suppress spermatozoal motility, and may actually improve this criterion. The percentage of morphologically normal sperm, however, has been reported to be reduced after seminal vesiculectomy.14, 15 Postsurgical complications described above for the bull are also applicable to the stallion. A surgical procedure found to be satisfactory for removal of the seminal vesicles of reproductively normal stallions16 may be an effective method for removing infected seminal vesicles of stallions refractory to medical treatment, but the technique, to our knowledge, has not been used clinically. For this technique, the horse is fasted for 2 days and then given mineral oil per os 24 hours before surgery to ensure low-bulk, soft stool in the early postsurgical period. The procedure is performed with the stallion sedated and standing in stocks. The seminal vesicles are catheterized and distended with fluid using a flexible endoscope, as described above. The perineal region is desensitized with caudal epidural anesthesia and appropriately draped. A sagittal incision is made along the caudal 10 cm of the ventral aspect of the rectum and through the anal sphincter and perineal body. Tissues beneath the rectum are dissected bluntly to the level of the seminal vesicles, and the rectal shelves are reflected to improve exposure of the surgical site. The fluid-distended seminal vesicles are identified and freed from the surrounding tissues using blunt and sharp dissection. The genital fold is penetrated and the abdominal cavity entered during the procedure. Each freed seminal vesicles is reflected caudally, ligated at its neck, and transected near the opening of the duct close to the pelvic urethra. During closure of the surgical site, the peritoneum (genital fold) is not repaired, but the other tissues are carefully apposed to prevent postsurgical infection.16 If seminal vesiculitis cannot be resolved, the stallion’s breeding life can sometimes be extended using minimum contamination breeding techniques. For artificial insemination programs, filtered semen can be placed in a semen extender containing antimicrobial drugs to which the bacteria from the vesicular glands are susceptible. An extended semen sample should be cultured for bacteria 15–30 minutes after it is exposed to the appropriate antimicrobial drugs. For natural service programs, semen extender (50–150 mL) can be instilled into the uterus immediately prior to breeding. These protocols have restored the fertility of a stallion with seminal vesiculitis caused by P. aeruginosa.17 Possible dissemination to the other accessory genital glands or locations within the excurrent duct system may occur.
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References 1. Varner DD, Schumacher J, Blanchard TL, Johnson L. Reproductive Anatomy and Physiology. In: Pratt PW (ed.), Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991, pp. 1–59. 2. Varner DD, Blanchard TL, Brinsko SP, Love CC, Taylor TS, Johnson L. Techniques for evaluating selected reproductive disorders of stallions. Anim Reprod Sci 2000;60–61:493– 509. 3. Love CC, Riera FL, Oristaglio-Turner RM, Kenney RM. Sperm occluded (plugged) ampullae in the stallion. In: Proceedings of the Annual Meeting of the Society of Theriogenology, San Antonio, Texas. August 14–15. 1992, pp. 117–27. 4. Turner RMO, McDonnell SM. Alkaline phosphatase in stallion semen: characterization and clinical applications. Theriogenology 2003;60:1–10. 5. Turner RM, Sertich PL. Use of alkaline phosphatase activity as a diagnostic tool in stallions with azoospermia and oligospermia. Anim Reprod Sci 2001;68(3–4):315–16. 6. Hughes JP. Significance of fertility evaluation of the stallion. In: Proceedings of the Annual Meeting of the Society of Theriogenology, Nashville, Tennessee. September 21–23. 1983, pp. 167–75. 7. Freestone JF, Paccamonti DL, Eilts BE, McClure JJ, Swiderski CE, Causey RC. Seminal vesiculitis as a cause of signs of colic in a stallion. J Am Vet Med Assoc 1993;203: 556–7. 8. Cooper WL. Methods of determining the site of bacterial infections in the stallion reproductive tract. In: Proceedings
9. 10.
11.
12.
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14.
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of the Annual Meeting of the Society of Theriogenology, Mobile, Alabama. September 13–14. 1979, pp. 1–4. Little TV, Woods GL. Ultrasonography of accessory sex glands in the stallion. J Reprod Fertil Suppl 1987;35:87–94. Malmgren L, Sussemilch B. Ultrasonography as a diagnostic tool in a stallion with seminal vesiculitis: a case report. Theriogenology 1992;37:935–8. Blanchard TL, Woods JA, Brinsko SP, Varner DD. Theriogenology. Question of the Month. J Am Vet Med Assoc 2002; 221:793–5. Blanchard TL, Varner DD, Hurtgen JP, Love CC, Cummings MR, Strezmienski PJ, Benson C, Kenney RM. Bilateral seminal vesiculitis and ampullitis in a stallion. J Am Vet Med Assoc 1988;192:525–6. Deegen E, Klug E, Lieske R, Freytag K, Martin J, Gunzel AR. Surgical treatment of chronic purulent seminal vesiculitis in the stallion. Dtsch Tierarztl Wochenschr 1979;86:140–4. ¨ Klug E, Deegen E, Liesk R, Freytag K, Martin JC, Gunzel AR, Bader H. The effect of vesiculectomy on seminal characteristics in the stallion. J Reprod Fertil Suppl 1979;27:61–6. Webb RL, Evans JW, Arns MJ, Webb GW, Taylor TS, Potter GD. Effects of vesiculectomy on stallion spermatozoa. J Equine Med Surg 1990;10:218–20. Taylor TS, Varner DD. Diseases of the accessory sex glands. In: Auer JA (ed.) Equine Surgery. Philadelphia: W.B. Saunders Co., 1992; pp. 723–7. Blanchard TL, Varner DD, Love CC, Hurtgen JP, Cummings MR, Kenney RM. Use of a semen extender containing antibiotic to improve the fertility of a stallion with seminal vesiculitis due to Pseudomonas aeruginosa. Theriogenology 1987;28:541–6.
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Chapter 108
Abnormalities of the Ejaculate Regina M.O. Turner
Pyospermia
Examination of the stallion
Pyospermia can be defined as the presence of purulent debris, including leukocytes, tissue debris, and/or pathogenic microorganisms, in the ejaculate. In humans, leukospermia (the presence of white blood cells in the ejaculate) is relatively common and is generally not considered to be a good marker for either the presence of pathological microorganisms or poor semen quality.1 In contrast, white blood cells are rarely seen in the ejaculate of normal stallions and, when present, typically indicate pathology. When present in large numbers, the problem can be identified grossly by the presence of purulent debris in the ejaculate. When present in lower numbers, the ejaculate may appear grossly normal. However, microscopic analysis of semen will readily permit the identification of leukocytes, usually together with tissue debris and an increased number of microorganisms. White blood cells, most often neutrophils, in the ejaculate of a stallion are indicative of inflammation in the reproductive or urinary tract. In some cases, spermatozoal motility may appear adequate even in the presence of large numbers of white blood cells. However, purulent debris will typically reduce longevity of spermatozoa and so reduce fertility of the stallion in many cases. Additionally, if an infectious cause underlies the inflammation, the potential for transmission of the infection to the mare during breeding or artificial insemination is present. Finally, even if antibiotics are added to the extended semen, the gross appearance of the ejaculate can be very disconcerting, particularly to mare owners or veterinarians preparing to inseminate client mares. When pyospermia is identified, the first approach to the problem is to identify the location and underlying cause of the inflammation. Appropriate treatment then can be instituted.
A careful examination of the external surface of the penis, the urethral fossa, and urethral process is indicated to determine whether a surface lesion is responsible for the problem. Swab samples of the external genitalia may be obtained for culture and cytological examination. Samples should be obtained from the unwashed surface of the penile shaft, the sheath, and the urethral fossa. However, these samples must be interpreted judiciously since a normal mixed population of microorganisms is present on the external genitalia. Normal flora of the stallion’s penis can include streptococci, staphylococci, coliforms including Escherichia coli, and Klebsiella, Pseudomonas spp., Corynebacterium spp., and occasionally yeast and fungi.2 The types of organisms present, as well as their growth patterns (pure growth vs mixed population) and numbers (heavy vs light growth), must all be taken into consideration. Pure, heavy growth of a suspected pathogen (e.g., Pseudomonas aeruginosa or Klebsiella pneumoniae) will warrant further investigation, while a mixed, light growth of a variety of commensal organisms is unlikely to be significant. It is important to keep in mind that, in the vast majority of cases, even suspected venereal pathogens such as Klebsiella and Pseudomonas typically do not result in clinical signs in the stallion.3 Thus, unless there is gross inflammation associated with the bacterial growth, the veterinarian should be suspicious of another possible source for the pyospermia other than the surface of the genitalia. However, these cultures often become more important if an infection is identified in the upper reproductive or urinary tract since pathogenic bacteria on the penile surface can serve as a source for ascending infection. Once the external surface of the genitalia has been sampled, the penis should be washed with clean water and dried with a clean cloth. Swab samples for culture and cytological examination can then be
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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obtained from the distal urethra both before and after ejaculation. Swab samples of semen for culture and cytological examination are also indicated. Heavy, pure growth of a microorganism from the pre-ejaculate urethra could be indicative of urethritis or could be evidence of cystitis. Heavy, pure growth of a microorganism from the post-ejaculate urethra and/or semen is more suggestive of an infection of the upper reproductive tract. Again, these cultures should be interpreted judiciously as one would expect growth of a mixed population of non-pathogenic bacteria from the urethra and ejaculate of normal stallions. If internal inflammation/infection is suspected, a urinalysis and urine culture may also be indicated to rule out cystitis. Contamination of the urethra with bladder contents can lead to pyospermia as a manifestation of cystitis. Free catch urine samples are not recommended since a lesion within the urethra could result in contamination of the sample and make definitive identification of the location of the inflammation more difficult. Additionally, free catch urine samples are more easily contaminated with nonspecific microorganisms that render interpretation of urine culture more difficult. Urinalysis and culture of an aseptically collected, catheterized urine sample generally facilitates interpretation of the results. The upper reproductive tract should also be examined if an internal infection is suspected. Although uncommon, seminal vesiculitis is the most frequently reported inflammatory/infectious pathology of the upper reproductive tract in stallions.4–10 Several cases of epididymitis have also been reported.11–17 Other reported sites of inflammation/infection include the ampullae, the testicles, and the pampiniform plexus.5, 13, 17 Transrectal ultrasonographic examination of the accessory sex glands and transcutaneous ultrasonographic examination of the scrotal contents can be an excellent diagnostic aid in localizing the problem. In severe cases, ultrasonography alone can lead to the identification of the location of the inflammation. In more mild cases, ultrasonographic findings can be interpreted in conjunction with results of culture, and physical and cytological examinations to arrive at a diagnosis. The ultrasonographic appearance of seminal vesiculitis, epididymitis, and orchitis are described in this text and elsewhere (see Chapter 151).13, 17, 18 Since different fractions of the stallion ejaculate contain fluid that is enriched for secretions from different parts of the reproductive tract, examination of each fraction individually can help the veterinarian better localize the source of the pyospermia. Swab samples from the whole ejaculate as well as samples from isolated fractions can be obtained for culture and for cytological examination.
r
r
r
r
Presperm fraction. If the penis is washed with warm water and the glans gently stimulated during washing, most stallions will emit a clear presperm fraction. This fraction largely contains secretions from the bulbourethral glands and will also serve to rinse the urethra.19 Identification of inflammatory cells in the presperm fraction therefore suggests inflammation of either the bulbourethral glands or urethra. Sperm-rich fraction. Approximately the first three jets of semen contain the sperm-rich fraction of the stallion ejaculate. The sperm-rich fraction is composed largely of fluid originating from the testes, epididymides, and ampullae with minimal contributions from the seminal vesicles, prostate, or bulbourethral glands. Identification of inflammatory cells in the sperm-rich fraction therefore suggests the presence of orchitis, epididymitis, or ampullitis.20 Sperm-poor fraction. The remaining jets of the ejaculate contain the sperm-poor fractions. These fractions are enriched with contributions from the seminal vesicles and prostate gland with lesser contributions from the testicles. Thus, identification of inflammatory cells in the sperm-poor fraction suggests the presence of seminal vesiculitis or prostatitis.20 Gel fraction. If the ejaculate is run through a filter, the gel fraction can be isolated. Since gel is contributed by the seminal vesicles,21 identification of inflammatory cells in this fraction suggests the presence of seminal vesiculitis. Alternatively, a sterile urinary catheter can be advanced through the urethra in a sexually stimulated (teased) stallion until it is just caudal to the seminal colliculus. The cuff is inflated and the vesicular glands can then be manually evacuated one at a time by gentle manual compression per rectum.22 If a sterile sample from the vesicular glands is needed, it can usually be obtained endoscopically. A small diameter (7–9 mm) endoscope can be passed through the urethra directly into the lumina of the vesicular glands. Once there, a sterile aspirate of the gland contents can be obtained.22 This technique can also be useful in treating diagnosed seminal vesiculitis by infusion of appropriate antibiotic preparations directly into the lumen of an infected gland.
Samples from the whole ejaculate and/or samples from the different semen fractions can be examined microscopically for the presence of inflammatory cells. Note that dismount semen samples collected following natural cover are not suitable for cytological analysis. These samples often normally contain large numbers of inflammatory cells and bacteria that originate from the mare’s vagina and therefore
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Figure 108.1 Microscopic appearance of cells consistent with white blood cells in a fixed, unstained sample of raw semen from a stallion with pyospermia. Black arrows point to the two cells whose appearance is consistent with that of inflammatory cells. A smear from this ejaculate was later stained and confirmed to contain large numbers of neutrophils. Note the large, homogeneous size of the cells in relation to the sperm. The third, smaller round cell in the frame is likely to be an immature spermatogenic cell based on its smaller size. Differential interference contrast image; 800× magnification.
cannot be used to diagnose inflammation of the stallion’s tract. In an unstained semen sample, inflammatory cells appear as large, round cells. Unlike immature spermatogenic cells, inflammatory cells typically are uniform in size (Figure 108.1). More definitive diagnosis of the somatic cell types present can be determined by examining stained samples. Swabs obtained for cytological examination should be gently rolled onto clean slides. Once the slides are dry, they can be stained with a Romanowsky stain such as Diff-Quik (International Medical Equipment, Inc., San Marcos, CA). In almost all cases, the inflammatory cells are neutrophils. These cells are characterized by their multilobulated nucleus with lobes connected by thin strands of nuclear material (Figure 108.2). If the underlying cause of the inflammation is infectious, large numbers of both phagocytized and extracellular bacteria will also often be seen on stained cytology samples. Cultures of samples from the whole ejaculate and/or from the different fractions can then be interpreted in light of the clinical cytology. Treatment Once pyospermia has been identified, the location of the inflammation confirmed, and the cause of the inflammation identified (e.g., infectious, traumatic, neoplastic), the problem can be treated accordingly. The exact treatment regimen and the prognosis will
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Figure 108.2 Microscopic appearance of neutrophils in a DiffQuik-stained smear of raw semen from a stallion with pyospermia. Note the dark-stained, multilobulated nuclei and the pale cytoplasm of the neutrophils. Significant numbers of extracellular chains of cocci are also present. Black arrow points to one of six neutrophils visible in this frame. Bright field image; 1000× magnification.
depend largely on the location and cause of the inflammation. Non-steroidal anti-inflammatory drugs typically are used. If the inflammation is bacterial in origin, systemic and/or topical antimicrobials are employed based on culture and sensitivity results.
Hemospermia Hemospermia is the presence of blood in the seminal fluid. The source of the blood can be anywhere in the reproductive tract, but most commonly originates from lesions in the urethra, on the urethral process, or on the penis itself. Depending on the nature of the lesion, the condition may be present in all ejaculates or may be intermittent. Etiology The causes of hemospermia in the stallion are varied. The condition can result from bleeding or inflammatory lesions anywhere in the reproductive tract from the testicles to the external surface of the penis. Some potential causes of hemospermia include epididymitis, blocked ampullae, seminal vesiculitis, urethral strictures, urethritis, urethral varicosities, neoplasia, and skin or mucosal surface lesions of the urethra, urethral process, glans penis, or penile shaft (Figure 108.3).4, 10,23–27 In one review of hemospermia in 18 stallions, the age of onset varied from 3 to 18 years. Although any breed can be affected, the condition may be more common in Quarter Horses.26 Trauma resulting in bleeding from the skin or mucosa of the reproductive tract is a relatively common cause of hemospermia. The urethral process is
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Stallion: Abnormalities of the Stallion’s Reproductive Tract liculus seminalis, rather than in the penile urethra. Viral urethritis has also been suspected as a possible cause of hemospermia.29 Clinical signs
Figure 108.3 Squamous cell carcinoma of the urethral fossa of a stallion (black arrow). Although the lesion is small, it is visibly ulcerated and this resulted in mild to moderate hemospermia. Because of its small size, the lesion was identified only after careful examination of the urethral fossa. It became more obvious if it was examined immediately after semen collection owing to the presence of fresh blood.
a common site of injury.22 Some of the more common traumatic injuries include kicks from a mare during breeding, lacerations of the penis or urethral process from tail hairs of the mare during breeding, and urethral strictures or skin lesions caused by the use of stallion rings. Iatrogenic traumatic hemospermia can occur following urinary bladder catheterization or urethroscopy. Idiopathic defects in the urethra that communicate with the underlying corpus spongiosum penis have been reported as relatively common causes of sometimes profuse hemospermia.25 These lesions can be quite small and most commonly are found near the pelvic ischium.22 Prior to the widespread use of ivermectin-based anthelmintics, cutaneous habronemiasis was a commonly reported cause of hemospermia. Lesions caused by Habronema are seen uncommonly in animals that are on proper anthelmintic programs. Stallions with blocked ampullae may exhibit hemospermia prior to or immediately following relief of the blockage.28 Discharge of chronically inspissated material from the ampullae may result in trauma to the ampullary mucosa and subsequent hemorrhage into the ejaculate. Infectious causes of hemospermia also have been reported. These include bacterial seminal vesiculitis, epididymitis, and urethritis. Additionally, active equine herpesvirus 3 infections can result in ulcerations on the skin surface of the penile shaft and glans penis that readily bleed during breeding. Bacterial urethritis has been reported as a relatively common cause of hemospermia in stallions. The most common organisms associated with bacterial urethritis include Streptococcus spp., E. coli, and Pseudomonas aeruginosa.26 This condition is seen more commonly in the pelvic urethra from the ischial arch to the col-
Infrequently, affected stallions may exhibit pain when they obtain a full erection or ejaculate.26 Stallions breeding by natural cover whose semen is not evaluated on a regular basis may present for infertility as it has been documented that significant whole blood contamination of the ejaculate will reduce pregnancy rates.30 The exact mechanism by which blood in the ejaculate reduces fertility is unclear as sperm motility, morphology, and numbers generally are not affected. One study suggested that the presence of red blood cells is responsible for the subfertility associated with hemospermia since 50% of subject mares bred with serum-contaminated semen became pregnant per cycle compared with only 7.7% of mares bred with whole blood-contaminated semen.30 In addition to infertility, blood may be seen at the mare’s vulva following breeding or it may be noted in the dismount sample.24 An accurate diagnosis of hemospermia generally can be made by collecting semen into an artificial vagina and observing blood in the ejaculate. Depending on the amount of blood contamination, the gross appearance of the ejaculate can range from pale pink (slight blood contamination) to that of frank blood (significant blood contamination). If the lesion communicates with the underlying cavernous tissue, bleeding can be quite dramatic.25 In more mild cases, microscopic examination of the ejaculate may be required to identify low numbers of red blood cells. Unless bleeding from a lesion is severe, signs typically are not seen when the stallion is at sexual rest. Hemospermia, like urospermia, can vary in severity from one ejaculate to another and may be sporadic. Thus, collection of a single, blood-free ejaculate does not rule out the problem. If hemospermia is suspected, multiple semen collections may be required to detect the condition. In particular, animals that are being examined following a period of sexual rest may not exhibit hemospermia in their first few ejaculates. Repeated ejaculations within a relatively narrow window of time (days) may be required to aggravate an old lesion sufficiently to cause recurrence. Diagnosis If hemospermia is suspected, semen should be collected in an artificial vagina and examined. Moderate to severe hemospermia can be diagnosed by gross inspection of the ejaculate. Milder hemospermia may require microscopic examination of the ejaculate. Microscopic examination of an unstained preparation
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Abnormalities of the Ejaculate
Figure 108.4 Microscopic appearance of red blood cells in a fixed, unstained sample of raw semen from a stallion with hemospermia. These cells were confirmed to be red blood cells after staining. In the unstained preparation, the cells appear round, homogeneous in size, and white and highly reflective. Note their small size in relation to the sperm. Black arrow points to one of four red blood cells visible in this frame. Differential interference contrast image; 800× magnification.
of semen will reveal the presence of round cells intermixed with the sperm. Red blood cells are much smaller than white blood cells and are homogeneous in size (Figure 108.4). Owing to their small size and homogeneous appearance, staining is usually not necessary. However, if in doubt as to the type of round cell present, a smear of the ejaculate can be prepared and stained with a Romanowsky-type stain (see Pyospermia). Red blood cells will stain a pale pink and lack a nucleus. When blood is indentified in the ejaculate, physical examination of the external surfaces of the penis, urethral fossa, and urethral process should be performed on both the flaccid and erect penis. Some superficial lesions are apparent on visual inspection (Figure 108.3). If a lesion is not identified, it may be helpful to collect semen and/or to fractionate semen using an open-ended artificial vagina. When appropriately filled, an open-ended artificial vagina allows the glans penis to protrude. This permits clear visualization of the glans during thrusting and during glans flare (‘belling’) and ejaculation. Some lesions that are not obvious in the flaccid penis become apparent when the cavernous tissues are engorged. Additionally, fractionation of the ejaculate may help to localize the source of the bleeding. Bleeding from the skin surface may be present in all jets of the ejaculate. Blood in only the first few jets could suggest a testicular or epididymal lesion (see Pyospermia) whereas blood in the later jets might suggest a problem with the accessory sex glands.
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If the skin, urethral fossa, or urethral process is not identified as the source of the bleeding, then other diagnostic techniques can be employed to examine higher areas of the reproductive tract. Palpation and ultrasonography of the scrotal contents may reveal abnormalities of the testes, epididymis, or spermatic cord. Transrectal palpation and ultrasonography of the internal accessory glands also help to identify pathology (e.g., blocked ampullae, seminal vesiculitis). Finally, urethroscopy with a flexible fiberoptic endoscope can be performed to fully examine the penile and pelvic urethra and to rule out the presence of urethritis or mucosal lesions that could be contributing to the hemospermia. Mucosal lesions can sometimes be quite small and difficult to identify. Performing the urethroscopic examination immediately following ejaculation may make the diagnosis more straightforward since blood may still be present in and around the lesion at this time.22 When performing urethroscopy, be aware that the procedure itself can readily cause iatrogenic urethral hyperemia. Contrast radiography has been suggested as another method of evaluating the urethra. Retrograde infusion of a contrast material into the urethra, followed by a series of radiographs, may help to identify strictures, masses, fistulae, and other similar urethral lesions.24 This method is used less commonly now that flexible endoscopy is readily available. Treatment The approach to treatment depends on the source of the hemorrhage, so accurate identification of the source of the bleeding is important. In this section, only the more general causes of hemospermia will be discussed. The reader is referred to other chapters for a discussion of related problems such as seminal vesiculitis, epididymitis, blocked ampullae, squamous cell carcinoma, and equine herpesvirus 3.3 Sexual rest is commonly employed as a nonspecific treatment for hemospermia arising from traumatic lesions, urethritis, or idiopathic urethral lesions. A minimum of several weeks, and often several months, of sexual rest may be required to allow the condition to heal. However, even with very prolonged periods of sexual rest, hemospermia may recur once the stallion returns to an active breeding program.26 Note that sexual rest should not include discouraging normal spontaneous erections and penile movements (also called masturbation). These actions are physiological and it has been well documented that aggressive methods of discouraging their occurrence can be harmful to the animal.31 As an alternative to conservative treatment with sexual rest alone, surgical approaches may be combined with sexual rest to help encourage resolution of hemospermia resulting from traumatic
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wounds or idiopathic lesions. Suturing of surface lacerations may be indicated to promote first intention healing. In the case of internal urethral lesions, we have attempted laser ablation of the ulcerated area with mixed results. A more aggressive method of treatment is to perform a temporary (approximately 4- to 6-week duration) subischial urethrotomy, thus allowing urine to bypass the distal aspects of the urethra entirely and also potentially reducing pressure in the corpus spongiosum penis at the end of urination. 25, 26 Temporary subischial urethrotomy may result in an increased likelihood of resolution of the problem than does sexual rest alone.26 However, this procedure is more costly and is associated with more sideeffects, including urethral strictures and fistulae.26, 32 Primary closure of these urethral lesions can be attempted if rest and subischial urethrotomy fail to resolve the problem. If the degree of hemospermia is mild, immediate extension of blood-contaminated ejaculates may be sufficient to allow pregnancies to occur.4 Alternatively, if a stallion is breeding by natural cover, extender can be placed into the mare’s uterus prior to breeding so that the stallion ejaculates into the extender and any blood contamination is diluted. These dilution techniques may not be sufficient in cases of more severe blood contamination. As a result of the underlying lesion, some animals may experience discomfort associated with ejaculation. If the pain is significant, it may eventually lead to ejaculatory problems and/or loss of libido. As such, animals exhibiting signs of pain in association with hemospermia may benefit from treatment with anti-inflammatory medications and other pain management modalities. Other reported treatments for hemospermia include intravenous formalin administration and urine acidification.29, 33 However, neither of these methods resulted in long-term cures for the hemospermia and so their use is questionable. Prognosis The prognosis for recovery, like the approach to treatment, varies based on the source of the hemospermia. Simple lacerations may heal with only rest. Puncture wounds may be more difficult to manage and may require surgical intervention. However, for traumatic lesions, the prognosis for full recovery is good if the animal is given an appropriate period of rest and wound care. If rest alone is insufficient, or if the wound is large, surgical debridement and closure may be required. Idiopathic ulcerated or rent-like lesions of the urethra can be more frustrating. Surgical correction and/or laser ablation combined with sexual rest may be helpful initially, but recurrence is possible once
the stallion is returned to active breeding. Temporary subischial urethrotomy will increase the chances of a permanent resolution; however, even when this treatment is used, recurrence is possible.
Urospermia Urospermia is defined as the presence of urine in the seminal fluid. The condition is reported infrequently in stallions as a whole.34 However, it appears to be fairly prevalent in stallions with ejaculatory problems since, in one report, 36% of all cases of ejaculatory dysfunction involved urospermia.35 Normal urine voiding and ejaculation Emission and ejaculation normally occur in a synchronized and controlled manner. In the normal stallion, emission occurs prior to ejaculation as the result of a thoracolumbar reflex arc that causes contraction of the smooth muscle of the ductus deferens and accessory glands. This same reflex arc is also responsible for the simultaneous contraction of the smooth muscle of the bladder trigone. As a result, ejaculatory fluids are discharged into the pelvic urethra and, at the same time, urine voiding is prevented. Both emission and bladder neck closure primarily involve α-adrenergic sympathetic fibers. Ejaculation follows emission and involves the forceful discharge of the seminal fluid anterograde through the urethra and out of the urethral orifice. Like emission, ejaculation is predominantly an α-adrenergic sympathetic event and is mediated primarily by a sacral reflex involving the pudendal nerve. The reflex initiates rhythmic contractions of the bulbocavernosus, ischiocavernosus, and urethralis muscles.35, 36 Sensory stimulation of the penis, usually together with psychic input from the brain, are the physiological triggers for the emission/ejaculation reflex. However, both emission and ejaculation can also occur without either sensory or psychic stimulation, as is the case with pharmacologically induced ejaculation.37, 38 Pathophysiology Any disease process that interferes with the synchronized reflex arc could result in urospermia, particularly if it interferes with reflex bladder neck closure. Neuritis of the cauda equina, equine herpesvirus 1, sorghum toxicosis, hyperkalemic periodic paralysis, and bacterial or sterile cystitis (as may be the case with urolithiasis) can all cause urospermia in stallions. Additionally, urospermia can arise secondarily due to fractures, osteomyelitis, and neoplasias that impinge on the nervous pathways of the lumbosacral region.34,39–42 With the exception of cystitis,
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Abnormalities of the Ejaculate these problems typically are not specific to the urinary tract and so stallions may demonstrate multiple neurological deficits in addition to urospermia. In addition to the above-listed causes, many cases of urospermia in stallions are idiopathic and the only presenting clinical sign is inappropriate urination during ejaculation. It seems likely that a nervous pathology that interferes with normal bladder neck closure during emission and ejaculation is to blame for many of these cases, but often the exact nature of the pathology cannot be identified. The result is inappropriate anterograde flow of urine through the urethra during ejaculation. In our clinic, animals with urospermia typically are monitored with 24-hour video surveillance in the stall. If carefully examined, many also demonstrate mild urinary incontinence and/or subtle bladder dysfunction (e.g., an inability to completely void the bladder during voluntary urination). Although less common than urospermia, retrograde ejaculation has also been reported in a stallion.43 This problem is relatively common in men but is an uncommon manifestation of bladder neck neuropathy in the horse. It seems likely that retrograde ejaculation and urospermia are different manifestations of similar problems. During retrograde ejaculation, semen passes in a retrograde fashion through the lax bladder neck and into the bladder, rather than moving anterograde through the penile urethra and urethral orifice.44 Stallions with retrograde ejaculation may show outward signs of normal ejaculation but then appear to produce little or no semen. A diagnosis can be easily made by obtaining a catheterized urine sample immediately following ejaculation. Although low numbers of sperm may be present in the urine of normal stallions (< 1 million sperm/mL), the urine of stallions experiencing retrograde ejaculation will contain much larger numbers.43 Regardless of the pathophysiology of urospermia or retrograde ejaculation, it has been documented that urine contamination of the equine ejaculate negatively affects sperm motility. This effect is most likely the result of changes in pH and osmolarity caused by the presence of urine. The more urine present in the ejaculate, the more profound will be its negative impact on sperm motility and subsequent fertility. Along these same lines, more concentrated urine is more detrimental to motility than is more dilute urine.45 Clinical signs and diagnosis Stallions breeding by natural cover whose semen is not evaluated on a regular basis may present for non-specific infertility.46, 47 A diagnosis of urospermia generally can be made by collecting semen into an ar-
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tificial vagina and observing gross urine contamination of the ejaculate. In more mild cases, the presence of urine in the semen may not be grossly apparent. However, even when urine contamination is not visibly evident, the presence of small amounts of urine will often alter the odor of the ejaculate and may increase its pH.48 Microscopic examination of the ejaculate may reveal a large number of urine crystals. Since sperm motility is severely adversely affected by the presence of urine,45 reduced sperm motility may be the most apparent problem in more mildly affected animals. In cases in which small, grossly undetectable amounts of urine are suspected to be contaminating the ejaculate, creatinine or urea nitrogen levels can be run on a sample of whole ejaculate. A creatinine level of over 2.0 mg/dL or a urea nitrogen level of over 30 mg/dL is suggestive of urospermia.49 These are simple tests that readily can detect the presence of even small amounts of urine contamination. As an alternative, commercially available test strips for the semiquantitative detection of urea nitrogen (Azostix, Miles, Inc.) or nitrite (Multistix, Miles, Inc.) can also be used. However, care must be taken when using these stall-side tests as false-positive results are common when the appropriate protocol is not strictly followed. Specifically, when using the Azostix test to detect urospermia, it is recommended that the reagent pad be rinsed with distilled water after only 10 seconds of horizontal exposure to the sample. After rinsing, the result can be read. A change in color of the test pad from yellow to green indicates the presence of urine. When using the Multistix test, the test pad should be placed in a horizontal position and exposed to the sample for 3.5 minutes before the test is read. A color change of the test pad from yellow to orange indicates the presence of urine.49 Urospermia is often intermittent and the amount of urine contamination can vary dramatically from ejaculate to ejaculate. Thus, collection of a single, urine-free ejaculate does not rule out the problem. If urospermia is suspected, collection and examination of multiple ejaculates, particularly when the stallion has a full bladder, may be required to arrive at a diagnosis. In cases where the stallion is breeding by natural cover, it may be possible to detect the presence of urine crystals in the mare’s uterus following breeding. When present in sufficient amounts, urine crystals appear as extremely echogenic accumulations within the mare’s uterus (A.O. McKinnon, personal communication, 2009). This can be used as a routine screening test as long as one keeps in mind that the sensitivity of this test is unlikely to be 100%, particularly when the volume of urine contamination is low.
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Stallion: Abnormalities of the Stallion’s Reproductive Tract
Observation and/or videotaping of the stallion in a box stall may also reveal related and subtler clinical signs. Affected animals may show evidence of neurological or infectious disorders of the urinary system including dysuria, polyuria, and/or stranguria. Additionally, videotaping of the stallion allows for ready evaluation of masturbation activity and penis control. Abnormalities in either of these areas may suggest the presence of either a localized or a systemic neurological problem involving the reproductive tract. Additional diagnostic tests Once a diagnosis of urospermia is made, it is recommended that the stallion have a full neurological examination, including examination of cranial nerve function, limb function, anal tone, anal reflex, tail tone, and defecation pattern and frequency. Standard neurological evaluations may reveal subtle deficits in other areas of the body, thus suggesting a more systemic cause for the urospermia. Additionally, the urinary tract should be carefully examined. Collection of a sterile urine sample via urinary catheter is typically indicated. Urinalysis and culture of the urine sample should be performed. Palpation and ultrasonographic examination of the bladder before and after voluntary urination may reveal an incomplete ability to fully evacuate the bladder contents. Additionally, the bladder wall and contents can be evaluated for lesions or for the presence of uroliths. An attempt can be made to express the bladder by palpation per rectum. The amount of pressure that is required to expel urine from the bladder can be used as an indirect measure of bladder sphincter and neck control. To evaluate the ability of the detrusor muscle to contract, bethanechol chloride, a cholinergic agent that normally causes evacuation of the bladder within 15 minutes of subcutaneous administration, can be given at a dose of 0.07 mg/kg bodyweight. Following urination, the bladder can be re-examined to determine whether complete evacuation has occurred. If a problem is identified, it should be addressed directly (e.g., systemic antibiotic administration for cases of bacterial cystitis) in the hope that correction of the underlying problem will improve or eliminate the urospermia. Treatment Limited therapeutic options are available for the treatment of idiopathic urospermia in stallions. And of these, none have been consistently effective. Treatment of urospermia often involves a commitment to long-term management changes that are designed to minimize the amount of urine present in the blad-
der at the time of semen collection or natural breeding and so decrease the likelihood of urine contamination of the ejaculate. Treatments addressed at correcting the suspected underlying cause(s) of the problem (e.g., a neurological condition) could, in theory, be more effective. However, since many cases of urospermia are idiopathic, this often is not an option. The simplest and most established method for the management of urospermia is to encourage the stallion to completely empty the bladder immediately prior to semen collection or natural breeding.50 This can usually be accomplished by moving the animal to a freshly bedded stall and allowing him to urinate voluntarily immediately before he is taken to the breeding shed. The addition of fresh manure from another stallion can also be placed in the stall with the subject stallion as an added stimulus to induce voluntary urination. As a final option, diuretics, such as furosemide, can be used to dilute the urine that is voided.51 In the author’s experience, these techniques can be quite helpful at reducing the frequency and severity of urospermia; however, they do not always result in a urine-free ejaculate. Many horses retain some variable amount of residual urine in the bladder after voluntary voiding that can still be expelled during ejaculation. Nonetheless, stimulating voluntary urination prior to ejaculation remains central to the management of urospermia. In animals in which voluntary voiding is not sufficient, or in cases in which a urine-free ejaculate must be obtained in a single session (e.g., for semen freezing), the stallion’s bladder can be completely emptied by passing a urinary catheter into the bladder to facilitate complete drainage.27 The bladder can then be lavaged with several liters of sterile physiological saline to insure that any residual bladder contents will not be harmful to the sperm should they be expelled during ejaculation. If a sufficient number of trained personnel are available, the bladder can be manually expressed completely per rectum following the final lavage and before the urinary catheter is removed.42 In highly tractable animals, the procedure can be performed in the breeding shed, after the stallion has obtained an erection, thus avoiding the need for sedatives or tranquilizers and allowing for semen collection to proceed as soon as the bladder is fully evacuated. This more extreme method of emptying the bladder can consistently result in urinefree ejaculation, but is not without risk. Repeated use of urinary bladder catheterization will predispose the stallion to urethritis and bacterial cystitis.27 The owners should be informed of the risk prior to using this technique and it may be advantageous to place the stallion on a prophylactic course of an
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Abnormalities of the Ejaculate appropriate antimicrobial if catheterization is to be performed routinely. Fractionation of the ejaculate can also aid in the management of urospermia. Stallions typically ejaculate in a series of seven or eight pulses of semen.52 Methods for separating an ejaculate by collecting individual pulses (fractions) have been described.53 Theoretically, by fractionating the ejaculate, it should be possible to collect only the urine-free portions of the ejaculate, thus avoiding the problem of urine contamination.27, 51, 54 However, in practice, the pattern of urine contamination can vary from one ejaculate to the next, even within the same stallion.54 Some ejaculates may be urine free, while, in other instances, urine contamination may precede, coincide with, or follow ejaculation. Although useful in some instances, the variable pattern of urine contamination seen in most stallions can make this approach frustrating. Depending on the amount of urine contamination present in an individual ejaculate, it may be possible to improve the longevity of sperm motility by immediately diluting the raw ejaculate with a standard semen extender. If the temperature of the extender is carefully controlled, semen extender can be placed directly into the collection bag of the artificial vagina so that the stallion ejaculates directly into the extender and semen is immediately diluted. In one study, the addition of semen extender to urinecontaminated semen restored sperm motility to values similar to uncontaminated control ejaculates.45 However, motility was followed only for 1 hour post contamination and no fertility trials were performed. It is likely that, even with immediate extension and with minimal urine contamination, the longevity of sperm motility in extended, urine-contaminated ejaculates will be reduced compared with urine-free ejaculates, particularly when urine contamination is moderate to heavy. Since bladder neck (trigone) closure in the stallion is predominantly an α-adrenergically controlled event,53 α-adrenergic agonists have been used to treat urospermia in the hope of increasing bladder neck tone and minimizing urine leakage during ejaculation. In general, these agents are not particularly useful in themselves.55 However, when combined with management practices aimed at evacuating the bladder prior to ejaculation, they may be useful adjunctive treatments. Examples of α-adrenergic agents that have been used to treat urospermia in stallions include imipramine hydrochloride (0.8 mg/kg administered orally 2–4 hours before semen collection), phenylpropanolamine (0.35 mg/kg orally, twice daily), and norepinephrine.42, 50 These agents must be used with caution and with careful
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attention to dosage. For example, in men, higher doses of imipramine have been implicated as causes of ejaculatory failure, erectile dysfunction, and impotence.56, 57 Other compounds that have been employed with limited or no success in the treatment of urospermia include bethanechol chloride (a muscarinic parasympathetic receptor agonist), flavoxate hydrochloride (a muscarinic parasympathetic antagonist), oxytocin (to aid bladder neck constriction), and β-blocking agents.27, 50, 55, 58 Pharmacological induction of ejaculation38 has been suggested as a possible means of obtaining a urine-free ejaculate.42 Ejaculation induced in this manner does not involve an increase in abdominal or intravesicular pressure and thus may minimize pressure on the urethral sphincter and bladder neck at the point of ejaculation. This has not been tested critically as a treatment for urospermia. When breed registries permit, cryopreservation of semen should be considered for affected animals. One reasonable management technique is to utilize more aggressive treatment modalities (e.g., bladder catheterization, lavage, α-adrenergic agonists) for a period of time sufficient to collect and cryopreserve a large bank of urine-free semen. Mares can then be bred when needed with frozen–thawed semen without the need for the stallion to produce a urine-free ejaculate on short notice or when labor or time is in short supply.
Prognosis In cases in which a cause for the urospermia can be identified, the prognosis for its resolution is related to the prognosis for the treatment and cure of the underlying cause. The prognosis for complete resolution of idiopathic urospermia is poor. In most cases, the condition is intermittent but chronic. Because the underlying pathophysiology of the disease is not clear, treatments are generally palliative, rather than curative. However, with meticulous management, and with a willingness to sometimes employ more aggressive treatment modalities (e.g., bladder catheterization and lavage), good pregnancy rates can be achieved even with severely affected stallions. Owners and managers should be aware that these management changes typically are long term and, even with ideal management, the condition can be very frustrating. In animals with more widespread neurological problems, the clinical signs may progress over time if the underlying problem cannot be identified and treated.
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References 1. Rodin DM, Larone D, Goldstein M. Relationship between semen cultures, leukospermia, and semen analysis in men undergoing fertility evaluation. Fertil Steril 2003;179 Suppl 3:1555–8. 2. Kenney RM, Hurtgen JP, Pierson RH, Witherspoon D, Simons J. Clinical fertility evaluation of the stallion. J Soc Theriogenol 1983;9:7–62. 3. Blanchard TL, Kenney RM, Timoney P. Venereal disease. Vet Clin North Am 1992;8:191–203. 4. Blanchard TL. Use of a semen extender containing antibiotics to improve the fertility of a stallion with seminal vesiculitis due to Pseudomonas aeruginosa. Theriogenology 1987;28:541–6. 5. Blanchard TL, Varner DD, Hurtgen JP, Love CC, Cummings MR, Strezmienski PJ, Benson C, Kenney RM. Bilateral seminal vesiculitis and ampullitis in a stallion. J Am Vet Med Assoc 1988;192:525–6. 6. Deegan E, Klug E, Lieske R, Freytag K, Martin J, Gunzel A-R. Surgical treatment of chronic purulent seminal vesiculitis in the stallion. Dtsch Tierarztl Wochenschr 1979;86:140–4. 7. Freestone JF, Paccamonti DL, Eilts BE, McClure JJ, Swiderski CE, Causey RC. Seminal vesiculitis as a cause of signs of colic in a stallion. J Am Vet Med Assoc 1993;203:556–7. 8. Holst W. Seminal vesiculitis in two stallions. Tijdschr Diergeneeskd 1976;101:375–7. 9. Malmgren L, Sussemilch BI. Ultrasonography as a diagnostic tool in a stallion with seminal vesiculitis: a case report. Theriogenology 1992;37:935–8. 10. Sojka JE, Carter GK. Hemospermia and seminal vesicle enlargement in a stallion. Compend Contin Educ Pract Vet 1985;7:S587–8. 11. Blue MG, McEntee K. Epididymal sperm granuloma in a stallion. Equine Vet J 1985;17:248–51. 12. Brinsko SP, Varner DD, Blanchard TL, Relford RL, Johnson L. Bilateral infectious epididymitis in a stallion. Equine Vet J 1992;24:325–8. 13. Gonzalez M, Tibary A, Sellon DC, Daniels J. Unilateral orchitis and epididymitis caused by Corynebacterium pseudotuberculosis in a stallion. Equine Vet Educ 2008;20:30–6. 14. Held JP, Adair S, McGavin MD, Adams WH, Toal R, Henton J. Bacterial epididymitis in two stallions. J Am Vet Med Assoc 1990;197:602–4. 15. Held JP, McCracken MD, Toal R, Latimer F. Epididymal swelling attributable to generalized lymphosarcoma in a stallion. J Am Vet Med Assoc 1992;201:1913–15. 16. Traub-Dargatz JL, Trotter GW, Kaser-Hotz B, Bennett DG, Kiper ML, Veeramachaneni DN, Squires E. Ultrasonographic detection of chronic epididymitis in a stallion. J Am Vet Med Assoc 1991;198:1417–20. 17. Wilson KE, Dascanio J, Duncan R, Delling U, Ladd SM. Orchitis, epididymitis and pampiniform phlebitis in a stallion. Equine Vet Educ 2007;19:239–43. 18. Turner RM. Ultrasonography of the genital tract of the stallion. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia: W.B. Saunders, 1998; pp. 446–79. 19. Setchell BP. Male reproductive organs and semen. In: Cupps PT (ed.) Reproduction in Domestic Animals. New York: Academic Press, 1991; p. 221. 20. Love CC. Semen collection techniques. Vet Clin North Am Equine Pract 1992;8:111–28. 21. Mann T, Luwak-Mann C. Male Reproductive Function and Semen. Berlin: Springer-Verlag, 1981.
22. Varner DD, Blanchard TL, Brinsko SP, Love CC, Taylor TS, Johnson L. Techniques for evaluating selected reproductive disorders of stallions. Anim Reprod Sci 2000;60–61:493–509. 23. Bedford SJ, McDonnell SM, Tulleners E, King D, Habecker P. Squamous cell carcinoma of the urethral process in a horse with hemospermia and self-mutilation behavior. J Am Vet Med Assoc 2000;216:551–3. 24. McKinnon AO, Voss JL, Trotter GW, Pickett BW, Shideler RK, Squires EL. Hemospermia of the stallion. Equine Pract 1988;10:17–23. 25. Schumacher J, Varner DD, Schmitz DG, Blanchard TL. Urethral defects in geldings with hematuria and stallions with hemospermia. Vet Surg 1995;24:250–4. 26. Sullins KE, Bertone JJ, Voss JL, Pederson SJ. Treatment of hemospermia in stallions: A discussion of 18 cases. Comp Cont Educ Pract Vet 1988;10:1396–403. 27. Varner D, Schumacher J, Blanchard T, Johnson L. Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991. 28. Love CC, Riera FL, Oristaglio (Turner) RM. Sperm occluded (plugged) ampullae in the stallion. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1992, pp. 117–25. 29. Voss JL, Pickett BW. Diagnosis and treatment of hemospermia in the stallion. J Reprod Fertil Suppl 1975;23:151–4. 30. Voss JL, Pickett BW, Shideler RK. The effect of hemospermia on fertility in horses. In: Proceedings of the 8th International Congress of Animal Reproduction and Artificial Insemination, 1976, pp. 1093–5. 31. McDonnell SM, Henry M, Bristol F. Spontaneous erection and masturbation in equids. In: Proceedings of the 5th International Symposium on Equine Reproduction, 1991, pp. 664–5. 32. Laverty S, Pascoe JR, Ling GV, Lavoie JP, Ruby AL. Urolithiasis in 68 horses. Vet Surg 1992;21:56–62. 33. Voss JL, Wotowey JL. Hemospermia. In: Proceedings of the 18th Annual Convention of the American Association of Equine Practitioners, 1973, pp. 103–12. 34. Blanchard TL, Varner DD, Bretzlaff KN. Diseases of the reproductive system: male reproductive disorders. In: Smith B (ed.) Large Animal Internal Medicine. St. Louis: CV Mosby Co., 1990; p. 1424. 35. McDonnell SM. Ejaculation. Physiology and dysfunction. Vet Clin North Am Equine Pract 1992;8:57–70. 36. Guyton AC. Textbook of Medical Physiology. Philadelphia: W.B. Saunders, 2000. 37. McDonnell SM. Oral imipramine and intravenous xylazine for pharmacologically-induced ex copula ejaculation in stallions. Anim Reprod Sci 2001;68:153–9. 38. McDonnell SM, Love CC. Xylazine-induced ex copula ejaculation in stallions. Theriogenology 1991;36:73. 39. Mayhew IG. Neurologic aspects of urospermia in the horse. Equine Vet Educ 1990;2:68–9. 40. McClure JJ. Paralytic bladder. In: Robinson ED (ed.) Current Therapy in Equine Medicine 2. Philadelphia: W.B. Saunders, 1987, pp. 712–13. 41. Naylor J, Nickel DD, Trimino G, Card C, Lightfoot K, Adams G. Hyperkalemic periodic paralysis in homozygous and heterozygous horses: a co-dominant genetic condition Equine Vet J 1999;31:153–9. 42. Turner RM, Love CC, McDonnell SM, Sweeney RW, Twitchell ED, Habecker PL, Reilly LK, Pozor MA, Kenney RM. Use of imipramine hydrochloride for treatment of urospermia in a stallion with a dysfunctional bladder. J Am Vet Med Assoc 1995;207:1602–6.
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Abnormalities of the Ejaculate 43. Brinsko SP. Retrograde ejaculation in a stallion. J Am Vet Med Assoc 2001;218:551–3. 44. Lakin MM. Diagnostic assessment of disorders of male sexual function. In: Montague DK (ed.) Disorders of Male Sexual Dysfunction. Chicago: Yearbook Medical, 1988; p. 26. 45. Griggers S, Paccamonti D, Thompson RA, Eilts BE. The effects of pH, osmolarity and urine contamination on equine spermatozoal motility. Theriogenology 2001;56:613–22. 46. Danek J, Wisniewski E, Krumrych W. A case of urospermia in a stallion. Med Weter 1994;50:129–31. 47. Lowe JN. Diagnosis and management of urospermia in a commercial Thoroughbred stallion. Equine Vet Educ 2001;13:4–7. 48. Hurtgen J. Evaluation of the stallion for breeding soundness. Vet Clin North Am 1992;8:149–68. 49. Althouse GC, Seager SWJ, Varner DD, Webb GW. Diagnostic aids for the detection of urine in the equine ejaculate. Theriogenology 1989;31:1141–8. 50. Leendertse IP, Asbury AC, Boening KJ, Saldern FC. Successful management of persistent urination during ejaculation in a Thoroughbred stallion. Equine Vet Educ 1990;2:62–4.
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51. Hurtgen J. Stallion genital abnormalities. In: Robinson ED (ed.) Current Therapy in Equine Medicine 2. Philadelphia: W.B. Saunders, 1987, p. 561. 52. Tischner M, Kosiniak K, Bielanski W. Analysis of the pattern of ejaculation in stallions. J Reprod Fertil 1974;41: 329–35. 53. Ellery JC. A modified equine artificial vagina for the collection of gel-free semen. J Am Vet Med Assoc 1971;158:765–6. 54. Nash Jr JG, Voss JL, Squires EL. Urination during ejaculation in a stallion. J Am Vet Med Assoc 1980;176:224–7. 55. Hoyos Sepulveda ML, Quiroz Rocha GF, Brumbaugh GW, Montiel J, Rodriguez S, Candanosa de Morales E. Lack of beneficial effects of bethanechol, imipramine or furosemide on seminal plasma of three stallions with urospermia. Reprod Domest Anim 1999;34:489–93. 56. Segraves RT. Sexual side-effects of psychiatric drugs. Int J Psychiatry Med 1988;18:243–52. 57. Segraves RT. Effects of psychiatric drugs on human erection and ejaculation. Arch Gen Psychiatry 1989;46:275–84. 58. Voss JL, McKinnon AO. Hemospermia and urospermia. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 864–70.
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Chapter 109
Abnormalities of the Penis and Prepuce James Schumacher and Dickson D. Varner
Penile and preputial lacerations Etiology Trauma to the penis or prepuce is the greatest indication for preputial and penile surgery.1 The penis or prepuce can be traumatized when the horse fails at jumping a barrier, during copulation from the mare’s tail hairs or a loosely tied breeding stitch, from contact with a cracked cover on a phantom, and from frost bite.1–4 Application of an undersized stallion ring can produce an erosion of the penile integument,1 and the shaft can be severely lacerated during castration, especially when one of the testes is located inguinally and the surgeon is inexperienced (Figure 109.1).5, 6 Most penile or preputial lacerations or erosions involve only the integument, but some extend into cavernous tissue or the urethra.5–9 Those that extend into the cavernous tissue produce severe hemorrhage during copulation or at the end of urination, and those that extend into the urethra cause necrosis of surrounding tissue from urine escaping through the laceration.5, 6, 8 Laceration of the urethra is invariably accompanied by laceration of the corpus spongiosum penis (CSP) because the CSP completely surrounds the urethra. Pathophysiology A preputial laceration or erosion, when left unattended, may become infected, and, because preputial areolar connective tissue is loosely adhered to the penis, infection has a tendency to migrate resulting in generalized cellulitis and inflammatory edema. When inflammatory edema becomes extensive, the penis and internal preputial lamina protrude through the preputial orifice causing gravitational edema, which perpetuates the swelling. Swelling may be so
severe that the horse becomes unable to retract its penis. Most penile and preputial wounds, when properly attended, heal without complication, but contraction of a large preputial wound may result in a cicatrix that restricts the normal telescoping action of the internal preputial lamina. Treatment Penile and preputial wounds should be debrided, lavaged, and sutured using either absorbable or nonabsorbable sutures. A laceration that appears to be infected should be cleansed and covered with an antimicrobial cream or ointment several times daily until its condition is suitable for delayed primary closure. If the wound is allowed to heal by second intention, it should be treated with a topically applied antimicrobial ointment or cream at least until it begins to develop granulation tissue. A laceration of the urethra is accompanied by laceration of the CSP and its overlying tunic, and, so, both the urethra and the lacerated tunica albuginea of the CSP should be sutured. Stenting the urethral lumen with a male urinary catheter simplifies suturing of the urethra and the tunica albuginea of the CSP. An unsutured, longitudinal, urethral laceration is likely to heal without stenosis, but an obstructing stricture is the likely outcome when a transverse urethral laceration is left to heal by second intention. If a laceration of the penis or internal preputial lamina is accompanied by severe preputial swelling, the penis and swollen prepuce should be supported against the abdomen or retained in the preputial cavity to facilitate venous and lymphatic circulation and to prevent the penis and prepuce from incurring more damage. Administration of a non-steroidal, anti-inflammatory drug is indicated to decrease inflammatory edema.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Abnormalities of the Penis and Prepuce
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is permitted to breed an improperly restrained mare or is pastured with other horses. A penile hematoma can also develop when a corporeal body or a subfascial vessel on the surface of the penis is torn when the penis is suddenly bent during a copulatory thrust. Penile hematomas usually arise from rupture of the extensive vascular plexus located subfascially on the surface of the penis, but a hematoma can sometimes occur from a tear of the tunica albuginea surrounding the corpus cavernosum penis (CCP) or CSP. 4, 10, 11 A penis whose CSP or CCP has ruptured is sometimes referred to as a fractured, broken, or ruptured penis.
Figure 109.1 The penis of this horse was transected at the scrotum during castration.
Partial phallectomy may be the most expedient means of treating a horse that has suffered complete urethral disruption distal to the preputial cavity, especially if the injured horse is a gelding.9 If the laceration into the urethra is caudal to the preputial cavity, a permanent urethrostomy can be created at the site of injury (J. Schumacher and D. Varner, personal observation).6 If the urethra is severely damaged during castration, the permanent urethrostomy can be created at the scrotal wound (Figure 109.2).
Penile hematomas Etiology A penile hematoma is most often caused by a blow to the penis and internal preputial lamina when the penis is erect and is most likely to occur when a stallion
Pathophysiology Tearing of the subfascial, vascular plexus or a corporeal body forces blood into the loose preputial fascia, and, within minutes of injury, preputial swelling may become so extreme that the horse is incapable of retracting its penis into the preputial cavity. Or, the horse may be unable to protrude its penis, if the penis, as it swells, resides within the preputial cavity. The hematoma may obstruct undamaged veins and lymphatic vessels, exacerbating the swelling,2 and the hematoma may interfere with urination by impinging on the urethra, especially if the hematoma has arisen from rupture of the portion of the tunica albuginea that surrounds the urethra.12 Rupture of the tunica albuginea may result in impotence caused by a vascular shunt created between the CCP and the subfascial vascular plexus on the surface of the penis. A hematoma confined within the CCP of a stallion’s penis by an intact tunica albuginea can result in deviation during erection, presumably by impeding blood flow through the corporeal tissue.13 Treatment
Figure 109.2 A new urethral opening (arrows) was created at the scrotum of this horse whose penis was transected during castration. This is a different horse than that seen in Figure 109.1.
A horse should be treated as soon as possible after a penile hematoma is recognized, and treatment should be directed toward minimizing hemorrhage and edema by applying a tight bandage to compress the internal preputial lamina against the underlying tunica albuginea, or the penis can be maintained against the abdomen with a bandage that encompasses both the penis and the abdomen. If the hematoma and edema continue to expand despite these measures, the tunica albuginea should be examined ultrasonographically, radiographically (using cavernosography), or surgically for a rent that extends into corporeal tissue. Suturing the rent stops progression of the hematoma and prevents formation of a shunt between corporeal tissue and the dorsal veins of the penis. If the horse is unable to
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retract its penis into the preputial cavity after several days when the likelihood of hemorrhage into fascial planes has diminished, the horse’s penis and internal preputial lamina should be retained within the preputial cavity with a retainer bottle created by removing the wide end of a plastic 500-mL bottle, which is suspended within the preputial cavity using a crupper and surcingle made of rubber tubing. Nylon netting or hosiery can also be used instead of a bottle. The penis and internal preputial lamina can be maintained within the preputial cavity indefinitely, without damage to the prepuce, using these devices. To avoid resumption of hemorrhage, a stallion that has incurred a penile hematoma should be isolated for a week to prevent sexual arousal. Sexual arousal after this time is unlikely to exacerbate hemorrhage and may prevent formation of adhesions between the penis and the damaged, loose areolar tissue.14 Exercise of affected horses should also be restricted for a week. Exercise initiated after this time, when hemostasis is assured, may decrease preputial edema.
Paraphimosis Etiology Paraphimosis, or the inability of the horse to retract its protruded penis into the preputial cavity, can develop from a variety of causes, the most common of which is edema of the internal preputial lamina resulting from preputial trauma (see Penile hematomas). Paraphimosis may accompany some systemic diseases, such as dourine and purpura hemorrhagica, that produce extensive subcutaneous edema that often affects the male external genitalia,2, 15 or it may accompany diseases that sometimes have a deleterious effect on penile innervation, such as equine herpes virus 1 and rabies.2, 16 Paraphimosis has also been reported as a complication of severe debilitation17 or severe exhaustion.18 Priapism, a persistent erection without sexual excitement that usually occurs in horses after administration of a phenothiazine-derivative tranquilizer, results in paraphimosis not only because the persistent erection prevents retraction of the penis but also because the persistent erection eventually produces penile paralysis (see Priapism).19, 20 Paraphimosis caused by penile paralysis unassociated with priapism can also occur as a side-effect of phenothiazine-derivative tranquilizers, most notably propiomazine (formerly termed propiopromazine).21 Penile paralysis has also occurred after administration of acepromazine, another phenothiazinederivative tranquilizer.1, 22
Pathophysiology The penis is maintained within the preputial cavity by the tonus of the retractor penis muscles and the smooth muscle of the cavernous spaces.17 Injury to the internal preputial lamina causes the loose areolar connective tissue between the penis and internal preputial lamina to become edematous, and the weight of edema produces muscular fatigue, which, in turn, causes the penis and internal preputial lamina to protrude from the preputial cavity. The association between paraphimosis and exhaustion or debilitation is obscure, but either may decrease muscular tonus, allowing the penis to protrude.17 Regardless of the cause of protrusion, blood pools and clots in the CCP within 2–5 hours, making the penis appear slightly rigid, rather than flaccid.23 This rigidity may cause the clinician to conclude erroneously that protrusion of the penis is caused by priapism. Chronic protrusion, regardless of the cause, may cause the pudendal nerves to become contused or stretched at the ischial arch, resulting in penile paralysis.19, 24, 25 The mechanism by which phenothiazinederivative tranquilizers produce penile paralysis is not understood, but the action of these tranquilizers on the smooth muscles of the penis is probably responsible for penile protrusion. Phenothiazinederivative tranquilizers likely exert their influence on penile position by blocking the α-adrenergic motor fibers that innervate the retractor penis muscles and those that innervate the smooth muscle of the cavernous tissue, allowing the cavernous tissue to dilate with blood.26 Decreased tonus of the retractor penis muscles and filling of the cavernous tissue with blood cause the penis to fall from the preputial cavity. Paralysis may ensue when prolonged protrusion results in damage to the pudendal nerves. Regardless of its initiating cause, paraphimosis becomes a self-perpetuating state because prolonged protrusion itself impairs venous and lymphatic drainage, thereby causing or exacerbating preputial edema. Swelling of the internal preputial lamina from edema causes the relatively inelastic preputial ring to act as a constricting band that further impairs venous and lymphatic drainage distal to it. The swollen penile and preputial epithelium, exposed to the elements and made fragile by edema, soon becomes damaged and inflamed, and defects in the epithelium allow bacteria to invade the loose areolar connective tissue. Fibrosis of the edematous and inflamed tissue ensues, permanently inhibiting the normal telescoping action of the prepuce.17 The fate of the penis is sealed when the pendulous weight of the penis and prepuce damages the pudendal nerves.21, 25 The protruded penis eventually
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Abnormalities of the Penis and Prepuce curves caudally. Paralysis has no effect on the horse’s ability to urinate or ejaculate,17, 21, 22 but the affected stallion loses erectile function.2
Treatment Treatment of a horse affected with paraphimosis should be aimed at resolving the cause of the condition, eliminating preputial edema, and preventing preputial trauma. If possible, the penis should be replaced into the preputial cavity because, by doing so, normal venous and lymphatic drainage is restored and the penis is protected against injury. A penis and internal preputial lamina can be maintained within the preputial cavity by occluding either the preputial orifice or preputial ring with towel clamps or sutures, but these devices damage the prepuce and exacerbate swelling at the site at which they are placed. The penis and internal preputial lamina may also be maintained within the preputial cavity as described above (see Penile hematomas, Treatment). The horse is able to urinate through these retaining devices without trapping urine. The penis and internal preputial lamina should be covered with an emollient, antimicrobial cream. A 2% testosterone cream, available by prescription from compounding pharmacies, added to an equal amount of emollient udder cream and applied once daily is effective in maintaining the health of the penile and preputial epithelium.23 If the prepuce is too swollen to permit placement of the penis into the preputial cavity, the swollen internal preputial lamina can be compressed against the penis with a tight elastic, non-adhesive bandage. Bandaging begins distally and proceeds proximally to the preputial orifice.14 The bandaged penis and prepuce are massaged until a decrease in size of the prepuce loosens the bandage. The procedure is repeated until swelling is resolved enough to permit the penis to be replaced into the preputial cavity. Two or three applications of bandaging and massage, performed over a period of 20–30 minutes, are usually required to remove most of the edema from the penis and internal preputial lamina.14 This procedure can be performed with the horse sedated. If this procedure provides insufficient resolution of swelling to permit replacement of the penis into the preputial cavity, the penis and internal preputial lamina can be elevated and compressed against the abdomen for several hours or more with a bandage. A hyperosmolar, hygroscopic agent, such as glycerin or jam, applied to the internal preputial lamina may increase the efficacy of the bandage in reducing edema.27 Lightly exercising the horse after its penis is fixed to the abdomen and administering a non-steroidal, antiinflammatory drug may also help reduce edema.
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If the inelastic, preputial ring prevents placement of the penis into the preputial cavity or if its strangulating effect is contributing to edema of the portion of the prepuce and penis distal to it, the ring can be incised longitudinally with the horse anesthetized or after administration of regional or local anesthesia. The incision is left unsutured to heal by second intention.16 A horse that has suffered severe preputial trauma, either as a cause or a result of paraphimosis, may be left with fibrosis and scarring of the internal preputial lamina that hampers normal telescoping action of the prepuce. Normal preputial function can sometimes be restored by segmental posthetomy (i.e., reefing) to removed restrictive scar tissue (see Chapter 163, Surgery of the Penis and Prepuce, for a description of this procedure). Horses with penile paralysis are not likely to recover the ability to retract the penis, or, in the case of stallions, erectile function.2, 22 A stallion with a paralyzed penis may retain its ability to ejaculate and so can sometimes be salvaged for breeding if it can be trained to ejaculate into an artificial vagina.2 A paralyzed penis can be permanently retracted into the preputial cavity by extensive posthetomy (i.e., the Adam’s procedure)18 or with sutures (i.e., the Bolz procedure),21 or its penis can be partially amputated.
Phimosis Etiology Phimosis, or the inability of the horse to protrude its penis from the preputial orifice or preputial ring, can be caused by congenital or acquired stenosis of the preputial orifice or preputial ring (Figure 109.3) or, more commonly, from a lesion on the penis or
Figure 109.3 This horse was unable to protrude its penis through a scarred preputial ring. Note the hole in the preputial fold proximal to the preputial ring (arrow).
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preputial lamina, such as a carcinoma, that prevents penile protrusion. Swelling of the internal preputial lamina of a retracted penis caused by trauma may also prevent the penis from protruding. The internal preputial lamina is fused to the free part of the penis during the first month after birth,28, 29 preventing foals from completely protruding their penis, but, discounting this normal phenomenon, congenital phimosis, to our knowledge, has never been reported to occur in horses. Entrapment of the penis in a rent in the suspensory ligament of the prepuce, apparently caused during castration, was a cause of phimosis in one horse.30 Regardless of its cause, phimosis may cause the horse to urinate into the preputial cavity, producing mucosal inflammation that may lead to fibrosis of the preputial lamina, compounding obstruction of the preputial orifice or preputial ring.
Treatment Phimosis caused by a lesion on the penis or internal preputial lamina, usually a carcinoma, is relieved when the lesion is resolved. Phimosis caused by stenosis of the preputial orifice can be relieved either by making a longitudinal incision that begins at the preputial orifice and extends caudally on the external preputial lamina or by excising a triangle of external preputial lamina, the base of which is the preputial orifice.31 The mucosal and cutaneous edges of the wound are apposed with interrupted sutures. Phimosis caused by stenosis of the preputial ring is similarly relieved either by making a longitudinal incision through the preputial ring or by removing a triangle, based at the preputial ring, from the internal preputial fold. The edges of the wound can be sutured or, after the penis is exteriorized, the preputial ring can be removed by segmental posthetomy (i.e., reefing). Phimosis caused by a tear in the suspensory ligament of the prepuce is relieved by suturing the tear.30
Priapism Priapism, or persistent erection without sexual excitement, is caused by failure of detumescence.32 Both stallions and geldings are affected,33, 34 but geldings are affected less commonly, perhaps because androgens influence the development of the condition, or perhaps because stallions develop erections more frequently.19, 35 The condition occurs infrequently in horses, but its occurrence in a breeding stallion is cause for grave concern, because, if it is not resolved, impotence is the final outcome.
Etiology Priapism in horses most commonly arises after administration of a phenothiazine-derivative tranquilizer, usually acepromazine.33, 34 The effect of phenothiazine-derivate tranquilizers in inducing priapism may be related to their potential for inducing peripheral α 1 -adrenergic blockade, directly interrupting sympathetic impulses for detumescence.36 Other reported causes of priapism in horses include general anesthesia,37, 38 a tumor in the pelvic canal,19 and migration of a nematode in the spinal cord.39
Pathophysiology Priapism occurs when arterial inflow or venous outflow to the CCP is disturbed, causing failure of detumescence.32 Only the CCP is involved in the persistent erection. Priapism is categorized as high flow, or arterial, when an increase in arterial inflow to the cavernous tissue of the penis overrides the venous outflow, and as low flow, or veno-occlusive, when the α-adrenergic impulses that bring about detumescence are blocked, such as occurs with phenothiazine-derivative tranquilizers or when primary vascular or hematological disease mechanically interferes with venous drainage.40 High-flow priapism of men is caused by an arterial–cavernosal shunt and is nearly always a consequence of trauma to the perineum or penis.41 Lowflow priapism occurs much more commonly in men than does high-flow priapism42, 43 and seems to be the type of priapism that occurs in horses. High-flow priapism was reported to occur in a stallion, based on response to treatment and comparison of blood gas values from peripheral arterial blood and blood aspirated from the CCP, but this stallion developed priapism after it received a phenothiazine-derivative tranquilizer, the most common cause of low-flow priapism in horses.44 Regardless of its etiology, low-flow priapism is characterized by stasis of blood within the CCP.32 Low-flow priapism in men is accompanied by a pH < 7.25, partial pressure of oxygen (Po2 ) < 30 mmHg, and partial pressure of carbon dioxide (Pco2 ) > 60 mmHg.41 Stasis of blood in the sinusoidal spaces of the CCP raises the Pco2 in the stagnant blood, causing erythrocytes to aggregate or sickle. The sickled erythrocytes occlude the venous outflow from the CCP, causing the trabeculae in the sinusoidal spaces to become edematous. The edematous trabeculae are eventually invaded by fibrous tissue, thereby inhibiting the ability of the cavernosal tissue to expand normally during erection. Arterial inflow to the CCP initially remains unobstructed, but, eventually, it also obstructs. The pendulous weight of the
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Abnormalities of the Penis and Prepuce horse’s penis and compression of the penis against the ischium may injure the pudendal nerves, causing penile paralysis.19 If priapism is not rapidly resolved, the horse’s penis loses erectile function and sensitivity, the result of which is impotence. Clinical signs The penis of an affected horse may be fully erect or it may appear deceptively flaccid, in which case turgidity of the CCP is identified when the penis is palpated. Turgidity of the CCP prevents the horse from retracting its penis into the preputial cavity. The internal preputial lamina becomes edematous, if the protruded penis is improperly cared for. A chronically affected penis may be insensitive to painful stimuli, and, when ultrasonographically examined, its CCP appears densely echogenic.19, 45 Uncommonly, the horse may have difficulty urinating.46 Treatment
Medical treatment Horses with priapism have been treated by massage of the penis and prepuce and elevation of the penis with slings,35 both of which are effective in preventing or ameliorating preputial edema, but these treatments have no effect on relieving the condition. Horses suffering from priapism caused by administration of an α-adrenergic drug that blocks detumescence, such as a phenothiazine-derivative tranquilizer, can sometimes be treated successfully by administration of an anticholinergic drug commonly used to treat people with Parkinson’s disease, benztropine mesylate.34, 38 An average-sized horse should receive 8 mg intravenously. Administration of terbutaline, a β 2 agonist, has been used successfully in affected men and may be of benefit in treating affected horses.47 Clenbuterol, a β 2 agonist commonly used in horses as a bronchodilator, might also be effective. These drugs are more likely to be effective when treatment is initiated soon after the onset of priapism. Men with low-flow priapism are often treated with an α-adrenergic drug, such as phenylephrine, ephedrine, etilefrine, or adrenaline, injected directly into the CCP.48 These drugs may bring about penile detumescence by enhancing contractility of smooth muscle in the cavernous tissue.48 Detumescence after injection of 10 mg of 1% phenylephrine, diluted in 10 mL physiological saline solution (PSS), directly into the erect CCP, is often immediate and dramatic, especially if the drug is administered soon after the onset of disease (J. Schumacher and D. Varner, personal observation). Similar treatment of horses with
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a longer duration of disease may provide only temporary detumescence.
Surgical treatment A horse that does not experience detumescence within a few hours after initiation of treatment should be treated by evacuating stagnant blood from the erect CCP by irrigating this cavernous body with PSS or balanced electrolyte solution to which heparin (10 IU heparin/mL) has been added. The CCP can be irrigated with the horse sedated, but it is mostly easily and safely irrigated with the horse anesthetized and in dorsal recumbency. The irrigation fluid is infused under pressure into the CCP just proximal to the glans penis through a 12- or 14-gauge needle, and exited, along with the stagnant blood, 10–15 cm distal to ischium, through a needle of the same size inserted into the CCP. Irrigation can be stopped when fresh blood appears in the efflux. Instilling 10 mg of 1% phenylephrine into the CCP at the end of irrigation may be useful in evacuating fluid in the CCP. Failure of bright red, arterial blood to appear at the exit portal after sludged blood has been evacuated from the CCP may signify permanent damage to the arterial supply to the CCP, the result of which is irreversible impotence. Guidelines have not been established for horses to indicate the time after the onset of erection at which irrigation of the CCP should be initiated because the time at which pathological changes in the CCP are permanent has not been determined. We believe that the pudendal nerves may be damaged long before irreversible changes to the CCP occur, and, therefore, we recommend irrigating the CCP within a few hours after onset of erection, if treatment by cholinergic blockade or α-adrenergic stimulation fails. Return of the erection after irrigation indicates that irrigation has failed to restore venous outflow and that arterial inflow remains unobstructed. If erection returns after irrigation, blood trapped in the CCP can be shunted to the CSP, which is not involved in priapism (see Chapter 163, Surgery of the Penis and Prepuce, for a description of the technique of cavernosal shunting).32, 49 Geldings affected with priapism can be treated by partial phallectomy if priapism does not resolve with medical therapy and irrigation.37 The horse does not experience greater than normal hemorrhage from the penile stump after phallectomy, but the stump remains erect after amputation. Tumescence gradually resolves as the remaining portion of the CCP fibroses. Erectile capability of men whose cavernous tissue has been damaged by priapism can be improved temporarily by injecting a vasoactive drug, such as phenoxybenzamine or phentolamine, into the CCP
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before copulation.32 Erectile function of a horse with damaged erectile tissue might also be enhanced before copulation by injecting one of these drugs into the CCP, but the doses of these drugs for injection into cavernosal tissue of horses have not been established. The copulatory capability of a stallion with diminished penile sensation and poor erectile function resulting from damage caused by priapism might be enhanced by decreasing the horse’s ejaculatory threshold by administering imipramine hydrochloride (2.2–3 mg/kg orally), 2 hours before breeding.46, 50 Testosterone-impregnated cream applied topically to the glans penis might increase penile sensation. A stallion unable to achieve a full erection might accomplish intromission if it is assisted in placing its penis into the vagina, or it might be trained to ejaculate into an artificial vagina. Some stallions can be induced to ejaculate without copulation, by administering imipramine hydrochloride orally to lower the ejaculatory threshold and then administering xylazine hydrochloride, a common sideeffect of which is emission of semen, usually within 15 minutes.46, 50 Semen obtained in this manner commonly has a high concentration of sperm in a low volume of semen, presumably because imipramine reduce contraction of the accessory sex glands, and because collection is not associated with sexual excitement.
Intersex Clinical features An intersex is an individual whose sexual identity is confusing because of contradictory morphological criteria of sex.51–53 A person’s or animal’s sex can be identified according to the genetic composition, the type of gonads, or the morphology of the accessory genital organs. The sex of a normal animal is the same in all three categories, but an intersex differs in one of these categories. An intersex results from errors in embryogenesis because the embryonic structures from which male and female reproductive organs form are common to both sexes.53 An intersex is sometimes referred to as a hermaphrodite, a term that implies that the animal or person is both fully male and fully female, a situation not recognized in horses or people. Regardless, a horse that has both testicular and ovarian tissue (i.e., either an ovary and a testis, or a combined gonad called an ovo-testis) is commonly termed a ‘true hermaphrodite’.51, 52, 54 True equine hermaphrodites are much less commonly encountered than are equine pseudohermaphrodites, which are horses that have the external genitalia of one sex but the gonads of the opposite sex.
A pseudohermaphrodite is defined as male or female according to its gonadal sex, and the most common type is the male pseudohermaphrodite, which typically has testes located at the external inguinal ring or within the inguinal canal.55 A phallus-like structure, which may resemble a large clitoris, and a rudimentary prepuce, which may resemble a vulva, are located on the ventral midline somewhere between the normal location of the preputial orifice and the ischial arch.31, 52, 55, 56 The mammary glands of a male pseudohermaphrodite may develop moderately and resemble those of a mare.31, 55 Although its external genitalia may resemble that of a mare, the non-castrated, male pseudohermaphrodite displays stallion-like behavior and can develop an erection of the rudimentary penis.57 Although equine male pseudohermaphrodites possess testes, most are actually genetic females (64XX).51–54,58
Treatment Treatment is aimed at decreasing the sexual ambiguity of the horse’s appearance. If the male pseudohermaphrodite’s external genitalia are located near the ischial arch, they can be altered to more closely resemble that of a female by amputating the phalluslike structure, creating a new urethral stoma, and constructing labia using the rudimentary prepuce.31 If the male pseudohermaphrodite’s external genitalia are less than 15 cm caudal to the nipples, they can be altered to more closely resemble that of a male by rotating the phallus-like structure and rudimentary prepuce cranially to a more normal position, slightly caudal to the umbilicus.55 The internal lamina proximal to the preputial ring is removed from the shaft of the rudimentary penis, and the penis is advanced cranially through a tunnel created beneath the skin to immerge through a circular incision cranial to the mammary glands. The preputial ring is sutured to the circular incision. The horse should be castrated at least 4 weeks before surgery.
Neoplasia Incidence and etiology Squamous cell carcinoma is by far the most common penile and preputial neoplasm.59, 60 Other occasionally encountered neoplasms of the penis and prepuce include squamous papillomas, sarcoids, and melanomas.59, 61 Squamous papillomas, the benign counterparts of squamous cell carcinoma, are sometimes found on the external genitalia of young horses62 and disappear spontaneously when the horse acquires immunity to the virus that causes
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Abnormalities of the Penis and Prepuce them.63 Melanomas are sometimes found on the external and internal preputial laminae of old gray horses.2 Although melanomas have a low rate of malignancy, they may interfere with normal protrusion and retraction of the penis. Old horses and geldings are more likely than young horses and stallions to develop penile and preputial lesions of squamous cell carcinoma.16, 61, 64, 65 Horses that typically have nonpigmented external genitalia, such as cremellos, palominos, American Paint Horses, and Appaloosas, are more likely to develop penile and preputial carcinomas because lack of preputial or penile pigmentation seems to predispose to carcinoma, even though the external genitalia are unexposed to direct sunlight.64 A study of the effects of equine smegma on the skin of mice concluded that smegma likely contains carcinogenic agents capable of initiating formation of preputial or penile squamous cell carcinoma,66 but another study of the cutaneous effects of human smegma on mice did not substantiate these findings. 67 Regardless of whether or not smegma contains carcinogens, chronic irritation caused by smegma may stimulate neoplastic transformation of penile and preputial integument. Geldings and old horses may be prone to developing squamous cell carcinoma of the external genitalia because they produce smegma in greater quantities than do stallions and young horses.64, 66 Penile and preputial squamous cell carcinomas generally have a low grade of malignancy, grow remarkably slowly for carcinomas, and metastasize to the inguinal lymph nodes late in the disease.60, 61, 65 In one study, only five out of 41 horses with penile or preputial squamous cell carcinoma had evidence of metastases of the neoplasm to the inguinal lymph nodes.60
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Figure 109.4 The glans and internal preputial lamina of this horse contain many small, precancerous, keratinized plaques.
contain necrotic, ulcerated areas. A large excrescence can interfere with protrusion and retraction of the penis or with urination by impinging on the urethra (Figure 109.7).68 Affected horses usually suffer from balanoposthitis.60, 65 Metastasis of penile or preputial squamous cell carcinoma is usually recognized by observing enlargement of the inguinal lymph nodes, but a carcinoma can metastasize to internal organs without causing grossly detectable enlargement of the
Diagnosis Affected horses are usually presented for examination shortly after the owner first observes lesions, but some may be presented because of a malodorous, blood-stained, preputial discharge.60, 65 The duration of disease is usually unknown because most owners seldom inspect their horse’s penis and prepuce.65 A precancerous lesion appears as a small, keratinized plaque (Figure 109.4), and a lesion that has undergone transformation into a neoplasm usually first appears as a shallow ulceration with an indurated base (Figure 109.5).61, 63, 65 Proliferating neoplastic cells may come to resemble granulation tissue (Figure 109.6). A long-standing carcinoma often develops a cauliflower-like appearance and may
Figure 109.5 A carcinoma usually first appears as a shallow ulceration with an indurated base.
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Surgical excision
Figure 109.6 Penile carcinomas may resemble granulation tissue.
inguinal lymph nodes.69 Slight enlargement of the inguinal lymph nodes from metastases may be difficult to distinguish from enlargement secondary to balanoposthitis caused by the penile or preputial carcinoma.60, 69
Small cancerous lesions on the external or internal preputial lamina can be excised with a scalpel or with a carbon dioxide laser.70 Using a laser to excise a neoplasm kills tumor cells at the base and cutaneous margins of the neoplasm and decreases postoperative swelling by sealing lymphatic vessels.71 Extensive neoplastic lesions on the external genitalia are removed by posthetomy (i.e., reefing) or by partial phallectomy. If the penis and prepuce are extensively affected by neoplasia, they and the inguinal lymph nodes may need be removed en bloc (see Chapter 163, Surgery of the Penis and Prepuce for a detailed description of surgical techniques for removal of neoplastic lesions).72
Cryotherapy Small lesions of squamous cell carcinoma can be removed using cryotherapy. Neoplastic lesions can be frozen using liquid nitrogen administered as a spray or using either liquid nitrogen or carbon dioxide administered through a cryoprobe.73 A fast freeze–slow thaw cycle, performed twice, is most effective in destroying neoplastic cells.74 Tissue should be frozen to –20◦ C to –30◦ C using thermocouple electrodes to measure the temperature of the tissue. Freezing too deeply could result in exposure of cavernous tissue and urethral damage. The primary disadvantage of freezing neoplastic lesions compared with surgical excision is that epithelialization of the wound resulting from freezing is prolonged.
Chemotherapy
Figure 109.7 A penile carcinoma, such as this one, can interfere with protrusion and retraction of the penis and with urination.
Horses with numerous, small, preputial and penile neoplastic and pre-neoplastic lesions have been treated successfully with 5% 5-fluorouracil cream applied to the penis and prepuce at 14-day intervals for up to seven treatments.75 More frequent application causes inflammatory edema and discomfort to the horse. A chemotherapeutic agent used successfully to treat horses with squamous carcinoma of the external genitalia is cisplatin, which is administered by intratumoral injection in an oily emulsion.76, 77 The drug is combined with sesame seed oil and sterile water (3.3 mg cisplatin/mL of emulsion), and the emulsion is administered into the tumor and a 1-cm margin of normal tissue at a ratio of 1 mg/cm3 tissue at 14-day intervals, usually until the horse has been treated four times. Intratumoral administration of cisplatin can be used as the sole treatment of horses with small tumors or combined with excision of the tumor for treatment of horses with
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Abnormalities of the Penis and Prepuce large tumors. Residual tumor cells along the margins of the wound after the tumor has been excised are stimulated into active proliferation and hence are most susceptible to chemotherapy immediately after surgery. Administering cisplatin intraoperatively and at 14-day intervals provides better results, at least for horses with tumors that have a high proliferation index, than does delaying chemotherapeutic treatment until the wound has healed.77 Injecting the tumor bed and a margin of normal tissue with cisplatin during surgery and periodically during healing does not have a clinically apparent detrimental effect on healing of wounds closed primarily or wounds left unsutured to heal by second intention.76, 77 Horses whose penile or preputial carcinoma has metastasized to internal organs can sometimes be treated with a systemically administered chemotherapeutic agent, such as doxorubicin, or a cyclooxygenase-inhibiting drug, such as piroxicam,78 but reports of the use of these drugs to treat horses for neoplasia are rare. Prognosis In a study of horses affected with penile or preputial carcinoma, about 81% of 45 affected horses survived at least 1 year after surgical treatment with no evidence of recurrence of disease,65 and, in another study, 64.5% of 48 horses were alive 18 months after surgical treatment.60 Horses in both studies had a poor prognosis for survival if the carcinoma had metastasized to the inguinal lymph nodes. Invasion of cavernous tissue by a carcinoma is an unfavorable prognostic factor for survival in men because neoplastic invasion into a corporeal body may be more likely to result in hematogenous metastasis of the neoplasm.79 In one study, three out of four horses that had metastases of penile or preputial carcinoma to internal organs had gross or histological evidence of neoplastic invasion of a corporeal body.65 Because horses whose corporeal tissue has been invaded by a carcinoma may have a high likelihood of abdominal metastases, owners of such horses should be forewarned, before treatment is undertaken, that corporeal invasion by the tumor may dramatically decrease the horse’s prognosis for survival. Horses should be examined periodically for recurrence of penile or preputial carcinoma after apparently successful treatment because squamous cell carcinoma sometimes recurs. Horses that have or have had squamous cell carcinoma of the external genitalia are predisposed to developing carcinomas in other regions, such as around the eyes, and, therefore, should receive frequent physical examination.
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Habronemiasis Etiology Cutaneous habronemiasis, also known as ‘summer sores’, ‘bursatti’, or ‘granular dermatitis’, is a granulomatous, cutaneous disease caused by infestation of wounds by the larvae of the equine stomach worm Habronema.63, 80 The adult parasites live in the stomach, and their larvae and eggs are expelled in the feces and are ingested by maggots of the intermediate host, the housefly or the stable fly. The maggot pupates into a feeding fly, which deposits the infective, third-stage larvae on a wound on the horse. Larvae are probably able to penetrate intact skin, and, consequently, lesions are often seen in chronically moist regions, such as the prepuce, to which flies are attracted.81 The disease appears in warm countries during warm months, when flies are prevalent, and may regress partially or completely during cold weather. The penis and prepuce are common sites of infestation by these larvae because flies, which are attracted to moisture on these structures, cannot be easily removed from the external genitalia by the tail hairs. Horses that are inclined to protrude their penis and horses that are infrequently administered anthelmintic drugs may be more prone to developing cutaneous habronemiasis of the penis and prepuce.80 Some horses appear to be more disposed to developing cutaneous habronemiasis and are afflicted with lesions in multiple sites and by yearly recurrence. Pathophysiology Deposited Habronema larvae may produce a substance that destroys tissue through which the larvae migrate,63 but local hypersensitivity to the larvae probably accounts for the granulomatous response.80 Horses with lesions of cutaneous habronemiasis are nearly always heavily parasitized by adult stomach worms, leading to the claim that the presence of adult worms in the stomach induces a generalized hypersensitivity to the larvae. Clinical signs Specific sites of genital infestation are the urethral process, the preputial ring, and the preputial orifice. Preputial lesions may appear as ulcerated areas filled with granulation tissue surrounded by depigmented integument.63 Lesions typically discharge a hemorrhagic or serosanguineous exudate and are mildly to severely pruritic. During close examination, the granulation is seen to contain numerous, small (i.e., 1 mm), hard, yellow, granules, called Bollinger’s
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granules, which are composed of eosinophils, nuclear remnants, and larvae.82 Stoma, or fistulous-like tracts, are commonly observed on the skin adjacent to the lesions.83 Chronic infestation of portions of the internal preputial laminae may result in fibrosis that hinders the telescoping action of the prepuce. Chronic infestation of the urethral process may cause periurethral fibrosis, which may be so severe that the horse becomes dysuric or sprays its limbs with urine.84 Erosion of a lesion into the corporeal tissue in the urethral process may result in hemorrhage at the end of urination or during ejaculation. Diagnosis A presumptive diagnosis of cutaneous habronemiasis is usually based on the typical appearance of lesions and clinical signs of disease. A granulating wound that enlarges or fails to heal and is accompanied by signs of pruritus is suggestive of the disease. Squeezing the lesion may cause granules to extrude. A marked eosinophilia is often noted during examination of a complete blood count. Definitive diagnosis requires cytological or histological identification of the larvae within the wound because a lesion on the genitalia or elsewhere can be confused with a sarcoid, squamous cell carcinoma, exuberant granulation tissue, or pythiosis.80, 82 Although failure to identify larvae during histological examination of a lesion prevents definitive diagnosis of cutaneous habronemiasis, other diseases, such as pythiosis and bacterial infection of a granulating wound, characterized by production of exuberant granulation tissue, or diseases, such as sarcoids and carcinomas, that produce tissue that resembles exuberant granulation, can often be excluded as a cause of the lesion. Motile larvae, the tails of which have a spiny process, can sometimes be identified during microscopic examination of deep scrapings or of exudate squeezed from the lesion onto a slide.82, 83 Cross-sections of larvae surrounded by an area of collagenolysis are often seen during histological examination of a lesion. Other histological characteristics of a lesion include a massive concentration of eosinophils and the presence of mast cells and multinucleated giant cells.82, 84, 85
corticosteroid.80, 82 Ivermectin, administered orally at 0.2 mg/kg, or moxidectin, administered orally at 0.4 mg/kg is effective in resolving lesions by killing the Habronema larvae.82 Organophosphates, such as runnel, administered orally have been effective in destroying the larvae and resolving lesions.86 Prednisolone or prednisone administered orally at 1–1.5 mg/kg, daily for 10–14 days and then at a reduced dosage for another 2 weeks is effective in resolving lesions by decreasing the horse’s inflammatory response to living larvae or larvae killed by a previously administered parasiticide. Daily topical application of a cream or ointment containing a corticosteroid, such as dexamethasone or triamcinolone acetonide, may bring about resolution of small, granulomatous lesions caused by cutaneous habronemiasis by dampening the body’s reaction to the larvae. These topically applied anti-inflammatory drugs are often combined with an organophosphate, such as trichlorfon or fenthion. Horses with genital lesions of habronemiasis can also be treated effectively by application of cryogen.87 Lesions can be frozen as a spray using liquid nitrogen or through a cryoprobe using liquid nitrogen or carbon dioxide. The lesions should be frozen twice using a fast freeze–slow thaw cycle. To prevent reinfestation with Habronema larvae, the life cycle of the nematode should be interrupted by controlling the fly population by hygienic disposal of manure and judicious use of fly sprays and repellents. Horses on the premises should regularly be administered an anthelmintic drug effective against the equine stomach worm. Surgical treatment Small lesions on the internal preputial lamina can be excised and the wound sutured. Surgical removal of large lesions may require segmental posthetomy. The most expedient means of eliminating lesions on the urethral process, especially lesions that result in dysuria, hematuria, or hemospermia, may be to amputate the urethral process (for details of these procedures, see Chapter 163).84
Urethral rents Etiology and pathophysiology
Treatment
Non-surgical treatment Lesions are resolved either by eliminating larvae from the wound with a parasiticide or by reducing the horse’s hypersensitivity to them with a
A rent in the proximal portion of the urethra has been identified as a cause of hematuria in geldings and hemospermia (or hematospermia) in stallions. The source of hemorrhage in the urine of affected geldings and the ejaculate of affected stallions is the CSP, which communicates with the lumen of the urethra
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Abnormalities of the Penis and Prepuce through the mucosal defect. Stallions typically hemorrhage from the CSP through the rent during or at the end of ejaculation when contraction of the bulbospongiosus muscle causes pressure within the CSP to increase from 17 to nearly 1000 mmHg.88 Geldings typically hemorrhage from the CSP through the rent at the end of urination when intraluminal urethral pressure decreases suddenly while the pressure in the CSP remains high. Hemorrhage was once thought to be caused by sudden increase in pressure in the CSP when the bulbospongiosus muscle contracted to expel urine from the urethra,89 but research has shown that the rise in pressure within the CSP associated with contraction of the bulbospongiosus muscle is slight.90 The rent in the urethra of stallions responsible for hemospermia is identical in appearance and location to the rent responsible for hematuria of geldings, but affected stallions rarely develop macroscopic hematuria.89, 91 The reason stallions that have a urethral rent rarely develop gross evidence of hematuria is that, during urination, pressure in the CSP of stallions is nearly half that of geldings.90 The pressure in the CSP of stallions is probably lower because the CSP of stallions is more voluminous than that of geldings. Stallions, like geldings, may hemorrhage into the urethra at the end of urination but in amounts too minute to be detected grossly. Even minute hemorrhage through the rent at the end of urination, however, may prevent the rent from healing. Although the nature of the forces that induce the rent is idiopathic, the constant location of the rent on the convex surface of the urethra caudal to the ischial arch might be explained by the anatomy of the urethra. At its origin, the urethra is 1–1.5 cm in diameter.92 It expands to a diameter of 3.5–5 cm in the pelvis, and then, as the urethra bends sharply where it passes around the ischial arch, it decreases dramatically to 1–1.5 cm. The acute bending and narrowing of the urethra in this region may expose the convex surface of the urethra at the level of the ischial arch to hydrodynamic forces not encountered by other portions of the urethra. Diagnosis Blood in the ejaculate is most easily recognized when the stallion’s ejaculate is collected with an artificial vagina. Semen of affected horses is usually pink to red, but microscopic examination of the semen may be necessary to detect hemospermia. Red blood cells in the ejaculate have a detrimental effect on spermatozoa, but the precise factors that cause infertility are not known.44, 93 Geldings that hemorrhage during urination through a urethral rent characteristically do so at the end of urination (i.e., terminal hematuria),
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Figure 109.8 Most rents, such as this one, appear endoscopically as a 5- to 10-mm-long, linear defect on the convex surface of the urethra, at the level of the ischial arch.
and some show signs of dysuria, such as tenesmus, at the end of urination.89 A urethral rent is identified by examining the urethra using a sterilized, flexible endoscope that is at least 100 cm long. Most rents appear as a 5- to 10-mm long, linear defect on the convex surface of the urethra, at the level of the ischial arch (Figure 109.8).89 Signs of inflammation around the defect are usually not detected. The location of the defect can be verified during the examination by observing the shaft of a 22- to 20-gauge hypodermic needle inserted percutaneously through the perineal raphe into the lumen of the urethra at the level of the ischial arch. Blood can sometimes be observed emanating from an otherwise undetectable rent by endoscopically examining the urethra of an affected gelding immediately after the gelding urinates or the urethra of an affected stallion immediately after the stallion ejaculates.94
Treatment A certain amount of blood in the ejaculate is compatible with fertility, and, so, fertility of stallions with slight hemorrhage in the ejaculate can sometimes be preserved by adding an extender to the semen to dilute the concentration, and hence effect, of red blood cells in the ejaculate.95, 96 Although a urethral rent of a stallion may heal during a long period of sexual abstinence and a urethral rent of a gelding may heal spontaneously,97 affected horses are treated most effectively by creating a temporary, perineal urethrostomy or by creating an ischial incision that extends into the CSP but does not enter the lumen of the urethra (see Chapter 163, Surgery of the Penis and Prepuce for a description of urethrostomy).89, 91, 98
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The horse may hemorrhage considerably from the perineal incision after surgery, especially at the end of urination, but hemorrhage from the urethral orifice during urination is no longer observed.89 Incising into the CSP through the ischial incision, regardless of whether the incision penetrates the urethral mucosa, resolves hematuria of geldings, without fail in our experience, presumably by reducing pressure in the CSP, thereby preventing blood from exiting through the rent at the end of urination. Eliminating hemorrhage into the urethra allows the rent to heal. Avoiding incising the urethral mucosa might reduce the uncommon risk of complications associated with perineal urethrostomy, such as development of a urethral fistula98 or stricture.99 Sexual abstinence, in addition to perineal urethrostomy, is also important in the treatment of stallions with hemospermia caused by a urethral rent because erection and contractions of the bulbospongiosus muscle during ejaculation prevent the rent from healing. Although urethrostomy is highly effective in resolving hematuria of geldings, hemospermia of stallions sometimes persists. If exposure of the CSP or urethral lumen through a perineal incision is ineffective in resolving hemospermia, the lesion can be closed primarily through a perineal incision (for a description of primary closure of the rent, see Chapter 163).
References 1. Vaughan JT. Surgery of the prepuce and penis. In: Proceedings of the 18th Annual Convention of the American Association of Equine Practitioners, 1972, pp. 19–40. 2. Neely DP. Physical examination and genital diseases of the stallion. In: Morrow DA (ed.) Current Therapy in Theriogenology. Philadelphia: W.B. Saunders, 1980, p. 694. 3. Perkins NR, Frazer GS. Reproductive emergencies in the stallion. Vet Clin North Am Equine Pract 1994;10:671–83. 4. Taylor NR. Traumatic balanoposthitis in a yearling Appaloosa colt. Vet Rec 1980;107:154–5. 5. Todhunter RJ, Parker JE. Surgical repair of urethral transection in a horse. J Am Vet Med Assoc 1988;193:1085–6. 6. Yovich JV, Turner AS. Treatment of postcastration urethral stricture by phallectomy in a gelding. Comp Cont Educ Pract Vet 1986;8:S393–9. 7. Munger RJ, Meagher DM. Surgical repair of a fistula of the urethral diverticulum in a horse. Vet Med Small Anim Clin 1976;71:96. 8. Pascoe RR. Rupture of the corpus cavernosum penis of a stallion. Aust Vet J 1971;47:610–11. 9. Perkins JD, Schumacher J, Waguespack RW, Hanrath M. Penile retroversion and partial phallectomy performed in a standing horse. Vet Rec 2003;153:184–5. 10. Boyer K, Jann HW, Dawson LJ, Cary M. Penile hematoma in a stallion resulting in proximal penile amputation. Equine Pract 1995;17:8–11. 11. Cox JE. Surgery of the Reproductive Tract in Large Animals. Liverpool, UK: Liverpool University Press, 1987.
12. Firth EC. Dissecting hematoma of corpus spongiosum and urinary bladder rupture in a stallion. J Am Vet Med Assoc 1976;169:800–1. 13. Hyland J, Church S. The use of ultrasonography in the diagnosis and treatment of a haematoma in the corpus cavernosum penis of a stallion. Aust Vet J 1995;72:468–9. 14. Boero MJ. A simple technique for conservative therapy of acute traumatic paraphimosis in the horse. In: Proceedings of the 36th Annual Convention of the American Association of Equine Practitioners, 1990, pp. 625–8. 15. Henning MW. Animal Diseases in South Africa, 3rd edn. Pretoria, South Africa: Central News Agency, 1956. 16. Walker DF. Surgery of the penis. In: Walker DF, Vaughan JT (eds) Bovine and Equine Urogenital Surgery. Philadelphia: Lea & Febiger, 1980; p. 10. 17. Simmons HA, Cox JE, Edwards GB, Neal PA, Urquhart KA. Paraphimosis in seven debilitated horses. Vet Rec 1985;116:126–7. 18. Guillaume A. Simplified surgical treatment of paralysis of penis in the horse. Vet J 1919;26:37–40. 19. Blanchard TL, Schumacher J, Edwards JF, Varner DD, Lewis RD, Everett K, Joyce JR. Priapism in a stallion with generalized malignant melanoma. J Am Vet Med Assoc 1991;198:1043–44. 20. Schumacher J, Hardin DK. Surgical treatment of priapism in a stallion. Vet Surg 1987;16:193–6. 21. Bolz W. The prophylaxis and therapy of prolapse and paralysis of the penis occurring in the horse after the administration of neuroleptics. Vet Med Rev Leverkusen 1970;4:255–63. 22. Wheat JD. Penile paralysis in stallions given propiopromazine. J Am Vet Med Assoc 1966;148:405–6. 23. McDonnell SM. Managing the paralyzed penis, priapism or paraphimosis in the horse. Equine Vet Educ 2005;17: 310–11. 24. Nie GJ, Pope KC. Persistent penile prolapse associated with acute blood loss and acepromazine maleate administration in a horse. J Am Vet Med Assoc 1997;211:587–9. 25. Teuscher H. Diseases of the male genital organs and hermaphroditism. In: Dietz O, Wiesner E (eds) Diseases of the Horse. New York: Karger, 1982; pp. 310–25. 26. de Lahunta A, Habel RE. Applied Veterinary Anatomy. Philadelphia: W.B. Saunders, 1986; pp. 287, 299. 27. Ryder-Davies P. Bad language? Vet Rec 1992;130:148. 28. Ashdown RR, Done SH. Color Atlas of Veterinary Anatomy. The Horse. Philadelphia: J.B. Lippincott, 1987; p. 8.26. 29. Faulkner LC, Pineda MH. Male reproduction. In: McDonald LE (ed.) Veterinary Endocrinology and Reproduction, 2nd edn. Philadelphia: Lea & Febiger, 1975; p. 212. 30. Swanstrom OG, Krahwinkel DJ. Preputial hernia in a horse. Vet Med Small Anim Clin 1974;69:870–1. 31. Frank ER. Veterinary Surgery, 7th edn. Minneapolis: Burgess, 1964, pp. 294–305. 32. Pohl J, Polt B, Kleinhans G. Priapism: a three phase concept of management according to aetiology and progress, Br J Urol 1986;58:113–18. 33. Lucke JN, Sansom J. Penile erection in the horse after acepromazine. Vet Rec 1979;105:21–2. 34. Sharrock AG. Reversal of drug-induced priapism in a gelding by medication. Aust Vet J 1982;58:39–40. 35. Pearson H, Weaver BMQ. Priapism after sedation, neuroleptanalgesia and anaesthesia in the horse. Equine Vet J 1978;10:85–90.
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Abnormalities of the Penis and Prepuce 36. Dorman WB, Schmidt JD. Association of priapism in phenothiazine therapy. J Urol 1976;116:51–3. 37. van Harreveld PD, Gaughan EM. Partial phallectomy to treat priapism in a horse. Aust Vet J 1999;77:167–9. 38. Wilson DV, Nickels FA, Williams MA. Pharmacologic treatment of priapism in two horses. J Am Vet Med Assoc 1991;119:1183–4. 39. Oyamada T, Miyajima K, Kimura Y, Kikuchi M, Nakanishi S, Yoshikawa H. Priapism possibly caused by spinal nematodiasis in a stallion. J Equine Sci 1997;8:101–7. 40. Rochat MC. Priapism: a review. Theriogenology 2001;56:713–22. 41. Stock KW, Jacob AL, Kummer M, Zimmermann U, Steinbrich W. High-flow priapism in a child: Treatment with superselective embolization. Am J Roentgenol 1996;166: 290–2. 42. Harmon WJ, Nehra A. Priapism: diagnosis and management. Mayo Clin Proc 1997;72:350–5. 43. Lue TF, Hellstrom WJG, McAninch JW, Tanagho EA. Priapism: a refined approach to diagnosis and treatment. J Urol 1986;136:104–8. ¨ 44. Boller M, Furst A, Ringer S, Dubs M, BettschartWolfensberger R. Complete recovery from long-standing priapism in a stallion after propionylpromazine/xylazine sedation. Equine Vet Educ 2005;17:305–11. 45. Love CC, McDonnell SM, Kenney RM. Manually assisted ejaculation in a stallion with erectile dysfunction subsequent to paraphimosis. J Am Vet Med Assoc 1992;200:1357–9. 46. Feary DJ, Moffett PD, Bruemmer JE, Southwood L, McCue P, Niswender KD, Dickinson C, Traub-Dargatz J. Chemical ejaculation and cryopreservation of semen from a breeding stallion with paraphimosis secondary to priapism and haemorrhagic colitis. Equine Vet Educ 2005;17:299–304. 47. Shantha TR, Finnerty DP, Rodriguez AP. Treatment of persistent penile erection and priapism using terbutaline. J Urol 1989;141:1427–9. 48. van Driel MF, Hesselink JW. [ Priapism in the stallion and in man]. Tijdschr Diergeneeskd 2003;128:255. 49. Schumacher J, Varner DD, Crabill MR, Blanchard TL. The effect of a surgically created shunt between the corpus cavernosum penis and corpus spongiosum penis of stallions on erectile and ejaculatory function. Vet Surg 1999;28:21–4. 50. McDonnell SM. Oral imipramine and intravenous xylazine for pharmacologically-induced ex copula ejaculation in stallions. Anim Reprod Sci 2001;68:153–9. 51. McIlwraith CW, Owen RA, Basrur PK. An equine cryptorchid with testicular and ovarian tissues. Equine Vet J 1976;8:156–60. 52. Roberts SJ. Veterinary Obstetrics and Genital Diseases. Ithaca, NY: Self-published, 1971; pp. 72–5. 53. Smith HA, Jones TC, Hunt RD. Pathology, 4th edn. Philadelphia: Lea & Febiger, 1972; pp. 330–1. 54. Stabenfeldt GH, Hughes JP. Reproduction in horses. In: Cole HH, Cupps PT (eds) Reproduction in Domestic Animals, 3rd edn. New York: Academic Press, 1977; p. 401. 55. Bracken FK, Wagner PC. Cosmetic surgery for equine pseudohermaphroditism. Vet Med Small Anim Clin 1983;78:879–81. 56. McFeely RA, Kanagawa H. Intersexuality. In: Hafez ESE (ed.) Reproduction in Farm Animals, 3rd edn. Philadelphia: Lea & Febiger, 1974. 57. Milliken JE, Paccamonti DL, Shoemaker S, Green WH. XX male pseudohermaphroditism in a horse. J Am Vet Med Assoc 1995;207:77–9.
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58. Fretz PB, Hare WC. A male pseudohermaphrodite horse with 63Xo?/64XX/65XXY mixoploidy. Equine Vet J 1976;8:130–2. 59. Cotchin E. Neoplasms of the Domesticated Mammals, 2nd edn. Wallingford, UK: Commonwealth Agricultural Bureau, 1956. 60. Howarth S, Lucke VM, Pearson H. Squamous cell carcinoma of the equine external genitalia: A review and assessment of penile amputation and urethrostomy as a surgical treatment. Equine Vet J 1991;23:53–8. 61. Jubb KVF, Kennedy PC. Pathology of Domestic Animals, vol. 1, 2nd edn. New York: Academic Press, 1970. 62. Junge RE, Sundberg JP, Lancaster WD. Papillomas and squamous cell carcinomas of horses. J Am Vet Med Assoc 1984;185:656–9. 63. Montes LF, Vaughan JT. Atlas of Skin Diseases of the Horse. Philadelphia: W.B. Saunders, 1983; pp. 91, 104. 64. Akerejola OO, Ayivor MD, Adams EW. Equine squamouscell carcinoma in Northern Nigeria. Vet Rec 1978;103: 336–7. 65. Mair TS, Walmsley JP, Phillips TJ. Surgical treatment of 45 horses affected by squamous cell carcinoma of the penis and prepuce. Equine Vet J 2000;32:406–10. 66. Plaut A, Kohn-Speyer AC. The carcinogenic action of smegma. Science 1947;105:391–2. 67. Reddy DG, Baruah IK. Carcinogenic action of human smegma. Arch Pathol 1963;75:414–20. 68. Scott EA. A technique for amputation of the equine penis. J Am Vet Med Assoc 1976;168:1047–51. 69. Patterson LJ, May SA, Baker JR. Skeletal metastasis of a penile squamous cell carcinoma. Vet Rec 1990;126:579– 80. 70. McCauley CT, Hawkins JF, Adams SB, Fessler JF. Use of a carbon dioxide laser for surgical management of cutaneous masses in horses: 32 cases (1993–2000). J Am Vet Med Assoc 2002;220:1192–7. 71. Palmer SE. Instrumentation and techniques for carbon dioxide laser in equine surgery. Vet Clin North Am Equine Pract 1996;12:397–414. 72. Markel MD, Wheat JD, Jones K. Genital neoplasms treated by en bloc resection and penile retroversion in horses: 10 cases (1977–1986). J Am Vet Med Assoc 1988;192:396–400. 73. Stick JA, Hoffer RE. Results of cryosurgical treatment of equine penile neoplasms. J Equine Med Surg 1978;2:505–7. 74. Joyce JR. Cryosurgery for removal of equine sarcoids. Vet Med Small Anim Clin 1975;70:200–3. 75. Fortier LA, MacHarg MA. Topical use of 5-fluorouracil for treatment of squamous cell carcinoma of the external genitalia of horses: 11 cases (1988–1992). J Am Vet Med Assoc 1994;205:1183–5. 76. Theon AP, Pascoe JR, Carlson GP, Krag DN. Intratumoral chemotherapy with cisplatin in oily emulsion in horses. J Am Vet Med Assoc 1993;202:261–7. 77. Theon AP, Pascoe JR, Galuppo LD, Fisher PE, Griffey SM, Madigan JE. Comparison of perioperative versus postoperative intratumoral administration of cisplatin for treatment of cutaneous sarcoids and squamous cell carcinomas in horses. J Am Vet Med Assoc 1999;215:1655–60. 78. Moore AS, Beam SL, Rassnick KM, Provost R. Longterm control of mucocutaneous squamous cell carcinoma and metastases in a horse using piroxicam. Equine Vet J 2003;35:715–18. 79. Soria JC, Theodore C, Gerbaulet A. Carcinome epidermoide de la verge. Bull Cancer 1998;85:773–84.
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80. McMullan W. Habronemiasis. In: Proceedings of the 22nd Annual Convention of the American Association of Equine Practitioners, 1976, pp. 295–8. 81. Soulsby E. Helminths, Arthropods and Protozoa of Domesticated Animals, 6th edn. Baltimore: Williams & Wilkins, 1968; pp. 270–3. 82. Rees C, Craig T. Cutaneous habronemiasis. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. Philadelphia: W.B. Saunders, 2003; pp. 195–7. 83. Rebhun WC. Observations on habronemiasis in horses. Equine Vet Educ 1996;8:188–91. 84. Stick JA. Amputation of the equine urethral process affected with habronemiasis. Vet Med Small Anim Clin 1979;74:1453–7. 85. Stick JA. Surgical management of genital habronemiasis. Vet Med Small Anim Clin 1981;76:410–14. 86. Herd RP, Donaham JC. Efficacy of ivermectin against cutaneous Drashia and Habronema infection (summer sores) in horses. Am J Vet Res 1981;42:1952–5. 87. Migioia S, Blanton AB, Davenport JW. Cryosurgical treatment of equine cutaneous habronemiasis. Vet Med Small Anim Clin 1978;73:1073–6. 88. Beckett SD, Walker DF, Hudson RS, Reynolds TM, Purohit RC. Corpus spongiosum penis pressure and penile muscle activity in the stallion during coitus. Am J Vet Res 1975;36:431–3. 89. Schumacher J, Varner DD, Schmitz DG, Blanchard TL. Urethral defects in geldings with hematuria and stallions with hemospermia. Vet Surg 1995;24:250–4. 90. Taintor J, Schumacher J, Schumacher J. Comparison of pressures in the corpus spongiosum penis during urina-
91. 92. 93.
94.
95.
96.
97.
98.
99.
tion between geldings and stallions. Equine Vet J 2004;36: 362–4. Schumacher J, Schumacher J, Schmitz D. Macroscopic haematuria of horses. Equine Vet Educ 2002;14:201–2. Sisson S, Grossman JD. The Anatomy of the Domestic Animals, 4th edn. Philadelphia: W.B. Saunders, 1953; p. 581. Pickett BW, Voss JL, Squires EL, Amann RP. Management of the Stallion for Maximum Reproductive Efficiency. General Series no. 1005. Fort Collins, CO: Colorado State University Experiment Station & Animal Reproduction Laboratory, 1981; pp. 79–84. Varner DD, Blanchard TL, Brinsko SP, Love CC, Taylor TS, Johnson L. Techniques for evaluating selected reproductive disorders of stallions. Anim Reprod Sci 2000;60:493– 509. Bedford SJ, McDonnell SM, Tulleners E, King D, Habecker P. Squamous cell carcinoma of the urethral process in a horse with hemospermia and self-mutilation behavior. J Am Vet Med Assoc 2000;216:551–3. Blanchard TL. Use of a semen extender containing antibiotics to improve the fertility of a stallion with seminal vesiculitis due to Pseudomonas aeruginosa. Theriogenology 1987;28:541–6. Schott HC. Hematuria. In: Reed SM, Bayly WM (eds) Equine Internal Medicine. Philadelphia: W.B. Saunders, 1988; pp. 890–5. Sullins KE, Bertone JJ, Voss JL, Pederson SJ. Treatment of hemospermia in stallions: a discussion of 18 cases. Comp Cont Educ Pract Vet 1988;10:S1396–400. Laverty S, Pascoe JR, Ling GV, Lavoie JP, Ruby AL. Urolithiasis in 68 horses. Vet Surg 1992;21:56–62.
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Chapter 110
Abnormalities of the Spermatic Cord James Schumacher and Dickson D. Varner
Anatomy and physiology The spermatic cord consists of structures carried ventrally with the testis as the testis passes through the inguinal canal from the abdomen to the scrotum. The constituents of the spermatic cord converge at the vaginal ring and terminate at the dorsal border of the testis. The structures that constitute the cord are the tunica vaginalis, the testicular blood and lymphatic vessels and nerves, and the ductus deferens. The inguinal canal is the slit-like passage in the abdominal wall through which the spermatic cord and overlying cremaster muscle traverse. The inguinal canal’s internal opening is the deep inguinal ring, and its external opening is the superficial inguinal ring. The deep inguinal ring is bordered cranially by the caudal edge of the internal abdominal oblique muscle, ventromedially by the rectus abdominis muscle and prepubic tendon, and caudolaterally by the inguinal ligament. The superficial inguinal ring is a slit in the aponeurosis of the external abdominal oblique muscle. The superficial inguinal ring is directed craniolaterally, and the deep inguinal ring is directed dorsolaterally, causing the lateral aspect of the rings to be widely divergent and the medial aspect of the rings to be in close proximity. The length of the canal in an average-sized adult horse, when measured along the spermatic cord, is about 15 cm.1 The tunica vaginalis, a diverticulum of the abdominal peritoneum, envelops the testis and its associated ducts and the constituents of the spermatic cord. It consists of the tunica vaginalis propria, or visceral tunic, and the tunica vaginalis communis, or parietal tunic. The visceral tunic is firmly adherent to the tunica albuginea of the testis, except at the dorsal border of the testis, where the constituents of the
spermatic cord enter or leave the testis. The parietal tunic is continuous with the parietal peritoneum of the abdomen at the deep inguinal ring and forms a sac, termed the vaginal sac or vaginal process, that lines the scrotal cavity and communicates with the peritoneal cavity. The cavity between the parietal and visceral tunics is the vaginal cavity, and the opening of the sac at the deep inguinal ring is termed the vaginal ring. The mesorchium is a fold of serous membrane, formed by a caudal invagination of the tunica vaginalis. It extends from the origin of the testicular vessels in the abdomen to the testis and envelops the testicular artery and veins, lymphatic vessels, and nerves. The ductus deferens and the deferential artery that accompanies it are enclosed by the mesoductus deferens, a medial fold of the mesorchium. The cremaster muscle is a striated muscle that originates from the internal abdominal oblique muscle and attaches to the tunica vaginalis at the caudolateral border of the spermatic cord. Because it lies outside the tunica vaginalis, it is not considered to be part of the cord. Contraction of the cremaster muscle retracts the testis by shortening the spermatic cord. The blood supply of the cremaster muscle is the cremasteric artery. The testicular artery, a branch of the aorta, descends in the mesorchium in the cranial aspect of the cord. The artery coils as it descends, and, when it reaches the cranial pole of the testis, it straightens. The testicular veins arise from the dorsal border of the testis, and, as they ascend, they coalesce into larger veins, which contain valves that prevent retrograde flow of blood. The veins divide and convolute to form the large, coiled pampiniform plexus, which constitutes the bulk of the spermatic cord and surrounds the coiled testicular artery. The veins coalesce within the abdomen to become the testicular vein.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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The lymphatic vessels of the testis and epididymis enter the medial iliac and lumbar lymph nodes. A plexus of autonomic and visceral sensory nerves accompany the blood vessels within the spermatic cord and innervate the testis and epididymis.
History and physical examination Information that may relate to problems of the spermatic cord should include history of fertility, unilateral or bilateral testicular retention, previous urogenital surgery, enlargement in the inguinal or scrotal areas, changes in testicular size, and signs of testicular pain. Physical examination of the spermatic cord and associated structures might include visual inspection, palpation, and ultrasonographic evaluation. The size, shape, texture, and temperature of the cords and testes should be observed, and right and left cords and testes should be compared. The horse should be observed for evidence of pain when the testes and cords are palpated. An enlargement in the spermatic cord or scrotum may suggest the presence of an inguinal hernia, hydrocele, hematocele, varicocele, or inflammation. To avoid evisceration after castration, the inguinal area of foals should be examined before castration for the presence of a congenital inguinal hernia. Palpation of an inguinal hernia may elicit a feeling of crepitus, and, occasionally, peristalsis of herniated intestine may cause movement of scrotal skin. Borborygmi may be heard during auscultation of the affected cord of a horse with an inguinal hernia. A ruptured inguinal hernia or an inguinal rupture should be suspected if the skin over an inguinal or scrotal swelling is cold and edematous or excoriated. A strangulated inguinal hernia or rupture should be suspected whenever a stallion develops signs of colic, especially if a testis and its spermatic cord are swollen and palpation elicits signs of pain. A hydrocele should be suspected if the scrotum and cord appear to be filled with fluid, especially if the testis is smaller than normal. The presence of fluid in the spermatic cord of a gelding is also suggestive of a hydrocele. A varicocele should be suspected if the spermatic cord appears enlarged and tortuous coils of vessels can be palpated. Examination per rectum Examining the vaginal rings and structures that pass through them, per rectum, may be useful in evaluating some abnormalities of the spermatic cord, such as inguinal cryptorchidism, inguinal or scrotal herniation, and septic funiculitis. The vaginal rings of a mature horse of average size are located about 6–8 cm cranial to the iliopectineal eminence and 10–12 cm
abaxial to the linea alba. The vaginal rings of stallions are generally large enough to accommodate one or two fingers, but those of geldings are generally palpable only as a slight depression. Before examining the vaginal rings per rectum, the value of information to be learned should be weighed against the risk of damaging the rectum. To palpate the vaginal rings, a hand is inserted into the rectum until the lateral aspect of the wrist rests on the symphyseal branch of the pubis. The tips of the flexed fingers are pressed against the abdominal wall and extended forward until a finger encounters the vaginal ring. Encountering a vaginal ring by flexing the fingers is difficult because the medial border of the deep inguinal ring tends to close, obscuring the vaginal ring. The ductus deferens can be palpated on the caudomedial aspect of the ring, provided that the testis or epididymis has descended. The testicular vasculature is difficult to palpate. The vaginal rings should be palpated per rectum for indication of intestinal incarceration when examining adult stallions with signs of intestinal obstruction. Palpating a distended loop of small intestine seemingly fixed to the body wall in the region where a vaginal ring should be found indicates that intestine has become incarcerated in the vaginal cavity. For horses that have developed infection of the spermatic cord after castration, enlargement of tissue surrounding the vaginal ring is evidence that infection in a cord has ascended close to the abdominal cavity. Palpation of the vaginal rings may help determine the location of an undescended testis (see Inguinal cryptorchidism). Other diagnostic tests Ultrasonographic examination may be useful in evaluating abnormalities of the spermatic cord or in identifying an inguinal testis. The echostructure of the spermatic cord can be examined adequately using a linear, 5-MHz transducer, but 7.5- or 10-MHz probes provide a more detailed examination.2 Color-flow Doppler ultrasonography is useful in evaluating distribution of blood vessels and measuring blood flow in the spermatic cord. Cytological examination of peritoneal fluid may be valuable in diagnosing certain diseases of the testis and spermatic cord, because changes in the fluid within the vaginal cavity may be reflected in the peritoneal fluid. For example, inspection of peritoneal fluid of a horse with a hematocele not associated with trauma of the testis or spermatic cord might reveal hemoperitoneum as the cause of hematocele. Sepsis of the spermatic cord might cause an increase in the nucleated cell count in the peritoneal fluid.
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Abnormalities of the Spermatic Cord
Inguinal herniation and rupture Inguinal herniation is the protrusion of intestine, usually a loop of the distal portion of the small intestine, or omentum through the vaginal ring into the vaginal cavity. If the herniated intestine descends into the scrotum, the condition is sometimes referred to as scrotal herniation, but the terms inguinal herniation and scrotal herniation are used interchangeably. Inguinal herniation is sometimes termed ‘indirect herniation’, a term used to describe a comparable condition in human beings.3 A ruptured inguinal hernia occurs when the vaginal sac ruptures, allowing the hernial contents of the vaginal cavity to pass subcutaneously into the scrotal or inguinal fascia. The intestine that protrudes through the rent may become strangulated or may adhere to subcutaneous tissue. Ruptured inguinal hernias occur primarily in foals. Inguinal rupture is the protrusion of intestine or omentum through a rent in the peritoneum and transverse fascia adjacent to the vaginal ring. Intestine invades fascia surrounding the vaginal sac. Inguinal rupture is sometimes erroneously referred to as ‘direct herniation’, a term used to describe herniation in human beings caused by weakening of the transverse fascia adjacent to the vaginal ring.3 The weakened fascia forms a peritoneum-lined sac. The alleged direct hernias of horses are actually ruptures, because tissue adjacent to the vaginal ring is torn, and the intestine passing through the tear is not surrounded by peritoneum.
Etiology Inguinal herniation and rupture occur predominately in stallions rather than in geldings because vaginal rings diminish in size soon after castration. Nevertheless, inguinal herniation of geldings has been reported.4, 5 Inguinal hernias of foals are congenital and usually hereditary.6 They may be caused by failure of the extra-abdominal part of the gubernaculum to regress, which results in an abnormally wide vaginal ring.1 Congenital inguinal hernias may occur unilaterally, in which case the left inguinal canal is most often involved, or bilaterally. They usually are accompanied by strangulation of the contents of the hernia, are reducible, and spontaneously resolve by the time the foal is 6 months old. A long-standing, congenital inguinal hernia may cause the testis to atrophy. In contrast, ruptured inguinal hernias of foals usually cannot be reduced, and the incarcerated intestine strangulates. A ruptured inguinal hernia is probably caused by compression of the foal’s abdomen during parturition.
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Inguinal hernias of adult stallions are usually acquired but result because the vaginal ring of the affected side is congenitally larger than normal. They often occur when the horse exercises or copulates, either of which activity may temporarily increase abdominal pressure and change the size or shape of the vaginal ring. Standardbreds and draft horses are reported to have a higher incidence of acquired inguinal herniation than other breeds,7, 8 and Tennessee walking horses and American Saddlebreds may also have an increased incidence of inguinal herniation. Inguinal rupture in males is traumatic in origin and, judging by the scarcity of reports of the condition, is quite rare.3 The intestine contained within an acquired inguinal hernia of an adult horse or that which has escaped through an inguinal rupture commonly strangulates causing a surgical emergency. Clinical signs associated with acquired inguinal hernias or inguinal ruptures, such as signs of severe pain, intestinal reflux into the stomach, and rectally palpable, distended small intestine, are similar to those caused by strangulating obstruction of the small intestine from other causes. Edema of the external genitalia and testicular degeneration or necrosis rapidly ensue. The constricting vaginal ring is responsible for strangulation of the herniated intestine and vasculature of the spermatic cord. Diagnosis Inguinal herniation or rupture can be diagnosed by history and clinical examination, including examination per rectum. The condition can be diagnosed definitively using ultrasonography by identifying loops of intestine within the vaginal cavity. The viability of the herniated intestine can be assessed ultrasonographically by examining the thickness of the intestinal wall and by observing the intestine for motility. Non-surgical management Congenital inguinal hernias often cause no distress, and most resolve spontaneously by the time the foal is 6 months old. Resolution may be hastened by frequent manual reduction of the hernial contents into the abdomen or by applying a truss after the hernia has been reduced. To apply a truss, the foal is sedated and positioned in dorsal recumbency. After reducing the hernia, a wad of rolled cotton is placed over the superficial inguinal ring of the affected side, and elastic gauze and tape are wrapped over the back and between the rear limbs in a figure-of-eight fashion. The bandage can be left in place for up to 1 week. Surgical reduction of a congenital inguinal hernia is necessary
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only if the hernial contents become incarcerated or if the hernia fails to resolve. Horses with an acquired inguinal hernia or an inguinal rupture nearly always require urgent surgical treatment because the intestine that has escaped from the abdominal cavity is likely to be strangulated. Traction on incarcerated intestine per rectum or external manipulation can be attempted to reduce an acquired inguinal hernia if herniation is diagnosed soon after its onset, but neither method of reduction should be attempted if the incarcerated intestine is likely to be non-viable, or if ruptured inguinal herniation or inguinal rupture cannot be ruled out as a cause of clinical signs. To replace the intestine contained within the spermatic cord back into the abdomen using external manipulation, the horse is sedated, and the testis of the affected side is grasped with one hand and pulled downward to tense the spermatic cord. The spermatic cord should be grasped with the other hand at the proximal aspect of the testis and should be slid proximally until the hernial contents are encountered. This hand maintains traction and compression on the cord to prevent intestine from moving distally. The testis is released by the first hand, which then grasps and compresses the cord above the second hand to force intestine proximally. The hands are exchanged in this manner until the contents of the hernia have been returned to the abdomen. Retracting the incarcerated loop of intestine into the abdomen by applying traction on it per rectum may be possible, particularly if only a small section of the incarcerated intestinal loop has entered the vaginal cavity and if herniation is diagnosed soon after its onset. The horse should be sedated, and local anesthetic solution administered into the epidural space to relax the rectum to reduce the risk of damaging the incarcerated intestine or the rectum. Systemic administration of N-butylscopolammonium bromide is also effective for relaxing the rectum. The horse should be observed closely for signs of septic peritonitis, in the event that the hernial contents were replaced into the abdomen without surgical intervention, because the reduced intestine may be non-viable. Intestinal viability should be monitored periodically by ultrasonographically examining the intestine for movement and thickness, by auscultating the abdomen for borborygmi, and by evaluating the horse’s peritoneal fluid obtained by abdominocentesis. Surgical management A congenital inguinal hernia must be surgically reduced if it fails to resolve, if it is so large that it is unlikely to resolve spontaneously, or if it becomes in-
carcerated within the vaginal cavity or within a tear in vaginal sac (i.e., a ruptured inguinal hernia). To surgically reduce an inguinal hernia, the foal is anesthetized and positioned in dorsal recumbency, and the inguinal region is prepared for surgery. The skin over the superficial inguinal ring of the affected side is incised, and, using blunt dissection, the vaginal sac, testis, and intestine contained within are bluntly isolated from surrounding fascia. The scrotal ligament, which attaches the vaginal sac to the scrotum, is transected to completely free the vaginal sac. The intestine contained within the vaginal sac is reduced into the abdomen. Tensing and twisting the spermatic cord may facilitate reduction of the intestine. The cord is ligated and transected close to the superficial inguinal ring. The ligature prevents intestine from escaping through the remnant of the spermatic cord. As an additional precaution against evisceration, the superficial inguinal ring can be sutured. The cutaneous incision can be sutured or left open to heal by secondary intention. Resolving congenital inguinal hernias of foals laparoscopically allows the foal to return rapidly to normal activity.9, 10 With the foal anesthetized and positioned in dorsal recumbency, and using laparoscopic equipment, herniated intestine and the testis are retracted from the vaginal sac into the abdominal cavity. The testis is removed, after the testicular vessels and ductus deferens are ligated and transected, and the vaginal ring is closed using a laparoscopic stapling device. The testis of the contralateral side can also be retracted into the abdominal cavity and removed laparoscopically at the same time. Ruptured inguinal hernias and inguinal ruptures must always be surgically corrected. Surgery is usually indicated to correct an acquired inguinal hernia and is always indicated if the viability of the testis or incarcerated intestine is doubtful because surgery allows these structures to be assessed visually. The horse should receive fluids intravenously in volumes sufficient to combat shock. The horse is positioned in dorsal recumbency, and the scrotal and inguinal areas and the ventral aspect of the abdomen are prepared for surgery. The skin over the superficial inguinal ring of the affected side is incised, and, using blunt dissection, the vaginal sac and its contents are bluntly isolated from surrounding fascia. The testis and herniated intestine are exposed through an incision in the vaginal sac. Applying traction to the intestine through a celiotomy and dilating or blindly incising the vaginal ring may be necessary to reduce the herniated intestine. Devitalized intestine is usually more easily transected and anastomosed through a celiotomy than through the incision in the vaginal sac. The testis should be excised if it appears to be infarcted. If the viability of the affected testis is
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Abnormalities of the Spermatic Cord questionable, the testis can be left in situ, even if the testis is totally infarcted, but re-herniation can be more easily prevented if the testis is removed.11, 12 Escape of intestine into the inguinal canal can be prevented by ligating the spermatic cord as close to the superficial inguinal ring as possible with heavy, absorbable sutures, and, for added safety, the superficial inguinal ring can be closed with heavy absorbable sutures. The cutaneous incision over the superficial inguinal ring can be sutured or left open to heal by second intention. The celiotomy is closed in a routine manner. To preserve a viable testis, the superficial inguinal ring can be partially sutured around the spermatic cord to prevent escape of intestine from the inguinal canal.13 The cranial half of the ring is sutured toward the caudally located spermatic cord. The ring must be closed adequately to prevent escape of intestine from the inguinal canal while still maintaining an adequate blood supply to the testis. A method of salvaging the testis, while at the same time preventing escape of intestine through the enlarged vaginal ring, is to laparoscopically implant a mesh beneath the peritoneum over the deep inguinal ring, with the horse anesthetized and in dorsal recumbency. This procedure is performed after the horse has recovered from previous surgery to reduce the hernia.14 Another laparoscopic technique of inguinal herniorrhaphy used in an attempt to salvage the testis involves inserting a coiled mesh into the vaginal ring, with the horse standing, and fixing the mesh to the ring with endoscopic staples.15 The vaginal ring is obliterated by fibrous reaction to the mesh.
Inguinal cryptorchidism Location of one or both testes other than in the scrotum is referred to as cryptorchidism. The undescended testis, which may be located abdominally or inguinally, is termed an undescended testis, and, by extension, a horse with, an undescended testis is referred to as a cryptorchid stallion. An inguinal cryptorchid is a horse with a testis retained within the inguinal canal. A colloquialism used to designate a horse with an inguinally located testis is ‘high flanker’. A retractile testis, which is a testis that can be palpated in the inguinal area but manipulated into the scrotum, should be distinguished from a truly inguinal undescended testis. Retraction of the cremaster muscle may maintain the retractile testis in a position away from the scrotum until puberty, when growth of the testis causes the testis to reside permanently in the scrotum. Stallions with two undescended testes are sterile because the seminiferous tubules of an undescended testis, regardless of its location, are hypoplastic and
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incapable of producing sperm. Exposure of the testis to a temperature higher than that of the scrotum is responsible for the hypoplasia. If only one testis is undescended, the stallion is fertile, even though the affected stallion usually produces less sperm than a stallion with two descended testes. A stallion with two undescended testes still exhibits libido because the androgen-producing cells of Leydig of an undescended testis remain functional and produce testosterone. Events that lead to failure of testicular descent in horses have not been determined, but, in boys, impaired intrauterine secretion of gonadotropins has been shown to result in cryptorchidism by leading to insufficient testosterone and dihydrotestosterone production.16 Production of androgens by the descending testes is responsible for rapid growth of the vaginal process, testicular vessels, and ductus deferens, and for changes in the gubernaculum, all of which are necessary for proper descent of the testes into the scrotum. Although cryptorchidism of horses is generally considered to be hereditary, no studies support a credible genetic mechanism, and the effects of nongenetic factors in the development of cryptorchidism have not been evaluated. A reviewer of this topic stated that one German study of cryptorchid stallions concluded that equine cryptorchidism is transmitted as an autosomal dominant gene, whereas another German study hypothesized that at least two genetic factors are involved, one of which is located on the sex chromosomes.17 The contradictory observations of the heritability of equine cryptorchidism suggest that the mechanism of inheritance is complex and may involve several genes. Given the data presented, the heritability of cryptorchidism remains an enigma. Prevalence and incidence An epidemiological investigation of the prevalence of equine cryptorchidism found that one out of six stallions aged 2–3 years presented to veterinary medical teaching hospitals had cryptorchidism.18 American Quarter Horses, Percherons, American Saddlebreds, ponies, and crossbred horses had the highest prevalence of cryptorchidism, and the Thoroughbred breed had the lowest prevalence. The right and left testes of horses fail to descend with nearly equal frequency.19–21 A study of a large number of cryptorchid horses found that 25% of left and 58% of right undescended testes were located inguinally.21 This study did not examine the effect of age on the incidence of right versus left testicular retention. The incidence of right retention may decrease as the population of cryptorchid horses ages because an increase in testicular size with the onset of puberty
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Stallion: Abnormalities of the Stallion’s Reproductive Tract
may cause an inguinally retained testis to descend into the scrotum.19, 20 Diagnosis Cryptorchidism is recognized by the absence of one or both testes in the scrotum, but, before declaring a horse to be a cryptorchid, especially a prepubescent stallion whose testes are often small and retractile, the scrotal and inguinal regions should be palpated carefully. To facilitate palpation of the inguinal region, the horse should receive a sedative to relax the cremaster muscles. A testis in the proximal part of the canal may escape detection during palpation because the canal of an average-sized stallion is approximately 15 cm deep. The vaginal rings can be palpated only by examination per rectum. Examining the vaginal rings per rectum may be useful in determining whether an undescended testis has entered the inguinal canal. Neither the vaginal ring nor the ductus deferens, which enters the vaginal ring, can be palpated in horses with complete abdominal retention of the ipsilateral testis. If the vaginal ring and the ductus deferens entering it can be palpated, the testis or at least its epididymis must have descended through the ring into the inguinal canal. Unfortunately, a horse with a testis that is retained within the abdomen but whose epididymis has descended through the vaginal ring cannot be distinguished from a horse whose testis has descended through the vaginal ring when examining the vaginal rings per rectum. An inguinal testis appears ultrasonographically as a homogeneous, granular, round structure surrounded by an echogenic capsule. To ultrasonographically image an inguinal testis, a 5-MHz rectal transducer is placed longitudinally over the superficial inguinal ring. When using a 5-MHz transducer, inguinal ultrasonography is unsuccessful in locating an abdominal testis, and transrectal ultrasonography is ineffective in locating an inguinal testis.22 Treatment Undescended inguinal testes of boys have been fixed in a scrotal position using sutures,23 but because cryptorchidism of horses is often considered to be hereditary, bringing about descent of an inguinal testis by surgical means is considered by most practitioners to be disreputable. There is good evidence that an impaired hypothalamic–pituitary–gonadal axis, which leads to insufficient production of testosterone and dihydrotestosterone, is the main cause of cryptorchidism in boys, and so boys with one or more undescended testes have been treated with good success using gonadotropin-releasing hormone (GnRH), human chorionic gonadotropin (hCG), or luteinizing hormone releasing hormone (LHRH).16, 24
Administering hormones to a cryptorchid horse to promote testicular descent seems to be a logical treatment, but using hormonal therapy to bring about testicular descent would seem no more ethical than using surgical therapy. Hormonal events involved in testicular descent in the horse are poorly understood, and so no specific hormonal therapy has been developed to bring about descent of an undescended testis. GnRH and hCG or LHRH have been administered separately or in combination to stallions to bring about testicular descent, but the efficacy of this therapy has not been established. Treatment with hCG was thought to facilitate descent of inguinally located testes in four out of eight colts in which one testis was identified in the inguinal canal before initiation of treatment.25 Increase in testicular size after the onset of puberty may cause an inguinal testis to descend, and this supports the rationale for administration of GnRH or hCG to horses with an inguinal testis. The optimal time to effect descent of an undescended testis in affected boys is the second year of life because, after this time, the seminiferous tubules of the testis may become irreparably damaged from the abnormally warm environment of the extrascrotal position.26 The age of the horse by which an inguinal testis must descend into the scrotum to avoid irreversible damage to the seminiferous tubules is not known. Hormonal therapy probably has no effect on descent of an abdominal testis because, by several weeks after birth, the size of the vaginal rings has diminished to the extent that the descent of an abdominal testis becomes impossible.27 An inguinal testis can be removed through a scrotal or an inguinal approach. The inguinal fascia is separated bluntly to expose the superficial inguinal ring. An inguinal testis is located easily during inguinal exploration, but an inexperienced surgeon may encounter and damage the shaft of the penis. The vaginal sac should always be opened and its contents examined closely to avoid mistaking the descended tail of the epididymis of an abdominal testis for a small inguinal testis. Rarely, one or both descended testes of a stallion may retract into the inguinal area, apparently from sustained contraction of the cremaster muscle caused by pain of genital or musculoskeletal origin. The position of the retracted testes can result in decreased spermatogenesis, which, in turn, may lead to subfertility or infertility. In an effort to restore the stallion’s semen quality, the testes can be returned to a more normal position in the scrotum by transecting the cremaster muscle which attaches to an affected spermatic cord. To transect the cremaster muscle(s), the horse is anesthetized, positioned in dorsal recumbency, and the inguinal regions are prepared for aseptic surgery. Each spermatic cord is exposed through a
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Abnormalities of the Spermatic Cord 5- to 6-cm cutaneous incision created over its longitudinal axis. A 3-cm section of cremaster muscle is bluntly separated from the parietal tunic of the cord and excised with scissors. The incision is closed in two layers. The effect of surgery on the position of the testes is immediate, and improved spermatogenesis may be observed in about 3 months.
Torsion of the spermatic cord Torsion of the spermatic cord, sometimes improperly referred to as testicular torsion, occurs when the testis rotates on its vertical axis. Extensive torsion of the cord results in occlusion of the testicular vasculature, leading to testicular congestion and edema if mild and complete testicular necrosis if severe. The entire spermatic cord may be twisted (i.e., extravaginal torsion) or the contents of the cord may twist within the vaginal tunic (i.e., intravaginal torsion). Although extravaginal torsion of the spermatic cord of a horse whose testes have descended into the scrotum has not been reported, we have identified, in some stallions, what appeared to be 180◦ extravaginal torsion of the spermatic cord. Torsion of the spermatic cord of the descended testis of horses occurs with highest frequency within the vaginal tunic and occurs because either the proper ligament of the testis or the ligament of the tail of the epididymis is unusually long. Contraction of the cremaster muscle may initiate torsion of a spermatic cord susceptible to torsion. The same ligament(s) of the contralateral testis may also be unusually long, placing that spermatic cord at risk of torsion. Clinically significant torsion of the spermatic cord in the horse is rare, based on the scarcity of reports of the condition. Diagnosis Torsion of the spermatic cord of stallions of 180◦ or less may cause no clinically apparent abnormalities, but ultrasonographic evaluation of testicular blood flow shows that torsion of a spermatic cord of 180◦ may cause retrograde blood flow, which indicates that torsion of 180◦ may affect testicular function detrimentally.28 Stallions with mild torsion of one or both spermatic cords (i.e., 180◦ ) have a lower gelfree and total seminal volumes and fewer spermatozoa in second ejaculates than do normal stallions, and motile spermatozoa and morphologically normal spermatozoa in first ejaculates are significantly depressed.29 The reason for fewer spermatozoa in the extragonadal reserves of sperm of stallions with 180◦ torsion of the spermatic cord is unknown. When the spermatic cord is twisted 180◦ , the tail of the epididymis lies in the cranial portion of the scrotum, rather than in its normal caudal position in the
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scrotum. The testes of young stallions are labile and hence are more apt to be rotated than are those of mature stallions (i.e., those ≥ 4 years old). If a testis of a mature stallion is observed to be rotated from its normal position, it usually remains in that position.29 Torsion of the spermatic cord of 360◦ or more causes the affected testis and cord to enlarge from edema and induces signs of acute pain, which may mimic signs of intestinal pain. Signs of pain may diminish as testicular necrosis progresses. When the spermatic cord is twisted 360◦ , the tail of the epididymis resides in its normal caudolateral position in the scrotum, but torsion of this degree may be accompanied by so much scrotal, testicular, and epididymal swelling that the epididymis cannot be palpated. Confirming the diagnosis of torsion of the spermatic cord or differentiating the condition from other causes of enlargement of the testis and cord may be difficult. If a horse shows signs of strangulating torsion of the cord, the cord should be surgically explored as soon as possible. Conditions that may resemble torsion of the spermatic cord include inguinal herniation and orchitis. A rare condition that may closely mimic torsion of the spermatic cord is thrombosis of the testicular artery. Torsion of the spermatic cord of an abdominal testis may cause the affected horse to show moderate to severe signs of abdominal pain. Some affected horses, however, may show no signs of abdominal pain, and, so, the condition may be discovered only when the necrotic or severely atrophied testis is removed from the abdomen during castration. Treatment Torsion of the cord of 360◦ usually requires removal of the affected testis, because the testis can rarely be restored to normal function. If the viability of the affected testis is questionable, the testis can be left in situ after the torsion is corrected and the testis fixed to the scrotum because sepsis occurs rarely, even if the testis is totally infarcted. One must be mindful, however, about the possibility of formation of anti-sperm antibodies that could adversely affect the function of the contralateral testis. Torsion of the contralateral spermatic cord can be avoided by permanently anchoring the contralateral testis in its normal position with non-absorbable sutures placed through the tunica albuginea and the dartos tissue at the caudal pole of the testis. The spermatic cord of an abdominal testis may be at more of a risk of developing torsion than is the spermatic cord of a descended testis, because the ligaments of the testis and epididymis of an abdominal testis are usually greatly elongated, allowing the testis freedom to rotate.
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Stallion: Abnormalities of the Stallion’s Reproductive Tract
Hydrocele (vaginocele) A hydrocele, or vaginocele, is an abnormal accumulation of serous fluid between the visceral and parietal layers of the tunica vaginalis. It may arise insidiously or acutely. Because the vaginal and peritoneal cavities communicate, some hydroceles may be caused when excess fluid in the abdominal cavity caused by disease descends into the vaginal cavity, but most hydroceles probably occur when fluid, which is normally secreted by the vaginal tunic, is produced at a rate greater than that at which it can be absorbed by the lymphatic vessels of the spermatic cord. The cause of the disparity between the rates of production and resorption of the fluid is usually idiopathic, but proposed causes of increased rate of production of fluid include viral-mediated serositis and traumainduced serositis. Migration of parasites through the vaginal cavity may interfere with lymphatic resorption of fluid. Other proposed causes of hydrocele include 180◦ torsion of the spermatic cord. Horses that live in a hot climate and that are denied exercise may be at risk of developing a hydrocele. Hot weather may contribute to formation of a hydrocele by causing the testes to descend maximally in the scrotum to dissipate heat, which, in turn, may impair lymphatic or venous circulation.30 A hydrocele may inhibit spermatogenesis by insulating the testis or by impinging on the testicular vasculature, and increased scrotal temperature caused by the hydrocele may depress spermatogenesis. Even if the hydrocele is unilateral, the insulating properties may be sufficient to impede spermatogenesis in the contralateral testis. Interestingly, one investigation found that hydroceles of stallions had no significant effect on semen quality.30 The prognosis for fertility may depend on the severity and duration of the condition.
Diagnosis Palpation of the affected spermatic cord and testis usually causes the horse no apparent discomfort. Ultrasonographic examination of the testis and spermatic cord is required to confirm the diagnosis. A hydrocele appears ultrasonographically as an abnormal concentration of anechoic fluid within the vaginal cavity. Ultrasonography may identify associated abnormalities in the testis or cord, such as a tumor, and can be used to differentiate a hydrocele from similar appearing abnormalities, such as varicocele, inguinal herniation, or hematocele. Anechoic fluid is observed surrounding the testis and epididymis during ultrasonographic examination of the scrotum and spermatic cord, and the involved testis of a chronically affected horse may appear to be abnormally small.
Horses chronically affected with hydrocele may develop fibrin strands within the vaginal cavity, giving the vaginal fluid a mixed, echogenic appearance during ultrasonographic examination. Clear, yellow fluid can be aspirated from the vaginal cavity, and this fluid often reforms quickly after aspiration. Clinical chemical analysis of the aspirated fluid rarely produces useful information, but we have diagnosed mesothelioma in two stallions with hydrocele, based on results of cytological examination of the aspirate. The diagnostic value of aspirating the vaginal cavity should be weighed against the slight risk of introducing infection into the vaginal cavity. Treatment A hydrocele may resolve spontaneously when the affected horse is moved to a cooler environment, or it may reduce temporarily in size when the horse exercises.30 Treatment of a stallion for hydrocele should be aimed at eliminating the inciting cause, but, because the cause is usually idiopathic, stallions affected with persistent, unilateral hydrocele are usually treated by removing the affected testis and as much of the spermatic cord as possible, before spermatogenesis of the contralateral testis becomes affected by the insulating effect of the hydrocele. A stallion that is persistently and bilaterally affected has a guarded prognosis for fertility, but some bilaterally affected stallions may be able to maintain normal spermatogenesis.30 Sclerotherapy, performed by injection of various concentrations of tetracycline or polidocanol into the vaginal cavity, and plication or removal of redundant portions of the vaginal tunic have been used successfully to resolve hydroceles in men,31, 32 but these forms of therapy have not been evaluated as a treatment for similarly affected stallions. Injection of a sclerosing agent into the vaginal cavity of men affected with hydrocele may result in decreased fertility.33 A hydrocele can be an uncommon, idiopathic complication of castration, and is more likely to occur with open castration because, with this technique of castration, the vaginal tunic is not removed. Mules may be at more risk than horses to develop a hydrocele after castration. Accumulation of fluid in the scrotum may cause the horse to appear to have an inguinal hernia or a scrotal testis. The fluid can often be squeezed from the vaginal sac into the abdomen. A hydrocele that forms after castration can usually be easily excised. Drainage relieves the condition only temporarily. To excise a hydrocele, the horse is anesthetized, positioned in dorsal recumbency, and prepared for surgery of the scrotal region. An elliptical portion of skin over the fluid-filled parietal
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Abnormalities of the Spermatic Cord tunic is excised, being careful not to penetrate the tunic. The parietal tunic is bluntly freed from fascia, and the scrotal ligament, which attaches the distal aspect of the parietal tunic to the scrotum, is severed. The spermatic cord proximal to the hydrocele is transected, using a scissors or an emasculator. The scrotal incision is sutured or left open to heal by secondary intention.
Hematocele A hematocele is a collection of blood within the vaginal cavity. Blood in the vaginal cavity most commonly occurs from testicular or scrotal trauma, but it can also enter from the abdominal cavity through the vaginal ring. A large blood clot in the vaginal cavity may cause adhesions to form between the visceral and parietal tunics. A hematocele may reduce spermatogenesis by insulating not only the testis of the affected side, but also the contralateral testis.
Diagnosis The condition can be differentiated from hydrocele or other causes of enlargement of the spermatic cord by history, ultrasonographic examination of the spermatic cord and testis, and aseptic aspiration of the contents of the vaginal cavity. During ultrasonographic examination of a hematocele, the vaginal cavity is seen to contain tissue that has a homogeneous echogenic appearance that increases in echogenicity as the blood clot organizes.
Treatment Aspirating blood from the vaginal cavity followed by lavage of the cavity with a balanced electrolyte solution may bring about resolution of the hematocele, provided that hemorrhage into the vaginal cavity has ceased. Hemorrhage from the testis caused by tearing of the tunica albuginea can be stopped by repairing the tear with sutures, but the testis should be excised if it has been severely damaged.
Varicocele A varicocele is dilation of the pampiniform plexus caused by increased hydrostatic pressure in this structure, which, in turn, is caused by incompetence of the testicular vein, where it empties into the vena cava or renal vein. The condition is found in a large percentage of men with normal fertility and in an even larger percentage of infertile men,34 but it is rarely observed in stallions. Stallions with a unilat-
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eral varicocele may have normal seminal quantity and quality. Although varicoceles are a common cause of poor sperm production and decreased semen quality in men,34 their effect on the quantity and quality of semen production of horses has not been evaluated. Varicoceles of men are associated with low total sperm count, but not with abnormal motility or morphology of spermatozoa. The mechanisms by which fertility is affected have not been satisfactorily explained. Spermatogenesis might be affected by impaired drainage of blood from the testis, leading to increased scrotal temperature, hypoxia, and increased testicular pressure.34
Diagnosis The affected spermatic cord has the texture of a ‘bag of worms’, and the affected spermatic cord may be wider than usual. Palpation of the affected spermatic cord does not cause the horse to display signs of pain. A varicocele appears ultrasonographically as circumscribed, uniform, anechoic areas within the spermatic cord. A characteristic reverse flow of blood in a varicocele may be demonstrated by color-flow Doppler ultrasonography.
Treatment A common treatment of men for infertility caused by a varicocele is ligation of the dilated vasculature,35 but there is no good evidence that treatment of subfertile men with a clinical varicocele improves the likelihood of conception.34 Treatment of subfertile stallions for varicocele has not been described, but treatment could entail excision of the affected testis and cord.
Septic funiculitis Septic funiculitis, or infection of the spermatic cord, is an uncommon complication of castration that can occur from extension of a scrotal infection or from a contaminated emasculator or ligature. The condition was evidently more commonly encountered before the advent of the emasculator, when hemorrhage from transection of the cord was controlled by clams. The infection is contained primarily within the vaginal cavity, and so the open method of castration, in which the parietal tunic is not removed, is far more likely than the closed method of castration to result in septic funiculitis. The condition is most commonly known as ‘scirrhous cord’.
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Diagnosis The infected cord fills with granulation tissue and exudate. The cord hardens and adheres to scrotal skin, exudate may discharge through cutaneous tracts, and the horse may suffer from preputial and scrotal edema and pyrexia. The horse may become lame if the stump of the cord expands to the extent that it mechanically interferes with locomotion. Rarely, the infection may progress proximally to involve the full length of the cord, causing a hard mass to be palpated per rectum at the deep inguinal ring. A chronically infected cord after castration has a heterogeneous, echogenic appearance, and the parietal tunic of the cord is thickened. The lesion may not become apparent for years after castration. Treatment Septic funiculitis is unlikely to resolve with antimicrobial therapy, and so removal of the infected stump is indicated. To remove the infected mass, the horse is anesthetized, positioned in dorsal recumbency, and prepared for surgery at the scrotum and affected inguinal region. Skin over the superficial inguinal ring is incised, and, using sharp and blunt dissection, the infected stump is isolated from adjacent tissue. Isolating an infected stump within a few weeks after castration may be no more difficult than isolating a spermatic cord at the time of castration, but isolating a chronically infected cord is often difficult and timeconsuming because the parietal tunic becomes adherent to surrounding fibrous tissue, and the stump becomes very vascular. The stump of the cord is transected proximal to the mass, using an emasculator, and the wound is left unsutured to heal by secondary intention. An e´ craseur is more easily applied proximal to a mass that extends deep into the inguinal canal than is an emasculator. The use of ligatures should be avoided.
References ¨ S. Anatomy of the Horse, 2nd 1. Budras K-D, Sack WO, Rock edn. London: Mosby-Wolfe, 1994. 2. Pozor MA. How to evaluate a stallion’s scrotum using ultrasound. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2005, pp. 288–93. 3. Cox JE. Hernias and ruptures: words to the heat of deeds. Equine Vet J 1988;20:1–156. 4. Bickers RJ, Lewis RD, Hays T, Noble JK. Acquired inguinal hernia in a gelding. Equine Pract 1998;20:28–9. 5. Schneider RK, Milne DW, Kohn CW. Acquired inguinal hernia in the horse: A review of 27 cases. J Am Vet Med Assoc 1982;180:317–20. 6. Roberts SJ. Veterinary Obstetrics and Genital Diseases, 2nd edn. Ithaca, NY: self-published, 1971.
7. Sembrat RF. The acute abdomen in the horse: Epidemiologic considerations. Arch Am Coll Vet Surg 1975;4:34–9. 8. Shoemaker R, Bailey J, Janzen E, Wilson DG. Routine castration in 568 draught colts: incidence of evisceration and omental herniation. Equine Vet J 2004;36:336–40. 9. Klohnen A, Wilson DG. Laparoscopic repair of scrotal hernia in two foals. Vet Surg 1996;25:414–16. 10. Marien T, van Hoeck F, Adriaenssen A, Segers L. Laparoscopic testis-sparing herniorrhaphy: a new approach for congenital inguinal hernia repair in the foal. Equine Vet Educ 2001;13:32–5. 11. Howard SS. Surgery of the scrotum and testes in childhood. In: Walsh PC, Retik AB, Stamey TA, Darracott Vaughan Jr E (eds) Campbell’s Urology, 2nd edn. Philadelphia: W.B. Saunders, 1992; pp. 1939–50. 12. Skoglund RW, McRoberts JW, Ragde H. Torsion of the spermatic cord: a review of the literature and an analysis of 70 new cases. J Urol 1970;104:604–7. 13. Vaughan JT. Surgery of the testes. In: Walker DF, Vaughan JT (eds) Bovine and Equine Urogenital Surgery. Philadelphia: Lea & Febiger, 1980; pp. 145–69. 14. Fischer Jr AT, Vachon AM, Klein SR. Laparoscopic inguinal herniorrhaphy in two stallions. J Am Vet Med Assoc 1995;207:1599–601. 15. Marien T. Standing laparoscopic herniorrhaphy in stallions using cylindrical polypropylene mesh prosthesis. Equine Vet J 2001;33:91–6. 16. Hadziselimovic F. Pathogenesis and treatment of undescended testes. Eur J Pediatr 1982;139:255–65. 17. Leipold HW, De Bowes RM, Bennett S, Cox JH, Clem MF. Cryptorchidism in the horse: genetic implications. In: Proceedings of the 31st Annual Convention of the American Association of Equine Practitioners, 1985, pp. 579–89. 18. Hayes HM. Epidemiological features of 5009 cases of equine cryptorchidism. Equine Vet J 1986;18:468–71. 19. Coryn M, De Moor A, Bouters R, Vandeplassche M. Clinical, morphological and endocrinological aspects of cryptorchidism in the horse. Theriogenology 1981;16:489–96. 20. Cox JE, Edwards GB, Neal PA. An analysis of 500 cases of equine cryptorchidism. Equine Vet J 1979;11:113–16. 21. Stickle RL, Fessler JF. Retrospective study of 350 cases of equine cryptorchidism. J Am Vet Med Assoc 1978;172:343–6. 22. Jann HW, Rains JR. Diagnostic ultrasonography for evaluation of cryptorchidism in horses. J Am Vet Med Assoc 1990;196:297–300. 23. Molenaar JC. Surgical treatment of undescended testes. Eur J Pediatr 1982;139:289–91. 24. Hagberg S, Westphal O. Treatment of undescended testes in intranasal application of synthetic LH-RH. Eur J Pediatr 1982;139:285–8. 25. Brendemuehl JP. Effects of repeated hCG administration on serum testosterone and testicular descent in prepubertal thoroughbred colts with cryptorchid testicles. In: Proceedings of the 52nd Annual Convention of the American Association of Equine Practitioners, 2006, pp. 381–3. 26. Bierich JR. Undescended testes: Treatment with gonadotrophin. Eur J Pediatr 1982;139; 275–9. 27. Bergin WC, Gier HT, Marion GB, Coffman JR. A developmental concept of equine cryptorchidism. Biol Reprod 1970;3:82–92. 28. Pozor MA, McDonnell SM. Color Doppler ultrasound evaluation of testicular blood flow in stallions. Theriogenology 2004;61:799–810. 29. Pickett BW, Voss JL, Bowen RA, Squires EL, McKinnon AO. Seminal characteristics and total scrotal width (T.S.W.) of
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Abnormalities of the Spermatic Cord normal and abnormal stallions. In: Proceedings of the 33rd Annual Convention of the American Association of Equine Practitioners, 1987, pp. 487–518. 30. Henry M, Amaral D, Tavares FF, Santos NR. Hydrocele of the vaginal cavity of stallions. J Reprod Fertil Suppl 2000;56:13. 31. Hu K-N, Khan AS, Gonder M. Sclerotherapy with tetracycline solution for hydrocele. Urology 1984;24:572–6. 32. Daehlin L, Tonder B, Kapstad L. Comparison of polidocanol and tetracycline in the sclerotherapy of testicu-
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lar hydrocele and epididymal cyst. Br J Urol 1997;80:468– 71. 33. Osegbe DN. Fertility after sclerotherapy for hydrocele. Lancet 1991;337:172. 34. Evers JL, Collins JA. Assessment of efficacy of varicocele repair for male subfertility: a systemic review. Lancet 2003;361:1838–9. 35. Szabo R, Kressler R. Hydrocele following internal spermatic vein ligation: A retrospective study and review of the literature, J Urol 1984;132:924–5.
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Chapter 111
Epididymal Abnormalities Aime K. Johnson and Charles C. Love
The epididymis consists of a single convoluted unbranched tubule 70–80 meters long that begins as the efferent ducts of the testis converge (rete testes) and ends as the ductus deferens. A complicated biochemical and physiological change occurs in the sperm cell as it passes through this single tubule, resulting in a cell capable of motility and fertilization.
Anatomy Grossly, the epididymis consists of three major portions: the head (caput), body (corpus), and tail (cauda). The stallion epididymis lies on the dorsolateral aspect of the testicle with the caput firmly attached to the cranial pole of the testicle, the corpus more loosely attached and oriented laterally across the dorsal surface of the testicle, and connecting to the cauda which is also firmly attached to the caudal pole of the testicle by the proper ligament of the testis, a remnant of the gubernaculum. Another gubernaculular remnant, the ligament of the tail of the epididymis, attaches the cauda epididymis to the parietal vaginal tunic.
Microscopic anatomy Histologically, the epididymis varies greatly between the three segments and is related to the function of each segment. The caput epididymis contains the distal segments of the 13–15 efferent ducts and is lined by ciliated and non-ciliated pseudostratified columnar epithelium. Ciliated cells are responsible for the movement of sperm cells from the efferent ducts into the ductus epididymis and contain concentrations of mitochondria in the apical portion of the cytoplasm.1 Specialized non-ciliated cells contain numerous small vesicles and multivesicular bodies along with a unique arrangement of rough endoplasmic reticulum
around the basal and perinuclear region, and smooth endoplasmic reticulum and Golgi bodies around the supranuclear regions.1, 2 The arrangement and density of these organelles are associated with a secretory function. The presence of endocytotic invaginations located in the apical membrane of the non-ciliated cells is consistent with an absorptive function. Peroxisomes were also observed near the rough endoplasmic reticulum and appear to be important in reducing toxic effects resulting from peroxides that are produced during the degradation of sperm cells.2 Phagocytosis of sperm cells has been noted during examination with electron microscopy.2, 3 This function and these organelles are unique to the caput epididymis.2 Transition from the efferent ductules and proximal caput into the corpus and remainder of the ductus epididymis is characterized by the disappearance of ciliated cells and an increased height of the columnar epithelial cells. Stereocilia covering the luminal surface increase surface area and absorptive capabilities of these columnar cells. The thin layer of smooth muscle surrounding the tubule in the corpus becomes progressively thicker distally, and is dramatically thickened in the cauda epididymis. The lumen of the proximal ductus is narrow owing to the tall columnar epithelium, but becomes progressively wider as the height of the epithelium decreases and allows the accumulation of sperm within the lumen (see Chapter 97).
Function The epididymis serves many functions other than simple spermatozoa transport from the testicle to ductus deferens. Sperm cells entering the caput epididymis are incapable of fertilization and must undergo a maturation process prior to ejaculation. Excess fluids necessary for carrying sperm cells from
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Epididymal Abnormalities the testicle’s efferent ducts into the epididymis are resorbed to concentrate the cells within the lumen. The combined secretory and absorptive properties of the corpus epididymis are essential for the maturation process of the spermatozoa. As the cells move from the efferent ductules through the epididymal duct, they undergo numerous changes including translocation of the cytoplasmic droplet from its proximal location next to the head to a distal location on the midpiece; motility acquisition; and specific changes in their plasma membrane enabling the sperm to undergo fertilization. A blood–epididymis barrier allows a specialized luminal environment ideal for sperm maturation.4 The spermatozoa are grossly similar throughout the epididymis, with the exception of the location of the cytoplasmic droplet (see Chapter 97). This droplet is consistently located proximally next to the head in the caput region. As the spermatozoa move distally through the corpus and into the proximal cauda, the droplet translocates distally and is usually associated with a transient bending of the midpiece. Most droplets are shed entirely prior to, or at, the time of ejaculation. Spermatozoa are transported through the epididymal duct primarily by smooth muscle contractions (longitudinal and circular) and are assisted by ciliary action. The time required for transit in the stallion averages 7.5–11 days (mean 10) but can be as short as 4 days (mean 4.9) with increased ejaculations.5 However, the transit time through the caput and corpus is constant and not affected by the frequency of ejaculation, age, or season. The cauda epididymis serves as extragonadal storage for spermatozoa and therefore the frequency of ejaculations affects the amount of time a sperm cell remains in this holding area. Almost 90% of extragonadal sperm reserves are located within the epididymides, with 60% concentrated in the cauda portion.5 Spermatozoa that are not ejaculated are intermittently released into the urethra and are voided with urine. Phagocytosis of spermatozoa within the cauda epididymis is not thought to occur.
Examination Palpation The epididymal head is closely apposed to the craniolateral testis surface and is soft and difficult to identify manually. The epididymal body, a firmer structure, is palpable along the dorsolateral aspect of the testicle. The epididymal tail is readily palpable as a distinct spherical bulge at the caudal-most aspect of the testis. The location of the cauda can be used to
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assist the clinician in determining the proper orientation of the testis in cases of suspected spermatic cord rotation. It should be noted that > 180◦ rotations may result in the ‘correct’ orientation of the tail, but this condition should also be associated with additional clinical signs (i.e., swelling and pain of the scrotum) (see Chapter 110). In the case of a 180◦ spermatic cord torsion, the cauda may be pulled dorsally owing to the pull of the ductus deferens that has been wrapped around the spermatic cord. While connected to both the proper ligament of the testis and the ligament of the tail of the epididymis, the epididymal tail may be either tightly or loosely attached to the testis. When loose, the tail may move considerably, especially dorsally, making its presence more difficult to determine. Ultrasonography The ultrasonographic appearance of the epididymal tail is a ‘Swiss cheese’ pattern owing to the convoluted ducts distended with sperm.6 Prominent dilation of this duct can be due to chromic accumulation of sperm or may indicate a disease process such as epididymitis.
Disease Congenital Congenital hypoplasia of the epididymis has been reported in many domesticated species. The animal generally presents for azoospermia owing to bilateral hypoplasia and incomplete patency of the duct. Although rare in stallions, this condition has been described in a stallion with normal libido and mounting behavior.7 Diagnosis of epididymal hypoplasia is made following confirmation of azoospermia with normal testicular histology. In addition, the tail will be difficult to palpate caudally, but this must be differentiated from a loose cauda that is just positioned dorsally. There is no treatment and it is suspected to have a hereditary component in bulls.8 Although rare in stallions, epididymal cysts may occur in blind-ended efferent tubules. Secretions accumulate causing dilation within the epididymis. Diagnosis can be made using palpation and confirmed by ultrasound. Cysts tend to occur around the head of the epididymis but can occur in other locations (Figures 111.1 and 111.2). Epididymal cysts range in size from non-detectable with the ultrasound to over 7 cm in diameter.9 No treatment is usually necessary; however, because these blind ductules communicate with the normal tubal system, they may become impacted with sperm and lead to formation of a sperm granuloma.9
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Figure 111.1 A cyst-like structure on the dorsomedial aspect of the right testis (arrow). These structures may or may not be continuous with the epididymis, but may appear contiguous when imaged by ultrasonography. These structures tend to be benign and do not alter epididymal function.
Acquired Acquired conditions involving the epididymides are more common than congenital problems, but still rare. Diagnosis of acquired diseases is best made using physical examination and genital palpation, ejaculate analysis including a stained smear to assess the presence of white blood cells, as well as ultrasonography of the external and internal genitalia. A complete blood count and serum chemistry should be used to rule out other systemic diseases. A fine needle aspiration or biopsy may be beneficial when an epididymal mass is present, but care should be taken to avoid complications associated with this procedure. Epididymitis Epididymitis is generally of bacterial etiology and rarely occurs as a primary condition. Commonly associated with orchitis, epididymitis may be ascend-
Figure 111.2 Cyst-like structures associated with the ductus deferens just distal to the tail of the epididymis (arrow). This structure did not alter sperm flow through the ductus deferens.
Figure 111.3 Enlarged epididymal tail of the left testis, which in this stallion was associated with epididymitis (arrow).
ing, originating from the bladder or accessory sex glands, or hematogenous. Clinical presentation includes recurrent signs of abdominal or inguinal pain, lethargy, and subfertility. Physical examination reveals an enlarged, painful epididymis. The cauda epididymis appears to be affected more often than the corpus or caput; however, the entire epididymis is often involved in chronic cases. Ultrasonographic findings include hypoechoic areas consistent with dilated ductile segments and the margins of the epididymis are often irregular. Areas of hyperechogenicity may be scattered throughout the epididymal tissue in cases of chronic epididymitis (Figures 111.3–111.5).10 Large numbers of inflammatory cells are present in the ejaculate, and bacterial cultures are generally positive. Streptococcus zooepidemicus is the organism most commonly isolated;11 however, one reported case of epididymitis caused by Proteus mirablis has been described.12 Treatment may be attempted in acute cases with antibiotic therapy. When choosing an antibiotic, the pH of the target tissue, liposolubility of the agent, and susceptibility of the organism should be taken into account. An antibiotic that has a high pKa and lipid solubility is ideal for penetration of the
Figure 111.4 Enlarged body (white arrow) and epididymal tail (black arrow) (testis on the right).
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Epididymal Abnormalities
Figure 111.5 White blood cells in an ejaculate from a stallion with epididymitis. Notice sperm that have been engulfed by the white blood cells (arrows).
epididymis. Trimethoprim–sulfamethoxazole (TMS) and chloramphenicol are common choices available in oral formulations for use in horses. Duration of therapy to achieve complete resolution is unknown, but a course of 3–4 weeks has been reported.11, 12 The prognosis for future fertility in cases of epididymitis is poor, and, even in unilaterally affected stallions that underwent hemicastration, a positive culture was still obtained several months following removal of the affected epididymis.11 If future fertility is to be preserved, early diagnosis and aggressive treatment is required. Sperm granuloma Although far more common in other domestic species, sperm granulomas can occur in stallions (Figure 111.6). Sperm granulomas have been associated with the parasitic migration of Strongylus edentatus,13, 14 but, with the introduction of improved anthelmentics, the incidence has decreased. Clinical presentation is similar to that of chronic epididymi-
Figure 111.6 Histology of a blind-ended pouch off the epididymal lumen. These outpouches have the potential to elicit an inflammatory reaction and subsequent granuloma formation.
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tis and induces firm enlargement of the epididymis, most commonly the tail region, and oligospermia. Bacterial cultures of the epididymis and ejaculated semen are negative, ruling out bacterial epididymitis. Histopathology of the enlarged epididymis consistently shows a granulomatous reaction to sperm cells located outside the tubules with an influx of macrophages and lymphocytes due to the disruption of the blood–epididymis barrier. In chronic cases, peritubular fibrosis is also present and can be detected as hyperechoic areas on ultrasonography. If the condition is unilateral, hemicastration is recommended to salvage the stallion’s breeding ability. If the condition is bilateral, prognosis for future fertility is poor.13, 14 Other conditions Neoplasia of the epididymis is rare, but has been reported. Held et al.15 described a case of lymphosarcoma in the epididymis of a stallion that presented for testicular swelling and infertility. Ultrasonography revealed several hyperechoic areas along with hypoechoic masses in the caput of both epididymides. Diagnosis was made following castration and led to the subsequent euthanasia of the stallion. The lymphosarcoma was generalized and affected several major organs, including the lymph nodes, spleen, liver, lung, bladder, and synovium.15 Stallions termed ‘accumulators’ store excessive sperm in the ampulla(e), also called ‘plugged ampullae’. This results in a variety of clinical manifestation (see Chapter 107). One clinical sign is distension of the one or both epididymal tails. Since the epididymal tails are upstream from the actual blockage, sperm also back up in the tail, which can be visualized easily by ultrasonography (Figures 111.7 and 111.8).
Figure 111.7 Ultrasonographic image of the cauda epididymis distended with sperm as a result of segmental aplasia of the ductus deferens. This appearance can also occur in stallions that accumulate (plugged ampullae) sperm.
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2.
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5. Figure 111.8 Leiomyoma ventral to the body of the epididymis of the left testis (arrow).
6.
Clinical use of epididymal sperm
7.
Recently, harvesting sperm directly from the cauda epididymis has been reported. This technique is valuable in preserving spermatozoa from stallions that die unexpectedly (see Chapter 317). Sperm can be collected by either flushing the epididymal duct or mincing the tissue to release the spermatozoa. Although pregnancies have been reported following the use of frozen–thawed epididymal spermatozoa, the in vivo fertility of epididymal sperm is lower than that of ejaculated sperm.16 Morris et al.16 achieved a 45% pregnancy rate when using 200 × 106 fresh epididymal sperm inseminated hysteroscopically. The pregnancy rate was reduced to 18% when the epididymal spermatozoa were used following cryopreservation. Fertility rates may be improved with the use of intracytoplasmic sperm injection (ICSI). If the testicles are at a remote site with no cryopreservation capabilities, it is currently recommended to cool the testicles for shipment. Neild et al.17 showed higher progressive motility as long as 24 hours after the death of the stallion when the testicles were cooled when compared with those transported at room temperature. This is an emerging area and much research is still to be performed.
8. 9.
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13. 14.
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References 1. Johnson L, Amann RP, Pickett BW. 1978. Scanning electron microscopy of the epithelium and spermatozoa in the
equine excurrent duct system. Am J Vet Res. 39(9):1428– 1434. Arrighi S, Romanello MG, Domeneghini C. Ultrastructure of the epithelium that lines the ductuli efferentes in domestic equidae, with particular reference to spermatophagy. Acta Anat 1994;149:174–84. Lopez Alvarez ML, Bustos Obregon E. Spermatophagy in the stallion epididymis: a scanning and transmission electron microscopy study. Acta Anat 1995;153:181–8. Lopez ML, Fuentez P, Retamal C, De Souza W. Regional differentiation of the blood-epididymis barrier in stallion (Equus Caballus). J Submicrosc Cytol Pathol 1997;29: 353–63. Gebauer MR, Pickett BW, Swierstra EE. Reproductive physiology of the stallion. III Extra-gonadal transit time and sperm reserves. J Anim Sci 1974;39:737–42. Love CC. Ultrasonographic evaluation of the testis, epididymis, and spermatic cord of the stallion. Vet Clin North Am Equine Pract 1992;8:167–81. Blanchard TL, Woods JA, Brinsko SP. Theriogenology question of the month. J Am Vet Med Assoc 2000;217:825–6. Roberts SJ. Veterinary Obstetrics and Genital Diseases. North Pomfret, VT: David and Charles, 1986; pp. 843–4. Varner DD, Schumacher J, Blanchard TL, Johnson L. Diseases of the epididymis. In Pratt P (ed.) Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991; pp. 233–40. Traub-Dargatz JL, Trotter GW, Kaser-Hotz B, Bennett DG, Kiper ML, Veeramachaneni NR, Squires E. Ultrasonographic detection of chronic epididymitis in a stallion. J Am Vet Med Assoc 1991;198:1417–20. Held JP, Adair S, McGavin MD, Adams WH, Toal R, Henton J. Bacterial epididymitis in two stallions. J Am Vet Med Assoc 1990;197:602–4. Brinsko SP, Varner DD, Blanchard TL, Relford RL, Johnson L. Bilateral infectious epididymitis in a stallion. Equine Vet J 1992;24:325–8. Blue MG, McEntee K. Epididymal sperm granuloma in a stallion. Equine Vet J 1985;17:248–51. Held JP, Prater P, Toal RL, Blackford JT, McCracken M. Sperm granuloma in a stallion. J Am Vet Med Assoc 1989;194:267–8. Held JP, McCracken MD, Toal R, Latimer F. Epididymal swelling attributable to generalized lymphosarcoma in a stallion. J Am Vet Med Assoc 1992;201:1913–15. Morris LH-A, Tiplady CA, Allen WR. Hysteroscopic insemination of mares with frozen-thawed epididymal spermatozoa. In: Havemeyer Foundation Monograph Series No. 6, 2001; pp. 38–40. Neild D, Miragaya M, Chaves G, Pinto M, Alonso A, Gambarotta M, Losinno L, Aguero A. Cryopreservation of cauda epididymis spermatozoa from slaughterhouse testicles 24h after ground transportation. Anim Reprod Sci 2006;94: 92–5.
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Chapter 112
Abnormalities of the Testicles Warren Beard
Introduction Sperm production requires normal function of the homeostatic mechanisms of the testes. Among the factors that influence spermatogenesis is testicular temperature. Normal scrotal temperature is less than 35◦ C and increases in scrotal temperature to 37◦ C for as little as 24 hours have been demonstrated to decrease sperm motility and sperm production for up to 60 days.1 Testicular injury, whether traumatic or infectious, will alter the thermoregulatory mechanisms of the testes and result in a temporary decrease in sperm production. If the injury is severe enough, or if appropriate medical treatment is not initiated in a timely manner, permanent testicular degeneration may result. Therefore, it is important to understand the pathophysiology of the various testicular conditions so that appropriate medical and/or surgical interventions may be made.
Testicular trauma Traumatic orchitis may be the most commonly encountered testicular abnormality in clinical practice. These injuries are largely preventable because the principal cause is inadequate restraint of the mare during breeding. The diagnosis is straightforward and evidenced by scrotal edema, dermal bruising, heat, and pain to palpation. Often such trauma may also result in injury to other parts of the external genitalia (Figure 112.1). Ultrasound examination is useful because scrotal edema is often severe enough to preclude adequate examination of the testes by manual palpation. Ultrasound examination allows objective assessment of the amount of scrotal edema; the testicular architecture, including rupture of the tunica albuginea; the presence of gas, which may indicate skin disruption; and the presence of either a hydrocele or hematocele.2 Serial ultrasound examination also allows objective assessment of the response to treatment.
Testicular trauma causes the production of cytokines that initiate the inflammatory process. Prostaglandins increase blood flow and vascular permeability and, along with cytokines, alter the hypothalamic thermoregulatory control to increase body temperature. The result is extravasation of fluid and protein from the vascular space into the interstitium, causing edema. Edema insulates the affected testicle and further interferes with thermoregulation. Local blood flow is increased and overwhelms the ability of the countercurrent mechanism of the spermatic cord to cool incoming blood. Fibrin extravasates from the vascular space into the vaginal cavity with inflammation and may persist and undergo conversion to fibrous connective tissue and form fibrous adhesions between the tunics that impair mobility and impair thermoregulation. The blood–testis barrier may be disrupted, exposing previously immunologically privileged sperm and sperm precursors to the immune system. The resulting autoimmune response can cause reduced fertility.3, 4 The therapeutic goal is to rapidly attenuate the inflammatory response because transient increases in testicular temperature of only a few degrees will have a negative impact on sperm production rate and semen quality. Phenylbutazone [4.4 mg/kg per os (p.o.) or intravenously (i.v.)] or flunixin meglumine (1.1 mg/kg i.v.) will inhibit prostaglandin synthesis and decrease blood flow, edema, and temperature. Strong consideration should be given to the use of dexamethasone (40 mg i.v. or intramuscularly (i.m.)] in cases in which sepsis is not likely. The antiinflammatory effects of steroids and non-steroidal medications are additive because of inhibition of the inflammatory cascade at separate points in the pathway. Scrotal edema may be reduced by the topical application of hyperosmotic ointments that osmotically draw edema fluid out of the interstitium. Systemic antimicrobial agents are indicated for horses
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Stallion: Abnormalities of the Stallion’s Reproductive Tract cal agent can be identified. Diagnosis of viral orchitis may be more difficult and would require virus isolation or polymerase chain reaction from nasal swabs or acute and convalescent serum titers. In cases of unilateral orchitis, unilateral castration may be warranted to save the contralateral testicle from permanent injury.
Hydrocele
Figure 112.1 Ruptured tunica albuginea secondary to trauma.
in which the skin is not intact and as prophylaxis against secondary bacterial infection for horses with severe edema and hemorrhage. Topical cold therapy is beneficial by providing analgesia, limiting edema, and decreasing blood flow. It is doubtful whether twice daily application of cold via cold water hydrotherapy is sufficient to decrease edema and appreciably lower testicular temperature. Consideration should be given to the continuous application of ice or cold packs held in place via a sling. Cold therapy is indicated as long as active inflammation is present. A single episode of trauma will result in increased blood flow and altered vascular permeability for 24–48 hours. A longer duration can be expected with more severe trauma or the presence of infection. Edema will persist after the active inflammation has resolved, and is resolved by lymphatic uptake of excess interstitial fluid. Once the active inflammation has resolved, exercise and hydrotherapy are beneficial in improving lymphatic drainage.
Infectious orchitis Infectious orchitis may result from trauma, hematogenous spread, extension from adjacent tissues, or peritonitis.5, 6 Infectious orchitis causes pain, edema, and leukospermia. The pathophysiology of infectious orchitis mirrors the pathway described for testicular trauma and early aggressive treatment is essential to prevent testicular degeneration. Non-specific therapy should include those treatments described for traumatic orchitis as well as specific treatment to target the infectious agent, if known. Methods of identifying the infectious agent may include bacterial culture of adjacent wounds from which testicular extension is expected; abdominocentesis; centesis of fluid within the vaginal cavity; or collection of an ejaculate. In cases of suspected bacterial orchitis, broad-spectrum antimicrobial therapy is indicated until a specific etiologi-
The vaginal cavity is the space between the parietal and visceral vaginal tunics and is continuous with the peritoneal cavity. It normally contains a small amount of fluid which is identical to peritoneal fluid, except with pathological conditions of the scrotal contents. Hydrocele is the term given to describe an abnormal accumulation of fluid in the vaginal cavity.7 The presence of excessive fluid is not pathological per se; rather, it is indicative of other underlying pathology. Abnormal accumulation of fluid may be detected by palpation of the scrotum if there is no scrotal edema. Ultrasound examination will allow measurement of the depth of the fluid surrounding the testicle, assessment of scrotal edema, and the ability to examine the testes for pathology.2 Some determination of the character of the fluid may be made via ultrasound. A hyperechoic fluid is likely the result of a high cell content and fibrin may be readily visible, possibly indicating sepsis. Hydroceles may be idiopathic or secondary to trauma, neoplasia, or any condition that decreases venous or lymphatic outflow. A hydrocele may occur as an incidental finding in geldings in which an open castration technique results in inadequate excision of the parietal tunic such that it seals over to accumulate fluid as healing occurs.8 Hydroceles resulting from trauma should be treated as outlined in the previous section. Treatment should be directed toward the inciting cause, although the cause can be difficult to identify. Unilateral castration may be successful in resolving the condition.
Hematocele Hematocele is defined as the accumulation of blood within the vaginal cavity.9 Trauma is the most common etiology. Ultrasound findings that are consistent with a hematocele include a homogeneous hyperechoic swirling fluid indicative of a highly cellular fluid (blood) and the presence of fibrin (Figure 112.2). Non-specific therapy for testicular trauma may be attempted. Centesis and drainage of the blood accompanied by lavage of the vaginal cavity can be performed in an attempt to prevent adhesions.10 Unresponsive cases or stallions which have testicular rupture are candidates for unilateral castration.
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Figure 112.2 Scrotal ultrasound of a hematocele. Courtesy of Dr. Laurie Beard.
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Figure 112.4 Cut surface of testicular seminoma from the stallion in Figure 112.3.
Monorchidism Neoplasia Testicular neoplasia is uncommon in horses, although the incidence is unknown because most males are castrated early in life. Neoplasia should be suspected when unilateral testicular enlargement occurs in the absence of trauma (Figure 112.3). The testicle is usually non-painful and firm to palpation and the contours of the normal testicle are lost as it enlarges. Ultrasound examination usually reveals the presence of abnormal testicular architecture.11 Histopathological examination of a biopsy of the affected area is usually diagnostic. Primary testicular neoplasia is much more common than metastatic neoplasia from other sites. Sertoli cell tumors, seminomas, Leydig cell tumors, and teratomas constitute the majority of cases of primary testicular neoplasia.12, 13 Tumors are usually locally invasive and infrequently metastasize. Unilateral castration is the treatment of choice if continued fertility is desired (Figure 112.4).
Figure 112.3 Scrotum of stallion with testicular seminoma.
Monorchidism is very uncommon in horses.14–17 The etiology is unknown; however, a vascular accident during embryological development that results in unilateral testicular agenesis is a popular theory that has some credence. The primary difficulty lies in distinguishing monorchidism from cryptorchidism because monorchidism is a rare condition that is never considered as a differential diagnosis until at least one abdominal exploration has failed to locate a cryptorchid testicle. Monorchidism can be confirmed after removal of the descended testicle by the failure of serum testosterone concentrations to increase following administration of human chorionic gonadotropin (hCG).
Torsion of the spermatic cord The testicle normally resides in the scrotum with the tail of the epididymis located caudally. The testicle may rotate up to 180◦ without clinical signs.18 Asymptomatic 180◦ torsion would be evident by an absence of the tail of the epididymis visible or palpable in the caudal scrotum, and no clinical signs of colic. Torsions of the spermatic cord greater than 180◦ will result in strangulation of the blood vessels in the spermatic cord. The thin-walled, low-pressure veins in the spermatic cord are more susceptible to occlusion than the thicker walled, high-pressure arteries. Venous outflow is decreased while some arterial inflow persists. This results in an increase in pressure, development of testicular and scrotal edema, and pain. Affected animals are presented for signs of colic.19, 20 The principal differential diagnoses would be a strangulating inguinal hernia and unilateral testicular trauma. Careful palpation should confirm the abnormal position of the testicle within the scrotum and ultrasonography plus rectal palpation can rule
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out a scrotal hernia by the absence of small intestine in the deep inguinal ring. Unilateral castration is indicated if the testicle is non-viable or if the damage is severe enough to result in testicular degeneration.
Scrotal hernia The deep inguinal ring is a slit-like opening of the parietal peritoneum into the vaginal tunic that allows the testicle to descend into the scrotum. The deep inguinal ring is normally small enough to allow passage only of the spermatic cord and ductus deferens. Scrotal herniation occurs when the deep inguinal ring is enlarged and allows small intestine to descend into the vaginal tunic. Congenital inguinal hernias are common in Standardbreds and draft breeds although they may occur in any breed.21 Affected foals may have unilateral or bilateral involvement and have no signs other than an enlarged scrotum (Figure 112.5). The condition is frequently self-correcting by several months of age and non-surgical management is the treatment of choice unless the foal develops signs of colic, the hernia contents are non-reducible, or the hernia has not corrected by several months of age. Conservative management consists of regular monitoring and frequent manual reduction of intestinal contents into the abdomen. Surgical correction is indicated for hernias that do not spontaneously resolve, if the foal develops a rent in the vaginal tunic, or if the contents become strangulated.22, 23 Closed castration is successful in resolving the hernias in the absence of strangulation. Immediate laparotomy is indicated for foals with a rent in the vaginal tunic and resection of the small intestine may be required (Figure 112.6). Inguinal hernias may be repaired without castration; however, a discussion with the owner about the possible heritability of this condition is warranted. Acquired inguinal hernias can occur in intact males. Common histories that accompany the onset of an acquired inguinal hernia include onset of colic
Figure 112.5 Weanling draft colt with a non-strangulating congenital inguinal hernia.
Figure 112.6 A 1-week-old Standardbred foal with a congenital inguinal hernia with a ruptured vaginal tunic. Notice the bruising of the inguinal skin and the large hernia that extends beyond the normal scrotal region.
after dismounting from a mare and onset of colic on the racetrack while training. Acquired inguinal hernias cause immediate signs of colic and should always be considered as a differential in any intact male.24, 25 The diagnosis is readily confirmed by the presence of small intestine entering the deep inguinal ring during rectal palpation. Immediate treatment is necessary to preserve the testicle, if desired, and to preclude the need for a small intestinal resection and anastomosis. With acute herniation, the hernia may be corrected by external manipulation of the scrotum under general anesthesia to squeeze the incarcerated bowel back into the abdomen.25 Concurrent per rectum manipulation of the affected portion of the intestine often helps in correcting the problem. Surgical correction is indicated if manual correction is unsuccessful or if the hernia contents are devitalized.7
Testicular degeneration Testicular degeneration is an acquired unilateral or bilateral atrophy of the seminiferous tubules, resulting in decreases in sperm production, spermatozoal quality, and testicular size. Testicular atrophy secondary to degeneration is distinguishable from testicular hypoplasia only by a known history of prior normal testicular size and function. The diagnosis of testicular degeneration is made by semen evaluation that may reveal oligospermia, asthenospermia, teratospermia, and azoospermia. Single semen evaluations may confirm decreased sperm production; however, serial sampling is necessary to confirm spermatogenic efficiency. Transient elevations of testicular temperature are capable of decreasing spermatogenesis for up to 60 days; therefore, serial sampling is required to determine whether the observed decrease in sperm output from a single determination is reversible or the result of permanent degeneration.
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Abnormalities of the Testicles Testicular biopsy and histopathological examination can definitively confirm testicular degeneration; however, there are risks associated with testicular biopsy.26, 27 The principal risks of testicular biopsy include hemorrhage, increased testicular pressure, adhesions between the parietal and visceral vaginal tunic, and disruption of the blood–testis barrier, resulting in an immune-mediated reaction. The preceding complications have the potential to induce testicular degeneration if it were not already present. Testicular degeneration is a secondary condition that usually is the result of one of the testicular conditions discussed previously. The key to preventing testicular degeneration in those instances in which it is preventable is early recognition of a testicular problem and timely aggressive treatment.
References 1. Freidman R, Scott M, Heath SE, Hughes JP, Daels PF, Tran TQ. The effects of increase testicular temperature on spermatogenesis in the stallion. J Reprod Fertil Suppl 1991;44:127–34. 2. Love CC. Ultrasonographic evaluation of the testis, epididymis, and spermatic cord of the stallion. Vet Clin North Am Equine Pract 1992;8:167–82. 3. Papa FO, Alvarenga MA, Lopes MD, Filho EP. Infertility of autoimmune origin in a stallion. Equine Vet J 1990;22:145–6. 4. Zhang J, Ricketts SW, Tanner SJ. Antisperm antibodies in the semen of a stallion following testicular trauma. Equine Vet J 1990;22:138–41. 5. Kasaback CM, Rashmir-Raven AM, Black SS. Theriogenology question of the month. Septic orchitis-periorchitis and epididymitis. J Am Vet Med Assoc 1999;215:787–9. 6. Estepa JC, Mayer-Valor R, Lopez I, Santisteban JM, Ruiz I, Aguilera-Tejero E. What is your diagnosis? Abscess developed as a result of scrotal and testicular lesions. J Am Vet Med Assoc 2006;228:515–16. 7. Schumacher J. Surgical disorders of the testis and associated structures. In: Auer JA, Stick (JA) Equine Surgery, 3rd edn. St. Louis, MO: Saunders, 2006; pp. 775–810. 8. Colbourne CM, Adkins AR, Yovich JV. Hydrocele formation after castration in 3 geldings. Aust Vet J 1996;73:156–7. 9. Gygax AP, Donawick WJ, Gledhill BL. Hematocoele in a stallion and recovery of fertility following unilateral castration. Equine Vet J 1973;5:128.
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10. Blanchard TL, Varner DD, Brinsko SP. Theriogenology question of the month: Scrotal Hematocele. J Am Vet Med Assoc 1996;209:2013–14. 11. Beck C, Charles JA, Maclean AA. Ultrasound appearance of an equine testicular seminoma. Vet Radiol Ultrasound 2001;42:355–7. 12. Caron JP, Barber SM, Bailey JV. Equine testicular neoplasia. Comp Cont Educ Pract Vet 1985;7:S53–9. 13. Brinsko SP Neoplasia of the male reproductive tract. Vet Clin North Am Equine Pract 1988;14:517–33. 14. Santschi EM, Juzwiak JS, Slone DE. Monorchidism in three colts. J Am Vet Med Assoc 1989;194:265–6. 15. Parks AH, Scott EA, Cox JE, Stick JA. Monorchidism in the horse. Equine Vet J 1989;21:215–17. 16. Strong M, Dart AJ, Malikides N, Hodgson DR. Monorchidism in two horses. Aust Vet J 1997;75:333–5. 17. Petrizzi L, Varasano V, Robbe D, Valbonetti L. Monorchidism in an appaloosa stallion. Vet Rec 2004;155: 424–5. 18. Pascoe JR, Ellenburg TV, Culbertson MR Jr, Meagher DM. Torsion of the spermatic cord in a horse. J Am Vet Med Assoc 1981;178:242–5. 19. Horney FD, Barker CA. Torsion of the testicle in a standardbred. Can Vet J 1975;16:272–3. 20. Threlfall WR, Carleton CL, Robertson J, Rosol T, Gabel A. Recurrent torsion of the spermatic cord and scrotal testis in a stallion. J Am Vet Med Assoc 1990;196:1641–3. 21. Shoemaker R, Bailey J, Janzen E, Wilson DG Routine castration in 568 draught colts: incidence of evisceration and omental herniation. Equine Vet J 2004;36:336–40. 22. Spurlock GH, Robertson JT. Congenital inguinal hernias associated with a rent in the common vaginal tunic in five foals. J Am Vet Med Assoc 1988;193:1087–8. 23. van der Velden MA. Ruptured inguinal hernia in new-born colt foals: a review of 14 cases. Equine Vet J 1988;20:178– 81. 24. Schneider RK, Milne DW, Kohn CW. Acquired inguinal hernia in the horse: a review of 27 cases. J Am Vet Med Assoc 1982;180:317–20. 25. van der Velden MA. Surgical Treatment of acquired inguinal hernia in the horse: A review of 51 cases. Equine Vet J 1988;20:173–8. 26. DelVento VR, Amann RP, Trotter GW, Veeramachaneni DN, Squires EL. Ultrasonographic and quantitative histologic assessment of sequelae to testicular biopsy in stallions. Am J Vet Res 1992;53:2094–101. 27. Threlfall WR, Lopate C. Testicular biopsy. In: McKinnon AOM, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 943–9.
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Chapter 113
Neoplasia of the Reproductive Tract John F. Edwards
Most male horses are castrated; thus few stallions, and even fewer old stallions, are available for study of neoplasia. Most neoplasms of the stallion are seen with increasing age, but some are characteristically seen in the young, or in cryptorchid horses. Regardless, many published cases of neoplasia in the stallion reproductive tract are single-case reports and are purely descriptive in nature. It is unfortunate that most cases do not include pathophysiological as well as morphological observations. Most neoplasms to be discussed involve the testis, penis or prepuce. The incidence of neoplasia in the stallion genital tract is poorly documented. Data for complete examinations of a large number of stallions in the population at large are not available. Kimura (quoted in McEntee1 ) reported 114 neoplasms in 77 224 horses slaughtered in Japan. This included 49 testicular neoplasms. The overall neoplasia incidence for horses is probably low because certain common neoplasms, such as pituitary pars intermedia adenomas and thyroid adenomas, of aged horses are not investigated at routine slaughter. The age and number of stallions slaughtered also were not included. An abattoir study in England reported one teratoma in 96 stallions.2 Neither of these studies mentioned neoplasms of the external genitalia. These data suggest that the incidence of stallion reproductive neoplasia is low. In a 10-year survey of 2250 equine neoplasms at one veterinary institution, a testicular teratoma and 11 preputial or penile squamous cell carcinomas were reported.3 Twenty-one equine neoplasms were reported from 687 necropsies performed over 5 years at another veterinary college, and while no testicular neoplasms were identified, 15 squamous cell carcinomas and 2 melanomas were diagnosed on male external genitalia.4 A study of 38 207 horses seen at 12 North American veterinary institutions reported
only 951 horses with neoplasms.5 Unfortunately, the incidence of genital neoplasia in stallions in this study is embedded in, and not separable from, the general data that cite 543 male horses and 419 mares having 97 genital neoplasms. In all surveys, no mention of neoplasia of accessory genital organs is made, thus indicating that there is a low incidence of bulbourethral, prostatic, and seminal vesicular neoplasia. Teratomas are the most common reproductive neoplasm of colts and stallions less than 2 years old.1 Teratomas can have germ tissue from embryological ectoderm, mesoderm, and endoderm. Most frequently, only two germ layers, ectoderm and mesoderm, are involved. Brain, glands, muscle, fat, bone, haired skin, and connective tissue are commonly found in these neoplasms.1,6–8 Some contain dermoid cysts, i.e., cysts lined by stratified, keratinizing, squamous epithelium with hair follicles and dermal adnexa.7, 8 It is important to sample from many areas in teratomas for adequate characterization. Teratomas occur more frequently in cryptorchid testes and often contain cysts ranging from several mm to 30 cm in diameter. Abdominal testicular teratomas have a broad size range, from small nodules that are difficult to identify to large and cystic or polycystic structures. Malignant teratomas are called teratocarcinomas, and one has been reported in a 1.5-year-old, bilateral cryptorchid.9 The author has seen two similar cases in yearlings. These animals had abdominal distention because the neoplasms metastasized widely to lymph nodes, liver and lung and implanted on the peritoneum. Seminomas are the most commonly reported testicular neoplasm of the mature stallion.6, 7,10–17 These neoplasms are derived from germ cells that migrate to the gonadal ridge from the yolk sac of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 113.1 The testicular parenchyma is almost totally replaced by a large, homogenous seminoma in this descended testis. An arrow indicates a remaining rim of dark, degenerate testis. The bar is 1 cm.
the embryo. Seminomas are soft and gray or light brown and can be large (see Figure 113.1). Although seen in descended testes, seminomas are reported most frequently in cryptorchid testes. Large scrotal or cryptorchid seminomas commonly are malignant and metastasize to the epididymides, spermatic cords, contralateral testis, inguinal, lumbar, iliac, aortic, tracheobronchial and mediastinal lymph nodes, liver, and lung, and they implant widely over peritoneal surfaces. The size of malignant seminomas and the high incidence of malignancy in cryptorchid testes may explain the high frequency of their reports in the literature; however, many seminomas are benign. Neoplastic seminoma cells have ample, gray cytoplasm with an indistinct cell border, and a hypochromatic, large, round nucleus with several large nucleoli and lacy chromatin. Anisokaryosis with karyomegaly is marked, and scattered, multinucleated cells are seen. Mitoses are common, and seminomas commonly have areas of lymphohistiocytic inflammatory infiltration. In fresh biopsies, variably sized, eosinophilic granules are noted in the cytoplasm of cells. Seminoma in situ and intratubular seminoma are terms used for early stages of the neoplasm when neoplastic cells are found only within tubules. Useful markers for the neoplastic cells in some species have been calretinin, vimentin, and OCT4 (POU5F1).18, 19 In human beings, placental alkaline phosphatase and neuron specific enolase (NSE), but not alpha-fetoprotein, can be stained in seminoma cells.19 Interstitial (Leydig) cell neoplasms (ICN) are seen less frequently and have been reported in horses from
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3 to 20 years old.20–24 The neoplasms are derived from the sex cord cells (sex cord stroma) that migrate into the gonad from the surface of the gonadal ridge in the embryo. In 9 of 10 published cases of ICN in cryptorchid testes, 6 of 10 animals had bilateral ICN, and two cases of bilateral neoplasms were in bilateral cryptorchid animals.15, 21 Further, one of the cryptorchid cases had segmental aplasia of the epididymis.22 It is interesting to note that surgically induced cryptorchidism induces ICN in mice.25 In three reported cases, ICN were associated with aggressive behavior, but no data on circulating hormones in these horses were available. In another, the ICN was associated with testicular torsion in a descended testis.22 While ICN are less commonly reported, it has been suggested that ICN are underdiagnosed because they may be small (Figure 113.2) and are not reported to metastasize.1 Interstitial cell neoplasms are described as pale gray, especially in cryptorchid testes, but also can be pale yellow, similar to the appearance of ICN in dogs and bulls. Neoplastic cells are polygonal to low cuboidal with eosinophilic and vacuolated or foamy cytoplasm and have an oval or indented, hypochromatic nucleus. Mitoses and multinucleated cells are rare. In cases of cryptorchid ICN, the neoplastic cells can be spindyloid, and these neoplasms have perivascular accumulations of lymphocytes and macrophages. Useful markers for ICN in some species include calretinin, inhibin, vimentin, and melan-A.18, 19 Sustentacular (Sertoli) cell neoplasms (SCN) are derived from sex cords and are uncommon.26–29
Figure 113.2 Two small, pale, coalescing nodules of interstitial cell neoplasia in a section of degenerate, dark hemitestis. The upper border is the tunica albuginea. The marker is 1 cm.
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Only four cases have been reported: two were in cryptorchid testes, and two were neoplasms in descended testes. One case in a cryptorchid testis and one in a descended testis were malignant and metastasized to the lumbar and hepatic lymph nodes, lungs, liver, spleen, and kidney.26, 27 Sertoli cell neoplasms are firm and pale gray, and usually the contralateral descended testis is atrophic. Animals in reports were presented for weight loss, loss of libido, and dyspnea (presumably due to pulmonary metastases). Histologically, they have cuboidal to columnar, indistinctly bordered, palisading cells with eosinophilic, lacy cytoplasm and an oval, hyperchromatic nucleus with a prominent nucleolus. Fibrous bands separate neoplastic lobules, and the mitotic index is regionally high. While staining for antimullerian hormone is most widely used to identify SCN, vimentin, cytokeratin, inhibin, NSE and GATA4 (a differentiation marker for sustentacular cells) also will stain the neoplastic cells.19 A neoplasm with characteristics of both ICN and SCN, a mixed sex cord-stromal tumor, has been reported in a descended testis of a 30-year-old stallion.30 Two mixed germ cell-sex cord stromal testicular neoplasms, neoplasms with characteristics of germ cell neoplasia and SCN or ICN, have been reported in stallions.31, 32 Both cases were in mature stallions (a 10-year-old and a 25-year-old) and one case had metastasized widely. Because of the pleomorphic histological pattern of these mixed neoplasms, it is important to sample these neoplasms from several areas to establish this diagnosis. A malignant testicular embryonal carcinoma originating from a scrotal testis was reported in a 7year-old stallion.33 These neoplastic cells have indistinctly bordered, basophilic cytoplasm and large, oval, hypochromatic nuclei with one to several nucleoli. Mitoses are common. Although bands of mature connective tissue separate lobules of this neoplasm, the primitive germ cells have not differentiated and form poorly differentiated nests, acini, and tubular structures in a loose, primitive mesenchyme. Positive staining for alpha-fetoprotein and demonstration of desmosomes between neoplastic cells are important features in establishing the diagnosis. Several other neoplasms have been reported in the stallion testis. The International Registry of Reproductive Pathology contains two cases of testicular lipomas and rete adenomas and a single case of a mast cell neoplasm.1 Bilateral testicular leiomyosarcomas were reported in a bilateral cryptorchid, 9month-old colt.34 This spindle cell neoplasm not only affected both testes and epididymides but also involved the popliteal, splenic and cervical lymph nodes, the masseter muscles, diaphragm, and caudal vena cava. The authors in that report acknowledged
the possibility that this was a congenital, multicentric neoplasm. Treatment of testicular neoplasia is usually early orchiectomy. In the case of cryptorchid neoplasia where there may be a genetic predisposition, it may be counseled to remove a contralateral, descended testis. Removal of a neoplastic, scrotal testis that is associated with degeneration or atrophy of the contralateral testicle may result in spontaneous recovery of function after removal of the neoplasm.20 Ultrasound-guided biopsies can be used to establish diagnosis in many instances, but sampling several points in a neoplastic testis after orchiectomy for diagnosis should be encouraged. Prognosis is more accurate if the spermatic cord is examined histologically for invasion of a neoplasm. Palpation and ultrasonography of lumbar lymph nodes and inguinal ring examination for metastasis are always indicated. Within a testis with neoplasia, the parenchyma adjacent to most neoplasms is degenerate. Although local treatment of testicular neoplasms has been undertaken in an effort to induce regression of the neoplasm and rescue surrounding parenchyma, it is difficult. One should remember that testicular neoplasia is sometimes bilateral, and examination of the testis contralateral to a neoplastic testis is critical. One should always take a preoperative serum sample and a follow-up serum sample to evaluate functionality of testicular neoplasms. Although this hormonal data has diagnostic significance in human medicine, it is uncommonly examined in equine cases.19, 35 Peritesticular neoplasia occurs. Mesotheliomas implanting in the peritoneal cavity can extend into the vaginal tunic, and we also have detected primary mesotheliomas of the scrotal tunics (Figure 113.3). A
Figure 113.3 Scrotal testis from a 15-year-old stallion with abdominal mesotheliomatosis. Numerous mesotheliomas are implanted over the testis and spermatic cord. Marker is 1 cm.
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Neoplasia of the Reproductive Tract leiomyoma of the testicular tunics in a stallion has been reported.36 These neoplasms are adherent bands of smooth muscle to the tunica albuginea and usually are an incidental finding noted after routine castrations. As stated above, neoplasms of the bulbourethral glands, prostate, seminal vesicle, ampullae, and epididymis are not reported; however, a case of generalized lymphosarcoma was reported to involve the epididymis.37 No doubt neoplasms of these organs have been seen and will be published as case reports. In our collection, we have one case of an ampullary carcinoma that obstructed the vasa deferentia and led to formation of secondary sperm granulomas in the epididymis. Neoplasia of the prepuce and/or penis occur in both stallions and geldings.3–5,38 In most published studies, the data related to neoplasia of the external genitalia combine the cases of geldings and stallions. Penile and preputial neoplasms reported most frequently are sarcoids, squamous cell papillomas, carcinomas and melanomas, and there are geographical differences in their incidence and sites of occurrence.24, 39 We have twice seen solitary lymphosarcoma limited to the prepuce. It should be remembered that nodular and ulcerated masses of the penis and prepuce may be non-neoplastic conditions such as mycoses and habronemiasis. Sarcoids are cutaneous fibrosarcomas that can affect the external male genitalia, but uncommonly cause problems in reproductive function. These neoplastic cells often contain bovine papillomavirus DNA sequences. The neoplasms are often multiple and can be verrucous, nodular, or may look like granulation tissue. They commonly occur at sites of trauma, and there is evidence that there is some genetic basis for susceptibility. They are more common in mules and donkeys, in which species there is some concern about increased post-castration, paragenital occurrence.40 A number of treatment modalities have been tried, and wide excision of small sarcoids is most successful when the neoplasm first occurs; however, large sarcoids and sarcoids that have reoccurred have a poorer prognosis. They usually do not metastasize except by local invasion (reviewed in Scott and Miller 2003).24 Squamous cell carcinoma (SCC) is the most common neoplasm of the stallion prepuce and penis.38, 41–43 It is a neoplasm of aged horses, and horse breeds with unpigmented skin are overly represented in the affected population. Solar keratosis or carcinoma in situ is observed in early lesions, and SCC often develops as a locally extensive, multicentric lesion. Although papillomavirus has been identified in equine papillomas, it has not been shown in SCC.44 The data related to papillomavirus and equine SCC is over 30 years old. In men, papillomavirus is associated with
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some forms of SCC and, using improved techniques, the agent is more frequently identified.45 Smegma has been shown experimentally to induce SCC. Geldings have a higher incidence of SCC of the penis and prepuce, probably because their population is greater than that of stallions, but perhaps also because geldings accumulate more smegma.38, 46 Our data agree with those of others. Of our last 25 paragenital SCC biopsy cases, 20 were geldings (3 preputial and 17 penile neoplasms; mean age 18.9 years, range 13 to 30 years) and 5 were stallions (5 penile neoplasms; mean age 17 years, range 10 to 24 years). Although squamous cell papilloma was not diagnosed, 4 carcinomas in situ were diagnosed. In the last 10 horses necropsied for SCC (not including those in the biopsy group above), all were geldings (17.8 years old, range 7 to 24 years), and all had lesions in the prepuce and penis. Six cases had metastasized to regional lymph nodes, and one had metastasized to the lung. Over the same period studied, one stallion and two geldings had preputial sarcoids, two geldings had severe preputial lymphosarcoma, and one gelding had a preputial SCC and a melanoma. Squamous cell carcinomas are difficult to treat, and although laser ablation, cryotherapy and chemotherapy of small/early lesions may be curative, larger and extensive lesions are managed with surgical therapy.41–43,47 Segmental posthectomy, phallectomy or en bloc resection of the penis, prepuce and local draining lymph nodes are the surgical options. SCC is locally aggressive and metastasizes slowly, especially to distant sites (lung, liver, and abdominal and thoracic lymph nodes). The best prognosis in men following SCC surgery is given when the neoplasm has not extended to the draining nodes,48 and this applies to SCC in the horse. Melanomas are multicentric in gray and white horses, and it is often unreasonable to remove all neoplasms, but if they interfere with reproductive function, an offending neoplasm can be removed. In the future, it is hoped that more data regarding neoplasia of the stallion reproductive tract will be available. When publishing archival data, as much information as possible should be included. Information derived from hormonal status of patients and the use of immunomarkers for testicular neoplasms will increase our understanding of neoplasia in stallions and may more firmly establish diagnoses. Longterm follow-up information is needed for evaluation of treatment modalities. Testing for papillomavirus using modern technology in SCC needs to be done in order to investigate its role in the pathogenesis and perhaps begin feasibility studies of developing a vaccine for SCC that might be used in young horses. We should strive to report new findings in collaborative groups whose pooled resources will assist thorough investigation.
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References 1. McEntee K. Scrotum, spermatic cord, and testis: proliferative lesions. In: Reproductive Pathology of Domestic Animals. San Diego, CA: Academic Press, 1990; pp. 279– 306. 2. Cotchin E, Baker-Smith J. Tumours in horses encountered in an abattoir survey. Vet Rec 1975;97(17):399. 3. Kerr KM, Alden CL. 1975. Equine neoplasia: a ten year survey. Proc 17th Ann Meet Am Assoc Vet Lab Diagn 1975; pp. 183–7. 4. Sundberg JP, Burnstein T, Page EH, Kikham WW, Robinson FR. Neoplasms of equidae. J Am Vet Med Assoc 1977;170(2):150–2. 5. Priester WA, Mantel N. Occurrence of tumors in domestic animals: data from 12 United States and Canadian colleges of veterinary medicine. J Natl Canc Instit 1980;47(6): 1333–4. 6. Aupperle H, Gerlac HK, Bartmann CP, Beerhenke S, Schoon HA. Histopathological findings in the cryptorchid testes of stallions. Pferdheikunde 1999;15(6):515–22. 7. Innes JRM. Neoplastic diseases of the testis in animals. J Pathol Bact 1942;54:485–98. 8. Stick JA. Teratoma and cyst formation of the equine cryptorchid testicle. J Am Vet Med Assoc 1980;176(3):211–14. 9. Shaw DP, Roth JE. Testicular teratocarcinoma in a horse. Vet Pathol 1986;23(3):327–8. 10. Becht JL, Thacker HL, Page EH. Malignant seminoma in a stallion. J Am Vet Med Assoc 1979;175(3):292–3. 11. Gibson GW. Malignant seminoma in a Welsh pony stallion. Comp Cont Educ Pract Vet 1984;6(5):S296–8. 12. Hunt RJ, Hay W, Collatos C, Welles E. Testicular seminoma associated with torsion of the spermatic cord in two cryptorchid stallions. J Am Vet Med Assoc 1990;197(11):484–6. 13. Hedjazi M, Naghshineh R. Un cas de seminome chez le cheval, metastase apres operation chirugicale [A case of seminoma in the horse, metastasis after surgery] Revu de Medecine Veterinaire 1981;132(2):143–5. 14. Knudsen O, Schantz R. Seminoma in the stallion: A clinical, cytological and pathologicoanatomical investigation. Cornell Vet 1963;53:395–403. 15. Smith BL, Morton LD, Watkins JP, Taylor TS, Storts RW. Malignant seminoma in a cryptorchid stallion. J Am Vet Med Assoc 1989;195(6):775–6. 16. Trigo FJ, Miller RA, Torbeck RL. Metastatic equine seminoma: report of two cases. Vet Pathol 1984;21(2): 259– 60. 17. Vaillancourt D, Fretz P, Orr JP. Seminoma in the horse:Report of two cases. J Eq Med Surg 1979;3(5):213– 18. 18. Owston MA, Ramos-Vara JA. Histologic and immunohistochemical characterization of a testicular mixed germ cell sex cord-stromal tumor and a Leydig cell tumor in a dog. Vet Pathol 2007;44(6):936–43. 19. Ro JY, Amin MB, Kim K-R, Ayala AG. Testis and paratesticular tissues. In: Fletcher CDM (ed.) Diagnostic Histopathology of Tumors vol. 1, 3rd edn. Churchill Livingstone Elsevier, 2007; pp. 812–60. 20. Bader H, Hoppen HO, Woekener A, Merkt H. Case studies in stallions with fertility problems: endocrine and spermatological aspects. Proc 11th Int Congr Anim Reprod Art Insem 1988; vol. 3, pp. 367–9. 21. Gelberg HB, McEntee K. Equine testicular interstitial cell tumors. Vet Pathol 1987;24(3):231–4.
22. Hay WP, Baskett A, Gregory CR. Testicular interstitial cell tumour and aplasia of the head of the epididymis in a cryptorchid stallion. Eq Vet Educ 1997;9(5):240–1. 23. May KA, Moll HD, Duncan RB, Pleasant RS, Purswell BJ. Unilateral Leydig cell tumour resulting in acute colic and scrotal swelling in a stallion with descended testes. Equine Vet J 1999;31(4):343–5. 24. Scott DW, Miller WH. Neoplastic and nonneoplastic tumors. In: Equine Dermatology. St Louis, MO: Elsevier Science, 2003; pp. 698–712. 25. Fowler KA, Gill K, Kirma N, Dillehay DL, Tekmal RR. Overexpression of aromatase leads to development of testicular Leydig cell tumors. Am J Pathol 2000;156(1):347– 53. 26. Duncan RB. Malignant Sertoli cell tumour in a horse. Equine Vet J 1998;30(4):355–7. 27. Pratt SM, Stacy BA, Whitcomb MB, Vidal JD, De Cock HEV, Wilson WD. Malignant Sertoli cell tumor in the retained abdominal testis of a unilaterally cryptorchid horse. J Am Vet Med Assoc 2003;222(4):486–90. 28. Rahaley RS, Gordon BJ, Leipold HW, Peter JS. Sertoli cell tumour in a horse. Equine Vet J 1983;15(1):68–70. 29. Stickle RL, Fessler JF. Retrospective study of 350 cases of equine cryptorchidism. J Am Vet Med Assoc 1978;172(3):343–6. 30. Zanghi A, Caton G, Marino G, De Vico G, Nicotina PA. Malignant mixed sex cord-stromal tumour in a stallion. Reprod Domest Anim 2004;39(5):376–9. 31. Cullen JM, Whiteside J, Umstead JA, Whitacre DM. A mixed germ cell-sex cord-stromal neoplasm of the testis in a stallion. Vet Pathol 1987;24(6):575–7. 32. Kollmann M, Friederike A, Hewicker-Trautwein M. Testikularer maligner keimzell-und gonadenstromatumor mit metastasiierung bei einem hengst. [Malignant germ cell and gonadostromal tumor with metastasis in a stallion.] Pferdeheilkunde 2006;22(2):177–80. 33. Valentine BA, Weinstock D. Metastatic testicular embryonal carcinoma in a horse. Vet Pathol 1986;23(1):92–6. 34. Allison N, Moeller RB Jr. Bilateral testicular leiomyosarcoma in a stallion. J Vet Diagn Invest 1999;11(2):179– 82. 35. Melo CM, Papa FO, Prestes NC, Alvaringa MA, LauferAmorim R. Bilateral Leydig cell tumor in stallion. J Eq Vet Sci 2007;27(10):450–3. 36. Johnson RC, Steinberg H. Leiomyoma of the tunica albuginea in a horse. J Comp Pathol 1989;100(4):465–8. 37. Held JP, McCracken MD, Toal R, Latimer F. Epididymal swelling attributable to generalized lymphosarcoma in a stallion. J Am Vet Med Assoc 1992;201(12):1913–15. 38. Brinkso SP. Neoplasia of the male reproductive tract. Vet Clin N. Am Eq Prac 1998;14(3):517–33. 39. Valentine BA. Survey of equine cutaneous neoplasia in the Pacific Northwest. J Vet Lab Invest 2006;18(1):123–6. 40. Reid SWJ, Mohammed HO. Longitudinal and crosssectional studies to evaluate the risk of sarcoid associated with castration. Can J Vet Res 1997;61(2):89–93. 41. Fortier LA, Mac Harg MA. Topical use of 5-fluorouracil for treatment of squamous cell carcinoma of the external genitalia of horses: 11 cases (1988–1992). J Am Vet Med Assoc 1994;205(8):1183–5. 42. Howarth S, Lucke VM, Pearson H. Squamous cell carcinoma of the equine external genitalia: a review and assessment of penile amputation and urethrostomy as a surgical treatment. Equine Vet J 1991;23(1):53–8.
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Neoplasia of the Reproductive Tract 43. Mair TS, Walmsley JP, Phillips TJ. Surgical treatment of 45 horses affected by squamous cell carcinoma of the penis and prepuce. Equine Vet J 2000;32(5):406–10. 44. Junge RE, Sundberg JP, Lancaster WD. Papillomas and squamous cell carcinomas of horses. J Am Vet Med Assoc 1984;185(6):656–9. 45. Velazquez EF, Cubilla AL. Penile squamous cell carcinoma: anatomic, pathologic and viral studies in Paraguay (1993–2007). Anal Quant Cytol Histol 2007;29(4):185– 98.
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46. Pratt-Thomas HR, Heins HC, Latham E, Dennis EJ, McIver FA. The carcinogenic effect of human smegma: an experimental study. Cancer 1956;9(4):671–80. 47. Markel MD, Wheat JD, Jones K. Genital neoplasms treated by en bloc resection and penile retroversion in horses: 10 cases (1977–1986). J Am Vet Med Assoc 1988;192(3):396– 400. 48. Novara G, Galfano A, De Marco V, Artibani W, Ficarra V. Prognostic factors in squamous cell carcinoma of the penis. Nature Clin Prac Urol 2007;4(3):140–6.
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Section (c): Management
114.
Drugs that Adversely Affect Spermatogenesis Rupert P. Amann
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115.
Dual Hemisphere Breeding Programs Norman W. Umphenour
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116.
Immunocastration Tom A.E. Stout and Ben Colenbrander
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Management of Stallions in Artificial Insemination H. Steve Conboy
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Management of Stallions in Natural-Service Programs Norman W. Umphenour, Phillip McCarthy and Terry L. Blanchard
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Nutrition and Exercise for Breeding Stallions Stephen G. Jackson
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Management of Subfertile Stallions Under Natural-Mating Conditions Terry L. Blanchard
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Venereal Disease Elizabeth S. Metcalf
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Chapter 114
Drugs that Adversely Affect Spermatogenesis Rupert P. Amann
Spermatogenesis and endocrine regulation of spermatogenesis are discussed in Chapters 96, 97, 99 and 101 and that information should be reviewed, if necessary, to understand the material in this chapter more clearly. Spermatogenesis is the most conspicuous aspect of testicular function, because disturbances of spermatogenesis often will be reflected in obvious changes in characteristics of semen from a stallion inadvertently or intentionally exposed to some noxious agent. Changes in spermatogenesis could be manifested as a reduction in number of spermatozoa produced, alteration of an attribute(s) of some or all of the spermatozoa produced, or concurrent changes of both types. Alternatively, the agent might affect epididymal function or pass into semen via secretions of the accessory sex glands and hence alter quality of the spermatozoa ejaculated without directly affecting the testes. Unfortunately, a change in seminal characteristics may not occur until 4–7 weeks after occurrence of a deleterious event, because of the time required for spermatogenesis and epididymal transit of sperm (Figure 114.1). Because spermatogonia or primary spermatocytes usually are affected once the agent is removed and normal primary spermatocytes are formed, an additional 4–6 weeks pass before normal spermatozoa can be ejaculated.1 Circumstances on many breeding farms will preclude detection of all disturbances of spermatogenesis. In severe cases, abnormal spermatogenesis will be evidenced by a reduction in size of one or both testes or an unusually firm or flaccid parenchyma. Both are detectable by palpation, provided earlier measurements or palpations of the testes of that stallion have established a baseline. Frequently, the first evidence of altered spermatogenesis will be a change in seminal characteristics. However, if seminal collections
are infrequent or procedures used to evaluate seminal quality are insensitive, seminal changes may remain undetected. Failure to detect an obvious change of seminal characteristics, testicular size or consistency, or hormone concentrations in blood serum does not mean that potential fertility has not changed. No laboratory test or combination of laboratory tests provides an accurate prediction of fertility of spermatozoa from a given male when they are used to inseminate a particular group of mares, either by natural mating or artificial insemination. The apparent fertility of a male is the product of his true fertility and that of the females with which he is mated. The implication of this fact is evident in several hypothetical cases (Table 114.1), in which fertility is expressed as a decimal fraction (0.64) rather than a percentage (64%). A 50% reduction in apparent fertility of a stallion (0.32 vs. 0.64) could be caused by a decline in his true fertility (case A) or to an unknown change in the population of mares to which he was bred (case B). In either case, to have a 90% probability of reaching the right conclusion that fertility was actually reduced, for whatever reason, each fertility estimate should be based on > 35 cycles in which timely inseminations occurred. The initial 25% reduction in fertility illustrated in case C might go undetected until the severity of the problem became worse. Detection of a decrease in fertility might be partially masked by insemination of an excessive number of spermatozoa.1, 2 On the other hand, insemination of an unusually high number of spermatozoa usually will not solve the problems of reduced fertility.3 Spermatogenesis places unique demands on the testis. A high rate of cell proliferation occurs in spermatogonia undergoing mitosis and spermatocytes undergoing meiosis. This imposes high demands for
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Stallion: Management Sperm Sm Sc
Se
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I
32
–39
II
IV
VI VII VIII
Cycle 4
I
II
45
–26
IV
VI VII VIII
57
Cycle 5 –14
0
Figure 114.1 A skeleton diagram of spermatogenesis in the stallion (see Figure 96.10 for details) to illustrate the intervals from death of specific types of germinal cells and when the effect might be anticipated in ejaculated semen. Cells designated A1 , A2 , A3 , B1 , and B2 are spermatogonia; P1, L, Z, P, and D are primary spermatocytes; 2◦ is a secondary spermatocyte; and SS , Se , Sc , and Sm are spermatids. An agent altering production of preleptotene spermatocytes (P1) will be detectable in terms of a decreased number of spermatozoa ejaculated, or altered spermatozoa, about 51 days later. However, an agent affecting newly formed spherical spermatids (Ss) might be detectable by altered seminal quality 34 days later. Similarly long intervals will be the minimum needed for restoration of normal spermatozoa once action of the agent stops. The duration of spermatogenesis is 57 days and it is assumed that 14 days are required for transport of spermatozoa through the epididymis.
delivery of glucose, amino acids, and lipid precursors to the interior of the seminiferous tubule, for energy and formation and remodeling of the germinal cells. Spermatogonia are in the basal compartment of the seminiferous tubule, and hence have relatively easy access to substrates in the interstitial fluid. However, more differentiated germinal cells must rely on Sertoli cells to provide all substrates and oxygen. Although data are not available for stallions, in rams a testis extracts about 15% of the glucose in testicular arterial blood and most of this is utilized by Sertoli cells.4 Much of this glucose is metabolized to lactate or pyruvate which, in turn, is oxidatively me-
tabolized by the developing germinal cells to provide the energy necessary for their development. The testis also has a high demand for oxygen, and the normal oxygen tension in the seminiferous tubule is relatively low. Thus, agents or events which depress delivery of glucose or oxygen to the seminiferous tubules will have a deleterious effect, even though general bodily functions remain unimpaired. Reproductive toxins may act by virtue of structure similar to an endogenous hormone, growth factor, or other compound involved in reproduction, because of their chemical reactivity to alkylate or chelate, because a metabolite of the agent exerts a toxic effect
Table 114.1 Role of the true fertility of groups of mares on the apparent fertility of stallions and the approximate numbers of mares needed to detect changes in true fertility of a stallion True fertility Case A B C
Stallion
Mares
Apparent fertilitya
0.80 0.40 0.80 0.80 0.80 0.60 0.30
0.80 0.80 0.80 0.40 0.70 0.70 0.70
0.64 0.32 0.64 0.32 0.56 0.42 0.21
Difference
Number of cycles to detect decreased fertilityb
0.32
35
0.32
35
0.14 0.35
100 33
Apparent fertility is the product of true fertility of the stallion and the mares to which he is mated: (0.40 × 0.80) = 0.32. The approximate number of cycles required to have a 90% chance of detecting this difference in apparent fertility is shown. This number of inseminations is necessary for establishing initial fertility and for determining if a reduction of fertility of the magnitude indicated has occurred.
a
b
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Drugs that Adversely Affect Spermatogenesis such as inhibiting or inducing an enzyme, or by a combined action.5 For a stallion, perhaps agents that mimic endogenous hormones are the greatest problem, because of intended administration. However, other types of agents should be considered in cases where administration of exogenous hormones can be excluded. Agents affecting delivery of spermatozoa, rather than production of normal spermatozoa and accessory sex gland fluids, should not be ignored. For example, drugs with α-adrenergic-blocking properties can result in failure of seminal emission or retrograde ejaculation in humans.6 However, combined administration of a β-adrenergic receptor blocker and an α agonist to stallions can induce emission in otherwise impotent stallions.7 Agents affecting spermatogenesis might act directly on the germinal cells by interfering with synthesis or function of flagellar elements.8 Such drugs will block cell division, so that primary spermatocytes are not formed, or differentiation of spermatids, so that any spermatozoa produced are malformed. Other drugs or reprotoxins alter function of Sertoli cells, and the lesion(s) on germinal cells are secondary to abnormal Sertoli cell function.1 Perhaps the most common clinical cause of abnormal spermatogenesis is a secondary response to abnormal function of the hypothalamic–hypophyseal–testes axis. Insufficient luteinizing hormone (LH) for stimulation of Leydig cells to secrete sufficient testosterone and to maintain the high concentration around the seminiferous tubules essential for normal Sertoli cell function, or insufficient follicle-stimulating hormone (FSH) to maintain Sertoli cell function will adversely affect spermatogenesis. Little information exists on effects of drugs or other agents on testicular function in stallions. Hence, the following discussion is based mainly on data for humans, mice, rats, and other animals. Data for stallions are identified when presented.
Agents that might act directly on germinal cells Germinal cells are actively dividing, and hence are susceptible to any drug which blocks cell division. Clearly γ radiation or x-rays can directly affect spermatogenesis, and the spermatogonia are much more sensitive to radiation than spermatocytes or spermatids.9, 10 Antineoplastic drugs (e.g., chlorambucil, cyclophosphamide, doxorubicin, and procarbizine) are selected because they kill rapidly dividing cells in tumors, but this property also targets them against spermatogonia. A decrease in number of spermatozoa ejaculated will be evident several weeks after initial treatment and the last spermatozoa produced may be malformed. The reserve spermatogonia are
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often spared because they rarely divide, and in many cases surviving reserve spermatogonia ultimately can repopulate the seminiferous epithelium. Alkylating agents (e.g., dibromochloropropane and ethylene dibromide, both used as pesticides or fumigants, or methyl chloride), colchicine, or the fungicide methyl 2-benzimidazolecarbamate (Carbendazim) all block division of germinal cells in several species.
Agents that might act on Sertoli cells During the evolution of most mammals, the Sertoli cells developed so that they produce necessary amounts of molecules needed by developing germinal cells only at a temperature 3–6◦ C below core body temperature. This is clearly evident from studies with cultured Sertoli cells, which have reduced secretory capacity at 37◦ C compared with that at 32◦ C.11 Studies are beginning to provide a molecular basis for the well-known, and clinically important, sensitivity of the seminiferous epithelium to elevated testicular temperature. Although pachytene spermatocytes and B-spermatogonia are the germinal cells most sensitive to elevated testicular temperature,4 the primary lesion causing their death and degeneration may be in Sertoli cells. In any case, summer heat or febrile conditions can induce a transitory decrease in number and quality of spermatozoa produced. Chemicals also can alter Sertoli cell function, and this probably is a common primary site of action for reprotoxins, the effect of which is seen secondarily by a decrease in number of normal spermatozoa ejaculated. For example, both di(2-ethylhexyl) phthalate and 1,3-dinitrobenzene act directly on Sertoli cells,12 and vacuolation of the cytoplasm and exfoliation of germ cells are characteristic sequelae. Di(2-ethylhexyl) phthalate is converted to an active ester by Sertoli cells, and this compound decreases response of Sertoli cells to FSH.12 Dinitrobenzene suppresses production of lactate and pyruvate by Sertoli cells, and this compromises the ability of Sertoli cells to nurture germinal cells.13 Pachytene spermatocytes and spermatids undergo degeneration. Drugs which modify hepatic function also might alter Sertoli cell function, because both hepatocytes and Sertoli cells secrete many transport proteins (e.g., transferrin and ceruloplasmin). Those produced by Sertoli cells are essential for spermatogenesis.
Agents that might act on Leydig cells Most alterations of Leydig cell function are a consequence of a change in the hypothalamic– hypophyseal–testes axis (as discussed in the next
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section), but some reprotoxins act directly on Leydig cells. For example, 2,3,7,8-tetrachlorodibenzop-dioxin blocks production of pregnenolone by Leydig cells,14 and hence testosterone cannot be produced (see steroidogenic pathway depicted in Figure 96.15). Ethane dimethanesulfonate apparently has a similar action.8, 15 In addition to environmental toxins, such as in the two examples, steroids or other drugs affecting androgen or estrogen synthesis can act directly on Leydig cells.
CNS T
Normal
GnRH T
Anterior pituitary
E Accessory sex glands
I LH
T Leydig cells
Agents that might affect the hypothalamic–hypophyseal–testes axis Normal spermatogenesis depends on the negative feedback regulation of the hypothalamic– hypophyseal–testes axis (Figure 114.2a). This provides sufficient FSH and testosterone to the seminiferous epithelium for maximum production of normal spermatozoa; testosterone in amounts necessary for normal function of accessory sex glands; steroids for expression of sexual behavior and achievement of erection of the penis at copulation; and negative feedback of testosterone, estradiol, and inhibin on the central nervous system, hypothalamus, and adenohypophysis (anterior pituitary gland). Unfortunately, both natural events and human intervention can alter this delicate hemostatic balance.
Hypothalamus
E
The most common pharmacological manipulation of spermatogenesis probably is the unintended sideeffect of administration of testosterone esters or anabolic steroids to stallions. Although the intent is to raise the concentration of androgen circulating in the blood and available to muscle tissue and bone, this therapy also raises the concentration of androgen impinging on the hypothalamus and anterior pituitary gland. As depicted in Figure 114.2b, the high concentration of androgen in systemic blood causes a depression or cessation of secretion of gonadotropinreleasing hormone (GnRH) and LH and probably a decrease in secretion of FSH. Because of unavailability of LH, secretion of testosterone by Leydig cells is decreased and the concentration of testosterone in the interstitial fluid bathing the seminiferous tubules is reduced to a concentration insufficient for normal spermatogenesis. In many cases, Leydig cells would produce no testosterone and the concentration of testosterone in interstitial fluid would be equal to that in peripheral blood rather than 20–100 times greater. Testicular production of estrogens also would be reduced or eliminated.
Sertoli cells
Germ cells
T
E
(a)
CNS T
Androgen
Hypothalamus
E GnRH T
Anterior pituitary
E Accessory sex glands
I LH
T
Leydig cells
Androgens
FSH
E
T
Sertoli cells
FSH Germ cells FEW or ABNORMAL
(b) Figure 114.2 The hypothalamic–hypophyseal–testes axis in a normal stallion (a) and a stallion receiving an injection(s) of androgen (b). Injection of androgen elevates (larger solid arrow) the concentration of androgen (T) in the blood, and this suppresses (smaller open arrow) secretion of GnRH by the hypothalamus and FSH and LH by the anterior pituitary gland. Secretion of testosterone and estrogens (E) declines. The effect on concentration of inhibin (I) is unknown, but an increase might occur. Although blood concentration of androgen is high, the intratesticular concentration of androgen is low, and Sertoli cells will receive insufficient stimulation of androgen and FSH. Consequently, with prolonged administration of androgen, a decrease or cessation of spermatogenesis will occur.
This is not just a hypothetical scenario. Countless clinical observations and direct experimentation show the severity of the problem in stallions16–20 and the potential for reversibility or restoration of normal spermatogenesis. In stallions receiving testosterone propionate intramuscularly (200 µg/kg every other day) for 88 days,17 a gradual decrease in
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Drugs that Adversely Affect Spermatogenesis
110 Control
105 100 95
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90 85 80
10
20
30 40 50 60 Days of treatment
70
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not shown) had recovered, because characteristics for stallions receiving 200 µg/kg testosterone propionate were similar to those for untreated (control) stallions (Table 114.2). Thus, although recovery from suppression of spermatogenesis occurred in stallions receiving 200 µg/kg testosterone propionate every other day, treatment for a longer interval or at a higher dose might have more severe or longer lasting effects. Anabolic steroids rather than testosterone esters frequently are administered to stallions, although this is not an approved use. Anabolic steroids have structures similar to those of testosterone or testosterone propionate (Figure 114.4), and thus they might be expected to affect spermatogenesis adversely. This was shown to be the case in a comprehensive study19 which compared changes in stallions treated with boldenone undecylenate (Equipose) (0.23 or 0.91 mg/kg body weight) or nandrolone decanoate (DecaDurabolin) (0.23 mg/kg body weight) by intramuscular injection once every third week for 15 weeks. Eight stallions received each treatment, and the experiment was conducted with three replicates, staggered so that the 15-week treatment period ended in September, October, or November. For all three groups of treated stallions, total scrotal width decreased compared with that for untreated (control) stallions (Figure 114.5). Characteristics of semen collected during weeks 13–15 of treatment were compared with pretreatment values. Total spermatozoa per ejaculate, percentage of motile spermatozoa, and percentage of normal spermatozoa all were greatly suppressed in the treated stallions, and these changes were greater than those attributable to season in the untreated stallions (Table 114.3). Severity of the damage induced by these anabolic steroids was evident in comparisons of testicular weight after 15 weeks of treatment (Table 114.3). Similar data were reported by Blanchard et al.,20 who studied pony stallions treated with stanozolol
115
Scrotal width (mm)
c114
90
Figure 114.3 Total scrotal width in control stallions and in stallions receiving testosterone propionate (200 µg/kg). (Modified from Pickett et al.16 )
testicular size was noted, determined as total scrotal width (Figure 114.3). Concentrations of LH in serum from treated stallions averaged approximately 50% of the values for control stallions. By 74–90 days after the start of treatment, spermatozoal concentration in ejaculated semen, total number of spermatozoa per ejaculate, percentage of motile spermatozoa, and percentage of normal spermatozoa were reduced from their pretreatment values; volumes of gel or gelfree semen were not significantly altered. The adverse effect of testosterone propionate is evident in data comparing the control and treated stallions (Table 114.2). Some of the stallions were castrated 90 days after starting the treatment and the remaining stallions were allowed 92 days to recover; semen was collected between days 166 and 180 when these stallions were castrated.18 Total sperm per ejaculate and testicular weight as well as spermatozoal production (data
Table 114.2 Effect of treatment of stallions with testosterone propionate for 90 days Days 74–90 (N = 8)
Gel-free semen (mL) Spermatozoa/mL (106 ) Spermatozoa/ejaculate (109 ) Motile spermatozoa (%) Normal spermatozoa (%) Testes weight (g)
Days 166–180 (N = 4)
Control
TPa
Difference (%)
Control
TP
Difference (%)
35 230 6.7 58 53 338
40 87 3.3 50 39 219
+14 –62 –51 –14 –26 –35
43 104 3.5 63 – 283
51 64 3.2 60 – 365
+19 –38 –8 –5 – +29
a Treated stallions received 200 µg/kg testosterone propionate (TP), intramuscularly every other day for 88 days. Four stallions per treatment group were castrated on day 90 and four were allowed to recover until day 180. Semen was collected every other day from days 74–90 and 166–180, after depleting extragonadal spermatozoal reserves. Adapted from Hoyer JH. The effect of testosterone on reproductive function in stallions. MS thesis, Colorado State University, 1978; and Squires et al.18
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O
Nandrolone decanoate
Figure 114.4 Structures of testosterone and related steroids sometimes administered to stallions, although administration of these compounds to stallions is not a recommended practice or an approved use. All have androgenic effects, and boldenone undecylenate and nandrolone decanoate have enhanced anabolic activity.
(Winstrol-V) or boldenone undecylenate (each tested at 0.55 or 1.11 mg/kg) for 13 weeks. Testes weight was reduced in all treated stallions, relative to the untreated (control) group, except those receiving 0.55 mg/kg stanozolol. As discussed elsewhere, administration of testosterone propionate or anabolic steroids appeared to have no beneficial effect on sexual behavior of stallions or growth rate of healthy geldings.16 Because such compounds do have adverse effects on spermatogenesis, androgenic compounds should not be administered to healthy stallions to be used for breeding. Human chorionic gonadotropin Occasionally a horse owner or veterinarian will consider administration of human chorionic gonadotropin (hCG) to a stallion, in the belief that an increase in testosterone might help some reproduc-
1D
75 PT
1
3
5 7 9 11 Week of treatment
13
15
Figure 114.5 Total scrotal width in control stallions and stallions receiving anabolic steroids before treatment (PT) and during the treatment period. Groups are designated as: C, control; 1E, 0.23 mg/kg boldenone undecylenate; 4E, 0.91 mg/kg boldenone undecylenate; and 1D, 0.23 mg/kg nandrolone decanoate. Stallions received steroid every 3 weeks. (Modified from Pickett et al.16 )
tive function. Concentration of testosterone in peripheral blood is doubled in < 1 hour after intravenous injection of hCG, and increased secretion of testosterone may continue for several days.21 Although both LH and hCG bind to LH receptors on Leydig cells, the two hormones are chemically and biologically different. Because hCG is more glycosylated, it is cleared more slowly from the blood and available to LH receptors on target cells far longer than LH. Target cells, and presumably stallion Leydig cells, process hCG–receptor complexes differently from LH–receptor complexes, further prolonging the stimulatory effect and interval of increased steroid secretion. Because concentration of testosterone in peripheral blood may be increased for days rather than hours after a single injection, changes in endocrine homeostasis may occur (Figure 114.6),
Table 114.3 Effect of treatment of stallions with anabolic steroids for 15 weeksa
Gel-free volume Spermatozoa/mL (106 ) Spermatozoa/ejaculate (109 ) Motile spermatozoa (%) Normal spermatozoa (%) Testes weight (g) a
Cb
1Ec
4Ed
1De
30 226∗ 5.0∗ 55∗ 68 285∗
22 117† 25† 34† 63 176†
17 60‡ 1.4† 34† 62 128‡
20 30‡ 05† 21‡ 53 114‡
Means for eight stallions per group. Treated animals were injected intramuscularly every 3 weeks. Semen was collected every other day for the last 22 days of the experiment. b Control stallions. c Stallions received 0.23 mg/kg boldenone undecylenate. d Stallions received 0.91 mg/kg boldenone undecylenate. e Stallions received 0.23 mg/kg nandrolone decanoate. Adapted from Squires et al.19
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Drugs that Adversely Affect Spermatogenesis
CNS T
Normal
Hypothalamus
E GnRH T
Anterior pituitary I
E Accessory sex glands
FSH
LH
T Leydig cells
Germ cells
Sertoli cells
T
E
CNS T
Hypothalamus
E GnRH T
Anterior pituitary I
E Accessory sex glands
LH
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Sertoli cells
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and hypothalamus cause suppression of GnRH secretion. The anterior pituitary gland, in turn, secretes less LH and FSH. The lack of LH is not a problem because the Leydig cells are receiving stimulation of hCG. However, the lack of FSH available to Sertoli cells could have deleterious effects on spermatogenesis if administration of hCG was prolonged. Both number of spermatozoa produced and spermatozoal structure could be affected, although no valid data exist for stallions showing sequelae of chronic administration of hCG. Repeated injection of a stallion with hCG is not recommended. Gonadotropin-releasing hormone
(a)
HCG
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FSH
Germ cells
T
(b)
Figure 114.6 The hypothalamic–hypophyseal–testes axis in a normal stallion (a) and in a stallion receiving an injection(s) of hCG (b). Injection of hCG stimulates (larger solid arrow) Leydig cells to secrete testosterone (T), and the high concentration of testosterone in blood reaching the hypothalamus and anterior pituitary gland suppresses (smaller open arrow) secretion of GnRH, FSH, and LH. Although there may be adequate testosterone and estrogens (E) secreted by Leydig cells, Sertoli cells will be deprived of FSH. Concentration of inhibin (I) probably would not change following a single injection of hCG. Consequently, with repeated administration of hCG, the seminiferous epithelium receives insufficient stimulation of FSH for normal spermatogenesis.
especially with a series of injections of hCG. Assuming that the endocrine profiles of a stallion are reasonably normal, injection of hCG causes increased production of testosterone by Leydig cells and abundant testosterone is available to the seminiferous epithelium; this is not deleterious to spermatogenesis. Secretion of estrogens by Leydig cells also increases. The increased concentrations of testosterone and estradiol impinging on the central nervous system
Use of GnRH to stimulate secretion of LH by the anterior pituitary gland and, thereby, secretion of testosterone by Leydig cells is more biologically normal than administration of hCG to stimulate secretion of testosterone. In a normal stallion, and many stallions with changes in the homeostatic balance of the hypophyseal–hypothalamic–testes axis, intravenous or subcutaneous injection of a bolus of GnRH is followed by increased secretion of FSH and especially LH. Availability of a transitory rise in LH impinging on the Leydig cells results in temporary elevations of concentrations of testosterone and estrogens in the interstitial fluid and blood. Thus, in contrast to injection of hCG, injection of GnRH results in an elevation of both FSH and, at least under some conditions, testosterone in fluid bathing the seminiferous tubules, rather than only testosterone. Furthermore, assuming appropriate dosage of GnRH is used (Chapter 147), the probability of down regulating LH or testosterone receptors on Leydig cells or Sertoli cells, respectively, is low whereas it is a virtual certainty after administration of hCG. The short interval of elevated blood concentrations of testosterone and estrogens after GnRH administration provides a relatively normal negative drive to the hypothalamic–hypophyseal axis. In theory, pulsatile administration of GnRH could be used to mimic the endogenous secretion of GnRH and hence drive a stallion’s endocrine system. Effects of this treatment have been studied in normal stallions. As anticipated, pulsatile injections of GnRH (1.25–5.0 µg/kg each 0.5 h or 10 µg/kg each 2 h, in the two experiments) resulted in pulsatile discharge of LH and FSH. However, in normal stallions the effect on testosterone secretion is more pronounced during the winter than summer,22 and this may explain why Blue et al.23 found no increased blood concentration of testosterone in normal stallions receiving pulsatile injections of GnRH. Pulsatile injection of GnRH into an abnormal stallion (with low serum concentrations of LH and testosterone similar to those of a normal
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CNS Hypogonadotrophic hypogonadism
T
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E GnRH T
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E Accessory sex glands
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FEW or ABNORMAL
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E GnRH T
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Figure 114.7 The hypothalamic–hypophyseal–testes axis in an abnormal stallion, with a central nervous system or hypothalamic abnormality resulting in a lack (small open arrow) of GnRH secretion (a) or a similar abnormal stallion receiving injections of GnRH (b). Pulsatile injection of GnRH stimulates the anterior pituitary gland to secrete LH and FSH (larger solid arrow). The LH, in turn, stimulates Leydig cells to secrete testosterone (T) in a pulsatile manner. Thus adequate amounts of FSH and testosterone might reach the seminiferous epithelium to stimulate restoration of spermatogenesis, and sufficient testosterone and estrogens (E) might reach other organs for normal expression of feedback regulation and other reproductive functions or behaviors. Although not studied, concentration of inhibin (I) might decline, especially if it had been elevated initially. Such GnRH therapy would have to continue for several months, because it might take several weeks of GnRH stimulation before normal function of Sertoli cells was restored, and the duration of spermatogenesis is 57 days in stallions.
stallion in the winter) probably would result in pulsatile secretion of LH and testosterone even in the spring or summer. Some stallions may have clinical symptoms of adult-onset hypogonadotropic hypogonadism, namely, reduced production of normal spermatozoa and low concentrations of LH and especially testos-
terone in peripheral blood (Figure 114.7a); blood concentration of FSH might be normal or elevated. In these stallions, the testicular malfunction is secondary to an insufficiency of GnRH. Diagnostic use of GnRH to distinguish these stallions from others with advanced or irreversible testicular degeneration is discussed in Chapter 147. In stallions where the primary problem is insufficient secretion of GnRH, the anterior pituitary gland should respond to an injection of GnRH with increased secretion of LH, and increased secretion of testosterone by Leydig cells is likely (Figure 114.7b); this indeed happens (T.M. Nett, personal communication). Appropriate pulsatile injection of GnRH may result in an improvement in spermatozoal production or quality after 1–2 months of treatment. Anecdotal reports of clinicians with an equine practice and published reports by physicians provide evidence that this treatment can be effective in some individuals, although others with an apparently similar clinical picture will not benefit.24–26
Acknowledgement Some individuals, and some manuscripts have wonderful insight and historical value. Such is the case with Dr. Rupert P. Amann and four chapters that he so ably produced for the first edition of Equine Reproduction. While Dr. Amann is now retired and has turned the baton over to others, these pieces of work have endured the test of time and continue to be an invaluable resource for the equine practitioner and academician. As such, we decided to reprint these chapters for you.
References 1. Amann RP. Detection of alterations in testicular and epididymal function in laboratory animals. Environ Health Perspect 1986;70:149–58. 2. Amann RP. Can the fertility potential of a semen sample be predicted accurately? J Androl 1989;10:89–98. 3. Pickett BW, Squires EL, McKinnon AU. Procedures for Collection, Evaluation and Utilization of Stallion Semen for Artificial Insemination. Animal Reproduction Laboratory Bulletin No. 03. Fort Collins: Colorado State University, 1987. 4. Waites GMH, Setchell BP. Physiology of the mammalian testis. In: Lamming GE (ed.) Marshall’s Physiology of Reproduction, 4th edn. London: Churchill Livingstone, 1990; pp. 1–105. 5. Mattison DR, Thomford PJ. Mechanism of action of reproductive toxicants. In: Working PK (ed.) Toxicity of the Male and Female Reproductive Systems. New York: Hemisphere, 1989; pp. 101–29. 6. Benson GS. Male sexual function. In: Knobil E, Neill JD (eds) The Physiology of Reproduction, vol. 1. New York: Raven Press, 1988; pp. 1121–39.
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Drugs that Adversely Affect Spermatogenesis 7. Klug E, Deegen E, Lazarz B, Rojem I, Merkt M. Effect of adrenergic neurotransmitters upon the ejaculatory process in the stallion. J Reprod Fertil Suppl 1982;32:31–4. 8. Linder RE, Klinefelter GR, Strader LF, Narotsky MG, Suarez JD, Roberts NL, Perreault SD. Dibromoacetic acid affects reproductive competence and sperm quality in the male rat. Fundam Appl Toxicol. 1995;28:9–17. 9. Meistrich ML. Interspecies comparison and quantitative extrapolation of toxicity to the human male reproductive system. In: Working PK (ed.) Toxicity of the Male and Female Reproductive Systems. New York: Hemisphere, 1989; pp. 303–21. 10. Oakberg EF. Irradiation damage to animals and its effect on their reproductive capacity. J Dairy Sci 1960;43(Suppl.):54–64. 11. Hagenas L, Ritzen EM, Suginami HL. Temperature dependence of Sertoli cell function. Int J Androl Suppl 1978;2:449–58. 12. Chapin RE, Foster PMD. Testis en plastique: use and abuse of in vitro systems. In: Working PK (ed.) Toxicity of the Male and Female Reproductive Systems. New York: Hemisphere, 1989; pp. 273–84. 13. Williams J, Foster PMD. The production of lactate and pyruvate as sensitive indices of altered rat Sertoli cell function in vitro following the addition of various testicular toxicants. Toxicol Appl Pharmacol 1988;9:160–70. 14. Kleeman JM, Moore RW, Peterson RE. Inhibition of testicular steroidogenesis in 2,3,7,8-tetrachlorodibenzo-p-dioxintreated rats: evidence that the key lesion occurs prior to or during pregnenolone formation. Toxicol Appl Pharmacol 1990;106:112–25. 15. Klinefelter GR, Laskey JW, Roberts NL. In vitro/in vivo effects of ethane dimethanesulfonate on Leydig cells of adult rats. Toxicol Appl Pharmacol 1991;107:460– 71. 16. Pickett BW, Amann RP, McKinnon AO, Squires EL, Voss JL. Management of the Stallion for Maximum Reproductive Effi-
17.
18.
19.
20.
21.
22.
23.
24.
25. 26.
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ciency, II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins: Colorado State University, 1989. Berndtson WE, Hoyer JH, Squires EL, Pickett BW. Influence of exogenous testosterone on sperm production, seminal quality and libido of stallions. J Reprod Fertil Suppl 1979;27:19–23. Squires EL, Berndtson WE, Hoyer JH, Pickett BW, Wallach SJ. Restoration of reproductive capacity in stallions after suppression with exogenous testosterone. J Anim Sci 1981;53:1351–9. Squires EL, Todter GE, Berndtson WE, Pickett BW. Effect of anabolic steroids on reproductive function of young stallions. J Anim Sci 1982;54:576–82. Blanchard TL, Elmore RG, Youngquist RS, Loch WE, Hardin DK, Bierschwal CJ, Ganjam VK, Balke JM, Ellersieck MR, Dawson LJ, Miner WS. The effects of stanozolol and boldenone undecylenate on scrotal width, testis weight, and sperm production in pony stallions. Theriogenology 1983;20:121–31. Amann RP, Ganjam VK. Effects of hemicastration or hCGtreatment on steroids in testicular vein and jugular vein blood of stallions. J Androl 1981;3:132–9. Roser JF, Hughes JP. Prolonged pulsatile administration of gonadotrophin-releasing hormone (GnRH) to fertile stallions. J Reprod Fertil Suppl 1992;44:155–68. Blue BJ, Pickett BW, Squires EL, McKinnon AO, Nett TM, Amann RP, Shiner KA. Effect of pulsatile and continuous administration of GnRH on reproductive function of stallions. J Reprod Fertil Suppl 1992;44:145–54. Baker HWG, Kovacs GT. Spontaneous improvement in semen quality: regression towards the mean. Int J Androl 1985;8:421–6. Wagner TOF. Pulsatile LHRH Therapy of the Male. Hamein: TM-Verlag, 1985. Winter SJ. Evaluation and management of hypogonadotropic hypogonadism. Serono Symp Rev 1989;20: 93–102.
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Chapter 115
Dual Hemisphere Breeding Programs Norman W. Umphenour
The process of shipping (shuttling) stallions from the northern to the southern hemisphere in order to breed two seasons in one calendar year began in the early 1970s. In the 1990s the number of stallions shuttling from the USA to the southern hemisphere for breeding started to increase, even though there had been some shuttling in the early 1980s. At the present time it has become commonplace for stallions to shuttle to Australia, New Zealand, South America, and South Africa, depending on the demand for certain bloodlines by the breeders and the strength of the sales and racing markets in these countries.
Some considerations for stallions selected for shuttling When contemplating the transport of a prospective shuttle stallion, there are factors to be considered in selecting such a stallion. These factors include inherited fertility, age, testicular size, sexual behavior, frequency of ejaculation, financial viability, season, and physical condition. Fertilization and pregnancy rates are terms commonly used to denote fertility. Many breeders contend that the only true measure of fertility is percentage of live foals. Obviously this is quite relevant. Unless a mare delivers a live, healthy foal, all other measurements are merely time consuming and expensive. However, for appropriate management of the stallion for maximum reproductive efficiency other aspects of fertility must be understood. In some areas and under certain circumstances, one may consider a foaling rate of 50% as satisfactory, while in another area anything less than a 65% foaling rate may be deemed unsatisfactory, depending on the individual perception and conditions determining
the perception. Therefore, fertility is a relative term that must be qualified to be meaningful. What should one expect for fertility from breeding to parturition (i.e., production of a live foal)? If a 10% embryonic loss rate is assumed from fertilization to Day 14, 15% from Day 14 to 50, and 10% from Day 50 to parturition, foaling rate would be in the 60–65% range for a normal, fertile stallion bred to normal fertile mares. There are numerous factors affecting fertility, which result in considerable variation in foaling rates among farms and stallions, but when a foaling rate of less than 65% is obtained, management, including animals, should be thoroughly evaluated to identify areas for improvement.1 Over the past few years, the number of mares bred per stallion has greatly increased. Formerly, a book of 40 mares was the industry standard, but more recently books of 120–150 mares or more have become the norm. Some data suggest that stallions that are bred to a book of 120 mares or more are likely to have a higher percentage of live foals (Table 115.1).2 This may be due to stimulation of the stallion’s reproductive system to maximum capacity as a result of exposure of the stallion to more mares; alternatively, the larger books usually are associated with increased servicefee stallions that are on well-managed breeding farms. This information should aid owners of shuttle stallions in determining if their stallion’s fertility is within expected ranges needed to handle a larger mare book, or year-round breeding. Most farms standing stallions keep sufficient records, many of which are computerized, to calculate fertility for each stallion. Pregnancy rate per cycle, particularly first cycle pregnancy rate, is one of the most important measures of fertility. Pregnancy rate by month and pregnancy rate by mare
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Dual Hemisphere Breeding Programs Table 115.1 Live foal rate achieved by Thoroughbred stallions grouped on the basis of the number of mares bred per stallion during each breeding season (book size). Live foal rate calculated as the number of foals born alive divided by the number of mares bred; values are reported as percentages.2 Book size Year
≤ 40
41–80
81–120
≥ 120
Total
1988 2003 2004 2005
54a 46a 45a 44a
67b 60b 58b 58b
71b 68c 67c 67b,c
NA 70c 73c 72c,d
55 47 47 46
NA, not applicable; no stallions bred > 120 mares. a to d within a row: values with different superscript letters differ significantly (P = 0.01; ANOVA). Table reproduced from Turner and McDonnell,2 by courtesy of the Journal of the American Veterinary Medical Association.
status can also be extremely helpful for diagnostic purposes when pregnancy rates vary consistently during any period of the breeding season. The number of spermatozoa needed for maximum reproductive efficiency depends on a stallion’s inherent fertility. Highly fertile stallions with good mare management can potentially get mares pregnant with 100 million sperm or less with natural service or artificial insemination (AI). The number of covers and the number of sperm delivered with each mating are two such factors to consider. The third factor is a stallion’s sperm longevity. Consequently, fewer covers are probably needed with stallions that have increased sperm longevity. With the use of ultrasonography for examination of mare reproductive status, sperm longevity in a mare’s reproductive tract can be determined by the interval from day of cover to ovulation. Mares that become pregnant when bred near ovulation but also produce additional follicles that ovulate at a later date will have different embryonic sizes when examined for pregnancy status by ultrasonography approximatley 13–15 days after service. I expect that most stallions of average fertility can produce spermatozoa with longevity of 96 to 120 hours in a reproductively sound mare. Some young, highly fertile stallions can produce spermatozoa that remain viable for up to 10 days (T.W Riddle et al., personal communication). With natural breeding, a conscientious effort is made to breed each mare only once during estrus. Fertility on farms in which mares are well managed is excellent. Fertility appears to increase as the number of mares in a stallion’s book increases for many fertile stallions. When considering the shuttling of stallions to the southern hemisphere, there is one factor that plays
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an important part in the breeding efficiency of the stallion. That factor is that 95% plus of the mares are generally kept on the same farm as the stallion in the southern hemisphere. Thus, with the mares being under one management program, they can be selected at the most ideal time for mating. In addition, the number of hours between covers is greater, i.e., approximately 6 hours, whereas in the northern hemisphere, the time between covers is shorter (4 hours) and mares being mated come from several different management programs. At one particular farm in central Kentucky, the stallions are covering mares that are transported from approximately 350 different satellite farms. So when considering a stallion for possible shuttle duty the following factors must be considered to determine if the prospective stallion can withstand the pressure of doing “back-to-back” breeding seasons in a calender year. 1. Stallion’s inherent fertility. To maintain maximum reproductive efficiency of stallions, particularly shuttle stallions, they must be well managed. Therefore, prior to the breeding season every stallion should have an evaluation of his seminal characteristics because they are related to fertility and will provide information for the stallion manager to alter management techniques to improve fertility. 2. Age and testicular size. The young stallion, 2–3 years of age, may not have sufficient testicular size or sperm output needed to efficiently handle a large book of mares (150–200) through the breeding season. In some instances, failure of a young stallion to cope with frequent covers may result in the need to reduce the stallion’s mare book. However, the older stallion (4–16 years) may have the testicular capacity and sperm output to handle a large book of mares, and in some situations additional mares may be added to his book. In general, the testicular capacity of normal stallions increases to about 11 years of age then begins to decrease between 11 and 14 years of age. Thus, using a 16-year-old stallion for shuttling must be seriously evaluated.1 3. Frequency of ejaculation and sexual behavior go hand in hand. Adequate, if not enthusiastic, expression of motivated behavior by a breeding stallion is essential for economic success of a breeding farm. A breeding farm suffers dramatically when a stallion’s mating behavior becomes erratic or unpredictable. More time and labor are spent for each breeding and fewer mares will become pregnant per cycle and over a breeding season. Stallion managers who have experienced the frustration of standing a ‘slow’ stallion at stud will appreciate the joy of handling a stallion that is sexually motivated. A responsive stallion will
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readily attain an erection upon presentation to a mare, mount immediately, copulate vigorously, and deliver a complete ejaculate without dismounting! The anxiety of an individual who is handling a slow stallion or one with abnormal sexual behavior or poor sex drive is exceeded only by the breeding manager’s fear that the stallion will not become sexually aroused and thus the mare(s) will not be bred at a time appropriate for maximizing the chance of becoming pregnant. Thus, frequency of ejaculation with a stallion of normal mating behavior is another important factor in sperm output for selecting a shuttle stallion. As the frequency of ejaculation increases, the number of spermatozoa per ejaculation decreases. Depending on testicular size and extragonadal sperm reserves, the stallion management will have to determine the number of times per day the stallion can be mated, which will dictate the number of mares the stallion can be bred to in the number of days for that particular breeding season.1 4. Season. Season has been shown to influence reproductive characteristics in stallions. This influence has been characterized by an increase in testicular size and weight, sperm production and output, sexual behavior, and concentration of reproductive hormones during the long days of the year because the stallion is a ‘long day’ breeder. It is generally assumed that the events triggering maximum reproductive capacity in stallions in May and June in the northern hemisphere occur in November and December in the southern hemisphere. Thus, the period of time from the ending of one breeding season (northern hemisphere) to the beginning of the next breeding season (southern hemisphere) will overlap. Perhaps the brief period of time when the stallion is again exposed to short daylength on return to the northern hemisphere in December-January is sufficient to ‘reset’ the photoperiodic control of his endocrine system so that he again experiences appropriate hormonal stimulation of sexual behavior and spermatogenesis. This may allow the production of reproductive hormones to maintain a high level. From the long days of light, as the southern hemisphere season comes to an end (December) the stallion is shuttled back to the northern hemisphere. This period of time from December to February with the ‘short days’ of light may allow the system for reproductive hormones to be ‘reset’. This area, however, has not been critically studied. The author does not currently recommend subjecting shuttle stallions to an artificial lighting program because of the possibility of the stallion becoming refractory to photoperiodic stimulation.
5. Physical condition. This is another factor that must be considered when selecting a shuttle stallion. Consider how sound he might be to withstand the rigors of shipping from one hemisphere to the other and how he can handle back-to-back breeding seasons. Horses that have racetrack injuries that may cause chronic pain, such as suspensory injuries, arthritic ankles/hocks, sore backs, cervical problems, or laminitis, may be poor risks for shuuttling. If the pain cannot be controlled with medication, shoeing, or some form of physical therapy, these conditions can greatly affect a stallion’s libido and ability to cover his mares. With the stress of shuttling and the changes in management practices, grooms, handlers, and environment, even apparently well-adjusted stallions can show signs of abnormal sexual behavior with continued shuttling. The most common signs will be: 1. Poor sex drive. 2. Easily distracted (in the breeding shed). 3. More frequent mounting without an erection or without a full erection. 4. Require multiple mounts per ejaculation. 5. Poor pelvic thrust when mating. 6. Dismount quickly before or after ejaculation. If before, you have a horse becoming a ‘cheater’. If after, the glans is ‘belled out’ so that there is a suction effect that pulls the ejaculate or semen out of the vaginal tract when dismounting. 7. Tendency to savage mares. 8. Failure to attain or maintain an erection. 9. Incomplete intromission. 10. Failure to ejaculate in spite of a complete, prolonged erection and multiple intromissions. 11. Show excellent sexual behavior only sporadically. 12. Stallions shuttling the first time, especially young first-year stallions, may not be able to handle the stress of change in management, etc. Such things as total loss of, or poor, sex drive or selfmutilation syndrome could develop.3 Regardless of the physiological and or psychological consequences for the stallion due to shuttling, or the economic ramifications in both hemispheres, the end result is more world-wide access to genetics of some superior Thoroughbred stallions that would otherwise not be available.
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Dual Hemisphere Breeding Programs
Regulations governing shipping To determine if a stallion is qualified to shuttle, there is another important item to be taken into consideration when getting the stallion from one country to another. Coordination with the company that is responsible for the actual transport (flying) of the stallion regarding US pre-export regulations for the country of destination can be extensive and time consuming, depending on the country of destination. The United States Department of Agriculture (USDA) and the shipping company must be responsible for enforcing the regulations for preexport quarantine to the satisfaction of the importing country’s veterinary authorities. This precaution minimizes the risk of transmission of infectious diseases from the batch of horses to be exported to the local horse population. It is most important to give these two agencies full cooperation in meeting and carrying out the requirements. Any mistake or non-compliance would put the batch of horses in quarantine at risk for delay of shipment or result in a cancelled shipment. The vaccination and testing schedule is too extensive to cover in this chapter, and the rules can change on a regular basis, depending on the country of destination. This is why it is important to contact the shipping agency and the USDA so that all pre-export requirements are fulfilled; for more information see the website of the USDA’s Animal and Plant Health Inspection Service (APHIS) (http://www.aphis. usda.gov/import export/animals/animal exports. shtml). It is best to contact the shipping agent and the USDA well in advance of any plans to export a stallion so that there is adequate time to prepare an approved Quarantine Area or to arrange shipment to an established pre-approved Quarantine Area. It is important that quarantine personnel be advised of a stallion’s habits, behavior, and his regular routine regarding handling and exercise. This is all very important in enabling the stallion to successfully handle the stress of quarantine and the shuttle process (Bob Davis, Ashford Stud, Versailles, KY, personal communication; Dan Considine, Considine Farm, Lexington, KY, personal communication; Pauline Cushing, International Racehorse Transportation, Chicago, IL, personal communication).
Care of the stallion during shipping Whenever possible, hostile or potentially hostile environmental conditions en route should be avoided. Adding extremes of heat or cold to the challenges inherent in transport is always undesirable.
r r r
r r r r r r
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Use competent staff, i.e., drivers, grooms, and specialist ‘flying grooms’. Ensure adequate ventilation. Avoid, as far as possible, prolonged stationary periods in traffic or at refueling stops. Sometimes, bad weather or equipment failure is the cause of delays that are unavoidable. Provide appropriate tack. Bring sufficient food and water. Have an effective means of restraint. Plan for rest or recovery periods. Check that veterinary help is available if required. Notify the point of arrival of the journey plan and any special requirements (Pauline Cushing, personal communication).
Prior to the actual flight, it is important that the horses are in good condition (good bodyweight, physically fit and healthy) to cope with the stress of flying, depending on the length of flight. Providing good health care for transported horses requires focus prior to travel, during the journey, and after arrival. For convenience, horse movements can reasonably be divided into short-, medium-, and long-haul journeys. In the context of air transport, short can be defined as less than 12 hours; medium as 12–24 hours; and long as greater than 24 hours. These divisions relate to door-to-door time, not just the air travel section. Focusing on the air travel section while ignoring the total journey time can be dangerously misleading. The time needed to transport horses from quarantine to the airport, the unloading and reloading into the jet stalls, weighing each of the stalls with horses, then loading the jet stalls into the aircraft in the proper order for weight distribution can all take a considerable time. This handling time must be incorporated when calculating the total journey time and assessing the potential stress for the horses. Prolongation of an event may result in stress, fatigue, and eventually illness. There is in horses, as in humans, great individual variation in their ability to tolerate travel. Journeys that seem to trouble some horses may cause little demonstrable effect in others. The term ‘transport stress’ has been utilized to describe some of the effects of travel. Stress is a term that is easily understood but is notoriously difficult to define. One of the most useful definitions of stress is that used by Fraser and his research colleagues in 1975.4 They stated that stress occurs when an animal (a horse) is required to make abnormal or extreme adjustments
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in its behavior or internal management (physiology) in order to cope with adverse aspects of its environment and management. This definition allows us to recognize that some adaptations to travel are normal and that we all have a role in optimizing the transport environment and our management of it.4 Preplanning is the key factor for a successful flight. Remember the quote, ‘preplanning prevents piss poor performance’. The amount of water and hay needed from departure at quarantine to the flight’s arrival at destination should be carefully calculated. Hay or perhaps the more dust-free ‘haylage’ should be available on an ad libitum basis thoughout the journey. Water should be offered every 4–6 hours. The addition of electrolytes to drinking water should be avoided as it may depress water intake. There is a belief that it is necessary to ‘pre-load’ horses with fluids and electrolytes prior to travel. This may well be unnecessary unless a horse has a demonstrable history of dehydration. Excessive or uncontrolled administration of electrolytes may actually have adverse effects on water and electrolyte balance. It is prudent to check that a horse that is to be transported has been drinking in accord with its own norm in the days leading up to transport and especially immediately prior to transport (Bob Davis, personal communication; Dan Considine, personal communication; Pauline Cushing, personal communication).4 Adequate blankets and sheets for each horse must be taken on the flight. When flying from the northern to the southern hemisphere the change of season must be considered. Horses leaving the USA in the summer and arriving in the southern hemisphere in the winter may need additional protection against the changes in temperatures over the short period of flying time. This will be the same on the return trip except temperature changes may be more severe depending on what part of the country they will be arriving at on the return flight (Pauline Cushing, personal communication). Bodyweight of the horse is another important factor on long flights. The amount of weight lost can range from 0.45 to 0.55% of total bodyweight (about 2.5 kg in a normal mature Thoroughbred horse) per hour of transport. This weight loss may reflect reduced dietary intake during travel, the evacuation of the contents of the rectum and muscular work associated with maintaining posture during movement, and respiratory vapor loss. It is not unusual for horses to lose 20 kg on an international flight, and horses with ‘shipping fever’ may lose 35 kg or more en route. However, horses that are accustomed to flying may not be affected. Weight lost in transit tends to be regained over the following 3–7 days in healthy horses and perhaps over longer periods in horses with shipping fever. There is thus much to be gained
from weighing horses prior to travel to establish a baseline for comparison with their weight status on arrival and the recovery period from the journey. Of course, there is no guarantee that any scales available for post-arrival weight measurement are identical with those used prior to departure. The bodyweight of the accompanying groom or some heavy piece of equipment or feed that has been transported with the horse should give an indication of the comparability of both sets of scales.4 However, if at all possible, the rechecking of bodyweight should be done in quarantine. The horse’s weight should be closely monitored until the weight lost during transport is regained. Specific health considerations Veterinary supplies such as sedatives, antibiotics, anti-inflammatory drugs, fluids, and other items necessary to cope with in-flight health problems and emergencies must be taken on the flight. Unnecessary medication should always be avoided. Adverse reactions are an ever-present hazard with all therapeutic substances. Tranquilizers should be administered by a veterinarian. Acepromazine, which seems to be widely available worldwide, is contraindicated in intact colts and stallions. Administration of this drug can lead to the possibility of paraphimosis/penile paralysis/priapism. The so-called ‘prophylactic’ use of antibiotics is questionable. Some veterinarians prefer to give antibiotics for 3 days prior to shipment, while others will give medication (antibiotics, anti-inflammatory drugs, and mineral oil) just prior to shipment from quarantine to the airport. This pre-shipment practice may lead to respiratory disease caused by bacteria that are resistant to the antibiotic that was administered in the mistaken belief that it would prevent disease. The use of immunostimulating agents has been avoided in the past because of possible adverse affects, such as depression, fever, and other undesirable signs that can be difficult to differentiate from shipping fever. However, there are some products presently available that are administered orally, and appear to have a beneficial effect either directly or indirectly on the horse’s immune system (e.g., APFTM Advanced Protection Formula; Auburn Laboratories, Inc., Penn Valley, CA). Administration of this product for 10 days or more prior to shipment to international equestrian competitions has been purported to be associated with an increase in water consumption during transit, a more rapid return to pre-shipment weight, and the ability to resume strenuous physical activity more quickly following transport (Mike Van Noy, Auburn Laboratories). Any substances that reduce the immune and inflammatory responses
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Dual Hemisphere Breeding Programs (e.g., non-steroidal anti-inflammatory drugs such as phenylbutazone, corticosteroids, etc.) also are contraindicated prior to shipment. However, these medications may have to be used when fever or colic is encountered during flights. There is a longstanding practice in the horse transport industry of seeking the administration of mineral/laxative oils prior to transport, in addition to feeding a light laxative diet on the night(s) immediately prior to transport. The latter may be more important than the former. Valiant attempts at restraint are necessary in some extreme circumstances. Any depression or injury in horses should be noted and appropriate first-aid action taken wherever possible. During any journey, environmental temperature, relative humidity, and the concentration of contaminants in the environment may change, particularly during travel by air. Monitoring temperature and humidity at regular intervals can be very helpful in deciding whether to cool or perhaps warm the environment in which the horses are being transported. It provides an objective means for doing so, and helps determine the success or failure of attempted improvement. It cannot be emphasized enough that horses travel better in cool conditions than in hot conditions. It must be understood that, when flying, a large number of horses in a closed aircraft can create a great amount of heat, and circulation of air can be critical. Temperatures in the 40s (◦ F; 5–10◦ C) are probably ideal if attainable. When stallions are subjected to higher environmental (cabin) temperatures and/or when the relative humidity is high or when horses are sweating, water should be offered more frequently than the norm. It is important to remember that the inevitable accumulation of feces and urine associated with confinement will lead to a build-up of environmental contaminants. These contaminants will be most deleterious when then plane is stationary. It is for this reason that it is recommended that stationary periods (refueling stops) are kept to a minimum. The removal of feces and urine is clearly impracticable during air travel. Shipping fever is the most common illness found in horses that are being, or have recently been, transported. It is a respiratory infection characterized by signs of depression, inappetance, fever, increased respiratory rate, nasal discharge, and coughing (often a distinctly soft, moist cough) that can rapidly progress to pleurisy and pneumonia. It can occur in as little as 4–6 hours after departure in all journeys. Initial signs of this condition may be seen relatively soon after takeoff in air journeys that have been preceded by long road journeys or by long delays spent in horse vans while awaiting loading onto the aircraft. In any
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shipment of 16 or more horses, it is reasonable to anticipate and provide for one or more instances of this disease complex. Not all cases of shipping fever are apparent during the journey or immediately after arrival. A substantial number of cases will be unapparent until the morning after arrival or days thereafter. On air journeys where a veterinarian is aboard the aircraft, clinical intervention should commence immediately. It has been shown that prompt intervention in this manner reduces the duration of therapy (and thus the severity of the illness) by 50%. Colic and all of the other unpredictable conditions that are inherent hazards of horse keeping can also occur in transit. As with shipping fever, all of the above should be dealt with immediately.4
Care on arrival After arrival at the destination, horses may be visibly excited by the unfamiliar surroundings. After long journeys, this initial excitement may be followed by apparent ‘tiredness’. If observed, fatigue should be monitored carefully. Horses should be allowed some light exercise after arrival, perhaps being led ‘in hand’ or turned out into a small paddock. They should show interest in hay and water within 2 hours of arrival. If they fail to do so, a thorough physical exam should be conducted. Taking of rectal temperatures immediately after arrival and twice daily for at least the first 3 days is essential after all long journeys. Fever (temperature > 100◦ F) or any of the aforementioned signs of shipping fever or any other disease, is an indication for immediate veterinary intervention.4 At least 3 days of recovery should be provided for after air journeys of 12 hours or more for all but exceptional individuals. Allow, where possible, a 7or 8-day recovery period to provide a window of opportunity for treatment of shipping fever, and its resolution.4
Summary In summary, when considering the possibility of shuttling a stallion to breed mares in two hemispheres in back-to-back breeding seasons, one must evaluate the horse carefully. Age, especially with the older horse (16 + years), his inherent fertility, and his breeding behavior are the first things to evaluate seriously. Anyone who is directly or indirectly responsible for setting up pre-quarantine facilities must be aware of all the regulatory requirements to be fulfilled for export to the country of destination. When directly involved, the time, expense, and responsibility must be considered as opposed to sending a stallion to
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a pre-approved facility. In the latter case, the horse must be in good health and body condition, and the quarantine personnel must be made aware of the horse’s behavior and normal habits. Last, but not least, be aware of the number of hours in transport time (from quarantine to the country of destination) so that proper planning can help reduce any possible stress. Some flights from door to door may take up to 70 hours, plus the quarantine, which can add up to a period of 6 weeks during which some horses may be under transport-related stress. This may be very important in determining the amount of time that the horse needs to recover after arrival at its destination (Pauline Cushing, personal communication).
References 1. Pickett BW, Voss JL. Management of shuttle stallions for maximum reproductive efficiency – Part 1. J Equine Vet Sci 1998;18:212–27 2. Turner RM, McDonnell SM. Mounting expectations for Thoroughbred stallions. J Am Vet Med Assoc 2007;230:1458–60. 3. Pickett BW, Voss JL. Management of shuttle stallions for maximum reproductive efficiency – Part 2. J Equine Vet Sci 1998;18:280–7. 4. Fraser D, Ritchie JS, Fraser AF. The term “stress” in a veterinary context. Br Vet J 1975;131:653–62;
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Chapter 116
Immunocastration Tom A.E. Stout and Ben Colenbrander
Introduction Immunocastration involves the use of a vaccine to induce antibodies that suppress reproductive activity and behavior. To date, the only such treatment described for use in the stallion is vaccination against gonadotropin-releasing hormone (GnRH), which results in suppression of the hypothalamic–pituitary–gonadal endocrine axis and, in particular, the luteinizing hormone (LH) arm of the axis. Nevertheless, since the launch of the first commercial GnRH vaccine for horses, EquityTM Oestrus Control, by CSL Animal Health (now part of Pfizer Animal Health) in 2002 in Australia, GnRH vaccination has become a practical proposition in equine practice. At present, EquityTM is registered only for ‘the prevention of estrus-related behavior in mares not subsequently intended for breeding’; however, it has also been tested for various applications in stallions. The most common reason for wanting to immunocastrate a stallion is to induce medium-term, preferably reversible suppression of reproductive or aggressive behavior. However, immunocastration has also been proposed as a means of resolving the equine virus arteritis ‘shedding’ state in breeding stallions, and as a contraceptive in feral horses. This chapter examines the rationale, method of action, and efficacy of GnRH vaccination in the horse, in comparison with other techniques for suppressing reproductive function.
Reasons for immunocastrating stallions The most common reason for wanting to immunocastrate a stallion is to prevent, inhibit, or ameliorate the expression of sexual or aggressive behavior in an animal currently used for purposes other than breeding, but for which a subsequent breeding career is envis-
aged.1, 2 In this context, it is important to bear in mind that not all components of sexual or aggressive behavior are dependent solely on testicular steroid hormones and it is, therefore, not always possible to rid a male horse of unwanted sexual or aggressive behavior either by surgical castration3, 4 or a pharmacological treatment that achieves a similar endocrinological end-point. Other reasons for wanting to suppress reproductive activity in stallions include temporary suppression of testosterone production to encourage elimination of equine arteritis virus from the accessory sex glands of animals that persistently ‘shed’ this pathogen,1 and the induction of infertility in ‘harem’ stallions as a means of contraception and population control in feral horse populations that have outgrown the carrying capacity of a wildlife reserve.5
Inhibition of reproductive steroid-driven behavior The expression of sexual behavior by intact male horses in training or competition is undesirable because it can negatively affect their level of performance and may endanger their rider, handler, or others. Fortunately, most young stallions can be taught not to exhibit sexual behavior in particular circumstances, or in response to specific cues. The potential downside to such ‘negative conditioning’ is that a stallion may be too confused or scared to mount a mare or phantom when he moves on to a breeding career, and may require appropriate remedial treatment to re-establish normal sexual behavior patterns.6 In addition, behavioral training is often more difficult or less effective in older, sexually mature stallions that have previously been used for breeding or are required to combine a breeding career with athletic competitions or shows. If training fails to curb unwanted behavior in an intact colt or stallion, the traditional approach is to
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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remove the source of androgenic steroids by surgical castration. Of course, this is not sensible if the animal may later be required as a breeding stallion, and it also entails surgical and anesthetic risks, particularly in mature stallions. Moreover, surgical castration is not always successful in preventing sexual or aggressive behavior in horses.3, 4 The failure of castration to ‘cure’ retired breeding stallions of behavioral patterns that they have learned by years of repetition is hardly surprising. However, a significant proportion (20–30%) of colts that are gelded before 2 years of age also go on to develop typical stallion-like aggressive and sexual behavior towards other horses, with 5% also becoming aggressive towards people.3 If surgical castration commonly fails to quieten behavior thought to be driven by testosterone, it should be no surprise that pharmacological treatments that aim to inhibit androgen production also sometimes fail to achieve the desired behavioral suppression. On the other hand, immunocastration may be a useful minimally invasive way of assessing the likely efficacy of surgical castration for calming an older stallion.
Suppressing reproductive behavior in stallions using a GnRH vaccine A GnRH vaccine typically comprises GnRH, or a modified form of the peptide, conjugated to a foreign protein that confers antigenicity, and combined with an adjuvant that enhances antibody formation.1, 7 The anti-GnRH antibodies raised are thought to bind to endogenous GnRH within the hypothalamic–pituitary portal vessels, preventing GnRH from binding to the receptors on the pituitary gonadotropes and thereby removing the stimulus to gonadotropin secretion (Figure 116.1). In most mammalian species, the GnRH vaccine-induced suppression particularly affects LH, and the resulting loss of stimulation to the Leydig cells results in a fall in testicular steroid hormone production and consequent reductions in sperm production and quality, and in the expression of sexual behavior.1, 2 Schanbacher and Pratt8 were the first to report successful GnRH immunization in a stallion when they suppressed testosterone secretion from the abdominal testis of a 3-year-old cryptorchid for 4 months using a series of five injections; they concluded that luteinizing hormone releasing hormone (LHRH) (i.e., GnRH) vaccination was a useful alternative to surgical castration for suppressing male behavior in cryptorchids. In the last 10–15 years, a handful of small studies have examined the effects of different GnRH vaccines on reproductive and behavioral parameters in stallions. While the efficacy of vaccination depends
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Figure 116.1 Schematic representations of the endocrinological control of reproductive processes in (a) a normal stallion and (b) following immunocastration by anti-gonadotropinreleasing hormone (GnRH) vaccination. In vaccinated stallions, GnRH antibodies primarily suppress luteinizing hormone (LH) release resulting in a dramatic fall in testosterone secretion by the Leydig cells; in young stallions, falling testosterone concentrations are reliably accompanied by reductions in libido and aggressive behavior. While follicle-stimulating hormone (FSH) secretion is also reduced, some autonomous FSH release continues such that spermatogenesis, while reduced, is not completely suppressed.
on the anti-GnRH antibody titer achieved and, therefore, on the exact composition of the vaccine used, the various studies do highlight some general patterns in responsiveness. In young stallions, vaccination reliably induces a rise in circulating anti-GnRH antibody titers, followed by reductions in testosterone concentrations (Figure 116.2), libido, and sperm production and quality.9–11 When mature stallions are vaccinated against GnRH, however, the clinical response (i.e., testosterone concentrations and behavior) is less reliable, more variable between individuals, and typically requires more booster injections to achieve an effect of relatively short duration. For example, only 50–80% of mature stallions show a GnRH vaccineinduced fall in testosterone concentrations.12–14 There
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(b) Figure 116.2 Graphs of the (a) gonadotropin-releasing hormone (GnRH) antibody and (b) testosterone responses for five 3- to 4year-old stallions immunized against GnRH using the vaccine described by Turkstra et al.11 All five stallions exhibited a rise in GnRH antibody titers, albeit of varying magnitudes, associated with a suppression of testosterone production lasting between 6 weeks and 6 months. Animals with the lowest GnRH antibody titers exhibited the shortest duration of reduced testosterone production.
are also suggestions that the incidence of injection reactions (local swelling and occasional systemic effects such as pyrexia or anaphylaxis) may be higher than in younger animals, although this may be a factor of the larger number of booster injections used,1, 12 and may not apply to all vaccine formulations.14 More significantly, although mature stallions that respond to vaccination exhibit the expected decreases in circulating testosterone concentrations, testicle size, sperm production, and sperm quality, the effect on behavioral
parameters is variable. While Janett et al.14 reported a marked reduction in libido in four of five adult (5–16 year old) stallions, other studies recorded little or no reduction in libido or aggressive behavior in mature animals.12, 13 In a field survey of 22 stallions vaccinated against GnRH for the resolution of behavioral problems, however, owners reported improvement in the majority of stallions presented for inappropriate sexual behavior (17 of 19), aggression (13 of 15), or distraction (seven of seven).15
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Alternative pharmacological treatments for suppressing sexual behavior in stallions Progestagens The major pharmacological alternative to GnRH vaccination for suppressing inappropriate sexual or aggressive behavior in stallions is progestagen administration.16 Indeed, until the advent of GnRH vaccines, the long-term repeated administration of progestagens was the only practical alternative to surgical castration. The major effect of progestagen administration in the stallion is assumed to be suppression of hypothalamic and anterior pituitary secretion of, respectively, GnRH and LH, leading to a fall in testicular steroid production and, as a result, diminished sexual and aggressive behavior.17 However, progestagens also appear to exert a direct ‘calming’ effect at a higher (central nervous system) level,16, 18 which presumably explains why progestagens such as altrenogest R (Regumate ) are also often effective in curbing unwanted sexual or aggressive behavior in geldings.19 Given how frequently progestagens are used on unruly young stallions, there is surprisingly little published data on dose rates, efficacy, and sideeffects. Nevertheless, there is some evidence that high doses of altrenogest (0.088 mg/kg daily) can induce the desired inhibition of both libido and aggressive behavior in young (2–4 years old) stallions.17 On the other hand, Miller et al.20 reported no diminution of sexual behavior in older (> 5 years old) stallions treated daily with 0.044 mg/kg altrenogest for 30 days, suggesting either that older stallions are less responsive, for example because the behavioral patterns have become ‘learned’, or that higher circulating concentrations of progestagen are needed to exert behavioral control. Long-acting progestagen implants or injections have also been anecdotally claimed to suppress sexual behavior in male horses (including delmadinone R ; a product used for treating hyperacetate, Tardak sexuality in dogs). However, effective doses and efficacy based on controlled trials have not been reported. In this respect, the ability of many of these products to achieve adequate serum progestagen concentrations or bind to progesterone receptors in horses is questionable, since a number of them (e.g., medroxyprogesterone acetate, hydroxyprogesterone hexanoate, megesterol acetate, norgestomet) are known to be incapable of rescuing early pregnancy in mares in which luteolysis has been induced.21 The major side-effect of progestagen administration is diminished spermatogenesis and semen quality secondary to reduced testicular steroid produc-
tion.17 In older stallions, this is unlikely to be a significant long-term problem, and Miller et al.20 reported no reduction in sperm production or quality in mature stallions treated with altrenogest for 30 days. Nevertheless, further studies are required to determine whether long-term administration of progestagens to peripubertal colts irreversibly damages spermatogenic potential. On a note of caution, it is also worth remembering that many sporting authorities consider progestagen administration to male horses a ‘doping offence’ because progestagens may be mildly anabolic. In addition, in countries where horses are classified as ‘foodproducing’ animals there may be further regulations that either prohibit the use of progestagens or limit the choice of agents that may be used, unless the horse is officially registered as ‘not for human consumption’. GnRH analogs Both GnRH antagonists and ‘overdoses’ of agonists have been examined for their ability to suppress reproductive behavior in stallions. GnRH antagonists suppress the release of gonadotropins by competitively occupying pituitary GnRH receptors. In stallions, GnRH antagonists have been shown to inhibit secretion of LH and testicular steroids.22, 23 As with GnRH vaccines and progestagens, however, the behavioral response appears to be age dependent, and while a large single dose of antarelix (100 µg/kg) was able to suppress libido in young stallions,22 daily administration of 10 µg/kg antarelix or cetrorelix for 35–37 days did not affect libido in mature stallions, despite a dramatic fall in circulating testosterone concentrations.23 Moreover, GnRH antagonists are currently too expensive to be seriously considered as a means of exerting behavioral control in clinical equine practice. In many species, chronic high-dose administration of a GnRH agonist leads, after an initial hyperstimulation, to decreased gonadotropin release as a result of pituitary desensitization. However, stallions appear to be relatively resistant to these downregulatory effects. Moreover, while Boyle et al.24 did report suppression of gonadotropin secretion, testicular steroid production, and spermatogenesis in mature stallions treated chronically with a GnRH agonist, libido was not affected. High doses of GnRH agonists do not, therefore, appear to be a useful way of suppressing reproductive behavior in stallions.
Elimination of the equine arteritis virus shedder status A second situation in which reducing circulating testosterone concentrations may be clinically useful
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Immunocastration is the stallion that persistently sheds equine arteritis virus (EAV) into its semen. Infection of a stallion with EVA is generally either asymptomatic or results in a transient vasculitis, with typical clinical signs including mild respiratory disease, conjunctivitis, and edema of the head and extremities; in mares, equine virus arteritis (EVA) infection is usually similarly mild. However, EVA is considered a significant threat because infection can cause abortion in pregnant mares or life-threatening illness in foals, particularly in horse populations where EVA is not endemic.25 Moreover, stallions are an important reservoir of infection because, while many completely clear the virus after acute infection, 30–60% become asymptomatic carriers in which the virus localizes in the accessory sex glands and is shed into the semen during ejaculation (see Chapters 121 and 246);26 EAV-naive mares can develop an active infection following insemination with virus-contaminated semen and further disseminate the virus. Many countries therefore insist that stallions are proven non-shedders before they can be registered for breeding, and certainly before their semen is approved for export. While vaccination against EAV should prevent the establishment of the shedder state,25 a licensed EAV vaccine is not available in all countries. Some shedder stallions spontaneously clear EAV from their accessory glands, but others continue to shed for many years26 and are effectively lost as breeding stallions. However, viral persistence in the stallion’s accessory sex glands appears to be testosterone dependent,27 and it has therefore been proposed that reversibly suppressing testosterone production via immunocastration may resolve the shedder state while also allowing subsequent full recovery of fertility.1 Indeed, preliminary clinical tri-
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als of the efficacy of GnRH vaccination for resolving the EAV shedding state are promising. Burger et al.15 reported the cessation of EAV shedding by four stallions within 6 months of a second injection of EquityTM (or the porcine version of the vaccine; ImprovacTM ). Importantly, none of the stallions subsequently resumed virus shedding, and test mating with seronegative mares did not result in seroconversion. The absence of recrudescence is important in the light of a study that demonstrated resumption of virus shedding in three of five stallions apparently cleared of virus shedding by daily treatment with a GnRH antagonist for 35–37 days.23 Equally importantly, Burger et al.15 reported the return to normal fertility of all four stallions cleared of EAV shedding by GnRH vaccination.
Reversibility of GnRH vaccination The major side-effect of using GnRH vaccines for behavioral control or resolution of EAV shedding is a profound suppression of sperm production (Figure 116.3). As for progestagen treatment, the degree to which testicular function recovers after the effects of vaccination have been removed is extremely relevant to potential clinical utility. In 2- to 4-year-old stallions yet to achieve full sexual maturity, spermatogenic capacity has been reported to recover to the level of untreated controls within 4 months of GnRH antibody titers declining.11 On the other hand, one of four mature stallions that responded to the same vaccine used in the above colts, and two of four treated with EquityTM , had failed to recover pre-treatment semen quality within 9 months13 or 12 months14 of the last injection. Given anecdotal reports of young Thoroughbred mares that fail to resume (or develop)
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Figure 116.3 Histological sections (hematoxylin and eosin stained) of testicle biopsies recovered from a 4-year-old pony stallion (a) before and (b) 1 month after vaccination against gonadotropin-releasing hormone. The post-vaccination testicular tissue exhibits clear shrinkage of the seminiferous tubules and disruption of spermatogenesis.
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normal ovarian cyclicity following treatment with EquityTM during their racing careers,28 the question of reversibility remains an important concern. Finally, although GnRH vaccines are not specifically included in lists of banned drugs, and testing for GnRH antibodies is not common, vaccination against GnRH for behavioral modulation does theoretically contravene the regulations of many sporting bodies. It would therefore be helpful if the appropriate authorities would issue clear guidelines on whether or not they consider immunocastration to be acceptable.
Conclusions Immunocastration appears to be a valuable potential tool for suppressing testosterone-dependent behavior (libido and aggression), or resolving EAV shedding, in a way that does not result in a permanent loss of breeding potential. With regard to behavioral suppression, however, it appears that GnRH vaccination is more reliable and effective in young (< 5 years old) than in older animals, both because the latter do not always mount an effective anti-GnRH antibody response and because the behavioral patterns in mature animals may have become ‘hard-wired’. Indeed, in recalcitrant cases, progestagens may have an advantage over immunocastration because they exert an extra calming effect at the level of the central nervous system. On the other hand, early field trials with a commercial GnRH vaccine have generated promising results with regard to both behavioral modulation and EAV shedding. Nevertheless, more widespread uptake in clinical practice would be easier to recommend if further studies were performed to clarify whether spermatogenesis is or is not permanently impaired in some treated animals.
References 1. Stout TAE, Colenbrander B. Suppressing reproductive activity in horses using GnRH vaccines, antagonists or agonists. Anim Reprod Sci 2004;82–83:633–43. 2. Stout TAE. Modulating reproductive activity in stallions: a review. Anim Reprod Sci 2005;89:93–103. 3. Line SW, Hart BL, Sanders L. Effect of prepubertal versus postpubertal castration on sexual and aggressive behaviour in male horses. J Am Vet Med Assoc 1985;186:249–51. 4. Cox JE. Behaviour of the false rig: causes and treatments. Vet Rec 1986;118:353–6. 5. Turner Jr JW, Kirkpatrick JF. Androgens, behaviour and fertility control in feral horses. J Reprod Fertil Suppl 1982;32:79–87. 6. McDonnell S, Murray SC. Bachelor and harem stallion behaviour and endocrinology. Biol Reprod Monogr 1995;1:577–90. 7. Thompson Jr DL. Immunization against GnRH in male species (comparative aspects). Anim Reprod Sci 2000; 60–61: 459–69.
8. Schanbacher BD, Pratt BR. Response of a cryptorchid stallion to vaccination against luteinising hormone releasing hormone. Vet Rec 1985;116:74–5. 9. Dowsett KF, Pattie WA, Knott LM, Jackson AE, Hoskinson RM, Rigby RP, Moss BA. A preliminary study of immunological castration in colts. J Reprod Fertil Suppl 1991;44:183–90. 10. Dowsett KF, Knott LM, Tshewang U, Jackson AE, Bodero DAV, Trigg TE. Suppression of testicular function using two dose rates of a reversible water-soluble gonadotrophin releasing hormone (GnRH) vaccine in colts. Aust Vet J 1996;74:228–35. 11. Turkstra JA, Van Der Meer FJUM, Knaap J, Rottier PJM, Teerds KJ, Colenbrander B, Meloen RH. Effects of GnRH immunization in sexually mature pony stallions. Anim Reprod Sci 2005;86:247–59. 12. Malmgren L, Andresen Ø, Dalin A-M. Effect of GnRH immunisation on hormonal levels, sexual behaviour, semen quality and testicular morphology in mature stallions. Equine Vet J 2001;33:75–83. 13. Clement F, Vidament M, Daels P, van der Meer F, Larry JL, Colenbrander B, Turkstra J. Immunocastration in stallions: Effect on spermatogenesis and behaviour. Anim Reprod Sci 2005;89:230–3. 14. Janett F, Stump R, Burger D, Thun R. Suppression of testicular function and sexual behaviour by vaccination against GnRH (EquityTM ) in the adult stallion. Anim Reprod Sci 2009;115:88–102. 15. Burger D, Janett F, Viadment M, Stump R, Fortier G, Imboden I, Thun R. Immunization against GnRH in adult stallions: Effects on semen characteristics, behaviour and shedding of equine arteritis virus. Anim Reprod Sci 2006;94:107–11. 16. Roberts SJ, Beaver BV. The use of progestagen for aggressive and hypersexual horses. In: Robinson NE (ed.) Current Therapy in Equine Medicine. London: W.B. Saunders, 1987; pp. 129–31. 17. Brady HA, Johnson NN, Whisnant S, Prien SD, Hellman JM. Effects of oral altrenogest on testicular parameters, steroidal profiles and seminal characteristics in young stallions. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 195–196. 18. Perkins NR. Reproductive management. In: Hinchcliff KW, Kaneps AJ, Geor RJ (eds) Equine Sports Medicine and Surgery. Philadelphia: Saunders, 2004; pp. 1193–200. 19. McDonnell S. Stallion behaviour problems. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. London: W.B. Saunders, 2003; pp. 317–19. 20. Miller CD, Varner DD, Blanchard TL, Thompson JA, Johnson L. Effects of altrenogest on behavior and reproductive function of stallions. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 197–198. 21. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32:83–5. 22. Hinojosa AM, Bloeser JR, Thomson SRM, Watson ED. The effect of a GnRH antagonist on endocrine and seminal parameters in stallions. Theriogenology 2001;56:903– 12. 23. Fortier G, Vidament M, DeCraene F, Ferry B, Daels PF. The effect of GnRH antagonist on testosterone secretion, spermatogenesis and viral excretion in EVA-virus excreting stallions. Theriogenology 2002;58:425–27.
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Immunocastration 24. Boyle MS, Skidmore J, Zhang J, Cox JE. The effects of continuous treatment of stallions with high levels of a potent GnRH analogue. J Reprod Fertil Suppl 1991;44:169– 82. 25. Timoney PJ, McCollum WH. Equine viral arteritis: essential facts about the disease. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 199–201.
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26. Timoney PJ, McCollum WH. Equine viral arteritis. Vet Clin North Am Equine Pract 1993;9:295–309. 27. McCollum WH, Little TV, Timoney PJ, Swerczek TW. Resistance of castrated male horses to attempted establishment of the carrier state with equine arteritis virus. J Comp Pathol 1994;111:383–8. 28. Robinson SJ, McKinnon AO. Prolonged ovarian inactivity in broodmares temporally associated with administration of Equity. Aust Equine Vet 2006;25:85–7.
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Chapter 117
Management of Stallions in Artificial Insemination H. Steve Conboy
Artificial insemination (AI), the man-made practice of placing semen into the female reproductive tract for the establishment of pregnancy, is a technique that has been used for many years. Its first use was reported in 1322 in Arab horses1 and many references of its development have been reported since that time.2 Early widespread use of AI in the USA was limited by many breed registries, and the technique is still not sanctioned by the Jockey Club. The most common reasons that the technique has been criticized are the perception that the offspring would be inferior to those conceived by actual copulation and because it creates an increased opportunity for indiscriminate and fraudulent mating practices. Many breeders feared that its use would more quickly express genetic problems within the breed by allowing stallions with unknown undesirable genetic traits or mutant genes to dominate the breed. This factor has, in fact, been a bona fide issue with several breeds in which a genetic defect has been rapidly disseminated through the extensive use of certain popular stallions. Examples of this are the hyperkalemic periodic paralysis (HYPP) and hereditary equine regional dermal asthenia (HERDA) syndromes in the American Quarter Horse.3 Other genetic defects that have this potential are the overo lethal white syndrome in the American Paint Horse4 and the equine epitheliogenesis imperfecta syndrome in the American Saddlebred.5 Within the horse industry, the argument of fraud has been significantly diminished by the ability to verify parentage with DNA testing. There is still the reservation that breeding fees might be devalued as well as the value of the resulting offspring with the ability to impregnate so many more mares by dividing each ejaculate. This argument has, in part, been neutralized by the current large increases
in numbers of mares bred by natural service (in many cases 200 plus mares in a season), through the use of ovulation-inducing drugs and other management practices, without reducing the breeding fees. There is, however, evidence that the greater number of offspring produced in a season has resulted in the market selecting the ‘top of the crop’ and devaluing the remainder. When all factors are considered, AI does have many significant advantages over natural cover, but, unfortunately, several of these advantages may be considered disadvantages by some: 1. It is safer for both the breeding personnel and the stallion. 2. It can reduce or prevent the spread of contagious and infectious diseases. 3. It affords one the opportunity to examine a representative sample of the ejaculate. 4. It facilities multiple impregnations from the same ejaculate for fertile stallions. 5. It provides the opportunity to extend the ejaculate with antibiotics and nutrients to extend the longevity of the spermatozoa. 6. It makes it possible to transport the semen over long distances prior to insemination. 7. It allows scheduling collection and storage of semen and insemination at a more convenient or appropriate time. 8. It eliminates the necessity of having the mare brought to the stallion for breeding. 9. It allows a stallion that has lameness or other physical issues to be used for breeding that may not be possible by natural cover. 10. Frozen semen can be used, which has additional advantages: (a) semen can be stored and used at any time, even after the stallion is deceased
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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(b) there are no time constraints on transporting semen (c) semen can be used while the stallion continues with his performance career (d) semen can be stored until a horse has been proven as a performance horse or a sire (e) distribution of semen for insemination can be used on a worldwide scale. There are also some notable disadvantages to the use of artificial insemination: 1. Special equipment is needed. 2. Skills and knowledge of the techniques are essential. 3. The stallion must be trained to ejaculate into an artificial vagina. 4. Fertility may be reduced with cool-transported semen from some stallions, and is usually reduced with frozen–thawed semen. 5. The additional cost for collection, processing and transporting can be significant.
Figure 117.2 Semen shipping and storage containers.
Equipment The basic equipment for collecting semen from the stallion for AI can be as simple as an artificial vagina, insemination pipette and syringe. Most breeding operations that utilize this technique in their breeding programs, however, maintain a laboratory stocked with the following basic equipment (Figures 117.1 and 117.2): 1. Artificial vagina (AV). 2. Binocular microscope (phase contrast and heated mechanical stage are ideal). 3. Instrument for measuring sperm concentration. (a) hemacytometer
4. 5. 6. 7. 8.
9. 10. 11. 12.
(b) computerized spermatozoal motility analyzer (which can also measure sperm concentration). (c) photometric counter (d) fluorescence-based sperm counter. Small incubator for maintaining glassware, plasticware, and extender at a warm temperature. Refrigerator. Insemination pipettes and plastic syringes. Specimen cups. A balance or small set of scales. Those veterinarians or breeders that plan to transport fresh or frozen semen will also need the following: Centrifuge that can accommodate 50-mL capacity tubes. Commercially available semen transport devices. Liquid nitrogen semen storage tank. Vapor nitrogen shipper.
While a mare in estrus can be used for the stallion to mount for semen collection, a ‘phantom mare’ is safer and more convenient. When a phantom mare is used for semen collection, it is usually necessary to provide an estrous mare in close proximity to the phantom to sexually stimulate the stallion. The mare should be typically stationed in a holding chute in front of the phantom (Figure 117.3).
Semen collection and processing
Figure 117.1 Semen laboratory equipment.
To collect semen, the stallion is trained to ejaculate into an artificial vagina (AV) that is fitted with a clean collection vessel (Figure 117.4). The collection vessel can be fitted with an internal filtration device or the semen can be filtered into a specimen cup through
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Figure 117.3 Mare in teasing chute in front of phantom.
several layers of sterile gauze. The artificial vagina must be heated to the proper temperature (approximately 115◦ F or 46◦ C) and adjusted to the proper pressure with hot water. Some stallions prefer that the temperature be hotter or colder and will not ejaculate readily unless the temperature is to their satisfaction. If the stallion fails to ejaculate after several attempts of entering the AV, the temperature and or pressure should be adjusted to determine his preference. When AV temperature or pressure is the reason for the failure to ejaculate, it is usually because the AV is too cold or too tight. Prior to collecting the semen, the AV should be lightly lubricated with a non-spermicidal lubricant. Caution should be taken to avoid excessive lubricant as it may be forced into the collection vessel, which may adversely affect the quality of the ejaculate. The collection vessel should be protected from temperature extremes and light, and the collected semen should be transported to the semen laboratory immediately after collection for further processing. In some cases, it may be beneficial for the stallion to ejaculate directly into warm extender. An ejaculate collected by this method can then be centrifuged if necessary to achieve a more desirable concentration.
Figure 117.4 Missouri artificial vagina with collection bottle.
If an in-line filter is not used during semen collection, the semen is filtered to remove debris and gel. The amount of gel will vary with the stallion, the amount of arousal (teasing) before collection, and the time of the year. The gel can be removed by filtering through gauze, by decanting from the collection vessel, or by aspiration with a syringe. The consistency of gel is similar to egg white in that it remains clumped together in the ejaculate. When decanting, as the vessel is tilted the gel will move to the top and separate itself from the remainder of the semen. To aspirate with a syringe, it is selectively drawn from the ejaculate by placing the tip of the syringe against the gel while aspirating. The gel will flow into the syringe until it is completely removed from the ejaculate. One should be aware that sperm losses are increased when this technique is used, as opposed to insertion of an in-line filter to separate gel from gel-free semen during the semen collection process. The extent of processing the semen is determined by the way it will be used for insemination. If a single mare is to be inseminated with the ejaculate, the semen can be simply filtered to remove particle debris and seminal gel and then inseminated directly into the mare without further processing. If semen extender is to be added, it is done following filtration and gel removal. A gel-free semen sample should be withdrawn for counting prior to initial extending at a ratio of 1:1. If the ejaculate is to be divided among several mares, it may be necessary to determine the sperm concentration, the total number of sperm in the ejaculate, and the viability before proceeding with the addition of more extender (see Counting semen below). If the semen is to be cooled for storage or for shipping, the extending process becomes more critical and centrifugation (see Centrifugation of semen below) may become necessary. A variety of semen extenders are commercially available, but several formulas can be easily prepared in the semen laboratory. The most common extenders are milk based and use non-fat dry milk solids (NFDMS) that are found in the grocery store (Table 117.1). They also contain an energy source, such as
Table 117.1 Example of a typical Non-fat Dry Milk Solids Glucose Extender (adapted from Kenny et al.6 ) Component
Amount
NFDMS Glucose Amikacin sulfate Sodium bicarbonate (NaHCO3 ) 8.4% Sterile deionized water
2.4 g 4.9 g 100 mg 2 mL 92 mL
NFDMS, non-fat dry milk solids.
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Management of Stallions in Artificial Insemination glucose, and antibiotics to prevent bacterial growth. The most commonly used antibiotics are gentamicin sulfate, amikacin sulfate, ticarcillin (ticarcillin containing clavulonic acid), and crystalline penicillin. When gentamicin or amikacin is used, a buffer (8.4% sodium bicarbonate) should be included for pH adjustment. Experience will determine which antibiotic performs best for a particular stallion’s semen. The diluent for the formula should be sterile deionized water (never saline). The osmolality of milk-based extenders should be between 300 and 400 mOsm (350 mOsm is ideal) and the pH should be 6.7–7.2.7 Filtered semen should be extended at a ratio of 1:1 or greater, depending on the semen concentration, as soon as the semen has been filtered. If the semen is to be cooled for storage or for shipping, a more precise dilution is required. It has been demonstrated that a sperm concentration of 25–50 × 106 maximizes spermatozoal survivability in cooled extenders.8
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and concentration (no less than 25 × 106 sperm/mL).8 When shipping cooled semen, a 50% attrition in PMS should be anticipated, and at least 1.0 × 109 PMS is generally sent for each insemination dose (S. Conboy, personal observation). A recent study13 demonstrated a deterioration of semen quality (based upon motility, velocity, and chromatin integrity) on all control samples stored both at room temperature (22–30◦ C) and at 5◦ C for up to 48 hours. This study compared the use of storage with a semen extender (Kenny’s) to a colloidal centrifugation process followed by dilution in Kenny’s extender.14 The authors reported that, when semen was centrifuged over the colloid and resuspended in Kenny’s extender, sperm quality was maintained for 24 hours at room temperature (22–30◦ C) or for at least 48 hours when cooled to 4◦ C. This technique suggests an alternative for those stallions whose semen does not tolerate cool storage.
Enumeration of sperm Volume and dosage The total volume of extended semen to be inseminated can be quite variable, depending upon the final concentration of extended semen. Some practitioners argue that large volumes of extended semen are undesirable as they tend to escape from the uterus through the open cervix. It has been demonstrated, however, that similar conception rates resulted in pony mares that received 30 or 120 mL of extended semen with a common sperm concentration of 50 × 106 .9 This would suggest that a large volume of semen does not adversely affect fertility, as long as the concentration is sufficient to provide the necessary number of viable sperm. Certainly, insemination with 250 × 106 progressively motile sperm (PMS) has been shown to produce a higher pregnancy rate when the insemination volume was 10 mL (i.e., sperm concentration of 25 × 106 /mL), as opposed to 100 mL (i.e., sperm concentration of 2.5 × 106 /mL).10 Studies have shown that good conception rates can be achieved with as few as 100 × 106 PMS but a dose of 500 × 106 PMS is accepted as the standard.11, 12 Breeding trials will dictate the minimum sperm dose that will optimize pregnancy rates for a particular stallion, as individual fertility is quite variable. If the extended semen will be used for immediate insemination, a minimum extended ratio of 1:1 is used. When shipping semen, a ratio greater than 1:1 is needed to preserve the quality of the semen. If the concentration of the filtered ejaculate intended to be extended for shipping is so low that it will cause the extended semen concentration to drop below 25 × 106 sperm/mL, it should first be concentrated by centrifugation and then extended to the desired ratio
An accurate measurement of the sperm concentration in an ejaculate is needed to accurately calculate the insemination dose. This is especially important when impregnating a relatively large number of mares with an ejaculate or when transporting semen. The gelfree volume must be recorded to determine the total number of sperm in the ejaculate. The concentration and total number of sperm can be derived using a variety of methods. The most accurate methods utilize either a hemacytometer (Table 117.2), which is inexpensive but time-consuming, or a photometric-based counter,15 a fluorescence-based sperm counter,16 or computerized spermatozoal motility analyzer. The fluorescence-based and computerized systems are expensive but provide instant and generally accurate results. There are numerous photometric instruments that are marketed to measure semen concentration. They give instant results, but can be inaccurate because they will read debris, blood cells, and other particulate matter in semen. They are limited to measuring gel-free semen that is not extended.
Centrifugation of semen Removal of the seminal plasma and replacing it with a semen extender is known to improve both motility and longevity of stored semen. Semen with extremely low sperm concentration and semen that fails to maintain good quality when cooled for storage or shipping may benefit from centrifugation to remove the seminal plasma followed by resuspension of the sperm pellet in warm (37◦ C) semen extender. A centrifuge with a swinging head and capacity for 50-mL
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Table 117.2 Using the hemacytometer, Spencer Bright-Line or Neubauer counting chamber to count sperm Diluent for counting semen in hemacytometer: Sodium bicarbonate (baking soda) 5 g Formalin 1 mL Distilled water q.s. 100 mL (alternatively, distilled water alone can be used for this purpose) Method 1: a. Diluent 3.9 mL to 0.1 mL gel-free or extended semen with a pipette b. Discharge several drops to clear the pipette of unmixed solution c. Mix and fill hemocytometer chamber d. Let stand for 10 minutes (prevent from drying by placing in refrigerator or humidified environment) e. Under 200–400× microscopic magnification, count five squares with in the center square f. Count only sperm heads g. Total of five squares × 2 × 106 = concentration h. Concentration × volume = total sperm number Method 2: red cell pipette a. Fill red cell pipette to 1.0 mark with gel-free semen or extended semen b. Dilute to 1.01 mark and mix c. Discharge several drops to clear the pipette of unmixed solution d. Fill counting chambers as for blood and let stand in refrigerator or humidified environment for 10 minutes e. Count five small squares in center large square (heads only) f. Five squares × 5 × 106 = concentration g. Concentration × volume = total sperm number Method 3: white blood cell (WBC) pipette for more dilute samples a. Fill WBC pipette to the 0.5 mark with gel-free semen or extended semen and fill to the 1.0 mark with diluent and mix b. Discharge several drops to clear the pipette of unmixed solution c. Fill chambers as for blood and let stand in refrigerator for 10 minutes d. Count the four large squares e. Four squares × 50 × 103 = concentration f. Concentration × volume = total sperm number Calculating the dose of semen Total sperm in dose (TS) Concentration of semen (C) Volume of dose (V) TS = C × V Example
Figure 117.5 Dynac II centrifuge and 50 mL tubes.
amount of damage. The following technique has been found to achieve this goal: 1. Dilute the gel-free semen with warn (37◦ C) milkbased extender 1:1 or greater. 2. Divide and balance in 50-mL centrifuge tubes (greater recovery rates have resulted when the column height is minimal; therefore, filling the 50-mL tubes to 20 mL will give the greatest sperm yield).18 3. Centrifuge at 400–500 × g for 10 minutes. 4. After centrifugation, carefully aspirate the supernatant and discard. Leave about 5 mL of extender in the centrifuge tube. 5. Resuspend the soft pellet in 10 mL of warm extender by gently swirling the tube. 6. After suspending the pellets the tubes can be combined. 7. The total concentration can be established by counting the sample with a hemacytometer, or fluorescence-based sperm counter (a 75% recovery rate is generally achieved by this technique and may be used as an estimate of the yield).17 8. After calculating the concentration the semen can be further extended to the desired concentration.
50 × 106 × 40 mL = 2.0 × 109 total sperm V = TS/C Example 50 × 106 × 40 mL = 2.0 × 109 total sperm 500 × 106 (total sperm desired) = 10 mLs (Volume of A1 dose) 50 × 106 (concentration of semen)
If the original ejaculate contained 6.00 × 109 sperm and the recovery was 75%, the new volume would contain 6.00 × 109 × 0.75 = 4.5 × 109 sperm. To determine the concentration divide, the total number of sperm by the volume: Yield = 75% × original number of sperm Concentration = yield/new volume
tubes is desirable for centrifuging large volumes of semen (Figure 117.5).17 When centrifuging stallion semen, the goal is to recover the maximum number of sperm with the least
The following formula can be used to calculate the gravitational force (g) when the radius (r) of the centrifuge arm in centimeters is known and the speed
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Management of Stallions in Artificial Insemination of the centrifuge in revolutions per minute (r.p.m.) is known: g = 11.18 × r × r.p.m.2 × 10−6 Example : g = 11.18 × 10 cm × 20002 × 10−6 g = 447
Using a ‘cushion’ with centrifugation This technique allows more vigorous and prolonged centrifugation (400–1000g for 20–25 minutes) with less sperm damage and results in recovery rates that approach 100%. A colloid medium (‘cushion’) is added to the centrifuge tube (3–4 mL) and the extended semen is then layered over the medium. After centrifugation a soft sperm pellet is concentrated between the cushion medium and the extender. The cushion is carefully aspirated by passing a pipette through the supernatant and past the sperm pellet. The supernatant is then removed and the sperm pellet is reconstituted in warm extender to the desired concentration.19, 20
Selection of mares for artificial insemination While one of the advantages of using AI is to reduce contamination and potential inflammation of the mare’s uterus during breeding, it remains important to evaluate and to attempt to correct any pathological conditions of the mare’s reproductive tract before proceeding with insemination. Pregnancy rates are significantly reduced when free uterine fluid and urovagina are not corrected prior to insemination. Mares that have pathological conditions frequently become pregnant only to abort later in gestation. Correcting urine pooling (urovagina) may be as easy as increasing the mare’s body score. If weight gain fails to eliminate urine pooling a urethral extension should be considered (see Chapter 263). Acupuncture appears to be helpful in some cases (S. Conboy, personal observations). Fluid accumulation in the uterus prior to insemination is extremely detrimental to establishment and maintenance of pregnancy and must, therefore, be corrected. Uterine fluid is diagnosed by ultrasound evaluation. The underlying cause must be identified followed by the appropriate treatment. The nature of the fluid must first be determined. It may be urine that results from urovagina and an open cervix, an exudate that results from a uterine infection, or a transudate that is related to uterine inflammation. Pluriparous mares with stretched uterine ligaments and older maiden mares whose cervices do not
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open during estrus are very susceptible to poor uterine clearance. When uterine fluid exceeds 1 cm in cross-sectional diameter during estrus, the mare should be treated with oxytocin prior to breeding and levels greater than 2 cm should be treated with drainage and appropriate diagnostic tests and treatments before breeding.21 Abnormal conformation of the vulva that contributes to vaginal or uterine fecal contamination or aspiration of air should be surgically corrected with a Caslick’s procedure. Abnormalities of the cervix such as adhesions, lacerations, and failure to dilate are frequently discovered during insemination. These conditions of the cervix must be corrected before a successful pregnancy can occur. Uterine infection diagnosed by cytology and culture should be treated with the indicated antibiotic and lavage.
Insemination technique The semen should be drawn up into a non-toxic syringe (typically one with a plastic-tipped plunger as opposed to one with a rubber-tipped plunger). The syringe and semen can be kept warm or from overheating by placing it in a suitable container or by wrapping it in several layers of paper towels while transporting it (locally) to the mare (Figure 117.6). The mare should be properly restrained in breeding stocks or in a stall doorway. The tail can be wrapped to keep the hair and dirt away from the perineal area or it can be skillfully held over the back. The vulva and adjacent perineal area are then scrubbed with warm water and soap. Good hygiene dictates that disposable gloves be worn and a disposable material, such as cotton or paper towels, be used for the scrub (Figure 117.7). Sponges or other non-disposable materials should not be used as they have the potential for spreading harmful bacteria and viruses. Special attention should be given to the clitoral fossa.
Figure 117.6 Artificial insemination materials.
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Figure 117.9 Clean cotton wipe after proper washing.
Figure 117.7 Washing mare’s vulva with cotton.
Care must be taken to insure that no soap or debris is left inside the vestibule (Figure 117.8). When the scrub is complete, a piece of clean moist cotton used to swipe over the vulva should remain clean (Figure 117.9). The inseminating hand should be gloved in a clean or sterile obstetrical sleeve and a sterile insemination pipette is then opened and grasped between the thumb and index finger. A small amount of sterile non-spermicidal obstetrical lubricant is placed on the knuckles (not the index finger). The fingers form a cone to protect the end of the pipette and to allow smooth atraumatic entry into the vestibule and then the vagina (Figure 117.10). The labia of the vulva can be parted slightly with the opposite hand or by an assistant to facilitate smooth entry. In the case of maiden mares, it may be necessary to slowly pen-
Figure 117.8 Cotton removing soap and debris from vestibule.
etrate an intact hymen. This is done by pressing the index finger against it until a small puncture results. The entire hand is then slowly forced into the vagina as the hymen stretches. When the hand is in the vagina the index finger is used to search for the cervix, which should be quite flaccid in the estrous mare. The maiden mare may be an exception, in which case the cervix is often semi-erect and will usually only accommodate one or two fingers at the external os. Mares that have had an episioplasty (Caslick’s surgery) may not have sufficient opening to allow the hand and arm to pass into the vagina. To deal with this situation, the vulva must either be opened (episiotomy) or the mare can be inseminated through a sterile vaginal speculum. When a speculum is used, the pipette is guided to the os of the cervix using a pen light. When the cervix is located either by the finger or visually the pipette is carefully passed into the uterine body. This may be difficult when using the speculum, as the tip of the pipette
Figure 117.10 Gloved hand cupping sterile pipette with lubricant on knuckles.
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Management of Stallions in Artificial Insemination often becomes entrapped in the flaccid cervix. Creating a bend in the end of a plastic pipette will often enable it to be manipulated into the body of the uterus. Another more difficult but useful technique is to manipulate the AI pipette through the cervix per rectum. If the hand can be entered into the vagina, it is helpful to compress the flaccid cervix around the pipette that has been placed in the body of the uterus to prevent excessive semen from draining immediately from the uterus as it is gently injected from the syringe. When very low volumes of semen are used, as when using frozen semen, it is desirable to perform a high tubular insemination. This is done by using a longer (76 cm) flexible pipettethat allows the pipette to be threaded up to the tip of the uterine horn in the direction of the dominant follicle. This can be effectively aided by manipulation with a hand per rectum. Also when inseminating with low volumes of semen it is necessary to deliver the entire dose from the pipette into the mare’s uterus. To insure that the complete dose is delivered, the semen should be drawn first into the pipette and any excess semen allowed to enter the syringe. A small volume of air forced into the pipette via the syringe will aid in pushing all of the semen from the pipette. The use of a small bore pipette also facilitates maximum transfer of the semen dose. A flexible pipette with a ¨ small bulb on the tip has been developed (Minitub, Tiefenbach, Germany) that can be felt per rectum as it passes through the uterine lumen. It features an internal stylet that will discharge a 0.5-mL semen straw and then retrieve the straw from the lumen of the pipette if additional straws are needed. Hysteroscope-guided insemination has been used to visually deposit low doses of semen directly at the papilla of the oviduct. This technique requires expensive equipment and multiple assistants, and the results are not reported to be better than the deep uterine procedure.22
Timing and frequency of insemination When using fresh semen, pregnancy rates similar to, or better than, natural cover should be expected. The ideal time of insemination is 12–24 hours prior to ovulation.23 Insemination 12–24 hours prior to ovulation allows time for sperm transport to the oviduct and capacitation before the egg enters the oviduct. The mare should generally be inseminated every 48 hours until ovulation occurs. Mares that are inseminated only after ovulation has occurred are known to have a significantly higher incidence of early embryonic loss.23–25 Stallions with exceptional semen quality often achieve acceptable pregnancy rates with longer intervals between inseminations. When good
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quality cooled transported semen is used within 24 hours of collection, pregnancy rates should be similar to those with fresh semen. Most cooled stored semen will begin to lose its quality after 24 hours of storage; however, pregnancies have been obtained from semen that had been stored for 3–4 days. Whenever possible, a sample of shipped semen should be retained to evaluate sperm viability when it will be stored for a period greater than 24 hours prior to insemination. Mares that are inseminated with cooled semen that do not ovulate within 24–48 hours after insemination should receive a second dose if possible. The need for this practice, however, is dependent on the intrinsic fertility of the stallion being used. When using frozen semen, the proximity to ovulation is more critical than with fresh or even fresh cooled semen. There are varying opinions on how to manage mares being bred with frozen semen. It is generally accepted in the USA that one dose of frozen semen should precede ovulation by no more than 12 hours and, if available, a second dose should be given no later than 6 hours post ovulation.26 The necessity for inseminating so close to ovulation is thought to be related to freeze–thaw damage to the sperm cells that reduces their survival in the oviduct. In Europe, where frozen semen has been used more extensively, there seems to be less concern of timing insemination closer than within 24 hours prior to ovulation. A timed insemination approach using two inseminations can be used to reduce the number of examinations required for breeding with frozen semen. The first dose is inseminated 24 hours after human chorionic gonadotropin (hCG) or deslorelin is administered. The second dose is inseminated 12–16 hours later. These two doses will generally cover the window required for optimal pregnancy rates (i.e., 12 hours before ovulation to 6 hours post ovulation).27 Regardless of whether using fresh or frozen semen, it is very helpful to use a routine schedule of palpation and ultrasound evaluation of the ovaries and the uterus to facilitate predicting ovulation. The use of ovulation-inducing medications is also advisable. This is especially important when using shipped or frozen semen that is limited in availability or supply. Those medications that are customarily used are hCG and the gonadotropin-releasing hormone (GnRH) agonist deslorelin. Either of these drugs offers predictable ovulation approximately 85% of the time. When administered to a mare with a mature follicle (35 mm or greater diameter) on the 3rd or 4th day of estrus, the treatment will usually result in ovulation in 24–48 hours with hCG28, 29 and in 36–48 hours with deslorelin.30
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Post-ovulation evaluation Following insemination, it is important to document ovulation by ultrasound (it is often difficult to distinguish a corpus hemorrhagicum from a follicle via palpation) and to evaluate the uterus for signs of failure to eliminate inflammatory insemination-induced fluids. It is normal for the uterus to respond with inflammatory fluids following insemination, whether it is with fresh, cooled, or frozen semen; however, the mare should evacuate this fluid from the uterus before the cervix closes as the mare enters diestrus. Failure to eliminate inflammatory fluids from the uterus is a phenomenon that is more common in older mares and in older maiden mares whose cervices do not dilate completely during estrus. Oxytocin is frequently given to susceptible mares 4–6 hours after insemination to aid in uterine clearance. Mares that have a history of chronic endometritis should be infused with antibiotics or lavaged with isotonic fluids following insemination and if possible limited to one insemination per estrus. If additional inseminations are required, treatment after each insemination is recommended. If 1–2 cm of fluid (cross-section diameter) remains in the uterus after ovulation it is common to treat the uterus with oxytocin. If greater than 2 cm of fluid remains after ovulation, lavage with isotonic fluids immediately prior to oxytocin administration is recommended. The addition of antibiotics to the lavage fluids may be indicated if the mare has a predisposition for uterine infection. It is safe to continue to lavage or infuse the uterus for up to 2 days post ovulation. After that period there is a danger of interfering with the development of the corpus luteum.21
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
References 1. Perry EJ. Historical. In: Perry EJ (ed.) The Artificial Insemination of Farm Animals. New Brunswick: Rutgers Press, 1945; pp. 3–4. 2. Brinsko SP, Varner DD. Artificial insemination. In: McKinnon A, Voss J (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 790–1. 3. Rudolf JA, Spiers SJ, Byrns G, Rojas CV, Bernoco D, Hoffman EP. Linkage of hyperkalemic periodic paralysis in Quarter Horses: a sodium channel mutation disseminated by selective breeding. Nature Genet 1992;2:114–47. 4. Vrotsos PD, Santchi EM, Michelson JR. The impact of mutant causing overo lethal white syndrome on white patterning horses. In: Proceedings of 47th Annual Convention of the American Association of Equine Practitioners, 2001, pp. 385–91. 5. Lieto LD, Swerczek TW, Cothran EG. Equine epitheliogenesis imperfecta in two American Saddlebred foals is a lamina lucida defect. Vet Pathol 2002;39:576–80. 6. Kenny RM, Bergman RV, Cooper WL, Mores GW. Minimal contamination technique and preliminary findings. In:
18.
19.
20.
21.
22.
23.
Proceedings of 21st Annual Convention of the American Association of Equine Practitioners, 1975, pp. 327–35. Varner DD. Composition of seminal extenders and its effect on motility of equine spermatozoa. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1991, pp. 146–50. Varner DD, Blanchard TL, Love CC, Garcia MC, Kenney RM. Effects of semen fractionation and dilution on equine spermatozoal motility parameters. Theriogenology 1987;28:709–24. Bedford SJ, Hinricks K. The effect of insemination volume on pregnancy rates of pony mares. Theriogenology 1994;42:571–8. Rowley HS, Squires EL, Pickett BW. Effect of insemination volume on embryo recovery rate in mares. J Equine Vet Sci 1990;4:298–300. Householder DD, Pickett BW, Voss JL, Olar TT. Effect of extender, number of spermatozoa and hcg on equine fertility. Equine Vet Sci 1981;1:9–11. Brinsko SP, Varner, DD, Blanchard TL. Cooled shipped semen-the two sides of the coin. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1999, pp. 91–107. Morrell JM, Johannisson A, Strutz H, Dalin A, RodriguezMartinez H. Colloidal centrifugation of stallion semen: changes in sperm motility, velocity, and chromatin integrity during storage. J Equine Vet Sci 2009;29:24–32. Morrell JM, Johannisson A, Dalin A, Hammar L, Sandebert T, Rodriguez-Martinex H. Sperm morphology and chromatin integrity in Swedish warmblood stallions and their relationship to pregnancy rates. Acta Vet Scand 2008;50:2. Pickett BW, Squires EL, McKinnon AO. Procedures for collection, evaluation, and utilization of stallion semen for artificial insemination. Colorado State University Bulletin No. 03, 1987, pp. 24–6. Comerford KL, Love CC, Brinsko SP, Emond AJ, White JA, Teague SR, Varner DD. Validation of a commercially available fluorescence-based instrument to evaluate stallion spermatozoal concentration. In: Proceedings of 54th Annual Convention of the American Association of Equine Practitioners, 2008, pp. 367–8. Vanderwall DK. How to process a dilute ejaculate of semen for cooled-transported insemination. In: Proceedings of 54th Annual Convention of the American Association of Equine Practitioners, 2008, pp. 369–73. Ferrer MS, Paccamonti DL, Dilts BE, Lyle SK, Aljarrah AH, Devireddy R. Improvement of sperm recovery rates after centrifugation of stallion semen. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 2004, p. 7. Loomis PR. Advanced methods for handling and preparation of stallion semen. Vet Clin North Am Equine Pract 2006;22:663–76 Waite JA, Mancill SS, Love CC, Brinsko SP, Teague SR, Salazar JJ, Varner DD. Factors impacting equine sperm recovery rate and quality following cushion centrifugation. Theriogenology 2008;70:704–14. Pycock JF. Therapy for mares with uterine fluid. In: Samper J, Pycock J, McKinnon A (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders, 2007; pp. 93–104. Matthews P, Morris L. Low-dose insemination techniques. In: Samper J, Pycock J, McKinnon A (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders, 2007; pp. 315–318. Woods J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic loss rate in mares. Equine Vet J 1990;22:410–15.
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Management of Stallions in Artificial Insemination 24. Koskinen E, Lindeberg H, Kuntsi H, et al. Fertility of mares after postovulation insemination. Zentralbl Veterinarmed A 1990;37:77–80. 25. Barbacini S, Gulden P, Marchi V, Zavaglia G. Incidence of embryonic loss in mares inseminated before or after ovulation. Equine Vet Educ 1999;11:251–4. 26. Pickett BW, Amann RP. Cryopreservation of semen. In: McKinnon A, Voss J (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 769–97. 27. Samper JC, Estrada AJ, McKinnon A. Insemination with frozen semen. In: Samper J, Pycock J, McKinnon A (eds)
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Current Therapy in Equine Reproduction. St. Louis: Saunders, 2007; pp. 285–8. 28. Ducamp G, Bour B, Combarnous Y, Palmer E. Alternative solutions to hcg induction of ovulation in the mare. Reprod Fertil Suppl 1987;35:221–8. 29. Davdison DF. The control of ovulation in the mare with reference to insemination of stored sperm. J Agric Sci 1947;37:287–90. 30. Blanchard TL, Brinsko SR, Rigby SL. Effects of deslorelin and hCG administration on reproductive performance. J Equine Vet Sci 2001;21:539–42.
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Chapter 118
Management of Stallions in Natural-Service Programs Norman W. Umphenour, Phillip McCarthy and Terry L. Blanchard
The overall goal of stallion management is to keep the breeding stallion content with respect to housing, management, breeding, exercise, and feeding. When this goal is achieved, the potential for the horse to be a joy to work with in the breeding shed is much greater. Overall management will affect libido, which is particularly important from the standpoint of intensive breeding schedules. Time is the critical factor limiting efficient operation of large breeding farms or stallion stations. Some stallions are required to breed three to five times daily for extended periods during seasonal peaks of breeding activity (see Table 118.1). Stallions that become slow to breed can delay breeding schedules throughout each day, reducing management efficiency and resulting in exhaustion of and frustration for breeding shed personnel. In some instances, breeding may have to be missed, thus reducing overall reproductive efficiency. Therefore, any management practice that will maintain excellent libido should be instituted in an effort to increase mating efficiency.
Selection of stallions for breeding Selection of most stallions for breeding is based primarily on their performance, conformation, and pedigree; virtually no consideration is given to reproductive potential. Consequently, the stallion manager’s task is to maximize the reproductive efficiency of each given stallion regardless of that horse’s inherent fertility. Veterinary expertise has reached a level of competence wherein owners may feel comfortable purchasing stallions with known reproductive problems. For example, some stallions that are unilateral cryptorchids or that otherwise ejaculate low numbers of normal motile sperm are purchased, syn-
dicated, and placed in service. Veterinarians are expected to manage such stallions in a manner to maximize the number of foals produced per year, although we know that this is not a sound management practice. Decisions made to breed subfertile horses today will likely have long-term, potentially catastrophic effects with regard to fertility of future offspring. Economic pressures, however, eventually take precedence over ethical decisions regarding breeding of subfertile horses. Veterinary skills are then sought to satisfy short-term economic demands of horse owners.
Syndication of stallions Syndications are designed for ease of marketing, not for maximizing reproductive efficiency; however, syndications affect how stallions are managed in the breeding shed. Decisions are made to breed a minimum number (book) of mares before any knowledge is obtained regarding the potential fertility of a stallion. The minimum book of mares for a novice stallion is extremely variable within the Thoroughbred industry, being affected by many factors including his racing performance, his pedigree, the price of the syndication agreement, and the location where the stallion stands. The number of breedings that can be sold or used is divided among the syndicate members, and may total three or four mares to be bred per share, with 45 to 47 shareholders in the syndicate. In addition to acceptance of his book of mares based on a given number of breeding rights per shareholder, there often are extra breeding rights that are provided to the stallion’s trainer and to the farm that stands the stallion, and complimentary breedings are often given to others. Therefore, it has become commonplace for popular Thoroughbred stallions standing in
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Management of Stallions in Natural-Service Programs Table 118.1 Number of days in season in which 0-5 covers (including doubles) are required from Thoroughbred stallions. 12 years of data using 29 stallions with 16 080 covers. Days in season with 0-5 covers Book Size
N
0
1
2
3
4
5
0-19 20-39 40-49 50-59 60-69 70-79 80-89 90-99 100-109 110-119 120-129 130-139 140-149 150-159 160-169 170-179 180-189 190-199
4 19 9 9 8 6 4 4 5 7 5 4 5 2 4 2 6 4
122 92 76 62 58 49 35 31 30 35 29 25 24 19 17 27 13 15
14 36 45 52 50 50 55 59 53 43 45 40 44 36 32 29 30 27
0 7 13 20 24 27 36 36 42 36 40 43 41 38 37 37 35 37
0 1 2 2 5 9 8 11 10 20 19 22 21 26 29 31 33 33
0 0 0 0 0 0 1 0 1 3 5 7 7 17 22 13 25 23
0 0 0 0 0 0 0 0 0 0 0 0 1 2 0 1 2 2
central Kentucky to be booked to 120 to 200 or more mares in their first season at stud.
Client relations The single greatest concern managers have with regard to effective breeding management of stallions is client relations. The stallion manager/veterinarian must be an effective communicator to keep clients and other veterinarians informed of what is happening with the particular stallion to which they are breeding their mare(s). The booking office usually requires a breeding shed form be submitted for each mare that is to be covered on a given day and time (e.g., Figure 118.1). In addition, communications to personnel responsible for scheduling date and time of matings must include the stallion’s breeding schedule (i.e., number of covers allowed per day), time of breeding shed operations, and what is expected in terms of mare management to maximize opportunities to achieve pregnancies on a single breeding (cover or service). The goal is to breed each mare in a manner to achieve a live foal from a single cover. The restrictive level of management becomes quite important when standing an aged stallion, as he must be protected even more from overuse. For example, the older Thoroughbred stallion may be restricted to only one or two covers per day with no ‘double’ covers permitted (i.e., no mares covered twice during one estrous period) in an attempt to avoid overbreeding,
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particularly if daily sperm output is declining. Clear communication with clients is necessary even if they are breeding mares to a young, highly fertile stallion for which breedings are not as closely monitored or restricted. In some cases, particular restrictions are made for ‘slow’ breeding stallions that the mare manager must be aware of. For example, a ‘slow’ breeding stallion may only be bred to the last mare presented for a breeding session so that activity (distractions) in the shed is minimized. Or, a stallion with an injury may require special assistance during mating that necessitates special mare handling during the breeding process. In rare instances, a stallion may have to be ‘taken off-line’ for a period of time (e.g., acute equine herpesvirus 3 infection; coital exanthema), and close communication regarding precautions, potential for transmission of an infection, and/or expected return of the stallion to active breeding duty are critical. Finally, a mare that will not stand safely for breeding may be rejected from the shed in order to avoid potential for injury to all involved. Communicating times during the next few breeding sessions that the stallion has open affords the mare manager some options for attempting to get the mare bred before ovulation is missed on that estrous cycle; plus, promptly communicating that every effort will be made to try again to get the mare safely bred maintains good relations with clients that have mares booked to a given stallion. To achieve maximum breeding success with a stallion, breeding records must be accurate and up to date. When central stallion stations are maintained with mares arriving from satellite farms, it is assumed that an accurate assessment of anticipated ovulation dates has been made so that stallions with heavy books are not overworked unnecessarily. An example wherein inadequate communication can adversely affect reproductive management is having too large a number of barren and maiden mares in a stallion’s book arriving ready to be bred during the first few days of the breeding season, most commonly due to veterinarians at satellite farms synchronizing estrus in these mares in an effort to get them bred and pregnant as early as possible in the season. In such a case, the majority of mares may need to be bred over a 3- to 5-day period. The stallion(s) might then remain sexually inactive for a couple of weeks, resulting in inefficient management. Some stallions might be unable to breed all synchronized mares at the ideal interval before ovulation. So, communications between those managing the mares and those managing the stallions must be especially thorough when large-scale synchronization programs are to be employed. Mare managers may be requested to stagger administration of hormones (i.e., to only two or three mares per group, with each group being treated
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2010 BREEDING SHED FORM (EXAMPLE)
2010
2010
Figure 118.1 Example of a breeding shed form to be presented to the stallion manager when a mare arrives at the breeding shed for mating.
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Management of Stallions in Natural-Service Programs 3 to 4 days apart) when synchronizing estrus in large groups of mares meant to be bred to a single Thoroughbred stallion. Mares may be located at satellite farms that have management systems ranging from poor to excellent; therefore, mares occasionally may be presented for breeding when pregnant, not in good standing estrus, or not near ovulation. Use of a teaser stallion with excellent libido is imperative to screen outside mares sent to be bred to minimize chances of injury to the stallion (see discussion of breeding shed management later in this chapter). Even some maiden mares in physiological estrus (i.e., large soft follicle with an edematous uterus and relaxed cervix) will still not allow mounting by the teaser stallion. Mating decisions, though they may be unpopular, will have to be made accordingly to avoid injury, or non-productive use of the stallion. In some cases, sedation will relieve nervousness sufficiently that a difficult mare will allow mounting by the teaser, and can then be safely bred by the intended stallion. However, safety for the stallion and breeding shed personnel must come first, so mares that will not safely stand for mounting should be rejected from the breeding shed. Promptly communicating with the personnel responsible for management of the rejected mare, assuring them that an effort will be made to find a spot for the mare if she does not ovulate and can be trained to stand safely for mounting with a teaser stallion, will improve the chances that the mare will be bred at the right time as well as instill confidence that the breeding shed personnel are doing their best to safely accommodate mare owner needs. Some horse owners avoid veterinary services and rely only on caretakers to inform them of their mare’s reproductive status. Sometimes owners attempt to manage their mares by themselves, and bring the mares to the shed on a given date when they assume they should be ready for breeding (e.g., on day 30 after foaling). Unless the stallion manager has confidence in their judgment (obtained from prior knowledge or experience with either these persons or the veterinarians performing the reproductive examinations at the satellite farm), he cannot assume such mares are ready for breeding. The stallion manager must make an effort to be aware of each management system where mares are kept. This can be a difficult undertaking, because a large stallion station may receive mares for breeding from 300 to 400 different farms per year, each with varying management conditions.
Usefulness of breeding records It is advantageous for the stallion manager to maintain accurate, up-to-date breeding records so that fer-
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tility data can be used to help detect potential problems quickly. A variety of record systems (manual or computer-based) can be used for this purpose as long as they permit easy and thorough evaluation of stallion performance at any given point during the breeding season. One must remember that every day of the breeding season is important, because only 135–140 days (mid-February through July 1) are typically available to get mares pregnant. When mares are returning for rebreeding on the second or third estrous cycle, it is important to check for potential problems to which the manager/owner of the mare could be alerted. This communication is usually appreciated and may afford prompt correction of a problem, which could prevent a barren year for the mare. Barren years also cost the stallion owner because breeding fees may be lost in some instances. The Jockey Club has a computerized record system (‘Horse Farm Management’) that does summarize many pertinent fertility end points that can be calculated at intervals during and after the season (see summary printouts by stallion, by month, and by session or cover of the day, in Tables 118.2 to 118.4). Examples of a manual record system utilizing two records are given – one for entry of pertinent data from each cover in chronological order (see Figure 118.2) and one that graphically presents pregnancy outcome that can be used to examine effects of mating frequency, beginning status of mares covered and their cycle of the year, and the cover of the day for the stallion (see Figure 118.3). Evaluating reproductive performance at regular intervals throughout the season can not only provide confidence that a given stallion’s fertility is good, but can alert the cognizant manager to potential problems that may require prompt intervention to restore reproductive performance to an acceptable level.
Exercise programs Young horses recently retired from racing have been accustomed to regular exercise during their athletic career. They are then expected to move to a career in the breeding shed for the rest of their life. Regular exercise is an important component of management programs designed to maximize the breeding longevity of these stallions. One must remember that the stallion may be kept until he is in his late twenties or even older, because the best stallions become progressively more desirable and acceptable from a genetic standpoint as their offspring are continually proved. Genetic value of some of these older stallions is proved regularly each year, and such stallions continue to command respectfully sized books and are bred as long as possible. The goal of the overall management program should be to have each stallion
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Stallion 3
Stallion 2
9 6 7 3
Maiden
Not Bred
Slipped
Unknown
6 10
Slipped
115
3 8 118
20
Maiden
Not Bred
Slipped Stallion Total
30
70
9 169
5
95
17
30
150
In-Foal
Total Book 118
1
14
Barren
Stallion Total
1
11
Maiden
Not Bred
Unknown
15
70
11
85
17
In-Foal
24
114
3
10
10
11
66
Barren
Total Book 115
77
45
In-Foal
14
1 9
0
0
7
1
8
0
1
1
0
4
2
9
0
1
3
0
4
1
8 108
2
20
63
15
105
1
10
5
11
64
14
72
3
7
5
8
42
7
0 10
1
0
7
2
6
0
0
1
0
4
1
5
0
0
1
1
3
0
No. In-Foal No. Barren
0 0
0
0
0
0
4
0
0
0
0
2
2
0
0
0
0
0
0
0
No. Unknown
100.00 % 91.53 %
66.67 %
100.00 %
90.00 %
88.24 %
91.30 %
100.00 %
100.00 %
83.33 %
100.00 %
91.43 %
82.35 %
93.51 %
100.00 %
100.00 %
83.33 %
88.89 %
93.33 %
100.00 %
% In-Foal
0.00 % 8.47 %
33.33 %
0.00 %
10.00 %
11.76 %
5.22 %
0.00 %
0.00 %
16.67 %
0.00 %
5.71 %
5.88 %
6.49 %
0.00 %
0.00 %
16.67 %
11.11 %
6.67 %
0.00 %
% Barren
0.00 % 0.00 %
0.00 %
0.00 %
0.00 %
0.00 %
3.48 %
0.00 %
0.00 %
0.00 %
0.00 %
2.86 %
11.76 %
0.00 %
0.00 %
0.00 %
0.00 %
0.00 %
0.00 %
0.00 %
% Unknown
1.125 1.361
1.500
1.500
1.317
1.467
1.267
1.000
1.400
1.400
1.364
1.188
1.429
1.444
1.000
1.429
2.000
1.250
1.357
2.000
Covers per Preg
(continued)
1.125 1.432
1.667
1.500
1.357
1.765
1.304
1.000
1.400
1.833
1.364
1.214
1.412
1.481
1.000
1.429
1.667
1.222
1.467
2.000
Covers per Mare
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Stallion Total
7
Total Book 78
Barren
No. Covers Double
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No. Mares
(By: Final Status)
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Beginning Status
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Table 118.2 Stallion fertility analysis by stallion
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Stallion 5
Barren In-Foal Maiden Not Bred Slipped Unknown Stallion Total
Barren In-Foal Maiden Not Bred Slipped Stallion Total
Total Book 50 7 33 3 5 2 50 Total Book 97 19 46 17 9 5 1 97
No. Mares
26 65 20 14 7 1 133
13 42 6 6 2 69 3 4 0 0 1 0 8
1 2 1 1 0 5
No. Covers Double
14 35 16 8 4 1 78
6 30 3 5 1 45 5 9 1 1 1 0 17
1 3 0 0 1 5
No. In-Foal No. Barren
0 2 0 0 0 0 2
0 0 0 0 0 0
No. Unknown
(By: Final Status)
73.68 % 76.09 % 94.12 % 88.89 % 80.00 % 100.00 % 80.41 %
85.71 % 90.91 % 100.00 % 100.00 % 50.00 % 90.00 %
% In-Foal
26.32 % 19.57 % 5.88 % 11.11 % 20.00 % 0.00 % 17.53 %
14.29 % 9.09 % 0.00 % 0.00 % 50.00 % 10.00 %
% Barren
0.00 % 4.35 % 0.00 % 0.00 % 0.00 % 0.00 % 2.06 %
0.00 % 0.00 % 0.00 % 0.00 % 0.00 % 0.00 %
% Unknown
1.000 1.286 1.188 1.375 1.500 1.000 1.231
1.500 1.267 2.000 1.200 1.000 1.333
Covers per Preg
1.368 1.413 1.176 1.556 1.400 1.000 1.371
1.857 1.273 2.000 1.200 1.000 1.380
Covers per Mare
Trim: 279mm X 216mm
Stallion 4
Beginning Status
EXAMPLE FARM
User: investigator
11:50
Season: 2008 Hemisphere: Northern
February 1, 2011
Company:
P2: SFK
Stallion Fertility Analysis by Stallion
BLBK232-McKinnon
Run Date: 7/12/2008 6:19 AM
c118
Table 118.2 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
1214
Stallion 4
Stallion 3
February March April May June Stallion Total
February March April May June Stallion Total
February March April May June Stallion Total
Total Book 50 6 19 13 19 7
Total Book 118 34 43 37 37 8
Total Book 115 20 40 41 35 11
Total Book 78 17 36 30 16 6
6 19 14 23 7 69
34 44 42 42 7 169
20 40 41 38 11 150
17 38 34 18 7 114
0 3 0 2 0 5
0 2 1 3 3 9
0 1 3 4 0 8
2 1 4 2 0 9
Double
4 14 8 14 5 45
24 28 24 26 6 108
13 30 31 25 6 105
9 25 20 14 4 72
No. In-Foal
66.67 % 73.68 % 61.54 % 73.68 % 71.43 % 90.00 %
70.59 % 65.12 % 64.86 % 70.27 % 75.00 % 91.53 %
65.00 % 75.00 % 75.61 % 71.43 % 54.55 % 91.30 %
52.94 % 69.44 % 66.67 % 87.50 % 66.67 % 93.51 %
% In-Foal
1.500 1.214 1.500 1.429 1.000 0.111
1.333 1.429 1.542 1.269 0.833 0.046
1.462 1.267 1.226 1.280 1.000 0.057
1.889 1.480 1.450 1.214 1.000 0.056
Covers per Preg
(continued)
1.000 1.000 1.077 1.211 1.000 1.380
1.000 1.023 1.135 1.135 0.875 1.432
1.000 1.000 1.000 1.086 1.000 1.304
1.000 1.056 1.133 1.125 1.167 1.481
Covers per Mare
Trim: 279mm X 216mm
Stallion 2
February March April May June Stallion Total
No. Covers
11:50
Stallion 1
No. Mares
(By: Final Status)
User: investigator
February 1, 2011
Month
EXAMPLE FARM
Season: 2008 Hemisphere: Northern
BLBK232-McKinnon
Company:
Stallion Fertility Analysis by Month
P2: SFK
Run Date: 7/12/2008 6:23 AM
c118
Table 118.3 Stallion fertility analysis by month
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
1215
Stallion 6
February March April May June Stallion Total
February March April May June Stallion Total Total Book 69 13 20 25 21 9
Total Book 97 20 35 31 34 10
No. Mares
13 22 26 25 10 96
20 37 31 35 10 133
No. Covers
1 2 2 2 2 9
1 1 4 2 0 8
Double
12 13 17 15 5 62
14 24 19 16 5 78
No. In-Foal
92.31 % 65.00 % 68.00 % 71.43 % 55.56 % 89.86 %
70.00 % 68.57 % 61.29 % 47.06 % 50.00 % 80.41 %
% In-Foal
(By: Final Status)
1.083 1.538 1.471 1.133 1.000 0.081
1.286 1.250 1.316 1.125 1.000 0.064
Covers per Preg
1.000 1.100 1.040 1.190 1.111 1.391
1.000 1.057 1.000 1.029 1.000 1.371
Covers per Mare
investigator
Trim: 279mm X 216mm
Stallion 5
Month
EXAMPLE FARM
User:
11:50
Season: 2008 Hemisphere: Northern
February 1, 2011
Company:
P2: SFK
Stallion Fertility Analysis by Month
BLBK232-McKinnon
Run Date: 7/12/2008 6:23 AM
c118
Table 118.3 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
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Stallion 2
115
150
91 43 15 1
1st Cover of Day 2nd Cover of Day 3rd Cover of Day 4th Cover of Day
Stallion Total
76 5 53 16
Session 1 Session 2 Session 3 Session 4
114
70 31 11 2
1st Cover of Day 2nd Cover of Day 3rd Cover of Day 4th Cover of Day
8
3 3 2 0
1 0 5 2
9
7 2 0 0
3 2 4 0 0
Double
105
60 32 12 1
50 3 40 12
72
43 18 9 2
27 5 30 9 1
No. In-Foal
6
4 1 1 0
3 0 2 1
5
4 1 0 0
3 0 2 0 0
No. Barren
4
4 0 0 0
4 0 0 0
0
0 0 0 0
0 0 0 0 0
No. Unknown
91.30 %
88.24 % 96.97 % 92.31 % 100.00 %
87.72 % 100.00 % 95.24 % 92.31 %
93.51 %
91.49 % 94.74 % 100.00 % 100.00 %
90.00 % 100.00 % 93.75 % 100.00 % 100.00 %
% In-Foal
5.22 %
5.88 % 3.03 % 7.69 % 0.00 %
5.26 % 0.00 % 4.76 % 7.69 %
6.49 %
8.51 % 5.26 % 0.00 % 0.00 %
10.00 % 0.00 % 6.25 % 0.00 % 0.00 %
% Barren
3.48 %
5.88 % 0.00 % 0.00 % 0.00 %
7.02 % 0.00 % 0.00 % 0.00 %
0.00 %
0.00 % 0.00 % 0.00 % 0.00 %
0.00 % 0.00 % 0.00 % 0.00 % 0.00 %
% Unknown
1.267
1.317 1.219 1.167 1.000
1.340 1.333 1.175 1.250
1.444
1.442 1.667 1.111 1.000
1.519 1.000 1.433 1.556 1.000
Covers per Preg
(continued)
1.304
1.338 1.303 1.154 1.000
1.333 1.667 1.262 1.231
1.481
1.489 1.632 1.222 1.000
1.467 1.000 1.531 1.667 1.000
Covers per Mare
Trim: 279mm X 216mm
Stallion Total
44 5 49 15 1
No. Covers
Session 1 Session 2 Session 3 Session 4 Session 5
77
No. Mares
(By: Final Status)
User: investigator 11:50
Session Cover
EXAMPLE FARM
Season: 2008 Hemisphere: Northern
February 1, 2011
Stallion 1
Company:
P2: SFK
Stallion Fertility Analysis by Session
BLBK232-McKinnon
Run Date: 7/12/2008 6:25 AM
c118
Table 118.4 Stallion fertility analysis by session
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
1217 118
69
Stallion Total
50
39 25 5 52 15 2
Session 1 Session 3 Session 4 1st Cover of Day 2nd Cover of Day 3rd Cover of Day
169
93 52 20 4
1st Cover of Day 2nd Cover of Day 3rd Cover of Day 4th Cover of Day
Stallion Total
75 9 65 20
Session 1 Session 2 Session 3 Session 4
No. Covers
5
2 2 1 4 1 0
9
4 3 1 1
2 0 5 2
Double
45
25 18 2 34 10 1
108
58 37 10 3
46 6 44 12
No. In-Foal
5
2 1 2 2 2 1
10
7 1 1 1
7 0 2 1
No. Barren
0
0 0 0 0 0 0
0
0 0 0 0
0 0 0 0
No. Unknown
90.00 %
92.59 % 94.74 % 50.00 % 94.44 % 83.33 % 50.00 %
91.53 %
89.23 % 97.37 % 90.91 % 75.00 %
86.79 % 100.00 % 95.65 % 92.31 %
% In-Foal
10.00 %
7.41 % 5.26 % 50.00 % 5.56 % 16.67 % 50.00 %
8.47 %
10.77 % 2.63 % 9.09 % 25.00 %
13.21 % 0.00 % 4.35 % 7.69 %
% Barren
0.00 %
0.00 % 0.00 % 0.00 % 0.00 % 0.00 % 0.00 %
0.00 %
0.00 % 0.00 % 0.00 % 0.00 %
0.00 % 0.00 % 0.00 % 0.00 %
% Unknown
1.333
1.280 1.333 2.000 1.324 1.400 1.000
1.361
1.345 1.351 1.600 1.000
1.370 1.500 1.341 1.333
Covers per Preg
1.380
1.444 1.316 1.250 1.444 1.250 1.000
1.432
1.431 1.368 1.818 1.000
1.415 1.500 1.413 1.538
Covers per Mare
Trim: 279mm X 216mm
Stallion 4
Stallion 3
No. Mares
(By: Final Status)
User: investigator
11:50
Session Cover
EXAMPLE FARM
Season: 2008 Hemisphere: Northern
February 1, 2011
Company:
P2: SFK
Stallion Fertility Analysis by Session
BLBK232-McKinnon
Run Date: 7/12/2008 6:25 AM
c118
Table 118.4 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
c118 BLBK232-McKinnon P2: SFK February 1, 2011 11:50 Trim: 279mm X 216mm
Figure 118.2 A manual recording system to be filled out chronologically with pertinent data for a given stallion after each cover is completed. Information to be filled in with each cover includes: date of cover, mare identification, farm of origin for the mare, beginning status of the mare (foaling, maiden, barren, not bred, slipped), cycle of breeding (1st, 2nd, etc.), cover of the day for the stallion (1st, 2nd, etc.), number of mounts required to complete service, whether the cover is a double during the same cycle or not, quality of the dismount sample (may include a scoring system for motility and concentration), whether reinforcement breeding was utilized on that cover or not, whether or not the mare had to be tranquilized to facilitate a safe cover, other comments thought to useful (e.g., was the mare difficult to cover and why, did the mare tolerate a twitch, were inflammatory cells noted in the dismount, etc.), and the outcome of the cover (pregnant or not pregnant).
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
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c118
P2: SFK
Color: 1C
BLBK232-McKinnon
February 1, 2011
11:50
Trim: 279mm X 216mm
Printer Name: Yet to Come
Figure 118.3 (continued)
P1: SFK/UKS
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P1: SFK/UKS c118
P2: SFK
Color: 1C
BLBK232-McKinnon
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February 1, 2011
11:50
Trim: 279mm X 216mm
Printer Name: Yet to Come
Stallion: Management
reach this age as fit and eager for breeding as during the first year he was at stud. Unless a young horse has suffered a serious racing injury or medical problem that necessitates an inactive lifestyle, the stallion manager must determine how to best exercise the horse daily. The determination will depend to a great extent on the individual horse’s disposition. Some stallions require no forced exercise. When they are turned out in a one- or twoacre paddock, they run and play all day, thus getting adequate exercise. Other horses, however, when turned out in a similar paddock will only graze or stand in one spot for most of the day, essentially getting no exercise. Such horses require some form of forced exercise to remain fit and healthy. Forced exercise may take the form of riding under tack. Lunging at a fast walk or trot for 15 to 30 minutes or an hour daily are other forms of forced exercise used on some breeding farms. Stallions can even be exercised just prior to breeding. We know of one well-managed Throroughbred breeding farm that has stallions ridden 6 days per week year-round. The horses are ridden at a slow canter for one or two miles daily. Exercise is felt to be the most important factor on that farm in maintaining proper weight, body condition, energy level, and breeding interest of the stallions. It is imperative that persons exercising stallions be good horsemen or horsewomen so that overworking and injuries can be avoided. If soreness develops, the exercise session should be discontinued and the stallion rested for an appropriate time period. The amount of time a horse is kept in the paddock must be tailored to his personality. Some
well-mannered stallions are best kept outdoors 20– 24 hours a day except during inclement weather. They are brought into the breeding shed for mating and then are turned back into the paddock. Other stallions may only be left in a paddock for 4 hours each day. Certain horses get too excited when they are outside during the day in a location where they can see other horses. A more suitable exercise program for such stallions is to keep them indoors in a stall during the day and turn them out at night when other activity is minimal. Some stallions tolerate being in a paddock adjacent to another stallion; others may not. Times when stallions are scheduled for paddock exercise must be coordinated with their individual preferences. Some stallion managers turn their stallions out with an old, mild-mannered mare for company in an effort to keep the stallion relatively quiet. We do not recommend this practice. We believe the flexible stallion manager using common sense can determine a more suitable exercise program that will better minimize chances of injury to the breeding stallion. In some cases, this may require locating the exercise paddock in a quiet, isolated area where little activity is visible to the stallion. Lastly, some stallions rub or kick at paddock fencing. One solution that may work for this problem is to board up the section or corner of the fence where this activity seems to be concentrated. We have noted this procedure works for a number of stallions, reducing the chance of self-injury and rapidly causing cessation of this behavior. Another approach
Figure 118.3 A manual recording system (chart) to be used to graphically represent a stallion’s performance (fertility) for a given month. Each box in the lower graph is filled out in order (from bottom up on each given date) as each mare is successfully covered. For example, if the cover was for a barren mare bred on her first cycle of the season, B1 would be placed in that block for that day. If the stallion then bred a foaling mare on her first cycle later on the same day, an F1 would be placed in the box just above the recording for the previous mare. As pregnancy data becomes available, blocks are highlighted with ‘green’ if the mare became pregnant, or ‘red’ if the mare did not become pregnant. The top graph is blocked in at the end of each day, with each column of blocks for a given date representing the total number of mares covered in the 3 days preceding that date. This graph provides an overall indication of how busy the stallion has been, and can then be used in conjunction with pregnant and open mares recorded as green or red in the lower graph, to examine the influence of frequency of mating on likelihood of establishing pregnancy. For example, if a stallion’s pregnancy rate declines substantially when he is averaging more than 3 covers each day (i.e., more than 9 blocks filled in the top graph), the stallion manager may wish to limit the maximum number of covers in any 3-day period to 9. Over time, glancing at the pattern of green vs red blocks will provide an impression of the stallion’s fertility. At the end of the month, fertility end points listed can be calculated. Evaluating these end points allows conclusions to be made about the stallion’s fertility, as both quality of mare and mating frequency can affect pregnancy rates. For example, if too many ‘problem’ mares are in the stallion’s book, the overall pregnancy rate may appear low, but may simply be due to having so many 2nd or 3rd cycle barren mares being covered. If the stallion’s pregnancy rates remain good for maiden and foaling mares, the stallion’s fertility may not be a concern. Additionally, the first cycle pregnancy rate is calculated only for those mares bred on their first cycle of the season. This percentage can remain good, despite a declining overall pregnancy rate (due to problem breeding mares, requiring more cycles of breeding, accumulating within a given month), thereby indicating acceptable fertility for the stallion. Finally, in addition to the discussion above on utilizing mating frequency data to see how it affects pregnancy rates, pregnancy rates can also be calculated for each cover of the day. If mating frequency is a problem, pregnancy rates are likely to dramatically decline on later covers of the day. Summarizing this information on a monthly basis can provide confidence that the stallion is maintaining acceptable fertility and management of the stallion needs not be altered. Alternatively, the stallion manager may be alerted to the need for further investigation into the cause of poor reproductive performance.
P1: SFK/UKS c118
P2: SFK
Color: 1C
BLBK232-McKinnon
February 1, 2011
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Trim: 279mm X 216mm
Printer Name: Yet to Come
Management of Stallions in Natural-Service Programs that we have seen some farms use successfully is to place electric tape around the top inside board of the paddock fence. Apparently, the stallion rapidly learns to respect the fencing. Once the behavior is corrected, electrical current to the tape may be turned off unless the behavior recurs.
Feeding programs Feeding stallions the best quality forages available will result in maintenance of excellent body condition, attitude, and breeding performance with fewer digestive upsets. It is best to purchase hay that has been tested accurately for nutrient content to ensure quality of the forage and to facilitate sound ration formulation. Energy (including fat), protein, and vitamin and mineral supplementation should be balanced according to NRC recommendations based on total nutrient consumption in the entire diet – thus the need for accurate forage nutrient analysis. Many farms employ nutritional consultants that rebalance rations as the source and quality of hay changes. There are many untested nutriceutical products on the market that are claimed to improve stallion libido and fertility. While certain supplements may someday be proven to be advantageous to stallion fertility, the stallion manager should be cautious about accepting manufacturer’s claims without scientific justification. Some specific nutrients known to be important in maintaining fertility in domestic animals include adequate amounts of the vitamins A (as beta-carotene), C and E, the mineral selenium, and appropriate ratios of omega-3 to omega-6 fatty acids in the ration. Current research pertaining to the horse is addressing the importance of antioxidants, immune system protective agents, and omega-3 fatty acids in diets. Again, the stallion manager is cautioned to not oversupplement fats, vitamins or minerals as detrimental health effects can follow. It is a good idea to weigh stallions monthly and to regularly monitor body condition and hair coats. Body condition scoring systems or weight tapes are useful in this regard. Recording amounts of each ingredient fed and appetite of the horse should be done daily so that adjustments can be made in the feeding program on a timely basis. Communication with farm personnel responsible for feeding, care, and handling of stallions is critical so that nutritional problems that may be associated with lackluster breeding performance are addressed in an efficient manner; making wholesale changes without knowing where the starting point is will just confuse the issue. Paradoxically, our biggest worry is getting breeding stallions overweight (obese). Overfeeding may predispose to digestive or feet problems that some-
1221
times lead to death or a decreased ability to use the stallion. In our experience, inadequate exercise is the principal cause of obesity in stallions. When overconditioning occurs, one must re-evaluate both feeding and exercise programs to correct this problem.
Preventive medicine In addition to sound exercise and feeding programs, preventive medicine programs should include adequate foot care, teeth care, vaccination, and deworming programs. The strategy of preventive medicine for the breeding stallion is predicated on the concept of having the stallion fit and healthy for the entire breeding season, which typically extends from February 14 or 15 through at least July 1 (northern hemisphere) of each year. During the breeding season, stallions must be ready to breed for 135 to 155 days. They cannot be sick or lame, and they must be capable of breeding every day. Hoof care An advantage of keeping stallions in grassy paddocks is that hoof problems are minimized. If a stallion has to be kept in a stall much of the time, his feet may dry out. Packing the feet daily or having the horse shod are procedures that will help to keep the hooves and soles flexible, thereby facilitating regular hoof trimming to minimize foot problems. Feet should be trimmed at regular 6- to 8-week intervals in most instances. Some stallions may require more frequent trimming. Any foot problem in a breeding stallion should be addressed promptly as the associated pain is likely to be worsened in the breeding shed, and may interfere with mating efficiency or even willingness to breed. Dental care Regular dental care (twice per year) is a must for stallions. If regular, thorough dental examination and preventive floating are initiated when young stallions arrive on the farm, dental problems will be minimized. Regular dental care helps ensure feeds are consumed properly and body condition is maintained, and reduces the occurrence of digestive upsets. Vaccinations Our strategy has been to vaccinate stallions for protection against all diseases they are at significant risk of contracting. We therefore vaccinate stallions against the following: tetanus, once yearly; influenza,
P1: SFK/UKS c118
P2: SFK
Color: 1C
BLBK232-McKinnon
1222
February 1, 2011
11:50
Trim: 279mm X 216mm
Printer Name: Yet to Come
Stallion: Management
every 2 months; rhinopneumonitis (EHV 1,4), every 2 months; botulism, twice the first year and then yearly thereafter if at risk; rabies, once yearly; Eastern, Western, and sometimes Venezuelan equine encephalitis (some countries prevent importation of horses that are seropositive for VEE – stallion managers should make sure that the owner(s) of the stallion do not have any intentions of shipping horses to these countries prior to vaccination for VEE); once yearly; West Nile virus infection, if using killed virus product – administer twice in the spring the first year, and again in late summer, then twice yearly thereafter – if using newer genetically engineered live vector vaccines, once yearly in the spring; equine arteritis virus, once yearly; Potomac horse fever, yearly if at significant risk. Some veterinarians do not vaccinate against Potomac horse fever since research studies with current vaccines have failed to demonstrate protection against contracting the disease or protection in the form of reducing severity of the disease once it is contracted. On January 1, each stallion is immunized against tetanus (toxoid), rabies, influenza, rhinopneumonitis, botulism, and Potomac horse fever (if used). On March 1, each stallion is immunized against rhinopneumonitis, influenza, Eastern and Western (and Venezuelan if given) equine encephalitis, and West Nile virus encephalomyelitis. This date is just prior to mosquito season in our area. Some farms prefer to administer rabies vaccines on March 1 instead of January if skunk (the primary carrier in our area) activity is minimal prior to this time. Remaining influenza and rhinopneumonitis vaccinations are administered on the first day of May, July, September, and November. Vaccination against equine viral arteritis should be guided by the requirements of the state in which stallions are standing at stud. For example, the veterinarian may be required to demonstrate a negative titer to equine viral arteritis prior to administration of vaccine, as vaccinal titers cannot be differentiated from those arising from natural infection. The incidence of fevers following vaccination against equine viral arteritis is apparently low, but one of the authors prefers to vaccinate against equine viral arteritis in late December, and to administer flunixin meglumine 2–4 hours prior to administering the equine viral arteritis vaccine, in order to avoid development of any vaccine-related fever that could result in transient thermal damage to testes. An interesting finding concerning botulism is given as a precautionary note. Even though botulism may not have occurred on a particular farm, new construction that results in soil disturbance can expose spores and lead to an outbreak of the disease. Therefore, if a new breeding shed, stallion barn or other building is being constructed, the manager may
wish to institute an immunization program against botulism. Deworming program We believe it is best to deworm and vaccinate all horses on the farm, including the stallions and teasers, on a tight schedule so that horses are not overlooked. This ensures that immune protection is adequate for all horses at risk of developing and transmitting infectious diseases, and parasite loads for all horses on the farm are reduced. Stallions are dewormed every 60 days. Broad-spectrum parasiticides, such as pyrantel pamoate, and ivermectin/avermectin, are used on a rotating basis in an effort to avoid development of parasite resistance. Praziquantel should be administered with an ivermectin product twice yearly, at 6-month intervals, to control tapeworm infestations that may lead to an increased incidence of spasmodic colic even when present in low numbers. Fecal flotation examinations (including estimates of eggs per gram) can be performed on each stallion’s manure monthly to monitor worm burdens. If stallions are limited to individual paddocks with no cross-contamination, and paddocks are harrowed at regular intervals, worm burdens usually remain very low (e.g., fecal flotations will yield 0–5 eggs per gram).
Breeding management The manager must ensure that all personnel in the breeding shed thoroughly understand their specific jobs and perform them in a professional, efficient manner. Such teamwork cannot be achieved on the first day of the breeding season unless the procedures have been practiced beforehand. Therefore, starting in early January, procedures are practiced to get the routine set for all personnel, as well as the stallions. The number of people, traffic, and noise in the area of the breeding shed should be limited during breeding sessions, because these variables will distract certain stallions. Additionally, we have observed some stallions that refuse to develop an erection and breed when women are present in the breeding shed, perhaps because they were trained and handled only by men in the past. Occasionally, the opposite is true, wherein a stallion will refuse to breed or may even become overly aggressive when men are present in the breeding shed. If patient retraining does not correct this behavior, handling by only men or only women may be necessary to breed such stallions successfully. Operation of large Thoroughbred stallion stations may necessitate the simultaneous use of two breeding sheds. The breeding operation can be viewed
P1: SFK/UKS c118
P2: SFK
Color: 1C
BLBK232-McKinnon
February 1, 2011
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Trim: 279mm X 216mm
Printer Name: Yet to Come
Management of Stallions in Natural-Service Programs as a volume operation necessitating extremely efficient use of time. Several breeding sessions occur every day, generally at relatively evenly spaced time intervals. For example, if three breeding sessions occur each day, stallions may be bred at 9 am, 2 pm and 7 pm. How critical it is for matings to be evenly spaced throughout the day is not known. Realizing that most stallions can deliver complete ejaculates following 1–2 hours of sexual rest, one manager instituted a program wherein stallions are bred up to five times per day between 7:30 am and 5 pm (e.g., 7:30 am, 10:00 am, 11:30 am, 2:30 pm and 5 pm). Pregnancy rates on different covers of the day appeared to be comparable to those achieved in previous years when matings were spaced further apart, between 7:30 am and 9 pm, yet the shorter-day schedule was less stressful on the breeding shed crew due to avoiding late evening matings. Stallions must be treated as individuals in that the stallion manager must be aware of idiosyncrasies among stallions. For example, some stallions may prefer breeding ‘uphill’ (i.e., off a breeding mound) or with two breeding rolls, whereas other stallions may prefer breeding on a flat surface. Individual stallion preferences should be accommodated in the interest of keeping the horse satisfied with the breeding process. In some instances, the stallion manager may find altering the location of breeding within the shed may occasionally improve libido and mating efficiency. For example, when a usually eager stallion is slow to express interest in a given mare, it may be helpful to move the mare to a different location within the shed for mating (flat vs mound; on rubber surface rather than mat surface; etc.; see Figure 118.4). Procedures for breeding the stallion by live cover can be viewed as activities that occur as a horse progresses through several ‘stations’: (1) the receiv-
Figure 118.4 A Thoroughbred breeding shed with shredded rubber surface throughout, except for a small breeding area with cocoa mat. Individual 4 × 6 cocoa mats can be placed behind or under mare’s hind feet to adjust height of hindquarters if stallion is too short or tall for a given mare. Mares can also be bred on level surface of shredded rubber floor.
1223
Figure 118.5 After the mare is unloaded, she is moved to a location for identification to ensure the correct mare is going to be bred to the correct stallion. Identification is checked against the breeding form, and a tag with the stallion’s name to which she is booked for breeding is attached to her halter.
ing area; (2) the teasing and identification area; (3) the mare preparation area; (4) the stallion preparation area; and (5) the breeding area. The receiving area must accommodate any type of vehicle that may bring mares to the breeding shed (e.g., vans, trailers, or trucks). This area must be designed so that mare transports may be moved in and out easily and mares may be unloaded and moved to their respective locations without the area becoming congested. The receiving area should be secure in case a mare escapes from her handler, to avoid injuries and to prevent the loose mare from disrupting other breeding procedures. The teasing and identification location (Figure 118.5) is an extremely critical area. The mare to be bred is moved to this location for identification to ensure the correct mare is going to be bred to the correct stallion. The owner or manager of the farm whence the mare came is responsible for ensuring that a neck strap or name plate on a halter is in place with the mare’s identification on it. A breeding form (Figure 118.1) that has been filled out describing the mare’s current status as maiden, barren, foaling (wet), not bred, or slipped, and indicating if the mare is imported (i.e., has not been in the United States for longer than a year), is examined to ensure the correct mare is being bred to a given stallion. The stallion to which the mare is to be bred is also indicated on the form. Farm personnel from the stallion station have already checked this information against their records, saving a great deal of time for breeding shed personnel. At this point, a tag that gives the mare’s status and the stallion to which she is to be bred is placed on the mare’s halter. Then the mare is teased (Figure 118.6) to ensure she is in firm, standing estrus. If the mare does not express convincing evidence that she will stand to be bred, she is
P1: SFK/UKS c118
P2: SFK
Color: 1C
BLBK232-McKinnon
1224
February 1, 2011
11:50
Trim: 279mm X 216mm
Printer Name: Yet to Come
Stallion: Management
Figure 118.6 The mare is turned free within a teasing stall and a window is opened that allows the teaser stallion to reach through to tease the mare for signs of standing estrus. If the mare is shy or noncommittal, she can be placed on a lead and forced next to the window so the stallion can nuzzle and rub her to better elicit estrus behavior.
mounted with a teaser stallion (Figure 118.7) that has been fitted with a breeding shield to prevent intromission. If she stands for the teaser, she is moved to the mare preparation area. The mare preparation area is a padded stock (Figure 118.8) where the mare’s tail is wrapped and the perineal and genital areas are washed with clean water, cotton, and a mild soap. A mild soap (coconut oil or Ivory soap) is used to avoid excessive displacement of normal flora. It is important that the person preparing the mare thoroughly rinse any residual soap off, and dry the area with paper towels, as soap and water can be toxic to sperm. Additionally, if a spray hose is used for this washing, it should never be directed forward, to avoid forcing water into the
Figure 118.7 This mare has been fitted with padded boots prior to being exposed to the teaser stallion, which has been fitted with a shield that completely covers his underside and genitals to prevent breeding. The stallion is encouraged to tease the mare strongly until she exhibits signs of standing estrus. He is then allowed to mount the mare and remain mounted for a short period of time. If the mare stands safely for the teaser stallion, this provides some confidence that the mare will stand safely for mating to the stallion to which she is booked.
Figure 118.8 In the preparation area, the mare’s tail is wrapped and the perineal and genital areas are then washed with clean water, cotton, and a mild soap.
vestibule or vagina. Such errors in preparing the mare may lower pregnancy rates. The mare’s vulvar lips and vestibular mucosa should be examined closely to ensure no active herpesvirus (EHV3 ) lesions (blisters, pustules, erosions) are present that could result in transmission of the virus to the stallion, and subsequently to other mares bred afterwards. It is also examined to ensure the stallion will have adequate space to insert his penis into the vulva and vagina. A restrictive Caslick suture may have to be opened, particularly in maiden mares that have been sutured at the race track. Occasionally, a breeding stitch will have to be removed because of improper placement. Finally, a small amount of sterile, nonspermicidal lubricant in placed on and just inside the vulvar lips, and the mare is then moved to the breeding shed proper where padded boots can be placed on her hind feet to reduce chances of a serious injury to the stallion should she inadvertently kick him (Figure 118.9). The breeding shed floor should be a non-slippery, cushioned, dust-free open area with padded walls
Figure 118.9 Padded boots are strapped onto a mare’s hind feet prior to being covered in the breeding shed. The padded boots lessen the risk of serious injury occurring to the stallion if he is kicked by the mare.
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Figure 118.10 Prior to being bred, the mare can be fitted with a protective cape to protect her from injury due to the stallion biting during the act of mating.
to ensure safety for all horses and personnel (Figure 118.4). At least four or five persons work together as the breeding shed team. Once the mare enters the breeding shed proper, this team controls the breeding procedure. No other personnel are permitted on the breeding shed floor, because the team knows the stallion’s habits and works together as an integral unit to accomplish the cover in a safe, efficient manner. Each member of the team has been trained to handle each job capably so they may be rotated among the various jobs as necessary. Prior to breeding, a cape or shield can be fitted over the mare’s neck and withers (see Figure 118.10) for protection from biting by the stallion. The cape also gives the stallion something to grasp onto and help maintain his balance during the cover, and also helps to prevent the mare from getting aggravated at the stallion for biting her. As the mare is positioned for breeding, the stallion is walked into the breeding shed and exposed to the mare until he develops an erection. He is backed into a safe area, sometimes padded, for washing (Figure 118.11). The stallion’s
Figure 118.11 The stallion can be situated in a padded area in order for the penis to be inspected and cleaned prior to intromission.
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Figure 118.12 The stallion’s penis is rinsed using cotton and plain warm tap water, and the penis is then dried. Disinfectants and soaps are not used, to prevent displacement of normal flora.
penis is examined, then rinsed using cotton and plain warm tap water, and dried (Figure 118.12). Disinfectants and soaps are not used, to prevent displacement of normal flora from the surface of the penis. Then the member of the team referred to as the ‘head-man’ places a twitch on the mare’s upper lip for restraint (see Figure 118.13). The head-man is responsible for informing team members if the mare’s disposition changes if he suspects the mare is about to kick, buck, or otherwise try to injure the stallion. The next member of the team, referred to as the ‘legman’, is responsible for placing a leg strap on the left front leg of the mare (Figure 118.13). The leg-man
Figure 118.13 The ‘head-man’ secures a twitch to the mare’s muzzle. In almost every instance, the mare will remain twitched by the head-man during the entire breeding procedure. The twitch will not be removed until the stallion has safely dismounted and has been backed to a safe distance from the mare. The ‘leg-man’ is responsible for placing a leg strap on the left front leg of the mare, pulling the leg up and wrapping the strap above the knee and around the lower leg. The leg is brought up just prior to permitting the stallion to mount the mare as an additional precaution against getting the stallion kicked.
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Figure 118.14 The stallion handler is responsible for controlling the stallion during the approach to the mare and during the mating process.
pulls the leg up, wrapping the strap above the knee and around the lower leg, just prior to permitting the stallion to mount the mare. Once the stallion has mounted the mare and is fully intromitted into her vagina, the stallion handler tells the leg-man to drop the leg to permit the mare to support herself under the stallion’s weight. In rare instances in which the mare does not stand firmly and moves from side to side, the leg may be held up during the entire mating until the stallion has safely dismounted and been backed away from the mare. In contrast to using a leg strap, some stallion managers prefer to use mare hobbles. The stallion handler is responsible for bringing the stallion to the mare’s hindquarters from the rear at a slight (30–45◦ ) angle (see Figure 118.14). As the stallion mounts the mare, the handler lifts his lead so the stallion’s front foot does not become entangled in it. If necessary, the handler uses pressure on the lead to ensure the stallion remains mounted in line with the mare. The ‘entering-man’ is responsible for assisting the stallion in inserting the penis into the mare’s vagina, ensuring the penis is inserted underneath the Caslick suture or breeding stitch (Figure 118.15). The ‘tail-man’ is positioned on the right side of the mare’s rump opposite the stallion’s approach. He is responsible for pulling the mare’s tail down and under the stallion’s right front foot when the stallion mounts the mare, and then holds the tail out of the way to give the entering-man full view of the mare’s vulva. This ensures tail hairs don’t get wound up on the stallion’s penis causing lacerations (Figure 118.16). He is also responsible for inserting a breeding roll between the mare’s perineal body and the dorsal surface of the stallion’s penis (Figure 118.17) if the mare is a maiden or if there is concern that the stallion’s penis is so large that injury may occur to the mare’s
Figure 118.15 The ‘entering-man’ is responsible for assisting the stallion in inserting the penis into the mare’s vagina, ensuring the penis is inserted underneath the Caslick suture or breeding stitch.
vagina. A breeding roll (sometimes two) can also be used to avoid tearing of vaginal or cervical tissues. Breeding rolls come in varying sizes, and can be covered with disposable plastic liners or rectal sleeves that are changed after each use to prevent transmission of infectious disease (e.g., EHV3). If necessary, the tail-man also helps steady the mare and stallion while the stallion is mounted on the mare. After the stallion’s tail flags or the stallion otherwise indicates he has ejaculated, he is held briefly in position on the mare to prevent a premature dismount while his glans penis remains engorged. This is done in an attempt to prevent the stallion from ‘dragging’ excessive semen from the vagina as he
Figure 118.16 The ‘tail-man’ is responsible for deflecting the tail to the side to prevent the penis from becoming entangled in tail hairs during intromission, and to help steady the mare during the breeding process. He is also responsible for inserting a breeding roll above the penis for selected mares to reduce penetration distance by the penis during mating.
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Figure 118.17 A breeding roll has been inserted between the mare’s perineum and the stallion, above his penis, to prevent the penis from being fully inserted. This procedure is most commonly done to prevent vaginal rupture in the maiden mare, but can also be used to protect against vaginal or cervical tearing following reconstructive surgery.
dismounts fully erect with an engorged glans (thereby acting as a plunger creating a vacuum when it is withdrawn). The stallion is permitted to dismount, at which time the mare’s hindquarters are swung away from the stallion (Figure 118.18). A dismount sample (i.e., ‘drippings’) of seminal fluid is collected from the stallion’s distal urethral process by the entering-man or attending veterinarian after the stallion dismounts (Figure 118.19). This sample is immediately taken to the semen laboratory, where it is examined under a phase-contrast microscope to confirm the presence of spermatozoa. If spermatozoa are not present in the sample, the stallion is brought back to the mare and the mare is covered again. When a second cover is required because the first attempt did not result in ejaculation, the stallion usually rapidly develops another erection and rebreeds the mare promptly.
Figure 118.18 As the stallion dismounts (and is backed away), the mare is vigorously turned inward to protect the stallion in case the mare kicks.
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Figure 118.19 Dismount drippings are collected from the end of the stallion’s penis for examination under a microscope to confirm that ejaculation has occurred.
After breeding, the stallion is backed into an area for rinsing the penis with warm tap water. The stallion is then returned to his stall or paddock. After removing breeding equipment from the mare, she is led from the shed to be transported to the farm of origin. The breeding process is then repeated for the next mare and stallion.
Requirements for imported mares The current code of practice requires that imported mares (mares in the United States less than 1 year) be bred last in line during the appropriate breeding session. After dismounting the mare, the stallion’s penis and sheath must be scrubbed with a 2% chlorhexidine solution, rinsed with tap water, and packed with nitrofurazone ointment. The stallion cannot be bred to another mare, either imported or domestic, that same session.
Summary Often, there is an adversarial relationship between senior management and livestock managers. It is important that the veterinarian responsible for breeding management and health care maintain open communications with senior managers. Although the veterinarian may recognize livestock management deficiencies that need changing, he or she must remain flexible within the constraints provided on the breeding farm. Years may be required to solidify this give-and-take relationship, and opinions on the value of any management procedures will change. The underpinnings of a successful stallion manager are open-mindedness, common sense, well-developed communication skills, and flexibility in deriving solutions to meet each individual stallion’s requirements.
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Chapter 119
Nutrition and Exercise for Breeding Stallions Stephen G. Jackson
Compared with other classes of horses, there has been little research done concerning the nutrient requirements of the stallion and the impact of nutrition on the breeding soundness of the stallion. In the absence of specific research data concerning nutrient requirements and nutritional management of the stallion, correct and effective feeding becomes more of an art than a strict science. Stallions are variable in their response to feeding and many times proper assessment of behavior is as important to meeting nutrient needs as bodyweight and other frequently used parameters for addressing nutrient needs. In the following discussion, research data will be used where possible to define nutrient needs but, in the absence of hard experimental data, management practices and feeding methods proven in the field will be used.
Basics of stallion nutritional management By far the most important factor to consider in feeding the breeding stallion is maintaining appropriate body condition. The biggest error made in feeding the stallion is feeding too much. Many stallions are allowed to become obese or, if not obese, then certainly overweight. This can have a much greater negative impact on behavior, longevity, libido, and other characteristics important to the stallion than any nutrient deficiency. Regulation of energy intake and maintaining appropriate body condition are the most important aspects of feeding and managing the breeding stallion. Stallions should be maintained in a hard, athletic condition and there is really no reason for great seasonal changes in body condition and weight. The most effective tool for the correct assessment of the stallion’s response to feeding and work is a weigh scales. There are a great many good farms
and stallion managers that have years of data on the weight of stallions and that use the weight and body condition of the horse to determine changes that are necessary in feed. Bodyweight is a very important piece of information that accurately describes the horse’s response to feed, number of covers, activity, and age. Once stallions reach maturity, e.g., by 4 or 5 years of age, and are ‘let down’ from their show or race careers, there is no real reason for large fluctuations in bodyweight. Total feed intake of the stallions should be based on the weight and condition of the stallion and on those characteristics that determine weight, condition, and health. There is no information that suggests that stallions that are maintained in a too-fat condition perform better in the breeding shed or have superior seminal characteristics or greater libido, but there is a great deal of information that suggests that fat stallions are more prone to colic, laminitis, behavioral problems, and decreased longevity. Finding the appropriate condition for a horse and making feed and management changes to maintain appropriate condition are far more important than requirements for individual nutrients. Carrying 50 kg of extra fat is going to have a far more deleterious effect on the horse than a deficiency of 10 g of calcium per day.
Exercise programs In addition to monitoring body condition and making appropriate changes in feed intake, mature stallions should be placed on a good exercise program if their skeletal and mental soundness allow. Exercise improves longevity, attitude, and behavior, and reduces health problems most associated with the fat horse. Exercise programs in use on better
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Nutrition and Exercise for Breeding Stallions farms include daily riding, longing, hand walking, use of mechanical horse walkers, swimming, and the use of underwater treadmill systems. The duration and intensity of exercise that is appropriate for each horse should be determined by soundness, body condition, age, historical performance during the breeding season, temperament, and environmental factors such as temperature. The exercise modality used to keep stallions fit is really not the important thing; the important thing is to make sure that the horses get some exercise and that the exercise program is appropriate for the condition and mental characteristics of the horse. Assess the condition of the horse, determine the correct body condition, and go about designing a program to maintain that condition. While it is important that physical activity be part of the stallion management program, there are some data which suggest that too much exercise may negatively impact on seminal characteristics. Taylor et al.1 conducted a small study involving five stallions to determine whether there was an effect of moderate exercise on seminal characteristics in stallions. Horses were exercised in a round pen for a period of 8 weeks and daily sperm output and seminal characteristics were determined. Exercise treatment included 1 minute walking, 2 minutes trotting, and cantering at a 165–180 beats per minute heart rate for 5 minutes, followed by 2 minutes trotting and 30 minutes on a mechanical horse walker. While the horses became fitter, based on recovery heart rate evaluation, there were some negative impacts of exercise on seminal characteristics. Though there tended to be differences between stallions in their response to exercise, there were consistent differences in pre- and post-exercise sperm morphological characteristics. There were no differences observed for pre- and post-exercise values for semen concentration, gel volume, or total sperm number in ejaculates. The results of this study suggest that higher work intensities may have a negative impact on seminal quality, but the changes were inconsistent among stallions.
Nutrient needs Energy The first and most important step in meeting the stallion’s nutrient requirements is to determine the energy required in the diet and the source of that energy. The concentration of all other nutrients in the diet is a function of how much feed is going to be fed. The lower the total daily feed intake, the higher the concentration of other nutrients in the diet must be in order to meet the requirements for those nutrients.
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Table 119.1 shows the energy requirements of mature breeding stallions of different weights in the breeding season, as well in the non-breeding season.2 The digestible energy (DE) requirement of the mature stallion is thought to be slightly higher than the DE requirement of the mature horse at maintenance. The DE requirement of the stallion in the nonbreeding season is derived from the formula: DE (Mcal/day) = 0.0363 × bodyweight (kg) For example, the 600-kg stallion in the non-breeding season would be 0.0363 × 600 = 21.78 Mcal/day. This may be compared with the mature horse at maintenance that requires 16.5 Mcal of DE. This is a good starting place for determining the energy required by the stallion in the period of time when he is not breeding mares. Individual variation in activity, response to feeding, temperature where housed, and age may elevate or decrease the actual amount of calories needed to maintain the horse’s bodyweight at an acceptable level. It is estimated that there is a 20% increase in the DE requirement of the stallion during the breeding season compared with requirements during the non-breeding season.2 Calculation of the DE requirement of the stallion during the breeding season is as follows: DE (Mcal/day) = 0.0363 × bodyweight (kg) × 1.2. Therefore, the mature 600-kg breeding stallion during the breeding season would require 0.0363 × 600 × 1.20 = 26.14 Mcal. This is roughly equivalent to the requirement of the horses at light work intensity. There are some stallions that will require less energy than this because they are not active in the season or are quiet in the stall or paddock. Likewise, horses that are very active in the presence of mares and fret in the stall or paddock or have a nervous disposition may lose weight when fed this amount of energy. Adjustment of energy intake based on the horse’s condition and response to the breeding of mares is critical to maintaining appropriate condition. While this is a good starting point for understanding the energy required by the stallion, there are numerous factors that influence the actual amount of energy horses require to maintain body condition during the breeding season and during the time they are not breeding mares. Siciliano et al.3 conducted a field study on a prominent Kentucky breeding farm and found that the average DE intake of stallions averaging 590 kg during the breeding season was 25 Mcal/day. Variation between stallions in DE intake was most closely correlated to number of covers in the season (Table 119.2).
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Table 119.1 Daily nutrient requirements of stallions of different bodyweights2 Bodyweight in kg∗
Nutrient DE (Mcal) Crude protein (g) Lysine (g) Calcium (g) Phosphorus (g) Magnesium (g) Potassium (g) Sodium (g) Chloride (g) Sulfur (g) Copper (mg) Iodine (mg) Iron (mg) Manganese (mg) Selenium (mg) Zinc (mg) Vitamin A (IU) Vitamin D (IU) Vitamin E (IU) Thiamine (mg) Riboflavin (mg)
200 NB
200 B
400 NB
400 B
500 NB
500 B
600 NB
600 B
900 NB
900 B
7.3 288 12.4 8.0 5.6 3.0 10.0 4.0 16.0 6.0 40 1.4 160 160 0.4 160 6000 1320 200 12 8
8.7 316 13.6 12.0 7.2 3.8 11.4 5.6 18.7 6.0 40 1.4 160 160 0.4 160 9000 1320 320 12 8
14.5 576 24.8 16.0 11.2 6.0 20.0 8.0 32.0 12.0 80.0 2.8 320 320 0.8 320 12 000 2640 400 24 16
17.4 631 27.1 24.0 14.4 7.6 22.8 11.0 37.3 12.0 80.0 2.8 320 320 0.8 320 18 000 2640 640 24 16
18.2 720 31.0 20.0 14.0 7.5 25.0 10.0 40.0 15.0 100 3.5 400 400 1.0 400 22 500 3300 500 30 20
21.8 789 33.9 30.0 18.0 9.5 28.5 13.9 46.6 15.0 100 3.5 400 400 1.0 400 25 000 3300 800 30 20
21.8 864 37.2 24.0 16.8 9.0 30.0 12.0 48.0 18.0 120 4.2 480 480 1.2 480 18 000 3960 600 36 24
26.1 947 40.7 36.0 21.6 11.4 34.2 16.7 56.0 18.0 120 4.2 480 480 1.2 480 27 000 3960 960 36 24
32.7 1296 55.7 36.0 25.2 13.5 45.0 18.0 72.0 27.0 180 6.3 720 720 1.8 720 27 000 5940 900 54 36
39.2 1421 61.1 54.0 32.4 17.1 51.3 25.0 83.9 27.0 180 6.3 720 720 1.8 720 40 500 5940 1440 54 36
∗
Bodyweight of stallions in kilograms. NB, non-breeding season; B, breeding season; DE, digestible energy; g, grams; mg, milligrams; Mcal, megacalories; IU, International units.
Other factors that influence the DE requirement of the stallion are activity, age, temperament, turnout time, and environmental conditions such as temperature, wind, and humidity. The best guide for determining the stallion’s response to energy intake is bodyweight. If the stallion is able to maintain bodyweight, then DE requirements are being met. If stallions lose weight, DE requirements are underestimated and feed intake should be increased; if stallions gain weight, energy requirements are being exceeded. In the absence of a weigh scales, routine assessment of body condition and assignment of a condition score that describes body condition may
be used very effectively. One of the more common condition scoring systems in use is the one developed by Henneke et al.4 This condition scoring system is shown in Table 119.3. There is general agreement that breeding stallions should be maintained in condition score 5 or 6 using the body condition scoring system proposed by Henneke et al.4 Repeatability of condition scores is sometimes not good and it may be just as effective to have only three scores; too thin, too fat, and just right. At any rate, routine measurement of condition and weight is the key to effectively meet the stallion’s DE requirement and to insure that it does not become too fat or too thin.
Table 119.2 Relationship between number of covers and digestible energy (DE) offered stallions on a commercial Thoroughbred farm in Kentucky3 Group number
Group 1
Group 2
Group 3
Group 4
Stallions/group Average total covers/season DE offered/ day (Mcal) Average bodyweight (kg)
7 70 25.4 598
12 94 25.6 587
10 110 26.1 594
4 144 29.9 576
G1, < 80 covers/season; G2, 80–100 covers/season; G3, 101–120 covers/season; G4, > 120 covers/season. DE, digestible energy.
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Table 119.3 Description of individual condition scores for horses4 Score
Description
1 Poor
Animal is extremely emaciated; spinous processes, ribs, tailhead, tuber coxae, and ischii projecting prominently; bone structure of withers, shoulders, and neck easily noticeable; no fatty tissue can be felt
2 Very thin
Animal emaciated; slight fat covering over base of spinous processes; transverse processes of lumbar vertebrae feel rounded; spinous processes, ribs, tailhead, tuber coxae, and ischii prominent; withers, shoulder, and neck structure faintly discernable
3 Thin
Fat build-up halfway on spinous processes; transverse processes cannot be felt; slight fat cover over the ribs; spinous processes and ribs easily discernable; tailhead prominent, but individual vertebrae cannot be identified visually; tuber coxae appear rounded but easily discernable; tuber ischii not distinguishable; withers, shoulder, and neck not obviously thin
4 Moderately thin
Slight ridge along back; faint outline of ribs discernable; tailhead prominence depends on conformation, fat can be felt around it; tuber coxae not discernable; withers, shoulder, and neck not obviously thin
5 Moderate
Back is flat with no crease or ridge; ribs not easily discernable but easily felt; fat around tailhead beginning to feel spongy; withers appear rounded over the spinous processes; shoulder and neck blend smoothly into body
6 Moderately fleshy
May have slight crease down the back; fat over the ribs is spongy; fat around the tailhead is soft; fat beginning to be deposited along the side of the withers, behind the shoulders, and along the sides of the neck
7 Fleshy
May have a crease down the back; individual ribs can be felt, but noticeable filling between ribs with fat; fat around tailhead soft; fat deposited along withers, behind shoulders, and along neck
8 Fat
Crease down the back; difficult to feel the ribs; fat around tailhead very soft; area along withers filled with fat; area behind shoulder filled with fat; noticeable thickening of neck; fat deposited along inner thighs
9 Extremely fat
Obvious crease down the back; patchy fat appearing over the ribs; bulging fat around tailhead, along withers, behind shoulders, and along neck; fat along inner thighs may rub together; flank filled with fat
Meeting energy needs Mature stallions in the breeding and non-breeding season should be capable of meeting their energy requirements with forage alone. Mature horses may consume a minimum of 2% and probably as much as 3.5% of their bodyweight per day in the form of forage dry matter (DM). This means that the 600-kg breeding stallion could consume roughly 12–21 kg of forage dry matter per day. To meet energy needs during the non-breeding season, the forage would need to contain 1.8–1.01 Mcal DE/kg of DM, and to meet energy needs during the breeding season the forage would need to contain 2.17–1.24 Mcal DE/kg of DM. For the purpose of discussion, the expected DE con-
centrations in some common forage types are shown in Table 119.4 along with the expected energy intake from 10 and 18 kg of forage dry matter. From Table 119.4, one would expect the stallion in the non-breeding season to be able to meet DE requirements from 10 kg of the example forages alone. During the breeding season, there would be a slight shortfall in DE intake from warm season hays or forages such as coastal Bermuda and cool-season pasture at 10 kg of forage dry matter per day; however, at 15 kg/day intake, all of the example forages would meet or exceed the digestible energy requirements of the 600-kg breeding stallion. Maximizing the contribution of forage in the feeding program of stallions does much to prevent many of the typical
Table 119.4 Approximate digestible energy concentrations in example forages and digestible energy (DE) intake from 18 and 10 kg/day of forage dry matter (DM)2
Forage Alfalfa, early bloom Alfalfa mid-bloom Pasture, mostly legume Pasture, cool season Coastal Bermuda pasture Coastal Bermuda hay Grass hay, cool season mature Oat hay
DE (Mcal/kg DM)
DE intake (18 kg DM/day)
DE intake (10 kg DM/day)
2.62 2.43 2.71 2.39 1.87 1.87 2.04 2.16
47.2 43.7 48.8 43.0 33.7 33.7 36.7 38.9
26.2 24.3 27.1 23.9 18.7 18.7 20.4 21.6
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problems associated with stallions. On all forage diets, we see significantly fewer cases of colic and laminitis, and basing the diet on forages reduces gastric ulcers and the development of stable vices. Horses are best evolved to handle high-forage, lowgrain diets, and do well when fed in this manner. Even though it is apparent that breeding stallions are able to meet energy requirements from forage alone, this is not the way stallions are typically fed. Most stallion managers are inclined to split DE intake between grain or ‘hard feed’ and forage since they may be unsure of the horse’s ability to consume forage or because the grain mix is being used as a carrier for other nutrients such as protein, vitamins, and minerals. Additionally, fat seems to be a pretty color among many horseman, and many stallions are fed not only to meet nutrient requirements but also to maintain a great deal of bloom and condition that is attractive to prospective breeders. Even so, maximizing forage intake and feeding only that amount of grain necessary to maintain peak condition is the safest, most effective way to feed the stallion.
Protein requirements The protein requirements for the breeding stallion are very little higher than those of the mature horse at maintenance. Protein or, more accurately, amino acids are required to maintain tissues such as muscle, and are components of hormones, enzymes, and most other tissues in the body. By definition, the mature horse needs only that amount of protein that is normally lost in the breakdown of tissues due to use. Most of the growth of muscle and bone is complete in the stallion and as such protein requirements are low when expressed as a percentage of diet or in terms of grams required per day. The most current publication of the National Research Council, Nutrient Requirements of Horses,2 suggests that the requirement for protein of the mature horse at maintenance is best described by the equation: Crude protein required = BW ×1.08 g CP/kg BW/day where CP is crude protein and BW is bodyweight in kilograms. The protein requirement of the breeding stallion during the non-breeding season is best expressed by the equation: Crude protein g/day = 1.44 × BW (kg) For example, a 600-kg breeding stallion would require 864 g of crude protein per day. Adjustments for activity and sweat loss are used to express the crude protein requirements of the breed-
ing stallion during the breeding season. Activity associated with the breeding season is thought to be equivalent to light work and sweat losses are thought to be equivalent to 0.25% of pre-exercise bodyweight. The thought being that, during exercise, additional muscle is synthesized and that synthesis requires nitrogen and that there is nitrogen lost in sweat that must be replaced for the horse to remain in zero nitrogen balance. Whether these two events actually occur in the breeding stallion is subject to some question, and there are few stallions that actually have a significantly increased requirement for protein during the breeding season. In any case, expressed as an increase in protein in the diet, the increase is small. The crude protein requirement of the breeding stallion during the breeding season is thought to be best estimated by the equation: Crude protein (g/day) = 1.44 × BW in kg +(sweat loss × 7.8 × 2/0.79) + 0.089 × BW where BW is weight in kg and sweat loss is 0.0025 × BW. For the 600-kg stallion this would equal: CP (g/day) = 1.44 × 600 + (0.0025(600) ×7.8 × 2/0.79) + 0.089 × 600 = 947 g/day Protein requirements for the 200-, 400-, 500-, 600-, and 900-kg stallion are shown in Table 119.1. There is little known about the specific amino acid requirements of the horse and less about the specific amino acid requirements of the breeding stallion. Most of the research done in amino acid nutrition of the horse has been done with lysine requirements. Lysine is the first limiting amino acid for growth in typical horse diets and, as such, there has been a greater focus on lysine requirements than on the requirements for other amino acids. In the absence of definitive data concerning the lysine requirements of the breeding stallion, the most current NRC2 guidelines express the lysine requirement of the breeding stallion as: Lysine (g/day) = 0.043 × crude protein requirement This would mean that the 600-kg breeding stallion in the non-breeding season would require: 1.44 × 600 × 0.043 g of lysine per day or 37.2 g/day. Similarly, the stallion in the breeding season would require: Lysine (g/day) = crude protein requirements (g) ×0.043 Lysine requirements for the breeding stallion are shown along with crude protein requirements in Table 119.1.
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Nutrition and Exercise for Breeding Stallions There are few practical diets fed to stallions that meet energy requirements but do not meet protein and lysine requirements. Alternatively, there are many cases when diets that meet energy requirements far exceed the protein and lysine requirements of the horse. As an example, if we assume that a 600-kg stallion in the non-breeding season is eating adequate alfalfa (lucerne) hay to meet requirements for energy and that alfalfa hay was 16% protein and 0.56% lysine, the calculations would be as follows. DE required = 21.78 Mcal/day/2.43 Mcal/kg (DE in alfalfa) = 8.96 kg alfalfa/day × 0.16% protein in the alfalfa = 1443 g crude protein per day. The crude protein required is 864 g/day, so the diet of alfalfa provides crude protein is 569 g in excess of the stallion’s requirement. Lysine intake from that diet would be 50 g/day and the horse requires 37.2 g/day. Similar calculations can be done for the breeding stallion during the breeding season and for other types of diets if the protein and lysine concentrations of the diets are known. There are very few instances when this author has evaluated stallion diets and thought that protein and lysine requirements were not fully met. On the other hand, there are practical diets fed to stallions that provide three or four times the requirement for protein with no noticeable problem. This does not mean that protein is a non-issue in the breeding stallion, but it does mean that it is rare that protein is a limiting factor in stallion diets. It is not uncommon for lysine to be a question if grass hay is being fed as the exclusive feed stuff. As such, determination of lysine and protein intake of the stallion is a necessary exercise if one is to be sure that nutrient requirements are being met.
Mineral requirements In practical feeding programs for stallions, there are far more cases when mineral intake is excessive than when there are actual mineral deficiencies. This is due to the way stallions are typically managed and to the fact that, compared with other classes of horses, mineral requirements are low. Many stallion managers supplement with unnecessary vitamin and mineral supplements. These supplemental sources of nutrients are, more times than not, additional to the intake of minerals that are already in excess in the basal diet. Similar to the case for other classes of nutrients, the mineral requirements of the stallion above those required for maintenance are low and there is little activity of the stallion that should result in greatly increased need for minerals over maintenance requirements. Since the breeding stallion’s requirements for nutrients are most like the mature idle horse, most of the requirements of the stallion pub-
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lished in the latest NRC edition2 are very similar to maintenance requirements for mature idle horses of like weights. The maintenance requirement for a mineral is defined as that amount of a specific mineral that is needed to balance daily endogenous mineral loss. The amount of mineral lost is different for each mineral and may also be affected by mineral balance in the diet. The mineral requirements of the breeding stallion in the non-breeding season are thought to be equal to the mineral requirements of the mature idle horse. While the current NRC guidelines2 suggest that mineral requirements of the stallion during the breeding season are marginally higher than that needed for maintenance, there is some question relative to the validity of this assumption. Current mineral allowances above maintenance during the breeding season are low and probably serve as some insurance in case there are greater needs associated with increased activity during the breeding season. It is interesting that in the last four NRC publications there has never been a decrease in the mineral requirements of the horse of any age or use but there have been numerous instances when mineral requirements have been increased. On a practical basis, we seldom see real mineral deficiencies in stallions. Care should be taken in balancing diets for stallions, and excesses of minerals should be avoided. The thought that ‘if a little is good a lot is better’ is a dangerous attitude when considering mineral nutrition. Calcium is the mineral in the greatest abundance in the body and, besides functioning to maintain the integrity of the skeleton, has major roles in nerve and muscle function, maintaining membrane integrity, and in blood clotting, just to name a few of the functions known for calcium. Calcium requirements may be affected by calcium source, concentration of other minerals in the diet, and by the presence of oxalates in the diet. High oxalate concentrations in the diet may significantly reduce the absorption of calcium. Calcium supplementation is inexpensive and most commercial rations are oversupplemented with calcium. In addition, most supplements on the market have additional calcium to make the label read better and the net effect is intake of calcium significantly greater than requirements. The requirement of the stallion for calcium in the non-breeding season is thought to be best described by the equation:2 Calcium requirement (g/day) = 0.04 × BW (kg) This would mean that the calcium requirement of the 600-kg stallion that is not breeding mares is: Calcium (g/day) = 0.04 × 600 = 24 g/day
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This calcium requirement is the same as the requirement of the mature horse at maintenance. It is suggested2 that the calcium requirement of the stallion in the breeding season is marginally higher than for maintenance. This increase in the amount of calcium required is to offset additional needs for work and sweat loss. The equation used by the NRC to express calcium requirements for the breeding stallion is calcium (g/day) = 0.06 × BW or, in the case of the 600-kg stallion, 0.06 × 600 = 36 g of calcium per day. There is little evidence to suggest that the stallion in the breeding season requires more calcium than the nonbreeding stallion but the small increase suggested is probably innocuous and may account for some calcium lost in sweat and also allows a margin for increased bone turnover. One could easily argue that a stallion maintained on predominantly legume hay needs no additional calcium in the grain mix. Stallions that consume 10 kg of alfalfa hay that contains 1.5% calcium would consume 150 g of calcium per day, which is five times the required intake of calcium. Adding additional calcium in the form of calcium-fortified feed or mineral supplements is not necessary. Using a liberal requirement of 36 g/day, calcium intakes seen in practical feeding programs for stallions are generally greater than requirements. The horse is capable of dealing with excesses of calcium and even with calcium marginal diets if the phosphorus intake is not greater than the calcium intake. The rare instances when we may see calcium deficiencies are when a great deal of non-fortified cereal grains, wheat bran, or rice bran are fed with low-quality grass hays or hays that contain a high concentration of oxalates. Tropical forages such as Guinea grass and Setaria contain high oxalate concentrations and horses maintained on these forages have shown a greater propensity for calcium deficiencies that may present as nutritional secondary hyperparathyroidism, also known as ‘big head disease’ or ‘miller’s disease’. In areas of the world where oxalate levels are known to be high, it is particularly important that the calcium status of the ration be evaluated and supplemental calcium be added if there are marginal or deficient calcium concentrations in the diet. Calcium is cheap and supplementation with ground limestone or di-calcium phosphate is the main way in which grain diets may be fortified to meet requirements. Phosphorus requirements of the stallion, like calcium requirements, are very little different than the requirements of the mature horse at maintenance. Rations should be evaluated and care should be taken that total daily phosphorus intake is not greater than the intake of calcium. There tends to be a great deal of discussion about the appropriate calcium–phosphorus ratio in horse diets but there
is little evidence to suggest that ratios as low as 1 part calcium to 1 part phosphorus or as high as 5:1 are a problem for mature horses if the basic requirements for the two mineral are met. Cereal grains such as oats and barley are reasonable sources of phosphorus, and wheat bran, rice bran, or byproducts of the rice and wheat milling industries are very good sources of available phosphorus. Additionally, rock sources of phosphorus such as di-calcium phosphate may be used to fortify diets deficient in phosphorus. The phosphorus requirement of the mature breeding stallion in the non-breeding season is best described by the equation:2 Phosphorus (g/day) = 0.028 × BW where BW is bodyweight in kg. This is the same as the phosphorus requirement of the mature horse at maintenance. Horses during the breeding season are thought to have a slightly elevated requirement and the requirement of the stallion in the breeding season is best described by the equation:2 Phosphorus (g/day) = 0.036 × BW Using these equations, the 600-kg breeding stallion would require 16.8 g of phosphorus in the nonbreeding season and 21.6 g in the breeding season. Salt, i.e., sodium and chloride, should be offered to horses free choice in the paddock and or stall. If adequate water is available the horse will not consume toxic amounts of salt and there is every reason to believe that the horse has some nutritional wisdom with respect to salt intake. Intake of freechoice salt varies with environmental temperature and with sweat loss of the horse, which is a function of temperature and activity. Most commercial horse feeds contain 0.5–1.0% salt. When feeding commercial rations, the actual intake of sodium and chloride varies depending on the intake of feed. Stallions usually do not consume adequate amounts of feed to meet their salt requirements and salt should be provided ad libitum. The sodium requirement (g/day) of the stallion in the breeding season is thought to be: 0.02 × BW + 3.1 × SL, where BW is weight in kg and SL is sweat loss. Sweat loss for light work is 0.0025 × BW.2 Sodium requirements of the stallion in the non-breeding season in grams per day are: 0.02 × BW. The chloride requirement of the breeding stallion (g/day) is 0.08 × BW + 5.3 × SL, where BW is bodyweight in kilograms and SL is sweat loss for light exercise.2 For stallions in the off-season, the chloride requirement (g/day) is: 0.08 × BW. Magnesium is usually adequate in practical forage diets depending on local magnesium concentrations in the herbage and soil. According to the most
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Nutrition and Exercise for Breeding Stallions recent NRC recommendations,2 the magnesium requirement (g/day) for the mature stallion in the nonbreeding season is: 0.015 × BW, and for the breeding stallion is 0.019 × BW. It is rare to see a magnesium deficiency in the mature horse and, most of the time, basal levels of magnesium in the diet are adequate to meet requirements. Magnesium oxide and magnesium sulfate may sometimes be used to supplement diets in areas that may be deficient or borderline deficient in magnesium. Potassium concentrations in forages are characteristically very high. Potassium concentrations range from greater than 1% to over 4% in grass and legume forages respectively. The current NRC guidelines2 estimate the potassium requirement (g/day) of the stallion in the non-breeding season to be: 0.05 × BW, and in the breeding season the requirement (g/day) is: 0.05 × BW + (1.4/0.5) × SL. As with other electrolytes, the increased potassium requirement of the breeding stallion during the breeding season is reflective of losses in the sweat. It is very rare that potassium supplementation is required when horses are maintained on forage diets. For example, if a 600kg stallion was consuming the minimum forage recommended, i.e., 1.5% of BW, and the forage were low in potassium, i.e., 1.5%, the potassium intake would be 600 × 0.015 × 0.015 or 135 g/day. The nominal requirement for potassium for the 600-kg stallion in the non-breeding season would be 30 g/day and, as such, potassium intake from the forage alone would be 105 g over the requirement. The only instance when these high intakes of potassium would pose a problem is if the stallion has hyperkalemic periodic paralysis (HYPP). In horses that have this genetic disorder, it is essential that dietary intake of potassium is limited as much as possible. In practical feeding programs, this means using grass hays that are lower in potassium and minimizing potassium in grain mixes. The HYPP problem is limited to Quarter Horse stallions with the stallion Impressive in their lineage. HYPP has not been identified in other breeds or in other lines of stallions in the Quarter Horse breed. If stallions other than HYPP stallions are on forage diets or if forages represent at least 50% of DE intake, then potassium intake will exceed requirements and further dietary supplementation is not called for. On the other hand, intuitively it makes sense that horses are able to handle very high potassium intakes since this is the definition of a forage-based diet which horses evolved to eat. As is the case for other nutrients, adding supplemental potassium to an already adequate diet makes no good sense. Besides the macrominerals discussed above, stallions have requirements for microminerals or trace minerals. There are established requirements for the trace minerals copper, zinc, manganese, selenium,
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iodine, cobalt, and iron. Some of the requirements now used are based on good data from horses and some are estimates. There are no studies specifically on the trace mineral requirements of the stallion even though the roles of the trace minerals in spermatogenesis and reproduction have been well elucidated. Supplementation with trace minerals over established requirements because there is proven function for that mineral in reproduction is not justified. For example, it is known that selenium is critical to the function of the antioxidant defense system via its role in the selenium-dependent enzyme glutathione peroxidase (GSH-Px). There is a significant amount of GSH-Px in the seminal fluid of bulls but apparently none in the seminal fluid of stallions. Supplementing with selenium in either organic or inorganic form has been shown to increase plasma and red blood cell selenium as well as to increase selenium in the seminal fluid or in the spermatozoa of several species.5 On the other hand there are no data that suggest that supplemental selenium has a positive impact on stallion semen quality. As such, meeting the requirement for selenium is important and selenium may play a crucial role in sperm function, but supplementing with selenium over and above the requirements is not necessary or prudent. The current requirements of the stallion are taken from data on mature horses at maintenance with an allowance for light work during the breeding season. There are no data that exist that would indicate that reproductive performance of the stallion or seminal characteristics would be enhanced by feeding levels over those recommended in the most recent NRC publication.2 Dietary requirements for copper, zinc, cobalt, iron, manganese, and selenium are shown in Table 119.1.
Vitamins In practice, there are far more instances when vitamins are overfed to the stallion than when stallions are deficient of vitamins. This is true for other classes of horses as well, but it seems like the tendency of horsemen to feed excessive vitamins is greater for stallions than for many other classes of horses. Luckily, water-soluble vitamins are generally thought to be excreted very efficiently when fed in excess, and horses have a fairly high tolerance for vitamin A, D, and E before the appearance of toxicity symptoms. Vitamin A in the diet is derived from beta-carotene found in forages and, to a lesser extent, in some grains and from synthetic sources of vitamin A such as retinyl palmitate that might be found in commercial concentrates and supplements that are added to the diet. The basic requirement for vitamin A is 30 IU × BW (kg) for the non-breeding stallion
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and 45 IU × BW(kg) for the stallion actively breeding mares. This would mean the 600-kg non-breeding stallion would require 30 × 600 or 18 000 IU/day, and the 600-kg stallion during the breeding season would require 45 × 600 or 27 000 IU of vitamin A per day. Most commercial supplement pellets contain 20–35 000 IU of vitamin A activity per kilogram and are designed or recommended to be fed at an intake of 1 kg/day. This would mean that a kilogram of these products alone would more than satisfy the stallion’s requirement for vitamin A with no reliance on beta-carotene from the forage component of the diet. Vitamin A equivalent of fresh forage is high and there is little reason to believe that there is additional vitamin A required for horses on pasture that is green and growing. Ralston et al.6 fed stallions a control diet of oats and hay containing no supplemental vitamin A and a treatment diet that provided 132 000 IU of vitamin A as retinyl palmitate. There were no treatment effects between the two diets for seminal characteristics, including gel-free seminal output, gel volume, total volume, percent progressively motile spermatozoa, number of spermatozoa per milliliter, number of spermatozoa per ejaculate, sperm morphology, and semen pH. Furthermore, there was no treatment effect on libido or behavioral characteristics of the stallions. This experiment indicated that, even though supplementation with retinyl palmitate resulted in higher plasma retinol, there was no positive effect on seminal characteristics or behavior that could be attributed to the addition of supplemental vitamin A to a diet that was adequate in beta-carotene to meet basic requirements. Horses on stored forage consisting of hay, hay cubes, or haylage may need additional vitamin A when access to fresh forage is limited but, even then, beta-carotene in the hay may be adequate to meet requirements. Some vitamin A activity in forage is lost during harvesting, storage, and preservation. Grass hays that are cut in late stages of maturity are low in beta-carotene and when hays such as oat hay, timothy, coastal Bermuda, and other grass hays are fed, there may be a need for additional supplementation of the grain mix that is fed. More times than not, when evaluation of diet is performed, vitamin A intake of all classes of horses far exceeds requirements. It is fortunate that intake of beta-carotene from hay or pasture does not contribute to a potential toxicity. The upper safe limit for vitamin A is thought to be 16 000 IU/kg DM/day.7 Vitamin D is involved with the assimilation of calcium and phosphorus in bone and with absorption of calcium and, as such, this vitamin gets a lot of attention in the formulation of rations and supplements. Vitamin D3 is the most commonly used supplemental source of vitamin D used in horse rations, but a far more important source of vitamin D is irradiation
of precursors in the skin via ultraviolet light. It is rare that we would see any symptoms of vitamin D deficiency in stallions and, even more so, in stallions that have exposure to sunlight. It is difficult to argue that the stallion housed outside or that has as little as 4 hours exposure to sunlight would have an additional dietary requirement for vitamin D. The current NRC guidelines2 recommend that horses receive 6.6 IU vitamin D per kilogram bodyweight per day but further state that the true dietary vitamin D requirement for horses exposed to sunlight is not known and that, in most practical feeding and management programs, vitamin D synthesized by irradiation of precursors in the skin should be adequate to meet requirements. In many cases, the concentration of vitamin D in commercial feeds is 800–1400 IU vitamin D/kg and supplements intended to be fed at 1 kg/day may contain 4000 IU of vitamin D/kg bodyweight. Assuming a requirement of 6.6 IU/kg bodyweight, the 600kg stallion would require roughly 3960 IU of vitamin D per day. This means that a kilogram per day of a ‘supplement pellet’ containing 4000 IU of vitamin D would exceed the requirement for vitamin D with no allowance for synthesis of vitamin D in the skin. Likewise, if a stallion were eating 3 kg of a feed containing 800 IU of vitamin D, vitamin D intake would be 2400 IU of vitamin D from the feed alone and would allow for some vitamin D synthesis and a modest level of vitamin D from hay. The upper safe level of dietary vitamin D is thought to be 44 IU vitamin D per kilogram bodyweight per day. For the 600-kg stallion this would be 26 000 IU of vitamin D per day. It is not common to see vitamin toxicity in horse diets but, if there is a chance of having toxic levels of intake of vitamins, the chance is greatest for vitamin D. Basic calculation of dietary vitamin D intake is essential. Vitamin D intake should not only be compared with the requirement but also with the upper safe level. Vitamin E is a potent biological antioxidant and, in recent times, has received a great deal of attention in human as well as in animal nutrition. The NRC publication for nutrient requirements of horses has recommended significant increases in the allowances for vitamin E in the last two editions published.2, 8 Vitamin E in equine diets is present in the form of naturally occurring tocopherols and may be supplemented as alpha-tocopheryl acetate. Natural vitamin E in feeds is subject to oxidative loss during storage so many commercially available diets are supplemented with vitamin E. The current NRC guidelines2 suggest a vitamin E allowance of 1 IU of vitamin E per kilogram of bodyweight per day for the stallion not breeding mares and 1.6 IU per kilogram bodyweight per day of vitamin E for the stallion in the breeding season.
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Nutrition and Exercise for Breeding Stallions There are few studies that directly measure the effect of vitamin E on stallions, but Rich et al.9 conducted an experiment with 20 stallions designed to measure the effect of vitamin E supplementation on seminal characteristics and sexual behavior of stallions. Stallions were split into two treatment groups: control (no supplemental vitamin E) and 5000 IU of supplemental vitamin E. The treatment group received supplemental vitamin E for 15 weeks. There were no differences in any seminal characteristics measured between the treatment and control group and there was no treatment effect on stallion sexual behavior. Unfortunately, the vitamin E content of the basal diet was not published. Authors did say that the basal control diet met nutrient requirements and, as such, this study only suggests that vitamin E supplementation over and above requirements does not have a positive impact on sexual behavior or seminal characteristics. Current recommendations for vitamin E are liberal and there is no reason to believe that additional vitamin E is necessary for the stallion if basal requirements are met. There are few published data dealing with the requirements of the horse for water-soluble vitamins and still less for the specific requirements of the stallion for the B vitamins and for vitamin C. Microbial synthesis of most B vitamins in the cecum is thought to be adequate to meet the B vitamin requirements of most classes of horses. There are no studies that have shown B vitamin deficiencies in the horse when they are fed practical diets, suggesting that, when taken together, B vitamin synthesis in the cecum and B vitamin concentrations in naturally occurring feedstuffs meet the basic requirements for the B vitamins for most classes of horses. The latest NRC guidelines2 do not list specific requirements for B vitamins for the stallion in the breeding and non-breeding season since there are no experiments that would suggest that stallions have an increased requirement for B vitamins over the requirements for maintenance. Table 119.1 shows the current recommendations for B vitamins for the horse for which requirements have been established and there is no reason to believe that these allowances are not more than adequate for all classes of horses. There are still no established requirements for most of the B vitamins or for vitamin C and there is every reason to believe that cecal and tissue synthesis of the water-soluble vitamins is adequate to meet requirements. Many commercially available diets are supplemented with higher levels of B vitamins than the basic requirements and it is highly unlikely that there would ever be a B vitamin deficiency for horses on anything but a purified diet or when horses are exposed to feeds or forages such as moldy sweet clover (vitamin K) or bracken fern (thiamin) that interfere with absorption or utilization
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of dietary sources of these vitamins. The B vitamins tend to be non-toxic at levels that might be fed to a horse but are expensive and not necessary.
Other nutrients or feed additives Feeding fat to all classes of horses has been popular over the last 20 years. Fats and oils are very readily utilized sources of calories and also a source of essential fatty acids. Most studies that address the utilization of fats or oils in horse diets have been conducted on horses in training or to establish the digestibility of various fat sources. Vegetable oils including rice, corn, soy, canola, and linseed oil, to name a few, have been utilized in the feeding of horses for 100 years. Vegetable oils are easily digested by the horse. Apparent digestibility of vegetable oils may range from 88% to 94% and, as such, they are readily utilized as a source of energy. Animal fats such as lard and tallow and choice white grease have been fed to horses but are utilized more poorly than are vegetable oils. There is little animal fat utilized in horse diets due to consumer reluctance to use these fat sources and to worries related to bovine spongiform encephalopathy (BSE) and, as such, these fat sources are not of real practical consideration. In addition, most animal fats are highly saturated and do not address essential fatty acid needs. Fat may be fed to the horse at levels exceeding 10% of the concentrate diet with no negative effects on the horse or on the digestion of other nutrients in the diet. When the fat content of a diet is over 6%, the fat is usually added as a source of calories to maintain or improve energy balance. Diets containing 3–4% fat are feeds that usually are not fat supplemented as many feed ingredients such as oats have 3–4% fat. Addition of fat at lower levels, 50–100 mL either in the feed or as a top-dress, is usually done to address the fatty acid requirements of the horse or to enhance hair quality. The stallion does not react to fat-supplemented diets any differently than do other classes of horses and, as such, supplemental fat can be very effectively used as a source of energy or to replace starch calories in the diet if horses are starch sensitive. There has been a great deal of interest recently in the use of omega-3 fatty acids in the diets of humans and horses. While there are some omega-3 fatty acids in corn oil and soy oil, the best plant source of omega-3 fatty acids is linseed oil or raw or boiled flax. Oil from cold water fishes is also a source of omega-3 fatty acids docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA). These two long-chain omega-3 fatty acids have been the subject of previous interest in enhancing boar fertility and are currently used in some supplements for breeding
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stallions. There have been several studies done with the stallion that suggest that dietary omega-3 fatty acids and especially DHA and EPA from marine sources may have positive effects on the seminal characteristics. Brinsko et al.10 have reported an improvement in motion characteristics of cooled and frozen–thawed semen from stallions fed a ‘DHA (docosahexaenoic)-rich’ nutriceutical. In the experiment, there was no effect on motion characteristics in fresh semen due to treatment. This product was a marine fatty acid source high in omega-3 fatty acids as well as antioxidants. Harris et al.11 fed stallions a similar fatty acid supplement at 550 g/day and determined stallion seminal characteristics in supplemented and unsupplemented stallions. They reported that there was an increase in the seminal membrane concentration of long-chain omega-3 fatty acids and there was an increase in daily spermatozoa output, but there was no impact of the supplemented fatty acids on seminal motion characteristics. At this point, it appears that there may be some benefit in adding omega-3 fatty acid sources to the feed of stallions if they are going to collected and semen is going to be cooled or frozen but there is a lack of evidence that suggests that seminal characteristics would be improved in horses that are used in natural-cover programs. Further research is needed before the recommendation that stallions be fed an omega-3 enriched supplement can be validated.
Feeding management Nutritional management of the stallion may be as important as insuring that specific nutrient requirements are met. Stallions should receive a minimum of 1.5% of their bodyweight per day in the form of fibrous feed. This feed may take the form of fresh pasture, stored forage in the form of long hay or haylage, or even forage cubes. Long hay or pasture provided to the horse will result in fewer diseases of the gastrointestinal tract than when the diet consists of predominantly concentrate feeds. Maximizing the contribution of forage in the diet and minimizing grain intake will make control of bodyweight easier because forages are nearly always lower in energy density than concentrates. The old adage of ‘little and often’ applied to feed further reduces chances of colic and laminitis because there is less of a starch challenge to the horse in any given meal. There are many farms where 3.5 kg (7.7 lbs) is the established working maximum concentrate feed intake per day and variation from this intake requires a special dispensation from the consulting nutritionist. Using a mean energy density for concentrates of 3.0 Mcal/kg this would
mean that DE intake from grain would be roughly 10.5 Mcal/day, which would represent roughly 40–50% of the stallion’s DE requirement depending on its bodyweight and if it was actively breeding mares or in the off-season. Using a working maximum grain intake encourages the use of higher quality forage if appropriate body condition and weight is to be maintained on a minimal concentrate feed intake. Split into two feedings per day this amount of concentrate does not generally exceed the ability of the horse to digest starch pre-cecally and reduces significantly the amount of starch that may enter the hindgut and be fermented. This is easier to achieve in areas of the world where there is high-quality hay and pasture available. Cecal impaction appears to be one of the predominant gut problems of the stallion. The higher the digestibility and more immature the forage, the lower the lignin content and the less the potential for cecal impaction. Using hays cut in early stages of maturity with lower lignin and higher digestibility results in fewer cases of impaction colic and should be considered when selecting hays for stallions. This is especially true for stallions that are confined to stalls and that receive little or no fresh forage. Selection of the species of forage fed as stored hay tends to be somewhat dictated by regional availability. Stallions do well on legume (e.g., alfalfa, clover, lespedeza), grass (e.g., timothy, orchard grass, Italian ryegrass, coastal Bermuda, and other Bermuda grass cultivars) and on mixtures of grasses and legumes. Because legume hays are higher in energy density, they are more appropriate when trying to use hay as the main source of calories or to induce weight gain without using much grain, and grass hays are easier to use when trying to keep a stallion from becoming too fat. Comments about quality and maturity apply to both types of hay as well as to mixtures of grasses and legumes. Physical form of the feed used for feeding stallions tends to be more of a human concern than an animal concern. Stallions do equally well on straight cereal grains such as oats and barley with appropriate mineral supplementation as they do on textured (sweet) feed or pelleted feeds. Stallions have also been fed and do well on extruded feeds. Extruded feeds may be particularly useful for stallions that are sensitive to starch in the cecum as the process of extrusion gelatinizes much of the starch in the feed making the starch more available pre-cecally. From a practical perspective, selection of the type of feed is more a question of the comfort level of an individual stallion manager than that of a stallion. Notwithstanding, the main consideration in feeding stallions is to keep them in a hard, athletic condition which will increase longevity and health. Remembering that most nutrient requirements of the
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Nutrition and Exercise for Breeding Stallions breeding stallion are no greater than the mature horse at maintenance will go a long way in making sure that stallions are in optimum condition and will also decrease nutrition-related problems.
References 1. Taylor MJ, Evans JW, Householder DD, Potter GD, Varner DD. Reproductive parameters of breeding stallions in response to a moderate physical conditioning program. In: Proceedings of the 15th Equine Nutrition and Physiology Symposium, 1997, pp. 104–8. 2. National Research Council. Nutrient Requirements of Horses, 6th edn. Washington, DC: National Academy Press, 2007. 3. Siciliano PD, Wood CH, Lawrence LM, Duren SD. Utilization of a field study to evaluate digestible energy requirements of breeding stallions. In: Proceedings of the 13th Equine Nutrition and Physiology Society Symposium, 1993, pp. 293–8. 4. Henneke DR, Potter GD, Kreider JL, Yates BF. Relationship between condition score, physical measurements and body fat percentage in mares. Equine Vet J 1983;15:371–2. 5. Saaranen M, Suistomaa M, Vanha-Perttula T. Semen selenium content and mitochondrial volume in human and some animal species. Hum Reprod 1989;4:504–8.
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6. Ralston SL, Jackson SA, Rich VA, Squires EL. Effect of vitamin A supplementation on the seminal characteristics and sexual behavior of stallions. In: Proceedings of the 9th Equine Nutrition and Physiology Symposium, 1985, pp. 74–7. 7. National Research Council. Vitamin Tolerance of Animals. Washington, DC: National Academy Press, 1987. 8. National Research Council. Nutrient Requirements of Horses, 5th edn. Washington, DC: National Academy Press, 1989. 9. Rich GA, McGlothlin DE, Lewis LD, Squires EL, Pickett BW. Effect of vitamin E supplementation on stallion seminal characteristics and sexual behavior. In: Proceedings of the 8th Equine Nutrition and Physiology Symposium, 1983, pp. 83–9. 10. Brinsko SP, Varner DD, Love CC, Blanchard TL, Day BC, Wilson ME. Effect of feeding a DHA-enriched nutriceutical on motion characteristics of cooled and frozen stallion semen. In: Proceedings of the 49th Annual Convention of the American Association of Equine Practitioners, 2003, pp. 350–52. 11. Harris M, Anderson C, Webel S, Godbee R, Sanders S, Schurg W, Baumgard L, Arns M. Effect of feeding an omega-3 rich supplement on the fatty acid composition and motion characteristics of stallion spermatozoa. In: Proceedings of the 19th Equine Science Society Symposium, 2005, p. 239.
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Chapter 120
Management of Subfertile Stallions Under Natural-Mating Conditions Terry L. Blanchard
Economic impact of lowered fertility A stallion that achieves suboptimal pregnancy rates greatly increases the cost of producing foals for mare owners and farm management. The increased costs associated with a stallion’s lowered fertility arise from: (i) increased mare expenses (e.g., extra covers, extra transport of mares to breeding sheds, extra veterinary examinations/treatments, and additional boarding fees required for mares left at a breeding farm/facility over repeated estrous cycles); (ii) wasted maintenance costs associated with support of mares that do not produce foals; (iii) decreased income (e.g., from sale of penalized late-born foals that arise from mares not becoming pregnant early in the year, lost income from failure to produce a foal, and lost income from non-productive stud fees), and (iv) labor associated with rebreeding mares and potentially lost service opportunities for the stallion that could have been breeding another mare. Thus, economic impact associated with breeding of a stallion with lowered fertility can be substantial. The following example is used to further illustrate the magnitude of losses that can occur with just a 20% difference in pregnancy rate (Table 120.1). If a stallion bred a book of 100 mares (using an estimate of 1.1 covers per cycle; a conservative 10% double rate) and achieved a 60% pregnancy rate per cycle over a total of three estrous cycles, the stallion would achieve a 93% seasonal pregnancy rate, requiring a total of 172 covers. If 85% of pregnancies (assuming a 15% embryonic and fetal loss rate) resulted in live foals, 79 foals would be produced. By contrast, if only a 40%
pregnancy rate per cycle was achieved, the stallion would achieve a 78% seasonal pregnancy rate requiring a total of 216 covers. If 85% of pregnancies resulted in live foals, 66 foals would be produced. The lower fertility would culminate in an extra 44 covers (40 estrous cycles × 1.1 covers/cycle) requiring board and veterinary expense over the course of the breeding season – yet produce 13 fewer foals and an additional 13 years of non-productive maintenance expense (13 additional barren mares). If the boarding fee at the breeding farm is $26/day, the 40 extra estrous cycles of breeding (21 days/cycle) result in an extra cost of $21 840 over that for the higher level of fertility. If veterinary fees average $250 for examinations and treatments per cycle, the 40 extra estrous cycles of breeding result in increased veterinary fees of $10 000 over that for the higher level of fertility. If transport fees for the mare to the breeding shed are $150 per round trip, the cost of transport fees for 44 extra trips totals $6600. The added cost of the lower fertility in this example totals $38 440, yet does not include lost income from 13 stud fees, the maintenance expenses for 13 extra barren mares for 1 year, or lost income from 13 foals that will not be produced or labor costs associated with the farm where the stallion is, or even the lost opportunity in breeding another mare when the stallion is heavily booked and may require separation of services to breed three times or more per day. While expenses, fees, and sales prices vary, this example serves to stress the importance of maximizing reproductive efficiency of the stallion.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Management of Subfertile Stallions Under Natural-Mating Conditions
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Table 120.1 Influence of two (40%, 60%) theoretical pregnancy rates per cycle (PR/cycle) on number of covers (no. covers) required to complete three estrous cycles of breeding and seasonal pregnancy rate (SPR) for a stallion mated to 100 mares by natural cover. Assuming 1.1 covers (i.e., 10% ‘double’ rate) are required per estrus, and 85% of pregnancies result in production of viable foals (i.e., 15% pregnancy loss rate), the lower level of fertility would result in 44 extra covers (40 extra cycles) throughout the season, yet produce 13 fewer foals (i.e., 13 more barren mares). Fertility achieved per cycle and per season with theoretical 40% and 60% PR/cycle No. mares bred per cycle × theoretical PR/cycle × covers/cycle = No. mares pregnant Lower theoretical fertility 100 mares bred 1st cycle × 40% PR/cycle × 1.1 covers/cycle = 40 mares pregnant on 1st cycle 60 mares bred 2nd cycle × 40% PR/cycle × 1.1 covers/cycle = 24 mares pregnant on 2nd cycle 36 mares bred 3rd cycle × 40% PR/cycle × 1.1 covers/cycle = 14 mares pregnant on 3rd cycle Total mares pregnant after 3 cycles of breeding = 78 No. foals produced = 78 – (78 × 15% loss rate) = 66 No. barren mares = 34 mares Higher theoretical fertility 100 mares bred 1st cycle × 60% PR/cycle × 1.1 covers/cycle = 60 mares pregnant on 1st cycle 40 mares bred 2nd cycle × 60% PR/cycle × 1.1 covers/cycle = 24 mares pregnant on 2nd cycle 16 mares bred 3rd cycle × 60% PR/cycle × 1.1 covers/cycle = 9 mares pregnant on 3rd cycle Total mares pregnant after 3 cycles of breeding = 93 No. foals produced = 93 – (93 × 15% loss rate) = 79 No. barren mares = 21
Some factors that affect stallion fertility Achievement of suboptimal pregnancy rates remains a serious problem within the horse industry. When performing breeding soundness examinations on stallions intended for stud service, over one-third of prospective breeding stallions fail the evaluation – most commonly due to ejaculation of low numbers of normal, motile sperm.1, 2 While there are certainly many other causes that contribute to lowered fertility of stallions, it is common for this finding to be present in stallions with lowered fertility. Unfortunately, the underlying cause for ejaculation of low numbers of normal, motile sperm by stallions is oftentimes undetermined and, thus, is often medically untreatable.3–5 Alterations in the breeding management of stallions that ejaculate low numbers of normal, motile sperm is therefore usually directed primarily at limiting the number of matings required of the stallion (i.e., limiting the number of mares booked to a stallion in a season, and the number of matings per day or week) to maximize the number of normal, motile sperm that are ejaculated with each breeding.4, 6 The veterinarian must remain cognizant that there are many factors that contribute to the overall fertility of a stallion, including inherent fertility of the stallion, inherent fertility of the mares bred by the stallion, and quality of management (e.g., nutrition and body condition, teasing and breeding management, level of veterinary care). Each contributing factor is capable of severely constraining the percentage and number of offspring produced each year by a given stallion. One should therefore not assume that lowered pregnancy rates must be due to a problem with
No. covers 110 covers 66 covers 36 covers Total no. covers for season = 216 covers
110 covers 44 covers 18 covers Total no. covers for season = 156 covers
stallion fertility unless record analyses and examination findings, along with a thorough evaluation of mating practices, support this conclusion. To this end it is often useful to examine fertility from stallions on the same breeding farm during the same time period. Investigation of suboptimal fertility achieved by a stallion should be directed toward identifying and correcting contributing factors. In some cases, treatment of a disease condition (e.g., infection, ejaculatory dysfunction) may improve the stallion’s fertility.7 In other cases, no treatment is indicated for the stallion; yet, reduced reproductive quality of the mare book will preclude significant improvement in fertility. More commonly, recommendations for altering breeding management practices can be identified that might improve the stallion’s fertility. The following discussion is provided to aid the veterinarian firstly in identifying some of the common factors contributing to lowered fertility of a given stallion and, secondly, to aid the veterinarian in developing a breeding management plan that could maximize the fertility of that stallion.
Historical considerations To assess potential for changes in reproductive management in an effort to enhance fertility of a given stallion, it is important to first obtain a meaningful history. An assessment of past breeding performance (assuming the stallion has been bred previously) is a good place to start, since previous fertility is one of the best predictors of future fertility.8 The number of seasons the stallion has been in service, and if/when any changes in fertility occurred, provide useful
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Stallion: Management
Table 120.2 Influence of declining quality of mare book on fertility achieved by a Thoroughbred stallion in two consecutive years. Seasonal pregnancy rate (PR/season) declined from 86% to 75%, and overall pregnancy rate/cycle (PR/cycle) declined from 66% to 57%. The lower than expected seasonal pregnancy rate in the second year was most likely due to two problems that are apparent from examining the records: (i) low fertility in the barren mare group in year 2, which constituted one-fourth of his book. Note that the PR/cycle was quite good in other classes of mares in year 2, and comparable in foaling and maiden mares (typically the most fertile classes of mares) among both years; and (ii) insufficient opportunity to rebreed mares that did not become pregnant. In this regard, note that only 53 cycles were used to cover the total book of 40 mares in year 2. The PR/cycle among other (than barren) classes of mares in year 2 varied from 67% to 73%; thus, if more opportunities had existed for covering those mares not becoming pregnant on the first cycle in those groups, seasonal pregnancy rate might not have been so low Year
Class of mare
1
Barren Foaling Maiden Not bred Slipped Total Barren Foaling Maiden Not bred Slipped Total
2
No. bred
No. pregnant
No. cycles
PR/cycle
PR/season
6 49 4 3 9 71 10 16 9 3 2 40
6 42 4 3 6 61 5 13 8 2 2 30
9 59 5 4 15 92 18 18 11 3 3 53
67% 71% 80% 75% 40% 66% 28% 72% 73% 67% 67% 57%
100% 86% 100% 100% 67% 86% 50% 81% 89% 67% 100% 75%
historical information. Congenital problems that result in poor fertility are often evident from the first season the stallion is used for breeding, while acquired problems (trauma, deterioration in physical condition, occurrence of disease(s), or management changes) often result in a decline in fertility when compared with previous seasons, or months within a given season. Age-related decline in a stallion’s fertility is often gradual in onset, although the difference in pregnancy rates achieved from one year to the next can be dramatic if due to progressive testicular degeneration.9 Owners or managers of stallions may only provide impressions based on a few salient observations (e.g., a low seasonal pregnancy rate), which are usually insufficient to support meaningful conclusions about fertility. It is therefore incumbent on the veterinarian to thoroughly evaluate breeding records to characterize the magnitude of the fertility problem. If breeding records do not exist, meaningful evaluation can seldom be performed. Poorly organized records can make the task arduous or even impossible. Even the more modern computerized record systems will require some reorganization and summarizing of salient fertility end points to ensure a given stallion’s fertility is adequately characterized.10 In addition, evaluation of fertility of other stallions bred to mares on the same farm in the same time period may help to identify that the stallion in question is not keeping up on a pregnancy per cycle basis compared with his peers. Do not hesitate to request records from previous years to substantiate whether a given stallion’s fertil-
ity has changed, and if a decline in fertility occurred as a result of deteriorating quality of the mare book. It is common for first-year stallions to have a demanding book, with many high-quality mares. After the first or second season, and until/unless the stallion’s offspring prove themselves, it becomes more difficult to fill his book. Thus, as the stallion’s popularity declines, his book tends to accumulate fewer young, fertile mares and an increasing proportion of mares with lower fertility (Table 120.2). This changing quality of the mares in his book many times will explain a decline in fertility.10 The number of mares bred by the stallion in each season is another important consideration in assessing potential causes of lowered fertility. Pickett et al.6 estimated a minimum of 40 mares must be bred to be able to adequately evaluate the stallion’s fertility. Many stallions do not breed a sufficient number of mares to adequately test their inherent fertility,11 so the veterinarian should be cognizant of this fact before arriving at a conclusion that a stallion breeding a limited number of mares does, in fact, have lowered fertility (Table 120.3). Regardless of the stallion’s fertility level, to achieve a high seasonal pregnancy rate, it is important for the stallion to have access to mares early in the season and for non-pregnant mares to be returned for service in a timely manner (i.e., so that return service opportunities are not missed10 ). Few breeding operations consist entirely of highly fertile mares that will likely become pregnant on one service. As a stallion’s popularity declines, his stud fee usually follows. With less at stake, many mare owners
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Management of Subfertile Stallions Under Natural-Mating Conditions Table 120.3 Fertility results from a 22-year-old Thoroughbred stallion with a book of 27 mares. The book of mares is small, limiting conclusions that can be made about the stallion’s inherent fertility. While the overall pregnancy rate (PR) per cycle was only 53%, his first cycle pregnancy rate achieved in foaling mares was 73% (14/19). Of the rest of his book of mares, only one (slipped) mare became pregnant on the first cover. This finding suggests that, if the stallion had a larger book of mares, of which more were known to be fertile (e.g., foaling mares), he could achieve better overall pregnancy rates per cycle and per season Class of mare
PR/season
PR/cycle
Barren Foaling Slipped All classes
2/3 (67%) 19/22 (86%) 2/2 (100%) 23/27 (85%)
2/7 (29%) 19/32 (59%) 2/4 (50%) 23/43 (53%)
are likely to take fewer precautions to ensure their mare is in good pre-breeding condition, is having regular estrous cycles with no reproductive problems, is bred at the optimal time during estrus, has good pre- and post-breeding veterinary care to optimize opportunity for pregnancy to be established, and has early pregnancy diagnosis performed to ensure that mares failing to become pregnant are returned for service promptly at their next estrus. When a stallion’s book of mares is afflicted with too many poorly managed mares, record analyses will reveal increased incidence of non-lactating (maiden, barren, slipped, and not-bred) mares that were being bred only once, often quite late in the season. Additionally, breeding intervals between services for mares failing to become pregnant may be extended beyond the normal 3-week (18–24 days) interovulation interval. By contrast, a popular stallion with a high stud fee will typ-
Distribution of Seasonal Covers by Month 30 Percentage of Covers
c120
25 20 15 10 5 0 Feb
Mar
Apr
May
Jun
Jul
Month Figure 120.1 Percentage of all covers by month of service. Represents data from 16 080 covers from 107 breeding seasons utilizing 29 different Thoroughbred stallions. Note that 60% of covers occur by the end of April, and 86% of covers occur by the end of May.
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Table 120.4 Influence of mating to ovulation interval for 902 estrous cycles of 414 Thoroughbred mares bred to one of 110 stallions Interval from mating to ovulation 1 day 2 days 3 days ≥ 4 days
Pregnancy rate/cycle 61.9% (206/333) 63.8% (326/511) 34.0% (16/47) 36.4% (4/11)
ically breed the majority of the non-lactating mares for the first time early in the breeding season (Figure 120.1);11 thus, having ample opportunity to get each of these mares pregnant if mating on the first estrus of the season is unsuccessful. Additionally, the breeding intervals for the majority of these mares failing to become pregnant on one service will usually experience a normal 18- to 24-day interovulation interval. Finally, good veterinary care will ensure mares are bred near to time of ovulation, thus resulting in a low percentage of mares in the book requiring multiple covers or inseminations per estrus. If the number of covers or inseminations per estrus is high, poor reproductive management (e.g., faulty teasing program, infrequent examinations to reliably predict optimal time for mating, failure to use ovulationinducing drugs to ensure ovulation near to time of mating and reduce number of repeat covers) should be suspected as a problem contributing to lowered fertility of a stallion (Table 120.4).
Mating frequency One question that often arises is whether diminishment of mating frequency (i.e., reducing the number of covers) would improve fertility of a stallion achieving suboptimal fertility. Because the influence of mating frequency on fertility achieved is a highly stallion-dependent phenomenon, no firm answer can be given without evaluation of the outcome achieved from a sufficient number of matings performed at different frequencies. Successful breeding seasons, in spite of the dramatic increase in Thoroughbred mare book sizes in the last decade, leads one to the conclusion that most stallions with normal testes size and semen quality can mate mares several times each day (assuming libido and mating ability are good) and still maintain good fertility (Figure 120.2 and Table 120.5). However, comparing pregnancy rates among days with differing numbers of covers may reveal whether breeding frequency alters fertility of a given stallion (i.e., provides justification for limiting the number of covers per day to maximize the opportunity for pregnancy establishment) (Table 120.6).
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Stallion: Management Book Size vs Total Number of Covers 350
300
250
Total Covers
c120
200
150
100 Total covers = 8.468 + (1.582 * No. mares) N = 107.000 R = 0.983 Rsqr = 0.967 Adj Rsqr = 0.966
50
0 0
25
50
75
100 125 Book Size
150
175
200
225
95% Confidence Interval Regression 95% Confidence Interval Population Figure 120.2 The number of covers required for a given book size is highly variable (being dependent on management level, size of the book, and fertility of both the stallion and the mares in his book). Data obtained from 107 stallion seasons.
General management strategies for stallions achieving suboptimal pregnancy rates Good breeding management practices that should be common among all breeding farms become critical when managing a stallion that achieves suboptimal pregnancy rates if a substantial number of mares are going to be bred. Mares should be in good body condition, free of genital infections, and have regular estrous cycles early in the breeding season so that nonlactating and early foaling mares can have opportunities to become pregnant early in the season, thus lessening the mating load later in the season when the majority of foaling mares are ready to breed. Frequent examinations (at least every other day initially, and once daily as the mare nears ovulation) and the use of ovulation-inducing hormones are essential in order to ensure breedings are not wasted, and thus maximize the number of sperm that are available for each mare covered. The goal is to breed each mare as near to the time of ovulation as possible. Post-breeding
examination should be performed no later than the day following mating to determine whether postbreeding endometritis/fluid accumulation is present and, if so, that it is promptly addressed. Additionally, ovulation date should be recorded, so that record review at a later date can be used to determine whether mares were more likely to get pregnant when they ovulated sooner after breeding (e.g., 1 day, 2 days, etc.). Management strategies sometimes employed with natural-service mating programs involving stallions experiencing lowered fertility include infusion of semen extender into the uterus of the mare immediately prior to cover, reinforcement breeding (infusion of an extended semen sample collected as drippings from the stallion’s urethra after dismount immediately following cover, or immediate aspiration of semen from the mare’s vagina after cover followed by infusion of this sample into the uterus), monitoring dismount samples for the presence of motile sperm and inflammatory cells to identify mares requiring an immediate second cover or post-breeding treatment, and purposeful multiple-mating (doubling, tripling) of mares.
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Management of Subfertile Stallions Under Natural-Mating Conditions Table 120.5 Number of days in season (includes doubles) with 0–5 covers (matings) that occur with varying book sizes (number of mares bred by stallion in a season). Data represent 60 080 covers by 29 stallions over a total of 107 stallion seasons Days in season with 0–5 covers Book size
N
0
1
2
3
4
5
0–19 20–39 40–49 50–59 60–69 70–79 80–89 90–99 100–109 110–119 120–129 130–139 140–149 150–159 160–169 170–179 180–189 190–199
4 19 9 9 8 6 4 4 5 7 5 4 5 2 4 2 6 4
122 92 76 62 58 49 35 31 30 35 29 25 24 19 17 27 13 15
14 36 45 52 50 50 55 59 53 43 45 40 44 36 32 29 30 27
0 7 13 20 24 27 36 36 42 36 40 43 41 38 37 37 35 37
0 1 2 2 5 9 8 11 10 20 19 22 21 26 29 31 33 33
0 0 0 0 0 0 1 0 1 3 5 7 7 17 22 13 25 23
0 0 0 0 0 0 0 0 0 0 0 0 1 2 0 1 2 2
Table 120.6 Influence of size of mare book and frequency of mating on fertility (pregnancy rate per cycle; PR/cycle) achieved by four Thoroughbred stallions during one breeding season. Stallion 3 appeared to suffer a decline in pregnancy rates on the third and fourth cover of the day. Stallion 4 (with a smaller book) appeared to suffer a decline in pregnancy rates when more than two mares were covered per day, and would benefit from limiting the number of covers per day to a maximum of three mares. High mating frequencies for stallions 1 and 2 (with larger books) did not appear to lower pregnancy rates to a level that would suggest they would benefit by constraining breeding frequency Stallion
Book size Seasonal PR Overall PR/cycle PR/cycle on days with 1 cover PR/cycle on days with 2 covers PR/cycle on days with 3 covers PR/cycle on days with 4 covers PR/cycle on 1st cover of day PR/cycle on 2nd cover of day PR/cycle on 3rd cover of day PR/cycle on 4th cover of day
1
2
3
4
184 95% 64% 80% 60% 67% 72% 72% 70% 60% 65%
170 95% 67% 69% 66% 68% 69% 75% 60% 60% 80%
114 85% 53% 61% 65% 47% 54% 60% 62% 41% 40%
88 80% 49% 48% 65% 44% 41% 49% 56% 44% 20%
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Monitoring of dismount semen samples Monitoring of dismount semen samples (collected as drippings from the stallion’s urethra and appropriately diluted in a pre-warmed semen extender) for the presence of motile sperm is a technique used by management personnel to confirm that ejaculation indeed occurred during a given cover. While it is rare for a stallion to exhibit behavioral and physiological signs of ejaculation yet fail to ejaculate, microscopic examination of the dismount semen sample will reveal few or no sperm present if ejaculation did not occur. Rarely, stallions may have retrograde ejaculation into the bladder.12 In most instances, stallions that did not ejaculate will promptly achieve another erection and cover the mare again within a few minutes. Repeating the mating sequence in these situations will thus improve the chance that the mare will become pregnant. The procedure is also useful for identifying incomplete ejaculations that some stallions occasionally experience. Usually, the stallion that only occasionally experiences incomplete ejaculation will promptly achieve another erection and cover the mare again. Routine examination of dismount samples is also useful as a technique for screening changing semen quality over the course of the season. If sperm concentration, motility, and morphology in dismount semen samples have consistently declined, breeding soundness evaluation of the stallion may be in order, and potential treatment or changes in breeding management might then be considered. The reader is cautioned against making judgments based on examination of only one or two dismount samples, as poor sperm concentration or motility may be related more to the quality of the sample collected than to the ejaculate. In such cases, the odds for development of pregnancy are just as good for the occasional cover with a poor quality dismount as with other covers for that stallion. It is the consistent decline in semen quality of dismount samples for a stallion that previously had good quality dismounts that should raise concern. Paying close attention to reports of pregnancy outcome by date of cover will confirm whether any concern should be investigated. Monitoring of dismount samples for presence of inflammatory cells is also useful. The occasional presence of a high number of neutrophils in a dismount semen sample suggests the mare as the source of inflammation. Occasionally, a mare is identified that has an underlying genital infection that was not detected by routine pre-breeding examination. The finding of a high number of neutrophils in dismount samples can therefore be used to alert the veterinarian to direct post-breeding treatment(s) to
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the affected mare. If there is a consistent increase in inflammatory cell numbers in virtually all dismount samples from a stallion mating mares of good reproductive quality, the potential for the stallion having an internal genital infection should be considered and explored. If semen collected in an artificial vagina likewise has increased numbers of inflammatory cells, further diagnostic procedures should be performed to determine location and extent of the infection, and to arrive at appropriate treatment modalities.7 Brief mention is made here of performing regularly scheduled cultures of material collected from the distal urethra, fossa glandis, and shaft of the penis of stallions used in natural-service programs. Since potential venereal pathogens are usually harbored only on the external genitalia, cultures should be collected before washing, and prior to covering the first mare of the day. Some Thoroughbred farms perform these screening cultures weekly, bi-weekly or even monthly. The reader is cautioned against incriminating the stallion as carrying a venereal pathogen simply because Pseudomonas aeruginosa or Klebsiella pneumoniae are recovered on culture, as these organisms are common environmental contaminants of soil, stall bedding, and breeding shed floors. There are many phage types of Pseudomonas and many capsule types of Klebsiella that are considered to be non-pathogenic, so the chance of the stallion being a carrier of a venereal pathogen should not be made based on simply recovering one of these organisms on culture without other confirmatory evidence.13 Certainly, any stallions from which these organisms are recovered should be re-cultured to determine whether true colonization of the external genitalia has occurred. The reader is referred elsewhere in this book and other review articles13 for discussion of diagnosis and treatment of venereal diseases in the stallion, including contagious equine metritis and equine viral arteritis.
Pre-breeding infusion of semen extender Infusion of semen extender containing antibiotics into the uterus of a mare immediately prior to cover has been proposed as a method to control/ prevent endometritis, particularly when potentially pathogenic bacteria are being shed in the semen (termed the minimum contamination technique14 ). Results of one clinical trial confirmed the technique was beneficial in preventing post-breeding endometritis and improving pregnancy rates when a stallion with an internal genital infection caused by P. aeruginosa (venereally transmitted) was mated to reproductively normal mares.15 Though unproven, it is also possible that infusion of pre-warmed semen
extender immediately prior to cover may be of value when a given stallion’s sperm have short livability in vitro [e.g., longevity of sperm motility is short (minutes for raw semen, < 1 hour for extended semen) when maintained at room temperature as raw and extended semen], or when ‘hostile’ fluid accumulations are present within a mare’s genital tract (e.g., from urine pooling, intrauterine fluid accumulation, or presence of numerous neutrophils within the genital tract). In this respect, recent research by Troedsson et al.16 demonstrated that sperm are bound to neutrophils in samples collected from uteri of mares with ongoing endometritis. It is common for sperm to be bound to neutrophils, and for sperm motility to otherwise be poor (perhaps also adversely affected by inflammatory cell products), when dismount samples are found to contain high numbers of neutrophils. If sufficient pre-warmed extender is immediately added to the dismount sample, sperm binding may decrease and sperm motility improve. It is also possible that pre-breeding infusion of extender might be beneficial when mares are mated on the first postpartum estrus. One study demonstrated a 17% improvement in foal-heat pregnancy rates in mares bred artificially with entire or half-ejaculates mixed with semen extender compared with mares bred by natural cover.17 That study did not determine what factors might have contributed to this finding (e.g., improved sperm motility, beneficial factors of antibiotic- containing semen extender, decreased genital tract contamination). The reader is cautioned that no scientific reports utilizing adequate numbers of mares and stallions exist that confirm pre-breeding infusion of extender into the mare uterus can be beneficial to fertility of either fertile or subfertile stallions. Additionally, determination of a suitable volume of extender for prebreeding infusions that would not be detrimental to fertility has not been made. Certainly, use of an excess volume could be detrimental since fluid deposited into the uterus is rapidly expelled (within 20 minutes) following breeding,18 and since the act of mating itself results in substantial release of endogenous oxytocin with subsequent promotion of uterine contractions.19 In addition, when mares were bred with the same number of spermatozoa in 10 mL or 100 mL of extender (a concentration of either 25 × 106 or 2.5 × 106 , respectively, pregnancy rates were significantly decreased in the mares bred with low concentration spermatozoa20 ). It seems logical that the volume of extender infused into the uterus should be limited for these reasons (e.g., perhaps to 35–50 mL, as larger volumes might predispose to premature expulsion of a significant portion of the ejaculate, or might prevent a significant number of ejaculated sperm from accessing the oviducts). Nevertheless, in clinical
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Management of Subfertile Stallions Under Natural-Mating Conditions practice, the judicious use of this technique has sometimes been claimed to improve pregnancy rates.
Reinforcement breeding Reinforcement breeding (often termed ‘impregnation’) is a process whereby the dismount semen sample (and/or aspiration of semen pooled in the vagina immediately after cover) is collected following natural cover, and is then infused into the uterus of the mare that was just covered. Since debris from the mating process and breeding shed floor can get into dismount samples, it is wise to first filter the dismount semen sample to remove as much debris or dirt as possible, and then to immediately mix the dismount semen sample with a suitable volume (e.g. 5–20 mL)of pre-warmed extender prior to infusion into the uterus of the mare. Alternatively, extender can be mixed with the dismount sample (or semen sample aspirated from the vagina) prior to filtering and infusion into the uterus of the mare. The process has two potential advantages that might improve fertility: (i) it results in an increased number of sperm placed directly into the uterus and (ii) the extender provides some protection to sperm that may improve their livability within the uterus until they can access the oviduct where fertilization occurs. Not all stallions will benefit from reinforcement breeding. Certainly, some stallions that have a tendency to dismount early after ejaculation (with the penis still erect, and sometimes with the glans penis still engorged with blood), or that have difficulty in remaining fully intromitted into the mare’s vagina during mating, are candidates that logically should benefit from the procedure because they are less likely to instill major portions of the ejaculate into the mare uterus during mating (Table 120.7). Other Table 120.7 Influence of reinforcement breedings on pregnancy outcome (pregnancy rate per cycle; PR/cycle) in two stallions used in natural-service programs. Both stallions tended to dismount prematurely. For reinforcement breedings, dismount semen samples were immediately filtered and mixed with pre-warmed semen extender. After evaluating sperm concentration and motility, the extended semen (10– 30 mL) was infused transcervically into the uterine body of the mare that was covered. Once reinforcement breedings were thought to result in improved fertility, attempts were made to utilize the protocol on all mares serviced by the stallions, regardless of cycle of cover PR/cycle
Stallion n 1 2
71 12
PR/cycle
Without reinforcement
n
44% 37%
114 56
With reinforcement Improvement 66% 64%
+22% +27%
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Table 120.8 Influence of reinforcement breedings on pregnancy outcome for mares mated on their second estrous cycle (pregnancy rate per second cycle; PR 2nd/cycle) for 13 stallions used in natural-service programs. For reinforcements, dismount semen samples were immediately filtered and mixed with pre-warmed semen extender. After evaluating sperm concentration and motility, the extended semen (10–30 mL) was infused transcervically into the uterine body of the mare that was covered. Pregnancy rate per second cycle appeared to be improved in all but two stallions, although sufficient numbers of second cycle matings are lacking for some stallions
Stallion 1 2 3 4 5 6 7 8 9 10 11 12 13 Total
PR 2nd/cycle without reinforcement
PR 2nd/cycle with reinforcement
5/10 (50%) 7/13 (54%) 1/3 (33%) 4/8 (50%) 1/2 (50%) 3/6 (50%) 6/9 (67%) 1/4 (25%) 3/12 (25%) 4/9 (44%) 1/2 (50%) 6/9 (67%) 0/3 (0%) 42/89 (47%)
28/41 (68%) 40/55 (73%) 11/19 (58%) 20/27 (74%) 1/3 (33%) 24/41 (59%) 42/59 (71%) 31/47 (70%) 29/46 (63%) 28/45 (62%) 20/26 (77%) 27/42 (64%) 4/7 (57%) 305/458 (67%)
stallion disorders that reinforcement breedings might aid include failure to ejaculate sufficient numbers of normal motile sperm (particularly if sperm motility rapidly declines in vitro), shedding of potential bacterial pathogens in semen, urospermia, or hemospermia. When significant amounts of urine (regardless of whether the stallion or mare is the source of urine) are present in dismounts, addition of extender may or may not restore motility,21 and reinforcement breeding with that sample is probably contraindicated. One management protocol employed for using reinforcment breedings is to ‘reinforce’ all mares that fail to become pregnant on the first estrous cycle when they return on later cycles for mating (Table 120.8). Once a sufficient number of mares have been reinforcement bred, pregnancy rates can be compared between ‘reinforced’ and ‘non-reinforced’ mares. If fertility seems to be improved, reinforcement breeding may be justified as a standard protocol for that stallion. If dismount samples are used for reinforcement breeding, it is important to collect as many sperm dripping from the urethra after the stallion dismounts from the mare as possible. Using a widemouth disposable cup is helpful in this regard (Figure 120.3). In a recent study utilizing multiple logistic regression to adjust for factors affecting fertility of Thoroughbred stallions used in a natural-service mating program, reinforcement breedings containing
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Stallion: Management motility when either cooled or frozen (presumably by improving sperm membrane stability). As yet, no significant potential problems have been noted when diets are supplemented with omega-3 fatty acids, so supplementing the diet of stallions ejaculating suboptimal numbers of normal motile sperm may prove to be a viable option for improving fertility of some stallions.
Summary
Figure 120.3 Using a wide-mouth disposable plastic cup to collect drippings from the urethra of a stallion after dismount.
≥ 200 million sperm increased pregnancy rates 11.7%, whereas reinforcement breedings containing < 200 million sperm increased pregnancy rates 6.3%.22 In rare instances when a stallion ejaculates a low number of normal motile sperm of short longevity (i.e., sperm motility rapidly declines), managing his book of mares for multiple covers just prior to and immediately after ovulation may improve pregnancy rates. The strategy proposed is directed toward maximizing the number of live normal sperm that can access the oviduct as near to the time of expected fertilization as possible (I. Liu, 2003, personal communication). By necessity, the mare book number must be reduced as the stallion may be mated two or three times to a given mare in a 24-hour period. This mating strategy was found to improve pregnancy rates of two aged Thoroughbred stallions with declining testes size and semen quality, with very short livability of sperm in vitro (Table 120.9). Recent research23–25 demonstrated that dietary supplementation with omega-3 fatty acids can increase the number of progressively motile, morphologically normal sperm in ejaculates of some stallions, and may increase longevity of progressive sperm Table 120.9 Results of planned multiple (double or triple) matings in two aged Thoroughbred stallions with declining fertility related to ejaculation of low numbers of normal, motile sperm that had short longevity of motility Pregnancy rate per cycle
1 cover/estrus 2 covers/estrus in 24 h ≥ 3 covers/estrus in 36 h
Stallion 1
Stallion 2
0% (0/6) 18% (7/38) 43% (12/28)
47% (20/43) 64% (9/14) –
Investigation of suboptimal fertility achieved by a stallion should be carefully investigated, and directed toward identifying and correcting contributing factors. When routine reproductive management practices and veterinary care are good, there still remains a number of simple techniques that can be applied in an effort to improve pregnancy rates. The value gained from increased production of valuable foals, and collection of more stud fees, will exceed the cost outlay for the increased management efforts required.
References 1. Pickett BW, Voss JL, Bowen RA, Squires EL, McKinnon AO. Seminal characteristics and total scrotal width (TSW) of normal and abnormal stallions. In: Proceedings of the 33rd Annual Convention of the American Association of Equine Practitioners, 1988, pp. 487–518. 2. Blanchard TL, Varner DD. Evaluating breeding soundness in stallions. 5. Predicting potential fertility. Vet Med 1997;Sept:815–18. 3. Roser JF. Idiopathic subfertility/infertility in stallions: new endocrine methods of diagnosis. In: Proceedings of the Annual Meeting of the Society for Theriogenologists, 1995, pp. 136–46. 4. Blanchard TL, Varner DD. Testicular degeneration. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 855–60. 5. Blanchard TL, Johnson L, Roser AJ. Increased germ cell loss rates and poor semen quality in stallions with idiopathic testicular degeneration. J Equine Vet Sci 2000;20:263–5. 6. Pickett BW, Squires EL, McKinnon AO. Influence of insemination volume and sperm number on fertility. In: Pickett BW, Squires EL, McKinnon AO (eds) Procedures for Collection, Evaluation and Utilization of Stallion Semen for Artificial Insemination. Bulletin No. 03:51. Fort Collins: Colorado State University Animal Reproduction Laboratory, 1989. 7. Varner DD, Schumacher J, Blanchard TL, Johnson L. Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991. 8. Kenney RM, Hurtgen JP, Pierson RH, Witherspoon D, Simons J. Manual for clinical fertility evaluation of the stallion. In: Proceedings of the Annual Meeting of the Society for Theriogenologists, 1983, pp. 1–100. 9. Douglas RH, Umphenour N. Endocrine abnormalities and hormonal therapy. Vet Clin North Am Equine Pract 1992;8:237–49. 10. Love CL. Evaluation of breeding records. In: Blanchard TL, Varner DD, Schumacher J, Love, CC, Brinsko SP, Rigby
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Management of Subfertile Stallions Under Natural-Mating Conditions
11.
12. 13. 14.
15.
16.
17.
18.
SL (eds) Manual of Equine Reproduction, 2nd edn. St. Louis: Mosby, 2003; pp. 229–37. Baker CB, Little TV, McDowell KJ. Studies of factors limiting reproductive success in horses and derivation of investigative strategies for identifying their causes. PhD dissertation, University of Kentucky, 1995. Brinsko SP. Retrograde ejaculation in a stallion. J Am Vet Med Assoc 2001;218:551. Blanchard TL, Kenney RM, Timoney PJ. Venereal disease. Vet Clin North Am Equine Pract 1992;8:191–203. Kenney RM, Bergman RV, Cooper WL, Morse G.W. Minimal contamination techniques for breeding mares: techniques and preliminary findings. In: Proceedings of the 21st Annual Convention of the American Association of Equine Practitioners, 1975, pp. 327–36. Blanchard TL, Varner DD, Love CL, J. Hurtgen, M. Cummings, R. Kenney. Use of a semen extender containing antibiotic to improve the fertility of a stallion with seminal vesiculitis due to Pseudomonas aeruginosa. Theriogenology 1987;28:541–6. Troedsson MHT, Loset K, Alghamdi AM, Dahms B, Crabo BG. Interaction between equine semen and the endometrium: the inflammatory response to semen. Anim Reprod Sci 2001;68:273–8. Blanchard TL, Thompson JA, Brinsko SP, Stich K, Varner DD, Rigby SL. Breeding mares on foal heat: a 5-year retrospective study. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2004, pp. 525–30. Katila T, Sankeri S, Makola O. Transport of spermatozoa in the genital tract studied by a scintigraphic method. In: Proceedings of the 7th International Symposium on Equine Reproduction, 1998, pp. 149–150.
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19. Madill S, Troedsson MHT, Alexander SL, Shand N, Santshi EM, Irvine CHG. Simultaneous recording of pituitary oxytocin secretion and myometrial activity in estrous mares exposed to various breeding stimuli. In: Proceedings of the 7th International Symposium on Equine Reproduction, J Reprod Fertil 2000, Suppl 56:351–361. 20. Squires EL, Barnes CK, Rowley HS, McKinnon AO, Pickett BW, Shideler RK. Effect of uterine fluid and volume of extender on fertility. In: Proceedings of the 35th Annual Convention of the American Association of Equine Practitioners, 1989, pp. 25–30. 21. Griggers S, Paccamonti DL, Thompson RA, Eilts BE. The effects of pH, osmolarity and urine contamination on equine spermatozoal motility. Theriogenology 2001;56:613–22. 22. Blanchard TL, Love CC, Thompson JA, Ramsey J, O’Meara A. Role of reinforcement breeding in a natural service mating program. In: Proceedings of the 52nd Annual Convention of the American Association of Equine Practitioners, 2006, pp. 385–7. 23. Brinsko SP, Varner DD, Blanchard TL, Day BC, Wilson ME. Effect of feeding a DHA-enriched nutriceutical on the quality of fresh, cooled and frozen stallion semen. Theriogenology 2005;63:1519–27. 24. Harris MA, Anderson CR, Webel SK, Godbee R, Sanders SR, Schurg WA, Baumgard LH, Arns MJ. Effects of feeding an omega-3 rich supplement on the fatty acid composition and motion characteristics of stallion spermatozoa. In: Proceedings of the 19th Annual Meeting of the Equine Science Society, 2005, p. 239. 25. Harris MA, Baumgard LH, Arns MJ, Webel SK. Stallion spermatozoa membrane phospholipids dynamics following dietary n-3 supplementation. Anim Reprod Sci 2005;89:234–7.
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Chapter 121
Venereal Disease Elizabeth S. Metcalf
Although there is considerable concern regarding the potential transmission of pathogenic organisms through horse breeding, high morbidity associated with equine venereal disease has been traced to very few organisms. The organisms of highest global significance that are a serious threat to the equine breeding industry include contagious equine metritis (CEM), equine viral arteritis (EVA), and dourine. These organisms, of bacterial, viral, and protozoal etiologies respectively, show potential for efficient venereal transmission that can have significant economic impact upon equine populations, through infertility, abortion, or death. There are, as well, other venereal microorganisms of less or questionable importance that can be isolated from the reproductive tract or semen of equine breeding stock. Since the stallion is often an asymptomatic carrier of venereal organisms, control of disease spread relies heavily on quarantine, testing, and vaccination (if available). Vigilant surveillance, consistent among countries that import and export both breeding horses and semen, as well as industry education contribute to management of disease transmission.
Contagious equine metritis Because of its large-scale distribution and often compromising effect on fertility, CEM is of significant interest in horse-breeding populations throughout the world. First documented in Europe in 1977,1 the disease quickly found its way into other countries, the USA included, harbored in the reproductive tract of Thoroughbred stallions.2, 3 Through intensive eradication and regulatory programs, until December 2008, the USA was considered to be a CEM-free country. At this time, eight out of 22 stallions on a Kentucky breeding farm were found to be infected with the organism and the status of the USA was evaluated for reclassification. Horses may be imported from CEM-affected countries following routine testing. Although some non-European coun-
tries are considered CEM-free, sporadic identification of imported breeding stock harboring this organism emphasizes the need for rigorous preventative and control programs. Taylorella equigenitalis, the causative agent for CEM, is a true sexually transmitted infection of horses, for its main recognized means of spread is via a venereal route by reciprocal transmission (i.e., it is spread both from the stallion to the mare and from the mare to the stallion). Infected fomites are suspected to act as a means of transmission as well. This highly contagious, Gram-negative coccobacillus appears to infect mares regardless of their susceptibility to endometritis. In the mare, signs of acute CEM exposure can range from a seemingly inapparent infection to an early return to estrus to a profound cervicitis, vaginitis, and metritis, coupled with a copious mucopurulent discharge from the vulva.4 These clinical signs persist in the mare for a relatively short period of time of less than 14 days. The disease may spontaneously resolve or the mare may become a persistent carrier. The bacteria have been cultured from the endometrium or, more commonly, from the clitoral fossa and sinuses of infected mares. During the acute phase of infection, T. equigenitalis can be readily cultured from the cervical swabs of infected mares (84%) and, less likely, from clitoral swabs (69%). In contrast, in chronically infected mares, the organism is more likely to be detectable from clitoral swabs (93%) than from cervical swabs (31%).5 Owing to the changing distribution of T. equigenitalis during the progression of infection, it is important to swab both sites. However, it has been estimated that, in 95% of mares infected for longer than 8 weeks, the organism should be detected by a single clitoral swab.6 Conversely, stallions are asymptomatic carriers. T. equigenitalis has been cultured from the sheath of the penis, the urethral fossa and sinus, and the distal urethra of contaminated stallions. The internal genitalia may be affected as well for CEM has also been
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Venereal Disease cultured from the seminal vesicles and epididymis of an infected stallion at necropsy.7 However, semen culture, thus far, is considered unreliable in accurate detection of the organism. Reliable diagnosis of T. equigenitalis can be challenging, not only because of its fastidious transport and growth requirements but also because of its changing predilection for certain locations, particularly within the mare, based on chronicity of the disease. A special, extremely small calcium alginate swab (Calgiswabs; Puritan Medical Products Co LLC, Guilford, ME) is required for clitoral sinus culture. To optimize isolation of T. equigenitalis and prevent contaminant bacterial overgrowth, swabs should be refrigerated, transported in Amies charcoal medium, and should be inoculated onto a chocolate agar plate within 48 hours of obtaining the sample. Two biotypes of T. equigenitalis are recognized – one that is streptomycin sensitive and one that is streptomycin resistant; therefore the agar culture plate must include appropriate antibiotics to prevent the overgrowth of contaminants. The plated agar is incubated in a microaerophilic environment under 5–10% CO2 at 37◦ C for a minimum of 7 days. Because T. equigenitalis is such a slow-growing bacterium and there is often overgrowth contamination on inoculated plates, the incidence of false-negative diagnoses is suspected to be high, even after 7 days of incubation under recommended conditions. Recent outbreaks of CEM in the USA were suspected to have originated from a failure to identify positive cultures from imported stallions. In the most recently reported outbreak of CEM in the USA, ultimately involving 11 CEM-infected stallions, more than 750 mares were exposed to this organism through artificial insemination. Although the index case was not identified, it was surmised that the spread of the organism among the stallions was caused by shared breeding equipment – wash bucket, phantom, or artificial vagina case. Fertility was not apparently compromised nor were there reports of copious post-breeding vaginal discharge that would raise suspicion of exposure. Initial screening of the bred mares did not yield positive cultures of the clitoral sinuses or fossa; however, five exposed mares ultimately yielded a positive clitoral culture. It is believed that the inclusion of antibiotics in semen extenders may have prevented massive spread and exposure. Laboratory identification of T. equigenitalis has become more sophisticated and reliable during the past decade, owing to the use of an amplified and highly sensitive polymerase chain reaction (PCR) assay for detection.8–10 In the early stages of testing the PCR assay for CEM, it appeared that the prevalence of the disease may be higher than expected, for initial anal-
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ysis identified far more horses that had been exposed to the organism than suspected.11 Furthermore, different strains of the organism continue to be identified that may vary in pathogenicity. As an example, in a recent study, Hayna et al.12 successfully transferred embryos from two donor mares that were bred with semen from stallions from which T. equigenitalis had been recovered from their external genitalia. Although repeated cultures from these stallions were positive, neither the inseminated donor mares nor the recipient mares showed any positive cultures for CEM. Other studies have shown that infection with a related organism, Taylorella asinigenitalis, has been reported mostly in jacks but also in a stallion; its significance is controversial and this variant may be falsely identified by culture as T. equigenitalis.13–17 Clearly, further research regarding the pathogenicity of Taylorella spp. is necessary and important in order to accurately identify carriers of the organism. T. equigenitalis is susceptible to a wide range of antibiotics. Early studies suggested that gentamicin, as opposed to ampicillin or penicillin derivatives, was most effective in the treatment of experimentally infected semen.18, 19 For more than 30 years, recommended treatment of the external genitalia has consisted of rigorous scrubbing of the prepuce, penis, urethral sinus, and fossa glandis with 4% chlorhexidine solution, followed by copious coating and packing of these areas with 2% nitrofurazone ointment for a minimum of 7 consecutive days. However, a more recent study20 has strongly advocated treating CEM carrier stallions with systemic as well as topical antibiotics. The author recommends initial treatment for 10 consecutive days with a 4% chlorhexidine wash of the external genitalia followed by the topical application of both a silver sulfadiazine cream and a solution of procaine penicillin G plus streptomycin. In this study of four CEM carrier stallions undergoing such treatment, the infection persisted in a single stallion until 10 days of systemic administration of trimethoprim–sulfamethoxazole was added to the treatment protocol. Therefore, the combination of both systemic and local treatment of carrier stallions may prove necessary to eliminate, or at least, reduce the risk of venereal transmission of T. equigenitalis.
Equine viral arteritis Sporadic outbreaks of EVA have caused significant economic damage to affected populations of breeding stock in many countries.21 EVA occurs sporadically in certain racing horse populations22 and has rapidly spread over the past three decades to show horse populations. This small, enveloped, single-stranded
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Stallion: Management
RNA virus primarily targets the endothelium of small blood vessels and macrophages, causing clinical signs that vary in severity, depending on the viral strain.23 Clinical manifestations of the disease range from subclinical infection to pyrexia, anorexia, nasal discharge, conjunctivitis, urticaria, dependent edema, and, most devastating, abortion. Abortion, due to equire arteritis virus (EAV), may be the result of placental compromise due to severe edema and necrosis of the fetal membranes rather than direct fetal infection. The respiratory signs caused by EAV are indistinguishable from other equine respiratory tract diseases. Congenital infections of foals may cause pneumonia during the first few days of life; acquired infections in older foals may cause both pneumonia and enteritis24 and can be fatal. In older animals, however, the disease is usually self-limiting. Treatment of respiratory signs consists of supportive therapy, non-steroidal anti-inflammatory agents, as well as antibiotic administration to reduce the chance of ensuing bacterial pneumonia. Other promising treatments are under investigation and hold promise in combating EAV infection. Recent results from an in vitro experiment demonstrated that EAV-infected cells were susceptible to a novel gene therapy consisting of RNA interference; EAV titer of equine cultured cells was significantly reduced by addition of interfering RNA oligonucleotides.25 Still, the major economic impact of EVA on the horse industry arises from abortion in mares and the resultant carrier state of stallions rather than from neonatal mortality. The EAV organism has been identified in the many areas of the stallion urogenital tract including the epididymis, vas deferens, ampullae, prostate, seminal vesicles, bladder, and urethra. Naive stallions infected with the organism will ‘shed’ the virus in their semen during the acute and convalescent phase of the infection. It is estimated that 30–70% of these stallions will progress to the carrier state and become persistently infected with EAV.24 These chronic ‘shedders’ will continue to shed the virus in their semen for months, years, or throughout their lifetime. Spontaneous cessation of shedding has been reported,26 and yet the mechanism by which this occurs is undetermined. The persistent infection is testosterone dependent;27 castration results in the cessation of viral shedding in the semen. However, the exogenous administration of testosterone to these castrated horses did not cause elimination of viral shedding. Other attempts to eliminate the virus from the ejaculate have been tested. Chemical castration through the administration of a gonadotropinrealeasing hormone (GnRH) antagonist has been only transiently effective in assuaging viral contamination of semen; it appears that viral excretion recommences
after treatment is withdrawn. Immunocastration using a GnRH vaccine may effectively lower testosterone production and therefore viral shedding, but it concomitantly impairs libido, testicular function, and semen characteristics;28 therefore, caution with these products should be exercised in consideration of the future fertility of these stallions. Further efforts to eliminate shedding have been aimed at ‘trapping’ the virus. Using density gradient centrifugation R (Equipure ) and a ‘swim-up’ protocol for processing semen, researchers were able to remove the arteritis virus from virus-laden semen. Although the viral burden of the ejaculate was significantly lowered following this sequential centrifugation, small numbers of viral particles still remained in the semen.29 Most recently, an antiviral strategy using lectin as the carbohydrate-linking moiety to bind small, enveloped viruses shows some promise as a strong in vitro inhibitor of arteriviridae.30 The ability to eliminate viral contamination of semen without compromising stallion fertility remains at the forefront of future research. As expected, fertility of the stallion is potentially compromised only during the acute phase of the infection, most likely due to the febrile period following exposure. Semen parameters and fertility remain unaffected by the carrier status of the chronically infected stallion.31, 32 Likewise, fertility of the mare does not appear to be affected when a naive mare is bred with EVA-positive semen from a carrier stallion. Although EVA is a venereal disease, the most common mode of transmission is through the respiratory route, as an aerosol33 or as a contaminant of bedding or fomites.34 Venereal transmission, although believed to occur less frequently, not only infects the mare through insemination, but the transmitted virus can then rapidly spread laterally throughout the herd by means of respiratory contact with the newly infected mare.32, 35 The incubation period of the disease ranges from 3 to 14 days, depending on the mode of transmission, following initial exposure to the arteritis virus.35 Up to 75% of a naive pregnant mare herd (3–10 months gestation) may abort 10–33 days after exposure; future fertility of these mares appears unaffected.35 Diagnosis of EVA consists of serological screening for EVA-specific antibodies and specific virus isolation. The virus neutralization test (VNT) is currently recognized as the gold standard test for antibody detection. It is important to realize that this widely accepted test for EVA may have some limitations. Contamination of serum by inoculation of horses with an inactivated EHV-1 vaccine has been reported to affect results of the VNT, as it may increase the number of false-positive reactors.36 The contamination is, however, limited to horses vaccinated with a specific
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Venereal Disease EHV-1 vaccine available only in the European Union and Australia. The development of a highly sensitive and specific enzyme-linked immunosorbent assay (ELISA) for detection of antibodies against the EVA envelope protein (GL ) has been reported but has not yet become widely available commercially37 and has not yet replaced the VNT. Apparently, the ELISA is not affected by serum cytotoxicity caused by the EHV-1 vaccine.36 Most recently, the development of a microsphere immunoassay, although less sensitive than the VNT, represents a rapid, economical screening test for the presence of EAV antibodies.38 To date, however, there is no ELISA test available with greater sensitivity and specificity than the VNT. Rising antibody titers are useful in diagnosis. Horses that are seronegative for EAV convert to the seropositive status following exposure to EAV through natural infection or vaccination. A fourfold increase in serological titer between two samples drawn from a horse 2–3 weeks apart is confirmatory of recent exposure of the naive horse. Antibody presence, especially from natural exposure to the virus, is long term. Until recently, it was believed that all stallions that persistently shed the virus in their semen would always test positive for the VN antibody.39 However, there is a single case report of a Swedish stallion that, in the absence of detectable EAV antibody levels, continued to shed the virus in semen.40 Presently, there are two means of diagnosis of the presence of EAV within the semen. Firstly, virus isolation (VI) using an RK-13 cell assay line41 is especially sensitive in detecting the virus in the semen of carrier stallions (Fig. 121.1). Secondly, and far more time-consuming and labor intensive, is test breeding of naive (seronegative) mares with subsequent identification of seroconversion by VNT. Lastly, a polymerase chain reaction test on nucleic acid is under development. In cases with EAV as the causal agent for an abortion, both the fetus and placenta contain large quantities of EVA viral particles. Submission of fetal spleen,
Figure 121.1 Electron micrograph of equine arteritis virus (EAV) in semen. Courtesy P. Timoney.
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lung, and kidney as well as fresh placenta is recommended for diagnosis. In the mare that has aborted, a fourfold increase in EVA titer over a 3-week period also supports a diagnosis of EVA-induced abortion. However, the time from EAV exposure to abortion ranges from 10 to 30 days, during which time EVA titers rapidly increase. By the time abortion occurs, antibody titers are already high. Therefore, unless the titer is measured at the time of initial infection, a titer at the time of abortion and 3 weeks later may fail to yield an accurate diagnosis. Control of the disease lies in testing, identification, and vaccination of both mares and stallions. Naive horses that have been exposed to the natural infection, through either venereal or respiratory exposure, will develop long, if not lifetime, immunity to EAV. A safe, highly effective, modified, live R ) provides long-lasting immunity in vaccine (Arvac naive breeding stock. A period of isolation should follow primary vaccination owing to the potential shedding of the attenuated virus through the nasopharynx26 and urine. An annual booster inoculation is currently recommended. It is important to recognize that the vaccine has not been approved by the manufacturer for use in pregnant mares. Despite this, the vaccine has been administered to thousands of pregnant mares during disease outbreaks with limited consequences of abortion due to vaccine virus.24 A recent EVA outbreak in the USA in 2006 led researchers to conclude that mares in the final 2 months of gestation may be the pregnant mare population most at risk for abortion due to the vaccine virus, for only a very few late-term mares aborted of all pregnant mares vaccinated.24 There are, as well, several other reports of an aborted fetus containing an identical gene sequence to the live vaccine with which the dam was vaccinated a few days earlier.42 A new generation of a recombinant EVA vaccine is currently under testing and investigation which may improve safety in vaccinating mares at all stages of gestation.43 In parts of Europe and other non-European R , countries, an inactivated EVA vaccine (Artevac Southampton, UK) is available. It does not appear as effective as a modified live vaccine, because several doses are recommended to stimulate protective levels of virus-neutralizing antibodies; unlike the modified live vaccine, efficacy and protection from the killed product remain questionable.44 Therefore, in order to prevent the venereal spread of this disease, all unexposed breeding stallions should be vaccinated with the modified live vaccine. Colts ranging from 6 to 12 months of age should be vaccinated in order to prevent a carrier state later in life. Naive mares intended to be bred with EAVinfected semen should be vaccinated as well. Documentation of the serologically negative status of
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a stallion at the time of vaccination is imperative and should remain with the stallion throughout its breeding lifetime. Some breed registries require proof of EVA status of their stallions. Other registries, such as the American Quarter Horse Association, have initiated a voluntary stallion vaccination reporting system whereby the information is available to mare owners. Within the USA, there are also some states that mandate individual stallion documentation of EVA status.
Dourine Dourine is a lethal, sexually transmitted disease of horses caused by Trypanosoma equiperdum. Currently, there are two recognized strains of T. equiperdum that are very closely related both antigenically and genetically. Although the disease is most often seen in horses in Africa, the Middle East, and South and Central America, sporadic cases have recently been recorded in Asia and Germany. Clinical signs of infection progress from intermittent pyrexia, weight loss, and copious purulent discharge from the urethra or vulva to raised coinsized skin plaques that are characteristic of the disease. These plaques become areas of skin depigmentation. Neurological signs (progressive ataxia, hindlimb paralysis) ensue and death eventually follows. There are rare reports of recovery. Diagnosis of the disease has relied upon the complement fixation test. However, because the test cannot reliably differentiate between T. equiperdum, Trypanosoma evansi and Trypanosoma brucei, current methods of diagnosis are under intense scrutiny. The ELISA and PCR assay are under current investigation but are not yet widely accepted for diagnosis.45–47
Other organisms Opportunistic bacterial invaders Because certain bacterial organisms are often associated with infectious endometritis in the mare, venereal transmission at coitus is suspected as a likely origin of infection. A wide variety of commensal bacteria colonize the stallion’s external genitalia; they are not considered to be pathogenic.48 However, alterations of this normal flora, perhaps caused by washing the penis with soap, housing the stallion in contaminated bedding, antibiotic treatment, or breeding an infected mare, can trigger the overgrowth of opportunistic bacteria not only on the external genitalia but also throughout the internal reproductive tract of the stallion. The agents of greatest concern for venereal spread include potentially pathogenic strains of Pseudomonas aeruginosa and Klebsiella
pneumoniae. P. aeruginosa has been shown to be detrimental to specific parameters of cooled semen and may, in turn, directly decrease fertility of semen49 rather than solely affect the fertility of the mare by inducing endometritis. It is important to differentiate between the capsular polysaccharide types of these organisms as only certain types are associated with reproductive disease in the horse. For instance, K. pneumoniae capsule types6 1, 2, 5, and 7 and type50 39 have been implicated as pathogens in mares and stallions. Unfortunately, few veterinary laboratories continue to offer identification of these capsule types at this time, but the development of a sensitive multiplex-PCR assay for the identification of K. pneumoniae isolates has been undertaken.51 Other potentially pathogenic bacteria that may be transmitted with breeding include strains of Streptococcus zooepidemicus and Escherichia coli. A normal mare that is resistant to persistent mating-induced endometritis usually clears these potential pathogens without veterinary intervention. Mares considered susceptible to endometritis and delayed uterine clearance are at highest risk of prolonged inflammation and subsequent infertility following inoculation with these organisms.52 There are numerous means of minimizing or eliminating the risk of disease transmission of these opportunistic bacteria. First, careful management of the breeding stallion with respect to bedding and appropriate cleansing of the penis with water before breeding or semen collection can reduce the risk of overgrowth of pathogenic bacteria on the penis. Secondly, by ensuring that mares bred by natural cover are not suffering from endometritis secondary to a pathogenic organism, the chance of disease transmission on subsequent covers by the stallion is lessened. Thirdly, the use of minimal contamination techniques in the artificial insemination of mares serves to lower the chances of concomitant uterine inoculation with a pathogenic organism. Lastly, the administration of antibiotics to which the pathogen is susceptible, as a prebreeding uterine infusion, a component of the semen extender, or a post-breeding uterine infusion(s), minimizes possible establishment of infection within the mare as well as the eventual spread of a sexually transmitted organism. Equine herpesviruses Equine coital exanthema (ECE) is a highly contagious venereal disease caused by equine herpesvirus type 3 (EHV-3). The clinical disease is recognized by the presence of painful lesions on the external genitalia of both the mare and the stallion, predominately on the stallion’s penis (Fig. 121.2) and the mare’s perineum and vulva (Fig. 121.3). The lesions often begin as
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Figure 121.2 Ulcerative lesion on the stallion penis caused by equine herpesvirus 3. Courtesy P. Timoney.
vesicles or papules and, within 10–14 days, erupt into ulcerative nodules. They eventually result in rounded, scabbed, depigmented areas. Clinical signs of the disease are limited to the distal reproductive tract; systemic signs of illness are not observed. Definitive diagnosis of infection is made by identifying inclusion bodies on histopathology or through the use of a highly sensitive and specific PCR assay.53 Electron microscopy can also be used
Figure 121.3 Areas of perineal depigmentation as a result of infection with equine herpes virus 3. Courtesy P. Timoney.
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to identify herpes viral particles on smears with negative contrast stain. Although usually transmitted by coitus, infected fomites have been implicated in the spread of EHV3. Therefore, artificial vaginas, rectal probes, and insemination equipment should be viewed as possible sources of infection. Changing rectal gloves should be a routine practice, but, in some institutions where multiple mares are presented to students at one time, outbreaks of coital exanthema occur quite regularly. Semen collection and artificial insemination have been shown to reduce the spread of EHV-3,54 but the pain associated with the lesions may dampen copulatory behavior. Ex copula ejaculation may be a less painful option for semen collection. Apart from reluctance to copulate, fertility appears to be uncompromised55 by EHV-3 infection. However, there may be exceptions in certain cases. Firstly, if inflammation of the genitalia and/or fever occurs with infection, the spermiogram may be affected. Secondly, pregnancy may be at risk because, under experimental conditions, intrauterine inoculation with the virus has reportedly caused an abortion in a mare at 7 months gestation.56 In contrast to other herpesviruses found in both equids and other species, reactivation of latency of EHV-3 remains controversial due to conflicting reports.6, 54, 57 In many local outbreaks, equine coital exanthema seemed to represent a self-limiting condition; affected animals usually recovered within 2–3 weeks of the initial clinical signs and overt clinical signs of reactivation of latency were not apparent. In contrast, however, recurrent EHV-3 infection among draft horses pastured together over several years in Japan has been documented whereby naive individuals introduced to the herd continued to exhibit clinical signs of ECE.54 Sporadic outbreaks in naive individuals introduced into a herd indicate that the virus is spread by certain carrier animals that fail to demonstrate overt clinical signs of disease. In an attempt to further investigate the latent nature and potential reactivation of both clinical signs and viral shedding of EHV-3, two previously infected mares were treated with high doses of corticosteroids for 3 days. Following treatment, one of the two mares developed a single small vulvar erosion and exhibited nasal viral shedding as well as a rise in viral serum neutralization titer. The other mare showed no clinical signs and her viral titer was decreased.58 Despite this disparity between studies and herd outbreaks, it must be assumed that EHV-3 undergoes neurolatency and clinical signs may recur. Further research is needed to determine the inconsistency in outcomes following infection with this α-herpesvirus. Interestingly, another herpesvirus, equine herpesvirus type 1 (EHV-1) has also been documented
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in stallion semen;59–61 its presence is of unknown significance. Different diagnostic tests for the organism yield conflicting results. EHV-1 has been found in the semen of both clinically normal, presumably latently infected, stallions as well as a Grevy’s zebra stallion that demonstrated acute signs of multisystemic disease related to EHV-1. In the zebra stallion, histopathological examination of the genital tract revealed EHV-1 infection of Leydig cells and germinal epithelium, suggesting that sperm parameters may be affected by the infection.60 In experimentally inoculated pony stallions, EHV-1-laden semen was associated with a decrease in normal sperm morphology as well as an influx of inflammatory cells in the sperm-rich fraction of the ejaculate.59 In contrast to the above studies, on a farm believed to be endemic for EHV-1, a stallion that was EHV-1 positive on a PCR analysis of a nasopharyngeal swab failed to reveal the presence of EHV-1 DNA in the ejaculate.62 Furthermore, in a study examining stallion fertility, PCR analysis of ejaculates from 50 stallions did not detect EHV-1 DNA in any samples.63 Since EHV-1 represents a leading cause of abortion in pregnant mares, and continues to cause devastating economic losses despite rigorous vaccination strategies, the identification of viral particles in semen is concerning. Experimental inoculation of equine embryos with EHV-1 particles has shown that the virus is difficult to remove from the embryos despite repeated washings.64 It has been estimated that > 50% of the horse population are latent carriers of the infection, and reactivation of latency of the virus can be responsible for clinical infection. No known outbreaks of clinical disease associated with venereal transmission have been reported.59
Piroplasmosis The potential venereal transmission of piroplasmosis receives attention only in cases of hemaspermia, or infected blood contamination of semen. Such transmission has never been reported. Piroplasmosis, a disease spread by ticks, is caused by Babesia equi or by a less pathogenic strain, Babesia caballi. Considered endemic in many areas of the world, the horse represents the reservoir for B. equi. Curative treatment is questionable and attempts are often unrewarding because recrudescence of the disease is frequently encountered.
Other potential pathogens Potential venereal spread of other pathogenic organisms isolated either from the external genitalia or from the ejaculate must be considered as well. Numerous species of yeasts and fungi, such as As-
pergillus spp. and Candida spp., can serve as potential pathogens, especially in mares susceptible to endometritis and infection. Likewise, Chlamydia spp. and Mycoplasma spp. have been isolated from semen from both clinically normal and subfertile stallions. Mycoplasma equigenitalium, a common genital tract organism, has been isolated from the clitoral fossa of mares65 as well as from the fossa glandis, penis, and semen of stallions.66 Chlamydial organisms have been recovered in cases of equine abortion. 67 Although their significance is unknown, these organisms have been, in some studies, associated with compromised fertility in the form of salpingitis,68, 69 endometritis, and abortion70 in mares as well as balanoposthitis in stallions. In contrast, another group of researchers who examined 80 semen samples and 19 vaginal swab samples by PCR analysis failed to find any mycoplasmal DNA in any of the samples.71 Therefore, both their presence and significance with respect to potential venereal transmission warrants future investigation. In addition to venereal spread, these organisms may affect not only equine fertility but also other systems and other species. Finally, the potential for zoonotic spread cannot be overlooked.
Conclusion The increasing ease of movement of equine breeding stock, gametes, and embryos enhances the potential risk for sexually transmitted disease. State and federal testing requirements often encourage the equine veterinary industry to regulate itself, as it becomes increasingly challenging to keep pace with disease diagnoses on a worldwide level. The codes of practice organized and annually updated for specific transmissible diseases in some European countries represent a means for the equine industry to constantly monitor emerging global diseases that place our equine population at risk. The inapparent carriers of disease represent the greatest obstacle, emphasizing the need for quarantine, serosurveillance, and prevention, by means of vaccination or treatment. The increasing sophistication, sensitivity, and specificity of diagnostic tests minimize the chances of false-positive and -negative reactors, thereby identifying carriers with greater accuracy.
References 1. Platt H, Atherton JG. Contagious equine metritis. Vet Rec 1977;101:434. 2. Timoney PJ, Ward J, Kelly P. A contagious genital infection of mares. Vet Rec 1977;101:103. 3. Knowles RC. Contagious equine metritis situation report. Proc Am Assoc Equine Pract 1983;29:295–9. 4. Timoney P. Contagious equine metritis. Comp Immunol Microb Infect Dis 1996;19:199.
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Venereal Disease 5. Wood JL, Kelly L, Cardwell JM, Park AW. Quantitative assessment of the risks of reducing the routine swabbing requirements for the detection of Taylorella equigenitalis. Vet Rec 2005;9; 157: 41–6. 6. Wood JL, Cardwell J, Castillo-Olivares J, Irwin V. Transmission of diseases through semen. In: Samper JC, Pycock J, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders, 2007; pp. 266–74. 7. Schluter H, Kuller HJ, Friedrich U, Selbitz JH, Marbitz T, Beyer C. Epizootiology and treatment of contagious equine metritis (CEM) with particular reference to treatment of infected stallions. Praktische Tierarzt 1991;72:503–11. 8. Parlevliet JM. Epidemiological aspects of Taylorella equigenitalis. Theriogenology 1997;47:1169–77. 9. Anzai T, Wada R, Okuda T, Aoki T. Evaluation of the field application of PCR in the eradication of contagious equine metritis from Japan. J Vet Med Sci 2002;64:999–1002. 10. Duquesne F, Pronost S, Laugier C, Petry S. Identification of Taylorella equigenitalis responsible for contagious equine metritis in equine genital swabs by direct polymerase chain reaction. Res Vet Sci 2007;82:47–9. 11. Parlevliet JM, Samper JC. Disease transmission through semen. In: Samper JC, Pycock J, McKinnon AO (eds) Equine Breeding Management and Artificial Insemination. Philadelphia: W.B. Saunders Co., 2000; pp. 133–40. 12. Hayna JT, Syverson CM, Dobrinsky JR. Embryo transfer during concurrentcontagious equine metritis infection. Reprod Fertil Dev 2008;20:157–8. 13. Jang SS, Donahue JM, Arata AB, Goris J, Hansen LM, Earley DL, Vandamme PA, Timoney PJ, Hirsh DC. Taylorella asinigenitalis sp. nov., a bacterium isolated from the genital tract of male donkeys (Equus asinus). Int J Syst Evol Microbiol 2001;51 (Pt 3): 971–6. 14. Premanandh J, George LV, Wernery U, Sasse J. Evaluation of a newly developed real-time PCR for the detection of Taylorella equigenitalis and discrimination from T. asinigenitalis. Vet Microbiol 2003;95:229–37. 15. Matsuda M, Moore JE. Recent advances in molecular epidemiology and detection of Taylorella equigenitalis associated with contagious equine metritis (CEM). Vet Microbiol 2003;97:111–22. ¨ C, Johansson KE. Isolation and iden16. B˚averud V, Nystrom tification of Taylorella asinigenitalis from the genital tract of a stallion, first case of a natural infection. Vet Microbiol 2006;116:294–300. 17. Wakeley PR, Errington J, Hannon S, Roest HI, Carson T, Hunt B, Sawyer J, Heath P. Development of a real time PCR for the detection of Taylorella equigenitalis directly from genital swabs and discrimination from Taylorella asinigenitalis. Vet Microbiol 2006;118:247–54. 18. O’Driscoll JG, Troy PT, Geoghegan FJ. An epidemic of venereal infection in thoroughbreds. Vet Rec 1977;101:359–60. 19. Timoney PJ, O’Reilly PJ, Harrington AM, McCormack R, McArdle JF. Survival of Haemophilus equigenitalis in different antibiotic-containing semen extenders. J Reprod Fertil Suppl 1979;27:377–81. 20. Kristula MA, Smith BI. Diagnosis and treatment of four stallions, carriers of the contagious metritis organism: case report. Theriogenology 2004;61:595–601. 21. MacLachlan N, Balasuriya U. Equine viral arteritis. Adv Exp Med Biol 2006;581:429–33. 22. Bryans JT, Doll ER, Knappenberger RE. An outbreak of abortion caused by the equine arteritis virus. Cornell Vet 1957;47:69–75. 23. Balasuriya U, Snijder E, Heidner H, Zhang J, ZevenhovenDobbe J, Boone J, McCollum W, Timoney P, MacLachlan N.
24.
25.
26.
27.
28.
29.
30.
31. 32.
33.
34.
35. 36.
37.
38.
39.
1257
Development and characterization of an infectious cDNA clone of the virulent Bucyrus strain of equine arteritis virus. J Gen Virol 2007;88:918–24. Holyoak GR, Balasuriya UBR, Broaddus CC, Timoney PJ. Equine viral arteritis: current status and prevention. Theriogenology 2008;70:403–14. Heinrich A, Riethmuller D, Gloger M, Schusser GF, Giese M, Ulbert S. RNA interference protects horse cells in vitro from infection with equine arteritis virus. Antiviral Res 2009;81:209–16. Timoney PJ, McCollum WH. Equine viral arteritis: further characterization of the carrier state in stallions. J Reprod Fertil Suppl 2000;56:3–11. Little TV, Holyoak GR, McCollum WH, Timoney PJ. 1992. Output of equine arteritis virus from persistently infected stallions is testosterone-dependent. In: Plowright W, Rossdale PD, Wade JF (eds) Proceedings of the 6th International Conference on Equine Infectious Diseases, Cambridge, 1991. Newmarket: R & W Publications, 1992; pp. 225– 229. Turkstra JA, van der Meer FJ, Knaap J, Rottier PJ, Teerds KJ, Colenbrander B, Meloen RH. Effects of GnRH immunization in sexually mature pony stallions. Anim Reprod Sci 2005;86(3–4): 247–59. Morrell JM, Geraghty RM. Effective removal of equine arteritis virus from stallion semen. Equine Vet J 2006;38: 224–9. van der Meer FJ, de Haan CA, Scuurman NM, Haijema BJ, Peumans WJ, Van Damme EJ, Delputte PL, Bamzarini J, Egberink. Antiviral activity of carbohydrate-binding agents against Nidovirales in cell culture. Antiviral Res 2007;76:21–9. Timoney PJ, McCollum WH. Equine viral arteritis. Vet Clin North Am Equine Pract 1993;9:295–309. Raz T, Corrigan M, Loomis P, Card C. Effects of equine arteritis-positive semen on mare fertility. Anim Repro Sci 2006;94:112–14. Timoney PJ. Aspects of the occurrence, diagnostics and control of selected venereal diseases of the stallion. In: Proceedings of the Annual Meeting of the Society of Theriogenology, Baltimore, MD, 1998, pp. 76–83. Guthrie AJ, Howell PG, Hedges JF, Bosman AM, Balasuriya UB, McCollum WH, Timoney PJ, MacLachlan NJ. Lateral transmission of equine arteritis virus among Lipizzaner stallions in South Africa. Equine Vet J 2003;35:596–600. Timoney PJ. Equine Viral Arteritis. Ithaca, NY: International Veterinary Information Service, 2002. Newton J, Geraghty R, Castillo-Olivares J, Cardwell J, Mumford J. Evidence that use of an inactivated equine herpesvirus vaccine induces serum cytotoxicity affecting the equine arteritis virus neutralisation test. Vaccine 2004;22 (29–30): 4117–23. Nugent J, Sinclair R, deVries AA, Eberhardt RY, CastilloOlivares J, Davis Poynter N, Rottier PJ, Mumford JA. Development and evaluation of ELISA procedures to detect antibodies against the major envelope protein (GL ) of equine arteritis virus. J Virol Methods 2000;90:167–83. Go Y, Wong S, Branscum A, Demarest V, Shuck K, Vickers M, Zhang J, McCollum W, Timoney P, Balasuriya U. Development of a fluorescent microsphere immunoassay for detection of antibodies specific to equine arteritis virus and comparison with the virus neutralization test. Clin Vaccine Immunol 2008;15:76–87. Timoney PJ, McCollum WH, Zhang J, Shuck KM. Diagnosis of the carrier state in stallions persistently infected with equine arteritis virus. In: Proceedings of the International
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41.
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43.
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48.
49.
50.
51.
52. 53.
54.
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Workshop on the Diagnosis of Equine Arteritis Virus Infection, Lexington, KY, October 20–21, 2004, p. 28. Treiberg Berndtsson L, Thoren P, Baule C, Kingeborn B, Belak S. A stallion without detectable humoral antibodies excreting equine viral arteritis in semen. In: Proceedings of the International Workshop on the Diagnosis of Equine Arteritis Virus Infection, Lexington, KY, October 20–21, 2004, p. 33. Timoney PJ. Equine viral arteritis. In: Commission in OS (ed.) Office International Epizooties: Manual of Standards Diagnostic Tests and Vaccines. Paris: Office International des Epizooties, 2000. Moore BD, Balasuriya U, Nurton JP, McCollum WH, Timoney PJ, Guthrie AJ, Maclachlan NJ. Differentiation of strains of equine arteritis virus of differing virulence to horses by growth in equine endothelial cells. Am J Vet Res 2003;64: 779. MacLachlan N, Balasuriya U, Davis N, Collier M, Johnston R, Ferraro G, Guthrie A. Experiences with new generation vaccines against equine viral arteritis, West Nile disease and African horse sickness. Vaccine 2007;25:5577–82. Cardwell JM, Newton JR Wood JL. NE Serologic response to equine arteritis vaccination of stallions. In: Proceedings of the British Equine Veterinary Association 41st Congress, Glasgow, September 11–14, 2002, p. 202. Clausen PH, Chuluun S, Sodnomdarjaa R, Greiner M, Noeckler K, Staak C, Zessin KH, Schein E. A field study to estimate the prevalence of Trypanosoma equiperdum in Mongolian horses. Vet Parasitol 2003;115:9–18. Zablotskij VT, Georgiu C, de Waal T, Clausen PH, Claes F, Touratier L. The current challenges of dourine: difficulties in differentiating Trypanosoma equiperdum within the subgenus Trypanozoon. Rev Sci Tech 2003;22:1087–96. ¨ Claes F, Buscher P, Touratier L, Goddeeris BM. Trypanosoma equiperdum: master of disguise or historical mistake? Trends Parasitol 2005;21:316–21. Aurich C, Spergser J, Nowotny N, Rosengarten R, Aurich JE. Prevalence of venereal transmissible disease and relevant potentially pathogenic bacteria in Austrian Noriker Draught horse stallions. Wien Tierarztyl Mschr 2003;90:124–30. Aurich C, Spergser J. Influence of genitally pathogenic bacteria and gentamicin on motility and membrane integrity of cooled-stored stallion spermatozoa. Anim Reprod Sci 2006;94:117–20. Kikuchi N, Iguchi I, Hiramune T. Capsule types of Klebsiella pneumoniae isolated from the genital tract of mares with metritis, extra-genital sites of healthy mares and the genital tract of stallions. Vet Microbiol 1987;15:219–28. Gierczyski R, Jagielski M, Rastawicki W, Kauzewski S. Multiplex-PCR assay for identification of Klebsiella pneumoniae isolates carrying the cps loci for K1 and K2 capsule biosynthesis. Pol J Microbiol 2007;56:153–6. Samper J, Tibary A. Disease transmission in horses. Theriogenology 2006;66:551–9. Kleiboeker SB, Chapman RK. Detection of equine herpes virus 3 in skin lesions by polymerase chain reaction. J Vet Diag Invest 2004;16:74–9. Seki Y, Seimiya YM, Yaegashi G, Kumagai S, Sentsui H, Nishimori T, Ishihara R. Occurrence of equine coital exanthema in pastured draft horses and isolation of equine herpesvirus 3 from progenital lesions. J Vet Med Sci 2004;66:1503–8.
55. Pascoe R. The effect of equine coital exanthema on the fertility of mares covered by stallions exhibiting the clinical disease. Aust Vet J 1981;57:111–44. 56. Gleeson LJ, Sullivan ND, Studdert MJ. Equine herpesviruses type 3 as an abortigenic agent. Aust Vet J 1976;52:349. 57. Lu KG, Morrese PR. Infectious diseases in breeding stallions. Clin Tech Equine Pract 2007;6:285–90. ˜ S, 58. Barrandeguy M, Vissani A, Olguin C, Becerra L, Mino Pereda A, Oriol J, Thiry E. Experimental reactivation of equine herpes virus-3 following corticosteroid treatment. Equine Vet J 2008;40:593–5. 59. Tearle JP, Smith KC, Boyle MS, Binns MM, Livesay GJ, Mumford JA. Replication of equid herpesvirus-1 (EHV-1) in the testes in epididymes of ponies and venereal shedding of infectious virus. J Comp Pathol 1996;115:385–97. 60. Blunden AS, Smith KC, Whitwell KE, Dunn KA. Systemic infection by equid herpesvirus-1 in a Grevy’s zebra stallion (Equus grevyi) with particular reference to genital pathology. J Comp Pathol 1998;119:485–93. 61. Carvalho R, Passos LM, Oliveira, AM, Henry M, Martins AS. Detection of equine herpesvirus 1 DNA in a single embryo and in horse semen by polymerase chain reaction. Arq Bras Vet Med Zoot 2000;47:351–9. 62. Brown JA, Mapes S, Ball BA, Hodder AD, Liu IK, Pusterla N. Prevalence of equine herpesvirus-1 infection among Thoroughbreds residing on a farm on which the virus was endemic. J Am Vet Med Assoc 2007;231:577–80. 63. Hodder AD, Brown J, Ball BA, Liu IKM, Leutenegger C, Pusterla N. Analysis of equine semen for equine herpes virus-1 using TAQMAN PCR. Theriogenology 2007;68: 506. 64. Hebia I, Fieni F, Duchamp G, Destrumelle S, Pellerin J, Zientara S. Potential risk of equine herpes virus 1 (EHV1) transmission by equine embryo transfer. Theriogenology 2007;67:1485–91. 65. Bermudez VM, Miller RB, Johnson WH, Rosendal S, Ruhnke L. Effect of sample freezing on the isolation of Mycoplasma spp. from the cliltoral fossa of the mare. Can J Vet Res 1988;52:147–8. 66. Spergser J, Aurich C, Aurich J, Rosengar R. High prevalence of mycoplasmas in the genital tract of asymptomatic stallions in Austria. Vet Microbiol 2002;87:119–29. 67. Szeredi L, Tenk M, Schiller I, R´ev´esz T. Study of the role of Chlamydia, Mycoplasma, Ureaplasma and other microaerophilic and aerobic bacteria in uterine infections of mares with reproductive disorders. Acta Vet Hung 2003;51:45–52. 68. Bermudez VM, Miller RB, Rosendal S, Fernando MA, Johnson WH, O’Brien PJ. Measurement of the cytotoxic effects of different strains of Mycoplasma equigeitalium on the equine uterine tube using a calmodulin assay. Can J Vet Res 1992;56:331–8. 69. Mendenbach K, Aupperle H, Schoon D Wittenbrink MM, Schoon D. Pathology of the equine salpynx. Pferdeheilkunde 1999;15:560–7. 70. Henning K, Sachse K, Sting R. Demonstration of Chlamydia from an equine abortion. Dtsch Tierarztl Wochenschr 2000;107:49–52. 71. Baczynska A, Fedder J, Schougaard H, Christiansen G. Prevalence of mycoplasmas in the semen and vaginal swabs of Danish stallions and mares. Vet Microbiol 2007;121:138–43.
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Historical Perspectives of Artificial Insemination John M. Bowen
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Semen Collection Techniques and Insemination Procedures Steven P. Brinsko
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Evaluation of Semen Julie Baumber-Skaife
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Spermatozoal Motility Aaron D.J. Hodder and Irwin K.M. Liu
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Spermatozoal Morphology D.N. Rao Veeramachaneni
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Principles of Cooled Semen James K. Graham
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Breeding With Cooled Transported Semen Juan C. Samper
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Experiences With a Large-Scale Cooled-Transported Semen Program Simon J. Robinson
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Containers for Transport of Equine Semen Terttu Katila
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Semen Extenders for Cooled Semen (Europe) Christine Aurich
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Semen Extenders for Cooled Semen (North America) Steven P. Brinsko
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Factors Affecting Sperm Production and Output Edward L. Squires and Bill W. Pickett
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Reproductive Parameters of Draft Horse, Friesian and Warmblood Stallions Tom A.E. Stout and Ben Colenbrander
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Reproductive Parameters from Light Horse Stallions Edward L. Squires
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Reproductive Parameters from Miniature Stallions Dale L. Paccamonti and Paul J. De Vries
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Chapter 122
Historical Perspectives of Artificial Insemination John M. Bowen
Domestication
Breeding
The history of the early domestication of horses and other Equidae is still being unraveled. It was originally thought that horses were first domesticated by the adoption of young foals by tribes following the hunting and killing of their dam for food. This was alleged to have been in the Caucasus region around 4000–3000 bc.1 More recent investigation has suggested that members of the equine species had also been domesticated in China. It is therefore not beyond the bounds of possibility to envisage that the domestication of horses took place almost simultaneously, around 3500–3000 bc, in four or five different places in the world.2 Young orphan foals would probably have followed, even been encouraged to follow nomadic tribes. Over time, they may have been used to carry goods and people, and eventually ridden into battle and used to draw wagons and chariots around 2000–1750 bc.3 As man settled into an agrarian lifestyle, the horse, along with oxen, became of greater use to the community. Breeding was not supervised and the animals would have been turned out at pasture and any mating happened spontaneously and without any input or control from the humans who were nominally in charge of the horses. Over time, there must have been some selection pressure from their human companions, as it is found that, in certain areas and regions, distinctive types of horses begin to appear. So much so, that four definite types of early horses and ponies are now recognized by paleobiologists. They are the Forest Horse, the Draft type, the Asian type, and the Tarpan. It is alleged that the Bedouin tribes kept records of the bloodlines of their horses, as did the nomads of the Mongolian steppes.2
In Europe it is known that the Irish monks, who settled in the Low Countries around 1100 ad, systematically set about improving the Draft Horses in that area, resulting in such breeds as the Percheron, the Belgian, and the Ardennes.4 Later in the eighteenth century, Robert Bakewell, along with others who promoted and led the British Agricultural Revolution, in addition to greatly and rapidly improving livestock production in the UK, improved the Draft Horse of the day, this was known as Bakewell’s Black Horse, which later became the Shire Horse of today.5 It is interesting to note that this greater interest in improving farm livestock had been preceded by an Islamic Agricultural Revolution during the eighth to thirteenth centuries, which led to modern farming methods of crop rotation and selective breeding of sheep, horses, and camels in the Islamic and Mediterranean regions.6 One factor which mitigated against good fertility in horses was the fact that many horse owners would rather use their mares for work, as opposed to breeding. This meant that many times a useful mare would not be considered as a brood mare until she was past her most fertile years. Putting a mare in foal would mean that that she would be unavailable for work for a considerable period of time. It also meant that land used for the brood mares could not be used to raise cattle and sheep, which were more profitable.7 In addition, mares that were used as brood mares were often those which were unsuitable for any other purpose. John Wilkinson, a renowned horse breeder, noted that these mares would be used ‘disregarding the fact that she may have a large head, ewe-neck, upright shoulders, calf-knees, long crooked pasterns, long slack back, weasel waist, short drooping
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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quarters, short thighs, curby or cow hocks, with dishing, speedy cutting or slouching gait’.8
Artificial insemination: early pioneers The history of artificial insemination in the genus Equus followed a rather long and convoluted pathway. It is surprising to realize that much of the early work on artificial breeding was first carried out in horses. It is alleged that the Arabs carried out artificial insemination in both humans and horses before the birth of Christ, but this cannot be verified. The first recorded incident was in 1322, when an Arab sheikh stole some semen from a rival’s stallion to breed one of his mares.9 The next milestone along this road was in 1780, when an Italian priest turned physiologist named Lazzaro Spallanzani conducted an experiment with frogs which showed that sperm were the essential component of semen required for fertilization. Following this, he inseminated a bitch with dog semen and successfully produced three puppies.10 Spallanzani later went on to successfully inseminate a mare. He also placed some dog semen in snow and found that, while it became immotile, motility was restored for several hours when the sample was warmed up to room temperature. These two experiments laid the foundation for the development of artificial insemination and the use of cooled semen in animal reproduction. Interestingly, the surgeon John Hunter in 1799 instructed a client to collect his ejaculate and inject it into his wife’s vagina, to overcome his infertility.11, 12 From the end of the eighteenth century to the 1870s very little progress was made, though it must be remembered that Europe and the USA were caught up in a series of conflicts during that time. In the 1880s, Walter Heape, the father of embryo transfer, carried out work on the artificial insemination of both dogs and horses. This prompted Professor Pearson, Veterinary Professor, University of Philadelphia, to write to Heape to recount his successes with artificial insemination in mares.9
Disease control In 1890 Repiquet, a French veterinarian, promoted the use of artificial insemination to counter what he referred to as ‘sterility’.13 As a result this practice was commenced in several studs around Europe, without much success. The earliest advocates of artificial insemination were those who favored its use for improving fertility and controlling sterility. Equine fertility in those days was poor, owing to many factors,
including a lack of understanding of the estrous cycle, the management of the mares, and poor estrus detection as well as infections, which included an insidious disease, dourine. Little was known about this disease or its cause, so contemporary literature tends to focus on the use of artificial insemination in an effort to improve fertility in the horse population. Dourine is a venereal disease spread by coitus, which can affect both the stallion and the mare, often to differing degrees, but can eventually result in the death of the animal. The cause was discovered by Rouget, and later confirmed by Schneider and Buffard that a trypanosome was the cause of this disease. They also found that it could be spread venereally.14 Once this was understood, it encouraged the use of artificial insemination in those regions where dourine was a particular problem. This happened to be in Eastern Europe and Russia, stimulating in this region great interest in artificial insemination and development of collection and semen-handling technology. Breed improvement It was not until 1902 when Sand and Stribolt in Denmark, reporting on their success with artificial insemination, promulgated the idea that artificial insemination should also be used as a tool for breed improvement.9 Traditionally the governments of Russia, Poland, Germany, Austria, and others had established equine breeding establishments, many of which were run by the military. They were responsible for producing and maintaining the standards for the desired characteristics of their particular breeds. With the more extensive and highly regulated use of artificial insemination in Eastern Europe to combat disease came the additional benefit of breed improvement. This made the acceptance of artificial insemination relatively straightforward. In contrast, the horse breeders in the USA and the UK took their lead from the Thoroughbred breeders of the time, who preferred their time-honored ways. The British government, unlike its continental counterparts, chose not to get involved in breeding horses. A Select Committee preferred the free market approach,7 leaving it to a nationwide traveling stallion program, in which retired racehorses traveled from town to town breeding mares. The Hunter Improvement Society and the Royal Agricultural Society of England were the two organizations that, in 1885, undertook this task.15 The owners of these stallions saw no need to change their ways as the farmers and landowners were content that their horses, which were being covered at little cost, produced foals which were eminently saleable.15 Indeed, many farmers preferred to have their Draft-type mares covered by a Thoroughbred stallion, as she would then produce a showier
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type of horse, which buyers seemed to prefer over a utilitarian horse.16 So pervasive was the influence of the racing fraternity that artificial insemination was not introduced into the Anglo Saxon sphere of influence until after World War II.
Sport horses, of which there were very few in the USA. As the freezing of stallion semen was not feasible at this time, the development of transported chilled semen came into vogue.
Development
Equipment
The real push to develop artificial insemination in the horse came from Il’ia Ivanovich Ivanov, who, in 1909, was asked by the Russian Royal Stud in St Petersburg to investigate the potential of artificial insemination.9, 17 He set about his task with great vigor, establishing veterinary laboratories under the Ministry of Agriculture, and, even though the initial results were poor, excepting where the stud was under his direct supervision, he persisted to improve the technique. Many people came to Ivanov’s laboratories to be instructed in the collection of semen and its insemination. After World War I he continued his work at a central experiment station.18 Interest in equine artificial insemination also developed in Japan and commenced in 1913 on a small scale. Between the two World Wars, artificial insemination was used extensively in Eastern Europe and Russia with some 300 000 mares being artificially bred in Russia in 1939.19 After World War II, horse breeding with artificial insemination recommenced in earnest, as horses were urgently needed for transportation.20 Farther east, workers began using artificial insemination for horse breeding in China and in Japan. Little was known about this work until Nishikawa21 and Cheng22 published details of the numbers of inseminations that were being carried out. In 1959 some 600 000 mares were inseminated in China.22 The United States Trotting Association permitted the use of artificial insemination, which was regulated in the 1950s to be an on-farm procedure, in which the stallion and the mare had to be present on the farm at the time of insemination (USTA 2008, personal communication). This led to a growth in the industry and to a radical change in the way Standardbred horses were bred. In 1963, the Quarter Horse Association (AQHA 2008, personal communication) also allowed on-farm insemination, an action which aided in the breed’s popularity. Since then, all manner of breed associations have bowed to owner demand for artificial insemination to be permitted within their breed. However the Thoroughbred community has rigidly and steadfastly held to their belief that the use of artificial insemination would be detrimental to their breeding programs and the use and diversity of their stallion population. During the 1970s and 1980s horse owners became interested in some of the Warmblood breeds and in
Around the turn of the century, Professor Hoffman in Stuttgart, Germany, was one of the first to describe how he obtained semen from the vagina of the mare, using a spoon-like scoop, and then using the collected fluid as the inoculum.9 His technique was to become the once popular ‘reinforcement’ of a cover, often used when the stallion was thought not to have completed the mating to the satisfaction of the observers. Others tried placing a sponge in the vagina to collect the ejaculate, but the resulting fluid was highly contaminated, yielding less than optimal results, yet this method persisted until the advent of the artificial vagina. Another early collection technique was the placement of a rubber bag within the vagina of the mare, into which the stallion would ejaculate. Unfortunately, when the stallion withdrew, he would often bring the rubber bag out with him and the ejaculate would be lost or severely contaminated. Many workers in Germany, Denmark, and the USA also tried using a rubber collection bag, often in conjunction with a vaginal retaining rubber ring, but this too was subject to failure. Modifications to this method were tried, but the development of the artificial vagina rendered these methods obsolete. The first artificial vagina was invented by Professor Amantea of Rome, Italy, for use in the collection of dog semen in 1914.9 For the stallion, Ivanov and his group, perhaps Milovanov, is credited with the development of the equine artificial vagina. Milovanov and his assistants developed the forerunners of today’s artificial vaginas in their own workshop, creating a much more suitable method of semen collection than the sponge or the rubber bag.23 It is also highly likely that they developed the concept of the phantom or dummy mare so frequently used today.19 Latex condoms were also designed and used for semen collection, but they were often removed in the act of coitus, or fell off the penis, with disastrous results, when the stallion dismounted. Following the development of the Russian artificial vagina, other models were developed in Hanover, Germany (Goetze), and Cambridge, UK (Walton). These large and somewhat clumsy models were followed by a light aluminum Japanese model (Nishikawa). In the USA, a Missouri model, which was light and self-contained, was produced as well as the heavy Colorado model (Pickett).24 The Missouri
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model, being light and inexpensive, has come to be the most commonly used artificial vagina. One advantage is that, owing to its low cost, it allows an artificial vagina to be purchased, marked, and retained for each stallion, thereby further reducing the chances of cross-infection due to inadequate cleansing. As artificial insemination became more widespread in Standardbred and Quarter Horse breeding, the use of a phantom or dummy to eliminate the need for a mount mare became desirable and safer for the stallion. Originally made from two oil drums suspended on posts, the phantom matured into a device into which the artificial vagina could be placed, or one in which the collection device was an integral part of the phantom.24
Handling of semen At first any semen which was collected was used without any extender, as raw semen. Michajilov25 had the bright idea of adding heated and cooled mare’s milk to the collected semen, as this allowed the semen to be split and used in several mares. Having found that this technique had no detrimental effect on the semen motility over a short period of time, this encouraged people to look for other possible diluents which might be capable of prolonging the viable life of the sperm. In addition it was found that diluting the semen gave better conception rates than could be obtained with less dilute samples.26 This was later recognized to be due to the dilution of bacterial pathogens. Pasteurized mare’s milk was replaced by the more available preheated cow’s milk, and this, with the addition of glucose, became a more common extender. Electrolytes such as sodium citrate, used to extend bull semen, have a detrimental effect on stallion spermatozoa13 and it was some time before workers realized that such an extender was not the one of choice for the stallion. The use of egg yolk appeared quite early on in the development of semen extenders and egg yolk/lactose and egg yolk/glucose were among the early semen extenders used.27 Vlachos and coworkers28, 29 in Greece did a great deal of comparative work with semen extenders and concluded that skim milk improved conception rates over the use of an egg yolk diluent. The advent of antibiotics brought about a marked improvement in bull fertility. Once this became known, the addition of antibiotics to semen extenders became widespread, both to reduce transmissible pathogens and to aid in preserving the extender. In horses, penicillin was the first to be used, then in conjunction with streptomycin. Later these were replaced by gentamicin and ticarcillin with
the intention that they would combat certain specific pathogens such as Klebsiella pneumoniae. Eventually the egg yolk extenders were replaced by skimmilk extenders, of which there are many, or by the cream/gel extender developed in California.30 The cream/gel extenders, while giving excellent conception rates, have the disadvantage that, once the semen is extended, it is impossible to evaluate the motility of the spermatozoa under a microscope. Today it is possible to buy pre-packaged skim-milk equine semen extenders, to which only sterile water need be added.24 The popularity of the use of chilled stallion semen came about from the early inability to reliably freeze a stallion’s semen. A physicist, at his wife’s behest, designed a container that would allow semen to cool slowly within the container, while at the same time protecting the contents from either extreme heat or extreme cold.31 The unit, the EquitainerTM ,32 is used extensively to transport chilled liquid stallion semen anywhere within and even between countries thanks to the development of the air freight courier services which came into being after 1971.33 Owners of sought-after stallions such as the Hanoverian and Trakehner breeds were able to offer, for a fee, doses of semen which could be collected and air freighted for same day insemination. It was the responsibility of the owners and their veterinarian to ascertain the optimal time to breed the mare. The results of such a service proved to be acceptable and this service is still available to those who need it. Polge and Parkes,34 working in Cambridge, had fortuitously found that glycerol would act as a cryoprotective agent. This discovery revolutionized the nascent cattle breeding industry.35 Glycerol was added to the existing effective extenders such as the egg yolk/citrate and the milk diluents, allowing bull studs and breeding centers to freeze semen with relative ease. This provided for the long-term storage of semen. All this was of little interest to the horse industry, which had regulated that artificial insemination could only be used on-farm, if it was used at all. Despite this philosophy, a few people persevered in trying to freeze stallion semen, but it was proving to be a much more difficult technical bridge to cross. Many of the successful extenders used to freeze bull semen proved to be unsatisfactory for stallion sperm. Again the reliance on the very well-developed techniques and extenders used to freeze bull semen proved to be a stumbling block for the stallion. The methods used to freeze bull semen did not yield acceptable post-thaw results and, when used with stallion semen, their ability to obtain pregnancies in mares was low.36 Some experiments into freezing bull semen at a higher freezing rate than had been the
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Historical Perspectives of Artificial Insemination norm led to the idea that perhaps stallion semen would benefit from a more rapid freezing rate. Another problem was that some stallions’ semen would seem to freeze well in one extender, while other stallions’ would not. Some stallions’ sperm demanded a slow freezing rate, whereas others were better when flash frozen.37 The earliest successes came with pelleted semen, in which semen was centrifuged to remove the seminal plasma and resuspended in a small quantity of cryoprotective extender. This condensed mixture was then dripped onto a block of dry ice, on which small indentations had been created by a heated, dimpled, metal plate or iron. The resulting pellets, which froze extremely rapidly on the dry ice surface, were then transferred into a shallow dish containing liquid nitrogen and then placed into marked plastic vials and capped, prior to being placed in a liquid nitrogen container.13 This method proved to have two problems. First, pelleted bull semen had been shown to deteriorate at a faster rate than bull semen stored in glass ampoules, and, second, it was difficult to mark and identify which stallion’s semen was frozen in the pellets.38 Workers in Germany later showed better storage results with pelleted stallion semen.39 Other methods of freezing stallion semen were attempted. Some, such as the use of sachets, were not successful (L.E.A. Rowson 1963, personal communication). Using a technique developed in Russia, small aluminum packets containing around 15 mL of extended semen were equilibrated in a refrigerator and then frozen in liquid nitrogen vapor in the neck of a wide neck liquid nitrogen container, before being plunged into the liquid nitrogen.40 Gradually, the French straw, ‘paillette Cassou’,41 supplanted the glass ampoule in the freezing of bull semen. This 0.5-mL or 0.25-mL plastic straw could be easily identified, as the straws could be of differing colors and printed with the name of the sire and other information. This, together with the different method used to freeze these straws, reawakened an interest in freezing stallion semen. Work was begun in both Germany and the USA. The Hanover group42 initially used standard 4.0-mL drinking straws with metal ball-bearings to seal the ends, whereas, in the USA, the 0.5-mL French straw was favored. It seems that, in spite of a great deal of effort, there is still considerable variability in the success rate with frozen stallion semen. Some of this is due to differences among stallions, while in other cases it can be blamed on poor semen transportation and handling. The uncertain freezability of stallion semen was partially overcome by the removal of seminal plasma and resuspension of the sperm in a seminal plasma-free extender.43, 44 Other work indicated that the spermatozoa themselves, in the semen of those stallions
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whose sperm do not freeze well, may be more susceptible to acrosomal damage.45 To make matters more confusing, it has been shown that, by removing the seminal plasma by centrifugation from a stallion whose sperm do not freeze well and placing these sperm in the seminal plasma of one whose sperm do freeze well, freezability will improve. This suggests that the seminal plasma has some effect on the freezability of spermatozoa.46 To add to our difficulties, the way frozen semen is thawed has greatly affected the success of its use. Many of those who are asked to inseminate a mare with frozen semen have little experience in handling material frozen and stored in liquid nitrogen. When thawing frozen semen of any kind, it is essential to follow the instructions given by the laboratory which froze that particular batch of semen, thus assuring that optimal results may be obtained. Mishandling the straws or pellets can be disastrous. The need to develop a reliable method of freezing stallion semen was in part stimulated by breeders becoming increasingly interested in the Warmblood breeds from continental Europe. The rigid regulations posed by the breed societies in Europe meant that it would be easier to import the semen than the sires, as well as greatly reducing the costs involved. Whether equine artificial insemination will reach the same level of acceptance and efficiency that cattle insemination has attained is still somewhat in doubt. There is no doubt that it is extremely useful for the international movement of stallion semen, but the difficulty in handling and thawing the semen has posed problems for the industry. It is unlikely to attain a comparable level of acceptance and use due to the uncertain results. Much work is still to be done in order to overcome the problems of freezing and thawing stallion semen successfully and on a routine basis.
Technique of insemination Techniques are a great deal more sophisticated than in the days when stallion semen was collected from the vagina with a spoon and placed into the vagina via a funnel and a flexible rubber tube. Another popular method used was to put 10–15 mL of semen into a large gelatin capsule, which was then manually placed in the vagina or in some cases introduced into the uterus through the mare’s generous and flaccid cervix.47 Others used a flexible rubber or soft plastic tube with a syringe or bulb on the end to administer the semen into the uterus of the mare.48 This could be introduced by means of a stylet, which would be removed to allow the passage of the semen.13 Still others would use a Vulsellum forceps to grasp the cervix
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and then, using a speculum, pass a catheter into the uterus to deposit the semen.49 As people became more familiar with the technique of insemination and appearance on the market of the disposable inseminating rod, it was very easy to adapt to using this rigid plastic rod by means of introducing an arm covered by a sterile plastic sleeve to dilate the cervix sufficiently to introduce the rod.50 The cattle rods proved to be too short, but, now that the equine breeding industry has accepted artificial insemination to a greater degree, longer equine inseminating rods are available. The site that the semen is deposited is typically in the uterine body This technique followed the method established for cattle insemination. All the time that horses have been bred by artificial insemination there has been a need, especially in the Standardbred and Quarter Horse breeds, to cover as many mares as possible with each semen collection. To this end, workers have been trying to reduce the number of live sperm that are needed to effect a pregnancy. In the 1970s work in Colorado suggested that a minimum number of live sperm per dose to obtain optimal pregnancy was approximately 500 million (500 × 106 ).51 However, Chinese reports from an earlier date had clearly shown that equine insemination rates could be improved by directing the semen into the uterine horn on the side where the follicle was maturing. This work showed that the live sperm dose could be reduced to 100 million (100 × 106 ) without a drop in the pregnancy rate.22
Newer developments With the arrival of newer equipment such as the hysterscope came the concept of depositing small discrete doses of semen directly at the utero-tubal junction. This technique has proved to be very successful. Workers in Newmarket, UK,52 have shown that normal fertility rates can be obtained with as few as 5 million live spermatozoa (5 × 106 ) and that pregnancy rates fall only when the numbers are reduced to 1 million sperm (1 × 106 ). This has affected the way stallion semen can be frozen. Now a full dose for one mare can be easily accommodated in a 0.5mL or 0.25-mL French straw. It does, however, require some sophisticated and expensive equipment not readily available on breeding farms. While this technique gives normal conception rates with fewer sperm numbers, it does not improve the fertility of infertile stallions,52 though it may prove to be of use in those stallions whose sperm do not possess great longevity. Apart from conservation of frozen semen from stallions with little semen available it has been useful in stallions that produce low numbers
of spermatozoa that have large numbers of mares to breed. This low-dose insemination technique is particularly suited for the frozen semen of certain prized stallions whose semen does not freeze well. Cases where the stallion’s post-thaw sperm numbers are particularly low, thus reducing the chances of obtaining pregnancies by the usual method of insemination, are clear candidates for this technique. This low-dose technique also lends itself to the deposition of sexsorted semen, as the number of spermatozoa available following treatment by flow cytometry are relatively small.53 Society and its equine enthusiasts have come a long way since that Bedouin stole semen from a much-prized stallion. Much physiology has been learnt and we have to thank Ivanov54 for his tenacity in developing artificial insemination to the point where it became a useful tool to combat disease and infertility. It is unfortunate that his later work in Africa brought ignominy on his memory.55 There are many others we have to thank for the many incremental steps which have brought us to where we are today. The future will no doubt bring further improvements and benefits to the horses we all admire.
References 1. Goodall DM. A History of Horse Breeding. London: Robert Hale, 1977. 2. Bennett D. Conquerors: The Roots of New World Horsemanship. Los Olivos, CA: Amigo Publications Inc., 1998 3. Kuznetsov PF. The emergence of Bronze Age chariots in eastern Europe. Antiquity 2006;80:638–45. 4. Fraser AF. The Behaviour of the Horse. Wallingford, UK: CAB International, 1974. 5. Bewick T. A General History of Quadrupeds. Trowbridge & London: Redwood Press Ltd, 1790 6. Stiles PR. Animal Husbandry. In: Meri JW (ed.) Mediaeval Islamic Civilization: An Encyclopedia. New York, NY: Taylor & Francis, 2006. 7. House of Lords, Select Committee 1873. Condition of this country with regard to horses. British Parliamentary Proceedings XIV, q 419. 8. Wilkinson J. On the supply of horses adapted to the requirements of the English army, with notes on remount system in the French army. J R Agric Soc Engl 1863;24:91–5. 9. Perry EJ. The Artificial Insemination of Farm Animals. New Brunswick, NJ: Rutgers University Press, 1968. 10. Spallanzani Abate. Dissertazioni di Fisica Animale, e Vegetabile. Modena: Presso La Societa Tipografica, 1780. 11. Guttmacher AF. Artificial insemination. Ann N Y Acad Sci 1962;97:623–31. 12. Foote RH. Artificial insemination from the origins up to today. In: Russo L, Dall’Olio S, Fontanesi L (eds) Proceedings of the International Symposium. Italy: Internet-First University Press, 1999; pp. 23–67. 13. Bowen JM. Artificial insemination in the horse. Equine Vet J 1969;1:98–110.
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Historical Perspectives of Artificial Insemination 14. Horace Hayes M. Veterinary Notes for Horse Owners, 6th edn. London: Hurst and Blackett, 1903. 15. Moore-Colyer R. Aspects of horse breeding and the supply of horses in Victorian Britain. Ag Hist Rev 1995;43: 47–52. 16. Youatt W. The Horse with a Treatise on Draught 1831. Published under the Superintendence of the Society for the Diffusion of Useful Knowledge; Baldwin & Cradock, London, 1831;30. 17. Zvereva GV. Professor I.I. Ivanov: founder of artificial insemination of animals. Veterinaria 1970;7:88– 90. 18. Ivanoff EI [Ivanov]. On the use of artificial insemination for zootechnical purposes in Russia. J Agric Sci 1922;12:244–56. 19. Milovanov VK. Artificial insemination of livestock in the U.S.S.R. Trans. by Birron A, Cole ZS, Monson S, Jerusalem Tech. Services. Washington, DC: US Dept. Commerce, 1964. 20. Tischner M. Artificial insemination in horses: a review of current status quo. Equine Vet Ed 1992;4:89–92. 21. Nishikawa Y. Studies on reproduction in horses. Kyoto: Koei, 1959. 22. Cheng PL. The present situation of artificial insemination in horses in China and some investigations on increasing conception rates of mares and breeding efficiency of stallions. Acta Vet Zootechnol Sen I 1962;5:29–34. 23. Foote RH. The history of artificial insemination: selected notes and notables. J. Anim Sci 2002;80:1–10. 24. Davies Morel MCG. Equine Artificial Insemination. Wallingford, UK: CAB International, 1999. 25. Michajilov NN. Sperm dilution in the milk. The CzechoSlovak Vet. Mag. J Am Vet Med Assoc 1950;117:337. 26. Arhipov G. The dosage of semen in the artificial insemination of mares. Konevodstvo 1957;27:33. 27. Phillips PH, Lardy HA. A yolk-buffer pabulum for the preservation of bull semen. J Dairy Sci 1940;23:399–404. 28. Vlachos K. Artificial insemination in mares in Greece and the use of glycerine for the conservation of stallion semen. Coll Vet Sci Budapest Summ Pap.sci.sess. 175th Anniv., 1962. 29. Vlachos K. Investigation of the raising of fertility of semen of Solipeds by the use of different diluents. Bull Physiol Pathol Reprod Artif Insem 1966;2:66. 30. Hughes JP, Loy RG. Artificial insemination in the equine. A comparison of natural breeding and artificial insemination of mares using semen from six stallions. Cornell Vet 1970;60:463–75. 31. Hamilton Research. Company history. See http://www. equitainer.com/Company/History.htm, 2008. 32. Douglas-Hamilton DH, Osol R, Osol G, Driscoll D, Noble H. A field study of the fertility of transported equine semen. Theriogenology 1984;22:291–304. 33. FedEx. Company history. See http://www.fedex.com/us/ about/today/history/, 2008. 34. Polge C, Smith AU, Parkes AS. Revival of spermatozoa after vitrification and dehydration at low temperatures. Nature 1949;164:666. 35. Smith AU, Polge C. Survival of spermatozoa at low temperatures. Nature 1950;166:668–9.
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36. Barker CAV, Gaudier JCC. Pregnancy in a mare resulted from frozen epididymal spermatozoa. Can J Comp Med Vet Sci 1957;21:47–53. 37. Loomis PR. The equine frozen semen industry. Anim Reprod Sci 2001;68(3–4): 191–200. 38. Nagase H, Soejima A, Niwa T, Oshida H, Sagara Y, Ishizaki N, Hoshi S. Studies on the freezing storage of stallion semen. Fertility results of semen in concentrated pellet form. Jpn J Anim Reprod 1966;12:48–51. 39. Merkt H, Klug E, Krause D, Bader H. Results of long-term storage of stallion semen frozen by the pellet method. J Reprod Fertil Suppl 1975;23:105–6. 40. Love CC, Loch WL, Bristol F, Garcia MC, Kenny RM. Comparison of pregnancy rates achieved with frozen semen using two packaging methods. Theriogenology 1989;31:613–22. 41. IMV Group. Our history: precursor for artificial insemination. See http://www.imvusa.com/dnn/Racine/Menu/ GroupeIMV HistoirePr%C3%A9curseurdelins%C3%A9minationanimale/tabid/79/language/en-US/default.aspx, 2008. 42. Klug E, Robbelen I, Kneissl S, Sieme H. Cryopreservation of stallion spermatozoa. Acta Vet Scand Suppl 1992;88:129–35. 43. Buell JR. A method for freezing stallions semen and tests for its fertility. Vet Rec 1963;75:900–2. 44. Rajamannan AHJ, Zemjanis R, Ellery J. Freezing and fertility studies with stallion semen. In: Proceedings of the 2nd International Congress on Animal reproduction and Artificial Insemination, 1968, pp. 1601–4. 45. Amann RP, Pickett BW. Principles of cryopreservation and a review of cryopreservation of stallion spermatozoa. Equine Vet J 1987;7:145–73. 46. Aurich JE, Kuhne A, Hoppe H, Aurich C. Seminal plasma affects membrane integrity and motility of equine spermatozoa after cryopreservation. Theriogenology 1996;46:791–7. 47. Swire PW. Artificial insemination in the horse. In: Maule JP (ed.) The Semen of Animals and Artificial Insemination. Farnham, UK: Commonwealth Agricultural Bureau, 1962; pp. 281–97 48. Berliner VR. Horses and jackstock. In: Perry EJ (ed.) The Artificial Insemination of Farm Animals. New Jersey: Rutgers University Press, 1947; pp. 99–132. 49. David J. Discussion at B.E.V.A. Annual Conference, Bristol 1968: artificial insemination in the horse. Equine Vet J 1969;1:107. 50. Day FT. The stallion and fertility. Vet Rec 1940;52:597–602. 51. Pickett BW, Back DG. Procedures for Preparation, Collection, Evaluation and Insemination of Stallion Semen. Bulletin of the Colorado State University 935. Fort Collins, CO: Animal Reproduction Laboratory General Services, Agricultural Experiment Station, 1973. 52. Morris LH, Allen WR. An overview of low dose insemination in the mare. Reprod Domest Anim 2002;37:206–10. 53. Morris LHA. Challenges facing sex preselection of stallion spermatozoa. Anim Reprod Sci 2005;89:147–57. 54. Wikipedia. Ilya Ivanovich Ivanov (biologist). See http:// en.wikipedia.org/wiki/Ilya Ivanovich Ivanov (biologist), 2008.
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Chapter 123
Semen Collection Techniques and Insemination Procedures Steven P. Brinsko
Introduction Collection of semen is an integral part of artificial insemination programs and breeding soundness evaluations for stallions. Proper collection techniques should be employed so as help ensure safety of the horse and personnel as well as to minimize iatrogenic or environmental alterations in semen quality. The history of semen collection and the development of semen collection methods have been reviewed previously.1
Semen collection methods A number of semen collection methods exist, all with advantages and disadvantages that are dependent on the situation at hand. These include (i) use of an artificial vagina, (ii) use of a condom, (iii) pharmacologically induced ejaculation, (iv) manual stimulation, and (v) epididymal sperm harvest. Although electroejaculation has been performed in horses, it requires heavy sedation or general anesthesia and has not been considered a desirable method of semen collection due to inconsistent results and a number of complicating factors that include urine contamination, retrograde ejaculation, poor sperm motility,2 and rectal perforation (R.M. Kenney, personal communication). However, an improved method that was very successful in eight out of nine Przewalski’s horses has been described.3 The one stallion that was not able to provide a sperm-rich fraction had undergone testicular trauma and probably was azoospermic.3 It is possible that in the future electroejaculation will be re-evaluated with improved results in domestic horses. When ejaculates are obtained with the use of an artificial vagina a variety of mount sources can be
used (estrous mare, ovariectomized mare, phantom), or no mount source may be employed in the case of pharmacological ejaculation or manual stimulation. Regardless of which method is used, the collection area should have minimal distractions, be dust-free, and provide adequate space and good footing for the horse and personnel.
The artificial vagina The most widely used method of semen collection from stallions is via the use of an artificial vagina (AV). There are several models of equine AVs available from various manufacturers. In the USA, the most commonly used AVs are the Missouri, Colorado State, and Japanese (Nishikawa)/HarVet models (Figure 123.1). The HarVet model closely resembles the Japanese model, which is no longer available commercially in the USA, but has a plastic rather than an aluminum casing. Descriptions of various models including details of their construction, advantages, disadvantages, and proper methods for cleaning and storage have been reviewed previously.1, 4 AVs should be cleaned and disinfected after each use. All components which come in contact with semen must be non-spermicidal; therefore, it is imperative that reusable items be chemically clean. Hot running water is usually sufficient to clean most AV liners, but, if the use of soap becomes necessary, it should be a non-residual type such as Alconox (Alconox Inc., New York, NY, USA). Disinfectant solutions or soaps containing disinfectants (e.g., povidone–iodine or chlorhexidine) may damage spermatozoa. Povidone–iodine concentrations as low as 0.05% have been shown to render spermatozoa completely immotile within 1 minute of contact.5 After the liners have been cleaned and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 123.1 Artificial vaginas (back to front): CSU model, Missouri model, and Japanese (Nishikawa) model. Figure 123.2 Missouri model artificial vagina (AV) with towel inserted between AV liner and case for added stimulation.
thoroughly rinsed, they should be submerged in ethyl or isopropyl alcohol for at least 30 minutes and allowed to air dry in a dust-free cabinet. Gas sterilization with ethylene oxide can be used, provided that the liners are allowed to air out for 48–72 hours prior to use. Sterile, non-toxic, disposable liners are available which fit most AVs. Disposable equipment simplifies cleaning and reduces risk of chemical contamination and horizontal transmission of disease with repeated use of the AV. However, some stallions dislike the texture of these disposable liners and will not readily breed the AV when they are in place. Virtually all AVs work on the same functional principle, in that they have a water jacket that can be filled with various amounts of water or air to adjust the temperature (typically 44–50◦ C) and pressure sufficient to stimulate the stallion’s penis to achieve ejaculation. Initially, the AV is overfilled in order to maintain temperature and the pressure is adjusted by releasing water and/or air just prior to semen collection. Before the AV is filled, a small amount of nonspermicidal lubricant is placed into the proximal end of the AV and the entire surface to be in contact with the penis is lightly coated to modify the amount of friction to the stallion’s penis. Excessive amounts of lubricant should be avoided as this can be detrimental to semen quality owing to alterations in pH and osmolarity. Ideally, an in-line, nylon micromesh filter (Animal Reproduction Systems, Chino, CA, USA) should be placed in the collection receptacle to trap debris and gel. Alternatively, the semen can be filtered in the laboratory, but this is less desirable as it results in increased exposure of the sperm-rich semen to contaminants and gel, as well as a greater potential loss of the sperm harvest. After the stallion has been teased to the mount source, the erect penis is washed with clean warm water, paying particular attention to remove smegma and debris from the fossa glandis, in order to reduce
contamination of the ejaculate. The stallion is then allowed to approach the mount source with the stallion handler and collector on the same side of the horse. Once the stallion has mounted, the penis is deflected into the AV, which is held at mid-hip level of the mount source at an angle parallel to the stallion’s abdomen. Allowing the distal end of the AV to drop downward can result in reduced thrusting and ejaculation failure. The warmth and pressure of the properly aligned AV will induce the stallion to thrust forward and couple with the mount source. Ideally, the glans penis should be as far as possible into the AV so that, when ejaculation occurs, the semen has minimal exposure to the elevated temperature of the water jacket. The AV should be held firmly against the mount source with one hand, maintaining the correct angle and minimizing movement, while the other hand is placed palm up at the base of the stallion’s penis, close to the scrotum. This allows the collector to feel the ejaculatory pulses as well as to apply gentle but firm base pressure to the penis of stallions that require additional stimulation. The design of the Missouri model AV allows the placement of a towel at the proximal end of the AV between the water jacket and the leather case (Figure 123.2). The towel can be saturated with hot water and used as a hot compress to provide added stimulation and base pressure for stallions that are otherwise hesitant to ejaculate. Care must be taken not to allow contamination of the ejaculate with water. Once the stallion begins to ejaculate the distal end of the AV is slowly lowered to facilitate drainage of the ejaculate into the semen receptacle. The AV is removed from the stallion’s penis and the in-line filter is removed promptly to minimize contamination of the sperm-rich semen with gel and contaminants. The semen is immediately taken to the laboratory for analysis and processing (see Chapter 124).
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Figure 123.3 Missouri model artificial vagina (AV) modified to mimic the Polish (Krakow) model AV.
Figure 123.4 Mount mare with protective shroud, tail tie, and padded boots.
For the vast majority of situations, a full-length AV is used. However, the Missouri and Colorado State model AVs can be modified by shortening them to mimic the Polish or Krakow model AV (Figure 123.3), which allows the glans penis to extend beyond the distal end of the AV during the collection process. This type of AV is useful for diagnostic purposes, such as determining when in the ejaculatory process conditions such as hemospermia or urospermia are occurring as well as facilitating fractionation of the ejaculate for semen freezing or other special processing needs. When using an open-ended AV, another person, in addition to the one handling the AV, is needed for observation and collection of the separate semen fractions.
mares may be intact, or ovariectomized. Ideally, mount mares should be mature, docile, and not be intimidated by aggressive, vocal stallions. They should stand firmly when mounted and tolerate a certain amount of biting and nipping by the stallion. Protective shrouds can be placed on the neck and withers of mares to minimize any pain or trauma associated with these activities (Figure 123.4). Breeding hobbles have long been used to inhibit the mare from kicking the stallion; however, they suffer from the drawback that the legs of the stallion can become entangled in the ropes of the hobbles if mounting or dismounting does not go smoothly. In addition, mares may react violently to the use of hobbles if not accustomed to their application, and can suffer injury. A safe alternative is to use heavily padded boots on the mare’s hind feet to soften the blow if the mare decides to kick (Figure 123.4). The mount mare’s tail should be wrapped to protect the stallion’s penis from lacerations as well as not to interfere with deflection and entry of the penis into the AV. It can also be tied snugly against the perineum to impede intromission (Figure 123.4). The disadvantage of relying on intact, cyclic mares is that one may not always be sexually receptive at the time that semen needs to be collected. In addition, not all mares in physiological or behavioral estrus will tolerate the restraint and other activities associated with the collection procedure. Wellselected ovariectomized mares have the advantage of being more predictable throughout the year in their behavior toward the stallion and the collection process. Only those mares that show desirable behavioral characteristics while intact and in estrus should be selected as ovariectomized mount sources. If selected properly, most ovariectomized mares rarely require exogenous estrogens to augment their sexual behavior. In fact, overuse of estrogens may lead to objectionable behavior. Mares used as a mount
The condom There are some stallions that may refuse to breed an AV or may back off the mount source when attempts are made to deflect the penis. Typically, these are older horses that have been used exclusively for natural service. In such cases it may be necessary to attempt semen collection with the stallion wearing a condom and while breeding an estrous mare. This is probably the least desirable method of semen collection. Placement of the condom on the erect penis and its subsequent removal can be challenging and there are a number of stallions who will not breed while wearing a condom. Ejaculates obtained using condoms are often heavily contaminated with bacteria and debris. Loss of the condom as well as semen is not uncommon.
Mount mares Mares displaying behavioral estrus are commonly used as mount sources for semen collection. The
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Figure 123.5 Breeding phantoms with single support post (a) and two support posts (b).
source are usually restrained with a twitch or a chain over the gums and mild sedation may occasionally be required.
involved in the collection procedure. Stallions can easily damage their penis on the mount source if it is not deflected quickly enough when they charge or begin to thrust vigorously.
Breeding phantom The breeding phantom or dummy is an excellent mount source for semen collection. When properly constructed, the phantom provides a consistent, stable, and safe mount for the stallion. The height and angle of the phantom should be adjustable to comfortably accommodate stallions of different heights and those with painful ailments such as hock or back problems. A single supporting post is preferable to two or more posts (Figure 123.5) because there is less chance of the stallion’s legs becoming entangled in the supports during dismount or if the stallion happens to fall off the phantom. The covering of the phantom should be durable, minimally abrasive, and easily cleaned. Most stallions can be taught to mount the phantom and many, especially those trained at a young age, recognize the phantom as a sexual object even without the presence of a mare. For the majority of stallions, having an estrous mare in close proximity, either in front of or alongside the phantom, increases sexual stimulation and enhances the desire of the horse to mount and breed. Here again, an ovariectomized mare can provide the consistent, reliable estrous behavior required for teasing stallions throughout and outside of the physiological breeding season. Whether a mare or the phantom is used as a mount source, the stallion should approach in a controlled manner and mount from the rear. Allowing the stallion to charge the mount source increases the risk of injury to the horses and personnel
Ground collection Certain situations or medical conditions may necessitate collecting semen from a stallion while he is standing on all four legs. Semen collection can be accomplished using an AV, with manual stimulation and a plastic bag or sleeve secured to the penis, or can be induced pharmacologically. When using the AV or manual stimulation for ground collection, the stallion can be brought into the breeding shed, or left in a stall. He is presented with a tease mare to achieve an erection, after which the penis is washed and dried. The teaser is maintained in close proximity and the AV is applied to the erect penis, parallel with the stallion’s abdomen. The right hand can be used to gently but firmly grasp the base of the penis ventrally for added stimulation as the stallion thrusts. The stallion handler should push on the stallion’s shoulder with his or her right hand to provide support to the horse. Ideally, the stallion should be against a smooth wall on his right side to help prevent lateral movement and a barrier should be in place between the stallion and the teaser to discourage mounting. As the stallion thrusts into the AV, he will typically adopt a head down, arched back posture (Figure 123.6). The procedure for manual stimulation is similar except that, instead of using an AV, a plastic bag or sleeve is secured to the stallion’s penis and warm water compresses are applied to both the glans and the base of the penis.6, 7
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Figure 123.6 Ground collection with stallion demonstrating typical head-down, arched back posture.
Pharmacological induction of ejaculation A number of drugs and drug combination protocols have been published for inducing ex copula ejaculation in stallions.8, 9 Xylazine [0.2–0.66 mg/kg intravenously (i.v.)] or combinations of xylazine and the tricyclic antidepressant imipramine hydrochloride (1.0–2.0 mg/kg i.v. or 100–500 mg orally, b.i.d.), which lowers the ejaculatory threshold, are used most commonly.4, 9 Other combinations that the author has found successful are detomidine (0.016–0.02 mg/kg i.v.) followed by butorphanol (0.01–0.02 mg/kg i.v.) with or without previous imipramine administration. Erection and masturbation prior to ejaculation can vary, based on the drugs used and the level of sedation. In general, erection and masturbation are more likely to occur with the use of imipramine and mild to moderate sedation, whereas heavy sedation with or without imipramine usually results in ejaculation without erection or masturbation. For optimal results, the administration of the drugs should be done as smoothly and quietly as possible and the stallion should be left undisturbed in a quiet stall until sufficient time has elapsed to determine whether the protocol was successful. Therefore, before sedating the horse, a plastic bag or sleeve can be placed over the penis and prepuce and held in place using a lightly applied flank strap (Figure 123.7). For stallions that do not tolerate a flank strap, it may be necessary to have an attendant in the stallion to collect semen in a receptacle attached to a long handle. In most successful cases, the stallion will ejaculate with 15–60 minutes following drug administration. Favorable response is very inconsistent and, even under optimal conditions, semen is generally obtained in only 25–30% of pharmacological induction attempts. Individual variation among stallions exists regarding
Figure 123.7 Stallion sedated for chemical induction of ejaculation. Note the plastic bag placed over the penis and prepuce using a 15-cm cross-stitch hoop and brown gauze as a flank strap.
success of drug-induced ejaculation. In certain stallions, the technique is repetitively successful.
Epididymal sperm harvest Harvesting sperm from the epididymides of stallions for freezing and future use is generally considered a method of last resort in order to preserve valuable genetic material when stallions suffer a catastrophic injury or die prematurely. It is also sometimes requested to be performed at the time of elective castration. Although the first reported pregnancy obtained with frozen–thawed stallion sperm was achieved using epididymal sperm,10 pregnancy rates using epididymal sperm are generally lower than those obtained with ejaculated, cryopreserved semen. The literature contains few reports of this process and optimal harvesting, processing, and preservation techniques for epididymal sperm have yet to be determined. The paired cauda epididymides and vas deferens can contain > 50 × 109 sperm.11 Therefore, when castrating a stallion with the intention of harvesting epididymal sperm, the epididymides, especially the tails, and as much of the vas deferens as possible must remain intact. The vas deferens should be transected as high as possible and its lumen mechanically occluded to minimize loss of sperm. There are two basic methods described for harvesting epididymal sperm: the retrograde flush method and the ‘float-up’ method. For the retrograde flush method, the lumen of the vas deferens is cannulated and the epididymis is transected near the corpus–cauda junction. The tissues are suspended over a collection vessel and the lumen is flushed with semen freezing extender or balanced, buffered salt
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Semen Collection Techniques and Insemination Procedures solution such as Tyrode’s solution. With the retrograde flush method there is no need to centrifuge the spermatozoal flush fraction if semen freezing extender is used to flush the vas deferens and cauda epididymis. The ‘float-up’ method involves placing the cauda epididymis and the vas deferens in a dish or tube with semen extender or other suitable solution and the tissues are incised in multiple (10–15) locations. The collection vessels are gently agitated for 10 minutes. One laboratory reported higher sperm recovery rates using the flush method,12 whereas another laboratory reported no difference in sperm recovery rate between the two methods and seemed to prefer the ‘float-up’ method.2 The need for addition of seminal plasma or the final concentration of seminal plasma to maximize post-thaw characteristics of cryopreserved epididymal sperm have yet to be elucidated.
General semen handling techniques Immediately after collection, semen should be quickly transported to the laboratory with minimal agitation, exposure to light or cold-shock. All materials, including the seminal extender, that will come in contact with the semen should be warmed to body temperature (37◦ C). If an in-line filter was not used during collection of the ejaculate, semen should be poured through a non-toxic filter to remove the gel fraction and debris. The gel fraction can also be removed by careful aspiration with a syringe. Loss of spermatozoa for breeding purposes is greater when using the latter two methods than when using an in-line filter. A commercially available in-line, nylon, micro-mesh filter (Equine Semen Filter, Animal Reproduction Systems, Chino, CA) is preferred in order to minimize spermatozoal retention (loss) during filtration.13 Spermatozoal concentration, volume, and color of the gel-free semen and the percentage of progressively motile spermatozoa should be determined and recorded. (See Chapter 124 for details.) Whether the semen is to be used immediately or preserved, it should always be mixed with an appropriate extender to protect spermatozoa from environmental injury prior to insemination. Properly formulated seminal extenders improve spermatozoal survival during the period between collection and insemination. The most commonly used equine seminal extenders are milk-based (see Chapters 131 and 132). An minimum dilution ratio of 1 : 1 to 1 : 2 (semen : extender) is recommended for semen to be used for insemination immediately. Warmed extender can be added to semen following its collection or placed in the seminal receptacle prior to collection, so that the spermatozoa come in contact with this supportive medium immediately af-
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ter ejaculation. To obtain accurate measurement of spermatozoal concentration in extended semen with photometric-type units, extenders must be optically clear. Otherwise, a hemocytometer, CASA system, or a NucleoCounterTM SP-100 (Chemometec, Denmark) must be used to quantify spermatozoa in the extended semen.
Insemination timing and frequency Historically, mares in many artificial insemination management systems were inseminated every other day, beginning the second or third day of estrus, until ovulation was detected or the mare no longer exhibited signs of behavioral estrus. However, the increased use of transrectal palpation and ultrasonography has increased our ability to predict ovulation. Ideally, transrectal ovarian palpation and ultrasonography should be used in a breeding program to more accurately predict ovulation, so that the number of required inseminations per estrous period is minimized. Limiting the number of inseminations improves the overall efficiency of the breeding program and reduces the risk of iatrogenic contamination of the mare’s reproductive tract. Reducing uterine contamination is especially important when breeding mares with an increased susceptibility to uterine infection or inflammation. Although acceptable pregnancy rates are obtained when mares are inseminated within 48 to 72 hours prior to ovulation with semen from fertile stallions,14 when using semen from subfertile stallions, daily or twice-daily inseminations until ovulation occurs may be required, to improve pregnancy rates.15 In one study,16 a single cycle pregnancy rate of 75% (9/12 mares) was obtained from single inseminations performed 3 days prior to ovulation. The highest pregnancy rate (88%; 7/8 mares) in that study16 was in the group of mares inseminated 1 day before ovulation. In mares inseminated within 6 hours after ovulation, it was found that pregnancy rates were comparable to those achieved with a single insemination before ovulation or two inseminations, one before and one after ovulation.17 Insemination within 6 hours after ovulation also resulted in pregnancy rates similar to those acheived from insemination performed 1–3 days before ovulation.16 However, a high embryonic loss rate (34%) occurred in the mares inseminated after ovulation.
Insemination dose (sperm number) Mares in an artificial insemination program are typically inseminated with 250 to 500 million progressively motile spermatozoa (PMS). However, it is clear
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from more recent studies, where insemination doses of 20 × 106 PMS,18 50 × 106 PMS19 or 100 × 106 PMS20 from fertile stallions did not reduce fertility, that an insemination dose of 500 × 106 PMS is not necessary to achieve acceptable pregnancy rates with fertile stallions. In addition, ample anecdotal information exists from breeding farms that have successfully used artificial insemination doses ranging from 200 to 400 × 106 PMS. The minimum effective insemination dose is probably stallion dependent. Under ideal conditions, the insemination dose may be reduced to 100 × 106 PMS without reducing fertility, when using semen from highly fertile stallions.21–23 However, inseminating mares with 500 × 106 PMS will help ensure that acceptable pregnancy rates are achieved, by allowing some margin for error in seminal evaluation and handling when conditions are less than optimal.
Insemination volume The number of spermatozoa in an insemination dose appears to be more critical than the volume of the inseminate. Although smaller or larger volumes can be used successfully, typical insemination volumes for extended equine semen range from 10 to 25 mL. When timed closely with ovulation, or using deep horn insemination techniques, volumes as low as 0.2 mL of semen have resulted in pregnancy (see Chapter 323). Care must be taken when small volumes of highly concentrated semen are used, so that a substantial portion of the inseminate, and therefore a large number of spermatozoa, do not remain in the insemination equipment. Colorado workers suggested that volume of inseminate within the range of 0.6 to 26.8 mL have no effect upon fertility24 but that insemination volumes 100 mL may be detrimental to fertility.25 However, others have reported that such a large volume is not detrimental when sperm concentration in the extended semen is adequate (i.e., 50 × 106 spermatozoa/mL).26 A minimal spermatozoal concentration for standard uterine body insemination is considered to be approximately 25× 106 spermatozoa/mL.27, 28 A spermatozoal concentration 5 × 106 /mL could yield lowered pregnancy rates.29 When a number of mares are to be inseminated with a single ejaculate, insemination volume can be calculated by dividing the desired number (e.g., 100 to 500 × 106 ) of PMS per insemination by the product of the spermatozoal concentration in the extended semen and the percentage of progressively motile spermatozoa in the ejaculate, i.e.:
Insemination volume =
Desired # of PMS [sperm conc.] × [% PMS]
For example: 10 mL = 500 (×106 )/(100 × 106 × 50%) 17.9 mL = 250 (×106 )/(35 × 106 × 40%)
Insemination procedure Sterile, non-toxic, disposable equipment should be used for artificial insemination procedures. Syringes with non-spermicidal, plastic plungers (e.g., Airtite, Vineland, NJ) are preferable for artificial insemination since rubber plungers may have spermicidal properties (Fig. 123.8).30–33 There appears to be great individual stallion variation regarding spermatozoal sensitivity to the toxic effects of syringes with rubber plunger tips. Toxic effects are apparent in semen from some stallions with as little as 1 minute of contact with syringe plungers (D. Driscoll and D.H. DouglasHamilton, personal communication, 1984). Washing and re-sterilization of syringes does not appear to affect spermatozoal motility.32 Insemination of the mare should be performed in accordance with the minimum contamination techniques described by Kenney et al.22 The mare should be adequately restrained, with her tail wrapped and diverted either off to the side or up over her rump (Fig. 123.9). The perineal area is thoroughly scrubbed and rinsed, paying particular attention to the vulva and making sure the caudal vestibule is free of fecal contamination (Fig. 123.10). A minimum of two to three scrubs with soap or a surgical scrub are recommended. Thorough rinsing is essential to eliminate any residual soap that can be spermicidal or that can irritate the mare’s genitalia. Once the mare is adequately prepared, the inseminator puts on a sterile sleeve or clean plastic sleeve over which a sterile,
Figure 123.8 All–plastic syringes (top) are preferred over syringes containing rubber plunger tips (bottom) because products from the rubber can leach into the semen and damage spermatozoa.
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Figure 123.11 A sterile rectal sleeve can be used for artificial insemination to guard against contamination of the mare’s reproductive tract with microorganisms at the time of insemination.
Figure 123.9 In preparation for artificial insemination, the mare’s tail is first wrapped with an impervious product, such as a plastic rectal sleeve, then lifted over the rump to expose the external genitalia for washing and rinsing.
disposable, latex glove can be worn (Fig. 123.11). The tip of the insemination pipette is covered in the gloved hand and a small amount of sterile, non-spermicidal lubricant is applied to the back of the glove (Fig. 123.12). The gloved hand and insemination pipette are passed between the vulvar lips to the cranial vaginal vault where the index finger identifies and penetrates the cervix (Fig. 123.13). The insemination pipette is advanced through the cervix to the uterine body where the semen is slowly deposited. When mares have had a Caslick operation performed, the cervix can be visualized and an
Figure 123.10 After washing and rinsing the mare’s external genitalia, a clean damp piece of cotton is used to check the vestibule for cleanliness.
Figure 123.12 A small amount of sterile non-spermicidal lubricant is placed on the back of the sleeve at the level of the fingertips to aid in entry into the mare’s vagina.
Figure 123.13 The inseminator passes an insemination pipette into the vaginal vault to the level of the cervix, then inserts the index finger and pipette into the cervical lumen and advances the tip of the pipette into the uterine body.
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(a)
(b)
Figure 123.14 (a) The inseminator can pass a pipette tip through the cervix for insemination when it is not possible to pass an arm into the vaginal vault because of a previous Caslick operation. (b) The pipette tip is placed into the cervical lumen and then advanced into the uterine body.
insemination pipette passed through it via a lighted tube speculum placed in the vagina (Fig. 123.14a,b). Placing a slight bend in the end of the pipette may aid in maneuvering it through the cervix with the aid of a tube speculum (Fig. 123.15).
Concluding remarks Artificial insemination is an effective technique for optimizing stallion utilization in a breeding program. When proper seminal handling and insemination procedures are used, excellent pregnancy rates can be achieved. Pregnancy rates may actually be improved over natural service when artificial insemination techniques are employed for mares and stallions with marginal fertility.
Figure 123.15 A slight bend can be created near the tip of the pipette to aid in passing it into the uterine body with a tube speculum.
References 1. Love CC. Semen collection techniques. Vet Clin North Am Equine Pract 1992;8:111–28. 2. Cary JA, Scott M, Farnsworth K, Hayna J, Duoos L, Fahning ML. A comparison of electroejaculation and epididymal sperm collection techniques in stallions. Can Vet J 2004; 45:35–41. 3. Collins CW, Songsasen N, Monfort SL, Bush M, Wolfe B, James SB, Wildt DE, Pukashenthi BS. Seminal traits in the Przewalski’s horse (Equus ferus przewalskii) following electroejaculation. Ann Reprod Sci 2006;94:46–9. 4. Hurtgen JP. Semen collection in stallions. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. Philadelphia: W.B. Saunders, 2000; pp. 81–90. 5. Brinsko SP, Varner DD, Blanchard TL, Meyers SA. The effect of postbreeding uterine lavage on pregnancy rate in mares. Theriogenology 1990;33:465–75. 6. Crump J, Crump J. Stallion ejaculation by manual stimulation of the penis. Theriogenology 1988;1:341–6. 7. McDonnell SM, Love CC. Manual stimulation collection of semen from stallions: Training time, sexual behavior and semen. Theriogenology 1990;33:1201–10. 8. Klug E, Deegan E, Lazarz B, Rojem I, Merkt M. Effect of andrenergic neurotransmitters upon the ejaculatory process in the stallion. J Reprod Fertil Suppl 1982;32:31–4. 9. McDonnell SM. Ejaculation physiology and dysfunction. Vet Clin North Am Equine Pract 1992;8:57–70. 10. Barker JAV, Gandier JCC. Pregnancy in a mare resulted from frozen epididymal spermatozoa. Can J Comp Med 1957;21:45–51. 11. Amann RP, Thompson DL, Squires EL, Pickett BW. Effects of age and frequency of ejaculation on sperm production and extragonadal sperm reserves in stallions. J Reprod Fertil Suppl 1979;27:1–6. 12. Bruemmer JE. Collecting and freezing of epididymal stallion sperm. Vet Clin North Am Equine Pract 2006;22:677–82. 13. Amann RP, Loomis PL, Pickett BW. Improved filter system for an equine artificial vagina. J Equine Vet Sci 1983;3:120–5.
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Semen Collection Techniques and Insemination Procedures 14. Palmer E. Factors affecting stallion semen survival and fertility. Proc 10th Int Cong Anim Reprod Artif Insem 1984;III:377. 15. Varner DD. Stallion utilization in artificial insemination programs. Proceedings of Society for Theriogenology 1987; pp. 118–39. 16. Woods J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic-loss rate in mares. Equine Vet J 19990;22:410–15. 17. Martin JC. Untersuchungen zur zyklusteuerung, insbesondere brunstsynchronisation im rahmen der instrumentellen samenubertragung biem pferd. Vet-med Diss, Hannover, 1980. 18. Woods J, Rigby S, Brinsko S, Stephens R, Varner D, Blanchard T. Effect of intrauterine treatment with prostaglandin E2 prior to insemination of mares in the uterine horn or body. Theriogenology 2000;53:1827–36. 19. Seime H, Bonk A, Hamann H, Klug E, Katila T. Effects of different artificial insemination techniques and sperm doses on fertility of normal mares and mares with abnormal reproductive history. Theriogenology 2004;62:915–28. 20. Rigby S, Hill J, Miller C, Thompson J, Varner D, Blanchard T. Administration of oxytocin immediately after insemination does not improve pregnancy rates in mares bred by fertile or subfertile stallions. Theriogenology 1999;51:1143–50. 21. Pickett BW, Burwash LD, Voss JL, Back DG. Effect of seminal extenders on equine fertility. J Anim Sci 1975;40:1136–43. 22. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: Technique and preliminary findings. Proceedings of American Association of Equine Practitioners 1975; pp. 327–36. 23. Demick DS, Voss JL, Pickett BW. Effect of cooling, storage, glycerolization and spermaotzoal numbers on equine fertility. J Anim Sci 1976;43:633–7.
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24. Pickett BW, Squires EL, McKinnon AO. Procedures for collection, evaluation and utilization of stallion semen for artificial insemination. Colorado State Univ. Exp. Sta. Anim. Reprod. Lab. Gen. Series Bull. 03, 1987. 25. Rowley HS, Squires EL, Pickett BW. Effect of insemination volume on embryo recovery in mares. Equine Vet Sci 1990;10:298–300. 26. Bedford SJ, Hinrichs K. The effect of insemination volume on pregnancy rates of pony mares. Theriogenology 1994;42:571–8. 27. Varner DD, Blanchard TL, Meyers PJ, Meyers SA. Fertilizing capacity of equine spermatozoa stored for 24 hours at 5 or 20◦ C. Theriogenology 1989;32:515–25. 28. Rigby SL, Brinsko SP, Cochran M, Blanchard TL, Love CC, Varner DD. Advances in cooled semen technologies: seminal plasma and semen extender. Anim Reprod Sci 2001;68:171–80. 29. Jasko DJ, Martin JM, Squires EL. Effect of insemination volume and concentration of spermatozoa on embryo recovery in mares. Theriogenology 1992;37:1233–9. 30. Petersen MC, Vine J, Ashley JJ, Nation RL. Leaching of 2-(2hydroxyethylmercapto) benzothiazole into contents of disposable syringes. J Pharm Sci 1981;70:1139–43. 31. Jones WE (ed.) Toxic agents in equine AI procedures. Equine Vet Data 1984;5:219. 32. Shull JW, Varner DD, Bretzlaff KN, Graham CW. Spermatozoal motility parameters of equine semen stored in plastic syringes. Proc Conf Res Workers Anim Dis 1986, p. 41. 33. Broussard JR, Goodeaux SD, Goodeaux LL, Thibodeaux JK, Moreau JD, Godke RA, Roussel JD. The effects of different types of syringes on equine spermatozoa. Theriogenology 1993;39:389–99.
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Chapter 124
Evaluation of Semen Julie Baumber-Skaife
There are many reasons why a stallion may be collected for a semen evaluation, including but not limited to the following:
r r r r r r
for a breeding soundness or pre-purchase examination; before the start of the breeding season; to determine if a stallion should be included in a breeding program; when fertility problems are suspected; to determine if the stallion will be a candidate for cooled or frozen semen; after semen collection, before artificial insemination or transport.
This chapter summarizes both the traditional and the more advanced techniques that a clinician can perform when conducting a semen evaluation. Routine tests include gross evaluation, determination of volume, concentration, motility, morphology, and pH. Intermediate tests include longevity, bacteriology and cytology, the hypo-osmotic swelling test, and live-dead staining. Advanced tests are those that rely on fluorescent technology (e.g., acrosomal status, capacitation, DNA integrity) or technical skill (zona binding). Of course, the true test of fertility is the pregnancy rate following breeding of multiple reproductively sound mares. However, this is not easily achieved since many stallions breed only a limited number of mares and the results are also dependent upon the fertility of the mare and her breeding management. Furthermore, this is a retrospective study. Ideally we would like to predict fertility before the start of a breeding career through physical examination and semen analysis. The current limitation of this approach is that, although a number of sperm function tests are available, their relationship to fertility has not been fully defined and guidelines for what is normal or acceptable are unavailable. Furthermore, the cause of subfertility is sometimes difficult to define because some stallions may
have normal semen characteristics yet still be subfertile under optimal management conditions. Generally, poor sperm quality can potentially indicate subfertility, whereas good semen quality is no guarantee of fertility. The main reason for this observation is that we have not fully identified all the elements of function a sperm requires to complete its destiny in fertilizing the egg. There are sperm characteristics still to be discovered and assays that remain to be developed. In the future, we hope to have available an array of tests that enable us to better predict the fertility of a stallion or quantify whether he will be a suitable candidate for cooled or frozen semen. Given the recent growth in the field of genomics it is feasible that prediction of fertility may be based upon a genetic test rather than a laboratory-based sperm function test. These advances will enable owners to select for fertility in their breeding programs and potentially optimize the financial investment in breeding horses.
Gross appearance A general evaluation of the color and appearance of the raw semen can provide valuable information and should not be ignored before continuing with the semen processing. Raw semen should be gray or white in color and the turbidity of the semen can give an initial indication of sperm concentration. Dilute, low-concentration ejaculates look gray and watery, whereas highly concentrated ejaculates look thicker and are creamier in appearance. Semen with a yellowish color and a urine-like smell indicates urospermia – the presence of urine in the ejaculate. Ejaculates that are red or pink in color indicate hemospermia – the presence of blood in the ejaculate. A sample that is blood-tinged and flocculent with clumps of purulent material may represent seminal vesiculitis. The causes of urospermia and hemospermia are covered in more detail in Chapter 108. A watery sample from
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Weight of a full ejaculate of raw semen vs volume (n = 21) 140 R2 = 0.99
120 100
Weight (g)
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80 60 40 20 0 0
20
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60 80 Volume (mL)
100
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Figure 124.1 Graph representing the relationship between volume and weight for full ejaculates of horse semen.
a stallion that usually has good sperm production may simply be a partial ejaculate or it may signify blockage of one or both of the ampullae. When released this can be observed as clumps within the gel filter and a very high concentration of cells of poor morphology, often detached heads, in the ejaculate.
Volume To calculate the total number of sperm in the ejaculate it is necessary to measure the volume of the semen and determine the sperm concentration. The volume of semen is generally divided into two fractions, gel volume and gel-free volume of the ejaculate. When using an in-line filter on the collection bottle the gel is separated from the sperm-rich fraction of the ejaculate. After collection the gel filter can be removed from the collection bottle and the volume of gel within the filter measured. The most accurate method for measuring gel-free volume is to pour the semen into a graduated cylinder and read off the volume from the bottom of the meniscus. The disadvantage of measuring volume with a graduated cylinder is that after use the container must be processed through a thorough cleaning protocol for disinfection followed by sufficient rinsing to ensure there is no detergent residue that could negatively impact semen quality. Many equine reproduction laboratories measure volume in disposable 50 mL conical tubes or specimen cups. However, one must acknowledge that this is just an approximation of volume and not an accurate measurement. Nonetheless for routine processing this method is generally sufficient. An alternative method for measuring volume is to use weight, based upon the premise that 1 mL of semen weighs 1 gram. Weight may be measured
in a tube or specimen cup or the empty collection bottle can be ‘zeroed’ before attaching to the artificial vagina and then weighed again after collection. Figure 124.1 is a graph showing the relationship between weight and volume of full ejaculates of raw semen (author’s unpublished results). Any equipment that comes into contact with raw semen must be sterile and warmed (37◦ C) before use so as not to cold-shock the sperm, or to contaminate or compromise the semen quality. Therefore, graduated cylinders or disposable plastic tubes or cups for measuring volume should be kept within the incubator so they are warmed before use. It is important to extend the semen as quickly as possible after collection; therefore determination of semen volume and concentration must be done in a timely manner.
Concentration Determining the concentration of spermatozoa in the gel-free fraction of an ejaculate is a fundamental procedure when performing a complete breeding soundness examination or preparing semen for artificial insemination. The sperm concentration (usually presented as millions/mL, e.g., 225 × 106 /mL) is multiplied by the gel-free volume (in mL) to give the total number of sperm cells in the ejaculate (designated as billions, e.g., 12.5 × 109 ). In equine reproduction laboratories the two most routinely used methods for sperm counting are spectrophotometer-based machines or the traditional hemocytometer. Because of the expense of the equipment, advanced counting methods utilizing flow cytometry have been limited to the research community; however, there are now convenient benchtop instruments incorporating fluR orescence technology, such as the NucleoCounter
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SP-100TM , that are marketed for clinical use. In addition, fluorescence-based counting applications are now available for computer-assisted sperm analysis (CASA) instruments.
Spectrophotometric methods A number of spectrophotometer-based counting instruments are available commercially, including the SpermaCue (Minitube) and Accuread (IMV), and the Densimeter or ‘blue box’ from Animal Reproduction Systems (ARS). The basic concept of these counting instruments is an optical system incorporating a beam of light that passes through the sample and is recorded by a photodetector. The signal from the photodetector is converted to an absorbance value, which is then electronically translated into a sperm concentration. The amount of light transmitted through a sample is inversely correlated with the sperm concentration in the samples. The machines are validated upon serial dilution of known concentrations performed on a hemocytometer and it is good practice to routinely test the calibration of the machine against a hemocytometer count. In the case of the ARS Densimeter, the protocol is as follows. A volume of 3.42 mL of 10% buffered formalin is aliquoted into a cuvette using an automatic dispenser. The dispenser must first be primed by discarding the first aliquot pumped from the bottle. The formalin within the arm of the dispenser evaporates over time and therefore the first aliquot will be short on volume. When determining sperm concentration the accuracy of pipetting volume is of strict priority. Handling the cuvette by the ribbed side only, it is placed into the machine and ‘zeroed’. The semen sample is mixed and a 180 µL aliquot removed and added to the cuvette. To improve the accuracy of pipetting the outside of the pipette tip is wiped with a Kimwipe to avoid the introduction of additional volume. The semen is mixed with the formalin in the cuvette and then replaced into the machine for a ‘count’. Many of these instruments also have the option for the technician to enter the volume and motility information for the ejaculate. The instrument then calculates the total sperm number, as well as the dose (volume) recommended for artificial insemination or cooled transport. Photometer-based instruments can only determine the sperm concentration of raw semen because, once the semen is diluted by the opaque extender, the extender contributes to the concentration read by the instrument, unless an optically clear extender is used. The following methods of sperm counting have the advantage of being able to determine sperm concentration in extended as well as raw samples.
Hemocytometer The hemocytometer was invented by Louis-Charles Malassez and was originally designed for counting of red blood cells. It consists of a thick glass microscope slide with two rectangular indentations or ‘chambers’ aligned vertically in the center of the slide. The chambers are engraved with a laser-etched grid of perpendicular lines. A diluted semen sample is loaded by capillary action into the chamber and the number of sperm within the counting grid determined. The counting grid has a known depth and size and consequently a specified volume; thus, the concentration of cells within this known volume can be translated into the concentration of the original sample. A number of different types of hemocytometer have been developed; the one most commonly used for counting sperm cells is the Improved Neubauer hemocytometer. A protocol for performing a hemocytometer count is presented below. Despite being considered the gold standard for determination of sperm concentration, the hemocytometer suffers from a number of limitations, such as the variation between different hemocytometer designs1 and the technical skill of the operator.2 Freund and Carol3 noted not only a considerable variation between technicians, but also variation between duplicate pipettes and duplicate chambers performed by the same technician. They anticipated mean differences of about 20% for duplicate determinations of sperm concentration by the same technician. Consequently it has been determined that at least four chambers must be counted in order to get a high level of precision and repeatability,1 and this is extremely time consuming.
Protocol for performing a hemocytometer count 1. Dilute the raw semen 1 : 100 (20 µL : 1.98 mL) in either 10% buffered formalin or 0.5 M sodium citrate. Ensure the semen sample is well mixed before removing an aliquot and wipe the tip of the pipette with a Kimwipe before transferring the semen into the diluent (so as to avoid introducing additional volume). 2. Ensure the hemocytometer and coverslip are clean and then place the coverslip over the hemocytometer chambers. 3. Mix the diluted semen and load 12 µL into a pipette. Balance the pipette tip within the grove of the hemocytometer chamber and slowly release the diluted semen into the chamber, allowing it to fill by capillary action. Be careful not to lift the coverslip while loading the chamber and do not overfill. Mix the diluted semen sample again and load the second chamber.
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Figure 124.2 (a) Photographic image of the 5 × 5 counting grid on a hemocytometer and (b) a close-up view of one of the 25 squares showing the three-line border to each 4 × 4 square.
4. Allow the sample to settle for a few minutes within the hemocytometer. It is preferable to let the hemocytometer settle within a humidity chamber so the sample does not dry out. A simple humidity chamber can be made by placing a piece of moist filter paper on the bottom of an embryo search dish and resting the hemocytometer upon two 0.5 mL straws used for frozen semen. 5. Place the hemocytometer under the microscope (using 20× objective) and locate the central grid. The central grid is a 5 × 5 arrangement of squares (Figure 124.2a), each square subsequently divided into 4 × 4 smaller squares (Figure 124.2b). 6. The borders of the 25 larger squares are defined by three lines (Figure 124.2b). The center line represents the boundary of the counting area. All sperm heads are counted that have more than half the sperm head within the shaded area. For sperm that lie with exactly half the sperm on the edge of the shaded area, count only those heads that lie on the top and right lines of the box; this prevents counting the same sperm twice in adjacent squares. 7. Sperm within all 25 squares that form the central grid are counted. The count is then repeated on the second chamber and the mean represents the number of sperm in the original raw semen sample in millions/mL. The result is obtained as follows. The 25 large squares represent an area of 1 square mm. The depth of the hemocytometer is 0.1 mm. Therefore, the volume above the 25 squares is 0.1 µL (1 × 1 × 0.1 = 0.1). To get the concentration in millions/mL of the original sample you multiply the number of sperm in 25 squares by 10 000 (1 mL = 1000 µL ÷ 0.1 µL = 10 000). For example, say the sperm count in 25 squares is 243.
R SP-100TM (ChemoMetec A/S, Figure 124.3 NucleoCounter Denmark) sperm counting machine.
100 before loading the hemocytometer. Therefore 2 430 000 × 100 (dilution factor) = 243 000 000 = 243 × 106 /mL. NucleoCounter R SP-100TM (ChemoMetec A/S, The NucleoCounter Denmark; Figure 124.3) is a relatively new instrument that offers innovative technology for the determination of sperm concentration. The basics of the methodology are as follows. The semen sample is first diluted into a detergent solution, Reagent S-100, which permeabilizes the sperm membrane. The diluted semen is then loaded into specially designed SP-1 cassettes (shown in Figure 124.4) which contain a microfluidic network. Within the channels of the microfluidic network is the fluorescent dye, propidium iodide, which intercalates with DNA and stains the nuclei of cells. Propidium iodide is a membraneimpermeable dye, typically used as a viability stain.
Number of sperm in 0.1 µ L = 243 Number of sperm in 1 mL = 2 430 000(243 × 10000) Then you must account for the dilution factor since the original sample was diluted 1 in
Figure 124.4 NucleoCounter SP-1 cassettes. The micro-fluidic channels contain the fluorescent probe, propidium iodide. Labeled cells are visualized by the NucleoCounter machine through the viewing window (square at top).
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By disrupting the membrane with dilution in S-100 reagent the dye is able to label the nucleus of every sperm cell. The integrated fluorescence microscope within the NucleoCounter detects the fluorescent signal from each sperm nucleus and the automated semen analysis software, SemenViewTM , converts this information into a value for sperm concentration. Hansen et al.4 compared the NucleoCounter SP100 to other cell counting methods in boar semen and determined the machine to have excellent repeatability (coefficient of variation (CV) of 3.1% as compared to 7.1% for hemocytometer) and to correlate well with the FACSCount flow cytometry-based counting system. The actual cell concentration did not correlate as well as expected with the hemocytometer, although it was noted that the presence of agglutinated sperm complicated hemocytometer counts. Research at Select Breeders Services Inc. found an excellent correlation (R2 = 0.97) between the hemocytometer and the NucleoCounter; however, this was only when four hemocytometer chambers were counted (author’s unpublished results). On average the NucleoCounter counted 15–20% fewer sperm than the ARS Densimeter, no doubt because the NucleoCounter is counting only sperm cells and not additional debris in the semen. For some stallions where a lot of debris was observed in the semen, the count using the NucleoCounter was half that of the ARS Densimeter; this has significant application when processing an accurate breeding dose. The advantage of the NucleoCounter is that it only counts DNA-containing cells within the semen sample and therefore provides a far more accurate sperm count than any photometer-based counter which can be biased by debris in the sample contributing to the sperm count. In addition to its superior accuracy and repeatability, the other advantages offered by the NucleoCounter compared to the hemocytometer are its ease of use and speed of counting (45 seconds). Disadvantages of the NucleoCounter include the expensive purchase price and the high cost per count for the SP-1 cassettes. Determination of the sperm concentration within the ‘viewing window’ of the cassette is sensitive to cell density; therefore samples must be diluted within an appropriate concentration range. This necessitates knowing in advance the approximate concentration of the sample so that it can be diluted accordingly. Most ejaculates (between 100 and 400 × 106 /mL) can be counted at a 1 : 100 dilution; however, low- and high-concentration ejaculates should be diluted differently as directed by the dilution chart that accompanies the machine. Finally, the accuracy of any counting method is directly dependent upon the precision of the technician using it. Therefore inappropriate pipetting and dilution, plus
failure to adequately the mix the semen sample before counting, mean that the superior accuracy of the machine can easily be lost to poor sample handling technique. Counting applications of CASA If the semen sample is analyzed by CASA using chambers of a known volume and depth, concentration can be calculated. The software can then be utilized to calculate total sperm, dilution volume, and dose volume for preparing semen for artificial insemination. This option has not been fully validated with equine semen and there is concern regarding correction for the Sergre-Silberberg (SS) effect, which describes fluid dynamics within a capillaryloaded chamber.5, 6 Furthermore, counting of debris in the sample can interfere with the sperm count. This is eliminated with a fluorescent application for the Hamilton-Thorne IVOS CASA machine, called IDENT. The IDENT stain is a DNA-specific fluorescent dye based upon Hoechst bisbenzimide; it only labels sperm cells and consequently debris is not counted. Use of the IDENT system for sperm counting has been described with human7 and rat8 spermatozoa. The more advanced option, VIADENT, enables determination of motility, concentration, and viability and has been validated in equine sperm.9
Analysis of sperm motility Sperm motility is a comprehensive topic that is discussed in further detail in Chapter 125. The author also refers the reader to excellent reviews on the physiology of sperm motion.10, 11 Evaluation of sperm motility is considered a fundamental test for assessing sperm quality and is probably the most quoted variable in a semen evaluation. This is because, at the current time, sperm motility is the only characteristic within the basic evaluation that can give an indication of the potential fertility of a stallion. A study by Jasko et al.12 compared conventional semen characteristics and fertility and showed that the percentage of progressive motility best corresponded to fertility. At least a billion progressively motile sperm are recommended for a stallion to pass a breeding soundness examination. Motility is also the primary determinant of whether an individual will be acceptable for breeding with cooled or frozen semen. A post-thaw progressive motility of 30% is recommended for commercial distribution of frozen semen. Unfortunately, a lack of uniformity between technicians in the estimation of visual motility, variability in CASA instrument settings, and the multitude of factors that can influence sperm motility make it difficult to define standard values for normal sperm motion.
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Furthermore, a stallion can have excellent motility and still be subfertile, so it is important to remember that sperm motility represents just one attribute of sperm function and, as a single test, it cannot predict fertility. Visual assessment Visual assessment of sperm motility is reported as total sperm motility (percentage of sperm exhibiting motility of any form), progressive motility (percentage of spermatozoa that exhibit rapid linear motion), and velocity (scored on an arbitrary scale of 0–4). It is a simple method that can be determined using a phase-contrast microscope (at 20× or 40×) by an experienced veterinarian or technician familiar with evaluating sperm motility. It is the most common method of motility analysis performed by the practicing clinician. However, it is a subjective method of analysis, and results are therefore susceptible to bias and can vary greatly between observers. The semen should be evaluated after being extended, as raw semen has a tendency to agglutinate and high sperm concentrations can prevent visualization of individual sperm trajectories. Dilution prior to analysis in a basic skim milk/glucose extender to a concentration of 25–50 × 106 /mL is preferable. Ideally, motility should be evaluated at 37◦ C; therefore, the microscope should have a heated stage and the slides and equipment should be warmed before use. Multiple fields across at least two suspensions should be examined and the fields should be located around the center of the slide. Motility is reduced at the edges where the sample dries out or accumulates cells because of drift. Sperm motility is particularly susceptible to environmental conditions; thus, it is critically important not to negatively impact motility by contamination with disinfectants, soaps or lubricants and to avoid excessive exposure to heat, cold, pH, and hyper- or hypo-osmotic conditions. Dirty slides and coverslips can also interfere with sperm motility. Computer-assisted sperm analysis Computer-assisted sperm analysis (CASA) for evaluation of sperm motility was developed in the 1980s. Over the years, analysis has evolved from multiexposure photomicrography, frame-by-frame playback of video recordings, to more recent semiautomated methods. Although expensive, these instruments offer a rapid and automated method of analysis with a greater degree of repeatability and less subjectivity than traditional visual estimates of motility. A number of instruments are available commercially. The most commonly used in equine re-
Figure 124.5 A typical computer-assisted sperm analysis (CASA) set-up.
production are the Hamilton-Thorne (Beverley, MA) CEROS (Figure 124.5) or IVOS sperm analyzers or the Minitube (Verona, WI) SpermVisionTM system. It is important to note that results obtained with different brands of CASA machines may not be comparable.13 A wealth of literature is available on the principles of CASA, its mode of operation, and the practical aspects that impact its use. The author refers readers to excellent reviews on these topics11, 14, 15 and presents just a brief overview here. The components of the CASA machine are a microscope, video camera, video-frame grabber card, a computer, and the motion analysis software. The image from the microscope is sent through the camera to the computer and is converted into a digital image. The computer perceives the sperm as white heads on a dark background by a dark-field or negative-highphase contrast image. The image is digitized and the computer identifies the sperm heads by size in pixels. There is a range of head size for each species and the expected maximum and minimum are defined to the computer, so that it identifies any object within this range as a sperm head. In addition to size the computer also uses the brightness of an object in order to discriminate sperm from non-sperm debris. All motile objects above a certain minimum size are presumed to be sperm and are tracked by the machine; the size and brightness of all motile objects is measured. The software then uses the mean size and mean brightness of the motile objects for each given field to determine which of the non-motile objects are sperm or non-sperm debris. User-defined gates for size and brightness are applied around this mean for all motile objects and any non-motile object that falls within this range is counted as non-motile sperm. The digitized sperm head images are tracked by the computer as x,y coordinates over sequential
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frames and then reconstructed into tracks representing the path the sperm swam during the time period of analysis for that particular field. Multiple fields are analyzed and the results presented as a mean. The reconstructed trajectory can be altered considerably by the number of images taken per second; the more frames, the more accurately the computer can reconstruct the track and this difference can alter the kinetic values for a trajectory.16 Therefore, it is important to standardize the frame rate in order to compare results. A typical frame rate is 60 frames/second for a total of 40 frames. The total number of frames analyzed should also be standardized, since an analysis of only a few frames will result in mostly linear tracks, whereas analysis of a larger number of frames provides more time for a sperm to deviate from a linear trajectory. As the number of fields and cells analyzed increases, so does the precision of the results. Therefore it is recommended that 300–500 motile cells per sample are evaluated17, 18 across multiple fields. The technician should be careful not to subjectively bias the result with field selection. In addition to quantifying total and progressive motility, CASA instruments are able to calculate more specific characteristics of sperm motion, for example, straight-line velocity (VSL), curvilinear velocity (VCL), average path velocity (VAP), linearity (LIN), straightness (STR), amplitude of lateral head displacement (ALH), and beat cross frequency (BCF). Total sperm motility is determined as the number of sperm moving at or above the minimum speed threshold defined in the CASA set-up (usually >20 µm/s for equine sperm). Sperm traveling at <20 µm/s are defined as slow cells and are usually classified as static. Slow cells generally result from drifting of non-motile cells within the semen sample or from slight movement of static cells caused by the flagellum of nearby motile sperm. By counting slow cells as static (non-motile) they do not contribute to the total motility. Progressive motility is also determined by thresholds for VAP (standard threshold value 50 µm/s) and STR (standard threshold value 75%) as defined in the CASA set-up. There is variability between laboratories in the thresholds used to define progressive motility; therefore it may not be possible to compare results for progressive motility between laboratories. Velocity values VAP, VSL, and VCL (expressed in µm/s) are calculated as shown in Figure 124.6. VCL is based upon the total distance that the sperm head travels during the period of analysis and it is therefore the highest value. The VSL is derived from the straight-line distance between the first and last points for the trajectory and is usually the lowest value. The VAP is the ‘average’ distance the sperm has traveled during the period of analysis. The average path is deter-
VCL VSL VAP
Figure 124.6 Diagram demonstrating how the velocity parameters, VAP (average path velocity), VSL (straight line velocity), and VCL (curvilinear velocity) are calculated from the sperm track captured by the CASA instrument.
mined by the CASA system using track smoothing. Basically for each point on the trajectory the computer takes an average of its x,y coordinates with the two points on either side of it. This gives a smoothed x,y coordinate for each track point and ultimately a smooth path. STR and LIN represent ratios (expressed as a percentage) of velocity parameters: LIN = (VSL/VCL) × 100 and STR = (VSL/VAP) × 100. ALH is calculated as the total width of the lateral movement of the sperm head and is based upon the smoothed path. The BCF is calculated as the number of times the curvilinear path crosses the average path per second (expressed in Hz). Kinetic parameters offer a great deal of additional information on sperm motion and can aid in the identification of subtle differences between treatments or individuals that may not be readily observed by visual analysis. For analysis by CASA, the raw semen should be diluted to ∼ 25 × 106 /mL.18 High sperm concentrations can interfere with the analysis since sperm collisions and path crossings lead to erroneous track interpretations. Semen must be diluted in media free of particles that are similar in size to the sperm head. Basic non-fat dried skim milk based extenders are fine, but freezing extenders that contain unclarified egg yolk interfere with CASA analysis because the tiny droplets of egg yolk are often counted as non-motile sperm. Despite the automatic nature of the analysis, care must still be taken to ensure correct set-up (e.g., settings are specific for each species) and validation of the settings. This is accomplished by making sure the computer software is accurately tracking each sperm along its trajectory and is not counting non-sperm cells or debris in the sample. After viewing the internal playback, subtle changes can be made by adjusting the gates for cell size and intensity under the QC plots accordingly. The computer software can incorrectly track cells when the field of view is not in focus or the illumination is insufficient. The illumination should be standardized before each reading under the optics set-up in order to ensure consistency across analyses.
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Evaluation of Semen It is also important to standardize the depth of the suspension. It can be difficult to track sperm motion at a depth greater than 20 µm, because sperm are constantly moving in and out of the plane of view, and at less than 10 µm depth sperm motion can be altered because the sperm head is not able to fully rotate. To maintain a standard depth the appropriate volume of sample for the size of the coverslip should be used (e.g., 10–12 µL for 22 mm2 ) and apply the coverslip carefully so as to avoid drift or the creation of areas under the coverslip that are deeper or thinner than others. Utilization of chambers with a fixed volume and depth would be advantageous for motility analysis in order to eliminate the variability with microscope slide preparation. DisposR R able fixed-depth chambers include Leja , Cell-Vu , R . Most of these chamMicroCell and Cell Vision bers were originally designed for sperm counting applications. Sperm motility can be significantly different between slide types and when compared to the standard microscope slide and coverslip (authors unpublished observations). Further investigation is required into the use of chambered slides for the analysis of equine sperm motility.
Morphology Sperm morphology is discussed in greater detail in Chapters 97 and 126. Only a brief introduction is therefore provided here. Abnormal sperm morphology has been associated with subfertility in many species. Evidence suggests that sperm with abnormal head morphology experience difficulty traveling through the female reproductive tract. Barriers to sperm transport, such as the cervix and cervical mucus in cattle and humans, are known to select for passage of morphologically normal sperm.19 The same may also be true for the uterotubal junction in the horse, although active sperm motility may also play a part here in transit. Nonetheless, sperm with misshapen heads and tails have impaired motility.20 Furthermore, it has been observed in vitro that sperm with abnormal heads were unable to penetrate the outer vestments of the ovum.21 There is limited data on the relationship between sperm morphology and fertility in the stallion (reviewed by Malmgren).22 However, it is a valuable test for sperm quality and can also be useful in identifying problems such as abnormal spermatogenesis, testicular degeneration, inflammation of the testis or epididymis, and heat stress, all of which result in a decline in sperm morphology. Sperm morphology is typically examined by bright-field microscopy (100× oil immersion) on airdried smears of stained semen. Background stains (e.g., eosin–nigrosin, India ink) are commonly used,
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in addition to general purpose cellular stains (e.g., Wright’s, Giemsa). Evaluation of sperm morphology on a wet mount of fixed cells viewed by phase-contrast or differential interference-contrast microscopy eliminates artifacts seen in stained smears.23, 24 At least 100–200 sperm should be evaluated and scored for the incidence of morphological defects. Abnormalities are usually broken down into the following categories: abnormal head, abnormal midpiece, abnormal tail, detached head, abnormal/ broken neck, proximal or distal droplets, and round (premature germ) cells. A sperm may be counted as having more than one type of defect; therefore outside of the percentage of normal sperm the numbers reflect the frequency of defects, not the actual number of abnormal cells. Technicians must be careful to ensure that semen preparation for morphological analysis does not introduce iatrogenic defects due to heat, cold (reflected midpieces), or hypo-osmotic shock (coiled tails). The preparation of stained smears may also cause physical damage to some cells.
pH The pH of gel-free semen can be measured using a correctly calibrated pH meter. The normal pH of stallion semen ranges from 7.2 to 7.7. Season of the year, frequency of ejaculation, and sperm concentration can influence the pH of semen. There is a reported negative correlation between seminal volume and pH and between the number of spermatozoa in the ejaculate and pH;25, 26 thus, as the number of spermatozoa in the ejaculate decreases, pH increases. Abnormally high semen pH can be associated with urospermia or infection of the internal genital tract.
Microbiology and cytology When collecting semen for bacterial culture, care must be taken not to contaminate the sample with bacteria from an external source. The penis and prepuce are ideal environments for bacterial growth and harbor a multitude of bacteria as part of their normal flora. Transfer of bacteria into the semen from these sites can be affected by poor hygiene protocols during semen collection, including insufficient sterilization of the artificial vaginas and the breeding phantom. Equine semen may contain a variety of bacteria, such as Streptococcus spp., Pseudomonas aeruginosa, Klebsiella pneumoniae, Acinobacter spp., Aerobacter spp., Proteus spp., Staphylococcus spp., and E. coli, and many of these organisms may not affect fertility.27–30 The majority of these bacteria are nonpathogenic, whereas others are capable of causing infection in mares. Usually, it is a positive culture in
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the mare that sends suspicion back to the stallion. Bacteria of concern are Pseudomonas aeruginosa, Klebsiella pneumoniae, E. coli, Taylorella equigenitalis, and beta-hemolytic streptococci. Indiscriminate washing of the penis with soaps can disturb the normal bacterial flora of the penis and encourage overgrowth of pathological forms. For artificial insemination, infected semen can be treated with antibiotics in the semen extender. Cytological examination is a very quick way to determine if the semen is contaminated with bacteria or if there is an infection in the male reproductive tract. An air-dried smear of raw semen is made upon a microscope slide and fixed with methanol. A number of differential staining kits are available commercially (e.g., Diff-Quik) that enable identification of white blood cell types. A count of more than six neutrophils per high-resolution field is indicative of infection.
Longevity of sperm motility Traditional tests of longevity involved incubating sperm in a dark environment at room temperature and removing aliquots for determination of sperm motility at regular intervals until 10% progressive motility remained. Longevity was then quoted as the number of hours it took to reach this cut-off. Today, with the widespread use of artificial insemination with cooled semen, longevity tests are now commonly determined at 5◦ C. Naturally, determination of the longevity or lifespan of a stallion’s sperm at 5◦ C is a critical test for those stallions intended for breeding with cool-shipped semen. The test for longevity also gives some indication of how well sperm can tolerate stress in their environment, which could relate to their lifespan within the female reproductive tract or their ability to withstand the stress of freezing; however, currently there are no data to support this. To determine longevity, raw semen is extended to 40–50 × 106 /mL and cooled (to 5◦ C) in a commercially available cooled semen shipper for at least 48 hours. An aliquot is removed and warmed (5 minutes) for motility analysis at 6, 12, 24, and 48 hours. Knowing the longevity of cooled semen allows a clinician to determine the optimal time period for shipping and insemination in addition to packaging an appropriate dose of semen, the industry standard being 500 × 106 progressively motile sperm at 24 h. Ideally, stallions in a cooled semen program should retain acceptable motility (> 30% progressive) for at least 24 h and preferably 48 h. Semen from some stallions may have a limited lifespan, necessitating same-day airline service for transport of cooled semen. Stallions with superior longevity semen can
maintain at least 30% progressive motility at 72 h. It must be remembered that the longevity of the sample may be influenced by a variety of factors including the dilution factor and final concentration of the extended semen, the extender formulation, and potentially the type of commercial shipper used. Furthermore, centrifugation and removal of seminal plasma may improve longevity of sperm motility for some stallions.
Hypo-osmotic swelling test The hypo-osmotic swelling (HOS) test was proposed by Jeyendran et al.31 as a relatively simple test to evaluate sperm membrane integrity. An important property of the sperm cell membrane is its ability to permit selective transport of molecules and adjust to the osmotic changes in its environment. When cells encounter hypo- or hypertonic environments they tend to swell or shrink, respectively, due to the influx or efflux of water during establishment of osmotic equilibrium. To perform the HOS test, a sample of raw semen (100 µL) is incubated for 60 minutes at 37◦ C in 1 mL of pre-warmed sucrose solution adjusted to 100 mOsm (1.712 g in 50 mL dH2 O).32 When exposed to the hypo-osmotic sucrose solution, sperm with an intact functional membrane will swell to obtain osmotic equilibrium with their environment. The swelling of the membrane causes the tail of the spermatozoa to coil. Those sperm with damaged membranes will retain straight tails. A minimum of 100 sperm are evaluated under bright-field microscopy and the percentage of them that show the typical coiled tail abnormality (denoted as HOS+) are counted. According to Jeyendran et al.,33 human ejaculates are considered normal when 60% or more HOS+ sperm are observed. In humans, the HOS test has been reported to correlate highly with other predictive tests for fertility, for example, hamster oocyte penetration33 and invitro fertilization.34 In boars, a modified HOS test correlated fairly well with pregnancy rates.35 While the HOS test has been validated for use on stallion sperm,32, 36 it is not yet clear how closely the results correlate with fertility. However, Jeyendran et al.33 have also suggested that the HOS can be used as a predictor for the success of cryopreservation. Magistrini et al.37 demonstrated a significant correlation between post-thaw motility of stallion semen and the results of the HOS in raw semen. Because the HOS test represents membrane integrity and the ability to regulate water transport, it makes sense that the test would relate predominantly to the evaluation of frozen-thawed semen, either in predicting freezability or in the assessment of post-thaw function.
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Live-dead staining of semen smears With this technique, the integrity of the plasma membrane is determined by the ability of the viable cell to exclude eosin dye, whereas the stain will passively cross the membranes of dead cells that have a compromised membrane. A drop of eosin stain is placed upon a microscope slide and beside it a small drop of diluted semen. The end of another microscope slide is used to mix the semen with the stain and then drag it across the slide to create a smear that is then left to air dry. The slide is evaluated under oil immersion at 100× and the number of live (unstained) and dead (stained purple) cells are counted and translated into a percentage. Trypan blue and Chicago sky blue are other stains that can be used for the determination of viability by light microscopy.38
Advanced sperm evaluation tests Tests are now available for an increasing range of sperm variables including viability, capacitation, acrosomal integrity, mitochondrial activity, and chromatin integrity. Most of these methods require the application of fluorescence microscopy or flow cytometry and, although the general clinician does not have access to the advanced equipment required to perform these techniques, such tests are finding their way into mainstream practice through referral to independent or university laboratories. The limitations of these assays are that they only assess a single attribute from a multitude of characteristics that a sperm must possess in order to fertilize an egg. A stallion may pass one test but fail another; therefore a combination of tests offers greater promise for the reliable identification of subfertile individuals. For this reason, flow cytometry has become the ideal tool because it allows the objective, rapid and simultaneous analysis of a number of sperm characteristics across a large population of sperm. Live-dead staining with fluorescent probes Sperm membrane integrity can also be determined by fluorescent probes. Usually a combination of two probes is used, one for staining live cells and one for staining dead cells, for example, the combination of carboxyfluorescein diacetate (CFDA) and propidium iodide39 or calcein-AM and ethidium homodimer.40 Calcein-AM and CFDA are non-fluorescent membrane-permeable fluorophores that undergo hydrolysis of their ester bonds by cellular esterases resulting in the formation of the non-membranepermeable products calcein or carboxyfluorescein that accumulate within the cell and exhibit a green fluorescence. The red fluorescent counter stains, propidium iodide and ethidium, are membrane-
Figure 124.7 Fluorescent micrograph showing equine sperm labeled with propidium iodide and Syber-14. Dead cells show red fluorescence with propidium iodide and live cells are stained green by Syber-14. (See color plate 41).
impermeable, nucleic acid-specific fluorophores; they only pass through the compromised membranes of dead cells and therefore do not stain the nuclei of live cells. A recently developed membranepermeant DNA stain for live cells, Syber-14, in combination with propidium iodide41 is available as a R LIVE/DEAD Sperm Viability Kit from Invitrogen (Carlsbad, CA) and is commonly used for the assessment of viability in equine sperm. Visualization of fluorescent live-dead stains can be performed on a fluorescent microscope (Figure 124.7) or quantified by flow cytometric analysis. Sperm–oocyte binding test The binding of sperm to the zona pellucida is an important preliminary step in fertilization. In humans it has been shown that an in vitro sperm zona binding assay using homologous oocytes can be used to predict the outcome of in vitro fertilization.42 Fazeli et al.43 found that batches of oocytes varied greatly in their ability to bind spermatozoa. To overcome this disadvantage, the use of matching halves of the zona pellucida from the same oocyte has been developed.44 In the hemi-zona assay (HZA) the number of sperm bound to a particular hemi-zona is only comparable to the number bound to the matching hemi-zona. Therefore a stallion of unknown fertility can be compared to a stallion of known fertility. Fazeli et al.43 and Meyers et al.45 reported that numbers of sperm bound to the zona pellucida were higher for fertile stallions than for subfertile individuals. A significant disadvantage of the HZA is the need for available equine oocytes and the technical skill required for bisecting the zonae. An alternative oocytebased approach to assessing sperm fertilizing capacity is the zona-free hamster oocyte penetration test.
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However, even studies that have reported an association between this assay and stallion fertility have noted anomalies such as subfertile stallions with high penetration rates.46 Capacitation After ejaculation, during transit through the female reproductive tract, the sperm cell must undergo a complex series of maturational changes, termed capacitation, that enable it to recognize and bind to the zona pellucida. Only capacitated sperm can bind to the zona pellucida and trigger the subsequent events that lead to fertilization.47 These changes include reorganization of membrane proteins, metabolism of membrane phospholipids, a reduction in membrane cholesterol levels, and hyperactivation.47 Spermatozoa can be tested for their ability to undergo capacitation in vitro in response to inducing agents (bicarbonate or cAMP), or the number of cells already capacitated in the ejaculate can be determined. The presence of capacitated cells in the fresh ejaculate suggests ‘premature capacitation’; these cells are believed to have reduced longevity.48 Assays for capacitation include chlortetracycline (CTC) staining and Merocyanine 540.49 CTC is a fluorescent antibiotic that binds to sperm cells in a calcium-dependent manner and distinguishes three different stages of sperm activation: non-capacitated, capacitated acrosome-intact, and capacitated acrosome-reacted. The assay is performed on fixed cells and is examined by fluorescent microscopy; it is a time-consuming protocol that also suffers the disadvantage of a limited understanding about how CTC interacts with the sperm membrane at the molecular level. Merocyanine 540 is a lipophilic fluorescent dye that stains cell membranes with increasing affinity as their lipid components become more disorganized, as is the case with sperm capacitation. It has the advantage of being analyzed by R flow cytometry and can be combined with Yo-Pro 1 for membrane integrity or fluorescent probes for acrosomal integrity.
of sperm with intact acrosomes at ejaculation.50 Once sperm have undergone the acrosome reaction they are no longer able to participate in fertilization. The acrosome reaction can be induced in vitro by incubating spermatozoa with calcium ionophore A2318751 or with physiological inducers like the zona pellucida,52 follicular fluid,53 or progesterone.54 In the study of Meyers et al.,54 fertile stallions showed higher rates of acrosome reaction in response to progesterone treatment than subfertile stallions. Furthermore, Rathi et al.55 found that the percentage of spermatozoa in an ejaculate with exposed progesterone receptors on their plasma membrane after incubation under capacitation conditions was highly correlated with the fertility of the donor stallion. Chlortetracycline can be used to differentiate acrosome-intact from acrosome-reacted spermatozoa; however, the most commonly used fluorescent probes are fluorescein isothiocyanate (FITC, green fluorescent fluorophore) conjugated peanut agglutinin (FITC-PNA)56 or FITC-conjugated Pisum sativum agglutinin (FITC-PSA)57 which offer the advantage over CTC of being analyzed by flow cytometry. PSA binds to α-mannose and α-galactose moieties of the acrosomal membrane; it cannot penetrate an intact acrosomal membrane, therefore only acrosomereacted or damaged sperm will stain. PNA binds to β-galactose moieties associated with the outer acrosomal membrane (OAM) of sperm, and ethanol permeabilization of the sperm membrane allows access of the probe to the OAM indicating acrosome-intact cells (Figure 124.8). These indicators of acrosomal status are usually combined with a viability probe, such as propidium iodide, so that four groups of spermatozoa can be identified: live acrosome-intact, live
Acrosome reaction Following capacitation and binding to the zona pellucida, the acrosome reaction, an exocytotic event that is required for penetration of the zona pellucida, is initiated. The number of acrosome-intact or -reacted spermatozoa can be determined in the fresh ejaculate or the number of sperm capable of responding to initiation of the acrosome reaction in vitro can be assayed. Visualization of sperm acrosome integrity may be beneficial in the evaluation of semen quality, since male infertility could be caused by a lack
Figure 124.8 Fluorescent micrograph showing permeabilized sperm labeled with FITC-conjugated peanut agglutinin and propidium iodide. Acrosome-intact sperm have a fluorescent green cap. (See color plate 42).
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Evaluation of Semen acrosome-reacted, dead acrosome-intact, and dead acrosome-reacted. The physiology of sperm capacitation and the acrosome reaction are discussed in further detail in Chapters 97 and 102. Evaluation of acrosomal function is described in Chapter 152. Mitochondrial function Correct functioning of the mitochondria and the associated generation of energy in the form of adenosine triphosphate (ATP) is required for sperm motility. Rhodamine 123 (R123), MitotrackerTM (MITO) and 5,5 ,6,6 -tetrachloro-1,1 ,3,3 -tetraethylbenzimidazolyl-carbo-cyanine iodide (JC-1) have been used to evaluate the mitochondrial function of spermatozoa. R123 and MITO are transported into the actively respiring mitochondria and their accumulation in the organelle causes them to fluoresce green. All functioning mitochondria stain green with these two fluorochromes, therefore no distinction can be made between spermatozoa exhibiting different respiratory rates. In contrast, JC-1 can distinguish between sperm with a low or high mitochondrial membrane potential. JC-1 is internalized by all functioning mitochondria and fluoresces green. In highly functioning mitochondria the concentration of JC-1 increases and the stain forms aggregates that fluoresce red-orange. When human sperm were divided into low and high membrane potential groups, based on JC-1 fluorescence, the in vitro fertilization rates were higher in the high potential group than in the low potential group.58 Garner et al.59 compared all three mitochondrial probes in bull spermatozoa and found that they correlated well with each other as well as cell viability and sperm motility. Sperm chromatin structure assay The sperm chromatin structure assay (SCSA) is discussed in more detail in Chapter 154 therefore, only a brief introduction will be given here. The chromatin in the sperm nucleus is highly condensed and normally resistant to denaturation. Abnormal sperm chromatin structure renders sperm susceptible to denaturation (i.e., DNA damage), and may also interfere with decondensation, a prerequisite for male pronucleus formation during fertilization. Fertilization with DNA-damaged spermatozoa could lead to paternal transmission of defective genetic material, with adverse consequences for embryonic development.60 In humans, reduced sperm chromatin stability has been related to recurrent abortion61 and may provide some explanation for the incidence (18%) of embryonic loss in normal mares between fertilization and Day 50 of pregnancy, as reviewed by Ball.62
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The SCSA uses the metachromatic properties of acridine orange to distinguish between denatured (red fluorescence = single stranded) and native (green fluorescence = double stranded) DNA.63 The results of the SCSA have been demonstrated to be highly correlated with fertility in both bull64 and boar.65 Kenney et al.66 reported higher rates of sperm chromatin denaturation in semen from subfertile compared to fertile stallions, and a negative correlation between denaturation score and seasonal pregnancy rate. In a population of fertile stallions it was found that fertility decreased as the percentage of sperm susceptible to denaturation increased.67 The SCSA offers the advantage of measuring a different characteristic of sperm function than traditional semen evaluation tests; therefore it may identify a reason for subfertility in a stallion with normal morphology and motility.
Conclusion Semen evaluation can be either a simple process using traditional methods or a complicated one, utilizing state-of-the-art equipment and the latest biomarkers available for sperm function. Currently, there is no single test that can predict the fertility of a stallion and it is unlikely, given the complexity of sperm function, that there will ever be just one test. The ability of a sperm to fertilize the egg is dependent upon many characteristics of sperm function, in addition to extrinsic factors that come into play, such as the seminal plasma, zona pellucida, and oviductal environment, not to mention the inherent fertility of the mare. The goal is to identify a battery of tests that together analyze the most important aspects of semen quality and sperm function in the hopes of identifying a characteristic that might be deficient. In this regard it is critical to know what is normal and what is acceptable. Herein lies a challenge since studies including large populations of stallions that can confidently determine the normal distribution or correlate a particular variable with fertility is very difficult to achieve. However, science and technology is constantly advancing our understanding of the physiology of sperm function and developing new tools for semen analysis, so the future looks bright.
References 1. Christensen P, Stryhn H, Hansen C. Discrepancies in the determination of sperm concentration using BurkerTurk, Thoma and Makler counting chambers. Theriogenology 2005;63:992–1003. 2. Brazil C, Swan SH, Tollner CR, Treece C, Drobnis EZ, Wang C, Redmon JB, Overstreet JW. Quality control of laboratory methods for semen evaluation in a multicenter research study. J Androl 2004;25:645–56.
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3. Freund M, Carol B. Factors affecting hemocytometer counts of sperm concentration in human semen. J Reprod Fertil 1964;8:149–55. 4. Hansen C, Vermeiden T, Vermeiden JPW, Simmet C, Day BC, Feitsma H. Comparison of FACSCount AF system, Improved Neubauer hemocytometer, Corning 254 photometer, SpermVision, UltiMate and NucleoCounter SP-100 for determination of sperm concentration of boar semen. Theriogenology 2006;66:2188–94. 5. Douglas-Hamilton DH, Smith NG, Kuster CE, Vermeiden JP, Althouse GC. Particle distribution in low-volume capillary-loaded chambers. J Androl 2005;26(1):107–14. 6. Douglas-Hamilton DH, Smith NG, Kuster CE, Vermeiden JP, Althouse GC. Capillary-loaded particle fluid dynamics: effect on estimation of sperm concentration. J Androl 2005;26(1):115–22. 7. Zinaman MJ, Uhler ML, Vertuno E, Fisher SG, Clegg ED. Evaluation of computer-assisted semen analysis (CASA) with IDENT stain to determine sperm concentration. J Androl 1996;17(3):288–92. 8. Strader LF, Linder RE, Perreault SD. Comparison of rat epididymal sperm counts by IVOS HTM-IDENT and hemacytometer. Reprod Toxicol 1996;10(6):529–533. 9. Wessel MT, Althouse GC. Validation of an objective approach for simultaneous assessment of viability and motility of fresh and cooled equine spermatozoa. Proceedings of the International Symposium on Equine Reproduction 2006. Anim Reprod Sci 2006;94:21–2. Abstract. 10. Gagnon C. Regulation of sperm motility at the axonemal level. Reprod Fertil Dev 1995;7(4):847–55. 11. Mortimer ST. A critical review of the physiological importance and analysis of sperm movement in mammals. Hum Reprod Update 1997;3(5):403–39. 12. Jasko DJ, Little TV, Lein DH, Foote RH. Comparison of spermatozoal movement and semen characteristics with fertility in stallions: 64 cases (1987–1988). J Am Vet Med Assoc 1992;200(7):979–85. 13. Jasko DJ, Lein DH, Foote RH. A comparison of two computer-automated semen analysis instruments for the evaluation of sperm motion characteristics in the stallion. J Androl 1990;11(5):453–9. 14. Boyers SP, Davis RO, Katz DF. Automated semen analysis. Curr Probl Obstet Gynecol Fertil 1989;I(5):167–200. 15. Mortimer ST. CASA: practical aspects. J Androl 2000;21(4):515–24. 16. Mortimer D, Serres C, Mortimer ST, Jouannet P. Influence of image sampling frequency on the perceived movement characteristics of progressively motile human spermatozoa. Gamete Res 1988;20(3):313–27. 17. Jasko DJ, Little TV, Smith K, Lein DH, Foote RH. Objective analysis of stallion sperm motility. Theriogenology 1988;30(6):1159–67. 18. Varner DD, Vaughan SD, Johnson L. Use of a computerized system for evaluation of equine spermatozoal motility. Am J Vet Res 1991;52(2):224–30. 19. Barros C, Vigil P, Herrera E, Arguello B, Walker R. Selection of morphologically abnormal sperm by human cervical mucus. Arch Androl 1984;12 (suppl):95–100. 20. Dresdner RD, Katz DF. Relationship of mammalian sperm motility and morphology to hydrodynamic aspects of cell function. Biol Reprod 1981;25:920–30. 21. Kot MC, Handel MA. Binding of abnormal sperm to mouse egg zonae pellucidae in vitro. Gamete Res 1987;18:57–63. 22. Malmgren L. Sperm morphology in stallions in relation to fertility. Acta Vet Scand 1992;88 (suppl) 52:281–7.
23. Ball L, Pickett BW, Gerbauer MR. Staining technique and stallion sperm morphology. J Anim Sci 1971;33:248. 24. Hurtgen JP, Johnson LA. Fertility of stallions with abnormalities of the sperm acrosome. J Reprod Fertil 1982;32 (Suppl):15–20. 25. Pickett BW, Voss JL, Bowen RA, Squires EL, McKinnon, AO. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. Proc. Am. Assoc. Eq. Prac. 1987;33:487–518. 26. Pickett BW, Voss JL, Bowen RA, Squires EL, McKinnon, AO. Comparison of seminal characteristiocs of stallions that passed or failed seminal evaluation. In: Proceedings of 11th International Congress on Animal Reproduction and Artificial Insemination 1988; vol 3, pp. 380. 27. Hughes JP, Asbury AC, Loy RG, Burd HE. The occurrence of Pseudomonas in the genital tract of stallions and its effects on fertility. Cornell Vet 1967;57:53–69. 28. Burns SJ, Simpson RB, Snell JR. Control of microflora in stallion semen with a semen extender. J Reprod Fertil Suppl 1975;(23):139–42. 29. Kenney RM. Clinical fertility evaluation of the stallion. In: Proceedings of the American Association of Equine Practitioners 1975; pp. 336–55. 30. Merkt H, Klug E, Bohm KH, Weiss R. Recent observations concerning Klebsiella infections in stallions. J Reprod Fertil Suppl. 1975;(23):143–5. 31. Jeyendran RS, van der Ven HH, Perez-Pelaez M, Crabo BG, Zaneveld LJ. Development of an assay to assess the functional integrity of the human sperm membrane and its relationship to other semen characteristics. J Reprod Fertil 1984;70(1):219–28. 32. Nie GJ, Wenzel JGW. Adaptation of the hypoosmotic swelling test to assess functional integrity of stallion spermatozoal membranes. Theriogenology 2001;55:1005–18. 33. Jeyendran RS, van der Venn HH, Zaneveld LJD. The hypoosmotic swelling test: an update. Arch Androl 1992; 29:105–16. 34. Van der Venn HH, Jeyendran RS, Al-Hasani S, P´erez-Pel`aez M, Diedrich K, Zaneveld LJD. Correlation between human sperm swelling in hypoosmotic medium (hypoosmotic swelling test) and in vitro fertilization. J Androl 1986;7:190–6. 35. P´erez-Llano B, Lorenzo JL, Yenes P, Trejo A, Garcia-Casado P. A short hypoosmotic swelling test for the prediction of boar sperm fertility. Theriogenology 2001;56:387–98. 36. Nield DM, Chaves MG, Flores MC, Mora NG, Beconi MT, ¨ Aguero A. Hypoosmotic swelling test in equine spermatozoa. Theriogenology 1999;51:721–7. 37. Magistrini M, Sattler M, Yvon JM, Vidament M. Freezability of stallion spermatozoa evaluated by motility, membrane integrity and ATP content. 34th Annual Meeting of the Society of Cryobiology, Barcelona, June 8–12. Cryobiology 1997;35(4):88, Abstract. ´ olgyi ¨ 38. Kutv G, Stefler J, Kov´acs A. Viability and acrosome staining of stallion spermatozoa by Chicago sky blue and Giemsa. Biotech Histochem 2006;81(4–6):109–17. 39. Garner DL, Pinkel D, Johnson LA, Pace MM. Assessment of spermatozoal function using dual fluorescent staining and flow cytometric analyses. Biol Reprod 1986;34(1): 127–38. 40. Althouse GC, Hopkins SM. Assessment of boar sperm viability using a combination of two fluorophores. Theriogenology 1995;43(3):595–603. 41. Garner DL, Johnson LA, Yue ST, Roth BL, Haugland RP. Dual DNA staining assessment of bovine sperm
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viability using SYBR-14 and propidium iodide. J Androl 1994;15(6):620–9. Liu DY, Lopata A, Johnston WI, Baker HW. A human sperm-zona pellucida binding test using oocytes that failed to fertilize in vitro. Fertil Steril 1988;50(5):782–8. Fazeli AR, Steenweg W, Bevers MM, Bracher V, Parlevliet J, Colenbrander B. Use of sperm binding to homologous hemizona pellucida to predict stallion fertility. Equine Vet J 1993;15(Suppl):57–9. Fazeli AR, Steenweg W, Bevers MM, van den Broek J, Bracher V, Parlevliet J, Colenbrander B. Relation between stallion sperm binding to homologous hemizonae and fertility. Theriogenology 1995;44(5):751–60. Meyers SA, Liu IK, Overstreet JW, Vadas S, Drobnis EZ. Zona pellucida binding and zona-induced acrosome reactions in horse spermatozoa: comparisons between fertile and subfertile stallions. Theriogenology 1996;46(7):1277–88. Wilhelm KM, Graham JK, Squires EL. Comparison of the fertility of cryopreserved stallion spermatozoa with sperm motion analyses, flow cytometric evaluation, and zona-free hamster oocyte penetration. Theriogenology 1996;46(4):559–78. Yanagimachi R. Mammalian fertilisation. In: Knobil E, Neill JD (eds) Physiology of Reproduction, 2nd edn. New York: Raven Press, 1994; pp. 189–317. Watson PF. Recent developments and concepts in the cryopreservation of spermatozoa and the assessment of their post-thawing function. Reprod Fertil Dev 1995;7(4):871–91. Rathi R, Colenbrander B, Bevers MM, Gadella BM. Evaluation of in vitro capacitation of stallion spermatozoa. Biol Reprod 2001;65(2):462–70. Zhang J, Boyle MS, Smith CA, Moore HD. Acrosome reaction of stallion spermatozoa evaluated with monoclonal antibody and zona-free hamster eggs. Mol Reprod Dev 1990;27(2):152–8. Zhang JJ, Muzs LZ, Boyle MS. Variations in structural and functional changes of stallion spermatozoa in response to calcium ionophore A23187. J Reprod Fertil Suppl 1991;44:199–205. Ellington JE, Ball BA, Yang X. Binding of stallion spermatozoa to the equine zona pellucida after coculture with oviductal epithelial cells. J Reprod Fertil 1993;98(1):203–8. Tarlatzis BC, Danglis J, Kolibianakis EM, Papadimas J, Bontis J, Lagos S, Mantalenakis S. Effect of follicular fluid on the kinetics of human sperm acrosome reaction in vitro. Arch Androl 1993;31(3):167–75. Meyers SA, Overstreet JW, Liu IK, Drobnis EZ. Capacitation in vitro of stallion spermatozoa: comparison of progesterone-induced acrosome reactions in fertile and subfertile males. J Androl 1995;16(1):47–54.
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55. Rathi R, Nielen M, Cheng FP, van Buiten A, Colenbrander B. Exposure of progesterone receptors on the plasma membranes of stallion spermatozoa as a parameter for prediction of fertility. J Reprod Fertil 2000;56 suppl:87–91. 56. Cheng FP, Fazeli A, Voorhout WF, Marks A, Bevers MM, Colenbrander B. Use of peanut agglutinin to assess the acrosomal status and the zona pellucida-induced acrosome reaction in stallion spermatozoa. J Androl 1996;17(6):674– 82. 57. Farlin ME, Jasko DJ, Graham JK, Squires EL. Assessment of Pisum sativum agglutinin in identifying acrosomal damage in stallion spermatozoa. Mol Reprod Dev 1992;32(1): 23–7. 58. Kasai T, Ogawa K, Mizuno K, Nagai S, Uchida Y, Ohta S, Fujie M, Suzuki K, Hirata S, Hoshi K. Relationship between sperm mitochondrial membrane potential, sperm motility, and fertility potential. Asian J Androl 2002;4(2):97– 103. 59. Garner DL, Thomas CA, Joerg HW, DeJarnette JM, Marshall CE. Fluorometric assessments of mitochondrial function and viability in cryopreserved bovine spermatozoa. Biol Reprod 1997;57(6):1401–6. 60. Aitken RJ, Krausz C. Oxidative stress, DNA damage and the Y chromosome. Reproduction 2001;122(4):497–506. 61. Ibrahim ME, Pedersen H. Acridine orange fluorescence as male fertility test. Arch Androl 1988;20(2):125–9. 62. Ball BA. Embryonic loss in mares: incidence, possible causes, and diagnostic considerations. Vet Clin N Am Equine Pract 1988;4(2):263–90. 63. Evenson DP, Jost LK, Marshall D, Zinaman MJ, Clegg E, Purvis K, de Angelis P, Claussen OP. Utility of the sperm chromatin structure assay as a diagnostic and prognostic tool in the human fertility clinic. Hum Reprod 1999;14(4):1039–49. 64. Ballachey BE, Evenson DP, Saacke RG. The sperm chromatin structure assay: relationship with alternate tests of semen quality and heterospermic performance of bulls. J Androl 1988;9(2):109–15. 65. Evenson DP, Thompson L, Jost L. Flow cytometric evaluation of boar semen by the sperm chromatin structure assay as related to cryopreservation and fertility. Theriogenology 1994;41(3):637–51. 66. Kenney RM, Evenson DP, Garcia MC, Love CC. Relationships between sperm chromatin structure, motility and morphology of ejaculated sperm and seasonal pregnancy rate. Biol Reprod monogr. ser. 1 (Equine Reprod) 1995;VI:647–53. 67. Love CC, Kenney RM. The relationship of increased susceptibility of sperm DNA to denaturation and fertility in the stallion. Theriogenology 1998;50(6):955–72.
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Chapter 125
Spermatozoal Motility Aaron D.J. Hodder and Irwin K.M. Liu
Introduction
Origin of sperm motility
Of the parameters examined when stallion semen is evaluated, assessment of sperm motility is considered by many to be one of the most important. Motility is generally considered to be a reflection of the viability of sperm in a given sample.1 Immotile or poorly motile sperm are considered incapable of fertilization without assisted reproductive techniques.2 Motility alone should never be used to estimate the fertility of a stallion, as spermatozoa must possess many characteristics to be able to fertilize an oocyte.3 The fertility of an individual stallion may be unsatisfactory in spite of satisfactory motility.4 There are many instances in which assessment of sperm motility should be conducted. Sperm motility should be determined in ejaculates from stallions undergoing breeding soundness evaluation, as well as those being examined for subfertility or infertility. Calculation of a breeding dose of semen, whether fresh or cooled transported, requires estimation of sperm motility to ensure that adequate numbers of progressively motile sperm are used. When a shipment of chilled semen is received for the purpose of breeding a mare, sperm motility should be assessed in a small aliquot of the semen to ensure that the sample is of sufficient quality to optimize the chances of conception and to identify any deterioration in the semen that may have occurred during shipment. Assessment of sperm motility over time in extended semen samples is commonly used to assess sperm longevity and to determine whether chilled, transported semen from a given stallion can be used to breed mares. When subjecting sperm to cryopreservation, sperm motility in a sample should be assessed both before freezing and after thawing to ensure that semen is of adequate quality to be frozen and that it maintains adequate post-thaw motility to be considered commercially acceptable.
Upon release from the seminiferous epithelium (spermiogenesis), spermatozoa are non-motile, and are propelled through the rete testis, efferent ductules, and epididymis by peristaltic contractions. During passage through the epididymis, equine sperm undergo a number of maturational changes, including the acquisition of motility. The motility of equine spermatozoa is first identified in the distal corpus, but is especially evident in the cauda epididymis.5
Types of motility Two types of physiological motility have been recognized in mammalian spermatozoa.6 Activated motility is typified by a symmetrical, lower amplitude flagellar beat and results in sperm movement in a relatively straight line. This kind of motility is seen in freshly ejaculated sperm and is thought to be necessary to move the sperm toward, and into, the oviduct. Activated motility is important for normal fertility as most immotile sperm are unable to reach the uterotubal junction. In contrast to activated motility, sperm with hyperactivated motility display an asymmetric flagellar beat with higher amplitude in aqueous media. This results in sperm traveling in circular or figure-ofeight paths. Hyperactivated motility is typically seen in sperm recovered from the site of fertilization and is thought to enable sperm to detach from the oviductal epithelium, reach the site of fertilization, and penetrate the cumulus and zona pellucida of the oocyte.
Structures responsible for sperm motility The structure responsible for providing a propulsive force in the spermatozoon is the axoneme, which
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Spermatozoal Motility extends throughout the flagellum (tail) of the sperm. The axoneme is composed of a central pair of microtubules, surrounded by a ring of nine outer microtubule doublets. These outer microtubule doublets are connected by nexin links, and a series of nine radial spokes that extend between the central pair of microtubules and the ring of the nine outer doublets. Each of the outer microtubule doublets possesses inner and outer dynein arms, which are responsible for generating flagellar movement. ATPase present in these dynein arms converts chemical energy to mechanical motion, which results in adjacent outer doublet microtubules moving relative to each other and causing flagellar bending. Other structural elements thought to assist sperm motility include the outer dense fibers and the fibrous sheath. Located outside each of the nine outer microtubule doublets, the outer dense fibers are thought to provide structural support to the flagellum. These fibers run from the neck of the sperm, through the midpiece and most of the principal piece, and are not present in the end piece. The fibrous sheath is present along the length of the principal piece and is also thought to provide rigid support to the flagellum. Recent evidence suggests a more active role for the fibrous sheath in sperm motility.7 Proteins involved in motility signaling pathways and metabolism have been identified within the fibrous sheath. The dynein arms located on the outer microtubule doublets require an energy source to generate flagellar movement. This energy is provided by ATP, which is produced by mitochondria via aerobic respiration and locally within the principal piece through glycolysis. Sperm mitochondria are found only in the mitochondrial sheath of the midpiece. The mitochondrial sheath is arranged in a helical fashion clearly visible with electron microscopy. A typical stallion spermatozoon has about 50 gyri, or helical turns, of mitochondria.3
Analysis of sperm motility Sperm motility should be assessed as soon as possible after semen collection.8 Whether using subjective or objective techniques, all objects and materials that come into contact with semen should be maintained at 37◦ C to avoid the detrimental effects of thermal shock on sperm motility.9 All equipment used should be clean, and, where possible, disposable.10 Using raw semen to evaluate sperm motility can be problematical if sperm concentration is high, as it may be difficult to examine individual sperm cells. It may also result in the underestimation of motility due to sperm agglutination and sperm cells adhering to the glass.10 It has, however, been suggested
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that sperm motility in a sample of raw semen be examined as a control for identifying any potentially detrimental effects of the semen extender and/or antibiotics on sperm motility in some stallions.11 Proficiency in assessing sperm motility in a highly concentrated sample may be beneficial when assessing post-thaw motility of frozen–thawed semen, which is typically frozen at high sperm concentrations. For accurate estimation, and to standardize semen motility evaluation, semen should be diluted with an appropriate extender to a concentration of between 25 and 50 million sperm cells/mL.10, 11 The extender should be maintained at a temperature of 37◦ C before addition to raw semen to avoid the detrimental effects of thermal shock on sperm motility. Likewise, chilled semen should be warmed to 37◦ C before evaluation of sperm motility; otherwise, values for sperm motility may be underestimated. Subjective The minimum requirements for subjective assessment of sperm motility include microscope slides, coverslips, semen extender, and a microscope. Ideally, sperm motility is assessed using a phase contrast microscope at 200× magnification.8 Subjective estimation of sperm motility should include total motility (motility in any form), progressive motility (sperm displaying rapid, linear movement), and velocity [subjectively graded on a scale of 0 (immotile) to 4 (rapidly motile)].11 Total and progressive motility are expressed as percentages of the total sperm population. Accurate, subjective assessment of sperm motility is gained through experience and consistency in the evaluation technique. A small drop (5–10 µL) of extended semen at 37◦ C is placed on a prewarmed microscope slide and a coverslip is placed on the sample. Care should be taken to avoid trapping air bubbles in the semen, as this may affect motility estimation.8, 10 It is important that the sample is maintained at 37◦ C throughout the evaluation. This can be achieved by using a stage warmer or other device designed to maintain a constant temperature. Sperm are examined near the center of the coverslip, taking care to avoid the edges where sperm motility will be underestimated owing to the negative effects of drying and exposure to air on the spermatozoa.10 Four to six fields are examined in a timely manner and the estimates for both total and progressive motility are averaged over the fields examined. Accuracy may be increased by using more than one examiner.8 Objective In an attempt to eliminate variations in motility estimates which may arise with subjective assessment
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of sperm motility, objective techniques have been developed. These techniques include computerized analysis, time-lapse photomicrography, frame-byframe playback videomicrography, and spectrophotometry.11 Computer-assisted sperm analysis (CASA) analyzes the tracks of a large number of sperm and, from this, determines a number of parameters, including percentage of motile sperm, percentage of progressively motile sperm, amplitude of lateral head displacement, average path velocity, and curvilinear velocity. The disadvantage of using these systems lies in the expense of the equipment needed, thereby limiting their routine use to facilities with a large caseload of stallion work. Subjective assessment remains a common practice in routine equine breeding management.
Factors affecting motility A number of factors have been identified that affect sperm motility in the stallion. These factors may be physiologic, pathologic, or as a result of poor semen collection and processing techniques. The clinician must consider these factors when attempting to identify the cause of reduced or absent sperm motility (asthenozoospermia) in a semen sample. Physiological factors A typical ejaculate from a peripubertal stallion will exhibit decreased sperm motility, decreased sperm concentration, and increased morphological defects in comparison with ejaculates later in life, or from older stallions.12 Stallions that have been sexually rested will often have a high number of sperm in the first ejaculate. Sperm from this first ejaculate typically display decreased motility in comparison with later collections when the stallion is on a regular semen collection schedule. The higher number of sperm and decreased motility typically seen in the first ejaculate of a sexually rested stallion is thought to be due to sperm storage in the tail of the epididymis and ductus deferens. Conflicting data exist regarding the effect of season on sperm motility, both in fresh semen and in frozen–thawed semen. Pickett et al.13 reported that the percentage of progressively motile sperm determined immediately after collection and extension was not affected by month. In contrast, Magistrini et al.14 found that the percentage of motile sperm was higher in the summer than in the winter. Pathological factors Pathological conditions affecting the testes and/or scrotum, or fever of any origin, often result in a fo-
cal or diffuse elevation in testicular temperature. As normal spermatogenesis in the stallion occurs below normal body temperature, any prolonged increase in temperature will disrupt normal spermatogenesis and result in testicular degeneration. Researchers have used scrotal insulation to elevate scrotal temperature and mimic pathological conditions such as fever and scrotal edema in the stallion. Friedman et al.15 examined several parameters in the semen of stallions following scrotal insulation for either 24 or 48 hours. Stallions undergoing scrotal insulation for 48 hours showed significant declines in both total and progressive motility beginning on the first day after insulation. The decline continued until reaching its lowest levels between 15 and 30 days after insulation. After this time, total and progressive motility increased, reaching pretreatment levels by day 50. While stallions exposed to scrotal insulation for only 24 hours also showed declines in total and progressive motility, these changes were not profound. Blanchard et al.16 subjected two stallions to 36 hours of scrotal insulation and observed a decrease in progressive motility by 1–2 weeks post insulation. Progressive motility reached its lowest point by 3–3.5 weeks post insulation, and returned to pretreatment levels by 7 weeks. Although testicular degeneration can result from a testicular insult, testicular degeneration may also occur in older stallions as a result of age. Age-related or idiopathic testicular degeneration results in a decline in overall semen quality, including percentages of motile sperm.17 While many potential treatment options have been investigated, none have proven successful. Instead, stallions with idiopathic testicular degeneration must be managed carefully to optimize the chances of pregnancy. Management factors Researchers in Texas examined the effect of feeding a docosahexaenoic acid (DHA)-enriched nutraceutical (as a source of omega-3 fatty acid) on the quality of fresh, cooled, and frozen stallion semen and found a significant improvement in motion characteristics of sperm (total motility, progressive motility, and rapid motility) after 48 hours of cooled storage, along with a significant improvement in motion characteristics of sperm following freeze–thaw.18 No difference was noted in sperm motion characteristics in fresh semen following use of the nutraceutical. The authors concluded that feeding a DHA-enriched diet for 14 weeks would be most beneficial for those stallions with marginal fertility whose sperm do not tolerate cooling and freezing well. Lubricants used in the semen collection process have been shown to affect the progressive motility of
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Spermatozoal Motility sperm in an ejaculate. Froman et al.19 demonstrated a reduction in sperm motility over time when raw semen was exposed to 10% sterile K-Y, non-sterile K-Y, or Maxilube jellies prior to addition of a semen extender. Samper et al.20 examined the effect of different lubricants on longevity of motility of stallion spermatozoa. Aliquots of extended semen were exposed to four different commercially available lubricants, many of which were considered nonspermicidal, and stored in a passive cooling device. While no differences in progressive motility were noted prior to storage, significant differences were noted beginning after 24 hours of storage. Of the lubricants examined, only Pre-SeedTM Equine did not significantly lower motility in a semen sample when compared with controls. Many different semen extenders are commercially available in the equine breeding industry. Most are composed of several basic ingredients, including non-fat dried milk solids, an energy source, and antibiotics. In addition, when semen is frozen, a cryoprotectant is added to aid in sperm stabilization during the freezing and thawing processes. These ingredients, if not used in the correct manner or at appropriate concentrations, may negatively affect sperm motility. This is especially evident when freezing semen, when use of an inappropriate cryoprotectant type or concentration may reduce post-thaw motility and fertility. Other factors during semen collection and processing have been studied in an attempt to examine their effect on many parameters, including sperm motility. Sieme et al.21 showed that, as the time to ejaculation and the number of mounts increased, progressive sperm motility after 24 hours of storage at 5◦ C decreased. Evaluation of the addition of antibiotics to semen extenders was examined by Varner et al.,22 who found that the mean percentages of motile, progressively motile, and rapidly motile sperm were significantly lower in semen exposed to polymyxin B than any other antibiotic. Other researchers have demonstrated that the addition of gentamicin to extender results in decreased spermatozoal motility and velocity.23 Hormonal treatments can be used to manage sexual and aggressive behavior in stallions. The effect of these treatments on reproductive parameters, including sperm motility, has been evaluated. Squires et al.24 showed a significant decrease in the mean progressive motility in stallions treated with altrenogest for 240 days, but not in those treated for only 150 days. Progressive motility returned to pretreatment levels in those stallions treated for 240 days by the end of the study. Immunization against gonadotropinreleasing hormone (GnRH) has also been studied as a method to suppress reproductive behavior in
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the stallion. Burger et al.25 demonstrated a continuous decrease in sperm motility after GnRH vaccination, with the lowest mean motility values recorded 6 months after the first vaccination.
Motility as it relates to fertility Stallion sperm require many characteristics to achieve successful fertilization. Sperm motility should never be used at the exclusion of other parameters to assess stallion fertility or the fertilizing capacity of semen. Fertility of an individual stallion may be unsatisfactory in spite of satisfactory motility.4 Investigations into apparent stallion infertility should also include assessment of the mare population and breeding management and practices. In spite of this, motility is commonly used as an estimate of the viability of a given population of sperm.1 Voss et al.4 reported that motility may be the best predictor of fertility in the stallion.
References 1. Varner DD, Schumacher J, Blanchard TL, Johnson L. Breeding soundness examination. In: Varner DD, Schumacher J, Blanchard TL, Johnson L (eds) Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991; pp. 61–96. 2. Turner RM. Tales from the tail: what do we really know about sperm motility? J Androl 2003;24:790–803. 3. Amann RP, Graham JK. Spermatozoal function. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Media, PA: Williams & Wilkins, 1993; pp. 715–45. 4. Voss JL, Pickett BW, Squires EL. Stallion spermatozoal morphology and motility and their relationship with fertility. J Am Vet Med Assoc 1981;178:287–9. 5. Johnson L, Amann RP, Pickett BW. Scanning electron microscopy of the epithelium and spermatozoa in the equine excurrent duct system. Am J Vet Res 1978;39:1428–34. 6. Turner RM. Moving to the beat: a review of mammalian sperm motility regulation. Reprod Fertil Dev 2006;18: 25–38. 7. Turner RM, Eriksson RL, Gerton GL, Moss SB. Relationship between sperm motility and the processing and tyrosine phosphorylation of two human sperm fibrous sheath proteins, pro-hAKAP82 and h-AKAP82. Mol Hum Reprod 1999;5:816–24. 8. Pickett BW. Collection and evaluation of stallion semen for artificial insemination. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Media, PA: Williams & Wilkins, 1993; pp. 705–14. 9. Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL. Semen collection and artificial insemination. In: Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL (eds) Manual of Equine Reproduction, 2nd edn. St. Louis: Mosby, 2003; pp. 131–42. 10. Estrada AJ, Samper JC. Evaluation of raw semen. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders, 2006; pp. 253–7. 11. Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL. Examination of the stallion for
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breeding soundness. In: Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL (eds) Manual of Equine Reproduction, 2nd edn. St. Louis: Mosby, 2003; pp. 143–64. Card C. Cellular associations and the differential spermiogram: making sense of stallion spermatozoal morphology. Theriogenology 2005;64:558–67. Pickett BW, Anderson EW, Roberts AD, Voss JL. Freezability of first and second ejaculates of stallion semen. In: Proceedings of the 9th International Congress on Animal Reproduction and Artificial Insemination, Madrid, Spain, vol. 5, 1980, pp. 339–47. Magistrini M, Chanteloube P, Palmer E. Influence of season and frequency of ejaculation on production of stallion semen for freezing. J Reprod Fertil Suppl 1987;35:127–33. Friedman R, Scott M, Heath SE, Hughes JP, Daels PF, Tran TQ. The effects of increase testicular temperature on spermatogenesis in the stallion. J Reprod Fertil Suppl 1991;44:127–34. Blanchard TL, Jorgensen JB, Varner DD, Forrest DW, Evans JW. Clinical observations on changes in concentrations of hormones in plasma of two stallions with thermally-induced testicular degeneration. J Equine Vet Sci 1996;16:195–201. Turner RMO. Pathogenesis, diagnosis, and management of testicular degeneration in stallions. Clin Tech Equine Pract 2007;6:278–84. Brinsko SP, Varner DD, Love CC, Blanchard TL, Day BC, Wilson ME. Effect of feeding a DHA-enriched nutriceutical
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on the quality of fresh, cooled and frozen stallion semen. Theriogenology 2005;63:1519–27. Froman DP, Amann RP. Inhibition of motility of bovine, canine and equine spermatozoa by artificial vagina lubricants. Theriogenology 1983;20:357–61. Samper JC, Garcia A, Burnett K. The effect of different lubricants on longevity of motility and velocity of stallion spermatozoa. Theriogenology 2007;68:496. Sieme H, Katila T, Klug E. Effect of semen collection practices on sperm characteristics before and after storage and on fertility of stallions. Theriogenology 2004;61:769– 84. Varner DD, Scanlan CM, Thompson JA, Brumbaugh GW, Blanchard TL, Carlton CM, Johnson L. Bacteriology of preserved stallion semen and antibiotics in semen extenders. Theriogenology 1998;50:559–73. Aurich C, Spergser J. Influence of bacteria and gentamicin on cooled-stored stallion spermatozoa. Theriogenology 2007;67:912–18. Squires EL, Badzinski SL, Amann RP, McCue PM, Nett TM. Effects of altrenogest on total scrotal width, seminal characteristics, concentrations of LH and testosterone and sexual behavior of stallions. Theriogenology 1997;48:313– 28. Burger D, Janett F, Vidament M, Stump R, Fortier G, Imboden I, Thun R. Immunization against GnRH in adult stallions: effects on semen characteristics, behavior and shedding of equine arteritis virus. Anim Reprod Sci 2006;94:107–11.
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Chapter 126
Spermatozoal Morphology D.N. Rao Veeramachaneni
Evaluation of spermatozoal morphology is one of the most common laboratory practices in the assessment of potential fertility of males. A basic understanding of the process of spermatogenesis (see Chapters 101 and 102) and anatomical features of a spermatozoon (see Figures 102.2 and 102.5) is essential for interpretation of the morphological aberrations found during such an evaluation and to optimally utilize that information for clinical management. A variety of classification schemes and criteria have been devised to describe and characterize the structural defects of spermatozoa: (i) based on their presumptive role in accomplishing fertilization – some defects are considered to play a major role while others are of minor importance, and hence are called major and minor types of spermatozoal defects, respectively; and (ii) based on their origin either during spermatogenesis or during post-testicular sojourn in the accessory sex organs, and hence called primary or secondary spermatozoal defects, respectively. It is important to recognize that an individual spermatozoon may have multiple defects (Figure 126.1) and that it can be difficult to judge which one of these abnormalities is more significant and deleterious to fertilization as there are no definitive studies. The reader is referred to a variety of anthologies and illustrated treatises dealing with sperm morphological evaluations.1–7 Themain purpose of this chapter is to present a new approach which emphasizes the use of a semen sample as biopsy material and relates the morphological defects to the various phases of spermatogenesis, spermiation, and epididymal transit, and to the sequelae resulting from the interactions with the admixture of accessory gland secretions in disease conditions. Successful fertilization requires that a sufficient number of sperm reach the site of fertilization, bind to and penetrate the vestments of an oocyte, and ultimately form a zygote that develops into a blastula and fetus. These functional events are accomplished, respectively, by middle piece and principal piece;
plasma membrane and acrosome; and nucleus. A variety of biochemical tests are available to assess the functional status of these sperm organelles. These include a mitochondria-specific fluorescent probe, JC-l,8, 9 to evaluate mitochondrial potential; flow cytometry using fluorescent nucleic acid stains R -14 in combination with propidium such as SYBR 10 iodide to detect structural integrity of the plasma R membrane; an acrosomal probe LysoTracker Green DND-26 that identifies living spermatozoa with intact acrosomes;11 and the sperm chromatin strucR 12 ), sperm chromatin dispersion ture assay (SCSA assay (Halosperm R kit),13 and TdT-mediated-dUTP nick end-labeling (TUNEL) assay,14, 15 all of which detect fragmentation or breakage of DNA strands. Although these assays are ideal for evaluation of functional status of multiple spermatozoal components in certain clinical situations, these functional analyses must be performed individually on freshly ejaculated cool-stored or frozen–thawed sperm,9, 16 and can require sophisticated instrumentation. Thus, the constraints of time, labor, and costs involved may be prohibitive in routine veterinary practice. As an alternative tool, inasmuch as structure likely reflects function, the normalcy and structural integrity of these pivotal spermatozoal organelles are oftentimes assessed by more simple, cost-effective morphological techniques. These include employment of phase contrast or differential interference contrast microscopy using either unstained wet semen preparations, or bright-field microscopy following differential staining of dry semen smears (Figure 126.2). For example, a variety of structural defects of nuclear and flagellar components could be easily identified using phase contrast or differential interference contrast microscopy. Similarly, the structural integrity of the plasma membrane over the sperm head can be assayed by simple dye exclusion tests, e.g., eosin,17 using bright-field microscopy. However, the integrity of acrosomal membrane can
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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be assayed only by fluorescence microscopy using fluorochrome-labeled lectins, e.g., pea or peanut agglutinins, that bind to acrosomal contents.18 Thus, although light microscopy is an important and useful component of routine breeding soundness evaluations of healthy stallions, use of light microscopy alone limits the inherent utility of a semen sample in identifying disease conditions contributing to subfertility or infertility. In this chapter, a relatively simple morphological procedure that facilitates a routine differential analysis, as well as a critical evaluation of multiple organelles (when necessary), is described; the former using a light microscope and the latter using a transmission electron microscope. Furthermore, this procedure can be useful in the diagnosis of existing degenerative and inflammatory conditions in the reproductive tract as it enables identification and characterization of any exfoliated cells along with the germ cells in the ejaculate.
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Figure 126.1 An individual spermatozoon may manifest more than one defect resulting from impact on various phases of spermatogenesis. It is not always possible to ascribe these multiple defects to a particular phase and determine which anomaly is more detrimental. (a) Four-headed spermatozoon with fused tails resulting from impaired spermiogenesis with confluence of a cohort of four spermatids. (b) A spermatozoon with a nuclear crater and defective middle piece enclosed in residual cytoplasm resulting from impaired spermiogenesis and spermiation.
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Processing of a semen sample The scope of this chapter precludes detailed description of the procedures involved in processing equine ejaculates for light and transmission electron microscopy; they can be found in publications from our laboratory.19–21 In brief, 5 mL of gel-free semen is fixed in 10 mL cacodylate-buffered 6% glutaraldehyde. The fixed sample is centrifuged, the
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Figure 126.2 Light microscopic techniques commonly used in spermatozoal morphology. The ability to identify subtle defects and the final outcome of an evaluation largely depends on the quality (resolution, numerical aperture) of the objective lens used. (a) Phase contrast microscopy. This technique is commonly used evaluate acrosomal abnormalities in wet smears; a knobbed acrosome (arrowheads) and an acrosome shared between two nuclei (arrow) resulting in fused heads are seen. (b) Differential interference contrast (Nomarski) microscopy. This technique, which provides a good contrast to evaluate cell surface, is used to assess integrity of acrosomes, nuclei, and flagellar components in wet smears. An acrosomal–nuclear defect (arrow) and coiled middle piece entrapping residual cytoplasm (arrowhead) are seen. (c) Bright-field microscopy. This technique is used to evaluate stained smears. With stains such as eosin (and nigrosin for background contrast) the plasma membrane integrity can be evaluated; intact plasma membrane is impermeable to eosin and hence sperm heads do not stain (white sperm head; bottom right).
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Spermatozoal Morphology supernatant is poured off, and the pellet is suspended in 0.1 M sodium cacodylate buffer. A 100-µL aliquot of the suspension is added to 1 mL of distilled water in a glass vial. A wet smear is freshly prepared by placing a drop of this suspension on a glass slide and placing a coverslip for phase contrast or differential interference contrast (Nomarski) microscopy. Dry smears are also prepared by placing a drop of the suspension on a glass slide and distributing the fluid evenly with a smooth glass rod or the tip of a long Pasteur pipette. The slide is placed on a heating block at 80◦ C, stained with toluidine blue, and coverslips mounted using a permanent mounting medium for use with bright-field microscopy. The remaining suspension is washed once more as described previously, the supernatant poured off, and the pellet processed for transmission electron microscopy. The pellet is post-fixed in a solution of 1% osmium tetroxide in 0.1 M cacodylate buffer and centrifuged. Osmicated pellets are dehydrated through a graded series of ethanol, rinsed in propylene oxide, and embedded in Poly/Bed 812 (Polysciences Inc., Warrington, PA). From these plastic tissue blocks, 1µm-thick sections are cut, placed on glass slides, and stained with toluidine blue for characterization of exfoliated cells using a light microscope. Then, ∼80-nm thin sections are cut, mounted on 300-mesh nickel grids, and stained with uranyl acetate and lead citrate for ultrastructural evaluation of spermatozoal organelles and other cellular constituents.
Light microscopy A light microscope to be used is equipped with planapochromatic as well as phase contrast objective lenses, together with a universal condenser that enables bright-field, phase contrast, and differential interference contrast microscopy. Utilization of toluidine blue-stained smears (bright-field microscopy), as well as unstained wet smears (differential interference contrast and phase contrast microscopy), complement each other by compensating for deficits inherent to these techniques (e.g., artifacts associated with smear preparation, resolution of subtle defects) and thus enhance critical evaluation. In our laboratory, the microscope is interfaced with a computerized imaging system (ImagePro, version 5.0) and a data collection spreadsheet (Microsoft Excel). One hundred spermatozoa from each of the wet and toluidine blue-stained smears are classified into normal or abnormal categories; although the precision of repeated estimates varies with the skill of the technician, little precision is gained by counting more than 100 spermatozoa when an experienced technician performs evaluation.22 The abnormalities are categorized as individual defective components –
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acrosome, nucleus, middle piece, principal piece, terminal piece, entire flagellum, retained cytoplasmic droplets at a proximal (neck) or distal (annulus) position, or diffuse residual cytoplasm. Tailless (detached) heads, multiple heads and/or tails, and miscellaneous forms, such as unique ‘Dag’ and ‘Diadem’ defects,1–3 are also enumerated. Examples of these abnormalities are shown in Figure 126.3. As noted earlier, because an individual spermatozoon tends to have multiple defects when various phases of spermatogenesis are affected simultaneously, every single defect associated with any of these structural components is individually recorded in a spreadsheet designed for this purpose (Figure 126.4). Formulae embedded in the spreadsheet compute the percentage of normal spermatozoa, spermatozoa with a single defect, and spermatozoa with multiple defects. Thus, the total number of sperm abnormalities could exceed 100% but the total of normal plus abnormal spermatozoa always equals 100%. This is in contrast to the routine practice of assigning each spermatozoon into only one category even when there are multiple defects. This method of assigning abnormalities to individual structural components of sperm and assimilation of their collective incidence facilitates identification of the origin of the defect(s) and assessment of the severity of damage, thereby aiding the examiner in pinpointing the phase in spermatogenesis and/or post-testicular sperm transit that is affected. In addition to spermatozoal defects, the presence of any prematurely exfoliated germ cells, teratoids, and other discernible somatic cells is recorded. When such denuded cells are noted in smears, 1-µm-thick sections of sperm pellets stained with toluidine blue are examined for definitive identification and characterization of various cell types, e.g., epithelial cells from excurrent ducts and accessory sex glands, urothelium, bacteria, and cells of hematological origin.
Transmission electron microscopy It is not uncommon to find subtle lesions of the plasma membrane, acrosome, nuclear chromatin, mitochondria, or axoneme in the so-called ‘idiopathic’ cases of infertility. Using a transmission electron microscope, thin sections stained with uranyl acetate and lead citrate can be evaluated to discern subtle lesions in these spermatozoal organelles. Although quantification of morphological defects is not possible, transmission electron microscopy can augment light microscopic findings by characterizing ultrastructural lesions such as abnormal plasma membranes, prematurely reacted acrosomes, unusual inclusions in the acrosomes, and cytoplasmic
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Figure 126.3 Differential interference contrast micrographs of normal and abnormal equine spermatozoa. Note that several spermatozoa shown have multiple defects but only some are highlighted to present a thematic overview of what could go wrong with individual components of sperm. (a) Frontal (left) and sagittal (right) planes of view of normal spermatozoa. The region of tail (flagellum) between the arrows is the middle piece. (b) Abnormal plasma membranes. Note the swollen and thickened appearance of the plasma membranes. Presence of inclusions and aberrant microtubule masses can be revealed by transmission electron microscopy as seen in Figures 126.5c and 126.6c. (c–h) Acrosomal defects. (c) Knobbed sperm showing an acrosomal cyst or vesicle. (d) Acrosomal invasion of nucleus resulting in acrosomal–nuclear dysplasia. (e) Acrosome separated from the nucleus. (f) Acrosomereacted sperm head. Normally acrosome reaction takes place in the vicinity of the egg. If the normal milieu of the seminal fluid is altered because of a possible infection or inflammation of the accessory sex organs, acrosomal reaction occurs prematurely. (g and h) Shared acrosomes. Acrosome shared between two sperm nuclei resulting in fused sperm (g). This developmental anomaly could be experimentally induced by exposure to environmental pollutants and the effect often lasts long after cessation of the exposure (for illustrated discussion of the morphogenesis of this lesion, see Veeramachaneni et al.20 ). Another form of acrosome shared between two sperm is shown in h; in this case, the fusion is only at the tip. (i) Two-headed sperm. Multiple headed sperm results very likely from failure of disintegration of the intercellular bridges between cohorts or fusion of the tails with common plasma membrane or mitochondrial sheath during spermiogenesis. This developmental anomaly is different from the shared acrosomes shown in (g) and (h). (j, k) Retained cytoplasmic droplets. As tubulobulbar complexes formed by spermatid and Sertoli cells disintegrate during spermiation, releasing sperm into the lumen of seminiferous tubules, a small remnant of spermatid cytoplasm remains attached to the neck. This cytoplasmic droplet translocates from a proximal to a distal position (relative to the neck or middle piece) during epididymal transit and is completely shed by the time of ejaculation. The presence of a retained droplet at a proximal position in ejaculated sperm indicates impairment of epididymal function, although involvement of impaired testicular function itself cannot be ruled out;5 (j) proximal cytoplasmic droplet; (k) distal cytoplasmic droplet. (l) Residual cytoplasm and pseudodroplets. Typically varying in size and more irregular than cytoplasmic droplets, residual cytoplasm may be associated with any region of the sperm. When residual cytoplasm is located in the region of middle piece (neck region of m) it may give the appearance of a pseudodroplet, the term commonly used to describe thick or swollen mitochondria (distal end of middle piece in the top profile of (p)). The presence of residual cytoplasm is an indication of impaired spermiogenesis and spermiation and may contain aberrant structures such as microtubule masses (see Figure 126.6c).
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Spermatozoal Morphology
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Figure 126.3 (Continued) (m–r) Middle piece defects. Mitochondria surrounding the outer dense fibers in the middle piece may be absent (aplasia; m) or swollen and irregular (n). Absence of mitochondrial sheath may be segmental or extend the entire length of the middle piece. Compare the thickness of the region between the arrows in (m) with that of (a). Mitochondrial aplasia is also evident in (f). Swollen or rough mitochondria may give rise to the appearance of a screw and hence the name corkscrew defect. A structurally deficient mitochondrial sheath may impair structural integrity of the axoneme and result in a variety of middle piece defects. Breaks at the neck region may result in middle pieces reflecting over the heads fusing with them (o) or rolled over (p), whereas breaks at the distal end may result in reflected middle pieces, sometimes called bent or hairpin tails (q). Occasionally, more than one middle piece is present, resulting in multiple tails; two middle pieces are seen in each of two sperm in (r). (s–v) Principal piece defects. Defective principal pieces may also accompany defective middle pieces. These abnormalities resulting from defective/missing axonemal components take a variety of forms – tight folding (s) or coiling (t) of proximal portions of the principal piece over the middle piece. In some only the terminal piece is coiled (u), while in others the entire tail is coiled (v; top profile). These coils may or may not entrap cytoplasmic droplets or residual cytoplasm. The bending of the middle piece together with coiling of the entire tail, as in (s and t), is commonly termed a Dag defect, after a Jersey bull named Dag in which it was first described. Some defects that look like sperm with tails coiled around the heads may actually be developmentally malformed germ cells called teratoids (v; bottom profile). (w) Detached or tailless head defect. An occasional detached head is not uncommon in any ejaculate but a high incidence is generally a sequel of ampullary spermiostasis or testicular degeneration and epididymal dysfunction. Detached heads (top profile) should be distinguished from tail stump defects (bottom profile) as the latter has been shown to be genetic in bulls. A tail stump sperm defect results from inhibition of axonemal formation as a consequence of an anomalous distal centriole. Note an implantation fossa is clearly seen in the detached head but not in the one with the tail stump. (x) Nuclear defects. There is some variation in the normal shape and size of sperm heads in any species; stallions are no exception. One of the common head shape defects is pyriform or pear-shaped nuclei. Note narrowness at the base of the heads in (m), (u), and bottom profile of (w) (compare with those of (a)). In addition to the defective shapes, vacuolations, and craters or pouches (invaginations of nuclear membrane into nuclear matrix) also occur in sperm nuclei. Several nuclear pouches are clearly seen as indentations in the top profile of (x). The occurrence of these pouches in an orderly fashion as in a string of beads at the post-acrosomal region in the equatorial segment of the nucleus (bottom profile) has been described in bulls as a diadem defect (as they look like a necklace). It may be difficult to discern defects such as these in stained smears using bright-field microscopy. Transmission electron microscopy reveals that nuclear malformations are often associated with dysplastic acrosomes and impaired chromatin condensation (see Figures 126.5c and 126.7b).
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A. Individual sperm scores Sperm no.
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PP
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PD
DD
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B. Summary %
Normal
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MHT
MISC
Wet smear Dry smear Average
ACR NCL FLG MP PP TP PD DD RC TLH MHT MISC
acrosome: missing, ruffled, knobbed/cystic; acrosomal dysgenesis: dysplasia, dysgenesis – shared; acrosome-reacted nucleus: pyriform, variations in size and shape, malformed – craters, apoptotic flagellum: entire tail – coiled, coiled around head, thin middle piece: abaxial, reflected, swollen, thin principal piece: bent, swollen, thin terminal piece: flayed, looped proximal droplet distal droplet residual cytoplasm (other than PD or DD) tailless head (detached head) multiple heads and/or tails miscellaneous: unique defects, e.g., ‘Dag’, ‘Diadem’, ‘tail stump’ etc.
Single Multiple
sperm having only one defect sperm having multiple defects
Record the presence of: Premature germ cells: spermatids, spermatocytes; teratoids Sertoli cell remnants, cell debris, bacteria, macrophages, erythrocytes, leukocytes Epithelial cells: originating from excurrent ducts, accessory glands, or urinary tract Figure 126.4 Data collection sheet for spermatozoal morphology. Data are entered into a computerized spreadsheet that computes the percentage of normal sperm, abnormal sperm with a single defect, and abnormal sperm with multiple defects. In addition, since all defects associated with individual structural components of a spermatozoon are recorded, an overview of the spreadsheet will provide valuable information pertaining to the predominant defect(s) and possible target/origin of the defect.
droplets (Figures 126.5 and 126.6). For example, the retained cytoplasmic droplets characterized at light microscopic level may not be simply cytoplasm, but may contain other structures such as microtubular masses (Figure 126.6c); the microtubular mass defect may involve proliferation of abnormal microtubules from sites associated with the manchette, a microtubular organelle important in shaping the spermatozoon. In disease conditions, it is not uncommon to find cells and cellular debris in ejaculates other than spermatozoa (Figures 126.7–126.10). Use of transmission electron microscopy facilitates identification of these constituents in pathological ejaculates, e.g., incomplete condensation of chromatin and its break down (Figure 126.7); premature germ cells (Figure 126.8); leukocytes, macrophages, and denuded cells from accessory sex organs (Figure 126.9); and even bacteria including rare pathogens such as spirochetes (Figure 126.10). Thus, electron
microscopy may help to identify etiological factors and the pathogenesis of underlying conditions.
Semen sample as biopsy material Testicular degeneration, inflammation, and subsequent repair processes in seminiferous epithelium impair spermiogenesis and spermiation, resulting in lesions as seen in Figures 126.8–126.10. In addition, when any of the components that contribute to seminal secretions – the testes, excurrent ducts, and accessory sex glands – are affected, the composition of the seminal mixture will be altered by the presence of cells and cellular debris of the affected organ. The presence of prematurely acrosome-reacted spermatozoa, as seen in Figure 126.5b, may indicate altered milieu in seminal fluid, which can result from an inflammatory process in the reproductive tract (Veeramachaneni D.N.R., unpublished observations).
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Figure 126.5 Transmission electron micrographs of sagittal sections of equine spermatozoal heads. (a) Normal sperm head. Note condensed chromatin (black) enveloped by the acrosome (gray) and plasma membrane (outermost layer). (b) Acrosome-reacted sperm heads. Left, note the acrosomic vesicles formed by the fusion of inner and outer acrosomal membranes. Right, note the acrosome completely reacted releasing its contents. Normally, acrosome reaction occurs in the female reproductive tract upon oocyte binding. Premature acrosomal reaction (because of altered milieu of seminal plasma) in the ejaculum itself renders the sperm nonfunctional. (c) Acrosomal dysgenesis. Note the dysplastic acrosome with cystic malformations. Also note inclusions between the acrosomal and plasma membranes. Such inclusions are barely discernible at a light microscopic level (compare with Figures 126.2b and 126.3b–d).
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Figure 126.6 All retained droplets observed at light microscopic level may not be the cytoplasmic droplets that normally translocate and shed during epididymal transit and ejaculation. (a) Differential interference contrast micrograph of proximal and distal cytoplasmic droplets. (b) Transmission electron micrograph of a proximal droplet containing cytoplasmic vesicles, membranous profiles, and vacuoles. (c) Transmission electron micrograph of a whorl of aberrant microtubule mass enclosed in plasma membrane. In contrast to the cytoplasmic droplet in (b), this structure involves proliferation of abnormal microtubules possibly originating from sites associated with the manchette, a microtubular organelle important in shaping the spermatozoon. Once found, this defect continues to manifest, suggesting that it is a developmental defect and may be associated with subfertility in stallions.
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Figure 126.7 Incomplete condensation and breakdown of nuclear chromatin. A variety of environmental factors can cause impaired chromatin condensation and breakage. (a) Photomicrograph of toluidine blue-stained smear. Note different intensities of nuclear staining among spermatozoa and darkly stained aggregates of particulate material. Without careful scrutiny, these aggregates are commonly dismissed as extraneous debris. (b) Transmission electron micrograph of four incompletely condensed sperm nuclei and a normally condensed sperm nucleus (lower right). Note ruptured plasma membranes surrounding the four nuclei; these nuclei are prone to breakage resulting in clumps of chromatin fragments (lower left). (See color plate 43).
(a)
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Figure 126.8 Seminal ejaculates contain cells and cellular debris other than spermatozoa. Identification of these cells is important in devising treatment strategies. (a) Differential interference contrast micrograph of a wet smear. Note the presence of a large proportion of exfoliated cells, some of which appear to be premature germ cells. Examination of wet smears alone cannot fully characterize these cells and therefore further histological processing is advocated. (b) Toluidine-blue stained thick section of a plastic-embedded cell pellet. Degenerate round spermatids (acrosomal vesicle is seen in profiles indicated by arrows) and epithelial fragments could be identified. The presence of these denuded cells in the seminal ejaculate reflects a variety of degenerative processes in the testis and excurrent ducts.
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Figure 126.9 Characterization of cellular debris provides useful information on the severity of disease. Whereas the components of the cellular debris could not be definitively identified using phase contrast microscopy (a), upon examination of a toluidine-stained thick section of the cell pellet of the same sample, they could be identified as neutrophils, denuded epithelial cells, and concretions from the reproductive tract. Further examination of thin sections using electron microscopy confirmed that they were epithelial cells lining the excurrent ducts (c) and macrophages (d). Note the presence of phagocytosed spermatozoal organelles. (See color plate 44).
Seminiferous epithelium is sensitive to a wide variety of environmental factors resulting in deleterious effects on germ cell differentiation that may perpetuate long after cessation of exposure to culprit toxicants.23 Acute insults are manifested as a spectrum of degenerative lesions, ranging from prematurely shed spermatids to exfoliation of aggregates of germinal epithelium, as seen in Figure 126.7. For example, incomplete condensation of chromatin (Figure 126.6b) or apoptosis of spermatozoal nuclei resulting from a variety of ailments may lead to breakage of chromatin resulting in flocculent aggregates that are not discernible by light microscopy (Figure 126.6a) and are likely to be ignored as debris.20 A variety of etiological factors, such as exposure to environmental toxicants and oxidative stress, are known to cause sperm DNA fragmentation and thus discernment of this cellular debris can help address the etiological factor(s). Ability to identify what is otherwise dubbed as ‘debris’ using lower resolution optics often provides information which might have valuable diagnostic and prognostic potential and thus renders a semen sample as useful biopsy material.
Summary The processing of a semen sample as described here facilitates routine light microscopic evaluation of spermatozoal morphology and also transmission electron microscopy, when needed, permitting critical evaluation of spermatozoal organelles as well as any exfoliated cells and pathogens in the ejaculate. Thus, by using semen as biopsy material, this technique can potentially circumvent the need for testicular biopsy or a battery of biochemical tests and serves as a valuable clinical tool. Compared with relatively expensive multiple functional assays used in andrology clinics, which assess one aspect of spermatozoal function at a time, this technique facilitates simultaneous evaluation of several attributes of an ejaculate.
Acknowledgement Technical assistance of Carol Moeller and Jennifer Palmer is gratefully acknowledged.
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Figure 126.10 In some instances, pathogens can also be identified. Bacteria can be seen embedded in flocculent cellular material in wet smears (a) as well as in stained dry smears (b). Electron microscopy reveals bacilli (c) and spirochetes (d). (See color plate 45).
References 1. Barth AD, Oko RJ. Abnormal Morphology of Bovine Spermatozoa. Ames: Iowa State University Press, 1989. 2. Blom E. Pathological conditions in genital organs and sperm as a cause for the rejection of breeding bulls for import into and export from Denmark (an andrologic retrospective, 1958–1982). Nord Vet Med 1983;35: 105–30. 3. Brito LFC. Evaluation of stallion sperm morphology. Clin Tech Equine Pract 2007;6:249–64. 4. Holstein AF, Roosen-Runge EC, Schirren C. Illustrated Pathology of Human Spermatogenesis. Berlin: Grosse Verlag, 1988. 5. Rao AR, Bane A, Gustafsson BK. Changes in the morphology of spermatozoa during their passage through the genital tract in dairy bulls with normal and impaired spermatogenesis. Theriogenology 1980;14:1–9. 6. Seed J, Chapin RE, Clegg ED, Dostal LA, Foote RH, Hurtt ME, Klinefelter GR, Makris SL, Perreault SD, Schrader S, Seyler D, Sprando R, Treinen KA, Veeramachaneni DNR, Wise LD. Methods for assessing sperm motility, morphology, and counts in the rat, rabbit, and dog: a consensus report. ILSI Risk Science Institute Expert Working Group on Sperm Evaluation. Reprod Toxicol 1996;10: 237–44. 7. Zamboni L. The ultrastructural pathology of the spermatozoon as a cause of infertility: the role of electron microscopy in the evaluation of semen quality. Fertil Steril 1987;48:711–34.
8. Gravance CG, Garner DL, Baumber J, Ball BA. Assessment of equine sperm mitochondrial function using JC-1. Theriogenology 2000;53:1691–1703. 9. Thomas CA, Garner DL, DeJarnette JM, Marshall CE. Effect of cryopreservation of bovine sperm organelle function and viability as determined by flow cytometry. Biol Reprod 1998;58:786–93. 10. Garner DL, Johnson LA, Yue ST, Roth BL, Haugland RP. Dual DNA staining assessment of bovine sperm viability using SYBR-14 and propidium iodide. J Androl 1994;15:620–9. 11. Thomas CA, Garner DL, DeJarnette JM, Marshall CE. Fluorometric assessments of acrosomal integrity and viability in cryopreserved bovine spermatozoa. Biol Reprod 1997;56:991–8. 12. Evenson DP, Jost LK, Marshall D, Zinaman MJ, Clegg E, Purvis K, de Angelis P, Claussen OP. Utility of the sperm chromatin structure assay as a diagnostic and prognostic tool in the human fertility clinic. Hum Reprod 1999;14:1039–49. 13. Fernandez JL, Muriel L, Goyanes V, Segrelles E, Gosalvez J, Enciso M, LaFromboise M, De Jonge C. Simple determination of human sperm DNA fragmentation with an improved sperm chromatin dispersion test. Fertil Steril 2005;84:833–42. 14. Heninger NL, Staub C, Blanchard TL, Johnson L, Varner DD, Forrest DW. Germ cell apoptosis in the testes of normal stallions. Theriogenology 2004;62:283–97. 15. Morte MI, Rodrigues AM, Soares D, Rodrigues AS, Gamboa S, Ramalho-Santos J. The quantification of lipid and
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17.
18.
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protein oxidation in stallion spermatozoa and seminal plasma: seasonal distinctions and correlations with DNA strand breaks, classical seminal parameters and stallion fertility. Anim Reprod Sci 2008;106:36–47. Kirk ES, Squires EL, Graham JK. Comparison of in vitro laboratory analyses with the fertility of cryopreserved stallion spermatozoa. Theriogenology 2005;64:1422–39. Dott HM, Foster GC. A technique for studying the morphology of mammalian spermatozoa which are eosinophilic in a differential ‘live-dead’ stain. J Reprod Fertil 1972;29:443–5. Farlin ME, Jasko DJ, Graham JK, Squires EL. Assessment of Pisum sativum agglutinin in identifying acrosomal damage in stallion spermatozoa. Mol Reprod Dev 1992;32: 23–7. Veeramachaneni DNR, Moeller CL, Pickett BW, Shiner KA, Sawyer HR. On processing and evaluation of equine
20.
21.
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seminal samples for cytopathology and fertility assessment: the utility of electron microscopy. J Equine Vet Sci 1993;13:207–15. Veeramachaneni DNR, Moeller CL, Sawyer HR. Sperm morphology in stallions: ultrastructure as a functional and diagnostic tool. Vet Clin North Am Equine Pract 2006;22:683–92. Veeramachaneni DNR, Sawyer HR. Use of semen as biopsy material for assessment of health status of the stallion reproductive tract. Vet Clin North Am Equine Pract 1996;12:101–10. Salisbury GW, Mercier E. The reliability of estimates of the proportion of morphologically abnormal spermatozoa in bull semen. J Anim Sci 1945;4:174–8. Veeramachaneni DNR. Impact of environmental pollutants on the male: effects on germ cell differentiation. Anim Reprod Sci 2008;105:144–57.
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Chapter 127
Principles of Cooled Semen James K. Graham
Introduction The use of cooled transported equine semen has become very popular in the industry as breed registries have accepted its use. Of course, the single major reason semen is cooled is to extend the life expectancy of the spermatozoa so that it can be transported, permitting insemination of mares located distant from the stallion; however, there are many other advantages as well as disadvantages to using cooled semen. Cooling cells exerts stresses on the spermatozoa that can be deleterious, and precautions must be taken to minimize these effects. In this chapter, the general principles involved in cooling stallion semen will be explained, including the advantages and disadvantages of its use, the biophysical responses stallion spermatozoa have to the cooling process itself, and how to ameliorate the deleterious aspects as much as possible. Discussion regarding specific cooling diluents (extenders) and the use and fertility of cooled stallion spermatozoa are topics addressed in other sections of this text.
Advantages and disadvantages of cooled semen Many of the advantages and disadvantages for using cooled equine semen have been addressed previously.1 In summary, these include:
1. lower cost to transport the semen than the mare 2. less stress on the mare to be inseminated 3. no additional boarding costs for mares to be inseminated 4. permits effective antimicrobial treatment of semen that may contain pathogenic organisms, thereby decreasing transmittance of diseases 5. increases desired genetic traits by using semen from only genetically superior stallions
6. increases the number of mares that can be bred to any particular stallion, including splitting an ejaculate to inseminate multiple mares 7. permits semen analysis prior to shipment While most of these advantages primarily affect the mare owners, stallion owners are also benefitted by being able to increase the number of mares booked to a particular stallion. With these advantages come some disadvantages as well. These include: 1. lower pregnancy rates from some stallions 2. additional expertise is required in the processes of semen collection, evaluation, and artificial insemination 3. additional costs are incurred for cooling diluents (extenders), and equipment to cool and transport the semen, as well as for managing and inseminating the mares 4. some stallions will produce semen that, for some reason, will not survive cooling; these stallion owners will not be able to participate in this technology. Most of these disadvantages can be minimized by careful management of the stallion, the semen, or the mares to be inseminated; therefore, the advantages to using cooled semen outweigh the disadvantages for most stallions and mares in most situations.
Principles of cooling stallion semen In order to understand how stallion semen should be cooled, including understanding why certain cooling diluents are used, what components within those diluents are important for the cooling process, and the exact mechanisms used to cool, store, warm, and utilize stallion spermatozoa, we need to know what happens to cells as they are cooled. In addition,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Extracellular
R P C C C
Figure 127.1 Structure of a phospholipid showing the threecarbon backbone, the phospho-head group (R), and the fatty acyl chains.
we need to understand how changes to cellular metabolism, membrane transport, and, very importantly, what membrane changes occur can ultimately affect cell survival.
Cytosolic (a) Extracellular
Z X
Cooling effects on cell membranes In 1972, the fluid mosaic model was presented as the structure of cell membranes.2 This model describes cell membranes as a dynamic bilayer of polar phospholipids, each of which is composed of a threecarbon backbone to which a phospho-head group is attached to an end carbon of the backbone and two fatty acyl chains are attached to the other carbons (Figure 127.1). The phospholipids in this bilayer are oriented with their hydrophilic head groups to the outside of the membrane (both extracellular and cytosolic compartments) and their hydrophobic fatty acyl chains to the membrane interior (Figure 127.2a). The phospholipid portion of the membrane is composed of many different phospholipid species, which are different from one another both in their phospo-containing head groups and in their fatty acyl chains. Although the most common lipid phospho-head group contains choline (phosphatidylcholine), head groups containing serine, ethanolamine, and inositol are also present in stallion spermatozoal membranes.3 These head groups can be glycosylated, have proteins, or have glycosylated proteins attached to them,4, 5 increasing the complexity of the membrane (Figure 127.2c). An important point regarding the phospholipids is that, while phosphatidylcholine is rather equally represented in both the inner and outer leaflet of the membrane, other phospholipids, such as phosphatidylserine, are located primarily in the inner leaflet. Flipping to the outer leaflet occurs only during membrane damage, such as cold shock or severe cooling damage, or during stages of a spermatozoon’s life when membrane changes must occur, such as capacitation and the acrosome reaction.6–13 Normally, however, phospholipid flipping from the inner leaflet to the outer or vice versa does not readily occur.
Y
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Cytosolic (c) Figure 127.2 General structure of a membrane showing (a) the phospholipid bilayer, (b) the lipid bilayer with integral (X) and peripheral (Y, Z) proteins, and (c) the bilayer with glycosylated lipids (∗) and proteins (P,R).
In addition to having different head groups, the fatty acyl chains constituting the phospholipids can be very different as well. The fatty acids contained in spermatozoal membranes range from 14 to 22 carbons in length and contain 0–6 unsaturated bonds.3 One of the important aspects of the fatty acyl chains, with regards to cooling spermatozoa, is that the fatty acyl chains will be a major determinant of whether
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the membrane is in a fluid or gel phase at any particular temperature.10 Another important component of the membrane are proteins, which constitute roughly 50% of membrane weight14 and which intermingle with the lipids (Figure 127.2b). Proteins constituting the membrane are considered to be either integral (essential for membrane structure and function) or peripheral (loosely attached to the membrane surface and can be removed). Integral proteins can serve as channels through the membrane, provide anchoring for the cytoskeleton, act as receptors for external molecules, and serve as second messenger signaling systems between the bilayers.14 Many of the proteins that extend onto the external surface of the membrane contain carbohydrate moieties that increase the complexity of protein receptors, provide the cell surface glycocalyx with a net negative charge, and help to absorb other proteins from the seminal plasma or surrounding medium to the membrane surface (Figure 127.2c).15 The last component of the membrane that is important for this discussion is cholesterol. Cholesterol is composed of four carbon rings and a carbon sidechain. The carbon rings are lipophilic, causing the majority of the molecule to intercalate into the core of the membrane associating with the fatty acyl chains, with only the molecule’s side-chain associating with the head groups of the phospholipid. Cholesterol is able to fill in any gaps created in the core of the membrane owing to adjacent phospholipids having fatty acyl chains of different carbon lengths or different degrees of unsaturation. Cholesterol, therefore, intercalates into the fatty acyl portion of the membrane, and helps to stabilize the membrane at body temperature. During spermatozoal capacitation, proteins in the female reproductive tract remove cholesterol from the plasma membrane, rendering the membrane less stable and more capable of undergoing fusion with the outer acrosomal membrane, causing the acrosome reaction.16 The ability of the phospholipids to move laterally throughout the membrane provides for its dynamic nature, which is important in overall membrane function. As the lipids and proteins move throughout the membrane they can interact with other lipids and proteins, and it is these interactions that permit ion channels, receptor signaling, and protein transporters to function normally. When the cell is cooled, the lipids undergo a phase transition from the fluid state into a gel, or solid state. Each fatty acyl chain undergoes this phase transition at a different temperature. Longer fatty acyl chains transition into the solid state at higher temperatures whereas shorter fatty acyl chains remain fluid to lower temperatures before they undergo phase transition. The number of unsaturated bonds in the fatty
acyl chain also affects the temperature at which the fatty acid undergoes the phase transition, and fatty acids with more unsaturated bonds exhibit lower transition temperatures than fatty acids of similar carbon number but with fewer unsaturated bonds. Synthetic membranes composed of a single phospholipid undergo the phase transition at distinct temperatures. For example, a membrane composed of phosphatidylcholine containing two palmitic (16-carbon) fatty acyl chains undergoes this phase transition at 42◦ C, while one composed of phosphatidylcholine containing two stearic (18-carbon) fatty acyl chains undergoes the phase transition at 54◦ C.17, 18 When the fatty acyl chains contain a single unsaturated bond, the transition temperature of the lipid is lower; the phase transition temperature of a membrane composed of phosphatidylcholine with 18-carbon fatty acyl chains decreases from 54◦ C when the fatty acids contain no unsaturated bonds to 11◦ C when the fatty acids contain one unsaturated bond.18 Natural membranes are composed of many different lipid species with different head groups and fatty acyl chains of different lengths and degrees of unsaturation. Therefore, natural membranes do not undergo the phase transition at a single temperature, but over a range of temperatures. In the case of stallion spermatozoa, this range is reported to be between 19◦ C and 8◦ C.19 When a membrane composed of many lipid species is cooled to the point that it reaches the highest temperature of the range at which the membrane undergoes the phase transition (19◦ C in the case of stallion spermatozoa), the lipids with the highest transition temperature will undergo the phase transition and will start to cluster as a solid lipid ‘iceberg’ in a sea of fluid lipids.15 Many of the lipids with the highest transition temperature associate with the membrane proteins and anchor the proteins in the membrane. As these lipids undergo the phase transition and coalesce into these ‘icebergs’, they leave their protein partners, which in turn coalesce to form protein clumps within the membrane, which significantly alters their functional capacities. As the temperature is reduced further, more and more lipid species undergo the phase transition and coalesce in these ‘icebergs’ of solid lipid, leaving a constantly smaller portion of fluid membrane. Finally, upon reaching the lowest temperature of the membrane range of transition, the entire membrane is in the ‘gel’ state. However, owing to significant rearrangements of both lipids and proteins from their preferred partners during the cooling process, membrane function may be altered not only in the gel state but also after warming, if normal lipid–lipid and lipid–protein associations cannot be reestablished.15 Physically, when a membrane is cooled, the rotational and vibrational movements of the fatty acyl chains of the membrane lipids slow, and, as the phase
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With cholesterol Fluid Membrane 37°C
Gel membrane 5°C
Fluid Membrane 5°C
Figure 127.3 Structure of membranes in the fluid state (at 37◦ C) and in the gel state (5◦ C), showing the effects that cholesterol has on lipid interactions.
transition occurs, van der Waals forces between the fatty acyl chains pull the fatty acyl chains of surrounding lipids (and entire lipid molecules) closer together (Figure 127.3). Therefore, in the gel state, the membrane is less fluid, and also possesses less surface area. Finally, cholesterol affects all of this. Although cholesterol intercalates into the fatty acyl portion of the membrane and helps stabilize the membrane at body temperature, at temperatures below the membrane’s phase transition cholesterol reduces the cohesive forces between adjacent fatty acyl chains and forces adjoining phospholipids apart; both of these processes make the membrane more fluid at these temperatures. In fact, when cholesterol is added to synthetic membranes, the phase transition temperature of the membrane is reduced; if sufficient cholesterol is included in the membrane, the membranes never undergo a phase transition at any temperature tested.20 Therefore, although cholesterol decreases membrane fluidity at high temperatures, it increases membrane fluidity at low temperatures, decreases the temperature at which the membrane undergoes a phase transition, and can even eliminate a phase transition altogether, which would permit more normal membrane composition (lipid–lipid and lipid–protein interactions) to be maintained to a higher degree (Figure 127.3). Cold shock and cooling damage Cellular injury, due to cooling, can be induced directly by altering or rupturing cellular structures, or indirectly by altering cell function.21 Direct damage to the plasma membrane induced by rapidly cooling stallion spermatozoa from 20◦ C to 5◦ C induces partially irreversible damage to the cell membrane, which is characterized by a loss of membrane lipids to the extracellular medium and a subsequent loss of
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intracellular ions and molecules. This causes a reduction in cellular metabolism and, ultimately, results in abnormal circular swimming patterns and rapid loss of motility.21 This significant damage is collectively referred to as ‘cold shock’ and results in rapid loss of spermatozoal viability.22 Much of the membrane damage is a direct result of the membrane undergoing an abrupt phase transition, resulting in significant membrane lipid and protein rearrangements, which ultimately result in aberrant membrane structure and function at 5◦ C or when the cells are warmed up to 38◦ C.15 Indirect cooling damage is more difficult to detect and often presents itself as early cell death after the cells have been rewarmed. Cooling damage likely results from the lipid and protein rearrangements that occur in the membrane as it undergoes the phase transition. Upon rewarming, although many of the lipid–lipid and lipid–protein interactions become reestablished, not all do so, and this results in membranes that maintain function but that still contain membrane aberrations; this can lead to problems in membrane structure and function over time and ultimately early cell death. Part of the difference between cold shock and cooling damage is the magnitude of membrane alterations. Cold shocked unextended spermatozoa incur greater membrane damage than spermatozoa that are cooled following dilution in an extender. This is because the lipid and lipoprotein additives in the spermatozoal diluent and a controlled cooling rate minimize damage when the membranes are within the temperature range at which they undergo the phase transition. However, although cooling spermatozoa increases the overall lifespan of a spermatozoon, even using the best techniques, cooled spermatozoa incur some cooling damage, which causes these cooled spermatozoa to exhibit a shorter lifespan when rewarmed than fresh spermatozoa. Cooling effects on cell metabolism Spermatozoa are terminally differentiated cells which have highly condensed chromatin and have lost most of their cellular organelles during spermatogenesis. Therefore, spermatozoa have very limited capacity for protein synthesis or cellular repair and, upon collection, have a relatively short lifespan. During normal spermatozoal metabolism, both anaerobic (glycolysis) and aerobic (tricarboxylic acid cycle) respiration convert monosaccharides into ATP, which is utilized for spermatozoal movement. Metabolic waste products – lactic acid and oxygen free radicals (hydrogen peroxide) – can irreparably damage the cells. Oxygen free radicals induce lipid peroxidation, which causes membrane permeability and metabolic enzyme dysfunction, leading to the loss of cell
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integrity, motility, and viability.23 Similarly, lactic acid production can reduce the pH of the medium, which decreases cell enzyme activity, resulting in the loss of ATP production and ultimately in cell death. Two methods can be utilized to reduce the production of oxygen free radicals and lactic acid. These include (i) suspending the spermatozoa in diluents that contain no oxygen, thereby forcing the cells to utilize anaerobic respiration, and (ii) cooling the cells. R diluent Oxygen-free diluents, such as the Caprogen in which the diluent is saturated with nitrogen gas to remove all oxygen, maintain spermatozoa for 3–6 days at room temperature, without affecting spermatozoal quality.24 Although this diluent is used extensively in New Zealand for storing bull spermatozoa, the need to use glass storage devices that are nonpermeable to gases has limited the use of this diluent elsewhere. The other method to reduce spermatozoal metabolism is to cool the spermatozoa. Cooling spermatozoa actually has two benefits. First, similar to anaerobic media described above, if spermatozoa are kept in an undisturbed diluent or a sealed container, aerobic metabolism quickly exhausts the oxygen dissolved in the medium and the cells are forced to rely on anaerobic metabolism only, resulting in lactic acid as the only metabolic end product. Second, reducing the temperature reduces the kinetics of the metabolic enzymes and metabolic requirements of the cells. A rule of thumb is that the rate of a reaction will double for every 10 K (Kelvin) increase in temperature.25 This projects that enzyme kinetics as well as metabolic needs decrease by approximately half with each 10◦ C reduction in temperature. Therefore, if the temperature of stallion spermatozoa is reduced from body temperature (38◦ C) to 5◦ C, overall metabolism will decrease to approximately 7% of that at body temperature (Figure 127.4). 100
80 Metabolic Rate (%)
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Therefore, the cells produce less lactic acid, there is less deterioration of cellular machinery, and the rate of lipid peroxidation by the few reactive oxygen species produced is slowed as well, extending the overall lifespan of the spermatozoa.
Minimizing cooling-induced damage to stallion spermatozoa Not only are metabolic rate and intracellular enzyme kinetics reduced at low temperature, but the rates of membrane transporters and the activity of membrane pores are also reduced. Therefore, the ability to move energy substrates into the cell and metabolic waste products out of the cell is reduced; this does not necessarily pose a problem, since overall cell metabolism is also reduced. However, if temperature affects one (metabolism or transport) more than the other, problems can arise, either in maintaining sufficient ATP in the cell or in eliminating waste products from the cells. Such a problem will likely not be observed during the cooling process itself, but will likely be manifested after long periods of storage. When stallion spermatozoa were held for 48 hours, higher percentages of motile spermatozoa were observed for spermatozoa held at lower temperatures until the temperature reached 2◦ C (Figure 127.5).19 At 0◦ C and 2◦ C, either membrane transport could not maintain the supply of monosaccharides necessary for cell metabolism or the removal of metabolic waste products, or cell metabolism was too low to support cell viability. Regardless, these data define an optimal temperature and a limiting temperature for maintaining stallion spermatozoa. These data refer only to stallion sperm, as differences in metabolic mechanisms and rates and differences in membrane composition and structure mean that optimal conditions to cool and store spermatozoa from other species will be different. A variety of compounds can be added to stallion spermatozoa to minimize the damage incurred by stallion spermatozoal membranes when they are cooled. Phillips and Lardy26 demonstrated that egg 50
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Figure 127.4 The relative percentage of stallion sperm metabolic rate at various temperatures.
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Temperature (°C)
Figure 127.5 The percentages of motile stallion sperm after storage at different temperatures for 48 hours.
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Principles of Cooled Semen yolk prevented bull spermatozoa from cold shock damage when the cells were cooled. Since then, egg yolk has proven useful to prevent cold shock in spermatozoa from many species, including stallion spermatozoa. Subsequent research has determined that it is the low-density lipoprotein fraction of the egg yolk,27, 28 and the phospholipid portion of the lipoproteins in particular,28–30 that benefit spermatozoa during cooling. Therefore, lipoproteins and phospholipids from other sources (milk, skim milk, cream, and soy products) have been used as diluent additives to cool stallion spermatozoa. However, the mechanism by which the phospholipids or lipoproteins affect the spermatozoon is not understood, as added phospholipids and lipoproteins do not incorporate into the spermatozoal membranes31 and their effects can be removed by washing them from the cells.28, 32 Changing the lipid composition of spermatozoa Studies using synthetic membranes delineate that membranes composed of lipids with more unsaturated fatty acyl chains (both in the number of unsaturated fatty acyl chains and in the number of unsaturated bonds in each fatty acyl chain) exhibit lower transition temperatures than membranes composed of lipids containing saturated fatty acyl chains. Similarly, membranes composed of lipids possessing short fatty acyl chains exhibit lower transition temperatures than those composed of lipids possessing long fatty acyl chains. However, increasing the amount of lipid with unsaturated fatty acyl chains is likely to be more beneficial than increasing the number of short-chain lipids, since adding short-chain lipids to spermatozoa can induce membrane fusion, resulting in the acrosome reaction at lower concentrations and cell death at higher lipid concentrations.33, 34 Finally, as already mentioned, adding cholesterol to membranes also lowers the transition temperature of membranes. Therefore, it is not surprising that spermatozoa from different species which exhibit very different lipid compositions also exhibit very different capacities to withstand cooling damage. When animals are fed diets containing elevated levels of unsaturated lipids, the composition of the lipid fatty acyl chains in the spermatozoon is altered after 4–6 weeks.35–40 However, even with higher levels of unsaturated fatty acyl chains in the sperm, most studies report either no improvement or only limited improvement in spermatozoa surviving cooling to 5◦ C or cryopreservation. Darin-Bennett and White41 reported that the amount of cholesterol spermatozoa from different species contained influenced their susceptibility to
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cold shock. In addition, cholesterol can be readily inserted into spermatozoal membranes in vitro, using cholesterol-loaded cyclodextrins. Increasing the cholesterol content of spermatozoa improves the survival of stallion spermatozoa after cooling to 5◦ C42 and after cryopreservation,43, 44 as well as the cryosurvival rates of bull45, 46 and ram47 spermatozoa. Adding cholesterol to spermatozoa did not improve the cryosurvival rates of rainbow trout48 or chicken (J. Graham, unpublished observations) spermatozoa. Cholesterol likely does not benefit these last species, as these spermatozoa do not exhibit ‘cold shock’, probably because of the spermatozoal membranes remaining fluid at 5◦ C.
Effect of cooling and warming rate As previously mentioned, rapidly cooling sperm from body temperature to 5◦ C results in cold shocking the spermatozoa. Moran et al.19 conducted an exhaustive series of experiments to determine the optimal rates for cooling stallion sperm, and when those rates needed to be applied. These experiments indicated that spermatozoa diluted in a skim-milk diluent can be cooled from body temperature or from room temperature to 19◦ C rapidly. However, upon reaching 19◦ C, the cells must be cooled slowly (–0.05◦ C/min) to 9◦ C for maximum retention of spermatozoa viability. Upon reaching 9◦ C, the spermatozoa can then be cooled rapidly, again, to 5◦ C. This ‘critical’ temperature range that requires slow cooling is also the temperature range in which the stallion spermatozoal membrane undergoes the phase transition from the fluid to the gel state. When one examines the cooling rates of passive cooling devices, the cooling rate through this critical temperature range is quite slow.49, 50 It should be noted, however, that the rate of cooling, even through the critical temperature range, is dependent upon the cooling diluent. Stallion spermatozoa diluted in egg yolk containing diluents can be cooled at faster rates than spermatozoa diluted in diluents containing only skim milk as the lipid/lipoprotein source (J. Graham, unpublished observations). Warming rates appear to be less critical for stallion spermatozoal survival, and cells warmed to body temperature at any reasonable rate appear to maintain cell function.
Effect of storage temperature As mentioned previously, the optimal temperature to store stallion spermatozoa should reduce spermatozoal metabolism effectively, but should not induce an imbalance between the energy still required to
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maintain cell viability and the capacity to transport nutrients into the cell and waste products out of the cell. When stallion spermatozoa were maintained at different temperatures for 24 or 48 hours, the percentage of motile spermatozoa in the samples after warming increased as the storage temperature decreased from 16◦ C to 4◦ C, but, when storage temperature was 0◦ C or 2◦ C, the percentages of motile spermatozoa in the samples was reduced (Figure 127.5). Therefore, the optimal storage temperature for stallion spermatozoa stored for 48 hours appears to be 4–6◦ C.19 At higher storage temperatures, metabolism is not sufficiently reduced, and, at 0–2◦ C, either ATP production is too low to maintain cell viability or there is an imbalance between metabolism and transport of nutrients or waste products.
Conclusions Cooling stallion spermatozoa to approximately 5◦ C can extend the overall longevity of the cells after collection. However, care must be taken to extend the cells in a proper diluent (one that contains lipids and/or lipoproteins), cool the cells at an appropriate rate (the specific cooling rate is dependent upon the diluent), and store the cells at the correct temperature (4–6◦ C). In addition, the spermatozoon’s biochemistry and physiology has been briefly described so that one can understand why these precautions need to be adhered to, and mechanisms to alter stallion spermatozoal biochemistry have been discussed to shed light on potential methodologies to improve the survival rates of cooled stallion spermatozoa. This is particularly important, as some stallions produce spermatozoa that, because of their specific biochemistry, do not survive cooling well, and such methodologies may prove useful in modifying spermatozoal membranes or metabolism sufficiently to permit these cells to survive the cooling and warming process.
References 1. Pickett BW. Seminal extenders and cooled semen. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 746–54. 2. Singer SJ, Nicolson GL. The fluid mosaic model of the structure of cell membranes. Science 1972;175:720–31. 3. Parks JP, Lynch DV. Lipid composition and thermotropic phase behavior of boar, bull, stallion, and rooster sperm membranes. Cryobiology 1992;29:255–66. 4. Aloia RC, Curtin CC, Gordon LM. Lipid Domains and the Relationship to Membrane Function. New York: Liss, 1988. 5. Rao KV. Role of Cell Surface in Development, vol. 1. Boca Raton: CRC Press, 1987. 6. Isrealachvili JN, Marcelja S, Horn RG. Physical principles of membranes organization. Q Rev Biophys 1980;13:121–200.
7. Kuypers FA, Roelofsen B, Berendsen W, Op den Kamp JAF, van Deenen LLM. Shape changes in human erythrocytes induced by replacement of the native phosphatidylcholine with species containing various fatty acids. J Cell Biol 1984;99:2260–6. 8. Gruner SM. Intrinsic curvature hypothesis for biomembrane lipid composition: a role for nonbilayer lipids. Proc Natl Acad Sci USA 1985;82:3665–9. 9. Tilcock CPS, Cullis PR. Lipid polymorphism. Ann N Y Acad Sci 1987;492:88–102. 10. Quinn PJ. Principles of membrane stability and phase behavior under extreme conditions. J Bioenerg Biomemb 1989;21:3–19. 11. Gadella BM, Miller NG, Colenbrander B, van Golde LM, Harrison RR. Flow cytometric detection of transbilayer movement of fluorescent phospholipid analogues across the boar sperm plasma membrane: elimination of labeling artifacts. Mol Reprod Dev 1999;53:108–25. 12. Gadella BM, Harrison RA. The capacitating agent bicarbonate induces protein kinase A-dependent changes in phospholipid transbilayer behavior in the sperm plasma membrane. Development 2000;127:2407–30. 13. Flesch FM, Brouwers JF, Nieverstein PF, Verkleij AJ, van Golde LM, Colenbrander B, Gadella BM Bicarbonate stimulated phospholipid scrambling induces cholesterol redistribution and enables cholesterol depletion in the sperm plasma membrane. J Cell Sci 2001;114(Pt 19): 3543–55. 14. Amann RP, Graham JK. Spermatozoal Function. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 715–45. 15. Hammerstedt RH, Graham JK, Nolan JP. Cryopreservation of mammalian sperm: what we ask them to survive. J Androl 1990;11:73–88. 16. Langlais J, Roberts KD. A molecular membrane model of sperm capacitation and the acrosome reaction of mammalian sperm. Gamete Res 1985;12:183–224. 17. Chapman D, Williams RM, Ladbrooke BD. Physical studies of phospholipids. VI. Thermotropic and lyotropic mesomorphism of some 1,2-diacyl-phosphatidylcholines (lecithin). Chem Phys Lipids 1967;1:445–75. 18. Caffrey M, Feigenson GW. Fluorescence quenching in model membranes. 3. Relationships between calcium adenosinetriphosphatase enzyme activity and the affinity of the protein for phosphatidylcholines with different acyl chain characteristics. Biochemistry 1981;20:1949–61. 19. Moran DM, Jasko DJ, Squires EL, Amann RP. Determination of temperature and cooling rate which induce cold shock in stallion spermatozoa. Theriogenology 1992;38:999–1012. 20. Ladbrooke BD, Williams RM, Chapman D. Studies on lecithin-cholesterol-water interactions by differential scanning calorimetry and X-ray diffraction. Biochim Biophys Acta 1968;150:333–40. 21. Watson PF. The effects of cold shock on sperm cell membranes. In: Morris GJ, Clark A (eds) Effects of Low Temperatures on Biological Membranes. New York: Academic Press, 1981; pp. 189–218. 22. Watson PF. The roles of lipid and protein in the protection of ram spermatozoa at 5-degrees-C by egg yolk lipoprotein. J Reprod Fertil 1981;62:483–92. 23. Hammerstedt RH. Maintenance of bioenergetic balance in sperm and prevention of lipid peroxidation: a review of the effect and design of storage preservation systems. Reprod Fertil Dev 1993;5:675–90.
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Principles of Cooled Semen 24. Verberckmoes S, Van Soom A, Dewulf J, de Kruir A. Comparison of three diluents for the storage of fresh bovine semen. Theriogenology 2005;63:912–22. 25. Houston PL. Chemical Kinetics and Reaction Dynamics. Mineola, NY: Dover Publications, 2001; pp. 34–90. 26. Phillips PH, Lardy HA. A yolk-buffer pablum for the preservation of bull semen. J Dairy Sci 1940;23:399–404. 27. Pace MM, Graham EF. Components of egg yolk which protect bovine spermatozoa during freezing. J Anim Sci 1974;39:1144–9. 28. Watson PF. The protection of ram and bull spermatozoa by the low density lipoprotein fraction of egg yolk during storage at 5◦ C and deep-freezing. J Thermal Biol 1976;1:137– 41. 29. Kampschmidt RF, Mayer DT, Herman HA. Lipid and lipoprotein constituents of egg yolk in the resistance and storage of bull spermatozoa. J Dairy Sci 1953;36:733–42. 30. Parks JE, Meacham TN, Saacke RG. Cholesterol and phospholipids of bovine spermatozoa. II. Effects of liposomes prepared from egg phosphatidylcholine and cholesterol on sperm cholesterol, phospholipids and viability at 4◦ C and 37◦ C. Biol Reprod 1981;24:399–404. 31. Foulkes JA. The separation of lipoproteins from egg yolk and their effect on the motility and integrity of bovine spermatozoa. J Reprod Fertil 1977;49:277–84. 32. Quinn PJ, Chow PYW, White IG. Evidence that phospholipid protects spermatozoa from cold shock at a plasma membrane site. J Reprod Fertil 1980;60:403–7. 33. Graham JK, Foote RH, Parrish JJ. Effect of dilauroylphosphatidylcholine on the acrosome reaction and subsequent penetration on bull spermatozoa into zona-free hamster eggs. Biol Reprod 1986;35:413–24. 34. Graham JK, Foote RH, Hough SR. In vitro penetration of zona-free hamster eggs by liposome treated sperm from the bull, ram, stallion and boar. Biol Reprod 1987;37:181–8. 35. Paulenz H, Taugbol O, Kommisrud E, Grevle IS. Effect of dietary supplementation with cod liver oil on cold shock and freezability of boar semen. Reprod Dom Anim 1999;34:431–5. 36. Rooke JA, Shao CC, Speake BK. Effects of feeding tuna oil on the lipid composition of pig spermatozoa and in vitro characteristics of semen. Reproduction 2001;121: 315–22. 37. Mitre R, Cheminade C, Allaume P, Legrand P, Legrand AB. Oral intake of shark liver oil modifies lipid composition and improves motility and velocity of boar sperm. Theriogenology 2004;62:1557–64.
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38. Brinsko SP, Varner DD, Love CC, Blanchard TL, Day BC, Wilson ME. Effect of feeding a DHA-enriched nutriceutical on the quality of fresh, cooled and frozen stallion semen. Theriogenology 2005;63:1519–27. 39. Amorim LS, Torres CAA, Amorim EAM, Graham J. Effect of dietary supplementation with salmon oil on cryopreservation of boar semen. Reprod Dom Anim 2008;43(Suppl. 3):124. 40. Kelso KA, Cerolini S, Noble RC, Sparks NH, Speake BK. The effects of dietary supplementation with docosahexaenioc acid on the phospholipid composition of avian spermatozoa. Comp Biochem Physiol B Biochim Mol Biol 1987;118:65–9. 41. Darin-Bennett A, White IG. Influence of the cholesterol content of mammalian spermatozoa on susceptibility to coldshock. Cryobiology 1977;14:466–70. 42. Kirk ES. Flow Cytometric Evaluation of Stallion Sperm. MS thesis. Colorado State University, CO, 2001. 43. Combes GB, Varner DD, Schroeder F, Burghardt RC, Blanchard TL. Effect of cholesterol on the motility and plasma membrane integrity of frozen equine spermatozoa after thawing. J Reprod Fertil 2000;56(Suppl):127–32. 44. Moore AI, Squires EL, Graham JK. Adding cholesterol to the stallion sperm plasma membrane improves cryosurvival. Cryobiology 2005;51:241–9. 45. Purdy PH, Graham JK. Effect of cholesterol-loaded cyclodextrin on the cryosurvival of bull sperm. Cryobiology 2004;48:36–45. 46. Moc´e E, Graham JK. Cholesterol-loaded cyclodextrins added to raw bull ejaculates improve sperm cryosurvival. J Anim Sci 2006;84:826–33. 47. Morrier A, Th´eriault M, Castonguay F, Bailey J. Effect of cholesterol loaded methyl-b-cyclodextrin on ram sperm during cryopreservation, cold-shock and artificial insemination. In: Proceedings of the 37th Meeting of the Society for the Study of Reproduction, 2004, p. 239. ¨ ¨ 48. Muller K, Muller P, Pincemy G, Kurz A, Labbe C. Characterization of sperm plasma membrane properties after cholesterol modification: consequences for cryopreservation of rainbow trout spermatozoa. Biol Reprod 2008;78:390–9. 49. Katila T, Combes GG, Varner DD, Blanchard TL. Comparison of three containers used for the transport of cooled stallion semen. Theriogenology 1997;48:1085–92. 50. McKinnon AO, Walker JB. Effect of ambient temperature and container on temperature of extended equine semen. World Equine Vet Rev 1998;3:5–11.
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Chapter 128
Breeding with Cooled Transported Semen Juan C. Samper
Proper artificial insemination (AI) in horses involves the deposition of an adequate number of sperm into the reproductive tract of the mare at the appropriate time. The technique is considered to be the oldest and most efficient reproductive technique for dissemination of genetics.1 AI can be done with semen collected at the farm for immediate use, but maximum utilization of this technology is achieved when mares are bred at considerable distances from where the stallion stands. To achieve this, semen either can be transported in its liquid state after passive cooling and deposited in the mare several hours (12–36) after collection or can be processed for freezing and maintained in liquid nitrogen until its use several weeks or years later. Three factors must be considered in order to achieve acceptable fertility results when inseminating mares with cooled shipped semen: the inherent fertility of the mare, the quality or fertility of the semen that is received, and the reproductive management of the mare before and after insemination. This chapter will focus on some of the factors that must be considered to maximize semen quality, as well as mare management procedures that must be taken into account to improve the fertility of mares bred with cooled/transported semen.
Semen quality Breeding stallions that are going to be used for AI should have a semen evaluation performed before each breeding season. It is virtually impossible to improve the inherent quality of semen from a stallion, so it is important for the mare owners and stud farm personnel to be aware of the stallion’s semen quality. A veterinarian working for clients who decide to stand a stallion or stallions for AI has the responsibility of overseeing that the clients are properly prepared for the services they are offering. This includes
proper stimulus for the stallion, including, if necessary, the availability of a mare in heat at all times during the breeding season; availability of shipping containers; and a laboratory for the proper evaluation and processing of the semen. It is also important to inform the stallion owner that a proper semen evaluation should be performed to determine semen quality and microbiological status of the genital integument and semen, including equine viral arteritis and Taylorella equigenitalis (the causative agent of contagious equine metritis). Stallion owners must realize that their stallion competes for a limited number of mares based on his genetic merit and/or show record. However, the service that is provided by the breeding farm plays a key role for some breeders in deciding which stallion to use. Some breeders would rather breed their mare to a second-choice stallion with the guarantee that they can order semen easily, the semen arrives on time, and is of good quality. Many stallions, particularly show or sport stallions, breed only a reduced number of mares, so they ejaculate only a few times during the breeding season. Some of these stallions have very large testicles and produce a high number of sperm that, if not collected on a regular basis, have the tendency to accumulate in the extragonadal ducts.2 Therefore, it is important to identify stallions that are ‘sperm accumulators’ since the accumulated sperm often have lower motility, shorter longevity, and poorer morphology than a stallion whose semen is collected on a regular basis.3 When such a stallion is offered for AI, it is important to collect semen from the stallion several times prior to breeding mares, or processing semen for cooled shipment or frozen storage. The number of ejaculates required to improve the semen quality will vary between stallions.3 Two consecutive ejaculates of similar quality should indicate that the stallion has voided the accumulated sperm. Recent evidence
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Breeding with Cooled Transported Semen suggests that dietary supplementation of stallions with certain nutraceuticals can result in an improvement in the quality of their cooled semen.4 In addition to the sexual activity of the stallion, the semen collection process can have a significant effect on the quality of the ejaculate that will be available for processing. The ideal ejaculate is one that has a low volume, a high sperm concentration, and a high number of motile sperm. Sperm concentration and semen volume are inversely proportional, i.e., the higher the volume the lower the sperm concentration. Semen volume can be affected by the length of teasing period for the stallion prior to semen collection and/or by the number of jumps the stallion has before ejaculation.5, 6 Stallions that do not ejaculate at the first attempt will also have a tendency to accumulate pre-ejaculatory (bulbourethral) fluid in the collection container. Exposure of sperm to concentrations of 10% pre-ejaculatory fluid will significantly reduce the longevity of sperm even after dilution with extender (J.C. Samper, unpublished observations). Therefore, unsuccessful semen collection attempts require that the collection bottle be emptied of this fluid.
Semen handling Raw semen is very fragile and should not be exposed to direct sunlight or sudden temperature changes. Once the stallion ejaculates, the semen should come into contact with a clean and warm container to prevent cold shock. The semen can be filtered in-line during ejaculation or filtered immediately after collection with nylon mesh or milk filter paper. This procedure is done to remove most extraneous particles and the gel fraction. However, filtration is not performed to compensate for the lack of effort in washing and cleaning the stallion prior to collection. Immediately after collection, the raw semen should be placed in an incubator at 37◦ C prior to mixing with an extender. The volume and color of the ejaculate should be recorded. In addition, the other physical characteristics of the ejaculate such as sperm movement, sperm concentration, and, if necessary, sperm morphology should be assessed from a small aliquot (1–2 mL) of raw semen that is removed prior to the addition of any diluents or extender. Once semen is diluted with a prewarmed extender it should be immediately removed from the incubator and cooled down to room temperature (20◦ C). An accurate assessment of the percentage of progressively motile sperm should be done when the semen has been diluted to 25–50 million sperm/mL.
Preservation of semen The processing of semen should be done according to the period of time that the semen is expected to re-
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main viable. It is recommended to always place the semen in an appropriate extender, even if it is going to be inseminated within a few minutes. There are several reasons why semen should be extended within a few minutes (i.e., less than 5 minutes) after collection. A proper extender buffers pH changes of the raw semen; maintains the osmolarity of the sample; provides energy and protein sources for sperm metabolism and membrane stabilization during thermal changes; reduces the detrimental effects of the seminal plasma; provides antibacterial properties by means of the antibiotics;7, 8 and maintains chromatin integrity.9 Semen can be preserved for varying periods of time. However, the state and the processing technique will be determined by the expected longevity of a given ejaculate. Fresh extended semen can be kept at 20◦ C by avoiding exposure to direct sunlight, without losing its fertilizing ability for several hours.10 If semen is intended to last 12 hours or more, either because the stallion will not be available or because the semen will be transported, it should be slowly cooled and maintained at 4–8◦ C.11 For semen that is intended to be stored for longer than 72 hours, it is recommended to process it for freezing. It is extremely important to insure that the extenders that are going to be mixed with the sperm are isothermal with the semen. In other words, extender that will be added to raw semen should be prewarmed. However, extender added to semen that has been cooled to room temperature should also be at room temperature. Failure to do so induces thermal shock to the sperm, which can result in irreversible reduction of the progressive motility and membrane damage, thereby reducing the fertilizing potential.12 Fresh semen Semen from stallions with good fertility that is collected and used immediately or up to 6 hours after collection does not necessarily have to be cooled and, in most cases, can be diluted with prewarmed extender at ratios of 1:1 to 1:3 (semen–extender) depending on the raw semen concentration and ejaculate volume. Immediately after extension, the semen should be removed from the incubator and can be cooled to room temperature (15–20◦ C) while avoiding exposure to direct light without loss of its fertilizing potential. Cooled semen Artificial insemination with cooled semen is still gaining popularity among breeders because more breed registries are accepting registration of foals resulting from the use of this method of breeding. Although there is a tremendous increase in its use, there is also a tremendous variability in the results
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achieved. In order to minimize the variability in the results, it is important that the handling and processing of the semen be standardized. When raw semen quality is good to excellent, the semen-handling and processing factors that could have the biggest impact on the quality of the semen shipped and the fertility of an insemination dose are (i) extender composition; (ii) cooling rates and container quality; (iii) dilution ratio, and (iv) the insemination process.13
Extenders Although there are several extenders that can be used for preserving semen in its liquid state, it appears that most researchers and clinicians favor the use of the non-fat, dried milk solids–glucose extender known as the Kenney extender.11 Several variations of this extender are commercially available that include primarily a change in the sugar content or the type of antibiotic added. Two types of sugars are primarily used – either glucose or sucrose, or a combination of the two. Some European countries such as Belgium, the Netherlands, and Germany use other types of extenders for routine cooling and shipping that are supplemented with egg yolk or, more recently, with no animal protein.7 In the last few years, the commerR cially available INRA-96 extender, which is a balanced salt solution enriched with phosphocaseinate, has gained popularity and is tolerated well by sperm of many stallions.14 All extenders should contain an antibiotic. The most common ones include a combination of potassium penicillin and amikacin sulfate, gentamicin, or ticarcillin. It is the author’s recommendation to avoid the use of polymyxin B since it has been show to have a deleterious effect on sperm motility.15 The bacterial flora of the stallion and the effect of the antibiotic on the sperm motion characteristics for the individual stallion should be determined because the most appropriate antibiotic(s) could vary between stallions.
Cooling rates and containers Spermatozoa suspended in an extender can be rapidly cooled from 37◦ C to 20◦ C, but require a slow cooling rate of –0.5 to –0.1◦ C/min between 20◦ C and 5◦ C to maximize spermatozoal viability. If sperm are cooled slowly, once they have reached 4–8◦ C, storage at this temperature is probably superior to storage at 20◦ C.16 It appears that the ideal storage temperature range for maintaining motility and fertility of cooled equine semen preserved for more than 12 hours in a Kenney-type extender is between 4◦ C and 6◦ C.17 However, Leboeuf et al.14 found that a storage temperature of 15◦ C was superior to 10◦ C or 4◦ C for maintaining spermatozoal motility when semen was
extended in INRA-96. Most stallions ejaculate more sperm than is needed to achieve maximal fertility; therefore, most shipments will have one or two doses containing 1 × 109 progressively motile sperm, with a final sperm concentration of 25–50 × 106 sperm/mL. Semen is slowly cooled to, and stored at, 4–8◦ C. However, with the popularity of some stallions, the total number of sperm needed has come under scrutiny, and it appears that many stallions could be able to achieve their maximal fertility with much less sperm than the 1 billion progressively motile sperm recommended in years past.18–20 There are several containers currently available commercially that are designed for cooling and transR is the most port of equine semen. The Equitainer widely used container for passive cooling and transporting of equine semen.21 Less expensive, disposable semen-transporting containers are also availR (Exodus Breeders able such as the Equine Express Corporation, York, PA, USA)or Equine Semen Transporter (ESTTM ) models (Plastilite Corporation, Omaha, NE, USA). These containers are also passivecooling transport devices, which provide variable rates of cooling. The main differences between the R and other semen-transporting systems Equitainer reside in that the former is less sensitive to environmental temperatures providing a more controlled rate at which the semen is cooled. In addition the R is more effective in maintaining the deEquitainer sired temperature for a longer period of time. A study conducted by Brinsko et al.22 to evaluate the effect of storage containers and environmental temperature indicated that sperm motility is adequately maintained in most commercial equine semen transport containers when the outside temperature is between 22◦ C and 37◦ C. However, an environmental temperature of –20◦ C for 6 hours resulted in a reduction in progressive sperm motility in most of the disposable containers. This study indicated that the R Equitainer was a better container to isolate the semen from the effect of environmental temperatures. In addition, sperm that are not cooled when extended in a Kenney-type extender are more susceptible to accelerated chromatin denaturation.9
Dilution ratio Although the processing and handling of the sperm is tolerated well by the semen from the majority of the stallions, some stallions have a significant and unexplainable reduction in semen quality when it is cooled and stored for over 12 hours. It appears that, for some of the ‘problem’ stallions, partial or total removal of the seminal plasma is important to maintain the percentage of progressively motile sperm following cooled storage. The ideal dilution ratio at which
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Breeding with Cooled Transported Semen sperm should be extended will vary according to the sperm concentration in the raw semen and the total number of progressively motile sperm. The author has found that semen with 100 million sperm should be diluted at a 1:3 ratio (semen–extender). Every increment of 50 million sperm/mL will increase the dilution ratio by 1; therefore, when the concentration of the ejaculate is between 450 and 500 million/mL the dilution rate would be 1:10. Extending the semen using this method will result in a final sperm concentration of 25–50 million/mL.13 The final volume to be transported will typically be calculated so that a total of 1 billion progressively motile sperm cells are sent to the mare. Contrary to what some owners believe, sending the entire ejaculate to provide more sperm in most cases is detrimental to semen quality. When using the skimmed milk–glucose extender, the packaging of the semen should be performed under anaerobic conditions. Semen can be packaged in preloaded syringes, disposable baby bottle liners, R or Whirl-Pak bags, provided that all the air is removed from the package prior to placing it in the container. If semen is to be packed and transported in syringes, they must be free of any product that could be toxic to spermatozoa. Air-tite syringes (HSW, Tuttingen, Germany) are proven to have no deleterious effect on semen quality.
Evaluation of cooled shipped semen It is expected that semen that is transported for several (6–24) hours will have a reduction in quality. The rate of deterioration is determined primarily by the quality of the stallion’s raw semen, the handling of the semen after collection, and the length of time it is transported. Poor-quality raw semen handled properly and transported for only a few hours can arrive in better condition than excellent raw semen that was improperly diluted and cooled and transported for over 24 hours. Every veterinarian or veterinary technician should evaluate motility as a rough indicator of semen quality prior to insemination. Although semen evaluation techniques are discussed in detail elsewhere in this text, there are a few considerations that should be taken into account when evaluating transported semen at the receiving end. Individuals evaluating cooled transported semen should be aware that evaluation is generally limited. The parameters that can be evaluated accurately are volume, motility, morphology, sperm concentration, and total number of sperm in the shipment. The evaluation should be carried out prior to insemination whenever possible. This gives the breeder the alternative of not inseminating should the semen be dead, or improving the quality by using gradient centrifugation techniques. If more than one
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dose is shipped and one dose is found to contain dead sperm, it should not be assumed that the other dose(s) are also of poor quality. However, it is also important to remember that some doses that receive poor scores on analysis will sometimes result in pregnancies. Therefore, it can be a challenge to choose whether to inseminate a mare with a poor dose of semen. If poor-quality semen arrives, the receiver should immediately contact the shipper to determine whether the semen was improperly handled at the processing center or during transport. For these types of claims the processing center can leave a sample at its facility in order to clarify these claims. Motility This analysis is often the only one done by receivers of cooled semen. The percentage of motile sperm is highly influenced by the temperature. There can be a dramatic reduction in progressive motility and velocity when the sample is not allowed to warm to body temperature (37◦ C). Also, temperatures over 37◦ C are detrimental to motility estimates.23 When evaluating motility there is no reason to warm the entire shipment, and, in fact, it is probably best to inseminate the mare with the cooled dose taken directly from the shipper. A small aliquot of the extended semen can be warmed separately from the insemination dose to 37◦ C prior to evaluating. Morphology Although morphological evaluations are not routinely performed by the receiver, it is part of the evaluation to consider if repeated inseminations are not resulting in pregnancies. The easiest and most accurate way to evaluate the morphology of a semen shipment is by placing one or two drops of semen in buffered formalin saline (BFS) and carrying out the evaluation on a wet mount using the oil immersion objective under phase contrast microscopy. Alternatively, conventional stains such as eosin–nigrosin can be used, as long as the sperm are fixed first in BFS and then stained from this solution. Concentration Like morphology, sperm concentration is not routinely done by many receiving cooled semen. If the technician or veterinarian feels the sample is too dilute, a count should be performed. Sperm counters that are based on estimation of light absorbance such as the DensimeterTM (Animal Reproduction Systems, Chino, CA, USA) or SpermaCueTM (Minitube of America, Verona, WI, USA) or a spectrophotometer are not appropriate for counting of extended semen because of interference by the extender. Therefore, the most reliable way of estimating concentration is by using a hemocytometer. An
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Stallion: Semen Collection, Evaluation and Artificial Insemination
JCS VETERINARY REPRODUCTIVE SERVICES LTD. Juan C. Samper DVM, MSc, PhD Diplomate American College of Theriogenologists 2943 216th Street, Langley BC V2Z 2E6 CANADA Tel: (604) 530 -0223 Fax: (604) 530 –0299 email: [email protected]
SEMEN COLLECTION AND DELIVERY RECORD DATE:_____________________
TIME OF COLLECTION:__________________
STALLION NAME:______________________________ OWNER:__________________________
BREED:_____________
SHIPPED TO: ___________________________
SHIPPED BY : AIR SAMEDAY _________________ OVERNIGHT___________________ Raw Semen Motility:__________________ Concentration:_______________Million/ml Extender Type:_____________ Antibiotic:___________ Dilution rate S/E:__________ Motility extended:________ Total Volume:__________ Volume per dose:_________ Motile sperm per dose:_________________ Number of Doses Shipped:___________ This shipment contains _______ insemination doses. The shipping container is not designed for long term storage. Spermatozoa in the oviduct of the mare can live for more than 96 hours, therefore it is suggested that the mare be inseminated with the entire contents upon arrival. If the semen does not look viable upon arrival please call us immediately to arrange for a replacement shipment. If we do not hear from you we will assume that all is fine.
______________________________ J.C. Samper DVM, MSc, PhD Diplomate A.C.T. Figure 128.1 Semen shipping form that is used in the authors practice. This form is intended to give the receiver a general idea of the semen quality as well as identification of the stallion that will be used to inseminate the mare.
alternative to the hemocytometer is the computerassisted semen analyzer (CASA) or the newly introduced fluorescence-based NucleoCounterTM SP-100 (Chemometec, Allerød, Denmark).24 It is imperative that individuals shipping cooled semen send an information sheet accompanying the shipment. Figure 128.1 shows an example of the information sheet that accompanies all shipments from the author’s practice. Although this form is not intended to be an exhaustive evaluation of the semen quality it is meant to inform the receiver of the proper identification of the stallion as well as the initial semen quality. Discepancies in stallion identification
should be clarified before insemination. In addition semen quality must be examined prior to insemination and compared with that of the sheet. Wide discrepancies in semen quality should be investigated further with the individual shipping the semen. Individuals involved in the insemination of mares with cooled transported semen should always evaluate the semen prior to inseminating the mare if possible. After properly warming, samples that have apparent poor-quality and/or low-motility semen can be centrifuged using a cushion preparation in order to reduce the volume of the inseminate to 1–2 mL. The sperm can be deposited by rectally guided
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Breeding with Cooled Transported Semen insemination to maximize the population of sperm that will reach the oviduct.25 To further enrich the live sperm population, the previously centrifuged sample can be recentrifuged using a gradient centrifugation technique to enrich the inseminate and select the best sperm population.26, 27 This low-dose inseminate should also be deposited by a rectally guided deephorn insemination technique.
Insemination of cooled semen Cooled shipped semen with motility > 35% and adequate concentration and numbers can be deposited in the body of the uterus using a conventional insemination technique. When two doses of semen are shipped of < 100 mL total volume, the author recommends inseminating the entire shipment at once. This is of particular importance when the stallion has suboptimal semen quality or its semen does not last long enough to be stored in a shipping container for 36–48 hours. Controversy exists regarding the number of insemination doses and the number of times that the mare should be inseminated.28, 29 In a retrospective study by Sieme et al.,30, 31 increasing the number of breedings per cycle had a significant effect on the pregnancy rate of mares bred with cooled semen. There was a 5% increase in fertility of mares that were inseminated twice compared with mares that were bred only once in the cycle. There was a further increase if mares were bred three or more times. Although the study analyzed over 100 000 mare cycles, it failed to analyze the fertility of the stallion and the interval between breeding and ovulation. It is well documented that breeding close to ovulation increases fertility.32 In the author’s opinion, there is no clear answer and it would depend on the quality of a stallion’s semen and the reproductive history of the mare. If the semen is of good quality and the mare is a young fertile mare she could be inseminated once or twice at 12- or 24-hour intervals. However, if the semen has poor quality, it would be important to breed the mare as close to ovulation with the entire dose. On the other hand, if the semen is of good quality but the mare has fertility problems or has a tendency to accumulate uterine fluid she should be bred only once within 24 hours prior to ovulation. The problem mare bred with poor-quality semen should be avoided since she will be a source of aggravation for the mare owner, the stallion owner, and the veterinarian. However, if there is no option available, these mares should be bred as close as possible to ovulation with all the semen that is provided. Regardless of the quality of the semen and the fertility potential of the mares, it is recommended that all mares be given an ovulation-inducing agent such as human chorionic gonadotropin or deslorelin to min-
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imize the insemination-to-ovulation interval,33 to reduce the costs of collection and transportation, and to reduce the amount of post-breeding inflammation in the mare’s uterus.
Fertility of cooled shipped semen The ability to achieve pregnancies with cooled semen has been documented for several years.11 In 1994, Heiskanen et al.34 reported on differences of semen between stallions cooled for several hours, with adequate fertility being retained up to 40 hours after collection. However, a minimum threshold of semen quality must be met in order to achieve satisfactory results. The total number of motile sperm in the shipment is estimated by multiplying the volume received by the sperm concentration per milliliter by the estimated motility of the sample. Shipments that have greater than 30% motility, with adequate sperm numbers and good sperm morphology, should achieve pregnancy rates of more than 50% per cycle when the mares are bred within 24 hours before ovulation. In a retrospective study reported by Metcalf,35 semen that was received with more than 40% motility, more than 50% morphologically normal sperm, and adequate concentration resulted in pregnancy rates over 50% per cycle (1.6 cycles per pregnancy) compared with a lower pregnancy rate for semen of lesser quality. The fertility of cooled semen is highly dependent also on the fertility of the mares being inseminated. Therefore, appropriate breeding management procedures, such as timing of insemination, diagnosis of endometritis, therapies prior to insemination, and identification of mares prone to developing persistent post-breeding endometritis, are critical. The implementation of appropriate therapies performed in a timely fashion to address issues that can affect fertility will have a significant impact on the pregnancy rates that can be achieved.
References 1. Basrur PK, King WA. Genetics then and now: breeding the best and biotechnology. Rev Sci Tech 2005;24:31–49. 2. Amann RP, Thompson DL, Squires EL, Pickett BW. Effects of age and frequency of ejaculation on sperm production and extragonadal sperm serves in stallions. J Reprod Fertil Suppl 1979;27:1–6. 3. Thompson JA, Love CC, Stich KL, Brinsko SP, Blanchard TL, Varner DD. A Bayesian approach to prediction of stallion daily sperm output. Theriogenology 2004;62:1607– 17. 4. Brinsko SP, Varner DD, Love CC, Blanchard TL, Day BC, Wilson ME. Effect of feeding a DHA-enriched nutriceutical on the quality of fresh, cooled and frozen stallion semen. Theriogenology 2005;63:1519–27.
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5. Bedford S, Hinrichs K. Effect of insemination volume on pregnancy rates of pony mares. Theriogenology 1994;42: 571–8. 6. Gahne S, G˚anheim A, Malmgren L. Effect of insemination dose on pregnancy rate in mares. Theriogenology 1998; 49:1071–4. ¨ ¨ 7. Aurich C, Seeber P, Muller-Schl osser F. Comparison of different extenders with defined protein composition for storage of stallion spermatozoa at 5 degrees C. Reprod Domest Anim 2007;42:445–8. 8. Pickett BW, Burwash LD, Voss JL, Back DG. Effect of seminal extenders on equine fertility. J Anim Sci 1975;40:1136–43. 9. Love CC, Thompson JA, Lowry VK, Varner DD. Effect of storage time and temperature on stallion sperm DNA and fertility. Theriogenology 2002;57:1135–42. 10. Batellier F, Duchamp G, Vidament M, Arnaud G, Palmer E, Magistrini M. Delayed insemination is successful with a new extender for storing fresh equine semen at 15 degrees C under aerobic conditions. Theriogenology 1998;50:229–36. 11. Varner DD, Blanchard TL, Meyers PJ, Meyers SA. Fertilizing capacity of equine spermatozoa stored for 24 hrs at 5 or 20 degrees C. Theriogenology 1989;32:515–25. 12. Watson PF, Plummer IM, Allen WE. Quantitative assessment of membrane damaged in cold shocked spermatozoa of stallions. J Reprod Fertil Suppl 1987;35:651–3. 13. Samper JC. Artificial insemination. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination, 2nd edn. St. Louis: Saunders, 2009; pp. 165–74. 14. Leboeuf B, Guillouet P, Batellier F, Bernelas D, Bonn´e JL, Forgerit Y, Renaud G, Magistrini M. Effect of native phosphocaseinate on the in vitro preservation of fresh semen. Theriogenology 2003;60:867–77 15. Aurich C, Spergser J. Influence of bacteria and gentamicin on cooled-stored stallion spermatozoa. Theriogenology 2007;67:912–18. 16. Rota A, Furzi C, Panzani D, Camillo F. Studies on motility and fertility of cooled stallion spermatozoa. Reprod Domest Anim 2004;39:103–9. 17. Akcay E, Reilas T, Andersson M, Katila T. Effect of seminal plasma fractions on stallion sperm survival after cooled storage. J Vet Med A Physiol Pathol Clin Med 2006;53:481–5. 18. Morris L. Advanced insemination techniques in mares. Vet Clin North Am Equine Pract 2006;22:693–703. 19. Lyle SK, Ferrer MS. Low-dose insemination: why, when and how. Theriogenology 2005;64:572–9. 20. Brinsko SP. Insemination doses: how low can we go? Theriogenology 2006;66:543–50. 21. Douglas-Hamilton DH, Osol R, Osol G, Driscoll D, Noble H. A field study of the fertility of transported equine semen. Theriogenology 1984;22:291–304.
22. Brinsko SP, Rowan KR, Varner DD, Blanchard TL. Effects of transport container and ambient storage temperature on motion characteristics of equine spermatozoa. Theriogenology 2000;53:1641–55. 23. Amann RP, Graham JK. Spermatozoal function. In: McKinnon AO, Voss JM (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 715–45. 24. Comerford KL, Love CC, Brinsko SP, Edmond AJ, Waite JA, Teague SR, Varner DD. Validation of a commercially available fluorescence based instrument to evaluate stallion spermatozoal concentration. Proc Am Assoc Equine Pract 2008;54:367–8. 25. Samper JC, Gomez I, Sanchez R. Rectally guided or hysteroscopic insemination: is there a difference? J Equine Vet Sci 2008;28:640–4. 26. Edmond AJ, Teague SR, Brinsko SP, Comerford KL, Waite JA, Mancill SS, Love CC, Varner DD. Effect of densitygradient centrifugation on quality and recovery rate of equine spermatozoa. Anim Reprod Sci 2008;107:318. 27. Morrell JM, Dalin AM, Rodriguez-Martinez H. Comparison of density gradient and single layer centrifugation of stallion spermatozoa: yield, motility and survival. Equine Vet J 2009;41:53–8. 28. Shore MD, Macpherson ML, Combes GB, Varner DD, Blanchard TL. Fertility comparison between breeding at 24 hours or at 24 and 48 hours after collection with cooled equine semen. Theriogenology 1998;50:693–8. 29. Squires EL, Brubaker JK, McCue PM, Pickett BW. Effect of sperm number and frequency of insemination on fertility of mares inseminated with cooled semen. Theriogenology 1998;49:743–9. 30. Sieme H, Bonk A, Hamann H, Klug E, Katila T. Effects of different artificial insemination techniques and sperm doses on fertility of normal mares and mares with abnormal reproductive history. Theriogenology 2004;62:915–28. 31. Sieme H, Sch¨afer T, Stout TA, Klug E, Waberski D. The effects of different insemination regimes on fertility in mares. Theriogenology 2003;60:1153–64. 32. Woods J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic loss in mares. Equine Vet J 1990;22:410–15. 33. Samper JC, Jensen S, Sargent J, Ruth L. Timed ovulation in mares using the Deslorelin Implant Ovuplant. J Equine Vet Sci 2002;65:121–4. 34. Heiskanen ML, Huhtinen M, Pirhonen A, M¨aenp¨aa¨ PH. Insemination results with slow-cooled stallion semen stored for approximately 40 hours. Acta Vet Scand 1994;35:257–62. 35. Metcalf ES. Pregnancy rates with cooled equine semen received in private practice. Proc Am Assoc Equine Pract 1998;44:16–18.
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Chapter 129
Experiences with a Large-Scale Cooled–Transported Semen Program Simon J. Robinson
The transportation and insemination of mares with cooled semen has been practiced in a number of horse breeds throughout the world for more than 20 years. Along with artificial insemination, it is the most widely adopted assisted reproductive technology used in horse breeding. In the Warmblood breeding industry throughout Europe and the Standardbred and Quarter Horse breeds in North America, Canada, Australia, and New Zealand, large-scale cooled–transported semen programs are now not only commonplace but form the basis of their breeding industries.1, 2 In the Standardbred breeding industries in Australia and New Zealand, thousands of breeding doses of semen are transported throughout and between these countries each year. Consequently, since the adoption of transported semen in the early 1990s, there has been a dramatic change in this breeding industry whereby the majority of mares are no longer domiciled on farms that stand the stallions. Rather, there has been an emergence of mare farms or ‘semen bases’ accommodating 50–100 mares. This change in the Standardbred breeding industry is illustrated by data from the largest Standardbred stud farm in Australia (Alabar Bloodstock, personal communication). In 1997–8, transported semen accounted for only 28% of mares bred with semen from stallions standing at Alabar’s Australian farm. The balance of the mares were maintained on the stud farm with the stallions and inseminated with fresh semen. By 2007-8, more than 85% of mares bred by the stallions standing at Alabar were inseminated with transported–cooled semen at distant locations throughout Australia and New Zealand.
Although it is widely accepted that an insemination dose of 500 million progressively motile spermatozoa (PMS) is the minimum required for achieving an acceptable per-cycle fertility rate, it should be recognized that this can often only be achieved if a dose of more than 1 billion PMS is actually dispatched. Furthermore, although this apparent safety margin of some 500 million PMS exists, technicians and stud managers should at all times try to optimize the conditions by which the semen is handled, extended, packaged, transported, and inseminated in order to achieve acceptable fertility in a large cooled–transported semen program. The wide adoption of this technology would suggest that the expertise and equipment requirements are not often limiting. However, there are a number of considerations and pitfalls which, in this author’s experience, have proven to be critical in a successful cooled semen program. This chapter addresses these practical considerations, with a view to optimizing the success and efficiency of a cooled–transported semen program.
Considerations and requirements for stallion owners and studs The adoption of any technology will be determined by economic factors such as demand and costs. Before embarking on a cooled–transported semen program, stallion owners and/or studs need to consider the likely market and demand for semen from a stallion. There are some fixed set-up costs involved in a cooled semen program (laboratory and semen
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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collection equipment and facility costs, transport containers, etc.). If the demand and/or service fee for the stallion are not sufficiently great to justify these initial set-up costs, stallion owners might consider either utilizing the facilities and expertise of a veterinary practitioner or, alternatively, frozen preservation of semen may prove to be more practical and cost-efficient, whereby semen can be stored and dispatched sporadically as the demand arises. Clearly, use of frozen semen as an alternative is reliant on the stallion’s semen being suitable for freezing. Once the decision to establish a cooled semen program has been made, the following logistical considerations are important in ensuring the program works efficiently and successfully and is convenient for mare owners:
3. 4.
5.
1. Availability of semen. Will semen be collected at a regularly scheduled times each week, or on demand only? 2. Semen ordering. How and when will a breeding dose need to be ordered so as to facilitate a timely collection and dispatch of the dose? 3. Availability of freight services. Stallion owners need to be aware of the relative locations of the stud and potential broodmares and whether freight services are available on a same-day or overnight basis or on a weekend. 4. Alteration of freight services during holiday periods. 6. The responsibility for freight charges is often a contentious area if not clearly addressed and understood by stallion and mare owners prior to semen being dispatched. There can be large variations in freight costs, depending on location and speed of delivery, so, when a number of options are available, this should be considered in light of the effects of time on semen quality and subsequent fertility. In most cases, overnight (next day) delivery of semen is satisfactory for achieving good fertility, while minimizing freight costs. Stallion and mare owners should also consider that a 1- or 2-hour drive to the nearest airport to dispatch or receive semen may save hundreds of dollars in freight costs and avoid the delay of next-day delivery when compared with door-to-door freight services. The minimum equipment and facility requirements for a cooled semen program are: 1. A phantom or mare in estrus for the stallion to mount to facilitate semen collection. 2. The availability of a tease mare in estrus at any time semen collection is performed. An ovariectomized mare is often the most convenient option as she will be receptive to the stallion through-
7.
out the year. Regular administration of an estrogen conjugate may also be useful to improve receptiveness to stallions. Clean area for evaluation and processing of semen. Equipment for evaluation and processing semen (microscope suitable for semen evaluation with a warmed stage, waterbath, or incubator set at 37◦ C, device for accurately determining the concentration of spermatozoa, sterile and chemically clean containers for measuring, extending, and packaging semen). Semen extender. There are a number of commercially available semen extenders. Many are based on a simple skim-milk and sugar formulation1–3 with various antibiotics added. Others include clear multisugar extenders such as EquiPro (Minitub, Germany) and more complex extenders which contain milk protein fractions such as INRA-96. 4 When selecting an extender it should be noted that there may be quarantine restrictions regarding certain extender components of animal origin. An alternative to utilizing commercially available extenders in a large-scale cooled semen program is to prepare extenders from raw ingredients using published formulae. Clearly, this will require additional laboratory equipment and the ability to closely monitor and control the quality. However, it is this author’s experience that this may be a practical and cost-beneficial alternative to commercial extenders. Semen packaging. There are various semen packages that are commonly used, including plastic R bags (baby bottle liners or Whirl-pak bags), a variety of screw-top disposable sterile plastic vials and tubes, and latex-free syringes. It is important to note that each has differing cooling profiles and responses to changes in ambient temperature, which are dependent on the thickness and material of packaging, volume, and surface area. Thus, testing each for satisfactory performance under the local conditions is recommended. It is the author’s experience that the plastic bags are less practical in large-scale programs as they are unable to easily stand up in racks on benches or in incubators, making them awkward to fill. If volume and transport container limitations permit, inseminators appreciate the convenience of pre-filled syringes, and these also avoid a further handling step in transferring the semen to a syringe at the place of insemination. Appropriate, permanent labeling of semen packaging is important in maintaining the integrity of the breeding program and in fulfilling quarantine requirements. Semen transport containers. The number of different types of cooled semen transport containers available has increased concomitantly with the
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Experiences with a Large-Scale Cooled–Transported Semen Program growth in adoption of this technology. Ideally, containers should cool the semen at a rate of –0.05 to –0.1◦ C per minute from room temperature (20◦ C) to 5◦ C and then maintain the semen at 4–6◦ C for up to 36 hours.5–8 The containers all provide passive, uncontrolled cooling followed by a variable period of fluctuating temperature regulation. In short, none of the commercially available containers are ideal, but most are satisfactory in their performance when taking into account cost, size, R (Hamilton and weight factors. The Equitainer Research, Inc., South Hamilton, MA, USA) has been shown in a number of studies to perform better than the various disposable polyfoam containers in terms of cooling rate and resistance to changes in ambient temperature.9–13 However, the R have size, weight, and cost of the Equitainer meant that it is uncommonly used in commercial programs in Australia, although this container remains quite popular in the USA. In contrast, the various polyfoam boxes are now relatively inexpensive and their size and weight keep freight costs (charged on a weight–volume basis) to a minimum. A further consideration with semen transport containers, especially in light of their relatively cheap cost, is whether mare owners are required to return the containers. While return of the container by postal service is inexpensive, biosecurity issues need to be addressed, particularly in countries/areas where outbreaks of viral diseases such as equine influenza exist.
Quarantine considerations If stallion owners are considering international transport of semen they should thoroughly investigate quarantine and health requirements of the importing country well before the start of the breeding season. There are often very specific requirements regarding the location and construction of stallion housing and semen collection facilities and health testing of stallions which must be performed some time before semen collection can commence. It is important to remember that, once a quarantine facility has been established for a breeding season, the quarantine requirements and health standards need to be adhered to at all times, even when semen is being collected for domestic use only. Any breaches in quarantine protocols may compromise the facility for the remaining period of the season. The costs associated with satisfying quarantine requirements may be prohibitive if the international demand for semen is not great. An alternative is to collect and freeze semen at an approved quarantine facility outside the breeding season, and make frozen semen, rather than cooled se-
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men, available to breeders located in other countries. A single shipment of a number of doses of frozen semen to other countries at the start of the breeding season may prove to be cost efficient. As stated above, the ability of semen from a given stallion to retain good fertility following freezing and thawing will dictate whether or not this is a logical managerial approach.
Suitability of stallions Most, but not all, stallions have semen that is suitable for a cooled–transported semen program. Before the start of the breeding season a series of ejaculates should be evaluated to determine whether the stallion’s sperm production and quality will be sufficient to satisfy the maximum number of mares that may require semen during the season. It should be recognized that the quality of spermatozoa in an ejaculate is affected by the frequency of ejaculation. Stallions which have small numbers of mares sporadically requiring semen may need to have semen collected two or three times a week regardless of demand, in order to avoid accumulation of aged spermatozoa in the ejaculate and to maintain quality. A laboratory evaluation of semen extended and cooled in a range of extenders (containing various antibiotics) should be performed prior to the breeding season, after stallions have been depleted of their extragonadal storage reserves. Such an evaluation should ideally be performed using the packaging and containers that are intended to be utilized, and conducted over a 24to 36-hour period with samples warmed to 37◦ C and evaluated at least every 6 hours. When evaluating cooled semen as described above, the semen should be inverted gently to evenly distribute spermatozoa within the sample as they will settle while stored in a cooled state. A small sample should be quickly removed and warmed for evaluating while minimizing temperature fluctuations in the balance of the extended semen. In cases where a stallion has been identified as having poor quality cooled semen, there are some techniques which may facilitate utilization of the semen, albeit in a limited transported semen program. In some cases, it is the presence of even small amounts of the stallion’s own seminal plasma which may be deleterious to the quality of the semen. The motility and subsequent fertility of semen from these stallions may be improved by centrifugation and resuspension of the spermatozoa in an extender containing up to 10% of seminal plasma from a donor stallion with proven satisfactory fertility.14–17 Alternatively, the semen can be centrifuged through a density gradient to facilitate isolation of a population
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of spermatozoa with a higher proportion of morphologically normal, motile cells which can then be extended appropriately, cooled, and transported.18–22 These techniques require increased levels of expertise, additional equipment, and longer processing times and are likely to be performed only at largescale breeding or specialist veterinary centers.
Documentation All semen shipments should be accompanied by a document that, in the least, confirms the identification of the stallion and mare to be inseminated. It is also useful for the stud or semen-collection center to provide some information regarding the quality of the dose at the time of dispatch (percentage of motile spermatozoa, dose volume, concentration, total number of spermatozoa). In a practical sense, the most useful document is one that encourages the inseminator to evaluate the semen at the time of insemination, and, more importantly, to provide prompt feedback regarding the quality and subsequent fertility of the semen. It is the timely receipt of such information that will be the key to identifying and addressing problems in a cooled–transported semen program.
Semen handling to optimize the quality of cooled semen The handling of semen during collection, evaluation, and extending is the primary determinant of the success of a cooled semen program. The optimum procedures for collection, handling, and extending semen have been thoroughly addressed in other chapters of this text. However, some key points are worthy of emphasis. Temperature regulation during semen handling is critical in optimizing the quality of cooled semen. At all times, technicians should be aware of the likely temperature of the semen and ensure that any surfaces, environments, or extenders that the semen contacts are at a similar temperature so that sudden fluctuations in semen temperature are avoided. Semen extenders, by virtue of protective components and dilution effect, provide a practical buffer against temperature fluctuations. As such, following removal of a small aliquot for concentration and motility evaluation, raw semen should be diluted at least 1:1 (semen–extender) with warmed (30–37◦ C) extender as soon as practical after collection. A dilution rate of 1:1 will afford protection against temperature changes and immediate dilution of seminal plasma (see earlier chapters for discussions on effects of seminal plasma on spermatozoal function), while still en-
abling further dilution to create a breeding dose. Semen that has been extended in this manner can then be safely kept at room temperature while the semen is evaluated and the calculation of further dilution requirements is made. It is important to be aware though that the diluted semen will equilibrate with the ambient temperature, thus any further dilutions should use extender which has been kept at ambient temperature rather than in a heated incubator or waterbath. There is a large amount of literature which addresses the optimal dilution of stallion semen for cooled transport.17, 18,23–30 A practical distillation of this information has enabled the following ‘rules’ for extending stallion semen to be devised: 1. semen should be extended at least 1:3 (semen–extender); 2. concentration should be 25–50 million spermatozoa per ml; 3. insemination volume should be 20–100 mL; 4. insemination dose should contain at least 500 million PMS. Although the insemination dose may be reduced to as low as 100 million PMS if supported by previous acceptable fertility data. These rules have been successfully applied by the author and colleagues in large cooled–transported semen programs over a number of years. Some stallions may produce ejaculates in which the concentration of spermatozoa is too low (less than 100 million per mL) to enable optimum dilution. In these cases, the semen should be diluted 1:1 (semen–extender), centrifuged at 300–500g for 10–15 minutes and the sperm-dense pellet resuspended with extender so as to satisfy the ‘rules’ outlined above. A 1- to 5-mL ‘cushion’ comprising a relatively dense material (Cushion FluidTM or OptiPrepTM ) underlying the diluted semen allows the semen to be centrifuged at higher speeds (up to 1000 g) and improves the recovery of spermatozoa.20
Considerations for mare owners or managers In general, all mares are suitable for insemination with cooled–transported semen. However, it is clear that mares that have trouble conceiving with natural service or insemination with fresh semen will require more intensive management prior to and after insemination with cooled semen. In most cooled semen programs, there are limited (usually only three) breeding opportunities per week, or, alternatively, there is a delay of up to 2 days between when semen
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Experiences with a Large-Scale Cooled–Transported Semen Program is ordered and when it is received. Thus, there is a greater requirement for the appropriate management of mares in order to facilitate timely insemination. Prior to examining and preparing mares for insemination, it is important to ascertain from the stud or collection center the following: 1. When is semen available? 2. How long before shipment do orders need to be placed? 3. How will the semen be delivered and how long does it take? 4. What time of day will the semen be delivered and when is the veterinarian or insemination technician available? It is only after these logistical factors are known that mare owners can schedule veterinary examinations and the veterinarian consider the mare for insemination. One of the consequences of having broodmare farms without stallions is that there is an absence of any ‘teaser’ effect, which is useful both early in the breeding season to stimulate mares to start cycling and also throughout the season in accurately identifying mares in heat. Thus, the use of a non-breeding ‘teaser’ stallion or gelding is recommended in combination with diligent examination of mares to identify those that are sufficiently in estrus to inseminate and respond to ovulation-inducing drugs. The use of ovulation-inducing drugs is critical to the success of cooled–transported semen programs. Owing to the limited breeding opportunities each week, the sometimes limited fertile life of the spermatozoa and uncontrollable factors that affect quality and delivery of the semen, it is important that mares are ‘programmed’ to ovulate within 24–48 hours after insemination (see Chapter 128). A number of studies have addressed the optimum time to inseminate mares with cooled semen to maximize fertility.31–33 The author has found that, on large mare farms, ovulation-inducing drugs (human chorionic gonadotropin or deslorelin acetate) are ideally administered 24 hours prior to insemination. Such a program allows mares to be examined every second day, with insemination being performed in between. Ovulation occurs approximately 12–18 hours after insemination and prior to the next examination of the mare. Of course, administration of ovulationinducing drugs prior to receipt of the semen is not sensible when delivery or quality of the semen are unreliable. The success of any cooled–transported semen program, regardless of scale, is dependent on a coopera-
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tive relationship between the stud (semen collection station) and the inseminator. The inseminator has key responsibilities in appropriately handling and inseminating the mare(s) and in providing prompt feedback to the stud regarding the quality and fertility of the semen. As soon as practical after delivery, the semen should be deposited into the mare using accepted minimal contamination breeding techniques. There is no requirement to warm cooled semen prior to insemination. Indeed, inappropriate warming is likely to be detrimental to the spermatozoa. It is not uncommon for more than one breeding dose or an excessively large volume or number of spermatozoa to be received. In these cases (and when accompanying documentation confirms that the delivery comprises more than one dose), it is appropriate to inseminate the mare upon receipt and again 12–24 hours later. However, the second insemination should not be performed after ovulation, and should be avoided in mares identified as problem breeders. If ovulation is imminent then normally all the semen is deposited unless the volume exceeds 100 mL. However, on occasion the mare may have an unexpectedly hostile uterine environment (uterine fluid) and it may be prudent to flush the mare’s uterus prior to insemination34 and keep semen for re-insemination at later time. The importance of evaluating semen at the time of insemination has been stated earlier in this chapter. It is equally important for the inseminator or mare owner to arrange and report the result of a pregnancy test in a timely manner.
Troubleshooting problems in a cooled–transported semen program Unsatisfactory fertility Early recognition of a problem is reliant on the availability of semen evaluation and pregnancy test data from mare owners or inseminators. Regular ‘inhouse’ evaluations of surplus semen which has been extended and stored under field conditions is another useful approach to identifying problems. Commonly encountered problems in a cooled–transported semen program can be categorized as stallion associated, laboratory or handling associated, transport associated, or inseminator associated. Stallionassociated problems are readily identified by a cooled semen evaluation performed over 36 hours as described above. Such an evaluation will also identify problems originating from laboratory or semenhandling procedures such as inappropriate or compromised extenders.
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Stallion: Semen Collection, Evaluation and Artifical Insemination
Table 129.1 Common problem areas in a cooled–transported semen program Problem identification technique and management Stallion-associated problems
Perform a cooled semen evaluation Bacteriological examination of stallion’s genital tract and semen Ensure adequate washing of stallion’s penis
Laboratory or semen-handling problems
Artificial vagina temperature, lubricant, and washing procedure Confirm accuracy of sperm concentration measuring devices Ensure packaging and handling vessels are sterile and non-spermicidal Ensure semen temperature fluctuations are minimized during handling Confirm that dilution rate and breeding dose are satisfactory Evaluate the effects of various extenders of semen quality
Transport-associated problems
Small infrared temperature logging devices used in food industries are inexpensive tools for monitoring temperature fluctuations inside semen transport containers Confirm that selected freight options minimize transport time and avoid exposing transport containers to chillers, excessive heating, or irradiation The correct ice packs (gel versus water based) should be used within the transport container
Inseminator-associated problems
Ensure appropriate mare and ovulation induction management so that mares are inseminated once in a timely manner Ensure correct semen handling prior to insemination and use of non-spermicidal equipment and lubricants Ensure semen is inseminated into mare soon after delivery
Table 129.1 is a checklist of common problem areas in a cooled–transported semen program and means by which they might be identified and addressed.
Summary The following points are critical to the success of a cooled–transported semen program: 1. Preparation a Evaluate stallions, extenders, and transport systems b Determine ordering and freight logistics.
2. Communication – maintain good communication between mare and stallion managers. 3. Semen handling a Temperature regulation b Dilution rate, concentration, insemination volume, total number of progressively motile spermatozoa. 4. Mare management a Use ovulation-inducing drugs b Aim to inseminate mares once per cycle at an optimal time. 5. Results a Encourage inseminators to evaluate semen at the time of insemination and provide feedback b Determine pregnancy results as early as possible c Evaluate semen throughout the breeding season to identify potential problems.
References 1. Aurich JE, Aurich C. Developments in European horse breeding and consequences for veterinarians in equine reproduction. Reprod Domest Anim 2006;41:275–9. 2. Barrelet FE. The use of artificial insemination in European horse breeding inductries. Equine Vet Educ 1992;4:225–8. 3. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: technique and preliminary findings. In: Proceedings of the 21st Annual Convention of the American Association of Equine Practitioners, 1975, pp. 327–35. 4. Batellier F, Duchamp G, Vidament M, Arnaud G, Palmer E, Magistrini M. Delayed insemination is successful with a new extender for storing fresh equine semen at 15◦ C. Theriogenology 1998;50:229–36. 5. Kayser JP, Amann RP, Shideler RK, Squires EL, Jasko DJ, Pickett BW. Effect of linear cooling rate on motion characteristics of stallion spermatozoa. Theriogenology 1992;38:601–14. 6. Moran DM, Jasko DJ, Squires EL, Amann RP. Determination of temperature and cooling rate which induce cold shock in stallion spermatozoa. Theriogenology 1992;38:999–1012. 7. Province CA, Squires EL, Pickett BW, Amann RP. Cooling rates, storage temperature and fertility of extended equine spermatozoa. Theriogenology 1985;23:925–34. 8. Varner DD, Blanchard TL, Love CC, Garcia MC, Kenney RM. Effects of cooling rate and storage temperature on equine spermatozoal motility parameters. Theriogenology 1988;29:1043–54. 9. Brinsko SP, Rowan KR, Varner DD, Blanchard TL. Effect of transport container and ambient storage temperature on motion characteristics of equine spermatozoa. Theriogenology 2000;53:1641–55. 10. Douglas-Hamilton DH, Osol R, Osol G, Driscoll D, Noble H. A field study of the fertility of transported equine semen. Theriogenology 1984;22:291–304. 11. Katila T, Combes GB, Varner DD, Blanchard TL. Comparison of three containers used for the transport of cooled stallion semen. Theriogenology 1997;48:1085–92. 12. Malmgren L. Effectiveness of two systems for transporting equine semen. Theriogenology 1998;50:833–9.
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Experiences with a Large-Scale Cooled–Transported Semen Program 13. McKinnon AO, Walker JW. Effect of ambient temperature and container on temperature of extended equine semen. World Equine Vet Rev 1998;3:5–11. 14. Brinsko SP, Crockett EC, Squires EL. Effect of centrifugation and partial removal of seminal plasma on equine spermatozoal motility after cooling and storage. Theriogenology 2000;54:136. 15. Jasko DJ, Moran DM, Farlin ME, Squires EL. Effect of seminal plasma dilution or removal on spermatozoal motion characteristics of cooled stallion semen. Theriogenology 1991;35:1059–67. 16. Love CC, Brinsko SP, Rigby SL, Thompson JA, Blanchard TL, Varner DD. Relationship of seminal plasma level and extender type to sperm motility and DNA integrity. Theriogenology 2005;63:1584–91. 17. Rigby SL, Brinsko SP, Cochran M, Blanchard TL, Love CC, Varner DD. Interactions between seminal plasma and semen extender: effects on motility and fertility of cooled-stored equine spermatozoa. Anim Reprod Sci 2001;68:171–80. 18. Loomis PR. Advanced methods for handling and preparation of stallion semen. Vet Clin North Am Equine Pract 2006;22:663–76. 19. Morrell JM, Dalin AM, Rodriguez-Martinez H. Comparison of density gradient and single layer centrifugation of stallion spermatozoa: yield, motility and survival. Equine Vet J 2009;41:53–8. 20. Waite JA, Love CC, Brinsko SP, Teague SR, Salazar Jr. JL, Mancill S, Varner DD. Factors impacting equine sperm recovery rate and quality following cushioned centrifugation. Theriogenology 2008;70:704–14. 21. Sieme H, Martinsson G, Rauterberg H, Walter K, Aurich C, Petzoldt R, Klug E. Application of techniques for sperm selection in fresh and frozen-thawed stallion semen. Reprod Domest Anim 2003;38:134–40. 22. Varner DD, Love CC, Brinsko SP, Blanchard TL, Hartman DL, Bliss SB, Carroll BS, Eslch MC. Semen processing for the subfertile stallion. J Equine Vet Sci 2008;28:677–85. 23. Brinsko SP. Insemination doses: how low can we go? Theriogenology 2006;66:543–50.
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24. Gahne S, Ganheim A, Malmgren L. Effect of insemination dose on pregnancy rate in mares. Theriogenology 1994;49:1071–4. 25. Householder DD, Pickett BW, Voss JL, Olar TT. Effect of extender, number of spematozoa and hCG on equine fertility. J Equine Vet Sci 1981;1:9–13. 26. Jasko DJ, Martin JM, Squires EL. Effect of insemination volume and concentration of spermatozoa on embryo recovery in mares. Theriogenology 1992;37:1233–9. 27. Newcombe JR, Lichtwark S, Wilson MC. Case report: The effect of sperm number, concentration, and volume of insemination dose of chilled, stored, and transported semen on pregnancy rate in standardbred mares. J Equine Vet Sci 2005;25:70–5. 28. Sieme H, Bonk A, Hamann H, Klug E, Katila T. Effects of different artificial insemination techniques and sperm doses on fertility of normal mares and mares with abnormal reproductive history. Theriogenology 2004;62: 915–28. 29. Squires EL, Brubaker JK, McCue PM, Pickett BW. Effect of sperm number and frequency of insemination on fertility of mares inseminated with cooled semen. Theriogenology 1998;49:743–9. 30. Varner DD, Blanchard TL, Love CC, Garcia MC, Kenney RM. Effects of semen fractionation and dilution ratio on equine spermatozoal motility parameters. Theriogenology 1987;28:709–23. 31. Shore MD, Macpherson ML, Combes GB, Varner DD, Blanchard TL. Fertility comparison between breeding at 24 hours or at 24 and 48 hours after collection with cooled equine semen. Theriogenology 1998;50:693–8. 32. Stone R. Timing of mating in relation to ovulation to achieve maximum reproductive efficiency in the horse. Equine Vet Educ 1994;6:29–31. 33. Woods J, Bergfelt DR, Guillouet P. Effect of time of insemination relative to ovulation on pregnancy rate and embryonic loss in mares. Equine Vet J 1990;22:410–15. 34. Vanderwall DK, Woods GL. Effect on fertility of uterine lavage performed immediately prior to insemination in mares. J Am Vet Med Assoc 2003;222:1108–10.
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Chapter 130
Containers for Transport of Equine Semen Terttu Katila
Widespread use of transported cooled stallion semen has become possible because of the development of superior semen extenders and transport containers. Equine semen transport escalated in the early 1980s and in many horse breeds it is now more popular than natural mating or on-site insemination. The transport time is usually 12–24 hours, but can be extended to ≥ 48 hours for stallions with good quality semen that tolerates cooled storage. Containers travel in cars, buses, trains, and airplanes and need to withstand rough handling. All commercial containers available today have a passive cooling system based on fluid that is frozen in a freezer and then gradually warms up by the heat delivered from the semen and air within the storage chamber. The ambient temperatures to which the containers are exposed vary from freezing cold to tropical temperatures. Usable transport containers must be capable of cooling the extended semen at the desired rate and maintaining the intended storage temperature in extreme ambient temperatures. In addition to the size, structure, and material of the container and the coolant, the cooling rate depends also on the initial temperature and volume of the semen dose. Manufacturers of most containers provide information regarding the semen volume range that will produce the best results.
Storage temperature In the incubator (+37◦ C), sperm motility is lost rapidly, as reported by Varner et al.1 In the same study, temperatures of 25◦ C and 4◦ C resulted in similar sperm motility following a 24-hour storage period, but 4◦ C yielded higher sperm motility for longer storage periods. When temperatures of 20◦ C and 5◦ C were compared in a subsequent study, motility decreased more rapidly at 20◦ C by 24 hours of storage.2
Semen can be stored at room temperature (+20◦ C) for 12 hours without significant loss of motility or fertilizing capacity.3–5 This storage temperature is not commonly used because the semen transport time often exceeds 12 hours. At room temperature, excessive bacterial growth may deteriorate semen quality, particularly if the semen is contaminated with high numbers of bacteria. If semen of a particular stallion is very sensitive to cooling and if the transport time does not exceed 12 hours, semen can be kept at room temperature. There are no commercial semen shipment containers available, however, that maintain a storage temperature of 20◦ C. For internal storage temperatures less than 20◦ C, containers equipped with cooling systems are required. Reduced storage temperature decreases metabolic rate, slows down chemical reactions, and retards bacterial growth, resulting in an extension of the fertile life of spermatozoa.5 A storage temperature of 15◦ C has been shown to maintain motility for 24 hours, but in the study by Householder et al.6 sperm motility decreased more rapidly at 15◦ C than at 5◦ C. The optimal storage temperature depends also on the semen extender.3 Semen diluted with the chemically defined extender INRA 96 (IMV Technologies, L’Aigle, France) and stored at 15◦ C under an aerobic atmosphere for 24 hours yielded a higher per cycle pregnancy rate than semen diluted with Kenney’s extender and stored at 4◦ C under anaerobic conditions.7 Satisfactory pregnancy rates were achieved even after storage for 72 hours.7, 8 The latter study demonstrated that INRA 96 extender is as efficient at 15◦ C as at 4◦ C after 24 hours of storage. Since practically all transport containers have been designed to cool semen < 10◦ C, the possibility of also using INRA 96 at lower temperatures will increase its use as a semen extender. Today, skim-milk
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Containers for Transport of Equine Semen extenders are most common and extenders containing egg yolk are also used, but INRA 96 is gaining in popularity. Storage temperatures of 4–6◦ C are generally considered to be ideal for stallion semen storage.1, 2,9–11 Temperatures of 0◦ C and 2◦ C result in lowered sperm motility after 48 hours of storage.11 The semen containers should keep the temperature at 4–6◦ C or at least < 10◦ C for 24 hours, and preferably for 48 hours.
Cooling rate Stallion spermatozoa tolerate rapid cooling down to 20◦ C. The spermatozoa are most sensitive to cold shock between 19◦ C and 8◦ C.11 Cold shock is characterized by decreased and circular motility and acrosome and plasma membrane damage. Rapid cooling can lead to a phase separation of the lipid and protein components, resulting in an irreversible change of membranes.9 The lipoproteins in egg yolk and milk are thought to produce structural modifications in cell membranes, thereby stabilizing them and allowing adaptation of sperm to low temperatures.1 During the sensitive phase, sperm should be cooled at ≤ –0.1◦ C/min and preferably at −0.05◦ C/min. At temperatures of 8◦ C downwards sperm can again be cooled more rapidly without deleterious effects.9–11 The semen transport containers cool semen passively in a curvilinear fashion with semen cooling at the greatest rate initially. The transport containers have been designed to cool semen slowly. In the EquitainerTM (Hamilton Research, Inc., South Hamilton, MA, USA), the initial cooling rate is −0.3◦ C/min and this has been shown to maintain the fertilizing capacity of spermatozoa.9 In some other containers, the cooling rate is higher, but this does not necessarily lower sperm motility during 24 hours of storage; however, cooling rates exceeding −1◦ C/min or rates below −0.001◦ C/min have shown to result in low sperm motility after storage.9
Transport containers The increasing popularity of shipped semen has created a market for semen transport containers. A number of them have been introduced during the last two decades. More are still to come, since there seem to be several patents pending. Some transport containers are designed to be inexpensive and disposable, but are still often used several times. Others are costly and are meant to last for years. The EquitainerTM was the first transport container for stallion semen that
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had been extensively studied and tested before marketing.9
Reusable transport containers
Equitainers When the EquitainerTM was launched in the early 1980s, it had a shape of a 5-gallon (18.9 L) cylindrical bucket. It was reshaped later, and today three types of EquitainerTM containers are available commercially (Hamilton Research, Inc., South Hamilton, MA, USA). The EquitainerTM I is designed to maintain the semen at a cooled temperature for 70 hours, provides X-ray protection, and is heavier than the EquitainerTM II, which maintains cooled internal temperature for 48 hours (http://www.equitainer.com). The walls of the container are made of molded polyethylene and copolymer designed to withstand rough handling for years. In the cup model, the semen sample is stored in a plastic bag and surrounded by thermal ballast bags (each containing 60 mL aqueous fluid). The number of ballast bags depends on the volume of the semen dose, since the total volume of stored fluid should be 120–180 mL. Heat is transferred through thermal impedance into the frozen canned refrigerant (previously kept in the freezer for 24 hours), and the isothermalizer contents are cooled at a controlled rate. There is also a new isothermalizer that accommodates two centrifuge tubes (maximum volume 50 mL each). If only one centrifuge tube is needed for semen transport, the remaining tube is filled with water, which serves as a replacement for the ballast bags in the older cup models of isothermalizers. The EquitainerTM Clipper is a disposable container which is adapted for use with 50-mL or 20-mL syringes. In all studies comparing cooling rates and holding time of transport containers, the EquitainerTM has shown its superiority to simpler inexpensive containers (Figure 130.1).12–17 During transport in different environmental and weather conditions, containers can be exposed to temperatures much higher or lower than room temperature. Brinsko et al.15 kept seven different transport containers (i) at 22◦ C for 24 or 72 hours, (ii) at −20◦ C for 6 hours followed by 22◦ C for 18 or 66 hours, and (iii) at 37◦ C for 24 or 72 hours. Ambient temperatures affected temperatures inside all containers, including the EquitainerTM I and EquitainerTM II. None of the Equitainer models cooled samples < 10◦ C when the ambient temperature was maintained at 37◦ C. Ambient temperature affected motility characteristics of spermatozoa stored in all containers except for the EquitainerTM II, in which motion characteristics remained high and similar
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Stallion: Semen Collection, Evaluation and Artificial Insemination 40 Equitainer Equine Express Expecta Foal
35 Temperature (°C)
30 25 20 15 10 5 0 −5 0
1
2
3
4
5
6
7
9 10 15 20 25 30 35 40 45 50 55 60
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Storage Time (hours) Figure 130.1 Comparison of cooling/warming rates for 40-mL aliquots of semen extender in three different equine semen transport containers. Reprinted from Katila et al.13 , with permission from Elsevier.
among all ambient temperatures, even at 37◦ C when the storage temperature of semen was 10–15◦ C (Figure 130.2). McKinnon and Walker17 exposed five different transport containers to the following conditions: room temperature (22–25◦ C), sealed car in the sun (maximum temperature > 50◦ C), and refrigerator (5–0◦ C). The semen volume was 100 mL or 10 mL. In the refrigerator, after 10 hours of storage, the semen temperature in the Equitainer decreased to 2◦ C, which was maintained for the observation period of 35 hours (Figure 130.3). In the hot car, the temperature of 100 mL of stored semen increased to 17◦ C during the first day and to > 20◦ C during the second day. In 10 mL of semen, even higher
temperatures were recorded: 20◦ C and 30◦ C. Despite this high temperature, progressive motility of sperm was maintained well for 24 hours ranging from 50% (10 mL in the car) to 60% (10 mL in the room). Webb et al.16 shipped semen-transport containers by air in winter and in summer. Ambient temperatures and internal temperatures of cooling devices and the cup containing semen were recorded for 24 hours including ground transportation and two flights. EquitainerTM I with two types of isothermalizers, cup and tube type, was one of the containers tested. At 22◦ C, the cooling rate of semen from 19◦ C to 10◦ C in the cup was −0.077◦ C/min and the minimum temperature 6.2◦ C, and in the tube type −0.033◦ C/min and 7.6◦ C, respectively. The minimum
30 25 20
15 T°C
c130
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5
0
Equit-I Equit-II Equit-III Expecta Bio Flite Lane Express
0
5
10 Time (hours)
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Figure 130.2 Cooling curves of equine semen extender for the 24-hour period after samples reached 20◦ C in containers stored at 37◦ C. Reprinted from Brinsko et al.15 , with permission from Elsevier.
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40 Equitainer
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Bio Flite Lane
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Expecta foal NZ
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Ideal
20 15 10 5 0 0
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20
30
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−5 Time Figure 130.3 Effect of container on temperature of extended semen (100 mL) stored under different environmental conditions refrigerator. Reprinted with permission from McKinnon and Walker.17
internal temperatures of cooling devices were 10.9◦ C in summer and 3.3◦ C in winter (Table 130.1). Progressive motility of spermatozoa at 24 hours was > 40% in all experiments. The conditions in the study by Webb et al.16 mimicked conditions that semen containers face during transportation. It is unlikely that a shipping container would have to stay in a hot car for 2 days as in the study by McKinnon and Walker17 unless placed near the engine during a long bus journey or for 24 hours at 37◦ C, as was performed in the study by Brinsko et al.15 The fact that semen stored in the EquitainerTM preserved satisfactory motility even in these extreme conditions demonstrates that it is a dependable transport container. Good motility does not necessarily reflect good fertilizing capacity, but, on the other hand, numerous mares have been inseminated over more than two decades with semen stored in the EquitainerTM .
EST XLTM H, Equine Semen Transporter The EST XLTM H, Equine Semen Transporter is a durable, double-walled polyethylene container for multicycle use. It accommodates two polyethylene jars for bagged semen or two 50- or 60-mL syringes. In both room winter and summer ambient temperatures, the EST XL H showed similar cooling curves to the EquitainerTM over 48 hours (http://www. barakafarm.com/est). No controlled studies with this container have been published.
Disposable containers The reusable EquitainerTM models are relatively expensive, and are rather big and heavy, thereby increasing transport costs. Because they are valuable, they must be returned back to the semen collection station, which means additional transport costs.
Table 130.1 Influence of ambient temperature (◦ C) during transport on internal temperature of commercial cooling devices; data are presented as means ± standard deviations. Reprinted from Webb et al.16 , with permission from Elsevier. Minimum internal temperature
Ambient temperature
Item
Equine Expressa
Equitainer Ib
Minimum
Maximum
Summer shipment Winter shipment
∗
∗
25.7 ± 4.3 5.7 ± 1.6
37.8 ± 6.7 25.3 ± 1.9
a b
†
10.0 ± 0.3 2.7 ± 0.3
†
10.9 ± 0.9 3.3 ± 0.3
Express Breeders Supply (York, PA). Hamilton Research Inc. (Beverly, MA). Means in the same column with different superscripts are different (P = 0.001).
∗†
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Therefore, less expensive, disposable transport containers have been developed.
(http://www.minitube.de). According to informa¨ Neopor boxes sell better than stytion from Minitub, rofoam boxes.
Equine Express and similar containers There are several disposable semen transport containers commercially available. Some of the styrofoam boxes are very similar in appearance, making it difficult to determine how they differ from each other. In the early 1990s, a Swedish stud farm introduced a disposable styrofoam box, designed originally to transport human samples, for stallion semen shipments. The container was termed the ‘Salsbro’ box, after the stud farm, but was reacquired and renamed the Equine ExpressTM .13, 14 This original styrofoam box accommodated two 20-mL syringes. A thin styrofoam plate was placed between the syringes and a rectangular frozen coolant. This container functioned well at room temperature. The cooling rate was similar to that of the EquitainerTM , and a temperature of 7–8◦ C was maintained for 24 hours at room temperature (Figure 130.1).13, 14, 16 Malmgren14 kept transport containers for 24 hours also in an incubator (37◦ C). There the temperature was significantly higher in the Equine ExpressTM (19.3◦ C) than in the EquitainerTM (12.2◦ C) at 24 hours. In both studies, sperm motility was similar in both containers at room temperature, but at 37◦ C sperm quality was significantly better in the EquitainerTM than in the Equine ExpressTM . Neither season nor container affected the percentage of progressively motile spermatozoa at 24 hours in the experiments by Webb et al.16 In the study by Brinsko et al.,15 the Equine ExpressTM kept in the freezer reached the lowest temperature of −1◦ C at 7 hours; under room temperature the lowest temperature of 7◦ C was recorded at 10 hours; and in the incubator 9◦ C at 13 hours. The temperature was maintained for 22 hours at < 10◦ C in the freezer, for 20 hours at room ambient temperature, and for 10 hours in the incubator (Figure 130.2). Total and progressive sperm motility were similar for semen stored in the Equine ExpressTM and in EquitainerTM models, except for freezer temperature, where semen in the Equine ExpressTM exhibited lower motility. This can be explained by the many hours of semen storage at < 4◦ C in the Equine ExpressTM (8 hours < 0◦ C, 12 hours < 2◦ C, 14 hours < 4◦ C), since this was shown to correlate with a reduction in spermatozoal motility. The Equine ExpressTM II is currently available with shipping capacities for 100 mL or 120 mL of extended semen. A Neopor box contains two ice packs and two syringes separated from each other by a Neopor ¨ Tiefenbach, Germany). This container wall (Minitub, cools semen similar to that of a styrofoam box for 35 hours, but a temperature < 0◦ C is preserved longer
NZ Semen Shipper McKinnon and Walker17 also tested this container under the three conditions. In the refrigerator, the temperature of 10 mL of semen was around 0◦ C during the observation period and in 100 mL of semen slightly above 0◦ C (Figure 130.3). In the room, the temperature was maintained between 10◦ C and 15◦ C. In the sealed car, the temperature rose to 15–20◦ C during the first hot day and to 30–40◦ C during the second day. As such, the progressive motility of spermatozoa was very low at 24 hours: 12% in 100 mL and 0% in 10 mL. Under the other two conditions, acceptable sperm motility was detected.
South American containers In one study, extended equine semen was stored in the Max-Semen ExpressTM (Agrofarma, Brazil) and the EquitainerTM at room ambient temperature for 24 hours and then the semen was subjected to cryopreservation.18 Motility of sperm stored in the EquitainerTM was higher than that of sperm stored in the Max-Semen ExpressTM , but plasma membrane integrity did not differ. The temperature inside the Max-Semen ExpressTM was higher (16◦ C) than in the EquitainerTM (6◦ C). Avanzi et al.19 studied four containers (EquitainerTM , BotutainerTM , Max-Semen ExpressTM , and BotuboxTM ), at ambient temperatures of 24◦ C and 40◦ C for 24 hours. At 24◦ C, the temperatures inside the Max Semen ExpressTM and the BotuboxTM were maintained at 15◦ C, whereas in the EquitainerTM and the BotutainerTM the internal temperatures were close to 5◦ C; no differences were detected in sperm motility. At 40◦ C, the temperatures in all containers started to rise after 6 hours, ending up between 10◦ C and 15◦ C at 24 hours in the EquitainerTM and the BotutainerTM , but in the BotuboxTM and Max-Semen ExpressTM the temperatures exceeded 25◦ C. This high temperature negatively affected both sperm motility and plasma membrane integrity at 24 hours, but not before. In the hot South American climate, the BotuboxTM and Max Semen ExpressTM can be used for short journeys, but for journeys ≥ 24 hours selection of the EquitainerTM or BotutainerTM is recommended.
Conclusions A variety of containers are available for cooled transport of stallion semen, and all can provide suitable
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Containers for Transport of Equine Semen results under selected conditions of storage. Persons wishing to pursue shipment of cooled semen are encouraged to examine the results of controlled trials with the available products, or to test semen under conditions appropriate for their veterinary practice before making a final decision on container selection. At many facilities, more than one type of container is used, to accommodate the varied needs of the facility.
References 1. Varner DD, Blanchard TL, Love CL, Garcia MC, Kenney RM. Effects of cooling rate and storage temperature on equine spermatozoal motility parameters. Theriogenology 1988;29:1043–54. 2. Varner DD, Blanchard TL, Meyers PJ, Meyers SA. Fertilizing capacity of equine spermatozoa stored for 24 hours at 5 or 20◦ C. Theriogenology 1989;32:515–25. 3. Province CA, Squires EL, Pickett BW, Amann RP. Cooling rates, storage temperatures and fertility of extended equine spermatozoa. Theriogenology 1985;23:925–34. 4. Francl AT, Amann RP, Squires EL, Pickett BW. Motility and fertility of equine spermatozoa in a milk extender after 12 or 24 hours at 20◦ C. Theriogenology 1987;27:517– 25. 5. Pickett BW, Amann RP. Extension and storage of stallion spermatozoa: a review. J Equine Vet Sci 1987;7:289– 302. 6. Householder DD, Pickett BW, Voss JL, Olar TT. Effect of extender, number of spermatozoa and hCG on equine fertility. J Equine Vet Sci 1981;1:9–13. 7. Batellier F, Duchamp G, Vidament M, Arnaud G, Palmer E, Magistrini M. Delayed insemination is successful with a new extended for storing fresh equine semen at 15◦ C under aerobic conditions. Theriogenology 1998;50:229– 36. 8. Batellier F, Vidament M, Fauguant J, Duchamp G, Arnaud G, Yvon JM, Magistrini M. Advances in cooled semen technology. Anim Reprod Sci 2001;68:181–90.
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9. Douglas-Hamilton DH, Osol R, Osol G, Driscoll D, Noble H. A field study of the fertility of transported equine semen. Theriogenology 1984;22:291–304. 10. Kayser JP, Amann RP, Shideler RK, Squires EL, Jasko DJ, Pickett BW. Effects of linear cooling rate on motion characteristics of stallion spermatozoa. Theriogenology 1992;38: 601–14. 11. Moran DM, Jasko DJ, Squires EL, Amann RP. Determination of temperature and cooling rate which induce cold shock in stallion spermatozoa. Theriogenology 1992;38: 999–1012. ¨ 12. Hueck C. Untersuchungen zur Flussigkonservierung von ¨ Pferdesperma unter Verwendung verschiedener Kuhlund Transportstysteme – Laborstudie [Laboratory studies on extended equine semen storage using different cooling and transport systems]. Dissertation, Tier¨arztliche Hochschule Hannover, 1990. 13. Katila T, Combes GB, Varner DD, Blanchard TL. Comparison of three containers used for the transport of cooled stallion semen. Theriogenology 1997;48:1085–92. 14. Malmgren L. Effectiveness of two systems for transporting equine semen. Theriogenology 1998;50:833–9. 15. Brinsko SP, Rowan KR, Varner DD, Blanchard TL. Effects of transport container and ambient storage temperature on motion characteristics of equine spermatozoa. Theriogenology 2000;53:1641–55. 16. Webb GW, Arns PMJ, Harris MA, Dekat CL. Technical note: Comparison of two containers used for shipment of stallion semen. Prof Anim Sci 2005;21:1–5. 17. McKinnon AO, Walker JB. Effect of ambient temperature and container on temperature of extended equine semen. World Equine Vet Rev 1998;3:5–11. 18. Melo CM, Alvarenga MA, Zahn FS, Martin I, Orlandi C, Trinque CLA, Dell’Aqua Jr JA, Papa FO. Effect of transport container and cryoprotectant on freezability of equine semen previously cooled for 24 h. Anim Reprod Sci 2006; 94:78–81. 19. Avanzi BR, Farr´as MC, Melo CM, Alvarenga MA, ´ GHM, Papa FO. EfDell’Aqua Jr JA, Medeiros ASL, Araujo ficiency of different cooling and storage systems for maintaining equine semen viability in a hot environment. Anim Reprod Sci 2006;94:152–4.
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Chapter 131
Semen Extenders for Cooled Semen (Europe) Christine Aurich
Introduction With the current semen-processing techniques, fertility of cooled stallion semen can generally be maintained no longer than 24–48 hours. After that time, pregnancy rates decrease dramatically. Despite the fact that new extenders are regularly advertised claiming to maintain fertility for longer time periods, an extender and a semen-processing protocol that provide a generally longer lifespan of spermatozoa have not been developed to date. During cooled storage, longevity of spermatozoal viability is influenced by extender composition, storage temperature, cooling rate, oxygen exposure, presence of bacteria, presence and type of antibiotics in the extender, and the concentration of seminal plasma. Consequently, the extender itself and the semen-processing protocol can have an impact on the result of semen preservation. Besides motility, the main factors that have to be preserved are the integrity of spermatozoal plasma membrane, the mitochondria, and the sperm chromatin (i.e., DNA and associated proteins). This is not possible without addition of semen extenders which have to fulfill at least the following requirements: osmolarity similar to or slightly higher than the semen (300–350 mOsm/L); pH that is slightly acidic (6.8) to neutral (7.0); components that provide for stabilization of pH and control of growth of microorganisms; a readily available energy source; properties which provide for stabilization of membranes and metabolic function of spermatozoa; properties which provide neutralization of metabolic substances; and, finally, must not interfere with semen evaluation results.
Osmolarity and pH of semen extenders Osmolarity of equine semen is approximately 300 mOsm/L. Equine spermatozoa tolerate a relatively wide variation in osmolarity; therefore, osmolarity of semen extenders can vary between 250 and 400 mOsm/L without causing significant problems. Slight hyperosmolarity of the extender appears to be beneficial for the preservation of semen function; best results of semen longevity are noted with an osmolarity of around 350 mOsm/L. The pH of stallion ejaculates physiologically range between 6.8 and 7. A pH of semen extenders in this range is tolerated without detrimental effects. Semen extenders should buffer the pH of processed semen in response to the production of metabolic substances by spermatozoa or contaminating bacteria. Main buffer substances added to extenders for equine semen are sodium bicarbonate (NaHCO3 ), sodium citrate, and HEPES (2-hydroxy-ethyl-1-piperazine ethane sulfonic acid). The efficiency of buffer substances is dependent on the environmental pH, with every buffer substance having an optimal range of pH. The most efficient pH of sodium bicarbonate (pH 6.8–7.2), HEPES (pH 7.0–8.0), and even citrate (around 8) is closest to the pH seen in equine semen during storage. Consequently, these buffers are successfully used in a variety of extenders. From a biochemical view, the successful use of EDTA (ethylenediamine tetraacetic acid) with equine semen is questionable, as the optimal buffering capacity of this product is in the range of 4–5. Milk products in extender also provide some buffering capability.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Semen Extenders for Cooled Semen (Europe)
Composition of semen extenders Extenders with a composition similar to the original formula containing non-fat dried skimmed milk and glucose (NFDSMG), as published by Kenney et al.,1 are the most popular and are used worldwide. They are inexpensive, easy to formulate, can be stored in frozen form, and result in good fertility rates. The composition of INRA-82, an extender of French origin,2 differs from NFDSMG in the aspect that, besides glucose, lactose and raffinose have been added as additional sugars, and a more complex buffer system, consisting of HEPES, potassium citrate, and sodium citrate, is used. A fractionated portion of milk, native phosphocaseinate, is also the only milk ingredient in this extender. All milk-based extenders can be used for centrifugation and storage and maintain semen motility, membrane integrity, and fertility for 24–48 hours at an ambient temperature of 4–6◦ C. Extenders based on egg yolk provide results comparable to milk-based extenders but are more complicated to process and do generally not result in better semen quality or fertility.3 The most popular egg yolk-based extender in Europe is a glycine–egg yolk extender (Dimitropoulos-extender D11).4 The nature of the positive effects of milk or egg yolk on longevity of spermatozoal viability following cooled storage is still more or less unknown. Egg yolk components seem to stabilize the sperm plasma membrane by neutralizing detrimental effects of extension and cooling. In ejaculated bull semen, seminal plasma proteins stimulate cholesterol and phospholipid efflux from the sperm membrane and thus facilitate premature capacitation that results in de-
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creased longevity and fertility.5 Components of semen extenders apparently delay this process. Similar mechanisms may also exist in stallion semen. Milkbased extenders also provide membrane protective activity; in addition, a positive effect on the antioxidative properties of equine semen could be demonstrated.6 Recently, more defined extender media have been developed to diminish detrimental components that might be included in complex biological substances, such as milk or egg yolk. Another aim is the development of extender media with a defined and constant composition.7 The French extender INRA-96 (IMV, L’Aigle, France) is a milk product-based extender containing phosphocaseinates only in the milk source. This extender provides optimal longevity of equine spermatozoal viability at 15◦ C,2 but it can also be used at 5◦ C. Storage at higher temperatures usually requires aerobic storage conditions, while at 5◦ C anaerobic conditions are recommended owing to the decreased metabolic activity of spermatozoa. ¨ Tiefenbach, Germany), anWhen EquiPro (Minitub, other milk-based extender with a closely related composition to INRA-96,8 was tested for cooled storage of equine semen at 5◦ C and 15◦ C under anaerobic or aerobic conditions, storage at 5◦ C tended to maintain better semen quality irrespective of oxygen exposure (Figure 131.1).9 A storage temperature of 15◦ C has not become popular because storage temperature has to be maintained during shipment. This requires controlled cooling and heating systems. These are substantiallymore expensive than transport containers designed to store semen at approximately 5◦ C.
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If insemination is performed within 12 hours after collection, semen can be stored at room temperature (20–24◦ C) after appropriate addition of extender medium.10, 11 The use of semen stored at room temperature with insemination occurring within a few hours after collection may be of special interest in the case of stallions with poor semen quality after cooling. Cooling of spermatozoa from 18◦ C to 8◦ C is a critical step and may result in ‘cold shock’, a feature indicative of specific damage of the sperm plasma membrane.12 Substitution of egg yolk by soybean lecithin has been performed with limited success. A soybeanbased extender is commercially available for the ¨ Tiefenbach, Germany). horse (AndroMed E, Minitub, However, the quality of cooled stored semen processed with AndroMed E is lower than semen processed with milk-based extenders if storage time exceeds 24 hours.13 Because high concentrations of seminal plasma in cooled stored stallion semen are detrimental to motility and fertility,14 centrifugation of semen has become a routine method for semen processing in most artificial insemination centers. Especially in ‘poor-cooler’ stallions, centrifugation and partial removal of seminal plasma result in an increase in the percentage of progressively motile spermatozoa in cooled stored semen.15 The adverse effects of seminal plasma on cooled spermatozoa may be partially attributed to endogenous lipase activity.16 However, when semen is extended with NFDSMG extender, residual seminal plasma (approximately 5–20%) is of importance for maintaining semen longevity.14 Especially in stallions with consistently poor post-cooling semen motility, the use of a modified Tyrode’s medium17 for resuspension of centrifuged semen may result in improved quality of cooled stored semen. In this case, complete removal of seminal plasma is recommended.18 A major problem of centrifugation is the loss of approximately 25% of spermatozoa in the supernatant.19 Higher centrifugation force, together with better recovery rates, has become possible by the introduction of a so-called ‘cushion’ centrifugation technique. This technique can be performed using INRA-96 extender in combination with iodixanol-based cushion medium.20
Control of bacterial growth Microorganisms may negatively affect semen longevity during storage.21 Contamination of semen during collection cannot be completely avoided,22 but extenders should be free of any bacteria. Bacteria in semen extender can originate from milk or egg yolk ingredients or from contamination of the extender during production, storage, or from its
addition to semen. The hygienic status of all extender components should be carefully considered during production and handling. Companies providing commercial extenders routinely test for bacterial contamination of these products; however, the handling of the extender itself is of prime importance. Extender bottles, once opened or warmed to body temperature, should not be stored for longer time periods; rather, these products should be used immediately or discarded. Waterbaths are a well-known source of bacteria, such as Pseudomonas aeruginosa.23 Handling of extender bottles stored in waterbaths has to be done with great caution to avoid any contamination of the extender and waterbaths should be frequently drained and dried. In most extenders, growth of microorganisms is routinely controlled by addition of antibiotics.10, 22 In Europe, penicillin and gentamicin are widely used. In contrast, amikacin, penicillin, and ticarcillin/clavulonic acid are often used in the USA. Amikacin has been usually reserved for use in human medicine, and because it is more expensive than gentamicin in Europe and has a similar spectrum of action it is not considered advantageous in Europe. A combination of penicillin and gentamicin effectively inhibits growth of many microorganisms found in processed equine semen under in vitro conditions and the combination has low spermatotoxic effects. However, the final concentration of antibiotics in extender medium is of importance to avoid any negative effects on semen longevity. Maximally tolerated concentrations seem to be related to the composition of the extender because gentamicin, at a concentration of 1 mg/mL, did not negatively affect semen longevity in Kenney extender,10 but was detrimental to semen longevity in another milk-based semen extender, EquiPro.21 After reduction of the gentamicin concentration to the still effective concentration of 250 µg/mL, no deleterious effects on semen quality were detected (Figure 131.2). Altogether, addition of antibiotics should be viewed critically. Exposing mares to antibiotics at insemination is a type of antibiotic exposure that may be an important factor in the development of antibiotic-resistant bacteria.24 Because antibiotics may be effective only at a temperature above 15◦ C, they may only exert their effects during cooling from collection to storage temperature, e.g., for a time period of only 3–6 hours. After reaching the storage temperature, no further effects can be expected. On the other hand, bacterial growth at 5◦ C is very low.9 It has to be borne in mind that many antibiotics do not kill bacteria but only inhibit further growth. Therefore, direct effects of existing bacteria or their toxins cannot be avoided, even in the presence of antibiotics.21 Irrespective of the use of antibiotics, optimal
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slightly increased maintenance of semen quality following cooled storage.28, 29
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Time of storage at 5 °C (h) Figure 131.2 Total motility of spermatozoa in EquiPro ex¨ tender (Minitub) with (closed symbols) and without (open symbols) addition of gentamicin (1 g/L) in the absence (control, circles) or presence (squares) of Pseudomonas aeruginosa (108 /mL) during cooled-storage of stallion semen at 5◦ C (Aurich and Spergser 2007, modified). ∗ Significant difference between groups with Ps. aeruginosa in comparison to control without bacteria (P < 0.05).
hygienic measures during semen collection and processing are important to keep bacterial contamination as low as possible.
Neutralization of metabolic substances Owing to its high content of unsaturated fatty acids, the sperm plasma membrane is highly sensitive to oxidative stress. In collected ejaculates and processed semen, reactive oxygen species (ROS) are mainly produced by leukocytes and spermatozoa with excess residual cytoplasm (e.g., spermatozoa with cytoplasmic droplets).25 In ejaculates from healthy fertile stallions, leukocytes are rarely found. Mature spermatozoa may also produce intracellular ROS as a result of flagellar activity.26 Excessive peroxidation will damage the sperm plasma membrane and result in loss of motility and fertilizing capacity of semen.25 As the mature spermatozoon has lost the majority of its cell organelles and DNA transcription has ceased, repair mechanisms are not available and damage to the sperm plasma membrane can result in irreversible loss of its functions.27 Semen extenders, therefore, should also possess the ability to neutralize metabolic substances that may result in damage of spermatozoa during storage. Dilution of semen with a milkbased extender alone results in a significant increase in the antioxidative capacity of extended semen, possibly due to a positive interaction between seminal plasma and extender.6 Addition of antioxidants, such as ascorbic acid or pyruvate, have been reported to
1. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: techniques and preliminary findings. In: Proceedings of the 21st Annual Convention of the American Association of Equine Practitioners, 1975; p. 327. 2. Battelier F, Duchamp G, Vidament M, Arnaud G, Palmer E, Magistrini M. Delayed insemination is successful with a ◦ new extender for storing fresh equine semen at 15 C under aerobic conditions. Theriogenology 1998;50, 229–36. ¨ 3. Malmgren L, Op den Kamp B, Wockener A, Boyle M, Colenbrander B. Motility, velocity and acrosome integrity of equine spermatozoa stored under different conditions. Reprod Dom Anim. 1994;29:469–476. ¨ 4. Wockener A, Colenbrander B. Liquid storage and freezing of semen from New Forest and Welsh Pony stallions. Dtsch Tier¨arztl Wschr 1993;100:125–6. 5. Bergeron A, Crete M-H, Brindle Y, Manjunath P. Lowdensity lipoprotein fraction from hen’s egg yolk decreases the binding of the major proteins from bovine seminal plasma to sperm and prevents lipid efflux from the sperm membrane. Biol Reprod 2004;70:708–17. 6. Kankofer M, Kolm G, Aurich JE, Aurich C. Activity of glutathione peroxidase, superoxide dismutase and catalase and lipid peroxidation intensity in stallion semen during storage at 5 ◦ C. Theriogenology 2005;63:1354–65. 7. Aurich C. Factors affecting the plasma membrane function of cooled-stored stallion spermatozoa. Anim Reprod Sci 2005;89:65–75. ¨ ¨ 8. Pagl R, Aurich JE, Muller-Schl osser F, Kankofer M, Aurich C. Comparison of an extender containing defined milk protein fractions with a skim milk-based extender for storage of equine semen at 5 degrees C. Theriogenology 2006;66:1115–22. 9. Price S, Aurich J, Davies-Morel M, Aurich C. Effects of oxygen exposure and gentamicin on stallion semen stored at 5 ◦ C and 15 ◦ C. Reprod Domest Anim 2008;43:261–266. 10. Varner DD, Scanlan CM, Thompson JA, Brumbaugh GW, Blanchard TL, Carlton TM, Johnson L. Bacteriology of preserved stallion semen and antibiotics in semen extender. Theriogenology 1998;50:559–73. 11. Love CC, Thompson JA, Lowry VK, Varner DD. Effect of storage time and temperature on stallion sperm DNA and fertility. Theriogenology 2002;57:1135–42. 12. Amann RP, Pickett BW. Principles of cryopreservation and a review of cryopreservation of stallion spermatozoa. J Equine Vet Sci 1987;7:145–73. 13. Aurich C, Seeber P, Mueller-Schloesser F. Comparison of different extenders with defined protein composition for storage of stallion spermatozoa. Reprod Domest Anim 2007;42:445–8. 14. Jasko DJ, Hathaway JA, Schaltenbrand VL, Simper WD, Squires EL. Effect of seminal plasma and egg yolk on motion characteristics of cooled stallion spermatozoa. Theriogenology 1992;37:1241–52. 15. Brinsko SP, Crockett EC, Squires EL. Effect of centrifugation and partial removal of seminal plasma on equine spermatozoa motility after cooling and storage. Theriogenology 2000;54:129–36.
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16. Carver DA, Ball BA. Lipase activity in stallion seminal plasma and the effect of lipase on stallion spermatozoa during storage at 5 ◦ C. Theriogenology 2002;58:1587–95. 17. Padilla AW, Foote RH. Extender and centrifugation effects on the motility patterns of slow-cooled stallion spermatozoa. J Anim Sci 1991;69:3308–13. 18. Rigby SL, Brinsko SP, Cochran M, Blanchard TL, Love CC, Varner DD. Advances in cooled-semen technologies: seminal plasma and semen extender. Anim Reprod Sci 2001;68:171–80. 19. Loomis PR. Advanced methods for handling and preparation of stallion semen. Vet Clin Equine 2006;22:663–76. 20. Waite JA, Love CC, Brinsko SP, Teague SR, Salazer JL Jr, Mancill SS, Varner DD. Factors impacting equine sperm recovery rate and quality following cushioned centrifugation. Theriogenology 2008;70:704–14. 21. Aurich C, Spergser J. Influence of bacteria and gentamicin on cooled-storage of stallion spermatozoa. Theriogenology 2007;67:912–18. 22. Cl´ement F, Vidament M, Gu´erin B. Microbial contamination of stallion semen. Biol Reprod Monogr 1995;1:779– 786.
23. Kenney RM, Cummings MR, Zierdt CH, Umphenour N. Pseudomonas aeruginosa – somatic typing of genital tract isolates and colonization of the stallion penis: significance, diagnosis, and treatment. In: Proceedings of the 38th Annual Convention of the American Association of Equine Practitioners, 1992, pp. 601–8. 24. Denver LA, Dermody TS. Mechanisms of bacterial resistance to antibiotics. Ann Intern Med 1991;151:886–95. 25. Aitken RJ, Baker MA. Oxidative stress and male reproductive biology. Reprod Fertil Dev 2004;16:581–8. 26. Gavella M, Lipovac V. NADH-dependent oxidoreductase (diaphorase) activity and isozyme pattern of sperm in infertile men. Arch Androl 1992;28:135–41. 27. Eddy EM, O’Brien DA. The spermatozoon. In: Knobil E, Neill JD (eds) The Physiology of Reproduction. New York: Raven Press, 1994; pp. 29–77. ¨ 28. Aurich JE, Schonherr U, Hoppe H, Aurich C. Effects of antioxidants on motility and membrane integrity of chilledstored stallion semen. Theriogenology 1997;48:185–92. 29. Bruemmer JE, Coy RC, Squires EL, Graham JK. Effect of pyruvate on the function of stallion spermatozoa stored for up to 48 hours. J Anim Sci 2002;80:12–18.
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Chapter 132
Semen Extenders for Cooled Semen (North America) Steven P. Brinsko
Breeding mares with cooled semen has become common practice and, except for Thoroughbreds, is widely accepted by breed registries in North America. In order to optimize the ability of stallion semen to withstand the rigors of cooling and storage, it must be properly diluted in an appropriate semen extender. Suitably formulated semen extenders contain ingredients that protect sperm from cold shock, provide metabolizable substrate(s), and inhibit or eliminate bacterial growth, while maintaining a pH of 6.7–7.2 and an osmotic pressure between 300 and 400 mOsmol/L. Over the years, many extender formulations have been developed for use with cooled equine semen, including base ingredients ranging from cream and gelatin to egg yolk to heated skim milk to non-fat dried milk solids. While some of these extenders were suitable for semen preservation, the nature of their formulations made preparation and handling cumbersome and accurate evaluation of sperm motility problematic. In 1975, the formulation of a non-fat dried milk solids–glucose extender (NFDMS-GLU), also known as non-fat dried skim milk–glucose (NFDSM-GLU) or skim milk–glucose (SKMG) extender, was published by Kenney et al.1 This convenient, reliable semen extender dramatically increased the use of artificial insemination in horses. Initially, this extender, and most others, was made from scratch at individual laboratories and critical attention had to be paid to water quality, pH, and osmolarity. Many equine semen extenders based on the NFDSM-GLC formulation, also known as ‘Kenney extenders,’ are now available commercially in North America (Figure 132.1). Most come in powdered form and a measured volume of distilled water is supplied to reconstitute the extender. The major differences among these commercial preparations are types of
antimicrobials offered or the inclusion of additional sugars such as sucrose. Most commercially available equine semen extenders are offered with or without antibiotics, so that the most appropriate antibiotic(s) for semen of a particular stallion can be included in the extender. Ideally, the optimal antibiotic in a semen extender eliminates all microbial growth in the stored semen while maintaining sperm quality. Texas workers found that, of seven antibiotics tested in coolstored semen, the combination of amikacin sulfate (1 mg/mL) and potassium penicillin G (1000 U/mL) was the most effective for controlling bacterial growth while optimizing sperm motion characteristics.2 It must be emphasized that, for certain antibiotics, for example amikacin sulfate and gentamicin sulfate, only reagent-grade products should be used because the preservatives present in injectable preparations are toxic to sperm. The equine semen extenders used most commonly in the USA are milk-based formulations. The composition of milk, however, is complex, with some components being beneficial to sperm survival and others being detrimental.3 French workers have determined that native phosphocaseinate is the milk component offering the most protective effects on cool-stored equine sperm.3 This and similar studies led to the development of an extender composed of Hank’s salts supplemented with HEPES, glucose, lactose, and native phosphocaseinate (HGLL-NPPC), R marketed as INRA 96 semen extender (IMV Technologies, L’Aigle Cedex, France), which is widely used in North America and Europe. However, the antibiotic levels in INRA 96 are insufficient to control bacterial growth in ejaculates of some stallions. Work in our laboratory indicates that supplementation of INRA 96 with TimentinTM (ticarcillin sodium
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 132.1 Common extenders used in North America for cool-transported equine semen. Left to right, Next Generation, INRA-96, EZ-Mixin.
plus clavulonic acid; 1 mg/mL) aids in controlling bacterial growth without impairing osmolality or pH of the extender (D.D. Varner, personal communication). A different group of investigators reported that another defined milk protein extender, stated to be R ¨ Tiefenbach, similar to INRA 96 (EquiPro ; Minitub, Germany), was superior to skim milk extender for preserving cooled semen.4 The exact composition of this extender has not been published. High levels of seminal plasma are detrimental to cool-stored equine semen.5 In order to reduce the adverse effects that seminal plasma exerts on cooled stallion semen, the semen can be diluted with at least four parts of extender to one part semen such that the level of seminal plasma does not exceed 20% by volume, and the final sperm concentration remains above 25 × 106 sperm/mL.6, 7 When milk-based extenders are used for cool-stored equine semen, some seminal plasma must be present to maintain semen quality.8 The minimum level of seminal plasma necessary to maintain semen quality has not been established, and may vary among stallions. As little as 5% seminal plasma (v/v) has been show to be effective,8 and this protocol has been used successfully with semen from many different stallions in our laboratory. Centrifugation and partial removal of seminal plasma can improve sperm motility in ejaculates that do not tolerate the rigors of cooling and storage following conventional processing techniques.9 For some stallions, however, the toxic effects of seminal plasma are so dramatic that exposure to even minimal amounts can result in significant reductions in sperm longevity. For these stallions, centrifugation and removal of virtually all the seminal plasma may be necessary to reduce or eliminate adverse effects of seminal plasma. This presents a conundrum because, when standard extenders are used, centrifugation
and removal of all seminal plasma results in reduced motility of cool-stored stallion sperm.8 Therefore, it appears that, when all of the seminal plasma is removed, it is necessary to supplement the extender with a ‘seminal plasma substitute’ in order to maintain sperm motility. Following centrifugation and seminal plasma removal, motility10, 11 and fertility11 of cool-stored stallion semen is maintained when it is diluted in a NFDMS-GLC extender supplemented with a high-potassium modified Tyrode’s medium (KMT). The DNA integrity of stallion sperm is also maintained at a higher level when seminal plasma is removed and ejaculates are cooled-stored in KMT extender.12 Modified Tyrode’s medium is tedious to prepare and requires high-quality reagents, laboratory techniques, and equipment. Commercially available phosphate-buffered saline supplemented with glucose and pyruvate may serve as a suitable substitute for modified Tyrode’s medium. In one study, sperm motility and acrosome integrity were maintained in cool-stored, seminal plasma-free semen that was diluted with SKMG extender supplemented with this medium.13 However, no fertility data were reported in this study.
References 1. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares. Technique and preliminary findings. In: Proceedings of the 21st Annual Convention American Association of Equine Practitioners, 1975, pp. 327–36. 2. Varner DD, Scanlan CM, Thompson JA, Brumbaugh GW, Blanchard TL, Carlton CM, Johnson L. Bacteriology of preserved stallion semen and antibiotics in semen extenders. Theriogenology 1998;50:559–73. 3. Batellier F, Magistrini M, Fauquant J, Palmer E. Effect of milk fractions on survival of equine spermatozoa. Theriogenology 1997;48:391–410. ¨ ¨ 4. Pagl R, Aurich JE, Muller-Schl osser F, Kankofer M, Aurich C. Comparison of an extender containing defined milk protein fractions with a skim milk-based extender for storage of equine semen at 5◦ C. Theriogenology 2006;66:1115–22. 5. Pickett BW, Sullivan JJ, Byers WW, Pace MM, Remmenga EE. Effect of centrifugation and seminal plasma on motility and fertility of stallion and bull spermatozoa. Fertil Steril 1975;26:167–74. 6. Varner DD, Blanchard TL, Love CC, Garcia MC, Kenney RM. Effects of semen fractionation and dilution ratio on equine spermatozoal motility parameters. Theriogenology 1987;28:709–18. 7. Jasko DJ, Moran DM, Farlin ME, Squires EL. Effect of seminal plasma dilution or removal on spermatozoal motion characteristics of cooled stallion semen. Theriogenology 1991;35:1059–67. 8. Jasko DJ, Hathaway JA, Schaltenbrand VL, Simper WD, Squires EL. Effect of seminal plasma and egg yolk on motion characteristics of cooled stallion spermatozoa. Theriogenology 1992;37:1241–52. 9. Brinsko SP, Crockett EC, Squires EL. Effect of centrifugation and partial removal of seminal plasma on equine
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Semen Extenders for Cooled Semen (North America) spermatozoal motility after cooling and storage. Theriogenology 2000;54:129–36. 10. Padilla AW, Foote RH. Extender and centrifugation effects on the motility patters of slow-cooled stallion spermatozoa. J Anim Sci 1991;69:3308–13. 11. Rigby SL, Brinsko SP, Cochran M, Blanchard TL, Love CC, Varner DD. Advances in cooled semen technologies: seminal plasma and semen extender. Anim Reprod Sci 2001; 68:171–80.
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12. Love CC, Brinsko SP, Rigby SL, Thompson JL, Blanchard TL, Varner DD. Relationship of seminal plasma level and extender type to sperm motility and DNA integrity. Theriogenology 2005;63:1584–91. 13. Dawson GR, Webb GW, Pruitt JA, Loughlin TM, Arns MJ. Effect of different processing techniques on motility and acrosomal integrity of cold-stored stallion spermatozoa. J Equine Vet Sci 2000;20:191–4.
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Chapter 133
Factors Affecting Sperm Production and Output Edward L. Squires and Bill W. Pickett
The primary job of any breeding stallion is to produce good-quality sperm of normal quantity for delivery to the mare or the artificial vagina. Presented in this chapter are the major factors that affect sperm output. Principal factors determining sperm output are season, testicular size, age, frequency of ejaculation, and sexual behavior. Although each factor will be discussed separately, they are so interrelated that the discussions will have some overlap. This should not be construed to mean these are the only factors that affect efficiency of sperm production; environmental factors, hormonal status, and drugs can also play important roles.
Season Both the mare and stallion are seasonal breeders. Generally, changes in the stallion’s seminal characteristics and sexual behavior, because of season, coincide with the natural breeding season of mares.1 Estrus can be induced in anestrous mares by artificial lights.2 From a practical standpoint, exposing mares to 16 h of light, beginning about mid-December, stimulates them to establish a normal cycle earlier in the breeding season, resulting in earlier foals.2 The effects of season on stallion seminal characteristics and sexual behavior have been reported.3 However, effects of artificial light on reproductive function of the stallion have received far less attention. Seasonal modulation of sperm production is largely the modulation of spermatocytogenesis. During the breeding season, there is a slight increase in number of A1 spermatogonia and less degeneration of spermatogonia. Degenation of spermatids occurs primarily in the non-breeding season. The number of Sertoli cells is low in young animals and increases in adults but eventually declines with advancing age. In
addition, the number of Sertoli cells is lower in the winter than in the summer.4 This seasonal variation in Sertoli cell number has not been reported for other species. Season and seminal characteristics The effect of season on seminal characteristics is important to know so that a potential fertility evaluation could be conducted on a stallion any time of the year and the results related to the breeding season. Furthermore, stallion owners need this information to manage their stallion(s) for maximum reproductive efficiency. First and second ejaculates of semen were collected with an artificial vagina (AV) from each of five aged Quarter Horse (QH) stallions each week from May of one year through May of the following year. The second ejaculate typically was collected one hour after the first.3 Presented in Figure 133.1 are monthly means of gel-free seminal volume. The average gel-free volume of first ejaculates was 58 mL, while that of second ejaculates was 50 mL. The volume of first ejaculates ranged from 45 mL in February to 81 mL in June. Second ejaculates varied from 42 mL in March to 63 mL in. June. This was an increase of approximately 40% from the low months to the peak of the breeding season. Total volume of first ejaculates averaged 66 mL. The range was from 45 mL in January to 104 mL in June. Second ejaculates averaged 52 mL, ranging from 42 mL in March to 68 mL in June (Figure 133.2). Total seminal volume was greatly influenced by season, but most of the difference between first and second ejaculates was the result of the greater volume of gel in first ejaculates.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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The mean number of sperm per first ejaculate was 15 billion and ranged from 10 billion in January to 22 billion in July. Second ejaculates averaged 8 billion and ranged from 5 billion in December to 12 billion in July (Figure 133.3). Because the five stallions were collected each Tuesday, these sperm numbers rep-
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Figure 133.3 Effect of season on mean number of spermatozoa per ejaculate (billions). First ejaculate, solid line; second ejaculate, dashed line. (Adapted from Pickett BW, et al. Reproductive physiology of the stallion. VI. Seminal and behavioral characteristics. J Anim Sci 1976;43:617–25.)3
resent several days’ production. Sperm output during the lowest months was approximately 50% of the higher month’s output (50% and 39% for first and second ejaculates, respectively). This dramatic seasonal decrease in sperm output reflected a decrease in spermatogenesis. Based on these data, when the breeding season starts (February), stallion managers may be working with a stallion that is producing only 50% of the sperm he will produce in May. Such stallions, whose mares are presented early in the breeding season, may have more difficulty settling them than when his mares are presented later in the season as a result of this sperm output issue. If the stallion is normal, the number of mares in an artificial insemination (AI) program and the frequency of ejaculation in natural service will be the principal factors determining fertility early in the season. The difference in sperm output between first and second ejaculates, collected approximately 1 h apart, was 54.7%. This relationship is important in the potential fertility evaluation of stallions and in determining the intervals at which a stallion may be used per day. Because the data presented in Figure 133.3 are from stallions collected twice a week, the number of sperm per first ejaculates does not represent what happens under practical conditions. Therefore, single ejaculates were collected daily for 10 weeks from two groups of mature stallions (Figure 133.4). Group I consisted of four Quarter Horse (QH) stallions, and group II of one Thoroughbred (TB) and six QH stallions. Seminal collections of group I stallions began April 27 and ended June 4. Because daily sperm output (DSO) may not have peaked on June 4, when the
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experiment had to be terminated, the study was repeated the following year. Seminal collections from stallions in group II were obtained daily from April 24 to July 2.5 Large numbers of sperm were obtained during the first few days of daily collections. The number of sperm collected per ejaculate stabilized by days 5 to 7 of the collections and for the following week varied between 2.6 and 3.6 billion.5 The range in DSO averaged 3.2 to 6.6 billion (mean 4.8 billion) and 3.3 to 5.8 billion (mean 4.3 billion) during daily collections for 10 weeks from stallions in groups I and II, respectively. The first seven ejaculates averaged 5.3 billion sperm per ejaculate in both studies. This difference of approximately 2 billion sperm per ejaculate between the first and second weeks was because epididymal sperm reserves became depleted the second week. After a stallion has had several days of sexual rest, he can breed a larger number of mares per day for a few days until his sperm reserves are again depleted.6 Jasko et al.7 also demonstrated the effect of season on seminal characteristics. They reported that sperm concentration in December was 147 and 217 in August. The number of sperm per ejaculate was 3.3 billion in December and 6.8 billion in August. They used the coefficient of variation (CV) as a measure of repeatability for the seminal characteristics. They reported that the CV was highest for gel-free volumes, sperm concentration and total sperm per ejaculate whereas CV was lowest for progressive motility (visual), morphology, and computer-assisted analysis for all motion characteristics. Johnson et al.8 examined the affects of season and age in 184 stallions, 4 to 20 years of age. They reported that parenchyma weight and daily sperm production increased with age. There was no effect of age on Ser-
toli cell numbers. In contrast, season contributed to the variation of Sertoli cell number. Stewart and Roser9 conducted a study to examine the effect of age, season, and fertility status on plasma and intratesticular hormone levels. Fifty-one normal stallions from 2 months to 25 years of age were involved. Samples were collected during the breeding and non-breeding seasons. The group of stallions included six fertile, three subfertile, and three infertile stallions. They demonstrated that inhibin, luteinizing hormone (LH), testosterone, estradiol, and estrogen conjugates increased with testicular maturation, with the greatest increase occurring between 1 year and 2 12 years. They did not detect a seasonal difference in plasma hormone levels but attributed this to only one sample being collected per month. Fertility status of the stallions did have an effect on hormone levels. Testosterone tended to decline in infertile stallions whereas follicle-stimulating hormone (FSH) and LH were higher in infertile stallions. Inhibin, estradiol, and estrogen conjugates were lower in infertile stallions. Hoffman and Landeck10 obtained semen samples from 11 Warmblood stallions, seven during the months of January to July and four during the months of November to February. He reported that estrone, estrone sulfate, and estradiol-17β peaked between April and July and that estrogens were lowest between December to February. Testicular steroids such as testosterone and estrone sulfate were 24 and 54 times higher in plasma than seminal plasma. Photoperiod and season It has been amply demonstrated that certain seminal and hormonal characteristics, and many aspects of sexual behavior, are affected by season. Because humans have imposed an artificial breeding period on domesticated horses, clinicians sometimes find it desirable to alter the stallion’s reproductive capacity to coincide with these demands. Therefore, the following are some basic concepts of photoperiodism that have been developed for other animals. An understanding of the concept of refractoriness is central to understanding circannual or seasonal rhythms in photoperiodic (seasonal breeding) animals such as the horse In general, the term ‘refractory’ is used to describe a condition in which an animal is no longer sensitive or responsive to some prior stimulus that initiated a response. With seasonal breeders, this prior stimulus can be long days or short days. For example, suppose the response to artificial light is controlled by an internal trigger-firing mechanism. Thus, a long day acts to pull the trigger and fire the mechanism(s) that sets into motion events leading to heightened reproductive capacity. However, after a period
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Factors Affecting Sperm Production and Output of time, these events will cease as the animal enters a period of reproductive quiescence. Until the trigger is reset, no amount of pulling, i.e., continued exposure to long days, will fire the mechanism(s) again. This is the condition of photorefractoriness and is why alternating periods of short and long days are essential. However, a further complication is that circannual rhythms in seasonal breeders are often endogenous, i.e., they are set internally. Therefore, the seasonal cycle will be expressed in the absence of changes in daylength, but the changes in daylength act to synchronize the cycle. Photosensitive animals can be classified according to the concepts of refractoriness. Long-day breeders, such as the stallion, often are classified into two categories: scotorefractory and photorefractory. The term ‘scotorefractory’ describes animals in which spontaneous gonadal recrudescence occurs when the animal is maintained continuously in inhibitory photoperiods. This means that short days followed by re-exposure to long days (stimulatory photoperiods) are required to abolish this refractory state. The term ‘photorefractory’ describes animals in which spontaneous gonadal regression occurs as a result of continuous, prolonged exposure to stimulatory photoperiods. In this case, long days followed by re-exposure to short days are necessary to abolish this photorefractoriness and sensitize the animal so that exposure to long days will result in gonadal recrudescence. It appears that the stallion fits the category of photorefractory. An experiment11 was conducted to determine: (i) if stallions exposed continuously to long days would become photorefractory and (ii) if stallions exposed to long or short days would continue to exhibit a seasonal reproductive cycle. A total of 21 stallions of light horse breeds, 4 to 15 years old, was assigned to one of three treatments (7 per group): (i) controls, exposed to natural daylength; (ii) exposed to 8 h light and 16 h dark (8 : 16) for 20 weeks beginning July 16, 1982, then switched to 16 : 8 on December 2, 1982, and exposed to 16 : 8 until March 1984 (S-L); (iii) exposed to an 8 : 16 photoperiod from July 16, 1982, until March 1984 (S-S; Figure 133.5). Before July 16, 1982, all stallions were housed in a non-lighted barn. On July 15, 1982, the 14 stallions in Groups S-L and S-S were moved to a barn containing two separate lightproof areas in which photoperiod could be controlled independently. Each stall was illuminated by two 300 W incandescent bulbs. The 7 control stallions remained in the original barn and were exposed to natural daylength. Temperature was not controlled, but it was measured and was similar for all three groups. Total scrotal width was measured every 4 weeks by two technicians three times each.
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Semen was collected during 10 collection periods: June, September, and November 1982, February, April, June, August, October, and December 1983, and February 1984. For each period, seven ejaculates were collected to deplete extragonadal sperm reserves, then an additional nine ejaculates to assess daily sperm output. Each ejaculate was evaluated for gel volume, gel-free seminal volume, sperm concentration (millions) per milliliter of gel-free semen, sperm per ejaculate (billions), pH of gel-free semen, and percentage of progressively motile sperm. On each of the last nine collection days, sexual behavior of stallions was assessed by timing intervals: (i) to erection, (ii) to first mount with an erection and intromission, and (iii) from erection to ejaculation. The number of mounts per ejaculation was also recorded. The exposure of S-L and S-S stallions to an 8 : 16 photoperiod in July did not hasten the normal seasonal regression of testicular size (Figure 133.6). However, the S-L stallions responded to the stimulatory 16 : 8 photoperiod with an increase in total scrotal width, so that by February they had significantly larger (P < 0.05) testes than either the control or S-S stallions. This trend continued through March. Thereafter, testicular size of S-L stallions regressed, which was presumably a consequence of photorefractoriness. A photorefractory state was also likely responsible for regression in testicular size exhibited by control stallions in mid- to late summer. The annual fluctuation of testicular size was similar in S-S and control stallions.
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During June (pretreatment), sperm output averaged 6.4, 7.8, and 6.8 billion per day (P > 0.05) for control, S-L, and S-S stallions, respectively (Figure 133.7). By September, sperm output was lower than in June for all groups, and means for all groups were similar (4.9 billion, 4.0 billion, and 4.1 billion; P > 0.05). Annual fluctuation in sperm output for S-S stallions was similar to that for controls. Within 2 months after switching S-L stallions to the 16 : 8 photoperiod, their sperm output (10.4 billion) was greater (P < 0.05) than that for controls (7.1 billion) and S-S stallions (6.0 billion). Thus, by exposing S-L stallions to short days,
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then an increased daylength, the researchers were able to hasten testicular recrudescence so that maximum sperm output occurred at least 2 months earlier than in control stallions. The increase in sperm output exhibited by S-L stallions occurred virtually coincident with an increase in testicular size. The continued exposure of S-L stallions to a 16 : 8 photoperiod did not prolong the cycle, and regression occurred, albeit somewhat earlier than in the other two groups. This observation supports the conclusion that induction of earlier sexual recrudescence in the stallion might result in an earlier regression, which is certainly different from the response of mares to light.2 These data clearly demonstrate a stimulatory effect of increased daylength, following 20 weeks of short days, on testicular size and daily sperm output of stallions. However, normal seasonal recrudescence, as exhibited by control stallions, did not appear to be initiated by long days. This is because recrudescence, leading to peak reproductive capacity in early summer or late spring, appeared to be initiated during the fall when daylength was declining. In this experiment, the nadir in sperm output occurred in September to October and gradually increased until May or June. Because testicular size and sperm output eventually decreased, stallions seemed to become photorefractory in a manner analogous to that described for birds.12 Based on this observation, photostimulation at, or after, the summer solstice would not likely be effective in prolonging heightened reproductive function or delaying testicular regression. The normal, physiological role of short days during the fall may not be to terminate the breeding season but rather to abolish a refractory condition and re-establish a mechanism whereby the stallion will again be sensitive to increasing daylength or long days. Thus, termination of photorefractoriness allows subsequent photoinduction. Gel-free seminal volume was lower (P < 0.05) for S-L and S-S stallions during November 1982 and February and June 1983 (Figure 133.8) compared with controls. The reason for consistently lower seminal volumes in ejaculates of treated stallions is not known. The clinician should recognize, however, that other factors, such as sexual stimulation, may influence ejaculate volume. Furthermore, low seminal volume is not necessarily indicative of depressed or enhanced reproductive capacity, because during February, S-L stallions had a higher sperm output than control stallions, yet mean gel-free seminal volume of ejaculates of S-L stallions was 36 mL compared with 50 mL for controls. Once again, this is evidence of the relative unimportance of seminal volume as an indicator of reproductive function. The percentage of progressively motile sperm was not
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Figure 133.9 Effect of photoperiod on stallion sperm mobility. Control, dotted line; S-L, dashed and dotted line; S-S, solid line. (Adapted from Clay CM, Squires EL, Amann RP, Pickett BW. Influences of season and artificial photoperiod on stallions: Testicular size, seminal characteristics and sexual behavior. J Anim Sci 1987;64:517–25.)11
affected by treatment (P > 0.05) nor did it fluctuate with season (Figure 133.9). Although not significant, a trend toward a shorter time interval to erection for S-L stallions in March and May was noted (Figure 133.10). A similar trend was noted for the mean time required to mount an estrous
mare. Furthermore, total time to ejaculation was reduced (P < 0.05) for S-L stallions in both March and May compared with control or S-S stallions. When compared with control stallions, a reduction (P < 0.05) in number of mounts per ejaculation in March was observed (Figure 133.10). Whether this reduction
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was actually caused by photostimulation is not clear, because this group was not different (P > 0.05) from S-S stallions, and in general, control stallions required more mounts to ejaculate than either S-S or S-L stallions. In any event, note that the improvement in libido, as assessed by a reduced time to ejaculation, was a result of photostimulation. The researchers concluded that stallions possess an endogenous circannual rhythm for which changes in daylength act only as the primary synchronizer. Thus, in the absence of photoperiodic changes, the endogenous cycle still is manifested. Without annual synchronization by short days, this annual cycle would probably ‘free-run’ eventually, resulting in a drift of the annual interval of maximum reproductive potential.13 Manipulation of photoperiod apparently could be used to increase reproductive capacity and sexual behavior of stallions early in the year. This would require exposure to a period of 8-h days before exposure to long days. In this experiment, a period of 20 weeks of 8-h days was used, although a shorter period may have been adequate. Increasing daylength can be used to maximize reproductive efficiency early in the breeding season (February to April). Exposure of stallions to 16 h of light and 8 h of dark, starting in mid-to late December in northern latitudes, and maintaining them on this photoperiod until natural daylength approximates 16 h (early June) will provide maximum stimulation. For this procedure to be effective, the stallions must be allowed to be exposed to the decreasing daylengths of fall, because stallions, like most seasonal breeders, require alternating periods of long and short days for proper synchronization of circannual cycles. In addition, the farm managers must decide whether this schedule fits their particular breeding program. If the majority of mares are to be bred between February and June, then a lighting program may be suitable. If, however, mares are going to be bred in the late spring or early summer, a lighting program will be unsatisfactory and the stallions should be allowed to cycle normally. Exposure of stallions to an artificial photoperiod in late winter and early spring will result in peaking of the breeding season earlier in the year.
Shuttle stallions The shipping (shuttling) of stallions for breeding from the northern to southern hemisphere, to breed two seasons in one calendar year, is now common place. However, what effect this has on the stallions’ sexual behavior as well as their semen quality and sperm output has not been studied extensively. Pickett and Voss14 and Pickett et al.15 published two non-refereed articles on the management of shuttle
stallions for maximum reproduction efficiency. The study indicated two major concerns: (i) not allowing the stallion to have enough exposure to short daylengths before being exposed to long days; and (ii) requiring the stallion to ejaculate on a regular basis for two entire breeding seasons. Fertility results were available on 20 stallions that were shuttled from the United States to the southern hemisphere in late 1992 through late 1995. The live foal percentage was 65% for shuttle stallions the year after being shuttled and 64% for non-shuttled stallions. The shuttling of stallions did not appear to affect their fertility when they returned to the United States. There was a direct relationship between the numbers of mares bred per stallion and the live foal percentage, i.e., the larger the mare book, the higher the pregnancy rate. These authors questioned the wisdom of placing stallions under artificial lights since the practice appears to ‘shut the stallion down’ approximately two months earlier in the breeding season. Obviously, more studies are needed to determine the number of short days needed to render the stallions capable of responding to long photoperiods. The authors went on to propose that it may be possible to shorten the period of photorefractoriness by feeding melatonin shortly before or around the time the stallion leaves the northern hemisphere. Feeding melatonin would continue until shortly before the stallion is needed to breed mares in the southern hemisphere. The stallions would then be switched to GnRH injections until their plasma testosterone levels reach normal values. Studies are needed to test the efficacy of these protocols
Testicular size Daily sperm production (DSP) is the number of sperm produced per day by the testes. Daily sperm output (DSO) is the number of sperm harvested per day, after extragonadal sperm reserves have been stabilized, expressed on a per day basis. The efficiency of sperm production is the number of sperm produced per day per gram of testicular tissue.16 The number of sperm that an animal can produce depends largely on the amount of functional testicular tissue. Testicular size is an important factor in selecting and managing a stallion for maximum reproductive efficiency, assuming testicular consistency is satisfactory. Testicular measurements can be used to estimate sperm production and output, to identify individual stallions with a potentially high or low output, and perhaps to enable the prediction of how many mares can be bred under natural or AI management programs. There are two methods of measuring stallion testes, calipers (Figure 133.11) and ultrasound.
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Factors Affecting Sperm Production and Output
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Figure 133.11 Calipers used at our laboratory to measure total scrotal width to the nearest millimeter. (From Pickett BW, et al. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989.)21
Testicular measurements are generally made from the left side of the horse. Both testes are forced down into the scrotum with the left hand (Figure 133.12). The calipers are positioned on the left and right sides of the scrotum so as to span the distance between the greatest curvatures of the left and right testes. The greatest curvature of the right testis is located with the little finger, and the greatest curvature of the left testis is determined visually (Figure 133.13). The distance between the two points is total scrotal width (TSW). A certain amount of replication is obviously necessary to obtain an accurate estimate of testicular size. Therefore to minimize error, at least three independent measurements should be taken by one individual, and these measurements should be averaged to obtain a value for TSW. Forty-three clinically normal stallions of light horse breeds, aged 2 to 16, were used in a study conducted at Colorado State University.17 The TSW and the width and length of each testis were measured with calipers five times by each of two technicians on each of 6 days. Immediately after removal at castration, each testis was weighed; the tunica albuginea was removed, cleaned, and weighed; and the parenchyma weight was determined by difference. DSP was determined from analyses of testicular parenchyma.18 Presented in Table 133.1 are average testicular measurements as well as means of other reproductive characteristics. Average TSW for all stallions, across age, was 102 mm. The correlation coefficients among testicular measurements, parenchymal weight, DSP, and DSO
Figure 133.12 Positioning the testes for measurement of total scrotal width. (From Pickett BW, et al. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989.)21
Figure 133.13 Measuring total scrotal width of a stallion. (From Pickett BW, et al. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989.)21
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Table 133.1 Reproductive characteristics of stallions∗ Mean†
Characteristics Measurement (mm) Total scrotal width Left width Right width Left length Right length Parenchyma weight, paired (g) Daily sperm Production (109 ) Output (109 )
102 ± 9.9 58 ± 5.2 56 ± 5.8 103 ± 8.2 108 ± 8.0 328 ± 104 4.55 ± 1.46 3.35 ± 1.41
∗
n = 43. Means are presented plus or minus standard deviation. (Adapted from Thompson DL Jr, Pickett BW, Squires EL, Amann RP. Testicular measurements and reproductive characteristics in stallions. J Reprod Fertil Suppl 1979;27:13–17)17 †
(Table 133.1) are presented in Table 133.2. These values provide an estimate of the relationship between two variables. For example, the highest correlation coefficient was found between scrotal width and parenchymal weight (r = 0.83). This means that TSW accounted for 69% (0.83 × 0.83 × 100) of the variation in weight of the testes. In addition, TSW accounted for approximately 55% of the variation in DSP. Measurements of length and width of the left and right testes, when evaluated singularly, did not provide better correlations with DSP and DSO than did measurement of TSW. These authors concluded the most useful application of measuring TSW is for predicting DSP in the intact animal. Testicular size is highly heritable in other species.19 The authors concluded that stallions with a scrotal width of two standard deviations less than the mean value reported in Table 133.1 should not be used for breeding, if maximum reproductive capacity is a desirable characteristic to promote
within a breed. Although TSW accounted for only 39% of the variation in DSO and it was not an accurate approach to estimate DSO (Table 133.2), in certain cases a general estimate of DSO may be useful. If semen is collected daily for 1 week or every other day for 2 weeks, DSO can be estimated to within ± 40%. The choice of technique to estimate DSO depends on the relative accuracy with which the estimate must be known and the amount of time and labor available for the estimate. Love and co-workers were the first to estimate testicular volume as a means to predict DSO in the stallion.20 These workers measured width, height, and length of each testis and used the formula of an ellipsoid to compute testicular volume of each testis. Measurements derived with ultrasonography were more accurate for predicting DSO than were measurements taken with hand calipers. The formula for computing testicular volume was: Volume = 4/3 π × width/2 × height/2 × length/2, where width, height, and length were measured in centimeters. Combining volumes of left and right testicular volumes yielded the total testicular volume (TTV). A regression formula was constructed for prediction of DSO: DSO (× 109 ) = 0.024 (TTV) – 1.26. With this technique of computing the testicular volume, the derived values were highly correlated with both testicular weight (r = 0.92) and DSO (r = 0.91). This correlation of DSO to testicular volume is much higher than that obtained with TSW. A small study was conducted to compare DSO of a stallion (Herb) with large testes (137 mm) and a stallion (BJ) with small testes (79 mm).21 Semen was collected from each stallion at a frequency of six times per week for 1 month (Table 133.3). Sperm output for Herb was 20.7 billion per ejaculate for the first 6 days, Table 133.3 Seminal volume and total spermatozoa per ejaculation at a frequency of one collection per day, 6 days per week
Table 133.2 Correlation coefficients between testicular measurements and reproductive characteristics∗ Measurement (mm) Total scrotal width Left width Right width Left length Right length
Parenchymal Weight (g) 0.83t 0.82 0.82 0.57 0.61
Stallion Herb∗
DSP
(109 )
0.75 0.68 0.76 0.52 0.59
DSO
BJ†
(109 )
0.55 0.50 0.61 0.34‡ 0.50
Weeks 1 2 3 4
Volume (mL)
Spermatozoa (109 )
Volume (mL)
Spermatozoa (109 )
85 74 90 75
20.7 12.9 12.0 11.7
76 72 63 56
5.9 3.7 2.6 1.8
∗
n = 43. r-value. ‡ p > 0.05; all others significant, p < 0.01. (Adapted from Thompson DL Jr, Pickett BW, Squires EL, Amann RP. Testicular measurements and reproductive characteristics in stallions. J Reprod Fertil Suppl 1979;27:13–17)17 †
∗
TSW= 137 mm. TSW = 79 mm. (From Pickett BW, et al. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989)21 †
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Factors Affecting Sperm Production and Output while extragonadal reserves (EGR) were being stabilized. During the following 3 weeks, weekly means were 12.9, 12.0, and 11.7 for a DSO of approximately 12 billion sperm. Assuming that 500 million motile sperm are needed per natural cover, how many mares could Herb breed? If 50% of the sperm ejaculated were progressively motile (a conservative estimate), Herb was ejaculating sufficient sperm to impregnate 12 mares per day by natural service. Obviously this is no problem with AI, but impossible under natural conditions, even if the covers were equally distributed (every 2 h) throughout the day. Therefore, in Herb’s case, sex drive becomes the limiting factor in number of mares that can be bred in a natural breeding program. If 100 million motile sperm were used in an AI program, Herb ejaculated sufficient sperm to inseminate 60 mares per day, or approximately 120 on an every-other-day collection schedule.22 On the other hand, BJ ejaculated 5.9 billion sperm per ejaculate during week 1 of collection, which decreased to a DSO of 1.8 billion in week 4. His DSO was so low by week 4 that BJ could impregnate fewer than two mares per day naturally or with AI, using 500 million motile sperm. When BJ was castrated and the testes were examined histologically, the testicular tissue was only 58% efficient, which was consistent with his low daily sperm output. Many stud managers believe that volume of ejaculate is important in fertility. However, little difference in volume ejaculated existed between Herb and BJ. Therefore, volume is not an important consideration in fertility of the stallion.22 Measuring TSW or volume will provide a general estimate of a stallion’s ability to produce and ejaculate sperm. The time interval required for formation (57 days) and transport (7 to 9 days), and ejaculation of sperm totals about 65 days. Estimation of DSO via scrotal measurements will not provide information on motility or morphology of sperm being produced.23 Moreover, stallions can have testes of normal size and consistency and yet be azoospermic.24 Thus, some type of seminal evaluation must be performed if fertility is to be predicted. Objective information obtained from testicular measurements can be used to identify horses with hypoplasia in contrast to those with a low efficiency of sperm production. Measurement of TSW or testicular volume is a recommended part of a routine potential fertility evaluation. Good stallion management should include measurements at least once each month and results plotted for each stallion. By maintaining and using such a plot, the clinician may be able to predict potential problems and take steps to alter management before a serious reduction in fertility occurs. In addition to measuring, each testis should be palpated for abnormalities and consistency.
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Age Determining the number of mares that a given stallion can breed, either naturally and/or through AI, is important for appropriate stallion management. All too often, a young horse is overused and an older stallion is underused. Age of stallion is one of the more important factors affecting sperm production and output and the number of mares that can be bred.17, 24, 25 Age not only affects the size of the testicles but also the number of sperm that are stored in the epididymides.
Sperm reserves The number of sperm ejaculated and quality of those sperm determine relative fertility of an ejaculate. The number of sperm available for ejaculation depends on reserves in the tails of the epididymides, deferent ducts, and ampullae.17, 24, 26 Sperm in the head and body of the epididymides are not available for ejaculation. The number available for ejaculation is profoundly influenced by interval since previous ejaculation(s), testicular size, and age. Stallions that have large sperm reserves in the tails of the epididymides can impregnate more mares in a shorter period of time than stallions with low reserves. A study was conducted to determine effect of age and ejaculation frequency on epididymal sperm reserves.24 In addition, the relationship between DSP and epididymal sperm reserves was investigated. Stallions of light horse breeds were sexually rested for at least 12 days, and then five ejaculates were collected at 1-h intervals or the stallions were ejaculated twice (1 h apart) every 4 days, once every 2 days, or once a day for at least 3 weeks. The total number of sperm recovered in each ejaculate was calculated. The 54 stallions were unilaterally or bilaterally castrated 1 to 2 h, 24 h, or 12 days after the last ejaculation. Sperm reserves in 71 epididymides were determined. DSP was calculated by analyzing testicular tissue. Presented in Table 133.4 are sperm reserves of 16 sexually rested stallions. Sperm reserves of sexually rested stallions increased with advancing age. Although the increase was not significant for the head, approximately twice the number of sperm was present in the body of the epididymis from 10- to 16year-old stallions (9.5 billion) than 2- to 4-year-olds (4.2 billion). The total number of sperm in sperm reserves of 2- to 4-, 5- to 9-, and 10- to 16-year-old stallions was 29, 41, and 45 billion, respectively. Thus, older stallions have a much larger storage capacity for sperm than 2- to 3-year-old stallions. Regardless of age, the tail of the epididymis contained about 62% of the sperm reserves and served as the major site
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Table 133.4 Effect of age on sperm reserves of sexually rested stallions∗
Table 133.5 Effect of age on testicular size (mm) in stallions age (years)
Spermatozoa Per Side (Billions)
Structure Efferent ducts Epididymis Head Body Tail Proximal deferent duct Total
2 to 4 Years (5)†
5 to 9 Years (5)
10 to 16 Years (6)
Percentage
0.3
0.1
0.1
0.6
4.0 4.2 18.7 1.2
5.0 7.8 25.8 2.1
6.4 9.5 28.0 1.4
14.3 19.4 61.7 4.0
28.5
40.8
45.4
Age (years) Measurements
∗
n = 16. Numbers in parentheses represent the number of stallions in that age group. (Adapted from Amann RP, Thompson DL Jr, Squires EL, Pickett BW. Effect of age and frequency of ejaculation on sperm production and extragonadal sperm reserves in stallions. J Reprod Fertil Suppl 1979;27:1–6)24 †
for sperm storage. Neither daily ejaculation nor a series of five ejaculations at hourly intervals reduced the number of sperm within the head or body of the epididymis. Thus, sperm transit through these portions of the epididymides, which are presumed to be involved in sperm maturation, is not altered by frequency of ejaculation. However, ejaculation either 24 or 48 h before a seminal collection reduced the number of sperm in the tail of the epididymis by 15 to 30%. Contrary to popular belief, ejaculation frequency has little or no effect on fertility of the individual spermatozoon ejaculated during mating. A common belief presumes that, if ejaculation occurs too frequently, the stallion will ejaculate immature sperm. No scientific evidence supports that belief. Testicular size, DSP, and EGR increased for several years after puberty. Based on these studies, the researchers concluded that DSP and EGR in a normal 3-year-old stallion should be adequate to permit use of the stallion at least once a day during the breeding season, provided testicular size is normal and sex drive is adequate. The reproductive capacity of older stallions should be adequate to permit use at least two or three times·a day during spring and summer. For most mature, normal stallions, sex drive is the limiting factor in number of mares that can be bred per day by natural service if they have normal size testes. It is not uncommon for stallions in natural breeding programs to be used three or more times per day.
Scrotal width Mean 67% of population 95% of population Left width Mean 67% of population 95% of population Right width Mean 67% of population 95% of population
2 to 3 (11)∗
4 to 6 (14)
≥7(18)
96† 88–103 81–111
100b 93–107 85–115
109C 102–117 95–124
55b 50–59 45–63
57b 53–62 49–66
61C 57–65 52–70
53b 48–58 43–62
55b 50–60 45–65
60C 55–64 50–69
∗
Numbers in parentheses represent the number of stallions in that age group. † Means with different superscripts differ (p < 0.05). (Adapted from Thompson DL Jr, Pickett BW, Squires EL, Amann RP. Testicular measurements and reproductive characteristics in stallions. J Reprod Fertil Suppl 1979;27:13–17)17
Testis size in relation to age A series of studies were conducted to determine the relationship of age of stallion with testicular measurements.17 A total of 48 stallions of light horse breeds, 2 to 16 years of age, were used. Testicular measurements were made from the left side of the stallion on six separate days by each of two technicians. Age of stallions significantly affected testicular size (Table 133.5). The testes of stallions 7 years or older were larger than those of younger horses. Average TSW for stallions 2 to 3, 4 to 6, and 7 years or older was 96, 100, and 109 mm, respectively. Ranges are presented that represent 67 and 95% of the population. Thus, in a given population of stallions, 66% should have a TSW within the first range of values. If 95% of the stallions are included, obviously the range is increased. These ranges should be considered when evaluating a stallion for breeding soundness. The correlation coefficient between TSW and age was 0.64 (P < 0.01) (Figure 133.14), which is another example of the interrelationships that affect sperm production and output. Approximately 40% of variation in testis size could be accounted for by age. Although scrotal width was correlated significantly with bodyweight, when bodyweight and age were considered together only age accounted for a significant amount of variation. Thus, for a given age, scrotal width was independent of bodyweight. In another experiment, five successive ejaculates were collected to determine the influence of age on
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Factors Affecting Sperm Production and Output 130
120
110 Scrotal width (mm)
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100
90 TSW (mm) = 92.0 mm + 1.35 x Age (years) r = .62
80
0
1
2
3
4
5
6
7 8 9 10 11 12 13 14 15 16 Age (year)
Figure 133.14 Total scrotal width as a function of age. TSW (mm) = 92 mm + 1.35 × Age (yr); r = 0.62. (Adapted from Pickett BW, et al. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989.)21
seminal characteristics.25 Two groups of clinically normal stallions of light horse breeds ranging in age from 2 to 16 years were used. The effect of age on seminal characteristics is presented in Table 133.6. Values for some characteristics declined markedly with successive ejaculates. Stallions in the 4- to 6and 9- to 16-year-old groups produced more gel-free semen and more total volume than 2- to 3-year-old stallions. The amount of gel in ejaculates of older stallions (13 mL) appeared to be greater than that for young stallions (2 mL), although the difference was not significant. The number of sperm per milliliter was similar for stallions in all three age groups (120
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million, 161 million, and 161 million, respectively). Fewer sperm were obtained in ejaculates from 2- to 3-year-old stallions than from 9- to 16-year-olds. Although the difference was not significant, twice the number of sperm was obtained in the average ejaculate from 4- to 6-year-old stallions (3.6 billion) than from 2- to 3-year-old stallions (1.8 billion). Number of sperm per ejaculate was similar for 4- to 6- and 9- to 16-year-old stallions. This is additional evidence that stallions do not become sexually mature until approximately 6 years of age or older, and sperm output in normal older stallions remains relatively high. Quality of sperm was similar for all age groups as evidenced by means of 55%, 63%, and 60% progressively motile sperm for 2- to 3-, 4- to 6-, and 9to 16-year-olds, respectively. This was not surprising because the stallions had been evaluated before initiation of the experiment and were classified as normal. Many stallion managers erroneously believe that seminal quality of a young stallion is inferior to a more mature stallion. The influence of age on number of sperm in successive ejaculates was most evident in first ejaculates collected 1 h apart (Figure 133.15). Major differences in sperm output were noted between 2- to 3-yearold stallions and the other groups, but not between 4- to 6- and 9- to 16-year-old stallions. Number of sperm per ejaculate was less (P < 0.05) for 2- to 3year-old stallions than from 9- to 16-year-olds in the first and second ejaculates but not in subsequent ejaculates. The first ejaculates from 4- to 6-year-old stallions contained more (P < 0.05) sperm than ejaculates from 2- to 3-year-old stallions, but slightly less than
Table 133.6 Effect of age on seminal characteristics of stallions
Per ejac Age (years) Characteristics∗ Seminal volume (ml) Gel Gel free Total Spermatozoa Concentration (106 /mL) Total (109 ) Motility (%) PH ∗
2 to 3(7)†
4 to 6 (16) 9 to 16 (21)
2.1 14.2 16.2
5.1 26.2 31.4
13.3 29.8 43.2
120.4 1.8 55.0 7.68
160.9 3.6 63.1 7.64
161.3 4.5 59.9 7.59
Means of five successive ejaculates. Numbers in parentheses represent the number of stallions in each age group. (adapted from Squires EL, Pickett BW, Amann RP. Effect of successive ejaculation on stallion seminal characteristics. J Reprod Fertil Suppl 1979;27:7–12)25 †
Sperm per ejaculate (109)
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Figure 133.15 Influence of age of stallion on sperm output in five successive ejaculates. 2- to 3-yr-olds, solid line; 4- to 6-yrolds, dashed line; 9- to 16-yr-olds, short and long dashed line. (Adapted from Pickett BW, et al. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989.)21
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Table 133.7 Effect of age and ejaculate number on total sperm per ejaculate (billion)
Table 133.8 Percentage of ejaculates with 200 million or less spermatozoa per ejaculate
Age (years) Ejaculate 1 2 3 4 5 Total
Age (years)
2 to 3 (7)∗
4 to 6 (16)
9 to 16(21)
4.5 2.4 0.9 0.6 0.5 8.9
9.5 3.5 2.6 1.3 1.1 18.0
11.4 5.5 2.4 1.8 1.2 22.3
Ejaculate 1 2 3 4 5
2 to 3 (7)∗
4 to 6 (16)
9 to 16 (21)
0 29 14 14 43
0 0 6 12 6
0 0 5 0 10
∗ ∗
Numbers in parentheses represent the number of stallions in that age group. (Adapted from Squires EL, Pickett BW, Amann RP. Effect of successive ejaculation on stallion seminal characteristics. J Reprod Fertil Suppl 1979;27:7–12)25
from 9- to 16-year-olds. Total number of sperm per ejaculate decreased with each successive ejaculate for 2- to 3-year-old stallions, but this decrease was not significant. Number of sperm obtained in five ejaculates from 2- to 3-year-olds (8.9 billion) was less (P < 0.05) than the 22.3 billion sperm obtained from 9to 16-year-old stallions. Sperm obtained in five ejaculates from 4- to 6-year-old stallions was intermediate at 18.0 billion (Table 133.7). By the fifth successive ejaculation, nearly half of the ejaculates from young stallions contained fewer than 200 million sperm. Thus 2- to 3-year-old stallions cannot be used with the same frequency as older stallions without lowering fertility. Because these stallions had at least 12 days of sexual rest, they probably could not be used daily in a natural service program. In contrast, all second ejaculates from 4- to 6- and 9to 16-year-old stallions contained more than 200 million sperm. These aged stallions probably could be used several times per day and not alter fertility in a natural service program. The clinician cannot assume that the fifth ejaculate from all aged stallions would provide satisfactory fertility, even after 12 days’ sexual rest, because of incomplete ejaculations. This is one of the advantages of an AI program, i.e., sperm are counted in each ejaculate to ensure that minimal numbers of motile sperm are available for breeding purposes. Based on data in Table 133.8, a normal, fertile, sexually rested, mature stallion could successfully impregnate five or more mares in one day by natural service. The willingness and ability of a normal aged stallion to mount and ejaculate may determine number of mares that can be bred on a given day. Stallions with a large reproductive capacity and excellent sex drive could have their breeding season shortened by synchronizing estrous cycles of mares, thus allowing a stallion to breed his mares in a shorter period. This
Numbers in parentheses represent the number of stallions in that age group. (From Pickett BW, et al. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989)21
may be extremely useful for a stallion with a heavy show or performance schedule. These data can be used to manage stallions for maximum reproductive efficiency. All too often, economic pressure and popularity determine the size of a stallion’s book, when it should be determined by reproductive capacity. The number of mares booked to some young stallions should be reduced, whereas in some cases, the number booked to older stallions could be increased without causing a decrease in fertility. Jones and Berndtson27 obtained testes from 47 stallions at castration or slaughter. Stallions ranged in age from 2 to 20 years. They reported that paired testis weight tended to increase until about 12 years of age. The numbers of Sertoli cells declined linearly from age 2 to 20 years. Sertoli cells numbers declined during this period of development but had a remarkable capacity to accommodate a greater number of developing cells. However, development of the reproductive organs proceeded until 12–13 years of age.
Frequency of ejaculation Frequency of ejaculation is one of the most important factors affecting sperm output. As frequency of ejaculation increases, number of sperm per ejaculation decreases, providing frequency exceeds one ejaculate every other day.6 If a stallion’s sex drive is sufficiently high, fertility may decline because of overuse, i.e., ejaculation of insufficient sperm, particularly early in the breeding season when sperm output may be only 50% as high as at the peak of the breeding season.3, 6 In fact, sex drive (libido) may be the limiting factor early in the breeding season when libido is normally low, and toward the end of the season when the stallion is sexually satiated.
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Factors Affecting Sperm Production and Output Two questions frequently asked are ‘How often can I use my stallion in a natural service program?’ and ‘How many mares can I book to my stallion in an AI program?’ To study the effects of frequency of ejaculation on some seminal and behavioral characteristics, nine Quarter Horse and Thoroughbred stallions were assigned randomly to three groups.6 The groups were assigned to a seminal collection frequency of one time (1 ×), three times (3 ×), or six times (6 ×) per week for 4 weeks (one collection period). Stallions in the 1 × group were collected once a week; 3 × stallions, every other day; and 6 × stallions, daily except Sunday. At the end of each collection period, stallions were sexually rested for 1 week, and groups were reassigned to a different collection period. This protocol was continued until each stallion had been collected at each frequency for 4 weeks. Evaluation of the results of this experiment was limited to data obtained during the last 2 weeks of the collection period, after EGR had stabilized (Table 133.9). No differences (P > 0.05) occurred in gel volume, gel-free seminal volume, or total seminal volume per ejaculate as a result of frequency of ejaculation. Volume of gel for the 1 ×, 3 ×, and 6 × frequencies averaged 18, 8, and 6 mL per ejaculate, respectively. Gel volume, gel-free seminal volume, and total volume per ejaculate were different (P < 0.01) among stallions. Although effect of frequency of ejac-
Table 133.9 Effect of frequency of ejaculation on mean stallion seminal characteristics over the last 2 weeks of a 4-week collection period Frequency of Ejaculation per Week Characteristic Volume/ejaculate (ml) Gel Semen, gel-free Total Spermatozoa Concentration/mL (106 ) Total/ejaculate (109 ) Total/week (109 ) Motility (%) PH Sexual behavior Mounts/ejaculate Reaction time (min)
1×
3×
6×
18 47 65
8 56 64
6 51 57
288a∗ 11.4a 11. 4a 54 7.56C
248a 11.7a 35.2a 52 7.48d
142b 5.9b 35.3a 56 7.58C
1.2 1.75
1.4 1.93
1.4 2.10
∗ Row means with different superscripts are significantly different: a,b, p < 0.01; c,d, p <0.05. (Adapted from Pickett BW, Sullivan JJ, Seidel GE Jr. Reproductive physiology of the stallion. V. Effect of frequency of ejaculation on seminal characteristics and spermatozoal output. J Anim Sci 1975;40:917–23)6
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ulation was not significant, the more frequently the animals were collected, the lower the gel volume. A total of 47, 56, and 51 mL of gel-free semen was collected at weekly frequencies of 1 ×, 3 ×, and 6 ×, respectively, but no differences in seminal volume caused by frequency of seminal collection were noted. Thus, a stallion can ejaculate large volumes of seminal plasma quite independently of concentration of sperm. Obviously, sperm number is more important than volume for maximum fertility. Total volumes per collection were 65 mL for 1 ×, 54 mL for 3 ×, and 57 mL for 6 × The differences were primarily the result of volume of gel, not gel-free semen. Sperm concentration averaged 288 million, 248 million, and 142 million per milliliter, respectively, for 1 ×, 3 ×, and 6 × treatments. In spite of two additional collections per week, the difference in sperm per milliliter between the 1 × and 3 × groups was only 40 million per milliliter. Although sperm concentration was similar when a stallion’s semen was collected once per week or every other day, when he was collected 6 × per week, the mean concentration was only 50% of the number obtained at the 1 × or 3 × frequencies. The stallions averaged 11.4 billion sperm per ejaculate at 1 × and 11.7 billion at 3 ×, but only 5.9 billion at 6 × (P < 0.05). Thus, a stallion being used every other day ejaculates as many sperm per cover as he would when used once or less per week. Total sperm output per week averaged 11.4 billion, 35.2 billion, and 35.3 billion at frequencies of 1 ×, 3 ×, and 6 ×, respectively. Thus, fewer sperm were collected at the 1 × frequency than 3 × or 6 ×. When EGR became stable, sperm output per week was identical at the 3 × and 6 × frequencies. This indicates that increasing frequency of collection above once every other day will not result in more sperm per week or per ejaculation. With an every-other-day seminal collection schedule, all sperm that are available for ejaculation are being collected. Therefore, for maximum use of the stallion, with a minimum amount of labor, stallions should be collected every other day. Percentage of progressively motile sperm averaged 54%, 52%, and 56%, respectively, for 1 ×, 3 ×, and 6 × frequencies. Thus, these frequencies of ejaculation had no detrimental effect on sperm quality as measured by motility. Reaction time and number of mounts per ejaculation were not affected by these frequencies of ejaculation. Thus, sex drive of a welladjusted stallion will remain as high on a daily collection schedule as on a schedule of once per week. In another experiment,6 eight mature stallions of the Arabian, Connemara, Quarter Horse, and Welsh Pony breeds were randomly assigned to ejaculation frequencies of two times (2 ×) or four times (4 ×) per week for 4 weeks (one collection period). For the lower frequency of ejaculation, one ejaculate was
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Table 133.10 Effect of frequency of ejaculation on stallion seminal characteristics Frequency of Ejaculation per Week Characteristic∗ Seminal volume (ml) Gel-free, 1st ejaculate Gel-free, 2nd ejaculate Total Spermatozoa ‘ Concentration/mL, 1st ejaculate (106 ) Concentration/ml, 2nd ejaculate (106 ) Mean concentration/ml (106 ) Total/1st ejaculate (109 ) Total/2nd ejaculate (109 ) Total/week (109 ) Sperm motility (%) 1st ejaculate 2nd ejaculate Mean Seminal pH 1st ejaculate 2nd ejaculate Mean
2×
4×
55 — 55
58 49 107
272 — 272 13.1 — 26.2
222 145 196 12.0 5.5 35.1
49 — 49
58 61 59
7.65 — 7.65
7.59 7.75 7.65
∗
Eight stallions collected from November 12 through February 21. (Adapted from Pickett BW, Sullivan JJ, Seidel GE Jr. Reproductive physiology of the stallion. V. Effect of frequency of ejaculation on seminal characteristics and spermatozoal output. J Anim Sci 1975;40:917–23)6
collected per day on Tuesday and Friday of each week, whereas for the higher frequency, two ejaculates were collected per day on Tuesday and Friday of each week. The study consisted of three 4-week collection periods. Thus, stallions in group I were collected at frequencies of 2 ×, 4 ×, and 2 × per week for the three collection periods. At the same time, stallions in group II were collected 4 ×, 2 ×, and 4 ×. Increasing ejaculation frequency from one to two ejaculates per day, twice a week, increased (P < 0.01) seminal volume obtained on 1 day. The volume of second ejaculates collected on the same day was lower (P < 0.01) than that of first ejaculates (58 vs. 49 mL) (Table 133.10). More sperm were collected in two ejaculates than in one, but mean concentration of sperm in the two ejaculates collected on the same day was lower (P < 0.05) than that of a single ejaculate. This was primarily caused by the lower (P < 0.01) concentration of sperm in the second ejaculate. As a result of lower seminal volume and concentration of sperm, number of sperm in the second ejaculate was 45.8% fewer (P < 0.01) than in first ejaculates collected the same day. Nevertheless, sperm per week for the 4 × frequency was 25.4% higher (P < 0.01) than at the 2 ×
frequency. Percent motile sperm in first and second ejaculates at the 4 × frequency tended to be higher than at the 2 × frequency (58 vs. 49%). Likewise, no difference (P > 0.05) occurred in percent motility between first and second ejaculates: Sperm output for stallions on the 3 × and 6 × frequencies (Table 133.9) was the same as the 4 × frequency. When two successive ejaculates were collected on the same day, number of sperm in the second ejaculate decreased by approximately 50% (45.8%). This relationship between first and second ejaculates is extremely valuable in evaluation of stallions for breeding soundness.3 A practical ejaculation frequency for stallions in an AI program, in breeds where semen cannot be stored, would be one ejaculate on alternate days. Two ejaculates per week might provide maximal number of sperm per ejaculate and be a practical ejaculation frequency, if semen were to be frozen and stored. Another advantage of these seminal collection frequencies, compared with daily collections, is that a smaller percentage of sperm are lost in the collection equipment and gel.28 This can be an important factor when a large number of mares are booked to a stallion. A total of 10 stallions of light horse breeds, between the ages of 3 and 23 years, were subjected to two frequencies of ejaculation in a double switchback experiment.29 Semen was collected from stallions either once (1 ×) or twice (2 ×) per day. Before beginning the experiment, all stallions were placed on a 1 × ejaculation frequency for 8 days. After the 8-day stabilization period, five stallions (group 1) were placed on a 1 × daily seminal collection schedule, while the other five stallions (group 2) were collected 2 × per day. This schedule was maintained for 14 days. At the end of the collection period, group 1 stallions were switched to 2 × and vice versa. During each collection period, the first 6 days were used as an additional stabilization period; data from the remaining 8 days were used to evaluate treatments. Second ejaculates from stallions on the 2 × schedule were collected approximately 8 h after first ejaculates. Total sperm available for insemination was lower (P < 0.001) for the 2 × frequency, because when semen was collected twice a day, more sperm were lost in the collection equipment and gel. However, sperm ejaculated were the same (7.5 billion vs. 7.4 billion) for the two frequencies of ejaculation, when ejaculations were corrected for sperm losses. Concentration of sperm per milliliter of gel-free semen was lower (P < 0.001) for the 2 × frequency (59 million vs. 134.4 million) because of the higher (P < 0.001) gel-free seminal volume. Presented in Table 133.11 is sperm output for the two groups of stallions at the two frequencies of ejaculation. In spite of a concerted effort to divide the stallions into two equal groups with
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Table 133.11 Mean 8-day sperm output in billions corrected for sperm losses
Table 133.12 Spermatozoal output (billions) the first 14 days of daily collection∗
Group
Day of Collection
1 2
Frequency of Ejaculation 1× 56.7 2× 63.3
2× 55.2 1× 63.4
1× 53.9 2× 64.5
(From Neil JR. Ejaculation frequency, sperm loss in collection equipment and training stallions to a phantom. M.S. thesis. Fort Collins, Colorado State University, 1983)28
respect to sperm output, a difference of 7.4 billion sperm in favor of group 2 was noted over the 8-day period. The most important observation in this experiment was the almost identical (7.5 billion vs. 7.4 billion) sperm output within groups, regardless of frequency of ejaculation.29 It must be concluded that increasing frequency of ejaculation from 1 × to 2 × per day did not result in an increase in sperm output. Thus, only so many sperm are available per day, and increasing frequency of breeding beyond once every other day will not increase daily sperm output. Therefore, for each mare to have an equal opportunity to become pregnant with natural service, the intervals between breedings must be long enough to ensure that each mare gets sufficient sperm for maximum reproductive efficiency. As any good stallion manager knows, each stallion must be treated as an individual when determining number of breedings per day, week, month, and season. To answer further the question ‘How often can I use my stallion?’, 11 stallions, ranging in age from 3 to 13 years old, were collected daily during the breeding season.30 Sperm output data for the first 14 days of collection are presented in Table 133.12. Based on number of sperm collected per ejaculate, EGR had stabilized by days 5 to 7 of consecutive daily collections. Thus, mean DSO by week 2 of daily collections should represent the true DSO of a stallion for that particular time of year. The large number of sperm collected during the first week of daily collections was attributed to accumulations in the EGR during the week of sexual rest. Thus, to obtain actual DSO for a particular time of year, semen must be collected from stallions daily for approximately 1 week before collecting sperm output data. After EGRs have been stabilized, accuracy of estimating actual DSO depends on number of consecutive days that ejaculates are collected. The number of sperm obtained on day 7 of daily collections should approximate daily sperm output. Thompson et al.31 further examined the precision of determining DSO using the 10-day, single ejaculate per –day
1 2 3 4 5 6 7 8 9 10 11 12 13 14
Spermatozoa (Billion)
Confidence Interval (95%)
8.8 6.9 6.9 4.4 3.8 3.9 2.6 4.1 2.7 3.4 2.9 3.0 3.6 3.5
±3.6 ±2.5 ±2.1 ±1.8 ±1.6 ±1.5 ±1.4 ±1.3 ±1.2 ±1.1 ±1.1 ±1.0 ±1.0 ±1.0
∗ Not corrected for spermatozoa lost in the collection equipment and gel. (Adapted from Gebauer MR, Pickett BW, Voss JL, Swierstra EE. Reproductive physiology of the stallion: Daily sperm output and testicular measurements. J Am Vet Med Assoc 1974;165:711–13)30
model. Ejaculation from 15 stallions collected for 10 days were analyzed using Bayesian modeling. There was an initial decline (1.54 billion per day) until a change point of 4.7 days, meaning it took approximately 5 days to stabilize sperm output. Testicular size accounted for 36.5% of the within-stallion variation in DSO. The number of mares that can be bred by natural service can only be rationally established by the individual(s) most familiar with that stallion, because in many cases sex drive is the limiting factor. For maximum reproductive efficiency the stallion must be managed to maintain maximum sex drive throughout the breeding season. His sex drive may change; thus, management must be realistic, intelligent, and observant. Several additional studies have been published on the effect of ejaculation frequency on daily sperm output. Magistrini et al.32 investigated whether frequency of ejaculation influenced the quality and freezability of stallion sperm. These investigators compared semen collection three times per week versus five times per week. Semen was evaluated fresh as well as frozen–thawed. They reported that, during the winter, ejaculates contained smaller volume of higher concentrated semen with slightly lower motility. The quality of frozen–thawed semen was not affected by ejaculation frequency. However, the best post-thaw motility was obtained in the winter. A study in Germany by Sieme et al.33 involved 71 stallions and 6,319 mares that were artificially inseminated within 12 hours of collection of the
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semen. The study was designed to evaluate the effect of frequency and interval of semen collection on sperm viability and fertility. They demonstrated that the ejaculates collected on the first mount had smaller volumes and higher concentrations. As the number of mounts increased before ejaculation, concentration of sperm decreased. Visual total motility and progressive motility was not affected by the number of mounts, but after 24 hours of storage, those ejaculates collected after three to six mounts had decreased progressive motility. As has been reported by others, fertility increased with the number of mares booked per stallion. In addition, stallions with superior progressive motility after 24 hours of storage had superior pregnancy rates. It stated that optimal fertility occurred when the frequency of collection was confined to one single daily collection for each stallion during the entire breeding season. Brinsko et al.34 evaluated semen quality of 10 stallions that were collected after at least 7 days of sexual rest at a frequency of two ejaculates on day 1, 1 h apart, and a third ejaculate the following day. Progressive motility was higher in the third (57%) than in the first (51%) ejaculate. This subtle but significant difference was likely due to the stallions being sexually rested for 7 days prior to the beginning of this experiment, with insufficient ejaculates collected in order to stabilize the extragonadal sperm reserves. One study, involving 15 stallions, revealed the number of daily ejaculates required to remove extragonadal sperm reserves and achieve daily sperm output ranged from 2 to 9.31 The exact ejaculation frequency that is needed to maximize the number of sperm per ejaculate and the quality of sperm per ejaculate for semen freezing has not been determined. Sieme et al.35 compared the quality of semen obtained from ejaculates collected twice in one day, 1 h apart at 48-h intervals versus those collected once per day. They concluded that double semen collection 1 h apart at 48-h intervals gave good or better results than daily collection. They suggested that every-other-day collections may be too infrequent for good semen quality from some stallions that are used in a frozen semen program.
References 1. Ginther OJ. Occurrence of anestrus, estrus, diestrus, and ovulation over a 12-month period in mares. Am J Vet Res 1974;35:1173–9. 2. Kooistra LH, Ginther OJ. Effect of photoperiod on reproductive activity and hair in mares. Am J Vet Res 1975;36:1413–19. 3. Pickett BW, Faulkner LC, Remmenga EE, Berndtson WE. Reproductive physiology of the stallion. VI. Seminal and behavioral characteristics. J Anim Sci 1976;43:617–25.
4. Johnson L, Blanchard TL, Varner DD, Scrutchfield WL. Factors affecting spermatogenesis in the stallion. Theriogenology 1997;48:1199–216. 5. Sullivan JJ, Pickett BW. Influence of ejaculation frequency of stallions on characteristics of semen and output of spermatozoa. J Reprod Fertil Suppl 1975;23:29–34. 6. Pickett BW, Sullivan JJ, Seidel GE Jr. Reproducive physiology of the stallion. V. Effect of frequency of ejaculation on seminal characteristics and spermatozoa output. J Anim Sci 1975;40:917–23. 7. Jasko DJ, Lein DH, Foote RH. The repeatability and effect of season on seminal characteristics and computeraided sperm analysis in the stallion. Theriogenology 1991;35:317–27. 8. Johnson L, Varner DD, Tatum ME, Scrutchfield WL. Season but not age affects Sertoli cell number in adult stallions. Biol Reprod 1991;45:404–10. 9. Stewart BL, Roser JF. Effects of age, season, and fertility status on plasma and intratesticular immunoreactive (IR) inhibin concentrations in stallions. Domest Anim Endocrinol 1998;15(2):129–39. 10. Hoffmann B, Landeck A. Testicular endocrine function, seasonality and semen quality of the stallion. Anim Reprod Sci 1999;57:89–98. 11. Clay CM, Squires EL, Amann RP, Pickett BW. Influences of season and artificial photoperiodon stallions: testicular size, seminal characteristics and sexual behavior. J Anim Sci 1987;64:517–25. 12. Follett BK. Photoperiodism and seasonal breeding in birds and mammals. In: Lamming GE, Crighton DB (eds) Control of Ovulation. London: Butterworth, 1978; pp. 267–93. 13. Davis EB. Endogenous cycles. In: Steinberger A, Steinberger E (eds) Testicular Development, Structure, and Function. New York: Raven Press, 1980; pp. 359–66. 14. Pickett BW, Voss JL. Management of shuttle stallions for maximum reproductive efficeincy, Part 1. J Equine Vet Sci 1998;18(4):212–27. 15. Pickett BW, Voss JL, Clay CM. Management of shuttle stallions for maximum reproductive efficiency, Part 2. J Equine Vet Sci 1998;18(5):280–7. 16. Amann RP. A critical review of methods for evaluation of spermatogenesis from seminal characteristics. J Androl 1981;2:37–58. 17. Thompson DL, Jr, Pickett BW, Squires EL, Amann RP. Testicular measurements and reproductive characteristics in stallions. J Reprod Fertil Suppl 1979;27:13–17. 18. Amann RP, Johnson L, Thompson DL, Jr, Pickett BW. Daily spermatozoal production, epididymal spermatozoal. reserves and transit time of spermatozoa through the epididymis of the Rhesus monkey. Biol Reprod 1976;15:586–92. 19. Coulter GH, Rounsaville TR, Foote RH. Heritability of testicular size and consistency in Holstein bulls. J Anim Sci 1976;43:9–12. 20. Love CC, Garcia MC, Riera FR, Kenney RM. Evaluation of measures taken by ultrasonography and caliper to estimate testicular volume and predict daily sperm output in the stallion. J Reprod Fertil Suppl 1991;44:99–105. 21. Pickett BW, Amann RP, McKinnon AO, Squires EL, Voss JL. Management of the Stallion for Maximum Reproductive Efficiency. II. Animal Reproduction Laboratory Bulletin No. 05. Fort Collins, Colorado State University, 1989. 22. Voss JL, Pickett BW. Reproductive Management of the Broodmare. Animal Reproduction Laboratory General Series Bulletin No. 961. Fort Collins, Colorado State University, 1976.
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Factors Affecting Sperm Production and Output 23. Voss JL, Pickett BW, Squires EL. Stallion spermatozoal morphology and motility and their relationship to fertility. J Am Vet Med Assoc 1981;178:287–9. 24. Amann RP, Thompson DL Jr, Squires EL, Pickett BW. Effect of age and frequency of ejaculation on sperm production and extragonadal sperm reserves in stallions. J Reprod Fertil Suppl 1979;27:1–6. 25. Squires EL, Pickett BW, Amann RP. Effect of successive ejaculation on stallion seminal characteristics. J Reprod Fertil Suppl 1979;27:7–12. 26. Gebauer MR, Pickett BW, Swierstra EE. Reproductive physiology of the stallion. III. Extra-gonadal transit time and sperm reserves. J Anim Sci 1974;39:737–42. 27. Jones LE, Berndtson WE. A quantitative study of Sertoli cell and germ cell population as related to sexual development and aging in the stallion. Biol Reprod 1986;35:138–48. 28. Neil JR. Ejaculation frequency, sperm loss in collection equipment and training stallions to a phantom. M.S. thesis. Fort Collins, Colorado State University, 1983. 29. Pickett BW, Neil JR, Squires EL. The effect of ejaculation frequency on stallion sperm output. Proceedings of Ninth Equine Nutritional Physiology Society Symposium, 1985, pp. 290–5.
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30. Gebauer MR, Pickett BW, Voss JL, Swierstra EE. Reproductive physiology of the stallion: Daily sperm output and testicular measurements. J Am Vet Med Assoc 1974;165:711– 13. 31. Thompson JA, Love CC, Stich KL, Brinsko SP, Blandard TL, Varner DD. 2004. A Bayesian approach to prediction of stallion sperm output. Theriogenology 2004;62:1607–17. 32. Magistrini M, Chanteloube PH, Palmer E. Influence of season and frequency of ejaculation on production of stallion semen for freezing. J Reprod Fertil Suppl 1987;35:127–33. 33. Sieme H, Echte A, Klug E. Effect of frequency and interval of semen collection on seminal parameters and fertility of stallions. Theriogenology 2002;58:313–16. 34. Brinsko SP, Spooner JA, Blanchard TL, Love CC, Varner DD. Relationships among sperm membrane integrity, motility, and morphology in first and third ejaculates of sexually rested stallions. Proceedings of Annual Convention of American Association of Equine Practitioners, 2004;50: 502–4. 35. Sieme H, Katila T, Klug E. Effect of semen collection practices on semen characteristics before and after storage and on fertility of stallions. Theriogenology 2004;61:769–784.
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Chapter 134
Reproductive Parameters of Draft Horse, Friesian and Warmblood Stallions Tom A.E. Stout and Ben Colenbrander
Introduction
Stallion fertility
Breeding stallions are selected primarily on the basis of pedigree, appearance, gait, and performance. In addition, many studbooks disqualify stallions that do not conform to certain breed norms (e.g., any white on a Friesian stallion) or that are affected by abnormalities known or suspected to be heritable (e.g., laryngeal hemiplegia, osteochondrosis, cryptorchidism). Fertility is not a primary selection criterion, but most European sport-horse studbooks do insist on pre-registration certification of ‘adequate breeding potential’ based either on actual fertility figures or on a breeding soundness evaluation (BSE). However, because conventional semen quality parameters correlate no more than moderately with in-field fertility, the BSE aims to do no more than exclude stallions obviously incapable of adequate fertility.1 In the Netherlands, the Royal Dutch Warmblood (KWPN), Friesian (KFPS), and draft horse (KVTH) studbooks all require stallions to pass a preregistration BSE performed at a recognized laboratory. On the other hand, each studbook stipulates different criteria for ‘acceptable’ semen quality, primarily because significant between-breed differences exist in reproductive parameters depending, for example, on body size, age/maturity, and degree of inbreeding, and also because the likely demands on registered stallions differ markedly between breeds (e.g., average mare book size for a draft horse is much smaller than for a Warmblood). This chapter will present normal baseline reproductive parameters for Warmblood, Friesian, and draft horse stallions.
In intensively managed Thoroughbreds, there are large differences in fertility between stallions; Morris and Allen2 reported per-cycle and end-of-season pregnancy rates ranging from 37% to 90% and from 69% to 100%, respectively. In a study of 107 Hanoverian Warmblood stallions used to mate more than 12 000 mares over nearly 110 000 estrous cycles, mean per-cycle pregnancy rates ranged between 31.9% and 46.7%, with a single outlier that achieved a pregnancy rate of only 9.8%.3 The relatively low absolute pregnancy rates for these Warmbloods presumably reflect less precise mare management in the field, since Sieme et al.4 reported lower per-cycle pregnancy rates for Hanoverian horses inseminated at field stations (39.1%) than for those monitored at the Celle State national stud (53.9%). In general, the apparent fertility of popular stallions used for artificial insemination (AI) also commonly suffers from the management decision to make ‘too many’ insemination doses, each containing a suboptimal number of normal sperm. In the Netherlands, only stallions with ‘adequate semen quality’ are registered for breeding, and there are regulations governing the number of sperm in an insemination dose; even so, mean annual foaling rates for stallions mating more than 10 mares have been reported to range from 36% to 83% for Warmblood5 and from 31% to 100% for Friesian stallions,6 even though seasonal rates generally mask marginal fertility. Although unbiased per-stallion early pregnancy data have not been published for these breeds, the closely related ‘non-return rate’ (the number of mares that fail to return for mating within 28 days of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 134.1 Testicular dimensions (mean ± SD) of draft horse, Friesian, and Warmblood stallions presented for breeding soundness examination Breed
Age (years)
Length (left)
Length (cm) (right)
Width (cm) (left)
Width (cm) (right)
Draft horse
2.5–4 (n = 107) 4–10 (n = 14) 10–16 (n = 7)
10.1 ± 1.3 10.4 ± 1.5 12.1 ± 1.6
9.7 ± 1.1 10.0 ± 1.6 12.4 ± 0.9
5.9 ± 0.7 6.3 ± 0.8 7.3 ± 1.3
5.7 ± 0.7 5.9 ± 0.8 7.6 ± 0.9
Friesian
2.5 (n = 977) 3.5 (n = 344)
7.9 ± 1.1 8.2 ± 1.0
7.9 ± 2.4 8.1 ± 1.1
4.8 ± 1.5 5.0 ± 0.7
4.7 ± 1.5 4.9 ± 0.7
a
Warmblood
2.5–3 (n = 399)
9.8 ± 0.9
9.8 ± 0.9
5.6 ± 0.8
5.7 ± 0.7
a
Data previously published by Parlevliet et al.20
AI) has been reported to range from 27% to 89% per cycle for Dutch Warmblood and from 48% to 83% for Friesian stallions.5, 7 While foaling and pregnancy data are useful indicators of reproductive performance, and provide the only definitive proof that a stallion is fertile, the data are retrospective and can be misleading because success rates are affected by factors extrinsic to the stallion, such as mare quality and breeding management.8–10 Moreover, most stallions submitted for registration by a studbook have yet to breed any mares. In lieu of fertility data, it is accepted that a combination of a thorough physical examination and conventional semen evaluation provides a useful indication of fertility potential.11
Breeding soundness evaluation A BSE covers many aspects of potential fertility, including libido, mating ability, testicle size, sperm production, and semen quality.12 While sperm production does not correlate closely with fertility per se,5 it gives valuable information about the number of mares a stallion can cover, or insemination doses he can produce, per day or week; exceeding these limits can negatively affect fertility. However, while a BSE is useful for identifying stallions incapable of adequate fertility, it is less useful for predicting the likely level of fertility; indeed, Jasko et al.9 found only modest correlations between in-field fertility and the percentages of motile, progressively motile, or morphologically normal sperm (0.40, 0.46, and 0.36, respectively).
tive groups of young (2.5–3.5 years old) Dutch Warmblood, Friesian, and draft horse stallions presented at Utrecht University for BSE are shown in Table 134.1. Testicles were measured using the thumb and forefinger as ‘calipers’ (Figure 134.1); although the resulting dimensions are not as precise or reliable as ultrasonographic measurements, they are quick, easy, and relatively safe (for both personnel and equipment) to perform, and require only the most fractious stallions to be sedated. The Dutch studbooks do not have guidelines for minimum testicle size or volume, since abnormally small testicles are almost invariably reflected in inadequate sperm production. On the other hand, guidelines do exist for the maximum discrepancy in size between the left and right testicles; if the length and width of the testicles differ markedly, the heights are measured and the volume calculated. A difference of more than 50% in volume is considered
Testis size Daily sperm production by a stallion is dependent on a combination of the quantity of functional testicular tissue and the efficiency of spermatogenesis. Testicle size is, therefore, a useful indicator of sperm production capacity. Testis width and length in representa-
Figure 134.1 Measurement of testicle dimensions using the thumb and forefinger as ‘calipers’. The dimensions are quantified by measuring the distance between the digits against a standard ruler and, assuming the testicle is not squeezed, are accurate to approximately ± 0.5 cm. This is considerably easier than using standard calipers.
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Table 134.2 Mean semen quality for 2.5- to 4-year-old draft horse, Friesian, and Warmblood stallions presented for breeding soundness examination (mean ± SD )
Breed
Ejaculate
Motile (%)
Normal morphology (%)
Abnormal acrosomes (%)
TNM (× 106 )
Draft horse 2.5–4 years old (n = 108)
1st 2nd
52.9 ± 16.0 59.6 ± 14.9
48.6 ± 17.8 51.0 ± 17.1
12.4 ± 11.1 12.9 ± 11.2
2939 ± 3384 1913 ± 1728
1st 2nd
67.8 ± 9.0 70.6 ± 8.3
65.2 ± 14.6 67.5 ± 14.4
6.4 ± 6.4
6881 ± 4398 3745 ± 2282
Friesian 2.5 years old (n = 975)
1st 2nd
63.5 ± 14.2 65.8 ± 13.3
51.7 ± 15.2 52.0 ± 18.4
23.3 ± 14.0 23.6 ± 14.2
3342 ± 3321 1732 ± 1732
Friesian 3.5 years old (n = 343)
1st 2nd
67.0 ± 12.0 69.0 ± 10.1
54.2 ± 16.0 54.9 ± 16.2
24.5 ± 14.8 24.1 ± 14.3
4396 ± 3503 1953 ± 1547
a
a
Warmblood 2.5–3 years old (n = 398)
Data previously published by Parlevliet et al.20
abnormal, and, in young stallions, is most commonly the result of delayed testicular descent leading to a flaccid testicle with poorly developed seminiferous tubules. A large discrepancy in testicle size is often accompanied by unsatisfactory semen quality but, even if it is not, most Dutch studbooks will not approve the stallion for breeding. The animal may, however, be re-submitted a year later by when the difference in testicular size may have resolved or become more pronounced. Despite similar body size, average testicle size is generally lower in Friesian than in Dutch Warmblood stallions, in part because Friesians are a late-maturing breed and are assessed in the autumn or winter of their third year when many are still obviously immature. Testicle size and sperm production increase with stallion age (Tables 134.1 and 134.2) and, as for other breeds, will continue to do so until the stallion is at least 8, and possibly as much as 13, years of age; this should be taken into account when providing negative BSE advice due to poor sperm production for an immature stallion.
Semen quality In the standard BSE, two ejaculates are collected approximately 1 hour apart; the major parameters recorded are the percentages of motile and morphologically normal sperm and the total number of morphologically normal motile sperm (TNM) per ejaculate. The various Dutch studbooks have different standards for ‘adequate semen quality’ (Table 134.3) and, in the early 2000s, the normal values for both the Warmblood and Friesian studbooks were relaxed to avoid rejection of stallions desirable on other grounds that might subsequently prove to be adequately fertile despite marginal semen quality. A standard BSE has obvious limitations with regard to the variable immediate pre-examination ejaculation history; in this respect, the two ejaculates collected are less representative of sperm production capacity or baseline semen quality than a single ejaculate collected after reducing a stallion to ‘daily sperm output’ by daily semen collection for 5–7 days.13 For example, while most stallions void sperm
Table 134.3 Minimal semen quality parameters for the Dutch draft horse, Friesian, and Warmblood studbooks
Studbook Drought Horse (KVTH) Warmblood (KWPN)a Acceptable Satisfactory Friesian (KFPS) 3 years old 4 years old a
Motile sperm (%)
Morphologically normal sperm (%)
TNM (million)
NA
NA
600
30 50
30 50
1000 2000
50 50
50 (45b ) 50
600 (1000b ) 1000
When registering stallions for breeding, the KWPN uses the first set of values to define stallions as ‘acceptable’ breeding prospects. The semen quality values for newly registered stallions are subsequently published alongside a table of ‘expected fertility’ levels in which only stallions achieving the higher standards are described as ‘satisfactory’ breeding prospects. b Stallions are passed as satisfactory with an average of ≥ 45% morphologically normal sperm per ejaculate as long as they also deliver a minimum of 1000 TNM.
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Reproductive Parameters of Draft Horse, Friesian and Warmblood Stallions in the urine during a period of sexual rest, some do not; during a BSE, such sperm accumulators produce unusually high concentrations and numbers of (usually) poor quality sperm. Maintenance of an extremely high sperm concentration in the second (or subsequent) ejaculates is an indication of such ‘spermiostasis’. In general, if a stallion narrowly fails to pass a BSE on the basis of sperm quality, daily semen collection for 5–7 days followed by 2–3 days of sexual rest prior to repeating the BSE offers the best chance of a positive outcome. In short, the standard BSE gives only an approximate indication of true semen quality.
Percentage of motile sperm During a routine BSE, sperm motility is usually estimated by light microscopic evaluation of a sample diluted at least 1:1 with a basic semen extender and maintained at 38◦ C. However, as computer-assisted sperm analyzers (CASA) become more widespread, and the settings standardized, this more objective and repeatable method of estimating the percentages of motile and progressively motile sperm will probably supersede the visual estimation. Mean percentages of motile sperm (visual estimation) in the first and second ejaculates of representative subpopulations of Dutch Warmblood, Friesian, and draft horse stallions submitted to Utrecht University for BSE during 1987–2003 are presented in Table 134.2. Motility tends to increase in the second ejaculate, which usually contains a higher proportion of recently produced sperm, and is generally lower in both ejaculates for draft horses.
Percentage of morphologically normal sperm In general, highly fertile stallions in regular use exhibit a low proportion of morphologically abnormal sperm.14 On the other hand, it is thought that a wide range of morphological sperm abnormalities are compatible with normal fertility,15 as long as the percentage of morphologically normal sperm does not drop below a certain threshold (around 30%).16 However, while some abnormalities may have little or no effect on fertility (e.g., a distal protoplasmic droplet), or it is thought that they are compensable by increasing the total number of sperm in an insemination dose (e.g., defects that impair sperm transit), it is likely that other defects are not compensable and will seriously compromise fertility even if sperm numbers are high (i.e., defects that impair fertilization or embryo development).16 During a conventional BSE, the precise percentage and type of morphological sperm defects recorded will be affected by recent ejaculation history, method
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of examination (stained smear versus wet mount; level of magnification), and method of recording (all defects vs most proximal or most deleterious defect only). In Table 134.2, selected figures for the morphological quality of semen from Warmblood, Friesian, and draft horse stallions examined at Utrecht University are presented; in all cases, morphological analysis was performed on smears stained with aniline blue–eosin at a magnification of 1000× (oil immersion lens), and involved examination of at least 100 unstained (‘live’) sperm and recording of only the most proximal defect. The percentage of morphologically normal sperm generally increases in the second ejaculate, and there are clear between-breed differences in the percentages of both morphologically normal sperm and of particular types of abnormality. Most striking is the high incidence of abnormal acrosomes in the semen of Friesian stallions. In most cases, these are the so-called ‘knobbed acrosomes’ that have been associated with reduced fertility and reduced ability of sperm to acrosome react in both boars and bulls.17 In the Friesian horse, it is similarly assumed, but not proven, that knobbed acrosomes are detrimental to fertility. Moreover, because a high incidence of knobbed acrosomes in Friesian horses is highly heritable (h = 0.62; T.A.E. Stout and B.J. Ducro unpublished data), a negative breeding prognosis for affected stallions is justified until evidence is generated to demonstrate that the defect is either not detrimental to fertility or compensable. The higher percentages of morphologically abnormal sperm recorded for both Friesian and draft horse stallions compared with Warmbloods may be related to average degrees of inbreeding. The KWPN studbook is ‘open’ and regularly approves other types of Warmblood or Thoroughbreds as breeding stallions, thereby maintaining low average levels of inbreeding. By contrast, the Dutch draft horse studbook is small, in spite of open exchange with the Belgian and Ardennes studbooks, while the Friesian studbook is closed and has suffered two severe ‘population bottlenecks’ resulting in a narrow population base and an average coefficient of inbreeding of approximately 16%.18 Although a definitive link between inbreeding and the high incidence of sperm abnormalities has not been proven for Friesians, it seems possible, given that higher coefficients of inbreeding are associated with increased percentages of abnormal sperm in Shetland ponies.19
Total number of morphologically normal motile sperm As stated previously, TNM values generated during a standard BSE are no more than an indicator of sperm production capacity. Nevertheless, the
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Table 134.4 Estimated heritability of semen quality parameters in Warmblood and Friesian stallions
Breed Friesian (n = 1146)
Warmblood (n = 208)
Semen quality parameter
Heritability (h2 ± SEM)
Motile sperm (%) Morphologically normal sperm (%) Abnormal acrosomes (%) TNM
0.26 ± 0.08 0.45 ± 0.10 0.62 ± 0.11 0.21 ± 0.07
Motile sperm (%) Abnormal acrosomes (%) TNM
0.23 ± 0.18 0.03 ± 0.17 0.21 ± 0.14
values presented in Table 134.2 show that mean TNM values differ markedly among Warmblood, Friesian, and draft horse stallions, largely because of between-breed differences in both testicle size and the mean percentages of morphologically normal sperm.
Conclusions Warmblood, Friesian, and draft horse stallions differ greatly in average semen production capacity and semen quality. These differences are reflected in the different arbitrary norms stipulated by the studbooks for ‘acceptable semen quality’ during a BSE. Moreover, because semen quality is not an accurate predictor of likely fertility, minimum thresholds are deliberately low and aim only to identify obviously unsuitable breeding prospects. Certainly, the thresholds employed are too low to constitute true ‘selection’ for semen quality. In theory, selection for improved semen quality should be possible because the percentages of both motile and morphologically normal sperm are moderately heritable in both Warmblood and Friesian stallions (Table 134.4). Despite differences in minimum BSE requirements between breeds, the proportion of stallions classified as ‘unsatisfactory breeding prospects’ also differs markedly. When the semen quality thresholds for Dutch Warmbloods were 50% motile sperm, 50% morphologically normal sperm and 2 billion TNM (mean over two ejaculates), only 8% of stallions were classified as ‘unsatisfactory’.20 For Friesian and draft horses, the target index for total sperm production is 600 million TNM (mean over two ejaculates); however, despite being allowed up to three attempts within a calendar year to meet the criteria, 30–40% of Friesian and 14% of draft horse stallions fail to make the grade.
References 1. Colenbrander B, Gadella BM, Stout TA. The predictive value of semen analysis in the evaluation of stallion fertility. Reprod Domest Anim 2003;38:305–11. 2. Morris LH, Allen WR. Reproductive efficiency of intensively managed Thoroughbred mares in Newmarket. Equine Vet J 2002;34:51–60. 3. Hamann H, Jude R, Sieme H, Mertens U, Topfer-Petersen E, Distl O, Leeb T. A polymorphism within the equine CRISP3 gene is associated with stallion fertility in Hanoverian warmblood horses. Anim Genet 2007;38:259–64. 4. Sieme H, Schafer T, Stout TA, Klug E, Waberski D. The effects of different insemination regimes on fertility in mares. Theriogenology 2003;60:1153–64. 5. Parlevliet JM, Colenbrander B. Prediction of first season stallion fertility of 3-year-old Dutch Warmbloods with prebreeding assessment of percentage of morphologically normal live sperm. Equine Vet J 1999;31:248–51. 6. Royal Dutch Friesian Studbook. Yearly Report. (Breeding and Foal Information). http://www.fps-studbook.com/, 2008. 7. van Buiten A, van den Broek J, Schukken YH, Colenbrander B. Validation of non-return rate as a parameter for stallion fertility. Livest Prod Sci 1999;60:13–19. 8. Sullivan JJ, Turner PC, Self LC, Gutteridge HB, Bartlett DE. Survey of reproductive efficiency in the Quarter-horse and Thoroughbred. J Reprod Fertil Suppl 1975;23:315–18. 9. Jasko DJ, Little TV, Lein DH, Foote RH. Comparison of spermatozoal movement and semen characteristics with fertility in stallions: 64 cases (1987–1988). J Am Vet Med Assoc 1992;200:979–85. 10. van Buiten A, Westers P, Colenbrander B. Male, female and management risk factors for non-return to service in Dutch mares. Prev Vet Med 2003;61:17–26. 11. Kenney RM, Hurtgen J, Pierson R, Witherspoon D, Simons J. Theriogenology and the equine. Part II. The stallion: semen examination. J Soc Theriogenol 1983;9:1100–10. 12. Colenbrander B, Puyk H, Zandee AR, Parlevliet J. Evaluation of the stallion for breeding. Acta Vet Scand Suppl 1992;88:29–37. 13. Thompson JA, Love CC, Stich KL, Brinsko SP, Blanchard TL, Varner DD. A Bayesian approach to prediction of stallion daily sperm output. Theriogenology 2004;62:1607–17. 14. Rousset H, Chanteloube P, Magistrini M, Palmer E. Assessment of fertility and semen evaluations of stallions. J Reprod Fertil Suppl 1987;35:25–31. 15. Kavak A, Lundeheim N, Aidnik M, Einarsson S. Sperm morphology in Estonian and Tori breed stallions. Acta Vet Scand 2004;45:11–18. 16. Sieme H. Semen evaluation. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination, 2nd edn. St. Louis, MO: Saunders Elsevier, 2009; pp. 57–74. 17. Chenoweth PJ. Genetic sperm defects. Theriogenology 2005;64:457–68. 18. Sevinga M, Vrijenhoek T, Hesselinks JW, Barkema HW, Groen AF. Effect of inbreeding on the incidence of retained placenta in Friesian horses. J Anim Sci 2004;82:982–6. 19. van Eldik P, van der Waaij EH, Ducro B, Kooper AW, Stout TA, Colenbrander B. Possible negative effects of inbreeding on semen quality in Shetland pony stallions. Theriogenology 2006;65:1159–70. 20. Parlevliet JM, Kemp B, Colenbrander B. Reproductive characteristics and semen quality in maiden Dutch Warmblood stallions. J Reprod Fertil 1994;101:183–7.
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Chapter 135
Reproductive Parameters from Light Horse Stallions Edward L. Squires
Introduction A commonly asked question is ‘What are the normal values for semen parameters of a stallion?’ This is certainly a difficult question to answer because the semen characteristics of a stallion are dramatically affected by factors such as age, breed, season, frequency of use, and testis size. These factors will be discussed in detail in a subsequent chapter. However, it is important for the veterinarian and breeder to have a basic knowledge as to what the published values are for semen characteristics of light-breed stallions.
Semen evaluation There are two types of semen evaluations that are typically conducted. One is to collect two ejaculates 1 hour apart and if sperm numbers are approximately 50% (range of 20–80%) in the second ejaculate compared to the first ejaculate, then both ejaculates are considered to be complete. The second type of evaluation includes collection of ejaculates daily for 6 or 7 days. This is a more complete evaluation and allows one to obtain more information useful for predicting daily sperm output. Generally on a well-managed farm that is performing artificial insemination, stallion semen is evaluated on each and every ejaculate. In the Thoroughbred industry, and other breeds that allow only natural mating, semen evaluation typically is done in the non-breeding season and dismount samples are obtained for examination after breeding mares during the breeding season. The importance of semen evaluation includes determining whether there are problems in semen characteristics or whether there have been significant changes from the previous year. Information also can
be used to predict the number of mares that can be booked to the stallion and perhaps even make some projection as to the possible fertility of the stallion. With the recent increase in use of cooledshipped semen and frozen-thawed semen, many semen evaluations now include ‘test cools’ and ‘test freezes.’ The data on seminal characteristics that one obtains after semen collection depend upon how well the collections were performed. In order to properly collect and evaluate semen, one must have a clean, safe facility that has adequate equipment for proper evaluation. Oftentimes seminal parameters are altered because of poor technique used for collection and/or evaluation. This might include using unclean or contaminated equipment, or perhaps improperly collecting semen from the stallion. Such things as too much lubricant in the artificial vagina, unclean liners, excessive pre-sperm or excessive number of mounts may impact the quality of the semen. The detailed procedures for collection of semen are presented in Chapter 123. Situations where stallions should have a reproductive evaluation include:
r r r r r r r r
before sale; before the breeding season; to estimate the number of covers or inseminations that can be made during the season; anytime lower fertility is suspected; anytime the breeder wants to increase the number of mares to be bred; when abnormal sexual behavior is observed; to determine if seminal quality or testicular consistency has changed from the last breeding season; when the stallion is suspected of harboring a pathogenic bacterium.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Parameters included in the reproductive evaluation often include:
r r r r r r r r
gel, gel-free, and total semen volumes; sperm concentration (million per mL of gel-free semen); spermatozoa per ejaculation (billions); percentage of motile and progressively motile spermatozoa; sperm morphology; number of mounts per ejaculation and sexual behavior; cultures from at least the prepuce, and possibly semen; measurement of testicular size (i.e., testicular volume or total scrotal size) and assessment of testicular consistency.
Studies on seminal characteristics There have been several significant published studies on reproductive parameters of light-breed stallions. It is difficult to compare these studies since they were done in different areas of the country with different techniques and various breeds. Thus data from these large studies will be presented independently. Pickett et al.1 presented the results of potential fertility evaluations on 1044 stallions. Most of the data was from two ejaculates collected the same day, approximately 1 hour apart. These investigators divided the stallions into those that were approved as sound breeders (passed) and those that were not approved (failed). In this study, however, the actual fertility of the stallions was not determined. The number of stallions per semen characteristic that passed ranged from 417 to 664 compared with a range of 260 to 373 for stallions that failed the seminal evaluations. In many evaluations, the first ejaculate was extremely good and the second extremely poor or vice versa. As a result, the clinicians passed the stallion in spite of an extremely poor single ejaculate. This resulted in rather low means for motility and morphology of passed stallions. Presented in Table 135.1 are the means and standard deviations of seminal characteristics and total scrotal width of stallions that passed or failed a semen evaluation. Those stallions that passed had greater gel-free semen volumes in first and second ejaculates than those that failed (45 vs 40 and 42 vs 38). However, differences in total semen volume between passed and failed stallions were small. Volume generally is considered unimportant as long as sperm concentration and total sperm numbers are adequate. Although Rowley et al.2 reported that insemination of volumes of 100 mL resulted in reduced fertility compared with volumes of 10 mL, this was later shown to be a result of low concentration for the high-volume insem-
inates. Generally there is an inverse relationship between seminal volume and sperm concentration. The most important characteristics are the number of sperm per ejaculate and the quality of semen in the ejaculate. This will determine the number of mares that can be bred per ejaculate. More spermatozoa were obtained in both the first and second ejaculate from stallions that passed than from those that failed. The number of spermatozoa in the second ejaculate from a sexually rested stallion will usually equal approximately 50% of the number obtained in the first ejaculate.3 The percentage in the study of Pickett et al. (1987)1 was 47.1% for stallions that passed compared to 50% for those that failed. If this relationship does not exist, one of the following should be suspected:
r r r r r
One ejaculate was incomplete. The stallion had abnormally low sperm reserves. The stallion’s sperm reserves had been depleted. The stallion was young, immature, or suffering from testicular dysfunction or had only one scrotal testis. The stallion was an accumulator (i.e., abnormally large extragonadal sperm reserves resulting in abnormally high sperm numbers in the first and sometimes second ejaculate).
Percent of progressive motility of sperm in the first and second ejaculates from passed stallion was 53 and 57%, respectively, compared with 30 and 34% in corresponding ejaculates from stallions that failed. As expected, second ejaculates contained a higher percentage of motile spermatozoa than first ejaculates since stallions were sexually rested. The number of morphologically normal sperm in both the first and second ejaculates of stallions that passed compared with those that failed were 51 and 57%, and 35 and 38%, respectively. Love4 reported that at least five to ten samples should be evaluated for morphology in order to get a representative sample. He suggested that there were severe limitations in evaluating only one or two samples. He indicated that if morphologies are done correctly on enough samples, then the results can be important in predicting fertility of the stallion. The pH of second ejaculates was higher than the pH of first ejaculates. This occurs because second ejaculates contain approximately 50% of the spermatozoa as in the first ejaculate and possibly a reduced quantity of fluid from the epididymides. These authors stated that pH is beneficial in determining when ejaculations are complete. The number of spermatozoa a stallion is capable of producing is directly related to the grams of testicular tissue. In one study, daily sperm output
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Table 135.1 Means and standard deviations of seminal characteristics and total scrotal width of stallions that passed and failed a seminal evaluation∗ Passed
Seminal volume (mL) Gel Gel-free Total Spermatozoa Concentration (106 /mL) Total/ejaculate (109 ) Motile (%) Morphology Normal Abnormal (%) Neck separated Head Midpiece Tail Proximal droplet Distal droplet Other Seminal pH Total scrotal width (mm)
Failed
Ejaculate
Mean
SD ±
Mean
SD ±
Significance†
1 2 1 2 1 2
10 5 45 42 54 47
23 16 30 28 42 33
10 6 40 38 50 44
23 17 30 28 41 34
NS NS 001 0.05 NS NS
1 2 1 2 1 2
335 160 11.9 5.6 53 57
232 131 9.0 6.7 15 15
251 130 8.4 4.2 30 34
215 131 8.7 4.9 21 22
0.01 0.01 0.01 0.01 0.01 0.01
1 2
51 57
15 14
35 38
18 19
0.01 0.01
1 2 1 2 1 2 1 2 1 2 1 2 1 2 1 2 1
5 5 9 9 11 10 20 19 3 2 3 4 0.14 0.07 7.52 7.52 108
6 6 8 7 11 10 13 12 5 3 4 5 1 1 0.18 0.17 12
11 12 17 17 12 11 27 25 3 3 4 3 0.12 0.15 7.47 7.55 98
16 19 15 16 12 12 16 15 5 4 7 5 1 1 0.21 0.19 19
0.01 0.01 0.01 0.01 0.05 0.10 0.01 0.01 NS NS 0.01 0.05 NS NS 0.01 0.05 0.01
∗
Based on 417 to 664 observations for passed stallions and 260 to 373 for failed stallions. Significance, less than or equal to the number presented. From Pickett BW et al. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. Proc Am Assoc Equine Pract 1987;33:485–518.1 †
and daily sperm production were highly correlated (r = 0.8) and daily sperm production and testicular weight were also highly correlated (r = 0.77). Stallions that passed an evaluation had a total scrotal width of 108 mm, compared to 98 mm for those that failed.1 Love4 reported on the use of calipers or ultrasound for determining testicular volume in stallions. This technique, if properly done, is accurate in determining the amount of testicular tissue present. Presented in Tables 135.2 and 135.3 are the means and standard deviations of seminal characteristics and total scrotal width by breed for stallions that passed or failed a seminal evaluation. Nineteen
breeds were represented, but only Appaloosa, Arabian, Paint Horse, Quarter Horse and Thoroughbred were in sufficient numbers to warrant presentation by breed. For stallions that passed the examination, Thoroughbred stallions appeared to ejaculate more spermatozoa in the first ejaculate than Paint Horses (13 vs 8.5 billion). This may have been a result of the large number of Thoroughbred stallions represented, while only 18 Paint Horse stallions were used in this study. However, sperm numbers appear to be consistent with total scrotal width, which was 111 mm for Thoroughbreds compared to 102 mm for Paint Horse stallions. Among stallions that failed a semen
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Table 135.2 The effect of breed on seminal characteristics and total scrotal width of stallions across month and age that passed a seminal evaluation Seminal volume (mL)
Gel Breed Quarter Horse Mean SD± Thoroughbred Mean SD± Arabian Mean SD± Appaloosa Mean SD± Paint Horse Mean SD± Overall mean n Mean SD± Significance‡
Gel-free
Spermatozoa
Total
Concentration per mL (106 )
Total (109 )
Motile (%)
Normal morphology (%)
1∗
2†
1
2
1
2
1
2
1
2
1
2
12 29
7 20
43 26
40 25
55 41
46 32
309 248
148 110
10.7 7.5
5.1 5.6
55 13
57 14
53 15
7 21
4 15
46 29
44 28
52 39
48 31
364 227
164 111
13.0 6.4
5.9 4.9
52 15
57 16
10 20
5 11
39 24
42 29
49 37
47 35
367 221
179 147
11.6 7.4
5.4 4.0
53 17
5 11
4 7
38 26
35 26
43 33
39 31
350 245
161 139
10.3 5.0
4.1 2.4
8 23
4 10
34 20
34 21
42 36
39 28
304 191
141 82
8.5 5.4
600 9 23 NS
563 5 16 NS
603 43 27 0.05
566 41 27 NS
600 52 39 NS
563 46 32 NS
591 342 234 0.10
554 161 121 NS
591 11.5 7.0 0.01
1
2
Seminal pH
Total scrotal width (mm)
1
2
57 14
7.44 0.18
7.53 0.19
106 11
48 14
54 13
7.42 0.19
7.52 0.15
111 11
58 14
54 16
59 16
7.40 0.18
7.49 0.16
107 13
56 14
61 12
52 15
58 11
7.43 0.17
7.57 0.15
103 7
4.3 3.0
53 14
56 11
55 14
61 10
7.42 0.18
7.55 0.17
102 12
554 5.3 4.9 NS
599 53 15 NS
561 57 14 NS
581 52 15 0.01
547 57 14 0.05
385 7.43 0.18 NS
368 7.53 0.17 NS
552 107 12 0.01
∗
First ejaculates. † Second ejaculates. At least one difference at the designated level of significance exists within this column. From Pickett BW et al. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. Proc Am Assoc Equine Pract 1987;33: 485–518.1 ‡
evaluation, Thoroughbred stallions ejaculated 11.1 billion and 5.2 billion spermatozoa in first and second ejaculates, compared with only 7.2 billion and 2.5 billion for Appaloosa stallions. Furthermore, total scrotal width was much higher for Thoroughbred stallions than Appaloosas. Percent normal morphology was lowest in Arabian horses and highest in the first and second ejaculates from Appaloosa stallions. Presented in Table 135.4 are the comparisons of semen characteristics and total scrotal width of stallions with one scrotal testis and stallions with two scrotal testes that passed a semen evaluation. Semen volumes were either larger or had a tendency to be larger for stallions with two testes. As suspected, sperm concentration and total sperm per ejaculate were higher in both ejaculates from stallions that had two scrotal testes. In addition, motile spermatozoa in the second ejaculate and the percentage of morphologically normal spermatozoa in both ejaculates were lower for stallions that had one scrotal testis. The total scrotal width of stallions with only one scrotal testis obviously was lower than stallions with two scrotal
testes (67 mm vs 108 mm). Compensatory testicular hypertrophy occurs in some species after one testis is removed before or near puberty. Testicular hypertrophy was not noted in the study of Pickett et al.1 The combined output for first and second ejaculates from normal stallions was 17.5 billion compared with 8 billion for stallions with one testis. A total of 37 (3.5%) of the 1044 stallions evaluated had a right testis (49%), left testis (51%), or both testes (11%) rotated. Stallions with at least one rotated testis (180◦ ) had lower gel-free and total seminal volume and fewer spermatozoa in second ejaculates than those that passed the reproductive evaluation. The ratio for the number of spermatozoa in the second compared to the first was only 39.2% for stallions with a rotated testis, compared to 47.1% for passed stallions. Percentages of motile sperm in both ejaculates and of morphologically normal sperm in the first ejaculate were lower in stallions with a rotated testis compared with stallions that passed a seminal examination. In addition, total scrotal width was smaller (104 mm vs 108 mm) when a testis was rotated.
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Table 135.3 The effect of breed on seminal characteristics and total scrotal width of stallions across month and age that failed a seminal evaluation Seminal volume (mL)
Gel Breed Quarter Horse Mean SD± Thoroughbred Mean SD± Arabian Mean SD± Appaloosa Mean SD± Paint Horse Mean SD± Overall mean n Mean SD± Significance‡
Gel-free
Spermatozoa
Conc. per mL (106 )
Total
Total (109 )
Motile (%)
Normal morphology (%)
1∗
2†
1
2
1
2
1
2
1
2
1
2
9 20
7 16
35 29
35 25
44 35
40 29
243 207
126 138
7.1 8.4
3.6 5.4
35 22
39 21
40 19
13 35
8 23
48 32
42 27
61 54
50 39
260 187
145 128
11.1 9.1
5.2 4.5
28 20
32 20
8 20
5 12
33 25
36 24
42 32
41 28
300 269
136 132
7.7 7.7
4.4 5.0
20 18
8 18
4 8
32 19
25 17
40 28
29 18
231 182
82 78
7.2 6.6
2.5 3.2
8 11
2 5
36 32
36 25
45 41
38 24
223 190
205 165
9.3 11.5
345 10 23 NS
310 6 17 NS
345 38 29 0.01
311 37 25 0.10
344 48 41 0.01
310 43 31 0.05
330 260 217 NS
294 134 133 NS
329 8.5 8.6 0.01
1
2
Seminal pH
Total scrotal width (mm)
1
2
43 20
7.49 0.23
7.57 0.19
93 20
36 15
38 16
7.48 0.17
7.52 0.20
104 17
23 20
23 15
27 17
7.44 0.20
7.56 0.16
101 15
38 22
38 25
42 18
51 20
7.46 0.26
7.57 0.22
85 20
4.5 3.8
37 28
45 27
39 15
44 15
7.37 0.15
7.53 0.17
97 18
294 4.2 5.0 NS
333 30 21 0.01
300 33 22 0.01
324 35 18 0.01
284 38 19 0.01
285 7.47 0.21 NS
238 7.55 0.19 NS
294 98 19 0.01
∗
First ejaculates. † Second ejaculates. At least one difference exists at the designated level of significance within this column. From Pickett BW et al. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. Proc Am Assoc Equine Pract 1987;33: 485–518.1 ‡
A total of 32 stallions were diagnosed by palpation as having testicular degeneration. Testicular degeneration was detrimental to semen characteristics and total scrotal width (Table 135.5). From clinical observations in the stallion, testicular degeneration appeared to follow one of two patterns: (i) testis becomes progressively softer than normal and may even increase in size temporarily but eventually shrinks to the point to which almost no tissue is found in the scrotum; and (ii) testis becomes lobulated, constricted or both and progressively more firm and smaller until a firm, smooth mass remains. From casual observation, only one testis is initially affected, then the other. Thus, degeneration in the first testis may be a causative factor for degeneration in the other. In some cases, it may be beneficial to remove the degenerating testis in order to extend the reproductive life of the stallion. Further discussion on testicular degeneration can be found in Chapter 112. The effect of bacteria on seminal characteristics also was examined in the study of Pickett et al.1 Nu-
merous studies have shown that equine semen contains a variety of bacteria, the majority of which are non-pathogenic. A total of 73 (7%) of the 1044 stallions failed their semen evaluation exclusively because a majority of the culture sites were positive for one or more of the following bacteria: Klebsiella pneumoniae, Pseudomonas aeruginosa, or beta-hemolytic Streptococcus. Of the 530 stallions that passed the bacteriological portion of the breeding soundness examination, 29% had at least one culture positive for a potential pathogenic organism. However, frequency of isolation or level of growth was insufficient to warrant failing those stallions. Klebsiella pneumoniae was the most commonly isolated organism. A total of 68 (12.8%) of stallions that passed had at least one positive culture for this organism, compared with 72 of the 73 (98.6%) of stallions that failed. Seminal characteristics of stallions that passed or failed due to the presence of bacteria were similar. Thus, these authors concluded that harboring potentially pathogenic bacteria did not affect the stallion’s semen characteristics. The percentage of motile spermatozoa was 54%
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Table 135.4 Seminal characteristics and total scrotal width of stallions that passed a seminal evaluation compared with stallions with one functional testis One functional testis
Passed Variable Seminal volume (mL) Gel Gel-free Total Spermatozoa Concentration/mL (106 ) Total/ejaculate (109 ) Motile (%) Morphology (%) Normal Abnormal Neck separated Head Midpiece kinked, coiled, or irregular Tail reversed, coiled, etc. Proximal droplet Distal droplet Other Seminal pH Total scrotal width (mm)
Ejaculate
n
Mean
1 2 1 2 1 2
661 619 664 622 661 619
10 5 45 42 54 47
32 28 32 28 32 28
7 6 33 34 40 41
NS NS 0.01 0.05 0.01 NS
1 2 1 2 1 2
651 609 651 609 659 615
335 160 11.9 5.6 53 57
29 24 29 24 30 28
184 88 6.2 2.6 46 47
0.01 0.01 0.01 0.01 NS 0.05
1 2
634 599
51 57
30 26
43 43
0.05 0.01
1 2 1 2 1 2 1 2 1 2 1 2 1 2 1 2
633 598 633 598 633 598 633 598 633 598 633 598 633 598 436 417 605
29 26 29 26 29 26 29 26 29 26 29 26 29 26 27 23 30
9 10 13 11 11 12 25 22 2 0.7 4 2 0 0 7.45 7.55 67
NS NS 0.10 NS NS NS 0.10 NS 0.01 0.01 NS 0.05 0.01 0.05 NS NS 0.01
5 5 9 9 11 10 20 19 3 2 3 4 0.14 0.07 7.42 7.52 108
n
Mean
Significance
From Pickett BW et al. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. Proc Am Assoc Equine Pract 1987;33:485– 518.1
for both groups. It is unlikely however that a stallion would fail an examination because of the presence of one of these three organism on the skin of the penis or prepuce. Dowsett and Pattie5 collected data over four consecutive breeding seasons from stallions used at a commercial breeding establishment. Presented in Table 135.6 are the mean and standard deviations of semen characteristics for 63 stallion seasons with normal parameters compared to 10 stallions with the poorest fertility. Total volume, gel-free volume, gel volume, concentration, sperm numbers, motility,
total live sperm per ejaculate were all lower for stallions with the poorest fertility. In a subsequent study, Dowsett and Knott6 obtained data during four consecutive breeding seasons on 222 stallions of 13 breeds. This included 648 semen collection attempts. Presented in Tables 290.7 and 290.8 are the overall means and standard deviations of semen and behavioral characteristics from 536 ejaculates collected from 168 stallions. These investigators reported a parameter entitled ‘density and mass activity.’ Immediately after collection, a drop of gel-free semen was placed on a warm
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Table 135.5 Comparison between means of seminal characteristics and total scrotal width of stallions with and without testicular degeneration Passed Variable Seminal volume (mL) Gel Gel-free Total Spermatozoa Concentration/mL (106 ) Total/ejaculate (109 ) Motile (%) Morphology (%) Normal Abnormal Neck separated Head Midpiece kinked, coiled, or irregular Tail reversed, coiled, etc. Proximal droplet Distal droplet Other Seminal pH Total scrotal width (mm)
Degenerating
Ejaculate
n
Mean
1 2 1 2 1 2
661 619 664 622 661 619
10 5 45 42 54 47
31 28 31 28 31 28
15 10 49 39 64 49
1 2 1 2 1 2
651 609 651 609 659 615
335 160 11.9 5.6 53 57
29 27 29 27 30 27
167 96 7.4 3.7 32 34
0.01 0.01 0.05 0.05 0.01 0.01
1 2
634 599
51 57
28 27
31 35
0.01 0.01
1 2 1 2 1 2 1 2 1 2 1 2 1 2 1 2
633 598 633 598 633 598 633 598 633 598 633 598 633 598 436 417 605
28 27 28 27 28 27 28 27 28 27 28 27 28 27 27 23 29
10 13 17 17 12 13 31 27 4 3 5 3 0.46 0.07 7.38 7.47 90
NS 0.10 0.05 0.01 NS NS 0.01 0.05 NS NS 0.10 NS NS NS NS NS 0.01
5 5 9 9 11 10 20 19 3 2 3 4 0.14 0.07 7.42 7.52 108
n
Mean
Significance
NS NS NS NS NS NS
From Pickett BW et al. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. Proc Am Assoc Equine Pract 1987;33:485– 518.1
microscope slide and viewed under low power. Density was a visual estimate of sperm concentration which was scored 0 to 3. Mass activity was a visual estimate of overall activity of a population of sperm. Scores were 0 to 3 (0 was azoospermic or necrospermia; 1 was < 30% motility and/or < 50 million/mL; 2 was 30–70% motility and/or 50–150 million/mL; 3 was > 70% motility and/or > 150 million/mL). The average mass activity score was 2.07 and density score was 2.03. Sperm concentration averaged 164 million and total sperm 6.34 billion. In this study, the highest sperm morphological abnormality was associated with the midpiece (12.9%). Presented in Table
135.9 are the least square means of standard errors of stallion semen characteristics for various breeds. Pony, Palomino and Appaloosa stallions in this study produced fewer sperm and lower total semen volumes than other breeds. Sperm concentration and total sperm numbers of Arabian stallions were considerably higher than those of all other breeds. The lowest sperm concentration was obtained from Standardbred and Appaloosa stallions. Arabian stallions had the lowest percentage of dead sperm, while Shetland pony stallions had the highest. Jasko7 presented a review on evaluation of stallion semen. Presented in Table 135.10 are the mean
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Table 135.6 Means and standard deviations of the semen characteristics from 63 stallion seasons and the 10 stallions with poorest fertility and the maximum likelihood analyses of the relationship between semen characteristics and fertility 62 Stallion seasons
10 Poorest stallions
Semen characteristic
Mean
SD
Mean
Total volume (mL) Gel-free volume (mL) Gel volume (mL) Sperm conc. (106 /mL) Total spermatozoa per ejaculate (106 ) Motile spermatozoa (%) Total live spermatozoa per ejaculate (106 ) Dead spermatozoa (%) Abnormal heads (%) Proximal droplets (%) Distal droplets (%) Abnormal tails (%) Loose heads (%)
63.6 45.3 16.8 178.0 7217.8 82.1 5795.1 21.2 7.1 13.1 8.1 10.9 2.6
45.9 30.9 24.7 168.0 6870.0 16.0 5456.4 13.7 5.9 10.3 11.7 9.2 4.8
43.4 28.4 15.0 157.6 4239.0 66.0 3804.0 28.9 11.4 11.7 4.1 10.5 4.6
Adapted from Dowsett and Pattie (1982).5
values for seminal characteristics from three studies. Also presented in Table 135.10 are the mean morphological features of stallion sperm reported in these same three studies. Percent normal sperm was only presented in two of the studies, but was quite similar (51 and 55%). One of the greatest abnormalities found in two of the three studies was associated with proximal and distal droplets. The percentage of midpiece abnormalities also was high, which is similar to that reported previously.1, 5, 6 However, in an exten-
Table 135.7 Overall means and standard deviations of semen and behavioral charactersitcs for 536 ejaculates collected from 168 stallions Characteristic
Mean
SD
Total volume (mL)a Gel-free volume (mL)a Mass activity (score 0 to 3) Non-motile spermatozoa (%) Dead spermatozoa (%) Color (score 0 to 6) Density (score 0 to 3) Sperm concentration (106 /mL) Total sperm number (106 ) PH of gel-free semenb No. of mounts No. of pulses
44.53 3370 2.88 2.07 23.57 17.44 2.83 164.13 6.34 7.62 1.44 6.42
2.33 2.13 5.09 0.79 1.75 1.93 1.11 39.35 1.93 0.31 0.15 2.55
sive study by Love and co-workers8 involving Thoroughbred stallions, the proportion of distal or proximal droplets identified in dismount semen samples was not a good predictor of fertility. Several changes have occurred in the last two decades as to how mares are bred or inseminated. In the Thoroughbred industry, a full book of mares used to be 40 to 50 and now there are many stallions that are breeding over 100 mares per year and some as many as 200 mares per year. In addition, artificial insemination initially was done with the mare and stallion on the same farm using raw semen or semen that was diluted and used immediately. Since that time, cooled-transported semen was introduced and is now readily used in most breeds
Table 135.8 Overall means and standard deviations of spermatozoal characteristics for 531 ejaculates collected from 168 sallions Characteristic Abnormal heads (%) Proximal droplets (%) Distal droplets (%) Abnormal midpieces (%) Abaxial midpieces (%)a Abnormal tails (%) Loose heads (%)
Mean
SD
5.40 6.66 4.79 0.35 12.92 7.54 1.51
1.91 2.55 2.69 2.31 2.31 2.20 2.93
a
Due to difficulties with separation of the gel fraction, 512 samples from 165 stallions were used. b A total of 234 samples from 92 stallions was used to measure pH. Adapted from Dowsett and Knott (1996).6
a
A total of 447 samples from 161 stallions was used to estimate the percentage of abaxial midpieces. Adapted from Dowsett and Knott (1996).6
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Table 135.9 Least-squares means and standard errors of stallion semen characteristics for various breeds Breed of stallion
Semen characteristic
TB n = 141
SB n = 111
Arabian n = 73
ASH n = 73
QH n = 30
Palomino n = 44
Pony n = 38
Shetland n=8
Appaloosa n = 16
Total volume (mL) SE Gel-free volume (mL) SE Gel volume (mL) SE Mass activity SE Non-motile sperm (%) SE Dead sperm (%) SE Density SE Sperm conc. (106 /mL) SE Total sperm number (× 106 /mL) SE
4.19 1.09 28.33 0.09 2.66 0.15 1.70 0.08 29.27 1.06 21.56 1.07 1.85 0.07 114.29 0.53 5.03 0.015
37.35 1.11 30.20 0.09 3.13 0.24 2.11 0.09 21.28 1.07 15.36 1.09 1.92 0.08 97.24 0.64 4.74 0.021
39.67 1.12 36.20 0.10 0.95 0.23 2.34 0.11 14.96 1.08 10.10 1.10 2.44 0.09 286.82 0.86 12.66 0.026
45.26 1.11 33.19 0.10 5.52 0.21 2.21 0.10 22.21 1.08 17.12 1.09 1.90 0.08 116.06 0.73 4.85 0.022
32.83 1.16 32.80 0.14 4.00 0.34 1.85 0.14 26.08 1.11 23.81 1.13 2.02 0.12 171.66 1.29 5.37 0.044
26.25 1.15 23.78 0.13 1.06 0.30 1.54 0.14 27.15 1.11 21.32 1.12 1.84 0.11 138.48 1.13 4.02 0.041
16.85 1.15 20.78 0.23 2.46 0.33 2.12 0.16 30.35 1.10 24.69 1.11 1.75 0.11 104.01 1.09 1.12 0.040
31.83 1.32 44.39 0.49 13.14 0.79 2.30 0.33 29.94 1.21 38.47 1.25 1.84 0.23 101.25 4.37 1.75 0.161
28.66 1.21 23.28 0.19 1.97 0.46 1.80 0.18 26.80 1.14 15.81 1.16 1.35 0.15 90.42 1.10 3.33 0.076
TB Thoroughbred; SB Standardbred; ASH Australian Stock Horse; QH Quarter Horse. Adapted from Dowsett and Knott (1996).6
except Thoroughbred. Furthermore, with the recent acceptance of frozen-thawed semen by the American Quarter Horse Association and the American Paint Horse Association, there is an increased usage of frozen-thawed sperm. Therefore, when one is evaluating semen, the survival of sperm after cooling and storage and/or freezing and thawing must be considered. Love et al.8 stated that the clinician should be able to evaluate all components of the sperm, such as integrity of the acrosome, plasma membrane, sperm chromatin and mitochondria, as well as the normal seminal parameters discussed previously. In fact, with the advent of fluorescent probes and the
use of flow cytometry, it is now possible to evaluate multiple attributes of the sperm in a matter of a few seconds. With multiple probes, the sperm can be evaluated for acrosome damage, sperm viability, and DNA structure. Table 135.11 shows sperm parameters based on different levels of stallion fertility. Those stallions with lower fertility had a lower percentage of motile sperm, low percentage of morphologically normal sperm, and a higher comp-α-T, which is an indication of DNA susceptibility to degradatoin. In addition, in Table 135.12, those stallions with lower fertility had a lower percentage of normal sperm and a higher
Table 135.10 Means (standard deviation) of stallion seminal characteristics reported in several studies Reference Characteristic
Pickett et al. (1987)1
Dowsett and Pattie (1982)5
Jasko (1992)7
Gel-free volume (mL) Sperm conc. (106 /mL) Total sperm/ejaculate (109 ) Motile sperm (%) Progressively motile sperm (%) Morphologically normal sperm (%)
45 (30) 335 (232) 11.9 (9.0) NR 53 (15) 51 (15)
45 (31) 178 (168) 7.2 (6.9) NR 72 (16) NR
58 (24) 173 (118) 9.2 (5.9) 70 (17) 53 (24) 55 (19)
NR, not reported. From Jasko (1992).7 Reprinted with the permission of Elsevier.
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Table 135.11 Sperm values based on different levels of stallion fertility Fertility groups
1
2
3
Seasonal pregnancy rate % Pregnant/cycle % Total motile % Morphologically normal % COMP-αt
93a 82a 84a 69a 12a
86b 55b 74b 46b 17b
83c 39c 63c 46b 25c
Superscripts in rows are different (P < 0.05). Adapted from Love (2002).4 Reprinted with the permission of the Society for Theriogenology.
a,b,c
percentage of abnormal sperm – detached heads, midpiece abnormalities, coiled tails, and premature germ cells. Voss et al.9 studied the relationship between seminal characteristics and fertility of Thoroughbred stallions for 2 years. During year 1 the number of matings per pregnancy was significantly related (r = 0.53) to midpiece abnormalities, whereas the relationship during year 2 was lower (r = 0.01). Love et al.8 examined the intra- and inter-stallion variation in morphology and their relationship with fertility. Semen samples were collected from Thoroughbred stallions during the breeding season to determine the variation in sperm morphology within and among stallions. The variation among ejaculates within the same stallion was approximately 20%, but this variation in sperm morphology from day to day did not appear to affect the fertility of the sperm. Sperm morphology between stallions (interstallion variation) had a higher variability (i.e., coefficient of variation 30%). This was true for the percentage of normal spermatozoa, abnormal heads, deTable 135.12 Morphological features related to fertility in a group of Thoroughbred stallions Fertility groups
High
Medium
Low
% Pregnant per cycle Normal Abnormal heads Detached heads Midpiece abnormalities Coiled tails Premature germ cells
66 66 7 1.4 2.9 1.0 0.2
51 56 12 2.6 3.7 2.1 0.3
37 23 26 4 11 5.4 1.2
Adapted from Love (2002).4 Reprinted with the permission of the Society for Theriogenology.
tached heads, general midpiece abnormalities, coiled tails, and premature germ cells. These authors used a statistical method entitled ‘odd ratios’ which determined which morphological feature in question affected pregnancy outcome. The intra-stallion variation in sperm morphology was not a significant predictor of fertility. That is, no morphological feature was a predictor of fertility for ejaculates within a stallion. In other words, the intra-stallion variation in morphology within stallions did not affect pregnancy outcome. In contrast, the inter-stallion variation in sperm morphology was a significant predictor of fertility. Morphological factors that were significant predictors of stallion fertility were percentage of normal sperm, abnormal heads, detached heads, midpiece abnormalities, coiled tails, and premature germ cells. These authors concluded that the use of dismount samples was a convenient, reliable method of obtaining a sperm sample from a stallion that was used exclusively for natural cover. The results indicated that changes in sperm morphological features can influence the likelihood of a mare becoming pregnant and that these morphological features can be used to predict and explain differences in fertility among stallions.
References 1. Pickett BW, Voss JL, Bowen RA, Squires EL, McKinnon AO. Seminal characteristics and total scrotal width (T.S.W.) of normal and abnormal stallions. Proceedings of the 33rd Annual Convention of the American Association of Equine Practitioners, 1987; pp. 487–518. 2. Rowley HS, Squires EL, Pickett BW. Effect of insemination volume on embryo recovery in mares. J Equine Vet Sci 1990;10:298–300. 3. Pickett BW, Faulkner LC, Seidel GE Jr, Berndtson WE, Voss JL. Reproductive physiology of the stallion. VI. Seminal and behavioral characteristics. J Anim Sci 1976;43:617–25. 4. Love CC. Stallion semen evaluation and interpretation. Proceedings of the Society for Theriogenology, 2002; pp. 93–101. 5. Dowsett KF, Pattie WA. Characteristics and fertility of stallion semen. J Reprod Fertil Suppl 1982;32:1–8. 6. Dowsett KF, Knott LM. The influence of age and breed on stallion semen. Theriogenology 1995;46:397–412. 7. Jasko DJ. Evaluation of stallion semen. Vet Clin N Am Equine Pract 1992;8:129–49. 8. Love CC, Varner DD, Thompson JA. 2000. Intra- and interstallion variation in sperm morphology and their relationship with fertility. J Reprod Fertil Suppl 2000;56:93–100. 9. Voss JL, Pickett BW, Loomis PR. The relationship between seminal characteristics and fertility in Thoroughbred stallions. J Reprod Fertil Suppl 1982;32:635–6.
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Chapter 136
Reproductive Parameters from Miniature Stallions Dale L. Paccamonti and Paul J. De Vries
The origin of miniature horses is open to debate; however, one thing is certain – they have been present for many years in a number of countries, and their popularity seems to be increasing. While the term ‘miniature horse’ encompasses a number of different breed associations, the common feature is the size of the horse. Many originated with ponies, in particular the Shetland. The Falabella breed has been attributed to Andalusian horses left to roam free in Argentina. Future work on the genetics of miniature horses, full-sized horses, and ponies may provide more information on their relationships. Little research has been done specifically with miniature horses in regards to reproduction and little published literature is available. Along with their increasing popularity, there has been an associated increase in economic value. With the increase in economic value comes a corresponding increase in the frequency of requests for breeding soundness and pre-purchase examinations. Likewise, an increased economic value necessitates valid criteria by which to evaluate miniature stallions. A breeding soundness evaluation in a miniature stallion consists of the same steps as an evaluation in a full-sized stallion. Semen is collected and semen volume, sperm concentration, sperm motility, and sperm morphology are assessed. The external genitalia are examined and the testes are measured. The internal genitalia are not routinely examined during the course of a breeding soundness evaluation, but an examination may be indicated during the work-up of conditions such as hemospermia, leukospermia, or reluctance to ejaculate. The stallion should also be routinely examined for diseases such as contagious equine metritis (CEM), equine infectious anemia (EIA) and equine viral arteritis (EVA) by
appropriate tests, and routine cultures obtained for microbiological interpretation .
Semen collection Standard artificial vaginas such as the Colorado model may not be ideal for semen collection with a miniature stallion. In addition to being unnecessarily large and cumbersome, the semen will be deposited within the warm water jacket of this artificial vagina due to the small size of the penis and the low volume of the ejaculate. The warm water jacket is usually 45–48◦ C and this temperature is detrimental to spermatozoa if they remain in there for any length of time. In addition, if a small-volume ejaculate is deposited in the middle of a Colorado-style artificial vagina, a large portion of the ejaculate may be lost by coating the liner before it gains access to the semen collection receptacle. Therefore, a short Missouri-style or a Hannover-style artificial vagina may be more appropriate when collecting semen from a miniature stallion. Alternatively, a large-sized condom made for R in the USA) can human use (e.g. Trojan Magnum be used on a miniature horse for semen collection. The condom is placed on the stallion’s penis after an erection is attained and the stallion is permitted to mount a mare. When the stallion dismounts, the condom is immediately removed and the semen is harvested. Unfortunately, the necessity to later remove gel without an in-line filter is a drawback with condom collection and results in spermatozoal losses. Other than the size of the artificial vagina and the size of the ‘jump’ mare or phantom that is used, semen collection from a miniature stallion is similar in all other regards to semen collection from a fullsized stallion. An appropriate-sized mare or dummy in proportion to the stallion will be necessary,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 136.1 Semen parameters of miniature stallions. Data are mean ± SD Stallion height at withers (cm)
Gel-free semen volume (mL)
Sperm concentration (× 106 /mL)
Total sperm (× 109 )
Progressive motility (%)
Morphology (% normal)
72–96a 97–104a ≤ 96 (presumed)b Full-sized stallionsc (1st/2nd collection)
20.7 ± 0.8 29.0 ± 1.3 13.5 ± 7.6 42/45
246.9 ± 16.3 215.0 ± 15.2 177.2 ± 139.0 335/160
4.59 ± 0.3 5.41 ± 0.3 2.01 11.9/5.6
65.4 ± 0.9 61.6 ± 1.3 48.8 ± 12.9 53/57
57.0 ± 1.3 50.4 ± 1.6 67.6 51/57
a
Paccamonti et al.1 Metcalf et al.2 c Pickett.4 b
unless ground collection can be performed. Ground collection is an option to consider if a miniature jump mare or mini-phantom is not available. Techniques for ground collection or collection using a jump mare or phantom are covered in Chapter 123.
Semen evaluation The most significant differences observed with the ejaculates of miniature stallions are with the volume of the ejaculate and total number of spermatozoa in the ejaculate (Table 136.1). In a study of 216 miniature stallions, ejaculate volume was lower in the miniature stallions than is commonly expected in full-sized stallions. Moreover, when grouped by height at the withers, volume was significantly lower in the group of smallest stallions (< 86.5 cm) than in the group of larger (97–104 cm) miniature stallions (19.5 ± 1.1 vs 29.0 ± 1.3 mL, mean ± SE, respectively; P < 0.05).1 Spermatozoal concentration in the ejaculates of miniature stallions was within the range of expected values for full-sized stallions. However, when calculating total sperm in an ejaculate, the lower volume results in significantly fewer total sperm than would be typical of a full-sized stallion. Fortunately for breeders of miniature horses, other parameters of semen quality, i.e., percent motile sperm and percent morphologically normal spermatozoa, are similar to that for a full-sized stallion. When reviewing the numbers in Table 136.1, it should be pointed out that, in the study by Paccamonti et al.,1 the reported values were derived from the average of two collections obtained 1–3 hours apart in sexually inexperienced stallions, while in the study by Metcalf et al.2 the stallions were collected for at least 5 consecutive days, after which they were collected for 2 days and the data from the last two collections were averaged.
External genitalia Regardless of the size of the stallion, the presence of two scrotal testicles is generally considered necessary
in order to pass a breeding soundness evaluation. Although some clinicians believe that cryptorchidism, or failure of complete descent of the testicles into the scrotum, is more common in ponies and miniature horses than in full-sized horses, there is little published information on the incidence of the condition. There are many variations of this condition, with the location of the retained testicle being anywhere from the abdominal cavity to the inguinal region near the scrotum. Temporary inguinal retention of the testes is reportedly most common in small breeds, and is usually seen when a testicle is small, e.g., weighing less than 40 g.3 This condition is termed temporary because most of these testes will reportedly grow and eventually assume a scrotal location by the time the stallion has reached 3 years of age.3 The right testicle is affected more often than the left, as temporary inguinal retention of a testicle is usually unilateral.3 Incomplete abdominal retention of a testis refers to a condition in which the testis is retained within the abdomen but the vaginal tunic and associated contents (cremaster muscle, portions of the epididymis) extend into the inguinal canal. Interestingly, it has been reported that in full-sized stallions the left testis is retained as often as the right, but in ponies the left side is affected twice as often as the right side.3 Guidelines for acceptable testicular measurements were developed using stallions of full-sized breeds. The most popular miniature stallions are usually the smallest ones and criteria for full-sized horses such as Thoroughbreds or Quarter Horses are not applicable to smaller breeds such as the Miniature Horse, Miniature Shetland, or Falabella. Obviously, when a fully mature stallion of these smaller breeds weighs only one-tenth of what a fully mature Thoroughbred might weigh, criteria used in full-sized stallions for testicular volume and scrotal width, expected daily sperm output, or total sperm in an ejaculate will not be appropriate. Published data on testicular measurements (Table 136.2) provide clear evidence that, compared with
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Reproductive Parameters from Miniature Stallions Table 136.2 Measurements of external genitalia in miniature stallions Stallion height (at withers) (cm) 72–96a 97–104a ≤ 96 (presumed)b Full-sized stallionsd
Scrotal width (cm) mean 7.25 7.95 – 10.8
Testicular volume (cm3 ) 91.6c 149.5c 62.5
a
Paccamonti et al.1 Metcalf et al.2 c Volume was calculated using testicular width as testicular height. d Pickett.4
b
full-sized horses, the gonads of miniature stallions are smaller. Metcalf et al.2 examined testicular sizes of 23 American Miniature Horse stallions. The heights of the stallions were not reported, but to meet American Miniature Horse Registry requirements these stallion must have been ≤ 96 cm. Testicular volumes were 30.6 cm3 and 31.9 cm3 for left and right testicles, respectively, with a combined testicular volume of 62.5 cm3 (Table 136.2). In a larger study of 216 miniature stallions, stallions ≤ 96 cm in height had a mean scrotal width of 7.25 cm with an estimated total testicular volume of 91.6 cm3 , while slightly larger stallions (97–104 cm) had a scrotal width of 7.95 cm and a slightly larger estimated testicular volume (149.5 cm3 ).1 In that study, testicular height was not measured directly; therefore, the measurement for width was used for height also. Pickett et al.,4 using total scrotal width (TSW) to estimate testicular volume and predict daily sperm production, stated that stallions with a total scrotal width less than 8.0 cm did not meet the standards set for being considered a satisfactory potential breeder regardless of their semen characteristics because of their potential for low daily sperm output. Using this criterion, very few miniature stallions would be considered acceptable. Miniature stallions must be assessed using information derived from miniature stallions. Daily sperm output (DSO) is often estimated using testicular volume (TV) with the formulas: DSO = (0.024)(TV) – 0.765 or DSO = (0.024) (TV) − 1.26.6 More recently, a Bayesian approach was used to estimate DSO and the formula derived was DSO = 0.79 + (0.018 × TV).8 Alternatively, a method used to determine DSO that is commonly accepted as more accurate, is to collect semen from a stallion daily to stabilize extragonadal sperm reserves, after which ejaculates are obtained for analysis of sperm numbers to determine daily sperm output. Studies have shown that daily collection for 5 days, especially in stallions with smaller testes, are sufficient
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to stabilize extragonadal reserves.7 In the study by Metcalf et al.,2 stallions were collected for at least 5 consecutive days before ejaculates were obtained for analysis. From those data, it was suggested that the formula normally used for full-sized stallions is not acceptable for miniature stallions. Using the presumed testicular volume from that study in the original formulas5, 6 would give an expected DSO of 0.24 or 0.74 × 109 , whereas 2 × 109 sperm were obtained after depleting extragonadal reserves. Using the more recently published formula8 would give an expected DSO of 1.9 × 109 spermatozoa. The difficulty lies in using a formula that subtracts a constant from a calculated figure. While suitable for a certain range of testicular volumes, if the volume drops too far below the ‘normal’ range that was used to develop the equation, the resulting expected DSO could easily be a negative number.
Internal genitalia Published data on the accessory sex glands in miniature stallions show a situation similar to that seen with testicular size. When compared with full-sized horses, the accessory glands are smaller in miniature stallions than those of larger horses such as Thoroughbreds (Table 136.3).9 Because the accessory glands are smaller, it would be expected that they would produce a lesser volume of secretions. Given that much of the volume of an ejaculate is from the accessory glands, it is not surprising that the volume of the ejaculate is less than is typical of full-sized stallions.
Breeding soundness Currently accepted guidelines for acceptable reproductive parameters have been developed using stallions of full-sized breeds. The total number of normal motile spermatozoa produced per time unit is a key criterion for deciding the reproductive potential of a stallion. Testicular mass, estimated by calculating testicular volume from measurements of the testes, is directly correlated with sperm production. The larger the testes, the greater amount of testicular parenchyma present and the more sperm produced per unit time. As such, larger testes generally produce more sperm. Given the direct correlation between testicular size and sperm production, total sperm in an ejaculate would be expected to be lower in miniature stallions with small testicles. On the other hand, assuming the testicular parenchyma is normal, parameters of semen quality such as percent motile sperm and percent morphologically normal spermatozoa would not be expected to differ between miniature stallions and full-sized stallions.
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Table 136.3 Measurements of accessory glands from miniature stallions and three breeds representative of full-sized stallions; mean (range) (from Pozor and McDonnell).9 Copyright Elsevier, 2002
Bulbourethral gland (mm) Length Height Prostate (mm) Lobe thickness Isthmus thickness (mm) Vesicular glands (mm) Total diameter Wall diameter Lumen diameter Ampulla (mm) Total diameter Wall diameter Lumen diameter Urethra (mm) Total diameter
Miniature Horse
Arabian
Standardbred
Thoroughbred
21 (16–27) 14 (11–19)
31 (17–42) 20 (12–29)
41 (30–54) 27 (14–32)
37 (28–59) 23 (15–39)
18 (11–29) 8 (4–11)
21 (16–28) 7 (4–11)
26 (10–35) 12 (6–22)
27 (15–40) 8 (4–17)
9 (5–15) 3 (1–6) 4 (2–11)
8 (4–14) 3 (1–5) 3 (0–7)
13 (5–20) 4 (1–8) 5 (1–11)
14 (4–19) 3 (1–7) 7 (1–10)
10 (5–20) 4 (2–10) 1 (0–3)
10 (4–13) 5 (2–8) 1 (0–2)
13 (16–18) 5 (2–7) 2 (0–5)
11 (7–15) 5 (2–7) 2 (0–6)
15 (10–20)
25 (15–35)
28 (19–33)
23 (14–31)
However, breed differences have been reported and may affect semen characteristics. Furthermore, a significant effect of inbreeding on semen quality has been reported, specifically in Shetland pony stallions, with a decrease in semen quality as the inbreeding coefficient rises.10 Even so, the overall percentage of morphologically abnormal spermatozoa is not appreciably different from that observed in full-sized stallions (Table 136.1). Even though sperm numbers in the ejaculate and the volume of the ejaculate are lower than is commonly expected from full-sized stallions, it must also be considered that the uterus of the miniature mare is smaller than that of a full-sized mare. The minimum number of sperm necessary to optimize pregnancy rates in horses is not exactly known. A figure of 500 × 106 progressively motile (normal?) spermatozoa is often cited as a recommended insemination dosage. However, evidence-based studies to support this figure are lacking. Furthermore, there is evidence that much lower numbers of spermatozoa are sufficient to ensure good fertility.11 The number of normal, motile sperm needed in a miniature mare with a much smaller tract is unknown. Similarly, the volume of ejaculate that could be inseminated into the uterus of a miniature mare without losing appreciable quantities through cervical reflux is unknown. It is highly likely that, if an entire ejaculate from a full-sized stallion were inseminated into a miniature mare, much of that volume would be expelled through the cervix. Because percentage of normal sperm and percentage of motile sperm are not different than full-sized stallions, the task of evaluating miniature stallions
is not dramatically different than is commonly practiced with full-sized stallions. Some modifications are necessary during the semen-collection process to obtain the ejaculate, but, once obtained, evaluation of the semen is routine. As long as an expectation of smaller testicular size and lower ejaculate volume, and therefore fewer total sperm, are taken into consideration when evaluating a miniature stallion, a fair assessment of breeding soundness can be accomplished.
References 1. Paccamonti DL, Buiten AV, Parlevliet JM, Colenbrander B. Reproductive parameters of miniature stallions. Theriogenology 1999;51:1343–9. 2. Metcalf ES, Ley WB, Love CC. Semen parameters of the American Miniature Horse stallion. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 202–3. 3. Cox JE. Cryptorchidism. In: Cox JE (ed.) Surgery of the Reproductive Tract in Large Animals. Liverpool: Liverpool University Press, 1987. 4. Pickett BW. Reproductive evaluation of the stallion. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 755–68. 5. Love CC, Garcia MC, Riera FR, Kenney RM. Use of testicular volume to predict daily sperm output in the stallion. In: Proceedings of the 36th Annual Convention of the American Association of Equine Practitioners, 1990, pp. 15–21. 6. Love CC, Garcia MC, Riera FR, Kenney RM. Evaluation of measures taken by ultrasonography and caliper to estimate testicular volume and predict daily sperm output in the stallion. J Reprod Fertil Suppl 1991;44:99–105. 7. Stich K, Brinsko S, Thompson J, Love C, Miller C, Blanchard T, Varner D. Stabilization of extragonadal sperm reserves
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Reproductive Parameters from Miniature Stallions in stallions: application for determination of daily sperm output. Theriogenology 2002;58:397–400. 8. Thompson JA, Love CC, Stich KL, Brinsko SP, Blanchard TL, Varner DD. A Bayesian approach to prediction of stallion daily sperm output. Theriogenology 2004;62:1607– 17. 9. Pozor MA, McDonnell SM. Ultrasonographic measurements of accessory sex glands, ampullae, and urethra
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of normal stallions of various size types. Theriogenology 2002;58:1425–33. 10. Van Eldik P, van der Waaij EH, Ducro B, Kooper AW, Stout TAE, Colenbrander B. Possible effects of inbreeding on semen quality in Shetland pony stallions. Theriogenology 2006;65:1159–70. 11. Brinsko SP. Insemination doses: How low can we go? Theriogenology 2006;66:543–50.
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Section (e): Sexual Behavior
137.
Normal Sexual Behavior Sue M. McDonnell
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Handling The Breeding Stallion Dickson D. Varner
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The Novice Breeding Stallion H. Steve Conboy
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Training a Stallion to the Phantom Glenn P. Blodgett
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Abnormal Sexual Behavior Sue M. McDonnell
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Pharmacological Manipulation of Ejaculation Sue M. McDonnell
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Pharmacological Manipulation of Stallion Behavior Sue M. McDonnell
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Specific Erection and Ejaculatory Dysfunctions Sue M. McDonnell
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Stereotypic Behavior of Stallions Katherine A. Houpt
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Chapter 137
Normal Sexual Behavior Sue M. McDonnell
Natural social organization and behavior Based on observational study of feral and semi-feral populations, the behavior of horses under natural social conditions has been described. The following summary points are useful to understanding sexual behavior of domestically bred stallions. Further details, including video illustrations, are available in recent reviews.1–4 General herd structure and size Natural horse herds include two types of relatively stable subgroups, called bands. One type is known as a harem band. These are the main family social units of a herd. The second common group is known as a bachelor band, composed of males that are not attached to a harem band. These all-male groups are cohesive and relatively stable. A herd typically includes multiple harem bands and one or more bachelor bands. A third type of subgroup which is sometimes seen, and which is less stable and often temporary, is an assemblage of young males and females in transition from their natal band. These are known in the literature by various names, including transitional bands, transitional gangs, or mixed sex peer-groups. Herd size and band size vary among the populations studied and over time, apparently because of available resources and other environmental conditions, with some herds numbering in the hundreds or more. Harem bands Each harem band consists of typically one mature breeding male known as the harem stallion, generally from one to a few mature breeding females, and their young offspring that have not yet left their natal band. Harem bands are generally stable in terms of membership and are cohesive year-round,
with some variation among populations studied. The harem stallion’s behavior appears aimed at keeping the adult mares from exposure to other stallions and to taking the lead in protection of the band from social challenges or predatory threat. Both the harem stallion and the adult mares play a role in protecting the young and in keeping the young from straying from the band or co-mingling with other bands. The harem stallion drives and directs the movement of the harem band whenever it is near other bands or when the band is threatened. The harem stallion performs ritualized posturing and marking responses to voided feces and urine of the mares, called elimination marking sequences. Sniffing and flehmen response appear to represent olfactory monitoring of mare feces and urine. Then immediate urination on top of the mare’s voided urine or feces appears to represent marking or masking behavior. Although the stallion often appears to human intruders as completely ‘in charge’ of all harem activities, his leadership and apparent ‘dominance’ within the band is typically limited to defense from intruders. Leadership of the harem band in terms of transition among ongoing maintenance activities or location of the band in the absense of a threat is characteristically the role of one or more mature mares. So, for example, treks to water, or movement during grazing, or periodic shifts from grazing to resting areas are typically led by one or more mature mares, with the other band members following with the harem stallion trailing at the rear. In established bands, the mares appear cohesive independent of the stallion’s harem maintenance efforts. Transitions of mature mares from one band to another appear limited to socially busy foaling and breeding periods or during periods of sparce resources, such as in late winter or in long dry seasons. Assistant harem stallions A relatively rare variation in harem structure is a second mature stallion, sometimes unrelated, sharing
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Stallion: Sexual Behavior
in harem maintenance activities. The second stallion appears to play the role of an assistant to the main breeding stallion in defense of the band. Typically, the assistant does not breed mature mares, but may breed fillies that remain within the natal band. There is evidence that two-stallion harems are more stable and often maintain a larger number of mares than singlestallion harems in the same herds. Bachelor bands Bachelor bands include predominantly younger stallions that have not yet attained a harem, along with fewer older stallions that have either never attained a harem or are former harem stallions that have been deposed or have lost their harem. Former harem stallions tend to remain loosely attached to the bachelors in a herd, rather than fully integrated into the band. They typically join the bachelors to collaborate in defense of the herd during times of significant threat. The bachelor bands tend to follow the harem bands as close as the harem stallions appear to allow. Bachelor stallions do have some limited opportunities for breeding. Most of their breeding opportunities are what is called ‘sneak breeding’. One example involves young fillies that typically are ‘released’ from protection of their harem stallion or mature mares when they become estrus. The harem stallions often tolerate bachelors interacting with these mares. ‘Sneak breeding’ opportunities also occur during periods of social upheaval and reorganization within the herd, for example when a harem stallion becomes disabled and his band is challenged by other males. A bachelor stallion that is sneak breeding within the vicinity of a harem is typically especially quiet, with limited vocalization that might attract attention of competing males. The stallion often appears especially vigilant to enviromental events. Transitional bands The young typically remain with their natal band for 1–3 or more years. The first year is characterized by a large amount of play behavior, including play sexual behavior that appears to be practicing and linking elements of precopulatory and copulatory behavior into organized adult sequences. Females mature reproductively at about 1 year old and, with good nutrition, may have their first offspring as early as 2 or 3 years old. Young males also mature at about 1–2 years old. The young typically experience a period of coming and going from the natal band, as opposed to an abrupt departure. They may gather with similar aged young from neighboring harem bands or linger near the bachelors during a period of transition from their natal harem band to their eventual adult group.
Although it is not typical, some harem stallions tend to harass their young males when they begin to mature sexually. This can happen in an abrupt explosive event, or gradually over a period of months. But most appear to leave on their own without any apparent encouragement from the harem stallion or adult mares and gradually over a period of a year or more of coming and going. Yearlings and 2-year-olds tend to form temporary affiliations with young from other harem bands. These temporary assemlages can grow into transitional bands of day-to-day variable membership1 of young males or of males and females that may come and go individually from their natal band for as long as a year or more before leaving permanently to join other bands or to form new bands. Fillies often first participate in transitional gangs when they come into their first estrus, at which time their harem stallion tends to ‘release’ them from his vigilant herding. Many fillies are bred by multiple transitional band males. When in estrus these young fillies may venture out from the natal band to contact bachelor band or transitional band males and, occasionally, other harem stallions for minutes to hours several times within a day, while others may join loosely with a transitional or other harem band for the duration of the estrous period. Most fillies return to their natal band for their initial pregnancy and foaling. Once out of estrus their natal harem stallion returns to protecting them as he does the other mares in his band. Affiliations within bands Within bands, there are often close pair-affiliations. Mares within a harem or stallions within a bachelor band may have ‘preferred’ neighbors or grooming partners. Bachelors often form two- and threepartner alliances and appear to cooperate in controlling limited resources, huddling for protection from cold or insects, sequestering a young filly, or fighting off intruders. Communication Horses are known for their multisensory alertness and instantaneous herd vigilance and reactivity to threat. A variety of subtle postures and expressions appear to be important visual elements of communication among the herd members. Within and between bands within the herd, subtle changes in ear or tail position appear to visually convey information about threat and non-threat status. Horses also respond to threat and alarm behavior of other species, including birds. Horses emit a variety of sounds and vocalizations that also appear to communicate within the herd. These include specific vocalizations and breath
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sounds, such as whinnies and neighs, snorts, squeals, grunts, groans, blows, sighs, and sniffs. Hoof sounds, such as pawing, stamping, or contact with the substrate and vegetation during locomotion, also appear to communicate information among horses. Chemical cues also likely play a large part in communication within and between groups of horses, as well as in perception of threatening predators. Stallions display conspicuous elimination and marking behavior sequences in which considerable time and energy are devoted to sniffing excretions of herd mates and of fecal pile accumulations of stallions from competing groups within a large herd. Whenever herd mates or strangers meet, they approach nose-to-nose with deliberate sniffing of one another’s exhaled breath. Many observers have speculated that exhaled breath may carry information about individual identity, relatedness, or social status that are improtant to social order and to reproduction. Among closely bonded band members, particularly mares and their foals, tactile communication no doubt is also important. An interesting behavior that likely includes elements of communication is mutual grooming. Two horses approach to stand facing one another, and then advance to stand parallel head-totail, nibbling insects or tufts of shedding coat from the neck or back of a partner as they advance. In addition to obvious grooming needs, this behavior has been proposed to reinforce trust and bonding among participants. Mutual grooming among bachelor stallions or among youngsters often precedes play bouts. So, in that context, it is considered a play-initiation behavior. Stallions, whether harem or bachelor, appear to be particularly alert and reactive to communication, both within and between species, and general environmental events. This is particularly the case for stallions during copulation, especially for bachelor stallions when ‘sneak breeding’.
head toss threat or threat gaze that involves a stare with the ears held back toward the neck and the head lowered. While the dominance heirarchy within bands is typically complex and situation dependent, dominance order among the bands within a herd often appears more linear and more consistent among situations and over time. Certain bands appear dominant over or subordinate to others in terms of leadership of activities and order of use of limited resources such as water, dustbathing bowls, shade, and shelter.
Dominance heirarchy
Annual reproductive patterns
Established order of dominance and submission are an important factor in the social order within and between bands within a herd. In well-established herds, social order within and between bands, though often situation-dependent and not always simple or linear, is clearly evident. There is little overt fighting even in the face of limited resources. Most competition for limited resources is settled with simple threats and retreats. For open plains species, an important element of avoiding fights is simple retreat from the threat. Submissive animals usually avoid attack by ‘keeping their distance’. Clearly dominant individuals can effectively direct the movement of other animals or can control a limited resource with a simple
Under natural conditions, foals are born at the time of year corresponding to best vegetation and weather conditions for survival of the young. In climates with distinct seasons, most foals are born in late spring. Exceptions in free-running populations have been noted in which there are examples of foals born in nearly every month, although out-of-season births and foal survival are apparently rare. With gestation of 11 months and ovulation at about 2–3 weeks post partum, most breeding occurs at more or less the same time of year as foaling. Mares within a band charcteristically foal and breed within a similar few weeks, during which the stallion is conspicuosuly more intently vigilant and protective of his band.
Foraging and feeding Horses are grazers that can also browse. They appear to prefer grasses and legumes when available, but will also browse shrubs, woody plants, leaves, and roots. For horses living on grasslands, the majority of their time is spent foraging and grazing. Depending upon forage quality, horses under natural conditions typically spend from half to three-quarters or more of their daily time budget grazing. Unlike ruminants, horses may graze continuously for many hours without stopping. When grazing, they move continuously – taking a few bites, taking a few steps forward, taking a few bites. Grazing bouts are usually interspersed with periods of loafing, and the entire herd simultaneously grazes or rests. Depending upon the terrain and distance from water, horses may go to water only every 2–3 days. In wet seasons with lush grasses, horses can obtain sufficient water from the vegetation. When water sources are freely available the usual drinking frequency is once or twice a day. The bands of a herd usually trek to water in single file, each band with a mare leading and the harem stallion herding at the rear. As the bands approach the watering site, there is typically a fixed order within each band and between the bands for approaching the water.
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Stallion: Sexual Behavior
Foals nurse most intensely from birth through 4–6 months of age. They continue nursing on a less intense schedule until the dam foals again, and some even beyond that. The reproductive life of the healthy mare in reasonable body condition is to foal annually, and to lactate most of the year. Horses under natural conditions commonly live up to 15–20 years, and many mares will have foals almost annually from 2–3 years of age until death. Courtship and copulation When a mare is in estrus, the harem stallion usually lingers nearby, grazing and resting as the nearest herd member, with increased attention and elimination marking response to her frequent urinations. This has been called ‘tending’ of the mare. Frequent bouts of sexual response and precopulatory interaction continue throughout the duration of estrus. Proceptive responses of the mare includes moderately increased locomotor activity, frequent approaches to the stallion, swinging of the hindquarters toward the stallion, lifting the tail, and frequent urinations with prolonged rhythmic eversion of the vulva exposing the clitoris and lighter internal membranes. Sudden jerky movements of the mare appear to stimulate bursts of interest arousal by the stallion. Stallions and mares breed many times a day when the mare is in estrus, and often there appears to be a rhythm of teasing and breeding at fairly regular intervals, as often as once hourly throughout the day and night. The female receptive mating stance includes a ‘sawhorse squat’, with the tail lifted off to the side of the perineum, the head turned back toward the stallion, and one foreleg flexed. This head-back posture with one forelimb flexed appears to be especially inviting for mounting, as if it signals to the stallion that she will not resist mounting by either kicking or moving forward. It is common for stallions to mount once or twice without erection, as if testing the receptivity of the mare, before mounting with erection and insertion. Ejaculation typically occurs on the first mount with erection and insertion, and the usual pattern includes one or more shallow tentative thrusts followed by seven to nine rhythmic committed deep pelvic thrusts. The mare may adjust her posture to guide insertion and step forward to accommodate thrusting. Animals conspicuously use terrain to overcome discrepant size. Ejaculation is evident with rhythmic rostral–caudal movement of the tail corresponding to urethral ejaculatory contractions along with relaxation of the hind limb and neck muscles. The mare is generally the first to move to disengage, which usually commences many seconds after ejaculation. Ejaculation usually occurs within 18–20 seconds after insertion and 20–22 seconds after mounting. Total
mount time is typically 25–35 seconds. Most copulatory interactions occur within less than a minute, and often are relatively quiet and may go unnoticed when the observer is not specifically scanning the herd each minute for breeding. The stallion and mare usually linger near one another quietly for several minutes following copulation. The stallion may sniff and perform flehmen over spilled vaginal fluids and ejaculate. The mare may urinate and expel vaginal fluids and semen. The stallion may investigate and urinate over those fluids, as if marking or covering them. A conspicuous feature of mating among free-running horses is the strong role the female appears to play in maintaining attention of the stallion, seemingly determining the time of copulation, as well as postural adjustment to facilitate successful mounting, insertion, and dismount.
Parturition guarding and parental behavior There is little observable change in behavior of the mare with pregnancy until parturition commences. Most parturition occurs in the evening or near dawn. Some especially vigilant stallions appear to recognize imminent parturition and sequester their harem away from other bands even for days before parturition commences, and may continue to remain at a distance from other bands and for the first days or week after parturition. Parturition is typically extraordinarily rapid under natural herd conditions, with typically only a few minutes from any observable sign of labor until the foal is fully expelled. As the mare initially shows signs of abdominal discomfort, getting up and down frequently, the harem stallion may appear stimulated to a heightened level of alertness and vigilance. Some harem stallions anxiously patrol the perimeter of the band at a trot, stopping to posture and mark areas with feces. The harem family members, including the harem stallion, other mature mares, and the young, may approach and/or gather around the foaling mare, sometimes nuzzling and vocalizing to the foal during and immediately after parturition. The dam and sometimes the stallion immediately attends the foal with licking that clears the birth membranes. Both the stallion and the dam sniff and may perform the flehmen response to the expelled fetal fluids. The harem stallion often appears urgent to move the band away from the birth site. He may paw, nudge, nip, or head-butt a recumbent dam with the nose while gesturing herding movements. In some cases, the stallion moves the band away from a birth site before the fetal membranes are passed, and then moves them on again after the membranes are passed. Unlike some species, it is rare for mares to ingest fetal membranes.
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Normal Sexual Behavior The harem stallion as well as all other band members participate in parenting and protection of the young. For the first week or two, the foal’s nearest neighbor is almost always the dam, but, should any disturbance occur, the harem stallion aids in protecting the foal. After about 2 weeks, the stallion takes on an increasing role in parenting the young, including retrieving straying foals and yearlings, and initiating or joining in group play of foals and older juveniles and young adult males. Incest avoidance Although the mechanisms are not understood, it is known that genetic diversity is reasonably well maintained in free-running equid populations. Where behavior observation and parentage data are available, it has been observed that related pairs do not breed when unrelated males are available and that harem stallions are generally not sexually responsive to fillies born within their band. Spontaneous erection and penile movements (masturbation) All male equids experience spontaneous erections and penile movements at regular intervals averaging 90 minutes throughout the day and night. This apparently involuntary behavior occurs during all quiet activities and quite reliably occurs after periods of rest or as a period of excitement resolves to calm. Ejaculation ocuurs only rarely.
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location and sociosexual exposure of various management conditions and breeding programs without significant problems. With good handling, most stallions can also transition with little difficulty between breeding methods, including pasture breeding, natural cover in-hand, or for semen collection with a mare, dummy mount, or on the ground. Also, in spite of seasonal variation in reproductive hormones and behavior, most stallions can breed in any month of the year. For most stallions, response latencies and refractory periods are conspicuously greater in fall and winter than in the late spring and summer natural breeding season. Pre- and post-copulatory behavior varies considerably in hand-bred stallions depending upon the level of access the stallion is given to mares in general and at the time of breeding as well as by the style and skill of the handler and the specific farm protocol for breeding. For example, some domestic stallions are more or less kept in isolation from mares except for the few minutes of breeding access that may be relatively infrequent. In general, these stallions tend to be highly animated and vocal for that very limited period of social exposure. On the other hand, stallions managed with ongoing opportunities for contact or communication with mares and frequent breeding tend to be less frenetic and urgent to breed when exposed to mares. Copulatory behavior of domestically managed stallions, on the other hand, is much less variable than pre-copulatory behavior and more closely approximates behavior of stallions bred under natural conditions. Table 137.1 summarizes measures of precopulatory and copulatory behavior of a
Development of sexual behavior Play sexual behavior commences as early as one day of age and continues to represent a significant percentage of play interactions through the first year. Normal adult precopulatory and copulatory sequences have been observed in colts as young as 8 months, with confirmed siring of foals as early as 10 months of age.
Domestic breeding conditions Modern management of domestic horses includes diverse management schemes that include diets, social and physical environments, housing, exercise, work, and other breeding husbandry practices that vary considerably from natural conditions and that likely influence reproductive behavior. Normal behavior and breeding expectations Most stallions readily adapt to the unnatural demands and often changing conditions inherent to domestic breeding. Most stallions can transition among
Table 137.1 Sexual response of normala stallions Range
Mean
Pre-copulatory end points (n = 100) Frequency of pre-copulatory responses before first mount: Sniff/nuzzle mare Lick mare Flehmen response Nip/bite mare Kick/strike mare Vocalize Time to erection (s) Time to first mount with erection (s)
0–80 0–20 0–10 0–25 0–10 0–35 0–500 10–540
6 1 2 1 1 3 67 100
Copulatory end points (n = 33) Number of mounts to ejaculation Mount time for ejaculatory mount (s) Number of thrusts on ejaculatory mounts Total time in breeding area (s)
1–3 17–45 2–12 30–240
1.4 27 7 150
a
Considered normal by owners and veterinarians. (Reproduced from McDonnell.6 )
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sample of hand-bred stallions that were considered by their managers and veterinarians as normal. Breeding frequency expectations vary widely between programs and individual stallions. In general, stallions bred under domestic conditions are not able to achieve or sustain the frequency of breeding that has been observed for harem stallions that under natural conditions breed almost hourly while their mares are receptive. Stallions breeding by artificial insemination are generally expected to ejaculate three to seven times per week. Successful breeding by natural cover generally requires more frequent ejaculation, especially during the peak breeding season months. Traditionally, two or three breedings per day, with efficient latencies to mount and ejaculation, for 4–7 days per week was considered good sustained libido. With the increased book size of the elite Thoroughbreds in recent years, many stallions have been achieving three to five breedings per day, 7 days per week, for much of the 4- to 5-month breeding season to serve 150 or more mares per season.5 Numerous studies have indicated that in domestic in-hand breeding, both for natural cover and for semen collection, stallions often require more than one mount to achieve ejaculation. Although more than one mount has been accepted as normal, it is different from pasture breeding in which stallions almost always ejaculate on the first mount with erection. The additional mounts for hand-bred horses likely reflects the suboptimal conditions inherent to domestic breeding, including less than ideal footing in some instances, suboptimal timing for breeding, less than ideal handling, and other factors. Spontaneous erection and penile movements are commonly called masturbation in domestic horses. It is similar to that of free-running horses. When undis-
turbed, domestic stallions exhibit erection and penile movements at approximately 90-minute intervals throughout the day and night. The rate, duration, and intensity of masturbation is unaffected by age, housing, exercise, or seasonal or breeding management. Factors influencing sexual behavior Within the range of normal, stallion behavior is conspicuously influenced by a number of factors, including breeding environment and method, the style of handling for breeding, behavior of the stimulus mare, sociosexual exposure,7 breeding schedule, season, exercise, and fitness. Casual observation suggests that sexual behavior, including general level of libido as well as specific pre-copulatory and copulatory style, are also genetically influenced.
References 1. Waring GH. Horse Behavior. Park Ridge, NJ: Noyes Publications, 2002. 2. McDonnell SM. The equid ethogram. In: McDonnell SM (ed.) A Practical Field Guide to Horse Behavior. Lexington, KY: Eclipse Press. Available with DVD video companion. See HorseBehaviorDVD.com, 2003 3. McGreevy P. Horse Behaviour: A Guide for Veterinarians and Animal Scientists. Philadelphia, PA: Saunders, 2005. 4. Mills DS, McDonnell SM. The Domestic Horse: The Origins, Development and Management of its Behaviour. Cambridge: Cambridge University Press, 2005. 5. Turner RM, McDonnell SM. Mounting expectations for Thoroughbred stallions. J Am Vet Med Assoc 2007; 230:1458–60. 6. McDonnell SM. Reproductive behavior of the stallion. Vet Clin North Am 1986;2:535–55. 7. McDonnell SM. Reproductive behavior of stallions and mares: comparison of free-running and domestic in-hand breeding. Anim Reprod Sci 2000;60–61:211–19.
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Chapter 138
Handling the Breeding Stallion∗ Dickson D. Varner
Effective communication Volumes have been written regarding methods for communicating with horses, and working clinics on the subject have been popularized in recent years. The concept is simple – use a mindful approach to interaction with your horse, realizing that it cannot speak your language but that it can respond in some manner to virtually everything that you say or do. This same approach is useful for handling the breeding stallion. Horse behavior is an acquired trait, and horsemanship skills are required to mold a horse’s behavior into one of favorable quality. To develop a unique language that both understand, we need to learn to think as they think, hear what they hear, and see what they see. This process begins by monitoring the actions of the horse in your presence, however subtle that they may be, so that you may refine these actions to capture the horse’s mind and body. As an end product, a stallion should demonstrate respect for a handler and be responsive to cues, while remaining vibrant, curious, and energetic, as opposed to subordinate or distrusting. These training methods are most easily applied when horses are young, when bad habits have not developed that require correction. Nonetheless, one can teach older horses new tricks. In my view, a well-behaved, yet vibrant, stallion is obtained through patience, persistence, and positive reinforcement on the part of the handler during the formative sessions. The term positive reinforcement is not applied correctly in the eyes of learned behaviorists. We use pressure and release tactics to train horses, i.e., negative-reinforcement training methods.
Nonetheless, positive-reinforcement methods, such as a pat on the forehead for a job well done, are interjected into the training process, to create a positive learning experience. Oftentimes, older horses have been allowed to repeatedly exhibit inappropriate behavior or have been subjected to excessive or overzealous reprimand for misconduct. Either of these scenarios generally requires significant retraining to erase the presenting problem while simultaneously creating a positive learning experience.
General principles for handling stallions First interaction First, one is ill advised to apply generalities to all stallions because each is, and should be treated as, an individual. The presenting behavioral traits of stallions can range from that of extreme timidness or withdrawal to that of unruliness or viciousness. Actions on the part of the handler should be based on the presenting behavior of the stallion. This is where the ‘feel’ component of horsemanship comes into play. Learn to read movements and gestures of a stallion when approaching him. Watch him in a paddock setting. Watch his behavior when entering his stall. Is he timid, frightened, anxious, or aggressive? Get some historical information regarding the stallion’s behavioral habits prior to your first encounter with him. Does he strike or bite at his handler? Does he wheel and gesture to kick when entering the stall with him? Has he been exposed to previous handling techniques and, if so, what kind? In other words,
∗
This chapter was reprinted with permission from the American Association of Equine Practitioners (Varner DD. Handling of the breeding stallion. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners, 2005, pp. 498–505).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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learn all you can about the stallion before directly interacting with him. Second, never be absolutely trusting of a stallion in any situation. Stallions have hurt scores of handlers. Some of these stallions were generally considered to be well mannered and trustworthy, but have attacked handlers without provocation. To avoid this situation, always be on guard around a stallion by taking a better-safe-than-sorry attitude. This does not imply that one should exert force with a stallion to show dominance. It only means that becoming too relaxed, lackadaisical, or distracted in the presence of a stallion can lead to disaster and can be averted in most instances by paying close attention to the stallion and the other activities occurring in close proximity to the stallion. Restraint A key element of stallion handling is minimal but effective restraint. My first choices in equipment are a well-fitted leather halter and a chain/leather shank. I recommend a 30-inch (0.8-m) chain attached to an 11-foot (3.4-m) leather lead strap with a leather button secured to the end of the shank. Most halters only have crown adjustment points. The crown strap should be adjusted so that the two stays connecting the nose strap, cheek pieces, and chin straps are immediately below the lower end of the facial crests. The halter (including the nose/chin straps and the throatlatch) should not be too loose or tight on the stallion’s head. I prefer stays that are made of round brass stock rather than flattened stock because it more readily accommodates a shank snap and does not impede the chain sliding through the stay (Figure 138.1). The chain shank is very versatile, as it can be snapped into the lead snap ring, or fitted under
(a)
(b)
Figure 138.1 Photographs from the right side of the stallion, demonstrating chain snap placement in (a) the right cheek ring or (b) the right stay. If the snap is placed in the right stay, the chain should first pass through the right cheek ring.
the chin, over the bridge of the nose, or through the mouth for increasing levels of restraint. The halter and attached lead should only be fitted to the stallion’s head when the stallion is in a confined area to prevent escape. In all instances, the handler should stand at the side of the stallion when securing the halter and attached shank. Unless the shank snap is simply secured to the lead snap ring, the chain should be placed though the left stay, then under the chin, over the nose or through the mouth, then through the right stay and affixed to the cheek ring on the right side, or through the right cheek ring and affixed to the right stay. If the stallion is unaccustomed to chain or bit placement in the mouth, or resists this effort because of previous negative experiences, it will be necessary to take the extra time to make this a comfortable experience for him. Repeatedly ask for him to open his mouth and accept the chain, by placing the left middle finger in the interdental space to open the mouth. In all instances, avoid forcing the chain into the mouth because this is inevitably met with resistance by the stallion, and the problem will only worsen with time. Some stallions are best handled by placing the chain in the mouth after it has first been left loose under the chin while the other side is fastened. This avoids dragging the chain through the mouth or attempting to fasten the chain to the cheek stay if the stallion resists a chain in his mouth. In addition, the snap part of the chain fastener should face away from the skin of the stallion so that it has less chance of being inadvertently opened. Some stallion handlers prefer Chifney bits for restraint. These bits are easily affixed to a halter or can be fitted with a head strap. I do not advocate use of these devices for breeding stallions, as the protrusion of the bit frame underneath the chin can be inadvertently bumped on a mare or dummy mount and distract the stallion when breeding. Many stallion handlers use various bits that can be fitted to the stays of a halter or that can be attached to a head strap. The mouthpiece can be broken or solid, beveled or round, steel, brass, or rubber, and some have players or rollers in the central portion of the mouthpiece. An advantage of bits over a chain is that they can be secured to the halter by standing only on the near (left) side of the horse, thereby reducing the risk of personal injury when passing to the right side of an aggressive or frightened stallion. In my view, it is unadvisable to use a rope halter and attached lead (like those sold for ‘natural horsemanship’) for breeding stallions unless they are extremely docile and their mannerisms in a breeding environment are well known. Remember, the only thing predictable about a stallion is its unpredictability.
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Handling the Breeding Stallion
Preparatory training methods A breeding stallion should readily exhibit two important maneuvers on command before engaging in the breeding process – ‘stop’ and ‘back’. The maneuvers should be taught in an area free of distractions, especially other horses. In fact, the rudimentary aspects of these maneuvers can be taught right in the stall. The stallion can then be walked into a more open space for more extensive training. The area should be free of obstacles to avoid injury to the stallion or personnel. Before beginning the training process, remember that the stallion will not learn well when distracted, tired, mad, hurt, or frightened. It is important to get the stallion in the proper frame of mind before initiating specific training techniques. When handling a stallion, allow him to work on a slack lead rather than applying constant contact. If he charges ahead of the handler, a sharp jerk on the shank should correct this behavior, but the shank tension should be immediately released. The process can be repeated until the stallion responds properly. Allow the stallion an opportunity to make, and learn from, mistakes. Once a stallion is under control at a walk, stop with the stallion on a loose lead, with the expectation that the stallion will stop simultaneously. If he continues to walk, apply a sharp jerk on the shank with sufficient force to stop the horse in his tracks. Repeat this procedure until the stallion stops immediately on a loose lead when the handler stops. This process teaches the stallion to read the body language of the handler and to respond accordingly. After the stallion has mastered the stop, work on the backing maneuver by turning toward the stallion and walking parallel to this left side. If he does not back, apply pressure with the shank to encourage him to back. This process should be repeated until the stallion backs on a loose lead when the handler turns and walks toward him. With time, these maneuvers become automatic to the stallion. Last, teach the stallion to back in a circle by walking toward his left shoulder and encouraging him to move the forequarters to the right (with the hindquarters moving to the left) while backing. This maneuver is very important because it prevents a stallion from circling with forward motion around the handler when excited in a breeding environment, and helps assure proper alignment of the stallion with the mare or breeding dummy prior to mounting. A stallion should never be out of control in a breeding shed. The two most common problems encountered are charging the mare or dummy mount and circling around the handler during approach to the mount. These two problems can be averted or corrected by ensuring that the stallion is responsive
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to the cues for stopping and backing, as described above. As with other animal training techniques, a key element is to avoid mixed signals during the training sessions. It also is important not to constantly ‘pick’ at the stallion, as it will dull its senses. Once a correction is made, leave the stallion alone to become comfortable in the ‘safe zone’ that you create for him. On a similar note, do not offer treats to a stallion during the training process. This practice, sometimes called a form of positive reinforcement, only encourages the stallion to enter ‘your space’ and to become ‘nippy’ or ‘mouthy’. The reward for the stallion should simply be release of pressure. The purpose for the preparatory training session(s) is to teach the stallion to listen to, and respond to, the handler in a relatively distraction-free setting. Some stallions, especially those that are fractious or overly aggressive, will require some roundpen training without a lead so that they may ‘connect’ with the handler. Stallions that are responsive in the round pen are universally more cooperative when in hand. It is also beneficial to teach socialization skills to stallions by applying the same groundwork techniques when in the vicinity of other horses. After many years of exposure to breeding stallions, I would consider western performance stallions (cutting, reining, roping, ranch, and reined-cow horses) to be the best behaved. It is customary for trainers to teach these stallions to maintain a cooperative attitude in the midst of other horses. This form of discipline is usually started a young age, i.e., as yearlings or 2-year-olds, and is instilled through daily exposure to this setting. Stallions managed in this manner usually display excellent obedience when breeding.
Breeding shed exposure Once a stallion has passed these initial tests, he can be introduced to the breeding environment. An important training tip is to introduce a stallion to one new thing at a time. For instance, allow a stallion to become accustomed to the floor surface of the breeding area, and then to any unique structures in the breeding area (such as the teasing stocks, breeding dummy, and mare breeding cape), before introducing him to a mare in estrus. The approach to training a breeding stallion depends on the stallion’s specific mannerisms in the breeding area. For instance, a timid novice stallion will be handled differently than a stallion that requires retraining because of overly aggressive behavior. Nonetheless, the objective is to train the stallion to be eager and confident, but immediately responsive to the commands of the trainer when breeding a mare or mounting a breeding dummy.
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Stallion: Sexual Behavior
Figure 138.2 For a stallion’s first exposure to an estrous mare, or if the stallion is expected to be unruly, it is wise to begin the approach process by first exposing the stallion to a mare protected within a stall, or by a padded teasing rail.
First, the stallion should demonstrate strong interest in a solicitous mare, but should yield to a command to control his approach to the mare. This response is paramount to the safety of the stallion, mare, and all personnel in the breeding area. To ensure that this feature is intact, it is wise to first approach an estrous mare that is in a stall or behind a padded tease rail, as opposed to one that is fully exposed to the stallion (Figure 138.2). When the stallion develops an erection, it is an excellent opportunity to teach the stallion to tolerate rinsing of the penis with water. To perform this procedure, the stallion should be backed into a padded corner for restraint and to reduce the likelihood of hind limb injury. The first penile washing should be done with very warm water, using a towel as opposed to a water hose, as the massaging nature of the towel on the glans penis stimulates sexual excitement. Stallions readily accept this procedure once they know it is a comfortable experience. Once any aggressive mannerisms of a stallion are controllable when in the presence of the mare, it is then permissible to place her on the breeding shed floor. If the stallion is to breed the mare, or if the mare is to be used as a mount source for semen collection in an artificial vagina, she should be properly outfitted and restrained to reduce the likelihood of injury to the stallion or the semen collector. Application of a lip twitch and breeding boots is recommended. Mare hobbles are sometimes used, but are inadvisable if a mare is not accustomed to wearing them. It is also wise to fit the mare with a breeding cape if one suspects that a stallion may savage the mare during the act of mating. Before allowing the stallion to mount a mare or breeding dummy, make certain that the area is free of obstructions and has a non-slippery surface and a
perimeter fence or wall. An outdoor paddock with a grass surface serves this purpose well. A breeding shed floor must have good footing for horses and personnel. Stone dust, clay, dirt, and tan bark are commonly used as floor surfaces. All furnish a non-slippery base but are difficult to clean. Artificial floors have been installed in some breeding sheds. These floors are usually made of a durable and easyto-clean rubberized material. Some types do not provide good footing, especially when wet. The stallion’s approach to a mare (or breeding dummy) should be controlled at all times. The preparatory training methods help ensure that control is not lost; however, the distraction of an estrous mare on the open floor increases the level of difficulty in commanding a stallion’s attention and respect. Begin the approach at a slight angle to the left of the mare’s hindquarters, and approximately 6–7.5 meters from the mare. Advance slowly, and frequently ask the stallion to stop and back if he becomes hurried. Do not allow the stallion to circle in an attempt to come within reach of the mare. If he begins to circle while advancing forward, back him in a circle by walking toward his shoulder, as was discussed above. Good libido is sought, so it is not appropriate to reprimand the stallion simply for vocalizing and becoming excited. Remember not to use continual tension on the lead shank. The objective is to ask the stallion to approach the mare in a deliberate, but controlled, fashion. Concentrate on knowing how and when to right wrongdoings. Timing and severity of corrections are critical to developing an energetic, but well-behaved stallion. Learn to read the stallion and correct misdeeds at their outset, but do not overdo punishment in the presence of the mare. If the stallion appears to be uncontrollable, remove him from the breeding environment, and go back to the basics of stopping and backing until you are convinced that the stallion is attuned and responsive to your body language. When you are in the breeding shed with the stallion, you should have the sensation that he has one eye on the mare and one eye on you, and is incessantly asking your permission to go forward. When the stallion establishes contact with the hindquarters of the mare without charging forward, allow him to mount the mare at will. Position both the mare and the stallion to help assure proper alignment during the mounting process. Do not tolerate any transgression, such as striking or vicious biting. Leniency with bad behavior will invariably lead to accentuated forms of the problem. When the stallion is mounted on a mare or dummy, maintain him on a loose lead as much as possible. If corrections are needed, such as with savagely biting the mare, apply disciplinary action, then release the pressure. Upon
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Handling the Breeding Stallion the completion of mating or semen collection, allow the stallion to dismount slowly, and then back him from the hindquarters of the mare. As discussed above, the basic concepts of stallion handling can be applied universally, but specific training strategies should be tailored to individual stallion behavioral traits. For instance, shy stallions may warrant some tolerance in misbehavior so as to not dampen their interest in breeding. As they become more aggressive breeders, one can fine-tune their mannerisms in the breeding shed. On the other hand, overly aggressive stallions may require more forceful reprimand. For instance, stallions that have developed tendencies to rear, strike, or bite at handlers when in the presence of a mare will require
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remedial training before reintroducing them to a mare.
Conclusions Reading script does not translate to immediate proficiency in stallion handling techniques but, with practice, adeptness will be forthcoming. Begin with a seasoned and well-mannered stallion to develop a feel for the desired product. Seek guidance from professional horsemen and stallioneers when perfecting your skills. The reward will be quite gratifying. You and your client will have a safer and more serviceable stallion. Make it fun, not fearful, for the horse and handler!
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Chapter 139
The Novice Breeding Stallion H. Steve Conboy
The stallion can be used to breed mares by two methods: pasture breeding, i.e., running free with a band of mares, or in hand and managed by man. When free to run with his band of mares, the stallion learns by trial and error when and how to breed his mares. While he appears to be the dominant member of the band, he has learned that when the mare is not in estrus he will be punished severely for making unwanted sexual advances towards her. During this learning process it is not uncommon for him to sustain significant injuries. He also learns that the mare will require him to be slow and gentle during the act of copulation. The young stallion that is to be hand bred is usually a valuable horse whose owner wants to protect from injury, thereby electing to closely control his breeding activities. The handler assumes the responsibility of protecting both the stallion and the mare(s) during the breeding process. It is easy to appreciate the time and patience that are required for training a well-mannered breeding stallion if you have ever seen an unruly stallion dragging his handler at the end of a lead shank as he charges a terrified mare in the breeding shed. This is not an uncommon occurrence when a novice horseman decides to stand a stallion. It creates a very dangerous situation for the personnel, the mare, and the stallion, and can be avoided by taking the time to properly train the novice stallion in the basics of breeding etiquette. If for no other reason than for safety and to reduce liability, all major breeding farms train their new young stallions well in advance of the commercial breeding season. There is also a great deal of pride in presenting a well-mannered stallion to the breeding program. Knowing how a breeding stallion is expected to behave enables the practitioner to influence the proper development of a young stallion. Prior to beginning a breeding career, it is desirable for the novice stallion to receive a period of ‘let down’ (several months if possible) from the routine of trav-
eling and training. This rest will allow time for his system to adjust to the different lifestyle he will experience as a breeding stallion. A mandatory quarantine period will apply in some cases. If not, a 30-day quarantine should be considered. While it is not uncommon for a successful show horse to be used for breeding while continuing his show career, it may be difficult for the stallion to focus on such a dual career. The young colt that is selected for breeding is usually one that has had a successful career as a performance or show horse and has often been subjected to medications and disciplinary techniques that alter libido. During the rest period he should be encouraged to demonstrate normal libido and should never be disciplined for developing an erection when in the presence of a mare. During the let-down period, routine health maintenance should be performed. Performance injuries and lameness should be addressed and treated. Vaccination, deworming, and dental care done during this period will prevent delays during the breeding season. Always remember that the time required for spermatogenesis is about 60 days, so anything that has the potential of causing a significant fever 60 days prior to the breeding season should be avoided. A titer for equine viral arteritis (EVA) should be taken as soon as possible. If the stallion has a negative titer, vaccination for EVA should be done and 28 days of isolation is recommended following vaccination.1 If he has a positive titer. without proof of a negative titer prior to vaccination, a semen virus isolation test must be done when a semen sample is available. A test for equine infectious anemia should also be conducted. Stallions that are subject to regulations for contagious equine metritis (CEM) testing must submit to such testing, as prescribed by the current regulations. In the absence of import requirements or code of practice regulations, it is a wise practice to collect a set of CEM cultures for testing (see Chapter 247) prior to the stallion being used for breeding.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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The Novice Breeding Stallion During routine grooming, the stallion should become accustomed to having his testicles and sheath touched, which will make cleaning, culturing, and evaluating the external genitalia easier. He should likewise become comfortable with having his temperature taken rectally. The goal of developing the novice stallion is to create a stallion that, when presented to an estrous mare or phantom, will demonstrate controlled enthusiasm, develop an erection quickly, mount gently, and ejaculate promptly. Each of these traits is achievable if proper training is conducted in advance of the breeding season. The basic concepts for training the novice stallion are no different than those applied to training a show or race horse. In order to get the best results, the colt must be given the opportunity to learn how to behave so as to prevent injury to himself, and others, during the act of breeding. Stallions that are out of control while breeding are dangerous to the personnel, as well as the mare being bred. Bad habits that are allowed to develop are difficult to correct because the stallion becomes so excited during the act of breeding that it is difficult to reason with him. All too often, the novice stallion is presented to a mare in need of breeding before he is properly trained. The objective then becomes to ‘just get the mare bred’, and the stallion is given free reign. By doing so, he is greatly rewarded for his behavior, which is usually uncontrolled chaos. There are several approaches to training the stallion in the basics of being handled, but the goal should be to develop his respect for the handler without creating animosity, fear, or shyness. The last two traits are a result of poor horsemanship and skill, and will result in a need for excessive restraint when handling the stallion. This situation is unpleasant for both the handler and the horse. A properly trained stallion will be enthusiastic when presented to the mare but will be responsive to the cues of the handler, waiting to be directed in the breeding process.
The breeding environment Regardless of whether the breeding area is a modern breeding shed, a barn aisleway, or a dedicated area outdoors, it must be of adequate size to turn the stallion, position the mare, and to safely work around both the mare and stallion. If a phantom mare is to be used, an area for the tease mare is necessary and an area to safely wash the stallion’s penis is needed. The area should be free of tools, vehicles, and other equipment that would be a hazard to the horses or personnel. The surface should provide good traction and should be as dust free as possible. Concrete and asphalt should be avoided. When a phantom is used,
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Figure 139.1 Stallion being led with loose shank.
it should be sturdy and easily adjusted. A single support base that is padded is preferable. The covering should be padded and free from torn material in which the stallion could get his foot caught. The phantom should also be free of squeaks and rattles that might distract the novice stallion.
Training to lead, stand, and back Training the novice stallion should begin with ensuring that he understands the basics of being handled, groomed, and walked under complete control. The first lessons should be directed at teaching him to walk quietly on a loose shank (Figure 139.1). The proper restraint for these training sessions requires a leather stallion halter and a brass chain shank, or bit if necessary. The chain can be placed in the mouth, over the nose, or under the chin; however, until it is ascertained that the stallion respects the handling, it is preferable to place the chain through the mouth. Proper use of the chain shank requires special attention to its free movement through the halter ring. If it is attached so that it remains tight when pressure is released, the stallion is likely to react violently by running backwards, rearing, or flipping over backwards, thereby increasing the risk of injuring himself and/or the handler (Figure 139.2). It is the handler’s responsibility to insure that all restraint devices be used humanely and with the least amount of pressure possible during the training process. Initial training should be done in an environment with limited distractions such as in an arena or barn aisle, because effective learning requires his undivided attention. As he masters these basics and becomes responsive to gentle pressure and soft voice commands, he should then be tested and worked where there are distractions. This will prepare him for the excitement of the breeding situations. If he pulls or strides ahead while being led from the shoulder, a sharp snatch on the loose shank will reprimand
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Figure 139.2 Stallion with a halter. Notice the round ring on the halter that allows the chain shank to slide freely without the potential of getting stuck.
him and a quick release reinforces the principle of loose control. He should become accustomed to voice commands for stopping and backing. The stallion should be taught to back by facing him and walking parallel to his left side. At first, pressure on the shank or a gentle jerk will be necessary but the goal is to have him learn to back on a loose shank when approached from the front and directed to back by soft voice command or by simply walking toward his shoulder (Figure 139.3). He should also be taught to respect the handler’s space by yielding when approached at the shoulder. While being held on a loose shank on his left side, he should be taught to turn away to the right by merely approaching his shoulder on an angle and to back in a circle. This maneuver will help to keep him from trying to circle out of control around the handler. This habit is quite annoying when trying to get him backed into a corner for rinsing the penis or when trying to get him to mount for breeding.2
Figure 139.4 Stallion with a leather muzzle. The leather muzzle will prevent the stallion from biting the handler or the mare.
When the stallion has mastered the basics of walking on a loose shank, standing quietly, and backing both straight back and in a circle, he should be taught to do so in the presence of a mare in behavioral estrus. When he is comfortable in the presence of a mare, he is ready to begin his training for the actual act of breeding. The novice handler is cautioned to never fully trust the stallion even though he seems to be completely safe. As he becomes more sexually mature his hormone levels may cause him to react in an aggressive manner spontaneously. The handler must always anticipate the potential for this. If the stallion does become unpredictably or persistently aggressive, it is advisable that he wear a leather muzzle to prevent him from savaging the handler or the mare (Figure 139.4). Full or half blinkers may be helpful in keeping the stallion focused and in preventing injury to the handler.
Training in the breeding shed
Figure 139.3 Teaching a stallion to back by approaching him from the front.
Prior to entering the breeding shed for training or breeding, the stallion should be groomed to remove loose hair and dirt. His shoes should be removed or covered to avoid injury to the mare or damage to the phantom. Depending on the stallion’s temperament, a long leather stallion shank with a 24- to 36-inch (60to 75-cm) brass chain should be passed through the mouth, over the nose, or under the chin. Avoid doubling the chain and leaving a loop that could entrap the stallion’s foot if he were to rear or strike out.
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The Novice Breeding Stallion
Figure 139.5 Mare in a teasing chute. A padded teasing chute is ideal to restrain the tease mare. If a phantom is used, it should be close to the tease mare.
Some stallioneers prefer to turn the novice stallion loose in the breeding shed to allow him to ‘mark’ his territory. It is important for him to become comfortable in the breeding environment. He should be led about the area, backed into the wash area, and allowed to smell the phantom, if one will be used. The next phase of training the novice stallion is to introduce him to an estrous mare in the breeding area. It is imperative that the mare be well behaved, in good estrus, and gentle. Ovariectomized mares treated with estrogens can be used for tease and mount mares. Maintaining such mares assures readily available and predictable mares for training novice stallions and for teasing in an artificial breeding program. It is very important for the stallion handler to be experienced in handling a novice stallion as it is at this point that the stallion’s breeding habits will be developed. Consequently, the handler must know when, and when not, to discipline the stallion. The first exposure should occur with a mare behind a teasing chute, in a stall or other solid partition that will protect the mare and stallion from injury (Figure 139.5). Vocalization, posturing, nipping the neck, body, and legs, and shoving with the head are natural means of ‘teasing’ to determine the receptiveness of the mare. The stallion should be allowed to conduct his teasing and testing of the mare as long as he does not become excessively aggressive or abusive or place himself or others in a dangerous position. If test results (CEM and EVA) are needed before breeding is permitted, the training session must terminate prior to actually breeding the mare. If a test mare is being used, or if the stallion is to be trained for semen collection with an artificial vagina (AV), the training session can continue. Even if the stallion is to be used for live cover, it is useful to train him to ejaculate into an AV. This should be done early in his
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training as stallions that become accustomed to natural cover may be difficult to train to an AV. If the stallion is to mount the mare, she should be properly restrained to prevent injury to the handler and stallion. When first training the novice stallion, a safe mare is very important. A twitch is usually applied and a leg strap may be necessary. Many breeding sheds use breeding boots that are put on the mare’s hind feet to reduce injury from kicking. A wrap is applied to her tail to keep his penis from becoming entrapped or cut by tail hair. After a period of teasing the mare, the stallion is allowed to approach her from her left side at a 45◦ angle. He should be permitted to smell and nip at her flank and perineum. Most young stallions will usually mount the mare after a few minutes of teasing. If one anticipates that the stallion will do so cautiously and gracefully, he should be permitted to mount. If, however, he charges or lunges at the mare, he should be directed to back away from her until he shows control. If he is under control, the novice stallion should never be reprimanded for mounting prematurely the first few times as it could significantly hinder his training program. Breeding form should not be an issue during the first few sessions as it should improve with experience. It is common for a novice stallion to mount without an erection or to lose his erection the first few times. This will be less of an issue as he develops confidence. If he becomes unruly he should be returned to his stall and re-schooled in the basic commands of backing and standing quietly before he is returned to the breeding shed. Ideally, the stallion should learn to tease the mare but to wait for a signal of approval from the handler before mounting her. Injury to the stallion can oftentimes be prevented by teaching him to mount only on command. Non-aggressive rearing in the presence of the mare should not be disciplined initially as it will often precede his mounting. When he develops confidence and skill in mounting, uncontrolled rearing should not be tolerated. Other bad habits, such as savaging the mare and kicking out, must not be tolerated as uncorrected bad habits will become quickly established. When the stallion successfully mounts the mare, his penis should be guided to the vulva to assist with intromission. When ejaculation is complete it is not uncommon for the novice stallion to ‘faint’ or fall off the mare. Care must be taken at the completion of ejaculation to insure a safe dismount. It may be necessary to pat or shake his head following the first ejaculation to prevent him from falling off the mare or phantom. After ejaculation is complete, he should be backed down slowly and backed into the wash area, where his penis is rinsed with clean water. While good manners are a must, during the first few
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Stallion: Sexual Behavior
sessions the novice stallion must be given some latitude on technique. When he accomplishes covering the mare, he will have more confidence during successive sessions, when technique can be perfected. If the stallion is being trained to mount a phantom, it may be necessary to let him ‘live’ cover a mare the first few training sessions. If, however, the stallion has good libido he may mount the phantom with a minimal amount of encouragement during the first training session. To encourage him to mount the phantom, the estrous mare should be positioned on the right side, parallel to the phantom. The stallion is then allowed to contact her over the phantom from the left side either perpendicular to the phantom or on a 45◦ angle to the rear of the phantom. When the stallion rears and tries to mount the mare, he is gently pulled over onto the phantom. If a chain shank is used to restrain the stallion, it is helpful to have a cotton shank attached to the halter along with the leather chain shank that can be used to pull the stallion so as to not cause discomfort with the chain shank. The novice stallion can be allowed to mount the phantom from either side on the first few attempts. After he has experienced an ejaculation he will mount more aggressively and can then be trained to mount properly from the rear or left side of the phantom. Although an erection is desirable at this time, if he tries to mount without an erection he should be allowed to do so. In this situation, he may thrust and lay on the mare or phantom for an extended time before dismounting. After several mounts, he will usually develop an erection. If the stallion is not inclined to rear to mount the mare, it is often helpful to lead him up to the rear of the phantom and cause his chest to bump the rear of the phantom (Figure 139.6). This bumping often stimulates him to mount. After teasing the mare to the point of developing an erection, he should be taught to back until he can be led forward to mount the phantom from the rear. During the learning process, it is very important that the young stallion has as many positive experiences as possible. If after a reasonable session (15–30 minutes) the stallion has not mounted the phantom, allowing him to mount the mare and to ejaculate will increase his confidence. If he is allowed to mount the mare, an effort should be made to have him ejaculate into the AV. When the stallion has successfully mounted the phantom or mare, his erect penis is directed into the AV. It is important to have an experienced semen collector involved in this procedure to insure a successful ejaculation. If the stallion successfully ejaculates in the AV, he will usually mount the phantom on future attempts without the mare being placed beside the phantom. When this stage is reached, the mare may be placed in a teasing chute in close proximity to the phantom.
If the stallion fails to produce an ejaculation following several breeding attempts, the temperature in the AV should be adjusted. If he still is unable to ejaculate, it may be necessary to let him enter the mare for ejaculation before he becomes too frustrated. The next session can be devoted to getting him to mount the phantom and ejaculating into the AV. Many novice stallions will mount the phantom during the first session and most stallions will ejaculate on the phantom by the second or third session.3 As he develops more confidence in breeding, the novice stallion may become more difficult to manage. At this point, the basic techniques of handling become even more important. The stallion may become so excited that he begins to try to mount before he has an erection or charges the phantom. He must not be allowed to mount until he is ready with a full erection. The two most common reasons for failure to ejaculate into an AV are (i) improper temperature (usually too cold) or (ii) incorrect pressure (usually too tight). If he has a complete erection but fails to ejaculate into the AV the temperature can be increased to 118–125◦ F (48–52◦ C) and the pressure can be increased or decreased to identify the stallion’s preference. If this adjustment fails to work, the addition of pulsating pressure to the glans penis through the AV with the left hand may be helpful. Some stallions refuse to ejaculate when a disposable AV liner is used or if it becomes wrinkled after several breeding attempts. It may be necessary to remove the disposable liner until the stallion has been collected successfully several times. If the stallion is timid and fails to attempt to mount on the first session it may be necessary to devote more time to exposing him to mares in estrus. Patience is essential at this stage of training. One cannot
Figure 139.6 Stallion bumping the rear of a phantom to encourage him to mount.
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The Novice Breeding Stallion make a stallion breed any more than he can be made to drink water. In the event that the stallion fails to show interest in the mare or fails to achieve an erection during the first training session, it may be helpful to allow him to watch another stallion perform in the breeding shed. In the absence of good libido, the stallion should be exposed to an estrous mare for 30 minutes daily in anticipation that he will develop interest in her. The use of diazepam (e.g., 10–20 mg by slow intravenous injection for a 500- to 550-kg stallion) 5–7 minutes prior to exposure to the mare has been helpful in some cases in which the stallion showed no interest in the mare. This pharmacological treatment is primarily indicated for those stallions that have had previous negative experiences with mares or handlers.4 Stallions that lack any interest in mares should be examined for a hormonal, physical, or neurological condition that may be affecting their libido. Occasionally, a stallion will be encountered that develops a firm erection but refuses to mount a mare or a phantom. If this phenomenon is due to fear or physical limitations, medications such as diazepam or non-steroidal analgesics may be helpful. Such stallions will frequently ejaculate into an AV while standing in the presence of a tease mare. There are in fact reports of breeders who train their stallions to ejaculate in this manner to avoid the need for a phantom.5 Stallions that will readily mount a mare but refuse to mount a phantom can be trained to mount the phantom by using full blinkers. The stallion is first allowed to mount the mare and semen is collected with an AV. When he becomes adept at this method he is blindfolded with the blinkers and encouraged to mount the mare and ejaculate into the AV. After he has learned to mount the mare blindfolded and ejaculates into the AV, the mare is replaced with a
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phantom and he is encouraged to mount and ejaculate while blindfolded. When he accepts the phantom as a mount the blinkers are removed.6 As the stallion learns to gracefully mount the mare or phantom and to ejaculate, he should be taught to quietly back into the wash area to have his penis rinsed before leaving the breeding shed. While teaching and enforcing these principles of behavior, positive reinforcement with praise for good behavior is helpful. Discipline should never be excessive and should be accomplished with the least amount of confusion and noise. Whips, twitches, brooms, and other objects used to punish should be avoided as they create fear rather than respect. When proper restraint is applied through a chain shank or bit, the intelligent stallion quickly learns that misbehavior will not be tolerated. With patience, perseverance, and good horsemanship a well-mannered stallion will be developed that will become a pleasure to manage.
References 1. Conboy, HS. Preventing contagious equine diseases. In Proceedings of the 51St Annual Convention of Am. Assoc. Equine Pract. 2005;439–445. 2. Varner DD. Handling the breeding stallion. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners, 2005, pp. 498–505. 3. Conboy HS. Training the novice stallion for artificial breeding. In: Blanchard TL, Varner DD (eds) Veterinary Clinics of North America: Equine Practice. Philadelphia: W.B. Saunders, 1992; pp. 101–9. 4. McDonnell SM, Garcia MC, Kenny RM. Pharmacological manipulation of sexual behavior in stallions. J Reprod Fertil Suppl 1987;35:45. 5. Crump J. Stallion ejaculation induced by manual stimulation of the penis. Theriogenology 1989;31:341. 6. Richardson G, Wenkoff M. Semen collection from a stallion using a dummy mount. Can Vet 1976;17:177.
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Chapter 140
Training a Stallion to the Phantom Glenn P. Blodgett
The proper training of a stallion to mount a phantom and subsequently ejaculate into an artificial vagina is an integral part of any artificial insemination program. Each year, young stallions go into artificial breeding programs, and older stallions previously exposed only to a natural live cover situation need to be trained to mount a phantom. This process requires a great deal of patience, and safety considerations for all participants should be maintained at all times. Improperly trained stallions can result in numerous episodes of handlers being dragged to the mare or phantom on the end of a stallion shank, a most embarrassing and frustrating experience.1 Also, the preparation of the stallion and selection of the proper facility and environment, tease mare, and the introduction of the stallion to the whole process is extremely important.
Facility or environment Suggested minimum dimensions for an enclosed semen collection area are: room size 9 × 9 m, entry door size 1.5 × 2.4 m, and minimum ceiling height 3.7 m. One should also be able to close gates/doors to secure an untrained stallion in case problems arise. The room or area utilized for semen collection should have good footing that does not become slippery when wet. The phantom should be placed where there is adequate room to safely maneuver completely around it. This room or area should also contain a designated area for a tease mare, which should be located toward the front of the phantom. The best surfaces include various rubber products, wood byproducts, or ground limestone. Concrete or asphalt should be avoided. If the facility will need to be inspected and certified by a government regulatory agency to qualify semen collection for
export shipment, the area in which the semen is collected should be easy to clean and disinfect. This requirement limits the surface to a rubberized or non-slippery synthetic surface over a concrete base. The area should be free of all obstacles such as buckets, shovels, manure forks, etc. Also, water faucets should be recessed into the wall or placed in a corner in such a manner to avoid potential injury to anyone. The phantom utilized should be well anchored and of sturdy construction, fully padded, and covered with a cleanable cover. The cover of preference is harness leather. It provides a durable, cleanable surface that provides less deleterious friction to the inside of the stallion’s forelimbs during his time on the phantom. For better hygiene when multiple stallions are collected, wrapping the mounting end of the phantom with high-performance stretch film plastic that is changed between stallions is preferable. The ideal phantom should have a single pedestal and have some means of adjusting the height (Figure 140.1). ‘Artificial hips’ built into the phantom also have an added benefit by providing a more ‘life-like’ experience for the stallion, enabling him to more easily clasp the phantom with the forelimbs during the act of breeding (Figure 140.2). A slight angle of 10–15◦ , with the lower end being where the stallion mounts, is desirable when constructing a phantom. A good length is 2.2 m. Phantoms that contain an opening to house the artificial vagina should be avoided if possible because these can lead to injury of the stallion’s penis. The phantom should be free of any unnecessary attachments and should make minimal or no unusual noises as the stallion is mounting or performing while mounted on the phantom. The area designed to restrain the tease mare in the semen collection area should be padded on her side as well as the stallion’s side for safety. It is the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Preparation and introduction of the stallion
Figure 140.1 A breeding phantom (dummy) with a single pedestal and attached hydraulic jack for adjusting the height.
author’s preference that there are no gates on either end of where the mare is positioned. Openings on both ends provide an environment for the tease mare that is less confining and safer if she bolts as the stallion approaches her or while he is teasing on her (Figures 140.3 and 140.4).
Figure 140.2 A mounting-end view of a breeding phantom that has built-in artificial hips and is wrapped with highperformance stretch film.
In preparation for the phantom-training phase, the stallion handler should become as familiar as possible with the stallion to be trained, including his behavior and response on the end of a lead shank. It may be necessary to spend some additional time with the stallion to make sure he responds appropriately to the handler and to determine what type of equipment may be necessary for controlling him when the training period begins. Timid stallions may require only a halter with a cotton lead, but more aggressive stallions may require a halter with a 76-cm chain and snap attached to a leather lead 3 m long. For more control, the chain may be placed in different positions. Some stallions are best controlled with the chain placed over the nose and others with the chain through the mouth or under the chin. A leather halter constructed with round brass rings at the junctions of the straps is best.2 It is very important to keep in mind that the goal is to achieve a safe, enjoyable, and satisfying experience for all participants. Prior to bringing the stallion to the semen collection area, a tease mare should be carefully selected and situated behind the teasing partition. The ideal tease mare should be gentle, easy to handle, medium sized, and exhibiting good signs of estrus. She should also be examined by transrectal palpation and/or ultrasound prior to the training session to ensure that her clinical behavioral signs of estrus match her reproductive tract examination. It is necessary to make this careful selection to avoid sudden objectionable outbursts by the mare to obtain the best overall experience for the novice breeding stallion. In some cases, ovariectomized mares are utilized as tease mares and estradiol can be administered to intensify estrous behavior in these mares. These ‘programmed’ tease mares are sometimes not the ideal candidates for use in training a new stallion as some become less tolerant to approaches by inexperienced stallions, and likely do not release pheromones appropriately for maximum sexual stimulation of the stallion. The tease mare is placed in her designated area prior to the stallion’s introduction. The stallion is led toward the mare in a cautious manner to avoid frightening her and also to determine, prior to the stallion getting very close, whether she will be receptive to his presence. After he arrives at her side, the stallion handler should allow him to nuzzle, smell, or lick the mare at his leisure. Some stallions will arrive in the semen collection area and develop a full erection and be rearing up prior to reaching the mare’s side. These individuals will need to be restrained and obviously will not require much time to proceed to the
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Stallion: Sexual Behavior
Figure 140.3 A stallion teasing a mare located behind a padded teasing wall.
next step. Others will require a moderate to considerable amount of time in direct contact with the mare to obtain an erection. Once satisfied that the mare is comfortable with the stallion’s presence, he should be allowed to approach the mare’s rear end from the side
Figure 140.4 A teasing wall, padded on both sides and top, and open on both ends.
without letting the stallion or the handler directly behind the mare. Some stallions become more aroused when allowed to approach the forequarters or head and neck region of the mare. Always be aware that the stallion may suddenly strike out with a front foot during the teasing process. Once the stallion becomes sexually aroused and his penis becomes erect, the penis should be washed with warm water. In the case of timid stallions, it may be appropriate to not wash the penis until after the stallion has been trained to mount, just in case penile washing slows his progression in the training process. It is preferable to put the water in a stainlesssteel bucket containing a disposable plastic liner. Disposable gloves should be worn by the person washing. Pieces of cotton presoaked in the warm water will facilitate washing to remove smegma and other debris on the shaft of the stallion’s penis. Following the washing of the penis, a soft paper towel is used to dry the penis. At the conclusion of the drying process, some stallions respond positively to a gentle massage of the glans penis. Timid stallions often have to be washed while continuing to tease the mare; however, many stallions that become excited and sexually aroused may need to be backed into a corner to wash or in the middle of the area away from the mare. If a corner is elected for this procedure, a padded wall is recommended because the stallion will often do some kicking with his hind legs during the washing procedure. While it is more desirable to wash the penis prior to collection in this training period, it is not mandatory. Some stallions are intimidated or frightened at first and some timid stallions are particularly sensitive to the washing procedure initially. Training this type of stallion may be better served without washing of the penis prior to
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Figure 140.5 A stallion being allowed to approach the breeding phantom with an erection.
attempting mounting and subsequent collection of semen.
Approaching the phantom and collection The tease mare should be twitched and positioned on the right side or opposite side of the phantom from the stallion with her rear end 15–30 cm from the rear end of the phantom. The stallion is allowed to approach the phantom, preferably with an erection, especially if he is rearing and extremely aggressive (Figure 140.5). It is more desirable for the stallion to have an erection when he mounts the phantom. Some
less aggressive, or more timid, stallions need more teasing of the mare over the rear end of the phantom. It is preferable to keep the stallion at the rear of the phantom with his chest touching the opposite rear corner of the phantom from the mare. This stimulation on the chest is often necessary to aid in the stimulus for the stallion to mount the phantom. Some stallions become more aroused if the mare’s head is turned toward the stallion while he is teasing. If the stallion fails to mount the phantom in a reasonable length of time, do not be afraid to put him back in his stall and try later in the day or even the next day. Some stallions respond very favorably during this second attempt. If at all possible, try to avoid using a
Figure 140.6 Forearm pressure from the collector is applied to the stallion to prevent him from moving up the side of the phantom during collection.
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mare as the mount source initially. This has to be done on some occasions, followed by semen collection attempts with the phantom as the mount source. Another technique that can be used in difficult stallions is to let the stallion follow the tease mare in a circle that ends up with the mare in the original position by the side of the phantom. This may help the stallion to feel more comfortable in placing his chest on the rear of the phantom. Once the stallion mounts the phantom, introduce his penis into the artificial vagina and proceed with the semen collection process to achieve ejaculation. If the stallion fails to mount the phantom in the proper way, try to reposition him on the phantom or pull him off the phantom. If you have to pull him off, do so with the minimum effort possible to avoid damaging his interest; however, safety considerations for all participants should be carefully maintained. Once the stallion has ejaculated, let him stay in position on the phantom as long as he wishes. The mare can be led away eventually if he fails to back off the phantom within a reasonable amount of time. Once dismounted, the stallion can be led back to the stall. If the stallion successfully ejaculated while on the phantom, the next time he is introduced things are likely to go much faster and easier. Many stallions eventually no longer need a tease mare next to the phantom and a few do not even need a mare in the
collection area at all once they become familiar with the routine. Some stallions, upon mounting the phantom, try to walk up the side of the phantom prior to, or after, entering the artificial vagina with their penis. The more aggressive stallions that walk up the side may require the collector to place forearm pressure above the stallion’s stifle in the mid-thigh region (Figure 140.6).
Summary Every breeding operation strives for maximum reproductive efficiency, and the performance of a safe, fast, smooth semen collection process is a vital component of artificial insemination programs. Every effort should be made to insure an enjoyable experience for the stallion each time he enters and leaves the collection area. The goal is to achieve a ‘one-jump collection’ in a timely, efficient, and safe manner.
References 1. Conboy HS. Training the novice stallion for artificial breeding. Vet Clin North Am Equine Pract 1992; 8: 101–9. 2. Varner DD. Handling the breeding stallion. AAEP Proc 2005; 51: 498–505.
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Chapter 141
Abnormal Sexual Behavior Sue M. McDonnell
There are several distinct types of abnormal and problem sexual behavior in domestically bred stallions. The most common are: (i) inadequate sexual interest and arousal or stamina including apparent preferences and aversions (commonly called ‘libido problems’); (ii) highly enthusiastic sexual behavior that challenges the skill of domestic handling and facilities; (iii) aggressive behavior; (iv) frenzied, frenetic behavior; and (v) self-mutilation. Specific erection and ejaculation dysfunctions are covered in Chapter 144.
Inadequate sexual interest and arousal (libido) Specific stallion libido problems include slowstarting novices, slow or sour experienced stallions, and specific aversions or preferences. Slow-starting novices Most stallions over the age of 2 have matured through puberty and are interested in, and will respond adequately to, a mare in estrus under natural conditions; however, it is not unusual for their behavior to appear immature in terms of copulatory organization and agility. Under domestic in-hand breeding conditions, there appears to be a greater percentage of stallions that experience difficulty as novice breeders. Much of this is likely attributable to suboptimal stimuli and handling conditions inherent to domestic breedings. Some stallions appear to be slow to mature behaviorally, possibly due to limited social interaction as a juvenile and pubertal colt. Signs of behavioral immaturity include playing with the tail of the mare as a foal does, as if ‘flossing his teeth’ with tail hairs, absence of vocalization, and assuming the nursing posture with a sucking motion known as champing.1 Many slow-starting novice breeding stallions show clear signs of experience-related suppression of
sexual behavior and often have a corresponding history, logically supporting the behavioral signs. These would include stallions that have been disciplined for showing sexual interest in mares during their performance careers, or discouraged from showing spontaneous erection and masturbation, or mishandled during breeding under halter. In horses, sexual behavior is particularly vulnerable to suppression as a result of associated negative experiences.2 Stallions that have been trained to subdue sexual response to mares during training often experience a difficult transition to a breeding career. When exposed to a mare for teasing, such stallions may simply stand quietly or may intermittently show bursts of interest and arousal or aggression. They may achieve a partial erection, and then withdraw again. They may appear anxious, confused, and/or easily distracted. Their behavior often suggests a state of what is known as approachavoidance conflict, or confusion as to whether to ‘behave’ or progress with sexual arousal. In this state, the animal may show sudden seemingly inappropriate aggression; the stallion may pin the ears and lunge to grasp onto the mane, or wheel around and kick out at the mare, or displace aggression onto the handler with sudden unexpected bites or strikes. The stallion may appear fearful. Fear and anxiety in horses are often unrecognized by handlers, and can easily be made worse with inappropriate response of the handler. Signs of fear include facial expression often described as fearful, anxious, or nervous (see Figure 141.1), defecation (which is not normal in a breeding situation), hyperalert gazing off into the distance, and hyper-reactivity to environmental sounds or visual cues. Other signs include inappropraite rubbing or self-grooming behavior, or yawning. In extreme anxiety, a horse may weave, even when being led in-halter. The stallion may regress into periods of licking the mare, or nipping incessantly, rather than progressing forward through the precopulatory and copulatory sequence. In what appears to be extreme
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 141.1 Facial expression of fear, anxiety, or nervousness.
conflict, a stallion may bite or nip at himself, as if in frustration, usually at the chest or shoulder (see Figure 141.2). These ‘out of context’ behaviors are known in animal behavior as displacement activities, and reflect a stressful underlying motivational conflict. Slow novice breeding stallions typically respond best to continued exposure to mares, in as natural a setting as possible. If possible, it is typically helpful to initially remove the human handling, allowing the stallion direct contact or fenceline access or housing within close proximity to mares. This will likely enable the stallion to gain confidence with sexual interaction and response with mares. Exposure to mares also typically leads to an increase in testicular hormone production to naturally drive sexual interest, response, and confidence.3 Once the stallion has bred a few times at pasture, or has gained confidence with fenceline access, human handling can be gradually reintroduced. Most stallions do best with a quiet, respectful, patient, non-confrontational style of handling. Care should be taken to minimize exposure to any other naturally threatening or learned threatening stimuli, such as exposure to a restrained non-receptive mare that may be explosively resistant to the advances of a stallion. Slow-starting stallions appear to respond favorably to reassurance for even
Figure 141.2 A stallion self-biting side-to-side at his chest and forelimbs in a state of apparent approach-avoidance conflict whenever led toward a breeding situation if near another stallion.
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Abnormal Sexual Behavior small increments of improvement. Tolerance of minor misbehavior, rather than punishment, is often a rewarding strategy. Progress may be more efficient if routines such as washing the penis or obtaining post-ejaculation semen samples are omitted initially and then introduced as the stallion gains trust of the handlers and enthusiasm for in-hand breeding. For a novice or timid breeder, the ideal stimulus mare is an intact mare in good natural estrus that requires almost no restraint or protective breeding gear that is presented without perineal washing and tail wrapping. In addition, handling of the mare in a manner that allows or even encourages appropriately timed natural movement and postures in an open area is typically most effective for novice and otherwise tentative breeders. The mare should be free to move in the short choppy steps typical of estrus and to swing the hindquarters toward the stallion as he approaches, rather than being forced to stand immobile. The most inviting mating posture of mares, which is common to quadruped females, includes extreme flexing of the neck, turning the head back toward the stallion with the nearside foreleg flexed. This is known in animal behavior as the ‘You are safe, I’m not going anywhere’ posture. For stallions that appear tentative to proceed with erection or mount, this posture typically evokes immediate progress in the form of the pre-mounting nicker and then mounting by the stallion. The preparation of the stimulus and mount mare for in-hand breeding is especially important to efficient progress of slow-starting or otherwise tentative stallions. Procedures that are used routinely in breeding sheds with experienced, confident breeding stallions should be carefully considered before use with a novice breeder. For example, the gear meant to protect the mare or the stallion can induce non-natural appearance and movements of the mare that can hinder progress. A lip or ear twitch, for example, alters the normal posture and movement. A twitch typically induces, especially when first applied, a nonnatural head-high, ears back, clamped tail, tense posture more similar to aggressive postures of the horse than to receptive or neutral postures. A twitch, while in some cases reducing the likelihood of kicking, typically inhibits the movements of the mare described earlier that are most stimulating to the stallion. Mares wearing breeeding boots meant to protect the stallion typically move with goose-like steps that are unnatural. Even experienced and confident breeding stallions that are unaccustomed to this behavior are typically distracted and sometimes fearful when first presented to mares wearing boots. Mares unfamiliar with the boots are often uncomfortable as well, and so appear to convey that tension to the stallion. Breeding capes and tail wraps can alter the appearance and mask naturally stimulating odors.
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Similarly, care to preserve the natural odors of the mare without introducing potentially negative odors, such as fly sprays or detergents, or disinfectant odors or the scents of other stallions can also be useful. For natural cover, residual odors of the teaser or test mount stallion on the mare can appear to be offputting to novice stallions until they gain confidence with the domestic protocol. Stallions appear to have a natural tendency to be reluctant to mount a mare that is not free to move forward as needed to accommodate thrusting. Some appear, especially as novices, to be hesitant to mount a mare that is positioned near a barrier such that she cannot move forward. Simply moving the mare into an open area can appear to release the stallion’s apparent inhibition. The breeding environment can greatly affect the confidence of a novice breeder. Stallions vary somewhat in what features of the environment appear to enhance or inhibit progress. In general, a quiet spacious area, either outdoors or with high ceilings, natural lighting, without shadows or window-like streaks or glass reflections, is best. Slow-starting stallions may appear inhibited or conflicted when in the presence of other stallions, which are naturally intimidating to young, inexperienced or bachelor stallions. The sight or sound of other stallions, even at a distance, can be distracting, as can be the residual scents of stallions in the breeding area. The horse may also be less inhibited in showing sexual interest, for example, if individuals who may have been involved in discouraging sexual behavior in show or performance training are not present. Similarly, the horse may be more sexually comfortable if the training for breeding is done in a new fresh enviroment away from the training environment. Horses, as an open plains grazing species, have evolved to respond to the behavior and physiological states of other species in their environment. Males appear particularly dependent upon alarm as well as safety cues from herdmates as well as species around them whenever placing themselves at risk during copulation. With the same abilities, horses can similarly appear especially responsive to the emotional states of humans, particularly in the breeding environment. In a sexual situation, horses appear to recognize and react accordingly to fear, anger, and impatience or agitation of humans. People who are especially effective at starting or rehabilitaing breeding stallions with behavior problems recognize this attribute of horses and strive for a calm and positive interaction among people and animals so as to avoid inhibiting the horses’ confidence. Some low libido problems appear to be associated with low gonadal hormone production, similar to that of young bachelor stallions under natural social conditions. These can improve with management
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aimed at increasing exposure to mares and reduced exposure to other, especially more dominant, stallions. As mentioned earlier, this will typically lead to increased androgen levels, general confidence, sexual interest and arousal. Slow experienced breeders Experienced breeders may suddenly or gradually develop slow breeding behavior, or even what appears to be a total disinterest in breeding. The problem may be intermittent. This is often in association with a heavy breeding schedule, physical discomfort, or with injudicious handling or a breeding accident. Specific preferences and aversions Stallions, whether novice or experienced, slow or enthusiastic, can exhibit differences in their response that appear to represent specific preferences or aversions to any number of aspects of the domestic breeding situation. Examples of preferences or aversions include characteristics of the mare such as size, breed, color, lactation status, proximity to ovulation; time of day; particular handlers; or particular breeding locations such as indoors or outdoors. Many of the preferences or aversions have logical natural explanations. For example, most stallions respond more enthusiastically with a mare in natural estrus than with an ovariectomized stimulus mare or a dummy mount. Alternatively, a horse’s history may suggest a learned or experience-related basis for the preference or aversion. For example, in the case of aversions, if a horse experiences slipping and falling when breeding in a particular location with poor footing, the horse may appear uninterested in breeding mares in that location or on that type of surface. The ability to adjust sexual interest and response as a result of experience in this regard has evolved as it is likely quite adaptive. If necessary, in many cases the horse can be retrained to overcome the aversion or preference, but in others, it may be simply more efficient to abide the apparent preference or aversion. Preferences and aversions are often more or less pronounced in association with overall libido. So with seasonal fluctuation in gonadal steroids, work, fatigue related to a heavy breeding schedule, or physical discomfort, the aversions are typiclly more conspicuous and problematic.
Problems with highly enthusiastic or aggressive breeding behavior Rowdy, ‘misbehaved’ or dangerously miscontrolled breeding stallions in most cases represent a human–animal interaction problem, that is in many cases aggravated by suboptimal domestic housing,
feeding, and management conditions that predispose a horse to high energy and short-tempered behavior. Most problems can,in large part, be overcome with judicious, skillful, respectful training for breeding that builds trust and patience with the handling, along with management aimed at identifying and relieving the stallion of the aggravating management factors. As with the training of pets, the key to successful behavior modification of unruly sexual behavior of a stallion is a change in the behavior of the handler to respectfully and positively redirect spontaneous undesirable behavior and to avoid inadvertent prompting of undesirable behavior. As such, the handler and breeding shed team typically may need more actual training time to acquire the necessary skills for positive interaction with the stallion. Successful modification of unruly behavior of a breeding stallion holds the special challenge of eliminating undesirable behaviors without suppressing normal sexual behavior. A misunderstanding of some stallion handlers is that vocalization, prancing gait, and normal sexual enthusiasm should be discouraged. Simple handler education, for example, that vocalization is not particularly associated with dangerous or undesirable behavior, can be productive. Some handlers benefit from systematic desensitization to the shrill sexual call of stallions. This can be done using audio/videotaped breeding or teasing of a vocal stallion that is being handled by an expert stallion handler. Even the most strong, vigorous, and ‘misbehaved’ breeding stallion can be retrained using simple positive and negative reinforcement, with very little or no punishment. It can be done in a safe and systematic manner without abuse or commotion, usually within a few brief sessions. Some of the most anxious and disorganized stallions may benefit from training that includes several breedings in rapid succession. With reduced urgency to breed, it is often easier to maintain the stallion’s attention for teaching the organized protocol and to give the horse the opportunity to learn that, by following the direction of the handler, the stallion will reach the goal of breeding. Our clinical approach to systematic behavior modification of unruly stallions and example case reports from our clinic have been detailed elsewhere.4 Examples of specific habits that commonly develop in breeding stallions that are considered undesirable, mostly for animal and human safety reasons, include biting the mare or handler, striking, rearing, charging or rushing the mare, as well as spinning around and kicking out. Handling procedures that are particularly problematic with overenthusiastic breeders and are to be avoided if possible are: circling the stallion around the handler, rather than
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Figure 141.3 Stallion rearing, inadvertently provoked by the handler applying sharp pressure on the mouth.
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kicks, and otherwise fight to the death as two stallions would fight or as a stallion would fight a small predator species. A less savage, yet dangerous, type of aggression of stallions toward people involves interacting with people as if they were another horse. So while the horse is not savagely attacking, it responds to inadvertent cues from people with the usual behavior that would be appropriate and safe in response to another horse, but is dangerous with people. An example is the horse that responds to touch of the forelimb or to the groin with the reflexive biting back toward the person, flexing the limb and dropping to the knee, or rearing and striking toward the person, as would be the normal interactive sequence among two stallions. While many young horses may react in this manner to initial handling by humans, most readily acclimate as if they distinguish human from conspecific interaction. Some of these animals that interact with people as they would with other horses have a history of bottle feeding as an orphan foal or otherwise intense human handling as neonates. It has been proposed that the interaction with humans as if they were horses is associated with becoming overly bonded to people as juveniles that is a fairly well-known phenomenon among hand-raised males in certain other species. For example, this phenomenon occurs commonly in male llamas, and is know as crazy male llama syndrome.
Frenzied or frenetic behavior backing away from the mare during teasing; jerking at head restraint or the mouth of the stallion, which may provoke rearing or unnecessarily confuse or discourage the stallion (see Figure 141.3); poking or punching at the mouth of the stallion, which tends to escalate inter-male fighting of stallions; and applying tension on the head restraint once the horse is mounted, which often distracts the horse from necessary focus to thrust and ejaculate.
Aggressive behvaior In contrast to highly enthusiastic urgent breeders, or inadequately directed, or dangerously ‘out of control’ stallions, a more concerning behavior problem is aggression directed toward people or savage aggression toward other animals. Truly savage attack of humans or other animals by stallions is rare, but does occur. The horse’s behavior seems aimed at killing the target animal. The attack is usually sustained, with the horse using its mouth and limbs to take the animal or person to the ground, pick it up and thrash or toss it, stomp it, or corner it against a barrier with hind leg
Distinct from simple rowdiness, some stallions, like some mares or geldings, are hyperactive or even ‘frenzied’ when kept in stalls or paddocks on breeding farms, especially when stimulated by, yet physically separated from, nearby mares or other stallions. They may pace and weave frenetically in the stall, and literally climb the walls in an apparent effort to gain social contact with mares or stallions in their environment. Reproductive hormones may or may not be a factor. Important factors are genetics, experience, sociosexual environment, housing, diet, and exercise. Many of these stallions become more relaxed when pastured with a mare, where they appear to settle into a focus on meaningful, normal harem maintenance activities with that mare. If it is not possible to arrange this scenerio of management, such stallions can often benefit from organized daily work or exercise, and consistent housing in a quiet area where they are not stimulated by horse activities. More roughage and less grain in the diet along with Ltryptophan supplementation (1–2 grams twice daily in feed) can have a measurable calming effect on such stallions.
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Self-mutilation A fairly rare, but highly problematic, condition of domestic stallions is self-directed injurious behavior, known as self-mutilation. There are at least three distinct categories of such self-injurious behavior, which have recently been classified and outlined.5 In summary, the most common cause of self-directed injurious behavior is likely physical discomfort that, for any number of reasons, may not be obvious. Another type appears to be self-directed inter-male aggressive interaction. The behavioral sequence includes those vocalizations, elimination marking behaviors, nips, bites, srikes and kicks, typical of the ritualized interaction that is normal among stallions, but in the absence of a cohort the stallion is his own target.1, 6 A third type of self-mutilation appears to be the stereotypic or ‘habit’ form of either of the two other types. An understanding of the differences and an appreci-
ation of the behavioral signs of each type are helpful to efficient diagnosis and treatment or management.
References 1. McDonnell SM. The Equid Ethogram: a practical field guide to horse behavior. Lexington, KY: Eclipse Publications, 2003. 2. McDonnell SM, Kenney RM, Meckley PE, Garcia MC. Conditioned suppression of sexual behavior in stallions and reversal with diazepam. Physiol Behav 1985;34(6):951–6. 3. McDonnell SM. Reproductive behavior of stallions and mares: comparison of free-running and domestic in-hand breeding. Anim Reprod Sci 2000;60–61:211–19. 4. McDonnell SM, Diehl NK, Oristaglio Turner RM. Modification of unruly breeding behavior in stallions. Compendium 1994;17(3):411–17. 5. McDonnell SM. Practical review of self-mutilation in horses. Anim Reprod Sci 2008;107(3–4:219–28. 6. McDonnell SM, Haviland JCS. Agonistic ethogram of the equid bachelor band. Appl Anim Behav Sci 1995;43:147–88.
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Chapter 142
Pharmacological Manipulation of Ejaculation Sue M. McDonnell
Ejaculation has two overlapping phases: (i) release of sperm from the ampullae and seminal fluids from the accessory sex glands, which is known as emission; and (ii) smooth muscle contractions of the pelvic and penile musculature (ischiocavernosus, urethralis, and bulbospongiosus muscles) resulting in forceful expulsion of semen, known as ejaculation proper. The onset of emission normally precedes and continues during ejaculation proper. During emission and ejaculation, the bladder neck normally contracts tightly to prevent movement of semen into the bladder or discharge of urine into the urethra. Specific ejaculatory problems in stallions include delayed ejaculation, ejaculation failure, inconsistent or incomplete ejaculation, retrograde ejaculation of semen into the bladder, urine discharge during ejaculation resulting in urospermia, and oligospermia or azoospermia due to emission failure or tubular blockages. In clinical practice, the most commonly presented ejaculation problems for horses breeding by natural service involve delayed ejaculation or anejaculation. Urospermia, oligospermia, or azoospermia are often recognized only when collecting semen in an artificial vagina for insemination purposes, or as a component of a breeding soundness examination. Ejaculation in copula is dependent upon erection, insertion, and thrusting. Therefore any deficits in sexual interest and arousal or breeding fitness may affect the ability to ejaculate. Even when arousal is within the range of normal, maximizing comfort and confidence, as well as increasing the urgency to breed can often improve ejaculatory function. Methods to enhance arousal, including simple management and handling changes, and optimal stimulus from mare and mount, have been addressed in Chapters 117, 118, 123, 137, 138, 139, and 140.
Ejaculation is principally an alpha-adrenergic mediated reflex event. A variety of agents with adrenergic properties are known to affect ejaculation. Details of the protocols that have been developed for inducing and enhancing ejaculation should be carefully reviewed before recommending such therapies.1 Most of the compounds employed have both inhibitory and facilitatory effects on both erection and ejaculation, depending upon the dose. Agents that enhance ejaculation may at certain doses inhibit erection processes (both tumescence and detumescence), and vice versa. As with other neuroactive agents, considerable variation in response is observed among individual stallions and within individuals, and is dependent upon the current state of arousal.
Pharmacologically induced ex copula ejaculation A number of specific protocols have been tested experimentally and have been used clinically for inducing ejaculation without sexual exposure or effort (reviewed in McDonnell, 2001).1 These include principally xylazine, detomidine, imipramine, and prostaglandins. Butorphanol has also been included in some protocols2 (Brinsko, personal communication). The protocols tested so far are unreliable, with typically no more than 50% of attempts resulting in an ejaculate, with the success rate often much lower. Careful titration of dosing for individual stallions has, in certain cases, resulted in higher rates of success. Semen characteristics and side-effects vary with the agents used. Ejaculates obtained with the most common xylazine hydrochloride protocol (0.66 mg/kg i.v.) typically have volume, gel, sperm
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Pharmacological Manipulation of Reproduction
concentration, and total number of sperm similar to that obtained in copula. Ejaculation typically occurs either within the first 3 to 5 minutes as the stallion is becoming sedate, or as the stallion is rousing from the sedation at about 12 to 15 minutes after administration. The addition of imipramine to a xylazine protocol (2.2 mg/kg orally 2 hours before xylazine i.v. 0.3 mg/kg) typically yields an ejaculate of lower volume, higher sperm concentration, and higher number of sperm compared to in copula ejaculates. With the prostaglandin protocol that has been tested (0.01–0.15 mg/kg i.m.), ejaculates typically occur between 10 and 60 minutes after administration, and are typically of greater volume, lower sperm concentration, greater volume of gel, and similar or higher number of sperm than in copula ejaculates. Sweating and abdominal cramping occurs. Enhancing strength of ejaculatory contractions A common ejaculation problem of stallions involves sperm accumulation within the ampullae, resulting in partial or complete blockages and sperm stasis.3 Diagnosis of this common condition has been detailed in Chapter 107. In such cases, oxytocin (10–20 IU intravenously) administered immediately before mounting, for each of a series of frequent ejaculations with high arousal, is oftentimes recommended as a first line of treatment for clearing ampullae. Lowering the ejaculatory threshold in copula For a variety of physical and behavioral problems, stallions may suffer reduced interest and arousal secondary to delayed ejaculation or intermittent ejaculatory failure, whether the cause is specific to the ejaculatory mechanism or secondary to musculoskeletal problems or arousal deficits. It is often judged helpful to pharmacologically lower the ejaculatory threshold for such stallions, to enable more consistent ejacula-
tion with less arousal and/or effort. Imipramine hydrochloride remains a well-tested choice for reducing the amount of effort and arousal required for the stallion to reach ejaculation for breeding or for semen collection in copula. We recommend 2.2 mg/kg orally 2 hours before breeding. In most instances, treatment with imipramine results in a low-volume, high-concentration ejaculate, with markedly reduced seminal plasma and little or gel. Xylazine, at a very low dose that does not impair musculoskeletal coordination and mental focus, can in some cases reduce the amount of effort required for ejaculation.
Reducing seminal plasma and gel in the ejaculate Imipramine treatment at 2.2 mg/kg orally 2 hours before semen collection typically reduces the contribution from the accessory sex glands to the ejaculate, and increases the contribution of sperm from the ampullae.1 This can be useful for avoiding exposure of sperm to gel, when other methods of semen fractionation are not available. It is also useful for obtaining semen with a higher concentration of sperm and total number of sperm, for example, for cryopreservation or for transporting of fresh semen in a limited volume. When using imipramine for this purpose we recommend periodic ejaculation without treatment to enable emptying of the vesicular glands so as to avoid over-accumulation of gel.
References 1. McDonnell SM. Oral imipramine and intravenous xylazine for pharmacologically-induced ex-copula ejaculations in stallions. Anim Reprod Sci 2001;68:153–9. 2. Johnson PF, DeLuca JL. Chemical ejaculation of stallions after the administration of oral imipramine followed by intravenous xylazine. Proc Am Assoc Equine Pract 1998;44:12–15. 3. Love CC, Riera FL, Oristaglio RM, Kenney RM. Sperm occluded (plugged) ampullae in the stallion. Proceedings of Society for Theriogenology, San Antonio, TX, 1992; pp. 117–23.
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Chapter 143
Pharmacological Manipulation of Stallion Behavior Sue M. McDonnell
The most efficient and effective approach to behavior modification of animal behavior, including stallions, typically and primarily involves changes in handling and management. Nonetheless, there are a number of pharmacological agents that can be helpful as an aid to manipulating stallion behavior, whether to enhance or suppress sexual and aggressive tendencies in stallions or to suppress residual stallion behavior in geldings.
Enhancing sexual interest and arousal A number of protocols have been developed in systematic studies and/or clinical experience that are useful to enhance sexual interest, arousal, and breeding stamina (libido) in stallions. These principally include analgesics, anxiolytics, and endocrine agents. Analgesics Musculoskeletal discomfort and other pain is likely the most common significant factor in inadequate sexual interest, response, and breeding efficiency of stallions. Specific musculoskeletal breeding behavior problems that commonly impact sexual behavior along with management and treatment strategies have been detailed in recent reviews (see Martin and McDonnell, 2003).1 In summary, treatment with phenylbutazone (2.2 mg/kg orally twice daily as tolerated) is widely used and appears to be quite effective for improving comfort in breeding stallions with any variety of musculoskeletal conditions. Most stallions respond better with long-term, as opposed
to short-term, treatment with phenylbutazone. Clinical experience suggests that full benefit for improved sexual behavior may not be reached until several days into treatment. Although fertility trials have not been done, clinical experience also suggests little if any adverse effects of long-term treatment with phenylbutazone on semen quality at levels tolerated gastrointestinally by the stallion. For stallions that tend to develop greater musculoskeletal problems during a busy breeding schedule, it is worth considering continuing treatment throughout the breeding season to help maintain comfort and efficient ejaculation, rather than only when discomfort is problematic. Such a strategy is aimed at avoiding the potential additive problems of delayed ejaculation, additional effort, and exacerbated lameness that often occurs in such stallions as the season progresses. Oral gabapentin has been used on a limited basis for breeding stallions where there was suggestion of a neurogenic component to discomfort, interfering with focused sexual behavior (McDonnell and Turner, unpublished clinical observations). Recent studies indicate that oral bioavailability of gabapentin is low compared to intravenous administration, but is well tolerated at 20 mg/kg.2 A COX-2 inhibitor analgesic formulation for horses (firocoxib) has been commercially available in the United States since 2006. Although not yet tested specifically in breeding stallions, in more than 250 horses with lameness associated with osteoarthritis, the proportion of patients with improved lameness scores at 2 weeks was similar for firocoxib treatment at 0.1 mg/kg orally once daily as for phenylbutazone treatment at 4.4 mg/kg orally once daily.3 For
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additional measures of pain on manipulation or palpation, joint circumference, and range of motion score, the proportion of horses improved at 2 weeks into treatment was greater for firocoxibtreated horses. Anxiolytic therapy For certain cases of slow-starting novice breeders, or for stallions with failure to respond in a novel environment or following a negative experience in a breeding context, such as rough handling, a painful event associated with breeding, or a breeding accident, intravenous diazepam (0.05 mg/kg, not to exceed 20 mg, injected slowly just before exposure to the mare) can be effective.4 The maximum effectiveness of this regimen appears to be at about 5 to 10 minutes after injection. Our clinical experience has been to expect about one-third of stallions that we judge would be good candidates for such therapy (based on history of behavioral signs of anxiety) to respond well immediately, about one-third to have no change with treatment, and about one-third to become quieter and less responsive. For horses that are quieted by this full dose, a slightly lower dose may be helpful. At much lower doses, the animal may experience a state of hyper-reactivity which is usually counterproductive to sexual arousal and response. Hormone therapy For most stallions, sexual arousal, response, and stamina can be enhanced by increasing circulating testosterone. Gonadotropin-releasing hormone (GnRH) can be used to induce a release of endogenous gonadotropins and testicular hormones. Independent of gonadal androgens, GnRH treatment has been shown to promote sexual behavior.5 A regimen of 50 µg of the simple GnRH decapeptide administered subcutaneously, 1 and 2 hours before each breeding, typically results in double the baseline concentration of testosterone at the time of breeding. Clinically, this regimen has been repeated as often as needed through a breeding season with no sign of adverse effect on fertility or significant downregulation of the endogenous hormone production. Should GnRH treatment not yield sufficient rise in testosterone or improvement in behavior, administration of testosterone in addition to GnRH may be useful. It is recommended that consideration be given to baseline endocrinology, the cost of potential downregulation and adverse effects on sperm production in light of the stallion’s breeding obligations, how dire the stallion’s breeding career situation has become, and the ability to manage a potential increase in potentially problematic aggressive behavior. Endocrinology is monitored before and during
treatment with the goal of testosterone not exceeding circulating concentrations 4 ng/mL (14 nmol/L), with fluctuations to remain within the range of 2 to 4 ng/mL. Traditionally, a variety of preparations of testosterone and the longer-stored esterified forms in aqueous and oil preparations for injection have been used for boosting circulating levels of testosterone. More recently, transdermal preparations of testosterone in oil have been applied to hairless perineal skin or directly to the penis for added benefit of improved genital sensitivity.6 For the typical stallion with baseline endogenous levels of 0.5 to 1 ng/mL testosterone, a dose for a light horse stallion of 80 to 100 mg testosterone every other day for injectable preparations, or daily or twice daily for transdermal application, will in most cases achieve serum testosterone levels that remain within the target range of 2 to 4 ng/mL. Improvement in sexual response typically lags as much as 5 to 8 days behind increased blood levels of testosterone. Aggressive behavior often increases before sexual arousal and response.
Suppressing stallion-like behavior Pharmacological strategies to assist in reducing sexual and aggressive behavior in stallions for general handling, training, work, performance situations, or in rare instances for handling for breeding, involve: (i) various approaches to disrupting endocrinology to reduce gonadal steroids; (ii) nutritional supplements to modify general temperament; and (iii) sedation. A calming pheromone that has been useful in other species is also under development for use in horses, and may have application in calming stallions that become frenzied under certain social challenges. Disrupting endocrinology
Progestagen administration In the United States, administration of progestagens (progesterone-like compounds) has been fairly commonly used for the purpose of quieting stallion-like aggressive and sexual behavior in training or performance situations where such behavior is problematic, either for intact stallions or for geldings showing residual stallion-like behavior. Oral altrenogest is the form of progestagen most widely available to the horse industry in North America in recent decades, and is likely the most commonly used and best tolerated form of progestagen for this purpose. In a 2002 survey of 200 Quarter Horse farms in which owners and trainers were asked specifically about oral altrenogest use in stallions, about one-third of 63 respondents reported using oral altrenogest in one or more stallions within that year, either for young colts or mature stallions in training, work or breeding.7
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Pharmacological Manipulation of Stallion Behavior In organized studies, with stallions of various ages, and with treatment regimens ranging from 0.044 to 0.088 mg/kg oral altrenogest daily for 8 weeks or longer, there has been consistent evidence of expected downregulation of the pituitary-gonadal axis hormones and consequent reduction in testicular size and sperm production.8–12 While reductions in sexual and aggressive behavior have been demonstrated, it is important to note that in general the changes in behavior, measured primarily in semen collection protocols, have been relatively modest. Certainly progestagen treatment, as tested in these studies, has not significantly or uniformly suppressed or eliminated sexual and aggressive behavior. Semen collection protocols in a breeding farm environment may not directly reflect the behavior changes sought or achieved for performance horses in training or work environments. So, as with many other treatments, without field efficacy studies it is difficult to judge the value of progestagen treatment. In studies to date, a treatment regimen has not been identified at which sexual and aggressive behaviors are effectively modified without inducing potentially fertility-limiting side-effects on semen. Adverse effects of progestagen treatment on spermatogenesis are expected to be temporary, at least in mature stallions, although this has not been adequately addressed.
GnRH vaccines, antagonists or agonists A second endocrine approach to modification of reproductive behavior and fertility in horses involves attempts to temporarily disrupt pituitary-gonadal hormone production by suppressing GnRH production. This is sometimes referred to as chemical castration. The specific methods of suppressing GnRH production have included administration of GnRH antagonists or agonists and, more recently, GnRH immunization. These approaches have been reviewed by Stout and Cooenbrander,13 who concluded that the most feasible current GnRH approaches for suppression of stallion behavior are immunization or use of antagonists, as stallions appear resistant to downregulation by administration of agonists. A GnRH vaccine is not yet available as approved for use in stallions. A GnRH vaccine developed in Australia for use in mares is available in some parts of the world, but not yet in North America. A GnRH vaccine developed for use in dogs, which is currently available in North America, has recently been tested for use in mares,14 but not in stallions. In that study, all six vaccinated mares stopped cycling within 2 weeks after a second vaccination at a 3-week interval using five times the canine dose. At the time of their report, data were not available on return to cycling.
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Nutritional supplements to quiet general temperament A number of dietary supplements are commercially available for calming horses. Horse-calming products are typically a formulation of two or more compounds, each with purported behavioral effects. While there are almost no published data on efficacy of particular dietary supplements for horses, some of the specific compounds included in horse-calming formulations have been used with horses for decades and judged helpful in many horses for improving general temperament and tractability. Some of the common ingredients, while not specifically tested in horses, do have basis for efficacy in theory or from research in other species. While most horse ‘calming’ supplements are not marketed specifically for male-type behavior, improved general temperament can be quite beneficial to overcoming common handling and management challenges of stallion behavior. For example, L-tryptophan, a common ingredient in commercial calming supplements, is the precursor of the behaviorally important neurotransmitter, serotonin. L-tryptophan supplementation has been found to induce calm and fatigue-like behavior in a number of species (reviewed by Grimmett and Sillence, 2005.15 We have evaluated the 24-hour time budgets of stallions with and without L-tryptophan supplementation. These protocols were used for a variety of behavior problems, including stereotypies, social hyperactivity in confined horses, as well as mild handling challenges in mares, geldings, and stallions (McDonnell, unpublished clinical case studies). We have found that supplementation with L-tryptophan at a rate of 1 to 2 grams once to twice daily typically has a mild calming effect on stalled horses, and is associated with less problem behavior and greater standing rest, as well as more normal cycles of eating and resting behavior beginning within 2 to 4 weeks after start of treatment.
Sedation Although not widely recommended, in rare instances it may be judged useful for a stallion to be lightly sedated for safe handling for breeding. Xylazine or detomidine may be the best choice among agents commonly available for use in horses. Many sedatives have variable and dose-dependent effects on erection and ejaculation, and so can compromise breeding efficiency. At sedative doses, xylazine and detomidine have been found to generally facilitate rather than inhibit ejaculation.16 When sedation is necessary for any reason for stallions, xylazine or detomidine are commonly recommended over acepromazine, due to greater risk of penile paralysis
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in horses with the phenothiazine derivative compounds. Calming pheromone In recent years, a synthetic preparation based on an appeasing pheromone identified in perimammary gland secretions of lactating females has been commercially available in Europe and North America as a behavior modification aid for a number of domesR tic species, including horses (Pherocalm in France, R Germany; Modipher EQ in USA). The target applications for the equine product have been general handling challenges that may involve fear or anxiety or stress. These would include problems loading and traveling for transportation, feet and leg manipulations, veterinary procedures, changes in work or environment, or stressful performance situations. While a pheromone approach to behavior modification is widely appealing as a more natural approach and perhaps quite promising, unfortunately efficacy of currently available products remains somewhat controversial in domestic species. For horses in particular, the efficacy claims based on sparsely available data with current formulations in horses have been formally refuted.17,18 In a clinical research setting, we have conducted over 200 controlled randomly switchback trials with mares, foals, stallions, and geldings, evaluating the effectiveness of the product as an aid to traditional behavior modification in a variety of common management and handling situations, using various application protocols and pheromone delivery methods provided to us by the product developer or obtained commercially (McDonnell and co-workers, unpublished). So far in those controlled blind studies, no statistically significant benefit of treatment has been evident. To the author’s knowledge, developers of the pheromone product have not proposed application to stallion behavior modification, and no data are available specifically for effects of a calming pheromone on problematic male-typical sexual behavior or aggression. Though often unrecognized by handlers, there is typically an element of fear and anxiety to stallion handling problems, either as the primary cause of colt or stallion ‘misbehavior’ or secondary to attempts by handlers to discipline the horse. Further investigation of a calming pheromone approach for use with stallion behavior modification, for breeding or otherwise, would be worthwhile.
References 1. Martin BB, McDonnell SM. Lameness in breeding stallions and broodmares, In Ross M (ed.) Equine Lameness. Philadelphia: Saunders, 2003; pp. 1077–84.
2. Terry RL, McDonnell SM, Soma LS, Liu Y, Uboh CE, Van Eps A, Moate P, Driessen B. Phramacokinetic profile and pharmacokinetic effects of gabapentin following intravenous and oral administration in the horse. BEVA Annual Meeting, September 2009, Birmingham, UK. 3. Doucet MY, Bertone AL, Hendrickson D, Hughes F, MacAllister C, McClure S, Reinemeyer C, Rossier Y, Sifferman R, Vrins AA, White G, Kunkle B, Alva R, Romano D, Hanson PD. Comparison of efficacy and safety of paste formulations of firocoxib and phenylbutazone in horses with naturally occurring osteoarthritis. J Am Vet Med Assoc 2008;232:91–7. 4. McDonnell SM, Kenney RM, Meckley PE, Garcia MC. Conditioned suppression of sexual behavior in stallions and reversal with diazepam. Physiol Behav 1985;34(6):951–6. 5. McDonnell SM, Diehl NK, Garcia MC, Kenney RM. Gonadotropin releasing hormone (GnRH) affects precopulatory behavior in testosterone-treated geldings. Physiol Behav 1989;45:145–9. 6. McDonnell SM, den Rooijen JWM, van IJzendoorn, JM. Transdermal application of supplementary testosterone to stallions. Voorjaarsdagen European Veterinary Congress, Amsterdam, 27–29 April 2007. 7. Goolsby HA, Brady HA, Prien SD. The off-label use of altrenogest in stallions: a survey. J Equine Vet Sci 2004;24: 72–4. 8. Brady H. Effects of oral altrenogest on testicular parameters and seminal characteristics in young stallions. Proc Am Assoc Equine Pract 1997;43:195. 9. Heninger NL, Brady HA, Herring AD, Prien SD, Guay KA, Janecka LA. The effects of oral altrenogest on hormonal, seminal and behavioral profiles of two-year-old stallions. Prof Anim Sci 1999;17:75–80. 10. Johnson NN, Brady HA, Whisnant CS, LaCasha PA. Effects of altrenogest on sexual/aggressive behavior, hormonal profiles, and testicular parameters in young stallions. Equine Vet J 1997;18:249–53. 11. Miller C. Effects of altrenogest on behavior and reproductive function of stallions. Proc Am Assoc Equine Pract 1997;43:197. 12. Squires EL, Badzinski SL, Amann RP, McCue PM, Nett TM. Effects of altrenogest on total scrotal width, seminal characteristics, concentrations of LH and testosterone, and sexual behavior of stallions. Theriogenology 1994;48: 313–28. 13. Stout TAE, Collenbrander B. Suppressing reproductive activity in horses using GnRH vaccines, antagonists or agonists. Anim Reprod Sci 2004;82–3:633–43. 14. Fiamengo T, Kutzler M. Suppression of estrus in mares using commercially available canine GnRH vaccine. Clinical Theriogenology Annual Conference Proceedings, 25–29 August 2009, p. 232. 15. Grimmett A, Sillence MN. Calmatives for the excitable horse: a review of L-tryptophan. Vet J 2005;170:24–32. 16. McDonnell SM. Oral imipramine and intravenous xylazine for pharmacologically-induced ex-copula ejaculations in stallions. Anim Reprod Sci 2001;68:153–9. 17. Falewee C, Gaultier E, Lafont C, Bougrat L, Pageat P. Effect of a synthetic equine maternal pheromone during a controlled fear-eliciting situation. Appl Anim Behav Sci 2006;101:144–53. 18. Dodman NH, Wrubel KH, Cottam N. Effect of a synthetic equine maternal pheromone during a controlled feareliciting situation: A critique on Falewee (2006). Appl Anim Behav Sci 2008;109:85–7.
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Chapter 144
Specific Erection and Ejaculatory Dysfunctions Sue M. McDonnell
Erection and ejaculation failure can be primarily due to inadequate interest and arousal or to musculoskeletal discomfort or a disability that impairs or distracts from necessary focused attention to the mare and thrusting effort. In contrast, there are certain cases of erection and ejaculation dysfunction in which arousal and musculoskeletal function appear normal, and so the cause of the dysfunction appears more likely specific to the erection and ejaculation process itself. Specific erection and ejaculation dysfunction are commonly the result of traumatic injury; vascular disease such as aortic iliac disease that compromises perfusion of the pelvic organs and hind limbs; neurological disease or genital tract infection that affects the pelvic structures and mechanisms serving erection and/or ejaculation; and cystic structures or sperm occlusions that interfere with emission of sperm and seminal fluids.
Erection dysfunction Deviations Stallions may suffer several types of inadequate or abnormal tumescence of the corpora cavernosa or spongiosa. Asymmetrical tumescence of the corpora cavernosa or localized injury to the overlying tunica albuginea can result in lateral deviations. Possibly, delayed tumescence of the distal relative to the proximal corpora cavernosa could result in failure of full erection. Deviation can be mild to severe enough to impair insertion for natural cover, even with assistance. Most deviations vary in degree with the level of arousal. Most become less pronounced with higher levels of arousal and greater tumescence. For example, some stallions may have a pronounced curvature before mounting, but upon mounting the
penis becomes more rigid and less deviated. For others, the curvature may increase with heightened tumescence. Deviations are commonly associated with a history of penile trauma, including breeding accidents and stallion rings or other constricting devices or mutilations applied in an effort to stop unwanted sexual response or spontaneous erections. Ultrasonographic examination of the erect penis can sometimes provide insight into the degree of damage to underlying tissues. In some, but certainly not all, cases of deviation, ejaculation may appear delayed, and there may be a suggestion that penile sensation and proprioception are impaired. Reduced sensation and proprioception Stallions may exhibit a variety of behavior patterns suggesting reduced sensation of the penis and/or proprioceptive ability to efficiently achieve and maintain insertion. These are often associated with a history of trauma or disease. Response to tactile or thermal stimuli may seem reduced. For example, during washing of the penis, the stallion may be less reactive than noted before the injury. Such stallions may be awkward with natural insertion, as if unaware of the position of the penis or unable to direct the penis efficiently. Once inserted, some may continue with seeking thrusts and postural adjustments rather than ‘locking in’ and committing to rhythmic pelvic thrusting, as if unaware that the penis is inserted. In some instances, tumescence of the glans penis may be delayed or absent in response to thermal and tactile stimulation. In some cases, the apparent deficits in response can be overcome by increasing intensity of tactile and thermal stimulation. In the case of semen collection, increased temperature and pressure
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of the artificial vagina may be helpful in eliciting an adequate response. In many cases the distal penis appears to be more affected, so focus on thermal and tactile stimulation of the most proximal aspect of the penis can often elicit adequate coupling and thrusting to achieve ejaculation. Premature tumescence of the glans penis In the usual sequence of erection and ejaculation events, complete tumescence of the corpus spongiosum occurs as emission and ejaculation commence. A somewhat common problem seen in stallions is a tendency for early tumescence of the corpus spongiosum glandis, such that the glans penis becomes too large for insertion. In some instances of premature tumescence, the sequence is maintained and so it is in essence premature ejaculation. In other instances, the tumescence is not immediately followed by ejaculation, and so it appears that there is a disturbance of the natural sequence. In such instances, even if insertion is achieved, ejaculation usually does not follow. In the author’s experience, such disturbances of the ejaculatory sequence have been most common in older stallions with neurological conditions, with other signs indicating impaired control of the pelvic neurovascular structures, for example, reduced anal and bladder tone, irregular and inconsistent pattern of ejaculatory urethral contractions and/or fraction constituents. Delayed detumescence Another irregularity of erection seen in stallions is delayed detumescence of the penis, such that the full tumescence of the corpus spongiosum glandis, with or without detumescence of the corpus cavernosum penis, persists for minutes after dismount. This typically occurs in stallions with other erection and ejaculation dysfunction, after an unsuccessful mount during which the stallion reached the point of imminent ejaculation but failed to ejaculate. While delayed detumescence can occur occasionally in some stallions, there are some stallions for which the problem occurs frequently. ‘Bent’ or ‘folded’ penis An often confusing type of erection dysfunction involves the folding back of the penis within the prepuce as tumescence commences. The behavioral hallmark of this situation is the appearance of arousal and readiness to mount, without the penis visibly entended. The stallion may also appear uncomfortable or intermittently distracted, pinning the ears, kicking toward the abdomen, and/or stepping gingerly
on the hind legs. Close visualization reveals a tightly stretched and rounded prepuce. The erection may not resolve without removing the stallion from the mare or breeding area. Once the penis detumesces and is fully withdrawn, application of a lubricating cream to the prepuce usually facilitates subsequent normal protrusion. The backfolding tends to repeat occasionally over time with particular stallions. Among the cases we have seen with repeated backfolding, there have been stallions with accumulation of sticky smegma, as well as stallions in which the penis and sheath is fully cleansed one or more times daily for breeding and appears especially ‘dry’.
Ejaculation dysfunction Premature ejaculation Once mounted, inserted, and well coupled, most stallions ejaculate within 10–15 seconds on the sixth, seventh, or eighth organized, committed thrust. There is appreciably normal variation among stallions in the rhythmic pattern, number, and vigor of copulatory thrusts. Similarly, the ejaculatory pattern, including the number and rhythmic sequence of urethral contraction, varies among stallions and to a lesser extent between occasions in the same stallion. The number of urethral contractions and corresponding pulses of semen and tail movements typically exceed the number of thrusts by 1 to 3. The volume and composition of the ejaculatory fractions also vary among stallions, but are typically quite consistent within stallion given similar conditions of breeding interval and exposure to mares. Rarely, some stallions routinely ejaculate with very little stimulation, either before insertion or with fewer than three thrusts. In the stallion, it would be reasonable to consider ejaculation premature if it occurs before well-coupled insertion that would ensure delivery of semen at the cervix of the mare. The glans penis typically fully engorges just as emission commences, such that for stallions that tend to ejaculate early with little stimulation, insertion may be impaired as a result of the glans tumescence and they may ejaculate outside the vagina. Another variation is for glans penis tumescence to actually occur prematurely, and out of the normal sequence, well before emission of semen rather than at the same time as emission. In this scenario, insertion is impaired and copulation is usually interrupted.
Delayed ejaculation and ejaculation failure It seems reasonable to propose that ejaculation requiring more than one mount with greater than 10
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Specific Erection and Ejaculatory Dysfunctions or more well-organized thrusts per mount, or greater than 12 thrusts on a single mount, should be considered delayed ejaculation worthy of monitoring and evaluation if persistent. This is sometimes explainable by suboptimal conditions inherent to the domestic breeding environment, or it may represent true ejaculatory dysfunction. We recommend trying to improve any suboptimal conditions with the goal of achieving one-mount ejaculation, and careful monitoring of the horse for any sources of discomfort that may be interfering with normal efficient one-mount ejaculation. This would include: (i) aggressive evaluation and treatment of any musculoskeletal problems including the hindlimbs, forelimbs, back, neck, and pelvis; (ii) evaluation of the pelvic tubular tract with transrectal ultrasonography1 and/or urethroscopy; (iii) evaluation of behavior and handling and breeding environment to identify any improvements that would reduce distraction. Clinical impression is that stallions with major muscle disease, such as hyperkalemic periodic paralysis, appear to be overrepresented in the population of stallions with ejaculation problems. Some affected stallions appear to sustain excellent thrusting effort, yet fail to reach emission or ejaculation. It is not known whether ejaculatory dysfunction in this class of stallions is primarily due to impairments of hindlimb skeletal muscles needed to sustain the mounting and thrusting efforts, or the detraction of muscular discomfort with thrusting effort, or whether the disease also impairs function of muscles of the pelvic ejaculatory apparatus.
Incomplete ejaculation The normal sequence of events of ejaculation commences with emission of the testicular contribution from the ampullae into the seminal colliculus as the smooth muscle contractions of the urethra commence, followed by emission of seminal fluids and gel from the accessory sex glands as contractions proceed. A variety of incomplete ejaculations have been observed in stallions. These include emission of semen, either just from the ampullae, or from some or all of the contributing glands, without the smooth muscle contractions known as ejaculation proper. In this instance, semen in various volumes dribbles from the urethra rather than appearing in forcefully expelled fractions. Emission may be incomplete, in that some glands may not contribute. It is difficult to know how often this may occur unnoticed. It becomes quite evident in stallions with advanced neurological disease affecting the pelvic organs, where the day-to-day variation in semen characteristics can be remarkable.
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Occluded ampullae A fairly common, and probably under-diagnosed, condition in stallions is occlusion of one or both of the ampullary ducts with accumulated sperm.2 This can result in an ejaculate with reduced or no sperm, or, in some cases, can result in failure to ejaculate in what appears to be discomfort just at the moment that emission and ejaculation would be expected to commence. In some instances, dense material within the distal ampulla can be appreciated on ultrasonographic imaging of the ampullae. Classically cited semen indications of accumulated sperm, including a large number of detached heads, high concentration of sperm, and clumps of inspissated sperm may or may not be evident, depending upon the degree and characteristics of the occlusion. Urospermia and retrograde ejaculation The bladder neck normally remains tightly closed during emission and ejaculation, such that all semen moves through the urethra and no urine is spilled from the bladder. In conditions in which the bladder neck is compromised by injury or infection, or in certain neurological conditions or states affecting the pelvic neural control, urine may spill into the ejaculate or some or all of the semen may be delivered into the bladder. The latter is known as retrograde ejaculation. Retrograde ejaculation is suspected when ejaculatory pulses are present, but semen volume is very low, and confirmed by recovery of significant numbers of sperm from urine collected from the bladder immediately after ejaculation. Retrograde ejaculation, which is quite common in men, has been confirmed (by recovery of significant numbers of sperm via catheterization of the bladder) in only a few instances in stallions (Brinsko, 2001;3 McDonnell, two unpublished cases; Varner and Love, two unpublished cases). Psychogenic erection and ejaculation dysfunction Some of the forms of specific erection and ejaculation dysfunction described here can occur transiently under unusual circumstances of sexual arousal that could logically result in disturbances of the finely coordinated sequence of sympathetic and parasympathetic balance supporting erection and ejaculation. For example, in highly aroused or urgent, yet approach-avoidance conflicted young novice breeders, it is not uncommon for premature or delayed ejaculation, incomplete ejaculation, or urine spillage to be evident in early sessions, and then resolve with experience. Similarly, certain specific conditions can also occur in experienced stallions that are managed
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or handled such that they are showing signs of strong approach-avoidance conflict. Such a state likely disturbs the delicate sympathetic-parasympathetic balance required for erection and ejaculation. A common example is the stallion that, through experience, anticipates being rushed by the handling team to dismount after ejaculation, and so develops a pattern of what appears to be delayed ejaculation in association with behavioral signs of anxiety (worried eye, ear movement of anticipation) just as emission and ejaculation appear imminent. Similar dysfunction has been long-appreciated and well-described in humans. In animals, such psychological distraction is challenging to distinguish from other forms of discomfort and other physical distractions. Medication-related erection and ejaculation dysfunction Unlike in human cases, there are few medications used routinely in breeding stallions that have adverse effects on erection and ejaculation. Exceptions worth
mentioning are drugs that have been used specifically to manipulate erection and ejaculation in stallions. The effective dose range for enhancing erection and ejaculation can be relatively narrow, and the dose–response relationship is not linear, such that both higher or lower doses can inhibit erection and/or ejaculation. In practice, the author has found that many of these medications are currently either prescribed by veterinarians or administered to stallions without veterinary advice at lower and higher doses than recommended.
References 1. Pozor M. Diagnostic applications of ultrasonography to stallion’s reproductive tract. Theriogenology 2005;64: 505–9. 2. Love CC, Riera FL, Oristaglio RM, Kenney RM. Sperm occluded (plugged) ampullae in the stallion. Proceedings of the Annual Conference of the Society for Theriogenology, San Antonio, TX, 1992; pp. 117–25. 3. Brinsko SP. Retrograde ejaculation in a stallion. J Am Vet Med Assoc 2001;218:251–3.
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Chapter 145
Stereotypic Behavior of Stallions Katherine A. Houpt
Equine stereotypies are common and not well understood. Formerly known as stable vices, they are now viewed as indicative of deficiencies in management and, therefore, signs of poor welfare. Whether or not stereotypies serve to help the horse cope with a poor environment is still unknown. There has been considerable research done on equine stereotypies and the management factors and breed predispositions that make an individual horse more likely to show the behavior. One group of investigators reported that the management factors that predispose to stereotypies are access to less than 6.8 kg/day of forage; use of bedding other than straw; stall designs that minimize contact between horses; and providing access to hay rather than other types of forage.1
Time spent cribbing Several groups have studied the time spent cribbing. The horses we have studied crib 15–21% of the time. Kusunose6 found that cribbing occurred at a very low rate when the horses had no feed available or when hay was available, but increased markedly when grain was fed. Gillham et al.7 measured cribbing for 30 minutes before and after various feeds were given. The cribbing rate rose from 2/5 minutes to 27/5 minutes when the horse was eating sweetened grain or pellets, but rose to only 8/5 minutes when alfalfa pellets were fed. McGreevy et al.8 found that air was not swallowed when a horse cribs, but the bolus of air could be pushed down with the food. Because cribbing is most likely to occur while the horse is eating, gastrointestinal consequences may occur.9
Oral stereotypies Prevalence of cribbing Cribbing Cribbing is addressed in Chapter 292, but a slightly different perspective is given here. Some studies indicate that males are more likely to crib. Luescher et al.2 found that males, whether intact or castrated, were more likely to crib. Mills et al.3 found that young males were most likely to crib. It is widely believed that horses learn cribbing by observing other horses crib, but there is no evidence for that. In fact, numerous attempts to have horses learn other activities by observing demonstrator horses have failed.4 We have recently conducted a survey of recreational horse owners. Of all the horses in 112 stables to which a cribbing horse was added, only 12 began to crib after the cribber arrived.5
The prevalence of cribbing has been determined by postal survey, telephone, or direct contact in different countries: the UK,1, 3, 10 Sweden,11 Australia,12 Canada,2 and the USA.5 The percentage of adult crib-biting horses varied from 0.4% of Swedish trotters11 to 3.8% in Prince Edward Island,13 to 8.3% of eventing horses.1 All the surveys in which more than one breed was observed found that Thoroughbreds have a higher incidence of crib biting1, 2, 5, 11 than other breeds, suggesting a heritable component (Table 145.1). Earlier studies of Japanese and Italian Thoroughbreds also indicated a heritable component.14, 15 The use to which the horse is put may influence the incidence of crib biting. Race horses and competition
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 145.1 Prevalence of stereotypies: percentage affected Author
Location
Crib
Weave
Stall walk
Bachmann et al.28 Luescher et al.2 McGreevy et al.1 Albright et al.5 Christie et al.13 Redbo et al.11 Vecchiotti and Galanti15 Prince29 McBride and Long10
Switzerland Canada (Ontario) UK USA Canada (Prince Edward Island) Sweden Italy UK UK
2.1 4.1 3.1–7.5 5.5 3.8 2.8 2.4 4.2 1.5–3.7
1.0 2.0 4–9.5
0.43 2.0 3.6–5.5
4.8 5.0 2.5 2.8 2.1–3.9
1.0 2.5 1.1 0.3–1.1
horses have a higher rate of cribbing than riding school, pleasure, or endurance horses,1, 10 but this may be confounded by the fact that virtually all race horses in the UK are Thoroughbred and show horses are often Thoroughbred or Thoroughbred crosses, such as Warmbloods, whereas endurance horses are usually Arabians. Heritability appears to be 61% in Thoroughbreds. Waters et al.16 has performed the only prospective study and found that 10% of weanling foals crib bite, a figure higher than reported in adult Thoroughbreds. This indicates either that cribbers die or that some of them stop crib biting.
Cribbing and ulcers The relationship of cribbing to gastric ulcers is interesting. Nicol et al.17 found that foals that cribbed had more severe gastric ulcers and a lower fecal pH than non-cribbing foals. These investigators hypothesized that cribbing may increase saliva production, which aids in buffering gastric acidity. This is a form of selfmedication which may occur in the animal world.18 Horses do not salivate more when they crib, but Nicol et al.,17 as well as Mills and Macleod,19 found a tendency for antacids to reduce crib biting. Maintenance on pasture and a diet without sweet feed but with antacids may be the best management strategy. Wood chewing Many lay people confuse wood chewing with cribbing, but they are very different behaviors. Cribbers grasp a piece of wood (or metal or plastic) and pull back, leaving only half-moon shaped marks on the wood. Wood chewers prehend, masticate, and ingest the wood. Wood chewing may be considered a stereotypic behavior, but appears to have a function – increasing roughage; in this case, non-digestible lignin. Wood chewing is most common in cold, wet weather when most grass would be buried under snow or very low in nutrients.
Wood chewing is very destructive because fences or barn walls can be ingested. Wood chewing can result in obstruction or a penetrating foreign body in the gastrointestinal tract. The treatment for wood chewing is relatively easy: increase the variety and amount of roughage. Surfaces can be treated with repellents or covered with metal, or a vitamin mineral supplement can be fed, but providing roughage is the best approach.
Locomotor stereotypies Stall walking and weaving are discussed in Chapter 292.
Self-mutilation Self-mutilation in stallions consists of biting at or actually biting the flanks, hind limb, and, less frequently, the chest or forelegs, often while spinning. This behavior is usually accompanied by squealing and kicking. Approximately 2% of stallions exhibit the behavior.2 McDonnell20 recognized three forms: (i) secondary to pain, (ii) self-directed male competitive aggression, and (iii) stereotypic, which is repetitive and directed to inanimate objects such as the stall walls. This behavior can occur in geldings, usually those housed with mares, especially their dams. The syndrome is less violent in geldings. A few mares have been reported to exhibit a similar syndrome. It is essential to eliminate the first form of selfmutilation through diagnostic work. In this type of self-mutilation, work or anticipation of work is the usual stimulus, presumably because the pain is greater when the horse exercises. An example of the first form is that of a 14-year-old Arabian stallion that began to self-mutilate at age 10 and had hemospermia. A squamous cell carcinoma was removed from the urethral process and the self-mutilation, as well as the hemospermia, ceased.21 The third form of selfmutilation is characterized by more repetitive, less
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Stereotypic Behavior of Stallions violent behavior, often confined to certain locations or times of day – especially feeding time. It often includes kicking at stall walls or fences and rarely results in serious wounding of the stallion. The usual treatment for stereotypic behavior – improvement in the environment and reduction of frustration – should reduce the occurrence of the behavior. The second form is the most serious and difficult to treat. It appears to be self-directed aggression because the actions mimic those of two stallions interacting before and during aggressive encounters.20 Bouts of self-mutilation can be initiated by visual stimuli – usually another horse – or the smell of another stallion’s feces or urine, or the sound of stallion vocalizations. These stimuli need not be from other stallions; a stallion’s own reflection in a mirror, the odor of his own feces, urine or body, or a tape-recording of his vocalizations can trigger selfmutilation, as can the sight of another horse. The behavior begins gradually and usually intensifies. It can occur in suckling foals when the social frustration is deprivation of the dam. Treatment of self-mutilation should concentrate on removing as many sources of frustration as possible, i.e., avoiding stimuli from other stallions. The optimal treatment is pasturing with mares, although sometimes this is not practical. Cradles, bibs, or muzzles can be used to prevent injury, but this is not changing the horse’s motivation. A variety of drugs have been used to treat self-mutilation, including opiate blocks,22 progestins, tryptophan, and tricyclic antidepressants.20 Castration eliminates self-mutilation in 30% of self-mutilators, but has no effect on another 30% with some improvement in the remainder.23 Increasing exercise and roughage and decreasing grain has been successful24 in the treatment of a case of self-mutilation. The stallion’s ration was reduced from 3.6 kg of sweet feed and 3.6 kg of oats to 0.25 kg of sweet feed; 4.5 kg of alfalfa was omitted from his diet and replaced with ad libitum hay. He was handwalked for 25 minutes per day. The self-mutilation stopped and, when the horse returned to the owners, round-pen training for 20 minutes four times a week and a companion goat were enough, with the dietary changes, to resolve the self-mutilating behavior.
Masturbation Masturbation is not a stereotypic behavior. It is a normal behavior of stallions, but is considered here because so many owners believe it is an abnormal behavior or, at least, a misbehavior. Geldings also masturbate, albeit at lower frequency. Masturbation consists of lifting the erect penis against the belly wall. Stallions rarely ejaculate while masturbating.
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An erection may last 30 minutes, whether masturbation occurs or not. Stallions masturbate two or three times a night for an average of 3 minutes. Masturbation is most frequent between midnight and 03:00, and frequently follows recumbent rest. Despite this nocturnal pattern, stallions masturbate at all times of the day and night.25
Headshaking Head shaking is probably not a stereotypic behavior, but, because it is frequently considered one by owners, it will be discussed here. Vertical head shaking can be so forceful that the horse is uncomfortable to ride. Males are more affected than females. There are numerous causes of head shaking and there are, no doubt, some unrecognized ones. Trigeminal irritation is often diagnosed and is associated with horses that head shake more in sunshine than in shadow. Other causes can be allergies, foreign bodies in the nasal cavity, or any abnormality of the head or upper neck. Rider-associated causes can be illfitting tack, over-collection, and simply poor riding technique. The horse should be observed in its stall and its turn-out area and then while being lunged or ridden. The passage of air over the membranes of the nose and pharynx seems to exacerbate head shaking. The treatments consist of removing the stimulus. If an airborne allergen is suspected, a nose mask should help to filter out the pollens, etc. If a photic reaction is suspected, then a face mask or a head band that serves to curtain the eyes should be useful. Drug treatment consists of cyproheptadine (0.3 mg/kg orally twice daily)26 and carbamazepine 4 mg/kg four times daily.27
References 1. McGreevy PD, Cripps PJ, French NP, Green LE, Nicol CJ. Management factors associated with stereotypic and redirected behaviors in the Thoroughbred horse. Equine Vet J 1995;27:86–91. 2. Luescher UA, McKeown DB, Dean H. A cross-sectional study on compulsive behavior (stable vices) in horses. Equine Vet J 1998;Suppl. 27:14–18. 3. Mills DS, Alston RD, Rogers V, Longford NT. Factors associated with the prevalence of stereotypic behavior amongst Thoroughbred horses passing through auctioneer sales. Appl Anim Behav Sci 200278:115–124. 4. Lindberg AC, Kelland A, Nicol CJ. Effects of observational learning on acquisition of an operant response in horses. Appl Anim Behav Sci 1999;61:187–9. 5. Albright JD, Mohammed HO, Heleski CL, Wickens CR, Houpt KA. Crib-biting in US horses: breed predispositions and owner perceptions of aetiology. Equine Vet J 2009;41:455–8. 6. Kusunose R. Diurnal pattern of cribbing in stabled horses. Jpn J Equine Sci 1992;3:173–6.
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7. Gillham SB, Dodman NH, Shuster L, Kream R, Rand W. The effect of diet on cribbing behavior and plasma βendorphin in horses. Appl Anim Behav Sci 1994;41:137. 8. McGreevy PD, Richardson JD, Nicol CJ, Lane JG. Radiographic and endoscopic study of horses performing an oral based stereotypy. Equine Vet J 1995;27:92–5. 9. Archer DC, Freeman DE, Doyle AJ, Proudman CJ, Edwards B. Association between cribbing and entrapment of the small intestine in the epiploic foramen in horses: 68 cases (1991–2002). J Am Vet Med Assoc 2004;224:562–4. 10. McBride SD, Long L. Management of horses showing stereotypic behavior, owner perception and the implications for welfare. Vet Rec 2001;148:799–802. ¨ 11. Redbo I, Redbo-Torstensson P, Odberg FO, Hedendahl A, Holm J. Factors affecting behavioural disturbances in racehorses. Anim Sci 1998;66:475–81. 12. Pell SM, McGreevy PD. A study of cortisol and betaendorphin levels in stereotypic and normal Thoroughbreds. Appl Anim Behav Sci 1999;64:81–90. 13. Christie JL, Hewson CJ, Riley CB, McNiven MA, Dohoo IR, Bate LA. Management factors affecting stereotypies and body condition score in nonracing horses in Prince Edward Island. Can Vet J 2006;47:136–43. 14. Hosoda T. On the heredity of wind-sucking dispositions of horses. Jpn J Zootech Sci 1950;21:2–28. 15. Vecchiotti G, Galanti R. Evidence of heredity of crib-biting, weaving and stall-walking in Thoroughbred horses. Livest Prod Sci 1986;14:91–5. 16. Waters AJ, Nicol CJ, French NP. Factors influencing the development of stereotypic and redirected behavior in young horses: the findings of a four year prospective epidemiological study. Equine Vet J 2002;34:572–9. 17. Nicol CJ, Davidson HPD, Harris PA, Waters AJ, Wilson AD. Study of crib-biting and gastric inflammation and ulceration in young horses. Vet Rec 2002;151:658–662.
18. Hart BL. The evolution of herbal medicine: behavioural perspectives. Anim Behav 2005;70:975–89. 19. Mills DS, MacLeod CA. The response of crib-biting and windsucking in horses to dietary supplementation with an antacid mixture. Ippologia 2002;13:1–9. 20. McDonnell SM. Practical review of self-mutilation in horses. Anim Reprod Sci 2008;107:219–28. 21. Bedford SJ, McDonnell SM, Tulleners E, King D, Habecker P. Squamous cell carcinoma of the urethral process in a horse with hemospermia and self-mutilation. J Am Vet Med Assoc 2000;216:551–3. 22. Dodman NH, Shuster L, Court MH, Dixon R. Investigation into the use of narcotic antagonists in the treatment of a stereotypic behavior pattern (crib-biting) in the horse. Am J Vet Res 1987;48:311–19. 23. Dodman NH, Normile JA, Shuster L, Rand W. Equine self-mutilation syndrome (57 cases). J Am Vet Med Assoc 1994;204:1219–23. 24. McClure SR, Chaffin MK, Beaver BV. Nonpharmacologic management of stereotypic self-mutilative behavior in a stallion. J Am Vet Med Assoc 1992;200:1975–7. 25. Wilcox S, Dusza K, Houpt KA. The relationship between recumbent rest and masturbation in stallions. Equine Vet Sci 1991;11:23–6. 26. Madigan JE, Bell SA. Owner survey of headshaking in horses. J Am Vet Med Assoc 2001;219:334–7. 27. Newton SA. Idiopathic headshaking in horses. J Equine Vet Educ 2005;17:79–82. 28. Bachmann I, Audig´e L, Stauffacher M. Risk factors associated with behavioural disorders of crib-biting, weaving, and box-walking in Swiss horses. Equine Vet J 2003;35: 158–63. 29. Prince D. Stable vices. In: McBane S (ed.) Behaviour Problems in Horses. North Pomfret, VT: David & Charles, Inc., 1987, pp. 115–22.
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Section (f): Specific Techniques Used in Reproductive Examination of the Stallion
146.
Historical Information Charles C. Love
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Diagnostics and Therapeutics for Stallions with Declining Fertility: An Endocrine– Paracrine–Autocrine Approach Janet F. Roser
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Endoscopy of the Internal Reproductive Tract Carla L. Carleton
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Examination of External Genitalia Patricia L. Sertich
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Cytogenetic Evaluation Keith W. Durkin, Terje Raudsepp and Bhanu P. Chowdhary
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Ultrasonography of the Genital Tract Regina M.O. Turner
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Chapter 146
Historical Information Charles C. Love
Fertility depends on mare, stallion, and management factors working simultaneously to maximize foaling rate. Fertility/subfertilty estimation often relies primarily on the evaluation of semen quality, with little diagnostic attention paid to the mare and the management of the mare and stallion. Often, the stallion is assumed to be the sole source of subfertility, simply because mare and management factors have not been evaluated. Primary causes of reduced fertility in the stallion include disease, especially those that elevate the body temperature for prolonged (> 24–48 hours) periods of time. Medications, notably anabolic steroids, and recently vaccination with GnRH analogs are two examples of iatrogenic causes of reduced testes size and poor libido. Individual stallions transported between hemispheres to accommodate two breeding seasons in one calendar year (i.e., shuttle stallions) may show transient changes in fertility in October (southern hemisphere breeding) especially as mare numbers increase due to the presentation of foaling mares. These stallions may have a stress response due to shipping 50–60 days earlier, resulting in decreased spermatogenesis. Mare factors include mare age, pre-breeding status (i.e. maiden, foaling, barren). This presentation describes our diagnostic methodology for describing and defining non-sperm factors and the role they play in understanding and diagnosing the primary cause(s) of reduced fertility. Assuming a stallion does not have a dramatic sperm-limiting factor such as extremely poor semen quality or very low sperm numbers, a working hypothesis is that semen quality accounts for 20–40% of the variability in the fertility of the average mare–stallion–management complex. This suggests, therefore, that 60–80% of fertility can be explained by non-sperm factors, broadly categorized as management and mare factors. The data
described in this chapter has been summarized using a self-designed computer program. Outlined below is a series of end points that we use to describe non-stallion factors when evaluating breeding farm fertility. The following end points are used as the framework for evaluation:
r r
Mare – identification of the mare; Beginning status – the reproductive status of the mare prior to the current breeding season. General classifications include maiden, barren, and foaling.
Maiden mares This group of mares have never been bred. They tend to be young; however, there are individual mares that will be older maiden mares and they can be much less fertile. In general, this group should be of high fertility. Mares that are recently retired from performance careers may not be cycling normally and may have difficulty getting pregnant initially.
Barren mares This group includes mares that were bred the previous year and are not in foal. It is often assumed these mares are reproductively unsound; however, their not being in foal may not be related to reproduction problems at all. One example includes mares not bred the previous breeding season, which occurs because the mare was not bred due to foaling late in the previous season, or for other reasons such as sickness or foal problems that reduced the opportunity to get pregnant. Another example is mares classified as slipped, which were diagnosed pregnant and subsequently lost the pregnancy, due to early embryonic loss or abortion.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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In-foal (or foaling) mares These are mares that have foaled in the current breeding season and will be rebred during the same season. They are the group of highest fertility along with the maiden mares. This group may exhibit reduced fertility if a large portion of the group foals late in the breeding season and has only one or two cycles in which to get pregnant. Because of limited exposure to the stallion they have not had adequate opportunities to get pregnant. A similar effect will occur if some event (injury/illness to the stallion) prematurely shortens the breeding season.
r r
The pregnancy status of each classification The pregnancy status of each mare classification can render important information about the fertility status of a stallion. As a general rule, maiden and foaling mares should be those mares that result in the highest fertility. Barren mares may be of similar or lesser fertility. If their barrenness is due to an inherent reproductive problem and there is a large number of this type of mare in a stallion’s book, this will have the effect of artificially (i.e., not the stallion’s fault) lowering a stallion’s fertility. Conversely, mares may be barren because they were not bred the previous year, in which case their fertility would be similar to the maiden and foaling groups. For diagnostic purposes, the fertility of the maiden mares represent the true intrinsic fertility of the stallion. In some cases the foaling mares may be the group of lowest fertility. This may be due to the fact that this group foaled late in the season and had only one or two estrus cycles to be bred before the breeding season ended. In this case, neither the stallion nor mares are directly at fault, but this is a result of a lack of mare exposure to the stallion.
r r r
Dates bred – identifies the calendar day(s) that the mare was bred; Date foaled – the date the mare foaled; Status after last examination – is the current pregnancy status: either in-foal or barren or slipped.
Using these data end points, the following fertility measures can be determined: Following is a list of those end points that should be measured. We currently use a self-designed computer program to enter and measure these end points.
r
Total number of mares bred – may have an impact on fertility. It is often assumed that as mare
number increases fertility declines. There is strong evidence, especially in the Thoroughbred industry, that the opposite may occur in many instances, primarily due to the positive influence of competent breeding management. Total number of mares pregnant. Seasonal pregnancy rate – this parameter is based on the number of mares diagnosed pregnant at a particular point in the breeding season divided by the total number of mares in a stallion’s book. This end point is very important economically, but is not a sensitive indicator of a stallion’s fertility because it does not reflect the total number of cycles that a mare is bred to achieve the pregnancy. A stallion can achieve a relatively low pregnancy rate per cycle yet end the season with a seasonal pregnancy rate similar to a stallion achieving a relatively high pregnancy rate per cycle simply, by being able to breed mares for more cycles, which typically means a longer breeding season. For example, if a stallion’s per cycle pregnancy rate is recorded at 60%/cycle then 84% of mares will be pregnant after two cycles. This figure (approximately) can be achieved with a stallion achieving 50% of mares in foal per cycle if he breeds three cycles, a stallion achieving 40% of mares in foal per cycle if he breeds for four cycles and a stallion achieving 30% of mares in foal per cycle if he breeds five cycles. Pregnancy rate is the fertility end point commonly talked about, but is not a sensitive end point and does not describe the source of reduced fertility.
There are several factors that can alter seasonal pregnancy rate:
r
r
The point in time seasonal pregnancy rate is determined (i.e., during the breeding season, shortly after the breeding season, or many months after the breeding season). As the point in time approaches the next breeding season, the value will more closely approximate the foaling rate and will decrease as the effects of early embryonic loss and abortion are taken into account. The number of estrous cycles that a mare was bred. It is common for the stallion manager/owner to add mares on to a stallion’s book as the end of the breeding season approaches. Adding mares on to a stallion’s book is often done when mares are switched from one stallion to another for a variety of reasons. These mares are included in the total number of mares that the stallion breeds, but in reality are given fewer opportunities to become pregnant (usually only one cycle) than other mares in the stallion’s book. If these mares do not
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Historical Information
r
r
become pregnant, the seasonal pregnancy rate for that stallion will be lower than expected. Therefore, if a stallion is presented because of a low seasonal pregnancy rate, it is important to determine that all mares were bred during an adequate number of estrous cycles to have a reasonable chance of becoming pregnant. Inclusion or exclusion of mares that exhibit early embryonic death (EED). The assumption can be made that the majority of embryonic deaths are due to mare and not stallion factors. If an accurate seasonal pregnancy rate is to be created to describe the stallion’s inherent fertility, all diagnosed pregnancies (which indicate that the stallion was able to accomplish fertilization in those mares) should be included in calculating the seasonal pregnancy rate. Yet, keep in mind that counting an embryonic death as a pregnancy when figuring seasonal pregnancy rate is of no economic relevance (i.e., stud fees will not be transferred). If embryonic deaths are included as pregnancies in calculating seasonal pregnancy rate, an inflated economic value for the seasonal pregnancy rate is created. Cycles/pregnancy (or pregnancy rate/cycle) – this measure is determined by counting all estrous cycles from all mares that a stallion has bred and dividing it by the number of pregnancies. Counting only estrous cycles from mares that become pregnant will artificially inflate the value. This is a more sensitive indication of a stallion’s fertility because it measures how efficient a stallion is at establishing pregnancies. The assumption is made that all mares were in normal estrus at the time of breeding and were bred near the time of ovulation. This assumption is not always correct, especially for breedings by artificial insemination. No doubt many mares are bred artificially when either not in estrus or near to the time of ovulation. Some mares are bred more than once (doubled) during an estrous period and should not be counted as a separate cycle. On well-managed Thoroughbred farms, the double rate rarely exceeds 10% (services per cycle of 1.10), but on many farms using artificial insemination it is not uncommon for the rate to be 20% or greater.
Non-stallion factors Mare age The age of the mare population to which a stallion is bred can be an important influence in that stallion’s fertility. As mares age and approach 12–14 years of age, as a population, fertility declines. Some stallions
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may accumulate a larger group of mares that are older than that of the average stallion, and will experience reduced fertility, usually because these mares tend to be barren.
Opportunity to become pregnant The ability of a mare to become pregnant is influenced, to a certain degree, simply by the number of estrous cycles that she is bred regardless of the intrinsic fertility of the stallion. A reproductively normal mare that is bred once to a stallion that has a 50% per cycle pregnancy rate has a 50% chance of getting pregnant; if bred two cycles she has a 75% chance; if bred three times, she has an 87.5% chance of getting pregnant. Opportunity is less important when highly fertile stallions (i.e., high per cycle pregnancy rate) are bred because they require fewer cycles to render a mare pregnant, making opportunity more important as the fertility of a stallion decreases.
Evaluation of opportunity to become pregnant Evaluation of opportunity can be determined using several estimates:
r
r
The average day of first breeding for each mare type. This is determined by averaging the first day of breeding (i.e., calendar year number) for all mares in each mare type. The intent of this value is to determine when mares in each group started to be bred in the breeding season. Maiden and barren mares have the potential to start on 15 February, since they enter the breeding season not pregnant. In contrast, foaling mares must wait until they foal before they can be rebred and therefore, as a group, will start the breeding season later. As a general rule the average day of first breeding for maiden and barren mares should be some time in the month of March, whereas foaling mares tend to start in mid to late April. Cycles/mare for each mare type – this end point determines the number of estrous cycles that each mare was bred. Maiden and foaling mares approximate 1.3–1.6 cycles/mare, with barren mares around 2.0. Maiden mares are low because they tend to be a highly fertile group and therefore get pregnant at an efficient rate and simply do not need to be bred more; whereas foaling mares are low because they must foal before they can be bred. In the latter case, the foaling mares have a lack of opportunity in some cases because the end of the breeding season occurs before they can be bred
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again. This can result in a low seasonal pregnancy rate in foaling mares, not because the stallion is at fault, but because the mares were not bred enough. Interestingly, in some cases barren mares will have a higher pregnancy rate than foaling mares, not because they are more fertile, but because they were bred more. Seasonality of pregnancy In North America, the breeding season officially starts on 15 Februaryand extends to the end of June early July, and therefore includes both nonphysiological (short daylengths) and physiological (longer daylengths) breeding season. The fertility of a stallion can vary considerably from month –to –month, based on a number of factors. Ideally mares that are anticipated to be bred early in the breeding season have been exposed to long daylength conditions using artificial lighting to initiate normal cyclicity early in the year. Sometimes while mares may have been exposed to lighting there may be an inadequate response, resulting in prolonged cycles or a transitional-like state. This will result in reduced fertility early in the breeding season. Another cause of a reduction in fertility early in the breeding season is a large book of barren mares and/or maiden mares that are subfertile. If the barren mares group suffers from poor inherent fertility they will continue to be bred and remain not pregnant until the end of the season, and therefore will result in poor fertility late in the breeding season as well.
Overbreeding Stallions may possess highly fertile sperm characteristics (excellent sperm motility, high percent of morphologically normal sperm), but may be limited by the ability to deliver a threshold level of total normal sperm for the number of mares in his book. The effects of overbreeding are usually manifest as a reduction in the efficiency (cycles/pregnancy) by which a stallion renders his mares pregnant, which, if dramatic enough, will result in a reduction in seasonal pregnancy rate.
Daily pregnancy rate based on the number of mares bred/day This value compares the fertility based on the number of mares that a stallion breeds per day. Theoretically, a stallion that breeds only one mare/day should deposit more sperm in the uterus than when he breeds two, three, or four mares/day. This value
simply compares the overall pregnancy rate on days when one mare is bred versus two versus three versus four, etc., and determines whether pregnancy rate declines as more mares are bred/day.
Pregnancy rate based on the order in which a mare is bred during the day This value is more sensitive than the daily pregnancy rate, because it specifically compares the pregnancy rate of those mares that were the stallion’s first cover of the day, with the mares that were the second, third, fourth, etc. If a stallion exhibits overbreeding, pregnancy rates will decline on the latter covers and will tend to be more obvious when intervals between breeding are less.
Decline in pregnancy rate coincidental with a rise in the number of mares bred in a week To identify this phenomenon, pregnancy data must be charted along with the number of mares bred in the week prior to the day the pregnancy data was gathered. The intent of these values is to determine if a stallion’s ability to deliver adequate numbers of sperm is depleted after a certain number of mares bred. Evaluating the effect of breeding frequency on fertility One factor that will modulate fertility during the breeding season is the frequency that a stallion is bred. For a farm that is using natural cover (usually Thoroughbred) the frequency includes the number of times a stallion is bred in a given time period, such as the number of times bred in a day and the number of times bred in a week. On farms where artificial breeding is used, the frequency is the number of mares bred following the splitting of the ejaculate. These frequencies evaluate a short-term and a long-term time interval and the resulting pregnancy outcome from those breeding frequencies. Figures 146.1 and 146.2 give examples of stallions of high and average fertility. These figures chronologically represent the dynamic nature of fertility throughout a breeding season. Four lines represent the following parameters:
r
Mares bred in last week – identifies the number of mares bred in the week previous to the day in question. This parameter is an evaluation of the long-term effect of frequent breedings and the intent is to determine whether there is a breeding frequency above which fertility declines. This
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Figure 146.1 Example of a highly fertile stallion. Fertility (cumulative pregnancy) increases throughout the breeding season with no declines in fertility.
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phenomenon occurs in many stallions and when it becomes extreme will result in clinically reduced fertility. Mares bred previous day – identifies the number of mares bred (either by natural or artificial breeding) on that day. Pregnancy differential – the date each mare is bred she is given a pregnancy score (+1 if preg-
r
nancy results, –1 if open). These scores are added for the week previous to a particular date and shows the fertility of a stallion for the previous week. Cumulative pregnancy – this value adds the pregnancy scores (+1 or –1) for the entire year leading up to the date of interest. The zero line represents a 50% per cycle pregnancy rate.
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Combined, these lines allow the diagnostician to compare the frequency of breeding with fertility outcome, to determine if overbreeding affects a stallion’s fertility. In addition, an overview of a stallion’s fertility can be determined throughout the breeding season to evaluate periods of high fertility and compare them to periods of low fertility.
References 1. Love CC. The role of breeding record evaluation in the evaluation of the stallion for breeding soundness. Proceedings of Annual Conference Society for Theriogenology, Columbus, OH, 2003; pp. 68–77. 2. Love CC. Evaluation of breeding records. In: Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL (eds) Manual of Equine Reproduction, 2nd edn. St Louis, MO: Mosby, 2003.
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Chapter 147
Diagnostics and Therapeutics for Stallions with Declining Fertility: An Endocrine– Paracrine–Autocrine Approach Janet F. Roser
Idiopathic subfertility and infertility in stallions: the nature of a hormone imbalance Subfertility or infertility in breeding stallions contributes to low pregnancy rates per cycle (< 20% to 30%/cycle; normal, >50%/cycle) and low pregnancy rates per season (< 60%/season; normal, > 80%/season), resulting in substantial financial loss in the equine industry.1, 2 There are a number of conditions associated with subfertility or infertility in stallions such as mismanagement, anabolic steroid treatment, infection, fever, tumors, toxins, injury, disease, behavior, nutrition deficiency, age, ejaculatory failure due to bilaterally blocked ampullae, testicular degeneration, and idiopathic testicular and hormone imbalance.1–16 A number of these problems can be diagnosed and treated appropriately. In addition, a stallion with declining fertility could be managed more carefully to increase pregnancy rates per cycle by breeding mares closer to the time of ovulation, breeding mares selected for fertility, and breeding a limited number of mares per season. However, the stallion with a hormone imbalance that is associated with testicular degeneration is difficult to manage and the condition is usually non-reversible, particu-
larly if the cause of the problem is unknown. These stallions come under the category of idiopathic subfertility/infertility. The exact nature of the condition and the hormone imbalance that ensues is not apparent. The hormone imbalance may be the cause or the result of testicular dysfunction and degeneration. Typically, these stallions with idiopathic testicular dysfunction initially display low sperm motility (≤ 20–30% progressive motility) and marginal sperm morphology (< 60% normal morphology) along with a slight change in testicular consistency.1, 17 These subfertile stallions may not show a circulating hormone imbalance at first, but if a testicular biopsy was taken would likely show a decline in intratesticular inhibin.18 As the condition progresses, clinical signs become more apparent and include decreasing testicular size, palpable softening or hardening of the testicular parenchyma, decreased sperm numbers in ejaculates, low daily sperm output (DSO) per volume of testicular parenchyma, the appearance of increasing numbers of immature round spermatogenic cells and/or multinucleate giant cells in the ejaculate, and an overall decline in semen quality.2, 3,19–22 These stallions are often presented with a decrease in circulating inhibin and estradiol-17β levels and an increase in follicle-stimulating hormone (FSH)
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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followed by luteinizing hormone (LH) levels.1, 17 Circulating testosterone can also decline, but appears to be the last hormone affected.1, 22
Endocrine–autocrine–paracrine malfunction in stallions with declining fertility (Figure 147.1)
Primary endocrine malfunction
A comprehensive understanding of the endocrine– paracrine–autocrine regulation of testicular function in the stallion would enable the clinician to identify, evaluate, and manage the idiopathic subfertile/infertile stallion. Such an understanding provides insight into the normal dynamics of the hypothalamic–pituitary–testicular axis (HPT) and the normal paracrine–autocrine interactions that modulate testicular function.15 This will permit the veterinarian to make informed decisions on abnormal mechanisms that may arise. A significant percentage of idiopathic subfertile/infertile stallions have been diagnosed with testicular degeneration.2, 3 These stallions make up a heterogeneous population of poor breeding performers with unknown pathophysiology. Changes in circulating reproductive hormones are undetectable and slow to change and then quickly become abnormal.3, 23
Endocrine-Paracrine-Autocrine Regulation of the Testis
Hypothalamus Pituitary GnRH
LH, FSH T, E2 ±
Inhibin –
A primary endocrine disorder or a primary paracrine–autocrine disorder: which comes first? (Figure 147.1)
± E2
Testis Germ Cells
Basal concentrations of circulating hormones Historically, clinicians have relied on baseline concentrations of reproductive hormones in the stallion to aid in evaluation of breeding soundness. For example, low levels of testosterone were observed in azoospermic stallions.24 Burns and Douglas13 made the first association between elevated plasma concentrations of FSH and subfertility in stallions. Two studies indicated that circulating plasma levels of FSH and estrogens, and not LH and testosterone, could be good markers to predict future changes in fertility,1, 25 whereas another group of investigators suggested that measurement of serum levels of estrogens and testosterone may help in the diagnosis of infertile stallions when other methods are not available.26 In addition to measuring basal levels of circulating hormone concentrations in subject stallions by radioimmunoassay (RIA) or enzyme immunoassay, it may be useful to measure bioactivity of these hormones by using radioreceptor assays, in vitro cell culture, or tissue culture techniques. It has been reported that the populations of LH and FSH are heterogeneous in nature, being composed of different isoforms.27 The heterogeneous nature of these gonadotropins is based on their carbohydrate sidechains and, in particular, their sialic acid residues. Different isoforms of LH or FSH can have an impact on receptor binding, signal transduction, and metabolic half-life. The reproductive state of the animal can change the population of different isoforms.27 Whitcomb and co-workers28 demonstrated the presence of what appears to be a circulating bioinactive LH isoform in stallions with poor breeding performance.
Leydig cells Sertoli cells Figure 147.1 Overview of the endocrine–paracrine–autocrine systems. The paracrine–autocrine systems modulate the actions of the endocrine system at the level of the testis. Endocrine factors are secreted by one organ to regulate actions in another organ (dashed arrows). Endocrine factors are present in the peripheral circulation. Intratesticular paracrine factors are secreted by one cell type in the testis to modulate the actions taking place in another testicular cell type (solid arrows). Intratesticular autocrine factors are secreted by one cell type to modulate the actions in the same cell type in the testis (dotted arrows). FSH, follicle-stimulating hormone; LH, luteinizing hormone; T, testosterone; E2 , estradiol.
Gonadotropin-releasing hormone challenge tests In addition to measuring basal levels of immunoactive/bioactive reproductive hormones in stallions with declining fertility, it may also be advisable to perform a gonadotropin-releasing hormone (GnRH) challenge test, whereby a bolus or intermittent doses of synthetic GnRH are given to diagnostically assess pituitary and testicular responsiveness Figures 147.2 and 147.3).25, 29 In humans, the GnRH challenge test appears to be of some value in revealing
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Diagnostics and Therapeutics for Stallions with Declining Fertility Temporal Changes in LH after GnRH in Fertile (F) and Subfertile (SF) Stallions in the Breeding (B) and Non-Breeding (NB) Season GnRH GnRH
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Time (min) Figure 147.2 Temporal changes in luteinizing hormone response, expressed in terms of percent of baseline levels, to pulses of gonadotropin-releasing hormone (GnRH) in the fertile (F; n = 8) and subfertile (S; n =6) group of stallions during the non-breeding (NB) and breeding (B) seasons. Arrows represent the time of administration of 5.0 µg GnRH in 1 ml saline. Representative time points include –5, 0, 5, and 10 minutes, every 10 minutes thereafter up to 180 minutes, and then 240 and 360 minutes. Note that GnRH was highest in fertile stallions during the non-breeding season and lowest in subfertile stallions during the non-breeding and breeding season. (Modified from Roser and Hughes.25 )
intrinsic differences in acute gonadotroph and Leydig cell stimulation between normal men and men with reproductive disorders.30–36 When stallions were challenged with repetitive pulses of exogenous GnRH, different pituitary and testicular responses were observed between fertile and subfertile stallions, which appeared to be dependent on season and gonadal status (Figure 147.2).26 The pituitary and testes of fertile stallions appeared to be more responTemporal Changes in T Response after a Single Injection of GnRH in Fertile and Subfertile Stallions T Response (Percent baseline)
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sive to pulsatile administration of GnRH in the nonbreeding season than in the breeding season.25 In contrast, the pituitary responsiveness to repetitive pulses of GnRH in subfertile stallions was significantly reduced during the non-breeding and breeding seasons.25 When subfertile stallions were subjected to a single dose of GnRH in a subsequent study (Figure 147.3) in the non-breeding season, responsiveness of pituitary LH was observed to be similar to that in the fertile stallions.29 However, the capacity of the Leydig cells to produce testosterone was shown to be significantly reduced in the subfertile group (Figure 147.3). The results of the above studies suggest that the low testosterone response in subfertile stallions may be due to a dysfunctional hypothalamic–pituitary axis producing bio-inactive LH or a primary testicular disorder. Primary intratesticular paracrine–autocrine malfunction Studies point to a primary testicular problem (Figure 147.4) in which a disrupter such as toxins or drugs affects the Sertoli cells by way of the intratesticular paracrine–autocrine system. The dysfunctional Sertoli cells via their paracrine system then affect the germ cells and the Leydig cells. Hormonal feedback affects changes involving the pituitary and hypothalamus. Bio-inactive gonadotropins are synthesized and secreted or an imbalance of the gonadotropins ensues. The process subsequently becomes cyclic.
Hypothesis Hypothesis for the etiology of idiopathic subfertility/infertility in the stallion Effectors?
(toxins, drugs, stress, steroids, nutrition, genetics)
Testicular Paracrine/Autocrine Disorder Sertoli Cell Dysfunction Leydig Cell Dysfunction
Germ Cell Dysfunction
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?
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Figure 147.3 Temporal changes in mean testosterone (T) response to a single dose of exogenous GnRH (25 µg) in 1 ml of saline expressed in terms of percent baseline in fertile (n = 5)and subfertile (n = 5) stallions. The study was carried out in the non-breeding season. Arrows represent the time of administration of GnRH. Although the luteinizing hormone response was similar (data not shown), the testosterone response was significantly different. Fertile stallions had a higher response than subfertile stallions. (Modified from Roser and Hughes.29 )
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↓ Inhibin ↑ LH ↓ Estrogen ↓ T ↑ FSH
Degeneration Subfertility/Infertility
Figure 147.4 Hypothesis of events that may occur following a decline in intratesticular paracrine–autocrine factors owing to effectors such as toxins, drugs, stress, steroids, a nutritional imbalance, or genetic abnormalities. The first cell affected in this cascade of events is proposed to be the Sertoli cell, which leads to dysfunctional germ cells and Leydig cells causing testicular dysfunction and possible degeneration. These changes then have an effect on the endocrine system which ends up being out of balance.23 FSH, follicle-stimulating hormone; LH, luteinizing hormone; T, testosterone. With permission from Elsevier.
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Hours from injection (hCG) Figure 147.5 Temporal changes in mean plasma concentrations in testosterone after a challenge of human chorionic gonadotropin (hCG; 10 000 IU in 10 ml saline) given at time 0 in fertile (n = 7), subfertile (n = 4), and infertile stallions (n = 3). Note that the fertile and subfertile stallions have similar testosterone (T) responses whereas the T response in infertile stallions is significantly lower. (Modified from Roser.17 )
In one study, a group of infertile stallions had a significantly lower testosterone response to a challenge of human chorionic gonadotropin (hCG; 10 000 IU) than the fertile or subfertile group (Figure 147.5).17 In another study, no significant differences were observed in pituitary function of fertile, subfertile, and infertile stallions 1 year after castration with or without steroid therapy or after GnRH treatment.37 Taken together, these studies suggest that the original malfunction leading to a decline in fertility may be at the level of the testes and not at the level of the hypothalamus or pituitary. To further explore testicular function in idiopathic subfertile and infertile stallions, two studies were carried out whereby testicular tissue from fertile, subfertile, and infertile stallions was assessed for LH receptor binding activity and inhibin concentrations. The first study demonstrated that the number of LH receptors and receptor affinity on Leydig cells did not change with fertility status, suggesting that a testicular disorder is most likely present at the post-receptor level22 or is due to a malfunction of other somatic cells such as the Sertoli cell. The second study indicated that, in stallions with poor fertility, intratesticular inhibin concentrations declined early on, before changes in intratesticular steroids and changes in circulating hormone were observed (Figure 147.6), suggesting that an early paracrine–autocrine dysfunction may occur at the level of the Sertoli cell, which produces inhibin.18 If inhibin acts as a paracrine factor influencing Leydig cell steroidogenesis and germ cell function, as suggested in the literature in rat and hamster models,38–40 a decrease in testicular inhibin may directly affect steroid production and eventually spermatogenesis in the horse. Other
paracrine–autocrine factors have been identified in equine testes such as estrogen, testosterone, transferrin, insulin-like growth factor 1 (IGF-1), insulin-like peptide 3 (INSL3), and β-endorphin.15 Neither intratesticular IGF-I nor circulating IGF-1 appear to be biomarkers for declining fertility.41 However, a slow decline in local factors such as intratesticular estrogen, inhibin, transferrin, and testosterone, which support spermatogenesis, may contribute to the decline in fertility.15, 42, 43 More research is needed to investigate paracrine–autocrine factors that may change in idiopathic subfertile and infertile stallions. In humans and other species, it appears to be a disruption in the testicular paracrine mechanisms that may initiate unexplained fertility.44
Diagnostic investigation of endocrine–paracrine–autocrine parameters in idiopathic subfertile/infertile stallions An approach to identifying the exact nature of the decline in fertility in idiopathic subfertile stallions might be to perform a semen analysis, take blood samples for basal hormone profiles, and perform GnRH challenges tests and an hCG stimulation test. As indicated above, idiopathic subfertility/infertility may be due to changes in paracrine–autocrine factors. These factors can be identified in testicular tissue only by implementing a testicular biopsy procedure. Evidence demonstrates that a testicular biopsy procedure can be a relatively safe diagnostic tool to identify testicular factors and function in stallions (Figure 147.7).45–47 Several parameters of the specimens collected by the biopsy procedure can be evaluated. These diagnostic tests can be useful in providing veterinarians with information on the state and stage of fertility in a breeding stallion. However, because this is an invasive procedure, it should only be carried out when other hormone test results are normal and yet there appears to be changes in the size and consistency of the testes along with changes in the semen parameters. Below are diagnostic protocols to evaluate endocrine–paracrine and autocrine disorders in the colt or stallion.
Diagnostics for breeding soundness Steroids and proteins in seminal plasma As part of a breeding soundness examination, raw semen may be centrifuged and the supernatant (seminal fluid) secured for evaluation of such steroids and proteins as androgens and estrogens,48, 49 transferrin,50 and alkaline phosphatase.4
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Figure 147.6 Effects of fertility status on intratesticular inhibin concentrations and other intratesticular paracrine–autocrine factors. Testicular extracts: after castration at the UC Davis Veterinary Clinic, the testes were kept on ice for 10 minutes while being transported to the laboratory. The tunica was removed and the testes weighed. Both testes were sliced into small pieces with a razor blade and homogenized in a 0.01 M NaPO4 buffer [0.1 M sucrose, 0.1% bovine serum albumin (BSA), 0.01% bacitracin, 0.01% benzamidine–HCl, 0.01% lima bean trypsin inhibitor, and 0.001% aprotinin, pH 7.5; Sigma] at 4◦ C for 30 minutes in a Warring blender. Homogenate was then spun for 30 minutes at 300g in a Sorvall RC-5B centrifuge. The resulting supernatant was filtered through two layers of cheesecloth and the volume measured. Five millimoles of magnesium chloride was added and the supernatant was centrifuged at 27 000g for 30 minutes at 4◦ C in a Beckman LS-70M ultracentrifuge. The supernatant was frozen and stored at –70◦ C until assayed for inhibin, E2 , estrogen conjugates (EC), and testosterone (T) by validated radioimmunoassay. To measure protein concentrations in all testicular extracts a Bio-Rad Protein Kit was used. ∗ P ≤ 0.10 compared with fertile stallions. ∗∗ P ≤ 0.05 compared with fertile and subfertile stallions. Note that only intratesticular levels of inhibin were different in the subfertile and infertile stallions. All other intratesticular factors (T, E2 , and EC were similar among the groups of the stallions. Not shown: only infertile stallions exhibited significant changes in circulating plasma levels of hormones. Inhibin and estradiol and estrogen conjugates (EC) were significantly lower while follicle-stimulating hormone and luteinizing hormone concentrations were significantly higher. Plasma concentrations of T were not significantly different among fertile, subfertile, or infertile stallions. (Modified from Stewart and Roser.18 ) E2 , estradiol; F, fertile; S, subfertile; I, infertile.
Serum or plasma concentrations of FSH, LH, estradiol, inhibin, and testosterone Secretion of hormones can be episodic throughout the day, so it is important to obtain an average baseline.51 Six venous blood samples are recommended, once every hour on the same day between 10 a.m. and 3 p.m.51 Alternatively, if only a rough estimate of baseline values is needed, a simpler approach is to take three blood samples, one each day for 3 days taken between 9 and 10 a.m. Usually a 10-ml blood sample is adequate. Either serum or plasma samples can be used. Variations in values can be caused by age, season, laboratory protocols, and fertility status. In our laboratory, using RIA techniques, the normal ranges of circulating hormones in the non-breeding/breeding seasons are as follows: LH, 1–10 ng/ml; FSH, 2–12 ng/ml; inhibin, 1.5–4 ng/ml; estradiol E2 , 32–62 pg/ml; estrogen conjugates (EC), 100–300 ng/ml; testosterone (T), 0.5–3.0 ng/ml. In our laboratory, circulating
hormone values in an idiopathic subfertile/infertile stallion usually show changes in the following order: decreasing inhibin and estradiol, increase in FSH levels followed by an increase in LH, and, finally, a decrease in testosterone. These changes may take a few months or a few years depending on the age of the stallion and the testicular disorder.
Three-pulse GnRH challenge test (Figure 147.2) To assess pituitary responsiveness, a series of small challenges of GnRH are needed. A single challenge is not sufficient to assess pituitary responsiveness in stallions with primary testicular disorder. Three small doses of GnRH should be given intravenously (i.v.), 5 µg/dose, 1 hour apart in the non-breeding season starting at 9 a.m. Blood samples should be obtained every 10 minutes starting at 30 minutes before the first injection, 0–60 minutes after the first
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Figure 147.7 Biopsy procedure. (a) Preparing to remove testicular tissue from a mildly sedated stallion using a spring-loaded Biopty instrument (Bard Inc., Covington, GA) attached to a 14-gauge split needle. Three biopsy punches were removed. (b) Area of incision: Testis was gently moved back caudally in the scrotum and held prior to incision. (c) The location (craniolateral quarter of left testis) from which to take testicular tissue. (d) Tissue obtained is dispensed into cryostat vials for freezing at –70◦ C prior to measuring intratesticular factors by validated radioimmunoassays for insulin-like growth factor-1 [IGF-1], inhibin, estradiol [E2], and estrogen conjugates [E2]). (e) Repetitive biopsy samples taken from the testis of 6-month-old, 2-year-old, and 19-year-old stallions. (f) Low power (200×) of a histological preparation of a testicular Biopty punch stained with hematoxylin and eosin. Note the seminiferous tubules (ST). With permission from Elsevier.
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hCG challenge (Figure 147.5) To assess only testicular responsiveness, a single dose of hCG (10 000 IU) should be given i.v. at 9 a.m. This procedure can be done in either the breeding or nonbreeding season. Blood samples should be collected 30 minutes before, immediately before, and every 30 minutes after the injection up to 3 hours after treatment. Blood samples should be analyzed for testosterone. A study on a small population of fertile and subfertile/infertile stallions indicated that only the infertile stallions had a poor testosterone and estradiol response to hCG.17 Testicular biopsy (Figure 147.7a–g)
(g) Figure 147.7 (Continued) (g) High power (400×) of a histological preparation of a testicular Biopty punch stained with hematoxylin and eosin. Note Leydig cells (LC) that surround the seminiferous tubule (ST).23, 37
injection, 0–60 minutes after the second injection, and 0–60 minutes after the third injection. The entire process should take 210 minutes. Blood samples should be analyzed for LH and testosterone. Compared with fertile stallions, subfertile stallions have a significantly lower response to the second and third injection of GnRH in the non-breeding season only.25 Single-pulse GnRH challenge test (Figure 147.3) To assess pituitary and testicular responsiveness, a single dose of 25 µg of GnRH should be given i.v. at 9 a.m. in the non-breeding season. An abnormal response to a single challenge may help in the diagnosis of a pituitary disorder in stallions with tumors or hypogonadotropic hypogonadism (low GnRH, LH, and FSH) or in stallions with testicular dysfunction. Blood samples should be collected 30 minutes before the injection, immediately before the injection, and every 30 minutes after the injection up to 120 minutes. Blood samples should be analyzed for LH and testosterone by RIA. Compared with fertile stallions, subfertile stallions have a similar LH response but the testosterone response is lower in the non-breeding season.29
This procedure provides a direct measurement of endocrine–paracrine–autocrine factors in testicular tissue.45, 47 Three biopsy punches are obtained using a spring-loaded Biopty instrument (Bard Inc., Covington, GA) attached to a 14-gauge split needle under mild sedation, using a stanchion to secure the stallion. After the scrotum is aseptically prepared with an iodine scrub and blocked using a 2% lidocaine–HCl solution, a small incision is made in the scrotum in the center of the craniolateral quarter of the most affected testis. The Biopty instrument is placed through the incision against the tunica vaginalis, subsequently fired with the needle projecting into the testicular parenchyma, and then removed. Two subsequent samples are collected through the same incision site but at slightly different angles. Two punches are each placed in 1.2 mL of phosphate buffered saline and snap-frozen in dry ice and alcohol or liquid nitrogen and stored frozen until processed for measurement of testicular factors. The third sample is placed in Bouin’s solution for 6 hours, transferred to 50% alcohol, and submitted for histological examination. Testicular fine needle aspiration (Figure 147.8a,b) Testicular fine needle aspiration (FNA) has been extensively described in human and veterinary medicine.52–59 This procedure is less invasive than the testicular biopsy method. Although it cannot be used to determine changes in intratesticular paracrine–autocrine factors, the aspirate can be placed on a slide and testicular cells stained for identification. The specimen acquired is sufficient to differentiate inflammatory diseases from noninflammatory diseases and also may allow diagnoses of neoplasia. Spermatogenic activity can be assessed by comparing the number of germinal cells with the number of Sertoli cells.58, 60 Aspiration biopsy of the
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Stallion: Specific Techniques Used in Reproductive Examination of the Stallion Testicular Fine Needle Aspiration
(a)
Testicular Fine Needle Cytology
(b)
Figure 147.8 Testicular fine needle aspiration. (a) Aspirating parenchymal tissue from the craniolateral quarter of the testis from a mildly sedated stallion using a 23-gauge needle attached to a 5-ml syringe. The needle is withdrawn and the material obtained is ejected onto a glass slide, air dried, and stained with Giemsa stain. (b) Testicular cell types by fine needle aspiration. Included in this cytology specimen are ES, early spermatid; LS, late spermatid; S2, secondary spermatocyte; SC, Sertoli cell; SG, spermatogonia; SP, spermatozoa. (Modified from Roser.23 )
testis can be performed using a 22- or 23-gauge needle and a 5-ml syringe. The horse is mildly sedated, and the scrotal skin is aseptically prepared with iodine scrub. The testicle is firmly immobilized, the needle is introduced into the parenchyma of the testicle, and the syringe plunger is retracted as two or three passes are made into the testicle. After careful release of the syringe plunger, the needle is withdrawn, and material obtained is ejected onto glass slides, air dried, smeared, and stained with a Giemsa stain. The smear is examined for Leydig cells, germ cells, and Sertoli cells under 1000–2000× magnification using light microscopy.47
Hormone therapy Attempts to develop an effective empirical hormone method to treat stallions of all ages with fertility problems have so far been largely unsuccessful.1, 61 Hormone therapies that have been attempted are presented below. Therapy with hormones such as GnRH, equine chorionic gonadotropin (eCG), hCG, or FSH have been clinically implemented, but with limited success.1,61–63 Although studies have not shown definitively that nutritional supplementation beyond that which is required enhances reproductive efficiency in the stallion,11 a recent study indicates that feeding a dihydroepiandrosterone (DHA) enriched nutriceutical increased sperm motility parameters in cooled stored semen and frozen–thawed semen.64 Until the pathophysiology of the problem is clearly understood, most therapies may have a higher risk than benefit. Currently, careful management may be
the only answer to getting the most out of an idiopathic subfertile/infertile stallion.
Gonadotropin-releasing hormone The use of GnRH as a ‘cure-all’ for stallions with idiopathic subfertility has been highly over-rated. Controlled studies have demonstrated that treatment with GnRH does not dramatically improve seminal characteristics or pregnancy rates, regardless of the therapeutic regimens used.63, 65, 66 Long-term treatment with a potent GnRH analog given to normal pony stallions had a short-term stimulatory effect followed by a limited suppression of the secretion of LH, FSH, testosterone, and estrone sulfate as well as a significant but reversible reduction in spermatogenesis.67 Although the evidence is equivocal, there is a possibility of downregulating the testis with GnRH,61 and downregulation has been observed in stallions and geldings given Ovuplant (GnRH agonist implant).68 Hypogonadotropic hypogonadism (low levels of GnRH, LH, and FSH) has not been reported in stallions, so it is unlikely that providing exogenous GnRH will improve fertility. In fact, many stallions have normal circulating concentrations of LH levels and testosterone but high concentrations of FSH, suggesting a problem in the feedback mechanism at the level of the testis. Thus, the efficacy of GnRH therapy to increase sperm production or motility in stallions is questionable. However, a study carried out by Sieme et al.69 demonstrated that GnRH therapy during the non-breeding season improved sexual behavior and increased the quality of frozen–thawed
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Diagnostics and Therapeutics for Stallions with Declining Fertility sperm in fertile stallions. This effect was comparable between good and poor freezer groups. However, stallions in the poor freezer group did not improve the quality of frozen–thawed spermatozoa enough to reach the accepted threshold of 30% progressive motility after thawing. In the same study, there were no observable motility changes found in the raw semen before freezing (M. Troedsson, personal communication). Immunization against GnRH caused a decrease in libido in adult stallions,70, 71 supporting the concept that GnRH and/or the gonadotropins affect sexual behavior either directly at the level of the brain or indirectly via testosterone release from the testis. GnRH has also been used to induce testicular descent in cryptorchid boys if one or both testes are already in the inguinal canal.72 The success rate is variable, but the study by Esposito and co-workers72 demonstrated that therapy with GnRH to 85 cryptorchid boys resulted in 25 boys experiencing testicular descent into the scrotum (29.4% success rate). Treatments that have been suggested to help induce testicular descent in young colts (< 2 years old) include 500 µg GnRH daily for up to 3 weeks.73 However the success rate is not expected to be high.
Human chorionic gonadotropin There is a report that hCG has been used to induce testicular descent in cryptorchids if one or both testes are in the inguinal canal.74 Brendemuehl74 treated normal pre-pubertal colts and colts whose testis had not descend into the scrotum with 2500 IU hCG twice daily for 4 weeks. In four of the eight (50%) colts in which one testicle was identified in the inguinal canal prior to initiation of treatment, the testicle was present in the scrotum after the last treatment. No change in testicle location was noted in the two colts with abdominal cryptorchid testicles. Other treatments that have been suggested to help induce testicular descent in young colts (< 2 years old) include 2500 IU hCG twice a week for up to 6 weeks.73 Administration of hCG to induce testosterone release from the testis is effective,17, 74 but, because it is a foreign protein to the horse, it can stimulate an immune response.75 Therefore, hCG as a therapeutic tool in colts or stallions must be used with great caution.
Growth hormone There is considerable evidence in mice, rats, and humans that growth hormone (GH) affects testicular function by modulating gonadal steroid synthesis and gametogenesis.76–79 In humans, a deficiency in GH can result in male infertility.79 It appears that GH can increase plasma IGF-I concentrations and restore
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sperm motility79 as well as induce spermatogenesis in patients with hypogonadotropin hypogonadism.80 In the stallion, treatment with somatostatin (inhibitor of GH) twice daily for 10 days caused a transient decrease in semen motility and hCG-induced testosterone release, suggesting that GH may play a role in the regulation of testicular function in the stallion. 81 A study by Champion and co-workers82 demonstrated that sperm incubated with GH or IGF-I increased the longevity of the sperm in vitro.82 GH treatment in vivo may improve fertility in the stallion with disturbances in GH output and impaired testicular function, but further research is needed before this therapeutic strategy can be embraced.
Steroids It has been known for decades that treatment with anabolic (androgens) steroids causes a severe depression of reproductive capacity in stallions,83 most likely because of the negative feedback effects on LH.84 It took about 90 days for stallions to recover (normal sperm count) from treatment with anabolic steroids given daily for 88 days.85 The therapeutic effect of treating stallions with estrogens has not been investigated. In some stallions with idiopathic subfertility/infertility, estrogens are low.1, 25 Estrogens have been shown to have a local effect on the testis of other species.86 Estrogens have been shown to enhance the GnRH-induced LH release from equine pituitary cells in culture87 but decrease circulating levels of FSH when given to geldings.88 An estrogen treatment protocol would have to be determined on a case-by-case basis. If estrogen is low, as determined by plasma and/or intratesticular levels, an innovative approach may be to insert estradiol-17β capsules in the tunica vaginalis, but research is needed before such treatment can be recommended.
Recombinant equine luteinizing hormone (eLH) and equine follicle-stimulating hormone (eFSH) In 1992, Douglas and Umphenour1 suggested that equine gonadotropins may be beneficial in stallions with poor fertility, reporting that injection with a pituitary extract resulted in significant increases in LH. Treatments with equine pituitary extracts (EPEs) as well as partially purified eFSH (Bioniche) have successfully induced superovulation in mares, but not consistently.89–91 There are many hormone contaminants in preparations of pituitary extracts or partially purified eFSH, but the major contaminate is
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LH91 (J.F. Roser, unpublished data). The material injected into mares has a fixed ratio of eLH to eFSH that changes from batch to batch, decreasing the chances of manipulating a desired outcome. The preparation of eFSH was more consistent in increasing the number of follicle and ovulation rates compared with controls, and the embryo to ovulation rates were the same.92 Pituitary extracts and partially purified eFSH have not been tried in stallions with poor fertility. In humans, recombinant human FSH has increased sperm production depending on the clinical diagnoses of oligospermia or azoospermia.93, 94 Recently, however, through a collaborative effort with Dr. I. Boime (Washington University, St. Louis, MI), recombinant eLH (reLH) and recombinant eFSH (reFSH) have been developed. Treatment with reLH in normal stallions induces a testosterone response.95 Incubation of cultured Sertoli cells with increasing amounts of reFSH induces a response release of estradiol-17β. 96 In the mare, treatment with reFSH promotes follicular development,97 and it will likely increase circulating estrogen and inhibin concentrations in the stallion. Further research with the use of reLH and reFSH in colts and stallions is needed. Therapy with these two hormones may be beneficial in stallions with low gonadotropin levels or endogenous gonadotropins that may be bio-inactive, as has been observed in infertile stallions.28 Recently, a clinical study has demonstrated that superovulating cycling mares with reFSH increases the pregnancy per cycle rate when breeding these mares to subfertile stallions (W. Zent, unpublished data). This treatment increases the number of oocytes that are available to be fertilized and does not modify pregnancy rate per ovulation The opportunity to treat stallions or mares with the individual gonadotropins enhances their manipulative power. Using cloned material ensures a constant and consistent supply of reLH and reFSH to the clinician.
Summary The reproductive function of the stallion is not only controlled by classical hormones such as LH, FSH, testosterone, estrogen, and inhibin but appears to be modulated by other hormones such as GH. In addition, a paracrine–autocrine system appears to be important in local testicular function. There are several diagnostic tests that can be applied to aid in determining at what level of the reproductive axis there may be a primary dysfunction. Of course, there is no silver bullet, but, as new therapeutic tools reach the marketplace, the veterinarian will be better equipped to manage the idiopathic subfertile/infertile stallion on a case-by-case basis.
References 1. Douglas RH, Umphenour N. Endocrine abnormalities and hormonal therapy. Vet Clin North Am Equine Pract 1992;8:237–49. 2. Blanchard TL, Varner DD. Testicular degeneration. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 855–60. 3. Turner RMO. Testicular abnormalities. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis, MO: Saunders Elsevier, 2007; pp. 195–204. 4. Turner RM, McDonnell SM. Alkaline phosphatase in stallion semen; characterization and clinical applications. Theriogenology 2003;60:1–10. 5. De Vries PJ. Diseases of the testes, penis, and related structures. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 878–84. 6. Pickett BW. Reproductive evaluation of the stallion. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 755–68. 7. Pickett BW. Sexual behavior. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 809–20. 8. Amann RP. Effects of drugs or toxin on spermatogenesis. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 831–9. 9. Pickett BW. Factors affecting sperm production and output. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 689–704. 10. McKinnon AO, Voss JL. Diseases of the stallion’s reproductive tract. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 845–94. 11. Hintz HF. Feeding the stallion. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 840–2. 12. McDonnell SM. Normal and abnormal sexual behavior. Vet Clin North Am Equine Pract 1992;8:71–89. 13. Burns PJ, Douglas RH. Reproductive hormone concentrations in stallions with breeding problems: case studies. J Equine Vet Sci 1985;5:40–2. 14. Wallach SJ, Pickett BW, Nett TM. Sexual behavior and serum concentrations of reproductive hormones in impotent stallions. Theriogenology 1983;19:833–40. 15. Roser JF. Regulation of testicular function in the stallion: an intricate network of endocrine, paracrine and autocrine systems. Anim Reprod Sci 2008;107:179–96. 16. Schumacher J, Varner DD. Neoplasia of the stallion’s reproductive tract. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 871–7. 17. Roser JF. Endocrine profiles in fertile, subfertile and infertile stallions: Testicular response to human chorionic gondadotropin in infertile stallions. Biol Reprod Monogr 1995;1:661–9. 18. Stewart BL, Roser JF. Effects of age, season and fertility status on plasma and intratesticular immunoreactive (Ir) inhibin concentrations in stallions. Domest Anim Endocrinol 1998;15:129–39. 19. Watson ED, Clarke CJ, Else RW, Dixon PM. Testicular degeneration in 3 stallions. Equine Vet J 1994;26:507–10. 20. Blanchard T, Johnson L, Roser JF. Increased germ cell loss rates and poor semen quality in stallions with idiopathic testicular degeneration. J Equine Vet Sci 2000;20: 263–5.
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Diagnostics and Therapeutics for Stallions with Declining Fertility 21. Blanchard TL, Johnson L. Increased germ cell and reduced germ cell:Sertoli cell ratio in stallions with low sperm production. Theriogenology 1997;47:655–77. 22. Motton DD, Roser JF. HCG binding to the testicular LH receptors is similar in fertile, subfertile and infertile stallions. J Androl 1997;18:411–16. 23. Roser JF. Endocrine diagnostics and therapeutics for the stallion with declining fertility. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis, MO: Saunders Elsevier, 2007; pp. 244–51. 24. Sato K, Miyake M, Tunoda N, Nakagawa A, Iwamura T. Relationship between the semen characteristics and serum testosterone and estrogen levels in three-year-old colts. Jpn J Zootech Sci 1981;52:447. 25. Roser JF, Hughes JP. Seasonal effects on seminal quality, plasma hormone concentrations and GnRH-induced LH response in fertile and subfertile stallions. J Androl 1992;13:214–23. 26. Inoue J, Cerbito WA, Oguri N, Matsuzawa T, Sato K. Serum levels of testosterone and oestrogens in normal and infertile stallions. Int J Androl 1993;16:155–8. 27. Lambert A, Talbot JA, Anobile CJ, Robertson WR. Gonadotropin heterogeneity and biopotency: implications for assisted reproduction. Mol Hum Reprod 1998;4:619–29. 28. Whitcomb RW, Schneyer AL, Roser JF, Hughes JP. Circulating antagonist of luteinizing hormone in association with infertility in stallions. Endocrinology 1991;128:2497–502. 29. Roser JF, Hughes JP. Dose-response effects of GnRH on gonadotropins and testicular steroids in fertile and subfertile stallions. J Androl 1992;13:543–50. 30. Marshall JC, Harsoulis P, Anderson DC, McNeilly AS, Besser GM, Hall R. Isolated pituitary gonadotrophin deficiency: gonadotrophin secretion after synthetic luteinizing hormone and follicle stimulating hormone-releasing hormone. BMJ 1972;4:643–5. 31. Bell J, Spitz I, Perlman A, Segal S, Palti Z, Rabinowitz D. Heterogeneity of gonadotropin response to LHRH in hypogonadotropic hypogonadism. J Clin Endocrinol Metab 1973;36:791–4. 32. Mortimer CH, Besser GM, McNeilly AS, Marshall JC, Harsoulis P, Tunbridge WM, Gomez-Pan A, Hall R. Luteinizing hormone and follicle stimulating hormone-releasing hormone test in patients with hypothalamic–pituitary–gonadal dysfunction. BMJ 1973; 4:73–7. 33. Mecklenburg RS, Sherins RJ. Gonadotropin response to luteinizing hormone-releasing hormone in men with germinal aplasia. J Clin Endocrinol Metab 1974;38:1005–8. 34. Turner D, Turner EA, Schwarzstein L, Aparicio NJ. Response of luteinizing hormone and follicle-stimulating hormone to different doses of synthetic luteinizing hormonereleasing hormone by intramuscular administration in normal and oligospermic men: preliminary report. Fertil Steril 1975;26:337–9. 35. Bain J, Moskowitz JP, Clapp JJ. LH and FSH response to gonadotropin releasing hormone (GnRH) in normospermic, oligospermic and azoospermic men. Arch Androl 1978;1:147–52. 36. Dony JMJ, Smals AGH, Rolland R, Fauser BCJ, Thomas CMG. Differential effect of luteinizing hormone-releasing hormone infusion on testicular steroids in normal men and patients with idiopathic oligospermia. Fertil Steril 1984;42:274–80. 37. Roser JF, Tarleton M, Belanger JM. Pituitary response to steroid replacement therapy in fertile, subfertile and
38.
39.
40.
41.
42.
43. 44.
45.
46.
47.
48.
49.
50.
51.
52. 53.
54.
55.
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infertile stallions after castration. J Reprod Fertil Suppl 2000;56:61–8. Hsueh AJ, Dahl KD, Vaughan J, Tucker E, Rivier J, Bardin CW, Vale W. Heterodimers and monodimers of inhibin subunits have different paracrine actions in the modulation of luteinizing hormone-stimulated androgen biosynthesis. Proc Natl Acad Sci USA 1987;84:5082–6. van Dissel-Emiliani FM, Grootenhuis AJ, de Jong FH, de Rooij DG. Inhibin reduces spermatogonial numbers in testes of adult mice and Chinese hamsters. Endocrinology 1989;125:1899–903. Hakovirta H, Kaipia A, Soder O, Parvinen M. Effects of activin-A, inhibin-A and transforming growth factorbeta 1 on stage specific deoxyribonucleic acid synthesis during rat seminiferous epithelial cycle. Endocrinology 1993;133:1664–8. Hess MF, Roser JF. The effects of age, season and fertility status on plasma and intratesticular insulin-like growth factor I concentrations in stallions. Theriogenology 2001;56:723–33. Niederberger CS, Shubhada S, Kim SJ, Lamb DJ. Paracrine factors and the regulation of spematogenesis. World J Urol 1993;11:120–8. Sharpe RM. Paracrine control of the testis. Clin Endocrinol Metab 1986;15:185–207. Fabbri A, Ulisse S, Moretti C, Gnessi L, Bonifacio V, Di Luigi L, Ridolfi M, Spera G, Isidori A. Testicular paracrine mechanisms and unexplained infertility. In: Spera S, Gnessi L (eds) Unexplained Fertility: Basic and Clinical Aspects. New York: Raven Press, 1989; pp. 111–20. Faber NF, Roser JF. Testicular biopsy in stallions; diagnostic potential and effects on prospective fertility. J Reprod Fertil Suppl 2000;56:31–42. Hillman RB, Casey JP, Kennedy PC. Testicular biopsy in stallions. In: Proceedings of the 6th International Symposium on Equine Reproduction, 1994, p. 145. Roser JF, Faber NF. Testicular biopsy. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis, MO: Saunders Elsevier, 2007; pp. 205–11. Lemazurier E, Moslemi S, Sourdaine P, Desjardins I, Plainfosse B, Seralini GE. Free and conjugated estrogens and androgens in stallion semen. Gen Comp Endocrinol 2002;125:272–82. Macpherson ML, Simmen RC, Simmen FA, Hernandez J, Sheerin BR, Varner DD, Loomis P, Cadario ME, Miller CD, Brinsko SP, Rigby S, Blanchard TL. Insulin-like growth factor I and insulin-like growth factor binding protein-2 and -5 in equine seminal plasma: association with sperm characteristics and fertility. Biol Reprod 2002;67:648–54. Parlevliet JM, Bidstrup LA, Roser JF. Transferrin concentrations in seminal fluid in horses: a marker for fertility. Anim Reprod Sci 2008;107:337–8. Nett TM. Reproductive endocrine function testing in stallions. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 689–704. Mallidis C, Baker HWG. Fine needle tissue aspiration biopsy of the testis. Fertil Steril 1994;61:367–75. Dahlbom M, Makinen A, Suominen J. Testicular fine needle aspiration cytology as a diagnostic tool in dog fertility. J Small Anim Pract 1997;38:506–12. Papa FO, Leme DP. Testicular fine needle aspiration cytology from a stallion with testicular degeneration after external genitalia trauma. J Equine Vet Sci 2002;22:121–4. Papic Z, Katona G, Skrabalo Z. The cytology identification and quantification of testicular cell subtypes:
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reproducibility and relation to histological findings in the diagnosis of male infertility. Acta Cytol 1988;32:697–706. Foresta C, Varotto A. Assessment of testicular cytology by fine needle aspiration as a diagnostic parameter in the evaluation of the oligospermic subject. Fertil Steril 1992;58:1028–33. Gottschalk-Sabag S, Glick T, Bar-On E, Weiss DB. Testicular fine needle aspiration as a diagnostic method. Fertil Steril 1993;59:1129–31. Leme DP, Papa FO. Cytological identification and quantification of testicular cell types using fine needle aspiration in horses. Equine Vet J 2000;32:444–6. Fazouliotis SJ, Sagran A, Porat-Katz A, Simon A, Laufer N, Lewin A. A high predictive value of the first testicular fine needle aspiration in patients with non-obstructive azoospermia for sperm recovery at the subsequent attempt. Hum Reprod 2002;17:139–42. Finco DR. Biopsy of the testicle. Vet Clin North Am 1974;4:377–81. Brinsko SP. GnRH therapy for subfertile stallions. Vet Clin North Am Equine Pract 1996;12:149–60. Dowsett KF. Seminal abnormalities. In: Robinson NE (ed.) Current Therapy in Equine Medicine 2. Philadelphia: W.B. Saunders, 1987; pp. 564–6. Evans JW, Finley M. GnRH therapy in a stallion of low fertility. J Equine Vet Sci 1990;10:182–6. Brinsko SP, Varner DD, Love CC, Blanchard TL, Day BC, Wilson ME. Effect of feeding a DHA-enriched nutriceutical on the quality of fresh, cooled and frozen stallion semen. Theriogenology 2005;63:1519–27. Blue BJ, Pickett BW, Squires EL, McKinnon AO, Nett TM, Amamr RP, Shiner KA. Effect of pulsatile or continuous administration of GnRH on reproductive function of stallions. J Reprod Fertil Suppl 1991;44:145–54. Roser JF, Hughes JP. Use of GnRH in stallions with poor fertility: a review. In: Proceedings of the 40th Annual Convention of the American Association of Equine Practitioners, 1994, pp. 23–5. Boyle MS, Skidmore J, Zhang J, Cox JE. The effects of continuous treatment of stallions with high levels of a potent GnRH analogue. J Reprod Fertil Suppl 1991;44:169–82. Johnson CA, Thompson Jr DL, Carmill JA. Effects of deslorelin acetate implants in horses: single implants in stallions and steroid-treated geldings and multiple implants in mares. J Anim Sci 2003;81:1300–7. Sieme H, Troedsson MHT, Weinrich S, Klug E. Influence of exogenous GnRH on sexual behavior and frozen/thawed semen viability in stallions during the non-breeding season. Theriogenology 2004;61:159–71. Turkstra JA, Van Der Meer FJ, Knaap J, Rottier PJ, Teerds KJ, Colenbrander B, Meloen RH. Effects of GnRH immunization in sexually mature pony stallions. Anim Reprod Sci 2005;86:247–59. Malmgren L, Andresen O, Dalin AM. Effects of GnRH immunization on hormone levels, sexual behaviour semen quality and testicular morphology in mature stallions. Equine Vet J 2005;33: 75–83. Esposito C, De Lucia A, Palmieri A, Centonze A, Damiano R, Savanelli A, Valerio G, Settimi A. Comparison of five different hormonal protocols for children with cryptorchidism. Scand J Urol Nephrol 2003;37:246–9. Ptaszynska M. Compendium of Animal Reproduction, 7th edn. Boxmeer: Intervet International BV, 2002; pp. 96–7. Brendemuehl JP. A comparison of the effects of repeated human chorionic gonadotropin administration of
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serum testosterone in prepubertal Thoroughbred colts with descended and cryptorchid testicles. Anim Reprod Sci 2006;94:155–7. Roser JF, Kiefer BL, Evans JW, Neely DP, Pacheco DA. The development of antibodies to human chorionic gonadotropin following its repeated injection in the cycling mare. J Reprod Fertil Suppl 1979;27:173–9. Zachmann M. Interrelations between growth hormone and sex hormones-physiology and therapeutic consequences. Horm Res 1992;38(Suppl. 1):1–8. Chandrashekar V, Bartke A. The role of growth hormone in the control of gonadotropin secretion in adult male rats. Endocrinology 1998;139:1067–74. Hull KL, Harvey S. Growth hormone: a reproductive Endocrine–paracrine regulator. Rev Reprod 2000;5:175–82. Breier BH, Vickers MH, Gravance CG, Casey PJ. Therapy with growth hormone: major prospects for the treatment of male subfertility? Endocr J Suppl 1998;45(Suppl.): S53–60. Shoham Z, Conway GS, Ostergaard H, Lahlou N, Bouchard P, Jacobs HS. Cotreatment with growth hormone for induction of spermatogenesis in patients with hypogonadotropic hypogonadism. Fertil Steril 1992;57:1044–51. Aurich JE, Kranski S, Parvizi N. Somatostatin treatment affects testicular function in stallions. Theriogenology 2003;60:163–74. Champion ZJ, Vickers MH, Gravance CG, Breier BH Casey PJ. Growth hormone or insulin-like growth factor-I extend longevity of equine spermatozoa in vitro. Theriogenology 2002;57:1793–800. Berndston WE, Hoyer JH, Squires EL, Pickett BW. Influence of exogenous testosterone on sperm production, seminal quality and libido of stallions. J Reprod Fertil Suppl 1979;27:19–23. Squires EL, Todter GE, Berndtson WE, Pickett BW. Effect of anabolic steroids on reproductive function of young stallions. J Anim Sci 1982;54:576–82. Squires EL, Berndston WE, Hoyer JH, Pickett BW, Wallach SJ. Restoration of reproductive capacity of stallions after suppression with exogenous testosterone. J Anim Sci 1981;53:1351–95. O’Donnell L, Robertson KM, Jones ME, Simpson ER. Estrogen and spermatogenesis. Endocr Rev 2001;22:289–318. Muyan M, Roser JF, Dybdal N, Baldwin DM. Modulation of gonadotropin-releasing hormone-stimulated luteinizing hormone release in cultured male equine anterior pituitary cells by gonadal steroids. Biol Reprod 1993;49:340–5. Thompson Jr DL, Pickett BW, Squires EL, Nett TM. Effect of testosterone and estradiol-17β alone and in combination on LH and FSH concentrations in blood serum and pituitary of geldings and in serum after administration of GnRH. Biol Reprod 1979;21:1231–7. Rosas CA, Algeria RH, Barcia JL, Agiler A, Chaves MG. Evaluation of two treatments in super ovulation of mares. Theriogenology 1998;49:1257–64. Scoggin CF, Meira C, McCue PM, Carnevale EM, Nett TM, Squires EL. Strategies to improve the ovarian response to equine pituitary extract in cyclic mares. Theriogenology 2002;58:151–64. Squires EL, McCue PM. Superovulation in mares. Anim Reprod Sci 2007;99:1–8. Niswender KD, Alvarenga MA, McCue PM, Hardy OP, Squires EL. Superovulation in cycling mares using equine follicle stimulating hormone (eFSH). J Equine Vet Sci 2003;23:497–500.
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Diagnostics and Therapeutics for Stallions with Declining Fertility 93. Foresta C, Bettella A, Merico M, Garolla A, Ferlin A, Rossato M. Use of recombinant human follicle stimulating hormone in the treatment of male factor infertility. Fertil Steril 2002;77:238–44. 94. Foresta C, Bettella A, Garola A, Ambrosini G, Ferlin A. Treatment of male idiopathic infertility with recombinant human follicle-stimulating hormone: a prospective, controlled, randomized clinical study. Fertil Steril 2005;84:654–60. 95. Jablonka-Shariff A, Roser JF, Bousfield GR, Wolfe MW, Sibley LE, Colgin M, Boie I. Expression and bioactivity of a single chain recombinant equine luteinizing hormone (reLH). Theriogenology 2007;67:311–20.
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96. Roser JF, Sibley LE, Jablonka-Shariff A, Daphna-Iken D, Boime I. Expression and bioactivity of single chain recombinant equine luteinizing hormone (eLH) and equine follicle stimulating hormone (eFSH). In: Proceedings of the 15th Annual Meeting of ICAR, 2004, p. 572. 97. Jennings MW, Boime I, Daphna-Iken D, Jablonka-Shariff A, Conley AJ, Colgin M, Bidstrp LA, Meyers-Brown GA, Famula TR, Roser JF. The efficacy of recombinant equine follicle stimulating hormone (reFSH) to promote follicular growth in mares using a follicular suppression model. Anim Reprod Sci 2009;116:291–307.
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Endoscopy of the Internal Reproductive Tract Carla L. Carleton
Overview With the advent of ultrasonography, detailed examination of soft-tissue density is possible and thus the frequency of use and necessity of endoscopy has diminished. However, endoscopy remains the best means of direct visualization of the lumen and walls of the urethra, internal surfaces and/or ducts of accessory glands, as well as the neck and lumen of the bladder. In addition, endoscopy aids in topical treatment of the internal reproductive tract, e.g., seminal vesicles. Abnormalities can be identified by transrectal palpation and ultrasonography, or may be suspected subsequent to pain, or observing an abnormal discharge. Site-specific, directed local therapy, e.g., of seminal vesicles or a urethral tear, can be accomplished by introduction of treatment via a channel of a flexible endoscope. Serial examinations using endoscopy assist in monitoring the progress of treatment of chronic conditions. Endoscopy is often a useful diagnostic tool to define the source of blood in cases of hemospermia or hematuria, of pain associated with ejaculation or urination, of cystitis (blood, cystic calculi, tumors, inflammatory cells), of urospermia or pyospermia, or as a follow-up if pain is elicited during transrectal palpation of the stallion’s accessory sex glands and other pelvic structures. Many of the conditions listed will present with conflicting clinical signs, including hematuria, urinary incontinence, or stranguria. Other non-specific signs of discomfort that may be related to the genitourinary tract include penile prolapse, weight loss, and recurrent colic. With the arrival of newer imaging modalities (ultrasonography, computed tomography (CT), magnetic resonance imaging (MRI)), endoscopy is used less often than a decade ago. However, if a breeding stallion’s history suggests abnormal cellular mate-
rial in the ejaculate (blood, inflammatory cells, tumor cells), endoscopy is a very effective and useful tool to visualize the luminal epithelium along its length, including the pelvic portion of the stallion’s urethra, the openings of the accessory sex glands that contribute to the stallion’s seminal fluid, and the bladder itself, including the trigone, ureters, neck, internal surface, and bladder contents. Additionally, by using specialized endoscopic instrumentation (sterilized catheters, suction, biopsy instruments), site-specific sampling is possible.
Features of flexible endoscopic or videoendoscopic equipment1
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Fiberoptic endoscopes are less expensive and are the original equipment used for equine endoscopy. Long, thin glass fiber bundles transmit light and images. Optic fibers run the length of the endoscope and can be damaged by dropping, bending, and aging. Fiber breakage compromises image quality, with broken fibers appearing as black dots in the image. The best resolution/image quality is via direct viewing through the eyepiece, as image quality is lost with the use of monitors and recorders. This technology is being replaced by videoendoscopy in referral and primary care facilities. Videoendoscopy yields images of superior quality. With recent advances in digital image capture, costs have dramatically decreased. Light is more efficiently transferred via wires in the flexible endoscope, signal conversion is accomplished in a processor, and the images are viewed on a video monitor. Image resolution far exceeds that of fiberoptic endoscopes, since digital images do not degrade between capture, viewing, and storage.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Digital capture also provides greater flexibility in managing and manipulating stored images. Portable digital units are available. Images recorded during an examination can be transferred to a monitor or stored on DVD, flash drives, or videotape. Video cameras can also be directly attached to the eyepiece. For most efficient use of equipment, if acquiring a flexible endoscope, purchase one capable of being used at more than one site, e.g., for stallion genital examinations as well as evaluations of respiratory or gastrointestinal (GI) systems and hysteroscopy. The maximum outer diameter for endoscopy is 9–11 mm with a 1-meter minimum length for use in light breed stallion genital examinations. A longer endoscope might be needed (up to 3 meters) for GI use. The longer endoscopes would be necessary for genital examinations of some Warmblood and draft breed stallions. To examine foals, miniature horses, and ponies, a smaller diameter is needed, e.g., a pediatric scope of no more than an 8 mm external diameter.
Specific equipment features
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A tight-bending radius to permit a wide viewing range within a small distance (maneuverability). Four-way distal tip deflection is essential to guide it around curves, into ejaculatory ducts, etc. An air valve. Allows air in or can be closed off to dilate the urethra; provides an unobstructed view of surfaces as the equipment is advanced. A flush (irrigation) channel to clear the distal tip of debris, irrigate the lumen, or conduct an instillation (administer treatment). At least one channel through which accessory equipment may be introduced (catheters, snares, brushes, biopsy instruments). Suction for removal of content for analysis and to eliminate air if distension has created discomfort during or upon completion of an examination. Attached rubber caps for the water/flush and air channels. If they have been lost, extension sets or three-way injection port stopcocks can be substituted; otherwise, luminal distension cannot be maintained during an examination. For flexible endoscopes, most of the function controls will be located in the handpiece. Equipment should include a focus (diopter) ring to adjust the eyepiece at the outset of any examination. Image quality can be poor because of failing to focus or it can be due to debris, mucus, or fluid across the lens. Reviewing which controls create which deflection prior to each insertion/examination can save time in orienting oneself to the images acquired.
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Note that some resistance will be encountered during an endoscopic examination of a stallion’s urethra because of the sharp reflection of the urethra at the level of the ischial arch. Proper deflection of the tip of the endoscope will facilitate passage at this point.
Equipment storage Custom storage cradles secure the proximal (eyepiece) end of the endoscope between uses. The cradle’s height must be sufficient to permit the endoscope to hang straight without any bending, eliminating kinks and coiling associated with storage in the original packing box or carrying case. Sufficient height also ensures that the endoscope and its channels can completely drain and dry while stored between uses. The storage cradle is best fixed either onto a tall stand or inside a protected closet to avoid inadvertent damage and fiber breakage. Fexible endoscopes and videoendoscopes should never be stored in their carrying cases.
Equipment cleaning and disinfection between uses
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The majority of equine endoscopy in clinical practice is of the respiratory tract. A myriad of pathogens and contaminants can be introduced into and carried by the endoscope during an equine respiratory tract evaluation or with frequent use in a multi-doctor busy veterinary hospital, and there is a likelihood of contamination with β-hemolytic streptococcal organisms, methicillin (meticillin)-resistant Staphylococcus aureus (MRSA), Pseudomonas aeruginosa, and nosocomial hospitalassociated infections increases. It is critical to avoid contaminating the genital tract of the stallion or mare with recently used equipment that has not been properly disinfected. Manufacturers’ recommendations for cleaning and disinfection should be followed as the first line for care. Cleaning brushes (appropriate size for the endoscope, reusable or disposable) facilitate maintenance of the endoscope’s channels. In the absence of specific instructions to the contrary, never assume that the eyepiece (the entire proximal end) of the endoscope can be submerged in any cleaning solution. Regardless of which procedure is used to clean and disinfect a flexible endoscope, both its external surface and all internal channels must be included in the cleaning protocol. Cleaning and flushing the channels is required after each procedure: first with water, then with an enzymatic cleaning solution, then water or sterile irrigation
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saline, and finally with air to dry the channels. More recently, the importance of drying the endoscope not only between uses but also between serial steps of cleaning and disinfection has been reported.2 Although the overall risk is low, wet or inadequately dried endoscopes pose a greater risk, and serve as a source of waterborne or nosocomial infection, than properly dried endoscopes (cleaned, disinfected, dried). The prior most common method of cold sterilization of the flexible portion of the endoscope was with R , Johnson & Johnson) a 2% gluteraldehyde (Cidex R solution. Once Cidex solution was charged (activator added), the solution was stable for 2 weeks. Maximum soaking time was not to exceed 30–45 minutes in preparation for a sterile procedure. If still used, it is essential to follow the manufacturer’s guidelines, as many disinfectants will erode/destroy the seals and other connections of the endoscope, resulting in leaks. Gluteraldehyde (GA) is a very irritating solution. To prevent hazards from GA exposure, closedsystem, fully automated washing machines are recommended, making it impractical for all but large hospitals.3, 4 It is critical that all GA meticulously be rinsed from the instrument before its introduction into any body cavity. Following submersion for cold sterilization, the endoscope is lifted from the solution tray with the flexible portion cradled by an assistant wearing sterile gloves. It is held in a manner that the distal tip is lower than the proximal end. Sterile water or saline can be used (3–4 liters minimum) to flush through all channels, as well as removing all GA from its external surface and, most importantly, the recesses of the endoscope’s distal tip.
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Figure 148.1 Dissected stallion genital tract with landmarks noted. (a) Free portion of penis and inner lamina of prepuce; (b) body of penis contained in preputial cavity; (c) right bulbourethral gland; (d) right prostate gland lobe; (e) right vesicular gland; (f) right ampulla; (g) bladder; (h) right ductus deferens; (i) left bulbourethral gland, (j) left ductus deferens.
bourethral glands and urethral glands, the prostatic ducts, and the colliculus seminalis and the slit-like openings on its papillae through which the contributions from the ampullae and seminal vesicles enter the urethra. Further cranial to the colliculus seminalis lie the muscular neck of the bladder (the internal urethral orifice) and the bladder proper. The penile urethra extends from the distal, free end of the glans penis to the ischial arch. The urethral process extends outward 1.5–2 cm within the glans penis (Figure 148.2c). While the urethra is by and large protected within the corpus spongiosum penis, traumatic injuries often include the glans penis and the urethral process. With the advent of ivermectin
Alternative methods of sterilization include ethylene oxide gas (EtOH) and steam cleaning. The time required to prepare the instrumentation for use with EtOH is greater (48–72 hours) than with wet sterilization. In an automated system, the EtOH exposure time is usually 2.5 hours, followed by an additional 8-hour aeration period. Steam cleaning is commonly practiced in larger referral hospitals and can be accomplished with a rapid turn-around time, often less than 1 hour.
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Anatomy review To understand the importance of abnormal cellular material observed in an ejaculate or gross anatomical abnormalities, knowledge of the stallion’s urogenital anatomy, as viewed internally, is essential (Figure 148.1). The structures characterized during an endoscopic examination include the urethra (penile and pelvic portions), the ducts entering from the bul-
Figure 148.2 Glans penis, demonstrating corona glandis (a), dorsal urethral sinus (b), and urethral process (c).
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Figure 148.3 Habronemiasis affecting the shaft of a stallion’s penis. This resolved following ivermectin treatment.
anthelmintics, Habronema lesions are now infrequent (Figure 148.3). Additional proliferative lesions to differentiate from habronemiasis include neoplasia and viral papillomatosis, and, less commonly, sarcoids (Figures 148.4 and Figures 148.5). The most common cancer of the stallion’s penis is squamous cell carcinoma. A biopsy and histopathology are necessary to provide a definitive diagnosis. To ensure a diagnostic sample, the biopsy sample should include a portion of the obvious lesion, to and through its margin, into normal tissue. The pelvic portion of the urethra is 10–20 cm long and extends from the neck of the bladder to the is-
Figure 148.5 Aged stallion, friable penis (same as in Figure 148.4), with hemorrhage post-collection.
chial arch. There is no line of demarcation or difference in size or structure between the pelvic urethra at its cranial portion and the neck of the bladder. At the point where the root of the penis attaches to the ischial arch, the urethra is relatively tight and introduction of the endoscope often meets with resistance. Once the urethral lumen at the level of the ischial arch has successfully been negotiated, the urethra widens within the pelvic area. It is within the pelvic portion of the urethra that the openings of the accessory sex glands are present. Bulbourethral glands The glands are situated on either side of the pelvic portion of the urethra at the level of the ischial arch (Figure 148.6). Each bulbourethral (B/U) gland has six to eight excretory ducts that open as small papillae caudal to the prostatic ducts, but close to the dorsal midline. The urethral gland openings are in the same vicinity, lying lateral to those of the B/U glands, both approximately 3–4 cm caudal to the colliculus seminalis. Both are small and cannot readily be canalized. Fortunately, the B/U and urethral glands are rarely affected by infectious or inflammatory conditions. Prostate and prostatic ducts
Figure 148.4 Aged stallion, multiple lesions on glans. Biopsies from multiple sites confirmed presence of squamous cell carcinoma, viral papillomatosis, sarcoid, and habronemiasis.
The prostate consists of a lobe lying to the left and right of the urethra with a connecting, dorsal isthmus.
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Figure 148.6 Immediately cranial to ischial arch, bulbourethral gland ducts (arrow).
The prostatic isthmus is not distinctly palpable transrectally, as dorsally it is covered by urethral muscle. Prostatic secretions enter the ejaculate through multiple (15–20) ducts to the left and right of the colliculus seminalis. Colliculus seminalis The central, dominant structure visible within the stallion’s pelvic region is the colliculus seminalis (Figure 148.7). Its rounded prominence presents as a dorsal papilla, approximately 5 cm caudal to the urinary bladder sphincter. Slightly tilted, horizontal ‘slits’ are visible on the left and right side of the prominence (Figure 148.8). Fractions of the ejaculate from the ductus deferentia and seminal vesicles enter at these
Figure 148.7 Bulbourethral ducts in the foreground; prominence of colliculus seminalis on the midline (arrow).
Figure 148.8 Colliculus seminalis, the two ejaculatory duct ‘slits’ are evident (arrow).
sites. Contributions from the left ampulla (the caudal, dilated portion of the ductus deferens) and the left seminal vesicle enter the urethra through the left ‘slit’ and those of the right ductus and seminal vesicle through the right ‘slit’. While there can be some variation, it is most common to find that the ducts from the paired (left or right) seminal vesicle and ampulla join immediately cranial to the common slit of the colliculus seminalis on the papilla. Rarely, separate ducts will be observed (i.e., one side single, opposite side separated, or all entering separately on the papilla) (Figure 148.9). Less commonly, the orifice of the uterus masculinus may be observed between, and slightly cranial to, the ejaculatory ducts, the ‘slits’, of the colliculus seminalis. This ‘potential’ opening is the posterior extrem¨ ity of a fetal remnant of the Mullerian ducts. It may present as a ‘small dot’; as a short, blind tube, possibly connected with one of the seminal vesicle ducts and thus be inapparent; or be absent. The pelvic urethra cranial to the prostate exhibits a dilation of its width to potentially 5–6 cm in the vicinity of the colliculus seminalis. As its name implies, it is the collection area for the seminal fractions as emission progresses prior to ejaculation. With serial, rapid contractions of the urethralis muscle during ejaculation, the seminal fluid is jetted out in a rhythmic fashion, usually described as pulses. Three ejaculatory fractions are generally described for the stallion. The first is from the B/U glands, the second from the ampullae and the prostate, and the third from the seminal vesicles.
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a mare in estrus for sexual stimulation prior to performing the examination.
Equipment and Procedure While an examination can be accomplished with a minimum of three persons, there is a distinct advantage to having four persons present to perform an endoscopic evaluation of a stallion.
r r r r Figure 148.9 Variation in colliculus seminalis appearance, more flattened, one ductus deferens (arrow) is separate from the seminal vesicle duct.
Patient preparation Endoscopy of the stallion’s genital tract is most often done with the horse standing and sedated. Restraint in stocks is essential to limit side-to-side movement. Sedation also aids in achieving penile relaxation and eases the stallion’s discomfort during passage of the endoscope into and through the urethra into the pelvic region. While not required, the stallion’s discomfort/resistance to passage of the endoscope can be lessened by using a lubricant containing lidocaine.5 Safe options for sedation/analgesia of stallions are xylazine (0.5 mg/kg intravenously (i.v.) or intramuscularly (i.m.)) or detomidine (0.02–0.04 mg/kg i.v. or i.m.; i.e., 20–40 µg/kg).6 Butorphanol (0.02–0.03 mg/kg) may also be used in combination with detomidine (0.01–0.02 mg/kg i.v. or i.m.) or xylazine (0.5–1 mg/kg). Keep in mind that spontaneous ejaculation may result with sedation with detomidine, or xylazine/butorphanol. Although the incidence of risk is low, the use of phenothiazine tranquilizers, e.g., acepromazine, is contraindicated because of their association with penile paralysis. Once sedated and in the stocks, the stallion’s tail can be wrapped and tied to one side. This keeps the tail from making contact with the operators and the endoscope during the procedures. If not already done, transrectal palpation and massage of the ampullae and seminal vesicles may serve to increase secretory activity and improve visualization of structures by both ultrasound and endoscopy. Alternatively, the stallion can be placed in the presence of
Participant A sedates the stallion and remains at his head, providing restraint as needed. Participant B passes the endoscope into the urethra. Participant C monitors and captures images, and directs the flexible tip as the endoscope is advanced into the urethra and beyond. Participant D becomes essential if additional procedures are necessary, such as for passing catheters or biopsy instruments, collecting and processing of samples by suction, etc. Transrectal massage of accessory sex glands during the procedure may result in increased secretions visible during endoscopy and, if they are abnormal, serve to highlight areas of concern or aid in sample retrieval.
Participant B wears sterile gloves and is responsible for holding the flexible portion of the sterilized endoscope, gently grasping the stallion’s penis, and introducing and passing the endoscope into the urethra. Participant B should be positioned alongside the horse, facing towards the rear leg to avoid the possible cow-kick. A firm grasp of the penis with one hand makes it easier for participant C to slightly distend the urethra as the endoscope is advanced. This modest introduction of air enhances the images captured on the way in before any irritation caused by the endoscope itself becomes apparent. Evidence of overdistension of the urethra and/or bladder may become apparent by discomfort or agitation of the stallion, e.g., by his posturing and attempting to urinate in the midst of the procedure. The endoscope is advanced only upon the direction of participant C, who manipulates the hand-controls, directs the tip of the endoscope, identifies anatomical landmarks, notes any abnormalities, and captures images.
Clinical findings External genitalia It is essential to begin with a general examination of both external and internal portions of the genital tract. The origin of hemospermia is often external, from the prepuce, penile shaft (including
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Figure 148.10 Hemospermia due to laceration of the penile shaft.
damage caused by poor management of stallion rings), glans penis, and/or urethral extension (Figures 148.10–148.12). Any abnormalities or diversions from normal anatomy must be noted, e.g., erosions, proliferative lesions, anatomical defects. Internal genitalia The mucosa of the penile portion of the urethra is pale pink and upon first entry will have none to few slightly darker pink linear striations (Figure 148.13). It is important to visualize the mucosa as the endoscope is advanced, as mere contact with the endoscope, as well as distension by air, will cause rapid changes in its appearance (Figures 148.14 and 148.15). Vessels in the surrounding corpus spongiosum ure-
Figure 148.11 Hemospermia due to equine herpesvirus 3 infection, acute exacerbation; bleeding ulcers.
Figure 148.12 Equine herpesvirus 3 infection; scars are evidence of prior vesicles and ulceration.
thra will result in hyperemia in a short time. The most notable aspect of the penile urethra is its uniform, unremarkable appearance. Therefore, any changes, such as strictures and ulcerations, inflammation (uniform or pervasive redness may indicate urethritis), and fissures in the mucosa, generally capture one’s attention. Urethral rents (single or multiple) are linear tears of the mucosa, and are most commonly observed in the vicinity of the ischial arch.7 The mucosal
Figure 148.13 Normal urethra.
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Figure 148.14 Urethra with early changes caused by the endoscope.
defects may first be suspected because of hemospermia and identified earlier with an artificial vagina collection than following live cover simply because the ejaculate is examined more completely. Resolution of these tears can be challenging. Treatment recommendations have ranged from sexual rest (infrequent success), to subischial urethrostomy, to laser surgery followed by extended supportive care. Within the pelvic portion of the urethra are situated the multiple ducts of the accessory sex glands. The B/U glands and urethral glands and their ducts are infrequently a concern, but noting their location provides a landmark as the endoscope is advanced cranially. The colliculus seminalis lies dorsally. With passage of the endoscope over the ischial arch and inversion of the image, the colliculus seminalis can appear to lie ventrally. As described earlier, its ejaculatory ducts appear as slits to the left and right of the midline, some appearing more elevated than others (Figures 148.8 and Figures 148.9). If there is a seminal vesiculitis, discharge observed at the ejaculatory duct
Figure 148.15 Urethra approximately 16 minutes after the beginning of the examination.
Figure 148.16 Hemorrhage/clot at the colliculus seminalis, seminal vesiculitis.
can range from none to bloody to mucopurulent (Figure 148.16). Flexible catheters can readily be passed through the biopsy channel and into the ejaculatory duct (Figure 148.17). If the endoscope is to be advanced into a seminal vesicle, its passage is made easier if, first, the catheter is placed into an ejaculatory duct and then the endoscope advanced over it. It is more likely that the catheters or biopsy instrument
Figure 148.17 Introduction of the biopsy instrument into an ejaculatory duct.
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Figure 148.18 Dissected pelvic portion of the genital tract; dorsal view. (a) prostate gland; (b) right vesicular gland; (c) right ampulla; (d) bladder; (e) right ductus deferens.
entering the common duct will enter the seminal vesicle than the ampulla on either side (Figures 148.18 and 148.19), but, with serial sampling, tissue from both seminal vesicle and the adjoining ampulla may be collected for cytology or histopathological analysis. If seminal vesiculitis has been confirmed, the same technique may be used to access the gland for flushing and instillation of treatment. The multiple, small prostatic ducts lie lateral to each ejaculatory duct. Prostatitis is infrequent in the stallion. Transrectal ultrasonography provides a better means of evaluating the normalcy of the prostate. With sexual stimulation, secretions will accumulate within the prostate, making evaluation of its function and sampling of its content easier. The central diameter of the pelvic urethra is dilated immediately cranial to the prostate, in the vicinity of the colliculus seminalis. Between the ischial arch and colliculus seminalis multiple sets of ducts are visible. The urethra’s most cranial portion terminates at the internal urethral orifice (urinary bladder sphincter). There is no anatomical line of demarcation
Figure 148.19 Pelvic portion of a stallion’s genital tract, incised ventrally. Polyethene tubing (a) inserted into one ejaculatory duct (b) can be identified passing directly through to the corresponding seminal vesicle (c). An ampulla can also be idendified (d).
between the pelvic urethra at its cranial portion and the neck of the bladder. The sphincter of the normal urinary bladder will resist passage of the endoscope. Insufflation and slight pressure are usually necessary to gain access to the bladder. If the bladder is full, removal of urine by aspiration or releasing pressure on the distal penis will allow the stallion to evacuate his bladder. A complete examination of the bladder requires removal of most of the urine. A cytological examination of the bladder contents for the presence of blood, inflammatory cells, spermatozoa (an indication of retrograde ejaculation), abnormal cells, etc., is essential for a complete evaluation. Cystic calculi may develop subsequent to cystitis, and may be associated with a nutritional imbalance or nephrolithiasis and/or some other renal dysfunction. Calculi may be identified by endoscopy or ultrasonography. If small pieces of cystic calculi can be retrieved, they should be submitted for analysis to determine their composition. Options for their subsequent removal include manual (endoscopic forceps) or surgical removal or lithotripsy of larger cystic calculi to decrease their size and facilitate their passage through the urethra. Increasing the horse’s water intake by adding salt to its diet may delay the formation of subsequent calculi. The ureters connect each renal pelvis to the bladder, each entering the bladder dorsally near its neck. The two ureteral orifices are little more than 2.5 cm apart. The internal urethral orifice is at the apex of the trigone of the bladder, approximately 4 cm caudal to the ureteral orifices. Ectopic ureters may enter the tubular genital tract caudal to the internal urethral orifice and result in urine dribbling. Local muscle hypertrophy of the ureter at the bladder may result in stenosis, with subsequent cystic dilation, hydroureter, and hydronephrosis. In cases of hydroureter, it may be possible to advance the endoscope into the ureter. If hydroureter is noted, regardless of symptoms displayed by the patient, a complete ultrasound examination of both kidneys is essential and a urinalysis highly recommended. Biopsy samples of proliferative lesions and from the margins of eroded areas can be taken through the biopsy channel. The color of normal bladder mucosa is usually a pale pink. In the presence of cystitis, the wall can be reddened, often extensively, throughout the entire bladder. As accessory sex gland function is testosterone dependent, if faced with a chronic urethral tear, seminal vesiculitis, or prostatitis, and treatment has failed, a last resort to salvage a horse and return it to health is castration. Following gelding, the accessory sex glands regress and return to a juvenile, quiescent state.
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Endoscopy of the Internal Reproductive Tract
References 1. Slovis NM. Endoscopic instrumentation. In: Slovis NM (ed.) Atlas of Equine Endoscopy. St. Louis, MO: Elsevier, Mosby, 2003. 2. Muscarella LF. Inconsistencies in endoscope-reprocessing and infection-control guidelines: the importance of endoscope drying. Am J Gastroenterol 2006;101:2147– 54. 3. Takigawa T, Endo Y. Effects of glutaraldehyde exposure on human health. J Occup Health 2006;48:75– 87.
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4. Omidbakhsh N. A new peroxide-based flexible endoscopecompatible high-level disinfectant. Am J Infect Control 2006;34:571–7. 5. Tibary A. Endoscopy of the reproductive tract in the stallion. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis, MO: Elsevier, 2007; pp. 214–19. 6. Plumb DC. Plumb’s Veterinary Drug Handbook, 6th edn. Oxford: Blackwell Publishing, 2008. 7. Schumacher J, Varner DD, Schmitz DG, Blanchard TL. Urethral defects in geldings with hematuria and stallions with hemospermia. Vet Surg 1995;24:250–4.
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Chapter 149
Examination of External Genitalia Patricia L. Sertich
Introduction Thorough clinical examination of the external genitalia can provide information about the breeding potential of a stallion and permit detection of possible problems. Comprehensive reviews of the anatomy and physiology of the stallion’s reproductive system should be consulted for detailed descriptions and diagrams of the genital tract (see Chapter 95) in this edition.1–3
Restraint There are different opinions regarding the restraint of stallions, but, for routine examination of the genital tract, a calm, confident, firm and steady approach is often quite effective. The stallion should be fitted with a well-adjusted stout halter with large cheek rings that permit a chain to slide easily. For examination of the scrotum, a shank may be simply attached on the bottom halter ring; however, with a stallion that is not known to the handler it is advised to place a chain through the stallion’s mouth. The stallion handler and the examiner should both stand on the same side of the horse, and normally the examination is performed from the near (left) side. The stallion may be most quiet in its own stall or grooming area. The examiner should stand in contact with the stallion’s left shoulder while leaning over to reach the inguinal area. The examiner should take care to keep his/her head up with body weight centered over the legs. It may be helpful to back a fidgety stallion into a padded corner. Initial examination should begin with the examiner grasping the mane at the level of the withers. For examination of the penis, the stallion may need to be teased to obtain an erection. This may re-
quire more restraint that can be achieved by having the handler place the shank’s chain either over the stallion’s nose or in the stallion’s mouth. The chain should be used as needed to get the stallion’s attention and care should be taken not to mindlessly jerk on the chain, rendering it less useful in controlling the stallion should the need for greater restraint arise. Should the stallion require sedation to safely perform the examination, one should not use a phenothiazine tranquilizer such as acepromazine as stallions may develop priapism or penile paralysis as a result of its use.4 Regardless of the type of drug administered, after completion of the examination care should be taken to confirm that the stallion has retracted the penis into the sheath and can safely swallow before returning it to the stall or transportation vehicle. Sexual excitement while under the influence of any tranquilizer should be avoided.
Scrotum The scrotum and its contents should be manually examined by direct palpation. The scrotal skin should be smooth, thin, and free of any abrasions. The layers constituting the wall of the scrotum include the skin, tunica dartos muscle, scrotal fascia, and the parietal vaginal tunic. The scrotum should be of uniform thickness, freely movable over, but in close contact with, the underlying testicles. A median raphe is present externally ventral to the median septum that separates the left and right scrotal sacs. The testicle is surrounded by a relatively thick tunica albuginea which, itself, is covered by the visceral vaginal tunic. The vaginal cavity is a potential space that permits the parietal vaginal tunic to slide freely over the visceral vaginal tunic. A small amount of fluid may
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Examination of External Genitalia accumulate in the vaginal cavity, localizing in the space surrounding the tail of the epididymis. The testicles should be ellipsoid, symmetrical in size and character, and have a turgid resilient consistency. The long axis of each testicle is horizontally positioned in the scrotum but may assume a more vertical position when the testicle is retracted. Total scrotal width is correlated to daily sperm production and daily sperm output.5 A stallion should have a total scrotal width of at least 8 cm to be considered a satisfactory prospective breeder.6, 7 Calipers can be used to obtain surface measurements of the testicles through the scrotum. Testicles may also be measured by transcutaneous ultrasound examination. Accurate estimation of testicular volume will allow improved prediction of daily sperm output.8 Using metric measurements (cm) consisting of the testicular width (W), height (H), and length (L), one can estimate the volume of each testicle by using the following equation: Volume = 4/3π × W/2 × H/2 × L/2 This equation reduces down to the simple formula: Volume = 0.523(W × H × L) The potential for the stallion’s daily sperm output (DSO) can be estimated using the sum of the volume of each testicle: Predicted DSO (×109 ) = (Total testicular volume × 0.024) − 0.76 Stallion managers should monitor their stallion’s testicles monthly so abnormalities can be detected promptly. The spermatic cord is positioned at the cranial aspect of the testicle and contains the testicular artery, pampiniform plexus, lymphatic vessels, deferent duct, and cremaster muscle. The deferent duct is not as readily palpable at the level of the spermatic cord in the stallion as it is in the bull and ram, but it can be felt medial and parallel to the body of the epididymis. The head of the epididymis is not readily discernable at the medial cranial aspect of the testicle. The body of the epididymis is positioned dorsally and lateral along the long aspect of the testicle. The tail of the epididymis is positioned at the caudal pole of the testicle. Care should be taken to always palpate the ligament of the tail of the epididymis. This ligament has a distinctive firm (∼1 cm flat) character and it is an obvious landmark that allows the examiner to clearly determine the position of the testicle in the scrotum. Incidentally, one may find a 180◦ rotation of the testicle in the scrotum which is not as-
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sociated with any clinical problems. Testicles that are a bit more spherical in shape (rather than ellipsoidal) seem to be more prone to spontaneous rotation. If the degree of twist in the spermatic cord is great enough to interfere with blood flow to or from the testicle the stallion may demonstrate acute signs of pain and be at risk for testicular compromise.
Penis Although a stallion caretaker may not actually observe discharge from the penis, its evidence may be seen on the dorsal aspect of the horse’s inner thighs. Stallions experiencing urinary incontinence in conjunction with pelvic neurological problems or urinary calculi may have dried urine crystals on the cranial surface of the hindlimbs. Horses with neoplasia of the penis, such as squamous cell carcinoma or melanoma, may have dried exudate or excessive smegma on the hindlimbs chronically. Using a gloved finger, one should gently probe the penile fossa and urethral sinus to remove any accumulated smegma (sometimes termed beans) and feel for irregularities that might be consistent with early neoplastic changes. The musculocavernous penis becomes rigid during tumescence, extending cranially and ventrally. The larger cavernous space, the corpus cavernosum, is positioned dorsally to the corpus spongiosum penis. The penile urethra is immediately surrounded by the corpus spongiosum penis, which extends distally to the glans and expands considerably during intromission as the corpus spongiosum glandis. During copulation, the distended engorged corona glandis,
Figure 149.1 Care should be taken to carefully examine the urethral process for any lesions or lacerations. The penile fossa should also be inspected for presence of accumulated smegma or mucosal irregularities.
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Figure 149.2 The integrity of the urethral process can be assessed during ejaculation by using an open-ended artificial vagina. Note the degree of dilation of the urethral process.
which may be referred to as the ‘bell,’ has a plungerlike effect to distend the anterior vagina. This may allow the distal end of the penile urethra, the urethral process, to be aligned with the open cervix, resulting in the ejaculate being deposited directly inside the uterus. The urethral process should be examined carefully for evidence of lacerations or abrasions (Figure 149.1). Lesions may not be obvious in the nonerect, or even the erect, penis because hemorrhage may only occur during ejaculation. A more thorough examination of the urethral process is indicated in stallions with hemospermia. This can be accomplished using an open-ended Polish-type artificial vagina that permits direct observation of the glans penis during ejaculation (Figure 149.2).
Figure 149.4 The internal lamina of the internal prepuce is adhered to the distal glans of a colt’s penis for the first few weeks of life. This adhesion does not interfere with urination but does prevent complete extension of the penis.
The urethral process distends extensively at the time of ejaculation, and hemorrhage from this location can be readily detected. The prepuce is thin and loosely attached to the shaft of the penis. In the retracted penis there is an external preputial fold at the interface with the skin of the sheath. The internal preputial fold is apparent midshaft in the erect penis. Signs of penile edema are often most pronounced in the internal and external lamina of the internal prepuce, which are located just distal and proximal to the internal preputial fold, respectively (Figure 149.3). Pronounced preputial swelling can usually be reduced significantly by the short-term intermittent use of an Eschmark rubber bandage to reduce the subcutaneous edema. It should be noted that the internal lamina of the internal prepuce is adhered to the distal glans of the penis in the male fetus. This attachment may persist for a few weeks after birth. This adhesion will not interfere with urination but will initially prevent neonatal colts from completely exposing the penis when posturing to urinate (Figure 149.4).
References
Figure 149.3 The more distally located internal lamina of the internal prepuce is more swollen than the external lamina of the internal prepuce. Note the dried exudate/smegma on the inner thigh of this horse.
1. Amman RP. A review of anatomy and physiology of the stallion. J Equine Vet Sci 1981;1:83–105. 2. Varner DD, Schumacher J, Blanchard TL, Johnson L. Reproductive anatomy and physiology. In: Pratt PW (ed.) Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991; p. 159. 3. Little TV, Holyoak GR. Reproductive anatomy and physiology of the stallion. Stallion Management. Vet Clin North Am Equine Pract 1992;8:1–29. 4. Rochat MC. Priapism: a review. Theriogenology 2001;56: 713–22.
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Examination of External Genitalia 5. Thompson Jr DL, Pickett BW, Squires EL, Amann RP. Testicular measurements and reproductive characteristics in stallions. J Reprod Fertil Suppl 1979;27:13–17. 6. Kenney R, Hurtgen J, Pierson R, Witherspoon D, Simons J. Theriogenology and the equine. Part II. The stallion. J Soc Theriogenol 1983;IX:50.
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7. Picket BW, Voss JL, Squires EL. Fertility evaluation of the stallion. Comp Cont Educ Pract Vet 1983;5(4):S194–S202. 8. Love CC, Garcia MC, Riera FR, Kenney RM. Use of testicular volume to predict daily sperm output in the stallion. In: Proceedings of Annual Convention of the American Association of Equine Practitioners, 1990, pp. 15–20.
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Chapter 150
Cytogenetic Evaluation Keith W. Durkin, Terje Raudsepp and Bhanu P. Chowdhary
Introduction Cytogenetic analysis has been used for decades for assessing normal fertility in animals and humans. The simple logic associated with the use of this approach is that it facilitates rapid analysis of the chromosome complement of an individual and aids in determining whether or not the chromosomes are structurally and/or numerically normal. Abnormalities of either type are a potential cause of reduced fertility or infertility because they disrupt normal pairing and segregation of chromosomes during meiosis. This leads to a reduction in the number of gametes that can establish viable pregnancies; furthermore, some defects can lead to a generation of progeny that further propagate the abnormal chromosome complement. Abnormal gametes have a direct impact on the survivability of the zygote or can cause a range of defects and disorders if the fetus is carried to full term. In instances where individuals carrying chromosomal abnormalities produce viable and apparently normal offspring, the proportion inheriting the abnormal chromosome complement will likely experience reduced fertility or infertility. As such, it would seem intuitive that all stallions and mares be subjected to chromosome analysis before being introduced into a breeding program. Such testing becomes particularly important for stallions because the equine industry uses a select group of stallions for breeding purposes. In this chapter, we first provide an outline of how cytogenetic evaluation is performed in horses and then give an overview of chromosome abnormalities found in stallions. A summary of how these abnormalities affect fertility is provided. Finally, the significance of establishing a mandatory program for cytogenetic evaluation of breeding stallions (as well as mares) is briefly discussed.
Standard approaches used to evaluate horse chromosomes Cytogenetic evaluation in the horse began in earnest following the first accurate count of the equine chromosomes (2n = 64) by Rothfels et al.1 Early studies relied on Giemsa staining to identify numerical aberrations but precluded definitive identification of the chromosomes involved.2 Development of improved cell culture and harvesting techniques, coupled with the introduction of chromosome banding techniques in the 1970s, allowed for the first time definitive identification of the horse chromosomes. This information, and a number of subsequent studies during the 1980s and 1990s, paved the way for the development of an International System for Cytogenetic Nomenclature of the Domestic Horse3 that is presently being used to evaluate horse chromosomes. Chromosome preparations are typically obtained from peripheral blood lymphocyte culture, although they can also be obtained from long-term tissue cultures. For short-term lymphocyte culture, 5–10 mL blood is aseptically collected from the jugular vein in heparinized tubes and allowed to settle. Lymphocytes are obtained from the ‘buffy coat’ and are cultured for 72 hours in medium supplemented with fetal calf serum, mitogen, and antibiotics. The cultures are harvested using standard approaches and the cells are subjected to hypotonic treatment.4 This is followed by four rounds of fixation, which are then followed by bursting the cells on a clean wet slide to obtain metaphase spreads (Figure 150.1a). The first step to evaluate the horse chromosomes involves counting the chromosome number and arranging the chromosomes into a karyotype to ensure that (i) the number of chromosome in each of the 20 spreads evaluated is 64XY for males or 64XX for the females; (ii) the chromosomes appear to be
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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(d) Figure 150.1 (a) A typical metaphase spread routinely imaged for analysis; (b and c) C-banding that usually stains the centromeres clearly shows heterochromatic regions (arrows) on the long arm of the X chromosome (b is a female = XX) and the majority of the Y chromosome (c is XY); (d) a G-banded karyotype of a normal stallion. Chromosomes are arranged according to the International System for Cytogenetic Nomenclature of the Domestic Horse.3
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Figure 150.2 Use of molecular cytogenetic approaches to define chromosome aberrations. (a) Fluorescent in situ hybridization (FISH) with two bacterial artificial chromosome (BAC) clones from the short arm of ECA5 show that the red-labeled BAC hybridizes to the marker chromosome (M) and the green-labeled probes hybridize to the derivative chromosome (D), indicating the precise break-/fusion-point region. (b) FISH with a BAC clone from ECA27 (red) and ECA28 (light blue) shows that the colt carries ECA27 trisomy. (See color plate 46).
structurally normal; and (iii) the genetic sex (as depicted by the sex chromosomes) corresponds to the phenotypic sex. To further verify the sex chromosomes, the preparations are stained by C-banding.5 The X chromosome shows a characteristic heterochromatic dark Giemsa band on the proximal part of the long arm, whereas the Y chromosome is small and largely dark staining because it is predominantly heterochromatic (Figure 150.1b,c). Each animal is also tested for the presence or absence of the sex-determining region Y (SRY) gene using molecular approaches. This routine analysis (which can generally be reported to the owners within 10 working days with an image of the chromosomes) is expanded if a structural abnormality is suspected or a numerical aberration is present. For this, the chromosomes are stained by G-banding analysis (trypsin digestion followed by Giemsa staining6 ) and the banding patterns of the homologues are compared (Figure 150.1d). If an aberration is found, it is further analyzed by molecular genetic techniques to precisely define losses or rearrangements (Figure 150.2). As a point of clarification, other techniques that can be used for karyotyping include Rbanding (the reverse of G-banding, in which the guanine/cytosine-rich euchromatic regions are dark and the thymine/adenine-rich heterochromatin regions are light); DAPI-banding (which utilizes a
fluorochrome 4 ,6-diamidino-2-phenylindole dihydrochloride and produces fluorescent G-band-like staining patterns); and C-banding (in which most of the chromatin is removed by hydrolysis; thus subsequent staining with Giemsa highlights only the most resistant heterochromatin at the centromeres). The C-banding technique is mainly used to identify equine sex chromosomes and is of little value for karyotyping.7
Chromosome abnormalities in stallions Traditionally, chromosome abnormalities are broadly classified as either numerical or structural. They are also classified as autosomal or sex chromosome type depending on the chromosomes involved. Here, we summarize various chromosomal aberrations detected to date in stallions. Numerical and structural abnormalities of the sex chromosomes will be discussed first followed by the abnormalities of the autosomes. Some chromosome abnormalities have not yet been reported in stallions and the occurrence of the others would appear to be considerably lower than that found in species such as humans, mice, pig, and cattle. These observations may lead one to a grossly misleading conclusion that chromosome abnormalities occur at a much lower frequency in the horse because far fewer cases are traditionally sent
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Cytogenetic Evaluation for chromosome analysis in the horse than in other species. Similarly, the number of stallions evaluated for chromosome abnormalities is significantly lower than the number of mares, a feature created by the mating structure within the equine industry where the number of males used at stud is limited. Within this limited population of stallions, traditionally very few have been evaluated for chromosome defects. This is in sharp contrast to pigs and cattle, where in a number of countries all breeding males are mandated to undergo chromosome analysis to ensure that chromosome abnormalities do not spread in the population. Abnormalities involving the X chromosome Among the few X chromosome abnormalities found up to now in stallions, all are numerical and involve one or two extra X chromosomes (di- or trisomies), resulting in 65XXY and 66XXXY karyotypes. Additionally, a few mosaic individuals have been described that have a mixture of normal (XY) and abnormal (XXY, XXXY, or XO) cell populations. The 65XXY stallions share phenotypic similarities to humans with Klinefelter’s syndrome, and to bulls and boars affected with a similar sex chromosome complement.8–10 To date, only two 65XXY stallions have been described in detail. The first was a draft horse11 that had no scrotum but two small, soft, and flaccid palpable gonads, unusually large epididymides, and a normal penis and sheath. The other animal was a French trotter12 that had relatively small, but normally descended, soft testes, a small penis, and a brownish azoospermic ejaculate. In a recent genotyping-based survey for sex chromosome abnormalities in a population of 17 471 foals,13 four new cases of 65XXY were identified. The authors concluded that, as only 60% of the studied animals had informative genotyping data, the actual frequency of sex chromosome abnormalities might be even higher. Men with multiple X chromosomes and a Y chromosome (e.g., XXXY, XXXXY) are typically infertile and often display additional somatic abnormalities.14 In horses, however, the only 66XXXY individual described so far15 had an intersex phenotype with underdeveloped external genitalia from both sexes, an infantile uterus, and gonads with abnormal seminiferous tubules. A few 65XXY mosaic (mixed population of cells with different chromosome numbers) horses have been described. The phenotype of these individuals was dependent on the proportion of male and female cells. An American Standardbred stallion with 60% 65XXY cells and 40% normal 64XY cells was phenotypically normal, but failed to produce a successful pregnancy, despite covering 27 mares,12 and was determined to be azoospermic. Animals in which
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65XXY cells are associated with 64XX cells or other numerically abnormal cells that lack a Y chromosome usually show an intersex phenotype.2 Intersex individuals with ambiguous genitalia might also be mosaics for 63XO (X monosomy) and Y chromosomecontaining cell lines.16–18 Abnormalities involving the Y chromosome Measurement of Y chromosomes in 31 male horses19 revealed significant interindividual size polymorphism both in the heterochromatic and euchromatic portions of the Y chromosome, although no associations between Y chromosome size and clinical abnormalities were detected. To date, no cases of pure 65XYY individuals have been reported in the horse. The karyotype has been seen in mosaic form only in conjunction with a 63XO cell line,20, 21 and is associated with an intersex phenotype. It is possible that 65XYY horses are present in the population but remain unidentified because they appear phenotypically and reproductively normal (as in humans22 ). Among structural abnormalities, an isochromosome Y has been described in a male pseudohermaphrodite (more correctly referred to as intersex) that had one cell line containing a Y isochromosome and another with a 63XO chromosome complement.23 Recent studies conducted in our laboratory indicate that minor deletions and/or rearrangements of the Y chromosome that usually go unnoticed by standard cytogenetic approaches can now be detected by molecular techniques. Sex reversal, a condition in which genetic/chromosomal sex does not agree with the phenotypic sex, can be detected only by chromosome analysis. Female sex reversal, where mares have a male (64XY) karyotype, is the second most common cytogenetic condition in horses and has been described elsewhere.2 These individuals show a range of intersex phenotypes – from phenotypic males with small undescended testicles and reduced libido to phenotypic females with abnormal testes and ovaries, an enlarged clitoris, and stallion-like behavior.2, 24 Initial studies show that an autosomal recessive mutation might be involved in 64XX sex reversal in horses.24 Abnormalities involving the autosomes The only known viable numerical abnormalities of horse autosomes are trisomies. Like other mammals, the smallest and most gene-poor chromosomes are typically involved in equine trisomies.2, 25 The five autosomal trisomies found in stallions involve four chromosomes (ECA23, ECA27, ECA28, and ECA31)
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and have been found in two Thoroughbred colts,26, 27 a Standardbred colt,28 a Standardbred gelding,29 and an Arabian colt.30 It is noteworthy that all trisomic individuals show a variety of developmental, reproductive, and behavioral abnormalities. Similarly, trisomies of small autosomes have been found in other species, of which the best known is the trisomy of chromosome 21 in human Down’s syndrome.31 To date, 10 documented cases of structural abnormalities have been described in horses,2 of which two deletions and two translocations have been found in stallions and all are associated with infertility or reduced fertility, as documented by rates of pregnancy establishment and pregnancy loss. The two autosomal deletions – del13qter in a Standardbred stallion32 and mosaicism for a deletion on ECA10 in an Arabian stallion33 – were identified because of fertility issues. A tandem-fusion translocation between chromosomes 1 and 30 has been described in a Thoroughbred stallion.34 The stallion had fertility problems.
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Mares covered by the stallion experienced a high incidence of early embryonic death with only a small number of pregnancies going to full term. More recently, cytogenetic analysis was carried out on another Thoroughbred stallion because only 34% of mares covered by the animal produced normal live foals. The stallion had small testicular size and a reduced sperm count. The stallion was phenotypically normal and had a successful career on the racetrack. A historical account of the stallion revealed that many mares mated to the stallion lost their conceptus during early pregnancy. Cytogenetic evaluation revealed a translocation between ECA5 and ECA16 and the presence of a small marker chromosome. Subsequent molecular cytogenetic analysis by fluorescent in situ hybridization (FISH) using bacterial artificial chromosome (BAC) probes from ECA5 showed that the marker chromosome was derived from the terminal 9 Mb of the euchromatic region of ECA5p (Figure 150.3). It is noteworthy
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(e) Figure 150.3 (a) Partial metaphase showing aberrant chromosomes: D, derivative chromosome ECA5+ECA16; M, marker chromosome. (b) Schematic representation of ECA5, ECA16, derivative, and the marker chromosomes in a translocation found in a stallion with poor fertility and a high rate of early embryonic death. (c) Possible pairing of normal and derivative chromosomes. (d) Alternate segregation produces a balanced chromosome complement (carries no duplications or deletions and has a viable pregnancy). (e) Adjacent 1 and 2 segregation produces a genetically unbalanced chromosome complement (carries duplications or deletions, non-viable pregnancy).
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that chromosome analysis of 10 resulting foals (six males and four females) revealed that only one colt and one filly had a normal karyotype while the remaining animals had the same translocation as their sire. Evidently, this is a classical case where a chromosome translocation is being spread out in a population because a carrier stallion is being used for breeding. The consequences of the translocation in the eight cytogenetically abnormal individuals will be felt by the owners at the time of breeding. Of the six types of gametes produced by the stallion, only one is normal (Figure 150.3); among the remaining gametes, only one is balanced (like the stallion) and the other four have deletions or duplications that lead to early embryonic death. Thus, over two-thirds of the gametes produced by the stallion are expected to cause either infertility or embryonic losses during the early stages of pregnancy and another proportion will recreate the abnormality in the offspring.
BACs spanning all equine chromosomes will very soon permit the generation of a tiling-path array that will be vital in the analysis of microscopic chromosome variations, including deletions, duplications, and copy number variations that may also contribute to reproduction and fertility problems. Once optimized, analysis of this type of array will add a new dimension to the cytogenetic analysis presently being offered. In our view, veterinarians, horse owners, potential buyers, insurance agencies, and the equine industry must initiate discussions regarding a mandate for basic cytogenetic analysis for all prospective breeding stallions (and mares). The test is simple and can readily be carried out from blood samples taken at any age. However, conducting the test at an early age will avoid financial losses incurred by all parties involved, and will also avoid spreading of a chromosomal abnormality.
Meiotic chromosomes
Acknowledgements
Cytogenetic analysis of meiotic chromosomes from stallion spermatocytes has been directed mainly at the examination of normal male meiosis.35–37 However, meiotic chromosomes from infertile equids have been analyzed by two groups: Chandley et al.35 examined meiosis in a mule and a hinny, whereas Power et al.39 investigated synaptonemal complexes in an infertile stallion carrying trisomy ECA28.26 The trisomic animal demonstrated a trivalent or a bivalent plus a univalent in primary spermatocytes which triggered meiotic arrest, with degeneration at both spermatocyte and spermatid levels, and was probably the main cause of observed azoospermia. Given that meiotic defects in humans (disorders in meiotic recombination, reduced recombination frequency, abnormal pairing, asynapsis, etc.) account for at least 8% of all cases of male infertility,40 analysis of meiotic chromosomes in stallions should be undertaken to develop an improved understanding of their contribution to reduced fertility.
Support for equine genome research, provided by Link Endowment, the Morris Animal Foundation, Grayson Jockey Club, TERRAC, and USDA (grant no. 2007-04801), is gratefully acknowledged.
Advances in equine cytogenetic evaluation and significance The horse genome has been sequenced and a highdensity genetic map of the horse is available. A range of other tools and resources, including chromosome paints and BAC clones for physically ordered markers on all autosomes and the sex chromosomes, enables the use of molecular techniques to analyze chromosomal abnormalities. This will permit detection of small aberrations that have been difficult to discover until now. Moreover, the availability of
References 1. Rothfels KH, Alexrad AA, Siminovitch L, McCulloch Parker RC. The origin of altered cell lines from mouse, monkey and man as indicated by chromosome and transplantation studies. In: Proceedings of the 3rd Canadian Cancer Research Conference, 1959, pp. 189–214. 2. Chowdhary BP, Raudsepp T. Cytogenetics and physical gene maps. In: Bowling AT, Ruvinsky A (eds) The Genetics of the Horse. Wallingford, UK: CABI, 2000; pp. 171– 242. 3. Bowling AT, Breen M, Chowdhary BP, Hirota K, Lear T, Millon LV, Ponce de Leon FA, Raudsepp T, Stranzinger G. Report of the Third International Committee for the Standardization of the Domestic Horse Karyotype, Davis, CA, USA, 1996. Chromosome Res 1997;5:433–43. 4. Raudsepp T, Chowdhary BP. FISH for mapping single copy genes. In: Murphy W (ed.) Phylogenomics: Methods in Molecular Biology. Totowa, NJ: Humana Press, 2007; p. 310. 5. Arrighi FE, Hsu TC. Localization of heterochromatin in human chromosomes. Cytogenetics 1971;10:81–6. 6. Seabright M. A rapid banding technique for human chromosomes. Lancet 1971;2:971–2. 7. Comings DE. Mechanisms of chromosome banding and implications for chromosome structure. Annu Rev Genet 1978;12:25–46. 8. M¨akinen A, Andersson M, Nikunen S. Detection of the X chromosomes in a Klinefelter boar using a whole human X chromosome painting probe. Anim Reprod Sci 1998;52:317–23. 9. Molteni L, De Giovanni Macchi A, Meggiolaro D, Sironi G, Enice F, Popescu P. New cases of XXY constitution in cattle. Anim Reprod Sci 1999;55:107–13.
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10. Slota E, Kozubska-Sobocinska A, Bugno M, Giemza-Marek A, Kulig B. Detection of the XXY trisomy in a bull by using sex chromosome painting probes. J Appl Genet 2003;42:59–64. 11. Kubien EM, Pozor MA, Tischner M. Clinical, cytogenetic and endocrine evaluation of a horse with a 65, XXY karyotype. Equine Vet J 1993;25:333–5. 12. M¨akinen A, Katila T, Andersson M, Gustavsson I. Two sterile stallions with XXY-syndrome. Equine Vet J 2000;32:358–60. 13. Kakoi H, Hirota K, Gawahara H, Kurosawa M, Kuwajima M. Genetic diagnosis of sex chromosome aberrations in horses based on parentage test by microsatellite DNA and analysis of X- and Y-linked markers. Equine Vet J 2005;37:143–7. 14. Miller OJ, Therman E. Human Chromosomes. New York: Springer, 2001. 15. Gluhovschi N, Bistriceanu M, Suciu A, Bratu M. A case of intersexuality in the horse with type 2A+ XXXY chromosome formula. Br Vet J 1970;126:522–5. 16. Chandley AC, Fletcher J, Rossdale PD, Peace CK, Ricketts SW, McEnery RJ, Thorne JP, Short RV, Allen WR. Chromosome abnormalities as a cause of infertility in mares. J Reprod Fertil Suppl 1975;23:377–83. 17. Bowling AT, Millon L, Hughes JP. An update of chromosomal abnormalities in mares. J Reprod Fertil Suppl 1987;35:149–55. 18. Long SE. Chromosome anomalies and infertility in the mare. Equine Vet J 1988;20:89–93. 19. Power MM. Y chromosome length variation and its significance in the horse. J Heredity 1988;79:311. 20. Hohn H, Klug E, Rieck GW. A 63, XO/65, XYY Mosaic in a case of questionable equine male pseudohermaphroditism. In: Proceedings of the 4th European Colloquium in Cytogenetics of Domestic Animals, 1980, pp. 82–92. 21. Paget S, Ducos A, Mignotte F, Raymond I, Pinton A, Seguela A, Berland HM, Brun-Baronnat C, Darre A, Darre R. 63, XO/65, XYY mosaicism in a case of equine male pseudohermaphroditism. Vet Rec 2001;148: 24–5. 22. Gonzalez-Merino E, Hans C, Abramowicz M, Englert Y, Emiliani S. Aneuploidy study in sperm and preimplantation embryos from nonmosaic 47, XYY men. Fertil Steril 2007;88:600–6. ¨ H, Klug E, Hecht W. Sex chromosome mo23. Herzog A, Hohn saic in male pseudohermaphroditism in a horse. Tier¨arztl Prax 1989;17:171–5. 24. Buoen LC, Zhang TQ, Weber AF, Ruth GR. SRY-negative, XX intersex horses: the need for pedigree studies to examine the mode of inheritance of the condition. Equine Vet J 2000;32:78–81.
25. Bugno M, Slota E, Koscielny M. Karyotype evaluation among young horse populations in Poland. Schweiz Arch Tierheilkd 2007;149:227–32. 26. Power MM. Equine half sibs with an unbalanced X; 15 translocation or trisomy 28. Cytogenet Cell Genet 1987;45:163–8. 27. Lear TL, Cox JH, Kennedy GA. Autosomal trisomy in a Thoroughbred colt: 65, XY, +31. Equine Vet J 1999;31:85–8. 28. Brito LFC, Sertich PL, Durkin K, Chowdhary BP, Turner RM, LM, Sue McDonnell S. Autosomic 27 trisomy in a standardbred colt. J. Equine Vet Sci 2008;28:431-436. 29. Klunder LR, McFeely RA, Beech J, McClune W. Autosomal trisomy in a Standardbred colt. Equine Vet J 1989;21:69–70. 30. Buoen LC, Zhang TQ, Weber AF, Turner T, Bellamy J, Ruth GR. Arthrogryposis in the foal and its possible relation to autosomal trisomy. Equine Vet J 1997;29:60–2. 31. Megarbane A, Ravel A, Mircher C, Sturtz F, Grattau Y, Rethore M-O, Delabar J-M, Mobley W. The 50th anniversary of the discovery of trisomy 21: the past, present, and future of research and treatment of Down syndrome. Genet Med 2009;11:611–16. 32. Halnan CRE, Watson JI. Detection by G-and C-band karyotyping of gonosome anomalies in horses of different breeds. J Reprod Fertil Suppl 1982;32:626–7. 33. McFeely RA, Klunder LR, Byars D. Equine infertility associated with autosomal mixoploidy. In: Proceedings of the 6th North American Colloquium on Domestic Animal Cytogenetics, 1989, p. 7. 34. Long SE. Tandem 1; 30 translocation: a new structural abnormality in the horse (Equus caballus). Cytogenet Cell Genet 1996;72:162–3. 35. Chandley AC, Jones RC, Dott HM, Allen WR, Short RV. Meiosis in interspecific equine hybrids. I. The male mule (Equus asinus X E. caballus) and hinny (E. caballus X E. asinus). Cytogenet Cell Genet 1974;13:330–41. 36. Scott IS, Long SE. An examination of chromosomes in the stallion (Equus caballus) during meiosis. Cytogenet Cell Genet 1980;26:7–13. 37. Safronova LD, Pimenova TI. [The karyotyping of bull (Bos taurus) and of horse (Equus caballus) on the basis of the synaptonemal complexes]. Genetika 1988;XXIV:709–15. 38. Nett von P, Jung HR, Stranzinger G. Neue Erkenntnisse uber die Chromosomenpaarung in der Meiose. Dtsch Tierarztl Wschr 1996;103:369–448. ¨ 39. Power MM, Gustavsson I, Switonski M, Ploen L. Synaptonemal complex analysis of an autosomal trisomy in the horse. Cytogenet Cell Genet 1992;61:202–7. 40. Topping D, Brown P, Hassold T. The immunocytogenetics of human male meiosis. In: Carrell DT (ed.) The Genetics of Male Infertility. Totowa, NJ: Humana Press, 2007; pp. 115–44.
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Chapter 151
Ultrasonography of the Genital Tract Regina M.O. Turner
Introduction Ultrasonography of the genital tract of the stallion is now used routinely as an adjunct to the routine breeding soundness examination. In addition, ultrasonography is very useful when evaluating suspected reproductive pathology. Because there can be variation in the ultrasonographic appearance of reproductive tract abnormalities, there is great benefit to becoming familiar with the normal appearance of the structures before proceeding to evaluation of abnormal states. This chapter will review the normal ultrasonographic appearance of the stallion reproductive tract and provide some examples of ultrasonographically identified pathology.
Examination of the scrotal contents Proper restraint is important to the safe examination of the stallion’s testicles, epididymis, and spermatic cord. One good location for the examination is with the stallion backed into a corner such that the animal’s right side and hindquarters are against the adjoining walls. This helps to limit lateral and backwards movement of the stallion and may discourage kicking. Stocks also can be used. However, they must be designed in a way that allows the veterinarian access to the scrotal contents (e.g., a removable side or a single horizontal bar). This examination causes no discomfort to the animal, so most stallions can be safely examined with only a chain shank over the nose or in the mouth. A twitch or chemical restraint may be necessary for nervous or intractable animals. When ultrasonographic examinations are performed as part of a breeding soundness examination, it may be advantageous to perform these examinations after the final semen collection. Most stallions will be more
relaxed and tolerant at this point than they would have been prior to semen collection.1 Ideally, the ultrasound machine should be placed on a cart that can be readily moved if the stallion misbehaves. If available, an assistant can be assigned the task of managing the ultrasound machine.
Technique for examination of the scrotal contents Prior to ultrasonographic examination, the testicles, epididymis, spermatic cord, and inguinal region should be visualized and palpated. This allows the stallion to become accustomed to the examination prior to introducing the transducer and also allows the veterinarian to identify any obvious abnormalities that may warrant more careful attention. The veterinarian should approach the horse’s left shoulder and, while keeping his or her body in contact with the stallion, reach under the stallion’s abdomen with the right hand. Gently patting or scratching the stallion on the ventral abdomen can help relax the stallion prior to palpation of the scrotal contents. Once the stallion accepts contact on the ventral abdomen, the veterinarian can then reach into the inguinal region for palpation of the scrotal contents. By reaching under the stallion with the right hand, the veterinarian should be able to keep the rest of his or her body out of reach of the stallion’s hind legs. If the stallion is standing quietly, the veterinarian can switch to examination with the left hand if preferred, however this will require that he or she stand closer to the stallion’s hind legs. After the scrotal contents have been fully examined by palpation and once the stallion is relaxed, ultrasonographic examination can begin.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Ultrasonographic examination will require the use of both hands simultaneously (one to hold the transducer and the other to stabilize the testes). This typically means that the veterinarian must crouch or kneel just in front of the stallion’s hind legs. Most stallions will tolerate this examination without objection. However, if you are in doubt about the stallion’s temperament, sedation is warranted to help insure safety. The ultrasound cart is placed in a location out of reach of the stallion but within easy visualization of the veterinarian. Generous amounts of an approved ultrasound coupling gel, methanol or ethanol, mineral oil or a water-soluble gel used for gynecological examinations should be used to ensure good contact between the transducer face and the scrotal skin. Most coupling agents should be removed by washing after the examination. A linear array or microconvex 7.5 MHz transducer is well suited for examination of the scrotal contents. The addition of a 10 MHz transducer may be useful for imaging details of more superficial structures like the epididymis. A 5 MHz transducer also is acceptable, although some superficial details of the scrotal contents may be lost. Sector scanners can be used, but proper placement of the transducer can be more difficult due to the curved footprint of the transducer. To begin the ultrasonographic examination, the left hand is used to push the right testicle up out of the way and extend and stabilize the left testicle to provide firm contact with the transducer. The transducer is held in the right hand and gel is applied to the face. The transducer face then is placed against the lateral surface of the testicle perpendicular to its long axis. The transducer then is slowly moved several times over the entire lateral surface of the testicle to view a series of cross-sectional images of the parenchyma (Figure 151.1). Sagittal (long axis) views of the testicular parenchyma can be obtained by moving the transducer over the cranial surface of the tes-
B
A
C
Figure 151.1 Methods for positioning the transducer to image the testicular parenchyma in short-axis cross-section (A) and the head (B) and tail (C) of the epididymis.
ticle and directing the beam caudally or by holding the transducer on the ventral surface of the testicle with the long axis of the transducer parallel to the long axis of the testicle. In most Thoroughbred type or larger breed stallions, the entire long axis of the testicle is too large to be seen in a single frame. The transducer can be moved around as needed to ensure that all areas of the testicular parenchyma have been visualized. The head, body, and tail of the epididymis can be more difficult to image due to their superficial locations along the surface of the curved testicle. The transducer can be placed either directly over each region of the epididymis (Figure 151.1) or, in the case of the epididymal tail, it can be placed over the testicle just cranial to the tail and the beam directed caudally toward the tail. After examination of the testicles and epididymides, the transducer should be moved further up into the inguinal region for examination of the spermatic cord. For this examination, the transducer is held in the right hand and positioned over the spermatic cord, parallel to the ground (i.e., providing cross-sections of the cord) while the left hand gently but firmly applies downward traction on the testicle. The transducer is slowly moved up and down the spermatic cord as far as can be safely examined. The entire process is then repeated with the right testicle while the left testicle is pushed up out of the way. Uncommonly, it may be necessary to rearrange the horse and the equipment to allow examination from the horse’s right side.
Ultrasonographic imaging of the scrotal contents Testicle The ultrasonographic appearance of the normal testicular parenchyma is homogeneous, gray, and granular (Figure 151.2). Any significant variation in echodensity of the parenchyma should be noted and examined more carefully for possible pathology. A small amount (typically < 5 mm) of anechoic fluid can normally be seen in the vaginal cavity between the visceral and parietal tunicae (Figure 151.2). The central vein of the testicle is visible as an anechoic line traversing the approximate center of the testicular parenchyma (Figure 151.2). The central vein is widest craniodorsally where it joins the spermatic cord and narrows dorsoventrally. Typical width measurements of the central vein vary from 1 mm to 4 mm depending on which area of the vein is imaged and the angle of the transducer. The parietal vaginal tunic and
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Figure 151.3 Ultrasonographic image of a short-axis crosssection of an arcuate artery on the dorsal surface of the testicle (arrow).
Figure 151.2 Ultrasonographic image of normal testicular parenchyma in short-axis cross-section with the transducer held as shown in Figure 151.1, A. Portions of the vaginal cavity are visible as anechoic borders, particularly along the dorsal surface of the testicle (black arrows). The central vein also is visible in the testicular parenchyma (white arrow). The parietal vaginal tunic is viewed as a thin white line outside the vaginal cavity.
its associated structures appear as soft-tissue density structures outside the vaginal cavity (Figure 151.2). When scanning with a 7.5 MHz or higher frequency transducer, the small branches of the testicular artery (arcuate arteries) typically are seen near the surface of the testicle in the tunica albuginea (Figure 151.3). These vessels are more difficult to discern when using a 5 MHz transducer due to their location and small size (usually less than 5 mm in diameter). During this examination, the veterinarian should measure width, height, and length of each individual testicle. Ultrasonographic measurement of testicular size is highly accurate2 and can be easier to perform than caliper measurements. For measurement of individual testicular width, the ultrasound transducer should be held on the lateral surface of one testicle while the contralateral testicle is pushed up and out of the way of the transducer. To minimize artifactual alterations of testicular measurements, the beam should be directed across the widest part of the testicle and perpendicular to its long axis (Figure 151.1, A). Height is measured by placing the transducer on the ventral surface of the testicle and directing the beam
dorsally at a 90◦ angle to the long axis of the testicle. Length can be determined by placing the transducer at the cranial aspect of the testicle and directing the beam caudally. However, in many stallions, testicular length is too large to fit on the view screen. In these animals, caliper measurements can be substituted. The epididymis should not be included in either ultrasonographic or caliper measurement of testicular dimensions. Normal testicular dimensions will vary depending on the age and breed of the stallion and the season of the year. For average sexually mature, Thoroughbred-type stallions, typical testicular length is 80 to 140 mm and width is 50 to 80 mm with height similar to width.3 Total scrotal width also should be measured. This is done by applying gentle, steady, downward traction on both testicles until they are side-by-side in the scrotum. With the testicles in this position, the beam of the transducer is directed across the widest part of the two testicles, perpendicular to their long axes. Total scrotal width is measured beginning at the lateral surface of the near testicle and ending at the lateral surface of the far testicle (Figure 151.4). A minimum total scrotal width of 8 cm is required for classification of a stallion as a satisfactory prospective breeder.4 These individual measurements can be valuable in themselves when used as a means of monitoring changes in testicular size over time for an individual stallion. Nonetheless, a stallion need not be labeled abnormal based on a single value that is marginally outside the normal range. More important information can be gained by using the length, width and height measurements to calculate total testicular volume. Testicular volume is calculated using the
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Testis
Epididymal Tail
Figure 151.4 Short-axis cross-sectional ultrasonographic image of normal testicles. The ultrasonographic calipers are delineating total scrotal width.
formula for the volume of an ellipsoid where: testicular volume (mL) = 4/3 pi × height/2 × width/2 × length/2. All testicular measurements should be in cm. Adding the volume of the left and right testicles together results in the value for total testicular volume in mL. Total testicular volume can then be used to predict daily sperm output.2 If a stallion does not produce normal sperm for his testicular volume, then that individual should be monitored closely and evaluated for possible testicular pathology. Epididymis Because the epididymis is composed of a single, highly convoluted, fluid-filled duct, ultrasonographically it appears to consist of numerous circular, anechoic areas distributed within a soft-tissue background. The lumen of the epididymal duct is narrowest in the epididymal head and widest in the tail. Because of the wider lumen in the epididymal tail, the tail is the easiest part of the epididymis to visualize and differentiate from the underlying testicular parenchyma5 (Figure 151.5).
Figure 151.5 Ultrasonographic image of the tail of the epididymis obtained with the transducer held as in Figure 151.1C. Note the numerous, round, anechoic cross-sections of the epididymal duct. A small portion of the testis is visible to the right of the image.
values now are available for normal stallions. A limited number of stallions with a variety of testicular pathologies also have been evaluated. These initial studies suggest that the technique may prove useful in the future for evaluation of testicular function.
Testicular abnormalities Cryptorchidism A cryptorchid horse is one in which one or both testicles have not descended into the scrotum. Retained testicles can be accurately identified, located, and measured ultrasonographically using either a 5
Spermatic cord When visualized in cross-section, the spermatic cord has a heterogeneous appearance with distinct anechoic circular areas surrounded by more hyperechoic regions (Figure 151.6). This appearance results from numerous cross-sections of the luminae of the testicular artery, the vessels in the pampiniform plexus, and the ductus deferens. In real time, blood flow can be seen within the vessels. Doppler and color Doppler ultrasonography have been used to evaluate blood flow in the vessels of the spermatic cord in stallions.6, 7 As a result, reference
Figure 151.6 Short-axis cross-sectional image of the spermatic cord. Note the numerous anechoic cross-sections of the vessels of the spermatic cord.
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Ultrasonography of the Genital Tract or 7.5 MHz linear array transducer transrectally (for abdominal testicles) or by scanning externally over the external inguinal ring (for inguinal testicles).8 Scanning over the external inguinal ring is easiest to accomplish using a sector scanner as the larger footprint of a linear transducer may make it harder to obtain a good image. A good approach to the identification of a retained testis is to first palpate and ultrasonographically examine the scrotum followed by the area adjacent to the external inguinal ring transcutaneously. Inguinal testes can usually be identified easily with ultrasound and some can be palpated. To search for an inguinal testicle, the horse should be safely restrained in a manner similar to ultrasonographic examination of the scrotal contents. Copious lubrication should be applied to the skin overlying the external inguinal ring. The face of the transducer is then pressed up into the inguinal region over the external inguinal ring to permit visualization of the inguinal canal. It may be necessary to apply firm pressure cautiously to the transducer to insure good contact with the skin. If testicular tissue is not identified by this approach, then a transrectal examination can be performed. Although abdominal testes can sometimes be palpated, they usually are easier to identify ultrasonographically. Abdominal testicles can be located anywhere from just caudal to the kidney to within the internal inguinal ring (i.e., anywhere along the path that the testis should follow during its normal descent into the scrotum). When searching for an abdominal testicle by palpation or ultrasonography, one approach is to first palpate the inguinal ring on the side of the missing testicle. The ring itself and the adjacent areas are then examined by additional palpation and ultrasonography until the presence or absence of the testicle is determined, as most abdominal testes are found in that area. Less commonly, the testicle may be located part way between the internal inguinal ring and the caudal pole of the kidney (the site of its embryonic origin). This area should be examined if the testicle is not located in the inguinal region. Starting at the inguinal ring, sweep the transducer back and forth slowly, gradually advancing towards the caudal pole of the kidney. If an abdominal testis is present it should be located somewhere between the internal inguinal ring and the kidney’s caudal pole. The parenchyma of a cryptorchid testicle is generally more hypoechoic than that of a descended testicle but is nonetheless readily identifiable. In some cases, the spermatic cord also may be identified within the abdomen. In these instances, the cord is typically more distinct than the testicle and can be used as a starting point from which the testicle can
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Figure 151.7 Ultrasonographic image of an abdominal testicle with its adjacent spermatic cord.
be found. Figure 151.7 shows the typical ultrasonographic appearance of an abdominal testis and its spermatic cord. Ultrasonographic identification of a retained testicle provides valuable information on the testicle’s location. In addition, teratomas and other neoplastic conditions occasionally are identified in the cryptorchid testicle.9 This can be extremely helpful in cases in which surgical removal is planned. Unlike endocrinological testing that can take several days, ultrasonography can provide an immediate answer as to whether or not a cryptorchid testis is present. Another condition where ultrasonography has been useful is in the diagnosis of male pseudohermaphroditism.9 Testicular enlargement Enlargement of the testicle is uncommon but can occur with orchitis, vascular or lymphatic stasis within the testicle, testicular abscess, testicular hematoma, or testicular neoplasia. A side-by-side ultrasonographic comparison of a normal and enlarged testis can be helpful when trying to assess whether or not the parenchyma in the affected testis is normal. Orchitis Orchitis, or inflammation of the testicular parenchyma, is a rare cause of testicular enlargement in the stallion. In horses, orchitis is often associated with trauma, however it can also be caused by infection, parasites or autoimmune disease.10 The affected testicle is typically enlarged, hot, edematous, and painful. Systemic signs such as fever, leukocytosis, and hyperfibrinogenemia may also be present. If semen is collected and evaluated, large numbers of white blood cells may be found and semen quality is often poor. Cultures of ejaculated semen may aid in identification of the causative organism in cases of infectious orchitis.
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(a)
(b)
Figure 151.8 Short-axis cross-sectional image of a normal testis (a) and a testis affected with orchitis (b). Note heterogeneity of the parenchyma and the interspersed areas of hyperechogenicity in the testis in panel B.
Chronic vascular stasis or lymphatic stasis, as in cases of chronic spermatic cord torsion, can cause testicular enlargement. If testicular enlargement is a result of spermatic cord torsion, abnormalities of the spermatic cord should also be evident ultrasonographically. Abnormalities of the spermatic cord will be discussed below.
enlarged, warm, painful testicle. Testicular abscesses can often be diagnosed with ultrasonography. A welldefined pocket of purulent fluid is generally visible within the testicular parenchyma. Abscess fluid typically contains large amounts of particulate debris that can sometimes be seen swirling in real time. In some instances, the fluid may be so echodense that it resembles soft tissue. In these cases, Doppler examination of the lesion will identify no blood flow, thus helping to rule out a soft-tissue lesion such as neoplasia. Fibrin tags and adhesions may be visible around the testicle and a hydrocele may be present. If the abscess ruptures, pyocele may result. The ultrasonographic character of the surrounding testicular parenchyma may be altered as a result of pressure from the abscess and a secondary orchitis. Figure 151.9 shows the ultrasonographic appearance of a well-defined, localized, abscess just under the surface of a testicle in a stallion.
Testicular abscess
Testicular hematoma
Testicular abscesses can arise as a result of a penetrating wound through the scrotum and testicle, testicular biopsy, a progression of orchitis, hematogenous spread, or a descending infection from peritonitis. Stallions may present as febrile with a unilaterally
Testicular hematomas can form either within the parenchyma or on the testicular surface and can arise from trauma or a penetrating wound or biopsy.11 Although small hematomas may not result in any noticeable scrotal enlargement, more sizeable
The ultrasonographic appearance of orchitis can be highly variable. The testicular parenchyma usually develops a heterogeneous or even granular appearance. Echogenicity of the parenchyma may be increased or decreased compared to normal. In cases where focal lesions are imaged, it may be difficult ultrasonographically to distinguish orchitis from neoplasia. Figure 151.8 shows one possible ultrasonographic appearance of orchitis.
Vascular and lymphatic stasis
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Figure 151.9 Ultrasonogram of a well-defined, localized, abscess just under the surface of a testicle. The appearance of this lesion also is consistent with a small testicular tumor. However, when the lesion was examined with Doppler ultrasonography, there was no evidence of blood flow, suggesting that it was comprised of fluid, rather than soft tissue. The testicular parenchyma itself is abnormally heterogeneous and is suspicious of a secondary orchitis. The calipers labeled ‘1’ delineate the abscess. The calipers labeled ‘2’ delineate the testis.
hematomas will. The stallion may demonstrate pain on palpation of the testicle. Ultrasonographically, acute testicular hematomas appear mottled grayish black to white, depending on the degree of consolidation (Figure 151.10). If hemorrhage is ongoing, large pockets of relatively hypoechoic, unclotted blood may also be seen swirling in real time. As the hematoma organizes, its ultrasonographic appearance becomes more echogenic, eventually appearing hyperechoic relative to the surrounding testicle. Fibrin tags and adhesions may also form in the affected area. Neoplasia Testicular neoplasia is uncommon in the stallion12 and, because most testicular tumors are relatively small, affected testes often are not noticeably enlarged. In fact, in some cases of testicular neoplasia, testicular degeneration is also present and the affected testicle may even be reduced in size. Testicular tumors can arise from either germinal or non-germinal cells. Germinal neoplasms are the most common tumor type found in stallions. Teratomas, seminomas, teratocarcinomas, and embryonic carcinomas all have been described in stallions.13, 14 Non-germinal tumors arise from stromal cells and include Sertoli and Leydig cell tumors.15 Mixed germ cell-sex cord-stromal tumors also have been
Figure 151.10 Ultrasonogram of a fresh testicular hematoma. The hematoma is heterogeneous and mottled, similar in appearance to a fresh corpus hemorrhagicum on a mare’s ovary. The testis itself is not visible in this image.
reported.16–18 Most, but not all, testicular tumors in the stallion are benign. Although ultrasonography may alert the clinician to the possible existence of a testicular tumor, at this time it is not possible to ultrasonographically differentiate the different tumor types. Tentative diagnosis of testicular neoplasia can be made based on ultrasonography; definitive diagnosis requires histopathological examination. Testicular neoplasia results in a localized heterogeneous appearance to the normally very homogeneous testicular parenchyma. It can sometimes be difficult to determine whether or not a focal lesion in the testis is soft tissue (as in the case of a neoplasia) or fluid (as in the case of a testicular abscess). Doppler ultrasonography is useful in these cases. By placing the color Doppler window over the lesion, one can determine if blood flow is present (indicating a soft-tissue lesion) or absent (suggesting fluid). Figure 151.11 illustrates one possible ultrasonographic appearance of testicular neoplasia and demonstrates the use of Doppler ultrasonography to determine if blood flow is present within the lesions. Although pictured here, multicentric
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T
T
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Figure 151.11 Ultrasonogram of multicentric testicular neoplasia in a stallion (A). Note the heterogeneity of the testicular parenchyma. Several fairly localized lesions are apparent (denoted by ‘T’ in the approximate center of each image). In panel B, Doppler ultrasonography identifies blood flow in one of the lesions, confirming it as a soft-tissue mass. The white rectangle is the Doppler window, placed to include one of the lesions and also a portion of the more normal surrounding parenchyma. The bright white dots are Doppler signals labeling cross-sections of vessels that contain active blood flow. These dots would appear blue or red in a full color image. (See color plate 47).
testicular neoplasia is uncommon. Most testicular neoplasms are single lesions; however, seminomas often appear to have multiple nodules.
at least once each year so that trends in testicular size can be determined. More frequent measurements are advisable when possible.
Testicular degeneration
Testicular cysts
Testicular degeneration (TD) is a common cause of acquired and often progressive infertility in stallions.19, 20 Although TD can arise secondarily to a known testicular insult, more commonly no underlying cause for the degeneration can be identified. This type of degeneration, also called idiopathic TD (ITD),21 is most often seen in middle-aged or older stallions, although less frequently it can occur in younger animals. Regardless of the age of onset, ITD is typically progressive and results in a steady decline in testicular size, semen quality, and subsequent fertility, sometimes ending in infertility. The ultrasonographic appearance of the testicular parenchyma in cases of ITD usually is no different from that of a normal testicle. The value of ultrasonography in evaluating animals with ITD is in the ability to reliably and regularly obtain measurements of the testicles. Measurements of length, width and height should be obtained and used to calculate total testicular volume. Any trends suggestive of ITD then can be identified early (e.g., decreasing testicular size and/or volume). ITD might be suspected if a stallion is producing low sperm numbers for his testicular volume.22 It is recommended that the testicles of breeding stallions be measured ultrasonographically
Structures that are ultrasonographically consistent with testicular cysts sometimes are found in stallions’ testes. In the author’s experience, these are usually found incidentally in fertile animals, e.g., during routine breeding soundness examinations. Thus, it appears that at least small testicular cysts do not adversely impact fertility. Ultrasonographically, these lesions appear as well-delineated, circular or oval, anechoic regions within the surrounding normal testicular parenchyma (Figure 151.12). This appearance closely resembles that described for simple testicular cysts in humans. In humans, true simple testicular cysts are rare and are considered benign.23, 24 In stallions, it can be easy to confuse these cysts with the central vein of the testis. However, Doppler ultrasonography will confirm the diagnosis, as there is no blood flow present within the cyst. Scrotal fluid: hydrocele, hematocele, and pyocele Although there are many possible causes for scrotal enlargement (see above), most cases of increased scrotal size are extratesticular in origin and are caused by an increase in volume in the vaginal space.
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Hydrocele
Testis Spermatic cord
Figure 151.13 Ultrasonographic image of a significant hydrocele outlining the spermatic cord and a portion of the testis.
Figure 151.12 Ultrasonogram of a testicular cyst in a fertile stallion. The cyst appears as a well-defined, roughly circular area within the testicular parenchyma and resembles the central vein of the testis.
Hydrocele Hydrocele is an increase in volume of the anechoic fluid that normally is present in the vaginal cavity surrounding the testis. Normally, the depth of fluid in the vaginal cavity is < 5 mm, with slightly larger pockets sometimes present around the tail of the epididymis. Since the vaginal cavity is continuous with the peritoneal cavity, any systemic disease that causes either increased production of peritoneal fluid or decreased drainage of peritoneal fluid also can result in hydrocele. In addition, trauma, testicular abscesses, orchitis, testicular torsion, neoplasia, or systemic disease resulting in dependent edema and/or pyrexia can result in hydrocele. Hydrocele can also be seen congenitally, most often in Shire horses. Congenital hydrocele is usually not detrimental to the stallion.25 Ultrasonographically, the increase in fluid volume that is seen in cases of hydrocele creates a more distinct ultrasonographic outline of the testis, spermatic cord and epididymis (Figure 151.13).
a corpus hemorrhagicum on a mare’s ovary (grayblack, mottled, irregular; Figure 151.15). The clot gradually increases in echogenicity as it organizes and fibroses. Hematocele can result from trauma, spermatic cord torsion, or secondarily to testicular biopsy. Purulent fluid generally has large amounts of particulate debris floating within it and, ultrasonographically, can be difficult to differentiate from hematocele. Pyocele can develop following a penetrating wound, secondarily to rupture of a testicular abscess or as a result of peritonitis. In cases of hematocele and pyocele, fibrin tags may form and may be visible ultrasonographically as grayish structures floating or waving in the surrounding fluid. Ultrasonography
Hematocele
Fibrin Tag
Testicle
Hematocele and pyocele Blood (hematocele) or purulent fluid (pyocele) can also be visualized ultrasonographically within the scrotum. Unclotted blood appears as heterogeneous fluid within the vaginal cavity (Figure 151.14). Clotted blood (hematoma) has an appearance similar to
Figure 151.14 Ultrasonographic image of a fresh hematocele. The fluid in the vaginal space is unclotted blood and is more echogenic and heterogeneous than is seen in cases of hydrocele. A fibrin tag is bisecting the fluid pocket.
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Figure 151.15 Ultrasonogram of a hematoma adjacent to the testicle. The injury was sustained as a result of a testicular biopsy that had been taken approximately 2 weeks earlier. Note the irregular edges of the hematoma and its heterogeneous appearance compared to the testicle.
can be a valuable tool for monitoring the progress of affected stallions and is often of prognostic value when used to assess the extent of the pathology. Inguinal and scrotal hernias An inguinal hernia occurs when a loop of bowel passes into the inguinal canal. A scrotal hernia is a progression of an inguinal hernia in which the bowel loop passes into the scrotum. The two terms are often used interchangeably. Inguinal or scrotal hernias may be congenital or acquired, uncomplicated (nonstrangulated) or complicated (strangulated). Acquired inguinal and scrotal hernias can occur in geldings or in stallions and may occur with a higher incidence in Standardbreds.26 Larger inguinal rings may predispose a horse to an acquired hernia.27 In some cases, acquired inguinal and scrotal hernias are associated with exercise or trauma.28 Most acquired hernias in horses are inguinal ruptures in which the intestine protrudes through a rent in the peritoneum and transverse fascia adjacent to the vaginal ring. As a result, the intestine becomes located immediately subcutaneously.29, 30 Small intestine is usually involved and most horses are presented for an enlarged scrotum, colic, or both.31 Ultrasonography can greatly aid in the examination of congenital and acquired inguinal or scrotal
0.47
CM
Figure 151.16 Ultrasonogram of a scrotal hernia. Bowel loops (labeled ‘gut’) are present in the scrotum next to the spermatic cord (labeled ‘cord’). The bowel wall thickness is slightly increased (delineated by calipers. It measures 0.47 cm). This bowel was edematous and compromised, but still viable when it was visualized during surgery.
hernias and in assessing bowel viability.32, 33 A 7.5 or 5.0 MHz linear array or sector scanner transducer can be used to evaluate scrotal contents or can be pressed high into the inguinal region to evaluate the external inguinal ring. When present, bowel loops are usually readily identifiable (Figure 151.16). In the case of scrotal hernias, hydrocele may also be present. If the hydrocele is severe, it can sometimes be difficult to find bowel loops when they are present. Ultrasonographic hallmarks of viable bowel include peristaltic waves and normal wall thickness. In cases of strangulating hernias, peristaltic waves are reduced or absent and the bowel wall appears thick and edematous. The type of bowel present in the hernia can also be determined using ultrasonography.31 Small intestine, as is usually present, can be identified by its small diameter and long mesentery.34 Epididymitis Most cases of epididymitis are bacterial in origin. The ultrasonographic appearance of an affected epididymis varies but, generally, increased frequency and size of hypoechoic regions within the epididymis are seen.35–37 These are presumably associated with accumulation of exudate and abscess formation. Scattered areas of increased echogenicity also may be seen and in some cases the epididymis appears consolidated, with loss of the normal hypoechoic lumen cross-sections (Figure 151.17). On physical examination, the epididymis is usually enlarged. This may be identified incidentally in non-breeding animals. In breeding stallions, reduced semen quality and white cells in the ejaculate may be detected. Some stallions may exhibit signs of
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Figure 151.17 (a) Ultrasonographic image of a normal epididymal body showing the typical pinpoint, anechoic cross-sections of the epididymal duct. (b) Ultrasonographic image consistent with epididymitis affecting the epididymal body. Note the two, large and irregular fluid-filled pockets and the loss of the normal surrounding architecture. White ovals delineate the approximate outlines of the two epididymides. Image courtesy of Dr Shana Chase.
discomfort during breeding or, alternatively, may present with signs of colic or lameness. Epididymal neoplasia Neoplasia of the epididymis is rare, but has been reported. The neoplastic epididymis reportedly had ultrasonographic characteristics similar to those of chronic epididymitis (heterogeneous appearance of both hypo- and hyperechoic areas, sometimes lobulated).38 A definitive diagnosis of epididymal neoplasia can be made following histopathological examination of the epididymis after orchidoepididymectomy. In cases where epididymal neoplasia is suspected, examination of the affected area with Doppler ultrasonography may help to confirm the presence of a solid soft tissue mass (e.g., neoplasia) by the presence of blood flow within the mass. Although ultrasonography is helpful in identifying epididymal pathology, it is unlikely that a definitive diagnosis can be made based on ultrasonographic examination alone. Histopathological examination of the epididymis following castration is usually required. Examination of ejaculated semen may help differentiate infectious epididymitis from sperm granuloma and neoplasia. Spermatic cord torsion Torsion of the spermatic cord may or may not result in vascular compromise to the testis. In the absence of vascular compromise, the condition may be entirely asymptomatic and non-pathogenic. However, when blood and/or lymphatic flow is compromised, clinical signs are usually apparent. Affected stallions classically present with signs of colic associated with an enlarged, painful testicle, enlarged scro-
tum, increased scrotal fluid, and an enlarged spermatic cord. In both symptomatic and asymptomatic cases of spermatic cord torsion, the torsion may be intermittent. Palpation and/or ultrasonographic identificaton of the location of the tail of the epididymis has been recommended as one means of identifying torsion of the spermatic cord. Normally, the epididymal tail should be located caudally. However, in cases of 180◦ torsion of the spermatic cord, the epididymal tail will be located cranially. Although the epididymal tail can be readily palpated or identified ultrasonographically in animals with asymptomatic cord torsions, it may be more difficult to identify in symptomatic cases where the scrotum is painful and significantly swollen.1 In cases of spermatic cord torsion in which the cord is compromised, ultrasonographic evaluation of the scrotal contents typically reveals dilated vessels in the spermatic cord and/or testicle and changes in the echogenicty of the cord and the testicular parenchyma (Figure 151.18). Either an increase or decrease in echogenicity may be seen. In addition, hydrocele is often present. Ultrasonographic evaluation of the testicular artery, arcuate arteries, central vein, and pampiniform plexus may allow the veterinarian to determine the presence or absence of blood flow to and from the testicle.39 If Doppler capability is available, it can be used to provide additional information on the presence or absence of blood flow in testicular vessels.
Varicocele Varicocele (abnormal dilation of the veins of the spermatic cord) is poorly documented and/or
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Figure 151.18 (a) Ultrasonographic appearance of a normal spermatic cord and (b) chronic spermatic cord torsion that resulted in a severely compromised testis. The stallion presented for pain and an enlarged scrotum. In this case, there are areas of increased echogenicity within the affected cord, suggesting that venous stasis has led to thrombus formation. The normal cross-sections of the vessels as seen in the left-hand image are not present in the right-hand image. Doppler examination of the abnormal cord revealed little blood flow. The white ovals approximately delineate the borders of the two spermatic cords. This is one possible way that cord torsion may manifest itself, but there are other ultrasonographic appearances that might be seen as well (e.g., dilated vessels, enlarged cord, edema).
uncommon in stallions.1 The effect of varicocele on stallion fertility is not known. It has been suggested that varicocele may cause a slight to moderate decrease in semen quality by interfering with thermoregulation of the testis.40 However, this has not yet been proven in the horse.41 In contrast, varicocele is common in men and there is evidence to support its association with decreased fertility.42–45 Experimentally induced varicocele also has been shown to affect testicular thermoregulation and cause a decrease in semen quality in dogs and monkeys.46, 47 In humans the disease is considered progressive and surgical correction generally is recommended for affected individuals with poor semen quality or abnormal testicular parameters.48–51 Additional studies will be needed to determine the effects, if any, of varicocele on stallion fertility. Ultrasonographically, varicocele in the stallion appears as well-circumscribed, heterogeneous anechoic regions within the spermatic cord. This is in contrast to the appearance of the normal spermatic cord in which the anechoic regions are typically homogeneous in size. In addition, it has been suggested that distention of the central vein in the ipsilateral testis may accompany varicocele in the stallion.1
Ultrasonographic imaging of the accessory sex glands Examination of the accessory sex glands of the stallion is performed per rectum. Stallions can be re-
strained and prepared for this examination similarly to mares. Stocks are preferable, since most stallions are not accustomed to this procedure. However, tractable animals also can be safely examined without stocks, using the same routine safety precautions that one would use with a mare. Sedation should be considered as most stallions are not used to and resent examination per rectum. Due to the superficial location of the accessory glands, a 7.5 MHz or higher frequency linear array transducer is preferred. If a 7.5 MHz transducer is not available, a 5 MHz transducer can be used, although some detail will be lost. The transducer is held, introduced, and advanced through the rectum as it is for ultrasonography of the mare. If a linear array transducer is not available, a microconvex transducer or a sector scanner with an off-line beam may suffice. The larger size of the sector scanner can make examination per rectum difficult and more dangerous and may preclude transrectal ultrasonographic examination in ponies or miniature horse stallions.
Ampullae The ampullae can be imaged individually in saggital sections with the transducer placed directly over each gland (Figure 151.19, A). With the transducer in this position, the ampullae are imaged as soft-tissue bands that extend from left to right across the upper edge of the ultrasound screen. The lumen of the ampulla may be visible as a thin black (fluid) or white
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Bladder
A
Ampulla
B C Seminal Vesicle
D
Prostatic Lobe Prostatic lsthmus Bulbourethral Gland E
Figure 151.19 Methods for positioning the transducer to image the accessory sex glands per rectum. (a) Position of the transducer for imaging the left ampulla in sagittal section. (b) Position of the transducer for imaging the left and right ampullae in short-axis cross-sections. (c) Position of the transducer for imaging the left seminal vesicle in sagittal section. (d) Position of the transducer for imaging the left lobe of the prostate gland in sagittal section. (e) Position of the transducer for imaging the left bulbourethral gland in sagittal section.
(no fluid present) line through the approximate center of the gland (Figure 151.20). In normal stallions, the lumen and total diameter of the ampulla will increase after sexual stimulation and decrease following ejaculation.55 Table 151.1 lists normal sizes for accessory glands.56 The ampullae can also be visualized in crosssection by turning the transducer at a 90◦ angle to the long axis of the glands (Figure 151.19, B). This view can be difficult or impossible to obtain with larger transducers or in smaller stallions as the pelvic diameter may limit proper positioning of the probe. In short-axis cross-section, both ampullae can be followed to their entrance to the urethra at the seminal colliculus52 (Figure 151.21). The pelvic urethra becomes visible in cross-section ventral and medial to the ampullae as the glands are followed caudally and approach the midline.53 When a higher resolution probe is employed, it may be possible to image the uterus masculinus in some animals. This structure is a remnant of the embryological paramesonephric (mullerian) ducts and, when present, is found medial to the ampullae just cranial to the prostatic isthmus.54 Ultrasonographically, the uterus masculinus appears as
Figure 151.20 Ultrasonogram of a normal ampulla in sagittal section. The image was obtained with the transducer held as in Figure 151.19A. The dashed black lines delineate the approximate edges of the gland. The ampullary lumen is not clearly visible in this image.
a thin-walled, oblong structure filled with anechoic fluid. Seminal vesicles The seminal vesicles are imaged by moving the ultrasound transducer slightly laterally and caudally to the ampullae (Figure 151.19, C). Similar to the ampullae, the seminal vesicles also can be imaged in cross-section by rotating the transducer 90◦ (Figure 151.21). When the stallion is at sexual rest and the glands are empty, the seminal vesicles appear as flattened, somewhat wavy, soft-tissue structures. Varying amounts of anechoic fluid may be present within the glandular lumen. As the fluid increases, the gland expands into a bladder-like shape (Figure 151.22). With increased sexual stimulation, the volume of intralumenal fluid will increase. In highly teased stallions, the glands can become quite large and appear as thin-walled, bladder-like structures containing large volumes of anechoic fluid. This fluid is usually, but not always, expressed as gel during ejaculation.55 See Table 151.1 for normal sizes.56
Prostatic isthmus and prostatic lobes The prostatic isthmus is located on the ventral midline of the pelvis, just caudal to the seminal colliculus. The lobes of the prostate extend outward to the right and left of the isthmus. The prostatic isthmus is imaged on the animal’s midline by moving the transducer caudally from the seminal vesicles, parallel to the long axis of the stallion’s body. The
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Table 151.1 Dimensions (mm; mean and range) of accessory sex glands, ampullae, and urethra in 102 stallions of various size types
Bulbourethral gland Length Height Prostate Lobe thickness Isthmus thickness Vesicular gland Total diameter Wall diameter Lumen diameter Ampulla Total diameter Wall diameter Lumen diameter Urethra Total diameter
Miniature horse (n = 7)
Pony (n = 27)
Light horse (n = 53)
Heavy horse (n = 15)
21a (16–27) 14a (11–19)
28b (21–39) 17a (10–24)
38c (17–59) 24b (12–39)
35c (30–44) 22b (16–33)
18a (11–29) 8a (4–11)
20a (14–27) 7a,b (2–16)
25b (10–40) 10b (4–25)
27b (18–39) 11b (5–31)
9a (5–15) 3a (1–6) 4a (2–11)
12a (3–30) 4a (1–10) 6a (1–26)
12a (4–20) 4a (1–8) 5a (0–11)
16b (8–43) 3a (1–4) 11b (2–39)
10a,b (5–20) 4a (2–10) 1a (0–3)
8a (4–12) 3a (1–5) 1a (0–2)
11b,c (4–18) 5b (2–8) 2a (0–6)
13c (6–17) 5b (3–7) 2a (0–4)
15a (10–20)
19b (13–24)
26c (14–35)
30d (24–38)
Values with different superscripts across different rows (a–d) differ significantly (P < 0.05). Reprinted from Pozor M, McDonnell SM.56 Ultrasonographic measurements of accessory sex glands, ampullae, and urethra of normal stallions of various size types. Theriogenology 2002;58(7):1425–33. Reprinted with permission from Elsevier.
prostatic lobes are viewed by moving the transducer laterally to the left and right of midline (Figure 151.19, D). In the absence of sexual stimulation, the prostatic parenchyma appears gray-white and heterogeneous, containing numerous, small, anechoic fluid spaces (Figure 151.23). The overall size of the prostate and size and number of the fluid-filled spaces increase following sexual stimulation.55 In a highly teased animal, the appearance of the gland can be quite dramatic with large, radiating anechoic fluid spokes extending throughout the parenchyma. See Table 151.1 for normal sizes.56
Bulbourethral glands The bulbourethral glands can be imaged caudal to the prostate to the right and left of the ventral midline (Figure 151.19, E). These glands are located just inside the anus. Thus, to properly place the transducer over these glands, the wrist and palm of the
E
B
B
A
A
Figure 151.21 Ultrasonogram of (A) normal ampullae and (B) normal fluid-containing seminal vesicles in short-axis crosssection near the region where the glands enter the seminal colliculus.
Figure 151.22 Ultrasonogram of a normal seminal vesicle in sagittal section. The image was obtained with the transducer held as in Figure 151.19C. The dashed black line delineates the approximate edges of the gland. This seminal vesicle contains a moderate amount of anechoic fluid. It also can be normal for the seminal vesicles to contain no fluid (in which case they can be difficult to identify ultrasonographically) or large amounts of fluid (if the stallion is highly teased).
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Abnormalities of the accessory glands Pathology of the accessory sex glands of the stallion is uncommon. Becoming familiar with the normal ultrasonographic appearance of the accessory glands may allow one to identify pathology should it occur. Routine evaluation of the accessory glands also has been recommended as part of the stallion fertility evaluation.4 Sperm accumulation within or occlusion of the ampullae Figure 151.23 Ultrasonogram of a normal prostatic lobe in sagittal section. The image was obtained with the transducer held as in Figure 151.19D. The dashed black line delineates the approximate edges of the gland. In this image, the stallion was not highly teased and the anechoic, fluid-filled spaces in the prostate are pinpoint. These spaces enlarge after teasing.
veterinarian’s hand must be withdrawn from the rectum leaving only the fingers holding the ultrasound transducer just within the anal sphincter. The bulbourethral glands are circular to oval soft-tissue structures (Figure 151.24). After sexual stimulation, the glands enlarge and small anechoic fluid-filled spaces may be visible within the parenchyma.55 See Table 151.1 for normal sizes.56
Figure 151.24 Ultrasonogram of a normal bulbourethral gland from a stallion in sagittal section. The image was obtained with the transducer held as in Figure 151.19E. The dashed black oval delineates the approximate edges of the gland. This stallion was not teased so the potential fluid-filled spaces in the gland are not apparent.
The most common pathologies of the ampullae in the stallion are sperm accumulation and sperm occlusion.57 Sperm occlusion results in a mechanical blockage that prevents testicular sperm from contributing to the ejaculate. When the condition is bilateral, affected horses are rendered azoospermic. Sperm accumulation seems likely to be a less severe variation of sperm occlusion. In these cases, a complete blockage is not present and sperm still appear in the ejaculate. However, because many or all of the ejaculated sperm have been trapped within the ampullae for extended periods (and thus exposed to body temperature), most or all of the ejaculated sperm are immotile and non-viable. The classic presentation also includes a high percentage of tailless heads in ejaculated semen. However, this is not found in all cases. Additionally, because accumulated sperm apparently are released in an inconsistent manner, ejaculates often contain highly variable and unpredictable total numbers of sperm. The ultrasonographic appearance of spermoccluded ampullae is highly variable and can be normal. When present, ultrasonographic abnormalities include abnormal amounts of echodense material (inspissated sperm) or echogenic or anechoic fluid within the ampullary lumen.58 Additionally, the glandular portion of the ampullae may appear more echodense and may contain hyperechoic spots throughout the parenchyma, again probably representing accumulated sperm (Figure 151.25). On the other hand, a dilated ampullary lumen in and of itself is not enough to make a diagnosis of sperm blockage. In normal, teased stallions, the lumen of the ampulla can get very large. In these cases, the diameter of the lumen should decrease following ejaculation when the gland is emptied of its contents. Because of the variable ultrasonographic appearances of affected ampullae, ultrasonography alone often cannot be used to definitively rule in or rule out this diagnosis. Sperm occlusion or accumulation should be suspected in stallions presenting with
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Figure 151.25 One potential ultrasonographic presentation of sperm occlusion of the ampulla. The ampulla is abnormally dilated and the edges of the lumen appeared echodense. A hyperechoic area (the blockage, presumably) was identified at the end of the dilatation (not visible in this picture). The edges of the gland are delineated with dashed lines. The arrow indicates the dilated lumen.
apparently normal testicles but very low percentages of motile/viable sperm or azoospermia. Seminal plasma alkaline phosphatase levels can be helpful in differentiating sperm occlusion from testicular origin azoospermia.59, 60 Obstruction of the seminal colliculus In addition to obstructive azoospermia associated with occluded ampullae, there is one report of obstructive azoospermia associated with a cystic mullerian duct remnant located at the seminal colliculus. This remnant was identified ultrasonographically as a hypoechoic structure located within the pelvic urethra at the level of the seminal colliculus.61 Seminal vesiculitis Seminal vesiculitis is reported uncommonly in stallions.62–64 Affected animals typically present for poor semen quality and/or subfertility, classically associated with a high percentage of neutrophils in the ejaculate. In some cases, stallions may present with signs of colic or pain during ejaculation that can lead to ejaculation failure.65 Most cases are suspected from gross observation of pus and debris associated with the gel of an ejaculate; however, microscopic examination of ejaculated semen often reveals all or any of the following: numerous neutrophils, bacteria (sometimes intracellular), red blood cells, and reduced sperm motility. These findings should prompt further examination of the stallion’s genitalia to determine the source of the problem. Palpation per rectum of affected glands may reveal no abnormalities. However, in some cases, the affected gland is enlarged, painful, and filled with fluid. Taken together with other clinical signs, ultrasonography can be very helpful in making a diag-
Figure 151.26 One potential ultrasonographic presentation of seminal vesiculitis. The seminal vesicles are imaged in shortaxis cross-section. The normal vesicle is on the left (A). Note the large, and notably asymmetric size of the abnormal, right vesicle and the relative echodensity of its contents (B). A small portion of the normal prostate also is visible in this image (C).
nosis of seminal vesiculitis.66, 67 Ultrasonographically, fluid within the affected gland(s) can vary from anechoic to relatively echogenic and often will contain particulate debris or fibrin tags (Figure 151.26). The lumen of the gland may be irregular and the luminal edges of the glandular tissue may be hyperechoic. In cases of chronic seminal vesiculitis, affected glands can become firm and often are almost devoid of fluid. Seminal vesicles in sexually stimulated stallions can normally become significantly enlarged and filled with anechoic fluid and thus it is important to be familiar with the normal appearance of seminal vesicles in both teased stallions and stallions following ejaculation in order to be able to identify pathology when present. Prostatic neoplasia Prostatic neoplasia is exceedingly rare. In the author’s limited experience (two cases: one stallion, one gelding), the tumors have been readily palpable on examination of the prostate per rectum. Ultrasonographically, the normally organized appearance of the prostate is lost and is largely replaced by the invasive mass (Figure 151.27). In the one case where histopathology was performed, the mass was confirmed to be a carcinoma. Bulbourethral gland cysts Cysts of the bulbourethral glands appear to have no adverse effects. Bulbourethral gland cysts may be identified incidentally on ultrasonographic examination of the accessory glands during a breeding soundness examination. Ultrasonographically, cysts
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Figure 151.27 Ultrasonogram of a prostatic tumor in an aged gelding. Note the heterogeneous appearance of the tumor compared to the organized appearance of the normal prostate in Figure 151.23. The fluid-filled spaces of the prostate appear to have been replaced by soft tissue in the abnormal gland. Note also the large, cystic cavity to the right (a). Ultrasonographic calipers delineate the height of the tumor.
of the bulbourethral glands appear as localized, circular, anechoic fluid-filled areas within the gland parenchyma.
Terminal aorta and iliac arteries The termination of the aorta and its branching into the internal and external iliac arteries occurs beneath the sacrum along the dorsal midline. During transrectal ultrasonographic examination of the accessory sex glands, the terminal aorta and iliac arteries are easily visualized by directing the beam of the transducer dorsally along the midline and advancing the transducer cranial to the pelvis. Aortic-iliac thrombosis (AIT) occurs in horses infrequently. In stallions, this condition has been associated with ejaculatory failure and weakness in the hindlimbs due to vascular compromise, disturbance of innervation and/or pain.68 The stallion may also show signs of pain in the hindlimbs, often becoming worse following exercise, and presumably in association with muscle hypoxia. On physical examination, the hindlimb on the affected side may be cool to the touch. In cases of AIT, ultrasonography can be used in the formulation both of a diagnosis and a prognosis.69, 70 Ultrasonography permits visualization and measurement of the thrombus and can aid in the assessment of blood flow through the iliac arteries to the hindlimbs and penis. The presence of a thrombus typically is discernible ultrasonographically. Normally, the walls of the vessels are smooth
Figure 151.28 Ultrasonogram of an aortic thrombus occupying approximately 75% of the lumen of the vessel. The aorta is seen in short-axis cross-section and is delineated by the dashed black line. The white dashed line outlines the thrombus. The thrombus projects into the lumen and partially blocks the flow of blood.
and blood flows steadily and smoothly within the lumen. When present, the gray-white thrombus can be seen extending from the wall of the artery into the lumen of the vessel where it interferes with the normally smooth flow of blood (Figure 151.28). If properly managed, stallions with ejaculatory dysfunction and/or weakness associated with AIT can return to breeding. Successful management involves pain control with phenylbutazone, appropriate sexual stimulation, and controlled exercise to encourage development of blood flow to the extremities.68 Pharmacologically induced ejaculation can also be attempted.
References 1. Love CC, Varner DD. Ultrasonography of the scrotal contents and penis of the stallion. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Williams, 1998; pp. 253–69. 2. Love CC, Garcia MC, Riera FR, Kenney RM. Evaluation of measures taken by ultrasonography and caliper to estimate testicular volume and predict daily sperm output in the stallion. J Reprod Fertil Suppl 1991;44:99–105. 3. Amann RP. Functional anatomy of the adult male. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 645–57. 4. Kenney RM, Hurtgen JP, Pierson RH, Witherspoon D, Simons J. Clinical fertility evaluation of the stallion. Theriogenology 1983;9 (issue): 7–62. 5. Weber J, Woods G. A technique of transrectal ultrasonography of stallions during ejaculation. Theriogenology 1991;36:831–6. 6. Pozor MA, McDonnell S. Doppler ultrasound measures of testicular blood flow in stallions. Theriogenology 2002;58:437–40. 7. Pozor MA, McDonnell S. Color doppler ultrasound evaluation of testicular blood flow in stallions. Theriogenology 2004;61:799–810.
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8. Jann HW, Rains JR. Diagnostic ultrasonography for evaluation of cryptorchidism in horses. J Am Vet Med Assoc 1990;196:297–300. 9. McKinnon AO. Reproductive ultrasonography. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Hoboken, NJ: Blackwell, 1998; pp. 79–102. 10. DeVries PJ. Diseases of the testes, penis and related structures. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; p. 878–84. 11. DelVento, VR, Amann RP, Trotter GW, Veeramachaneni DN, Squires EL. Ultrasonographic and quantitative histologic assessment of sequelae to testicular biopsy in stallions. Am J Vet Res 1992;53:2094–101. 12. Moulton JE. Tumors of the genital system. In: Moulton JE (ed.) Tumors in Domestic Animals. Berkeley, CA: Davis Press, 1978, p. 309. 13. Beck C, Charles JA, Maclean AA. Ultrasound appearance of an equine testicular seminoma. Vet Radiol Ultrasound 2001;42:355–7. 14. Ladd PW. The male genital system. In: Jubb KVF, Kennedy PC, Palmer N (eds) Pathology of Domestic Animals. New York: Academic Press, 1985; pp. 409–59. 15. Caron JP, Barber SM, Bailey JV. Equine testicular neoplasia. Comp Contin Educ Pract Vet 1985;7:53–9. 16. Brito L, Engiles J, Turner RM, Getman L, Ebling A. Bilateral testicular mixed germ cell-sex cord-stromal tumors in a stallion. Reprod Domest Anim 2009;44:846–51. 17. Cullen JM, Whiteside J, Umstead JA, Whitacre MD. A mixed germ cell-sex cord-stromal neoplasm of the testis in a stallion. Vet Pathol 1987;24:575–7. 18. Zanghi A, Catone G, Marino G, De Vico G, Nicotina PA. Malignant mixed sex cord-stromal tumour in a stallion. Reprod Domest Anim 2004;39:376–9. 19. Blanchard TL, Varner DD, Bretzlaff KN, Elmore RG. Testicular degeneration in large animals: identification and treatment. Vet Med 1991;86:537–42. 20. Turner RM. Pathogenesis, diagnosis and management of testicular degeneration in stallions. Curr Tech Equine Pract 2007;6:278–84. 21. Blanchard T, Johnson L, Roser AJ. Increased germ cell loss rates and poor semen quality in stallions with idiopathic testicular degeneration. J Equine Vet Sci 2000;20: 263–5. 22. Blanchard TL, Johnson L, Varner D, Rigby S, Brinsko S, Love CC, Miller C. Low daily sperm output per ml of testis as a diagnostic criteria for testicular degeneration in stallions. J Equine Vet Sci 2001;21:11–35. 23. Rifkin MD, Jacobs JA. Simple testicular cyst diagnosed preoperatively by ultrasound. J Urol 1983;129:982–3. 24. Tosi SE, Richardson JR Jr. Simple cyst of the testis: case report and review of the literature. J Urol 1975;114:473–5. 25. Cox JE. Developmental abnormalities of the male reproductive tract. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 895–904. 26. Sembrat RF. The acute abdomen in the horse: epidemiologic considerations. Arch Am Coll Vet Surg 1975;4:34–8. 27. Ashdown RR. The anatomy of the inguinal canal in the domesticated mammals. Vet Rec 1963;75:1345–51. 28. Vasey JR. Simultaneous presence of a direct and an indirect inguinal hernia in a stallion. Aust Vet J 1981;57:418–21. 29. Cox JE. Hernias and ruptures: words to the heat of deeds. Equine Vet J 1988;20:155–6. 30. Schumacher J. Testis. In: Auer JA, Stick JA (eds) Equine Surgery. St Louis, MO: Saunders Elsevier, 2006; pp. 775–810.
31. Schneider RK, Milne DW, Kohn CW. Acquired inguinal hernia in the horse: a review of 27 cases. J Am Vet Med Assoc 1982;180:317–20. 32. Reef VB. Equine pediatric ultrasonography. Comp Contin Educ Pract Vet 1991;13:1277–95. 33. Reef VB. The use of diagnostic ultrasound in the horse. Ultrasound Quart 1991;9:1–34. 34. Reef VB. Adult abdominal ultrasonography. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia: Saunders, 1998; pp. 273–363. 35. Brinsko SP, Varner DD, Blanchard TL, Relford RL, Johnson L. Bilateral infectious epididymitis in a stallion. Equine Vet J 1992;24:325–8. 36. Held JP, Adair S, McGavin MD, Adams WH, Toal, R, Henton J. Bacterial epididymitis in two stallions. J Am Vet Med Assoc 1990;97:602–4. 37. Traub-Dargatz JL, Trotter GW, Kaser-Hotz B, Bennett DG, Kiper ML, Veeramachaneni DN, Squires E. Ultrasonographic detection of chronic epididymitis in a stallion. J Am Vet Med Assoc 1991;198:1417–20. 38. Held JP, McCracken MD, Toal R, Latimer F. Epididymal swelling attributable to generalized lymphosarcoma in a stallion. J Am Vet Med Assoc 1992;201:1913–15. 39. Love CC. Ultrasonographic evaluation of the testis, epididymis, and spermatic cord of the stallion. Vet Clin N Am Equine Pract 1992;8:167–82. 40. Roberts SJ. Veterinary Obstetrics and Genital Diseases: Theriogenology, 3rd edn. Woodstock, VT: SJ Roberts, 1986. 41. Turner RM. Ultrasonography of the genital tract of the stallion. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia: Saunders, 1998; pp. 446–79. 42. Gorelick JI, Goldstein M. Loss of fertility in men with varicocele. Fertil Steril 1993;59:613–16. 43. Green KF, Turner TT, Howards SS. Varicocele: reversal of the testicular blood flow and temperature effects by varicocele repair. J Urol 1984;131:1208–11. 44. Saypol DC, Howards SS, Turner TT, Miller ED Jr. Influence of surgically induced varicocele on testicular blood flow, temperature, and histology in adult rats and dogs. J Clin Invest 1981;68:39–45. 45. Witt MA, Lipshultz LI. Varicocele: a progressive or static lesion? Urology 1993;42:541–3. 46. Fussell EN, Lewis RW, Roberts JA, Harrison RM. Early ultrastructural findings in experimentally produced varicocele in the monkey testis. J Androl 1981;2:111–19. 47. Shafik A, Wali MA, Abdel-Azis YE, el-Kateb S, el-Sharkawy AG, Olfat E, Saleh M. Experimental model of varicocele. Eur Urol 1989;16:298–303. 48. Chehval MJ, Purcell MH. Deterioration of semen parameters over time in men with untreated varicocele: evidence of progressive testicular damage. Fertil Steril 1992;57:174–7. 49. Khera M, Lipshultz LI. Evolving approach to the varicocele. Urol Clin N Am 2008;35:183–9,viii. 50. Lipshultz LJ, Corriere JN. Progressive testicular atrophy in the varicocele patient. J Urol 1977;117:175–6. 51. Zorba UO, Sanli OM, Tezer M, Erdemir F, Shavakhabov S, Kadioglu, A. Effect of infertility duration on postvaricocelectomy sperm counts and pregnancy rates. Urology 2009;73:767–71. 52. Little TV, Woods GL. Ultrasonography of accessory sex glands in the stallion. J Reprod Fertil Suppl 1987;35:87–94. 53. Schmidt AR. Transrectal ultrasonography of the caudal portion of abdominal and pelvic cavities in horses. J Am Vet Med Assoc 1989;194:365–71.
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Ultrasonography of the Genital Tract 54. Little TV, Holyoak GR. Reproductive anatomy and physiology of the stallion. Vet Clin N Am Equine Pract 1992;8: 1–29. 55. Weber J, Woods G. Transrectal ultrasonography for the evaluation of stallion accessory sex glands. Vet Clin N Am Equine Pract 1992;8:183–90. 56. Pozor MA, McDonnell SM. Ultrasonographic measurements of accessory sex glands, ampullae, and urethra of normal stallions of various size types. Theriogenology 2002;58:1425–33. 57. Love CC. Examination of the male reproductive tract: evaluation of potential breeding soundness. In: Youngquist RS (ed.) Current Therapy in Large Animal Theriogenology. Philadelphia: Saunders, 1997; pp. 12–15. 58. Love CC, Riera FR, Oristaglio Turner RM, Kenney RM. Sperm occluded (plugged) ampullae in the stallion. In: Proceedings of the Annual Meeting of the Society for Theriogenology 1992; pp. 117–25. 59. Turner RM, McDonnell SM. Alkaline phosphatase in stallion semen: characterization and clinical applications. Theriogenology 2003;60:1–10. 60. Turner RM, Sertich PL. Use of alkaline phosphatase activity as a diagnostic tool in stallions with azoospermia and oligospermia. Third International Symposium on Stallion Reproduction, Fort Collins, Colorado, 2000; p. 35. 61. Little TV. Accessory sex gland and internal reproductive tract evaluation. In: Rantanen NW, McKinnon AO (eds)
62.
63.
64. 65.
66.
67.
68.
69. 70.
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Equine Diagnostic Ultrasonography. Baltimore: Williams & Williams, 1998; pp. 271–87. Blanchard TL, Varner DD, Hurtgen JP, Love CC, Cummings MR, Strezmienski PJ, Benson C, Kenney RM. Bilateral seminal vesiculitis and ampullitis in a stallion. J Am Vet Med Assoc 1988;192:525–6. Deegan E, Klug E, Lieske R, Freytag K, Martin J, Gunzel A-R. Surgical treatment of chronic purulent seminal vesiculitis in the stallion. Deut Tierarz Wochen 1979;86:140–4. Holst W. Seminal vesiculitis in two stallions. Tijdschrift Diergeneeskunde 1976;101:375–7. Freestone JF, Paccamonti DL, Eilts BE, McClure JJ, Swiderski CE, Causey RC. Seminal vesiculitis as a cause of signs of colic in a stallion. J Am Vet Med Assoc 1993;203:556–7. Malmgren L, Sussemilch BI. Ultrasonography as a diagnostic tool in a stallion with seminal vesiculitis: a case report. Theriogenology 1992;37:935–8. Varner DD, Blanchard TL, Brinsko SP, Love CC, Taylor TS, Johnson L. Techniques for evaluating selected reproductive disorders of stallions. Anim Reprod Sci 2000;60–61:493–509. McDonnell SM, Love CC, Martin BB, Reef VB, Kenney RM. Ejaculatory failure associated with aortic-iliac thrombosis in two stallions. J Am Vet Med Assoc 1992;200:954–7. Barrelet A. Aorto-iliac thrombosis in a breeding stallion and an eventer mare. Equine Vet Educ 1993;5:86–9. Tithof PK, Rebhun WC, Dietze AE. Ultrasonographic diagnosis of aorto-iliac thrombosis. Cornell Vet 1985;75:540–4.
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Acrosomal Function Stuart A. Meyers
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Evaluation of the Plasma Membrane Ben Colenbrander and Tom A.E. Stout
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Sperm Chromatin Structure Assay Aime K. Johnson and Charles C. Love
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Spermatozoal Viability Stuart A. Meyers
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Chapter 152
Acrosomal Function Stuart A. Meyers
The acrosome The acrosome is a single organelle of the sperm anterior head1 and is located between the plasma membrane and the nuclear envelope (Figures 102.2 and 102.5). Acrosomes have a unique set of membranes:1 (i) the inner acrosomal membrane (IAM), which is separate from and bordering the nuclear envelope, and (ii) the outer acrosomal membrane (OAM), which is located underneath the plasma membrane and fuses with that structure during acrosomal exocytosis, also termed the acrosome reaction. The acrosomal matrix is located between the OAM and the IAM and consists of a protein matrix core that contains numerous hydrolytic and glycolytic enzymes. Although not described for stallion sperm specifically, the most well-known acrosomal enzymes are proacrosin-acrosin, hyaluronidase, β-galactosidase, various proteinases, neuraminidases, esterases, arylsulfatase, and phospholipases A and C, as well as numerous phosphatases and regulatory enzymes and proteins. The acrosome is derived from the Golgi complex of the spermatid as that organelle is reconfigured during spermiogenesis. Upon exocytosis of the acrosome, the contents of the acrosome are thought to contribute to the sperm’s ability to traverse the cumulus-oocyte complex (COC) and penetrate the oocyte during the process of fertilization.1 The acrosomal matrix contains a number of proteins with galactosyl residues and can be detected using a range of fluoresceinated lectins as well as antibodies raised against acrosomal proteins. It is this matrix of proteins and carbohydrates that is the target for most methods of acrosomal detection and these methods will be discussed later in this chapter. In particular, the fluorescently labeled pea lectin, Pisum sativum agglutinin (PSA) and the peanut lectin Arachis hypogaea agglutinin (PNA) have been used with fluorescence microscopy and flow cytometry2–4 to accurately detect stallion sperm acrosomes. Coomassie blue and
Triple stain are non-fluorescent dyes that have been used to detect acrosomes in horses5 and other mammalian species.
Acrosomal exocytosis (the acrosome reaction) For fertilization to occur, the acrosome must first undergo exocytosis (Figure 152.1) whereby the contents of the acrosome (matrix) are expelled into the vicinity of the oocyte and zona pellucida (ZP). The ensuing loss of the plasma membrane overlying the sperm head exposes the IAM and allows the sperm to penetrate the acellular zona pellucida and eventually enables the sperm plasma membrane to fuse with, and penetrate, the oocyte plasma membrane (also termed the oolemma or oocyte plasmalemma) to complete fertilization. Following transit through the ZP, the inner acrosomal membrane of the sperm recognizes, and subsequently fuses with, the oocyte plasmalemma, a process that requires specific ligand–receptor interactions between the sperm and oocyte surfaces, as well as fusion proteins. In this process, the sperm head becomes engulfed by the oolemma.6, 7 In order for the inner acrosomal membrane to contact the oocyte plasma membrane, the sperm must first shed its plasma membrane and outer acrosomal membrane. This occurs in vivo by the exocytotic event known as the ‘acrosome reaction’, whereby this Golgi-derived structure undergoes specifically regulated calcium-requiring membrane events that lead to an increased fusogenicity of the OAM and plasma membrane, resulting in release of the acrosomal contents (Figures 152.1 and 152.2). The acrosome reaction of the fertilizing sperm must take place at, or near, the ZP and the natural stimulus for this event may involve molecules associated with the oocyte. In particular, the ZP of the egg is known to induce the acrosome reaction in a speciesspecific manner for all species studied.6, 7 The ZP is
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 152.1 Morphology of the acrosome reaction (A–D). Acrosomal membrane (AM), plasma membrane (PM), as modified by Wasserman et al., 20056 from an original drawing from Yanagimachi, 1994.7 Reprinted with permission from Elsevier.
Figure 152.2 Mammalian fertilization. Within the female reproductive tract sperm undergo a series of surface and intracellular transformations collectively termed capacitation (i) which enables them to bind (ii) to the zona pellucida (ZP) and undergo the acrosome reaction (iii). The release of hydrolytic enzymes from the acrosome (iv) facilitates sperm passage through the ZP and fusion with the oolemma (v). Reprinted from Nixon et al., 2007.30 With permission from Springer.
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Acrosomal Function comprised of three major glycoproteins designated as ZP1, ZP2, and ZP3; a fourth glycoprotein has been described for some species under certain conditions (reviewed by Sinowatz et al.).8 ZP3 has wide acceptance by researchers as the physiological inducer of the acrosome reaction and has been shown to be bifunctional: it has both recognition of the sperm and induction of the acrosome reaction as its main functions. Because inappropriate acrosome reactions in the absence of ZP may render sperm infertile, prolonged acrosomal stability of sperm in vivo may ensure that sperm reaching the oocyte are competent to fertilize the oocyte. This is particularly important in the horse because of the long estrus duration and receptivity to the stallion relative to ovulation, and the potentially long residence of sperm within the female reproductive tract (reportedly up to 6 days). Acrosomal exocytosis is possible only following the process of capacitation (see Chapter 102) that occurs within the upper female genital tract in vivo, but can be induced to occur in vitro. Capacitation is a final maturation process that prepares a sperm to undergo the acrosome reaction and fuse with the oocyte. It is largely undefined but includes primarily a number of changes to the plasma membrane including loss of cholesterol and phospholipid rearrangement, but increases in intracellular calcium and pH changes are required before acrosomal exocytosis can proceed.7, 9 As the capacitated sperm approaches the oocyte and penetrates the egg investments (cumulus oophorus and its associated extracellular matrix), a sperm cell ligand, that is not well described, interacts with the acellular zona pellucida surrounding the oocyte. This occurs as a result of binding to the glycoprotein ZP3 molecule and results in sperm surface receptor aggregation.10 A signal transduction cascade follows receptor aggregation that likely involves membrane- and cytosol-associated complexes including guanosyl nucleotide regulating protein (G protein), cyclic adenosine monophosphate (cAMP), phospholipase C (PLC), and protein kinase A (PKA). The stimulation of cAMP activates protein kinase A that begins to phosphorylate a number of proteins11–14 that are largely unknown. Within 1 minute of sperm stimulation, there is a great influx of calcium ions into the sperm cell. This is followed closely in time by swelling of the acrosomal matrix in what appears to be a prelude to fusion and vesiculation of the plasma membrane and outer acrosomal membrane. The acrosomal matrix is thought to leak from the matrix area as pores form between areas of vesicle formation between the OAM and plasma membrane, resulting in the release of acrosomal enzymes and structural elements into the immediate vicinity of the anterior sperm head and acrosome (Figures 152.1 and 152.2).
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A number of substances have been demonstrated to induce acrosome reactions in vitro in a range of vertebrate species, including the horse. The question of whether these inducers stimulate a physiological acrosome reaction has not been fully determined; however, there have been elements of both pharmacological and non-physiological inductions of acrosomal exocytosis. For instance, the calcium ionophores, A23187 and ionomycin, have been used to induce acrosomal exocytosis in many species, but because these mediators of calcium influx are so potent in stimulating a very large and rapid influx of calcium, a rapid exocytotic reaction may occur. This overwhelming response involves large numbers of cells (> 80%)and is thus considered by many to be non-physiological. In comparison, the generally accepted physiological inducer, homologous zona pellucida, typically induces approximately 20–30% of capacitated cells to undergo exocytosis. Nevertheless, the ionophores allow calcium influx and acrosome reactions to occur, although the cells are mostly rendered non-viable. It has been demonstrated that stallion sperm could be induced in vitro to capacitate using a modified human in vitro fertilization medium (e.g., Tyrode’s medium) and could then undergo the acrosome reaction when stimulated by progesterone, equine zona pellucida, or porcine zona pellucida.15–20 Progesterone treatment following in vitro capacitation resulted in increased acrosome reactions in live sperm from the fertile stallions as compared to control-treated sperm.16 A non-genomic progesterone receptor located on or near the sperm plasma membrane may mediate these events in human spermatozoa21, 22 and this has also been demonstrated for equine sperm.20, 23 Cheng et al.23 demonstrated the presence of a membraneassociated progesterone receptor in stallion sperm. This receptor becomes exposed as capacitation proceeds such that more progesterone receptor is available for binding progesterone and stimulating induction of the acrosome reaction. Further, it has been shown that the action of the progesterone receptor is linked with phosphorylation of tyrosine residues during in vitro capacitation.20
Clinical application of acrosome reactions in stallion sperm In several studies, stallion subfertility was associated with a diminished ability of ejaculated sperm to capacitate in vitro and respond to progesterone- and zona pellucida-stimulated acrosome reactions.16–18 Similar studies had been reported for human sperm subfertility. Progesterone was evaluated for its ability to stimulate acrosome reactions because the equine
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preovulatory follicle has been reported to secrete progesterone before ovulation and luteinization24 and is present in oviductal fluids and within the cumulus of ovulated oocytes of mares in vivo. Similarly, bovine and human male fertility has been correlated to acrosome reaction inducibility by progesterone, chondroitin sulfates, and dilauroylphosphatidylcholine liposomes.25, 26 A number of stallions have been identified with histories of subfertility that failed to acrosome-react following in vitro treatment with progesterone or the calcium ionophore, A23187.16, 27, 28 Other spermatozoal motility and morphological parameters were unable to explain the subfertility in these horses. In these studies, the acrosome reaction was stimulated over a 2- to 3-hour period in seven subfertile stallions. The calcium ionophore A23187 stimulated 60–80% of acrosome-reacted sperm over this time period in control stallions with history of normal fertility while the subfertile stallions failed to demonstrate any significant percentages of cells with acrosome reactions.27, 28 Although this method is considered by some to be non-physiological because the A23187 causes an extensive influx of calcium through altered membranes, this acrosome responsiveness assay (ASA) has been gaining utility as a diagnostic test in several equine andrology laboratories that may distinguish between stallions with normal and abnormal acrosomal function. A companion study using the same stallions recently demonstrated that the molar ratio of cholesterol to phospholipids was notably higher in the sperm and seminal plasma of these subfertile stallions, suggesting a cellular mechanism for poor acrosomal function in this subset of subfertile stallions.29
Assays for acrosomal status and acrosome reactions In certain species, the acrosome can be visualized using simple light microscopy without staining, such as bright-field, phase contrast, or differential interference contrast (DIC) at 1000× magnification. In the rabbit and guinea pig, in particular, the acrosome is located over the anterior sperm head and its protrusion over the edges of the posterior sperm head allows visible assessment of acrosomal intactness. Sperm in which the acrosome is present are termed ‘acrosome intact’ while the absence of this organelle is termed ‘acrosome reacted or acrosome lost.’ In species with smaller sperm heads, including human, bull, horse, and dog, the acrosome cannot be visualized without some type of staining that sets the acrosome region apart from the rest of the cell. However, scanning electron microscopy (SEM) or transmission electron microscopy (TEM) can be used
for acrosomal status and is often called the ‘gold standard’ for this assay because of the high degree of accuracy in defining the acrosome with EM techniques. This method, while accurate and precise, is expensive, time-consuming, labor intensive, and requires experience. As such, EM has not become routine in a clinical setting for stallions, even at referral hospitals. A number of bright-field stains have been used to stain the acrosomes of stallion sperm and these include Wright’s stain, Giemsa, Coomassie blue, Triple stain, Trypan blue, Chicago sky blue, eosin-nigrosin, and India ink, the latter two being background, or, exclusion, type stains. The former stains have allowed some visualization of acrosomes but, while useful for sperm morphology, can be difficult to use for acrosome detection because the posterior acrosome borders may be ambiguous and lack contrast. These cells are usually dried flattened to a glass slide or cover slip surface. Brum et al.5 validated the Coomassie blue acrosomal stain against a fluorescent acrosome stain and reported a strong correlation between Coomassie blue-stained acrosomes and those stained with a fluoresceinated lectin (FITC-PSA, discussed below). Very few studies have validated these stains against electron microscopy (EM), which, as mentioned above, has long been considered the gold standard for acrosome detection. A fluorescence method using the antibiotic chlortetracycline (CTC) has been used in the mouse primarily to detect capacitating and acrosome reacting sperm. This assay determines a series of sperm cell labeling patterns that some have associated with changes in intracellular calcium concentrations giving a graded response during capacitation conditions. Its usefulness has also been reported in stallion sperm.15 The CTC assay for acrosomal status, although simple to perform, has not become clinically useful in large part because sperm capacitation has not been worked out well in the horse model, making validation of this assay difficult at this time. In contrast, fluorescence staining of acrosomes has become useful and often routine in some clinical settings as an adjunctive test for the evaluation of breeding soundness, cryopreservation survival, and subfertility in stallions. A fluorophore is a functional portion of a molecule that can be excited at one specific wavelength to emit fluorescent light at another specific wavelength. Most fluorochromes (fluorescent dyes) used in biological imaging are conjugated to derivatives of isothiocyanate (ICT) joined to some other protein, lipid, or carbohydrate molecule that has binding activity. Common derivatives are fluorescein (FITC) that fluoresces green around a maximum wavelength of 520 nm when excited at 488 nm and rhodamine isothiocyanate (TRITC) that
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Acrosomal Function fluoresces red at 576 nm when excited at 505 nm. In general, when an intact acrosome is present, the plasma membrane is intact and therefore prevents the fluorochrome from accessing the interior of the acrosomal matrix. Intact sperm, as such, usually do not bind or fluoresce with most fluorochromes and require permeabilization that allows the fluorochrome to access and bind the acrosomal matrix. The fluoresceinated (fluoroisothiocyanite, FITC) lectins (e.g., PSA and PNA) discussed above bind to carbohydrate or sugar moieties within the acrosome as their mechanism of action. When the acrosome is lost, reacted, or damaged, the sugar moieties have been lost and the fluoresceinated lectin or antibody fails to bind and fluoresce. Several reports using acrosome-directed antibodies along with secondary fluorescent-conjugated antibodies have also been reported for detecting stallion acrosomes but these methods are not as efficient as the single-step fluorescence methods discussed17, 18 and are not in routine use. This method allows detection of acrosome reactions when sperm are bound to oocytes because the lectin methods tend to label the zona as well as the sperm, making detection difficult or impossible. Sperm cells are generally fixed using a buffered formaldehyde-based fixative such as 2–4% paraformaldehyde, 2% glutaraldehyde, or 1–10% formalin prior to labeling with a fluorescent dye. Such fixed cells are more stable and take up fluorescent dyes more evenly than unfixed cells. These fixatives are highly toxic to living sperm cells so should not be used in the vicinity of semen used for breeding. Permeabilization is usually performed using 95% ethanol or methanol briefly followed by extensive cell washing in phosphate-buffered saline (PBS). The fluoresceinated lectins have relatively short fluorescence lifespan, in that they tend to degenerate in fluorescence over the time in which they are excited. A newer class of fluorochromes, those conjugated to Alexa Fluors, demonstrate excitation and emission properties similar to FITC and TRITC but have been developed to have more stability, brighter intensity, and less pH sensitivity, making this group of fluorochromes very useful in sperm biology laboratories for acrosomal detection and a number of other cell functions including mitochondrial function, membrane potential, and cell viability/membrane integrity assays. To increase visualization of acrosomal labeling, several laboratories have added propidium iodide (PI), which fluoresces in the red wavelengths, or bisbenzimidine (Hoeschst) dyes, which fluoresce in the blue ultraviolet (UV) range, to provide contrast to the green fluorescence of fluorescein-conjugated or Alexinated lectins. These are DNA dyes that enter the nuclear compartment and complex with com-
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Figure 152.3 Two acrosome intact stallion sperm demonstrating double fluorescence labeling of the nuclear compartment by propidium iodide (red) and the acrosomes by peanut lectin Arachis hypogaea agglutinin (PNA) conjugated to FITC (yellowgreen). (See color plate 48).
pacted sperm DNA. Propidium iodide is membraneimpermeable and only gains access to sperm DNA in cells that have disrupted plasma membranes, or have been fixed or permeabilized with ethanol. So, if the cells have been fixed, all cells treated with PI will fluoresce red, in high contrast to the green fluorescence of Alexa or fluorescein (see Figures 152.3 and 152.4). Ethidium homodimer is a fluorescent chemical similar to PI and binds to sperm DNA and has been used to provide red fluorescence contrast to acrosomal staining.3, 4 There are two principal Hoeschst dyes, No. 33258 and No. 33242. The latter dye is membrane permeable and thus enters all cells to complex with DNA and fluoresce. The other Hoeschst dye (33258) behaves much like PI, in that it only enters cells with membrane damage, but in
Figure 152.4 Two acrosome-reacted sperm are shown with primarily propidium iodide-stained nuclei with an absence of FITC-PNA staining because the acrosomes are lost. The other two sperm are acrosome-intact. (See color plate 49).
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the case of H33258, fluorescence is blue (UV). Both the 33258 and PI therefore behave as indicators of membrane integrity (viability) and provide for four distinct cell types with regard to acrosomal status: (i) acrosome intact, live, (ii) acrosome intact, dead, (iii) acrosome reacted, live, and (iv) acrosome reacted, dead.2 Figure 152.3 demonstrates the use of FITCconjugated PSA with acrosome-intact stallion sperm. The yellow-green FITC-PNSA label defines the acrosome in a freshly ejaculated normal sperm while the red area of the posterior sperm head defines the nuclear compartment as defined by propidium iodide. Figure 152.4 shows the appearance of two acrosomeintact sperm in addition to two acrosome-reacted sperm lacking yellow-green fluorescence over the anterior sperm head indicating the loss of acrosomes in those cells. These categories can be very helpful when attempting to distinguish sperm cell health and function in freshly ejaculated or even cryopreserved sperm cells.
Summary The dynamic process of acrosomal exocytosis is a powerful sperm function that is essential for sperm to bind, penetrate, and ultimately fuse with oocytes. This cellular process is complex and events leading to it are poorly understood. It is not yet clear to what extent stallion fertility is dependent on acrosomal status although several studies indicate that acrosomal integrity may be responsible for subfertility in at least some stallions. Nonetheless, a number of methods to detect this important process have emerged and vary from bright-field staining methods to sophisticated fluorescence and flow cytometric methods. Many of these diagnostic methods can be performed in the field with routine microscopic equipment while others require relatively expensive fluorescence microscopes and laser-based flow cytometers. However, as with most computer-based technologies, the costs are coming down rapidly and these recently developed diagnostic techniques are seen in a growing number of referral stallion facilities worldwide.
References 1. Eddy EM, O’Brien DA. The Spermatozoon. In: Knobil E, Neill JD (eds) The Physiology of Reproduction. Vol 1, 2nd edn. New York: Raven Press, 1994, pp. 29–77. 2. Casey PJ, Hillman RB, Robertson KR, Yudin AI, Liu IKM, Drobnis EZ. Validation of an acrosomal stain for equine sperm that differentiates between living and dead sperm. J Androl 1993;14(4):289–97. 3. Magistrini M, Guitton E, Levern Y, Nicolle JC, Vidament M, Kerboeuf D, Palmer E. New staining methods for sperm evaluation estimated by microscopy and flow cytometry. Theriogenology 1997;48:1229–35.
4. Cheng F-P, Fazeli A, Voorhout WF, Marks A, Bevers MM, Colenbrander B. Use of PNA (Peanut Agglutinin) to assess the acrosomal status and the zona pellucida induced acrosome reaction in stallion spermatozoa. J Androl 1996;17:674–82. 5. Brum AM, Thomas AD, Sabeur K, Ball BA. Evaluation of Coomassie blue staining of the acrosome of equine and canine spermatozoa. Am J Vet Res 2006;67(2):358–62. 6. Wassarman PM, Jovine L, Qi H, Williams Z, Darie C, Litscher ES. Recent aspects of mammalian fertilization research. Mol Cell Endocrinol 2005;234:95–103. 7. Yanagimachi, R. Mammalian fertilization. In: Knobil E, Neill JD (eds) The Physiology of Reproduction. Vol 1, 2nd edn. New York: Raven Press, 1994, pp. 189–317. 8. Sinowatz F, Topfer-Petersen E, Kolle S, Palma G. Functional morphology of the zona pellucida. Anatomia Histologia Embryologia J Vet Med Series C 2001;30(5):257–63. 9. Flesch FM, Gadella BM. Dynamics of the mammalian sperm plasma membrane in the process of fertilization. Biochim Biophys Acta 2000;1469:197–235. 10. Leyton L, Saling P. Evidence that aggregation of mouse sperm receptors by ZP3 triggers the acrosome reaction. J Cell Biol 1989;108:2163–8. 11. Breitbart H, Naor Z. Protein kinases in mammalian sperm capacitation and the acrosome reaction. Rev Reprod 1999;4:151–9. 12. Visconti PE, Ning X, Fornes MW, Alvarez JG, Stein P, Connors SA, Kopf GS. Cholesterol efflux-mediated signal transduction in mammalian sperm: cholesterol release signals an increase in protein tyrosine phosphorylation during mouse sperm capacitation. Dev Biol 1999;214:429–43. 13. Pommer AC, Rutllant J, Meyers S. Phosphorylation of protein tyrosine residues in fresh and cryopreserved stallion spermatozoa under capacitating conditions. Biol Reprod 2003;68:1208–14. 14. McPartlin LA, Littell J, Mark E, Nelson JL, Travis AJ, Bedford-Guaus SJ. A defined medium supports changes consistent with capacitation in stallion sperm, as evidenced by increases in protein tyrosine phosphorylation and high rates of acrosomal exocytosis. Theriogenology 2008;69:639–50. 15. Varner DD, Bowen JA, Johnson L. Effect of heparin on capacitation/acrosome reaction of equine sperm. Arch Androl 1993;31:199–207. 16. Meyers SA, Overstreet JW, Liu IK, Drobnis EZ. Capacitation in-vitro of stallion spermatozoa: comparison of progesterone-induced acrosome reactions in fertile and subfertile males. J Androl 1995;16:47–54. 17. Meyers S, Liu IKM, Overstreet JW, Drobnis EZ. Induction of acrosome reactions in stallion sperm by equine zona pellucida, porcine zona pellucida, and progesterone. Biol Reprod 1995;Monogr 1:739–44. 18. Meyers S, Liu IKM, Overstreet JW, Drobnis EZ. Spermzona pellucida binding and zona-induced acrosome reactions in the horse: comparisons between fertile and subfertile males. Theriogenology 1996;46:1277–88. 19. Varner DD, Thompson JA, Blanchard TL, Heck R, Love CC, Brinsko SP, Johnson L. Induction of the acrosome reaction in stallion spermatozoa: effects of incubation temperature, incubation time, and ionophore concentration. Theriogenology 2002;58:303–6. 20. Rathi R, Colenbrander B, Stout TAE, Gadella BM. Progesterone induces acrosome reaction in stallion spermatozoa via a protein tyrosine kinase dependent pathway. Molec Reprod Devel 2003;64(1):120–8.
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Acrosomal Function 21. Meizel S, Turner K. Progesterone acts at the plasma membrane of human sperm. Mol Cell Endocrin 1991;11:R1– R5,65. 22. Melendrez C, Meizel S, Berger T. Comparison of the ability of progesterone and heat solubilized porcine zona pellucida to intiate the porcine sperm acrosome reaction in vitro. Mol Reprod Dev 1994;39:433–8. 23. Cheng FP, Gadella BM, Voorhout WF, Fazeli AR, Bevers MM, Colenbrander B. Progesterone-induced acrosome reaction in stallion spermatozoa is mediated by a plasma membrane progesterone receptor. Biol Reprod 1998;59(4):733–42. 24. Linford R, McCue P, Montavon S, Lasley BL. Longterm cannulation of the ovarian vein in mares. Am J Vet Res 1992;53:1589–93. 25. Tesarik J, Mendoza C. Defective function of a nongenomic progesterone receptor as a sole sperm anomaly in infertile patients. Fertil Steri1 1992;58:793–7.
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26. Ax RL, Dickson K, Lenz RW. Induction of acrosome reactions by chondroitin sulfates in vitro corresponds to nonreturn rates of dairy bulls. J Dairy Sci 1985;68:387–90. 27. Bosard T, Love C, Brinsko S, Blanchard T, Thompson J, Varner D. Evaluation and diagnosis of acrosome function/dysfunction in the stallion. Anim Reprod Sci 2005;89(1–4):215–17. 28. Love C, Bosard TS, Brinsko SP, Carrell BP, Varner DD. Acrosomal dysfunction in stallion spermatozoa: what is known. Havemeyer Workshop, Bandera, TX, Sept 22–25, 2006. 29. Brinsko SP, Love CC, Bauer JE, Macpherson ML, Varner DD. Cholesterol to phospholipid ratio in sperm of stallions with unexplained subfertility. Anim Reprod Sci 2005;89(1–4):217–19. 30. Nixon B, Aitken RJ, McLaughlin EA. New insights into the molecular mechanisms of sperm-egg interaction. Cell Mol Life Sci 2007;64:1805–23.
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Chapter 153
Evaluation of the Plasma Membrane Ben Colenbrander and Tom A.E. Stout
Introduction Sperm cells are entirely surrounded by a plasma membrane that not only contains and protects the various organelles but also plays vital roles in many aspects of sperm function. With regard to protection, the plasma membrane presents a physical barrier to the environmental hazards to which a sperm may be exposed during its passage through the female genital tract, or during ex vivo handling and storage for artificial insemination. While the sperm plasma membrane has the same basic structure as that surrounding a somatic cell, it differs significantly in the precise composition and distribution of lipids, proteins, and surface sugars (glycocalyx).1 Moreover, the sperm plasma membrane is homogeneous neither across its full extent nor over time; instead, certain areas undergo precise, sequentially regulated changes in either composition and/or distribution of specific components in preparation for their roles in processes such as temporary sperm binding to the oviductal epithelium (sperm reservoir), binding to the zona pellucida, the acrosome reaction, and fusion with the oolemma during fertilization.2, 3 The resting sperm plasma membrane has a characteristic polarized morphology. Its surface is laterally heterogeneous and divided into ‘membrane domains’ that undergo further reorganization during sperm maturation and activation. The most fundamental division is that between the plasma membranes of the sperm head, midpiece, and tail, which are distinct and remain so because the posterior and annular ring structures function as barriers to lipid exchange. The plasma membrane of the sperm head shows further marked regional heterogeneity, with regional differences reflecting the different roles of specific areas of the sperm head.
At least four sperm head surface regions can be distinguished: the pre-equatorial, equatorial, and post-equatorial areas, and the apical ridge (Figure 153.1). The apical ridge is involved in primary sperm–zona binding (Figure 153.2) and, following capacitation, specific zona-binding proteins accumulate here. Zona binding via the apical ridge in turn triggers fusion of the sperm plasma membrane over the pre-equatorial region with the underlying outer acrosomal membrane at multiple points, via Soluble N-ethylmaleimide-sensitive fusion Attachment protein REceptor (SNARE) protein-mediated vesicle formation events.2 The resulting formation of ‘mixed vesicles’ bounded by a combination of plasma membrane and outer acrosomal membrane (Figures 153.1 and 153.2) is the key event in the acrosome reaction, and results in the release of the acrosomal contents (mostly proteolytic enzymes) that enable zona pellucida (ZP) penetration. The acrosome reaction (AR), therefore, involves loss of the plasma and outer acrosomal membranes over the pre-equatorial region, leaving the inner acrosomal membrane as a ‘surrogate’ plasma membrane, while a ‘hairpin’ membrane structure remains at the equator where the plasma membrane and outer and inner acrosomal membranes meet; it is via this hairpin structure that the sperm fuses to the oolemma (Figure 153.2). An intact and functional plasma membrane is therefore vital to both sperm survival and fertilizing potential. As such, tests of both plasma membrane integrity and function, where the latter are based on the regionalized changes in lipid and protein organization or membrane permeability that occur during capacitation or the acrosome reaction, may give valuable information about sperm quality and ‘fertilizing potential’.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 153.1 Schematic representations of a stallion sperm. (a) Cross-section; 1, plasma membrane; 2, outer acrosomal membrane; 3, acrosomal contents; 4, inner acrosomal membrane; 5, nuclear envelope; 6, nucleus; 7, nuclear ring; 8, mitochondria; 9, midpiece; 10, annular ring; 11, flagellum (principal and end pieces); 12, fibrous sheath. (b) Plasma membrane subdomains of the sperm head; 13, apical ridge; 14, pre-equatorial region; 15, equatorial area; 16, post-equatorial region. (c) Acrosome reaction; 17, ‘mixed vesicles’ formed by fusion of the plasma membrane and outer acrosomal membrane. Adapted from Gadella et al.41 with permission from Anim Reprod Sci.
Plasma membrane integrity Staining of sperm with dyes that cannot cross an intact plasma membrane (‘vital staining’ using ‘membrane-impermeant dyes’) has long been recognized as a useful way of quantifying the proportion of viable sperm in a sample. Under field conditions, this can be achieved by light microscopic evaluation of a simple ‘smear’ preparation stained with a dye combination that includes a membrane-impermeant component (e.g., eosin or trypan blue); commonly used combinations include aniline blue–eosin, nigrosin–eosin, William’s stain (carbol fuchsin–eosin), and Giemsa–trypan blue.4–6 Following vital staining, cells that exclude the membrane-impermeant dye (i.e., remain white) are considered viable (or at least membrane intact), whereas those that take up the stain are classified as dead (Figure 153.3). Vital staining techniques are simple, rapid indicators of plasma membrane integrity that do not require expensive or complex equipment and, when performed under conditions of stable ambient humidity and temperature, may offer useful information about the proportion of viable sperm in a sample. In recent years, these traditional vital stains have largely been superseded by an array of
membrane-impermeant fluorescent DNA dyes such as Hoechst 33258, YoPro-1, propidium iodide, ethidium homodimer-1, ToPro-3, and TOTO.6–11 When a fluorescent dye is used, evaluation of sperm viability is performed using either a fluorescence microscope or, preferably, a fluorescence-activated cell sorter (FACS/flow cytometer). Flow cytometric analysis has the considerable advantage of allowing assessment of dye uptake for thousands of sperm within seconds, thereby generating an accurate and objective assessment of viability. It is, nevertheless, worth noting that, while the data generated using a given fluorescent viability stain broadly correlate with those generated using conventional vital stains and other fluorescent dyes, results can differ. For example, conventional vital stains have been reported to overestimate the proportion of viable sperm in frozen–thawed stallion semen.12 An alternative group of fluorescent dyes used to examine membrane integrity are the acylated DNA dyes (e.g., SYBR-14, fluorescein diacetate and its carboxy(methyl) derivatives). These dyes pass through the plasma membrane into living sperm, but, once inside the cell, are deacylated by intracellular esterases and thereby rendered impermeant (no longer able to cross an intact membrane) such that the dye accumulates within membrane-intact, but not
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Figure 153.4 Viability staining of stallion sperm with the fluorescent dyes SYBR-14 and propidium iodide (PI). Live sperm accumulate SYBR-14 and, therefore, stain intensely green, whereas the dead sperm allow entry of PI and thus fluoresce red. (See color plate 51).
Figure 153.2 Diagrammatic representation of the roles of sperm membrane subdomains in sperm–oocyte interaction and fertilization. 1, The sperm binds to the zona pellucida (ZP) via its apical ridge; 2, the acrosome reaction involves fusion of the outer acrosomal membrane with the plasma membrane over the apical ridge and pre-equatorial region; 3, the equatorial membrane ‘hairpin structure’ remains intact during penetration of the sperm through the ZP; 4, sperm binding and fusion with the oolemma takes place via the hairpin structure; 5, sperm cytosolic factors activate the oocyte; 6, secretion of cortical granules induces the definitive block to polyspermy. Adapted from Gadella et al.41 with permission from Anim Reprod Sci.
membrane-damaged, sperm; as a result, membraneintact sperm fluoresce more intensely. In fact, combinations of the two types of fluorescent dye are now commonly used to improve the sensitivity of sperm viability testing (Figure 153.4), e.g. SYBR-14 to stain live sperm (green) combined with ethidium homodimer or propidium iodide to simultaneously stain dead sperm (red).7 Such a combination of dyes greatly improves the accuracy of flow cytometric viability analysis in semen that has been cryopreserved, or for other reasons contains large amounts of nonsperm particles, such as egg yolk or milk fat droplets. When a combined viability stain is used, analysis can be performed immediately without processing to remove the non-sperm particles because the latter can instead be ‘gated out’ during FACS analysis because they do not contain DNA and, therefore, do not take up either dye.10 By contrast, if only a membrane-impermeant dye is used for viability staining of frozen–thawed semen it can be difficult to flow cytometrically distinguish live sperm from egg yolk particles.
Plasma membrane osmoregulation
Figure 153.3 Vital staining of stallion sperm with aniline blue–eosin. The two unstained sperm (white) would be classified as viable and the stained sperm (red) as non-viable. The sperm on the far left has both an obviously abnormal acrosome and a protoplasmic droplet on the proximal part of the midpiece. (See color plate 50).
The sperm plasma membrane is not simply a physical barrier, but performs many essential cellular functions, including osmoregulation. The osmoregulatory capacity of the plasma membrane can be tested using the hypo-osmotic swelling test (HOS).13–15 When exposed to hypo-osmotic conditions, sperm with an intact and functional plasma membrane will equilibrate with the extracellular environment by actively pumping in water. In the tail region, there is little room to accommodate the influx of water, and the result is visible expansion and coiling of the
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portant role in synchronizing sperm capacitation and migration to the site of fertilization with ovulation. Moreover, the ability of sperm to bind to oviduct epithelium in vivo and in vitro and the quality of the bound sperm appear to be influenced by stallion and mare fertility.18 In particular, poor stallion fertility has been associated with reduced capacity of sperm to associate with cultured oviduct epithelial cells, and a consequent reduced mean survival time.19, 20 In practice, however, sperm–oviduct epithelial cell binding tests are laborious.
Figure 153.5 The effects of the hypoosmotic swelling test on stallion sperm. The two sperm with coiled tails would be classified as having an intact, osmotically active plasma membrane whereas the sperm with a straight tail would be classified as having a defective plasma membrane.
flagellum, beginning at the distal end and progressing towards the midpiece; this produces a characteristic spectrum of sperm with swollen or coiled tails (Figure 153.5). The HOS test is thus a simple method (no need for a fluorescence microscope or flow cytometer) of testing both plasma membrane integrity and osmotic function and, as such, is considered a useful adjunct to routine semen analysis.14 Stallions for which less than 40% of sperm exhibit a positive HOS reaction have been reported to be of doubtful fertility.14, 16 The HOS test can even be automated, by using an electronic counter to measure sperm-cell volume, so that it becomes a rapid objective test of plasma membrane integrity and osmoregulatory function.17
Interaction of sperm with the oviductal epithelium With the exception of deliberate post-ovulatory insemination of frozen–thawed semen, sperm generally arrive in the mare’s reproductive tract before ovulation. A relatively small population is then apparently selected, by some unknown mechanism, for normality or fertilizing potential and establishes a sperm ‘reservoir’ in the caudal isthmus of the oviduct.18 Sperm in the reservoir bind to oviduct epithelial cells via plasma membrane lectins, and this binding maintains the sperm in a noncapacitated state and thereby prolongs their viability.19 Close to the time of ovulation, signals within the oviductal environment (thought to include progesterone) stimulate the sperm to detach from the oviduct epithelial cells, capacitate, and move toward the newly arrived cumulus–oocyte complex. While sperm–oviduct binding does not appear to be essential to fertilization per se, it does appear to play an im-
Capacitation-related changes in the sperm plasma membrane Capacitation is the process by which a sperm is prepared for its role in ZP binding, the acrosome reaction, and fertilization. Broadly speaking, capacitation is thought to consist of the removal of ‘decapacitating factors’ that coat the sperm within the epididymis and up until shortly after ejaculation, accompanied by modifications in plasma membrane organization. In particular, the plasma membrane phospholipids of ‘resting’ sperm are distributed asymmetrically across the lipid bilayer with the choline phospholipids, sphingomyelin (SM) and phosphatidylcholine (PC), located predominantly in the outer lipid leaflet and the aminophospholipids, phosphatidylethanolamine (PE) and phosphatidylserine (PS), located primarily in the inner leaflet.21 This asymmetrical distribution of lipids is maintained by enzymatic activity, until the onset of capacitation. One of the early events in sperm capacitation is the loss of the asymmetrical phospholipid distribution (‘membrane scrambling’).3 The resulting more random distribution of phospholipids across the bilayer is associated with an increase in membrane fluidity that enables the depletion of cholesterol from the membrane,22 allows increased regional membrane heterogeneity (i.e., establishment of functional ‘membrane domains’), and promotes activation of membrane tyrosine kinases.23 While capacitation is an essential step in the preparation for fertilization, capacitated sperm have a fairly limited lifespan, and premature capacitation is therefore undesirable. As a result, analysis of capacitation has two major aspects: (i) detection of ‘prematurely’ capacitated sperm and (ii) assessing the proportion of sperm able to undergo capacitation-like processes in response to appropriate stimuli. A critical and possibly rate-limiting step in capacitation is the depletion of membrane cholesterol, and the relatively high cholesterol content of stallion sperm presumably explains why capacitation is relatively slow in this species. The amount and location
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of cholesterol in the sperm plasma membrane can be visualized using filipin, a fluorescent antibiotic that complexes with unesterified cholesterol. Tracking the filipin–cholesterol complexes using ultraviolet fluorescence microscopy has shown that cholesterol is initially distributed evenly over the entire sperm head plasma membrane, but that, once capacitation is initiated by exposing the sperm to a high concentration of bicarbonate (15 mM HCO3 − ), the cholesterol accumulates in the apical ridge. If the sperm are additionally exposed to a cholesterol acceptor, such as albumin, around 45% of the cholesterol is depleted from the plasma membrane, which as a result is ‘destabilized’. 22 Because of the importance of cholesterol in maintaining membrane stability, it has been suggested that artificially maintaining high cholesterol concentrations may improve survival during storage; in this respect, addition of cholesterol-loaded cyclodextrins to a semen freezing extender have been reported to improve cryosurvival of stallion sperm.24, 25 Nevertheless, further studies are needed to examine the consequences of cholesterol loading on subsequent cholesterol depletion, membrane function, and in-field fertility. An early and sensitive indicator of plasma membrane reorganization during capacitation is increased membrane fluidity or disorder as a result of ‘lipid scrambling’; this increased membrane fluidity can be detected using the fluorescent probe merocyanine 540,26 which cannot intercalate into the ordered, asymmetric plasma membrane of non-capacitated sperm, but is incorporated into the more fluid membrane of capacitated sperm, which therefore fluoresce.27 However, while merocyanin 540 staining has been described as a sensitive flow cytometric assay for assessing the capacitation status of stallion sperm,27, 28 a number of laboratories have struggled to validate its use and it is, therefore, not currently in routine use. Other fluorescence-labeled phospholipid probes have been described for assessing increased plasma membrane disorder in capacitated sperm. For example, fluorescein-conjugated annexin V binds to phosphatidylserine, which is ‘hidden’ in the inner leaflet of the plasma membrane bilayer of non-activated sperm, but is ‘visible’ in the outer leaflet following capacitation.3 As for merocyanin 540, however, although annexin V detection of the capacitation status of stallion sperm has been described,28, 29 the assay is difficult to validate and is not yet in widespread use. A simpler, but less well-understood technique for assessing the capacitation status of stallion sperm is staining with chlortetracycline (CTC). Chlortetracycline is a fluorescent antibiotic that binds to the plasma membrane in a calcium-dependent fashion. If CTC staining (green) is combined with a membrane-
impermeant viability stain such as Hoechst 33258 (blue), sperm can be divided into four categories; namely, dead (blue fluorescent nucleus), live noncapacitated (uniform bright green fluorescence over entire sperm head), live capacitated (green fluorescence over pre-equatorial region only), or live acrosome reacted (mottled green fluorescence over the head, green fluorescence only in the post-acrosomal region, or no fluorescence on the head).30, 31 A major advantage of CTC staining is, thus, that it allows simultaneous assessment of capacitation status and acrosome integrity.
Sperm progesterone receptor exposure Another major aspect of capacitation is the removal of so-called ‘decapacitation factors’ that coat the outer surface and inhibit further activation; their removal alters the surface glycocalyx and allows exposure and interaction of previously ‘hidden’ surface molecules. For example, inducing capacitation of stallion sperm leads to the exposure of a specific plasma membrane progesterone receptor that is thought to play an important role in the physiological AR.32, 33 Indeed, it is thought that progesterone in follicle fluid and/or secreted by the cumulus oophorus enhances both sperm–zona binding and the ZPmediated AR.34 Moreover, subfertility in some stallions has been correlated to an inability of their sperm to undergo the AR in response to progesterone,33 and/or a low percentage of capacitated sperm with detectable progesterone receptors.27 Progesterone receptor exposure can be detected by fluorescence microscopy or FACS analysis after labeling with progesterone conjugated to a fluorescent ligand, such as fluorescein isothiocyanate (FITC).32
Zona pellucida binding After leaving the sperm reservoir in the oviduct and undergoing capacitation, sperm cells head for the cumulus–oocyte complex with the aim of reaching, binding to, and penetrating the ZP. This process requires an intact plasma membrane with zona binding capacity and the ability to acrosome react. As for other species, the capacity of stallion sperm to bind to or penetrate the ZP correlates with fertility. However, because conventional in vitro fertilization is only poorly successful with horse gametes, in vitro fertilization is not a suitable test of sperm quality, and the significance of binding to, or penetration of, nonequine oocytes is unclear. Binding of stallion sperm to equine ZP does yield useful information but, because individual zonae differ significantly in their
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Figure 153.6 Detection of acrosome integrity in stallion sperm by staining with peanut agglutinin conjugated with fluorescein isothiocyanate (PNA–FITC; green), combined with the membrane-impermeant DNA dye ethidium homodimer (red) to indicate viability status. (a) Acrosome-reacting sperm show mottled fluorescence over the acrosomal cap region. (b) Acrosome-reacted sperm show fluorescence only along the equatorial band. (c) A red nucleus indicates a non-viable sperm. Live, acrosome-intact sperm do not take up either fluorescent dye and can thus only be visualized by conventional (not fluorescence) microscopy. (See color plate 52).
sperm-binding capacity, the most efficient way to obtain interpretable data is to use a ‘hemi-zona’ assay. In the hemi-zona assay, an oocyte is bisected and the binding capacity of sperm from a stallion of interest to the evacuated hemi-zona is compared with that of a stallion of known fertility. Although subfertile stallions show reduced sperm–ZP binding affinity in the hemi-zona assay,35 it is a laborious test that allows only limited between-stallion comparison.
The acrosome reaction The AR is an irreversible calcium-dependent process that involves fusion of the plasma membrane with the outer acrosomal membrane at multiple points, resulting in release of the acrosomal enzymes that aid ZP penetration. Several non-fluorescent and fluorescent staining combinations have been developed to assess the acrosome integrity of fresh and fixed sperm samples using microscopy, fluorometry, or flow cytometry.9, 15, 36, 37 The most commonly used stains are lectins that bind to glycoconjugates on the outer acrosomal membrane, such as peanut agglutinin (Arachis hypogaea; PNA) or Pisum sativum agglutinin (PSA).38, 39 For detection purposes, these lectins are conjugated to a fluorescent molecule such as FITC, and combined with a viability stain such as propidium iodide to allow simultaneous evaluation of viability and acrosomal integrity by fluorescence microscopy or flow cytometry. A sperm with an
intact acrosome will not show PNA- or PSA-specific fluorescence; acrosome ‘reacting’ sperm demonstrate a mottled fluorescent appearance over the entire sperm head; and sperm that have completed the acrosome reaction will demonstrate fluorescence in the equatorial region only (Figure 153.6). During flow cytometric analysis, any increase in PSA- or PNA-specific fluorescence suggests that the AR has been initiated; however, care is needed with interpretation of flow cytometric results if a high proportion of immature sperm are present in a sample (e.g., epididymal sperm) because the protoplasmic droplet also stains positively for both PNA and PSA (Figure 153.7). In the absence of fluorescence optics (or electron microscopy), Coomassie blue staining has been proposed as a useful technique for distinguishing acrosome-intact from acrosome-damaged or reacted sperm.40 After fixation with 4% paraformaldehyde to allow access of the dye to the acrosomal compartment, acrosome-intact sperm show blue staining over the entire pre-equatorial region, whereas acrosome-damaged or reacted sperm show either patchy staining or no staining. Acrosome integrity analysis by Coomassie blue staining appears to correlate well with FITC PNA analysis,40 and has the advantage of allowing analysis with a light microscope. If Coomassie blue staining can be successfully combined with a viability stain, which may be technically challenging given the need for sperm fixation prior
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Stallion: Specific Techniques Used in Semen Evaluation of the Stallion status, as well as integrity. Some of the viability and membrane function (HOS) tests can be assessed using a simple light microscope, whereas other tests employ fluorescent markers and require either a fluorescence microscope or a flow cytometer for analysis. Indeed, because flow cytometry allows rapid and objective analysis of multiple properties for many thousands of sperm, it has become the technique of choice for analyzing sperm quality. By contrast, while the various sperm-binding assays described (cultured oviductal epithelial cells or hemi-zonae) yield interesting and clinically relevant information, they are laborious and prone to experimental variation. Similarly, while assays for plasma membrane and acrosome integrity are well established, validation of a ‘user-friendly’ and reliable assay to detect capacitation of stallion sperm has proven more elusive, and further studies are required to establish exactly what happens in the plasma membrane of stallion sperm during capacitation and how this can best be monitored. In addition, while it is reasonable to assume that the ability of sperm to capacitate or acrosome react in vitro in response to either pharmacological or relatively physiological stimuli gives valuable information about ‘fertilizing potential’, there is a need for large-scale surveys to determine how commonly problems in these processes arise, and at what levels and to what extent they affect fertility in the field.
Figure 153.7 Stallion sperm stained with peanut agglutinin conjugated with fluorescein isothiocyanate (PNA–FITC; green), demonstrating that the protoplasmic droplet also takes up this fluorescent dye. This is relevant for flow cytometric analysis because sperm exhibiting PNA–FITC staining need to be differentiated into those that have actually acrosome reacted, and those that show green fluorescence only because they have a protoplasmic droplet. (See color plate 53).
to acrosome staining, it may become a useful method for assessing acrosome status in the field.
Conclusions The sperm plasma membrane plays critical roles in sperm survival and function and, presumably because of its great diversity of functions, shows regional specialization characterized by a heterogeneous distribution of lipid and protein components, which becomes even more marked following capacitation. The critical importance of the plasma membrane to sperm survival and function make it an important subject of tests for semen quality, while the regional and temporal differences in plasma membrane composition have made it possible to develop a range of tests for assessing membrane activation
References 1. Gadella BM. Sperm membrane physiology and relevance for fertilization. Anim Reprod Sci 2008;107:229–36. 2. Boerke A, Tsai PS, Garcia-Gil N, Brewis IA, Gadella BM. Capacitation-dependent reorganization of microdomains in the apical sperm head plasma membrane: functional relationship with zona binding and the zona-induced acrosome reaction. Theriogenology 2008;70:1188–96. 3. Gadella BM, Tsai PS, Boerke A, Brewis IA. Sperm head membrane reorganisation during capacitation. Int J Dev Biol 2008;52:473–80. 4. Dott HM, Foster GC. A technique for studying the morphology of mammalian spermatozoa which are eosinophilic in a differential ‘life-dead’ stain. J Reprod Fertil 1972;29:443–5. 5. Kenney RM, Huertgen JJ, Pierson R, Witherspoon D, Simons J. Theriogenology and the equine. Part II. The stallion, semen examination. J Soc Theriogenol 1983;9:1100–10. 6. Varner DD. Developments in stallion semen evaluation. Theriogenology 2008;70:448–62. 7. Garner DL, Johnson LA. Viability assessment of mammalian sperm using SYBR-14 and propidium iodide. Biol Reprod 1995;53:276–84. 8. Garner DL, Thomas CA, Joerg HW, DeJarnette JM, Marshall CE. Fluorometric assessments of mitochondrial function and viability in cryopreserved bovine spermatozoa. Biol Reprod 1997;57:1401–6. 9. Graham JK. Assessment of sperm quality: a flow cytometric approach. Anim Reprod Sci 2001;68:239–47.
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Evaluation of the Plasma Membrane 10. Nagy S, Jansen J, Topper EK, Gadella BM. A triple-stain flow cytometric method to assess plasma- and acrosomemembrane integrity of cryopreserved bovine sperm immediately after thawing in presence of egg-yolk particles. Biol Reprod 2003;68:1828–35. 11. Sieme H. Semen evaluation, In: Samper J (ed.) Equine Breeding Management and Artificial Insemination. St. Louis, MO: Saunders, 2009; pp. 57–74. 12. Merkies K, Chenier T, Plante C, Buhr MM. Assessment of stallion spermatozoa viability by flow cytometry and light microscope analysis. Theriogenology 2000;54:1215–24. 13. Jeyendran RS, Van der Ven HH, Perez-Pelaez M, Crabo BG, Zaneveld LJ. Development of an assay to assess the functional integrity of the human sperm membrane and its relationship to other semen characteristics. J Reprod Fertil 1984;70:219–28. 14. Neild D, Chaves G, Flores M, Mora N, Beconi M, Aguero A. Hypoosmotic test in equine spermatozoa. Theriogenology 1999;51:721–7. 15. Colenbrander B, Gadella BM, Stout TA. The predictive value of semen analysis in the evaluation of stallion fertility. Reprod Domest Anim 2003;38:305–11. 16. Nie GJ, Wenzel JG. Adaptation of the hypoosmotic swelling test to assess functional integrity of stallion spermatozoal plasma membranes. Theriogenology 2001;55:1005–18. 17. Petrunkina AM, Petzoldt R, Stahlberg S, Pfeilsticker J, Beyerbach M, Bader H, Topfer-Petersen E. Sperm-cell volumetric measurements as parameters in bull semen function evaluation: correlation with nonreturn rate. Andrologia 2001;33:360–7. 18. Troedsson MH, Liu IK, Crabo BG. Sperm transport and survival in the mare: a review. Theriogenology 1998;50:807–18. 19. Ellington JE, Samper JC, Jones AE, Oliver SA, Burnett KM, Wright RW. In vitro interactions of cryopreserved stallion spermatozoa and oviduct (uterine tube) epithelial cells or their secretory products. Anim Reprod Sci 1999;56:51–65. 20. Samper JC, Hellander JC, Crabo BG. Relationship between the fertility of fresh and frozen stallion semen and semen quality. J Reprod Fertil Suppl 1991;44:107–14. 21. Gadella BM, Miller NG, Colenbrander B, van Golde LM, Harrison RA. Flow cytometric detection of transbilayer movement of fluorescent phospholipid analogues across the boar sperm plasma membrane: elimination of labeling artifacts. Mol Reprod Dev 1999;53:108–25. 22. Colenbrander B, Brouwers JFMH, Neild DM, Stout TAE, da Silva P, Gadella BM. Capacitation dependent lipid rearrangements in the plasma membrane of equine sperm. Theriogenology 2002;58:341–5. 23. Piehler E, Petrunkina AM, Ekhlasi-Hundrieser M, TopferPetersen E. Dynamic quantification of the tyrosine phosphorylation of the sperm surface proteins during capacitation. Cytometry A 2006;69:1062–70. 24. Combes GB, Varner DD, Schroeder F, Burghardt RC, Blanchard TL. Effect of cholesterol on the motility and plasma membrane integrity of frozen equine spermatozoa after thawing. J Reprod Fertil Suppl 2000;56:127–32. 25. Moore AI, Squires EL, Graham JK. Adding cholesterol to
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the stallion sperm plasma membrane improves cryosurvival. Cryobiology 2005;51:241–9. Harrison RA, Miller NG. cAMP-dependent protein kinase control of plasma membrane lipid architecture in boar sperm. Mol Reprod Dev 2000;55:220–8. Rathi R, Colenbrander B, Bevers MM, Gadella BM. Evaluation of in vitro capacitation of stallion spermatozoa. Biol Reprod 2001;65:462–70. Thomas AD, Meyers SA, Ball BA. Capacitation-like changes in equine spermatozoa following cryopreservation. Theriogenology 2006;65:1531–50. Johannisson A, Morrell JM, Thoren J, Jonsson M, Dalin AM, Rodriguez-Martinez H. Colloidal centrifugation with Androcoll-E prolongs stallion sperm motility, viability and chromatin integrity. Anim Reprod Sci 2009;116:119–28. Varner DD, Bowen JA, Johnson L. Effect of heparin on capacitation/acrosome reaction of equine sperm. Arch Androl 1993;31:199–207. Neild DM, Gadella BM, Chaves MG, Miragaya MH, Colenbrander B, Aguero A. Membrane changes during different stages of a freeze-thaw protocol for equine semen cryopreservation. Theriogenology 2003;59:1693–705. Cheng FP, Gadella BM, Voorhout WF, Fazeli A, Bevers MM, Colenbrander B. Progesterone-induced acrosome reaction in stallion spermatozoa is mediated by a plasma membrane progesterone receptor. Biol Reprod 1998;59:733–42. Meyers SA, Overstreet JW, Liu IK, Drobnis EZ. Capacitation in vitro of stallion spermatozoa: comparison of progesterone-induced acrosome reactions in fertile and subfertile males. J Androl 1995;16:47–54. Cheng FP, Fazeli AR, Voorhout WF, Tremoleda JL, Bevers MM, Colenbrander B. Progesterone in mare follicular fluid induces the acrosome reaction in stallion spermatozoa and enhances in vitro binding to the zona pellucida. Int J Androl 1998;21:57–66. Fazeli AR, Steenweg W, Bevers MM, van den Broek J, Bracher V, Parlevliet J, Colenbrander B. Relation between stallion sperm binding to homologous hemizonae and fertility. Theriogenology 1995;44:751–60. Cross NL, Meizel S. Methods for evaluating the acrosomal status of mammalian sperm. Biol Reprod 1989;41:635–41. Silva PF, Gadella BM. Detection of damage in mammalian sperm cells. Theriogenology 2006;65:958–78. Farlin ME, Jasko DJ, Graham JK, Squires EL. Assessment of Pisum sativum agglutinin in identifying acrosomal damage in stallion spermatozoa. Mol Reprod Dev 1992;32:23–7. Cheng FP, Fazeli A, Voorhout WF, Marks A, Bevers MM, Colenbrander B. Use of peanut agglutinin to assess the acrosomal status and the zona pellucida-induced acrosome reaction in stallion spermatozoa. J Androl 1996;17:674– 82. Brum AM, Thomas AD, Sabeur K, Ball BA. Evaluation of Coomassie blue staining of the acrosome of equine and canine spermatozoa. Am J Vet Res 2006;67:358–62. Gadella BM, Rathi R, Brouwers JF, Stout TA, Colenbrander B. Capacitation and the acrosome reaction in equine spermatozoa. Anim Reprod Sci 2001;68:249–65.
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Chapter 154
Sperm Chromatin Structure Assay Aime K. Johnson and Charles C. Love
Introduction The ability to predict stallion fertility is vital in the equine industry and is often based on semen quality. Although numerous studies have attempted to find a single parameter that predicts fertility, a single test cannot accurately assess all factors involved in producing an embryo or a live foal. Sperm motility, sperm morphology, longevity of sperm motility, and determination of total sperm number are routinely performed in stallions to evaluate semen quality. Adjunct tests include acrosome assessment and membrane integrity. Flow cytometric analysis of individual sperm has been used to evaluate various sperm components, including DNA quality. The sperm chromatin structure assay (SCSA), introduced by Evenson et al.1 and Ballachey et al.,2 uses the metachromatic dye acridine orange (AO) to evaluate the proportion of single- and double-stranded DNA in sperm. While chromatin (DNA plus associated nucleoproteins) is in the title of the assay, only the DNA is actually evaluated. Depending on the type of DNA to which it is bound, AO will fluoresce either red (single-stranded DNA) or green (doublestranded DNA). Single-stranded DNA is associated with sperm unable to maintain a double helix. The SCSA can evaluate any type (fresh or frozen) of sperm. Evenson et al.1 refer to an important analytical end point as the DNA fragmentation index (DFI) to further clarify the function of the test. Briefly, a small aliquot of the sperm is exposed to acidic conditions, which tests the ability of the DNA to maintain the normal double helix. Following this stress, the addition of AO allows the measurement of the amount of green (normal DNA) and red (fragmented DNA) emitted from each spermatozoon. The fluorescent data are transformed into a scattergram with the rel-
ative amount of green fluorescence on the y-axis, and red on the x-axis (Figure 154.1). The majority of normal sperm (main population) will emit mostly green fluorescence and are located in an elliptical group in the upper left-hand portion of the graph. As the proportion of single-stranded DNA increases, sperm nuclei will emit more red fluorescence and be scattered across the x-axis outside the main population. The percentage of cells containing abnormal DNA (single-stranded) is represented as the percentage of cells outside the main population (%COMP) or DFI. The term alpha-t (αt) is created from the fluorescence values (green and red) from each sperm, and therefore uniquely identifies each spermatozoon based on its metachromic emission of those two colors. Figure 154.2 illustrates an actual scattergram from three different stallions showing low (10%), moderate (20%), and high (30%) COMPαt . The SCSA requires the use of a flow cytometer, thereby limiting routine field use. However, semen samples can be easily preserved (frozen) in the field and then transported to a reference laboratory for analysis, so the SCSA is an easily applied adjunctive test of sperm quality. Samples (0.5–1.0 mL) should be snap-frozen immediately after collection on dry ice or liquid nitrogen and shipped to the laboratory. Conventional straws used for commercial freezing are a space-efficient alternative. Common household freezers should be avoided for long-term storage because freeze–thaw cycles may iatrogenically damage sperm DNA. Fertility in stallions is often described as a bellshaped curve, with the majority of stallions in the population having a 45–75% per-cycle pregnancy rate (Table 154.1). Stallions in this majority may have similar semen parameters (morphology and motility), but have markedly different per-cycle pregnancy rates.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 154.1 The level of total sperm motility, percentage morphologically normal sperm, and percentage of cells outside the main population (%COMP), for three different fertility levels based on seasonal pregnancy rate and percentage pregnant per cycle (superscripts within rows are different; P < 0.05)
Scattergram (SCSA) Main population
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93a 82a 84a 69a 12a
86b 55b 74b 46b 17b
73c 39c 63c 46b 25c
Red fluorescence Figure 154.1 Characteristic scattergram used to evaluate the DNA quality using the SCSA. The main population identifies sperm with ‘normal’ DNA, while the percentage of cells outside the main population (%COMP) region identifies those sperm with easily denatured (‘abnormal’) DNA.
When the SCSA is utilized, a difference in %COMPαt between these groups indicates an increase in damaged DNA which may be the underlying cause of the fertility difference, despite other similar semen parameters.3, 4 This suggests that SCSA is useful in determining differences in stallions with varying fertility success and similar semen parameters. While DNA quality tends to decline along with other sperm quality measures with storage time or stress, these measures do not appear to decline at the same rate, suggesting that different sperm compartments have different sensitivities to stress. For example, sperm motility can decline as a result of both reversible (cooling) and irreversible (toxic) causes, yet morphology and DNA quality may remain high.
Interpretation of results
and %COMPαt , total motility, and percentage of morphologically normal sperm are listed for each of the three groups. The highly fertile stallion group (group 1) has an average %COMPαt of 12%, which is lower than groups 2 (17%) or 3 (25%). The fertility of groups 2 and 3 has the most potential to exhibit a further decline owing to environmental factors (semen handling) or intrinsic susceptibility to stress. Therefore, these stallions may not be as tolerant to variable semen handling and may benefit from a higher level of quality control that could result in higher fertility.
Use of SCSA for evaluation of cooled transported sperm The use of cooled transported semen is now widely available and is accepted by most breed registries. There is a wide variability between stallions with regard to fertility with the use of cooled transported semen. Reduced fertility in stallions following cooled transported semen may be due to a number of factors ranging from mare subfertility/infertility or mismanagement to poor semen quality or handling. We use the SCSA routinely when evaluating the suitability of a stallion’s semen for cooled transport. This 1000 GREEN 600 400
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Figure 154.2 Scattergrams representative of the fertility groups in Table 154.1. The percentage of cells outside the main population (%COMP) is 10%, 20%, and 35%, respectively.
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testing compares fresh extended semen against the same sample after 24 and/or 48 hours under cooled storage conditions. The expected result, if the semen has been processed optimally, is maintenance of DNA quality after 24–48 hours of cooled storage. If, however, the %COMP increases, the clinician should attempt to diagnose and treat the cause. A variety of factors have been reported to affect sperm parameters with cooled stored semen, including extender type, antibiotic type, storage temperature, and seminal plasma level. In most cases, as DNA quality declines, so does sperm motility; however, there are individuals that maintain sperm motility in the face of declining DNA quality.5 Therefore, in these cases of reduced fertility, the cause may be diagnosed only following SCSA evaluation. A common reason for a rise in the %COMP is an elevated (i.e., > 10–20%) seminal plasma level following dilution with an extender. Seminal plasma is a complex mix of substances designed to activate motility and protect the spermatozoa from damage from reactive oxygen species. However, prolonged storage in high concentrations of seminal plasma is detrimental to the DNA quality. This effect may be magnified in stallions in which semen quality is already compromised. The relationship between sperm motility and DNA quality was evaluated using ejaculates of known fertile stallions, and cooling them in varying concentrations of seminal plasma.5 After storage for 48 hours, the motility was not significantly different between samples stored in 0%, 10%, and 20% seminal plasma. However, the quality of the DNA declined as the amount of seminal plasma increased. Optimum seminal plasma levels in cooled transported semen may vary for each individual, but should be a minimum of 5% and should not exceed 20% in any stallion. In stallions that produce dilute ejaculates, centrifugation is necessary to increase sperm concentration and reduce the amount of seminal plasma; otherwise, the sperm from these individuals may be exposed to very high levels of seminal plasma. When evaluating multiple treatments (effect of extender, etc.) or ejaculates from the same stallion, our laboratory charges a flat fee for the stallion rather than a per sample fee. We feel this is important to determine the correct clinical approach to diagnosing a stallion’s fertility-limiting factor.
the study were significantly lower (i.e., high-quality DNA) than those from couples who achieved pregnancies in months 4–12 of the study or not at all. Additionally, all couples who had an SCSA value > 30% experienced delayed or no pregnancy. More recently, Bungum et al.7 reported that %COMPαt > 30% was associated with significantly lower fertility than SCSA values < 30%. There was also a statistical difference between fertilization rates in the use of intracytoplasmic sperm injection (ICSI) or in vitro fertilization (IVF) in men whose SCSA value was > 30%. In the stallion, our experience is modified by conditions that may not apply to the human model. While fertility certainly declines as %COMP reaches 30%, the deficit in DNA quality can, in some cases, be compensated for by increasing the insemination dose. In addition, while samples with < 30% COMP tend to exhibit higher fertility, we do detect a decline in fertility as %COMP goes from 10% to 20% to 30% (Table 154.1). Although stallions with 20% COMP may be considered ‘clinically fertile’, they tend to be less efficient in terms of cycles per pregnancy than stallions with 10% COMP. Also, stallions with a higher %COMP are also at more risk of being less tolerant of the cooling process as well as more susceptible to the detrimental effects of seminal plasma.
SCSA and embryo quality Although advances in assisted reproductive techniques (ARTs) have made the production of offspring from subfertile males possible, if the sperm DNA is damaged embryo quality and development is compromised. It appears that ARTs are able to compensate for poor sperm quality, including damaged DNA. However, normal fertilization (and cleavage rate) does not ensure that the spermatozoa contained normal DNA. In human males, those with higher SCSA values had a higher degree of embryo loss and miscarriage than normal fertile men.6, 8 The same is observed in the horse. Stallions with higher %COMPαt may have lower embryo survival and higher early embryonic loss both in vitro and in vivo than normal stallions (Hartman D., personal communication). Therefore, any stallion utilized for ARTs should have an SCSA as a routine part of the semen analysis to determine the potential for an increased rate of embryonic death.
SCSA and predicting fertility As assisted reproductive techniques are becoming more available, the ability to predict fertility is increasingly important. Evenson et al.6 used SCSA as a diagnostic tool to predict fertility in human males. In their study, SCSA values (%COMPαt ) from couples who achieved pregnancies in the first 3 months of
Conclusion The SCSA is an important adjunct sperm quality test when one is presented with a stallion with subfertile or reduced fertility. This test appears to correlate well with fertility and embryo development and may be independent, in some cases, of other parameters
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Sperm Chromatin Structure Assay such as sperm morphology and sperm motility. Low (normal) values for %COMPαt are not directly predictive of fertility potential, and analysis of other sperm quality parameters should be conducted (e.g., sperm motility, sperm morphology, longevity of sperm motility). Nonetheless, high (abnormal) values for %COMPαt are predictive of poor fertility, and, when using ARTs, reduced embryo survival. Because fertility depends on many factors, including sperm quality, mare quality, mare and stallion management, and sample handling, a single test may not identify the cause of reduced fertility. Sperm quality tests tend to be considered useful only when they correlate with the expected level of fertility; however, when sperm quality is normal in the face of reduced fertility/subfertility, it strongly suggests another source such as management or mare factors.
References 1. Evenson DP, Darzynkiewicz Z, Melamed MR. Relation of mammalian sperm chromatin heterogeneity to fertility. Science 1980;240:1131–3.
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2. Ballachey BE, Evenson DP, Saacke RG. The sperm chromatin structure assay: relationship with alternate tests of semen quality and heterospermic performance of bulls. J Androl 1988;9:109–15. 3. Love CC, Kenney RM. The relationship of increased susceptibility of sperm DNA to denaturation and fertility in the stallion. Theriogenology 1998;50:955–72. 4. Love CC. The sperm chromatin structure assay: a review of clinical applications. Anim Reprod Sci 2005;89:39– 45. 5. Love CC, Brinsko SP, Rigby SL, Thompson JA, Blanchard TL, Varner DD. Relationship of seminal plasma level and extender type to sperm motility and DNA integrity. Theriogenology 2005;63:1584–91. 6. Evenson DP, Jost LK, Marshall D, Zinaman MJ, Clegg E, Purvis K, de Angelis P, Claussen OP. Utility of the sperm chromatin structure assay as a diagnostic and prognostic tool in the human fertility clinic. Hum Reprod 1999;14: 1039–49. 7. Bungum M, Humaidan P, Axmon A, Spano M, Bungum L, Erenpreiss J, Giwercman A. Sperm DNA integrity assessment in prediction of assisted reproduction technology outcome. Hum Reprod 2007;22:174–9. 8. Acharyya S, Kanjilal S, Bhattacharyya AK. Does human sperm nuclear DNA integrity affect embryo quality? Indian J Exp Biol 2005;43:1016–22.
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Chapter 155
Spermatozoal Viability Stuart A. Meyers
In its most basic sense, a viable cell is a cell with an intact membrane utilizing oxygen and energy, and expiring carbon dioxide and other waste products. There is a multitude of other cell functions attributable to living cells, including the ability to maintain membrane potential, ion channel function and ion fluxes characteristic of a particular cell’s metabolism, production of ATP, and normal membrane integrity. A living cell is too complex for a single parameter to be used as a definitive viability marker. This is especially true in light of the sperm’s largely inactive nucleus. Clearly, should cellular processes stop, a series of changes attributable to apoptotic and/or necrotic cell death would result in a cell’s loss of viability. However, the loss of a detectable parameter may, or may not, reflect immediate cell death. For sperm, loss of flagellar motility leading to the cell’s inability to participate in fertilization in vivo essentially renders the sperm non-viable. However, a non-motile sperm could still be used to fertilize an oocyte in vitro using the technique of intracytoplasmic sperm injection (ICSI) and, as such, remains capable of fusion with the egg. However, certain parameters of cell viability may not coincide with the loss of sperm motility. Membrane and mitochondrial integrity may remain intact despite the loss of motility, thus assessment of these parameters will indicate the cells are viable. This is a very common scenario for cooled or frozen-thawed semen. For many somatic cell types and sperm, definitions of cell death may vary with different important parameters. Towards the end of a cell’s life cycle (for reasons of normal or induced apoptotic or necrotic insult) there are characteristic membrane changes that prevent the membrane from protecting the inner portions of the cell from the external environment. These changes include but are not limited to decreased membrane-associated energy utilization, ATP loss, glycolytic pathway failure, exchange of inner leaflet-associated phosphatidylserine to the
outer membrane leaflet, and phospholipid disorder. Thus, for the purpose of this chapter, the terms ‘viability’, ‘membrane integrity’, and ‘live’ will be used interchangeably since the concepts are overlapping and, ultimately, complex. Cell ‘viability’ then, as determined for a population of sperm cells, refers to the subpopulation of cells, within a larger population, that are ‘alive’, respiring, or otherwise functioning. Staining of living cells outside the body is known as ‘supravital’ staining and this method can be used to detect both living and non-viable cells. In the case of a stallion ejaculate, we are interested in the total number of ‘live’ cells within the entire sperm cell population. Clearly, if sperm cells are motile, they are viable; however, there are typically varying numbers of cells that are immotile yet demonstrate one or more other viability parameters. The difference between these two subpopulations may be quite considerable in the case of certain stallions whose sperm possess poor cooling or freezing ability. For these breeding stallions, it may be advantageous to determine viability using more than one method, focusing on different sperm structural or functional compartments. Cell viability is usually represented as a percentage of cells within a population, whether using microscopic individual cell counting or other automated methods of counting. For most cell viability determinations, the observer must use a method that distinguishes ‘live’ from ‘dead’ cells. Most current methods of determining cell viability are actually aimed at membrane integrity, since breakdown or loss of integrity of the membrane is a common marker for cell death. As an endpoint, however, it is not clear whether the loss of membrane integrity is an early or late change relative to loss of other cell functions. Clearly, cells undergoing early stages of cell death may still have adequate membrane integrity and would likely result in some falsepositive cells. In contrast, cells undergoing a loss of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Spermatozoal Viability membrane structure and function may display their loss of ‘viability’ prior to the complete loss of cell function, providing a false-negative population of cells. In either case, these may not fully reflect the amount of ensuing damage to a population of cells. In summary, the loss of membrane integrity is used to indicate the loss of a cell’s viability, but may not be wholly representative of cell viability. There are many commercially available methods for detection of membrane integrity as a measure of viability. Sperm viability can be a useful measure of functionally competent sperm when used to determine the size of the population of potentially viable cells within an ejaculate. For breeding or insemination purposes, sperm motility is widely considered the clinical ‘gold standard’ of cell viability since motile cells are essential for fertilization to occur in the mare’s genital tract. However, motility has not been highly correlated to fertility in all stallions and may not be the ultimate indicator of spermatozoal competence.1, 2 However, sperm viability/membrane intactness could be very useful to estimate the impact of a treatment, such as a cooling or freezing protocol. 2 Noxious exposures could also compromise sperm function and this category includes contamination from soap or water during preparation of an artificial vagina, bleach or iodine contamination, or even cold or osmotic shock. Although sperm motility would likely be affected by these exposures, membrane permeability or intactness could be a subclinically important finding since the population of motile cells is most certainly a subpopulation of the viable cells. Three basic methods for assessing sperm cell viability have been reported from numerous laboratories worldwide: (i) a cell smear air-dried on a glass slide, stained and viewed using light microscopy; (ii) fluorescence microscopy as a wet- (cover-slipped) or dry-mounted glass slide(without coverslips); or (iii) flow cytometry, the latter a more recent development. The wet mount preparations are most simple to use, particularly in the field or in the semen laboratory, while flow cytometry requires sophisticated equipment containing laser optics as well as extensive training on such equipment. Frequently, a hemacytometer-based cell count is performed in conjunction with viability stains, especially with fluorescence microscopy. The dried cell or smear method is also simple to perform but results can vary with conditions such as age of sample, humidity, temperature, and staining pH.
Non-fluorescence methods Most supravital viability staining has been based on ‘exclusion’ in that the dye may only enter cells if there is damage to the membrane or to vital cellular
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processes. A very common method for determining cell viability in vitro is that of trypan blue exclusion staining. Normal viable cells exclude the anionic trypan blue from crossing the plasma membrane, thus ‘dead’ cells are blue in color because the cell’s cytosolic compartment has been exposed to the dye. The dye requires a damaged membrane to gain entrance to a cell’s cytoplasmic area. This method has been used routinely in cell culture systems for an enormous variety of somatic cell types and gametes. One problem with this method is that there is considerable variation in staining sperm, and therefore subjectivity, making interpretation of ‘viable’ or ‘non-viable’ sometimes difficult; trypan blue often overestimates true viability. A combination trypan blue viability dye and Giemsa acrosomal/morphology dye has been reported for use in stallions and appears to differentiate between live and dead cells, as well as being relative easy to use.3, 4 These authors also used the anion dye, Chicago sky blue, that is similar to trypan blue in structure, with comparable results. The well-known eosin–nigrosin (Hancock’s) cell viability stain has been shown to be a reliable indicator of sperm viability, in addition to having strong utility as a morphological stain. Differential staining of live and dead sperm has been used for decades in the bovine AI industry and was first reported for bovine spermatozoa in 1942.5 Blom6 and Swanson and Bearden7 worked out the combination of eosin and nigrosin as a useful method. It is not clear from the literature when this method was adapted for equine sperm viability but several studies have demonstrated its utility in this species.8–10 The method has been shown to be less reliable when used with diluted or cryopreserved stallion sperm likely due to its incomplete staining characteristics.9, 10 The latter contribute to varying degrees of subjectivity when interpreting or scoring staining patterns. Subsequently, a number of other dyes and fluorescence staining methods have emerged that utilize differential fluorescence between ‘live’ and ‘dead’ cells.
Fluorescence methods While bright-field microscopy is inexpensive and simple to use in a field situation, fluorescence microscopy has provided a method by which the basic exclusion assay can utilize a variety of fluorescent probes, thus providing the opportunity to assess large numbers of cells and several cell compartments simultaneously. Fluorescence probes can be used either with microscopy or with flow cytometry. A number of these probes intercalate with nucleic acids and fluoresce, accessing the nucleus by way of a damaged plasma membrane. The ultraviolet (UV) spectrum is utilized by a class of fluorescence dyes
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Stallion: Specific Techniques Used in the Semen Evaluation of the Stallion
Figure 155.1 Stallion spermatozoa stained using Hoechst 33258 dye under UV illumination. Non-viable cells take up the dye while viable cells remain unstained. (1000× oil immersion.) (See color plate 54).
Figure 155.2 Stallion spermatozoa stained using SYBR14 (green) and propidium iodide (red). In this method, all cells take up the SYBR14 and fluoresce green while non-viable cells fluoresce red or orange due to the influence of PI. (1000× oil immersion.) (See color plate 55).
known as the bis-benzimidines, or Hoechst stains, and the class varies by degree of cell membrane permeability. Hoechst No. 33258 is an impermeant UV dye that is excited at approximately 350nm UV light and emits a blue color at a maximum emission of 461nm (Figure 155.1). Hoechst No. 33342 has an identical excitation/emission spectrum but it contains an extra ethyl group that imparts lipophilic qualities that allow the dye to cross cell membranes. The former dye is well suited to cell viability/membrane integrity studies as it gains access to DNA only when the membrane is compromised, while the latter is more suited to staining all cells that contain DNA, live as well as dead cells. Both Hoechst 33258 and 33342 have a strong tendency to leak back out of stained cells within minutes to hours so cell evaluation must be performed immediately after staining. Fixation and storage at 4◦ C can minimize loss of nuclear staining for longer periods, but alternative dyes have more recently emerged with improved staining efficiency over time. One of these latter dyes is propidium iodide (PI) and it has been demonstrated to identify viable/non-viable cells (Figure 155.2) because PI stains only structures containing double helix nucleic acids.11 Papaioannou et al.12 determined that PI staining of stallion sperm was essentially equivalent to that of Hoechst 33258 but remained inside the cell longer. Propidium iodide may be more stable than the Hoechst dyes and remains in cell nuclei for days to weeks following staining depending on storage conditions. Storage at 4◦ C provides optimum stability in unfixed cells for typically 1–2 weeks; fixed cells maintain longer fluorescence if fixation occurs immediately following staining. Fixatives such as paraformaldehyde, glutaraldehyde, 10% buffered formalin, or 95% ethanol have been used for sperm fixation. An additional advantage of
PI over Hoechst stain is that, with flow cytometry, PI–stained cells can be evaluated using a routinely available argon ion laser flow cytometer. Hoechst stains require an ultraviolet laser that is significantly more costly and less commonly available than the argon ion laser typical of most flow cytometer facilities. The flow cytometer method for differentiating sperm viability is preferable to counting cells manually because it allows for many thousands of cells (> 10 000) to be evaluated with statistical precision and accuracy while fluorescence microscopy typically allows smaller numbers of cells (e.g., 100–500) to be manually counted and assessed. As such, flow cytometric assessment of sperm cell viability (as well as other sperm parameters) has become an increasingly useful tool for reference laboratories and for institutional semen laboratories, although some veterinary practices have added this technology to their diagnostic armamentarium. In addition to the established viability probes, there are numerous newer and commercially available dyes with varying permeabilities and fluorescence spectra that demonstrate promise for cell viability assessment. Double- or triple-staining fluorescence methods, in which viable and non-viable cells can be identified simultaneously, have also been reported for use in stallion sperm. This staining method can be used both for microscopic and flow cytometric evaluations.13, 14 The most common combination is the membrane-permeant SYBR14 in combination with the membrane-impermeant PI dye. This combination is commercially available individually or as a kit (Invitrogen, Inc.). SYBR14 permeates all sperm and fluoresces in the green color spectrum while the PI permeates only membrane-damaged/dead sperm and fluoresces in the red. This has been used by numerous authors for studies in stallion sperm.9, 10, 15
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Spermatozoal Viability When incorporated into cells, the PI overpowers and displaces the green SYBR14 such that only the live cells are displayed as green while only the dead cells are displayed as red or orange in appearance (Figure 155.2). Further, another method has been used to determine the relationship between equine sperm viability and mitochondrial function. Since sperm motility is reliant upon mitochondrial function, several fluorescent stains have been utilized to assess the mitochondrial compartment of sperm. In particular, the fluorochrome rhodamine (R123) was used to demonstrate that only functional mitochondria may accumulate this dye in an energy-requiring process. Papaioannou et al.12 further demonstrated that this method is highly reliable for determining that spermatozoa with functioning mitochondria were very highly correlated with the percentage of viable (PI negative) sperm cells. The use of SYBR14 and PI has been coupled with the use of another mitochondrial fluorescence stain, JC-1, in order to determine a flow cytometric relationship amongst motile sperm cells and cell viability/membrane integrity. JC-1 (5,50,6,60-tetrachloro1,10,3,30-tetraethylbenzimidazolyl carbocyanine iodide) labels mitochondria differentially with high and low mitochondrial membrane potential (MMP) with differing colors (red-orange and/or green fluorescence). Although mitochondria can have areas that have both high and low MMP, flow cytometry can be used to quantitate the summation of the two colors and characterize cells as either an overall high MMP or low MMP.16 Since motile cells cannot be determined as such using the flow cytometer, Love and coworkers17 determined that high MMP correlates very highly with sperm motility in both fresh and cooled, extended stallion semen. An additional mechanism for identifying viable cells is the use of esterase substrate dyes. These can be used in combination with PI or other nucleic acid stains for sperm cell viability. Since all living cells contain esterase enzymes, a number of dyes that contain esterase substrates freely diffuse into cells. Once past the plasma membrane, these non-fluorescent substrates are converted by non-specific intracellular esterases into fluorescent products that are retained by cells with intact plasma membranes. Most of the esterase dyes have not been well worked out for use in stallion sperm and are reliant on cell loading properties such as permeability, pH, and temperature for optimum effects. Examples of these dyes are calceinAM (CAM), carboxyfluorescein diacetate (CFD), fluorescein diacetate (FD), and chloromethyl fluorescein diacetate (Cell Tracker Green, CFDA). Calcein-AM has been used with PI for viability of frozen stallion
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sperm and shown to be a reliable and useful method to distinguish viable cells.18
References 1. Jasko DJ, Little TV, Lein DH, Foote RI-I. Comparison of spermatozoal movement and semen characteristics with fertility in stallions: 64 cases (1987–1988). J Am Vet Med Assoc 1992;200:979–85. 2. Kirk ES, Squires EL, Graham JK. Comparison of in vitro laboratory analyses with the fertility of cryopreserved stallion spermatozoa. Theriogenology 2005;64(6):1422–39. 3. Kovacs A, Foote RH. Viability and acrosome staining of bull, boar and rabbit spermatozoa. Biotech Histochem 1992;67:119–24. 4. Kutvolgyi G, Stefler J, Kovacs A. Viability and acrosome staining of stallion spermatozoa by Chicago sky blue and Giemsa. Biotechnic Histochem 2006;81(4–6):109–17. 5. Lasley JF, Easley GT, McKenzie FF. A staining method for the differentiation of live and dead spermatozoa. I. Applicability to the staining of ram spermatozoa. Anat Rec 1942;82:167. 6. Blom E. A one-minute live-dead sperm stain by means of eosin–nigrosin. Fertil Steril 1950;1(2):176–7. 7. Swanson EW, Bearden HJ. An eosin–nigrosin stain for differentiating live and dead bovine spermatozoa. J Anim Sci 1951;10:981–7. 8. Lagares MA, Petzoldt R, Sieme H, Klug E. Assessing equine sperm-membrane integrity. Andrologia 2000;32(3):163–7. 9. Merkies K, Chenier T, Plante C, Buhr M. Assessment of stallion spermatozoa viability by flow cytometry and light microscope analysis. Theriogenology 2000;54:1215–24. 10. Merkies K, Buhr MM. Stallion spermatozoa viability: comparison of flow cytometry with other methods. In: Heide Schatten (ed.) Germ Cell Protocols: Volume 2: Molecular Embryo Analysis, Live Imaging, Transgenesis, and Cloning. Humana Press, 2004; pp. 49–59. 11. Biggionera M, Biggionera F. Ethidium bromide and propidium iodide -PTA staining of nucleic acids at the electron microscopic level. J Histochem Cytochem 1989;37:1161–6. 12. Papaioannou KZ, Murphy RP, Monks RS, Hynes N, Ryan MP, Boland MP, Rachel JF. Assessment of viability and mitochondrial function of equine spermatozoa using double staining and flow cytometry. Theriogenology 1997;48:299–312. 13. Garner DL, Pinkel D, Johnson LA, Pace MM. Assessment of spermatozoa1 function using dual fluorescent staining and flow cytometric analysis. Biol Reprod 1986;34:127–38. 14. Garner DL, Johnson LA. Viability assessment of mammalian sperm using SYBR14 and propidium iodide. Biol Reprod 1995;53:276–84. 15. Linfor JJ, Meyers SA. Assessing DNA integrity of cooled and frozen-thawed equine sperm using single cell gel electrophoresis. J Androl 2002;23(1):107–13. 16. Gravance CG, Garner DL, Baumber J, Ball BA. Assessment of equine sperm mitochondrial function using JC-1. Theriogenology 2000;53(9):1691–703. 17. Love CC, Thompson JA, Brinsko SP, Rigby SL, Blanchard TL, Lowry VK, Varner DD. Relationship between stallion sperm motility and viability as detected by two fluorescence staining techniques using flow cytometry. Theriogenology 2003;60(6):1127–38. 18. Akcay E, Reilas T, Andersson M, Katila T. Effect of seminal plasma fractions on stallion sperm survival after cooled storage. J Vet Med A 2006;53:481–5.
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Section (h): Surgical Procedures of the Male Reproductive Tract
156.
Clinical Uses of Testicular Biopsy Terry L. Blanchard
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Testicular Biopsy Walter R. Threlfall
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Cryptorchid Castration Thomas M. Russell and Patrick J. Pollock
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Inguinal Hernia Patrick J. Pollock and Thomas M. Russell
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Laparoscopic Surgery David G. Wilson
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Laser Assisted Castration Scott E. Palmer
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Normal Field Castration Glenn P. Blodgett
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Surgery of the Penis and Prepuce James Schumacher and Dickson D. Varner
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Chapter 156
Clinical Uses of Testicular Biopsy Terry L. Blanchard
Introduction Testicular biopsy has been recommended by some investigators as a routine diagnostic procedure to investigate certain types of reproductive problems in men. This includes screening for tumors in retained and scrotal testes, post-testicular obstruction causing azoospermia or oligospermia, and hypogonadism, including testicular degeneration.1, 2 Other laboratory methods often identify the cause(s) of reproductive dysfunction, so testicular biopsy may only have value for confirming an untreatable condition and thus be of academic interest only.1 Certainly, the strength of this diagnostic technique lies in the ability to actually visualize, or perform specific tests directly on, testicular parenchyma, which can be provided only by sampling the testis. Appropriately fixed specimens allow one to determine qualitatively whether spermatogenesis is complete by observing formation of elongated spermatids or the presence of stage VIII seminiferous tubules with elongated spermatids lining the lumen and in the process of being released.3 Critical quantitative analysis requires large sections of testicular parenchyma that can be obtained by a surgically obtained incisional biopsy. Parenchymal samples obtained with punch biopsy instruments are usually too small to allow accurate quantitation of spermatogenesis, because insufficient tubules are obtained to adequately represent the cycles of the seminiferous epithelium during spermatogenesis. Nevertheless, punch-type testicular biopsies can be used to qualitatively evaluate the level of spermatogenesis and determine which components (e.g., interstitium and Leydig cells, seminiferous tubules with Sertoli cells and spermatogenic cells) may be abnormal.
Punch biopsy specimens have been demonstrated to be of sufficient size and quality to permit assay of testicular parenchyma for proteins, testosterone, estrogens, inhibin, and insulin-like growth factor 1.4 Roser4 proposed that determination of actual hormone and growth factor concentrations in the testis may elucidate mechanisms of testicular function that could otherwise not be studied. Assay of parenchymal specimens and blood harvested from stallions at abattoirs has shown that intratesticular concentrations of testosterone in the testis greatly exceeds that in the peripheral circulation,5 and intratesticular concentration of testosterone is better associated with level of spermatogenesis than is circulating testosterone level.5, 6 While Roser4 suggested that a single punch biopsy sample provides protein and hormone concentrations that represent the entire testis, sampling parenchyma from three areas of each testis obtained from pony stallions at castration yielded values too variable (both between locations within the same testis and between testes) in concentrations of testosterone and estradiol-17β to be useful in correlating them to the level of spermatogenesis and germ cell degeneration rate (T.L. Blanchard, D.D. Varner, L. Johnson, 1998, unpublished observations). Potential adverse side-effects that sometimes occur as a result of testicular biopsy include hemorrhage, hematoma formation, orchitis, localized testicular inflammation, fibrosis, adhesion formation between testicular tunics, sperm granuloma, degeneration of germ cells in seminiferous tubules, and perhaps formation of sperm antibodies.1, 2, 7, 8 In a study of short-term (27 days) effects in stallions, testicular biopsy caused a transient degeneration of type B spermatogonia and preleptotene spermatocytes.7 A decrease in the size of the biopsied testis was also
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Stallion: Surgical Procedures of the Male Reproductive Tract
noted when compared with the non-biopsied testis. The stallions were castrated at the end of the study. Histopathological examination revealed damage to spermatogenic cells in the seminiferous epithelium, which correlated with the timing of the biopsy. Threlfall and Lopate8 reviewed the literature on the outcome of testicular biopsy in animals and men and suggested that the main complication would be a decrease in sperm output (for several weeks or months), but predicted seminal quality would return to pre-biopsy levels given adequate time. Brief mention is made of aspiration biopsy, which is purported to be potentially less damaging than punch biopsy, but will not provide as much information regarding spermatogenesis. Aspiration biopsy may be helpful in differentiating between causes of testicular enlargement (e.g., neoplasia, trauma, or septic orchitis). The technique, which uses a 23- or 25-gauge needle and 12-mL syringe, has been described by Threlfall and Lopate.8 Criteria used for identifying testicular cells obtained by fine needle aspirates have been published by Leme and colleagues.9, 10 Owing to the simplicity and relative safety in procuring a punch biopsy from the testis and the quality of parenchyma that can be obtained for evaluation, the author prefers this technique for obtaining testicular biopsies from stallions.
Technique for obtaining punch biopsy from the testis The procedure is done with the stallion standing, having first been sedated with xylazine hydrochloride and butorphanol tartrate. After aseptic preparation of the scrotal skin, the testis to be biopsied is positioned laterally with the scrotal skin stretched tightly over its surface, and with the testis isolated entirely within one hand of the operator. A springloaded biopsy instrument (Bard Monopty Biopsy Instrument, C. R. Bard, Inc., Covington, GA; Figures 156.1–156.3) is removed from its sterile packaging and the needle is placed against the scrotal skin in
Figure 156.2 End of biopsy needle of the Bard biopsy instrument, with the biopsy window shown.
the desired location for biopsy procurement (Figure 156.4). Needle location for insertion should be in the center of the craniolateral quarter to third of the testis to avoid lacerating a testicular artery (Figures 156.4 and 156.5).11 The biopsy needle guide is then forcefully pushed through the scrotal skin and testicular tunics (a pronounced ‘popping’ sensation can be felt), just into the outer testicular parenchyma. The thumb is used to push the button trigger of the biopsy instrument, which will quickly cause the notched needle to penetrate into the center of the parenchyma and retrieve tissue automatically back into the needle guide. It is important to hold the testis firmly throughout the procedure so the parenchyma is not pushed away during procurement of the biopsy. The biopsy instrument is then withdrawn from the testis, tunics, and scrotum, and firm pressure is placed on the biopsy site for approximately 30 seconds to control any hemorrhage. The biopsy specimen should be immersed into fixative solution and gently teased off the instrument so that tissue architecture is minimally affected. A variety of fixatives can be used, depending on the choice of the examiner. The author routinely uses Bouin’s solution for 24 hours, followed by immersion in 10% formalin if shipped to a diagnostic laboratory, or in alcohol if it is taken immediately to a histology laboratory. If special studies (e.g., terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) stain for apoptosis) are to be performed, it is best to fix the tissue in 4% paraformaldehyde to improve nick-end labeling of damaged DNA. After fixation, the tissue is embedded in paraffin,
Bard Monopty Biopsy Instrument C. R. Bard, Inc. Covington, GA 30014 800-526-4455
14 ga × 16 cm length; penetration 22 mm; sample notch 1.7 cm Figure 156.1 Bard biopsy instrument used for procurement of testis biopsy.
Figure 156.3 Testicular parenchymal sample obtained during the biopsy procedure with the Bard biopsy instrument.
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Clinical Uses of Testicular Biopsy
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biopsy site
Figure 156.4 Left testis removed by castration. The testis is positioned with the lateral aspect uppermost, and the tail of the epididymis to the right. Site for biopsy procurement is indicated, and should be in an area with few superficial blood vessels.
sectioned, and stained (most commonly) with periodic acid–Schiff–hematoxylin (PAS–hematoxylin). Light microscopic examination of a normal testicular biopsy will reveal Leydig cells in the interstitium and seminiferous tubules of normal appearance (basal lamina lined with Sertoli cells and spermatogonia, with typical early and late primary spermatocytes, round spermatids, and elongating spermatids evident as germ cells progress toward the lumen of the tubule) (Figure 156.6). Stage VIII seminiferous tubules (i.e., lumen of tubules lined with elongated spermatids being released; Figure 156.7) represent up to 15% of tubules in normal stallion testes,12 and their presence can be used to confirm that spermatogenesis is proceeding to completion. TUNEL staining for apoptotic germ cells can be performed on testes thought to be undergoing degeneration (Figure 156.8). The incidence of apoptotic germ cells has been
Figure 156.5 Testis and biopsy instrument positioned for collection of testicular biopsy.
Figure 156.6 Testis biopsy obtained with the Bard biopsy instrument. Periodic acid–Schiff–hematoxylin stain. (See color plate 56).
shown to be well correlated with the rate of degeneration of germ cells in the stallion.13
Some indications for procurement of testicular biopsy The most common conditions viewed by the author that may dictate the need for a testicular biopsy are azoospermia or oligospermia. Azoospermia can be a diagnostic dilemma, as the most common cause of this condition is failure to ejaculate. Diagnostic efforts are directed first toward determining whether the stallion is actually ejaculating. In the stallion with normal libido and breeding behavior, indirect evidence for ejaculation includes engorgement of the glans penis (usually occurs at maximal erection just prior to and after ejaculation), urethral pulsations (typically 3–9 pronounced pulsations of the urethra palpable on the ventrum of the base of the penis), flagging of the tail, and the presence of gel in the sample collected.14, 15 After removal of the gel portion, the
Figure 156.7 Normal stage VIII seminiferous tubule with elongated spermatids lining the lumen in preparation for release. (See color plate 57).
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Stallion: Surgical Procedures of the Male Reproductive Tract
Figure 156.8 Seminiferous tubule of a young stallion treated with dexamethasone, and biopsied at intervals to monitor the incidence of apoptotic germ cells (brown staining). TUNEL stain. (See color plate 58).
gel-free liquid is examined by using a phase-contrast microscope at 200–400× magnification for the presence of spermatozoa and premature germ cells. If no spermatozoa are present, assay of the gel-free liquid for alkaline phosphatase is performed to confirm the presence of epididymal/testicular fluids (6913–28, 180 U/L in ejaculates of normal stallions).16 High alkaline phosphatase values confirm the presence of epididymal/testicular secretions in the seminal fluid, and thereby suggest a testicular problem. Testicular biopsy can then be performed to determine whether spermatogenesis fails to proceed to completion, as in spermatogenic failure. Severe testicular hypoplasia or degeneration can cause azoospermia due to failure of spermatozoal production by the seminiferous epithelium (i.e., spermatogenesis may not progress through spermiogenesis, resulting in failure to produce elongated spermatids) (Figure 156.9). Low alkaline phosphatase levels do not rule out a testicular problem, but confirm that testicular and epididymal secretions are not entering the ejaculate, perhaps because of obstruction of the extragonadal duct lumina for spermatozoal passage, and failure of emission (release of sperm and accessory gland fluids into the pelvic urethra) or ejaculation (forceful expulsion of the combined fluids from the urethra).14, 15 Complete obstruction of the epididymal or deferent ducts also results in azoospermia, as sperm produced by the testes cannot pass through the excurrent duct system to enter the urethra for ejaculation. Spermatozoa occasionally become impacted within the deferent duct lumina to the point that they are not released by the strong muscular contractions of the ducts at the time of ejaculation. The condition is often referred to as ‘plugged ampullae’ when ampullary luminae become obstructed. Perhaps the most common cause
Figure 156.9 Testicular parenchyma from azoospermic stallion with ejaculates containing only premature germ cells. Spermatogenesis was not proceeding through spermiogenesis. Some tubules were present that contained a few round spermatids, but elongating spermatids were not seen. Tissue fixed in 2% glutaraldehyde and stained with Periodic acid–Schiff–hematoxylin.
of epididymal obstruction is chronic epididymitis, which usually is detectable by palpation of firm, enlarged, sometimes irregular epididymides that are adhered to testicular tunics. Trans-scrotal ultrasonography typically reveals enlarged distended hypoechoic epididymal tubules proximal to the obstruction, and is frequently accompanied by mild hydrocele. Inflammatory cells may not be detected in ejaculates of horses with chronic epididymitis, in contrast to the numerous degenerate neutrophils with relatively few spermatozoa in ejaculates of horses with acute epididymitis. In such cases, testicular biopsy could be used to confirm that the problem does not lie in the testes, as spermatogenesis would be proceeding to completion at normal rates. Oligospermia, in conjunction with a high incidence of abnormal sperm and premature germ cells in the ejaculate, is common in cases of advanced testicular degeneration.14, 17 Testicular biopsy will confirm the degeneration (Figure 156.10) and reduced spermatogenic efficiency, which sometimes is so severe that only Sertoli cells and spermatogonia (in reduced numbers) remain lining the seminiferous tubules.
Case illustrations To illustrate this approach, three cases of stallions with azoospermia are presented. A 5-year-old stallion with slightly small testes appeared to ejaculate normally with gel being present in his ejaculates, but the ejaculate contained only premature germ cells (0.5 million/mL). Assay of seminal plasma from this
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Figure 156.10 Testis biopsy from a stallion with severe oligospermia and a high incidence of premature germ cells present in the ejaculate. Seminiferous tubule is prominently vacuolated with degenerating germ cells present, some being desquamated into the lumen. Some tubules were present that contained a few elongated spermatids. (See color plate 59).
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per rectum revealed normal size and location of the paired ductus deferens and ampullae, seminal vesicles and lobes of the prostate.20 A testicular biopsy was taken and revealed seminiferous tubules of normal appearance (basal lamina lined with Sertoli cells and spermatogonia, with normal appearing early and late primary spermatocytes, round spermatids, and elongating spermatids present as germ cells advanced in development toward the lumen of the tubule). Several stage VIII seminiferous tubules were present. The finding that spermatogenesis proceeded to completion, in conjunction with the azoospermia and low concentration of alkaline phosphatase in the gel-free semen, suggested obstruction of the extragonadal ducts, preventing sperm from entering the ejaculates. The presence of the extremely small epididymides in a young stallion likewise suggested epididymal hypoplasia (failure of epididymides to develop to normal size).
Summary stallion yielded an alkaline phosphatase concentration of 1130 U/L, indicating testicular/epididymal contribution to the ejaculate. Testicular biopsy revealed spermatogenesis was not proceeding to completion (seminiferous tubules of smaller than normal diameter with no elongating or elongated spermatids, and only a few round spermatids were present). Another 7-year-old stallion with small testes ejaculated gel and gel-free semen containing only premature germ cells (6.5 million/mL). Biopsy confirmed spermatogenesis was not proceeding to completion and the horse was castrated. Histomorphometric analyses of fixed testicular parenchyma18 revealed that spermatogenic arrest occurred between meiosis and spermiogenesis (i.e., counts of late primary spermatocytes yielded a daily sperm production estimate of 30 million/g, while greatly reduced numbers of round spermatids yielded an estimate for daily sperm production of less than 2 million/g). Only occasional elongating spermatids were seen in the seminiferous tubules of this horse. Finally, a 3-year-old stallion with slightly small testes was examined that ejaculated gel and gel-free semen with no sperm or germ cells in the ejaculate. Assay of seminal plasma from this stallion yielded only 43 U/L alkaline phosphatase, confirming that testicular/epididymal secretions were absent from the ejaculate. The epididymides were quite small, particularly where the body (corpus epididymis) joins the tail (cauda epididymis), and were uniformly echogenic – without the ‘Swiss cheese’ appearance attributed to the convoluted epididymal duct containing fluid19 on ultrasound examination. Palpation
While clinical findings and other laboratory methods often identify the cause of reproductive dysfunction in the stallion, in some cases testicular biopsy may provide useful information regarding the contribution of the testes to the problem. Besides providing a diagnosis, testicular biopsy analysis can aid one in determining whether a condition is likely to be treatable or reversible.
References 1. Glezerman M. Testicular biopsy. In: Bandehauer K, Frick J (eds) Disturbances in Male Fertility. Berlin: Springer-Verlag, 1982; pp. 215–23. 2. Colbern M, Wheeler T, Lipshultz L. Testicular biopsy. Its use and limitations. Urol Clin North Am 1987;14:551–61. 3. Blanchard TL, Varner DD. Evaluating breeding soundness in stallions. 4. Hormonal assay and testicular biopsy. Vet Med 1996;91:358–365. 4. Roser JF. Idiopathic subfertility/infertility in stallions: new endocrine methods of diagnosis. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1995, pp. 136–46. 5. Johnson L, Thompson Jr DL. Effect of seasonal changes in Leydig cell number on the volume of smooth endoplasmic reticulum in Leydig cells and intratesticular testosterone content in stallions. J Reprod Fertil 1987;81:227–32. 6. Blanchard TL, Johnson L. Increased germ cell degeneration and reduced germ cell:Sertoli cell ratio in stallions with low sperm production. Theriogenology 1997;47:665–78. 7. DelVento VR, Amann RP, Trotter GW, Veeramachaneni DNR, Squires EL. Ultrasonographic and quantitative histologic assessment of sequelae to testicular biopsy in stallions. Am J Vet Res 1992;53:2094–101. 8. Threlfall WR, Lopate C. Testicular biopsy. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 943–9.
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9. Leme DP, Papa FO. Cytologic identification and quantification of testicular cell types using fine needle aspiration in horses. Equine Vet J 2000;32:444–6. 10. Leme DP, Papa FO, Trinca LA. Assessment of seasonal influence on equine spermatogenic cells and Sertoli cells by testicular fine needle aspiration cytology. Proceedings of the 8th International Symposium on Equine Reproduction. Theriogenology 2002;58:269–72. 11. Smith JA. Biopsy and the testicular artery of the horse. Equine Vet J 1974;6:81–3. 12. Swierstra EE, Pickett BW, Gebauer MR. Spermatogenesis and duration of transit of spermatozoa through the excurrent ducts of stallions. J Reprod Fertil Suppl 1975;23:53–7. 13. Forrest DW, Blanchard TL, Heninger NL, Varner D, Roser JF, Johnson L. Relationship of germ cell apoptosis to spermatogenic efficiency in stallions with reduced semen quality. Anim Reprod Sci 2006;94:140–3. 14. Varner DD, Schumacher J, Blanchard TL, Johnson L. Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991; pp. 54, 166–8.
15. McDonnell SM. Ejaculation: physiology and dysfunction. Vet Clin North Am Equine Pract 1992;8:57–70. 16. Turner RM. Alkaline phosphatase activity in stallion semen: characterization and clinical implications. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1996; pp. 284–93. 17. Blanchard TL, Johnson L, Varner DD, Rigby SL, Brinsko SP, Love CC, Miller C. Low daily sperm output per ml of testis as a diagnostic criteria for testicular degeneration in stallions. J Equine Vet Sci 2001;21:33–5. 18. Johnson L, Varner DD, Thompson Jr DL. Effect of age and season on the establishment of spermatogenesis in the horse. J Reprod Fertil Suppl 1991;44:87–97. 19. Love CC. Ultrasonographic evaluation of the testis, epididymis, and spermatic cord of the stallion. Vet Clin North Am Equine Pract 1992;8:167–82. 20. Kenney RM, Hurtgen JP, Pierson R. Theriogenology and the equine. Part II. The stallion. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1983, pp. 1– 100.
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Chapter 157
Testicular Biopsy Walter R. Threlfall
Reports on the causes of infertility and subfertility of a general nature are not available for most domestic species. However, in humans, Tomaszewski1 reported that 40% of infertility problems are of male origin, 50% of female origin, and 10% are a combination of male and female disorders. The percentage of domestic animals with infertility or subfertility caused by the male is more variable and often is very obvious when the male breeds large numbers of females. Many factors can cause infertility or subfertility in the male. One method of classification is by pretesticular, testicular, and post-testicular causes.2–4 Examples of pre-testicular causes include maturation arrest, hypospermatogenesis, chromosomal abnormalities, cryptorchidism, and orchitis. Post-testicular causes include blockage of ducts leaving the testis, defective spermatozoal maturation or storage in the epididymis, and biochemically abnormal seminal plasma. A blockage close to the testis increases the severity of the lesion.5 In addition, extratesticular causes of reduced fertility can occur, which include infections, excessive heating of the genital tract, and toxic chemicals.2, 3 Veterinarians are generally asked to examine the subfertile stallion following failure of mares to conceive. The sequence of the evaluation involves obtaining a complete history and then performing a breeding soundness examination. Medical history includes viral and bacterial infections, traumatic injury, and general health status (past and present). History of fertility includes past pregnancy rates, seminal evaluations, and libido. Medications administered might include those related to past or present fertility treatment, systemic or localized body infections, use of androgenic steroids, or use of the antigonadotropin-releasing hormone vaccine. Palpation of the testes, epididymides, vas deferentia, spermatic cords, accessory sex glands, and penis should be per-
formed. Seminal analysis should be performed on more than one occasion. Blood chemistries and serum hormonal concentrations, if indicated, should be performed. The semen examination includes assessment of spermatozoal motility, concentration, morphology, and volume of the ejaculate, plus notation of any aberrant cellular or fluid component. Depending on the results of this examination and the endocrine profiles, sequential examinations over as long as a 2-month period, testicular ultrasonography, spermatozoal antibody titers, karyotyping, and testicular biopsy may be indicated. Any of the aforementioned tests may be indicated either individually or in combination, depending on the abnormalities found.
Methods of evaluation Endocrine profiles Endocrine profiles should include multiple samples over time for quantitative testosterone assays, estrogens, inhibin, luteinizing hormone (LH), folliclestimulating hormone (FSH), and thyroid hormones. Serum concentrations of FSH and inhibin provide the most useful information regarding testicular disorders. In cases of azoospermia in man, FSH concentrations are elevated in patients exhibiting signs of tubular hyalinization and Sertoli cell-only syndrome and in 50% of patients showing maturation arrest. In azoospermatic human patients with normal testicular biopsies and normal FSH concentrations, complete ductular obstruction is suspected.6, 7 Partial excurrent duct obstruction is likely if FSH concentrations and the testicular biopsy are normal in an oligospermic patient.6, 7 In cases of severe oligospermia (0.5 million cells/mL) with elevated FSH concentrations, severe hypospermatogenesis is the most frequently seen lesion and the prognosis
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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is poor.6–9 Obstructive causes of infertility in the human can often be surgically repaired.7 The relationship between circulating hormonal values and fertility has been reported for stallions. Circulating concentrations of FSH tend to increase with a reduction in fertility, whereas circulating concentrations of estradiol, total estrogens, and inhibin tend to decrease with reducing fertility.10, 11 The lack of commercially available FSH assays for most domestic species is a definite disadvantage in male fertility examinations; however, these determinations are available at some diagnostic and teaching institutions. Ultrasonography The use of ultrasonography to delineate lesions within the testis has been described and may be beneficial in the stallion. Lesions such as sperm granulomas, spermatoceles, and neoplasia are easily visualized with B-mode ultrasonography. This technique is always indicated prior to more invasive procedures. Spermatozoal antibody determination If autoimmune orchitis is suspected, spermatozoal antibody titers should be determined. This determination is not easily accomplished owing to the lack of commercial availability of the test for use in horses. Karyotyping If genetic mutation is suspected, then karyotyping is indicated. Mutations can be associated with a visibly gross alteration in the normal appearance of the genital tract. Testicular biopsy Testicular biopsy has been used with seminal evaluation to help diagnose and classify human males with varying degrees of testicular failure.5, 12, 13 This combination is especially useful in veterinary medicine, because the commercial availability and economic practicality of hormonal assays is often limited. Testicular biopsy was first recommended in man to diagnose cases of obstructive versus non-obstructive azoospermia, to classify oligospermia, and to aid in the determination of treatment and prognosis. Hotchkiss performed the first clinical testicular biopsy in man in 1944.14 Testicular biopsy can aid in the determination of many disorders, including, but not limited to, germinal cell aplasia, germinal cell arrest, hypogonadotropic eunuchoidism, hypospermatogenesis, Klinefelter’s syndrome, and hypogonadotropism.2, 5, 14 Biopsy can also help dis-
criminate among inflammatory disorders, noninflammatory disorders, and neoplasia..2, 15 Biopsy may be indicated in cases of cryptorchidism, because testes allowed to remain retained tend to become dysfunctional and often fibrotic or neoplastic.2 For this reason, in man, at the time of orchiopexy, biopsy is often recommended. Orchiopexy as a treatment for cryptorchidism is not recommended in the stallion because of the possible inheritance of this condition. Many researchers have reported that, if the testes are of equal size and consistency, unilateral biopsy is sufficient.5, 13, 16, 17 In such cases in man, biopsy specimens as small as 10 mg from the right testis have provided representative quantitative values for both testes, and the quality and quantity of sperm production was found to be uniformly distributed throughout the testicular parenchyma.18 Shakkebaek and Heller19 found no significant difference between left and right testes in man with quantitative histological analysis. Unilateral biopsy of the left testis has provided accurate information in 85% of the men regarding sperm production in the opposite testis because of the uniformity of spermatogenesis from one testis to the other, if size and consistency are similar.5 Therefore, a biopsy sample from one testis provides an accurate histological diagnosis that represents the status of both testes in most but not all situations.18 However, bilateral biopsy provides pathologists with an idea of the generalization or focality of a lesion.
Histometric analysis The use of quantitative histometric analysis with testicular biopsy has shown significant positive correlation between the number of germ cells per seminiferous tubule or the number of Sertoli cells per tubule and the sperm concentration in the human ejaculate.18–20 There are two methods whereby histomorphometrics can be utilized. The first is the Sertoli cell-ratio method that allows quantification by comparison of the germinal epithelium with a reference cell, the Sertoli cell.19 The number of Sertoli cells per tubule rarely changes because of tissue fixation or preparation or because of pathological or therapeutic circumstances (i.e., hormone treatment or irradiation). In laboratory animals and man, Sertoli cells do not divide after sexual maturity. Therefore, comparison of germ cell with Sertoli cell numbers can provide an indication of dysfunction in spermatogenesis or spermiogenesis. This method provides less accurate results in situations in which tubules are so severely damaged that Sertoli cells are also damaged.19, 20 The second method is the cell-association method and permits quantification of germinal epithelium
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compared with seminiferous tubular circumference.20 This method can be used in the more severely damaged tubules with greater precision than can the Sertoli cell-ratio method. Comparison of numbers of each type of germ cell (and Sertoli cells) present with known standards of germ cells present in normal testes has permitted localization of a lesion to a specific cell type in the tubular epithelium. Correlations between spermatozoal output and patterns of spermatogenesis can also be made.
Value of testicular biopsy Biopsy has been shown to be of value in the determination of the cause, severity, and prognosis of a low spermatozoal count.5, 19, 20 Blockage may also be the cause of spermiostasis in stallions. Normal spermatogenesis in a tissue sample does not reflect the spermatozoal count.2, 19, 20 In man, histological changes ranging from normal spermatogenesis to Sertoli cellonly syndrome was observed in tissue sections of patients clinically demonstrating conditions from aspermia to oligospermia.17, 19, 20 Normal spermatogenesis can also be seen with surrounding abnormal supporting tissue.16 The relationship of testicular biopsy to stallion fertility has been reported. A 5-year-old azoospermic stallion with small testes had testicular biopsy results indicating that the seminiferous tubules had a smaller than normal diameter with no elongation of spermatids and few round spermatids present.21 Another stallion, aged 7 years, had small testes and only premature cells in the ejaculate. Histomorphometric analysis of the testes indicated disruption of spermatogenesis between meiosis and spermiogenesis.22 A third stallion with slightly small testes and small epididymides was biopsied, which revealed seminiferous tubules of normal appearance. Spermatogenesis proceeded to completion.21 In mink, testicular palpation, seminal examination, and testicular biopsy were all found to be practical and suitable methods of assessing male reproductive capacity. Serum testosterone quantities were found to be poorly correlated.23
Figure 157.1 Position of the testicle within the scrotum.
cutaneous tissue only with a locally acting anesthetic agent (Figure 157.1). An incision is made through the scrotum and tunica albuginea. The incision into the tunic should not be larger than 0.5 cm to minimize the trauma to the testis and its vasculature (Figure 157.2).13 The internal pressure pushes a small section of testicular tissue through the incision in the tunic. This tissue is gently cut with a razor blade and placed immediately into a fixative solution (Figure 157.3). The tunic and scrotum are closed with an appropriate suture (Figure 157.4). Split needle The second type of biopsy is the split needle method.12, 24 The scrotal area is clipped (optional) and
Biopsy techniques Three different biopsy techniques are used commonly in man and certain domestic and laboratory animals. Incisional or open The first and most common technique in human medicine is the incisional or open biopsy.2, 5, 20 This technique is also commonly used in small animals. The scrotal area is clipped and scrubbed. Anesthesia is accomplished by infiltration of the skin and sub-
Figure 157.2 Incision of tunica propria.
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Figure 157.3 Removal of testicular tissue that bulges through the tunica propria incision. Figure 157.5 Scrotal skin incision for passage of a split needle.
scrubbed. In this procedure, local anesthesia is appropriate for the scrotal skin and subcutaneous tissue only. The skin incision is 0.5 cm in length. The scrotum is incised with the point of a no. 10 blade, and the split needle is gently pushed through the tunic albuginea and into the testicular tissue (Figures 157.5 and 157.6). Puncture of the tunic is painless because no sensory neural pain fibers, which would respond to cutting normally, exist between the dermal layer and the tunica albuginea.14 In addition, extraction of the testicular tissue sample is also relatively painless because of the lack of innervation within the testicular parenchyma. The testicular tissue is obtained with the split needle and immediately and carefully placed in fixative using a small gauge hypodermic needle (Figures 157.7 and 157.8). The scrotal incision is medicated with a topical tissue adhesive, but not sutured. No systemic antibiotics are necessary after this procedure if asepsis was maintained. The split needle biopsy will provide tissue samples of approximately 1.0 mm in diameter and between 2.0 and 6.0 mm in length.14
Figure 157.4 Surgical closure of the tunica propria.
Fine needle aspiration The third type of biopsy is known as fine needle aspiration biopsy. It is the most recently described technique and is currently used in human medicine clinically and to some extent in veterinary medicine.5, 15, 25 After surgical preparation and anesthesia, the testis is percutaneously punctured with a 20- to 23-gauge needle.25, 26 A small portion of tissue is aspirated into a 5-mL syringe. The aspirate can be placed on a slide for cytological examination or placed in a fixative for histological examination. Technique summary The open biopsy technique provides the most tissue and, therefore, the most tubules for examination.5, 15 The split needle biopsy usually provides adequate tissue; however, the interstitium may be disrupted.5, 14, 15 The fine needle aspiration biopsy will usually provide few intact tubules; however, this technique provides germinal cells of all types,
Figure 157.6 Placement of the split needle biopsy instrument into the testicle.
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Figure 157.7 Advancement of the inner needle into the testicular tissue.
including Sertoli cells, and allows the diagnosis of the conditions of normal spermatogenesis and spermatogenic arrest.5, 15, 25, 26 A high correlation exists between the results of cytological examination and histological examination in regard to spermatogenic activity.26 Histological examination allows the distinction of interstitial changes whereas cytological examination does not.26 Fine needle aspiration biopsy is the least invasive of the three techniques, split needle biopsy is mildly invasive, and open biopsy is moderately invasive. The testicular biopsy technique recommended by the author for use in the stallion is as follows. The stallion is first tranquilized intravenously with 0.8 mg/kg of xylazine hydrochloride. Approximately 5 minutes after this administration, 0.66 mg/kg morphine or 0.22 mg/kg butorphanol tartrate is administered intravenously. The primary consideration of a testicular biopsy in any species is that of asepsis.
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Proper preparation of the surgical site is of extreme importance such that no bacteria are introduced into the testicle. The surgical area is prepared by first thoroughly cleansing with a detergent, cotton, and clean water. Because the procedure is performed in the standing position, the clinician must thoroughly cleanse the entire scrotum and the inner surfaces of the legs in the inguinal area. After cleansing, a surgical scrub is performed using providine–iodine or a similar preparation and cotton. After an approximate 10-minute scrub of the area, it is rinsed thoroughly with clean water, then rinsed a final time with an alcohol solution. The testicle to be biopsied is isolated within the scrotum and grasped firmly with a sterile gloved hand so that the skin over the testis is tight. The area of the skin incision, which generally is in the midportion of the testis on the free or most ventral border, is infiltrated with 2–3 mL of a local anesthetic. Anesthetic is placed in the skin and subcutaneous area, with a 23- to 25-gauge needle. No anesthetic should be placed within the testicle, because doing so can cause testicular degeneration. Once the anesthetic has been administered, the point of a no. 10 surgical blade is used to incise the anesthetized skin (Figure 157.5). The approximate length of the incision is 0.5 cm. A Tru-Cut (Travenol Laboratories, Inc., Deerfield, IL), a Monoject (Sherwood Medical, St. Louis, MO), or Bard (Bard Urology, Covington, GA) split needle biopsy instrument is used. The needle is inserted through the skin incision and forced with sharp pressure through the tunica albuginea surrounding the testis. The biopsy instrument will pop through the tunic and then progress inward easily (Figure 157.6). The inner portion of the needle is advanced deeper into the posterior portion of the testis, away from the anterior aspect and the area of the head of the epididymis (Figure 157.7). Then the outer portion of the biopsy instrument is advanced to cover the inner needle, which provides the biopsy sample (Figure 157.8). The instrument is removed with the outer portion closed over the inner needle. The biopsy sample must be gently teased off the instrument using a small hypodermic needle into a fixative.
Complications
Figure 157.8 Advancement of the outer needle to separate the biopsy sample and maintain it in the biopsy instrument channel.
Early reports in the bull indicated that incisional biopsy induced a decrease in spermatozoal count, an increase in the number of abnormal spermatozoa, tubular degenerative changes, adhesions between the tunics (which was believed to deleteriously affect spermatogenesis), and hemorrhage present in the ejaculate for 2 weeks to 4 months post biopsy.27 In 1960, clinicians reported that split needle biopsy of
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the bull testis did not produce any long-term effects on the testis, although a transient decrease in spermatozoal number occurred. More recently, Galina27 evaluated the effects of split needle biopsy in the boar, bull, ram, and stallion. In the boar, no significant differences in spermatozoal counts were found. In the bull, the ejaculate had a transient discoloration 1 week after biopsy, but, within 3 weeks, the color had returned to normal. Two of four bulls had a line of fibrotic demarcation within the testes. The volume and motility of the bulls with normal fertility before biopsy were normal after biopsy. Rams had no histological change or alteration in spermatozoal count. In the horse, no histological lesions were found after biopsy. This has also been reported by Threlfall and Lopate.28 Seminal evaluation was not performed in any of the stallions used in the previous two reports. Because open biopsy may temporarily be detrimental to spermatogenesis in some species, split needle biopsy is a more desirable and effective method to evaluate abnormalities of the testes in these animals.27 Considerable research has been conducted in man, and little in the domestic species, on the effect of biopsy on seminal quality and output. Although conflicting results have been reported, a transient decrease in spermatozoal output for several weeks to months after incisional biopsy seems likely, but seminal values return to normal given adequate time.3, 9, 15, 16, 18, 27, 29, 30 In man, spermatozoal counts were decreased (42% average median) by 3 weeks after biopsy and had returned to normal within 10–18 weeks.30 In the same study, researchers found that the higher the spermatozoal count was before biopsy, the more profound the decrease in number after biopsy. In the dog, heat and swelling lasted for more than a week after incisional biopsy, and these signs were associated with decreased concentration and increased morphologically abnormal spermatozoa. How long these signs will persist or whether these effects are permanent is unknown.5 The transient decrease in spermatozoal counts can be attributed to several factors: (i) inflammation induced by the procedure, (ii) heating of the testis as a result of the inflammatory response, or (iii) autoimmune reaction to spermatozoa.15 Careful handling of testicular tissue prevents artifacts in the histological preparation.1, 5, 14, 31 Clinicians have reported that the reason testicular biopsy was slow to gain universal acceptance in human medicine was because of the poor quality of the tissue sample slides. Excessive pressure on the testis must be avoided. The surgeon must handle the testis gently so as not to disrupt normal tissue architecture. Trauma can be inflicted on the tissue by using a blade that is too thick or dull. The tissue sample
should be gently nudged into the fixative utilizing a fine-gauge needle. If the specimen is manipulated excessively, the tubular and interstitial architecture will be disrupted. The tissue should never be handled by forceps or placed onto any surface before fixation to prevent crush artifact. The choice of fixative will also determine the quality of the histopathological slide. Bouin’s solution is the preferred fixative, because it causes the least tissue shrinkage and preserves the greatest nuclear detail.13, 14, 31 Markewitz et al.31 recommended that the sample be allowed to fix for 6–12 hours before further processing. The sample cannot be left in Bouin’s fixative longer than 24 hours because it will become too hard to cut with the microtome. Zenker’s formol is also an acceptable fixative.13, 14 However, it causes the dissociation of architecture between tubules, which may be mistakenly interpreted as sloughing.31 A less desirable choice is 10% formalin, because it causes tissue shrinkage, provides poor nuclear images, and can result in a loss of mature spermatozoa.2, 12, 13, 31 Many authors allude to the disadvantages or complications of testicular biopsy but have presented little evidence to justify this concern. Cahill32 reported that testicular biopsies were not considered by owners of famous Kentucky stallions with infertility problems to aid in the diagnosis of the problem ‘for obvious reasons’. These reasons are not obvious, and if in fact they do exist, they should be elucidated. If the reasons do not exist, factual information should be presented. Stallion testicular biopsies are becoming more frequently used since fact has replaced these previous concerns. Several considerations are necessary before performing a testicular biopsy. Hematomas occurring between the testicular tissue and the tunics, or between the tunics and scrotum, are the most common complication.13 They usually result from vigorous manipulation of the testicle. The biopsy must be performed gently without excessive tension or torsion on the testicle, the spermatic cord, or the vasculature to the testicle. In most cases, during incisional or split needle biopsy, hemorrhage can be controlled if it does occur by exerting gentle pressure on the tunic with a sterile sponge for 1–2 minutes.13, 15, 29 If the surgeon is sure that bleeding is controlled, then hematomas rarely result. The testicular artery partially embedded in the tunica albuginea passes around the caudal pole of the testis and then branches on the ventral surface into a medial and lateral branch.33 These branches then pass craniad. In addition, lateral branches may originate dorsally to the testis from the testicular artery (medial and lateral branches), which supplies the posterior portion of the testicle. The biopsy should be taken to avoid the main branches of the testicular artery. No permanent
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Testicular Biopsy side-effects as a result of testicular biopsy in the stallion have been reported. However, in one case in which a large hematoma between the testicle and vaginal tunic occurred as a result of a lateral approach in a pony stallion, it resolved without adverse effects on that stallion’s fertility.27 Adhesions are seen commonly between the two tunics as sequelae to incisional biopsy of the canine testicle.5 The adhesions are not related to hemorrhage but rather to apposition of the incised tunics during healing.5 Careful suturing of the tunica vaginalis will reduce the probability of adhesions. Adhesions between the tunica vaginalis and tunica albuginea have been rarely observed in the human, but when present were asymptomatic in regard to testicular function.13 Other possible complications induced by biopsy are inflammation, increased intratesticular pressure, and the induction of an immune reaction against the sperm.9 These are uncommon problems in most species and can usually be prevented by using a careful technique and by maintaining sterile conditions.5, 14 A decrease in testicular size has been noted following incisional biopsy.34 This decrease may be the result of hemorrhage and connective tissue deposition at the site of the biopsy.15, 34 The formation of a connective tissue reaction to the tissue insult results in a contraction of the tissue, which in turn causes a decrease in testicular size. In the rat, both split needle and fine needle aspiration biopsy induced secondary lesions in the testicle.35 Tubules with degenerate germinal epithelium were found to be focally located at the biopsy site and in the surrounding parenchyma. These lesions were found 8 weeks after biopsy, at which time the rats were castrated. This finding indicates that the biopsy procedure may damage the testis. This is at least in part due to the small size of the testes and the size of the biopsy taken. The large size of the equine testes in relation to a split needle biopsy instrument greatly reduces this concern. If a tumor or abscess is suspected, removal of the affected testis is advised and biopsy is contraindicated. In a study consisting of eight serial split needle biopsies on Beagle dogs, slight scrotal swelling and polycythemia vera (primary increase in red blood cells in the circulating blood) were noted for 3 days after biopsy.36 In addition, at 3 days, cellular degeneration and necrosis were present at the biopsy site. This condition had changed to focal interstitial fibrosis and tubular atrophy 2 weeks later. The adjoining tissue was histologically normal. Testicular size and/or seminal characteristics did not change overall. The researchers concluded that serial split needle biopsy had no effect on reproductive functionability of these dogs.
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Clearly, differences between species concerning the method of biopsy that will provide the most valuable information and the least detrimental effects on the testicle must be considered. The majority of the research has been done in man. This emphasis has resulted in a large population of patients from which to evaluate data and techniques. In domestic and laboratory animals, accurate information regarding the effects of each type of biopsy on the different species is scarce.
Summary The following information is a summary of data collected through the use of split needle testicular biopsy on client-owned and research stallions over the past 30 years. Split needle biopsies have been performed on research stallions and have indicated no decrease in total spermatozoa per ejaculate following biopsy. The lesions induced to the testis also have not appeared to affect adversely the functional capability of the biopsied testis. Biopsies performed on client animals have been beneficial in the diagnosis and prognosis of many infertility and subfertility conditions affecting the stallion. Biopsies from 50% of the infertile stallions indicated testicular degeneration had occurred. The remainder of the stallions had normal testicular tissues present. Stallions with severe disruption or no normal germinal epithelium remaining within the tubules are candidates for retirement from breeding. These animals cannot respond to any type of current therapy and to attempt such is economically unrewarding to the owners. Animals with normal germinal epithelium in the absence of mature spermatozoal production or with low sperm production may be candidates for attempted hormonal therapy. These animals may respond to therapy. The use of testicular biopsy is advocated in the human male to have complete obtainable information on the patient and to determine which patients are potentially treatable. Stallions with normal germinal epithelium and spermatozoal production but with no spermatozoa in the ejaculate should be examined for obstruction of spermatozoal transport. In conclusion, testicular biopsy is not without disadvantages. However, the clinical significance of the trauma induced to the testis appears to be insignificant in regard to the total productivity of that testis when the split needle biopsy technique is used. Testicular biopsy may be useful in clinical situations, when it can provide the clinician with additional information about the spermatogenic state of an infertile or subfertile stallion that otherwise would not be possible. Testicular biopsy can be used to help determine the cause, the prognosis, and the potential for
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therapy of infertile or subfertile stallions when used with fertility records and other diagnostic techniques.
References 1. Tomaszewski JE. Male and female infertility. Paper presented at the 76th Annual Meeting of the International Academy of Pathology, Short Course 25. Chicago, March, 1987. 2. Levin HS. Testicular biopsy in the study of male infertility. Hum Pathol 1979;10:569–84. 3. Scott RR, Sinclair J, Rourke A, Chowdhury S, Yates A, Shaba J. The results of 100 small tissue biopsies of testis in male infertile patients. Postgrad Med J 1976;52:693–8. 4. Wong TW, Straus FH, Jones TM, Warner NE. Pathological aspects of the infertile testis. Urol Clin North Am 1978;5:503–9. 5. Larsen RE. Evaluation of fertility problems in the male dog. Vet Clin North Am 1977;7:735–45. 6. Anthony S, Leong Y, Matthews CD. The role of testicular biopsy in the investigation of male infertility. Pathology 1982;14:205–9. 7. de Kretser DM. Testicular biopsy in the management of male infertility. Int J Urol 1982;5:449–56. 8. Curtis D, Thomas A, Williams JL, Lenton E, Cooke ID. Cytogenetic and histological studies in a series of subfertile males. Int J Urol 1982;5:113–19. 9. Hjort T, Linnet L, Shakkebaek NE. Testicular biopsy: indications and complications. Eur J Pediatr 1982;138:23–31. 10. Douglas RH, Umphenour N. Endocrine abnormalities and hormonal therapy. In: Blanchard TL, Varner DD (eds) Veterinary Clinics of North America: Equine Practice: Stallion Management, vol. 8. Philadelphia: W.B. Saunders, 1992; pp. 237–49. 11. Stewart BL, Roser JF. Effects of age, season, and fertility status on plasma and intratesticular immunoreactive (IR) inhibin concentrations in stallions. Domest Anim Endocrinol 1998;15:129–39. 12. Netto NR. Needle method of testicular biopsy. Int Surg 1974;59:172–80. 13. Rowley MJ, Heller CG. The testicular biopsy: surgical procedure, fixation, and staining techniques. Fertil Steril 1966;17:177–86. 14. Cohen MS, Frye S, Warner RS, Leiter E. Testicular needle biopsy in diagnosis of infertility. Urology 1984;24:439–51. 15. Finco DR. Biopsy of the testicle. Vet Clin North Am 1974;4:377–81. 16. Brannen GE, Roth RR. Testicular abnormalities of the subfertile male. J Urol 1979;122:757–62. 17. Meinhard E, McRae CU, Chisholm GD. Testicular biopsy in evaluation of male infertility. Br Med J 1973;3:577–81. 18. Johnson L, Petty CS, Neaves WB. The relationship of biopsy evaluations and testicular measurements to overall daily sperm production in human testis. Fertil Steril 1980;34:36–40.
19. Shakkebaek NE, Heller CS. Quantification of human seminiferous epithelium. I. Histological studies in twenty-one fertile men with normal chromosome complements. J Reprod Fertil 1973;32:379–89. 20. Zuckerman Z, Rodriguez-Rigau LJ, Weiss DB, Chowdhury AK, Smith KD, Steinberger E. Quantitative analysis of the seminiferous epithelium in human testicular biopsies, and the relation of spermatogenesis to sperm density. Fertil Steril 1978;30:448–55. 21. Varner DD, Blanchard TL, Brinsko SP, Love CC, Taylor TS, Johnson L. Techniques for evaluating selected reproductive disorders of stallions. Anim Reprod Sci 2000;11:493– 509. 22. Johnson L, Varner DD, Thompson Jr DL. Effect of age and season on the establishment of spermatogenesis in the horse. J Reprod Fertil Suppl 1991;44: 87–97. 23. Sundqvist C, Toppari J, Parvinen M, Fagerstroem R, Lukola A. Elimination of infertile male mink from breeding using sperm test, testicular palpation, testosterone test and fine needle aspiration biopsy of the testis. Anim Reprod Sci 1986;11:295–305. 24. Wilson GP. Surgery of the male reproductive tract. Vet Clin North Am 1975;5:537–50. 25. Nseyo UD, Englander LS, Huben RP, Pontes JE. Aspiration biopsy of testis: another method for histologic examination. Fertil Steril 1984;42:281–4. 26. Persson PS, Ahren C, Obrant KD. Aspiration biopsy smear of testis in azoospermia. Scand J Urol Nephrol 1971;5:22– 9. 27. Galina CS. An evaluation of testicular biopsy in farm animals. Vet Rec 1971;88:628–31. 28. Threlfall WR, Lopate C. Testicular biopsy. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1987, pp. 65–73. 29. Paufler SK, Foote RH. Semen quality and testicular function in rabbits following repeated testicular biopsy and unilateral castration. Fertil Steril 1969;20:618–25. 30. Rowley MJ, O’Keefe KB, Heiler CG. Decreases in sperm concentration due to testicular biopsy procedure in man. J Urol 1969;101:347–56. 31. Markewitz M, Sommers SC, Veenema RJ, Butler MD. Testicular biopsy artifacts resulting from improper tissue processing. J Urol 1968;100:44–51. 32. Cahill C. Immature spermatozoa in Thoroughbred stallion. Southwestern Vet 1975;28:13–17. 33. Smith JA. Biopsy and the testicular artery of the horse. Equine Vet J 1974;6:81–3. 34. Lopate C, Threlfall WR, Rosol TJ. Histopathologic and gross effects of testicular biopsy in the dog. Theriogenology 1989;32:585–602. ¨ 35. Schwedes U, Usadel KH, Obert I, Schoffling K. Morphologic testicular changes in rats after biopsy or puncture. Acta Endocrinol 1973;173:149. 36. James RW, Heywood R, Fowler DJ. Serial percutaneous testicular biopsy in the Beagle dog. J Small Anim Pract 1979;20:219–28.
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Chapter 158
Cryptorchid Castration Thomas M. Russell and Patrick J. Pollock
Introduction Cryptorchidism (‘hidden testis’) refers to the failure of descent of one or both testes from the fetal position in the sublumbar region close to the kidney to the normal adult position in the scrotum. It is the most common non-lethal developmental defect in the horse, occurring in approximately 1% of all male births.1 The term cryptorchid is used to describe the nondescended testis and, by extension, affected horses. Colloquial terms for the condition include rig, rigling, and original.2 Three distinct groups of cryptorchids are described, which depend on the position of the affected testis: 1. complete abdominal cryptorchid – the testis and epididymis are located within the abdomen; 2. partial abdominal cryptorchid – the testis is located in the abdomen and the epididymis is located in the inguinal canal; 3. inguinal cryptorchid – the testis and epididymis are located in the inguinal canal, or adjacent to the superficial inguinal ring within the inguinal fascia; these animals are sometimes referred to as ‘high flankers’.3 As a result of the increased temperature of the abdomen, relative to the scrotum, the seminiferous tubules of the abdominally retained testis undergo hypoplasia and affected testes do not produce spermatozoa. As a result, retained testes are usually smaller and softer. An exception to this generalization may occur when the normal testis of a unilateral abdominal cryptorchid is removed. The abdominal testis may then undergo compensatory hypertrophy, complicating surgical removal.4 Unilateral cryptorchids have normal fertility but reduced spermatozoal numbers, and bilaterally affected horses are infertile.5, 6 The Leydig cells are
still functional, albeit producing a reduced amount of testosterone, and therefore affected horses display normal sexual behavior.5, 6 Many breed associations do not allow registration of affected stallions, as cryptorchidism is considered to be an inherited condition. The mechanism of inheritance is poorly understood and most likely involves several different genes.7 Although the condition has been reported in many different breeds, those most commonly affected include Percherons, Saddlebreds, Quarter Horses, native breed ponies, and cross-bred horses. Thoroughbreds appear to have the lowest prevalence.2
Etiology In the normal horse, the fetal gonad differentiates from the embryological mesonephros into a testis by day 37 of gestation.6 By day 45 of gestation, the gubernaculum can be detected, surrounded by an outgrowth of peritoneum that eventually becomes the vaginal process.8 The gubernaculum is a fibrous cord which runs from the testis, to the epididymis, through the inguinal canal, and attaches to the scrotum. During a period of high circulating estrogen in the mare, the interstitial cell numbers in the testis increase until the combined testes weigh approximately 70 g in horses and 50 g in ponies at 7–8 months of gestation. This gonadal hypertrophy is unique to Equidae. With reducing estrogen levels in the mare, each testis reduces in size again to about 5–10 g in the horse and 2.5 g in the pony foal at birth. The gubernaculum increases in size, thus enlarging the inguinal canal, and then reduces in size, undergoing shrinkage and fibrosis, pulling the testis into and through the inguinal canal. The right testis is often smaller than the left, and usually precedes the left into the scrotum.9 Descent of the testis usually occurs in the final month of gestation, or in the first 2 weeks of life. The epididymis descends first.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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The fetal gubernaculum is divided into three parts: (i) a portion connecting the testis to the tail of the epididymis – the proper ligament of the testis, (ii) a portion between the tail of the epididymis and vaginal process – the caudal ligament of the epididymis, and (iii) the caudal part of the gubernaculum, running from the parietal peritoneum (vaginal process) to the scrotum, which becomes the scrotal ligament (Figure 97.1). In the cryptorchid, the scrotal ligament is referred to as the inguinal extension of the gubernaculum testis (IEGT), and is an important landmark in the surgical retrieval of abdominal testes. Failure of this portion of the gubernaculum to enlarge may lead to failure of the vaginal ring to expand sufficiently to allow normal descent. Similarly, overenlargement of the gubernaculum and/or failure of regression may also inhibit passage of the testis into the inguinal canal.6, 10 Endocrinological events controlling development of the gubernaculum are poorly understood. Abnormalities of the testis may also play a role in failure of descent, particularly large testes, persistence of the cranial suspensory ligament of the testis, or testes affected by cysts or teratomata that may be too large to pass through the inguinal canal.11–13 A number of large retrospective studies have investigated the incidence of right versus left cryptorchidism, and indicate that failure of descent occurs with nearly equal frequency. However, the location of the retained testis may vary with side. One study found abdominal cryptorchidism to be left sided in 75% of cases, while right-sided abdominal retention was 42%. This difference is most likely due to the fact that the left testis is larger during development; therefore, if the gubernaculum does not sufficiently enlarge the vaginal ring, the larger left testis has less chance of passing successfully through it.14–16
Clinical signs and diagnosis History More often than not, the history is vague. However, if the animal was bred and reared by the owner, its behavioral and castration history is of value. Frequently, this information is not available and the condition is noticed only when the animal is presented for castration.
Clinical examination Visual inspection of the scrotum is not sufficient, and palpation must be performed in every case. The examiner should stand on the same side as the handler,
and, while facing the thorax of the horse, pass a hand under the abdomen and into the inguinal region. Palpation should include the scrotum, inguinal region, and external inguinal ring. Sedation may allow relaxation of the cremaster muscle, and subsequent descent of an inguinal testicle into palpable reach. It also reduces the inherent danger of this procedure in poorly handled animals. It should be noted that palpation of the scrotum to determine testicular descent is unreliable in neonatal foals, as the bulb of the gubernaculum can feel like a testis for up to 3–4 weeks after birth.17 The equine scrotum is often asymmetrical, owing to the longer left spermatic cord; however, in cryptorchids, the affected side is poorly developed. An attempt should be made to identify the presence of scars that may indicate a previous castration.
Examination per rectum Some workers suggest that examination per rectum should be performed on horses suspected of being cryptorchid.2 The technique involves palpation of the vaginal rings, which are located 6–8 cm cranial to the iliopectineal eminence and 10–12 cm abaxial to midline in an average 500-kg horse. In geldings, the vaginal ring is palpable as a slight depression; in stallions, it is possible to insert a finger into the ring. Infrequently, an abdominally placed testis or testicular abnormality may be identified. It should be recognized that inadvertent rectal perforation is a risk of rectal palpation. The risk is increased in smaller and immature animals, and in male horses in general, which are less used to the examination and tend to resist more than mares.18–21 Examination per rectum is rarely necessary in the investigation of cryptorchidism if the history of the horse is known.
Diagnostic imaging Recently, the use of transabdominal ultrasound has been described for the identification of abdominally retained testis. In horses with a short hair coat, transabdominal ultrasonography proved to be an accurate method of locating a cryptorchid testis.22 The presence of the testicle was confirmed by hyperechoic tunica albuginea, and either a central vein or the epididymis of the testicle. This discriminates the testis from other abdominal structures such as empty small intestinal loops, the density of which was sometimes identical to that of a cryptorchid testis. Most abdominal testes (18/22) were visualized on the ventral abdominal wall within a few centimeters of, or against, the urinary bladder.
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Cryptorchid Castration Transrectal ultrasonography is also an excellent method to detect abdominal cryptorchid testes21 and is encouraged as long as examiners are experienced in transrectal palpation. Utilizing this technique also avoids the surprise of finding a testicular cyst or teratoma at subsequent exploratory surgery.
Hormonal assays The plasma or serum androgen concentration can be used to determine whether extrascrotal testicular tissue is present. Stallions and cryptorchid stallions have significantly higher levels than geldings. In a number of studies, resting testosterone concentrations were generally less than 40 pg/mL in geldings, compared with concentrations greater than 100 pg/mL in stallions.23, 24 Unfortunately, the literature suggests that there is a wide variation in basal concentrations of testosterone. It seems that the age of the horse, season, and individual variation have a significant part to play, and as such this technique can lead to confusion. A human chorionic gonadotropin (hCG) stimulation test can be used to improve the accuracy of detecting testicular tissue. The technique involves measuring the basal concentration of testosterone before and 60–120 minutes after the administration of 6000–12 000 units of hCG. Horses are considered to be cryptorchids if the basal and post-stimulation concentration of testosterone is above 100 pg/mL, and to be geldings if the concentration remains below 40 pg/mL. Variations reported in the test include obtaining post-stimulation samples 24 hours after injection or even 2–3 days later. It should be noted that several studies suggest that horses sampled during the winter and those sampled when younger than 18 months had a poor response to the hCG test. Despite this, a 94.6% accuracy has been described.23, 24 Plasma or serum concentration of conjugated estrogens has also been used for the purpose of identifying cryptorchid animals. This technique appears to be most effective when used for detecting cryptorchids older than 3 years, and was unreliable when used for detecting cryptorchidism in donkeys. Geldings were found to have basal concentrations of estrone sulfate below 50 pg/mL, whereas cryptorchids had concentrations in excess of 400 pg/mL.24, 25 Many laboratories offer a group of tests to determine whether a horse is a cryptorchid, and knowledge of the normal values for the laboratory is important. While false negatives are a considerable problem, clinicians can be reasonably confident that, if a test indicates the presence of testicular tissue, it is indeed present.
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Treatment of cryptorchidism Pre-surgical considerations Prior to surgery, horses should undergo a full clinical examination to rule out the presence of underlying disease. The inguinal region should be thoroughly investigated for the presence of a testis, and the presence of inguinal herniation, which may influence choice of technique. If general anesthesia is to be induced, the horse should be placed in dorsal recumbency for optimal access to the surgical site. Following anesthetic induction, the inguinal region should be visually and manually investigated for any evidence of previous surgery, suggesting prior removal of one or both testes. Manual examination, in the form of deep palpation of the inguinal region and superficial inguinal ring, may reveal a testis that was not apparent during standing examination. For laparoscopic and flank cryptorchid castration, the horse is best restrained in stocks and sedated. Incremental doses of sedatives can be used effectively; however, the use of a continuous infusion of a sedative combination is the method of choice, as this provides a steady plane of sedation, reducing ataxia, and therefore unwanted movement. A useful technique has been described26 for standing surgery. Detomidine is administered as an initial bolus loading dose of 7.5 µg/kg. Detomidine is added to Hartmann’s solution to make a solution of 25 µg/ml. The initial infusion rate is 0.6 µg/kg/min, and this is halved every 15 minutes.
Surgical techniques
Inguinal approach (Figure 158.1) General anesthesia is essential for an inguinal approach to the cryptorchid horse. A variety of approaches are described, including an incision made directly over the superficial inguinal ring, an elliptical incision, or a scrotal approach. The scrotal approach has the advantage that both scrotal and cryptorchid testes can be removed through the same incision. To expose the superficial inguinal ring, blunt dissection is used to separate the inguinal fascia. Once the correct tissue plane is identified, the tissue should separate with moderate digital pressure. Failure to identify the correct tissue plane greatly increases the difficulty of dissection and identification of the testis. Care should be taken not to damage any of the large pudendal vessels which traverse the region. If the testis is retained inguinally, it is easily identified at this point beside the superficial inguinal ring. Once the vaginal sac is identified, it should always be
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Figure 158.1 Approaches for cryptorchidectomy. (a) Inguinal; (b) parainguinal; (c) paramedian; and (d) scrotal.
opened to ensure that it contains a testis. In this way accidental mis-identification, and removal, of the epididymal tail of a partially abdominal testis as an inguinal testis can be avoided. To confirm that the testis has already been removed, it is necessary to identify the stump of the spermatic cord as it exits the inguinal canal. The stump should contain remnants of the cremaster muscle, ductus deferens, and testicular vessels.2 The stump is often encountered partially adhered to the surrounding subcutaneous tissue. If the testis is not readily identified within the inguinal region, an abdominal or partially abdominal testis can be retrieved using the following noninvasive technique. Careful exposure of the superficial inguinal ring using appropriate retraction allows the identification of the scrotal ligament or inguinal extension of the gubernaculum testis (IEGT).27 The IEGT is a cord of tissue located by carefully examining the margin of the superficial inguinal ring. It is usually found at the junction of the middle and cranial third of the ring and can be medial or lateral. By gently grasping and retracting the loose fascia around the ring with one hand and palpating within the ring with the other hand, the IEGT can usually be identified, as it causes the vaginal process to evert into the inguinal canal. The IEGT can be differentiated from the surrounding fascia, which usually tears. Care should be taken not to confuse the genitofemoral nerve, which exits the canal more caudally, with the IEGT.
Following identification and traction on the IEGT, the common vaginal tunic or vaginal process everts into the canal and can often be visualized as a white glistening structure deep within the canal. A hypoplastic cremaster muscle may be evident attached to the vaginal process. A curved sponge forceps introduced into the canal can be used to grasp the vaginal process, and, with gentle traction, evert more of it. At this point, the vaginal process is stripped of fascia and incised using a small scalpel blade (the curved no. 12 works well). Upon incising the vaginal process, peritoneal fluid is often encountered. A hemostat should be introduced into the vaginal process to grasp the epididymis contained within. Traction on the epididymis to exteriorize the tail of the epididymis, which is connected to the testis via the proper ligament, allows the testis to be pulled through the vaginal ring. At this stage patience is required to gently pull the testis through the ring. In some horses, particularly mature stallions, dilation of the ring is required to exteriorize the testis. To facilitate dilation of the ring, a finger is passed through the ring to the first knuckle and gently flexed to disrupt the ring. Using this technique, even very large testes can be removed through a relatively small vaginal ring. Following exposure of the testis, it is removed using standard techniques. An assistant can be asked to apply pressure to the abdominal wall, to facilitate emasculation, if the vascular pedicle is very short. Once the testis has been removed, the vaginal ring should once again be palpated to ensure that it has not become excessively dilated, thereby increasing the risk of eventration. Generally, if only two fingers can be passed into the ring, then no further action is required, and the horse can be recovered and treated as a normal castration. The inguinal fascia can be sutured and the wound left to heal by secondary intention. If the ring is found to be excessively dilated, several techniques have been described to prevent eventration, including packing the inguinal canal with sterile gauze for 24–36 hours. This technique is not generally recommended as it does not allow the vaginal ring to contract, and the risk of eventration is still present following removal, especially if the intestine or greater omentum becomes adherent to the packs. It also requires further manipulation, and therefore increased risk of contamination of the surgical site. The technique of choice involves closure or partial closure of the superficial inguinal ring using heavy gauge monofilament absorbable suture material in a simple continuous or interrupted pattern. A large needle is required to perform this technically demanding procedure. As a further safeguard, the scrotum sac and skin may also be closed.
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Cryptorchid Castration Following suturing, the horse should be confined to a stable for 48 hours and should not be allowed unrestricted exercise for 3 weeks. Horses subjected to this technique may display evidence of increased discomfort and hind limb stiffness following recovery from anesthesia; this is usually transient.
Parainguinal approach (Figure 158.1) Some surgeons prefer not to disrupt the inguinal ring, and in rare cases in which the vaginal process cannot be identified, a paramedian laparotomy may be performed.28 A 4-cm incision is made parallel and approximately 2 cm medial to the cranial portion of superficial inguinal ring. The incision is carried through the aponeurosis of the external abdominal oblique muscle and blunt dissection used to enter the abdomen. The incision need be large enough only to insert two fingers in a caudolateral direction, in order to identify and grasp the vaginal process and then part of the epididymis or gubernaculum. If these structures cannot be identified, the incision should be enlarged to accommodate a hand. The hand should be passed into the abdomen flat against the body wall and swept around until the testis is encountered. It is surprising how similar a loop of small intestine feels to an abdominal testis, and patience may be required for identification. Once the testis has been identified and exteriorized, it is emasculated in the usual fashion. The aponeurosis of the external abdominal oblique muscle is closed using a simple continuous or interrupted pattern with heavy gauge monofilament absorbable suture material. The subcutaneous tissue and the skin can be closed, or left to heal by secondary intention. If the testis was retrieved by finger palpation alone, through a short incision, the horse is confined to a box for to 2–4 days, followed by restricted exercise for a period of 3–4 weeks. Horses requiring a larger paramedian laparotomy require a longer period of box confinement.
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inguinal ring to identify the testis. If the testis cannot be readily identified, the ductus deferens is located in the genital fold of the bladder and traced to the testis. Previous reports have suggested that the testes of bilateral abdominal cryptorchids can be retrieved through one incision;2 however, it can be difficult to exteriorize the contralateral testis, necessitating ligation and transection within the abdominal cavity. The abdominal tunic, subcutis, and skin are closed separately and the horse is allowed to recover. Following laparotomy, the horse must be confined to a stable for 3–4 weeks, followed by a further 8 weeks of rest in a small paddock. The horse should not be allowed unrestricted exercise for approximately 3 months.
Flank approach With the horse either standing and sedated using instillation of local anesthetic solution or recumbent under general anesthesia, a 10- to 15-cm vertical incision is made in the paralumbar fossa on the affected side. The external abdominal oblique muscle is incised in the same direction as the skin incision. The internal abdominal oblique and transverse abdominal muscles are divided in the direction of their fibers. Finally, the peritoneum and retroperitoneal fat are bluntly divided, using fingers or an instrument, to access the peritoneal cavity.20 Locating, exteriorizing, and emasculating the testis is as described for the paramedian approach. Closure of the peritoneum and transverse abdominal muscle is challenging and unnecessary. The internal and external abdominal muscles, subcutaneous tissue, and skin are closed separately using a continuous or interrupted technique with monofilament absorbable material. The use of a stent applied over the wound may reduce post-operative swelling. Stall rest is generally unnecessary.
Laparoscopic approach Suprapubic paramedian approach (Figure 158.1) The horse is anesthetized and placed in dorsal recumbency. A 10- to 20-cm incision is made approximately 5–10 cm lateral, and in some cases obliquely, to the ventral midline. The incision begins at the level of the preputial orifice and extends caudally. The abdominal tunic and sheath of the rectus abdominis muscle are incised in the same direction as the skin incision and the underlying rectus abdominis muscle is bluntly separated to expose the rectus sheath, retroperitoneal fat, and peritoneum, which can be bluntly penetrated with a finger or instrument.29 A hand is passed into the abdomen towards the internal
Laparoscopic removal of a testis is increasingly employed for the treatment of cryptorchid horses.30, 31 The technique can be used with the animal standing and sedated, or during general anesthesia. Fractious stallions and small stallions should be subjected to general anesthesia;30 however, the use of continuous infusion of a sedative mixture greatly facilitates the standing procedure in most animals. To date, the authors have encountered very few animals, including miniature horses, in which a standing laparoscopic cryptorchidectomy could not be performed. Whether the procedure is performed standing or under general anesthesia, food should be withheld for 12–24 hours to reduce the amount of intestinal
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contents. This increases surgical visibility and maneuverability, as well as reducing the risk of inadvertent puncture of viscera.32 With the horse sedated and restrained in stocks, the tail is tied to prevent contamination of the surgical field. The paralumbar fossa is prepared for surgery in routine fashion. The presence of the spleen on the left side reduces the chance of inadvertent damage to viscera during introduction of trocars and insufflation. The right paralumbar fossa is prepared in cases of right-sided cryptorchidism, bilaterally retained testis, or in horses in which the location of the testis cannot be determined prior to surgery. Three laparoscopic ports are required, and infiltrated with local anesthetic solution prior to incision. Alternatively an inverted L, line infiltration, or paravertebral block can be performed. The authors prefer to insert a safety trocar unit into the paralumbar fossa for insufflation, whereas other surgeons have reported the use of a Verres needle or trocar placed via the ventral midline. Whichever method is employed, it is crucial to ensure that the retroperitoneal space is not inadvertently insufflated, otherwise the procedure is difficult or impossible to perform. Laparoscopic castration requires the use of three or four 15-cm by 10-mm trocars. A 15-mm incision is made through the skin and fascia of the external abdominal oblique muscle, at the level of the tuber coxae, midway between the last rib and tuber coxae. The authors generally place the laparoscope into the abdomen prior to insufflation to further confirm correct placement. The first trocar unit is boldly inserted in a slightly ventral, caudal direction. Correct placement is confirmed by the sound of a rush of air entering the abdomen. Insufflation of the abdomen with carbon dioxide is made to a pressure of 15 mmHg. Following insufflation, a further two or three trocar units are placed (Figure 158.2). The testis is located by inspecting the region around the internal inguinal ring. The testis can be located hanging from a curtain of mesorchium, which also contains the testicular vasculature and vas deferens (Figure 158.3). The mesorchium courses from the area of the kidney caudally to the inguinal ring, and, although an abdominal testis can be located anywhere between these two structures, it is most frequently encountered approximately 10–15 cm cranioventral to the internal inguinal ring. The testis can be anesthetized by injection of local anesthetic solution into the testis or mesorchium using a laparoscopic injection device, or by the use of high epidural anesthesia prior to the start of surgery. Laparoscopic surgery is based on the technique of triangulation. The mesorchium and testis (if present) is grasped with a laparoscopic grasping forceps and manipulated to ensure the entire testis and associated structures are within the abdomen, and that none of the
Figure 158.2 Laparoscopic removal of an abdominal testicle.
structures are located within the inguinal canal. Various techniques of hemostasis have been described including the use of suture material and extracorporeal knotting techniques, proprietary ligatures, and bipolar electrocautery. More recently, controlled feedback bipolar cautery has been described using a LigaSureTM device that provides sealing for up to a 7mm-diameter vessel without a proximal thrombus and with little tissue dissection. Following successful hemostasis, the testicle is amputated with the use of either laparoscopic scissors or an electrocautery unit. In bilaterally affected horses, the same procedure is carried out on the contralateral side. Alternatively, a technique has been described whereby both testes are retrieved from the same side; however, this is technically more demanding.30 In order that abdominal pressure is not lost, neither testis is retrieved from the abdomen until both have been amputated and the stumps examined for evidence of hemorrhage. Sub-
T
Figure 158.3 An abdominally located testis (T) viewed laparoscopically. The arrows indicate the elongated proper ligament of the testis running into the inguinal canal.
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Cryptorchid Castration sequently, the ventral-most trocar incision is enlarged to facilitate removal of the testis. Some surgeons prefer to use a laparoscopic bag for retrieval of the testis. Partial abdominal testes can be pulled back into the abdomen with the use of a grasping device. This may require enlargement of the internal inguinal ring by the use of electrocautery. In the case of horses with a scrotally situated contralateral testis, several options for removal have been described: standard standing castration; short general anesthesia; and pulling the testis back into the abdomen for ligation and amputation. The last is technically demanding. Closure of laparoscopic portals is performed in one layer (skin) with monofilament suture material, except for the enlarged portal used for testicular removal, which is closed in two layers (external abdominal oblique muscle and the skin). Following laparoscopic castration, post-operative care is as for standard castration. Selection of approach Laparoscopic cryptorchid castration is the treatment of choice, as it allows the procedure to be carried out with the horse standing and sedated, and is not affected by the position of the testis. However, the technique requires considerable expertise and the use of expensive equipment, and in many cases an increase in surgical time. In the authors’ opinion, horses subjected to laparoscopic castration return to function more rapidly and have reduced morbidity. Using the inguinal approach described above, a testis can be located no matter where it is positioned. It is very rarely necessary to perform a laparotomy as, in most cases, even an abdominally retained testis can be retrieved using the inguinal technique. If necessary, a small parainguinal incision can be made if the testis cannot be identified. Paramedian, parainguinal, and flank laparotomies are appropriate only for the retrieval of abdominally retained testes. The longer incisions associated with these approaches are associated with an increased risk of complications and longer convalescence.
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tus deferens are found, then it is confirmed that the animal was previously castrated.
Monorchidism The term monorchidism is used to describe animals with the complete absence of one testis. This is the result of surgical removal of one testis, atresia of one of the testis, or degeneration, possibly as a result of vascular damage to the testis.34 Monorchidism can be congenital, or may develop following torsion of the spermatic cord and subsequent degeneration of the affected testis. Determining the cause of monorchidism is difficult; however, some reports have suggested that the presence of a vaginal process within the inguinal canal indicates that it was acquired. The presence of cremaster muscle remnants, testicular vessels, and ductus deferens within the inguinal region suggests surgical excision.
Incomplete cryptorchid castration In some cases, the tail of the epididymis of a partial abdominal cryptorchid may lie in the inguinal region owing to excessive length of the proper ligament of the testis. The epididymal tail within a well-developed vaginal process can be mistaken for a testis by an inexperienced surgeon, who may mistake it for a hypoplastic inguinal testis (Figure 158.4). In these cases, the horse will continue to display stallion-like behavior. To avoid this mistake, the vaginal process should always be opened and the contents investigated to ensure the presence of an epididymis and testis, prior to proceeding with castration.2
B A
False cryptorchids These are castrated males that show signs of male behavior. Around 5% of animals submitted as cryptorchids fall into this category. These animals often present in the springtime.33 Investigation of false cryptorchids includes the use of the hormonal assays previously described, followed, where necessary, by exploratory surgery, generally in the inguinal area. If two spermatic cords with tunica vaginalis, cremaster remnants, and duc-
C
Figure 158.4 (a) Normal testis. (b) Cryptorchid testis. (c) Epididymal tail with elongated proper ligament of the testis.
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Summary Treatment of cryptorchids is a challenge for any surgeon, and it is often said that no two cryptorchidectomies are the same. The introduction of standing laparoscopic surgery has certainly revolutionized the treatment of these horses, and no doubt will become even more widespread in the future. However, given the requirement for considerable training and expensive equipment, more traditional methods are still frequently employed. The keys to effective treatment are a good history, thorough clinical examination, and an in-depth knowledge of the events surrounding testicular descent, the surgical anatomy of the region, and the variations that may be encountered. Recognition of the remnants of a partially removed testis, amputated epididymal tail, or congenital abnormality may be essential to success. In 1875 Degive,35 a well-known cryptorchid surgeon, suggested that ‘When one has done one’s hundredth cryptorchidectomy, one hasn’t done one’s hundred and first’. This is as true today as ever it was.
References 1. Hayes HM. Epidemiological features of 5009 cases of equine cryptorchidism. Equine Vet J 1986;18:467–71. 2. Schumacher J. Testis. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St. Louis, MO: Saunders Elsevier, 2006; pp. 775–810. 3. Bishop MWH, David JSE, Messervy A. Some observations on cryptorchidism in the horse. Vet Rec 1964;76:1041– 8. 4. Arthur GH. The surgery of the equine cryptorchid. Vet Rec 1961;73:385–9. 5. Foster RA, Ladds PW. Male genital system: testis and epididymis. In: Maxie MG (ed.) Jubb, Kennedy and Palmer: Pathology of the Domestic Animals, 5th edn. Edinburgh: Saunders Elsevier, 2007; pp. 572–5. 6. Bergin WC, Gier HT, Marion GB, Coffman JR. A developmental concept of equine cryptorchism. Biol Reprod 1970;3:82–92. 7. Leipold HW, DeBowes RM, Bennett BS, Cox JH. Cryptorchidism in the horse: genetic implications. In: Proceedings of the 32nd Annual Convention of the American Association of Equine Practitioners, 1986, pp. 579–83. 8. Wensing CJG, Colenbrander B. Normal and abnormal testicular descent. In: Clarke J (ed.) Oxford Reviews: Reproductive Biology, vol. 8. Oxford: Clarendon Press, 1986; pp. 125–30. 9. Smith JA. The development and descent of the testis in the horse. Vet Ann 1975;15:156–61. 10. Ellenport CR. General urogenital system. In: Getty R. (ed.) Sisson and Grossman The Anatomy of the Domestic Animals. Philadelphia: W.B. Saunders, 1975; pp. 145–9. 11. Wilson DG, Nixon AJ. Case of equine cryptorchidism resulting from persistence of the suspensory ligament of the gonad. Equine Vet J 1986;18:412–13.
12. Allison N, Moeller Jr RB. Bilateral testicular leiomyosarcoma in a stallion. J Vet Diagn Invest 1999;11:179– 82. 13. Pollock PJ, Prendergast M, Skelly C, Callanan JJ. Testicular teratoma in a three day-old Thoroughbred foal. Vet Rec 2002;150:348–9. 14. Stickle RL, Fessler JF. Retrospective study of 350 cases of equine cryptorchidism. J Am Vet Med Assoc 1978;172: 343–6. 15. Cox JE, Edwards GB, Neal PA. An analysis of 500 cases of equine cryptorchidism. Equine Vet J 1979;11:113– 16. 16. Coryn M, De Morr A, Bouters R, Vandeplassche M. Clinical, morphological and endocrinological aspects of cryptorchidism in the horse. Theriogenology 1981;16:489– 96. 17. Adams SB, Fessler JF. Noninvasive inguinal cryptorchidectomy. In: Adams SB, Fessler JF (eds) Atlas of Equine Surgery. Philadelphia: W.B. Saunders, 2000; pp. 215–18. 18. Greatorex JC. Rectal exploration as an aid to the diagnosis of some medical conditions in the horse. Equine Vet J 1968;1:26–9. 19. Stauffer VD. Equine rectal tears: a malpractice problem. J Am Vet Med Assoc 1981;178:798–9. 20. Searle D, Dart AJ, Dart CM, Hodgson DR. Equine castration: review of anatomy, approaches, techniques and complications in normal, cryptorchid and monorchid horses. Aust Vet J 1999;77:428–34. 21. Jann HW, Rains JR. Diagnostic ultrasonography for evaluation of cryptorchidism in horses. J Am Vet Med Assoc 1999;196:297–300. 22. Schambourg MA, Farley JA, Marcoux M, Laverty S. Use of transabdominal ultrasonography to determine the location of cryptorchid testes in the horse. Equine Vet J. 2006;38:242–5. 23. Cox JE. Experiences with a diagnostic test for equine cryptorchidism. Equine Vet J 1975;7:179–81. 24. Cox JE, Redhead PH, Dawson FE. Comparison of the measurement of plasma testosterone and plasma oestrogens for the diagnosis of cryptorchidism in the horse. Equine Vet J 1986;18:179–82. 25. Ganjam VK. An inexpensive, yet precise, laboratory diagnostic method to confirm cryptorchidism in the horse. In: Proceedings of the 23rd Annual Convention of the American Association of Equine Practitioners, 1977, pp. 245– 53. 26. Wilson DG, Bohart GV, Evans AT, Robertson S, Rondenay Y. Retrospective analysis of detomidine infusion for standing chemical restraint in 51 horses. Vet Anaesth Analg 2002;29:54–7. 27. Valdez H, Taylor TS, McLaughlin SA, Martin MT. Abdominal cryptorchidectomy in the horse, using inguinal extension of the gubernaculum testis. J Am Vet Med Assoc 1979;174:1110–12. 28. Wilson DG, Reinertson EL. A modified parainguinal approach for cryptorchidectomy in horses. An evaluation in 107 horses. Vet Surg 1987;16:1–4. 29. Cox JE, Neal PA, Edwards GB. Suprapubic paramedian laparotomy for equine abdominal cryptorchidism. J Am Vet Med Assoc 1978;173:680–2. 30. Fischer Jr AT. Standing laparoscopic surgery. Vet Clin North Am Equine Pract 1991;7:641–7. 31. Fischer AT, Vachon AM. Laparoscopic intra-abdominal ligation and removal of cryptorchid testes in horses. Equine Vet J 1998;30:105–8.
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Cryptorchid Castration 32. Hendrickson DA. Standing laparoscopic cryptorchidectomy. In: Fischer Jr AT (ed.) Equine Diagnostic and Surgical Laparoscopy. Philadelphia: W.B. Saunders 2002; pp. 155–61. 33. Cox JE. Behaviour of the false rig: Causes and treatments. Vet Rec 1986;118:353–6.
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34. Parks AH, Scott EA, Cox JE, Stick JA. Monorchidism in the horse. Equine Vet J 1989;21:215–17. 35. Degive A. De la castration d’animaux cryptorchides. Ann Med Vet 1875;11:629–720.
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Chapter 159
Inguinal Hernia Patrick J. Pollock and Thomas M. Russell
Introduction Inguinal hernia describes the passage of abdominal contents into the inguinal canal. Most commonly, hernias involve the passage of jejunum or ileum; however, herniation involving portions of the large colon, small colon, bladder, and omentum have been described in previous reports.1–3 If the herniated material passes through the superficial inguinal ring and enters the scrotum, the condition may be defined as a scrotal hernia; however, it should be noted that these definitions are often used interchangeably.4
Etiology Adults Although inguinal herniation and ruptures have been described in male and female horses, they are much more commonly encountered in stallions.1, 2, 5 Although geldings are affected, the reduction in size of the internal inguinal ring after castration leads to a reduced incidence.5 In adult horses inguinal hernias and ruptures commonly occur during heavy exercise or copulation,1 or as a result of trauma.
Terminology The terminology used to describe inguinal hernias is somewhat confusing owing to the use of terms borrowed from the description of similar conditions in human beings.4 In an indirect hernia (Figure 159.1), abdominal viscera passes through the vaginal ring and lies within the vaginal cavity, adjacent to the normal contents. A ruptured inguinal hernia (Figure 159.2) occurs when the herniated material then passes through a rent in the parietal tunic and scrotal fascia, to a lie in a subcutaneous position in the inguinal or scrotal region. An inguinal rupture (Figure 159.3) is used to describe the situation where viscera passes through a tear in the peritoneum and body wall adjacent to the vaginal ring, and comes to lie extraperitoneally in the subcutaneous tissue of the inguinal or scrotal region. Inguinal ruptures have been termed direct hernias; however, this terminology is incorrect and refers to a condition in humans whereby the body wall adjacent to the vaginal ring is weakened and protrudes, allowing abdominal viscera to lie within a peritoneum-lined sac.5
Foals In foals the condition is congenital, although it may not be immediately apparent at birth. Most are considered to be hereditary.6 Hernias may be unilateral or bilateral, although studies suggest that left inguinal herniation is more common than right.5 Mechanisms for the development of herniation are poorly understood; however, excessive enlargement of the extra-abdominal portion of the gubernaculum may lead to widening of the mouth of the vaginal process.7 Compression of the abdomen during parturition has been suggested as a cause of ruptured inguinal hernias, as has excessive straining in the first few days after birth due to meconium impaction.8, 9 Foals are more commonly affected by ruptured inguinal hernias (Figure 159.2) than adults, whereas in adults inguinal ruptures (Figure 159.3) are more common. Inguinal herniation and ruptures have been documented in a variety of breeds of horse; however, there is evidence to suggest that the incidence is
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 159.1 An indirect hernia.
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Figure 159.4 Congenital hernias are identified as a fluctuant, non-painful, reducible swelling in the inguinal region.
highest in Standardbreds and European Warmblood horses, with anecdotal evidence in Tennessee walking horses, American Saddlebreds, and Arabians.10 In foals, the heavier draft breeds seem to be overrepresented.11
Clinical signs, diagnosis, and treatment Congenital inguinal hernia
Figure 159.2 A ruptured inguinal hernia
Figure 159.3 An inguinal rupture
Congenital inguinal hernias are usually recognized shortly after birth as a fluctuant, non-painful, reducible swelling in the inguinal region (Figure 159.4). In some cases, they can become quite large and may interfere with locomotion, lymphatic drainage, and blood supply. Testicular atrophy has been documented in foals with longstanding untreated hernias.12 Some inguinal hernias resolve spontaneously within several days, and the majority by 3–6 months of age. Owners and managers should be encouraged to frequently manually reduce the hernia until such time as the vaginal ring reduces in size and prevents herniation. The use of a truss has been advocated to speed resolution, and for treatment of larger hernias. With the foal sedated and placed in dorsal recumbency, the hernia is reduced and a roll of cottonwool or gauze is packed into the region of the superficial inguinal ring. The cottonwool or gauze is held in place with an adhesive bandage placed in a figure-of-eight pattern over the back and both inguinal regions. Generally, the truss is kept in place for approximately 1 week.5 Medical records should be kept for colts affected by inguinal herniation, as the incidence of post-castration evisceration may be higher than in the general equine population. In foals affected by ruptured inguinal hernias or inguinal ruptures, the swelling is often irreducible
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and can be very large. Individual loops of intestine may be apparent under the skin and, in some cases, viscera adheres to the subcutaneous tissue. Sloughing of the thin skin may occur due to pressure necrosis. With a ruptured inguinal hernia or inguinal rupture, the intestine may become strangulated, leading to clinical signs of colic. These cases are surgical emergencies. Ultrasonography may be useful in determining the presence of intestine and the viability of the testis.13 Acquired inguinal hernia Acquired inguinal hernias and ruptures are more commonly encountered in adult horses and generally lead to acute strangulation of intestine. Clinical signs are referable to the compromised intestine and include tachycardia, nasogastric reflux, sweating, rolling, and other signs of colic. One side of the scrotum may be enlarged and firm. Distended loops of small intestine may be palpable per rectum and it may be possible to identify intestine entering the internal inguinal ring. The use of ultrasonography may be helpful in determining whether intestine is present, differentiating the condition from testicular torsion, and assessing the viability of the gonad.13 The presence of strangulated intestine within the vaginal sac may lead to obstruction of the testicular vasculature and associated swelling of the testis. In most cases, these animals require emergency surgical treatment; however, two techniques for non-surgical reduction of acquired inguinal hernias have been described. The first involves grasping the incarcerated loop of intestine per rectum as it enters the inguinal canal and applying traction to pull the intestine back into the abdomen.14 The risk of inadvertent perforation of the rectum must be taken into account with this technique and this may be reduced by the prior administration of epidural anesthesia or intravenous administration of butylscopolamine bromide (0.15–0.3 mg/kg) or propanthelene bromide (0.04– 0.06 mg/kg). With the second technique, the horse is sedated, and downward pressure is applied to the testis on the affected side, while the intestine is squeezed upwards using a one-hand-over-another technique, until the intestine passes through the vaginal ring and back into the abdomen.15 It is crucial that these techniques only be used if the horse is presented prior to strangulation of intestine. They must not be attempted if there is any doubt about the viability of the affected intestine, or if a ruptured inguinal hernia or inguinal rupture is present. Horses treated in this way should be carefully observed after replacement for any evidence of colic or depression which might
indicate non-viable intestine. Performing these procedures during standing laparoscopy allows visual evaluation of the viability of the affected intestine.
Surgical treatment General surgical considerations Horses presented with signs referable to strangulated intestine require emergency surgery, and open surgical correction is always indicated if intestinal or testicular viability is in doubt. Administration of intravenous fluids, non-steroidal anti-inflammatory drugs, and broad spectrum antimicrobials should be undertaken prior to induction of anesthesia. Nasogastric intubation and gastric decompression is recommended prior to induction of anesthesia. Anesthetic techniques should be carefully selected to minimize cardiovascular and respiratory compromise. The horse is placed in dorsal recumbency and prepared for aseptic surgery. The penis is placed into the prepuce, which is packed with gauze and closed with either a purse-string suture or towel clamps. Inguinal, scrotal, and ventral midline regions are prepared in anticipation of inguinal and ventral midline surgical incisions. Care should be taken during the preparation of the skin in young foals, as the tissue is easily traumatized, particularly in foals affected by ruptured inguinal hernias and inguinal ruptures where the intestine is subcutaneous.
Inguinal herniorrhaphy in foals Surgery is indicated for foals with clinical signs of strangulated intestine, those affected by hernias that are not manually reducible, those with enlarging hernias or hernias that are so large that spontaneous resolution is unlikely,16 and for foals with ruptured inguinal hernias and inguinal ruptures. Surgical techniques include open inguinal and ventral midline approaches or laparoscopic approaches in the absence of strangulated intestine.
Inguinal herniorrhaphy in adult horses Adult horses are most commonly presented for repair of inguinal hernias and ruptures following incarceration of segments of intestine. In these cases, clinical signs are attributable to the presence of strangulated intestine. Careful examination of the testis and intestine largely determines surgical decision-making. Open repair of inguinal hernias and ruptures An incision is made over the superficial inguinal ring and blunt dissection is used to isolate the vaginal sac.
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Inguinal Hernia The scrotal ligament, running from the vaginal sac to the scrotum, is transected. The testis is gripped in one hand and traction is applied to the spermatic cord, while the second hand ‘milks’ the intestine back through the inguinal canal. Twisting the spermatic cord greatly facilitates this procedure. A transfixation ligature is then applied to the cord as proximal as possible and the cord is emasculated and the testis removed. Some surgeons prefer to make a small incision in the vaginal tunic prior to ligation and emasculation to ensure that no herniated viscera remains. Ligation of the tunic should prevent re-herniation; however, for added security or if the superficial inguinal ring is particularly large, it can be closed using a continuous suture of heavy gauge absorbable suture. Surgical treatment of ruptured inguinal hernias is similar, except that the intestine is encountered in a subcutaneous position. The tear in the parietal vaginal tunic is identified and the herniated intestine is replaced through it prior to being ‘milked’ back into the abdominal cavity. The tear is most often located within the inguinal canal.2 It may take some time to replace the intestine through a rent in the tunic and patience is required using a ‘last out, first in’ approach.17 The tunic is friable in foals, and care should be taken during manipulation, as further tearing or loss of this tissue may make secure abdominal closure difficult or impossible. Some surgeons advocate packing the inguinal canal to minimize the chance of eventration.5 We believe that this slows contracture of the vaginal ring, and increases the chance of post-operative infection. We recommend primary closure of tunic and skin, wherever possible. In horses affected by inguinal rupture, the intestine must be replaced through the rent in the body wall. An attempt should then be made to close the rent by suturing the aponeurosis of the external abdominal oblique muscle18 using a simple continuous or interrupted pattern with heavy gauge monofilament absorbable suture material. Closure of the deeper layers is difficult, time-consuming, and adds little, if anything, to the security of the closure. In cases with strangulated intestine, it is possible to resect devitalized tissue via an inguinal incision; however, it is generally considered easier to perform this surgery through a ventral midline celiotomy or parainguinal or paramedian incision. Patience may be required to return compromised, edematous intestine through the inguinal canal. Manual dilation of the vaginal ring, and traction on the intestines from within the abdominal cavity, may be required. A second surgeon greatly facilitates this procedure. Alternatively, the affected intestine can be resected and the stumps inverted either by oversewing or using a sur-
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gical stapler, prior to being returned to the abdomen, where an anastomosis is performed. In cases involving the terminal ileum, the only option is to pull the affected section of intestine back into the abdomen. While the anastomosis is performed, the second surgeon, if available, performs a castration on both sides. Depending on the viability of the skin, the inguinal wound may be left open to heal by second intention or closed primarily. Laparoscopic inguinal herniorrhaphy Non-strangulating inguinal herniation can also be managed laparoscopically. In foals these techniques are performed with the foal anesthetized; in adults, both recumbent and standing techniques can be used. A number of techniques are described for hernia repair, including direct stapling of the vaginal ring after removal of the testis, direct stapling of the vaginal ring without testis removal, laparoscopic suturing of the vaginal ring, transabdominal pre-peritoneal mesh repair, placement of a cylindrical mesh within the inguinal canal, and repair using the transposition of a flap of peritoneum.10,19–23 The choice of technique depends largely on the age of the horse, the size of the vaginal ring, and the intended use of the horse. In general, if the horse is not intended for breeding, the testes are pulled back into the abdomen and castration is performed at the same time as hernia repair. If the procedure is performed under general anesthesia, the Trendelenburg position is used, and this usually will lead to spontaneous resolution of the hernia. In the standing position, the contents of the hernia are grasped laparoscopically and pulled back into the abdomen.
Hemicastration and testicle-sparing techniques Testicle-sparing inguinal herniorrhaphy techniques are used when the owner wishes to maintain the animal for breeding. Prior to undertaking these procedures, the owner should be carefully counseled regarding the heritable nature of the condition, and the possibility of vascular or pressure damage to one or both testes leading to infertility. Both open and laparoscopic techniques have been described. The testis can be left in situ only if there is no doubt regarding viability. Using the open technique, the hernia is reduced using the procedure described above, and then the superficial inguinal ring is partially closed. Suturing begins at the cranial aspect of the ring and ends close to the middle of the ring, to give a snug fit around the spermatic cord but not so tight as to affect blood flow.5
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Laparoscopic techniques have been described in anesthetized and standing horses. Following reduction of the hernia, the deep inguinal ring is partially closed using laparoscopic staples or sutures, closed with a flap of peritoneum elevated from the underlying muscles, or closed with a mesh placed across the entrance to the canal. Another technique whereby the mesh is inserted into the canal has also been described.10,19–22 Simple reduction in size of the internal inguinal ring, while able to prevent re-herniation in foals, may not be sufficient in stallions owing to larger intra-abdominal forces. The use of peritoneal flaps and mesh are therefore considered superior. The use of mesh carries a risk of increased likelihood of abdominal adhesions, and mesh is placed extraperitoneally in humans for this reason. However, this technique is technically demanding and has not been evaluated in the horse to date. Placement and fixation of a cylindrical mesh within the inguinal canal, which is then allowed to unfurl and hence plug the canal, has been described and offers the advantage that it can be performed with the horse standing and sedated. Risks with this technique include adhesions within the inguinal canal and the potential for compromise of the testicular vasculature, and the possibility of displacement of the mesh into the scrotum.
Conclusions The prognosis for foals affected by congenital inguinal hernia without the presence of intestinal strangulation is generally excellent. Many of these hernias will resolve with conservative management. Foals treated conservatively should be subjected to closed castration techniques so that these animals are not put at increased risk of abdominal eventration. Foals and adult horses with evidence of strangulated intestine are emergencies and require prompt referral to a surgical facility. Expedient surgery and aftercare, including post-operative supportive care and treatment of ileus, are integral to achieving good survival rates. The development of laparoscopic techniques has greatly enhanced the anatomical understanding of the inguinal region, and as a result has increased the number of options for dealing with horses affected by inguinal hernias and ruptures. The number of techniques available is likely to increase with time, as the equipment, facilities, and those experienced in their use become more accessible. The prognosis with the use of testicle-sparing techniques is good providing the gonads remain viable. Nevertheless, owners should be counseled in the her-
itable nature of the condition prior to undertaking these procedures.
References 1. Schneider RK, Milne DW, Kohn CW. Acquired inguinal hernia in the horse: a review of 27 cases. J Am Vet Med Assoc 1982;180:317–20. 2. Van der Velden MA. Surgical treatment of acquired inguinal hernia in the horse: a review of 51 cases. Equine Vet J 1988;20:173–7. 3. Robertson JT, Embertson RM. Surgical management of congenital and perinatal abnormalities of the urogenital tract. Vet Clin North Am Equine Pract 1988;4:359–79. 4. Cox JE. Hernias and ruptures: words to the heat of deeds. Equine Vet J 1988;20:155–6. 5. Schumacher J. Testis. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St. Louis, MO: Saunders Elsevier, 2006; pp. 775–810. 6. Roberts SJ. The inherited or genetic anomalies or malformations in domestic animals. In: Veterinary Obstetrics and Genital Diseases. Newton Abbot: David and Charles, 1986; pp. 54–74. 7. Wensing CJG, Colenbrander B. Normal and abnormal testicular descent. In: Clarke JR (ed.) Oxford Reviews: Reproductive Biology, vol. 8. Oxford: Clarendon Press, 1986; pp. 125–30. 8. Van der Velden MA. Ruptured inguinal hernia in new-born colt foals: a review of 14 cases. Equine Vet J 1988;20:178– 81. 9. Spurlock GH, Robertson JT. Congenital inguinal hernias associated with a rent in the common vaginal tunic in five foals. J Am Vet Med Assoc 1988;193:1087–8. 10. Rossignol F, Perrin R, Boening KJ. Laparoscopic hernioplasty in recumbent horses using transposition of a peritoneal flap. Equine Vet J 2007;36:557–62. 11. Knottenbelt D, Holdstock N, Madigan JE. Neonatal syndromes: conditions associated with colic and tenesmus. In: Knottenbelt D, Holdstock N, Madigan JE (eds) Equine Neonatology Medicine and Surgery. Edinburgh: Saunders, 2004; pp. 247–9. 12. Wright JG. The surgery of the inguinal canal in animals. Vet Rec 1963;75:1352–8. 13. Blackford JT, Toal RL, Latimer FG, Blackford LW. Percutaneous ultrasonographic diagnosis of suspected inguinal and scrotal herniation in horses. In: Proceedings of the 38th Annual Convention of the American Association of Equine Practitioners, 1992, pp. 357–74. 14. Fischer AT, Vachon AM, Klein SR. Laparoscopic inguinal herniorrhaphy in two stallions. J Am Vet Med Assoc 1995;207:1599–1601. 15. Ragle CA, Southwood LL, Schneider RK. Injury to abdominal wall vessels during laparoscopy in three horses. J Am Vet Med Assoc 1998;212:87–9. 16. Mari¨en T, Van Hoeck F, Adriaenssen A, Segers L. Laparoscopic testis-sparing herniorrhaphy: A new approach for congenital inguinal hernia repair in the foal. Equine Vet Educ 2000;13:32–5. 17. Adams SB, Fessler JF. Neonatal inguinal herniorrhaphy. In: Adams SB, Fessler JF (eds) Atlas of Equine Surgery. Philadelphia: W.B. Saunders, 2000; pp. 219–21. 18. Adams SB. Surgical approaches to and exploration of the equine abdomen. Vet Clin North Am Equine Pract 1982;4:89–104.
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Inguinal Hernia 19. Mari¨en T. Standing laparoscopic herniorrhaphy in stallions using cylindrical polypropylene mesh prosthesis. Equine Vet J 2001;33:91–6. 20. Fischer AT. Laparoscopic inguinal herniorrhaphy. In: Fischer Jr AT (ed.) Equine Diagnostic and Surgical Laparoscopy. Philadelphia: W.B. Saunders, 2002; pp. 171–6. 21. Klohnen A. Laparoscopic inguinal herniorrhaphy (nontesticle-sparing). In: Fischer Jr AT (ed.) Equine Diagnostic
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and Surgical Laparoscopy. Philadelphia: W.B. Saunders, 2002; pp. 177–80. 22. Caron JP, Brakenhoff J. Intracorporeal suture closure of the inguinal and vaginal ring in foals and horses. Vet Surg 2008;37:126–31. 23. Klohnen A, Wilson DG. Laparoscopic repair of scrotal hernia in two foals. Vet Surg 1996;25:414–16.
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Chapter 160
Laparoscopic Surgery David G. Wilson
Laparoscopic surgery offers minimally invasive approaches for cryptorchidectomy, castration, and inguinal herniorrhaphy. This chapter details the equipment necessary to accomplish these techniques and procedure-specific descriptions of the techniques.
Instrumentation Laparoscopic observation and manipulation was first reported in the eighteenth century. Unlike arthroscopic surgery in which a single individual can often perform the procedure, operative laparoscopy requires at least one assistant, and both surgeon and assistant must be able to simultaneously view the operative field. The development of the video cameras that allowed the projection of the laparoscopic image to a video monitor revolutionized operative laparoscopy in human medicine in the late 1970s. The first use of laparoscopy in the management of the equine cryptorchid involved confirming the presence or absence of abdominal testis in the hemicastrate.1 With the development and popularization of laparoscopic cholecystectomy in humans came laparoscopic instrumentation that facilitated intra-abdominal manipulation, and the first laparoscopic cryptorchidectomy procedures in the horse were reported.2 The limitations of human instrumentation, specifically length, have largely been resolved with the development of instrumentation designed specifically for use in the horse. Laparoscopic cryptorchidectomy can be performed using a standard length (33 cm) 10 mm diameter 0◦ or 30◦ viewing scope. Angled view laparoscopes are more versatile and allow direct visual observation of instrument cannula insertion. Laparoscopes of 50 cm or more allow visual observation of both sides of the abdomen in standing horses. Video tower placement is greatly facilitated by using a longer than standard light guide; the author uses a 3-m light guide.
A wide assortment of video chip cameras are available commercially. High-quality affordable cameras are widely available through reputable secondary sources. While a standard television can be used, the image is much better on purpose-designed video monitors. Automated light intensity is available on matched systems; however, manual control of light intensity in unmatched systems (camera and light source from different manufacturers) does not present a significant problem. The equine abdomen is a relatively large cavity and 300-watt xenon light sources offer optimal illumination. Vendors claiming that their 175-watt light source is just as bright as a competitor’s 300-watt source unit should be viewed with skepticism. Abdominal insufflation is essential to visual observation and manipulation within the abdomen. Highflow (> 15 L/min) CO2 insufflators are essential to allow abdominal distension and rapid re-establishment of distension during laparoscopic manipulation. Laparoscopic cannulas allow maintenance of distension during the procedure. Disposable single-use cannula systems work well, but, for economic reasons, most surgeons utilize reusable systems. Standard length cannulas (10 cm) work well in the recumbent horse, but longer systems (20 cm) are needed in the standing horse. The author prefers a cannula with a guarded trocar for the first insertion in the recumbent horse. A wide assortment of instrumentation is available. Minimal instrumentation for laparoscopic cryptorchidectomy should include: an endolaparoscopic Babock forceps, a right-angle forceps, an acute claw grasper, scissors, and ligating loops. In the author’s experience, disposable scissors can be used repeatedly and perform better than most reusable scissors. Additional equipment that falls into the ‘nice to have’ category includes: a standard bipolar cautery forceps (Conmed; Linvatec, Largo, FL), a tripolar cautery forceps (ACMI, Southborough, MA) or
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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capacitance-monitored bipolar cautery device (LigaSureTM , Tyco Healthcare Group LP, Boulder, CO). The reader is referred to Chapters 211 and 265 for a more complete list of equipment, including photographs.
Laparoscopic cryptorchidectomy Patient selection and patient management Laparoscopic cryptorchidectomy offers a minimally invasive approach for the surgical management of the abdominal cryptorchid. The technique is especially well suited to the management of the hemicastrate and the apparent gelding that is exhibiting stallionlike behavior. The procedure can be performed in the standing or recumbent horse. Factors considered in approach selection include surgeon preference and experience as well as the horse’s temperament. The testis is easier to locate in the standing horse, but, owing to the size of the paralumbar fossa, instrument manipulation is more difficult than in the recumbent horse. To facilitate visibility within the abdomen, horses are fasted for a minimum of 12 hours for standing procedures and 36–48 hours is preferred by the author in the recumbent horse. Standing cryptorchidectomy Bilateral cryptorchids and hemicastrates with unknown castration histories are generally approached initially through the left paralumbar fossa. After confirming the status on the left side, a 50-cm or longer laparoscope can be inserted and passed ventral to the small colon to allow visual observation of the right internal inguinal area. If the right testis is retained, it can be removed from the left flank if instrumentation of appropriate length is available. In the event that the retained testis is on the right side, the procedure is performed through the right paralumbar fossa. If possible, the flank area is clipped and prepared for aseptic surgery before the horse is sedated. The author typically sedates the horse with xylazine hydrochloride (0.5 mg/kg intravenously (i.v.)) and butorphanol (0.04 mg/kg i.v.). Controlled rate infusion using a combination of detomidine and butorphanol also works well. Local anesthetic can be infused at each insertion site; however, local anesthetic infused in an inverted-L pattern is preferred by the author. A final surgical scrub is performed and the surgical site is draped. Until recently, the author routinely insufflated the abdomen before laparoscope placement using a 12-Fr thoracic catheter. Although the approach was effective, inadvertent insufflation of the retroperi-
Figure 160.1 Laparoscope and instrument cannula positioning for standing laparoscopy. x, laparoscope; o, instrument cannulas.
toneal space was a common complication, especially when surgical trainees placed the thoracic catheter. More recently, the author has adopted the technique of inserting the laparoscope cannula with its blunt obturator prior to insufflation (D.A. Hendrickson 2006, personal communication). Specifically, a 1-cm skin incision is made at the dorsal margin of the internal abdominal oblique muscle at the cranial–caudal midpoint of the paralumbar fossa. The incision is carried through the fascia of the external abdominal oblique muscle to minimize frictional resistance during insertion. A 20-cm-long cannula with a blunt obturator is inserted into the incision. Little resistance is encountered as the cannula passes through the abdominal muscles and into the retroperitoneal space. The resistance of the peritoneum is overcome with a controlled thrust similar to when performing abdominal paracentesis with a teat cannula. The blunt obturator is removed and the laparoscope is placed and insufflation is started once intra-abdominal placement of the scope has been confirmed. When the abdomen has been distended to a pressure of approximately 15 mmHg, the internal inguinal area is visually assessed. If an abdominal testis is observed, instrument cannulas are inserted cranial and caudal to the laparoscope insertion site (Figure 160.1). Another author prefers a vertical alignment of instrument cannula positioning.3 The abdominal testis is readily seen immediately ventral to the vaginal ring (Figure 160.2). An endolaparoscopic Babcock or right-angled dissecting forceps is placed through the cranial instrument cannula. A ligating loop is inserted through the caudal instrument cannula and the grasping instrument is positioned through the ligating loop. The testis is grasped and the ligating loop is manipulated onto the vascular pedicle and tightened. The free end of the ligature is cut with scissors and the vascular
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B M
T
Figure 160.2 Left abdominal cryptorchid in standing horse. B, bladder; M, mesorchium; T, testis; arrow, vaginal ring.
a reluctance to enlarge an instrument cannula incision before attempting testis removal resulted in the occasional dropped testis and, commonly, lodging of the testis in the abdominal wall. This complication is eliminated by effective enlargement of the skin and external abdominal oblique external fascia incisions. If a testis is lost after being freed from its vascular supply, avascular necrosis is likely and, although leaving the testis in the abdomen has not been evaluated, it is likely to undergo necrosis similar to that of avascular ovaries left in the abdomen.4 Post-operative management should include nonsteroidal anti-inflammatory drugs for 48–72 hours. Horses should be restricted to stall confinement for 3–5 days. Return to exercise and athletic pursuits can resume as early as 10 days after surgery. Recumbent cryptorchidectomy
structures and vas deferens are transected. The vascular stump is observed for hemorrhage. With continued development of hemostatic techniques, many surgeons have abandoned the use of the ligating loop in favor of electrocoagulation. The capacitancemonitored bipolar cautery device is user friendly and well suited to this procedure. Once the testis is free, it is transferred to an acute claw grasper placed through the caudal instrument cannula. The incision in the skin and the external fascia of the external abdominal oblique is enlarged to allow removal of the testis. At this point, the abdomen is deflated and the cannula incisions are routinely closed. Bilateral abdominal cryptorchids are commonly approached first from the left side to allow removal of the left testis and subsequently approached from the right side to remove the right testis. If a long laparoscope and long instrumentation are available, it is possible to remove the right testis from the left flank. In these cases, the right testis is identified and freed from its attachments as described above. To avoid loss of abdominal distension, removal of the right testis is delayed until the left testis has been similarly freed. To avoid losing the right testis, the author places a third instrument cannula and leaves the testis in the grasp of an acute claw grasper until the left testis is freed and removed from the abdomen. Alternatively, the right testis can be fixed to the caudal abdominal wall using an endoscopic staple. Abdominal insufflation is commonly lost during removal of the left testis and can usually be temporarily re-established by inserting a coned hand into the enlarged incision while the right testis is transferred from its grasping forceps to a forceps inserted through the enlarged incision. Complications with standing laparoscopic cryptorchidectomy are rare. Early in the author’s career,
The horse is placed under general anesthesia and positioned in dorsal recumbency. The ventral abdomen is clipped, prepared, and draped for aseptic surgery. A 1-cm skin incision is made at the umbilicus and a number 15 scalpel blade is used to make a stab incision through the linea alba. A teat cannula is inserted into the abdomen and the abdomen is distended with CO2 to a pressure of 10–15 mmHg. The teat cannula is removed and a laparoscope cannula with a safety trocar placed. The trocar is withdrawn and replaced with a 30◦ view standard length 10-mm-diameter laparoscope. When intra-abdominal placement has been visually confirmed, the hindquarters are elevated into the Trendelenburg position. As the abdominal viscera displaces cranially, the internal inguinal area generally comes into view. In the horse with a normally descended testis, the testicular vessels and vas deferens
M
Figure 160.3 Normal right vaginal ring in horse that is in dorsal recumbency. M, mesorchium; arrow, vaginal ring; short arrows, vas deferens.
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Laparoscopic Surgery enter the vaginal ring (Figure 160.3). Any departure from this appearance should cause the surgeon to suspect an abdominal testis. Although the testis may be covered by the bowel or bladder, the pampiniform plexus or the attachment of the ligament to the tail of epididymis to the vaginal process are often visible. Most surgeons employ two instrument cannulas to facilitate removal of abdominal testes. A 10- to 12-mm instrument cannula is inserted 15 mm lateral to the midline, midway between the external inguinal ring and the umbilicus. Ideally, the cannula should enter the abdomen in an area that is devoid of retroperitoneal fat because this approach may decrease the likelihood of vascular injury during insertion. To avoid inadvertent injury to abdominal viscera, the instrument cannulas are inserted with blunt obturators and under direct visual observation. When both instrument cannulas are in place, an endoscopic Babcock forceps is placed through one cannula and a right-angle dissecting forceps is inserted through the contralateral instrument cannula. The mesorchium and testicular vessels or the ligament to the tail of the epididymis is grasped and the testis is brought into view using both instruments in a ‘handover-hand’ fashion. Once the testis has been drawn from beneath the viscera, it generally remains in view. Complete assessment of both internal inguinal areas should be completed before beginning the removal of a testis. The surgeon should know whether the horse is a bilateral abdominal cryptorchid before proceeding. Over the past 20 years, the author has found two abdominal testes in more than a few horses that had reportedly already had at least one testis removed. The testis must be freed of its attachments in preparation for removal from the abdomen. Methods of achieving hemostasis include ligation, electrocoagulation (monopolar or bipolar), or capacitancemonitored bipolar electrocoagulation. Electrocoagulation using standard bipolar forceps requires switching of the cautery forceps and scissors in a repeating fashion and, though effective, the approach is time-consuming. The introduction of the capacitancemonitored electrocautery device with its cutting feature is an improvement, but there are significant costs associated with the use of the device. For these reasons, the author continues to use ligation. An endolaparoscopic ligating loop is placed through the ipsilateral instrument cannula. A Babcock forceps, placed through the contralateral cannula, is passed through the loop before the testis is grasped. The ligating loop is manipulated proximally on the vascular pedicle and tightened by advancing the knot with the pusher (Figure 160.4). The free end of the ligature is cut with scissors and the vascular pedicle and
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T
Figure 160.4 Recumbent left side cryptorchidectomy ligation. T, testis; arrow, vaginal ring.
testicular attachments are severed. The scissors are withdrawn and replaced with an acute claw grasper. The testis is transferred from the Babcock to the acute claw grasper and removed from the abdomen after enlarging the cannula incision. If the horse is a bilateral abdominal cryptorchid, removal of the testis from the abdomen is delayed until both testes have been freed of their attachments. In this case, abdominal distension is commonly lost during removal of the first testis, but can easily be temporarily restored by inserting a coned hand into the enlarged cannula incision. Alternatively, one of the instrument cannulas can be replaced with a 33-mm-diameter tissue cannula (Ethicon Endo-Surgery, Cincinnati, OH) and each testis can be removed as it is freed of its attachment. The abdomen is decompressed by opening one or more cannula. Care should be taken to ensure that no small intestine has entered a cannula or incision during the decompression process. The enlarged instrument cannula portal is closed in three layers (rectus fascia, subcutaneous tissue, and skin). Other incisions are typically closed in two layers (linea alba or rectus fascia and skin). Skin closures are routinely achieved using synthetic absorbable sutures placed in a subcuticular pattern. Post-operatively, horses are maintained on nonsteroidal anti-inflammatory drugs for 3 days. Horses can return to normal exercise in as few as 10 days post-operatively.
Laparoscopic castration Laparoscopic castration can be performed in any stallion; however, the technique is best suited to the foal, the juvenile stallion (< 1 year of age), and the inguinal cryptorchid. Approaches to laparoscopic castration include in situ ligation and laparoscopic removal of the testes.
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Laparoscopic castration in foals and juvenile stallion: testis removed Laparoscopic castration with removal of the testis can be performed in any stallion; however, the procedure is especially well suited to the foal and the stallion less than 1 year of age because their testes can readily be returned to the abdomen without enlarging the vaginal ring. In essence, the laparoscopic approach to castration involves rendering the horse an abdominal cryptorchid and then removing the testes from the abdomen. The foal is placed under general anesthesia and positioned in dorsal recumbency. The ventral abdomen is clipped, prepared, and draped for aseptic surgery. The laparoscopic approach is identical to that described for recumbent cryptorchidectomy. A right-angle forceps is introduced through one instrument cannula and a Babcock is placed through the other cannula. The mesorchium and spermatic vessels are grasped and a ‘hand-over-hand’ technique is used to draw the testis into the abdomen. Bipolar cautery is used to electrocoagulate the ligament to the tail of the epididymis and scissors are used to divide that structure, allowing the inverted vaginal tunic to return to the scrotum. The mesorchium is dissected to allow placement of a ligating loop on the vascular pedicle and scissors are used to free the testis from its attachments. To avoid any possibility of inguinal hernia, endolaparoscopic staples are used to close the vaginal ring. The procedure is repeated on the contralateral side and both testes are removed from the abdomen as described for recumbent cryptorchid castration. Post-operatively, there is no expectation of scrotal swelling. Non-steroidal anti-inflammatory drugs are administered for 3 days. Horses undergoing laparoscopic castration tend to be more comfortable than horses that have had primary closure castrations.
10 minutes (considerably faster than doing a scrotal approach with primary closure).
Laparoscopic in situ castration Castration can be achieved using in situ ligation and transection of vascular supply to the testis.5 This approach is well suited to the inguinal cryptorchid undergoing standing laparoscopy but can also be applied in the recumbent horse. The laparoscopic approach is identical to that for either standing or recumbent cryptorchidectomy. If hemostasis is achieved by ligation, a third instrument cannula facilitates the procedure. A ligating loop is placed through one instrument cannula and a right-angled dissecting forceps is placed into the abdomen and through the ligating loop. The spermatic vessel and the vas deferens are grasped in the right-angle forceps. Bipolar cautery forceps are introduced through the third instrument cannula and the vascular structures and the vas deferens are electrocoagulated distal to the right-angled forceps. The vascular structures and vas deferens are transected and the ligating loop is manipulated over the cut end and tightened. The author now uses the capacitance-monitored electrocoagulation device for this purpose and no longer uses a ligating loop. Although this technique was effective in an experimental study,5 there are two reports of failure to effect castration.6, 7 To date, it seems that the technique may fail in up to 10% of patients.7 Even though the author would not intentionally set forth to perform in situ castration in a horse discovered to be an inguinal cryptorchid during laparoscopy, the procedure offers an alternative to an open approach in either the standing or recumbent horse. The small percentage of horses that are not effectively castrated, based on hormonal criteria, are easily remedied with a standard approach to inguinal castration.
Laparoscopic castration in mature stallions: testes removed The testes can be removed laparoscopically in any stallion. The procedure is similar to the approach in foals, but the vaginal ring must be enlarged to allow the testis to be pulled into the abdomen.5 Enlarging the vaginal ring is readily accomplished with scissors and, once the testis is in the abdomen, removal is routine, although the abdominal incision must be enlarged to accommodate the large testis. That said, the technique holds no advantage over primary closure castration of the normally descended stallion. On the other hand, the author can remove a descended testis laparoscopically from a horse that is a unilateral abdominal cryptorchid in approximately
Inguinal herniorrhaphy Inguinal herniation is most common as a congenital condition in foals, although the condition does occur in the breeding stallion. In either case, the intestine (usually small intestine) exits the abdomen through the vaginal ring and is usually contained within the vaginal tunics. Intestinal strangulation is rare in foals, but common in mature stallions. Laparoscopic herniorrhaphy offers a minimally invasive approach to managing the condition. Laparoscopic techniques most commonly include concomitant castration; however, testis-sparing techniques have also been described.
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Laparoscopic Surgery Laparoscopic inguinal herniorrhaphy in foals with castration In the absence of intestinal strangulation, inguinal herniorrhaphy can be a scheduled elective procedure. The foal is placed under general anesthesia and positioned in dorsal recumbency. The ventral abdomen and inguinal area is clipped, prepared, and draped for aseptic surgery. The laparoscopic approach is similar to that described for laparoscopic cryptorchidectomy in the recumbent horse. In most cases, the hernia will reduce spontaneously when the foal is placed in dorsal recumbency. If that does not occur, atraumatic forceps are used in a ‘hand-over-hand’ fashion to return the herniated intestine to the abdomen. One clear advantage of laparoscopic herniorrhaphy over the more traditional open approaches is the ability to visually assess the viability of the involved intestine. Once the intestine has been returned to the abdomen, the remainder of the procedure is identical to that described for laparoscopic castration in foals and thus endolaparoscopic staples are used to close the vaginal ring. Irrespective of whether the condition was unilateral or bilateral, the author routinely performs the procedure bilaterally. Post-operatively, exercise is usually restricted for 10 days. Post-operative swelling is limited to the cannula insertion sites. Distension of the vaginal tunic that can occur during surgery generally resolves within 24 hours of surgery. Complications are rare and foals are considerably more comfortable after surgery than when traditional open herniorrhaphy is performed.
Laparoscopic inguinal herniorrhaphy in foals without castration The ethics of performing testes-sparing inguinal herniorrhaphy is called into question, given that there is some evidence that the condition is heritable. In spite of this concern, some owners choose to discount the heritability concerns and opt to spare the testes. Any herniorrhaphy technique that involves sparing the testis carries an increased risk of recurrence of the hernia because of the need to maintain some of the diameter of the inguinal canal. The approach to testis-sparing laparoscopic herniorrhaphy is identical to the inguinal herniorrhaphy approach described above, with the exception that the testes are left in place. To reduce the likelihood of reherniation, some surgeons partially close the vaginal ring with endolaparoscopic staples. Care must be taken not to compromise the vas deferens or the spermatic vessels when performing this procedure. More recently, one surgeon has described the use of intracorporeal suturing to partially close the vaginal ring
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(L. Boure 2003, personal communication). In this instance, the surgeon must be conscious of the risks of overly aggressive suture bites encountering structures deep to the ventromedial aspect of the vaginal ring, the area known in human medicine as the ‘triangle of doom’. The triangle of doom is demarcated by vas deferens medially, spermatic vessels laterally, and external iliac vessels inferiorly. This triangle contains the external iliac artery and vein, the deep circumflex iliac vein, the genital branch of the genitofemoral nerve, and the femoral nerve, which is hidden by fascia. Laparoscopic inguinal herniorrhaphy in mature stallions without castration: retroperitoneal mesh augmentation Laparoscopic herniorrhaphy can be performed in the mature stallion. Fischer et al.8 described meshaugmented herniorrhaphy in two stallions as an elective procedure several days after resolution of strangulated hernias. Patient selection is critical and laparoscopic examination of the abdomen may be contraindicated in horses with abdominal distension. The author has operated one stallion with an acute onset strangulated inguinal hernia. In that case, the abdomen was not distended and the strangulated small intestine was readily returned to the abdomen by applying traction with atraumatic grasping forceps. Unfortunately, when the hernia had been resolved, visual access to the internal inguinal area was compromised by overall intestinal fill and mesh-augmented herniorrhaphy was not possible. Mesh-augmented herniorrhaphy is performed with the horse under general anesthesia. To facilitate visual access, horses should be fasted for a minimum of 24 hours, although 48 hours of fasting further enhances exposure. The horse is placed in dorsal recumbency and then the ventral abdomen and external inguinal areas are clipped, prepared, and draped for aseptic surgery. The laparoscope is placed into the abdomen at the umbilicus as previously described. The hindquarters are elevated into the Trendelenburg position and two instrument cannulas are placed on the side of the hernia. Using a combination of a grasping forceps and scissors, the peritoneum is dissected and elevated beginning cranial to the vaginal ring and extending caudally to essentially develop a caudally based peritoneal flap with 360◦ exposure of the mesorchium, testicular vessels, and the vas deferens. An oval piece of mesh with a V-shaped cutout is manipulated over the mesorchium with the V directed caudally. The mesh is fixed to the underlying muscle with endolaparoscopic staples. At this point, the peritoneal flap is repositioned and stapled, thereby securing the mesh in a retroperitoneal position.
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Post-operative management should include stall rest with hand walking for a period of 30 days. Horses should not return to breeding until 60 days after surgery. Potential complications include recurrence of the hernia, possible testicular compromise due to exposure to elevated temperature during the time of herniation and possibly vascular compromise to the testicle induced by mesh placement, or possible fibrous scarring. Laparoscopic inguinal herniorrhaphy in mature stallions without castration: cylindrical mesh prosthesis Testis-sparing laparoscopic inguinal herniorrhaphy can entail placing a piece of polypropylene mesh within the inguinal canal to stimulate fibrosis of the inguinal canal to prevent herniation.9 This technique can be performed as an elective procedure in stallions with a history of inguinal hernia. Horses should be fasted for a minimum of 24 hours. The ipsilateral flank area is clipped, prepared, and draped for aseptic surgery. The laparoscope is inserted as described for standing laparoscopic cryptorchidectomy and two instrument cannulas are inserted. A 6 × 8 cm piece of polypropylene mesh is rolled into a cylinder along its long axis and maintained in that shape with a suture at either end. The cranioventral margin of the vaginal ring is grasped with a forceps and the cylinder of mesh is inserted into the inguinal canal. The proximal suture in the mesh is cut to allow the mesh to expand within the canal. Two endolaparoscopic staples are used to anchor the proximal part of the mesh to the peritoneum of the vaginal ring.
Post-operative management should include stall rest with hand walking for a period of 30 days. Horses should not return to breeding until 60 days after surgery. Some surgeons have noted difficulty in placing the mesh owing to instrument length limitations, although that issue is readily resolved with the availability of longer instrumentation (≥ 40 cm).
References 1. Wilson DG. Laparoscopy as an aid in the surgical management of the equine hemicastrate. In: Proceedings of the 35th Annual Convention of the American Association of Equine Practitioners, 1989; pp. 347–353. 2. Fischer AT, Vachon AM. Laparoscopic cryptorchidectomy in horses. J Am Vet Med Assoc 1992;201:1705–8. 3. Hendrickson DA, Wilson DG. Laparoscopic cryptorchid castration in standing horses. Vet Surg 1997;26:335–9. 4. Shoemaker RW, Read EK, Duke T, Wilson DG. In situ coagulation and transection of the ovarian pedicle: an alternative to laparoscopic ovariectomy in juvenile horses. Can J Vet Res 2004;68:27–32. 5. Wilson DG, Hendrickson DA, Cooley AJ, DegraveMadigan E. Laparoscopic methods of castration for equids. J Am Vet Med Assoc 1996;209:112–14. 6. Bergeron JA, Hendrickson DA, McCue PM. Viability of an inguinal testis after laparoscopic cauterization and transaction of its blood supply. J Am Vet Med Assoc 1998;213:1303–4. 7. Voermans M, Rijkenhuizen ABM, Van Der Velden MA. The complex blood supply to the equine testis as a cause of failure in laparoscopic castration. Equine Vet J 2006;38:35–9. 8. Fischer AT, Vachon AM, Klein SR. Laparoscopic inguinal herniorrhaphy in two stallions. J Am Vet Med Assoc 1995;207:1599–1601. 9. Marien T. Standing laparoscopic herniorrhaphy in stallions using cylindrical polypropylene mesh prosthesis. Equine Vet J 2001;33:91–6.
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Chapter 161
Laser Assisted Castration Scott E. Palmer
Introduction
Preoperative considerations
Castration is one of the most common surgical procedures performed on the male horse. Although the most common indication for castration of normal horses is the elimination of aggressive male behavior, castration is also performed as a treatment for cryptorchidism, inguinal or scrotal hernia repair, and less commonly as treatment of testicular disease, neoplasia, or injury. Castration of the horse may be performed at any age and may be safely accomplished with sedation and local anesthesia or with the horse placed under general anesthesia. The choice of surgical technique is determined by a number of factors, including temperament of the horse, the specific indication for castration, the owner’s or surgeon’s preference, available facilities and cost.1–3 Complications related to open castration of the horse are common and account for a large percentage of malpractice claims in equine practice. Common complications include hemorrhage, swelling, infection, and evisceration. Uncommon complications include hydrocele, scirrhous cord formation, and damage to the penis. Use of the carbon dioxide laser and scrotal-ablation technique for castration of the horse eliminates most of the complications typically associated with traditional open surgical techniques. Incisions appropriately made with the carbon dioxide laser and good surgical technique are characterized by good hemostasis. This effect is achieved by creation of a controlled zone of coagulation at the margin of the incision. Superpulse mode and moderate power settings are used for skin incisions to limit the zone of coagulation so that there is no significant delay in healing of the incision postoperatively. Continuous mode and higher powers are used to divide subcutaneous tissues in order to achieve maximum hemostasis.
Preoperative organization of the operating room includes pre-placement of the laser and a dedicated laser smoke evacuation system in the room. The laser should be tested by the surgeon or laser safety officer to insure nominal operation of the laser prior to surgery. Laser warning signs should be placed outside the operating room and appropriate eyewear protection must be provided to personnel working within the nominal hazard zone of the laser. A thorough physical examination and routine hematology are performed prior to surgery. Preoperative antimicrobial, anti-inflammatory and tetanus prophylaxis are administered at the discretion of the surgeon. Anesthesia is induced and the horse is placed in dorsal recumbency under general anesthesia. The inguinal region is routinely prepared and draped for aseptic surgery. A sterile 125 mm handpiece is attached to the articulating arm of the laser and the arm is draped with a sterile plastic sleeve. Smoke evacuation tubing is placed in the surgical field and attached to the smoke evacuator unit (Figure 161.1).
Surgical technique for routine castration The power of the laser is set to 15 watts and superpulse mode for creation of the skin incision. With the skin placed under tension, a midline elliptical incision is created along the greater curvature of the testicles by keeping the focusing guide of the handpiece in light contact with the skin surface as the light beam is passed smoothly along the path of the incision (Figure 161.2). Once the skin incision is complete, the laser is placed in standby mode and the power is increased
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 161.1 The carbon dioxide laser is fitted with a sterile 125 mm handpiece and the articulating arm of the laser is draped with a sterile plastic sleeve. Smoke evacuation tubing is placed adjacent to the incision to remove the laser plume.
to 40 watts and the mode is switched to continuous mode. With the laser hand piece held approximately 2 cm from the tissue surface, the elliptical incision is continued through the tunica dartos and the scrotal fascia. This portion of the incision is accomplished by placing a titanium laser rod beneath the target tissue and elevating the fascia, creating tension on the tissue plane. The laser beam is then passed over the fascia to cleanly divide the tissue. The use of the laser rod prevents the laser beam from penetrating the parietal vaginal tunic (Figure 161.3). The testicles are freed from the scrotal fascia by blunt and laser dissection, making sure that the spermatic cord is completely separated from the adjacent fascia (Figure 161.4). If the level of anesthesia is not adequate at this point, some horses will resist traction placed on the cord by the surgeon. In order to provide for maximum length of spermatic cord removal, a small amount of local anesthetic may be injected into the cord, or 5 mg of butorphanol tartrate may be administered intravenously to provide appropriate analgesia.
Figure 161.2 An elliptical skin incision is made with the carbon dioxide laser, using moderate power and superpulse mode.
Figure 161.3 Subcutaneous tissues and the dartos fascia are divided with the carbon dioxide laser, using high power and continuous mode. A titanium laser rod is used to ‘backstop’ the laser beam.
The median raphae and the base of the scrotum are removed by placing the median raphae under tension with one hand and passing the laser handpiece along the length of the median raphae at a distance of 2 cm from the tissue surface (Figure 161.5). Forty watts and continuous mode of power are used for this incision. Moist gauze is placed behind the median raphae in the plane of the incision to act as an optical backstop for the laser beam. There are numerous blood vessels found within the median raphae. Vessels less than 1 mm diameter may be coagulated with the laser, but all vessels larger than 1 mm must be ligated since laser coagulation of these larger vessels is unreliable postoperatively. The testicle and spermatic cord are placed under tension and Doyen forceps are placed transversely across the spermatic cord at the level of the external inguinal ring. A Carmalt forcep is placed across the spermatic cord distal to the Doyen forceps in order to provide a ‘seat’ for ligation of the cord. Two transfixation sutures of #2 polydioxanone suture are used to ligate the spermatic cord (Figure 161.6).
Figure 161.4 The testicles and spermatic cord are isolated by blunt and laser dissection, making sure that the spermatic cord is completely separated from the adjacent fascia.
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Laser Assisted Castration
Figure 161.5 The median raphae and the base of the scrotum are removed by placing the median raphae under tension with one hand and passing the laser handpiece along the length of the median raphae at a distance of 2 cm from the tissue surface.
An emasculator is then placed just distal to the ligatures to divide the spermatic cord (Figure 161.7). Once the emasculator is removed, the stump of the spermatic cord is cleaned of crushed muscle tissue with a moist gauze and the Doyen forceps are removed, allowing the spermatic cord remnant to retract into the inguinal ring. The second testicle and spermatic cord are removed in similar fashion. The incision is lavaged with sterile saline and all surfaces are carefully inspected for hemorrhage. The incision is closed with a simple continuous suture pattern, using #1 polydioxanone suture material. This suture engages subcutaneous and deep fascia on either side of the incision and includes the remnant of the median raphae to close dead space (Figure 161.8).
Surgical technique for cryptorchid castration Cryptorchid castration is accomplished using the same surgical approach and techniques described above, with the following modifications. The midline
Figure 161.6 Two transfixation sutures of #2 polydioxanone suture are used to ligate the spermatic cord.
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Figure 161.7 The spermatic cords are divided with an emasculator placed immediately distal to the transfixation ligatures.
elliptical skin incision is created over the greater curvature of the single dependent testicle that is positioned on the midline as the skin incision is created. Once the dependent testicle is removed, the incision is repositioned over the external inguinal ring of the cryptorchid testicle and the external inguinal ring is explored for remnants of the spermatic cord. The retained testicle is located and removed using previously described techniques and non-invasive technique whenever possible.4, 5 Closure of the skin incision is the same as for routine castration.
Postoperative care Postoperative systemic antimicrobial treatment may be continued for 3 days at the discretion of the surgeon. Phenylbutazone (4.4 mg/kg i.v. or p.o. q 24 h) is administered for 5 days. In uncomplicated cases, the horse is walked in hand for 7 days, then jogged for another week prior to a gradual return to normal daily exercise. If the castration is performed in conjunction with an orthopedic procedure such as
Figure 161.8 The incision is closed with a simple continuous suture pattern that incorporates the subcutaneous and deep fascia on either side of the incision and includes the remnant of the median raphae to close dead space.
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Stallion: Surgical Procedures of the Male Reproductive Tract
arthroscopy or internal fixation of a fracture, the aftercare for the orthopedic procedure takes precedence. Horses that are confined to the stall may develop some degree of swelling of the sheath postoperatively. This swelling is usually transient and does not require specific treatment.
Complications In the author’s experience, immediate postoperative complications following laser-assisted castration of the horse are limited to transient pain following recovery from anesthesia and swelling of the sheath in some horses. Infrequently, horses castrated with a scrotal-ablation technique will develop a seroma at the surgical site. These seromas may develop within 2 to 3 weeks of surgery and are resolved by making a stab incision on either side of the midline
incision with a # 10 scalpel blade. The incision is lavaged with sterile saline and horses are kept in light exercise.
References 1. Palmer SE. Castration of the horse using a primary closure technique. Proceedings of the Annual Meeting of the American Association of Equine Practitioners 1984; pp. 17–20. 2. Palmer SE, Passmore JL. Midline scrotal ablation technique for unilateral cryptorchid castration in horses. J Am Vet Med Assoc 1987;190(3):283–5. 3. Palmer SE. Use of lasers in urogenital surgery. In: Wolfe DF, Moll HD (eds) Large Animal Urogenital Surgery. Baltimore: Williams & Wilkins, 1999; pp. 165–78. 4. Stickle RL, Fessler JF. Retrospective study of 350 cases of equine cryptorchidism. J Am Vet Med Assoc 1987;172:343–6. 5. Valdez H, Taylor TS, McLaughlin SA, Martin MT. Abdominal cryptorchidectomy in the horse, using inguinal extension of the gubernaculum testis. J Am Vet Med Assoc 1979;174:1110–12.
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Chapter 162
Normal Field Castration Glenn P. Blodgett
Castration, orchiectomy, emasculation, ‘cutting’, and ‘gelding’ are terms used for the surgical removal of testicles. It is probably the most common surgical procedure performed on male horses. While the procedure is considered to be one of the simplistic of surgical procedures, complications from castration are a common source of malpractice actions against veterinarians.1 Castration prevents or decreases sexual behavior or aggressive temperament that may be objectionable in males not destined to become breeding stallions. Other indications include testicular neoplasia, spermatic cord trauma, hydrocele, cryptorchidism, or excessive trauma or infection of the testicles or surrounding tissues.2, 3 Also, testicles are frequently removed during the repair of inguinal and scrotal hernias. Prior to castration, all horses should be examined for any of the above described conditions and to insure the presence of both testicles in the scrotum. Most commonly, this procedure is performed when the horse is 1–2 years old. There have been reports of castration of stallions as newborns; however, the inguinal rings are often wide open at this time and herniation may be a complication. In some cases, the time is delayed to adequately assess the stallion’s potential as a sire.
Standing castration In the case of standing castration, the veterinarian is advised to first assess the stallion’s behavior in response to a manual examination of the scrotum and surrounding area as well as his calmness and willingness to cooperate in a quiet manner. Horses that respond in a negative fashion should be castrated in lateral recumbency under general anesthesia. Small breeds and younger animals are usually not good candidates for standing castration.
Standing castration is best performed under a combination of physical/mechanical (twitch) and chemical restraint along with local anesthesia. Detomidine (0.011–0.022 mg/kg intravenously (i.v.)), alone or in combination with butorphanol (0.011–0.022 mg/kg i.v.) is a good choice of chemical restraint. The right-handed surgeon should stand on the left side of the stallion. A surgical preparation of the area of the scrotum is performed. The spermatic cords are grasped with the left hand and 10–20 mL of lidocaine is injected in and around each cord. As an alternative, some may elect to inject 10–25 mL of lidocaine directly into the parenchyma of each testicle. Either of these injections can be performed with a 19- to 20-gauge 1 12 -inch needle. Anesthesia of the skin can be performed by infiltrating lidocaine subcutaneously along each side of the median raphe. After a few minutes the surgeon should grasp the two spermatic cords with the left hand and fully descend both testicles down into the scrotum. Next an incision can be made through the skin all the way into the body of each testicle in a craniocaudal direction. These incisions should be equidistant from and parallel to the median raphe and into the midline longitudinally of each testicle. The testicles are then removed one at a time with a pair of emasculators carefully placed proximal to the testicle and associated structures. The spermatic cord and as much vaginal tunic as possible should be transected in this procedure. In the case of older stallions or stallions with larger testicles it is desirable to bluntly perforate or tear through the mesorchium to separate the spermatic cord and ductus deferens from the vaginal tunic and attached cremaster muscle. Once separated, these structures can be transected and removed separately with emasculators before moving on to the next testicle. The above described procedure is an open standing castration and the incisions are left to drain and subsequently heal by secondary intention.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 162.1 The upper hind limb is secured to the base of the horse’s neck with a large diameter cotton rope.
Castration in lateral recumbency Castration of stallions in lateral recumbency under general anesthesia is considered to be the safest approach for personnel performing the procedure. For this approach, the horse is administered xylazine (1.0 mg/kg i.v.) as a pre-anesthetic followed in 5–10 minutes by a combination of ketamine (2.0 mg/kg) and diazepam (0.03–0.05 mg/kg) mixed in the same syringe and administered intravenously. This anesthetic regimen will generally provide 12–18 minutes of analgesia during the procedure and also provides a smoother induction and recovery than alternative choices of short-acting anesthetic combinations. The horse is positioned with the upper hind limb drawn cranially toward the shoulder by a rope secured around the base of the horse’s neck to the pastern. The rope is used to encircle the foot and sometimes the hock to insure the leg stays in a flexed position (Figure 162.1). As an alternative in the presence of an able-bodied assistant, the upper hind limb can be flexed and held with the assistant assuming a crouched position (Figure 162.2) in a frog-like fashion. For the right-handed surgeon the procedure is most easily performed with the horse lying on his right side. In the event the surgeon is unsure about the security of the leg rope or the assistant restraining the hind limb, the castration procedure may be more safely performed by reaching over the back or flank of the horse. In this case, the horse is best positioned on his left side for a right-handed surgeon. The scrotum is prepared for aseptic surgery. Removal of the bottom of the scrotum is performed in our practice to gain best access to the testicles and allows for good post-surgical drainage. This procedure may be performed by placing towel clamps in the skin centrally on the median raphe in the scrotal wall. The bottom of the scrotum is retracted away from the body wall in a tent-like fashion with the
Figure 162.2 The upper hind limb is secured in a flexed position by an able-bodied assistant and the scrotum is aseptically prepared for surgery.
attached towel clamp. Three or four small skin incisions are made with a scalpel blade midway between the attachment of the towel clamp and the base of the scrotum (Figure 162.3). These incisions are connected while traction is applied to the towel clamp, allowing
Figure 162.3 The scrotum is retracted in a tent-like fashion by a towel clamp centrally placed on the median raphe and three or four incisions are made and subsequently connected to excise the scrotum.
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Normal Field Castration
Figure 162.4 Placement of a towel clamp in the distal body of the testis will aid in blunt dissection and exposure of structures to be removed.
the bottom of the scrotum to be completely excised. Another commonly used technique for castration is to make two parallel incisions, with each incision approximately 1 cm either side of the median raphe. The tissue between the two incisions can also be removed. In the author’s opinion, fewer complications and better drainage occur if the entire ventral portion of scrotum is resected. The incision is carried on beneath the skin through the tunica dartos and underlying scrotal fascia. An alternative approach is to force the testicle to the bottom of the scrotum and incise directly through the skin and vaginal tunic into the testicle body (if an open castration is planned) similar to the standing castration method. If an open castration technique is performed, the common vaginal (parietal) tunic is also incised and the caudal ligament (remnant of the gubernaculum which connects the common vaginal tunic to the epididymal tail and attached testes) is severed. The spermatic cord is exposed and transected with an emasculator close to the external (superficial) inguinal ring. In an open castration, the common vaginal tunic is typically not removed. In the case of a closed castration, the common vaginal tunic is not excised until the spermatic cord is transected. The common vaginal tunic, along with its contents (which include the testis, epididymis, and spermatic cord as well as the attached cremaster muscle), is retracted and freed from the surrounding scrotal fascia by blunt dissection. The placement of a pair of towel clamps into the distal body of the testis will aid in retraction and blunt dissection of the structures to be removed (Figure 162.4). The cord can be transected with conventional emasculators, or a Henderson castration instrument can be utilized to facilitate the twisting of the spermatic cord and attached cremaster muscle until detachment occurs with the aid of an attached variable-speed drill (Figure 162.5).
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Figure 162.5 A Henderson castration instrument can be attached to the spermatic cord proximal to the testis to facilitate twisting and severing of the spermatic cord.
If an emasculator (Figure 162.6) is utilized to crush and transect the spermatic cord, care must be taken to insure proper placement of the instrument and that it is in good working condition. The crushing component of the emasculator should be closest to the body of the horse and the cutting component should be closest to the body of the testicle. For quick reference, the visible nut or screw seen on a pair of emasculators should be placed closest to the body of the testicle (often paraphrased as ‘nut to nut’). Placement of this instrument during the emasculation process is at a right angle to the tissues to be transected. The instrument should be left in the closed position for 30–60 seconds. This time may be increased in horses aged 2 years or older. In the case of a large cord, the common vaginal tunic and cremaster muscle can be separated and transected separately. If the Henderson instrument is utilized, it should be securely attached to the chuck of a 14.4-volt variable-speed electric drill with a 3/8 chuck. A cordless drill with a detachable chuck is preferred. The Henderson instrument, which resembles a large
Figure 162.6 (1) Serra emasculator. (2) Henderson equine castrating instrument. (3) White’s improved emasculator.
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Stallion: Surgical Procedures of the Male Reproductive Tract the wound is to be sutured, complete hemostasis is also required and the cord would need to be ligated proximal to the point of transection. For this purpose, the skin incision is closed with a synthetic absorbable suture material, utilizing a continuous intradermal pattern. No attempt is made to close dead space.
Post-operative management
Figure 162.7 Initially upon rotation of the drill, the testis is allowed to partially retract into the scrotum.
curved forceps or clamp, is placed across the entire cord and associated cremaster muscle. The clamping portion of the instrument is placed just above the main body of the testis. The drill and instrument should be held in a straight line with the cord. For this procedure, the drill motor should be started slowly to initiate twisting of the spermatic cord. Initially upon rotation, the testis will retract partially into the scrotum (Figure 162.7). After 5–10 revolutions, the speed of the drill can be increased in concert with gradual traction on the drill away from the body wall. After 20–25 revolutions, the cord and attached structures will be severed by this cord-twisting mechanism (Figure 162.8). In the author’s opinion, utilization of the Henderson instrument will substantially reduce bleeding associated with castration. Whatever method of castration is utilized, care should be taken to avoid incorporation of scrotal skin during transection of the spermatic cord. Following removal of the testicles, some fascia or tunic may protrude from the incision. This may be removed with surgical scissors or emasculators. The scrotum wound is usually left unsutured to allow drainage. If
Figure 162.8 After 20–25 revolutions, the cord and attached structures will be severed in a tightly twisted fashion.
All horses should receive, or be current with regard to, tetanus immunization. Some form of regulated exercise is recommended for several days following castration to reduce localized swelling. Antibiotics are usually unnecessary unless conditions under which the procedure was performed so dictate. The application of a fly repellant product surrounding the resulting wound may be seasonally indicated.
Complications of castration The most common post-operative complication is excessive hemorrhage. The spermatic vessels, especially the spermatic artery, may not be crushed properly. Some vessels associated with the skin or large pudendal vessels in the area may also be compromised. The spermatic cords of older horses are usually larger and may require special handling, such as double emasculation, ligature placement above the site of emasculation, or transaction. The horse will likely require some form of additional sedation and, in some cases, general anesthesia to stop the uncontrolled hemorrhage. If the severed ends of the vessel(s) are not accessible, rolled gauze may be temporarily sutured into the wound for 12–24 hours and then removed to stop the excessive hemorrhage. Edema or swelling of the prepuce and scrotum may occur post-operatively. This may occur because of lack of exercise or inadequate drainage of the wound. If the bottom of the scrotum is excised instead of making two single incisions without the tissue between them resected, swelling is less likely to occur. Hydrotherapy and manual establishment of wound drainage are often the treatment of choice, along with controlled exercise. Sepsis of the area may also cause swelling and can be addressed with antimicrobials in addition to the above described supportive treatment. Clostridial infections, although rare, can be particularly dangerous and may require aggressive treatments with penicillin, non-steroidal anti-inflammatory agents, and other supportive therapy. Some infections may lead to peritonitis and require a lavage of the peritoneal cavity. If infections of this area are left untreated, the stump may become enlarged and develop abscessation. In some cases, a piece of the vaginal tunic
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Normal Field Castration may be incorporated into the enlarged infected stump and is sometimes referred to as a scirrhous cord. The scirrhous cord becomes adhered to the scrotal skin and draining tracts are often present. Surgical removal of the scirrhous cord is necessary. This procedure should be performed under general anesthesia in dorsal recumbency. An incision is made over the mass and the entire infected portion is bluntly dissected away from the surrounding structures and transected as far proximally as possible above the involved area. This transaction is best performed with an emasculator. The wound is left open to heal by second intention. Post-operative management is the same as routine castration with the addition of antimicrobials for several days. A hydrocele, also called a vaginocele or ‘water seed’, may appear a few months following castration. It may appear as a circumscribed fluid-filled painless swelling and often resembles a swollen testis or inguinal hernia. Hydrocele can occur in stallions as well as in castrated horses. This swelling is the result of the accumulation of fluid within the vaginal cavity. The highest incidence appears in castrated mules. Open castration predisposes the horse to this condition when a portion of the vaginal tunic is not removed and the treatment is removal of this vaginal tunic remnant. It is recommended that when mules are castrated this vaginal tunic be removed. Evisceration is an uncommon complication that may have a breed association, particularly of Standardbreds. Most evisceration occurs within 4–6 hours of castration; however, longer time intervals have been reported. Evisceration is a true emergency and results in bowel contamination and strangulation. Rapid replacement requires general anesthesia and the horse may need referral and transport to a facility where a cleaner environment, experienced surgeons, and the necessary intraoperative support personnel are provided. Prior to referral the horse should
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have the bowel contained in the scrotum or continued eventration will occur. This may necessitate a rapid anesthetic prior to referral. Evisceration should be distinguished from prolapse of the greater omentum. This condition can often be corrected by transaction of the omentum followed by tension replacement by rectal palpation. Continued stallion-like behavior is a reported complication of castration. These individuals may display both masculine and breeding behavior. It has been suggested that if a portion of the epididymis were not removed at castration it may lead to male behavior. The epididymis and spermatic cord remnants are incapable of producing androgens, because testosterone is synthesized in the Leydig cells of the testis.4
Conclusions Castration is a common procedure that clients and veterinarians often take for granted. The complication rate (swelling and or infection) may be as high as 10% of all open castrations. Good surgical technique and good post-operative management are important in reducing complications.
References 1. Vaughan JT. professional liability in equine practice, risk management, and loss control. Proc Am Assoc Equine Pract 2008;54;110–19. 2. Walker DF, Vaughan JT. Surgery of the testes. In: Walker DF, Vaughan JT (eds) Bovine and Equine Urogenital Surgery. Philadelphia: Lea & Febiger, 1980; pp. 145–69. 3. Varner DD, Schumacher J, Blanchard TL, Johnson L (eds). Castration Techniques in Diseases and Management of Breeding Stallions. Goleta, CA: American Veterinary Publications, 1991; pp. 143–58. 4. Dohle GR, Smit M, Weber RFA. Androgens and male fertility. World J Urol 2003;21:341–5.
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Chapter 163
Surgery of the Penis and Prepuce James Schumacher and Dickson D. Varner
Indications for surgery of the penis and prepuce include penile and preputial trauma, paraphimosis, phimosis, priapism, genital neoplasia, cutaneous habronemiasis, genital anomalies, urolithiasis, and urethral rents causing hematuria and hemospermia. The most common indication for preputial and penile surgery is genital trauma, and the next most common indication is genital neoplasia.1 Genital trauma and neoplasia account for over 50% of penile and preputial surgery performed.
Segmental posthetomy Indications Segmental posthetomy, or excision of a circumferential segment of the internal preputial lamina, is indicated to remove preputial lesions, such as carcinomas or granulomas caused by cutaneous habronemiasis, so extensive that the lesions cannot be simply excised, provided that the preputial lesions do not extend into the underlying tunica albuginea. By removing nearly the entire internal preputial lamina (i.e., Adam’s operation), a paralyzed penis can be permanently retained within the preputial cavity.2 Other names for segmental posthetomy include reefing, posthioplasty, or circumcision. Surgical technique Segmental posthetomy is most easily and safely performed with the horse anesthetized and in dorsal or lateral recumbency, but the procedure can be performed with the horse sedated after desensitizing the internal preputial laminae by anesthetizing the pudendal nerves at the ischial arch. The sites of injection are on the right and left sides of the anus, about
2 cm dorsal to the ischial arch and 2 cm lateral to the anus. A 21- or 22-gauge, 3.81-cm (1 12 -inch) needle is inserted at each site and angled ventrally toward the midline, until the point of the needle contacts the ischial arch. Local anesthetic solution (5–10 mL) is deposited adjacent to each nerve. The penis usually protrudes within 5 minutes. A catheter is inserted into the lumen of the urethra, and the penis and prepuce are prepared for surgery. The penis is extended by traction on gauze looped around the collum glandis, and a tourniquet is positioned around it proximal to the lesions to decrease hemorrhage. Two parallel, circumferential incisions through the internal preputial lamina are created, one distal and one proximal to the lesion(s), taking care to avoid transecting the large, longitudinal, branches of the external pudendal vessels that lie in the loose adventitia beneath the preputial lamina. The amount of internal preputial lamina that can be removed from a stallion without inhibiting copulatory ability has not been determined. If segmental posthetomy is performed to retain a paralyzed penis within the preputial cavity (i.e., the Adam’s procedure), the proximal, circumferential preputial incision should be made close to the preputial orifice, and the distal, circumferential preputial incision should be made close to where the internal preputial lamina inserts on the free body of the penis.2 The preputial lamina is incised longitudinally to connect the two parallel incisions, and the segment of prepuce between the circumferential incisions is freed from the loose adventitia with scissors, being careful to avoid transecting the large, subcutaneous vessels. Integument to be apposed is maintained in proper orientation by inserting a suture at the dorsal and the ventral margin of each
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 163.1 Segmental posthetomy or reefing procedure. Two circumferential incisions through the integument of the internal preputial lamina are created, and the incisions are connected with a longitudinal incision. The segment of prepuce between the circumferential incisions is freed from the loose adventitia, and the adventitia is apposed with interrupted absorbable sutures.
circumferential incision before the segment of tissue is removed. The tourniquet is removed, and bleeding vessels are cauterized or ligated with absorbable suture. Adventitia is apposed with interrupted no. 2–0 or 3–0 absorbable sutures (Figure 163.1), and the integument is apposed with no. 2–0 absorbable or nonabsorbable sutures. When removing nearly all the internal lamina of the prepuce to permanently retain a paralyzed penis in the preputial cavity, the distal circumferential incision is sometimes difficult to suture to the much larger, proximal circumferential incision.2 In this case, two triangles of epithelium, each about 3 cm wide and 4 cm long, whose base is the circumferential incision, are removed proximal to the posthetomy, on each side of the penis. Suturing the sides of the triangles decreases the circumference of the proximal incision so that it better matches that of the smaller, distal incision. Aftercare Stallions should be isolated from mares for 2–4 weeks and should wear a stallion ring during this time. Regular light exercise during this period of confinement may reduce edema. Non-absorbable sutures are removed at 12–14 days.
Bolz technique of phallopexy Indications The Bolz technique of phallopexy is performed to permanently retain a paralyzed penis in the preputial cavity so that partial phallectomy is avoided.3 Phallopexy is contraindicated if the penis or internal
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Figure 163.2 Bolz technique of phallopexy. The penis is exposed through a 10- to 12-cm incision on the perineal raphe that begins caudal to the scrotum or scrotal scar and extends caudally.
preputial lamina is severely damaged, unless the damaged segment of the internal preputial lamina can be excised at the same time. The horse can be castrated at the same time phallopexy is performed, but the scrotal incision should be sutured so that the likelihood of infection at the adjacent site of phallopexy is reduced. Surgical technique The procedure is performed with the horse anesthetized and in dorsal recumbency. The urethra is catheterized so that it is easily identified during the procedure. The shaft of the penis is exposed through a longitudinal incision on the perineal raphe that begins just caudal to the base of the scrotum or scrotal scar and extends 10–12 cm caudally (Figure 163.2). The penis is bluntly separated from fascia, taking care to avoid damaging the large pudendal vessels that surround it. The penis is retracted into the preputial cavity through the incision so that the annular ring at the reflection of the internal preputial lamina onto the free body of the penis is visible at the cranial aspect of the incision (Figure 163.3). The penis is fixed in the preputial cavity with two heavy, non-absorbable sutures that pass through the skin, one on each side of the incision, and then through the annular ring, one on each side of the penis (Figure 163.4). Each suture is inserted through the skin at the level of the center of the incision, 2–4 cm abaxial to the margin of the incision. Each suture is passed through the annular ring on the ipsilateral lateral surface of the penis, taking care to avoid entering the preputial cavity, the urethra, or a corporeal body. The fornix of the preputial cavity should be palpated by an assistant during placement of each suture through the annular ring to confirm that the suture does not pierce the preputial epithelium. Each suture should exit the skin
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Figure 163.3 Bolz technique of phallopexy. The penis is bluntly separated from fascia and retracted through the incision so that the free portion of the penis is retracted into the preputial cavity.
2–3 cm from its entry point. Each suture is tied over a large button or gauze roll, to prevent the suture from cutting through the skin from pressure necrosis (Figure 163.5). The sutures are tightened until the distal end of the penis is situated at the level of the preputial orifice. The incision is closed in two layers. The anchoring sutures are removed after 12–14 days, at which time fibrous adhesions of adequate strength to maintain the penis in its retracted position have formed.1 Skin beneath the buttons or gauze rolls inevitably necroses, but the percutaneous placement of the sutures allows tension on the sutures to be adjusted, if necessary, so that the penis can be repositioned. Necrosis of skin can be avoided by using two heavy, absorbable sutures that anchor the annular ring to the subcutaneous fascia, rather than to skin,4 but these sutures cannot be adjusted after surgery. When insufficient tension is applied to the anchoring sutures, the penis may protrude, imparting an unac-
Figure 163.5 Bolz technique of phallopexy. Each suture is tied over a gauze roll, to prevent the suture from cutting through the skin from pressure necrosis. By the time these anchoring sutures are removed after 12–14 days, fibrous adhesions have formed, preventing the penis from extruding from the preputial cavity.
ceptable cosmetic appearance to the horse, and when excessive tension is applied, the horse may urinate into the preputial cavity causing posthitis. Retracting the penis into the preputial cavity produces acute bends in the penis, but urination and penile blood supply are not impeded. Aftercare The horse should be walked frequently to reduce swelling at the surgery site and can be allowed unrestricted exercise after 4–5 weeks.
Cavernosal shunting Indications Cavernosal shunting is a technique used to treat horses for priapism, a condition characterized by persistent erection without sexual excitement, after medical therapy and irrigation of the corpus cavernosum penis (CCP) have failed to bring about detumescence of the penis. Circulation through the CCP is shunted through the corpus spongiosum (CSP), which is not involved in the persistent erection. Surgical technique
Figure 163.4 Bolz technique of phallopexy. The penis is fixed in the preputial cavity with two heavy, non-absorbable sutures, one on each side of the penis. Each suture passes through the skin and the annular ring at the reflection of the internal preputial lamina onto the free body of the penis.
The shunt is created with the horse anesthetized and in dorsal recumbency. A stallion catheter is inserted into the urethra, and the horse’s penis and perineum are prepared for surgery. The shaft of the penis is exposed through a longitudinal incision on the perineal raphe that begins about 5 cm caudal to the base of the
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Figure 163.6 Cross-sectional and three-dimensional views demonstrating creation of a shunt between the corpus spongiosum penis (CSP) and corpus cavernosum penis (CCP). 1, Retractor penis muscle (not shown in three-dimensional view); 2, bulbospongiosus muscle; 3, CSP surrounding the urethra; 4, CCP; 5, tunica albuginea of CSP; 6 tunica albuginea of CCP. The right or left border of the bulbospongiosus muscle is separated and elevated from the edge of the urethral groove to expose a segment of the tunica albuginea surrounding the CSP. The tunica albuginea of the CCP and that of the CSP adjacent to the edge of the urethral groove are incised longitudinally. The abaxial edge of the incision in the CCP is sutured to the axial edge of the incision in the CSP. The edge of the bulbospongiosus muscle is sutured to its origin on the tunica albuginea at the urethral groove. (Modified from Schumacher J: The penis and prepuce. In: Auer JA, Stick JA (eds): Textbook of Equine Surgery. 3rd ed., page 821, W.B. Saunders Co. Philadelphia, © Elsevier 2006).
scrotum (or scrotal scar) and extends 12–15 cm caudally. Care should be taken to avoid damaging the large pudendal vessels that surround the shaft. The bulbospongiosus muscle, which covers the ventral aspect of the CSP, is identified, and its right or left border is separated and elevated from the edge of the urethral groove to which it attaches to expose 4–5 cm of the CSP, which is surrounded by a thin tunica albuginea. The CSP and its tunic are easily identified because the CSP surrounds the catheterized urethra. A 3- to 4-cm longitudinal incision is made through the tunica albuginea of the CCP about 5–7 mm adjacent to the edge of the urethral groove, and dark, sludged blood is evacuated from the CCP through this incision by infusing irrigation solution, under pressure, into the CCP just proximal to the glans penis through a 12- or 14-gauge needle (Figure 163.6). A 3- to 4-cm longitudinal incision is made through the tunica albuginea of the CSP adjacent to the incision in the CCP, taking extreme care to avoid exposing the lumen and the catheterized urethra. Bright red blood flows heavily through the incision into the CSP, making suction a necessity at this point in the procedure. The shunt is completed by suturing the abaxial edge of the incision in the CCP to the axial edge of the incision in the CSP with a no. 2–0 monofilament absorbable suture placed in a simple continuous pattern. The edge of the bulbospongiosus muscle is sutured to its origin on the tunica albuginea at the
urethral groove with the same suture using a simple continuous pattern. The subcutaneous tissue is similarly sutured, and the skin incision is closed with the same suture using a continuous pattern placed intradermally. Horses show no signs of discomfort after surgery, and swelling at the surgical site is slight. Complications Men who receive cavernosal shunting as treatment for priapism sometimes develop complications, which include urethrocavernous or urethrocutaneous fistula, penile gangrene, cavernosal infection, pain during erection, and impotency.5, 6 Impotency may develop from failure to attain cavernosal pressure required for intromission, but this failure may be due to damage to corporeal tissue rather than from the shunt itself. The shunt may close when blood outflow in the CCP returns to normal, but closure may not be necessary for return to potency, provided that the trabeculae in the CCP have not been damaged severely. In a study using stallions with normal erectile and ejaculatory function, a shunt created between the CCP and the CSP of the stallions did not interfere with erection and ejaculation, even if the shunt remained patent.7 Based on the results of this study, investigators concluded that impotency after creation of a cavernosal shunt for treatment of stallions
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affected by priapism is most likely the result of trabecular fibrosis, rather the shunt.
can no longer dilate when the CSP becomes engorged with blood.
Amputation of the urethral process
Partial phallectomy
Indications
Indications
The urethral process is sometimes excised to remove a neoplastic lesion or a granuloma caused by cutaneous habronemiasis, when the lesion fails to resolve using more conservative therapy.
The most common indication for partial phallectomy of horses is treatment for penile or preputial neoplasia that is so extensive that more conservative methods of treatment, such as cryosurgery, local excision, or segmental posthetomy, are impractical. Partial phallectomy is also performed as a treatment for horses whose penis is permanently paralyzed, especially if the penis is irreparably damaged. Stallions that receive partial phallectomy must usually be used for purposes other than breeding, but amputation of only the glans penis may not interfere with copulation.8
Patient preparation The urethral process can be amputated with the horse sedated after infiltrating the base of the urethral process with local anesthetic solution or anesthetizing the pudendal nerves at the ischial arch, but the process is most easily and safely amputated with the horse anesthetized and in lateral or dorsal recumbency. The distal end of the penis is prepared for surgery, and a stallion catheter is inserted into the urethra. Traction is placed on the catheterized urethral process with Allis tissue forceps, and two smallgauge needles (e.g., 23-gauge) are placed perpendicular to each other through the urethral process and the catheter, proximal to the diseased portion of the process. These needles anchor the urethral process to the catheter, making the incised margin of the process more accessible for suturing. Surgical technique The urethral process is amputated by circumferentially incising all its layers, proximal to the lesion and distal to the anchoring hypodermic needles. The incision extends through integument, CSP, and urethral mucosa. The stump of the process is closed with a simple interrupted or simple continuous pattern using a no. 4–0 or 5–0 absorbable suture that passes through the integument, the CSP, and urethral mucosa. The sutures should be closely spaced to compress the cavernosal tissue of the CSP. A simple continuous suture pattern is probably more effective in compressing the erectile tissue of the CSP, and thus reducing hemorrhage from this structure, than is the simple interrupted suture pattern. Aftercare Stallions should be isolated from mares for at least 3 weeks after the process has been amputated. Hemorrhage from the remnant of the process, especially at the end of urination, should be expected for at least several days. Removal of the urethral process may result in fibrosis at the site of excision, and this fibrous tissue may tear at the time of breeding because it
Patient preparation A stallion should be castrated at least 3 weeks before receiving partial phallectomy, if possible, to avoid post-operative erection, which may lead to dehiscence or excessive hemorrhage. The penis sometimes can be amputated with the horse sedated after anesthetizing the pudendal nerves at the ischial arch, but the penis is most easily and safely amputated with the horse anesthetized and in dorsal recumbency. The urethra is catheterized with a stallion urinary catheter, and a tourniquet is placed proximal to the proposed site of transection. The penis is extended slightly with gauze looped around the collum glandis. Surgical techniques
Vinsot’s technique of partial phallectomy Vinsot’s technique is the simplest technique of partial phallectomy. Using this technique, a triangular section of tissue, the apex of which points distally and which includes the integument and underlying bulbospongiosus muscle and CSP, is excised from the ventral aspect of the penis about 4–5 cm proximal to the proposed site of transection. Care should be taken not to incise into the urethral lumen. The exposed triangular section of urethral mucosa is incised from the center of its base to its apex, and all layers of the triangle are apposed with no. 3–0 or 2–0 absorbable sutures using a simple interrupted or simple continuous suture pattern (Figure 163.7). Sutures should include the tunica albuginea of the CSP, and should be closely spaced to compress the corporeal tissue of the CSP. A simple continuous suture pattern is probably more effective in compressing the CSP, and thus
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Figure 163.7 Vinsot’s technique of partial phallectomy. A triangular section of integument, bulbospongiosus muscle, and corpus spongiosum penis is excised from the ventral aspect of the penis proximal to the proposed site of transection. The triangular section of urethral mucosa is incised from the center of its base to its apex, and the mucosa is sutured to the sides of the triangle. To prevent hemorrhage, a ligature of heavy suture material is placed around the penis distal to the apex of the triangle, and the penis is severed distal to this ligature.
decreasing hemorrhage, than is a simple interrupted suture pattern. The diseased portion of the penis is excised 4–5 cm distal to the newly created urethral stoma with a wedge-shaped incision. Large branches of the external pudendal vessels on the dorsal and lateral aspects of the penis are ligated with absorbable suture, and the corporeal bodies are compressed with no. 0 or 1 absorbable sutures placed through the tunica albuginea using an everting or appositional pattern. The penile or preputial integument is sutured in an everting or appositional pattern using a no. 2–0 or 0 absorbable or non-absorbable sutures. Rather than closing the end of the penile stump primarily, the surgeon can leave the stump unsutured to heal by secondary intention. To prevent the cavernous tissue and vessels on the dorsal and lateral aspects of the penis from hemorrhaging, a tight ligature of heavy suture material, such as umbilical tape, is placed around the penis 2–3 cm distal to the apex of the triangle before the penis is severed 1–2 cm distal to this ligature (Figure 163.7). Vinsot’s technique can be further simplified by creating a 4- to 5-cm longitudinal incision on the ventral midline of the penis into the urethral lumen rather than removing a triangular section of tissue (J.R. Joyce, personal communication). The incised urethral mucosa and penile or preputial integument are apposed with no. 3–0 or 2–0 absorbable sutures using a simple interrupted or simple continuous pattern. These sutures should include and compress the cavernous tissue of the CSP. A complication of Vinsot’s technique, or its modifications, is the tendency of the urethral stoma to stricture.
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Figure 163.8 Williams’ technique of partial phallectomy. A triangular section of integument, underlying bulbospongiosus muscle, and corpus spongiosum penis is removed from the ventrum of the penis. The triangular section of urethral mucosa is incised from the center of its base to its apex, and the mucosa is sutured to the sides of the triangle. The penis is transected obliquely at the base of the triangle, so that the dorsal border of the penile stump is slightly longer than the ventral border.
Williams’ technique of partial phallectomy The likelihood of urethral stricture is decreased when the penis is amputated using Williams’ technique.8 Using this technique, a triangular section of integument and underlying bulbospongiosus muscle and CSP, the dimensions of which are similar to that described in Vinsot’s technique, is removed from the ventrum of the penis, but the triangle’s apex is directed proximally, rather than distally, and the base of the triangle is the site of penile amputation (Figure 163.8). Removing a triangular section of tissue exposes the urethra, which is incised from the center of the base to the apex of the triangle. As described by Williams, the incised edge of the urethra and the triangle’s integument are apposed with no. 3–0 or 2–0 absorbable sutures using a simple interrupted or simple continuous suture pattern. These sutures include and compress the cavernous tissue of the CSP. A simple continuous suture pattern is probably more effective in compressing the CSP, and thus decreasing hemorrhage, than is a simple interrupted suture pattern. The base of the triangle is temporarily left unsutured. To provide better hemostasis, the Williams technique can be modified by suturing the urethral mucosa to the tunic of the urethral groove to compress the transected end of the CSP, using a continuous suture pattern, before the urethral mucosal is sutured to the integument. Compressing the CSP at the base of the triangle with sutures placed in a simple continuous pattern though the urethral mucosa, CSP, and overlying tunica albuginea also decreases postoperative hemorrhage.
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Figure 163.9 Williams’ technique of partial phallectomy. As described by Williams,8 the end of the stump is closed with heavy sutures, placed in a simple interrupted pattern, that pass through the urethral mucosa, the tunica albuginea of the urethral groove, the tunica albuginea of the dorsal and lateral aspects of the corpus cavernosum penis, and the penile or preputial integument.
The urethral catheter is removed, and the penis is transected obliquely at a 30–45◦ angle, in a craniodorsal direction, at the base of the triangle, so that the dorsal border of the penile stump is slightly longer than the ventral border (Figure 163.8). The transected ends of the large branches of the external pudendal vessels that reside in loose adventitia on the dorsal and lateral aspects of the penile stump are readily apparent at the end of the stump. These vessels are ligated with multiple no. 2–0 or 0 absorbable sutures. As described by Williams, the end of the stump is closed with no. 0 or 1 absorbable or non-absorbable sutures, placed in a simple interrupted pattern, that pass through the urethral mucosa, the tunica albuginea of the urethral groove, the tunica albuginea of the dorsal and lateral aspects of the CCP, and the penile or preputial integument (Figure 163.9). Preplacing the sutures at equidistant intervals provides an even closure. Tightening the pre-placed sutures compresses the corporeal bodies and apposes the penile or preputial integument to the urethral mucosa. Closing the end of the stump in two layers provides better hemostasis and better apposition of urethral mucosa to the integument. To close the end of the stump in two layers, the tunica albuginea of the urethral groove is sutured to the tunica albuginea of the dorsum of the CCP with interrupted no. 0 or 1 absorbable sutures placed at bisecting intervals. The penile or preputial integument is then sutured to the urethral mucosa with interrupted no. 2–0 absorbable or non-absorbable sutures.
Scott’s technique of partial phallectomy With this technique, the penile or preputial integument is incised circumferentially at the intended site
Figure 163.10 Scott’s technique of partial phallectomy. The penile or preputial integument is incised circumferentially at the intended site of amputation, and, after ligating the vessels that overlie the tunica albuginea, the incision is continued through the corpus cavernosum penis (CCP) and the corpus spongiosum penis (CSP) to expose the urethra. A 4- to 5-cm segment of urethra is dissected from the excised portion of penis and transected.
of amputation.9 After identifying and ligating the large branches of the external pudendal vessels that overlie the tunica albuginea, the incision is continued perpendicular to the long axis of the penis through the CCP to the urethral groove. The CSP, which encircles the urethra, is incised circumferentially to expose the urethral mucosa, which is easily recognized by the urinary catheter within its lumen, and a 4- to 5-cm segment of urethra is dissected from the excised portion of penis and transected (Figure 163.10). The sinusoidal tissue at the base of the urethral stump is compressed by suturing the tunica albuginea that surrounds the CSP to the submucosa at the base of the urethral tube with interrupted no. 3–0 or 2–0 absorbable sutures pre-placed at equidistant intervals (Figure 163.11). The sinusoidal spaces
Figure 163.11 Scott’s technique of partial phallectomy. Sinusoidal tissue of the corpus cavernosum penis (CCP) is compressed by suturing the tunica albuginea surrounding the CCP to the tunica albuginea of the urethral groove.
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Figure 163.12 Scott’s technique of partial phallectomy. The tube of urethral mucosa can be folded over the end of the penile stump and sutured to the penile or preputial epithelium.
of the CCP are compressed by suturing the dorsal and lateral aspects of the tunica albuginea surrounding the CCP to the tunica albuginea of the urethral groove with interrupted no. 0 or 1 absorbable sutures (Figure 163.11). The urethral tube is divided into three equal-sized triangular segments, the apices of which point distally. These three triangles of urethra are interjoined with and apposed to three corresponding triangular segments of penile or preputial integument using simple interrupted no. 3–0 or #-0 absorbable or non-absorbable sutures. Sutures should include underlying tunica albuginea. Rather than dividing the urethral tube, as described by Scott, 9 the highly dilatable tube can be folded back over the end of the penile stump, where it is sutured to the penile or preputial epithelium. Sutures should include underlying tunica albuginea (Figure 163.12).
Partial phallectomy by en bloc resection with penile retroversion The free portion of the penis, the internal and external lamina of the prepuce, and the inguinal lymph nodes are excised when these structures are extensively affected by neoplasia.10 To prepare the horse for removal of these structures, the horse is anesthetized, positioned in dorsal recumbency, and the external genitalia and subischial region are prepared for surgery. After inserting a stallion catheter into the urethra, a fusiform incision is created around the preputial orifice. The incision begins 6 cm cranial to the orifice and ends 10 cm caudal to it. The incision is extended through the deep fascia to the abdominal tunic, and, if neoplasia has metastasized to the superficial lymph nodes, dissection is continued caudally and abaxially through this plane to expose the superficial inguinal lymph nodes at the superficial inguinal rings. Neoplastic lymph nodes are removed, the shaft of the penis is transected approximately 6–8 cm caudal to the fornix of the preputial cavity, and the prepuce and amputated portion of the penis are removed en bloc. As described by Markel et al.,10
Figure 163.13 Partial phallectomy by en bloc resection with penile retroversion. A subischial incision is created on the perineal raphe, and the stump of the penis is retroverted through this incision, so that the stump points caudally and extends a few centimeters beyond the subischial incision.
the penis is transected using Scott’s technique of partial phallectomy, so that a 4-cm tube of urethra protrudes from the penile stump.9 The procedure can be modified by using Williams’ or Vinsot’s technique of partial phallectomy to amputate the penis.11 A 6-cm-long subischial incision is created on the perineal raphe approximately 20 cm ventral to the anus on a horse of average size, and, using a hand introduced into the incision, penile fascia is bluntly separated until the stump of the penis can be grasped. The stump of the penis is retroverted through this incision, so that the stump points caudally and extends a few centimeters beyond the subischial incision (Figure 163.13). The tunica albuginea of the ventrally positioned CCP is sutured to the subcutaneous tissue of the ventral aspect of the subischial incision, and loose dorsal and lateral fascia of the penis is sutured to the subcutaneous tissue of the lateral and dorsal aspect of the subischial incision. If the penis was transected using Scott’s technique of partial phallectomy, the ventral border of the urethral tube is incised longitudinally to its base, and the margin of the urethra is sutured to the surrounding margin of the incised subischial skin. If the penis was transected using Vinsot’s or Williams’ technique of partial phallectomy, the margin of the urethra, which was sutured to surrounding tunica albuginea before the penis was retroflexed, is sutured to the surrounding margin of the incised subischial skin. The subcutaneous tissue and skin of the cranial incision are each closed separately after placing Penrose drains deeply within the incision. En bloc penile and preputial resection has been combined with subischial amputation of the penis,12
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but retroversion allows the flow of urine to be directed caudally rather than ventrally. The subischial incision must be positioned at the site where urine is most likely to be diverted caudally, rather than ventrally, and so this site should be identified and marked before the horse is anesthetized. Horses that have received the procedure lift their tails and squat like mares to urinate. The 180◦ flexure of the penis causes no impediment to the flow of urine.
Partial phallectomy by en bloc resection without penile retroversion This technique of partial phallectomy is similar to partial phallectomy by en bloc resection with penile retroversion, but, with this technique, the stump of the penis is retained at the site where it was transected.13 To prepare the horse for this technique of partial phallectomy, the horse is anesthetized, positioned in dorsal recumbency, and the external genitalia and inguinal regions are prepared for surgery. After inserting a stallion catheter into the urethra, a fusiform incision that begins at the umbilicus is created around the preputial orifice. The incision extends caudally on the midline to a point 10 cm caudal to the preputial orifice. If neoplasia has metastasized to the inguinal lymph nodes, the incision is extended caudally and deepened to expose the nodes, which are removed. The external lamina of the prepuce is bluntly dissected from the abdominal wall, ligating large vessels as they are encountered. After the shaft of the penis is exposed, dissection is directed caudally to expose at least 10 cm of the penile shaft. A tourniquet is applied around the shaft of the penis proximal to the site of amputation, and the dorsal arteries and veins of the penis are ligated. The penis is amputated just caudal to the fornix of the prepuce, using a technique similar to that described by Williams,8 and the tourniquet is removed. The stump of the penis is fixed to the midline of the body wall with heavy, absorbable, interrupted sutures, and subcutaneous tissue surrounding the penile shaft is apposed. The mucosa and submucosa of the new triangular urethral stoma are sutured to the cranial end of the incision, after first trimming the cranial end of the cutaneous incision so that it interjoins with the triangular urethral stoma. Skin cranial and caudal to the new urethral stoma is sutured. The technique of en bloc resection without retroversion allows removal of the prepuce, the regional lymph nodes, and a large portion of the penis, without altering the horse’s appearance as much as does the technique of en bloc resection with retroversion, and, because it requires less dissection of tissue, the sutured incision is less likely to undergo dehis-
cence.13 Care should be taken to avoid placing excessive tension on the penis when anchoring it to the body wall because excessive tension may cause severe post-operative pain. Care of the horse after partial phallectomy The horse should not be allowed heavy exercise for several weeks after undergoing partial phallectomy but should be walked daily to decrease swelling at the surgical site. Because partial phallectomy of stallions is usually performed to salvage the horse for purposes other than breeding, stallions should be castrated at least several weeks prior to phallectomy to decrease the risk of the horse developing an erection in the penile stump. Stallions and recently castrated geldings should be isolated from mares for 2–3 weeks. Complications of partial phallectomy Hemorrhage from the penile stump should be expected for at least a few days after partial phallectomy. Hemorrhage usually emanates from the CSP and is greatest at the end of urination. Hemorrhage results in minor dehiscence, which is usually inconsequential, but excessive hemorrhage may result in complete dehiscence of the stump and healing by second intention, which could lead to stricture of the urethral stoma (Figure 163.14). Other complications of partial phallectomy include signs of pain; infection at the site of surgery; excessive preputial edema, which may lead to paraphimosis; chronic cystitis; urine scalding; dysuria caused by urethral edema or stricture of the urethral stoma; neoplasia at the site of amputation; and neoplastic metastases to inguinal lymph nodes and internal organs.
Figure 163.14 A urethral stricture after partial phallectomy.
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Surgery of the Penis and Prepuce
Temporary perineal urethrostomy Indications Temporary urethrostomy at the ischial arch is indicated to treat horses affected with hemospermia or hematuria, to provide access to small cystic calculi, and to redirect the flow of urine for such conditions as urethral laceration or urethral urolithiasis. Surgical technique Temporary perineal urethrostomy is best performed with the horse standing, after being sedated, and administering epidural anesthesia or infiltrating local anesthetic solution into the tissue at the proposed site of incision. A 6- to 8-cm longitudinal incision, centered on the ischial arch, which is usually 2–3 cm below the anus, is created on the perineal raphe. The incision extends through the skin, the retractor penis and bulbospongiosus muscle, the CSP, and the urethral mucosa. The length of the perineal incision decreases as the incision extends deeper, so that the incision in the urethra is about 3 cm. Large branches of the external pudendal artery can be lacerated if the incision strays from the midline, and a periurethral hematoma can result in urinary obstruction (J. Schumacher and D.D. Varner, personal observation). A large-bore stallion catheter inserted into the urethra before surgery aids recognition of the urethra and helps to keep the incision on the midline. The urethrotomy is allowed to close by secondary intention and is usually healed within 2 weeks. Development of clinically apparent urethral stenosis after urethrotomy is rare.
Suturing a urethral rent Indications A rent in the urethra at the level of the ischial arch responsible for hematuria or hemospermia can be closed primarily through a perineal incision if temporary perineal urethrostomy or perineal incision into the CSP is ineffective in bringing about healing of the rent by second intention. Surgical technique In preparation for suturing a urethral rent, the insertion tube of an endoscope is inserted in the urethra
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so that the defect can be visualized. Two 20-gauge needles are placed through the skin in the perineal area and advanced into the urethral lumen so that the shafts of the needles emerge at the proximal and distal margins of the urethral defect. A perineal incision is created as if a perineal urethrostomy was to be performed, but the incision extends only through the tunica albuginea surrounding the CSP and not through the urethral mucosa. The rent is identified between the shafts of the needle, and its edges are apposed with a no. 3–0 absorbable suture using a simple continuous suture pattern. The tunica albuginea of the CSP, the bulbospongiosus muscle, subcutaneous fascia, and skin are each sutured separately. Only rents located at the level of the ischial arch can be sutured, but rents located in other areas of the urethra have not been reported.
References 1. Vaughan JT. Surgery of the prepuce and penis. In: Proceedings of the 18th Annual Convention of the American Association of Equine Practitioners, 1972, p. 19. 2. Guillaume A. Simplified surgical treatment of paralysis of penis in the horse. Vet J 1919;26:37–40. 3. Bolz W. The prophylaxis and therapy of prolapse and paralysis of the penis occurring in the horse after the administration of neuroleptics. Vet Med Rev Leverkusen 1970;4:255. 4. Kersjes AW, Nemeth F, Rutgers LJE. Atlas of Large Animal Surgery. Baltimore: Williams & Wilkins, 1985; p. 51. 5. Cosgrove MD, La Rocque MA. Shunt surgery for priapism. Urology 1974;4:1. 6. Pohl J, Polt B, Kleinhans G. Priapism: a three phase concept of management according to aetiology and progress. Br J Urol 1986;58:113. 7. Schumacher J, Varner DD, Crabill MR, Blanchard TL. The effect of a surgically created shunt between the corpus cavernosum penis and corpus spongiosum penis of stallions on erectile and ejaculatory function. Vet Surg 1999;28:21–4. 8. Williams WL. The Diseases of the Genital Organs of Domestic Animals. Worcester, MA: Ethel Williams Plimpton, 1943; pp. 201, 436. 9. Scott EA. A technique for amputation of the equine penis. J Am Vet Med Assoc 1976;168:1047. 10. Markel MD, Wheat JD, Jones K. Genital neoplasms treated by en bloc resection and penile retroversion in horses: 10 cases (1977–1986). J Am Vet Med Assoc 1988;192:396. 11. Archer DC, Edwards GB. En bloc resection of the penis in five geldings. Equine Vet Educ 2004;16:12–17. 12. Vaughan JT. Surgery of the penis and prepuce. In: Walker DF, Vaughan JT (eds) Bovine and Equine Urogenital Surgery. Philadelphia: Lea & Febiger, 1980; p. 137. 13. Doles J, Williams JW, Yarbrough TB. Penile amputation and sheath ablation in the horse. Vet Surg 2001;30:327–31.
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PART III
The Mare
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External Reproductive Anatomy John J. Dascanio
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Internal Reproductive Anatomy Robert A. Kainer
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Chapter 164
External Reproductive Anatomy John J. Dascanio
Evaluation of the external perineal conformation is often the first step in assessing the reproductive attributes of the mare. The perineum is defined as the area from the pubic symphysis to the coccyx and relates to the opening of the pelvic canal. It includes the caudal rectum, anus, vestibule, vulva, clitoris, and the superficial muscles, nerves, and skin. Anatomically, the perineum consists of muscles, bone, and adipose tissue and their associated nerve, lymph, and blood vessels. The ischium of the pelvis provides an anchor for attachment of the constrictor vulva muscles and the constrictor vestibuli muscles. Part of the external anal sphincter is continuous with the constrictor vulva muscle and helps to form the perineal body – the tissue that extends from the dorsal commissure of the vulva to the anus. The constrictor vestibuli muscle encircles the vulva, but is not complete dorsally. Motor and sensory innervation to the perineum and genitalia is supplied by the pudendal and caudal rectal nerves.1 The caudal cutaneous femoral nerve may also provide some sensory fibers to the perineal periphery. Anesthesia of this area may be achieved by epidural anesthesia or by infiltration of local anesthetics into the perineal body and/or labia.1–4 The external and internal pudendal and the urogenital vessels supply blood to the vulva and vestibule.5 There are three main anatomical barriers to contamination of the female reproductive tract: the vulva, the vestibulovaginal fold, and the cervix. The reproductive tissue from the vulvar labia to the remnant of the hymen/urethral orifice is considered the vestibule.6 From the remnant of the hymen/urethral orifice to the cervix is considered the vagina. For this chapter the dorsal transverse fold of tissue between the vestibule and vagina, a partial remnant of the hymen, is termed the vestibulovaginal fold. The
vulva and vestibule originate embryologically from the urogenital sinus. In contrast, there are reproductive tissues that are thought of as common bacterial reservoirs such as the clitoral fossa, clitoral sinuses, and the vagina. This chapter will concentrate on evaluation of the external genitalia of the mare with special attention to functional characteristics. Assessment of the reproductive tract first begins with examination of the vulvar labia. The vulva should be upright, without significant tilt to its vertical orientation (Figure 164.1). The vulvar labia meet dorsally at a sharp angle to form the dorsal commissure and ventrally at a rounded angle to form the ventral commissure. As the vulva departs from vertical, there is an increased risk of fecal contamination of the vestibule as there is increased residual contact of the vulvar labia to discharged fecal material.7 A measurement system was described for vulvar evaluation, termed the Caslick index.7 In this system the angulation of the vulva from vertical (angle of declination) is multiplied by the length of the vulva that is above the floor of the ischium. As an example, a vulvar angle of 30 degrees from vertical and a length of 4 cm from the dorsal commissure to the ischial floor would equal a Caslick index of 120. A value of < 100 would indicate a normal mare (Group 1), a value > 150 (Group 3) would indicate the need for a Caslick operation (vulvoplasty), and a value between 100 and 150 (Group 2) would be intermediate, with the mare possibly needing a Caslick operation depending on other concurrent issues. Mares with a Caslick index of < 150 had an increased pregnancy rate compared to those > 150. Mares in Group 1 had, on average, 2–3 cm between the dorsal commissure of the vulva and the pelvic floor. Group 2 mares had about 6–7 cm effective vulvar length above the floor, and Group 3 mares averaged 5–9 cm
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Dorsal Commissure Level of Pelvic Brim
Ventral Commissure Figure 164.1 Cross-section of a normal mare’s perineum. The vulvar lips are vertically oriented, with two-thirds of the opening below the level of the pelvic brim. Drawing by Terry Lawrence, Virginia-Maryland Regional College of Veterinary Medicine.
(Figure 164.2). The total vulvar opening is usually 12 to 15 cm in length.8 It has been suggested that no more than 4 cm of the vulvar opening be above the pelvic floor,6 and the vulva be no more than 10 degrees from horizontal.7 In reality, measurements are not often made and the clinician uses a combination of relative vulvar angulation, estimation of the length of the vulvar opening in relation to the pelvic floor, a history of reproductive problems, previous vulvar trauma, combined with their judgment to determine the need for a Caslick operation. In mares with a low body condition score or those with low body fat,5 such as young mares in work, there may be less intrapelvic deposition of fat causing the rectum
Dorsal Commissure Angle of Declination Level of Pelvic Brim Ventral Commissure
Figure 164.2 Cross-section of a mare with an abnormal perineal conformation. The vulvar lips are not vertically oriented, as indicated by the angle of declination. Less than two-thirds of the vulvar opening is below the level of the pelvic brim. Drawing by Terry Lawrence, Virginia-Maryland Regional College of Veterinary Medicine.
Figure 164.3 Vulvar angulation in an aged mare. Approximately half of the vulva is above the pelvic floor and the vulvar opening is angled, predisposing this mare to reproductive tract contamination. This mare would be a good candidate for vulvoplasty.
to be drawn slightly inward and the vulvar labia to have more angulation. By increasing bodyweight and intrapelvic fat, the angulation of the vulva may become more upright. In older mares with a decrease in ventral abdominal muscle tone and stretching of ligamentous attachments, the anus may acquire a more cranial orientation and the vulva become more angled (Figure 164.3). Mares that have a flat croup or an elevated tail head may also be predisposed to more vulvar angulation.5 Thoroughbred and American Saddlebred horses tend to have more vulvar angulation due to their flat croup, while Quarter Horse and Standardbred breeds tend to have a lower tail set and more slope to their croup resulting in a more vertical vulva.5 The lips of the vulva should meet evenly throughout from the dorsal to the ventral commissures, insuring a good mucosal seal. With previous trauma or past Caslick operations the lips may meet unevenly, potentially increasing the ability of air/fluids/solids to enter the caudal reproductive tract. This is due to irregularity in the mucosal membrane contact that normally forms a seal at the vulvar lips. Sometimes in cases of vulvar involvement with perineal tumors, such as melanomas, the vulva may not seal appropriately. The vulvar labia and associated musculature may become more toned under the influence of progesterone and less toned under estrogen.7 Hence, evaluation of a mare for vulvar angulation and tissue apposition should be made during estrus when tissue relaxation is potentially at maximum. With pregnancy, the vulva often appears to elongate and
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External Reproductive Anatomy
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C D A B
B E
Figure 164.4 The clitoris, everted in a mare in estrus. (A) median clitoral sinus; (B) lateral clitoral sinuses; (C) transverse frenular fold; (D) lateral recesses; (E) clitoral fossa.
become relaxed, especially in the last few weeks of gestation. Discharges may be evident on the ventral vulvar labia or the dependent perineum and hind legs of the mare. Occasionally, mares may have a grayish-white chalky substance on the ventral labia that is a normal residue from urination. The high calcium and mucus content of the urine is responsible for this residue. Discharges that are purulent, hemorrhagic, or otherwise discolored are abnormal. Most mares that are passively discharging liquid/mucus from their vulva will have a corresponding discharge in their tail, as the tail is down against the vulva. With urination, the tail is raised, preventing it from becoming soiled. The clitoris is located at the ventral commissure of the vulva (Figure 164.4). It is analogous to the penis of the stallion. The clitoris, or glans clitoridis, may be everted by placing a finger on either side of the ventral vulvar labia and slightly pushing inward. ‘Winking’, or eversion of the clitoris during estrus, is effected by contraction of the constrictor vulva muscle, the constrictor vestibuli muscle, and the retractor clitoridis muscle. The skin overlying the clitoris is wrinkled and there are three sinuses almost hidden in the dorsal aspect: a median sinus and two lateral sinuses. The median sinus is the deepest of the three sinuses. The lateral sinuses may be variable in depth and presence. The clitoris and associated structures may be a reservoir for bacteria or fungi, and they are routinely cultured as part of a surveillance program for contagious equine metritis. A clitoral sinusectomy may be performed in an attempt to limit this reservoir for bacteria.9 It has been suggested that only the median sinus is deep enough to establish the proper
anaerobic environment to harbor Taylorella equigenitalis, the causative agent for contagious equine metritis.10 All three clitoral sinuses may exude smegma with digital pressure. The glans clitoridis is bordered dorsally by the transverse frenular fold and ventrally by the clitoral fossa. The transverse frenular fold has a close attachment to the dorsal clitoris. There are lateral recesses on either side of the clitoris. The clitoris may become hypertrophied in mares given anabolic steroids,11 or become enlarged due to an intersex condition.12 In other cases intersex conditions may affect the urethral opening causing it to be located ventral to its normal location in a mare, midway between where the mare’s vulva and the stallion’s prepuce would be located.13 Diagnosis of an intersex condition may be made based on anatomical abnormalities, the presence of opposite sex gonads, and karyotyping. The opening to the vestibule should be below the floor of the pelvis, so that there is an upward orientation to the vestibule toward the vestibulovaginal fold/urethral orifice. This conformation will also serve as a barrier to contaminants entering the cranial reproductive tract. The pelvic floor may be determined by pushing a finger on either side of the vulva to palpate the deeper tissues. The bony edge of the ischium is easily determined by this method. An estimate may then be made as to the amount of the vulva below the pelvic floor in relationship to the operator’s finger. Two-thirds of the vulvar opening should be below the ischium. Others view the normal condition as being when an estimated 70% of the vulva is below the floor of the pelvis.14 Mares with a direct horizontal communication between the vulva, vestibule, and
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Figure 164.5 Separation of the vulvar labia to expose the vestibulovaginal fold. The picture on the left demonstrates a noncompetent vestibulovaginal fold with visualization to the vagina (‘windsucker’). The picture on the right demonstrates a normal vestibulovaginal fold.
vagina are more likely to have reproductive tract contamination issues. The vestibule possesses glands on its ventrolateral walls. The vulvar lips may be separated, exposing the vestibule and vestibulovaginal fold (windsucker test). There should be no inrush of air into the vagina when the lips are parted. If this occurs it may indicate a poor seal at the level of the vestibulovaginal fold or a more horizontal communication between the vestibule and vagina. With some mares, the seal is disrupted to the point of being able almost to visualize the cervix through the parted vulvar lips (Figure 164.5). Pneumovagina may lead to pneumouterus and an increase in inflammatory cells within the uterus, especially eosinophils.15 Mares who have significant pneumovagina/pneumouterus may exhibit air discharge from the vulva during urination, defecation, or exercise.5 During estrus, there may be decreased integrity of the vestibulovaginal fold with relaxation of supporting structures. Thus, assessments made during diestrus versus estrus may vary. The demarcation between the mucosa of the vestibule and the labial skin can be seen often as a slightly raised line. This is an important landmark, as the mucosa is incised when performing a Caslick operation (vulvoplasty).6 Vulvoplasty (Caslick operation) is a surgical procedure whereby the dorsal vulvar lips are co-joined.16 Most often a small strip of vestibular mucosa is removed just inside the vulvar skin and the two sides are sutured together. An alternative procedure is where no strip of mucosa is removed and skin sta-
ples are placed in the vulvar lips effectively to join the two sides together. This is performed as a temporary vulvoplasty during times where entry into the reproductive tract is needed through a larger opening; consequently, staples may be removed and replaced as needed. Once treatment or breeding has been completed, a permanent vulvoplasty may be performed. Surgical procedures have also been described to correct anatomical abnormalities associated with vulvar angulation.6, 17, 18 Pouret’s operation involves a surgical incision in the perineum between the vulva and anus, with reconstruction to reorient the vulva into a horizontal orientation, while the Gadd technique involves incision of the vestibule along with a Caslick operation to reorient the vulva. The Pouret procedure will not decrease the effective vulvar opening, while the Gadd technique will. Trauma may occur to the vulva at the time of foaling or breeding, or due to unknown factors (Figure 164.6). Depending on the degree of tissue damage, contamination to any wounds and any subsequent swelling/edema, possible surgical correction may or may not be instituted immediately. Often a more appropriate closure is done once the wound has been debrided of any dead tissue and edema has waned. This may take 10–14 days, and vulvar repair would potentially coincide with vulvoplasty. Mares may become constipated, avoiding defecation due to pain associated with the caudal reproductive tract. Irregularities of pigmentation in the vulvar labia may occur secondary to infectious agents such as equine herpes virus 3 infection.19 Nodules or masses
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External Reproductive Anatomy
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5. 6.
7.
8.
9. 10.
11.
12. Figure 164.6 Vestibule and vulvar trauma post-foaling. The skin overlying the vulvar labia is abraded from tail movement over the swollen tissue.
13.
14.
may occur that may affect the vulva, including melanoma, squamous cell carcinoma, and papilloma.5, 20 The ideal mare would have vertically oriented normal vulvar lips, a good mucosal seal, two-thirds of the vulvar opening below the ischium, and no air aspiration into the vagina with separation of the vulvar lips.
15.
16.
17.
References 1. Schumacher J, Bratton GR, Williams JW. Pudendal and caudal rectal nerve blocks in the horse – An anesthetic procedure for reproductive surgery. Theriogenology 1985;24:457–464. 2. LeBlanc PH, Caron JP, Patterson JS, Brown M, Matta MA. Epidural injection of xylazine for perineal analgesia in horses. J Am Vet Med Assoc 1988;193:1405–8. 3. Grubb TL, Riebold TW, Huber MJ. Comparison of lidocaine, xylazine, and xylazine/lidocaine for caudal
18. 19. 20.
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epidural analgesia in horses. J Am Vet Med Assoc 1992;201: 1187–90. DeRossi R, Sampaio BF, Varela JV, Junqueira AL. Perineal analgesia and hemodynamic effects of the epidural administration of meperidine or hyperbaric bupivacaine in conscious horses. Can Vet J 2004;45:42–7. Roberts SJ. Veterinary Obstetrics and Genital Diseases, 3rd edn. Woodstock: Stephen J. Roberts, 1986. Trotter GW, McKinnon AO. Surgery for abnormal vulvar and perineal conformation in the mare. Vet Clin N Am Equine Pract 1988;4:389–405. Pascoe RR. Observations on the length and angle of declination of the vulva and its relation to fertility in the mare. J Reprod Fertil Suppl 1979;27:299–305. Easley J. External perineal conformation. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 20–4. Digby NJ, Ricketts SW. A method for clitoral sinusectomy in mares. In Practice 1982;4:145–7. McAllister RA, Sack WO. Identification of anatomic features of the equine clitoris as potential growth sites for Taylorella equigenitalis. J Am Vet Med Assoc 1990;196: 1965–6. Maher JM, Squires EL, Voss JL, Shideler RK. Effect of anabolic steroids on reproductive function of young mares. J Am Vet Med Assoc 1983;183:519–24. Moreno-Millan M, Delgado Bermejo JV, Lopez Castillo G. An intersex horse with X chromosome trisomy. Vet Rec 1989;124:169–70. Basrur PK, Kanagawa H, Gilman JP. An equine intersex with unilateral gonadal agenesis. Can J Comp Med 1969;33:297–306. Evans TJ, Constantinescu GM, Ganjam VK. Clinical reproductive anatomy and physiology of the mare. In: Youngquist RS, Threlfall WR (eds) Current Therapy in Large Animal Theriogenology 2. Philadelphia: Elsevier, 2006; pp. 47–67. Slusher SH, Freeman KP, Roszel JF. Eosinophils in equine uterine cytology and histology specimens. J Am Vet Med Assoc 1984;184:665–70. Caslick EA. The vulva and vulvovaginal orifice and its relation to genital health of the Thoroughbred mare. Cornell Vet 1937;27:178–87. Gadd JD. The relationship of bacterial cultures, microscopic smear examination and medical treatment to surgical correction of barren mares. Proc Am Assoc Eq Pract 1975;21: 362. Pouret EJ. Surgical technique for the correction of pneumoand arovagina. Equine Vet J 1982;14:249–50. Pascoe RR, Spradbrow PB, Bagust TJ. Equine coital exanthema. Aust Vet J 1968;44:485. Junge RE, Sundberg JP, Lancaster WD. Papillomas and squamous cell carcinomas of horses. J Am Vet Med Assoc 1984;185:656–9.
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Chapter 165
Internal Reproductive Anatomy Robert A. Kainer
Neuroendocrine contributors Neural and endocrine functions of three regions of the diencephalon – pineal gland, hypothalami, and pituitary gland – contribute to the development and cyclic physiology of the reproductive system of the mare. Interactions with other parts of the nervous system and with hormones produced by organs of the reproductive tract influence sexual behavior, pregnancy, parturition, and lactation. Pineal gland Unlike its human counterpart, which is shaped somewhat like a pinecone (hence, pineal), the equine pineal gland is a dark brownish red, ovoid projection from the epithalamus occupying a median position between the rostral colliculi and thalami. Its short stalk contains the pineal recess of the third ventricle, and a suprapineal recess covers the gland dorsally (Figure 165.1).1 Septa extending into the pineal gland from a capsule of pia mater separate masses of epithelial cells derived from ependyma. Melatonin secreted by these cells is considered to exert an antigonadotropic effect in the control of reproductive photoperiodicity via action on the hypothalamic – pituitary – gonadal axis.2 Light inhibits the organ’s activity, thus stimulating reproduction. In a report on pony mares, pineal activity was highest during late fall and winter and lowest during spring, summer, and early fall, and the percentage of ovulatory mares was inversely related to pineal hydroxyindole-O-methyltransferase activity.3 Blood supply to the pineal gland is from branches of the deep cerebral arteries that terminate in the formation of a rete pinealis. Venous blood drains into the ventral longitudinal sinuses.4 Nerve fibers in the stroma of the gland are probably of sympathetic ori-
gin. It is believed that impulses from the retina travel via the rostral accessory optic tract to the hypothalamic tegmental tract, then to the thoracic spinal cord, and up to the cranial cervical ganglion.4 Nerve fibers from the latter follow arteries to end on pineal epithelial cells.4
Hypothalami Contralateral hypothalami face each other across the third ventricle. They join basally to form the tuber cinereum, and caudal to this region, the fused mainnilliary bodies (Figure 165.1). The tuber cinereum is continuous with the infundibulum of the hypophysis. Nuclei of nerve cell bodies occur in each hypothalamus. Supraoptic, paraventricular, preoptic, and rostra! hypothalamic nuclei contain cell bodies of neurosecretory cells. Large, magnocellular neurosecretory cells in supraoptic nuclei (dorsal to the optic chiasm) are the source of vasopressin (antidiuretic hormone), whereas magnocellular cells in the paraventricular nuclei (in lateral walls of the third ventricle) produce both vasopressin and oxytocin. Processes of magnocellular secretory cells project to the neurohypophysis, terminating in its neural lobe. Smaller, parvicellular neurosecretory cell bodies in pre-optic nuclei (rostral to the optic chiasm) produce gonadotropin-releasing hormone; parvicellular neurosecretory cell bodies in rostral hypothalamic nuclei (caudal to preoptic nuclei) produce thyrotropinreleasing hormone.5 These and other releasing hormones are carried by processes of parvicellular neurosecretory cells to the median eminence region of the hypophysial infundibulum, where they end in association with dorsal and ventral capillary plexuses.6
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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(b) Figure 165.1 (a) Neuroendocrine contributors of the diencephalon to the reproductive system. (b) Enlargement of a median section through the diencephalon. 1, Pineal gland; 2, hypothalamus; 3, hypophysis cerebri; Ill, third ventricle; 4, suprapineal recess of III; 5, pineal recess of III; 6, tuber cinereum; 7, infundibular recess of III; 8, median eminence; 9, infundibular stalk; 10, neural lobe; (8, 9, 10 = neurohypophysis); 11, pars tuberalis; 12, pars distalis; 13, pars intermedia; (11, 12, 13 = adenohypophysis).
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Hypothalamic blood supply is derived from rostrodorsal hypophysial (infundibular) arteries and branches from caudal communicating and internal carotid arteries.7 Venous blood drains into cavernous and intercavernous sinuses. Central nervous system connections to the hypothalami include input from the limbic system, cerebral cortex, thalami, and reticular-activating system. The hypothalami have autonomic connection with the spinal cord and direct input from the accessory optic tract. Hypophysis (pituitary gland) Most of the hypophysis lies within the hypophysial fossa of the Turkish saddle (sella turcica) of the basisphenoid bone. In the horse, the long axis of this dorsoventrally flattened, oval mass is almost horizontal to the hypothalamic base (Figure 165.1). The hypophysis is held in place by the sellar diaphragm of dura mater. A foramen in the diaphragm admits passage of the hypophysial stalk. Two major parts constitute the hypophysis – the neurohypophysis, derived from the developing diencephaion, and the adenohypophysis, derived from a pouch of oral ectoderm. The infundibulum of the neurohypophysis descends from the hypothalamic tuber cinereum. An outpocketing of the third ventricle, the infundibular recess, extends into the proximal part of the infundibular stalk. The well-vascularized median eminence region of the infundibulum is continued distad by the infundibular stalk, which terminates in an expanded neural lobe (human ‘posterior pituitary’). The bulk of the adenohypophysis, the pars distalis (human ‘anterior pituitary’), and the contiguous pars intermedia enclose nearly all of the neural lobe with only a small area of the caudal surface of the latter exposed. The pars tuberalis of the adenohypophysis ascends to enclose the infundibular stalk and the median eminence. Together the infundibular stalk and the tuberal part constitute the hypophysial stalk. Axons and telodendria of parvicellular neurosecretory cells, vessels of the primary capillary plexus, and origins of hypophysial portal veins are present in the median eminence and in the proximal part of the infundibular stalk. Axons of magnocellular neurosecretory cells pass through the median eminence and infundibular stalk on their way to terminate in the neural lobe. Supportive neuroglial cells are present in the nellrohypophysis. The tuberal part of the adenohypophysis consists of follicles or cords of weakly basophilic cells and numerous hypophysial portal veins. Clumps of basophilic cells in the pars intermedia are the source of melanocyte-stimulating hormone.
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Figure 165.2 Blood supply to and venous drainage from the hypophysis. Dorsal view of hypophysis and optic chiasm. 1, Rostroventral hypophysial arteries; 2a, internal carotid artery emerging from cavernous sinus; 2b, internal carotid artery within cavernous sinus; 3, caudal communicating branch; 4, caudal hypophysial arteries; 5, intercarotid artery within intercavernous sinus; 6, hypophysial veins; 7, ventral petrosal sinus; 8a, cavernous sinus; 8b, cavernous sinus exiting through orbital fissure. (Adapted in part from Vitums.6 )
The pars distalis consists of clumps and cords of granular acidophils and basophils and agranular chromophobes. On proper stimulation, specific cell types secrete somatotropin, prolactin, adrenocorticotropic hormone, follicle-stimulating hormone, luteinizing hormone, thyrotropin, and lipoproteins. Sinusoidal capillaries and smaller efferent capillaries are present among the groups of cells.
Vascular relationships The internal carotid arteries give origin to rostroventral infundibular (hypophysial) arteries, which supply the ventral region of the primary capillary plexus in the median eminence (Figure 165.2). The caudal communicating branches detach rostrodorsal infundibular arteries, which supply the dorsal region of the plexus. Superficial and deep, looped capillaries are linked together in the median eminence and extend caudad as far as the end of the infundibular recess. Ventral and dorsal groups of hypophysial
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Figure 165.3 Hypothalamic–hypophysial relationships. 1, Paraventricular nucleus; 2, pre-optic nucleus; 3, rostral hypothalamic nucleus; 4, supraoptic nucleus; 5, processes of magnocellular neurosecretory cells; 6, processes of parvicellular neurosecretory cells; 7, branch of rostrodorsal hypophysial (infundibular) artery; 8, branch of rostroventral hypophysial (infundibular) artery; 9, primary capillary plexus within median eminence; 10, dorsal group of hypophysial portal vessels; 11, ventral group of hypophysial portal vessels; 12, sinusoidal capillary network within pars distalis; 13, branches of caudal hypophysial arteries; 14, capillaries of neural lobe; 15, hypophysial veins. (Adapted in part from Vitums6 and Krieger.5 )
portal vessels, descending from the primary capillary plexus through the tuberal part, constitute the sole blood supply to the distal part of the adenohypophysis. There the portal vessels terminate in a sinusoidal capillary network (Figure 165.3). It has been suggested, but not substantiated, that ventral and dorsal groups of portal vessels are functionally related to specific zones of the median eminence and to certain regions of the distal part of the adenohypophysis.6 Right and left caudal infundibular arteries arise from the caudal intercarotid artery (or from internal carotid arteries) within the intercavernous sinus. They pierce the wall of the sinus and run in the dural sheath, sending branches to the dura mater and capsule of the hypophysis, and terminate in capillaries of the neural lobe and distal infundibular stalk.6 Venous drainage is mainly from the caudal aspect of the hypophysis to the right and left cavernous sinuses. From each sinus, two courses exist: (i) a caudoventral course to the ventral petrosal sinus and thence to the foramen lacerum and (ii) a rostrolateral course to emissary veins in the orbital fissure – the ophthalmic vein and deep facial vein.8 Continuity with the deep facial vein provides a route for
catheterization to obtain samples of blood coming from the hypophysis. A catheter may be passed into the facial vein, then through the deep facial vein into the cavernous sinus.
Hypothalamic – hypophysial connections In the median eminence, neurohemal organs – junctions between telodendria at the ends of processes of parvicellular neurosecretory cells descending from nuclei in the hypothalamus and capillaries of the primary plexus – are the sites of transfer of releasing and inhibitory hormones to the blood. Hypophysial portal veins coursing distad from the primary capillary plexus carry blood through the tuberal part to the distal part of the adenohypophysis where the veins terminate by branching into the sinusoidal (secondary) capillary plexus (Figure 165.3). Releasing and inhibitory hormones leave the capillaries and contact target cells in the distal part of the adenohypophysis. Hormones produced by the target cells enter efferent capillaries that carry blood to veins draining to the cavernous sinuses. Axons of magnocellular neurosecretory cells descend from hypothalamic nuclei through the median
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eminence and infundibular stalk to the neural lobe of the neurohypophysis (Figure 165.3). Secretory granules (Herring bodies), consisting of prohormones, enzymes, and carrier molecules (neurophysins), migrate distad through the axons to neurohemal organs where hormones (oxytocin and antidiuretic hormone) are transferred through capillary walls into the blood.5 Veins drain to the cavernous sinuses.
Reproductive tract of the mare General relationships The mare’s reproductive tract – ovaries, uterine tubes, uterus, vagina, and vulva – extends from the approximate position of the ovaries, ventrally to the fourth or fifth lumbar vertebra and caudally through the pelvic cavity to the vulva (Figures 165.4 and 165.5). Contralateral ovaries, uterine tubes, and uterine horns lie on either side of the descending colon and its mesocolon; the uterine body and cervix are ventral to the terminal part of the descending colon and the rectum. The bladder is located ventrally to the uterine body and cervix with the urethra coursing caudad ventral to the vagina and emptying into the vestibule at the urethral orifice. The vagina is related bilaterally to the sacrotuberous ligaments, dorsally to the rectum, and ventrally to the urethra. The vestibule of the vagina, according to the Nomina Anatomica Veterinaria9 is related dorsally to the anus and extends caudoventrad over the ischiatic arch to the vulva. The vestibule and vulva are related dorsally to the muscles and fascia of the perineal body. Laterally the vestibule and vulva are covered by the perineal fascia in apposition with the semimembranosus muscles.10 Variations in distension and the movement of the intestines and the bladder as well as pregnancy influence the positions of the internal genitalia. In the non-pregnant mare, ovaries, uterine tubes, and uterine horns may be in contact with the dorsal abdominal wall or may be located among intestinal coils. Distension of the bladder or descending colon can displace the uterine body to one side. It has been suggested that intestinal temperature and motility may affect the function of the reproductive organs in contact with the intestines.11 When the intestinal mass is removed, the internal genitalia droop into the emptied abdominal cavity, stretching the broad ligament on each side (Figure 165.5). Broad ligaments are connecting double peritoneal sheets extending along lateral sublumbar and lateral pelvic walls from the third or fourth lumbar vertebra to the level of the fourth sacral vertebra. They become continuous with the serous tunics enclosing the ovaries, uterine tubes, and uterus. The
portion of the broad ligament suspending the ovary, the mesovarium, detaches the mesosalpinx from its lateral surface. The mesosalpinx suspends the uterine tube (oviduct or salpinx). Most of the broad ligament is mesometrium, the connecting peritoneum extending to and continuous with the perimetrium, the serous tunic enclosing the uterus. In their abnormally suspended position, uterine horns are slung in a V-shaped configuration with the mesometrial attachment dorsally. Each stretched broad ligament provides for visualization of the contained ovarian vessels near its cranial edge, uterine vessels in its middle, and the round ligament of the uterus, on its lateral aspect. The round ligament of the uterus, the homologue of the gubernaculum testis, extends from its rounded, blunt end to the internal inguinal ring. Also contained between the two layers of the broad ligament are lymphatic vessels, loose collagenous and adipose tissues, and sheets of smooth muscle continuous with the outer muscular layer of the uterine tube and uterus. During pregnancy, thickening of the broad ligaments restraining the descending gravid uterus is a result of enlargement of blood and lymphatic vessels and an increase in the smooth muscle mass. Cystic remnants of mesonephric tubules and ducts ¨ (epoophoron and paraoophoron) occur commonly in the mesovarium and mesosalpinx. They tend to regress with age.12 The long broad ligaments of the mare permit exteriorization of the ovaries, uterine tubes, and uterine horns through a midventral celiotomy.11 An ovary may be exteriorized through an incision in the paralumbar fossa.13 Caudal to where the broad ligaments become continuous with the lateral ligaments of the bladder, two dorsoventrally flattened peritoneal pouches project into the pelvic cavity. Peritoneum reflects from the rectum onto the vagina, forming the rectogenital pouch, and from the bladder onto the vagina, forming the vesicogenital pouch that extends farther caudad. The extent of the vagina covered varies inversely with distension of the rectum and bladder.14 When the latter are nearly empty, enough peritoneum (8– 10 cm) covers the vagina so that the rectogenital pouch can be entered surgically through the dorsolateral wall of the vaginal fornix in the procedure of ovariectomy via colpotomy.’15 Farther caudally, a rectovaginal septum separates the rectum and vagina. Ovaries As described above, the position of the ovaries is variable largely because of the extensive mesovarium that permits the ovary a wide range of passive movement. An ovary may lie on top of the intestinal mass in contact with the dorsal abdominal wall or it may be
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(b) Figure 165.4 (a) Left lateral view of mare’s reproductive tract with intestines in situ. Left aterocaudal abdominal and lateral pelvic walls removed. 1, Left ovary; 2, left uterine horn; 3, uterine body; 4, bladder; 5, ampulla of rectum; 6, vagina; 7, urethra; 8, pelvic brim; 9, descending colon; 10, left ventral colon; 11, external anal sphincter muscle; 12, vestibular constrictor muscle; 13, clitoral retractor muscle; 14, vulvar constrictor muscle. (b) Left lateral view of mare’s reproductive tract with intestinal mass removed. Rectum, uterine cervix, bladder, urethra, vaginal vestibule, and clitoris in median section. 1, Ovarian artery; 2, uterine artery; 3, vaginal artery; 4, round ligament of uterus; 5, left ovary; 6, left uterine tube; 7, left uterine horn; 8, uterine body; 9, uterine cervix; 10, bladder; 11, ampulla of rectum; 12, vagina proper; 13, urethra; 14, pelvic brim; 15, vaginal vestibule; 16, corpus cavernosum clitoridis.
situated among the intestinal coils roughly on a plane through the fourth or fifth lumbar vertebra. The left ovary is caudal to the right, but it is closer to the ipsilateral kidney. The distance from the ovaries to tips of the uterine horns also varies. During pregnancy, the
ovaries are pulled cranioventrad as well as mediad, an important position to recall when palpating per rectum. The large ovaries (averaging 7–8 cm along the long axis) of the mare are reniform with a convex dorsal
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Figure 165.5 Frontolateral view of a mare’s reproductive tract with intestinal mass removed. 1, Mesocolon; 2, descending colon; 3, ampulla of rectum; 4, mesovarium; 5, ovarian artery; 6, round ligament of uterus; 7, left ovary; 8, left uterine tube; 9, right ovary; 10, proper ligament of ovary; 11, uterine artery; 12, mesometrium; 13, right uterine horn; 14, uterine body; 15, left lateral ligament of bladder; 16, bladder; 17, rectogenital pouch; 18, vesicogenital pouch.
Figure 165.6 Left ovary, uterine tube, ovarian bursa, and opened tip of left uterine horn. 1, Ovarian branch of left ovarian artery; 2, uterine branch of left ovarian artery; 3, adrenocortical nodules; 4, ampulla of left uterine tube; 5, infundibuurn of uterine tube covering ovulation fossa; 6, isthmus of uterine tube; 7, rnesosalpinx; 8, ovarian bursa; 9, left proper ligament of ovary; 10, uterine ostium of uterine tube (slightly exaggerated).
border attached to the mesovarium and a sharply concave ventral free border indented by an ovulation fossa (ovarian fossa9 ). Medial and lateral surfaces are covered by visceral peritoneum continuous with the mesovarium and contain prominent parallel branches from the ovarian vessels. Yellow, grape-like clusters of adrenocortical nodules (1–2 mm in diameter) may be present in loose connective tissue on the surface of the ovary.16 A variable amount of adipose tissue also occurs in the loosely adherent serosa. From the caudal pole of the ovary, the proper ligament of the ovary – a band of smooth muscle within the broad ligament – extends to the tip of the uterine horn. The cranial pole of the ovary is attached, in part, to the uterine tube’s infundibulum, which overlays the ovulation fossa (Figure 165.6). Polygonal cells of the superficial epithelium (‘germinal epithelium’) lining the ovulation fossa impart a darker color than the contiguous mesothelial cells covering the rest of the organ.1, 14 The histological structure of the equine ovary is unique among domestic mammals in that it has a peripheral collagenous connective tissue vascular
zone around a central parenchymatous zone, containing developing and atretic ovarian follicles, corpora lutea, and corpora albicantia. The parenchymatous zone surfaces at the ovulation fossa, and during folliculogenesis, maturing follicles move toward the fossa, enlarging to a diameter of 4.5–6 cm. Mature ovarian follicles become less turgid immediately before ovulation. A corpus hemorrhagicum commonly forms following ovulation. The developing reddish corpus luteum does not bulge from the ovary, but it may project slightly into the ovulation fossa. Later, the corpus luteum becomes yellow. The primary corpus luteum of pregnancy is supplemented by secondary corpora lutea that begin to form around day 40 of gestation, developing from follicles that continue to grow rapidly during the first couple of months of gestation. Luteinization takes place in follicles that have ovulated and in anovulatory follicles.11 Secondary corpora lutea are spherical, irregular, or gourd-like with a tract leading to the ovulation fossa. Many corpora lutea have a central cavity.17 Corpora lutea continue to function until the sixth month of gestation, and then regression occurs.
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Internal Reproductive Anatomy The mare’s large ovaries are eclipsed in size by the developing fetal ovaries at midgestation. Fetal ovaries later regress, and, by the time of parturition, they are one-tenth their greatest fetal size.1 Their shape at birth is oval, with superficial (‘germinal’) epithelium covering the entire free surface. Observed as early as 5 months of age,18 indentation of the free border to form the ovulation fossa continues until puberty when the growth of the organ is accelerated. The ovary is larger in young mares, and it tends to become more fibrous in older mares.
Uterine tubes (oviducts or salpinges) Each uterine tube consists of an expansive infundibulum covering the ovary’s ovulation fossa, a highly tortuous ampulla about 6 mm in diameter, and a less tortuous isthmus half the diameter of the ampulla (Figure 165.6). The isthmus terminates at a small uterine ostium on a papilla within the end of the uterine horn. Based on measurements made with the loops effaced and free of the suspending mesosalpinx, equine uterine tubes are from 20 to 30 cm lon with the ampulla constituting about half the length.1, 14 Irregular fimbriae are present along the margin of the funnel-shaped infundibulum. Some are attached to the cranial pole of the ovary, allowing the rest of the infundibulum to spread over the ventral aspect and cover the ovulation fossa. At the edges of the fimbriae, their serosal covering is contiguous to the mucous membrane. The latter is highly folded, especially in the ampulla where secondary and tertiary ridges branch from lon, itudinal folds. The lining of simple columnar epithelium is intermittently ciliated. Ciliogenesis and ciliary motion (toward the uterus) reflect stages of the sexual cycle. A thin, wellvascularized lamina propria supports the epithelium. Inner, circularly disposed smooth muscle fibers are covered by outer, longitudinally arranged fibers that continue into the mesosalpinx. The abdominal ostium in the center of the infundibulum is about 6 mm in diameter; the uterine ostium of the tube, 2–3 mm in diameter.14 The inner circular muscle increases to form a sphincter at the tubo-uterine junction. Unfertilized ova are retained for a considerable duration in the uterine tubes (up to several months), parthogenetic cleavage occurring in some of them.19 The ovarian bursa of the mare is a peritoneal pouch extending from the ovulation fossa caudad to the cranial aspect of the uterine horn. Laterally it is bounded by the uterine tube and mesosalpinx. A fold of broad ligament containing the proper ligament of the ovary forms the medial wall of the ovarian bursa.
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Uterus Two uterine horns, diverging sharply from a small intercornual ligament, are located entirely within the abdominal cavity, lying on or intermingling with intestinal coils. The body of the uterus becomes continuous caudally with the uterine cervix in the cranial part of the pelvic cavity. Length of the uterine horns ranges from 20–25 cm. They are asymmetrical in parous mares. The diameter of the horns increases from their blunt tips to their junction with the body. The body averages 18–20 cm in length; the cervix, 5–7 cm.14 Mesometrium attaches to the dorsal borders of the horns and follows to the lateral aspects of the body and cervix. In situ before manipulation, the walls of the horns and body of the uterus are flaccid and intestine-like. Handling causes the uterus to contract rapidly, the decrease in length and increase in diameter imparting the sausage shape of the horns observed after removal of the uterus from the abdominal cavity.11 Internally, the uterine ostium of a uterine tube opens on a small papilla in the endometrium in the tip of each horn. The sphincter of circular muscle at the tubo-uterine junction serves as a valve, preventing reflux of uterine contents through the uterine ostium. A short median septum is present where the uterine horns join the body. The lumen of the uterus is practically obliterated by its flaccid walls and 12–15 large longitudinal folds of reddish brown endometrium with collagenous connective tissue cores. Only capillary spaces occur between the endometrial folds.20 The epithelial layer of the endometrium of the uterine body and horns is simple columnar epithelium (pseudostratified during estrus) with cuboidal to tall cells, depending on stage of the estrous cycle. Fewer than half of the cells are ciliated. Branched tubular glands extend from the surface through a compact layer of connective tissue cells and reticular fibers into a fibrous and vascularized spongy layer of lamina propria.20 The myometrium of the uterine wall consists of an inner circular layer of smooth muscle, a middle vascular layer, and an outer longitudinal layer of smooth muscle. The last two layers are coextensive with the vasculature and musculature of the mesometrium. Enclosing visceral peritoneum, the perimetrium, is continuous with the two peritoneal sheets of the mesometrium (Figure 165.7). The constricted, thick-walled cervix is traversed by the cervical canal, which extends from the internal uterine orifice and opens into the body to the external uterine orifice in the protruding vaginal portion of the cervix (Figure 165.8). The wall contains an increased amount of coilgenous connective tissue and a sphincter of smooth muscle derived from the inner
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Figure 165.7 Cross-section of uterine horn. 1, Mesometrium; 2, perimetrium; 3, myometrium; 4, endometrium; 5, collagenous connective tissue core; 6, surface epithelium; 7, compact layer of lamina propria; 8, spongy layer of lamina propria; 9, endometrial glands; 10, internal (circular) layer of myometrium; 11, middle (vascular) layer of myometrium; 12, external (longitudinal) layer of myometrium.
layer of myometrium. Longitudinal mucosal folds in the cervical canal are continuous with those in the endometrium of the body and tend to bulge caudad at the external uterine orifice. A fold may continue onto the vaginal floor as a frenulum.11 The endometrium of the cervical canal is grossly paler. Tubular glands do not project from the epithelial layer, which consists of both ciliated and mucigenous cells.
Cyclic uterine anatomy
Uterine anatomy during estrus During estrus, the uterine wall thickens, muscular tone increases, vascularity becomes greater, and proliferated endometrial glands are active. Vaginoscopic examinations of the vaginal portion of the cervix reveal functional anatomic variations that correlate with ovarian activity and behavioral changes during the estrous cycle.21 In pro-estrus, the cervix and its folds begin to enlarge and the mucous membrane becomes unevenly congested. During estrus, the cervix is completely relaxed and droops ventrad; the folds protrude farther from the enlarging external uterine orifice, which will eventually admit three or four fingers (Figure 165.9a). Mucosal congestion is even and deep, and cervical secretions are abundant.21
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11 12 13 Figure 165.8 Dorsal view of opened vagina and vulva. 1, Vaginal artery; 2, vaginal portion of cervix; 3, vaginal fornix; 4, ventral vaginal wall; 5, transverse fold; 6, urethral orifice; 7, vestibular constrictor muscle; 8, orifices of vestibular glands; 9, locus of submucosal vestibular bulb; 10, vulvar constrictor muscle; 11, vulvar labium; 12, transverse frenular fold (clitoral prepuce); 13, clitoral glans.
Manipulation of the cervix tends to restore some of its turgidity.
Uterine anatomy during diestrus In diestrus, the thickness of the uterine wall decreases and muscular tone and endometrial glandular activity subside. The cervical canal constricts as the cervix becomes firmer and protrudes caudad (Figure 165.9b). It is occluded by a plug of mucus. Mucosal folds protrude less and are no longer congested.16
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Figure 165.9 Vaginal portion of uterine cervix. (a) During estrus and (b) during diestrus. 1, Vaginoscopic view; 2, median section.
Uterine anatomy during pregnancy
Vagina
During pregnancy, the developing embryo, and then fetus, causes enlargement of the horn, then the body, and finally, the contralateral horn. The mesometrium, enlarging in response to distension of vessels and increased muscular mass, exerts tension on the enlarging horns, causing them to flex somewhat. As the uterus descends to the abdominal floor, the myometrium becomes thinner, except in the then-firm cervix, which is pulled craniad out of the pelvic cavity. The cervical canal is filled with a thick mucous plug, and the external uterine orifice is contracted to a slit. The arrangement of collagenous, elastic, and smooth muscle fibers in the wall permit extreme dilation of the cervix, accommodating passage of the foal during parturition. It has been suggested that collagenase is involved in the process of labor.22 The endometrium becomes thicker and more vascular during the formation of the yolk-sac placenta and then the diffuse, microcotyledonous, epitheliochorial placenta of the mare is formed. A ring of endometrial cups develops, starting about day 37 of gestation. Slow regression begins around day 70 and is complete by 130 days.23 Endometrial cups are formed by hypertrophy of endometrial glands and the subsequent invasion of the endometrium by chorionic epithelial cells.24 These cells develop into cup cells, which produce pregnant mare serum gonadotropin (equine chorionic gonadotropin).
The vaginal fornix is an anular recess formed by the junction of the cranial vaginal walls with the caudally projecting vaginal portion of the cervix. The vagina, including its vestibule,9 extends caudad from the fornix to the labia of the vulva (Figure 165.8). Some authors include the vestibule as part of the vulva in keeping with the designation of the shallow human vestibule and the embryonic origin of these parts of the reproductive tract. The thin-walled vagina proper (averaging about 20 cm in length) ends caudally at the transverse fold above the external urethral orifice. This fold is a remnant of the hymen, which partitioned the vestibule from the vagina proper. It may extend for a variable distance up the lateral walls, and occasionally a persistent hymen is present.11 Stratified squamous epithelium covers the wellvascularized lamina propria of the longitudinally folded vaginal mucous membrane. An abrupt junction exists with the simple columnar epithelium on the protruding cervical folds at the external uterine orifice. During estrus, an increased amount of mucus on the pale mucosal surface comes from the cervical canal. The equine vagina proper is aglandular. Cornification of the superficial layer of the epithelium is minimal during estrus and cannot be used as a cytological criterion for determining different stages of the estrous cycle.
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Longitudinal folds of the vaginal mucous membrane are effaced when the organ is distended. Exposure to air tends to darken the mucous membrane. The fibroelastic tissue and smooth muscle in the wall permit distension of the vagina to the limits of the pelvic wall during passage of the foal. At other times, compression from feces in the rectal ampulla causes dorsoventral collapse of the vaginal wall, reducing the lumen to a horizontal slit. The vagina is enclosed by a loose collagenous connective tissue adventitia, containing numerous sympathetic ganglia, a venous plexus, and adipose tissue. As noted earlier, the extent of the cranial part of the vagina covered by peritoneum is related inversely to the fullness of the rectum and bladder. Vestibule The vestibule (whether vaginal or vulvar) has a short, straight dorsal wall and a longer (10–12 cm), sloping ventral wall extending to the labia of the vulva just beyond the ischiatic arch. From the ventral commissure of the labia inward, this configuration gives the vestibular vault a ventrodorsal slope (Figure 165.4b). Parting the vulvar labia reveals a pink to brownish red, folded mucous membrane of the vestibule (Figure 165.10). Stratified squamous epithelium continues as the epithelial lamina of the mucous membrane. A layer of encircling smooth muscle fibers lies deep to the proper lamina. The latter contains elastic fibers and lymph nodules.25 A ventral row and a lateral row of openings of branched tubular, mucous vestibular glands extend along each side. A darker region of mucous membrane on each lateral wall just cranial to the vulvar labium lies over a vestibular bulb, a mass of erectile tissue approximately 7 × 3 cm (Figures 165.8 and 165.10). The vestibular bulb is related laterally to the constrictor muscle of the vestibule. The constrictor muscle of the vestibule arises from the levator fascia and the lateral edges of the perineal septum, which covers the roof of the vestibule.10 Some of the muscle’s fibers attach to the retractor muscle of the clitoris (suspensory ligament of the anus). Here the clitoral retractor is a band of smooth muscle descending from a decussation ventral to the subanal loop of the muscle and extending dorsocaudal to the vestibular constrictor toward the vulvar labium.10 The sides and ventral aspect of the vestibule are covered by the vestibular constrictor muscle that blends caudally with the vulvar constrictor muscle (Figure 165.4a). Vulva Projecting caudad over the ischiatic arch, the vulva consists of two labia and the clitoris. The labia are
Figure 165.10 View of vulva and vestibule through parted vulvar labia. 1, Dorsal commissure; 2, orifices of vestibular glands; 3, location of vestibular bulb (beneath darker mucous membrane); 4, vulvar labium; 5, transverse frenular fold (clitoral prepuce); 6, clitoral glans; 7, ventral commissure. Inset: clitoral glans. 8, Transverse frenular fold retracted; 9, frenulum; 10, openings of lateral sinuses; 11, opening of median sinus; 12, locations of frenular sinuses; 13, clitoral fossa; 14, labial recesses; 15, locations of ventral sinuses. (Locations of sinuses adapted in part from McAllister and Sack.26 ). With permission from American Veterinary Medical Association.
in apposition at the vulvar cleft, uniting at the sharp angle of the dorsal cominissure and at the rounded ventral commissure. At the mucocutaneous junction, the aglandular mucous membrane meets the smooth, highly glandular, usually pigmented skin covering the labia. The mucocutaneous junction continues onto the transverse frenular fold (clitoral prepuce9 ). The vulvar constrictor muscle and, on each side, the smooth muscle of the clitoral retractor are contained within the labia. Dorsally, the vulvar constrictor muscle is continuous with the caudal part of the external anal sphincter muscle; ventrally, it sweeps under the clitoris. On its way toward the clitoris, each band of the clitoral retractor muscle is covered by superficial fascia and a few fibers of the vulvar constrictor muscle.10 Contractions of the vulvar constrictor function with the vestibular constrictor during copulation. The former muscle also causes eversion of the clitoris (‘winking’) following urination or during the flow of fluids during estrus.11 The arrangement of the tissues within the labia permit their wide expansion during delivery of the foal.
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Internal Reproductive Anatomy The wrinkled, creased skin of the large clitoral glans is observed when the labia are parted at the ventral commissure (Figure 165.10). The clitoral glans presents a central part separated partially by grooves from two lateral parts. A clitoral fossa surrounds the sides and yentral aspect of the glans, and the transverse frenular fold of integument covers the glans dorsally. A median frenulum extends from the clitoral glans to the transverse frenular fold. A constant median clitoral sinus (about 1 cm deep) and two shallow (2–3 mm deep) lateral sinuses present in 75% of clitorides are located in creases in clitoral skin immediately ventral to the transverse frenular fold (Figure 165.10). In 30% of clitorides examined, the orifice of the median sinus could be seen without displacing the transverse frenular fold.26 Frenular recesses at the junction of the transverse frenular fold with the medial labial walls and the glans and ventral recesses in the deepest part of the clitoral fossa are 7–8 mm deep. Shallower labial recesses, 2 mm deep, may be observed in the medial labial walls. All of these invaginations contain a variable amount of smegma (from sebaceous glands in clitoral skin). The significance of this detailed anatomy is that, although smegma may be sponged from frenular, ventral, and labial recesses, it is retained in the median and lateral sinuses. But the lateral sinuses are considered too shallow to support the growth of anaerobic bacteria, in particular Taylorella equigenitalis.26
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The body of the mare’s large clitoris contains an erectile tissue mass, the corpus cavernosum clitoridis. Two crura extend caudad to attach on the ischiatic arch.
Vascular and neural relationships of the reproductive tract
Arterial supply and venous drainage The ovary is supplied by the tortuous ovarian branch of the ovarian artery coursing ventrolaterad from the abdominal aorta. A smaller uterine branch of this artery anastomoses with the cranial branch of the main blood supply to the uterus, the uterine artery, descending from the external iliac artery. These vessels supply blood to the uterine tube and cranial part of the uterine horn. The caudal branch of the uterine artery anastomoses with the uterine branch of the vaginal artery (from the internal pudendal artery), providing the blood supply to the caudal part of the uterine horn and the uterine body (Figures 165.11 and 165.12). Blood from the ovary drains through several smaller veins that converge to form the ovarian branch of the ovarian vein. While satellite veins accompany the arteries, the main venous drainage of the uterus is via the ovarian vein. Interconnecting veins from the ovary and cranial part of the uterus form an extensive utero-ovarian plexus, but the close
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Figure 165.11 Dorsal view of latex casts of arteries to uterine horn and ovary in the mare. Insert: ventral view of ovary. 1, Caudal branch of uterine artery; 2, cranial branch of uterine artery; 3, uterine branch of ovarian artery; 4, uterine artery; 5, ovarian artery; 6, ovarian branch of ovarian artery. (From Ginther et al.27 ). With permission from American Veterinary Medical Association.
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Figure 165.12 Dorsal view of intervascular relationships of the vessels to the ovary and uterine horn. Latex casts of main arteries (gray) and veins (black). Venous branches to uterine horn incompletely injected. Insert: Ventral view of blood supply to ovary. 1, Uterine horn; 2, ovary; 3, ovarian branch of ovarian artery; 4, uterine branch of ovarian artery; 5, ovarian branch of ovarian vein; 6, ovarian vein; 7, ovarian artery; 8, uterine branch of ovarian vein. (From Ginther et al.27 ). With permission from American Veterinary Medical Association.
intertwining of the ovarian vein and artery observed in other species (e.g., the ewe) does not exist in the mare.27 The main blood supply to the caudal part of the reproductive tract is from the internal pudendal artery.28 A small branch from the umbilical artery courses craniad along the ureter into the broad ligament. The first part of the uterine branch of the vaginal artery (which may also arise from the umbilical or cranial gluteal artery) supplies the vagina, and its vestibular branch supplies the rest of the vagina and courses ventrad around the vestibule. The next branch, the ventral perineal artery, gives a dorsal labial branch to the vulva (Figure 165.13) and occasionally sends one to the vestibular bulb. The termination of the internal pudendal artery is the artery of the vestibular bulb. It courses to the ventral surface of the vulva, providing small branches to the vestibular bulb. A second blood supply to this region, the
Figure 165.13 Nerve and blood supply to vulvar labium. 1, 2, and 3, Superficial perineal nerves; 4, ventral perineal artery and vein. Size of nerves and vessels slightly exaggerated. (Adapted in part from Habel.10 )
obturator artery, terminates by entering the root of the clitoris, dividing into deep and dorsal arteries of the clitoris. Satellite veins carry blood from the caudal part of the reproductive tract. Veins from the clitoris communicate with the vestibular bulb.
Lymphatic drainage Lymph flows from the ovaries, uterine tubes, and uterus to lumbar aortic lymph nodes.29 Lymphatic vessels from the uterus, vagina, and vulva are afferent to medial iliac lymph nodes. A uterine lymph node occurs occasionally in the broad ligament.
Innervation Renal, aortic, uterine, and pelvic plexuses supply sympathetic fibers to the ovaries, uterine tubes,
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Internal Reproductive Anatomy
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Figure 165.14 Gross anatomy of the equine udder. (a) Udder of a virgin mare. (b) Lactating udder. 1, Intermammary groove. (c) Sagittal section of a lactating udder. 1, Cranial papillary duct (streak canal); 2, caudal papillary duct; 3, papillary part of caudal lactiferous sinus (teat cistern); 4, glandular part of cranial lactiferous sinus (gland cistern).
uterus, and vagina.14 Pudendal and caudal rectal nerves (derived from the third and fourth sacral nerves) innervate the muscles of the vestibule and vulva and the vulvar skin. They both contribute to the formation of superficial permeal nerves, which emerge ventrolateral to the anus and supply the labia (Figure 165.13).10
The mare’s udder The small, inconspicuous udder of the virgin mare lies caudal to two prominent folds of abdominal skin extending to the bases of two small, laterally flattened teats. In some mares each fold of skin continues caudad lateral to the teat (Figure 165.14). The longitudinal intermammary groove dividing the right and left halves of the organ contains wrinkled skin covered with a layer of dark colored sebum that can be peeled away. In very young fillies, a few hairs protrude from the orifices at the tips of the teats, but they soon regress. As the ducts, secretory alveoli and their enclosing myoepithelial cells, and supportive tissues develop during gestation, the udder enlarges, effacing cutaneous folds. The teats of the roughly flattened hemispherical halves (mainmac) of the udder elongate and become conical. Two (sometimes three) ostia (orifices) – one caudal to the other – open into a depression at the apex of the teat. Each orifice leads to
a separate papillary duct (streak canal). Continuing from each papillary duct, a separate papillary part (teat cistern) leads into the glandular part (gland cistern) of the lactiferous sinus. Thus, there are two (or three) closely associated lactiferous duct systems in the glandular complex of each half of the mare’s udder with the cranial system being the largest (Figure 165.14). At its junction with the teat cistern, the stratified squamous epithelium of the folded lining the papillary duct gradually changes to the bistratified columnar epithelium, which lines most of the lactiferous duct system. Smaller ducts leading from the alveoli are formed by simple columnar epithelium. Smooth muscle, fibroelastic tissue, and blood vessels constitute the bulk of the teat wall. The apex of the teat lacks an organized sphincter.30 Support for each half of the udder is provided by a fibrous lateral suspensory ligament and a more elastic medial ligament. The medial suspensory ligament is separated from the contralateral ligament by loose collagenous connective tissue.14 Supporting trabeculae projecting inward from the ligaments enclose illdefined lobes and branch into finer lobular connective tissue surrounding the ducts and alveoli. The sparsely haired skin of the udder, including the teats, is slightly thicker than the average for equine skin.31 Numerous sebaceous glands and apocrine tubular sweat glands are present, increasing toward the apices of the teats. Sebum in the
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intermamrnary groove protects the halves of the udder against friction during locomotion.30 Concentrations of sebaceous glands occur peripheral to eachpapillary duct ostium. ‘Waxing’ at the tips of the teats is a phenomenon presaging impending parturition. It is caused essentially by increased secretion of sebum plus small amounts of cellular debris and colostrum. Each external pudendal artery descends through an inguinal canal and enters the caudal part of the udder, branching into cranial and caudal mammary arteries. An interconnecting venous plexus on either side of the base of the udder drains principally to contralateral external pudendal veins. Blood also drains from the plexus craniad in the caudal superficial epigastric vein, which connects with a superficial vein of the thoracic wall (especially during the first lactation1 ). The plexus also connects caudally with the obturator vein or with branches of the internal pudendal vein.30 Lymph from the udder flows through lymphatic vessels afferent to a conglomerate of superficial inguinal (mammary) lymph nodes at the base of the udder. Afferents may come from the subcutis to accessory mammary lymph nodes that drain to main mammary nodes.29 Genitofemoral nerves coursing through the inguinal canals provide the main innervation to the udder. The skin of the udder is supplied by nerves of the flank and a descending branch from the pudendal nerve.1 An average-sized lactating mare produces around 10 L of milk daily.30 The milk fat content of equine milk is suite low (1.6%); the lactose content is high (6.1%).32
References 1. Dyce KM, Sack WO, Wensing, CJG. Textbook of Veterinary Anatomy. Philadelphia: W.B. Saunders, 1987. 2. Reiter RJ. Pineal control of reproduction. In: Vidrio EA, Galina MA (eds) Eleventh International Congress of Anatomy: Advances in the Morphology of Cells and Tissues. New York: Alan R. Liss, 1981; pp. 349–55. 3. Wesson JA, Orr EL, Quay WB, Ginther OJ. Seasonal relationship between pineal hydroxindole-O-methyltransferase (HIOMT) activity and reproductive activity in the pony. Gen Comp Endocrinol, 1979;38:46–52. 4. Venzke WG. Endocrinology. In: Getty R (ed.) Sisson and Grossman’s The Anatomy of the Domestic Animals, vol. 1, 5th edn. Philadelphia: W.B. Saunders, 1975; pp. 550–51. 5. Krieger DT. The hypothalamus and neuroendocrinology. In: Krieger DT, Hughes JC (eds) Neuroendocrinology. Sunderland: Sinauer Associates, 1980; pp. 3–12. 6. Vitums A. Observations on the equine hypophysial portal system. Anat Histol Embryol, 1975;4:149–161. 7. Nanda BS. Blood supply to the brain. In: Getty R (ed.) Sisson and Grossman’s The Anatomy of the Domestic Animals, vol. 1. 5th edn. Philadelphia: W.B. Saunders, 1975; p. 579.
8. Vitums A. Development of the venous drainage of the equine hypophysis cerebri. Anat Histol Embryol 1978;7: 120–8. 9. Nomina Anatoniica Veterinaria, 3rd edn. Ithaca: World Association of Veterinary Anatomists, 1983. 10. Habel R. The perineum of the mare. Cornell Vet 1953;43:249–78. 11. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects. Ann Arbor: McNaughton, Gunn, 1979. 12. Warszawsky LF, Parker WG, First NL, Ginther OJ. Gross changes of the internal genitalia during the estrous cycle in the mare. Am J Vet Res 1972;33:19–26. 13. Witherspoon DM. The site of ovulation in the mare. J Reprod Fertil Suppl 1975;23:329–330. 14. Sisson S. Female genital organs. In: Getty R (ed.) Sisson and Grossman’s The Anatomy of the Domestic Animals, vol. 1, 5th edn. Philadelphia: W.B. Saunders, 1975; pp. 542–9. 15. Scott EA, Kunze DJ. Ovariectomy in the mare: Presurgical, surgical and postsurgical considerations. J Equine Med Surg 1977;1:5–12. 16. Ono H, Satoh H, Miyake M, Fujimoto Y. On the development of ‘ovarian adrenocortical cell nodule’ in the horse. Exp Reprod Equine Health Lab 1969;6:59–90. 17. Squires EL, Douglas RH, Steffenhagen WP, Ginther OJ. Ovarian changes during the estrous cycle and pregnancy in mares. J Anim Sci 1974;38:330–8. 18. Prickett ME. Pathology of the equine ovary. Proc Am Assoc Equine Pract 1966; pp. 145–154. 19. Van Niekerk CH, Gerneke WH. Persistence and parthogenetic cleavage of tubal ova in the mare. Onderstepoort J Vet Res 1966;33:195–232. 20. Kenney RM. Cyclic and pathologic changes in the mare endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1978;172:241–62. 21. Lieux P. Relationship between the appearance of the cervix and the heat cycle in the mare. Vet Med Small Anim Clin 1970;65:859–66. 22. Hafez ESE. Reproduction in Farm Animals. 3rd edn. Philadelphia: Lea & Febiger, 1974; p. 51. 23. Clegg MT, Boda JM, Cole HH. The endometrial cups and allantochorionic pouches in the mare with emphasis on the source of equine gonadotropin. Endocrinology 1954;54:448–63. 24. Allen WR, Hamilton EW, Moor RM. Origin of equine endometrial cups. II. Invasion of the endometrium by trophoblast. Anat. Rec 1973;177:485–502. 25. Krolling O, Grau H. Lehrbuch der Histologie und vergleichenden mikroskopischen Anatomie der Haustiere. Berlin: Paul Parey, 1960. 26. McAllister RA, Sack WO. Identification of anatomic features of the equine clitoris as potential growth sites for Taylorella equigenitalis. J Am Vet Med Assoc 1990;196:1965–6. 27. Ginther OJ, Garcia MC, Squires EL, Steffenhagen WP. Anatomy of the vasculature of the uterus and ovaries in the mare. Am J Vet Res 1972;33:1561–8. 28. Goshal NG. Equine heart and arteries. In: Getty R. (ed.) Sisson and Grossman’s The Anatomy of the Domestic Animals, vol. 1, 5th ed,. Philadelphia: W.B. Saunders, 1975: pp. 605– 7. 29. Saar LI, Getty R. Equine lymphatic system. In: Getty R (ed.) Sisson and Grossman’s The Anatomy of the Domestic Animals, vol. 1. 5th edn. Philadelphia: W.B. Saunders, 1975; pp. 625–7. 30. Schummer A, Wilkens H, Vollmerhaus B, Habermehi K-H. In: Nickel R, Schummer A, Seiferle’s E (eds) Anatomy of the
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Internal Reproductive Anatomy Domestic Animals. vol. 3. The Circulatory System, the Skin, and the Cutaneous Organs of the Domestic Mammals. New York: Paul Parey, 1981; pp. 538–40. 31. Talukdar AH, Calhoun ML, Stinson AW. Microscopic anatomy of the skin of the horse. Am J Vet Res 1972;33:2365–90.
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32. Jacobson NL, McGilliard AD. The mammary gland and lactation. In Swenson MJ (ed.) Dukes’ Physiology of Domestic Animals, 10th edn. Ithaca: Comstock Publishing Associates, 1984, p. 871.
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Section (b): Physiology and Endocrinology
166.
How Hormones Work Peter R. Morresey
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167.
GnRH Susan L. Alexander and Clifford H.G. Irvine
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168.
FSH and LH Susan L. Alexander and Clifford H.G. Irvine
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169.
Estrogens Bruce W. Christensen
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170.
Progesterone Dirk K. Vanderwall
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Prostaglandins Tom A.E. Stout
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Equine Chorionic Gonadotropin (eCG) Amanda M. de Mestre, Douglas F. Antczak and W.R. (Twink) Allen
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Adrenal Steroids Valerie J. Linse
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Melatonin Daniel C. Sharp
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Oxytocin, Inhibin, Activin, Relaxin and Prolactin Peter R. Morresey
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Chapter 166
How Hormones Work Peter R. Morresey
Introduction The endocrine system is an example of a humoral control system in the body. The messengers of this system, hormones, are classically broadly defined as chemical messengers produced by a ductless endocrine gland that are transported via the bloodstream to a distant target organ to effect changes in biochemical processes. This definition, however, is incomplete as it does not include those substances that do not enter the bloodstream, and those that are secreted by ducted glands.
History Starling in 1905 is credited with the first application of the term hormone from the Greek verb hormoa, to excite.1 This term was used in the context of describing substances which ‘have to be carried from the organ where they are produced to the organ which they affect by means of the blood stream and the continually recurring physiological needs of the organism must determine their repeated production and circulation throughout the body’.2
Methods of hormonal signaling
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Endocrine: the hormone is secreted by a ductless gland, is distributed in the blood, and binds to distant target cells. Example: gonadotropin (folliclestimulating hormone, FSH; luteinizing hormone, LH) action stimulating ovarian steroid production from granulosa and theca cells. Paracrine: the target cell is physically close to the hormone-releasing cell. The hormone acts locally by diffusing from its source to target cells in the neighborhood. Examples: histamine, neurotransmitters, and neurohormones. Autocrine: the hormone acts on the same secretory cell that produced it. Examples: growth factors, cytokines.
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Neuroendocrine: cells acting in this manner provide a linkage between the nervous and endocrine systems. Hormones are released into the circulation in response to a neural stimulus. Example: epinephrine release from the chromaffin cells of the adrenal medulla. Juxtacrine: intercellular communication requires physical contact between cells, with transmission via oligosaccharide, lipid, or protein components of the cell membrane. Adjacent cells must possess broad patches of closely opposed plasma membrane linked by transmembrane channels known as connexons. Examples: growth factors, cytokine and chemokine cellular signals. Intracrine: refers to the action of the hormone, in this case being intracellular, being directed towards the cell of origin following retention or occurring after internalization. Examples: some peptide hormones and growth factors. Exocrine: the hormone or secretory product is directed outwards via a duct system. Examples: within the reproductive system the prostate and mammary glands. Outside the reproductive system many examples exist, including the exocrine pancreas, sweat glands, and salivary glands.
Some hormones can be classified as working in more than one of the above fashions. Example: testosterone has both paracrine actions (stimulates spermatogenesis in local seminiferous tubules) and endocrine actions (secondary sexual characteristics and muscle deposition). A pulsatile pattern of secretion is seen for virtually all hormones, with variations in pulse characteristics that reflect specific physiological states. In addition to the short-term pulses, longer-term temporal oscillations or endocrine rhythms are also commonly observed and are undoubtedly important in both normal and pathological states. Differential secretion of gonadotropin-releasing hormone (GnRH) with respect to pulse frequency
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Physiology and Endocrinology
and amplitude has profound effects on reproductive function in both the female and male.3 In the female, a slow GnRH pulse frequency favors increased secretion of FSH, resulting in the development of the ovarian follicle during the follicular phase of the estrous cycle. The onset of high frequency and amplitude GnRH pulses results in a sudden increase in LH leading to ovulation, marking the onset of the luteal phase. Conversely, constant GnRH secretion has been shown to suppress pituitary LH and FSH secretion which is only restored once pulsatile secretion recommences. More in depth information on the physiology of these hormones is available in Chapters 167 and 168.
Hormone classes Steroid hormones Steroid hormones are derived from cholesterol and include products of the adrenal cortex (glucocorticoids, mineralocorticoids, androgens), the gonads (estrogen, progesterone, testosterone), and vitamin D (see Table 166.1). All adrenal and gonadal steroids have the same basic ring structure, the cyclopentanophenanthrene ring, and the same atomic numbering system as
cholesterol. Specificity results from differences in side chains and spatial orientation. The conversion of cholesterol to pregnenolone is the first and rate-limiting step in the synthesis of all steroid hormones. The eventual conversion of 27carbon cholesterol to the 21-, 19-, and 18-carboncontaining steroid hormones begins with cleavage of a 6-carbon residue from cholesterol, producing pregnenolone (C21 ) plus isocaproaldehyde. Pregnenolone is formed on the inner membrane of mitochondria by the action of cytochrome P450scc. Repeated passage between the mitochondria and the endoplasmic reticulum allows further enzymatic transformations to form the various steroid hormones. Pregnanes contain 21 carbon atoms, androstanes contain 19, and estranes contain 18 carbon atoms. Vitamin D differs from the other steroid hormones in that it is formed directly from cholesterol, not pregnenolone. Hydroxylation of cholecalciferol and ergocalciferol occurs first in the liver followed by the kidney, bone or placenta to form the active form 1,25 dihydrocholecalciferol (calcitriol). Increases in secretion of steroid hormones reflect accelerated rates of synthesis. Following appropriate stimulus, enzymes can rapidly act to generate the desired product. Cholesterol for steroid synthesis is ubiquitous and does not require gene expression, rather the presence of specific enzymes that convert cholesterol into the appropriate steroid. Different
Table 166.1 How hormones work Class
Examples
Method of synthesis
Action
Target tissues
Steroid
- estrogen, progesterone, testosterone - cortisol - aldosterone, vitamin D
Enzymatic transformation of cholesterol under influence of trophic peptide hormone
Alteration of gene expression
- genitalia, secondary sex characteristics - metabolically active tissue - kidney
Peptide and protein
- GnRH, ACTH - FSH, LH - insulin
Synthesized as pre-prohormones in endoplasmic reticulum. Considerable post synthetic modification
Binding to cell transmembrane receptors initiates intracellular enzyme cascade
- pituitary - gonads - multiple energy storage and expending tissues (muscle, fat, liver)
Amino acid derivatives
- epinephrine, thyroxine - norepinephrine - serotonin, melatonin
Conjugation or modification of amino acids tyrosine and tryptophan
Transmembrane receptors initiate intracellular second messenger production (except thyroxine – intracellular receptors)
- multiple energy storage and expending tissues (muscle, fat, liver) - neurotransmitter - brain
Fatty acid derivatives (eicosanoids)
- prostaglandins - leukotrienes - thromboxanes
Metabolism of arachidonic acid, a cell membrane fatty acid
Intracellular second messenger production (cyclic nucleotides, calcium)
Act locally due to rapid inactivation, alter intracellular enzyme cascades - bronchoconstriction, vasodilation, vasoconstriction, platelet aggregation - vascular permeability, vasoconstriction - platelet aggregation
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How Hormones Work steroid-secreting cells express different enzymes, and are controlled by tropic hormones and other factors. Steroid hormones are rapidly secreted from the cell, with little if any storage occurring. Once secreted, steroids non-specifically bind to plasma proteins (albumin) with low affinity; however, some are transported by specific binding proteins. These binding proteins affect steroid half-life and rate of elimination. Steroid hormones are predominantly eliminated by metabolic inactivation and excretion in urine or bile. Peptide and protein hormones Peptide and protein hormones vary considerably in size and post-translational modifications, ranging from peptides as short as three amino acids to large, multiple subunit glycoproteins with complex sugars attached. Examples of peptide hormones include GnRH, the gonadotropins, prolactin, adrenocorticotropic hormone (ACTH), growth hormone, and oxytocin (see Table 166.1). Peptide and protein hormones are synthesized in endoplasmic reticulum, the initial protein generated being unlikely to be the final secreted molecule. These hormones can be synthesized as larger (pre)prohormones, then reduced in the endoplasmic reticulum by proteolysis to generate an active molecule. Multiple cleavages may be necessary to release a hormone embedded within the sequence of a large precursor. Hormones are then transferred to the Golgi apparatus for further cleavage and packaging into secretory vesicles. Peptide and protein hormones can be secreted by one of two pathways. Regulated secretion is the most commonly used pathway. Cells store hormones in vesicles which are released in bursts following stimulation, allowing cells to secrete a large amount of hormone over a short period of time. With constitutive secretion, the cell does not store hormone but secretes it from secretory vesicles as it is synthesized. Once released, the half-life of circulating peptide hormones is only a few minutes. Peptide and protein hormones are cleaved by circulating proteases, or degraded within the target cell following interaction with the receptor. Amino acid derivatives Hormones derived from amino acids have tyrosine and tryptophan as their basis. Tyrosine yields the catecholamines epinephrine and norepinephrine, the latter also functioning as a neurotransmitter. The conjugation of two tyrosine molecules with incorporation of iodine results in production of the thyroid hormones. Tryptophan is the precursor of both serotonin and melatonin (see Table 166.1).
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Once secreted by the adrenal medulla catecholamines are rapidly degraded by metabolism, with circulating half-lives of only minutes. The circulating half-life of thyroid hormones ranges from hours to a few days, being inactivated by intracellular deiodinases. Fatty acid derivatives (eicosanoids) Eicosanoids are derived from a number of 20-carbon polyunsaturated fatty acids making up cell and nuclear membranes known as essential fatty acids. They consist of the omega-6 (arachidonic acid, eicosapentanoic acid) and omega-3 (lipotechoic) groups. Eicosanoids derived from the omega-6 fatty acids are generally proinflammatory, with the omega-3 derived compounds markedly less so with purported anti-inflammatory properties. All mammalian cells except erythrocytes synthesize eicosanoids and these compounds cause profound physiological effects at very low concentrations. Eicosanoids function locally at the site of synthesis via cell membraneassociated G-protein-linked signaling pathways (see Table 166.1). The principal groups of hormones of the eicosanoids class are prostaglandins, prostacyclins, leukotrienes, and thromboxanes. Prostaglandins were first shown to be synthesized in the prostate gland, leukotrienes were derived from leukocytes, and thromboxanes from platelets (thrombocytes) leading to the derivation of their names. Eicosanoids are responsible for a wide range of biological actions, including inflammation, the intensity and duration of pain and fever, and on reproductive function (luteolysis, parturition). They also regulate blood pressure through vasodilation or vasoconstriction, inhibit or activate platelet aggregation and thrombosis, and modulate gastric acid secretion. Arachidonic acid is the principal source of the biologically relevant eicosanoids, with the minor eicosanoids derived from eicosapentaenoic acid. Linoleic acid (arachidonic acid precursor) and αlinolenic acid (eicosapentaenoic acid precursor) are essential fatty acids obtained from the diet. The major source of arachidonic acid is through its release from cell membrane lipid stores by the action of phospholipase A2 (PLA2 ). Numerous stimuli are responsible for the activation of PLA2 , including epinephrine, inflammatory mediators such as histamine, and components of the coagulation system such as thrombin. Cell production of specific eicosanoids is determined by the enzymes expressed in that cell with two main pathways being involved: the prostaglandins and thromboxanes are synthesized by the cyclic pathway, the leukotrienes by the linear pathway. The
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cyclic pathway involves the action of prostaglandin endoperoxide synthetase (PGS), this enzyme having both cyclooxygenase (COX) and peroxidase activities. Cyclooxygenase-1 (COX-1) activity is present in gastric mucosa, kidney, platelets, and vascular endothelial cells in a constitutive role. The inducible COX-2 is expressed by macrophages and monocytes in response to inflammation. Lipoxygenase (LOX) initiates the linear pathway and occurs in three forms: 5-LOX, 12-LOX, and 15-LOX. The leukotrienes result from the action of 5-LOX in leukocytes, mast cells, lung, spleen, and heart. Eicosanoids are rapidly inactivated by metabolism, and are typically active for only a few seconds. Continued effect is therefore dependent on continued production.
Transport of hormones Steroid and thyroid hormones are less soluble in aqueous solution than protein and peptide hormones, with the great majority circulating in blood bound to specific plasma globulins or albumin. Most binding proteins are synthesized in the liver. Alterations in serum concentrations of these proteins alter total serum concentrations of hormones; however, this has much less effect on the concentrations of free hormone. Free hormone is biologically active, with hormone binding delaying metabolism and providing a circulating reservoir. Specific binding globulins appear not to be solely passive transporters, but may also interact with membrane receptors affecting signal transduction. Recently, binding proteins for several protein and peptide hormones as well as some growth factors have been identified. Amino acid derivatives upon release from the source gland into the circulation may bind specifically to a number of dedicated plasma proteins, or non-specifically to albumin. Fatty acid derivatives (eicosanoids) are unlike most hormones in that, although they act as hormones, they do not conform to the classic definition as they are generally not produced in a central location before transport to the target organ. Eicosanoids act locally in the source cells and disappear quickly due to inactivation and extremely short half-lives.
Feedback/control mechanisms Feedback circuits are the basis of most control mechanisms in physiology, and are particularly prominent in the endocrine system. Negative feedback is seen when the output of a pathway inhibits inputs to the pathway providing a means to achieve equilibrium. Instances of positive feedback occur, but negative feedback is the predominant mechanism of control.
The effects of hormones on target cell physiology are controlled by the concentration delivered, this being determined by the rate of production, rate of delivery, and rate of elimination of the hormone itself. Production and secretion of hormones are the most highly regulated aspect of the endocrine system, with control occurring by positive and negative feedback loops. Rate of delivery is controlled by blood flow to a target organ or group of target cells, with higher blood flow delivering more hormone and therefore more response is generated than occurs with low blood flow. Hormones have predictable rates of decay, and are eliminated (metabolized and excreted) from the body by a number of routes. The half-life of the hormone is also a factor in persistence of effect. The half-life is defined as the period of time necessary for clearance of one half of the maximal concentration of the hormone released. Stopping secretion of a hormone with a short half-life causes circulating hormone levels to rapidly decrease, allowing attenuation of effect. A long half-life, however, allows effective concentrations to remain in circulation for an extended period, which prolongs the hormone’s effect for some time after secretion stops. The rate of metabolism of specific hormones in the circulation varies; as a generalization, catecholamines from the adrenal medulla have a half-life of seconds, protein and peptide hormones have a half-life of minutes, and steroid and thyroid hormones have their half-lives measured in hours. An example of feedback is the pituitary–ovarian axis. Subsequent to ovulation of the ovarian follicle and formation of the corpus luteum, high levels of ovarian steroids (estrogen and progesterone) feed back to the hypothalamus inhibiting GnRH pulses and therefore both LH and FSH secretion. Following demise of the corpus luteum, the fall in ovarian steroids restarts the estrous cycle by allowing the return of the slow frequency GnRH pulses required for FSH release and recruitment of the next follicle.
Hormone–cell interaction: receptors and second messengers Hormones alter the structure and function of their receptors thereby causing an effect. Alterations in cell permeability, activation of an intracellular enzyme cascade, or activation of gene transcription can be thought of as the outcomes of hormone–cell interaction. Receptors Hormones generally do not affect intracellular processes directly, but instead first combine with a
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How Hormones Work receptor located on the cell membrane or within the cytoplasm of the cell. This initiates a cascade of amplifying reactions within the cell. Receptors tend to be large proteins that are found in great numbers on target cells. They tend to be very specific for the hormone acting on the target tissue. Receptor location within the target cell varies according to the hormone involved. Hormones can be further classified depending on the location of their receptor: the lipid-soluble hormones (steroids, thyroxine) which can diffuse through plasma membranes to directly interact with their cytoplasmic receptors, and the water-soluble hormones (peptides, proteins, and the catecholamines) which have transmembrane receptors located within the membrane or on the surface of the cell. The number of receptors on the target cell is dynamic. Hormone binding decreases the number of active receptors available as well as altering receptor protein generation. Control of hormone–receptor interaction also happens through internalization of cell surface receptors, this in many cases being stimulated by hormone binding. Internalization occurs by endocytosis, with the resulting endosomes subsequently fusing with lysosomes. This allows destruction of the hormone–receptor complex. Alternately, the hormone may dissociate from the receptor, freeing the receptor for recycling back to the cell membrane. Downregulation is the process whereby continuous high levels of hormone decrease the population of available receptors. This is the most common response to continued hormone secretion and is seen in nearly all feedback systems. Upregulation does occur rarely, one example being an increase in follicular FSH receptors in response to continued FSH secretion during the early follicular phase. Second messengers Binding of hormones to cell membrane receptors initiates a series of events which culminate in the generation of second messengers within the cell (by definition the hormone becomes the first messenger). The second messengers then initiate a series of molecular interactions that change the physiological state of the cell. Multiple hormones can utilize the same second messenger system, and individual hormones can utilize more than one system. By the action of second messengers, a small signal can be amplified sufficiently to cause a physiological change in the target cell. Second messenger initiated cascades are necessarily transient events to enable restoration of normal unstimulated cell function to occur once hormonal stimulus is no longer required. A number of second messenger systems are recognized and these can be placed in three categories.
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Cyclic nucleotides: cyclic adenosine monophosphate (cAMP) and cyclic guanosine monophosphate (cGMP) The stimulating hormone first binds to its receptor and activates a G protein which, in turn, activates adenylyl cyclase.4 The resulting increase in cAMP activates the appropriate response in the cell, being a change in the molecular activities in the cytosol as a result of protein kinase A (PKA) phosphorylation of target proteins, or a new gene transcription is begun. Hormones using this pathway include LH, epinephrine, and glucagon. Cyclic GMP is synthesized from the nucleotide guanosine triphosphate (GTP) using the enzyme guanylyl cyclase following hormone receptor interaction. Effects of cGMP are mediated through phosphorylation of proteins by protein kinase G (PKG), a cGMP-dependent protein kinase. Nitric oxide and atrial natriuretic peptide are among those hormones that use cGMP as a second messenger.
Membrane phospholipid breakdown products (phophatidylinositol system) The activation of G protein-coupled receptors (GPCR) by hormone binding activates the intracellular enzyme phospholipase C (PLC), leading to hydrolysis of the phospholipid phosphatidylinositol4,5-bisphosphate (PIP2 ) which is found in the inner layer of the plasma membrane.5 Breakdown of PIP2 yields two products: diacylglycerol (DAG) and inositol-1,4,5-trisphosphate (IP3 ). Protein kinase C (PKC), a calcium-dependent kinase that phosphorylates other proteins, is recruited by DAG which remains in the inner layer of the plasma membrane. Calcium ions are made available by IP3 which binds to receptors on the endoplasmic reticulum causing release of calcium ions into the cytosol. Peptide and protein hormones, neuronal and immune cell signaling use aspects of this system.
Calcium/calmodulin Calcium ions can be considered the most widely used intracellular messenger.6 They rise in concentration in the cell by the action of voltage-gated channels, receptor-controlled channels, release of intracellular stores (endoplasmic reticulum, mitochondria) and the action of GPCR. Calcium influences a great number of biological processes including cell motility, muscle contraction, axonal flow, cytoplasmic streaming, chromosome movement, neurotransmitter release, endocytosis, exocytosis, and glycogen metabolism.7 Calmodulin is an intracellular protein that undergoes a conformational change following binding with
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calcium ions.8 This leads to activation of calmodulin which in turn activates enzyme cascades separate from cAMP. Calcium has the ability to regulate its own cellular concentration by interacting with calmodulin.9 Calcium is extracted from the cytoplasm by the action of Ca2+ -ATPase. This enzyme is activated by an increase in intracellular calcium binding to calmodulin which causes calcium to be released from the cytoplasm. This allows a low level of intracellular calcium through self-regulation.
Mechanisms of hormone action Steroid and thyroid hormones: intracellular receptors Receptors for steroid and thyroid hormones are located inside target cells, in either the cytoplasm or nucleus, and are members of a superfamily of transcription factors.10 Steroid hormones, being lipophilic, enter the cell by simple diffusion across the plasma membrane. Thyroid hormones enter by facilitated diffusion. Multiple forms of a receptor may be expressed, allowing a complex response to stimulation. Receptors are composed of a single polypeptide chain with three distinct domains. The aminoterminus is involved in stimulating transcription by interacting with components of the transcriptional apparatus. This region is highly variable among different receptors. The DNA-binding domain contains amino acids allowing binding of the receptor to specific DNA sequences. The carboxy-terminus (also known as the ligand-binding domain) binds the hormone. Two other important regions of the receptor protein are the nuclear localization sequence, which targets the protein to the nucleus, and a dimerization domain, which attaches two receptors together to enable binding of DNA. Hormone binding to the receptor initiates a set series of events.11 Conformational changes in the receptor are induced by hormone binding, this being termed receptor activation. This allows the receptor to bind DNA. Activated receptors then bind to short specific sequences of DNA (hormone response elements) which are located in promoter regions of hormone-responsive genes. Most commonly, receptor binding stimulates or inhibits transcription from those genes thereby modulating gene expression in target cells. By selectively affecting gene transcription, concentrations of proteins are altered with effects on the phenotype of the cell. The hormone–receptor complex thus functions as a transcription factor. Effects on the action of nuclear factor kappa B (NF-κB), a major promoter of gene expression ubiquitous in the cell population,12, 13 have also been re-
ported for the steroid and other classes of hormones. Both inhibitory and stimulatory effects are known to occur. The primary mechanism of NF-κB control is by regulating the ability to bind DNA. Inhibitor κB (IκB) proteins bind NF-κB dimers, preventing DNA binding and nuclear accumulation, resulting in large stores of suppressed cytoplasmic NF-κB bound to IκB proteins.14 Hormonal stimulation results in release of the NF-κB-bound IκB protein, allowing NF-κB to relocate to the nucleus and bind DNA.15 In addition to the classical theory of steroid hormone signaling, it has recently become apparent that genomic responses to steroid hormones are mediated by not only the formation of a complex of the hormone and its associated steroid-hormone nuclear receptor. It appears that rapid responses are mediated by a variety of plasma membrane-associated receptors, including a membrane-associated nuclear receptor.16 Both modes of action appear necessary to explain the complex actions of steroid hormones on target cells.17 Protein and peptide hormones: second messengers Protein and peptide hormones are water soluble and hence cannot penetrate the lipid cell membrane (in contrast to steroid hormones which enter the cell). Binding of these hormones allows cell change by a process called signal transduction. Cell surface receptors for protein and peptide hormones are themselves proteins integrated into the plasma membrane.18 Each has three basic domains: extracellular, transmembrane, and intracellular. Extracellular domains are protein residues exposed to the surface of the cell that directly interact with the hormone. Transmembrane domains are hydrophobic chains of amino acids anchoring the receptor within the cell membrane lipid bilayer. Intracellular (cytoplasmic) domains are the effector regions of the receptor, interacting with molecules in the cell cytoplasm to change cell physiology. Variations in cell membrane receptor structures occur. Receptors can take the form of proteins passing once through the cell membrane, can be composed of more than one subunit, or thread through the cell membrane several times. The majority of protein and peptide hormone receptors are G-protein linked receptors. Hormone–receptor interactions cause dissociation of the associated intracellular trimeric G proteins, causing either the opening of membrane ion channels or activation of membrane-bound enzymes that stimulate or inhibit second messenger production leading to activation of serine/threonine kinases or phosphatases.
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How Hormones Work Amino acid derivatives: second messengers Binding of amino acid derivatives to the cell receptor initiates second messenger production, either the cyclic nucleotides or the PIP2 derivatives.5, 19 The possibility of direct membrane effects of the amino acid derivatives in addition to their known intracellular actions has been demonstrated.20 Fatty acid derivatives (eicosanoids): second messengers In general, eicosanoids bind to receptors on target cell plasma membranes in various tissues and stimulate or inhibit the synthesis of other messengers. Activation of PKC and increases in intracellular calcium ions are reported.21 Intercellular range of effect depends on chemical stability of the individual eicosanoid due to a lack of protective carriers, and a selective interaction of responsive cells with the range of metabolites locally present. Uptake and metabolism by surrounding cells also affects duration of eiconsanoid activity.
Summary Hormonal signaling pathways consist of a finely orchestrated interplay between hormone production, receptor regulation, and feedback mechanisms. These do not act in a manner isolated from the physiology of the entire animal. The ever-changing interactions within a hormonal feedback loop and the relationship between seemingly disparate physiological processes should be carefully considered whenever therapeutic hormonal intervention is contemplated.
References 1. Henderson J. Ernest Starling and ‘Hormones’: an historical commentary. J Endocrinol 2005;184(1):5–10. 2. Starling E. Croonian Lecture: on the chemical correlation of the functions of the body I. Lancet 1905;2:339–41. 3. Ferris HA, Shupnik MA. 2006. Mechanisms for pulsatile regulation of the gonadotropin subunit genes by GNRH1. Biol Reprod 2006;74(6):993–8.
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4. Gilman AG. G proteins: transducers of receptor-generated signals. Annu Rev Biochem 1987;56:615–49. 5. Weiss ER, Kelleher DJ, Woon CW, Soparkar S, Osawa S, Heasley LE, Johnson GL. Receptor activation of G proteins. FASEB J 1988;2(13):2841–8. 6. Brown BL, Walker SW, Tomlinson S. Calcium calmodulin and hormone secretion. Clin Endocrinol (Oxf) 1985;23(2):201–18. 7. Wang JH, Pallen CJ, Sharma RK, Adachi AM, Adachi K. The calmodulin regulatory system. Curr Top Cell Regul 1985;27:419–36. 8. MacNeil S, Lakey T, Tomlinson S. Calmodulin regulation of adenylate cyclase activity. Cell Calcium 1985;6(3):213–16. 9. Cheung WY. Calmodulin plays a pivotal role in cellular regulation. Science 1980;207(4426):19–27. 10. Whitfield GK, Jurutka PW, Haussler CA, Haussler MR. Steroid hormone receptors: evolution, ligands, and molecular basis of biologic function. J Cell Biochem 1999;Suppl 32–33:110–22. 11. Gronemeyer H. Control of transcription activation by steroid hormone receptors. FASEB J 1992;6(8):2524–9. 12. Sen R, Baltimore D. Inducibility of kappa immunoglobulin enhancer-binding protein NF-kappa B by a posttranslational mechanism. Cell 1986;47(6):921–8. 13. Sen R, Baltimore D. Multiple nuclear factors interact with the immunoglobulin enhancer sequences. Cell 1986;46(5):705–16. 14. Baeuerle PA, Baltimore D. I kappa B: a specific inhibitor of the NF-kappa B transcription factor. Science 1988;242(4878):540–6. 15. Hoffmann A, Natoli G, Ghosh G. Transcriptional regulation via the NF-kappa B signaling module. Oncogene 2006;25(51):6706–16. 16. Norman AW, Mizwicki MT, Norman DPG. Steroidhormone rapid actions, membrane receptors and a conformational ensemble model. Nat Rev Drug Discov 2004;3(1):27–41. 17. Walters MR, Nemere I. Receptors for steroid hormones: membrane-associated and nuclear forms. Cell Mol Life Sci 2004;61(18):2309–21. 18. Catt KJ, Dufau ML. Peptide hormone receptors. Annu Rev Physiol 1977;39:529–57. 19. Baxter JD, Funder JW. Hormone receptors. N Engl J Med 1979;301(21):1149–61. 20. Grollman EF, Lee G, Ambesi-Impiombato FS, Meldolesi MF, Aloj SM, Coon HG, Kaback HR, Kohn LD. Effects of thyrotropin on the thyroid cell membrane: hyperpolarization induced by hormone-receptor interaction. Proc Natl Acad Sci USA 1977;74(6):2352–6. 21. Harris RH, Ramwell PW, Gilmer PJ. Cellular mechanisms of prostaglandin action. Annu Rev Physiol 1979;41:653–68.
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Chapter 167
GnRH Susan L. Alexander and Clifford H.G. Irvine
The hypothalamus exerts control of reproduction by secreting gonadotropin-releasing hormone (GnRH), which regulates the synthesis and secretion of the gonadotropins, luteinizing hormone (LH), and folliclestimulating hormone (FSH). This discovery revolutionized the clinical management of reproduction in the horse, human, and other species, and applications of this knowledge continue to expand today. The objectives of this chapter are to present current views on the molecular biology and physiology of GnRH, gained largely from work in non-equine species, to spur research in the horse and to enable GnRH to be used more rationally in equine reproductive management. ’
Molecular biology Structure and synthesis Gonadotropin-releasing hormone was first extracted and sequenced from porcine hypothalami in 1971, in a Nobel Prize-winning effort by Andrew Schally and colleagues. It was found to consist of 10 amino acids (pyroGlu-His-Trp-Ser-Tyr-Gly-LeuArg-Pro-Gly·NH2 ).1 Because it induced marked release of LH from the pituitary gland in all species tested it was initially called luteinizing hormonereleasing hormone (LHRH). Subsequently it was found also to induce FSH release in many species including horses,2 and so today the decapeptide is usually termed gonadotropin-releasing hormone. Gonadotropin-releasing hormone is synthesized as part of a larger molecule called prepro-GnRH. The gene encoding prepro-GnRH has been cloned in number of species and is approximately 4300 base pairs with four short exons separated by three large introns. A brief review of the control of GnRH transcription can be found in Clarke and Pompolo.3
Once prepro-GnRH has been synthesized and before secretion, GnRH is split off from the remainder of the molecule, a 56-amino-acid peptide called gonadotropin-releasing hormone-associated peptide (GAP), which is released concurrently with the decapeptide.4 The function of GAP was once a subject of intense research, and still remains unknown. Natural and manmade analogs Although the GnRH isolated from porcine hypothalami was originally thought to be a unique structure, 23 different forms of the decapeptide have been discovered in vertebrate and protochordate species.5 In most vertebrates there are at least two, and usually three, forms of the GnRH decapeptide (reviewed by Millar5 ), which differ in amino acid sequence, gene location, embryonic origins, and distribution within the brain and peripheral tissues. The existence of different forms of GnRH within a given species has suggested to researchers that one variant may be the long sought after FSH-releasing hormone (FSHRH). A specific FSH-RH was predicted by Geoffrey Harris, the father of neuroendocrinology, who proposed that each pituitary hormone would be regulated by its own hypothalamic hormone.6 Moreover, it is clear that the pituitary can secrete the two gonadotropins differentially, as discussed in Chapter 168. The most ubiquitous variant of GnRH was first isolated from chicken brains in 1984,7 and differs from the GnRH described by Schally’s group at amino acids 5, 7, and 8 (His5, Trp7, Tyr8). The structure of chicken GnRH is completely conserved from bony fishes to humans, and so it is believed to be the first form of the hormone to evolve.5 This primordial GnRH has been designated GnRH II, while Schally’s GnRH is called GnRH I. Although GnRH II can stimulate LH secretion in a variety of mammalian species, it has approximately 2% of the potency of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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GnRH GnRH I, and most recent studies have found it not to have selective FSH-releasing activity (reviewed by Kauffman8 ). GnRH II is therefore unlikely to be either a primary regulator of gonadotropin secretion or the FSH-RH. Rather, myriad central and peripheral actions have been proposed for GnRH II in higher mammals, although not yet the horse. These include promotion of sexual behavior in poorly fed females, modification of short-term food intake, and regulation of cell proliferation and hormone secretion in the ovary and placenta in an autocrine/paracrine manner (reviewed by Kauffman8 ). The current favorite for being the FSH-RH is an isoform of GnRH isolated from lampreys,8 now termed GnRH III, which differs from GnRH I at amino acids 5 through 8 (His5, Asp6, Trp7, Lys8).5 GnRH III has been detected in the hypothalamus of rodents, primates,8 and some species of birds.9 It stimulates preferential FSH release in cattle,10 in some11 but not other12 studies in rats and not in chickens.9 The existence and role, if any, of GnRH III in horses awaits investigation. This chapter will focus on GnRH I, unless otherwise noted. The 23 natural variants of GnRH found to date have common sequences at the amino-terminus (pGlu-His-Trp-Ser) and the carboxy-terminus (ProGly NH2 ), which have therefore been conserved for more than 500 million years of evolution.5 This implies that these sequences are critical for receptor binding and activation, a deduction that has been confirmed by structure/activity data from thousands of GnRH analogs developed primarily on an empirical basis.5 These manmade analogs have shed light on the binding requirements of the GnRH receptor, and many have been found to have clinically useful agonist or antagonist properties. The mammalian pituitary GnRH receptor has stringent requirements for ligand conformation (reviewed in Millar5 ). When GnRH binds to its receptor, the conserved amino- and carboxy-terminal domains are closely apposed, with the molecule bending around the flexible glycine in position 6. Substitution with d-amino acids in position six stabilizes the folded conformation, increasing binding affinity and decreasing metabolic clearance. This feature is incorporated in all agonist and antagonist analogs. Removal of the terminal Gly10 with amidation of Pro9 also increases agonist activity.13 Some GnRH agonists are up to 100 times more potent than native GnRH. Examples of GnRH agonists that have been used in horses are buserelin (t-butyl-Ser6)14 and deslorelin (d-Trp6, N-ethyl-l-prolineamide9, deglycineamide10).15 The amino-terminus alone is involved in receptor activation, and substitutions in this region produce antagonists that bind to the GnRH receptor without activating gonadotropin se-
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cretion (see Millar5 ). At appropriate doses these analogs can compete with natural GnRH for its receptors, and thus suppress its action. A GnRH antagonist that is effective in horses is cetrorelix (substitutions of amino acids at positions 1, 2, 3, 6, and 10).16 The GnRH receptor, signal transduction, and desensitization The amino acid sequence of the mammalian GnRH receptor was first deduced by cloning the receptor from a mouse pituitary αT3 gonadotrope cell line.17 Pituitary GnRH receptors were subsequently cloned from other mammalian species, including rats, humans, sheep, cows, pigs, guinea pigs, and horses (reviewed by Rispoli and Nett18 ) and were found to share over 83% amino acid identity. The mammalian GnRH receptor is a member of the G-protein coupled receptor (GPCR) superfamily and is a glycosylated 327 (rodent) or 328 amino acid protein with seven αhelical transmembrane domains connected by three extracellular and three intracellular loops. Through use of spot mutations, the regions of the receptor that are crucial for binding GnRH, for orientating the receptor within the membrane, and for coupling occupation to signal transduction have been characterized (reviewed by Millar5 and Rispoli and Nett18 ). The GnRH receptor resides in specialized microdomains of the plasma membrane.19 When the receptor binds GnRH, its conformation alters and a complex is formed consisting of GnRH, the receptor, a G-protein (generally Gα q/11 ), and possibly other proteins (reviewed by Millar5 and Bliss et al.19 ). This activates a range of intracellular signaling systems, including phospholipase Cβ (with subsequent elaboration of the second messengers, phosphatidylinositol-3-phosphate and diacylglycerol, and thence a sharp rise in intracellular calcium concentrations) and the mitogen-activated protein kinase pathways. A description of the complex series of intracellular events triggered by GnRH is beyond the scope of this chapter. For more information see Call and Wolfe,20 Kraus et al.,21 Hapgood et al.,22 and Bliss et al.19 Activation of these intracellular pathways ultimately regulates the synthesis and secretion of LH and FSH. The density of GnRH receptors on gonadotropes determines their ability to respond to GnRH.23 Receptor density varies during the ovulatory cycle, peaking before ovulation, and is regulated by estradiol, inhibin, activin (see below), and by GnRH itself.18 Some GnRH input is essential to maintain the full complement of GnRH receptors, and increasing GnRH pulse frequency within the physiological range stimulates the expression of the GnRH receptor gene.18
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However, sustained stimulation with high doses of GnRH or its superactive agonist analogs causes a drop in gonadotropin secretion and then a decline in gonadal function. The equine gonadotrope seems to be relatively resistant to this process, as will be discussed below. In other species, the loss of gonadotrope responsiveness to GnRH is associated with decreases in the amounts of mRNA for, and the numbers of, GnRH receptors; a process called downregulation. The decline in receptor numbers appears to occur because receptors are internalized and degraded faster than they can be replaced.18 For most Gprotein coupled receptors the process of internalization is triggered by phosphorylation, often within the carboxy-terminal tail of the receptor. The consequent binding of β-arrestin not only targets the receptor for internalization but also prevents it from activating its G-protein (termed receptor ‘desensitization’).24 However, a unique feature of the mammalian pituitary GnRH receptor is the absence of a carboxy-terminal tail, which is present in all other GPCRs and in all of the non-mammalian GnRH receptors.5, 18, 24 Mammalian GnRH receptors apparently do not undergo agonist-induced phosphorylation or bind β-arrestin. Accordingly, they are resistant to receptor desensitization and internalize slowly. The prompt reduction in gonadotropin secretion that occurs with continuous activation of GnRH receptors is therefore thought to involve additional adaptive responses distal to the receptor (for more detail see McArdle et al.24 ). Degradation After its release, GnRH can be degraded by several membrane-bound and plasma peptidases. Thimet oligopeptidase (EC 3.4.24.15) is the only one of the putative GnRH-metabolizing peptidases to have been sequenced and cloned.25 It is a neutral thiolactivated zinc metallopeptidase that cleaves GnRH between the fifth and sixth amino acids. However, it acts very slowly unless the C-terminal glycinamide has first been removed by a prolylendopeptidase (PEP) (reviewed by Smith et al.25 ). Angiotensinconverting enzyme (ACE) can also cleave GnRH at two sites.26 Although these and other unidentified enzymes can degrade GnRH in vitro and are located at sites of GnRH release, their function in vivo is unclear. Several studies have suggested that the rate of GnRH degradation is affected by the gonadal steroid milieu, raising the possibility that GnRH-metabolizing peptidases play a physiological role in regulating the amount of GnRH reaching gonadotrope receptors (reviewed by Lew et al.26 ). However, when specific inhibitors to thimet oligopeptidase, PEP, or ACE are infused into the third cerebral
ventricle of ovariectomized ewes, the pattern of LH release is unaltered.26 This implies that these enzymes do not have a major influence on the control of GnRH signaling to the pituitary in vivo. Degradation of GnRH is faster at 37◦ C in horse plasma in vitro than in other species tested, with a half-disappearance time of less than 15 min, compared with 240 min in sera from humans and sheep.27 Degradation in vitro can be inhibited by quick chilling and adding bacitracin to a final concentration of 1 mM (authors’ unpublished observations).
Physiology Localization of GnRH neurons In horses,28 as in other mammals, GnRH cell bodies are located in the rostral preoptic nuclei, the organum vasculosum of the lamina terminalis (OVLT), and in a ventral continuum extending from the medial septum to the tuberoinfundibular region that includes the arcuate nucleus. The majority of GnRH cell bodies are concentrated in the medial basal hypothalamus, and most GnRH fibers project to the external capillary plexus of the median eminence. Here they store their secretory product in terminal granules. When the neuron is depolarized, GnRH is discharged and enters fenestrated capillaries for transport via the long portal vessels to the anterior pituitary. In other species it has been shown that some GnRH fibers traveling to the median eminence have branches that terminate in higher centers of the brain and presumably release GnRH simultaneously in both areas.29 Moreover, approximately 20–30% of GnRH neurons project only to higher centers.8 These pathways probably exist in horses too.28 It has been proposed that the GnRH released in these higher brain centers may be involved in mediating sexual behavior,8, 30 although GnRH II may also modulate these behaviors in some circumstances (reviewed by Kauffman8 ). In geldings given subeffective doses of testosterone, injection of GnRH causes a significant increase in copulatory behavior.31 Thus, endocrine and behavioral sexual responses may be coordinated by one GnRH neuronal system, which would be a design of elegant economy.
The GnRH pulse generator In the horse the reproductive axis is driven by a few thousand GnRH neurons.28 In all species studied, including horses, GnRH cells form a loosely connected group scattered through the medial basal hypothalamus and called the GnRH pulse generator.32
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GnRH Each GnRH neuron has the ability to discharge spontaneously, like myocardial, myometrial, and gut muscle cells. Also, like these other cell types, GnRH cells have functional connections that permit a degree of synchronization proportional to the intimacy of connections. In vitro human and rat hypothalami discharge GnRH approximately once to twice per hour.33, 34 This activity, harnessed by other neuronal connections, provides the fundamental driving force to the reproductive axis. Methods for monitoring GnRH secretion Monitoring GnRH secretion patterns has proved challenging because the portal vessels into which GnRH is released lie deep within the cranial cavity and travel directly to the pituitary. Unfortunately, hypothalamic GnRH secretion cannot be detected in peripheral blood because the picomolar concentrations in portal blood are diluted approximately 500-fold to immeasurably low levels in the jugular vein.27 It is therefore necessary to sample blood close to the pituitary. The portal vessels are almost inaccessible; nevertheless surgical techniques for collecting portal blood have been developed in the rat,35 monkey,36 and sheep37 but not the horse. These techniques entail removal of part of the skull, exposure then cutting of the pituitary stalk or incision of the portal vessels, and collection by suction of the blood that leaks from them. The development of the ovine model by Clarke and Cummins37 was a landmark in neuroendocrinology because it allowed portal blood to be collected from conscious animals without major disruption of pituitary function, as well as simultaneous sampling of peripheral blood. The size and thickness of the equine skull make it difficult to apply to the horse the ovine model for collecting portal blood. Nevertheless Sharp and Grubaugh38 have described a roughly similar approach for estimating GnRH secretion in horses through push-pull perfusion of the medial basal hypothalamus. This involves surgical implantation of the perfusion device, but sampling can be done in conscious animals. The authors of this chapter have devised an alternative technique that exploits the unique venous anatomy of equids.39 This method is non-surgical and involves introducing a cannula via percutaneous puncture into the facial vein and advancing it along a venous pathway until its tip reaches the opening of the paired pituitary veins into the intercavernous sinus. The pituitary venous blood collected by a properly placed cannula contains readily measurable concentrations of GnRH and other hypothalamic hormones, as well as very high concentrations of pituitary hormones. This method has some
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advantages, not the least of which is its non-invasive nature. Other advantages are:
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sufficient blood can be collected in 30 s to measure multiple hypothalamic and pituitary hormones; sampling can continue for hours or over several days thus enabling secretion patterns and hormone interactions to be defined with great precision; hormone secretion rates can be calculated; sampling can be done without restraint and under a variety of physiological conditions, e.g. during teasing40 or exercise.41
Secretion patterns of GnRH in mares Experiments using the ovine model showed for the first time that GnRH is secreted in pulses that are concurrent with pulses of LH in peripheral blood.37 This finding is cited as final proof that GnRH is the major endogenous factor stimulating LH secretion.3 In horses as in sheep GnRH secretion is pulsatile, with pulse frequency varying with stage of cycle and season. The most detailed information on GnRH secretion patterns in horses comes from measurements in pituitary venous blood: In the midluteal phase mare GnRH pulses are high in amplitude but low in frequency42 (Figure 167.1). They occur synchronously with large bursts of FSH and LH secretion42 (Figure 167.1). Concentrations of GnRH generally remain immeasurably low between large pulses; however, low-amplitude pulses occur sporadically. Approximately a third of the small GnRH pulses are unaccompanied by gonadotropin pulses and may therefore be artifacts of the assays or the pulse detection algorithm used. However, in luteal phase ewes a similar percentage of GnRH pulses in portal blood are unassociated with peripheral LH pulses and are thought to serve to stimulate gonadotropin synthesis.43 This may be the case in the mare. During the ovulatory LH surge GnRH pulses are high frequency but low amplitude44 (Figure 167.2). They accompany LH and FSH pulses with significant coincidence, and GnRH secretion can be detected in over 75% of samples. The moment-to-moment GnRH secretion pattern appears erratic unless the data are smoothed by a 5-min moving average whereupon a relatively rhythmic pattern emerges (Figure 167.2). The irregularity of GnRH secretion and its detection in the majority of pituitary venous blood samples collected during the surge may be caused by desynchronization of the firing of GnRH neurons under the influence of high plasma estradiol. This would lead to
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Time (minutes) Figure 167.1 Secretion rates of gonadotropin-releasing hormone (GnRH), follicle-stimulating hormone (FSH), and luteinizing hormone (LH) calculated from samples of pituitary venous blood collected continuously and divided into 30-second segments from a typical luteal phase mare. Adapted from Irvine and Alexander.42
fairly continuous secretion, as has been suggested for the ewe.45 These results from pituitary venous blood are qualitatively similar to those obtained from pushpull perfusion of the medial basal hypothalamus, which show GnRH secretion to be irregularly episodic with intervening periods of no secretion in anestrous, transitional, and luteal phase mares. However, GnRH secretion is continuous during estrus.38 Thus the two methods of monitoring GnRH secretion agree that in the horse both continuous and discontinuous modes of GnRH secretion occur.
Figure 167.2 Secretion rates of gonadotropin-releasing hormone (GnRH), follicle-stimulating hormone (FSH), and luteinizing hormone (LH) calculated from samples of pituitary venous blood collected continuously and divided into 30-sec segments during the ovulatory surge of a typical mare. The solid line in the top panel shows the GnRH data smoothed using a 10-point (i.e. 5-minute) moving average. Modified from Irvine and Alexander.44 Reproduced by permission of the Society for Endocrinology.
Secretion rates Secretion rates of GnRH can be calculated from measurements in pituitary blood39, 46 (Figure 167.3). Mean GnRH secretion rates in cyclic mares are less than 100 ng per day, with mean rates at estrus approximately thrice those in the luteal phase (Figure 167.3).44 The increased secretion rate at estrus is due entirely to accelerated pulse frequency, since pulse amplitude is lower than that during the major episodes in the
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Figure 167.3 Geometric mean (± SEM) daily secretion rates of GnRH during estrous (n = 6) and luteal phase (n = 4) mares. Estrous and luteal phase values were compared by Student’s t-test. Adapted from Irvine and Alexander.44
luteal phase. In these latter, secretion rates can briefly exceed 0.5 ng/min (authors’ unpublished observations). Exogenous administration Extrapolation from the log:linear GnRH dose: gonadotropin response curve has shown that a gonadotropin pulse similar to a major luteal phase episode would be induced by intrajugular bolus injection of 7 µg GnRH.47 Assuming rapid distribution in the extracellular space this dose would induce peak GnRH concentrations of 56 pg/mL, which is in the ballpark of those measured in pituitary venous blood during a major luteal phase episode.42 Note that pituitary venous blood, while derived mainly from portal blood is diluted by blood from other sources to an extent depending on the placement of the collecting cannula.46 Pulsatility is an important part of the GnRH signal, with the frequency of the pulses affecting how the gonadotropes respond. For example, numerous experiments have shown that the GnRH pulse frequency is a key factor in regulating the ratio of FSH and LH synthesized and released.48 As will be discussed in Chapter 177, when anestrous mares are given pulses of GnRH to replicate the frequencies observed during an ovulatory cycle, the normal cyclic profiles of FSH and LH are induced, as are folliculogenesis and ovulation.49 However, folliculogenesis, ovulation, and approximately normal cyclic patterns in plasma FSH and LH can be generated in anestrous mares by GnRH doses and administration protocols that are far from physiological, including continuous infusion50 and implants that release potent GnRH agonists constantly over several weeks.14, 49, 51, 52 These observations are surprising because it is almost dogma that sustained stimulation with GnRH
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reduces GnRH receptor numbers thereby lowering the responsiveness of gonadotropes and causing plasma gonadotropin concentrations to fall, as discussed above. However, the dose of GnRH used as well as the mode of its administration may be a factor in producing receptor downregulation and desensitization.53, 54 It should also be noted that endogenous GnRH secretion has been demonstrated to have a low-level continuous component during the ovulatory surge in both ewes and mares. Indeed, ovulation and normal luteal function can be induced in seasonally acyclic ewes by continuous infusion of GnRH at the low dose rates of 125 ng/h55 or 250 ng/h.56 Conversely, plasma LH concentrations can be suppressed and estrous cycles disrupted in cyclic mares by implants that release the GnRH agonist, gosarelin, over 28 days at dose rates several times greater than those that induce ovulation out of season.57 While it is difficult to compare studies in different species, it appears that equine gonadotropes may tolerate higher levels of continuous GnRH stimulation than other gonadotropes.58 A partial explanation for this is that equine gonadotropes have the unique ability to provide GnRH receptors to the cell surface membrane in the face of continuous GnRH stimulation. Furthermore,the equine GnRH receptor is internalized more slowly than those in other species.59 The mechanisms underlying these observations are unknown.
Factors affecting GnRH secretion
Gonadal steroids, neurotransmitters, and neuromodulators The synthesis and secretion of GnRH are regulated by feedback from gonadal steroids (reviewed by Clarke and Pompolo3 ). In mares and ewes progesterone reduces GnRH pulse frequency to that observed in the luteal phase. In ewes withdrawal of progesterone and treatment with estradiol produces first a negative feedback phase in which GnRH secretion progressively declines, with pulse frequency rising but amplitude falling. This is followed by a GnRH surge as in the normal cycle. In mares GnRH secretion has not been monitored after estradiol administration. Gonadotropin measurements suggest that responses in mares may differ from those in ewes as will be discussed in Chapter 168. Gonadotropin-releasing hormone neurons do not express progesterone or androgen receptors, or the estradiol receptor α, which appears to be the subtype most involved in the feedback control of GnRH (reviewed by Clarke and Pompolo3 ). This means that for the most part gonadal steroids must exert their effect on GnRH indirectly via steroid-responsive neurons that synapse with GnRH neurons. This indirect effect
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consists of both stimulatory and inhibitory inputs. The pathways involved and the neurotransmitters used by the steroid-responsive neurons have not been conclusively determined in any species. Most experiments have entailed administering various neurotransmitters or their antagonists and observing the effect on plasma LH concentrations in vivo (as an index of GnRH secretion) or hypothalamic GnRH mRNA levels in vitro. It should be emphasized that results vary with species, according to whether the end point is GnRH synthesis or secretion, and, as might be expected, the steroid status of the subjects. A recent review of this topic can be found in Clarke and Pompolo.3 Neurotransmitters that predominantly inhibit GnRH neurons include gamma-aminobutyric acid (GABA) and the endogenous opioids, while those that predominantly stimulate GnRH neurons include nitric oxide and the aptly named kisspeptin.60 Neurotransmitters that can stimulate or inhibit GnRH neurons depending on the steroid milieu are: noradrenaline, neuropeptide Y (NPY), dopamine, glutamate (i.e. the carboxylate anion of glutamic acid), and serotonin. Data from horses are relatively scant and most work in the area has focused on the endogenous opioids, glutamate, and dopamine. Administration of the broad-acting opioid receptor blocker, naloxone, to luteal phase mares immediately triggers a major episode of GnRH, LH, and FSH secretion,42 indicating that opioids had been restraining the GnRH neurons. Naloxone is less effective in inducing GnRH and gonadotropin release in estrous mares,61 which suggests that reduced opioid input may be a factor in generating an ovulatory LH surge. Endogenous opioids are also involved in the seasonal rhythm of gonadotropin secretion, with the degree of opioid inhibition increasing in the winter in mares61, 62 and stallions.63 Glutamate appears to stimulate GnRH neurons in horses as in other species with the effect depending on the steroid milieu. For example, administration of the glutamate agonist Nmethyl-d,l-aspartic acid (NMA) causes a prompt rise in plasma LH concentrations in intact mares and stallions but not in geldings64 or long-term ovariectomized (OVX) mares.65 Responses also vary during the ovulatory cycle, with the percentage of mares responding to NMA higher in the luteal phase than at estrus.66 The effect of dopamine on GnRH in horses is controversial. Daily treatment of anestrous mares with a dopamine D2 receptor antagonist, either sulpiride67 or domperidone,68 can advance the first ovulation of the season. This suggests that dopamine inhibition plays a role in regulating seasonal breeding in horses as in sheep.69 However, it is not clear in horses whether this dopamine inhibition is exerted at the level of the GnRH neuron. Domperidone, un-
like sulpiride, does not cross the blood–brain barrier, but the OVLT and some parts of the arcuate nucleus (both sites of GnRH cell bodies in horses) as well as the median eminence (where synapses between dopamine and GnRH neurons have been observed in sheep70 ) are outside this barrier.71 Daily sulpiride treatment has been reported to accelerate FSH pulse frequency and elevate mean plasma concentrations of FSH (but not LH).67 Other workers have observed no effect of daily sulpiride treatment on mean plasma FSH or LH in anestrous mares72 or of acute sulpiride injection on plasma LH in stallions either inside or outside the breeding season.63 Daily domperidone treatment has been found not to alter plasma FSH concentrations, despite stimulating folliculogenesis, and to raise plasma LH concentrations but only close to the time of ovulation.68 In contrast to the inconsistency observed in their effects on gonadotropins, both sulpiride and domperidone always elevate plasma prolactin concentrations. Prolactin has been proposed to stimulate follicular development during the vernal transition.73 It has also been suggested that dopamine may act directly at the ovary.74
Environmental influences One of the most exciting areas of current research is into the mechanisms by which the environment can affect GnRH synthesis and secretion, and hence reproduction. Some of the environmental influences being studied are: photoperiod and seasonal breeding, energy balance and nutritional status, and stress. The first of these environmental signals, photoperiod, is the best documented and will be discussed in Chapter 183. The following examples are given to illustrate the complex interactions that are being discovered to occur between endocrine axes and to emphasize that the study of the control of reproduction now extends beyond the hypothalamo–pituitary–gonadal axis. Energy balance and reproduction The close link between bodyweight and fertility has long been recognized in many species, but the neuroendocrine networks comprising the link have only recently begun to be unraveled.75 Leptin and insulin are two of numerous metabolic factors (for information on other factors see Fernandez-Fernandez et al.75 ) that signal nutritional status to the hypothalamus and are being studied in horses in relation to reproduction.76–78 Leptin is a 167-amino-acid protein secreted by white adipose tissue.79 One of its roles appears to be informing the GnRH neurons when body condition is adequate to support reproduction. In horses as in other species greater adiposity
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GnRH is associated with higher plasma leptin concentrations.76 GnRH neurons do not express leptin receptors,80 so raised leptin levels must influence GnRH secretion indirectly. It has been proposed that one way in which leptin acts is by inhibiting an NPYsecreting pathway in the arcuate nucleus, thus freeing the GnRH neurons from an inhibitory (in the absence of estradiol) input and so increasing GnRH secretion.81 Insulin secretion rises after a meal in horses,82 as in other species, and overfed, obese animals can become insulin resistant, with chronically elevated plasma insulin levels.78 Obesity and insulin resistance are associated with reproductive abnormalities, for example polycystic ovary syndrome in women, and longer estrous cycles and continuous reproductive activity through winter in mares.78 In some species, insulin is thought to stimulate GnRH secretion either directly or, like leptin, through inhibiting NPY neurons.81 This may not be the case in horses since induced transient insulin resistance does not alter pulsatile patterns in plasma LH or FSH or overall LH concentrations during the ovulatory surge.77, 78 Rather, insulin may act at the ovary, where it, together with insulinlike growth factor, promotes follicular growth and modulates ovarian hormone secretion (reviewed by Donadeu and Ginther83 and Vick et al.78 ). Stress and reproduction Chronic stress, whether physical, psychological, or nutritional, can interrupt reproductive function in many species including horses.84 All stressors activate the hypothalamo–pituitary–adrenal (HPA) axis, the end result of which is an increase in plasma concentrations of the glucocorticoids, cortisol, and/or corticosterone. Prolonged administration of the synthetic glucocorticoid, dexamethasone, to cyclic mares lowers plasma LH concentrations, disrupts follicular growth, and decreases estrous behavior and the incidence of ovulation.85 The effect of endogenously or exogenously raised cortisol levels on the minuteto-minute gonadotropin secretion patterns has not been studied in horses. In OVX ewes even a brief elevation in plasma cortisol reduces the amplitude of LH pulses but does not alter pulsatile GnRH secretion.86 Cortisol must therefore be acting at the pituitary to lower the responsiveness to GnRH.86 In follicular phase ewes cortisol administration reduces LH pulse frequency, which suggests that in intact animals cortisol may inhibit GnRH secretion possibly via an ovarian steroid-responsive input to the GnRH neurons.86 Stress may also suppress reproduction by increasing the release of corticotropinreleasing hormone (CRH), a hypothalamic hormone that stimulates the pituitary to secrete adrenocorticotropic hormone (ACTH), which then induces the
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adrenal glands to release cortisol. In rats CRH administration inhibits GnRH secretion and acutely lowers plasma LH concentrations.87 The suppression of GnRH is achieved at least in part through a CRHactivated opioidergic pathway.88 . The effect of acute stresses in horses is debatable and would be a fruitful area of research. Several perturbations, for example transportation89 or sexual arousal,90 have been shown to raise plasma levels of both LH and cortisol. Indeed, our observations in pituitary blood suggest that the minute-to-minute secretion patterns of ACTH and LH are usually concurrent, both at rest and during acute stress (authors’ unpublished observations).
References 1. Matsuo H, Baba Y, Nair RMG, Arimura A, Schally AV. Structure of the proposed porcine luteinizing hormoneand follicle-stimulating hormone-releasing hormone. Biochem Biophys Res Commun 1971;43:1334–9. 2. Evans MJ, Irvine CHG. Measurement of equine follicle stimulating hormone and luteinizing hormone: Response of anestrous mares to gonadotropin releasing hormone. Biol Reprod 1976;15:477–84. 3. Clarke IJ, Pompolo S. Synthesis and secretion of GnRH. Anim Reprod Sci 2005;88:29–55. 4. Mason AJ, Nikolics K, Stewart TA, Seeburg PH. The mammalian GnRH gene: a central role in mammalian reproduction. In: Imura H (ed.) Neuroendocrine Control of the Hypothalamo-pituitary System. Tokyo: Karger, 1988; pp. 3–11. 5. Millar RP. GnRHs and GnRH receptors. Anim Reprod Sci 2005;88:5–28. 6. Harris GW. Neural Control of the Pituitary Gland. London: Arnold, 1955. 7. Miyamoto K, Hasegawa Y, Nomura M. Identification of the second gonadotropin-releasing hormone in chicken hypothalamus: evidence that gonadotropin secretion is probably controlled by two distinct gonadotropin-releasing hormones in avian species. P Natl Acad Sci USA 1984;81:3874–8. 8. Kauffman AS. Emerging functions of gonadotropinreleasing hormone II in mammalian physiology and behaviour. J Neuroendocrinol 2004;16:794–806. 9. Proudman JA, Scanes CG, Johannsen SA, Berghman LR, Camp MJ. Comparison of the ability of the three endogenous GnRHs to stimulate release of follicle-stimulating hormone and luteinizing hormone in chickens. Domest Anim Endocrin 2005;31:141–53. 10. Dees WL, Dearth RK, Hooper RN, Brinsko SP, Romano JE, Rahe H, Yu WH, McCann SM. Lamprey gonadotropinreleasing hormone-III selectively releases follicle stimulating hormone in the bovine. Domest Anim Endocrin 2001;20:279–88. 11. Yu WH, Karanth S, Walczewska A, Sower SA, McCann SM. A hypothalamic follicle-stimulating hormone-releasing decapeptide in the rat. P Natl Acad Sci USA 1997;94:9499–503. 12. Kovacs M, Seprodi J, Koppan M, Horvath JE, Vincze B, Teplan I, Flerko B. Lamprey gonadotropin-releasing hormone-III has no selective follicle-stimulating hormonereleasing effect in rats. J Neuroendocrinol 2002;14:647–55. 13. Sandow J. et al. Structure-activity relationships in the LHRH molecule. In: Crighton DB, Haynes NB, Foxcroft GR,
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Lamming GE (eds) Control of Ovulation. London: Butterworth, 1978; pp. 49–70. Harrison LA, Squires EL, Nett TM, McKinnon AO. Use of gonadotropin-releasing hormone for hastening ovulation in transitional mares. J Anim Sci 1990;68:690–9. McKinnon AO, Nobelius AM, del Marmol Figueroa ST, Skidmore J, Vasey JR, Trigg TE. Predictable ovulation in mares treated with an implant of the GnRH analogue deslorelin. Equine Vet J 1993;25:321–3. Evans MJ, Kitson NE, Alexander SL, Irvine CHG, Turner JE, Perkins NR, Livesey JH. Effectiveness of an antagonist to gonadotrophin releasing hormone on the FSH and LH response to GnRH in perfused equine pituitary cells, and in seasonally acyclic mares. Anim Reprod Sci 2002;73:37– 51. Tsutsumi M, Zhou W, Millar RP, Mellon PL, Roberts JL, Flanagan CA, Dong K, Gillo B, Sealfon SC. Cloning and functional expression of a mouse gonadotropin-releasing hormone receptor. Mol Endocrinol 1992;6:1163–9. Rispoli LA, Nett TM. Pituitary gonadotropin-releasing hormone (GnRH) receptor: Structure, distribution and regulation of expression. Anim Reprod Sci 2005;88:57–74. Bliss SP, Navratil AM, Breed M, Skinner DC, Clay CM, Roberson MS. Signaling complexes associated with the type I GnRH receptor: Colocalization of the ERK2 and GnRH receptor within membrane rafts. Mol Endocrinol 2007;21:538–54. Call GB, Wolfe MW. Gonadotropin-releasing hormone activates the equine luteinizing hormone beta promoter through a protein kinase C/mitogen-activated protein kinase pathway. Biol Reprod 1999;61:715–23. Kraus S, Naor Z, Seger R. Intracellular signaling pathways mediated by the gonadotropin-releasing hormone (GnRH) receptor. Arch Med Res 2001;32:499–509. Hapgood JP, Sadie H, van Biljon W, Ronacher K. Regulation of expression of mammalian gonadotrophin-releasing hormone receptor genes. J Neuroendocrinol 2005;17:619–38. Wise ME, Nieman D, Stewart J, Nett TM. Effect of number of receptors for gonadotropin-releasing hormone on the release of luteinizing hormone. Biol Reprod 1984;31:1007–13. McArdle CA, Franklin J, Green L, Hislop JN. The gonadotrophin-releasing hormone receptor: signalling, cycling and desensitisation. Arch Physiol Biochem 2002; 110:113–122. Smith AI, Shrimpton CN, Norman UM, Clarke IJ, Wolfson AJ, Lew RA. Neuropeptidases regulating gonadal function. Biochem Soc Trans 2000;28:430–4. Lew RA, Cowley M, Clarke IJ, Smith AI. Peptidases that degrade gonadotropin-releasing hormone in the ewe. J Neuroendocrinol 1997;9:707–12. Nett TM, Adams TE. Further studies on the radioimmunoassay of gonadotropin-releasing hormone: effect of radioiodination, antiserum and unextracted serum on levels of immunoreactivity in serum. Endocrinology 1977;101:1135–44. Melrose PA, Pickel C, Cheramie HS, Henk WG, LittlefieldChabaud MA, French DD. Distribution and morphology of immunoreactive gonadotropin-releasing hormone (GnRH) neurons in the basal forebrain of ponies. J Comp Neurol 1994;339:268–87. LeBlanc P, Crumeyrolle M, Latouche J, Jordan D, Fillion G, l’Heritier A, Kordon C, Dussaillant M, Rostene W, Haour F. Characterization and distribution of receptors for gonadotropin-releasing hormone in the rat hippocampus. Neuroendocrinology 1988;48:482–8.
30. Moss RL. Actions of hypothalamic hormones on the brain. Ann Rev Physiol 1979;41:617–31. 31. Pozor MA, McDonnell SM, Kenney RM, Tischner M. GnRH facilitates copulatory behavior in geldings treated with testosterone. J Reprod Fertil Suppl 1991;44:666–7. 32. Knobil E. The electrophysiology of the GnRH pulse generator in the rhesus monkey. J Steroid Biochem 1989;33:669–71. 33. Rasmussen DD, Gambacciani M, Swartz W, Tueros VS, Yen SSC. Pulsatile gonadotropin-releasing hormone release from the human mediobasal hypothalamus in vitro: opiate receptor-mediated suppression. Neuroendocrinology 1989;49:150–6. 34. Rasmussen DD. Episodic gonadotropin-releasing hormone release from the rat isolated median eminence in vitro. Neuroendocrinology 1993;58:511–18. 35. Porter JC, Smith KR. Collection of hypophysial stalk blood in rats. Endocrinology 1967;81:1182–7. 36. Carmel PW, Antunes JL, Ferin M. Collection of blood from pituitary stalk and portal veins in monkeys and from the pituitary sinusoidal system of monkeys and man. J Neurosurg 1979;50:75–80. 37. Clarke IJ. Cummins JT. The temporal relationship between gonadotropin-releasing hormone and luteinizing hormone secretion in ovariectomized ewes. Endocrinology 1982;111:1737–9. 38. Sharp DC, Grubaugh WR. Use of push-pull perfusion techniques in studies of gonadotrophin-releasing hormone secretion in mares. J Reprod Fertil Suppl 1987;35:289–96. 39. Irvine CHG, Alexander SL. A novel technique for measuring hypothalamic and pituitary hormone secretion rates from collection of pituitary venous effluent in the normal horse. J Endocrinol 1987;113:183–92. 40. Shand N, Alexander SL, Irvine CHG. Oxytocin secretion patterns in normal stallions as measured in pituitary venous blood: Correlation with gonadotropin secretion and effect of sexual arousal. Biol Reprod Monogr 1995;1:565– 75. 41. Alexander SL, Irvine CHG, Ellis MJ, Donald RA. The effect of acute exercise on the secretion of corticotropin-releasing factor, arginine vasopressin and adrenocorticotropin as measured in pituitary venous blood from normal horses. Endocrinology 1991;128:65–72. 42. Irvine CHG, Alexander SL. Secretory patterns and rates of gonadotropin-releasing hormone, follicle-stimulating hormone, and luteinizing hormone revealed by intensive sample of pituitary venous blood in the luteal phase mare. Endocrinology 1993;132:212–18. 43. Clarke IJ, Cummins JT. The significance of small pulses of GnRH. J Endocrinol 1987;113:413–18. 44. Irvine CHG, Alexander SL. The dynamics of gonadotrophin-releasing hormone, LH and FSH secretion during the spontaneous ovulatory surge of the mare as revealed by intensive sampling of pituitary venous blood. J Endocrinol 1994;140:283–95. 45. Moenter SM, Brand RC, Karsch FJ. Dynamics of gonadotropin-releasing hormone (GnRH) secretion during the GnRH surge: insights into the mechanism of GnRH surge induction. Endocrinology 1992;130:2978–84. 46. Alexander SL, Irvine CHG. Secretion rates and short-term patterns of gonadotrophin-releasing hormone, FSH and LH throughout the periovulatory period in the mare. J Endocrinol 1987;114:351–62. 47. Alexander SL, Irvine CHG. Effect of graded doses of gonadotrophin-releasing hormone on serum LH concentrations in mares in various reproductive states: comparison
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with endogenously generated LH pulses. J Endocrinol 1986;110:19–26. Ferris HA, Shupnik MA. Mechanisms for pulsatile regulation of the gonadotropin subunit genes by GnRH1. Biol Reprod 2006;74:993–8. Turner JE, Irvine CHG. The effect of various gonadotrophin-releasing hormone regimens on gonadotrophins, follicular growth and ovulation in deeply anoestrous mares. J Reprod Fertil Suppl 1991;44:213–22. Ainsworth CGV, Hyland JH. Continuous infusion of gonadotrophin-releasing hormone (GnRH) advances the onset of oestrous cycles in Thoroughbred mares on Australian studfarms. J Reprod Fertil Suppl 1991;44:235–40. Allen WR, Sanderson MW, Greenwood RES, Ellis DR, Crowhurst JS, Simpson DJ, Rossdale PD. Induction of ovulation in anoestrous mares with a slow-release implant of a GnRH analogue (ICI 118 630). J Reprod Fertil Suppl 1987;36:469–78. Fitzgerald BP, Meyer SL, Affleck KJ, Silvia PJ. Effect of constant administration of a gonadotropin-releasing hormone agonist on reproductive activity in mares: Induction of ovulation during seasonal anestrus. Am J Vet Res 1993;54:1735–45. Weiss J, Cote CR, Jameson JL, Crowley WF Jr. Homologous desensitization of gonadotropin-releasing hormone (GnRH)-stimulated luteinizing hormone secretion in vitro occurs within the duration of an endogenous GnRH pulse. Endocrinology 1995;136:138–43. Vizcarra JA, Wettemann RP, Braden TD, Turzillo AM, Nett TM. Effect of gonadotropin-releasing hormone (GnRH) pulse frequency on serum and pituitary concentrations of luteinizing and follicle-stimulating hormone, GnRH receptors, and messenger ribonucleic acid for gonadotropin subunits in cows. Endocrinology 1997;138:594–601. McNatty KP, Hudson NL, Ball K, Forbes S. Treatment of seasonally anestrous Romney ewes with continuous infusion of low doses of GnRH: Effects on estrus, ovulation and plasma progesterone concentration. Theriogenology 1988;30:953–60. Fray MD, Lamming GE, Haresign W. Inter-breed variation in the postpartum ewe response to continuous low dose infusion of GnRH. Theriogenology 1996;45:1047–64. Fitzgerald BP, Peterson KD, Silvia PJ. Effect of constant administration of a gonadotropin-releasing hormone agonist on reproductive activity in mares: Preliminary evidence on suppression of ovulation during the breeding season. Am J Vet Res 1993;54:1746–51. Porter MB, Cleaver BD, Peltier M, Robinson G, Thatcher WW, Sharp DC. Comparative study between pony mares and ewes evaluating gonadotrophic response to administration of gonadotrophin-releasing hormone. J Reprod Fertil 1997;110:219–29. Porter MB, Sharp DC. Gonadotropin-releasing hormone receptor trafficking may explain the relative resistance to pituitary desensitization in mares. Theriogenology 2002;58:523–6. Gottsch ML, Clifton DK, Steiner RA. Kisspeptin-GPR54 signaling in the neuroendocrine reproductive axis. Mol Cell Endocrinol 2006;.254–255:91–6. Turner JE, Irvine CHG, Alexander SL. Regulation of seasonal breeding by endogenous opioids in mares. Biol Reprod Monogr 1995;1:443–8. Davison LA, McManus CJ, Fitzgerald BP. Gonadotropin response to naloxone in the mare: effect of time of year and reproductive status. Biol Reprod 1998;59:1195–9.
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63. Aurich C, Gerlach T, Aurich JE, Hoppen HO, Lange J, Parvizi N. Dopaminergic and opioidergic regulation of gonadotropin and prolactin release in stallions. Reprod Domest Anim 2002;37:335–40. 64. Sticker LS, Thompson DL Jr, Gentry LR. Pituitary hormone and insulin response to infusion of amino acids and N-methyl-d,l-aspartate in horses. J Anim Sci 2001;79:735– 44. 65. Fitzgerald BP. Effects of administration of N-methyld,l-aspartate (NMA) on gonadotropin secretion in untreated and steroid-treated ovariectomized mares during the breeding season and in intact and ovariectomized mares during anestrus. Domest Anim Endocrin 1996;13:211– 18. 66. Fitzgerald BP, Davison LA. Comparison of the effects of n-methyl-D,L-aspartic acid on gonadotropin and prolactin secretion in anestrous mares and mares exhibiting estrous cycles during anestrus. Biol Reprod 1997;57:36–42. 67. Besognet B, Hansen BS, Daels PF. Dopaminergic regulation of gonadotrophin secretion in seasonally anoestrous mares. J Reprod Fertil 1996;108:55–61. 68. Brendemuehl JP, Cross DL. Influence of the dopamine antagonist domperidone on the vernal transition in seasonally anoestrous mares. J Reprod Fertil Suppl 2000;56:185– 93. 69. Havern RI, Whisnant CS, Goodman RL. Hypothalamic sites of catecholamine inhibition of luteinizing hormone in the ewe. Biol Reprod 1991;44:476–82. 70. Kuljis RO, Advis JP. Immunocytochemical and physiological evidence of synapes between dopamine- and luteinizing hormone releasing hormone-containing neurons in the ewe median eminence. Endocrinology 1989;124:1579–81. ¨ uc ¨ ¸ B, Meister B. Protein components of the 71. Norsted E, Gom blood-brain barrier (BBB) in the mediobasal hypothalamus. J Chem Neuroanat 2008;36:107–21. 72. Donadeu FX, Thompson DL Jr. Administration of sulpiride to anovulatory mares in winter: effects on prolactin and gonadotropin concentrations, ovarian activity, ovulation and hair shedding. Theriogenology 2002;57:963–76. 73. Nequin LG, King SS, Johnson AL, Gow GM, FerreiraDias GM. Prolactin may play a role in stimulating the equine ovary during the spring transition. J Equine Vet Sci 1993;13:631–5. 74. King SS, Jones KL, Mullenix BA, Heath DT, Everson KA, Arbogast LA. Evidence for dopaminergic activity in equine follicles and ovarian germinal epithelium. Anim Reprod Sci 2006;94:175–8. 75. Fernandez-Fernandez R, Martini AC, Navarro VM, Castellano JM, Dieguez C, Aguilar E, Pinilla L, Tena-Sempere M. Novel signals for the integration of energy balance and reproduction. Mol Cell Endocrinol 2006;254–255:127–32. 76. Fitzgerald BP, McManus CJ. Photoperiodic versus metabolic signals as determinants of seasonal anestrus in the mare. Biol Reprod 2000;63:335–40. 77. Sessions DR, Reedy SF, Vick MM, Murphy BA, Fitzgerald BP. Development of a model for inducing transient insulin resistance in the mare: Preliminary implications regarding the estrous cycle. J Anim Sci 2004;82:2321–8. 78. Vick MM, Sessions DR, Murphy BA, Kennedy EL, Reedy SE, Fitzgerald BP. Obesity is associated with altered metabolic and reproductive activity in the mare: effects of metformin on insulin sensitivity and reproductive cyclicity. Reprod Fertil Dev 2006;18:609–17. 79. Misra A, Garg A. Leptin, its receptor and obesity. J Invest Med 1996;44:540–8.
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80. Cunningham MJ, Clifton DK, Steiner RA. Leptin’s actions on the reproductive axis: perspectives and mechanisms. Biol Reprod 1999;60:616–20. 81. Gamba M, Pralong FP. Control of GnRH neuronal activity by metabolic factors: The role of leptin and insulin. Mol Cell Endocrinol 2006; 254–255: 133–9. 82. Depew CL, Thompson DL Jr, Fernandez JM, Sticker LS, Burleigh DW. Changes in concentrations of hormones, metabolites, and amino acids in plasma of adult horses relative to overnight feed deprivation followed by a pellet-hay meal fed at noon. J Anim Sci 1994;72:1530–9. 83. Donadeu FX, Ginther OJ. Changes in concentrations of follicular fluid factors during follicle selection in mares. Biol Reprod 2002;66:1111–18. 84. Plotka ED, Eagle TC, Gaulke SJ, Tester JR, Siniff DB. Hematologic and blood chemical characteristics of feral horses from three management areas. J Wildlife Dis 1988;24:231– 39. 85. Asa CS, Ginther OJ. Glucocorticoid suppression of oestrus, follicles, LH and ovulation in the mare. J Reprod Fertil Suppl 1982;32:247–51.
86. Breen KM, Karsch FJ. New insights regarding glucocorticoids, stress and gonadotropin suppression. Front Neuroendocrin 2006;27:233–45. 87. Petraglia F, Sutton S, Vale W, Plotsky P. Corticotropinreleasing factor decreases plasma luteinizing hormone levels in female rats by inhibiting gonadotropin-releasing hormone release into the hypothalamic-portal circulation. Endocrinology 1987;120:1083–8. 88. Almeida OF, Nikolarakis KE, Herz A. Evidence for the involvement of endogenous opioids in the inhibition of luteinizing hormone by corticotropin-releasing factor. Endocrinology 1988;122:1034–41. 89. Baucus KL, Squires EL, Ralston SL, McKinnon AO, Nett TM. Effect of transportation on the estrous cycle and concentrations of hormones in mares. J Anim Sci 1990;68:419–26. 90. Rabb MH, Thompson DL Jr, Barry BE, Colborn DR, Garza F Jr, Hehnke KE. Effect of sexual stimulation with and without ejaculation on serum concentrations of LH, FSH, testosterone, cortisol and prolactin in stallions. J Anim Sci 1989;67:2724–9.
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Chapter 168
FSH and LH Susan L. Alexander and Clifford H.G. Irvine
The gonadotropins, follicle-stimulating hormone (FSH) and luteinizing hormone (LH), are considered together in this chapter, because they are structurally similar, are regulated similarly, and work in concert to achieve fertility in both males and females. These hormones are products of a single cell type, the gonadotrope, found in the pars distalis and pars tuberalis of the pituitary gland. Gonadotropins, and FSH in particular, have multiple actions on the gonads, at different stages stimulating cell proliferation, growth, differentiation, hormone synthesis and secretion, and ovulation. Coordination of the events that control follicle growth, selection, and subsequent ovulation at the correct time of the year is crucial for the wellbeing of the foal and the survival of the species. This depends on the precise regulation of FSH secretion to control follicle numbers and of LH secretion to time ovulation when ova are mature and conditions favorable. This chapter will consider the unique chemistry of the equine gonadotropins and the factors involved in regulating their secretion.
Chemistry Structure The gonadotropins belong to a glycoprotein hormone family that also includes thyroid-stimulating hormone (TSH) and equine chorionic gonadotropin (eCG). They consist of two dissimilar subunits, called α and β, which are non-covalently associated. The αsubunit is common to all members of the family and is encoded by a single gene. The β-subunits are specific for each hormone and are products of different genes with the exception of the β-subunits for LH and eCG, which have identical amino acid sequences and are encoded by the same gene.1 Both subunits are involved in the binding of the hormone to its receptor.2 Equine FSH and LH have been isolated, purified, and sequenced. Early studies showed that the two
hormones have approximately the same molecular weight (i.e., 34 000 kDa).3 More recently, the cDNA has been cloned for the common α-subunit,4 the LH/eCG β-subunit,5 and the FSH β-subunit.6 Considerable work has also been done to characterize the carbohydrate side-chains (e.g., refs 7 & 8). The αsubunit is composed of 96 amino acids and has two complex-type N-linked carbohydrate chains located at asparagines, Asn56 and Asn82. The FSH β-subunit consists of 111 amino acids with N-linked glycosylation sites at Asn7 and Asn24. Unlike the human FSH β-subunit, which is glycosylated in an all-ornone manner, the equine FSH β-subunit can be glycosylated either at Asn24 alone or at both potential sites.7 In both subunits, the carbohydrate chains are mainly biantennary and can end in either sialic acid, sulfate, or one of each.8 Equine LH is unique amongst mammalian LHs in several respects. Firstly, the β-subunit of equine LH is composed of 149 amino acids, whereas those of most mammalian LHs consist of 121 amino acids.7 Equine LH is extended at the C-terminal by 28 amino acids. It had previously been thought that such C-terminal extensions were restricted to chorionic gonadotropins.9 The C-terminal extension of equine LH is the consequence of the fact that the same gene encodes both it and eCG, as discussed above. In primates separate genes encode LH and CG,10 indicating that primate CG evolved differently to equine CG.7 Secondly, equine LH contains more carbohydrate than other LHs. It is approximately 30% carbohydrate by weight.11 The equine LH β-subunit has one N-linked carbohydrate at Asn13, but the majority of the carbohydrate side-chains are attached to the C-terminal extension. Twelve of the 28 C-terminal amino acids are O-glycosylated.12 While some of the carbohydrate chains terminate in sialic acid,7 more than 72% of them are sulfated.13 The additional carbohydrate residues on equine LH make it more acidic, retard its electrophoretic mobility,7 and lengthen its
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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circulatory half-life (5 h compared with 25 min for ovine LH14 ) when compared with other LHs. Note that equine CG is even more heavily glycosylated and sialylated than is equine LH.15 Thirdly, equine LH (like eCG) has FSH activity in other species, although not in the horse.16 This anomaly has been pinpointed to a couple of amino acid residues in the β-subunit and a single amino acid residue in the α-subunit.2 How these specific substitutions induce FSH bioactivity is still unknown. Nevertheless, the anomaly provides a valuable clue to the relationship between gonadotropin structure and function, which is one of the reasons that the equine gonadotropins are the most studied aspect of equine reproductive endocrinology. Polymorphism and biological activity The carbohydrate side-chains on the gonadotropins occupy a significant portion of the molecules. For example, the dimensions of the protein skeleton of ˚ human chorionic gonadotropin are 75 × 35 × 30 A, while the length of a typical N-linked carbohy˚ (reviewed by Dias17 ). It is drate side-chain is 30 A therefore not surprising that the composition of the carbohydrate side-chains, particularly the terminal negatively charged sialic acid and sulfate residues, greatly affects the functional properties of the gonadotropins, including: subunit association, receptor binding, antigenicity, signal transduction, circulatory half-life, and the final biological response of the target cell (reviewed by Dalpathado et al.8 ). However, unlike the amino acid sequences, which are constant within a species, the carbohydrate chains assembled at each glycosylation site can be highly variable (see, e.g., ref. 8). Thus, each equine gonadotropin exists in the pituitary as a family of molecular forms with the same amino acid skeleton but different carbohydrate side-chains. These isoforms can be separated on the basis of charge,14 and early studies showed them to differ in relative receptor binding and in vitro biological potencies.14, 18, 19 Gonadotropin polymorphism might be of purely academic interest if multiple isoforms were found only in the pituitary and merely represented various stages in the synthetic process, with just one isoform being secreted. However, there is abundant evidence in many species, including horses, that both FSH and LH circulate in multiple forms. Furthermore, the relative distribution of isoforms in circulation as well as in the pituitary is altered by the prevailing gonadal steroid milieu or by gonadotropin-releasing hormone (GnRH) stimulation (reviewed by Ulloa-Aguirre et al.20 ). Likewise, in horses the forms of circulating LH21, 22 and FSH23 vary during the ovulatory cycle. In other species pituitary and circulating gonadotropins are less acidic
when plasma estrogen levels are high. These forms show greater bioactivity in vitro and have a shorter metabolic half-life in vivo than do acidic isoforms. One mechanism by which estrogens can alter the charge distribution of gonadotropin isoforms is by reducing the mRNA expression of sialyltransferase, an enzyme involved in attaching terminal sialic acid residues to the carbohydrate side-chains.24 The situation may be slightly different in mares in the high-estrogen state of estrus, in which less acidic isoforms of LH (but not FSH) appear to be selected for secretion; i.e., secreted LH is much less acidic than the bulk of the hormone found in the pituitary.25 The selection of the less acidic LH isoforms occurs at the time of secretion, as shown by subjecting pituitary venous effluent to chromatofocusing, which separates proteins primarily on the basis of charge.25 These observations suggest that the gonadotropic signal can vary in quality as well as quantity, and that the polymorphism of LH and FSH has physiological relevance. Indeed, some cases of infertility in humans (reviewed by Ulloa-Aguirre et al.20 and Bergendah and Veldhuis26 ) and subfertility in horses (Whitcomb et al.27 ) are associated with the circulation of aberrant gonadotropin isoforms. In fact, some isoforms have been shown to act as antagonists, binding to the receptor without activating it.27, 28 However, the full physiological significance of gonadotropin polymorphism remains to be elucidated. As mentioned above, both gonadotropins have several physiological roles, and it is possible that the various isoforms subserve different functions. One way that this might be accomplished is if the gonadotropin isoforms acted through correspondingly variant gonadotropin receptor populations (reviewed by UlloaAguirre et al.20 ). Testing these hypotheses requires new analytical tools. For example, pure preparations of each major circulating isoform are needed,8 as well as better functional assays in which to evaluate their activity.20 It is especially important that the bioactivity of equine gonadotropins be assessed in equine tissues rather than the rodent tissues used in many studies, including our own.21 This is because equine gonadotropin receptors have a unique structure that affects ligand–receptor interactions, as will be discussed in the next section. Receptors Gonadotropin receptors are expressed primarily in the gonads. The LH receptor is found on Leydig cells in the testis, and thecal, granulosa, and luteal cells in the ovary. The FSH receptor is found on Sertoli cells in the testis, and granulosa cells in the ovary. Both receptors belong to a subfamily of the G-proteinassociated seven-transmembrane-domain receptors. They have a large concave extracellular domain and
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FSH and LH a short cytoplasmic tail (for more details see Lin et al.29 ). Most importantly, the receptors, like the gonadotropins, are glycoproteins. The equine LH receptor has been cloned, and the mature protein consists of 672 amino acids.30 The transmembrane domain is composed of 266 amino acids, including seven hydrophobic segments potentially spanning the cytoplasmic membrane, followed by a 71-amino-acid carboxy-terminal intracellular domain.30 The equine LH receptor shows approximately 90% amino acid homology with LH receptors of other species; however, the equine receptor has seven N-glycosylation sites, whereas those of other species have six.30 The equine LH receptor binds eCG with 2–4% of the affinity of equine LH, while the porcine LH receptor binds both hormones with equal affinity. This difference between porcine and equine receptors has been attributed to the extra carbohydrate side-chain on the equine receptor.30 The equine FSH receptor has been cloned, and the mature protein consists of 677 amino acids.31 It shows approximately 90% homology with other mammalian FSH receptors, but has 10 unique amino acid replacements in the extracellular domain and a unique fourth N-glycosylation site.31 These slight structural differences seem to confer on the equine FSH receptor a high degree of specificity, since it alone of all mammalian FSH receptors studied does not bind eCG.32 How the gonadotropins interact with their receptors and the way that the carbohydrate side-chains on both hormone and receptor influence this process are active topics of research but beyond the scope of this chapter. For some recent work see Bousfield et al.33 and Lin et al.29 An intriguing aspect of gonadotropin receptors is that they, like the gonadotropins, can exist in several forms. Receptor polymorphism can be genetic, arising from mutations such as single nucleotide polymorphisms (SNPs).34 Such SNPs have been discovered in the human FSH receptor gene, meaning that the nucleotide sequence can differ between individuals. The resultant slight change in amino acid sequence appears to affect primarily the function of the receptor, influencing a woman’s sensitivity to FSH during the menstrual cycle as well as cycle length (reviewed by Gromoli and Simoni34 ). It has therefore been recommended that the nucleotide sequence of the FSH receptor gene be checked in women undergoing assisted reproduction techniques.34 Receptor polymorphism can also arise from alternate splicing of the gene, which leads to the synthesis of forms of the receptor differing in size and/or amino acid sequence within an individual.35 For example, four subtypes of the FSH receptor have been identified in gonadal tissue from sheep and mice.35 Each subtype has the structural potential to mediate a different aspect of FSH function, and the relative prevalence
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of subtypes may vary during folliculogenesis.35 This would provide a previously unsuspected mechanism for modulating the gonadal response to gonadotropic stimulation. The extent to which gonadotropin receptor polymorphism occurs in horses is not known and would be an extremely interesting area of research.
Physiology Patterns throughout the ovulatory cycle of the mare In the early 1970s radioimmunoassays were developed to measure equine LH36 and FSH,37 which were then used to determine the profiles of these hormones during the ovulatory cycle, using samples of jugular blood collected once daily or thrice weekly. For LH, concentrations are low during the midluteal phase, but rise in a prolonged ovulatory surge beginning a few days before the onset of estrus, peaking usually on the day after ovulation and returning over several days to mid-luteal phase values (Figure 168.1). Thus, the ovulatory LH surge can last well over a week, which is uniquely long compared with other studied mammal species. For FSH, several different profiles have been reported during the cycle. All studies agree that FSH concentrations reach a nadir in early estrus and are high at some point during the luteal phase. The original observation was that FSH has a biphasic pattern during the cycle, with surges at 10- to 12-day intervals.37 One surge occurs just after ovulation, the second during the mid-luteal phase approximately 10 days before the next ovulation (Figure 168.1). While some subsequent studies have also found two FSH surges per cycle, others have observed one or three surges 25 20
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Figure 168.1 Schematic representation of concentrations of luteinizing hormone (LH; solid line) and follicle-stimulating hormone (FSH; dashed line) in jugular blood samples collected daily during the ovulatory cycle of the mare. The day of ovulation is marked by an arrow. The duration of behavioral estrus is shown by the horizontal bar. Adapted from Evans and Irvine.37
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(reviewed by Ginther38 ). The variability in reported FSH profiles has been attributed to various factors, including season, age, breed, and individual (reviewed by Ginther38 ). Another possibility is that some or all of the variability is an artifact, caused by the mode of gonadotropin secretion and inadequate sampling frequency.39 Early workers assumed that the gonadotropins were secreted continuously so that infrequent blood samples would be adequate to define the pituitary signal to the gonads. However, we now know that in horses as in other species the gonadotropins are secreted episodically. These episodes or pulses reflect the mode of GnRH secretion, as discussed in Chapter 167. Pulse frequency is slow in the luteal phase,40, 41 seasonal acyclicity and the transitional period into breeding season.42, 43 In these states, synchronous LH and FSH pulses can be detected in jugular blood and occur fewer-than-one to four times daily (Figure 168.2). When blood is collected close to the pituitary,44 these large gonadotropin pulses are greatly amplified Pituitary venous blood 150
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Figure 168.2 Concentrations of luteinizing hormone (LH; filled squares) and follicle-stimulating hormone (FSH; open circles) in pituitary venous blood collected every 5 min (upper panel) and jugular blood collected every 15 min (lower panel) from a luteal phase mare 12 days after ovulation. Note the 10-fold difference in the y-axis scale between panels.
(Figure 168.2), and when pituitary blood is sampled every 30 seconds the gonadotropin pulses can be seen to be the summation of a train of 4–6 secretory bursts of decreasing amplitude41 (see Figure 167.1). The total duration of secretion in one large pulse is approximately 1 hour. Overall, LH and FSH secretion patterns are closely correlated, but the amplitude of the slow luteal phase FSH pulses far exceeds that of the LH pulses (Figure 168.2). This means that once-a-day sampling is particularly misleading for determining FSH profiles.39 This became evident when six mares were bled every 4 hours throughout an ovulatory cycle (Figure 168.3). All mares studied had large, slow FSH pulses during the luteal phase but different profiles could be obtained when each day’s value was determined from samples collected at only one time point, thus simulating once-a-day sampling. No mare had a ‘surge’ of FSH; however, FSH pulse amplitude (but not frequency) tended to wax and wane. A period of higher-amplitude FSH pulses preceded ovulation by approximately 10 days.39 This timing is significant. In mares follicles can develop up to 1 cm in diameter without pituitary hormones. However, further development to 2.5 to 3 cm in diameter requires FSH. The ovulatory follicle emerges in a cohort of developing follicles midway in the luteal phase (reviewed by Ginther et al.45 ). Its emergence coincides with elevated plasma FSH concentrations, but the selective development of the dominant ovulatory follicle begins as FSH concentrations start to decline about 8 days before ovulation (reviewed by Ginther et al.45 ). Increased plasma FSH is necessary to initiate this process, since if mid-luteal phase concentrations are prematurely lowered to estrous nadir values an ovulatory follicle does not develop. Once the block on FSH secretion is removed follicular development resumes and ovulation occurs in approximately 10 days.46 While daily sampling suggested that the emergence of the dominant follicle was linked to a surge of FSH spanning several days (e.g., Bergfelt and Ginther47 ), it now appears that the actual FSH signal is 2–8 days of rhythmic, high-amplitude pulses.39 This indicates that high FSH concentrations need be attained only intermittently to develop an ovulatory follicle. The fall in FSH concentrations as estrus approaches is important because it restricts the number of ovulatory follicles selected. Preventing this decline by administering FSH results in the development of multiple pre-ovulatory-sized follicles.48 As plasma FSH concentrations decrease the pulses detected with 4-hourly blood sampling eventually disappear.39 Likewise for LH, which at the same time is rising in the ovulatory surge, pulses cannot be convincingly demonstrated in jugular blood,39, 49
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Figure 168.3 Jugular venous follicle-stimulating hormone (FSH) and luteinizing hormone (LH) concentrations in two mares (a,b) bled at 4-h intervals during an ovulatory cycle. Squares mark the maxima of pulses detected by the Cluster program. The heavy solid line shows the daily mean concentration of each hormone. The day of ovulation is marked by an arrow, and the duration of behavioral estrus is shown by the solid bar. Note that the profile of daily mean FSH concentrations of the mare in (a) is similar to that originally reported by Evans and Irvine,37 while the profile of the mare in (b) is different. Reproduced from Irvine et al.39 with permission of the Society for Reproduction and Fertility.
even when sampled at 5-minute intervals50 (Figure 168.4). The apparent stability of jugular gonadotropin concentrations during estrus means that once-a-day sampling is sufficient to determine FSH and LH profiles at this time. However, the failure to detect gonadotropin pulses in jugular blood during the ovulatory surge does not mean that secretion is not pulsatile. Collection of blood from close to the pituitary during the ovulatory surge clearly shows that FSH and LH are, in fact, secreted in concurrent pulses51, 52 (see Figure 168.4). When pituitary blood is sampled every 30 seconds, the FSH and LH pulses can be seen to be brief (5–7 min) and rapid (two to five per hour) with frequency tending to increase as ovulation approaches52 (see Figure 167.2). Secretion can be detected during 72% (LH) or 43% (FSH) of the total sampling period. The most likely reason that pulses cannot be detected in jugular blood is that the long circulatory half-life of the equine gonadotropins maintains a large peripheral pool when secretion rate is rapid, which serves to dampen individual pulses.
Control of secretion The secretion of both FSH and LH is stimulated by gonadotropin-releasing hormone (GnRH) from the hypothalamus (see Chapter 167). This is shown by the fact that GnRH administration increases plasma concentrations of FSH and LH in horses,53 and the observation of a tight synchrony between GnRH, FSH, and LH secretion patterns in pituitary venous blood from mares in the luteal phase41 and during the ovulatory surge.52 However, active immunization of intact mares against GnRH reduces plasma LH (but not FSH) concentrations to undetectably low levels.54 Likewise, administration of GnRH antagonists to estrous mares decreases plasma LH dramatically without affecting FSH.55 In luteal phase mares, treatment over several days with the GnRH antagonist, cetrorelix, does lower plasma FSH but not to the assay’s detection limit.46 These observations imply that some FSH secretion may occur independently of GnRH stimulation. In sheep, a substantial component of FSH secretion is
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pulses is slightly but not significantly greater than that of LH pulses (86% vs 75%; p = 0.07 Fisher’s exact test;41 S.L. Alexander and C.H.G Irvine, unpublished observations). Thus there is no preferential dissociation of FSH from GnRH pulses that would be suggestive of the presence of an FSH releasing factor. However, non-pulsatile FSH and LH secretion continues at a low level during approximately 50% of the period between large pulses in luteal phase mares.41 This gonadotropin secretion appears to be unrelated to GnRH, although this may be a function of the sensitivity of the GnRH assay. Alternatively, it may indicate that there is a small constitutive component to the secretion of both FSH and LH in the horse.
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The ovulatory cycle of the mare is characterized by periods of differential FSH and LH secretion (Figure 168.1). In the luteal phase, FSH secretion predominates. During the ovulatory surge, LH secretion prevails. If there is no FSH releasing factor, how can one gonadotropin-releasing hormone regulate the differential secretion observed during the cycle? One answer to this riddle is that the frequency of GnRH pulses can signal to the gonadotropes to synthesize and secrete FSH or LH preferentially. This effect is independent of ovarian feedback. Slow GnRH pulses, as occur during the luteal phase, favor FSH release59 and transcription of the FSH β-subunit (reviewed by Burger et al.60 ). Fast GnRH pulses, as occur during the ovulatory surge, favor LH release59 and transcription of the LH β-subunit (reviewed by Burger et al.60 ). This phenomenon can be demonstrated in seasonally acyclic mares, in which the ovaries are small and inactive, and gonadotropin pulse frequency is profoundly suppressed. Under these conditions, administering GnRH pulses every 45 minutes causes predominantly LH secretion, whereas GnRH pulses given every 6 hours increase FSH secretion.61
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constitutive and does not require any hypothalamic input. In addition, sampling close to the pituitary shows that approximately one-third of FSH pulses (but no LH pulses) in ovariectomized sheep are unaccompanied by a GnRH pulse (reviewed by Padmanabhan and McNeilly56 ). Moreover, FSH but not LH pulsatility persists during treatment with a GnRH antagonist.57 While GnRH-less pulses of FSH may arise from waxing and waning of local pituitary and peripheral feedback factors (see later), they may also be stimulated by a specific hypothalamic FSH-releasing factor (reviewed by Padmanabhan and McNeilly56 ). The existence of such a factor has been postulated for decades (e.g., Igarashi and McCann58 ). More recent work proposes that one of the naturally occurring variants of the GnRH decapeptide may preferentially release FSH (see Chapter 167). However, to date no factor has been proven to be a physiological FSH releasing factor. In pituitary venous blood from luteal phase mares, the percentage of FSH pulses associated with GnRH
Regulation by ovarian hormone feedback Changes in GnRH pulse frequency are not the only means by which LH and FSH may be differentially regulated. The ovaries produce both steroid and protein hormones; the most widely studied of these in mares are the steroids, estradiol and progesterone, and the protein hormone, inhibin. These hormones act at the pituitary and/or hypothalamus to affect FSH and LH secretion, often in differential ways. They will be discussed in more detail in subsequent chapters.
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FSH and LH Ovarian proteins Growing follicles synthesize the proteins inhibin, activin, and follistatin, which are involved in regulating gonadotropin, particularly FSH, secretion. Inhibin and activin are structurally related members of the transforming growth factor β superfamily. They are dimers, consisting of two of three possible subunits: inhibin α, βA, or βB. Inhibin pairs an α-subunit with a β-subunit, so that two types of inhibin exist: inhibin A (α βA) and inhibin B (α βB). Activin pairs two β-subunits, so that three types of activin exist: activin A (βA βA), activin AB (βA βB), and activin B (βB βB). Follistatin is a protein that binds the βsubunits. Inhibin acts at the pituitary to suppress FSH secretion, an effect that is independent of GnRH input.62 By contrast, activin stimulates FSH production and secretion (reviewed by Padmanabhan et al.63 ). Follistatin interferes with the binding of activin to its receptor and so inactivates it (reviewed by Padmanabhan et al.63 ). It has a lesser effect on inhibin64 so that follistatin predominantly inhibits FSH. While the major action of these proteins is on FSH, they may also have a minor positive influence on LH secretion, particularly when GnRH input from the hypothalamus is low, through modulating GnRH receptor numbers,65 and in the case of activin increasing GnRH release.66 Only inhibin has been proven to be an endocrine signal from the ovary to the pituitary. In mares there is a reciprocal relationship during the cycle between plasma concentrations of FSH and immunoactive inhibin.67 This profile is also seen when inhibin A68 or inhibin pro-αC, a precursor protein of the inhibin α-subunit,69 are specifically measured. Immunization, whether passive70 or active immunization against the inhibin α-subunit,71–73 raises plasma FSH,70, 73 and increases follicle numbers73 or ovulation rates in mares.70–72 Likewise, administration of a proteinaceous steroid-free fraction of follicular fluid lowers FSH,74 an effect that is enhanced by coadministration of estradiol (E2).75 However, during the normal cycle, it appears that the suppression of circulating FSH at the beginning of the deviation of the ovulatory follicle is a function of circulating inhibin, with no indication that changes in plasma E2 are also required.76 A unique feature of the plasma inhibin profile in mares is that there is a sharp spike in concentration on the day of ovulation, whether inhibin A,68 total immunoactive inhibin, or pro-αC69 are measured. This spike is thought to be due to the release of follicular fluid into the peritoneal cavity upon ovulation and the subsequent adsorption of inhibin into the bloodstream.77 The inhibins, activins, and follistatins that are present in follicular fluid from mares are thought to play a role in follicle maturation and steroidogenesis (reviewed by Ginther et al.78 and
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Beg and Ginther79 ). In other species they have also been found in tissues outside the ovary including the pituitary itself, where they may act in an autocrine and paracrine manner to regulate FSH synthesis and secretion (reviewed by Padmanabhan et al.63 ).
Ovarian steroids Progesterone is produced by the corpus luteum. Its plasma concentrations roughly parallel those of FSH, being high during the luteal phase and low during estrus. In sheep progesterone has been shown to act at the hypothalamus to slow GnRH pulse frequency and does not affect the responsiveness of the pituitary to GnRH stimulation.80 However, in rats progesterone plays a role in the phenomenon of GnRH self-priming, in which the LH response to the second of two successive identical GnRH pulses is greater compared with the response to the first pulse.81 Self-priming occurs within the gonadotrope, depends on estradiol being present, and involves indirect activation of the progesterone receptor by GnRH.81 In mares, as in sheep, lowering plasma progesterone by lysing the corpus luteum increases the frequency of FSH and LH pulses.82, 83 The effect seems to be titrated rather than all-ornone, since gonadotropin pulse frequency rises progressively as progesterone levels fall.83 In ovariectomized (ovx) mares, progesterone administration, or combined treatment with progesterone and estradiol, lowers LH levels, as assessed by daily blood samples, in the breeding but not in the non-breeding season.84 The seasonal difference in response may stem from the fact that gonadotropin pulse frequency in ovx mares is much higher in the breeding (approximately 13 pulses per day) than the non-breeding (approximately 1 pulse per day) season.85 Obviously, the effect of progesterone-induced slowing of gonadotropin pulses would be expected to be more marked when the pretreatment frequency is rapid. In contrast to LH, FSH concentrations tend to be elevated by progesterone or progestagen treatment of ovx mares in the breeding season,84 or of intact mares in shallow acyclicity.43, 86 This is most likely due to the fact that slow GnRH pulse frequencies favor FSH secretion over LH as discussed above. Estradiol (E2) is produced by maturing follicles and possibly the corpus luteum (reviewed by Ginther et al.45 ). Its plasma concentrations roughly parallel those of LH, being low in the luteal phase and rising during estrus to a peak approximately 2 days before ovulation. In other species, the effect of E2 on LH secretion is complex and can be negative or positive depending on the concentration attained and the duration of exposure. It is believed that E2 acts at the hypothalamus to alter GnRH secretion. In sheep
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withdrawal of progesterone followed by E2 administration firstly causes a progressive decline in GnRH secretion. This lull is then followed by a sustained burst of GnRH secretion that drives the massive increase in LH during the ovine ovulatory surge (reviewed by Clarke and Pompolo87 ). Estradiol also acts directly at the pituitary in multiple ways to influence the responsiveness to GnRH (reviewed by Padmanabhan et al.63 and Nett et al.88 ). In sheep and monkeys deprived of hypothalamic input but with LH secretion sustained by pulsatile GnRH replacement, the initial response to E2 administration is a fall in plasma LH levels (reviewed by Clarke89 ). However, estradiol can also enhance the LH response to GnRH. One way that E2 does this is by increasing the synthesis and insertion of GnRH receptors into gonadotrope membranes.88 As discussed in Chapter 167, the response of gonadotropes to GnRH is directly related to the number of GnRH receptors on the cell surface. Since the GnRH receptor gene does not appear to have an estrogen response element (ERE), this effect is presumed to be mediated by an as yet unidentified factor. Interestingly, the first detectable increase in GnRH receptor mRNA after luteolysis in ewes occurs before the pre-ovulatory rise in E2 and is now thought to be stimulated by the increase in GnRH pulse frequency as progesterone falls (reviewed by Nett et al.88 ). This physiological increase in GnRH input induces the synthesis of its own receptor (see Chapter 167). Another way that E2 affects gonadotrope responsiveness is by altering FSH and LH synthesis but in opposite ways: increasing LH, decreasing FSH. The effect on LH is thought to be direct since there is an ERE upstream of the coding region in the rat LH β-subunit gene.90 The effect on FSH is probably indirect since the FSH β-subunit gene lacks an ERE, at least in sheep, and may involve local inhibition of the transcription of the activin βB subunit. The resultant drop in activin would lead to a reduction in FSH synthesis (reviewed by Nett et al.88 ). Estradiol also influences the heterogeneity of both FSH and LH, promoting the production of more alkaline, more biologically active isoforms, as discussed above. Lastly, E2 regulates the second messenger systems in the gonadotrope as well as the function of ion channels in the plasma membrane.91 What is known about the effect of E2 on gonadotropins in mares? Long-term administration of E2 to ovx mares raises LH for the duration of treatment in both the breeding92 and non-breeding seasons.84 Likewise, more recent work using periods of 15-minute blood sampling found that E2 elevates plasma LH, with no evidence for initial suppression, when given in the breeding season to ovx pony mares.93 The E2 effect is dose-dependent, with the largest E2 dose used replicating estrous levels and in-
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Figure 168.5 Geometric mean (± SEM) luteinizing hormone (LH) responsiveness to endogenous gonadotropin-releasing hormone (GnRH) during estrus (n = 6) and the luteal phase (n = 4). Responsiveness was estimated as the mean ratio between newly secreted LH (i.e., the difference between LH concentrations in pituitary venous and jugular blood) and pituitary venous GnRH concentrations. Estrous and luteal phase values were compared by Student’s t-test. Adapted from Irvine and Alexander.52
ducing a fivefold increase in plasma LH concentrations,93 which is similar to the increment observed in some mares during a spontaneous ovulatory surge (e.g., Figure 168.3b). In contrast, studies using follicle ablation and ovariectomy suggest that the ovaries only inhibit LH and FSH secretion; i.e., no positive influence could be demonstrated at any time during the cycle.45 How these conflicting results can be reconciled is unclear. With respect to plasma FSH, E2 administration to ovx mares either has no effect84 or lowers plasma FSH.92, 94 In mares there is a marked rise in LH responsiveness to endogenous GnRH during the spontaneous ovulatory surge (Figure 168.5).52 This may be due at least in part to increased E2 levels, since many (but not all: see Greaves et al.93 ) studies show that E2 administration augments the LH response to GnRH in intact estrous, acyclic, or ovx mares (reviewed by Baldwin et al.95 ) and in equine pituitary cells in vitro.95 However, this effect may not be due to increased GnRH receptors as in other species since receptor concentrations do not differ in equine pituitaries collected during anestrus, early or late vernal transition, or estrus.96, 97 Rather E2 may act by stimulating LH synthesis. It has been shown in horses that the amount of LH released in response to GnRH is a function of the amount of LH in the pituitary.98 Estradiol treatment increases the LH content of the pituitary.92, 99 Studies in ovx, pituitary stalk-sectioned mares indicate that E2 treatment maintains the expression of the α-subunit but not the LH β-subunit gene.100 Thus, the mare differs from other species in which E2 stimulates transcription of the LH β-subunit gene. In mares, again unlike other species, E2 treatment increases92 or tends to increase99 the
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FSH and LH FSH content of the pituitary, which is consistent with an E2 effect on the transcription of the α-subunit gene. However, in gonadectomized horses FSH responsiveness to GnRH varies with the duration of E2 treatment, increasing initially and then decreasing (reviewed by Thompson et al.92 ). Likewise, GnRHinduced FSH release decreases as the breeding season approaches despite constant pituitary FSH concentrations.98 However, since the mares in the study of Silvia et al.98 were intact, the declining FSH responsiveness to GnRH could have been due to inhibition by inhibin produced by growing follicles.98 There is some evidence that E2 in concert with GnRH can influence the form of circulating LH in the mare. Treatment of seasonally acyclic mares with E2 and hourly pulses of GnRH, thus mimicking hormonal events during the periovulatory period, results in a sustained increase in the relative in vitro biological activity of plasma LH.101
References 1. Sherman GB, Wolfe MW, Farmerie TA, Clay CM, Threadgill DS, Sharp DC, Nilson JH. A single gene encodes the β-subunits of equine luteinizing hormone and chorionic gonadotropin. Mol Endocrinol 1992;6:951–9. 2. Chopineau M, Martinat N, Gibrat JF, Galet C, Lecompte F, Foulon-Gauze F, Pourchet C, Guillou F, Combarnous Y. Identification of amino-acids in the a-subunit first and third loops that are crucial for the heterospecific follicle-stimulating hormone activity of equid luteinizing hormone/choriogonadotropin. Eur J Endocrinol 2004; 150:877–84. 3. Braselton WE, McShan WH Jr. Purification and properties of follicle-stimulating and luteinizing hormones from horse pituitary glands. Arch Biochem Biophys 1970;139:45–58. 4. Stewart F, Thomson JA, Leigh SE, Warwick JM. Nucleotide (cDNA) sequence encoding the horse gonadotrophin α-subunit. J Endocrinol 1987;115:341–6. 5. Min KS, Shinozaki M, Miyazawa K, Miyazawa K, Nishimura R, Sasaki N, Shiota K, Ogawa T. Nucleotide sequence of eCG α-subunit cDNA and its expression in the equine placenta. J Reprod Dev 1994;40:301–5. 6. Saneyoshi T, Min K-S, Ma XJ, Nambo Y, Hiyama T, Tanaka S, Shiota K. Equine follicle-stimulating hormone: Molecular cloning of β-subunit and biological role of the asparagine-linked oligosaccharide at asparagine56 of α subunit. Biol Reprod 2001;65:1686–90. 7. Bousfield GR, Butnev VY, Gotschall RR, Baker VL, Moore WT. Structural features of mammalian gonadotropins. Mol Cell Endocrinol 1996;125:3–19. 8. Dalpathado DS, Irungu J, Go EP, Butnev VY, Norton K, Bousfield GR, Desaire H. Comparative glycomics of the glycoprotein follicle stimulating hormone: Glycopeptide analysis of isolates from two mammalian species. Biochemistry 2006;45:8665–73. 9. Bousfield GR, Liu W-K, Sugino H, Ward DN. Structural studies on equine glycoprotein hormones: amino acid sequence of equine lutropin β-subunit. J Biol Chem 1987; 262:8610–20.
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10. Talmadge K, Boorstein WR, Fiddes JC. The human genome contains seven genes for the β-subunit of chorionic gonadotropin but only one gene for the β-subunit of luteinizing hormone. DNA Cell Biol 1983;2:281–9. 11. Roser JF, Papkoff H, Murthy HM, Chang YS, Chloupek RC, Potes JA. Chemical, biological and immunological properties of pituitary gonadotropins from the donkey (Equus asinus): comparison with the horse (Equus caballus). Biol Reprod 1984;30:1253–62. 12. Bousfield GR, Butnev VY, Butnev VY. Identification of twelve-O-glycosylation sites in equine chorionic gonadotropin β and equine luteinizing hormone β by solidphase Edman degradation. Biol Reprod 2001;64:136–47. 13. Smith PL, Bousfield GR, Kumar S, Fiete D, Baenziger JU. Equine lutropin and chorionic gonadotropin bear oligosaccharides terminating with SO4 –4-GalNAc and Siaα 2,3 Gal, respectively. J Biol Chem 1993;268:795–802. 14. Irvine CHG. Kinetics of gonadotrophins in the mare. J Reprod Fertil Suppl 1979;27:131–41. 15. Legardiniere S, Duonor-Cerutti M, Devauchelle G, Combarnous Y, Cahoreau C. Biological activities of recombinant equine luteinizing hormone/chorionic gonadotropin (eLH/CG) expressed in Sf9 and Mimic insect cell lines. J Mol Endocrinol 2005;34:47–60. 16. Stewart F, Allen WR. The binding of FSH, LH and PMSG to equine gonadal tissues. J Reprod Fertil Suppl 1979;27: 431–40. 17. Dias JA. Is there any physiological role for gonadotrophin oligosaccharide heterogeneity in humans? II. A biochemical point of view.” Hum Reprod 2001;16: 825–30. 18. Matteri RL, Papkoff H. Characterization of equine luteinizing hormone by chromatofocusing. Biol Reprod 1987; 36:262–9. 19. Matteri RL, Papkoff H. Microheterogeneity of equine follicle-stimulating hormone. Biol Reprod 1988;38:324–31. 20. Ulloa-Aguirre A, Timossi C, Mendez JP. Is there any physiological role for gonadotrophin oligosaccharide heterogeneity in humans? I. Gonadotrophins are synthesized and released in multiple molecular forms. A matter of fact. Hum Reprod 2001;16:599–604. 21. Alexander SL, Irvine CHG. Radioimmunoassay and invitro bioassay of serum LH throughout the equine oestrous cycle. J Reprod Fertil Suppl 1982;32:253–60. 22. Pantke P, Hyland J, Galloway DB, MacLean AA, Hoppen H-O. Changes in luteinizing hormone bioactivity associated with gonadotrophin pulses in the cycling mare. J Reprod Fertil Suppl 1991;44:13–18. 23. Alexander SL, Irvine CHG, Turner JE. Comparison by three different radioimmunoassay systems of the polymorphism of plasma FSH in mares in various reproductive states. J Reprod Fertil Suppl 1987;35:9–18. 24. Damian-Matsumura P, Zaga V, Maldonado A, SanchezHernandez C, Timossi C, Ulloa-Aguirre A. Oestrogens regulate pituitary α2,3-sialyltransferase messenger ribonucleic acid levels in the female rat. J Mol Endocrinol 1999;23:153–65. 25. Shand N, Alexander SL, Irvine CHG. Comparison of the microheterogeneity of horse LH and FSH in the pituitary with that secreted into pituitary venous blood at oestrus. J Reprod Fertil Suppl 1991;44:1–11. 26. Bergendah M, Veldhuis JD. Is there a physiological role for gonadotrophin heterogeneity in humans? III. Luteinizing hormone heterogeneity: a medical physiologist’s perspective. Hum Reprod 2001;16:1058–64.
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27. Whitcomb RW, Schneyer AL, Roser JF, Hughes JP. Circulating antagonist of luteinizing hormone in association with infertility in stallions. Endocrinology 1991;128:2497–502. 28. Barrios-De-Tomasi J, Timossi C, Merchant H, Quintanar A, Avalos JM, Yding Andersen C, Ulloa-Aguirre A. Assessment of the in vitro and in vivo biological activities of the human follicle-stimulating isohormones. Mol Cell Endocrinol 2002;186:189–98. 29. Lin W, Bernard MP, Cao D, Myers RV, Kerrigan JE, Moyle WR. Follitropin receptors contain cryptic ligand binding sites. Mol Cell Endocrinol 2007;260–262:83–92. 30. Saint-Dizier M, Foulon-Gauze F, Lecompte F, Combarnous Y, Chopineau M. Cloning and functional expression of the equine luteinizing hormone/chorionic gonadotrophin receptor. J Endocrinol 2004;183:551–9. 31. Robert P, Amsellem S, Christophe S, Benifla J-L, Bellet D, Koman A, Bidart J-M. Cloning and sequencing of the equine testicular follitropin receptor. Biochem Biophys Res Commun 1994;201:201–7. 32. Richard F, Martinat N, Remy JJ, Salesse R, Combarnous Y. Cloning, sequencing and in vitro functional expression of recombinant donkey follicle-stimulating hormone receptor: a new insight into the binding specificity of gonadotrophin receptors. J Mol Endocrinol 1997;18:193– 202. 33. Bousfield GR, Butnev VY, Butnev VY, Nguyen VT, Gray CM, Dias JA, MacColl R, Eisele L, Harvey DJ. Differential effects of α subunit asparagine56 oligosaccharide structure on equine lutropin and follitropin hybrid conformation and receptor-binding activity. Biochemistry 2004;43:10817–33. 34. Gromoli J, Simoni M. Genetic complexity of FSH receptor function. Trends Endocrinol Metab 2005;16:368–73. 35. Sairam MR, Babu PS. The tale of follitropin receptor diversity: A recipe for fine tuning gonadal responses? Mol Cell Endocrinol 2007;260–262:163–71. 36. Whitmore HL, Wentworth BC, Ginther OJ. Circulating concentrations of luteinizing hormone during estrous cycles of mares as determined by radioimmunoassay. Am J Vet Res 1973;34:631–6. 37. Evans MJ, Irvine CHG. Serum concentrations of FSH, LH and progesterone during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1975;23:193–200. 38. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992. 39. Irvine CHG, Turner JE, Alexander SL, Shand N, van Noordt S. Gonadotrophin profiles and dioestrous pulsatile release patterns in mares as determined by collection of jugular blood at 4 h intervals throughout an oestrous cycle. J Reprod Fertil 1998;113:315–22. 40. Thompson DL Jr, McNeill DR, Wiest JJ, St George RL, Jones LS, Garza F Jr. Secretion of luteinizing hormone and follicle stimulating hormone in intact and ovariectomized mares in summer and winter. J Anim Sci 1987;63:247–53. 41. Irvine CHG, Alexander SL. Secretory patterns and rates of gonadotropin-releasing hormone, follicle-stimulating hormone, and luteinizing hormone revealed by intensive sampling of pituitary venous blood in the luteal phase mare. Endocrinology 1993;132:212–18. 42. Fitzgerald BP, Affleck KJ, Barrows SP, Murdoch WL, Barker KB, Loy RG. Changes in LH pulse frequency and
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amplitude in intact mares during the transition into the breeding season. J Reprod Fertil 1987;79:485–93. Alexander SL, Irvine CHG. Control of onset of breeding season in the mare and its artificial regulation by progesterone treatment. J Reprod Fertil Suppl 1991;44:307–19. Irvine CHG, Alexander SL. A novel technique for measuring hypothalamic and pituitary hormone secretion rates from collection of pituitary venous effluent in the normal horse. J Endocrinol 1987;113:183–92. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Regulation of circulating gonadotropins by the negative effects of ovarian hormones in mares. Biol Reprod 2005;73:315–23. Evans MJ, Alexander SL, Irvine CHG, Taylor TB, Kitson NE, Livesey JH. Administration of the GnRH antagonist cetrorelix during the luteal phase of the mare. Theriogenology 2002;58:527–32. Bergfelt DR, Ginther OJ. Relationships between FSH surges and follicular waves during the estrous cycle in mares. Theriogenology 1993;39:781–96. Irvine CHG. Endocrinology of the estrous cycle of the mare: Applications to embryo transfer. Theriogenology 1981; 15:85–104. Fitzgerald BP, I’Anson H, Legan SL, Loy RG. Changes in patterns of luteinizing hormone secretion before and after the first ovulation in the post-partum mare. Biol Reprod 1985;33:316–23. Alexander SL, Irvine CHG. Effect of graded doses of gonadotrophin-releasing hormone on serum LH concentrations in mares in various reproductive states: comparison with endogenously-generated LH pulses. J Endocrinol 1986;110:19–26. Alexander SL, Irvine CHG. Secretion rates and shortterm patterns of gonadotrophin-releasing hormone, FSH and LH throughout the periovulatory period of the mare. J Endocrinol 1987;114:351–62. Irvine CHG, Alexander SL. The dynamics of gonadotrophin-releasing hormone, LH and FSH secretion during the spontaneous ovulatory surge of the mare as revealed by intensive sampling of pituitary venous blood. J Endocrinol 1994;140:283–95. Evans MJ, Irvine CHG. Measurement of equine FSH and LH: Response of anestrous mares to gonadotropinreleasing hormone. Biol Reprod 1976;15:477–84. Garza F, Thompson DL, French DD, Wiest JJ, St George RL, Ashley KB, Jones LS, Mitchell PS, McNeill DR. Active immunization of intact mares against gonadotropinreleasing hormone: Differential effects on secretion of luteinizing hormone and follicle-stimulating hormone. Biol Reprod 1986;35:347–52. Guillaume D, Bruneau B, Briant CC. Comparison of the effects of two GnRH antagonists on LH and FSH secretion, follicular growth and ovulation in the mare. Reprod Nutr Dev 2002;42:251–64. Padmanabhan V, McNeilly AS. Is there an FSH-releasing factor? Reproduction 2001;121:21–30. Padmanabhan V, Brown MB, Dahl GE, Evans NP, Karsch FJ, Mauger DT, Neill JD, Van Cleeff J. Neuroendocrine control of follicle-stimulating hormone (FSH) secretion: III. Is there a gonadotropin-releasing hormoneindependent component of episodic FSH secretion in ovariectomized and luteal phase ewes? Endocrinology 2003;144:1380–92. Igarashi M, McCann SM. A hypothalamic follicle stimulating hormone releasing factor. Endocrinology 1964;74:446–52.
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FSH and LH 59. Clarke IJ, Cummins JT, Findlay JK, Burman KJ, Doughton BW. Effects on plasma luteinizing hormone and follicle-stimulating hormone of varying the frequency and amplitude of gonadotropin-releasing hormone pulses in ovariectomized ewes with hypothalamopituitary disconnection. Neuroendocrinology 1984; 39:214–21. 60. Burger LL, Haisenleder DJ, Dalkin AC, Marshall JC. Regulation of gonadotropin subunit gene transcription. J Mol Endocrinol 2004;33:559–84. 61. Turner JE, Irvine CHG. The effect of various gonadotrophin-releasing hormone regimens on gonadotrophins, follicular growth and ovulation in deeply anoestrous mares. J Reprod Fertil Suppl 1991;44:213–25. 62. De Kretser DM, Hedger MP, Loveland KL, Phillips DJ. Inhibins, activins and follistatin in reproduction. Hum Reprod Update 2002;8:529–41. 63. Padmanabhan V, Karsch FJ, Lee JS. Hypothalamic, pituitary and gonadal regulation of FSH. Reprod Suppl 2002; 59:67–82. 64. Knight PG. Roles of inhibins, activins, and follistatin in the female reproductive system. Front Neuroendocrin 1996;17:476–509. 65. Phillips DJ. Activins, inhibins and follistatins in the large domestic species. Domest Anim Endocrinol 2005;28:1–16. 66. Gregory SJ, Kaiser UB. Regulation of gonadotropins by inhibin and activin. Semin Reprod Med 2004;22:253–67. 67. Roser JF, McCue PM, Hoye E. Inhibin activity in the mare and stallion. Domest Anim Endocrinol 1994;11:87–100. 68. Medan MS, Nambo Y, Nagamine N, Shinbo H, Watanabe G, Groome NP, Taya K. Plasma concentrations of irinhibin, inhibin A, inhibin pro-αC, FSH, and estradiol17β during estrous cycle in mares and their relationship with follicular growth. Endocrine 2004;25:7–14. 69. Nagaoka K, Nambo Y, Nagamine N, Nagata S-I, Tanaka Y, Shinbo H, Tsunoda N, Taniyama H, Watanabe G, Groome NP, Taya K. A selective increase in circulating inhibin and inhibin pro-α C at the time of ovulation in the mare. Am J Physiol 1999;277:E870–E875. 70. Nambo Y, Kaneko H, Nagata S, Oikawa M, Yoshihara T, Nagamine N, Watanabe G, Taya K. Effect of passive immunization against inhibin on FSH secretion, folliculogenesis and ovulation rate during the follicular phase of the estrous cycle in mares. Theriogenology 1998;50:545–57. 71. McCue PM, Carney NJ, Hughes JP, Rivier J, Vale W, Lasley BL. Ovulation and embryo recovery rates following immunization of mares against an inhibin alphasubunit fragment. Theriogenology 1992;38:823–31. 72. McKinnon AO, Brown RW, Pashen RL, Greenwood PE, Vasey JR. Increased ovulation rates in mares after immunisation against recombinant bovine inhibin alphasubunit. Equine Vet J 1992;24:144–6. 73. Derar RI, Maeda Y, Hoque SM, Osawa T, Watanabe G, Taya K, Miyake Y. Effect of active immunization of pony mares against recombinant porcine inhibin alpha subunit on ovarian follicular development and plasma steroids and gonadotropins. J Vet Med Sci 2004;66:31–5. 74. Miller KF, Wesson JA, Ginther OJ. Changes in concentrations of circulating gonadotropins following administration of equine follicular fluid to ovariectomized mares. Biol Reprod 1979;21:867–72. 75. Miller KF, Wesson JA, Ginther OJ. Interaction of estradiol and a nonsteroidal follicular fluid substance in the regulation of gonadotropin secretion in the mare. Biol Reprod 1981;24:354–8.
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76. Donadeu FX, Ginther OJ. Interrelationships of estradiol, inhibin, and gonadotropins during follicle deviation in pony mares. Theriogenology 2004;61:1395–405. 77. Nambo Y, Hashimoto K, Nakai R, Asai Y, Watanabe G, Taya K. Biological significance of follicular fluid discharged into the peritoneal cavity at ovulation in mares. Anim Reprod Sci 2006;94:228–31. 78. Ginther OJ, Gastal EL, Gastal MO, Beg MA. In vivo effects of pregnancy-associated plasma protein-A, activinA and vascular endothelial growth factor on other follicular-fluid factors during follicle deviation in mares. Reproduction 2005;129:489–96. 79. Beg MA, Ginther OJ. Follicle selection in cattle and horses: role of intrafollicular factors. Reproduction 2006;132:365–77. 80. Karsch FJ, Cummins JT, Thomas GB, Clarke IJ. Steroid feedback inhibition of pulsatile secretion of gonadotropin-releasing hormone in the ewe. Biol Reprod 1987; 36:1207–18. 81. Waring DW, Turgeon JL. A pathway for luteinizing hormone-releasing hormone self-potentiation: cross talk with the progesterone receptor. Endocrinology 1992; 130:3275–82. 82. Silvia PJ, Meyer SL, Fitzgerald BP. Pulsatile gonadotropin secretion determined by frequent sampling from the intercavernous sinus of the mare: possible modulatory role of progesterone during luteolysis. Biol Reprod 1995;53:438–46. 83. Irvine CHG, Alexander SL. Patterns of secretion of GnRH, LH and FSH during the postovulatory period in mares: mechanisms prolonging the LH surge. J Reprod Fertil 1997;109:263–71. 84. Garcia MC, Freedman LJ, Ginther OJ. Interaction of seasonal ovarian factors in the regulation of LH and FSH secretion in the mare. J Reprod Fertil Suppl 1979;27:103–11. 85. Fitzgerald, BP, I’Anson H, Loy RG, Legan SJ. Evidence that changes in LH pulse frequency may regulate the seasonal modulation of LH secretion in ovariectomized mares. J Reprod Fertil 1983;69:685–92. 86. Turner DD, Garcia MC, Webel SK, Ginther OJ. Influence of follicular size on the response of mares to allyl trenbolone given before the onset of the ovulatory season. Theriogenology 1981;16:73–84. 87. Clarke IJ, Pompolo S. Synthesis and secretion of GnRH. Anim Reprod Sci 2005;88:29–55. 88. Nett TM, Turzillo AM, Baratta M, Rispoli LA. Pituitary effects of steroid hormones on secretion of folliclestimulating hormone and luteinizing hormone. Domest Anim Endocrin 2002;23:33–42. 89. Clarke IJ. GnRH and ovarian hormone feedback. Oxford Rev Reprod Biol 1987;9:54–95. 90. Shupnik MA, Rosenzweig BA. Identification of an estrogen-responsive element in the rat LH beta gene. DNA-estrogen receptor interactions and functional analysis. J Biol Chem 1991;266:17084–91. 91. Clarke IJ. Multifarious effects of estrogen on the pituitary gonadotrope with special emphasis on studies in the ovine species. Arch Physiol Biochem 2002;110:62–73. 92. Thompson DL Jr, Garza F Jr, St George RL, Rabb MH, Barry BE, French DD. Relationships among LH, FSH and prolactin secretion, storage and response to secretagogue and hypothalamic GnRH content in ovariectomized pony mares administered testosterone, dihydrotestosterone, estradiol, progesterone, dexamethasone or follicular fluid. Domest Anim Endocrin 1991;8:189–99.
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93. Greaves HE, Porter MB, Sharp DC. Effect of oestradiol on LH secretion and pituitary responsiveness to GnRH in ovariectomized mares. J Reprod Fertil Suppl 2000;56:227–37. 94. Garza F Jr, Thompson DL Jr, St George RL, French DD. Androgen and estradiol effects on gonadotropin secretion and response to GnRH in ovariectomized pony mares. J Anim Sci 1980;62:1654–9. 95. Baldwin DM, Roser JF, Muyan M, Lasley B, Dybdal N. Direct effects of free and conjugated steroids on GnRH stimulated LH release in cultured equine anterior pituitary cells. J Reprod Fertil Suppl 1991;44:327–32. 96. Hart PJ, Squires EL, Imel KJ, Nett TM. Seasonal variation in hypothalamic content of gonadotropin releasing hormone (GnRH), pituitary receptors for GnRH, and pituitary content of luteinizing hormone and follicle stimulation hormone in the mare. Biol Reprod 1984;30:1055– 62. 97. Silvia PJ, Squires EL, Nett TM. Changes in the hypothalamo-hypophyseal axis of mares associated with seasonal reproductive recrudescence. Biol Reprod 1986;35:897–905.
98. Silvia PJ, Squires EL, Nett TM. Pituitary responsiveness of mares challenged with GnRH at various stages of the transition into the breeding season. J Anim Sci 1987;64:790–6. 99. Sharp DC, Grubaugh WR, Weithenhaur J, Davis SD, Wilcox CJ. Effects of steroid administration on pituitary luteinizing hormone and follicle stimulating hormone in ovariectomized pony mares in the early spring: pituitary responsiveness to gonadotropin-releasing hormone and pituitary gonadotropin content. Biol Reprod 1991;44:983–90. 100. Robinson G, Porter MB, Peltier MR, Cleaver BC, Farmerie TA, Wolfe MW, Nilson JH, Sharp DC. Regulation of luteinizing hormone β and α messenger ribonucleic acid by estradiol or gonadotropin-releasing hormone following pituitary stalk section in ovariectomized pony mares. Biol Reprod Monogr 1995;1:373–83. 101. Alexander SL, Irvine CHG. Alteration of relative bioimmunopotency of serum LH by GnRH and estradiol treatment in the mare. In: Proceedings of the Tenth International Congress of Animal Reproduction and Artificial Insemination, 1984;2:1–4.
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Chapter 169
Estrogens Bruce W. Christensen
Steroidogenesis Estrogens are steroid hormones, and, as with all steroid hormones, are derivatives of cholesterol. Estrogens are produced primarily by the ovaries, though smaller amounts are produced by the zona reticularis of the adrenal glands. Investigations in other species have elucidated the steroidogenic pathway of estrogen in both adrenal and ovarian tissue.1 The ovarian steroid pathway involves both theca and granulosa cells (Figure 169.1). Cholesterol crosses the theca cell plasma membrane bound to a lipoprotein, where it is then stored in cytoplasmic vacuoles and transported to the outer mitochondrial membrane. Luteinizing hormone (LH) is released in a pulsatile manner from the anterior pituitary gland and binds to its receptor on the theca cell membrane, mobilizing cholesterol into the steroidogenic pathway. Steroid acute regulatory protein (StAR) aides in transferring cholesterol to the inner mitochondrial membrane where the cholesterol side-chain enzyme system (CYP11A1) cleaves cholesterol into pregnenalone.2 Pregnenalone is further converted to androstenedione. After conversion from cholesterol in the theca cells, androstenedione is transported across the basement membrane to the granulosa cells. Follicle-stimulating hormone (FSH) supports the development of granulosa cells and the steroidogenic pathway within the granulosa cells that further converts androstenedione to estradiol. In the mare, the corpora lutea (CL), and not just the follicles, are significant sources of ovarian estrogen.3–5 An active CL is necessary for a rise in estrogen between days 35 and 50 of gestation (see below). The biosynthetic pathway of estrogen steroidogenesis in the equine CL is not yet completely understood. It has been demonstrated that equine chorionic gonadotropin (eCG) stimulates increased androgen and estrogen production by the CL.4 Specifically, eCG stimulates the activity of 17,20-lyase, a key enzyme in the steroidogenic conversion of pregnenolone to an-
drostenedione, which is then converted by aromatase to estradiol.5 The production of androgens appears to be compartmentalized to the small cells of the CL.6 The result of this increase in enzymatic activity is that the equine CL produces not only progesterone, but also estrogens.
Estrous cycle Estrogens begin to rise approximately 6–8 days prior to ovulation and peak around 2 days prior to ovulation.7 The beginning of this rise is due to estrous follicular growth and is roughly correlated with estrous behavior in the mare. The length of this period is seasonably variable, with longer estrous periods occurring earlier in the breeding season and shorter periods at the peak of the season. High concentrations of estrogens during estrus are positively correlated with increased stromal estrogen and progesterone receptors in the endometrium.8 Estrogen concentrations begin falling prior to ovulation. A dramatic drop in estradiol occurs within 6 hours of ovulation, and basal concentrations are reached within one or two days after ovulation.9–11 Both stromal estrogen and progesterone receptors steadily decline in numbers throughout the luteal phase.8 Estrogen conjugates begin to rise again starting 3 days after ovulation, possibly due either to production from diestral follicular growth or to corpus luteal secretion.3 Possible clinical effects of this secondary peak of estrogens include increased uterine tone around Day 6 post-ovulation and a ‘slight but significant’ increase in uterine edema, noted with transrectal ultrasound evaluation, in early diestrus.10 Estrogen conjugate concentrations return to baseline by the end of the diestral period, around Day 14–15, due either to regression of diestral follicles or luteolysis.9 At this time, corresponding with luteolysis in the cyclic, non-pregnant mare, estrogen and progesterone receptors begin to rise again, in glandular and luminal epithelial cells as well as stromal tissue. The progressive loss of progesterone
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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AC Cholesterol StAR CYP11A
cAMP Pregnenolone
+
CYP17 17OH-pregnenolone CYP17 HSD3B2
Androstenedione
DHEA
Estradiol
Estrone HSD17B1
Androstenedione CYP19
+ cAMP
AC
Figure 169.1 Follicular phase steroid biosynthesis in the ovary illustrating the two-cell/two-gonadotropin theory. It should be noted that there is some species variation in steroidogenic pathways within the ovary and the pathway provided in this figure is characteristic of humans. AC, adenylate cyclase; FSH, follicle-stimulating hormone; LH, luteinizing hormone; StAR, steroidogenic acute regulatory protein; CYP11A, cholesterol side-chain cleavage; CYP17, 17α-hydroxylase,17,20-lyase; HSD3B2, 3β-hydroxysteroid dehydrogenase type II; HSD17B1, 17β-hydroxysteroid dehydrogenase type I; CYP19, aromatase. Modified from Havelock et al., Ovarian granulosa cell lines. Molecular and Cell Endocrinology 2004; 228:67–78. With permission.
receptors during the luteal phase may remove the progesterone block to estrogen receptor synthesis.8 The progressive rise of the effects of estrogens due to the increased estrogen receptor synthesis (by Day 17) allows for increased progesterone receptor synthesis at the end of the luteal phase (Day 20).8
Early pregnancy Pre-implantation Types of estrogens secreted during pregnancy are many and can be grouped into specific free (unconjugated) estrogens (e.g., estradiol-17β and estrone)
and those bound to sulfates (conjugated). These latter types represent up to 100 times greater concentrations in the plasma or urine than the unconjugated forms. Both plasma and urine may be used to assay estrogens. While estradiol is the primary estrogen secreted by follicular activity, the range of estradiol concentrations during peak activity is so narrow that it makes assays of estradiol very difficult to interpret. Estrone sulfate concentrations in the urine, however, are many times more concentrated during this time and measurements of these conjugates can be very useful clinically.9 There is a mechanism, not fully understood in the mare, by which unfertilized ova are retained in the
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Estrogens oviducts whereas embryos proceed along the oviductal tract and are deposited in the uterus between day 5.5 and day 6. Prostaglandin E2 (PGE2 ) is thought to play a role in the transport of the oviductal embryo, but more investigation is necessary to elucidate the entire mechanism.12 In the rat, estrogens have been shown to play a key role in regulating estrogen β2 receptors, which further mediate protein–protein interactions between both gametes and embryos with oviductal papilla, resulting in passage through the oviduct.13 Betteridge14 has postulated that a similar mechanism may exist in the horse. Specific investigations have yet to be performed. The pre-implantation equine conceptus secretes large amounts of estrogens. Specific roles of estrogens in early pregnancy are not determined yet. Steroidogenesis begins as early as day 6 in the conceptus, based on the evidence of 5-3β-hydroxysteroid dehydrogenase activity.15 The presence of intracellular estrogen receptors within the conceptus as early as day 10, with associated protein expression by day 14,16 suggests a role in conceptus growth modulation. Estrogens are detectable from the conceptus as early as day 12.17 These estrogens are not detectable in the maternal circulation,11 and are therefore thought to exert a strictly local effect. Both progesterone and estrogen receptors in the endometrium decrease significantly after day 14 in the pregnant endometrium, compared with the estrous endometrium, suggesting a functionally divergent time period (days 14–17) between the pregnant and estrous endometrium, either as a response to an undiscovered signal for maternal recognition of pregnancy or to the fall in progesterone concentrations.8 It is thought that the secretion of estrogens during this period plays an integral role in signaling the presence of the conceptus and modulating maternal recognition of pregnancy,18, 19 though this has not been clearly proven.20 In pigs, the role of estrogen secretion by the conceptuses to signal maternal recognition of pregnancy is well established.21 In that species, the concentration of luminal epithelial estrogen receptors remains very high during the critical time for maternal recognition of pregnancy, whereas glandular epithelial estrogen receptor concentrations fall.22 In the mare, by contrast, there is no such maintenance of luminal epithelial estrogen receptors, which in fact disappear during this critical time period.8 In addition, the rise in estrogens in the pig is transient and confined to the critical period of maternal recognition of pregnancy,23 where the estrogens serve an indirect luteotropic function by redirecting the secretion of prostaglandins away from the endometrial venous drainage (and therefore away from the corpora lutea).21 In the mare, the rise in estrogens has a much longer, sustained period,
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with a marked rise in estrogens within the yolk sac from day 10 through day 16, after which time they plateau.24 It has been postulated that the estrogens at this time may serve to aid in increasing endometrial vascular permeability and increased vascularization of the conceptus mesoderm and thus maintain the viability of the early conceptus, which relies on simple diffusion for exchange of nutrients and oxygen with the maternal endometrium in the pre-implantation stages.24 It should be noted that the mere presence of increased concentrations of estradiol and its conjugates does not address the bioavailability and effect of each individual molecule type. These actions must be individually elucidated. There is little doubt that estrogens play an integral role in establishment and maintenance of equine pregnancies, but the specific actions are still being investigated. Beyond maternal recognition of pregnancy, it has been suggested that estrogens and their metabolites at this early stage of pregnancy may play roles in embryo growth and fixation.24
Post-implantation In the pregnant mare, starting around day 34, a surge in estrogens begins that is ovarian in origin and is stimulated by endometrial cup secretion of eCG, as discussed above. Evidence suggests that at this time the estrogens are sourced from the corpora lutea, rather than the follicles.3 Another postulated source for these estrogens is the feto-placental unit, though experiments seem to support the former rather than the latter.9 After day 40 of gestation, there appears to be a preferential rise of estrone over estradiol.9 This rise exceeds the pre-ovulatory rise in estrogens (800–900 pg/mL) and then plateaus at around 3 ng/mL.
Late pregnancy Estrogens are produced jointly by the fetal gonads and equine placenta from around day 70 of gestation and peak between days 150 and 270 of gestation.25–27 The proportional size of the fetal gonads during this time is much larger than at any other time in the life of the horse and positively corresponds with estrogen output (Figure 169.2). Using a biochemical pathway similar to that shown for ovarian steroidogenesis in Figure 169.1, the enlarged fetal gonads use the same enzymes to convert cholesterol to large amounts of dehydroepiandrosterone (DHEA), which is secreted into the umbilical artery. 28 In the placenta the DHEA is aromatized into estrone, estadiol-17α, and estradiol-17β.27, 29 Different
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Mare: Physiology and Endocrinology Primary CL
Endometrial cups
Fetal gonads
Progesterone 20
10
400
150
30
Oestrogens 300
100 Parturition
200
eCG 50 100 FSH
0
0
0 0
40
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Plasma conjugated oestrogens (ng ml–1)
Accessory CL
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Figure 169.2 Summary of major endocrinological events throughout pregnancy in the mare. The endometrial cups are present in the uterus between days 40 and 120 after ovulation, and the gonadotropin they secrete (eCG) ovulates or luteinizes secondary follicles that are stimulated to develop in the maternal ovaries by continuous 10–12-day surges of FSH released from the pituitary gland. During the second half of gestation, C19 steroid precursors secreted by the enlarged fetal gonads are aromatized by the placenta to produce very high concentrations of phenolic and ring β-unsaturated estrogens in maternal blood and urine. Maternal plasma progestagen concentrations remain low owing to direct endometrial metabolism of the progestagens synthesized by the diffuse placenta, but they increase during the final 4–6 weeks as a result of additional pregnenolone secreted by the maturing fetal adrenal glands. CL, corpus luteum; eCG, equine chorionic gonadotropin; FSH, follicle-stimulating hormone. Reprinted from Allen,20 with permission.
androgenic precursors, similarly secreted by the fetal gonads into the umbilical artery, are aromatized by the placenta into equilin and equilenin.27 Estrone increases dramatically during the fourth month of gestation and remains high until it decreases around the eighth or ninth month. This rise and fall reflects the growth and regression in size of the fetal gonads.30, 31 Equilin also increases during this time, but remains elevated until the last month of gestation.32 Estrogen concentrations decrease steadily in the mare in the final four months leading up to parturition, more dramatically in some breeds (e.g., Iceland pony) than others.33 Estrogen concentrations in late pregnancy are also lower in multiparous than primiparous mares,32 and are affected by the amount of environmental light to which the mares are exposed, more so than the actual gestational length.34 Differences in estrogen concentrations were not detected between mares carrying colt versus filly fetuses.35
Parturition Estrogens play an important role at parturition by supporting endometrial contractions. In other species, it has been demonstrated that estrogens help to increase myometrial contractility by stimulating
prostaglandin production and increasing concentrations of both oxytocin receptors and myometrial gap junctions.36, 37 A study in the mare demonstrated that removing fetal gonads resulted in a drop in maternal plasma total estrogen concentrations, lower 13, 14-dihydro-15-oxo-PGF-2 alpha (PGFM) concentrations, and weaker myometrial contractions compared with sham-operated mares.30 In non-human primates, increased estrogen concentrations are associated with a change in the amplitude and frequency of oxytocin pulses.38 Earlier in pregnancy, myometrial contractions are low-amplitude and irregular waves, resulting in a tonic uterus. At term, higher estrogen levels mediate increased oxytocin pulses, resulting in strong, progressive uterine contractions that are a part of labor. This illustrates that even though overall estrogens are decreasing late in pregnancy, local estrogen effects may be increasing. Parturition in mares more often than not happens during the night.39 Mares show increased myometrial contractility in the evenings for the 6 days prior to parturition.40 Estradiol-17β plasma concentrations are correspondingly elevated during the evening hours, with this pattern more evident in the final 6 days prior to parturition.41 This pattern suggests a role of estradiol-17β in evening myometrial contractions and thus a significant contribution to the phenomenon of night-time foaling.
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Estrogens
Estrogen assay diagnostics Measurement of serum estrogen concentrations, or measurement of estrogen metabolites in urine42, 43 or feces,44 can be used to diagnose pregnancy in cases when using ultrasound or palpation is not practical or possible, as may be the case in poorer areas or when dealing with wild equid species that will not allow transrectal evaluations.45 Monitoring urinary estrone sulfate concentrations has also been reported as a measure of fetal distress.46, 47
References 1. Ghayee HK, Auchus RJ. Basic concepts and recent developments in human steroid hormone biosynthesis. Rev Endocr Metab Disord 2007;8:289–300. 2. Stocco DM, Clark BJ. Role of the steroidogenic acute regulatory protein (StAR) in steroidogenesis. Biochem Pharmacol 1996;51:197–205. 3. Daels PF, Demoraes JJ, Stabenfeldt GH, Hughes JP, Lasley BL. The corpus luteum: source of oestrogen during early pregnancy in the mare. J Reprod Fertil Suppl 1991;44:501–8. 4. Daels PF, Albrecht BA, Mohammed HO. Equine chorionic gonadotropin regulates luteal steroidogenesis in pregnant mares. Biol Reprod 1998;59:1062–8. 5. Albrecht BA, MacLeod JN, Daels PF. Expression of 3betahydroxysteroid dehydrogenase, cytochrome p450 17alphahydroxylase/17,20-lyase and cytochrome p450 aromatase enzymes in corpora lutea of diestrous and early pregnant mares. Theriogenology 2001;55:551–61. 6. Watson ED. Compartmentalization of steroidogenesis by the equine corpus luteum. Theriogenology 2000;53:1459–66. 7. Makawiti DW, Allen WE, Kilpatrick MJ. Changes in oestrone sulphate concentrations in peripheral plasma of pony mares associated with follicular growth, ovulation and early pregnancy. J Reprod Fertil 1983;68:481–7. 8. Hartt LS, Carling SJ, Joyce MM, Johnson GA, Vanderwall DK, Ott TL. Temporal and spatial associations of oestrogen receptor alpha and progesterone receptor in the endometrium of cyclic and early pregnant mares. Reproduction 2005;130:241–50. 9. Daels PF, Ammon DC, Stabenfeldt GH, Liu IK, Hughes JP, Lasley BL. Urinary and plasma estrogen conjugates, estradiol, and estrone concentrations in nonpregnant and early pregnant mares. Theriogenology 1991;35:1001–17. 10. Ginther OJ. Endocrinology of the ovulatory season. In: Reproductive Biology of the Mare: Basic and Applied Aspects. Chapter 7. Cross Plains, WI: Equiservices, 1992; pp. 233–90. 11. Terqui M, Palmer E. Oestrogen pattern during early pregnancy in the mare. J Reprod Fertil Suppl 1979;27:441–6. 12. Betteridge KJ. Comparative aspects of equine embryonic development. Anim Reprod Sci 2001;60/61:691–702. 13. Shao R, Weijdegard B, Fernandez-Rodriguez J, Egecioglu E, Andersson N, Thurin-Kjellberg A, et al. Ciliated epithelialspecific and regional-specific expression and regulation of the estrogen receptor beta-2 in the fallopian tubes of immature rates. Am J Physiol Endocrinol Metab 2007;293(1): E147–58. 14. Betteridge KJ. Equine embryology: an inventory of unanswered questions. Theriogenology 2007;68S:S9–S21. 15. Paulo E, Tischner M. Activity of 5-3β-hydroxysteroid dehydrogenase and steroid hormone content in early
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20. 21.
22.
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preimplantation horse embryos. Folia Histochim Cytobiol 1985;23:81–4. Rambags BPB, van Tol HTA, van den Eng MM, Colenbrander B, Stout TAE. Expression of progesterone and oestrogen receptors by early intrauterine equine conceptuses. Theriogenology 2008;69:366–75. Choi SJ, Anderson GB, Roser JF. Production of free estrogens and estrogen conjugates by the preimplantation equine embryo. Theriogenology 1997;47:457–66. Marsan C, Goff AK, Sirois J, Betteridge KJ. Steroid secretion by different cell types of the horse conceptus. J Reprod Fertil Suppl 1987;35:363–9. Walters KW, Roser JF, Anderson GB. Maternal-conceptus signaling during early pregnancy in mares: oestrogen and insulin-like growth factor I. Reproduction 2001;121, 331–8. Allen TW. Fetomaternal interactions and influences during equine pregnancy. Reproduction 2001;121:513–27. Bazer FW, Thatcher WW. Theory of maternal recognition of pregnancy in swine based on estrogen controlled endocrine versus exocrine secretion of prostaglandin F2α by the uterine endometrium. Prostaglandins 1977;14:397–400. Geisert RD, Brenner RM, Moffatt RJ, Harney JP, Yellin T, Bazer FW. Changes in oestrogen receptor protein, mRNA expression and localization in the endometrium of cyclic and pregnany gilts. Reprod Fertil Develop 1993;5:247–60. Gadsby JE, Heap RB, Burton RD. Oestrogen production by blastocyst and early embryonic tissue of various species. J Reprod Fertil 1980;60:409–17. Raeside JI, Christie HL, Renaud RL, Waelchli RO, Betteridge KJ. Estrogen metabolism in the equine conceptus and endometrium during early pregnancy in relation to estrogen concentrations in yolk-sac fluid. Biol Reprod 2004; 71:1120–7. Ginther OJ. Endocrinology of pregnancy. In: Reproductive Biology of the Mare. Basic and Applied Aspects. Chapter 10. Cross Plains, WI: Equiservices, 1992; pp. 419–56. Hoffman B, Gentz F, Failing K. Investigations into the course of progesterone-, oestrogen- and eCGconcentrations during normal and impaired pregnancy in the mare. Reprod Domest Anim 1996;31:717–23. Ousey JC. Peripartal endocrinology in the mare and foetus. Reprod Domest Anim 2004;39:222–31. Raeside JI, Gofton N, Liptrap RM, Milne FJ. Isolation and identification of steroids from gonadal vein blood of the fetal horse. J Reprod Fertil Suppl 1982;32:383–7. Pashen RL. Maternal and foetal endocrinology during late pregnancy and parturition in the mare. Equine Vet J 1984; 16:233–8. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J Reprod Fertil Suppl 1979;27: 499–509. Raeside JI, Renaud RL, Christie HL. Postnatal decline in gondal secretion of dehydroepiandrosterone and 3bhydroxyandrosta-5-7-dien-17-one in the newborn foal. J Endocrinol 1997;155:277–82. Cox JE. Oestrone and equilin in the plasma of the pregnant mare. J Reprod Fert Suppl 1975;23:463–8. ¨ E. Faecal oestrogens Palme R, Entenfellner U, Hoi H, Mostl and progesterone metabolites in mares of different breeds during the last trimester of pregnancy. Reprod Domest Anim 2001;36:273–7. Haluska GJ, Currie WB. Variation in plasma concentrations of oestradiol-17β and their relationship to those of progesterone, 13,14-dihydro-15-keto-prostaglandin F-2α and
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oxytocin across pregnancy and at parturition in pony mares. J Reprod Fertil 1988;84:635–46. Raeside JI, Liptrap RM. Patterns of urinary estrogen excretion in individual pregnancy mares. J Reprod Fertil Suppl 1975;23:469–75. Silver M. Prenatal maturation, the timing of birth and how it may be regulated in domestic animals. Exp Physiol 1990;75:285–307. Egarter CH, Husslein P. Biochemistry of myometrial contractility. In: Elder MG (ed.) Bailli`ere’s Clinical Obstetrics and Gynaecology – Prostaglandins. London: Bailli`ere Tindall, 1992; pp. 755–69. Nathanielsz PW, Giussani DA, Mecenas CA, Wu W, Winter JA, Garcia-Villar R, Baguma-Nibasheka M, Honnebier BOM, McDonald TJ. Regulation of the switch from myometrial contractures to contractions in late pregnancy: studies in the pregnant sheep and monkey. Reprod Fertil Dev 1995;7:293–300. Bain AM, Howey WP. Observations on the time of foaling in Thoroughbred mares in Australia. J Reprod Fertil Suppl 1975;23:545. Henry J. Patterns of uterine myoelectrical activity in late gestation pony mares and in those with induced ascending placentitis. University of Florida, Master’s thesis, 2002. O’Donnell LJ, Sheerin BR, Hendry JM, Thatcher MJ, Thatcher WW, LeBlanc MM. 24-hour secretion patterns of
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plasma oestradiol 17β in pony mares in late gestation. Reprod Domest Anim 2003;38:233–5. Lasley B, Ammon D, Daels P, Hughes J, Munro C, Stebenfeldt G. Estrogen conjugate concentrations in plasma and urine reflect estrogen secretion in the nonpregnant and pregnant mare: a review. J Equine Vet Sci 1990;10:444– 8. Evans KL, Kasman LH, Hughes JP, Couto M, Lasley BL. Pregnancy diagnosis in the domestic horse through direct urinary estrone sulfate conjugate analysis. Theriogenology 1984;22:615–20. ¨ E, Bamberg E, Lorin D, Arbeiter K. SicherPalme R, Mostl ¨ heit der Trachtigkeitsdiagnose bei der Stute mittels Ostrogenbestimmung im Kot. Prakt Tierarzt 1989;6:43–4. Monfort SL, Arthur NP, Wildt DE. Monitoring ovarian function and pregnancy by evaluating excretion of urinary oestrogen conjugates in semi-free-ranging Przewalski’s horses (Equus przewalskii). J Reprod Fert 1991;91:155– 64. Douglas RH. Endocrine diagnostics in the broodmare: what you need to know about progestins and estrogens. In: Proceedings for the Society for Theriogenology Annual Conference. Lexington, KY, 2004, pp. 106–15. Kasman LH, Hughes JP, Stabenfeldt GH, Starr MD, Lasley BL. Estrone sulfate concentrations as an indicator of fetal demise in horses. Am J Vet Res 1988;49:184–7.
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Chapter 170
Progesterone Dirk K. Vanderwall
Progesterone is aptly named owing to the fact that an adequate level of progesterone, and/or related progestins, is essential for maintenance of pregnancy in the mare (like most mammals), making progesterone the ultimate ‘pro’-gestational hormone. Changes in the systemic level of progesterone are temporally and causally associated with dramatic changes in behavior and physiological parameters within the hypothalamic–pituitary–ovarian axis and tubular genitalia that are critical for normal reproductive activity and optimum fertility in mares. Some of the major physiological factors regulated by progesterone include gonadotropin secretion from the anterior pituitary gland; uterine and cervical tone; endometrial edema; and secretory activity within the tubular genitalia. In parallel with the role that endogenous progesterone plays in regulating reproductive events in the mare, exogenous progesterone and other synthetic progestins are extensively utilized clinically in both non-pregnant and pregnant mares; the use of exogenous progesterone/progestins is covered in Chapters 184 and 189.
Chemistry and mechanism of action Steroid hormones such as progesterone are derived from cholesterol. The primary site of progesterone synthesis is within the cells of the corpus luteum; however, in pregnant mares the fetal–placental unit becomes the predominant source of progesterone/progestin synthesis and secretion.1, 2 Synthesis of progesterone within steroidogenic cells begins in the mitochondria, where cholesterol is enzymatically converted to pregnenolone by cholesterol side-chain cleavage cytochrome P450 complex.3 Pregnenolone is then transported to the smooth endoplasmic reticulum, where 3β-hydroxysteroid dehydrogenase converts it to progesterone, which then diffuses from the cell and enters the circulatory system. Once progesterone enters the bloodstream it is distributed systemically throughout the body, where
it influences activity at ‘target’ cells containing the appropriate progesterone receptor. The classic paradigm that describes the mechanism of action of progesterone (and other steroid hormones) at its target cells is via regulation of gene expression (i.e., protein synthesis). At target cells, progesterone binds with its specific nuclear receptor and the hormone–receptor complex then acts as a transcription factor that binds with progesterone response elements on the DNA, either turning on the expression of specific genes, resulting in progesterone-induced protein synthesis, or turning off the expression of specific genes, which inhibits protein synthesis in a progesterone-dependent manner.3 Changes in gene expression/protein synthesis induced by progesterone have profound effects on cellular activity, and, depending on the specific target cell/tissue, the altered activity may have local effects (e.g., secretory activity into the uterine lumen) or systemic effects (e.g., altered secretory activity of the hypothalamic–pituitary axis). In contrast to the classic ‘genomic’ model of steroid hormone action, there is evidence that progesterone (and other steroid hormones) can act through rapid, non-genomic pathways that do not alter gene expression/protein synthesis;4 however, the extent and potential importance of such non-genomic actions of progesterone in the mare are not currently known. It is difficult to discuss progesterone without at least a cursory discussion of the other predominant steroid hormone, estrogen, since these two hormones can have synergistic and/or antagonistic actions at their target tissues. For example, estrogen stimulates the formation of progesterone receptors (synergistic effect), whereas progesterone can downregulate estrogen receptors (antagonistic effect).3 Of the two hormones, progesterone is the dominant hormone; a phenomenon readily demonstrated by the concurrent administration of both hormones to seasonally anovulatory or ovariectomized mares, which results in predominantly progestational effects.5, 6
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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10 Progesterone (ng/ml)
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Figure 170.1 Typical concentrations of progesterone in the systemic circulation of mares during the estrous cycle and early pregnancy. Black bar represents estrus.
Sources of progesterone synthesis and secretion In reproductively cycling, non-pregnant mares, the corpus luteum that forms after ovulation is the principal source of progesterone synthesis and secretion during the approximately 2-week diestrous phase of the estrous cycle. Diestrus ends when corpus luteum function (i.e., progesterone secretion) ceases owing to the luteolytic action of prostaglandin F2α (PGF2α ) that is secreted from the endometrium on days 14–16 post ovulation, resulting in regression of the corpus luteum (Figure 170.1).1 The fall in progesterone to basal levels (< 1.0 ng/mL) that occurs during luteolysis allows mares to return to estrus. In pregnant mares, the initial events of corpus luteum formation and progesterone secretion are the same as in non-pregnant mares; however, the presence of the conceptus within the uterine lumen inhibits the secretion of PGF2α from the endometrium at the expected time of luteolysis, which results in continued secretion of progesterone from what is then referred to as the primary corpus luteum of pregnancy (Figure 170.1).1 The primary corpus luteum of pregnancy is the principal source of progesterone secretion until approximately day 40 of gestation when additional corpora lutea begin to form on the ovaries as a result of the combined actions of follicle-stimulating hormone (FSH) from the anterior pituitary gland and equine chorionic gonadotropin secretion from the developing endometrial cups in the uterus.2 These supplemental corpora lutea arise from ovulated follicles (referred to as secondary corpora lutea) or from luteinization of unovulated follicles (referred to as accessory corpora lutea), and collectively these supplemental corpora lutea contribute to a resurgence of progesterone secretion.7 The source of progesterone
synthesis and secretion in pregnant mares becomes even more complex when the fetal–placental unit starts secreting progesterone and related progestagens in appreciable quantities around day 70 of gestation,2 the levels of which are solely adequate to maintain pregnancy (i.e., without the assistance of ovarian luteal progesterone) by approximately day 100 of gestation.8 Physiologically, the transition from luteal to fetal–placental progestational support of pregnancy is complete by approximately day 180 of gestation, at which time all of the corpora lutea (primary and supplemental) regress.7
Biological functions of progesterone Behavior The most readily observable effect of progesterone is its profound influence on reproductive behavior.5, 9 As shown in Figure 170.1, during estrus progesterone is at a basal level < 1.0 ng/mL, which has a ‘permissive’ effect allowing mares to exhibit estrogeninduced estrous behavior that is characterized by receptivity to a stallion. Within 24–48 hours after ovulation, concurrently with the onset of luteal development, the level of progesterone rises above 1.0 ng/mL, which is sufficient to block estrous behavior.10 Non-pregnant mares remain non-receptive to a stallion throughout diestrus while progesterone levels are elevated, and they return to estrus when the progesterone concentration falls to a basal level as a result of luteolysis. In contrast, continued secretion of progesterone, and later fetal–placental progestins, in pregnant mares blocks estrous behavior throughout gestation.
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Progesterone Gonadotropin secretion Because both of the predominant ovarian steroid hormones, estrogen and progesterone, play important and intertwined roles in positive and/or negative feedback effects on the hypothalamic–pituitary axis (i.e., gonadotropin secretion) that collectively regulate numerous reproductive events that are important for normal fertility, the role of both hormones in that capacity will be briefly discussed. Although their specific effects are dose and time (i.e., duration) dependent, in general, estrogen has a stimulatory effect on secretion of luteinizing hormone (LH),11, 12 whereas progesterone has an inhibitory effect on LH secretion.11, 13 In contrast, estrogen has an inhibitory effect on secretion of FSH,12 whereas progesterone does not appear to appreciably affect secretion of FSH.11 The interplay between the hypothalamic–pituitary–ovarian axis and positive/ negative feedback effects of the gonadal steroid hormones on that axis along with numerous other neural and/or hormonal factors (e.g., melatonin, inhibin) results in the ultimate orchestration of the fundamental behavioral and physiological events in non-pregnant and pregnant mares that are necessary for normal reproductive activity and fertility. In addition, the effects of estrogen and progesterone on gonadotropin secretion have important clinical implications for the reproductive management of mares, in particular hormonal methods used for estrus synchronization. For example, because progesterone does not appreciably alter FSH secretion, when exogenous progesterone/progestins are used alone for estrus synchronization there is minimal or no suppression of follicular development, which tends to cause a more variable response to treatment among mares (i.e., less synchrony).14 In contrast, concurrent administration of estrogen and progesterone effectively suppresses follicular growth leading to more uniform synchrony of estrus/ovulation in treated mares.14 Estrus synchronization procedures and protocols are more fully discussed in Chapter 197.
Uterine tone and endometrial edema Progesterone has dramatic effects on the tubular genitalia in both non-pregnant and pregnant mares that can be readily assessed with transrectal palpation and ultrasonography. During estrus when progesterone is at its basal level, the uterus has minimal (i.e., flaccid) tone and the endometrial folds are prominent ultrasonographically owing to accumulation of interstitial fluid (i.e., edema) in the endometrial tissue.6,15–18 Although estrogen clearly plays a role in the development of these uterine characteristics during estrus, a crucial contributing factor is the absence of proges-
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terone.6, 18 When progesterone levels increase during diestrus, uterine tone increases and the endometrial edema dissipates causing the uterine tissue to take on a homogeneous ‘ground-glass’ appearance ultrasonographically because the individual endometrial folds are not discernible (though the folds themselves remain).6,15–18 In pregnant mares, there is a dramatic increase in uterine tone above that associated with diestrus beginning on days 14–16 of gestation that is due to continued progesterone secretion from the corpus luteum that is apparently augmented by estrogen of embryonic and/or maternal origin.6 Collectively, the effects of progesterone on the uterus during early gestation are essential for the dynamic physical interactions between the embryonic vesicle and uterus that are involved with mobility, fixation, orientation, and continued survival of the conceptus.19
Uterine secretion Of all the roles that progesterone plays in regulating various reproductive events throughout the body, the one that may be the most fundamentally important for its ‘pro’-gestational effects is the temporal regulation of the uterine luminal environment, since the developing conceptus is completely dependent on uterine secretions for all of its nutritional and metabolic needs (until the placenta assumes that role). The uterine lumen contains a mixture of numerous biologically active molecules (proteins, carbohydrates, steroids, prostaglandins, ions, etc.) that are collectively referred to as uterine histotrophe. Proposed functions of histotrophe include: (i) enabling spermatozoa to survive and ascend the tubular genitalia, (ii) providing a system for metabolic exchange of nutrients and waste products between the conceptus and the endometrium (until the placenta takes over those functions), (iii) maintaining the appropriate physical–chemical environment (e.g., osmolarity), (iv) fulfilling immunological and antibacterial requirements, and (v) providing lubrication for embryo mobility and later providing for adhesiveness to aid maintenance of conceptus position and orientation.7 The composition of uterine histotrophe changes throughout the estrous cycle, and during pregnancy the uterine luminal milieu also reflects the secretory and metabolic activity of the developing conceptus. Research investigating the composition of the uterine luminal environment during the estrous cycle and early pregnancy in mares has primarily focused on the presence and characteristics of uterine secretory proteins. In 1978, Zavy et al.20 described a non-surgical technique for collection of uterine fluid from mares to quantify changes in uterine luminal proteins throughout the estrous cycle. The
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total amount of protein obtained from the uterine lumen increased significantly during the course of the estrous cycle, with the highest amounts present in the mid- to late diestrous period (peak on day 16 post ovulation), which was followed by a precipitous decline on days 18 and 20 in non-pregnant mares.20 In a subsequent report,21 column chromatography was used to begin characterization of the proteins recovered from the uterine lumen throughout the estrous cycle in non-pregnant mares. Three out of five protein fractions recovered from the uterus had the same characteristics as those found in blood plasma, suggesting that these proteins enter the uterine lumen via transudation from the bloodstream; however, two other fractions consisted of proteins that differed from blood plasma.21 In addition, Zavy et al. described the presence of a specific protein with acid phosphatase activity (subsequently identified as uteroferrin) that changed significantly during the estrous cycle in non-pregnant mares; maximal values were found in the late luteal phase on days 16–18 post ovulation, followed by a rapid decline on day 20.21 These investigators then went on to characterize and compare the uterine luminal proteins in non-pregnant and pregnant mares.22 As Zavy et al. demonstrated in their prior work,20, 21 the total protein in the uterine fluid from non-pregnant mares peaked in mid-diestrus and then declined, whereas somewhat surprisingly in pregnant mares there was a significant depression in total recoverable protein on days 8, 12, 14, and 16 of gestation that the authors surmised may have been due to increased utilization by the conceptus to meet its nutritional and metabolic needs.22 Acid phosphatase activity (i.e., uteroferrin) peaked in the fluid from non-pregnant mares on day 14 and then decreased, whereas it continued to increase in pregnant mares.22 In a subsequent report,23 two-dimensional gel electrophoresis was used to further characterize and compare the uterine luminal proteins in non-pregnant and pregnant mares, and investigate the uterine luminal proteins present in ovariectomized mares treated with progesterone, estrogen, or estrogen followed by progesterone. Based on differing electrophoretic patterns, several acidic and basic proteins were identified that were not found in blood serum and, although the exact pattern of when the various individual proteins were identified in the uterine fluid varied, in general they were evident on or around day 12 post ovulation in both non-pregnant and pregnant mares; however, the levels then decreased in non-pregnant mares, whereas they continued to increase in the pregnant mares.23 The same proteins were also identified in the uterine fluid recovered from ovariectomized mares treated with progesterone or estrogen followed by progesterone, but not in mares treated
only with estrogen, demonstrating the progesterone dependence of these uterine secretory products.23 As in their prior work,21, 22 a specific protein with acid phosphatase activity was identified that, based upon its electrophoretic properties, appeared to be identical to the porcine uterine protein uteroferrin that appears to be involved in iron transport from the endometrium to the conceptus.23 In a follow-up study, McDowell et al.24 confirmed that the equine uterine acid phosphatase is uteroferrin. Further studies using steroid-treated ovariectomized mares have substantiated the primary role that progesterone (augmented by estrogen) plays in regulating quantitative and qualitative changes in uterine secretory proteins;25, 26 additional progesterone-dependent uterine proteins have been identified and their roles during pregnancy continue to be actively investigated.27–32
References 1. Allen WR. The physiology of early pregnancy in the mare. In: Proceedings of the 46th Annual Convention of the American Association of Equine Practitioners, 2000, pp. 338–54. 2. Allen WR. The physiology of later pregnancy in the mare. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 2000, pp. 3–16. 3. Niswender GD, Juengel JL, Silva PJ, Rollyson MK, McIntush EW. Mechanisms controlling the function and life span of the corpus luteum. Physiol Rev 2000;80:1–29. 4. Bishop CV, Stormshak F. Non-genomic actions of progesterone and estrogens in regulating reproductive events in domestic animals. Vet J 2008;176:270–80. 5. Asa CS, Goldfoot DA, Garcia MC, Ginther OJ. The effect of estradiol and progesterone on the sexual behavior of ovariectomized mares. Physiol Behav 1984;33:681–6. 6. Hayes KEN, Ginther OJ. Role of progesterone and estrogen in development of uterine tone in mares. Theriogenology 1986;25:581–90. 7. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992; pp. 317, 324–328, 438–45. 8. Holtan DW, Squires EL, Lapin DR, Ginther OJ. Effect of ovariectomy on pregnancy in mares. J Reprod Fertil Suppl 1979;27:457–63. 9. Crowell-Davis SL. Sexual behavior of mares. Horm Behav 2007;52:12–17. 10. Loy RG, Swan SM. Effects of exogenous progestogens on reproductive phenomena in mares. J Anim Sci 1966;25:821–6. 11. Garcia MC, Freedman LJ, Ginther OJ. Interaction of seasonal and ovarian factors in the regulation of LH and FSH secretion in the mare. J Reprod Fertil Suppl 1979;27:103–11. 12. Burns PJ, Douglas RH. Effects of daily administration of estradiol-17β on follicular growth, ovulation, and plasma hormones in mares. Biol Reprod 1981;24:1026–31. 13. Ginther OJ, Utt MD, Bergfelt DR, Beg MA. Controlling interrelationships of progesterone/LH and estradiol/LH in mares. Anim Reprod Sci 2006;95:144–50. 14. Lofstedt RM. Control of the estrous cycle in the mare. In: Van Camp SD (ed.) Veterinary Clinics of North America:
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Progesterone
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Equine Practice, vol. 4, no. 2. Philadelphia: W.B. Saunders Co., 1988; pp. 177–96. Ginther OJ, Pierson RA. Ultrasonic anatomy and pathology of the equine uterus. Theriogenology 1984;21:505–15. Hayes KEN, Pierson RA, Scraba ST, Ginther OJ. Effects of estrous cycle and season on ultrasonic uterine anatomy in mares. Theriogenology 1985;24:465–77. Griffin PG, Ginther OJ. Dynamics of uterine diameter and endometrial morphology during the estrous cycle and early pregnancy in mares. Anim Reprod Sci 1991;25: 133–42. Pycock JF, Dieleman S, Drifjhout P, van der Brug Y, Oei C, Van Der Weijden GC. Correlation of plasma concentrations of progesterone and oestradiol with ultrasound characteristics of the uterus and duration of oestrus behaviour in the cycling mare. Reprod Dom Anim 1995;30:224–7. Kastelic JP, Adams GP, Ginther OJ. Role of progesterone in mobility, fixation, orientation, and survival of the equine embryonic vesicle. Theriogenology 1987;27:655–63. Zavy MT, Bazer FW, Sharp DC. A non-surgical technique for the collection of uterine fluid from the mare. J Anim Sci 1978;47:672–6. Zavy MT, Bazer FW, Sharp DC, Wilcox CJ. Uterine luminal proteins in the cycling mare. Biol Reprod 1979;20: 689–98. Zavy MT, Mayer R, Vernon MW, Bazer FW, Sharp DC. An investigation of the uterine luminal environment of nonpregnant and pregnant pony mares. J Reprod Fertil Suppl 1979;27:403–11. Zavy MT, Sharp DC, Bazer FW, Fazleabas A, Sessions F, Roberts RM. Identification of stage-specific and hormonally induced polypeptides in the uterine protein secretions of the mare during the oestrous cycle and pregnancy. J Reprod Fertil Suppl 1982;64:199–207.
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24. McDowell KJ, Sharp DC, Fazleabas A, Roberts RM, Bazer FW. Partial characterization of the equine uteroferrin-like protein. J Reprod Fertil Suppl 1982;32:329–34. 25. McDowell KJ, Sharp DC, Grubaugh W. Comparison of progesterone and progesterone +oestrogen on total and specific uterine proteins in pony mares. J Reprod Fertil Suppl 1987;35:335–342. 26. Hinrichs K, Kenney RM, Sharp DC. Differences in protein content of uterine fluid related to duration of progesterone treatment in ovariectomised mares used as embryo recipients. Equine Vet J 1989; Suppl. 8: 49–55. 27. McDowell KJ, Adams MH, Smith TL, Baker CB. Retinolbinding protein in equine endometrium. Equine Vet J 1993; Suppl. 15: 19–23. 28. Crossett B, Suire S, Herrler A, Allen WR, Stewart F. Transfer of a uterine lipocalin from the endometrium of the mare to the developing equine conceptus. Biol Reprod 1998;59:483–90. 29. Stewart F, Gerstenberg C, Suire S, Allen WR. Immunolocalization of a novel protein (P19) in the endometrium of fertile and subfertile mares. J Reprod Fertil Suppl 2000;56:593–9. 30. Muller-Schottle F, Bogusz A, Grotzinger J, Herrler A, Krusche CA, Beier-Hellwig K, Beier HM. Full-length complementary DNA and the derived amino acid sequence of horse uteroglobin. Biol Reprod 2002;66:1723–8. 31. Ellenberger C, Wilsher S, Allen WR, Hoffmann C, Kolling M, Bazer FW, Klug J, Schoon D, Schoon HA. Immunolocalisation of the uterine secretory proteins uterocalin, uteroferrin and uteroglobin in the mare’s uterus and placenta throughout pregnancy. Theriogenology 2008;70:746–57. 32. Hayes MA, Quinn BA, Keirstead ND, Katavolos P, Waelchli RO, Betteridge KJ. Proteins associated with the early intrauterine equine conceptus. Reprod Dom Anim 2008;43 Suppl. 2: 232–7.
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Chapter 171
Prostaglandins Tom A.E. Stout
Introduction Since their initial description in the 1930s as seminal plasma components capable of eliciting smooth muscle contractions,1 the prostaglandins have been shown to play critical roles in a large and diverse range of physiological processes. Prostaglandins are 20-carbon atom derivatives of the fatty acids arachadonic acid and di-homo-γ -linolenic acid; most are labile molecules that are metabolized rapidly in the lungs2 and, therefore, have a short circulatory half-life. As a result, prostaglandins are most commonly involved in local (paracrine or autocrine) signaling pathways. Nevertheless, some of the betterknown reproductive functions of prostaglandins involve their action as classical endocrine hormones, with a primary effect at a target organ distant to the site of production, as typified by the role of prostaglandin F2α (PGF2α ) in cyclical luteolysis. From a research or diagnostic perspective, the lability of the prostaglandins complicates examination of endogenous secretion; in most cases, in vivo secretion is therefore assumed, or estimated by measuring concentrations of more stable circulating prostaglandin metabolites; for example, endogenous PGF2α secretion is generally estimated by measuring plasma concentrations of its major circulating metabolite, 13,14dihydro-15-keto-PGF2α (PGF metabolite or PGFM).3 In the horse, initial interest in prostaglandins was stimulated by the demonstration during the early 1970s that PGF2α produced by the endometrium was responsible for cyclical luteolysis in domestic ruminant species.4 Not surprisingly, therefore, the first studies into the role of a prostaglandin in equine reproduction focused on determining whether exogenous administration of the recently developed PGF2α analogs would induce premature destruction of the corpus luteum followed by a predictably timed return to estrus and ovulation.5, 6 Subsequently, it has become clear that prostaglandins play critical
roles in many other aspects of female equine reproductive physiology including ovulation, passage of the early embryo through the oviduct, intrauterine migration of the conceptus during maternal recognition of pregnancy, and cervical relaxation and uterine contraction during parturition. As a result, analogs of the most clinically relevant prostaglandins (E2 and F2α ), and treatments aimed at either preventing or augmenting endogenous prostaglandin release, have become important tools in the management or manipulation of a range of reproductive processes. In addition, recent advances in molecular biology have aided the discovery of additional, predominantly paracrine or autocrine, roles of prostaglandins in reproductive pathways, while technological advances have refined the prostaglandin analogs or inhibitors available (e.g., more potent, or more specific and with fewer side-effects) and led to the development of novel ways of delivering the prostaglandins to the desired site of action. This chapter summarizes our current understanding of the major physiological roles of prostaglandins in mare reproduction; therapeutic uses are discussed in more detail in Chapter 187.
Ovulation Prostaglandins produced by the follicle wall are believed to be essential for ovulation in a wide range of mammalian species; in particular, prostaglandins are thought to play an obligatory role in follicle wall rupture.7 Indeed, in the mare, there is good reason to believe that LH (or exogenous hCG) triggers ovulation by upregulating follicle wall expression of cyclooxygenase 2 (COX-2),8 the key rate-limiting enzyme in prostaglandin synthesis. Indeed, COX-2 expression in the wall of the pre-ovulatory follicle rises sharply 30–33 hours after hCG injection, followed closely (36–39 hours post-hCG) by a profound increase in
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Prostaglandins follicular fluid concentrations of both PGE2 and PGF2α .9 That this increase in prostaglandin secretion is instrumental to ovulation in the mare was suggested by the postponement of ovulation that followed injection of the prostaglandin synthesis inhibitor, indomethacin, into the lumen of pre-ovulatory follicles.10 Moreover, in other species, indomethacin-induced prevention of ovulation can be overcome by intrafollicular administration of PGE2 or PGF2α , thereby confirming a pivotal role for prostaglandins in the ovulatory process (for review see Weems et al.).7 While the exact mechanisms by which follicular prostaglandins induce ovulation are not known, induction of follicle wall ischemia and collagenolysis appear to be important contributory components.7 In practical terms, the central role of prostaglandins in physiological ovulation may in large part explain why administration of PGF2α analogs has been reported to induce ovulation,11 albeit not as reliably as hCG or long-acting GnRH products. It may also explain why large diestrus follicles (> 36 mm) often ovulate within 48 h after administration of a large dose of the PGF2α analog cloprostenol (e.g., 250–625 µg) to induce luteolysis,12 i.e., even before progesterone concentrations have reached baseline levels and the mare has returned to estrus. Indeed, Newcombe et al. were able to show that both increasing dose of PGF2α analog and larger follicle diameter at the time of administration were associated with a shortened interval to ovulation. With respect to the possible ovulatory effects of exogenous PGF2α , it is undoubtedly also significant that administration of a PGF2α analog to a diestrous mare results in a rapid increase in circulating LH concentrations, which are maintained above the levels in control mares for at least 72 h.13
Selective oviductal transport In the mare, only developing embryos are reliably transported into the uterus; unfertilized oocytes generally remain in the ampulla of the oviduct where they slowly degenerate.14 Moreover, the effector of this selective recognition and transport mechanism is embryonic PGE2 .15 In a comprehensive series of experiments, Weber and colleagues demonstrated that equine embryos not only secrete PGE2 (and PGF2α ) from around day 4–5 after ovulation,16, 17 but that intra-oviductal administration of PGE2 on day 4 induces premature uterine entry of the embryo.18 As a result of these and related findings, it is now accepted that early embryos are initially trapped in the ampullary region of the oviduct by tonic contraction of the isthmic circular smooth muscle. Once a developing embryo has reached the compact morula stage, however, it is able to secrete sufficient PGE2 to relax
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the isthmic ‘sphincter’, thereby allowing the embryo to pass rapidly (within a matter of hours) through the isthmus and into the uterus. Since unfertilized oocytes are unable to secrete the prostaglandins required to stimulate isthmic dilation, they rarely enter the uterus unless accompanying an embryo (e.g., from a subsequent cycle). Since the initial demonstration that locally applied PGE2 is able to induce the ‘emptying’ of oviductal contents into the uterus, laparoscopic application of PGE2 gel onto one or both oviducts has been described as a means of either hastening oviductal passage of early-stage embryos suitable for manipulations such as cryopreservation,19 or for reestablishing oviductal patency in subfertile mares in which the problem, by a process of eliminating all other likely causes, is presumed to be blockage of the oviduct with degenerating oocytes and/or inspissated follicular fluid.20
Cyclical luteolysis and the maternal recognition of pregnancy During the late 1960s and early 1970s it became clear that PGF2α was the uterine ‘luteolysin’ that destroyed the corpus luteum and initiated a new estrous period in the poly-estrous farm animal species.4 Moreover, the central role of PGF2α in this pre-programmed system for terminating diestrus made it the obvious physiological target for a pregnancy recognition signal, given that a developing conceptus must somehow avert cyclical luteolysis if it is to ensure the prolongation of the progesterone secretion needed to ensure continuation of a healthy pregnancy.21 Cyclical luteolysis The role of uterine PGF2α in the initiation of luteolysis was the first, and remains arguably the most clinically relevant, function proven for a prostaglandin in equine reproduction. During the 35 years since PGF2α was first proposed as the uterine ‘luteolysin’, it has become clear that cyclical luteolysis in the mare is indeed initiated by pulses of PGF2α released from the endometrium during approximately days 14–16 after ovulation.3, 22 Somewhat unexpectedly, it subsequently transpired that mares lack the close anatomical intertwining of uterine vein and utero-ovarian artery23 that in the ruminant species enables direct ‘counter-current’ transfer of PGF2α from the blood exiting the uterine horn to that coursing to the ipsilateral ovary, and thereby allows the luteolysin to reach the corpus luteum without a passage through the lungs that would render more than 95% of the hormone inactive.2 In the mare, as compared to the ruminants, PGF2α must instead reach the ovary via the systemic circulation, a feat that appears to be made
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possible by a much lower proportion of the luteolysin being destroyed during passage through the lungs (approximately 60%), and because the mare is more sensitive to the luteolytic effects of PGF2α . In the last 15 years, it has also become clear that oxytocin plays an important role in (cyclical) luteolytic PGF2α secretion, and it is now generally accepted that oxytocin of hypothalamic-pituitary24 and/or endometrial25–27 origin both stimulates uterine PGF2α release28 and is itself released in response to PGF2α ,24 thereby completing a positive feedback loop that is almost certainly instrumental in ensuring the generation of the multiple large pulses of PGF2α that are required to complete luteolysis. In short, there appear to be two critical aspects to cyclical luteolysis in the mare: (i) development of uterine oxytocin receptors and the ability to release PGF2α in response to oxytocin, which begins at around day 10 after ovulation28, 29 ; and (ii) upregulation of COX-2 expression in the endometrium, and thereby the ability to produce prostaglandins, which occurs between days 13 and 15 after ovulation.30 While it is assumed that progesterone, estrogen, and their respective endometrial receptors are instrumental in establishing the various components of the luteolytic cascade,31 it is not yet clear how they do so; neither is it clear how the luteolytic PGF2α -oxytocin feedback loop is initially set in motion. Furthermore, while PGF2α is clearly instrumental to the initiation of corpus luteum destruction, it is not known exactly how it exerts its luteolytic effects; it is, however, believed that the salient effects include vasoconstriction, reduced intracellular (luteal cell) cholesterol transport, and stimulation of an influx of leukocytes that, in turn, secrete cytokines capable of triggering luteal cell apoptosis and luteal matrix remodelling (for review see Weems et al.).7 Maternal recognition of pregnancy Short (1969)21 coined the term ‘maternal recognition of pregnancy’ (MRP) to describe the process by which a female mammal determines that a conceptus is present in her uterus and makes the appropriate physiological changes to ensure that an adequate environment is established for the further development of that conceptus. Since the primary determinant of an ‘adequate environment’ is a continuing supply of progesterone, MRP is now more commonly taken to refer specifically to the conceptus-initiated events that result in the prolonged lifespan and secretory activity of the primary corpus luteum. In the mare, MRP involves an absolute suppression of the endometrium’s ability to release PGF2α . In analogy to the establishment of the luteolytic pathway, MRP appears to involve at least two components: (i) inhi-
bition of the endometrium’s ability to secrete PGF2α in response to oxytocin32, 33 ; and (ii) suppression of COX-2 expression and thereby the ability of the endometrium to secrete prostaglandins per se.30 Given its critical importance to pregnancy maintenance, it is surprising that the equine conceptus MRP signal has still not been identified. In this respect, molecules proven to function as MRP signals in the other large domestic species, such as interferon-τ in the ruminants and estrogens in the pig (for review see Spencer et al.)34 are either not secreted by equine conceptuses (interferon-τ )35 or are secreted but do not reliably extend corpus luteum lifespan (estrogens).36 Recently, interest in the conceptus factor(s) responsible for prolonging corpus luteum lifespan in the pregnant mare has been reawakened by reports that corpus luteum lifespan can be reliably prolonged in cycling mares by continuous29 or repeated37 administration of oxytocin for 10 days starting on day 7 or 8 after ovulation, intrauterine insertion of a plastic ball early in diestrus,38 or a single intrauterine administration of a plant oil between days 8 and 12 after ovulation.39 However, while studies investigating the size of the conceptus product that inhibits PGF2α secretion by equine endometrium have concluded that the equine MRP signal must have a molecular weight of 1–6 kDa40 or 3–10 kDa,41 further characterization has proven elusive. On the other hand, MRPstage equine conceptuses have been shown to secrete a variety of hormones, including PGI2 , PGE2 , and PGF2α ,42, 43 IGF-1,44 and estrogens45, 46 which, although they may not be directly involved in inhibiting endometrial PGF2α secretion, almost certainly play significant roles in processes essential to pregnancy maintenance, such as conceptus migration,36 increased uterine vascularity,47 and qualitative and quantitative alterations in the composition of the uterine secretions that make up the protein-rich histotrophe or ‘uterine milk’.48 Indeed, since administration of the prostaglandin synthesis inhibitor, flunixin meglumine, temporarily halts conceptus migration, it is clear that prostaglandins produced by either conceptus or uterus are critical in generating the myometrial contractions that drive conceptus migration during days 10–16 after ovulation.36 In turn, this conceptus migration is thought to aid conceptus harvesting of nutrients present in the histotrophe,49 and is known to play a critical role in enabling adequate distribution of the MRP signal. Indeed, when McDowell et al.50 surgically restricted horse conceptuses to a single uterine horn, PGF2α release was not inhibited sufficiently to block luteolysis and, as a result, the pregnancies failed. It is now accepted that conceptus migration during days 10–16 after ovulation is a critical component of MRP signaling and, in clinical practice, it is thought that large intraluminal uterine cysts or
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Prostaglandins accumulations of cysts may impede conceptus migration to a degree that endangers MRP. The possible effect on conceptus migration is one reason for recommending removal of large intraluminal endometrial cysts detected during the pre-breeding examination of a broodmare. However, the utility of cyst removal is disputed (see Eilts et al.)51 since current evidence supporting improved fertility following cyst ablation ¨ (e.g., Kollmann et al.)52 is largely anecdotal.
Parturition Prostaglandins also play critical roles during parturition in the mare. In particular, PGF2α appears to be instrumental in eliciting the myometrial contractions (for review see Ousey)53 involved in all three stages of labor, i.e., initial cervical dilation and rupture of the allantois, expulsion of the foal, and subsequent expulsion of the fetal membranes. In this respect, PGF2α release has been reported to increase significantly some 1–2 h prior to parturition, and to peak between approximately 15 minutes before and very soon after spontaneous foaling.54 While oxytocin also clearly plays an important role in stimulating uterine contractions during equine fetal and placental expulsion, it is not yet clear whether the oxytocin is of exclusively hypothalamic-pituitary origin, or is supplemented by an endometrial source, nor is it clear exactly how or whether oxytocin and PGF2α release are related or coordinated during these processes. PGE2 is also thought to be involved in parturition and, in particular, in promoting cervical ‘ripening’ and relaxation in preparation for delivery. However, this is currently an assumption based largely on studies in women and ruminant species, in which PGE2 has been shown to induce cervical softening via changes in the collagen matrix that resemble those seen during spontaneous parturition,55 combined with a few reports that intracervical application of PGE2 accelerates cervical relaxation during induced parturition in mares.56 There is as yet, however, no definitive proof that PGE2 induces cervical relaxation during spontaneous foaling in the mare, or for that matter in any other species. On the other hand, both PGE2 and PGF2α are known to be synthesized by utero-placental tissues and can be detected in both maternal and fetal plasma, and the allantoic fluid, during late gestation.57 Indeed, uteroplacental PGF2α production capacity probably begins during mid-gestation, at which time it is thought that the initiation of pre-term labor is prevented by endometrial production of the enzyme 15-hydroxyprostaglandin dehydrogenase (PGDH; from approximately day 150 of gestation), which rapidly metabolizes any PGF2α into inactive metabolites.58 If this is the case, a precipitous fall in placental proges-
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terone production at term could suppress PGDH activity and thereby permit the rapid, approximately 50-fold, increase in PGF2α secretion that accompanies birth. Moreover, maintenance of PGDH concentrations may help to explain how and why exogenous progesterone administration may help prevent premature labor in mares.53 Nevertheless, the exact pattern of utero-placental prostaglandin production during late gestation in the mare, the relationship of prostaglandin secretion and activity to the other endocrinological changes that occur around parturition, and the way in which the prostaglandins, and other hormones such as relaxin and oxytocin, contribute to cervical relaxation and uterine contraction in the mare have yet to be fully explained.
Conclusions Prostaglandins are known to play important roles in a wide range of reproductive processes in the mare and, as a result, prostaglandin analogs or inhibitors are used to modulate a number of reproductive events. However, for a number of the physiological processes in which prostaglandins are thought to participate, we do not yet fully understand how endogenous prostaglandins exert their effects or, therefore, whether exogenous prostaglandin analogs or inhibitors are the most physiological or practical way to intervene or interrupt these processes.
References 1. von Euler US. On the specific vaso-dilating and plain muscle stimulating substances from accessory genital glands in man and certain animals (prostaglandin and vesiglandin). J Physiol 1936;88:13–234. 2. Piper P, Vane J, Wylie JH. Inactivation of prostaglandins by the lungs. Nature 1970;225:600–4. 3. Kindahl H, Knudsen O, Madej A, Edqvist L-E. Progesterone, prostaglandin F2a , PMSG and oestrone sulphate during early pregnancy in the mare. J Reprod Fertil Suppl 1982;32:353–9. 4. McCracken JA, Carlson JC, Glew MC, Goding JR, Baird DT, Green K, Samuelsson B. Prostaglandin F2a identified as a luteolytic hormone in the sheep. Nature 1972;238:129–34. 5. Douglas RH, Ginther OJ. Effect of prostaglandin F2alpha on length of diestrus in mares. Prostaglandins 1972;2:265–8. 6. Allen WR, Rowson LE. Control of the mare’s oestrous cycle by prostaglandins. J Reprod Fertil 1973;33:539–43. 7. Weems CW, Weems YS, Randel RD. Prostaglandins and reproduction in female farm animals. Vet J 2006;171:206–28. 8. Boerboom D, Sirois J. Molecular characterization of equine prostaglandin G/H synthase-2 and regulation of its messenger ribonucleic acid in preovulatory follicles. Endocrinology 1998;139:1662–70. 9. Sirois J, Dor´e M. The late induction of prostaglandin G/H synthase-2 in equine preovulatory follicles supports its role as a determinant of the ovulatory process. Endocrinology 1997;138:4427–34.
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10. Watson ED, Sertich PL. Concentrations of arachidonic metabolites, steroids and histamine in preovulatory horse follicles after administration of human chorionic gonadotropin and the effect of intrafollicular injection of indomethacin. J Endocrinol 1991;129:131–9. 11. Savage NC, Liptrap RM. Induction of ovulation in cyclic mares by administration of a synthetic prostaglandin, fenprostalene, during oestrus. J Reprod Fertil Suppl 1987;35:239–43. ¨ 12. Newcombe JR, Jochle W, Cuervo-Arango J. Effect of dose of cloprostenol on the interval to ovulation in the diestrous mare: A retrospective. J Equine Vet Sci 2008;28: 532–9. 13. Ginther OJ, Gastal EL, Gastal MO, Beg ME. Effect of prostaglandin F2a on ovarian, adrenal, and pituitary hormones and on luteal blood flow in mares. Domest Anim Endocrinol 2007;32:315–28. 14. van Niekerk CH, Gerneke WH. Persistence and parthenogenetic cleavage of tubal ova in the mare. Onderstepoort J Vet Res 1966;31:195–232. 15. Weber JA, Woods GL, Lichtenwalner AB. Relaxatory effect of prostaglandin E2 on circular smooth muscle isolated from the equine oviductal isthmus. Biol Reprod Mono 1995;1:125–30. 16. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 secretion by oviductal transport-stage equine embryos Biol Reprod 1991;45:540–3. 17. Freeman DA, Woods GL, Vanderwall DK, Weber JA. Embryo-initiated oviductal transport in mares. J Reprod Fertil 1992;75:535–8. 18. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 hastens oviductal transport of equine embryos. Biol Reprod 1991;45:544–6. 19. Robinson SJ, Neal H, Allen WR. Modulation of oviductal transport in mares by local application of prostaglandin E2 . J Reprod Fertil Suppl 2000;56:587–92. 20. Allen WR, Wilsher S, Morris L, Crowhurst JS, Hillyer MH, Neal HN. Laparoscopic application of PGE2 to re-establish oviducal patency and fertility in infertile mares: a preliminary study. Equine Vet J 2006;38:454–9. 21. Short RV. Implantation and the maternal recognition of pregnancy. In: Wolstenholme GEW, O’Connor M (eds) Ciba Foundation Symposium on Foetal Autonomy. London: J & A Churchill Ltd, 1969; pp. 2–26. 22. Douglas RH, Ginther OJ. Concentration of prostaglandin F in uterine venous plasma of anaesthetised mares during the oestrous cycle and early pregnancy. Prostaglandins 1976;11:251–60. 23. Ginther OJ, Garcia MC, Squires EL, Steffenhagen WP. Anatomy of vasculature of uterus and ovaries in the mare. Am J Vet Res 1972;33:1561–8. 24. Vanderwall DK, Silvia WJ, Fitzgerald BP. Concentrations of oxytocin in the intercavernous sinus of mares during luteolysis: temporal relationship with concentrations of 13,14-dihydro-15-keto-prostaglandin F2a . J Reprod Fert 1998;112:337–46. 25. Behrendt-Adam CY, Adams MH, Simpson KS, McDowell KJ. The effect of steroids on endometrial oxytocin mRNA production. J Reprod Fertil Suppl 2000;56:297–304. 26. Stout TAE, Lamming GE, Allen WR. The uterus as a source of oxytocin in cyclic mares. J Reprod Fertil Suppl 2000;56:281–7. 27. Watson ED, Buckingham J, Bjorksten TS, Nikolakopoulos E. Immunolocalisation of oxytocin in the uterus of the mare. J Reprod Fertil Suppl 2000;56:289–96.
28. Goff AK, Sirois J, Pontbriand D. Oxytocin stimulation of plasma 15-keto-13,14-dihydro prostaglandin F2a during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1987;35:253–60. 29. Stout TAE, Lamming GE, Allen WR. Oxytocin administration prolongs luteal function in cycling mares. J Reprod Fertil 1999;116:315–20. 30. Boerboom D, Brown KA, Vaillancourt D, Poitras P, Goff AK, Watanabe K, Dor´e M, Sirois J. Expression of key prostaglandin synthases in equine endometrium during late diestrus and early pregnancy. Biol Reprod 2004;70:391–9. 31. McDowell KJ, Adams MH, Adam CY, Simpson KS. Changes in equine endometrial oestrogen receptor alpha and progesterone receptor mRNAs during the oestrous cycle, early pregnancy and after treatment with exogenous steroids. J Reprod Fertil 1999;117:135–42. 32. Sharp DC, Thatcher M-J, Salute ME, Fuchs AR. Relationship between endometrial oxytocin receptors and oxytocininduced prostaglandin F2a release during the oestrous cycle and early pregnancy in pony mares. J Reprod Fertil 1997;109:137–44. 33. Starbuck GR, Stout TAE, Lamming GE, Allen WR, Flint APF. Endometrial oxytocin receptor and uterine prostaglandin secretion in mares during the oestrous cycle and early pregnancy. J Reprod Fertil 1998;113:173–9. 34. Spencer TE, Burghardt RC, Johnson GA, Bazer FW. Conceptus signals for establishment and maintenance of pregnancy. Anim Reprod Sci 2004; 82–83:537–50. 35. Sharp DC, McDowell KJ, Weithenauer J, Thatcher WW. The continuum of events leading to maternal recognition of pregnancy in mares. J Reprod Fertil Suppl 1989;37:101–7. 36. Stout TAE, Allen WR. Role of prostaglandins in intrauterine migration of the equine conceptus. Reproduction 2001;121:771–5. 37. Vanderwall DK, Rasmussen DM, Woods GL. Effect of repeated administration of oxytocin during diestrus on duration of function of corpora lutea in mares. J Am Vet Med Assoc 2007;231:1864–7. 38. Rivera Del Alamo MM, Reilas T, Kindahl H, Katila K. Mechanisms behind intrauterine device-induced luteal persistence in mares. Anim Reprod Sci 2008;107:94–106. 39. Wilsher S, Allen WR. Maternal recognition of pregnancy in the mare: Inhibition of luteolysis by intrauterine administration of plant oils. Pferdeheilkunde 2008;24:118 (abstract). 40. Weithenauer J, Sharp DC, McDowell KJ, Seroussi M, Sheerin P. Characterisation of the equine conceptus prostaglandin inhibitory product. Biol Reprod 1987;36 Suppl. 1, Abstract 329 41. Ababneh MM, Troedsson MHT, Michelson JR, Seguin BE. Partial characterization of an equine conceptus prostaglandin inhibitory factor. J Reprod Fertil Suppl 2000;56:607–13. 42. Watson ED, Sertich PL. Prostaglandin production by horse embryos and the effect of co-culture of embryos with endometrium from pregnant mares. J Reprod Fertil 1989;87:331–6. 43. Stout TAE, Allen WR., Prostaglandin E2 and F2a production by equine conceptuses and concentrations in conceptus fluids and uterine flushings recovered from early pregnant and dioestrous mares. Reproduction 2002;123:261–8. 44. Walters KW, Roser JF, Anderson GB. Maternal-conceptus signalling during early pregnancy in mares: oestrogen and insulin-like growth factor I. Reproduction 2001;121:331–8. 45. Zavy MT, Mayer R, Vernon MW, Bazer FW, Sharp DC. An investigation of the uterine luminal environment of
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Prostaglandins
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non-pregnant and pregnant pony mares. J Reprod Fertil Suppl 1979;27:403–11. Heap RB, Hamon MH, Allen WR. Oestrogen production by the preimplantation donkey conceptus compared with that of the horse and the effect of between-species embryo transfer. J Reprod Fertil 1991;93:141–7. Silva LA, Gastal EL, Beg MA, Ginther OJ. Changes in vascular perfusion of the endometrium in association with changes in location of the embryonic vesicle in mares. Biol Reprod 2005;72:755–61. Zavy MT, Sharp DC, Bazer FW, Fazleabas A, Sessions F, Roberts RM. Identification of stage-specific hormonally induced polypeptides in the uterine protein secretions of the mare during the oestrous cycle and pregnancy. J Reprod Fertil 1982;64:199–207. Allen WR, Stewart F. Equine placentation. Reprod Fertil Dev 2001;13:623–34. McDowell KJ, Sharp DC, Peck LS, Cheves LL. Effect of restricted conceptus mobility on maternal recognition of pregnancy in mares. Equine Vet J Suppl 1985;3: 23–4. Eilts BE, Scholl DT, Paccamonti DL, Causey R, Klimczak JC, Corley JC. Prevalence of endometrial cysts and their effect on fertility. Biol Reprod Mono 1995;1:527–32. ¨ Kollmann M, Bartmann CP, Schiemann V, Klug E, Ellenberger C, Schoon H-A. Hysteroscopic removal of uterine
53. 54.
55.
56.
57.
58.
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cysts in mares II: Follow-up and long-term fertility analysis with regard to patho-histological findings. Pferdeheilkunde 2008;24:35–7. Ousey J. Hormone profiles and treatments in the late pregnant mare. Vet Clin Equine 2006;22:727–47. Vivrette SL, Kindahl H, Munro CJ, Roser JF, Stabenfeldt GH. Oxytocin release and its relationship to dihydro-15keto PG F2a and arginine vasopressin release during partuition and to suckling in postpartum mares. J Reprod Fertil 2000;119:347–57. Word RA, Li XH, Hnat M, Carrick K. Dynamics of cervical remodeling during pregnancy and parturition: mechanisms and current concepts. Semin Reprod Med 2007;25:69–79. Rigby S, Love C, Carpenter K, Varner D, Blanchard T. Use of prostaglandin E2 to ripen the cervix of the mare prior to induction of parturition. Theriogenology 1998;50:897–904. Silver M, Barnes RJ, Comline RS, Fowden AL, Clover L, Mitchell MD. Prostaglandins in maternal and fetal plasma and in allantoic fluid during the second half of gestation in the mare. J Reprod Fertil Suppl 1979;27:531–9. Han X, Rossdale PD, Ousey J, Holdstock N, Allen WR, Silver M, Fowden AL, McGladdery AJ, Labrie F, Belanger A, Ensor CM, Tai H-H, Challis JRG. Localisation of 15-hydroxy prostaglandin dehydrogenase (PGDH) and steroidogenic enzymes in the equine placenta. Equine Vet J 1995;27:334–9.
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Chapter 172
Equine Chorionic Gonadotropin (eCG) Amanda M. de Mestre, Douglas F. Antczak and W.R. (Twink) Allen
Introduction One can imagine the air of excitement that would have pervaded the Department of Animal Science of the University of California at Davis in 1930 when Harold Cole and George Hart first discovered the existence of Pregnant mare serum gonadotropin (PMSG),1 nowadays more accurately termed equine chorionic gonadotropin (eCG). All the more so when it became apparent that eCG, in contrast to human chorionic gonadotropin (hCG),2 exhibited a mixture of both follicle-stimulating hormone (FSH) and luteinizing hormone (LH) biological activities and could therefore stimulate both follicle growth and ovulation of mature follicles in the ovaries of rats injected only once with a small volume of serum recovered from a pregnant mare between 40 and 100 days of gestation.3 In their classic paper announcing the discovery of eCG, Cole and Hart1 demonstrated, with remarkable accuracy, the general pattern of secretion of the hormone, with gonadotropic activity first appearing in peripheral serum between 37 and 41 days after ovulation and with levels rising steeply to reach a variable peak between days 60 and 80; thereafter, falling again steadily to disappear completely from the serum between days 120 and 160 (Figure 172.1).1 Cole and his colleagues in Davis remained active and pre-eminent in the general field of eCG during the next 30 years. They modified, refined, and increased the accuracy, sensitivity, and speed of their in vivo bioassay method to measure concentrations of eCG in serum and partially purified extracts of hormone;4–6 investigated thoroughly the essential physicochemical7 and biological3, 8, 9 properties of the hormone; examined the relationships between the secretion of eCG and other morphological and endocrinological events in the pregnant mare;10–13 and
studied, and eventually discovered, the source of eCG during pregnancy.14, 15 They also determined the half-life of the hormone in horse blood16, 17 and confirmed the finding of the team led by the late Professor Wadslaw Bielanski18 in Poland of the profound influence of fetal genotype on the rate and duration of eCG production in early pregnancy.19 This chapter is dedicated to the memory of Professor Harold H. Cole and his merry band of co-workers who made such valuable use of their good fortune to have lived and worked in the golden age of endocrinological discovery.
Physicochemical properties of eCG eCG is a high molecular weight (72 000) acidic glycoprotein with an abnormally high carbohydrate content (45%), consisting mainly of sialic acid, galactose, and glucosamine.8,20–41 It has a low isoelectric point of only 2.4 and is remarkably resistant to acid hydrolysis.21, 22 Owing to its high degree of glycosylation, eCG is highly negatively charged. eCG was traditionally extracted from pregnant mares’ serum by initial precipitation with metaphosphoric acid or acetone, followed by further precipitation ‘cuts’ with graded concentrations of alcohol.20, 24 The biological value of the extracted protein and its scale of production has since been improved using direct absorption onto anionic exchange resins after initial conditioning of the plasma to reduce its pH and ionic strength.42 Further purification using cation exchange column chromatography yields a final highly purified extract which exhibits a specific gonadotropic potency of 13 500–15 000 IU/mg, or 4000 IU/mg protein.22, 24, 29, 31, 32, 42 Serum purified eCG is available commercially in many continents of the world marketed as Folligon (Intervet Laboratories, Boxmere, The Netherlands) and Pregnecol 6000
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 172.1 Concentrations of equine chorionic gonadotropin (eCG) measured by hemagglutination-inhibition assay in the serum of 9 pregnant pony mares.
(Bioniche, Belleville, Ontario, Canada). Although recombinant eCG has not yet been synthesized in commercial quantities, significant progress has been made in recent years, which suggests that it may well be produced in the not too distant future.43, 44 Recombinant eLH/CG can be produced in both mammalian and insect cells. The mammalian cells produce a recombinant protein which possesses both FSH and LH biological activities,45 but the total amount of protein produced is somewhat limited. Furthermore, eCG produced in insect cells displays a reduced molecular weight and degree of glycosylation compared with the native protein, and therefore lacks biological activity in vivo.44 The recognized international standard for eGG is a partially purified extract from horse serum maintained at the National Centre for Medical Research at Mill Hill in London. It was prepared in the early 1960s and assayed by 16 different laboratories using a variety of in vivo bioassay methods, and the product was assigned a potency of 1600 IU per vial containing 1.6 mg protein.23 A new batch of partially purified eCG (eCG NZY-01) was assayed by eight laboratories in 1998 and is currently available in vials of 500 IU.46 Pituitary and chorionic gonadotropins and thyroid-stimulating hormone (TSH) are heterodimeric glycoproteins that consist of α and β subunits joined non-covalently.26 The α subunits are identical within each animal species, whereas the β subunits differ, indicating that the specificity of action of gonadotropins resides in the β subunit. The equine α subunit is encoded by a single gene,40 which arose originally from gene duplication of a common α/β gene.47 The equine α subunit gene encodes for a 96-amino-acid protein that shows 68–79% homology
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with the other available mammalian α subunit sequences. It contains two N-linked oligosaccharide chains at positions Asn56 and Asn82 and possesses several unique substitutions, including transposition between tyrosine and histidine at positions 87 and 93,30 which may be related to some of the unusual properties of the equine gonadotropins.40 In general, each β subunit is encoded by a single gene which, together, form a family of closely related genes that have also arisen through gene duplication.47 Equids are an exception, with a single gene encoding for the β subunit of both eLH and eCG.39, 41 This indicates that modifications, not duplication, has led to placental expression of eCG. Importantly, this is in contrast to the β subunit of hCG, which is encoded by four highly homologous placental CG β subunit genes, CGB, CGB5, CGB7, and CGB8. An additional two hCG β subunit genes are located within this genome cluster, but the function of these truncated gene copies are unknown.48 These genes are all distinct from the pituitary expressed gene that encodes for human LHβ. The equine LH/CGβ gene encodes for a protein of 169 amino acids that shows 56–78% homology with other mammalian β subunit sequences. Processing results in the cleavage of a 20-amino-acid signal peptide at the N-terminus, resulting in the mature 149amino-acid protein. Horse and human CG β subunits are longer at the C-terminal by 30 residues than all of the pituitary β subunits. These C-terminal peptide (CTP) extensions have arisen as a result of gene mutations (a single base deletion in primates and a 10-base pair (bp) deletion in equids) that have occurred in the same area of the gene in each case. The function of the β subunit C-terminal extension in primates and equids remains a mystery but it seems likely to be related in some way to expression and/or secretion of the hormone in placental tissues. It has been shown to confer FSH bioactivity to the protein. As mentioned, eCG contains a much higher carbohydrate content than any of the other known mammalian pituitary and CGs, including hCG and eLH.20, 23, 28 Recent studies have shown that around 20% of the α subunit and > 50% of the β subunit of eCG comprises carbohydrate moieties, each having a mixture of N- and O-linked carbohydrate chains.49–51 In addition to having a higher carbohydrate content, a much higher proportion of this carbohydrate in eCG is in the form of sialic acid than in the gonadotropins of other species.28 Although eCG and eLH are encoded by the same genes, their β subunits display quite different chemical compositions (Table 172.1). The eCG β subunit contains a single N-linked glycosylation site at Asn13 , and 12 O-glycosylation sites in the CTP, four of which are highly glycosylated at Thr133 , Ser137 , Ser140 , and Ser141 .
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Table 172.1 Physiochemical characteristics of equine luteinizing hormone (eLH) and equine chorionic gonadotropin (eCG) β subunits Characteristics
eCG
eLH
Amino acids Molecular weight CHO content CHO moieties C-terminal peptide N-glycosylation O-glycosylation
149 44 kDa 45% Sialylated 30 amino acids Asn13 12 sites
149 34 kDa 30% Sulfated and sialylated 30 amino acids Asn13 12 sites
% O-glycosylation
eCG increased compared with LH at Thr133 , Ser137 , Ser140 , Ser142
O-glycan oligosaccharides FSH:LH LH receptor binding Half-life
GlcNac 1.0 Low affinity 6 days
GalNac, Gal, GalNac 1.0 High affinity (×10) 5 hours
FSH, follicle-stimulating hormone.
Biological properties of eCG Perhaps the most unusual and interesting feature of the eCG molecule is its unique ability to express both FSH-like and LH-like biological activities when administered to other mammalian species.1, 8 This property is in contrast to hCG and the CGs of other primate species, which can induce only LH-like responses.52 Human menopausal gonadotropin (hMG) likewise expresses dual FSH-like and LH-like activities in vivo, but this arises from a mixture of pituitary FSH and LH which are secreted in large quantities when the pituitary gland escapes in older age from the negative feedback effects of ovarian steroids.53 The first indication that donkey CG also exhibits FSH-like activity in non-equine species came with the development of the very useful radioreceptor assay technique. Employing rat testicular FSH and LH receptors and a variety of radioactively labeled gonadotropins,54 donkey CG was shown to possess significant, but considerably less, FSH receptor binding activity than eCG. For example, the FSH:LH ratio of eCG is approximately 1.0 compared with 0.1 for donkey CG, and the horse × donkey hybrid conceptuses (mules and hinnies) produce an FSH:LH ratio midway between the two parental extremes (i.e., around 0.5–0.7).55–57 A number of studies have confirmed that donkey CG expresses much less FSH-like activity than eCG,34, 55, 58 and that zebra CG is more like donkey CG than eCG. Donkey LH has also been shown to be chemically and biologically similar to donkey CG,56, 57 which is not surprising because they are almost certainly, as in the horse, encoded by the same genes.59 Donkey CG has a C-terminal extension
similar in length to eCG58 but, owing to a frameshift in the DNA sequence, shows only 21% amino acid sequence homology. The structural basis for the dual biological, and therefore dual receptor binding, activities of eCG (and LH) remains unclear. It presumably lies within the β subunit, which, apart from the C-terminal extension, is understandably LH-like, showing approximately 70% homology with other species’ LH β sequences, including hCG. Its ability to bind to FSH as well as LH receptors in non-equine species is, therefore, probably the result of minor amino acid differences within the β subunit. Although the carbohydrate moieties of the gonadotropin molecules contribute to biological half-life, and possibly receptor binding affinity and/or signal transduction, evidence does not exist to support their involvement in receptor binding specificity (Table 172.1). Donkey and horse eCG/LH possesses differing degrees of FSH activity and this property has been exploited to identify the amino acids responsible for FSH activity. It was initially thought that the β subunit alone conferred FSH activity,45 but more recent studies suggest that both α and β subunits are required for the FSH activity of eCG/LH.60 Chimeric and mutagenesis studies have identified residues Val102 –Phe103 –Arg104 and the CTP as key amino acids required for FSH activity45, 60, 61 possessed by eCG/ LH, as mutations in these sites result in a five- to eightfold loss of FSH activity. Biological activity is also negatively regulated by low pH, a decrease in non-covalent links or sialic acid content, and/or modifications by oxygen free radicals.62, 63 The relatively constant levels of serum eCG can be attributed to both the long half-life of the hormone and the constant release of eCG by endometrial cup cells. Previously, the CTP was thought to contribute to the half-life, but more recent evidence does not support this. Rather, it is the molecular weight, negative charge, and sialic acid content that confers the 6-day half-life of eCG. The FSH-like properties of eCG have resulted in the widescale use of partially purified extracts of the hormone as the cheapest and most readily available form of exogenous gonadotropin to stimulate follicular growth and ovulation in laboratory animals and in the larger domestic and farm animal species. It can be used at lower doses as an adjunct to progestagen withdrawal or other forms of synchronization of estrus to induce normal rates of ovulation in adult or prepubertal animals. Alternatively, when administered in higher doses, its FSH-like component stimulates the development of multiple follicles, which are then ovulated by the LH-like component to give the superovulation that is required in embryo recovery and transfer programs.64, 65 In this latter use,
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Equine Chorionic Gonadotropin (eCG) however, the exceptionally long half-life of the eCG molecule can be disadvantageous in that a single injection of hormone can stimulate two or more asynchronous waves of follicular growth in the animal’s ovaries. Furthermore, the LH-like component of the eCG may ovulate a partly mature follicle existing in the ovaries of the recipient at the time of the eCG injection, and the progesterone secreted by the resulting corpus luteum can interfere with ovulation of the subsequent wave of follicles stimulated by the FSHlike component of the eCG.66 Because of these difficulties, most modern embryo transfer programs in cattle prefer to treat donor animals during days 10–14 of diestrus with repeated injections over a 4-day period of lower doses of purified pituitary FSH, with a luteolytic dose of prostaglandin F2α given 48 hours after starting the gonadotropin therapy.67, 68
Source of eCG in the mare Cole’s group discovered the source of eCG in the pregnant mare, the unique and still puzzling placental outgrowths now known universally as endometrial cups,15 but this important finding was not made without some problems along the way. Two of Cole’s graduate students, Hubert Catchpole and Bill Lyons, carried out an exhaustive study over 2 years that involved painstaking dissections of pregnant horse uteri recovered from abattoirs and crude saline extracts of known weights of fetal, placental, and endometrial tissues taken from various parts of the uterus (H.R. Catchpole, personal communication). They submitted the extracts to their rat utero-ovarian weight bioassay to measure gonadotropic activity and found higher concentrations of hormone at 40–80 days of gestation in the endometrium of the gravid uterine horn in direct contact with the developing fetal membranes than in the empty non-gravid horn. Furthermore, because they did not specifically identify and assay the endometrial cups within the selected samples of endometrium, they tended to find higher levels of gonadotropic activity in extracts of the allantochorion than in those of the endometrium. They concluded, quite reasonably, that, like hCG in human pregnancy,69 eCG was probably secreted by the trophoblast cells of the developing placenta to be taken up by, and accumulated selectively in, the underlying endometrium.14 The subject rested at that point until, 9 years later, Cole came again to the fore with his ‘rediscovery’ of the endometrial cups. Some 40 years previously the German anatomist W. Schauder had mentioned the existence of a series of saucer-shaped endometrial protuberances in the gravid uterine horn of mares
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during the first half of pregnancy,70 which appeared to have no physical connection to the overlying fetal membranes but which accumulated a honey-colored, sticky secretion in their central crater-like depression. Schauder gave them the name endometrial cups. Cole and Goss looked afresh at Schauder’s endometrial cups and, to their surprise, found that the cup tissue itself and the associated exocrine secretion both contained concentrations of eCG that were orders of magnitude higher than those in the adjacent normal endometrium, or in any parts of the fetal placenta.15 They, therefore, concluded that the endometrial cups were the real source of eCG and that the hormone entered the maternal bloodstream via a complex of large lymph sinuses which developed in the endometrial stroma beneath each cup.15 Thus, it now seemed that eCG, unlike hCG in primates, was actually maternal in origin. Two further elucidative errors were to be made before the real source of eCG was to come to light. In their next substantive paper on the subject, the Davis group gave a detailed morphological description of the development and regression of the endometrial cups between days 40 and 150 of gestation. They noted that each mature cup at around days 60–70 consisted of a mass of large epithelioidtype cells that were similar in appearance to the large glycogen-filled decidual cells of rodent and human placentas,71 occasional blood vessels and distended endometrial glands that were interspersed throughout the tight knot of decidual-like cells. Because the cups appeared to have no direct physical connection to the overlying epitheliochorial placenta, Clegg and colleagues72 concluded that the decidual-like cells were maternal in origin, probably having differentiated from endometrial stromal cells in a manner similar to the development of the decidual reaction in pregnant or pseudopregnant rodent uteri. Furthermore, the epithelial cells lining the dilated endometrial glands within each cup and the exocrine secretion within these glands and accumulated on the surface of the cup stained intensely with periodic acid–Schiff (PAS), thought at that time to be specific for glycoproteins. Since eCG was known to be a large glycoprotein molecule with an unusually high carbohydrate content, the Davis group concluded, very reasonably, that eCG was probably secreted by the epithelium of the dilated glands within the cup. Twenty years later, the true picture at last emerged when W.R. Allen and his colleague R.M. Moor dissected chorionic girdle tissue from horse conceptuses recovered intact at 34–36 days of gestation by surgical hysterotomy, and cultured in vitro the minced girdle tissue as simple explants. To their surprise, this produced stable colonies of large, epithelioid, binucleated cells that secreted high concentrations of eCG
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into the culture medium for > 180 days. The cultured girdle cells showed all the morphological characteristics of endometrial cup cells in vivo and they transformed similarly when transplanted to extrauterine sites such as the testis.73 Subsequently, a light microscopic and ultrastructural study of the placental interface carried out in mares between 34 and 40 days of gestation demonstrated invasion and migration of the specialized chorionic girdle cells into the maternal endometrium during days 36–38, followed by their rapid transformation into the large eCGsecreting decidual-like cells of the endometrial cup.74 Thus, the original conclusions of Catchpole and Lyons14 were vindicated. Fetal trophoblast cells do indeed secrete eCG, but only those specialized ones of the chorionic girdle after they have migrated into the endometrium around day 36 to form the endometrial cups.
Development and invasion of the chorionic girdle The chorionic girdle was first described by Cossar Ewart in 1897 as constituting a ‘complex whitish band, nearly a quarter of an inch in width, placed
21 DAYS
nearly equatorially on the embryonic sac which is concerned in fixing the embryo to the uterine surface and which may be a means of absorbing additional nourishment’.75 The girdle was not really mentioned again in the literature until remarked on by van Niekerk in 1965,76 and was then described in more detail by van Niekerk and Allen77 following the discovery of its role in endometrial cup formation. The girdle comprises a discrete thickening of the trophoblast that develops around the circumference of the spherical conceptus at the point where the enlarging allantoic and regressing yolk sac membranes abut each other (Figures 172.2 and 172.3). It first appears histologically at around day 25 after ovulation as a series of shallow ridges in the single-cell layer of trophoblast (Figure 172.4a). By days 30–32 of gestation, the trophoblast cells on the tops of these folds are proliferating rapidly, leading to the formation of ridges and intervening grooves that grow and deepen to form elongated flap-like projections (Figure 172.4b). These tend to become flattened on their apical surface by the pressure of the opposing endometrium, thereby creating a thickened, stratified type of trophoblast some 8–10 cells deep that forms simple gland-like structures created by the surface 28 DAYS
Bilaminar omphalopleure
Sinus terminalis
Bilaminar omphalopleure
Sinus terminalis Yolk sac
Trilaminar omphalopleure
Trilaminar omphalopleure
Progenitor chorionic girdle
Yolk sac
Allantois
Allantois Amnion 35 DAYS Chorionic girdle Yolk sac
Allantochorion Figure 172.2 Diagrammatic representation of the development and differentiation of the extraembryonic membranes of the equine conceptus between days 21 and 35 after ovulation. Open arrows show direction of growth of the allantois. Adapted with permission from Steven128 with permission from Elsevier.
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Equine Chorionic Gonadotropin (eCG)
Figure 172.3 A horse conceptus on day 34 after ovulation. The chorionic girdle (CG) is seen as the pale annulate thickening of the trophoblast around the conceptus between the enlarging allantochorion (ALC) and regressing yolk sac (YS). The sinus terminalis and persisting region of non-vascularized bilaminar omphalopleure (BO) lies beneath the fetus (F).
epithelium of adjacent folds (Figure 172.4c). These simple glands appear to secrete a mucoid exocrine material that may help to bind the luminal surface of the girdle to the overlying endometrium. From
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around day 32, the rapidly proliferating uninucleate trophoblast cells of the chorionic girdle begin to differentiate into non-dividing binucleate cells that are capable of synthesizing eCG (Figure 172.4d).78, 79 Between days 36 and 38, the hyperplastic, now binucleate, trophoblast cells on the surface of the girdle begin to vigorously invade and destroy the luminal epithelium of the endometrium by extending blunt pseudopodia-like processes that force their way between, and sometimes straight through, the maternal epithelial cells. When they reach the basement membrane, they continue to migrate down the mouths of endometrial glands, dislodging the glandular epithelium from its basement membrane as they go (Figure 172.5a). After a brief delay the girdle then breaches the basement membrane by the same method of pseudopodium extrusion and the girdle cells stream off into the endometrial stroma, both vertically downwards from the luminal surface and laterally from the lumenae of the glands (Figure 172.5b). The migrating girdle cells disrupt lymphatic vessels, but there is surprisingly little disruption to the rich subepithelial network of endometrial capillaries.80 Within as little as 24–48 hours after entering the stroma, the girdle cells cease to migrate, and
(a)
(b)
(c)
(d)
Figure 172.4 Histological sections showing the development of the chorionic girdle, progenitor of the endometrial cups on days (a) 25, (b) 30, (c) 34 and (d) 36 after ovulation. Note the intense hyperplasia of the trophoblast cells at the tips of the elongated folds in (d). (See color plate 60).
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(a)
(c)
(e)
(b)
(d)
(f)
Figure 172.5 Histological sections of endometrial cups at different stages of development. (a) Day 38; showing the chorionic girdle attached to, and beginning to invade, the endometrium; the luminal epithelium and the epithelium of some glands have been dislodged and replaced by invading girdle cells. Lymphocytes are already accumulating in the subepithelial stroma (100×). (b) Day 40; the invading binucleate girdle cells have broken through the basement membranes of the luminal epithelium and endometrial glands and are now streaming out into the endometrial stroma. (c) Day 50; the young cup is now a solid mass of large equine chorionic gonadotropin-secreting binucleate cells and the luminal surface is already covered by a regenerated layer of endometrial epithelium; some endometrial glands are beginning to become distended with accumulated exocrine secretion and appreciable numbers of lymphocytes are accumulated at the periphery of the cup (260×). (d) Day 65; some endometrial cup cells are just beginning to degenerate at the luminal surface of the cup, where a small amount of exocrine secretion is trapped beneath the overlying allantochorion; lymphocytes and plasma cells are forming a definite band at the periphery of the cup (100×). (e) Day 93; the accumulated leukocytes are moving into the cup tissue and actively killing the large cup cells at the base of the cup. (f) Day 120, a distinct line of separation shows the completely necrotic cup about to be sloughed off the surface of the endometrium. (See color plate 61).
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Equine Chorionic Gonadotropin (eCG) round up, enlarge greatly, and become tightly packed together in the endometrial stroma, to create the typical endometrial cups (Figure 172.5c).74 Exactly how the tightly controlled processes of proliferation, differentiation, and subsequent migration of chorionic girdle trophoblast cells are regulated is still unclear.80 Several growth factors are expressed by the tissues that abut the chorionic girdle and thus are well positioned to regulate its development. Hepatocyte growth factor-scatter factor (HGF-SF) is expressed exclusively by allantoic mesenchyme and mesothelial cells that underlie the chorion and chorionic girdle,81 and epidermal growth factor (EGF), vascular endothelial growth factor (VEGF) and transforming growth factor β (TGF-β) are all expressed by pregnant endometrium.82–84 Stewart et al.81 proposed a model for development of the chorionic girdle whereby secretion of the highly mitogenic and motogenic HGF-SF exclusively by allantoic mesoderm stimulates multiplication of the overlying chorionic trophoblast cells (Figure 172.6). Where these are firmly attached to the basement membrane of the allantochorion, expansion of the allantochorion occurs and this portion of the conceptus enlarges relative to the underlying choriovitelline membrane. At the junction of these chorioallantoic and choriovitelline membranes, the chorion and trophoblast cells are similarly stimulated by HGF-SF secreted by the nearby allantoic mesoderm and, since they are not firmly attached to underlying allantoic tissue, they multiply and pile up on one another, rather than expanding laterally, thereby giving rise to the thickened chorionic girdle (Figure 172.3). Furthermore, the mitogenic properties of the HGF-SF impose an invasive phenotype upon the rapidly multiplying trophoblast cells of the chorionic girdle, thereby driving the invasion of the chorionic girdle cells into the endometrium to form the endometrial cups. This theory of chorionic girdle formation was greatly supported by the finding of high concentrations of c-met, the active receptor for HGF-SF, on chorionic girdle cells (Figure 172.6).81 Furthermore, metalloproteinases, degradative enzymes that confer a migratory phenotype in other cell systems, are secreted by the chorionic girdle.85 Recently, the authors described a new method for culturing chorionic girdle cells that promotes the formation of three-dimensional spherical vesicles.86 The cells in the outermost layer of these vesicles are polarized, columnar in shape, and possess microvillous projections, all of which are morphological features displayed by chorionic girdle cells just prior to their invasion of the endometrium. The chorionic girdle has also received attention because of its immunological features. Just prior to invasion of the endometrium the trophoblast cells of the girdle express high levels of both paternally and
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ac HGF-SF c-met t
am
f
alc cg
al
am c
ysc t
Figure 172.6 Diagrammatic representation showing the hypothesized secretion of hepatocyte growth factor-scatter factor (HGF-SF) by the allantoic mesoderm (am) acting upon the attached trophoblast (t) of the allantochorion (alc) to stimulate conceptus expansion and diffusing across to stimulate the unattached chorion above the vitelline membrane (yolk sac) to cause the chorion cells (c) to pile up on themselves to form the thickened chorionic girdle (cg) which expresses high concentrations of c-met, the receptor for HGF-SF. Allantois (al) and yolk sac cavity (ysc). Reproduced with permission from Stewart et al.81 , with permission from the Society for the Study of Reproduction.
maternally derived polymorphic major histocompatibility complex (MHC) class I molecules.87–90 Furthermore, shortly after invasion of the fetal girdle cells into the maternal endometrium, the mare routinely mounts strong humoral antibody responses to these paternally derived MHC alloantigens.91 This apparently unique ability of chorionic girdle cells to actively simulate the maternal immune system remains one of the paradoxes of equine pregnancy. It is in marked contrast to trophoblast cells of other
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mammalian species which primarily express nonpolymorphic MHC molecules and are therefore able to evade immunological detection more readily.
Development and regression of the endometrial cups The young endometrial cup around day 50 of gestation now consists of a dense mass of the large, trophoblast-derived binucleate cup cells with little or no stroma between them and only occasional blood and lymph vessels (Figure 172.5c).72, 80, 92 The apical portions of the endometrial glands have been obliterated during the original invasion by the chorionic girdle but the fundic regions of the glands remain intact and they become increasingly distended throughout the lifespan of the cup. This glandular enlargement seems to occur both as a result of blockage of the apical outlets to cause accumulation of the excessive secretions, and a very pronounced hyperstimulation of the secretory epithelium. Large quantities of carbohydrate-rich, PAS-positive material accumulate within the gland lumenae before being released onto the surface of the cup when the latter begins to degenerate after about day 70 of gestation.92 Between days 55 and 75, the mature endometrial cups form a circle or horseshoe of pale, raised plaques on the surface of the endometrium at the base of the gravid uterine horn. Individual cups vary tremendously in size, ranging from small isolated mounds measuring as little as 1.0 cm in diameter, to long, unbroken ‘ribbons’ of cup tissue measuring 1.0–1.5 cm in width and > 10 cm in length (Figure 172.7a,b). This variability occurs within and between mares and it seems to have two interacting causes. First, there is appreciable variation between conceptuses in the width and overall development of the progenitor chorionic girdle which, in turn, affects the breadth and overall sizes of the endometrial cups that develop following invasion of the endometrium by the chorionic girdle around days 36–38.92, 93 These differences in girdle width and general development are most marked when mating across the equine species to produce hybrid interspecific conceptuses, and when using embryo transfer to create true extraspecific pregnancies.93, 94 However, there is also circumstantial evidence of genetic factors that may influence chorionic girdle development and the size of the resulting endometrial cups within normal intraspecific horse matings.95 A second factor influencing endometrial cup size, and hence the amount of eCG secreted into maternal blood, is the surface architecture and tone of the maternal endometrium at the time of girdle invasion. The chorionic girdle is held closely against the endometrium by the dual actions of increased myome-
trial tone and expansion of the developing fluid-filled conceptus. The surface of the endometrium to which the girdle becomes attached can vary from highly folded and ridged (antimesometrial) to flattened and almost smooth (mesometrial). Should the girdle lie apposed to the ridged surface at the time of invasion it will be able to gain access only at the apices of the ridges, thereby leaving the troughs between them uninvaded. Subsequently, as the uterus expands further with increasing gestation, the ridges will tend to flatten out, so leaving isolated, discrete cups (Figure 172.7b). On the other hand, should the chorionic girdle invade an area of smooth endometrium at day 36, unbroken lengths of girdle will invade, giving rise to continuous unbroken ribbons of endometrial cup tissue (Figure 172.7a).93 Not surprisingly, mares which conceive at the 9-day foal heat are noted to have higher serum concentrations of eCG than foaling mares that conceive at subsequent estrous periods.96 Incomplete involution of the uterus, and, subsequently, an enlarged endometrial surface area at the time of chorionic girdle invasion, may explain this observation. In the mature endometrial cup at around days 60–75, the transformed eCG-secreting fetal cup cells are very large, essentially round, and they are all binucleated. These nuclei are large euchromatic bodies that contain dense, characteristic nucleoli (Figure 172.5c). The cytoplasm is filled with short profiles of endoplasmic reticulum which resemble granules, some lipid droplets are present, and a few mitochondria are scattered throughout the cell. The Golgi apparatus is a significant structure and many smooth-surfaced Golgi-associated vesicles are evident. Nonetheless, the cup cells contain no obvious dense secretory granules and secretion, therefore, appears to be constitutive rather than regulated like the pituitary gonadotropins. It is of interest that chorionic girdle cells recovered from mares prior to their invasion of the endometrium and cultured in vitro appear to undergo the same transformation processes as in vivo. They form monolayer colonies of binucleated cells which show the same morphological features as mature endometrial cup cells in the mare.86, 97 With increasing time in culture some of the binucleate cells will coalesce to form a type of multinucleated syncytium.73 A striking feature of the endometrial cup reaction is the associated maternal leukocyte response. This is first observed at the time of the initial invasion of the chorionic girdle and is predominantly composed of lymphocytes.89, 98 While the cups are young and immature, the accumulated lymphocytes lie between the migrating trophoblast cells as well as the periphery, where they are located in clusters that surround the endometrial glands (Figure 172.5c). The
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Equine Chorionic Gonadotropin (eCG)
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Figure 172.7 (a) A continuous band of young endometrial cup in the gravid uterine horn of a pony mare on day 45 of gestation; the cup appears as a pale, raised plaque on the endometrial surface. (b) A mass of mature endometrial cups at day 60 of gestation; note the great variations in size and shape of the individual cups. (c) Mature endometrial cups at day 73 of gestation; the allantochorion is pulled back to expose the saucer-shaped cups, the central depressions of which are filling with accumulated exocrine secretion mixed with cell debris. (d) Degenerating endometrial cups at day 95 of gestation; the cups are pale and ‘cheesy’ in appearance and significant quantities of their sticky, honey-colored, equine chorionic gonadotropin-rich exocrine secretion are adhered to the overlying allantochorion. (e) The fetal surface of the placenta of a mare at term showing two pendulous allantochorionic pouches (arrow) filled with necrotic endometrial cup tissue and inspissated endometrial cup secretion.
lymphocyte populations associated with young endometrial cups primarily consist of CD4+ lymphocytes (55–65%), CD8+ lymphocytes (15–20%), and B cells (10–15%).96, 98 As gestation proceeds, the lymphocytes become confined to the periphery of the
cups and are reduced in numbers. Small clusters of B cells and the intermittent macrophages are also present during these developmental stages. As the cups begin to degenerate and regress, peripherally located CD4+ and CD8+ lymphocytes are joined by
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clusters of eosinophils and larger clusters of B lymphocytes and, collectively, may begin to infiltrate the dying cup tissue (Figure 172.5d,e). In focused areas of necrosis, neutrophils can also be seen.98 As this combination of degenerative changes progresses, sloughing of necrotic cup cells from the luminal surface increasingly unblocks the outlets of distended endometrial glands. The accumulated secretion exudes from the glands and, mixing with the disintegrating cup cells, becomes increasingly rich in eCG activity and progressively more inspissated, glutinous, and sticky. It fills the central depression on the surface of the cup and adheres to the overlying allantochorion (Figure 172.7c,d). A distinct line of separation forms between the periphery of the necrotic cup tissue and the endometrial stroma packed with accumulated leukocytes (Figure 172.5f) until, eventually, between days 100 and 140, but with considerable variation, both between mares and between individual cups within the same mare, the necrotic lump of cup debris and admixed glutinous cup secretion is sloughed completely from the endometrial surface. In areas of the uterine horn in which the dehisced material lies above the conceptus, gravity causes the lump to invaginate into the allantochorion to give rise to the pedunculated structures termed allantochorionic pouches.72 These hang into the allantoic cavity (Figure 172.7e). Historically, the inability of the endometrial cups to resist the strong maternal immune cell infiltration was thought to be the main explanation for the restricted lifespan of the cups. However, recent studies suggest that the mechanisms involved are more complex than this. First, endometrial cup cells downregulate the MHC class I molecules expressed by their girdle progenitors so that little MHC expression is detectable beyond day 44 of gestation;99 the cup cells do not express MHC class II molecules at any stage.88 This distinct absence of detectable paternal alloantigen after establishment of the cups assists the cup cells to evade antigen-specific killing by lymphocytes. Second, the CD4+ lymphocytic infiltration around the endometrial cups includes not only traditional T helper lymphocytes, but also regulatory T lymphocytes that possess the ability to functionally suppress immune responses.100 Third, the fetal endometrial cup cells possess an inherent ability to modulate immunity; secretory products of chorionic girdle cells suppress lymphocyte proliferation and cytokine expression.101 In addition, chorionic girdle, in the form of either pieces of tissue or cultured chorionic girdle vesicles, survives after allogeneic transplantation into sites outside the uterus in non-pregnant mares.86, 102 Independent of their location, transplanted chorionic girdle cells share striking similarities to endometrial cups, including their
secretion of eCG and the ability to resist destruction by cell-mediated and humoral antibody responses amounted by the recipient. This ability to survive and function outside the pregnant endometrium, and independent of the hormonal milieu of pregnancy, suggests that binucleate cells are ‘master’ regulators, ultimately controlling their own fate. In summary, the equine endometrial cups comprise, in effect, a precisely timed injection of specialized trophoblast cells into the maternal endometrium where they secrete large quantities of gonadotropic hormone and stimulate strong humoral and cellmediated maternal immune responses. Yet, despite this strong sensitization of the mare to foreign antigens expressed by this invasive component of the fetal placenta,103 no similar response is generated against the non-invasive trophoblast of the allantochorion, which allows the conceptus to be carried safely to term.104
Functions of eCG in the mare Cole and his colleagues drew attention to the close relationship between the secretion of eCG and the considerable degree of secondary luteal development that occurs in the mare’s ovaries between 40 and 150 days of gestation.10 Amoroso et al.105 described the same phenomenon and Allen106 counted as many as 35 distinct luteal structures in the ovaries of one mare on day 140 of gestation. This coincidental development, persistence, and regression of endometrial cups and secondary corpora lutea in mares between 40 and 150 days of gestation led to a general assumption that eCG provided both the gonadotropic and the luteotrophic stimuli for these accessory luteal structures. However, subsequent studies showed that the continued 10- to 12-day rhythmical releases of pituitary FSH which continue in the mare during early pregnancy, just as during the estrous cycle,107, 108 stimulate waves of follicular growth in her ovaries, and it is just the LH-like component of eCG which brings about the ovulation and/or luteinization of the most advanced follicle in each wave. In this way, a crop of accessory luteal structures is built up as gestation advances (Figure 172.8). More recent studies have shed fresh light on the function of eCG during equine pregnancy. The gonadotropin stimulates the luteal cells of both primary and secondary corpora lutea to secrete both progesterone and estrogen.109, 110 Thus, concurrent with the onset of eCG secretion, an increase in P45017α and a decrease in P450(arom) secretion rates in the luteal cells promote luteal secretion of estrogens. eCG exhibits a much lower capacity to bind to horse gonadal receptors (< 2%) than does either horse or human pituitary FSH and LH, although it
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Figure 172.8 Peripheral plasma progesterone concentrations measured in two pony mares during the first 120 days of gestation. The period of equine chorionic gonadotropin secretion is shown diagrammatically in hatched profile. Note the accumulative rises in progesterone concentrations, beginning around day 40, from successive development of secondary corpora lutea.
binds with equal avidity to them and other pituitary gonadotropins to the gonadal tissues of other species.111 Thus, instead of gross stimulation occurring around day 60 of gestation when eCG concentrations in serum are at their peak, only the single ovulations described above, which result essentially from the pituitary FSH-stimulated waves of follicular growth, continue to occur. A noteworthy exception to this norm occurs when the Jenny donkey carries either an interspecies hinny conceptus, or a transferred extraspecies horse-in-donkey conceptus. In both these situations, endometrial cup size, eCG secretion rate, secondary luteal development in the ovaries, and plasma progesterone concentrations between days 40 and 120 of gestation are all much higher than in donkeys carrying normal intraspecific donkey conceptuses.93,112–114 However, this breakthrough in ovarian responsiveness is explained not just by the increased amounts of gonadotropin produced by the larger-than-normal endometrial cups in the unusual pregnancies. Rather, it relates to the much higher ratio of FSH:LH in the CG produced by the hinny conceptus than in a normal donkey conceptus.55 Thus, when the female donkey, the ovaries of which are prepared genetically to encounter low levels of a CG containing very little FSH-like activity, are suddenly faced with greatly increased levels of CG with a much higher FSH-like content produced by the hinny and horse conceptuses, the normal protective barrier in the ovaries is overcome.113 One further point in relation to the function of eCG in the pregnant mare is its possible role in stimulating the growth of, and steroid production by, the fe-
Figure 172.9 The grossly enlarged fetal ovaries and inactive maternal ovaries recovered from a mare at day 253 of gestation.
tal gonads. As highlighted originally by Cole et al., 12 and studied by many others since then, the gonads of both male and female equine fetuses undergo tremendous hypertrophy between days 100 and 240 of gestation, followed by regression and shrinkage to normal size at birth (Figure 172.9).115 This is mirrored by an equally striking increase and subsequent decline in the concentrations of both phenolic (estrone and estradiol) and ring B unsaturated (equilin and equilenin) estrogen concentrations in maternal blood and urine,116, 117 and firm evidence now exists that the enlarged gonads secrete 19-carbon steroid precursors, such as dehydroepiandrosterone (DHEA) and 7α hydroxyDHEA, which are then aromatized to estrogens by the placenta.118–121 It would be easy to assume that eCG might be the essential gonadotropic stimulus for this enormous fetal gonadal growth, but limited available evidence suggests that this is unlikely to be the case. The gonads really only begin to enlarge significantly from days 100–120, when eCG levels in maternal blood have already fallen dramatically. Furthermore, as shown originally by Cole and Saunders,122 and confirmed subsequently by Rowlands,123 eCG is selectively excluded from placental absorption and this hormone remains virtually undetectable in fetal blood and fluids. On the other hand, the more recent studies124 have shown that the eLH/CG receptor is expressed by fetal gonad cells as early as day 44 of gestation, which re-ignites discussion on whether or not eCG may play a role in the transition of steroidogenesis from the corpus luteum to the fetoplacental unit.
Abnormal persistence of eCG secretion For many decades conventional wisdom has held that the demise of the endometrial cups in normal intraspecies horse pregnancy occurs between
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Figure 172.10 Videoendoscopic views of (a) the detached and bloodless allantochorion in the uterus of a mare undergoing pregnancy failure at 77 days after ovulation; (b,c) the endometrial cups persisting in the previously gravid uterine horn of the same mare at, respectively, 150 and 189 days after ovulation; (d) peripheral serum progestagen (open circles) and equine chorionic gonadotropin (eCG) (closed circles) profiles in the same mare that underwent fetal death around day 77 of gestation. PG, intramuscular injection of 250 µ g cloprostenol to induce luteolysis.
100 and 140 days of gestation.72, 92 This was based on the histological appearance of cups examined in that time period and measurement of eCG levels in the blood of the pregnant mares. Cup morphology showed degeneration and widespread cell death and eCG concentrations fell to undetectable levels. Surprisingly, the results of recent experimental studies and field observations have established that the lifespan of the cups can be extended dramatically beyond 120 days.125, 126 Allen and colleagues followed a mare in which fetal death occurred spontaneously on day 77 of gestation in which the endometrial cup and eCG secretion persisted to beyond the equivalent of day 200 (Figure 172.10).127 Field observations support this experimental finding.126 This phenomenon of persisting endometrial cups had not been de-
scribed previously, probably because of its rarity – perhaps only 1–2% of mares produce endometrial cups that live beyond 140 days of gestation. It is not clear whether the persisting cups live longer than expected because the mare fails to destroy them, or whether the cups themselves fail to trigger a self-destructing mechanism, such as apoptosis. However, the observations from the late John Steiner and his colleagues in Lexington, Kentucky (personal communication) suggest that the endometrial cups can sometimes persist throughout gestation and into the post-parturient period. This is evidence for a condition of prolonged fetal–maternal chimerism that can affect the reproductive status of the mare. Mares with persisting cups fail to cycle normally and therefore remain infertile as long as the cups persist.
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References 1. Cole HH, Hart GH. The potency of blood serum of mares in progressive stages of pregnancy in effecting the sexual maturity of the immature rat. Am J Physiol 1930;83:57–68. 2. Ascheim S, Zondek B. Die Schwangerschaftsdiagnose aus dem Harn durch Nachweis des Hypophysenvordeslapperhormons. Klin Wochenschr 1928;7:1404–11. 3. Saunders FJ, Cole HH. Two gonadotrophic substances in mare serum. Proc Soc Exp Biol Med 1935;32:1476–8. 4. Goss H, Cole HH. Sex hormones in the blood of mares. III. Some chemical properties of the ovary-stimulating principle. Endocrinology 1931;15:214–24. 5. Saunders FJ, Cole HH. Means of augmenting the ovarian response to gonadotrophic substances. Proc. Soc. Exp. Biol. Med. 1936;33:505–8. 6. Cole HH, Erway J. 48-hour test for equine gonadotropin with results expressed in international units. Endocrinology 1941;29:514–19. 7. Cole HH, Guilbert HR, Goss H. Further considerations of the properties of the gonad-stimulating principle of mare serum. Am Physiol 1932;102:227–40. 8. Cole HH. On the biological properties of mare gonadotropic hormone. Am Anat 1936;59:299–332. 9. Goss H, Cole HH. Further studies on the purification of mares gonadotropic hormone. Endocrinology 1940;26: 244–9. 10. Cole HH. The changes occurring in the ovary of the mare during pregnancy. Anat Rec 1931;49:199–209. 11. Cole HH. On the biological properties of highly purified gonadotropin from pregnant mares serum. Endocrinology 1940;27:548–53. 12. Cole HH, Hart GH, Lyons WR, Catchpole HR. The development and hormonal content of fetal horse gonads. Anat Rec 1933;56:275–93. 13. Cole HH, Hart GH. Concerning gonadotropic substances in mare serum. Proc Soc Exp Biol Med 1934;32:370–3. 14. Catchpole HR, Lyons WR. The gonad stimulating hormone of pregnant mares. Am Anat 1934;55:167–227. 15. Cole HH, Goss H. The source of equine gonadotrophin. In: Essays in Honor of Herbert M. Evans. Berkley: University of California Press, 1943; pp. 107–19. 16. Catchpole HR, Cole HH, Pearson PB. Studies on the rate of disappearance and fate of mare gonadotropic hormone following intravenous injection. Am Physiol 1935;112:21–6. 17. Cole HH, Bigelow M, Finkel J, Rupp GR. Biological half-life of endogenous PMS following hysterectomy and studies on losses in urine and milk. Endocrinology 1967;81:27–30. 18. Bielanski W, Ewy Z, Pigoniowa H. Differences in the level of gonadotrophin in the serum of pregnant mares. In: Proceedings of the 3rd International Congress on Animal Reproduction and Artificial Insemination, Cambridge, 1956, pp. 110–11. 19. Clegg MT, Cole HH, Howard CB, Pigon H. The influence of foetal genotype on equine gonadotrophin secretion. J Endocrinol 1962;25:245–8. 20. Rimington C, Rowlands IW. Serum gonadotrophin: preparation of a stable concentrate from pregnant mare’s serum. Biochem J 1941;35:736–48. 21. Rimington C, Rowlands IW. Serum gonadotrophin. 2. Further purification of the active material. Biochem J 1944;38:54–60. 22. Raacke ID, Lostroh AJ, Boda JM, Li CH. Some aspects of the characterization of pregnant mare serum
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
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36.
37.
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39.
40.
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gonadotrophin. Acta Endocrinol (Copenh) 1957;26:377– 87. Bangham DR, Woodward PM. The second international standard for serum gonadotrophin. Bull World Health Org 1966;35:761–73. Gospodarowicz D, Papkoff H. A simple method for the isolation of pregnant mare serum gonadotropin. Endocrinology 1967;80:699–702. Gospodarowicz D. Purification and physico-chemical properties of the pregnant mare serum gonadotrophin (PMSG). Endocrinology 1971;91:101–6. Papkoff H. Chemical and biological properties of the subunits of pregnant mare serum gonadotropin. Biochem Biophys Res Commun 1974;58:397–404. Papkoff H, Bewley TA, Ramachandran J. Physicochemical and biological characterizations of pregnant mare serum gonadotropin and its subunits. Biochim Biophys Acta 1978;532:185–94. Christakos S, Bahl OP. Pregnant mare serum gonadotropin. Purification and physicochemical, biological, and immunological characterization. J Biol Chem 1979;254:4253–61. Farmer SW, Papkoff H. Immunochemical studies with pregnant mare serum gonadotropin. Biol Reprod 1979;21:425–31. Moore WT, Burleigh BD, Ward DN. Chorionic gonadotrophins: comparative studies and comments on relationships to other glycoprotein hormones. In: Segal S (ed.) Chorionic Gonadotrophins. New York: Plenum Press, 1980; pp. 89–126. Moore Jr WT, Ward DN. Pregnant mare serum gonadotropin. Rapid chromatographic procedures for the purification of intact hormone and isolation of subunits. J Biol Chem 1980;255:6923–9. Aggarwal BB, Papkoff H. Relationship of sialic acid residues to in vitro biological and immunological activities of equine gonadotropins. Biol Reprod 1981;24:1082–7. Moudgal NR, Papkoff H. Equine luteinizing hormone possesses follicle-stimulating hormone activity in hypophysectomized female rats. Biol Reprod 1982;26:935–42. Combarnous Y, Guillou F, Martinat N. Comparison of in vitro follicle-stimulating hormone (FSH) activity of equine gonadotropins (luteinizing hormone, FSH, and chorionic gonadotropin) in male and female rats. Endocrinology 1984;115:1821–7. Talmadge K, Vamvakopoulos NC, Fiddes JC. Evolution of the genes for the beta subunits of human chorionic gonadotropin and luteinizing hormone. Nature 1984;307:37–40. Bousfield GR, Sugino H, Ward DN. Demonstration of a COOH-terminal extension on equine lutropin by means of a common acid-labile bond in equine lutropin and equine chorionic gonadotropin. J Biol Chem 1985;260:9531–3. Crawford RJ, Tregear GW, Niall HD. The nucleotide sequences of baboon chorionic gonadotropin beta-subunit genes have diverged from the human. Gene 1986;46:161–9. Policastro PF, Daniels-McQueen S, Carle G, Boime I. A map of the hCG beta-LH beta gene cluster. J Biol Chem 1986;261:5907–16. Bousfield GR, Liu WK, Sugino H, Ward DN. Structural studies on equine glycoprotein hormones. Amino acid sequence of equine lutropin beta-subunit. J Biol Chem 1987;262:8610–20. Stewart F, Thomson JA, Leigh SE, Warwick JM. Nucleotide (cDNA) sequence encoding the horse
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gonadotrophin alpha-subunit. J Endocrinol 1987;115: 341–6. Sugino H, Bousfield GR, Moore Jr WT, Ward DN. Structural studies on equine glycoprotein hormones. Amino acid sequence of equine chorionic gonadotropin betasubunit. J Biol Chem 1987;262:8603–9. Gonzalez G, Castro B, Massaldi H. Extraction of equine chorionic gonadotrophin from pregnant mare plasma by direct adsorption on chromatographic media. Biotechnol Bioeng 1997;57:22–5. Legardinier S, Duonor-Cerutti M, Devauchelle G, Combarnous Y, Cahoreau C. Biological activities of recombinant equine luteinizing hormone/chorionic gonadotropin (eLH/CG) expressed in Sf9 and Mimic insect cell lines. J Mol Endocrinol 2005;34:47–60. Legardinier S, Klett D, Poirier JC, Combarnous Y, Cahoreau C. Mammalian-like nonsialyl complex-type Nglycosylation of equine gonadotropins in Mimic insect cells. Glycobiology 2005;15:776–90. Chopineau M, Martinat N, Troispoux C, Marichatou H, Combarnous Y, Stewart F, Guillou F. Expression of horse and donkey LH in COS-7 cells: evidence for low FSH activity in donkey LH compared with horse LH. J Endocrinol 1997;152:371–7. Lecompte F, Roy F, Combarnous Y. International collaborative study of a preparation of equine chorionic gonadotrophin (eCG NZY-01) proposed as a new standard. J Reprod Fertil 1998;113:145–50. Li MD, Ford JJ. A comprehensive evolutionary analysis based on nucleotide and amino acid sequences of the alpha- and beta-subunits of glycoprotein hormone gene family. J Endocrinol 1998;156:529–42. Bo M, Boime I. Identification of the transcriptionally active genes of the chorionic gonadotropin gene cluster in vivo. J Biol Chem 1992;3179–84. Bahl OP, Thotakura NR, Anumula KR. Biological role of carbohydrates in gonadotropins. In: Saxena BB (ed.) Hormone Receptors in Growth and Reproduction. New York: Raven Press, 1984; pp. 165–82. Bahl OP, Waugh PV. Characterization of glycoproteins: carbohydrate structures of glycoprotein hormones. Adv Exp Med Biol 1986;205:1–51. Damm JB, Hard K, Kamerling JP, Van Dedem GW, Vliegenthart JF. Structure determination of the major N- and O-linked carbohydrate chains of the beta subunit from equine chorionic gonadotropin. Eur J Biochem 1990;189:175–83. Catchpole HR. Physiology of the gonadotropic hormones. In: Cole HH (ed.) Gonadotropins: Their Chemical and Biological Properties and Secretory Control. San Francisco: W.H. Freeman, 1963; pp. 40–70. Butt WR, Crooke AC, Cunningham FJ. Studies on human urinary and pituitary gonadotrophins. Biochem J 1961;81:596–605. Stewart F, Allen WR, Moor RM. Pregnant mare serum gonadotrophin; Ratio of follicle-stimulating hormone and luteinising hormone activities measured by radioreceptor assay. J Endocrinol 1976;71:371–82. Stewart F, Allen WR, Moor RM. Influence of foetal genotype on the follicle-stimulating hormone:luteinising hormone ratio of pregnant mare serum gonadotrophin. J Endocrinol 1977;73:419–25. Aggarwal BB, Farmer SW, Papkoff H, Stewart F, Allen WR. Purification and characterization of donkey chorionic gonadotrophin. J Endocrinol 1980;85:449–55.
57. Roser JF, Papkoff H, Murthy HM, Chang YS, Chloupek RC, Potes JA. Chemical, biological and immunological properties of pituitary gonadotropins from the donkey (Equus asinus): comparison with the horse (Equus caballus). Biol Reprod 1984;30:1253–62. 58. Leigh SE, Stewart F. Partial cDNA sequence for the donkey chorionic gonadotrophin-beta subunit suggests evolution from an ancestral LH-beta gene. J Mol Endocrinol 1990;4:143–50. 59. Stewart F, Maher JK. Analysis of horse and donkey gonadotrophin genes using Southern blotting and DNA hybridization techniques. J Reprod Fertil Suppl 1991;44:19–25. 60. Chopineau M, Martinat N, Gibrat JF, Galet C, Lecompte F, Foulon-Gauze F, Pourchet C, Guillou F, Combarnous Y. Identification of amino-acids in the alpha-subunit first and third loops that are crucial for the heterospecific follicle-stimulating hormone activity of equid luteinizing hormone/chorionic-gonadotropin. Eur J Endocrinol 2004;150:877–84. 61. Min KS, Hattori N, Aikawa J, Shiota K, Ogawa T. Sitedirected mutagenesis of recombinant equine chorionic gonadotropin/luteinizing hormone: differential role of oligosaccharides in luteinizing hormone- and folliclestimulating hormone-like activities. Endocr J 1996;43:585– 93. 62. Martinuk SD, Manning AW, Black WD, Murphy BD. Effects of carbohydrates on the pharmacokinetics and biological activity of equine chorionic gonadotropin in vivo. Biol Reprod 1991;45:598–604. 63. Ortega-Camarillo C, Guzman-Grenfell AM, Hicks JJ. Oxidation of gonadotrophin (PMSG) by oxygen free radicals alters its structure and hormonal activity. Mol Reprod Dev 1999;52:264–8. 64. Rowson LEA. The role of research in animal reproduction. Vet Res 1974;95:276–6. 65. Saumande J, Chupin D. Superovulation: a limit to egg transfer in cattle. Theriogenology 1977;7:141–9. 66. Christie WB, Newcomb R, Rowson LE. Ovulation rate and egg recovery in cattle treated repeatedly with pregnant mare serum gonadotrophin and prostaglandin. Vet Rec 1979;104:281–3. 67. Elsden RP, Nelson LD, Seidel GE. Superovulation of cows with follicle stimulating hormone and pregnant mare’s serum gonadotrophin. Theriogenology 1978;9:17–25. 68. Gordon I. Synchronization of estrus and superovulation in cattle. In: Adams CE (ed.) Mammalian Egg Transfer. Boca Raton: CRC Press, 1982; pp. 63–80. 69. Zondek B. Hormonelle Schwangerschaftstreaktion aus dem Harn bei Mensch und Tier. Klin Wochenschr 1930;9:2285–9. 70. Schauder W. Untersuchungen uber die Eihaute und Embryotrophe des Pferdes. Arch Anat Physiol 1912; 259–302. 71. Mossman HW. Comparative morphogenesis of the fetal membranes and accessory uterine structures. Contrib Embryol 1937;26:129–246. 72. Clegg MT, Boda JM, Cole HH. The endometrial cups and allantochorionic pouches in the mare with emphasis on the source of equine gonadotrophin. Endocrinology 1954;54:448–63. 73. Allen WR, Moor RM. The origin of the equine endometrial cups. I. Production of PMSG by fetal trophoblast cells. J Reprod Fertil 1972;29:313–16. 74. Allen WR, Hamilton DW, Moor RM. The origin of equine endometrial cups. II. Invasion of the endometrium by trophoblast. Anat Rec 1973;177:485–501.
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Equine Chorionic Gonadotropin (eCG) 75. Ewart JC. A Critical Period in the Development of the Horse. London: Adam & Charles Black, 1897. 76. Van Niekerk JC. The early diagnosis of pregnancy, the development of the foetal membranes and nidation in the mare. S Afr Vet Med Assoc 1965;36:483–8. 77. Van Niekerk CH, Allen WR. Early embryonic development in the horse. J Reprod Fertil Suppl 1975;23:495–8. 78. Wooding FB, Morgan G, Fowden AL, Allen WR. A structural and immunological study of chorionic gonadotrophin production by equine trophoblast girdle and cup cells. Placenta 2001;22:749–67. 79. de Mestre AM, Miller D, Roberson MS, Liford J, Chizmar LC, McLaughin, KE, Antczak DF. Glial cells missing homologue 1 is induced in differentiating equine chorionic girdle trophoblast cells. Biol Reprod. 2009;80:227–34. 80. Enders AC, Lantz KC, Schlafke S, Liu IKM. New cells and old vessels: the remodeling of the endometrial vasculature during establishment of endometrial cups. Biol Reprod Monogr 1995;1:181–90. 81. Stewart F, Lennard SN, Allen WR. Mechanisms controlling formation of the equine chorionic girdle. Biol Reprod Monogr 1995;1:151–9. 82. Stewart F, Power CA, Lennard SN, Allen WR, Amet L, Edwards RM. Identification of the horse epidermal growth factor (EGF) coding sequence and its use in monitoring EGF gene expression in the endometrium of the pregnant mare. J Mol Endocrinol 1994;12:341–50. 83. Lennard SN, Stewart F, Allen WR. Transforming growth factor beta 1 expression in the endometrium of the mare during placentation. Mol Reprod Dev 1995;42:131–40. 84. Allen WR, Gower S, Wilsher S. Immunohistochemical localization of vascular endothelial growth factor (VEGF) and its two receptors (Flt-I and KDR) in the endometrium and placenta of the mare during the oestrous cycle and pregnancy. Reprod Dom Anim 2007;42:516–26. 85. Vagnoni KE, Ginther OJ, Lunn DP. Metalloproteinase activity has a role in equine chorionic girdle cell invasion. Biol Reprod 1995;53:800–5. 86. De Mestre AM, Bacon SJ, Costa CC, Leadbeater JC, Noronha LE, Stewart F, Antczak DF. Modeling trophoblast differentiation using equine chorionic girdle vesicles. Placenta 2008;29:158–69. 87. Crump A, Donaldson WL, Miller J, Kydd JH, Allen WR, Antczak DF. Expression of major histocompatibility complex (MHC) antigens on horse trophoblast. J Reprod Fertil Suppl 1987;35:379–88. 88. Donaldson WL, Zhang CH, Oriol JG, Antczak DF. Invasive equine trophoblast expresses conventional class I major histocompatibility complex antigens. Development 1990;110:63–71. 89. Kydd JH, Butcher GW, Antczak DF, Allen WR. Expression of major histocompatibility complex (MHC) class 1 molecules on early trophoblast. J Reprod Fertil Suppl 1991;44:463–77. 90. Maher JK, Tresnan DB, Deacon S, Hannah L, Antczak DF. Analysis of MHC class I expression in equine trophoblast cells using in situ hybridization. Placenta 1996;17:351– 9. 91. Antczak DF, Miller JM, Remick LH. Lymphocyte alloantigens of the horse. II. Antibodies to ELA antigens produced during equine pregnancy. J Reprod Immunol 1984;6:283– 97. 92. Allen WR. The influence of fetal genotype upon endometrial cup development and PMSG and progestagen production in equids. J Reprod Fertil Suppl 1975;23:405–13.
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93. Allen WR. Immunological aspects of the endometrial cup reaction and the effect of xenogeneic pregnancy in horses and donkeys. J Reprod Fertil Suppl 1982;31:57–94. 94. Allen WR, Skidmore JA, Stewart F, Antczak DF. Effects of fetal genotype and uterine environment on placental development in equids. J Reprod Fertil 1993;98:55–60. 95. Martinuk SD, Bristol F, Murphy BD. Effects of the dam on equine chorionic gonadotropin concentrations during pregnancy. Dom Anim Endocrinol 1990;7:551–7. 96. Bell RJ, Bristol FM. Equine chorionic gonadotrophin in mares that conceive at foal oestrus. J Reprod Fertil Suppl 1991;44:719–21. 97. Moor RM, Allen WR, Hamilton DW. Origin and histogenesis of equine endometrial cups. J Reprod Fertil Suppl 1975;23:391–5. 98. Grunig G, Triplett L, Canady LK, Allen WR, Antczak DF. The maternal leucocyte response to the endometrial cups in horses is correlated with the developmental stages of the invasive trophoblast cells. Placenta 1995;16:539–59. 99. Donaldson WL, Oriol JG, Plavin A, Antczak DF. Developmental regulation of class I major histocompatibility complex antigen expression by equine trophoblastic cells. Differentiation 1992;52:69–78. 100. De Mestre AM, Noronha L, Wagner B, Antczak DF. Split immunological tolerance to trophoblast. Int J Dev Biol 2010;54:445–55. 101. Flaminio MJ, Antczak DF. Inhibition of lymphocyte proliferation and activation: a mechanism used by equine invasive trophoblast to escape the maternal immune response. Placenta 2005;26:148–59. 102. Adams AP, Antczak DF. Ectopic transplantation of equine invasive trophoblast. Biol Reprod 2001;64:753–63. 103. Antczak DF, Bright SM, Remick LH, Bauman BE. Lymphocyte alloantigens of the horse. I. Serologic and genetic studies. Tissue Antigens 1982;20:172–87. 104. Antczak DF, Allen WR. Invasive trophoblast in the genus Equus. Ann Immunol (Paris) 1984;135D:325–31. 105. Amoroso EC, Hancock JL, Rowlands IW. Ovarian activity in the pregnant mare. Nature 1948;161:353–6. 106. Allen WR. Equine gonadotrophins. PhD thesis, University of Cambridge, 1970. 107. Evans MJ, Irvine CH. Serum concentrations of FSH, LH and progesterone during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1975;23:193–200. 108. Urwin VE, Allen WR. Pituitary and chorionic gonadotrophic control of ovarian function during early pregnancy in equids. J Reprod Fertil Suppl 1982;32:371–81. 109. Sirois J, Kimmich TL, Fortune JE. Developmental changes in steroidogenesis by equine preovulatory follicles: effects of equine LH, FSH, and CG. Endocrinology 1990;127:2423–30. 110. Daels PF, Albrecht BA, Mohammed HO. Equine chorionic gonadotropin regulates luteal steroidogenesis in pregnant mares. Biol Reprod 1998;59:1062–8. 111. Stewart F, Allen WR. The binding of FSH, LH and PMSG to equine gonadal tissues. J Reprod Fertil Suppl 1979;27:431–40. 112. Allen WR. Factors influencing pregnant mare serum gonadotrophin production. Nature 1969;223:64–5. 113. Stewart F, Allen WR. Biological functions and receptor binding activities of equine chorionic gonadotrophins. J Reprod Fertil 1981;62:527–36. 114. Allen WR, Kydd JH, Antczak DF. Xenogeneic donkeyin-horse pregnancy created by embryo transfer:
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immunological aspects of a model of early abortion. In: Chauoat G (ed.) Immunology of the Fetus. Boca Raton: CRC Press, 1990; pp. 267–83. Hay MF, Allen WR. An ultrastructural and histochemical study of the interstitial cells in the gonads of the fetal horse. J Reprod Fertil Suppl 1975;23:557–61. Cox JE. Oestrone and equilin in the plasma of the pregnant mare. J Reprod Fertil Suppl 1975;23:463–8. Raeside JI, Liptrap RM. Patterns of urinary oestrogen excretion in individual pregnant mares. J Reprod Fertil Suppl 1975;23:649–75. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J Reprod Fertil Suppl 1979;27:499–509. Bhavnani BR. Oestrogen biosynthesis in the pregnant mare. J Endocrinol 1981;89 Suppl: 19P–32P. Pashen RL, Sheldrick EL, Allen WR, Flint AP. Dehydroepiandrosterone synthesis by the fetal foal and its importance as an oestrogen precursor. J Reprod Fertil Suppl 1982;32:389–97. Tait AD, Hodge LC, Allen WR. The biosynthesis of 3 betahydroxy-5,7-androstadien-17-one by the horse fetal gonad. FEBS Lett 1985;182:107–10. Cole HH, Saunders FJ. The concentration of gonadstimulating hormone in blood serum of oestrin in the
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urine throughout pregnancy in the mare. Endocrinology 1935;19:199–208. Rowlands IW. Levels of gonadotropins in tissues and fluids with emphasis on domestic animals. In: Cole HH (ed.) Gonadotropins: Their Chemical and Biological Properties and Secretory Control. San Francisco: W.H. Freeman, 1963; pp. 75–107. Saint-Dizier M, Chopineau M, Dupont J, Combarnous Y. Expression of the full-length and alternatively spliced equine luteinizing hormone/chorionic gonadotropin receptor mRNAs in the primary corpus luteum and fetal gonads during pregnancy. Reproduction 2004;128:219– 28. Mathias SS. Maternal immune responses to equine trophoblast. PhD thesis, University of Cambridge, 1997. Steiner J, Antczak DF, Wolfsdorf K, Saville K, Brooks S, Miller D, Barley E, Zent, W. Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. Allen WR, Kolling M, Wilsher S. An interesting case of early pregnancy loss in a mare with persistent endometrial cups. Equine Vet Educ 2007;19:539–44. Steven DH. Development of the foetal membranes. In: Steven DH (ed.) Comparative Placentation Essays in Structure and Function. London: Academic Press, 1975; p. 67.
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Chapter 173
Adrenal Steroids Valerie J. Linse
The adrenal glands are located retroperitoneally, at the cranial pole of each kidney. Each adrenal gland is composed of two parts, the outer cortex and the inner medulla. The cortex synthesizes a variety of steroid hormones collectively known as corticosteroids. The adrenal cortex comprises three histologically distinct regions: the zona glomerulosa, which produces mineralocorticoids (aldosterone); the zona fasciculata, which secretes glucocorticoids; and the zona reticularis, which produces androgens.1 The adrenal steroids are biosynthesized from cholesterol through a number of transformations. The rate-limiting first step in the process is the conversion of cholesterol to pregnenolone, and this conversion is accelerated by the stimulation of the adrenal cortex by adrenocorticotropic hormone (ACTH). After pregnenolone is formed, it can be readily transformed into a variety of hormones, including progesterone, corticosterone, aldosterone, cortisol, androstenedione, testosterone, or estradiol.
Functions of adrenal steroids Aldosterone secretion is increased in response to hyperkalemia and hyponatremia. Aldosterone increases renal excretion of potassium and reabsorption of sodium in the distal convoluted tubules, thus performing the vital function of maintaining fluid and electrolyte balance within the body. Glucocorticoids are produced by the adrenal gland in response to ACTH secreted from the pituitary, and exogenous administration of ACTH will increase glucocorticoid concentration in the blood.2, 3 Cortisol is the primary glucocorticoid in circulation in the horse,4 although other glucocorticoids such as corticosterone, cortisone, and deoxycorticosterone are also produced in the adrenal cortex.5 Cortisol is responsible for a wide array of physiological processes, including gluconeogenesis, inhibition of glucose uptake, suppression of the immune system, and
inhibition of the inflammatory response.6 Under normal physiological conditions, the glucocorticoids exert negative feedback on the production of ACTH, resulting in a circadian rhythm of secretion such that concentrations of glucocorticoids are high during early morning, plateau during the day, decrease during late afternoon and evening, and finally reach a nadir around midnight.7–11 In addition to the diurnal pattern, cortisol also appears to be secreted in a pulsatile pattern in the horse, with mean intervals of 31 to 105 minutes between pulses.10, 12 Caution is warranted in the interpretation of measured cortisol levels. The total cortisol in circulation is composed of a portion (87%)13 that is bound to the protein corticosteroid-binding globulin (CBG), and an unbound, or ‘free’ portion. While a horse’s measured total cortisol may not rise due to stress, a decrease in the binding capacity of CBG may occur, resulting in a decrease in the percentage of bound compared with unbound cortisol.14 This change allows a larger amount of free cortisol to be available for movement out of the capillaries and into biological activity.15, 16 Therefore, in addition to measurement of free cortisol and total cortisol concentrations, accurate assessment of the responsiveness of the adrenal axis requires the evaluation of the binding capacity of CBG.
Reproductive effects of adrenal steroids Pituitary pars intermedia dysfunction (PPID) In the healthy horse, the hypothalamic hormones thyrotropin-releasing hormone (TRH) and corticotropin-releasing hormone (CRH) cause release of the precursor protein pro-opiomelanocortin (POMC) from the pituitary. Dopamine, also secreted from the hypothalamus, inhibits the production
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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of prolactin (PRL) and POMC. The melanotropes that form the intermediate pituitary and the corticotropes comprising the anterior pituitary both secrete POMC. POMC is a precursor protein that must be cleaved into smaller proteins before it becomes metabolically active. POMC produced by the melanotropes in the intermediate pituitary is cleaved into α-melanocyte-stimulating hormone (α-MSH), β-MSH, corticotropin-like intermediate lobe peptide (CLIP), and β endorphin (βEND). In contrast, the POMC produced in the corticotropes in the anterior pituitary is preferentially cleaved into ACTH.17 The ACTH produced by the corticotropes is released into the bloodstream and carried to the adrenal gland, where it acts on the three zones of the adrenal cortex to stimulate the production of mineralocorticoids, glucocorticoids, and androgens. In horses with PPID, the secretion of dopamine from the hypothalamus is decreased and consequently ACTH secretion from the pituitary is increased. The negative feedback mechanism is disrupted, glucocorticoids are secreted at much higher levels, and the diurnal pattern of secretion is lost. The decrease in dopamine inhibition of the pituitary leads to hypertrophy of the melanotropes, increased production of POMC, and consequently a large increases in synthesis of α-MSH and βEND from the melanotropes. Comparatively small increases in ACTH from the corticotropes also occur.11, 18 While this small increase in ACTH synthesis can lead to increased glucocorticoid production from the adrenal cortex, it is not likely sufficient to cause clinical signs. The problem arises because of the large increases in α-MSH and βEND peptides, which potentiate a sixfold increase in steroidogenic actions of ACTH,19, 20 resulting in continuous glucocorticoid production and a loss of the circadian rhythm.18 The persistently high cortisol levels that ensue lead to a variety of clinical signs in the horse, including muscle wasting, hirsutism, and obesity, with bulging supraorbital fat pads, cresty neck, and fat deposits along the dorsum.21 The high cortisol concentrations predispose the horse to complications such as chronic laminitis,22 infertility, and chronic infections, both parasitic and bacterial. The precise mechanism causing infertility has yet to be elucidated, but has been hypothesized to involve compression of the anterior pituitary by the intermediate pituitary with subsequent reduction in LH and FSH secretion. In addition, LH secretion in humans is stimulated by α-MSH and inhibited by endogenous opioids.23 Although equine research into the effects of α-MSH on gonadotropin secretion has not been published, it is logical to surmise that if the same process occurs in the horse, perturbation of the pulsatile secretion of LH would significantly alter the reproductive cycle of the animal.
The adrenal cortex is also capable of producing the sex steroids progesterone, testosterone, and estradiol.24 Administration of ACTH has been shown to increase secretion of progesterone, androstendione, testosterone, and cortisol in intact and ovariectomized mares.25 While the secretion of sex steroids from the adrenal cortex is normally low, if the adrenal cortex is excessively stimulated and produces above-normal amounts, the sex steroids can have deleterious effects on the reproductive system through inhibition of gonadotropin secretion and consequent disruption of gamete development. Mares Stress induced by transportation of mares and the consequent increased cortisol production has not been shown significantly to alter reproductive parameters, including pre-ovulatory LH and estradiol surges, duration of estrus, time of ovulation, or pregnancy rates.26, 27 However, administration of the synthetic glucocorticoid dexamethasone has been shown to inhibit estrous behavior in ovariectomized mare28 and to decrease LH concentrations, pre-ovulatory follicular growth, and ovulation in intact mares.29 At the ovarian level, dexamethasone inhibits LH-induced synthesis of steroidogenic acute regulatory protein (StAR) and progesterone production in cultured rat pre-ovulatory follicles. This effect is blocked with the addition of RU-486, a glucocorticoid receptor antagonist, suggesting that the effects of dexamethasone are mediated by the glucocorticoid receptor on the ovary.30 Stallions In the stallion, administration of ACTH has been shown to increase serum concentrations of cortisol and decrease concentrations of testosterone, but not affect concentrations of LH.3 However, administration of dexamethasone (30 mg/stallion/day for 7 days) lowered testosterone levels and increased concentrations of LH, and tended to increase sperm midpiece defects in two of five stallions.31 Exerciseinduced stress in stallions has been shown to increase serum concentrations of cortisol and testosterone. Sperm motility and viability were decreased during strenuous exercise, and major seminal defects were increased in the 8 weeks following exercise.32 Pregnancy and the fetus Healthy function of the fetal hypothalamic– pituitary–adrenal (HPA) axis is essential for stimulation of parturition and for maturation of organs necessary for postnatal life.33–35 While pregnant cows
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Adrenal Steroids and ewes are exquisitely sensitive to administration of exogenous cortisol, induction of parturition in the mare can require high doses of dexamethasone (100 mg every 24 h, i.m. for four consecutive days33 ) late in gestation.36 Intrafetal injection of ACTH can lead to precocious maturation of the fetus and decreased length of gestation.37 As parturition approaches in the healthy pregnancy, the equine fetal pituitary begins to produce more ACTH, and the adrenal gland also becomes more responsive to ACTH. In addition, the equine fetal adrenal gland doubles in weight during the last 5% of gestation.38 Concentrations of fetal plasma cortisol concentrations have been shown to increase in response to exogenous ACTH by 304 ± 3 days gestation, and by 313 ± 2 days 50–60% increases in cortisol were elicited.39 The enzyme 17α-hydroxylase, which converts progesterone to cortisol, also becomes detectable at higher levels.40 Consequently, fetal cortisol levels begin to rise sharply 2–3 days before parturition.41, 42 Although the fetal adrenal gland is able to produce more cortisol as parturition approaches, it does not assume a diurnal pattern of cortisol secretion until the foal is approximately 2 months of age.43, 44 In cases of placental pathology, increased fetal adrenocortical activity and maternal plasma concentrations of progestagen have been linked to increased production of cortisol by the fetus.45 Plasma concentrations of progestagen normally rise beginning approximately 20 days before parturition, and injection of the fetus with ACTH, CRH, or betamethasone at 250–320 days gestation has been shown to elicit rises in fetal cortisol and increased concentrations of maternal progestagens.46 Caution is warranted in administering high doses of glucocorticoids to pregnant mares in early to mid-gestation, as fetal growth retardation, low birthweight, and insulin resistance have been associated with their use in pregnant sheep and humans.47–49
References 1. Cameron EHD, Grant JK. Biochemistry of histologically defined zones in the adrenal cortex: cortisol synthesis in the horse. J Endocrinol 1967;37:413–20. 2. MacHarg MA, Bottoms GD, Carter GK, Johnson MA. Effects of multiple intramuscular injections and doses of dexamethasone on plasma cortisol concentration and adrenal responses to ACTH in horses. Am J Vet Res 1985;46: 2285–7. 3. Wiest JJ, Thompson DL, McNeill-Wiest DR, Garza F, Mitchell PS. Effect of administration of adrenocorticotropic hormone on plasma concentrations of testosterone, luteinizing hormone, follicle stimulating hormone and cortisol in stallions. Equine Vet Sci 1988;8:168–70. 4. James VHT, Horner MW, Moss MS, Rippon AV. Adrenal cortical function in the horse. J Endocrinol 1970;48:319–35.
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5. Zolovick A, Upson DW, Eleftheriou BE. Diurnal variation in plasma glucocorticosteroid levels in the horse (Equus caballus). J Endocrinol 1966;35:249–53. 6. Martin PA, Crump MH. The adrenal gland. In: Pineda MH, Dooley MP (eds) McDonald’s Veterinary Endocrinology and Reproduction, 5th edn. Ames: Iowa State University Press, 2003; pp. 165–200. 7. Bottoms GD, Roesel OF, Rausch FD, Akins AL. Circadian variation in plasma cortisol and corticosterone in pigs and mares. Am J Vet Res 1972;33:785–90. 8. Kumar MSA, Liao TF, Chen CL. Diurnal variation in serum cortisol in ponies. J Anim Sci 1976;42:1360. 9. Larsson M, Edqvist LE, Ekman L, Persson S. Plasma cortisol in the horse, diurnal rhythm and effects of exogenous ACTH. Acta Vet Scand 1979;20:16–24. 10. Toutain PL, Oukessou M, Autefage A, Alvinerie M. Diurnal and episodic variation of plasma hydrocortisone concentrations in horses. Domest Anim Endocrinol 1988;5:55–9. 11. Dybdal NO, Hargreaves KM, Madigan JE, et al. Diagnostic testing for pituitary pars intermedia dysfunction in horses. J Am Vet Med Assoc 1994;204:627–32. 12. Evans JW, Winget CM, DeRoshia C, Holley DC. Rhythmic cortisol secretion in the equine: Analysis and physiological mechanisms. J Interdiscipl Cycle Res 1977;8:111–21. 13. Gayrard V, Alvinirie M, Toutain PL. Interspecies variations of corticosteroid-binding globulin parameters. Domest Anim Endocrinol 1996;13:35–45. 14. Alexander SL, Irvine CHG. The effect of social stress on adrenal axis activity in horses: the importance of monitoring corticosteroid-binding globulin capacity. J Endocrinol 1998;157:425–32. 15. Rosner W. The functions of corticosteroid-binding globulin and sex hormone binding globulin: recent advances. Endocr Rev 1990;11:80–91. 16. Rosner W. Plasma steroid-binding proteins. Endocrin Metab Clin 1991;20:697–720. 17. Eipper BA, Mains RE. Structure and biosynthesis of proadrenocorticotropin/endorphin and related peptides. Endocr Rev 1980;1:1–28. 18. Wilson MG, Nicholson WE, Holscher MA, et al. Proopiolipomelanocortin peptides in normal pituitary, pituitary tumor, and plasma of normal and Cushing’s horses. Endocrinology 1982;110:941–54. 19. Shanker G, Sharma A. Beta-endorphin stimulates corticosterone synthesis. Biochem Biophys Res Commun 1979;86:1–5. 20. Seger M, Bennett H. Structure and bioactivity of the amino-terminal of proopiomelanocortin. J Steroid Biochem 1986;25:703–10. 21. Van der Kolk JH, Kalsbeek HC, van Garderen E, et al. Equine pituitary neoplasia: a clinical report of 21 cases (1990–1992). Vet Rec 1993;133:594–7. 22. Hillyer MH, Taylor FRG, Mair TS, et al. Diagnosis of hyperadrenocortism in the horse. Equine Vet Educ 1992;4:131–4. 23. Limone P, Calvelli P, Altare F, Ajmone-Catt P, Lima T, Molinatti GM. Evidence for an interaction between alphaMSH and opioids in the regulation of gonadotropin secretion in man. J Endocrinol Invest 1997;20:207–10. 24. Silberzahn P, Rashed F, Zwain I, Leymarie P. Androstenedione and testosterone biosynthesis by the adrenal cortex of the horse. Steroids 1984;43:147–52. 25. Hedberg Y, Dalin AM, Forsberg M, Lundeheim N, Sandh G, Hoffmann B, Ludwig C, Kindahl H. Effect of ACTH (tetracosactide) on steroid hormone levels in the mare. Part B: effect in ovariectomized mares (including estrous behavior). Anim Reprod Sci 2007;100:92–106.
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26. Baucus KL, Squires EL, Ralston SL, McKinnon AO, Nett TM. Effect of transportation on the estrous cycle and concentrations of hormones in mares. J Anim Sci 1990;68:419–26. 27. Berghold P, Mostl E, Aurich C. Effects of reproductive status and management on cortisol secretion and fertility of oestrous horse mares. Anim Reprod Sci 2007;102:276–85. 28. Asa CS, Goldfoot DA, Garcia MC, Ginther OJ. Dexamethasone suppression of sexual behavior in the ovariectomized mare. Horm Behav 1980;14:55–64. 29. Asa CS, Ginther OJ. Glucocorticoid suppression of oestrus, follicles, LH and ovulation in the mare. J Reprod Fert Suppl 1982;32:247–51. 30. Huang T-J, Li PS. Dexamethasone inhibits luteinizing hormone-induced synthesis of steroidogenic acute regulatory protein in cultured rat preovulatory follicles. Biol Reprod 2001;64:163–70. 31. Juhasz J, Vidament M, Nagy P, Kulcsar M, Guillaume D, Magistrini M, Huszenicza Gy. Effect of glucocorticoid treatment on fresh and deep frozen semen and on endocrine parameters in the stallion. Havemeyer Foundation Monograph Series 1999;1:36–7. 32. Janett F, Burkhardt C, Burger D, Imboden I, H¨assig M, Thun R. Influence of repeated treadmill exercise on quality and freezability of stallion semen. Theriogenology 2006;65:1737–49. 33. Alm CC, Sullivan JJ, First NL. Induction of premature parturition by parenteral administration of dexamethasone in the mare. J Am Vet Med Assoc 1974;165:721–2. 34. Webb PD, Leadon DP, Rossdale PD, Jeffcott LB. Studies on equine prematurity 5: Histology of the adrenal cortex of the premature newborn foal. Equine Vet J 1984;16:297–9. 35. Giussani DA, Forhead AJ, Fowden AL. Development of cardiovascular function in the horse fetus. J Physiol 2005;565:1019–30. 36. Ousey JC, Kolling M, Allen WR. The effects of maternal dexamethasone treatment on gestation length and foal maturation in Thoroughbred mares. Anim Reprod Sci 2006;94:436–8. 37. Ousey JC, Rossdale PD, Dudan FE, Fowden AL. The effects of intrafetal ACTH administration on the outcome of pregnancy in the mare. Reprod Fertil Dev 1998;10:359–67. 38. Fowden AL, Silver M. Comparative development of the pituitary-adrenal axis in the fetal foal and lamb. Reprod Domest Anim 1995;30:170–7.
39. Silver M, Fowden AL. Prepartum adrenocortical maturation in the fetal foal: responses to ACTH. J Endocrinol 1994;142:417–25. 40. Mason JI, Hinshelwood MM, Murray BA, et al. Tissue specific expression of steroid 17α-hydroxylase/C-17,20 lyase, 3β-hydroxysteroid dehydrogenase and aromatase in the fetal horse. In: Proceedings of the Society for Gynecological Investigation Meeting, 1993, p. 357. 41. Silver M. Prenatal maturation, the timing of birth and how it may be regulated in domestic animals. Exp Physiol 1990;75:285–307. 42. Cudd TA, LeBlanc M, Silver M, Norman W, Madison J, Keller-Wood M, Wood CE. Ontogeny and ultradian rhythms of adrenocorticotropin and cortisol in the late-gestation fetal horse. J Endocrinol 1995;144:271– 83. 43. Komosa M, Flisinska-Bojanowska A, Gill J. Development of a diurnal rhythm in some metabolic parameters in foals. Comp Biochem Physiol 1990;95:549–52. 44. Hart KA, Ferguson DC, Heusner GL, Barton MH. Evaluation of low-and high-dose synthetic adrenocorticotropic hormone stimulation tests in healthy neonatal foals. In: P Annu Conv Am Assoc Equine 2007;53:360–7. 45. Rossdale PD, Ousey JC, Cottrill CM, Chavatte P, Allen WR, McGladdery AJ. Effects of placental pathology on maternal plasma progestagen and mammary secretion calcium concentrations and on neonatal adrenocortical function in the horse. J Reprod Fertil Suppl 1991;44:579–90. 46. Rossdale PD, McGladdery AJ, Ousey JC, Holdstock N, Grainger L, Houghton E. Increase in plasma progestagen concentrations in the mare after foetal injection with CRH, ACTH or betamethasone in late gestation. Equine Vet J 1992;24:347–50. 47. Jobe AH, Wada N, Berry LM, Ikegami M, Ervin MG. Single and repetitive maternal glucocorticoid exposures reduce fetal growth in sheep. Am J Obstet Gynecol 1998;178:880– 5. 48. Moss TJ, Sloboda DM, Gurrin LC, Harding R, Challis JRG, Newnham JP. Programming effects in sheep of prenatal growth restriction and glucocorticoid exposure. Am J Physiol 2001;281:960–70. 49. Sloboda DM, Moss TJ, Gurrin L-C, Newnham J, Challis JRG. The effect of prenatal betamethasone administration on postnatal ovine hypothalamic-pituitary-adrenal function. J Endocrinol 2002;172:71–81.
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Chapter 174
Melatonin Daniel C. Sharp
Introduction The early Greek anatomists were struck by the fact that the unpaired pineal is situated close to the fourth ventricle of the brain which channels cerebrospinal fluid (CSF) out of the brain and down the spinal cord. These Greek philosophers thought the pineal acted as a valve to control the flow of ‘thought’ (CSF). Rene Descartes expanded on this idea by proposing that the pineal translated messages entering the brain into musculoskeletal actions. That perfectly fits the definition of a neuroendocrine transducer, a description that was entirely prescient and agrees well with our current understanding of how the pineal functions. Centuries passed with little further elaboration on Descartes’ quaint concept of pineal function until 1917, when McCord and Allen observed that injection of extracts from bovine pineals caused blanching of the skin of tadpoles.1 In 1958, a dermatologist and biochemist interested in pigmentation isolated the factor that caused skin blanching in frog skin.2 The compound was 5-methoxytryptamine, or melatonin. The name is unfortunate because current understanding of the role of melatonin has little to do with skin blanching. Furthermore, the name is unfortunate because of its similarity to melanocytestimulating hormone (MSH), which is an entirely different hormone with entirely different functions. It has been suggested that melatonin may have been originally part of the retinal shedding regulatory mechanism. Photoreceptor cells of the retina reside within an outer segment connected to the inner segment of the retina by a thin stalk. To maintain the integrity of these cells, there is a continuous shedding and replacement process in which the older cells become phagocytized and newer material synthesized and transported to the outer segment.3 This process of retinal disc morphogenesis has been shown to be temporally associated with melatonin secretion, suggesting a causal relationship. Rufiange
et al.4 examined electroretinograms (ERGs) and salivary melatonin concentrations in humans, and reported that ERG amplitude was lowest at times when melatonin concentrations were highest. These authors stated that melatonin, from the retina itself and/or from the pineal gland, is necessary for the normal processing of visual information. Thus, melatonin’s involvement in the visual system likely made it a good candidate for evolutionary reassignment to photic measurement capabilities. Although the action of melatonin on skin pigmentation changes in frogs was of scientific interest, research in the involvement of the pineal gland in reproductive function grew out of clinical cases of precocious puberty that were associated with pineal tumors in children.5, 6 Tumors in the pineal area have also been associated with delayed puberty.7 This apparent dichotomy has been explained by the hypothesis that some tumors secrete excess melatonin and hence suppress normal reproductive development, whereas other tumors in the pineal area actually result in the destruction of the normally inhibitory effects of the pineal, thus enabling premature gonadal development. Our current understanding of the role of melatonin is twofold. Melatonin affects the timing of circadian rhythms through a direct action at the biological clock, adjusting phase of the rhythm. More importantly for horse breeders, melatonin regulates the timing of the seasonal reproductive rhythm by informing the central nervous system of the changes in daylength. This information is processed in both the hypothalamus and in the pars tuberalis of the pituitary.8 For many years, studies of melatonin focused on the rhythm of melatonin in the peripheral circulation,9 but there was little understanding of where melatonin exerted its actions. Discovery and description of two high-affinity melatonin receptors in the late 1980s opened new areas of investigation and led
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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to the current appreciation that melatonin receptors are located throughout the hypothalamus, suprachiasmatic nucleus (SCN), and the pars tuberalis.10 The melatonin receptors, named MT1 and MT2, are both members of the seven transmembrane, G-proteincoupled cell surface receptor family. The two receptors appear to exhibit some redundancy, as targeted deletion of the MT1 receptor may be partially compensated by the presence of the MT2 receptor, at least in rodents.8 As might be anticipated, expression of the MT1 receptor and iodo-melatonin binding exhibit daily variations in rodents, with elevated expression and binding during the photophase.8 Furthermore, light exposure during the scotophase increases these end points. This suggests that ligand availablility is at least partially responsible for regulation of the receptors. Little is known of the melatonin receptors in equids, with the exception of the report of Nonno et al.11 in which iodo-melatonin binding was observed only in the pars tuberalis of the horse pituitary. No evidence of binding in the hypothalamus was cited. Smith et al.12 stated the possibility that melatonin signaling alters KiSS-I gene expression because melatonin receptors are known to exist in the same regions as the KiSS-1 gene (caudal arcuate nucleus). Because the pineal is thought to be a long-term timer, it seems reasonable to speculate that it could play a role in such long-term events such as initiation of puberty and onset of reproductive function following seasonal anestrus. Whether melatonin plays a role in regulating these events through an interaction with the KiSS-1 gene in horses is yet to be determined, but is an exciting possibility.
Circadian timekeeping system In order to understand the role of the pineal gland and melatonin as a timekeeping system, it is worth discussing the circadian timekeeping system in general. The word circadian is Latin and means ‘about a day’. Therefore, the term circadian describes events that take place with a periodicity of about 24 hours. To be truly circadian, a rhythm must adhere to two principles: (i) the rhythm must be genetically inherent and expressed under constant conditions with a period or tau that is close to, but not necessarily exactly, 24 hours; and (ii) the rhythm must be shown to be entrainable to exactly 24 hours in a normal photoperiod. Care should be taken not to refer to a biological rhythm as being circadian unless it is known that the rhythm adheres to these two principles. Even events that take place regularly, such as every day (quotidian) may not be derived from the circadian timekeeping system, even though they appear to occur regularly. The circadian timekeeping system is
an important synchronizer of a great variety of physiological events in the interest of maintaining homeostasis. For instance, in rodents the time of ovulation is precisely regulated to occur at night. Because these animals are nocturnally active, ovulation occurring during the day would not likely result in breeding at the appropriate time. Likewise, the growing field of chronopharmacology indicates the importance of the circadian system; the efficacy of many drugs has been shown to vary with time-of-day of administration. In many cases, the range of efficacy is from a better response to the drug, to complete absence of any effect. The existence of a biological timekeeping system is the real-life version of the quackery-based biorhythm charts you can purchase for a small price. While such nostrums are of little value, our current understanding of the biological clock has improved tremendously and is making inroads into medical and veterinary practice. It is satisfying to note that in the original version of this chapter, the authors evoked the suprachiasmatic nuclei (SCN) as ‘the potential source of the clock’, and it can now be stated accurately that the SCN is the master clock in virtually all mammals in which studies have been performed.13 As we live on a planet that rotates, all living things are exposed to daily changes in illumination that provide both visual information and a temporal signal. The temporal signal, alternating dark and light, became a useful tool in coordinating activities of living things. For instance, the dark phase (scotophase) of the daily photoperiod is advantageous for prey animals to move about, as predation is more difficult then. It is thought that the role of melatonin in retinal function became adapted to the pineal gland as a reliable indicator of time. In mammals, information about environmental lighting conditions is perceived by the retina of the eyes and is converted into the nocturnal elevation of the hormone melatonin. In mammals, in which the pineal is not directly photoreceptive, this information depends on the master circadian pacemaker located in the hypothalamus, the SCN. The science of circadian rhythms has grown exponentially in the last 10 years, and our understanding of the timekeeping function of the pineal has changed. The ability to sustain a rhythm of ‘about a day’ yet be influenced (entrained) by external environmental lighting reflects a transcription/translation-based autoregulatory feedback system. This consists of a number of ‘clock genes’ interacting to inhibit or stimulate other clock genes such that the outcome of their interaction creates a cycle of about a day. Furthermore, this set of clock genes can be influenced or entrained by photoperiod (see Chapter 183).
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Melatonin In order to understand the complex system evolved in mammals, it is necessary to first discuss the function of lower vertebrate animals. Vertebrate fishes, lacertilian lizards, and some species of birds have direct photoreceptive cells in the pineal gland cells (pinealocytes) and adhere to the principles of a circadian clock system. That is, they have: (i) a self-sustained oscillator or rhythm circuit, (ii) a photic input pathway that can ‘tune’ the oscillator, and (iii) an output pathway or signal (melatonin). In these species, removal of the pineal from the animal and establishing the pinealocytes in an in vitro culture system in a photoperiodic environment results in robust maintenance of the circadian clock system.14 Such cells in culture not only continue to produce melatonin in a rhythmic fashion representing the environmental photoperiod, but also demonstrate the ability to be phase-shifted by exposure to light at the beginning of subjective night. This description of non-mammalian pinealocytes is similar to the existence of a transcription/translationbased autoregulatory feedback system found in the suprachiasmatic nucleus of mammals, and now known to be the site of the biological clock. The course of evolution in mammals has apparently utilized a similar circadian time-keeping system, but has eliminated, for whatever reason, the direct photoreception. Instead, the mammalian melatonin-producing system relies on retinal photoreception and transmission of photic information to the pineal gland via the SCN. The SCN serves to modify or interpret the photic information relative to its own clock gene timing. In contrast, melatonin may also provide feedback to the SCN to inform or modify the SCN clock gene timing as well.15 In a marvelous revelation, it was recently discovered that pinealocytes derived from day-old rats and cultured in vitro exhibited direct photoreceptive properties and resembled pinealocytes of sauropsids (crocodiles, turtles, snakes, and birds)and anamniotes (frogs and lizards).16 This development toward the more primitive neuronal photoreceptor lineage is thought to reflect the fact that day-old rat pups had not yet been exposed to extrinsic factors such as norepinephrine (NE) as rats are altricial and the sympathetic innervation of the pineal gland matures postnatally. HIOMT (see below) and melatonin formation do not occur until postnatal days 1 and 5, respectively in rats. This agrees with the work of Kilmer et al.,17 who reported that pony foals did not exhibit robust melatonin rhythms until 7–11 weeks of age. Mammalian pineal cells have been shown to contain all major clock genes of the endogenously oscillating clockwork of the SCN, but it is unclear whether these clock genes participate in daily waves of gene expression, or whether they are atavistic genes that
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never underwent activation. This is indeed a fascinating aspect of pineal function and one that deserves to be followed closely as it will no doubt provide further clues as to pineal regulation, and, thus, hints as to how we might control pineal function exogenously.
Biosynthetic pathway Elucidation of the chemical structure of melatonin as an indole amine (5-methoxy-N-acetyltryptamine) attracted the interest of many researchers, including Julius Axelrod, winner of the 1970 Nobel prize for physiology. Axelrod’s interest in methylation reactions led him to isolate and purify an enzyme from bovine pineal glands that O-methylated several indoles. These researchers found that of all the hydroxyindoles that served as substrate for the O-methylating enzymes, N-acetylserotonin was by far the best substrate, and the enzyme was named hydroxyindole-O-methyltransferase (HIOMT). This enzyme was later found to be highly localized in the pineal gland, leading to the speculation that melatonin synthesis was essentially unique to the pineal gland. In the course of their studies, another enzyme which converted serotonin to N-acetylserotonin, the substrate for HIOMT, was isolated from pineal glands. Axelrod and his colleagues proposed a biosynthetic pathway for melatonin originating with the amino acid tryptophan18 (see Figure 174.1).
Anatomic pathway An association between pineal function and environmental lighting had long been suspected, based on diverse observations. Some species of birds also
TRYPTOPHAN
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Figure 174.1 Melatonin biosynthetic pathway. The enzyme arylalkylamine N-acetyltransferase (AA-NAT) is thought to be the rate-limiting step in the synthesis. Importantly, AA-NAT is expressed in the pineal gland in response to norepinephrine release from postganglionic fibers of the sympathetic chain only during darkness.
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Figure 174.2 Coronal section of horse brain showing presence of color reaction in left, but not right, SCN. III Vent, third ventricle; SCN, suprachiasmatic nucleus; OC, optic chiasm. Arrow indicates the stained SCN.
have direct pineal photoreception.19 In fact the parietal eye, or so-called third eye as the pineal was sometimes named, has recently been shown to have many functions in common with the retina. As the mammalian pineal timekeeping system is not directly photoreceptive the retina of the eye is the primary photoreceptor and an elaborate mechanism of photoadaptation has evolved which depends on ocular photoreception. Photoadaptation consists of three dominant mechanisms: (i) reduction of retinal illumination by pupillary light reflex; (ii) direct modulation of neuroendocrine function; and (iii) entrainment of the circadian timekeeping system.20 Complete loss of rod and cone function in humans and in animal models does not eliminate photoresponsiveness, whereas experimental bilateral enucleation abolishes all ocular light responses. This suggests that a novel set of ocular photoreceptors, essentially independent of the rods and cones, exists. Evidence for such intrinsically photosensitive photoreceptors has only recently been discovered. Hattar et al.21 reported the existence of an opsin-like protein called melanopsin located in retinal ganglion cells which project monosynaptically to the suprachiasmatic nuclei (SCN). This direct connection to the SCN is called the retinohypothalamic tract (RHT) and has been shown to be present in all mammals studied, including horses22 (see Figure 174.2). Presence of the RHT was demonstrated by administration of horseradish peroxidase into the aqueous humor of horses and then tracing its anterograde flow to brain structures using the tetramethylbenzidene procedure to generate a color reaction. Figure 174.2 shows a coronal section of horse brain at the level of the optic chiasm. The horseradish peroxidase color reaction can be seen adjacent to the third ventricle, ventrally
(arrow). Of interest, the horseradish peroxidase was administered into the right eye, and the color reaction was observed entirely in the left SCN, although the crossover (decussation) of optical fibers at the chiasm was reported to be 80.8%.23 This observation suggests that the photoregulatory system consists of non-image-forming fibers which project from the retina to the SCN. Thus, the melanopsin-containing retinal ganglion fibers are capable of conveying photic information directly to the SCN where adaptation to changes in ambient light is regulated by the SCN. This photoregulatory pathway permits the apparently paradoxical situation in which an animal can be completely visually impaired, i.e. rods and cones absent, yet be perfectly capable of adapting to photoperiodic changes. Thus visually blind horses may not be incapable of adjusting to season changes in daylength, depending on the specific cause of the blindness. From the SCN, fibers, presumably containing adjusted information from the SCN, pass through the paraventricular nuclei and midbrain bundle to the anteriomedial-lateral cell column of the spinal cord and exit the brain in the sympathetic trunk where preganglionic fibers enter the superior cervical ganglia adjacent to the common carotid artery near the caudal border of the mandibles (Viborg’s triangle). From the superior cervical ganglia, postganglionic fibers re-enter the brain as the nervi conarii and terminate as terminal axons closely associated with pinealocytes.24, 25 Interruption of this pathway at any point from retina to pineal can lead to disruption of circadian timekeeping. From the standpoint of equine reproduction, that can lead to disruption of the secretion of melatonin, and ultimately to the disruption of seasonal breeding rhythms. Spinal cord injury at the level of C1–6 disrupts the nerve pathway, leading to nyctohemeral rhythms of melatonin secretion. In humans, this condition has been shown to lead to deleterious interruption of the sleep-wake rhythm,26 and nocturnal supplementation with melatonin has been suggested as important therapy for such patients. As indicated in the above discussion, melatonin is secreted in a circadian rhythm with peak secretion during the scotophase or dark phase of the photoperiod (see Figure 174.3). This reflects the activity of the rate-limiting enzyme arylalkylamine Nacetyltransferase (AA-NAT) which is driven by the endogenous master oscillator located in the SCN. The SCN are themselves entrained to ambient photoperiod through the action of melanopsin and the retinal ganglion cells which constitute the retinohypothalamic tract. Expression of the gene encoding AA-NAT is stimulated by the neurotransmitter norepinephrine released by postganglionic terminals impinging directly on the pinealocytes. Of interest and potentially
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Melatonin
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Figure 174.3 Melatonin concentration (pg/mL) in peripheral plasma throughout a 24-hour period in mares. Note the rise in melatonin at the beginning of the scotophase.
practical value, it has been suggested that AA-NAT and other pineal genes which are induced by cyclic adenosine monophosphate (cAMP) may become imprinted by the photoperiod of prior entrained cycles.27 This means that storage of photoperiodic information which is manifested as the daily profile of melatonin secretion represents the photoperiodic memory of the master clock, the SCN. If that is the case, then peripheral secretion of melatonin should continue to display some form of rhythm under constant conditions, i.e., continuous darkness. When sheep were exposed to constant light, the melatonin rhythm was completely abolished, but when they were exposed to constant darkness, the melatonin rhythms persisted, albeit reduced in magnitude.28 In an experiment designed to test the possible endogenous nature of melatonin secretion in mares, melatonin was measured at 2-hour intervals for 72 hours after a 3-week exposure to a 14:10 LD photoperiod in four mares. In an attempt to monitor other well-known circadian rhythms, water intake was measured by means of metered automatic watering devices, and locomotor activity was monitored via halter-mounted hiking pedometers. The mares were then exposed to constant darkness (L:D 0:24) for an additional 3 weeks, and melatonin, locomotor activity, and water intake were again monitored at 2-hour intervals for 72 hours. All three measurements displayed regularly recurring periodicity, with melatonin concentrations rising immediately with onset of darkness, and locomotor activity and water intake rhythms exhibiting peaks early in the photophase (subjective day).29 Importantly, all mares were well synchronized in the timing of peak events for all three rhythms. However, after exposure to constant darkness for 3 weeks, all three rhythms were disrupted, both within mares, and among mares. That is, the rhythms either did not exhibit peak times (locomotor activity and water intake) or exhibited marked times of peak and nadir activity, but no longer associated
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with subjective night (melatonin). Furthermore, the melatonin rhythms were no longer temporally coordinated among mares, with times of peak secretion varying from mare to mare (see Figure 174.4a,b). The practical significance of this observation is possibly twofold: (i) changes in the melatonin secretory rhythms in response to changes in photic exposure may require some time before entrainment to the new photoperiod becomes ‘permanent’; (ii) the natural occurrence of renewed reproductive function of mares in the late winter and spring may reflect such photoperiodic imprinting from the master clock, as opposed to an immediate, direct response to increasing daylength. Thus, it is interesting to contemplate the possibility that naturally occurring sexual recrudescence in mares reflects the slow, spontaneous response to the photic history of the master oscillator (SCN), whereas stimulation of early sexual recrudescence by exposure to artificial lighting may reflect active re-entrainment (likely phase advance) of the master clock. In further studies of melatonin secretion Sharp and Grubaugh30 demonstrated that the rhythm of melatonin in the peripheral circulation was entrainable to different photoperiods. Because of high variability experienced in sampling mares at 1-hour intervals, jugular vein samples were collected at 10-minute intervals for 30 hours from pony mares under L:D 16:8 or L:D 8:16. The results of this experiment indicated that the secretion of melatonin was highly episodic. The episodes of elevated melatonin in the peripheral circulation were often short-lived, sometimes confined to one or two samples, and were represented in equal numbers during the photophase and the scotophase. High pressure liquid chromatography was used to isolate fractions in several of the episodic samples which co-migrated with authentic melatonin. The fractions were subsequently re-assayed using radioimmunoassay with the same results, so it is clear that the episodes of elevated melatonin were genuine. The term episodic is defined as ‘a noteworthy happening, for which the time can usually be fixed, occurring in the course of ordinary events’. In contrast, the term pulsatile is defined as being ‘characterized by a regularly recurring pulsation, as in the heart beat’. The difference between the two terms is not trivial, as the underlying mechanisms regulating secretory patterns may be different. A key feature of a pulsatile rhythm is that it is regularly recurring. That means that, having observed three or four events, the timing of the next subsequent event can easily and accurately be predicted. Certainly, the regularly recurring component of a pulsatile rhythm is important in that it makes analysis of the rhythmic pattern simpler. The irregularly occurring
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Figure 174.4 (a) Melatonin, water intake, and locomotor activity in an individual mare exposed for 3 weeks to L:D 14:10. The data represent the last 3 days of the 3-week period. Note the regular occurrence of water intake and locomotor activity early in the subjective day and the nocturnal rise in melatonin at first darkness. (b) Melatonin, water intake, and locomotor activity in an individual mare exposed for 3 weeks to L:D 0:24. The data represent the last 3 days of the 3-week period. Note the divergence in rhythms from mares in L:D 14:10, as well each rhythm from the others. Also note the higher amplitude of melatonin concentrations. (From Berglund et al. 1981.)29
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Melatonin secretion of melatonin, observed when frequent samples were collected, seems a poor biological signal, as predictability of subsequent events was not possible. However, it is thought that the duration of melatonin secretion is what conveys the major photic information. The more important observation of this experiment was that, despite the presence of episodic bursts of melatonin secretion, the mean concentrations of melatonin faithfully reflected the length of the scotophase. Of further interest, the magnitude of melatonin secretion (mean concentration during scotophase), was considerably higher in mares subjected to L:D 8:16 compared with mares subjected to L:D 16:8 (185.7 ± 4.1 vs 47.4 ± 7.3 pg/mL, respectively) (see also Berglund et al.).29 Measurement of peripheral melatonin concentrations throughout a 24-hour period once monthly for an entire year also revealed similar observations. That is, the nocturnal rise of melatonin faithfully reflected the time of sunset and the duration of the scotophase. However, the mean nocturnal melatonin concentrations were greatly elevated during the scotophase of the winter months compared with the mean melatonin concentrations during scotophase of the summer months. In support of these observations, Wesson et al.31 obtained equine pineal glands from a nearby slaughter plant at monthly intervals throughout the year and analyzed the HIOMT activity in them. These authors reported significant seasonal changes in pineal HIOMT activity with peak activity during the months of November, December, and January. Although these data do not necessarily indicate a need for higher melatonin concentrations during times of prolonged scotophase, they do support the notion that prolonged darkness enhances HIOMT abundance, and, thus, makes possible greater melatonin production during the scotophase. The significance of this latter observation is not clear. Little information is available on the potential physiological effects of higher melatonin concentrations because most research attention has been drawn to the clear role of melatonin as a chronobiotic. However, the observation of greatly elevated melatonin concentrations during times of either constant darkness or prolonged scotophase in equids is novel, and is certainly worth follow-on studies.
Effects of melatonin administration If melatonin plays such an important role in reproduction, one might ask why a practical way of manipulating it for breeding purposes has not yet been developed. The most obvious answer is that there is already a practical way to manipulate melatonin secretion; by manipulating the photoperiod. The time of greatest melatonin secretion is the time of reproductive quiescence leading to the natural
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conclusion that manipulation of melatonin secretion to enhance breeding potential means to reduce the amount and/or timing of melatonin secretion. Secretion of melatonin is coupled to darkness, and the first rise in melatonin during the scotophase begins almost immediately with darkness. Therefore, by exposing mares to prolonged daylight (or by mimicking daylight with artificial light), one effectively postpones the nocturnal secretion of melatonin. Therefore, the simplest form of melatonin secretion is to extend day by a few hours, thus postponing the nocturnal secretion of melatonin (see Chapter 183). As previously stated, the current dogma is that the duration of melatonin serves as the major photic signal. However, the alternative hypothesis, that the first few hours of melatonin secretion in the scotophase may be in coincidence with a photosensitive phase of the biological clock, has not been well tested. As this is discussed at length in Chapter 183, the following is presented as evidence for the negative aspects of melatonin on reproduction in horses. It is important to consider that there are species differences in the use of melatonin as a time-keeping signal, and data from other species should not be extrapolated for use in horses without extensive research. For instance, ewes are signaled to begin their breeding season in autumn, when night-time melatonin secretion begins to increase, and studies have shown that subcutaneous implants of melatonin can in fact stimulate an early onset of the breeding season in sheep.32 In early studies, melatonin was administered in horses by subcutaneous implants of beeswax containing melatonin, during the summer.33 After 4 weeks, the hypothalami were harvested and analyzed for gonadotropin releasing hormone (GnRH) by radioimmunoassay. These authors reported that the GnRH content of the hypothalamus was significantly reduced, and exhibited GnRH content similar to mares during the winter anestrus. Cleaver et al.34 reported that exposure of mares to constant light significantly reduced the concentration of circulating melatonin and increased hypothalamic GnRH content. Thus the inverse relationship between melatonin and the hypothalamic-pituitary axis in horses began to emerge. To study the effect of melatonin removal on luteinizing hormone (LH) and GnRH secretion, Cleaver et al.35 housed two groups of ovariectomized mares in photoperiod rooms for 31 days beginning in October (autumn). One group (control) was maintained in a photoperiod similar to the ambient photoperiod (L:D 12:12) and the other group (constant light) was exposed to L:D 24:0). Melatonin secretion in the control mares exhibited the normally expected nyctohemeral rhythm, with daytime
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concentrations extremely low and night-time concentrations of 100–150 pg/mL. Mares in the constant light group did not exhibit nyctohemeral melatonin rhythms with melatonin concentrations throughout the entire 24-hour period equivalent to daytime in the control group. Thus, constant light exposure was an effective means of removing the effects of melatonin. After 2 weeks of constant light exposure, mean LH concentrations as well as mean follicle-stimulating hormone (FSH) rose significantly and remained elevated until 4 weeks of exposure to constant light when both declined somewhat. Hypothalamic GnRH content was elevated at 4 weeks following exposure to constant light compared with control mares, but the difference only approached significance at P = 0.07. As GnRH was a one-time measurement, and as it coincided with the small decline in both LH and FSH, it is possible that GnRH was also significantly elevated, by 2 and 3 weeks of constant light exposure, but the experimental design did not permit assessment of GnRH throughout the experiment. The slight reduction in LH and FSH following their initial increase is unexplained currently, but not unexpected. Sharp et al.36 reported significantly elevated GnRH secretion using push-pull perfusion techniques following exposure to long day. These researchers observed a highly significant increase in GnRH after 2 weeks of increased photoperiod, but not 1 week (Figure 174.5; data not shown for 1 week). Interestingly, this initial increase was somewhat reduced at 3 weeks, but was still highly significantly increased over that of mares exposed to short day. Furthermore, FSH was also increased by 3 weeks of exposure to long day (see Figure 174.6), presumably in response to the increased gonadotropin releasing
hormone. However, LH did not increase in the relatively short duration of the experiment. This was also not unexpected as Sherman et al.37 had shown that the genes encoding LH subunit synthesis were undetectable throughout the winter anestrus. Further evidence of the inhibitory effects of melatonin on gonadotropin secretion was presented by Cleaver et al.35 These researchers utilized a model in which melatonin was administered to mares exposed to constant light for 3 weeks and blood samples collected at 30-minute intervals for 12 hours at 0, 7, and 14 days of exposure. Melatonin was administered orally in sugar cubes (20 mg/cube) at 2-hour intervals. Three groups of mares were utilized; group 1 received melatonin so as to elevate circulating melatonin for 12 hours (6 cubes), group 2 received melatonin so as to elevate circulating melatonin concentrations for 8 hours (4 cubes), and a third group also received melatonin so as to elevate circulating melatonin concentrations for 8 hours (4 cubes) but this group was previously pinealectomized and had no endogenous melatonin. FSH was elevated in the peripheral circulation by 14 days of treatment in pinealectomized mares receiving 8 hours of melatonin compared with mares in constant light receiving melatonin for 12 hours. Episodes of gonadotropin secretion were evaluated via frequent sampling regimes. The number of LH episodes/12-h period was relatively low (1.8 ± 1.1) and was not different among groups or sampling times. The number of FSH episodes/12-h period was somewhat higher (4.6 ± 1.9) and was not different between pineal intact and pinealectomized mares receiving 8 hours of melatonin treatment. At the first sampling period, however, the number of FSH
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episodes was significantly lower in pineal-intact mares receiving 12 hours of melatonin treatment. Pinealectomized mares exhibited overall higher LH and FSH concentrations, suggesting absence of any inhibitory effects of endogenous melatonin. Pinealectomy was performed approximately 1 month prior to the experiment, and it is possible that the elevated gonadotropins in pinealectomized mares reflected the longer time without melatonin inhibition. Evidence for the direct role of melatonin in conveying photic information important in regulating timing of the annual reproductive rhythm was provided by an experiment in which melatonin and photoperiod information were in conflict. Cheves et al.38 used anestrous mares to test the hypothesis that exogenous melatonin administration would block the stimulatory effects of long-day photoperiod on date of the first ovulation of the year. Four experimental groups were established (n = 5 mares per group): group 1, short day, control (ambient light, about L:D 9:15); group 2, short day, melatonin administration; group 3, long-day, melatonin administration, and group 4, long day, control. Long day photoperiod consisted of ambient light exposure, with additional 2.5 hours of artificial light (approximately 100 lux) beginning at sunset. The rationale behind this experiment was that melatonin secretion has been shown to be closely coupled to first darkness.30 Therefore, we reasoned that the 2.5 hours of additional photoperiod would delay the secretion of melatonin by approximately 2.5 hours. Melatonin administration was timed to provide elevated circulating melatonin concentrations during the 2.5 hours that the mares in group 3 were exposed
to artificial light. Melatonin (25 mg/animal) was administered orally in a small bolus of grain fed approximately 4 hours before elevated melatonin concentrations were desired. Mares were monitored daily for follicular development and ovulation, and the major end point of the experiment was the date of the first ovulation of the year. The dates of first ovulation of the year were similar for mares in groups 1, 2, and 3: 93.0, 101.3, and 126.0 (Julian dates), respectively. The date of the first ovulation of the year for mares in group 4 was significantly earlier (Julian day 85.0) as would be expected from exposure to the stimulatory photoperiod provided by adding 2.5 hours additional lighting artificially at the time of sunset. These data indicate that the stimulatory effects of long day were blocked by the presence of melatonin at the comparable time of day in group 3.
Practical considerations The desire to regulate melatonin secretion for the purpose of accelerating onset of the breeding season is the most common purpose for horse breeders. However, unlike in sheep, the operative need for accelerating onset of the breeding season in mares is to alter the timing of melatonin secretion. While there are exciting new advances taking place currently using melatonin receptor antagonists, the time-tested practice of delaying melatonin secretion at night with artificial light is likely the most effective for the expense. As the reader will appreciate, removing melatonin completely is not desirable. Rather, the daily rhythm of melatonin secretion must be adjusted, just as occurs naturally with changing photoperiod.
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References 1. McCord CP, Allen FP. Evidences associating pineal gland function with alterations in pigmentation. J Exp Zool 1917;23:207–24. 2. Lerner AB. Isolation of melatonin, the pineal factor that lightens melanocytes. J Am Chem Soc 1958;80:2587. 3. Boesze-Battaglia K, Goldberg AF. Photoreceptor renewal: a role for peripherin/rds. Int Rev Cytol 2002;217; 183–225. 4. Rufiange M, Dumont M, Lachapelle P. Correlating retinal function with melatonin secretion in subjects with an early or late circadian phase. Invest Ophthalmol Vis Sci 2002;43:2491–9. 5. Borit A, Schmidek HH. Pineal tumors and their treatment. In: Reiter RJ (ed.) The Pineal Gland. New York: Raven Press, 1984; pp. 323–44. 6. Heubner O. Tumor der glandula pinealis. Dtsch Med Wochenstr 1898;24:214. 7. Kitay J. Pineal lesions and precocious puberty: A review. J Clin Endocrinol Metab 1954;14:622–5. 8. Von Gall C, Stehl JH, Weaver DR. Mammalian melatonin receptors: molecular biology and signal transduction. Cell Tissue Res 2002;309:151–62. 9. Arendt J. Melatonin assays in body fluids. J Neural Transm Suppl 1987;13:265–78. 10. Weaver DR, Rivkees SA, Reppert SM. Localization and characterization of melatonin receptors in rodent brain by in vitro autoradiography. J Neurosci 1989;9:2581–90. 11. Nonno R, Capsoni S, Lucini V, Møller M, Fraschini F, Stankov B. Distribution and characterization of the melatonin receptors in the hypothalamus and pituitary gland of three domestic ungulates. J Pineal Res 1995;18(4):207–16. 12. Smith JT, Clay CM, Caraty A, Clark IJ. KiSS-1 messenger ribonucleic acid expression in the hypothalamus of the ewe is regulated by sex steroids and season. Endocrinology 2007;148:1150–7. 13. Brown TM, Piggins HD. Electrophysiology of the suprachiasmatic circadian clock. Progr Neurobiol 2007;82:229–55. 14. Okano T, Fukada Y. Photoreception and circadian clock system of the chicken pineal gland. Microsc Res Tech 2001;53:72–80. 15. Challet E. Minireview: entrainment of the suprachiasmatic clockwork in diurnal and nocturnal mammals. Endocrinology 2007;148:5648–55. 16. Maronde E, Stehl JH. The mammalian pineal gland: known facts, unknown facets. Trends Endocrinol Metab 2007;18:142–50. 17. Kilmer DM, Sharp DC, Berglund LA, Grubaugh W, McDowell KJ, Peck LS. Melatonin rhythms in pony mares and foals. J Reprod Fertil Suppl 1982;32:303–7. 18. Axelrod J, Weissbach H. Enzymatic O-methylation of N-acetylserotonin to melatonin. Science 1961;131:1312. 19. Ekstrom P, Meissl H. Evolution of photosensory pineal organs in new light: the fate of neuroendocrine photoreceptors. Phil Trans R Soc Lond 2002;358:1679–700. 20. Nayak SK, Jegla T, Panda S. Role of a novel photopigment, melanopsin, in behavioral adaptation to light. Cell Mol Life Sci 2007;64:144–54. 21. Hattar S, Liao HW, Takao M, Berson DM, Yau KW. Melanopsin-containing retinal ganglion cells: architecture, projections, and intrinsic photosensitivity. Science 2002;295(5557):955–7.
22. Sharp DC, Grubaugh WR, Gum GG, Wirsig CR. Demonstration of a direct retino-hypothalamic projection in the mare. Biol Reprod 1984;(Suppl 1)30:156. 23. Herron MA, Martin JE, Joyce JR. Quantitative study of the decussating optic axons in the pony, cow, sheep, and pig. Am J Vet Res 1978;39(7):1137–9. 24. Inouye ST, Turek FW. Horizontal knife cuts either ventral or dorsal to the hypothalamic paraventricular nucleus block testicular regression in golden hamsters maintained in short days. Brain Res 1986;370(1):102–7. 25. Anderson E. The anatomy of bovine and ovine pineals. light and electron microscopic studies. J Ultrastruct Res 1965;Suppl 8:1–80. 26. Scheer FAJL, Zeitzer JM, Ayas NT, Brown R, Czeisler CA, Shea SA. Reduced sleep efficiency in cervical spinal cord injury; association with abolished night time melatonin secretion. Spinal Cord 2006;44:78–81. 27. Spessert R, Gupta BBP, Rohleder N, Gerhold S, Engel L. Cyclic AMP-inducible genes respond uniformly to seasonal lighting conditions in the rat pineal gland. Neuroscience 2006;143:607–13. 28. Rollag MD, Niswender GD. Radioimmunoassay of serum concentrations of melatonin in sheep exposed to different lighting regimens. Endocrinology 1976;98(2):482–9. 29. Berglund LA, Sharp DC, Grubaugh W. Effects of constant darkness on melatonin rhythms in pony mares. Biol Reprod 1981;(Suppl. 1)24:71. 30. Sharp DC, Grubaugh WR. Pulsatile secretion of melatonin during the scotophase in mares. Biol Reprod 1983;(Suppl. 1)28:100. 31. Wesson JA, Orr EL, Quay WB, Ginther OJ. Seasonal relationship between pineal hydroxyindole-Omethyltransferase (HIOMT) activity and reproductive status in the pony. Gen Comp Endocrinol 1979;38:46–52. 32. Eldon J. Effect of exogenous melatonin and exposure to a ram on the time of onset and duration of the breeding season in Icelandic sheep. J Reprod Fertil 1993;99(1):1–6. 33. Strauss SS, Chen CL, Kalra SP, Sharp DC. Depletion of hypothalamic gonadotropin releasing hormone (GnRH) in ovariectomized mares following melatonin implants. Annual Meeting of the Federation of American Societies for Experimental Biology, Atlantic City, NJ, 1978. 34. Cleaver BD, Davis SD, Grubaugh WR, Sheerin PC, Sharp DC. The role of constant light on circulating gonadotropin levels and on hypothalamic gonadotropin-releasing hormone (GnRH) in ovariectomized (OVX) mares. Biol Reprod 1990;(Suppl. 1)42:150. 35. Cleaver BD, Davis SD, Sharp DC. Effect of pinealectomy (PNX) and melatonin replacement on seasonal regulation of gonadotropin secretion in the pony mare. Biol Reprod 1992;(Suppl. 1)46:60. 36. Sharp DC, Grubaugh WR, Weithenauer J, Sheerin P. Exposure to long photoperiod results in increased GnRH secretion in anestrous pony mares. Biol Reprod 1988;Suppl. 1:39. 37. Sherman GB, Wolfe MW, Farmerie TA, Clay CM, Threadgill DS, Sharp DC, Nilson JH. A single gene encodes the β subunits of equine luteinizing hormone and chorionic gonadotropin. Mol Endocrinol 1992;6:951–9. 38. Cheves LL, Sharp DC. The effects of oral melatonin on the onset of the breeding season of pony mares under artificially lengthened photoperiod. 5th Florida Neuroendocrine Symposium, Tallahassee, FL, 22 February 1985.
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Chapter 175
Oxytocin, Inhibin, Activin, Relaxin and Prolactin Peter R. Morresey
Knowledge of the peptide hormones involved in equine reproduction is comparatively sparse. Application of knowledge and principles found in other species is necessary to understand the many functions peptide hormones are responsible for within the horse.
Oxytocin Oxytocin is a neuropeptide with a wide range of central and peripheral effects, being a neurohormone, neurotransmitter, and neuromodulator. Typical actions of oxytocin include stimulation of uterine smooth muscle contraction during labor and milk ejection during lactation. Oxytocin is considered in the mare, as in the cow, to have a role in maternal recognition of pregnancy, being involved in the luteolytic release of prostaglandins should signaling from the conceptus fail.1 It also is ascribed a wide range of behavioral effects, including maternal, in a number of species.2 Structure Oxytocin is a nine amino acid peptide elucidated in 1953 that is structurally similar to the closely related peptide, arginine vasopressin (AVP). Oxytocin differs from the antidiuretic hormone AVP in two of the nine amino acid residues. These two neurohypophysial hormones display a high degree of evolutionary stability in vertebrate species.3 Source Classically oxytocin is recognized as being produced in the hypothalamus and posterior pituitary; however, it has also been suggested to be produced from
the gonads, placenta, and in some non-reproductive tissues.4 Although oxytocin is present in small quantities in extracts of corpus luteum and follicles during the estrous cycle, these are not considered a source of circulating oxytocin in mares, as cyclic variation in the release of oxytocin is not seen.5 In the mare, oxytocin-mRNA has been identified in the endometrium, with oxytocin and associated proteins detected in the uterus. Endometrial biopsy specimens collected during estrus showed oxytocin by immunostaining in the secretory cells of the endometrium. Vesicles containing immunoreactive oxytocin were present on the luminal surface, suggesting that oxytocin is secreted into the uterine lumen by apical exocytosis. Oxytocin was not present in ciliated epithelial cells of the uterine lumen and endometrial glands, stromal cells, or basal endometrial glands.4 Physiology Oxytocin acts via a G protein-coupled receptor. Receptors have been identified in a wide range of tissues including kidney, heart, thymus, pancreas, and adipocytes.6 Oxytocin likely acts in a paracrine manner, with a high number of oxytocin receptors within the uteroplacental tissues being functionally more important than elevated circulating oxytocin. It is likely that oxytocin receptors are present in high concentrations in utero-placental tissue of the mare due to the marked sensitivity of the uterus to low doses of oxytocin at the time of parturition.7, 8 Baseline oxytocin levels fluctuate, suggesting a possible circadian rhythm. Oxytocin is secreted in a pulsatile manner and may be involved in estrous
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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cycle regulation by promoting luteolysis via uterinederived PGF2α .9, 10 Endometrial oxytocin mRNA presence changes with the day of the estrous cycle or pregnancy, with levels at estrus being higher than at any other day.11 Oxytocin mRNA levels have been shown to be negatively correlated with serum progesterone and positively correlated with serum estradiol in non-bred mares.11 A priming effect of estrogen on endometrial-derived oxytocin has been shown.12 Stimulation of the genital tract and uterine distension during breeding activity have been shown to increase circulating oxytocin concentrations.13 Oxytocin may be involved in the shortening of the luteal phase in response to cervical manipulation.14 It is suspected that locally synthesized uterine oxytocin affects uterine contractility, in addition to its role in luteolysis in the mare.4 The concentration of endometrial oxytocin receptors determines uterine secretion of prostaglandin (PG) F2α in cyclic mares. A role in pregnancy recognition is suggested as endometrial oxytocin receptor concentrations are reduced in early pregnancy by the secretory products of the conceptus.15 Oxytocin concentrations are persistently low throughout gestation, but increase markedly during parturition. Levels remain low until rupture of the allantochorion, peaking during fetal expulsion.16
ulating contractions of the reproductive tract, aiding sperm release. Oxytocin is synthesized within the testis, epididymis, and prostate. Receptors throughout the reproductive tract suggest a paracrine role in stimulating contractility of the seminiferous tubules, epididymis, and prostate gland. Oxytocin has also been shown to modulate androgen levels in these tissues via stimulation of 5α-reductase.17, 18 Both directly and via interaction with androgen metabolism, oxytocin affects prostate growth. Of clinical relevance, oxytocin has been found useful in the clearance of ampullary blockages with sperm that may render a stallion azoospermic.19, 20 Recent studies also demonstrate the ability of repeated oxytocin injection during the early to middiestrus period to block luteolysis and extend the lifespan of the corpus luteum.21 This is proposed as a practical means of long-term estrus suppression in mares.
Inhibin Inhibin was initially identified as a glycoprotein hormone involved in the regulation of folliclestimulating hormone (FSH) release from the pituitary gland.22 In the early 1930s, testicular extracts and follicular fluid had been found to be associated with lower circulating levels of FSH. The existence of inhibin was only proven in the mid-1980s.23
Function Oxytocin has been long considered a hormone that predominantly functions in females. It appears to be involved in sexual and maternal behaviors. Parturition is dependent upon endogenous release of oxytocin in response to cervical and vaginal dilation (Fergusson reflex). Therapeutically administered oxytocin is useful to enhance uterine clearance and facilitate controlled induction of parturition. Various treatment regimens are used. In one parturition study, injection of a daily low dose of oxytocin to those mares diagnosed ready to foal by mammary secretion calcium content offered an advantage over conventional induction protocols, with delivery only occurring in mares carrying a mature fetus in which foaling was imminent, the majority of which foaled within 120 minutes.7 Milk letdown is facilitated by contraction of the myoepithelial cells lining the alveoli and ducts of the mammary gland. A neuroendocrine reflex involving stimulation by suckling causes production and release of oxytocin. This also occurs in response to visual and olfactory cues. Oxytocin has been assigned both endocrine and paracrine roles in male reproduction. At ejaculation, oxytocin is released from the neurohypophysis, stim-
Structure Inhibins belong to the transforming growth factorβ (TGF-β) superfamily. The α- and β-subunits of inhibin are both synthesized as large precursor proteins joined by one or more disulfide bonds.24 Different combinations of the subunits yield molecules with differing activities. Two inhibins are recognized being produced by the combination of the α and either a βA- or a βB-subunit (forming inhibin A and inhibin B, respectively).23 Also, differential post-translational processing of the α precursor gives rise to different size variants of inhibin A and B.25
Source Inhibins are gonadal glycoproteins produced by the granulosa and Sertoli cells under the influence of FSH. Inhibin α subunits have been demonstrated in granulosa cells of small and large follicles and in the theca interna cells of large follicles, with inhibin βA and βB subunits localized to the granulosa and theca interna cells of large follicles.26 Levels of inhibins in maternal circulation were very low compared to fetal levels, therefore fetoplacental inhibins are likely
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Oxytocin, Inhibin, Activin, Relaxin and Prolactin produced in the fetal gonads.27 Inhibin subunit proteins can be found in interstitial cells of the equine fetal ovary.26 The equine fetal testes also secrete large amounts of inhibins including dimeric inhibin A, possibly inhibin B, and activins into the fetal circulation. This suggests that the inhibins play an important role in the development of fetal testes during gestation.28 In the adult male, inhibin is sourced from the Sertoli and Leydig cells.29, 30 Physiology Concentrations of inhibins are low when minimal follicular activity is present in the ovaries, during deep winter anestrus. Concentrations of inhibins are higher during growth of anovulatory follicles in spring transition than during winter anestrus. Plasma concentrations of inhibin A are significantly higher during the growth of pre-ovulatory follicles than during the growth of transitional anovulatory follicles. However, concentrations of inhibin pro-αC isoforms, precursor molecules for the α subunit, do not differ between the two types of follicle. Plasma inhibin proαC isoforms but not inhibin A peak on the day of ovulation. Inhibins likely contribute to negative feedback on the release of FSH from the pituitary gland, during both the transitional period and the breeding season.31 There is an inverse relationship between concentrations of FSH and the inhibins throughout the equine estrous cycle.26 An ovulatory surge in the level of inhibin in the mare occurs. This is the result of the release of follicular fluid into the abdominal cavity when the follicle ruptures. These findings also suggest that circulating inhibin pro-αC may be useful for determining the time of ovulation in the mare, as the time of increase corresponded to ovulation detection by ultrasonography.32 Concentrations of inhibin and related compounds are elevated in fetal compared to maternal circulation between days 100 and 250 of gestation, with fetal ovaries also containing large amounts of inhibin proαC and inhibin A. It is thought inhibin has local regulatory input into the developing gonad.33 These inhibin forms are undetectable in maternal ovaries and the placenta.26
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mRNA has been detected in the placenta, pituitary, adrenal, bone marrow, kidney, spinal cord, and brain. The presence of inhibin RNA in the brain and spinal cord suggests neuroregulatory functions in the central and peripheral nervous systems.34 Inhibin A, but not the pro- and α-C isoforms, is significantly higher in pre-ovulatory follicles during both spring transition and regular cyclicity than in dominant anovulatory transitional follicles. High follicular levels of inhibin A are associated with continued follicle growth and ovulation.35 In the stallion, both FSH and inhibin exhibit similar seasonal changes; peripheral FSH and inhibin increase in the spring and decrease in the fall.29, 36 Inhibin feeds back to the pituitary in the stallion to inhibit release of FSH, and likely acts in conjunction with estrogen and testosterone. Inhibin therefore controls androgen secretion and germ cell proliferation.37 Discussion on pharmacological manipulation of the mare’s reproductive cycle associated with inhibin are to be found in Chapter 195.
Activin Activins are the dimer of β (A or B) inhibin subunits.23 These dimers act as multifunctional growth factors, with roles in feedback within the reproductive system and regulation of both reproductive and non-reproductive organs.38 Structure Dimers of the β-subunits of inhibin are called activin A or activin B (homodimers) and activin AB (heterodimer).22 They are joined together by disulfide bonds.39 Source Cumulus granulosa cells have been found to express high levels of inhibin α, βA, and βB subunits, with oocytes expressing activin receptors.25 Expression of inhibin α-subunit was not observed in the placenta, with inhibin/activin βA-subunit and its mRNA confined to maternal endometrial glands. Activin A in the fetal placental tissue diffuses across from the endometrial glands. The βB-subunit is not present in either endometrial glands or microcotyledons. The large amounts of inhibin subunits in fetal circulation are secreted from fetal gonads.27
Function Inhibin acts to decrease the synthesis and release of FSH, regulating the growth of spring anovulatory and pre-ovulatory follicles. Inhibin subunits have been demonstrated in both gonadal and extragonadal tissues. In addition to the ovary and testis, inhibin
Physiology Activins are locally produced and perform local regulatory roles in a wide range of tissues, including the pituitary, gonads, and placenta.40 Activins of ovarian origin are unlikely to exert endocrine effects on the
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pituitary, as all activin present in the peripheral circulation is tightly bound to and neutralized by follistatin, an activin-binding protein.41 In both maternal endometrial and fetal placental tissues, activin A levels have been demonstrated to decrease as pregnancy progresses.27 Function Activin essentially has the opposite biological effects to inhibin.42 Experimentally, activin A induces an increase in FSH secretion from the pituitary, an increase in ovarian and uterine weight, and increased ovarian secretion of inhibin and estradiol.43 Activin induces granulosa cell proliferation; increases FSH receptor expression, granulosa cell steroidogenesis, basal and gonadotropin-stimulated aromatase activity and estradiol production; activin suppresses thecal androgen production, and delays the onset of luteinization and follicular atresia.25 In several species, cumulus-derived activins affect nuclear and cytoplasmic maturation of oocytes. Although activin has been shown to positively modulate the release of FSH at the level of the pituitary in many species, its action in the horse is largely unknown;30 however, placental function and development are considered to be at least partly regulated by locally produced activin in a paracrine or autocrine fashion.27 A role in placental attachment during early equine pregnancy is suggested by the relatively high concentrations of activin A in the placental tissues at days 100–140, followed by a subsequent decrease.27 Activin has not been identified in the testicle of the stallion. In other species, activin has a paracrine role in partially inhibiting steroidogenesis in Leydig cells.44 Activin stimulates follistatin and inhibin in an autocrine fashion from Sertoli cells.45 Experimentally, activin has age-dependent effects on Sertoli cell division and affects the development of male germ cells.46
Relaxin Relaxin was discovered in 1926, due to the ability of serum from pregnant guinea pigs and rabbits to relax the pubic symphysis of virgin guinea pigs when injected immediately following estrus.47 Structure Relaxin has shown to be identical to insulin with respect to its disulfide bond distribution, suggesting that relaxin and insulin were derived from a common ancestral gene.48 The structure of relaxin is highly variable between species, this being reflected
in its functional diversity and variable potency with a standard assay. However, in spite of an average 50% sequence difference between relaxins from various species, a considerable degree of biological crossreactivity is present.49 Relaxin consists of two polypeptide chains with one intrachain and two interchain disulfide links, identical to those present in insulin.48 In addition to the cystines, five sequence identities occur between these two hormones. The B-chain contains two CysGly residues which are essential structural elements enabling a β- turn, allowing the B-chain to fold back onto the molecular surface. Source The source of relaxin varies among species. Relaxin is produced in the corpora lutea (rats, mice, pigs), the placenta (rabbits, hamsters), and in the uterus (guinea pigs).47 In the equine species, relaxin is produced by the trophoblastic cells of the placenta.50, 51 Using in situ hybridization employing a cRNA probe, tissues of the placenta–endometrium interface recovered between 33 and 153 days of gestation, from mares carrying intraspecific horse, interspecific mule and extraspecific donkey conceptuses, were investigated.52 Relaxin mRNA and relaxin were both present in the single-cell non-invasive trophoblast layer of the allantochorion between 45 and 153 days of gestation in all three types of equine pregnancy examined. Relaxin mRNA and relaxin were both absent from invasive chorionic girdle and differentiated trophoblast cells of the endometrial cups throughout their period of development and regression.52 Physiology Relaxin is not detected in the first part of gestation and appears around day 80. A progressive rise to peak levels around 200 days of gestation is followed by a decline over the next 60 days.53 Maternal concentrations of relaxin rise again in late gestation and increase further during parturition.54 Postpartum relaxin concentrations decrease gradually following placenta expulsion, but remain elevated if placental retention occurs. Substantial between-breed differences in maternal plasma relaxin levels have been demonstrated; however, this seems to be unrelated to placental size and may indicate differences in physiology between horse breeds.55 Function Relaxin has been credited with remodeling of the pelvic girdle, softening of the cervix, and vaginal
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Oxytocin, Inhibin, Activin, Relaxin and Prolactin changes allowing passage of the fetus. However all these functions are not present in all species. Relaxin has been shown to contribute to myometrial relaxation, although relaxin concentrations increase substantially during labor. A period of myometrial quiescence is observed 2 to 4 hours before delivery; this is potentially overcome by high concentrations of PGF2α that occur during second-stage labor.56 Additionally, relaxin may reflect placental function with evidence that maternal plasma relaxin concentrations decline before abortion.55, 57 Decreasing levels are present with poor microcotyledon development; relaxin may, therefore, be useful as an indication of placental viability.50, 57 A recent study shows a positive relationship between low circulating relaxin and adverse pregnancy outcome in mares with compromised placental function.58 Although of value in identifying mares with high-risk pregnancies associated with placental dysfunction, relaxin was not a good indicator of response to therapeutic intervention in this study. Relaxin has also been shown to be present in mare ovaries at different stages of the estrous cycle. In follicular fluid, relaxin has been demonstrated to increase with increasing follicular size. Relaxin has also been found in granulosa and theca cells of preovulatory follicles, although this is dependent on the test used. Messenger RNA for relaxin has been shown in all stages of corpora lutea development, but not in corpora hemorrhagica or corpora albicans. Mature corpora lutea have been found to contain mRNA and relaxin in the large luteal cells.59 It is thought that relaxin participates in remodeling of the reproductive organs by regulating proteolytic enzyme activity. By this action, relaxin is involved in ovarian follicle development and ovulation. As equine follicles are found in the ovarian cortex at the center of the ovary, it is suggested that relaxin may contribute to the considerable expansion of the follicle and migration to the ovulation fossa by modulating the degradation of extracellular matrix in ovarian stromal tissue.59, 60 Relaxin in the male has been detected in testicular and accessory sex gland tissue of a number of species.47 The function of relaxin in the male is not yet proven; however, a lack affects development of the male reproductive tract and a role in testicular descent has been surmised due to experimental changes in gubernacular growth in deprived laboratory species.47
Prolactin The history of the isolation of prolactin has been reviewed.61 In the 1920s, minimal knowledge existed about the anterior pituitary. Extracts of the ante-
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rior pituitary were shown to contain substances that had dramatic effects on a wide range of organ systems, including the reproductive organs. Milk secretion by the mammary gland could be stimulated. Using the pigeon crop-sac assay, Riddle62 in 1933 was able to report the isolation and assay of prolactin using fractions of anterior pituitary extracts. This work established that there existed a milk-secreting substance distinct from growth-promoting and gonadstimulating properties of anterior pituitary extracts. Radioimmunoassay was developed in the 1970s, specific for prolactin, that enabled elucidation of the patterns of prolactin secretion in blood. Structure Prolactin is synthesized as a single polypeptide including a signal peptide, which is cleaved to leave three intrachain disulfide bridges. Source Examination of the pituitary has revealed two cell populations containing only prolactin (type I and II lactotropes), one cell population containing only growth hormone (somatotropes), and one further group of cells containing both growth hormone and prolactin in the same secretory granules (mammosomatotropes).63 Physiology Of the anterior pituitary hormones, prolactin is unique, being under predominantly dopaminergic inhibitory control by the hypothalamus. Prolactin secretion is relatively constant, with only minor surges above a high baseline.64 Plasma prolactin concentrations in most species are positively correlated with daylength, with concentrations higher during summer than during winter.65 Prolactin release is elevated during the breeding season of the horse.66 Exercise and growth hormone levels have both been shown to affect prolactin levels, and resting levels are higher in intact male and female horses.64 Thyroidreleasing hormone has also been shown to stimulate prolactin release, independent of the season.67 In the mare, photoperiod plays an important role in controlling circulating levels of prolactin. Other seasonal factors, such as ambient temperature, are involved in modulating the seasonal rhythm of serum prolactin release.68 In the stallion, opioidergic regulation and dopaminergic inhibition of prolactin secretion undergoes seasonal variations.69 Secretion of prolactin has been shown to increase as the stallion increases in age.66
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Function Prolactin has been shown to have an effect on many body tissues in many species. These vary from involvement in fluid homeostasis, participation in metamorphosis, and support of corpus luteum function. In the mare, effects on mare mammary development and milk secretion have been shown, in agreement with other species.70 A rise in prolactin accompanies the rise in luteinizing hormone preceding the onset of the ovulatory season, suggesting a role for prolactin in the termination of the spring transition period.71 Increased secretion of prolactin, whether endogenous or stimulated by dopamine antagonists, has been associated with spring transitional follicle growth.72 Dopamine receptors are also present throughout ovarian tissue, suggesting a direct effect of dopamine on ovaries is possible in addition to the well-known endocrine pathway involving the hypothalamic–pituitary–gonadal axis.73 During the autumn transition, a failure of PGF2α release occurs with greater frequency than at other times during the reproductive season, this failure being associated with prolongation of function of the corpus luteum. Prolactin receptors have been demonstrated to occur in the corpus luteum of the mare.74 The luteolytic process, whether endogenous or induced, is associated with prolactin secretion; however, prolactin secretion does not appear to be affected in estrous cycles where prolongation of corpus luteum function occurs.75 In the male of a number of species, prolactin has stimulatory effects on androgen-sensitive tissues. Together with LH and growth hormone, it controls synthesis of LH receptors in the testes, activates androgen synthesis, and affects spermatogenesis.76, 77 Specific information in the stallion is lacking.
19.
References
20.
1. Hansel W, Dowd JP. Hammond memorial lecture. New concepts of the control of corpus luteum function. J Reprod Fertil 1986;78(2):755–68. 2. Neumann ID. Brain oxytocin: a key regulator of emotional and social behaviours in both females and males. J Neuroendocrinol 2008;20(6):858–65. 3. Murphy D, Si-Hoe SL, Brenner S, Venkatesh B. Something fishy in the rat brain: molecular genetics of the hypothalamo-neurohypophysial system. Bioessays 1998;20(9):741–9. 4. Bae SE, Watson ED. A light microscopic and ultrastructural study on the presence and location of oxytocin in the equine endometrium. Theriogenology 2003;60(5):909–21. 5. Stevenson KR, Parkinson TJ, Wathes DC. Measurement of oxytocin concentrations in plasma and ovarian ex-
6. 7.
8. 9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
21.
22.
23.
24.
tracts during the oestrous cycle of mares. J Reprod Fertil 1991;93(2):437–41. Kiss A, Mikkelsen JD. Oxytocin – anatomy and functional assignments: a minireview. Endocr Regul 2005;39(3):97–105. Camillo F, Marmorini P, Romagnoli S, Cela M, Duchamp G, Palmer E. Clinical studies on daily low dose oxytocin in mares at term. Equine Vet J 2000;32(4):307–10. Pashen RL. Low doses of oxytocin can induce foaling at term. Equine Vet J 1980;12(2):85–7. Stout TA, Lamming GE, Allen WR. Oxytocin administration prolongs luteal function in cyclic mares. J Reprod Fertil 1999;116(2):315–20. Tetzke TA, Ismail S, Mikuckis G, Evans JW. Patterns of oxytocin secretion during the oestrous cycle of the mare. J Reprod Fertil Suppl 1987;35:245–52. Behrendt-Adam CY, Adams MH, Simpson KS, McDowell KJ. Oxytocin-neurophysin I mRNA abundance in equine uterine endometrium. Domest Anim Endocrinol 1999;16(3):183–92. Behrendt-Adam CY, Adams MH, Simpson KS, McDowell KJ. Effects of steroids on endometrial oxytocin mRNA production. J Reprod Fertil Suppl 2000;56:297–304. Nikolakopoulos E, Kindahl H, Gilbert CL, Goode J, Watson ED. Release of oxytocin and prostaglandin f(2alpha) around teasing, natural service and associated events in the mare. Anim Reprod Sci 2000;63(1–2):89–99. Handler J, Konigshofer M, Kindahl H, Schams D, Aurich C. Secretion patterns of oxytocin and PGF2alpha-metabolite in response to cervical dilatation in cyclic mares. Theriogenology 2003;59(5–6):1381–91. Starbuck GR, Stout TA, Lamming GE, Allen WR, Flint AP. Endometrial oxytocin receptor and uterine prostaglandin secretion in mares during the oestrous cycle and early pregnancy. J Reprod Fertil 1998;113(2):173–9. Vivrette SL, Kindahl H, Munro CJ, Roser JF, Stabenfeldt GH. Oxytocin release and its relationship to dihydro-15keto PGF2alpha and arginine vasopressin release during parturition and to suckling in postpartum mares. J Reprod Fertil 2000;119(2):347–57. Ivell R, Balvers M, Rust W, Bathgate R, Einspanier A. Oxytocin and male reproductive function. Adv Exp Med Biol 1997;424:253–64. Thackare H, Nicholson HD, Whittington K. Oxytocin: its role in male reproduction and new potential therapeutic uses. Hum Reprod Update 2006;12(4):437–48. Varner DD, Blanchard TL, Brinsko SP, Love CC, Taylor TS, Johnson L. Techniques for evaluating selected reproductive disorders of stallions. Anim Reprod Sci 2000;(60–61):493–509. Estrada A, Samper JC, Lillich JD, Rathi RR, Brault LS, Albrecht BB, Imel MM, Senne EM. Azoospermia associated with bilateral segmental aplasia of the ductus deferens in a stallion. J Am Vet Med Assoc 2003;222(12):1740–2, 1707. Vanderwall DK, Rasmussen DM, Woods GL. Effect of repeated administration of oxytocin during diestrus on duration of function of corpora lutea in mares. J Am Vet Med Assoc 2007;231(12):1864–7. Ying SY. Inhibins, activins, and follistatins: gonadal proteins modulating the secretion of follicle-stimulating hormone. Endocr Rev 1988;9(2):267–93. Moore KH, Dunbar BS, Bousfield GR, Ward DM. Initial characterization of equine inhibin. Biol Reprod 1994;51(1):63–71. Forage RG, Ring JM, Brown RW, McInerney BV, Cobon GS, Gregson RP, Robertson DM, Morgan FJ, Hearn MT, Findlay
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25.
26.
27.
28.
29.
30. 31.
32.
33.
34.
35.
36. 37.
38.
39.
JK. Cloning and sequence analysis of cDNA species coding for the two subunits of inhibin from bovine follicular fluid. Proc Natl Acad Sci USA 1986;83(10):3091–5. Knight PG, Glister C. Potential local regulatory functions of inhibins, activins and follistatin in the ovary. Reproduction 2001;121(4):503–12. Nagamine N, Nambo Y, Nagata S, Nagaoka K, Tsunoda N, Taniyama H, Tanaka Y, Tohei A, Watanabe G, Taya K. Inhibin secretion in the mare: localization of inhibin alpha, betaA, and betaB subunits in the ovary. Biol Reprod 1998;59(6):1392–8. Arai KY, Tanaka Y, Taniyama H, Tsunoda N, Nambo Y, Nagamine N, Watanabe G, Taya K. Expression of inhibins, activins, insulin-like growth factor-I and steroidogenic enzymes in the equine placenta. Domest Anim Endocrinol 2006;31(1):19–34. Tanaka Y, Taniyama H, Tsunoda N, Shinbo H, Nagamine N, Nambo Y, Nagata S, Watanabe G, Herath CB, Groome NP, Taya K. The testis as a major source of circulating inhibins in the male equine fetus during the second half of gestation. J Androl 2002;23(2):229–36. Nagata S, Tsunoda N, Nagamine N, Tanaka Y, Taniyama H, Nambo Y, Watanabe G, Taya K. Testicular inhibin in the stallion: cellular source and seasonal changes in its secretion. Biol Reprod 1998;59(1):62–8. Roser JF. Endocrine and paracrine control of sperm production in stallions. Anim Reprod Sci 2001;68(3–4):139–51. Watson ED, Heald M, Tsigos A, Leask R, Steele M, Groome NP, Riley SC. Plasma FSH, inhibin A and inhibin isoforms containing pro- and -alphaC during winter anoestrus, spring transition and the breeding season in mares. Reproduction 2002;123(4):535–42. Nambo Y, Nagaoka K, Tanaka Y, Nagamine N, Shinbo H, Nagata S, Yoshihara T, Watanabe G, Groome NP, Taya K. Mechanisms responsible for increase in circulating inhibin levels at the time of ovulation in mares. Theriogenology 2002;57(6):1707–17. Yamanouchi K, Hirasawa K, Hondo E, Hasegawa T, Ikeda A, Sugawara Y, Matsuyama S, Miyazawa K, Sawasaki T, Tojo H, Tachi C, Takahashi M. Expression and cellular localization of inhibin alpha-subunit mRNA in equine fetal gonads. J Vet Med Sci 1997;59(7):569–73. Meunier H, Rivier C, Evans RM, Vale W. Gonadal and extragonadal expression of inhibin alpha, beta A, and beta B subunits in various tissues predicts diverse functions. Proc Natl Acad Sci USA 1988;85(1):247–51. Watson ED, Thomassen R, Steele M, Heald M, Leask R, Groome NP, Riley SC. Concentrations of inhibin, progesterone and oestradiol in fluid from dominant and subordinate follicles from mares during spring transition and the breeding season. Anim Reprod Sci 2002;74(1–2): 55–67. Roser JF, McCue PM, Hoye E. Inhibin activity in the mare and stallion. Domest Anim Endocrinol 1994;11(1):87–100. Dissel-Emiliani FM, Grootenhuis AJ, de Jong FH, de Rooij DG. Inhibin reduces spermatogonial numbers in testes of adult mice and Chinese hamsters. Endocrinology 1989;125(4):1899–903. Mather JP, Moore A, Li RH. Activins, inhibins, and follistatins: further thoughts on a growing family of regulators. Proc Soc Exp Biol Med 1997;215(3):209–22. Ling N, Ying SY, Ueno N, Shimasaki S, Esch F, Hotta M, Guillemin R. A homodimer of the beta-subunits of inhibin A stimulates the secretion of pituitary follicle stimulating hormone. Biochem Biophys Res Commun 1986;138(3): 1129–37.
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40. Besecke LM, Guendner MJ, Sluss PA, Polak AG, Woodruff TK, Jameson JL, Bauer-Dantoin AC, Weiss J. Pituitary follistatin regulates activin-mediated production of folliclestimulating hormone during the rat estrous cycle. Endocrinology 1997;138(7):2841–8. 41. Woodruff TK. Regulation of cellular and system function by activin. Biochem Pharmacol 1998;55(7):953–63. 42. Ling N, Ying SY, Ueno N, Shimasaki S, Esch F, Hotta M, Guillemin R. Pituitary FSH is released by a heterodimer of the beta-subunits from the two forms of inhibin. Nature 1986;321(6072):779–82. 43. Doi M, Igarashi M, Hasegawa Y, Eto Y, Shibai H, Miura T, Ibuki Y. In vivo action of activin-A on pituitary-gonadal system. Endocrinology 1992;130(1):139–44. 44. Lejeune H, Chuzel F, Sanchez P, Durand P, Mather JP, Saez JM. Stimulating effect of both human recombinant inhibin A and activin A on immature porcine Leydig cell functions in vitro. Endocrinology 1997;138(11):4783–91. 45. de Kretser DM, Buzzard JJ, Okuma Y, O’Connor AE, Hayashi T, Lin SY, Morrison JR, Loveland KL, Hedger MP. The role of activin, follistatin and inhibin in testicular physiology. Mol Cell Endocrinol 2004;225(1–2):57–64. 46. Meehan T, Schlatt S, O’Bryan MK, de Kretser DM, Loveland KL. Regulation of germ cell and Sertoli cell development by activin, follistatin, and FSH. Dev Biol 2000;220(2):225–37. 47. Sherwood OD. Relaxin’s physiological roles and other diverse actions. Endocr Rev 2004;25(2):205–34. 48. Schwabe C, McDonald JK. Relaxin: a disulfide homolog of insulin. Science 1977;197(4306):914–15. 49. Schwabe C, Bullesbach EE. Relaxin: structures, functions, promises, and nonevolution. FASEB J 1994;8(14):1152–60. 50. Klonisch T, Hombach-Klonisch S. Review: Relaxin Expressed at the Feto-Maternal Interface. Reprod Domest Anim 2000;35(3–4):149-52. 51. Stewart DR, Stabenfeldt GH, Hughes JP, Meagher DM. Determination of the source of equine relaxin. Biol Reprod 1982;27(1):17–24. 52. Klonisch T, Mathias S, Cambridge G, Hombach-Klonisch S, Ryan PL, Allen WR. Placental localization of relaxin in the pregnant mare. Placenta 1997;18(2–3): 121–8. 53. Stewart DR, Stabenfeldt G. Relaxin activity in the pregnant mare. Biol Reprod 1981;25(2):281–9. 54. Stewart DR, Kindahl H, Stabenfeldt GH, Hughes JP. Concentrations of 15-keto-13, 14-dihydro-prostaglandin F2 alpha in the mare during spontaneous and oxytocin induced foaling. Equine Vet J 1984;16(4):270–4. 55. Stewart DR, Addiego LA, Pascoe DR, Haluska GJ, Pashen R. Breed differences in circulating equine relaxin. Biol Reprod 1992;46(4):648–52. 56. Haluska GJ, Currie WB. Variation in plasma concentrations of oestradiol-17 beta and their relationship to those of progesterone, 13,14-dihydro-15-keto-prostaglandin F-2 alpha and oxytocin across pregnancy and at parturition in pony mares. J Reprod Fertil 1988;84(2):635–46. 57. Ryan PL, Vaala WE, Bagnell CA. Evidence that equine relaxin is a good indicator of placental insufficiency in the mare. Proceedings of the 44th annual convention of the American Association of Equine Practitioners, 1998; pp. 62–3. 58. Ryan PL, Christiansen DL, Hopper RM, Bagnell CA, Vaala WE, Leblanc MM. Evaluation of systemic relaxin blood profiles in horses as a means of assessing placental function in high-risk pregnancies and responsiveness to therapeutic strategies. Ann N Y Acad Sci 2009;1160: 169–78.
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59. Ryan PL, Klonisch T, Yamashiro S, Renaud RL, Wasnidge C, Porter DG. Expression and localization of relaxin in the ovary of the mare. J Reprod Fertil 1997;110(2):329–38. 60. Song L, Ryan PL, Porter DG, Coomber BL. Effects of relaxin on matrix remodeling enzyme activity of cultured equine ovarian stromal cells. Anim Reprod Sci 2001;66(3–4):239–55. 61. Smith MS. Anterior pituitary hormones: development of a bioassay leading to the discovery of prolactin. AJP Endocrinol Metab 2004;287(5):E813–14. 62. Riddle O, Bates RW, Dykshorn SW. The preparation, identification and assay of prolactin: a hormone of the anterior pituitary. Am J Physiol 1933;105:191–216. 63. Rahmanian MS, Thompson DL Jr, Melrose PA. Immunocytochemical localization of prolactin and growth hormone in the equine pituitary. J Anim Sci 1997;75(11): 3010–18. 64. Thompson DL Jr, DePew CL, Ortiz A, Sticker LS, Rahmanian MS. Growth hormone and prolactin concentrations in plasma of horses: sex differences and the effects of acute exercise and administration of growth hormone-releasing hormone. J Anim Sci 1994;72(11):2911–18. 65. Worthy K, Colquhoun K, Escreet R, Dunlop M, Renton JP, Douglas TA. Plasma prolactin concentrations in nonpregnant mares at different times of the year and in relation to events in the cycle. J Reprod Fertil Suppl 1987;35:269–76. 66. Thompson DL Jr, Johnson L, Wiest JJ. Effects of month and age on prolactin concentrations in stallion serum. J Reprod Fertil Suppl 1987;35:67–70. 67. Thompson FN, Caudle AB, Kemppainen RJ, Nett TM, Brown J, Williams DJ. Thyroidal and prolactin secretion in agalactic mares. Theriogenology 1986;25(4):575–80. 68. Johnson AL. Seasonal and photoperiod-induced changes in serum prolactin and pituitary responsiveness to thyrotropin-releasing hormone in the mare. Proc Soc Exp Biol Med 1987;184(1):118–22.
69. Aurich C, Gerlach T, Aurich JE, Hoppen HO, Lange J, Parvizi N. Dopaminergic and opioidergic regulation of gonadotropin and prolactin release in stallions. Reprod Domest Anim 2002;37(6):335–40. 70. Worthy K, Escreet R, Renton JP, Eckersall PD, Douglas TA, Flint DJ. Plasma prolactin concentrations and cyclic activity in pony mares during parturition and early lactation. J Reprod Fertil 1986;77(2):569–74. 71. Ginther OJ. Reproductive seasonality. In: Ginther OJ (ed.) Reproductive Biology of the Mare: Basic and Applied Aspects. Cross Plains, WI: Equiservices, 1992; pp. 105–34. 72. Nequin LG, King SS, Johnson AL, Gow GM, Ferreira-Dias GM. Prolactin may play a role in stimulating the equine ovary during the spring reproductive transition. J Equine Vet Sci 1993;13(11):631–5. 73. King SS, Campbell AG, Dille EA, Roser JF, Murphy LL, Jones KL. Dopamine receptors in equine ovarian tissues. Domest Anim Endocr 2005;28:405–15. 74. King SS, Jones KL, Dille EA, Roser JF. Prolactin receptors in the corpus luteum of mares. Proceedings of 15th International Congress on Animal Reproduction, 2004, Abstract 39. 75. King SS, Douglas BL, Roser JF, Silvia WJ, Jones KL. Differential luteolytic function between the physiological breeding season, autumn transition and persistent winter cyclicity in the mare. Anim Reprod Sci 2010;117: 232–40. 76. Jedlinska M, Rozewiecka L, Ziecik AJ. Effect of hypoprolactinaemia and hyperprolactinaemia on LH secretion, endocrine function of testes and structure of seminiferous tubules in boars. J Reprod Fertil 1995;103(2):265–72. 77. Regisford EG, Katz LS. Effects of bromocriptine-induced hypoprolactinaemia on gonadotrophin secretion and testicular function in rams (Ovis aries) during two seasons. J Reprod Fertil 1993;99(2):529–37.
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176.
Puberty Bruce E. Eilts
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Anestrus Donald L. Thompson, Jr.
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Vernal Transition into the Breeding Season Daniel C. Sharp
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Estrus Patrick M. McCue, Charles F. Scoggin and Alicia R.G. Lindholm
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Diestrus ¨ Robert M. Lofstedt
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Autumnal Transition Out of the Breeding Season Sheryl S. King
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The Abnormal Estrous Cycle Patrick M. McCue and Ryan A. Ferris
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Chapter 176
Puberty Bruce E. Eilts
Introduction The onset of puberty in most production livestock species is integral in maximizing profitability. Earlier puberty means shorter generation intervals, therefore allowing improved genetics to be introduced sooner. Attaining early puberty in the mare has not been as important as in cattle and swine, therefore the study and pursuit of earlier puberty onset in the mare has not been as intense as in other livestock species. Performance history plays a major role in the selection of which mares will be bred, therefore most mares will have attained puberty by the time their performance history has been completed. In breeds in which pedigree is nearly as important as a dam’s performance history in selecting matings, breeding regulations may prevent the use of many assisted reproductive techniques that could be employed on peripubertal fillies. Rarely have horse owners requested information on how to obtain offspring from peripubertal fillies. The Assisted Reproduction Technology program at the Animal Reproduction and Biotechnology Laboratory at Colorado State University, as of June 2007, had had no requests for assisted reproductive techniques to be performed in young, peripubertal fillies (E.M. Carnevale, personal communication), whereas it is not an uncommon procedure in cattle. It has been shown that successful embryo collections could be performed on Haflinger pony mares as young as 12–16 months of age; such collections resulted in embryo recovery rates and pregnancy rates no different from 2- and 3-year-old donors.1 There has been interest, however, in the potential need to harvest oocytes after the death of a valuable filly. If colts and fillies are commingled, knowing the approximate age of puberty onset in the fillies may be vital so that the sexes can be separated well in advance of time of puberty to prevent unwanted matings. If unwanted matings do occur, knowledge of the pregnancy risks of early puberty matings enables a more scientific approach
to decisions about how to handle the pregnancy. The importance of puberty in the mare, the age of onset, and factors that influence puberty onset will be presented in this chapter. Many papers state birth dates, first estrus dates, and date of first ovulation; however, some papers do not definitively state the birth date, age at pubertal estrus, or age of pubertal ovulation. Estimates and calculations may have been performed to extrapolate data from some cited papers in an attempt to compile a more complete data set in writing this chapter. Since some cited studies were performed in the northern hemisphere and some in the southern hemisphere, this chapter will standardize the definitions across papers written in the northern and southern hemisphere (Table 176.1). All birth dates will be referred to as days before (negative) or days after (positive) the summer solstice, and puberty date will be referred to as months from the winter solstice of the birth year. For example, a filly born in April in the northern hemisphere will be said to be born 60 days (–60) before the summer solstice, as would a filly born in October in the southern hemisphere. Similarly, a mare that first ovulates in May in the northern hemisphere will be said to have attained puberty 5 months after the winter solstice after its birth year, as would a filly ovulating in November in the southern hemisphere.
Age at puberty Age at puberty (in months) in fillies housed in traditional horse-rearing settings, summarized from 14 papers, ranged from 7.8 to 37 months with a median of 15 months (see Table 176.2 for a complete listing). These studies differed widely in breeds, birth date, locales, management, and puberty definition of the fillies studied, thus resulting in much variation in the ‘average’ onset of puberty in the mare.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: The Estrous Cycle Table 176.1 Seasonal months in the northern vs southern hemispheres Northern
Southern
Months from winter solstice
Month
Months from winter solstice
1 2 3 4 5 6 7 8 9 10 11 0
January February March 21 April May June 21 July August September 21 October November December 21
7 8 9 10 11 0 1 2 3 4 5 6
Spring equinox
Summer solstice
Fall equinox
Winter solstice
Pubertal estrus Puberty is often defined as the onset of the first estrus. Criteria used to identify the pubertal estrus were the first rise in plasma progesterone,2–9 the initial estrous behavior at teasing,10, 11 the first observed estrus (S.M. McDonnell, unpublished data for the New Bolton Center semi-feral herda ) or the presence of a corpus luteum at slaughter.12 In 13 studies that defined puberty using the first estrus or first ovulation based upon a rise in progesterone, the age in months to attain puberty ranged from 7.8 to 27.0 months with a median of 14.3 months (see Table 176.2 for a complete listing). When both the age at first estrus and age at first ovulation were reported, there was little difference seen, as seven of eight pony fillies ovulated during the first noted estrus and one of eight ovulated during the subsequent full estrus.5 It is interesting to note that there seemed to be minimal transitional estrous periods preceding the initial pubertal estrus in the ponies studied.5 Hormonal stimulus of pubertal estrus An interesting, simple overview of female puberty in domestic animals has been proposed by Senger and a
SM McDonnell: unpublished data on the New Bolton Center semi-feral herd of 50 to 80 Shetland ponies on approximately 20 hectares. The herd is continuously in a natural environment (light and weather) and natural social conditions with only natural shelter. Foals are born in the enclosure and never get out until they leave the project. Supplemental hay of the same grass pasture mix is scattered in hard winters. The animals have salt and mineral blocks, but no other supplements. The animals get no vaccinations for reproductive diseases, only Eastern and Western encephalitis, tetanus, and rabies. They are on a strategic deworming protocol (fecal egg counts and they are dewormed as needed). The herd is observed for a minimum of 2 hours per day 365 days per year, in one-hour sessions in the morning and evening.
Fall equinox
Winter solstice
Spring equinox
Summer solstice
is outlined below.13 Before puberty, the tonic centers in the hypothalamus have a high sensitivity to the negative feedback of low concentrations of estrogen, which prevents sufficient follicle-stimulating hormone (FSH) and luteinizing hormone (LH) release for adequate follicular development or ovulation. At puberty, the negative feedback sensitivity on the tonic center decreases and allows the release of more gonadotropin-releasing hormone (GnRH), which leads to increased FSH production. Increased FSH stimulates follicles to produce high estrogen concentrations. The high estrogen has a positive feedback on the surge center and causes sufficient LH release to cause ovulation. Factors that influence the age at which puberty is attained include attainment of a threshold body size or fat composition, environmental cues such as season of birth and photoperiod, and social cues such as the presence of other females or males. Once the proper interaction of these factors is in place to initiate puberty, the surge center in the hypothalamus releases sufficient quantities of GnRH, when stimulated by estradiol from follicles on the ovary, to cause a surge of LH from the anterior pituitary. This LH surge then causes ovulation of the estradiol-producing follicle. While this first ovulation is considered by many as the onset of puberty, three definitions of puberty are presented by Senger.13 These definitions are: (i) the age at first estrus; (ii) the age at first ovulation; and (iii) the age at which a female can support pregnancy without deleterious effects. The exact stimuli that initiate puberty in the mare, as defined by the first estrus or ovulation, are not totally known. The LH and FSH secretion patterns in prepubertal pony fillies,14 peripubertal pony fillies5, 15 and horse fillies11 have been examined. Pony fillies in Wisconsin born 2 to 3 months before the summer solstice (April) had basal LH concentrations
Brazil
Italy Italy Slovenia Slovenia Wisconsin Wisconsin
Wisconsin Wisconsin
Wisconsin England England France
France
Colorado Australia New Zealand New Zealand New Zealand Brazil Pennsylvania
Traditional
Traditional Traditional Traditional Traditional Traditional Traditional
Traditional Traditional
Traditional Traditional Traditional Traditional
Traditional
Traditional Traditional Traditional
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b
a
6 6 11 13 10 6 5 4 4 2 35 33 10 54
43◦ 43◦ 43◦ 53◦ 53◦ 47◦ 47◦ 40◦ 27◦ 40◦ 40◦ 40◦ 23◦ 26 39◦
Unpublished data, personal communication. Not stated.
4 4 8 7 3 5
43◦ 43◦ 45◦ 40 45◦ 40 43◦ 43◦
TB Ponies, Shetland
TB
TB
Quarter Horses Stock horses TB
Ponies, Welsh
Ponies Ponies, New Forest Ponies, New Forest Ponies, Welsh
Ponies Ponies
−124 −43
−53
−46
30 −50 66
−20
−173 −51 −51 −141
−58 −81
−49 −132 −96 −96 4 −38
NSb
−50 −23 −27 NSb
Stud farm Natural social organization enclosed
Fescue pasture
Fescue pasture
Drylot Paddock Fescue pasture
Stable
Barn – natural light Pasture Pasture Stable
Barn – natural light Slaughter
Paddock Paddock Free stable Free stable Barn – natural light Barn – artificial light
NSb
Free range Free range Free range NSb
Progesterone Estrus/foaling
Progesterone
Progesterone
Tease/Ultrsound Progesterone Progesterone
Progesterone
Progesterone Tease Tease Progesterone
Progesterone CL
Progesterone Progesterone Progesterone Progesterone Progesterone Progesterone
Progesterone
Foaling Foaling Foaling Progesterone
Puberty basis
14.4 12.0
11.7
11.5
22.5 13.1 7.8
27.0
11.8 15.0 14.5
13.6 15.2
12.1 14.3 15.3 25.3 11.3 13.2
25.0
37.0 25.0 36.0 15.0
127 134
117
117
522 162 119
607
xx 121 215 111
169 192
131 114 179 480 162 177
NSb
877 544 870 NSb
Nogueira et al.9 McDonnella
Brown-Douglas et al.8
Brown-Douglas et al.8
Wesson and Ginther5 Wesson and Ginther20 Wesson and Ginther5 Wesson and Ginther12 Wesson and Ginther5 Ellis and Lawrence10 Ellis and Lawrence10 Palmer and Driancourt6 Palmer and Driancourt6 Naden et al.11 Skelton et al.7 Brown-Douglas et al.8
Camillo et al.3 Camillo et al.3 ˇ Cebulj-Kadunc et al.4 ˇCebulj-Kadunc et al.4
Souza et al.2
Keiper and Houpt19 17 Dyrmundsson ´ 18 Lucas et al. Souza et al.2
Reference
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Traditional Free roam
20
20◦ 43
Assateague Icelandic horse Sable horse Brasileiro de Hipismo Brasileiro de Hipismo Haflinger Haflinger Lippizan Lippizan Ponies Ponies
Housing type
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93 136 50 20
38◦ 64◦ 44◦ 20◦ 43
Breed
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Maryland Iceland Nova Scotia Brazil
Feral Feral Feral Traditional
n
Latitude
Days to puberty from first winter solstice
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First estrus (months)
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Days ± summer solstice from birth date
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Table 176.2 Summary of puberty data in mares
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to 10 months of age, but FSH rose at 2–3 months of age and at 1 month past the summer solstice (June and July) to a high at 3–4 months of age (2–3 months past the summer solstice in August and September) and then fell to lows at 6–8 months of age (6 months past the summer solstice – October to early December) and then began to rise again at 8–9 months of age (1 month past the winter solstice – December and January).14 Fillies were not followed to the first ovulation, but the pattern appeared similar to that seen in other studies when pony mares were followed to puberty. In a similar study, pony fillies in Wisconsin born 2–3 months before the summer solstice (April) or around the summer solstice (June and July) had LH and FSH profiles similar to post-pubertal mares, which followed daylength patterns (low around the winter solstice and higher around the summer solstice).15 Mares in Colorado born 1–2 months after the summer solstice (July and August) had basal LH concentrations until an initial rise at 7 months of age (1–2 months after the winter solstice – January to March) that peaked at 10 months of age (4–6 months past the winter solstice – April to June) and then declined at 16 months of age (5 months past the summer solstice – November).11 The FSH had an initial rise at 7 months of age (1 month after the winter solstice – January) and peaked at 10 months of age (4–6 months past the winter solstice – April to June) and then remained elevated.11 These Quarter Horse fillies were also challenged with GnRH and were capable of responding with significant increases in LH at 7 months of age (2 months past the winter solstice – February) to 13 months of age (4 months past the winter solstice – April), but the response declined at 16.8 months of age (4 months past the summer solstice – October). The FSH response to a GnRH challenge was an increase at 7.4 to 13 months of age (3–6 months past the winter solstice – March to June), a decrease at 15 months of age (3 months past the summer solstice – September), and a rise again at 16.8 months of age (4 months past the summer solstice – October) then a decline at 22.4 months of age (1–2 months past the summer solstice – July and August).11 The age of puberty was 22.5 months (4 months after the second winter solstice of their birth – April).11
Puberty as defined by foaling The definition of puberty as the time of first estrus or ovulation seems to be much different than that of first foaling. The first fertile estrus, based on foals born 11 months later was seen at 12–14 months old in mixed breed fillies,16 but not seen until 25,17 36,18 and 3719 months in feral herds in Iceland, Nova Scotia, and Maryland, respectively. Foaling rates were only 4.1% (3/73) in mares bred as yearlings in feral Sable Island Horses in Nova Scotia, and rose with age to 62.5%
(25/40) in those potentially bred as 3-year-olds.18 Interestingly, of 10 mares that were presumed pregnant by fecal estrogen assays when bred as yearlings, seven appeared to have lost their pregnancies.18 A similar finding was noted earlier in 12–14-monthold mixed breed fillies by Mitchell and Allen,16 where 69% of those mares lost their pregnancies by 5 months of gestation and 55% had stillborn foals. This is in contrast to the first estrus (observed and based on foaling) at 12 months in 53/54 (98%) ponies housed in a semi-feral herd at the University of Pennsylvania New Bolton Center (described above). These young foaling ponies tended to have small but mature foals, but accounted for all three of the dystocias in the 175 births observed. Maternal behavior in the primiparous mares was sometimes lax, but the foals usually kept pace with the family, or were tended by older mares and the harem stallion. When comparing other criteria influencing the age of puberty using foaling as the definition, it does not appear that location or latitude caused differences. In Iceland at 64◦ N, 136 Icelandic horses had the first fertile estrus at around 25 months;17 in Nova Scotia at 44◦ N, 50 Sable horses had the first fertile estrus at around 36 months;18 and in Maryland at 38◦ N, 93 Assateague horses had the first fertile estrus at 37 months.19 It appears then that the 54 ponies observed by McDonnell in Pennsylvania at 39◦ N had the earliest fertile estrus at 12 months. The pubertal fertile estrus occurred at 69% (90 kg) of the mature weight (mature weight was 131 kg). Of course a confounding factor in the McDonnell herd was that it was not completely feral, and was supplemented with grass hay in the winter. This could have given the fillies more energy to grow and mature than their wild range counterparts. Another study of 13 New Forest ponies raised in a pasture setting in England at 53◦ N under good or poor nutrition, showed the first pubertal estrus was at 12 and 15 months, respectively.10 Although these fillies were not feral and did not have foals, the first pubertal estrus was similar to those observed by McDonnell. This would indicate that the nutritional status of the fillies observed by McDonnell was not a major factor in attaining puberty earlier than the other breeds in the feral or semi-feral herds. It appears that the only studies actually looking at first foaling (and then calculating back to the first fertile estrus) indicate that breed is probably a major factor in attaining puberty.
Puberty as defined by first estrus McDonnell’s observation certainly confirms that the pubertal estrus can be fertile in pony fillies; however, the presumed prenatal loss in the Sable Island Horses in Nova Scotia18 and Canada mixed breed mares16 prevents this from being a universal statement. The
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Puberty first pubertal estrus cannot be equated with the first time a filly is capable of carrying a pregnancy to term in all breeds. Although breedings at the pubertal estrus and subsequent pregnancy are only documented in ponies in a semi-feral herd (as described by McDonnell; see above), these breedings probably have occurred countless times in the history of human domestication of the horse, but are not well documented in the scientific literature. The first pubertal estrus is therefore the only criterion to enable direct comparisons of the onset of puberty between different breeds and locales. This may not be true if the definition of puberty is taken to be the age at which a mare can first sustain pregnancy, however. In examining breeds, ponies from six studies had their first estrus at ages ranging from 11 to 27 months, with a median of 14 months (see Table 176.2 for a complete listing). The age of pubertal estrus in ponies does not seem appreciably different from the age of pubertal estrus in eight studies of large mares, which ranged from 8 to 25 months with a median of 14 months (see Table 176.2 for a complete listing). Birth date and pubertal estrus The main difference in the age at attaining puberty in the larger breeds was primarily birth date and not nutritional status. Haflingers born 49 days before the summer solstice had their first estrus 131 days after the winter solstice of their birth year while those born 132 days before the summer solstice had their first estrus 114 days after the winter solstice of their birth year.3 Even though the early born mares were older at puberty, the date at puberty was not different between the two groups. Similarly this was seen in Thoroughbred mares in New Zealand. Mares born 66 days after the summer solstice attained puberty at 119 days after the winter solstice of their birth year while those born 53 days before the summer solstice attained puberty 117 days after the winter solstice of their birth year.8 As in the Italian study on Haflingers,3 even though one group of mares was older, they attained puberty at the same time in relation to the first winter solstice of their birth year. The estrous cycles subsequent to the pubertal estrus have not been extensively studied. In late born ponies (4 days after the summer solstice), the number of cycles post-puberty tended to be fewer, with fewer ovulations and a shorter breeding season than the early born (58 days before the summer solstice).5 Estrus accompanied 91% of ovulations.5 Daylength and pubertal estrus Another factor that could influence the age of pubertal estrus or ovulation is length of daylight. This does not seem to be supported by the data on latitude. In
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ponies, the latitude, age at pubertal estrus (months), and days to pubertal estrus after the winter solstice (Figure 176.1 and Table 176.2) were: 39◦ , 12, 134 (S.M. McDonnell, unpublished); 43◦ , 11, 162;5 43◦ , 13, 177;20 43◦ , 14, 169;5 43◦ , 15, 192;12 47◦ , 15, 111;6 47◦ , 27, 607;6 53◦ , 12, 121.2;10 and 53◦ , 15, 215.4.10 In horses it was 23◦ 26 , 14, 127;9 27◦ , 13, 162;7 40◦ , 8, 119;8 40◦ , 12, 117;9 40◦ , 23, 522;11 43◦ , 12, 131;3 43◦ , 14, 114;3 45◦ 40 , 15, 179;4 and 45◦ 40 , 25, 480.4 The absence of effect of daylength on puberty was also seen experimentally when it was shown that the addition of 16 hours of artificial light actually interfered with the pubertal estrus in ponies and resulted in fewer ponies ovulating, fewer ovulations, and fewer estrous periods per filly.20 Bodyweight and pubertal estrus Limited studies have been done to examine the bodyweight at the pubertal estrus. Pubertal estrus was seen on average at 49% (280 kg) of mature (575 kg) weight in Thoroughbreds in New Zealand if the correct photo cues were in place.8 The weight at pubertal estrus in ponies in Wisconsin was 135 kg; however, the adult weight and percentage of adult weight at the first pubertal estrus were not stated.5 This is in contrast to the age of onset of first ovulation not being correlated with bodyweights in Brasilieiro de Hipismos fillies in Brazil.2 Fillies had their first ovulation at 15 months and weighed 338 ± 8 kg, while those having their first ovulation in their second season at 25 months weighed 411 ± 20 kg.2 As stated, if the correct photo cues are in place, pubertal estrus probably occurs at approximately 50 to 60% of mature weight.
Social cues affecting puberty The least known aspects regarding puberty in the mare are the social cues that may be involved. It is known in swine and cattle that different social conditions can hasten or delay puberty. As Senger summarized,13 gilts raised in small groups have delayed puberty compared to those raised in large groups. In addition, the presence of a boar, either in direct contact or in close proximity, hastened puberty in gilts raised in small groups. Senger13 also noted that bulls hastened the onset of puberty in heifers. Social cues and their effect on puberty have not been directly investigated in the equine. In fact, none of the published papers investigating puberty in the horse clearly describes the potential social interactions of colts and fillies or of stallions and fillies. Papers generally state that animals were kept in a pasture or paddock and imply that sexes were not commingled, but do not state if colts or stallions were in direct or indirect contact. Traditional horse-rearing practices
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Shetland Ponies 134 d Quarter Horses 521 d
New Forest Ponies 121, 215 d
Icelandic Horse 543 d
Ponies 162, 177, 169, 192 d Sable Horse 870 Assateague 877 d
Lipizzan 178, 479 d Welsh Ponies 111, 607 d
Haflinger 131, 114 d
Stock Horses 162 d
Thoroughbreds 127 d
Thoroughbreds 117, 119 d
Figure 176.1 Map depicting locations of cited studies describing mare puberty. Square boxes represent mares raised in traditional manners, octagons represent feral mares. Ponies are listed in italics. Days refers to the days after the first winter solstice in life that puberty was attained.
are to commingle colts and fillies to weanling age (150–210 days) and then separate them. There is usually no direct stallion contact, although indirect stallion exposure may occur. This is completely different than wild horses, which run in bands with fillies directly exposed to colts, mares, and stallions. The first fertile estrus in the semi-feral ponies that had the colts and the fillies commingled did not seem to result in a clear-cut earlier onset of estrus. In the six papers studying ponies, the age of pubertal estrus ranged from 11 to 27 months with a median of 14 months, and the days past the winter solstice ranged from 111 to 607 days with a median of 169 days (see Table 176.2 for a complete listing). The feral Icelandic horses, Sable horses and Assateagues had their first estrus 544,17 870,18 and 87719 days after the first winter solstice of life. Valid direct comparisons between these different breeds and conditions of foal sexes that were commingled make it impossible to draw any real conclusions on the effect of commingling versus other effects on the initiation of puberty. The effect of an adult male on age of puberty is probably greatly overshadowed by the photo cues that are essential in stimulating FSH and LH before initial puberty.
Summary In summary, there has been much less work done on investigating puberty in the mare than in the other domestic species. This is probably because traditional breeding practices have not required breeding mares as young as possible to reduce generation intervals. Since the studies vary widely as to puberty definition,
breed, management practices and locale, it is difficult to make universal statements regarding puberty in the mare. Ponies in a semi-feral situation routinely attained a fertile pubertal estrus after the first winter solstice following their birth and foaled after the second winter solstice following their birth. Feral horses appeared to attain a fertile pubertal estrus approximately 1 year older than ponies. This may be a result of breed or management conditions. Large horses often have their pubertal estrus by the first winter solstice after their birth; however, depending on breed, this may not occur until after the second winter solstice of their birth. No critical studies have looked at the effect of breeding larger horses on pubertal estrus; however, this management practice is generally discouraged to allow mares to attain full body maturity before breeding. Management interventions of breeding age and breeding season make retrospective studies of breeding records impossible for studying puberty in the mare. It does appear that if yearlings are commingled or oocyte collection is desired, the majority of mares have a pubertal estrus in the first year after the first winter solstice following their birth.
References 1. Panzani D, Rota A, Pacini M, Vannozzi I, Camillo F. One year old fillies can be successfully used as embryo donors. Theriogenology 2007;67:367–71. 2. Souza JAT, Gacek F, Oliveira JV, Augusto C. Growth and onset of puberty in female Brazilian Sport Horses. Rev Bras Reprod Anim 1997;21:117–20. 3. Camillo F, Vannozzi I, Rota A, Panzani D, Illuzzi A, Guillaume D. Age at puberty, cyclicity, clinical response to PGF2α, hCG and GnRH and embryo recovery rate in yearling mares. Theriogenology 2002;58:627–30.
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Puberty ˇ 4. Cebulj-Kadunc N, Cestnik V, Kosec M. Onset of puberty and duration of seasonal cyclicity in Lipizzan fillies. Equine Vet J 2006;38:350–3. 5. Wesson JA, Ginther OJ. Puberty in the female pony: reproductive behavior, ovulation, and plasma gonadotropin concentrations. Biol Reprod 1981;24:977–86. 6. Palmer E, Driancourt MA. Some interactions of season of foaling, photoperiod, and ovarian activity in the equine. Livest Prod Sci 1983;10:197–210. 7. Skelton KV, Dowsett KF, McMeniman NP. Ovarian activity in fillies treated with anabolic steroids prior to the onset of puberty. J Reprod Fertil Suppl 1991;44:351–6. 8. Brown-Douglas CG, Firth EC, Parkinson TJ, Fennessy PF. Onset of puberty in pasture-raised Thoroughbreds born in southern hemisphere spring and autumn. Equine Vet J 2004;36:499–504. 9. Nogueira GP, Barnabe RC, Verreschi ITN. Puberty and growth rate in Thoroughbred fillies. Theriogenology 1997;48:581–8. 10. Ellis RNW, Lawrence LJ. Energy under-nutrition in the weanling filly foal. I. Effects on subsequent live-weight gains and onset of oestrus. Brit Vet J 1978;134:205–11. 11. Naden J, Squires EL, Nett TM. Effect of maternal treatment with altrenogest on age at puberty, hormone concentrations, pituitary response to exogenous GnRH, oestrous cycle characteristics and fertility of fillies. J Reprod Fertil 1990;88:185–95.
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12. Wesson JA, Ginther OJ. Influence of season and age on reproductive activity in pony mares on the basis of a slaughterhouse survey. J Anim Sci 1981;52:119–29. 13. Senger PL. Puberty. In: Senger PL (ed.) Pathways To Pregnancy and Parturition, 2nd edn. Ephrata, PA: Cadmus Professional Curriculum, 2003; pp. 128–43. 14. Wesson JA, Ginther OJ. Plasma gonadotropin levels in intact and ovariectomized prepubertal ponies. Biol Reprod 1979;20:1099–104. 15. Wesson JA, Ginther OJ. Plasma gonadotropin concentrations in intact female and intact and castrated male prepubertal ponies. Biol Reprod 1980;22:541–9. 16. Mitchell D, Allen WR. Observations on reproductive performance in the yearling mare. J Reprod Fertil Suppl 1975;23:531–6. 17. Dyrmundsson OR. Reproduction of Icelandic horses with ´ special reference to seasonal sexual activity. Buv´ ´ ısindi 1994;8:51–7. 18. Lucas Z, Raeside JI, Betteridge KJ. Non-invasive assessment of the incidences of pregnancy and pregnancy loss in the feral horses of Sable Island. J Reprod Fertil Suppl 1991;44:479–88. 19. Keiper R, Houpt K. Reproduction in feral horses: An eightyear study. Am J Vet Res 1984;45:991–5. 20. Wesson JA, Ginther OJ. Influence of photoperiod on puberty in the female pony. J Reprod Fertil Suppl 1982; 32:269–74.
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Chapter 177
Anestrus Donald L. Thompson, Jr.
Anestrus, by definition, is the lack of display of estrus, or sexual receptivity to the stallion, by the mare for an extended period of time. Typically, mares are not suspected to be anestrous unless they do not show any signs of heat for at least 21 days, or for one full estrous cycle. A missed estrus period due to poor heat detection or unfamiliarity with the mare (especially if she has only 1 or 2 days of heat) is not uncommon, and should be one of the first things considered. Due to the multitude of possible underlying causes for anestrus, the term in itself relays only partial information to the clinician or producer. That is, anestrus may be accompanied by anovulation, such as in the case of seasonal anestrus, when the ovaries become small and inactive due to insufficient gonadotropin stimulus.1–3 Alternatively, anestrus may occur in the presence of normal ovarian cyclicity with follicular development and ovulation. Thus, the first step in diagnosing the cause of anestrus should involve an initial assessment of ovarian size and follicular activity. Small, inactive ovaries with no follicles greater than 10 mm is a different problem for the breeder than large, normal appearing ovaries with active folliculogenesis. In the first case, an organic cause with several possible etiologies presents itself, whereas in the latter case, the culprit may be organic (such as high progesterone due to pregnancy or retained luteal tissue) or it may be psychological (Figure 177.1).
Anestrus with ovarian activity For a mare that has not shown any signs of estrus for at least 21 days but has significant follicular activity on her ovaries, the first question to be asked is whether the mare has been bred in the recent past or within the past 11 months. Non-return to estrus is a primary indicator of pregnancy, thus pregnancy status should be determined before proceeding fur-
ther. Pregnant mares have normal follicular waves on the ovaries during the first several months of pregnancy,4 as well as at least one corpus luteum.3 After the demise of all luteal structures (primary and supplementary) around day 180 of gestation, the ovaries regress and become inactive, and remain in this state through parturition.3 Whether the mare is in early or late gestation, a thorough ultrasonographic exam should reveal if she is indeed pregnant. Upon evaluation of the ovaries, if significant ovarian size and follicular activity are detected, yet the mare still has no history of estrus periods, two likely possibilities exist as to the underlying cause. The first is the presence of progesterone, either from luteal tissue, the feto-placental unit, or some ectopic source, sufficient to maintain the mare in a constant diestrous-like state. Measurement of progesterone in a jugular blood sample should provide the necessary information for confirmation. Although corpora lutea can generally be detected by ultrasonographic exam,5 this is not always the case. Concentrations of progesterone of 1 ng/mL or higher usually will keep a mare from showing estrus, and will in fact cause her to show the typical signs of diestrus: aggressive behavior towards the stallion, ears back, kicking, and a swishing of the tail in an irritated manner.6 Under the influence of high progesterone, the cervix will be closed, firm, and elongated, and the uterus will normally have a ‘garden hose’ firmness (tone) upon palpation and no edema.3 If progesterone is indeed high in the mare’s jugular plasma, and it has been confirmed that she is not pregnant, a luteolytic dose of prostaglandin F2α (PGF2α ) should cause lysis of any luteal tissue, if present, and the mare should return to estrus within 3 to 4 days.7 Failure to come into heat after 5 to 6 days may merit another PGF2α injection, and lack of response to a second injection would confirm some non-luteal source of the high progesterone. Other than the feto-placental unit in mid- to late pregnancy,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Anestrus Pregnant Retained luteal tissue
Constant high progesterone
Ectopic luteal tissue or tumor
Active ovaries No progesterone or Cyclic
Behavioral Estrogen insensitivity
Anestrus
Seasonal anestrus Nutritional/seasonal anestrus (extended beyond seasonal) Lactational anestrus
Small, inactive ovaries
Post-Ovuplant anestrus Hypopituitarism GnRH vaccines Senescence Idiopathic
Figure 177.1 Schematic outlining the various forms of anestrus. It is intended to guide diagnosis of the underlying cause(s) of anestrus in clinical and farm settings.
non-luteal sources of progesterone are rare, and may require extensive investigation to elucidate. If progesterone in the initial blood sample is low to undetectable, and significant follicular activity is present, the mare may either be (i) cyclic and the sample was drawn during the follicular phase, or (ii) the mare is acyclic due to some failure of ovulation. These two cases can be differentiated from one another with a second blood sampling 10 to 12 days after the first. High progesterone in the second sample would indicate formation of luteal tissue and cyclicity. Low progesterone in the second sample would indicate lack of cyclicity. If the mare is cyclic, from the standpoint of progesterone levels, and still not displaying estrus, then a psychological cause would be suspected. Mares protective of their foals may display such lack of estrous behavior, even though they have normal follicular activity and estrogen levels in the blood.8 Such mares will often show subtle differences in behavior when in the follicular phase, especially if they can be gradually entrained to brief separation periods from their foals. A few mares may not. In the most stubborn cases, regular ultrasonographic exams and prediction of ovulation based on follicular size and
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morphology may be the only way to know when the mare should be bred. Obviously, artificial insemination (AI) of such mares greatly simplifies the breeding process. If AI is not possible or allowed, such as with Thoroughbred mares, an alternate, more creative, means of getting her bred will be required. In the case of mares with low progesterone in both samples, failure to ovulate accompanied by significant follicular activity is common during the transitional phase from the non-breeding season to the breeding season.3, 9 However, such cases typically display some form of estrous behavior, albeit often irregular in duration or inconsistent from day to day. A mare displaying no signs of estrus for an extended period but having large follicles on her ovaries is rare, and again the cause may be some type of psychological disorder. One might question whether the follicles present are producing sufficient estrogen to cause estrous displays, or whether the mare has a neural lesion preventing her from responding to normal estrogen stimulation. One test to differentiate between these two possibilities would be to administer 100 mg estradiol cypionate i.m., which will increase estradiol levels in the mare to approximately 15–20 pg/mL for 14 days; alternatively, two injections of 10 mg estradiol benzoate in oil 2 days apart will increase estradiol levels to the same degree, but levels will return to undetectable within 2 to 3 days after the second injection.10 If the mare has the ability to respond to estrogen, she should show signs of estrus within 1 to 3 days. Failure to show estrus within 10 days would be indicative of neural abnormality. This diagnosis, coupled with a failure to ovulate over an extended period of time in the presence of large ovulatory-size follicles, would lead to a grim prognosis. For mares showing signs of estrus in response to the estradiol injection, insufficient estrogen production by the follicles would be concluded. Such a state is normal in transitional mares, prior to development of a true dominant follicle.11 However, for a mare in the breeding season to display such symptoms for an extended period of time is rare, and may indicate an insufficiency of luteinizing hormone (LH) secretion by the pituitary gland. Administration of human chorionic gonadotropin (hCG) may provide an appropriate stimulus for follicular maturation and ovulation; alternatively, a gonadotropin-releasing hormone (GnRH) analog, such as deslorelin, in an extended-release formulation, may be tried.
Anestrus with no ovarian activity in winter Anestrus accompanied by anovulation and small inactive ovaries is common in mares in the
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non-breeding season.3 Folliculogenesis on the equine ovary is thought to proceed in the absence of gonadotropins up to a follicular diameter of approximately 2 mm.12, 13 Beyond that size, the follicles are dependent on follicle-stimulating hormone (FSH) in the blood to stimulate further development, and eventually on LH for final maturation and ovulation.14 Because seasonally anovulatory mares often have some, albeit lowered, circulating FSH levels in peripheral blood, they may attain follicular sizes greater than 10 mm, but usually less than 20 to 25 mm due to the absence of LH.3 Pituitary content and secretion of LH is reduced in the winter to a greater extent than FSH; this differential effect on the two gonadotropins was also observed in mares actively immunized against GnRH.15 There appears to be a constitutive component of FSH production and secretion in horses that persists in the face of low (perhaps nil) GnRH input from the hypothalamus.15–17 In contrast, LH production and secretion is highly dependent upon a ‘normal’ level of GnRH input to the pituitary gland.15–17 Seasonal anestrus and anovulation is well documented as to its characteristics and causes. Other chapters in this text describe the transition into the anovulatory state in the fall (see Chapter 181) and back into the breeding season in the spring (see Chapter 178). The seasonal anestrous state is more appropriately called seasonal anovulation, due to the fact that some mares will exhibit signs of estrus, albeit often weak and irregular, even though they have small, inactive ovaries.3 Somewhat paradoxically, mares with very low body condition scores (around 3)18 may even exhibit strong signs of estrus during the winter.19 The duration of the seasonal anovulatory state, as well as the degree to which the ovaries regress, can be greatly influenced by body condition and nutritional plane. Mares with body condition scores of 6 to 9 often continue to cycle, or exhibit significant follicular activity, throughout the winter, especially in temperate climates.19–21 Mares with low body fat content, if kept on a maintenance diet, will tend to experience a more profound regression of the ovaries and a longer period of anovulation.22 As in many cases of reproductive inactivity across species, the hypothalamus appears to be the central control center responsible for the lack of ovarian activity. Seasonally anovulatory mares have lower hypothalamic GnRH content in the winter compared with the breeding season.1 Research has shown that active immunization against GnRH, and consequent reduction in the amount of GnRH reaching the pituitary gland, greatly reduces the amount of LH, and to a lesser extent FSH, in the gland.15, 17 Secretion of LH is low to non-existent in seasonally anovula-
tory mares, as evidenced by circulating blood levels, whereas secretion of FSH is reduced by at least 50%, but usually not to undetectable levels.23, 24 Administration of GnRH or one of its analogs during the nonbreeding season results in little LH response, whereas the FSH response is similar to that of a diestrous mare in the breeding season.25, 26 The mechanisms by which seasonal anovulation is controlled have been studied, and in most instances are similar to those described for other seasonally breeding species. The shorter duration of daylight in the winter appears to be sensed by the horse via a pineal-melatonin signal, similar to other species.9, 27, 28 In some way not yet elucidated, this melatonin signal is interpreted by the hypothalamus, and GnRH production and secretion is altered accordingly (decreased in winter, increased with increasing day length). The pituitary gland and ovaries can be thought of as downstream obligatory responders, because once the GnRH pulse generator29 is turned on, they have no choice but to respond. That is, the increased GnRH input to the pituitary stimulates LH and FSH production and secretion, and the increased circulating LH and FSH concentrations stimulate the ovaries to produce follicles greater than 10 mm, eventually resulting in ovulation. This obligatory status is confirmed by the fact that hourly pulsatile GnRH administration to seasonally anovulatory mares in winter can produce the same downstream events as the naturally occurring increase in activity of the GnRH pulse generator.30
Light treatment Procedures for reversing anestrus and anovulation associated with season include the use of artificial photoperiod and various hormonal manipulations. The stimulatory effect of long daylength on ovarian activity in mares was first reported in 1949 by Burkhardt;31 numerous studies followed, and various forms of long-day photoperiods or their equivalent have been used in the industry since the early 1970s.9, 32, 33 Typically, mares are subjected to a total daylength of 16 h, usually by adding additional light at the end of the day, starting at least 30 days before the desired stimulation in ovarian activity. Because mares stimulated with long days will experience the same transition period-related problems as mares going through normal transition (see Chapter 183), a starting date of mid-December should allow for enough time for stimulation of the ovaries and eventual ovulation for mid-February breeding. Many of the transitional period problems can be avoided by the use of altrenogest, or some similar progestin
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Anestrus alternative, to synchronize and normalize the mare once she achieves significant follicular growth (25 to 30 mm) on her ovaries.34 As stated earlier, body condition profoundly affects the degree of ovarian regression and the duration of the anovulatory state due to season. Most studies with artificial photoperiod-induced ovarian activity in winter were carried out with mares of moderate to high body condition. Stimulation of thin mares in the same manner may not be as effective, and may not be a good idea otherwise. Putting thin mares on a rising plane of nutrition during light stimulation may counterbalance the thin body condition and increase the chances of success, based on similar situations in other large animal species. Hormonal treatments In spite of the on-farm efficacy of photoperiod treatments, they do require a fair amount of resources to enact, both in terms of labor and in facilities. Thus, research into a hormonal ‘silver bullet’ has continued in an effort to find a simple but effective method of stimulating ovarian activity in seasonally anovulatory mares. Successful hormonal treatments to stimulate ovarian activity in seasonally anovulatory mares have included: (i) treatment with GnRH or one of its analogs to mimic the turned-on GnRH pulse generator, and (ii) administration of prolactin, or alternatively, dopaminergic antagonists that stimulate endogenous prolactin secretion. A flurry of research activity on the practical use of GnRH agonists occurred in the late 1980s, and various degrees of success were obtained with multiple formulations.35–37 In spite of the successes, this approach has not gained popular acceptance and use due to the lack of availability of the technology to most producers. Early research with the dopamine antagonist sulpiride revealed that this drug could stimulate endogenous prolactin in the winter,38, 39 and stimulation of prolactin at this time tended to result in increased follicular growth and, in some studies, ovulation.38 Although sulpiride treatment required multiple injections of the compound, which is undesirable from a practical standpoint, further research with the orally active dopamine antagonist, domperidone, revealed that ovulation could in fact be obtained in a practical setting.40 The success with domperidone has resulted in the development of a commercial product (Equidone; Dechra Pharmaceuticals PLC; www.dechraplc.com), which is currently pending approval. The success in inducing ovarian activity and ovulation in winter with dopamine antagonists has varied from experiment to experiment. The assumption
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that these compounds are producing their stimulatory effects via an increase in prolactin secretion is supported by the fact that administration of recombinant bovine prolactin itself stimulated ovarian activity in seasonally anovulatory mares,41 and daily administration of recombinant porcine prolactin to pony mares in winter resulted in ovulation (as determined by elevated progesterone levels) at an average of 22 days later.42 Moreover, Kelley et al.43 reported that estradiol pretreatment of seasonally anovulatory mares with estradiol benzoate greatly enhanced the prolactin response to sulpiride treatment, and resulted in eight of nine mares ovulating an average of 29 days after onset of treatment. Thus, lowdose estrogen pretreatment may be a useful means of increasing the success rate of dopamine antagonist treatments that would otherwise be borderline successful or unsuccessful.
Anestrus with no ovarian activity in the breeding season Nutritional anestrus Even though thin mares take longer (more breedings) to be rebred after foaling,44 and experience more profound and longer periods of seasonal anestrus,22 they eventually do return to estrus in spite of their thin condition (at body condition scores of 3 to 4). Whether thinner or emaciated mares display normal estrous cycles, can get pregnant, or can carry foals to term may be interesting biological questions; however, such information has little practical value for horse breeders. Lactational anestrus Although the majority of mares display a foal heat within 10 days after foaling, lactational anestrus is occasionally encountered, occurring in 3–4% of foaling mares of average or good body condition,45 and in up to 16% under less than ideal conditions.46 Mares in poor body condition at foaling will likely exhibit a higher incidence of lactational anestrus.44, 46 Mares foaling early in the year may experience a foal heat (or have a silent foal heat) and then enter a period of anestrus. Such mares will begin to have follicular activity and will eventually ovulate once the daylength gets to a normal stimulatory duration. This condition is likely a seasonal, rather than lactational, anestrus, and should be treated as such. Mares foaling during the normal ovulatory season and nursing a foal may exhibit a true lactational anestrus. The question is whether they are experiencing a lack of follicular activity and ovulation, or are just failing to display estrous behavior. Examination
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of the ovaries should be performed, and if the mare is found to have significant follicular activity, several potential solutions may be tried: 1. altrenogest feeding to normalize her estrous cycle and synchronize ovulation; 2. prostaglandin F2α administration if she is suspected of having luteal tissue (perhaps a prolonged corpus luteum); or 3. regular ultrasound evaluation without other treatment until she develops a 30-mm follicle or greater, after which hCG or deslorelin injection can be used to induce and help pinpoint ovulation. If the mare develops an ovulatory sized follicle but still does not display estrus, artificial insemination can be used to breed to the follicle at the appropriate time (an approach unfortunately not available for Thoroughbred breeders). Mares not having significant follicular activity on their ovaries present a unique problem. Very thin mares should respond to an appropriate increasing plane of nutrition, although it may take several weeks or more to increase body condition to a level sufficient for ovarian activity.44 For mares of adequate body condition, the cause may be a paucity or lack of hypothalamic input of GnRH to the pituitary gland (such as causes seasonal anestrus), and treatment with GnRH or one of its analogs is a possible solution. Williams et al.47 reported limited success (three of eight mares in 14 days; 80% or better overall) with osmotic minipumps loaded with GnRH in mares with lactational anestrus. Even with average body condition at foaling, metabolic demands of lactation may be a problem in some mares, thus an increase in nutrient intake may be helpful. These mares may also respond to treatment with equine FSH to stimulate follicular growth followed by hCG.48 GnRH implant-induced anestrus Following reports from the field that the commercially available deslorelin implant, Ovuplant (Dechra Veterinary Products; www.dechra-eu.com), was causing a delay in return to estrus in some mares, and complete anestrus in others, Johnson et al.49–51 did a series of experiments to better describe what was happening to the pituitary–gonadal axis in those mares. Although the implant, used to hasten ovulation, caused a temporary downregulation of pituitary LH and FSH secretion in all mares, all but a few recovered within 14 days to normal ovarian activity and gonadotropin secretion and responsiveness to GnRH.49, 50 Two mares in those studies had extended periods of anestrus, and one mare did not recycle until the following spring. In that mare, pituitary func-
tion appeared to recover after 14 to 21 days; however, rather than experiencing subsequent cycles in ovarian activity, her ovaries went inactive. A continual, gradual rise in LH and FSH concentrations in her blood indicated that she was in an ovariectomizedlike state as far as her pituitary function, indicating that the lesion was at the ovarian level. That is, her pituitary had recovered, but her ovaries had been altered and were not responsive to the gonadotropins in her blood (analogous to the situation in postmenopausal women). The mare was turned out to pasture until the next spring, at which time she was followed through one full estrous cycle to confirm that she was indeed recovered and cyclic. Administration of a single Ovuplant on the next estrus resulted in a repeat of her previous ovarian shutdown, which again lasted through the breeding season. It is concluded that a small percentage of mares are overly sensitive to continual high deslorelin stimulation, resulting in ovarian regression. Whether other potent GnRH analogs, or whether other types of release formulations (e.g., liquids rather than implants), might cause the same effect is unknown. However, it has been demonstrated that the effects can be ameliorated by removing implants after ovulation.52 Hypopituitarism Hypopituitarism is a naturally occurring lack of normal pituitary hormone production and secretion. Panhypopituitarism usually affects most or all of the pituitary hormones,53 thus abnormalities in growth and metabolism would be noticed first, and a failure to undergo puberty would likely result in external appearances somewhat different than the normal mare. Measurement of LH and FSH levels in the blood, before and after a standard GnRH injection (1 µg/kg bodyweight25 ), should confirm either normal or abnormal (no) gonadotropin secretion. A lack of response to GnRH would indicate a paucity of gonadotropin in the pituitary; the prognosis for such a mare is poor. Although GnRH therapy, similar to that used in seasonally anovulatory mares, might return pituitary function to normal, the possibility of an underlying genetic cause for the condition would preclude using the mare for breeding. Moreover, any defect in the pituitary that led to an absence of LH or FSH production would not lend itself to reversal by GnRH therapy. Direct stimulation of the ovaries with exogenous FSH and LH (or hCG) may indicate that the mare’s ovaries are normal, but again, the practicality of such therapy is questionable. GnRH vaccines A potential cause of anestrus in mares, similar to hypopituitarism but not naturally occurring, is the use
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Anestrus of GnRH vaccines, such as Equity, earlier in a mare’s lifetime.54 These vaccines are used to suppress the reproductive system of young mares, thereby avoiding behavioral problems sometimes associated with estrus.55 As mentioned previously, immunoneutralization of endogenous GnRH by active immunization against GnRH conjugated to a foreign protein resulted in undetectable LH concentrations in plasma of mares in the breeding season, and a reduction of FSH concentrations to about 50% of normal.15 Those immunized mares had no follicles on their ovaries greater than 10 mm, and were acyclic throughout the breeding season. Rabb et al.,17 using pony geldings as a model, showed that the pituitary LH content was reduced by 91% by active immunization against GnRH. Although the antibody titers needed to effectively immunoneutralize GnRH usually wane with time (typically 6 months to 1 year), it is possible that a small proportion of mares could retain sufficient titers over several years to keep them from cycling. These mares would have small, inactive ovaries with follicles no greater than about 10 mm in diameter. Diagnosis for this condition should be possible by comparing the LH and FSH responses to GnRH with those after injection of a GnRH agonist, such as deslorelin or historelin. That is, challenge with an agonist (e.g., 5 µg historelin i.v.), which should not cross-react with the anti-GnRH antibodies, would result in a very low or no LH response, but a normal or only slightly reduced FSH response. In comparison, injection of an equivalent dose of GnRH (250 µg i.v.) would result in little or no response in either gonadotropin, due to the immunoneutralization by the anti-GnRH antibodies. For mares testing positive for anti-GnRH antibodies, it may be possible to stimulate the pituitary gland and induce ovarian activity with frequent injections of GnRH agonist; however, even if a mare were induced to ovulate, the same therapy might also be needed to maintain CL function until endometrial cup formation; an alternative would be altrenogest therapy from ovulation to cup formation. Obviously, the practicality of such an approach would be low. Senescence Most mares display estrous cycles throughout their last years of life, and numerous studies have documented age-related decreases in fertility. Failure to experience cyclic follicular waves and ovulations in older mares has been less well documented. The decrease in follicular numbers and number of ovulations in aged versus young mares was well summarized by Carnevale56 and Vanderwall et al.57 Although the number of follicles decrease with age, depletion of follicles per se is not thought to occur in
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mares as it does in some other species. Whatever the cause of cessation of cyclicity, the prognosis would seem poor based on the age of the mare and the fact that embryos from aged mares, when obtained, tend to have a lower success rate when transferred to young recipients.56 Idiopathic Various undocumented causes of anestrus accompanied by small, inactive ovaries could include primary hypogonadism, in which a defect is localized in the ovaries themselves. In such cases, assuming no other complicating defects, concentrations of LH and FSH would be expected to be high, similar to those in ovariectomized mares. Again, the prognosis for such a mare would be poor. Various causes of primary hypogonadism in humans have been described, and often are the result of gene defects affecting important proteins such as hormonal receptors58 or key enzyme systems.59 Although interesting from a biological standpoint, treatment of such conditions would be impractical.
References 1. Hart PJ, Squires EL, Imel KJ, Nett TM. Seasonal variation in hypothalamic content of gonadotropin-releasing hormone (GnRH), pituitary receptors for GnRH, and pituitary content of luteinizing hormone and follicle-stimulating hormone in the mare. Biol Reprod 1984;30:1055–62. 2. Silvia PJ, Squires EL, Nett TM. Changes in the hypothalamic hypophyseal axis of mares associated with seasonal reproductive recrudescence. Biol Reprod 1986;35:897– 905. 3. Ginther OJ. 1992. Reproductive Biology of the Mare. 2nd ed. Cross Plains, WI: Equiservices 4. Ginther OJ, Bergfelt DR. Associations between FSH concentrations and major and minor follicular waves in pregnant mares. Theriogenology 1992;38:807–21. 5. Ginther OJ. Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains, WI: Equiservices, 1986. 6. Back DG, Pickett BW, Voss JL, Seidel GE Jr. Observations on the sexual behavior of nonlactating mares. J Am Vet Med Assoc 1974;165:717–20. 7. Oxender WD, Noden PA, Bolenbaugh DL, Hafs HD. Control of estrus with prostaglandin F2alpha in mares: Minimal effective dose and stage of estrous cycle. Am J Vet Res 1975;36:1145–7. 8. Nett TM, Holtan DW, Estergreen VL. Levels of LH, prolactin and oestrogens in the serum of post-partum mares. J Reprod Fertil Suppl 1975;23:201–6. 9. Sharp DC. Environmental influences on reproduction in horses. In: Hughes JP (ed.) Veterinary Clinics of North America: Large Animal Practice. Philadelphia: WB Saunders, 1980; pp. 207–73. 10. Thompson DL Jr, Mitcham PB, Runles ML, Burns PJ, Gilley RM. Prolactin and gonadotropin responses in geldings to injections of estradiol benzoate in oil, estradiol benzoate in biodegradable microspheres, and estradiol cypionate. J Equine Vet Sci 2008;28:232–7.
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11. Davis SD, Sharp DC. Intra-follicular and peripheral steroid characteristics during vernal transition in the pony mare. J Reprod Fertil Suppl 1991;44:333–40. 12. Bergfelt DR, Ginther OJ. Delayed follicular development and ovulation following inhibition of FSH with equine follicular fluid in the mare. Theriogenology 1985;24:99– 108. 13. Ginther OJ, Bergfelt DR. Growth of small follicles and concentrations of FSH during the equine oestrous cycle. J Reprod Fertil 1993;99:105–11. 14. Irvine CHG. Endocrinology of the estrous cycle of the mare: Applications to embryo transfer. Theriogenology 1981; 15:85–104. 15. Garza F Jr, Thompson DL Jr, French DD, Wiest JJ, St George RL, Ashley KB, Jones LS, Mitchell PS, McNeill DR. Active immunization of intact mares against gonadotropinreleasing hormone: Differential effects on secretion of luteinizing hormone and follicle-stimulating hormone. Biol Reprod 1986;35:347–52. 16. Garza F Jr, Thompson DL Jr, Mitchell PS, Wiest JJ. Effects of active immunization against gonadotropin releasing hormone on gonadotropin secretion after ovariectomy and testosterone propionate administration to mares. J Anim Sci 1988;66:479–86. 17. Rabb MH, Thompson DL Jr, Barry BE, Colborn DR, Hehnke KE, Garza F Jr. Effects of active immunization against GnRH on LH, FSH and prolactin storage, secretion and response to secretagogue in pony geldings. J Anim Sci 1990;68:3322–9. 18. Henneke DR, Potter GD, Kreider JL, Yeates BF. Relationship between condition score, physical measurements and body fat percentage in mares. Equine Vet J 1983;15:371–2. 19. Gentry LR, Thompson DL Jr, Gentry GT Jr, Davis KA, Godke RA, Cartmill JA. The relationship between body condition, leptin, and reproductive and hormonal characteristics of mares during the seasonal anovulatory period. J Anim Sci 2002;80:2695–703. 20. Fitzgerald BP, CJ McManus. Photoperiodic versus metabolic signals as determinants of seasonal anestrus in the mare. Biol Reprod 2000;63:335–40. 21. Fitzgerald BP, Reedy SE, Sessions DR, Powell DM, McManus CJ. Potential signals mediating the maintenance of reproductive activity during the non-breeding season of the mare. J Reprod Fertil Suppl 2002; 59:115–29. 22. Gentry LR, Thompson DL Jr, Gentry GT Jr, Davis KA, Godke RA. High versus low body condition in mares: Interactions with responses to somatotropin, GnRH analog, and dexamethasone. J Anim Sci 2002;80:3277– 85. 23. Thompson DL Jr, Johnson L, St George RL, Garza F Jr. Concentrations of prolactin, luteinizing hormone and follicle stimulating hormone in pituitary and serum of horses: Effect of sex, season and reproductive state. J Anim Sci 1986;63:854–60. 24. Thompson DL Jr, McNeill DR, Wiest JJ, St George RL, Jones LS, Garza F Jr. Secretion of luteinizing hormone and follicle stimulating hormone in intact and ovariectomized mares in summer and winter. J Anim Sci 1987;64:247–53. 25. Foster JP, Evans MJ, Irvine CHG. Differential release of LH and FSH in cyclic mares in response to synthetic GnRH. J Reprod Fertil 1979;56:567–72. 26. Gentry LR, Thompson DL Jr, Stelzer AM. Treatment of seasonally anovulatory mares with thyrotropin releasing hormone and(or) gonadotropin releasing hormone analog. J Anim Sci 2002;80:208–13.
27. Grubaugh W, Sharp DC, Berglund LA, McDowell KJ, Kilmer DM, Peck LS, Seamans KW. Effects of pinealectomy in pony mares. J Reprod Fertil Suppl 1982;32: 293–5. 28. Kilmer DM, Sharp DC, Berglund LA, Grubaugh W, McDowell KJ, Peck LS. Melatonin rhythms in Pony mares and foals. J Reprod Fertil Suppl 1982;32:303–7. 29. Hastings MH, Herbert J, Martensz ND, Roberts AC. Annual reproductive rhythms in mammals: Mechanisms of light synchronization. Ann N Y Acad Sci 1985;453:182– 204. 30. Johnson AL. Gonadotropin-releasing hormone treatment induces follicular growth and ovulation in seasonally anestrous mares. Biol Reprod 1987;36:1199–206. 31. Burkhardt J. Transition from anoestrus in the mare and the effects of artificial lighting. J Agric Sci 1947;37:64–8. 32. Kooistra LH, Ginther OJ. Effect of photoperiod on reproductive activity and hair in mares. Am J Vet Res 1975;36: 1413–19. 33. Oxender WD, Noden PA, Hafs HD. Estrus, ovulation, and serum progesterone, estradiol, and LH concentrations in mares after an increased photoperiod during winter. Am J Vet Res 1977;38:203–7. 34. Webel SK, Squires EL. Control of the oestrous cycle in mares with altrenogest. J Reprod Fertil Suppl 1982;32:193–8. 35. Allen WR, Sanderson MW, Greenwood RES, Ellis DR, Crowhurst JS, Simpson DJ, Rossdale PD. Induction of ovulation in anoestrous mares with a slow-release implant of a GnRH analogue (ICI 118 630). J Reprod Fertil Suppl 1987;35:469–78. 36. Harrison LA, Squires EL, Nett TM, McKinnon AO. Use of gonadotropin-releasing hormone for hastening ovulation in transitional mares. J Anim Sci 1990;68:690–9. 37. Turner JE, Irvine CHG. The effect of various gonadotrophin-releasing hormone regimens on gonadotrophins, follicular growth and ovulation in deeply anoestrous mares. J Reprod Fertil Suppl 1991;44:213–25. 38. Besognet B, Hansen BS, Daels PF. Induction of reproductive function in anestrous mares using a dopamine antagonist. Theriogenology 1997;47:467–80. 39. Donadeu FX, Thompson DL Jr. Administration of sulpiride to anovulatory mares in winter: Effects on prolactin and gonadotropin concentrations, ovarian activity, ovulation and hair shedding. Theriogenology 2002;57:963–76. 40. Brendemuehl JP, Cross DL. Influence of the dopamine antagonist domperidone on the vernal transition in seasonally anoestrous mares. J Reprod Fertil Suppl 2000;56:185– 93. 41. Nequin LG, King SS, Johnson AL, Gow GM, Ferreira-Dias GM. Prolactin may play a role in stimulating the equine ovary during the spring transition. J Equine Vet Sci 1993; 13:631–5. 42. Thompson DL Jr, Hoffman R, DePew CL. Prolactin administration to seasonally anestrous mares: Reproductive, metabolic, and hair shedding responses. J Anim Sci 1997; 75:1092–9. 43. Kelley KK, Thompson DL Jr, Storer WA, Mitcham PB, Gilley RM, Burns PJ. Estradiol interactions with dopamine antagonists in mares: Prolactin secretion and reproductive traits. J Equine Vet Sci 2006;26:517–28. 44. Henneke DR, Potter GD, Kreider JL. Body condition during pregnancy and lactation and reproductive efficiency of mares. Theriogenology 1984;21:897–909. 45. Heidler B, Aurich JE, Pohl W, Aurich C. Body weight of mares and foals, estrous cycles and plasma glucose
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concentration in lactating and non-lactating Lipizzaner mares. Theriogenology 2004;61:883–93. Nagy P, Huszenicza G, Juh´asz J, Kulcs´ar M, Solti L, Reiczigel J, Abav´ary K. Factors influencing ovarian activity and sexual behavior of postpartum mares under farm conditions. Theriogenology 1998;50:1109–19. Williams GL, Amstalden M, Garcia M R, Williams SW, Ward J, Blodgett GP, Quirk KS, Unnerstall DA. Control of lactational and idiopathic anovulation in mares with low-dose, native GnRH: Anterior pituitary and ovarian responses to continuous, subcutaneous infusion. Biol Reprod 2000;62 (Suppl. 1):302 McCue PM, Squires EL, LeBlanc MM. eFSH in clinical equine practice. Theriogenology 2007;68:429–33. Johnson CA, Thompson DL Jr, Kulinski KM, Guitreau AM. Prolonged interovulatory interval and hormonal changes in mares following the use of Ovuplant to hasten ovulation. J Equine Vet Sci 2000;20:331–6. Johnson CA, Thompson DL Jr, Cartmill JA. Pituitary responsiveness to GnRH in mares following deslorelin acetate implantation to hasten ovulation. J Anim Sci 2002; 80:2681–7. Johnson CA, Thompson DL Jr, Cartmill JA. Effects of deslorelin acetate implantation in horses: Single implants in stallions and steroid-treated geldings and multiple implants in mares. J Anim Sci 2003;81:1300–7.
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52. McCue PM, Farquhar VJ, Carnevale EM, Squires EL. Removal of deslorelin (Ovuplant) implant 48 h after administration results in normal interovulatory intervals in mares. Theriogenology 2002;58:865–70. 53. Guyton AC, Hall JE. Textbook of Medical Physiology, 9th edn. Philadelphia: W.B. Saunders, 2000. 54. Robinson SJ, McKinnon AO. Prolonged ovarian inactivity in broodmares temporally associated with administration of Equity. Aust Equine Vet 2006;25:85–7. 55. Elhay M, Newbold A, Britton A, Turley P, Dowsett K, Walker J. Suppression of behavioural and physiological oestrus in the mare by vaccination against GnRH. Aust Vet J 2007;85:39–45. 56. Carnevale EM. Reproductive disorders. In: Bertone JJ (ed.) Equine Geriatric Medicine and Surgery. New York: Elsevier Health Sciences, 2005; pp. 194–200. 57. Vanderwall DK, Woods GL, Freeman DA, Weber JA, Rock RW, Tester DF. Ovarian follicles, ovulations and progesterone concentrations in aged versus young mares. Theriogenology 1993;40:21–32. 58. Latronico AC, Arnhold IJ. Inactivating mutations of LH and FSH receptors-from genotype to phenotype. Pediatr Endocrinol Rev 2006;4:28–31. 59. Timmreck LS, Reindollar RH. Contemporary issues in primary amenorrhea. Obstet Gynecol Clin N Am 2003;30: 287–302.
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Chapter 178
Vernal Transition into the Breeding Season Daniel C. Sharp
Introduction and mechanisms of seasonality Because our world is tilted on its vertical axis, the annual journey around the sun results in differential heating or cooling of the earth. Mares that foaled during the cold time of year, when food sources were scarce, were in jeopardy of not passing on their genes. Therefore, through the random process of evolutionary selection, mechanisms were developed to assure the breeding season was timed so as to minimize the chances of foaling during the winter. Thus, the annual reproductive rhythm of horses was established. An important concept of seasonality is that the mechanisms were developed to assure the time of foaling, not necessarily the time of breeding. This means regulation of breeding to a time of year that precedes the foaling season by approximately one gestation length. This is a clue to the concept that seasonality involves precise anticipation of environmental factors a gestation length later. The environmental factor that provides that much precision is daylength. In temperate zones where horses evolved, important environmental cues are provided by the changing daylength from the winter solstice to the summer solstice and back to the next winter solstice. Exactly what the nature of those cues is remains a mystery. Arguments have been proposed for absolute length of day (hours of daylight versus hours of darkness), rate of change of daylength, change of direction of daylength (i.e., decreasing then increasing daylength), and photic history as the primary cues for regulating the annual reproductive rhythm. However, definitive data remain elusive. For instance, it has been speculated that the ‘spontaneous’ return to estrous cyclicity in the spring reflects a response to increasing daylength directly and it has also been
proposed that it reflects refractoriness to the inhibitory effects of short day. The concept of photorefractoriness has been well researched in other photoresponsive species, such as sheep and birds,1, 2 but there is little formal research on the potential role of photorefractoriness in mares. Photorefractoriness is defined as the failure of a photic signal to maintain a given state. In terms of seasonality in horses, it would refer to the inability of short winter day photoperiods to maintain a sexually inactive state, and, thus, mares may ‘escape’ the negative stimulus of short winter photoperiod. Of interest, photorefractoriness in birds has been equated to the loss of hypothalamic GnRH,2 a condition of anestrus in mares as well. On the other hand, there is good evidence for the existence, in sheep, of a mechanism in the pars tuberalis for a circannual rhythm in photoresponsiveness.1, 3 While a detailed description of such a mechanism is beyond the scope of this chapter, it may be summarized as the existence of Clock genes (see Chapter 183) in the pars tuberalis of the pituitary, that owe their temporal expression to photic changes through the pineal hormone, melatonin. This is an area of interest to equine practitioners and breeders alike, as it promises to provide a clearer understanding of the impact of photoperiod on the annual reproductive rhythm of mares. While the existence of Clock genes in the pars tuberalis of horses remains to be demonstrated, there is clear evidence that melatonin receptors are found there.4 The fact that melatonin binding occurs in the pars tuberalis of the pituitary strongly suggests that temporal control of seasonal reproduction may be regulated there in horses as well. At least, it is strong incentive for targeted research in that area. Furthermore, there is a possibility that our understanding of the regulatory factors behind seasonality may be clouded by the existence of dual mechanisms.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Vernal Transition into the Breeding Season Naturally occurring reproductive recrudescence in the springtime may be effected by one mechanism involving photic history, as described above, whereas the stimulation of early recrudescence by enhanced environmental lighting may act through somewhat different mechanisms. This apparent dichotomy will become clearer as the events of vernal transition are discussed below, and questions are raised as to how some early vernal events could reflect increased daylength when they occur so early after the winter solstice that changes in daylength are minimal. Again no definitive answer is currently available. The following discussion will address the events of vernal transition to formulate a working hypothesis of how reproductive recrudescence is effected, and to offer suggestions for potential management of vernal transition in a broodmare operation.
Phases of the annual reproductive rhythm The annual reproductive rhythm can be partitioned, for discussion, into four phases or stages, each with functional characteristics.
Breeding season proper The breeding season is defined as the time of year characterized by regularly recurring estrous cycles. The start date of the breeding season is defined as the first ovulation of the year. This definition also serves as the end date of vernal transition. The beginning of the breeding season proper (first ovulation of the year) is remarkably precise among mares in a herd from year to year, with average date of the first ovulation in horse mares of April 7 ± 9.1 days and May 7 ± 21.1 in pony mares in the northern hemisphere.5 The difference in start dates between horse mares and pony mares appears to reflect breed type differences, but the underlying principles are likely similar. It is important to keep in mind that the above-mentioned dates reflect the average over several years, with markedly differing environmental conditions of temperature, rainfall, and pasture growth. Despite these differences, the date of the first ovulation of the year remains remarkably stable, providing strong support for the role of a highly consistent environmental zeitgeber or time-giver. Only photoperiod provides that kind of precision. An equally important consideration is the observation that the end of the breeding season, defined as the last ovulation of the year, does not display such precision among mares, with a range of dates extending from September to December in the northern hemisphere. To have such precision at the begin-
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ning of an event and extreme imprecision regulating the end of the event may seem strange at first consideration. However, it is quite reasonable to speculate that evolutionary pressure to regulate the first opportunities to become pregnant might have been high, whereas under natural conditions in the wild, mares that were bred late, or were not bred at all, may have dropped out of the evolutionary gene pool. Thus, selection for a precise end of the breeding season may not have been effective. Little further discussion of the breeding season is required here, as it will be well covered in other chapters of this book. Autumnal transition As indicated above, the autumnal transition is highly variable among mares, and there is speculation about various ways reproduction fails at this time. Studies of the mechanisms of autumnal transition are difficult because of the wide range of dates of occurrence. This makes studies extremely time-consuming, and time-confounded, as environmental conditions change drastically over the duration of transition. Furthermore, autumnal transition does not enjoy the commercial importance of vernal transition, hence fewer researchers become interested in understanding the mechanisms. Mares entering this transition are commonly seen to appear to exhibit estrus, albeit with reduced vigor, develop a large preovulatory-like follicle, suggesting adequate follicle-stimulating hormone (FSH) concentrations, but fail to ovulate, suggesting reduced luteinizing hormone (LH).6 Another pattern has also been detected wherein the mare ovulates and fails to develop further follicles and a persistent corpus luteum is ultrasonically detected, despite low progesterone levels for an extended period of time. Both patterns probably reflect declining gonadotropinreleasing hormone (GnRH) during the autumn, associated with the failure of LH secretion.7 It has been shown that hypothalamic GnRH is reduced in the winter,8, 9 but the course of decline in the fall has not been studied. Anestrus Anestrus is defined as the time of the annual reproductive rhythm during which reproduction comes to a complete halt. The failure of LH secretion during autumnal transition is manifested in anestrus as a marked depletion of LH from the pituitary10 and, indeed, loss of the ability to synthesize LH subunits reflected in inability to detect the messenger ribonucleic acid (mRNA) for the subunits alpha and LH beta in pituitaries during anestrus.11 GnRH content in the
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hypothalamus is diminished during anestrus as well as its regional distribution within the hypothalamus.8 Additionally, GnRH secretion, as measured by pushpull perfusion12 or by cavernous sinus cannulation,13 is diminished severely during anestrus. Pituitary content of FSH does not appear to change throughout the year and remains high during anestrus;10 however, there are no reports of which this author is aware of studies of pituitary FSH beta mRNA at this time. Importantly, several reports have indicated that FSH secretion into the peripheral circulation is also markedly diminished during anestrus.6, 14 Therefore, these data suggest a dual mechanism of gonadotropin control in which the reduced secretion of LH reflects the diminished synthetic capacity of the pituitary, but also reflects the relative lack of GnRH. The reduced secretion of FSH, on the other hand, likely reflects only the lack of GnRH stimulation. The importance of this will become evident when the events of vernal transition are discussed below. Vernal transition Vernal transition is characterized by a series of events, likely obligatory, that serve to reverse the wintertime decline in reproductive capacity. This phase of the annual reproductive rhythm, more than any of the others, is a source of reproductive inefficiency leading to commercial anxiety. Part of the problem lies in the fact that mares exhibit ambiguous signs in vernal transition that are easy to misinterpret, such as large, but anovulatory, follicles and estrous behavior. Because of the commercial emphasis on early-born foals, breeders may be tempted to rush the transition phase and breed mares before they actually enter the breeding season proper. At best, this activity is futile, resulting in a low conception rate or a high rate of early embryonic death associated with inadequate corpus luteum maintenance.15 At worst, inappropriate breeding during the vernal transition can lead to uterine contamination that requires extensive treatment. Timing of vernal transition is also the most precise phase of the annual reproductive rhythm. As stated above, the signal to begin the process of sexual recrudescence is unknown, but the process apparently begins soon after the shortest day of the year, December 21 in the northern hemisphere. These will be outlined below in the discussion of events of vernal transition.
Gonadotropin-releasing hormone In one study, GnRH was measured in pituitary venous effluent via cavernous sinus cannulation and it
was reported that GnRH was elevated by 2 weeks after the winter solstice.16 Similar results were reported by Sharp and Grubaugh,12 who measured GnRH secretion via push-pull perfusion in anestrous mares at weekly intervals after exposure to artificially increased photoperiod. These researchers observed that GnRH was markedly elevated in mares exposed to artificially lengthened day for only 2 weeks (see also Chapter 174). These data suggest that the first event of vernal transition is increased GnRH. The next set of events, increased FSH secretion and follicular development, support that notion.
Follicle-stimulating hormone In apparent response to renewed GnRH secretion, FSH begins to be secreted early in January,6, 14, 17 and remains highly variable throughout the vernal transition. In a study of the relationship between FSH secretion, follicle formation and inhibin secretion during vernal transition, Watson et al.18 reported that the emergence of each follicular wave was preceded by an increase in FSH secretion. This agrees well with the report of Donadeu and Ginther19 who suggested that the temporal relationship between FSH secretion and follicle waves during vernal transition reflected a causal relationship. Watson et al.18 also stated their belief that inhibins ‘contributed to negative feedback on the release of FSH from the pituitary gland both during the transition period and the breeding season in mares.’ Watson et al.18 also reported the observation that vernal transition was characterized by an average of 3.7 follicular waves; each wave reflecting a large follicle (> 30 mm), with approximately 10 days intervening between successive waves. These data are remarkably similar to the report of Davis et al.20 who found an average of 3.7 ± 0.9 consecutive large follicles (> 30 mm) with an interval between first detection of each new follicle ≥ 30 mm of approximately 12 days. Thus, it seems clear that the follicular response to renewed FSH secretion after the shortest day of the year is the first readily observed indication of sexual recrudescence in mares. With an average of 3.7 successive follicle waves, at an interval of approximately 12 days each, it is also apparent that progress toward sexual competence in the vernal transition is slow indeed. The arithmetic indicates that approximately 45 days is taken up with these successive large but anovulatory follicles before the first ovulation of the year occurs. By definition, that first ovulation of the year signals the end of vernal transition and the beginning of the breeding season proper. The subject of vernal transition follicles deserves much more discussion. For one thing, the occurrence of these large follicles, often coupled with estrous display, lures many a breeder to send mares to the
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Vernal Transition into the Breeding Season breeding shed prematurely. As previously stated, this can produce unfavorable consequences, especially if the mare has a tendency to develop post-breeding endometritis. Premature breeding during vernal transition in an attempt to save a few weeks can lead to multiple more problems and expenses such as inappropriate use of stallions and staff labor. In addition, if mares are treated with agents that force the induction of ovulation too early during the vernal transition, then although ovulation occurs and pregnancy may ensure, the corpus luteum is not maintained in all mares and a high rate of embryonic death has been reported.15
Follicle steroidogenesis during vernal transition Perhaps the first suggestion that vernal transition follicles were not normal steroidogenically came from Seamans and Sharp21 who reported on the changes in follicular aromatase activity throughout vernal transition and showed that estradiol concentrations in the peripheral plasma remained low throughout most of vernal transition, rising only a few days prior to the first ovulation of the year. They also showed that follicular tissues (granulosa and theca) were capable of converting either progesterone or androstenedione to estradiol and/or estrone, thus indicating that the lack of estrogen production by early transition follicles reflects a block or a deficiency earlier in the steroidogenic pathway. Further evidence demonstrating inability of early transition follicles to synthesize androgens and estrogens was offered by Davis and Sharp.22 In this study, follicular dynamics were monitored to detect the first, second, third, or fourth consecutive follicle with a diameter ≥ 30 mm (transition follicle), or the first ovulatory follicle of the breeding season. Twenty-four hours after a large follicle was detected, theca (T) and granulosa (G) tissues were harvested. Separate and co-incubations of these tissues were conducted to determine steroid production in early transition (first or second consecutive vernal transition follicle; ET), late transition (third consecutive vernal transition follicle; LT), and preovulatory (OV) follicles. Peripheral plasma and follicular fluid steroids and gonadotropins also were assayed. Circulating and follicular fluid estradiol concentrations were significantly lower at the time of the first and second consecutive ≥ 30 mm follicle, compared with the third consecutive ≥ 30 mm follicle or with the OV follicle. Androstenedione and estradiol production in vitro did not increase until the time of the first ovulation of the year. Testosterone increased somewhat, but not significantly until the time of the post-luteal follicle, whereas
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progesterone production showed no change. These authors suggested that the early transition follicles lacked the 17-alpha steroid hydroxylase (17-OH) enzyme. This observation was later supported by Tucker et al.23 who evaluated the abundance of steroidogenic enzymes in follicular tissue obtained from the first, second, third, or OV follicles of vernal transition. These authors indicated that 17-OH was indeed undetected in first or second vernal transition follicles, but was highly abundant in the third vernal transition follicle of the year. These results were further supported by the report of Watson et al.24 who used in situ hybridization to evaluate steroid acute regulatory protein (StAR), side chain cleavage enzyme, (P450scc; CYP11A1), 3-beta hydroxysteroid dehydrogenase (3β-HSD), 17-alpha hydroxylase, 1720 lyase (CYP17), and aromatase (P450arom (CYP19). Although they did not discriminate among first, second, third, or ovulatory follicles, their results support the notion that steroid-forming enzymes, especially side chain cleavage and 17-OH, are minimally abundant in the first-developed follicles of vernal transition, and therefore steroidogenesis by these follicles is incompetent. In addition to the steroidogenic incompetence of vernal transition follicles, other characteristics mark them as abnormal. Although early vernal transition follicles achieve diameters of 30 mm or larger, the rate at which they grow is significantly slower.23 This observation can be of practical importance when faced with an incomplete history on a mare in the springtime. If the opportunity arises, repeat examinations over a few days may be revealing; follicles in early transition (first and second follicle) grow at a much reduced daily rate (2 mm/day) compared with follicles in late transition or during the breeding season (6 to 8 mm/day).23 Follicles obtained from the early transition period were less richly vascularized than later follicles,23 which agrees with the report of Channing25 and Kenney et al.26 that more vascular follicles were shown to have higher steroid levels. Additionally, Tucker et al.23 demonstrated that the number of mural granulosa cells is significantly lower in first and second follicles compared with third or OV follicles. Apparently these abnormal vernal transition follicles reflect regulatory events that result in their growth, but not their functional development. Moreover, it is probably instructive to think of these follicles as merely a passive by-product of ongoing changes in the hypothalamic–pituitary axis, rather than to regard them as vital events in the reestablishment of reproductive competence. A recurring observation during vernal transition is that mares in early-to-late transition, when their vernal transition follicles are not yet capable of significant estrogen production, often display strong
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behavioral signs of estrus, either sporadically or continuously. While this apparent dichotomy is difficult to reconcile, an intriguing observation from Seamans and Sharp could offer a possible explanation. Seamans and Sharp21 used radiolabeled steroids in their tissue incubation studies and assessed the fate of radiolabel following all their incubations with recrystallization and high-performance liquid chromatography (HPLC). They reported that a reasonably high percentage (as high as 15%) of radiolabel remained in the water-soluble phase after incubation, suggesting some conversion of free steroid to conjugated or more highly polar steroids. These fractions were not characterized. However, Younglai and Short27 reported 6α-hydroxyestradiol in follicular fluid of mares, indicating the potential presence of more polar steroid compounds. Therefore, it is reasonable to hypothesize that a small amount of steroidogenesis might occur during early vernal transition, but without detection through analysis of free steroids. Furthermore, such postulated steroid would be expected to have little effect on the pituitary, as Baldwin et al.28 have shown that equine pituitary cells in culture do not respond to sulfated forms of estrogen whereas they do respond to estradiol or estrone. Whether production of small amounts of conjugated steroids could account for the behavioral changes in mares remains to be tested.
Luteinizing hormone The question that naturally arises is: what is the mechanism behind development of the abnormal vernal transition follicles? So far the changes in FSH after the winter solstice have been discussed, but little has been said about LH. As mentioned above, pituitary LH content becomes depleted during anestrus10 and that appears to be because the mRNA encoding LH subunit formation is virtually undetectable in the pituitary.11 Throughout much of the early and midvernal transition, LH secretion remains low14, 29 as pituitary LH content10 and LH subunit mRNA11, 30 remain low until some time in April. Therefore, the development of follicles in early and mid-vernal transition takes place in an environment that is FSH-rich, but LH-poor. It seems clear that such an environment does not favor normal follicle development or function. The follicles that develop in early pregnancy, prior to production of equine chorionic gonadotropin (eCG), may be good models with which to study vernal transition follicles, as the former also exist in an environment rich in FSH and poor in LH. It is true that little estrogen production can be detected during early pregnancy,6, 31 and this likely reflects the relative lack of LH. Further evidence of this lack was revealed by Albrecht et al.32
who demonstrated that with secretion of eCG, estrogen secretion was detected, in the corpora lutea of pregnant mares. These authors further demonstrated that transcription of the 17-OH enzyme gene was greatly increased after onset of eCG secretion. Thus, the hallmark of vernal transition is the continuing failure of LH secretion, leading to repeated development of large, anovulatory follicles that are steroidogenically incompetent. Hence, the crux of sexual recrudescence is the re-establishment of the gene(s) for LH subunit production. The succession of anovulatory, large (> 30 mm) follicles ends with the first ovulation of the year, and that event is preceded by only a few days by a pronounced increase in intrafollicular and peripheral plasma estrogen and by plasma LH. The temporal association between secretion of estrogen by vernal transitional follicles and secretion of LH is intriguing, and naturally leads to the speculation that the two events are causally related. Indeed, it has been shown clearly that administration of estrogen to ovariectomized mares at an equivalent time of early vernal transition can increase LH secretion.17 Furthermore, administration of estrogen for 6 to 9 days to ovariectomized mares at an equivalent time of early vernal transition has also been shown to increase pituitary mRNA encoding both alpha and LH/CG beta subunit expression.30 Thus, it is tempting to suggest that estrogen from the first steroidogenically competent follicle of the year is a signal for renewal of pituitary LH synthesis and secretion. However, caution is urged at that speculation for several reasons. First, estrogen and LH rise essentially simultaneously, raising the question of whether either one is a necessary stimulus for the other. Second, and most damaging to the above speculation, it is clear that estrogen is not necessary for re-establishment of LH synthesis and secretion as LH has been shown by several authors to rise spontaneously in the springtime in ovariectomized mares.6, 14, 17, 33 In the report of Peltier et al.33 groups of intact and ovariectomized mares were studied from the fall through the following summer, and their LH and estradiol secretory patterns monitored closely. They reported that the timing of the rise in circulating LH in intact and in ovariectomized mares was not significantly different. Blood samples for LH were collected daily and the timing of vernal transition was estimated by fitting a third-order regression equation to the data. The vernal transition was considered to be ended when the first order derivative of the regression equation increased above 4 ng/mL. The inflection point of the LH rise occurred at essentially the same time for intact mares and for ovariectomized mares. These data indicate that estrogen may not play a critical role in the timing of LH synthesis and secretion
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Vernal Transition into the Breeding Season Table 178.1 Critical events of vernal transition in the northern hemisphere Time
Events
Shortly after winter solstice
Increased hypothalamic GnRH
Early January
Increased FSH secretion
Late January–early February
Onset of follicular development
Throughout early springtime
Approximately 3.7 sequential anovulatory follicles @ 10- to 12-day intervals
Throughout early springtime
Failure of LH secretion
Throughout early springtime
Failure of follicular steroidogenesis
Approximately 5 days before first ovulation of the year
Follicular steroidogenesis begins, characterized by surge in estradiol
Approximately 5 days before first ovulation of the year
LH secretion begins, characterized by increased mRNA in pituitary and ovulatory surge of LH in periphery
Termination of transition
First ovulation of the year
FSH, follicle-stimulating hormone; GnRH, gonadotropin-releasing hormone, LH, luteinizing hormone.
renewal, although Peltier et al.33 reported that LH secretion rose faster in ovariectomized mares than it did in intact mares. The latter point suggests that ovarian estrogens might instead moderate LH secretion. Overall, the temporal agreement in initiation of LH secretion in vernal transition between ovarian intact and ovariectomized mares suggests that factors other than ovarian input, possibly a direct environmental effect or an indirect, long-term timing effect, is responsible for the initiation of LH synthesis and secretion in the springtime. The events of vernal transition are listed in Table 178.1. The impact of the foregoing is that the crux of the problem of transition from anestrus into the breeding season is the initiation of LH synthesis and secretion. Until we learn precisely what mechanisms control this process, attempts to manage entry into the breeding season will likely be met with variable success.
Manipulation of vernal transition The holy grail of hormonal reproductive manipulation in horses is the ability to regulate the timing of the first ovulation of the year. To date, most attempts have gained little utility, indicating that we have yet to understand completely the mechanisms underlying the transition from reproductive incompetence to reproductive capability. The following discussion includes several attempts to regulate the timing of first
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ovulation of the year, but the reader should be advised that manipulation of this event by artificial photoperiod remains the most successful and universal way to do so. A caveat of using artificial photoperiod is that it does not shorten or truncate the process of vernal transition; it simply initiates the process earlier. Therefore, adequate time to complete the process must be allowed in order to benefit from this system (see Chapter 183). GnRH administration As GnRH hypothalamic content and secretory rate are much reduced during winter anestrus, and as an early event in the beginning of vernal transition is the re-initiation of GnRH secretion, it seems reasonable to attempt to hasten the time of first ovulation of the year by administering GnRH to mares in the springtime. Several such attempts have been made34–38 and success reported, but the lack of prominence of GnRH treatment during vernal transition clinically currently attests to the inefficacy of this approach. Method and timing of administration of GnRH, as well as structure of the molecule have been varied, further confusing the issue. Hyland et al.39 utilized miniosmotic pumps to deliver GnRH at 0, 50, or 100 ng/kg body weight/h for 28 days to mares in deep seasonal anestrus. These authors reported that 0/10, 3/10, and 7/10 mares ovulated, respectively. These authors also reported treating mares in shallow anestrus similarly, with ovulation occurring at 25 days after initiation of treatment in GnRH-treated mares, and 42 days in control mares. The recognition that there might be differences between mares identified as being in ‘deep’ anestrus or ‘shallow’ anestrus points to one of the problems interpreting such experimental protocols. As there appears to be a continuum of changing events throughout vernal transition, it is imperative that experimental designs take into account the stage or degree of transition. It is highly recommended that mares assigned to experimental treatments be evaluated for presence of follicular development and/or corpora lutea over a 30-day period prior to beginning such an experiment. The presence of corpora lutea would immediately disqualify a mare from such experiments, as it is obvious that she would be cyclic. Presence of follicles greater than, say, 20 mm might also indicate that the process of FSH-induced follicular development has already begun. The reports of McKinnon et al.37, 38 support the notion that GnRH or its analogs (deslorelin in the latter reports) result in advanced date of first ovulation of the year when treatment assignment is based on follicular development and other physiological signs (i.e., endometrial edema) indicating that the mare is well into the vernal transition.
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Figure 178.1 Ultrasonogram of ovaries of a mare administered control vehicle throughout the vernal transition. The left ovary exhibits no follicles > 6 mm. (+ = calipers).
Figure 178.2 Ultrasonogram of ovaries of a mare administered des-gly throughout the vernal transition. The left ovary exhibits a follicle = 41 mm. (+ = calipers)
Most of the above reports used native GnRH (GnRH-A; amino acid sequence gln-his-ser-tyr-glyleu-arg-pro-gly), which has a relatively short half-life. Interest in administration of longer-acting analogs prompted other experimental designs, as in the reports of McKinnon et al.37, 38 In these latter reports, deslorelin, a GnRH analog with tryptophan substituted for glycine at the sixth amino acid position, was used. It would be useful to conduct studies of the binding kinetics of native GnRH and various analogs to compare their impact on pituitary function. For instance, it has been shown that continuous administration of native GnRH does not result in extinction of LH secretion (i.e., downregulation of receptors),40 whereas continuous administration of deslorelin, for instance, does.41 Davis et al.42 utilized several different GnRH analogs in either once-daily or twice-daily administration protocols. Native GnRH (GnRH-A) and d-Ala6 , and des-Gly10 respectively were administered twice daily (GnRH-A and d-Ala6 ) or once daily (des-Gly10 ) to mares in deep anestrus (experiment started January 20). LH and FSH were measured in blood samples obtained daily, and follicular development was monitored daily via ultrasound. FSH secretion and follicular development were increased in all three groups, but LH secretion was not changed. Consequently treated mares underwent development of large, anovulatory follicles for a prolonged period without ovulating (see Figures 178.1 and 178.2). Mares that received d-Ala6 for instance exhibited an average of 10 consecutive large anovulatory follicles (mean interval 12 days) prior to development of the first ovulatory follicle of the year. These results indicate the problem of vernal transition; that, although FSH appears to be available for release from the pituitary, LH is not necessarily available. Sherman et al.11 reported that, using Northern Blot analysis of equine pituitary total RNA, the mRNA encoding the LH subunits was undetectable during the winter months. Thus, in retrospect, all attempts to
stimulate an early onset of the breeding season with GnRH are likely doomed to failure, unless the status of pituitary LH subunit message is known. In fact, the crux of the problem of resumption of full reproductive activity appears to be the re-establishment of the LH gene. Estrogen As previously stated, the first ovulatory follicle of the year becomes steroidogenically competent just before ovulation. This has prompted the suggestion that the preovulatory surge of estradiol may be important in the final re-establishment of the LH gene, and therefore might suggest a clinically practical way to surmount the problem of LH unavailability. Mumford et al.35 used estrogen alone or in combination with GnRH in attempts to stimulate releasable LH in mares in early transition. These authors reported no effect of estrogen alone, but there was an increase in LH response to either GnRH or GnRH in combination with estrogen. However, they concluded that estradiol promoted neither the folliculogenesis nor the increased circulating LH required for ovulation. One interpretation of these results is that the acute administration of estrogen and/or GnRH may have caused the release of a small amount of releasable LH in the pituitary, but neither treatment was sufficient to stimulate expression of the mRNA encoding the LH subunits for sustained LH production. Sharp et al.17 reported that estrogen administration to ovariectomized mares at an equivalent time of early in transition (start date, March 10) resulted in increased circulating LH, increased LH responsiveness to exogenous GnRH administration challenge, and increased pituitary LH content (see Figure 178.3). FSH secretion tended to decrease, the FSH response to exogenous GnRH was greatly reduced, but pituitary FSH content did not appear to change.17 These results indicate that estrogen administration to mares during
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Vernal Transition into the Breeding Season 25
LH NG/ML
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CONTROL ESTROGEN
20 15 10 5 0 0
2
4
6
8
10
DAYS
14
CONTROL ESTROGEN-TREATED
12 10 8 6 4 2 0 3
6 DAYS
9
Figure 178.3 Circulating LH in control and estrogen-treated, ovariectomized mares during vernal transition time. Tests for homogeneity of regression indicated that the two lines were significantly different, as seen in the elevated LH concentrations in estrogen-treated mares.
Figure 178.5 Abundance of mRNA specific for LH beta subunit expression in control and estrogen-treated, ovariectomized mares during vernal transition time. There was significantly more LH beta mRNA by 9 days of estrogen treatment.
vernal transition can enhance LH secretion and increase pituitary LH content. In a more specific experiment, ovariectomized mares (n = 4 per group) were treated with estrogen or control vehicle for 3, 6, or 9 days at an equivalent time of early vernal transition (start date, February 2). Pituitaries were harvested from each group at the designated day of treatment, and pituitary total RNA was evaluated by Northern Blot for specific mRNA encoding equine alpha and LH beta subunit expression.31 As with the previous experiment described above, LH concentrations were increased in the peripheral circulation in estrogentreated, but not in control vehicle-treated mares (Figure 178.3). Importantly, mRNA encoding equine alpha subunit was significantly elevated in pituitaries from estrogen-treated mares on day 9 of treatment, as was mRNA encoding LH beta (see Figures 178.4 and 178.5). Thus, these data indicate that renewal of the gene regulating LH synthesis and secretion can be stimulated by estrogen administration.
It might be easy, then, to assume that the naturally occurring surge in estradiol from the first steroidogenically competent follicle of the year has a causal relationship with the simultaneous secretion of LH at that time. However, it is not likely that simple. For one thing, estradiol secretion and LH secretion just preceding the first ovulation of the year are essentially simultaneous, raising the question of whether one is causally related to the other. Furthermore, these data are not reconciled by the multiple observations that the increase in LH secretion in ovariectomized mares occurs in the absence of estrogens.14, 33, 43 Of these reports, Peltier et al.33 is the most convincing that LH increases in the peripheral circulation at the same time of year in ovarian intact and ovariectomized mares (see Figure 178.6). Therefore, it is imprudent to state that estrogen stimulation of mRNA expression of equine alpha and LH beta is the naturally occurring mechanism. Rather, it may be a pharmacological response, but the clinical value of that response is somewhat questionable, for, although estrogen administration surely increases LH gene expression and secretion, it also has an inhibitory effect on FSH17 and follicular development.44 Therefore, estrogen treatment as a method of advancing the onset of the breeding season is not promising. However, development of protocols in which estrogen could be used to re-establish the LH gene, at the expense of follicular development, might be followed by treatments (or time) to permit FSH concentrations to rise and stimulate a new cohort of follicles. A further complication to attempts to use estrogen to advance the onset of the breeding season is that apparently a photic stimulus is necessary before estrogen has even a pharmacological effect on pituitary subunit mRNA expression. Cleaver and
16 CG beta mRNA RELATIVE ABUNDANCE
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CG beta mRNA RELATIVE ABUNDANCE
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12 10 8 6 4 2 0 3
6 DAYS
9
Figure 178.4 Abundance of mRNA specific for LH alpha subunit expression in control and estrogen-treated, ovariectomized mares during vernal transition time. There was significantly more alpha mRNA by 9 days of estrogen treatment.
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80 70
(a)
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60 50 40 30 20 10 20 (b) 15 LH (ng/ml)
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Non OV
10
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0 −60
−50
−40
−30
−20
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Day Relative to Ovulation
Figure 178.6 Circulating LH and estradiol concentrations prior to the first ovulation of the year. The vertical bar denotes the day of ovulation. Note that LH and estradiol rise at essentially the same time (from Peltier et al., 1998.)33
Sharp45, 46 reported results of experiments to test the importance of photic exposure and/or estrogen treatment on releasable LH. In the first experiment, anestrous mares were exposed to short day photoperiod (L:D 12:12) or long day photoperiod (L:D 24:0) for 28 days. Blood samples were collected daily and at 15-minute intervals for 12 hours on days 14, 21, and 28. On day 29, hypothalami were harvested for analysis of GnRH content.45 LH and GnRH were significantly elevated by day 14 in mares exposed to L:D 24:0, but not to L:D 12:12. In the second experiment, four groups of anestrous mares (n = 4 each; start date, November 23) were exposed to photostimulation (2 groups; 2.5 hours additional light beginning at sunset) or ambient photoperiod (2 groups; approximately L:D 10.5:13.5). The groups were further partitioned such that each group of mares in each photic group received either control vehicle or 5 mg estradiol twice daily for 14 days, beginning after
14 days of photic exposure. LH concentrations were measured daily throughout the 4-week experiment, and GnRH challenge was performed at the end of the experiment. LH concentrations tended (P = 0.061) to be higher in the mares that received both photic stimulation and estrogen compared with the other three groups of mares (see Table 178.2). Furthermore, the LH response to GnRH challenge was significantly (P = 0.007) greater in mares that were exposed to both photostimulation and estrogen (see Figure 178.7). These results indicate that however estrogen acts to induce the gene for LH subunit synthesis, there is a requirement for photostimulation to either precede or accompany the estrogen treatment. This is the concept of a ‘photic gate’, meaning that exposure to stimulatory photoperiod is necessary to adjust central nervous events, allowing biochemical changes such as expression of the mRNA encoding LH subunits. This may add focus to comments about the time of year experimental treatments are begun, and the ‘depth’ of anestrus of a given mare. Interpretation of results of attempts to alter the vernal transition phase should always take into account the potential existence of a previous photostimulation. This is an exciting concept and clearly deserves further research.
Dopamine Several studies on seasonality in sheep have suggested that the neurotransmitter, dopamine, acts on the median eminence, either directly, or through its regulation of prolactin, to inhibit GnRH.47 Because of studies like these in sheep, there has been interest in inhibiting dopamine in horses in attempts to hasten the onset of the breeding season. The two dopamine antagonists that have been evaluated are sulpiride and domperidone and both are dopamine D-2 receptor antagonists. Besognet et al.48 injected two groups of mares (n = 9 each; start date, February 5) with sulpiride (100 mg ∼ 2 mg/kg) daily until the first ovulation was detected, or for a maximum of 58 days. These authors reported that circulating prolactin concentrations were elevated, and that the date of the first ovulation of the year was significantly advanced in sulpiride-treated mares (day 77.3 ± 7.9 vs 110.0 ± 6.8). In contrast, Donadeu and
Table 178.2 LH concentrations following two weeks of exposure to either ambient photoperiod or photostimulation (2.5 artificial light beginning at sunset), and two additional weeks of either control vehicle or estrogen (5 mg) administration twice daily. LH was increased (P = 0.061) only after exposure to both increased photoperiod and estrogen Ambient photoperiod; control vehicle
Ambient photoperiod; estrogen administration
Photostimulation; control vehicle
Photostimulation; estrogen administration
2.42 ± 0.6
2.39 ± 0.6
2.42 ± 0.8
3.96 ± 0.6
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Vernal Transition into the Breeding Season 30 Group 1 Group 2
Group 3 Group 4
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These reports are few in number, and in some cases, contradictory, leaving the subject of dopamine antagonists incomplete. This interesting area deserves further research, especially in the underlying mechanisms of dopamine and/or prolactin in the control of GnRH. Progestins
10
0 GnRH Ĉhallenge Figure 178.7 Circulating LH concentrations in mares following a GnRH challenge. Group 1 = ambient light, control vehicle; Group 2 = ambient light, 5 mg estradiol twice daily for 14 days; Group 3 = photostimulation, control vehicle; and Group 4 = photostimulation, 5 mg estradiol twice daily for 14 days. There was a significantly greater response to GnRH in mares exposed to photostimulation and estradiol compared with all other mares, indicating that the LH stimulatory effects of estrogen require a prior exposure to stimulatory photoperiod. (From Cleaver et al., 1995.)46
Thompson49 administered sulpiride (1 mg/kg body weight) to seasonally anovulatory mares daily from January 14 to February 14 and reported that, although prolactin was elevated, there were no changes in LH or FSH, nor were there any differences in follicular development between control and treated groups. Dixon et al.50 administered domperidone (1.1 mg/kg) or control vehicle daily to anestrous mares (n = 5 each; start date, January 20). At the time of the first and second consecutive follicle of the year (≥ 30 mm) follicular fluid was aspirated via transvaginal ultrasound-guided folliculocentesis, and the fluid analyzed for progesterone and estradiol concentrations. Mean date of first ovulation was not different among groups, but the follicular fluid estradiol concentrations were significantly elevated in the mares receiving domperidone, suggesting potential effects of domperidone on the follicle, either directly or indirectly through its action on dopamine. These results were supported by the report of Brendemuehl and Cross51 who administered domperidone to anestrous mares (n = 8; start date, January 15) until the first ovulation of the year. LH and conjugated estrogens were significantly elevated, although these authors did not find any significant difference in FSH. They concluded that greater follicular development and estrogen conjugate concentrations in the absence of any detectable change in FSH indicated that either domperidone or prolactin acted at the level of the ovary.
The idea of administering progestins to advance the timing of entry into the breeding season has a relatively long history, perhaps dating to a report by Evans et al. (1977)52 in which they reported induction of increased LH and FSH secretion in acyclic mares. They stated that progesterone (150 mg in oil daily for approximately 10 days before administration of GnRH. The rationale for this pretreatment with progesterone was to ‘simulate luteal phase levels,’ ‘since pituitary and/or hypothalamic activity is also modulated by ovarian steroids.’ This rationale seems strange, as the naturally occurring vernal transition is not associated with detectable circulating progesterone concentrations until after the first ovulation of the year and corpus luteum formation. Thus, attempts to induce an earlier ovulation with progesterone treatment must be considered pharmacological. Perhaps the most widespread application of this idea has been administration of the synthetic progestin, altrenogest (RegumateTM ), as it is orally active and easily administered by farm personnel. The report of Squires et al.53 provides focus on this technique as they showed that administration of altrenogest to mares at two different times of spring was ineffective (i.e., no difference between treated and control mares with respect to date of first ovulation) early (February) and more effective (i.e., advanced date of first ovulation vs control mares) later in the spring (April). Furthermore, they pointed out that of 31 mares in the early group, only 3 exhibited follicular development, whereas of 32 mares in the late group, 28 exhibited follicular activity. Thus, one interpretation of these data is that progestin administration to mares in vernal transition with the idea of accelerating the time of first ovulation of the year would only be expected to succeed if the mares were already well into the mechanisms of sexual recrudescence. In further support of this, Alexander and Irvine administered progesterone to mares in transition when their ovaries exhibited some degree of follicular development, and monitored further follicular development and LH and FSH secretion closely. They concluded ‘that exogenous progesterone has no consistent effect on follicular development and gonadotrophin secretion patterns in transition phase mares, and in the present study it did not advance the mean date of
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the first ovulation of the breeding season compared with untreated control mares’.54 Thus, progesterone treatment to mares in vernal transition may simply mask the underlying process already under way, and evidence for accelerated reproductive activity after withdrawal of the progesterone may simply reflect culmination of vernal transition anyway.
Conclusion It is clear from the foregoing that the events of vernal transition represent a rich source of material with which to understand the process of sexual recrudescence. As stated, the crux of vernal transition and re-establishment of reproductive capability is the renewal of mRNA encoding the LH subunits. Understanding the forces that result in renewed LH synthesis and secretion will help develop new and rational regimes for advancing the onset of the breeding season.
References 1. Lincoln GA Andersson H, Hazlerigg D. Clock Genes and the long-term regulation of prolactin secretion: Evidence for a photoperiod/circannual times in the pars tuuberalis. J Neuroendocrinol 2003;15:390–7. 2. Dawson A, Sharp PJ. Photorefractoriness in birds – Photoperiodic and non-photoperiodic control. Gen Comp Endocrinol 2007;153:378–84. 3. MacGregor D, Lincoln GA. A physiological model of a circannual oscillator. J Biol Rhythms 2008;23:252–64. 4. Nonno R, Capsoni S, Lucini V, Møller M, Fraschini F, Stankov B. Distribution and characterization of the melatonin receptors in the hypothalamus and pituitary gland of three domestic ungulates. J Pineal Res 1995;18(4):207–16. 5. Sharp DC. The effects of artificial lighting on reproduction. In: Robinson NE (ed.) Current Therapy in Equine Medicine. Philadelphia: Saunders, 1983; pp. 399–401. 6. Ginther O.J. Reproductive Biology of the Mare. Cross Plains, WI: EquiServices, 1992. 7. King SS, Nequin LG. Spontaneously prolonged corpus luteum activity is a common route into anestrus in the mare. Proceedings of the Twelfth Equine Nutrition and Physiology Sympopsium, University of Calgary, 1991. 8. Strauss SS, Chen CL, Kalra SP, Sharp DC. Localization of gonadotropin releasing hormone in the hypothalamus of ovariectomized pony mares. J Reprod Fertil Suppl 1979;27: 123–9. 9. Sharp DC, Grubaugh W. Seasonal patterns of GnRH secretion in the horses assessed by push-pull perfusion. Biol Reprod 1986;(Suppl 1)34:143. 10. Hart PJ, Squires EL, Imel KJ, Nett TM. Seasonal variation in hypothalamic content of gonadotropin-releasing hormone (GnRH), pituitary receptors for GnRH, and pituitary content of luteinizing hormone and follicle-stimulating hormone in the mare. Biol Reprod 1984;30(5):1055–62. 11. Sherman GB, Wolfe MW, Farmerie TA, Clay CM, Threadgill DS, Sharp DC, Nilson JH. A single gene encodes the β subunits of equine luteinizing hormone and chorionic gonadotropin. Mol Endocrinol 1992;6:951–9.
12. Sharp DC, Grubaugh WR. Use of push-pull perfusion techniques in studies of gonadotrophin-releasing hormone secretion in mares. J Reprod Fertil Suppl 1987;35:289–96. 13. Silvia PJ, Meyer SL, Fitzgerald BP. Pulsatile gonadotropin secretion determined by frequent sampling from the intercavernous sinus of the mare: possible modulatory role of progesterone during luteolysis. Biol Reprod 1995;53(2): 438–46. 14. Freedman LJ, Garcia MC, Ginther OJ. Influence of photoperiod and ovaries on seasonal reproductive activity in mares. Biol Reprod 1979;20(3):567–74. 15. Bergfelt DR, Ginther OJ. Embryo loss following GnRHinduced ovulation in anovulatory mares. Theriogenology 1992;38:33–43. 16. Silvia PJ, Johnson L, Fitzgerald BP. Changes in the hypothalamic-hypophyseal axis of mares in relation to the winter solstice. J Reprod Fertil 1992;96(1):195–202. 17. Sharp DC, Grubaugh,WR, Davis SD, Weithenauer J, Wilcox. CJ. Effects of steroid administration on luteinizing hormone and follicle stimulating hormone in ovariectomized pony mares in the springtime: pituitary responsiveness to gonadotropin releasing hormone (GnRH), circulating gonadotropin concentrations, and pituitary gonadotropin content. Biol Reprod 1991;44:983–990. 18. Watson ED, Thomassen R, Steele M, Heald M, Leask R, Groome NP, Riley SC. Concentrations of inhibin, progesterone and oestradiol in fluid from dominant and subordinate follicles from mares during spring transition and the breeding season. Anim Reprod Sci 2002;74(1–2): 55–67. 19. Donadeu FX, Ginther OJ. Follicular waves and circulating concentrations of gonadotrophins, inhibin and oestradiol during the anovulatory season in mares. Reproduction 2002;124(6):875–85. 20. Davis SD, Grubaugh W, Weithenauer J. Follicle integrity and serum estradiol 17β patterns during sexual recrudescence in the mare. Biol Reprod 1987;Suppl 1:143. 21. Seamans KW, Sharp DC. Changes in follicular aromatase activity during transition from winter anestrus. J Reprod Fertil Suppl 1982;32:225–33. 22. Davis SD, Sharp DC. Intra-follicular and peripheral steroid characteristics during vernal transition in the pony mare. J Reprod Fertil Suppl 1991;44, 333–40. 23. Tucker KE, Cleaver BD, Sharp DC. Does resumption of follicular estradiol synthesis during vernal transition in mares involve a shift in steroidogenic pathways? Biol Reprod 1993;Suppl 1:519. 24. Watson ED, Bae SE, Steele M, Thomassen R, Pedersen HG, Bramley T, Hogg CO, Armstrong DG. Expression of messenger ribonucleic acid encoding for steroidogenic acute regulatory protein and enzymes, and luteinizing hormone receptor during the spring transitional season in equine follicles. Domest Anim Endocrinol 2004;26(3):215–30. 25. Channing CP. Studies on tissue culture of equine ovarian cell types: pathways of steroidogenesis. J Endocrinol 1969; 43:403–14. 26. Kenney RM, Condon W, Ganjam VK, Channing C. Morphological and biochemical correlates of equine ovarian follicles as a function of their state of viability or atresia. J Reprod Fertil Suppl 1979;(27):163–71. 27. Younglai EV, Short RV. Pathways of steroid biosynthesis in the intact Graafian follicle of mares in estrus. J. Endocrinol 1970;47:321–31. 28. Baldwin DM, Roser JF, Muyan M, Lasley B, Dybdal N. Direct effects of free and conjugated steroids on GnRH
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stimulated LH release in cultured equine anterior pituitary cells. J Reprod Fertil Suppl 1991;44:327–32. Sharp DC, Garcia MC, Ginther OJ. Luteinizing hormone during sexual maturation in pony mares. Am J Vet Res 1979; 40:548. Sharp DC, Wolfe MW, Cleaver BD, Nilson J. Effects of estradiol-17ß administration on steady-state messenger ribonucleic acid (mRNA) encoding equine and LH/CGß subunits in pituitaries of ovariectomized pony mares. Theriogenology 2001;55:1083–93. Albrecht BA, MacLeod JN, Daels PF. Differential transcription of steroidogenic enzymes in the equine primary corpus luteum during diestrus and early pregnancy. Biol Reprod 1997;56(4):821–9. Albrecht BA, MacLeod JN, Daels PF. Expression of 3beta-hydroxysteroid dehydrogenase, cytochrome p450 17alpha-hydroxylase/17,20-lyase and cytochrome p450 aromatase enzymes in corpora lutea of diestrous and early pregnant mares. Theriogenology 2001;55(2):551–61. Peltier MR, Robinson G, Sharp DC. Effects of melatonin implantation in pony mares. 2. Long term effects. Theriogenology 1998;49:1126–42. Gentry LR, Thompson DL Jr, Stelzer AM. Responses of seasonally anovulatory mares to daily administration of thyrotropin releasing hormone and(or) gonadotropin releasing hormone analog. J Anim Sci 80:208–13. Mumford EL, Squires EL, Jasko DJ, Nett TM. Use of gonadotropin-releasing hormone, estrogen, or a combination to increase releasable pituitary luteinizing hormone in early transitional mares. J Anim Sci 1994;72:174–7. Evans MJ, Irvine CHG. Measurement of equine follicle stimulating hormone and luteinizing hormone: response of anestrous mares to gonadotropin releasing hormone. Biol Reprod 1976;15:477–84. McKinnon AO, Vasey JR, Lescun TB, Trigg TE. Repeated use of a GnRH analogue deslorelin (Ovuplant) for hastening ovulation in the transitional mare. Equine Vet J 1997; 29(2):153–5. McKinnon AO, Nobelius AM, del Marmol Figueroa ST, Skidmore J, Vasey JR, Trigg TE. Predictable ovulation in mares treated with an implant of the GnRH analogue Deslorelin. Equine Vet J. 1993;25(4):321–3. Hyland JH, Wright PJ, Clarke IJ, Carson RS, Langsford DA, Jeffcott LB. Infusion of gonadotropin-releasing hormone (GnRH) induces ovulation and fertile oestrus in mares during seasonal anestrus. J Reprod Fertil Suppl 1987;35:211–20. Porter MB, Cleaver BD, Peltier M, Robinson G, Thatcher WW, Sharp DC. Comparative study between pony mares and ewes evaluating gonadotrophic response to administration of gonadotrophin-releasing hormone. J Reprod Fertil 1997;110:219–29.
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41. Johnson CA, Thompson DL Jr, Cartmill JA. Pituitary responsiveness to GnRH in mares following deslorelin acetate implantation to hasten ovulation. J Anim Sci 2002; 80(10):2681–7. 42. Davis SD, Sharp DC, Grubaugh WR. Stimulation of LH by LHRH analogs in anestrous pony mares. Biol Reprod 1986(Suppl 1)34:143. 43. Garcia MC, Freedman LJ, Ginther OJ. Interaction of seasonal and ovarian factors in the regulation of LH and FSH secretion in the mare. J Reprod Fertil Suppl 1979;(27):103–11. 44. Woodley SL, Burns PJ, Douglas RH, Oxender WD. Prolonged interovulatory interval after oestradiol treatment in mares. J Reprod Fertil Suppl 1979;27:205–9. 45. Cleaver BD, Davis SD, Grubaugh WR, Franklin K, Sheerin PC, Sharp DC. The effect of constant light exposure on circulating gonadotropin levels and hypothalamic gonadotropin-releasing hormone (GnRH) content in the ovariectomized (OVX) pony mare. J Reprod Fertil Suppl 1991;44:259–66. 46. Cleaver BD, Sharp DC. LH secretion in anestrous mares exposed to artificially lengthened photoperiod and treated with estradiol. Biol Reprod 1995; Monograph Series No. 1:449–457. 47. Thiery JC, Chemineau P, Hernandez X, Migaud M, Malpaux B. Neuroendocrine interactions and seasonality. Domest Anim Endocrinol 2002;23:87–100. 48. Besognet B, Hansen BS, Daels PF. Induction of reproductive function in anestrous mares using a dopamine antagonist. Theriogenology 1997;15;47(2):467–8. 49. Donadeu FX, Thompson DL Jr. Administration of sulpiride to anovulatory mares in winter: effects on prolactin and gonadotropin concentrations, ovarian activity, ovulation and hair shedding. Theriogenology 2002;15;57(2):963–76. 50. Dixon LN, Sharp DC, Porter MB, Pelehach LM, Cross DL. The effects of domperidone and hCG treatment on follicular steroidogenesis in vernal transitional pony mares. Biol Reprod 2000;Suppl 1:137. 51. Brendemuehl JP, Cross DL. Influence of the dopamine antagonist domperidone on the vernal transition in seasonally anoestrous mares. J Reprod Fertil Suppl 2000;56:185–93. 52. Evans MJ, Irvine CH. Induction of follicular development, maturation and ovulation by gonadotropin releasing hormone administration to acyclic mares. Biol Reprod 1977; 16(4):452–62. 53. Squires EL, Heesemann CP, Webel SK, Shideler RK, Voss JL. Relationship of altrenogest to ovarian activity, hormone concentrations and fertility of mares. J Anim Sci 1983;56(4): 901–10. 54. Alexander SL, Irvine CH. Control of onset of breeding season in the mare and its artificial regulation by progesterone treatment. Reprod Fertil Suppl 1991;44:307–18.
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Chapter 179
Estrus Patrick M. McCue, Charles F. Scoggin and Alicia R.G. Lindholm
Mares ovulate at approximately 21-day intervals throughout the breeding season. The equine estrous cycle can be divided into phases based either on sexual receptivity (estrus versus diestrus) or physiology (follicular phase versus luteal phase). The two categories largely, but not completely, overlap with behavioral estrus occurring primarily during the follicular phase and behavioral diestrus occurring almost exclusively during the luteal phase.
Definition of estrus Estrus is the period during which a mare is sexually receptive to the advances of a stallion. Behavioral estrus is stimulated by increasing levels of estradiol produced by the developing dominant follicle in the absence of progesterone. The average length of estrus has been reported to be 6.5 days, with a range of 4.5 to 8.9 days.1 Duration of estrus is related to the follicle size at the end of diestrus, growth rate of the dominant follicle, and the ultimate size of the follicle prior to ovulation. Ovulation typically occurs 24 to 48 hours before the end of estrus. An increase in progesterone from the developing corpus luteum is responsible for the cessation of behavioral estrus.
Sexual behavior involves a complex sequence of interactions between a stallion and mare.2, 3 A mare should be teased with a stallion that exhibits good libido in order to successfully evaluate estrous cycle stage. Adequate time should be taken to allow shy or nervous mares to express behavioral estrus, and knowledge of the mare’s previous behavioral patterns may be helpful. Expression of estrus does not always indicate that a mare is in the follicular phase of the estrous cycle. Seasonally anovulatory mares, ovariectomized mares, and pregnant mares have all been reported to occasionally show signs of estrus when teased with a stallion.4, 5
Silent estrus Maiden mares may not show heat well, and foaling mares may not show heat unless the foal is restrained and/or safely away from the stallion. Subordinate mares may be inhibited from expressing estrus in the presence of a dominant mare.2 In addition, a mare may have a preference for or an aversion toward an individual stallion.6 Mares with ‘silent estrus’ may have lower concentrations of estradiol-17β than mares expressing normal estrus.7
Estrus detection
Individual teasing
The common use of ultrasonography and artificial insemination has reduced the dependency on teasing or estrus detection in many breeding programs. However, in breeds that require natural service or programs that rely on natural service, and in cases where ultrasonographic techniques return ambiguous results, teasing is still an important part of reproductive management.
An individual mare should be exposed to a stallion for a interval of time that is long enough for her to show estrous or diestrous types of behavior. Mares that remain indifferent may need to be teased longer, teased with a different stallion, or may just show more subtle signs. Mares may be reluctant initially and subsequently show frank estrous behavior (i.e., ‘break down’). Sometimes full behavioral estrus is
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 179.1 Common behavioral characteristics of mares in estrus Tail raised and arched or deviated to one side Rhythmic eversion of the labia and exposure of clitoris (‘winking’) Passive urination Ears relaxed and either held forward or in a neutral position Rear limbs slightly abducted (i.e. wide-based stance) Stifles and hocks flexed Lowering of the pelvis (i.e. ‘squatting’) Leaning into fence or gate Vocalization (squealing) Calm behavior; does not try to bite or strike stallion
Figure 179.1 Photograph of a mare in estrus (i.e. teasing in heat). Note the base-wide stance, raised tail, and urination.
only expressed within a few hours of ovulation. It is also not unusual for a mare to fail to show signs of estrus while being directly teased to a stallion, and then break down as the stallion leaves. Mares may also display estrus at the mere sound or sight of a stallion.
the mare exhibits overt, subtle, or no signs of estrus throughout a cycle (Table 179.2).
Follicular development and selection of the dominant follicle Group teasing A stallion may be used to tease more than one mare concurrently if he is brought to the edge of a pen or turned out adjacent to a group of mares. Mares are allowed to approach the stallion at will in such a teasing program. However, some mares will not approach the stallion and will not express estrus when teased as part of a group. It may be necessary to tease such mares individually. It is often not very efficient to tease mares as a group, since often the only mares that come to the fence or tease rail are assertive mares in heat or mares that want to attack the stallion. One may not be able to determine the heat status of mares that remain a distance from the stallion. It is generally more effective, but certainly more time consuming, to tease mares individually.2 There are many systems used for teasing mares, including chutes, rails, fences, pens, and paddocks. Keys to successful teasing are patience, persistence and knowing the behavioral characteristics of each mare. Consequently, it is advantageous for the same individual(s) to tease mares each day, so that slight variations in individual responses can be recognized. Common behavioral responses of mares in estrus when teased with a stallion include raising of the tail, passive urination, repeated eversion of the labia with exposure of the clitoris (winking), and assuming a mating posture with the back legs spread apart (Table 179.1 and Figure 179.1). Maintaining an accurate record of teasing behavior will be helpful when monitoring the estrous cycle of a mare. Notations can be made as to whether or not
Mares utilize an active process of follicular selection to limit the number of ovulations in most estrous cycles to one.8 The mechanism of follicle selection involves a complex, closely associated interaction between pituitary gonadotropins and ovarian hormones. The major or primary follicular wave of the equine estrous cycle, or the wave that ends with ovulation during estrus, originates in diestrus approximately 7–8 days after ovulation when a cohort of follicles emerge from a pool of smaller follicles over a period of several days.9 After emergence, follicles in the wave go through a common-growth phase for approximately 6–7 days, during which all follicles increase in size at approximately the same rate (3 mm/day). Deviation occurs at the end of the common-growth phase and represents the point at which the future dominant follicle continues to develop and smaller subordinate follicles regress. Follicle-stimulating hormone (FSH) produced by the anterior pituitary gland provides the drive for Table 179.2 Abbreviations for responses of a mare to a stallion (i.e., teasing behavior)
Abbreviation
Levels: option 1
Levels: option 2
H O H/O
+ – +/–
+++ – ++
In heat Out of heat Coming into or going out of heat
+
Indifferent
I
Behavior
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emergence and initial growth. Small follicles in the follicular wave require FSH for initial growth and development. The future dominant follicle usually emerges approximately 1 day earlier than the other follicles in the wave.9 Consequently, this follicle is slightly larger than the other follicles in the cohort at the end of the common-growth phase. Circulating levels of FSH peak when the largest follicle is approximately 13 mm in diameter.9–11 Levels of FSH subsequently decline as, first, inhibin and then estradiol are produced by the larger follicles within the wave. Deviation takes place approximately 3 days after the FSH peak as levels of FSH are declining8 (Figure 179.3).
Estrus detection on a Thoroughbred breeding farm Teasing of mares is an invaluable resource in the breeding management of mares on a large Thoroughbred farm. In the vast majority of cases, estrus behavior is readily apparent. However, it is important to recognize individual characteristics or unique signs of estrus in certain mares. Consequently, obtaining and maintaining a detailed reproductive history on all mares can optimize breeding efficiency. Detection of estrus is accomplished by: (i) observation of a mare’s behavior when in the presence of a stallion (teasing); (ii) observation of a mare’s behavior to other mares that she is kept with out in a pasture or paddock; and (iii) simulation of breeding.
Teasing mares Mares are teased early in the morning (∼ 5 a.m.). At least two individuals are present to assist with the process. One person handles the stallion, while the other person evaluates the mare before and after she is introduced to the stallion. If the mare has a foal at side, care is taken to ensure that the foal is standing and not in harm’s way. Starting with the first stall, the stall door is opened, and the stallion is allowed to place his head into the stall. He is allowed to nicker at and nuzzle on the mare. Mares that are in behavioral estrus will often demonstrate one or more of the following signs: nuzzling of the stallion, raising of their tail, posturing in lordosis, urinating, and eversion of the clitoris (‘winking’). Mares that are not in heat (i.e., in diestrus) will immediately display their displeasure at the presence of the stallion by pinning their ears back, lunging, and/or spinning and kicking at the stallion. The stallion is then taken to another stall where he is allowed to tease another mare, and the
process is continued until the stallion has teased all the mares in the barn. The individual not handling the stallion will observe the mare for a short period of time (15–30 seconds) after the stallion has moved to another stall to see if the mare will display any outward signs of estrus. Such a method is useful for mares that are highly protective of a foal (i.e., ‘foal proud’), anxious, or shy of the stallion. In some cases, the behavioral state of a mare may not be readily apparent; that is, a mare may seem relatively indifferent to the presence of the stallion. In these cases, the mare is backed out of the stall and positioned such that half of her body is within the stall and the other half within the barn alley. The stallion is then led towards the flank of the mare and allowed to sniff and nuzzle her flank and perineum, but not allowed to mount the mare. If a mare tolerates this process, then she is judged to be in heat. Mares may also be teased in pastures or paddocks. However, some shy mares may not show estrus well when teased in a group, and a dominant mare may prevent a subordinate mare from interacting with a stallion in a group setting. Stallions may be placed in specialized teasing areas adjacent to or within a paddock of mares.
Observation in pasture All mares are observed in their pastures or paddocks at least twice daily for signs of estrus (e.g., nuzzling, squealing, posturing in lordosis, and, occasionally, mounting other mares). This technique has proven particularly useful for maiden mares, which are often either shy of or anxious in the presence of a teaser stallion. If a mare is observed to be teasing another mare, both mares will be placed on the veterinary examination list to be further evaluated by palpation and ultrasonography. Behavior of mares will also be observed as the teaser stallion is led either through or next to a pasture housing mares as he is being returned to his paddock. Occasionally, mares will display signs of estrus as the stallion is led past, despite not being in close proximity to the stallion. These mares will also be placed on the veterinary examination list to determine if they are suitable for mating.
Evaluation of past teasing and reproductive history Approximately 80% of mares consistently demonstrate typical signs of estrus when they are close to or ready for breeding. The remaining 20% of mares will either be shy, anxious, afraid, indifferent, or quiet to a stallion when teased.
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Estrus Evaluation of a mare’s past teasing and reproductive history can help determine when she is suitable for mating. In certain cases, a mare may display unique and individual signs of estrus. For instance, some mares simply become quiet when they are ready for breeding; more specifically, these mares will stand quietly at the stall door when introduced to the teaser stallion, yet not display any outward signs of estrus. Often this type of behavior has been observed in the past and subsequently recorded for this particular mare. In other cases, an individual mare, when in heat, may stand with her rear end to the door upon arrival of the stallion, whereas when she was not in heat, she would stand as far off in the corner as possible, as if attempting to avoid the stallion. This mare may be described as ‘changing’ in that her behavior towards the teasing process has changed to where she has now shown her own unique sign of behavioral estrus. Again, this particular mare’s behavior has likely been observed in the past and recorded as such, thereby indicating that she may be ready for breeding even though she may never display more characteristic signs of estrus. Other cases in which a past reproductive history has proved to be useful are post-foaling mares. As a routine, all mares are evaluated with rectal palpation and ultrasonography 28 days post-foaling for possible breeding on their 30-day heat cycle or three to four days after PGF2α administration on day 16–18 after parturition. This examination is performed regardless of whether or not they displayed signs of estrus when teased. Such a method is useful for those mares that are overly protective of their foal or do not consistently display signs of estrus even when not having a foal by their side. It is also useful for determining when the mare should be evaluated next and when to tentatively book her to a stallion.
Simulation of breeding (‘test jumping’) Teaser stallions should have good libido and an even temperament. One teaser is used exclusively at the breeding shed. In addition to teasing mares through a partition, he will also be asked to simulate the act of breeding (i.e., ‘test jump’) certain mares prior to the actual act of breeding by a commercial stallion. Test jumps are routinely performed on non-resident maiden mares that are to be bred for the first time. Such a process is performed to gauge both the temperament and breeding behavior of the maiden mare, as well as to avoid any undue harm to a standing stallion. The act of breeding is new to maiden mares, and they can become anxious, fractious, or even aggres-
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Figure 179.2 Teaser stallion wearing a leather apron during a test jump.
sive when they first arrive to be bred. The teaser must therefore also demonstrate sensitivity, perseverance, tenacity, and courage when test jumping maiden mares. A large, leather ‘apron’ or ‘shield’ may be secured around his ventral abdomen to prevent accidental intromission during a test jump (Figure 179.2). Test jumps are performed as many times as needed until the stallion foreman is satisfied that the mare will not harm the stallion to which she will ultimately be mated. As a routine, resident maiden mares are also ‘test jumped’ prior to departing for the breeding shed. The test jump appears to desensitize them to the process of being mounted by a stallion. Furthermore, it allows for the evaluation of their breeding behavior, especially if they did not display overt signs of estrus, but were deemed to have suitable follicular development, uterine edema, and cervical relaxation to warrant being bred. Teaser stallions are allowed to breed nurse mares that were purchased to raise orphan foals. While they may only breed one or two mares a year, it does appear to maintain their interest and dampen any perceived sexual frustration. Follicular deviation in horse mares occurs when the largest follicle is approximately 21 to 23 mm in diameter.8, 11 The process of deviation represents selection against future development of subordinate follicles by inhibin and estradiol produced by the dominant follicle.9, 10 The developing dominant follicle becomes increasingly responsive to FSH and luteinizing hormone (LH) as compared to other follicles in the wave, due in part to an increase in the number of gonadotropin receptors in the dominant follicle, stimulated by elevated intrafollicular estradiol levels.10, 13 Consequently, the larger follicle continues to develop in the face of declining circulating FSH levels, while subordinate follicles begin to regress.
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Mare: The Estrous Cycle GRAAFIAN FOLLICLE
Primary Ovulatory Wave
(a)
35
Diameter (mm)
30
Dominant follicle
25 20 day 2–6 Largest subordinate follicle
15
day 8–12
day 16–18
day 20–22
OVULATION
OVULATION
(ng/ml)
10 5
ESTRADIOL
50
LH
40
(b)
40 LH
Inhibin
8
36
FSH
30 20
32
FSH
6
28 4 Estradiol
24
Inhibin (ng/ml)
FSH (ng/ml) and estradiol (pg/ml)
10
20
2
10 0
ESTRUS 0
ESTRUS 2
4
6
8
10 12 DAYS
14 16 18
20
Figure 179.4 Hormone patterns during estrus in mares (modified from Daels and Hughes, 1993).19
16 0 40
(c)
10 9
32
8
28
LH
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24
5 4
20
3 16
2 Progesterone
12
Progesterone (ng/ml)
36
7 LH (ng/ml)
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FSH
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8 10
12
14 16 18 20 Days after ovulation
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Figure 179.3 Hormone profiles during follicular development and estrus in mares (From Bergfelt and Adams, 2007).12
An increase in luteinizing hormone levels occurs concurrently with an increase in estradiol. Estradiol produced by the developing dominant follicle may be responsible for stimulating the increase in LH secretion from the anterior pituitary.14, 15 Luteinizing hormone is responsible for the final maturation of the follicle and induction of ovulation.16 The profile of LH secretion during estrus is unique in the mare. In contrast to other domestic animal species, mares have a broad LH surge which peaks 1 to 2 days after ovulation17, 18 (Figure 179.4).
Estradiol-17β and inhibin concentrations during estrus Estradiol levels peak 1 to 2 days prior to ovulation.18, 20 The decrease in estradiol prior to ovula-
tion is likely due to luteinization of granulosa cells, and is reflected by a synchronous decrease in uterine edema. Estradiol-17β is responsible for a variety of physical, endocrine, and behavioral changes in the mare. These include relaxation of the cervix, stimulation of edema formation in the uterus, increase in the character and volume of uterine secretions, stimulation of pituitary LH release, and induction of behavioral estrus. Inhibin is a glycoprotein hormone produced by granulosa cells of the developing dominant follicle. Concentrations of inhibin are positively correlated with estradiol and inversely correlated with FSH.20–22 The primary physiological role of inhibin is the specific suppression of pituitary FSH secretion, and therefore the process of follicle selection.23 Inhibin levels increase during estrus, peak on the day of ovulation, and decline to low levels during diestrus20–22 (Figure 179.5).
Prediction of ovulation Factors that can be used to predict ovulation are listed in Table 179.3. The dominant follicle often changes from a spherical structure prior to ovulation to a more irregular shape just prior to ovulation. In addition, a follicle typically becomes softer within 12 hours prior to ovulation. Transrectal ultrasonography may reveal a thickened echogenic border around the pre-ovulatory follicle (Figure 179.6) and a decrease in uterine edema from that noted the previous day.
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0.7 0.6 Inhibin (ng/ml)
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0.5 0.4 0.3 0.2 0.1 –2
0
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6 8 10 12 14 16 18 20 22 Estrous Cycle Day
Figure 179.5 Pattern of inhibin secretion during the equine estrous cycle. The arrow indicates the day of ovulation (McCue, 1993).24
A majority of mares of a given breed ovulate follicles of approximately the same diameter during most estrous cycles. Individual mares occasionally ovulate a follicle significantly smaller than predicted, and a breeding opportunity may be missed. In general, larger mares ovulate follicles of a greater diameter than smaller mares. Within a breed, follicle size at ovulation is smaller in aged mares, postpartum mares, transitional mares, and mares with multiple dominant follicles.18 Mares administered an ovulation-inducing agent may ovulate a smaller follicle than if they were left to ovulate spontaneously.
Palpation and ultrasonography of the estrous mare Palpation Transrectal palpation of the reproductive tract is often performed in conjunction with ultrasonography. Manual palpation can identify certain features of the
Figure 179.6 Ultrasonographic image of a preovulatory follicle with a thick echogenic border.
reproductive tract that cannot be easily detected by ultrasonography. These include tone in the uterus and cervix, softness of an ovarian follicle, sensitivity of the ovary to touch and pressure, and parovarian cysts. Ovarian structures that may be noted during palpation include developing and pre-ovulatory follicles (Figure 179.7), a recent ovulation, and a corpus hemorrhagicum. Mares may exhibit a significant pain response upon palpation of a fresh ovulation site, and the collapsed follicular wall can often be detected. A corpus hemorrhagicum has a tactile feel of a wet sponge. In many cases it may be necessary to examine each ovary by palpation with the contralateral hand to facilitate observations of mare behavior and to evaluate follicle softness, follicle collapse, etc. In contrast, a mature corpus luteum cannot be detected by transrectal palpation in the mare. The uterus is generally large and edematous during estrus and has a characteristic heavy or ‘doughy’ feel. The cervix of a mare in heat is generally soft and
Table 179.3 Factors used to predict ovulation in mares Previous history of mare Mare size and breed Follicle size Growth pattern of follicle Number of days in estrus Shape of follicle Tone/Softness of follicle Thickness of follicle border measured using ultrasonography Decrease in uterine edema Administration of an ovulation inducing agent Time of the year
Figure 179.7 Large pre-ovulatory equine follicle.
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Figure 179.8 Ultrasonographic image of a large preovulatory follicle in a mare.
relaxed, and in some instances may be difficult to distinguish. Maiden mares tend to have tighter cervices when assessed by rectal palpation compared to older mares in the same stage of estrus. After ovulation, the increase in progesterone causes the uterus and cervix to increase in tone. Ultrasonography Ovarian follicles contain clear yellow-orange colored follicular fluid, which shows up as a black image ultrasonographically (Figure 179.8). Sequential ultrasonographic examinations of a mare in estrus will allow for evaluation of follicular growth, changes in shape, characteristics of the follicular wall and follicular fluid, and evaluation of the pattern of uterine edema. The dominant follicle will increase in diameter at a rate of 3 to 5 mm per day. Maximal follicle size is often reached 1 to 2 days prior to ovulation. Ovulation can be recognized as a rapid decrease in follicular size, the absence of a previous large follicle, and/or the presence of a collapsed structure (Figure 179.9).25, 26 Ovulation has been reported to occur over a period of less than 60 seconds.25, 27 Clinically, it is advantageous to record the size of all follicles > 30 mm on each ovary, as well as the size of the largest follicle < 30 mm on each ovary. This will allow the practitioner to track follicular growth, monitor number of ovulations, and predict the potential for twin pregnancies. In addition, it is recommended that all follicles > 30 mm present at ovulation of the dominant follicle be monitored for an additional 1 to 4 days to
Figure 179.9 Ultrasonographic image of a collapsed follicle (i.e. a fresh ovulation).
determine whether or not they continue to develop and ovulate, or regress. After ovulation, the follicle lumen fills with blood, forming the corpus hemorrhagicum. This is initially detected as a structure with an echogenic rim and a hypoechoic to anechoic center that may develop strands of fibrin as clotting occurs) on ultrasonography (Figure 179.10). The corpus hemorrhagicum matures into the corpus luteum over the next few days and takes on a more echogenic (whiter) ultrasonographic appearance (Figure 179.11). Ultrasound can also help differentiate abnormal
Figure 179.10 Ultrasonographic photograph of a corpus hemorrhagicum.
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Figure 179.11 Ultrasonographic image of a mature corpus luteum.
conditions of the ovary, such as anovulatory follicles (Figs 179.12 and 132.13) and ovarian tumors. Ultrasonography of the uterus is an important part of a reproductive examination. Mares in estrus usually have edema present in the endometrium, yielding a characteristic ‘sand dollar’, ‘sliced orange’, or
Figure 179.13 Ultrasound image of a large (80 mm) nonluteinized anovulatory follicle.
‘spoke wheel’ appearance. Edema is caused by the presence of elevated estrogen and low progesterone levels. The pattern of edema changes over the course of estrus, and scoring systems have been used to monitor the pattern of uterine edema (Table 179.4).28 Edema scores typically increase with growth of the dominant follicle, peak 1 to 2 days prior to ovulation, and decrease the day prior to ovulation. In contrast, the uterus of a mare in diestrus appears tubular and homogeneous without edema. Care in interpretation of uterine edema is necessary as inflammatory changes can also cause edema. A small volume of clear (non-echogenic) fluid may be present in the uterus of a normal mare in estrus. However, an increased volume of echogenic fluid is suggestive of inflammation, infection, or urine in the uterus (Figure 179.14). On occasion this fluid may not be easily detected until uterine edema subsides.
Vaginal examination of an estrous mare
Figure 179.12 Ultrasound image of a hemorrhagic anovulatory follicle prior to luteinization.
A white plaque is often observed on the ventral aspect of the vulva of a mare in estrus (Figure 179.15). The plaque is comprised of calcium carbonate crystals adhered to the vulva by estrual mucus. This should be differentiated from purulent discharge from a mare with endometritis.
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Table 179.4 A scoring system used to evaluate uterine edema in mares at Colorado State University Edema score
Edema amount
Description
0
None
No edema present; individual endometrial folds not discernable; homogenous echotexture typical of diestrus or anestrus
1
Slight
Endometrial folds easily observed in a light ‘spoke-wheel’ pattern; edema may be more evident in uterine horns than uterine body. Typical of early estrus as dominant follicle is developing or late estrus prior to or at time of ovulation
2
Moderate
Endometrial folds increased in thickness; edema pattern obvious throughout uterus. Typical of mid-estrus and usually represents the peak estrogen effect noted 1 to 2 days prior to ovulation
3
Heavy
Large distended endometrial folds; exaggerated degree of edema; not typical of a normal mare in estrus. May be associated with uterine inflammation
Ultrasonographic image
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Table 179.5 Comparison of cervical characteristics of a mare in estrus versus diestrus Characteristic
Estrus
Diestrus
Location
Draped on floor of vagina
High on cranial wall of vagina
Color Moistness Lumen Edema
Pink Moist May be open Edematous
Pale to white Dry Tightly closed Not edematous
A vaginal speculum examination may be performed to evaluate the vaginal vault and external os of the cervix, determine the stage of the estrous cycle, and to detect pathological conditions such as urine pooling, cervical discharge, and trauma. A sterile vaginal speculum is lubricated with a sterile water-based lubricant, inserted between the labia, and advanced forward and upward at a 45◦ angle until the speculum has passed through the vestibular-vaginal fold. The speculum is then advanced horizontally through the vestibular-vaginal fold toward the cranial end of the vagina. A pen-light or other light source is used to illuminate the vaginal vault. The external os of the cervix is identified and notations made regarding location, shape, tone,
color, and presence of secretions. The presence or absence of urine in the cranial vagina is also recorded. The visual examination is continued as the speculum is withdrawn to identify abnormalities of the caudal vagina and vestibule. Appearance of the external cervical os varies with season, stage of the estrous cycle (Table 179.5), pregnancy status, and presence of infection. When a mare is in winter anestrus, the cervix appears pale, dry, and may be open. During estrus, the cervix relaxes onto the floor of the cranial vagina, becomes edematous, pink, and is moistened with clear mucus (Figure 179.16). In diestrus, the cervix is high on the cranial wall of the vagina, pale, dry, and may be covered with a viscous, sticky mucus (Figure 179.17). The cervix of a pregnant mare is tightly closed and may have a thick mucus cap. In the presence of infection, the cervix becomes hyperemic, or reddened, and may have a purulent discharge. Urine pooling is recognized by an accumulation of cloudy yellow fluid in the cranial vagina. Pooling of urine is most common in older mares with poor perineal conformation. Multiple examinations throughout the estrous cycle may be necessary to detect urine pooling, as some mares only accumulate urine in the vagina during estrus. Relaxation of the cervix during estrus may allow for a direct flow of urine into the
Figure 179.15 White plaque at the ventral commissure of the vulva of a mare in estrus.
Figure 179.16 Photograph of a pink, moist, relaxed cervix of a mare in estrus.
Figure 179.14 Large volume of echogenic fluid in a mare 24 hours after insemination.
Speculum examination
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Figure 179.17 Photograph of a dry, tight, pale cervix of a mare in diestrus.
Figure 179.20 Cervical adhesions in a mare.
uterine lumen, resulting in significant inflammation and a reduction in fertility. A persistent hymen may be encountered during speculum examination of maiden mares (Figure 179.18). The hymen is located at the junction of the vestibule and vagina, approximately 6 to 8 cm inside the vulva. The hymen may be complete, occluding the entire vestibular-vaginal opening, resulting in an accumulation of mucus and cellular debris cranial to the hymen, or the hymen may be a partial membrane. The hymen should be manually transected or opened prior to breeding. Vaginal varicose veins may be 0.5 cm or larger in diameter and are located in the region of the vestibular-vaginal junction (Figure 179.19). These blood vessels may be a source of intermittent bleeding from the vulva. Digital examination of the cervix
Figure 179.18 Persistent hymen in a mare.
After the speculum examination is completed, the cervix may be examined manually for patency and the presence of abnormalities. Wearing a sterile obstetrical sleeve, the examiner inserts an index finger into the external cervical os, and the cervix is examined manually between the thumb and index finger for tone, patency, and the presence of adhesions, lacerations, or other abnormalities (Figure 179.20). During estrus, it may be possible to insert two to three fingers into the cervix, while it may prove difficult to insert a single finger into the cervical os during diestrus due to the increased tone stimulated by high progesterone levels.
References
Figure 179.19 Vaginal varicose veins in a mare.
1. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 1st edn. Cross Plains, WI: Equiservices, 1979. 2. Crowell-Davis SL. Sexual behavior of mares. Horm Behav 2007;52:12–17. 3. Curry MR, Eady PE, Mills DS. Reflections on mare behavior: social and sexual perspectives. J Vet Behav 2007;2:149–57.
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Estrus 4. Asa CS, Goldfoot DA, Garcia MC, Ginther OJ. Sexual behavior in ovariectomized and seasonally anovulatory mares (Equus caballus). Horm Behav 1980;14:46–54. 5. Asa CS, Goldfoot DA, Ginther OJ. Assessment of the sexual behavior of pregnant mares. Horm Behav 1983;17:405–13. 6. Pickerel TM, Crowell-Davis SL, Caudle AB, Estep DQ. Sexual preferences of mares (Equus caballus) for individual stallions. Appl Anim Behav Sci 1993;38:1–13. 7. Nelson EM, Kiefer BL, Roser JF, Evans JW. Serum estradiol17β concentrations during spontaneous silent estrus and after prostaglandin treatment in the mare. Theriogenology 1985;23:241–62. 8. Ginther OJ, Beg MA, Bergfelt DR, Donadeu FX, Kot K. Follicle selection in monovular species. Biol Reprod 2001;65:638–47. 9. Gastal EL, Gastal MO, Bergfelt DR, Ginther OJ. Role of diameter differences among follicles in selection of a future dominant follicle in mares. Biol Reprod 1997;57: 1320–7. 10. Donadeu FX, Ginther OJ. Interrelationships of estradiol, inhibin, and gonadotropins during follicle deviation in pony mares. Theriogenology 2004;61:1395–405. 11. Ginther OJ, Beg MA, Gastal MO, Gastal EL. Follicle dynamics and selection in mares. Anim Reprod 2004;1:45–63. 12. Bergfelt DR, GP Adams GP. Ovulation and corpus luteum development. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 1–13. 13. Gastal EL, Gastal MO, Wiltbank MC, Ginther OJ. Follicle deviation and intrafollicular and systemic estradiol concentrations in mares. Biol Reprod 1999;61:31–9. 14. Garcia MC, Ginther OJ. Regulation of plasma LH by estradiol and progesterone in ovariectomized mares. Biol Reprod 1978;19:447–53. 15. Pattison ML, Chen CL, Kelley ST, Brandt GW. Luteinizing hormone and estradiol in peripheral blood of mares during estrous cycle. Biol Reprod 1974;11:245–50. 16. Alexander SL, Irvine CHG. FSH and LH. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Baltimore: Williams & Wilkins, 1993, pp. 45–56.
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17. Alexander SL, Irvine CHG. Radioimmunoassay and in vitro bioassay of serum LH throughout the equine oestrus cycle. J Reprod Fertil 1982; Suppl 32:253–60. 18. Ginther OJ. Reproductive Biology of the Mare, Basic and Applied Aspects. Cross Plains, WI: Equiservices, 1992; pp. 41–74. 19. Daels PF, Hughes JP. The normal estrous cycle. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Baltimore: Williams & Wilkins, 1993; pp. 121–32. 20. Medan MS, Nambo Y, Nagamine N, Shinbo H, Watanabe G, Groome N, Taya K. Plasma concentrations of Ir-inhibin, Inhibin A, Inhibin pro-αC, FSH, and estradiol-17β during estrous cycle in mares and their relationship with follicular growth. Endocrine 2004;25:7–14. 21. Bergfelt DR, Mann BG, Schwartz NB, Ginther OJ. Circulating concentrations of immunoreactive inhibin and FSH during the estrous cycle of mares. J Equine Vet Sci 1991;11:319–22. 22. Roser JF, McCue PM, Hoye E. Inhibin activity in the mare and stallion. Domest Anim Endocrinol 1994;11:87–100. 23. de Kretser DM, Robertson DM. The isolation and physiology of inhibin and related proteins. Biol Reprod 1989;40:33–47. 24. McCue PM. Pathophysiology of follicular development in the mare. PhD Dissertation, University of California, Davis, 1993; pp. 106–47. 25. Carnevale EM, McKinnon AO, Squires EL, Voss JL. Ultrasonographic characteristics of the preovulatory follicle preceding and during ovulation in mares. J Equine Vet Sci 1988;8(6):428–31. 26. Ginther OJ. Ultrasonic imaging of equine ovarian follicles and corpora lutea. Vet Clin North Am Equine Pract 1988;4:197–213. 27. Townson DH, Ginther OJ. Duration and pattern of follicular evacuation during ovulation in the mare. Anim Reprod Sci 1987;15:131–8. 28. Samper JC, Pycock JF. The normal uterus in estrus. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 32–5.
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Chapter 180
Diestrus ¨ Robert M. Lofstedt
Introduction The term diestrus means, or at least intimates, ‘a period of sexual quiescence between two estrus periods’. But truly, diestrus is anything but quiescent. In fact, the ovaries are very active during diestrus in all mammals. Incidentally, the term diestrus is a noun, the adjective ‘diestrous’ being used to describe events that occur during diestrus. These terms are often misused. The characteristics of diestrus have been eloquently and substantially described elsewhere, especially by O.J. Ginther in his 1992 publication Reproductive Biology of the Mare: Basic and Applied Aspects.1 Therefore, not all aspects of diestrus are discussed here, mostly updates and condensations of current knowledge, especially as they apply to daily events in stud practice.
The first diestrus of the breeding season In equine stud management the first diestrous period of the year is a welcome event after the frustrating transition1 that a mare experiences between anestrus and the onset of her normal estrus cycles. This is described as frustrating because there is nothing practical one can do to hasten the onset of normal estrous cycles once transition has begun. This applies even if transition begins earlier than normal after light treatment has been used. It is true that progress has been made with regard to the induction of early season estrous cycles; first with pulsatile administration of gonadotropin-releasing hormone (GnRH),2 and more recently with the use of dopamine antagonists and prolactin.3 However, until these techniques or others have been refined for practical use, we will still have to wait patiently for the first ovulation of the breeding season. Only then can the mare become productive
on the studfarm. Of course, this does not apply in the case of a foaling mare (a mare due to foal) because these mares show estrus and ovulate within a short time after foaling, irrespective of suckling or season. In our practice, when ultrasonography reveals the typical ovarian features of transition (multiple small to medium-sized follicles in both ovaries and an absence of luteal tissue) we inform mare owners that they should return to the studfarm after a somewhat arbitrary period, usually about 2–3 weeks. After that time, we re-examine these mares for evidence of ovulation, i.e., being in diestrus. The evidence we seek is luteal tissue within either of the ovaries and an absence of the multiple follicles that characterize transition. This is done using ultrasonography. We might also assay a blood sample for an elevated concentration of progesterone if we are uncertain as to the presence of luteal tissue on ultrasonography. A progesterone serum concentration of more than 2–3 ng/mL is taken to indicate the presence of a functional corpus luteum (CL).1 Mare-side progesterone assays are also useful for this purpose.4 During late transition, the first time that uterine edema is noticed on ultrasonography has been found to be an unreliable indicator that the first ovulation of the breeding season is about to occur.5 Nevertheless, uterine edema is useful for predicting ovulation once estrous cycles have been established. If a mare owner has a stallion and is familiar with proper teasing methods, that mare can be teased daily using a stallion before she is brought to the studfarm in springtime. In that regard, behavior will usually change from disinterest or more or less persistent estrus during transition, to a strong negative response to the stallion after an ovulation has occurred.1 When we are sure that mares on our studfarms have begun to have estrous cycles, they are left there for breeding and their ovaries are monitored for the next ovulation of the season.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Diestrus If steps are not taken to ascertain the onset of normal estrous cycles, mares may spend unnecessarily long periods of time on the studfarm, waiting to be bred, incurring boarding and veterinary fees and being exposed to threats of disease. If owners to breed a mare at the first opportunity of the season, minimizing potential delays in the birth date of the foal, the limitations of the transitional period should be explained to them beforehand.
Characteristics of diestrus By definition, a mare in diestrus has a functional CL. Serum progesterone concentrations rise rapidly from about 1–2 ng/mL shortly after ovulation, to maximum concentrations of 8–16 ng/mL when the corpora lutea are mature. Approximately 2 weeks after ovulation, concentrations begin to decline and reach baseline concentrations again at the onset of the next estrous period, about 15 to 16 days after ovulation.1 This serves to remind one that diestrus is short in mares, shorter than in other farm animals despite the fact that a mare’s estrous cycle is 21 to 22 days long. This of course is the result of having a relatively long estrous period (∼ 6 days) compared to other farm animals. The relatively long luteal phases in cows, goats, and even sheep, give one proportionally more time to manipulate the estrous cycles using prostaglandins in those species than in mares, i.e., mares have a luteal phase that is only about half the length of the estrous cycle, which is substantially less than in other farm animals. Other factors also limit the value of prostaglandin treatment in mares (see below). Mares are not usually sexually receptive during diestrus. However, as intuition suggests, and as Ginther has pointed out in his reviews, the interface between diestrus and estrus is not black and white, but shades of gray. For example, mares with high serum progesterone concentrations may occasionally show overt estrus.1
Shortening diestrus After ovulation, oocytes are fertile for little more than 18 hours.6 Therefore, when an ovulation has been missed due to an oversight, a serious error in management has occurred on the studfarm. The error is serious because post-ovulation breeding frequently fails, especially when more than a day has elapsed since ovulation. In addition, the next ovulation is often more than 2 weeks away (see below). Estrous cycles cannot be monitored closely enough to predict the exact time of ovulation, especially if a studfarm is only visited every two or three days. Therefore, when one is in doubt as to the possibility of ovulation in the
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next 48 hours, it is best to breed the mare when possible. As mentioned earlier, the next ovulation after a missed ovulation will often occur up to 3 weeks later, and a foal arising from that breeding or the next, may be noticeably smaller that its cohorts in the weanling sales ring, commanding a lower price as a result. Missed ovulations are also important when one is facing the end of the breeding season after numerous fruitless inseminations. At a certain point, in fact, late season breeding is no longer feasible. For these reasons the stud veterinarian often attempts to shortcycle mares after missed ovulations as is done with cows. Prostaglandins are often used to “short-cycle” mares. Although the next ovulation will sometimes occur sooner than normal, it often does not. This is because there is a major difference in follicle growth patterns between mares and cows, depriving exogenous prostaglandins of much of the utility they have in cattle. Cattle usually have two or three waves of follicle development during their estrous cycles7 and, occasionally, even four (M.C. Wiltbank, personal communication). When luteolysis occurs (naturally or pharmacologically) serum progesterone concentrations decline and serum concentrations of LH increase. At that time, a dominant follicle within one of the follicle waves is recruited and begins to mature. Then it induces a surge of LH and it ovulates. Therefore the estrous cycle can be shortened significantly by using prostaglandin treatment in cattle. By contrast, most mares and ponies only have a single follicle wave during their estrous cycles.1,8–11 Therefore a dominant follicle is not always available for ovulation at the time of prostaglandin treatment and the interovulatory interval cannot always be shortened by using prostaglandin treatment in mares. A dominant follicle(s) can only be recruited from a cohort of growing follicles; it cannot materialize quickly from a group of dormant follicles due to prostaglandin treatment, irrespective of the absence or presence of a corpus luteum. In mares with a single follicle wave during their estrous cycles, the follicle destined to ovulate can be first identified on ultrasound about 10 days before ovulation. Its diameter increases by approximately 3 mm a day, occasionally slowing in growth for about 48 hours until it ovulates late in estrus.1 Interestingly, it is only at this time, i.e., at the end of its growth phase, that progesterone can have had a significant effect on the equine follicle, by stopping its maturation and preventing ovulation. Therefore the presence or absence (due to treatment) of a CL is of little importance in follicle development. Together with the usual phenomenon of a single follicle wave, this explains
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why prostaglandin-induced luteolysis have no effect in hastening the time of ovulation in mares. For example, in one study when prostaglandin was given to mares on day 7, 10, or 13 of the estrous cycle, estrus began within 3 or 4 days in all cases but ovulation still occurred between 18 and 24 days, not shortening the mean length of their interovulatory intervals.12 Despite this limitation, prostaglandins are sometimes effective in shortening the interestrus interval. For example, in a group of 28 Thoroughbred mares treated with prostaglandin 6 days after ovulation, 85% ovulated again between 12 and 16 days after treatment, i.e., about a week sooner than normal. Similar results were seen in Appaloosa and Quarter Horse mares treated early in the luteal phase.13, 14 In fact, even before prostaglandins became widely used in horses, it was noticed that the interovulatory interval was shortened in some mares with endometritis or after intrauterine saline infusions; yet this did not occur in others.1,8–10 The reason for the reduced interovulatory interval in some mares appears to be another follicle wave that occurs during early diestrus, a situation comparable to that in cows. In this regard, Thoroughbreds, Trotting Standardbreds, and Appaloosas may be more likely to have estrous cycles with two follicle waves, while this is less likely in Pacing Standardbreds, pony breeds, Quarter Horses, and other breeds.1, 15 Therefore, within individual mares, and probably even within the months of the breeding season, the character of follicle waves will predict responses to prostaglandin treatment.
When to use prostaglandins during diestrus Prostaglandin analogs can cause luteolysis in some mares as early as 1 or 2 days after ovulation but luteolysis only becomes predictable from about 4 days onward.13, 14, 16 Therefore, when one is attempting to “short-cycle” a mare, prostaglandin treatment should probably be administered as early as possible in the luteal phase; no later than 6 days after ovulation. Otherwise, a follicle may not be recruited from an early diestrus follicle wave (when present). This is evident from the study on Thoroughbreds mentioned in the previous paragraph, where the interovulatory period was shortened when treatment occurred at 6 days after ovulation. This did not occur in mares treated between 6 and 8 days after ovulation. Importantly, a secondary wave will not yet be obvious on ultrasonography early in the estrous cycle, therefore one can only proceed with treatment on the assumption that a second follicle wave will indeed occur.
Controlling follicle development during the diestrous period Progesterone has only minor effects on the growth rate of follicles.1 We see this with ultrasound in dayto-day stud practice. For example, large follicles are often seen during early pregnancy when serum progesterone concentrations are high. Some mares even ovulate during diestrus.17 Follicles also grow and ovulate during pregnancy when serum progesterone concentrations are also high. This explains why neither altrenogest nor high doses of progesterone are able to inhibit follicle development, i.e., to control follicle dynamics. After treatments in either case, a wide range of follicles sizes is present and again, some mares even ovulate during treatment. We have seen this in practice and it is also the subject of several studies.11,18–20 Therefore one should not rely on progesterone or other progestogens to control the estrous cycles of mares. Although progesterone cannot control follicle dynamics effectively for management, estradiol can. As reviewed by Kelley et al.,3 estradiol is able to suppress follicle-stimulating hormone (FSH) secretion and, therefore, to suppress follicle development as well. This substantiates the findings of Loy and his colleagues when they began to examine the use of progesterone and estradiol for the control of follicle growth.21 Their studies led to the widespread use of a combination of progesterone and unconjugated 17β-estradiol (colloquially called P&E) for controlling the time of ovulation in mares. This combination of steroids is administered to mares daily for about 10 days, during which time ovarian follicular development is significantly suppressed. At the end of treatment, follicles are seldom larger than 18–22 mm in diameter (R. Lofstedt, unpublished observations). Interestingly, the continued growth of large pre-existing follicles is not impeded during progesterone/estradiol (P&E) treatment, presumably because they have bound enough FSH to develop to completion. In fact, some of these follicles may ovulate during P&E treatment (R.G. Loy, personal communication). Once again, this demonstrates the lack of control that progesterone has over follicle development. Because ovulation can indeed occur during P&E treatment, mares are treated with prostaglandins on the last day of P&E treatment. In the event that ovulation does occur late in P&E treatment, prostaglandin treatment should probably be delayed for a few days after the P&E treatment in all cases but this is not common practice. Also, it is quite likely that estradiol alone would work as well as P&E for follicle control in mares. These subjects remain to be studied.
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Diestrus After P&E treatment ends, follicle development progresses at approximately the same rate as it does in a normal estrous cycle. If human chorionic gonadotropin is given 8 days after the end of treatment, ovulation is extremely predictable, usually occurring on the morning of day 9 after the end of P&E treatment.
Synchronization of estrous cycles using two injections of prostaglandin As stated earlier, the interovulatory period can only be manipulated effectively when the mares being treated have two follicle waves during each estrous cycle. Therefore, if double injection techniques (similar to those used in cattle) are to be used in mares, the breed of horse may be important. Although the nature of follicle waves in mares seriously limits the value of double injection techniques in this species, the subject has received no attention.
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References 1. Ginther OJ. Sexual behavior (Chapter 3); Characteristics of the ovulatory season (Chapter 6). In: Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Wisconsin: Equiservices Publishing, 1992. 2. Becker SE, Johnson AL. Effects of gonadotropin-releasing hormone infused in a pulsatile or continuous fashion on serum gonadotropin concentrations and ovulation in the mare. J Anim Sci 1992;70:1208–15. 3. Kelley KK, Thompson DL, Storer WA, Mitcham PB, Gilley RM, Burns PJ. Estradiol interactions with dopamine antagonists in mares: prolactin secretion and reproductive traits J Equine Vet Sci 2006;26:517–28. 4. Relave R, LeFebre RC, Beaudoin S, Price C. Accuracy of a rapid enzyme-linked immunosorbent assay to measure progesterone in mares. Can Vet J 2007;48:823–6. 5. Watson ED, Thomassen R, Nikolakopoulos E. Association of uterine edema with follicle waves around the onset of the breeding season in pony mares. Theriogenology 2002;59:1181–7. 6. Woods J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic-loss rate in mares. Equine Vet J 1990;22:410–15. 7. Burns SD, Jimenez-Krassel F, Ireland JLH, Knight PG, Ireland JJ. Numbers of antral follicles during follicular waves in cattle: evidence for high variation among animals, very
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high repeatability in individuals, and an inverse association with serum follicle-stimulating hormone concentrations. Biol Reprod 2005;73:54–62 Loy RG, Buel JR 1st, Stevenson W, Hamm D. Sources of variation in response intervals after prostaglandin treatment in mares with functional corpora lutea. J Reprod Fertil Suppl 1979;27:229–35. Ginther OJ. Major and minor follicular waves during the equine oestrous cycle. J Equine Vet Sci 1993;13:18–25. Ginther OJ. Selection of the dominant follicle in cattle and horse. In: 14th International Congress on Animal Reproduction, Stockholm, Sweden, 2000, pp. 60–79. ¨ Handler J, Schonlieba S, Hoppenc H-O, Aurichb C. Seasonal effects on attempts to synchronize estrus and ovulation by intravaginal application of progesterone-releasing device (PRIDJ) in mares. Theriogenology 2006;65:1145– 58. Douglas RH, Ginther OJ. Effects of prostaglandin F2 alpha on the estrous cycle and pregnancy in mares. J Reprod Fertil Suppl 1975;23:257–61. Douglas RH, Ginther OJ. Effects of prostaglandin F2 alpha on length of diestrus in mares. Prostaglandins 1972;2:265–8. Brendemuehl JP. Effect of oxytocin and PGF2 alpha on luteal formation, function, and pregnancy rates in mares. 47th Proc Annu Conv Am Assoc Equine Pract 2001; 239–41. Shirazi A, Gharagozloo F, Niasari-Naslaji A, Bolourchi M. Ovarian follicular dynamics in Caspian mares. Equine Vet J 2002;22:208–11. Bergfelt DR, Pierson RA, Ginther OJ. Regression and resurgence of the CL following PGF2α treatment 3 days after ovulation in mares. Theriogenology 2006;65:1605–19. Hughes JP, Couto MA, Stabenfeldt GH. Luteal phase ovulations: What are the options? In: Proceedings of the Annual Meeting of the Society of Theriogenology 1985, pp. 123–5. Holtan DW, Douglas RH, Ginther OJ. Estrus ovulation and conception following synchronization with progesterone, prostaglandin F-2 alpha and human chorionic gonadotropin in pony mares. J Anim Sci 1977;44:431–7. ¨ Lofstedt RM, Patel JH. Evaluation of the ability of altrenogest to control the equine estrous cycle. J Am Vet Assoc 1989;194:361–4 Daels PF, McCue PM, DeMoraes MJ, Hughes JP. Persistence of the luteal phase following ovulation during altrenogest treatment in mares. Theriogenology 1996;46:799– 811. Taylor TB, Pemstein P, Loy RG. Control of ovulation in mares in the early breeding season with ovarian steroids and prostaglandin. J Reprod Fertil Suppl 1982;32:219–24. Palmer E, Jousset B. Synchronization of oestrus in mares with a prostaglandin analogue and HCG. J Reprod Fertil Suppl 1975;23:269–73. Hyland JH, Bristol F. Synchronization of oestus and timed insemination of mares. J Reprod Fertil Suppl 1979;27:251–5.
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Chapter 181
Autumnal Transition Out of the Breeding Season Sheryl S. King
The autumnal transition from estrous cyclicity to anestrus is an intriguing process that does not appear to be a simple reversal of the anestrus-to-cyclicity spring transitional process. During the spring transition, the hypothalamic–pituitary axis plays a major controlling role. Detailed examination of the endocrine and ovarian events associated with the autumnal transition indicates that the ovary rather than the brain is often the first responder to the seasonal change, and that the mare’s recognition and response to the change in season begins quite early – shortly after the summer solstice. The fall transition into anestrus is both longer and appears to be less under endocrine control than the vernal transition into cyclicity. In fact, a significant number of mares fail to transit into acyclicity and continue ovulatory estrous cycles throughout the winter. For the purposes of this chapter, anestrus is defined as that period during which the mare’s ovary is structurally and hormonally quiescent. That is, the time during which follicles do not grow to > 20 mm in diameter, circulating estradiol remains at basal concentrations, there is no active luteal structure, and circulating progesterone concentrations are consistently < 1 ng/mL. Care should be taken when reading the existing literature concerning the fall transition to understand the definition being used by different authors to describe reproductive states during this period. A number of researchers choose the time of the last ovulation of the season as the end point to the autumnal transition, and other events are plotted against this reference point. However, as we will see in this chapter, the final ovulation of the year often does not mark the end of ovarian structural or hormonal activity and cannot be considered the beginning of anestrus. Thus, a mare can be, and for a period of time often is, anovulatory without being
anestrus. In fact, some mares never achieve anestrus by this definition, but experience only an anovulatory period of variable length with persistent but anovulatory follicular growth followed by recrudescence and the first ovulation of the next season.1, 2 Events characterizing the autumnal transition include a number of reproductive anomalies, including spontaneously prolonged luteal function, abbreviated luteal function, anovulatory follicular growth, hemorrhagic (‘autumn’) follicles, and silent as well as inappropriate estrus, that are seen only rarely during the physiological breeding season. In fact, in one study,3 over 60% of the cycles which occurred after the autumnal equinox demonstrated behavioral, ovarian, and/or hormonal events different from those typical of the summer breeding season. The relatively frequent occurrence of these anomalous events during the autumnal transition presents an additional set of challenges to practitioners treating reproductive conditions – particularly if they are relying on repeated estrous cycles as an ally in treatment – and for horse breeders that are covering mares during this season. Additionally, we have found that some of the common ‘remedies’ that are used for amelioration of these anomalies during the spring, such as human chorionic gonadotropin (hCG), exogenous progesterone, and dopamine antagonists, are not effective when used in the fall (S. King, unpublished observations). Scientists should be cognizant of the increasing asynchrony of reproductive events as well as the changing endocrinology of this period that begins as early as late summer and to take this into account when designing experiments, particularly when those studies rely upon ‘normal’ cyclicity to persist into August and September (early autumn in the northern hemisphere).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Autumnal Transition Out of the Breeding Season Further research into this period of seasonal equine reproductive function is needed. Discoveries will certainly help us to understand the complex and unique interactions between the environment and the hypothalamic–pituitary–gonadal axis in the mare. In addition, and perhaps as importantly on a practical level, a thorough understanding of fall transitional events could well lead to a method whereby producers can maintain reproductive cyclicity in mares yearround, thus providing a more reliable way to have mares cycling and ready for conception early in the year.
Control over the fall transition Photoperiod Horses appear to have an innate circannual reproductive rhythm. When photoperiodic cues were removed through a full year of exposure to long days (15.5 hours light/day), horses continued to demonstrate a season of reproductive activity followed by a season of reproductive quiescence,4 although this seasonal pattern did not conform exactly to the pattern adopted by mares under natural photoperiod. Superimposed on their innate circannual rhythm, mares use environmental cues to refine their reproductive periodicity to conform to the climate they occupy. The primary zeitgeber used to entrain this rhythm to the environment is photoperiod. Since it appears that horses would likely be seasonal breeders without photoperiodic cues, teasing out the effects of photoperiod from the horse’s natural tendencies is difficult, particularly where this pertains to the autumnal transition wherein typical photoperiodic mediators such as melatonin appear to be ignored by the mare. Although changes in photoperiodicity can reliably alter the timing of the spring transition in the majority of mares,5 the level of influence that photoperiod has over the fall transition is much less clear. From the limited work that has been accomplished to date, it appears that the process of transition into acyclicity is less strongly entrained to photoperiodic signals. Although many references state that the autumnal transition occurs between September and November, careful examination of the limited number of studies conducted during this period indicates that anestrus occurs anywhere between August and February6–10 and that the autumnal transition is often prolonged well past the winter solstice (December 21 in the northern hemisphere). This puts the mare in the curious situation of reproductive decline during increasing photoperiod. In one study, the time of entry into
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anestrus was equally distributed between November (31%), December (25%), and January (37.5%), with one mare completing her transition in February.7 This makes the duration of the fall transition twice as long as the spring transition. Much of the descriptive work defining the events of the autumnal transition has been conducted at temperate latitudes in the northern and southern hemispheres (Southern Illinois, Central Kentucky ∼38◦ N; Southern Wisconsin, 43◦ N; Central Hungary, 47◦ N; South-Central New Zealand, 43.6◦ S), where photoperiodic cues are rather distinct. Differences between maximum and minimum hours of daylight in these regions are 6–8 hours. Information available from studies conducted in torrid zones (Mexico, 15–22◦ N; Venezuela, 10◦ N) indicates that, although horses near the equator demonstrate seasonality, the characteristics of the transition between reproductive states varies from those observed at higher latitudes. It is difficult to determine whether horses are responding to the very modest fluctuation in daylength (1 hour, 15 minutes excursion between winter and summer solstice) at these equatorial latitudes or whether they are simply responding to their own innate circannual clock. Kooistra and Ginther4 showed that ponies exposed to stimulatory (15.5 hours light/day) photoperiods year round experienced seasonal reproductive intervals no different from ambient-lighted controls in Wisconsin. However, when pony mares were exposed to this maximum daylength only from the summer (beginning June 22, July 1, or August 1) through the fall and winter, they experienced a delay in the autumnal transition plus an unusually high incidence (29% average in two studies) of persistent winter cyclicity.4, 11 It appears that pony mares can escape from photoperiodic entrainment when there is no seasonal variation in daylength. Since horses are less seasonal than ponies, one might assume an even higher degree of photoperiodic independence. Causes for failure of an autumnal transition, resulting in persistent winter cyclicity, remain undefined. This rarely happens in ponies, but many studies have reported a variable percentage of horse mares that do not transition into anestrus in the fall.7, 8, 12, 13 The degree of fluctuation in photoperiod between seasons appears to have an influence on the percentage of mares that recognize and respond to the oncoming winter season. Review of data for horse mares monitored over the entire fall transition and winter anestrous periods (to account for late transition past the winter solstice) resulted in an average winter cyclicity incidence of 27% at latitudes between 30◦ and 50◦ ,3, 10, 14 while on average over 55% of mares between 10◦ and 30◦ latitude did not experience anestrus.2,15–18
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Environmental factors, nutrition, and management The horse remains largely tied to its evolutionary heritage, which includes intrinsic systems designed to maximize its chances of survival during harsh periods (e.g., winter) by minimizing physical demands on its body extraneous to its immediate survival (e.g., breeding). It should not be surprising, therefore, to find that the horse responds to its short-term environment with modifications in its longer term patterns. Modifiers including temperature, age, nutrition, and stress can influence reproductive seasonality, and scientists need to understand how they operate in order to help maximize optimal management conditions for horse breeders. Unseasonal weather is a stressor for the horse, and like most stressors is likely mediated through autonomic nervous signaling pathways involving the sympathetic nervous system and possibly the opioidergic system. The sympathetic neurotransmitter dopamine has been found to play a role in seasonal reproduction in the horse.7,19–23 Although most investigations have involved dopaminergic action on stimulating early cyclicity following anestrus, one study126 reported results from administration of a dopamine antagonist, domperidone, throughout the autumnal transition. Domperidone treatments did not delay the fall transition; however, it tended to increase the incidence of winter cyclicity and improved luteolytic function. No significant changes in other endocrine patterns (progesterone, prolactin) were noted. Likewise, endogenous opioid peptides are known mediators of mental, physical, and environmental stresses. Opioids appear to act directly on the hypothalamus to decrease gonadotropin-releasing hormone (GnRH) release in other species, resulting in a decline in luteinizing hormone (LH) secretion.24 Endogenous opioid peptides may play a role in the development of seasonal acyclicity in mares insofar as the level of opioid inhibition of the gonadotropin axis was higher during anestrus than during cyclicity.25, 26 Endogenous opioids also increase the release of the catecholamines, dopamine, and norepinephrine.27 Evidence suggests that dopamine has a direct action on the ovary,23, 28, 52, 96 and not on the hypothalamic–pituitary axis,20, 29 although the specific nature of this action is as yet undefined. Taken together, negative environmental stimuli (temperature, nutrition, other stressors) could suppress reproduction centrally through depression of gonadotropins via the opioidergic system and at the ovaries through the dopaminergic system. Hopefully, the nature of the feedback between environment and
the reproductive system in the horse will be the subject of future research. Factors such as the amount of body fat stores a horse possesses in preparation for the upcoming winter may alter the timing of the autumnal transition. Mares entering the fall season in high body condition have a higher incidence of autumnal transitional failure and persistent winter cyclicity.1,30–32 The adipose tissue hormone leptin is thought to be the primary signaler of body condition to the brain. High leptin concentrations are associated with high body fat stores. Circulating leptin displays a seasonal pattern in mares, with the lowest levels occurring in winter, despite good body condition.1, 30 Leptin and body condition have been implicated as factors in the control of time of transition into anestrus and in persistent winter cyclicity. Leptin concentrations decreased during the autumnal transition and were lower in mares that achieved anestrus than in those with persistent winter cyclicity.32 Studies designed to modify circulating leptin concentrations in mares to either provoke or prevent winter acyclicity have met with ambiguous results. Manipulation of body fat during the autumn months through feed restriction33 or pharmacological treatments33, 34 led to decreased leptin concentrations in treated mares in all studies; whether this translated into persistent winter cyclicity, however, differed between studies. There was no change in the timecourse of the fall transition or the proportion of mares that continued estrous cyclicity during the winter in the two Kentucky studies.34, 35 However in one of these studies,34 it was noted that both treated and control groups lost weight during the experimental period, which could have affected the results. The Louisiana group1 avoided this problem and demonstrated a difference in leptin concentrations as well as a higher rate of winter reproductive activity (i.e., failure of transition to anestrus) between mares of high compared with low body condition. Administration of exogenous recombinant equine leptin resulted in weight loss in mares but no change in seasonal reproductive patterns,36 suggesting that leptin does not have direct antigonadal effects. Leptin appears to modulate body condition and may therefore have an indirect effect on seasonal reproduction. Mature horse mares do not reliably enter anestrus; however, young mares (2–3 years) achieve anestrus at or near a 100% rate.12, 37 Young (< 5 years old) mares were also found to complete the transition into anestrus earlier than adult mares.30, 38, 39 Although this may be an effect of age and a relative immaturity in the reproductive axis, differences in body weight and body condition could also be a controlling factor. Young mares generally have lower body fat, and
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Autumnal Transition Out of the Breeding Season therefore lower leptin concentrations. Young mares also do not display a strong seasonal variation in leptin concentrations.30 Metabolic state during the breeding season appears to influence the transition into anestrus. Foaling mares, regardless of whether they were still lactating during the late summer and into the fall transition, reliably entered anestrus during their foaling year.12, 37 Fitzgerald et al.35 assert that mares that are left unbred for successive years are the ones most likely to continue estrous cyclicity over the winter. Whether the process of pregnancy, parturition, and/or lactation influences the mare’s sensitivity to seasonal cues is currently unknown. Late pregnancy and lactation certainly affect body condition; therefore, it is reasonable to hypothesize that it is not the maternal condition per se, but rather the negative energy balance engendered by this condition that influences the transition into anestrus. In a study of melatonin effects on transition and anestrus, Guillaume et al.37 reported weight losses between –0.44 and –0.25 kg/day in their groups of 17 foaling draft mares. Although the extent of individual weight losses could not be correlated with time of onset of transition or anestrus, weight loss could nevertheless be a contributing factor to the achievement of anestrus.
Ovarian events characterizing the fall transition Many equine reproductive physiologists focus intently on the endocrinology of the hypothalamus and pituitary, assuming that it is this endocrine unit that drives all other reproductive activity. This is unfortunate because they tend to overlook other regulatory systems, including the ovary and nerves, or give them only cursory consideration. Particularly when examining the events associated with the autumnal transition into anestrus in the mare, it is important to sort out leading versus following effects; that is, which events come first and drive others. This is especially germane to examination of causeand-effect between hypothalamic–pituitary–ovarian interactions during the autumnal transition. The prelude to achievement of full anestrus (ovarian structural and hormonal inactivity) is elimination of corpora lutea and progesterone, cessation of subsequent estradiol secretion and ovulations, as well as failure of follicular development beyond a rudimentary size (generally considered to be 20–25 mm in diameter). As we will see, the sequence of these events can be changed depending on the individual mare. Cessation of ovarian function is not abrupt – at minimum it appears to take 60 days or more for the transition into anestrus to occur. The time at which transition occurs and the mode through which anestrus is
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achieved varies – individual mares do not experience the same mode of transit out of cyclicity from year to year, and are not consistent as to the time of transition between years (S. King, unpublished observation). Some transitional ovarian events are seen quite predictably (e.g., progesterone decline), others more frequently in particular mares (e.g., prolonged corpus luteum, hemorrhagic anovulatory follicles), and still other anomalies appear to have a random distribution.
Corpus luteum One of the obvious differences between the fall transition out of cyclicity and the spring transition into cyclicity is that luteal structures are present only during the former. Luteal function is the first change to be noted with seasonal advancement. Both luteotropic and luteolytic function evidence decreased efficiency and an increased incidence of either partial or full failure. Luteal steroidogenesis Early during the autumnal transition, mares begin to demonstrate subtle changes in their luteal activity. From the estrous cycle following the summer solstice and throughout subsequent estrous cycles, the average progesterone concentration during the luteal phase decreases in unbred mares.40 Possible effects of pregnancy on this phenomenon have not been studied. A number of studies have documented this phenomenon,7, 9, 14, 40, 41 and appear to differ in only one aspect; one study40 detected a decline in average progesterone only in mares destined to become acyclic during that winter (14 of 20) and not in mares who remained cyclic year-round (6 of 20), whereas two later studies7, 14 found no difference between these two groups. Average luteal phase progesterone production declines in a linear or quadratic fashion40 throughout the late summer and fall. Although the steroidogenic capacity of the late-season corpora lutea (CLs) appears to decline by approximately 50% compared with summer CLs, complete failure of luteal development (defined as an ovulation without subsequent CL formation and progesterone remaining ≤ 1 ng/mL) during the autumn has been observed on only one occasion (S. King, unpublished observation; Figure 181.1). The cause for this decline in steroidogenic capacity of the CL in relation to season is unknown. Ultrasonography revealed no differences in the size of the CLs produced or in the echogenic structure between autumn CLs with decreased progesterone production and spring/summer CLs.7 Macroscopic inspection of autumn CLs from abattoir-collected ovaries revealed no noticeable differences compared with CLs during
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Figure 181.1 Failure of luteal development following the collapse (presumed ovulation) of an ovulatory-sized follicle. Note the elongated follicular growth phase during the final two cycles. This mare demonstrated a gradual reduction in LH over two cycles.9
spring and summer (S. King, unpublished observation). This suggests that the change in CL steroid production is not due to differences in the proliferation of luteal cells within the CL. Histological comparison of autumn CLs with those typical of summer is required to confirm this, however. Diminution of luteotropic support with advancement of the season could explain the progesterone decline. Unfortunately, luteotropic factors for the mare remain largely undefined. It has been suggested that one factor contributing to initial luteal development, if not its sustenance, is LH.5, 42 However, waning progesterone is established far in advance of detectible declines in circulating LH concentrations.9 Hart et al.43 reported that LH content of the pituitary gland decreased between the middle of the physiological breeding season in July and the time of the autumnal transition in October, to reach its lowest concentrations during anestrus in December and into March. Once again, the decline in luteal phase progesterone initiated in July or August appears to precede this pituitary event. Additionally, the decline in pituitary LH content was not reflected in a corresponding decrease in circulating LH secretion, which remained unchanged during the late summer and up to the third-to-last estrous cycle in the fall.9 A comprehensive study of LH receptor populations on the CL throughout the entire ovulatory season, and including the autumnal transition, has not been reported; therefore, possible seasonal changes in the CL’s ability to respond to LH through its receptor are unknown. Prolactin is known to be luteotropic in some species,44 but this role has yet to be studied in the
mare. The seasonal decline in prolactin concentrations45 (see Prolactin below) temporally mimics the seasonal decline in progesterone. Moreover, prolactin receptors have been identified on the equine CL.46 The possibility that changes in luteal progesterone production can be influenced through prolactin in the mare deserves further study. Luteal lifespan Luteal lifespan is also affected during the autumnal transition. Weedman et al.3 reported a high incidence (29%) of abnormally abbreviated luteal periods (12.0 ± 0.8 days in autumn versus 14.8 ± 0.3 days during summer), particularly approaching the final luteal phase of the autumnal transition. Some periods of progesterone secretion were extremely abbreviated, lasting only 1 or 2 days. In one study, there was one incidence of what we termed a ‘collapsed’ follicle;9 that is, an ovulatory-sized follicle disappeared from one consecutive ultrasound day to the next, yet an ultrasonographically detectible CL did not develop and progesterone concentrations did not increase following this event (Figure 181.1). We speculate that this constitutes failure of luteal development following ovulation (S. King, unpublished observation); however, further documentation is required to distinguish this phenomenon from a single aberrant episode. Irvine et al.41 also documented a progressive, incremental shortening in average luteal phase length over the final three luteal periods in the fall (11.4 ± 0.8 days third-last; 9.5 ± 0.4 days secondlast; 8.5 ± 0.8 days last luteal period). A likely causative candidate for this phenomenon could be the diminution of luteotropic factors,
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Autumnal Transition Out of the Breeding Season such as LH, that is known to occur during this period9, 41, 47 and/or other factors as yet unknown. Not only was the mean peak value of the LH surge decreased over the last two or more cycles prior to anovulation,9, 41, 47 Irvine et al.41 reported a shortened period for the LH surge during the final ovulatory cycle of the year. The persistence of LH into the luteal phase is thought to contribute luteotropic support. Loss of luteotropic support could result in a nonviable CL of shortened lifespan. If this is so, however, it is not reflected in decreased progesterone production capacity in short-lived CLs. Although both types of autumn CLs produced less progesterone than summer CLs, circulating progesterone concentrations between shortened and normal length luteal phases (3.8 ± 0.8 and 5.3 ± 0.4 ng/mL, respectively) did not;3 the possible effects of pregnancy have not been studied. Luteolysis is negatively affected by the autumnal transition, resulting in prolongation of the luteal phase. Numerous studies3,7–9,14, 17, 18, 48 report an increased incidence of spontaneously prolonged corpus luteum (SPCL) activity during the autumnal transition (20–25% of cycles) compared with the peak of the natural breeding season (8–10% of cycles). In retrospect, even the seminal articles first describing the SPCL condition49–51 generated data during the autumn season (August–November), although the authors were not aware of the import of season on their discovery at the time. Spontaneously prolonged CLs are differentiated from prolonged CLs of different origin [such as a late diestrous ovulation or endometrial prostaglandin (PG) F2α insufficiency] in that they arise from a single ovulatory event during an estrous cycle (i.e., no additional luteal phase ovulations were present) from a reproductively healthy (i.e., no evidence of endometritis), unbred (i.e., no embryonic influence) mare. Autumn SPCLs appear to be similar to summer SPCLs in duration; minimum duration of autumn SPCLs was 55 days,3 similar to the average 63-day duration for prolonged CLs reported by Stabenfeldt et al.49 SPCLs appear to respond to the same seasonal effects as CLs of normal lifespan; average progesterone concentrations from autumn SPCLs were lower than those from summer SPCLs.14 The precipitating cause for SPCLs also appears to be the same between summer and autumn transition. By monitoring a group of mares from mid-summer through the autumnal transition using ultrasonography and frequent blood sampling during the luteolytic period, King et al.14 showed that both summer and autumn transitional SPCLs are created through a failure of the release of PGF2α at the appropriate time for normal luteolysis. The fact that these SPCLs regressed following intrauterine saline infusion implies that SPCLs possess
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receptors for PGF2α and that the ability of the uterus to produce PGF2α upon stimulation appears uncompromised. Concentrations of 13,14-dihydro,15-keto PGF2α (PGFM) measured as an indicator of PGF2α secretion were not different between normal length summer or autumn cycles or in response to saline infusion during the summer or autumn in normal length cycles or SPCL cycles.14 This suggests that the factors influencing the production and release of PGF2α are influenced by seasonal changes in ovarian and/or uterine function. Once again, the mechanism by which PGF2α is controlled in the mare is currently unknown and would be a fertile field of research. One factor that appears to play some role in the differential seasonal incidence of SPCL development is dopamine and/or prolactin. This was discovered as part of an experiment designed to investigate the ability of domperidone, a specific dopamine D2 receptor antagonist, to delay or prevent the fall transition.14 Three groups of mares (control, monthly domperidone treatment for 7 consecutive days once every 30 days, or daily domperidone) were monitored from July through anestrus. Although control mares had a typical (24%) incidence of autumn SPCL activity, domperidone treatment decreased the autumn SPCL incidence to 6.7% and 4.4% in the monthly and daily treatment groups, respectively. This SPCL frequency is more typically seen during the height of the breeding season. The mechanism by which dopamine may act to influence luteolysis is currently unknown. It is possible that dopamine has a direct effect on some aspect of luteal function; in this regard, dopamine D1 and D2 receptors and receptor mRNA have been identified in the equine CLs.28, 52 Dopamine is inversely related to prolactin, and some preliminary evidence53 suggests that prolactin may play some role in the luteolytic process (see Prolactin below).
Folliculogenesis Considerable attention has been devoted recently toward describing the physical changes that occur during receding follicular development in the autumnal transition. Investigations of functional changes in autumn follicles have been fewer. In fact, the first discernable change in folliculogenesis during the autumnal transition involves a functional rather than a structural change in the form of reduced steroidogenic capacity of autumn follicles. Along the path toward anestrus, some interesting follicle-related phenomena occur. Follicular steroidogenesis The steroidogenic capacity of the dominant follicle begins to diminish around the time of the autumn equinox (time of equal day and night) and generally
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Figure 181.2 Entry into anestrus via anovulatory follicular growth waves. Note decreased estrogenic capacity from late transitional follicles. FSH, follicle stimulating hormone; E2, estradiol-17β.9
before changes in follicular growth patterns are noticeable.3, 9 Two studies that followed mares through the entire autumnal transition reported a similar decrease in the correlation between size of the dominant follicle and concomitant circulating estradiol-17β concentrations. Nequin et al.9 reported loss of a positive relation between follicle size and estradiol-17β concentration during the final two cycles prior to anestrus while Weedman et al.3 detected a significant shift in estradiol production by the thirdto-last cycle (Figure 181.2). Irvine et al.41 measured estrone conjugates in fall transitional mares and reported a decline between the second-to-last and last cycle of the season. The precise time of emergence of the decline in estrone conjugate concentrations could not be determined because data were not available to compare values from second-to-last fall cycles with earlier fall or summer cycles. In total, fall transitional follicles possess a lower estradiol-17β producing ability per millimeter diameter compared with summer follicles.3 The decline in follicular estrogenic capacity preceded anovulatory follicular growth waves,3, 9, 41 continued into anovulatory follicular growth waves,2, 9, 41, 54 and could persist well into anestrus. Even small follicles (≤ 12 mm) had lower levels of estradiol during anestrus than follicles of comparable size during the spring or summer.55 Hemorrhagic anovulatory follicles Another ovarian anomaly occurring with greater frequency during the autumnal transition than at other
times of the year is development of large, anovulatory, hemorrhagic (‘autumn’) follicles. The hemorrhagic anovulatory follicle (HAF) was first described clinically by Burkhardt in 1948.56 HAFs have been reported during the spring and the autumn transition periods, although the incidence during the fall is much higher (5–8% versus 23%, respectively).57 The incidence of this problem increases with age, and tends to occur repeatedly in some (44%) mares.58 HAF cycles appear to be infertile.58 The future HAF is recruited with a cohort and grows consistent with the typical follicular growth wave pattern. However, after it becomes the dominant follicle and achieves ovulatory size, the HAF fails to ovulate; instead, its lumen fills with blood. During this process, the HAF often continues to grow, sometimes to large proportions; average peak diameters of approximately 51 and 60 mm have been reported.58, 59 Postmortem inspection of the HAF describes the follicular lumen to be filled with a bloody, liquid-to-gelatinous material.60 Ultrasonic evaluation reveals these follicles developing normally to the point of ovulation, then acquiring echoic specks progressing to fibrin strands within the antrum, followed by filling with echogenic, hemorrhagic material.58, 59 Evaluation with color-Doppler ultrasonography indicates increased vascularization in the wall of follicles destined to become HAFs compared with normal ovulatory follicles.59 It is this hemorrhagic appearance on ultrasonography, likely caused by delayed clotting within the follicle,59 that distinguishes the HAF from a luteinized follicle. Unlike the seasonal
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Autumnal Transition Out of the Breeding Season variation with HAFs, the incidence of luteinized follicles has not been reported to change with season. Most HAFs appear to develop luteal tissue, since they produce circulating progesterone concentrations similar to the normal corpus luteum58, 59, 61 and respond to the luteolytic effect of exogenous PGF2α .58 A small percentage (14%) of spring transition HAFs did not develop luteal tissue, as they maintained circulating progesterone concentrations ≤ 1ng/mL.58 These data were gathered in studies performed on HAFs that developed early in the breeding season; HAFs formed during the autumnal transition may function differently. Although most HAFs appear to be capable of responding to exogenously administered PGF2α , the abnormally prolonged average interovulatory interval of 38.5 ± 15 days reported in one study58 suggests that the normal luteolytic mechanism is not triggered during HAF cycles. Prolonged interovulatory intervals are not always the case, however, as Ginther et al.61 reported a normal interovulatory interval of 22 days in seven cases of HAFs during the height of the breeding season. Perhaps this difference is due to the season of the year in which the HAF was formed. The cause of the HAF phenomenon is unknown, although it is tempting to hypothesize that these follicles fail to ovulate due to inadequate LH concentrations, and the subsequent physical strains within the follicle result in internal hemorrhage. In light of their formation around the time of declining LH in the autumnal transition, this may be the case for the larger proportion of HAFs forming during the autumn transitional period, and needs to be investigated. However, a recent evaluation of events surrounding spring and summer HAFs revealed no difference between LH concentrations associated with HAF follicular phases and normal ovulatory follicular phases.59 Estradiol concentrations in plasma59 and follicular fluid60 were significantly higher when associated with a spring–summer HAF follicle than when associated with a normally ovulating follicle. Whether these endocrine patterns are the same for HAFs formed in the fall and what role elevated estradiol may have in the formation of HAFs deserves further investigation. Follicular growth patterns/waves General follicular growth wave patterns include luteal phase recruitment of a cohort of small (2–5 mm) follicles that grow to medium (< 20 mm) size (minor waves) followed by dominant follicle(s) selection (major waves) concurrent with subordinate follicle regression, continued growth, and, finally, ovulation of the dominant follicle(s) during estrus.62 This general pattern changes little in the autumn until ovu-
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lation failure54 prior to full anestrus. In fact, the first follicular wave in the transition that fails to end in ovulation produces a pre-ovulatory follicle of comparable size to that of the previous, ovulatory wave.63 Slight modifications occur in the form of fewer small (2–5 mm) and large (> 20 mm) follicles in the autumn compared with May–July.64 Following the first anovulatory follicular growth wave, there is a progressive decline in the maximum size of the largest and second largest follicles generated from subsequent follicular growth waves leading to cessation of growth with anestrus.63, 65 Also, there is a slowing of the growth rate of the dominant follicle during the autumnal transition (Figures 181.1 and 181.3). This increases the length of the follicular phase of fall transitional cycles compared with summer cycles.3,9,17,41 The extent of follicular phase elongation increased with each estrous cycle approaching anestrus from the third-to-last ovulatory cycle of the year to the last.3,9 There are conflicting reports as to the size of the dominant follicle at ovulation in autumn estrous cycles. Weedman et al.3 described significantly larger average diameters for autumn ovulations (39–42 mm) than those in the summer (35 mm); however, Nequin et al.9 found no difference in average ovulatory size between seasons and Pierson and Ginther64 reported larger average diameters at ovulation in May–July than in August–October. In an early descriptive study of the mare’s estrous cycle, Hughes et al.66 predicted that mares may enter into anestrus from several different ovarian states. Data gathered from September to the first ovulation in the spring for 2 years7 formed the basis for the conclusion that mares are capable of entering anestrus from three different ovarian states. 1. Following the failed ovulation of a dominant follicle during a follicular phase (progesterone ≤ 1 ng/mL); there may be several of these follicular growth-and-atresia episodes prior to ovarian quiescence Figure 181.2). 2. Following a normal luteal phase wherein lysis of the CL is not followed by significant follicular growth (Figure 181.4). 3. Following a SPCL wherein the eventual decline of progesterone to ≤ 1 ng/mL is the last activity on the ovary (Figure 181.3). These distinctions are important when considering the hormonal milieu under which the mare completes her final act of reproductive transition. The majority (56% or 42%) of mares entered anestrus via follicular failure, while 25% entered following normal luteolysis, and 20% or 33% by termination of a
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Figure 181.3 Entry into anestrus via a spontaneously prolonged corpus luteum (SPCL). No follicle growth > 20 mm followed SPCL resolution. Note prolonged follicular growth phase during the ovulatory cycle. This mare demonstrated a sudden LH decline at the final ovulation of the season. P4, progesterone; LH, luteinizing hormone.9
SPCL.7, 9 The distribution of mares within these groups appears to vary. In two studies by Nunes et al.,2, 63 43% and 66% of mares entered anestrus via the first method, and 57% and 33% via the second method, respectively (no SPCL activity was reported in either study). Pony mares may vary from this pattern in that fewer (33%) appear to experience persistent follicular waves past anovulation and more (66%) experience an abrupt cessation of major folli-
cle development during the final luteal phase of the season.65 Relative to the first method of transition into anestrus, most mares experience a variable period following the final ovulation of the season during which waves of significant (maximum diameter ≥ 25 mm) follicular growth persist. Ginther5 has termed this period the receding phase. Many of these post-ovulatory follicular waves produce large,
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Figure 181.4 Entry into anestrus via a normal luteal phase. Note lack of follicle growth > 25 mm following luteolysis despite persistent follicle-stimulating hormone (FSH) surges. FSH, follicle stimulating hormone; P4, progesterone.9
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Autumnal Transition Out of the Breeding Season pre-ovulatory-sized follicles that fail to ovulate and subsequently regress. A series of studies in which follicle ablation was used to elucidate the growth patterns of major and minor follicular waves determined that, in many cases, major and minor follicular waves continue to be generated past the time of the final ovulation of the season, and may then progress into the generation of a series of only minor waves.67, 68 Minor follicular waves may continue during anestrus; however, these follicles appear to have no endocrinological consequence.54, 69
Ovarian size As mares enter their anestrous period and ovarian function declines, the overall size of the ovary often decreases as well.70 The greatest changes in ovarian size and weight appear to occur during the final stages of, or even following achievement of, anestrus. Secondary to a study of seasonal hypothalamic function, Hart et al.43 reported ovarian weight decline between July and October, but a greater decline occurred between October and December. Equine practitioners rarely palpate ovaries during the autumn decline in reproductive function, but most are quite familiar with the unwelcome feel of the small, hard, inactive ovaries characteristic of deep winter anestrus when evaluating mares for early spring breeding.
Endocrine events characterizing the autumnal transition The autumnal seasonal transition provides us with a fascinating model for exceptions to endocrinological dogma. As we have seen, the ovary appears to respond to seasonal cues remarkably early in the physiological breeding season. The first and most striking characteristic of the autumnal transition to anestrus is that ovarian-derived endocrine changes appear to precede pituitary endocrine changes. Alteration in ovarian steroid production is apparent weeks, and in some cases months, before changes in gonadotropin secretion are detected. Moreover, complete cessation of some gonadotropin secretion is never achieved; LH secretion declines during the autumn transition and is eventually extinguished in anestrus, but follicle-stimulating hormone (FSH) secretion persists. Curiously, follicular development beyond a very rudimentary stage ceases despite persistent FSH secretion. Even more perplexing, it appears that the gonadotropin changes that do occur during the autumnal transition are not directly mediated by GnRH. Other hormones of anterior pituitary origin, namely prolactin, demonstrate a far more predictable
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seasonal effect than the gonadotropins. Although it appears to be somehow linked to follicular growth during the spring transition, prolactin does not have the same effect in the fall. The influence of melatonin on all of these factors continues to elude researchers. This begs the question, if the ovarian response appears to be independent of known pituitary signals, by what mechanism does the ovary sense seasonal change? Perhaps the autumnal transition involves a reversal of pituitary–gonadal communication wherein the decline in gonadal function drives a reduction in positive feedback to the pituitary, subsequently affecting such hormones as LH and prolactin. The ensuing reduction in these pituitary hormones might further reduce and finally eliminate ovarian function. Perhaps the early seasonal message to the ovaries is neuronal in origin. Little is known about control of the equine ovary through autonomic innervation, although this is an area of substantial research in other species. The number of these seemingly anomalous endocrine events lends a degree of fascination to the study of the autumn transitional period, and, if for no other reason, should stimulate further research out of pure curiosity.
Gonadotropin-releasing hormone The precise method by which GnRH signals the pituitary to secrete gonadotropins in the mare is still a subject of conjecture. We know even less about possible seasonal responses in GnRH production and secretion. A common assumption has been that seasonal factors, presumably photoperiod, act to diminish GnRH production and secretion during the autumnal transition, to eventually attain a sufficiently low state that it ceases to stimulate pituitary gonadotrophs, resulting in anestrus. A consequent decline in LH, and possibly FSH, would be the end result. The problem with this widely held hypothesis is that it is contradicted by the body of data that is available. The first investigations of seasonal GnRH activity involved postmortem assay of hypothalamic GnRH content. Hart et al.43 reported no change in GnRH content during the time of the autumnal transition between July and October. Most mares are well into their fall transition by October, both progesterone and estradiol concentrations have declined by this period, and many mares have experienced a sufficient decline in LH to complete their final ovulation of the year.9 If GnRH drives these events, it should have demonstrated a decline by this time. GnRH content did decline between October and December.43 Since the recession in reproductive activity would have
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occurred prior to this reduction in GnRH, care should be taken about interpreting the GnRH decline as a cause of winter acyclicity. Interestingly, a subsequent study71 using the same methods and conducted by the same laboratory group reported no difference in hypothalamic GnRH content or distribution between summer cycling and anestrous mares. Likewise, no significant difference in hypothalamic GnRH concentrations between summer and winter was found in long-term ovariectomized pony mares.72 This contradiction between results illustrates the confusing nature of neuroendocrine study in the mare. The idea assumed by some equine reproductive physiologists is that horses are similar to other seasonal species in that seasonal GnRH signals are encoded by the frequency and amplitude with which GnRH is released from the hypothalamus. This has not been proven in the mare. A basic pulsatile pattern of GnRH secretion was demonstrated in the cycling mare48, 73 and GnRH pulse frequency changed with stage of the estrous cycle.73 Presumably, this means that ovarian steroids can alter GnRH secretion, a fact that has been established in other species74 but not in the horse. There remains a question as to whether the anterior pituitary responds to a pulsatile GnRH signal, and also whether that signal is altered by season. The fact that long-term, non-pulsatile administration of GnRH, given in several forms, during anestrus75–77 can elicit a LH response in the same manner as endogenous, pulsatile GnRH does during cyclicity suggests that equine pituitary gonadotrophs can respond to a non-pulsatile signal. Continuous infusion of GnRH during cyclicity also did not disrupt cyclic events,78 indicating that the pituitary is not desensitized by constant exposure to a nonpulsatile GnRH message. A complete picture is far from clear; for example, high doses (10–20 mg) of the synthetic GnRH analog deslorelin acetate can arrest cyclicity (A.O. McKinnon, unpublished observations) and gonadotropin concentrations can be negatively affected79 following deslorelin implantation. On balance, the evidence does call the physiological significance of GnRH pulsatility into question. Direct measurement of GnRH pulses in vivo has been hampered by the difficulty in accessing the hypothalamic effluent, particularly for repeated measurement of the same animal under different experimental conditions. The development of research techniques including cannulation of the intercavernous sinus80 and push–pull perfusion of the hypothalamic region81 has given us some information on GnRH production and secretion in mares during all seasonal phases. Until recently, however, repeated measures on the same animals using these techniques have not been possible. This greatly limits the impact of data gathered from limited numbers of horses, as horses
are notorious for their individual endocrine variability. The picture that emerges from the small number of in vivo experiments is far from clear. The most compelling work suggests that GnRH does not control seasonal reproduction in the mare, at least not in a manner similar to its action in other seasonal species. Use of push–pull brain perfusion allowed for one-time (November–December) in vivo sampling of small numbers of pony mares.81 Although autumn transitional mares were not studied (September–November), hypothalamic secretion of GnRH was significantly lower during anestrus than during cyclicity, suggesting that fall would be the time of hypothalamic conversion from active secretion to relative quiescence. Extremely large variations in basal GnRH secretion appeared to exist between mares, making evaluation of data based on only three individuals used in this study very tenuous. These results conflict with a recent, well-executed experiment utilizing direct pituitary measurements in vivo through sampling from the intercavernous sinus; this generated concurrent measures of GnRH, LH (lutelizing hormone), and FSH from the same seven mares during all four seasons (fall and spring transitions, ovulatory season, and anestrus). The results of this study indicate that GnRH does not change its secretion dynamics, whether measured as mean concentrations, pulse amplitude, or pulse frequency, between seasons.48 In fact, in at least one case, baseline GnRH concentrations were higher during the autumnal transition and anestrus than during the ovulatory period. Like the push–pull results,81 this study also found a significant degree of individual variation in baseline GnRH concentrations between mares. Obviating this wide individual variation by using the same mares during all four seasons strengthens the veracity of this study’s findings when compared with data generated from different horses in a one-time evaluation. An additional piece of compelling evidence that GnRH does not control the fall transition out of cyclicity was recently reported by Collins et al. 82 Constant GnRH infusion was administered to still-cycling mares during the autumnal transition and through the winter in an attempt to prevent or at least delay the transition into anestrus. No effect on transitional events was noted; GnRHtreated mares achieved anestrus at the same time and in the same proportions as untreated mares. GnRH administration also did not affect circulating LH or FSH concentrations during the experimental period. Certainly, there is a need for further investigation on the action of GnRH in seasonal reproductive control in the mare. The most persuasive information
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Autumnal Transition Out of the Breeding Season suggests that GnRH does not directly mediate the seasonal change in pituitary gonadotroph (primarily LH) secretion. If GnRH is not the source of seasonal control over LH, then the decline in LH during the autumnal transition and cessation during anestrus may be mediated at the level of the anterior pituitary. Several possible scenarios could explain the lack of pituitary responsiveness to a continuing GnRH message. (i) LH decline develops owing to loss of positive steroid feedback to the pituitary. The temporal relationship between steroidogenic decline and transitional hypothalamic–pituitary effects reported by Nequin et al.9 support this theory. Some evidence gathered during the vernal transition suggests that estradiol may positively affect the pituitary to stimulate GnRH sensitivity in the mare. Strauss et al.83 demonstrated that vernal increases in estradiol levels precede resumption of LH secretion by the pituitary. When estradiol was administered to ovariectomized pony mares, or given in conjunction with light stimulation to anestrous mares, a subsequent increase in LH84 was measured that was not evidenced in untreated controls. If estrogenic competency must precede GnRH regulation of LH in the spring, then the reverse is also plausible: that the loss of estrogenic capacity in the fall that precedes the decline in LH9 actually causes GnRH decline. Certainly, estrogensecreting ability is preserved and cyclicity is retained in the subpopulation of mares that continue to cycle year-round. Since LH surges continue in winter cycling mares (S. King, unpublished observation),85 it is logical to suppose that pituitary sensitivity to GnRH has also been retained (although this has not been confirmed). Winter cycling mares continue estrous cyclicity despite receiving normal short-day photoperiodic information in the form of melatonin fluctuations,30 ruling out failure of the melatonin signal as the reason for persistent winter cyclicity (see below). Therefore, the single most distinctive difference between mares experiencing winter cyclicity and those that undergo anestrus appears to be retention of steroidogenic capacity and gonadotropic capacity in the former. (ii) Loss of GnRH receptors on the pituitary gonadotrophs during the fall transition could also explain a theoretical disconnect between persistent GnRH and declining LH. Two studies, however, have shown that pituitary GnRH receptor concentrations do not change by season.43, 71 (iii) Lack of pituitary responsiveness to GnRH could be mediated through the action of other elements such as the recently discovered gonadotropin-inhibitory hormone (GnIH).86 GnIH has been identified in mammals, and has the ability to inhibit GnRH-induced LH secretion. Moreover, its production is regulated by melatonin, suggesting that it may be linked to photoperiodic stimuli.
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Luteinizing hormone The influence of photoperiod as a driver of the LH decline during the autumnal transition is called into question by the results generated from a year-long study of ovariectomized pony mares87 in which the LH profile in response to supplemental light (16 light hours/day) was compared with ambient light. Although a seasonal difference was apparent in the spring, there was no difference in the timing of the decline in LH during the fall. Several studies have noted that the time of the final ovulation of the year can be quite variable between mares, not infrequently occurring as late as January or February in the northern hemisphere,1, 7, 8, 88 which would put the seasonal decline in LH secretion in the unusual position of occurring at a time of increasing photoperiod. If neither photoperiod nor GnRH provide signaling for the autumnal transition, the question becomes, what does? It is doubtful that the fall decline simply occurs at a predetermined interval relative to the spring increase, since supplemental lighting to advance the spring transition does not affect the timing of the autumnal transition.5 LH does decrease during the autumnal transition to acyclicity. In fact, current data suggest diminished LH secretion precedes changes, if they occur, in GnRH. Pituitary LH content is higher in summer than in winter,89 suggesting that the decline in pituitary LH occurs during the fall. Hart et al.43 documented a significant decrease in pituitary LH content between July and October, followed by a much larger decline between October and December; pituitary GnRH content did not decrease until December in this study. Several researchers have documented a decline in circulating LH levels between the summer and winter.5, 89 Fewer have analyzed the timecourse and method of this change during the autumnal transition. The annual decline in LH coincides with ovulation failure in the fall,5 although the extent of the decline precipitating anovulation varies widely between mares (S. King, unpublished observation). Most studies reported data from only a segment of the fall transition, generally from one cycle prior to the final ovulatory cycle of the year,41, 47 and showed a decline in the average peak of the LH surge between the second-to-last and last ovulatory cycles. Irvine et al.41 continued data collection into the anovulatory period to show a lack of LH surge at the time of the next expected ovulation. In a study encompassing the entire autumnal transition, Nequin et al.9 profiled two patterns of LH activity: (i) gradual decline over two to three cycles, culminating in a depleted LH surge insufficient to stimulate ovulation (Figure 181.1) and (ii) sudden abatement of the LH surge from normal LH concentrations during the final ovulatory cycle
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to drastically (> 50%) decreased concentrations occurring during the next (anovulatory) follicular wave (Figure 181.3). Pituitary venous blood sampling during the autumnal transition indicated that LH pulse frequency remained unchanged between seasons (approximately 11 pulses/8 hours or 1.4 pulses/hour), although mean concentrations and pulse amplitudes declined during the autumn transitional and anestrous periods.48 In contrast, a very slow LH pulse frequency, approximately 1 pulse/day, was reported from ovariectomized mares during winter.90 Lack of the estradiol influence in these mares that would have been present in many of the intact mares of the previous study may account for this difference in secretion rates. Follicular dynamics appeared to be altered prior to the time of LH decline in that estrogen concentrations declined prior to9 or concurrent with41 decreases in the LH surge in autumn transitional mares. In a complex follicle ablation protocol, Nunes et al.2 showed that, when estradiol concentrations increased during final ovulatory cycles, it was followed approximately 2 days later by an increase in LH; however, when ablation resulted in low estradiol concentrations, LH did not increase. These data suggest that estradiol feedback to the pituitary might diminish positive feedback enough to drive the reduction in LH secretion. We know that estradiol enhances LH secretion in mares.84, 91 In ovariectomized pony mares in which the hypothalamic input to the pituitary had been destroyed, replacement of estradiol increased transcription of the α subunit of the LH molecule.92 This theory is difficult to reconcile with studies showing that ovariectomized pony mares displayed the same timing of LH decline (albeit different peak concentrations) as ovary-intact mares in the fall.87, 93 Continued research is required to determine the nature of estradiol feedback effects on the pituitary at this time. Follicle stimulating hormone The mare is unusual in that she is capable of differentially secreting LH and FSH. The exact method by which this is achieved is unknown, although it has been suggested to be through differential GnRH pulse frequency ‘messages’ from the hypothalamus.94 In fact, not only do FSH and LH surge at different times during the estrous cycle, gonadotropin secretion is independently affected during the seasonal transition. The LH signal from the pituitary declines and is extinguished weeks and sometimes months before the FSH signal is affected in the autumn.5, 9 The nature of the episodic secretion pattern for FSH has not been unequivocally resolved.5 Relative to the autumnal transition, it has been reported
that FSH secretion changes from a double peak (first peak at late estrus to early diestrus and second peak at mid- to late diestrus) during the early portion of the ovulatory season and a single peak as the season progresses.93 Whether there are one, two, or more peaks during the height of the breeding season, FSH secretion appears to resolve into a single peak occurring during mid- to late diestrus through the autumn and early winter.9 There appears to be little photoperiodic influence over plasma FSH concentrations in the autumn. The same FSH pattern of decline was displayed by ovaryintact93 or ovariectomized87, 95 mares regardless of whether they were light supplemented or under the influence of ambient light. Pituitary FSH content also exhibits no seasonal variation.43, 71, 89 Mid-diestrous FSH surges are displayed throughout the autumnal transition, continuing past the time of anovulation and often persisting into acyclicity.9, 41, 54 In fact, FSH secretion can continue unabated in some mares throughout the entire acyclic period.68, 69, 93 In those mares wherein FSH secretion does cease sometime in anestrus, it characteristically does so by way of surges of gradually decreasing amplitude until it dissipates altogether.9 The most interesting aspect of this FSH profile is that major (> 25 mm), and often minor, waves of follicular growth cease in the face of this persistent FSH secretion.54, 68 The mechanism through which follicles fail to recognize the FSH stimulatory signal has not been studied. A simple explanation would be a seasonally related decline and eventual loss of FSH receptors on small, pre-ovulatory follicles. Although this has not been specifically studied in the fall, FSH receptor mRNA was expressed in significantly lower quantity in anestrous ovaries than in summer cycling ovaries,96 suggesting that FSH receptor proliferation may be diminished during the autumnal transition. Variability in the extent of FSH receptor loss in individual mares, and whether receptor sensitivity is lost in accordance with the depth of anestrus, would go far in explaining some of the inconsistent responses to the administration of GnRH, its analogs, and even pituitary extracts, to deeply anestrous mares.97 Another theory to account for the autumn/winter loss of follicular sensitivity to the stimulatory effects of FSH could be that a change in the nature of the pulsatile release of FSH renders it less stimulatory to the follicle.41 FSH pulse frequency as measured by jugular sampling every 6 hours was about half as frequent (0.7 pulses/day) during the autumn transitional period41 compared with summer cycles.94 However, the slowed FSH pulses noted in the fall were similar to those reported for deep winter anestrous mares95, 98 wherein follicular growth had ceased. It is difficult to understand how this pulse frequency can support
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Autumnal Transition Out of the Breeding Season ovulatory follicular growth waves during the fall and also be the driving force behind suspension of follicular growth later in the season. It was noted that excursion in FSH concentrations (the difference between peak and trough values) did decrease during the autumn anovulatory surges.41 Whether the body is sensitive to this difference is not known. Detection of FSH pulses through pituitary venous sampling every 5 minutes for 8 hours from the same mares during each of four separate seasons48 allowed for a very straightforward comparison of FSH pulsatility between seasons while eliminating the considerable variability between mares. Moreover, the fortuitous timing of the autumn sampling protocol spread the seven subject mares into three transitional categories: transitional and ovulatory (one within 5 days of the final ovulation of the year, one > 60 days from final ovulation); major follicular growth waves (≥ 35 mm) without ovulation; anovulatory with no significant follicular growth (i.e., anestrus). FSH concentrations and pulse amplitudes were not different between autumn and the other seasons. FSH pulse frequency was slightly slower during the autumn transitional period (∼10 pulses/8 hours or 1.25 pulses/hour) than during other seasons (∼12 pulses/8 hours or 1.5 pulses/hour). It is not known whether the mare is sensitive to such a subtle difference in pulse frequency. Since at least 29% of the fall transitional mares were cyclical at the time of sampling, and therefore presumably affected by other hormonal feedback, the time during the cycle at which intensive sampling was conducted could affect FSH pulse frequency. Although stage of cycle was taken into account during the spring and summer sampling protocol, it did not appear that was considered during the fall sampling period. This may account for the fact that FSH pulse frequency was different in the anovulatory subgroup of mares compared with the cycling and still-ovulatory mares. It is interesting that FSH pulse frequency during winter anestrus was not different from spring or summer in this study, and not different from the anovulatory group of autumn transitional mares. Although confusing and somewhat conflicting, the results of these two studies suggest that differential pulse frequency messages are not the means by which follicular growth is suspended following the autumnal transition. The possible influence of ovarian steroids on FSH has not been specifically studied during the autumnal transition, but comparisons can be made between cyclicity and anestrus. Progesterone did not significantly change FSH secretion when administered to ovariectomized mares during winter or summer.99 Conflicting data have been reported for estradiol effects on FSH during the winter. Administration of estradiol to ovariectomized mares in winter had no
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effect on FSH in one study,99 but reportedly decreased it in another.100 Apparently, FSH operates rather autonomously in the fall. It does not appear to be influenced by photoperiod; therefore, it does not appear to be closely controlled by GnRH at this time, and it also appears little influenced by ovarian steroids. FSH is affected by inhibin produced by the antral follicles, the effects of which are inhibitory in nature. Inhibin concentrations persist as long as ovulatory follicular growth waves are produced during the autumnal transition41 and may thus help to regulate the character of the FSH surges at this time. What regulates the initiation of a FSH surge during the late transition and into the seasonal anovulatory period is still unknown. Prolactin Prolactin is a well-known player in seasonal reproduction in other species, although its role in the horse is still under investigation. Circulating and pituitary prolactin concentrations display a decidedly seasonal pattern, with highest concentrations in the summer and lowest in winter.89, 101, 102 The circannual change in prolactin concentrations are closely aligned to photoperiod,102 although prolactin’s mode of regulation appears to differ between the spring and the fall transitions. Apparently, studies have not been conducted investigating whether photoperiodic manipulation can alter seasonal prolactin profiles during the autumnal transition. Prolactin is the first of the pituitary hormones to decline in the fall. Fitzgerald et al.45 reported the first change from summer peak levels between June and July with further decreases occurring until nadir in November. The autumn prolactin pattern has been verified by numerous studies.89, 101, 102, 126 Circulating prolactin profiles appear to reflect the state of prolactin stores within the pituitary, which are significantly lower during the winter than summer.89 Lack of sufficient prolactin stores from which to draw for secretion rather than an inhibition of secretion from adequate pituitary stores could explain some of the confusing dichotomy in prolactin secretion dynamics between the vernal and autumnal transitional seasons. It is interesting to note that the pituitary is capable of responding quite early to seasonal change by modifying prolactin secretion, yet its seasonal gonadotropin response is delayed by 1–3 months or more. This likely reflects the differential brain control mechanisms for prolactin compared with the GnRH control over LH and FSH secretion. Dopamine is generally acknowledged to serve as a prolactin-inhibiting factor in the mare and other species, and prolactin is controlled primarily
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through inhibition of its release from pituitary stores.103 Dopamine measured from the cerebrospinal fluid (CSF) displays a seasonal variation consistent with circannual prolactin changes; that is, CSF dopamine concentrations are highest in winter and lowest in summer.104 Short-term19, 20, 105, 106 and longer lasting22, 107 increases in prolactin concentrations can be achieved during the winter anestrus and vernal transition through administration of a number of dopamine antagonists. Many studies have reported success with stimulating early follicular recrudescence and ovulation through administration of dopamine antagonists during anestrus.19–22,108 In contrast, dopamine antagonist treatment beginning in September and continuing throughout the autumn and winter was incapable of delaying the seasonal prolactin decline, and had no effect on follicular growth patterns.126 In fact, a number of mares continue reproductive cyclicity through the winter when basal prolactin concentrations are low,109 making a possible role for prolactin in folliculogenesis difficult to resolve. Dopamine agonist administration was also unsuccessful in provoking a further decline in seasonal prolactin concentrations during the nonbreeding season,107 suggesting that its seasonal nadir levels are reflective of lack of prolactin production by the pituitary lactotrophs. Thyrotropin-releasing hormone is a prolactin secretagogue.29, 102, 107 Its efficacy in stimulating prolactin appears to be restricted to the physiological breeding season.102 This may be related to the reproductive status of the mare, since Gentry et al.29 showed a significantly higher prolactin response to TRH administered in January to a population of mares that continued estrous cyclicity over the winter than when TRH was administered to mares in anestrus. Eliminating the stimulatory action of TRH on prolactin through thyroidectomy did not alter the time of anestrus in horses.110 Collectively, these data suggest that the method of prolactin regulation is different between the spring recrudescence and the autumn recession in reproductive activity. Prolactin regulation during the autumnal transition does not appear to be controlled through the inhibition of prolactin through the dopamine D2 receptor. The seasonal decline may be a reflection of the state of pituitary prolactin production and storage rather than regulation (i.e., inhibition) of secretion. This possibility awaits further study. A putative prolactin effect on reproductive function does not appear to be mediated through the hypothalamic–pituitary gonadotropic axis. Dopamine antagonist treatments during anestrus did not affect LH or FSH secretion.20, 108 In addition to its inhibitory effect on prolactin, dopamine is also a mediator of opioidergic function,
and interactive effects between dopamine antagonists and opioid antagonists during the fall have been reported.106 Opioids mediate the body’s response to adverse environmental stimuli; it is possible that prolactin works synergistically with opioids to modify endogenous reproductive rhythms to adapt to specific environmental conditions. For example, the body fat stores a mare possesses when entering the autumnal transition can certainly influence how well she is likely to survive the privations of winter. Conservation of energy would be more important to a mare with low body condition than one entering autumn with high body condition, and one way to conserve energy is to eliminate estrous cyclicity. Indeed, plasma prolactin concentrations in mares with low body condition scores were lower than high body conditioned mares during the autumnal transition.29 Prolactin is well known to be positively influenced by estradiol.111 Estradiol treatment of ovariectomized pony mares increased prolactin secretion and prolactin content in the pituitary.112 In another study, however, prolactin secretion in estrogentreated ovariectomized mares could be stimulated only in concert with the use of naloxone, an opioid antagonist.106 The interaction of photoperiod and ovarian steroids could influence the nature of seasonal prolactin secretion. Dopamine concentrations in the CSF were higher in long-term ovariectomized mares than intact mares, and ovariectomy appeared to remove the dopamine response to seasonal cues.104 Prolactin concentrations were increased, and the stimulatory effects on anestrous ovaries produced by dopamine antagonist administration were enhanced by estradiol pretreatment prior to administration of a dopamine antagonist.113 However, this positive effect of estradiol may only be operating under conditions of the vernal transition and summer cyclicity, since persistent estrous cycles that generate estradiol-17β peaks persist well past the time of prolactin decline,9 and sometimes during its winter nadir without an apparent modulating effect on circulating prolactin concentrations. Prolactin is negatively influenced by melatonin in a seasonally sensitive pattern. Specifically, the mare responds to melatonin with a decrease in circulating prolactin concentrations during the spring and summer months, but appears refractory to the suppressive effects of melatonin during the fall and winter.45 The end-of-season decline in prolactin proceeds regardless of the melatonin input. This indicates that the seasonal decline in prolactin is not regulated through melatonin, and is therefore perhaps an endogenous effect. The local effect of prolactin on ovarian function has not been defined, although its involvement has been implicated in luteolysis and folliculogenesis.
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Autumnal Transition Out of the Breeding Season Episodic prolactin releases occur around the time of luteolysis to early estrus,14, 53, 109, 114 leading researchers to speculate as to its possible role in luteolysis. In one experiment, administration of exogenous PGF2α increased episodic secretion of prolactin,53 suggesting a link between these two hormones. As discussed previously (see Ovarian events characterizing the fall transition), the rate of failure of luteolytic release of PGF2α , resulting in a spontaneously prolonged corpus luteum, increases during the autumnal transition.14 If prolactin is found to be involved with the luteolytic process, perhaps through a link to PGF2α , its decreased availability during the autumnal transition would provide a theoretical mechanism to explain the increased incidence of luteolytic failure in the fall. This episodic prolactin secretion appears to persist at least as long as estrous cycles persist into the fall and winter,41, 109 although pulses are superimposed on a much diminished baseline.41 Increasing prolactin availability to the ovary through dopamine inhibition19–22,115 or exogenous prolactin administration108, 115 during winter anestrus results in follicular recrudescence. Consideration should be given to the possibility that stimulation of follicular growth during anestrus following administration of crude pituitary extracts116 might be attributable, at least in part, to prolactin residues within the extract. Prolactin has also been identified in the follicular fluid of the mare in concentrations proportional to the size of the follicle and appears to have a seasonal component.117 Comparably sized autumn follicles contain lower levels of follicular fluid prolactin than their summer counterparts.117 The significance of this discovery to the reproductive physiology of the mare is not known. In summary, prolactin appears to have a rigidly entrained, endogenously driven rhythm in the fall, wherein circulating prolactin concentrations decline starting shortly after the summer solstice to reach a nadir prior to the winter solstice. This decline does not appear to be regulated through activity of photoperiodic mediators or activity of prolactin inhibiting factor, or to be sensitive to prolactin-stimulating factors. It is likely reflective of a sharp decrease in pituitary prolactin production, resulting in depleted prolactin stores available for release. In this respect, although prolactin appears to play some role in reproduction at the ovarian level in the mare, that role does not include events driving the autumnal transition. Melatonin Melatonin is the hormone through which photoperiodic information is translated into a chemical mes-
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sage to the body. Melatonin is released from the pineal during periods of dark.118, 119 Therefore, it is assumed that the progressive prolongation of melatonin secretion in response to the increasingly long nights of autumn triggers a number of endocrine events associated with the recession in reproductive activity at this time. This does not appear to be the case, however; like the less-than-straightforward situation we find with other hormones during the autumnal transition, melatonin’s activities during this period are enigmatic. In a general sense, melatonin is released over a longer period and is therefore circulating in higher quantities during the autumn and winter than in the spring and summer.120 Individually, however, there is a substantial variability in mares’ melatonin responses to decreased illumination; melatonin frequently failed to increase in response to the dark phase.8, 121 When episodic melatonin concentrations were measured every 2 hours for 48 hours during the spring, summer, fall, and winter periods, Diekman et al.121 found that responses did not correlate with light/dark phases during any seasonal period except summer. Increased melatonin production was reported to be temporally associated with decreased reproductive activity.5, 120 However, a cause-and-effect relationship does not exist, at least during the autumn and winter. In two studies encompassing the autumnal transition and the winter anestrous period,8, 121 no association between melatonin concentrations and cyclic (or acyclic) reproductive conditions was found. Mares that continued winter cyclicity generally did so despite elevated melatonin concentrations.8, 30, 34, 37, 121 In fact, mares cycling in December had higher melatonin concentrations than anestrous mares.121 These researchers concluded that melatonin is not used to regulate cyclic activity in the mare throughout the seasons. Efforts to manipulate seasonal reproduction through influencing melatonin concentrations have generally met with disappointing results. Supplemental melatonin delivered by implants, oral supplements, and multiple injections failed to reliably change seasonal reproductive patterns in mares, particularly when researchers sought to change the autumn-to-winter reproductive transition. Constant melatonin administered during the summer and through the autumn/winter had no effect on the timing of the fall transition.8, 30, 34, 37 Guillaume et al.37 did report a forward shift in the onset of anestrus in response to melatonin implants in a population of very young mares (1.5 years old) undergoing their first anestrous period. This suggests that melatonin acts as a signaling device that may be used in concert with other factors such as age and body condition
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(see Control over the fall transition, Environmental factors, nutrition, and management) to inform the mare about her environment and modify the endogenous seasonal reproductive pattern. The elimination of any melatonin response through pinealectomy or superior cervical ganglionectomy also does not abolish reproductive seasonality in the horse.122–124 Although the autumnal transition was not specifically studied, altered mares achieved seasonal anestrus. Melatonin has been suggested to decrease hypothalamic GnRH content,83, 124 and its influence on seasonal reproduction is mediated through GnRH availability. We have seen, however, that GnRH does not appear to have a strong role in leading the autumnal transition in the mare (see Endocrine events characterizing the autumnal transition, Gonadotropinreleasing hormone). Therefore, as far as the autumnal transition is concerned, melatonin’s possible effects on GnRH are moot. The melatonin axis is activated through catecholaminergic nerve fibers.97 Although the α-adrenergic agonist isoproterenol was capable of stimulating melatonin secretion,125 neither it nor administration of a α 2 -adrenergic agonist34, 126 were successful in changing autumn seasonal reproductive profiles. The bulk of the work on melatonin underscores the existence of a strong endogenous circannual reproductive rhythm in the mare that is likely the primary driver of the autumnal transition. Collectively, these results suggest that melatonin, perhaps in concert with other factors such as opioids, may help to inform and entrain that endogenous reproductive rhythm, but it does not exert a direct influence on the pituitary or the ovary.
Behavioral changes with the fall transition Those who follow mares’ estrous cycles through the late summer and into the autumn are well aware of the numerous anomalous reproductive behaviors in transitional mares. This shift to unreliable reproductive behaviors appears to be random; that is, it occurs as frequently in mares who display strong, consistent, appropriate sexual behaviors during the summer as it does in those who are known to have aberrant behaviors even during the height of the breeding season (S. King, personal observation). Anyone attempting to breed or treat mares for reproductive problems during this time of year would do well to keep this in mind, and to adopt the policy of regular ovarian assessment over relying on behavior as an indicator of reproductive status.
Up to 25% of autumnal cycles127 were characterized by follicular growth, estradiol secretion, and ovulation without accompanying estrous behavior (silent estrus). This behavioral shift may be influenced by environmental factors, as the incidence of silent estrus did not appear to increase between seasons at 10◦ latitude.17 Silent estrus was strongly associated with the final ovulation of the season.9, 47, 127 Mean peak estradiol levels during silent estrus (5.3 ± 0.9 pg/mL) were lower than for summer (see Follicular steroidogenesis) but not different from autumnal cycles in which estrus was displayed, perhaps indicating a changing brain sensitivity to the hormone in addition to lowered follicular estradiol production.9 Inappropriate or paradoxical estrus is estrus unaccompanied by significant follicle size (≥ 25 mm) or increased circulating estrogens, and has been reported during the autumnal transition and winter anesrus.1, 3, 127 Perhaps because this demonstration of estrus is not driven by ovarian or hormonal events, it is generally transient and often discontinuous. Estrous behavior was displayed by one autumntransitional mare during a period of spontaneously prolonged corpus luteum activity.127 This mare developed a pre-ovulatory follicle and increased estradiol concentrations in the face of persistent progesterone secretion, a situation that can occur during normal length luteal phases,5 but has not been associated with estrous behavior. As mares progress further into the autumn transition toward anestrus, not only the incidence but also the intensity of estrous behavior declines (S. King, personal observation).47, 128 Anestrus is typified by an attitude of indifference by the mare toward the stallion; she neither actively repulses nor invites his attention.128 This indifferent behavior pattern appears to develop, at least in some mares, gradually during the transition and into anestrus (S. King, personal observation).
The four autumnal transitional phases Based on results from data compiled over 5 years of autumn transitional studies7, 9, 14, 23, 40 we have defined four phases of the transitionary process that occur over an approximately 4- to 5-month period (Figure 181.5). Phase 1 encompasses a period of normal appearing estrous cycles during the early autumn wherein prolactin concentrations and luteal function (progesterone secretion and luteolytic ability) decline with few outward signs of change in estrous cyclicity. Spontaneously prolonged corpora lutea tend to be formed during this phase. Phase 2 defines a
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Progesterone
phase 3 of their transition and move into the vernal transition toward renewed reproductive function.
FSH
LH
The free-running autumnal transition
OV Follicle Diameter
OV OV
OV
OV
Estradiol
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18
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13 27 Nov
11 Dec
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Figure 181.5 Gonadotropins, ovarian steroids, follicular diameters, and ovulations compiled from representative mares during the fall transition to illustrate the four transitional phases. Phase 1, normal estrous cycles despite declining ovarian steroid production. Phase 2, ovulatory cycles with prolonged follicular growth phases and shortened luteal phases. Follicular estradiol production is greatly decreased. Many mares demonstrate a progressive decline in luteinizing hormone (LH) during this period. Phase 3, ovulation failure due to low LH secretion. Major and minor follicular waves may persist for weeks. Folliclestimulating hormone (FSH) surges often persist past this phase. Phase 4, full ovarian quiescence, sometimes despite continued FSH surges. Data modified from Nequin et al.9 With permission from Nottingham University Press.
period of aberrant but ovulatory cycles occurring during the late autumn wherein the pace of follicular growth slows, follicular steroidogenesis declines, and LH may decrease over a period of one to three estrous cycles. Silent estrus episodes appear more frequently during this phase, as do shortened luteal phases and prolonged follicular phases. Phase 3 marks the anovulatory period of the transition wherein LH declines to the point of ovulatory failure. Waves of significant follicular growth and atresia often persist for a variable period (30 days or longer) following initial ovulatory failure and gradually subside into phase 4, which is defined by total reproductive quiescence (anestrus), during which follicular growth does not exceed 20 mm and ovarian steroid as well as gonadotropin secretion ceases. Some mares never achieve this phase of ‘deep’ anestrus; they reach
Taken in total, the data available on the autumnal transition into anestrus indicate that the mare responds to an endogenous rhythmicity that drives her seasonal reproductive downturn. This endogenous rhythm appears to be open to modest modification by a number of factors, many of which we understand only slightly. Some of these factors are photoperiod (although this is not interpreted through melatonin), temperature and possibly other environmental stressors, age, and body condition. The autumnal transition proceeds at a leisurely pace, at least in horse mares from temperate and torrid regions. The first adjustments to the advancing season are initiated shortly following the summer solstice, and complete reproductive quiescence may not be accomplished until after the winter solstice, if at all. In light of the horse breeding industry’s incentives to produce early season foals before the time of the natural breeding season, a more thorough understanding of the autumnal transition could be instrumental in devising a reliable method by which seasonality can be blocked and breeders freed to breed at any time of the year.
References 1. Gentry LR, Thompson DL, Gentry GT, Davis KA, Godke RA, Cartmill JA. The relationship between body condition, leptin, and reproductive and hormonal characteristics of mares during the seasonal anovulatory period. J Anim Sci 2002;80:2695–703. 2. Nunes MM, Gastal EL, Gastal MO, Rocha Filho AN, Mellagi AP, Ginther OJ. Follicle and gonadotropin relationships during the beginning of the anovulatory season in mares. Anim Reprod 2005;2:41–9. 3. Weedman BJ, King SS, Neumann KR, Nequin LG. Comparison of circulating estradiol-17β and folliculogenesis during the breeding season, autumn transition and anestrus in the mare. J Equine Vet Sci 1993;13:502–5. 4. Kooistra LH, Ginther OJ. Effect of photoperiod on reproductive activity and hair in mares. Am J Vet Res 1975;24:1413–19. 5. Ginther OJ. Reproductive Biology of the Mare; Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992; pp. 135–72. 6. Ginther OJ. Occurrence of anestrus, estrus, diestrus and ovulation over a 12-month period in mares. Am J Vet Res 1974;35:1173–9. 7. King SS, Neumann KR, Nequin LG, Weedman BJ. Time of onset and ovarian state prior to entry into winter anestrus. J Equine Vet Sci 1993;13:512–15. 8. Fitzgerald BP, Schmidt MJ. Absence of an association between melatonin and reproductive activity in mares
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9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
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during the nonbreeding season. Biol Reprod Monogr 1995;1:425–34. Nequin LG, King SS, Roser JF, Stoderstrom BL, Carnevale EM, Neumann KR. Uncoupling of the equine reproductive axes during transition into anoestrus. J Reprod Fertil Suppl 2000;56:153–61. Huszenicza G, Nagy P, Juhasz J, Korodi P, Kulcsar M, Reiczigel J, Guillaume D, Rudas P, Solti L. The relationship between thyroid function and expression of seasonal reproductive activity in mares. J Reprod Fertil Suppl 2000;56:163–72. Scraba ST, Ginther OJ. Effects of lighting programs on onset of the ovulatory season in mares. Theriogenology 1985;24:667–79. Palmer E, Driancourt MA. Some interactions of seasons of foaling, photoperiod and ovarian activity in the equine. Livest Prod Sci 1983;10:197–210. Colquhoun KM, Eckersall PD, Renton JP, Douglas TA. Control of breeding in the mare. Equine Vet J 1987;19:138–42. King SS, Douglas BL, Roser JF, Silvia WJ, Jones KL. Differential luteolytic function between the physiological breeding season, autumn transition and persistent winter cyclicity in the mare. Anim Reprod Sci 2010;117:232–40. Saltiel A, Calderon A, Garcia N, Hurley DP. Ovarian activity in the mare between latitude 15◦ and 22◦ N. J Reprod Fertil Suppl 1982;32:261–7. Dowsett KF, Knott LM, Woodward RA, Bodero DA. Seasonal variation in the estrous cycle of mares in the subtropics. Theriogenology 1993;39:631–53. Quintero B, Manzo M, Diaz T, Verde O, Benacchio N, Sifontes L. Seasonal changes in ovarian activity and estrous behavior of Thoroughbred mares in a tropical environment. Biol Reprod Monogr 1995;1:469–74. Boeta M, Porras A, Zarco LA, Aguirre-Hernandez R. Ovarian activity of the mare during winter and spring at a latitude of 19◦ 21’ north. J Equine Vet Sci 2006;26:55–8. Nequin LG, King SS. A dopaminergic neural system inhibits ovarian function during anestrus in the mare. Biol Reprod Suppl 1988;1:114. Besognet B, Hansen BS, Daels PF. Induction of reproductive function in anestrous mares using a dopamine antagonist. Theriogenology 1997;47:467–80. Bennett-Wimbush K, Loch WE, Plata-Madrid J, Evans T. The effects of perphenazine and bromocriptine on follicular dynamics and endocrine profiles in anestrous pony mares. Theriogenology 1998;49, 717–33. Brendemuehl JP, Cross DL. Influence of the dopamine antagonist domperidone on the vernal transition in seasonally anoestrous mares. J Reprod Fertil Suppl 2000;56:185–93. King SS, Jones KL, Nequin LG, Murphy LL, Campbell AG. Evidence for a local dopamine-driven regulatory system in the mare. Theriogenology 2002;58:619–22. Calogero AE, Gagdy G, D’Agata R. Mechanisms of stress on reproduction. Evidence for a complex intrahypothalamic circuit. Ann N Y Acad Sci 1998;851:364–70. Irvine CHG, Alexander SL, Turner JE. Differential effect of graded doses of naloxone on the reproductive and adrenal axes in seasonally acyclic mares. Endocrine 1994;2:913–19. Turner JE, Irvine CHG, Alexander SL. Regulation of seasonal breeding by endogenous opioids in mares. Biol Reprod Monogr 1995;1:443–8. Fichna J, Janecka A, Costentin J, Do Rego J-C. The endomorphin system and its evolving neurophysiological role. Pharmacol Rev 2007;59:88–123.
28. King SS, Campbell AG, Dille EA, Roser JF, Murphy LL, Jones KL. Dopamine receptors in equine ovarian tissues. Dom Anim Endocrinol 2005;28:405–15. 29. Gentry LR, Thompson DL, Stelzer AM. Responses of seasonally anovulatory mares to daily administration of thyrotropin-releasing hormone and(or) gonadotropinreleasing hormone analog. J Anim Sci 2002;80:208–13. 30. Fitzgerald BP, McManus CJ. Photoperiodic versus metabolic signals as determinants of seasonal anestrus in the mare. Biol Reprod 2000;63:335–40. 31. Guillaume D, Duchamp G, Salazar-Ortiz J, Nagy P. Nutrition influences the winter ovarian inactivity in mares. Theriogenology Suppl 2002;58:593–7. 32. Ferreira-Dias G, Claudino F, Carvalho H, Agricola R, Alpoim-Moreira J, Robalo Silva J. Seasonal reproduction in the mare: possible role of plasma leptin, body weight and immune status. Dom Anim Endocrinol 2005;29:203–13. 33. Vick MM, Sessions DR, Murphy BA, Kennedy EL, Reedy SE, Fitzgerald BP. Obesity is associated with altered metabolic and reproductive activity in the mare: effects of metformin on insulin sensitivity and reproductive cyclicity. Reprod Fertil Dev 2006;18:609–17. 34. McManus CJ, Fitzgerald BP. Effect of daily clenbuterol and exogenous melatonin treatment on body fat, serum leptin and the expression of seasonal anestrus in the mare. Anim Reprod Sci 2003;76:217–30. 35. Fitzgerald BP, Reedy SE, Sessions DR, Powell DM, McManus CJ. Potential signals mediating the maintenance of reproductive activity during the non-breeding season of the mare. Reproduction Suppl 2002;59:115–29. 36. Mitcham PB, Thompson DL, Storer WA, Gentry LR, Godke RA, Gertler A. Metabolic and reproductive effects of recombinant equine leptin on seasonally anovulatory mares. In: Proceedings of the 20th Symposium of the Equine Science Society, 2007, pp. 165–6. 37. Guillaume D, Arnaud G, Camillo F, Duchamp G, Palmer E. Effect of melatonin implants on reproductive status of mares. Biol Reprod Monogr 1995;1:435–42. 38. Wesson JA, Ginther OJ . Influence of season and age on reproductive activity in pony mares on the basis of a slaughterhouse survey. J Anim Sci 1981;52:119–29. 39. Wesson JA, Ginther OJ . Puberty in the female pony: reproductive behavior, ovulation, and plasma gonadotropin concentrations. Biol Reprod 1981;24:977–86. 40. King SS, Nequin LG, Drake S, Hebner TS, Roser JF, Evans JW. Progesterone levels correlate with impending anestrus in the mare. J Equine Vet Sci 1988;8:109–11. 41. Irvine CHG, Alexander SL, McKinnon AO. Reproductive hormone profiles in mares during the autumn transition as determined by collection of jugular blood at 6 h intervals throughout ovulatory and anovulatory cycles. J Reprod Fertil 2000;118:101–9. 42. Evans MJ, Irvine CHG. Serum concentrations of FSH, LH and progesterone during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1977;23:193–200. 43. Hart PJ, Squires EL, Imel KJ, Nett TM. Seasonal variation in hypothalamic content of gonadotropin-releasing hormone (GnRH), pituitary receptors for GnRH, and pituitary content of luteinizing hormone and follicle-stimulating hormone in the mare. Biol Reprod 1984;30:1055–62. 44. Murphy BD, Rajkumar K. Prolactin as a luteotrophin. Can J Physiol Pharm 1985;63:257–64. 45. Fitzgerald BP, Davison LA, McManus CJ. Evidence for a seasonal variation in the ability of exogenous melatonin
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to suppress prolactin secretion in the mare. Dom Anim Endocrinol 2000;18:395–408. King SS, Jones KL, Dille EA, Roser JF. Prolactin receptors in the corpus luteum of mares. In: Proceedings of the 15th International Congress on Animal Reproduction, 2004, p. 39. Snyder DA, Turner DD, Miller KF, Garcia MC, Ginther OJ. Follicular and gonadotrophic changes during transition from ovulatory to anovulatory seasons. J Reprod Fertil Suppl 1979;27:95–101. Cooper DA. Reproductive neuroendocrine function in the mare as reflected in the intercavernous sinus during ovulatory, anovulatory, and transitional seasons. MS thesis, Texas A&M University, 2006. Stabenfeldt GH, Hughes JP, Evans JW, Neely DP. Spontaneous prolongation of luteal activity in the mare. Equine Vet J 1974;6:158–63. Neely DP, Kindahl H, Stabenfeldt GH, Edqvist LE, Hughes JP. Prostaglandin release patterns in the mare: physiological, pathophysiological, and therapeutic responses. J Reprod Fertil Suppl 1979;27:181–9. Stabenfeldt GH, Hughes JP, Neely DP, Kindahl H, Edqvist LE. Physiologic and pathophysiologic aspects of prostaglandin F2 alpha during the reproductive cycle. J Am Vet Med Assoc 1980;176:1187–94. Heath DT, Welsh CM, Jones KL, King SS. Immunohistochemical localization of the dopamine D2 receptor in ovarian tissues during anestrus and cyclicity. In: Proceedings of the 20th Symposium of the Equine Science Society, 2007, pp. 50–1. Shand N, Irvine CHG, Turner JE, Alexander SL. A detailed study of hormonal profiles in mares at luteolysis. J Reprod Fertil Suppl 2000;56: 271–9. Ginther OJ, Woods BG, Meira C, Beg MA, Bergfelt DR. Hormonal mechanism of follicle deviation as indicated by major versus minor follicular waves during the transition into the anovulatory season in mares. Reproduction 2003;126:653–60. Donadeu FX. Early indicators of follicular growth during the anovulatory season in mares. Anim Reprod Sci 2006;94:179–81. Burkhardt J. Some clinical problems of horse breeding. Vet Rec 1948;60:243–8. Gastal EL, Gastal MO, Ginther OJ. The suitability of echotexture characteristic of the follicular wall for identifying the optimal breeding day in mares. Theriogenology 1998;50:1025–38. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Incidence, endocrinology, vascularity, and morphology of hemorrhagic anovulatory follicles in mares. J Equine Vet Sci 2007;27:130–9. Knudsen O, Velle W. Ovarian oestrogen levels in the non-pregnant mare: Relationship to histological appearance of the uterus and to clinical status. J Reprod Fertil 1961;2:130–7. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Conversion of a viable preovulatory follicle into a hemorrhagic anovulatory follicle in mares. Anim Reprod 2006;3:29–40. Gastal EL, Gastal MO, Bergfelt DR, Ginther OJ. Role of diameter differences among follicles in selection of a future dominant follicle in mares. Biol Reprod 1997;57:1320–7. Nunes MM, Gastal EL, Gastal MO, Rocha AN. Influence of the autumn transitional phase on follicu-
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69.
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72.
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lar development in mares. Theriogenology Suppl 2002;58: 603–6. Pierson RA, Ginther OJ. Follicular population dynamics during the estrous cycle of the mare. Anim Reprod Sci 1987;14:219–31. Ginther OJ, Beg MA, Donadeu FX, Bergfelt DR . Mechanism of follicle deviation in monovular farm species. Anim Reprod Sci 2003;78:239–57. Hughes JP, Stabelfeldt GH, Evans JW. Clinical and endocrine aspects of the estrous cycle of the mare. In: Proceedings of the 18th Annual Convention of the American Association of Equine Practitioners, 1972, pp. 119–48. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Seasonal influence on equine follicle dynamics. Anim Reprod 2004;1:31–44. Donadeu FX, Ginther OJ. Follicular waves and circulating concentrations of gonadotrophins, inhibin and oestradiol during the anovulatory season in mares. Reproduction 2002;124 875–85. Donadeu FX, Ginther OJ. Interactions of follicular factors and season in the regulation of circulating concentrations of gonadotrophins in mares. Reproduction 2003;125:743–50. Van Niekerk CH, Gerneke WH, van Heerden JS. Anatomical and histological observations on the reproductive tract of mares with abnormal oestrous cycles. J South Afr Vet Med Assoc 197344:141–152. Silvia PJ, Squires EL, Nett TM. Changes in the hypothalamus-hypophyseal axis of mares associated with seasonal reproductive recrudescence. Biol Reprod 1986;35: 897–905. Strauss SS, Chen CL, Kalra SP, Sharp DC . Localization of gonadotrophin releasing hormone (GnRH) in the hypothalamus of ovariectomized pony mares by season. J Reprod Fertil Suppl 1979;27:123–9. Alexander SL, Irvine CHG. Secretion rates and short-term patterns of GnRH, FSH and LH throughout the periovulatory period in the mare. J Endocrinol 1987;114:351–62. Herbison AE. Multimodal influence of estrogen upon gonadotropin-releasing hormone neurons. Endocr Rev 1998;19:302–30. Hyland JH, Wright PJ, Clarke IJ, Carson RS, Langsford DA, Jeffcott LB. Infusion of gonadotrophin releasing hormone (GnRH) induces ovulation and fertile oestrus in mares during seasonal anoestrus. J Reprod Fertil Suppl 1987;35:211–20. Allen WR, Sanderson MW, Greenwood RES, Ellis DR, Crowhurst JS, Simpson DJ, Rossdale PD. Induction of ovulation in anoestrous mares with a slow-release implant of a GnRH analogue (ICI 118 630). J Reprod Fertil Suppl 1987;35:469–78. Turner JE, Irvine CHG. The effect of various gonadotrophin-releasing hormone regimens on gonadotrophins, follicular growth and ovulation in deeply anoestrous mares. J Reprod Fertil Suppl 1992;44:213–25. Williams GL, Amstalden M, Blodgett GP, Ward JE, Unnerstall DA, Quirk KS. Continuous administration of lowdose GnRH in mares I. Control of persistent anovulation during the ovulatory season. Theriogenology 2007;68:67– 75. Johnnson CA, Thompson DL, Cartmill JA. Pituitary responsiveness to GnRh in mares following deslorelin acetate implantation to hasten ovulation. J Anim Sci 2002;80:2681–7. Irvine CHG, Alexander SL. A novel technique for measuring hypothalamic and pituitary hormone secretion rates
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from collection of pituitary venous effluent in the normal horse. J Endocrinol 1987;113:183–92. Sharp DC, Grubaugh WR. Use of push-pull perfusion techniques in studies of gonadotrophin-releasing hormone secretion in mares. J Reprod Fertil Suppl 1987;35:289–96. Collins SM, Zieba DA, Williams GL. Continuous administration of low-dose GnRH in mares II. Pituitary and ovarian responses to uninterrupted treatment beginning near the autumnal equinox and continuing throughout the anovulatory season. Theriogenology 2007;68: 673–81. Strauss SS, Chen CL, Kalra SP, Sharp DC. Depletion of hypothalamic gonadotropin releasing hormone (GnRH) in ovariectomized mares following melatonin implants. Fed Proc 1979;37:225. Sharp DC, Grubaugh WR, Weithernauer J, Davis SD, Wilcox CJ. Effects of steroid administration on pituitary luteinizing hormone and follicle-stimulating hormone in ovariectomized pony mares in the early spring: pituitary responsiveness to gonadotropin-releasing hormone and pituitary gonadotropin content. Biol Reprod 1991;44:983–90. Davison LA, McManus CJ, Fitzgerald BP. Gonadotropin response to naloxone in the mare: effect of time of year and reproductive status. Biol Reprod 1998;59:1195–9. Bentley GE, Kriegsfeld LJ, Osugi T, Ukena K, O’Brien S, Perfito N, Moore IT, Tsutsui K, Wingfield JD. Interactions of gonadotropin-releasing hormone (GnRH) and gonadotropin-inhibitory hormone (GnIH) in birds and mammals. J Exp Zool 2006;305A:807–14. Freedman LJ, Garcia MC, Ginther OJ. Influence of photoperiod and ovaries on seasonal reproductive activity in mares. Biol Reprod 1979;20:567–74. Nagy P, Huszenicza G, Juhasz J, Solsti L, Kulcsar M. Diagnostic problems associated with ovarian activity n barren and postpartum mares early in the breeding season. Reprod Dom Anim 1998;33:187–92. Thompson DL, Johnson L, St. George EI, Garza F. Concentrations of prolactin, luteinizing hormone and follicle stimulating hormone in pituitary and serum of horses: effect of sex, season and reproductive state. J Anim Sci 1986;63:854–960. Fitzgerald BP, l’Anson H, Loy RG, Legan SJ. Evidence that changes in LH pulse frequency may regulate the seasonal modulation of LH secretion in ovariectomized mares. J Reprod Fertil 1983;69:685–92. Aurich C, Daels PF, Ball BA, Aurich JE. Effects of gonadal steroids on the opioid regulation of LH and prolactin release in ovariectomized pony mares. J Endocrinol 1995;147:195–202. Robinson G, Porter MB, Peliter MR, Cleaver BC, Farmerie TA, Wolfe MW, Nilson JH, Sharp DC. Regulation of luteinizing hormone b and a messenger ribonucleic acid by estradiol or gonadotropin-releasing hormone following pituitary stalk section in ovariectomized pony mares. Biol Reprod Monogr 1995;1:373–83. Turner DD, Garcia MC, Ginther OJ. Follicular and gonadotropic changes throughout the year in pony mares. Am J Vet Res 1979;40:1694–700. Irvine CHG, Alexander SL. FSH and LH. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 50–1. Thompson DL, McNeill DR, Wiest JJ, St. George RL, Jones LS, Garza F. Secretion of luteinizing hormone and follicle
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stimulating hormone in intact and ovariectomized mares in summer and winter. J Anim Sci 1987;64:247–53. King SS, Jones KL, Mullenix BA, Heath DT. Seasonal relationships between dopamine D1 and D2 receptor and equine FSH receptor mRNA in equine ovarian epithelium. Anim Reprod Sci 2008;108: 259–66. Nagy P, Guillaume D, Daels P. Seasonality in mares. Anim Reprod Sci 2000; 60–61:245–62. Alexander SL, Irvine CHG. Control of onset of breeding season in the mare and its artificial regulation by progesterone treatment. J Reprod Fertil Suppl 1991;44:307–19. Garcia MC, Freedman LJ, Ginther OJ. Interaction of seasonal ovarian factors in the regulation of LH and FSH secretion in the mare. J Reprod Fertil Suppl 1979;27:103–11. Garza F, Thompson DL, St. George RL, French DD. Androgen and estradiol effects on gonadotropin secretion and response to GnRH in ovariectomized pony mares. J Anim Sci 1980;62:1654–9. Johnson AL. Serum concentrations of prolactin, thyroxine and triodothyronine relative to season and the estrous cycle in the mare. J Anim Sci 1986;62:1012–20. Evans MJ, Alexander SL, Irvine CHG, Livesey JH, Donald RA. In vitro and in vivo studies of equine prolactin secretion throughout the year. J Reprod Fertil Suppl 1991;44:27–35. Ben-Jonathan N, Hnasko R. Dopamine as a prolactin (PRL) inhibitor. Endocr Rev 2001;22:724–63. Melrose PA, Walker RF, Douglas RH. Dopamine in the cerebrospinal fluid of prepubertal and adult horses. Brain Behav Evol 1990;35:98–106. Thompson DL, DePew CL. Prolactin, gonadotropin, and hair shedding responses to daily sulpiride administration in geldings in winter. J Anim Sci 1997;75:1087–91. Aurich C, Parvizi N, Brunklaus D, Hoppen H-O, Aurich JE. Opioidergic and dopaminergic effects on LH and prolactin release in pony mares at different times of the year. J Reprod Fertil Suppl 2000;56:195–203. Johnson AL, Becker SE. Effects of physiologic and pharmacologic agents on serum prolactin concentrations in the nonpregnant mare. J Anim Sci 1987;65:1292–7. Nequin LG, King SS, Johnson AL, Gow GM, Ferreira-Dias GM. Prolactin may play a role in stimulating the equine ovary during the spring reproductive transition. J Equine Vet Sci 1993;13:631–5. Worthy K, Colquhoun K, Escreet R, Dunlop M, Renton JP, Douglas TA. Plasma prolactin concentrations in non-pregnant mares at different times of the year and in relation to events in the cycle. J Reprod Fertil Suppl 1987;35:269–76. Porter MB, Cleaver B, Robinson G, Peltier M, Shearer LC, Dahl GE, Sharp DC. A comparative study examining the role of the thyroid in seasonal reproduction in pony mares and ewes. Biol Reprod 1995;52 Suppl 1:134. Lamberts SWH, MacLeod RM. Regulation of prolactin secretion at the level of the lactotroph. Physiol Rev 1990;70:279–318. Thompson DL, Garza F, St. George RL, Rabb MH, Barry BE, French DD. Relationships among LH, FSH and prolactin secretion, storage and response to secretagogue and hypothalamic GnRH content in ovariectomized pony mares administered testosterone, dihydrotestosterone, estradiol, progesterone, dexamethasone or follicular fluid. Dom Anim Endocrinol 1991;8:189–99. Kelley KK, Thompson DL, Storer WA, Mitcham PB, Gilley RM, Burns PJ. Estradiol interactions with dopamine
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antagonists in mares: Prolactin secretion and reproductive traits. J Equine Vet Sci 2006;26:517–28. Roser JF, O’Sullivan J, Evans JW, Swedlow J, Papkoff H. Episodic release of prolactin in the cyclic mare. J Reprod Fertil Suppl 1987;35:687–8. Thompson DL, Hoffman R, DePew CL. Prolactin administration to seasonally anestrous mares: Reproductive, metabolic, and hair shedding responses. J Anim Sci 1997;75:1092–9. Lapin DR, Ginther OJ. Induction of ovulation and multiple ovulations in seasonally anovulatory and ovulatory mares with an equine pituitary extract. J Anim Sci 1977;44:834–42. King SS, Roser JF, Jones KL . Follicular fluid prolactin and the periovulatory prolactin surge in the mare. J Equine Vet Sci 2008;28:468–72. Grubaugh W, Sharp DC, Berglund LA, McDowell KJ, Kilmer DM, Peck LS, Seamans KW. Effects of pinealectomy in pony mares. J Reprod Fertil Suppl 1982;32:293–5. Kilmer DM, Sharp DC, Berglund LA, Grubaugh W, McDowell KJ, Peck LS. Melatonin rhythms in pony mares and foals. J Reprod Fertil Suppl 1982;32:303–7. Wesson JA, Orr EL, Quay WB, Ginther OJ. Seasonal relationship between pineal hydroxyindole-Omethyltransferase (HIOMT) activity and reproductive status in the pony. Gen Comp Endocrinol 1979;38:46–52. Diekman MA, Braun W, Peter D, Cook D. Seasonal serum concentrations of melatonin in cycling and noncycling mares. J Anim Sci 2002;80:2949–52.
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122. Sharp DC, Vernon MW, Zavy MT. Alteration of seasonal reproductive patterns in mares following superior cervical ganglionectomy. J Reprod Fertil Suppl 1979;27: 87–93. 123. Cheves LL, Sharp DC. The effect of oral melatonin on the onset of the breeding season in pony mares under artificially lengthened photoperiod. In: Proceedings of the 9th Equine Nutrition and Physiology Symposium, 1985, pp. 330–4. 124. Sharp D, Grubaugh W, Berglund LA, Seamans KW, McDowell KJ, Kilmer DM, Peck LS. The interaction of photoperiod and pineal gland on seasonal reproductive patterns in mares. In: Photoperiodism and Reproduction in Vertebrates. Proc INRA Colloq 1981;6:201–29. 125. Sharp DC, Grubaugh WR, Berglund LA, Seamans KW. Isoproterenol-stimulation of melatonin release in mares. J Anim Sci 1980;51 Suppl 1:534. 126. King SS, Roser JF, Cross DL, Jones KL . Dopamine antagonist affects luteal function but not cyclicity during the autumn transition. J Equine Vet Sci 2008;28: 345–50. 127. Weedman BJ. Relationships between folliculogenesis and circulating estradiol-17b during autumnal transition in the mare. Master’s thesis, Southern Illinois University, Carbondale, 1993. 128. Sharp DC, Davis SD. Vernal transition. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993, p. 134.
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Chapter 182
The Abnormal Estrous Cycle Patrick M. McCue and Ryan A. Ferris
Introduction Abnormalities of the equine estrous cycle can be categorized into conditions associated with follicular development, ovulation, luteal function, and sexual behavior. Some abnormalities are transient physiological conditions and others are congenital or acquired pathological problems that may be permanent. A correct diagnosis is central to the design and implementation of an optimal management or therapeutic plan. Acquisition of a complete medical and reproductive history is critically important. A thorough physical examination and reproductive evaluation should be performed to identify general medical and reproductive abnormalities. It should be noted that a combination of issues may be present in a mare exhibiting abnormal estrous cycles.
Abnormalities of follicular development The causes of abnormal follicular development are listed in Table 182.1.
Season Approximately 80% or more of mares enter a prolonged anovulatory phase during the winter months.1 Factors that influence seasonal reproductive activity in mares include ambient photoperiod, environmental conditions (i.e., temperature, weather), nutrition, age, breed, and housing. Young mares and mares that gave birth the previous season have the highest probability of becoming anestrous, whereas barren mares housed indoors have a lower probability of becoming anovulatory during the late fall and winter.2, 3 Nutrition plays a role in seasonality as mares in poor body condition going into the fall
and winter months are more likely to become acyclic than mares with higher body condition scores.4 Increasing the plane of nutrition5 and allowing mares to graze on green grass pasture6, 7 has been reported to stimulate ovulation earlier in the year. Breed is also a factor that may affect seasonality in mares as pony mares are more likely to become anestrous in the winter and begin cycling later in the spring than horse mares.8, 9 A progressive increase in the duration of ambient photoperiod stimulation in the spring modulates the production of melatonin from the pineal gland. Melatonin, a protein hormone released during the hours of darkness, inhibits secretion of GnRH from the hypothalamus of the mare. A decrease in duration of melatonin exposure in the vernal transition leads to an increase in GnRH production and secretion.1 The anterior pituitary increases secretion of gonadotropins and ovarian follicular development begins in response to the increased GnRH output.10 Mares may exhibit multiple waves of follicular growth and regression during the transition period.11, 12 Pony mares have been noted to develop 3.7 ± 0.9 waves of follicles 30 mm in diameter during transition.11 The total number of follicles present on each ovary increases during the transition, and ultrasound examination classically reveals a ‘grape-like’ cluster of small to medium-sized follicles. The transition period may last for 50 to 70 days or more prior to the first ovulation of the season. Once a mare has ovulated, she will generally continue to cycle at regular intervals. Body condition Poor body condition has been associated with altered reproductive performance in mares, including a delay in the onset of the first ovulation of the breeding season, decreased pregnancy rates, increased embryonic loss, and increased gestation rates.13, 14 In
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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The Abnormal Estrous Cycle Table 182.1 Abnormalities of follicular development Common causes
Less common causes
Season Poor body condition Advanced age Equine Cushing disease Iatrogenic therapy
Chromosomal abnormalities Granulosa cell tumor Postpartum anestrus Persistent endometrial cups
contrast, mares with higher body condition scores tend to cycle throughout the winter.4 In general, reproductive efficiency is enhanced when mares are maintained at a body condition score of 5.0 or above.4,13–15 Advanced age Geriatric mares may have reduced reproductive performance due to age-related changes in ovarian function, uterine health, perineal conformation, and other factors.16 Older mares (> 20 years of age) may experience a delay in their initial ovulation of the year by an average of 2 weeks.17 Geriatric mares may have a longer interovulatory interval than younger mares due to a longer follicular phase.18, 19 A lengthening of the follicular phase in association with elevated gonadotropin concentrations may indicate impending reproductive senescence in older mares.20 Complete ovulation failure or ovarian senescence has been observed in aged mares and may be due to an insufficient number of primordial follicles. The incidence of equine Cushing disease is also increased in older mares and may contribute to poor reproductive performance. The sporadic nature of estrous cycles and age-related changes in uterine health of older mares necessitates enhanced reproductive management to optimize success. Equine Cushing disease Equine Cushing disease (ECD) or pituitary pars intermedia dysfunction (PPID) affects 0.075% to 0.5% of the equine population.21, 22 The physiological basis for ECD may be a loss of dopaminergic inhibition by the hypothalamus, resulting in hypertrophy, hyperplasia, or adenoma formation of the pituitary pars intermedia. A majority of horses diagnosed with ECD are older, with the average age being approximately 20 years. Consequently, the decrease in reproductive efficiency in mares with ECD may be partly due to advanced age. Clinical signs of ECD include hirsutism and abnormal hair-coat shedding patterns, polyuria, polydipsia, and hyperhidrosis.23, 24 Reproductive abnormalities reported in mares with
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ECD have included abnormal estrus activity, estrus suppression, and reduced fertility.25–27 Diagnostic tests for ECD include measurements of serum glucose, insulin, adrenocorticotropic hormone (ACTH) and cortisol levels and dexamethasone suppression, ACTH stimulation, and thyrotropin-releasing hormone response tests. Medical management of ECD may include administration of pergolide mesylate, a dopamine receptor agonist, at a dosage of 0.5–2 mg q 24 h per adult horse, or cyproheptadine, a serotonin antagonist, at a dose of dose of 0.25 mg/kg q 24 h, given once in the morning.28 Iatrogenic therapy Administration of anabolic steroids, glucocorticoids, and gonadal steroids all inhibit follicular development. In addition, vaccination against GnRH or administration of a potent GnRH agonist can adversely affect ovarian function. The common denominator of these iatrogenic treatments is suppression of pituitary gonadotropin secretion. Anabolic steroid administration at low doses may cause aggressive or stallion-like behavior, while high doses may inhibit ovarian activity and result in failure of follicular development and ovulation.29 Progestins given to cycling mares for the suppression of estrus or synchronization of ovulation will suppress luteinizing hormone (LH) secretion, but will have a limited effect on levels of follicle stimulating hormone (FSH). Consequently, mares treated with progestins will still have active ovaries. Addition of estradiol to the progestin regimen will result in suppression of both FSH and LH, and is associated with a profound inhibition of follicular development. Agonists of gonadotropin releasing hormone (GnRH) have been used to stimulate follicular development in seasonally anestrous mares and induce ovulation in cycling mares. An implant (OvuplantTM , Ft. Dodge Animal Health, Ft. Dodge, Iowa) containing the GnRH agonist deslorelin acetate approved for induction of ovulation in mares was reported to cause a decrease in follicular development and an increase in the interovulatory interval in some mares.30, 31 This effect is caused by prolonged suppression or downregulation of pituitary FSH secretion. Administration of prostaglandins 7–8 days after deslorelininduced ovulation increases the risk of delayed follicular development. Removal of the deslorelin implant 48 hours after administration has been reported to prevent the adverse effects on pituitary function.32 For ease of removal, the implant may be inserted subcutaneously by using the implant device provided by the manufacturer in a region of the vulva that has been infused with a small amount of local anesthetic.32 After ovulation, the implant may be removed
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through the original tract created by the implant device by using gentle topical pressure. Interestingly, administration of high doses of GnRH agonists to mares with the intent of inducing pituitary downregulation and blocking follicular development has not been universally successful in mares.33 Vaccination of mares against GnRH is potentially an effective and eventually reversible technique for estrus suppression and/or contraception. Vaccination against GnRH results in production of antiGnRH antibodies that block the biological activity of GnRH released from the hypothalamus and a subsequent reduction of LH secretion from the anterior pituitary. A commercial GnRH vaccine has been licensed in Australia (EquityTM , Pfizer Animal Health, Australia) ‘for use in the control of estrus and estrusrelated behavior in fillies and mares not intended for breeding’. The vaccine is administered as a two-dose series, with the second dose given 4 weeks after the initial dose. Peak antibody response occurs approximately 2 weeks after the second dose.34, 35 Immunization results in a reduction of ovarian activity, with the ovaries in most mares resembling those of seasonally anovulatory mares within 4 weeks after the second dose, and a corresponding reduction in estrus-related behavior. An increase in ovarian activity and a return to estrus may be observed in a majority of vaccinated mares as the concentrations of anti-GnRH antibodies in circulation decrease over time. Card and co-workers36 reported that live foal rates were similar between mares receiving a placebo treatment and mares vaccinated against GnRH in subsequent years after the effects of the vaccine had disappeared. However, it has also been noted that some young mares vaccinated against GnRH when they were in training have subsequently failed to develop follicles or ovulate for 2 to 3 years or more after the end of their performance career.37 The incidence rate of prolonged failure of follicular development following vaccination against GnRH has not been determined. Chromosomal abnormalities Chromosomal abnormalities, especially of the sex chromosomes, have been associated with infertility in the horse. The prevalence of sex chromosome abnormalities in the mare has been reported to be less than 3%.38 A chromosomal abnormality may be suspected in a mare of breeding age with primary infertility and gonadal hypoplasia, especially if the mare is presented during the physiological breeding season and has not been administered exogenous hormones. The most commonly reported chromosomal abnormality of the horse is 63,X gonadal dysgenesis, in which only a single sex chromosome is present.39 The condition may occur when the sex chromosome
pair fails to separate during meiosis, producing one gamete without a sex chromosome and another with two sex chromosomes. The equine condition is analogous to Turner syndrome in humans. The 63,X (or XO) condition has been detected in most domestic horse breeds, including draft and Miniature breeds. Horses with gonadal dysgenesis develop as phenotypic females because of the absence of a Y sex chromosome, or more specifically absence of the sexdetermining region (Sry) normally present on the Y chromosome. Affected horses are often small in size for their age and breed and have small ovaries with limited or no follicular development. The uterus and cervix are generally small and flaccid and the endometrial glands are hypoplastic. The external genitalia are female, but the vulva may be smaller than normal and there is no clitoral hypertrophy. XO mares may exhibit anestrus or irregular estrous behavior and occasionally stand to be mated. True XO mares are considered to be sterile. However, mares with a mosaic or chimeric karyotype (i.e., 63,XO/64,XX) are not always small in stature and some have been reported to produce a foal. Mosaic mares account for approximately 15–30% of all cases of gonadal dysgenesis. Numerous other chromosomal abnormalities have been reported in the mare in addition to XO gonadal dysgenesis.40,41 Mares with chromosomal abnormalities may present with failure of follicular development, subfertility, repeated early embryonic loss, anatomical (phenotypical) abnormalities or other reproductive conditions.39–41 Diagnosis of a chromosomal abnormality is made by chromosome analysis or karyotyping. A fresh blood sample collected into acid citrate dextrose or heparin may be sent by overnight courier to a laboratory specializing in animal karyotyping. Granulosa cell tumors Granulosa cell tumors (GCT) are almost always unilateral, slow growing, and benign.42 On transrectal ultrasound examination, the affected ovary may appear multicystic or honeycombed, or it may present as a solid mass or as a single large cyst. The key to clinical diagnosis is the status of the contralateral ovary, which is usually small and inactive in the presence of a GCT. Mares with a GCT may exhibit prolonged anestrus, aggressive or stallion-like behavior, or persistent estrus.43–45 Granulosa cell tumors are hormonally active, and clinical diagnostic assays for detection of a GCT include measurement of inhibin, testosterone, and progesterone. Inhibin is elevated in approximately 90% of the mares with a GCT. Testosterone is elevated in approximately 50–60% of affected mares and is usually associated with stallionlike behavior. Progesterone concentrations in mares
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The Abnormal Estrous Cycle with a GCT are almost always below 1 ng/mL, since normal follicular development, ovulation, and corpus luteum formation do not occur. Treatment is surgical removal of the affected ovary. Ovulation from the remaining ovary will occur approximately 6–8 months after tumor removal. Anecdotal reports indicate that attempts at inducing follicular development in the remaining ovary with GnRH or equine follicle stimulating hormone have had limited success.
Postpartum anestrus Mares may exhibit a temporary failure of follicular development or ovulation after foaling. A majority of mares develop follicles and ovulate early in the postpartum period and continue to cycle if they do not become pregnant at a foal heat breeding. Alternatively, a foal heat ovulation may be followed by a variable period of anestrus or anovulation until the mare resumes normal cyclic activity. Finally, some postpartum mares have no significant follicular development or may exhibit moderate follicular development without ovulation. Mares in the latter groups may remain anestrus or anovulatory for weeks or months before cyclic ovarian activity is initiated. A majority of mares that do not cycle after giving birth are mares that foal early in the year. ‘Lactational anestrus’ may be caused by the combined effects of lactation, body condition, and season. Maintenance of late-term pregnant mares due to foal between January and March (northern hemisphere) under a stimulatory artificial photoperiod for the last 2–3 months of pregnancy may result in a decreased length of gestation,46 decreased incidence of postpartum anestrus,3 and a shortened interval from foaling to first ovulation.47
Persistent endometrial cups Persistence of endometrial cups has recently been described in mares with an early termination of pregnancy (i.e., abortion) and mares that carried a normal pregnancy to term.48–50 Persistent endometrial cups have been associated with failure to show behavioral estrus, complete ovarian inactivity, sporadic follicular growth, and luteinization of partly developed follicles.49, 50 Diagnosis is based on observation of endometrial cups on ultrasonography per rectum, videoendoscopic evaluation of the uterine lumen, and measurement of circulating equine chorionic gonadotropin (eCG). The functional lifespan of the persistent cups has been estimated to be 6 to 30 months.49
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Table 182.2 Abnormalities of ovulation Common causes
Less common causes
Anovulatory follicles Stress Equine Cushing disease Age
Failure of ovulation-inducing agents Iatrogenic therapy
Abnormalities of ovulation The causes of abnormalities of ovulation are listed in Table 182.2. Anovulatory follicles Ovulation failure is a significant cause of reproductive inefficiency and economic loss in the mare. Since affected mares do not ovulate, fertilization and pregnancy cannot occur. In addition, ovulation failure results in a prolonged interovulatory interval, which translates into more days not pregnant and higher subsequent costs. The incidence of ovulation failure has been reported to range from 3.1 to 8.2% of all equine estrous cycles. Older horses have a higher incidence of ovulation failure, as approximately 13% of mares greater than 15 years of age experience ovulation failure at least once during a given breeding season.51 A high percentage of mares that develop one anovulatory follicle will experience other cycles culminating in ovulation failure during the same breeding season. An occasional mare will develop anovulatory follicles during repeated estrous cycles throughout a breeding season. It is difficult with our existing diagnostic techniques to predict if a follicle is destined for ovulation or ovulation failure when a mare first comes into heat. The initial growth patterns of follicles that ultimately fail to ovulate appear normal and affected mares exhibit behavioral estrus and have normal patterns of uterine edema in a majority of cases. Two distinct types of anovulatory follicles exist in the mare. Approximately 15% of anovulatory follicles remain ‘cystic’ and do not luteinize. These ‘persistent anovulatory follicles’ (PAF) may remain for several weeks and will eventually regress spontaneously. Attempts to induce ovulation or luteinization of affected follicles with human chorionic gonadotropin (hCG) or GnRH agonists have not been successful.51 Administration of prostaglandins to a mare with a non-luteinized anovulatory follicle will have limited efficacy. Approximately 85% of anovulatory follicles luteinize over a period of a few days. The process usually begins with hemorrhage into the follicular lumen, which can be detected ultrasonically as
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Mare: The Estrous Cycle mares, which may ovulate early in the breeding season within a few weeks of arrival and then fail to develop significant follicular activity for 2 or more months. It is recommended that maiden mares be segregated early in the breeding season and individually managed with respect to nutrition, temperature, and social stress. Equine Cushing disease
Figure 182.1 Ultrasound photograph of a luteinized anovulatory follicle.
scattered free-floating echogenic spots within the follicular fluid (previously termed anovulatory hemorrhagic follicles, AHF). A progression from echogenic particles, to strands traversing the follicular lumen, to complete infiltration of the follicular lumen with echogenic material may be observed ultrasonographically (Figure 182.1). Mares with ‘luteinized anovulatory follicles’ (LAF) with no evidence of bleeding were previously referred to as luteinized unruptured follicles (LUF). Both types will have elevated serum progesterone levels. Administration of prostaglandins will result in destruction of the luteal tissue, a rapid decline in serum progesterone levels, and a return to estrus. It is recommended that prostaglandins be administered 7 to 10 days after initial detection of echogenic material within the follicular lumen or after complete infiltration has occurred. The optimal timing of prostaglandin therapy may vary between mares, as the rate of luteinization and subsequent progesterone production may be delayed in some mares.
Failure of ovulation in mares affected by ECD may be related to alterations in pituitary gonadotropin secretion secondary to destruction of gonadotrophs or increased levels of glucocorticoids and/or androgens produced by the adrenal cortex. Treatment of affected mares with the dopamine agonist pergolide mesylate has resulted in clinical improvement medically28 and anecdotally has resulted in enhanced reproductive performance. Age Older mares have an increased incidence of spontaneous ovulation failure51 and a decreased response to administration of an ovulation-inducing agent.56, 57 Iatrogenic therapy Prolonged administration of glucocorticoids58, 59 and progestins60 during the follicular phase of the estrous cycle have been associated with suppression of estrus, reduction of plasma LH, and an increased incidence of ovulation failure. Short-term administration of dexamethasone to manage mares with matinginduced endometritis is unlikely to adversely affect pituitary function or ovulation.61, 62
Abnormalities of luteal function The causes of shortened luteal phase are listed in Table 182.3.
Stress Stress due to medical conditions, adverse weather, transportation, a new environment, housing, social rank, overcrowding, fighting, and other factors can all lead to alterations of the estrous cycle in domestic animals.52 The adverse effects of stress are presumably mediated through the hypothalmo– pituitary–adrenocortical axis, ultimately manifested through an increase in cortisol levels.53–55 While experimental ‘transport stress’ models have not clearly demonstrated adverse effects of stress on reproductive parameters, it is not unusual to see a new mare arrive on a breeding farm and have an altered estrous cycle (i.e., follicular regression without ovulation). This is most commonly observed in maiden
Endometritis The luteal phase refers to the period of the estrous cycle dominated by the corpus luteum. It begins with formation of the corpus luteum after ovulation and ends when luteolysis occurs 13 to 15 days later. Premature destruction of the corpus luteum is associated Table 182.3 Shortened luteal phase Common causes
Less common causes
Acute endometritis Iatrogenic therapy
Endotoxemia Cervical or uterine manipulations
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The Abnormal Estrous Cycle with a return to estrus and a decrease in the interovulatory interval. The most common cause of premature luteolysis in the mare is endometritis. Acute bacterial infections of the uterus can cause sufficient synthesis and release of prostaglandins by the endometrium to cause luteal regression.45 Inoculation of bacteria into the uterus during diestrus was reported to result in a decrease in the interovulatory interval.63 The clinical significance is that any mare that exhibits a shortened diestrus or an early return to estrus should be examined for infectious endometritis. A cytological examination, culture, and/or biopsy of the uterus would be indicated.
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Table 182.4 Prolonged luteal phase Common causes
Less common causes
Pregnancy
Chronic degenerative endometritis
Embryonic loss after maternal recognition of pregnancy
Iatrogenic therapy
Fetal loss after endometrial cup formation Luteinized anovulatory follicles Idiopathic (insufficient PGF release) Autumnal transition Diestrus ovulation
Endotoxemia Medical conditions that result in endotoxemia, such as gastrointestinal disease, colitis, pleuropneumonia, retained placenta, and peritonitis have the potential to alter luteal function in a mare. Bacterial endotoxins induce the release of prostaglandins, which can result in regression of a corpus luteum and a decrease in the length of the estrous cycle.64 In pregnant mares, endotoxemia can result in fetal death mediated through a PGF2α -induced reduction in luteal function.65 Iatrogenic therapy Administration of prostaglandins to a mare with a mature corpus luteum (i.e., on day 5 post-ovulation or later) will result in complete luteolysis, as noted by a rapid decrease in progesterone, and a return to estrus. Administration of prostaglandins on the day of ovulation or during the first 1 or 2 days after ovulation may adversely affect development of a corpus luteum.66, 67 Progesterone concentrations are commonly reduced and pregnancy rates may be affected. Cervical or uterine manipulation Reproductive techniques that involve manipulation of the cervix and/or uterus during the luteal phase, such as collection of a uterine biopsy or non-surgical transfer of an embryo, may occasionally result in luteolysis.68–70 Administration of non-steroidal antiinflammatory drugs may inhibit the inflammatory response and reduce prostaglandin release in mares following embryo transfer.71 Pregnancy The causes of prolonged luteal phase are listed in Table 182.4. The most common cause of prolongation of the lifespan of the corpus luteum that forms after ovulation is pregnancy. A signal from the developing embryo as it migrates through the uterus during the
critical period of ‘maternal recognition of pregnancy’ results in an alteration of endometrial prostaglandin secretion. This allows the initial corpus luteum to be maintained. A corpus luteum in a non-pregnant mare that fails to regress after 14 or 15 days is considered to be abnormally retained (i.e., a persistent corpus luteum) and the condition is commonly but erroneously referred to as pseudopregnancy. The most common causes of a persistent corpus luteum: are (i) ovulations late in diestrus; (ii) embryonic loss after the time of maternal recognition of pregnancy; and (iii) chronic uterine infections. Inadequate prostaglandin release from the endometrium has been suggested as a cause of spontaneous persistence of the corpus luteum, but documentation is limited. Diestrus ovulation Ovulations anywhere in the luteal phase may result in formation of a persistent corpus luteum.72 The later a corpus luteum forms as a diestrus ovulation the less responsive it is to prostaglandins that are released from the endometrium at the end of the initial luteal phase. Embryonic loss The presence of an embryo during the time of maternal recognition of pregnancy will result in altered prostaglandin release and preservation of the corpus luteum. Mares that lose their pregnancy after the period of maternal recognition will often retain their corpus luteum.73 In addition, mares that experience pregnancy loss after endometrial cup formation may continue to produce progesterone from accessory corpora lutea in response to continued secretion of equine chorionic gonadotropin (eCG).
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Chronic degenerative endometritis Mares with severe, chronic degenerative endometritis, as noted in mares with pyometra, may have an increased incidence of persistence of the corpus luteum.74, 75 The cause is a decreased ability of the endometrium to release sufficient prostaglandins to induce luteolysis. Infusion of saline into the uterus of mares with severe degenerative endometritis resulted in lower release of uterine PGF2α than observed in normal mares.75 Luteinized anovulatory follicles Luteinized anovulatory follicles may persist for several weeks and result in a markedly prolonged interovulatory interval.51 Administration of prostaglandins will result in regression of a luteinized anovulatory follicle, if administered after the structure is completely luteinized. It is recommended that prostaglandins be given > 9 days after the first recognition of echogenic spots in the follicular lumen or visualization of luteal tissue on the periphery of the LAF. Iatrogenic therapy Iatrogenic causes of prolongation of luteal function include insertion of an intrauterine device, chronic administration of oxytocin during the luteal phase of the estrous cycle, and ovulations during administration of progestins. Nie and others76 reported on the efficacy of a glass ball inserted into the uterus of mares for suppression of estrus. A 35 mm sterilized glass marble was placed into the body of the uterus after ovulation and left in utero (Figure 182.2). Luteal function was extended in 7 of 18 mares (39%) in which the glass ball was retained. The mean duration of luteal lifespan in mares with extended luteal function was 87 days (range, 76 to 109 days). Re-
Figure 182.2 Ultrasound photograph of a marble in the uterus of a mare.
Figure 182.3 Marble removed from a uterus of a mare.
moval of the glass ball resulted in an eventual return to estrus (Figure 182.3). In the breeding season following removal of the glass ball, 17 of 23 mares (74%) conceived. It was hypothesized that the prolonged suppression of estrus was due to maintenance of the corpus luteum (pseudopregnancy) caused by an alteration in prostaglandin secretion from the endometrium. In another study, water-filled plastic balls 20 mm in diameter were placed into the uterus 2 to 4 days after ovulation.77 The intrauterine device (IUD) induced prolongation of the luteal phase in 9 of 12 mares (75%) treated. The mean length of diestrus was 57.0 days. Evaluation of PGF2α metabolite levels suggested that the IUD may have prevented the endometrial cells from releasing PGF2α . Chronic administration of oxytocin to mares during diestrus, either by a subcutaneous minipump78 or intramuscular injections every 12 hours,79 has been reported to result in prolongation of luteal function. Stout and co-workers78 noted that luteolysis was prevented in 4 of 5 mares treated by continuous infusion of oxytocin from day 8 to 20 after ovulation. It was hypothesized that treatment suppressed the upregulation of oxytocin receptors in the endometrium, which altered the normal physiological pathway toward prostaglandin release. Vanderwall and colleagues79 reported that administration of 60 units of oxytocin every 12 hours from day 7 through day 14 resulted in prolonged luteal function in 6 of 6 mares treated. All 6 oxytocin-treated mares maintained progesterone concentrations above 1.0 ng/mL through day 30 post-ovulation. It was suggested that this regimen could be used to induce long-term suppression of estrus in mares that would be reversible by administration of prostaglandins. Altrenogest treatment, when initiated late in diestrus or early in estrus, has been associated with a high incidence of persistence of the corpus luteum.80, 81 The mechanism responsible for the failure of luteolysis in these studies is not clear.
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The Abnormal Estrous Cycle Diagnosis of a persistent corpus luteum may be made by behavioral observations, palpation, ultrasonography, analysis of plasma progesterone concentrations or clinical response to prostaglandin administration. Progesterone concentrations > 1.0 ng/mL are indicative of the presence of luteal activity. Mares with a persistent corpus luteum will have good tone in the cervix and uterus on palpation, and the cervix will appear tight and dry on vaginal speculum examination. A retained corpus luteum is almost always visible on ultrasound. Persistence of the corpus luteum may be resolved by administration of a single dose of prostaglandins (PGF2α, 10 mg; or cloprostenol, 250 µg). However, it is critical that a mare be examined by ultrasound prior to administration of prostaglandins to confirm that the mare is not, in fact, actually pregnant and that no other reproductive abnormalities are present. Inadvertent or inappropriate administration of prostaglandins to a pregnant mare will result in loss of the pregnancy. Autumnal transition Occasionally, mares will experience a prolonged period of progesterone secretion at the end of the physiological breeding season associated with either a persistent corpus luteum or a hemorrhagic anovulatory follicle.
Behavioral abnormalities Mares are seasonal breeders, with regular estrous cycles occurring during the spring and summer months. Ovarian follicular activity is minimal in most mares throughout the winter when length of daylight is short. The lack of estrogen production results in behavioral anestrus in most mares during the winter. An increase in daylength in the spring stimulates ovarian activity. The spring or vernal transition period is associated with estrogen production during each wave of follicular growth82 and results in expression of behavioral estrus. Regression of follicles is associated with a decrease in estrogens and a decrease in estrous activity. The transition period is marked behaviorally by irregular or, in some instances, prolonged expression of estrus. Reproductive behavior during the physiological breeding season is dictated by the effects of estradiol and progesterone. In ovariectomized mares, administration of estradiol will stimulate estrous behavior, while administration of progesterone inhibits estrous behavior.83 In intact cycling mares, the presence of estradiol produced by the developing dominant follicle, in the absence of progesterone, will elicit behavioral estrus. Progesterone produced by the corpus
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Table 182.5 Failure to exhibit estrus Common causes
Less common causes
Seasonal anestrus
Failure of follicular development (see above)
Silent heat (maiden, foaling) Persistent corpus luteum Luteinized anovulatory follicle Pregnancy Iatrogenic therapy
luteum that forms after ovulation will result in behavioral diestrus, or active rejection of the advances of a stallion. It is generally considered that mares will be in estrus for 5 to 7 days and out of heat (diestrus) for approximately 14 days out of a 21-day estrous cycle.84 One research study reported that the average (± standard deviation) duration of estrus and diestrus were 7.7 ± 2.9 and 15.8 ± 2.5 days, respectively.85 Abnormalities of reproductive behavior in mares can be classified as failure to exhibit estrus (Table 182.5), persistent estrus (Table 182.6), and aggressive or stallion-like behavior (Table 182.7). Failure to exhibit estrus Failure to show heat can be a significant management problem for mares in a natural service breeding program. Any mare that does not come into estrus or return to estrus within 14 to 16 days after she goes out of heat should be examined (Figure 182.4). Pregnancy Pregnancy is often overlooked as a cause of failure to return to estrus. Almost every year we are presented with a mare to examine for breeding that is already pregnant, and in those situations, the owner is often unaware that the mare had been bred. Seasonal anestrus Mares in ‘deep’ seasonal anestrus have minimal ovarian follicular development (i.e., follicles < 20 mm in Table 182.6 Persistent or inappropriate estrus Common causes
Less common causes
Spring transition
Granulosa cell tumor Adrenal steroid production Pregnancy Phytoestrogens Iatrogenic therapy
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Table 182.7 Aggressive or stallion-like behavior Common causes
Less common causes
None
Pregnancy Estrus Granulosa-theca cell tumor Iatrogenic therapy Developmental abnormality
progesterone exposure. Treatment for both a persistent corpus luteum and luteinized anovulatory follicle consists of administration of a single dose of prostaglandins. It is recommended that a mare be examined to confirm that she is not pregnant prior to administration of prostaglandins. Mares should return to estrus within 4 to 5 days after treatment.
Iatrogenic therapy diameter) and low levels of estradiol (< 5 pg/mL). Behavior of mares with seasonal ovarian inactivity may range from unreceptive (teasing out), to indifferent (passive), to receptive (teasing in).9, 86 Adrenal steroids may be responsible for expression of estrus in winter acyclic mares.87 Silent heat Occasionally mares will develop a dominant follicle and ovulate without showing estrus, a condition referred to as ‘silent heat’. The common factors associated with silent heat are teasing an inexperienced maiden mare and teasing a mare with a foal at side. Other factors noted are inhibition of behavior of a subordinate mare by a dominant mare, preferences between individual stallions and mares, type of teasing technique, and inexperienced personnel teasing or interpreting mare behavior.84, 88 Persistent corpus luteum Another cause of failure to show heat is persistence of the corpus lutum (pseudopregnancy).72 Non-pregnant mares normally have a luteal phase approximately 14 days in duration. Failure of luteal regression results in continued secretion of progesterone from the corpus luteum for up to 2 to 3 months. Development of a luteinized anovulatory follicle can also result in a prolonged period of
In some instances, failure to show heat is due to iatrogenic treatment with one or more medications during training or performance. It is quite possible that the medications may have been administered without direct knowledge or consent of the mare owner. Consequently, the medical history provided for a mare with failure to show heat or failure to cycle may not accurately reflect the entire story. Possibilities include, but are not limited to, administration of progesterone, certain progestins (e.g., altrenogest), or anabolic steroids; vaccination against GnRH; and/or insertion of an intrauterine device (e.g., a marble). Perhaps the most difficult cause of failure to exhibit estrus to confirm is previous administration of a vaccine against GnRH. In one study, nine mares were immunized twice at an interval of 4 weeks.35 All mares ceased cycling within 8 weeks of the first vaccination. Signs of estrus were absent in four treated mares during the period of ovarian suppression. The other five mares showed sporadic or prolonged estrus not obviously related to follicular activity. Follicular activity in one mare was suppressed for more than 100 weeks after the first vaccination. There are currently no diagnostic tests readily available to practitioners to detect the presence of anti-GnRH antibodies. Herbal supplements (e.g., chaste tree berries) have also been administered in an attempt to suppress estrus in performance mares.89 Medications used unsuccessfully in an attempt to suppress estrus or maintain pregnancy in mares include medroxyprogesterone acetate and other synthetic progestins90–92 or cattle implants containing low levels of progesterone and estrogen.93
Failure of follicular development Failure of follicular development should also be considered as a potential cause of failure to show heat. Causes of failure of follicular development are reviewed elsewhere in this chapter.
Persistent or inappropriate estrus Figure 182.4 A mare in diestrus.
Expression of behavioral estrus, or heat, in mares can have a profound negative effect on training
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in mares with GCTs and not always clearly related to the predominant behavior expressed.45, 97 In fact, estradiol levels in most mares with GCTs are not elevated above the range of normal cycling mares (20 to 45 pg/mL). Iatrogenic therapy
Figure 182.5 Mare exhibiting estrus when teased to a stallion.
and performance (Figure 182.5). Examples of adverse behaviors noted in performance mares when in heat include attitude changes, tail swishing, difficulty in training, squealing, horsing, excessive urination, kicking, decrease in performance, and colic-like discomfort. It has been estimated that 5 to 10% of pregnant mares exhibit estrous behavior.94 It has been reported that the incidence of behavioral estrus in pregnant mares carrying a female conceptus is higher than mares carrying a male conceptus.95 In addition, it was noted that a majority of estrus signs were exhibited on days 12 to 20 of pregnancy. Consequently, expression of sexual receptivity does not always indicate that a mare is open or non-pregnant. Whenever possible, pregnancy status should be confirmed by transrectal palpation and ultrasonography. It is not uncommon for seasonally acyclic mares, ovariectomized mares, or mares with a chromosomal abnormality (i.e., 63,XO, gonadal dysgenesis) to occasionally exhibit behavioral estrus and stand to be mated. It has been estimated that up to 90% of ovariectomized mares and 80% of seasonally anovulatory mares show signs of estrus.96 The physiological trigger for behavioral estrus in a mare lacking a dominant follicle is likely to be secretion of steroids from the adrenal glands. Administration of dexamethasone was noted to suppress the various manifestations of behavioral estrus in ovariectomized mares.87 Dietary sources of estrogens (i.e., phytoestrogens) have also been proposed as a potential cause of behavioral estrus in acyclic mares, but no controlled research has been reported in this area.
Administration of estrogens such as estradiol 17β or conjugated estrogens has been used for many years to induce behavioral estrus in ‘tease’ or ‘jump’ mares in artificial insemination programs. Any mare without a functional corpus luteum (i.e., seasonally acyclic mares and ovariectomized mares) should begin to develop uterine edema and express behavioral estrus within 1 to 2 days after estrogen administration. Aggressive or stallion-like behavior Display of true stallion-like behavior is considered rare in mares.98 In some instances, such behavior may be due to an elevation in endogenous testosterone of either gonadal or adrenal origin.99 Estrus Elevated production of testosterone from the ovary during estrus has been implicated in stallion-like behavior in mares.98 Observations over a 3-year period on a herd of research horses showed that mounting behavior occurred during approximately 0.6% of estrous cycles. In most instances the mare standing was in estrus, and the mare mounting showed signs of estrus when exposed to a stallion, even though she showed only male behavior when exposed to other mares. Testosterone concentrations were higher in mounting mares (17.7 ± 2.3 pg/mL) than in standing mares (10.9 ± 0.5 pg/mL). Concentrations of testosterone increased during estrus and reached peak levels from 2 days before to 1 day after ovulation.98
Granulosa cell tumor Persistent estrus is one of the classic behavioral signs exhibited by a mare with a granulosa cell tumor. Meagher and co-workers44 noted that 14 of 63 (22.2%) mares with GCT showed continuous estrus. However, plasma concentrations of estradiol are variable
Figure 182.6 A granulosa-theca cell tumor.
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Parturition Figure 182.7 A mare with a granulosa theca cell tumor mounting a normal mare in estrous. (Courtesy of University of California, Davis).
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Granulosa cell tumor One of the most commonly recognized causes of stallion-like behavior in mares is the presence of a granulosa-theca cell tumor (Figure 182.6). The presence of thecal cells within the interstitial tissue of the tumor is associated with an increase in blood testosterone levels (Jubb et al., 1985 [91]).100 Testosterone concentrations were reported to be elevated in 40 to 50% of mares with ‘granulosa cell tumors’.45 Meagher and co-workers44 reported that 29 of 63 (46%) affected mares showed stallion-like behavior (Figure 182.7). Mares expressing stallion-like behavior often have testosterone levels above 100 pg/mL, whereas normal cycling mares have testosterone levels of 20 to 45 pg/mL.42, 45 Surgical removal of the affected ovary results in a rapid decline in testosterone levels and an eventual return to normal behavior. The hallmark for clinical diagnosis is a unilaterally enlarged ovary and a small, inactive contralateral ovary. Endocrine tests may reveal an elevation in plasma inhibin in approximately 90% of mares with a granulosa cell tumor.42 Pregnancy Pregnant mares may occasionally exhibit stallionlike behavior. McDonnell101 noted that the abnormal behavior was first noted during the fourth or fifth month of pregnancy, continued until term, and then ceased. Testosterone levels in the blood of pregnant mares increases to peak concentrations by the sixth or seventh month of gestation and then declines to basal levels by the date of foaling102 (Figure 182.8). The source of the elevated testosterone is presumed to be the fetal gonads. Clinically, we have evaluated mares that referred for a combination of stallion-like behavior, elevated testosterone, and
Figure 182.8 Testosterone levels in mares throughout pregnancy (adapted from Silberzahn et al.102 ; see Ginther,1 page 427).
bilaterally enlarged ovaries only to determine that they were pregnant with multiple corpora lutea on each ovary. Developmental abnormalities Developmental abnormalities may result in the presence of testes and elevated testosterone in a phenotypic mare. The most common such developmental abnormality of horses is male pseudohermaphroditism (XY sex reversal), in which the horse has a male (64,XY) karyotype, cryptorchid testes, absence of female internal genitalia, and an external phenotype that is either female or somewhat ambiguous103 (Figure 182.9). Cases of stallionlike behavior associated with a variety of other chromosomal or developmental abnormalities have also been reported.103, 104 In some instances, geneticbased developmental abnormalities appear to be inherited.105–107 Iatrogenic therapy Administration of exogenous androgens or anabolic steroids has also been reported to result in alterations of normal behavior. Long-term administration of anabolic steroids (boldenone undecylenate or nandrolone decanoate) resulted in a suppression of normal estrous behavior and an increase in mounting behavior and aggressiveness.29, 108 Another study also reported that anabolic steroid (boldenone undecylenate) treatment suppressed visible signs of estrous behavior in young fillies and resulted in aggression
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Figure 182.9 Enlarged clitoris in a male pseudohermaphrodite horse. Adapted from Silbersahn et al. 1984.102
between horses and stallion-like herding behavior.109 In addition, administration of anabolic steroids was noted to be associated with an increased incidence of ‘contact agonistic behaviors’, such as biting and kicking.110 A decrease in reproductive performance and an increase in clitoral size have also been noted following anabolic steroid therapy.29, 108, 109
References 1. Ginther OJ. Reproductive Biology of the Mare. Cross Plains WI: Equiservices, 1992. 2. Nagy P, Guillaume D, Daels P. Seasonality in mares. Anim Reprod Sci 2000;60–61:245–62. 3. Palmer E, Driancourt MA. Some interactions of foaling, photoperiod and ovarian activity in the equine. Livestock Prod Sci 1983;10:197–210. 4. Gentry LR, Thompson DL Jr, Gentry GT Jr, Davis KA, Godke RA, Cartmill JA. The relationship between body condition, leptin, and reproductive and hormonal characteristics of mares during the seasonal anovulatory period. J Anim Sci 2002;80:2695–703. 5. Van Niekerk CH, Van Heeden JS. Nutrition and ovarian activity of mares early in the breeding season. J S Afr Vet Med Assoc 1972;45:351–60. 6. Allen WR. Control of oestrous and ovulation in the mare. In: Crighton DB, Haynes NB, Foxcroft GR, Lammings GE. (eds) Control of Ovulation. Oxford: Butterworth, 1978; pp. 453–68.
1765
7. Carnevale EM, Hermenet MJ, Ginther OJ. Age and pasture effects on vernal transition in mares. Theriogenology 1997;47:1009–18. 8. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Seasonal influence on equine follicle dynamics. Anim Reprod 2004;1:31–44. 9. Sharp DC. Environmental influences on reproduction in horses. Vet Clin N Am Large Anim Pract 1980;2:207–23. 10. Sharp DC. Transition into the breeding season: clues to the mechanisms of seasonality. Equine Vet J 1988;20:159– 61. 11. Davis SD, Grubaugh WR, Weithenauer J. Follicle integrity and serum estradiol 17β patterns during sexually recrudescence in the mare. Biol Reprod Suppl 1987;36:121. 12. Donadeu FX, Ginther OJ. Follicular waves and circulating concentrations of gonadotrophins, inhibin and oestradiol during the anovulatory season in mares. Reproduction 2002;124:875–85. 13. Henneke DR, Potter GD, Kreider JL. Body condition during pregnancy and lactation and reproductive efficiency of mares. Theriogenology 1984;21:897–909. 14. Hines KK, Hodge SL, Kreider JL, Potter GD, Harms PG. Relationship between body condition and levels of serum luteinizing hormone in postpartum mares. Theriogenology 1987;28:815–25. 15. Henneke DR, Potter GD, Krieder JL, Yeates BF. Relationship between body condition score, physical measurements and body fat percentage in mares. Equine Vet J 1983;15:371–2. 16. Madill S. Reproductive considerations: mare and stallion. Vet Clin N Am Equine Pract 2002;18:591–619. 17. Vanderwall DK, Woods GL. Age-related subfertility in the mare. Proc Am Assoc Equine Pract 19990;36:85–9. 18. Carnevale EM, Bergfelt DR, Ginther OJ. Aging effects on follicular activity and concentrations of FSH, LH and progesterone in mares. Anim Reprod Sci 1993;31:287–99. 19. Vanderwall DK, Peyrot LM, Weber JS, Woods GL. eproductive efficiency of the aged mare. Proc Soc Theriogenology 1989;153–6. 20. Carnevale EM, Bergfelt DR, Ginther OJ. Follicular activity and concentrations of FSH and LH associated with senescence in mares. Anim Reprod Sci 1994;35:231–46. 21. Evans DR. The recognition and diagnosis of a pituitary tumor in the horse. 18th Proc Am Assoc Equine Pract 1972, pp. 417–19. 22. Van Der Kolk JH, Kalsbeek HC, van Garderen E, Wensing T, Breukink HJ. Equine pituitary neoplasia: a clinical report of 21 cases (1990–1992). Vet Rec 1993;24:1123–7. 23. Dybdal N. Pituitary pars intermedia dysfunction (equine Cushing’s-like diease). In: Robinson NE (ed.) Current Therapy in Equine Medicine, 4th edn. Philadelphia: Saunders, 1997; pp. 499–501. 24. McCue PM. Equine Cushing’s disease. Vet Clin Equine 2002;18:533–43. 25. Love S. Equine Cushing’s disease. Br Vet J 1993; 149:139–53. 26. Beech J. Evaluation of thyroid, adrenal, and pituitary function. Vet Clin N Am Equine Pract 1987;3:649–60. 27. Dybdal NO, Hargreaves KH, Madigan JE, Gribble DH, Kennedy PC, Stabenfeldt GH. Diagnostic testing for pituitary pars intermedia dysfunction in horses. J Am Vet Med Assoc 1994;204:627–32. 28. Messer NT IV, Johnson PJ. Evidence-based literature pertaining to thyroid dysfunction and Cushing’s syndrome in the horse. Vet Clin Equine 2007;23:329–64.
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29. Maher JM, Squires EL, Voss JL, Shideler RK. Effect of anabolic steroids on reproductive function of young mares. J Am Vet Med Assoc 1983;183:519–24. 30. Farquhar VJ, McCue PM, Nett TM, Squires EL. Effect of deslorelin acetate on gonadotropin secretion and ovarian follicle development in cycling mares. J Am Vet Med Assoc 2001;218:749–52. 31. Johnson CA, Thompson DL, Kulinski KM, Guitreau AM. (2000) Prolonged interovulatory interval and hormonal changes in mares following the use of Ovuplant to hasten ovulation. J Equine Vet Sci 2000;20:331–6. 32. McCue PM, Farquhar VJ, Carnevale EM, Squires EL. Removal of deslorelin (Ovuplant) implant 48 h after administration results in normal interovulatory intervals in mares. Theriogenology 2002;58:865–70. 33. Montovan SM, Daels PP, Rivier J, Hughes JP, Stabenfeldt GH, Lasley BL. The effect of a potent GnRH agonist on gonadal and sexual activity in the horse. Theriogenology 1990;33:1305–21. 34. Elhay M, Newbold A, Britton A, Turley P, Dowsett K, Walker J. Suppression of behavioural and physiological oestrus in the mare by vaccination against GnRH. Austr Vet J 2007;85:39–45. 35. Imboden I, Janett F, Burger D, Crowe MA, Hassig M, Thun R. Influence of immunization against GnRH on reproductive cyclicity and estrous behavior in the mare. Theriogenology 2006;66:1866–75. 36. Card C, Raz T, LeHeiget R, Sibert G. GnRF immunization in Mares: ovarian function, return to cycling, and fertility. Proc Am Assoc Equine Pract 2007;53:576–7. 37. Robinson SJ, McKinnon AO. Prolonged ovarian inactivity in broodmares temporally associated with administration of Equity. Aust Equine Vet 2006;25:85–7. 38. Nie GJ, Momont HW, Buoen L. A survey of sex chromosome abnormalities in 204 mares selected for breeding. J Equine Vet Sci 1993;13:456–9. 39. Hughes JP, Benirschke K, Kennedy PC, Trommershausen-Smith A. Gonadal dysgenesis in the mare. J Reprod Fertil Suppl 1975;23:385–90. 40. Lear TL, Bailey E (2008) Equine clinical cytogenetics: the past and future. Cytogenet Genome Res 120:42– 49. 41. Zhang TQ, Buoen LC, Weber AF, Ruth GR. Variety of cytogenetic anomalies diagnosed in 240 infertile equine. Proceedings of the 12th International Congress on Animal Reproduction and Artificial Insemination, 1992; pp. 1939– 41. 42. McCue PM, Roser JF, Munro CJ, Liu IKM, Lasley BL. Granulosa cell tumors of the equine ovary. Vet Clin Equine 2006;22:799–817. 43. McCue PM. Equine granulosa cell tumors. Proc Am Assoc Equine Pract 1992;38:587–93. 44. Meagher DM, Wheat JD, Hughes JP, Stabenfeldt GH, Harris BA. Granulosa cell tumors in mares – a review of 78 cases. Proc Am Assoc Equine Pract 1977;133. 45. Stabenfeldt GH, Hughes JP, Kennedy PC, Meagher DM, Neely DP. Clinical findings, pathological changes, and endocrinological secretory patterns in mares with ovarian tumors. J Reprod Fertil Suppl 1979;27:277. 46. Hodge SL, Kreider JL, Potter GD, Harms PG, Fleeger JL. Influence of photoperiod on the pregnant and postpartum mare. Am J Vet Res 1982;43:1752–5. 47. Koskinen E, Kurki E, Katila T. Onset of luteal activity in foaling and seasonally anoestrous mares treated with artificial light. Acta Vet Scand 1991;32:307–12.
48. Willis LA, Riddle WT. Theriogenology question of the month. J Am Vet Med Assoc 2005;226:877–9. 49. Steiner J, Antczak DF, Wolfsdorf K, Saville K, Brooks S, Miller D, Bailey E, Zent W. (2006) Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. 50. Allen WR, Kolling M, Wilsher S. An interesting case of early pregnancy loss in a mare with persistent endometrial cups. Equine Vet Educ 2007;19:539–44. 51. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. 52. Liptrap RM. Stress and reproduction in domestic animals. Ann N Y Acad Sci 1993;697:275–84. 53. Alexander SL, Irvine CHG. The effect of social stress on adrenal axis activity in horses: the importance of monitoring corticosteroid-binding globulin capacity. J Endocrinol 1998;157:425–32. 54. Baucus KL, Squires EL, Ralston SL, McKinnon AO, Nett TM. J Anim Sci 1990;68:419–26. 55. Nambo Y, Oikawa M, Yoshihara T, Kuwano A, Hobo S, Nagata S, Watanabe G, Taya K. Effects of transport stress on concentrations of LH and FSH in plasma of mares: a preliminary study. J Equine Sci 1996;7:1–5. 56. Barbacini S, Zavaglia G, Gulden P, Marchi V, Necchi D. Retrospective study on the efficacy of hCG in an equine artificial insemination programme using frozen semen. Equine Vet Educ 2000;12:312–17. 57. Farquhar VJ, McCue PM, Vanderwall DK, Squires EL. Efficacy of the GnRH agonist deslorelin acetate for inducing ovulation in mares relative to age of mare and season. J Equine Vet Sci 2000;20:722–5. 58. Asa CS, Ginther OJ. Glucocorticoid suppression of oestrus, follicles, LH and ovulation in the mare. J Reprod Fertil Suppl 1982;32:247–51. 59. Ferris RA, McCue PM. The effects of dexamethasone and prednisolone on pituitary and ovarian function in the mare. Clin Theriogenol 2009;1:225. 60. Squires EL, Heesemann CP, Webel SK, Shideler RK, Voss J. Relationship of altrenogest to ovarian activity, hormone concentrations and fertility of mares. J Anim Sci 1983;4:901–10. 61. Bucca S, Carli A, Buckley T, Dolci G, Fogarty U. The use of dexamethasone administered to mares at breeding time in the modulation of persistent mating induced endometritis. Theriogenology 2008;70:1093–100. 62. McKinnon AO, Perriam WJ, Lescun TB, Walker J, Vasey JR, Trigg TE. Effect of a GnRH analogue (Ovuplant), hCG and dexamethasone on time to ovulation in cycling mares. World Equine Vet Rev 1997;2:16–18. 63. Hughes JP, Loy RG. Investigations on the effect on intrauterine inoculation of Streptococcus zooepidemicus in the mare. Proc Am Assoc Equine Pract 1969;15:289–92. 64. Fredriksson G, Kindahl H, Stabelfeldt GH. Endotoxininduced and prostaglandin-mediated effects on corpus luteum function in the mare. Theriogenology 1986;25:309–16. 65. Daels PF, Starr M, Kindahl H, Fredriksson G, Hughes JP, Stabenfeldt GH. Effect of Salmonella typhimurium endotoxin on PGF-2 alpha release and fetal death in the mare. J Reprod Fertil Suppl 1987;35:485–92. 66. Gunthle LM, McCue PM, Farquhar VJ, Foglia RA. Effect of prostaglandin administration post ovulation on corpus luteum formation in the mare. In: Proceedings of the Society of Theriogenology 2000; p. 139. 67. Troedsson MH, Ababneh MM, Ohlgren AF, Madill S, Vetscher N, Gregas M. Effect of periovulatory
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68.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84. 85.
prostaglandin F2alpha on pregnancy rates and luteal function in the mare. Theriogenology 2001;55:1891–9. Baker CB, Newton DI, Mather EC, Oxender WD. Luteolysis in mares after endometrial biopsy. Am J Vet Res 1981;42:1816–18. Hurtgen JP, Whitmore HL. Effects of endometrial biopsy, uterine culture, and cervical dilatation on the equine estrous cycle. J Am Vet Med Assoc 1978;173:97–100. McCue P, Deluca C, Patten M, Squires EL. Effect of sham transcervical embryo transfer and/or Altrenogest administration on plasma progesterone concentrations in recipient mares. In: Proceedings of International Symposium on Equine Embryo Transfer, 2008; p. 73. Koblischke P, Kindahl H, Budik S, Aurich J, Palm F, Walter I, Kolodziejek J, Nowotny N, Hoppen HO, Aurich C. Embryo transfer induces a subclinical endometritis in recipient mares which can be prevented by treatment with non-steroid anti-inflammatory drugs. Theriogenology 2008;70:1147–58. Stabenfeldt GH, Hughes JP, Evans JW, Neely DP. Spontaneous prolongation of luteal activity in the mare. Equine Vet J 1974;6:158–63. Bergfelt DR, Woods JA, Ginther OJ. Role of the embryonic vesicle and progesterone in embryonic loss in mares. J Reprod Fertil 1992;95:339–47. Hughes JP, Stabenfeldt GH, Kindahl H, Kennedy PC, Edqvist LE, Neely DP, Schalm OW. Pyometra in the mare. J Reprod Fertil Suppl 1979;27:321–9. Neely DP, Kindahl H, Stabenfeldt GH, Edqvist LE, Hughes JP. Prostaglandin release patterns in the mare: physiological, pathophysiological, and therapeutic responses. J Reprod Fertil Suppl 1979;27:181–9. Nie GJ, Johnson KE, Branden TD, Wenzel JGW. Use of an intra-uterine glass ball protocol to extend luteal function in mares. J Equine Vet Sci 2003;23:266–73. Rivera del Alamo MM, Reilas T, Kindahl H, Katila T. Mechanisms behind intrauterine device-induced luteal persistence in mares. Anim Reprod Sci 2008;107: 94–106. Stout TAE, Lamming GE, Allen WR. Oxytocin administration prolongs luteal function in cyclic mares. J Reprod Fertil 1999;116:315–20. Vanderwall DK, Rasmussen DM, Woods G.L. Effect of repeated administration of oxytocin during diestrus on duration of function of corpora lutea in mares. J Am Vet Med Assoc 2007;231:1864–7. Daels PF, McCue PM, DeMoraes MJ, Hughes JP. Persistence of the luteal phase following ovulation during altrenogest treatment in mares. Theriogenology 1996;46:799–811. Hofferer S, Duchamp G, Palmer E. Ovarian response in mares to prolonged treatment with exogenous equine pituitary gonadotrophins. J Reprod Fertil Suppl 1991;44:341–9. McCue PM, Figueiredo E, Lasley BL. Estrogen dynamics during the transition period in the mare. Biol Reprod1992;46 (Suppl 1):157. Asa CS, Goldfoot DA, Garcia MC, Ginther OJ. The effect of estradiol and progesterone on the sexual behavior of ovariectomized mares. Phys Behav 1984;33:681–6. Crowell-Davis SL. Sexual behavior of mares. Horm Behav 2007;52:12–17. Ginther OJ, Whitmore HL, Squires EL. Characteristics of estrus, diestrus, and ovulation in mares and effects of season and nursing. Am J Vet Res 1972;33:1935–9.
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86. Sharp DC, Davis SD. Vernal transition. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 133–43. 87. Asa CS, Goldfoot DA, Garcia MC, Ginther OJ. Dexamethasone suppression of sexual behavior in the ovariectomized mare. Horm Behav 1980;14:55–64. 88. McDonnell SM. Normal and abnormal sexual behavior. Vet Clin N Am Equine Pract 1992;8:71–89. 89. McCue PM. Estrus suppression in performance mares. J Equine Vet Sci 2003;23:342–4. 90. Gee EK, DeLuca C, Stylski JL, McCue PM. Efficacy of medroxyprogesterone acetate in suppression of estrus in cycling mares. J Equine Vet Sci 2009;29:140–5. 91. McKinnon AO, Figueroa STD, Nobelius AM, Hyland JH, Vasey JR. Failure of hydroxyprogesterone caproate to maintain pregnancy in ovariectomized mares. Equine Vet J 1993;25:158–60. 92. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32:83–5. 93. McCue PM, Lemons SS, Squires EL, Vanderwall DK. EfR implants in suppression of estrus ficacy of Synovex-S in the mare. J Equine Vet Sci 1997;17:327–9. 94. Asa CS, Goldfoot DA, Ginther OJ. Assessment of the sexual behavior of pregnant mares. Horm Behav 1983;17:405–13. 95. Hayes KEN, Ginther OJ. Relationship between estrous behavior in pregnant mares and the presence of a female conceptus. J Equine Vet Sci 1989;9:316–18. 96. Asa CS, Goldfoot DA, Garcia MC, Ginther OJ. Sexual behavior in ovariectomized and seasonally anovulatory pony mares (Equus caballus). Phys Behav 1980;14:46–54. 97. Meinecke B, Gips H. Steroid hormone secretary patterns in a mares with granulosa cell tumors. J Vet Med Assoc 1987;34:545–60. 98. Gastal MO, Gastal EL, Beg MA, Ginther OJ. Stallion-like behavior in mares: review of incidence, characteristics, ovarian activity, and role of testosterone. J Equine Vet Sci 2007;27:390–3. 99. Beaver BV, Amoss Jr MS. Aggressive behaviour associated with naturally elevated serum testosterone in mares. Appl Anim Ethol 1982;8:425–8. 100. Jubb KVF, Kennedy PC, Palmer N. Pathology of Domestic Animals, 3rd edn. New York: Academic Press, 1985; pp. 305–23. 101. McDonnell SM. Sexual behavior dysfunction in mares. In: Robinson NE (ed) Equine Medicine vol.3. Philadelphia: Saunders, 1992; pp. 633–7. 102. Silberzahn P, Zwain I, Martin B. Concentration increase of unbound testosterone in plasma of the mare throughout pregnancy. Endocrinology 1984;115:416–19. 103. Bowling AT, Hughes JP. Cytogentic abnormalities. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 258–65. 104. Lear TL, Bailey E. Equine clinical cytogenetics: the past and future. Cytogenet Genome Res 2008;120; 42–9. 105. Cammaert S, Coryn M, De Kruif A, Spincemaille J, Delbeke FT, Debackere M. Three full sister mares with stallion-like behaviour and a high blood testosterone concentration: A case report. J Equine Vet Sci 1993;13:220–2. 106. Kieffer NM. Male pseudohermaphroditism of the testicular feminizing type in a horse. Equine Vet J 1976;8:38– 41.
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107. McKinnon AO. Reproductive ultrasonography. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore, MD: Williams & Wilkins, 1998; pp. 79–102. 108. Squires EL, Voss JL, Maher JM, Shideler RK. Fertility of young mares after long-term anabolic steroid treatment. J Am Vet Med Assoc 1985;186:583–6.
109. Skelton KV, Dowsett KF, McMeniman NP. Ovarian activity in fillies treated with anabolic steroids prior to the onset of puberty. J Reprod Fertil Suppl 1991;44: 351–6. 110. Schumacher EMA, Blackshaw JK, Skelton KV. The behavorial outcomes of anabolic steroid administration to female horses. Equine Pract 1987;9:11–15.
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Photoperiod Daniel C. Sharp
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Chapter 183
Photoperiod Daniel C. Sharp
Introduction The world is filled with cyclic events, the result of the daily rotation of the Earth and its annual revolution around the sun. Faced with such cyclic phenomena, most living things developed temporal organization or time-keeping systems with which to keep pace and fit into their environments as harmoniously as possible. The daily rotation of the Earth creates clocklike timing of events, such as the rising and setting of the sun, and lunar cycles. The annual change in seasons reflects the Earth’s tilt on its axis and changing angle of solar radiation as the Earth orbits our sun. The influence of these changes on our environment is, of course, dramatic, and could be disruptive of attempts at homeostasis if we were not able to anticipate and coordinate our reactions to them. Fortunately, evolutionary pressures to adjust to the environment apparently were in effect, because most species that evolved in a temperate zone where such changes were prominent developed a circadian timekeeping system that enabled them to predict and adjust to the environment. Thus, the circadian timekeeping system utilizes the daily changes in photoperiod to adjust homeostasis of multiple organ systems. Further understanding of the circadian time-keeping system will aid in the continuing effort to regulate the circadian clock of our animals and ourselves.
The circadian time-keeping system The term circadian comes from Latin for around a day (circa, around; dies, a day) and describes a very specific rhythmic regulatory system that provides an ongoing, internal time-keeping mechanism, yet one that is capable of reacting and adjusting to changing environmental conditions. Thus, to be truly circadian, a biological rhythm must adhere to the following conditions: (i) it must be an endogenously generated rhythm with a documentable period or tau of about,
but not precisely, 24 hours and (ii) it must be entrainable to a specific photoperiod, such as the solar day. The expression of the endogenous rhythm in the absence of entrainment is called free-run. This name is derived from early experiments in which locomotor activity of hamsters was recorded in running wheels. Hence, when the hamsters expressed their endogenous rhythms in the lack of entrainment, it was ‘free run’. Just because a rhythm occurs, even if it occurs every day, does not automatically mean that it is a circadian rhythm, and care must be used in describing biological rhythms as circadian unless it is known that the rhythm satisfies the above criteria. Other rhythms that deserve definition are as follows. 1. Diurnal. Diurnal means simply occurring during the day and implies no circadian basis at all, although a diurnal rhythm may indeed be circadian. 2. In contrast, nocturnal means a rhythm that occurs during the night. 3. Quotidian. Quotidian means occurring every day. 4. For the purist, the term nyctohemeral refers to a rhythm occurring both day and night. It would be correct, for instance, to refer to the melatonin rhythm as nyctohemeral, and, as it turns out, probably circadian as well. Another important aspect of circadian rhythms is that, once entrained by some environmental factor (daylength, food intake, and temperature are among common entraining factors), not only is the frequency of the environmental factor imposed on the internal clock but also the inherent rhythm is phased appropriately to local time.1 Anatomical pathway for photoperiodism While birds and lower vertebrates can apparently monitor changes in photoperiod directly through the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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pineal gland, in mammals the eyes constitute the primary photoreceptor. Complete loss of rod and cone function in humans and in animal models does not eliminate photoresponsiveness, whereas experimental bilateral enucleation abolishes all ocular light responses. In fact, several aspects of photoperiodism, such as inhibition of nocturnal melatonin, pupillary constrictor responses, and light-induced phase shifts of the circadian clock, have all been shown to be mediated by a non-rod, non-cone system. Evidence for such intrinsically photosensitive photoreceptors was reported by Hattar et al.,2 who demonstrated the existence of an opsin-like protein called melanopsin located in retinal ganglion cells which project monosynaptically to the ventrolateral portion of the suprachiasmatic nuclei (SCN). This direct connection to the SCN is called the retinohypothalamic tract (RHT) and has been shown to be present in all mammals studied, including horses (see Chapter 174). Monosynaptic projections from the retinal ganglion cells travel through this anatomical pathway to neurons in the ventrolateral SCN where the photic input is processed.3 Glutamate and pituitary adenyl cyclase activating peptide (PACAP) are the principal neurotransmitters of the retinohypothalamic tract, with PACAP apparently acting as a potent modulator of the glutamatergic signaling within the SCN.4, 5 Clock genes in the SCN The past decade has been one of exciting advances in our understanding of photoperiodism and functioning of the circadian clock.6 Critical breakthroughs in understanding the molecular and biochemical basis of circadian time-keeping have revolutionized the concepts of SCN input and output mechanisms. It is now quite clear that circadian time-keeping is the result of a self-sustaining master oscillator (the SCN) that controls numerous peripheral circadian oscillators (slave oscillators) in ways that remain to be completely understood. The biochemical mechanism within the SCN that creates and maintains clock function of the SCN is the autoregulatory interaction of positive and negative transcriptional/translational feedback loops that result in recurring rhythms of RNA and protein levels of key clock components.4 Specifically, two basic helix–loop–helix (bHLH) transcription factors, CLOCK and BMAL1, dimerize and activate transcription through binding to E box enhancers. This is the so-called positive loop of the clock mechanism. Transcriptional activation by CLOCK–BMAL1 leads to activation of three period genes (mPer1, mPer2, and Mper3) in mice, and two cryptochrome genes (mCry1 and mCry2). The mPer and mCry proteins translocate back into the nucleus and inhibit further transcription through their negative
interaction with CLOCK and BMAL1. RNA levels of BMAL1 peak approximately 12 hours out of phase with the RNA of mPer and mCry. These genes interact in an autoregulatory manner with a periodicity of approximately 24 hours, thus creating a self-sustaining oscillation reminiscent of the quartz tuning fork in an electric watch. These clock genes are induced in the ventrolateral SCN. It has been proposed that a light-unresponsive oscillator exists in the dorsomedial SCN, and that the two oscillators, ventrolateral SCN (light responsive) and dorsomedial SCN (light unresponsive), may be synchronized through afferent fibers originating in the ventrolateral SCN.7 Generation of action potentials by SCN neurons provides a way of transmitting the timing signal through neurotransmitters and secreted factors. The SCN-derived inhibitory or stimulatory factors have been shown to drive activity and sleep/wake rhythms in rodents.4 One known secreted factor, prokineticin-2 (PK2), a peptide whose RNA levels are regulated by CLOCK and BMAL1, appears to contribute to the circadian control of behavior. Intracerebroventricular administration of PK2 at night, when its levels are normally low, results in reduction in nighttime behavioral activity, suggesting that PK2 may act during daylight hours to limit activity in rodents.8 The SCN acts as a master oscillator but there appear to be auxillary or ‘slave’ oscillators throughout the body. How the master oscillator regulates multiple slave oscillators outside the brain is less well understood. It seems likely that neural outputs from the SCN to peripheral oscillators by way of the autonomic nervous system could account for some coordination. Hormonal signals from the brain are capable of entraining peripheral oscillators as well, as evidenced by a shift in peripheral oscillators in mice by glucocorticoid agonists.9 Finally, the SCN may regulate peripheral rhythms by virtue of its direct regulation of activity rhythms. By regulating activity rhythms, for example, timing of biochemical events in the liver is also regulated because the timing of feeding behavior is altered. Regulation of the clock by light As discussed, the SCN acts as a self-sustaining oscillator, providing a continuous and precise ‘tuning fork’ against which biological organisms can adjust their homeostatic needs. How then does light impinge on this system to apprise the organism of the need to adjust? Glutamatergic terminals from the melanopsin-containing cells trigger a calciumdependent cascade, involving mitogen-activated protein (MAP) kinases, resulting in activation of the cyclic adenosine monophosphate (cAMP) response element (CRE)-binding protein (CREB). However,
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Photoperiod response to glutamate is circadian time dependent (clock gated), in that the response to light delivered during the circadian day is ineffective. On the other hand, when light is delivered during the circadian night, when SCN activity is relatively quiet, the above described phosphorylation, calcium/cAMP cascade is highly effective, and genes for Per1 and Per2 are rapidly induced. Furthermore, gene expression of Per1 is rapidly induced after light exposure at either the beginning or the end of the dark phase (scotophase). On the other hand, light exposure at the beginning of scotophase results in a robust expression of the gene for Per2, but light exposure later in the scotophase does not lead to any detectable change in expression of the gene for Per2. The significance of these findings is that Per2 appears to regulate earlynight responses (phase delay of the circadian clock), whereas Per1 appears to be more a mediator of circadian clock advances (light exposure late in the scotophase).4 In rodents and in humans, light exposure at the beginning and at the end of the scotophase has been shown to result in circadian clock phase delays and advances, respectively.10 Health implications of circadian time-keeping As previously stated, the purpose of the circadian time-keeping system is to assist in homeostasis by predicting future environmental quality and adjusting to such changes by phase advance or phase delay.11 The importance of this system to maintaining good health is just beginning to be appreciated. Cardiovascular and cerebrovascular crises are most common; for instance, in the early morning hours when the clock upregulates cardiovascular activity. Rotating shift workers have a higher risk of developing cancers, and, in mice, experimental ablation of the SCN or altering light schedules frequently also facilitate tumor development. Over- or underexpression of Per1 can reduce or increase the rate of apoptosis in human cancer cells.12 The foregoing has used mostly experimental animal models or human clinical evidence to point out the function of clock genes and their importance in health and disease. The importance of clock gene regulation to equine health and disease, and more specifically to reproductive function, has been little studied. Recently, equine genes encoding clock genes ARNTL (BMAL1), Per2, CLOCK, and Cry1 have been sequenced and identified,13 providing tools with which to begin understanding the role of circadian clock genes in equine reproduction. A recent study of the effects of systemic inflammation on peripheral blood clock genes suggested that Per2 and BMAL1 were upregulated following administration
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of lipopolysaccharide (LPS) to induce a febrile condition experimentally. Furthermore, co-administration of LPS and phenylbutazone, a non-steroidal antiinflammatory drug known to inhibit prostaglandin synthesis, prevented both the febrile response and the synchronized increase in the two clock genes.13 These data provide a glimpse of the potential role of clock genes in equine health and disease.
Nature of photoperiodic signals in horses One of the major questions of photoperiodic regulation of reproduction is whether the duration of photic exposure or the temporal coincidence of photic exposure with specific clock events is the critical regulator of circadian events. As applied to animal studies of seasonal reproduction, this question is generally framed around the response of circulating melatonin to different experimental or commercial photoperiods in attempts to explain how a changing melatonin signal might convey the appropriate information as to whether a given animal should reduce or increase reproductive activity. The coincidence hy¨ pothesis, originally proposed by Bunning in 1936,14 stated that light had two roles: (i) entrainment of the circadian system and (ii) activation of a particular light-sensitive phase (photosensitive phase) of the circadian clock by coincidence of that phase with ambient illumination. Thus, according to this theory, an animal exposed to changing daylength would not experience a change in circadian timing until the daylength was sufficiently long so as to coincide with the ‘photosensitive phase’ of the circadian ¨ uz ¨ and Stetson15, 16 recently provided evclock. Gund idence in support of the coincidence theory in hamsters. The foregoing discussion of the effects of light exposure on clock genes such as Per1 and Per2 put into perspective how such a mechanism might be regulated. On the other hand, Lees put forth a contrasting theory in 195014 known as the hourglass or duration hypothesis. This hypothesis essentially states that, as daylength (or nightlength) changes with season, a critical nightlength (CNL) is reached that evokes physiological changes. Although these two apparently contrasting theories have not yet been resolved, it is worth contemplating their potential involvement in setting up photoperiod systems for horses.
Quality and quantity of light required to affect the circadian system of horses One of the most frequently asked questions by breeders is ‘How much light is enough?’ It is a good question; one that has not yet been resolved. Burkhardt17
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Table 183.1 Aperture readings and light intensity based on an ASA setting of 400 and a shutter speed of 1/60. A plain white Styrofoam cup placed over the lens acts as a diffuser Aperture reading F8 F5.6 F4 F3.5 F2
Table 183.2 Aperture readings and light intensity based on an ASA setting of 400 and a shutter speed of 1/30. The camera meter readings result from reflection from a neutral gray card
Intensity (FT-CDLS)
Intensity (Lux)
Luminance
45 12 10 5 3
485 129 108 54 32
cd/ft2
Footlamberts
Footcandles
Lux
0.08 0.17 0.52 0.67 1.34 2.67 5.35 10.70 21.40 43 86 171 342
0.26 0.52 1.05 2.10 4.20 8.40 17 34 67 135 270 540 1075
1.5 3 6 12 23 47 93 190 375 750 1500 3000 6000
16 32 63 125 250 500 1000 2000 4000 8000 16000 32150 64300
The light intensity and aperture reading representing an approximate threshold for reproductive stimulus is shown in bold font.
used a simple incandescent light and was able to induce early sexual recrudescence in anestrous mares. As a result of his work and that of others, it has become widely accepted that a plain 100-W incandescent bulb in a 12 × 12-feet 3.6m × 3.6m box stall is sufficient to elicit a reproductive response. This information, however useful, does not fill the void of unanswered questions about what kind of light and what exposure patterns are best. For instance, what role does bright moonlight play, if any. Is there a deleterious effect of the neighbor’s security light up on a pole 100 meters or more distant? What is the minimum intensity of light required to generate a reproductive response? The average intensity of Burkhardt’s 100-W bulb was approximately 10 footcandles (ft-cdls) or 107 lux. Many breeders who are preparing a light program for their mares face the problem of testing to see whether their installation does indeed generate about 100 lux. Many lightbulb manufacturers are happy to supply average or expected luminance of a given bulb in a given fixture. Luminance is a measure of the amount of light emitted from a given source, and impinges on a particular area. The terms footcandle and lux have largely been supplanted by the currently accepted International System of Units (SI), which uses candela per square meter (cd/m2 ) ( Table 183.1). Perhaps the best way to determine whether your light installation is sufficient is to obtain a photometer and measure the intensity directly. To use such a system, set the DIN (ASA) reading of the camera to 400 and the shutter speed to 1/4 second. Because the metering system of most cameras is sensitive to a focal point of light, it is important to diffuse the light entering the camera so as to estimate the general luminance. This can be done by holding a plain white Styrofoam cup over the lens. With this homemade diffuser in place, hold the camera as if to take a picture, at approximately the mare’s eye level, and walk around the area you wish to test. The meter reading should be f4 or higher in order to have a light program of sufficient intensity to stimulate reproduc-
1.0 1.4 2.0 2.8 4.0 5.6 8.0 11.0 16.0 22.0 32.0 45.0 64.0
Illuminance
tion. A website from Kodak describes a similar way to estimate luminance (http://www.kodak.com/ cluster/global/en/consumer/products/techInfo/ am105/am105kic.shtml). The Kodak system is different in that it uses reflected light off a neutral gray card, and a shutter speed of 1/30 second. Despite these differences, the system is similar (Table 183.2). The relatively lower values obtained with the Styrofoam cup and using a faster shutter speed (1/60 second) have always been associated with successful lighting systems, so there appears to be a margin of safety in using the Styrofoam cup system. When to begin lighting An oversight that often occurs when instituting a lighting program is to begin the lighting regime too late in the year to obtain a reproductive stimulation. Many breeders are not aware, or forget, that the springtime transition from anestrus into the breeding season requires several weeks or even months for completion. During this transition phase, despite many indications to the contrary, most mares are not fertile (see Chapter 178). The effect of a stimulatory photoperiod is not to shorten the transition phase, but to begin it earlier. Therefore, in order to accomplish a reproductive stimulus, and have mares begin to cycle considerably earlier in the year than they would under ambient light, it is necessary to begin the lighting program in the late fall. It is recommended that a program of artificial lighting be instituted no later than December 1. In that regard, a good practice is to couple the beginning of the
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Photoperiod
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DAY LENGTH BY LATITUDE 16 15
25 degrees N 45 degrees N
14 DAY LENGTH hr
c183
13 12 11 10 9 8 7 6 Jan
Feb
Mar
Apr
May
Jun
Figure 183.1 Changes in daylength by latitude. The rate of change in daylength at these two latitudes is different, yet the date of first ovulation of the year is remarkably similar.
artificial light program with an easily remembered date, such as November 25 (Thanksgiving). Remember that the response to changes in photoperiod may be a complex interaction of direct experience, i.e., the length of day today, and the history of experience, i.e., the photoperiod of the past 6 months or even 12 months. The importance of this is that it is not likely wise to maintain a long day photoperiod in the hopes that it will keep mares reproductively stimulated throughout the year. It may be necessary for them to experience a period of daylength decline before experiencing a period of increasing daylength. Further work is needed in this area, but the successes currently enjoyed with beginning stimulatory photoperiod around the time of Thanksgiving is hard to argue. Lighting systems The decision to use artificial lighting in a breeding system is likely to require some construction and/or changes to the physical plant of a farm. If the existing management system has mares stalled overnight, then simply adding an incandescent or fluorescent light fixture in each stall may be all that is required. More likely, stalls already have lighting fixtures. If that is the case, then it may only be a matter of having a relatively inexpensive electronic timer installed on the lighting circuit so that all stalls can be controlled. If, on the other hand, mares are not brought into a stall at night, adequate lighting systems can be developed for paddocks. It is a good idea to consult a lighting specialist before constructing such a paddock, as a specialist should be able to provide estimates for the
number and size of fixtures required to provide adequate illumination. The size of the paddock and the number of mares using it are judgments best left to the breeding manager. For obvious reasons, it is best not to crowd too many animals into a relatively small area. One management system that works well will be described below. Briefly, this involves bringing mares into the paddock just before sunset, with the lights coming on at the time of sunset, exposing the mares to 2.5–3 hours of artificial light, then turning them out to pasture. This accomplishes the goal of providing a reliable photostimulation, and avoids the cost and labor associated with stalling them overnight. Required length of photostimulation Classically, the length of day thought to be photostimulatory in a variety of seasonal animals is 14.5 hours or longer. However, this axiom may not be entirely true. Consider, for example, that mares exposed to natural conditions in the temperate zone of North America (approximately 25–47◦ N) enter the breeding season at approximately the same time of year, yet are exposed to large differences in daylength (Figure 183.1). The longest day of the year in Florida, for example, is just about 14 hours of daylight, yet the timing of the annual reproductive rhythm in mares in Florida (30◦ N latitude) is almost precisely the same as in Wisconsin (47◦ N), where the longest day of the year is about 15.4 hours. Therefore, the photic signal that regulates onset and offset of the breeding season in mares may not be as simple as total length of daylight. Likewise, the rate of change, i.e., the rate at
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which day lengthens from, say, December 21 to June 22, is much faster at the more extreme latitudes; yet, in mares within these extremes, timing of the equine breeding season is essentially the same (Figure 183.1). If the photic signal is not absolute length of day, or the rate at which daylength changes, what is the nature of the signal? That question remains to be answered, but the following possibilities are open for consideration.
NIGHT DAY
NIGHT DAY
SHORT DAY (December 22)
LONG DAY (June 22) (a)
Light in the evening only Sharp and Seamans18 reported that an early onset of the breeding season could be stimulated by exposing anestrous mares to only 2.5 hours of additional artificial light beginning at the time of sunset. The artificial lighting regime was begun at sunset every evening and the time of ‘lights on’ was updated every week or so, to match the natural time of sunset as it changed. This regime was started in late October. Importantly, the experiment examined the effects of additional artificial light applied at sunset, as mentioned, prior to sunrise, and at both time points (actually 5 hours additional lighting). The results of that experiment indicated that only those mares exposed to light in the evening were photostimulated and experienced an earlier onset of the breeding season (Figure 183.2). Furthermore, when mares were exposed to light at both morning and evening, their breeding season was not significantly earlier than in mares experiencing additional light in the evening only. This suggests that the timing of light experienced in the evening is different from that experienced in the morning. That is to say, the data indicate that the effects of light exposure are clock-gated as discussed above. This is an exciting concept, as some of the equine clock genes, Per2 in particular, have been cloned19 and are available as tools to undertake the studies needed to clarify this mechanism.
Night interruption, or ‘pulse lighting’ Palmer et al.20 reported that early onset of the breeding season could also be stimulated by exposing anestrous mares to a 1-hour pulse of artificial light if the pulse were timed appropriately. They reported that the pulse of light exposure was effective only when it occurred 9.5–10.5 hours after the onset of darkness (Figure 183.3). This lighting regime seems to have practical application, primarily in management systems in which mares are housed in stalls overnight. Application of this system to outdoor paddocks, however, is of lesser utility because it eliminates one advantage of paddock lighting, the short time of confinement. The night interruption program also raises questions of how it works. If the equine
NIGHT DAY
NIGHT DAY PM
*
NIGHT DAY
NIGHT DAY AM
AM
PM
* (b) Figure 183.2 (a) Representation of daylength for the shortest day of the year (December 22) and the longest day of the year (June 22), showing that the main difference between these times of year occurs at sunrise and sunset. (b) Experimental design of an experiment to determine whether light at sunrise or sunset is more effective. The dotted lines represent the timing of artificial light added. Asterisks denote groups in which date of first ovulation was significantly advanced.
clock genes function as understood in other mammals, light exposure late in the scotophase might not affect Per2, but might affect expression of Per1. By extrapolating from rodent studies, enhancement of Per1 gene expression with light exposure late in the
Figure 183.3 The top bar represents a typical winter time short day. The second bar represents the successful ‘night interruption’ program in which mares are exposed to a 1-hour pulse of light at 9.5–10.5 hours after onset of darkness. The third and fourth bars represent photoperiods that did not result in a reproductive stimulus. Adapted from Palmer and Driancourt, 1981.20
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Photoperiod scotophase could lead to a circadian clock phase advance. That is, experiencing a light pulse late in the dark phase could lead to a shift in timing of the circadian clock so that a photosensitive phase, if one exists in horses, might be shifted to a time when light is present. Do not forget that, with a night interruption program, animals are really exposed to dawn and dusk twice in a 24-hour period, and it becomes difficult to state with confidence where the circadian clock is set unless experiments to establish a phase response curve are performed. Therefore, light exposure in the early evening and late in the dark phase both appear to be successful methods for advancing the onset of the breeding season, but it is likely that they do so by somewhat different mechanisms. At the surface, it appears that evening light exposure may impinge on a photosensitive phase directly, and a night interruption light exposure may shift the circadian clock so that a photosensitive phase is moved to a new circadian time. Caution is urged at these interpretations, as no one has yet performed the proper experiments to explain the underlying mechanisms. As pointed out above, knowledge of the equine clock genes may now permit us to advance our understanding of how light works to regulate the annual reproductive rhythm of mares, and permit development of rational management schemes.
References 1. Aschoff J. Circadian rhythms: General features and endocrinological aspects. In: Kreiger DT (ed.) Endocrine rhythms. New York: Raven Press, 1979; pp. 1–61. 2. . Hattar S. Melanopsin-containing retinal ganglion cells: architecture, projections and intrinsic photosensitivity. Science 2002; 295: 1065–70. 3. Kalsbeek A, Palm IF, La Fleur SE, Scheer FAJL, PerreauLenz S, Ruiter M, Kreier F, Cailotto C, Buijs RM. SCN outputs and the hypothalamic balance of life. J Biol Rhythms 2006; 21: 458–69. 4. Reppert SM, Weaver DR. Coordination of circadian timing in mammals. Nature 2002; 418: 935–41. 5. Michel S, Itri J, Han JH, Gniotczynski K, Colwell CS. Regulation of glutamatergic signalling by PACAP in the mammalian suprachiasmatic nucleus. BMC Neurosci 2006; 16: 15.
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6. Takahashi JS. Finding new clock components: past and future. J Biol Rhythms 2004; 19: 339–47. 7. Nagano M, Adachi A, Nakahama K, Nakamura T, Tamada M, Meyer-Bernstein E, Sehgal A, Shigeyoshi Y. An abrupt shift in the day/night cycle causes desynchrony in the mammalian circadian center. J Neurosci 2003; 23: 6141–51. 8. Cheng MY, Bullock CM, Li C, Lee AG, Bermak JC, Belluzzi J, Weaver DR, Leslie FM, Zhou QY. Prokineticin 2 transmits the behavioural circadian rhythm of the suprachiasmatic nucleus. Nature 2002; 417: 405–10. 9. Balsalobre A, Brown SA, Marcacci L, Tronche F, Kellendonk ¨ G, Schibler U. Resetting of circaC, Reichardt HM, Schutz dian time in peripheral tissues by glucocorticoid signaling. Science 2000; 289: 2344–7. 10. Khalsa SB, Jewett ME, Cajochen C, Czeisler CA. A phase response curve to single bright light pulses in human subjects. J Physiol 2003; 549: 945–52. 11. Kalsbeek A, Kreier F, Fliers E, Sauerwein HP, Romijn JA, Buijs RM. Minireview: Circadian control of metabolism by the suprachiasmatic nuclei. Endocrinology 2007; 148: 5635–9. 12. Hastings M, O’Neill JS, Maywood ES. Circadian clocks: regulators of endocrine and metabolic rhythms. J Endocrinol 2007; 195: 187–98. 13. Murphy BA, Vick MM, Sessions DR, Cook RF, Fitzgerald BP. Acute systemic inflammation transiently synchronizes clock gene expression in equine peripheral blood. Brain Behav Immun 2007; 21: 467–76. ¨ 14. Saunders DS. Erwin Bunning and Tony Lees, two giants of chronobiology, and the problem of time measurement in insect photoperiodism. J Insect Physiol 2005; 51: 599–608. ¨ uz ¨ B, Stetson MH. A test of the coincidence and dura15. Gund tion models of melatonin action in Siberian hamsters: the effects of 1-hr melatonin infusions on testicular development in intact and pinealectomized prepubertal Phodopus sungorus. J Pineal Res 2001; 30: 97–107. ¨ uz ¨ B, Stetson MH. A test of the coincidence and du16. Gund ration models of melatonin action in Siberian hamsters. II. The effects of 4- and 8-hr melatonin infusions on testicular development of pinealectomized juvenile Siberian hamsters (Phodopus sungorus). J Pineal Res 2001; 30: 56–64. 17. Burkhardt J. Transition from anestrous in the mare and the effects of artificial lighting. J Agric Sci 1947; 37: 64–8. 18. Sharp DC, Seamans KW. Effect of time of day on photostimulation of the breeding season in mares. In: Proceedings of the 72nd Annual Meeting of the American Society of Animal Science, 1980. 19. Murphy BA, Lear TL, Adelson DL, Fitzgerald BP. Chromosomal assignments and sequences for the equine core circadian clock genes. Anim Genet 2007; 38: 84–5. 20. Palmer E, Driancourt MA, Ortavant R. Photoperiodic stimulation of the mare during winter anoestrus. J Reprod Fertil Suppl 1982; 32: 275–82.
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Chapter 184
Progesterone Edward L. Squires
Introduction
Progestin treatment
‘Transition period’ is defined as the time between winter anestrus and normal cyclicity. This is generally characterized by long, erratic estrous periods accompanied by anovulatory follicles. By definition, the first ovulation of the year is the end of the transition period. Failure of mares to ovulate during the transition period is a result of low levels of pituitary luteinizing hormone (LH) and low circulating levels of LH. Concentrations of follicle-stimulating hormone (FSH) in the pituitary are high and peripheral concentrations of FSH are elevated during the transition period.1 It is not until LH increases and FSH decreases that the mare ovulates the first time in the breeding season. Duration of estrus can be as long as 15–30 days in the transition period and mares may develop several anovulatory follicles before an ovulatory follicle is obtained. This makes it extremely difficult to manage mares for breeding. One option is to examine the mare’s ovary by rectal palpation and ultrasonography twice a week during the transition period and, once follicles > 30 mm are obtained, then more frequent examinations can be performed. Eventually the mare will develop a large pre-ovulatory follicle that warrants breeding or insemination. Obviously this is expensive and can be unproductive. The other alternative is to suppress these long, erratic estrous periods during the transition season using some form of progestin or to use gonadotropinreleasing hormone (GnRH) or perhaps LH compounds to hasten the first ovulation of the year. Information about the use of GnRH or LH products to hasten the first ovulation of the year are presented in Chapters 185 and 190. This chapter will focus on the use of exogenous progestins (progesterone-like compounds) in transitional mares.
The intent of using exogenous progestins in transitional mares is to suppress estrus for 12–14 days, then allow mares to return to estrus after the end of the progestin treatment. There is some controversy as to whether or not progestins actually hasten the first ovulation of the year or simply synchronize the first ovulation of the year. Many breeders use progestin treatment in transitional mares to ‘program the mare into the breeding shed’. Mares are put on and taken off progestin treatment so that they can be scheduled into the breeding shed when the stallion is available. Studies examining various durations of progestin treatment have not been done, but it would appear that treatment of transitional mares anywhere from 10 to 20 days is an effective means of suppressing estrus and scheduling mares for breeding after the end of treatment. Altrenogest is an orally active progestin compound that has been available since the early 1980s. Many of the studies on suppression of estrus in transitional mares have been performed with altrenogest since this is the only drug approved in the USA for suppression of estrus. Webel and Squires2 were the first to report on the use of altrenogest for estrus and ovulation control in mares. Since that time, there have been numerous reports on the oral progestin altrenogest for regulation of estrus in mares. These studies have demonstrated that prerequisites must be met before mares are responsive to exogenous progesterone during the transition period. That is, for best response, mares should be in midto late transition.3–5 This is generally determined by the follicular population on the mare’s ovaries. Mares having follicles 20–25 mm in size on both ovaries generally are classified as mid-transitional mares.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Progesterone Studies have demonstrated that altrenogest was ineffective for inducing estrus and ovulation in mares in deep anestrus. Only mares that are in mid- to late transition respond favorably to altrenogest treatment.4–6 Typically, mares are treated for 12–14 days during the transition period. Follicular growth is inhibited during treatment with progestin. A hypothesis is that LH content in the pituitary is increased during the time of progestin treatment, resulting in a rebound of secretion of gonadotropins after the cessation of treatment. Upon cessation of treatment, only one pre-ovulatory follicle develops and ovulation occurs as a result of either endogenous or exogenous LH. McCue et al.7 conducted studies that did not support this hypothesis. They examined the effects of prolonged administration of altrenogest in combination with GnRH on baseline concentrations of LH and the release of LH in response to a GnRH challenge in transitional mares. Twenty-four light horse mares were used between April 18 and May 18 (late transition) in the northern hemisphere. Mares with follicles 20–25 mm in diameter were randomly assigned to one of four groups of six mares each: group 1, 1 ml saline; group 2, 22 mg altrenogest orally daily; group 3, 250 µg native GnRH every 12 h; and group 4, 22 mg altrenogest daily and 250 µg GnRH every 12 h. On the day before the first exogenous hormone treatment and the day after the last treatment, mares were challenged with an injection of 1 mg of native GnRH. Blood samples were then collected at 0, 0.5, 1, 1.5, 2, 3, and 4 h post-GnRH. Levels of serum LH were measured by radioimmunoassay. There was no difference in GnRH-induced LH response before the onset of exogenous hormone therapy. During the 12-day treatment period, concentrations of LH were lower in altrenogest-treated mares than in either controls or GnRH-treated mares. At the end of the 12-day treatment period, LH release was significantly greater in control and GnRH-treated mares than altrenogesttreated mares or altrenogest + GnRH-treated mares. There was no difference in interval from the end of treatment to ovulation among groups. There was no evidence in this study that administration of altrenogest to transitional mares resulted in accumulation of LH in the anterior pituitary or that LH release is enhanced when altrenogest treatment is discontinued. Subjecting mares to an artificial photoperiod allows the transition period to occur earlier in the year. Generally, once mares have been exposed to 16 h of photoperiod for 60 days they are in mid- to late transition. A study was conducted in which the combined effects of artificial light and altrenogest were evaluated.6 Mares were exposed to a 16-h photoperiod for 60 days, then altrenogest for an additional 12 days.
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Within treatment groups, one-half of the mares received human chorionic gonadotropin (hCG) on day 2 of estrus and the other half received saline (control). All progestin-treated mares exhibited estrus within 12 days after the end of treatment. The average interval to estrus was 3.4 days. Administration of hCG on day 2 of estrus shortened duration of estrus in both progestin-treated and control mares. However, more progestin-treated mares ovulated within 12 days after treatment than controls. It was concluded that treatment with altrenogest in combination with an artificial photoperiod was effective in regulation of estrus and ovulation and assisted in establishment of normal estrous cycles early in the year. Fertility of mares bred after altrenogest treatment does not appear to be altered. A large clinical trial was conducted in which 441 mature broodmares at 17 different locations were administered altrenogest for 14 days during the months of January to May.2 This represented the entire transition period. During treatment, estrus was suppressed in 94% of the mares. Mares were divided into two groups according to the date of the initial treatment (i.e., before or after March 15). Those mares that were treated before March 15 did not exhibit estrus or ovulate sooner than untreated controls. However, treatment with altrenogest during the late transition phase resulted in shorter post-treatment estrus and a shorter interval to pregnancy for treated versus control mares. This further suggested that altrenogest assisted in normalizing the estrous cycle during late transition. Mares with active ovaries (i.e., follicles > 20 mm) responded with shorter post-treatment estrous periods than those with inactive ovaries. Squires et al.4, 5 reported that the larger the follicle before treatment, the shorter the post-treatment estrus and the more favorable the post-treatment response. Therefore, mares should be examined by rectal palpation and ultrasonography to determine sizes of follicles on the ovaries before treatment. Only those mares with multiple follicles > 20 mm should be selected for treatment. An alternative to the use of an oral progestin is injectable progesterone for control of estrus in transitional mares. Researchers8, 9 administered 150 mg progesterone plus 10 mg estradiol-17β intramuscularly daily. Treated mares also were administered 10 mg prostaglandin F2α (PGF2α ) on the last day of steroid treatment. The authors concluded that the combined steroid–prostaglandin regime provided satisfactory control of ovulation in mares early in the breeding season. However, Wiepz et al.10 conducted a trial to investigate the effect of altrenogest alone or in combination with estradiol on induction of estrus and ovulation in late transitional mares. Treatments were
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initiated during the months of April and May and were given for 15 days. Both altrenogest treatments suppressed estrous behavior and follicular growth compared with controls. However, suppression of follicular activity was significantly greater for the combined steroid treatment. Unfortunately, the interval to estrus and ovulation was longer for mares given altrenogest plus estradiol than for those given altrenogest alone. These authors concluded that the greater suppression of follicular activity with combined steroids was of no advantage over altrenogest alone for induction of estrus and ovulation in late transitional mares. The combined progesterone and estradiol injectable treatment, commonly called P+, still remains a favorable treatment for programming mares in the Thoroughbred country in North America. Vaginal progesterone devices also have been used to control estrus in transitional and cycling mares. Newcombe and colleagues11, 12 reported on the use of CIDR-B vaginal implants containing 1.9 g progesterone inserted for 12 days during late February to early April in the United Kingdom. At the end of treatment, mares with follicles > 30 mm were given Ovuplant, hCG, or no ovulatory agent. At the time of treatment, mares were grouped into those having follicles < 10 mm, 11–20 mm, 21–30 mm, and > 30 mm. Follicle diameter increased in all groups of mares with the insertion of CIDR-B devices. Follicles > 30 mm were obtained in 4 days or less and ovulation occurred in approximately 5.3 days. This study demonstrated the folliclestimulating effects of low doses of progesterone. Although these authors did report vaginitis in a significant number of mares, this did not appear to have any adverse effect on fertility. Unfortunately, the CIDR devices in this study remained in the mares for a variable length of time, depending on how quickly follicles grew, and there were no untreated controls. Foglia et al.,13 at Colorado State University, evaluated the effects of CIDR devices in 26 transitional mares. Mares were assigned to either a control group or CIDR-B devices for 12 days. Deslorelin was given after removal of the CIDR devices. Diameter of the largest follicle at the onset of treatment was 26 mm in both groups; however, at the end of treatment, the diameter of the largest follicle was 39 mm for treated mares and 29 mm for controls. Ten of 13 (77%) of the treated mares had follicles > 35 mm at the time of removal of the CIDR devices compared with only one of 13 controls. Card et al.14 conducted a study on cycling mares using CIDR-B devices. Mares were given either CIDR-B devices alone or CIDR-B devices combined with single injection of estradiol on either day 0 or
day 5. A control group of PGF2α -treated mares was included. Vaginal discharge was noted in 35–50% of the mares. However, 8 days of the CIDR device resulted in estrous synchronization and ovulation rates similar to PGF2α -treated mares. Administration of estradiol did not influence the outcome of the CIDR devices. Another vaginal device (PRID, Santa Fe Animal Health) containing 1.55 g of progesterone and 10 mg of estradiol-17β was evaluated by Newcombe et al.12 The device remained in the mares from 6 to 20 days with a mean duration of 10.9 days. Ovulation occurred an average of 6.6 days after removal of CIDR devices in 92% of the mares. Vaginitis was a relatively common occurrence in mares given the PRID device. However, vaginitis was resolved quickly and had no effect on pregnancy rates or the incidence of endometritis. These authors concluded that this device was a simple, safe, inexpensive method of regulating estrus in transitional mares. Recently, Storer et al.15 evaluated the effectiveness of three sustained release injectable formulations of altrenogest and also evaluated the effectiveness of medroxyprogesterone acetate in cycling mares. Thirty-one normal, cycling mares were assigned to the following groups: (i) 1.5 mL altrenogest, 225 mg altrenogest in a sustained release solution (LA 150); (ii) LA 150 (3 mL altrenogest, 450 mg); (iii) MP 500 (altrenogest, lactide–glycolide microparticles, 500 mg); and (iv) medroxyprogesterone, 1000 mg. The objective was to determine which of these progesterone formulations would delay estrus and ovulation. The LA 1.5 and 3 mL increased days to estrus and ovulation. The MP 500 treatment had the longest suppression of estrus and days to ovulation. Medroxyprogesterone had no effect on days to estrus or ovulation. Values for mares given medroxyprogesterone were similar to untreated controls. They concluded that the LA 150 1.5 mL or the LA 150 3 mL could be used to suppress estrus for 12–14 days, whereas the MP 500 altrenogest treatment could be used to delay estrus for up to 30–40 days. Burns et al.16 measured progesterone levels of 18 anovulatory mares administered either LA 300 (1500 mg), LA 300 (3000 mg), or LA 150 (1500 mg). This product was designed to elevate progesterone for 10–12 days. All of the treatments were successful in elevating progesterone for 10 days. In a subsequent study by these same authors,16 60 non-cycling mares that had not responded to artificial photoperiod were given an injection of LA 300. Seventeen of 30 (57%) of LA 300-treated mares ovulated in 4 weeks compared with only two of 30 controls. They concluded that this product may be used to hasten ovulation in transitional mares that failed to respond to artificial photoperiod.
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Progesterone
Conclusion In conclusion, there would appear to be several types of progestins and progesterone devices that can be used in transitional mares to suppress estrus, synchronize estrus, and perhaps even hasten the onset of the first ovulation of the year.
References 1. Silvia PJ, Squires EL, Nett TM. Changes in the hypothalamic-hypophyseal axis of mares associated with seasonal reproductive recrudescence. Biol Reprod 1986;35:897–905. 2. Webel SK, Squires EL. Control of the estrous cycle in mares with altrenogest. J Reprod Fertil Suppl 1982;32:193–8. 3. Allen WR, Urwin V, Simpson DJ, Greenwood RE, Crowhurst RC, Ellis DR, Ricketts SW, Hunt MD, Digby NJ. Preliminary studies on the use of an oral progestagen to induce estrus and ovulation in seasonally anestrous Thoroughbred mares. Equine Vet J 1980;12:141–5. 4. Squires EL, Heesemann CP, Webel SK, Shideler RK, Voss JL. Relationship of altrenogest to ovarian activity, hormone concentrations and fertility of mares. J Anim Sci 1983;56:901–10. 5. Squires EL, Shideler RK, Voss JL, Webel SK. Clinical applications of progestins in mares. Comp Cont Educ Pract Vet 1983;5:S16–S22. 6. Squires EL, Stephens WB, McGlothlin DE, Pickett BW. Effect of an oral progestin on the estrous cycle and fertility of mares. J Anim Sci 1979;49:729–35. 7. McCue PM, Nickerson KC, Squires EL, Farquhar VJ, Nett TM. Effect of altrenogest on luteinizing hormone concentration in mares during the transition period. In: Proceedings of the 47th Annual Convention of the American Association of Equine Practitioners, 2001, pp. 249–51.
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8. Loy RG, Pemstein R, O’Canna D, Douglas RH. Control of ovulation in cycling mares with ovarian steroids and prostaglandin. Theriogenology 1981;15:191–200. 9. Taylor TB, Pemstein R, Loy RG. Control of ovulation in mares in the early breeding season with ovarian steroids and prostaglandins. J Reprod Fertil Suppl 1982;32:219–24. 10. Wiepz GJ, Squires EL, Chapman PL. Effect of norgestomet, altrenogest and/or estradiol on follicular and hormonal characteristics of late transitional mares. Theriogenology 1988;30:181–93. 11. Newcombe JR. Field observations on the use of a progesterone-releasing intravaginal device to induce estrus and ovulation in seasonally anestrous mares. J Equine Vet Sci 2002;22:378–82. ¨ 12. Newcombe JR, Handler J, Klug E, Meyers PJ, Jochle W. Treatment of transition-phase mares with progesterone intravaginally and with deslorelin or hCG to assist ovulation. J Equine Vet Sci 2002;22:57–64. ¨ 13. Foglia RA, McCue PM, Squires EL, Jochle W. Stimulation of follicular development in transitional mares using a proR ) . In: Proceedings of the gesterone vaginal insert (CIDR-B Annual Conference of the Society for Theriogenology, 1999, p. 33. 14. Card CE, Bruemmer J, Moffett P, Helvight U, Pledger A, Jamison S, Longson G, Loomis P, Squires E. Comparison of R , estrous synchronization in mares treated with CIDR-B estradiol-17β, estradiol cypionate and prostaglandin F2 α. In: Proceedings of the Annual Conference of the Society for Theriogenology, 2003, p. 39. 15. Storer WA, Thompson Jr DL, Gilley RM, Burns PJ. Evaluation of injectable sustained-release progestin formulations for suppression of estrus and ovulation in mares. J Equine Vet Sci 2009;39:33–6. 16. Burns PJ, Morrow C, Abraham J. Evaluation of Biorelease P4 LA 300 in the mare. In: Proceedings of the 7th Equine Embryo Transfer Symposium, Cambridge, UK, 2008, p. 71.
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Chapter 185
GnRH John Hyland
Seasonal anestrus in the mare The mare is seasonally polyestrus, with reproductive activity cued to increasing daylength. The anestrous period is usually subdivided into deep and shallow (transitional) components, the latter occurring during recrudescence.1 Deep anestrus is characterized by a lack of ovarian activity and occurs during winter and early spring. Transitional anestrus occurs typically after the winter solstice and is the time during which follicular activity commences on the ovaries and irregular displays of estrus may be exhibited by mares. Waves of ovarian follicles develop and regress prior to the first ovulation of the season. The transitional stage lasts from several weeks to several months. The importance of seasonal anestrus in mares is that man has imposed an artificial birthday on most breeds to conform to show and race events. The birthday for Thoroughbreds is August 1 in the southern hemisphere and January 1 in the northern hemisphere. For Standardbreds the birthdays are September 1 and January 1 for the southern and northern hemispheres, respectively. Given that the mare has a 340-day gestation, breeding must start on or around September 1 each year in the southern hemisphere and February 1 in the northern hemisphere to maintain a generation interval of 12 months. Standardbred breeders in Australia routinely commence breeding on October 1. Most breeders prefer to have foals born as close to the official birthday as possible as they believe earlier foals have growth, maturity, and weight advantages over their later born contemporaries. This may be translated into bigger and more profitable yearlings at national sales and provide performance advantages in racing as 2-year-olds and as show horses. However, because the imposed breeding season does not completely overlap the physiological breeding season, which is also related to latitude, only 20–25% of mares are cycling when breeding commences.2
This imposes financial and management constraints and is a significant cost to the horse breeding industry.
Physiological control of seasonality in the mare It is now well established that photoperiod drives the seasonal reproductive cycle of the mare.3, 4 It is less clear how this signal is translated into the first ovulation of the season. Sharp and his co-workers in Florida undertook a series of ground-breaking experiments in ponies to show that the hypothalamic content of gonadotropin-releasing hormone (GnRH) was higher in summer than it was in winter5 and that, when mares were treated with melatonin implants, the summer content of GnRH in the hypothalamus was lower than in untreated control mares. These experiments implicate melatonin and GnRH in the recrudescence from anestrus. It is now accepted in a number of species with seasonal traits that photic information is relayed to the pineal gland through a multisynaptic pathway that involves the suprachiasmatic nucleus (SCN).6 This information is converted into a diurnal endocrine signal in the form of melatonin production. The photic information is first conveyed to the SCN, located in the neurons of the anteroventral hypothalamus. There is evidence that the central diurnal pacemaker resides in these nuclei. The pathway connecting the SCN to the pineal gland involves the paraventricular nucleus in the hypothalamus, cell columns in the thoracic spinal cord, and the superior cervical ganglion. The information is then transmitted to the pineal gland.7 Melatonin is produced during darkness and this appears to be under the control of norepinephrine, whose nocturnal secretion is 100fold that of daytime in the rat.8 Melatonin appears to provide an endocrine signal of daylength. This
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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GnRH has been demonstrated by replacement experiments in pinealectomized sheep, in which provision of long-day or short-day melatonin results in inhibition or stimulation, respectively, of luteinizing hormone (LH) secretion.9, 10 It is now accepted that the role of melatonin is the alteration of GnRH secretion at the level of the central nervous system, in the animals studied to date (sheep;11 hamster12 ). Whether the same physiology applies to the mare is not known, but the results of Strauss et al.5 and Hart et al.,13 in which hypothalamic concentrations of GnRH in ponies and horses were higher in summer than in winter, supports this hypothesis. By contrast, Silvia et al.14 recorded lower hypothalamic GnRH concentrations 12 weeks after the winter solstice than those measured at the winter solstice. These authors concluded that GnRH secretion was maximal at this time, resulting in low hypothalamic concentrations. Release of GnRH results in stimulation of the gonadotropins, LH, and folliclestimulating hormone (FSH). The effect on FSH is not as clear as on LH, as FSH concentrations increase only slowly during recrudescence.14 However, it is likely that ovarian secretions modify LH and FSH levels during anestrus and cyclicity. Ovariectomy results in an increase of both hormones during the breeding season compared with intact control mares, with a similar pattern to normal cycling mares.15 The studies described above lead to the conclusion that daylength is the driver of events in certain species with seasonal reproduction. This seems to be the case in short-day breeders (sheep) or longday breeders (horses). Photic events are translated by melatonin into neuroendocrine events that are poorly understood. The rhythm of melatonin secretion informs the animal about the time of year, owing to the length of the melatonin signal. The melatonin signal is translated into endocrine responses either directly or indirectly through neural circuits. Why the melatonin signal has different effects in short-day breeders and long-day breeders is not known. Presumably these effects are genetically predetermined, but other factors such as nutrition, temperature, and social interactions combine with photoperiod to finetune seasonality.
Gonadotropin-releasing hormone GnRH is the master hormone of reproduction. Its existence was inferred by early physicians who observed hypogonadism and obesity in patients with hypothalamic injury.16 The hormone was the second of the hypothalamic releasing hormones to be isolated and identified.17 The hormone is a decapeptide with the linear structure pyro-Glu1 -His2 -Trp3 -Ser4 -
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Tyr5 -Gly6 -Leu7 -Arg8 -Pro9 -Gly10 -NH2 . Substitution of amino acids at various positions along its length results in hormones of changed affinity for the GnRH receptor.18 GnRH is released from specialized neurons in the mediobasal hypothalamus. The cell bodies are aligned along a pathway in the mammalian forebrain in the shape of an inverted Y, with the greatest concentration rostrally around the organum vasculosum of the lamina terminalis above the optic chiasm.19 The cell bodies exhibit predominantly a bipolar morphology with relatively few synaptic contacts compared with neighboring neurons. There are, however, some GnRH neurons that form cytoplasmic bridges with other GnRH neurons and may act to coordinate activity.20 There is extensive interconnection between dendrites of the GnRH neurons, with estimations that each GnRH neuron may be influenced directly or indirectly by 5 000 000 other neurons.19 GnRH neurons release GnRH in a pulsatile manner in culture.21 Not all GnRH neurons exhibit pulsatility and the way in which GnRH neurons communicate to produce synchronous activity is not understood. There are several ways this could occur.19
r r r
Leader neuron model. A small number of neurons generate a synchronous pulse, which is conveyed to the other neurons. Integrated model. In this model, synchronicity arises from extensive interconnectivity of the GnRH neurons. Slave model. This model assumes that there is an external pacemaker which imposes pulsatility on the GnRH neurons, which have only limited interactions with each other.
It is now well established that the pre-ovulatory LH surge is generated by GnRH release into the hypophysial portal vessels in sheep.22, 23 The release is modified by steroid inputs from the ovary and typically involves a change from slow episodic release to a faster release frequency with increased pulse amplitude. The mare appears to be similar to the ewe in that GnRH pulse frequency and amplitude increase around the LH surge.24, 25 The response of FSH to GnRH is less well established than that of LH. It has been shown that GnRH pulses every 2 hours release more FSH mRNA in rat pituitary cell cultures than pulse frequencies of 30 minutes, 1 hour, or 4 hours.26 The differential release of LH and FSH message has been hypothesized due to a change in frequency of GnRH release. At low frequencies, FSH is produced preferentially, while at high frequencies the FSHβ gene is suppressed, with
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no effect on the α gonadotropin subunit gene or the βLH gene.27
Hormonal events during seasonal anestrus in the mare It is accepted that transition into the breeding season is driven by GnRH in the mare in response to increasing photoperiod.28 Strauss et al.5 showed that the concentration of GnRH in the hypothalamus of pony mares was higher in summer than in winter, although this observation was not repeated in a later study.29 Measurement of GnRH secretion directly, using a push–pull technique via a catheter implanted in the sella turcica revealed a non-significant increase in GnRH concentration in extracted fluid between deep anestrus and transitional anestrus, with a greater increase in estrous mares.25 Irvine and Alexander30 also measured GnRH directly by sampling pituitary venous blood in mares in various reproductive states and concluded that GnRH secretion was lowest during seasonal anestrus. During transition concentrations of LH and FSH increase gradually, although LH remains low until just before the first ovulation of the season. This is probably due to low pituitary stores of LH, which increase in response to changes in daylength.31 This situation has been summarized by Sharp,32 wherein increasing daylength results in an increase in GnRH secretion. Increased GnRH causes FSH release and the growth of waves of follicles which are steroidogenically incompetent. Positive feedback on LH secretion does not occur until a follicular wave releases sufficient estrogen to cause an LH surge, resulting in ovulation and the onset of regular cyclicity. Recent studies have confirmed the stimulatory effect of FSH on follicular development in anestrous mares.33–35 Further, Donadeu and Ginther36 divided the latter part of the anovulatory period into mid-anovulatory and transitional periods based on differences in follicular activity. There is a decrease in mean circulating concentrations of FSH between the onset and the termination of the anovulatory season.29, 37, 38 This is due to a decrease in pulse amplitude and may reflect the reported reduction in FSH release from the pituitary in response to GnRH during the transitional period.29 Donadeu and Ginther36 measured the diameters of the second to sixth largest follicle on the ovaries of mares from the end of January to the first ovulation of the season, in the northern hemisphere. Using a statistical model they found that the anovulatory period could be divided into an early period of minimal follicle growth, where the largest follicle did not exceed 21 mm in diameter (mid-anovulatory period).
This was followed by a surge in follicle growth, in a series of waves (transitional period), followed by the pre-ovulatory follicle wave before the first ovulation of the season. To reduce variability, the largest follicle in a wave and any follicle > 28 mm were not included in the data. The transition period began 31–93 days before ovulation and contained two or three waves. There was no association between surges of FSH and the emergence of follicular waves. Mean daily FSH concentrations were similar during midanovulatory and transitional periods, and decreased just before the beginning of the pre-ovulatory period. The number of FSH surges was highest during the mid-anovulatory period, decreasing thereafter into the transition and pre-ovulatory periods. Only 31% of all FSH surges were associated with a follicular wave during the mid-anovulatory and transitional periods, which is in agreement with Watson et al.39 However, there was direct association between a FSH surge and the emergence of a follicular wave during the midanovulatory and transitional periods. The mean follicle diameter at wave emergence increased from midanovulatory to late transition, although FSH profiles were similar. The increase in follicle diameters cannot therefore be attributed to changes in FSH. It is possible other ovarian or growth factors are involved,40 and this is a fertile area for study. Donadeu and Ginther35 used follicle ablation to eliminate the overlapping of sequential follicular waves during the mid-anovulatory, early transition, late transition, and ovulatory periods. This technique allowed more precise monitoring of wave events and revealed that follicular activity increased from midanovulatory to the transition, although the waveassociated surges of FSH were similar. There was a positive seasonal effect on FSH secretion, which was counteracted by an increased production of inhibin during all periods. A seasonal increase in FSH secretion has been previously identified in ovariectomized mares around the time of transition from the anovulatory to ovulatory season.37 The FSH increase appears to be mediated by an increase in GnRH.13, 25 More work needs to be done to clarify the role of FSH in the development of follicular waves during recrudescence from deep anestrus in the mare. Estradiol is implicated in follicular growth and ovulation. It is known that the follicles which develop and regress within the follicular waves generated during the mid-anovulatory period and transitional period may reach similar sizes to their ovulatory counterparts, but produce considerably lower levels of steroid hormones.41 The theory of recrudescence from the anovulatory season is that the first ovulation of the season does not occur until follicular estrogen production is sufficient to induce an LH surge.42 However, Donadeu and Ginther35 recorded a rise in
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GnRH LH in the breeding season in mares in which all follicles > 5 mm had been ablated from the ovaries. This occurred despite tonically low estradiol concentrations, suggesting that an estradiol surge is not necessary to induce a LH surge in these mares. More work needs to be done on the hormonal control of the first ovulation of the season in mares. LH is required for ovulation and luteal maintenance in the mare.43 Plasma concentrations of LH remain low after the last ovulation of the breeding season, but begin a gradual rise after commencement of the transition.36 The reduction in LH during the anovulatory season appears to be due to cessation of production by the pituitary gland.13, 29 The size of the largest follicle produced during transitional anovulatory follicular waves is related to the plasma concentration of LH.44, 45 Therefore, LH appears to be necessary for the final growth spurt of the dominant follicle. Consistent with this theory, Watson et al.39 measured lower levels of LH receptor mRNA in transitional follicles than in pre-ovulatory follicles. There is therefore some evidence to support the proposition that, despite minimal secretion, LH is an important component of the complex events preceding the first ovulation of the season.
Using GnRH to induce cyclicity in seasonally anestrus mares Given that GnRH is the master hormone of reproduction, a number of authors have used this information in devising treatment strategies to induce early cyclicity in acyclic mares. Evans and Irvine46 appear to have been the first authors to administer GnRH (twice daily at 10-day intervals for three or four injections) during transitional anestrus and induce ovulation in 5/5 mares. A subsequent experiment47 in deeply anestrous mares using similar methods was not so successful, with 3/4 mares ovulating without producing a corpus luteum. Johnson48, 49 infused GnRH hourly to mares kept in a non-stimulatory photoperiod (8L:16D) and induced ovulation in 4/4 mares receiving 50 or 250 µg/h,48 and in 13/13 mares given 2 or 20 µg/h.49 In the latter experiment the ovulation rate was 1.2 and 3.0, respectively. Subsequent experiments performed by various scientists confirmed that GnRH or its agonists, given by multiple injection or infusion for several days, could be used to induce cyclicity in anestrous mares.50–54 The above studies terminated treatment after certain end points had been attained. Other workers utilized osmotic minipumps to administer GnRH at doses of 2.5–225 µg/h for periods of 14–28 days to mares at various times during the anovulatory season.50, 55 Although results varied, all these studies
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recorded an increase in ovulatory cycles in response to treatment. The success in inducing ovulation was related to GnRH dose rate55 and follicular development at the commencement of treatment. In the work by Hyland et al.50 there was a rapid decline in progesterone concentrations in deeply anestrous mares after GnRH delivery by osmotic minipumps ceased. This suggested that support of the induced corpus luteum was lost when GnRH release was too low to maintain luteotrophic LH concentrations. This observation was extended when Bergfelt and Ginther53 treated mares with a GnRH analog (200 µg twice daily). Mares with inactive ovaries (follicles < 15 mm diameter) ovulated earlier than control mares, but showed a rapid decline in circulating concentrations of progesterone and lower LH concentrations than controls. Although initial pregnancy rates in the two groups were similar (64% and 70% for treated and control mares, respectively) embryo loss rate to day 40 after ovulation was much higher in the treated mares (64% vs 7% for treated and control mares, respectively). The above studies emphasize that, although mares can be induced to ovulate during seasonal anestrus, the effects of treatment cannot override the inherent seasonal suppression of ovarian activity. The suppression of corpus luteum activity occurred before antiluteolytic effects of the embryo were apparent. Analogues of GnRH have been used in attempts to induce fertile estrus in anestrous mares. Several studies utilized commercial implants of goserelin,56 deslorelin,57 buserelin,58 or buserelin injections (40 µg twice daily).59 The implants advanced the day of ovulation, although most results were not sufficiently impressive to be commercial. Several of the studies did not include contemporary controls, so the results are difficult to interpret. It is also apparent from these reports that the best response occurred in mares treated later in the year. This was demonstrated by McKinnon et al.,60 showing that deslorelin implants (Ovuplant Peptech, Sydney, Australia) given to anestrous mares every 2 days were more effective in inducing ovulation late in transition than earlier in the year (mean time to ovulation, 3.7 vs 24.6 days, for late and early transition, respectively). There appears to be a difference in response of anestrous mares to GnRH treatment depending on whether native hormone or analogs are used. Accordingly, several papers from the same laboratory, describing the use of native GnRH delivered by pulsatile infusion (2–100 µg/h) to anestrous mares or mares in an artificial inhibitory photoperiod, record 54/55 (97%) of mares responding to treatment by ovulating within the designated experimental period.48, 49, 52, 55 In contrast, different GnRH analogs, delivered as implants55, 56 to anestrous mares resulted in 53/162 (28%) ovulating within the experimental period. Therefore, the
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depth of anestrus of individual mares and possibly the preparation of GnRH used will determine the response to treatment.
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Future developments Because of the financial impediments associated with seasonal anestrus, a number of commercial companies have shown interest in developing an implant or controlled release injection to stimulate cyclicity in anestrous mares. At the time of writing, the dopamine antagonist domperidone seems to have taken the place of GnRH in the management of seasonal anestrus.61 Domperidone needs to be administered daily and is not effective in deeply anestrous mares.62 Therefore, any product based on a single administration, containing GnRH or one of its analogs, will be a large step forward in the management of reproduction in the mare.
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References 1. Ginther OJ. Reproduction in the Mare, 2nd edn. Cross Plains, WI: Equiservices, 1990. 2. Osborne VE. An analysis of the pattern of ovulation as it occurs in the annual reproductive cycle of the mare in Australia. Aust Vet J 1966;42:149–54. 3. Burkhardt J. Transition from anoestrus in the mare and the effects of artificial lighting. J Agric Sci 1947;37:64–8. 4. Loy R. Effects of artificial lighting regimes on reproductive patterns in mares. In: Proceedings of the Annual Convention of the 14th American Association of Equine Practitioners, 1968, p. 159. 5. Strauss SS, Chen CL, Kalra SP, Sharp DC. Localisation of gonadotrophin releasing hormone (GnRH) in the hypothalamus of ovariectomised pony mares by season. J Reprod Fertil Suppl 1979;27:123–9. 6. Moore RY. Entrainment pathways and the functional organization of the circadian system. Prog Brain Res 1996;111:103–19. 7. Reiter RJ. The mammalian pineal gland: structure and function. Am J Anat 1981;162:287–313. 8. Drijfhout WJ, van der Linde AG, Kooi SE, Grol CJ, Westerink BH. Norepinephrine release in the rat pineal gland: the input from the biological clock measured by in vivo microdialysis. J Neurochem 1996;66:748–55. 9. Bittman EL, Dempsey RJ, Karsch FJ. Pineal melatonin secretion drives the reproductive response to daylength in the ewe. Endocrinology 1983;113: 2276–83. 10. Bittman EL, Karsch FJ. Nightly duration of pineal melatonin secretion determines the reproductive response to inhibitory daylength in the ewe. Biol Reprod 1984;30:585–93. 11. Malpaux B, Daveau A, Maurice-Mandon F, Duarte G, Chemineau P. Evidence that melatonin acts in the premammillary hypothalamic area to control reproduction in the ewe: presence of binding sites and stimulation of luteinising hormone secretion by in situ microimplant delivery. Endocrinology 1998;139:1508–16. 12. Maywood ES, Hastings MH. Lesions of the iodo-melatonin binding sites of the mediobasal hypothalamus spare the lactotropic but block the gonadotropic response of male
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
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Syrian hamsters to short photoperiod and melatonin. Endocrinology 1995;136:144–53. Hart PJ, Squires EL, Imel KJ, Nett TM. Seasonal variation in hypothalamic content of gonadotrophin releasing hormone (GnRH), pituitary receptors for GnRH, and pituitary content of luteinising hormone and follicle stimulating hormone in the mare. Biol Reprod 1984;30:1055–62. Silvia PJ, Johnson L, Fitzgerald BP. Changes in the hypothalamic-hypophysial axis of mares in relation to the winter solstice. J Reprod Fertil 1992;96:195–202. Freedman LJ, Garcia MC, Ginther OJ. Influence of photoperiod and ovaries on seasonal reproductive activity in mares. Biol Reprod 1979;20:567–74 Everett JW. Pituitary and hypothalamus: perspectives and overview. In: Neill JD (ed.) Knobil and Neill’s Physiology of Reproduction, 3rd edn. Sydney: Elsevier, 2006; pp. 1289– 307. Schally AV, Arimura A, Kastin AJ. Hypothalamic regulatory hormones. Science 1973;179:341–9. Conn PM, McArdle CA, Andrews WV, Huckle WR. The molecular basis of gonadotrophin releasing hormone (GnRH) action in the pituitary gonadotrope. Biol Reprod 1987;36:17–35. Herbison AE 2006. Physiology of the gonadotrophin releasing hormone neuronal network. In: Neill JD (ed.) Knobil and Neill’s Physiology of Reproduction, 3rd edn. Sydney: Elsevier, 2006; pp. 1415–82. Witkin JW, O’Sullivan H, Silverman AJ. Novel associations among gonadotrophin releasing hormone neurons. Endocrinology 1995;136:4323–30. Duittoz AH, Batailler M. Pulsatile GnRH secretion from primary cultures of sheep olfactory placode explants. J Reprod Fertil 2000;120:391–6. Clarke IJ, Cummins JT. The temporal relationship between gonadotrophin releasing hormone (GnRH) and luteinising hormone (LH) secretion in ovariectomised ewes. Endocrinology 1982;111:1737–49. Moenter SM, Brand RM, Midgely AR, Karsch FJ. Dynamics of gonadotrophin releasing hormone release during a pulse. Endocrinology 1992;130:503–10. Vivrette SL, Irvine CHG. Interaction of oestradiol and gonadotrophin-releasing hormone on LH release in the mare. J Reprod Fertil Suppl 1979;27; 151–5. Sharp DC, Grubaugh WR. Use of push-pull perfusion techniques in studies of gonadotrophin-releasing hormone secretion in mares. J Reprod Fertil Suppl 1987;35:289–96. Kaiser UB, Jakubowiak A, Steinberger A, Chin WW. Differential effects of gonadotropin-releasing hormone (GnRH) pulse frequency on gonadotropin subunit and GnRH receptor messenger ribonucleic acid levels in vitro. Endocrinology 1997;136:1224–31. Jeong K-H, Kaiser UB. Gonadotrophin-releasing hormone regulation of gonadotrophin biosynthesis and secretion. In: Neill JD (ed.) Knobil and Neill’s Physiology of Reproduction, 3rd edn. Sydney: Elsevier, 2006; pp. 1635–701. Irvine CHG. The nonpregnant mare: a review of some current research and of the last 25 years of endocrinology. Biol Reprod Monogr 1995;1:343–60. Silvia PJ, Squires EL, Nett TM. Pituitary responsiveness of mares challenged with GnRH at various stages of the transition into the breeding season. J Anim Sci 1987;64:790–6. Irvine CHG, Alexander SL. A novel technique for measuring hypothalamic and pituitary hormone secretion rates from collection of pituitary venous effluent in the normal horse. J Endocrinol 1987;113:183–92.
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GnRH 31. Alexander S, Irvine CHG. Control of onset of breeding season in the mare and its artificial regulation by progesterone treatment. J Reprod Fertil Suppl 1991;44:307–18. 32. Sharp DC. Transition into the breeding season: clues to the mechanism of seasonality. Equine Vet J 1988;20:159–61. 33. Fitzgerald BP, Affleck KJ, Barrows SP, Murdoch WL, Barker KB, Loy RG. Changes in LH pulse frequency and amplitude in intact mares during the transition into the breeding season. J Reprod Fertil 1987;79:485–93. 34. Alexander SL, Irvine CHG. Control of the onset of the breeding season of the mare and its artificial regulation by progesterone treatment. Reproduction Suppl 1991;44:307–18. 35. Donadeu FX, Ginther OJ. Interactions of follicular factors and season in the regulation of circulating concentrations of gonadotrophins in mares. Reproduction 2003;125:743–50. 36. Donadeu FX, Ginther OJ. Follicular waves and circulating concentrations of gonadotrophins, inhibin and oestradiol during the anovulatory season in mares. Reproduction 2002;124:875–85. 37. Freedman IJ, Garcia MC, Ginther OJ. Influence of ovaries and photoperiod on seasonal reproductive activity in mares. Biol Reprod 1979;20:567–74. 38. Hines KK, Affleck KJ, Barrows SP, Murdoch WL, Fitzgerald BP, Loy RG. Follicle Stimulating Hormone pulse amplitude decreases with the onset of the breeding season in the mare. Biol Reprod 1991;44:516–21. 39. Watson ED, Heald M, Tsigos A, Leask R, Steele M, Groome NP, Riley SC. Plasma FSH, inhibin A and inhibin isoforms containing pro- and αC during winter anoestrus, spring transition and the breeding season in mares. Reproduction 2002;123:535–42. 40. Donadeu FX, Watson ED. Seasonal changes in ovarian activity: Lessons learnt from the horse. Anim Reprod Sci 2007;100:225–42. 41. Watson ED, Thomassen R, Steele M, Groome NP, Riley SC. Concentrations of inhibin, progesterone and oestradiol in fluid from dominant and subordinate follicles from mares during spring transition and the breeding season. Anim Reprod Sci 2002;74:55–67. 42. Sharp DC. Environmental influences on reproduction in horses. Vet Clin North Am 1980;2:207–23. 43. Evans MJ, Irvine CHG. Serum concentrations of FSH, LH and progesterone during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1975;23:193–200. 44. Bergfelt DR, Gastal EL, Ginther OJ. Response of estradiol and inhibin to experimentally reduced luteinising hormone during follicle deviation in mares. Biol Reprod 2001;65:426–32. 45. Ginther OJ, Beg MA, Donadeu FX, Bergfelt DR. Mechanism of follicular deviation in monovular farm species. Anim Reprod Sci 2003;78:239–57. 46. Evans MJ, Irvine CHG. Induction of follicular development, maturation and ovulation by gonadotrophin releasing hormone administration to acyclic mares. Biol Reprod 1977;16:452–62. 47. Evans MJ, Irvine CHG. Induction of follicular development and ovulation in seasonally acyclic mares using gonadotrophin releasing hormone and progesterone. J Reprod Fertil Suppl 1979;27:113–21. 48. Johnson AL. Induction of ovulation in anoestrous mares with pulsatile administration of gonadotrophin releasing hormone. Am J Vet Res 1986;47:983–6.
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49. Johnson AL. Gonadotrophin releasing hormone treatment induces follicular growth and ovulation in seasonally anoestrous mares. Biol Reprod 1987;36:1199–206. 50. Hyland JH, Wright PJ, Clarke, IJ, Carson RS, Langsford DA, Jeffcott, LB. Infusion of gonadotrophin-releasing hormone (GnRH) induces ovulation and fertile oestrus in mares during seasonal anoestrus. Proceedings of the 4th International Symposium on Equine Reproduction. J Reprod Fertil Suppl 1987;35:211–20. 51. Ainsworth CGV, Hyland JH. Continuous infusion of gonadotrophin-releasing hormone (GnRH) advances the onset of oestrous cycles in Thoroughbred mares on Australian studfarms. Proceedings of the 5th International Symposium on Equine Reproduction. J Reprod Fertil Suppl 1990;44:235–40. 52. Williams GL, Amstalden M, Blodgett GP, Ward JE, Unnerstall DA, Quirk KS. Continuous administration of low-dose GnRH in mares. I. Control of persisitent anovulation during the ovulatory season. Theriogenology 2007;68:67–75. 53. Bergfelt DR, Ginther OJ. Embryo loss following GnRHinduced ovulation in anovulatory mares. Theriogenology 1992;38:33–43. 54. Palmer E, Quellier P. Uses of LHRH and its analogues: In: Proceedings of the 11th International Congress on Animal Reproduction and Artificial Insemination, vol. 5, 1988, pp. 339–46. 55. Fitzgerald BP, Affleck KJ, Pemstein R, Loy RG. Investigation of the potential of LHRH or an agonist to induce ovulation in seasonally anoestrous mares with observations on the use of the agonist in problem acyclic mares. J Reprod Fertil Suppl 1987;35:683–4. 56. Allen WR, Sanderson MW, Greenwood RES, Ellis DR, Crowhurst JS, Simpson DJ, Rossdale PD. Induction of ovulation in anoestrous mares with a slow release implant of a GnRH analogue (ICI 118630). J Reprod Fertil Suppl 1987;35:469–78. 57. Mumford EL, Squires EL, Peterson KD, Nett TM, Jasko DJ. Effect of various doses of a gonadotrophin releasing hormone analogue on induction of ovulation in anoestrous mares. J Anim Sci 1994;72:178–83. 58. Fitzgerald BP, Meyer SL, Affleck KJ, Silvia PJ. Effect of constant administration of a gonadotropin-releasing hormone agonist on reproductive activity in mares: induction of ovulation during seasonal anestrus. Am J Vet Res 1993;54:1735–45. 59. Camillo F, Pacini M, Panzani D, Vannozzi I, Rota A, Aria G. Clinical use of twice daily injections of buserelin acetate to induce ovulation in the mare. Vet Res Commun 2004;28:169–72. 60. McKinnon AO, Vasey JR, Lescun TB, Trigg TE. Repeated use of a GnRH analogue deslorelin (Ovuplant) for hastening ovulation in the transitional mare. Equine Vet J 1997;29:153–5. 61. Brendemuehl JP, Cross DL. Influence of the dopamine antagonist domperidone on the vernal transition in seasonally anoestrous mares. J Reprod Fertil Suppl 2000;56: 185–93. 62. McCue PM, Buchanan BR, Farquar VJ, Squires EL, Cross DL. Efficacy of domperidone on induction of ovulation in anoestrus and transitional mares. In: Proceedings of the 45th Annual Convention of the American Association of Equine Practitioners, 1999, pp. 217–18.
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Chapter 186
Dopamine Antagonists Ahmed Tibary
One of the most common challenges in equine reproduction practice, particularly in the thoroughbred industry, is the pressure to produce foals in the northern hemisphere as close to January 1 as possible due to the implications age may have on athletic performance.1 It is well established that the most important regulatory mechanism of seasonality of reproduction in horses is photoperiod.2 The equine hypothalamo–pituitary–gonadal axis is subject to a circannual endogenous rhythm that is primarily regulated by daylength. Other factors such as age, body condition, lactation, temperature, and nutrition affect the response to changing photoperiod such that the onset of the first ovulation of the year varies from one mare to another and even within a mare from one year to another.3 Artificial photoperiod has been used for decades to manipulate the breeding season in mares and advance the transition period.4 Response to photoperiod stimulation depends on time of initiation of the stimulus as well as its duration and intensity. When photoperiod stimulation is initiated too early (late fall/autumn) the response is often less dramatic than when the treatment is initiated in early winter. The length of daylight and intensity has been extensively studied but most studies today agree on the use of a minimum of 14.5 hours of light and 107 lux (equivalent to 100 Watt bulb or 10 footcandles). However, it is possible that as little as 50 watt bulb per stall may produce similar results. Research has shown that the key factor in response to light is the duration of the darkness. Mares placed in darkness during 9.5 hours and subsequently exposed to light for 1 hour and then returned to natural darkness until dawn respond in the same fashion as mares exposed to 14.5 hours of continuous light (see Chapter 183). Adequate stimulation can be obtained in practice by exposing mares to 2 hours of light starting 9 hours after onset of darkness. Although the minimum du-
ration of the photoperiod treatment that produces a shift in the transition period is 35 days, our recommendation is to use at least 60 to 70 days’ treatment. There is a considerable variation in the interval from the start of treatment to first ovulation in individual mares. It is important to realize that photoperiod treatment merely displaces events that precede the first ovulation to an earlier date in the year. Therefore, mares still have to experience a period of transition from anestrus to cyclicity that is often variable. This period of transition is characterized by resumption of sexual function, onset of follicular development, and behavioral estrus in response to increased secretion of pituitary gonadotropins and ovarian hormones. However there is a lack of ovulation despite development of several large follicles.5 Mares with large ovaries and follicles ranging 25–35 mm in late winter and early spring are considered to in the transitional phase.6 The first ovulation of the season is often delayed by cold weather and in mares with low body condition score (< 5 on a scale of 1 to 9). This effect of temperature and body condition is experienced in mares even in areas where the seasonal variation of photoperiod is not very large. Mares that are on lush pasture seem to respond better to photoperiod treatment than mares kept inside.3 Other clues related to body fat and leptin levels in particular may play a role in seasonality and depth of anestrus.7–9 In some situations the length of photoperiod treatment and the variability of results render treatment cumbersome or impractical. Several pharmacological protocols alone or in conjunction with photoperiodic manipulation have been utilized to shorten the period of treatment and reduce the variability of the response. These include the use of equine folliclestimulating hormone (eFSH), pituitary extracts, GnRH, and progestogens.6 In recent years more focus has been on the use of dopamine antagonists.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Dopamine Antagonists HO
NH2
HO Figure 186.1 Structure of dopamine.
Role of dopamine in reproductive seasonality Studies in several species displaying seasonal reproductive patterns have shown that this phenomenon is in part regulated by dopamine D2 receptors as evidenced by the presence of synapses between dopaminergic and GnRH neurons in the median eminence. Inhibition of D2 dopamine receptors was found to be effective in increasing luteinizing hormone (LH) secretion during anestrus, possibly through a reduction in the negative feedback of estradiol on GnRH.10, 11 In mares, dopamine concentration in cerebrospinal fluid is higher during the anovulatory period compared to the breeding season12 and it is inversely related to prolactin plasma levels.13 The exact mechanism of action of catecholamines, dopamine (Figure 186.1) in particular, on follicular dynamics is not known but is believed to involve regulation of prolactin as pituitary production of prolactin is primarily regulated through inhibition by the neurotransmitter dopamine.14 In mares, circulating levels of prolactin are depressed in autumn and winter and increase with increasing follicular activity in spring and summer.13 In addition, plasma levels of prolactin were found to be highly correlated with follicular diameter during the spring transition period and administration of ovine or recombinant porcine prolactin to seasonally anovultory mares has been shown to advance the date of the first ovulation.15, 16 In one study, rapid follicular growth was observed within 3 days of administration of ovine prolactin in three out of four treated anestrous mares.15 Mares receiving recombinant porcine prolactin (4 mg s.c. s.i.d.) starting on January 15 in the northern hemisphere had their first ovulation on average 4 weeks (February 6 ± 3 days vs March 14 ± 6 days) earlier than placebo-treated mares.17 It is important to note that FSH levels are not affected in treated mares, suggesting that the effect of prolactin is not due to stimulation of pituitary gonadotropin secretion. Recently dopamine receptors have been detected on both theca and granulosa cells and dopamine seasonal variation was found to be very well correlated with high prolactin levels in fluid of follicles >20 mm in diameter.18, 19 In addition, dopamine D1 and D2 receptor proteins have been detected in equine corpus luteum tissues,18 and primordial and/or primary and secondary preantral follicles
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are suspected to have a great ability to produce D2 receptors as indicated by the elevated mRNA levels for D2 receptor in the equine ovarian cortex.20 Based on these observations and the lack of FSH increase in prolactin-stimulated mares, it has been suggested that dopamine could act through D2 receptor protein to inhibit ovarian follicular growth. This would explain why the dopamine antagonists domperidone and sulpiride, which have D2 receptor activity, would have a stimulatory effect on follicular development in anestrous mares. This hypothesis is validated by the observation that sulpiride and domperidone treatment does not stimulate episodic FSH secretion in anestrous mares, suggesting that their stimulatory effect on follicular development does not operate through a stimulation of the pituitary but may act directly on the ovary through regulation of the FSH receptor population.15, 17, 19, 21, 22
Use of dopamine antagonists Several dopamine antagonists are available and differ primarily by the type of receptor they target. In the equine, the most commonly used dopamine antagonists are domperidone21 and sulpiride.22–25 Fluphenazine (a long-acting D2 receptor antagonist)15 and perphenazine26, 27 have been used to a lesser extent. Research has demonstrated that all these compounds are able to stimulate follicular growth and/or ovulation in seasonally anovulatory mares to some extent. However, the effects of dopamine antagonists on the onset of reproductive activity varies greatly21 between experiments,28, 29 and a great individual variability in response exists.6 Photoperiod, temperature, nutrition, and body condition could possibly influence the impact of dopamine antagonists on the time of first ovulation. Differences in results among studies may be due to differences in vehicle, dosage, and/or frequency of administration.22 Domperidone Domperidone (C22 H24 ClN5 O2 , mol.wt 425.911) (Figure 186.2) has strong affinity for D2 and D3 dopamine receptors. It is generally administered in oral or intravenous form and is primarily metabolized in the liver and intestine (first pass) with a O O
HN N
NH N
N
Figure 186.2 Structure of domperidone.
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half-life of 7 hours. Domperidone has been used extensively to counteract the effects of fescue toxicosis and to treat agalactia and is also used to promote follicular growth in transitional mares. Treatment (1.1 mg/kg, per os s.i.d.) is started at the end of the second week of photoperiod stimulation. Mares are maintained under lights and domperidone administered until occurrence of the first ovulation. Administration of domperidone (1.1 mg/kg) to seasonally anestrous mares maintained under natural photoperiod from January 15 stimulated follicular development within 14 days of treatment and hastened mean date of ovulation by 78 days.21, 30 Twothirds of the mares that ovulated continued to cycle normally whereas the others reverted back to anestrus. Treatment with domperidone resulted in increased prolactin concentration but had no effect on FSH and LH secretion. In contrast, other studies found no positive effect on follicular development or ovulation in anestrous mares housed outdoors under natural photoperiod and treated with domperidone (1.1 or 2.2 mg/kg bodyweight s.i.d.) for 60 days.29 The same authors failed to show any effect of domperidone in mares housed indoors under long photoperiod.31 Sulpiride Sulpiride (C15 H23 N3 O4 S, mol.wt 341.427) (Figure 186.3) is a selective dopamine antagonist at postsynaptic D2 receptors and has a mild sedative effect at high doses. It is absorbed slowly from the gastrointestinal tract and has very limited bioavailabilty (25–35%) through this route with a half-life of 7 hours. In mares, sulpiride is available only through compounding pharmacies. Doses used vary from 200 mg/mare s.i.d.24 to 1.0 mg/kg bodyweight s.i.d.23 Although using a limited number of mares, one study reported that administration of sulpiride to anestrous mares hastened ovulation by 33 days.24 In addition, mares showed a typical winter hair coat shedding following an increase in prolactin. Ovulating mares showed normal luteal phase with normal length and progesterone levels and normal pregnancy rate. Daels et al., using 0.5 mg/kg bodyweight b.i.d., showed a decrease in the interval to first ovulation.32, 33 In another study where mares were matched
O
O H 2N
by reproductive history, body condition, and bodyweight, administration of sulpiride (1 mg/kg bodyweight, b.i.d.) for a maximum of 21 days starting 14 days after initiation of a 14.5 hours light : 9.5 hours dark photoperiod, resulted in an ovulation 16.7 days earlier than in control mares.25 In contrast, other studies failed to show any effect on date of first ovulation despite an increase in prolactin in anestrous mares treated with sulpiride (1 mg/kg bodyweight, s.i.d.) from January 14 through February 14 in absence of a photoperiod treatment.22, 34 One possible explanation of the discrepancies between studies is the differences in depth of anestrus of the mares used in trials, resulting in various levels of FSH and/or prolactin.33, 35 When sulpiride and domperidone were compared under the same experimental conditions, only sulpiride administration was able to significantly hasten the transitional period and the first ovulation in deep anestrous mares.36 A recent study28 showed that treatment of seasonally anovulatory mares with sulpiride increased prolactin concentrations, but not to the degree reported for orally administered domperidone.21 The prolactin response to the first sulpiride injection was much greater than it was after subsequent injections, even 2 days later.28 A similar finding was reported earlier by Thompson and Depew16 who found a rapid drop in response after the first day of treatment. These authors concluded that sulpiride stimulated prolactin secretion, but did not sustain a long-term production. Other dopamine antagonists Fluphenazine (C22 H26 F3 N3 OS, mol.wt 437.523) (Figure 186.4) is a dopamine D2 receptor blocker with an oral availability of 40 to 50% and is metabolized in the liver with a half-life of 15 to 30 hours. In mares, administration of fluphenazine decanoate (178.6 µg/kg, i.m. s.i.d. for 3 weeks) during anestrus accelerated follicular growth but did not result in an advancement of the first ovulation.15 Perphenazine (C21 H26 ClN3 OS, mol.wt 403.97) (Figure 186.5) has an oral bioavailibity of 40% with a half-life of 8 to 20 hours. Administration of perpherazine (0.375 mg/kg, per os b.i.d.) to anestrous
O
N
S N H
N
N
N
CF3
O
S Figure 186.3 Structure of sulpiride.
Figure 186.4 Structure of fluphenazine.
OH
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Dopamine Antagonists HO N
Cl
N N S
Figure 186.5 Structure of perphenazine.
mares hastened follicular growth and subsequent ovulation.27
Effect of estradiol pretreatment One of the proposed mechanisms for failure of dopamine antagonists to stimulate ovarian follicular activity is the lack of estrogen in seasonally anestrous mares. Estradiol was shown to significantly increase prolactin secretion and pituitary content in ovariectomized pony mares during the breeding season.37 Also, LH secretion was shown to be positively affected by estradiol 17ß (1 mg in oil daily for 16 days) in ovariectomized mares in winter.38 Recently, it was shown that estradiol pretreatment of seasonally anovulatory mares (11 mg i.m. every other day for 10 injections) enhances prolactin and ovarian responses to daily injection of sulpiride starting on day 11 of the treatment and continued until ovulation or 45 days.28 Prolactin levels in mares pretreated with estradiol benzoate then receiving sulpiride was twice that found in mares receiving sulpiride without estradiol pretreatment.28 The potentiating effect of estradiol on prolactin response to sulpiride treatment seems to last more than 24 hours, resulting in several-fold increase of prolactin compared to untreated mares.28 The effect of estradiol continued for several days. Pretreatment with estradiol was found to significantly increase LH concentration to levels comparable to those observed in the normal preovulatory surge.28 This may explain the observed effect of sulpiride on hastening ovulation. However, previous studies with recombinant porcine prolactin treatment alone showed that ovulation and/or luteal formation occurs about 22 days after onset of treatment with no or little increase in LH.17 This observation has led to development of the hypothesis that hastening of ovulation in dopamine antagonist-treated mares is not necessarily a result of increased LH but is due in great part to the increased sensitivity of the ovary (follicle) to LH through increased receptor numbers as a result of prolactin increase.39, 40 In one experiment, estradiol-treated mares ovulated on the 40th day of the year on average (February 9).28 If this response is shown to be repeatable, it
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would be an ideal approach for breeding programs intending to start breeding in mid-February. A similar pattern of response has been found with a single injection of domperidone microparticles following estradiol benzoate treatment. However, further studies are needed in anestrous mares.28 In summary, advancement of the first ovulation can be achieved by pretreatment of mares with estradiol benzoate (11 mg i.m. every other day) for a total of 10 injections followed on day 11 with a series of sulpiride injections (250 mg/mare s.c.) until ovulation. Sulpiride treatment could be substituted with a single injection of domperidone (3 grams in microparticles).
Conclusion All dopamine antagonists (sulpiride,23, 36 domperidone,21, 36 perphenazine,15 and fluphenazine27 ) seem to stimulate follicular growth; however, only sulpiride seems to be capable of inducing ovulation (larger size follicle). A major difference between sulpiride and domperidone is that the former crosses the blood–brain barrier.41, 42 The inability of domperidone to cross the blood–brain barrier may explain its lower activity. The effect of sulpiride on the hypothalamus and/or the pituitary has been suspected based on an increase in LH secretion pattern in sulpiride-treated mares but needs to be studied further.24, 36 The increase in LH could also be mediated by an increase in prolactin. Another hypothesis is that sulpiride may exert an effect on GnRH secretion during anestrus via a decrease in dopaminergic inhibition of GnRH neurons, resulting in an increased frequency of FSH pulses.24 The state of our knowledge at this point is that dopamine antagonists can be used in conjunction with light to reduce the length of photoperiod treatment and to shorten the spring transition period. In this system, exposure of mares to long photoperiod (14.5 hours) could be started in mid-January rather that the traditional late November starting date. In the case of sulpiride (0.5 to 1 mg/kg i.m. b.i.d.) treatment should commence in combination with exposure to 14.5 hours of light and continued until occurrence of ovulation or for a maximum of 21 days. The induced ovulation following sulpiride treatment results in normal cycle length.36 Further, the pregnancy and foaling rates of sulpiride-treated mares was reportedly normal with normal return to estrus for non-pregnant mares.23, 33, 36 Similar results should be achievable with the use of domperidone (1 mg/kg per os daily). Success rate is likely to be higher in transitional mares and mares maintained indoors under a stimulatory artificial photoperiod and in good
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body condition. Sulpiride could be combined with other treatment such as progestogen or eFSH for advancing and shortening the transitional phase and the first ovulation.31, 43, 44
References 1. Langlois B, Blouin C. Effect of a horse’s month of birth on its future performances. Proceedings of the 47th Annual Meeting of EAAP, 1996. 2. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects. Cross Plain, WI: Equiservices 1992, pp. 105–34. 3. Carnevale EM, Hermenet MJ, Ginther OJ. 1997. Age and pasture effects on vernal transition in mares. Theriogenology 1997;47(5):1009–18. 4. Palmer E, Guillaume D. 1992. Photoperiodism in the equine species: what is a long night? Anim Reprod Sci 1992;28(4):21–30. 5. Sharp DC, Davis SD. Vernal transition. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 133–43. 6. Nagy P, Guillaume D, Daels P. Seasonality in mares. Anim Reprod Sci 2000;60(special issue):245–62. 7. Cavinder CA, Vogelsang MM, Gibbs PG, Forrest DW, Schmitz DG. Endocrine profile comparison of fat versus moderately conditioned mares following parturition. J Equine Vet Sci 2007;27:72–9. 8. Fitzgerald BP, Reedy SE, Sessions DR, Powell DM, McManus CJ. Potential signals mediating the maintenance of reproductive activity during the non-breeding season of the mare. Reproduction Suppl. 2002;59:115–29. 9. Waller CA, Thompson DL, Cartmill JA, Storer WA, Huff NK. Reproduction in high body condition mares with high versus low leptin concentrations. Theriogenology 2006;66(4):923–8. 10. Ciechanowska M, Lapot M, Malewski T, Mateusiak K, Misztal T, Przekop F. Implication of dopaminergic systems on GnRH and GnRHR genes expression in the hypothalamus and GnRH-R gene expression in the anterior pituitary gland of anestrous ewes. Exp Clin Endocrinol Diab 2008;116(6):357–62. 11. Thiery JC, Malpaux B. Seasonal regulation of reproductive activity in sheep: modulation of access of sex steroids to the brain. Steroids Nerv Sys 2003;1007:169–75. 12. Melrose PA, Walker RF, Douglas RH. Dopamine in the cerebrospinal fluid of prepubertal and adult horses. Brain Behav Evol 1990;35:98–106. 13. Johnson AL. Serum concentrations of prolactin, thyroxine and triiodothyronine relative to season and the estrous cycle in the mare. J Anim Sci 1986;62(4):1012–20. 14. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995;73(3):899–908. 15. Nequin L.G., King S.S., Johnson A.L., Gow G.M., Ferreiradias GM. Prolactin may play a role in stimulating the equine ovary during the spring reproductive transition. J Equine Vet Sci 1993;13(11):631–5. 16. Thompson DLJ, DePew CL. Prolactin, gonadotropin, and hair shedding responses to daily sulpiride administration in geldings in winter. J Anim Sci 1997;75(4):1087–91. 17. Thompson DLJ, Hoffman R, DePew CL. Prolactin administration to seasonally anestrous mares: reproductive, metabolic, and hair-shedding responses. J Anim Sci 1997;75(4):1092–9.
18. King SS, Campbell AG, Dille EA, Roser JF, Murphy LL, Jones KL. Dopamine receptors in equine ovarian tissues. Domest Anim Endocrinol 2005;28:405–15. 19. King SS, Jones KL, Mullenix BA, Heath DT. Seasonal relationships between dopamine D1 and D2 receptor and equine FSH receptor mRNA in equine ovarian epithelium. Anim Reprod Sci 2008;108(4):259–66. 20. King SS, Roser JF, Jones KL. Follicular fluid prolactin and the periovulatory prolactin surge in the mare. J Equine Vet Sci 2008;28(8):468–72. 21. Brendemuehl JP, Cross DL. Influence of dopamine antagonist domperidone on the vernal transition in seasonally anoestrous mares. J Reprod Fertil Suppl 2000;56:185– 93. 22. Donadeu FX, Thompson DL. Administration of sulpiride to anovulatory mares in winter: effects on prolactin and gonadotropin concentrations, ovarian activity, ovulation and hair shedding. Theriogenology 2002;57(2):963–76. 23. Besognet B, Hansen BH, Daels PF. Dopaminergic regulation of gonadotrophin secretion in seasonally anoestrous mares. J Reprod Fertil 1996;108(1):55–61. 24. Besognet B, Hansen BS, Daels PF. Induction of reproductive function in anestrous mares using a dopamine antagonist. Theriogenology 1997;47(2):467–80. 25. Duchamp G, Daels PF. Combined effect of sulpiride and light treatment on the onset of cyclicity in anestrous mares. Theriogenology 2002;58:599–602. 26. Bennett-Wimbush K, Loch WE. A preliminary study of the efficacy of fluphenazine as a treatment for fescue toxicosis in gravid pony mares. J Equine Vet Sci 1998;18(3):169–74. 27. Bennett-Wimbush K, Loch WE, Plata-Madrid H, Evans T. The effects of perphenazine and bromocriptine on follicular dynamics and endocrine profiles in anestrous pony mares. Theriogenology 1998;49(4):717–33. 28. Kelley KK, Thompson DL, Storer WA, Mitcham PB, Gilley RM, Burns PJ. Estradiol interactions with dopamine antagonists in mares: prolactin secretion and reproductive traits. J Equine Vet Sci 2006;26(11):517–28. 29. McCue PM, Buchanan BR, Farquhar VJ, Squires EL, Cross DL. Efficacy of domperidone on induction of ovulation in anestrous and transitional mares. Proc Am Assoc Equine Pract 1999;45:217–18. 30. Brendemuehl JP, Cross DL. Effects of the dopamine antagonist domperidone on follicles, time to ovulation and luteal function in seasonally anestrus mares. In: Proceedings of the Society for Theriogenology, Kansas City, 1996; pp. 304–11. 31. McCue PM, Logan NL, Magee C. Management of the transition period: hormone therapy. Equine Vet Educ 2007;19(4):215–21. 32. Daels PF. Seasonal anestrus in mares: physiology, diagnosis and treatment. In: Proceedings of the Society for Theriogenology, Hastings, NB, 2000; pp. 189–96. 33. Daels PF, Fatone S, Hansen BS, Concannon PW. Dopamine antagonist-induced reproductive function in anoestrous mares: gonadotrophin secretion and the effects of environmental cues. J Reprod Fert Suppl 2000;56:173–83. 34. Nagy P, Bruneau B, Duchamp G, Daels PF, Guillaume D. Dopaminergic control of seasonal reproduction in mares. Reprod Dom Anim Suppl 1999;6:33. 35. King SS, Jones KL, Roser JF, Nequin LG, Murphy LL, Campbell AG. Evidence for a local dopamine-driven ovarian regulatory system in the mare. Theriogenology 2002;58(2–4):619–22. 36. Mari G, Morganti M, Merlo B, Castagnetti C, Parmeggiani F, Govoni N, Galeati G, Tamanini C. Administration of
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Dopamine Antagonists sulpiride or domperidone for advancing the first ovulation in deep anestrous mares. Theriogenology 2009;71(6):959– 65. 37. Thompson DLJ, Garza FJ, St George RL, Rabb MH, Barry BE, French DD. Relationships among LH, FSH and prolactin secretion, storage and response to secretagogue and hypothalamic GnRH content in ovariectomized pony mares administered testosterone, dihydrotestosterone, estradiol, progesterone, dexamethasone or follicular fluid. Domest Anim Endocrinol 1991;8(2):189–99. 38. Garcia MC, Ginther OJ. Regulation of plasma LH by estradiol and progesterone in ovariectomized mares. Biol Reprod 1978;19(2):447–53. 39. Advis JP, White SS, Ojeda SR. Delayed puberty induced by chronic suppression of prolactin release in the female rat. Endocrinology 1981;109(5):1321–30.
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40. Chowdhury M, Kabir SN, Pal AK, Pakrashi A. Modulation of luteinizing hormone receptors: effect of an inhibitor of prolactin secretion, p-coumaric acid. J Endocrinol 1983;98(3):307–11. 41. Champion MC, Hartnett M, Yen M. Domperidone, a new dopamine antagonist. Can Med Assoc J 1986;135:457–61. 42. Olin RB, Hebel SK, Connel SI. Drug Facts and Comparisons. St Louis, MO: Lippincott, 1991; p. 2561. 43. Rutten DR, Chaffaux S, Valon M, Deletang F, De Haas V. Progesterone therapy in mares with abnormal oestrous cycles. Vet Rec 1986;119(23):569–71. 44. Allen WR, Urwin V, Simpson DJ, Greenwood RE, Crowhurst RC, Ellis DR. Preliminary studies on the use of oral progestogen to induce oestrus and ovulation in seasonally anoestrous Thoroughbred mares. Equine Vet J 1980;12:141–5.
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Section (e): Pharmacological Manipulation of Reproduction
187.
Prostaglandins Simon A. Staempfli
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188.
Human Chorionic Gonadotropin John R. Newcombe
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189.
Progestagens and Progesterone Carlos R.F. Pinto
1811
190.
Gonadotropin-Releasing Hormones Edward L. Squires
1820
191.
Estrogen Therapy Ahmed Tibary
1825
192.
Oxytocin Michelle M. LeBlanc
1830
193.
Superovulation Edward L. Squires and Patrick M. McCue
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194.
Estrus Suppression Dirk K. Vanderwall and Gary J. Nie
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195.
Inhibin Immunotherapy Patrick M. McCue
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196.
Induction of Ovulation Angus O. McKinnon and Patrick M. McCue
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Synchronization of Ovulation Etta A. Bradecamp
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198.
New Techniques in Hormonal Delivery Patrick J. Burns
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199.
Contraception Jay F. Kirkpatrick and John W. Turner
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Chapter 187
Prostaglandins Simon A. Staempfli
Prostaglandin F2α (PGF2α ) In 1972, PGF2α was first reported to cause lysis of the corpus luteum, thereby allowing cycling mares to return to estrus 3 to 4 days after treatment.1 Since then, PGF2α has been extensively used in equine stud medicine to manipulate normal and abnormal estrous cycles, terminate early pregnancies, synchronize horses for breeding or embryo transfer programs, induce gonadotropin secretion, and treat uterine infections. The most frequently administered prostaglandins in equine practice are the naturally occurring PGF2α , dinoprost tromethamine R ; Pfizer Animal Health, New York), (Lutalyse and the synthetic prostaglandin analog for cattle, R ; Schering-Plough Animal cloprostenol (Estrumate R Health Corp., Union, NJ). Luprostiol (Prosolvin ; Virbac Animal Health, Bury St Edmunds, UK) and the recently developed PGF2α analog d-cloprostenol R ; Dr E. Graeub AG, Bern, Switzerland) (Genestran are two additional products registered in Europe for use in horses. Other PGF2α analogs such as fluprostenol, fenprostalene, or alfaprostenol have been used in some studies cited in this chapter, but are no longer commercially available.
Dosage and administration The standard recommended dose for a 500-kg horse is 5–10 mg for dinoprost tromethamine (10– 20 µg/kg), 7.5 mg for luprostiol (15 µg/kg), 250 µg for cloprostenol (0.55 µg/kg), and 22.5–37.5 µg for d-cloprostenol (0.045–0.075 µg/kg). Early studies showed that the minimal effective luteolytic dose rate of a single injection of free acid PGF2α in ponies was 6 µg/kg and in horses 9 µg/kg in comparison to 144 µg/kg for the ewe or the cow.2 The reason for the higher susceptibility of the equine corpus luteum to prostaglandin is because of the unique vascular anatomy of the mare’s reproductive tract. The mare
does not possess the characteristic intertwining of the uterine vein and the ovarian artery in the ovarian pedicle, as occurs in the ewe and other farm animals.3 Therefore, when endometrial PGF2α is released during Days 14 to 17 in the cycling mare, it can only reach the ovary by passing through the cardiovascular system, where it is subject to rapid metabolism in the lungs and other organs. The lack of a local utero-ovarian transport system in the mare results in lower peripheral prostaglandin levels during luteolysis, requiring an increased sensitivity of the corpus luteum to prostaglandin compared to other species, on a dose per bodyweight basis.4 Additionally, it is suspected that there is a slower peripheral degradation of PGF2α in the tissues of the horse, especially in the lungs, which permits systemic distribution and recirculation.5 Administration of PGF2α is equally effective by intramuscular, intravenous, intrauterine, subcutaneous, and intraluteal routes,4, 6, 7 although the intramuscular route is usually preferred for simplicity and reduced side-effects. However, even when prostaglandins are given at recommended doses by intramuscular injection, about 10% of horses experience some side-effects including sweating, diarrhea, or abdominal discomfort for up to 20 minutes.5 Early studies using varying doses of PGF2α showed dose-dependent side-effects, particularly in the frequency and severity of sweating and hyperthermia.6 Very large doses of PGF2α (i.e., up to 500 µg/kg PGF2α given intramuscularly or subcutaneously) produced ataxia, increased gastrointestinal motility, protracted diarrhea, hyperthermia, labored respiration, and changes in blood chemistry and hematological parameters. Nevertheless, no mares have reportedly died or appeared to sustain adverse effects following recovery.6 Although even very large doses are not life threatening, and the standard clinical dose appears to cause only transient discomfort in some horses, several recent studies have aimed to minimize the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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unpleasant side-effects by reducing the dose administered. Research published in 2001 showed that a microdose of 25 µg cloprostenol administered intramuscularly was as effective at lysing the equine corpus luteum as the larger standard dose (250 µg).8 Two other studies induced luteolysis with a single administration of 1.25 mg dinoprost,9, 10 and one publication reported successful luteolysis with administration of two injections of 0.5 mg of dinoprost 24 hours apart.11 In all four studies, the investigators did not observe the occurrence of PGF2α -induced side-effects in treated mares.
Indications
Termination of a normal luteal phase In a high proportion of normally cycling mares, the corpus luteum is not responsive to one single injection of PGF2α until day 5 after ovulation.12 However, recent studies have discovered that treatment during early diestrus is capable of inducing functional regression of the corpus luteum. Bergfelt and co-workers investigated the response of the corpus luteum in the mare after injection of a single dose of PGF2α (10 mg/mare) at 3 days after ovulation.13 They observed a functional and structural regression of the corpus luteum, based on an abrupt decrease in circulating concentrations of progesterone, a decrease in luteal gland diameter, and an increase in luteal tissue echogenicity. Three days after treatment, functional resurgence of the corpus luteum was detected in 75% of the mares (12 of 16), as indicated by a transient major (six mares) and minor (six mares) increase in progesterone. Despite the functional resurgence in progesterone, the mean length of the interovulatory interval in all mares treated on day 3 after ovulation was significantly shortened (13.2 ± 0.9 days), compared to mares injected on Day 10 after ovulation (19.2 ± 0.7 days).13 When PGF2α (2.5 mg/mare) was administered once daily on Days 2, 3, and 4 after ovulation, 6 out of 10 mares underwent complete luteolysis.14 The six mares that underwent complete luteolysis ovulated 9.4 ± 1.36 days after treatment. The remaining four mares underwent partial luteolysis, followed by resurgence in circulating progesterone concentrations. Interestingly, three of the mares undergoing partial luteolysis ovulated prematurely with high concentrations of circulating progesterone at days 9, 15, and 16 after treatment.14 When 10 mg of PGF2α was administered twice daily on days 0, 1, and 2 post-ovulation, the mean plasma progesterone concentration on day 4 remained < 1.0 ng/mL, and four of six mares ovulated 7.0 ± 1.8 days after treatment.15 Unlike the horse, the donkey corpus luteum seems to be susceptible earlier to prostaglandin-
induced luteolysis. A recent study using 22 Martina Franca jennies showed that PGF2α administered on day 3 post-ovulation caused complete structural and functional regression of the corpus luteum, with induction of estrus within 4 days in 95.5% of treated animals.16 Timing of prostaglandin administration during diestrus affects outcome. On average, mares return to estrus 3 to 4 days after administration, and ovulation occurs 8 to 10 days after treatment.17 It is important to remember that elevated blood progesterone concentrations do not suppress increases in blood concentration of follicle-stimulating hormone (FSH) in the horse; therefore follicular development continues during diestrus. Consequently, there are often follicles of various sizes and stages of growth during the luteal phase. The size of the follicle at the time of prostaglandin administration is very important for predicting the interval to estrus and ovulation. When only small follicles are present at the time of prostaglandin administration, a new follicular wave has to emerge and ovulation of a dominant follicle occurs 7 to 12 days after treatment.18 Mares with medium-sized follicles (2.5 to 3.5 cm) will show signs of estrus soon after PGF2α administration and ovulate within 3 to 6 days. When there is a large growing follicle present (3.5 to 5 cm) at the time of treatment, ovulation is possible within 24 to 72 hours after treatment, without overt signs of estrus (standing heat, relaxation of the cervix, endometrial edema).19, 20 Conversely, if the follicle has reached its maximal diameter during the progesterone-dominated luteal phase, this follicle is destined for atresia and will inevitably regress and a new cohort of follicles will have to be recruited, therefore estrus and ovulation will be delayed.21 If a large growing follicle is present, administration of microdoses of prostaglandin (1.25 mg dinoprost) several times daily until regression of the corpus luteum, has been successful in bringing mares slowly into estrus, and thereby preventing the dominant follicle from ovulating too rapidly (A.O. McKinnon, personal communication, 2008). The degree of endometrial edema can be used to predict the response to treatment, as mares with a large, growing follicle can have substantial endometrial edema within 24 hours after treatment. However, mares with large atretic follicles do not exhibit endometrial edema until 3 to 5 days after treatment.22
Termination of a persistent luteal state The corpus that forms after ovulation is usually functional for 14 to 15 days in the non-pregnant mare,23 after which time luteolysis occurs as a result of prostaglandin release from the endometrium.24
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Prostaglandins Corpora lutea that fail to regress at the normal time post-ovulation are considered to be pathologically persistent.25 The most common causes of a persistent luteal phase are ovulations late in diestrus, resulting in immature corpora lutea (< 5 days old) at the time of prostaglandin release,25 embryonic loss after the time of maternal recognition of pregnancy, chronic uterine infections with destruction of the endometrium and therefore diminished prostaglandin release, and formation of a luteinized anovulatory follicle.26 The persistent luteal stage may persist for 2 to 3 months, during which time the mare will not express behavioral estrus. Regardless of the cause of the persistent luteal state, administration of a single intramuscular dose of dinoprost tromethamine (5–10 mg) or cloprostenol (250 µg) will result in luteolysis and subsequent estrus in more than 80% of cases.5
Estrous synchronization Synchronization of estrus and ovulation may allow mares to be bred or inseminated at a predetermined time, thus simplifying breeding management and providing more efficient use of heavily booked stallions over a predetermined time period. Additionally, estrous synchronization is an integral part of equine embryo transfer programs, where a close synchrony between donor and recipient mare is crucial for survival of the transferred embryo. Synchronization of estrus has been used successfully as a management tool in the cattle industry, where the administration of two doses of prostaglandin 11 to 12 days apart results in more than 80% of the cows exhibiting estrus within 2 to 5 days after the second injection.27 Although administration of two doses of prostaglandin 14 days apart has been described as a protocol for estrus synchronization in mares, it is not a very reliable method of ensuring that a recipient mare will be synchronized with a donor mare.19 After two injections of prostaglandin, ovulation occurs on average 7 to 10 days after the second injection, but ovulation can potentially occur anywhere from 0 to 17 days.28 Therefore it was concluded that one would need to treat at least 10 recipient mares to have an 80% chance that at least one recipient would ovulate within 24 hours of the donor.29 Due to the large variation in time of ovulation from administration of prostaglandin, a double injection scheme is not very useful when used alone in the synchronization of the estrous cycles in mares. However, if examination via transrectal ultrasonography is used to eliminate those mares that will not respond appropriately to the administration of prostaglandin, tightening of the date of ovulation can be achieved,
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especially when ovulation induction is included in the protocol.
Induction of gonadotropin secretion Experiments in farm animals have shown that PGF2α can increase blood luteinizing hormone (LH) concentration. Treatment with PGF2α increased LH concentration in bulls,30, 31 diestrous mares,32, 33 and transitional anestrous mares.34 In transitional anestrous mares with low circulating progesterone concentrations, the PGF2α analog luprostiol increased LH concentration in the pituitary venous blood. The initial rise in LH concentration was attributed to a direct action on the pituitary gland. Following the rise in LH, there was an increase in gonadotropin-releasing hormone (GnRH), which in turn sustained the elevation of LH for 2 hours post-treatment.34 Similarly, in early diestrus intravenous cloprostenol caused an immediate marked increase in FSH and LH secretion, which lasted 1 to 2 hours and was not associated with any increase in GnRH concentration.5 Unlike in anestrus or diestrus, PGF2α and its analogs failed to induce an increase in LH and FSH concentration during estrus.35, 36 It is suspected that hypothalamic or pituitary prostaglandin synthetic capabilities and receptor numbers are markedly different in diestrous and transitional anestrous mares compared to estrous mares. Despite these findings, one study reported that the PGF2α analog, fenprostalene, induced ovulation in 81% of treated mares within 48 hours following treatment compared to 31% of control mares.37 The exact mechanism by which fenprostalene was able to hasten ovulation remains unknown. The authors suggested that the site of action was either along the hypothalamic–pituitary axis or at the ovary.
Induction of abortion Prostaglandins are the preferred method of inducing abortion of the early equine pregnancy. As described previously, the corpus luteum is not particularly responsive during the first 5 days after ovulation. Thereafter administration of a single injection of dinoprost (5–10 mg) or cloprostenol (250 µg) will result in luteolysis, with subsequent loss of the conceptus from day 5 to day 30 of gestation,38 although in some cases after maternal recognition of pregnancy two or more consecutive injections may be necessary to lyse the primary corpus luteum.39 Abortion can be expected 2 to 5 days after treatment.40 The endometrial cups, a band of specialized trophoblastic cells invading the endometrium, are established during days 28 to 35 of pregnancy.3 The cup cells produce equine chorionic gonadotropin (eCG),
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which stimulates the primary corpus luteum and the formation of secondary or accessory corpora lutea (jointly known as supplemental corpora lutea).39 Therefore, multiple injections of prostaglandin are necessary for successful termination of pregnancy after establishment of the endometrial cups. Two studies showed that all mares given 2.5 to 5.0 mg PGF2α or 250 µg of fluprostenol, once or twice a day, aborted 3 to 5 days after starting treatment.41, 42 Although the pregnancy is terminated, the endometrial cups remain functional until they are removed by the maternal immune system approximately 60–80 days after establishment.43 As long as the cups are functional and eCG is produced, the mare will not return to normal estrous cycles after pregnancy termination. Elective termination of pregnancy beyond the first trimester may be complicated by dystocia, retained fetal membranes, or trauma to the genital tract. Although multiple injections of PGF2α have resulted in abortions in mares between 100 and 245 days of gestation,44 two injections of fluprostenol or prostaglandin administered between 160 and 320 days of gestation failed to induce abortion.45 Therefore PGF2α is not the preferred method of inducing abortion after the first trimester of pregnancy, and other methods such as large-volume saline infusions should be considered.46
Induction of parturition Induction of parturition with PGF2α is possible,47, 48 but no longer recommended because of an increased incidence of premature placental separation and decreased foal viability associated with its use. Detailed information on induction of parturition is presented in Chapter 232.
Treatment of uterine infections Progesterone predisposes the uterus to infection by constricting the cervix and inhibiting uterine contractions, whereas estradiol has the opposite effect on the uterus and also stimulates the local immune system.5 Administration of PGF2α to diestrous mares allows them to come into heat, thereby increasing estrogens and relaxing the cervix. More importantly, PGF2α has a direct effect on the myometrium.49–51 Prostaglandins have been demonstrated to be released very early in mares with endometritis, and this has subsequently been shown to play a useful role in increasing myometrial activity and assisting in uterine clearance.52 While oxytocin induces strong uterine contractions for 30 to 50 minutes, PGF2α produces low-amplitude contractions that persist for 4 to 5 hours.53 Oxytocin clears radiocolloid more rapidly than prostaglandins,54 but prostaglandin-
induced uterine contractions may assist in lymphatic flow as lymph drainage relies on myometrial contractions.55 Cloprostenol produces a more consistent response than PGF2α and fenprostalene.54 The suggested dose of cloprostenol is 250 µg given at 12 and 24 hours after ovulation. Cloprostenol needs to be administered with caution after ovulation because administration after Day 2 from ovulation is associated with decreased plasma progesterone concentrations on Days 3 through 7 of the estrous cycle. Although plasma progesterone concentrations rebounded by Days 7 to 9 of the estrous cycle, two reports showed decreased pregnancy rates when cloprostenol was given at 500 µg until Day 2 after ovulation.56, 57 When 250 µg was given every 24 hours from 4 hours after breeding until Day 2 post-ovulation, pregnancy rates were similar to those of mares treated post-ovulation with oxytocin.58
Prostaglandin E2 (PGE2 ) Dinoprostone, a naturally occurring PGE2 , has been widely used in human gynecology as a vaginal suppository for induction of abortion, labor, and initiation of cervical ripening near term.59–61 Although dinoprostone is not approved for use in horses, the effects of PGE2 on the reproductive tract in the mare and other species have been extensively investigated in research studies over the last 20 years. As in human medicine, it is thought that PGE2 induces cervical relaxation in the mare near term.62 Additionally, PGE2 appears to play a role in gamete and embryo transport through the oviduct.63 One study showed the potential role of PGE2 as a stimulator of uterine contractions during the embryo mobility phase of the first two weeks of pregnancy.64 Indications
Accelerated spermatozoal transport through the reproductive tract PGE2 may play an important role in gamete and embryo transport through the oviduct, by increasing oviductal smooth muscle contraction. Electromyographic activity of the equine oviduct is stimulated by intramuscular administration of 10 mg PGE2 ,53 and locally applied PGE2 induces relaxation of circular smooth muscle of the oviductal isthmus.65 Smooth muscle contractions at the utero-tubular junction (UTJ) may play a role in propulsion of spermatozoa through the UTJ into the isthmus in the pig.66 Addition of PGE2 to ram semen increased pregnancy rates after artificial insemination in ewes.67 Although PGE2 has been shown to promote Ca2+ influx and acrosome reaction in human spermatozoa,68 it remains
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Prostaglandins unknown whether increased pregnancy rates after addition of PGE2 to the inseminate in some studies were caused by accelerated spermatozoal transport through the reproductive tract or by direct effects of PGE2 on spermatozoa. The intrauterine infusion of 0.25 mg PGE2 2 hours prior to the insemination of good-quality semen improved pregnancy rates in a study of 34 light-breed mares; however, no improvement in pregnancy rates was observed when a stallion with poor semen quality was used.69 Further research is needed to determine the effects of locally applied PGE2 and pregnancy rates achieved by subfertile stallions.
Re-establishment of oviductal patency In 2006 Allen and co-workers reported laparoscopic application of PGE2 to re-establish oviductal patency in infertile mares.70 The study was performed on 15 infertile mares, with no identifiable pathology of the reproductive tract, that consistently failed to conceive when bred to fertile stallions during one to four breeding seasons. The authors hypothesized that the oviducts of these mares were temporarily obstructed by naturally occurring debris (oviductal masses), thus preventing oviductal transport of the embryo to the uterus. After application of 0.2 mg of PGE2 onto the external surface of the oviduct, 93% (14/15) of the mares conceived within the same or the subsequent breeding season. Although these results are very promising, this study is limited by a lack of controls and no direct evidence that oviductal blockage was the cause of infertility.
Cervical ripening prior to induction of parturition Intracervical administration of PGE2 in women results in cervical ripening, a term used in human medicine to describe softening and dilation of the cervix.61 It is believed that PGE2 leads to a relaxation of the cervical smooth muscle fibers, resulting in a dilated cervix, which allows passage of the fetus through the birth canal.60 In women, intracervical deposition of PGE2 has been associated with a shorter induction-to-delivery interval, a lower number of required caesarean sections, and a decreased maximum dosage of oxytocin required to induce parturition.71, 72 In cattle, intravaginal application of 2 mg dinoprostone increased cervical mucus secretion, and significantly relaxed the cervix within 6 to 10 hours after application.73 In a controlled equine study, Rigby and co-workers deposited 2.0 to 2.5 mg PGE2 intracervically 6 hours prior to induction of parturition at term, and observed shorter delivery periods, facilitated fetal delivery, and increased
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foal vigor in treated mares.62 PGE2 has also been combined with estrogens (estradiol cypionate, 6 mg, given 24 hours before induction) to dilate the cervix in a protocol for oxytocin-induced parturition.74 To the author’s knowledge there are no published controlled studies on the use of PGE2 or PGE1 for cervical dilation in mares with impaired cervical drainage and subsequent uterine fluid accumulation during estrus. Topical application of PGE1 4 to 6 hours before natural mating or artificial insemination, was reported to relax the cervix for up to 8 hours after treatment.75 In some cases multiple applications may be necessary (A.O. McKinnon, personal communication, 2008), although repeated use has been associated with inflammation of the cervical and vaginal mucosa.75 However, success of PGE2 treatment in older maiden mares with a tight cervix remains variable, due to a decrease in elastic tissue and an increase in the amount of fibrotic tissue in those mares.74
Summary Despite a wide range of possible applications of prostaglandins in equine stud medicine, the major indication for PGF2α treatment remains luteolysis. Recent studies suggest that doses smaller than those recommended by the manufacturer are effective in inducing complete luteolysis by 5 days after ovulation. The subsequent induced estrus is characterized by normal ovarian dynamics and conception rates.
References 1. Douglas RH, Ginther OJ. Effect of prostaglandin F2α on length of diestrus in mares. Prostaglandins 1972;2:265–8. 2. Oxender WD, Noden PA, Bolenbaugh DL, Hafs HD. Control of estrus with prostaglandin F2α in mares: minimal effective dose and stage of estrous cycle. Am J Vet Res 1975;36:1145–7. 3. Ginther OJ. Equine pregnancy: physical interactions between the uterus and conceptus. P Annu Conv Am Assoc Equine 1998;44:73–104. 4. Douglas RH, Ginther OJ. Route of prostaglandin F2α injection and luteolysis in mares. P Soc Exp Biol Med 1975;148:263–9. 5. Irvine CH. Prostaglandins. In: McKinnon AO, Voss J (eds) Equine Reproduction. London: Lea & Febiger, 1993; pp. 319– 24 6. Lauderdale J, Miller P. Regulation of reproduction in mares with prostaglandins. P Annu Conv Am Assoc Equine 1975;21:235–44. 7. Weber JA, Causey RC, Emmans EE. Induction of luteolysis in mares by ultrasound-guided intraluteal treatment with PGF2α. Theriogenology 2001;55:1769–76. 8. Nie GJ, Goodin AN, Braden TD, Wenzel JGW. Luteal and clinical response following administration of dinoprost tromethamine or cloprostenol at standard intramuscular sites or at the lumbosacral acupuncture point in mares. Am J Vet Res 2001;62:1285–9.
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9. Wuestenhagen A, Handler J, Kindahl H, Aurich C. Luteal function and estrous cycle characteristics in mares treated with subtherapeutic doses of prostaglandin F2α. Theriogenology 2002;58:537–9. 10. Barker C, Echeverria K, Davis M, Whisnant CS, Pinto CRF. Effects of different doses of PGF2α on luteal function and on the subsequent estrous cycle. Anim Reprod Sci 2006;94:207–9. 11. Irvine CHG, McKeough VL, Turner JE, Alexander SL, Taylor TB. Effectiveness of a two-dose regimen of prostaglandin administration in inducing luteolysis without adverse side effects in mares. Equine Vet J 2002;34:191–4. 12. Allen WR, Rowson LEA. Control of the mare’s oestrous cycle by prostaglandins. J Reprod Fertil 1973;33:539–43. 13. Bergfelt DR, Pierson RA, Ginther OJ. Regression and resurgence of the CL following PGF2α treatment 3 days after ovulation in mares. Theriogenology 2006;65:1605–19. 14. Holland B, Pinto CRF. Luteal function and ovulation in mares treated with PGF2α during early and mid-diestrus. Reprod Domest Anim 2008;43:111. 15. Rubio C, Pinto CRF, Holland BE, Da Silva SA, Layne SA, Heaton LH, Whisnant CS. Anti-luteogenic and luteolytic effects of PGF2α during the post-ovulatory period in mares. Theriogenology 2008;70:587. 16. Carluccio A, Panzani S, Contri A, Tosi U, De Amicis I, Veronesi MC. Luteal function in jennies following PGF2α treatment 3 days after ovulation. Theriogenology 2008;70:121–5. 17. Loy RG,. Buell JR, Stevenson W, Hamm D. Sources of variation in response intervals after prostaglandin treatment in mares with functional corpora lutea. J Reprod Fertil Suppl 1979;27:229–35. 18. McKinnon AO, Squires EL. Effect of follicular status on response time following prostaglandin. Unpublished work cited in Asbury AC. P Annu Conv Am Assoc Equine, 1988, pp. 191–6. 19. Bristol F. Control of the mare’s reproductive cycle. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1987, pp. 309–21. 20. Asbury AC. The use of prostaglandins in broodmare practice. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners 1988, pp. 191–196. 21. Samper JC. Induction of estrus and ovulation: Why some mares respond and others do not. Theriogenology 2008;70:445–7. 22. Samper JC, Geertsema H, Hearn P. Rate of luteolysis, folliculogenesis and interval to ovulation of mares treated with a prostaglandin analogue on d 6 or 10 of the estrous cycle. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 1993, pp. 169–71. 23. Ginther OJ, Pierson RA. Regular and irregular characteristics of ovulation and the interovulatory interval in mares. J Equine Vet Sci 1989;9:4–12. 24. Neely DP, Kindahl H, Stabenfeldt GH. Prostaglandin release patterns in the mare: Physiological, pathophysiological, and therapeutic responses. J Reprod Fertil Suppl 1979;27:181–9. 25. Stabenfeldt GH, Hughes JP, Evans JW, Neely DP. Spontaneous prolongation of luteal activity in the mare. Equine Vet J 1974;6:158–63. 26. McCue PM. Review of ovarian abnormalities in the mare. P Annu Conv Am Assoc Equine 1998;44:125–33. 27. Wright PJ, Malmo J. Pharmacologic manipulation of fertility. Vet Clin N Am-Food A 1992;8:57–89.
28. Bradecamp EA. Estrous synchronization. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 22–5. 29. Irvine CHG. Endocrinology of the estrous cycle of the mare: Applications to the embryo transfer. Theriogenology 1982;15:895. 30. Haynes NB, Kiser TE, Hafs HD, Marks JD. Prostaglandin F2α overcomes blockade of episodic LH secretion with testosterone, melengestrol acetate or aspirin in bulls. Biol Reprod 1977;17:723–8. 31. Haynes NB, Collier RJ, Kiser TE, Hafs HD. Effect of prostaglandin E2 and F2α on serum luteinizing hormone, testosterone, and prolactin in bulls. J Anim Sci 1978;47:923–6. 32. Noden PA, Oxender WD, Hafs HD. Early changes in serum progesterone, estradiol, and LH during prostaglandin F2α induced lutolysis in mares. J Anim Sci 1978;47:666– 71. 33. Nett TM, Pickett BW, Squires EL. Effects of equimate (ICI81008) on levels of luteinizing hormone, follicle stimulating hormone, and progesterone during the estrous cycle of the mare. J Anim Sci 1979;48:69–75. 34. Jochle W, Irvine CHG. Release of LH, FSH and GnRH into pituitary venous blood in mares treated with PGF analogue, luprositol, during the transition period. J Reprod Fertil Suppl 1987;35:261–7. 35. Harrison L, Squires E, McKinnon AO. Acute effects of luprositol on LH and FSH duing estrus in cycling mares. In: Proceedings of the 10th Equine Nutrition and Physiology Symposium, 1987, pp. 265–9. 36. Crickman JA, Momont HW, Al-Hassan MJ. Plasma LH concentration in the mare following administration of alfaprostenol during oestrus. J Reprod Fertil Suppl 1991;44:686–8. 37. Savage NC, Liptrap RM. Induction of ovulation in cyclic mares by administration of a synthetic prostaglandin, fenprostalene, during oestrus. J Reprod Fertil Suppl 1987;35:239–43. 38. Penzhorn BL, Bertschinger HJ, Coubrough RI. Reconception of mares following termination of pregnancy with prostaglandin F2α before and after day 35 of pregnancy. Equine Vet J 1986;18:215–17. 39. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992. 40. Baucus KL, Squires EL, Morris R, McKinnon AO. The effect of stage of gestation and frequency of prostaglandin injection on induction of abortion in mares. In: Proceedings of the 10th Equine Nutrition and Physiology Symposium, 1987, 255–8. 41. Squires EL, Hillman RB, Pickett BW, Nett TM. Induction of abortion in mares with equimate: effect on secretion of progesterone, PMSG and reproductive performance. J Anim Sci 1980;50:490–5. 42. Rathwell AC, Asbury AC, Hansen PJ, Archbald LF. Reproductive function of mares given daily injections of prostaglandin F2α beginning at day 42 of pregnancy. Theriogenology 1987;27:621–30. 43. Allen WR. Immunological aspects of the endometrial cup reaction and the effect of xenogenic pregnancy in horses and donkeys. J Reprod Fertil Suppl 1982;31:57–94. 44. Madej A, Kindahl H, Nydahl C, Edqvist LE, Stewart DR. Hormonal changes associated with induced late abortions in the mare. J Reprod Fertil Suppl 1987;35:479–84. 45. Leadon DP, Rossdale PD, Jeffcott LB, Allen WR. A comparison of agents for inducing parturition in mares in the
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pre-viable and premature periods of gestation. J Reprod Fertil Suppl 1982;32:597–602. Paccamonti DL. Elective termination of pregnancy in mares. J Am Vet Med Assoc 1991;198:683–9. Jeffcott LB, Rossdale PD. A critical review of current methods for induction of parturition in the mare. Equine Vet J 1977;9:208–15. Ley WB, Hoffman JL, Crisman MV, Meacham TN, Kiracofe RL, Sullivan TL. Daytime foaling management of the mare 2: Induction of parturition. J Equine Vet Sci 1989;9:95–9. Goddard PJ, Allen WE. Genital tract pressures in mares II. Changes induced by oxytocin and prostaglandin F2α. Theriogenology 1985;24:35–44. Cross DT, Ginther OJ. The effect of estrogen, progesterone and prostaglandin F2α on uterine contractions in seasonally anovulatory mares. Domest Anim Endocrinol 1987;4:271–8. Cross DT, Threlfall WR, Rogers RC, Kline RC. Uterine electromyographic activity in horse mares as measured by radiotelemetry. Theriogenology 1991;35:591–601. Cadario ME, Merritt AM, Archbald LF, Thatcher WW, LeBlanc MM. Changes in intrauterine pressure after oxytocin administration in reproductively normal mares and in those with a delay in uterine clearance. Theriogenology 1999;51:1017–25. Troedsson MHT, Liu IKM, Ing M, Pascoe JR. Smooth muscle electrical activity in the oviduct, and the effect of oxytocin, prostaglandin F2α, prostaglandin E2 on the myometrium and the oviduct of the cycling mare. Biol Reprod Monogr 1995;1:475–88. Combs GB, LeBlanc MM, Neuwirth L, Tran TQ. Effects of prostaglandin F2α, cloprostenol and fenprostalene on uterine clearance of radiocolloid in the mare. Theriogenology 1995;45:1449–55. LeBlanc MM. Persistent mating induced endometritis in the mare: pathogenesis, diagnosis and treatment. Electronic citation. International Veterinary Information Service (www.ivis.org) 2003. Troedsson MHT, Ababneh MM, Ohlgren AF, Madill S, Vetscher N, Gregas M. Effect of periovulatory prostaglandin F2α on pregnancy rates and luteal function in the mare. Theriogenology 2001;55:1891–9. Brendemuehl JP. Effect of oxytocin and cloprostenol on luteal formation, function and pregnancy rates in mares. Theriogenology 2002;58:623–6. Nie GJ, Johnson KE, Wenzel JGW, Braden TD. Effect of administering oxytocin or cloprostenol in the periovulatory period on pregnancy outcome and luteal function in mares. Theriogenology 2003;60:1111–18. Thiery M. Induction of labor with prostaglandins. In: Keirse MJNC, Anderson ABM, Gravenhorst JB (eds) Human Parturition. Boston. Martinus Nijhoff, 1979; pp. 155–64. Bryman I, Lindblom B, Norstrom A. Extreme sensitivity of cervical musculature to prostaglandin E2 in early pregnancy. Lancet 1982;ii:1471.
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61. Trofatter KF, Bowers D, Gall SA, Killam AP. Preinduction cervical ripening with prostaglandin E2 gel. Am J Obstet Gynecol 1985;153:268–71. 62. Rigby S, Love C, Carpenter K, Varner D, Blanchard T. Use of prostaglandin E2 to ripen the cervix of the mare prior to induction of parturition. Theriogenology 1998;50:897– 904. 63. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 hastens oviductal transport of equine embryos. Biol Reprod 1991;45:544–6. 64. Gastal MO, Gastal EL, Torres CAA, Ginther OJ. Effect of PGE2 on uterine contractility and tone in mares. Theriogenology 1998;50:989–99. 65. Weber JA, Woods GL, Lichtenwalner AB. Relaxatory effect of prostaglandin E2 on circular smooth muscle isolated from the equine oviductal isthmus. Biol Reprod Monogr 1995;1:125–30. 66. Persson E, Rodriguez-Martinez H. The ligamentum infundibulo-cornuale in the pig: morphological and physiological studies on the smooth muscle component. Acta Anat 1990;138:111–20. 67. Dimov V, Georgiev F. Ram semen prostaglandin concentration and its effect on fertility. J Anim Sci 1977;44: 1050–4. 68. Shaefer M, Hofmann T, Schultz G, Gudermann T. A new prostaglandin E receptor mediates calcium influx and acrosome reaction in human spermatozoa. Cell Biol 1998;95:3008–13. 69. Woods J, Rigby S, Brinsko S, Stephens R, Varner D, Blanchard T. Effect of intrauterine treatment with prostaglandin E2 prior to insemination of mares in the uterine horn or body. Theriogenology 2000;53:1827–36. 70. Allen WR, Wilsher S, Morris L, Crowhurst JS, Hillyer MH, Neal HN. Laparoscopic application of PGE2 to re-establish oviductal patency and fertility in infertile mares: a preliminary study. Equine Vet J 2006;38:454–9. 71. O’Herlihy C, MacDonald HN. Influence of preinduction prostaglandin E2 vaginal gel on cervical ripening and labor. Obstet Gynecol 1979;54:708–10. 72. Ferguson JE, Ueland FR, Stevenson DK, Ueland K. Oxytocin-induced labor characteristics and uterine activity after preinduction cervical priming with prostaglandin E2 intracervical gel. Obstet Gynecol 1988;72:739– 45. 73. Duchens M, Fredriksson G, Kindahl H, Aiumlamai S. Effect of intracervical administration of a prostaglandin E2 gel in pregnant and non-pregnant heifers. Vet Rec 1993;133: 546–9. 74. Estrada AJ, Samper JC. Cervical failure to dilate. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders, Elsevier, 2007; pp. 134–6 75. LeBlanc MM. Reproduction deduction – part 2. In: Proceedings of the North American Veterinary Conference 20, 2006, pp. 139–42.
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Human Chorionic Gonadotropin John R. Newcombe
Introduction The use of human chorionic gonadotropin (hCG) to advance ovulation in mares was first suggested by Fred Day, a practitioner, in 1939. It has been widely used in broodmare practice since then and, although various gonadotropin-releasing hormone (GnRH) agonists are now available, it is probably still the drug of choice for most practitioners. HCG is a protein consisting of two peptide chains containing galactose and hexosamine. Its molecular weight is 30,000 and it has a half-life of between 8 and 12 hours.1 It is formed in the chorionic villi of the human placenta and is detectable from only a few days after conception, peaking at about 35–50 days. In primates it is an essential luteotrophin to maintain and stimulate luteal function until the placenta is sufficiently developed to maintain the pregnancy, which is at least the seventh week in women.2 In the human it is not only luteotrophic but possibly it is also luteoprotective against endogenous prostaglandin release, binding to the luteinizing hormone (LH) receptors. It has LH-like activity in most species and promotes ovulation and development of the corpus luteum (CL).3 When injected into mares in mid- to late estrus, it accelerates follicle maturation and hence ovulation occurs sooner, while estrus is also shortened. Its role in the development of the CL is not clear since, routinely, its activity will have waned before the time of ovulation.
Preparation and presentation HCG is available commercially as a freeze-dried preparation of 1500, 2000, 5000, or 10 000 international units (IU) supplied with a solvent. It is
intended that the powder be resuspended immediately prior to administration and discarded after 24 hours. The manufacturers of Chorulon (Intervet UK Ltd) suggest doses of 1500 or 3000 IU regardless of bodyweight. They warn of rare cases of anaphylactoid reaction.
Use Although it is unlikely that hCG will promote the ovulation of follicles that would not otherwise ovulate spontaneously, it allows for a more predictable timing of ovulation and may even reduce the interval between ovulations of mares destined to have multiple ovulations. Therefore, it is possible to delay the mating or insemination of mares until relatively close to ovulation without the need for further inseminations. When demand for semen is high and when a stallion may have 100 or more mares to mate in a season, it is essential during the busiest part of the breeding season that no mare should need to be mated more than once per estrous. A recent development in broodmare practice is ‘appointment mating’. The mare resides at the owner’s farm or locally but is given an appointment (time and date) to visit the stallion, which may be a few, but is often several hundred, miles away. The mare is transported to the stallion farm, mated, and returned home directly. Although this system saves money in keep/agistment charges, transport to meet an appointment several hundred miles away can be very expensive. It is therefore highly desirable that, by the next examination after mating, the mare is found to have ovulated and does not have to visit the stallion again in that same estrous period.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Human Chorionic Gonadotropin When fresh cooled semen has to be ordered in advance and then transported overnight across continents, it is important that it arrives within 24 hours, or preferably even nearer, of ovulation, since the longevity of the semen prior to some stallions can be seriously impaired by cooling. Similarly frozen/ thawed semen needs to be inseminated within 6–12 hours either side of ovulation for optimum results. The use of hCG greatly assists these timings.
Timing and follicle diameter Developing follicles are treated to accelerate and define the time of ovulation. HCG will not induce ovulation of atretic follicles. These are most commonly found in the spring transition but may also be found in cyclic mares in early to mid-estrus. The latter are follicles which have developed to near pre-ovulatory diameters during diestrus but become atretic at, or just after, luteolysis. At examination of a mare for the first time in early estrus it will not be known whether such follicles are atretic or whether they have developed since luteolysis without overt estrus. Knowledge of the interval since the previous ovulation is helpful in recognizing such follicles. However, large follicles still developing at the time of luteolysis may ovulate rapidly, resulting in a short follicular phase, a shortened interovulatory interval, and a very short period of estrus. Atretic follicles may or may not show echogenic particles floating in the antral fluid but some also show palpable softening. If it were known when a follicle would ovulate spontaneously then hCG could be given 3–4 days ahead. Unfortunately, this is not known and therefore the time of ovulation has to be predicted on the basis of clinical judgment and experience.
Diameter at ovulation and after hCG Evans et al.4 showed that the immediate preovulatory diameter was smaller after hCG treatment (38.7 mm) than controls (44.9 mm). In fact, the follicles of hCG-treated mares grew only about 3 mm in 42 hours (1.7 mm/day), whereas in the controls they continued to grow at about 2.25 mm/day for 4 days. This observation that, after hCG, follicle growth slows was confirmed.5 A recent study suggested there was little or no growth following hCG in follicles which subsequently ovulated within 48 hours.6
Efficacy There is no evidence that ovulations induced by hCG (or any GnRH preparation) are any less fertile than spontaneous ovulations. Some reports suggest enhanced fertility,7–11 but others suggest no advantage
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in normally cyclic mares.12–21 One report13 suggested that, although hCG mares may fail to conceive at the first estrus, it enhances their conception rate at subsequent cycles without further treatment. Given at the correct time, when a developing ovulatory follicle is about 2–4 days away from ovulation, ovulation can normally be assured in the period 36–48 hours after treatment. However, most reports suggest that, in practice, only about 85% of follicles ovulate within 48 hours. Brugmans22 reported 91% ovulating between 24 and 48 hours when follicles of 35–40 mm were treated with 1500 IU intravenously. Barbacini et al.23 reported that only 76% of mares ovulated in the period between 25 and 48 hours when mares with 35 mm or larger follicles were given 2000 IU intravenously. Another 16% ovulated within the first 24 hours, whereas 9% took more than 48 hours. It is noteworthy that in this report only 42% ovulated within the period 37–48 hours. Some of these observations may have varied owing to breed (see below) and experience of the authors. Other studies clearly show at least 90% of mares ovulating in 36 ± 4 hours under controlled conditions.24, 25 The ovulation of follicles which are already destined to ovulate within 0–36 hours from the time of treatment may not be accelerated. When hCG is given, it must be remembered that ovulation can occur before 36 hours. It should be noted that, in the report by Barbacini et al.,23 50% of mares ovulated before 36 hours. However, experienced practitioners will be aware of this possibility, either from previous experience with a particular mare or from the various other palpable and ultrasonic parameters, apart from diameter, that are used to predict the time of ovulation. Although it is considered routine to treat with hCG when a follicle has grown to 35 mm or larger, some mares will ovulate from diameters considerably smaller than this, especially in summer and when multiple follicles are developing. Also there are individual mares and breeds of horses which regularly ovulate from diameters of 50 mm or greater. Draught and Friesian horses frequently ovulate from follicles > 50 mm. Larger than average diameters have been noted in Welsh cobs (J. Newcombe, unpublished observation), whereas Thoroughbreds only infrequently reach such diameters. In a study of over 700 ovulatory follicles in Standardbred mares, 41% of spontaneous ovulations were from follicles < 40 mm while 9.8% ovulated from those > 50 mm (J. Newcombe, unpublished observations). In the latter mares, a 35-mm follicle could be 5–7 days from ovulation and still be too immature to ‘respond’ to hCG. Treatment must therefore be delayed until the follicle has grown to 40 mm or even larger. Samper et al.,26 using 2500 IU
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intravenously, found that even by delaying treatment until the follicle was over 40 mm only 83% ovulated within 48 hours, with 8% taking between 72 and 96 hours. Sauberli et al.27 found that the mean time to ovulation was only 6 hours shorter when treatment was delayed until follicles were 40–44 mm than when treatment was carried out at 35–39 mm. The most erratic responses are experienced during the spring transition when it is well known that follicles can grow to the normal pre-ovulatory diameter or even larger before regressing.28 It is doubtful that such steroidally incompetent follicles can be induced to ovulate with hCG. However, Bour and colleagues29 reported success giving multiple 200-IU doses in the transitional period. The most rapidly growing follicles at this stage, especially when accompanied by a strong endometrial edema pattern, are the ones most likely to ovulate and therefore to ‘respond’ to hCG treatment. Carnevale et al.30 induced ovulation in 17 out of 19 > 40-mm transitional follicles with 3300 units given intravenously, whereas a control group took from 2 to 30 days to ovulate. Although mares can be induced to develop pre-ovulatory size follicles using various GnRH regimes, treatment with hCG can never guarantee ovulation. Mares treated with progesterone (by injection or intravaginally) or progestagen (orally) if started at the right stage of the spring transition will ovulate most reliably. However, if progesterone treatment is started too soon, although follicles of over 35 mm may develop, they may not respond to hCG treatment.31
Effect of hCG on corpus luteum development and progesterone concentrations One report32 showed that treatment with 1000 IU on days 3, 4, and 5 after estrus resulted in enhanced progesterone (P4) concentrations from day 7 to day 14. Paccamonti et al.33 found that 1000 IU on days 0–2 did not increase P4 until day 7. Carnevale et al.30 found no effect of hCG treatment before ovulation on CL formation or function. However, Michel et al.9 found that hCG-treated mares had an earlier rise in P4 concentration 72 hours after treatment (i.e., about 30 hours after ovulation) than GnRH- or salinetreated mares.
Interval from treatment to ovulation Most investigators have found that, when mares are examined daily following hCG treatment, ovulation has occurred by the time of the 48-hour examination. A small percentage of follicles are found very close to ovulation at this time, suggesting that in
some cases the ‘response’ interval is a little longer than 48 hours. Follicles destined to ovulate within 24 hours of treatment will of course have already ovulated by the 24-hour examination. Frequent examinations have determined the peak time of ovulation more closely. Evans et al.4 with 6-hourly examinations found 96% and 95% of mares treated with 2500 and 1500 IU, respectively, had ovulated within 48 hours, with means of 43.5 and 44 hours, respectively. Ginther34 reported that 65% of 55 mares given 2000 units intramuscularly and examined every 6 hours had ovulated in the period 36–42 hours. Newcombe,35 however, examining mares every 8 hours found that most mares ovulated between 34 and 45 hours when given 1500 IU subcutaneously.
Effect of mare’s age Erratic response to hCG seems to be more prevalent in older mares.36, 37 Whether this is due to interference from antibodies generated by previous hCG treatment or to delayed follicular development has not been established. McCue and colleagues38 described a clear relationship between age and delayed ovulation, whereas Green et al.39 found none.
Effect of dose and route Reports describe the successful use of hCG by either the intravenous or intramuscular route. This author has found the subcutaneous route to be equally successful. Dosage is empirical and successful reports vary from 1000 IU to 5000 IU. Most recent reports find 1500–2500 IU to be as effective as larger doses. Day40 found 80% of mares ovulated from 1500 IU in about 40 hours. There is little evidence that the larger doses act more successfully on slightly less mature follicles or that the interval from treatment to ovulation is any shorter. Grimmet and Perkins11 found no difference between 1500, 3000, and 6000 IU. Evans and colleagues4 studied LH levels after 500, 1500, and 2500 IU given intravenously and found no difference between 1500 and 2500. After 500 IU however, LH was slower to rise and reached only 10 ng/mL at 40 hours compared with 14 ng/mL with the other two dosages. The mean time to ovulation was similar to controls (83 hours) compared with 43.5 hours and 44 hours for 1500 and 2500 IU, respectively (6-hourly examinations). At ovulation (83 hours and 98 hours), levels of LH had reached 13 and 14 ng/mL in the 500-IU and control groups, respectively. J. Newcombe (unpublished observations) found an equal response rate to follicles of 40–50 mm with
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Human Chorionic Gonadotropin either 1250 IU or 2500 IU intravenously (82%) but the larger dose was more effective at diameters of less than 40 mm (76% vs 63%). In this study of Standardbred mares, most spontaneous ovulations were from diameters of 45–60 mm. In a field study, Thoroughbred mares were examined every 48 hours and the decision when to advise mating was based on various parameters acquired by clinical experience and by other management factors, but not necessarily on follicular diameter. Secondary follicles that regressed were omitted from the data, but data of the smaller follicle in sets of twins which ovulated up to 144 hours after treatment were included. Some 327 single follicles (mean diameter 39.7 mm, range 27–62 mm) and 294 multiple follicles (mean diameter for larger follicles 38.6 mm, range 27–50 mm; and for smaller 33.3 mm, range 21–45 mm) were treated with either 3000 IU intravenously or 1500 or 750 IU of Chorulon subcutaneously on the day of, or the day after, mating. Ovulation occurred within 48 hours in 93.3%, 96%, and 89.4% of the three treatment groups, respectively. Mean diameters at treatment of follicles which failed to ovulate within 48 hours were: single, 36.9 mm (range 28–45 mm); larger twin follicles, 39.6 mm (34–42 mm); and smaller twin follicles 29.8 mm (range 21–40 mm). The percentage of follicles which hemorrhaged without collapse was 0.3%, 1.4%, and 2.0% for the three groups (J. Newcombe, unpublished data). In another field trial, mares with single follicles were mated on the same basis and Chorulon given on a random basis, either 750 IU (n = 283) or 1500 IU subcutaneously (n = 437), or 3000 IU intravenously (n = 112). Response rate (ovulated within 48 hours) for mares treated at follicle diameters of 26–30 mm, 31–35 mm, 36–40 mm, and 41–45 mm for the three doses were 72% (750 IU), 85.7% (1500 IU), and 62.5% (3000 IU) for 26–30 mm; 90.1%, 89.7%, and 86.5% for 31–35 mm; 96.3%, 94.3%, and 95.3% for 36–40 mm; and 98%, 96.6%, and 100% for 41–45 mm. Other than the better response of 1500 IU on follicles of 26–30 mm, there was little difference between dose rates regardless of follicular diameter (J. Newcombe, unpublished data).
HCG in diestrus It has been clearly demonstrated in the cow that hCG is luteotrophic41–44 and induces accessory CL formation.45 The data sheet for Chorulon suggests that in cattle its use on day 12 both enhances the lifespan of the CL and increases P4 concentration. This use has not been investigated in mares. Repeated doses of hCG in pregnant mares before day 38 may cause luteolysis and pregnancy loss,46 but
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since there is little value in administering hCG at this time, this finding, although curious, appears unimportant. Aruda and Fleury47 reported an increase in plasma P4 after hCG on day 7, while Hendricks et al.48 found no effect on P4 concentration or on bloodflow in the uterine arteries when 3000 IU was given intravenously on day 7.
Synchronization of ovulations (reduction of spread between multiple ovulations) About half of all spontaneous multiple ovulations are not synchronous but occur over a period of more than 12 hours, often 2–3 days. hCG will induce follicles that may otherwise have ovulated more than 12 hours apart to ovulate synchronously (within a few hours). When the longevity of the sperm may have been too short for sufficient viable sperm to be still present in the fallopian tubes to fertilize a second oocyte, then only a single embryo can be expected. Synchronization of ovulation increases the chance that at least one oocyte, if not both, will be fertilized and therefore increases the chance of a pregnancy. Multiple embryos must of course be efficiently reduced to ensure a singleton pregnancy. However, a recent study21 found that, although hCG tended to increase the synchronicity, there was no increase in pregnancy or multiple pregnancy rate.
Antibody formation That repeated dosage with hCG causes antibody formation is no longer disputed. There is, however, much conflicting evidence of whether formation of antibodies affects the efficacy of hCG or even delays ovulation in untreated estrous periods. Sullivan and colleagues12 reported that, by the third treated estrus, ovulation was delayed by 1.2 days (based on 15 treated mares). Duchamp and colleagues49 found that none of 10 immunized mares ovulated within 72 hours. Roser et al.50 treated mares at each cycle from March to October. Five out of 12 mares developed significant antibody levels after two to five injections, and the half-life of antibodies ranged from 1 to several months. However, their production did not cause ovulatory refractiveness owing to absence of cross-reaction with endogenous LH. Evans et al.4 found no effect on LH concentrations after three hCG treatments. Nor did Blanchard et al.51 find any loss of efficacy in mares treated up to four times in the same season. Wilson and colleagues52 found antibodies, but no effect on time to ovulation until the fifth treatment. However, in the next year, the same mares had mean time to ovulation intervals from 53 hours at the sixth
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treatment (including five in the previous year) to 62 hours at the seventh treatment, and to 88 hours in three mares that received 10 treatments. Studies of antibodies, however, showed a very diverse mare response and high antibody levels did not correlate with longer time to ovulation. Green and colleagues39 found that 80% of 580 mares ovulated within 48 hours up to two treatments, but only 58% (n = 109) after the third. However, 67% (n = 18) ovulated after six to eight treatments. Loy and Hughes8 did not notice any loss of efficiency following repeated doses. In a retrospective study of the records of Standardbred mares over seven seasons,53 32 individual mares were found which had been given 2500 IU intravenously on 5–16 occasions, with a mean number of treatments of 8.3 times over 3.9 seasons. Mares were examined on Mondays, Wednesdays, and Fridays so that a ‘response’ to hCG treatment was found after either 48 or 72 hours; 80.2% of follicles (mean diameter 42.1 mm) had ovulated within 48 or 72 hours. When the data were divided into mares receiving either 5–8 (n = 16) treatments or 9–16 treatments (n = 16) the 48/72-hour response rate was 83.5% and 83.2%, respectively, after follicles which regressed or hemorrhaged had been excluded. The mean pre-ovulatory follicular diameter in the group treated 9–16 times was 41.6 mm, only 0.9 mm smaller than the 5–8 treatments group, and the percentage of follicles which took more than 48/72 hours to ovulate was also less, 10.2% compared with 12.7%. One mare treated 16 times over seven seasons had a 100% response rate from follicles of 39.2 mm mean diameter. McCue and colleagues,38 in a retrospective analysis of 1321 cycles treated with 2500 IU intravenously, thought that there was clear evidence of reduced efficacy even by the time of the second treatment. When follicles which did not ovulate were excluded from the data, they found that the number of mares ovulating within the 24- to 48-hour period fell from 71% (n = 839) after the first treatment to 55% by the third treatment (106) in the same season. However, at the same time, the number of mares ovulating in less than 24 hours increased from 17% to 28%. Therefore, the proportion which ovulated within the first 48 hours fell only from 88% to 83% owing mainly to a 4% increase in the numbers ovulating within the 48to 72-hour period. Although the number of ovulations within 48– 72 hours increased from 7% to 20% by the fifth treatment (n = 15), those mares taking at least five cycles to conceive were more likely to be the older ‘problem’ mares which were less likely to respond reliably.23 These data clearly showed an inverse age–response relationship with 8%, 14%, 18%, 21%, and 26% of mares aged < 6, 6–10, 11–15, 16–20, and > 20 years, respectively, failing to ovulate within 48 hours. The
percentage of mares ovulating within 0–72 hours actually increased from 92% to 93% by the fifth cycle. A similar explanation could account for the results of Green and colleagues,39 who started with 357 mares and were down to 43 by the fourth treatment; surely, 43 older and ‘problem’ mares would not be expected to respond as reliably as the 314. A further study by the author involved mares examined every 8 hours so that the time of ovulation was known to within ± 4 hours.54 In that study, 23 mares were treated four times in the same season at mean follicle diameters of 39.6 ± 1.1 mm. The interval to ovulation did not vary significantly with the number of treatments, namely 45.1, 44.1, 43.1, and 47.8 hours for the first to fourth treatment respectively. The response in mares given repeated doses over several seasons was analyzed. There was no effect of hCG dose number given successively in the same season (P = 0.24) nor an accumulative effect of season (P = 0.89), but a significant effect of follicle diameter at the time of treatment (P < 0.001) and of mare (P = 0.002). While there is still some debate on this question, practitioners will continue to make their own choice of ovulatory agent and dose based on experience.
References 1. McDonald LE. Hormones of the pituitary gland. In: Booth NH, McDonald LE Veterinary Pharmacology and Therapeutics, 6th edn. Ames: Iowa State University Press, 1988; p. 590. 2. Stewart F, Allen WR. Comparative aspects of the evolution and function of the chorionic gonadotrophins. Reprod Dom Anim 1995;30:231–9. 3. Patton PE, Stouffer RL. Current understanding of the corpus luteum of women and non-human primates. Clin Obstet Gynecol 1991;34:127–43. 4. Evans MJ, Gastal EL, Silva LA, Gastal MO, Kitson NE, Alexander SL, Irvine CHG. Plasma LH concentrations after administration of human chorionic gonadotrophin to oestrous mares. Anim Reprod Sci 2006;94:191–4. 5. Gastal MO, Gastal EL, Ginther OJ. Effects of hCG on characteristics of the wall of the developing preovulatory follicle evaluated by B-mode and colour-Doppler ultrasonography and interrelationships with systemic oestradiol concentrations in mares. Anim Reprod Sci 2006;94:195–8. 6. Cuervo-Arango J, Newcombe JR. Repeatability of preovulatory follicle diameter and uterine oedema pattern in two consecutive cycles in the mare and how they are influenced by induction treatments. Theriogenology 2008;69:681–687. 7. Day FT. Ovulation and descent of the ovum in the fallopian tube of the mare after treatment with gonadotrophic hormones. J Agric Sci Camb 1939;29:459–69. 8. Loy RG, Hughes JP. The effects of human chorionic gonadotrophin on ovulation, length of oestrus and fertility in the mare. Cornell Vet 1966;56:41–50. 9. Michel TH, Rossdale PD, Cash RSG. Efficacy of human chorionic gonadotrophin and gonadotrophin releasing hormone for hastening ovulation in Thoroughbred mares. Equine Vet J 1986;18:438–42.
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Human Chorionic Gonadotropin 10. Kloppe LH, Varner DD, Elmore RG, Bretslaff KN, Shull JW. Effect of insemination timing on the fertilising capacity of frozen/thawed equine spermatozoa. Theriogenology 1988;29:429–39. 11. Grimmet JB, Perkins NR. Human chorionic gonadotrophin (hCG): the effect of dose on ovulation and pregnancy rate in Thoroughbred mares experiencing their first ovulation of the breeding season. N Z Vet J 2001;49:88–93. 12. Sullivan JJ, Parker WG, Larson LL. Duration of oestrus and ovulation time in nonlactating mares given human chorionic gonadotrophin during three successive oestrous periods. J Am Vet Assoc 1973;162:895–8. 13. Voss JL, Pickett BW, Burwash LD, Daniels WH. Effect of human chorionic gonadotrophin on duration of oestrous cycle and fertility of normally cycling, nonlactating mares. J Am Vet Assoc 1974;165:704–6. 14. Woods J, Bergfelt DR, Ginther OJ. Effects of time insemination relative to ovulation on pregnancy rate and embryonic loss rate in mares. Equine Vet J 1990;22:410–15. 15. Hyland JH, Bristol F. Synchronisation of oestrus and timed insemination of mares. J Reprod Fertil Suppl 1979;27: 251–5. 16. Squires EL, Stevens WB, McGlothlin DE, Pickett BW. Effect of an oral progestin on the oestrous cycle and fertility of mares. J Anim Sci 1979;49:729–35. 17. Holtan DW, Douglas RH, Ginther OJ. Oestrus, ovulation and conception following synchronisation with progesterone, prostaglandin F2a and human chorionic gonadotrophin in pony mares. J Anim Sci 1977;44:431–7. 18. Shilova AV, Platov EM, Lebedev SG. The use of human chorionic gonadotrophin for ovulation date regulation in mares. In: Proceedings of the International Congress on Animal Reproduction and Artificial Insemination, 1976, p. 331. 19. Palmer E, Jousset B. Synchronisation of oestrus in mares with prostaglandin analogue and hCG. J Reprod Fertil Suppl 1975;23:269–74. 20. Squires EL, Wallace RA, Voss JL, Pickett BW, Shideler RK. The effectiveness of PGF2a, hCG and GnRH for appointment breeding of mares. J Equine Vet Sci 1981;1:5–9. 21. Davies Morel MC, Newcombe JR. The efficacy of different hCG dose rates and the effect of hCG treatment on ovarian activity: Ovulation, multiple ovulation, pregnancy, multiple pregnancy, synchrony of multiple ovulation in the mare. Anim Reprod Sci 2008;109:89–99. 22. Brugmans AC. Investigation on the efficiency of hCG, deslorelin, luprostinol and dinoprost on the termination of ovulation in mares and on pregnancy rates following time interval defined artificial insemination after induction of ovulation with hCG and deslorelin. D. Vet. Med. dissertation, Hannover University, 1997. 23. Barbacini S, Zavaglia G, Gulden P, Marchi V, Necchi D. Retrospective study on the efficiency of hCG in an equine artificial insemination programme using frozen semen. Equine Vet Educ 2000;12:312–17. 24. McKinnon AO, Nobelius AM, Figueroa STD, Skidmore J, Vasey JR, Trigg TE. Predictable ovulation in mares treated with an implant of the GNRH analogue Deslorelin. Equine Vet J 1993;25:321–3. 25. McKinnon AO, Perriam WJ, Lescun TB, Walker J, Vasey JR. Effect of a GNRH analogue (Ovuplant), hCG and dexamethasone on the time to ovulation in cycling mares. World Equine Vet Rev 1997;2(3):16–18. 26. Samper JC, Jensen S, Sergeant J, Estrada A. Timing of induction of ovulation in mares treated with Ovuplant or Chorulon. J Equine Vet Sci 2002;22:320–3.
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27. Sauberli DS, Lyman JT, Lock TF. Comparison of hCG and deslorelin use on two commercial horse farms: a retrospective study. 2001. 28. Webel SK, Franklin V, Harland B, Dzuik PJ. Fertility, ovulation and maturation of eggs of mares injected with hCG. J Reprod Fertil 1977;51:337. 29. Bour B, Palmer E, Driancourt M-C. Stimulation of ovarian activity in the pony mare during winter anoestrus. Endocrine causes of seasonal and lactational anoestrus in farm animals. A seminar in the CEC Programme of Co-ordination of Research on Livestock Productivity and Management, Institut fur Tierzuchtund Tierverhalten, Mariensee, Bundes forschung san stalt fur Landwirt, 1985, p. 85. 30. Carnevale EM, Squires EL, McKinnon AO, Harrison LA. Effect of human chorionic gonadotrophin on time to ovulation and luteal function in transitional mares. J Equine Vet Sci 1989;9:27–9. 31. Newcombe JR. Field observations on the use of a progesterone releasing intravaginal device to induce oestrus and ovulation in seasonally anoestrous mares. J Equine Vet Sci 2002;22:378–82. 32. Kelly CM, Hoyer PB, Wise ME. In-vitro and in-vivo responsiveness of the corpus luteum of the mare to gonadotrophin stimulation. J Reprod Fertil 1988;84:593–600. 33. Paccamonti DL, Rodriguez HF, Myers MW, Eilits BE, Godke RA. Effects of PGF2a or hCG administration during the early luteal phase on progesterone secretion in the mare. In: Proceedings of 37th American Association of Equine Practitioners, 1991, pp. 151–9. 34. Ginther OJ. Reproductive Biology of the Mare, 2nd edn. Cross Plains, WI: Equiservices, 1992; pp. 279–80. 35. Newcombe JR. Time to ovulation following 1500 units of hCG given subcutaneously. Unpublished. 36. Barbacini S, Necci D, Postinger G, Parmeggiani F, Squires E. What to expect when barren mares are inseminated with frozen-thawed semen. In: Proceedings of the 54th Annual Convention of Equine Practitioners, 2008, pp. 274– 5. 37. Carnevale EM, Coutinho da Silva MA, Panzani D, Stokes JE, Squires EL. Factors affecting the success of oocyte transfer in a clinical program for subfertile mares Theriogenology 2005;64:519–27. 38. McCue PM, Hudson JJ, Bruemmer JE, Squires EL. Efficacy of hCG at inducing ovulation: a new look at an old issue. Proc AAEP 50:510–513, 2004. 39. Green JM, Raz T, Epp T, Carley SD, Card CE. Relationships between utero-ovarian parameters and ovulatory response to human chorionic gonadotrophin (hCG) in mares. In: Proceedings of the 53rd Annual Convention of Equine Practitioners, 2007, pp. 563–7. 40. Day FT. The veterinarians approach to breeding problems in the mare. Vet Rec 1957;69:1258–67. 41. Wiltbank JN, Rothlisberger JA, Zimmerman DR. Effect of human chorionic gonadotrophin on maintenance of the corpus luteum and embryonic survival in the cow. J Anim Sci 1961;20:827–9. 42. Christie WB, Newcomb R, Rowson LEA. Embryo survival in heifers after transfer of an egg to the uterine horn contralateral to the corpus luteum and the effect of treatments with progesterone or hCG on pregnancy rates. J Reprod Fertil 1979;56:701–6. 43. Walton JS, Herbert GW, Robinson NA, Leslie KE. Effects of progesterone and human chorionic gonadotrophin administration, five days after insemination on plasma and
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milk concentrations of progesterone and pregnancy rates of normal and repeat breeder cows. Can J Vet Res 1990;54:305– 8. Hansel W, Blair RM. Bovine corpus luteum: a historic overview and implications for future research. Theriogenology 1996;45:1267–96. Escalera-Martinez JF, Rangel-Santos Rodriguez-De Lara R, Garcia-Muniz JG, Sanchez-Del Real C. Effect of human chorionic gonadotrophin (hCG) administration on ovarian activity and fertility of Holstein cows. Theriogenology 2006;66:676. Allen WE. Pregnancy failure induced by human chorionic gonadotrophin in pony mares. Vet Rec 1975;94:505. Arruda RP, Fleury PDC. Pregnancy rates and plasma progesterone concentrations in embryo recipient mares receiving hormone treatment. In: Proceedings of the 14th Havemeyer Foundation Monograph Series, 2004, pp. 95– 6. Hendricks WK, Colenbrander B, Stout TAE. Effects of administering PGF2a or hCG on day 7 after ovulation on ovar-
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ian, uterine and luteal blood flow in the mare. Anim Reprod Sci 2006;94:223–5. Duchamp G, Bour B, Combarnous Y, Palmer E. Alternative solutions to hCG induction of ovulation in the mare. J Reprod Fertil Suppl 1987;35:221–8. Roser JF, Kiefer EL, Evans JW, Neely DP, Pacheco CA. The development of antibodies to human chorionic gonadotrophin following its repeated injection in the cyclic mare. J Reprod Fertil Suppl 1979;27:173–9. Blanchard TL, Varner DD, Schumacher J. Manual of Equine Reproduction, 2nd edn. St. Louis: Mosby, 2003. Wilson CG, Downie CR, Hughes JP, Roser JF. Effects of repeated hCG injections on reproductive efficiency in mares. J Equine Vet Sci 1990;10:301–8. Newcombe JR, Wilson MC. The effect of repeated treatment with human chorionic gonadotrophin to induce ovulation in mares. Proceedings of the 46th Congress British Equine Veterinary Association, 2007, p. 291. Newcombe JR, Cuervo-Aranga J. A retrospective study of the use of hCG in a fertility clinic. 2007, Unpublished.
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Chapter 189
Progestagens and Progesterone Carlos R.F. Pinto
Introduction The term ‘progestagens’ refers to a group of naturally occurring or synthetically derived steroids with 21 carbons (C21) that, just like progesterone, bind to progesterone receptors and, therefore, exert progestational effects. Other terminology such as progestins, progestagens, gestagens, and gestogens have all been used when referring collectively to progestational compounds. Progesterone is the steroid hormone produced by the corpus luteum responsible for suppressing estrous behavior. In equids, luteal progesterone is the main hormone responsible for maintaining pregnancy during the first trimester. For the remainder of the pregnancy, placental production of progestagens – primarily 5α- pregnanes – become the primary source of progestational support of the pregnancy. Administration of progestational compounds in the cycling mare suppresses estrous behavior by interfering with ovarian follicular development and release of gonadotropin. The availability of progestational compounds (natural and synthetic) presents an excellent approach to control the mare’s estrous cycle and to induce pregnancy-like changes in the endometrium to support pregnancies in non-cycling mares (without a functional corpus luteum), or in those with signs of or at risk for luteal insufficiency. Progestational compounds differ in their ability to bind to progesterone receptors (PRs). Receptors for progesterone belong to the steroid receptor superfamily and act as a ligand-dependent transcription factor. There are two PR isoforms, A and B, which possess distinct pharmacological and biological significance. The PR-A isoform acts as a transcriptional inhibitor and modulator for other steroid receptors including PR-B, whereas typical progestational effects of progesterone and derivatives occur
after binding to PR-B.1 Although the expression of progesterone receptors A and B has been demonstrated in many vertebrates including primates and rodents, to date no information on the expression of progesterone receptor isoforms is available for horses. If they exist in horses, it remains unknown what effects the administration of exogenous progestational compounds would have on the expression of progesterone receptors A and B. This discussion will focus on the applications of progestagens and natural progesterone for the non-pregnant and pregnant mare, but first we will describe the different types of progestational compounds currently available for the equine practitioner. In this chapter, the terminology progestagens will be used to indicate synthetic progestational compounds, whereas the pleonasm ‘natural progesterone’ will be used to denote the difference.
Progestagens Altrenogest Altrenogest is a synthetic progestagen and the only approved progestational drug for horses in many countries. Chemically, altrenogest is a 17α-allyl-17β-hydroxy-estra-4,9,11-trien-3-one compound called allyltrenbolone. This is a derivative of the androgenic–anabolic steroid (AAS) C19-steroid nortestosterone (also known as nandrolone) differing only by two double bonds in positions 9 and 11 and an allyl group at the carbon 17. Altrenogest is also structurally related to the AAS trenbolone for having a 3-keto-4,9,11-triene steroid nucleus. Whereas minor potential androgenic–anabolic effects of altrenogest have been suggested for pigs,2 no significant effects
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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were found in studies that investigated its effects on the growth of young stallions and on the body mass and body condition score of mature mares when both groups were treated with altrenogest for 8 weeks.3 Altrenogest treatment does lower concentrations of plasma testosterone and decrease overall sexual functions (behavior and semen production), especially in young, growing animals.2, 4, 5 Although the potential anabolic effects of altrenogest have not been critically investigated, it appears unlikely that mares treated according to the manufacturer’s recommendations would experience any anabolic–androgenic effect. Treatment of altrenogest during pregnancy had no effect on hormonal profiles and fertility of fillies.6 However, foals born from mares treated with altrenogest during late pregnancy until term had lower respiratory rates than control foals and overall tended to take longer in adapting to the initial hours of life.7 In that study, altrenogest-treated mares tended to have shorter gestation length than control mares and had the second stage of parturition increased. Administration of altrenogest to lactating mares did not interfere with the mare’s fertility, milk production, and foal growth.8 Altrenogest is also the only known available progestagen bioavailable after oral administration in horses, and recent pharmacokinetics analyses obtained from 10 geldings receiving 44 µg/mL once daily for 5 days have been reported.9 Maximum concentrations (Cmax ) of altrenogest were reached at 15–180 minutes after administration and mean concentration of Cmax values was 35 ng/mL. At day 5, Cmax ranged between 24 and 52 ng/mL with a mean of 31 ng/mL; baseline concentrations were achieved within 72 hours after the last administration. For the 10 animals, the mean plasma clearance of altrenogest was 356 mL/h/kg at day 1 and 264 mL/h/kg at day 5. The plasma elimination half-lives were 2.6 and 3.7 hours at day 1 and day 5, respectively. The effects of the administration of altrenogest on the mare’s estrous cycle were first reported in 1975 by Webel.10 Following that report, research and clinical observations in France, the USA, and the UK corroborated the efficacy of altrenogest in suppressing estrus and its ability to regulate estrus synchronization/induction in cycling and transitional mares. AlR trenogest (Regu-Mate equine solution) is available in an oil oral solution that should be given once daily directly in the mouth or mixed in the grain ration. Fertility of mares treated with oral altrenogest has been reported to be approximately 80% after two cycles,11 indicating that treating a mare with altrenogest should not affect its inherent fertility. In a recent study by Storer et al.,12 long-acting formulations of injectable altrenogest were pre-
pared and administered to cycling mares immediately after being treated with luteolytic doses of prostaglandin (PG) F2α mares. Mares were treated with injectable solutions of altrenogest containing 225 mg (10-day formulation), 450 mg (15-day formulation), and 500 mg (30-day formulation) of altrenogest as lactide–glycolide microparticles and the effects on prolongation of the estrous cycle and interovulatory intervals were compared with control mares and mares receiving 1000 mg of medroxyprogesterone acetate. All single injection altrenogest formulations were significantly effective at extending the return to estrus and interovulatory interval in mares when compared with controls or mares treated with medroxyprogesterone acetate. The 30-day altrenogest formulation had the longest effect at suppressing estrus and inhibiting ovulation. The mean return to estrus was 32.8 ± 4.8 days compared with 3.9 ± 0.5 days in the control mares. The other two long-acting altrenogest formulations (10- and 15-day formulations) also effectively extended the return to estrus following treatments for 12.7 ± 0.4 days and 15.8 ± 1.5 days, respectively. In that study, medroxyprogesterone acetate treatment was not effective at delaying estrus or ovulation, thus corroborating previous studies that found no other progestagen other than altrenogest to have meaningful effect on the mare’s estrous cycle. Based on that study, long-acting (12 and 30 days) injectable formulation of altrenogest (BioRelease Altrenogest LA 150 mg/mL, 1.5 mL/intramuscular dose) produced by a compounding pharmacy in the USA (BET Reproductive Laboratories, Lexington, KY) can be ordered by a veterinary prescription and provides an interesting alternative to daily treatments, which are typically needed when an oral solution of altrenogest is used. Other synthetic progestagens Several other progestagens have been studied or used to control the mare’s reproductive cycle or behavior (Table 189.1); however, no established efficacy has been scientifically demonstrated for these compounds. One report considered successful the use of proligestone, a long-acting injectable progestagen, as 68% of 47 transitional mares treated with 1500 mg of proligestone as a bolus intramuscular dose ovulated within 24 days and 67% of 39 mares became pregnant after the first ovulation; however, the study regretfully did not have adequate control mares for comparison.13 The authors further acknowledged that, despite the interesting findings reported, the absence of control animals, both those untreated and those dosed with other progestagens, made the interpretation of the data difficult. More recently, the efficacy
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Progestagens and Progesterone
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Table 189.1 Progestational compounds. Note that, with the exception of natural progesterone and altrenogest, all other progestational compounds have not conclusively been shown to effectively affect reproductive phenomena in the mare Progestational compound
Efficacy
Route of administration
Natural progesterone Altrenogest
High High
Proligestone Melengestrol acetate Chlormadinome acetate Norgestomet Hydroxyprogesterone caproate or hexanoate Medroxyprogesterone acetate
Low (possibly dose dependent) Low (possibly dose dependent) None None None None
Injectable or intravaginal Oral Injectable (long-acting) Injectable Oral Oral Implant Injectable Injectable (long-acting)
of melengestrol acetate (MGA) to shorten the vernal transition of mares by synchronizing and accelerating the first ovulation of the year was investigated.14 Mares were exposed to artificial lights during the winter for 60 days before 16 mares in late transition were fed 150 mg/mare/day (group I) or 100 mg/mare/day (group II) of MGA for 10 days. A luteolytic dose of prostaglandin was administered to each mare 1 day after the end of MGA treatment. The presence and duration of estrus, follicular growth, uterine edema, and presence of ovulation were monitored by ultrasonography and the cervical tone was evaluated by rectal palpation. Ovulation rate was 87.5% and 62.5% for mares in groups I and II, respectively, which was significantly greater than the rate for control mares (20%). The mean ovulation interval for mares receiving the higher doses was significantly shorter (13.1 days) than mares receiving the lower dose of MGA (25.6 days). The authors concluded that MGA can be used for synchronizing and hastening the first ovulation of the year in mares. In a previous report by Loy and Swan,15 oral administration of 10 or 20 mg of MGA to cycling and non-cycling mares produced no significant effects in controlling or affecting estrous behavior and ovulation. In a review of therapy with progestational compounds by Neely,16 it was mentioned that equine practitioners working with race or performance horses would advocate the use of 60–200 mg of MGA per day to suppress signs of behavioral estrus. Thus, it is possible that the use of significantly high doses of some progestagens may explain the anecdotal evidence favoring the use of progestagens such as MGA. In ovariectomized mares, whereas daily administration of 0.044 mg/kg altrenogest was found to support pregnancy,17 administration of 1000 mg of hydroxyprogesterone caproate did not maintain pregnancy.18 In another study designed to investigate the ability of synthetic progestagens to maintain pregnancy in the mare, five groups of five mares were treated with medroxyprogesterone acetate, hydrox-
yprogesterone hexanoate, norgestomet, megesterol acetate, or altrenogest.19 After being treated for 2 days, all mares received a luteolytic dose of PGF2α at 18 days of gestation. All but the mares treated with altrenogest experienced embryo losses within 8 days following the administration of PGF2α on day 18 after ovulation, thus providing clinical evidence that altrenogest was the only progestagen tested that could maintain pregnancy in the absence of a progesterone-secreting corpus luteum. It has been suggested that the inability of these progestagens to suppress estrus and its associated behavior may be related to inadequate binding of these progestagens to the equine progesterone receptor. Norgestomet was found to have poor binding activity to equine hypothalamic, hypophyseal, and uterine tissues compared with that of progesterone and altrenogest.20 In a more detailed report in a thesis by Nobelius21 at the Monash University in Australia, 13 progestagens were tested and the binding affinities for only seven progestagens were documented (including the wellknown altrenogest, progesterone, and proligestone) while the remaining six progestagens did not show any affinity detectable at the range of the assay employed in that study. Among the six progestagens that did not show affinity for the progesterone receptor were norgestomet, hydroxyprogesterone caproate, and medroxyprogesterone acetate. Despite the lack of scientific evidence attesting the efficacy of the aforementioned progestagens in controlling the mare’s ovarian function and behavior, many horse owners and equine practitioners still use them as a last resort in an attempt to correct undesirable behavior especially because some of these progestagens are available as implants or depot preparations. The development of a technology to compound long-acting formulations of an injectable altrenogest solution may prove of value to horse owners wishing to reduce the labor associated with daily treatments of oral altrenogest or injectable natural progesterone.
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Progesterone
Intravaginal devices to deliver progesterone
Injectable progesterone – natural progesterone prepared as an injectable oil solution – has been extensively used to control the mare’s estrous cycle and to promote pregnancy-like changes in the uterus. Natural progesterone obviously offers excellent results as it has, in contrast with many other synthetic progestagens, high affinity for its own receptors. As in many countries there are no commercially available preparations of natural progesterone approved for use in horses, natural progesterone is typically prepared by compounding pharmacies. For many years, the main disadvantage was the need for daily injections that would invariably cause local swelling and inflammation. Most mares resent daily injections and long-term treatments are difficult, in addition to the risk of prolonged swellings at the injection sites leading to formation of seromas, abscesses, and fibrosis. The need for daily injections of progesterone to achieve therapeutic blood levels was shown by Ganjam et al.22 when studying the biological halflife of progesterone in ovariectomized mares. The authors reported the half-life of progesterone to be very short (2.5 minutes for the first component of the disappearance rate in plasma, and 20 minutes for the second component); for mares treated with daily doses of 150 mg, concentrations of plasma progesterone would increase from 2.1 to a maximum of 5.9 ng/mL only between days 10 and 25 after treatment. When mares were treated with a daily dose of 300 mg of progesterone, concentrations of plasma progesterone rose more rapidly, from 2.0 to 7.0 between days 2 and 14. Therefore, based on these findings, the authors refuted the practice of treating pregnant mares only once every 2 or 4 weeks to prevent abortion. To circumvent this problem, several pharmacies in the USA now compound highly concentrated and long-acting preparations of progesterone. One of these pharmacies (BET Labs, Lexington, KY) has developed the preparation of a biocompatible controlled release progesterone-based product that delivers an adequate amount of progesterone to maintain serum concentrations of progesterone ≥ 2 ng/mL for periods of 7–10 days following one intramuscular injection. The controlled biorelease system used is a proprietary low-viscosity nonaqueous liquid system that uses an injectable solution designed to deliver the entire dose of progesterone at a high concentration (300 mg/mL) at a controlled rate for a period of 7–10 days. These longacting preparations of progesterone despite being non-aqueous, do not induce local inflammatory and tissue swelling at the injection site as often as traditional oil preparations of injectable progesterone do.
Intravaginal progesterone-releasing devices (PRIDTM , progesterone-releasing intravaginal device; R CIDR , controlled intravaginal drug releasing deR vice; Cue-Mate ) are commonly used in livestock and have also been successfully used in the mare, albeit with variable results. Some authors found that intravaginal progesterone-releasing devices accelerated the occurrence of estrus and first ovulation in anestrous and transitional mares23 and in cycling mares,24 whereas no significant effects of treatment were seen in anestrous mares nor an accurate synchronization of ovulation was documented in cycling mares.25, 26 Newcombe23 reported that 89% of anestrous/transitional mares treated with PRIDTM ovulated within 10 days after its removal. R is another intravaginal device develCue-Mate oped for cattle that has been adapted for use in R . This device contains mares; it is called Cue-Mare 1.72 g of progesterone impregnated into a silicone matrix (treatment ‘pods’) secured in a plastic carrier body shaped as a ‘wishbone.’ In the original report R , concentrations describing the use of Cue-Mare of plasma progesterone were above 1.0 ng/mL in all mares that had the device placed intravaginally for 10 days.24 Of 34 mares that were artificially inseminated following the device removal, 18 (52.9%) became pregnant. In a more recently study, Hanlon27 R reported that 85% of 151 mares treated Cue-Mare in New Zealand were in estrus within 5 days and 88% had ovulated within 7 days. The treated mares were pregnant (90%) 18 days earlier than control mares, resulting in a pregnancy rate of 84%. Taken together, these reports certainly indicate that intravaginal progesterone slow-releasing devices have the capability to deliver adequate amounts of progesterone to promote an artificial luteal stage in the mare followed by fertile ovulation. While the estrus and ovulation synchronization using these devices alone may not be optimal, the association of ovulation-inducing agents upon device removal may produce acceptable success rates for commercial use. These devices are not approved for use in R horses (except Cue-Mare in New Zealand) and there are many veterinarians and horse owners that oppose to their use because of the transient mild to moderate vaginitis induced by the intravaginal device. The local inflammation apparently occurs in all mares, but vaginal discharges are seen in only a small percentage of treated mares. Mares with overt signs of vaginitis do not show signs of distress or discomfort. Most authors have reported the vaginal inflammation completely resolves within 2 days and fertility of mares artificially inseminated or mated on the induced estrus has varied from 60%
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Progestagens and Progesterone to 90%, thus attesting the absence of significant and permanent deleterious effects on the reproductive tract. Future development of synthetic intravaginal devices specifically designed for use in mares may help to minimize the degree of vaginal inflammation and pacify concerns about these side-effects.
Indications for progestagen therapy in mares Transitional mares The transition period that mares experience from the winter anestrous, anovulatory season into the physiological cycling (breeding) season is characterized by frequent, erratic, estrual behavior without ovulation. More in-depth discussion of the hormonal events implicated in the nature of this phenomenon (vernal transition) in mares is discussed elsewhere in this book. Veterinarians are often asked to prescribe hormonal treatments to advance the onset of the breeding season because many horse breeders want to have their mares bred early in the year. While protocols utilizing progestagens to control the mare’s estrous cycles during the transition period and to advance the first ovulation have provided acceptable success rates, they do not produce straightforward results. Several studies have found that progestagens do not cause significant effects in anestrous mares (deep anestrus) but do have the ability to suppress or prevent the occurrence of long or erratic estrous periods common during the vernal transition period. Progestagen treatment has also been shown to advance the first ovulation of the year. For mares classified as being in transition, the best results following progestagen therapy have been documented when altrenogest was used in mares found in mid- to late transition, displaying signs of behavioral estrus for several days, and having one or multiple follicles > 20 mm in diameter. When these criteria are met, these studies have reported a significantly greater number of mares exhibiting estrus and ovulating than untreated control mares.28 There is controversy as to whether progestagen treatment of transitional mares hastens the first ovulation of the year or synchronizes the first ovulation. Recently, some promising results with the use of intravaginal progesteronereleasing devices have been reported.25, 29 For many countries, oral administration of altrenogest is the only approved therapy for use in transitional mares.
tion of progestational compounds can start at any stage of the estrous cycle and it should keep mares in a diestrus-like stage for the duration of treatment.11 In one study, mares were treated with altrenogest during estrus and early and mid-diestrus; estrus was successfully blocked in all treated groups. Mares in estrus had their signs of behavioral estrus blocked within 2–3 days. There are apparently no long-term deleterious effects on the mare’s general or reproductive health when treated with altrenogest for prolonged periods (months). In one study by Shideler et al.,17 mares were treated for 88 days with doses up to five times greater (0.22 mg/kg) than the recommended label dose (0.044 mg/kg) and showed no signs of toxicity nor effects on their general health condition. In a subsequent study,11 mares were treated with oral altrenogest daily for 15, 30, and 60 days so that the treatment ended in all groups on March 31. No effects on mare’s fertility (first-cycle pregnancy rate) were documented for all groups. Interestingly, mares treated the longest became pregnant sooner than control mares. Therefore, performance mares can be safely treated with altrenogest for prolonged periods and may return to their original reproductive function once treatment is discontinued. However, one should not treat mares that are susceptible to uterine infection.
Estrus synchronization Controlling the estrous cycle may prove beneficial to horse owners experiencing the following scenarios.
r
r
Cycling mares
The performance mare Suppression of estrual behavior may be desirable in mares that are competing or showing. Administra-
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Accommodating the stallion’s breeding schedule – a mare may be treated with progestational compounds to induce her to be in estrus when the stallion is available for breeding. For example, mating or semen collection may not be possible if stallions are experiencing poor reproductive performance or any other health problem, traveling for shows or competitions, or if the horse owner’s or manager’s schedules preclude them from collecting semen from the stallion. Maximizing the number of mares being artificially inseminated with one single ejaculate – an average single ejaculate from a fertile stallion can be divided into many potential breeding doses, depending on the stallion’s particular fertility and sperm output. Even stallions with poor sperm count may benefit from spaced, scheduled collections or matings to prevent further decrease in sperm count caused by excessive breeding activity. Projecting the foaling season – synchronized breeding may help the horse owner to concentrate the birth of foals during a specific time of the year
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when plans for assisted foaling or hiring additional help can be made. Estrus synchronization in artificial insemination and embryo transfer programs – two common approaches to synchronize estrus and ovulation in cycling mares are (i) to administer oral altrenogest (0.044 mg/kg bodyweight) for 10–14 days or (ii) to administer intramuscular injections of progesterone and estradiol (P&E) for 10 days. With either treatment, PGF2α is typically administered (5–10 mg of dinoprost intramuscularly) on the last day of treatment. Shorter treatments of altrenogest or P&E for 7 or 8 days have also been used with success.
The benefit of administering P&E was originally described by Loy and co-workers.30 The protocol involves the administrations of intramuscular injections of progesterone (150 mg) combined with estradiol-17β (10 mg) in oil for 10 consecutive days with a single injection of PGF2α administered on the 10th day of treatment. After the combined steroid treatment regimen is discontinued, administration of human chorionic gonadotropin (hCG) (2500 IU intravenously) when a 35-mm follicle is detected will further improve ovulation synchrony. The reason for adding estradiol to the P&E protocol relies on the fact that the presence of estradiol causes a stronger negative feedback on follicle-stimulating hormone (FSH) secretion and consequently on follicular development than progesterone alone. Once treatment is discontinued, it is expected that at day 8 post-treatment, the majority of the mares will have a preovulatory follicle greater than 30 mm in diameter. At that time, the decision to treat the mare with an ovulation-inducing agent may be made. Long-acting formulations of P&E have been anecdotally reported to not provide the same degree of synchrony that daily injection preparations do. Postpartum mares Although mares naturally come into estrus shortly after foaling (foal heat) and have the potential for normal fertility, many husbandry factors may contribute to a relatively lower fertility of mares mated or artificially inseminated during the foal heat than that of mares that are beyond 30 days postpartum or non-lactating mares. One strategy to circumvent the potential for decreased fertility associated with the foal heat has been to treat mares with progestagens beginning no later than the first day postpartum and continued for at least 8 days. The goal of this approach is to provide more time for the mare to involute her uterus, thus creating a more favorable uterine environment to withstand potential contam-
inations during mating and to establish a successful pregnancy. Daily injectable progesterone or oral altrenogest treatment have been shown to delay the first ovulation and improve fertility rates when compared with control mares. In one study, mares ovulating after 15 days postpartum had greater pregnancy rates than mares ovulating before day 15, 82% versus 50%, respectively.31 Altrenogest is given orally (0.044 mg/kg/day) for 8–15 days after foaling followed by PGF2α on the last day of treatment. Prostaglandin is given because progestagen therapy alone may not prevent ovulation even though estrus is suppressed. If ovulation has occurred during altrenogest treatment, the resulting luteal tissue will continue to produce progesterone despite withdrawal of the exogenous progestagen. Treatment with PGF2α ensures that progesterone concentration will decrease and permit the mare to return to estrus, usually within 2–7 days. A combination of P&E as described earlier also delays the onset of foal heat and ovulation. Recently, a study was designed to study the effects of treating postpartum mares with 300 mg of progesterone and 20 mg of estradiol-17β administered within 24 hours after foaling.32 Mares were examined daily by palpation per rectum and transrectal ultrasonography to determine the day of ovulation. No treated mare (0/20) ovulated before day 10 after postpartum, whereas six out of 30 control mares did; however, for mares bred on the first postpartum estrus pregnancy rates did not differ between treatment groups (16/18, 89%) and controls (22/30, 81%). Delay of foal heat with hormone therapy until after day 10 postpartum allows more complete uterine involution to occur, as evidenced by a greater proliferation of endometrial glands and increased endometrial gland density and ciliation of luminal epithelium.30, 33 Delaying foal heat with progestational compounds likely improves the uterine environment and endometrial secretions necessary to sustain conceptus development. Pregnant mares Horse owners often seek hormonal supplementation for mares with a history of undiagnosed abortion or even for those mares displaying infertility after a few cycles exposed to semen of good fertility. Although low concentrations of progesterone have been found in association with early embryonic loss in the mare, true primary luteal insufficiency seems to be an uncommon event. It is likely that, in most cases, luteal insufficiency (concentrations of progesterone < 2 ng/mL) is occurring in association with a uterine inflammatory process that releases PGF2α , and, consequently, luteolysis. It is a fact that luteal
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Progestagens and Progesterone progesterone is needed for maintenance of pregnancy during the first trimester as mares that were ovariectomized before day 70 of gestation aborted.34 Other studies have demonstrated the ability of altrenogest or progesterone to support pregnancy in the absence of a luteal source of progesterone (ovariectomized mares) or mares treated with luteolytic doses of PGF2α . Acyclic, non-ovulating mares have also been successfully used as embryo transfer recipients after being primed with estradiol and progesterone, and kept on progestagen treatment until formation of supplementary corpora lutea is observed (days 45–90 of pregnancy) or until the placenta begins to take over the maintenance of pregnancy at approximately 100–120 days of gestation.17,35–37 Taken together it seems that pregnant mares treated with progestagens may have their treatment discontinued once the placenta takes over the maintenance of pregnancy. In an original report by Hinrichs et al.38 on the efficacy of altrenogest to maintain pregnancy in ovariectomized mares, altrenogest at a dose of approximately 44 µg/kg was not as effective in maintaining pregnancy as a dose of approximately 88 µg/kg was. High-risk pregnant mares are typically treated with 88 µg/kg of altrenogest, which corresponds to twice the recommended dose by the manufacturer.39 High-risk pregnant mares are treated when signs of disease or threatened abortion are identified until near term. Whether beneficial effects of administering altrenogest to high-risk pregnant mares are related to its ability to directly affect myometrial contractions or indirectly by interfering with PG-induced myometrial contractions is not known. Despite the administration of altrenogest to mares in late gestation classified as high-risk pregnancies being common practice, this apparent success has been questioned by a recent study that reported treatment with altrenogest (88 µg/kg) until term did not prevent parturition.7 Furthermore, foals born from treated mares exhibited delayed time to stand up, nurse, etc. Based on these findings, the premise that the purported effectiveness of altrenogest to prevent abortion or to treat preterm foalings in mares with disturbed pregnancies should be questioned.
Other indications
Stress Any source of stress to pregnant mares has the potential to induce fetal losses. Until the source of stress is identified, controlled, or removed, exogenous administrations of progestational compounds are indicated.40 In one study by Baucus et al.,41 the effects of transporting mares during the third and fifth week of
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gestation on cortisol progesterone and ascorbic acid were investigated. No differences in progesterone concentrations were documented, whereas transported mares had significantly higher concentrations of cortisol. However, the incidence of early embryonic death (EED) did not increase when mares were transported for 472 km for approximately 9 hours. In that study, the incidence of EED was 13.3% in transported mares and 12.5% in non-transported mares. Mares may be at risk for maintaining pregnancy when experiencing gastrointestinal colic, laminitis, or septicemia. Endotoxemia associated with various clinical conditions could result in pregnancy wastage, and release of prostaglandins in association with inflammatory processes could lead to premature myometrial activity that causes abortion or premature labor. In the study by Daels et al.,42 pregnant mares were treated with endotoxin and oral altrenogest. Injection of endotoxin resulted in secretion of PGF2α , as determined by the plasma 15-keto-13,14-dihydroPGF2α concentrations, characterized by a biphasic pattern, with an initial peak at 30 minutes followed by a second, larger peak at 105 minutes after endotoxin injection. Concentrations of plasma progesterone decreased in all mares to less than 1 ng/mL within 24 hours after endotoxin injection. For mares that were treated with altrenogest until day 70, pregnancy was successfully maintained despite the presence of low concentrations of plasma progesterone. Taken together, because stressful conditions (e.g., transport, endotoxemia) and clinical diseases (gastrointestinal colic, navicular disease, babesiosis, etc.) can directly result in pregnancy losses,40 it is recommended to administer progestational compounds to mares experiencing clinical conditions deemed stressful or potentially capable of resulting in toxemia.
References 1. Giangrande PH, McDonnell DP. The A and B isoforms of the human progesterone receptor: two functionally different transcription factors encoded by a single gene. Recent Prog Horm Res 1999;54:291–313. 2. Kluber EF 3rd, Pollmann DS, Davis DL, Stevenson JS. Body growth and testicular characteristics of boars fed a synthetic progestogen, altrenogest. J Anim Sci 1985;61: 1441–7. 3. Hodgson D, Howe S, Jeffcott L, Reid S, Mellor D, Higgins A. Effect of prolonged use of altrenogest on behaviour in mares. Vet J 2005;169:322–5. 4. Squires EL, Badzinski SL, Amann RP, McCue PM, Nett TM. Effects of altrenogest on total scrotal width, seminal characteristics, concentrations of LH and testosterone and sexual behavior of stallions. Theriogenology 1997;48: 313–28.
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5. Johnson NN, Brady HA, Whisnant CS, LaCasha PA. Effects of oral altrenogest on sexual and aggressive behaviors and seminal parameters in young stallions. J Equine Vet Sci 1998;18:249–53. 6. Naden J, Squires EL, Nett TM. Effect of maternal treatment with altrenogest on age at puberty, hormone concentrations, pituitary response to exogenous GnRH, oestrous cycle characteristics and fertility of fillies. J Reprod Fertil 1990;88:185–95. 7. Neuhauser S, Palm F, Ambuehl F, Aurich C. Effects of altrenogest treatment of mares in late pregnancy on parturition and on neonatal viability of their foals. Exp Clin Endocrinol Diabetes 2008;116:423–8. 8. Sigler DH, Ericson DE, Gibbs PG, Kiracofe GH, Stevenson JS. Reproductive traits, lactation and foal growth in mares fed altrenogest. J Anim Sci 1989;67:1154–9. 9. Machnik M, Hegger I, Kietzmann M, Thevis M, Guddat S, Sch¨anzer W. Pharmacokinetics of altrenogest in horses. J Vet Pharmacol Ther 2007;30:86–90. 10. Webel SK. Estrus control in horses with a progestin. J Anim Sci 1975;41:385. 11. Squires EL, Heesemann CP, Webel SK, Shideler RK, Voss JL. Relationship of altrenogest to ovarian activity, hormone concentrations and fertility of mares. J Anim Sci 1983;56:901–10. 12. Storer WA, Thompson DL, Gilley RM, Burns PJ. Evaluation of injectable sustained release progestin formulations for suppression of estrus and ovulation in mares. J Equine Vet Sci 2009;29:33–6. 13. Van der Holst W, van Laar PH, Oldenkamp EP. Prolonged spring oestrus in mares: the use of progestogens with specific reference to proligestone. Theriogenology 1985: 24; 609–17. ´ 14. Lopez-Bayghen C, Zozaya H, Ocampo L, Brumbaugh GW, Sumano H. Melengestrol acetate as a tool for inducing early ovulation in transitional mares. Acta Vet Hung 2008;56:125–31. 15. Loy RG, Swan SM. Effects of exogenous progestogens on reproductive phenomena in mares. J Anim Sci 1966;25:821–6. 16. Neely DP. Progesterone/progestin therapy in the broodmare. In: Proceedings of the 34th Annual Convention of the American Association of Equine Practitioners, 1988, pp. 203–18. 17. Shideler RK, Squires EL, Voss JL, Eikenberry DJ, Pickett BW. Progestagen therapy of ovariectomized pregnant mares. J Reprod Fertil 1982;32:Suppl 459–64. 18. McKinnon AO, Tarrida del Marmol Figueroa S, Nobelius AM, Hyland JH, Vasey JR. Failure of hydroxyprogesterone caproate to maintain pregnancy in ovariectomised mares. Equine Vet J 1993;25:158–60. 19. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32:83–5. 20. Wiepz GJ, Squires EL, Chapman PL. Effects of norgestomet, altrenogest, and/or estradiol on follicular and hormonal characteristics of late transitional mares. Theriogenology 1988;30:181–93. 21. Nobelius, AM. Gestagens in the mare: an appraisal of the effects of gestagenic steroids on suppression of oestrus in mares. MSc thesis, Monash University, Australia, 1994. 22. Ganjam VK, Kenney RM, Flickinger G. Effect of exogenous progesterone on its endogenous levels: biological half-life
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of progesterone and lack of progesterone binding in mares. J Reprod Fertil Suppl 1975;23:183–8. Newcombe JR. Field observations on the use of a progesterone-releasing intravaginal device to induce estrus and ovulation in seasonally anestrous mares. J Equine Vet Sci 2002;22:378–82. Grimmett JB, Hanlon DW, Duirs GF, Jochle WA. A new intra-vaginal progesterone-releasing device (Cue-Mare) for controlling the estrous cycle in mares. Theriogenology 2002;58:585–7. ¨ Handler J, Schonlieb S, Hoppen HO, Aurich C. Seasonal effects on attempts to synchronize estrus and ovulation by intravaginal application of progesteronereleasing device (PRID) in mares. Theriogenology 2006;65: 1145–58. ¨ Handler J, Schonlieb S, Hoppen HO, Aurich C. Influence of reproductive stage at PRID insertion on synchronization of estrus and ovulation in mares. Anim Reprod Sci 2007;97:382–93. Hanlon DW. Treatment of transitional thoroughbred mares with an intravaginal progesterone-releasing device. In: Proceedings of the New Zealand Equine Veterinary Association Annual Conference, Palmerston North, New Zealand, 2007, pp. 41–4. Webel SK, Squires EL. Control of the estrous cycle in mares with altrenogest. J Reprod Fertil Suppl 1982;32:193–8. Norman ST, Larsen JE, Morton JM. Oestrous response and follicular development in mares after treatment with an intravaginal progesterone releasing device in association with single injections of oestradiol benzoate and PGF2alpha. Aust Vet J 2006;84:47–9. Loy RG, Evans, MJ, Pemstein R, Taylor TB. Effects of injected ovarian steroids on reproductive patterns and performance in post-partum mares. J Reprod Fertil Suppl 1982;32:199–204. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproductive tract of mares after parturition: effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed first postpartum ovulatory cycles. J Am Vet Med Assoc 1988;192:350–3. Bruemmer JE, Brady HA, Blanchard TL. Uterine involution, day and variance of first postpartum ovulation in mares treated with progesterone and estradiol-17beta for 1 or 2 days postpartum. Theriogenology 2002;57:989–95. Loy RG, Hughes JP, Richards WPC, Swan SM. Effects of progesterone on reproductive function in mares after parturition. J Reprod Fertil 1975 Suppl;23:291–5. Holtan DW, Squires EL, Lapin DR, Ginther OJ. Effect of ovariectomy on pregnancy in mares. J Reprod Fertil Suppl 1979;27:457–63. Squires EL, Ginther OJ. Collection technique and progesterone concentration of ovarian and uterine venous blood in mares. J Anim Sci 1975;40:275–81. Hinrichs K, Sertich PL, Palmer E, Kenney RM. Establishment and maintenance of pregnancy after embryo transfer in ovariectomized mares treated with progesterone. J Reprod Fertil 1987;80:395–401. Knowles JE, Squires EL, Shideler RK, Tarr SF, Nett TM. Progestins in mid- to late-pregnant mares. J Equine Vet Sci 1994;14:659–63. Hinrichs K, Sertich PL, Kenney RM. Use of altrenogest to prepare ovariectomized mares as embryo transfer recipients. Theriogenology 1986;26:455–60.
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Progestagens and Progesterone 39. Macpherson ML, Bailey CS. A clinical approach to managing the mare with placentitis. Theriogenology 2008;70: 435–40. 40. Van Niekerk CH, Morgenthal JC. Fetal loss and the effect of stress on plasma progestagen levels in pregnant Thoroughbred mares. J Reprod Fertil Suppl 1982;32: 453–7.
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41. Baucus KL, Ralston SL, Nockels CF, McKinnon AO, Squires EL. Effects of transportation on early embryonic death in mares. J Anim Sci 1990;68:345–51. 42. Daels PF, Stabenfeldt GH, Hughes JP, Odensvik K, Kindahl H. Evaluation of progesterone deficiency as a cause of fetal death in mares with experimentally induced endotoxemia. Am J Vet Res 1991;52:282–8.
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Chapter 190
Gonadotropin-Releasing Hormones Edward L. Squires
Out of all the reproductive hormones available for use in broodmare practice, gonadotropin-releasing hormone (GnRH) and its analogs are of extreme value. Since the chapter written by Irvine in the first edition of this textbook in 1993, there have been tremendous advancements in the area of clinical uses of GnRH. Irvine1 predicted that there would be an increased usage of analogs if they could be developed and given in a practical manner. He stated that the major role for GnRH is to stimulate the secretion of follicle-stimulating hormone (FSH) and luteinizing hormone (LH). Thus, any place where this would be of use is where GnRH could be used in a broodmare practice.
Early studies GnRH was first given to anestrous and estrous mares by Ginther and Wentworth,2 who reported plasma LH concentrations doubled after GnRH administration. In 1976, Evans and Irvine3 showed that GnRH induced FSH and LH release in anestrous mares. They were the first to devise a protocol for repeated injections of GnRH to induce follicular development and ovulation in anestrous mares.4 Johnson5 induced follicular development, ovulation, and corpus luteum formation in mares treated hourly with 2 or 20 µg of native GnRH. In a similar study, Turner and Irvine6 injected a group of mares with 25 µg three times daily for 28 days, and, by day 21, 11 of 12 mares had pre-ovulatory follicles. Allen et al.7 induced ovulation within 18 days in 15 of 20 anestrous mares with a single subcutaneous insertion of an implant, which released the GnRH analog ICI 118 630 at a rate of 30 µg/day for 28 days. Using implants that released the GnRH analog buserelin, Harrison et al.8 induced ovulation in 56%
of transitional mares in 28 days compared with 0% of controls. Turner and Irvine6 found that a release rate of 180 µg/day induced a high rate of follicular development and a 50–75% ovulation rate in anestrous mares.
Osmotic mini-pumps Ainsworth and Hyland9 investigated the use of osmotic mini-pumps to advance the onset of ovulation in Thoroughbred mares on an Australian stud farm. GnRH was delivered via subcutaneous osmotic implants at dose rates of 100 ng/kg/h and 200 ng/kg/h. They reported a continuous infusion of GnRH induced ovulation in a high percentage of anestrous mares. However, others have reported that pulsatile administration of GnRH may be advantageous over continuous infusion of GnRH.10 Becker and Johnson11 compared pulsatile with constant infusion of GnRH at 2 or 20 µg of GnRH per hour continuously or 20 µg of GnRH per hour in a pulsatile fashion. Mares administered pulsatile GnRH ovulated 2.8 days earlier than those receiving 20 µg of GnRH per hour continuously. Fitzgerald et al.12 evaluated the use of subcutaneous implants containing goserelin acetate for inducing ovulation in seasonal and anestrous mares. Mares were selected that had less than 20-mm follicles. These authors reported tremendous variation from year to year, but, generally, a high percentage of mares ovulated in response to osmotic mini-pumps containing goserelin acetate. More recently, two papers were published on the administration of a low dose of GnRH continuously using osmotic mini-pumps. The objective of the first study13 was to confirm the lack of pituitary desensitization to continuous infusion of low-dose GnRH. The osmotic mini-pumps were inserted after day 12
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Gonadotropin-Releasing Hormones of ovulation and released 5 µg of GnRH per hour for 14 days. Continuous administration of GnRH for 14 days in a cycling mare increased progesterone levels and tended to increase LH levels, but did not affect the interovulatory interval, indicating that there was no desensitization of the pituitary–hypothalamic axis. In the second study, the endocrine response of persistently anovulatory mares to continuous low-dose administration of GnRH was evaluated. Mares were selected that failed to develop 35-mm follicles or ovulate by April 1. They were then administered GnRH continuously at a rate of 2.5 µg/h for 14 days and then 5 µg/h for 14 days. GnRH increased LH in all mares, and all treated mares ovulated compared with none of the controls. In a subsequent paper from this group,14 they questioned whether continuous subcutaneous treatment with low-dose GnRH to mares from late September/early October to March would prevent anovulation. Osmotic pumps were inserted to deliver 2.5 µg/h for the first 60 days then 5 µg/h thereafter. Continuous infusion of low-dose GnRH beginning soon after the fall equinox and continuing until March failed to prevent the occurrence of anovulation or to hasten ovulation. Irvine1 stated, in the first edition of this textbook, that, of the treatments available, the use of longacting implants provides reasonable efficacy with ease of treatment and minimal animal handling. He further stated that it seemed highly likely that a suitable dose of analog will be developed that could offer a useful alternative to human chorionic gonadotropin (hCG). He certainly demonstrated some foresight in this statement because currently injections of the GnRH analog deslorelin are used as a means of inducing ovulation in transitional and cycling mares.
Ovuplant The LH surge in the mare is known to encompass several days and very often occurs after the time of ovulation. Thus, in designing a single-dose protocol for administering GnRH an injection that would last several days and result in a prolonged LH surge around the time of ovulation was important. Thus, an implant was developed for induction of ovulation that contained 2.1 mg of deslorelin acetate (OvuplantTM , Peptide Technologies, Sydney, Australia). Initially, the implant was inserted under the skin in the area near the neck. Although the implant is no longer available in the USA, it is used quite often in Canada, Australia, and other countries where it is available. A summary of the studies regarding Ovuplant can be found in Jochle and Trigg.15 These authors sum-
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marized the efficacy and safety of Ovuplant from studies conducted in 1990–4. These studies contained results from 1294 mares administered Ovuplant in Australia, Canada, Germany, Sweden, Brazil, Argentina, and the USA. Various doses of deslorelin in the implant were evaluated for efficacy. It was determined that 2.1–2.2 mg was the most effective dose. This was based on a study carried out by McKinnon et al.,16 in which they evaluated doses of 1.3, 1.6, and 2.2 mg versus 3000 units of hCG. The number of mares ovulating in 48 hours was similar in the 2.2-mg group to that in the hCG-treated group. In the summary provided by Jochle and Trigg,15 the efficacy of Ovuplant for inducing ovulation within 48 hours was 88% compared with 37% for untreated controls. They also reported no adverse effect on pregnancy rates. Mumford et al.17 reported on the effect of repeated treatments with Ovuplant on consecutive cycles. They found no negative effect from repeated treatments with Ovuplant. Mumford et al.18 also evaluated mares given 0, 1×, 3×, or 5× implants of Ovuplant (1× = 2.2 mg). The interval to ovulation was shortened with Ovuplant treatment and the percentage of mares ovulating within 48 hours was 20%, 83%, 73%, and 86% for 0, 1×, 3×, and 5× treatments, respectively. As recorded by several people, the diameter of the largest follicle at ovulation was smaller in Ovuplanttreated mares than in controls. In addition, in the study by Mumford et al.18 the interovulatory interval was increased by the mares given the 5× implants, indicating that there was some downregulation with the higher doses used in this study. After a few years of Ovuplant use in broodmares within the USA, it was noticed that a small percentage of mares that did not become pregnant after natural mating or insemination had a longer interovulatory interval if given Ovuplant during estrus. Morehead and Blanchard19 in Kentucky and Johnson et al.20 at Louisiana State University (LSU) reported a longer interovulatory interval for mares given Ovuplant. In the LSU study, 12 mares were assigned to either an Ovuplant treatment or controls. On days 1, 4, 7, and 10 after ovulation, 50 µg of native GnRH was given as a challenge to the pituitary. The interovulatory interval was longer in treated mares than in controls. In addition, the concentration of LH and FSH in treated mares was lower than in controls on days 1–5 and on day 7. Both the LH and FSH response was suppressed in treated compared with control mares. These authors indicated a temporary downregulation of the pituitary gland. A series of studies was conducted at Colorado State University by Farquhar et al.21, 22 to demonstrate the reason for the longer interovulatory interval for
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the mares given Ovuplant. They demonstrated that FSH secretion was suppressed in early diestrus after Ovuplant administration and follicular development was also suppressed. In addition, their studies indicated that administration of prostaglandin to mares in mid-diestrus that had previously been given Ovuplant had an even longer interovulatory interval. Lastly, they went on to demonstrate that removal of Ovuplant implants within 1 or 2 days after ovulation prevented the depressed FSH secretion and the increased interovulatory interval.23 Thus, if Ovuplant is used in the USA and other countries, it is now recommended to place it in the lips of the vulva and remove it 1–2 days after ovulation, to prevent any possible increase in the interovulatory interval. McKinnon et al.24 evaluated the response of transitional mares to Ovuplant. In the first experiment, 21 early transitional mares were given an implant every other day until ovulation or a maximum of six implants. In a later experiment, 20 late transitional mares (≥ 30-mm follicles) were administered Ovuplant. In the early transition, six of 11 treated mares ovulated compared with zero of 10 controls. The average time to ovulation was 24 days, which unfortunately did not differ from the control group . In later transitional mares, the implants hastened ovulation in eight of 10 mares within 3 days with an average of 2.1 implants being given to transitional mares.
Injectable GnRH analogs Once Ovuplant was removed from the market in the USA, the need for an injectable GnRH was quite apparent. Therefore, several companies began to provide a compounded GnRH agonist, deslorelin, as an injectable form. One of the earliest forms of deslorelin included the BioRelease formulation from BET Laboratories, Lexington, KY. This particular slowrelease formulation allowed the deslorelin to be released over a couple of days. Stich et al.25 evaluated the short-term release deslorelin from BET Laboratories in foaling mares. Beginning 5–6 days post partum, 33 mares were assigned to either 1.5 mg of the BioRelease deslorelin or controls. Deslorelin increased the number of ovulations within 48 hours (75% versus 7% for controls). There was no effect of fertility and the number of covers per conception was decreased in deslorelintreated mares (1.6 versus 2.9). More recently, there have been an increased number of deslorelin injectable products made available on the market. Many of these are suspended in saline or sterile water and do not contain any slow-release mechanisms. It is somewhat surprising that deslorelin in an aqueous solution is able to produce ovu-
lation in mares. However, McCue et al.26 compared various formulations of deslorelin and reported all formulations tested in their study resulted in a shortening of the follicular phase, hastening of ovulation, and a similar response to hCG. However, these studies were carried out in the middle of the breeding season, and the results could have been different if given early in the breeding season. Although not examined critically, there does not appear to be any adverse effect on FSH secretion or follicular development in mares that do not become pregnant after being given injectable deslorelin. More than likely, since the injectable deslorelin products are cleared from circulation faster than deslorelin from Ovuplant, this results in less suppression on follicular development and FSH secretion. Injectable deslorelin is not only used in cycling mares but has also been used in studies to hasten the first ovulation of the year in transitional mares. Raz et al.27 evaluated the effect of injectable deslorelin on ovulation in transitional mares and compared this with administration of equine FSH. Transitional mares (20- to 25-mm follicles) were assigned to equine FSH (eFSH) injection twice daily or 63 µg of deslorelin acetate twice daily. Transitional mares were assigned to one of the two groups until 35-mm follicles were obtained. hCG was given to induce ovulation once the pre-ovulatory follicles were obtained. The mean number of days to ovulation was similar between the two groups – 8.2 and 7.2 days for eFSH and deslorelin, respectively. However, the number of ovulations was greater in the eFSHtreated mares (3.4 versus 1.1 for deslorelin-treated mares). In addition, the number of embryos obtained 8 days after ovulation was 2.6 for eFSH-treated mares and 0.4 for deslorelin-treated mares. Furthermore, the number of embryos per ovulation was greater for eFSH-treated mares than for deslorelin -treated mares. Based on this study, it would appear that either twice daily injections of deslorelin or twice daily injections of eFSH can be used in transitional mares to hasten the first ovulation of the year. In summary, injectable deslorelin is currently being used to hasten ovulation in cycling mares and stimulate follicular development in transitional and seasonal anestrous mares.
Other GnRH analogs Although deslorelin is the most common GnRH analog used in broodmares, others, such as buserelin, have been evaluated. In one study,28 40 transitional mares (March) were assigned to one of three groups: controls; 40 mg buserelin injected twice daily; and subcutaneous implants containing buserelin (100 mg/day for 28 days). The number ovulating
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Gonadotropin-Releasing Hormones within 30 days was zero of 15, seven of 15, and nine of 15 for groups 1, 2, and 3, respectively. Practitioners in central Kentucky have also utilized the twice daily injection of 40 mg buserelin as a means of hastening the first ovulation of the year in transitional mares (KE Wolfsdorf, personal communication, 2008). Generally, only mares with follicles of size 20–25 mm at the onset of treatment respond, and 5–7 days of treatment are required. Buserelin is also used as a means of inducing ovulation in cycling mares. Barrier-Battut et al.29 reported buserelin was equally as effective as hCG in inducing ovulation. However, in their study, buserelin was given two to four times per day for 2 days. In contrast, Camillo et al.30 found that hCG resulted in a greater number of mares ovulating within 48 hours than those given twice daily injections of 40 mg buserelin. Use of a commercial buserelin (Suprefact) in France31 resulted in a similar ovulation rate within 48 hours to hCG controls and significantly less time to ovulation than in untreated controls. This product contains 1.05 mg/mL and 6 mL was given subcutaneously. Buserelin has also been given during early diestrus to pregnant mares as a means of improving pregnancy rates.32, 33 Based on a study involving 2346 mares administered 20–40 mg buserelin between days 8 and 12, post-ovulation pregnancy rates was increased by approximately 10 percentage points. Kanitz et al.33 detected higher levels of LH in mares given buserelin on day 10, but progesterone levels were similar between controls and treated mares. In their study involving 191 mares, buserelin given during diestrus increased pregnancy rates by 10 percentage points, but the differences were not statistically different. The exact mechanism as to how GnRH increases pregnancy rates is unclear since progesterone does not appear to be increased.
References 1. Irvine CHG. GnRH clinical application. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 329–33. 2. Ginther OJ, Wentworth BC. Effect of a synthetic gonadotropin-releasing hormone on plasma concentrations of luteinizing hormones in ponies. Am J Vet Res 1974;35:79–81. 3. Evans MJ, Irvine CHG. Measurement of equine follicle stimulating hormone and luteinizing hormone: response of anestrous mares to gonadotropin releasing hormone. Biol Reprod 1976;15:477–84. 4. Evan MJ, Irvine CHG. Induction of follicular development maturation and ovulation by gonadotropin releasing hormone to acyclic mares. Biol Reprod 1977;16:452–6. 5. Johnson AL. Gonadotropin releasing hormone treatment induces follicular growth and ovulation in seasonally anestrous mares. Biol Reprod 1987;36:1199–206.
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6. Turner JE, Irvine CHG. The effect of various gonadotropinreleasing hormone regimens on gonadotropins, follicular growth and ovulation in deeply anestrous mares. J Reprod Fertil Suppl 1991;44:213–25. 7. Allen WR, Sanderson MW, Greenwood RES, Ellis DR, Crowhurst JS, Simpson DJ, Rossdale PD. Induction of ovulation in anestrous mares with a slow-release implant with a GnRH analogue (ICI 118 630). J Reprod Fertil Suppl 1987;35:409–78. 8. Harrison LA, Squires EL, Nett TM, McKinnon AO. Use of gonadotropin-releasing hormone for hastening ovulation in transitional mares. J Anim Sci 1990;68:690–9. 9. Ainsworth CG, Hyland JH. Continuous infusion of gonadotropin-releasing hormone (GnRH) advances the onset of oestrus cycles in thoroughbred mares on Australian stud farms. J Reprod Fertil Suppl 1991;44:235–40. 10. Johnson AL. Pulsatile administration of gonadotropin releasing hormone advances ovulation in cycling mares. Biol Reprod 1986;35:1123–30. 11. Becker SE, Johnson AL. Effect of gonadotropin-releasing hormone infused in a pulsatile or continuous fashion on serum gonadotropin concentrations and ovulation in the mare. J Anim Sci 1992; 70: 1208–15. 12. Fitzgerald BP, Meyer SL, Affleck KJ, Silvia PJ. Effect of constant administration of a gonadotropin-releasing hormone agonist on reproductive activity in mares: induction of ovulation during seasonal anestrus. Am J Vet Res 1993;54:1735–45. 13. Williams GL, Amstalden M, Blodgett GP, Ward JE, Unnerstall DA, Quirk KS. Continuous administration of lowdose GnRH in mares. I. Control of persistent anovulation during the ovulatory season. Theriogenology 2007;68: 67–75. 14. Collins SM, Zieba DA, Williams GL. Continuous administration of low-dose GnRH in mares II. Pituitary and ovarian responses to uninterrupted treatment beginning near the autumnal equinox and continuing throughout the anovulatory season. Theriogenology 2007;68:673–81. 15. Jochle W, Trigg TE. Control of ovulation in the mare with Ovuplant. A short-term release implant (STI) containing the GnRH analogue deslorelin acetate: studies from 1990 to 1994. J Equine Vet Sci 1994;14:632–44. 16. McKinnon AO, Nobelius AM, Tarrida del Marmol Figueroa S, Skidmore J, Vasey JR, Trigg TE. Predictable ovulation in mares treated with an implant of the GnRH analogue deslorelin. Equine Vet J 1993;25:321–3. 17. Mumford EL, Squires EL, Jochle E, Harrison LA, Nett TM, Trigg TE. Use of deslorelin short-term implants to induce ovulation in cycling mares during three consecutive estrous cycles. Anim Reprod Sci 1995;39:129–40. 18. Mumford EL, Squires EL, Peterson KD, Nett TM, Jasko DJ. Effect of various doses of a gonadotropin-releasing hormone analogue on induction of ovulation in anestrous mares. J Anim Sci 1994;72:178–83. 19. Morehead JA, Blanchard TL. Clinical experience with deslorelin (Ovuplant) in a Kentucky Thoroughbred practice. J Equine Vet Sci 1999;20:358–402. 20. Johnson CA, Thompson Jr DL, Cartmill JA. Pituitary responsiveness to GnRH in mares following deslorelin acetate implantation to hasten ovulation. J Anim Sci 2002;80:2681–7. 21. Farquhar VJ, McCue PM, Nett TM, Squires EL. Effect of deslorelin acetate on gonadotropin secretion and ovarian follicle development in cycling mares. J Am Vet Med Assoc 2001;218:749–52.
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22. Farquhar VJ, McCue PM, Carneval EM, Nett TM, Squires EL. Deslorelin acetate (Ovuplant) therapy in cycling mares: effect of implant removal on FSH secretion and ovarian function. Equine Vet J 2002;34:417–20. 23. McCue PM, Nisinender KD, Farquhar VJ. Induction of ovulation with deslorelin: option to facilitate implant removal. J Equine Vet Sci 2002;22:54–5. 24. McKinnon AO, Vasey JR, Lescun TB, Trigg TE. Repeated use of a GnRH analogue deslorelin (Ovuplant) for hastening ovulation in the transitional mare. Equine Vet J 1996;29:153–5. 25. Stich KL, Wendt KM, Blanchard TL, Brinsko SP. Effects of a new injectable short-term release deslorelin in foal-heat mares. Theriogenology 2004;62:831–6. 26. McCue PM, Magee C, Gee EK. Comparison of compounded deslorelin and hCG for induction of ovulation in mares. J Equine Vet Sci 2007;27:58–61. 27. Raz T, Carley S, Card C. Comparison of the effects of eFSH and deslorelin treatment regimes on ovarian stimulation and embryo production of donor mares in early vernal transition. Theriogenology 2009;71:1358–66. 28. Briant C, Ottogalli M, Guillaume D. Attempt to control the day of ovulation in cycling pony mares by associat-
29.
30.
31.
32.
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ing a GnRH antagonist with hCG. Domest Anim Endocrinol 2004;27:165–78. Barrier-Battut I, Le Poutre N, Trocherie E, Hecht S, Grandchamp des Rauz A, Nicaise JL, V´erin X, Bertrand J, Fi´eni F, Hoier R, Renault A, Egron L, Tainturier D, Bruyas JF. Use of buserelin to induce ovulation in the cyclic mare. Theriogenology 2001;55:1679–95. Camillo F, Pacini M, Panzani D, Vannozzi I, Rota AI, Aria G. Clinical use of twice daily injections of buserelin acetate to induce ovulation in the mare. Vet Res Commun 2004;28:169–72. Levy I, Duchamp G. A single subcutaneous administration of buserelin induces ovulation in the mare: field data. Reprod Domest Anim 2007;42:550–4. Newcombe JR, Martinez TA, Peters AR. The effect of the gonadotropin-releasing hormone analog, buserelin, on pregnancy rates in horse and pony mares. Theriogenology 2001;55:1691–31. Kanitz W, Schneider F, Hoppen HO, Unger C, Nurmberg G, Becker K. Pregnancy rates, LH and progesterone concentrations in mares treated with GnRH agonist. Anim Reprod Sci 2007;97:55–62.
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Chapter 191
Estrogen Therapy Ahmed Tibary
Introduction Estrogens play an important role in the reproductive function of both the mare and the stallion. Estrogens are also a key factor in maternal recognition of pregnancy as the equine embryo has a high level of aromatizing activity. Estrogens have been traditionally used for the induction of estrous behavior in ovariectomized or non-cyclic mares used as jump mares. Estradiol is a potent inhibitor of FSH secretion in horses1, 2 and, when given at sufficiently high doses, prevents follicular growth.3 The discovery of the role of estrogen in the regulation of FSH secretion and follicular growth opened the way for the use of these hormones for estrous/ovulation synchronization in cyclic as well as transitional mares. In addition, the role estrogens play in mid to late pregnancy may justify their use in the management of high-risk pregnancies. Estrogens commonly used in equine reproduction include estradiol cypionate (ECP; Figure 191.3), estradiol benzoate (EB; Figure 191.2), and 17β-estradiol (Figure 191.1). The pharmacokinetics of these products vary primarily by the duration of their effect, with ECP having the longest duration and 17βestradiol having the shortest duration. Long-acting formulations relying on special vehicles are available for various estrogens. It is important to point out that, depending on the country, most if not all these products are only available through compounding pharmacies. It is also important to remember that not all estrogens will target the reproductive tissue in the same manner.
Use of estrogens in teaser mares It is well established that, in absence of a source of progesterone, minimal doses of estrogen will produce estrous behavior in mares. However, the exact
mechanism of activation of this behavior by estrogens is not known. In laboratory animals, it has been suggested that both non-genomic and genomic actions of estrogens interact in a synergistic manner to induce specific behavioral postures associated with receptivity. These mechanims are likely involved in the hormonal feedback on neural systems that regulate gonadotropin secretion and may also be present in male behavior. Estrogens may also operate directly on the central nervous system and may play a role in aggressive behavior.4 Administration of various estrogen products induces estrous behavior in ovariectomized or noncyclic mares. Typically, the onset of behavioral signs of estrus occurs 4 to 12 hours after injection and lasts for variable periods of time depending on the product used (48 hours to 2 weeks). ECP has been used at doses ranging from 1 to 3 mg i.m. and 17βestradiol at doses of 0.5 to 1 mg i.m. for this purpose. 17β-Estradiol given to anestrous or ovariectomized mares will produce estrous behavior as early as 4 hours post-treatment. Even at high doses (10 to 20 mg) estradiol will not produce estrous behavior in mares if luteal tissue (progesterone equal or greater than 1 ng/mL) is present. It is recommended that ovariectomized mares used as jump mares not be given estrogen too frequently as this practice may lead to increased aggressiveness towards stallions and other mares. A protocol used by most practitioners consists of administering 1 mg of ECP i.m. every 2 to 3 weeks during the period of use of the ovariectomized mare. This protocol does not seem to have as many adverse effects. There are no studies on the long-term effects of estrogen administration on the welfare of jump mares. Ovariectomized jump mares tend to develop crested necks after a certain number of years and have a tendency to become obese. This author has observed one anaphylactic reaction in an ovariectomized mare after administration of ECP.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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OH H
H H
CH3 O
O
H
H H
HO
H
HO
Figure 191.1 17β-Estradiol.
Figure 191.3 Estradiol cypionate.
Synchronization of follicular wave Reliable control of follicular dynamic is an important condition for the success of artificial insemination with cooled or frozen semen as well as assisted reproduction biotechnologies, which require a near perfect synchronization of follicular growth. It has long been known that progesterone does not inhibit follicular growth in mares and therefore synchronization protocols based on the use of progestagens alone (injectable, oral, and intravaginal) followed by administration of PGF2α give limited control of follicular development and may only delay ovulation. Estradiol has been shown to have a suppressive effect on FSH release in the mare.5, 6 Administration of a combination of progesterone and estradiol (specifically 17β-estradiol) produces a high degree of control of follicular dynamics in mares.7 The discovery of this effect of estradiol has led to the development of the progesterone/estradiol regimen for estrus synchronization in mares. Daily intramuscular injection of progesterone (150 mg) and 17β-estradiol (10 mg) for 10 days, followed by an injection of a luteolytic dose of PGF2α or analog produces a uniform response return to estrus and development of follicles. Ovulation occurs in about 80% of mares when given hCG or deslorelin 9 days after the end of the treatment.8 This standard dose of progesterone and estrogen works well for mares weighing 450–550 kg but may need to be adjusted for heavier horses. Long-acting estrogens (ECP and EB) are not suitable for this protocol as they produce a more variable response in terms of inhibition of follicular development and resumption of subsequent follicular waves.
CH3 H O
H O
Figure 191.2 Estradiol benzoate.
OH
To circumvent the inconvenience of daily injections for 10 days, biorelease formulations have been proposed but they produce less reliable response than the daily injections. Estradiol and progesterone delivered as a single microencapsulated injection (1.25 g progesterone, 100 mg estradiol) followed by PGF2α or analog, produces similar results as the daily i.m. injections but this formulation is not available commercially.9, 10 Although there are anecdotal reports of the effectiveness of these steroids orally, this route has not been thoroughly investigated. In some countries, intravaginal drug-releasing devices (e.g., sponges, silastic CIDR, PRIDTM or CueMareTM ) have been used to deliver progestagen and produce the same results as progesterone/estradiol injection when estradiol is administered at the time of insertion of the device.11 A single injection of R estradiol at the time of CIDR-B insertion was found to exert a suppressing effect on follicle growth and may also cause a supportive FSH release afR removal. This treatment results in a ter CIDR-B high degree of synchrony and quicker onset of estrus after treatment compared to injections of progesterone/estradiol with 95% of mares in estrus within 2.0 to 2.4 days.11 Although a transient vaginitis is present with these vaginal pessaries, it usually resolves spontaneously within 48 hours and does not seem to have any effect on fertility.
Management of the transitional mare Recently, it was shown that estradiol pretreatment of seasonally anovulatory mares (11 mg i.m. every other day for 10 injections) enhances prolactin and ovarian responses to daily injection of sulpiride starting on day 11 of the treatment and continued until ovulation or 45 days.12 (This treatment is discussed in more detail in Chapter 000 [Use of Dopamine antagonists in the transitional mare].)
H
High-risk pregnancy Many forms of estrogens are present in the pregnant mare and include (in order of magnitude)
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Estrogen Therapy estrone, equilin, equilenin, 17β-estradiol, and 17αestradiol. All these compounds are present in the maternal plasma, urine, and allantoic fluid. Hydroxyderivatives (e.g., 17β-hydroxyequilenin) and sulfoconjugates (e.g., estrone sulfate) are also present. Most estrogens are produced by aromatization in the placenta of the precursor dehydroepiandrosterone (DHA) and 5,7-androstadiene-3,17-diol and 7-dehydro-DHA produced by the large equine fetal gonad and excreted in the umbilical artery.13 In pregnant mares, estrogen levels increase from day 100 of gestation and remain high until the last 2 months of pregnancy, and have been used for decades for diagnosis of pregnancy and evaluation of fetal viability.14, 15 Between day 150 and 310 of gestation, the mean estrogen concentration remains above 1000 pg/mL. Clinical observations suggest that estrogen concentrations below 1000 pg/mL are associated with placentitis.16 The addition of estrogen therapy to the treatment of mares with placentitis has been evaluated in the field. Mares presumed to have placentitis and receiving either estradiol cypionate or 17β-estradiol at a dose of 10 to 20 mg s.i.d. or b.i.d. intramuscularly, in addition to antimicrobials, had better outcome (delivery of live foals) than control mares with placentitis receiving only antimicrobial therapy. The foaling rate in normal mares, placentitis mares receiving estrogens, and placentitis mares without estrogen therapy was 87%, 70%, and 20%, respectively.16 Irrespective of treatment, mares with estrogen level of lower than 300 pg/mL had a less than 15% live foal rate. These results provide some new insights on the medical management of mares with placentitis but need further investigation.
Postpartum The conception rate following breeding on foal heat has long been a subject of debate in the equine breeding industry. Early retrospective studies have shown that pregnancy rates are decreased and early pregnancy loss is increased in mares ovulating before day 10 postpartum.17 This reduced fertility was suspected to be due to insufficient gross involution of the uterus and histological repair of the endometrium. The administration of progesterone and estrogen (P/E) has been examined as a method to delay the first ovulation postpartum.18–20 Injection of P/E in oil for 5 days starting on the day of parturition was found to delay the onset of foal heat by 4.8 days18 and to delay the first postpartum ovulation by 5 to 6 days.18, 20 Several treatment regimens were investigated over the years ranging in length from a single injection on the day of parturition to daily injections for 10 days.18,20–22 Bristol et al.21 found that 92% of
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mares ovulated 6 to 10 days after the end of treatments with P/E starting 18 hours after foaling. The length of treatment (1 to 5 days vs 6 to 10 days) did not affect response significantly. The time of initiation of the P/E treatment in postpartum mares seems to play an important role in the suppression of follicular activity.20, 21 Loy et al.20 delayed first postpartum ovulation by 5.3 days when treatments began within 12 hours of foaling. Lower efficacy in control of follicular wave development was observed when treatments was initiated more than 24 hours postpartum. It is important to point out that these treatments do not necessarily enhance uterine involution.23, 24 A single or double dose of progesterone (300 mg) and 17β-estradiol (20 mg) given within 12 hours of foaling does not affect the rate of uterine involution but delays the first postpartum ovulation.23 In one study, none of the treated mares ovulated before day 10 whereas 6/30 control mares ovulated before day 10. In addition the variance in days to ovulation was lower in treated mares (3.73 days) compared to control (7.64 days).23 This treatment regime may be advantageous in mares foaling later in the breeding season, which tend to exhibit foal heat and ovulation at shorter intervals postpartum. In a study of postpartum mares they were injected with biodegradable poly (DL-lactide) microspheres containing 100 mg 17β-estradiol. Estradiol treatment did not hasten uterine involution, increase duration of estrus, delay ovulation, or increase fertility in these postpartum mares.25
Use in oocyte and embryo recipients Pretreatment of ovariectomized mares with estradiol prior to initiation of progesterone therapy has been used to mimic the endocrinological steroidal events taking place immediately before and after ovulation in the hope that it will prime the uterus for pregnancy and increase pregnancy rate following transfer of embryos. However, results comparing estrogen-treated mares to those treated with progesterone alone did not show any advantage to using estrogens.26 Estrogen therapy has been used to prepare nonovulating oocyte recipients and to eliminate the need for aspiration of follicles from the recipient prior to oocyte transfer.27 Mares receive 17β-estradiol (3.3 mg i.m.) once daily for 1 to 3 days followed by the same dose twice daily for 2 days, with the last dose given the day before transfer. Mares are inseminated in the evening before oocyte transfer.27
Other uses of estrogen Estrogens have been used for the prevention and/ or treatment of uterine infection. The primary
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justification for this use is the role estrogen may play in enhancement of the uterine defense mechanism and increased phagocytic activity in the peripheral circulation.28 Also, estrogens are believed to enhance blood flow to the uterus and improve resistance to infection. However, a recent study showed that estradiol benzoate (5 mg) given at various times of the estrous cycle did not have any effect on blood flow resistance in the mare even during the period where the concentration of estrogen receptor is at a maximum.29 It is possible that these contradictory effects may be due to the type of estrogen used (estradiol benzoate vs 17β-estradiol).30 Estrogen (estradiol benzoate, 50 mg/500 kg mare, single intramuscular injection) is used with progestagen and sulpiride for the induction of lactation in nurse mares. (This use is discussed in more detail in Chapter 234.) Estrogen therapy has also been used empirically to promote cervical dilation or enhance oxytocin action on myometrial contraction but these claims are largely unsubstantiated. Although rare, estrogen-responsive urinary incontinence has been described in the mare. A welldocumented case was described in an 18-year-old Shetland pony with urinary incontinence and chronic urine scalding which responded to estradiol cypionate treatment. Urine dribbling stopped within 3 days of administration of 2 mg of ECP and the mare remained continent as long as the therapy was continued every other day. The incontinence reoccurred within 10 days after stopping the treatment.31 Estrogen plays an important role in sperm production as evidenced by the rather large amounts of this hormone produced by the stallion testis. Estrogens appear to play a role in Leydig cell function, modulation of steroidogenesis, and functional maturation of the spermatozoa.32 Estrogen treatment has been shown play some role in sexual behavior since estrogen injections alone restore sex drive in recent castrates.2 However, there have been no studies on the effect of exogenous estrogen on modulation of stallion reproductive function.
References 1. Garza FJ, Thompson DLJ, St George RL., French DD. Androgen and estradiol effects on gonadotropin secretion and response to GnRH in ovariectomized pony mares. J Anim Sci 1986;62:1654–9. 2. Thompson DLJ, Pickett BW, Squires EL, Nett TM. Effect of testosterone and estradiol 17β alone and in combination on LH and FSH concentrations in pituitary and blood serum of geldings and in serum after GnRH. Biol Reprod 1979;21:1231–7. 3. Woodley SL, Burns PJ, Douglas RH, Oxender WD. Prolonged interovulatory interval after oestradiol treatment in mares. J Reprod Fertil Suppl 1979;27:205–9.
4. Cornil CA, Ball GF, Balthazart J. Functional significance of the rapid regulation of brain estrogen action: Where do the estrogens come from? Brain Res 2006;1126:2–26. 5. Burns PJ, Douglas RM. 1981. Effects of daily administration of estradiol-17ß on follicular growth, ovulation and plasma hormones in mares. Biol Reprod 1981;24:1026. 6. Evans M, Loy RG, Taylor TB, Barrows SP. Effects of steroids on serum FSH and LH, and on follicular development in cyclic mares. J Reprod Fertil Suppl 1982;32:205–12. 7. Lofstedt RM. Control of the estrous cycle of the mare. Vet Clin N Am Equine Pract 1988;4:219–41. 8. Loy RG, Pemstein R, O’Canna D, Douglas RH. Control of ovulation in cycling mares with ovarian steroids and prostaglandin. Theriogenology 1981;15:191–200. 9. Burns PJ, Steiner JV, Sertich PL, Pozar M, Tice TF, Mason D, Love DF. Evaluation of biodegradable microspheres for the controlled release of progesterone and estradiol in an ovulation control program for cycling mares. J Equine Vet Sci 1993;19:521–4. 10. Jasko DF, Farlin ME, Hutchinson H, Moran DM, Squires E, Burns PJ. Progesterone and estradiol in biodegradable microspheres for control of estrus and ovulation in mares. Theriogenology 1993;40:465–78. 11. Klug E, Jochle W. Advances in synchronizing estrus and ovulations in the mare: A mini review. J Equine Vet Sci 2001;21:474–9. 12. Kelley KK, Thompson DL, Storer WA, Mitcham PB, Gilley RM, Burns PJ. Estradiol interactions with dopamine antagonists in mares: Prolactin secretion and reproductive traits. J Equine Vet Sci 2006;26(11): 517–28. 13. Ousey JC. Peripartal endocrinology in the mare and foetus. Reprod Dom Anim 2004;39:222–31. 14. Allen WE, Wilsher S, Stewardt F, Ousey JC, Fowden A. The influence of maternal size on placental, fetal and postnatal growth in the horse. II. Endocrinology of pregnancy. J Endocrinol 2002;172:237–46. 15. Kashman LH, Hughes JP, Stabenfeldt GH, Starr MD, Lasley BL. Estrone sulfate concentrations as an indicator of fetal demise in horses. Am J Vet Res 1988;49:184–7. 16. Douglas RH. Endocrine diagnostics in the broodmare: What you need to know about progestins and estrogens. In: Proceedings of the Annual Conference of the Society for Theriogenology, Aug 4–7, 2004, Lexington, Kentucky, pp. 106–15. 17. Loy RG. Characteristics of postpartum reproduction in the mare. Vet Clin N Am Large Anim Pract 1980;2:345–9. 18. Bell RJ, Bristol F. Fertility and pregnancy loss after delay of foal estrus with progesterone and estradiol-17β. J Reprod Fertil 1987;35:667–8. 19. Burns PJ, Douglas RH. Effect of exogenous steroids on follicular growth, ovulation and FSH secretions in postpartum mares. In: Proceedings of the 73rd annual meeting of the American Society of Animal Science, Raleigh, NC, 1981; p. 433. 20. Loy RG, Evans G, Pemstein R, Taylor TB. Effects of injected ovarian steroids on reproductive patterns and performance in postpartum mares. J Reprod Fertil 1982;32:199–204. 21. Bristol F, Jacobs KA. Synchronization of estrus in postpartum mares with progesterone and estradiol 17b. Theriogenology 1983;19:779–85. 22. Sexton PE, Bristol FM. Uterine involution in mares treated with progesterone and estradiol 17b. J Am Vet Med Assoc 1985;186:250–6. 23. Bruemmer JE, Brady HA, Blanchard TL. Uterine involution, day and variance of first postpartum ovulation in mares
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24.
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26.
27.
treated with progesterone and estradiol-17 beta for 1 or 2 days postpartum. Theriogenology 2002;57(2): 989–95. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproductive tract of mares after parturition: effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed first postpartum ovulatory cycles. J Am Vet Med Assoc 1988;192:350–3. Arrott C, Macpherson M, Blanchard TL, Varner D, Thompson DL, Simpson B, Bruemmer JE, Vogelsang S, Fernandez M, Fleet T, Burns PJ. Biodegradable estradiol microspheres do not affect uterine involution or characteristics of postpartum estrus in mares. Theriogenology 1994;42:371– 84. McKinnon AO, Squires EL, Carnevale EM, Hermenet MJ. Ovariectomized steroid-treated mares as embryo transfer recipients and as a model to study the role of progestins in pregnancy maintenance. Theriogenology 1988;29:1055–63. Hinrichs K, Provost PJ, Torello EM. Treatments resulting in pregnancy in nonovulating, hormone-treated
28.
29.
30.
31.
32.
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oocyte recipient mares. Theriogenology 2000;54(8): 1285– 93. Washburn SM, Klesius PH, Ganjam VK, Brown BG. Effect of oestrogen and progesterone on the phagocytic response of ovariectomized mares infected with beta-hemolytic streptocci. Am J Vet Res 1982;43:1367–70. Bollwein H, Kolberg B, Stolla R. The effect of exogenous estradiol benzoate and altrenogest on uterine and ovarian blood flow during the estrous cycle in mares. Theriogenology 2004;61(6): 1137–46. Varner DD, Blanchard TL, Brinsko SP. Estrogens, oxytocin and ergot alkaloids – Uses in reproductive management of mares. In: Proceedings of the American Association of Equine Practitioners, 1988; pp. 219–41. Madison JB. Estrogen-responsive urinary incontinence in an aged pony mare. Comp Cont Educ Pract Vet 1984;6(7): S390–2. Roser JF. Regulation of testicular function in the stallion: An intricate network of endocrine paracrine and autocrine systems. Anim Reprod Sci 2008;107:179–96.
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Chapter 192
Oxytocin Michelle M. LeBlanc
Introduction Oxytocin is a neuropeptide synthesized as part of a large precursor molecule, neurophysin, in the supraoptic nucleus of the hypothalamus and is transported in small vesicles down the hypothalamichypophyseal nerve axons.1 Oxytocin is comprised of nine amino acids that appear to be cleaved from neurophysin during transport from cell bodies in the hypothalamic nuclei to nerve terminals in the posterior pituitary. Stimulation of oxytocin neurons in the hypothalamus result in its release from nerve terminals in the posterior pituitary. Oxytocin has a number of functions in reproduction. During the follicular phase of the estrous cycle and during the late stages of gestation, oxytocin stimulates uterine contractions. During estrus, uterine contractions facilitate sperm transport to the oviduct and assist in clearing inflammatory debris associated with mating from the uterine lumen. During parturition, oxytocin plays a pivotal role in the instigation and continuation of uterine contractions until the foal is delivered and fetal membranes are dropped. The best-known action of oxytocin is the reflex release of milk from the mammary gland. Oxytocin causes contraction of myoepithelial cells that surround the alveoli in the mammary gland, resulting in milk letdown1 when a lactating mare is stimulated tactically or visually. Oxytocin also contributes to luteal function. Late in diestrus, oxytocin acts on the endometrium to induce prostaglandin F2α release leading to regression of the corpus luteum.2 If given before day 10 after ovulation, continual or repeated daily administration results in prolongation of the corpus luteum.3 Clinical uses for oxytocin include promotion of uterine contractions to clear uterine fluids, loosen retained fetal membranes, and induce parturition. It has been given to mares to promote maternal behavior, induce lactation, and recently as a method for prolonging the life span of the corpus luteum to prevent estrus behavior. In the stallion, it is given for
blocked ampulla, to increase sexual arousal, and to induce a more complete ejaculation.4 This chapter will discuss the use of oxytocin for promoting uterine contractions, inducing parturition, and prolonging the life span of the corpus luteum.
Promoting uterine contractions in mares during estrus Persistent mating-induced endometritis or delayed uterine clearance is a major cause of reduced fertility in mares and affects approximately 15% of Thoroughbred mares5 in clinical practice. Oxytocin in combination with uterine irrigation after breeding is the current treatment; however, until the early 1990s, oxytocin was primarily used for inducing parturition in the mare. Prior to that time, it was thought that the uterus would only respond to oxytocin in the first few days after foaling and that, if given at other times of the cycle, it would cause mares to colic. In 1991, studies conducted in New Zealand indicated that oxytocin given during estrus increased uterine myometrial electrical activity.6 These studies were followed by landmark work by Dr W.E. Allen who showed through repeated ultrasonographic evaluations of the uterus that oxytocin promoted uterine drainage in mares with defective uterine clearance.7 Uterine scintigraphy studies confirmed this finding.8 Mares that cleared less than 10% of a radiocolloid infused into the uterine lumen within 2 h, cleared more than 95% of the colloid within 30 minutes of intravenous injection of 20 IU of oxytocin.8 A rapid succession of clinical studies then followed, indicating that oxytocin improved pregnancy rates in mares with delayed uterine clearance of intrauterine fluid.9–11
Dose The length and strength of uterine contractions induced by oxytocin varies by dose, when it is given
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Oxytocin during the estrous cycle, and by route of administration. When given during estrus and in the 48h period after ovulation, intravenous doses of oxytocin ranging from 5 to 20 IU induce high-amplitude uterine contractions for approximately 30 to 45 minutes.8,12–14 A dose of 10 IU of oxytocin given during estrus but before ovulation caused the uterus to contract longer than when 5 or 2.5 IU were given intravenously (24 minutes, 16.3 minutes, and 17.6 minutes, respectively).15 After ovulation, the effect of oxytocin on uterine contractility diminishes so doses of 20 to 25 IU are recommended.13 The decrease in response after ovulation appears to be inversely related to the concentration of progesterone.12 High doses of oxytocin, 30 IU or greater given intravenously during estrus, appear to decrease uterine contractions and inhibit emptying.16 High doses may cause a sustained contraction rather than an increased frequency of contractions, thereby interfering with uterine emptying. Intramuscular administration of oxytocin after breeding produces a slightly different response than intravenous administration. Mean time to onset of uterine contractions after intramuscular administration is 11.8 ± 1.0 minutes whereas response to treatment after intravenous administration is 0.5 to 1 minute.15, 17 Duration of uterine contractions also vary by intramuscular dose. Low doses of oxytocin given intramuscularly, 5 or 10 IU, are associated with 40 to 50 minutes of myometrial activity while higher doses, 20 to 40 IU, are associated with 90 to 100 minutes of activity.17 The half-life of oxytocin is 6.8 minutes.13 A longacting synthetic oxytocin analog, carbetocin, has recently become available in Europe, Canada, and Mexico. It was well tolerated in a group of horses following intravenous administration of 175 µg. The half-life of carbetocin is about 17 minutes, or 2.5 times that of oxytocin. The drug may be of benefit in mares where more prolonged uterine contractions are needed.18 Oxytocin does not appear to be effective in all mares that accumulate intrauterine fluid. Some mares appear to be refractory to low doses of oxytocin (20 IU given intravenously or intramuscularly) but will respond dramatically to higher doses (50–80 IU) when given as an intrauterine infusion.19 Factors that may affect response include age of the mare, concurrent drug administration, endometrial oxytocin receptor status, a pendulous uterus, inappropriate dose, abnormal propagation of uterine contractions, and inflammatory conditions in the uterus. Propagation of uterine contractions has been found to differ between reproductively normal mares and mares with delayed uterine clearance when mares were given a tranquilizer before receiving oxytocin in an experimental model.20 Following administration of
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detomidine or detomidine and oxytocin, intraluminal uterine pressure first increased in the uterine horn and then in the uterine body in reproductively normal mares. This pattern was rarely seen in susceptible mares. They responded with an inverted contraction pattern with the first increase in pressure occurring near the cervix, followed by contractions in the uterine horn. This inverted pattern could interfere with flow of luminal fluids into the vagina and may result in drug failure in some mares with delayed uterine clearance. It is not known if these mares would respond differently to cloprostenol, a prostaglandin analog that induces uterine contractions.
Relationship between oxytocin and prostaglandin Administration of oxytocin to mares during estrus stimulates prostaglandin F2α release and uterine emptying.21, 22 However, the mechanism that controls myometrial contractions is not known. In women and rats, occupation of myometrial oxytocin receptors directly stimulates the myometrial cell to contract while occupation of endometrial receptors indirectly stimulates the myometrial cell and therefore uterine contractions.23 Occupied myometrial oxytocin receptors induce contractions by causing an influx of calcium into the cell through calcium channels. Occupied endometrial oxytocin receptors activate the phosphatidylinositol pathway via an activation of phospholipase C. Associated with the increase in inositol triphosphate and diacylglycerol is an increase in calcium and mobilization of arachidonic acid that leads to PGE2 and PGF2α secretion. The resultant prostaglandins then diffuse from the endometrium into the adjacent myometrium to act on the myometrial cell.23, 24 In both pathways, calcium is released from the sarcoplasmic reticulum, and the contractile mechanism is activated.25, 26 The mare also has oxytocin receptors in the myometrium and endometrium27 and uterine contractions may be induced both directly through contraction of myometrial cells and indirectly through prostaglandin release from the endometrium. Administration of oxytocin to reproductively normal mares causes a dramatic clearance of uterine radiocolloid within 15 minutes and an acute increase in plasma PGFM.28 Uterine clearance of radiocolloid and the rise in plasma PGFM concentrations are inhibited, however, when mares received 2 g of phenylbutazone. The inhibitory effect of phenylbutazone on uterine clearance of radiocolloid can be overridden by administration of 20 IU of oxytocin, resulting in rapid clearance of radiocolloid without a concurrent rise in PGFM. As it is not uncommon for broodmares to receive phenylbutazone for
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musculoskeletal problems, mares may accumulate intrauterine fluids after breeding.21, 29 Thus one should consider administering oxytocin to these mares 4 to 8 h after breeding to stimulate uterine contractions and clearance of inflammatory debris associated with breeding.
Clinical use of oxytocin in post-mating endometritis Mares with post-mating induced endometritis accumulate intrauterine fluid after mating. The timing of post-breeding treatment has been designed to mimic the inflammatory response and the rate of uterine clearance observed in reproductively normal mares. After mating, there is a rapid, dramatic influx of neutrophils into the uterine lumen, with concentrations peaking around 8 h.30 By 12 h post-mating, neutrophil numbers are on the decline. This decline in neutrophil numbers is not seen in mares prone to post-mating endometritis, instead there is a steady rise in neutrophils within the uterine lumen. In an experimental model of endometritis, reproductively healthy mares and mares susceptible to endometritis received an intrauterine inoculation of β-hemolytic streptococcus during estrus and clearance of the bacteria was monitored. Bacterial numbers within the uterine lumen for the two groups were similar until 9 h, when clearance rates diverged. Reproductively normal mares had virtually no bacteria remaining in the uterine lumen after 9 h whereas susceptible mares exhibited logarithmic growth in bacteria, with numbers peaking 48 h after inoculation.31 Clinical use of oxytocin after breeding is based on the results of these experimental findings. Oxytocin is first given between 4 and 8 h after mating. This gives sperm time to travel to the oviduct but not enough time for bacteria to begin a logarithmic growth phase.31, 32 Delaying oxytocin treatment post-mating results in lower pregnancy rates.33 Oxytocin may be administered repeatedly at 4- to 6-hour intervals for 2 to 3 days to assist in clearance of uterine fluid. Uterine lavage is commonly added to oxytocin therapy especially if the mare has ≥ 2 cm of fluid in the uterine lumen.33–36 All susceptible mares should be re-evaluated 24 hours after breeding to determine if there is intrauterine fluid as additional treatments such as uterine irrigation, ecbolics, or antibiotics may be needed.
Induction of parturition Inducing parturition should never be undertaken lightly as an induced delivery can result in critical problems for both the mare and foal. Final maturation of the foal occurs only in the last 5 to 7 days of gestation37–39 and, as normal gestation length varies
from 325 to 360 days, it is difficult to predict when a mare will foal. If parturition is induced outside the narrow window of final maturation, the foal may be premature at birth and may die even after extensive neonatal care. Problems that may occur in the mare include premature placental separation, dystocia, and retained fetal membranes. Criteria used for determining if a mare can be induced include a minimum gestation length of 330 days, colostrum in the udder, mammary secretion calcium concentrations > 40 mg/dL (10 mmol/L (conversion of mg/dL to mmol/L is × 0.25) and mammary secretion potassium concentration greater than the sodium concentration.40, 41 It has also recommended that the mare have a relaxed cervix and softening of the cervical plug.42 Parturition has been induced in the mare with oxytocin, prostaglandin or dexamethasone.43–46 Of the three drugs, oxytocin is used most often. Various doses of oxytocin have been used and range from 2.5 to 140 IU. These doses have been given as an intravenous bolus, an intramuscular injection, or a slow intravenous drip. Early reports recommended administering 100 to 140 IU to a 450 kg mare as a single intramuscular injection.44 Mares induced with these doses showed signs of marked discomfort and sweating in 10 to 15 minutes and most delivered the foal within 30 minutes of injection. They also exhibited significant abdominal discomfort and many foals were hypoxic at delivery.44 Therefore, lower doses ranging from 40 to 60 IU are advocated and are given either as an intravenous bolus or added to 1 liter of physiological saline that is administered over 15 minutes. Alternatively, 10 to 15 IU of oxytocin can be administered intramuscularly or intravenously every 20 minutes until rupture of the chorioallantois (subject to a maximum total dose of 75 IU).47 Time from oxytocin injection to delivery adversely affects foal viability.42 Mares given small doses of oxytocin (15 IU) intramuscularly every 15 minutes for a total dose of 75 IU took longer to deliver their foals than mares induced with a intravenous oxytocin drip (75 IU of oxytocin in 1 L of physiological saline solution). More foals were abnormal when the interval from oxytocin administration to delivery was > 60 minutes and these foals took longer to stand and suckle than normal foals.42 Daily low dose of oxytocin for induction of parturition in mares at term has recently been recommended as a useful, safe method for simplifying the management of foaling mares. The major advantage of injecting low-dose oxytocin appears to be that such a low dose induces delivery only in mares carrying a mature fetus and which are ready to foal. Two studies have been reported48, 49 using the technique. In the first study, 38 near-term Haflinger mares were given
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Oxytocin a single injection of 2.5 IU oxytocin intravenously between 1700 and 1900 h when mammary secretion calcium carbonate concentrations were ≥ 200 ppm (14 mmol/L), while ten mares were given a placebo.48 The test kit used in the study measured only calcium carbonate and not magnesium. Many mammary secretion electrolyte test kits test for divalent cations which includes magnesium as well as calcium. Magnesium rises in mammary secretions earlier than calcium, complicating interpretation of results50 (see Chapters 232 and 286). Of the treated mares, 95% delivered a healthy, viable foal within 120 minutes; 24 of 38 (63%) foaled after the first injection of oxytocin, 9 of 38 (24%) after the second injection, and 3 of 38 (8%) foaled after the third treatment. In the second study, conducted on a large commercial Standardbred farm, 174 mares were given 3.5 IU of oxytocin intramuscularly when mammary secretion calcium carbonate concentrations were ≥ 200 ppm while 176 mares were allowed to foal spontaneously.49 Of the treated mares, 70% delivered a viable, healthy foal within 120 minutes of oxytocin administration with 51% responding to the first injection, 14% responding to the second, and 3.4% responding to the third injection. Duration of foaling, incidence of dystocia, or premature placental separation did not differ between the two groups. Physical and behavioral characteristics were normal in foals from both groups.
Retained fetal membranes Fetal membranes are normally expelled within 30 minutes to 3 hours in the mare.51 However, it is not uncommon for the fetal membranes to remain in the mare for up to 12 to 16 h without clinical signs of illness. Oxytocin used alone or in combination with other treatments is the most common therapy for retained fetal membranes.52, 53 Initial treatment is usually begun 5 to 8 h after foaling and consists of repeated doses of oxytocin either intravenously or intramuscularly. There are no scientific reports that establish the best dosing interval, the optimal dose or route of administration for the treatment of retained placenta with oxytocin. Recommended doses range from 10 to 40 IU, given at intervals of 30 minutes to 4 hours, in multiple repeated intravenous or intramuscular injections or as a slow intravenous drip diluted in physiological saline. It has been suggested that slow intravenous infusion of 30 to 60 IU diluted in 1 L of physiological saline solution administered over 1 h provides a more physiological response to treatment than repeated bolus injections.53 In refractory cases, administration of 1 L of lactated Ringer solution containing 100 to 150 mL of 23% calcium gluconate followed by 20 IU of oxytocin intravenously has
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resulted in rapid release of membranes (LeBlanc, unpublished observations). The postpartum mare has more available oxytocin receptors than the cyclic mare and therefore can respond with abdominal discomfort if high doses are given frequently. Preliminary work from our laboratory showed that postpartum pony mares fitted with uterine myometrial electrodes and given 10 IU of oxytocin intravenously on days 1,3, 5, and 7 post-foaling exhibited waves of uterine contractions for 3 h, the length of each recording session, after administration of oxytocin (C Ellis and M LeBlanc, unpublished observations). However, none of the pony mares had retained their fetal membranes.
Postpartum conditions Oxytocin is given in combination with systemic antibiotics, uterine lavage, and non-steroidal antiinflammatory drugs for the treatment of metritis. Doses are similar to that used for retained fetal membranes. One dose is always given after large volume uterine lavage and remaining doses are given at 4- to 6-h intervals. Mares that are endotoxic or dehydrated may also require intravenous fluids so oxytocin can be added to the fluids (40 to 60 IU/5 L lactated Ringer solution). Oxytocin has been used to stimulate uterine involution in mares after dystocia, retention of fetal membranes, or vestibule-vaginal reflux. It has also been recommended for mares that are confined to a stall because their foal has medical or orthopedic problems.54 Recommended doses range from 20 to 40 IU and are given at 6- to 8-h intervals. Oxytocin therapy is often combined with large volume uterine lavage to remove lochia and debris in addition to stimulating uterine involution. Oxytocin does not appear to improve the rate of uterine involution or pregnancy rate in mares that do not experience foaling difficulties or postpartum problems. Arabian mares administered either oxytocin (30 IU) or cloprostenol (250 µg) within 12 h of foaling had similar foal heat pregnancy rates and no difference in cross-sectional diameter of the gravid uterine horn.55 Oxytocin is also included as part of the medical management for prolapsed uterus and uterine tears. Oxytocin is given after a prolapsed uterus has been replaced and the uterine horns are everted and not before, because administration before replacement increases uterine tone, making it difficult to evert the uterine horns. Uterine tears occur most commonly at the tip of the pregnant horn. Uterine tears may be difficult to identify in the first few days after foaling if they are small and they are difficult to palpate rectally or visualize ultrasonographically.
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Uterine tears have been treated both medically or surgically. Frequent injections of oxytocin have been used as part of the therapy. Doses ranging from 20 to 50 IU have been recommended and given intravenously, intramuscularly or contained within 1 L of lactated Ringer solution and given slowly intravenously.
Blocking luteolysis and estrous behavior Many owners request veterinarians to provide a method of blocking estrous behavior in their performance mares when that behavior is deemed undesirable or interferes with intended activities. Intramuscular injections of progesterone, oral administration of altrenogest, insertion of growth-promoting implants containing progesterone and estradiol, and intrauterine insertion of a glass ball to extend corpus luteum function are all methods that have been used. Each method has varying degrees of success and complications including swelling at injection site, inconvenience of daily oral administration, premature loss or splintering of a glass ball in the uterus. Recent work indicates that intramuscular administration of 60 IU of oxytocin twice daily on days 7 to 14 after ovulation is an efficacious method of inhibiting luteolysis and extending corpus luteum function in mares.3 All of six mares administered oxytocin retained their corpus luteum and had plasma progesterone concentrations > 1.0 ng/mL for 30 or more days. Although the exact mechanism of action is not known, it is hypothesized that repeated or continuous administration of oxytocin initiated before day 10 after ovulation prevents luteolysis by inhibiting the increase in endometrial oxytocin receptor concentration that permits oxytocin-induced PGF2α secretion at the time of luteolysis.56
References 1. Hafez ESE, Jainudeen MR, Rosnina Y. Hormones, growth factors and reproduction. In: Hafez B, Hafez ESE (eds) Reproduction in Farm Animals, 7th edn. Philadephia: Lippincott Williams & Wilkins, 2000; pp. 33–54. 2. Starbuck GR, Stout TA, Lamming GE, Allen WR, Flint AP. Endometrial oxytocin receptor and uterine prostaglandin secretion in mares during the oestrous cycle and early pregnancy. J Reprod Fertil 1998;113:173–9. 3. Vanderwall DK, Rasmussen DM, Woods GL. Effect of repeated administration of oxytocin during diestrus on duration of function of corpora lutea in mares. J Am Vet Med Assoc 2007;231:1864–7. 4. Tibary A. Stallion reproductive behavior. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 174– 84.
5. Zent WW, Troeddsson MHT. Postbreeding uterine fluid accumulation in a normal population of thoroughbred mares: a field study. Proc Am Assoc Equine Pract 1998;64–5. 6. Jones DM, Fielden ED, Carr DH. Some physiological and pharmacological factors affecting uterine motility as measured by electromyography in the mare. J Reprod Fertil 1991;Suppl 44:357–68. 7. Allen WE. Investigations into the use of exogenous oxytocin for promoting uterine drainage in mares susceptible to endometritis. Vet Rec 1991;128:593–4. 8. LeBlanc M, Neuwirth L, Mauragis D, Klapstein E, Tran T. Oxytocin enhances clearance of radiocolloid from the uterine lumen of reproductively normal mares and mares susceptible to endometritis. Equine Vet J 1994;26:279–82. 9. LeBlanc MM. Oxytocin – the new wonder drug for treatment of endometritis? Equine Vet Ed 1994;6:39–43. 10. Pycock JF. Assessment of oxytocin and intrauterine antibiotics on intrauterine fluid and pregnancy rates in mares. Proc Am Assoc Equine Pract 1994;19–20. 11. Pycock JF, Newcombe JR. Assessment of the effect of three treatments to remove intrauterine fluid on pregnancy rate in the mare. Vet Rec 1996;138:320–3. 12. Gutjahr S, Paccamonti DL, Pycock JF, Taverne MA, Dieleman SJ, van der Weijden GC. Effect of dose and day of treatment on uterine response to oxytocin in mares. Theriogenology 2000;54:447–56. 13. Paccamonti DL, Pycock JF, Taverne MA, Bevers M, Van Der Weijden GC, Gutjahr S, Schams D, Blouin D. PGFM response to exogenous oxytocin and determination of the half-life of oxytocin in nonpregnant mares. Equine Vet J 1999;31:285–8. 14. Troedsson MHT, Liu IKM, Ing M, Pascoe J. Smooth muscle electrical activity in the oviduct, and the effect of oxytocin, prostaglandin F2α , and prostaglandin E2 on the myometrium and the oviduct of the cycling mare. Biol Reprod 1995;Mono 1:475–88. 15. Cadario ME, Merritt AM, Archbald LF, Thatcher WW, LeBlanc MM. Changes in intrauterine pressure after oxytocin administration in reproductively normal mares and in those with a delay in uterine clearance. Theriogenology 1999;51:1017–25. 16. Campbell MLH, England GCW. A comparison of the ecbolic efficacy of intravenous and intrauterine oxytocin treatments. Theriogenology 2002;58:473–7. 17. Madill S, Troeddsson MHT, Santschi EM, Malone ED. Dose-response effect of intramuscular oxytocin treatment on myometrial contraction of reproductively normal mares during estrus. Theriogenology 2002;58:479–81. 18. Schramme AR, Pinto CR, Davis J, Whisnant CS, Whitacre MD. Pharmacokinetics of carbetocin, a long-acting oxytocin analogue, following intravenous administration in horses. Equine Vet J 2008;40:658–61. 19. McKinnon AO. Breeding the problem mare. Proceedings of the 11th South African Equine Veterinary Association Congress, Kwazulu Natal, 2009; pp. 192–227. 20. von Reitzenstein M, Callahan MA, Hansen PJ, LeBlanc MM. Aberrations in uterine contractile patterns in mares with delayed uterine clearance after administration of detomidine and oxytocin. Theriogenology 2002;58:887–98. 21. Cadario ME, Thatcher WW, Klapstein E, Merrit AM, Archbald LF, Thatcher MJ, LeBlanc MM. Dynamics of prostaglandin secretion, intrauterine fluid and uterine clearance in reproductively normal mares and mares with delayed uterine clearance. Theriogenology 1999;52:1181– 92.
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Oxytocin 22. Goff AK, Pontbriand D, Sirois J. Oxytocin stimulation of plasma 15-keto-13,14-dihydro-prostaglandin F2α during the estrous cycle and early pregnancy in the mare. J Reprod Fertil 1987;Suppl 35:253–60. 23. Fuchs AR. Prostaglandin F2α and oxytocin interaction in ovarian and uterine function. J Steroid Biochem 1987;27: 1073–80. 24. Ghaudhuri G. Biosynthesis and function of econsanoids in the uterus. In: Carsten ME, Miller JD (eds) Uterine Function: Molecular and Cellular Aspects. New York: Plenum Press, 1990; pp. 423–48. 25. Carsten ME. Hormonal regulation of myometrial caclium transport. Gynecol Invest 1974;5:269–75. 26. Carsten ME, Miller JD. Effect of prostaglandin and oxytocin on calcium release from a uterine microsomal fraction. J Biol Chem 1977;252:1576–81. 27. Stull CL, Evans JW. Oxytocin binding in the uterus of the cycling mare. J Equine Vet Sci 1986;6:14–19. 28. Cadario ME, Thatcher MD, LeBlanc MM. Relationship between prostaglandin and uterine clearance of radicolloid in the mare. Biol Reprod 1995;Monograph Series 1:495–500. 29. LeBlanc MM. Effects of oxytocin, prostaglandin and phenylbutazone on uterine clearance of radiocolloid. Pferdeheilkunde 1997;13:483–5. 30. Katila T. Onset and duration of uterine inflammatory response of mares after insemination with fresh semen. Biol Reprod 1995;Monograph Series 1:515–17. 31. Williamson P, Munyua S, Martin R, Penhale WJ. Dynamics of the acute uterine response to infection, endotoxin infusion and physical manipulation of the reproductive tract in the mare. J Reprod Fertil 1987;Supplement 35:317–25. 32. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post insemination on pregnancy rate in mares. Theriogenology 1991;35:1111–19. 33. Knutti B, Pycock JF, van der Weijden GC, Kupper U. The influence of early postbreeding uterine lavage on pregnancy rate in mares with intrauterine fluid accumulations after breeding. Equine Vet Ed 2000;5:346–9. 34. Brinsko S, Varner DD, Blanchard TL, Meyers SA. The effect of postbreeding uterine lavage on pregnancy rate in mares. Theriogenology 1990;33:465–475. 35. Rasch K, Schoon HA, Sieme H, Klug E. Histomorphological endometrial status and influence of oxytocin on the uterine drainage and pregnancy rate in mares. Equine Vet J 1996;28:455–60. 36. Troedsson MH, Scott MA, Liu IK. Comparative treatment of mares susceptible to chronic uterine infection. Am J Vet Res 1995;56:468–72. 37. Cudd TA, LeBlanc M, Silver M, Norman W, Madison J, Keller-Wood M, Wood CE. Ontogeny and ultradian rhythms of adrenocorticotropin and cortisol in the lategestation fetal horse. J Endocrinol 1995;144:271–83. 38. Rossdale PD, Ousey JC, Chavatte P. Readiness for birth: an endocrinological duet between fetal foal and mare. Equine Vet J Suppl 1997: 96–99. 39. Silver M, Fowden AL. Pre-partum adrenocortical maturation in the fetal foal: response to ACTH1–24. J Endocrinol 1994;142:417–25.
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40. LeBlanc MM. Induction of labor in the mare: assessment of readiness for birth. In: Koterba A, Drummond W (eds) Manual of Equine Neonatal Care. Philadelphia: Lea & Febiger, 1990; pp. 34–40. 41. Ousey JC, Dudan F, Rossdale PD. Preliminary studies on mammary secretions in the mare to assess foetal readiness for birth. Equine Vet J 1984;16:259–63. 42. Macpherson ML, Chaffin MK, Carroll GL, Jorgensen J, Arrott C, Varner DD, Blanchard TL. Three methods of oxytocin-induced parturition and their effects on foals. J Am Vet Med Assoc 1997;210:799–803. 43. Alm CC, Sullivan JJ, First NL. Induction of premature parturition by parenteral administration of dexamethasone in the mare. J Am Vet Med Assoc 1975;165:721– 2. 44. Jeffcott LB, Rossdale PD. A critical review of current methods for induction of parturition in the mare. Equine Vet J 1977;9:208–15. 45. Ousey JC, Kolling M, Allen WR. The effects of maternal dexamethasone treatment on gestation lenght and foal maturation in Thoroughbred mares. Anim Reprod Sci 2006;94:436–8. 46. Rossdale PD, Pashen RL, Jeffcott LB. The use of synthetic prostaglandin analogue (fluprostenol) to induce foaling. J Reprod Fertil 1979;Suppl 27:521–9. 47. Lopate C, LeBlanc MM, Pascoe RR, Knottenbelt DC. Parturition. In: Knottenbelt DC, Le Blanc MM, Lopate C, Pascoe RR (eds) Equine Stud Farm Medicine and Surgery. Edinburgh: Saunders, 2003; pp. 269–324. 48. Camillo F, Marmorini P, Romagnoli SE, Cela M, Duchamp G, Palmer E. Clinical studies on daily low dose oxytocin in mares at term. Equine Vet J 2000;32:307–10. 49. Villani M, Romano G. Induction of parturition with daily low-dose oxytocin injections in pregnant mares at term: clinical applications and limitations. Reprod Domest Anim 2008;43:481–3. 50. Paccamonti D. Milk electrolytes and induction of parturition. Pferdeheilkunde 2001;17:616–18. 51. Roberts SJ. Veterinary Obstetrics and Genital Disease (Theriogenology), 3rd edn. Woodstock, VT: published by the author, 1986. 52. Threlfall WR, Provencher R, Carleton CL. Retained fetal membranes in the mare. Proc Am Assoc Equine Pract 1987;649–56. 53. Vandeplassche M, Spincemaille J, Bouters R. Aetiology, pathogenesis and treatment of retained placenta in the mare. Equine Vet J 1971;3:144–7. 54. Matthews P, Samper JC. Breeding the post-partum mare. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination, 2nd edn. St Louis, MO: Saunders Elsevier, 2009; pp. 277–9. 55. Gunduz MC, Kasikci G, Kaya HH. The effect of oxytocin and PGF2alpha on the uterine involution and pregnancy rates in postpartum Arabian mares. Amin Reprod Sci 2008;104:257–63. 56. Stout TAE, Lamming GE, Allen WR. Oxytocin administration prolongs luteal function in cyclic mares. J Reprod Fertil 1999;116:315–20.
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Chapter 193
Superovulation Edward L. Squires and Patrick M. McCue
Introduction Superovulation has been a subject of investigation for the past 30 years. Numerous products have been evaluated over the past three decades including gonadotropin-releasing hormone (GnRH), various gonadotropins, and inhibin immunoneutralization. Superovulation has the potential for increasing embryo recovery and decreasing the cost of embryo transfer. It also is a means of providing extra embryos for embryo freezing or vitrification. The following equation demonstrates the success of obtaining a pregnancy on a per cycle basis from an embryo donor. Success is the product of embryo recovery rate multiplied by pregnancy rate after transfer of the embryo. If one uses a 60% embryo recovery rate multiplied by a 75% pregnancy rate after embryo transfer, this means that, on a given cycle, one has a 45% chance of obtaining a pregnant recipient. The only part of this equation that can be increased dramatically is embryo recovery rate through superovulation. If one superovulates a mare and obtains two or three embryos per flush, then the pregnancy rate per cycle is well over 100%. It has been stated in the literature that superovulation cannot occur in the mare since the ovary is ‘inside out’ compared with the cow. It is certainly true that cattle and other farm animals respond to superovulatory drugs much more readily than the horse. The superovulated cow provides an average of six transferable embryos. However, nearly all studies that have been conducted have shown an increase in embryo recovery with superovulation in mares.1 Historical perspective It is not the intent of this review to provide a detailed description of all the treatments that have been used for superovulation of mares in the past since these have appeared in recent reviews.1, 2 The majority of this chapter will focus on the more recent
use of equine follicle-stimulating hormone (FSH) for superovulation of mares. There have been numerous approaches to superovulation of mares, including injection of porcine FSH, inhibin vaccines, equine chorionic gonadotropin (eCG), and GnRH.1, 2 Even though eCG is involved in development of multiple secondary corpora lutea in pregnant mares, Allen3 injected very large quantities of eCG and was unsuccessful in inducing superovulation in mares. The levels of eCG during gestation are extremely high in the mare and those high levels, in combination with FSH, may be responsible for the multiple follicular development, ovulation, and luteinization in pregnant mares. Investigators have shown that injection of GnRH or GnRH agonists in the transitional mare result in multiple ovulations.4 However, twice daily injections of GnRH in cycling mares did not result in multiple ovulation.5 Although GnRH analogs currently are being used for induction of follicular development and ovulation in transitional mares, this generally requires twice daily injections for several weeks and, thus, is impractical. Secretion of inhibin from the dominant ovarian follicle suppresses FSH secretion.6 Therefore, inhibin antibodies have been used to decrease inhibin bioactivity and increase the concentration of endogenous FSH. Active immunization of mares against inhibin resulted in doubling the ovulation rate in two studies.7, 8 Multiple inoculations over several weeks were required. In addition, occasional adverse reactions at the injection sites have prevented the commercial use of inhibin vaccine for superovulation. With the advances in vaccine technology, it would seem reasonable to reinvestigate the possibility of using inhibin vaccine as a means of superovulation in mares. Studies have examined the use of porcine FSH for induction of multiple ovulations in mares. Squires et al.9 compared the response of mares to injection of equine pituitary extract (EPE) with that to injection of porcine FSH. Mares were given 150 mg of porcine
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Superovulation FSH for 6.8 days and only 1.6 ovulations on average were obtained, compared with 2.2 ovulations for mares receiving EPE. Fortune and Gimmich10 evaluated various doses of porcine FSH for induction of multiple ovulations in mares. Mares were given 8, 16, or 32 mg of porcine FSH twice daily for approximately 6 days. The ovulation rate was 1.5–1.8 and the percentage of mares with double ovulations was 50–83%. Embryo recovery was not attempted in this particular study. Krekeler et al.11 injected 10 or 25 mg of porcine FSH (Folltropin, Bioniche, Bellville, CA) to evaluate superovulation. Both doses of porcine FSH increased ovulation rates (2.3, 2.3; 25 and 10 mg, respectively) above controls (1.1). However, only the 25-mg dose improved embryo recovery (1.3, 25 mg; 0.7, 10 mg). Further studies are needed to evaluate this product for superovulating mares.
Equine pituitary extract The majority of studies on superovulation in the mare have utilized EPE prepared according to the procedures of Braselton and McShan12 and later by Guillou and Combarnous.13 Many of the early studies were done in Wisconsin on pony mares, which have a low incidence of spontaneous natural double ovulations. Douglas14 reported a 2.3 ovulation rate for pony mares given EPE once daily for 7 days during anestrus compared with one ovulation per mare for untreated controls. Woods and Ginther15 reported an average of three ovulations per mare in 111 cycling mares treated with EPE. Seventy percent of the mares experienced two or more ovulations. A series of studies were conducted at Colorado State University during the mid- to late-1980s on light-horse mares using EPE. Squires et al.16 reported 3.8 ovulations for EPE-treated mares and two embryos recovered per mare. This compared with 1.2 ovulations and 0.65 embryos per cycle from untreated control mares. The embryo recovery per ovulation was similar between treated and control mares. Summarized over numerous studies conducted at Colorado State University, the average ovulation rate after EPE treatment ranged between 2.3 and 3.9 ovulations per mare. Of 170 mares treated with EPE, an average of 3.2 ovulations was detected and 1.96 embryos were recovered per mare compared with 0.65 embryos recovered from untreated control mares. EPE was primarily used in research animals since there was no commercially available product that could be utilized in client-owned mares. This was problematic since there was minimal quality control in the preparation of EPE and the potency of EPE varied tremendously depending on the laboratory, source of pituitaries, and season during which the pituitaries were obtained. Although the majority of
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mares injected with EPE twice daily had no sideeffects, there was a low incidence of injection-site reactions, as well as an occasional incidence of colic. Thus, in 2003, the procedures for preparation of EPE were given to Bioniche Animal Health, which produces a more purified, standardized pituitary extract that contains a ratio of FSH to luteinizing hormone (LH) of 10:1. The majority of this chapter will deal with studies that have been performed with this commercial product. Unfortunately, this product was distributed only in the USA.
Equine follicle-stimulating hormone The initial study with equine follicle-stimulating hormone (eFSH) was conducted by Niswender et al.17 Forty-nine normally cycling mares were used to evaluate one of four treatments: (i) control, (ii) eFSH – 25 mg twice daily plus GnRH agonist (deslorelin, Ovuplant 2.1 mg), (iii) eFSH – 12 mg twice daily plus human chorionic gonadotropin (hCG) 2500 IU, and (iv) eFSH – 12 mg twice daily plus deslorelin. Treatment was initiated beginning 5 or 6 days after ovulation. Mares received 250 µg of cloprostenol on the second day of eFSH treatment. Administration of eFSH continued until the majority of follicles reached a diameter of 35 mm, at which time either a deslorelin implant (Ovuplant, Fort Dodge Animal Health, Fort Dodge, IA) or hCG (Chorulon, Intervet, Millsboro, DE) was administered. Mares in all groups were bred with semen from one of four stallions and pregnancy status was determined 14–16 days after ovulation. The number of pre-ovulatory follicles (follicles <35 mm) was greater for all eFSH-treated groups, as well as the number of ovulations. Twice daily injections of 25 mg of eFSH resulted in a greater number of pre-ovulatory follicles (6.7) than the other two eFSH groups (3.4 and 3.8). However, the number of pregnancies was similar among controls and the 25 mg eFSH group. Mares administered 12 mg eFSH twice daily followed by hCG resulted in 1.8 pregnancies compared with 0.6 for controls. Administration of hCG appeared to provide a better ovulation response than administration of deslorelin for mares given the 12 mg of eFSH twice daily. Based on this study, administration of 12 mg eFSH twice daily followed by hCG was used for a subsequent study conducted on 16 mares in Brazil. On the first cycle, mares served as controls, and, on the second cycle, mares were administered 12 mg of eFSH twice daily until a majority of follicles were 35 mm in diameter, at which time hCG was administered. Mares were inseminated on both cycles and embryos collected 7 or 8 days after ovulation. The number of ovulations during the cycle on which mares were treated with eFSH was 3.6 compared with 1.0 for
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control cycles. The average number of embryos recovered per mare for the eFSH cycle (1.9) was greater than the embryo recovery rate for the control cycle (0.5). This study demonstrated that eFSH could be used to enhance embryo recovery nearly fourfold.
Factors affecting response to eFSH Frequency of injection In the first report of successful superovulation in mares EPE was administered twice daily in seasonally anovulatory mares,18 but a similar response was also noted using EPE once daily. Scoggin et al.19 conducted a series of experiments to improve the ovarian response to EPE. They evaluated the effect of increasing the frequency or dose of EPE treatment on the ovarian response. Mares received 25 mg of EPE once daily, 50 mg EPE once daily, 12.5 mg EPE twice daily, or 25 mg EPE twice daily. All mares began EPE treatment 5 days after detection of ovulation. More ovulations were detected from mares receiving 25 mg of EPE twice daily than from those receiving either 25 mg of EPE once daily or 12.5 mg of EPE twice daily. However, the embryo recovery rate per mare was greater in the mares that received 12.5 mg of EPE twice daily than in those that received 25 mg of EPE once daily. It was concluded that increasing the frequency of EPE treatment, but not necessarily the dose, improved embryo collection per mare and per ovulation. Welch et al.20 utilized 40 cycling mares to evaluate the effect of once versus twice daily injection of eFSH. The ovulation rate was higher in mares administered 12.5 mg of eFSH twice daily than in mares administered 25 mg once a day. Based on these studies most superovulatory protocols incorporate the use of equine FSH twice daily. It may be that in the future equine FSH could be suspended in a slow release formulation that would allow administration of eFSH only once per day. Purification of eFSH The original EPE had a much higher concentration of LH than the more purified commercial eFSH product. The eFSH product contains approximately 10 parts FSH to one part LH. There was some suggestion that the more highly purified the FSH, the less the ovulatory response. Therefore, Welch et al.20 compared the standard 12 mg of eFSH twice daily with a treatment in which 12.5 mg of eFSH was given twice daily for 3.5 days and then mares were subsequently given a non-commercial preparation containing a greater amount of LH (5:1 ratio of FSH to LH) at a dose of 12.5 mg twice per day until half the cohort of follicles was greater than 35 mm. The number
of pre-ovulatory follicles was less for the group that contained the addition of the non-commercial product compared with eFSH (3 vs 4.1 follicles >35 mm). However, the ovulation rate was similar between the two groups, as well as the embryo recovery rate. It would appear that the ratio of FSH to LH is not nearly as critical in the horse as it might be in the cow and that a good ovulatory response is possible with a ratio of 10:1 to 5:1 (FSH to LH). Coasting Coasting is defined as a period of no gonadotropin administration after initial stimulation and before administration of an agent to induce ovulation. The benefits of coasting are, theoretically, to prevent ovarian hyperstimulation, limit the occurrence of large anovulatory follicles, and shorten the treatment duration, thereby decreasing the cost of a superovulation cycle. Coasting has been attempted in various species. D’Occhio et al.21 first proposed the idea of delaying the endogenous LH surge of superovulated cows as a means to improve the ovulatory response. The pre-ovulatory surge was blocked with a GnRH agonist and ovulation was induced by injection of LH. Coasting protocols have also been used in women in an attempt to minimize ovarian hyperstimulation.22 Welch et al.20 conducted a study to determine the effect of follicular coasting before administration of hCG. They reported a higher (P < 0.05) embryo recovery per mare for those mares that received 12.5 mg of eFSH twice daily in which hCG was delayed 42–50 hours after the last eFSH treatment than for those that were given hCG once follicles were 35 mm in size following eFSH treatment. Timing of eFSH treatment Ultrasonography studies have demonstrated that the majority of mares have one follicular wave during the diestrous phase of the cycle.23 During the follicular wave a cohort of follicles are initiated and grow at a similar rate. The first follicle to acquire a size of approximately 23 mm becomes dominant and continues to grow, while subordinate follicles have a slower growth rate and eventually regress. If mares are experiencing a one-wave cycle, then the dominant follicle will go on to ovulate, but, during a twowave cycle, the dominant follicle in the first wave becomes anovulatory and eventually regresses and a second cohort of follicles is initiated later in the cycle. An FSH surge occurs at the beginning of each follicular wave and peaks about the time follicles are 13.5 mm in size. The dominant follicle generally has the advantage of being slightly larger during the common growth phase and, once the FSH peaks and
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Superovulation declines, it continues to develop. Subordinate follicles are smaller and cannot continue to develop as FSH levels decline. The dominant follicle also eventually acquires LH receptors, which makes it responsive to rising levels of LH. As with any species, exogenous FSH should begin prior to the formation of the dominant follicle. Administering exogenous FSH causes circulating FSH levels to remain high and therefore the subordinate follicles can continue to grow. One of the biggest problems in horses is determining when a follicular wave has been initiated. In general there are two methods to make this determination. One is by using ultrasound on a daily basis in early diestrus to identify the beginning of the follicular wave and assess follicular development each day to determine when to start exogenous FSH. The alternative is to use a technique to induce a follicular wave. In cattle, the most common method of inducing a follicular wave is administration of estrogen, followed by insertion of a progesterone vaginal device. Approximately 3–4 days after administration of estrogen, a new follicular wave is initiated and one can schedule the FSH treatment.24 Several studies have been conducted to determine whether progesterone and estradiol can be used to synchronize follicular waves in mares and to simplify the onset of eFSH treatment. Logan et al.25 examined the effect of pretreatment with progesterone and estradiol on subsequent follicular response to eFSH. Mares were pretreated with progesterone and estradiol (P and E; 150 mg progesterone and 10 mg of estradiol-17β intramuscularly (i.m.)) once daily for 10 days beginning 5–10 days after ovulation. Prostaglandin was administered on the final day of P and E treatment. Mares were examined by transrectal ultrasonography every other day beginning 3–5 days after the end of P and E treatment. Treatment with eFSH was initiated when the largest follicle was 20–25 mm in diameter. Pretreatment with progesterone and estradiol decreased the number of preovulatory follicles, number of ovulations, and number of embryos per flush compared with mares not pretreated with progesterone and estradiol and given eFSH. Raz et al.26 administered progesterone and estradiol (150 mg of progesterone and 10 mg of estradiol) once daily for 10 days followed by prostaglandin injection on the last day of P and E treatment. In controls, an injection of prostaglandin was administered on day 5 after ovulation. In both groups when a follicle >20 mm in diameter was detected, eFSH was administered twice daily until one or more follicles was ≥35 mm in diameter. The P and E-treated mares had a longer interval from prostaglandin to ovulation than the prostaglandin-treated mares. The mean number of ovulations tended to be higher in
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the prostaglandin-treated group than in the P and Etreated group of mares. Based on these two studies pretreatment with progesterone and estradiol prior to eFSH had no advantage. Further studies are needed to determine whether exogenous estradiol can effectively suppress FSH in the mare and initiate a new follicular wave. Other ways of controlling follicular development in the mare include aspiration of the dominant follicle and/or the use of GnRH agonist for suppression of follicular development. Aspiration of the dominant follicle in cattle results in initiation of a new follicular wave in approximately 2–3 days.24 Aspiration of the dominant follicle with ultrasound has been conducted in the horse and has been shown to be effective in eliminating the dominant follicle and allowing subordinate follicles to continue.27 Unfortunately, in both cattle and horses aspiration of the dominant follicle with a transvaginal ultrasound probe is not practical for a practice setting. Administration of a GnRH agonist has been successfully used in both humans and cattle to prevent spontaneous LH surges and the recovery of multiple oocytes and embryos following ovarian hyperstimulation with FSH. In the horse, studies have shown decreased pituitary gonadotropin secretion in mares treated with a GnRH agonist implant (Ovuplant, deslorelin acetate, 2.1 mg/dose)28 and suppression of follicular growth in mares. Scoggin et al.19 evaluated pretreating mares with a GnRH agonist before EPE stimulation. Twenty normally cycling mares were randomly assigned to one of two treatments. Group 1 mares were allowed to ovulate spontaneously and were subsequently administered 25 mg of EPE twice daily beginning 5 days after ovulation. Group 2 mares received two doses of GnRH agonist (Ovuplant) prior to ovulation and 25 mg of EPE twice daily beginning 5 days post-ovulation. After EPE treatment, a greater number of follicles >35 mm were detected for mares in the EPE-only group than for mares in the GnRH agonist group. Treatment with EPE was longer for mares in the GnRH agonist group than in those in the EPE-only group. Unfortunately, these mares were not bred and therefore no embryo collection attempts were performed. Based on follicular data obtained in this study, pretreatment with GnRH agonist did not improve the superovulatory response with EPE. Follicular activity was suppressed in mares following treatment with two doses of Ovuplant, even during EPE administration.
Alternative protocols Because of the expense of eFSH, protocols that utilize a lower dose of FSH or fewer injections of FSH have been evaluated. In cows with spontaneous double
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ovulations, FSH levels are higher prior to the time of selection of the dominant follicle than in singleovulating cows.29 It has been suggested that exogenous FSH given immediately prior to selection of the dominant follicle would result in a higher incidence of double ovulations. McCue et al.30 determined whether short-term (3 day) treatment with eFSH initiated prior to the selection of the dominant follicle would result in similar ovulation and embryo recovery rates to those of the standard eFSH protocol. In 2005, mares were assigned to one of three treatment groups: (i) untreated controls, (ii) standard eFSH treatment, and (iii) 3-day eFSH treatment. Beginning 5–7 days after ovulation, group 2 mares were examined daily to assess follicular size. Once one or more follicles 20–25 mm in size were detected, 12.5 mg of eFSH was administered twice daily. On the second day of treatment mares were given prostaglandin F2 α (PGF2 α) to induce luteolysis. Daily treatment with eFSH was continued until the majority of follicles were ≥35 mm. Thirty-six hours after the last eFSH treatment mares were given 2500 IU of hCG and were inseminated. Group 3 mares were handled identically to those in group 2, except eFSH was given for only 3 days beginning when one or more follicles were 20–25 mm in diameter. In 2006, these same treatments were repeated. In the first experiment (2005) the standard eFSH treatment resulted in a greater number of pre-ovulatory follicles, ovulations, and embryos per flush than did the 3-day treatment. In contrast, in 2006, the standard treatment resulted in a similar number of preovulatory follicles and ovulations to the 3-day treatment. However, in both 2005 and 2006, the standard treatment resulted in more embryos per flush than the 3-day treatment. The number of days mares were treated in the standard eFSH treatment was 4.9 days in 2005 and 3.5 days in 2006. This was due to the technician waiting longer to initiate eFSH treatment in 2006. Thus, it was difficult to show a difference between the standard and 3-day treatments in 2006, since the number of days treated was 3.5 and 3.0 days, respectively. By waiting until the largest follicles approach 25 mm, one is able to decrease the number of days of injections to approximately 4, which significantly lowers the cost of superovulation of mares. One other method of decreasing the cost of superovulation would be to use a decreasing dose of administration, similar to the technique used in cattle, or giving mares approximately half the recommended dose. McCue et al.30 compared a step-down protocol of eFSH with the standard equine FSH protocol. The standard eFSH protocol was 12.5 mg of eFSH twice daily until greater than 50% of the developing follicles reached 35 mm in diameter. Treat-
ment was ceased and all mares were given recombinant LH or hCG 24 hours later. For the step-down protocol mares were given 12.5 mg of eFSH twice a day on the first day of treatment, 8 mg twice a day on day 2, and 4 mg twice a day on day 3. The ovulatory agent was given as described for the standard protocol. The standard eFSH protocol resulted in a greater number of pre-ovulatory follicles and ovulations, and tended to increase the number of embryos per flush compared with mares given the step-down protocol. The step-down protocol provided a similar number of embryos per flush to the controls, whereas embryo recovery for the standard eFSH protocol was significantly greater than that for controls. The standard protocol used in the experiments reported by McCue et al.30 differed in relation to the timing of prostaglandin administration. In the 2005 experiment,30 prostaglandin was given on the second day of eFSH treatment, whereas in experiment 2 (2006) prostaglandin was given between days 5 and 7 post-ovulation and eFSH treatment was initiated once mares developed follicles 22–25 mm in diameter. It would appear that delaying administration of prostaglandins until day 2 of eFSH treatment may have resulted in greater follicular stimulation and a greater number of ovulations. Giving prostaglandins and allowing the mare to come into heat prior to initiation of eFSH treatment was not as efficacious as starting the eFSH during diestrus and giving prostaglandins on day 2 of the eFSH treatment. This concurs with some of the earlier studies in which initiation of EPE treatment late in the estrous cycle was not as efficacious as starting eFSH treatment early in the cycle. Once prostaglandin is administered and the mare returns to estrus LH is no longer suppressed by progesterone and LH begins to rise during the time of eFSH treatment. Thus, the best protocol would certainly be to administer prostaglandins on the second day of eFSH treatment during the time that a mare has a fully formed, functional corpus luteum. Similar to cattle, individual mares have a varying sensitivity to eFSH. Some mares given a 12.5 mg dose of eFSH overstimulate because of the increased sensitivity to this exogenous hormone. It is likely that, similar to cattle, practitioners that routinely superovulate the same mare will very quickly determine the appropriate dose for that individual mare. Logan et al.25 demonstrated that a similar ovarian response could be obtained by administering 6.25 mg of eFSH twice a day to that obtained by administering the standard 12.5 mg of eFSH twice a day. Although the number of ovulations was higher for mares given the standard dose of eFSH, the number of embryos recovered per flush was similar between those given 6.25 mg of eFSH and those given 12.5 mg of eFSH.
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Ovulatory agent In order for the majority of ovulations to occur on one day, generally an ovulatory agent, such as hCG, deslorelin, or recombinant LH (reLH), is given to induce synchronous ovulation. This simplifies the scheduling of embryo recovery and management of recipient mares. Unfortunately, no recent studies have been reported in which an ovulatory agent was not given. Thus, most of the data concern comparison of various ovulatory agents for induction of ovulation. In the study by Niswender et al.17 it appeared that hCG was more efficacious in inducing ovulation in superovulated mares than the GnRH agonist deslorelin. Although based on a small number of mares (n = 5 per group) the number of ovulations per mare and number of pregnancies per mare were significantly higher for mares given eFSH followed by hCG than those given eFSH followed by deslorelin. Based on this study, additional studies done at Colorado State University have generally used hCG as the ovulatory agent. Obviously, additional studies are needed to determine whether there is a true difference in ovulatory response between deslorelin and hCG. Recently, a new recombinant equine LH was made available to the equine practitioner (EquiPure LH, AspenBio, Castle Rock, CO). McCue et al.30 compared the efficacy of hCG and recombinant eLH (reLH). Mares were assigned to one of four groups: (i) control, (ii) standard eFSH protocol (12.5 mg twice a day (b.i.d.)), (iii) 3-day protocol, and (iv) step-down protocol. Mares were given either reLH (150 µg) or hCG (2500 IU) 24 hours after cessation of treatment. The percentage of control mares ovulating within 48 hours in response to hCG (88%) or reLH (100%) was similar. In contrast, a higher (P < 0.05) percentage of eFSH-treated mares ovulated within 48 hours in response to reLH (92%) than to hCG (71%). Unfortunately, the recombinant LH is not available at the present time, and thus hCG would appear to be the hormone of choice for induction of ovulation in superovulated mares. The one exception would be mares that have received numerous injections of hCG during a given breeding season and mares that are older than 16 years (E.M. Carnevale, unpublished observations). Both of these categories of mares have been shown to be less responsive to hCG administration.
Recombinant FSH Although eFSH appears to be a very effective hormone for superovulation in mares, the raw material (pituitary glands) is not readily available since all of the slaughterhouses were closed within the USA.
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In addition, gonadotropin extraction is labor intensive and, therefore, expensive, and the biosecurity of crude pituitary extract is a cause for concern. The purification procedure for eFSH preserves glycoprotein hormone activity but probably has a limited capacity for destroying viruses and prion proteins. In addition, since this is a pituitary extract, the composition of each batch of eFSH may vary in respect to the ratio of FSH to LH activity and overall potency. Initial studies in horses that examined whether further purification of EPE to increase the FSH:LH ratio would improve the magnitude or consistency of the superovulatory response did not indicate any significant advantage.31, 32 More recently Briant et al.33 found that an FSH-enriched pituitary extract resulted in failed or disturbed ovulations. They suggested that the exact FSH:LH ratio needed for maximum superovulation in the mare is not truly known. In a recombinant FSH preparation, such as recombinant human FSH, the amount of LH present is extremely minimal. Recombinant FSH is commonly used in human in vitro fertilization programs. Recently, the effect of recombinant human FSH on follicular development and embryo production in mares has been reported.34 Five mares were treated twice daily with 400 IU of recombinant human FSH starting on day 6 after ovulation coincident with PGF2 α analog administration. Five control mares were treated similarly but with saline instead of FSH. When the dominant follicle exceeded 35 mm, ovulation was induced with hCG and embryos were recovered 7 days after ovulation. Unfortunately, with the dose of human recombinant FSH used in this study, there was no stimulatory effect on follicular development and ovulation. Niswender et al.35 evaluated the superovulatory response of mares administered recombinant equine FSH. The FSH was produced in a similar way to the human FSH using novel single-chain recombinant equine gonadotropin production.36 Thirteen mares were randomly assigned to one of three groups and follicular development post-ovulation was suppressed with 14 days of treatment with 150 mg of progesterone and 10 mg of estradiol in oil. To induce luteolysis, mares received PGF2 α on the sixth day of P and E treatment. During the last 7 days of P and E treatment mares were treated with either 850 or 500 µg of recombinant FSH or saline. When a 35mm follicle was present, mares received 750 µg of recombinant LH to induce ovulation. Treatment with 850 µg and 500 µg of recombinant FSH resulted in follicular growth 5.6 and 4.4 days earlier than controls. The mean number of follicles 20–29 mm was 5.6 after 850 µg of recombinant FSH and 4.8 after 500 µg of recombinant FSH, which was significantly greater than controls. This demonstrated that the recombinant
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FSH was effective in stimulating follicular development. The number of ovulations detected for the three groups was 2.8, 3.0, and 2.0, respectively. In a second experiment 20 cycling mares were randomly assigned to one of four treatment groups and administered 850 µg of recombinant FSH twice daily, 850 µg recombinant FSH once daily, 500 µg recombinant FSH twice daily, or 12.5 mg of eFSH twice daily (positive control). Mares received 1.5 mg of recombinant LH intravenously to induce ovulation about 38 hours after the last FSH treatment. The number of pre-ovulatory follicles and the number of ovulations was similar among groups, with all FSHtreated mares having a greater number of ovulations than controls. Of all parameters measured, values for the recombinant FSH treatment group were similar to the positive control group (eFSH). They concluded that recombinant FSH was effective in stimulating multiple pre-ovulatory follicles and multiple ovulations. In 2007, an unpublished study was conducted at Colorado State University to further compare recombinant FSH with eFSH for induction of multiple ovulation in cycling mares. This work was based on the preliminary data available at that time. The response to recombinant FSH in our study was similar to that of Niswender et al.35 That is, the number of pre-ovulatory follicles, number of ovulations, and embryo recovery rate were similar in mares administered recombinant FSH to those receiving eFSH. Once this product becomes commercially available, it may serve as an alternative to the use of EPE.
Challenges One of the main challenges in superovulation is the extreme variability in number of ovulations and number of embryos recovered from superovulated mares. This is not unique to the horse, since the same variability appears in superovulation of cattle. The dogma in the cattle world is that 30% of donor cows provide 70% of the embryos. In addition, in cattle 25% of embryo recovery attempts result in no fertilized embryos being recovered. This is similar to the 25–30% negative flushes that occur in the mare.37 However, it is not uncommon for mares to provide an average of three or four embryos per flush, and some mares have provided as many as 10 embryos in a given flush. In a retrospective study Squires et al.37 examined some of the factors affecting response to administration of eFSH. Data from 40 mares that were treated with eFSH during two estrous cycles in 2004 and 40 mares that were treated with eFSH on two to six cycles in 2005 were examined. As expected, the
correlation between the number of pre-ovulatory follicles and the number of ovulations was high. Unfortunately, the correlation between the number of ovulations and the number of embryos was not nearly as high. Mares in this study were retrospectively divided into those that were considered to be good donors (provided an average of ≥1.5 embryos per flush) and mares considered to be poor donors (provided an average of <1.5 embryos per flush). Mares that were good donors had a greater number of follicles in the category of 11–15 mm and 16–20 mm at the onset of treatment. Also mares in the good donor category had more pre-ovulatory follicles and more ovulations. These characteristics could be used to select mares that are more likely to respond to eFSH treatment. One of the other major challenges of superovulation is the formation of anovulatory follicles after administration of eFSH. This occurs in approximately 10% of treated cycles. If some follicles ovulate and others either luteinize or fail to ovulate, embryo recovery is going to be compromised. Obviously, if all follicles fail to ovulate, then no embryos will be recovered and one cannot recover the expense of the eFSH. Many times it is difficult to determine whether superstimulated follicles will ovulate, luteinize, or become hemorrhagic. The cause of these anovulatory follicles has not been determined. However, Ginther et al.38 suggested that the only difference between a follicle that ovulates and one that fails to ovulate is a higher level of estrogen in those follicles that fail to ovulate. One other suggestion by Briant et al.33 was that, when too high a dose of eFSH is administered, the LH content in the eFSH product results in increased progesterone levels prior to ovulation which may be responsible for inducing anovulatory follicles. Lastly one of the major challenges to superovulation does not concern the mare, eFSH, or physiology, but is due to the attitude of the breeder. Unlike cattle, in which a donor cow can be bred to one valuable bull and an average of six transferable embryos obtained, resulting in high satisfaction of the owner of the donor cow, in mares generally the breeder wants to obtain one embryo from a specific genetic combination and then switch stallions in order to obtain an embryo from a second genetic combination. Therefore, breeders are reluctant to superovulate mares and obtain multiple embryos which, if transferred successfully, would require that multiple stud fees be paid for a specific genetic combination. Equine breeders seem to be satisfied with a 50–60% embryo recovery rate per flush and generally are willing to have the mare rebred and flushed additional times during a breeding season in order to get the right genetic combinations for the embryos.
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Future studies Some of the challenges that need to be addressed in future studies are as follows. First, developing methods of initiating follicular waves in the mare so that daily ultrasound examinations are not needed in order to initiate treatment. Second, minimizing or preventing the occurrence of anovulatory follicles. Third, investigating the proper timing of prostaglandin administration during an eFSH treatment regime.
Conclusions It is quite clear that equine FSH can be used to successfully superovulate mares and improve embryo recovery. The standard protocol that is now being recommended is to begin examining the mare with ultrasound 5–7 days after ovulation. Once the mare obtains multiple follicles that are 20–24 mm in size, then eFSH treatment is initiated (12.5 mg twice daily). Prostaglandin is administered on the second day of eFSH treatment. Equine FSH treatment is continued twice daily until the majority of developing follicles are 32–35 mm in size. Equine FSH treatment is then discontinued and hCG given 24–36 hours later. On the average with this treatment approximately three or four ovulations will be obtained and 2–2.5 embryos per flush will be recovered. Based on studies done at Colorado State University during the past 5 years with eFSH the quality of embryos recovered from superovulated mares is the same as those from non-stimulated mares. In addition, pregnancy rates for embryos from superovulated mares appear to be identical to those from untreated controls.
References 1. McCue PM. Superovulation. Vet Clin North Am Equine Pract 1996;12:1–11. 2. Squires EL, McCue PM. Superovulation in mares. Anim Reprod Sci 2007;99:1–8. 3. Allen WR. Embryo transfer in the horse. In: Adams CE (ed.) Mammalian Egg Transfer. Boca Raton, FL: CRC Press, 1982; pp. 135–54. 4. Ginther OJ, Bergfelt DR. Effect of GnRH treatment during the anovulatory season on multiple ovulation rate and on follicular development during the ensuing pregnancy in mares. J Reprod Fertil 1990;88:119. 5. Squires EL, Rowley HR, Nett TM. Comparison of pulsatile versus constant release of GnRH for superovulation in mares. Proc Am Soc Anim Sci 1989;67 Suppl. 1:391–2. 6. Ginther OJ, Beg MA, Bergfelt DR, Donadeu FX, Kot K. Follicular selection in monovular species. Biol Reprod 2001;65:638–47. 7. McKinnon AO, Brown RW, Pashen RL. Increased ovulation rate in mares after immunization against recombinant bovine inhibin-alpha-subunit. Equine Vet J 1992;24:144–6.
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8. McCue PM, Hughes JP, Lashley BL. Effect on ovulation rate of passive immunization of mares against inhibin. Equine Vet J 1993;15 Suppl.:103–6. 9. Squires EL, Garcia RH, Ginther OJ, Voss JL, Seidel Jr GE. Comparison of equine pituitary extract and follicle stimulating hormone for superovulating mares. Theriogenology 1986;26:661–70. 10. Fortune JE, Gimmich TL. Purified pig FSH increases the rate of double ovulation in mares. Equine Vet J 1993;15 Suppl.:95. 11. Krekeler N, Hollinshead FK, Fortier LA, Volkmann DH. Improved ovulation and embryo recovery rates in mares treated with porcine FSH. Theriogenology 2006;66:663–87. 12. Braselton WE, McShan WH. Purification and properties of follicle stimulating and luteinizing hormone from horse pituitary glands. Arch Biochem Biophys 1970;139:45–8. 13. Guillou F, Combarnous Y. Purification of equine gonadotropins and comparative study of the acid-dissociation and receptor-binding specificity. Biochim Biophys Acta 1983;755:229–36. 14. Douglas RH. Review of induction of superovulation and embryo transfer in the equine. Theriogenology 1979;11:33–46. 15. Woods GL, Ginther OJ. Induction of multiple ovulation during the ovulatory season in mares. Theriogenology 1983;20:347–55. 16. Squires EL, McKinnon AO, Carnevale EM, Morris RP, Nett TM. Reproductive characteristics of spontaneous single and double ovulating mares and superovulated mares. J Reprod Fertil 1987;35 Suppl.:399–403. 17. Niswender KD, Alvarenga MA, McCue PM, Hardy QP, Squires EL. Superovulation in cycling mares using equine follicle stimulating hormone (eFSH). J Equine Vet Sci 2003;23:497–500. 18. Douglas RH, Nuti L, Ginther OJ. Induction of ovulation and multiple ovulation in seasonally anovulatory mares with equine pituitary fractions. Theriogenology 1974;2:133–42. 19. Scoggin CF, Meira C, McCue PM, Carnevale EM, Nett TM, Squires EL. Strategies to improve the ovarian response to equine pituitary extract in cyclic mares. Theriogenology 2002;58:151–64. 20. Welch SA, Denniston DJ, Hudson JJ, Bruemmer JE, McCue PM, Squires EL. Exogenous eFSH, follicle coasting, and hCG as a novel superovulation regimen in mares. J Equine Vet Sci 2006;26:262–70. 21. D’Occhio MJ, Sudha G, Jillella D, Whyte T, Maclellan LJ, Walsh J, Trigg TE, Miller D. Use of GnRH agonist to prevent the endogenous LH surge and injection of exogenous LH to induce ovulation in heifers superstimulated with FSH: a new model for superovulation. Theriogenology 1997;47:601–13. 22. Flucker MR, Hooper WM, Yuzpe AA. Withholding gonadotropin (coasting) to minimize the risk of ovarian hyperstimulation during superovulation and in vitro fertilization – embryo transfer cycles. Fertil Steril 1999;71:294– 301. 23. Ginther OJ, Gastal EL, Gastal MO, Bergfelt DR, Baerwald AR, Pierson RA. Comparative study of the dynamics of follicular waves in mares and women. Biol Reprod 2004;71:1195–201. 24. Bo GA, Guerrero DC, Adams GP. Alternative approaches to setting up donor cows for superovulation. Theriogenology 2008;69:81–7. 25. Logan NL, McCue PM, Alonso MA, Squires EL. Evaluation of three equine FSH superovulation protocols in mares. Anim Reprod Sci 2007;102:48–55.
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26. Raz T, Carley S, Green J, Corrigan M, Card C. Effect of prostaglandin in early diestrus or progesterone and estradiol administration on equine FSH-treated donor mare embryo recovery and recipient pregnancy rate. In: Proceedings of the 51st American Association of Equine Practitioners, 2005, pp. 204–8. 27. Dippert KD. Follicular development and superovulation in mares. PhD dissertation, Colorado State University, Fort Collins, CO, 1995. 28. Farquhar VJ, McCue PM, Nett TM, Squires EL. Effect of deslorelin acetate on gonadotropin secretion and ovarian follicle development in cycling mares. J Am Vet Med Assoc 2001;218:749–52. 29. Lopes H, Satori R, Wiltbank MC. Reproductive hormones and follicular growth during one or multiple dominant follicles in cattle. Biol Reprod 2005;72:788–95. 30. McCue PM, Patten M, Denniston DD, Bruemmer JE, Squires EL. Strategies for using eFSH for superovulating mares. J Equine Vet Sci 2008;28:91–6. 31. Hofferer S, Lecompte F, Magallon T, Palmer E, Combarnous Y. Induction of ovulation and superovulation in mares using equine LH and FSH separated by hydrophobic interaction chromatography. J Reprod Fertil 1993;98:597–602. 32. Rosas CA, Alberio RH, Baranao JL, Aguero A, Chaves MG. Evaluation of two treatments in superovulation of mares. Theriogenology 1998;49:1257–64.
33. Briant C, Toutain PL, Ottogalli M, Magallon T, Guillaume D. Kinetic studies and production rates of equine FSH in ovariectomized pony mares: Application to the determination of a dosage regime for equine FSH in a superovulation treatment. J Endocrinol 2004;182:43–54. 34. Tharasanit T, Colenbrander B, Bevers MM, Stout TAE. Effects of recombinant human follicle stimulating hormone on follicle development and ovulation in the mare. Theriogenology 2006;65:1071–81. 35. Niswender KD, Jennings M, Boime I, Colgin M, Roser JF. In vivo activity of recombinant equine follicle stimulating hormone in cycling mares. In: Proceedings of the 53rd American Association of Equine Practitioners, 2007, pp. 561–2. 36. Yoon MJ, Boime I, Colgin M, Niswender KD, King SS, Alvarenga M, Jablonka-Shariff A, Pearl CA, Roser JF. The efficacy of a single chain recombinant equine luteinizing hormone (reLH) in mares: Induction of ovulation, hormone profiles and interovulatory intervals. Dom Anim Endocrinol 2007;33:470–9. 37. Squires EL, Logan N, Welch S, McCue PM. Factors affecting the response to administration of equine FSH. Anim Reprod Sci 2006;94:408–10. 38. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Incidence, endocrinology, vascularity and morphology of hemorrhagic anovulatory follicles in mares. J Equine Vet Sci 2007; 27:130–9.
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Chapter 194
Estrus Suppression Dirk K. Vanderwall and Gary J. Nie
Mares are long-day, seasonally polyestrous animals that undergo cyclic reproductive activity during the spring and summer months of the ovulatory season. During each estrous cycle of approximately 21– 22 days, behavioral signs of estrus generally last for 5 to 7 days, during which time estrogen is elevated and progesterone is at basal levels. Within 24 to 48 hours after ovulation, progesterone levels have increased sufficiently to block behavioral signs of estrus for about a 2-week diestrus phase of the cycle. In addition to the cyclical pattern of estrous behavior displayed during the ovulatory season, mares also exhibit estrous behavior during the spring ‘transitional’ period that marks the annual recrudescence of hypothalamic-pituitary-ovarian activity at the end of the winter anovulatory season. During the transitional period there is a progressive increase in ovarian follicular activity that ultimately leads to the development of one to three anovulatory follicular waves typified by the formation of a large (> 38-mm) dominant follicle that fails to ovulate due to insufficient levels of luteinizing hormone (LH).1, 2 Behaviorally, the spring transition period is characterized by erratic and/or prolonged periods of estrous behavior associated with the progressive increase in ovarian follicular activity.2, 3 Once LH levels have increased sufficiently,1, 4 the first ovulation of the year occurs, which by definition marks the beginning of the ovulatory season. Mares are unique among domestic animals, because in addition to the ovarian-derived estrogeninduced signs of estrous behavior described above, many seasonally anovulatory (and ovariectomized) mares exhibit paradoxical estrous behavior associated with hormone secretion from the adrenal cortex.5, 6 The intensity of this type of ‘unseasonable’ estrous behavior was judged to be equivalent to the behavior that intact cycling mares display during the initial and terminal days of estrus, but less intense than the behavior displayed near ovulation.5 Such
behavioral receptivity to a stallion outside the breeding/ovulatory season that is independent of ovarian estrogen secretion may have developed as a means of maintaining social bonds between a harem stallion and his mares.5, 7 This phenomenon has important implications for the clinical management of estrous behavior in mares, since simply suppressing ovarian follicular activity (or removing the ovaries) and its attendant estrogen production may not ensure the elimination of estrous behavior.
Indications for suppressing estrus Owners, trainers, managers, and veterinarians, from time to time, may feel it is necessary to suppress behavioral signs of estrus in their mares. The primary indications for suppressing estrous behavior include cycle-related behavior/performance problems and pain/colic during estrus. Based on a survey completed by over 750 veterinarians, Jorgensen et al.8 reported that approximately 90% of the veterinarians had the clinical impression that the estrous cycle impacted the performance of mares. The most frequently reported clinical sign associated with an effect of the estrous cycle on performance was attitude change, while other signs included tail swishing, difficulty to train, squealing, ‘horsing’, excess urination, kicking, and a decrease in performance. It is important to note that some problematic behaviors displayed by mares that are thought to be associated with estrus are in fact not estrous behaviors, such as signs of (i) submissive behavior, (ii) urogenital discomfort, or (iii) stallion-like behavior.9, 10 Of these, submissive behavior may be most easily confused with estrous behavior. Submissive behavior includes leaning away from perceived threats, swishing/ringing the tail, and actively squirting urine, which collectively can give the impression of estrus.10 In contrast to submissive behavior, true estrous behavior includes leaning towards the stallion
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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(or other stimulus), relaxed lifting motion of the tail, stationary/squatting stance, and passive urination (full stream or small amounts in spurts).10 In an effort to evaluate the potential for an effect of stage of estrous cycle on the behavior of mares, Hedberg et al.11 conducted two behavior tests (novel object and isolation) on 12 mares, once when they were in estrus and once during diestrus in a crossover design. Five of the mares served as controls, while the other seven mares were classified as ‘problem’ mares based on their owner’s perception of estrousrelated behavioral problems. There were no significant differences in the behavioral responses between estrus and diestrus within the control and ‘problem’ groups, nor were there differences in the behavioral responses between the two groups of horses; however, as the author’s note, the sample size was small and a crossover study design may not have been the most appropriate experimental design. Therefore, further work in this area is warranted. Although it has not been conclusively documented in the horse, it is plausible that changing levels of estrogen and progesterone throughout the estrous cycle may influence non-reproductive body systems, such as the musculoskeletal system, since such effects have been observed in women throughout the menstrual cycle.12 If estrogen and progesterone have differing effects on the musculoskeletal system in mares, it is plausible that a subtle lameness could be exacerbated at specific times during the estrous cycle. Clearly, this is an area that warrants further investigation. When evaluating an owner/trainer complaint of an estrous cycle-related behavior/performance problem in a mare, the veterinarian should first determine if the problematic behavior is or is not related to a specific phase of the estrous cycle. In order to evaluate the mare thoroughly, additional expertise may be needed in the form of consultation with or referrals to behavior and/or reproduction experts. A team approach to evaluating and solving the problem may be most successful.13 There are some problem behaviors that are truly estrous cyclerelated.9, 10 For example, some mares display such intense behavioral signs of estrus that they directly interfere with training/performance. In other mares, the condition may be much more subtle, simply causing owners and trainers to report that the mare is less cooperative or attentive during estrus.14 Some mares experience pain in the periovulatory period, which ranges from (i) increased sensitivity/pain during manual transrectal evaluation of the ovary; to (ii) sensitivity to weight and/or manipulation of the back; to (iii) overt colic-like symptoms of pain.12,15–17 Of these, sensitivity during manual evaluation of the ovary is most common and does
not warrant therapy or other intervention, since it only occurs during transrectal manipulation of the ovulatory ovary. In contrast, mares that experience back pain and/or colic-like symptoms in the periovulatory period may benefit from the use of ovulatory agents to reduce the time a large pre-ovulatory follicle is present on the ovary or they may benefit from suppression of cyclical reproductive activity.
Methods of suppressing estrus A number of methods have been used to suppress behavioral signs of estrus in mares. Some methods employ approved products with well-documented efficacy, while others are not approved and their use is based entirely on anecdotal information. Each has advantages and disadvantages. The primary methods that have been used for suppression of estrous behavior include:
r r r r
administration of exogenous progesterone/ progestins; extending the duration of corpora luteal function; suppressing ovarian follicular activity; ovariectomy.
Administration of exogenous progesterone/progestins
Progesterone Daily intramuscular administration of 100 mg progesterone in oil (0.2 mg/kg) effectively suppresses signs of estrus in mares;18 however, the need for daily administration and the potential for soreness at the site of injection are limitations to its use. Progesterone in oil is available from several compounding pharmacies. It was recently demonstrated that intramuscular administration of a compounded long-acting formulation of progesterone containing a total dose of 1.5 g progesterone will maintain blood levels of progesterone above 1.0 ng/mL for approximately 10 days,19 which is a level of progesterone sufficient to block estrous behavior;18, 20 however, the potential for soreness at the injection site is a limitation to its use especially in the performance animal.
Altrenogest R Altrenogest (Regu-Mate , Intervet, Millsboro, DE) is a synthetic progestin approved for use in horses for suppressing estrus, and can probably be considered the current ‘gold standard’ for methods of suppressing estrous behavior in mares. Daily oral administration of altrenogest at a dose of 0.044 mg/kg (1 mL per 50 kg [110 lb] of bodyweight) is highly
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Estrus Suppression effective for suppressing estrus in mares.21, 22 Although altrenogest is effective for estrus suppression there are a number of disadvantages to its use. As an oil solution, it can be messy to administer, and owners often complain about the expense and inconvenience of daily dosing. Human safety must also be considered for individuals handling the solution since it is readily absorbed through the skin. In addition, it has a labeled contraindication for use in mares with uterine inflammation.
Medroxyprogesterone acetate Manufactured and marketed as a human contraceptive, medroxyprogesterone acetate (Depo-Provera, Pharmacia) suppresses ovulation in women for 3 months. Although there are anecdotal reports on the use of medroxyprogesterone acetate for estrus suppression in mares, it was recently reported that intramuscular administration of an initial dose of 1600 mg medroxyprogesterone acetate followed by 400 mg once weekly for 5 weeks did not suppress estrous behavior in mares.23 On a related note, medroxyprogesterone acetate was unable to maintain pregnancy and failed to block the return to estrus in mares following prostaglandin-induced luteolysis when 1000 mg was administered intramuscularly every 7 days.24
Hydroxyprogesterone caproate Hydroxyprogesterone caproate is a synthetic progestin available through human and veterinary pharmacies under a number of trade names, the most recognized being Hyproval (Solvay). McKinnon et al.25 demonstrated that intramuscular administration of 500 mg hydroxyprogesterone caproate every other day was unable to maintain pregnancy and failed to block the return to estrus in mares that were unilaterally ovariectomized to remove the primary corpus luteum (CL) of pregnancy. The same synthetic progestin, repackaged as hydroxyprogesterone hexanoate (Gesteron-500, Illium Veterinary Products), though labeled for pregnancy maintenance was also unable to maintain pregnancy and failed to block the return to estrus in mares when 500 mg was administered intramuscularly every 4 days following prostaglandin-induced luteolysis.24
Melengestrol acetate Melengestrol acetate (MGA, Pfizer), a synthetic progestin, is labeled as a feed additive for estrus suppression in cattle. However, when fed to mares at 10 or 20 mg/day for up to 15 days, it failed to sup-
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press estrus.18 More recently, higher doses of MGA (100 mg/day and 150 mg/day) were shown to hasten the onset of ovulation in spring transitional mares compared to untreated control mares; therefore, investigating the effect of higher doses on estrous behavior may be warranted.26
Norgestomet Norgestomet is a synthetic progestin used for estrus synchronization in ruminants that is generally administered as an implant (discussed below). There is one report describing the use of an injectable formulation of norgestomet in cottonseed oil in mares; in that study,27 intramuscular administration of 1.5 mg or 3.0 mg norgestomet daily for 15 days did not suppress estrous behavior.
Implants A variety of implants containing various progestins, labeled and marketed for use in other species, have been used in mares with the intent of suppressing estrus. The most widely used has been Synovex S (Fort Dodge Animal Health) implants. Labeled for increasing feed efficiency in steers, each dose of eight pellets contains 200 mg progesterone and 20 mg estradiol benzoate, which is slowly absorbed over months. McCue et al.28 were unable to suppress estrus or cyclicity in mares with 80 Synovex S pellets (10 times the labeled dose for steers). This is understandable, considering that Loy and Swan reported18 that 100 mg of progesterone in oil was required on a daily basis reliably to suppress estrus in mares; absorption of progesterone from Synovex S implants over extended periods will provide far less than the equivalent of 100 mg per day. Scheffrahn et al.29 reported that following subcutaneous placement of a implant containing 6.0 mg norgestomet in six mares, two of the mares displayed estrous behavior 2 days before the implants were removed 10 days after placement, although the implant was not critically tested for its ability to suppress estrous behavior. On a related note, placement of five subcutaneous implants containing a total of 15 mg norgestomet was unable to maintain pregnancy and failed to block the return to estrus in mares following prostaglandin-induced luteolysis.24 Extending corpora luteal (CL) function
Intrauterine glass marble In contrast to using exogenous progesterone/ progestins to block estrus, CL function can be extended allowing continued secretion of endogenous
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progesterone to block estrus. One method of extending CL function involves placing a sterile glass marble into the uterine lumen; Nie et al.30 reported that placement of a 25- or 35-mm sterile glass ball into the uterine body immediately after ovulation resulted in prolonged CL function in 7 of 18 (39%) mares that retained the glass marble (6 of 12 mares expelled the 25-mm marble). Luteal function was maintained for approximately 90 days before mares cycled again. Follow-up work showed that there was no detrimental effect on the uterus or fertility from the glass marble. The biggest disadvantage is that it did not appear to work in the majority of mares. In fact, the efficacy of a glass marble for extending luteal function appears only to be moderate. However, if it does work in a given mare it offers several advantages. First, it takes advantage of natural progesterone production. Second, it is inexpensive. Finally, and most importantly, the luteal function appears to last for approximately 3 months. The protocol for using a glass marble to suppress estrus in a mare is relatively simple. As with any long-term progestin protocol, the mare should be evaluated for uterine inflammation prior to treating. First, a glass marble (www.glassmarbles.com), 35 mm in diameter, is sterilized by autoclaving. Avoid breakage by autoclaving at a temperature of 121◦ C (250◦ F) and pressure of 1.1 bar (16 psi), without a pre-vacuum or dry-cycle and a slow cool-down phase. The mare is positioned in late estrus or within 24 hours after ovulation, while the cervix is relaxed. The perineum is prepared as for any clean procedure. A sterilized sleeve is donned and a small amount of sterile lubricant applied to the back of the hand. The marble is manually carried through the vagina to the caudal os of the cervix and placed in the lumen. Using a finger, the marble is pushed forward to the caudal uterine body. After removing the hand from the vagina, locate the marble per rectum and push it forward to the horn–body junction, if it has not already moved to that position. After placing the marble, the uterus can be infused with antibiotics and the mare treated with oxytocin, if there is concern about uterine contamination during the procedure. When removal of the glass marble is desired the procedure should be performed when the mare is in peak estrus. Occasionally a mare may require sedation during the removal procedure. Mares with pendulous horns present the most difficulty. Removal is accomplished by manipulating the glass marble, per rectum, caudally toward the cervix, through the cervix, and then to the vulva for retrieval. If the cervix is not fully dilated, a gloved hand is taken per vagina to the caudal cervical os and the glass marble retrieved from the lumen. It is important to note that if it is not removed, some mares may retain the glass
marble for markedly prolonged periods of time (i.e., years), such that its presence in the uterine lumen may be unbeknown to individuals working with the animal. This has led to situations in which attempts have been made to breed a mare when an intrauterine glass marble has been present, and although the presence of the glass marble will likely cause iatrogenic infertility and interfere with the establishment of pregnancy, there are anecdotal reports of mares actually becoming pregnant despite of the presence of the glass marble in the uterine lumen with subsequent compromise and loss of the pregnancy due to the presence of the glass marble. In addition, there are anecdotal reports of glass marbles that have spontaneously fragmented in the uterine lumen leaving behind glass shards that could permanently impair fertility; therefore, there is the potential for severe unintended consequences if a glass marble is not eventually removed from the uterine lumen, so their use should be carefully considered. In a recent study, Rivera del Alamo et al.31 examined the effect on the duration of CL function in mares of intrauterine placement of a 20-mm water-filled polypropylene ball, with the specific objective of investigating two potential mechanisms by which CL function is extended: (i) whether the intrauterine device induces mild endometrial inflammation that completely blocks or markedly attenuates prostaglandin (PG) F2α secretion (i.e., prevents high-magnitude luteolytic pulses); or (ii) whether the physical presence of the device (movement and/or contact with the endometrium) directly mimics the inhibitory effect of a conceptus on PGF2α secretion. Corpora luteal function was extended in 9 of 12 mares (75%), with an average duration of 57 days, compared to 0 of 12 control mares, in which the average duration of CL function was 16 days. In six of the nine mares with extended CL function, small accumulations of intrauterine fluid (≤ 10 × 20 mm) were identified during the luteal phase, but no neutrophils or bacteria were recovered on uterine swabs when they were examined during the subsequent estrus. In addition, changes in uterine biopsy scores for inflammation and glandular dilation pre- and post-treatment were similar for control and uterine device mares (with or without extended CL function); therefore, there was no evidence the intrauterine device induced an inflammatory response in the uterus. Based on intensive blood sampling and measurement of PGF2α metabolite (PGFM) levels in the systemic circulation on Days 11 to 16 postovulation in four control and eight uterine device mares, PGF2α secretion was attenuated in mares with prolonged CL function, with the exception of two mares; one mare showed a single PGFM peak and another showed two isolated PGFM peaks. Because
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Estrus Suppression there was no evidence of inflammatory changes caused by the intrauterine device, the authors concluded that the physical presence of the device in the uterine lumen somehow mimicked the effect of a conceptus by impairing endometrial secretion of PGF2α ; however, the exact mechanism remains unknown.
Administration of exogenous oxytocin In contrast to using an intrauterine device to extend CL function, administration of exogenous oxytocin during diestrus is an alternative method of blocking luteolysis to prolong CL function. Endogenous oxytocin secretion is involved in regulating prostaglandin PGF2α secretion from the endometrium during spontaneous luteolysis in the mare,32, 33 and although administration of exogenous oxytocin to mares around the time of luteolysis (i.e., Days 11 to 15 post-ovulation) stimulates an acute onset of PGF2α secretion,34–36 when oxytocin is administered in the mid-luteal phase prior to the expected time of luteolysis (i.e., before Day 10 post-ovulation) it does not induce PGF2α secretion and often disrupts luteolysis causing prolonged CL function.35 Experimentally, continuous infusion of oxytocin using a subcutaneous osmotic minipump from Day 8 to 20 post-ovulation blocked luteolysis in four of five mares, whereas luteolysis occurred at the expected time in all four control mares that received saline infusion.37 Although it successfully induced prolonged CL function, continuous infusion of oxytocin to disrupt luteolysis would not be a practical method of long-term suppression of estrous behavior. More recently, Vanderwall et al.38 showed that twice daily intramuscular administration of 60 units of oxytocin on Days 7 to 14 post-ovulation was an efficacious method of disrupting luteolysis, since it caused prolonged corpora luteal function through Day 30 post-ovulation in six treated mares, whereas six saline-treated control mares underwent luteolysis by Day 16 post-ovulation. Progesterone levels fell below 1.0 ng/mL between Days 30 and 40 in two of the mares with prolonged CL function, while the other four mares maintained progesterone levels above 3.0 ng/mL through Day 40, when blood sampling was discontinued. The cessation of CL function before Day 40 in two mares may have reflected a seasonal effect on CL function, since the study was completed at the end of the physiological breeding season when gonadotropin secretion wanes, and CL function (i.e., progesterone secretion) is dependent upon adequate support from endogenous gonadotropin secretion.39–41 Further work is needed to characterize more fully the duration of CL function when exogenous oxytocin is
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used to inhibit luteolysis earlier in the physiological breeding season. In a follow-up study (D.K. Vanderwall et al., unpublished), once daily intramuscular administration of 60 units oxytocin was as effective as twice daily administration in inducing prolonged CL function in mares, which simplifies the treatment protocol. An advantage of using exogenous oxytocin to prolong CL function is that it can be readily reversed by the administration of a luteolytic dose of PGF2α , in contrast to the need physically to remove an intrauterine device as described above.
Inducing late-diestrus ovulation In a recent report, Hedberg et al.42 described the results of a preliminary study in which their objective was to prolong the luteal phase in mares by using human chorionic gonadotropin (hCG) to induce a late-diestrus ovulation to produce a new CL that would be too immature to respond to the luteolytic effects of endogenous PGF2α secretion at the end of diestrus (i.e., Day 14 to 15 after the initial ovulation). Mares were randomly assigned to control (n = 4) and experimental groups (n = 5), and beginning on approximately Day 8 after ovulation (or last signs of estrus in three mares) their ovaries were examined with transrectal ultrasonography every other day to determine the size(s) of their diestrus follicles. When a diestrus follicle ≥ 30 mm was detected, control mares were treated with saline and experimental mares were treated with 3000 IU hCG intramuscularly. After treatment the mares were followed with transrectal ultrasonography for up to 72 hours or until ovulation was detected, and then once weekly for 3 weeks. Beginning on the day of treatment, blood samples were collected twice weekly for at least 1 month and then once weekly for another 2 to 4 months for progesterone determination. If a mare did not develop a diestrus follicle ≥ 30 mm during the first diestrus period, they were monitored for a second, and if necessary a third, diestrus period. Three of the nine mares developed a follicle ≥ 30 mm during the first diestrus period, four mares during the second diestrus period, and one mare in the third diestrus period. One experimental mare never developed a diestrus follicle that was ≥ 30 mm during the three diestrus periods that were monitored, and therefore could not be treated with hCG. Overall, three out of the four (75%) experimental mares treated with hCG ovulated within 72 hours after treatment with hCG, which resulted in luteal phases that lasted for 58 to 82 days after treatment. None of the control mares ovulated during the luteal phase; however, one control mare had a spontaneously prolonged luteal phase during both a nontreated cycle in which she never developed a diestrus
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follicle ≥ 30 mm (CL function was terminated with exogenous PGF2α ) and during the subsequent cycle in which she was treated with saline when she had a large diestrus follicle (that did not ovulate). Although, based on this study, the use of hCG to induce a late-diestrus ovulation looks promising for prolonging CL function, it is important to note that as described above, for some mares (five out of nine) it required multiple estrous cycles to develop a diestrus follicle ≥ 30 mm. In addition, one mare never developed a large diestrus follicle during the three cycles that were monitored, which precluded her from receiving the hCG treatment. Therefore, in addition to the effort (and expense) of monitoring mares in order to evaluate their suitability for treatment, the fact that some mares may not develop a large enough diestrus follicle to warrant treatment, the use of hCG to induce a late-diestrus ovulation does not appear to be a reliable ‘on-demand’ method of blocking estrous behavior in mares. It is interesting to note that although the use of hCG was apparently efficacious for inducing ovulation of diestrus follicles ≥ 30 mm in diameter in this study by Hedberg et al.,42 previous work by Glazar et al.43 demonstrated that administration of the gonadotropin-releasing hormone (GnRH) agonist deslorelin acetate failed to induce ovulation and/or luteinization of diestrus follicles > 30 mm in diameter.
Pregnancy Pregnancy is another means of suspending cyclicity by taking advantage of the natural ability of the conceptus to block luteolysis and maintain CL function/progesterone secretion. Although efficacious, this method has obvious disadvantages that may make it undesirable for many horse owners. In addition to the time and expense necessary to establish pregnancy, is the need to eventually terminate pregnancy (unless an offspring is ultimately desired). Lefranc and Allen44 reported that manual transrectal rupture of the conceptus between Days 16 and 22 of gestation in 11 mares resulted in continued CL function for at least 60 days in all of the mares, during which time they did not display estrous behavior. Although efficacious, as noted above, terminating a normal, healthy pregnancy may be untenable to many horse owners. Suppressing ovarian follicular activity
Downregulation with gonadotropinreleasing hormone (GnRH) analogs Although a subcutaneous implant containing the potent GnRH analog deslorelin acetate (Ovuplant,
Fort Dodge Animal Health) was initially developed and found to be efficacious for inducing timed ovulation for the breeding management of mares,45, 46 it soon became evident that the implant caused prolonged anovulatory periods in some mares.47–49 Subsequent work demonstrated the deslorelin implant caused reduced circulating concentrations of follicle-stimulating hormone (FSH) and absence of the mid-cycle FSH peak that was associated with a prolonged interovulatory interval.50 Although problematic for the breeding management of mares, the downregulating effect of the deslorelin implant on pituitary function has been used clinically to suppress ovarian activity; for example, placement of two deslorelin implants suppressed follicular development in mares used as recipients for oocyte transfer.51 Fitzgerald et al.52 reported suppressing ovulation for 30–90 days in 15/20 mares that were treated with a subcutaneous implant containing another GnRH analogue (goserelin acetate). Collectively, these data indicate that treating mares with a potent GnRH analog may be efficacious for suspending cyclicity and suppressing estrus in mares for extended periods of time. However, as noted previously, since anovulatory mares can show estrous behavior, suppressing follicular activity may not ensure a complete absence of estrous behavior.
Immunological approaches Immunizing a mare against GnRH eliminates gonadotropin release from the pituitary resulting in suspension of cyclicity. Tshewang et al.53 reported successfully suspending ovarian activity and suppressing estrus for 25–30 weeks in mares after treating with a GnRH vaccine. All of the vaccinated mares recovered from the effect of immunization with normal cyclicity and estrous behavior. Over the next two seasons, each mare also conceived and produced a normal foal. Other studies54–56 have similarly demonstrated the efficacy of vaccination against GnRH for suppressing ovarian follicular activity; however, not surprisingly, some of the vaccinated mares continued to show estrous behavior in spite of their anovulatory state.54, 55 It is important to note that in one study,54 a vaccinated mare had not resumed normal ovarian activity 2 years after initial vaccination, and in another report57 10 young Thoroughbred mares that had been anovulatory for one to three breeding seasons after retirement from racing were retrospectively identified as having been vaccinated (prior to retirement from racing) two or more times against GnRH; therefore, although most mares appear to regain ovarian activity within a reasonable period of time post-vaccination, some mares
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Estrus Suppression may not, which could be particularly problematic if subsequent use as a broodmare is desired. Although a GnRH vaccine is not currently available in North America, commercial preparations are available in Europe and Australia (Equity, CSL Animal Health).
Ovariectomy Bilateral ovariectomy may be warranted in some circumstances for permanently eliminating ovarian activity in mares. Although the ovaries can be removed via ventral abdominal or flank laparotomy; laparoscopy; or colpotomy, at this time, the latter two methods are most commonly utilized.16, 17 In an initial report, Hooper et al.16 described the outcome of the use of bilateral ovariectomy for treatment of objectionable behavior during estrus in 17 mares; they found that after surgery, owners reported that behavior was no longer a problem in 14 mares (82%), while the remaining three mares continued to show estrous behavior. Of the 23 mares in the study16 that underwent bilateral ovariectomy for any reason, eight (35%) continued to show estrous behavior after surgery. In a recent report, Kamm and Hendrickson17 evaluated clients’ perspectives on the outcome following the use of laparoscopic ovariectomy for behavioral and medical problems in mares. Overall, client satisfaction (rated as very satisfied or satisfied) with bilateral ovariectomy as a treatment for behavioral problems was 78% (18 of 23 of mares). Assessment of outcomes for specific behavioral problems showed that aggression problems improved in 86% of cases; general disagreeable demeanor improved in 81%; excitability improved in 75%; kicking and biting at other horses improved in 73%; problems in training improved in 72%; frequent urination improved in 64%; and problems with other horses improved in 64% of cases. The most common source of dissatisfaction for owners of patients with behavioral problems was lack of behavioral change after surgery, including continued signs of estrous behavior. Although ovariectomy offers the advantage of being a potentially permanent solution to a cycle-related behavior/performance problem, there are significant disadvantages as well. The procedure is relatively expensive, though when compared to repeated hormonal treatments, the cost may in fact be comparable. There are also risks associated with the surgical procedure, though newer laparoscopic procedures have significantly reduced them.16, 17 The permanency of the procedure can also be a significant disadvantage because all possibility for future reproduction is eliminated. And as noted above, if the behavior/performance problem is truly related to estrus, removing the ovaries may not solve
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the problem if the mare continues to display estrous behavior. Therefore, this option should be weighed carefully before proceeding. One way of evaluating the potential effectiveness of ovariectomy for a behavioral problem is to evaluate the mare’s behavior during either a natural (i.e., seasonal) or induced anovulatory state (e.g., downregulation with a GnRH agonist); if the problem is not resolved or at least significantly improved during the anovulatory state, it is unlikely ovariectomy will be any more efficacious.
Summary The primary indications for suppressing estrous behavior in mares include cycle-related behavior/ performance problems and pain/colic during estrus. When evaluating an owner/trainer complaint of an estrous cycle-related problem in a mare, the veterinarian should first determine if the problematic behavior is or is not related to a specific phase of the estrous cycle. In order to thoroughly evaluate the mare, additional expertise may be needed in the form of consultation with or referrals to behavior and/or reproduction experts. All non-cycle-related problems should be identified and treated to avoid attempts to mask more serious sources of pain or irritation in a mare simply for human benefit. If a behavior/ performance/ovarian pain problem is confidently defined as estrous cycle-related, the previously discussed methods (with proven efficacy) of suppressing estrous behavior can be considered for use. Each method has advantages and potential disadvantages that should be weighed for each individual animal/owner/trainer.
References 1. Freedman LJ, Garcia MC, Ginther OJ. Influence of photoperiod and ovaries on seasonal reproductive activity in mares. Biol Reprod 1979;20:567–74. 2. Ginther OJ. Folliculogenesis during the transitional period and early ovulatory season in mares. J Reprod Fertil 1990;90:311–20. 3. Ginther OJ. Occurrence of anestrus, estrus, diestrus, and ovulation over a 12-month period in mares. Am J Vet Res 1974;35:1173–9. 4. Hart PJ, Squires EL, Imel KJ, Nett TM. Seasonal variation in hypothalamic content of gonadotropin- releasing hormone (GnRH), pituitary receptors for GnRH, and pituitary content of luteinizing hormone and follicle-stimulating hormone in the mare. Biol Reprod 1984;30:1055–62. 5. Asa CS, Goldfoot DA, Garcia MC, Ginther OJ. Sexual behavior in ovariectomized and seasonally anovulatory pony mares (Equus caballus). Horm Behav 1980;14:46–54. 6. Asa CS, Goldfoot DA, Carcia MC, Ginther OJ. Dexamethasone suppression of sexual behavior in the ovariectomized mare. Horm Behav 1980;14:55–64. 7. Crowell–Davis SL. Sexual behavior of mares. Horm Behav 2007;52:12–17.
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8. Jorgensen JS, Vivrette S, Correa M, Mansmann RA. Significance of the estrous cycle on athletic performance in mares. Proc Annu Conv Am Assoc Equine Practnr 1996;42:98– 100. 9. McDonnell SM. Evaluation and modification of mare behavior problems. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1993; pp. 185–9. 10. McDonnell S. Performance problems in mares. The Horse 2000;April:61–70. 11. Hedberg Y, Dalin AM, Ohagen P, Holm KR, Kindahl H. Effect of oestrous-cycle stage on the response of mares in a novel object test and isolation test. Reprod Domest Anim 2005;40:480–8. 12. Pryor P, Tibary A. Management of estrus in the performance mare. Clin Tech Equine Pract 2005;4:197–209. 13. McDonnell SM. Is it psychological, physical, or both? Proc Annu Conv Am Assoc Equine Practnr 2005;51:231–8. 14. McDonnell SM. Estrus cycle-related performance problems. J Equine Vet Sci 1997;17:196. 15. Cox JH, DeBowes RM. Colic-like discomfort associated with ovulation in two mares. J Am Vet Med Assoc 1987;191:1451–2. 16. Hooper RN, Taylor TS, Varner DD, Blanchard TL. Effects of bilateral ovariectomy via colpotomy in mares: 23 cases (1984–1990). J Am Vet Med Assoc 1993;203:1043–6. 17. Kamm JL, Hendrickson DA. Client’s perspectives on the effects of laparoscopic ovariectomy on equine behavioral and medical problems. J Equine Vet Sci 2007;27:435–8. 18. Loy RG, Swan SM. Effects of exogenous progestogens on reproductive phenomena in mares. J Anim Sci 1966;25:821–6. 19. Vanderwall DK, Marquardt JL, Woods GL. Use of a compounded long-acting progesterone formulation for equine pregnancy maintenance. J Equine Vet Sci 2007;27: 62–6. 20. Hawkins DL, Neely DP, Stabenfeldt GH. Plasma progesterone concentrations derived from the administration of exogenous progesterone to ovariectomized mares. J Reprod Fertil Suppl 1979;27:211–16. 21. Squires EL, Stevens WB, McGlothlin DE, Pickett BW. Effect of an oral progestin on the estrous cycle and fertility of mares. J Anim Sci 1979;49:729–35. 22. Webel SK, Squires EL. Control of the oestrous cycle in mares with altrenogest. J Reprod Fertil Suppl 1982;32:193–8. 23. Gee EK, DeLuca C, Stylski JL, McCue PM. Efficacy of medroxyprogesterone acetate in suppression of estrus in cycling mares. J Equine Vet Sci 2009;29:140–5. 24. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32:83–5. 25. McKinnon AO, Tarrida Del Marmol Figueroa S, Nobelius AM, Hyland JH, Vasey JR. Failure of hydroxyprogesterone caproate to maintain pregnancy in ovariectomized mares. Equine Vet J 1993;25:158–60. ´ 26. Lopez-Bayghen C, Zozaya H, Ocampo L, Brumbaugh GW, Sumano H. Melengestrol acetate as a tool for inducing early ovulation in transitional mares. Acta Veterinaria Hungarica 2008;56:125–31. 27. Wiepz GJ, Squires EL, Chapman PL. Effects of norgestomet, altrenogest, and/or estradiol on follicular and hormonal characteristics of late transitional mares. Theriogenology 1988;30:181–93. 28. McCue PM, Lemons SS, Squires EL, Vanderwall DK. Efficacy of synovex-S implants in suppression of estrus in the mare. J Equine Vet Sci 1997;17:327–9.
29. Scheffrahn NS, Wiseman BS, Vincent DL, Harrison PC, Kesler DJ. Reproductive hormone secretions in pony mares subsequent to ovulation control during late winter. Theriogenology 1982;17:571–85. 30. Nie GJ, Johnson KE, Braden TD, Wenzel JGW. Use of an intra-uterine glass ball protocol to extend luteal function in mares. J Equine Vet Sci 2003;23:266–73. 31. Rivera del Alamo MM, Reilas T, Kindahl H, Katila T. Mechanisms behind intrauterine device-induced luteal persistence in mares. Anim Reprod Sci 2008;107:94–106. 32. Vanderwall DK, Silvia WJ, Fitzgerald BP. Concentrations of oxytocin in the intercavernous sinus of mares during luteolysis: temporal relationship with concentrations of 13,14-dihydro-15-keto-prostaglandin F2α . J Reprod Fertil 1998;112:337–46. 33. Shand N, Irvine CHG, Turner JE, Alexander SL. A detailed study of hormonal profiles in mares at luteolysis. J Reprod Fertil Suppl 2000;56:271–9. 34. Betteridge KJ, Renard A, Goff AK. Uterine prostaglandin release relative to embryo collection, transfer procedures and maintenance of the corpus luteum. Equine Vet J 1985;Suppl. 3:25–33. 35. Goff AK, Pontbriand D, Sirois J. Oxytocin stimulation of plasma 15-keto-13,14-dihydro prostaglandin F-2α during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1987;35:253–60. 36. Starbuck GR, Stout TA, Lamming GE, Allen WR, Flint AP. Endometrial oxytocin receptor and uterine prostaglandin secretion in mares during the oestrous cycle and early pregnancy. J Reprod Fertil 1998;113:173–9. 37. Stout TAE, Lamming GE, Allen WR. Oxytocin administration prolongs luteal function in cyclic mares. J Reprod Fertil 1999;116:315–20. 38. Vanderwall DK, Rasmussen DM, Woods GL. Effect of repeated administration of oxytocin during diestrus on duration of function of corpora lutea in mares. J Am Vet Med Assoc 2007;231:1864–7. 39. Pineda MH, Ginther OJ, McShan WH. Regression of corpus luteum in mares treated with an antiserum against an equine pituitary fraction. Am J Vet Res 1972;33:1767–73. 40. Pineda MH, Garcia MC, Ginther OJ. Effect of antiserum against an equine pituitary fraction on corpus luteum and follicles in mares during diestrus. Am J Vet Res 1973;34:181–3. 41. Bergfelt DR, Ginther OJ. Embryo loss following GnRHinduced ovulation in anovulatory mares. Theriogenology 1992;38:33–43. 42. Hedberg Y, Dalin AM, Santesson M, Kindahl H. A preliminary study on the induction of dioestrous ovulation in the mare – a possible method for inducing prolonged luteal phase. Acta Vet Scand 2006;48:12. 43. Glazar BS, McCue PM, Bruemmer JE, Squires EL. Deslorelin on Day 8 or 12 postovulation does not luteinize follicles during an artificially maintained diestrous phase in the mare. Theriogenology 2004;62:57–64. 44. Lefranc AC, Allen WR. Nonpharmacological suppression of oestrus in the mare. Equine Vet J 2004;36:183–5. 45. Meinert C, Silva JFS, Kroetz I, Klug E, Trigg TE, Hoppen ¨ HO, Jochle W. Advancing the time of ovulation in the mare with a short-term implant releasing the GnRH analogue deslorelin. Equine Vet J 1993;25:65–8. 46. McKinnon AO, Nobelius AM, Tarrida Del Marmol Figueroa S, Skidmore J, Vasey JR, Trigg TE. Predictable ovulation in mares treated with an implant of the GnRH analogue deslorelin. Equine Vet J 1993;25:321–3.
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Estrus Suppression 47. Johnson CA, Thompson DL Jr, Kulinski KM, Guitreau AM. Prolonged interovulatory interval and hormonal changes in mares following the use of OvuplantTM to hasten ovulation. J Equine Vet Sci 2000;20:331–6. 48. Morehead JA, Blanchard TL. Clinical experience with deslorelin (OvuplantTM ) in a Kentucky Thoroughbred broodmare practice (1999). J Equine Vet Sci 2000;20:358–402. 49. Vanderwall DK, Juergens TD, Woods GL. Reproductive performance of commercial broodmares after induction of ovulation with HCG or OvuplantTM (Deslorelin). J Equine Vet Sci 2001;21:539–42. 50. Farquhar VJ, McCue PM, Nett TM, Squires EL. Effect of deslorelin acetate on gonadotropin secretion and ovarian follicle development in cycling mares. J Am Vet Med Assoc 2001;218:749–52. 51. Carnevale EM, Checura CH, Coutinho da Silva MA, Maclellan LJ, Squires EL. Use of deslorelin acetate to suppress follicular activity in mares used as recipients for oocyte transfer. Theriogenology 2001;55:358 (abstract). 52. Fitzgerald BP, Peterson KD, Silvia PJ. Effect of constant administration of a gonadotropin-releasing hormone agonist
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on reproductive activity in mares: preliminary evidence on suppression of ovulation during the breeding season. Am J Vet Res 1993;54:1746–51. Tshewang U, Dowsett KF, Knott LM, Trigg TE. Preliminary study of ovarian activity in fillies treated with a GnRH vaccine. Aust Vet J 1997;75:663–7. Imboden I, Janett F, Burger D, Crowe MA, Hassig M, Thun R. Influence of immunization against GnRH on reproductive cyclicity and estrous behavior in the mare. Theriogenology 2006;66:1866–75. Dalin A-M, Andresen Ø, Malmgren L. Immunization against GnRH in mature mares: antibody titres, ovarian function, hormonal levels and oestrous behaviour. J Vet Med 2002;49:125–31. Elhay M, Newbold A, Britton A, Turley P, Dowsett K, Walker J. Suppression of behavioural and physiological oestrus in the mare by vaccination against GnRH. Aust Vet J 2007;85:39–45. Robinson SJ, McKinnon AO. Prolonged ovarian inactivity in broodmares temporally associated with administration of Equity. Aust Eq Vet 2006;25:85–7.
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Chapter 195
Inhibin Immunotherapy Patrick M. McCue
Introduction Inhibin is a heterodimeric glycoprotein hormone consisting of α and β subunits linked by disulfide bonds.1 The primary role of inhibin is the specific suppression of follicle-stimulating hormone (FSH) secretion from the anterior pituitary gland.2 Inhibin plays an integral and active role in the complex process of follicle selection and regulation of the number of ovulations in the mare. The process begins when a cohort of follicles is recruited and emerges from a pool of smaller follicles 7 to 8 days after ovulation. These follicles subsequently proceed through a common-growth phase for approximately 6 to 7 days.3, 4 Follicle-stimulating hormone produced by the anterior pituitary gland provides the drive for emergence and initial growth and development of all follicles in the follicular wave. The future dominant follicle is usually one that emerges early in the wave and is slightly larger at the end of the common-growth phase than other follicles in the cohort. Deviation occurs at the end of the common-growth phase and represents the point at which the future dominant follicle continues to develop and smaller subordinate follicles regress. Production of inhibin and estradiol by granulosa cells of the developing dominant follicle results in suppression of circulating FSH levels. The dominant follicle is more responsive to FSH and luteinizing hormone (LH) than smaller follicles in the wave, due in part to an increase in the number of gonadotropin receptors stimulated by elevated intrafollicular estradiol levels. Consequently, the larger follicle continues to develop in the face of declining FSH levels while subordinate follicles, which are still critically dependent on FSH for continued development, begin to regress. Deviation in follicular development in mares occurs when the largest follicle is 22 to 25 mm in diameter.5 A prolonged elevation of LH is responsible for continued
development of the dominant follicle after deviation and subsequent ovulation. Levels of circulating inhibin in mares have been reported to be positively correlated with estradiol levels and inversely correlated with FSH concentrations.6, 7 Inhibin concentrations increase during estrus, are highest on the day of ovulation, and decline rapidly thereafter.7 In contrast, concentrations of FSH in mares peak in mid-diestrus and then decline as levels of inhibin and estradiol increase during estrus.
Granulosa cell tumors Studies of ovarian pathophysiology have provided valuable insight into control of pituitary gonadotropin secretion, regulation of follicle development, follicular wave dynamics, and control of ovulation rate in the mare. Mares with ovarian granulosa cell tumors have high circulating concentrations of inhibin8–11 and reduced FSH levels.12 The lack of FSH support results in suppression of follicular development in the otherwise normal contralateral ovary.10, 13, 14 Follicular development and ovulation in the contralateral ovary does not usually resume until 6 to 8 months after the neoplastic ovary is surgically removed.
Inhibin administration Administration of charcoal-treated follicular fluid has been used as a technique to determine the effects of exogenous inhibin on gonadotropin secretion and follicular development.15, 16 Charcoal extraction is used to remove steroids from the follicular fluid. Treatment of ovariectomized,15 intact cycling,16–18 and intact pregnant mares19 with steroid-free follicular fluid resulted in suppression of circulating FSH levels. In addition, follicular development was reduced in intact cycling mares and early pregnant
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Inhibin Immunotherapy mares. In vitro, the suppressive effect of steroid-free equine follicular fluid on FSH secretion from cultured rat pituitary cells was reversed by the addition of inhibin antiserum.20 Collectively, these studies confirm the suppressive effects of inhibin on FSH secretion in the mare.
Active immunization against inhibin Since an elevation in circulating inhibin is associated with a reduction of pituitary FSH secretion and subsequent suppression of follicular development, it follows that blockage of inhibin bioactivity would allow for an increase in FSH levels and enhanced follicular development. Active immunization of mares against inhibin was originally reported using a synthetic porcine α-subunit fragment9 or a recombinant bovine α-subunit.21 In one study, the antigen was the first 26 amino acids of the porcine inhibin α-subunit conjugated to bovine serum albumin (BSA), and mares were immunized five times at 3-week intervals.9 In the other study, the intact recombinant bovine α-subunit was the immunogen, and mares were vaccinated twice at an interval of approximately 5 weeks.21 Immunization of mares with the anti-inhibin vaccines resulted in seroconversion and development of high antibody titers in most mares. In general, antibody titers increased further after booster immunizations. The overall antibody response to immunization varied between mares within each study. Ovulation rates of mares actively immunized against inhibin were increased in both studies. McCue et al.9 noted an ovulation rate of 2.8 ± 1.1 after completion of the immunization series, compared to 1.1 ± 0.1 for non-immunized control mares. Ovulation rate following immunization was not significantly correlated with the serum antibody titer. McKinnon et al.21 recorded 2.3 ± 0.2 ovulations per cycle after booster immunization as compared to 1.2 ± 0.1 ovulations per cycle for the same mares prior to immunization. The embryo collection rate for mares immunized against inhibin (1.6 ± 0.5 embryos per flush) was noted to be significantly greater than that of control mares (0.7 ± 0.2 embryos per flush).9 There was no significant difference in embryo recovery rate per ovulation between immunized and control mares (0.58 and 0.57, respectively). Two additional studies were subsequently reported in which mares were actively immunized against inhibin.22, 23 Terhaar and colleagues22 used a recombinant human α-subunit as the vaccine antigen. Immunization did not alter the pattern of FSH secretion or change ovulation rate. It was further noted
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that three of the four immunized mares exhibited estrous cycle abnormalities, including delayed ovulations, follicular atresia, formation of anovulatory hemorrhagic follicles, and ovulations independent of behavioral estrus. Meyers-Brown and co-workers23 vaccinated mares five times with a porcine inhibin α-subunit conjugated to keyhole limpet hemocyanin (KLH). Ovulation and embryo recovery rates were noted to be increased in comparison to control mares.
Passive immunization against inhibin Passive transfer of inhibin antibodies in mares has been reported in three studies. McCue et al.24 administered 2 L of hyperimmune anti-inhibin plasma to five mares on Day 10 after ovulation and, to another group of five mares, 1 L on Day 10 and a further 1 L on Day 15 after ovulation. There was no effect of passive immunization on circulating levels of FSH detected, but 8 of 10 treated mares developed multiple large (≥ 35 mm) pre-ovulatory follicles, and 5 of 10 mares had spontaneous multiple ovulations. The overall ovulation rate increased from 1.0 ± 0.0 ovulations in control cycles to 1.6 ± 0.2 ovulations in treated cycles. The ovulation rate of mares passively immunized against inhibin was lower than that of mares actively immunized as reported by the same research group. This may have been due to a lower total amount of circulating anti-inhibin antibodies available following plasma transfusion than that present following active immunization. Nambo and colleagues20 administered inhibin antiserum raised in a castrated goat to a total of eight mares 4 days after prostaglandin administration. Treatment resulted in an increase in plasma FSH and estradiol levels, whereas the level of LH was not affected. A dose-dependent increase in ovulation rate was noted, with mares receiving 100 mL or 200 mL of antiserum exhibiting ovulation rates of 3.8 ± 0.6 and 4.5 ± 0.7, respectively, as compared to an ovulation rate of 1.3 ± 0.3 in mares receiving control serum. However, it was reported that ovulations in antiserum-treated mares were not simultaneous, but separated over a period of 3 to 7 days. Mares in this study were not administered an ovulation-inducing agent, such as human chorionic gonadotropin (hCG). Administration of hCG may have resulted in more synchronized ovulations. Meyers-Brown and colleagues23 reported that passive immunization of mares with a purified and concentrated inhibin antibody preparation once daily for 2–4 days beginning 14 days after ovulation resulted in increased serum FSH concentrations. Passive immunization was associated with an increased number
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of large (> 35 mm) follicles and higher embryo recovery rates versus control cycles.
Summary Both active and passive immunization of mares against inhibin have been used as techniques to increase endogenous FSH levels by neutralizing the bioactivity of the primary factor that suppresses FSH secretion. Clinically, the ultimate goal of increasing endogenous FSH is to enhance ovulation rates for embryo transfer or other reproductive purposes. The main potential advantages of active immunization against inhibin are that it could be an easy and relatively inexpensive technique to increase ovulation rates in mares, and embryos recovered following active immunization have been noted to be normal in morphology and grade. There are also several important disadvantages to inhibin immunotherapy. First, there is no commercially available inhibin vaccine. Second, the immune response after active immunization varies greatly between mares. In addition, active immunization cannot be used ‘on demand’ to alter ovulation rate during a specific estrous cycle, and the antibodymediated effect cannot be turned off when the effect on ovarian function is no longer desired. Finally, potential adverse effects of inhibin immunotherapy (i.e., immune-mediated ovarian pathology) should be investigated before this technique is ever utilized on privately owned mares. Passive immunization against inhibin was reported to be effective in increasing ovulation rate during a specific estrous cycle in mares. Ultimately, however, it is unlikely that blood products, especially those from another species, will be approved for use as a technique to modulate the estrous cycle of mares. It is possible that future development of nonimmunological inhibin antagonists may provide an alternative to active or passive immunization against inhibin. Recent advances in superovulation with purified equine FSH and recombinant equine FSH have resulted in a decreased interest in the use of inhibin immunotherapy for enhancement of ovulation rates in mares.
References 1. Burger HG, Igarashi M. Inhibin: definition and nomenclature, including related substances. Endocrinology 1988; 122:1701–2. 2. de Kretser DM, Robertson DM. The isolation and physiology of inhibin and related proteins. Biol Reprod 1989;40: 33–47.
3. Ginther OJ. Major and minor follicular waves during the equine estrous cycle. J Equine Vet Sci 1993;13:18–25. 4. Gastal EL, Gastal MO, Beg MA, Ginther OJ. Interrelationships among follicles during the common-growth phase of a follicular wave and capacity of individual follicles for dominance in mares. Reproduction 2004;128:417–22. 5. Ginther OJ, Beg MA, Donadeu FX, Bergfelt DR. Mechanism of follicular deviation in monovular farm species. Anim Reprod Sci 2003;78:239–57. 6. Bergfelt DR, Mann BG, Schwartz NB, Ginther OJ. Circulating concentrations of immunoreactive inhibin and FSH during the estrous cycle of mares. J Equine Vet Sci 1991;11:319–22. 7. Roser JF, McCue PM, Hoye E. Inhibin activity in the mare and stallion. Domest Anim Endocrin 1994;11:87–100. 8. Bailey MT, Troedsson MHT, Wheaton JE. Inhibin concentrations in mares with granulosa cell tumors. Theriogenology 2002;57:1885–95. 9. McCue PM, Carney NJ, Hughes JP, Rivier J, Vale W, Lasley BL. Ovulation and embryo recovery rates following immunization of mares against an inhibin alpha-subunit fragment. Theriogenology 1992;38:823–31. 10. McCue PM, Roser JF, Munro CJ, Liu IKM, Lasley BL. Granulosa cell tumors of the equine ovary. Vet Clin Equine Pract 2006;22:799–817. 11. Watson ED, Heald M, Leask R, Groome NP, Riley SC. Detection of high circulating concentrations of inhibin proand -αC immunoreactivity in mares with granulosa-theca cell tumours. Equine Vet J 2002;34:203–6. 12. McCue PM. Equine granulosa cell tumors. In: Proceedings of the 38th Annual Convention of the American Association of Equine Practitioners, 1992; 38: 587–93. 13. Meagher DM, Wheat JD, Hughes JP, Stabenfeldt GH, Harris BA. Granulosa cell tumors in mares – a review of 78 cases. Annual Convention of the American Association of Equine Practitioners 1978;23:133–43. 14. Stabenfeldt GH, Hughes JP, Kennedy PC, Meagher DM, Neely DP. Clinical findings, pathological changes and endocrinological secretory patterns in mares with ovarian tumors. J Reprod Fertil Suppl 1979;27:277–85. 15. Miller KF, Wesson JA, Ginther OJ. Changes in concentrations of circulating gonadotropins following administration of equine follicular fluid to ovariectomized mares. Biol Reprod 1979;21:867–72. 16. Miller KF, Wesson JA, Ginther OJ. Interaction of estradiol and nonsteroidal follicular fluid substance in the regulation of gonadotropin secretion in the mare. Biol Reprod 1981;24:354–8. 17. Bergfelt DR, OJ Ginther. Delayed follicular development and ovulation following inhibition of FSH with equine follicular fluid in the mare. Theriogenology 1985;24:99–108. 18. Plata-Madrid H, Loch WE, Youngquist RS, Thompson DL Jr, Bennett-Wimbush KG, Wilkerson C, Bouchard G, Smith MF, Braun WF, Aveiro JJ. Control of FSH, follicular development and estrus synchronization in the mare with steroid-free follicular fluid. Theriogenology 1992;37:817–38. 19. Bergfelt DR, Ginther OJ. Follicular populations following inhibition of follicle stimulating hormone with equine follicular fluid during early pregnancy in the mare. Theriogenology 1986;26:733–47. 20. Nambo Y, Kaneko H, Nagata S, Oikawa M, Yoshihara T, Nagamine N, Watanabe G, Taya K. Effect of passive immunization against inhibin on FSH secretion, folliculogenesis and ovulation rate during the follicular phase of the estrous cycle in mares. Theriogenology 1998;50:545–57.
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Inhibin Immunotherapy 21. McKinnon AO, Brown RW, Pashen RL, Greenwood PE, Vasey JR. Increased ovulation rates in mares after immunization against recombinant bovine inhibin α-subunit. Equine Vet J 1992;24:144–6. 22. Terhaar H-J, Schlote S, Hoppen H-O, Hennies M, Holtz W, Merkt H, Bader H. Active immunization of mares against the recombinant human inhibin α-subunit. Reprod Domest Anim 1997;32:243–50.
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23. Meyers-Brown GA, Whisnant CS, Brown BS, Hower MA. The effects of active versus passive immunization of mares against inhibin on embryo quantity and quality. In: Proceedings of the Equine Nutrition and Physiology Society, 1997, pp 288–93. 24. McCue PM, Hughes JP, Lasley BL. Effect on ovulation rate of passive immunisation of mares against inhibin. Equine Vet J 1993; Suppl. 15: 103–6.
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Chapter 196
Induction of Ovulation Angus O. McKinnon and Patrick M. McCue
Introduction Induction of ovulation is a controlled advancement of an event that would have been expected to have occurred anyway. In that regard, the term ‘induction’ of ovulation is a misnomer and the term we prefer is ‘programming’ mares to ovulate within a specific time interval. Typically, a maturing dominant follicle that would have ovulated in 3–5 days can undergo earlier maturation with ovulation within a reasonably predictable timeframe of 24–48 hours after treatment. For agents to demonstrate efficacy in controlling time to ovulation, mares must be in estrus, and apart from follicle size, attention to teasing patterns, a softening cervix, and endometrial edema are all important.1 Programming time of ovulation is a common procedure in most veterinary practices that use good management. However, the mechanisms of action of pharmacological agents and reasons for failure are still areas that need clarification. Induction of ovulation will not occur in a precise manner when agents are administered with a follicle not mature enough to respond. It is expected that ovulation will occur in 24–48 hours after an ovulation-inducing agent is administered. When ovulation occurs earlier than 24 hours it is likely the follicle was already destined to ovulate, before the agent was administered. When a follicle ovulates more than 48 hours after administration, it is likely that the agent was not administered when a suitably mature follicle was present. Typically, a follicle will respond to an ovulating agent when the mare has been in estrus for 2–3 days and the follicle has reached a minimum critical size that is characteristic for the type or breed of mare (Figure 196.1). The overall goal when programming a mare is to administer the ovulating agent to a mare with a dominant follicle of an appropriate size and maturation status early enough in estrus to advance the time of
ovulation to a period that can be predicted (i.e., the ‘ovulation window’). In general, administration of an ovulating agent will advance the ovulation by 1 to 3 days, as compared to when the mare would have ovulated spontaneously. Recognition of estrus is paramount and, all too often, mares are programmed to ovulate without reference to teasing patterns and a softening cervix as more and more reliance is placed on ultrasonography. This happens more frequently on smaller farms that may not have access to a teaser stallion and on farms that rely heavily on artificial insemination (AI). Failure to tease properly may result in mares presented for breeding that are not in estrus, and some mares may even have a corpus luteum present. The size of a follicle that can respond to an ovulating agent will vary according to the time of the year and the breed. Typically, mares will need two or three follicular waves to develop and subside before the first ovulation from the transitional period to the ovulatory season. The response to ovulating agents in the early transition follicular waves is very poor. In the regular ovulatory season (once the first ovulation has occurred) most Thoroughbred and Standardbred mares could be expected to have a predictable response to programming when follicles reach 35–40 mm, and most Warmbloods 40–45 mm. However, many smaller breeds (ponies and some Quarter Horses) will have already ovulated before the follicles have reached that size and will have much smaller follicles that are capable of a predictable response to programming. Draft horses and Friesians are at the other extreme, and will usually not respond to an ovulating agent until the dominant follicle is greater than 50 mm in diameter. When using care and attention to detail, most clinicians should expect a response rate (defined as ovulating between 24–48 hours after administration) of at least 95%. The ability to achieve this will vary with the skill of the person programming, the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 196.1 Expected time interval to ovulation in response to programming versus the unpredictable ovulation pattern when mares are left to spontaneously ovulate. The vertical arrow is the time of programming and the horizontal arrows the expected and possible ovulation window for programmed and spontaneously ovulating mares, respectively. Adapted from Ginther, 1992.
frequency of examination prior to programming, the use of PGF2α to initiate estrus, the time of year, and the agents used. On larger breeding farms where clients are very discerning of veterinary abilities, and similar to the ability of predicting the response to prostaglandin administration, the response of programming mares to ovulate is an easy target for farm personnel to identify and monitor and to some extent judge the veterinarian. This chapter will outline the reason for use, agents available, techniques used to assess follicle maturation, and reasons for failure when programming mares to ovulate.
Reasons for programming ovulation Routine use
Transition The transition period (see Chapter 178) is a difficult time to predict ovulation. Multiple follicular waves are characteristic of the transitional period. In general, one can predict how far the mare is through the transition period by the size of the largest follicle. However, it is common for mares to have multiple follicles that are > 35 mm that often do not progress to ovulate and then regress, to be followed by recrudescence or growth of other follicles. It is difficult to predict not only when a follicle will ovulate but also which follicles are far enough advanced to respond to an ovulating agent. Despite this, the only way to accurately control the transitional period and avoid multiple rebreeds is to use ovulating agents. Commonly a combination of lights, progestagens or dopamine antagonists, and specific ovulatory agents work well. However, when presented with a late transitional mare that has not been under lights and has a large follicle and signs of estrus, it is well worthwhile to use a specific ovulatory agent (see be-
low). A gradual rather than rapid transition from anestrus to normal cyclicity is preferable, as it has been clearly demonstrated that ovulation from deep anestrus is associated with a high incidence of corpus luteum (CL) failure2 and early embryonic death.3
Cycling season Natural service and artificial insemination of fresh semen In any breeding situation with a goal of a single breeding or insemination, the use of agents to predictably hasten ovulation is critical to management. Despite training, experience and skill, it is not always possible with once-daily or every-other-day examinations of mares with rectal palpation and ultrasonography to reliably predict ovulation within a 24-hour period. It is possible to be reasonably accurate, but not at the required level to prevent rebreeding on many occasions if the mare has not ovulated within 48 hours from breeding. It is ideal to aim at a breeding-per-cycle ratio of around 1.05.4 That means that for every 100 cycles, five mares will need to be rebred during a given cycle if they have not ovulated 48 hours after breeding. A recent study of 6123 mare cycles (3105 mares) on Thoroughbred farms reported that utilizing natural service it was possible to achieve a ratio of 1.03. In this study, mares were examined 6–7 times per week and rebreeding was performed if the mare had not ovulated 48 hours after service.5 To achieve a low percentage of rebreeding, ovulating agents are administered at the time of breeding. Some mares will ovulate earlier than expected, but most will ovulate according to the time period expected from the ovulatory agents (see below). The recognition that ovulatory agents improve breeding efficiency has meant a shift in focus in our practice
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from breeding mares with natural service at a time that they are expected to be close to ovulation (predicting ovulation) to a time that they are expected to respond to an ovulatory agent. This paradigm shift has meant that very few mares are accidentally missed in a cycle due to earlier-than-expected ovulation before breeding. Another approach that is often used is to administer ovulatory agents the day before (∼ 24 hours) prior to breeding.6 In a natural service program, we do not favor this for three reasons. First, the early programming of the mare limits our abilities to manipulate the time of breeding, for example, to delay breeding of an individual when a stallion has six or seven mares all waiting to be bred on the one day or the stallion availability has been reduced due to injury, sickness or other generally unforeseen circumstances. Second, our aim is to identify mares that are suitable for programming rather than breeding, so administration one day before breeding may result in mares with insufficient maturity of the dominant follicle to respond appropriately. Finally, some maiden mares are difficult to breed due to an unwillingness to show behavioral estrus in an unfamiliar and stressful environment (for all concerned). These mares often take time to demonstrate proper sexual receptivity and may show best breeding behavior close to a natural ovulation. As a consequence, these mares are examined very frequently before service to more accurately predict ovulation of a follicle not programmed to ovulate at a specific time. This protocol avoids the unfair and unprofessional experience of maidens being physically restrained with excessive force. On Thoroughbred farms, reducing the number of times a stallion breeds during the season is of benefit when large numbers of mares (> 150) are to be bred. Busy stallions with defined breeding times (number of weeks) or services per season (some shuttle stallions) may be especially prone to libido problems and on occasion low sperm numbers, both of which may improve with decreased numbers of breedings and increased intervals between breeding. In addition, a reduction in services limits bacterial challenge to the mares’ reproductive tract, decreases expenses from labor, and decreases transport charges when mares are bred by appointment at remote facilities. Reasons for programming are applicable with AI of fresh semen as well. However, in large AI centers where semen may be readily available to inseminate mares on the farm every other day, many choose to let the mare ovulate naturally. This is also more common where economic restraints are important, as demonstrated with a breeding per cycle of 2.21 in a population of 1330 Standardbred mares over 2086 cycles.4 Interestingly, the fertility of the Standardbred
mares was 62% per cycle, which was significantly less than the Thoroughbred population of 70.4% per cycle.4 In addition, more mares were pregnant per cycle (72%, 1189/1651) within the Thoroughbred population when ovulatory agents were used as compared to not used (64.5%, 527/817).4 Chilled transported semen The difficulties and expense of getting chilled transported semen to be delivered on time in a suitable condition to achieve good fertility suggest that it is imperative to use an ovulatory agent to avoid transporting semen a second time (see Chapters 128 and 129). The decision to use an ovulatory agent before the semen arrives, or immediately after the mare is bred, is up to the discretion of the veterinarian. In general, unless timely delivery and excellent-quality semen can be expected, we prefer to use an ovulatory agent when the semen arrives, as if the semen does not have good motility parameters or does not arrive on time we can potentially order more semen if it can be delivered within a 24–36-hour period. Frozen semen The well-documented decrease in longevity of frozen semen necessitates either multiple inseminations every 12–24 hours or single insemination within 12 hours pre-ovulation up to 6 hours after ovulation.7 Advantages of using an ovulatory agent are to (i) decrease the total number of examinations, (ii) concentrate the reproductive examinations around the time of predicted ovulation, (iii) choose timing of ovulation to match staffing requirements and (iv) limit the number of inseminations when mares have not ovulated within12 hours after breeding. Specific programs are discussed below.
Assessment of follicle maturation Apart from actually being in estrus, the most important aspect for determining when a follicle is mature enough to respond to ovulatory agents is the follicle size. Mares should be in estrus for ≥ 2 days and should have a soft moist cervix and ultrasonographic identification of uterine edema grade 2 or 3. Uterine edema is graded from 0 to 3, with 3 being the most obvious. Uterine edema of grade 4 is considered pathological and associated with inflammation (see Chapters 179 and 223). The most predicable responses are when mares with follicles ≤ 25 mm are given PGF2α and monitored frequently enough to attain follicle sizes listed in Table 196.1. The optimal follicle size at which to administer an ovulating agent varies according to the breed and even the number of potential
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Table 196.1 Follicle sizes recommended for programming mares to ovulate within a 24–48-h time interval according to breed and during the cycling season. The follicle sizes are guidelines only as mares do not always respond exactly as expected
Breed Thoroughbred and Standardbred Quarter Horse, Riding ponies and Arabians Miniatures Draught horse Friesian
Follicle size (mm) below which mares are unlikely to respond
Ideal follicle size (mm) to respond
Mature follicle size (mm) and may ovulate spontaneously or not respond predictably
<35
35–45
>45
<30
30–40
>40
<30 <40 <45
30–35 40–55 45–55
>35 >55 >60
pre-ovulatory follicles. Mares in estrus for several days, with large mature follicles, may spontaneously ovulate sooner than expected based solely on response to an ovulating agent. Table 196.1 lists suggested follicle size by breed for either (i) ideal size to program, (ii) unlikely to respond, or (iii) follicles that may already be destined to ovulate imminently. A specific managerial problem occurs in mares that are given PGF2α when large follicles (≥ 35 mm) are present in mid-diestrus. These follicles may ovulate quite quickly (< 48 hours), often before attaining characteristics that allow accurate programming. In 1999 we (AOM) identified a technique of using multiple low doses (1/8–1/6 typical dose given 4 times per day until uterine edema is visible) of PGF2α . The idea was to mimic the more natural endogenous prostaglandin release seen in mares in late diestrus and avoid the large and rapid drop in progesterone associated with luteal regression and exogenous PGF2α . This technique allows a more natural return to estrus and commonly allows programming of follicles once estrus is demonstrated and uterine edema identified, but before the follicle development has progressed so rapidly that ovulation is imminent or occurs unexpectedly. Another specific problem is the presence of more than one large follicle, which can make it difficult to predict if one, two, or three follicles will ovulate, or which is the dominant follicle and which is destined to ovulate first. It is usually more accurate to program using the largest follicle, unless the rapid growth of another smaller follicle has been detected. Use of ovulating agents increases the number of twins detected,4, 6 possibly by reducing the time interval between ovulations rather than ‘forcing’ more follicles to ovulate (see Chapter 241). Occasionally, a mare does not ovulate in response to an agent and subsequently develops and ovulates another follicle 4–8 days later after the original dom-
inant follicle regresses. It is likely that these mares were programmed too early to respond initially. It is difficult to detect these mares clinically before the problem occurs. Teasing patterns are often useful in this situation, and mares that normally tease well should be programmed with caution if the teasing pattern does not match the follicular size.
Ovulatory agents Only human chorionic gonadotropin (hCG) and deslorelin acetate (an analog of gonadotropinreleasing hormone; GnRH) are available in many countries and commercially as ovulatory agents. Many other agents mentioned below have demonstrated some efficacy for hastening ovulation and are briefly discussed. Human chorionic gonadotropin hCG is a large glycoprotein hormone obtained from the urine of pregnant women and freeze dried after extraction. In the horse, hCG mimics luteinizing hormone (LH) and actually increases LH after administration8 and thus can be used to control ovulation in mares and stimulate testosterone secretion in stallions.
Time of ovulation hCG will control ovulation to a time period of 36 ± 4 hours after administration to mares with a responsive follicle.9–11 It should be noted that if the percentage of mares ovulating due to hCG is normally distributed then one standard deviation means that 68% of ovulations can be expected with ± 4 hours (32–40 hours). Two standard deviations would be expected to have 95% of ovulations ± 8 hours (28–44 hours).
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Dose hCG is typically available commercially in vials of 1500, 2000, 5000, and 10 000 IU and is reconstituted immediately before use (see Chapter 188). Commonly used dose rates are 1500–3000 IU, either intravenous or intramuscular, with larger doses (> 5000 IU) associated with a decrease in fertility.12, 13 Some clinicians reduce the dose to 1500 IU as the breeding season advances. Mares have been reported to respond to doses as low as 750 IU.14
Transitional mares hCG has been demonstrated to be useful in controlling ovulation in the transitional mare.15 However, it works best with other forms of stimulation such as a combination of lights and progestagens16, 17 or dopamine antagonists. In general, hCG is not as robust as deslorelin for programming ovulation in the transition mare17 and must be used at doses of at least 2500 IU.
Frozen semen When hCG is used to control ovulations in mares bred with frozen semen, the time of administration is recommended to be approximately 12:00 midnight. Ovulation is then expected between 8 a.m. and 4 p.m. when maximum numbers of staff are available. An alternative is to administer hCG at approximately 12:00 (noon); ovulation would be expected approximately 36 hours later (12:00 midnight the following day). The mare would initially be inseminated the evening prior to the expected ovulation, and possibly a second time the next morning after ovulation has been confirmed. Cost and availability of semen will often dictate the regime chosen. When mares are bred prior to ovulation, as in the second regime above, the semen dose may be wasted if the mare does not ovulate on time or develops an abnormal follicle (see below and Chapter 222).
Advantages Advantages of hCG are a reduced cost when compared with deslorelin and a high clinical efficacy.18 hCG works very effectively when used the first time in the breeding season. It is less effective when used on multiple cycles.1, 19, 20 There is an association with increased antibody formation and multiple administration; however, it has not been clearly demonstrated that the decreased efficacy noted is due to antibody formation.21 hCG can be used to control ovulation in the late transition mare.15 Our recommendations are to use hCG on alternate cycles with
deslorelin (see below) in the normally cycling mare. Mares that are likely to respond to ovulatory agents are administered hCG first, and then if not pregnant and are rebred again, are given deslorelin the next breeding cycle.
Disadvantages Disadvantages of the use of hCG are, first, that it is of human origin. Potentially this could impact supply and purity. Second, it is less efficacious after the first administration in the breeding season. Third, it is associated with a poor response in older mares.22 GnRH GnRH is a small molecular weight decapeptide with little immunogenic properties (see Chapter 167). Most studies on use of GnRH have been performed with the analog, deslorelin acetate; however, buserelin acetate has been used with some success in cycling mares.23–25 GnRH stimulates the release of LH and follicle-stimulating hormone (FSH) from the anterior pituitary gland. Deslorelin is commercially available as OvuplantTM (2.1 mg deslorelin in a subcutaneous implant) and has been the subject of many research projects;10,26–33 more recently it has become available as compounded injectable products in slow-release formulations.34, 35 Other GnRH analogs have been demonstrated to be effective in controlling the time of ovulation but have not been released as commercial products for horses.36–42
Time of ovulation A study comparing time from administration of hCG or deslorelin to ovulation11 demonstrated that the time to ovulation was significantly different (P < 0.001) between hCG (35.9 h ± 3.8 h) compared to Ovuplant (40.7 ± 3.2 h) (Figure 196.2). These studies involved ultrasonography approximately every 2 hours until ovulation occurred. The clinical relevance of the precise timing of ovulation is associated with breeding mares with frozen semen (see below).
Dose Studies examining the dose of deslorelin have concluded that the most consistent dose was 2.2 mg. Time to first ovulation was shorter (P < 0.001) for treated mares in groups receiving 1.6 mg or 2.2 mg deslorelin, or 3000 IU hCG compared to controls (Table 196.2).10 Follicle size at assignment was not different; however, follicle size at ovulation was
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Time to ovulation 10 Number of mares ovulating
c196
hCG
8
Ovuplant
6 4 2 0 −2
23
25
27
29
31
33
35
37
39
41
43
45
47
49
Hours Figure 196.2 Number of mares ovulating in response to either hCG or Ovuplant as a function of time (values grouped into 2-hourly increments and the curve smoothed).
significantly larger in control mares (P < 0.01) (Table 196.2). Number of mares ovulating within 48 hours of treatment was greater in groups with 1.3 mg, 1.6 mg, and 2.2 mg (deslorelin) and 3000 IU hCG (19/25, 15/25, 22/23, 23/23, respectively) compared to controls (4/25) (P < 0.001). In another report, the mean time to ovulation was 68, 49, 48, 47, and 44 h in experiment 1, and 91, 66, 58, 46, and 58 h in experiment 2, respectively, for mares each given a 0, 1.2, 1.7, 2.2, or 2.7 mg deslorelin implant in the two trials. Averaged for both experiments, the proportion of mares ovulating within 48 h of treatment was 40, 75, 85, 90, and 90% for 0, 1.2, 1.7, 2.2, and 2.7 mg/mare. For both experiments, there was no effect of GnRH on pregnancy rate and there was no advantage of doses higher than 2.2 mg/mare.43 Similar results were seen in Germany with Hanoverian mares.30
Interovulatory interval Administration of deslorelin was not associated with a reduction in efficacy when used over multiple cy-
cles.44 Fertility was not affected by higher doses; however, when mares were treated with 11 mg deslorelin, a increased interovulatory interval was detected.44 Later it was identified in clinical practice that an increased interovulatory interval was occasionally identified after treatment with the normal 2.1 mg dose .45, 46 This effect has been linked to temporary downregulation of pituitary gonadotropin secretion and suppression of ovarian follicular development,47–49 and was more commonly encountered when mares were short cycled with prostaglandin after ovulation had been programmed with deslorelin, such as is commonly performed in embryo transfer programs. The prolonged interovulatory interval could be avoided by implant removal.50–52 Administration of the implant in the lip of the vulva facilitated subsequent removal 48 hours later after ovulation was detected.51 Problems of delayed interovulatory intervals have not been identified with injectable deslorelin,53 which suggested that delayed or prolonged release of deslorelin from the implants was causing the problem.
Table 196.2 Effect of treatment on mean follicle size, time to ovulation and number of mares ovulating in 48 hours after treatment with 1.3 mg deslorelin, 1.6 mg deslorelin, 2.2 mg deslorelin, 3000 IU hCG, or controls10
Group 1.3 mg deslorelin 1.6 mg deslorelin 2.2 mg deslorelin 3000IU hCG Controls
Follicle size (mm) at assignment
Follicle size (mm) at ovulation
Time to ovulation (days)
Percent of mares ovulating in 48 hours
33.5 34.1 33.0 33.8 33.0
41.8bc 38.3ac 38.5ac 37.5a 43.4b
2.68ab 2.20a 1.98a 1.88a 3.52b
76% (19/25)a 60% (15/25)ac 96% (22/23)ad 100% (23/23)ad 16% (4/25)b
abc (P < 0.01)
ab (P < 0.001)
ab (P < 0.001) cd (P < 0.02)
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Transitional mares Final maturation and control of ovulation can be achieved in the transitional mare with deslorelin.54 In one study using mares in the early transition period (follicles < 15 mm), an apparent effect of administration of 2.1 mg deslorelin every other day in hastening ovulation (ovulation in < 10 days) in treated mares (6/11) versus controls (0/10) was demonstrated. However, the mean ovulation date was not different between groups, so there was no overall advancement of the transition.54 The same research group reported the ability of the implants to hasten ovulation late in the transition, when mares were treated every other day after reaching a follicle size of 30 mm. The average number of implants needed was 2.1 and 8/10 mares ovulated within 4 days of first deslorelin implant versus none of the controls. 54 Treating mares with deslorelin during the transition period is not associated with an increase in the number of multiple ovulations.55 Other regimes involving deslorelin in combination with progestagens have been reported.56–58 It is not recommended to use deslorelin to ‘force’ mares through the transition period from anestrus or the early transition, as such treatment is associated with poor results59 and failure of CL maintenance.
When not restricted by frozen semen availability it may be easier (without sacrificing pregnancy rates) to breed twice per cycle: first at 24 hours after the ovulatory agent is given; then second at 42 hours, after ovulation has been confirmed. If the mare ovulates 18–54 hours after administration of the ovulatory agent this will result in semen being inside the mare and available to fertilize from 12 hours prior to 6 hours after ovulation.60
Advantages
r r r r r r r
Predictable response; Robust treatment that probably overcomes some poor programming decisions; Ovulates multiple follicles closer to each other and therefore improves detection of twins or ability to obtain multiple embryos; Synthetic and inexpensive to manufacture; Can be used in consecutive cycles or multiple cycles during a breeding season; Can be used the following cycle after hCG; Different mechanism of action than hCG; stimulates secretion of endogenous LH which subsequently induces follicle maturation and ovulation.
Disadvantages Frozen semen It is advantageous to have ovulation occur in a predictable time period when breeding mares with frozen semen. Two options for deslorelin administration and subsequent insemination are suggested. First, one may administer deslorelin at 7:00 p.m. (1900 h) and expect an ovulation approximately 40–41 hours later (approximately 12:00 noon; 1200 h). A second option is to administer deslorelin at approximately 8:00 a.m. (0800 h) and expect an ovulation around 12:00 midnight (2400 h) the following day. Insemination(s) can be performed prior to the predicted ovulation and/or after ovulation is confirmed. Of course the programming of mares to ovulate at a precise time does not obviate the veterinarian from continuing vigilant monitoring. Not all mares respond as predicted. Breeding with frozen semen once immediately after ovulation occurs is advantageous when one dose of semen is available for that cycle and when there is a possibility that mares may not ovulate normally, such as when anovulatory hemorrhagic follicles occur (AHFs, also termed persistent anovulatory follicles orPAFs) (see Chapter 222). In such cases it may not be prudent to breed before ovulation has been confirmed.
r r
Expensive to purchase commercially relative to hCG; Prolonged interovulatory interval noted with the implant in some mares.
Other agents Other agents that may have efficacy in controlling the time of ovulation include:
Prostaglandins Prostaglandin F2α (PGF2α ) was reported to be capable of inducing ovulation.61 Fenprostalene (250 µg) or saline was given at 60 h after the onset of estrus in alternate estrous periods to eight mares for four cycles, and the interval from treatment to ovulation was significantly decreased (41.25 vs 73.50 h; P < 0.001) as was the duration of estrus (5.63 vs 6.88 days; P < 0.005).61 Despite these encouraging results, further work has failed to find any other form of PGF2α that could perform similarly.62 It is possible that the long action of the particular form of PGF2α used (Fenprostalene) was responsible for the result, as PGF2α (Luprostiol) has subsequently been shown to induce an LH release from the anterior pituitary gland.63
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Induction of Ovulation Recombinant equine luteinizing hormone An exciting new development is recombinant equine luteinizing hormone (reLH) which is a novel singlechain LH analog in which the equine α-subunit is directly linked to the equine LH β-subunit.64 In-vitro studies demonstrated that the biological activity of the recombinant hormone is similar to that of natural equine LH, as incubation of equine Leydig cell cultures with reLH stimulated a dose-dependent increase in testosterone production.65 Administration of reLH to stallions caused an increase in plasma testosterone concentrations within 15 minutes of treatment.65 Testosterone levels returned to baseline within 24 to 48 hours after treatment. Repeated administration of reLH to a small number of mares was not associated with development of antibodies.65 Increasing doses of reLH (0.3, 0.6, 0.75, and 0.9 mg) showed increasing effectiveness at inducing ovulation in mares within 48 h of treatment. Treatments with the 0.3, 0.6, 0.75, and 0.9 mg doses of reLH resulted in 28.6% (2/7), 50% (10/20), 90% 9/10), and 80% 16/20) ovulation rates within 48 hours, respectively, which were similar to 2500 IU hCG treatment (85.7%, 6/7).64 It was concluded that reLH was a reliable and effective ovulatory agent that does not significantly alter endogenous hormone profiles or affect interovulatory intervals.64 Any synthetic product that can be produced inexpensively that removes the need of using human supply sources (hCG-urine from pregnant women) offers much potential.
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of equine reproductive physiology and even treatment regimes to induce ovulation in cycling mares, and perhaps techniques to truncate the transitional breeding season.
Equine pituitary extract Historically treatment with equine pituitary extract (EPE) has been shown to increase multiple follicular development and ovulation rates69–71 and induction of ovulation.72 EPE is a mixture of LH and FSH in varying concentrations at different times of the year, and extracted from pituitaries collected from abattoirs. It was replaced by a recombinant form of FSH (eFSH)73–75 for superovulation but is currently not being produced commercially. Some problems with using crude equine pituitary extract for induction of ovulation are, first, an approved pituitary extract is not commercially available. Second, there is potential for prion transmission as transmissible spongiform encephalopathies (TSEs), for example, bovine spongiform encephalopathies (BSE), scrapie in sheep and kuru in humans,76–78 are associated with animal to animal, animal to human, and human to human transmission of neural proteins. There is much we do no understand about prion transmission and further experiments using EPE should be regarded as controversial.
Factors that may affect results Kisspeptin
Time of year
It has recently been revealed that the hormone/neuropeptide, kisspeptin, exerts a major influence in the control of GnRH secretion. This molecule, first described as having a crucial role in triggering the onset of puberty, is involved in all phases of reproductive life. Kisspeptin (KiSS) strongly induces the secretion of gonadotropins in many species, mainly through stimulation of GnRH secretion.66 Few studies have been reported in the horse; however, one report demonstrated a hastening of ovulation (defined as ovulating within 48 h of treatment) in pony mares when treated with 10 mg kisspeptin versus saline controls (6/8 versus 2/8, P < 0.07).67 Another report identified that administration of 500 µg and 1.0 mg kisspeptin (rKP-10) elicited peak, mean, and area under the curve LH and FSH responses indistinguishable to that of 25 µg GnRH, although a single intravenous injection of 1.0 mg kisspeptin was insufficient to induce ovulation in the estrous mare.68 It is possible that further research with kisspeptin may result in a better understanding
The most consistent response is when mares are programmed to ovulate during the natural cycling season. Restrictions from breed registries that encourage foals to be born from breeding of mares not in the normal physiological cycling season mean that many programming events occur during transitional estrus. Once mares have had their first ovulation of the year to enter the cycling season, their response to programming agents will be more predictable. In addition, one study demonstrated mares that responded better to deslorelin as the season progressed.79 Ovulatory agent There are considerable differences in ability of ovulatory agents to achieve the desired effect that are related to dose and specific ovulatory agent. In general, deslorelin is more reliable than hCG, and smaller follicles will respond to deslorelin better than hCG. In addition, mares with a late transitional follicle are more likely to respond when deslorelin is
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administered rather than hCG. Specific studies are lacking for compounded injectable deslorelin.
Age of the mare Older mares are less likely to respond accurately to any ovulatory agent, and both hCG and deslorelin have been demonstrated not as efficacious in older mares.22, 79
Size and maturity of the follicle Size of the follicle does not always equate to maturity, of which a specific example is the large middiestrus follicle that will not respond to ovulating agents while progesterone levels are elevated.80 Often mares are presented with large follicles and minimal uterine edema visible with ultrasonography. It is useful with these mares to examine the vagina for mucosal color and degree of softness of the cervix. However, teasing behavior is critical to making accurate decisions with these mares. Occasionally, mares do not ovulate despite ultrasonographic recognition of uterine edema, a soft cervix, and a typical sized follicle that could be expected to respond. Often in this situation, the follicle will continue to grow and will respond to administration of the same or a different ovulatory agent 48 hours or more after the initial dose. Mares presenting with small follicles that have good uterine edema and have been demonstrating good teasing behavior are also difficult to program. Prior history of the mare’s cycle may be useful in determining if the mare is likely to ovulate a small follicle and thus respond to an ovulatory agent. On occasion, mares in estrus are examined that already have large dominant follicles. It may be difficult to predict the ovulation response of these mares to either hCG or deslorelin. These mares may ovulate spontaneously within a few hours, or may ovulate in the normal interval in response to an ovulating agent. In these instances we would normally program the mare, as it is surprising how often they will ovulate within the predicted ‘ovulatory window’. In any case, more examinations after programming are necessary if frozen semen is being used to ensure mares are bred within 12 hours before and 6 hours after ovulation.
Programming time We visit most breeding farms at the same or a similar time every day. As deslorelin is most likely to cause mares to ovulate within a 41 ± 3 hour window,11 if mares are programmed when bred in the
evening (7 p.m.) and then examined two days later in the morning (7 a.m.), only 36 hours have elapsed since programming and many mares will have not ovulated at that time. Enthusiastic farm managers often need to be restrained from wanting to rebreed the mare that has not ovulated at 36 hours. The wishes of stud managers to rebreed every 48 hours is annoying when the stallion’s fertility is normal, especially if compounded by a lack of understanding due to communication breakdown. Fertility of normal stallions can be expected to be maintained even if the mare does not ovulate until 3 days after last breeding (see Chapter 118). It is not uncommon for stallions to achieve pregnancies when the interval from breeding to ovulation is as long as 5 or more days. Multiple follicles It is not always possible to predict which one or all of two or three large follicles will ovulate when mares are programmed. On occasion, a mare in estrus with good uterine folds will have a 30-mm follicle on one ovary and two 35- to 40-mm follicles on the other ovary, and yet the smaller follicle will ovulate within 48 hours of programming and the others will not ovulate and gradually recede. More often than not, this occurs in cases where we do not have the ability to examine the mare frequently enough to identify the accurate growth pattern of follicles. Another similarly frustrating scenario is when mares are programmed and bred and then do not ovulate the intended follicle, which then gradually decreases in size and, despite the mare staying in estrus, the next follicle may take a week to grow to a size that the mare can be rebred. Fortunately those problems do not happen very often, as most mares conform to the expected patterns of ovulation.
References 1. Samper JC. Induction of estrus and ovulation: why some mares respond and others do not. Theriogenology 2008;70(3):445–7. 2. McCue PM, Troedsson MH, Liu IK, Stabenfeldt GH, Hughes JP, Lasley BL. Follicular and endocrine responses of anoestrous mares to administration of native GnRH or a GnRH agonist. J Reprod Fertil Suppl 1991;44:227–33. 3. Bergfelt DR, Ginther OJ. Embryo loss following GnRHinduced ovulation in anovulatory mares. Theriogenology 1992;38:33–43. 4. McKinnon AO. Reproductive efficiency of horses in Australia. http://www.gvequine.com.au/breeding efficiency.htm. 2000. 5. Nath LC, Anderson GA, McKinnon AO. Reproductive efficiency of Thoroughbred and Standardbred horses in northeast Victoria. Aust Vet J 2010;88:169–75. 6. Perkins NR, Grimmett JB. Pregnancy and twinning rates in Thoroughbred mares following the administration
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of human chorionic gonadotropin (HCG). N Z Vet J 2001;49(3):94–100. Loomis PR, Squires EL. Frozen semen management in equine breeding programs. Theriogenology 2005;64(3):480–91. Ginther OJ, Almamun M, Shahiduzzaman AK, Beg MA. Disruption of the periovulatory LH surge by a transient increase in circulating 17beta-estradiol at the time of ovulation in mares. Anim Reprod Sci 2010;117(1–2):178–82. Sullivan JJ, Parker WG, Larson LL. Duration of estrus and ovulation time in nonlactating mares given HCG during three successive estrous periods. J Am Vet Med Assoc 1973;163:895–8. McKinnon AO, Nobelius AM, Figueroa STD, Skidmore J, Vasey JR, Trigg TE. Predictable ovulation in mares treated with an implant of the GnRH analog deslorelin. Equine Vet J 1993;25:321–3. McKinnon AO, Perriam WJ, Lescun TB, Walker J, Vasey JR. Effect of a GnRH analogue (Ovuplant), hCG and dexamethasone on time to ovulation in cycling mares. World Equine Vet Rev 1997;2(3):16–18. Voss JL. Human chorionic gonadotropin. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, London: Lea & Febiger, 1993: pp. 325–8. Shilova AV, Platov EM, Lebedev SG. The use of human chorionic gonadotrophin for ovulation date regulations in mares. Int Cong Anim Reprod Artif Insem 1976;3:204–7. Davies Morel MC, Newcombe JR. The efficacy of different hCG dose rates and the effect of hCG treatment on ovarian activity: ovulation, multiple ovulation, pregnancy, multiple pregnancy, synchrony of multiple ovulation in the mare. Anim Reprod Sci 2008;109(1–4):189–99. Carnevale EM, Squires EL, McKinnon AO, Harrison LA. Effect of human chorionic gonadotropin on time to ovulation and luteal function in transitional mares. J Equine Vet Sci 1989;9:27–9. Colbern GT, Squires EL, Voss JL. Use of altrenogest and human chorionic gonadotropin to induce normal ovarian cyclicity in transitional mares. J Equine Vet Sci 1987;7:69–72. Cuervo-Arango J, Clark A. The first ovulation of the breeding season in the mare: the effect of progesterone priming on pregnancy rate and breeding management (hCG response rate and number of services per cycle and mare). Anim Reprod Sci 2010;118(2–4):265–9. Barbacini S, Zavaglia G, Gulden P, Marchi V, Necchi D. Retrospective study on the efficacy of hCG in an equine artificial insemination programme using frozen semen. Equine Vet Educ 2000;12(6):312–17. McCue PM, Hudson JJ, Bruemmer JE, Squires EL. Efficacy of hCG at inducing ovulation: a new look at an old issue. Proc Am Assoc Equine Pract 2004;50:510–13. Wilson CG, Downie CR, Hughes JP, Roser JF. Effects of repeated hCG injections on reproductive efficiency in mares. J Equine Vet Sci 1990;10:301–8. Roser JF, Kiefer BL, Evans JW, Neely DP, Pacheco DA. The development of antibodies to human chorionic gonadotrophin following its repeated injection in the cyclic mare. J Reprod Fertil Suppl 1979;( 27): 173–9. Carnevale EM. The mare model for follicular maturation and reproductive aging in the woman. Theriogenology 2008;69(1):23–30. Camillo F, Pacini M, Panzani D, Vannozzi I, Rota A, Aria G. Clinical use of twice daily injections of buserelin acetate to induce ovulation in the mare. Vet Res Commun 2004;28(Suppl 1):169–72.
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24. Barrier-Battut I, Le Poutre N, Trocherie E, Hecht S, Grandchamp des Raux A, Nicaise JL, V´erin X, Bertrand J, Fi´eni F, Hoier R, Renault A, Egron L, Tainturier D, Bruyas JF. Use of buserelin to induce ovulation in the cyclic mare. Theriogenology 2001;55(8):1679–95. 25. Levy I, Duchamp G. A single subcutaneous administration of buserelin induces ovulation in the mare: field data. Reprod Domest Anim 2007;42(5):550–4. 26. Blanchar TL, Brinsko SP, Rigby SL. Effects of deslorelin or hCG administration on reproductive performance in first postpartum estrus mares. Theriogenology 2002;58(1):165–9. 27. Farquhar VJ, McCue PM, Vanderwall DK, Squires EL. Efficacy of the GnRH agonist deslorelin acetate for inducing ovulation in mares relative to age of mare and season. J Equine Vet Sci 2000;20(11):722–5. 28. Hemberg E, Lundeheim N, Einarsson S. Successful timing of ovulation using deslorelin (Ovuplant) is labour-saving in mares aimed for single AI with frozen semen. Reprod Domest Anim 2006;41(6):535–7. ¨ 29. Jochle W, Trigg TE. Control of ovulation in the mare with OvuplantTM : a short-term release implant (STI) containing the GnRH analog deslorelin acetate – studies from 1990 to 1994. J Equine Vet Sci 1994;14:632–44. 30. Meinert C, Silva JFS, Kroetz I, Klug E, Trigg TE, Hoppen ¨ HO, Jochle W. Advancing the time of ovulation in the mare with a short-time implant releasing the GnRH analogue deslorelin. Equine Vet J 1993;25:65–8. ¨ 31. Meyers PJ, Jochle W, Trigg TE. Acceleration of ovulation in the mare with the use of the GnRH analog, deslorelin acetate. Proc Am Assoc Equine Pract 1994;40:13–14. 32. Morehead JA, Blanchard TL. Clinical experience with deslorelin (OvuplantTM ) in a Kentucky Thoroughbred broodmare practice (1999). J Equine Vet Sci 2000;20(6):358. 33. Stich KL, Wendt KM, Blanchard TL, Brinsko SP. Effects of a new injectable short-term release deslorelin in foal-heat mares. Theriogenology 2004;62(5):831–6. 34. Burns PJ, Fleury JJ, Gibson J, Sullivan S, Tipton A. Evaluation of the SaberTM delivery system for the controlledrelease of deslorelin acetate for advancing ovulation in the mare – effects of formulation and dose. Theriogenology 1998;49(1):256. 35. Donadeu FJ, Thompson DL, Burns PJ, Tipton AJ, Gibson JW, Swaim RP, Kincaid LA, Leise BS. Pharmacodynamic evaluation of the SABERTM delivery system for the controlled release of the GnRH analog, deslorelin acetate, for the control of ovulation in the mare. Proceedings of the Fifteenth Equine Nutrition & Physiology Symposium 1997; pp. 102–3. 36. Allen WR, Sanderson MW, Greenwood RE, Ellis DR, Crowhurst JS, Simpson DJ, Rossdale PD. Induction of ovulation in anoestrous mares with a slow-release implant of a GnRH analogue (ICI 118 630). J Reprod Fertil Suppl 1987;35:469–78. 37. Cordeiro JM, Fonseca FA, Motta VAF, Espeschit CJB, Torres CAA. Effect of a GnRH analog (buserelin) and hCG on estrus, ovulation and fertility of crossbred mares after treatment with PGF-2-alpha. Revista Brasileira De Reproducao Animal 1989;13:143–50. 38. Fitzgerald BP, Meyer SL, Affleck KJ, Silvia PJ. Effect of constant administration of a gonadotropin-releasinghormone agonist on reproductive activity in mares: induction of ovulation during seasonal anestrus. Am J Vet Res 1993;54:1735–45. 39. Greaves HE, Kalariotes V, Cleaver BD, Porter MB, Sharp DC. Effects of ovarian input on GnRH and LH secretion
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immediately postovulation in pony mares. Theriogenology 2001;55(5):1095–106. Harrison LA, Squires EL, Nett TM, McKinnon AO. Use of gonadotropin-releasing hormone for hastening ovulation in transitional mares. J Anim Sci 1990;68:690–9. Irvine DS, Downey BR, Parker WG, Sullivan JJ. Duration of oestrus and time of ovulation in mares treated with synthetic Gn-RH (AY-24,031). J Reprod Fertil Suppl 1975;23:279–83. Mumford EL, Squires EL, Peterson KD, Nett TM, Jasko DJ. Effect of various doses of a gonadotropin-releasinghormone analog on induction of ovulation in anestrous mares. J Anim Sci 1994;72:178–83. Squires EL, Moran DM, Farlin ME, Jasko DJ, Keefe TJ, ¨ Meyers SA, Figueiredo E, McCue PM, Jochle W. Effect of dose of GnRH analog on ovulation in mares. Theriogenology 1994;41(3):757–69. ¨ Mumford EL, Squires EL, Jochle E, Harrison LA, Nett TM, Trigg TE. Use of deslorelin short-term implants to induce ovulation in cycling mares during 3 consecutive estrous cycles. Anim Reprod Sci 1995;39:129–40. Morehead JA, Blanchard TL. Clinical experience with deslorelin (OvuplantTM ) in a Kentucky Thoroughbred broodmare practice (1999). J Equine Vet Sci 2000;20(6): 358. Vanderwall DK, Juergens TD, Woods GL. Reproductive performance of commercial broodmares after induction of ovulation with HCG or OvuplantTM (deslorelin). J Equine Vet Sci 2001;21(11):539–42. Farquhar VJ, McCue PM, Nett TM, Squires EL. Effect of deslorelin acetate on gonadotropin secretion and ovarian follicle development in cycling mares. J Am Vet Med Assoc 2001;218(5):749–52. Johnson CA, Thompson DL, Kulinski KM, Guitreau AM. Prolonged interovulatory interval and hormonal changes in mares following the use of OvuplantTM to hasten ovulation. J Equine Vet Sci 2000;20(5):331–6. Johnson CA, Thompson DL, Jr., Cartmill JA. Pituitary responsiveness to GnRH in mares following deslorelin acetate implantation to hasten ovulation. J Anim Sci 2002;80(10):2681–7. Farquhar VJ, McCue PM, Carnevale EM, Nett TM, Squires EL. Deslorelin acetate (Ovuplant) therapy in cycling mares: effect of implant removal on FSH secretion and ovarian function. Equine Vet J 2002;34(4):417–20. McCue PM, Niswender KD, Farquhar VJ. Induction of ovulation with deslorelin: Options to facilitate implant removal. J Equine Vet Sci 2002;22(2):54–5. McCue PM, Farquhar VJ, Carnevale EM, Squires EL. Removal of deslorelin (Ovuplant) implant 48 h after administration results in normal interovulatory intervals in mares. Theriogenology 2002;58(5):865–70. McCue PM, McGee CD, Gee EK. Comparison of compounded deslorelin and hCG for induction of ovulation in mares. J Equine Vet Sci 2007;27:58–61. McKinnon AO, Vasey JR, Lescun TB, Trigg TE. Repeated use of a GnRH analogue deslorelin (Ovuplant) for hastening ovulation in the transitional mare. Equine Vet J 1997;29(2):153–155. Raz T, Carley S, Card C. Comparison of the effects of eFSH and deslorelin treatment regimes on ovarian stimulation and embryo production of donor mares in early vernal transition. Theriogenology 2009;71(9):1358–66. ¨ Arbeiter K, Barth U, Jochle W. Observations on the use of progesterone intravaginally and of deslorelin STI in
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acyclic mares for induction of ovulation. J Equine Vet Sci 1994;14:21–5. ¨ Newcombe JR, Handler J, Klug E, Meyers PJ, Jochle W. Treatment of transition phase mares with progesterone intravaginally and with deslorelin or HCG to assist ovulations. J Equine Vet Sci 2002;22(2):57–64. Newcombe JR. Anestrus in mares – experiences using Ovuplant, a short-term implant of the GnRH analog deslorelin, following implantation with Cidr-B, a progesterone releasing intravaginal device. Irish Vet J 1998;51(11): 582–5. Nickerson KC, McCue PM, Squires EL, Nett TM. Comparison of 2 dosage regimens of the GnRH agonist deslorelin acetate on inducing ovulation in seasonally anestrous mares. J Equine Vet Sci 1998;18(2):121–4. Loomis PR. The equine frozen semen industry. Anim Reprod Sci 2001;68(3–4):191–200. Savage NC, Liptrap RM. Induction of ovulation in cyclic mares by administration of a synthetic prostaglandin, fenprostalene, during oestrus. J Reprod Fertil Suppl 1987;35:239–43. Harrison LA, Squires EL, McKinnon AO. Comparison of hCG, Burserelin and Luprostiol for induction of ovulation in cycling mares. J Equine Vet Sci 1991;11:163–6. ¨ Jochle W, Irvine CH, Alexander SL, Newby TJ. Release of LH, FSH and GnRH into pituitary venous blood in mares treated with a PGF analogue, luprostiol, during the transition period. J Reprod Fertil Suppl 1987;35:261–7. Yoon MJ, Boime I, Colgin M, Niswender KD, King SS, Alvarenga M, Jablonka-Shariff A, Pearl CA, Roser JF. The efficacy of a single chain recombinant equine luteinizing hormone (reLH) in mares: induction of ovulation, hormone profiles, and inter-ovulatory intervals. Domest Anim Endocrinol 2007;33(4):470–9. Jablonka-Shariff A, Roser JF, Bousfield GR, Wolfe MW, Sibley LE, Colgin M, Boime I. Expression and bioactivity of a single chain recombinant equine luteinizing hormone (reLH). Theriogenology 2007;67(2):311–20. Caraty A, Franceschini I. Basic aspects of the control of GnRH and LH secretions by kisspeptin: potential applications for better control of fertility in females. Reprod Domest Anim 2008;43 Suppl 2:172–8. Briant C, Schneider J, Guillaume D, Ottogalli M, Duchamp G, Bruneau B, Caraty A. Kisspeptin induces ovulation in cycling Welsh pony mares. Anim Reprod Sci 2006;94:217–19. Magee C, Foradori CD, Bruemmer JE, Arreguin-Arevalo JA, McCue PM, Handa RJ, Squires EL, Clay CM. Biological and anatomical evidence for kisspeptin regulation of the hypothalamic-pituitary-gonadal axis of estrous horse mares. Biomed Tech (Berlin) 2009;150(6):2813–21. Alvarenga MA, McCue PM, Bruemmer J, Neves N, Jr., Squire EL. Ovarian superstimulatory response and embryo production in mares treated with equine pituitary extract twice daily. Theriogenology 2001;56(5):879–87. Palmer E, Hajmeli G. Superovulation in mares. Recueil De Medecine Veterinaire De L’Ecole D’Alfort 1992;168:897–905. Squires EL, Garcia RH, Ginther OJ, Voss JL, Seidel GEJ. Comparison of equine pituitary extract and FSH for superovulating mares. Theriogenology 1986;26:661–70. Duchamp G, Bour B, Combarnous Y, Palmer E. Alternative solutions to hCG induction of ovulation in the mare. J Reprod Fertil Suppl 1987;35:221–8. Logan NL, McCue PM, Alonso MA, Squires EL. Evaluation of three equine FSH superovulation protocols in mares. Anim Reprod Sci 2007;102(1–2):48–55.
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Induction of Ovulation 74. McCue PM, LeBlanc MM, Squires EL. eFSH in clinical equine practice. Theriogenology 2007;68(3):429– 33. 75. Squires EL, McCue PM. Superovulation in mares. Anim Reprod Sci 2007;99(1–2):1–8. 76. Zou WQ, Gambetti P. Prion: the chameleon protein. Cell Mol Life Sci 2007;64(24):3266–70. 77. Chakraborty C, Nandi S, Jana S. Prion disease: a deadly disease for protein misfolding. Curr Pharm Biotechnol 2005;6(2):167–77.
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78. Goldfarb LG, Cervenakova L, Gajdusek DC. Genetic studies in relation to kuru: an overview. Curr Mol Med 2004;4(4):375–84. 79. Farquhar VJ, McCue PM, Vanderwall DK, Squires EL. Efficacy of the GnRH agonist deslorelin acetate for inducing ovulation in mares relative to age of mare and season. J Equine Vet Sci 2000;20(11):722–5. 80. Glazar BS, McCue PM, Bruemmer JE, Squires EL. Deslorelin on Day 8 or 12 postovulation does not luteinize follicles during an artificially maintained diestrous phase in the mare. Theriogenology 2004;62(1–2):57–64.
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Chapter 197
Synchronization of Ovulation Etta A. Bradecamp
Introduction Manipulation of the equine estrous cycle to control the timing of estrus and subsequently ovulation is performed most frequently for breeding management purposes. Unlike the bovine industry in which the synchronization of estrus is utilized to induce simultaneous estrus in a large number of cows, in the equine industry the necessity to control the estrous cycle in an individual or small number of mares is much more common. The cow’s short estrus is closely associated with ovulation and lends itself well to estrus synchronization programs that reliably yield a large percentage of the treated animals displaying estrus and ovulating within a narrow window of time. This close synchronization allows for the use of timed insemination protocols, and the ability to breed a large number of cows over a very short period of time, thereby minimizing labor and time. Owing to the longer period of estrus and the greater variability in time of ovulation as it relates to estrus, the mare’s reproductive cycle does not lend itself to close synchronization within a large group of animals. There are fewer synchronization protocols available for the mare than for the cow, and a complete understanding of the mare’s estrous cycle is necessary to obtain reliable results.
ing bred with cooled transported semen to minimize the number of semen shipments required and minimize the costs incurred by repeat collections, shipment, and inseminations, (v) managing mares being bred with frozen semen to reduce the labor and costs associated with the repeat examinations required to insure correct timing of insemination as it relates to ovulation, (vi) managing mares that benefit from a single insemination timed closely to ovulation owing to their tendency to develop postbreeding endometritis, and (vii) allowing insemination of a group of mares over a short period of time to minimize labor and costs.1–3 Embryo transfer programs In equine reproduction, a common use of ovulation synchronization among a group of mares is in embryo transfer programs. If a large recipient herd providing mares that have ovulated in synchrony with the donor is not available, then a small group of recipient mares must be synchronized with the donor mare. To be suitable recipients, these mares must ovulate from 1 day before to 3 days after the date of ovulation of the donor mare. There is one report in the literature that recipient mares that ovulated up to 5 days after the donor have been used successfully in a large embryo transfer program.4
Indications for estrus synchronization
Stallion availability
The most common applications of estrus synchronization in the mare include (i) synchronization of donor and recipient mares in an embryo transfer program, (ii) scheduling breedings to a stallion with limited availability, (iii) scheduling the breedings of a large number of mares to a stallion with a full book of mares to minimize the number of covers per mare and thereby maximize the number of mares that can be bred to a given stallion, (iv) managing mares be-
A second use of estrus synchronization is to synchronize a group of mares for breeding since the stallion’s availability is limited to only specific periods of time during the breeding season (e.g., owing to a show schedule). Unlike in the bovine industry, in which synchronization of estrus has been used successfully to time the breeding of a large group of cows over a very short period of time, synchronization of the mare’s estrous cycle is much more difficult.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Synchronization of Ovulation Compared with the cow, the length of estrus in the mare is much longer (5–7 days) and the time of ovulation relative to the end of estrus varies with ovulation occurring on the last 2 days of estrus in 69% of mares and after the end of estrus in 14% of mares.5 These factors make the development of a synchronization program that allows for a single timed breeding of a group of mares difficult. However, combined with ultrasonographic monitoring of the mare’s reproductive tract there are established hormone therapy protocols that are useful in controlling the onset of estrus and time of ovulation so that timed breedings can occur. Semen availability In addition to synchronization of estrus in a group of mares, pharmacological agents can be used to induce ovulation to allow for the synchronization of breeding with ovulation. With the use of both frozen and cooled transported semen it is often desirable to use only one dose of semen; therefore, it is essential that ovulation occurs no more than 24 hours after breeding with cooled transported semen and no more than 12 hours before to 6 hours after ovulation when frozen semen is used. When live cover of mares is performed and the stallion has a full book of mares, it is also desirable to breed the mare only once during a cycle, thereby maximizing stallion power. With close monitoring of the mare’s estrous cycle via transrectal ultrasonography and the use of pharmacological agents the synchronization of ovulation with breeding can be achieved.
Methods of estrus synchronization There are four basic methods used to achieve estrus synchronization in the mare: (i) termination of the luteal phase, (ii) lengthening of the luteal phase, (iii) induction of ovulation, and (iv) inhibition of the follicular phase. The pharmacological agents most commonly used to accomplish these methods are prostaglandins, progestins, human chorionic gonadotropin (hCG; Chorulon, Intervet Inc., Millsboro, DE), deslorelin acetate implant (Ovuplant; Ovuplant, Fort Dodge Animal Health, Fort Dodge, IA), deslorelin injectable, recombinant equine luteinizing hormone (reLH), or estradiol-17β. The most reliable and successful estrus synchronization programs are achieved when a combination of these methods and pharmacological agents are used. Hormonal therapies
Prostaglandins Prostaglandin F2α (PGF2α ) is used to terminate the luteal phase by causing lysis of the corpus luteum
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(CL), thereby allowing the return to estrus. The most common prostaglandins used are dinoprost (Lutalyse, Pharmacia Animal Health, Kalamazoo, MI) and cloprostenol (Estrumate, Schering-Plough Animal Health Corporation, Union, NJ). Dinoprost is a naturally occurring PGF2α and cloprostenol is a synthetic prostaglandin analog. For prostaglandins to effectively terminate the luteal phase a mature CL must be present that is responsive to prostaglandin. The equine CL exhibits incomplete sensitivity to PGF2α until approximately days 5–6 after ovulation in most mares.6–9 The standard recommended dose of dinoprost is 9 µg/kg (5–10 mg per 1000 lb/454 kg horse). The recommended dose of cloprostenol is 250 µg per 1000 lb/454 kg horse (0.55 µg/kg). More recent research has shown that much lower doses of prostaglandin are just as effective at lysing the CL with fewer negative side-effects. As early as 1975 Douglas and Ginther10 demonstrated that doses of the prostaglandin dinoprost as low as 1.25 mg were as effective at inducing luteolysis as more standard doses. In 2001 Nie et al.11 showed that a microdose of cloprostenol (25 µg) was just as effective at lysing the equine CL as the larger standard dose. One year later in 2002, Irvine et al.12 demonstrated that two doses of 0.5 mg of the prostaglandin dinoprost, 24 hours apart, caused lysis of the corpus luteum in 100% of treated mares. On average, mares return to estrus and ovulate 5–7 and 9–11 days, respectively, after administration of prostaglandin in the presence of a mature CL. However, since the follicular phase plays a greater role in controlling the total length of the estrous cycle in the mare, the length of time from administration of prostaglandin to onset of estrus and ovulation can be quite variable depending on the size and status of follicles present on the ovaries at the time of administration.13 The time of ovulation after PGF2α -induced estrus may vary as much as 2–15 days post treatment.7–9,14–16 Administration of prostaglandin when a large follicle is present may result in the mare returning to estrus and ovulating in 2–3 days. Some of these mares never develop overt signs of standing heat, endometrial folds, or relaxation of the cervix prior to ovulation.15, 17, 18 If a large follicle destined for atresia is present on the ovary at the time of prostaglandin administration, the follicle may not ovulate and instead regress as a new wave of follicles develop. This scenario will result in the time from prostaglandin administration to ovulation being more prolonged as a new wave of follicles is recruited. In a number of farm species prostaglandins have been utilized to synchronize estrous cycles. The protocol is to administer prostaglandin to all of the females simultaneously without any knowledge
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of their stages in the estrous cycle.13 Only those females that have mature, responsive corpora lutea will respond and return to estrus. Enough time is allowed for this group of females to ovulate and develop mature corpora lutea prior to the second injection of prostaglandin being administered. By this time the rest of the females that did not respond to the first injection of prostaglandin should have mature responsive corpora lutea.13 Theoretically, all of the females should respond to the second dose of prostaglandin and return to estrus. In cattle, the administration of two doses of prostaglandin 11– 12 days apart results in approximately 80% exhibiting estrus within 2–5 days after the second injection.19–22 Since cows exhibit estrus approximately 24 hours prior to ovulation this protocol provides fairly tight synchrony. While cattle respond to luteolytic doses of prostaglandin over more than half of their cycle length,23 mares typically respond to only a single luteolytic dose of prostaglandin over approximately 30% of their cycle, from approximately day 6 to day 14.7–9,13 While there have been reports of mares treated with prostaglandins 1–2 days after ovulation having decreased progesterone levels (and sometimes returning to estrus), this does not occur in all mares. This suggests that the CL can be responsive to prostaglandins in some mares earlier than traditionally perceived and when multiple doses are used.12, 24, 25 Since cattle are responsive to prostaglandin over a longer period of time during their estrous cycle this protocol works well for synchronizing estrus in a herd of cattle; however, owing to the mare’s narrow range of responsiveness, only a small amount of variation in response to the first injection can be accommodated for at the time of the second injection.13 A mare that responds very quickly to the first injection or has just ovulated and therefore does not respond at all may already be returning to estrus on its own at the time of the second injection, whereas a mare that takes 11 days to ovulate after the first injection of prostaglandin will not have a responsive CL at the time of the second injection.13 If this method of estrus synchronizaton is utilized in mares, an interval of 14–15 days between prostaglandin injections has been found to be optimal to achieve the most reliable results.14 An interval that is any shorter than this will result in some mares not being responsive to either injection, and an interval greater than 15 days results in a group of mares that were too early to respond to the first injection and are already in estrus and getting close to ovulating on their own at the time of the second injection. Approximately 90% of mares will display signs of estrus by day 6 after the second injection.26, 27 While the administration of two doses of prostaglandin 14 days apart has been described
in the literature as a protocol for estrus synchronization in mares, it is not a very reliable method of ensuring that a recipient mare will be synchronized with a donor.14 Research has shown that, after two injections of prostaglandin, on average ovulation occurs between 7 and 10 days after the second injection,8, 14, 28 with ovulation potentially occurring anywhere between 0 and 17 days.7, 14, 28 In a review of the literature, it was stated that, when this protocol was administered, one would need to treat at least 10 recipient mares to have an 80% chance that at least one recipient would ovulate within 24 hours of the donor.29 Obviously, this is not ideal, especially when in most small embryo transfer programs there are only a few mares available to be used as recipients. Owing to the large variation in time of ovulation from administration of prostaglandin, it is not very useful when used alone in the synchronization of estrous cycles in mares. Utilization of transrectal ultrasonographic examination of these mares at the time of the second injection of prostaglandin and subsequently thereafter when the mares return to estrus and the addition of hCG or deslorelin to the protocol should maximize the synchrony of ovulation.13, 27
Progestins Exogenous progestins are commonly used in estrus synchronization programs to lengthen the luteal phase. The two most common forms of progestins used are progesterone in oil and altrenogest (allyltrenbolone; Regumate, Intervet Inc., Millsboro, DE), a synthetic progestin. Progesterone or a synthetic progestin administered alone or in combination with estradiol-17β will effectively lengthen the luteal phase indefinitely until their administration is discontinued. Progesterone has an inhibitory effect on luteinizing hormone (LH) release from the anterior pituitary; therefore, when administered for an extended period of time (15–18 days) the CL should regress and the only source of progesterone to suppress estrus is the exogenous progesterone. However, even the endogenous level of progesterone present during the midluteal phase is not sufficient to suppress gonadotropin release, follicular development, and ovulation.1 Studies have also shown that the synthetic progestin altrenogest, administered at 0.044 mg/kg orally once a day, fails to suppress follicular development.30 When using progestins for estrus synchronization it is recommended that either altrenogest (0.044 mg/kg p.o. q. 24 hours) or progesterone in oil (150 mg/day i.m.) be administered for 10 days combined with the administration of PGF2α on the last day of progestin therapy to lyse any CLs present. Owing to the varying size of follicles that
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Synchronization of Ovulation may be present at the time of prostaglandin administration the time to ovulation can be quite variable and may range from 3 to 11 days. Recently, injectable sustained release progestin formulations have been described. In a report published by Storer and colleagues31 in 2009 three sustained release formulations containing altrenogest were compared for their ability to suppress estrus and ovulation in mares. BioRelease LA 150 altrenogest solutions (BET Pharm LLC, Lexington, KY) containing 225 mg (1.5 mL LA 150) and 450 mg (3.0 mL LA 300) of altrenogest inhibited the onset of estrus for 12.7 and 15.8 days, respectively, with ovulation occurring in a mean of 16.5 and 21.2 days. The coefficient of variations for the mean time of ovulation were 11% (15–20 days) and 6% (20–23 days), respectively, for those two groups, demonstrating the potential usefulness of these compounds in estrus synchronization protocols. In the same study, when six mares were treated with a formulation of 500 mg altrenogest encapsulated in lactide–glycolide microparticles designed to release 16.6 mg/day for a 30-day period (MP 500; BioRelease Technologies LLC, Birmingham, AL), estrus and ovulation were delayed for an average of 32.8 and 34.7 days, respectively; however, four of the mares did not display estrus around their first ovulation after treatment. This finding reinforces that the threshold concentration of altrenogest necessary to inhibit behavioral estrus is lower than that to suppress LH secretion and ovulation. The MP 500 formulation did not provide as tightly synchronized ovulations, yielding mares ovulating 28–48 days from treatment with a coefficient of variation of 27%. Further studies with these products will reveal their role in estrus synchronization protocols.31
Progesterone and estradiol To improve estrus synchronization in randomly cycling mares, a combination of progesterone and estrogen, rather than progesterone alone, may be administered. The combined effect of the two hormones provides a more profound negative feedback on gonadotropin release than does progesterone alone, which results in a more uniform inhibition of follicular development. When the exogenous progesterone/estrogen therapy is discontinued, there is less diversity in follicular maturation and the date ovulation occurs can be more closely synchronized. The recommended protocol involves intramuscular injections of progesterone (150 mg) combined with estradiol-17β (10 mg) in oil for 10 consecutive days with a single injection of PGF2α administered on the 10th day of treatment.32 To further tighten the synchronization of ovulation, hCG [2500 interna-
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tional units (IU)/kg i.v.], deslorelin acetate, or reLH is administered when a 35-mm follicle is present. With this protocol, more than 70% of mares ovulate between 10 and 12 days after the last steroid injection with a range of ovulations occurring from 8 to 17 days.33 Currently, there is no commercial preparation containing progesterone and estradiol-17β combined; therefore, this formulation must be ordered from a compounding pharmacy.32, 33
Ovulation-inducing agents Induction of ovulation is an integral part of an estrus/ovulation synchronization program. The three most common pharmacological agents used to induce ovulation are human chorionic gonadotropin (hCG), deslorelin acetate, and reLH. hCG is a protein consisting of two peptide chains that, although chemically different from pituitary LH, has primarily LHlike activity. hCG is produced by cytotrophoblasts of the chorionic villi of the human placenta and appears in the urine of pregnant women a few weeks after conception. While there has been a range of dose rates published for hCG from 1000 to 3300 IU,34–37 a recent study published in 2007 by Davies Morel and Newcomb38 demonstrated that a hCG dose of 750 IU is just as effective in inducing ovulation within 48 hours as 1500 IU. Deslorelin acetate is a potent synthetic gonadotropin releasing hormone (GnRH) analog. Deslorelin is a small peptide that is administered subcutaneously in the form of an implant (Ovuplant) containing 2.1 mg of the active ingredient or intramuscularly in a biorelease formulation. The advantage of using deslorelin over hCG to induce ovulation is that its smaller molecular weight makes it less antigenic, thereby reducing the chance that antibodies will be developed against it after multiple administrations in a season. However, it must be noted that administration of deslorelin implants in a small percentage of mares does result in a prolonged interovulatory interval owing to downregulation of the pituitary by the continued release of deslorelin from the implant after ovulation has occurred.39–41 This downregulation is most pronounced when prostaglandins are administered early in diestrus after the administration of deslorelin, compared with mares that are allowed to return to estrus naturally. This side-effect can be avoided by removing the implant after ovulation has occurred.41 Currently, Ovuplant is not available in the USA; however, an injectable formulation of deslorelin, BioRelease Deslorelin Injection (BET Pharm, Lexington, KY), or numerous compounded formulations have become available. This deslorelin product is administered intramuscularly at the recommended dose of 1.5 mg
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when a 35-mm or greater follicle is present and the mare is in estrus. Based on the results of recent research, this injectable formulation of deslorelin does not cause the prolonged interovulatory interval seen with the administration of deslorelin implants.42 Administration of hCG when a 35-mm or greater follicle is present in an estrual mare results in ovulation in 36 ± 5 hours in more than 80% of mares, and deslorelin acetate administration with the presence of a 30-mm or greater follicle in an estrual mare results in ovulation in 41 ± 3 hours in 88.2% of mares.43 Recombinant equine LH may also be used in the same manner as hCG and deslorelin acetate to induce ovulation in the mare. Unlike hCG, studies have shown that mares do not appear to develop antibodies to reLH, and therefore should not become refractory to its use over time. The recommended dose of reLH is 750 µg administered intravenously when there is a 35-mm or greater follicle present and the mare is in estrus. reLH has been shown to reliably induce ovulation within 48 hours of administration.44 hCG, deslorelin, and reLH are most commonly used to induce ovulation within 48 hours of breeding. In addition to their application in programs utilizing fresh and cooled semen, these pharmacological agents are used extensively in frozen semen programs. Owing to the decreased longevity of frozen semen, insemination must occur within the time frame of 12 hours before to 6 hours after ovulation. If two doses of semen are available, a timed insemination program can be utilized. In this program, once a 35-mm follicle is detected in an estrual mare, hCG, deslorelin, or reLH is administered that day. The mare is then examined via transrectal ultrasonography and inseminated at 24 and 40 hours. If only one dose of semen is available, the same basic plan is followed, except the mare is not inseminated at the 24-hour examination, but is then examined via transrectal ultrasound every 6 hours until ovulation is detected. Once ovulation is detected, the mare is inseminated within 6 hours after ovulation.
Progesterone-releasing intravaginal devices In addition to the pharmacological agents previously described there are other products and estrus synchronization programs described in the literature that warrant discussion. Numerous reports in the literature discuss the use of an intravaginal progesteronereleasing device originally designed for cattle (CIDRB) in mares to synchronize estrus. These devices have been used with and without the administration of estradiol (10 mg, i.m. or intravaginal) on the day of
CIDR-B placement. Results of these studies were similar to those with progesterone in oil or altrenogest. Without the addition of the estradiol, CIDR-B administration resulted in synchronization of estrus but not of ovulations.45–47 The CIDR was first produced in New Zealand and was approved in May 2002 for use in the USA, where it is marketed under R Cattle Insert (Pharthe name Eazi-BreedTM CIDR macia Animal Health, Kalamazoo, MI) and contains 1.38 g progesterone. In Canada, this product contains 1.9 g progesterone. These products are not labeled ¨ for use in horses. In 1994 Lubbecke et al.45 reported on using CIDR-B and deslorelin to synchronize estrus and ovulation in mares. In the study CIDR-Bs were inserted into the vaginas of 40 normally cyclic Hanoverian mares on day 0 and were removed on day 12; all mares were treated with prostaglandin at the time of CIDR-B removal. Fifteen of the mares were treated with a short-term implant containing 2.2 mg of deslorelin when the largest ovarian follicle reached 40 mm in diameter whereas a second group of 15 mares received the implant on day 3 of estrus regardless of the follicular diameter. The remaining 10 mares served as controls. Ovulations occurred in 20% of the mares while the CIDR-Bs were in place. The mean day of ovulation for groups A, B, and C was 18.5 ± 2.2 days (range 16–21 days), 18.4 ± 1.3 days (range 16–21), and 19.1 ± 2.3 days (range 16–23 days), respectively. Based on the results of this study, treatment of cyclic mares with the CIDR-B for 12 days synchronized the onset of estrus; however, it did not synchronize ovulations tightly enough to allow for single timed inseminations owing to the progesterone treatment not sufficiently controlling follicular growth during administration.45 In 1994 Arbeiter et al.46 reported on the use of CIDR-Bs in combination with subcutaneous implants of deslorelin in acyclic mares to induce follicular growth and ovulation. The CIDR-Bs were inserted in 29 acyclic mares and were removed in 21 mares on day 10 and in eight mares on day 12 after insertion. Twenty-seven out of 29 exhibited an increase in ovarian activity while the progesterone-releasing device was in place. Twenty-two mares received a subcutaneous implant containing deslorelin (1.5 or 2.25 mg) at a mean follicular size of 37.9 mm. Ovulation occurred 2.77 ± 1.66 days after placement of the implant. Two of the mares treated with the implant did not ovulate and developed atretic follicles. Only three of the seven mares that did not receive deslorelin implants ovulated spontaneously after removal of the CIDR-B. Results of this study demonstrated that CIDR-B implants could be used in combination with deslorelin implants to induce follicular growth and ovulation in acyclic or anestrous mares.46
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Synchronization of Ovulation In contrast to the 1994 study by Arbeiter et al.,46 Handler et al.48 reported that the use of a progesterone-releasing device (PRIDTM , Sanofi, France) was not satisfactory in synchronizing estrus and ovulation in anestrus mares treated in January. Mares in this study were treated for 11 days with a PRIDTM and cloprostenol was administered at the time of PRIDTM removal. There was also a high variability of interval from the time of PRIDTM removal to ovulation, 10–22 days in January, 7–11 days in March, 1–9 days in June, and 5–14 days in October.48 It should be noted that no ovulation-inducing agents were used in this study that may have decreased this variability in the interval from PRIDTM removal to ovulation. A more recent study by Card and Green49 further reinforced the previous findings that CIDR-Bs or other forms of progestin, in combination with an ovulation-inducing agent such as deslorelin, synchronize estrus but not ovulation owing to the uncontrolled patterns of follicular development. To address the problem of uncontrolled follicu47 ¨ lar development, Klug and Jochle administered 10 mg of estradiol on day 1 of CIDR-B placement, prostaglandin on day 12 at the time of CIDR-B removal, and deslorelin implant when a 40-mm follicle was detected. This protocol resulted in tighter synchronization of estrus and ovulation owing to what appears to be a damping effect of the estradiol on follicular growth and a more uniform population of follicles at the time of prostaglandin administration.47 For completion it should be noted that in the early 1980s reports of using progestin-coated sponges or plastic spirals to deliver progesterone for a duration of 8–19 days were described.50, 51 The results of these studies did not yield consistent follicular and ovulatory responses. In addition, Thompson et al. 52, 53 published data demonstrating that insertion of unmedicated polyurethane sponges into the vaginas of acyclic mares during seasonal anestrus resulted in the immediate release of follicle-stimulating hormone (FSH) in 50% of repeatedly treated mares. This FSH surge was accompanied by follicular development with diameters of greater than 20 mm and short LH elevations in some mares. A similar intravaginal progesterone-releasing device designed for mares, Cue-MareTM (Duirs PfarmAg, Hamilton, New Zealand), has been described for the use of estrus synchronization in transitional mares. After a 10-day application of the CueMareTM in 38 transitional mares, all of the mares exhibited signs of estrus 3 days after removal of the devices. Further research is needed to determine the value of this product in estrus synchronization programs.54 One advantage to intravaginal progesterone-releasing devices is that they do not re-
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quire the labor associated with daily administration of progesterone in oil or altrenogest. However, mild to moderate vaginitis is observed in most mares that have an intravaginal device administered.
Follicular ablation The use of ultrasound-guided transvaginal follicle ablation for synchronizing ovarian function in the mare has been described. In one study, on day 0 all follicles greater than or equal to 10 mm were ablated via ultrasound-guided transvaginal follicular aspiration. Four days later, two doses of cloprostenol (250 µg i.m.) were given 12 hours apart followed by hCG administration 6 days after that. Results of this study showed that 96% (22/23) of mares ovulated within 48 hours of hCG treatment.55 When compared with conventional steroid regimes in which the interval from beginning of treatment to ovulation can be up to 20–25 days with56 or without57 hCG administration, this protocol represents an approximate 50% reduction in the interval from follicle ablation to ovulation. In addition, utilization of this procedure avoids the labor associated with administration of pharmacological agents daily. However, more research is needed to assess the effects of this procedure on pregnancy rates, its practicality in a practice setting, and its effects on the health of the mares’ ovaries.55 In 2007 Bergfelt et al.58 reported on a study in which follicular ablation was compared with a regimen of progesterone plus estradiol (P&E) as a method of ovulation synchronization. On day 1 of the study, mares in the ablation group were lightly sedated and all follicles greater than 10 mm in diameter were removed by transvaginal ultrasound-guided follicle aspiration. PGF2α was administered at a dose of 10 mg per mare 12 hours apart on day 5 and hCG was administered at a fixed time on day 6 after prostaglandin administration. In the P&E group, a combination of progesterone (150 mg) and estradiol (10 mg) prepared in safflower oil was administered i.m. for 10 days. A single dose of prostaglandin was given on day 10 and hCG was administered 10 days after the prostaglandin. Results of this study demonstrated that the mean interval from the start of the study and from prostaglandin administration to ovulation was significantly shorter in the ablation group: 13.7 and 9.7 days, respectively, in the ablation group compared with 22.3 and 13.2 days in the P&E group. However, there was no significant difference in the degree of ovulation synchronization between the ablation and P&E mares within a 2-day (56% and 70%) or 4-day period (83% and 90%).58 While follicular ablation does result in a significant reduction of 39% in the interval from the start of the
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synchronization regimen to ovulation as compared with the P&E regimen, there is no improvement in the degree of ovulation synchronization when there is fixed-time administration of hCG after prostaglandin treatment.58 This study demonstrated that follicular ablation is a feasible and efficacious method of ovulation synchronization of mares; however, it does require the necessary equipment and skills to perform and may not be practical in all cases.
Estrus synchronization in an embryo transfer program To achieve the best results, at least three recipient mares should be synchronized with the donor mare. The administration of progesterone or progesterone and estradiol should be initiated in both the donor and recipient mares on the same day. Approximately 8–10 days later discontinuation of the progestin therapy and administration of prostaglandin should occur in all of the mares. Transrectal ultrasonography should be performed 3–4 days after prostaglandin administration to evaluate the follicular status in the mares. Since it is ideal for the recipient mares to ovulate 1–3 days after the donor mare, administration of ovulation-inducing agents should be timed to achieve these results. It is important to realize that all of the recipients may not synchronize with the donor, nor may they be ideal recipients at the time of transfer owing to the presence of uterine fluid or edema, poor cervical tone, or inadequate CL formation. For this reason, it is important to synchronize at least three recipients with each donor to maximize the chances of having a suitable recipient at the time of transfer.
Conclusion Estrus synchronization affords the veterinarian the ability to control the estrous cycle in a manner to optimize the time of insemination as it relates to ovulation. This becomes especially prudent when there is limited availability of the stallion or when cooled transported or frozen semen is being used. Estrus synchronization is also an important part of embryo and gamete transfer programs. The use of a combination of pharmacological agents, most commonly progesterone and estradiol-17β, followed by the administration of PGF2α and hCG, deslorelin, or reLH, yields the most reliable results. However, research is ongoing in this field, and current knowledge of the latest findings will afford the veterinarian the best options when developing an estrus synchronization program.
References 1. Meyers PJ. Control and synchronization of the estrous cycle and ovulation. In: Youngvist RS (ed.) Current Therapy in Large Animal Theriogenology. Philadelphia: WB Saunders, 1997; pp. 97–102. 2. Shiner K. A review of clinical applications of hCG on breeding farms. In: Proceedings of the 34th Annual Convention of the American Association of Equine Practitioners, 1988;197. 3. Neely DP. Hormone therapy. In: Hughes JP (ed.) Equine Reproduction. Trenton, NJ: Veterinary Learning Systems, 1983; p. 24. 4. Foss R, Wirth N, Schiltz P, Jones J. Nonsurgical embryo transfer in a private practice. In: Proceedings of the 45th Annual Convention of the American Association of Equine Practitioners, 1999, pp. 210–12. 5. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1993; pp. 173–232. 6. Allen WR, Rowson LEA. Control of the mare’s estrous cycle by prostaglandins. J Reprod Fertil 1973;33:539–43. 7. Douglas RH, Ginther OJ. Effects of prostaglandin F2 alpha on length of diestrus in mares. Prostaglandins 1972;2: 265. 8. Holtan DW, Douglas RH, Ginther OJ. Estrus, ovulation and conception following synchronization with progesterone, prostaglandin F2 alpha and human chorionic gonadotrophin in pony mares. J Anim Sci 1977;44:431. 9. Oxender WD, Noden PA, Bolenbaugh DL, Hafs HD. Control of estrus with prostaglandin F2 alpha in mares: minimal effective dose and stage of estrous cycle. Am J Vet Res 1975;136:1145. 10. Douglas RH, Ginther OJ. Route of prostaglandin F2 α injection and luteolysis in mares. Proc Soc Biol Med 1975;148: 263–9. 11. Nie GJ, Goodin AN, Braden TD, Wenzel JGW. Luteal and clinical response following administration of dinoprost tromethamine or cloprostenol at standard intramuscular sites or at the lumbosacral acupuncture points in mares. Am J Vet Res 2001;62:1285–9. 12. Irvine CHG, McKeough VL, Turner JE, Alexander SL, Taylor TB. Effectiveness of a two-dose regimen of prostaglandin administration in inducing luteolysis without adverse side effects in mares. Equine Vet J 2002; 34:191– 194. 13. Lofstedt RM. Control of the estrous cycle in the mare. Vet Clin North Am Equine Pract 1988;4:177–96. 14. Bristol F. Studies on estrous synchronization in mares. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1981, p. 258. 15. Hughs JP, Stabenfeldt GH, Evans JW. Clinical and endocrine aspects of the estrous cycle of the mare. In: Proceedings of the 18th Annual Convention of the American Association of Equine Practitioners, 1972, p. 229. 16. Spincemaille J, Coryn M, Vanderkerckhobe D, Vandeplassche M. The use of prostaglandin F2 alpha in controlling the oestrous cycle of the mare and steroid changes in the peripheral blood. J Reprod Fertil Suppl 1975;23:263. 17. Ginther OJ. Occurrence of anestrous, estrus and diestrus and ovulation over a 12-month period in mares. Am J Vet Res 1974;35:1173. 18. Loy RG, Buell JR, Stevenson W, Hamm D. Sources of variation in response intervals after prostaglandin treatment in mares with functional corpora lutea. J Reprod Fertil Suppl 1979;27:229.
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Synchronization of Ovulation 19. Wenzel JGW. Estrous cycle synchronization. In: Youngvist RS (ed.) Current Therapy in Large Animal Theriogenology. Philadelphia: WB Saunders, 1997; p. 291. 20. Seguin BE, Momont H, Baumann L. Cloprostenol and dinoprost tromethamine in experimental and field trials treating unobserved estrus in dairy cows. Bovine Pract 1985; 20:85. 21. Seguin BE. Role of prostaglandins in bovine reproduction. J Am Vet Med Assoc 1980;176:1178. 22. MacMillan KL. Prostaglandin responses in dairy herd breeding programmes. N Z Vet J 1983;31:110. 23. Momont HW, Sequin BE. Influence of day of estrous cycle in response to PGF2 alpha products: implications for AI programs for dairy cattle. In: Proceedings of the 10th International Congress Animal Reproduction and Artificial Insemination, 1984, vol. 2, p. 336. 24. Nie GJ, Johnson KE, Wenzel JGW, Braden TD. Luteal function in mares following administration of oxytocin, cloprostenol or saline on day 0, 1 or 2 post-ovulation. Theriogenology 2003;60:1119–25. 25. Brendemuehl JP. Effect of oxytocin and cloprostenol on luteal formation. Theriogenology 2002;58:623–6. 26. Hyland JH, Bristol F. Synchronization of oestrus and timed insemination of mares. J Reprod Fertil Suppl 1979;27: 251. 27. Squires EL, Wallace RA, Voss JL, Pickett BW, Shideler RK. The effectiveness of PGF2 alpha, hCG and GnRH for appointment breeding in mares. J Equine Vet Sci 1981; 1:5. 28. Voss JL, Wallace RA, Squires EL, Pickett BW, Shideler RK. Effects of synchronization and frequency of insemination on fertility. J Reprod Fertil 1979;27:Suppl 256. 29. Irvine CHG. Endocrinology of the estrous cycle of the mare: applications to the embryo transfer. Theriogenology 1981;15: 895. 30. Lofstedt RM, Patel JH. Evaluation of the ability of altrenogest to control the equine estrous cycle. J Am Vet Med Assoc 1989;194:361. 31. Storer WA, Thompson Jr DL, Gilley RM, Burns PJ. Evaluation of injectable sustained release progestin formulations for suppression of estrus and ovulation in mares. J Equine Vet Sci 2009;29:33–6. 32. Loy RG. Characteristics of postpartum reproduction in mares. Vet Clin North Am Large Anim Pract 1980;2:345– 58. 33. Blanchard TL, Varner DD, Schumacher J. Manual of Equine Reproduction. St. Louis, MO: Mosby, 1998; pp. 22–3. 34. McCue PM. Induction of ovulation. In: Robinson NE (ed.) Current Therapy in Equine Reproduction. Philadelphia: Saunders, 2003; p. 240. 35. Loy RG, Hughes JP. The effect of human chorionic gonadotrophin on ovulation, length of estrus and fertility in the mare. Cornell Vet 1966;56:41. 36. Michel TH, Rossdale PD. Efficacy of human chorionic gonadotrophin and gonadotrophin releasing hormone for hastening ovulation in Thoroughbred mares. J Equine Vet Sci 1986;18:438. 37. Duchamp G, Bour B, Combarnous Y, Palmer E. Alternate solutions to hCG induction of ovulation in the mare. J Reprod Fertil 1979;27:Suppl 173. 38. Davies Morel MC, Newcomb JR. The efficacy of different hCG dose rates and the effect of hCG treatment on ovarian activity: ovulation, multiple ovulation, pregnancy, multiple pregnancy, synchrony of multiple ovulation; in the mare. Anim Reprod Sci 2008;109:189–99.
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39. Farquar VJ, McCue PM, Nett TM, Squires EL. Effect of deslorelin acetate on gonadotropin secretion and ovarian follicle development in cycling mares. J Am Vet Med Assoc 2001;218:749–52. 40. Farquar VJ, McCue PM, Carnevale EM, Nett TM, Squires EL. Deslorelin acetate (OvuplantTM ) therapy in cycling mares: effect of implant removal on FSH secretion and ovarian function. Equine Vet J 2002;34:417–20. 41. McCue PM, Farquar VJ, Carnevale EM, Squires EL. Removal of deslorelin (OvuplantTM ) implant 48 hours after administration results in normal interovulatory intervals in mares. Theriogenology 2002;58:865–70. 42. Stich KL, Wendt KM, Blanchard TL, Brinsko SP. Effects of a new injectable short-term release deslorelin in foal-heat mares. Theriogenology 2004;62:831–6. 43. McKinnon AO, Perriam WJ, Lescun TB, Walker J, Vasey JP. Effect of a GnRH analog (Ovuplant), hCG and dexamethasone on time to ovulation in cycling mares. World Equine Vet Rev 1997;2:16–18. 44. Yoon MJ, Boime I, Colgin M, Niswender KD, King SS, Alvarenga M, Jablonka-Shariff A, Pearl CA, Roser JF. The efficacy of a single chain recombinant equine luteinising hormone (reLH) in mares: induction of ovulation, hormone profiles and inter-ovulatory intervals. Domest Anim Endocrinol 2007;33:470–9. ¨ ¨ 45. Lubbecke M, Klug E, Hoppen HO, Jochle W. Attempts to synchronize estrus and ovulation in mares using progesterone (CIDR-B) and GnRH-analog deslorelin. Reprod Dom Anim 1994;29:305–14. ¨ 46. Arbeiter K, Barth U, Jochle W. Observations on the use of progesterone intravaginally and of deslorelin STI in acyclic mares for induction of ovulation. J Equine Vet Sci 1994;14:21–5. ¨ 47. Klug E, Jochle W. Advances in synchronizing estrus and ovulations in the mare: a mini review. J Equine Vet Sci 2001;21:474–9. ¨ 48. Handler J, Schonlieb S, Hoppen H Aurich C. Seasonal effects on attempts to synchronize estrus by intravaginal application of progesterone-releasing device (PRIDTM ) in mares. Theriogenology 2006;65:1145–58. 49. Card C, Green J. Comparison of pregnancy rates by week from stallion exposure and overall pregnancy rates in pasture-bred mares synchronized with CIDR and/or prostaglandin F2 alpha. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2004, p. 1493. 50. Dinger JE, Noiles EE, Bates MJ. Effect of progesterone impregnated vaginal sponges and PMSG administration on oestrous synchronization in mares. Theriogenology 1981; 16:231–3. 51. Rutten DRS, Chaffaux M, Valon F, Deletang F, De Hass V. Progesterone therapy in mares with abnormal estrous cycles. Vet Rec 1986;119:569–71. 52. Thompson Jr DL, Godke RA, Nett TM. Effects of melatonin and thyrotropin releasing hormone in mares during the nonbreeding season. J Anim Sci 1983;56:668– 77. 53. Thompson Jr DL, Reville SI, Derrick DJ, Walker MP. Effects of placement of intravaginal sponges on LH, FSH, estrus and ovarian activity in mares during the nonbreeding season. J Anim Sci 1984;58:159–64. ¨ 54. Grimmett JB, Hanlon DW, Duirs GF, Jochle W. A new intra-vaginal progesterone-releasing device (Cue-MareTM ) for controlling the estrous cycle in mares. Theriogenology 2002;58:585–7.
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55. Bergfelt DR, Adams GP. Ovulation synchrony after follicle ablation in mares. J Reprod Fertil Suppl 2000;56:257– 69. 56. Varner DD, Blanchard TL, Brinsko SP. Estrogen, oxytocin and ergot alkaloids: uses in reproductive management of mares. In: Proceedings of the 35th Annual Convention of the American Association of Equine Practitioners, 1988, pp. 219–41.
57. Loy RG, Pemstein R, O’Canna D, Douglas RH. Control of ovulation in cycling mares with ovarian steroids and prostaglandin. Theriogenology 1981;15:191–7. 58. Bergfelt DR, Meira C, Fleury JJ, Fleury PD, Dell’Aqua JA, Adams GP. Ovulation synchronization following commercial application of ultrasound-guided follicle ablation during the estrous cycle in mares. Theriogenology 2007;68: 1183–91.
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Chapter 198
New Techniques in Hormonal Delivery Patrick J. Burns
There is a need to develop and capitalize on new mare management programs within the equine breeding industry that are based on recent advances in effective hormonal control treatments. Success of such programs will be attributable in large part to recent advances in biodegradable controlled release drug delivery systems that will allow single administration products to replace prolonged daily treatment protocols.1, 2 These formulations will reduce labor and the associated handling stress to the mares and producers and offer veterinarians an important means of maintaining effective compliance rates on farms with wide varieties of management systems. In this chapter, we will discuss new controlled release treatments used for reproductive management in modern equine breeding programs.
Controlled release progestin formulations Progestins are among the most widely used reproductive drugs in veterinary medicine. Therefore, controlled release hormone treatment formulations for the delivery of progestins are highly valued by equine practitioners. This is because most progestin treatment protocols require 1–2 weeks or longer and daily treatments are inconvenient, time-consuming, and prone to missed dosings. Both injectable and intravaginal progestin delivery systems have been examined in horses.
Intravaginal progestin delivery Palmer3 conducted the initial intravaginal progestin delivery studies in which he demonstrated that suppression of estrus and ovulation for > 20 days
was possible with 500–1000 mg of altrenogest. The polyurethane sponges were also coated with antibiotics to prevent bacterial vaginitis. It was noted that sponge retention was excellent. However, some vaginal adhesions and irritation were observed. Over the next 20 years no intravaginal products were formulated for horses, but the use of the commercially available PRID and CIDER-B products have been successfully explored and reviewed.4 Recently, progesterone delivery by the Cue-MareTM , an intravaginal delivery device formulated for horses, was shown to be effective in transitional mares with a history of estrus behavior of longer than 10 days and follicles less than 25 mm.5 The authors reported an impressive 87% ovulation response rate within 7 days of device removal and a 52.9% conception rate for mares bred after device removal. They also reported the device was capable of maintaining blood levels of > 2 ng/mL for 5 days and levels above 1 ng/mL for 10 days. Intramuscular progestin delivery The first injectable long-acting progestin was formulated as a 50 mg/mL suspension in a propylene glycol base (repositol progesterone) that required a dose of 2000 mg to achieve consistent plasma levels above 2 ng/mL for a 7-day period.6 It should be noted that the 2000-mg dose would require a 40-mL injection, which may be one reason the product left the marketplace more than 20 years ago, shortly after the introduction of the oral progestin altrenogest. Biodegradable poly(dl-lactide) microspheres were the next progestin delivery system to be examined in horses. The microspheres were produced using a new solvent extraction process2, 7 and were successfully evaluated for maintenance of pregnancy8
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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and synchronization of estrus in cyclic mares when combined with estradiol microspheres.9–14 Blanchard et al.12 demonstrated that using the combination of progesterone and estradiol microspheres when followed by prostaglandin treatment at the end of the 14-day steroid treatment was superior to progesterone microspheres followed by prostaglandin treatment for the control of estrus and ovulation in mares. Fleury et al.13 demonstrated that the 100-mg dose of estradiol microspheres produced optimal control of estrus and ovulation when combined with progesterone microspheres. The clinical efficacy of progesterone and estradiol microspheres to control estrus and ovulation in late transitional non-cyclic mares, as well as in cyclic mares, has been demonstrated in a controlled multicentered clinical trial using a total of 135 mares.11 Unfortunately, scaling up commercial microsphere production to meet the demands of the domesticated livestock market is a very complex process requiring costly specialized facilities, water systems, and equipment that may not be economically feasible for most veterinary products.1 Therefore, progesterone formulation studies using the new more cost-effective Saber Delivery System followed the biodegradable microsphere polymer systems. The Saber Delivery System uses a unique highviscosity base compound to provide controlled release of active compounds over a specified period of time. Upon addition of a small amount of solvent, the high-viscosity component, sucrose acetate isobutyrate (SAIB), converts to an easily injectable liquid. Once inside the body, the solvent dissipates and a semisolid, biodegradable implant is formed. The use of different solvents and additives can alter the duration of release from the Saber Delivery System from
hours to months. As opposed to microspheres, production of Saber formulations involve a simple mix and fill operation. Burns et al.15 demonstrated the superiority of a progesterone and estradiol combination Saber formulation compared with progesterone alone. The authors noted that the two Saber formulations provided similar serum progesterone levels; however, the addition of estradiol to the formulation delayed ovulation by 6.5 days, allowing for a normal estrus to accompany ovulation. Fleury et al.13 reported that addition of the same 100-mg dose of estradiol to the same 1.25g progesterone dose as a microsphere formulation also delayed ovulation by 6.5 days compared with progesterone microspheres alone. The results confirm earlier observations with microspheres that the combination of progesterone and estradiol acts to provide a unique contribution to controlled release formulations for control of estrus and ovulation which cannot be achieved with progesterone alone. Recently, several new compounded controlled release progestin formulations have become available to veterinarians by prescription. The formulation BioRelease P4 LA150 (BET Pharm, Lexington, KY) has been shown to maintain mean blood levels above 2 ng/mL for approximately 8–10 days.16 Vanderwall et al.17 demonstrated that the formulation was effective at maintaining pregnancy between days 18 and 45 when given every 7 days to pregnant mares whose endogenous luteal progesterone had been removed by injecting cloprostenol. Burns, et al.19 recently reported that a more concentrated progesterone formulation, BioRelease P4 LA300, produced similar blood levels to the earlier P4 LA150 formulation, but required only half the injection volume, as shown in Figure 198.1.
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Figure 198.1 Serum progesterone concentrations (ng/mL) after intramuscular injection of 5 mL of BioRelease P4 LA300, 10 mL of BioRelease P4 LA300, or 10 mL of BioRelease P4 LA150 (adapted from Burns et al.19 ).
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New Techniques in Hormonal Delivery Table 198.1 Days to estrus and ovulation following lutalyse and sustained release progestin treatment (adapted from Burns et al.18 ) Formulation
N
Days to estrus
Days to ovulation
LA150 1.5-mL LA150 3 mL MP 500 Medroxyprogesterone Controls
6 6 6 6 7
12.7 ± 0.42b 15.8 ± 1.51b 32.8 ± 4.80c 6.2 ± 1.38a 3.9 ± 0.51a
17.3 ± 0.75b 21.5 ± 0.54b 34.2 ± 3.72c 11.0 ± 2.14a 7.2 ± 1.17a
Means ± SE; means with different superscripts differ (P < 0.05).
Recently Burns et al.19 reported on the use of the compounded BioRelease P4 LA300 (2 mL) in nonresponding ‘lighted’ mares that had been lighted > 50 days but had follicles < 15 mm prior to treatment. In this study, 57% of the P4 LA300 mares ovulated within 28 days after injection compared with 7% in control mares. Several new compounded controlled release injectable altrenogest formulations using either the BioRelease liquid delivery system or the longer acting lactide-co-glycolide microparticle (MP) system have been evaluated.18, 20 Results are summarized in Table 198.1. The more potent progestin altrenogest offers the advantage of smaller injection volumes ranging down to 1.5 mL for 12 days of suppression of estrus. All injectable altrenogest formulations were effective at extending the interovulatory interval and delaying estrus. The altrenogest MP 500 microparticle formulation had the longest inhibitory effect. This formulation should be beneficial for the performance horse by inhibiting estrous behavior for a period of 30 days and could be administered repeatedly as desired to maintain reproductive quiescence or anestrous behavior. The altrenogest LA150/1.5-mL formulation was also very effective and would be valuable when shorter periods of suppression of estrus (12–14 days) are desired, such as in transitional mares to establish normal cycles or for estrus and ovulation synchronization programs where the degree of estrus and ovulation synchronization compares favorably with daily altrenogest treatment21 or progesterone with estradiol treatments.10 Lastly, the authors confirmed the observations of McKinnon et al.,22 demonstrating that doses of long-acting medroxyprogesterone acetate did not suppress estrus or ovulation in mares. Since progestin-treated non-cyclic mares are frequently used as embryo recipients in large commercial programs, the new once-weekly controlled release formulations are advantageous in such programs. Weekly treatment with BioRelease altrenogest
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(225 or 450 mg) was effective at maintaining pregnancy in prostaglandin-treated mares with no significant differences between the two doses studied. 23 However, based on extensive clinical experience with BioRelease P4 LA150 or 300 for pregnancy maintenance in recipient mares, the authors recommend the use of natural progesterone formulations until additional larger studies with BioRelease altrenogest LA150 have been completed. Many commercial embryo transfer programs routinely supplement cyclic recipient mares with longacting progestin treatments or daily altrenogest treatment following transfer. The effectiveness of such treatments remains to be critically demonstrated. A recent study by Squires et al.24 indicated that 53.3% of pregnant embryo transfer recipient mares on daily oral altrenogest treatment (22 mg) showed greater than a 33% drop in progesterone following transfer and that 10% dropped below 2 ng/mL. Additional studies to determine the effect of non-surgical transfer and/or altrenogest supplementation on progesterone secretion are needed. Daels et al.25 reported that a daily double dose of oral altrenogest (44 mg) during pregnancy significantly decreased progesterone levels, with 80% of altrenogest-treated mares exhibiting progesterone levels < 2 ng/mL. The treated mares also tended to show a delay and reduction in the development of a secondary corpus luteum. Recently, McCue et al.26 observed significant reductions in progesterone blood levels in altrenogest-treated pregnant mares compared with untreated control pregnant mares on days 14, 15, 17, 18, and 21 of the study. They noted that eight out of nine treated mares had decreasing levels of < 4 ng/mL with one mare dropping to 1.5 ng/mL. Such observations suggest that progestin therapy during early pregnancy may be contraindicated until further data show otherwise.
Controlled release estrogen formulations Based on observations that mares under estrogen influence are more capable of eliminating uterine infection than mares under the influence of progesterone,27–30 interest in long-acting estrogen formulations has been increasing in recent years. Cook et al.31 reported that estradiol microsphere treatment with doses as high as 100 mg had no effect on fertility in normal mares during treatment (75% vs 75% in vehicle and treated). Johnson et al.32 demonstrated that estradiol could be delivered equally as effectively using the more cost-efficient Saber delivery system as using poly(dl-lactide) microspheres for a period of 14–18 days.
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Pinto et al.33 compared various estrogens in terms of serum estradiol concentrations following a single intramuscular (i.m.) injection of a 50-mg dose of BioRelease estradiol cypionate with two different estradiol-17β formulations. The results illustrated that peak serum concentrations of estradiol-17β were 10-fold higher on day 1 (between 300 and 350 pg/mL) in mares treated with BioRelease E2-17β than mares treated with BioRelease estradiol cypionate treatment which averaged 15–25 pg/mL but remain elevated for 10–12 days. More recently, Thompson et al.34 measured estradiol levels after various estrogen treatments with the goal of determining a blood profile similar to every other day estradiol benzoate injection in oil. Pharmacodynamic analysis indicated that a single 100-mg BioRelease estradiol cypionate injection could replace 10 injections of estradiol benzoate in oil every other day in the treatment regimen reported by Kelly et al.35 for stimulating ovulation in deep seasonally anovulatory mares when paired with a dopamine D2 antagonist such as domperidone. Douglas36 recommended the shorter acting BioRelease estradiol-17β formulation at doses between 20 and 100 mg/day (based on estradiol blood levels) along with appropriate antibiotic therapy for treatment of pregnant mares with placentitis (R.H. Douglas, personal communication).
Controlled release gonadotropin-releasing hormone and its analogs The ability of gonadotropin-releasing hormone (GnRH) or its analogs to stimulate luteinizing hormone (LH) release has led to their use for ovulation induction in anovulatory mares using frequent injections,37, 38 pulsatile infusion,39, 40 continuous infusion,41 or controlled release implants.42–45 Similar to progestins, the most successful responses to treatment occur as the time of year advances or the diameter of the largest follicle at the initiation of treatment increases. In addition, GnRH treatment of mares with small follicles (< 15 mm) is unpredictable and is associated with a high rate of early pregnancy loss (58%) owing to inadequate luteal function.38 Furthermore, recent reports of suppression of follicular growth and increased interovulatory intervals in cyclic mares with goserelin controlled release implants46, 47 or commercial deslorelin implants48 point out the complexity of working in this area and the need for careful pharmacodynamic research in GnRH/GnRH analog formulation development. It may be that long-acting GnRH analog formulations ultimately are used to suppress reproductive function in mares and stallions.
Induction or hastening of ovulation in cyclic mares has been a major goal of researchers for many years. The discovery that human chorionic gonadotropin (hCG) given to mares would shorten estrus and hasten ovulation49–51 led to its widespread use, even though repeated usage was associated with decreased response and antibody formation.52–54 The ability of GnRH55 or its analogs56, 57 to stimulate LH release led to research into their use for induction of ovulation, as their small size renders them less antigenic. However, a single injection of GnRH or its analogs produced inconsistent results,58–60 and injections every 12 hours, although effective,57 are generally considered impractical. Deslorelin, a potent GnRH analog, delivered via a short-term release (OvuplantTM ) implant61, 62 or via Saber Mate E, which uses the Saber Delivery System, 63, 64 has been shown to consistently advance ovulation within 48 hours in estrous mares having follicles 30–40 mm in diameter. However, in its first 2 years of commercial use, anecdotal reports suggested that, sporadically, mares with Ovuplant-induced ovulations experience delayed return to estrus and prolonged interovulatory intervals.65, 66 Delayed return to estrus and prolonged interovulatory intervals have also been reported after prostaglandin-induced luteal regression, commonly used in embryo transfer programs when Ovuplant was used.48 In contrast, studies15, 63, 64, 67 with Saber Mate E revealed no extended interovulatory intervals, even at higher threefold, fivefold, and 10-fold doses. The reason for the difference in response of mares to Ovuplant or Saber Mate E may be related to the deslorelin release rate in the two products. The Saber formulation released nearly twice as much deslorelin in the first 24 hours as Ovuplant.63 Moreover, blood levels of deslorelin in mares given the Saber formulation had returned to baseline by 24 hours after treatment, while blood levels of deslorelin in Ovuplant-implanted mares remained significantly elevated compared with the saline group at 24 and 36 hours.15 This suggests that continued deslorelin release past 24–48 hours might increase interovulatory intervals and lead to hyposecretion of LH and folliclestimulating hormone (FSH), as observed by Johnson et al.48 In support of this concept, Farquhar et al.68 and McCue et al.69 observed that if the Ovuplant implants were removed at 48 hours after implantation the increased interovulatory interval in mares short-cycled with progesterone (PG) F2α could be prevented. More recently, a new compounded deslorelin formulation containing 1.5 mg/mL has become available to veterinarians by prescription. The formulation BioRelease deslorelin (Bet Pharm, Lexington, KY) has been reported to effectively induce ovulation in cyclic mares with follicles of 30 to > 40 mm.70, 71 Kolling and
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New Techniques in Hormonal Delivery Allen72 recently compared BioRelease deslorelin with other current ovulation induction treatments including hCG, Ovuplant or CPE (equine pituitary extract; INRA). All four of the treatments were > 97% effective at producing ovulation within 48 hours, including a half dose of the BioRelease deslorelin (750 µg). Reported embryo recovery rates were 37.5% (15/40) for CPE, 53% (26/53) for Ovuplant, 55% (44/80) for hCG, and 68.5% (63/92) or 69.2% (126/182) for the 0.75-mg and 1.5-mg doses of the BioRelease formulations, respectively. Although not statistically examined in their paper,72 statistical analysis of the reported embryo recovery data show increased embryo recovery for BioRelease deslorelin (1.5 mg) compared with CPE (P < 0.0002), hCG (P < 0.0270), or Ovuplant (P < 0.0071). The reason for the 14.2–31.7% increase in embryo recovery for the BioRelease formulations was not discussed, but may be related to its ability to stimulate a larger LH surge capable of ovulating smaller follicles, thereby allowing for more multiple embryo recoveries even when a half dose was used. The authors did indicate that there were no adverse effects on follicular growth or prolongation of interovulatory intervals in mares treated for several consecutive cycles with either dose of the BioRelease formulation, as had been previously observed with Ovuplant treatment.48 McCue et al.73 reported that a short-acting compounded deslorelin was effective in stimulating ovulation in mares with mean follicular diameters > 40 mm.
Controlled release dopamine antagonist In the last decade, treatment with dopamine D2 antagonists such as domperidone and sulpiride have been shown to induce cyclicity in anovulatory mares in some studies74–76 but not in others.77, 78 As with progestins or GnRH/GnRH analogs, it appears that the treatment is most effective in later transitional mares, mares in locations with mild winter climates, 76 or mares that have received 28 days of increased photostimulation.79 Daels et al.80 has also reported that sulpiride treatment of mares maintained indoors in locations with cold climates was more effective than in mares maintained outdoors at the same location. Nequin et al.81, 82 and Thompson et al.83 suggested that the dopamine antagonist could be acting through prolactin at the ovarian level. Recently, Kelly et al.35 reported on the use of domperidone or sulpiride for stimulating prolactin secretion in cyclic and deeply anestrous non-lighted mares. Importantly, both dopamine antagonists can be used
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in combination with an estrogen to greatly enhance prolactin secretion. The combination of estradiol benzoate and sulpiride was also effective at inducing ovulation in deeply anestrous non-lighted mares. Mares receiving the estradiol and sulpiride ovulated 45 days earlier than mares receiving sulpiride alone (29 days vs 73.6 days). Kelly et al.35 also demonstrated that a single long-acting BioRelease domperidone lactide-co-glycolide microparticle injection stimulated prolactin in estradiol benzoate-treated cyclic mares for several weeks and Thompson et al.34 demonstrated that a single injection containing 100 mg of BioRelease estradiol cypionate could replace 10 injections of estradiol benzoate in oil every other day in the treatment regimen. Thus, now just two injections using controlled release formulations can replace an average of 39 treatments in the estrogen/dopamine D2 antagonists treatment regimen reported by Kelly et al.35 Mitcham et al.84 recently confirmed the effectiveness of the two-injection controlled release estrogen and domperidone treatment in a multisite study.
Thyroid hormones: thyroxine (T4) An accurate figure for the incidence of equine hypothyroidism is difficult to estimate but it has been reported to be high, especially in young horses and broodmares.85, 86 Lowe et al.87 failed to observe reproductive problems in thyroidectomized mares. Despite limited scientific evidence to support it, many practitioners continue to report clinical improvements in fertility after thyroid hormone supplementation. Nachreiner and Hyland88 advocate using baseline total T3 and T4 as the most economical and practical diagnostic approach and suggest 5–10 mg l-thyroxine/500 kg bodyweight per day as a good starting dose if thyroxine replacement is initiated. Because thyroid supplementation requires daily treatment for long periods of time, a controlled release injection lasting 3–4 weeks has been frequently requested. Recently, Burns et al.89 reported that a 50-mg dose of the new solvent-free extrusion compounded formulation, sterilized by gamma radiation, produced biocompatible T4 microparticles capable of delivering T4 for at least 30 days, as illustrated in Figure 198.2.
Conclusions The future offers significant opportunities to improve reproductive performance within the equine industry. This chapter has attempted to review historical and new pharmacological treatments for reproductive management in mares, with special reference
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T4 Microparticles
T4 Microparticles 2.5 MRads Gama Sterlization
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Days Post-Injection Figure 198.2 Serum thyroxine (T4) concentrations (ng/mL) above baseline after intramuscular injection of 50-mg dose of compounded thyroxine microparticles (adapted from Burns and Gilley90 ).
to the benefits of new biodegradable drug delivery systems. These benefits include the ability to integrate the administration of products to control estrus around farm and veterinary schedules and the reduction of stress by decreasing the number of times animals must be handled, thereby enhancing the financial benefits realized from effective hormone therapy.
References 1. Burns PJ. Enhanced reproductive management of domesticated livestock using biodegradable controlled release drug delivery formulations. In: Proceedings of the 26th International Symposium on Controlled Release Bioactive Materials, Boston, MA, 1999, pp. 64–5. 2. Rathbone MJ, Burns PJ, Ogle CR, Burggraaf S, Bunt CR. Controlled release drug delivery systems for estrous control of domesticated livestock. In: Rathbone MJ, Gurney R (eds) Controlled Release Veterinary Drug Delivery. Amsterdam: Elsevier Science, 2000; pp. 201–28. 3. Palmer E. Reproductive management of mares without detection of oestrus. J Reprod Fertil Suppl 1979;27:263–70. 4. Rathbone MJ, Macmillian KL, Jochle W, Boland MP, Inskeep RK. Controlled-release products for the control of the estrous cycle in cattle, sheep, goats, deer, pigs, and horses. Crit Rev Ther Drug Carrier Syst 1998;15:285–379. 5. Grimmett JB, Hanlon DW, Duirs GF, Jochle W. A new intra-vaginal progesterone-releasing device (Cue-MareTM ) for controlling the estrous cycle in the mare. Theriogenology 2002; Suppl 58:585–7. 6. Hawkins DL, Neely DP, Stabenfeldt GH. Plasma progesterone concentrations derived from the administration of exogenous progesterone to ovariectomized mares. J Reprod Fertil Suppl 1979;27:211–16. 7. Burns PJ, Tice TR, Mason DW, Love DF, Ferrell T, Gibson FJ, Dippert K, Squires EL. Evaluation of a new process for preparation of progesterone and estradiol microspheres: an in-vivo pharmacokinetics study. In: Proceedings of the
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Controlled Release Society Conference: Advances in Controlled Delivery, Baltimore, MD, 1996, abstract 639. Ball BA, Wilker C, Daels PF, Burns PJ. Use of progesterone in microspheres for maintenance of pregnancy in mares. Am J Vet Res 1992;53:1294–7. Burns PJ, Ball BA, Tice TR, Mason DW, Love DF. A preliminary report on the efficiency of biodegradable microspheres for the controlled release of progesterone and estradiol for synchronization of ovulation in mares. Theriogenology 1990;33:202. Burns PJ, Steiner JV, Sertich PL, Pozar M, Tice TR, Mason D, Love D. Evaluation of biodegradable microspheres for the controlled release of progesterone and estradiol in an ovulation control program for cycling mares. J Equine Vet Sci 1993;13:521–4. Burns PJ, Foss R, Sarver F, Woods JA, Sissener TR, Heitland AV, Wilhelm K, Farlin M, Squires EL. Control of estrus and ovulation in mares using progesterone and estradiol microspheres in a multicenter trial. In: Proceedings of the 16th Equine Nutrition and Physiology Symposium, Raleigh, NC, 1999, pp. 194–9. Blanchard TL, Varner DD, Burns PJ, Everett KA, Brinsko SP, Boehnke L. Regulation of estrus and ovulation in cyclic mares with progesterone or progesterone plus estradiol biodegradable microspheres with or without PGF2α. Theriogenology 1992;38:1091–106. Fleury JJ, Costa-Neto JM, Burns PJ. Regulation of estrus and ovulation with progesterone and estradiol microspheres: effect of different doses of estradiol. J Equine Vet Sci 1993;13:525–8. Jasko DJ, Farlin ME, Hutchinson H, Moran DM, Squires EL, Burns PJ. Use of progesterone and estradiol in biodegradable microspheres for control of estrus and ovulation in mares. Theriogenology 1993;40:465–78. Burns PJ, Simon BW, Gilley RM, Vanderwall DK, Woods G, Squires EL. Development of new hormonal therapies for mares. In: Proceedings of the Annual Meeting of the Society for Theriogenology, San Antonio, TX, 2000, pp. 197–209. Bringel BA, Jacob JCF, Zimmerman M, Alverenga M, Douglas RH. Biorelease progesterone LA 150 and its application
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to overcome effects of premature luteolysis on progesterone levels in mares. Brasil J Anim Reprod 2003;27:498–500. Vanderwall DK, Marquardt J, Woods GL. Use of a compounded long-acting progesterone formulation for equine pregnancy maintenance of pregnancy in mares. J Equine Vet Sci 2007;27:2:62–6. Burns PJ, Thompson Jr DL, Storer WA, Gilley RM. Evaluation of sustained release progestin formulations in mares. In: Proceedings of the 7th International Symposium on Equine Embryo Transfer, Cambridge, UK, 2008, pp. 71–2. Burns PJ, Morrow CM, Abraham J. Evaluation of Biorelease P4 LA300 in the mare. In: Proceedings of the 7th International Symposium on Equine Embryo Transfer, Cambridge, UK, 2008, pp. 82–3. Storer WA, Thompson Jr DL, Gilley RM, Burns PJ. Evaluation of sustained release progestin formulations in mares. J Equine Vet Sci 2009;29:1:33–6. Squires EL, Martin JM, Jasko DJ. Reproductive response of mares after treatment with progestogens with and without the addition of estradiol. J Equine Vet Sci 1992;12:28– 32. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32:83–5. Morrow CM, Burns PJ. Evaluation of BioRelease altrenogest LA 150 for maintenance of pregnancy in mares. In: Proceedings of the Conferencia International de Caballos de Deporte (CICADE), San Jose, Costa Rica, 2007. Squires EL, Patten ML, Zumbrunnen J, Bruemmer JE, Denniston DD, McCue PM. Variation in progesterone in early gestation after non-surgical embryo transfer. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007, pp. 573–5. Daels PF, Jorge-De Moraes M, Stabenfeldt GH, Hughes JP. The effect of altrenogest on the development of secondary corpora lutea and progesterone secretion. In: Proceedings of the 12th International Congress on Animal Reproduction, vol. 4, 1992, pp. 1855–7. McCue PM, DeLuca C, Patten M, Squires EL. Effect of a sham transcervical embryo transfer and/or altrenogest administration on plasma progesterone concentrations in recipient mares. In: Proceedings of the 7th International Symposium on Equine Embryo Transfer, Cambridge, UK, 2008, pp. 73–4. Ganjam VK, McLeod C, Kiesius PH, Washburn SM, Kwapien S, Brown B, Fazeli MH. Effect of ovarian hormones on phagocytic response of ovariectomized mares. J Reprod Fertil Suppl 1982;32:169–74. Evans MJ, Hamer J, Ganson LM, Graham CS, Asbury AC, Irvine CHC. Clearance of Bacteria and non-antigenic markers following intra-uterine inoculation into maiden mares: effect of steroid hormone environment. Theriogenology 1986;26:37–50. Evans MJ, Hamer JM, Ganson LM, Graham CS, Irvine CHG. Factors effecting uterine clearance of inoculated materials in mares. J Reprod Fertil Suppl 1987;35:327–34. Bracher S, Mathias V, Sheldrick S. Estrogens and progesterone receptor concentrations in the endometrium of normal and subfertile mares. J Reprod Fertil Abstract Series 1991;7:42 (abstract 7). Cook N, Cook VM, Squires EL, Burns PJ. Effect of exogenous estradiol treatment in cyclic mares following PGF2α induced luteal regression. In: Proceedings of the 13th Equine Nutrition and Physiology Symposium, Gainesville, FL, 1993, pp. 334–9.
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32. Johnson CA, Thompson Jr DL, Sullivan SA, Gibson JW, Tipton AJ, Simon BW, Burns PJ. Biodegradable delivery systems for estradiol: comparison between poly-DL-lactide microspheres and the SABER delivery system. In: Proceedings of the 26th International Symposium on Controlled Release Bioactive Materials, Boston, MA, 1999, pp. 74–5. 33. Pinto CR, Burns PJ, Whisnant S. Effects of a single oestradiol administration on follicular dynamics and luteal function of cyclic mares. In: Proceedings of the 6th International Symposium on Equine Embryo Transfer, Havemeyer Series No. 14, 2004, pp. 92–4. 34. Thompson Jr DL, Mitcham PB, Runles ML, Burns PJ, Gilley RM. Prolactin and gonadotropin responses in geldings to injections of estradiol benzoate in oil, estradiol benzoate in biodegradable microspheres, and estradiol cypionate. J Equine Vet Sci 2008;28:232–7. 35. Kelley KK, Thompson Jr DL, Storer WA, Mitcham PB, Gilley RM, Burns PJ. Estradiol interactions with dopamine antagonists in mares: prolactin secretion and reproductive traits. J Equine Vet Sci 2006;26:517–28. 36. Douglas RH. Endocrine diagnostics in the broodmare: what you need to know. In: Proceedings of the Annual Conference of the SFT/ACT, 2004, pp. 106–15. 37. Bailey VE, Douglas RH. Induction of ovulation in seasonally-anovulatory mares with GnRH. In: Proceedings of the Annual Meeting of American Society of Animal Science, Madison, WI, 1978, p. 336. 38. Bergfelt DR, Ginther OJ. Embryo loss following GnRHinduced ovulation in anovulatory mares. Theriogenology 1992;38:33–43. 39. Johnson AL. GnRH treatment induces follicular growth and ovulation in seasonally anestrous mares. Biol Reprod 1987;36:1199. 40. Becker SE, Johnson AL. Effects of gonadotropin-releasing hormone infused in a pulsatile or continuous fashion on serum gonadotropin concentrations and ovulation in the mare. J Anim Sci 1992;70:1203. 41. Hyland JH, Wright PJ, Clarke IJ, Carson RS, Langsford DA, Jeffcott LB. Infusion of gonadotrophin-releasing hormone (GnRH) induces ovulation and fertile oestrus in mares during seasonal anoestrus. J Reprod Fertil Suppl 1987;35:211. 42. Allen WR, Sanderson MW, Greenwood RES, Ellis DR, Crowhurst JS, Simpson DJ, Rossdale PD. Induction of ovulation in anoestrous mares with a slow-release implant of a GnRH analogue (ICI 118 630). J Reprod Fertil Suppl 1987; 35:469–78. 43. Harrison LA, Squires EL, Nett TM, McKinnon AO. Use of gonadotropin-releasing hormone for hastening ovulation in transitional mares. J Anim Sci 1990;68:690–9. 44. Meyer SL, Affleck KJ, Fitzgerald BP. Induction of ovulation in anestrous mares by a continuous-release depot of a gonadotropin-releasing hormone agonist (ICH 690030). In: Proceedings of the American Association of Equine Practitioners, 1990, p. 47. 45. Mumford EL, Squires EL, Peterson KD, Nett TM, Jasko DJ. Effect of various doses of a gonadotropin-releasing hormone analogue on induction of ovulation in anestrous mares. J Anim Sci 1994;72:178. 46. Fitzgerald BP, Meyer SL, Affleck KJ, Silvia PJ. Effect of constant administration of a gonadotropin-releasing hormone agonist on reproductive activity in mares: induction of ovulation during seasonal anestrus. Am J Vet Res 1993;54:1735–45. 47. Fitzgerald BP, Peterson KD, Silvia PJ. Effect of constant administration of a gonadotropin-releasing hormone agonist
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on reproductive activity in mares: Preliminary evidence of suppression of ovulation during the breeding season. Am J Vet Res 1993;54:1746–51. Johnson CA, Thompson Jr DL, Kulinski KM, Guitreau AM. Prolonged interovulatory interval and hormonal changes in mares following the use of OvuplantTM to hasten ovulation. J Equine Vet Sci 2000;20:331–6. Loy RG, Hughes JP. The effects of human chorionic gonadotropin on length of estrus and fertility in the mare. Cornell Vet 1966;56:41–50. Sullivan JJ, Parker WG, Larson LL. Duration of estrus and ovulation time in nonlactating mares given hCG during three successive estrous periods. J Am Vet Med Assoc 1973;63:895–8. Voss JL, Sullivan JJ, Pickett BW, Parker WC, Burwash LD, Larson LL. The effect of hCG on duration of oestrus, ovulation time and fertility in mares. J Reprod Fertil Suppl 1975;23:297–301. Roser J, Kiefer FB, Evans JW, Neely DP, Pacheco CA. The development of antibodies to hCG following its repeated injection in the cyclic mare. J Reprod Fertil Suppl 1979; 27:173–9. Wilson CG, Downie CR, Hughes JP, Roser JF. Effects of repeated hCG injections on reproductive efficiency in mares. J Equine Vet Sci 1990;10:301–8. McCue PM, Hudson JJ, Bruemmer JE, Squires EL. Efficacy of hCG at inducing ovulation: a new look at an old issue. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2004, pp. 510– 13. Ginther OJ, Wentworth BC. Effect of a synthetic gonadotropin-releasing hormone on plasma concentrations of LH in ponies. Am J Vet Res 1974;35:79–81. Squires EL, Slade NP, Nett TM. Effect of GnRH analogue on duration of estrus and secretion of gonadotropins. In: Proceedings of the 8th Equine Nutrition and Physiology Symposium, 1983, pp. 273–9. Harrison LA, Squires EL, McKinnon AO. Comparison of hCG, buserelin, and luprostiol for induction of ovulation in cycling mares. J Equine Vet Sci 1991;11:163–6. Irvine DS, Downey BR, Parker WG, Sullivan JJ. Duration of oestrus and time of ovulation in mares treated with synthetic GnRH (AY-24,031). J Reprod Fertil Suppl 1975;23:279–83. Wallace RA, Squires EL, Voss JL, Pickett BW. Effectiveness of GnRH or GnRH analogues in inducing ovulation and shortening estrus in mares. In: Proceedings of the 69th Annual Meeting of American Society of Animal Science, Madison, WI, 1978, p. 215. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992, p. 281. McKinnon AO, Nobelius AM, Del Marmol T, Fiqueroa S, Skidmore J, Vasey JR, Trigg TE. Predictable ovulation in mares treated with an implant of the GnRH analogue deslorelin. Equine Vet J 1993;25:321–3. Jochle W, Trigg TE. Control of ovulation in the mare with OvuplantTM , a short-term releasing implant (STI) containing the GnRH analogue Deslorelin acetate. J Equine Vet Sci 1994;14:632–44. Burns PJ, Thompson Jr DL, Donadue F, Kincald L, Leise B, Gibson J, Swaim R, Tipton AJ. Pharmacodynamic evaluation of the SABERTM Delivery System for the controlled release of deslorelin acetate for advancing ovulation in cyclic mares. In: Proceedings of the 24th International Sympo-
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sium on Controlled Release of Bioactive Materials, Stockholm, Sweden, 1997, pp. 737–8. Fleury JJ, Squires EL, Betschart R, Gibson J, Sullivan S, Tipton A, Burns PJ. Evaluation of the SABERTM Delivery System for the controlled release of deslorelin acetate for advancing ovulation in the mare: effects of formulation and dose. In: Proceedings of the 16th Equine Nutrition and Physiology Symposium, Raleigh, NC, 1999, pp. 343–8. ¨ Mumford EL, Squires EL, Jochle W, Harrison LA, Nett TM, Trigg TE. Use of deslorelin short-term implants to induce ovulation in cycling mares during three consecutive estrous cycles. Anim Reprod Sci 1995;39:129–40. Morehead JA, Blanchard TL. Clinical experience with deslorelin (OVUPLANTTM ) in a Kentucky Thoroughbred Broodmare Practice (1999). J Equine Vet Sci 2000;20:358. Burns PJ, Betschart R, Fleury JJ, Squires EL, Nett T, Gibson J, Sullivan S, Tipton A. Evaluation of the SABERTM Delivery System for the controlled release of deslorelin acetate for advancing ovulation in the mare: effect of gamma radiation. In: Proceedings of the 16th Equine Nutrition and Physiology Symposium, Raleigh, NC, 1999, pp. 338–34 Farquhar VJ, McCue PM, Carnevale EM, Squires EL. Interovulatory intervals of embryo donor mares administered deslorelin acetate to induce ovulation. Theriogenology 2001;55:362. McCue PM, Farquhar VJ, Carnevale EM, Squires EL. Removal of deslorelin (OvuplantTM ) implants 48 h after administration results in normal interovulatory intervals in mares. Theriogenology 2002;58:865–70. Fleury JJ, Fleury P, de Sousa FA, Gilley RM. Preliminary evaluation of a BioRelease delivery system for the controlled release of deslorelin (Des) for advancing ovulation in the mare: effect of dose. Brasil J Anim Reprod 2003;27: 501–2. Fleury P, Alonso MA, Alvarenga MA, Douglas RH. Intervals to ovulation after treatment with estradiol cypionate (ECP)and BioRelease Deslorelin (BRT-Des). In: Proceedings of the 6th International Symposium on Equine Embryo Transfer, Havemeyer Series No. 14, 2004, p. 89. Kolling M, Allen WR. Ovulation induction for embryo transfer: hCG versus GnRH analogue. In: Proceedings of the International Equine Gametes Group Workshop II, Havemeyer Monograph Series No. 18, 2005, pp. 54–8. McCue PM, Magee C, Gee EK. Comparison of compounded deslorelin and hCG for induction of ovulation in mares. J Equine Vet Sci 2007;27:58–61. Besognet B, Hansen B, Daels PF. Dopaminergic regulation of gonadotrophin secretion in seasonal anestrous mares. J Reprod Fertil 1996;108:55–61. Besognet B, Hansen BS, Daels PF. Induction of reproductive function in anestrous mares using a dopamine antagonist. Theriogenology 1997;47:467–80. Brendemuehl JP, Cross DL. Influence of the dopamine antagonist domperidone on the vernal transition in seasonally anoestrous mares J Reprod Fertil Suppl 2000;56:185– 93. McCue PM, Buchanan B, Farquhar V, Squires EL, Cross DL. Efficacy of domperidone on induction of ovulation in anestrous and transitional mares. In: Proceedings of the 45th Annual Convention of the American Association of Equine Practitioners, 1999, pp. 217–18. Donadeu FX, Thompson Jr DL. Administration of sulpiride to anovulatory mares in winter: effects on prolactin & gonadotropin concentrations, ovarian activity, ovulation and hair shedding. Theriogenology 2002;57:963–76.
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New Techniques in Hormonal Delivery 79. Nagy P, Bruneau B, Duchamp G, Daels PF, Guillaume D. Dopaminergic control of seasonal reproduction in mares. In: Proceedings of the 3rd Meeting of the European Society for Domestic Animal Reproduction, 1999, p. 53. 80. Daels PF, Fatone S, Hansen BS, Concannon PW. Dopamine antagonist-induced reproductive function in anoestrous mares: gonadotropin secretion and effect of environmental cues. J Reprod Fertil Suppl 2000;56:173–84. 81. Nequin LG, King SS, Ferreira-Dias GM, Gow GM. Possible role for prolactin during the equine spring reproductive transition. In: Proceedings of the 13th Equine Nutrition and Physiology Symposium, Gainesville, FL, 1993, p. 358. 82. Nequin LG, King SS, Johnson AL, Gow GM, Ferreira-Dias GM. Prolactin may play a role in stimulating the equine ovary during the spring reproductive transition. J Equine Vet Sci 1993;13:631–5. 83. Thompson DL, Hoffman R, DePew C. Prolactin administration to seasonally anestrous mares: reproductive, metabolic, and hair-shedding responses. J Anim Sci 1997;75: 1092–9. 84. Mitcham PB, Thompson Jr DL, Thompson T, Bennett SD, Burns PJ, Caltabilota TJ. Estradiol and domperidone stim-
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ulation of ovulation in mares in winter: dose and combination studies. J Equine Vet Sci 2009;29:341–2. Irving CHG, Evans MJ. Post natal changes in total and free thyroxine and triiodothyronine in foal serum. J Reprod Fertil 1975;23:709. Chen CL, Riley AM. Serum thyroxine and triiodothyronine concentrations in neonatal foals and mature horses. Am J Vet Res 1981;42:1415–17. Lowe JE, Baldwin BH, Foote RH, Hillman RB, Kallefelz FA. Equine hypothyroidism: the long term effects of thyroidectomy on metabolism and growth in mares and stallions. Cornell Vet 1974;64:267–95. Nachriener RF, Hyland JH. Reproductive endocrine function testing in mares. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp 303–10. Burns PJ, Gilley RM. Pharmacodynamic evaluation of biodegradable DL-lactide-glycolide microparticles for a 30 day controlled release thyroxine formulation using a new solvent free extrusion process. In: Proceedings of the Conferencia International de Caballos de Deporte (CICADE), San Jose, Costa Rica, 2007.
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Chapter 199
Contraception Jay F. Kirkpatrick and John W. Turner
Downregulation of fertility in the horse is a limited subject. Historically, equine fertility control has focused on castration of the stallion. Most often, this common procedure is carried out not only to limit reproduction but also to eliminate steroid production and accompanying untoward behavior. Beginning in the 1970s, interest in equine contraception increased, largely because of uncontrolled populations of freeranging wild and feral equids, and the need for management of captive exotic equids in zoological parks.
Contraception of the stallion Steroids It has long been known that exogenous androgens can exert a downregulation of endogenous androgens and sperm production in the stallion.1 Increased doses of exogenous androgens cause a significant reduction in spermatogenesis.2 This phenomenon is caused by negative feedback on hypothalamic gonadotropin-releasing hormone (GnRH),3 and subsequent downregulation of pituitary luteinizing hormone (LH) and follicle-stimulating hormone (FSH),4 and ultimately a decline in gonadotropin-dependent spermatogenesis.5 To achieve oligospermia to the degree necessary to reduce fertility, the androgens had to be given frequently. For example, testosterone propionate (200 µg/kg) had to be administered every other day for 88 days to cause significant depression in motile sperm.6 These antifertility effects of androgen treatment were largely reversible.7 This fundamental biological strategy led to the first attempts at actual contraception aimed at free-ranging wild horses. Trials with domestic pony stallions examined the effects of six repeated monthly injections of testosterone propionate (TP) and 17-alpha-ethinylestradiol, 3-cyclopentyl ether (quinestrol), at doses of 1.7 g/100 kg. Both regimens resulted in significant degrees of oligospermia and
decreased sperm motility.8 The practicality for application in the field, however, was limited because of the need for repeated treatments. Thus, testosterone propionate was microencapsulated in a polymer of dl-lactide (mTP) (Southern Research Institute, Birmingham, AL), permitting a sustained release for up to 6 months after intramuscular injection. On contact with intercellular water, the lactide coating erodes and releases the active steroid inside. The lactide coating is converted to carbon dioxide and lactic acid. Seven experimental and eight control stallions in Idaho were immobilized from a helicopter with 300 mg succinylcholine and received injections of 5.0, 7.5, or 10 g of mTP in the hip. Stallion libido and quantitative aspects of sexual behavior, based on elimination marking behavior,9 were unaffected and breeding took place, but there was an 83% reduction in foal production compared with mares bred by control stallions. There were no differences in fertility rates between the three doses administered.10 There were concerns for the safety of stallions and the dangers and high costs associated with helicopter use and immobilizing drugs, and these led to a second field trial in which the microencapsulated TP was delivered remotely, by dart. Wild harem stallions on Assateague Island National Seashore, MD (ASIS), were given 3 g of microencapsulated TP administered with barbless darts, from the ground and without immobilization. The pharmacological success of the mTP was evident, with a 28.9% fertility rate for the mares accompanying the treated stallions and a 45% fertility rate among the mares accompanying untreated stallions. Unfortunately, the logistics of delivering 3.0-g microcapsules in four separate doses to each stallion was daunting and impractical for routine use. Vasectomy has also been tested in order to limit population growth in wild horses. In an unpublished study, two wild stallions in the Pryor Mountain National Wild Horse Range were vasectomized,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Contraception and for 2 years following the surgery, no foals appeared among accompanying mares (J.F. Kirkpatrick, unpublished data). Equally important, the stallions exhibited normal sexual behaviors, a requisite for maintaining harem groups. This experiment was later repeated with larger numbers of wild stallions in Nevada, and the results were much the same, except that it was shown that not all breeding is done by dominant harem stallions.11 Immunocontraception Interest turned to immunological approaches to contraception in the stallion. The first approach tested used an anti-GnRH vaccine (Peptide Technology Ltd, Sydney, Australia).12 Because of the molecule’s homology across mammalian species, there were difficulties in producing a strongly immunogenic vaccine against GnRH. Further studies led to a conjugated form of GnRH,13 in which the GnRH molecule was conjugated to ovalbumin. At doses of 200–400 mg there was a significant reduction in testicular dimensions, testosterone production, and daily sperm production for up to 30 weeks. More recently, Janett et al.14 injected eight adult stallions with three inoculations of 200 µg of a GnRHprotein conjugate (Equity; CSL Limited, Australia), over an 8-week period. Testosterone values, scrotal width, semen quality, and libido decreased significantly over the 8 months, but results varied among individual animals.
Contraception of the mare Steroids Logistical difficulties in treating stallions with steroids, concerns about steroid-related pathologies, and a general concern that the treatment of wild stallions would have serious genetic consequences for the gene flow in free-ranging herds turned the focus of contraception in horses to the mare. Based on experience with persistent corpora lutea15 and data that indicated that plasma progesterone concentrations in excess of 0.5–1.0 ng/mL inhibited ovulation in mares,16–18 attempts were made to administer contraceptive doses of progestins to wild horses.19 Two-gram doses of northisterone (norethindrone), microencapsulated in a slow-release lactide matrix similar to that described above for testosterone propionate, were delivered remotely to six wild mares on Assateague Island. All six treated mares produced a foal one year later, a highly improbable event among Assateague mares, where normal foaling rates seldom exceed 55%.20 In a second steroid contraceptive experiment, groups of 30 captive wild mares in Nevada were
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each implanted with silastic rods containing various doses of estradiol (E, 4–12 g) and/or progesterone (P, 8–24 g) or no hormone.21 Fewer mares receiving 8 g E, 12 g P plus 4 g E, or 8 g E plus 8 g P displayed behavioral estrus, but all animals displaying estrus, treated or control, ovulated. These data indicated a rapid decline in plasma steroid concentrations within 5 weeks of implantation and suggested increased metabolic clearance of the steroid. Because of the rapid decline in estradiol and progesterone concentrations, silastic implants containing the synthetic estrogen ethinylestradiol (EE2) or EE2 plus P were placed in captive wild horses.22 Animals pregnant at the time of implantation delivered healthy foals, and contraceptive efficacy ranged from 88% to 100% through two breeding seasons. Endocrine studies of these mares suggested that contraception was affected by blocking ovulation and/or implantation. In a similar study, intraperitoneal implants of EE2 alone also resulted in contraceptive efficacy of 75% to 100% through two breeding seasons, and rates of EE2 decline in the plasma suggested a contraceptive life of 16, 26, and 48–60 months, for 1.5 g, 3.0 g, and 8.0 g of EE2, respectively.23 A third experiment aimed at using steroids to inhibit fertility in mares utilized estradiol and progesterone incorporated in biodegradable poly (dllactide) microspheres.24 Mares were given 0.625 g progesterone plus 50 mg estradiol, 1.25 g progesterone plus 100 mg estradiol, or 1.875 g progesterone plus 150 mg estradiol. While the intervals between estrus and ovulation were lengthened by the various treatments, rates for embryo recovery (performed by uterine lavage) did not differ between groups. Results achieved with estradiol, progesterone, and ethinylestradiol in mares highlight advantages and disadvantages of natural versus synthetic steroids for contraceptive purposes in the horse. Steroids native to the mare, such as estradiol and progesterone, are required in impractically large doses due to their rapid enzymatic degradation in vivo. The use of some long-acting synthetic steroids such as ethinylestradiol may delay metabolic degradation and permit more sustained contraception. But the potential risk, however small, of the passage of these synthetic steroids to humans or non-target wildlife has made acceptance by regulatory agencies [Food and Drug Administration (FDA), the United States Department of Agriculture (USDA), or the Environmental Protection Agency (EPA)] unlikely. Immunocontraception Because of the difficulties with delivering large masses of microencapsulated steroids, dangers associated with capture and restraint of horses, surgical
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procedures associated with intraperitoneal implants, concern over long-term effects of steroid contraception, and passage of synthetic steroids through the food chain, attention turned to immunocontraception. One immunologically based contraceptive strategy involves blocking the release of gonadotropinreleasing hormone (GnRH) or its attachment to the pituitary target cells responsible for the release of luteinizing hormone (LH) and follicle-stimulating hormone (FSH), with subsequent abolition of tropic actions on the ovary or testis.25 Immunization of domestic mares against GnRH blocked ovulation in three of five mares for 4 months.26 Each mare was inoculated with 2.0 mg GnRH conjugated to human serum albumin emulsified in Freund’s complete adjuvant (FCA). Analysis of plasma LH revealed a lack of pulsatile secretion, which was correlated to antibody titers. The high variability in the mare’s response to the antigen and the subsequent variability in antibody titers suggested this approach was unreliable. When a dose of 200 or 400 mg of conjugated GnRH-ovalbumin was injected into four mares,13 ovarian activity ceased 4 weeks after preliminary treatment and caused significantly decreased progesterone secretion for up to 20 weeks. In another study,27 Goodloe et al. immunized domestic mares with GnRH conjugated to keyhole limpet hemocyanin (KLH), using as an adjuvant either aluminum hydroxide (alum) or Triple Adjuvant (TA) consisting of monophosphoryl lipid A/trehalose 6,6-dimycolate/BCG cell wall skeleton. Those mares immunized with GnRH plus TA had higher antibody titers and significantly less ovarian follicular activity. The vaccine was field tested on 29 feral mares on Cumberland Island National Seashore, GA. The vaccine was freeze-dried, sequestered in a solid biodegradable 0.25-caliber bullet, and administered by an air-powered gun (Ballistivet, Inc., White Bear Lake, MN). After embedding in the muscle of the target mare, the compressed compound forming the biobullet degrades over 24 h, releasing the antigen. A total of 25 treated mares survived until the next foaling season and 17 (68%) produced foals, which was not significantly different from control foaling rates. A more recent attempt to immunize wild horses against GnRH was carried out in Nevada.28 Mares received either 1800 or 2800 µg GnRH vaccine (National Wildlife Research Center, Fort Collins, CO) with Adjuvac adjuvant, which is a dilution of a commercial Johne’s disease vaccine (Fort Dodge, Ames, IA). Following a single breeding season, 0 of 18 mares, in both treatment groups, were pregnant on the basis of ultrasound evaluations. All GnRHtreated mares had low concentrations of serum
estrogen and progesterone, which is consistent with the predicted actions of a GnRH vaccine. Imboden et al.29 immunized domestic mares with a GnRHprotein conjugate and aqueous adjuvant (Improvac; CSL Limited, Australia). Each of nine mares received a primer and booster inoculation of 400 µg. Eight of nine mares had injection site reactions (including swelling, pain, stiffness, pyrexia, and apathy) but these subsided by 4–5 days post-injection. All test mares ceased ovarian cyclicity for 23–100 weeks, and reproductive behaviors were partially or completely inhibited in seven of nine mares. Botha et al.30 applied the same vaccine in doses of 400 µg to 55 mares, 35 days apart, and 175 days later none of the treated mares remained in anestrus, while 50% of controls were pregnant. In a similar study, Elhay et al.31 injected mares with the GnRH-protein conjugate Equity, at doses of 200 µg, and a saponin-based adjuvant. Inoculations were given at Day 0 and Day 28. Serum progesterone, estradiol, and estrus-related behavior all declined over a minimum period of 3 months. Robinson and McKinnon32 reported on prolonged (2 years) failure to express recrudescence to cyclicity in a population of adult mares in horses treated with 200 µg of Equity for racing and presented later for breeding. A second immunological approach to equine contraception is based on the identification of antibodies directed against the zona pellucida of the ovum in naturally infertile mares,33 and immunological cross-reactivity of equine zona-positive antisera and porcine sperm binding.34 Liu et al.35 immunized 10 captive wild mares and four domestic mares with the protein equivalent of 2000 to 5000 porcine zonae pellucidae (PZP). Freund’s complete adjuvant was used for the first inoculation and Freund’s incomplete adjuvant (FIA) for the 3-monthly booster inoculations that followed. Of the 14 treated mares, 13 failed to conceive during the first year. The four domestic mares all conceived during the second year, after antibody titers had decreased. A field test of the PZP vaccine was carried out on Assateague Island National Seashore (ASIS).36 Twenty-six free-ranging wild mares were remotely inoculated with the protein equivalent of 5000 PZP (65 µg) and FCA in March 1988 by means of barbless darts. The mares all received a second inoculation, with FIA, 2 weeks later, and 18 of the 26 mares received a third inoculation with FIA a month later. No foals were produced among the treated mares a year later whereas 50% of the six sham-injected control mares produced foals. Of the 26 PZP-treated mares, 14 were pregnant at the time of inoculation and all 14 produced healthy foals; thus the PZP vaccine had no effect on pregnancies in progress or the health of the foals. Once antigen recognition had taken place,
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Contraception a single annual booster inoculation was sufficient to maintain contraceptive levels of antibodies,37 and animals that did not receive booster inoculations returned to normal fertility.35, 37 Studies of PZP contraception in wild mares examined the long-term effects on ovarian endocrine function and reversibility of contraceptive action, through the use of urinary and fecal steroid metabolite analysis.38 These studies examined urinary estrone conjugates and immunoreactive pregnanediol glucuronide-like progesterone metabolites (iPDG) in mares treated from one through seven consecutive years with the PZP vaccine. Reversibility occurred in all durations of treatment except the 7-year group,39 which failed to ovulate and remained infertile over a 12-year period. Prolonged treatment (3 years or more), however, did temporally depress urinary estrogens.40, 41 Continued multi-year application of the PZP vaccine to ASIS horses, for management purposes, resulted in zero population growth within 2 years, and a marked reduction in herd population.42, 43 Associated with these long-term treatments were decreased mortality rates, increased body condition scores, and increased longevity among treated mares.44 Turner and Kirkpatrick42 observed normal foal survival and no out-of-season births for these mares,45 while Powell46 reported absence of behavioral side-effects, and Powell and Monfort47 found normal ovarian cyclicity. Additional and successful contraceptive studies with PZP and other species of equids were mounted with feral burros,48 Przewalski’s horses,49 and zebra.50 The non-seasonal nature of the burros’ and zebras’ breeding season necessitated booster inoculations more frequent than annually. Following the success of multi-injection field applications of PZP contraception in wild horses, modifications to the original protocol were tested toward maximizing vaccine (i) delivery efficiency, (ii) duration, and (iii) cost-effectiveness. Among these modifications was the substitution of Freund’s modified adjuvant (Calbiochem Inc., La Jolla, CA) for FCA51 with no loss of immunogenicity. Regarding vaccine delivery, Willis et al.52 employed the biobullet technology described above. The PZP was lyophilized and incorporated into a biodegradable 25-caliber biobullet composed of calcium carbonate and hydroxypropylcellulose. Each bullet weighed 575 mg and contained 200 µg of PZP and was adjuvanted with 500 µg of synthetic trehalose dicorynomycolate (TDCM) glycolipid and squalene oil. Each of three domestic mares was given two inoculations. Serum antibody titers were detectable for 40 weeks, and two of the three mares remained infertile. However, use of the same approach with free-ranging horses in New Zealand yielded disappointing results. Each of 26
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mares received 400 µg of PZP in biobullets, and 25 became pregnant following treatment.53 The causes for the failure were multiple. The TCDM adjuvant was likely inadequate for mounting an adequate immune response to the PZP. Because an earlier study with captive New Zealand horses, using this adjuvant resulted in highly variable antibody titers at all doses.54 Additionally, the limited range of the biobullets and the inability to determine if they had struck the intended target probably reduced effectiveness. The difficulty in delivering PZP remotely, to hundreds or thousands of wild horses in the western USA, led to the study of one-inoculation, long-acting forms of the vaccine. In an initial trial, 132 adult wild mares in Nevada were captured and freeze branded.55 Mares received one of the following treatments: (i) saline and adjuvant (FCA); (ii) two inoculations of 65 µg PZP, with FCA for the first inoculation and FIA for the second, 2 weeks later; or (iii) 65 µg of PZP + FCA along with 65-µg PZP boosters sequestered in controlled-release biodegradable lactide-glycolide microspheres, as described by Wang et al.56 and prepared by the School of Pharmacy at the University of Iowa. The microspheres, designed to release across approximately 1 month, were suspended in the PZP/adjuvant emulsion for injection. Control mares had a 54.5% foaling rate a year later, the mares receiving the standard two-inoculation treatment had a 4.5% foaling rate, and the mares receiving the one-inoculation, microsphere-delivered PZP had a 20% foaling rate. In a larger-scale test of the microspheres,57 267 Nevada wild mares were captured, freeze branded, and treated with (i) the standard two-inoculation PZP treatment, with FCA/FIA adjuvants; or (ii) a single inoculation of 65 µg PZP + FCA + 60–80 µg PZP in microspheres with no adjuvant; or (iii) 65 µg PZP + FCA + 60–80 µg PZP in microspheres with a carbomer adjuvant (Carbopol 934; B.F. Goodrich, Cleveland, OH,). Control mares had a pregnancy rate of 62.5% a year later, while 12.8% of the two-inoculation group and 11.3% of the microsphere + carbomer group became pregnant. Clearly the lactide-glycolide microsphere technology was effective in delivering a one-inoculation PZP contraceptive. However, difficulty in injecting this vaccine was encountered, because the suspended microspheres settle out over time in syringes and darts. This led to experiments with lactide-glycolide pellets, small enough to fit into a 14-gauge needle of a syringe or dart.58 The pellet-containing PZP vaccine was originally tested in domestic mares and yielded anti-PZP titer patterns across 45 weeks, which were similar to those in mares given the standard two-inoculation PZP treatment.57 The controlled-release pellet vaccine was then field tested as a one-inoculation, 1-year duration treatment on free-roaming wild mares in
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three different herds in Nevada. Fertility rates among untreated mares were 46–62% across herds, and among treated mares were 11–26% across herds. The lower fertility rates were associated with herds in which mares had been captured and hand-injected, while the higher fertility rates occurred in mares corralled and darted. As stated previously, the higher fertility rates with dart delivery likely reflected incomplete delivery of vaccine. In 2000, 96 adult wild mares in Nevada were captured and treated with a primary dose of 100 µg aqueous PZP + booster doses of PZP incorporated in controlled-release lactideglycolide polymer pellets. While previous pelletcontaining vaccine tests employed two pellets releasing at approximately 1 and 3 months, this study included a third pellet, releasing at approximately 12 months, to serve as an annual booster. Reproductive success from 2001 through 2004 was, respectively, 5.9%, 14.0%, 32.0%, and 47.5%, while untreated mares on the same range had a 53.8 ± 1.3% reproductive success rate.58 Finally, two attempts have been made to inhibit fertility in horses using intrauterine devices (IUDs). In the initial attempt, silastic O-ring-shaped IUDs (silastic 382; Dow Corning, Midland, TX) were placed in six mares that were then turned out with 12 nontreated mares and a fertile stallion.59 None of the IUD-treated mares became pregnant, while all 12 non-treated mares became pregnant. Reversibility of contraception was 100% a year later, after the IUDs were removed. Mild endometritis occurred in most of the treated mares, but disappeared after removal of the IUDs. In a second trial,28, 60 copper-containing Tcu-380A IUDs (Ortho-McNeil, Raritan, NJ) were placed in 15 mares. Each IUD contained a total surface area of 380 mm2 of copper. Although 3 of these 15 mares became pregnant, it appears that contraception failed in these animals because the IUDs were not retained. The major drawback with this approach, is the need to capture horses, and to know their pregnancy status before application.
References 1. Blanchard TL. Some effects of anabolic steroids: especially on stallions. The Compendium 1984;7:1–8. 2. Ludwig DJ. The effect of androgen on spermatogenesis. Endocrinology 1950;46:453–81. 3. Holma P, Aldercreutz H. Effect of an anabolic steroid (Metadienone) on plasma, LH, FSH, and testosterone on the response to intravenous administration of LRH. Acta Endocrinolica (Copenh) 1976;83:856–64. 4. Squires EL, Berndston WE, Hoyer JH. Restoration of reproductive capacity of stallions after suppression with exogenous testosterone. J Anim Sci 1981;53:1351–9. 5. Steinberger E. Current status of research on hormonal contraception in the male. Research Frontiers in Fertility. Regulation 1980;1:1–12.
6. Berndston WE, Hoyer JH, Squires EL, Pickett BW. Influence of exogenous testosterone on sperm production, seminal quality, and libido of stallions. J Reprod Fertil Suppl 1979;27:19–23. 7. Berndston WE, Squires EL, Thompson DL. Spermatogenesis, testicular composition and the concentration of testosterone in the equine as influenced by season. Theriogenology 1983;20:449–57. 8. Turner JW, Kirkpatrick JF. Androgens, behaviour and fertility control in feral stallions. J Reprod Fertil Suppl 1982;32:79–87. 9. Turner JW, Perkins A, Kirkpatrick JF.. Elimination marking behavior in feral horses. Can J Zool 1981;59:1561–6. 10. Kirkpatrick JF, Perkins A, Turner JW. Reversible fertility control in feral horses. J Equine Vet Sci 1982;2:114–18. 11. Asa CS. Male reproductive success in free-ranging feral horses. Behav Ecol Sociobiol 1999;47:89–93. 12. Dowsett KF, Pattie WA, Knott LM, Jackson AE, Hoskinson RM, Rigby RPG, Moss BA. A preliminary study of immunocastration in colts. J Reprod Fertil Suppl 1991;44: 183–90. 13. Dowsett KF, Tshewang U, Knott LM, Jackson AE, Trigg TE. Immunocastration of colts and immunospaying of fillies. Immunol Cell Biol 1993;71:501–8. 14. Janett F, Stump R, Burger D, Thun R. Suppression of testicular function and sexual behavior by vaccination against GnRH (Equity) in the adult stallion. Anim Reprod Sci 2009 115(1–4):88–102. 15. Stabenfeldt GH, Hughes JP, Evans JW, Neely DP. Spontaneous prolongation of luteal activity in the mare. Equine Vet J 1974;6:158–63. 16. Squires EL, Wentworth BC, Ginther OJ. Progesterone concentration in blood of mares during the estrous cycle, pregnancy and after hysterectomy. J Anim Sci 1974;39:759– 67. 17. Noden PA, Oxender WD, Hafs HD. Early changes in serum progesterone, estradiol, and LH during prostaglandin F2αinduced luteolysis in mares. J Anim Sci 1978;47:666–761. 18. Palmer E, Jousett B. Urinary oestrogen and plasma progesterone levels in non-pregnant mares. J Reprod Fertil Suppl 1975;23:213–21. 19. Turner JW, Kirkpatrick JF. New methods for selective contraception of wild animals. In: Cohn PN, Plotka ED, Seal US (eds) Contraception in Wildlife. Lewiston, NY: Edwin Mellen Press, 1996; pp. 191–208. 20. Keiper R, Houpt K. Reproduction in feral horses: An eightyear study. Am J Vet Res 1984;45:991–5. 21. Plotka ED, Eagle TC, Vevea DN, Koller AL, Siniff DB, Tester JR, Seal US. Effects of hormone implants on estrus and ovulation in feral mares. J Wildlife Dis 1988;24:507–14. 22. Eagle TC, Plotka ED, Garrott RA, Siniff DB, Tester JR. Efficacy of chemical contraception in feral mares. Wildlife Soc Bull 1992;20:211–16. 23. Plotka ED, Vevea DN. Serum ethinylestradiol (EE2) concentrations in feral mares following hormonal contraception with homogenous implants. Biol Reprod 1990;42 (Suppl. 1): 43. 24. Jasco DJ, Farlin MS, Hutchinson H, Moran DM, Squires EL, Burns PJ. Progesterone and estradiol in biodegradable microspheres for control of estrus and ovulation in mares. Theriogenology 1993;40:465–78. 25. Schanbacher BD. Active immunization against LH-RH in the male. In: Crighton DB (ed.) Immunological Aspects of Reproduction in Mammals. London: Butterworths, 1984; pp. 345–62.
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Contraception 26. Safir JM, Loy RG, Fitzgerald BP. Inhibition of ovulation in the mare by active immunization against LHRH. J Reprod Fertil Suppl 1987;35:229–37. 27. Goodloe RB, Warren RJ, Sharp DC. Sterilization of feral and captive horses: A preliminary report. In: Cohn PN, Plotka ED, Seal US (eds) Contraception in Wildlife. Lewiston, NY: Edwin Mellon Press, 1997; pp. 229–46. 28. Killian G, Miller LA, Diehl NK, Rhyan J, Thain D. Evaluation of three contraceptive approaches for population control of wild horses. In: Tirron RM, Gorenzel WP (eds) Proceedings of the 21st Vertebrate Pest Conference. Davis, CA: University of California, Davis, 2004; pp. 263–8. 29. Imboden I, Janett F, Burger D, Crowe MA, Hassig M, Thun R. Influence of immunization against GnRH on reproductive cyclicity and estrous behavior in the mare. Theriogenology 2006;66:1866–75. 30. Botha AE, Schulman ML, Bertschinger HJ, Guthrie AJ, Annendale CH, Hughes SB. The use of a GnRH vaccine to suppress mare ovarian activity in a large group of mares under field conditions. Wildlife Res 2008;35:548–54. 31. Elhay M, Newbold A, Britton A, Turley P, Dowsett K, Walker J. Suppression of behavioural and physiological oestrus in the mare by vaccination against GnRH. Aust Vet J 2007;85:39–45. 32. Robinson SJ, McKinnon AO. Prolonged ovarian inactivity in broodmares temporarily associated with administration of Equity. Australian Equine Veterinarian 2006;25:84–6. 33. Liu IKM, Shivers CA. Antibodies to the zona pellucida in mares. J Reprod Fertil Suppl 1982;32:309–13. 34. Shivers CA, Liu IKM. Inhibition of sperm binding to porcine ova by antibodies to equine zona pellucidae. J Reprod Fertil Suppl 1982;32:315–18. 35. Liu IKM, Bernoco M, Feldman M. Contraception in mares heteroimmunized with porcine zonae pellucidae. J Reprod Fertil 1989;85:19–29. 36. Kirkpatrick JF, Liu IKM, Turner JW. Remotely-delivered immunocontraception in feral horses. Wildlife Soc Bull 1990;18:326–30. 37. Kirkpatrick JF, Liu IKM, Turner JW, Bernoco M. Antigen recognition in mares previously immunized with porcine zonae pellucidae. J Reprod Fertil Suppl 1991;44:321–5. 38. Lasley BL, Kirkpatrick JF. Monitoring ovarian function in captive exotic species and free-roaming wildlife by means of urinary and fecal steroids. J Zoo Wildlife Med 1991;22:23–31. 39. Kirkpatrick JF, Turner A. Reversibility of action and safety during pregnancy of immunizing against porcine zona pellucida in wild mares (Equus caballus). Reproduction 2002; (Suppl. 60): 197–202. 40. Kirkpatrick JF, Liu IKM, Turner JW, Naugle R, Keiper R. Long-term effects of porcine zonae pellucidae immunocontraception on ovarian function of feral horses (Equus caballus). J Reprod Fertil 1992;94:437–44. 41. Kirkpatrick JF, Naugle R, Liu IKM, Bernoco M, Turner JW. Effects of seven consecutive years of porcine zona pellucida contraception on ovarian function in feral mares. Biol Reprod Monogr Series 1: Equine Reproduction VI: 1995; pp. 411–18. 42. Turner A, Kirkpatrick JF. Effects of immunocontraception on population, longevity and body condition in wild mares (Equus caballus). Reproduction 2002; (Suppl. 60): 187–95. 43. Kirkpatrick JF, Turner A. Achieving population goals in a long-lived wildlife species (Equus caballus) with contraception. Wildlife Res 2008;35:513–19.
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44. Kirkpatrick JF, Turner A. Immunocontraception and increased longevity in equids. Zoo Biol 2007;25:237–44. 45. Kirkpatrick JF, Turner A. Absence of effects from immunocontraception on seasonal birth patterns and foal survival among barrier island horses. J Appl Anim Welf Sci 2003;6:301–8. 46. Powell DM. Preliminary evaluation of porcine zona pellucida (PZP) immunocontraception for behavioral effects in feral horses (Equus caballus). J Appl Anim Welf Sci 1999;2:321–35. 47. Powell DM, Monfort SL. Assessment of effects of porcine zona pellucida immunocontraception on estrous cyclicity in feral horses. J Appl Anim Welf Sci 2001;4:271–84. 48. Turner JW, Liu IKM, Kirkpatrick JF. Remotely-delivered immunocontraception in free-roaming feral burros. J Reprod Fertil 1996;107:31–5. 49. Kirkpatrick JF, Zimmermann W, Kolter L, Liu IKM, Turner JW. Immunocontraception of captive exotic species. I. Przewalski’s horse (Equus przewalskii) and banteng (Bos javanacus). Zoo Biol 1995;14:403–13. 50. Frank KM, Lyda RO, Kirkpatrick JF. Immunocontraception of captive exotic species. IV. Species differences in response to the porcine zona pellucida vaccine and the timing of booster inoculations. Zoo Biol 2005;24:349–58. 51. Lyda RO, Hall R, Kirkpatrick JF. A comparison of Freund’s Complete and Freund’s Modified adjuvant used with a contraceptive vaccine in wild horses. J Zoo Wildlife Med 2005;36:610–16. 52. Willis P, Heusner GL, Warren RJ, Kessler D, Fayrer-Hosken RA. Equine immunocontraception using porcine zona pellucida: A new method for remote delivery and characterization of the immune response. J Equine Vet Sci 1994;14:364–70. 53. Cameron EZ, Linklater WL, Minot EO, Stafford KJ. Population Dynamics 1994–1998, and Management of the Kaimanawa Wild Horses. Science Conservation Bulletin 171. Wellington, NZ: Department of Conservation, 2001. 54. Stafford KJ, Minot EO, Linklater WL, Cameron EZ, Todd SE. Use of an immunocontraceptive vaccine in feral Kaimanawa mares. Conservation Advisory Science Notes 330. Wellington, NZ: Department of Conservation, 2001. 55. Turner JW, Liu IKM, Rutberg AT, Kirkpatrick JF. Immunocontraception limits foal production in free-roaming feral horses in Nevada. J Wildlife Manage 1997;61:873– 80. 56. Wang HT, Palmer H, Linhardt RJ, Flanagan DR, Schmitt E. Influence of formulation methods in the in vitro controlled release of protein from polyester microspheres. J Control Release 1991;17:23–32. 57. Turner JW, Liu IKM, Flanagan DR, Rutberg AT, Kirkpatrick JF. Immunocontraception in feral horses: One inoculation provides one year of infertility. J Wildlife Manage 2001;65:235–41. 58. Turner JW, Liu IKM, Flanagan DR, Rutberg AT, Kirkpatrick JF. Immunocontraception in wild horses: One inoculation provides two years of infertility. J Wildlife Manage 2007;71:662–7. 59. Daels PF, Hughes JP. Fertility control using intrauterine devices: an alternative for population control in wild horses. Theriogenology 1995;44:629–39. 60. Killian G, Thain D, Diehl NK, Rhyan J, Miller LA. Four-year contraceptive rates of mares treated with a single-injection porcine zona pellucida and GnRH vaccines and intrauterine devices. Wildlife Res 2008;35:531–9.
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Section (f): Techniques in Reproductive Examination
200.
History Walter W. Zent
1897
201.
Vaginal Examination Walter W. Zent and John V. Steiner
1900
202.
Direct Rectal Palpation Thomas R. Bowman, Jr.
1904
203.
Ultrasonography Jonathan F. Pycock
1914
204.
Uterine Cytology Michelle M. LeBlanc
1922
205.
Endometrial Biopsy Charles C. Love
1929
206.
Endoscopic Examination Claire E. Card
1940
207.
Cytogenetic Evaluation of Mares and Foals Teri L. Lear and Daniel A.F. Villagomez
1951
208.
Uterine and Clitoral Cultures Sidney W. Ricketts
1963
209.
Use of Chromogenic Agar to Diagnose Reproductive Pathogens Angus O. McKinnon and David P. Beehan
1979
210.
Evaluation of Uterine Tubal Patency William B. Ley
1988
211.
Laparoscopy Dean A. Hendrickson and David G. Wilson
1991
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Chapter 200
History Walter W. Zent
Introduction
Age
Obtaining a complete and accurate reproductive history of the mares under your care, at the beginning of the breeding season, is an important first step to a successful outcome at the end of the season. This is true whether dealing with a new mare entering the herd or continuing to manage mares that have been in your care for years. Reviewing their past reproductive performance is the first step in insuring a successful outcome to the coming season. Whether the mare is new to the farm or is a longtime resident, thinking about her and anticipating her good and bad points is a very useful exercise before you see her for the first time during the breeding season. The review of a mare’s history will allow you to formulate a plan of attack for her and may allow you to correct or evaluate some of her issues before the season starts. Remember that in any breeding program, time is your enemy and the more that can be done before the season the less time will be wasted once the season has begun. The evaluation of a mare that is being seen for the first time cannot be adequately performed without a complete and accurate history of all of the factors that affect a mare’s reproductive performance.
Age may play a very important role in the evaluation of the history of a mare. Young mares are generally thought to be more fertile than older mares. This may be generally true but there are many instances in which older mares remain extremely fertile, and also instances when young mares are not. Mares that have not reached maturity may be less fertile than their older counterparts. There is also a definite breed influence as regards age at maturity, with some breeds maturing at a later age than others. It is important to consider these factors when planning the management of mares entering the broodmare band.
Factors that can relate to reproductive potential Breed The breed of the animal can affect the fertility; some breeds are predisposed to certain fertility problems, which are thus less common in other breeds. Breed associations also regulate the ability to use advanced reproductive techniques, so in some instances reproductive performance cannot be enhanced by the use of artificial insemination, embryo transfer, or any other advanced techniques that may improve the opportunity for some problem mares to become pregnant.
Status Mares are defined in terms that have very specific applied meanings, which may be different in different breeds. The following are some of the terms and the definitions as they are used in the Thoroughbred in Kentucky. A maiden mare is a mare that is entering the breeding band for the first time and has never been bred. A maiden foaling mare is carrying her first foal, and a maiden barren mare is a mare that was bred for the first time the year before and didn’t become pregnant. A barren mare is a mare that has had foals, been bred and is not pregnant at the beginning of the season. A mare may also be listed as ‘not bred’, which would mean that she has been pregnant in the past but was not mated the previous season. A foaling mare would be considered a mare that is pregnant, and a wet mare would be a mare that is nursing a foal. These terms are for the most part self-explanatory but nevertheless should be defined so that all parties are talking about the same thing.
Past breeding records The records of mares that have been pregnant or are currently pregnant can be very helpful in the management of the mare in the future.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Techniques in Reproductive Examination
Previous foaling data The lengths of a mare’s previous gestations are important to know as mares will generally behave in a similar fashion in subsequent gestations. Mares that carry their fetuses for 320 days will tend to repeat a similar gestation length, and conversely mares that have a gestation of 360 days will continue to do likewise. Both will have normal foals that will perform well even though their gestation lengths are not within the range that is considered normal (335 to 342 days). The parity of the mare is important as the frequency and regularity of foaling is a good indicator of the mare’s fertility and may help to predict her future performance. Mares that have produced foals only every other year may have a tendency to continue to perform in that manner. Foaling histories need to be examined. Has the mare had a dystocia, is she prone to having retained placentas, or is she a difficult mare to foal? – these are questions that need to be answered. Mares that have had a difficult foaling are more apt to have vaginal and cervical damage – injuries that, even though they are pregnant, could impair future fertility. If any history of these conditions exists the mare should be examined after foaling to make sure that there is no residual damage. Some mares have unusual foaling habits – they foal standing up, savage their foal, or do other unusual or dangerous things. Many of these habits will be repeated, and so knowledge of them will assist new management in handling the mare. Any reproductive surgical procedures should be noted. Urethral extensions, perineal lacerations and correction, and even the presence of a Caslick procedure are important facts to be aware of when managing a broodmare. The removal of an ovary or the fact that a cesarean was performed, even though the mare has completely recovered, are important bits of information that can affect the performance of a mare during the breeding season. When a Caslick procedure has been performed the mare must have her vulva opened before foaling to prevent damage. Mares tend to repeat their cycling patterns. Mares that have long estrous cycles, exhibit strong signs of estrus, or no outward signs of estrus, tend to continue the same patterns, and knowledge of these behavioral traits can be very helpful. Some mares are more prone to develop anovulatory follicles or hemorrhagic follicles, and some mares may be prone to double ovulations. Knowing these behaviors ahead of time will allow the mare to be managed more effectively. If the mare has a history of uterine infection it would be useful to know what the contaminating or-
ganism was and the treatment history and outcome. Past biopsy and cytology findings should also be included in the history so that if a pattern develops it can be noted. Recent ultrasonographic finding such as the presence of uterine cyst, and the presecne of pre- and post-breeding uterine fluid, would help the new management manage the mare in a proactive manner.
Pregnancy history Mares that have a history of having twin pregnancies often will have double ovulations repeatedly. Such knowledge is beneficial, as mares with such a history are apt to have double ovulations and twin embryos again – forewarned is to be forearmed. Pregnancy losses should also be categorized. If a mare habitually loses her pregnancy before 35 days and endometrial cup formation, she will be handled differently than if she aborts after cup formation. The cause of abortion near term should be identified to enable closer monitoring of the mare in subsequent pregnancies, in an attempt to prevent similar problems from occurring. It is also important to ascertain if the mare has been managed as a high-risk pregnancy before or if she has had placentitis.
Previous athletic performance Mares that return to the broodmare band from other occupations such as showing or racing may come with a different set of problems than the non-performance mare that is put into the broodmare band. Performance horses are often given performance-enhancing drugs such as anabolic steroids, which can significantly affect their ability to cycle. Most of these products are rather transitory in their effect but can cause a considerable amount of frustration if the veterinarian at the breeding farm is unaware of the history of their use. A new antigonadotropin-releasing hormone (anti-GNRH) vaccine, which is now available in some parts of the world, can cause an even longer delay in a mare’s return to normal cycling behavior. Knowledge of this product’s use would be very important before entering the mare into the breeding herd. Animals that are retired because of injury may also become a management challenge depending on how much pain or how severe the injury. In the Thoroughbred, injuries that are so significant as to make natural covering potentially dangerous will delay breeding until the mare has recovered. In other breeds in which artificial insemination is used, the delay might not be as long.
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History
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General health
General management
An animal’s previous illnesses and treatments are important pieces of history that should be noted for any horse arriving at a farm, and certainly the broodmare is no exception. Mares in poor body condition may be that way because of lack of care or illness, and these conditions should be noted, and the animal’s health history examined.
It is often very helpful to know how a mare has been managed previously. Was she in a large group of mares or handled more individually? Was she stalled or did she live outside? Was she under artificial light? How is she teased? Is she used to being in stocks? Answers to these questions can make a mare’s transition to a new environment less stressful and in turn help the mare perform better.
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Chapter 201
Vaginal Examination Walter W. Zent and John V. Steiner
Anatomy The vagina is anatomically the passage which extends horizontally through the pelvic cavity from the neck of the uterus to the vestibule.1 The structure is tubular and approximately 15–20 cm in length and 10–12 cm in diameter when slightly distended. The pelvic wall limits the amount of distention possible. Most of the vagina is retroperitoneal and is surrounded by loose connective tissue, a variable amount of fat and a venous plexus.1 Usually, the vaginal cavity is obliterated by the walls of the vagina and is only a potential space. The anterior portion of the vagina is largely occupied by the vaginal part of the neck of the cervix. The posterior part is directly continuous with the vestibule without a definitive line of demarcation except for the transverse fold which covers the urethral orifice. In young animals, this fold is often continuous and forms the hymen. The arteries that supply the vagina are branches of the internal pudic arteries and the veins drain to the internal pudic veins. Examination of the vagina should be an essential part of a routine reproductive examination of the mare. When properly done, the examination can provide information about the presence of abnormalities and physiological states of the estrous period and cycle.2 The examination should include palpation and ultrasonography of the cervix and vagina per rectum, as well as vaginoscopic visual examination of the vagina and cervix. In addition, internal palpation of the vagina and cervix is warranted.
Equipment There are numerous types of glass, plastic, cardboard and metal vaginoscopes available for viewing the vagina (Figure 201.1). The Caslick speculum (metal) has advantages over many of the other speculums available. It provides
excellent visibility of the anterior vagina and cervix. It provides a wide view of the anterior vagina and the entire mucosa can be visualized by rotating the instrument. However, it is slightly more difficult to sterilize because of its size and more expensive than other types of speculums. However, its advantages outweigh the disadvantages. Glass speculums are somewhat outdated, but still useful. They are usually 40 cm long and 4 cm in diameter. They are less expensive and relatively easy to sterilize and clean. The glass should be at least 3 mm in thickness. Most common speculums used for routine examination in equine practice are disposable cardboard speculums that are usually foil lined (Figure 201.2). These are individually packaged and sterile. They do not provide as good a view of the vaginal mucosa but are very adequate for viewing the cervix. All of these types of speculum require some form of light source to illuminate the vagina for visualization. These include penlights of various kinds, flashlights or halogen-type illuminators. Flashlights are not as bright as the others and are not as desirable to use.
Preparation and technique Proper restraint of the mare is of primary importance when conducting an examination of the vagina.2 Some mares object to the examination and therefore protection of the examiner is important. A proper set of stocks is ideal, but backing the mare partially through a stall door with a lip twitch applied is also satisfactory. Occasionally, a mare may have to be sedated to facilitate the examination. The mare’s tail should be wrapped and held off to one side or tied to the side using the tail wrap end around the horse’s neck. This will prevent tail hairs from interfering with the examination and will also prevent contamination of the external perineal area once the mare is properly prepared.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Vaginal Examination
Figure 201.1 Metal and cardboard speculums commonly used for mare examination.
The cervix and vagina are usually assessed per rectum with palpation and ultrasonography. This is usually carried out prior to speculum examination since conducting this examination first will often cause the vagina to balloon with air and possibly interfere with the rectal examination. Generally the cervix can be evaluated for size and tone very easily with palpation per rectum. By pressing downward with the hand positioned over the cervix one can determine tone and size. Evaluation of the vaginia with ultrasonography is usually uneventful. After the rectal assessment of the vagina and cervix, the perineal area should be thoroughly scrubbed with a non-irritating soap of choice. The area should then be thoroughly rinsed with clear, clean water and blotted dry. Clean, moist cotton should be used to swab the inner lips of the vulva and clitoral fossa where bacteria and debris can reside.3 Insertion of the vaginal speculum is facilitated by lubrication of the speculum – sterile lubricants are preferred. The speculum is introduced into the vagina without undue force or irritation. It should be
Figure 201.2 Insertion of a disposable cardboard speculum.
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initially directed slightly upward and slightly rotated to avoid the opening to the urethra. Slight rotation and angulation will then be dictated by the position of the vagina. When fully inserted, the speculum should be very close to the cervix. When using the Caslick speculum, the instrument is inserted in a closed position with the handle facing laterally. Once it has been fully inserted, the Caslick speculum is rotated back into an upright position. It is then spread open to allow visualization of the vagina.2 Insertion of the vaginal speculum is usually quite easy in most normal mares. Obstructions such as a persistent hymen and acquired adhesions or constrictions are occasionally encountered and diagnosed during the examination procedure.
Examination Once the speculum is in position, the examination of the vagina should proceed in a systematic manner. The external os of the cervix should be located and visualized first. The vagina and vaginal mucosa are inspected during withdrawal of the speculum. Appearance, color, and position of the external os of the cervix are noted. In addition, the color and appearance of the vaginal mucosa is noted. Any discharges, blood or fluids are also noted. Fluid inside the vagina should be cultured for bacteria and a cytology made from the swab. If bleeding is detected then the source of the bleeding determined. Manual examination of the external os of the cervix and vagina should also be an integral part of a complete examination.4 A sterile disposable shoulder-length sleeve that is adequately lubricated with sterile lubricant should be employed. The mare’s labia should be parted slightly with the opposite hand to avoid introducing any contaminants. This portion of the examination will better facilitate the diagnosis of cervical and vaginal tears, small fistulas and adhesions, which often can be more readily felt than visualized. The appearance of the external os of the cervix and the vaginal mucosa can be correlated with certain stages of the estrus cycle. The normal vaginal mucosa in diestrus is pale pink and dry (Figure 201.3). When the speculum is initially inserted, air enters the vagina which is necessary for visualization, but allows the mucosa to become congested. Therefore, the color of the vaginal mucosa should be noted immediately upon insertion of the speculum so as to avoid artificially induced color change. Also in diestrus, the external os of the cervix is tight, erect, and off the floor of the vagina. During estrus, the vaginal mucosa is a deeper pink and glistening (moist) (Figure 201.4). There may be a small amount of clear secretion on the floor of the
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Mare: Techniques in Reproductive Examination (urovagina) is seen in older, multiparous mares. There is often a marked downward and forward deviation of the vaginal floor. Some of these mares exhibit this condition only while in estrus and estrogen levels are highest. At this time, the perineal area is most relaxed. In severe cases, the urine can flow into the uterus through an open cervix and cause endometritis and cervicitis. Various treatments have been used depending on the severity of the condition. These include swabbing urine from the vagina with sterile gauze, acupuncture, and surgery. Scars, adhesions, lacerations, and tumors
Figure 201.3 Appearance of a cervix in diestrus.
anterior vagina. The external os of the cervix is typically relaxed, edematous, slightly reddened and lies on or closer to the floor of the vagina. During pregnancy, the cervix is pale and tight, and the mucosa of the vagina in late pregnancy is coated with a thick, sticky exudate.
Abnormalities detected during vaginal examination Persistent hymen This is usually incomplete but can be complete. Usually, the hymen is thin and can easily be broken down manually. Inflammation Inflammation may be caused by pneumovagina. A complete examination of the vulva conformation will help diagnose this condition. In the absence of infection, suturing of the vulva lips is all that is needed (Caslick procedure) to take care of this condition. Occasionally, the retention of urine in the vagina
Figure 201.4 Appearance of a cervix in estrus. Note the soft and moist appearance.
These may involve the vaginal vault and/or the cervix. Small adhesions can usually be broken down manually. Lacerations, abrasions, and scars are often the result of foaling trauma and trauma during natural mating. Many of these heal spontaneously with minimal treatment depending on their location and severity. Tumors, although rare, can occur in the vaginal, cervical area.
Cervical lacerations These may involve the external os or the entire length of the cervix. These should be diagnosed and evaluated both visually and digitally. Small tears in the external os are usually of minor importance as long as the cervix has the ability to open and close normally. Large lacerations involving the external os or the entire cervix are of a more dire nature and may require surgery to repair. Many of these types of mare become relegated to an embryo transfer program if the breed registry permits. Exudates The vaginal examination can usually determine the source of the exudate. Purulent material seen exiting the cervix during estrus is a general sign of endometritis. The vaginal mucosa and cervix are generally hyperemic and the cervix is relaxed and open. Palpation of the uterus, cervix, and vagina will also enhance the diagnosis. A speculum examination should be conducted at least once during each estrous cycle for the large amount of information that can be obtained. However, continual, routine use during each normal estrus cycle is usually not warranted unless a specific condition is being monitored. Repeated speculum examination will expose the vagina and uterus to air and its subsequent inflammatory effects. This is especially important in older mares with less resistance to infection. The use of a sterile disposable speculum is recommended when routine evaluation of the vagina
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Vaginal Examination and cervix is done for determining if the mare is appropriate for breeding.
References 1. Sisson S, Grossman JD. The Anatomy of the Domestic Animals, 4th edn. Philadelphia: Saunders, 1953; p. 612.
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2. Zemjanis R. Diagnostic and Therapeutic Techniques in Animal Reproduction. Baltimore, MD: Williams & Wilkins, 1962; pp. 94–100. 3. McKinnon AO, Voss JL. Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 221–4. 4. Morrow DA. Current Therapy in Theriogenology: Diagnosis, Treatment and Prevention of Reproductive Disease in Animals. Philadelphia: Saunders, 1980; pp. 717–18.
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Chapter 202
Direct Rectal Palpation Thomas R. Bowman, Jr.
Manual palpation of the equine internal reproductive tract, per rectum, provides important information concerning both form and function. As the younger generation of veterinary graduates have become increasingly reliant upon ultrasonography for tract evaluation, less emphasis has been placed upon acquiring the skill necessary for competent manual palpation. However, when performed by an experienced palpator this procedure is an efficient cost-effective means of gaining valuable knowledge about reproductive status and breeding soundness. It provides the basis for decisions concerning further diagnostic procedures, laboratory tests, and treatment modalities. Although rectal palpation is considered a routine procedure it should never be conducted in a casual manner. The safety of all involved is of major concern. It is important to conduct each palpation in a systematic manner. Accurate record-keeping is essential for individual mare assessment as well as optimal herd breeding management. Effective interpretation of knowledge gained during palpation requires comprehensive awareness of relevant anatomy and physiology. This discussion refers to expected features of the ovaries and tubular reproductive tract in a healthy, non-pregnant mare. Reference is made to pregnancy and abnormal conditions only as it applies to the interpretation of palpation results. For detailed discussions of pregnancy and specific pathological conditions the reader is referred to appropriate sections of this text.
General comments Rectal palpation per se is a straightforward procedure. The objective is not simply to locate and identify the reproductive organs, but to assess breeding soundness and endocrine status through a
reasonable interpretation of palpation findings. The interpretation may be conclusive, such as a pregnancy diagnosis subsequent to palpating a live fetus. Or it may be presumptive, such as a pregnancy diagnosis subsequent to palpating a compatible uterine enlargement. Accuracy of interpretation is greatly enhanced by the knowledge and experience of the palpator. Palpation of the physical properties of the reproductive system provides an opportunity to evaluate the functional status of its component parts. One must be able to appreciate the synchrony of physiological events necessary for optimal reproductive performance. Inexperienced individuals should regularly palpate a group of mares through several estrous cycles. Repeated palpations will allow one to become comfortable and therefore confident with the procedure. Using the same group of mares allows one to recognize cyclic changes that occur among all mares as well as differences between individual ones. The dynamic nature of the system, seasonal effects, and individual variations make providing a reasonable interpretation challenging. Issues concerning management practices, body condition, and general health should be properly addressed. Mares that are ill, injured, in poor body condition, or suffering chronic pain cannot be considered sound for breeding. It is important to develop a pattern of thorough examination that will provide maximum information in an efficient manner. Assessment should include size and location of the ovaries and each segment of the tubular tract. Estimates of size can be obtained by relating the palpable structures to known measurements pertaining to hand width, finger width, and finger length. With practice the reliability of such estimates will become quite dependable. Absolute values are best obtained using
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Direct Rectal Palpation ultrasonography. Tissue tonicity, as well as apparent sensitivity, should also be ascertained. Information gathered in subsequent examinations will allow for rapid evaluation of seasonal and cyclic changes. This is especially important when scheduling breedings and estimating time of ovulation. There is a tendency to visualize the female reproductive tract as being suspended by supporting structures within the pelvis and posterior abdomen. Much of the reproductive tract is loosely supported by a network of broad ligamentous fibers, which originate from the sublumbar region and areas of the lateral pelvic wall. These broad ligaments attach on the dorsal aspect of the reproductive organs and converge over the body of the uterus and cervix. However, in the non-pregnant mare most support is actually provided by ventral structures – the floor of the bony pelvis and loops of bowel in the posterior abdomen (Figure 202.1). This flexible framework allows for considerable variation in the location of specific components. In young non-parous mares
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the tubular tract is often found entirely within the pelvic cavity. As the uterus enlarges it will be located primarily in the region of the pelvic brim and posterior abdominal cavity. With advancing pregnancy the uterus tends to descend towards the ventral abdomen and the broad ligaments assume a more supportive role. The cervix will be pulled forward, on, or anterior to, the brim of the pelvis. An increase in uterine size and position also affects ovarian location. As the uterus moves anteroventrally the ovaries are pulled ventromedially. During mid-pregnancy the ovaries may descend to near the level of the pelvic brim and be in close apposition. An unexpected location of various parts of the system is frequently noted during routine palpation. Because of the supple nature of the entire tract, parts are subject to be displaced by surrounding structures (distended urinary bladder, pelvic hematomas, engorged bowel, etc.). Understanding the cause and significance of such positional changes requires a working knowledge of the entire regional anatomy.
Record keeping Conclusions and decisions derived from palpation are enhanced by consistent, accurate record-keeping. Multiple-entry record sheets should be maintained to provide information concerning sequential palpations, as well as pertinent health and reproductive data. It is helpful to maximize the information on individual records by establishing alphabetical or numerical codes for various observations (cervical relaxation, uterine tone, follicular turgidity, etc.). The size of relevant structures should be recorded at each examination. Detailed records allow for rapid assessment of changes in normal structures, pre-existing abnormalities, and ongoing pathological conditions.
Abnormalities
Figure 202.1 Relationship of anterior tract to intestines: mare suspended in standing position, view from right side. Dissection of musculature has allowed abdominal contents to sag creating artifactual dorsal space (sp). A, marker for tuber coxae; B, marker for last rib; int, intestine; ov, ovary; mo, mesovarium; uh, uterine horn.
Particular attention should be directed toward observations considered to be abnormal. Abnormalities of size, structure, and location are frequently encountered. Aberrations from the expected are not always of functional significance. They may affect form, but not function; be permanent or transient; have developed recently or be relatively longstanding (Figure 202.2). Palpation alone may not suffice to conclusively identify and assess abnormal situations but clearly serves as a primary source of recognition. Careful palpation will often allow for presumptive identification of abnormal findings during an initial examination. Frequently decisions concerning the purchase, insuring, or breeding of mares must be based upon readily available information. When abnormalities are recognized, the skill, knowledge, and
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Safety and restraint
Figure 202.2 Ultrasonogram: example of longstanding uterine abnormality that does not affect reproductive performance. Left: fibrotic mass in uterine wall (margins designated by arrows). fib, fibrotic mass; en, endometrium. Right: normal pregnancy. emb, embryo (indicated by arrow); en, endometrium.
deductive ability of the palpator become the cornerstone of such decision-making. There are several non-specific conditions that affect the equine tract. Hematomas primarily involve the uterus and ovaries but are not limited to the reproductive organs. They tend to be large, spreading between fascial layers and along the extensive vascular network of the anterior tract and pelvic soft tissues. During the acute phase they are fluctuant and poorly demarcated. Palpation will often elicit a painful response. Clot formation and regression eventually result in the deposition of areas of dense, irregular, insensitive fibrous tissue. Aggressive palpation of recent hematomas should be avoided. Abscesses may also develop in any area of the tract. They can be sequelae of direct trauma, a complication from surgery, or an extension of peripheral infection. Acute abscessation results in areas of increased density that demonstrate varying degrees of tissue sensitivity. If the infective process is abated, palpable evidence will diminish and may eventually be completely resolved. Adhesions develop following primary trauma to the tract or secondary to inflammation of surrounding structures. They cannot always be identified by palpation. However, adhesion formation should be suspected whenever a portion of the tract cannot be readily differentiated from surrounding structures, or lacks the expected mobility during palpation. Cysts may also be associated with any portion of the internal system. The functional significance of cystic structures is unclear.1 However, recognition of these fluctuant structures helps to avoid confusion when evaluating follicular development or determining early pregnancy.
Rectal palpation can be accomplished in numerous farm or clinical settings. Although convenience and efficiency are important considerations, safety must be the primary objective. Concerns for safety should not only be directed toward the person performing the palpation but also include the handler and mare as well. When deciding on the location to be used, consideration must be given to the temperament of the mare and the experience level of the handler. Even well-behaved mares may become fractious when introduced into an unfamiliar setting. The site used for palpation should be well lit and dry. Insect repellent should be readily available. The objective is to adequately control mare movement, while minimizing the ability to forcefully respond in the direction of the palpator. Restraining stocks designed specifically for rectal palpation should be used whenever possible. They need to control movement without being so confining as to cause panic. The back door, including hinges and latch, should be durable enough to withstand the force of direct blows. It needs to be low enough to allow access to the mare while being high enough to protect the examiner. Problems do arise because of the restrictive structure of stocks. If a mare abruptly squats while being examined, the palpator’s arm is subject to injury. If a mare panics and attempts to escape she may collide with overhead obstacles, come to rest partially on top of the stock, or lose her footing and become wedged within it. These situations lead to further struggling and a cycle of self-destruction may ensue. The potential for serious injury to the mare can be lessened by constructing the stock with safety as a priority. Support beams and overhead structures should be minimized and well-positioned. The front, back, and sides should be of durable material and preferably extend completely to the floor. In addition to the rear door, a second exit option is advisable. Padding can be installed, especially over the top edge and corners. Further restrictive devices designed to squeeze or tie down the mare should be avoided (Figures 202.3 and 202.4). When stocks are not used, a means of restricting lateral movement is important. A stall door frame works well but the end of any solid wall, even the outside corner of a building, will suffice. It is important that the mare be made to stand close to the frame or wall on the same side as the examiner (Figure 202.5). Some mares are calm enough to be examined in an unrestricted setting. In this instance the handler should be positioned on the same side as the palpator. It is tempting to put objects behind the mare. These may actually increase the risk of injury to the
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Direct Rectal Palpation
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Figure 202.3 Padded palpation stock. Note the side exit door, and walls and doors extending completely to the floor. Figure 202.5 Palpation using stall door frame to restrict lateral movement.
Figure 202.4 Palpation stock in use. Note the metal reinforced latch and doors.
palpator. Solid objects can, when kicked, fragment or be launched backwards. Bales of straw are frequently used. Because of their thickness they present a different concern. One must lean well forward in order to reach over the bales. This off-balance position, with head and shoulders extended forward, makes one particularly vulnerable to injury. Additional means of restraint are advisable. Serious injuries are not always caused by ill-tempered mares. Mares that are straining, frightened, offbalance, or simply responding to an insect bite can react in a dangerous manner. Supplemental restraint, usually a lip twitch or chain over the upper gum, helps to avoid possible injury. The chain is well tolerated by most mares and is easier to apply than a twitch. Occasionally chemical restraint is required. Tranquilizers and sedatives, either singly or in combination, may be used. Some examples are acepromazine maleate, xylazine hydrochloride, detomidine hydrochloride, and butorphanol tartrate. A low-dose combination of detomidine and butorphanol provides adequate sedation while reducing the kick response. Consideration should be given to other effects generated by these drugs. Alterations
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Mare: Techniques in Reproductive Examination
in tissue tone and sensitivity can make evaluation of function difficult. Care should be taken to assess the known risks to pregnancy for any chemical used. In a farm setting, clients should be advised of potential problems if a sedated mare is returned to a herd or field situation.
Supplies Supplies necessary for palpation are minimal. Disposable plastic sleeves are available in various sizes and thicknesses. Some palpators prefer to fit an additional examination glove over the sleeved hand. Other protective apparel, such as coveralls, aprons, and shoulder protectors, is also available. Lubricants prepared especially for internal palpation may be purchased from numerous suppliers. The primary ingredient, carboxymethylcellulose, is water soluble and is easily cleaned from the perineal area. However, contact with loose, watery fecal material may serve to negate the lubricating properties of commercial products. Thicker preparations using an available powder will help overcome this problem (e.g. J-Lube; Jorgensen Laboratories, Loveland, CO). Other non-irritating liquids such as mineral oil or a mild soap solution may also be used. Mineral oil is readily available and provides superior lubrication. It is not water soluble, thereby providing better residual lubrication than commercial carboxymethylcellulose preparations.
Technique Palpation should proceed only after the mare has been properly positioned and restrained. Care should be taken to avoid abrupt or alarming movement. Most mares respond favorably when handled in a calm and reassuring manner. Approach should be from the side rather than directly from behind. If a stock is not being used it is advisable to place the nonpalpating hand high on the mare’s hindquarters in the area of the tuber coxae. This will enable the palpator to quickly push away if difficulty ensues (Figure 202.6). It is preferable to immobilize the mare’s tail prior to palpation. Ideally, an assistant will be available; otherwise the palpator can firmly grip the tail with the free hand. Liberal amounts of lubricant should be applied to the sleeved arm and the mare’s anal area. With the hand in a ‘cupped’ position, it is slowly advanced through the anal sphincter. In some instances this is met with considerable resistance and the hand needs to be introduced in progressive fashion, one or two fingers at a time. The rectal cavity should be evacuated of fecal material. To
Figure 202.6 Palpation in unrestricted setting. Note the position of the free hand.
avoid mucosal irritation the hand should always be advanced with fingers in close apposition. It is not unusual to encounter strong, prolonged waves of peristalsis. These may subside if efforts to palpate are ceased and the arm remains passively in position. Alternately, withdrawing from the rectum and waiting a few seconds while providing additional lubrication to the sleeve may achieve the same objective. Occasionally a mare will continue to strain during efforts to resume palpation. Attempts to forcefully overcome this straining by ‘fighting’ the mare should be avoided. Not only is effective palpation impossible, but serious injury to the rectal wall may result. There are several means of addressing this problem. Increasing the force of restraint may adequately divert the mare’s attention. Increasing the dosage of chemical agents may render the mare less capable of resisting. There are also chemicals that act directly upon the gut wall. Lidocaine can be applied locally to the rectal mucosa. Using a hand-guided insemination pipette, 20 mL of lidocaine is deposited into the evacuated rectal lumen and immediately applied to as much mucosa as possible. Probanthine bromide may be given and is reported to provide good relaxation.2 Injectable N-butylscopolamine bromide also effectively reduces peristalsis. This
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Direct Rectal Palpation potent anticholinergic agent is effective at levels onehalf or less the recommended therapeutic dosage (T. Bowman, personal observations). These chemicals will quickly result in lowered resistance. But it should be recognized that less resistance may actually heighten the potential for tissue damage by allowing more aggressive palpation. Chemicals should never be used as a substitute for patience and proper technique.
Palpation
Cervix The cervix is located by pressing an open hand ventrally against the bony pelvis and ‘sweeping’ from side to side until the cervix’s tubular structure is identified. Features such as thickness, length, and tone provide information relative to anatomical and physiological status. During early pregnancy the cervix is firm, relatively long and narrow, and well demarcated from the uterine body. During diestrus these characteristics are less distinct, but its tubular nature is still easily recognized. During estrus the relaxed, edematous cervix may be difficult to identify. It is less tubular and similar in tone and consistency to that of the uterine body. It is suggested that the cervix be assessed before proceeding anteriorly. Palpable properties of the cervix suggest findings for the rest of the tract. When cervical properties do not correlate with teasing behavior, or appear out of synchrony with palpation of the anterior tract, an explanation of the disparity should be determined. For example, there are instances where a mare will vigorously reject a teaser stallion despite having a palpably relaxed cervix. Further investigation may demonstrate significant follicular activity and the presence of uterine edema, indicating the mare’s teasing behavior to be unreliable. However, it is not unusual for a mare in early pregnancy to demonstrate signs of returning to estrus. In such case a firm, elongated, ‘pregnant’ cervix would rule out aggressive uterine palpation until further pregnancy evaluation can be completed. At times interpreting cervical palpation is difficult. Relaxation is not always indicative of estrogen related effects. Relaxation is often observed in anestrus mares. In this instance it is presumed to reflect the absence of endocrine stimulation rather than cyclic change. Significant injury can destroy tissue integrity to the extent that the cervix is difficult to identify by palpation. Cervicitis, severe expansion during parturition, or delayed postpartum involution will also result in a poorly defined, apparently relaxed cervix. Conversely in many maiden mares the cervix will seem ‘closed’ similar to the diestrous older mare,
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despite other reliable physiological signs of estrus. However, with experience the subtle softening of a maiden’s cervix is usually discernible. When cervical relaxation cannot be appreciated by rectal palpation, especially in maiden mares, direct palpation per vagina will provide additional information.
Uterus The uterus is located by first advancing anterior to the pelvic brim. With apposed fingers directed ventrally the hand is slowly withdrawn until the free, ventral border of the uterine body is engaged. The non-gravid uterus is located partially within the posterior abdominal cavity. Initial evaluation is achieved by suspending the uterus in a cupped hand with the ventral surface supported by the fingertips. By sliding from the tip of one horn to the tip of the other, a general impression of size and symmetry can quickly be gained. Horn and body diameter can be estimated by encircling each section within thumb and forefinger (Figure 202.7). A disparity in horn size is usually detectable in all but maiden mares. Tone and consistency are best appreciated by gently capturing the uterus between thumb and forefinger and systematically evaluating the entire organ. Endocrine status directly affects uterine tone and consistency. During anestrus the uterus is thinwalled, flaccid, and often difficult to define. During diestrus the progesterone-dominated uterus is firm, tubular, and easily identified. These properties may be accentuated during early pregnancy.3 During proestrus and estrus, estrogen-related edematous changes cause the uterine wall to relax and thicken. Although still well defined it will seem softer, more pliable, and less tubular. Gentle palpation allows for recognition of subtle regional inconsistencies, abnormal structures, or fluid accumulations. Deeper
Figure 202.7 Palpation of the uterine horn. uh, uterine horn; bl, broad ligament.
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more aggressive palpation will yield additional information. Endometrial folds can be identified as they slip between compressed thumb and forefingers. Endometrial cysts are often recognized during deep palpation. Aggressive palpation and compression of the uterus should not be attempted until one is absolutely certain the mare is not pregnant.
Oviduct The firm, convoluted oviducts may be identified as they course along the broad ligament from ovary to uterine horn. Abnormalities of the equine oviduct are rare and difficult to detect by palpation.4 However, identification can serve as a useful landmark when locating an ovary proves to be difficult.
Ovaries Each kidney-bean-shaped ovary is located lateral and slightly dorsal to its corresponding uterine horn. Each is normally positioned immediately cranial to the shaft of the ileum. A portion of the broad ligament, the mesovarian, encloses the dorsal convex border and functions as a supportive structure. In the non-pregnant or early pregnant mare the ovaries lie on top of intestines and the mesovarian is not fully extended. This loose attachment allows considerable mobility during palpation. The ovary should be completely examined along its entire surface and all structures noted according to location, size, consistency, tone, and apparent sensitivity. By grasping an ovary in the palm of the hand, the palpator can use fingers and thumb to search the anterior half of its surface (Figure 202.8). It is then rotated 180 degrees and the opposite half examined in similar fashion. For orientation purposes, two structures are particularly useful. First the cranial half of the ovary is attached to the cranial or free edge of the mesovarian. Second the ovulation fossa is located on the convex ventral ovarian surface. Examination of the ovary may be impeded by its ligamentous attachment, and it is often necessary to rotate it craniomedially in order to gain unencumbered access to its entire surface (Figure 202.9). When repeated attempts to locate an ovary are unsuccessful the possibility of previous ovariectomy should be considered. The equine ovary is a dynamic structure that exhibits marked seasonal and cyclic change. Each is about 7–8 cm long, 3–4 cm thick, and one is usually larger than the other.5 A classification of equine ovaries involving seasonal and structural variation has been described.6 Anestrus is associated with little, if any, palpable activity. The ovaries are small and firm. During transition, increased activity is characterized by overall enlargement and decreased
Figure 202.8 Palpation of the right ovary using the left hand. ov, ovary; mo, mesovarium.
firmness. Numerous follicles develop in seemingly random patterns of growth and regression. These follicles will often occur in groups or ‘clusters’, and the ovary may seem abnormally large. Eventually one or more ovulate and regular estrous cycles begin. When evaluating ovarian structure and activity, consideration should be given to all influencing factors including body condition, foaling history, previous cyclic activity, and seasonality.
Figure 202.9 Craniomedial rotation of the right ovary (indicated by arrow). ov, ovary; mo, mesovarium.
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Direct Rectal Palpation Ovarian palpation is most often used for evaluating follicular development and estimating time of ovulation. Ovulation occurs at the ovulation fossa but initial follicular growth is apparent on the surrounding convex surface of the ovary. A follicle is first identified as a firm, round structure projecting above the ovarian surface. Although several follicles may be present, one or two will rapidly enlarge with the onset of estrus. The average growth rate is about 3 mm/day for the 7 days preceding ovulation.7 Preovulatory follicles develop in a predictable pattern of maturation that can be likened to the ripening of a smooth-skinned fruit. Initially the follicle increases in size while remaining firm and symmetrical. In light horse breeds the mean diameter of ovulatory follicles is between 40 and 45 mm. However, maximum follicle diameter is affected by many factors.8 Prior to ovulation, growth ceases,9 the ‘ripened’ follicle becomes fluctuant and loses symmetry. However, not all follicles undergo appreciable lessening of tone prior to ovulation.10 When a follicle is palpated soon before ovulation the mare will often exhibit signs of tenderness or increased sensitivity. Ovulation is signaled by a rapid collapse of the follicle. The remaining depression slowly fills with capillary blood and eventually becomes distended above the ovarian surface. This fluctuant structure, the corpus hemorrhagicum, is easily mistaken for a mature pre-ovulatory follicle. Careful palpation will demonstrate a pulpy consistency to the ‘mistaken’ follicle due to fibrin organization within the developing clot. Additionally the corpus hemorrhagicum is usually smaller than the follicle that it has replaced. The corpus hemorrhagicum rapidly organizes and retracts into a firmer structure, the corpus luteum. The corpus luteum is difficult to identify by palpation for more than a few days post-ovulation.11 The palpable characteristics of a developing corpus luteum can be similar to those of a maturing pre-ovulatory follicle. When there is cause for doubt these structures may be differentiated using ultrasonography. Detailed record-keeping will enhance one’s ability to predict ovulation. Changes in follicular size and turgidity become easier to recognize. This is especially important when a mare does not ‘follow the rules’ and ovulates a small follicle, a firm follicle, or maintains a large soft follicle for several days. Most mares will follow the same ovulatory pattern through successive estrus cycles (T. Bowman, unpublished data). Prediction becomes more dependable when previously recorded information is readily accessible. Ovarian palpation may reveal structural changes inconsistent with normal ovulatory function. Large, persistent anovulatory follicles are associated with spring and fall transition, but may also occur during
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the fertile breeding season. Although ovulation does not occur, these follicles frequently fill with blood and most develop into luteal tissue.12 Ovarian hematomas result from excessive bleeding following ovulation. Palpable evidence of a regressing hematoma may persist for several months. Ovarian tumors are rare, with the granulosa cell type being most common. This benign tumor should be suspected when a palpable ovulation fossa cannot be located. The opposite ovary will be small, hard, and inactive. Ultrasonography is an essential diagnostic tool when dealing with ovarian abnormalities.
Ultrasonography Diagnostic ultrasonography has emerged as a routine procedure for reproductive evaluation in the broodmare. The usefulness and advantages of ultrasound are undeniable. However, reliance upon the use of ultrasound has diminished the perceived need to become proficient at manual palpation. This is unfortunate because skilled palpation can provide information not discernible by other means. Tissue tone, texture, and sensitivity cannot be appreciated without tactile evaluation (Figure 202.10a–c). Restrictions of mobility, especially resulting from adhesion formation, may go unnoticed during ultrasound examination. Recent ovulations, not apparent by ultrasound, can be detected by palpation. When parts of the tract are obscured by surrounding structures, smaller than expected, or in an abnormal location, they may be ‘out of view’ using ultrasound. Significant structures may not be visible despite being detectable manually. Structures that seem out of place during ultrasonography may be better identified by palpation. Such would be the case for an apparent follicle that proves to be a periovarian cyst. Finally, there are instances where an ultrasound machine is unavailable or inoperable. A skilled palpator should be able to proceed without technological assistance.
Suggestions The author feels compelled to issue the following words of caution. They are especially directed toward veterinary students and recent graduates. There are times when proceeding with palpation is not wise. Reasonable working conditions are necessary to conduct a thorough palpation. Unsafe facilities predispose to serious accidents and devastating injuries. Overzealous attempts at palpation may have disastrous consequences and lead to costly litigation. 1. Work to become equally proficient palpating with either hand. Numerous circumstances may
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(c) Figure 202.10 Series of ultrasonograms demonstrating early pregnancy loss (indicated by initial palpation): (a) Apparently normal 14-day pregnancy (uterus less tubular than expected). ev, embryonic vesicle; en, endometrium. (b) Two days later: vesicle appears normal but has not enlarged appreciably; echogenic fluid is visible around the vesicle; uterine tone is markedly decreased. ev, embryonic vesicle; en, endometrium; fl, fluid. (c) Three days later: the vesicle is no longer visible; fluid has markedly increased, and the uterus is flaccid. en, endometrium; fl, fluid.
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indicate the preferential use of one hand versus the other. Facilities may dictate palpation in a position favoring one side of the mare. Anatomical abnormalities may be easier to identify or ‘work around’. Palpating using the backhand position might put the palpator at more risk for shoulder and elbow injuries. Personal injury may necessitate ‘switching hands’ in order to carry on with routine daily palpations. Obtain a proper history for the mare. Quite often mares are sent to farms or clinics with notes concerning dangerous behavior, previous injuries, or pertinent health conditions. Pay attention to this information. If there is a perceived danger due to unsafe facilities, lack of capable assistance, or an unmanageable mare, one should re-evaluate the situation. Often a simple change is all that is necessary. If the mare is too small to allow safe passage of hand and arm into the rectum it is best to seek someone more suitable for the task. Vigorous attempts to overcome this problem can result in serious injury to the mare. There are instances where even experienced palpators cannot ‘find’ a part of the tract. When this is caused by a transient condition, such as a distended urinary bladder, or displaced loop of bowel, it is prudent to wait until the situation is resolved. If blood is seen at any time, during or after palpation, the source should be determined and appropriate action taken immediately. Repeated palpation should not be attempted until the origin of bleeding has been identified. A disposable vaginal speculum is useful for viewing the rectal mucosa. No matter how inocuous the injury may seem, it should never be attended to in a casual manner.
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References 1. Holyoak GR, Ley WB. Management regimens for uterine cysts. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; p. 122–5. 2. Baker CB, Kenney RM. Systematic approach to the diagnosis of the infertile and subfertile mare. In: Morrow DA (ed.) Current Therapy in Theriogenology. Philadelphia, London, Toronto: W.B. Saunders, 1980; p. 721–36. 3. Ginther OJ. Maternal aspects of pregnancy. Reproductive Biology of the Mare. Ann Arbor: McNaughton and Gunn, Inc., 1979; p. 217–54. 4. Bennett SD. Diagnosis of oviductal disorders and diagnostic techniques. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; p. 78–82. 5. Sisson S. The Anatomy of the Domestic Animals, 4th edn (revised by Grossman JD). Philadelphia, London: W.B. Saunders, 1964; pp. 606–7. 6. Baker CB, Kenney RM. Systematic approach to the diagnosis of the infertile and subfertile mare. In: Morrow DA (ed.) Current Therapy in Theriogenology. Philadelphia, London, Toronto: W.B. Saunders Co., 1980; pp. 724–6. 7. Ginther OJ. Follicles Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains: Equiservices, 1986; p. 133– 54. 8. Newcombe JR. The follicle: practical aspects of follicle control. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; p. 14–21. 9. Bergfelt DR, Adams GP. Ovulation and corpus luteum development. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; p. 1–13. 10. Newcombe JR. Ovulation and formation of the corpus luteum. J Equine Vet Sci 1996;16:48–52. 11. Ginther OJ. Characteristics of the ovulatory season. In: Reproductive Biology of the Mare. Ann Arbor: McNaughton and Gunn, Inc., 1979; p. 133–68. 12. Rickets S, Troedsson MHT. Fertility expectations and management for optimal fertility. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; p. 57.
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Chapter 203
Ultrasonography Jonathan F. Pycock
Ultrasonographic examination of the reproductive tract of the mare is an essential component of the routine gynecological investigation of the mare. The technique has advanced our understanding of both the normal and the abnormal and given both clinicians and researchers a tool to examine the uterus and ovaries non-invasively. As the ultrasonographic machines increase in their sophistication and portability, it is likely our knowledge will continue to grow and develop. It remains important to perform an initial rectal examination. This ensures removal of all fecal material, facilitates rapid location of the tract during scanning, and provides information on texture of structures. The many applications of diagnostic ultrasonography of the mare’s reproductive tract are listed below.
Ovaries
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Assessment of follicular size and number of preovulatory follicle(s); Assessment of follicular shape; Observation of ultrasonographic changes in the preovulatory follicle(s); Examination of the appearance, development, location, and morphology of the corpus luteum; Diagnosis of ovulation or ovulation failure; Diagnosis of ovarian neoplasia, periovarian cysts, and other developmental abnormalities.
Uterus
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Estimation of stage of estrous cycle; Pregnancy diagnosis; Management of twins; Monitoring fetal development; Fetal sexing; Diagnosis of imminent or actual early embryonic death;
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Diagnosis of uterine pathological processes including intrauterine fluid, uterine cysts, air, debris, hematoma, and neoplasia.
Principles The ultrasound transducer contains piezoelectric crystals which, when electrically stimulated, generate a series of high-frequency sound waves which are transmitted into the tissue immediately below it. Different tissue interfaces reflect a proportion of the sound waves back to the transducer where the same crystals convert the sound waves into electrical energy, creating an image through a series of white dots which are displayed on a screen. Different gray scales are then assigned to the returning echoes to build up an image of the structure. The ultrasound signal reflected varies in strength and is processed by the ultrasound machine into a series of dots which forms the displayed image. The brightness of each dot is proportional to the amplitude of the returning echo. In general, the greater the density of tissues, such as muscle or bone, the greater the strength of the echo produced. These dense tissues reflect most of the ultrasonic beam and the image on the screen appears white, termed echoic. Fluid provides little impedance to the ultrasound signal until an interface with an adjacent tissue of different density is encountered. Fluid-filled structures appear black or anechoic on the screen. Depending upon their ability to reflect sound waves, other tissues are seen in various shades of gray. Somewhat confusingly, gas appears white or hyperechoic because it causes a strong echo. Ultrasound measures difference in conductivity from a ‘normal’ or ‘typical’ soft tissue value of 1540 m/s, which is an enormous difference to air which conducts the sound waves at 340 m/s. Diagnostic ultrasonography in mare reproductive examinations typically employs high-frequency
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Ultrasonography sound waves, usually between 2 and 10 megahertz (MHz), or millions of cycles per second. Higherfrequency sound waves are attenuated more than lower-frequency sound waves, but they generate an image with better resolution. Resolution can be simply defined as the ability to detect small differences in tissue density. In reality, resolution is a complicated feature of ultrasonography, involving an in-depth understanding of the physics of ultrasound. Resolution can be broken down into detail, contrast, and temporal resolution and for a comprehensive review of resolution in particular and the physics of ultrasound in general, readers are referred to Reef1 and Powis.2 Higher-frequency transducers (7.5 MHz) are typically used for routine mare gynecological examinations where the structures requiring detailed study are close to the transducer. Lower-frequency transducers (3 MHz) are used for viewing larger structures at a greater distance from the transducer such as following the development of an older fetus by transabdominal scanning. There are various modes of display of the image and mare reproductive examinations use a Bmode display. B-mode refers to brightness mode (gray scale) and is a two-dimensional display of the returning echoes as white dots (pixels) produced on a screen. The greater the amplitude, or power, of the returning echo, the brighter the dot on the screen. The tissue within the scan plane is imaged as a crosssection and the location of the dot on the screen corresponds to the location of the echo reflector within the tissue. This cross-section can be viewed as a single frozen image (static B-mode) or, more commonly, several frames are displayed within 1 second, producing a continual visual image of tissues. By refreshing the frame each time the next echo pulse sweeps the cross-section, the structure and motion are observed in real time. B-mode, real-time, is the typical modality of imaging the reproductive tract of the mare.
Equipment B-mode, real-time ultrasonographic transducers can be divided into linear, including curved (convex) linear-array, and sectoral. Linear-array transducers are the preferred design for mare reproductive examinations. They are solid-state transducers with the ultrasound crystals arranged in rows forming a rectangular image corresponding to the length of the active, crystallized portion of the transducer. The number of crystals within the transducer varies between manufacturers. A linear-array transducer is usually held within the rectum in the longitudinal plane. Since the uterus of the mare is ‘T-shaped’, the uterine body appears as a rectangular image in the longitudinal
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A/D = Analogue to Digital Converter Figure 203.1 Diagrammatic representation of difference between digital and analog machines. (Courtesy of BCF Technology Ltd, Livingstone, Scotland, UK).
plane. To image the uterine horns and ovaries the transducer should be rotated slowly to the right and then the left side. Therefore, the uterine horns appear as circular images in cross-section. Curved lineararray transducers display the image on the screen as a curved pie-shaped (or sector-like) image. Sectorscanner transducers produce a beam that is triangular in shape and penetrates further into the tissue than linear-array transducers of the same frequency. Sector-scanner transducers are large and difficult to insert into the rectum of the mare and orientate correctly to obtain a meaningful image. For this reason they are not widely used in mare reproductive examinations. The last five years has seen a change from analog images to digital images being displayed on the screen. Any scanner purchased for mare reproductive examinations should be a digital machine. In a digital ultrasound system, the signal is processed into a digital image as soon as it is received by the electronics within the machine (Figure 203.1). Digital images can be reproduced and manipulated with no loss of quality, unlike earlier analog images. A digital image has clear and sharp resolution compared with an analog image which is ‘noisy’ (more artifacts) and less focused (Figure 203.2). As an analogy, think of the comparison of cassette to CD quality music and the associated ‘hiss’ and other interference you hear with audio tapes. Of course, digital images are easy to be saved and stored on laptops and PCs, but it is important to understand that, in the first place, digital refers to a processing of the image rather than its subsequent management and storage.
Machine functions The quality of the ultrasound image is dependent upon the operator. It is vital that the user understands how to optimize the quality of the ultrasound image using the variety of controls on the ultrasound machine. The instrument controls can be divided into primary and secondary controls.
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Display
Display Digital Image Clear and sharp resolution
Analogue Image Noisy and less focused
Figure 203.2 Diagrammatic representation of digital and analog images. (Courtesy of BCF Technology Ltd, Livingstone, Scotland, UK).
Primary instrument controls The primary functions of the machine are as follows:
Brightness and contrast The optimum brightness is determined by the lighting conditions. The contrast should be adjusted so that all the shades of the gray bar (usually located on the left of the screen) can be seen. The gray scale or, in technical terms, the dynamic range, can be changed by the operator on most ultrasound machines.
Power Power is the voltage delivered to the crystals in the transducer. Increasing the power increases the amplitude of the outgoing echoes. However, the higher the power setting, the more artefact (noise) compared with lower power settings. This is due to distortion of the signal.
justed so that the furthest-away structure of interest is towards the bottom of the screen. This ensures maximum magnification and the best-focused image (Figure 203.4). Secondary instrument controls The secondary functions of the machine are as follows:
Frequency The operator should begin the examination by selecting the highest frequency that allows enough penetration for the furthest structure of interest. Many ultrasound transducers only operate at a single frequency. More recent machines often have widebandwidth (multi-frequency) transducers. These
Gain The returning ultrasound signals to the transducer need to be converted into larger voltages that can be processed by the ultrasound machine. The gain is used to amplify the returning echoes and therefore alter the intensity level on the screen. The gain controls should be used to obtain optimal image quality rather than increasing power. Most machines allow the gain to be preferentially increased in particular sections of the displayed image. There is usually a set of sliding buttons which allow the screen to be segmented for selectively increasing the gain (Figure 203.3). The gain increases can be: Overall; Near (top half of the screen); Far (bottom half of the screen). Gain increases make the picture look brighter and decreasing the gain gives a darker image. Increasing the gain amplifies all signals, including background ‘noise’ (artefact).
Scale and depth Markers on the side of the ultrasound screen nearly always represent 1 cm. These controls should be ad-
Figure 203.3 Sliding buttons used to selectively increase the gain. (Courtesy of BCF Technology Ltd, Livingstone, Scotland, UK).
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Figure 203.4 (a) Left image has optimum magnification and focus as the structure fills the screen compared with (b) the right image of the same structure viewed on a larger scale. (Courtesy of BCF Technology Ltd, Livingstone, Scotland, UK).
machines can have up to three different switchable frequencies. These are sometimes labeled as:
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Penetrating (lower frequency) General (medium frequency) Resolution (higher frequency)
Variable focusing Focusing the beam reduces beam diameter, improving the resolution. There are two different methods of focusing the image on ultrasound machines:
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Distinct focus points (typically up to four) (Figure 203.5) Dynamic focus bands (a feature of digital ultrasound machines) (Figure 203.6)
tronic transducers. Increasing the degree of focusing reduces the number of frames per second and causes staggering of the image.
Dynamic range The dynamic range is the ratio of the largest to smallest amplitude that the system can process.3 The dynamic range is a contrast function of the ultrasound machine (not the screen). Most ultrasound scanners display 256 shades of gray. Each shade of gray is assigned to an ultrasound level of the reflected image. The relationship between these levels can then be varied and this changes how black and white the image looks. In general, high contrast images are good for mare reproductive examinations whereas a smoother, less contrasted image with lots of grays is better for late fetal assessment.
The focal zone is fixed in mechanical sectorscanner transducers and can only be adjusted on elec-
Figure 203.5 Arrows circled indicate distinct focus points. (Courtesy of BCF Technology Ltd, Livingstone, Scotland, UK).
Figure 203.6 Line circled indicates a dynamic focus band. (Courtesy of BCF Technology Ltd, Livingstone, Scotland, UK).
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Frame rate The frame rate is the number of frames displayed per second on the ultrasound screen. The frame rate is shown as frames per second on the ultrasonographic display (Figure 203.7) and is a function of:
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focus points dynamic range frame rate frame averaging.
Imaging modes amount of focus viewing area depth of imaging.
The lower the frame rate, the greater the resolution and the higher the quality of the image. A low frame rate is only acceptable when there is little or no movement of the structures being imaged. A low frame rate can result in the inability to detect fetal heart beats. High frame rates are needed to resolve motion in real time. If hand movement is too rapid across the structures being imaged, blurring or ‘smearing’ of the image may occur and this can be removed by increasing the frame rate (or slowing hand movement). Many modern ultrasound machines use a process known as frame averaging or correlation in which data is stored from previous images (up to three) and a weighted average value calculated. This reduces random background noise, and static structures are enhanced. If the structures of interest are moving rapidly then frame averaging will cause ‘smearing’ of the image. Frame averaging should be maximized for static structures and reduced where quick movement is present. In conclusion, it is important to take the time to set up your ultrasound machine properly as this will make a big difference to your imaging. All of the following need to be considered:
r r r r
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brightness and contrast suitable gain levels scale or depth of view frequency
Figure 203.7 Frame rate (circled) is shown as frames per second.
B-mode or brightness mode (2-dimensional, real-time) where the image is displayed as dots on the screen, with stronger signals being represented by brighter dots, is the commonest imaging mode in mare reproductive examinations. Other imaging modes are:
A-mode: this is amplitude mode where the returning ultrasound echo is displayed as a spike on a graph. M-mode: this is time-motion mode where the image is displayed in high resolution moving over time and is commonly used in assessment of fetal heart rate. D-mode: this is Doppler ultrasonography which relies on detecting Doppler-shift frequencies reflected from red cells moving either away or towards the ultrasound transducer.
Doppler ultrasonography in equine reproduction Doppler imaging allows the study of blood flow and complements the information from conventional B-mode scanning. Doppler ultrasonography is an emerging technology in the field of equine reproduction and has been excellently reviewed by Ginther and Utt4 and more recently an entire textbook has been devoted to the subject.5 Astutely, these authors recognized that ‘Doppler technology has the potential for providing information on the status and future success of a structure.’ This would appear to be confirmed by the literature published to date. For example, Gastal et al.,6 Silva et al.,7 and Palmer et al.8 all concluded that the measurement of blood flow parameters in the follicular wall using color Doppler techniques was a promising tool to predict ovulation. Blood flow is color coded when using the technique of color Doppler ultrasonography with pixels being assigned color based on their mean velocity, thereby displaying the direction of blood flow. Red usually indicates blood flow towards the transducer whereas blue indicates blood flow away from the transducer (Figure 203.8). Color-Doppler ultrasonography has also been used to study the early development of the equine pregnancy.9 This study discovered an early vascular indicator of the future position of the embryo proper.
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Ultrasonography
Figure 203.8 Color Doppler: red flow towards transducer, blue flow away from transducer. (Courtesy of BCF Technology Ltd, Livingstone, Scotland, UK). (See color plate 62).
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Figure 203.9 Note column of relatively echoic appearance immediately below the follicle and column of relatively anechoic appearance at the edge of the follicle.
Acoustic enhancement artifacts As such work continues, early indicators of impending embryonic death may become apparent.
Image storage Ultrasound machines used for mare reproductive examinations should be capable of readily storing images and video clips in a digital format which can be easily transferred to a computer for referral and data management. The ability to store video clips directly from the ultrasound machine for later transfer is a very useful feature and should be given strong consideration when purchasing a machine.
Artifacts There are many artifacts, or distortions, that can occur during routine mare reproductive examinations. Ultrasound waves are like any other sound waves and can be reflected, refracted (bent), scattered, bounced back and forth several times (known as reverberation), become attenuated, or even completely blocked. Artifacts and their production are a massive subject, comprising a whole chapter in one of the standard texts on equine ultrasonography.10 It is important not to mistake these artifacts for normal or pathological changes. Artifacts can be caused by electromagnetic interference and something as simple as turning off the equipment determined to be causing the interference can eliminate the problem. The most important artifacts in mare reproductive examinations are caused by fluid-filled structures such as follicles, embryonic vesicles, and uterine cysts.
When ultrasound travels through a fluid-filled structure such as a follicle, there is very little attenuation of the beam and echoes are enhanced beneath the follicle compared with adjacent tissue (Figure 203.9). Therefore, a brighter echo exists beneath the fluidfilled structure, in this case a follicle, when compared with echoes of corresponding depth beneath adjacent tissues. Refraction artifacts These artifacts are generated when the ultrasound beam hits a curved edge of a structure, meaning it does not return to the transducer, which diminishes the returning ultrasound beam’s intensity. There is a shadowing or lack of echo formation beyond the site of refraction. When follicles are imaged, refraction artifacts can be frequently seen as columns of relatively anechoic (black) appearance at the edge of the follicle (Figure 203.9). Specular reflections When an ultrasonic beam hits the upper and lower surface of a perfectly spherical, fluid-filled structure, an intensely echogenic reflection is produced on the screen. Specular reflections are typically seen when the early conceptus is imaged (Figure 203.10). It is important to note that, if a perfectly spherical cyst was imaged, then specular reflections would be seen (Figure 203.11). At one stage specular reflections were thought to be diagnostic of the early pregnancy, but this is now known not to be the case. Specular reflections are an artifact formed when any perfectly spherical fluid-filled structure is imaged.
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Figure 203.10 14-Day conceptus showing specular reflections.
The ideal ultrasound machine In the author’s opinion, the ideal ultrasound machine for mare reproductive examinations will possess the following features.
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Digital: crisp, sharp imaging with dynamic focusing; High density transducer; Small and portable; Battery powered; Very high frame rates; Easy image and video clip storage and transfer.
Procedure To carry out ultrasonographic examinations safely, mares should be suitably restrained. Ideally, one should have a set of stocks approximately 75 cm (30 inches) wide and just longer than an average mare. This is adequate for most animals and will even accommodate large draft mares. In a few cases,
Figure 203.11 Perfectly spherical uterine cyst with specular reflections.
a twitch may be required to provide additional restraint. Foals should be restrained in front of, or to the side of, the mare. Tying the tail to one side keeps it out of the way, and prevents hairs entering the rectum. Precautions necessary for rectal examinations also apply to ultrasound examinations and rectal examination should always precede the ultrasound examination. The scanner should be as close to eye level as practicable and the control panel of the machine within easy reach of the operator. The scanner can be placed either side of the mare. Where the operator’s left hand holds the transducer, the scanner is placed obliquely to the right side of the mare’s hindquarters, allowing the right hand to make any notes or adjustments to the controls. To facilitate correct orientation of the transducer, a groove for the finger of the operator is usually located on the transducer, on the opposite side to the working face. The fingers should always be in front of the transducer as it is being introduced and later manipulated rather than pushing the transducer on ahead. For reasons of hygiene, it may be desirable to have the transducer in a plastic sleeve. Coupling gel should be used to exclude air from between the transducer and its protective cover. Using copious amounts of lubricant, which also acts as a coupling medium to ensure good contact and prevent air interference, the transducer and hand are gently inserted into the rectum. Both air and gas are poor propagators of ultrasonic signals and cause marked attenuation of the beam. Close contact of the transducer with the tissue to be examined is, therefore, essential. The gel provides a medium for the ultrasound wave to travel into the tissue by removing the air gap. Should the mare strain, the examination should be stopped and one should wait for the rectum to relax. However, straining is usually not a significant problem. It is best to examine the reproductive tract systematically and to scan the entire uterus and both ovaries at least twice. The transducer is usually held within the rectum in the longitudinal plane. Since the uterus of the mare is ‘T-shaped’, the uterine body appears as a rectangular image in the longitudinal plane. When scanning the uterine body, it is important to move the transducer forwards and backwards and from side to side so that no feature is missed. It is important to move the transducer slowly at all times. To image the uterine horns and ovaries the transducer should be rotated slowly to the right and then the left side. Therefore, the uterine horns appear as circular images in cross-section. If difficulties are encountered with finding a structure, the transducer can be withdrawn a short distance and the structure located by palpation. Ultrasonographic examination can then be resumed.
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Ultrasonography
References 1. Reef VB. Physics and instrumentation. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia, London: Saunders, 1998; pp. 1–23. 2. Powis RL. Ultrasound science for the veterinarian. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore, London: Williams & Wilkins, 1998; pp. 1–18. 3. Kremkau FW Diagnostic Ultrasound: Principles and Instruments, 4th edn. Philadelphia: Saunders, 1993. 4. Ginther OJ, Utt MD. Doppler ultrasound in equine reproduction: principles, techniques, and potential. J Eq Vet Sci 2004; 24:516–26. 5. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Color Doppler Ultrasonography. Book 4. Wisconsin: Equiservices Publishing, 2008. 6. Gastal EL, Gastal MO, Ginther OJ. Relationships of changes in B-mode echotexture and colour-Doppler signals in
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the wall of the preovulatory follicle to changes in systemic oestradiol concentrations and the effects of human chorionic gonadotrophin in mares. Reproduction 2006; 131:699–709. Silva LA, Gastal EL, Gastal MO, Beg MA, Ginther OJ. Relationship between vascularity of the preovulatory follicle and establishment of pregnancy in mares. Anim Reprod 2006; 3:339–46. Palmer E, Chavatte-Palmer P, Verdonck E. Field trial of Doppler ultrasonography of the preovulatory follicle in the mare. Anim Reprod Sci 2006; 94:182–5. Silva LA, Ginther OJ. An early endometrial vascular indicator of completed orientation of the embryo and the role of dorsal endometrial encroachment in mares. Biol Reprod 2006; 74:337–43. Reef VB. Artifacts. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia, London: Saunders, 1998; pp. 24– 38.
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Chapter 204
Uterine Cytology Michelle M. LeBlanc
Introduction Endometrial cytology has become a routine tool for diagnosing endometrial inflammation in equine reproduction since it was first described in the 1960s.1 When used in conjunction with a thorough reproductive examination, mares with active endometritis can be identified quickly and decisions regarding reproductive management can be made without having to wait for culture results. Recent work indicates that not all uterine pathogens are associated with an inflammatory cytological specimen so it is imperative that laboratory findings are correlated with clinical findings before instituting treatment.2–5 Various collection, staining, and interpretation techniques have been described. Advantages and disadvantages of each are discussed below.
Collection of cytology specimen Cytological specimens have been collected from a gloved finger, a uterine culture swab or cap attached to the end of a double-guarded culture instrument, a modified human cytology brush, uterine biopsy specimens or uterine lavage fluids.3,6–13 Whatever method is chosen, a sufficient number of uterine cells need to be collected without contaminating the specimen with vaginal secretions. A culture swab, cap of a guarded uterine culture instrument or cytology brush is most commonly used for collecting samples. Types of swabs used include dry or moistened, sterile or non-sterile, cotton or calcium alginate. Calcium alginate swabs are preferred over cotton swabs as cotton fibers can shed on the slide.14 Advantages of swabs, caps and brushes include commercially available instrumentation and rapid results that are fairly accurate. The major disadvantage is that only a small surface area of the uterine body is sampled and focal infections may be missed. A small volume uterine flush (50 to 100 mL) evaluates a larger en-
dometrial surface area; however, it is cumbersome as the sample requires extra processing.3, 8 The recovered uterine fluid needs to be centrifuged or the collection tube stood upright for a few hours for particulate matter to settle. Furthermore, cell numbers are diluted (may affect results if sample not centrifuged), and debris in a centrifuged sample may interfere with neutrophil identification. For these reasons, the uterine flush technique is usually reserved for mares with chronic infertility. Cytological samples are most commonly collected at the beginning to middle of estrus.11, 14 Uterine defenses are believed to be maximal at that time, so if bacteria or neutrophils are present, a significant problem is likely to exist. False-negatives can occur if cultures are taken in diestrus or during a prolonged transitional estrus.15 It is also possible to induce an iatrogenic infection if cytological specimens or cultures are obtained during diestrus and the luteal phase is not shortened with prostaglandin. To obtain a sample, the veterinarian should wear a sterile, shoulder-length plastic sleeve and place a small amount of sterile non-bacteriostatic obstetric lubricant on the back of the gloved hand.14 The instrument used for cytology needs to be guarded by the gloved hand as it is guided through the vulva and vagina to avoid contamination. The external os of the cervix is identified and the index finger is used to gently find the opening. Both the index finger and the guarded culture instrument are advanced into the body of the uterus. Once inside the uterus, the inner plastic rod (when using a guarded swab or cytology brush) is advanced and the tip rubbed or rolled along the surface of the endometrium for a minimum of 15 seconds. If the cap of a Kalayjian swab is used to collect the specimen, it is either simultaneously rotated as the culture is being obtained or after the culture swab has been withdrawn into the outer sheath. Some veterinarians prefer to pass a culture/cytology instrument through a sterile
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Staining of cytological specimens Cytology specimens obtained off a swab, cytology brush, or cap of a Kalayjian culture instrument should be rolled immediately onto a clean glass microscope slide. It is best to make the cytological smear immediately to prevent cells drying on the swab or brush. Slides can be either air-dried or fixed with ethanol or other suitable cytology fixative. Air-dried slides have less cell loss than fixed slides; however, cellular detail is sacrificed. Samples that are heavily contaminated by mucus when air-dried stain darkly and the mucus may mask cell identification. Wet fixation tends to float mucoid secretions off the slide, thereby preserving cellular detail. Uterine biopsy specimens can be smeared on a microscope glass slide, usually after the tissue is smeared onto blood agar for bacterial culture, or the cytology sample can be obtained by rolling a swab over the tissue specimen and then onto a microscope slide.4 Small volume uterine flush fluids can be centrifuged and the pellet sampled or fluids can be allowed to settle and the cytological specimen obtained by placing a swab in the bottom of the tube. The swab
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is then rolled onto a glass microscope slide. Slides can be stained with modified Wrights stain, Diff-Quik (American Scientific Products, McGaw Park, IL) or R -system (Merck, Glostrup, with the Haemacolour Denmark). Slides can also be stained with Gram’s stain or new methylene blue stain for demonstrating bacteria or fungi.
Evaluation of cytological smear The primary clinical use of cytological smears is to determine the presence or absence of an active inflammatory response in the uterus. Cytology smears should be assessed for total cellularity, cell morphology, cell type, and background content.3, 11, 17, 18 Specimens should first be evaluated at low magnification (100×) under bright-field microscopy and then at high dry magnification (400×). Cellularity is assessed by counting all cells in 5 to 10 high-power fields (400×) and averaging the results or by qualitatively assessing cellularity and categorizing the cellularity as hypocellular, mild, moderate or heavy. Epithelial cell architecture should be evaluated and classified as intact, distorted, fragmented or degenerate. Background can be graded as clear, containing red blood cells, mucus or debris. The amount of debris should be recorded as light (< 25% of slide taking up stain), moderate (25–75% of slide taking up stain) or heavy (> 75% of slide taking up stain). Moderate to heavy debris in cytological smears obtained from small volume flush is correlated with isolation of bacteria from the sample.3 The presence of rods, cocci, hyphae, spores, or urine crystals should be noted and are best evaluated under oil immersion.
Cell types, mucus and debris Epithelial cells are the most commonly seen cell in normal uterine cytological smears. They range from cubiodal in anestrus to tall columnar during the estrous cycle.18 Most epithelial cells are not ciliated; however, ciliated epithelial cells can be seen in diestrus, during the autumn and in chronic infection (Figure 204.1). Collecting cytological smears with a gloved finger or swab may distort epithelial cells, as the rolling of the swab or finger on the slide elongates the cells. Squamous epithelial cells are rare and usually represent cervical contamination. They may also be seen in postpartum smears and in mares refluxing urine in the uterus.19–21 Degenerate epithelial cells are frequently seen in poorly fixed or air-dried smears and in chronic infections with heavy exudate. Neutrophils are the predominate inflammatory cell in mares with endometritis (Figures 204.2 and 204.3). They are rarely present in reproductively healthy mares when smears are obtained during
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Figure 204.3 Severe inflammation. Cytology smear obtained from mare with Streptococcus equi subsp. zooepidemicus endometritis. (See color plate 65).
Figure 204.1 Ciliated epithelial cell and moderate debris collected from small volume uterine lavage fluids. (See color plate 63).
estrus and before breeding. Neutrophils can be found in smears taken in the two to three days after breeding,16 after uterine lavage, and during the early postpartum period.20 Neutrophils may appear well preserved in cellular architecture or may vary from well-preserved to degenerate. Nuclear hypersegmentation is a frequent observation in mares with an active inflammatory responses (Figures 204.2 and 204.3).
Figure 204.2 Epithelial cells, neutrophils, and mucus from a cytological smear collected off the cap of a Kalajyian guarded culture instrument. Many neutrophils are hypersegmented. (See color plate 64).
Lymphocytes, eosinophils, and macrophages are not commonly seen. Lymphocytes are small, relatively dark cells with scant cytoplasm, and may be present in chronic, longstanding infection or in mares that reflux urine into their uterus. Eosinophils stain pink and contain large, round intracytoplasmic granules and have a lobulated nucleus (Figure 204.4). They can be seen in mares with chronic yeast infections, severe pneumovagina, or vestibulo-vaginal
Figure 204.4 Eosinophil and hypersegmented neutrophils recovered from small volume uterine flush. Mare was refluxing urine in her uterus. (See color plate 66).
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Figure 204.7 Heavy debris. Cytological smear was made from centrifuged uterine lavage fluids. An E. coli was isolated. (See color plate 69).
Figure 204.5 Macrophage collected from small volume uterine lavage fluids. Mare had chronic Pseudomonas endometritis. (See color plate 67).
reflux.21 Eosinophils may also be seen in mares infused with an irritating compound or in rare cases of allergies to ampicillin or penicillin. Macrophages are large cells with foamy gray to light blue cytoplasm. Vacuoles and bacteria may be present in the cytoplasm (Figure 204.5). They are commonly seen in smears taken in the first few days after foaling and in rare cases of chronic severe endometritis. Red blood cells occasionally occur in post-foaling smears, in severe acute endometritis or as a result of rough sampling technique. Calcium carbonate crystals may be seen occasionally in mares that reflux urine17 (Figure 204.6). Mucus is usually present on cytological smears as relatively thin, streaks that stain light blue.14 Mucus
Figure 204.6 Urine crystals collected from small volume flush fluids of chronically infertile mare. No urine was observed in vagina. (See color plate 68).
becomes thick, heavy, granular and dark staining in chronically barren mares.3 The mucus may be infiltrated with debris or clumps of degenerated cellular material (Figure 204.7). Bacteria, rods, and cocci can be visualized in smears stained with Diff Quik stain under oil immersion. They are readily visible if smears are stained with Gram’s stain. Fungal elements, including branching hyphae, may be the only indication of a fungal infection as some fungi are difficult to grow under normal laboratory conditions (Figure 204.8). Hyphae may be missed in smears with heavy, thick debris and may be confused with lubricant. Yeast spores may vary in size, shape, and color (Figures 204.9 and 204.10). They are usually blue, black or dark brown with a hint of gold coloring in the capsule.
Figure 204.8 Severe neutrophilic inflammatory response with hyphae. An Aspergillus was isolated. (See color plate 70).
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Figure 204.9 Yeast spores (dark purple, round structures), epithelial cells and mucus from mare with a Mucor infection, a mold that can cause endometritis. (See color plate 71).
Interpretation Endometrial inflammation has been quantified by a number of methodologies. A commonly used technique is to count the number of neutrophils in 10 high power fields (400×) and express the results as a percentage of neutrophils to epithelial cells. Many laboratories use a ratio of 1 neutrophil to 40 epithelial cells (approximately 2%) as an indicator of active inflammation. However, the percentage of neutrophils that constitute active inflammation varies between authors, with 0.5% to 5% neutrophils reported as a positive indicator of inflammation.8, 19,22–24 These studies obtained their results using swabs, the cap off a guarded culture instrument, or uterine flush fluids. If samples are collected by cytology brush, a neutrophil to epithelial cells of 1:20 should be used as an indicator of acute inflammation as cytology brushes yielded significantly more epithelial cells
Figure 204.10 Yeast spores, neutrophils, and mucus. Note the different sizes and uptake of stain as compared to Figure 252.9. (See color plate 72).
per high-power field (hpf) than cotton swabs (44.5 ± 11.2 versus 21.0 ± 12.4, respectively); however, the number of neutrophils per high-power field did not differ.9 Bourke et al also reported that cotton swabs produced significantly more distorted epithelial cells and clumping of epithelial cells than brushes while cytology brush samples tended to have more proteinaceous material and red blood cells, possibly due to the brush collecting more material or to trauma from the bristles. A second method of interpreting cytological smears is to count the number of neutrophils in 10 high-power fields (hpf) and categorize the degree of inflammation.1, 7, 12, 25 Mares with 0–2 neutrophils/ hpf are classified as normal; 3 to 5 neutrophils are classified as having moderate inflammation while mares with > 5 neutrophils/hpf are classified as having severe inflammation. This method of interpretation was evaluated retrospectively in a large Thoroughbred practice in central Kentucky and relationships between 2123 paired uterine cytology, culture and pregnancy rates were compared.12 Endometrial cytology identified twice as many mares with acute endometritis than did endometrial culture. Approximately 20% (423/2123) of cytology samples were positive for inflammation (> 2 neutrophils), whereas approximately 11% (n = 231) of cultures had bacteria recovered. This is in agreement with Wingfield-Digby and Ricketts who showed that 91% of mares with clinical evidence of persistent endometritis had cytological evidence of acute endometritis but only 45% had significant bacteriological findings.13 The degree of inflammation in cytological specimens adversely affected pregnancy rates in the Kentucky study. Mares with positive cytology (2 or more neutrophils/hpf) or bacteria isolated on culture had lower pregnancy rates than mares with no bacteria isolated or negative cytological smears. Lowest pregnancy rates were recorded for mares with severe endometrial inflammation (21%, vs moderate inflammation 48%, vs normal findings 60%). Isolation of bacteria was associated with reduced pregnancy rates (36%), even in the apparent absence of inflammation. Unexpectedly, recovery of a pathogen was not always associated with an inflammatory cytology.12 Streptococcus equi subsp. zooepidemicus was associated with more positive cytologies (65%) than E. coli (55%) or Pseudomonas (52%). Correlation between inflammatory endometrial cytology smears and recovery of pathogenic bacteria in early reports ranged from 76% to –88%.13, 25, 26 However, these investigators did not separate data by organism recovered. Different organisms likely induce different uterine inflammatory responses. Ricketts isolated anaerobes from
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Figure 204.11 Epithelial cells and moderately heavy mucus from a cytological smear made from centrifuged uterine lavage fluids. (See color plate 73).
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84 uterine swabs obtained from 263 Thoroughbred mares; however, only 12% (10/84) were associated with cytological evidence of acute endometritis.23 Recent work shows that isolation of E. coli from culture swabs or endometrial biopsies was less likely to be associated with a positive cytology than when other bacterial species were recovered.2, 5, 12 E. coli isolated from small volume flushes obtained from barren mares was associated with moderate to heavy debris in cytological smears (Figure 204.11), while Streptococcus zooepidemicus was not associated with heavy debris. These studies emphasize the importance of basing a diagnosis of endometritis on clinical findings and not solely on diagnostic results.
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Summary Examination of endometrial smears is relatively easy to perform and produces rapid results. Decisions on whether or not to cover a mare or treat for subclinical endometritis can be made quickly without having to wait for culture results. Cytological smears are more accurate than uterine culture in identifying mares with acute endometritis as almost twice as many mares exhibited inflammation on cytological smears than had bacteria isolated from their uterus. Not all uterine pathogens elicit a strong inflammatory response so results should always be taken in context with clinical findings.
References 1. Knudsen O. Endometrial cytology as a diagnostic aid in mares. Cornell Vet 1964;54:414–22. 2. Bindslev MM, Villumsen MH, Petersen MM, Bogh IB, Bojesen AM. Genetic diversity of S. equi ssp. zooepidemicus
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and E. coli isolated from the reproductive tract of the mare. Reprod Domest Anim 2008;43:110–11. LeBlanc MM, Magsig J, Stromberg AJ. Use of a low-volume uterine flush for diagnosing endometritis in chronically infertile mares. Theriogenology 2007;68:403–12. Nielsen JM. Endometritis in the mare: a diagnostic study comparing cultures from swab and biopsy. Theriogenology 2005;64:510–18. Nielsen JM, Troedsson MH, Zent WW. Results of bacteriological and cytological examinations of the endometrium of mares in a practice in Denmark and in central Kentucky. In: LeBlanc MM (ed.) The Havemeyer Foundation: The Chronically Infertile Mare. Workshop held at Hilton Head Island, SC, 5–8 November 2008; p. 34. Aguilar J, Hanks M, Shaw DJ, Else R, Watson E. Importance of using guarded techniques for the preparation of endometrial cytology smears in mares. Theriogenology 2006;66:423–30. Asbury AC. Bacterial endometritis. In: Robinson NE (ed.) Current Therapy in Equine Medicine. Philadelphia: Saunders, 1983; pp. 410–14. Ball BA, Shin SJ, Patten WH, Lein DH, Woods GL. Use of a low-volume uterine flush for microbiologic and cytologic examination of the mare’s endometrium. Theriogenology 1988;29:1269–83. Bourke M, Mills JN, Barnes AL. Collection of endometrial cells in the mare. Aust Vet J 1997;75:755–8. Brook D. Cytological and bacteriological examination of the mare’s endometrium. Equine Vet Sci 1985;5:16–22. Brook D. Uterine cytology. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 246–53. Riddle WT, LeBlanc MM, Stromberg AJ. Relationships between uterine culture, cytology and pregnancy rates in a Thoroughbred practice. Theriogenology 2007;68:395– 402. Wingfield-Digby NJ, Ricketts SW. Results of concurrent bacteriological and cytological examinations of the endometrium of mares in routine stud farm practice. J Reprod Fertil Suppl 1982;32:181–5. Ley WB, Digrassie WA, Holyoak GR, Slusher SH. Endometrium. In: Cowell RL, Tyler RD (eds) Diagnostic Cytology and Hematology of the Horse, 2nd edn. St Louis, MO: Mosby, 2002; pp. 180–6. Liu IK. Uterine defense mechanisms in the mare. Vet Clin N Am Equine Pract 1988;4:221–8. Card C. Post-breeding inflammation and endometrial cytology in mares. Theriogenology 2005;64:580–8. Freeman KP, Roszel JF, Slusher SH. Equine endometrial cytologic smear patterns. Comp Cont Educ Pract Vet 1986;8:349–60. Roszel JF, Freeman KP. Equine endometrial cytology. Vet Clin N Am Large Anim Pract 1988;247–62. Couto MS, Hughes JP. Technique and interpretation of cervical and endometrial cytology in the mare. Equine Vet Sci 1984;4:265–73. Saltiel A, Gutierrez A, de Buen-Llado N, Sosa C. Cervicoendometrial cytology and physiological aspects of the postpartum mare. J Reprod Fertil Suppl 1987;35:305–9. Slusher SH, Freeman KP, Roszel JF. Eosinophils in equine uterine cytology and histology specimens. J Am Vet Med Assoc 1984;184:665–70. LaCour A, Sprinkle TA. Relationship of endometrial cytology and fertility in the broodmare. Equine Practice 1985;7:27–36.
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23. Ricketts SW, Mackintosh ME. Role of anaerobic bacteria in equine endometritis. J Reprod Fertil Suppl 1987;35: 343–51. 24. Wingfield-Digby NJ. Studies of endometrial cytology in mares. Equine Vet J 1978;10:167–70.
25. Brook D. Uterine culture in mares. Mod Vet Pract 1984; 65:A3–8. 26. Reiswig JD, Threlfall WR, Rosol TJ. A comparison of endometrial biopsy, culture and cytology during oestrus and dioestrus in the horse. Equine Vet J 1993;25:240–1.
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Chapter 205
Endometrial Biopsy Charles C. Love
Introduction Histological evaluation of the mare endometrium has become a routine and essential aspect of a mare’s breeding soundness evaluation. Historically, histological descriptions of the mare endometrium have been made1 with Seaborn (1925)2 one of the first to describe histological changes in abattoir-derived uteri, but these were not used for diagnostic or prognostic purposes. Currently, it is the only test that evaluates the structural integrity of the lining of the uterus (endometrium) with respect to inflammatory infiltrates, fibrotic changes, and dilatation of both endometrial glands and lymphatic vessels. In addition, seasonal and cyclic changes consistent with the normal endometrium can also be evaluated. Numerous reviews and studies3–5 have described the relationship between fertility and specific pathological changes in the mare endometrium. A recent article by Schlafer6 reviewed the literature since the 1980s including studies related to endometrial function, measures to quantitate endometrial structure and function, as well as pathology. This chapter will review the basic aspects of collection, evaluation, and interpretation of the mare endometrial biopsy. In addition, eight cases are presented that highlight pathological changes, categorical interpretation, and suggested therapies.
Contraindication The primary contraindication is removal of endometrial tissue in a pregnant or recently bred mare due to the disruption of the fetal membranes as well as the release of prostaglandin and blood into the uterine lumen. Therefore, the clinician should thoroughly determine the recent breeding history as well as examine the mare’s uterus per rectum using manual and ultrasonographic techniques to assure that the biopsy will be taken from a non-pregnant uterus. The recent (<10–14 days prior to collection) breed-
ing history is particularly important because this is the time period, if the mare was bred, when pregnancy would be missed with current diagnostic capabilities. Therefore, the clinician must determine when breeding took place and especially when ovulation occurred as well as when the mare went out of estrus. A mare should never be assumed to be non-pregnant prior to biopsy removal. Indications for collection of the biopsy have been described (Chapter 272) and are summarized below.
Indications
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Mares with suspected abnormalities of the endometrium, such as aged mares, or mares with a history of reproductive complications. Pre-purchase breeding soundness evaluation, particularly in older mares with unknown fertility histories or a history of recent barrenness in spite of adequate management and being bred to fertile stallions. Non-pregnant mares that are anestrous during the normal physiological breeding season (April 15 and August 15 in the northern hemisphere), suggesting physiological asynchrony along the hypothalamic–pituitary–gonadal axis. Older mares that are to undergo reproductive surgery (cervical repair, urine pooling, or endometrial cyst removal). It is recommended that a biopsy be evaluated prior to surgery to determine if the quality of the endometrium is sufficient such that the mare’s chance of carrying a foal to term will be improved by the surgery alone. If the endometrium has significant pathology the mare’s chances of carrying a foal to term may not be enhanced by the surgery. The owner should be informed of the prognosis prior to surgery.
The histological anatomy has been previously described4 and consists of luminal epithelium plus
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Cilia Luminal epithelium
Duct
Stratum compactum
Vein Mid gland Stratum spongiosum
Lamina Endometrium propria
Basal gland Artery Inner circular muscle Vascular layer
Myometrium
Outer longitudinal muscle Perimetrium
Figure 205.1 Cross-section of the mare uterus, with the endometrium enhanced to show detail, highlighting those stuctures evaluated. Reprinted from Kenney, 1987, with permission.4
lamina propria stratum compactum (SC) and stratum spongiosum (SS); the lamina propria consists primarily of endometrial glands and vessels (arterioles, venules, lymphatics) all connected by a collagen matrix (Figure 205.1). The glands themselves open into the luminal surface and extend and branch deep into the stratum spongiosum. Beneath the endometrium lies the myometrium which is seldom seen on biopsy.
Biopsy acquisition An endometrial biopsy should only be removed following confirmation of the non-pregnant status of the mare. An appropriate-sized piece of endometrium should be removed so proper tissue evaluation can be performed. This requires a biopsy instrument with a large basket size (20 × 4× 3 mm) that is sharp to minimize tearing and tissue distortion. The specimen can be removed from any location in the uterine body, but should focus on areas of gross pathology that may have been identified following manual and ultrasonographic evaluation. The most convenient location to remove a biopsy is from the dorsal uterine body or base of one horn, but samples can be removed ventrally by cupping the
hand around the uterine body and pushing the endometrium up into the biopsy basket. The biopsy instrument is passed manually, with the jaws in the closed position, through the vestibule, vagina, and cervix into the uterine lumen. Once the basket of the instrument is in the uterine lumen the hand is removed from the reproductive tract and passed per rectum over the site where the jaws are located. The punch is rotated so the jaws are positioned horizontally and the handle released, opening the jaws. The endometrium is pressed firmly into the jaws, closed, and the tissue excised and removed (which is facilitated by a quick tug on the instrument). The tissue should be immediately and gently placed into fixative for subsequent evaluation. The fixative of choice is Bouin solution, a combination of formalin and picric acid. It should be noted that this fixative has come under scrutiny as a potentially hazardous material and therefore should be handled with caution. Nevertheless as a fixative it is superior to formalin because it firms the tissue prior to cutting, thereby minimizing artifactual changes that commonly occur with a softer fixative such as formalin. After 12–24 hours of fixation, the tissue is removed and placed in 70% ethanol to prevent
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Endometrial Biopsy further hardening of the tissue. While one sample is considered representative of the whole surface area (850–1350 cm2 ), multiple samples can be removed if the clinician is interested in specific areas of the uterus. There is minimal bleeding following removal, but clients may see small amounts of blood several days following the procedure. While uncommon, there have been cases where bleeding has filled the entire uterus, possibly due to a blood clotting disorder or laceration of a deeper artery.
Evaluation of the biopsy Stage of the estrus cycle Histological endometrial findings can determine the stage of the mare’s estrous cycle. These findings are important to the clinician when determining synchrony between ovarian, cervical, uterine, and behavioral findings following the examination. It has been reported that during the period of transition to normal cyclicity endometrial changes lag behind ovulation onset. Pathologically some mares
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continue to ovulate in the face of an anestrus-like endometrium during the physiological breeding season. Typically the anestrus epithelium is cuboidal and short and the gland density sparse, while the estrus epithelium is tall and the glands may be spread out due to edema. The diestrus epithelium may also be tall but, because of the loss of edema, gland density may appear increased. Endometrial pathology Categorization of the endometrium is based on the evaluation and quantitation of pathological changes within the endometrium. The endometrial biopsy is a unique diagnostic tool for the clinician evaluating the potential fertility status of the mare because, in contrast to culture and cytology techniques that primarily evaluate superficial inflammatory causes or changes, the biopsy evaluates deeper infiltrates and periglandular fibrosis, gland distension (cystic gland distension), lymphatic stasis, gland density, as well as appropriate and inappropriate seasonal changes (Figure 205.2).7 Epithelial pleomorphism
Pregnancy
Thickened basement membrane Lymphocyte stasis
Margination of P.M.N.s
Lymphatic lacuna Chronic reaction
Periglandular fibrosis
Cystic dilatation
Periglandular fibrosis
Figure 205.2 Cross-section of the mare uterus, highlighting the location and appearance of pathological changes. Reprinted from Kenney, 1977 with permission.7
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Inflammation Inflammatory changes tend to occur more commonly in the SC than the SS portion of the endometrium, suggesting that most antigenic stimuli originate in the uterine lumen. While less common, inflammation does occur in the deeper layers of the endometrium and may be systemic in origin. The location of the inflammation is important diagnostically as it may affect the type of treatment instituted (i.e., luminal for superficial vs systemic for deeper layers). In most mares inflammatory cells, usually lymphocytes, can be identified in small numbers. The clinical question, however, is what degree of inflammation is clinically relevant, warranting therapy. The answer to this question can be quite subjective and may be based on the presenting history of the mare. In addition, it is common for ‘significant’ inflammation to not be associated with the recovery of organisms following culture of endometrial swabbings. This suggests that swabbing techniques are not sensitive enough or, more likely, that these mares are more susceptible to antigenic stimuli that ‘normal’ mares are able to resist. This theory fits well with the concept that some mares are inherently ‘resistant’ and some are ‘susceptible’ to infection. Therefore, barren mares with both a negative cytology and culture are not necessarily ‘clean’ until confirmed by biopsy.
This change is not pathological and must be differentiated from PGF.
Lymphatic lacunae Lymphatic lacunae (LL) originate from dilated lymphatic vessels that normally occupy the lamina propria of the endometrium. Lymphatic lacunae can be differentiated from artifactual edema caused by the biopsy instrument by identifying the endothelial layer lining the lacunae. Lymphatic lacunae can be commonly identified, but are considered clinically significant when they are large and particularly when palpable changes (thickened endometrial folds) can be identified. When lacunae become prominent, grossly they can form cysts which, when large enough, have the potential to obstruct embryonic migration, resulting in early embryonic loss.
Cystic gland distension Cystic gland distension (CGD) may or may not occur in the presence of PGF, but tends to be more severe due to the constriction imposed on the gland. Mild CGD occurs during the anestrus period, probably due to the relative inactivity of the myometrium, resulting in slight fluid accumulation.
Periglandular fibrosis Periglandular fibrosis (PGF) represents the degree of ‘scar tissue’ in the endometrium and is determined by evaluating the degree of ‘nesting’ as well as the number of individual glands that are affected. Fibrotic nests occur when excessive collagen forms around the basal portion of individual glands, contracting them into a bundle or nest. Nest numbers are enumerated at a low magnification with two to four nests considered moderate and five or more nests considered severe resulting in a category III classification. The number of rings that surround individual glands is classified as slight (1–3), moderate (4–10), and severe (>10). Periglandular fibrosis predominates around the glands located in the SS and becomes more severe as individual glands form larger ‘nests’ that include large portions of individual glands. Periglandular fibrosis is considered irreversible with increased severity associated with aged mares. It is not clear what factor associated with aging causes the increase in PGF; however, it should not be assumed that inflammation is the primary cause. Non-pathogenic (non-fibrotic) nesting also occurs when glands aggregate without the fibrotic rings characteristic of PGF and tends to occur during the ‘proestrus period’ as a result of edema accumulation.
Focal and cellular changes
r
r
r
White blood cells: when outside the vessels in sufficient quantities, white blood cells are considered inflammatory, with neutrophils and lymphocytes associated with acute and chronic changes, respectively. Inflammation, to some degree, is the most common change reported. However, not all inflammation is considered clinically relevant (i.e., treatable). Clinical relevance is subjective and tends to be based on other factors, such as a mare’s age, years barren, ultrasonographic identification of uterine fluid, and history of reproductive difficulty. Siderophages: these result from macrophages that consume hemosiderin, which is degraded from hemoglobin. This cell type is commonly associated with recent hemorrhage events in the uterus such as parturition and abortion, but may also result from recent caustic insults to the endometrium. Plasma cells: if present in sufficient numbers, plasma cells suggest chronic antigenic exposure of the endometrium, caused by inadequate perineal conformation leading to aspiration of air and debris.
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Eosinophils: the presence of these cells has been associated with the aspiration of air (Slusher 1984)8 into the uterine lumen, suggesting poor perineal conformation.
Categorization Various methods of endometrial evaluation including categorization4, 5,9–11 have been described. The intent of categorization is to summarize endometrial findings and relate the severity of changes to fertility. Following complete evaluation of the biopsy and reproductive history, a mare is placed into one of four categories based on the pathological findings (or lack thereof). Three categories (I, II, and III) were introduced by Kenney)4 based on the degree of the pathological change in the endometrium. Category I is considered a normal endometrium in which few, if any, changes are present and in which the epicrisis would comment that ‘there are no changes present that should reduce this mare’s ability to carry a foal to term’. Category III has the highest level of pathological change and these mares have a ‘low probably of carrying a foal to term if the endometrium remains in its current state’. Mares in this category tend to be older and have higher levels of PGF, but this is not always the case since widespread inflammation can also be considered category III. Prognostically, these would be two very different mares, with the first unlikely to change due to the irreversible nature of the PGF, but the second has the potential to improve to a categoryI or II if the inflammation resolves. Category II is a broad category including those mares close to category I and those close to category III. Therefore, it was subsequently modified10, 11 to include two categories, category IIA (close to I) and category IIB (close to III). This is a dynamic category since most mares fall into this category and many of these mares will likely benefit from some form of intrauterine or system therapy or modification of their reproductive anatomy (e.g., surgeries to prevent air and fecal material aspiration, torn cervix, urine pooling, or perineal reconstruction).
Fertility and the endometrial biopsy The goal of most diagnostic tests related to reproduction is their relationship to fertility and ideally their ability to ‘predict’ fertility. Kenney clarified this by stating the relationship of biopsy findings to the ‘ability of a mare to carry a foal to term’, not simply whether she was diagnosed pregnant at some point. This clarification is important because the health of the endometrium is critical for the entire gestation
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length. Therefore, it is possible for a mare to be diagnosed pregnant early in gestation only to lose the pregnancy at a later time due to an endometrial deficiency. All of the studies4, 7, 9, 10,12–14 relating biopsy grade to fertility have shown that as biopsy grade increases foaling rate declines. The foaling rates of category I, II, and III mares have ranged from 61–92%, 24–67%, and 0–36%, respectively (reviewed by Doig and Waelchli 1992).15 While there is considerable range within and between these categories, the biopsy grades provide useful information regarding a mare’s potential fertility based on the health status of the endometrium. Possible reasons for variation in foaling rate among studies include: (i) variation in the clinician/pathologist evaluating and interpreting the sample; (ii) variation in the history of the mare recorded (mare age and years barren can be important modifiers in the prognosis); (iii) variation in the mare’s management and in stallion fertility (these have a large impact on whether a fertile mare is ‘barren’). In two separate studies Kenney found considerable variation in his categorization scheme and felt these differences were due primarily to the management of the mares.4 One study, in which management factors were ‘heterogeneous’, had a lower foaling rate for category I and II mares (68% and 51%) compared to the second study in which management was consistent and of high quality (92% and 67%).4 Category III mares were low in both studies (11% and 4.3%), suggesting that the changes associated with this category, in most cases, cannot be compensated for by improved management. Nevertheless, it should be noted that category III mares are not sterile, but the likelihood of carrying a foal to term is low. Two additional studies10, 14 compared foaling rates using the four-category system. The comparison of the two studies found mares in category-I and III to be high fertility and very low fertility groups, respectively; but there was considerable variation when comparing categories IIA and IIB,15 highlighting the fact that category II is a dynamic category and can be influenced by the individual performing the evaluation as well as the recommendations and treatment instituted at the recommendation of the epicrisis. Doig 198110 (Table 205.1), in a large group of mares (700) reclassified category II into IIA and IIB primarily based on the degree of PGF. In addition, he reported that mares with moderate PGF under ‘strict veterinary supervision’, which included artificial insemination with a gentamicin-containing extender, had a higher foaling rate than similar mares bred naturally or without extender. This study also reported that mares with similar biopsy categories, but barren two or more years, had lower foaling
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Table 205.1 Foaling rates (percent foaling) from two studies using similar four-category rating systems Number foaling/number bred (%) Category
Doig et al.∗
Waelchli†
I IIA 11B 111 Overall
47/57 (82) 155/209 (74) 59/129 (46) 0/8 (0) 261/403 (65)
70/100 70) 24/57 (42) 5/27 (18) 0/8 (0) 99/192 (52)
∗
Doig PA, McKnight JD, Miller RB. The use of endometrial biopsy in the infertile mare. Can Vet J 1981;22:72–6. † Waelchli RO. Endometrial biopsy in mares under non-uniform breeding management conditions: prognostic value and relationship with age. Can Vet J 1990;31:379–84.
rates than mares barren only one year, highlighting the importance of a mare’s reproductive history in the diagnosis. These findings also suggest other factors related to foaling rate than simply the degree of PGF in categories IIA and IIB. Waelchli14 (Table 205.1) studied the relationship between mare age and biopsy category. Within biopsy categories I, IIA, and IIB, mares above the median age for that category tended to have lower foaling rates than mares below, suggesting that factors associated with increasing age, irrespective of the pathological changes in the endometrium, affect mare fertility. This is interesting in light of the difficulty that older maiden mares have in becoming pregnant. These mares tend to have category I or IIA endometrium scores, suggesting that factor(s) other than endometrial quality influence the ability of these mares to become pregnant. Epicrisis As introduced by Kenney, the epicrisis ‘is the examiner’s interpretation of all the findings of the case’, which ‘takes into consideration the history and the physical, behavioral, and microscopic changes, as well as microbiologic and endocrinologic findings, and putting them together in meaningful way for obtaining appropriate diagnoses as well as making prognoses of the ability of the mare to carry a foal to term’. The intent is for the clinician to be aware of all aspects of the mare’s history; particularly those related to her reproductive status and not just record the histological findings. Since endometrial biopsies are usually taken by practitioners from mares that have a reproductive deficiency(ies), the practitioner is interested in the pathology of the sample, but more so what to do about it. Specifically, are the endome-
trial changes responsible for the mare’s barrenness; if so are they reversible; and what is the recommended treatment? Therefore, oftentimes at the objection of the pathologist, Kenney promoted the concept that the mare reproductive specialist should be trained in the evaluation and interpretation of endometrial biopsies, and because of their clinical training in all aspects of mare reproduction, are best qualified to write the epicrisis. Recent research findings Since the introduction of the biopsy technique there have been numerous studies using specific markers to study mating-induced endometritis (MIE) to identify those mares susceptible and those that are resistant to inflammation. Lactoferrin, an estrogen regulated glycoprotein with antibacterial and immunogenic properties, is higher (5500 times) during estrus than diestrus and early pregnancy and, in mares with MIE, levels were higher in the proestrus period.16 It may become possible in the future to use biopsy to study the biochemical make-up of the endometrium in addition to the histological characteristics. Aupperle17 identified atypical co-expression of vimentin and desmin and suggested that staining for these intermediate filaments may be useful in measuring endometrial health. In this study 151 mares were evaluated and co-expression of vimentin and desmin was identified in fibrotic glands and mares with ‘pathologically inactive endometria’ as well as older barren mares. Associations were made between the severity of PGF and levels of 11β-hydroxysteroid dehydrogenase (11β-HSD2 ) and transforming growth factor-β 1 (TGF-β 1 ) which, interestingly, also play a role in perivascular fibrosis of the human heart.18 Walter19 found glycoprotein levels were different between endometria with and without degenerative changes and these differences were not estrogen/progesterone dependent. Walter20 also identified calcium-containing structures in the lumen of glands affected by PGF, suggesting these structures may play a role in subsequent cystic gland distension. In an additional study Walter21 determined that matrix metalloproteinase (MMP-2) and tissue transglutaminase (TG-2) play a role in extracellular matrix regulation in areas of periglandular fibrosis. The endometrial biopsy is a useful clinical tool for determining endometrial health. While it provides information about the inflammatory status of the endometrium, additional pathological findings are also evaluated.
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Case 1a Shows a moderately frequent to frequent, diffuse amount of chronic inflammation (lymphocytes) in the stratum compactum. If consistent throughout the sample would be classified as category IIB with potential to upgrade to category I or IIA if the inflammation is resolved. This finding may or may not be associated with a positive culture, but requires intrauterine therapy as well as treatment of any physical reproductive limitations such as poor perineal conformation. (See color plate 74).
Case 1b Shows slight to diffuse amount of chronic inflammation (lymphocytes) in the stratum compactum. Could be considered marginal as to whether treatment is warranted and would depend on distribution and mare reproductive history. Solid arrows identify some but not all lymphocytes. Also notice siderophages (yellow cells – dashed line) which result from a recent hemorrhagic event such as parturition or abortion. (See color plate 74).
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Case 2 Shows one to two layers of periglandular fibrosis (slight) forming nests (solid arrow). Notice slight cystic gland distension (solid arrow) associated with those glands, suggesting that the PGF is causing constriction of the gland. This degree of change is commonly associated with a middle-aged mare. Depending on the frequency of the PGF this mare could be classified as category IIA or IIB. (See color plate 75).
Case 3 Shows large PGF nest with cystic gland distension and inspissation (notice cellular debris, possible degenerated neutrophils), and endometrial atrophy (solid arrow, notice cuboidal endometrium) changes consistent with a category III endometrium. The PGF is irreversible, but intrauterine lavage with warm saline may reduce the degree of CGD and upgrade the category depending on the severity of PGF. Notice flattening of glandular epithelium (dashed arrow), an early sign of degeneration/atrophy due to long-term CGD and PGF, compared to normal epithelium (white arrow). (See color plate 76).
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Case 4 Shows slight widespread CGD not associated with PGF. This change can be associated with anestrous mares, and if identified during the physiological breeding season, suggests inadequate uterine tone and contractility, warranting intrauterine lavage. Lavage may only be a temporary treatment as the endometrium from such mares may have insufficient uterine contractility, resulting in recurrence of the condition. Therefore, it may be important to breed these mares immediately following treatment. (See color plate 77).
Case 5 Shows small gland density associated with seasonal atrophy. Notice cuboidal luminal epithelium (dashed arrow). A similar change occurring during the physiological breeding season is abnormal and suggestive of inadequate hormonal stimulation. (See color plate 78).
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Case 6 Shows lymphatic lacunae (widespread) in the stratum spongiosum. This change can be differentiated from staining artifact and estrual edema, by identifying the endothelial nuclei (solid arrows) that line the lymphatic lacunae. The vacuolation (dashed arrow) is a staining/processing artifact. Lymphatic lacunae become particularly significant if associated with a palpably flabby, poorly toned uterus. Recommended treatment includes warm saline lavage to stimulate uterine contractility. This treatment, however, may not be long-lasting and an immediate attempt should be made to get the mare pregnant, if desired. (See color plate 79).
Case 7 Shows cervical (left) and vaginal (right) samples. The cervix is characterized by an undulating epithelium while the vagina is characterized by a stratified squamous epithelium. Both areas are devoid of glands beneath the epithelium. The cervix should not be confused with complete endometrial glandular atrophy. Cervical and vaginal samples result from incomplete passage of the biopsy instrument through the cervical canal. (See color plate 80).
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Case 8 Shows the ‘string of pearls’ appearance of the glands, a normal finding associated with increased gland tortuosity during diestrus. (See color plate 81).
References 1. van Niekerk CH, Gerneke WH, van Heerden JS. Anatomical and histological observations on the reproductive tract of mares with abnormal oestrous cycles. J S Afr Vet Assoc 1973;44:141–52. 2. Seaborn E. The estrous cycle in the mare and some associated phenomena. Anat Rec 1925;30:277–87. 3. Van Camp SD. Endometrial biopsy of the mare: a review and update. Vet Clin N Am Equine Pract 1988;4:229– 45. 4. Kenney RM. Cyclic and pathologic changes of the mare endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1978;172:241–62. 5. Ricketts SW. The technique and clinical application of endometrial biopsy in the mare. Equine Vet J 1975;7:102– 7.
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6. Schlafer DH. Equine endometrial biopsy: enhancement of clinical value by more extensive histopathology and application of new diagnostic techniques? Theriogenology 2007;68 (3): 413–22. 7. Kenney RM. Clinical aspects of endometrial biopsy in fertility evaluation of the mare. Proc Am Assoc Equine Pract 1977; 105–22. 8. Slusher SH, Freeman KP, Roszel JF. Eosinophils in equine uterine cytology and histology specimens. J Am Vet Med Assoc 1984;184:665–70. 9. Shideler RK, McChesney AE, Morrow GL, Voss JL, Nash JG. Endometrial biopsy. Proc Am Ass Equine Pract 1977; 97– 104. 10. Doig PA, McKnight JD, Miller RB. The use of endometrial biopsy in the infertile mare. Can Vet J 1981;22:72–6. 11. Kenney RM, Doig PA. Equine endometrial biopsy. In: Current Therapy in Theriogenology 2. Philadelphia: Saunders, 1986; pp. 723–9. 12. Asbury AC. Some observations on the relationship of histologic inflammation in the endometrium of mares to fertility. Proc Am Assoc Equine Pract 1982; 401–4. 13. de la Concha-Bermejillo, Kennedy PC. Prognostic value of endometrial biopsy in the mare: a retrospective analysis. J Am Vet Med Assoc 1982;181:680–1. 14. Waelchli RO. Endometrial biopsy in mares under nonuniform breeding management conditions: prognostic value and relationship with age. Can Vet J 1990;31:379– 84. 15. Doig PA, Waelchli RO. Endometrial biopsy. In: McKinnon AO (ed.) Equine Reproduction. Philadelphia: Lea & Febiger, 1992; pp. 225–33. 16. Kolm G, Klein D, Knapp E, Watanabe K, Walter I. Lactoferrin expression in the horse endometrium: relevance in persisting mating-induced endometritis. Vet Immun Immunopath 2006;114:159–67. 17. Aupperle H, Schoon D, Schoon H-A. Physiological and pathological expression of intermediate filaments in the equine endometrium. Res Vet Sci 2004;76:249– 55. 18. Ganjam VK, Evans TJ. Equine endometrial fibrosis correlates with 11β-HSD2 , TGF- β 1 and ACE activities. Mole Cell Endocrinol 2006;248:104–8. 19. Walter I, Klein M, Handler J, Aurich JE, Reifinger M, Aurich C. Lectin binding patterns of uterine glands in mares with chronic endometrial degeneration. Am J Vet Res 2001;62:840–5. 20. Walter I, Helmreich M, Aurich C. Mineralised deposits in the uterine glands of mares with chronic endometrial degeneration. Vet Rec 2003;153:708–10. 21. Walter I, Handler J, Miller I, Aurich C. Matrix metalloproteinase (MMP-2) and tissue transglutaminase (TG-2) are expressed in periglandular fibrosis in horse mares with endometrosis. Histol Histopath 2005;20:1105–13.
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Chapter 206
Endoscopic Examination Claire E. Card
Indications for endoscopic examination of the mare Endoscopic examination of the mare’s reproductive tract is used as a diagnostic aid for unusual rectal and uterine conditions, such as intractable infections, tumors, abscesses, foreign bodies, adhesions, and congenital abnormalities.1, 2 Endoscopy is also used as an adjunct to a breeding soundness examination or is used as a clinical tool for assisted reproductive procedures such as hysteroscopic insemination.3–5
Routine anatomic features viewed using endoscopy The routine anatomic structures of the mare’s reproductive tract that may be viewed using endoscopy include the vagina, vestibulovaginal fold, hymen in maiden mares, cervix, uterine body, bifurcation, horn, uterotubal papilla and ostium.6, 7 Other common structures in the endometrium include endometrial cysts and endometrial lymphatic cysts. Endoscopy is used to find foreign bodies, determine the health of the endometrium, and evaluate tumors, uterine hematomas, acquired anomalies such as adhesions, and congenital abnormalities.1, 8, 9 In the urogenital system the urethral tubercle, urethra, bladder, and openings of the ureters may also be evaluated endoscopically.10 The interior of the rectum is sometimes examined to evaluate iatrogenic trauma or foaling injuries. Vagina, vestibulovaginal fold, and cervix Endoscopic evaluation of the vagina is indicated when a speculum examination alone provides insufficient information. The cervix is a dynamic organ which changes shape and position based on seasonal factors, mare status (postpartum, barren, maiden, pregnant), and stage of the estrous cycle. Large clear vaginal speculums or a Caslick (three-spoon) specu-
lum are frequently used for vaginal examinations in mares. Endoscopy may be used to evaluate the cervix in the cranial vagina, and the external os protrudes into the vagina (Figure 206.1a). The free edge of the cervical os should not be adherent to the vaginal wall. Anatomically there is a vertically oriented fold of tissue where the two halves of the mullerian system fused. This tissue fold leads to the cervix. The interior of the cervix appears smooth (Figure 206.1b). Hysteroscopy – hysteroscopic anatomy An endoscopic examination of the interior of the uterus is called hysteroscopy. Hysteroscopy is used often as an adjunct technique during a breeding soundness examination, and has specific applications for the diagnosis of certain conditions. The uterus in the mare is Y-shaped with a short body (Figure 206.2a). There are at least a dozen longitudinal folds of endometrial mucosa that form ridges which run from the body into the uterine horns. The region of the uterus where the two horns diverge is called the bifurcation (Figure 206.2b). The uterine horns are 15–20 cm in length and 4–7.5 cm in width. The longitudinal folds lead to the end of the uterine horn. The uterotubal papilla is located dorsally and eccentrically at the end of the horn.6–8 (see Figures 206.2c and 206.2d). Hysteroscopic procedure To prepare a mare for a hysteroscopic examination empty the rectum of manure, wrap the tail, and cleanse the perineum. Sedate the mare using alpha agonists, combined with butorphanol to minimize movement and any discomfort. Position the mare so that following sedation her airway is unobstructed and she is positioned near the back of the stocks. The endoscopic equipment should be gas or chemically sterilized and the disinfectant removed using
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 206.1 (a) View of the external cervical os in early estrus. (b) Smooth interior of the cervical canal.
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Figure 206.2 (a) Hysteroscopic view of the uterine body. Dark areas represent bruising in the area of the internal cervical os. (b) Hysteroscopic view of the bifurcation of the uterine horns. (c) Hysteroscopic view of the uterotubal papilla visible (arrow) as a small mound at the end of the uterine horn. (d) Close-up hysteroscopic view of the uterotubal papilla indicated by the arrows.
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alcohol and sterile water or saline. Generally two people are required to perform a hysteroscopic examination. An examiner wearing a rectal sleeve and sterile surgical glove passes the equipment through the vagina and cervix into different uterine locations, and an operator directs the tip of the endoscope when required. A one-meter gastroscope or a videoendoscope is used. The examiner shields the end of the endoscope in their gloved and lubricated hand and carries it forward to the cervix. The cervical canal is dilated by the examiner’s finger and the endoscope is passed so the tip just enters the uterine body. The cervix is then held closed around the endoscope and the uterus is insufflated.8, 11 Typically with fully functional endoscopic equipment, complete uterine insufflation should take less than 20 seconds using room air. External insufflation sources may be used. Excessive uterine insufflation results in gas being released through the cervix.12 Full visualization of intrauterine structures has been reported with mean pressures of 17.8–30 mmHg, with excessive pressures causing discomfort and elevated heart rates.12 The insufflation of the uterus is maintained using intermittent air flow during the hysteroscopic examination. The uterine body typically is situated in the pelvis, with the uterine bifurcation located cranioventrally. The cranioventral curve must be followed to atraumatically enter and evaluate the uterine horns. Minimal manipulation of the tip of the endoscope is used to pass the endoscope into the uterine horns. The examiner, using their hand in the mare’s vagina and another on the endoscope, feeds and directs the endoscope into the uterine horns. Once a uterine horn has been entered the directional turning knobs of the endoscope may be locked by the operator and the tip of the scope directed using fine manipulation as needed. Samples such as cultures, fluid aspirates, or guided endometrial biopsies may be obtained through the biopsy port of the endoscope.13 Once the entire contents of the uterus have been viewed, the air flow is turned off and the biopsy port opened to allow the gas to be released from the uterus. Transrectal palpation may be used to encourage the emptying of air. Maintenance protocols for cleaning the equipment following each procedure help keep the endoscope functional, and reduce the potential for mechanical transfer of pathogens between patients. Iatrogenic infection is a risk of the procedure.14, 15
Figure 206.3 Hysteroscopic view of edematous endometrial folds.
In estrus, there will be glistening mucus present on the surface of the endometrium and endometrial edema will be prominent. The folds of the uterus will be visibly edematous (see Figure 206.3). Endometrial glandular and lymphatic cysts Endometrial glandular cysts are small structures usually less than 1 mm in diameter. Endometrial glandular cysts correspond histologically to dilated endometrial glands and contain glandular secretions (see Figure 206.4). Endometrial lymphatic cysts vary in size and number, and may be identified as solitary cysts or clusters. They may be larger than 6 cm in diameter. Endometrial lymphatic cysts may have compartments, or may be found in clusters. They may or may not be spherical like embryos (see Figure 206.5). Structures such as endometrial glandular cysts or endometrial lymphatic cysts by themselves are not clinically significant.16, 17 A few cysts of either type are not associated with fertility problems in mares. These cysts may be present in subfertile and fertile mares.1, 17 Very large endometrial lymphatic cysts are sometimes removed.18, 19 Larger endometrial lymphatic cysts may be confused with embryos during
The healthy endometrium The healthy endometrium of the mare is pale pink and appears to have a homogenous texture.6–8 The uterus should be evaluated for color, texture, the presence of discharge, foreign matter, endometrial cysts, lymphatic cysts, and presence of adhesions.1
Figure 206.4 Hysteroscopic view of a uterine horn. The arrow indicates a small endometrial glandular cyst.
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Figure 206.5 Hysteroscopic view of endometrial lymphatic cysts indicated by the arrows in (a) and (b).
an ultrasound examination, or result in the diagnosis of a twin pregnancy, when a cyst and an embryo are present. Therefore clinically it is important to note the location and size of endometrial cysts in the uterus. Endometrial cysts grow very slowly, may be found year after year, and do not develop heartbeats, unlike embryos. For this reason if there is no previous examination history, a mare should not be diagnosed in foal until an embryo with a heartbeat is identified, or prior to that time a structure ‘compatible with an embryonic vesicle’ should be recorded.
dure13 (see Figure 206.7a). Iatrogenic bleeding at the site of an endometrial biopsy is a normal finding (Figure 206.7b). Uterine biopsies where large strips of tissue come away, when the instrument is dull, or when biopsies are taken from mares with fibrotic endometrial changes are associated with more excessive bleeding. The presence of bleeding after a biopsy procedure is performed should be taken into account when performing a hysteroscopic procedure after a biopsy. Hysteroscopic insemination
Postpartum uterus The process of uterine involution in the mare is rapid, and in an uncomplicated foaling the uterus of the mare will be one and half times its non-pregnant size by 12 hours postpartum. The normal lochia of the mare is low volume, thick, and reddish in color. The postpartum uterus due to its larger size and stretched cervix may be difficult to insufflate (see Figure 206.6). Endometrial biopsy Hysteroscopy may be used as an means to directly visualize the endometrium during a biopsy proce-
A hysteroscopic approach was developed for cannulation of the uterotubal ostium and low-dose insemination. In the normal mare the volume of the uterine tube is 300 µL, and small volumes may be injected into the uterine tube by cannulating the uterotubal ostium. In a mare with patent uterine tubes the injection will occur with some pressure and may be used as a means of evaluating tubal patency; however, the procedure is technically difficult.20 Hysteroscopic insemination is a low-dose insemination technology. A reduced number of sperm and minute insemination volume are used in this technique. The side where the dominant follicle is located is determined. The endoscope is directed up the uterine horn ipsilateral to the dominant follicle. A small catheter is passed through the biopsy channel of the endoscope and advanced so it touches the uterotubal papilla.4, 20, 21 The inseminate is then injected onto the surface of the papilla (Figure 206.8). This technique has been successful in achieving pregnancies with low numbers of fresh, frozen thawed, and flow-sorted sperm.5, 22
Congenital abnormalities
Figure 206.6 Hysteroscopic view of the postpartum uterus.
Congenital abnormalities of the vagina, cervix, and uterus in the mare have reported. Imperforate hymens have been reported, and mucoid material may be present behind the septum. In athletic mares the
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Figure 206.7 (a) Hysteroscopically guided endometrial biopsy. (b) Hysteroscopic view of bleeding from an endometrial biopsy site.
hymen may bulge out through the vulva during exercise. A paramesonephric duct remnant (persistent mullerian) remnant in the mare sometimes forms a longitudinal septum-like division of the vagina. Cervical aplasia has also been reported. A double cervix either communicating or not communicating with a uterus, or two separate uterine horns, has been reported. Uterus unicornis and uterus bicollis have been reported in the mare. These conditions are frequently investigated using hysteroscopy to confirm the nature of the problem. Mares with the 63XO genotype have a small flaccid cervix and uterus. Ectopic ureters that empty into the vagina have also been reported in the mare.
predisposing factor for urovagina is pneumovagina. Pneumovagina is caused by poor perineal conformation and may occur post-foaling due to stretching of the reproductive tract, and the opening of Caslick for foaling in mares with poor perineal conformation. The urine should be removed from the vagina of the mare using a speculum or cotton before a hysteroscopic examination is performed. Persisent urovagina may be due to poor perineal conformation, a dysfunction or injury to the bladder, or urethra, or may be due to an ectopic ureter.10 Vaginal hematomas and abscesses
Pathological conditions of the vagina
Vaginal hematomas occur primarily as a result of an injury during foaling. Hematomas may become secondarily infected. The depth of an abscess may sometimes be determined using an endoscopic approach.
Urovagina
Vaginal lacerations
Urovagina is a condition where urine accumulates in the vagina and causes a chemical inflammation. A
Foaling trauma may cause severe vaginal inflammation, scar formation, abscess formation, hematoma
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Figure 206.8 (a) Polyethylene catheter exiting the biopsy channel near the uterotubal papilla. (b) Hysteroscopic view of the uterotubal papilla following deposition of an inseminate.
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Endoscopic Examination formation, and adhesions. Rectovaginal fistula may be visualized by examining the dorsal wall of the vagina using an endoscope. Vaginal lacerations may occur at the time of breeding. Certain stallions known to cause vaginal injuries to mares may undergo semen collection for artificial insemination, or a breeding roll is used to prevent deep penile penetration. The extent of the vaginal laceration or injury may be explored using endoscopy. Seminoperitoneum has been reported due to a breeding injury.
Vaginal fusion Vaginal fusion is typically diagnosed following severe foaling trauma or a difficult dystocia. Miniature horses, due to their small size, and the subsequent manual difficulty encountered in managing their obstetrical problems, are prone to this foaling complication. In cases of vaginal fusion the mucosa is injured and inflamed resulting in the sides of the vagina literally adhering and fusing together. Urination keeps a small passageway open from the bladder to the vulva. Endoscopy of the narrow channel may be used to assess the extent of the vaginal fusion.
Varicose veins Pregnant mares with a history of intermittent vaginal bleeding may have varicose veins. Varicose veins are tortuous venous structures that may bleed during pregnancy, or in some cases in non-pregnant mares. Varicose veins in the vagina may be hard to find because they are often located dorsally in the region of the vestibulovaginal fold. These veins are more prominent in older mares and may cause significant intermittent bleeding. Vaginal dryness or irritation from pneumovagina is a predisposing factor and may result in vascular fragility leading to subsequent hemorrhage. Endoscopy may be used to help locate the offending bleeding vessels. A Caslick may solve the mare’s problem by eliminating the vaginal dryness after a week or more as the vaginal environment becomes restored to a moister state, or the vessels may need to be ligated, cauterized or injected with a sclerosing solution. Persistent vaginal bleeding has also been associated with uterine tumors in a few mares.23, 24
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Cervical lacerations Cervical lacerations typically occur as a consequence of a foaling injury. Endoscopy may be used to evaluate the extent of the injury. Lacerations that involve only the external cervical os carry a better prognosis than injuries that penetrate the cervical canal. Cervical adhesions Foaling injuries or injuries to the cervix during fetotomy procedures may result in significant cervicitis that results in the formation of transluminal adhesions. Transluminal cervical adhesions are associated with the development of pyometra in mares. Tumors of the cervix Tumors of the cervix are usually leiomyomas or leiomyosarcomas. The tumors arise from smooth muscle cells. These tumors are usually slow-growing and benign. Intravaginal ultrasonographic examination may be used to determine the diameter of the tumor. Smooth muscle tumors are usually more commonly found in older mares.
Non-infectious pathological conditions of the uterus Endometrial lymphatic cysts Endometrial lymphatic cysts may reach an excessively large size and occasionally require removal. Rarely an endometrial lymphatic cyst is deemed to be large enough to interfere with embryonic migration or sperm transport. The endometrial lymphatic cyst may be manually ruptured, or ruptured using a laser through a hysteroscopic approach, and then removed at its base.1, 18, 19, 25 Lymphstasis The endometrium of subfertile mares, with a significant degree of lymphstasis, has a reticular congested pattern1, 11 (see Figure 206.9). Histologically the reticular pattern corresponds to lymphstasis and endometrial gland necrosis. These subfertile mares are prone to intrauterine fluid accumulation during breeding and have a defect in uterine contractility.1
Pathological conditions of the cervix Cervical trauma from lacerations, excessive prolonged pressure, or infections, causing cervicitis, may be evaluated using an endoscope. Cervical masses such as tumors or abscesses may also be visualized.
Aembryonic (trophoblastic) vesicle Rarely an aembryonic vesicle forms where no embryonic body is identified. These pregnancies are self-limiting and are eventually spontaneously
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Mare: Techniques in Reproductive Examination smelling watery uterine discharge is associated with post-foaling endometritis. Normal lochia is thick and low in volume. Chronic endometritis
Figure 206.9 Hysteroscopic view of a subfertile mare with lymphstasis. A reticular pattern is present in the endometrium.
eliminated at around 30 days. The mare may be returned to estrus using prostaglandin.
Infectious pathological conditions of the uterus Acute endometritis Acute uterine inflammation associated with endometritis may be identified in mares using endoscopy. In mares physiological processes such as breeding, or foaling cause only transient, short-lived inflammation. Persistent inflammation of the endometrium is associated with lower fertility and uterine subinvolution in foaling mares.1 Acute endometritis is characterized by the presence of cloudy, free intrauterine fluid that is neutrophil laden (Figure 206.10). A vaginal discharge may also be evident. Uterine discharge is usually caused by bacterial infection, and should be evaluated using cytology and culture. In postpartum mares, high-volume, foul-
Chronic endometritis may cause a change in the texture of the mucosa of the endometrium from a smooth to a rough or pitted appearance. In mares with chronic endometritis there may be ulcers, erosions, adhesions, and/or plaque-like lesions present. The plaque-like lesions may form a nidus for intractable uterine infections. Microbial organisms are believed to colonize the plaque, forming a pathological biofilm. Pathogens in these biofilms may reactivate and cause a subsequent endometritis or placentitis (Figure 206.11). Lymphstasis may also be present in the diseased endometrium. Pyometra Pyometra is a term that describes a persistent condition where the uterus fills with pus. A true pyometra in a mare usually involves a cervical injury, often resulting in adhesion formation, that mechanically interferes with cervical dilation or patency. The endometrium is often diseased as well so that it has lost its ability to secrete prostaglandin, resulting in retained luteal tissue and prolonged diestrous periods. The diseased endometrium may lose its normal smooth appearance (Figure 206.12). The chronic retention of pus in the mare’s uterus may result in mild anemia or no systemic change in the blood count. The fluid may or may not grow bacteria when cultured. Persistent endometrial cups Persistent endometrial cups are a rare finding in mares that fail to establish normal estrus cycles. These mares may have experienced a pregnancy loss or are post-foaling. Levels of the hormone eCG remain high. There is a failure of the endometrial cup tissue to regress. The cup tissue may be visualized using endoscopy. The secretion of eCG is associated with follicular luteinzation rather than ovulation. Persistent anovulatory follicles may be a feature of these mares. Transluminal adhesions
Figure 206.10 Hysteroscopic view of free intrauterine fluid in a cycling mare.
Transluminal uterine adhesions may result from trauma and inflammation following dystocia, hysterotomy, or intrauterine infusion of caustic materials. Transendoscopic electrosurgery and laser has been used to reduce adhesions26, 27 (Figure 206.13).
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(c) Figure 206.11 (a) Plaque (indicated by the arrow) on the endometrium of a mare with a history of chronic intractable fungal endometritis. (b) Ulcers (indicated by the black arrows) on the endometrium of a mare with chronic endometritis. (c) Focal endometrial lesion (indicated by the black arrow) in a mare with a chronic uterine infection.
Tumors of the uterus The differential diagnoses of a mass in the uterus includes tumor and intramural hematoma formation. Intramural hematomas occur following foaling, and generally slowly resolve over time. The most common uterine tumor is a leiomyoma (Figure 206.14a).9
Figure 206.12 Hysteroscopic view of the uterine bifurcation in a mare with chronic infectious endometritis that has resulted in chronic endometrial disease.
These are slow-growing tumors that form from the proliferation of cells that resemble smooth muscle. They have been reported to range in size from 2 to 5 cm, and may be pedunculated. If other tissue elements are more active, or the tumor is more invasive,
Figure 206.13 Diseased endometrium of a mare with a pyometra. The pyometra was related to cervical adhesions that formed after a fetotomy procedure. Transluminal adhesion indicated by the arrow as a vertical band on the left side of the image.
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Figure 206.14 Hysteroscopic view of a uterine a uterine leiomyoma (a)(white arrows) and a leiomyosarcoma (b) (white arrows) with superficial necrosis (indicated by the black arrow).
the tumor may be called a fibroleiomyoma, or leiomyosarcoma. Small tumors of this classification may be followed and may not interfere with fertility. Larger tumors may obstruct the uterine lumen, become necrotic on the surface as they outgrow their blood supply, and interfere with fertility (Figure 206.14b). This type of tumor is removed using a surgical approach.9, 23, 24 A mare must have 70% of her uterus remaining following surgical removal of the tumor for her to recognize the presence of the embryo and carry her own pregnancy to term. Uterine adenocarcinomas have also been reported and result in significant tissue damage, and frequently uterine discharge. Lymphosarcoma and melanoma may also metastasize to the uterus.28 Foreign bodies Foreign bodies in the mare’s uterus often involve broken parts of culture swabs including parts of the rod and the sample tip (Figure 206.15). The broken ends
Figure 206.15 Hysteroscopic view of the broken tip of a culture swab shown by the arrow in the uterus of a mare.
of culture swabs are often retrieved manually by dilating the cervix and retrieving the missing piece, or by using uterine lavage to flush out the pieces. In some cases implements such as endoscopic biopsy tools or a biopsy forceps may be used to dislodge a foreign body that is embedded or trapped in the endometrium. A tool may be used to drag the piece near the cervix for retrieval or may be used to hold onto the foreign body as the endoscope is withdrawn from the uterus. Fetal maceration or mummification Occasionally a fetus dies and the mummified or macerated bones are retained in the mare’s uterus. The bones may be adherent to the endometrium and require manipulations such as bringing the mare into heat and using manual cervical dilation or use of tools through the endoscope’s biopsy port to dislodge them. Uterine lacerations The uterus of the mare may be injured during foaling. Full-thickness lacerations of the dorsal portion of the uterine horn are the most common injury. These mares present with progressive depression postpartum. Some advocate the use of transrectal palpation and transrectal/transabdominal ultrasonography to identify uterine subinvolution, and focal hematoma formation or disruption at the site of trauma, combined with hysteroscopy to visualize the tear or to confirm the diagnosis. The postpartum uterus is difficult to insufflate due to its large volume, large cervix, and excessive endometrial folds, and a laceration further increases the difficulty in achieving good insufflation. Therefore, it is typically not possible to fully evaluate the subinvoluted postpartum
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Endoscopic Examination uterus using either ultrasonography or hysteroscopy. Abdominocentesis should be performed to determine if septic peritonitis is present when an uterine tear is suspected. Retained fetal membranes Mares may retain all or part of their fetal membranes. Retention of the tip of fetal membranes in the nonpregnant uterine horn is one of the most common presentations. This is why foaling attendants should evaluate the fetal membranes to determine if all of it has been delivered. Mares following cesarean section may have the fetal membranes inadvertently sutured into the hysterotomy closure. Mares with retained fetal membranes usually show clinical signs by 12–24 hours postpartum. Subinvolution of the uterus, foulsmelling fluid discharge, and echogenic material in the uterus are all indications of retained fetal membranes. Hysteroscopy is sometimes used to evaluate the post-foaling mare, or to identify the location of a piece of the fetal membranes.
Pathological conditions of the rectum Rectal stricture Mares that strain excessively during a dystocia, particularly as a result of excessive traction on the fetus, may prolapse their rectum and develop rectal injuries. The mesocolic artery in the mare is the main blood supply to the caudal rectum. Prolapse of the rectum may result in rupture or damage to this artery and subsequent tissue perfusion injury, leading to devitalization and necrosis of the rectum. The examiner may not be able to perform a rectal examination due to a stricture of the devitalized tissue. The rectum may be evaluated by placing a clear speculum into the rectum, a Caslick-type speculum into the rectum, or through endoscopy. A yellowish necrotic area is clearly seen in the rectum when this injury occurs. This condition requires surgical correction to save the life of the mare, and the prognosis is poor due to the difficulty in accessing this area surgically, inability to resect the rectum, and complications from peritonitis. Rectal tears Rectal tears may occur as a consequence of foaling. Small bowel or small intestine often prolapse through these rectal tears. Iatrogenic rectal tears in the mare are a major cause of professional liability claims. Rectal injury is a well-recognized risk of rectal examination. The overall risk of injury is low, but the
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consequences after a full-thickness rectal injury are often disastrous. If a popping or yielding sensation was felt during a rectal examination, or blood was found on the examiner’s rectal sleeve following the examination, it should trigger further investigation. Bleeding may be caused by rectal mucosal irritation, mucosal injury, mucosal and submucosal injury, and full-thickness injuries. Speculum examination of the rectum, palpation of the lesion, and endoscopy are often used to determine if a rectal injury extends from the submucosa into the abdomen. Endoscopy is particularly useful in cases where the lesion dissects away from the rectal luminal injury. Visualization of bowel is confirmation of a full-thickness injury. Incision of the rectum for the purpose of visualizing the full-thickness rectal injury, followed by manual prolapse to exteriorize and repair the injury, has recently been reported.29
References 1. Bracher V, Mathias S, Allen WR. Videoendoscopic evaluation of the mare’s uterus: II. Findings in subfertile mares. Equine Vet J 1992;24(4):279–84. 2. Wilson GL. Equine hysteroscopy: a window to the internal reproductive tract. Vet Med Small Anim Clinician 1983;78(9):1455–9,1462–6. 3. Ball BA. Hysteroscopic and low-dose insemination techniques in the mare. Ippologia 2006;17(3):25–30. 4. Morris LHA, Allen WR. An overview of low dose insemination in the mare. In 6th Annual Meeting of the European Society for Domestic Animal Reproduction, University of Parma, Italy, 12–14 September 2002, pp. 206–10. 5. Lindsey AC, Schenk J, Graham J, Bruemmer J, Squires EL. Hysteroscopic insemination of low numbers of flow sorted fresh and frozen/thawed stallion spermatozoa. Equine Vet J 2002;34(2):121–7. 6. Wilson GL. Equine hysteroscopy: normal intrauterine anatomy. Vet Med Small Anim Clinician 1984;79(11):1388– 93. 7. Bracher V, Allen WR. Videoendoscopic evaluation of the mare’s uterus: 1. Findings in normal fertile mares. Equine Vet J 1992;24(4):274–8. 8. Leidl W, Schallenberger-Pottiez U. Hysteroscopy in the mare. Vet Med Rev 1976(2):203–17. 9. Berezowski C. Diagnosis of a uterine leiomyoma using hysteroscopy and a partial ovariohysterectomy in a mare. Can Vet J 2002;43(12):968–70. 10. Menzies-Gow N. Diagnostic endoscopy of the urinary tract of the horse. In Practice 2007;29(4):208–13. 11. Wilson GL. Hysteroscopic examination of mares. Vet Med Small Anim Clinician 1983;78(4):568–78. 12. Bartmann CP, Schiemann V. Establishment of a standardized intrauterine distension pressure for the hysteroscopy in the horse. Deutsche Tierarztliche Wochenschr 2003;110(2):43–8. 13. Hecker C, Hospes R. Endometrial biopsy in the mare – ‘blind’ sampling of tissue or under hysteroscopic control? Pferdeheilkunde 2006;22(2):140–4. 14. Schiemann V, Bartmann CP, Kirpal G, von Reiswitz A, Schoon HA, Klug E. Diagnostic hysteroscopy in the
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mare – uterine contamination and endometrial reaction. In: 3rd International Conference on Equine Reproductive Medicine, Leipzig, Germany, 27–28 October 2001, pp. 557–64. Ferrer MS, Lyle SK, Eilts BE, Godke RA, Paccamonti DL. Post-mating endometritis after low dose hysteroscopic insemination in the mare. Theriogenology 2005;64(3): 781–4. Kaspar B, Kahn W, Laging C, Leidl W. Endometrial cysts in mares. I. Post mortem studies, occurrence and morphology. Tierarztliche Praxis 1987;15(2):161–6. Leidl W, Kaspar B, Kahn W. Endometrial cysts in the mare. 2. Clinical examinations: frequency and importance. Tierarztliche Praxis 1987;15(3):281–9. Bartmann CP, Komann M, Schiemann V, Stief B, Schoon HA, Klug E. Hysteroscopic removal of uterine cysts in mares I – Hysteroscopy and surgical procedures. In: Fifth International Conference on Equine Reproductive Medicine. Leipzig, Germany, 24–25 November 2007, pp. 31–4. Hollerrieder J, Toth J. Minimal-invasive surgical removal of endometrium cysts in mares. Magyar Allatorvosok Lapja 2003;125(1):5–10. Manning ST, Bowman PA, Fraser LM, Card CE. Development of hysteroscopic insemination of the uterine tube in the mare. In: Proceedings of the American Association of Equine Practitioners, 1998; pp. 84–5. Vasquez JJ, Medina V, Liu IKM, Ball BA, Scott MA. Nonsurgical uterotubal insemination in the mare. In: Proceedings of the American Association of Equine Practitioners, 1998; pp. 68–9. Lindsey AC, Morris LHA, Allen WR, Schenk JL, Squires EL, Bruemmer JE. Hysteroscopic insemination of mares with
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low numbers of nonsorted or flow sorted spermatozoa. Equine Vet J 2002;34(2):128–32. Hollerrieder J, Toth J. Minimal invasive surgery of a bleeding uterine tumor in a mare using a wire cerclage. Tierarztliche Praxis (Ausgabe G, Grosstiere/Nutztiere), 2002;30(6):404–7. Janicek JC, Rodgerson DH, Boone BL. Use of a handassisted laparoscopic technique for removal of a uterine leiomyoma in a standing mare. J Am Vet Med Assoc 2004;225(6):911–14. Kollmann M, Bartmann CP, Schiemann V, Klug E, Ellenberger C, Schoon HA. Hysteroscopic removal of uterine cysts in mares II – Follow-up and long term fertility analysis with regard to patho-histological findings. In: Fifth International Conference on Equine Reproductive Medicine, Leipzig, Germany, 24–25 November 2007, pp. 35–7. Schiemann V, Bartmann CP. Transendoscopic synechiolysis of extensive intrauterine adhesions by repeated operative hysteroscopy in a mare – a case report. Pferdeheilkunde 2003;19(6):661–5. Bartmann CP, Brickwedel I, Klug E. Trans-endoscopic high frequency electrosurgery for minimal invasive treatment of intrauterine adhesions in mares. Tierarztliche Praxis (Ausgabe G, Grosstiere/Nutztiere), 2000;28(4):233–9. Neufeld JL. Lymphosarcoma in a mare and review of cases at the Ontario Veterinary College. Can Vet J 1973;14(7):149– 53. Kay AT, Spirito MA, Rodgerson DH, Brown SE. Surgical technique to repair grade IV rectal tears in post-parturient mares. Proceedings of the 51st Annual Convention Proceedings of the American Association of Equine Practitioners, Seattle, Washington, USA, 3–7 December 2005, pp. 345–9.
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Chapter 207
Cytogenetic Evaluation of Mares and Foals Teri L. Lear and Daniel A.F. Villagomez
Cytogenetics is the study of chromosomes, which are threadlike complexes of DNA and proteins present in all nucleated cells. The genes and regulatory elements residing on each chromosome control cellular functions and development. Chromosomal loss or aberrations can disrupt normal cellular functions resulting in embryonic death, congenital abnormalities, or infertility. In humans, chromosome abnormalities are well documented as causes of congenital disorders, infertility, or subfertility. Overall, chromosome abnormalities can be identified in more than 0.5% of human newborns.1 In humans, a myriad of chromosome abnormalities cause a wide variety of phenotypes. Chromosome abnormalities also occur in horses but they have not been studied as extensively as in humans. However, as chromosome analysis techniques have improved over the last 50 years and more horse samples have been submitted for study, more chromosome abnormalities have been identified that cause infertility or congenital abnormalities.
The most common chromosome abnormalities identified in horses are associated with an abnormal number of sex chromosomes. Most of these occur spontaneously and are not inherited. Some abnormalities that may be inherited are identified as inconsistencies between gonadal sex and phenotypic sex. Both of these conditions cause infertility. Abnormalities of other chromosomes cause repeated early embryonic loss, which appears as subfertility of one of the parents. Since the chromosomal aberration can be inherited from the sire or dam, offspring of these horses should be karyotyped to determine if they have a normal chromosome complement or if they carry the same abnormality. This will avoid wasting valuable time and resources trying to breed these animals. While most foals with severe congenital abnormalities are not karyotyped, it should be kept in mind that abnormalities in chromosome number can cause severe birth defects. Identifying a chromosome abnormality could prevent undue suffering of a foal for which there is no curative treatment.
Horse karyotyping All domestic horses, regardless of breed, have a diploid chromosome number of 64 (2n = 64). The correct number of horse chromosomes was identified by Rothfels and co-workers in 1959.2 Since that time it has been shown that chromosome abnormalities cause infertility, early embryonic loss, and congenital abnormalities in horses. Chromosome abnormalities may be present in approximately 2–3% of the living horse population,3 with many abnormalities going undetected until a horse goes to the breeding shed. Some chromosome abnormalities are never identified, as in the case of foals that are euthanized because of severe congenital abnormalities.
Chromosome terminology, karyotype standards, procedures Chromosome terminology When reading a clinical cytogenetics report, chromosome terminology can be confusing. A basic knowledge of this terminology can be helpful when interpreting a report and communicating the results to colleagues and clients.
Chromosome morphology Chromosomes are identified primarily by their size and shape relative to their centromere (Figure 207.1).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 207.1 Images of horse chromosomes 2, 4, and 24 representing horse chromosome morphology classification by centromere position (→), arm length, and size. Chromosome short arms are designated ‘p’ and long arms ‘q’.
The centromere is a constricted region of the chromosome where the mitotic apparatus attaches to pull the chromosomes apart during mitosis. A chromosome having a centromere located in or near the middle of the chromosome is described as metacentric or submetacentric. One in which the centromere is located at or near the end is said to be acrocentric or telocentric. Chromosome arms are designated ‘p’ for the short arm and ‘q’ for the long arm. Chromosomes are lined up for comparison in a diagram, or karyotype, based on their size, arm length, centromere position, and banding pattern.
Figure 207.2 G-banded horse chromosome 1 and its ideogram. Ideogram adapted from ISCNH.8
Interphase versus metaphase analysis Autosomes and sex chromosomes Most genes that are important for sexual development reside on the sex chromosomes, X and Y. Normal females have two X chromosomes and normal males have one X and one Y chromosome. All other chromosomes are referred to as the autosomes.
Chromosome banding Chemical and enzymatic treatments that create chromosome-specific staining patterns result in chromosome banding. The chemical make-up of each chromosome (the proteins present, the nature of the DNA along the chromosome) determines the banding pattern on each chromosome. Each chromosome has a unique banding pattern, making it possible for cytogeneticists to determine if the chromosome is present, lost, duplicated, or structurally altered. Chromosome pairs with the same banding pattern are numbered based on their position in the karyotype. Also, bands are given numbers so regions of the chromosome that are deleted, duplicated, or translocated can be identified. Band numbers are represented in ideograms, which are diagrams representing the chromosome’s morphology and banding pattern. An example of a banded horse X chromosome with its ideogram is shown in Figure 207.2.
Most of the life cycle of a cell is spent in a state called interphase, when the cell is not undergoing cell division although DNA synthesis is occurring. At interphase the chromosomes within the nucleus are not condensed, therefore they are not visible under the microscope. At the beginning of cell division the cells enter metaphase. During metaphase the chromosomes become highly condensed prior to cell division, and they are clearly visible. Both interphase and metaphase chromosomes are useful for cytogenetic analysis. Metaphase chromosomes are used for karyotypes. Interphase nuclei are very useful for molecular cytogenetic analysis when a large number of cells need to be scored to rule out mosaicism, where an individual has two or more cell lines. In these cases, interphase cells are used to determine the percentage of normal versus abnormal cells. In many cases a higher ratio of normal to abnormal cells is associated with a less severe phenotype. Karyotype standard The current karyotype standard for the horse, the International Standard for Cytogenetic Nomenclature for the Horse (ISCNH), was completed in 1997 as part of the Horse Genome Mapping Workshop. Equine genetics researchers realized that a high-quality karyotype standard was needed to enable all scientists
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Cytogenetic Evaluation of Mares and Foals to consistently identify the same chromosomes based on their morphology and banding patterns. This allowed scientists to use the same standard when locating genes on chromosomes to develop a horse genetic map. The standard consists of karyotypes, ideograms, and written descriptions of each chromosome’s banding patterns for two of the most commonly used chromosome identification methods.
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immediately to the laboratory. The biopsy is then minced and digested with collagenase prior to placing the cells in culture medium. After several days the cells attach to the culture flask and begin to divide. When the cells have reached sufficient density they are removed from the flasks and placed in new flasks at a lower cell density. Approximately 16–48 hours later the cells are harvested using techniques similar to those described for lymphocytes.
Procedures
Sample submission Veterinarians who wish to submit samples for cytogenetic testing should check with their laboratory of choice to see what requirements the laboratory may have for sample submission. Sample types Most laboratories prefer working with blood samples, although chromosomes can be obtained from skin fibroblasts and even amniotic fluid. However, blood sample collection is the least invasive method for obtaining nucleated cells. It is also easier to transport and culture and results are obtained faster than if other tissues are used. Peripheral blood is collected in sodium heparin, which is the best preservative for nucleated lymphocytes. The blood should be sent to the laboratory as quickly as possible. Living cells are needed for culture, thus the blood should not be subjected to extremes of heat or cold. White blood cells are isolated from the blood samples and incubated in special tissue culture medium containing mitogenic compounds that stimulate cell division. After 70–72 hours of culture, cell division is arrested by the addition of colcemid. The cells are treated with a hypotonic solution that causes the cellular nuclei to swell. Next, the cells are fixed in several changes of methanol and acetic acid. And finally, the cells are dropped onto microscope slides causing the cells to burst open allowing the chromosomes from individual nuclei to spread onto the glass slide. At this point the slides can be treated with the appropriate chemicals or enzymes to produce the bands before staining with dyes as described below. The chromosomes are then ready for viewing under the microscope. Usually fibroblast cultures are obtained from skin biopsies using a sterile biopsy punch, or skin and organ samples are collected at necropsy. The skin should be shaved and prepped as for surgery. All traces of iodine-containing agents should be thoroughly cleaned off and the skin rigorously cleaned with 70% alcohol. Whenever possible the biopsy should be placed in sterile collection medium obtained from the cytogenetics laboratory and shipped
Chromosome banding Numerous types of chromosome banding methods can be used by a cytogenetics laboratory to identify each chromosome pair. Each method offers specific information. The choice of banding method may be up to the veterinarian, thus it is important to understand what method would provide the most information for a particular case. Nearly all laboratories use Giemsa stain (Figure 207.3) for a first look at horse chromosomes. Giemsa stains each chromosome and enables the chromosome number and morphology to be determined. However, it does not enable the identification of each chromosome pair. Giemsa stain is best used when only a chromosome count is needed, such as in cases of infertility and gonadal dysgenesis or ambiguous genitalia. For confirmation of sex chromosomes C-banding (Figure 207.4) can be used.4, 5 This method identifies chromosome centromeres, a band on the X chromosome, and the majority of the Y chromosome. These regions are composed of highly repetitive DNA called heterochromatin. This is especially useful for identifying the Y chromosome, which is almost entirely heterochromatic. Most X chromosomes can be identified by the heterochromatic band on Xq; however, this band may vary in size making some X chromosomes difficult to identify. In these cases it is best to do other more precise types of banding. G-banding (Giemsa trypsin banding or GTGbanding) (Figure 207.5a,b) produces a banding pattern specific for each chromosome following treatment with trypsin and Giemsa stain.6 In this way each chromosome can be paired and their banding patterns compared in order to identify any gain or loss of genetic material or any structural rearrangement. G-banding is useful for all types of suspected chromosome abnormalities and should definitely be done in cases of repeated early embryonic loss. R-banding (reverse banding), more complicated than G-banding, creates a reverse pattern of G-banding.7 Many European laboratories use R-banding while G-banding has been the method of choice at North American laboratories. Both methods are equally informative.
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Figure 207.3 Giemsa-stained karyotype from a normal mare.
Whichever method is used, the chromosomes are paired together in a karyotype based on their relative order of size and morphology. The karyotype should follow the most recent standard for the horse.8
Molecular tests Other types of testing may need to be performed in order to more accurately to characterize a chromosome abnormality. For instance, to rule out the presence of a Y chromosome or to characterize the chro-
Figure 207.4 Partial metaphase showing C-banding patterns of the X and Y chromosomes.
mosome breakpoints involved in a structural rearrangement, fluorescence in situ hybridization (FISH) can be used. FISH involves tagging a cloned piece of DNA with a fluorescent molecule creating a ‘probe’. The probe is hybridized to the specific chromosomal site containing homologous DNA. The probe will illuminate under a fluorescent microscope revealing the presence of a gene. The process begins by cloning a piece of DNA, and growing it up in a vector, such as a bacterial artificial chromosome (BAC). The BAC contains a piece of horse DNA for a known gene. BAC DNA is purified and tagged with a fluorescent reporter molecule. By taking advantage of the double-stranded nature of DNA the probe is denatured and allowed to bind to the denatured chromosomal DNA or ‘target’. If the two DNAs share complementary sequences they will anneal, or ‘hybridize’, to each other and whatever does not anneal will be washed off. The probe will be illuminated under a fluorescent microscope allowing the cytogeneticist to determine if the gene is present and its position on the chromosome (Figure 207.6). Polymerase chain reaction (PCR) is a useful technique for identifying the presence or absence of specific genes, for example genes involved in sex
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Cytogenetic Evaluation of Mares and Foals
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Figure 207.5 G-banded metaphase spread from a normal mare as it would appear under the microscope (a) and karyotyped (b).
determination such as the sex-determining region Y gene (SRY). The technique involves using a DNA polymerase to amplify a piece of DNA of interest via an enzymatic replication reaction.9 Horse DNA (template) is combined with the polymerase and nucleotides in the presence of DNA oligonucleotides (primers), which are required to initiate DNA syn-
thesis. If the DNA sequence specific for the primers is present in the template DNA the primers will bind the template and DNA synthesis will occur. The reaction products can be observed by agarose gel electrophoresis with the appropriate DNA fragment size standards and controls and the presence or absence of a particular gene noted (Figure 207.7).
Clinical reports A karyotype report should contain any information associated with the animal such as name, laboratory identification number, referring veterinarian, number of cells analyzed, chromosome number associated with the cells analyzed, banding techniques used, at least one karyotype, and the diagnosis. In cases showing a chromosome abnormality a brief description of the cause of the abnormality, if known, could be useful.
Clinical cases involving infertility or subfertility
Figure 207.6 An example of fluorescence in situ hybridization (FISH): two bacterial artificial chromosome (BAC) probes mapped next to each other (arrow) just above the centromere at horse chromosome Xp in a stallion. (See color plate 82).
Most cases submitted for karyotyping are from horses that are infertile or subfertile. Karyotyping can identify a chromosomal cause of infertility and differentiate the type of chromosomal aberration. The majority of cases involve abnormalities of the sex chromosomes.
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Figure 207.7 Polymerase chain reaction (PCR) results for sexdetermining region Y gene SRY. Lane 1: DNA fragment size ladder. Lane 2: normal SRY-negative mare. Lane 3: normal SRYpositive stallion.
(0.5 × 0.5 × 1.0 cm), with a flaccid uterus and cervix, should be karyotyped to rule out X chromosome monosomy. Monosomy is a term meaning that only one copy of a chromosome is present; as noted above, the normal karyotype has two copies of all chromosomes with the exception of the sex chromosomes in males. This is the most common chromosome abnormality of mares10 (Figure 207.8). These mares have only one X chromosome instead of the two X chromosomes seen in normal mares. In humans, this condition is known as Turner syndrome and causes primary infertility. Sometimes the condition is referred to as ‘equine Turner syndrome’, although the phenotypic effects on horses are not as severe as those seen in humans. In humans, a large proportion of embryos with Turner syndrome are aborted.11 The degree to which X monosomy contributes to early embryonic loss in the horse is unknown. In addition to primary infertility, mares with only one X chromosome may have angular limb deformities, small stature, and overall poor conformation.12 These mares are always infertile.
63,X mosaicism Mares with normal external genitalia, gonadal dysgenesis, primary infertility, and in some cases small stature
63,X Mares that exhibit primary infertility, abnormal or absent estrous cycles, small inactive ovaries
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Mares with X monosomy may also be ‘mosaics’. Mosaicism is the presence of at least two different cell populations within a single individual. This may occur when one X chromosome is lost during one of the very early embryonic cell divisions, resulting in a population of cells with a normal set of chromosomes (64,XX) and one with an abnormal set of
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Figure 207.8 (a) A 10-month-old Thoroughbred filly with growth retardation and monosomy X. (b) Giemsa-stained monosomy X karyotype from the same filly.
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64,X,i(Xq) One report described a Standardbred filly of small stature with poorly developed pelvic musculature, small ovaries, enlarged frontal sinuses, a head tilt, vacuoles in her eye lenses resulting in night blindness, and a wrinkled muzzle.17 Karyotyping showed this filly to have a deletion of the short arm (Xp) and a duplication of the long arm of the X chromosome (Xq). The two X long arms had fused at the centromere creating a mirror image, called an isochromosome (i). Figure 207.9 C-banded X chromosomes demonstrating a deletion of Xp (arrow) in a mare with gonadal dysgenesis.
chromosomes (63,X). Other complicated variations have been described in mosaic animals. For example, individuals with three different cell types, would present with three different karyotypes during cellular analysis, 63,X/64,XX/65,XXX. The degree of fertility is correlated with the percentage of normal cells present. A large percentage of mares with X monosomy described in the literature were mosaic forms.10, 13 Nearly all of these mares were infertile.
64,X,delXp Mares with deletions (del) of the short arm of the X chromosome (Xp) are indistinguishable from mares with X monosomy and are also infertile14 (Figure 207.9). This may be due to the deletion of genes along the short arm of the X chromosome, which are critical for normal sexual development.
64,XX,delXq/64,XY A 4-year-old Standardbred mare with normal external genitalia and normal estrous cycles but extremely masculine behavior was shown to be a mosaic.15 She had a cell line with a deletion of the long arm of the X chromosome (Xq) and another normal male cell line. It was reported that she produced a foal.
65,XXX Some individuals have been identified with three copies of the X chromosome.14, 16 Three copies of a chromosome are referred to as trisomy. While human individuals with trisomy X appear to be fertile, this is not the case in horses. Horses with X trisomy also exhibit normal external genitalia but gonadal dysgenesis.
Mares with normal external genitalia, gonads, and estrous cycles but repeated early embryonic loss There are numerous causes of early embryonic loss in mares, some of which involve chromosome abnormalities. Chromosome abnormalities that involve a change in the DNA organization can cause infertility or subfertility. When a portion of a chromosome is transferred or exchanged with another chromosome a translocation has occurred. This is demonstrated in Figure 207.10, where a portion of horse chromosome 1 has broken and reattached to chromosome 16. If the translocation does not result in a gain or loss of genetic material it is called a balanced translocation. If a gain or loss of material occurs then it is an unbalanced translocation. When all the genetic material is present the horse will appear as a normal healthy individual. However, during the formation of gametes, the abnormal chromosomes may segregate to the gamete causing them to be unbalanced. The abnormal genetic complement results in a non-viable embryo. The production of unbalanced gametes depends on the severity of the chromosome rearrangement. For example, mares with a poor reproductive record that experience repeated early embryonic loss prior to day 65 of gestation may carry balanced translocations even if they have produced one or more normal foals.
64,XX,t(1;3) 64,XX,t(1;16) 64,XX,t(1;21) 64,XX,t(16;22) 64,XX,t(4;13) Only five mares, all Thoroughbreds, with balanced chromosome translocations involving only the autosomes have been reported in the literature.18–20 All of the mares were submitted for karyotyping due to repeated early embryonic loss. In all except the t(1;3) case, both G-banding and FISH were utilized to identify and characterize the structural rearrangements constituting the translocations. The ability to produce a live foal varied greatly among these mares and may be dependent on the type of
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Mare: Techniques in Reproductive Examination mation, a stiff gait, and mental dullness, was shown to have a chromosome abnormality.21 She carried a deletion of the short arm of one X chromosome (Xp). An extra copy part of chromosome 15 (trisomy 15) was translocated to the deleted X (t(X;15)). While mares missing the short arm of X are viable, it is unclear how this mare survived trisomy for even a small portion of chromosome 15.
Multiple congenital abnormalities with or without growth retardation
Figure 207.10 G-banded karyotype of a phenotypically normal Thoroughbred colt with the same chromosome translocation as his dam. A portion of chromosome 1q is translocated to the terminal end of chromosome 21 [64,XY,t(1;21)] (see arrows).
chromosome abnormality present. In one case a phenotypically normal colt carried the same translocation as his dam (Figure 207.10), thus demonstrating the heritable nature of structural chromosome rearrangements. In the case of his dam, molecular FISH studies with BAC clones mapping around the proposed breakpoint on chromosome 1 were done in order to characterize the chromosome breakpoints and confirm the chromosomes involved in the translocation (Figure 207.11).20
64,X,-X,t(X;15) An infertile Thoroughbred mare with normal external genitalia and reproductive tract, but poor confor-
Horses with multiple congenital anomalies and/or growth retardation should be karyotyped to rule out the presence of an extra autosome, a condition called autosomal trisomy. The most familiar form of trisomy in humans is Down syndrome, which affects children born with an extra copy of chromosome 21. Although autosomal trisomy in horses is rare, trisomy of chromosome 31 has been reported in both pure and mosaic forms. Nearly all cases of pure trisomy have involved the smallest chromosomes in the horse karyotype. Trisomy of larger autosomes is incompatible with life and may contribute to embryonic loss. Trisomy is not confined to one breed but has been reported in many different breeds. The phenotypes of all cases of trisomy reported are summarized below:
65,XY,+23 This was a Standardbred colt with rough hair coat, poor body condition, facial asymmetry, dorsomedial strabismus, a dysmetric gait, mildly uncoordinated with head and tail tilt, and small testicles.22
Figure 207.11 In order to determine the chromosome breakpoint and confirm the chromosomes proposed to be involved in the translocation, four BAC clones known to map on horse chromosome 1 near the breakpoint were FISH mapped. All four BAC clones, RET and GHITM and FES and EFTUD1 were present on the normal chromosome 1. However, RET and GHITM (most proximal to the centromere) were present on the deleted chromosome 1, while FES and EFTUD1 (more distal to the centromere) transferred with the broken chromosome 1 fragment and fused with the terminal end of chromosome 21. (Adapted from Lear et al.20 ).
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Figure 207.12 (a) A Thoroughbred colt with severe congenital abnormalities.(b) The same colt’s karyotype demonstrating trisomy 31 (see arrow).
65,XX,+26
65,XY,+31
This case was a Thoroughbred mare with angular limb deformities of all four limbs, a stiff gait, physically inactive, poor conformation, mentally dull, reproductively normal with normal maternal behavior, and intolerant of pain or physical restraint.23
This Thoroughbred colt (Figure 207.12) was dysmature, had a fine hair coat, an inability to suck, a domed skull, brachygnathia inferior, superficial and deep flexor contracture of both front limbs with carpal and fetlock valgus angular deformity of the left front limb, scoliosis, laxity of flexor tendons in the rear limbs, a malformed prepuce, a small right testis, no discernible scrotum, neurological deficits, metabolic disorders, partial anhidrosis, and a head tilt.26 Two cases of mosaic trisomy 31 have been reported. The first horse was a Thoroughbred mare with 93% normal and 7% abnormal cells, with no phenotypic abnormalities.3 The second case was a Thoroughbred filly with 40% normal and 60% abnormal cells, with severe angular limb deformities, growth retardation, and neurological deficits (T.L. Lear, unpublished data).
65,XY,+27 This case was a Quarter Horse colt with bilateral carpal flexural deformity of its front limbs and inability to suck.24
65,XY,+28 This Thoroughbred colt was described with small stature, unilateral cryptorchidism, unstimulated serum testosterone of 25 pg/mL, stimulated serum testosterone of 85 pg/mL, azoospermia, and somewhat abnormal behavior.21
Disorders of sexual development of the equine embryo
65,XX,+30 This case was an Arabian filly with small stature, severe angular deviation of the front legs with mild polydactyly, and joint rigidity.23 A case of mosaic trisomy 30 involved a Polish pony mare with 26% normal and 74% abnormal cells; it showed no phenotypic abnormalities.25
Normal male sexual differentiation Each embryo has bisexual potential and each gonad can develop into either an ovary or testis. The genetics of sexual development is complex. A plethora of genes appear to be involved with many affecting gonadal precursor tissues.27, 28 However, if male sexual
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development is to take place, testicular determinant genes must be present and functional. The primary switch for male sexual differentiation appears to be a gene on the Y chromosome, SRY (sex-determining region Y). However, SRY does not appear to be necessary for testis development. This may be the role of SOX-9 (SRY HMG-box-related gene 9).28 In any case, normal testes must produce testosterone, dihy¨ drotestosterone (DHT), and Mullerian inhibiting substance (MIS) for normal male sexual development to occur. In addition, androgen receptors must be functional for normal masculinization to occur. Normal female sexual differentiation When testicular determinant genes are not present or do not function properly the embryo will begin to develop as a female regardless of genetic sex.27, 28 The development of both female internal and external genitalia occurs in the absence of ovarian hor¨ mones. Mullerian development continues unless MIS is present, and without androgens the external genitalia will develop into a normal female. While many genes are known that play a part in male sexual differentiation, there is no known ovarian differentiation factor. Development of the intersex Intersexuality is a general clinical term for referring to ambiguous stages of internal and external reproductive organs. The disagreement between the genetic sex (XX or XY sex chromosome constitution) and sexual phenotype (female or male respectively) defines the sex-reversal syndrome in mammals. The occurrence of individuals in which the genetic sex does not agree with the expected phenotypic sex has been repeatedly documented in horses. In mammals, it is generally accepted that the sex chromosomes, X and Y chromosomes, determine the sex of an individual; however, deletions or mutations of genes controlling sexual development can result in the intersex state. Defects in a part of the developmental pathway lead to the intersex condition.29 The action of androgen is dependent on its ability to bind to a cytoplasmic protein receptor in target cells. If the androgen receptors (AR) are absent or defective, the target tissue becomes completely or partially insensitive to androgens, which results in deficient masculinization, also known as androgen insensitivity syndrome (AIS) and testicular feminization syndrome.30 However, in humans, in utero exposure of the genetic female fetus to exogenous androgens can cause virilization of the external genitalia. These androgens can come from ovarian tumors or from the ingestion
of androgens or progestins by the mother during critical stages of fetal sexual differentiation. Enzymatic defects in the cortisol pathway, called congenital adrenal hyperplasia (CAH), can produce androgens, which cause virilization of the female fetus.29, 31 None of these situations have been shown conclusively to cause abnormal sexual development in the horse, although research is currently underway. One gene involved in testis determination is SRY. Studies in humans have shown that mutations in SRY can result in gonadal dysgenesis among genotypic males (46,XY). Without normal Sertoli cells and Leydig cells female genitalia develop, wolffian ducts regress, and normal female genitalia develop.32 However, while the mechanisms of action are unclear, other factors (genes) are known to influence normal sexual development. Moreover, not all of these genes reside on the sex chromosomes. The majority of this research has been done in mice and humans. It is beyond the scope of this chapter to review the research in this area; however, two excellent reviews by MacLaughlin and Donahoe (2004)27 and Blecher and Erickson (2007)28 are available. Disorders of sexual development such as pseudohermaphroditism and true hermaphroditism in the horse are complex and not completely understood. It is difficult to categorize the disorders unless it is possible to combine the results from karyotyping, molecular, endocrine, biochemical, and surgical or ultrasound studies.
Normal female genitalia, absence of cycles, small ovaries, gonadal dysgenesis or abdominal testes, possible masculine behavior/large stature
Equine XY sex-reversal syndromes Equine XY sex-reversal occurs in phenotypic females that are genetic males. The syndrome was first described in the Arabian horse breed but has since been shown to exist in other breeds.14, 33, 34 These horses have normal female genitalia. They can be of normal or large stature. Some mares may demonstrate masculine behavior. Testosterone levels can vary greatly, and higher levels have been correlated with masculine behavior. These horses have a normal male karyotype (64,XY). In some Arabian horse families with a higher frequency of XY sex-reversal mares, pedigree analysis suggests that different hereditary patterns exist for this condition. When tested by PCR for the presence of SRY, most of these horses were negative.35–39 The SRY-negative XY sex-reversal horses are characterized by remarkably underdeveloped gonads, which have been interpreted as gonadal dysgenesis. Rectal palpation and
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Cytogenetic Evaluation of Mares and Foals ultrasonography may reveal a hypoplastic female reproductive track. Cases of equine XY sex-reversal SRY-positive horses are rare. Only one case has been reported in the literature.40 However, another similar case was reported for which SRY testing was not done.41 Both horses exhibited extreme masculine behavior and abdominal testes. In the case of the SRY-positive horse, it was concluded that the phenotype may be caused by androgen insensitivity resulting in the female phenotype. Ambiguous genitalia, elongated and fused vulva, enlarged clitoris
Equine XX sex-reversal syndromes Cases of ambiguous external genitalia occur most often in horses that have an enlarged clitoris and an elongated, fused vulva (Figure 207.13).38, 42 Horses may present as female pseudohermaphrodites with ambiguous external genitalia and a normal female karyotype (64,XX).43, 44 Molecular studies have shown that they are negative for the SRY gene. It is suspected that these individuals may have an equine form of the human autosomal recessive disorder, congenital adrenal hyperplasia (CAH). A true hermaphrodite with ovotestes was described in an Arabian filly.38 This filly had a normal
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karyotype (64,XX) and was SRY-negative. Recent literature on similar cases seen in humans has shown that testicular development can occur even in the absence of the SRY gene, but the genes involved have not been determined.45
Conclusions Chromosome analysis is an informative tool in the equine veterinary practice when trying to determine a cause of infertility and congenital abnormalities. Clearly, chromosome abnormalities have an impact on production and health. Conventional karyotyping combined with molecular studies using PCR and FISH can provide a wealth of information for the equine practitioner. In 2007 the horse genome sequence was completed. As a result of this, new molecular tools have become available that promise to provide higher-resolution analysis – ‘molecular karyotypes’ – of each horse chromosome than previously attained by conventional cytogenetics.46, 47 The information obtained using molecular karyotyping could shed light on how even the smallest changes in the genome can cause genetic disease in the horse.
Acknowledgements The authors would like to thank Dr Ernie Bailey for critically reading the manuscript. This chapter is in connection with a project of the University of Kentucky Agricultural Experiment Station and is published as paper 10-14-076.
References
Figure 207.13 Elongated, fused vulva, enlarged clitoris in a 64,XX, SRY-negative horse.
1. Gelehrter TD, Collins FS, Ginsburg D. Principles of Medical Genetics, 2nd edn. Baltimore: Williams & Wilkins, 1998. 2. Rothfels KH, Alexrad AA, Siminovitch L, McCulloch Parker RC. The origin of altered cell lines from mouse, monkey and man, as indicated by chromosome and transplantation studies. Proc Can Cancer Res Conf 1959;3:189–214. 3. Bugno M, Słota E, Ko´scielny M. Karyotype evaluation among young horse populations in Poland. Schweiz Arch Tierheilkd 2007;149:227–32. 4. Arrighi FE, Hsu TC. Localization of heterochromatin in human chromosomes. Cytogenetics 1971;10:81–6. 5. Sumner AT, Evans HJ, Buckland RA. A new technique for distinguishing between human chromosomes. Nature New Biology 1971;232:31–2. 6. Seabright M. A rapid banding technique for human chromosomes. Lancet 1971;ii:971–2. 7. Dutrillaux B, Lejeune J. Sur une novelle technique d’analyse du caryotype human. C R Acad Sci Paris 1971;272: 2638–40. 8. ISCNH Committee. International system for cytogenetic nomenclature of the domestic horse. Chromosome Res 1997; 5:433–43. 9. Mullis KB, Faloona FA, Scharf SJ, Saiki RK, Horn GT, Erlich HA. Specific enzymatic amplification of DNA in
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10.
11. 12. 13. 14.
15. 16.
17. 18.
19.
20.
21.
22.
23.
24.
25. 26. 27. 28. 29.
30.
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vitro: the polymerase chain reaction. Cold Spring Harb Sym 1986;51:263–73. Power MM. Chromosomes of the horse. In: McFeely RA (ed.) Advances in Veterinary Science and Comparative Medicine, Domestic Animal Cytogenetics, vol. 34. San Diego: Academic Press, 1990; pp. 131–67. Jones KL. Smith’s Recognizable Patterns of Human Malformation, 6th edn. Philadelphia: Elsevier, 2006; pp. 76–81. Bowling AT. Horse Genetics. Wallingford: CAB International, 1996. Lear TL, Bailey E. Equine clinical cytogenetics: the past and future. Cytogenet Genome Res 2008;120:42–9. Bowling AT, Millon L, Hughes JP. An update of chromosomal abnormalities in mares. J Reprod Fertil Suppl 1987; 35:149–55. Halnan CRE. Cytogenetics of Animals. Wallingford: CAB International, 1989. Chandley AC, Fletcher J, Rossdale PD, Peace CK, Ricketts SW, McEnery RJ, Thorne JP, Short RV, Allen WR. Chromosome abnormalities as a cause of infertility in mares. J Reprod Fertil Suppl 1975;23:377–83. M¨akel¨a O, Gustavsson I, Hollm´en T. A 64,X,i(Xq) karyotype in a Standardbred filly. Equine Vet J 1994;26:251–4. Power MM. The first description of a balanced reciprocal translocation [t(1q;3q)] and its clinical effects in a mare. Equine Vet J 1991;23:146–9. Lear TL, Layton G. Use of Zoo-FISH to characterize a reciprocal translocation in a Thoroughbred mare: t(1;16)q16; q21.3). Equine Vet J 2002;34:207–9. Lear TL, Lundquist JM, Zent WW, Fishback WD Jr, Clark A. Three autosomal chromosome translocations associated with repeated early embryonic loss in the domestic horse (Equus caballus). Cytogenet Genome Res 2008;120:117–22. Power MM. Equine half sibs with an unbalanced X;15 translocation or trisomy 28. Cytogenet Cell Genet 1987;45: 163–8. Klunder LR, McFeely RA, Beech J, McClune W, Bilinski WF. Autosomal trisomy in a Standardbred colt. Equine Vet J 1989;21:69–70. Bowling AT, Millon LV. Two autosomal trisomies in the horse: 64,XX,-26,+(26q26q) and 65,XX,+30. Genome 1990; 33:679–82. Buoen LC, Zhang TQ, Weber AF, Turner T, Bellamy J, Ruth GR. Arthrogryposis in the foal and its possible relation to autosomal trisomy. Equine Vet J 1997;29:60–2. Kubien´ E, Tischner M. Reproductive success of a mare 64,XX/65,XX,+30. Equine Vet J 2002;34:99–100. Lear TL, Cox JH, Kennedy GA. Autosomal trisomy in a Thoroughbred colt: 65,XY,+31. Equine Vet J 1999;31:85–8. MacLaughlin DT, Donahoe PK. Sex determination and differentiation. New Engl J Med 2004;350:367–78. Blecher SR, Erickson RP. Genetics of sexual development: A new paradigm. Am J Med Genet A 2007;143A:3054–68. Hutchinson J, Snyder HM III. Ambiguous genitalia and intersexuality. 2006; www.emedicine.com/ped/ TOPIC1492.HTM. Holterhus PM, Werner R, Hoppe U, Bassler J, Korsch E, ¨ HG, Hiort O. Molecular features and clinRanke MB, Dorr ical phenotypes in androgen insensitivity syndrome in the absence and presence of androgen receptor gene mutations. J Mol Med 2005;83:1005–13.
31. Nussbaum RL, McInnes RR, Willard HF. Clinical cytogenetics: disorders of the autosomes and sex chromosomes. In: Thompson & Thompson Genetics in Medicine, 6th edn. Philadelphia: W.B. Saunders, 2001; pp. 157–79. 32. Migeon CJ, Wisniewski AB. Sexual differentiation: from genes to gender. Hormone Res 1998; 50:245–51. 33. Kieffer NM, Burns SJ, Judge NG. Male pseudohermaphroditism of the testicular feminizing type in a horse. Equine Vet J 1976;8:38–41. 34. Kent MG, Shoffner RN, Buoen L, Weber AF. XY sexreversal syndrome in the domestic horse. Cytogenet Cell Genet 1986;42:8–18. 35. Pailhoux E, Cribiu EP, Parma P, Cotinot C. Molecular analysis of an XY mare with gonadal dysgenesis. Hereditas 1995;122:109–12. 36. Abe S, Miyake Y-I, Kageyama S-I, Watanabe G, Taya K, Kawakura K. Deletion of the Sry region on the Y chromosome detected in a case of equine gonadal hypoplasia (XY female) with abnormal hormonal profiles. Equine Vet J 1999;31:336–8. ´ ´ 37. Bugno M, Klukowska J, Słota E, Tischner M, Swito nski M. A sporadic case of the sex-reversed mare (64,XY;SRYnegative): molecular and cytogenetic studies of the Y chromosome. Theriogenology 2003;59:1597–603. ´ 38. Villagomez DAF, Lear TL, Chenier T, Lee S, Cahill J, Reyes E, King WA. Equine XY and XX sex-reversal syndrome in SRY positive and negative mares. In: Proceedings of the 18th International Colloquium on Animal Cytogenetics and Gene Mapping, 8–10 June 2008, University of Agriculture Sciences, Bucharest, Romania. 39. Makinen A, Hasegawa T, Makila M, Katila T. Infertility in two mares with xY and XXX sex chromosomes. Equine Vet J 1999;30:346–9. ´ ´ 40. Swito nski M, Chmurzynska A, Szczerbal I, Lipczynski A, Yang F, Nowicka-Posłuszna A. Sex reversal syndrome (64,XY;SRY-positive) in a mare demonstrating masculine behavior. J Anim Breed Genet 2005;122:60–3. 41. Howden KJ. Androgen insensitivity syndrome in a Thoroughbred mare (64,XY-testicular feminization). Can Vet J 2004;45:501–3. 42. Buoen LC, Zhang TQ, Weber AF, Ruth GR. SRY-negative, XX intersex horses: the need for pedigree studies to examine the mode of inheritance of the condition. Equine Vet J 2000;32:78–81. 43. Vaughan L, Schofield W, Ennis S. SRY-negative XX sex reversal in a pony: a case report. Theriogenology 2001;15;55(5): 1051–7. 44. Bannasch D, Rinaldo C, Millon L, Latson K, Spangler T, Hubberty S, Galuppo L, Lowenstine L. SRY negative 64,XX intersex phenotype in an American Saddlebred horse. Vet J 2007;173:437–9. 45. Temel SG, Gulten T, Yakut T, Saglam H, Kilic N, Bausch E, Jin WJ, Leipoldt M, Scherer G. Extended pedigree with multiple cases of XX sex reversal in the absence of SRY and of a mutation at the SOX9 locus. Sex Dev 2007. 1:24–34. 46. Peiffer DA, Le JM, Steemers FJ, Chang W, Jenniges T, Garcia F, Haden K, Li J, Shaw CA, Belmont J, Cheung SW, Shen RM, Barker D, Gunderson KL. High-resolution genomic profiling of chromosomal aberrations using Infinium whole-genome genotyping. Genome Res 2006;16:1136–48. 47. Ashkenas J. Cytogeneticists embrace molecular karyotyping. Scientist 2007;19:33.
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Chapter 208
Uterine and Clitoral Cultures Sidney W. Ricketts
Dimmock and Snyder1 and Dimmock and Edwards2 concluded that, if correctly performed during estrus, bacteriological examination of the mare’s cervix is a reliable method of detecting uterine infection. They found a good correlation between the presence of uterine infection and infertility. These authors and, in the 1960s, Lord Porchester’s committee,3 drew attention to Klebsiella pneumoniae as a potential cause of equine venereal disease. Both studies concluded that bacteriological examinations of the cervix should be routine aids to equine studfarm preventive medicine, and surveys of equine cervical bacteriological examinations were published from all over the world.4–8 Nevertheless, these recommendations were not universally accepted into routine studfarm practice until the outbreak of contagious equine metritis (CEM), which was first reported in Newmarket during 1977. This new equine venereal disease, which was subsequently diagnosed in many other countries, highlighted the effect that epidemic venereal disease could have, particularly on the international Thoroughbred breeding industries.9, 10 Simpson and Eaton-Evans11, 12 defined the clitoral fossa and sinuses as potential areas of chronic symptomless CEM carrier status, and clitoral swabbing became incorporated into routine studfarm preventive medicine programs. In response, some horse breeding industries formulated ‘codes of practice’ primarily for control of CEM. Where applied quickly and rigorously, these have been highly successful and are an excellent example of veterinary preventive medicine working well. In the UK, the Horserace Betting Levy Board’s (HBLB) Codes of Practice have:
monly in temporary transit quarantine isolation, in horses from mainland Europe destined for Australasia. 2. Reduced the incidence of epidemic venereal disease caused by K. pneumoniae and Pseudomonas aeruginosa (see below). 3. Resulted in improved management and communications within the horse breeding industry. 4. Provided standard operating procedures for the management of reproductive disease control on studfarms.
1. Eradicated CEM (Taylorella equigenitalis has not been isolated in epidemic form in the UK Thoroughbred horse population since January 1986).13 Since 1986 it has been isolated sporadically in imported non-Thoroughbred horses, most com-
1. swabs must be taken from the endometrium, through the open estrous cervix, for results to be meaningful;17 2. guarded swabs collect fewer insignificant contaminant organisms;18
The HBLB Codes of Practice, now including codes for equine herpesvirus-1 and -4 abortion and coital exanthema (EHV-3), equine viral arteritis (EVA), and equine infectious anemia (EIA), and guidelines on strangles (Streptococcus equi) infection, are now in their 32nd year of publication.14 They are reviewed annually by delegates from France, Germany, Italy, Ireland, and the UK and are adopted in similar form in each of these countries. In some continental European countries where similar control measures were not applied in time, CEM has become endemic in many non-Thoroughbred horse populations, now apparently in a more insidious clinical form. The basic principles regarding the pathogenesis of equine acute endometritis were established in the 1960s and 1970s.15, 16 From these principles, it followed that mares should be screened for acute endometritis before mating, in order to protect the stallion and other mares that he mates, and to maximize the potential for successful conception and pregnancy for the individual mare. For the pre-mating screening of mares for acute endometritis it is recognized that:
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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3. concurrent cytological examinations markedly improve diagnostic accuracy;19 4. anaerobic bacteria may play a significant role.20
Screening for acute endometritis Genital swab samples are collected from mares, for microbiological examinations, for the following important reasons: 1. Venereal disease screening. Before the start of breeding operations, swab samples are collected from the vaginal vestibule, clitoral fossa, and sinuses21 at any stage of the estrous cycle, and from the endometrium22 during early estrus. These swab samples are cultured aerobically to screen for the presence of P. aeruginosa and K. pneumoniae (capsule types 1, 2, or 5) and microaerophilically for the presence of T. equigenitalis.23 If these organisms are isolated, the mare is not presented to the stallion until she has been specifically treated and a series of follow-up swab samples (usually three sets taken at appropriate intervals, i.e., 7 days apart for clitoral swabs and during three consecutive estrous periods for endometrial swabs) have confirmed successful elimination. The objective of these screening cultures is to prevent contamination of the stallion’s genitalia by mares that are either symptomless carriers of, or actively infected with, potential venereal disease organisms. 2. Non-venereal acute endometritis screening. Samples are collected before the start of breeding operations and subsequently if and each time the mare returns to estrus after mating. Concurrent endometrial swab and smear samples are collected during early estrus,24 for bacteriological culture22 and cytological examination, screening for the presence [more than the occasional polymorphonuclear leukocyte (PMN)] or the absence (less than the occasional PMN) of acute endometritis.19, 20, 24 These authors classified PMN numbers in endometrial smears as ± (the occasional PMN, i.e., less than 0.5% cells seen), 1 + (small numbers of PMNs, i.e., 0.5–5% cells seen), 2 + (moderate numbers of PMNs, i.e., 5–30% cells seen), and 3 + (large numbers of PMNs, i.e., greater than 30% of cells seen) and use this system for routine reporting of endometrial smear results to clinicians. In addition to routine venereal disease screening, as described above, the swab samples are cultured aerobically to screen for the presence of other potential non-venereal acute endometritis producers (see below). If there are cytological signs of acute endometritis then the bacteriological isolates are examined and appropriate antibiotic treatment is applied.20, 25, 26 The mare is re-examined at the next
estrus with repeat smear and swab examinations to prove successful resolution. If potential venereal disease-producing organisms [T. equigenitalis, P. aeruginosa, K. pneumoniae (capsule types 1, 2 or 5)] are isolated, the mare is managed and treated as described above, irrespective of the cytological results. The objectives of these procedures are: (i) to prevent potential venereal disease carrier mares, or mares with acute infections, from contaminating the stallion’s genitalia; and (ii) routinely to identify mares with non-venereal acute endometritis so that this can be resolved before mating and the chances for successful conception and gestation for the individual mare can be maximized. 3. Examination/investigation of genital abnormalities. Bacteriological examinations are an essential part of the complete gynecological examination of a mare and should be performed in mares who fail to conceive, suffer early fetal death or abortion, show signs of genital abnormality, or are being routinely examined as barren mares after the end of the mating season.25, 27 Swab samples are collected from the vestibule, clitoral fossa, and sinuses at any stage of the estrus cycle and from the endometrium during estrus. If the mare is not in estrus, a uterine aspirate, washing, and/or endometrial biopsy specimen may be collected and cultured. In addition to routine venereal disease screening, as described above, endometrial swab samples are cultured aerobically and sometimes anaerobically to screen for the presence of other potential non-venereal acute endometritis producers (see below).20 At the same time, complete gynecological examinations are performed, including the collection of endometrial biopsy specimens. If there are histological signs of acute endometritis, the bacteriological isolates are examined to make sure that appropriate antibiotic treatment is applied. A follow-up examination is made to assess the response to treatment at the next estrus or, if performed after the mating season, following an appropriate period of rest. If potential venereal disease-producing organisms are isolated then the mare is managed and treated as described above, irrespective of the histological results. The objectives of these procedures are: (i) to prevent potential venereal disease carrier mares, or mares with acute infections, from contaminating the stallion’s genitalia; and (ii) to identify mares with acute endometritis so that this can be resolved to improve the chances for successful conception and gestation for the individual mare, either before the next estrus or, for routine barren mare examinations, before the next mating season.
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Techniques Preparation of the mare For all gynecological examinations, the mare should be bridled and restrained with her hindquarters at a doorframe or, ideally, in stocks, in a manner that facilitates a safe, thorough, and relaxed examination26, 28 (Figure 208.1). For fractious mares, a twitch or sedation may be required. The tail should be bandaged and held to one side. The perineum should be washed with warm, clean water. Disinfectants should not be used before taking clitoral/vestibular swabs. Before collection of endometrial swab samples, the perineum is again thoroughly washed with warm, clean water. Disinfectants are usually unnecessary and, if used regularly and excessively, may encourage the colonization of the clitoral fossa with resistant organisms such as P. aeruginosa and K. pneumoniae.29 Collection of samples Laboratory results are only as good as the samples examined allow them to be.30 Swab samples should be collected with appropriate and careful technique, using suitable equipment (www.rosdale.com/ pages.php?page=000182).
Clitoral/vestibular swabs
Figure 208.2 Standard hospital-type swabs, with Amies charcoal transport medium, used to swab the clitoral sinus (narrowtipped; above) and clitoral fossa (standard-tipped; below) of mares.
and the clitoris is everted. A sterile swab is placed in the ventral aspect of the vestibule to pick up material before thoroughly swabbing all areas of the clitoral fossa (Figure 208.3). A narrow-tipped (pediatric-type) sterile swab is then placed in the central clitoral sinus (Figure 208.4) and, if present, the lateral sinuses, and rotated to pick up smegma. For routine (non-export) screening of mares, many clinicians swab both the clitoral sinus and fossa with a single narrow-tipped swab, in the interests of efficiency and economics. The swabs are immediately placed into the transport medium and clearly labeled with the name of the
Standard hospital-type swabs with Amies charcoal transport medium (Technical Service Consultants Ltd, Heywood, Lancashire, UK) may be used for this purpose (Figure 208.2). With a gloved hand, using the thumb and index finger either side of the ventral commissure of the vulva and with the second finger under the base of the vulva, the vulval lips are parted
Figure 208.1 Endometrial swab/smear being collected, via a sterile disposable speculum lit with a pen torch, from an estrous mare, restrained in stocks with tail bandaged and held to one side following hygienic perineal preparation.
Figure 208.3 A sterile swab is used to sample thoroughly all areas of the mare’s digitally inverted clitoral fossa.
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Figure 208.5 Standard hospital-type swabs, with (for bacteriological culture) and without (for cytological examination) Amies charcoal transport medium, one shown extended with an insemination pipette, and a disposable speculum and penlight torch.
Figure 208.4 A sterile narrow-tipped (pediatric-type) swab is used to sample the mare’s central clitoral sinus.
mare and the date and site of swabbing. It has been suggested that the lateral sinuses may be too shallow to support the growth of T. equigenitalis.31
Non-guarded endometrial swabs Standard hospital-type swabs (Figure 208.5) with Amies charcoal transport medium, extended (with a sterile insemination pipette or purpose-made sterile rod), may be used for non-guarded endometrial swabs, providing concurrent endometrial smear tests are used. A sterile and disposable speculum is passed into the vagina and the cervix is viewed using a penlight torch. If the cervix is sufficiently relaxed, i.e., the mare is in estrus, the extended swab may be carefully passed through the sterile speculum (see Figure 208.1) and then through the relaxed cervix into the uterine body, where it is rotated to pick up secretions.22 The swab is then carefully removed from the uterus, through the cervix and the sterile speculum, and placed immediately into the transport medium. The swabs must be clearly labeled with the mare’s name and the date and site of swabbing. Another swab is taken immediately, in a similar manner, but is not placed into transport medium.
This swab is used for making an endometrial smear for cytological examination.19, 24 Providing concurrent swab and smear samples are collected, this simple and economical technique is perfectly acceptable for useful results and has stood the test of time in clinical practice.
Guarded/semi-guarded swabs Over the years, some clinicians have preferred to use a variety of specially designed swabs using various guarding techniques. These more elaborate, time-consuming, and expensive swabs are passed through the relaxed estrous cervix into the uterine lumen, either via a sterile vaginal speculum as described above or manually, guarded in a sterile gloved hand. Double-gloving and double-guarded techniques have been described.18 Specific technical details depend on the swab used and preferences for speculum or manual passage. Separate transfer to Amies charcoal transport medium is then usually necessary. The objective is to obtain a true endometrial sample, without contamination by vaginal or cervical material, by exposing the swab tip only when it is inside the uterine lumen. The swab is retracted into its sterile sheath before careful removal from the uterus, and is placed immediately into transport medium. The swabs must be clearly labeled with the mare’s name and the date and site of swabbing. Another swab is taken immediately, in a similar manner, but is not placed into transport medium.
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Uterine and Clitoral Cultures This swab is used for making an endometrial smear for cytological examination. Cytological examinations may be performed on material harvested by special brushes incorporated in dual-system devices, in or on the cap or swab sheath of some guarded swabs.
Uterine aspirations and washings If the uterine lumen contains fluid or purulent material, a sample may be aspirated using a sterile insemination pipette, via a sterile vaginal speculum or by manual manipulation, guided if necessary by ultrasonographic examination. Alternatively, and if the uterus does not contain fluid or pus, a uterine washing may be performed. Sterile saline (20–50 mL) may be introduced into the uterus via an insemination pipette, and the end of the pipette may be used to gently dislodge epithelial cells into the lumen before aspirating the fluid.32 Another technique is to place a sterile uterine flush catheter (Har-Vet, Spring Valley, WI, USA; Dairymac Animal Health Products, Wickham, Hampshire, UK) through the cervix and to inflate its bulb just inside the uterine lumen. A 250- or 500-mL pack of sterile saline solution (Arnolds Veterinary Products, Harlescott, Shrewsbury, UK) is then connected to the catheter and the fluid is passed into the uterus by gravity flow. While maintaining the airtight connection between the empty saline bag and the catheter, the uterus may be massaged per rectum, and as the bag is lowered the saline refills the bag, which is then sealed with a clamp. The sedimented washings may be examined bacteriologically and cytologically. A cytocentrifuge (Cytospin 4, Thermo Scientific, Runcorn, Cheshire, UK) provides an excellent sample for cytological staining. If there is to be a delay in processing, the fluid should be swabbed in an aseptic manner and placed into bacteriological transport medium; another portion should be fixed in Cytospin collection fluid (Thermo Scientific, Runcorn, Cheshire, UK) (50:50) for cytological preservation.
Endometrial biopsy Using aseptic techniques, tissue samples may be retrieved from the jaws of biopsy punches (Jørgen Kruuse, Langeskov, Denmark), following endometrial biopsy. If there is to be a delay in processing, the tissue should be carefully swabbed and placed into bacteriological transport medium. Handling of samples The most meticulously collected samples can be rendered useless by improper handling.
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Transport media If there is any delay between swab collection and processing, the use of bacteriological transport medium is essential. In 1977, studies clearly demonstrated that T. equigenitalis is a fragile organism, particularly temperature sensitive, and that bacteriological transport medium was required to prevent these organisms from becoming non-viable during transit to the laboratory and providing false-negative results.10, 11 This organism was also found to be sensitive to light, and thus Amies transport medium, with added charcoal, became the standard all-purpose bacteriological transport medium for equine gynecological practice.
Transport to the laboratory Ideally, there should be minimal delay between swab collection and laboratory processing. In practice, arrangements should be made so that swabs, in transport medium, reach the laboratory with 48 hours of collection, even though studies suggest that longer viability is possible.33 Swabs should be transported in insulated packaging to protect them from extremes of temperature and from direct sunlight. Packaging should be safe and secure to prevent breakages and to satisfy postal regulations. Laboratory techniques Equine gynecological bacteriology is a specialized and demanding subject, which requires experience, care, and attention to detail.23 Important clinical decisions may be made directly on the basis of results, the significance of which should be fully understood by laboratory personnel. The standard of facilities and technical staff must be sufficient to allow accuracy and efficiency, which are essential to provide the clinician with useful results. In UK, the HBLB maintains and publishes a list of designated veterinary laboratories for their Code of Practice scheme by biannual quality assurance testing.14 A plentiful and reliable supply of fresh, goodquality agar plates is essential. Media may be purchased in dehydrated form and made up as required, or as pre-poured plates. Agar plates have a limited shelf life and should be used when as fresh as possible or stored only for short periods. If they are to be stored before use they should be refrigerated and tightly sealed in plastic to prevent drying out. Incubators must be reliable and functioning at the required temperature and atmospheric conditions. Swabs should be inoculated onto agar plates in an environment that is clean and free from sources of plate contamination, using careful standard plating techniques.34
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Aerobic culture For all routine equine gynecological bacteriological examinations, swabs are plated directly onto blood agar (Oxoid Ltd, Basingstoke, Hampshire, UK) and either MacConkey agar (Oxoid Ltd) or cystinelactose-electrolyte deficient (CLED) agar (Oxoid Ltd), using standard laboratory techniques. Both are incubated under normal atmospheric (aerobic) conditions at 37◦ C for up to 48 hours. Cultures are examined at 24 and 48 hours.
Microaerophilic culture For all routine gynecological cultures, swabs are plated onto a modified Eugon agar (Oxoid Ltd) or enriched heated (chocolate) blood agar with CEMO agar base (Lab M or Mast Laboratories), one with and one without the addition of 200 mg/L streptomycin, both with the addition of 5 mg/L amphotericin B. These plates are cultured under microaerophilic conditions for at least 7 days to screen for both the streptomycin-resistant and the streptomycinsensitive strains of T. equigenitalis.35 Swabs can also be plated onto clindamycin/ trimethoprim agar.36 Each swab is inoculated onto an enriched heated (chocolate) blood agar plate containing 5 µg/mL clindamycin, 1 µg/mL trimethoprim, and 5 µg/mL amphotericin. This agar will support the growth of both strains of T. equigenitalis but will eliminate many of the commensal microflora. If T. equigenitalis is isolated, further tests need to be performed to determine the strain type. Microaerophilic cultures are examined at 2, 3, 4, and 7 days. A control strain of T. equigenitalis (provided for HBLB designated laboratories in UK but also available from the National Collection of Type Cultures, Central Public Health Laboratory, Colindale, London, UK) should be cultured in parallel with each batch of swabs to ensure satisfactory media and incubation conditions.
Anaerobic culture For anaerobic culture, we recommend that swabs are directly inoculated onto two plates of Wilkins–Chalgren anaerobe agar (Oxoid Ltd) containing 5% horse blood. One plate contains 70 µg/mL neomycin, which is a good general purpose selective agent for anaerobes. A 5-µg metronidazole disk is placed on each plate in the area of heavy inoculum. Obligate anaerobes are sensitive to metronidazole and so this acts as a valuable aid for discriminating between obligate (grows only under anaerobic conditions) and facultative (may grow under a variety of culture conditions) anaerobic colonies on
both the selective and non-selective plates. These are incubated in an atmosphere of 10% hydrogen, 10% carbon dioxide and 80% nitrogen at 37◦ C for 48 hours. Gas-generating envelopes are available commercially (AnaeroGen, Oxoid Ltd) and are a convenient and reliable method for generating an anaerobic environment. The AnaeroGen sachet is placed in a sealed anaerobic jar or plastic pouch (AnaeroGen Compact, Oxoid Ltd) and will produce an optimal gaseous atmosphere for the growth of anaerobic bacteria within 30 min. Swabs may be incubated in an enrichment medium of Robertson’s meat granules (Lab M, Amersham International, Lancashire, UK) for 24 hours before inoculation onto Wilkins–Chalgren anaerobe agar as described above.
Fungal culture Pathogenic fungi such as Candida spp. and Aspergillus spp. will often grow on blood agar incubated under aerobic conditions. For specific fungal culture, we recommend that swabs are inoculated directly onto plates containing Sabouraud’s dextrose agar (Oxoid Ltd) and incubated under normal atmospheric conditions at 37◦ C for up to 7 days.
Identification Using the cultural conditions recommended above, an experienced technician should be able to identify the majority of equine aerobic pathogens by their colonial appearance on blood and MacConkey agar. To facilitate this, good inoculation technique is required to allow adequate colony separation. For well-collected endometrial swabs, visual identification from the primary inoculum is usually straightforward, but for clitoral swabs, which often grow luxuriant mixed cultures, more detailed investigation is required. Aerobic pathogens P. aeruginosa (Figure 208.6) produces large (3–4 mm) flat gray-green colonies with a characteristic fruity/musty odor, on blood agar. Most strains are hemolytic on blood agar. P. aeruginosa produces large, pale colonies on MacConkey agar (it does not ferment lactose) with a greenish pigment. Gram’s stain reveals Gram-negative rods. P. aeruginosa may be differentiated from P. fluorescens and the other pseudomonads by pyocin production, its failure to grow at 4◦ C, its ability to grow at 41–42◦ C (only P. aeruginosa is able to grow at this temperature) and/or by the use of the API 20NE (bioM´erieux UK) biochemical identification system. Some isolates take 48 hours to grow on blood agar at 37◦ C, only
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Figure 208.6 Pseudomonas aeruginosa growing on blood agar after 24 hours of aerobic culture showing greenish coloration. (See color plate 83).
Figure 208.8 Klebsiella pneumoniae growing on MacConkey agar after 24 hours of aerobic culture showing large, coalescing, mucoid pink colonies. (See color plate 84).
producing small colonies; therefore, all aerobic culture plates should be examined again at 48 hours. Most strains of P. aeruginosa fluoresce under a Wood’s lamp (Figure 208.7). K. pneumoniae (Figure 208.8) produces large, nonhemolytic, mucoid colonies on blood agar after 24 hours of aerobic culture. It grows readily and luxuriantly on MacConkey agar and, being a lactose fermenter, it produces pink colonies that are mucoid. Gram’s stain reveals Gram-negative rods. Isolates should be confirmed by biochemical typing (see below) before being further differentiated by capsular typing (see below). Streptococcus zooepidemicus produces pinpoint colonies, with a clear zone of beta-hemolysis, on blood agar after 24 hours of aerobic culture. It does not grow on MacConkey agar. Gram’s stain reveals Gram-positive cocci. Staphylococcus aureus produces cream, yellow, to gold colonies, sometimes with a
clear zone of beta-hemolysis, on blood agar after 24 hours of aerobic culture. It does grow on MacConkey agar. Gram’s stain reveals Gram-positive cocci. Escherichia coli produces creamy colonies, sometimes with a clear zone of beta-hemolysis, on blood agar after 24 hours of aerobic culture. It grows readily on MacConkey agar and, being a lactose fermenter, it produces pink colonies. Gram’s stain reveals Gram-negative rods.
Figure 208.7 Pseudomonas aeruginosa growing on specialized Pseudomonas agar and showing recognizable fluorescence under a Wood’s lamp.
Figure 208.9 Taylorella equigenitalis growing on enriched hemolyzed blood agar after 4 days of microaerophilic culture. (See color plate 85).
Microaerophilic pathogens All small, pale, translucent colonies are investigated first for their oxidase and catalase activities. Taylorella equigenitalis (Figure 208.9) will produce fast (< 10 s) positive oxidase and catalase reactions. If both are positive, a Gram’s stain is performed. If this reveals
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a small Gram-negative coccobacillus, a phosphatase test is performed to look for a positive reaction. A slide agglutination test is then performed, using rabbit T. equigenitalis antiserum (Monotayl, Bionor, Skien, Norway) (Figure 208.10). Unfortunately,
some other bacteria, for example, Oligella urethralis, will grow on hemolysed blood agar under microaerophilic conditions and may give a positive reaction to the Monotayl test. These organisms can be differentiated from T. equigenitalis by their ability to grow aerobically on ordinary blood agar when subcultured, by the use of more extensive biochemical testing (API 20E, BioM´erieux), and sometimes by their cultural appearance. T. equigenitalis infection is legally notifiable in France, Ireland, and the UK. If T. equigenitalis is isolated in the UK, this must be reported by the isolating laboratory to a Divisional Veterinary Manager (DVM) of the Department for Environment, Food and Rural Affairs (Defra) who will investigate both the isolate and, if reconfirmed, the clinical circumstances further.14 Occasionally, aerobic pathogens, such as P. aeruginosa, will grow under microaerophilic conditions, providing a useful additional ‘screen’. Anaerobic pathogens Obligate anaerobes are differentiated from facultative anaerobes by their growth inhibition around the metronidazole disk. Isolates are purified and identified using standard methods.37 Fungal pathogens Candida spp. produce cream/white, raised, glistening, soft and creamy colonies with a ‘beer-like’ odor after 48 hours of incubation at 37◦ C on blood or Sabouraud’s agar. Aspergillus spp. grow on Sabouraud’s agar. These fungi can be confirmed by examining colony smears mounted in lactophenol cotton blue, and species differentiation can be made using standard methods.38
(a)
(b) Figure 208.10 (a) Monotayl specific rabbit Taylorella equigenitalis antiserum slide agglutination test kit. (b) Monotayl specific rabbit Taylorella equigenitalis antiserum slide agglutination test reaction showing positive agglutination in the test sample, confirming T. equigenitalis, and a negative control reaction.
Subculture Where identification from the primary inoculation is not straightforward, often because mixed cultures or overgrowth of a contaminant non-pathogen has occurred, purification by subculture is essential. A single well-isolated colony should be transferred onto a solid non-selective medium identical to that on which the original isolation was made, and culture should be performed under identical temperature and atmospheric conditions. Gram’s stain Gram’s stain is performed using standard methods.38 For reliable results, the culture smear must not be too thick and the Gram-positive organisms must not be over-decolorized with acetone. Biochemical tests Commercially available, prepacked biochemical reaction systems are widely used in clinical bacteriology
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and examined under a phase-contrast microscope. The absorption of specific antibody onto the polysaccharide of the organism’s capsule results in a change in the refractive index of the capsule, causing it to appear to swell and darken. Immunofluorescent staining with specific fluorescein isothiocyanate (FITC) labeled antisera39 may also be used. First, the isolate must be subcultured for purity and then grown on Worfel–Ferguson medium (Difco Ltd) to swell the capsules. Capsular types 1, 2, 4, 7, and 68 are most commonly isolated from equine genital swab samples,39 and only types 1, 2, and 5 have been isolated in association with outbreaks of true venereal disease.40 P. aeruginosa serotypes may be differentiated but no correlation has been found between serotype and potential venereal pathogenicity.41
Polymerase chain reaction (PCR) tests
Figure 208.11 Examples of prepacked biochemical reaction systems (API, bioM´erieux UK) for the identification of bacterial isolates.
for their accuracy, ease, and convenience of use (Figure 208.11). Systems are available for differentiation of Gram-negative rods (API 20E, API bioM´erieux, Vercieu, France), streptococci (Streptococcal Grouping Kit, Oxoid Ltd), and staphylococci (API Staph, API bioM´erieux). These systems are simple to use and provide excellent results if manufacturers’ instructions are followed meticulously. Clearly, it is essential that only single colonies or pure cultures are used to inoculate reaction compartments or spurious results will be produced. The API 20E system is designed for results to be interpreted with the aid of an ‘API Quick Index’, which lists over 900 profiles and is sufficient for most cases. It differentiates some 51 bacteria on a plus/minus scoring system. When this is insufficient, users have free telephone access to the API 20E computer service. Capsule/serotyping K. pneumoniae capsule types may be differentiated by the quellung test, which involves capsular swelling. Firstly, the isolate is grown on a high-carbohydratecontaining agar in order to enhance capsule production. A suspension of organisms in saline solution is then mixed with dilute capsule-specific antiserum
A real-time PCR test for T. equigenitalis is now commercially available (Qiagen UK) and an extensive field trial conducted by Beaufort Cottage Laboratories42 has shown complete concordance of results between microaerophilic culture and PCR results for over 2000 mare and stallion genital swab samples. The PCR test differentiates T. equigenitalis from T. asinigenitalis and O. urethralis. It detects T. equigenitalis DNA in swab samples where the bacteria have been inactivated by time delay or temperature stress. No ‘false positive’ results were obtained. In addition, the main advantage for the horse breeding industries will be a faster turnround time (24–48 hours for PCR as compared with at least 7 days for negative confirmation by conventional microaerophilic culture). The shorter testing time means that, stallion managers will have PCR results for T. equigenitalis alongside aerobic culture results for K. pneumoniae and P. aeruginosa. This will be very helpful, particularly for ‘walking in’ mares and for semen shipped for artificial insemination (AI), and the test will overcome many difficulties regarding microaerophilic cultures. This PCR test appears robust enough to be used as a rapid screening test for T. equigenitalis. Positive PCR screening test results will need to be investigated by conventional microaerophilic cultures, and a positive result in the UK is reportable to Defra under the law.14
Antibiotic sensitivity tests Specialized agar medium (Sensitest Agar, Oxoid Ltd) should be used to overcome sulfonamide and trimethoprim antagonists, and fresh blood need be added only for particularly fastidious organisms. Selective media should not be used for antibiotic sensitivity tests.
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One medium-sized colony should be emulsified in 5 mL sterile medium, distilled water, or nutrient broth; a swab is dipped into this and drained before being evenly spread over the agar plate. Tests run on direct inocula may give next-day, but spurious, results, unless the culture is pure and the inoculum size is satisfactory. Standard or personalized antibiotic-impregnated disks can be used; the choice depends on clinical preferences and antibiotic availability. Although eight disks are commonly used per plate, six may be preferable to avoid close proximity interactions between antibiotic diffusion in agar leading to misinterpretation. These tests are qualitative and not quantitative, and zone sizes cannot be correlated with degree of sensitivity or insensitivity. Disk sensitivity tests run on organisms that require more than 48 hours for visible growth, such as T. equigenitalis, are not practical.
Figure 208.13 Endometrial smear (Pollack’s Trichrome, ×40) from an estrous mare with acute endometritis demonstrating degenerate polymorphonuclear leukocytes (PMNs) within stringy mucous and endometrial epithelial cells. (See color plate 87).
Interpretation of results
Laboratory results
It is the clinician’s responsibility to interpret the results provided, aided, where necessary, by the diagnostic laboratory. Interpretation is based not only on the organisms isolated but also on clinical history of the mare and results of other gynecological examinations. During the last 30 years, concurrent endometrial cytological examinations have been routinely used to improve the accuracy of interpretation of routine pre-mating endometrial bacteriological results19, 20, 24 (Figures 208.12 and 208.13). It is now recognized that without concurrent cytopathological data, the interpretation of the significance of non-venereal bacteria isolated from endometrial swab samples taken by whatever technique is pure guesswork.
Table 208.1 presents the aerobic and microaerophilic bacterial species isolated from 1000 routine equine clitoral swabs cultured at Beaufort Cottage Laboratories during 1990.43 Table 208.2 presents the T. equigenitalis, K. pneumoniae, and P. aeruginosa isolates obtained from 1025 clitoral swabs processed at Beaufort Cottage Laboratories during 2007. No T. equigenitalis isolates were made from routine clitoral swabs submitted during either of these studies. K. pneumoniae was isolated from 0.6% and 0.5% of clitoral swabs, and P. aeruginosa was isolated from 1.0%
Figure 208.12 Endometrial smear (Pollack’s Trichrome, ×40) from a normal estrous mare with sheets of normal endometrial epithelia cells and no leukocytes. (See color plate 86).
Table 208.1 Aerobic and microaerophilic bacterial species isolated from 1000 routine equine clitoral swabs processed at Beaufort Cottage Laboratories during 199043
Bacteria isolated
Number of samples
Percent
Taylorella equigenitalis Klebsiella pneumoniae Pseudomonas aeruginosa Escherichia coli Streptococcus faecalis Corynebacterium spp. Staphylococcus aureus Beta-hemolytic streptococci Staphylococcus albus Acinetobacter spp. Bacillus spp. Enterobacter aerogenes Pasteurella spp. Proteus spp. Alpha-hemolytic streptococci Non-hemolytic streptococci Citrobacter spp.
0 6 10 724 318 228 202 172 106 86 30 7 6 6 6 6 1
0 0.6 1.0 72.4 31.8 22.8 20.2 17.2 10.6 8.6 3.0 0.7 0.6 0.6 0.6 0.6 0.1
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Uterine and Clitoral Cultures Table 208.2 Aerobic and microaerophilic bacterial species isolated from 1025 routine equine clitoral swabs processed at Beaufort Cottage Laboratories during 2007
1973
Table 208.4 Bacterial and fungal species isolated following aerobic culture of 14 874 equine endometrial swab samples at Beaufort Cottage Laboratories (2002–07)
Bacteria isolated
Number of samples
Percent
Bacteria isolated
Number of samples
Percent
Taylorella equigenitalis Klebsiella pneumoniae Pseudomonas aeruginosa No significant growtha
0 5 2 1081
0 0.5 0.2 99.32
Beta-hemolytic streptococci Escherichia coli Staphylococcus aureus Alpha-hemolytic streptococci Coagulase-negative staphylococci Non-hemolytic streptococci Acinetobacter spp. Bacillus spp. Streptococcus faecalis Proteus spp. Klebsiella pneumoniae Pseudomonas aeruginosa Candida spp. Enterobacter agglomerans Enterobacter aerogenes Aspergillus spp. Klebsiella oxytoca Pasteurella spp. Corynebacterium spp. Serratia spp.
3006 1183 531 231 91 46 46 44 33 24 17 12 10 8 6 5 3 2 1 1
20.21 7.95 3.57 1.55 0.61 0.31 0.31 0.30 0.22 0.16 0.11 0.08 0.07 0.05 0.04 0.03 0.02 0.01 0.01 0.01
a For the purposes of this table, no significant growth means no growth of T. equigenitalis, K. pneumoniae, or P. aeruginosa.
and 0.2% of clitoral swabs during 1990 and 2007 respectively. Tables 208.3 to 208.6 present the results of examinations of 14 874 sets of concurrently collected endometrial swab and smear samples19, 20 at Beaufort Cottage Laboratories during the years 2002 to 2007. Microaerophilic culture was performed on all of these swabs and no isolates of T. equigenitalis were made. Table 208.3 shows that no organisms were cultured from 71% of these 14 874 endometrial swab samples, 23% cultured a pure growth of bacteria or fungi, and 6% cultured a mixed growth of bacteria or fungi. Although non-guarded swabbing techniques were used similar to those for which results (1973–89) were reported in 1993,43 there had been a marked increase in the no-growth results (39% to 71%). Laboratory handling and culture techniques have not changed significantly between these periods. The 1973–89 results included swab samples referred to Beaufort Cottage Laboratories from many veterinary practices through the postal services whereas the 2002–07 samples were paired swab and smear samples collected by Rossdale & Partners clinicians and therefore not subjected to postal delays. Also, closer investigation of the 2002–07 results reveals a trend of annual increase in no-growth results, from 60% in 2002 to 75% in 2007, which is statistically significant (p > 0.01). Therefore, improvements in the standard of swabbing technique and/or the gynecological status of the mare population examined may have occurred between these two periods. Table 208.3 Bacterial growth obtained from 14 874 equine endometrial swab samples at Beaufort Cottage Laboratories (2002–07)
Bacteria isolated No growth Mixed growth Pure growth
Number of samples 10 497 885 3492
Table 208.4 shows the species of bacteria and fungi isolated from these 14 874 endometrial swab samples. As in the 1973–89 series, beta-hemolytic streptococci (20% vs 18% previously), Escherichia coli (8% vs 15% previously) and Staphylococcus aureus (4% vs 9% previously) were the prominent isolates. Alphahemolytic streptococci were the most frequently isolated ‘non-pathogens’ (2% vs 4% previously). Significance of isolates Bacteriological examinations of endometrial swab samples, taken with whatever equipment and by whatever techniques, cannot be accurately interpreted without concurrent endometrial cytological data.19 Table 208.5 shows that of 14 874 pairs of concurrently taken endometrial swab and smear Table 208.5 Acute endometritis diagnoses (more than small numbers of PMNs, i.e., more than 5% of cells seen) made following cytological examination and bacterial culture of 14 874 pairs of equine endometrial swab and smear samples at Beaufort Cottage Laboratories (2002–07)
Percent 70.57 5.95 23.48
Diagnosis
Number of samples
Percent
Acute endometritis No acute endometritis
1242 13 632
8.4 91.6
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Table 208.6 Bacterial and fungal species isolated following culture of 1242 equine endometrial swab samples with cytological evidence of acute endometritis (more than small numbers of PMNs, i.e., more than 5% of cells seen) at Beaufort Cottage Laboratories (2002–07)
Bacteria isolated
Number of samples
Percent
No growth of bacteria or fungi Bacterial and/or fungal isolates Beta-hemolytic streptococci Escherichia coli Staphylococcus aureus Alpha-hemolytic streptococci Coagulase-negative staphylococci Acinetobacter spp. Non-hemolytic streptococci Klebsiella pneumoniae Pseudomonas aeruginosa Bacillus spp. Streptococcus faecalis Proteus spp. Candida spp. Enterobacter agglomerans Enterobacter aerogenes Klebsiella oxytoca Pasteurella spp. Aspergillus spp.
360 882 701 183 123 31 9 7 5 4 3 3 3 3 3 1 1 1 1 1
28.9 71.1 56.4 14.7 9.9 2.5 0.7 0.6 0.4 0.3 0.2 0.2 0.2 0.2 0.2 0.1 0.1 0.1 0.1 0.1
samples, only 1242 (8%) showed cytological evidence of acute endometritis (more than small numbers of PMNs, i.e., more than 5% of cells seen). In the 1973–89 series, 21% of the 16 386 paired swab and smear samples revealed cytological evidence of acute endometritis, suggesting improvements in the gynecological status of the mare population examined. Table 208.6 shows the bacterial and fungal isolates from 1242 pairs of endometrial swab and smear samples with cytological evidence of acute endometritis (more than small numbers of PMNs, i.e., more than 5% of cells seen). Three hundred and sixty swabs (29%), with concurrent cytological evidence of acute endometritis, revealed no aerobic or microaerophilic growth of organisms. Anaerobic bacterial cultures were not routinely performed for these swabs. From the 882 swabs (71%) with bacterial isolates, beta-hemolytic streptococci (56%), E. coli (15%), and Staph. aureus (10%) were the most frequent isolates associated with acute endometritis, supporting the opinion that these are the three most important non-venereal potential pathogens of the equine uterus. K. pneumoniae (n = 4; 0.3%) and P. aeruginosa (n = 3; 0.2%) were isolated infrequently from mares with cytological evidence of acute endometritis. Table 208.4 shows that K. pneumoniae (n = 17; 0.11%) and P. aeruginosa (n = 12; 0.08%) were isolated less fre-
quently from mares overall than in the 1973–89 series (n = 192; 0.6% and n = 99; 0.3% respectively).43 This is believed to reflect the success of the HBLB Codes of Practice in reducing the incidence of acute endometritis caused by these two potential equine venereal disease pathogens, by encouraging routine pre-mating clitoral and endometrial screening. This conclusion is also supported by a comparison of Tables 208.1 and 208.2, where the percentage isolation of K. pneumoniae and P. aeruginosa in approximately 1000 routine clitoral swabs collected in 1990 and 2007 has decreased from 0.6% to 0.5% and 1.0% to 0.2% respectively. It is clear that any bacterial species may be isolated in association with acute endometritis in individual mares but, in general terms, some bacterial species may be more likely to be opportunist pathogens than others and may therefore be classified as outlined below.
Potential venereal disease-producing organisms Only T. equigenitalis, P. aeruginosa, and K. pneumoniae (capsule types 1, 2, and 5) have been isolated as causes of true venereal disease in mares. True equine venereal disease is defined by infection of the stallion with a specific pathogen by an infected or clitoral carrier mare, followed by production of epidemic acute endometritis caused by the same pathogen in subsequent mares mated by this stallion. Infected mares have normal or abnormal local uterine immunological defense mechanisms. If potential venereal disease-producing bacteria, i.e., T. equigenitalis, P. aeruginosa, and K. pneumoniae (capsule types 1, 2, or 5), are isolated from either vestibular/clitoral swabs or from endometrial swabs, with or without concurrent cytological evidence of acute endometritis, the mare in question should be identified as being unsuitable for natural mating until treated and proven free of infection or carrier status. Two strains of T. equigenitalis, i.e., streptomycinsensitive and streptomycin-resistant strains, have been isolated from mares with venereal acute endometritis. The sensitive strain was first isolated in Newmarket in 1977, causing copious vaginal discharge, and the hitherto unknown disease was called contagious equine metritis (CEM). The resistant strain was subsequently isolated in Kentucky but has since been isolated from non-Thoroughbred horses imported into the UK from mainland, principally eastern, Europe. Both strains appear capable of producing epidemic venereal disease in mares. When first diagnosed in Newmarket in 1977, the organism produced severe acute endometritis, with most mares developing copious vaginal discharge. It
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Uterine and Clitoral Cultures is now reported that the disease, which appears to be endemic in many non-Thoroughbred horse populations of mainland Europe, is much more insidious, producing minimal clinical signs, although the organism itself is indistinguishable from that originally isolated. Towards the end of 2008, T. equigenitalis was isolated in Kentucky, and further investigations revealed predominantly non-Thoroughbred positive contacts. A bulletin from the US Department of Agriculture (USDA) on 8 May 2009 reported that the outbreak involved 820 positive and ‘exposed’ horses in 48 states, of which 18 stallions in six states and five mares in three states had been confirmed positive. Up to May 2009, all isolates were of the streptomycinresistant strain and at least half were noted to have had a vulvar discharge. Four experimentally exposed mares also displayed evidence of a cervicitis, vaginitis, and a vulvar discharge (P. Timoney, personal communication). All strains of T. equigenitalis should be considered potential venereal pathogens, and if isolated, the following standard procedures should be adopted: 1. isolation should be notified to the appropriate authority (in the UK the isolation of T. equigenitalis is notifiable by law to Defra14 ); 2. all sexual contacts should be traced and investigated; 3. mares with isolates should be treated and proven to be clear from both infection and a symptomless carrier state before they are presented again for mating. A distinct but related organism, Taylorella asinigenitalis, has been isolated from donkeys and horses without recognizable clinical signs of infection.44 The two organisms must be differentiated by specific PCR testing.45 Different serotypes of P. aeruginosa have been isolated from horses but no reliable association with venereal pathogenicity has been identified.41 Although it is recognized that some isolates may not behave as true venereal pathogens, mechanical contamination of the stallion’s penile and preputial smegma may lead to infection of susceptible mares, and treatment can be difficult, owing to the resistance of this organism to antimicrobial drugs and to its ability to reappear following apparent clearance. Therefore, no alternative exists but to consider all isolates to be potential venereal pathogens and to take appropriate action, as described above. Experience suggests that small, slow-growing colonies (see above) are not isolated under circumstances where true venereal disease occurs, but this is not recommended as a prognostically reliable characteristic.
1975
Different capsular types of K. pneumoniae have been isolated from horses and have been correlated with potential venereal pathogenicity.40 Capsular types 1, 2, and 5 have been isolated in association with true epidemic venereal disease in horses; therefore, these capsular types should be considered potential venereal pathogens and appropriate action should be taken, as described above. Capsular types 7 and 68, among others, are occasionally isolated from penile, clitoral, and fecal swabs but have not been isolated in association with true epidemic venereal disease in horses and, therefore, should not be considered potential venereal pathogens.
Non-venereal ‘opportunist’ acute endometritis-producing organisms The external genitalia of the normal mare, including the vestibular and clitoral areas, irrespective of age and reproductive status, possess a mixed aerobic (see Table 208.1) and obligate anaerobic microflora.20 Isolation of bacteria other than the potential venereal disease-producing bacteria (see Table 208.2) discussed above, from these sites is, therefore, of no significance. It has been stated that the normal equine uterus is bacteriologically sterile or has a temporary, nonresident, rather than a resident microflora.46 Studies have since shown that even in maiden mares with no cytological evidence of acute endometritis, obligate anaerobic bacteria can inhabit the uterus as surface commensals.20 Even using double-guarded occluded swabs, following repeated external genital scrubbing with disinfectants, bacteria were isolated from 15 out of 48 (31%) endometrial swabs taken from mares with no evidence of acute endometritis.47 During the breeding season, parturition and then natural mating results in repeated contamination of the uterine lumen with environmental and external genital microflora. Transient endometritis is, therefore, an inevitable sequel to parturition and coitus, and at those times the normal, genitally healthy mare produces an efficient transient acute endometritis, which resolves within 48 to 72 hours.15, 16 Mares with genital abnormality, for example, pneumovagina or vulval, rectovaginal, or cervical injury, or mares that have impaired local immune mechanisms, produce an inefficient, persistent acute endometritis. Thus non-venereal ‘opportunist’ endometritis is more a reflection of the individual mare’s genital abnormalities than the microbial organisms isolated. Virtually any bacterial species may be involved, but experience suggests that the aerobes (or more correctly, facultative anaerobes) beta-hemolytic streptococci, E. coli and Staph. aureus (see Tables 208.4 and 208.6) and the anaerobe Bacteroides fragilis20 are the most commonly
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isolated species, both in pure and mixed cultures, from mares with acute endometritis. All species may be isolated from endometrial swab samples either as significant acute endometritis producers or as insignificant contaminants or epithelial surface commensals. Thus for accurate interpretation, clinical (vaginal discharge, cervicitis, uterine fluid pooling, pyometritis) and/or concurrent endometrial cytological data (active PMN with endometrial epithelial cells) must be used to determine significance.19
Contaminant/commensal organisms Contaminant/commensal organisms are most commonly the normal microflora of the external genitalia and perineal skin. In effect this means all aerobic bacterial isolates other than T. equigenitalis, P. aeruginosa, and K. pneumoniae. Contaminants/commensals are of no primary significance when isolated from clitoral/vestibular swab samples but may, of course, become opportunist uterine pathogens, as discussed above. Acute endometritis associated with these organisms can only be ruled out on the basis of the results of concurrent endometrial cytological data.19 Correlation with concurrent endometrial cytological examinations A survey of concurrent endometrial cytological and aerobic bacteriological examinations performed on mares under routine studfarm practice conditions during the course of four breeding seasons19 showed that bacteria were isolated from 65% of the (nonguarded) swabs examined. A total of 18% of mares had cytological evidence of acute endometritis, comprising 10% who had bacteriological isolates that would have traditionally been interpreted as significant (on the basis of growth purity, intensity, and species isolated), 4% who had non-significant bacteriological isolates, and 4% who yielded no bacterial growth. Without concurrent cytological data, the latter 4% would have gone undetected. Conversely, 13% of the mares would have been falsely interpreted as having significant bacterial isolates when concurrent cytological results revealed no evidence of acute endometritis. Another, smaller survey that included anaerobic bacterial cultures performed on mares under similar conditions20 isolated bacteria from 61% of the swabs examined. In that study, 20% of mares had cytological evidence of acute endometritis, comprising 8% who had mixed aerobic and anaerobic isolates, 5% who had anaerobic isolates alone, and 2% who yielded no bacterial growth. Table 208.6 suggests that no aerobic or microaerophilic growth of organisms is obtained in as many as 29% of mares with concurrent cytological evidence of acute endometri-
tis. These mares would not be diagnosed as having acute endometritis without the benefit of smear tests and their fertility potential, when mated, would be jeopardised. This study showed that from the 71% of swabs with bacterial isolates, beta-hemolytic streptococci (56%), E. coli (15%), and Staph. aureus (10%) were the most frequent isolates associated with acute endometritis. All these findings and the now considerable field experience of using these techniques in practice have confirmed that for diagnosis of non-venereal acute endometritis in mares, cytological rather than bacteriological data are of primary importance.
Conclusions Routine bacteriological examinations should be performed as follows.
1. Vestibular/clitoral swabs should be collected before breeding commences at the start of each season to screen for potential venereal disease carrier status, with aerobic and microaerophilic cultures, screening specifically for T. equigenitalis, P. aeruginosa, and K. pneumoniae (capsule types 1, 2, and 5), in order that such mares are not presented to the stallion for natural mating. Vestibular/clitoral swabs should also be taken from sexual contacts if any of these organisms are subsequently isolated from mares after breeding has commenced or if there is clinical evidence of epidemic acute endometritis, or as part of the complete gynecological examination of a barren mare after the end of the breeding season. Although CEM has been eradicated in most Thoroughbred horse populations, the organism is still found in the non-Thoroughbred horse populations of many mainland European countries and in the USA, and vigilant screening must be maintained internationally to protect equine fertility and export/import movement of horses. 2. Concurrent endometrial swabs and smears should be collected during estrus before breeding commences, and at each subsequent estrous period where conception has not occurred, to screen for acute endometritis, in order that such cases can be treated and resolved before mating to maximize the chances for conception and normal pregnancy for individual mares. The diagnosis of acute endometritis should be made on the basis of cytological evidence from endometrial smear samples and not solely on the basis of bacterial isolates (with the exception of T. equigenitalis, P. aeruginosa, and K. pneumoniae). Bacterial isolates should be used to maintain screening for the potential venereal
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Uterine and Clitoral Cultures disease-producing organisms and to make sure that when acute endometritis is diagnosed, appropriate treatment is applied, with appropriate antibiotic sensitivity testing. 3. Endometrial aspirates, flushings, and/or biopsies should be collected and cultured under aerobic, microaerophilic, and anaerobic conditions and examined mycologically, in addition to being examined cytologically and/or histologically, in cases of diagnosed uterine abnormality (rectal palpation, ultrasonographic examination) and as part of the complete gynecological examination of a barren mare after the end of the breeding season. 4. Stallion managers should require negative clitoral and endometrial cultures for T. equigenitalis, P. aeruginosa, and K. pneumoniae (capsule types 1, 2, and 5) before they initially accept a mare for natural mating. Provided that these organisms have been excluded by certification, ‘clean smears’ should satisfy their requirement for ‘clean swabs’.47 There is little doubt that true epidemic equine venereal disease associated with T. equigenitalis, P. aeruginosa, and K. pneumoniae is now rare in comparison to the 1960s and 70s, and that standards of mare swabbing and the gynecological management of mares have improved, resulting in a reduced incidence of opportunist acute endometritis. This must be at least partly associated with the beneficial effects of the HBLB’s Codes of Practice,14 which are updated and republished annually and have become standard operating procedures for equine breeding and veterinary management.
Acknowledgements I am indebted to my colleagues at Beaufort Cottage Laboratories, particularly Robert Cash, Kath Gifkins, Kevin Grimes, Lorraine Palmer and Adam Fletcher, not only for their technical skills but also for help with the data collection for this article.
References 1. Dimmock WW, Snyder EM. Bacteria of the genital tract of mares and the semen of stallions and their relation to breeding efficiency. J Am Vet Med Assoc 1923;64:288–98. 2. Dimmock WW, Edwards PR. Pathology and bacteriology of the reproductive organs of mares in relation to sterility. In: Research Bulletin of the Kentucky Agricultural Experimental Stallion, Lexington. No. 286, 1928. 3. Porchester, Lord. Uterine infections in mares. Vet Rec 1965;77:110–11. 4. Farrely BY, Mullaney PE. Cervical and uterine infections in Thoroughbred mares. Ir Vet J 1964;18:201–12. 5. Collins SM. A study of the incidence of uterine infections in Thoroughbred mares in Ireland. Vet Rec 1964;76:673–6.
1977
6. Bain AM. The role of infection in fertility in the Thoroughbred mare. Vet Rec 1966;78:168–75. 7. Elliott REW, Callaghan EJ, Smith BL. The microflora of the cervix of the Thoroughbred mare: A clinical and bacteriologic survey in a large practice in Hastings. NZ Vet J 1971;199:291–302. 8. Scott P, Daley P, Baird GG, Sturgess S, Frost AJ. The aerobic bacterial flora of the reproductive tract of the mare. Vet Rec 1971;88:58–61. 9. Platt H, Atherton JG, Simpson DJ, Taylor CE, Rosenthal RO, Brown DF, Wreghitt TG. Genital infections in mares. Vet Rec 1977;101:20. 10. David JSE, Frank CJ, Powell DG. Contagious metritis 1977. Vet Rec 1977;101:189–90. 11. Simpson DJ, Eaton-Evans WE. Developments in contagious equine metritis. Vet Rec 1978;102:19–20. 12. Simpson DJ, Eaton-Evans WE. Sites of CEM infection. Vet Rec 1978;102:488. 13. Meldrum KC. The UK Contagious Equine Metritis (C.E.M.). Crown Copyright BL 5888. Tolworth, UK: Ministry of Agriculture, Fisheries and Food, 1989. 14. Horserace Betting Levy Board. Codes of Practice on Contagious Equine Metritis (CEM), Klebsiella pneumoniae, Pseudomonas aeruginosa; Equine Viral Arteritis (EVA); Equine Herpesvirus (EHV); Equine Infectious Anaemia (EIA) and Guidelines on Strangles. London: HBLB, 2010 (www.hblb.org.uk/sndFile.php?fileID=58). 15. Peterson FB, McFeely RA, David, JSE. Studies on the pathogenesis of endometritis in the mare. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 1969; pp. 279–82. 16. Hughes JP, Loy RG. The relation of infection to infertility in the mare and stallion. Equine Vet J 1974;7:155–9. 17. Greenhof GR, Kenney RM. Evaluation of reproductive status of nonpregnant mares. J Am Vet Med Assoc 1975;167:449–58. 18. Blanchard TL, Cummings MR, Garcia MC, Hurtgen JP, Kenney RM. Comparison of two techniques for obtaining endometrial bacteriologic cultures in the mare. Theriogenology 1981;16:85–93. 19. Wingfield Digby NJ, Ricketts SW. Results of concurrent bacteriologic and cytologic examinations of the endometrium of mares in routine stud farm practice 1978–1981. J Reprod Fertil Suppl 1982;32:181–5. 20. Ricketts SW, Mackintosh ME. Role of anaerobic bacteria in equine endometritis. J Reprod Fertil Suppl 1987;35:343–51. 21. Ricketts SW. Contagious equine metritis. Equine Vet Educ 1996;8:166–70. 22. Ricketts SW. Bacteriological examinations of the mare’s cervix: Techniques and interpretation of results. Vet Rec 1981;108:46–51. 23. Mackintosh ME. Bacteriological techniques in the diagnosis of equine genital infections. Vet Rec 1981;108:52–5. 24. Wingfield Digby NJ. Studies of endometrial cytology in mares. Equine Vet J 1978;10:167–70. 25. Ricketts SW. The barren mare. Diagnosis, prognosis, prophylaxis and treatment for genital abnormality. Part 1. In Practice 1989;11:119–25. 26. Ricketts SW. The treatment of equine endometritis in studfarm practice. Pferdeheilkunde 1999;6:588–93. 27. Ricketts SW. The barren mare. Diagnosis, prognosis, prophylaxis and treatment for genital abnormality. Part 2. In Practice 1989;11:156–64. 28. Rossdale PD, Ricketts SW. Equine Studfarm Medicine, 2nd edn. London: Bailli`ere Tindall, 1980.
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29. Ricketts SW. Klebsiella aerogenes in mares. Vet Rec 1976;99:489–90. 30. Ricketts SW. The laboratory as an aid to clinical diagnosis. Vet Clin N Am Equine Pract 1987;3:445–60. 31. McAllister RA, Sack WO. Identification of anatomical features of the equine clitoris as potential growth sites of Taylorella equigenitalis. J Am Vet Med Assoc 1990;196:1965–6. 32. Freeman KP, Johnston JM. Collaboration of a cytopathologist and practitioner using equine endometrial cytology in a private broodmare practice. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 1987, pp. 629–39. 33. Atherton JG. Isolation of the C.E.M. organism. Vet Rec 1978;102:67. 34. Cowan ST, Steel KJ. Manual for the Identification of Medical Bacteria, 2nd edn. Cambridge: Cambridge University Press, 1974. 35. Taylor CED, Rosenthal RO, Brown DF, Lapage SP, Hill LR, Legros RM. The causative organism of contagious equine metritis, 1977 – Proposal for a new species to be known as Haemophilus equigenitalis. Equine Vet J 1978;10:136–44. 36. Timoney PJ, Shin SJ, Jacobson RH. Improved selective medium for isolation of Contagious Equine Metritis organism. Vet Rec 1978;111:107–8. 37. Willis AT. Anaerobic Bacteriology: Clinical and Laboratory Practice, 3rd edn. London: Butterworths, 1977. 38. Cottral GE (ed.). In: Manual of Standardized Methods for Veterinary Microbiology. Ithaca: Cornell University Press, 1978, pp. 583–7. 39. Riser E, Noone P, Bonnet ML. A new serotyping method for Klebsiella species: Development of the technique. J Clin Pathol 1976;29:305–8.
40. Platt H, Atherton JG, Ørskov I. Klebsiella and Enterobacter organisms isolated from horses. J Hyg Camb 1977;77: 401–8. 41. Atherton JG, Pitt TL. Types of Pseudomonas aeruginosa isolated from horses. Equine Vet J 1982;14:329– 32. 42. Ousey JC, Palmer L, Cash RSG, Grimes KJ, Fletcher AP, Barrelet A, Foote AK, Manning FM, Ricketts SW. An investigation into the suitability of a commercial realtime PCR assay to screen for Taylorella equigenitalis in routine prebreeding equine genital swabs. Equine Vet J 2009;41:878–882. 43. Ricketts SW, Young A, Medici EB. Uterine and clitoral cultures. In: McKinnon AO, Voss JL (eds) Equine Reproduction, 1st edn. Lea & Febiger, 1993; pp. 234–45. 44. Jang SS, Donahue JM, Arata AB, Goris J, Hansen LM, Earley DL, Vandamme PA, Timoney PJ, Hirsh DC. Taylorella asinigenitalis sp. nov., a bacterium isolated from the genital tract of male donkeys (Equus asinus). Int J Syst Evol Microbiol 2001;51:971–6. 45. Wakeley PR, Errington J, Hannon S, Roest HI, Carson T, Hunt B, Sawyer J, Heath P. Development of a real time PCR for the detection of Taylorella equigenitalis directly from genital swabs and discrimination from Taylorella asinigenitalis. Vet Microbiol 2006;118:247– 54. 46. Woolcock JB. Equine bacterial endometritis. Vet Clin N Am Large Anim Pract 1980;2:241–51. 47. Hinrichs K, et al. Bacteria recovered from the reproductive tracts of normal mares. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners 1989; pp. 11–16.
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Chapter 209
Use of Chromogenic Agar to Diagnose Reproductive Pathogens Angus O. McKinnon and David P. Beehan
Introduction The traditional approach to the detection of bacteria associated with the reproductive tract has been to culture either the mare or stallion and then inoculate one or more general-purpose media such as horse blood agar (HBA) and MacConkey agar, followed by aerobic incubation at 37◦ C for 24–48 h. Additional culturing techniques are used for anaerobic1 and microaerophillic pathogens.2, 3 Identification of the suspect pathogens has been based on colony size, morphology, pigmentation, hemolytic patterns, and Gram stain, which rarely permit more than a presumptive identification. Biochemical and/or serological tests are then used for definitive diagnosis. This approach frequently necessitates the testing of commensal bacteria that resemble pathogens,4 which is often more expensive and labor intensive. This is especially true in a veterinary practice wishing for accurate and expedient bacterial diagnoses. Since 1990, a wide range of chromogenic culture media has been made commercially available, providing useful tools for diagnostic clinical microbiology. These chromogenic substrates release specific colored dyes when hydrolyzed by enzymes of pathogenic microorganisms, thus resulting in readily identifiable colored colonies.5 The principle of chromogenic agar is the use of substrates incorporated into the agar that reveal genus- or speciesspecific enzyme activities of microorganisms.6 By the inclusion of chromogenic enzyme substrates targeting microbial enzymes, these media are able to target certain pathogens with high specificity. The inclusion of multiple chromogenic substrates into culture
media facilitates the differentiation of polymicrobial cultures, thus allowing for diagnosis of clitoral sinus cultures that, if taken correctly, should be highly contaminated with multiple different colonies. Selective chromogenic medium was first used in 1976 for the direct identification of Escherichia coli in the primary culture of urine.7 Since then, a wide range of chromogenic culture media has been developed for use in diagnostic clinical microbiology. For example, in human medicine a chrome agar has been developed that allows the rapid and highly specific identification of methicillin-resistant Staphylococcus aureus (MRSA). A positive result can now be available as early as 18 to 24 h. This early diagnosis allows improvement of infection control measures and reduces the cost associated with MRSA in human8 and veterinary hospitals.9 Other examples of interest are isolation of Salmonella spp.4, 10, 11 which are typically hard to isolate and yet thought to be commonly associated with equine diarrhea, and for enhanced discrimination of yeasts such as Candida spp.4 which are slow growing. The most common microbiological pathogens of the reproductive tract in the horse are Streptococcus spp., E. coli, Klebsiella pneumoniae, S. aureus, and Pseudomonas aeruginosa. Specific bacterial venereal pathogens of horses are K. pneumoniae, P. aeruginosa and Taylorella equigenitalius (see Chapter 208). Some of the previously mentioned bacteria (Streptococcus spp., E. coli, K. pneumoniae, and P. aeruginosa) are also common pathogens of the human urinary tract. A number of commercially prepared chromogenic agars have been developed for the accurate and rapid identification of pathogens of the human urinary tract. Urine cultures in human hospitals
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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contribute greatly to the daily workload of a microbiological laboratory,12 hence there has been much research devoted to the development of highly specific chromogenic agars that have the same ability to detect urine pathogens as the combination of traditional blood agar and MacConkey agars, and to reduce laboratory workloads. Brilliance Urinary Tract Infection (UTI) Agar (Oxoid Ltd, Basingstoke, Hampshire, UK) is one such commercially prepared medium and is the chromogenic medium used at the Goulburn Valley Equine Hospital (GVEH) to detect potential reproductive tract pathogens. The potential of chromogenic agar for use in our veterinary practice was first realized after a ‘Google’ search was performed by one of us (AOM) in 2006 to find a practical, ‘in house’ rapid and accurate laboratory technique to diagnose P. aeruginosa and K. pneumoniae. The use of chromogenic agar had also been reported to allow identification of other, less common, pathogens of the human urinary tract including Streptococcus spp., Staphylococcus spp., Enterobacter aerogenes, Enterococcus fecalis, Citrobacter spp., Proteus spp., and Candida spp. Similarly these pathogens are also isolated from the reproductive tract of horses and recently we reported on the use of chromogenic agar in the rapid identification of these pathogens.13, 14
Detection of potential equine reproductive tract pathogens A total of 1374 endometrial swabs and 1158 clitoral swabs were collected from Thoroughbred mares on farms serviced by practice veterinarians at the Goulburn Valley Equine Hospital and from other regional veterinary practices.14 Endometrial samples were taken using double-guarded swabs to minimize vaginal or perineal contamination. The swabs were transported to the laboratory in transport medium. On arrival the name of the mare, the breeding farm, the stallion farm, the date and time were recorded and the swabs were then streaked on chromogenic agar, HBA and MacConkey split agar plates (Oxoid Ltd, Basingstoke, Hampshire, UK). Plates were incubated aerobically at 37◦ C, and any bacterial growth was registered at 24 h and 48 h. Examination at 24 h was required to identify heavy mixed cultures that may have required subculturing, so individual colonies could then be identified at 48 h. Also at 24 h, presumptive identification of pure uterine cultures permitted antimicrobial sensitivity testing to begin, with results often available at 48 h. All plates were then re-examined at 48 h for a final evaluation. A negative result was only confirmed at 48 h. Any fur-
Table 209.1 Table 162-1 Common reproductive tract bacteria identified by the RapIDTM SS/U test Pseudomonas spp. Klebsiella spp. Escherichia coli Staphylococcus spp. Enterobacter spp. Citrobacter spp. Enterococcus spp. Proteus spp. Candida albicans
ther subculturing or antimicrobial sensitivity testing was performed at this time. When all microorganism identification was complete and antimicrobial sensitivities were analyzed, a complete report was made available to both the stallion farm and broodmare farm. In general, for reasons of discretion, a positive result was only made available to the broodmare farm. Using chromogenic agar, microorganisms are generally identified by their color and Gram stain appearance; however, to confirm the presence of P. aeruginosa and K. pneumoniae, biochemical testing is always performed. The three biochemical test kits used at the GVEH practice laboratory are the RapIDTM NF Plus, the RapIDTM ONE, and the RapIDTM SS/U identification tests (Remel, Lenexa, KS). These tests are designed to be easy to set up, easy to interpret, accurate, and allow same-day results, as these tests only require 2–4-hour incubation periods.15, 16 The RapIDTM NF Plus test was used for the diagnosis of the medically important glucose nonfermenting (catalase positive and oxidase variable) Gram-negative rods (e.g., P. aeruginosa), the RapIDTM ONE test for the identification of medically important Enterobacteriaceae and other selected, oxidase negative, Gram-negative bacilli (e.g., K. pneumoniae), and the RapIDTM SS/U test was for general identification of microorganisms commonly isolated from the reproductive tract when it was not clear that either of the above-mentioned tests were more appropriate (Table 209.1). The color and colony appearance guide provided in Table 209.2 was used to identify the bacteria isolated. The identification of microorganisms is made at both 24 h and 48 h. The similar appearance of Staphylococcus spp. colonies and those of P. aeruginosa required an oxidase test to be performed. Oxidase Test Strips (Oxoid Ltd, Basingstoke, Hampshire) were used to perform this test (Figure 209.1). The diagnosis of P. aeruginosa is suggested based on colony appearance (Figure 209.2a,b), a positive oxidase test, identification by Gram stain of medium-sized Gramnegative rods and then, after subculturing to ensure
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Table 209.2 Colony identification guide and percentage of isolates from reproductive cultures at the Goulburn Valley Equine Hospital Species
Isolation % at GVEH
Pseudomonas aeruginosa
1.17
Klebsiella pneumoniae Escherichia coli Streptococcus spp. Staphylococcus spp. Enterobacter aerogenes Citrobacter spp. Enterococcus spp. Proteus spp. Candida spp.
1.21 33.58 27.76 18.02 3.72 1.41 5.41 7.60 0.12
Appearance of colonies on Chrome UTI Agar Variable. Usually cream/pale green or tan/brownish. Oxidase positive. Good but slow growth. 2.5–4 mm Good growth, large purple/deep blue/metallic navy colonies. 2–4 mm Good growth, darkish pink colonies. 2–3 mm Good growth, light blue colonies. 0.5–1 mm Good growth, white colonies. Oxidase negative, always catalase positive. 2–3 mm Dark blue/navy colonies. Similar but subjectively smaller than Klebsiella spp. 2–4 mm Dark blue/navy colonies. Usually smaller than Klebsiella spp. and Enterobacter spp. Good growth, light blue colonies. 0.5–1 mm Good growth, brown colonies. Swarming Good but slow growth. Typically blue/white colonies
pure colonies, a RapIDTM NF Plus identification test is performed as the final confirmation. It was noted that the emergence of P. aeruginosa colonies were somewhat slower than other tested microorganisms, and therefore a 48 h examination of incubated plates, despite a previous presumptive diagnosis, was required in all cases. Also noted was the unique effect that pyocyanin and pyoverdin (Figure 209.2c), a blue-green pigment typically produced by P. aeruginosa, had on sensitivity plates. Another interesting aspect of P. aeruginosa is fluorescence in dark conditions when exposed to an ultraviolet light (e.g., Wood’s UV light) (Figure 209.2d). This affect can also be seen on the vulvar labia of affected mares (Figure 209.2e). The diagnosis of K. pneumoniae (Figure 209.3a) required the differentiation of these colonies from those of Enterobacter spp. (Figure 209.3b) and Citrobacter spp. as their similar color appearance on chromogenic agar (Figure 209.3c) prevented differentiation among them.5 They were differentiated by subculturing when necessary to isolate single colonies, a Gram stain to confirm a Gram-negative rod, and then the RapIDTM One test was performed for final identification (Figure 209.3d). The presumptive diagnosis of the other main offending bacteria (i.e., E. coli, Streptococcus spp., and Staphylococcus spp.) was made by using the colony
Figure 209.1 A positive oxidase test result. A purple color within 30 seconds is a positive result. Oxidase tests are performed by lightly rubbing the suspect colony on the test strip. Typically the color change occurs in < 10 seconds. (See color plate 88).
identification guide shown in Table 209.2. E. coli grew rapidly and was the only bacterium that produced pink colonies (Figure 209.4a,b) on the chromogenic agar. The most significant Streptococcus spp. associated with endometrial disease are the β-hemolytic Streptococcus spp. (e.g., Streptococcus equi var. zooepidemicus). To diagnose the presence of a β-hemolytic Streptococcus spp., the hemolytic pattern must be taken from the bacterial growth on HBA as chromogenic agar will not show any hemolytic patterns. This also allows the differentiation from the nonhemolytic and α-hemolytic Streptococcus spp. The presence of β-hemolytic pinpoint colonies that are only present on HBA (Figure 209.5a), in conjunction with the presence of light blue colonies on chromogenic agar (Figure 209.5b) allows the presumptive diagnosis of β-hemolytic Streptococcus spp. Enterococcus spp. (Gram-positive cocci formerly classified in the Streptococcus group) is the other main differential for the growth of light blue colonies on chromogenic agar. However, Enterococcus spp. are unique in that they have the ability to grow on both HBA and MacConkey agar, thus aiding in the differentiation from Streptococcus spp. (which do not grow on MacConkey agar). Staphylococcus spp. can be differentiated from P. aeruginosa, as it is oxidase negative and catalase positive (Figure 209.6). The catalase test is another simple test that can be used to identify a species of bacterium. It is especially useful for distinguishing between Staphylococcus spp. and Streptococcus spp. when both may show hemolysis on blood agar and both are coccoid on Gram stain. (Typically, Streptococcus spp. are ovoid as they divide only in one plane and are present in chains, while Staphylococcus spp. are perfectly round and are present in clusters as they divide in two planes.) The catalase test is performed by placing a drop of 3% hydrogen peroxide on a microscope slide, and smearing a bacterial colony
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(e) Figure 209.2 (a) Pseudomonas aeruginosa. The cream to light brown colored colonies are the most common colony types. (b) Pseudomonas aeruginosa on a HBA/MacConkey split plate (c) Pseudomonas aeruginosa, demonstrating effects of pyocyanin and pyoverdin on a sensitivity plate. (d) Left: Klebsiella demonstrating no fluorescence with UV stimulation; right: Pseudomonas aeruginosa demonstrating fluorescence. (e) Pseudomonas aeruginosa on the vulval lips of a mare demonstrating fluorescence with UV stimulation. (See color plate 89).
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Figure 209.3 (a) Klebsiella pneumoniae. (b) Enterobacter aerogenes. (c) Enterobacter aerogenes (left) and Klebsiella pneumoniae (right). (d) RapIDTM One test with various color reactions (Klebsiella pneumoniae). (See color plate 90).
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Figure 209.4 (a) The pink colonies of Escherichia coli. (b) Large pink colonies are indicative of E. coli in this example taken from the clitoris, which also demonstrates multiple other colonies that can be presumptively identified from the color guide. (See color plate 91).
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Figure 209.5 (a) β-hemolytic Streptococcus spp. on HBA/MacConkey split plate. (b) Streptococcus spp. on chromogenic agar. (See color plate 92).
from chrome agar through the drop. A positive result is bubbling or frothing associated with generation of oxygen (Figure 209.6). Staphylococcus spp. are strongly catalase positive and Streptococcus spp. and Enterococcus spp. are catalase negative. The catalase test should not be performed on blood agar as it contains catalase. To confirm the presence of Staphylococcus aureus, a coagulase test must be performed. Coagulase is an enzyme produced by particular Staphylococcus spp. and correlates with pathogenicity. The test is performed by mixing a suspension of staphylococci with rabbit plasma on a slide. A positive reaction is indicated by clumping of bacteria within 1–2 minutes. S. aureus is coagulase positive, whereas many commensal Staphylococcus spp. are negative. This test is not routinely performed at the GVEH. Proteus spp. are a reasonably common isolate and can be recognized by ‘swarming’ of the
colonies and brown pigment on chromogenic agar (Figure 209.7). Candida spp. and Staphylococcus spp. can have similar color colony formation on both HBA/ MacConkey Agar and chromogenic agar (Figure 209.8a,b) and can occasionally confuse the diagnosis as both may produce white or pale colored colonies that are oxidase negative. A Gram stain easily differentiates these microorganisms and Staphylococcus spp. invariably are catalase positive. A specific chromogenic agar is also available for yeast identification (Oxoid Ltd) (Figure 209.8c,d) and a biochemical test (RapID yeast plus) (Oxoid Ltd) can separate the identities of up to 304 clinically significant yeasts.17 The value of chromogenic agar compared to HBA/MacConkey traditional plates for rapid identification of potential pathogens can be appreciated when examining multiple colonies in heavy
Figure 209.6 A catalase test is performed by placing a drop of 3% hydrogen peroxide on a microscope slide, smearing a bacterial colony from chrome agar into the hydrogen peroxide, and a positive result is to observe frothing.
Figure 209.7 Proteus spp. can be recognized by ‘swarming’ of the colonies and brown pigment on chromogenic agar. (See color plate 93).
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Figure 209.8 Staphylococcus spp. (left side of plate) and Candida spp. (right side of plate) can have similar color colony formation on both HBA/MacConkey Agar (a) and chromogenic agar (b). (c) Candida albicans on specialized yeast medium. (d) Candida tropicalis on specialized yeast medium. (See color plate 94).
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Figure 209.9 Multiple colonies in heavy concentrations from clitoral sinus cultures on HBA/MacConkey (a) and chromogenic agar (b). (See color plate 95).
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concentrations from clitoral sinus cultures (Figure 209.4a,b, Figure 209.9a,b). In our laboratory an endometrial culture is considered positive if five colony-forming units per plate are identified, and had no more than three different bacterial pathogens present. Any positive endometrial swabs that have four or more different bacterial colonies are presumed to be contaminated, and re-swabbing of the affected mare is advised. If an endometrial swab result has a positive growth, antimicrobial sensitivity testing is performed, using colonies taken from the chromogenic agar. Some antibiotics tested at our facility for microbial sensitivity are penicillin, ticarcillin, gentamicin, neomycin, ceftiofur, and enrofloxacin (Figure 209.10a,b). Quality control measures are used to ensure that results are accurate and reliable. These consist of standard pure cultures of bacterial specimens that have been previously validated by an accredited microbiology laboratory, and can be used to compare color and biochemical tests results. In-house biochemical testing is routinely used on all bacteria that are from positive endometrial cultures. As shown in Table 209.1, the RapIDTM SS/U test can be used to identify many of the bacteria that are expected to be reproductive pathogens. During our first year using the new procedure, independent diagnosis of bacteria was also confirmed on selected samples by an accredited microbiological laboratory.
(a)
Potential use of chromogenic agar in routine reproductive practice The use of chromogenic agar is a new and exciting technique to help identify the pathogens of the equine reproductive tract. It has revolutionized our practice’s microbiological management due to rapid and accurate identification of common reproductive tract pathogens. It has a number of advantages over traditional methods of microbial identification. These include the reduction in the time required for the laboratory processing of samples; the ease of colony identification; the low level of skill required to interpret results; the reliability of colonies taken from chromogenic agar for pathogen identification and antimicrobial sensitivity testing; and the proven high level of sensitivity of chromogenic agar in identifying pathogens.15, 16 The slightly higher cost of chromogenic agar per plate (A$1.15) compared to conventional HBA/MacConkey split plate media (A$0.58) was offset by a reduced need for complementary reagents and less labor associated with the processing of culture plates and suspect pathogens.4 In addition, the technique removed the often identified prob-
(b) Figure 209.10 Antimicrobial sensitivity testing of a bacterium (a) and a yeast (Candida albicans) (b). The clear zone in the center of Figure 209.10(b) is 3% hydrogen peroxide. (See Chapter 272). (See color plate 96).
lems associated with samples submitted immediately prior to the weekend. The disadvantages of this technique include: the slightly higher cost of chromogenic agar plates, especially in facilities processing small numbers of samples; chromogenic agar is of no benefit for the screening and diagnosis of Taylorella equigenitalis, the causative agent of contagious equine metritis (CEM),
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Use of Chromogenic Agar to Diagnose Reproductive Pathogens another potential venereal pathogen that recently has emerged again in the USA (see Chapter 247). The use of chromogenic agars does not differentiate reproductive pathogenic from non-pathogenic bacteria, and any interpretation of the significance of uterine bacteria isolated must be related to endometrial cytology, endometrial biopsy, or other forms of clinical examination. In summary, due to a limited availability of regional laboratory facilities that offer a rapid, accurate and continuous weekly screening service in many areas, the introduction of chromogenic agar has been very useful. It has resulted in a fast and accurate service which was especially important when samples are collected from mares close to the weekend, when traditional services are not able to provide a report at a time suitable to facilitate breeding, when mares are required to be presented for breeding with a negative culture for potential reproductive pathogens.
Acknowledgements John F Prescott, Professor, Department of Pathobiology, University of Guelph; Lucy Cudmore BVSc, Ellen Oostelaar DVM, and Amy Williamson BVSc from the Goulburn Valley Equine Hospital.
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References 1. Ricketts SW, Mackintosh ME. Role of anaerobic bacteria in equine endometritis. J Reprod Fertil Suppl 1987;35:343–51. 2. Ricketts SW. Contagious equine metritis (CEM). Equine Vet Educ 1996;8 (3): 166–70. 3. Timoney PJ. Contagious equine metritis. Comp Immunol Microbiol Infect 1996;19(3):199–204. 4. Perry JD, Freydiere AM. The application of chromogenic media in clinical microbiology. J Appl Microbiol 2007;103(6):2046–55. 5. Samra Z, Heifetz M, Talmor J, Bain E, Bahar J. Evaluation of use of a new chromogenic agar in detection of urinary tract pathogens. J Clin Microbiol 1998;36(4):990–4. 6. Carricajo A, Boiste S, Thore J, Aubert G, Gille Y, Freydiere AM. Comparative evaluation of five chromogenic
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media for detection, enumeration and identification of urinary tract pathogens. Eur J Clin Microbiol Infect Dis 1999;18(11):796–803. Kilian M, Bulow P. Rapid diagnosis of Enterobacteriaceae. I. Detection of bacterial glycosidases. Acta Pathol Microbiol Scand B 1976;84B(5):245–51. Krishna BV, Smith M, McIndeor A, Gibb AP, Dave J. Evaluation of chromogenic MRSA medium, MRSA select and Oxacillin Resistance Screening Agar for the detection of methicillin-resistant Staphylococcus aureus. J Clin Pathol 2008;61(7):841–3. Van den Eerde A, Martens A, Lipinska U, Struelens M, Deplano A, Denis O, Haesebrouck F, Gasthuys F, Hermans K. High occurrence of methicillin-resistant Staphylococcus aureus ST398 in equine nasal samples. Vet Microbiol 2009;133(1–2):138–44. Eigner U, Reissbrodt R, Hammann R, Fahr AM. Evaluation of a new chromogenic medium for the isolation and presumptive identification of Salmonella species from stool specimens. Eur J Clin Microbiol Infect Dis 2001;20(8):558–65. O’Neill W, Cooke RP, Plumb H, Kennedy P. ABC chromogenic agar: a cost-effective alternative to standard enteric media for Salmonella spp. isolation from routine stool samples. Br J Biomed Sci 2003;60(4):187–90. Fallon D, Ackland G, Andrews N, Frodsham D, Howe S, Howells K, Nye KJ, Warren RE. A comparison of the performance of commercially available chromogenic agars for the isolation and presumptive identification of organisms from urine. J Clin Pathol 2003;56(8):608–12. McKinnon AO, Beehan DP. A novel technique using Chrome Agar to diagnose equine reproductive tract pathogens. Proc 41st Annual Congress of the South African Equine Veterinary Association. J. Marais (Ed). Kwa Zulu Natal: 41:264–267, 2009 Beehan DP, McKinnon AO. How to diagnose common reproductive tract pathogens using chromogenic agar. Proc Am Assoc Equine Pract 2009;55:320–5. Kitch TT, Jacobs MR, Appelbaum PC. Evaluation of the 4-hour RapID NF Plus method for identification of 345 gram-negative nonfermentative rods. J Clin Microbiol 1992;30(5):1267–70. Kitch TT, Jacobs MR, Appelbaum PC. Evaluation of RapID ONE system for identification of 379 strains in the family Enterobacteriaceae and oxidase-negative, gram-negative nonfermenters. J Clin Microbiol 1994;32(4):931–4. Kitch TT, Jacobs MR, Mcginnis MR, Appelbaum PC. Ability of RapID Yeast Plus System to identify 304 clinically significant yeasts within 5 hours. J Clin Microbiol 1996;34(5):1069–71.
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Chapter 210
Evaluation of Uterine Tubal Patency William B. Ley
Undoubtedly the mare can experience reproductive tract pathology resulting in infertility. Embryo transfer (ET) has been offered in such instances as a potential means of producing offspring. However, a limited number of mares that have demonstrable failure to conceive may possess uterine tubal abnormality (e.g., blockage of sperm transport or blockage of embryo descent into the uterus). Successful ET requires that uterine tubal blockage is not contributing to the mare’s infertility. Assisted reproductive technologies (ART) such as in vivo oocyte recovery for in vitro fertilization (IVF), intractyoplasmic sperm injection (ICSI), gamete intrafallopian transfer (GIFT), or oocyte transfer (OT) have been developed as an additional means of obtaining offspring from mares that fail to provide embryos. ART are comparatively expensive to perform, and require technical expertise. GIFT or OT and ICSI are viable options when the donor is suspected of having uterine tubal occlusion. Oviductal pathology has received limited investigation in equine reproduction. A tubal blockage cannot readily be diagnosed through routine rectal palpation, ultrasonography, or laparoscopic examination unless a grossly obvious condition exists.
Diagnosis: the starch granule test Diagnosis of tubal patency Clinical diagnostics to demonstrate uterine tubal patency range from the straightforward and less invasive, to the more complicated and surgically involved. The starch granule test was first described by Allen et al. in 1979.1 A solution of starch granules is deposited onto the ovarian surface via a flank or paralumbar fossa approach with a long needle. Twentyfour hours later the uterus and cervix are lavaged for
sample collection by a vaginal approach. The recovered fluid is stained with 2% Lugol’s iodine and is examined microscopically to look for the presence of any starch granules that may have been transported to the uterine or cervical environment by oviductal transport. This test must be repeated to evaluate left versus right uterine tubal patency, one side per procedure. The use of laparotomy/laparoscopy to allow direct visualization of the tubal structures in question and their catheterization for flushing with nonreactive dyes has more recently been described in people2 and horses.3–5 These techniques are comparatively more invasive, and more expensive to perform. The use of ultrasound-guided transvaginal follicular aspiration for recovery of equine oocytes has been reported.6–8 By adaptation of this clinically accepted procedure, the author and co-workers9 investigated a standing diagnostic procedure that would discriminate left from right uterine tubal transport, and thus demonstrate patency of the mare’s uterine tubes in a single procedure. Thirteen barren mares were used for the study. These mares were used in laboratories for student instruction, and thus their ages, reproductive histories, and endometrial histomorphological architectures were known. These mares had been barren for at least the previous 2 years, and the majority for at least 5 years. The mares were examined by daily rectal palpation and transrectal ultrasonography for 3 days prior to their use in this clinical trial to determine ovarian activity. Blood samples were drawn on the day of each procedure to determine concentrations of progesterone. Mares were scheduled for tubal patency evaluation when they were in diestrus. Mares were restrained in stocks, the tail wrapped and perineal area prepared and scrubbed as for collection of per vaginal endometrial cytology and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Evaluation of Uterine Tubal Patency culture.10 Each mare received a vaginal douche with 500 mL sterile saline containing diluted 0.05% (vol./vol.) inorganic iodine solution followed by 450 mL sterile saline with 50 mL 2% lidocaine added. A cold-sterilized, transvaginal ultrasound probe (Aloka 500V, 5-MHz curvilinear transvaginal probe; Corometrics Medical Systems, North Wallingford, CT) pre-rinsed in sterile saline was then introduced per vaginum using a sterile gloved hand and passed forward to the anterior vagina. The probe remained in this location while the gloved hand was removed and then introduced into the rectum. One ovary (left or right) was identified, retracted, and stabilized near the probe head. Using a 60-cm, 18-Ga, singlechannel sterile needle introduced through the biopsy channel of the ultrasound probe, 2 million (15-µm) FluoresbriteTM flow cytometry microspheres (Polysciences Inc., Warrenton, PA) in 5 mL sterile saline were deposited over the surface of the ovary. Uterine lavage using 1 L sterile saline was performed at one of two intervals (24 and 48 hours) following deposition of the microspheres. Four mares were used in a cross-over design to determine the most suitable interval between microsphere deposition on the ovary and uterine lavage for recovery of transported microspheres, with a minimum of 14 days between each repetition. The optimum interval for uterine lavage following microsphere deposition on the ovaries was determined to be 48 hours. In four mares, no microspheres were recovered at 24 hours, but they were present in uterine lavage fluids recovered at 48 hours. In one mare, microspheres were recovered at 24 hours, but were not present at 48 hours. In another mare, microspheres were recovered at both time intervals (i.e., 24 and 48 hours postdeposition). The recovered uterine lavage fluid was either centrifuged at 1500 rpm for 10 min at 5◦ C, the supernatant decanted, and the pellet resuspended in 5 mL sterile saline, or was filtered through a 2-µm filter to separate microspheres and other particulate debris from the larger volume of recovered lavage fluid. The centrifuged or filtered samples were then tested by flow cytometry (Epics V; Coulter Diagnostics, Hialeah, FL) for microsphere presence and fluorescence pattern (green and red). The recovery of microspheres and the color of fluorescent microsphere beads recovered from left and right ovary deposition was recorded for each mare. No attempt was made to count the number of microspheres recovered. Mares received ultrasound-guided microsphere bead deposition on both ovaries at least twice, separated by a minimum 14-day interval between successive procedures. Eight (61.5%) of the 13 mares studied demonstrated patency of both uterine tubes on at least one
1989
of two attempts. One mare demonstrated patency of the left oviduct only on two attempts, and one mare demonstrated patency of the right oviduct only after two attempts. Three mares did not transport the fluorescent microsphere beads on either of two attempts separated by an interval of at least 14 days. When mares received the second of their two procedures, the respective color of the microsphere bead was deposited on the ovary opposite to that of the first procedure (i.e., cross-over fashion). In all attempts for all mares, the red-labeled fluorescent microsphere beads were less successful in demonstrating transport (26% for red vs 74% for green). When side of the reproductive tract was considered for all mares and all attempts, the left oviduct demonstrated microsphere bead transport 52% of the time whereas the right demonstrated transport 60% of the time. As all mares were part of an instructional herd none were sacrificed to determine whether oviductal pathology was present or not.
Incidence of impaired tubal patency Retention of uterine tubal eggs in the mare has been documented to occur either alone as degenerating masses or in association with proteinaceous globular masses.11, 12 Liu et al.13 examined the oviducts of 43 mares with known reproductive history and found that 59.1% of mares in their older study group (i.e., n = 22; > 7 years in age) had oviductal masses. Large masses were found to occupy the entire lumen of the oviduct and appeared to cause physical occlusion. Masses were greatest in number and size in the oviducts from mares aged 21 to 22 years. The mare’s oviduct has been considered well protected from uterine pathological conditions due to the strong muscular papillae at the utero-tubal junction and to the dorsal position of the oviduct in the abdominal cavity. Vandeplassche and Henry14 examined the genital tracts from 700 mares and concluded that salpingitis did occur in the mare as determined by histopathological criteria. Inflammation was found to occur more frequently in the ampullary region than in the isthmus. Vandeplassche and Henry14 further suggested a positive correlation between salpingitis and endometritis. Abnormalities occurred to a greater degree in the right side versus the left. Saltiel et al.15 examined 325 mares macroscopically, and a subset (n = 124) of these microscopically. This study found that 87.7% of these mares had at least one macroscopic lesion, and 93.5% had at least one microscopic lesion.
Summary Evaluation of uterine tubal patency by starch granule deposition as described by Allen et al.1 is relatively
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simple and requires little in the way of advanced technology or training. This author has used that technique clinically and with good results; however, the lack of discrimination between left and right ovaries is less than desirable. Allen et al.1 reported that starch granules from 6 to 56 µm in diameter were transported in all mares with patent oviducts as recovered at 24 hours. Based on this we selected a microsphere bead diameter that was within that range (i.e., 15 µm). Initial attempts at using a larger bead diameter were unsuccessful due to limitations related to the flow cytometer. A larger bead diameter would be feasible if the method of detection employed fluorescence microscopy rather than flow cytometry. Allen et al.1 further demonstrated that starch granules were transported in all mares regardless of the mare’s stage of estrous. However, when starch granules were deposited on an ovary that had ovulated a follicle on that same day, starch granule transport could not be demonstrated until 4 to 7 days after that ovulation. For this reason in our study we elected to deposit microsphere beads when mares were determined to be in diestrus. Evaluation of uterine tubal patency as performed by laparoscopy/laparotomy is comparatively involved and costly, and requires surgical training and an aseptic setting. Expense for the owner, healing time post-surgery, and potential postsurgical complications, some of which may be life-threatening, are serious considerations making such invasive techniques less than optimal as diagnostic tests of tubal patency. Ultrasound-guided microsphere deposition is a modified oviductal patency test that is less expensive for the mare owner, and is less invasive for the mare than laparoscopy/laparotomy. Furthermore, this is a standing diagnostic procedure that has the potential both to discriminate left from right uterine tubal transport, and to demonstrate patency of the mare’s uterine tubes in a single procedure. Successful treatment of suspected oviductal blockage has recently been described by Allen et al.16 using laparoscopic application of prostaglandin E2 (PGE2 ) gel directly onto the oviductal surface. Fifteen mares with known reproductive histories of conception failure for inexplicable reasons were subjected to the treatment protocol. Fourteen of these mares conceived within the same or subsequent breeding season. The authors describe their procedure as ‘remarkably straightforward and apparently successful,
although definitive pretreatment diagnosis remains problematic.’ It is unfortunate that they had not sufficiently reviewed the literature and applied the diagnostic technique described by Ley et al.9 to their treatment group prior to application of PGE2 gel to the oviducts.
References 1. Allen WE, Kessy BM, Noakes DE. Evaluation of uterine tube function in pony mares. Vet Rec 1979;105:364–6. 2. Bevan CD, Johal BJ, Mumtaz G, Ridgway GL, Siddle NC. Clinical, laparoscopic and microbiological findings in acute salpingitis: report on a United Kingdom cohort. Brit J Obstet Gynaecol 1995;102:407–14. 3. Fischer AT, Lloyd K, Carlson GP, Madigan JE. Diagnostic laparoscopy in the horse. J Am Vet Med Assoc 1986;189: 289–92. 4. Fischer AT. Standing laparoscopic surgery. Vet Clin N Am Equine 1991;7(3):641–7. 5. Witherspoon DM, Kraemer DC, Seager S. Laparoscopy in the horse. In: Harrison RM, Wildt DE (eds) Animal Laparoscopy. Baltimore: Williams & Wilkins, 1980; pp. 157–67. 6. Squires EL, Cook NL. Transvaginal aspiration. Vet Clin N Am Equine 1996;12:13–29. 7. Carnevale E, Ginther O. Use of a linear ultrasonic transducer for the transvaginal aspiration and transfer of oocytes. J Equine Vet Sci 1993;13:331–3. 8. Ray B, Squires EL, Cook NL, Tarr SF, Jasko DJ, Hossner KL. Pregnancy following gamete intrafallopian transfer in the mare. J Equine Vet Sci 1994;14:27–30. 9. Ley WB, Bowen JM, Purswell BJ, Dascanio JJ, Parker NA, Bailey TL, DiGrassie WA. Modified technique to evaluate uterine tubal patency in the mare. P Annu Conv Am Assoc Equine Pract 1998;44:56–9. 10. Ley WB. Current thoughts on the diagnosis and treatment of acute endometritis in mares. Vet Med 1994;89: 648–60. 11. David JSE. A survey of eggs in the oviduct of mares. J Reprod Fertil Suppl 1975;23:513–17. 12. Onuma H, Ohnami Y. Retention of tubal eggs in mares. J Reprod Fertil Suppl 1975;23:507–11. 13. Liu IKM, Lantz KC, Schlafke MA, Bowers JM, Enders AC. Clinical observations of oviductal masses in the mare. P Annu Conv Am Assoc Equine 1991;36:41–5. 14. Vandeplassche M, Henry M. Salpingitis in the mare. P Annu Conv Am Assoc Equine 1977;23:123–31. 15. Saltiel A, Paramo R, Murcia C, Tolosa J. Pathologic findings in the oviducts of mares. Am J Vet Res 1986;47: 594–7. 16. Allen WR, Wilsher S, Morris L, Crowhurst JS, Hillyer MH, Neal HN. Laparoscopic application of PGE2 to re-establish oviductal patency and fertility in infertile mares: a preliminary study. Equine Vet J 2006;38:454–9.
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Chapter 211
Laparoscopy Dean A. Hendrickson and David G. Wilson
Introduction The primary indication for laparoscopic surgery in the mare is for removal of ovaries. The most common procedures performed are removal of granulosa thecal cell tumors and ovarian removal for behavior modification in the performance horse. Traditionally, ovariectomy was performed in the standing horse via colopotomy or paralumbar fossa celiotomy, and in the anesthetized mare though ventral midline, paramedian, or flank incisions. Laparoscopic observation of the mare’s reproductive tract was described as early as 1983.1 Coincident with the development of the video camera for use in laparoscopy and of laparoscopic instrumentation, in the early 1990s equine surgeons began to develop minimally invasive approaches to removal of normal and abnormal ovaries.2 Laparoscopic ovariectomy is now routinely performed in both the standing and recumbent mare. Since its original introduction, most publications on the technique have focused on the development and evaluation of alternative techniques for hemostasis. This review is intended to provide a contemporary summation of the state of laparoscopic ovariectomy in the horse as well as a brief introduction to the techniques of oviductal ligation and oocyte transfer.
Laparoscopic instrumentation Operative laparoscopy was limited until the development of the video camera, which allowed projection of the laparoscopic image to a video monitor to allow the surgical team to simultaneously view the surgical field. Standard equipment needed to accomplish laparoscopic ovariectomy includes: a laparoscope, a light source, a camera and video monitor, an insufflator, and an assortment of laparoscopic instrumentation. A list of required and helpful instrumentation can be found in Table 211.1. Most surgeons use a 10-mm laparoscope with either a 0◦ or 30o view. Laparoscopic ovariectomy
can be accomplished with either view; however, the angled view offers more utility and facilitates direct visual observation of cannula insertions that might not be possible with a straight-ahead viewing instrument. Standard-length laparoscopes in the neighborhood of 33 cm long offer sufficient length to accomplish the procedure. Although 33-cm scopes can be sufficient, longer scopes (57-cm) allow the surgeon to explore the abdomen more completely, especially the contralateral side. The light source is one of the most critical components of the system. The abdomen of the horse is a large cavity and illumination requires considerable light power. In general, brighter is better and most systems used for laparoscopic surgery in the horse are 300-watt xenon light sources. When purchasing a light source for use in laparoscopy, vendors who suggest that their 175-watt light source provides just as much light as a competitor’s 300-watt source should be viewed with skepticism. Virtually any quality computer chip camera and monitor will suffice. Systems that offer threedimensional capacity looked promising upon entry to the market place, but have not enjoyed much popularity to date. Flexibility of positioning of the video tower complex is enhanced if a longer than standard light guide is available. The author uses a 10foot (3-m) light cord, as at times, it can be difficult for everyone involved in the procedure to be strategically placed to view the video monitor. The largest diameter light cord available should be used, as the increased number of fibers will transmit more light into the abdomen. The use of multiple monitors and the fairly recent introduction of affordable head-mounted virtual reality video goggles greatly improve the operative environment. Abdominal insufflation is essential to allow visual exposure within the abdomen. A variety of CO2 insufflators are available. Adequate insufflation of the adult abdomen can require 30–40 liters of CO2 .
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 211.1 A list of instrumentation for laparoscopic ovariectomy (instruments that are available through only a limited source are so identified) Type
Number
Description
Basic instruments needed to perform laparoscopy Laparoscope 1 or 57 cm × 10 mm 30◦ or 0◦ 1 30 cm × 10 mm 30◦ or 0◦ Light source 1 300 watt xenon with large-diameter light cord Video camera and monitor 1 Insufflator 1 At least 6 L/min (20 L/min preferred) Trocars/cannulas 3 minimum 10–11-mm diameter × 20 cm long with one blunt trocar and one sharp pyramidal-shaped trocar 6 preferred Cannula reducer 1–2 5-mm reducer Instruments 1 Babcock atraumatic grasping forceps (45 cm long preferred) 2–3 Acute claw grasping forceps 45 cm long 1 Serrated scissors 45 cm long is good Injection needle 1–2 33 cm or 45 cm (preferred) Ligation instruments Knot pusher LigaSure device Bipolar cautery unit Ultrasonic shears
1–2 or 1 or 1 or 1
35 cm or 40 cm (preferred) (Surgical Direct, Deland, FL) Atlas shears
Optional equipment Instruments Needle holders Digital recorder Hard copy imager
1 2 1 1
Ligaclip applier for medium/large clips 10-mm/45 cm long (Surgical Direct, Deland, FL)
Insufflators capable of delivering flows in a range of 10–20 L/min facilitate rapid initial distension of the abdomen as well as maintenance of distension during the procedure. Most surgeons utilize laparoscopic cannula systems. Longer cannulas (15–20 cm) are needed for standing laparoscopy (Figure 211.1). A minimum of three cannulas are required; however, additional cannulas may be used when performing bilateral standing ovariectomy, or ovariectomy in the recumbent mare. Given a choice, the authors prefer cannulas with manual valves that allow the valve to be held open during withdrawal of instrumentation. Standard-diameter cannulas range from 10 to 12 mm. Reducer systems should be available to allow the use of both 5-mm diameter and 10-mm diameter instruments (Figure 211.2). Surgical manipulation during laparoscopic ovariectomy can be accomplished using a variety of instruments. Disposable single-use instrumentation works well and can often be cleaned and reused. Reusable instrumentation offers long-term cost savings. One of the authors (D.G.W.) uses single-use disposable scissors for tissue dissection, while the other (D.A.H.) uses disposable scissors only for suture transection. Basic instrumentation should include: an endolaparoscopic Babcock forceps (Figure
211.3), a right-angle dissecting forceps (Figure 211.4), at least two acute claw graspers (Figure 211.5) and some type of ligating device such as a knot pusher or a bipolar cautery delivery system (Figure 211.6).
Figure 211.1 Reusable laparoscopic cannulas and trocars showing 11-mm diameter cannulas in 10.5- and 20-cm lengths (Karl Storz Endoscopy, Goleta, CA).
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Laparoscopy
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Ovarian anatomy
(a)
(b) Figure 211.2 (a) Laparoscopic cannula (11-mm diameter) with 5-mm diameter reducer valve (Karl Storz Endoscopy, Goleta, CA). (b) A 5-mm reducer that can be used as a knot protector as well (Surgical Direct, Deland, FL).
Figure 211.3 Endolaparoscopic Babcock forceps (10-mm diameter) 37 cm long with pistol grip handle (Dr Fritz, LLC Louisville, KY).
Figure 211.4 Endolaparoscopic right-angle forceps (10-mm diameter) 47 cm long with in-line box lock handle (Dr Fritz, LLC Louisville, KY).
Figure 211.5 Claw grasper (10-mm diameter) 47 cm long (Dr Fritz, LLC Louisville, KY).
Figure 211.6 Klepinger 5-mm diameter bipolar cautery forceps (CONMED Linvatec, Largo, FL).
The ovary is suspended from the dorsal body wall by a complex of attachments commonly referred to as the ovarian pedicle. Lateral attachments that must be divided to allow removal of the ovary include the mesosalpinx, which forms the lateral border of the ovarian bursa, and the uterine ovarian ligament, which connects the tip of the uterine horn to the caudal margin of the mesosalpinx and the oviduct. Cranially, the suspensory ligament of the ovary is essentially confluent with the mesosalpinx laterally and the mesovarium medially. This medial component of the pedicle is thicker than the lateral component, and the ovarian artery and vein are transmitted in the more cranial part. The proper ligament of the ovary connects the caudal aspect of the ovary to the tip of the uterine horn and is essentially a caudal continuation of the mesovarium. This is the most dense of the attachments.
Laparoscopic ovariectomy in the standing horse To facilitate visual observation within the abdomen, the mare is fasted for a minimum of 24 hours up to 48 hours. The mare is restrained in a stocks and the paralumbar fossa is prepared for aseptic surgery. Unilateral ovariectomy is approached from the ipsilateral side while bilateral ovariectomy requires bilateral approaches. Standing sedation can be achieved using a variety of drug combinations including xylazine (0.5–1.0 mg/kg) and butorphenol (0.02 mg/kg) or detomidine (2–10 g/kg) and butorphenol (0.02 mg/kg).3 Owing to the duration of the typical standing ovariectomy, many surgeons use a continuous-rate infusion of detomidine supplemented with butorphenol. Local anesthesia can be accomplished with an inverted ‘L’ in the paralumbar fossa, or by instilling local anesthetic at each portal site. A xylazine epidural can be performed at the S5–C1 or C1–C2 space (0.18 mg/kg xylazine q.s. to 10 mL with saline). In the authors’ experience, this approach allows ovarian manipulation without instilling local anesthetic into the ovarian pedicle in approximately 50% of horses. Detomidine epidurals have also been used (40–60 µg/kg q.s. to 10 mL with saline); however, this approach will provide significant sedation requiring dose modification of any supplemental systemic sedation protocol.4 Trocar insertion is probably safer if the abdomen is insufflated before insertion; however, inadvertent insufflation of the retroperitoneal spaces is a frequent complication, especially for the inexperienced surgeon. Insufflation can be performed through a ventral midline-placed teat cannula or through a cannula
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placed in the paralumbar fossa. Some surgeons use specially designed insertion cannulas (requires 0◦ viewing laparoscope) that allow visual observation during insertion (Visiport Optical Trocar; Covidien, Mansfield, MA) to gain entry into the uninsufflated abdomen.5 Others place the initial trocar without insufflation using a sharp or blunt trocar. For over 15 years, one author (D.G.W.) routinely insufflated the abdomen before trocar insertion using a 12-Fr chest trocar catheter (12-Fr Medicut Argyle Catheter; Sherwood Medical, St Louis, MO) placed into the abdomen approximately 1 cm ventral to the dorsal margin of the internal abdominal oblique muscle, with few complications. Developing the right feel for complication-free insertion of the chest trocar is difficult to teach. More recently, both authors have adopted the technique of inserting the initial 11-mm diameter 20-cm cannula with a blunt trocar without insufflation. Specifically, a 1-cm skin incision is made slightly below the dorsal aspect of internal abdominal oblique muscle at the cranial-to-caudal midpoint of the paralumbar fossa. The incision is carried through the external fascia of the external abdominal oblique muscle to allow reasonably friction free passage of the cannula to the level of the peritoneum. The cannula with a blunt trocar is inserted until the resistance of the peritoneum is encountered and advanced into the abdomen with a controlled thrust similar to the technique for collection of peritoneal fluid with a teat cannula. The blunt obturator is withdrawn and the laparoscope is inserted to confirm intra-abdominal placement. At this point, the abdomen is insufflated to a pressure of between 10 and 20 mmHg. The authors have slightly different approaches to further cannula placement. D.G.W. prefers to continue cannula placement by inserting an instrument cannula cranioventral to the initial insertion. A second instrument cannula is inserted caudoventral to the la-
Figure 211.8 Standing laparoscopy from the left paralumbar fossa showing left ovary. O, ovary; black arrow, suspensory ligament of the ovary; white arrow, mesosalpinx.
paroscope (Figure 211.7). Alternatively, the cranial cannula can be placed through the last intercostal space to allow instrument separation. D.A.H. prefers to place the telescope and the two instrument cannulas in a more vertical configuration (Figure 211.7). The ovary is identified and the suspensory ligament is grasped at the cranial pole (Figure 211.8). To ensure desensitization of the ovary, local anesthetic (10–30 mL) is injected directly into the ovarian pedicle in multiple locations (Figure 211.9). Injection of local anesthetic directly into the ovary is less effective.6 The specific manipulative approach from
(a)
(b)
Figure 211.7 Laparoscope and instrument cannula positioning for standing laparoscopy. D.G.W.: ×, laparoscope; ◦, instrument cannulas. D.A.H.: 1, laparoscope, 2 and 3, instrument cannulas.
Figure 211.9 (a) Injection needle. (b) Intra-abdominal picture showing injection of the ovarian pedicle.
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Laparoscopy this point forward is dependent upon the selected approach for hemostasis. Ligation of the ovarian pedicle Ligation of the ovarian pedicle is substantially more difficult than ligation of the typical cryptorchid testis. In the first instance it can be difficult to manipulate the ligating loop over the ovary, especially in the case of large granulosa thecal cell tumors. Secondly, the very mass of the pedicle can preclude effective closure of the ligature. Although simply placing a large-diameter suture ligature over the ovarian pedicle has been described,7 ligature slippage/failure has also been reported.8 The authors have never been comfortable with block ligation of the ovarian pedicle. Ligature failure can largely be avoided if the ovarian pedicle is dissected before ligature placement. Pedicle dissection is achieved by stabilizing the ovary with a Babcock or acute claw grasping forceps placed through the cranial instrument cannula (D.G.W.) or instrument cannula 2 (D.A.H.) (Figure 211.10). Endolaparoscopic scissors are used to divide the uterine ovarian ligament, oviduct, and mesosalpinx in a cranial direction. The proper ligament of the ovary is partially divided beginning at the tip of the uterine horn and when combined with division of the suspensory ligament of the ovary, the mass of the remaining ovarian pedicle is substantially reduced to allow secure placement of two ligating loops (Figure 211.10).9 Ligature options include commercially available size 0 polydioxinone suture (PDS) loops (Endoloop; Ethicon Endo-Surgery Inc., Johnson & Johnson Co., New Brunswick, NJ) and hand-tied loops. A variety of knot configurations have been described, but the 4S modified Roeder knot using size #1 Maxon provides the greatest security.10, 11 Ligature using polyamide tie-rap (similar to cable ties) has been reported.12 To a large extent
1995
ovarian pedicle ligation has been supplanted by the development of other hemostatic techniques, most often electrocoagulation. Electrocoagulation of the ovarian pedicle Electrocoagulation is used extensively as a sole method of hemostasis in laparoscopic surgery in humans. Monopolar electrocautery is widely available and has been used to achieve hemostasis in laparoscopic ovariectomy. Unfortunately, inadvertent capacitance discharge can result in iatrogenic injury, and most surgeons prefer to avoid that potential complication by using bipolar cautery. The suspensory ligament of the ovary is grasped with a Babcock forceps or some other endoscopic grasping forceps. Electrocautery is applied, beginning proximal to the cranial pole of the ovary using a bipolar cautery forceps (Figure 211.6). Adequate coagulation is confirmed when the tissue blanches and shrinks. The cautery forceps is withdrawn and the coagulated tissue is transected with endolaparoscopic scissors. Sequential coagulation and cutting is continued until the ovary is freed from its pedicle.13 The ovarian bursa is entered early in the sequential process allowing the avascular lateral part of the pedicle (mesosalpinx) to be sharply divided, terminating with division of the oviduct and the uterine ovarian ligament. The vascular structures of importance reside in the thicker medial component of the pedicle (the mesovarium proper).13 The sequential electrocoagulation and cutting process is continued in a cranial to caudal direction along the medial part of the pedicle (mesovarium and proper ligament of the ovary) to free the ovary. From an efficiency standpoint, having to alternate the cautery forceps with scissors is cumbersome, and a bipolar cautery instrument with a built-in blade is available in both 5-mm diameter and 10-mm diameter instruments, 32 cm in length (Figure 211.11). Technological advances have provided a feedback-controlled bipolar cautery device known as the LigaSureTM (Valleylab, Boulder, CO). This device monitors tissue capacitance during the application of bipolar current to create a consistent controlled coagulation. In its current iteration, this laparoscopic instrument has a deployable blade between the halves of the bipolar cautery allowing sequential coagulation and cutting with the same instrument (Figure 211.12). This device represents the state of the art for achieving hemostasis for laparoscopic ovariectomy.14 Ultrasonic coagulation of the ovarian pedicle
Figure 211.10 Intra-abdominal picture showing the sharply dissected ovarian pedicle.
The Harmonic Scalpel (Ethicon Endo-Surgery, Inc., Cincinnati, OH) is an ultrasonic instrument that cuts
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Figure 211.11 Tripolar cutting forceps (10-mm diameter) 32 cm long (ACMI, Southborough, MA).
and coagulates tissue. Though relatively expensive to purchase, this instrument has been used successfully to achieve hemostasis during laparoscopic ovariectomy in standing horses.15 A variety of applicators are available, but the laparoscopic coagulation shears offer the best option for grasping and coagulating the tissue of the ovarian pedicle. Unfortunately, this instrument may fail to achieve hemostasis during
the first application, and manipulation of the pedicle may be required to locate sites of hemostatic failure to allow reapplication of the device.15 Other authors reported similar complications with a high-powered ultrasonic shears (United States Surgical, Norwalk, CT).16
Laser coagulation of the ovarian pedicle
(a)
The first laparoscopic ovariectomy was described by Palmer in 1993.2 He used an Nd:Yag laser to divide the ovarian pedicle. Effective hemostasis was achieved, but some breakthrough bleeding occurred, necessitating reapplication of the laser or the application of ligating clips. While the laser can be used effectively in this application, cost considerations are significant and electrocoagulation techniques offer better and more efficient hemostasis.
Endo-GIA facilitated division of the ovarian pedicle
(b) Figure 211.12 (a) LigaSure vessel sealing device with Atlas Wand (Covidien, Mansfield, MA). (b) Intra-abdominal picture of LigaSure application.
Surgical stapling equipment has routinely been used to achieve hemostasis during ovariectomy performed via ventral celiotomy approaches. When applied across the ovarian pedicle, the TA-90 (ThoracoAbdominal Stapler, Covidien, Manchester, MA) produces effective hemostasis. The development of endolaparoscopic stapling equipment has allowed similar applications in laparoscopic surgery. The EndoGIA has been used to simultaneously staple and transect the ovarian pedicle.17 Prior to application of the stapling device, the ovarian pedicle was dissected, as described above for placement of ligatures, to reduce the mass of the vessel containing part of the pedicle. In some cases, part of the pedicle escaped the device,
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Laparoscopy requiring repositioning. There were instances where electrocoagulation was used to augment hemostasis. Though effective, the use of this device requires a larger diameter cannula and pedicle dissection, and staple cartridges add significantly to the cost of the procedure.
Removing the ovary from the abdomen in the standing horse When performing bilateral ovariectomy in the standing horse, most surgeons begin on the left side. The left ovary can be removed at the point that it becomes free in the abdomen by enlarging one of the instrument portals. When that is done, abdominal insufflation is lost and the enlarged incision must be closed to allow insufflation to be re-established before proceeding to the right side of the abdomen. For this reason, when the left ovary has been freed from its attachments, surgeons commonly leave it attached to an instrument and move to the right side to proceed with transection of the right ovarian pedicle. When both ovaries have been freed, they can be removed from their respective sides after enlarging one of the instrument portals. To lessen the chance of dropping the ovary during removal, an acute claw grasper is used (Figure 211.5). The complication of dropping the ovary in the abdomen or having the ovary lodge part-way through the abdominal wall can largely be avoided by sharply incising the fascia of the external abdominal oblique muscle and using finger dissection to enlarge the opening in the peritoneum. Alternatively, the right ovary can be passed under the mesocolon to the left side of the abdomen and then both ovaries can be removed through a single enlarged instrument portal. In this case, insufflation is lost with removal of the first ovary, but can be temporarily restored by blocking the outflow by inserting coned fingers into the enlarged incision. This allows visual observation of the transfer of the second ovary to the instrument placed through the enlarged incision. The use of a large disposable tissue cannula has been described (Ethicon EndoSurgery, Johnson & Johnson Co., New Brunswick, NJ), but the 33-mm diameter is often too small to accommodate the ovary. In the authors’ experience, this large cannula frequently is spontaneously expelled from the distended abdomen, due to its short length, unless an assistant physically holds it in place. Incisions that were enlarged to allow removal of the ovaries should be closed in two to three layers (external abdominal oblique fascia, subcutaneous tissue in some cases, and skin). Skin sutures are sufficient for the other incisions.
1997
Hand-assisted ovariectomy in the standing horse Laparoscopic removal of granulosa thecal cell tumors has been reported in both the recumbent horse, using ligating loops for hemostasis,18 and the standing horse, where a linear staple instrument19 or a vesselsealing device were used.20 Both approaches were effective, and many surgeons now remove granulosa thecal cell tumors using electrocoagulation techniques. Irrespective of the laparoscopic approach, manipulation of the enlarged ovary can be difficult. A hand-assisted laparoscopic technique has been developed to facilitate removal of granulosa thecal cell tumors in standing mares.21 The paralumbar fossa on the side of the affected ovary is prepared for laparoscopic surgery. An inverted ‘L’ block or direct infiltration with local anesthetic along the proposed incision is performed. A 12–20-cm vertical skin incision is made at the cranialto-caudal midpoint of the paralumbar fossa. The incision begins 3–6 cm ventral to the dorsal margin of the internal abdominal oblique muscle and extends ventrally. The external abdominal oblique muscle is sharply divided and the internal abdominal oblique muscle and the transverse abdominis muscle are divided in the direction of their fibers. The peritoneum is punctured and enlarged to allow placement of a hand into the abdomen. A 1.5-cm skin incision is made immediately dorsal to the internal abdominal oblique muscle, and a laparoscopic cannula with a blunt trocar is introduced into the abdomen while using the intra-abdominal hand to guard against visceral injury. The trocar is withdrawn and a 10-mm laparoscope is inserted. Ovarian pedicle anesthesia is achieved under laparoscopic observation using a needle placed through the large abdominal incision. Hemostasis is achieved using a 90-mm thoracoabdominal staple instrument with 4.8-mm staples (TA90 stapler; United States Surgical Corp., Norwalk, CT). The jaw end of the stapler is placed into the abdomen and the handle remains outside. The stapler is manipulated over the ovarian pedicle in a caudal-tocranial direction and closed on the tissue. The laparoscope is used to visually confirm correct placement of the device and the staples are fired. If the entire pedicle was not enclosed in the jaws, an overlapping line of staples is placed in a cranial-to-caudal direction. At this point, the ovarian pedicle is severed distal to the line of staples using laparoscopic scissors introduced through the incision and observed through the laparoscope. Any residual hemorrhage is controlled with ligating clips or electrocoagulation. When the ovary is free, it is removed from the abdomen. Removal of particularly large ovaries is facilitated by placing them in a tissue bag before removal from
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the abdomen. Tissue macerators are available but are not commonly used in veterinary medicine. Ovaries with large cysts are easier to remove if they are decompressed with a needle before removal. One of the authors (D.A.H.) has removed ovaries up to 18 cm in diameter using a standard laparoscopic approach and a LigaSure for vessel ligation and amputation. The laparoscope incision is closed with skin sutures. The individual layers of the larger abdominal incision are closed with simple continuous sutures using absorbable sutures.
Laparoscopic ovariectomy in the recumbent mare Laparoscopic ovariectomy can be performed with the mare in dorsal recumbency. This approach ensures patient cooperation and manipulation of instruments is easier owing to the separation between insertions. In addition, bilateral manipulation can be performed. Unfortunately, locating the ovaries is more difficult than in the standing horse, and gaining access to the ovarian pedicle can be complicated by overlying viscera. Mares should be fasted for 48 hours before surgery. With the horse under general anesthesia, the ventral abdomen is clipped and prepared for aseptic surgery. A 1-cm skin incision is made immediately caudal to the umbilicus and a #15 blade is used to make a stab incision through the linea alba. A teat cannula is inserted into the abdomen and the abdomen is insufflated to a pressure of 15–20 mmHg. The teat cannula is withdrawn and a laparoscopic cannula with a sharp trocar is inserted into the abdomen. For this first insertion, one author (D.G.W.) prefers a system with a guarded trocar (Figure 211.13) while the other author uses a blunt trocar similar to standing flank placement. The trocar is removed and the laparoscope is inserted. When intra-abdominal positioning of the scope is confirmed, the hindquarters are elevated into the Trendelenburg position to facilitate cranial displacement of the viscera. Instrument cannulas are placed approximately 15 cm lateral to the midline at a level approximately 10 cm cranial to the udder. Instrument cannulas should be placed
Figure 211.13 Laparoscopic cannula (12.5-mm diameter) 15 cm long with safety trocar (Dr Fritz, LLC Louisville, KY).
under direct laparoscopic visual observation to avoid inadvertent injury to the viscera. Placement of the cannula into the abdomen in an area where there does not appear to be retroperitoneal fat may reduce the incidence of vascular injury during insertion. Using a blunt trocar during this insertion process also reduces the likelihood of vascular injury. Two instrument cannulas are usually sufficient if electrocoagulation is used for hemostasis. If the ovary is not in view, the body of the uterus should be located dorsal to the bladder and a hand-over-hand manipulation using two grasping instruments is used to locate the ovary. If the surgeon chooses to use ligating loops, two additional instrument cannulas facilitate the process.22 These cannulas are inserted 15 cm off the midline midway between the umbilicus and the initial instrument cannulas. In this case, hemostasis is achieved with block ligation of the ovarian pedicle.22 Prior to moving to electrocoagulation, David G. Wilson performed dissection similar to that described for standing ovariectomy with ligature to ensure ligature stability. With the development of electrocoagulation techniques for hemostasis, the use of ligatures has been largely abandoned. One of the instrument cannulas is enlarged and the ovaries are routinely removed. An acute claw grasper is used to reduce the likelihood of dropping the ovary during the removal process. The enlarged incision is routinely closed in three layers (external rectus fascia, subcutaneous tissue, and skin). The other cannula incisions are most often closed with skin sutures; however, the authors prefer to place a single cruciate suture in the rectus fascia or linea alba before placing skin sutures.
In situ coagulation and transection of the ovarian pedicle Removing the ovary from the abdomen during laparoscopic ovarietomy can be difficult and on occasion ovaries may be dropped and lost necessitating open laparotomy to allow the misplaced ovary to be recovered. Shoemaker and co-workers described a technique whereby the ovarian pedicle was laparoscopically transected and the freed ovaries were released into the abomen.23 In juvenile horses, the avascular ovary most frequently became adhered to the omentum in the ventral aspect of the abdomen. Ovarian parenchyma was necrotic and all 10 horses were effectively ovariectomized. There were no surgical complications. To date, this technique has not found favor in the surgical community; however, several surgeons have commented that given the knowledge that transected ovaries undergo necrosis,
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Laparoscopy the occasional misplaced ovary can now simply be left in the abdomen.
In situ ovariectomy In situ ovariectomy has been described by Rijkenhuizen.24 Ligatures are placed around the ovarian pedicle with a goal of creating an avascular ovary. Though potentially effective, consistent occlusion of both the ovarian artery and vein is difficult. Furthermore, mares subjected to this technique often exhibit colic during the immediate postoperative period. For these reasons, in situ ovariectomy has not found favor in the surgical community.
Other uses of laparoscopy Laparoscopy has been described for oviductal ligation to prepare mares for gamete intrafallopian transfer.25 Mares underwent standing laparoscopy for unilateral oviductal ligation. They were then followed to monitor ovulation and breeding from each ovary. None of the mares became pregnant when they ovulated on the side of the ligation. This is a simple and straightforward technique. The same approach as for standing ovariectomy is used. The oviduct is identified, and two ligating clips are applied. Scissors are used to transect the oviduct between the ligating clips. The skin is closed with size 2–0 nylon using a cruciate pattern. Another approach that shows promise is for laparoscopic oocyte or embryo transfer. While the technique needs some refinement, it may offer an approach that allows multiple surgical incisions on recipient mares without the resultant large scars that occur after flank laparotomy.
Summary Since their beginnings in the 1990s, laparoscopic ovariectomy techniques have evolved to a point when laparoscopic ovariectomy is now favored over traditional open approaches. Whether the procedure is performed in the standing or recumbent mare, the preferred method of hemostasis is now electrocoagulation, and the best bipolar cautery implement available at the present time is the capacitance-controlled vessel-sealing device (LigaSure). In the hands of experienced laparoscopists, these minimally invasive approaches have a low complication rate and horses can return to their intended uses significantly sooner than when ovariectomy is performed using traditional open approaches.
1999
References 1. Wilson GL. Laparoscopic examination of mares. Vet Med Sm Anim Clin 1983;78:1629–33. 2. Palmer SE. Standing laparoscopic laser technique for ovariectomy in five mares. J Am Vet Med Assoc 1993;203:279–83. 3. Peroni JF, Rondenay Y. Analgesia and anesthesia for equine laparoscopy and thoracoscopy. In: Fisher AT (ed.) Equine Diagnostic and Surgical Laparoscopy. Philadelphia: WB Saunders, 2002; pp. 119–28. 4. Hendrickson DA. Minimally invasive surgery of the reproductive system in large animals. In: Freeman LJ (ed.) Veterinary Endosurgery. Baltimore: Mosby, 1998; pp. 217–25. 5. Desmaizieres LM, Martinot S, Lepage OM, Bareiss E, Cador´e JL. Complications associated with cannula insertion techniques used for laparoscopy in standing horses. Vet Surg 2003;32:501–6. 6. Farstvedt EG, Hendrickson DA. Intraoperative pain responses following intraovarian versus mesovarian injection of lidocaine in mares undergoing laparoscopic ovariectomy. J Am Vet Med Assoc 2005;227:593–6. 7. Boure L, Marcoux M, Laverty S. Paralumbar fossa laparoscopic ovariectomy in horses with use of endoloop ligatures. Vet Surg 1997;26:478–83. 8. Rodgerson DH, Hanson RR. Ligature slippage during standing laparoscopic ovariectomy in a mare. Can Vet J 2000;41:395–7. 9. Hanson CA, Galuppo LD. Bilateral laparoscopic ovariectomy in standing mares: 22 cases. Vet Surg 1999;28:106–12. 10. Shettko DL, Frisbie DD, Hendrickson DA. A comparison of knot security of commonly used hand-tied laparoscopic slipknots. Vet Surg 2004;33:521–4. 11. Carpenter EM, Hendrickson DA, Franke C, Frisbie D, Trostle S, Wilson D. A mechanical study of ligature security of commercially available pre-tied ligatures versus hand tied ligatures for use in equine laparoscopy. Vet Surg 2006;35:55–9. 12. Cokelaere SM, Martens AM, Wiemer P. Laparoscopic ovariectomy in mares using a polyamide tie-rap. Vet Surg 2005;34:651–6. 13. Rodgerson DH, Belknap JK, Wilson DA. Laparoscopic ovariectomy using sequential electrocoagulation and sharp dissection of the equine mesovarium. Vet Surg 2001;30:572–9. 14. Hand R, Rakestraw P, Taylor T. Evaluation of a vesselsealing device for use in laparoscopic ovariectomy in mares. Vet Surg 2002;31:240–4. 15. Dusterdieck KF, Pleasant RS, Lanz OI, Saunders G, Howard RD. Evaluation of the Harmonic Scalpel for laparoscopic bilateral ovariectomy in standing horses. Vet Surg 2003;32:242–50. 16. Alldredge JG, Hendrickson DA. Use of high-power ultrasonic shears for laparoscopic ovariectomy in mares. J Am Vet Med Assoc 2004;225:1578–80. 17. Van Hoogmoed LM, Galuppo LD. Laparoscopic ovariectomy using the Endo-GIA stapling device and Endo-Catch pouches and evaluation of analgesic efficacy of epidural morphine sulfate in 10 mares. Vet Surg 2005;34:646–50. 18. Ragle CA, Southwood LL, Hopper SA, Buote PL. Laparoscopic ovariectomy in two horses with granulosa cell tumors. J Am Vet Med Assoc 1996;209:1121–4. 19. Rocken M. Laparoscopic cryptorchidectomy and ovariectomy in the standing horse. Praktische Tierarzt 1998;79:113–19.
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20. Hubert JD, Burba DJ, Moore RM. Evaluation of a vesselsealing device for laparoscopic granulosa cell tumor removal in standing horses. Vet Surg 2006;35:324–9. 21. Rodgerson DH, Brown MP, Watt BC, Keoughan CG, Hanrath M. Hand-assisted laparoscopic technique for removal of ovarian tumors in mares. J Am Vet Med Assoc 2002;220:1503–7. 22. Ragle CA, Schneider RK. Ventral abdominal approach for laparoscopic ovariectomy in horses. Vet Surg 1995;24:482–97.
23. Shoemaker RW, Read EK, Duke T, Wilson DG. In situ coagulation and transection of the ovarian pedicle: an alternative to laparoscopic ovariectomy in juvenile horses. Can J Vet Res 2004;68:27–32. 24. Rijkenhuizen A, Jonker H. Laparoscopic castration of the mare by ligation of the blood supply to the ovaries. Vet Surg 2000;29:283. 25. McCue PM, Hendrickson DA, Hess MB. Fertility of mares after unilateral laparoscopic tubal ligation. Vet Surg 2000;29:543–5.
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Section (g): Diagnostic Ultrasonography
212.
Normal Anatomy Elaine M. Carnevale and Lawrence M. Olsen
2003
213.
Folliculogenesis Mohd A. Beg and Don R. Bergfelt
2009
214.
Ovulation: Part 1. Follicle Development and Endocrinology During the Periovulatory Period Eduardo L. Gastal
2020
215.
Ovulation: Part 2. Ultrasonographic Morphology of the Preovulatory Follicle Eduardo L. Gastal
2032
216.
Luteal Development Don R. Bergfelt and Gregg P. Adams
2055
217.
Pregnancy Don R. Bergfelt and Gregg P. Adams
2065
218.
Equine Fetal Sex Determination Between 55 and 150 Days Richard D. Holder
2080
219.
Fetal Gender Determination from Mid to Advanced Gestation Stefania Bucca
2094
220.
Management of Twins Angus O. McKinnon
2099
221.
Early Embryonic Loss Dirk K. Vanderwall
2118
222.
Ovarian Abnormalities Patrick M. McCue and Angus O. McKinnon
2123
223.
Uterine Abnormalities Angus O. McKinnon and Patrick M. McCue
2137
224.
Mammary Gland Bryan M. Waldridge
2162
2001
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Chapter 212
Normal Anatomy Elaine M. Carnevale and Lawrence M. Olsen
Ultrasonography of the reproductive tract Transrectal ultrasonography should be considered a routine part of a mare’s reproductive examination. This procedure provides valuable information concerning form and function of the reproductive tract. The transrectal ultrasonographic examination requires the same precautions as rectal palpation. The mare is restrained in a set of stocks or with a nose twitch and held by an experienced handler. Rectal palpation should immediately precede the ultrasound examination to ensure that the reproductive tract is oriented normally and fecal material is cleared from the rectum before insertion of the ultrasound probe.1 This manual examination facilitates assessment of tone and size of the uterus and ovaries as well as supportive structures.2 The ultrasound probe or transducer is lubricated and protected by the examiner’s hand to prevent trauma to the rectal wall. Good contact between probe and rectal wall is imperative for good image quality. The examiner should develop a repeatable, standard routine for the ultrasound examination so that all areas of the uterus, ovaries, and supportive structures are viewed to ensure that scanning errors do not occur.3 A common scanning routine might be anterior uterine body, right uterine horn, right ovary, right uterine horn, anterior uterine body, left uterine horn, left ovary, left uterine horn, uterine body, and cervix.4 To evaluate and interpret ultrasound images of the reproductive tract accurately, an understanding of reproductive physiology and knowledge of reproductive anatomy are necessary. Ultrasound images reflect the dynamic nature of ovarian hormones and the effect of these hormones on the physiological state of the uterus and ovaries. These images facilitate assessment of stage of the reproductive cycle in relation to follicular development, ovulation, and formation of the corpus luteum.
Most veterinary ultrasound machines designed for reproductive evaluations in animals utilize a linear-array (5- to 7.5-MHz bandwidth) transducer and many have adjustable bandwidth probes. The probe is usually held in a sagittal plane or parallel to the long axis of the horse, with the image of the cervix and uterine body longitudinally oriented. The image orientation of craniad on the right of the screen and caudad on the left of the screen is often used as the accepted standard. The direction of the image can be changed on most ultrasound machines; an arrow or circle usually marks the anterior aspect of the probe on the image.
Cervix and uterus The uterine horns are imaged in cross-section and the uterine body is imaged in a longitudinal section during the examination (Figure 212.1). The cervix is located caudal to the body of the uterus. The tone of the cervix reflects the endocrine status of the mare. The cervix can be palpated, and cervical tone can be graded: 1 (one-finger width, toned, diestrus or pregnant); 2 (two-finger width, moderate tone, early estrus or early diestrus or maiden mare); and 3 (three-finger width, relaxed, estrus). With ultrasound, the cervix is imaged in longitudinal section (Figure 212.2). During estrus the cervix is hypoechoic and may be difficult to differentiate from adjacent structures although echogenic lines are occasionally imaged. In contrast, the diestrous or pregnant cervix is imaged by horizontal echogenic parallel lines represented by the peripheral muscular boundaries.5 The vagina can be imaged caudad to the cervix as a thin horizontal line of moderate echogenicity; however, the vagina is usually not notable unless pathology, such as pooled urine or air, is present. The uterus of the mare changes in appearance with hormonal influences during the anovulatory
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 212.1 Cross-section of the uterine horn (left) and longitudinal section of the uterine body (right) in a diestrous mare. The bladder (bl) is imaged below the uterus. A small cyst is noted as an anechoic area in the uterine horn.
period, the estrous cycle, or pregnancy. These changes can be imaged by routine or serial ultrasonographic examinations. During anestrus, the crosssection of the uterine horns and the longitudinal section of the uterine body are often compressed or
Figure 212.2 The cervix (cx) is imaged in longitudinal section as a slightly more echogenic structure than the posterior uterine body (pub). The bladder (bl) is located ventral to the uterus. Note the arrow above the ultrasound image indicating that the anterior aspect of the transducer is facing to the right. Lines at the top and lefthand side of the image represent 10 mm.
flattened and irregular and may orient closely with the bladder or intestine (Figure 212.3). During estrus, the uterine horns and body commonly have an interdigitated pattern of alternating echogenic and anechogenic areas.4 The areas of altered echogenicity correspond to edema within the endometrial folds; the edema is an effect of estrogen secreted from developing ovarian follicles during estrus.6 Ultrasound images of the uterus during estrus have an echotexture that resembles a sliced orange, giving the uterine horns a ‘cartwheel’ pattern in cross-section (Figures 212.4 and 212.5). A change in endometrial edema during the estrous cycle is a normal physiological event. Edema in the endometrial folds can be observed at the end of diestrus, as progesterone declines and estrogen increases. The edema becomes pronounced during early estrus and diminishes or disappears 24 hours before ovulation.2 Echotexture of the endometrial folds can be graded from 0 (no ultrasonographic evidence of endometrial folds) to 3 (prominent endometrial folds).4 Declining edema can be used as a predictor of impending ovulation.4 Endometrial folds are less defined or not discernible (grade 0) during diestrus under the influence of progesterone secreted from the ovarian corpus luteum (Figure 212.1). The ultrasonographic image of the uterine lumen in the body of the uterus often appears as a hyperechogenic white line due to the apposition of the dorsal and ventral endometrial surfaces. During diestrus the uterus is well circumscribed and defined with ultrasound. The uterus is toned, and the cervix is closed.
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Normal Anatomy
2005
Figure 212.3 (a) The uterus (ut) during anestrus is flaccid with a poorly defined shape as it lies on the bladder (bl) and intestine (int). (b) An inactive ovary with no follicular development.
Figure 212.4 Ultrasound images from a mare in early estrus. The dominant follicle (left) is approximately 32 mm in diameter. Edema in the uterine body (right) is causing anechoic areas within the endometrial folds of the uterus; this pattern is characteristic of estrus.
Figure 212.5 Ultrasound images from a mare in late estrus. The dominant follicle can be measured by internal ultrasound calipers, and the corresponding measurements are present. The uterine horn is imaged in cross-section, with subtle edema and a slight texture.
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Mare: Diagnostic Ultrasonography
Ovaries The ovaries are dynamic, with changes ranging from development of large pre-ovulatory follicles to no follicular activity during anestrus3 (Figures 212.3b, 212.4 and 212.5). Ultrasonography is useful for rapid, real-time assessments of ovarian follicles and luteal structures. The right and left ovaries are routinely imaged as a continuation of the ultrasound examination of the uterus by rotating the probe slightly laterally from the end of the corresponding uterine horn. A more craniad position with the probe may have to be adopted in some mares depending on the individual positioning of the ovary, which should be evaluated during the initial rectal palpation. The mare ovary is kidney-bean-shaped with a prominent depression, the ovulation fossa, on the ventral concave border. Average size of the equine ovary is 50 × 30 × 30 mm, but considerable variation occurs with changes in follicular activity and stage of the reproductive cycle.5 The mare’s ovary is unusual compared to other animals in that the ovarian medulla is superficial and the cortex is interior except at the ovulation fossa, where ovulation occurs. A corpus luteum cannot readily be detected by palpation but can be imaged with ultrasound. The two major endocrine structures on the equine ovaries are follicles7 and the corpus luteum.8 Sequential monitoring of dynamic changes in a follicular population during the estrous cycle is a valuable management tool to follow cyclic activity and to predict time of breeding and ovulation. The follicles are fluid-filled, non-echogenic structures that appear as black and roughly circumscribed ultrasonographic images viewed within the confines of the ovarian stroma (see Figures 212.4 and 212.5). During anestrus,
inactive ovaries are readily differentiated from active ovaries by the absence of a corpus luteum and small or no developing follicles (Figure 212.3b). During the period of hormonal transition, the ovaries may have multiple, large follicles (Figure 212.6). In the cyclic mare, the approximate follicular growth rate is 3 mm per day.9 The size of the follicle at ovulation varies, although many light-horse mares will ovulate a follicle of approximately 45 mm in diameter. As ovulation approaches, pre-ovulatory follicles become soft and painful. A thickening of the follicular wall can be imaged, and the follicle will often change shape, from spherical to oblong or pearshaped (Figure 212.7). Ovulation can be determined with ultrasound with the disappearance of the large, fluid-filled, non-echogenic structure (follicle) and the appearance of an echogenic area (ovulation site).10 Immediately after ovulation, the ovulation site may be amorphous and difficult to image (Figure 212.8). Upon palpation the ovary is usually painful, and an edematous indentation may be felt. The corpus luteum early in diestrus will often be hyperechoic (Figure 212.9). The corpus luteum is echogenic because of its tissue-dense structure, formed from packed luteal cells producing progesterone. Approximately 50% of ovulations result in the formation of an immediate corpus luteum with uniform echogenicity, and 50% of ovulations result in the formation of a corpus hemorrhagicum with a central non-echogenic core for a short period.8 The mature corpus luteum usually has a well-delineated shape and is moderate in echogenicity (Figure 212.10a). The corpus luteum is located centrally in the equine ovary; therefore, palpation of the structure is unsuccessful, although the uterus and cervix should have good tone consistent with diestrus. Growing and regressing follicles are
Figure 212.6 Multiple small follicles are found on the ovaries of mares during the spring transition. Uterine edema is inconsistent, and the mares will often show periods of estrus without ovulation.
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Figure 212.7 The appearance of the follicle changes as it approaches ovulation. The follicle on the left is slightly oblong with a thickened border. The thickening of the follicular wall occurs with changes in the granulosa layer of the follicle; the darker ring surrounding the follicle represents a vascular layer. The follicle on the right is irregular in shape, suggestive of imminent ovulation.
Figure 212.8 Immediately after ovulation, ovulation sites (arrows) may be difficult to image, because the shape and echogenicity of the site is poorly differentiated from ovarian stroma and surrounding tissue.
Figure 212.9 One day after ovulation, this developing corpus luteum (arrows) is hyperechoic with a projection (large arrow) toward the ovulation fossa. The uterus (right image) has a homogeneous echotexture.
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Figure 212.10 (a) The mature corpus luteum (left) is moderately echoic with defined borders. A large, diestrous follicle (f) is present on the same ovary. (b) At the end of diestrus, the regressing corpus luteum is associated with a decrease in size. An anechoic border and hyperechoic central area may be observed.
consistently observed on the ovary during diestrus. The corpus luteum will regress at the end of diestrus, usually in association with a growing population of follicles. The regressing corpus luteum becomes smaller in size (Figure 212.10b). A small, hyperechoic structure, the corpus albicans, may be imaged as the non-functional remnant of the corpus luteum.
Conclusions Transrectal ultrasonography provides a method for real-time imaging of the mare’s reproductive tract. Ovarian structures and uterine morphology can be assessed to determine stage the estrous cycle.
References 1. Torbeck RL. Diagnostic ultrasound in equine reproduction. Vet Clin N Am-Equine 1986;2:227–52. 2. McKinnon AO, Carnevale EM. Ultrasonography. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, PA: Lea & Febiger, 1993; pp. 211–20.
3. Ginther OJ. Techniques: Orientation of the reproductive tract. In: Ginther OJ (ed.) Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains, WI: Equiservices Publishing, 1986; pp. 99–113. 4. McKinnon AO. Reproductive ultrasonography. In: Rantanen NR, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore, MD: Williams and Wilkins, 1998; pp. 79–102. 5. Sertich PL. Ultrasonography of the genital tract of the mare. In: Reef VB (ed.) Equine Diagnostic Ultrasound. Philadelphia, PA: WB Saunders Co, 1998; pp. 405–24. 6. Ginther OJ, Pierson RA. Ultrasonic anatomy and pathology of the equine uterus. Theriogenology 1984;21:505–15. 7. Pierson RA, Ginther OJ. Ultrasonic evaluation of the preovulatory follicle in the mare. Theriogenology 1985;24:359–68. 8. Pierson RA, Ginther OJ. Ultrasonic evaluation of the corpus luteum of the mare. Theriogenology 1985;23:795–806. 9. Ginther OJ. Characteristics of the ovulatory season. In: Ginther OJ (ed.) Reproductive Biology of the Mare. Cross Plains, WI:Equiservices Publishing, 1992; pp. 173– 232. 10. McKinnon AO, Squires EL, Voss JL. Ultrasound evaluation of the mare’s reproductive tract – Part I. Comp Cont Educ Pract 1987;9:336–49.
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Chapter 213
Folliculogenesis Mohd A. Beg and Don R. Bergfelt
Introduction The ovarian follicles in mares were first visualized in situ and studied with ultrasound in 1980.1 Since then, the application of real-time ultrasonography as a non-invasive, repeatable research and clinical tool has significantly advanced our understanding of ovarian function in domestic farm species, especially the mare. The sequential monitoring of growth and regression of individual follicles in the same animal has opened new areas of research and led to discoveries into the nature of folliculogenesis. For example, the real-time image obtained by ultrasonography has been used as a guide for intrafollicular injection, follicular fluid sampling, ovarian biopsy, and collection of granulosa cells and ova for basic and applied purposes (see review by Beg and Ginther2 ). Recently, more refined and improved techniques of ultrasonography such as computer-assisted image analysis and mathematical modeling; duplex ultrasonography; three- and fourdimensional ultrasonography; spectral, color-flow, and power Doppler ultrasonography; and photoacoustic imaging and contrast-enhanced ultrasonography have become available (see http://en.wikipedia. org/wiki/Medical ultrasonography). Although this chapter primarily focuses on the use of B-mode and color-Doppler ultrasonography, these exciting new capabilities have the potential to bridge the gap between structure and function of ovarian dynamics, and act as a catalyst for research activities to enhance further our understanding of follicular development during various reproductive states and improve clinical and breeding farm management practices in mares. The ovaries of mares, like all farm species, contain a large reserve of primordial follicles and a more limited population of preantral and antral follicles. In general, a young adult mare may have a pool of about 35 000 primordial or non-growing follicles
and 100 growing follicles in each of her ovaries.3 The comparable number in other monovulatory species is about 120 000 non-growing and 350 growing follicles in cows,4 and about 300 000 non-growing follicles in women.5 A primordial follicle takes several months and a preantral follicle requires several weeks to grow to a pre-ovulatory size.6 The study of folliculogenesis can be partitioned into three main periods in the reproductive life of an individual – fetal, prepubertal, and adult. The fetal and prepubertal periods will be discussed briefly to provide context for the adult period, which is the reproductive life-stage most conducive for the use of ultrasonography in the mare. Although several reviews6–11 and recent publications12–16 are available on the use of ultrasonography to detect and monitor follicular development during the seasonal transitional periods, estrous cycle, and pregnancy in the mare, the intent of this chapter is to compile and provide an overview of that information with emphasis on the principal aspects of folliculogenesis to create a single contemporary source to aid equine practitioners, clinicians, breeding-farm managers, and researchers in reproductive management and discovery.
Technique The procedures and techniques involved in detecting, measuring, and monitoring follicular structures during an ultrasound examination of the ovaries in mares have been thoroughly validated, described, and reviewed in several publications.17–22 Briefly, transrectal ultrasonography is the route of choice for detecting ovarian structures in mares because of the ovary’s close approximation to the rectum, and the large diameter and good maneuverability per rectum for presentation to ensure a clear image and accurate measurements. Transcutaneous (abdominal or flank) scanning of the ovaries is of limited value due to the distance between the ultrasound transducer
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 213.1 Ultrasonogram of a 25-mm follicle taken with B-mode (panel A) or Doppler mode (panel B) ultrasonography. Note the shape change primarily as a result of impingement of another follicle from above. The red color signals in the follicle wall (panel B) are localized to the theca cell layer and represent blood flow. (See color plate 97).
and ovaries in the abdominal cavity and intervening intestinal structures that cannot be readily cleared from the path of scanning. Antral follicles appear on an ultrasound monitor as distinct, black (anechoic), circumscribed structures. The ultrasound morphology of an equine ovarian follicle is shown in Figure 213.1 using B-mode and color-Doppler ultrasonography. A single follicle is usually spherical, but maturity and the presence of multiple follicles or a corpus luteum can impinge on adjacent follicles creating irregular shapes. Follicle size is generally expressed as a measure of the cross-sectional diameter that can be estimated using the incremental grid on the ultrasound monitor; alternatively, electronic built-in clippers can be used. From a still or frozen image of a follicle’s maximum cross-sectional image, an average of two measurements, usually perpendicular to one another, are taken across the antrum, beginning at the interface of the echogenic follicle wall and non-echogenic follicular fluid, from one side of the follicle to the other. For follicles that are less spherical, an average of the greatest distances across the antrum in two directions is determined. The minimal diameter that can be accurately measured on a regular basis is approximately 2 mm using a 5.0–7.5-MHz transducer; therefore, primordial, preantral, and early antral follicles are beyond the scope of current ultrasound machines used in veterinary medicine. Depending on the objective, follicle size may also be expressed in terms of volume or surface area to represent better the functional and secretory activities. Although follicle volume and surface area can be calculated as a function of diameter of a sphere by a formula (4/3πr3 for volume and 4πr2 for surface area; r = radius), most contemporary ultrasound machines have a built-in function to calculate the volume and surface area of a structure.
In addition to diameter, the most common end points that are recorded during an ultrasound examination to determine follicular development with B-mode ultrasonography are thickness of the follicle wall, thickness of the granulosa cell layer, echogenicity of the granulosa layer, and prominence and extent of circumferential involvement of the anechoic layer beneath the granulosa.17, 18 With color-Doppler ultrasonography, end points such as peak systolic velocity, time-averaged maximum velocity, end-diastolic velocity, resistance index, and pulsatility index are some of the most common measurements (built-in) taken for evaluating blood flow associated with follicle development in the mare.19 In addition, the percentage of follicle wall area with blood flow is either recorded subjectively as a percentage of color signals in the follicle wall from an image on the ultrasound monitor, or objectively by capturing a frozen colorDoppler image of the follicle using a video-editing program. Color spots or pixel aggregates are selected from the images, extracted and saved using an imageediting program as described by Ginther and Utt.23 ImageJ software (National Institutes of Health, USA; http://rsb.info.nih.gov/ij/) is one such program that is free to the public and can be used for calculation of the total number of colored pixels for each image or selected aspects of an image.
Definitions The terminology that is commonly used to describe the morphology and physiology of follicular dynamics since the advent of ultrasonography in mares has been presented in several publications and texts cited throughout this chapter, and is summarized in Table 213.1.
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Table 213.1 Terminology commonly used in association with follicular dynamics in mares Characteristic
Definition
Follicular wave
A cohort or group of antral follicles (mean 7 to 11) that are first detected (≥ 6 mm in diameter) within 1 to 2 days of one another and increase in diameter similarly until follicle deviation
Follicle or wave emergence
The earliest ultrasonic detection of follicles (≥ 6 mm) of a wave compatible with tracking individual growing and regressing follicles in a reliable and consistent manner. Emergence of a wave is a useful reference point for aligning the emergence of waves that occur at different times among mares
Common growth phase
The period from emergence of a major follicular wave to beginning of deviation (6 to 7 days). All follicles of the wave grow at a similar rate (about 3 mm/day) during the common growth phase
Follicle deviation
The dissociation in diameter between the largest follicle and next largest follicles of a major wave that begins, on average, when the largest follicle reaches 22.5 mm. The largest follicle continues to grow whereas the next largest follicles cease growth and regress after the beginning of deviation. The beginning of deviation is calculated to occur at the ultrasound examination preceding the first examination with an apparent change in diameter difference between two largest follicles. The beginning of deviation is a useful reference point for aligning the growth profiles of follicles of a wave for comparing structural and functional characteristics within and among mares
Dominant follicle
The largest follicle of a major wave that continues growth after deviation, reaches ≥ 28 mm and becomes ovulatory or anovulatory
Subordinate follicles
The next largest follicles of a major wave relative to a dominant follicle, that cease growth after deviation, reach < 28 mm and eventually regress
Dominance phase
The period from the beginning of deviation until the beginning of the pre-ovulatory phase (approximately 7 to 8 days). Growth of all subordinate follicles of the wave ceases during this period and typically no new follicular waves emerge
Pre-ovulatory phase
The period beginning when the dominant follicle of the major primary wave attains ≥ 30 mm and until ovulation or formation of a hemorrhagic follicle. The pre-ovulatory follicle attains the developmental capacity to respond to an endogenous or exogenous ovulatory stimulus
Major follicular wave
A follicular wave that develops at least one dominant follicle (≥ 28 mm) and corresponding subordinate follicles (< 28 mm) during the transitional period, estrous cycle, and early pregnancy
Minor follicular wave
A follicular wave that does not develop a dominant follicle. Instead, the largest follicle of the wave reaches < 28 mm during the transitional period, estrous cycle, and early pregnancy
Primary follicular wave
A major follicular wave that emerges during late diestrus where the dominant follicle becomes pre-ovulatory and results in the primary ovulation at the end of estrus
Secondary follicular wave
A major wave that emerges during late estrus of the previous estrous cycle or early diestrus where the dominant follicle either is anovulatory and regresses or forms an anovulatory hemorrhagic follicle or results in secondary ovulation during mid-diestrus
Folliculogenesis during the fetal period Sexual differentiation in the developing equine embryo appears to occur between 39 and 45 days of gestation (see review by Ginther24 ). The gonads at this time are in the form of germinal ridges on the surface of the mesonephros. By 70 to 80 days of gestation, large numbers of oogonia and oocytes are present in the fetal ovaries,25, 26 and between 100 to 120 days there is a massive amount of germ-cell degeneration, which slows considerably by about 150 days, concomitant with the first appearance of primordial follicles. The follicle-contained oocytes are soon arrested in meiotic prophase by about 180 and 200 days of gestation and, thereafter, primordial follicles may occasionally grow and increase to antral
diameters of about 200 µm. However, antral size follicles greater than 1 mm are rarely seen in prenatal life or until about 15 to 21 days of neonatal life, when they may reach 3 to 4 mm. Apart from research applications, ultrasonography has limited diagnostic value for detecting and monitoring follicular dynamics during fetal and early neonatal life. Moreover, logistical application of ultrasonography would be limited and impractical at this time.
Folliculogenesis during the prepubertal period From a slaughterhouse study of excised ovaries involving 69 filly ponies ranging in age from 4 to 21 months, the average diameter of the largest
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follicle did not exceed 10 mm during the first summer through the first winter.27 By the second summer and fall, when the fillies were over 1 year of age, almost all of the fillies had developed an ovulatory follicle and had ovulated, as indicated by evidence of apparent luteal glands within the ovaries. The first ultrasound study in prepubertal mares involved 10 pony fillies beginning from a mean age of 35 days until 12 months.28 Ultrasonic imaging resulted in the detection and monitoring of antral follicles during the prepubertal period ranging from 6 to 9 mm in diameter in 6 of 10 fillies at a mean of 35 days after birth. More than 6 weeks later, follicles greater than 6 mm were observed in the remaining four fillies. Follicles greater than 10 mm were not observed until 2 months of age but started appearing in some fillies at 3 or 4 months. By about 10 months of age, a mean of seven follicles ranging from 10 to 14 mm in diameter and a mean of three follicles greater than 14 mm were detected. Curiously, the 10 largest follicles were found more frequently in the left than in the right ovary. A monthly wave pattern of follicle growth was identified in all fillies; however, successive waves did not partition into dominant and subordinate follicles (see Table 213.1), as indicated by small differences in diameter between the two largest follicles of a wave. Hence, follicular wave development during the prepubertal period appeared to be similar to minor follicular waves (Table 213.1) that occur during the estrous cycle, pregnancy, and fall and spring transition in adult mares.29 Near the end of the prepubertal period or during the peripubertal period, 12 to 15 months after birth,28 follicular wave development appeared comparable to that preceding the first ovulation of the season in adult mares such that there was successive development of follicles greater than 30 mm in 50% of the fillies, which eventually resulted in ovulations. Corresponding to the spring transitional period, there was a prolonged period of about 21 days in which follicles > 25 mm in diameter were present prior to the first ovulation in fillies. At the time of ovulation, the diameter of the pre-ovulatory follicle ranged from 30 to 47 mm, with a mean of 37 mm. Thus, the onset of puberty as marked by the first ovulation in the reproductive life of fillies occurred, on average, 346 days or about 11.5 months after birth. Ultrasonographic examination of the reproductive organs in fillies or yearlings may not be warranted for immediate breeding purposes but may be required to clarify a healthy and sound reproductive tract or to establish that puberty has been reached prior to sale and for future breeding purposes. In regard to the latter, however, there may be logistical challenges
for the use of ultrasonic imaging in young mares by the clinician. In general, it was found28 that after 10 months of age, the ultrasound transducer may be accompanied and guided by the hand into the rectum, whereas before 10 months the transducer may be affixed to a semirigid material for external manipulation of the transducer similar to that used for prepubertal cattle,30 sheep,31 goats,32 and llamas.17
Folliculogenesis during the adult period Estrous cycle The onset of puberty (first ovulation 12 to 15 months after birth) is a milestone event that leads to sexual maturity characterized by successive and continuous estrous cycles that may be interrupted by season, pregnancy, senility, or pathology of the reproductive tract. The estrous cycle, or more precisely the interovulatory interval, begins at ovulation associated with an estrus and ends at ovulation associated with the next estrus. The mean length of an interovulatory interval is typically 22 or 24 days in horses or ponies, respectively. The estrous cycle or interovulatory interval is characterized behaviorally by diestrus (12 to 16 days) and estrus (5 to 9 days) or, physiologically, by luteal and follicular phases (for review see Ginther24 ). The introduction of ultrasonography in the 1980s for detecting and tracking individual antral follicles > 5 mm in diameter confirmed that in a majority of the estrous cycles a single major follicular wave develops.33–36 In general, and in reference to Table 213.1, a follicular wave refers to a cohort or group of follicles that are first detected within 1 to 2 days of one another37 and increase in diameter similarly until follicle selection or regression occurs. Profiles of day-to-day changes in follicle diameters in various diameter categories,34 and tracking of individual follicles35, 36 with ultrasonography have established that the ovulatory follicle originates from a major follicular wave that is first detected during late diestrus or 13 to 14 days before ovulation. In a major wave, the largest follicle reaches ≥ 28 mm in diameter and is referred to as a dominant follicle, whereas in minor waves the maximum diameter of the largest follicle is < 28 mm.29, 38, 39 Major follicular waves include primary and secondary waves (Table 213.1). A primary major wave emerges during late diestrus and gives rise to a dominant follicle that becomes a pre-ovulatory follicle, and a secondary major wave emerges before the last ovulation of the previous interovulatory interval and gives rise to a dominant follicle that is either anovulatory and regresses, or forms an anovulatory hemorrhagic follicle, or ovulates.29 In
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MARE A Ovulatory Major anovulatory Minor
ov
MARE B ov
2013
ov
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ov
MARE D
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ov
ov
35 25 15 5 2 4 6 8 10 12 14 16 18 20 22 24
-2 0 2 4 6 8 10 12 14 16 18 20
Days from first ovulation of interovulatory interval Figure 213.2 Combinations of major and minor follicular waves during an interovulatory interval in four mares as illustrated by day-to-day diameters of the largest follicles of each wave type. Note, most secondary major anovulatory waves emerge before the last ovulation of the previous interovulatory interval and minor waves emerge before or after ovulatory waves. Ov = ovulation. Adapted from Ginther et al.9
regard to the latter, the incidence of secondary ovulations during diestrus varies widely among breeds, from 18 to 21% in Standardbreds and Thoroughbreds, 4% in Trotters, and close to 0% in Quarter Horses, ponies,24 and Bretons.39 Other ultrasonographic differences between a major primary and secondary follicular wave are that with a secondary major wave the mean maximum diameter of the dominant follicle is smaller (37 vs 46 mm) and the uterus remains characteristic of diestrus.29 Examples of the types of follicular waves in four individual mares are illustrated in Figure 213.2 using the day-to-day changes in diameters of the largest follicle of each wave type. While a single major primary wave is characteristic of most estrous cycles, some breeds exhibit a major secondary wave or minor wave before the primary wave. In ponies, usually only the major primary wave is detected, while in Quarter Horses and Brazilian Bretons a major secondary anovulatory wave has been detected 24%29 and 20%39 of the time, respectively. An even higher frequency of major secondary waves may be observed in Thoroughbreds. Once follicles leave the primordial or ‘resting’ pool, they progress through a series of developmental changes before reaching the pre-ovulatory stage. The entry of follicles into a growth and differentiation phase is an irreversible commitment that ends in either atresia or ovulation. Atresia or regression is the fate of about 99% of all follicles. The mean daily diameter profiles for the six largest follicles
without regard to individual identity during an interovulatory interval in mares is shown in Figure 213.3. Although the non-identity approach resulted in a unimodal follicular wave profile, minor or secondary major waves could have been masked by averaging the diameters of regressing and growing follicles that overlap between minor or secondary waves preceding the primary waves. The development of antral follicles as detected ultrasonographically can be divided into functional divisions for describing the events that occur during the estrous
40 Diameter (mm)
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35 30 25 20 15 10 0
2
4
6 8 10 12 14 16 18 20 22 0 Days after ovulation
Figure 213.3 Mean (± SEM) profile of the six largest follicles normalized to the mean length of the interovulatory interval (n = 9 mares). Diameters of the follicles are for the largest follicles each day without regard to tracking day-to-day identity. Note the dissociation of the follicles into a single growing (dominant) follicle and five regressing (subordinate) follicles. Mean day of the beginning of deviation is shown by a vertical broken line. Adapted from Ginther et al.58
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cycle in association with major follicular waves. The divisions are: (i) emergence and common growth phase, during which time many follicles of the wave emerge and grow together as a cohort or group; (ii) deviation and dominance phase, during which time one follicle of the cohort continues to grow and becomes dominant and other follicles cease growth and regress; and (iii) pre-ovulatory phase, during which time the dominant follicle increases to maximal size and undergoes morphological and physiological changes in preparation for ovulation.
35
FOLLICLES Dominant follicle
30 Diameter (mm)
c213
25 Subordinate follicle
20 15 10 5 -6
(a)
Follicle emergence and common growth phase The emergence of follicles of a wave refers to the earliest ultrasonic detection compatible with tracking of individual follicles (Table 213.1). In studies that have used follicle tracking, the time of follicle emergence was based on retrospective examination of the largest follicle when it first reached 15 mm in diameter.35, 40, 41 When follicle ablation was used to remove the interfering follicles from the previous wave and induce a new wave, follicle emergence for the new wave was consistently and reliably studied by detection and tracking of follicles beginning at 6 mm.12, 42 Although tracking of individual follicles may not be necessary from a practical perspective until they reach greater than 15 mm, hypothesis testing may require tracking of individual follicles beginning at approximately 6 mm as a reference for follicle or wave emergence. Follicles that emerge within 2 days of one another are included in the cohort of follicles of the wave in which the mean number of antral follicles ranges from 7 to 11 at the time of first detection or wave emergence. In some instances, follicles emerge more than 2 days apart from one another and reach a small maximum diameter of approximately 12 mm compared to the maximum diameter of the largest and dominant follicles of minor and major waves, respectively, before regressing. This type of follicular growth and regression underlying minor and major waves occurs intermittently throughout the estrous cycle.12 On average, the largest follicle of a wave is first detected about 1 day before the next largest follicle, with a size advantage of about 3 mm or about 1 day’s growth.12, 42 Correspondingly, the growth rates of the cohort of follicles of a wave are similar at approximately 3 mm/day from emergence until the two largest follicles begin to dissociate in diameter in association with the follicle deviation phenomenon (Table 213.1). Hence, the time from wave emergence to beginning of follicle deviation (6 to 7 days) in association with major waves is referred to as the common growth phase.43 The mean growth profile (centralized to the beginning of deviation) of the two
35
-5 -4 -3 -2 -1 0 1 2 3 Days from beginning of deviation
m Do
25 20 15 10
4
DEVIATION HYPOTHESIS
30 Diameter (mm)
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as ph
e
e
as ph h t w ro ay -g /d on m m m m 8 2. Co
Subordinate follicle
Deviation
5 Emergence
(b)
0 1 2 3 4 5 6 7 8 9 10 Days from future dominant follicle 6 mm
Figure 213.4 Mean (± SEM) profile of dominant and subordinate follicles centralized to the beginning of deviation (a) and diagrammatic illustration of common growth and dominance phases encompassing deviation (b). For the deviation hypothesis, the larger follicle (future dominant) emerges earlier and, therefore, has a size advantage reaching the critical stage (black bar) first since the two follicles grow at a similar rate after emergence. At the critical stage, the smaller follicle (future subordinate) is inhibited through a deviation mechanism that involves the dominant follicle so that a similar diameter is not reached (white bar). Adapted from Ginther et al.9
largest follicles from wave emergence until the largest follicle was 35 mm is shown in Figure 213.4a. Although the cohort of follicles of the wave may grow similarly during the common growth phase, the mean frequency with which the largest follicle will become the future dominant follicle at the end of the common growth phase is approximately 61% compared to approximately 25% for the second-largest follicle.12 The frequency of other smaller follicles of the wave becoming the future dominant follicle at the end of the common growth phase diminishes accordingly. As reported for individual mares,12 the second-largest follicle replaces the largest follicle as the future dominant follicle 33% of the time towards
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Folliculogenesis the end of the common growth phase. Nonetheless, it has been proposed that follicle-size hierarchy at the time of emergence of a major wave is a component of the selection process that allows the largest follicle to establish its destiny as the future dominant follicle during the common growth phase before the second-largest follicle reaches a similar diameter, culminating in follicle deviation at the end of the common growth phase.43 An illustration of the deviation hypothesis is shown in Figure 213.4b. Follicle deviation and dominance phase Follicle deviation refers to the dissociation in diameter between follicles at the end of the common growth phase as one or occasionally two follicles continue growth and other follicles of the wave cease growth and eventually regress.8 Follicle deviation is a key component of the follicle selection process and is thought to be a mechanism to prevent multiple follicles from becoming co-dominant in monovular species, especially since many follicles of a wave are capable of becoming dominant.43 Based on ultrasonic imaging, the beginning of follicle deviation can be expected to occur when the diameter of the largest follicles reaches 22 to 25 mm (mean 22.5 mm),12, 15, 16, 39, 42 and the second-largest follicle is an average of 19 mm. Follicle deviation is a conspicuous and often abrupt event recognized by a sudden decrease in the growth rate of the secondlargest follicle. Despite the reduced growth rate after follicle deviation, the second-largest follicle retains the capacity to become dominant12 or co-dominant13 for about two days after the beginning of deviation before it is committed to regression. The capacity for subordinate follicles to become dominant after beginning of deviation is supported by experimental observation. In 67% of mares, the second-largest follicle resumed growth and became dominant when the largest follicle was ablated 1 or 2 days after the beginning of deviation.12 In addition, intrafollicular treatment with insulin-like growth factor 1 (IGF1) to all but the largest follicle within 0 to 2 days after the expected time of deviation resulted in multiple dominant follicles and ovulations.13 Reportedly,44 if the difference in diameter between the two largest follicles around the time of deviation is ≥ 6 mm, subordinate follicles are more likely to become regressive than dominant after the ablation of the largest follicle. In this regard, follicle diameter and diameter differences around the beginning of follicle deviation have practical importance for the timing of exogenous gonadotropin treatments for enhancing multiple dominant follicles and ovulations for embryo transfer as discussed in Chapter 168 on FSH. In addition to follicle diameters and diameter differences between the two largest follicles of a major
2015
wave, other B-mode and color-Doppler ultrasonographic changes can be used to indicate the beginning of follicle deviation. Thickness and echogenicity of the granulosa cell layer, and prominence and extent of circumferential involvement of the anechoic layer beneath the granulosa of the follicle were all greater for the future dominant follicle compared to future subordinate follicles, as shown in Figure 213.5.45 In addition, follicle wall area with color signals for blood flow, and blood flow velocities as determined by color-Doppler ultrasonography, were greater for the future dominant follicle than subordinate follicles before the beginning of deviation.46 The dominance phase is commonly referred to as the period from the beginning of deviation until beginning of the pre-ovulatory phase (approximately 8 days) in association with major waves, especially the primary wave.8 Dominance can be indicated morphologically by ultrasonic imaging of follicle diameter and diameter differences between the two largest follicles, and physiologically by assay techniques to determine the capacity for the differential development of increased amounts of hormone receptors, such as luteinizing hormone (LH) receptors, in granulosa cells, and/or differential increase in amounts of reproductive hormones and growth factors (estradiol, inhibin, and free IGF1) in follicular fluid.8 A greater number of LH receptors makes the dominant follicle responsive to the stimulatory effects of LH; higher IGF1 promotes mitogenesis and growth of the follicle; and increased estradiol and inhibin suppress systemic follicle-stimulating hormone (FSH) and halt the development of subordinate follicles as discussed in more detail in Chapter 168 on FSH. The dominant follicle, as indicated by a diameter ≥ 28 mm, continues growth after deviation at about 3 mm/day and either regresses, becomes hemorrhagic, or ovulates depending, in part, on whether it is associated with a secondary or primary major wave and time of season. Pre-ovulatory phase The pre-ovulatory phase refers to the developmental capacity of the dominant follicle to respond to an endogenous or exogenous ovulatory stimulus. Although there are no direct studies to indicate the minimal diameter at which the dominant follicle enters the pre-ovulatory phase, it is generally considered to be when the follicle reaches ≥ 30 mm, since follicles < 30 mm rarely ovulate spontaneously (for review see Ginther24 ) or in response to exogenous stimulation.47 There are, however, distinctive follicle characteristics that have been documented using Bmode48 and color-Doppler49 ultrasonography to evaluate the ovulatory potential and estimate the time to ovulation, as reviewed and discussed in detail in
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Figure 213.5 Ultrasonograms illustrating the echotextural appearance of the follicle wall in a future dominant follicle (a–c) and a subordinate follicle (d–f) over time as the time to deviation approaches. The anechoic layer beneath the granulosa cell layer (arrows) is more pronounced over time for the future dominant follicle than the subordinate follicle. The graduation marks represent 10-mm increments. Adapted from Gastal et al.45
Chapter 214 on ovulation. Ultrasonographic characteristics have also been documented for distinguishing between ovulatory and anovulatory follicles as reviewed (see Ginther19 ) and discussed in detail in Chapter 214–216 on ovulation. In brief, the prominence of an anechoic layer beneath the granulosa cell layer, increased thickness and echogenicity of the follicle wall as detected by B-mode ultrasonography,48, 50 and increased color signals for blood flow in the follicle wall as detected by color-Doppler ultrasonography49 are characteristics of the pre-ovulatory follicle. Two common irregularities associated with folliculogenesis are double dominant, or co-dominant, follicles and hemorrhagic anovulatory follicles. These aberrations of folliculogenesis are discussed in detail in Chapters 214 and 222 on ovulation and ovarian abnormalities. In brief, double dominant follicles occur as a result of disruption or failure of the deviation mechanism between the two largest follicles at the end of the common growth phase such that the second-largest follicle also continues to grow and
becomes dominant. The incidence of double dominant follicles in some recent reports ranged from 20 to 38%.15, 16, 39, 51 Spontaneous double dominant follicles frequently result in double ovulations. In one study,14 the incidence of double dominant follicles resulting in double ovulations was 43%. Follicular waves with double dominant follicles not resulting in double ovulations undergo a second deviation in diameter (between two dominant follicles), on average, 2 days after the first deviation.16 As observed, the first deviation in double dominant waves occurs between the largest and third largest follicles. The incidence of spontaneous triple dominant follicles is very low (0.6 to 4.0%15, 24 ) and the incidence of more than three spontaneous dominant follicles is not known. Hemorrhagic anovulatory follicles result as a consequence of blood entering the antrum of an impending ovulatory follicle, which subsequently fails to ovulate.52 Hemorrhagic anovulatory follicles are more commonly seen in old mares (36% in mares ≥ 20
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Days before first ovulation Figure 213.6 Daily diameter profiles of the six largest follicles without regard to tracking day-to-day identity during transition from the anovulatory period to the ovulatory period in three mares. Note the consistent pattern of minor waves that precede the major waves, and that in some instances a major anovulatory wave precedes the first ovulatory wave (mares B and C). anov = anovulatory; ov = ovulation. Adapted from Donadeu and Ginther.55
years53 ) and have about a 50% incidence of repeatability in individual mares.52 The incidence of hemorrhagic anovulatory follicles is approximately 5% during the early ovulatory season and 20% during the late ovulatory season.54 The only differentiating feature capable of serving as a potential indicator of impending anovulation seems to be the increase in percentage of color signals for blood flow (i.e., enhanced vascularization) in the follicle wall near the apical area of expected ovulation as determined by color-Doppler ultrasonography.52 Early pregnancy and spring transitional season In general, follicular development during early pregnancy [< 30 days after ovulation before equine chorionic gonadotropin (eCG) production] and late spring transition occurs similarly involving development of minor, major, or a combination of both wave types in succession in which the dominant follicle of major waves (≥ 28 mm) is anovulatory. Although the mechanism may be different between the two reproductive states, reduced circulating concentrations of LH appear to be common to both states. As a consequence of reduced LH, the largest follicles of minor waves fail to reach ≥ 28 mm (i.e., dominance) and dominant follicles of major waves fail to ovulate. Neverthless, during early pregnancy, endometrial cups form and eCG (with LH-like activity) production begins after about 30 days of gestation (for review see Ginther24 ). As a result, dominant follicles of major waves during early pregnancy are either ovulatory or anovulatory forming supplemental corpora lutea as discussed in Chapter 217 on pregnancy. The timing of ovulation of dominant follicles of major waves during spring transition is discussed in Chapter 178 on vernal transition. Briefly, however, during the middle of the anovulatory season
(i.e., two midwinter months), follicle activity is minimal and only small follicles (< 15 mm) are present. As day length gradually increases during the spring transitional period, follicles grow to increasingly larger diameters culminating in the development of dominant, ovulatory follicles.55 In some instances, however, the latter part of the transitional phase consists of periods in which the largest follicle of each sequential wave approaches or is similar to the diameter of a dominant follicle (≥ 28 mm) but does not ovulate.36, 56 Daily diameter profiles of the six largest follicles in three mares during the spring transitional period are illustrated in Figure 213.6. Most striking and of importance to equine practitioners and clinicians and breeding-farm managers is the development of major anovulatory waves during the transitional period. Dominant-sized transitional anovulatory follicles have been found to be dysfunctional as a result of a deficient follicular hormonal milieu.56, 57 A marked morphological difference that may be exploited to distinguish anovulatory from ovulatory follicles during the spring transitional period is reduced blood flow (i.e., decreased colorDoppler signals) associated with anovulaory follicles as determined by color-Doppler ultrasonography.57 Recall, this is in contrast to the enhanced blood flow (i.e., increased color-Doppler signals) associated with anovulatory follicles during the ovulatory season. As illustrated in Figure 213.7, follicle wall area with color signals for blood flow was less for dominant-sized anovulatory follicles than for ovulatory follicles during the transitional period, starting when the follicles attained approximately 25 mm in diameter.
Summary Ultrasonic imaging provides a non-invasive and repeatable approach for visualizing the ovaries in
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DOMINANT FOLLICLES
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Figure 213.7 Mean (± SEM) diameter profiles of dominant ovulatory and anovulatory follicles (a) and follicle wall area with blood flow associated with each follicle type (b). Note diameters between two dominant-follicle types were statistically different after the follicles reached about 30 mm (asterisks), whereas follicle wall area with blood flow between the two dominant-follicle types was statistically different when the follicles were about 25 mm (asterisks); blood flow was greater for the ovulatory follicle than the anovulatory follicle. Adapted from Acosta et al.57
real time for evaluation of follicular development in mares. Follicle characteristics can be quantitated and sequentially monitored using B-mode and colorDoppler ultrasonography for studying the dynamic interactions within the population of follicles; these data can be used for basic and applied purposes as well as for practical purposes to determine the opportune time to start gonadotropin treatment relative to follicle dominance and differentiating between ovulatory and anovulatory follicles during the estrous cycle and fall and spring transitional periods.
Acknowledgements The authors are grateful to Dr. O.J. Ginther and his laboratory for support in preparation of this chapter. In addition, the authors are thankful to Drs E.L. Gastal, M.O. Gastal, R.R. Araujo, and K.R. Girish for their help with preparation of figures.
1. Palmer E, Driancourt MA. Use of ultrasound echography in equine gynecology. Theriogenology 1980;13:203–16. 2. Beg MA, Ginther OJ. Follicle selection in cattle and horses: role of intrafollicular factors. Reproduction 2006;132:365–77. 3. Driancourt MA, Paris A, Roux C, Mariana JC, Palmer E. Ovarian follicular populations in pony and saddle-type mares. Reprod Nutr Dev 1982;22:1035–47. 4. Erickson BH. Development and senescence of postnatal bovine ovary. J Anim Sci 1966;23:800–5. 5. McGee EA, Hsueh AJW. Initial and cyclic recruitment of ovarian follicles. Endocrine Rev 2000;21:200–14. 6. Driancourt MA. Regulation of ovarian follicular dynamics in farm animals. Implication for manipulation of reproduction. Theriogenology 2001;55:1211–39. 7. Evans ACO. Characteristics of ovarian follicle development in domestic animals. Reprod Domest Anim 2003;38:240–6. 8. Ginther OJ, Beg MA, Donadeu FX, Bergfelt DR. Mechanism of follicle deviation in monovular farm species. Anim Reprod Sci 2003;78:239–57. 9. Ginther OJ, Beg MA, Gastal MO, Gastal EL. Follicle dynamics and selection in mares. Anim Reprod 2004;1:45–63. 10. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Seasonal influence on equine follicle dynamics. Anim Reprod 2004;1:32–45. 11. Donadeu FX, Pedersen HG. Follicle development in mares. Reprod Domest Anim 2008;43 (Suppl. 2): 224–31. 12. Gastal EL, Gastal MO, Beg MA, Ginther OJ. Interrelationships among follicles during the common-growth phase of a follicular wave and capacity of individual follicles for dominance in mares. Reproduction 2004;128:417–22. 13. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Intrafollicular effect of IGF1 on development of follicle dominance in mares. Anim Reprod Sci 2008;105:417–23. 14. Ginther OJ, Gastal EL, Rodrigues BL, Gastal MO, Beg MA. Follicle diameters and hormone concentrations in development of single versus double ovulations in mares. Theriogenology 2008;69:583–90. 15. Ginther OJ, Jacob JC, Gastal MO, Gastal EL, Beg MA. Development of one versus multiple ovulatory follicles and associated systemic hormone concentrations in mares. Reprod Domest Anim 2009;44:441–9. 16. Jacob JC, Gastal EL, Gastal MO, Carvalho GR, Beg MA, Ginther OJ. Follicle deviation in ovulatory follicle waves with one or two dominant follicles in mares. Reprod Domest Anim 2009;44:248–54. 17. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Fundamentals, Book 1. Cross Plains, WI: Equiservices Publishing, 1995. 18. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Horses, Book 2. Cross Plains, WI: Equiservices Publishing, 1995. 19. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Color-Doppler Ultrasonography, Book 4. Cross Plains, WI: Equiservices Publishing. 2007. 20. McKinnon AO. Reproductive ultrasonography – the mare. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Philadelphia: Lea & Febiger, 1998; pp. 125–251. 21. Reef VB. Equine Diagnostic Ultrasound. St Louis: Saunders, 1998. 22. Wolfgang K. Ultrasonography in the mare. In: Veterinary Reproductive Ultrasonography. Hannover: Schlutersche Verlagsgesellschaft mbH, 2004; pp. 1–80.
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Folliculogenesis 23. Ginther OJ, Utt MD. Doppler ultrasound in equine reproduction: principles, techniques, and potential. J Equine Vet Sci 2004;24:516–26. 24. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices Publishing, 1992. 25. Deanesly R. Germ cell development and meiotic prophase in fetal horse ovary. J Reprod Fertil Suppl 1975;23:547–52. 26. Deanesly R. Germ cell proliferations in the fetal ovary. Cell Tissue Res 1977;185:361–71. 27. Wesson JA, Ginther OJ. Influence of season and age on reproductive activity in pony mares on the basis of a slaughterhouse survey. J Anim Sci 1981;52:119–29. 28. Nogueira GP, Ginther OJ. Dynamics of follicle populations and gonadotropin concentrations in fillies age two to ten months. Equine Vet J 2000;32:482–8. 29. Ginther OJ. Major and minor follicular waves during the equine estrous cycle. J Equine Vet Sci 1993;13:18–25. 30. Evans AC, Adams GP, Rawlings NC. Follicular and hormonal development in prepubertal heifers from 2 to 36 weeks of age. J Reprod Fertil 1994;102:463–70. 31. Schrick FN, Surface RA, Pritchard JY, Daily RA, Townsend EC, Inskeep EK. Ovarian structures during the estrous cycle and early pregnancy in ewes. Biol Reprod 1993;49:1133–40. 32. Fernandez-Moro D, Veiga-Lopez G, Ariznavarreta C, Tresguerres JAF, Encinas T, Gonzalez-Bulnes A. Preovulatory follicle development in goats following oestrous synchronization with progestagens and prostaglandins. Reprod Domest Anim 2008;43:9–14. 33. Palmer E. New results on follicular growth and ovulation in mare. In: Roche JF, O’Callaghan D (eds) Follicle Growth and Ovulation in Farm Animals. Dordrecht: Martinus Nijhoff Publishers, 1987; pp. 237–55. 34. Pierson RA, Ginther OJ. Follicular population dynamics during the estrous cycle of the mare. Anim Reprod Sci 1987;14:219–31. 35. Sirois J, Ball BA, Fortune JE. Patterns of growth and regression of ovarian follicles during the oestrous cycle and after hemiovariectomy in mares. Equine Vet J (Suppl.) 1989;8:43–8. 36. Ginther OJ. Folliculogenesis during the transitional period and early ovulatory season in mares. J Reprod Fertil 1990;90:311–20. 37. Gastal EL, Gastal MO, Nogueira GP, Bergfelt DR, Ginther OJ. Temporal interrelationships among luteolysis, FSH and LH concentrations and follicle deviation in mares. Theriogenology 2000;53:925–40. 38. Ginther OJ, Bergfelt DR. Associations between FSH concentrations and major and minor follicular waves in pregnant mares. Theriogenology 1992;38:807–21. 39. Ginther OJ, Gastal EL, Gastal MO, Bergfelt DR, Baerwald AR, Pierson RA. Comparative study of the dynamics of follicular waves in mares and women. Biol Reprod 2004;71:1195–201. 40. Bergfelt DR, Ginther OJ. Relationships between circulating concentrations of FSH and follicular waves during early pregnancy in mares. J Equine Vet Sci 1992;12:274–79. 41. Bergfelt DR, Ginther OJ. Relationships between FSH surges and follicular waves during the estrous cycle in mares. Theriogenology 1993;39:781–96. 42. Gastal EL, Gastal MO, Bergfelt DR, Ginther OJ. Role of diameter differences among follicles in selection of a
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future dominant follicle in mares. Biol Reprod 1997;57: 1320–7. Ginther OJ, Beg MA, Bergfelt DR, Donadeu FX, Kot K. Follicle selection in monovular species. Biol Reprod 2001;65:638–47. Gastal EL, Gastal MO, Ginther OJ. Experimental assumption of dominance by a smaller follicle and associated hormonal changes in mares. Biol Reprod 1999;61:724–30. Gastal EL, Donadeu FX, Gastal MO, Ginther OJ. Echotextural changes in the follicular wall during follicle deviation in mares. Theriogenology 1999;52:803–14. Acosta TJ, Gastal EL, Gastal MO, Beg MA, Ginther OJ. Differential blood flow changes between the future dominant and subordinate follicles precede diameter changes during follicle selection in mares. Biol Reprod 2004;71:501–7. Bergfelt DR, Meira C, Fleury PDC, Dell’Aqua JA, Adams GP. Ovulation synchronization following commercial application of ultrasound-guided follicle ablation during the estrous cycle in mares. Theriogenology 2007;68:1183–91. Gastal EL, Gastal MO, Ginther OJ. The suitability of echotextural characteristics of the follicular wall for identifying the optimal breeding day in mares. Theriogenology 1998;50:1025–38. Gastal EL, Gastal MO, Ginther OJ. Relationships of changes in B-mode echotexture and colour-Doppler signals in the wall of the preovulatory follicle to changes in systemic estradiol concentrations and effect of human chorionic gonadotropin in mares. Reproduction 2006;131:699– 709. Pierson RA, Ginther OJ. Ultrasonic evaluation of the preovulatory follicle in the mare. Theriogenology 1985;24: 359–68. Ginther OJ, Jacob JC, Gastal MO, Gastal EL, Beg MA. Follicle and systemic hormone interrelationships during spontaneous and ablation-induced ovulatory waves in mares. Anim Reprod Sci 2008;106:181–7. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Conversion of a viable preovulatory follicle into hemorrhagic anovulatory follicle in mares. Anim Reprod 2006;3:29–40. Carnevale EM. Folliculogenesis and ovulation. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasound. Baltimore, MD: Williams & Wilkins, 1998; pp 201–11. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Incidence, endocrinology, vascularity, and morphology of hemorrhagic anovulatory follicles in mares. J Equine Vet Sci 2007;27:130–9. Donadeu FX, Ginther OJ. Follicular waves and circulating concentrations of gonadotropins, inhibin and oestradiol during anovulatory season in mares. Reproduction 2002;124:875–85. Watson ED, Heald M, Tsigos A, Leask R, Steele M, Groome NP, Riley SC. Plasma FSH, inhibin A and inhibin isoforms containing pro and -αC during winter anoestrus, spring transition and the breeding season in mares. Reproduction 2002;123:535–42. Acosta TJ, Beg MA, Ginther OJ. Aberrant blood flow area and plasma gonadotropin concentrations during the development of dominant-sized transitional anovulatory follicles in mares. Biol Reprod 2004;71:637–42. Ginther OJ, Utt MD, Beg MA. Follicle deviation and diurnal variation in circulating hormone concentrations in mares. Anim Reprod Sci 2007;100:197–203.
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Chapter 214
Ovulation: Part 1. Follicle Development and Endocrinology During the ∗ Periovulatory Period Eduardo L. Gastal
Introduction In many species, the most extensively studied aspects of folliculogenesis are the morphological, hormonal, and biochemical changes during the preovulatory stage in preparation for ovulation. However, regulatory mechanisms of ovulation in mares are not well known, especially the role of the long secretory pattern of luteinizing hormone (LH) during the periovulatory period. The biochemical and biophysical aspects of ovulation in the mare have also been neglected research areas. Changes in concentrations of circulating hormones during the estrous cycle began to be characterized in the early 1970s; however, the classical descriptions are still being progressively refined. In mares, only a few studies have considered the relationships between follicle diameter and hormonal changes during the periovulatory period. Attention to this aspect of reproductive physiology is needed due to the pivotal role of ovulation in cyclicity and in reproductive management. Our temporal and mechanistic studies performed recently (2005 to 2009) have elucidated intriguing relationships among the preovulatory follicle and circulating hormones. The focus of this chapter will be the aspects related to the development of the ovulatory follicle and the associated systemic and intrafollicular hormones. Biochemical, molecular, and cellular mechanisms involved in ovulation1 and synchronization and ∗
induction of ovulation (see Chapters 196 and 197) will not be addressed herein. This review emphasizes the contribution of the recent findings and is directed to equine theriogenologists and scientists who are involved in monitoring, managing, and manipulating the mare’s reproductive system during the periovulatory period. The topics addressed in this chapter are: development of one versus two dominant follicles with a single ovulation; development of one versus multiple ovulatory follicles; conversion of double dominant follicles to double ovulations; the role of hormones in development of double ovulations; the interrelationships of periovulatory reproductive hormones; repeatability of preovulatory follicle diameter and hormones; and factors that affect preovulatory follicle diameter and hormones, such as breeds and types, season, body condition, and aging. Ultrasonographic characteristics of the preovulatory follicle preceding normal and atypical ovulations is the topic of the following chapter (see Chapter 215).
Preovulatory follicle development and endocrinology Ovarian follicle development in the mare is characterized by waves of several follicles that emerge and initially grow in synchrony. Various numbers and types of follicular waves develop during an equine
Portions of this chapter have been adapted from Gastal E.L. 2009. Recent advances and new concepts on follicle and endocrine dynamics during the equine periovulatory period. Animal Reproduction 6:144–58.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Ovulation: Part 1. Follicle Development and Endocrinology interovulatory interval (IOI).2 In a major wave, the largest follicle attains the diameter of a dominant follicle (≥ 28–30 mm), whereas in minor waves the largest follicle does not become dominant. The ovulatory wave emerges midway during an IOI of 21– 24 days. After emergence at 6 mm,3 the follicles of a wave develop in a common-growth phase for several days.4 At the end of the common-growth phase, a distinctive change in growth rates begins. This process is called deviation, and in mares it begins when the diameters of the two largest follicles on average are 22.5 mm and 19.0 mm.3, 5, 6 The deviation in growth rates between the future dominant and subordinate follicles is a key event during the selection of the ovulatory follicle. After deviation, the developing dominant follicle maintains a constant growth rate until 1–2 days before ovulation7, 8 and the remaining follicles (subordinate follicles) grow at a reduced rate and regress. More general aspects as well as specific details about folliculogenesis in mares and associated reproductive hormones may be found in Chapter 213 of this book and in recent reviews.6, 9
Development of one versus two dominant follicles with a single ovulation In the monovular species (horses, cattle, women) usually only one dominant follicle develops as a result of the deviation mechanism, but double dominant follicles also occur, based on two follicles reaching ≥ 28–30 mm in mares.3, 6, 10, 11 The incidence of two dominant follicles in an ovulatory wave has varied from 20% in Breton horses12 to about 30% in large ponies.13, 14 This incidence seems to be lower in Miniature mares, since no double dominant follicles (≥ 28 mm) were observed in 36 preovulatory periods.15 Studies have not found a hormonal basis for the development of two dominant follicles in mares. Waves with one versus two dominant follicles (no double ovulations) had similar plasma concentrations of LH, estradiol, and immunoreactive (ir)-inhibin; however, concentrations of folliclestimulating hormone (FSH) were lower in mares with two dominant follicles.14 Therefore, the factors that lead to the development of two dominant follicles are still unknown in mares.
Development of one versus multiple ovulatory follicles In a recent study using ablation-induced waves, comparisons were made between waves with one (33 waves, 72%) and multiple (13 waves, 28%) ovulatory follicles in large pony mares.13 Multiple ovulatory follicles had later emergence and were preceded by
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Figure 214.1 Means (± SEM) for systemic concentrations of follicle-stimulating hormone (FSH), luteinizing hormone (LH), and estradiol in induced ovulatory waves with one versus multiple ovulations. Data were normalized to the beginning of deviation. Numbers of waves for the one and multiple ovulations are 33 and 13 for FSH and LH, and 12 and 9 for estradiol, respectively. Data adapted from Ginther et al.13
more follicles ≥ 20 mm at the beginning of deviation, had a higher LH concentration preceding deviation, lower concentrations of FSH on the day of deviation and thereafter, and higher estradiol concentrations by 2 days after deviation (Figure 214.1). During the periovulatory period, systemic hormone concentrations for waves with multiple ovulations involved higher estradiol before ovulation, lower FSH before and after ovulation, and both higher progesterone and lower LH beginning on the day after ovulation (Figure 214.2). These findings are compatible with the concept that a higher LH concentration in certain
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4
Figure 214.2 Means (± SEM) for systemic concentrations of follicle-stimulating hormone (FSH), luteinizing hormone (LH), estradiol, and progesterone in induced ovulatory waves with one versus multiple ovulations. Data were normalized to the first ovulation. Numbers of waves for the one and multiple ovulations are 33 and 13 for FSH and LH, 12 and 9 for estradiol, and 20 and 12 for progesterone, respectively. Data adapted from Ginther et al.13
mares preceding deviation favors the development of multiple estrogen-competent follicles ≥ 20 mm at deviation, leading to the development of multiple ovulations. Conversion of double dominant follicles to double ovulations In addition to breed, the variability in doubleovulation rate reflects the effects of reproductive status, age, and repeatability in individuals. In a review16 of many reports on equine double ovulations, it was concluded that the incidence varied extensively but was lower in ponies (2%) than in horses
(7–25%). This incidence seems to be very low in Miniature mares, since no double ovulations were observed in any of 36 preovulatory periods.15 The percentage of double dominant follicles that spontaneously become double ovulations apparently has not been reported previously for any monovular species, except for recent reports in mares. In six cases of double dominant follicles in Breton horses, ovulation occurred from either one follicle (five mares) or both follicles (one mare).12 The incidence of double dominant follicles becoming double ovulations has varied from 9% to 43% in large pony mares.14, 17, 18 A combination of an ovulation and a hemorrhagic anovulatory follicle (HAF) from the largest remaining follicle can also occur in some mares with double dominant follicles. Recent studies have found that the incidence of double or multiple dominant follicles was similar between ablation-induced and spontaneous ovulatory waves; however, double or multiple dominant follicles ovulated more frequently in induced waves.13, 14, 18 In the induced waves, both the day of the FSH surge and day of deviation were more synchronized, FSH and LH concentrations were greater before and after deviation, and estradiol concentrations were greater after deviation.18 FSH, LH, and estradiol concentrations did not differ between induced and spontaneous waves before ovulation. It can be speculated that the uniform prominent wavestimulating FSH surge or the post-ablation LH increase during the common-growth phase in induced waves enhanced the ovulatory capabilities of the future dominant follicles. This concept is compatible with the greater estradiol production of the double follicles before dominance (≥ 28 mm) was established, leading to increased ovulatory capacity of both dominant follicles. However, the hormonal basis for an apparent greater rate of conversion of double dominant follicles to double ovulations in induced waves than in spontaneous waves is still unknown. Role of hormones in development of double ovulations The relationship between periovulatory circulating hormone concentrations and double preovulatory follicles and the subsequent double ovulations has not been widely studied in mares. In addition, it is not known whether in any of the monovular species such a relationship would reflect a role for the hormones in development of preovulatory follicles for double ovulations or an effect of the follicles on the hormones. Previous studies19, 20 in mares did not report significant differences in gonadotropin concentrations between single and double ovulators; however, due to either the experimental design utilized
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Single ovulator
30 25 -2
-1
0 1 Days from ovulation
2
3
Figure 214.3 Means (± SEM) for diameters of ovulatory follicles for single ovulating mares and double ovulating mares. Data are organized for the 2.5 days after the first follicle (F1) reached ≥ 30 mm and for the 2.5 days before ovulation to correspond to a mean 5-day interval between a ≥ 30-mm follicle and ovulation. The ovulating follicle for single ovulators first became significantly larger than the largest follicle in double ovulators on Day – 2.5. Data adapted from Ginther et al.17
or the data presented, the results seemed to be inconclusive. Recently, the relationships between follicle diameters and hormone concentrations have been compared between single ovulator and double ovulator mares17 (Figures 214.3 and 214.4). In double ovulators, compared to single ovulators, the largest follicle was smaller, FSH was lower, and estradiol was higher on most days, but LH and ir-inhibin were not different. A relatively greater depression of FSH after double ovulations than after one ovulation was seen and may have been caused by inhibin from multiple follicles. Lower concentrations of LH were detected after double ovulations and likely reflected, at least in part, the greater concentrations of progesterone on the corresponding days. Results from this study indicated that smaller preovulatory follicles in double ovulators are a response to lower FSH, owing to higher estradiol from two preovulatory follicles, and that the preovulatory differences in hormone concentrations between single and double ovulators are an effect rather than a cause of the double ovulations. The length of the IOI does not differ between single ovulators and double ovulators.17, 19 The diameter of the preovulatory follicle is larger for single than for double ovulations,2 and the growth rate of the preovulatory follicle preceding a single or double ovulation diminishes on the day before ovulation in mares.7, 8, 17 The mechanism involved in the reduced growth of the follicle before ovulation apparently begins when the ovulatory LH surge reaches a critical level; this has been suggested by an immediate reduction in follicle growth and estradiol production when an ovulatory dose of human chorionic gonadotropin (hCG) was given.7, 8 Although the small diameter of double preovulatory follicles in one ovary can be attributed, speculatively, to a crowding effect, a reduction in diameter also occurs with bilateral ovulations, indicating the involvement of other factors.
Concentration (pg/ml)
Double ovulator
35
0 1 2 Days after 30-mm F1
2023
10 Estradiol
40
8
Double ovulators
6 Single ovulators
4
12 FSH Concentration (ng/ml)
45 Diameter (mm)
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10
8
6
12 Concentration (ng/ml)
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LH
10 8 6 4 2 0 1 2 Days after 30-mm F1
-2
-1
0 1 Days from ovulation
2
3
Figure 214.4 Means (± SEM) for concentrations of estradiol, follicle-stimulating hormone (FSH), and luteinizing hormone (LH) in single and double ovulators. Data were analyzed for 2.5 days after the first follicle (F1) reached ≥ 30 mm and separately for the 2.5 days before ovulation and 3 days after ovulation to correspond to a mean 5-day interval between a ≥ 30mm follicle and ovulation. Owing to a negative effect of estradiol, FSH was lower throughout the preovulatory period, resulting in smaller follicles in the double ovulators. Adapted from Ginther et al.17
Interrelationships of periovulatory reproductive hormones Recent studies have contributed considerably to the understanding of the hormonal changes during the periovulatory period in mares.13, 17, 18, 21, 22 Briefly, as depicted in Figure 214.5, the concentrations of LH during the ovulatory LH surge initially increase slowly, followed by a more rapid increase beginning on Day – 2 (ovulation = Day 0). The surge reaches a peak on Day 1. Concentrations of FSH begin increasing on Day – 2, forming an early diestrous surge that precedes a surge that stimulates emergence of the next ovulatory follicular wave. The preovulatory
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1 16 LH, FSH, P4 (ng/ml), E2 (pg/ml x 2)
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5 4
14 12 E2
10 8
FSH
6 LH
4 2
P4
0 -4
-3
-2
-1 0 1 2 Days from ovulation
3
4
Figure 214.5 Means (± SEM) for periovulatory systemic concentrations of estradiol (E2), luteinizing hormone (LH), folliclestimulating hormone (FSH), and progesterone (P4). Estradiol is shown at twice the actual concentrations. A circle around a mean indicates the day of a transition from a significant increase to a significant decrease (LH, E2) or from a decrease to an increase (FSH). A square indicates the day of change in the rate of a significant increase (LH) or rate of decrease (E2). A triangle indicates the beginning of a significant increase (P4). The circled numbers refer to the following: (1) days of continuous growth of the preovulatory follicle, decreasing FSH concentrations, and reduced rate of increase in LH, owing to a negative effect of the increasing E2; (2) day of acquisition of a negative effect of LH on E2 and apparently on cessation of follicle growth at the peak of E2; (3) days of rapidly decreasing E2 from the increasing negative effect of the rapidly increasing LH, resulting in turn in a lessening of the negative effect of E2 on LH and FSH and therefore a greater rate of production of LH and an increase in FSH; (4) day of transition between increasing and decreasing LH when P4 reaches a critical concentration; and (5) days of reduction in LH from increasing P4, with a lessening of the negative effect of LH on E2 and therefore a slower decrease in E2. Data adapted from Jacob et al.22
estradiol surge reaches a peak on Day – 2. Progesterone increases slightly but significantly by Day 0. The interrelationships among the changing hormone concentrations during the periovulatory period have been considered in two studies discussed below. In a study in which hormone concentrations were evaluated every 24 hours from a large number of periovulatory periods (n = 66), mean concentrations of FSH began to increase and estradiol began to decrease in synchrony on Day – 2 (see Figure 214.5).22 These two changes are likely related, given that exogenous estradiol has a negative effect on FSH in mares.23 The FSH increase between Days – 2 and – 1 was followed by a transient suspension in the
increase or a plateau in FSH between Days – 1 and 0, consistent with a discharge of hormone-laden follicular fluid into the peritoneal cavity during ovulation that occurred sometime between Day – 1 and Day 0. In this regard, an increase in ir-inhibin concentrations on Day 024–26 has been related to absorption of inhibins into the circulation after the discharge of follicular fluid into the abdomen.26 A similar indication that estradiol of the discharged follicular fluid alters the profile of plasma estradiol has not been apparent. Thus, the absorption of inhibin from the follicular fluid in the peritoneal cavity accounts at least partially for the transient suspension in the FSH increase between Days – 1 and 0. An increase, apparently a rebound, in FSH occurred between Days 0 and 1. This FSH response likely reflects the release of FSH from the suppressing effects of inhibin during the absorption of inhibin from the peritoneal cavity. In this regard, a rebound in FSH began 24 hours after administration of a proteinaceous fraction of follicular fluid in ovariectomized mares27 and ovarian-intact mares.28 The rapid decrease in estradiol concentrations on Days – 2 to 1 and a slower decrease on Days 1 to 4 resulted in an abrupt change in the rate of decrease on Day 1 in synchrony with the mean day of the LH peak. The rapid decrease in estradiol is attributable to a negative effect of the rapidly increasing LH on estradiol,7, 17 and the slower decrease is attributable to a diminishing negative effect of the decreasing LH. The LH decrease after the peak of the surge on Day 1 is related to a negative effect of the postovulatory increase in progesterone. In another recent study, the periovulatory hormonal interrelationships were considered using collection of blood samples and ultrasound scanning every 12 hours in 18 single-ovulating mares.17 Results for Days – 1 to 2 are relevant to the previous report,22 and are depicted in Figure 214.6. The gradual preovulatory mean increase in each gonadotropin was temporarily disrupted at ovulation. Concentrations of LH and FSH increased significantly between Days – 1 to – 0.5 and Days 0.5 to 1 but not between Days – 0.5 to 0.5. The concentration of estradiol decreased significantly between Days – 1 and – 0.5 and between Days 0 and 0.5, but the decrease between Days – 0.5 and 0 was not significant. In this regard, estradiol and inhibin have a synergistic effect on suppression of FSH,23, 29 and estradiol has a negative effect on LH.29, 30 Therefore, the estradiol content of the discharged follicular fluid seems to contribute to the disruption in the LH increase at the time of ovulation, as well as to the FSH disruption. The estradiol content of a 44-mm follicle (80 µg)21 is adequate for disruption of the LH surge, based on a titration study,30 and contributes to disruption of the FSH increase.
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Concentration FSH and LH (ng/ml)
12
10
Ovulation
8 FSH
6 LH
4 180
10
170
Estradiol
8 160 6
150 140
4 130
Concentration inhibin (pg/ml)
Concentration estradiol (pg/ml)
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Inhibin
2
120 -1.0
-0.5
0.0 0.5 1.0 1.5 Days from ovulation
2.0
Figure 214.6 Means (± SEM) for periovulatory concentrations of follicle-stimulating hormone (FSH), luteinizing hormone (LH), estradiol, and immunoreactive inhibin. The increases in concentrations of FSH and LH are disrupted in temporal association with ovulation. Significant increases in FSH and LH occurred between Days – 1 and – 0.5 and between Days 0.5 and 1, but not between – 0.5 and 0.5. A significant decrease in estradiol occurred between adjacent half-days, except between Days – 0.5 and 0. Inhibin reached a peak on Day 0. These data are consistent with the discharge of follicular fluid with its estradiol and inhibin content into the peritoneal cavity, resulting in a disruption in the gonadotropin surges. Data for FSH, LH, and E2 (Ginther et al.17 ) and for inhibin (Bergfelt et al.24 ) were adapted from published reports. Modified from Ginther et al.21
Repeatability of preovulatory follicle diameter and hormones Repeatability of follicle diameters and hormone concentrations has been recently studied in mares.13, 22 Several end points were significantly correlated between consecutive ovulatory waves within mares, indicating animal repeatability. Significant correlations were found for the diameter of the preovulatory follicle during the 3 days before ovulation in spontaneous waves22 and for the maximal diameter of the preovulatory follicle in induced waves.13 When induced waves with only one ovulation were considered, there were stronger and significant correlations for diameter of the preovulatory follicle at maximum (r = + 0.70) and on the day before ovulation (r = + 0.66). Significant correlations as indicators of re-
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peatability in concentrations of FSH and LH were found at specific events throughout the induced ovulatory waves13 and are consistent with the previously reported repeatability on many days during the estrous cycle.22 Repeatability within mares in both LH concentrations and double ovulations is compatible with the concept that mares with relatively high concentrations of LH are more likely to develop double ovulations. Clinically, predicting the time of occurrence of an event (e.g., ovulation) on the basis of another (e.g., attainment of a given LH concentration) is not likely to be successful without knowledge of the history for each individual. The finding that the preovulatory follicle tends to reach a diameter that is characteristic of the mare may be useful knowledge in equine breeding programs. The significant positive correlations for diameter of the preovulatory follicle recently found occurred on the 3 days before ovulation. These days encompass the day that the follicle first reaches ≥ 35 mm in approximately 70% of the IOIs, and ≥ 35 mm is a common diameter for administration of an ovulation-inducing drug. These observations suggest that knowledge of the mare’s history concerning the diameter preceding ovulation may be useful for estimating the optimal follicle diameter for a given mare for ovulation induction, as well as for the optimal time for breeding before spontaneous ovulation.
Factors that affect preovulatory follicle diameter and hormones Breeds and types Preovulatory follicles in horses and large ponies generally reach 40–45 mm the day before ovulation. The preovulatory follicle is larger in Quarter Horses (43 mm) than in Arabians (40 mm),31 larger in one type of Thoroughbred than in another (44 vs 41 mm),32 and 3.1 mm larger for Thoroughbreds in Australia than in England.33 The diameter or growth rate of the ovulatory follicle before ovulation is similar between ponies and Quarter Horses.34 However, the maximum diameter of the preovulatory follicle is approximately 3 mm larger in French saddle horses than in Welsh ponies.35 Limited data suggest that preovulatory follicles are about 5 mm smaller in Miniature mares and 10 mm larger in Clydesdales than in Quarter Horses and large ponies.2 In a recent study, follicular dynamics was considered in Miniature mares.15 The mean diameter of the ovulatory follicle at maximum (37.3 ± 0.5 mm) was greater than on Day – 1 (35.9 ± 0.6 mm). A reduction in growth rate of the ovulatory follicle between maximum diameter (1 or 2 days before
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Table 214.1 Mean (± SEM) number of follicles per ovulatory wave and diameter and growth rate of the ovulatory follicle in Miniature ponies, large ponies, and horse mares† . End points
Miniature ponies
Large ponies
Breton horses
Bodyweight (kg, range) Age (years, range) Interovulatory interval (days) Number of growing follicles ≥ 10 mm/ovulatory wave Number of ovulatory waves with only one follicle ≥ 10 mm Diameter of ovulatory follicle (mm) At maximum On Day – 1 Growth rates (mm/day) from: Days – 6 to – 3 Days – 3 to – 1
110–136 4–12 23.3 ± 0.9a 1.5 ± 0.3a
300–400 4–13 23.9 ± 0.5a 9.8 ± 0.8b
400–530 6–13 20.3 ± 0.7b 5.8 ± 0.9c
6/11a
0/12b
0/12b
38.3 ± 0.7a 36.8 ± 1.2a
40.7 ± 1.4a 40.4 ± 1.3ab
44.5 ± 1.4b 44.0 ± 1.3b
2.9 ± 0.3X 1.0 ± 0.6a,Y
3.8 ± 0.2X 2.6 ± 0.4b,Y
3.0 ± 0.4 1.9 ± 0.4ab,#
†
n = 12 mares per group, except for 11 Miniature mares for number of waves with one follicle. Ovulation = Day 0. Means with different letters within rows are different (P < 0.05 to P < 0.0004). XY Means with different letters within columns are different (P < 0.02). # Mean on Days – 3 to – 1 tends to be different (P < 0.07) from the mean on Days – 6 to – 3. Adapted from Gastal et al.15 abc
ovulation) and ovulation was seen in the Miniature ponies and is consistent with that reported for large ponies7, 17 and horses.8, 36, 37 A hormonal mechanism possibly involved in the reduction in growth rate of the preovulatory follicle before ovulation has been discussed earlier in this chapter. The growth profile of the ovulatory follicle from Days – 6 to – 1 did not differ between left and right ovaries. However, the frequency of an ovulatory follicle and future corpus luteum (CL) in the right ovary (61%) tended to be greater than in the left ovary (39%). This latter result in the Miniature mares seems to conflict with the findings for larger breeds where ovulation was more frequent from the left ovary, especially in maiden mares.16 These apparently contrasting results indicate that Miniature ponies are a potential comparative model for studying the factors that favor ovulation from the ovary on a given side of the body. Knowledge of the diameter of the preovulatory follicle in Miniature mares is useful in comparisons of breeds and types, considering the extremely small body size. In the same study15, 38 described above, a preliminary comparison was made among Miniature ponies, large ponies, and Breton horses (Table 214.1). The Miniature ponies and the large ponies had a longer IOI than the Breton horses, agreeing with previous comparisons between large ponies and horses (reviewed in Ref. 16). The Miniature ponies had fewer growing follicles ≥ 10 mm per ovulatory wave and more ovulatory waves with only one growing follicle ≥ 10 mm compared with large ponies and horses. The diameter of the ovulatory follicle was smaller at
maximum and on Day – 1 in the Miniature ponies than in the horses, but was not different from the diameter in large ponies (Figure 214.7). The growth rate (approximately 3 mm/day) of the preovulatory follicle, prior to the cessation or reduction in growth, was similar among the three types of mares (see Figure 214.7) and agrees with reported studies in ponies and horses (reviewed in Ref. 2). Therefore, differences in diameter of the preovulatory follicle among breeds and types of mares have to be considered and incorporated in the reproductive management. These findings demonstrate that when body size difference is large (e.g., Miniature and draft horses), follicle size difference might be more pronounced among breeds and types of mares. However, these differences are
45 Ovulatory follicle
Horses Ponies
40 Diameter (mm)
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35
Miniature ponies
30 25 20 -6
-5 -4 -3 -2 Days before ovulation
-1
Figure 214.7 Means (± SEM) for diameter of the ovulatory follicle from Days – 6 to – 1 in Miniature ponies, large ponies, and horses. n = 12 mares per type. Adapted from Gastal et al.15
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Ovulation: Part 1. Follicle Development and Endocrinology small when contrasted to the great difference in body size. Season Mares are seasonally ovulatory, with the transition between the anovulatory and ovulatory seasons occurring in the spring. The spring transition has been studied extensively because of its practical importance and because comparisons between the events preceding the first ovulation and a later ovulation of the year provide insights into preovulatory endocrinology (reviewed in Refs 9 and 16). The diameter of the preovulatory follicle on the day before ovulation is about 5 mm greater before the first than before the second ovulation of the year.39, 40 Mean and peak plasma estradiol concentrations are similar between the first and second preovulatory periods of the year from the day the preovulatory follicle reaches 25 mm and the day of ovulation.41 The LH surge associated with the first ovulation of the year is of lower magnitude than the surge associated with subsequent ovulations.42–45 It has been concluded that the attenuated LH surge before the first
2027
ovulation is a consequence of incomplete recrudescence of the hypothalamic–pituitary axis.46, 47 Considering the profound differences in magnitude of the LH surge associated with the first versus later ovulations of the year, more studies are needed to determine if differences in LH concentrations between the two ovulations are associated with differences in estradiol concentrations. Such information may be related to the effects of estradiol on LH or the effects of LH on estradiol,7, 30 as discussed earlier. The temporal relationships among LH, estradiol, and follicle vascularization preceding the first versus a later ovulation of the year have been studied recently in 40 pony mares for 6 days preceding ovulation.48 Diameter of the preovulatory follicle was greater for the first ovulation of the year than for the second ovulation and for a later ovulation (Figure 214.8). Concentrations of LH were greater during the second preovulatory period than during the first preovulatory period, whereas concentrations of estradiol were not different between periods. The blood-flow area, based on color-Doppler signals, in the wall of the preovulatory follicle increased at a reduced rate during the first preovulatory period. The
Study 1 45 Diameter
First ovulation
mm
40
Study 2
35 45 Diameter
30
First ovulation
Second ovulation
40 mm
Concentration (ng/ml)
25
Concentration (pg/ml)
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7 LH 6 5
35 Later ovulation
30
4
25
3 2
2.5 Blood flow
1
2.0
Area (cm 2)
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7 Estradiol 6
1.5 1.0
5 0.5
4
-6
3
-5 -4 -3 -2 Days before ovulation
-1
2 1 -6
-5 -4 -3 -2 Days before ovulation
-1
Figure 214.8 Means (± SEM) for diameter of the ovulatory follicle and plasma concentrations of luteinizing hormone (LH) and estradiol for the 6 days preceding the first (n = 8) and second (n = 6) ovulation of the year (left panel; study 1) and for diameter of the ovulatory follicle and area of the follicle wall with color-Doppler signals for blood flow for the 6 days preceding the first ovulation of the year (spring; n = 14) and a later (summer; n = 12) ovulation (right panel; study 2). Adapted from Gastal et al.48
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vascularized area was similar between the preovulatory periods on Day – 6, but the reduced rate of increase before the first ovulation resulted in a lower area on Day – 1 than before a later ovulation. In conclusion, the first preovulatory period of the year has a lower magnitude of LH surge, similar estradiol concentrations, and lower rate of vascularization of the follicle wall than a later preovulatory period. These findings demonstrate that factors other than estradiol are involved in the differences between first and later ovulations of the year in the magnitude of the LH surge and the extent of follicle vascularization. Body condition The effects of inadequate nutrition or poor body condition on follicle dynamics and reproductive hormones during the equine ovulatory season are not fully understood. However, there is evidence that inadequate nutrition or body condition has been associated with delayed onset of the breeding season, decreased pregnancy rate, increased embryo loss, and increased gestation length in mares.49–51 During the winter, mares with low body condition had fewer medium (11–19 mm) and large (≥ 20 mm) follicles than mares with high body condition.52 The mechanisms by which feed restriction and low body condition modify the reproductive axis are also not well known. Apparently, glucose, insulin, leptin, growth hormone, and fatty acids seem to be involved, at some level, in the regulation of the reproductive axis. A study in our laboratory compared follicle activity and gonadotropin concentrations between mares of similar age with low and high body conditions.53 Examinations began during the anovulatory season (14 August, Southern Hemisphere) and continued until the second ovulation of the year. Low body condition, compared to high body condition, was associated significantly with the following: longer interval to first ovulation; smaller diameter of the preovulatory follicle at maximum (45 vs 51 mm, combined for both ovulations) and at Day – 1 (45 vs 50 mm); small diameter of the four largest follicles; fewer medium (11–19 mm) and large (≥ 20 mm) follicles; and a smaller total number of follicles ≥ 5 mm. The body condition score was positively correlated with the preovulatory follicle at maximum diameter and at Day – 1. There were no differences between groups in growth rate of the ovulatory follicle or in concentrations of FSH and LH preceding both ovulations. The finding for the gonadotropins agrees with previous studies in mares subjected to feed restriction or with low body condition. Acute54 or chronic52 feed restriction did not alter the plasma concentrations of FSH and LH. However, low plasma concen-
trations of leptin, IGF1 (insulin-like growth factor 1), and prolactin were observed in mares with low body condition score during the middle of the anovulatory season.52 Results from the previous study53 indicated for the first time in mares that low body condition was associated with reduced follicle development, including diameter of the ovulatory follicle, during the transition between the anovulatory and ovulatory seasons and during the first interovulatory interval of the ovulatory season. These results were not attributable to altered circulating concentrations of FSH and LH. More studies are needed to evaluate the interrelationships between systemic and intrafollicular hormones and body condition in mares. These studies should be specifically designed and take into account also the age of the mares as a factor, since recent studies55, 56 have shown the influence of aging in follicle dynamics and reproductive hormones (see discussion ahead).
Aging Equine and bovine comparative models have been advocated for the study of reproductive aging in women.57, 58 Mares are potentially useful research models for studying humans because of similarities in the number and nature of follicular waves and a constant relative diameter of the largest follicle between the two species at definable events throughout the ovulatory wave.12 A recent study of spontaneous ovulatory waves in mares compared the effects of age (5–6 years, young; 10–14 years, intermediate; ≥ 18 years, old) on follicle and hormone dynamics during an interovulatory interval (IOI; n = 46) and on concentrations of follicular-fluid factors.55 The old mares were not approaching senescence, as indicated by regular lengths of IOIs. The duration of the IOI was extended by about 1 day in the old group compared with the younger groups, and this was attributable to slower growth rate of the ovulatory follicle and lower LH concentrations (Figure 214.9). The old group had diminished follicle activity, as indicated by significantly smaller and fewer follicles. The diameters of the preovulatory follicle on Day – 1 and at maximum were smallest in the old group. In an early study of age in mares,59 the preovulatory follicle grew more rapidly in young mares (5–7 years) than in mares ≥ 15 years. The results of the recent study are also consistent with the finding of fewer follicles ≤ 20 mm in older ponies,59, 60 although some of the mares in the early reports were approaching senescence. Concentrations of FSH did not differ among age groups (see Figure 214.9), except that the maximum concentration was greater in the
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20
_
16
Young (5 6 yr) _ Intermediate (10 14 yr) Old (≥ 18 yr)
12 8 4 0 24
Estradiol 20
2029
the greater number of follicles in the young group reflected a greater follicle reserve and that greater LH concentrations throughout the ovulatory surge in the young group reflected a more positive response to an extraovarian/environmental influence after removal of the negative effect of progesterone. These studies indicate the importance of investigating the role of the mare as a comparative research model for age effects, for consideration of age as a potential confounding factor in equine research protocols, and for consideration of age in development of theriogenology programs (e.g., optimal time to breed and superovulation regimes).
24 LH
Concentration LH and FSH (ng/ml); estradiol (3 x pg/ml)
c214
FSH
16
Acknowledgements
12 8 4 0 -18 -16 -14 -12 -10 -8 6
8
-6 -4
-2
0
2
4
10 12 14 16 18 20 22 Days relative to ovulation
Figure 214.9 Means (± SEM) for luteinizing hormone (LH), follicle-stimulating hormone (FSH), and estradiol concentrations for young (n = 14), intermediate (n = 16), and old (n = 16) groups of mares. Greater LH concentrations in the young group was seen throughout the LH surge. Concentrations of estradiol are shown at three times the actual means to improve the presentation balance between estradiol and FSH. Estradiol concentration at the peak was greatest in the young group. Adapted from Ginther et al.55
old group. A striking finding was a greater concentration of LH in the young group than in the other two groups throughout the ovulatory LH surge (Figure 214.9), and this may have played a role in a shorter interval from maximum diameter of the preovulatory follicle to ovulation. In a previous study, LH was lower on the day of the peak of the ovulatory surge in mares approaching senescence (≥ 20 years) than in younger mares (≤ 19 years) only during one ovulatory period but not during the previous period.59 In the recent study, maximum circulating concentration of estradiol during the preovulatory surge was greatest in the young group (Figure 214.9). Concentrations of ovarian steroids in the preovulatory follicular fluid were not affected by mare age, but the concentrations of free IGF1 were greater in the old group. In another study, we have used the induction of an ovulatory wave with prostaglandin F2α treatment and ablation of follicles on Day 10 to evaluate the effects of age on the induced wave using the three previous groups of mares.56 Results suggested that
I would like to thank the Eutheria Foundation, Cross Plains, WI, and University of Wisconsin, Madison, WI, for financial support. Appreciation is expressed to Dr. Melba Gastal for helping with the figures for this chapter. Thanks are also due to Dr. O.J. Ginther for his scientific guidance and to the following colleagues who have participated during the collection of data in many of the experiments: T.J. Acosta, M.A. Beg, D. Cooper, F.X. Donadeu, M.O. Gastal, J.C. Jacob, A.P. Neves, B.L. Rodrigues, M.A. Siddiqui, L.A. Silva, and M.D. Utt.
References 1. Hunter RHF. Physiology of the Graafian Follicle and Ovulation. Cambridge, UK: Cambridge University Press, 2003. 2. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Book 2, Horses. Cross Plains, WI: Equiservices Publishing, 1995. 3. Gastal EL, Gastal MO, Bergfelt DR, Ginther OJ. Role of diameter differences among follicles in selection of a future dominant follicle in mares. Biol Reprod 1997;57:1320–7. 4. Gastal EL, Gastal MO, Beg MA, Ginther OJ. Interrelationships among follicles during the common-growth phase of a follicular wave and capacity of individual follicles for dominance in mares. Reproduction 2004;128:417–22. 5. Gastal EL, Bergfelt DR, Nogueira GP, Gastal MO, Ginther OJ. Role of luteinizing hormone in follicle deviation based on manipulating progesterone concentrations in mares. Biol Reprod 1999;61:1492–8. 6. Ginther OJ, Beg MA, Gastal MO, Gastal EL. Follicle dynamics and selection in mares. Anim Reprod 2004;1:45–63. 7. Gastal EL, Gastal MO, Ginther OJ. Relationships of changes in B-mode echotexture and colour-Doppler signals in the wall of the preovulatory follicle to changes in systemic oestradiol concentrations and the effects of human chorionic gonadotrophin in mares. Reproduction 2006;131:699–709. 8. Gastal EL, Silva LA, Gastal MO, Evans MJ. Effect of different doses of hCG on diameter of the preovulatory follicle and interval to ovulation in mares. Anim Reprod Sci 2006;94:186–90. 9. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Seasonal influence on equine follicle dynamics. Anim Reprod 2004;1:31–44.
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10. Ginther OJ. Major and minor follicular waves during the equine estrous cycle. J Equine Vet Sci 1993;13:18–25. 11. Ginther OJ, Wiltbank MC, Fricke PM, Gibbons JR, Kot K. Selection of the dominant follicle in cattle. Biol Reprod 1996;55:1187–94. 12. Ginther OJ, Gastal EL, Gastal MO, Bergfelt DR, Baerwald AR, Pierson RA. Comparative study of the dynamics of follicular waves in mares and women. Biol Reprod 2004;71:1195–201. 13. Ginther OJ, Jacob JC, Gastal MO, Gastal EL, Beg MA. Development of one vs multiple ovulatory follicles and associated systemic hormone concentrations in mares. Reprod Domest Anim 2009;44:441–9. 14. Jacob JC, Gastal EL, Gastal MO, Carvalho GR, Beg MA, Ginther OJ. Follicle deviation in ovulatory follicular waves with one or two dominant follicles in mares. Reprod Domest Anim 2009;44:248–54. 15. Gastal EL, Neves AP, Mattos RC, Petrucci BPL, Gastal MO, Ginther OJ. Miniature ponies: 1. Follicular, luteal, and endometrial dynamics during the oestrous cycle. Reprod Fert Develop 2008;20:376–85. 16. Ginther OJ. Reproductive Biology of the Mare, Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices Publishing, 1992. 17. Ginther OJ, Gastal EL, Rodrigues BL, Gastal MO, Beg MA. Follicle diameters and hormone concentrations in the development of single versus double ovulations in mares. Theriogenology 2008;69:583–90. 18. Ginther OJ, Jacob JC, Gastal MO, Gastal EL, Beg MA. Follicle and systemic hormone interrelationships during spontaneous and ablation-induced ovulatory waves in mares. Anim Reprod Sci 2008;106:181–7. 19. Urwin VE, Allen WR. Follicle stimulating hormone, luteinising hormone and progesterone concentrations in the blood of Thoroughbred mares exhibiting single and twin ovulations. Equine Vet J 1983;15:325–9. 20. Squires EL, McKinnon AO, Carnevale EM, Morris R, Nett TM. Reproductive characteristics of spontaneous single and double ovulating mares and superovulated mares. J Reprod Fertil Suppl 1987;35:399–403. 21. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Passage of postovulatory follicular fluid into the peritoneal cavity and the effect on concentrations of circulating hormones in mares. Anim Reprod Sci 2008;107:1–8. 22. Jacob JC, Gastal EL, Gastal MO, Carvalho GR, Beg MA, Ginther OJ. Temporal relationships and repeatability of follicle diameters and hormone concentrations within individuals in mares. Reprod Domest Anim 2009;44:92–9. 23. Donadeu FX, Ginther OJ. Suppression of circulating concentrations of FSH and LH by inhibin and estradiol during the initiation of follicle deviation in mares. Theriogenology 2003;60:1423–34. 24. Bergfelt DR, Mann BG, Schwartz NB, Ginther OJ. Circulating concentrations of immunoreactive inhibin and FSH during the estrous cycle of mares. J Equine Vet Sci 1991;11:319–22. 25. Roser JF, McCue PM, Hoye E. Inhibin activity in the mare and stallion. Domest Anim Endocrin 1994;11:87–100. 26. Nambo Y, Nagaoka K, Tanaka Y, Nagamine N, Shinbo H, Nagata S, Yoshihara T, Watanabe G, Groome N, Taya K. Mechanisms responsible for increase in circulating inhibin levels at the time of ovulation in mares. Theriogenology 2002;57:1707–17. 27. Miller KF, Wesson JA, Ginther OJ. Changes in concentrations of circulating gonadotropins following administra-
28.
29.
30.
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32.
33.
34.
35.
36. 37.
38.
39.
40.
41.
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tion of equine follicular fluid to ovariectomized mares. Biol Reprod 1979;21:867–72. Bergfelt DR, Ginther OJ. Delayed follicular development and ovulation following inhibition of FSH with equine follicular fluid in the mare. Theriogenology 1985;24:99–108. Miller KF, Wesson JA, Ginther OJ. Interaction of estradiol and a nonsteroidal follicular fluid substance in the regulation of gonadotropin secretion in the mare. Biol Reprod 1981;24:354–8. Ginther OJ, Utt MD, Beg MA, Gastal EL, Gastal MO. Negative effect of estradiol on LH throughout the ovulatory LH surge in mares. Biol Reprod 2007;77:543–50. Dimmick MA, Gimenez T, Schalager RL. Ovarian follicular dynamics and duration of estrus and diestrus in Arabian vs Quarter Horse mares. Anim Reprod Sci 1993;31:123–9. Vivo RR, Vinuesa SM. Diameter of the preovulatory follicle and early embryonic vesicle in Arab and Spanish Thoroughbreds. Arch Zootec 1993;42:263–7. Newcombe JR. A comparison of ovarian follicular diameter in Thoroughbred mares between Australia and the UK. J Equine Vet Sci 1994;14:653–4. Bergfelt DR, Ginther OJ. Ovarian, uterine and embryo dynamics in horses versus ponies. J Equine Vet Sci 1996;16:66–72. Palmer E. New results on follicular growth and ovulation in the mare. In: Roche JF, O’Callaghan D (eds.), Follicular Growth and Ovulation Rate in Farm Animals. Dordrecht: Martinus Nijhoff Publishers, 1987; pp. 237–55. Palmer E, Driancourt MA. Use of ultrasonic echography in equine gynecology. Theriogenology 1980;13:203–16. Koskinen E, Kuntsi H, Lindeberg H, Katila T. Predicting ovulation in the mare on the basis of follicular growth and serum oestrone sulphate and progesterone levels. Zbl Vet Med A 1989;36:299–304. Gastal EL, Gastal MO, Beg MA, Neves AP, Petrucci BPL, Mattos RC, Ginther OJ. Miniature ponies: similarities and differences from larger breeds in follicles and hormones during the estrous cycle. J Equine Vet Sci 2008;28: 508–17. Pierson RA, Ginther OJ. Follicular population dynamics during the estrous cycle of the mare. Anim Reprod Sci 1987;14:219–31. Ginther OJ. Folliculogenesis during the transitional period and early ovulatory season in mares. J Reprod Fertil 1990;90:311–20. Watson ED, Thomassen R, Nikolakopoulos E. Association of uterine edema with follicle waves around the onset of the breeding season in pony mares. Theriogenology 2003;59:1181–7. Oxender WD, Noden PA, Hafs HD. Estrus, ovulation, and serum progesterone, estradiol, and LH concentrations in mares after an increased photoperiod during winter. Am J Vet Res 1977;38:203–7. Freedman LJ, Garcia MC, Ginther OJ. Influence of photoperiod and ovaries on seasonal reproductive activity in mares. Biol Reprod 1979;20:567–74. Fitzgerald BP, Affleck KJ, Barrows SP, Murdoch WL, Barker KB, Loy RG. Changes in LH pulse frequency and amplitude in intact mares during the transition into the breeding season. J Reprod Fertil 1987;79:485–93. Sharp DC, Grubaugh WR, Weithenauer J, Davis SD, Wilcox CJ. Effects of steroid administration on pituitary luteinizing hormone and follicle-stimulating hormone in ovariectomized pony mares in the early spring: pituitary responsiveness to gonadotropin-releasing hormone
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and pituitary gonadotropin content. Biol Reprod 1991;44: 983–90. Silvia PJ, Squires EL, Nett TM. Changes in the hypothalamic-hypophyseal axis of mares associated with seasonal reproductive recrudescence. Biol Reprod 1986;35:897–905. Silvia PJ, Fitzgerald BP. Determinants of attenuated LH-release associated with the first ovulation of the equine breeding season. Domest Anim Endocrin 1991;8: 255–64. Gastal EL, Gastal MO, Donadeu FX, Acosta TJ, Beg MA, Ginther OJ. Temporal relationships among LH, estradiol, and follicle vascularization preceding the first compared with later ovulations during the year in mares. Anim Reprod Sci 2007;102:314–21. Henneke DR, Potter GD, Kreider JL, Yeats BF. Relationship between body condition score, physical measurements and body fat percentage in mares. Equine Vet J 1983;15: 371–2. Henneke DR, Potter GD, Kreider JL. Body condition during pregnancy and lactation and reproductive efficiency of mares. Theriogenology 1984;21:897–909. Hines KK, Hodge SL, Kreider JL, Potter GD, Harms PG. Relationship between body condition and levels of serum luteinizing hormone in postpartum mares. Theriogenology 1987;28:815–25. Gentry LR, Thompson DL Jr, Gentry GT Jr, Davis KA, Godke RA, Cartmill JA. The relationship between body condition, leptin, and reproductive and hormonal charac-
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teristics of mares during the seasonal anovulatory period. J Anim Sci 2002;80:2695–703. Gastal MO, Gastal EL, Spinelli V, Ginther OJ. Relationships between body condition and follicle development in mares. Anim Reprod 2004;1:115–21. McManus CJ, Fitzgerald BP. Effects of a single day of feed restriction on changes in serum leptin, gonadotropins, prolactin, and metabolites in aged and young mares. Domest Anim Endocrin 2000;19:1–13. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Siddiqui MA, Beg MA. Effects of age on follicle and hormone dynamics during the oestrous cycle in mares. Reprod Fertil Dev 2008;20:955–63. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Beg MA. Age-related dynamics of follicles and hormones during an induced ovulatory follicular wave in mares. Theriogenology 2009;71:780–8. Malhi PS, Adams GP, Singh J. Bovine model for the study of reproductive aging in women: follicular, luteal, and endocrine characteristics. Biol Reprod 2005;73:45–53. Carnevale EM. The mare model for follicular maturation and reproductive aging in the woman. Theriogenology 2008;69:23–30. Carnevale EM, Bergfelt DR, Ginther OJ. Aging effects on follicular activity and concentrations of FSH, LH, and progesterone in mares. Anim Reprod Sci 1993;31:287–99. Wesson JA, Ginther OJ. Influence of season and age on reproductive activity in pony mares on the basis of a slaughterhouse survey. J Anim Sci 1981;52:119–29.
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Ovulation: Part 2. Ultrasonographic Morphology of the Preovulatory Follicle Eduardo L. Gastal
Introduction Ovulation in the mare seems to be a culmination of a complex series of events, under elevating luteinizing hormone (LH) concentrations (see Chapter 214), that leads to rupture of a preovulatory follicle (e.g., > 30 mm) at the ovulation fossa and the extrusion of follicular fluid, granulosa cells, and the cumulus–oocyte complex. This process usually occurs 1 or 2 days before the end of the estrus. Controlling or predicting the time of ovulation within hours is important, and has practical implications for the equine industry and for research protocols that require a close interval between breeding and ovulation. Insemination close to the time of ovulation (e.g., within 6 hours) is important to prevent rebreeding mares and to maximize conception rate when using frozen semen or perhaps semen from stallions with limited sperm longevity. In addition, prediction of the time of ovulation is desired for equine oocyte retrieval programs that use assisted reproductive technologies. During recent years the mare has become an increasingly productive research model in the area of folliculogenesis (reviewed in Gastal;1 see also Chapter 214). The striking similarities between mares and women in follicle dynamics and hormonal changes during the interovulatory interval and the ovulatory follicular wave,2–6 in ultrasonographic changes of the preovulatory follicle before ovulation,7–11 and in reproductive aging processes12–14 provide the rationale for the use of, and highlights the importance of, the mare as an experimental model for the study of folliculogenesis in women. The equine model allows
hypothesis testing using invasive technologies and may provide additional information that can also be considered useful for other farm animal species and in human clinical medicine. The focus of this chapter will be on the Bmode (gray scale) and color-Doppler ultrasonographic morphological changes of the preovulatory follicle as ovulation approaches. Biochemical, molecular, and cellular mechanisms involved in ovulation15 and synchronization and induction of ovulation (Chapters 196 and 197) will not be addressed herein. This chapter is directed to those who are interested in monitoring, managing, and manipulating the mare reproductive system during the periovulatory period. Details about other aspects of ovulation in mares not presented herein can be found in previous reviews.16–21
Ultrasonographic characteristics of the preovulatory follicle Ultrasonography has brought a powerful dimension to the evaluation of the equine preovulatory follicle. High-resolution ultrasonographic machines with B-mode (gray scale) and color- and power-Doppler modes have been used recently in studies of the preovulatory follicle characteristics with different goals.10, 11,22–27 These technologies have the potential for providing clinical information on the status and future success of a follicle and its oocyte (see below). In mammals, a preovulatory follicle consists of a fluid-filled antrum encapsulated progressively (from inside to outside) by granulosa, theca interna, and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Ovulation: Part 2. Ultrasonographic Morphology of the Preovulatory Follicle theca externa.15 The avascular stratum granulosum is separated from the vascular thecal layers by a basement membrane. The theca interna is a layer of glandular cells mingled with a rich capillary plexus. The theca externa is more fibrous but contains a prominent network of arterioles and venules. As the follicle matures, expansion of the theca vessels and capillaries involves formation of new vessels as well as dilation of existing vessels.28 The main ultrasonographic characteristics of the equine preovulatory follicle with clinical and scientific importance will be addressed in the next sections. B-mode echotextural changes of the follicle wall Ultrasonographically detectable changes in grayscale echotexture and color-Doppler signals of blood flow can be detected in the wall of the future dominant follicle as early as 1 or 2 days before the beginning of follicle diameter deviation or selection in mares.29, 30 Thereafter, as the follicle matures and ovulation approaches, several ultrasonographic changes can be seen in the wall of the preovulatory follicle (Figure 215.1). Changes in follicle shape from spherical to non-spherical can be noticed from 3 days before
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ovulation, with the highest frequency occurring within 24 to 12 hours before ovulation.9 In a recent study, loss of spherical shape was associated with decreased follicle turgidity, indicated during transducer pressure, and occurred mostly between the last 24 to 12 hours before ovulation.11 The subjectivity aspect for evaluation of the previous characteristics can produce different results between operators that can be at least partly attributable to the criteria used to differentiate between spherical and non-spherical follicles and variations with transducer pressure. In regard to the follicular wall, the granulosa layer is identifiable as an echoic band enclosing the antrum9 (Figure 215.1). The two layers of theca can be assumed on the basis of position, but the boundary between the theca externa and interna is indistinct. In the initial equine B-mode ultrasonographic study,31 granulosa thickness increased as the interval to ovulation decreased. In another study,20 mean scores of echogenicity and thickness of the granulosa increased during the 24 hours before ovulation, and changes occurred in 60% and 70% of individuals, respectively. In addition to an increase in echogenicity of the granulosa, prominence of an anechoic band (Figure 215.2) in the expected area of the theca layers increased daily and progressively over 3 days before
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Figure 215.1 Sequential B-mode ultrasonograms of preovulatory follicles of one mare at 0 (a), 12 (b), 36 (c), and 40 (d) hours after the largest follicle reached ≥ 35 mm and at 41 (a), 29 (b), 5 (c), and 1 (d) hours before ovulation. As ovulation approached (top to bottom), the granulosa layer (large opposing arrows) became thicker and more echogenic. Segments of the follicular wall with an anechoic band (single short arrows) are shown. Note the decreased thickness of the granulosa at the apical area (dashed line), when compared with the thicker opposite granulosa. The distance between graduation marks (left margin) on this and the following ultrasonograms is 10 mm. Adapted from Gastal et al.10
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Figure 215.2 (a and b) B-mode ultrasonograms of preovulatory follicles of two mares at 2 days after the largest follicle reached ≥ 35 mm and 1 day before ovulation. Note the presence of the anechoic band (arrows). Images have been magnified to improve visibility of the anechoic band. Adapted from Gastal et al.10
ovulation.9, 32 Follicle diameter and the two echotexture characteristics were more prominent early than late in the season.9 Early in the season, both characteristics were at the maximal score in 70% (33/47) of follicles on day –1 (ovulation = day 0). The anechoic band is located in the area of the thecal layers and its characteristics have been described.9, 29 A similar anechoic band has been described in women.33, 34 Color-Doppler signals of blood flow are dispersed within the anechoic band, indicating that the band includes the fluid of blood vessels, presumably venules as well as arterioles (Figure 215.3). In another study,35 several mean pixel values increased in an approximately linear fashion during the 14 hours before ovulation in human chorionic gonadotropin (hCG)-treated mares. In a more recent study, mares with and without hCG treatment have been com-
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pared10 (Figure 215.4). An increase in granulosa thickness and echogenicity and a decrease in circulating estradiol during the 36 hours post-treatment were greater in the hCG group than in the controls. During 36 to 12 hours before ovulation, the granulosa thickness and echogenicity, and percentage of follicle wall with color-flow signals and prominence of the signals increased in both groups. However, 4 hours before ovulation, the two groups showed similar decreases in prominence and percentage of wall with an anechoic band and prominence and percentage of wall with color-flow signals. This study indicated that the ultrasonographic changes of the wall of the preovulatory follicle were not associated temporally with changes in estradiol concentrations and prominence of an anechoic band, and color-Doppler signals decreased during the few hours before ovulation. In
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Figure 215.3 Color-Doppler ultrasonograms of preovulatory follicles of two mares at 4 days after the largest follicle reached ≥ 35 mm. Images were taken at 2 days (a) and 3 hours (b) before ovulation. Note the anechoic band (arrows) and color-Doppler signals located within or extending beyond the width of the anechoic band. Images have been magnified to improve visibility of the anechoic band. Adapted from Gastal et al.10 (See color plate 98).
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Figure 215.4 Means (± s.e.m.) for B-mode and color-Doppler ultrasonographic end points for the wall of the preovulatory follicle in control (•) and hCG-treated (◦) groups during the preovulatory period (hours –36 to –12; n = 8–14), and impendingovulation period (hours –4 to –1; n = 4–8). For each ultrasound end point, excluding diameter, the hour effect was significant for both intervals before ovulation with no difference between groups. Adapted from Gastal et al.10
regard to age-related effects on preovulatory follicle echotexture and vascularity, a recent study has not found differences among young, intermediate, and old mares for granulosa thickness and echogenicity, anechoic band prominence, and blood flow during 4 days before ovulation.14
Recently, transrectal Doppler ultrasonography has been utilized increasingly for research and clinical studies of ovarian and follicle hemodynamics in large farm animals.18,36–38 Follicle blood flow assessment by Doppler ultrasonography has been used in mares to study follicle selection,30 anovulation during transitional seasons,39 first versus later ovulations of the year,22 follicle maturity and proximity to ovulation,10, 11, 25, 40 oocyte recovery rate, maturity, and quality,26 effects of circulatory hCG antibodies on follicle and oocyte,27 age-related effects,14 and pregnancy establishment.24, 41 The use of color-Doppler technology to evaluate the vascularity of the follicle wall in mares started in 2004. In a study of the vascular changes associated with the beginning of follicle deviation, the future dominant follicle was evaluated by transrectal color-Doppler ultrasonography until the follicle was about 30 mm (4 days before ovulation).30 The results demonstrated that deviation in the blood flow between the two largest follicles occurred 1 or 2 days before diameter deviation during follicle selection in mares. This conclusion is compatible with an earlier demonstration that an anechoic band surrounding the granulosa of the dominant follicle begins to expand differentially between the two largest follicles 1 day before the beginning of diameter deviation.29 These results provided the first evidence in any species that differential blood flow changes between future dominant and subordinate follicles begin early in the ovulatory wave and precede diameter deviation during follicle selection. Color-Doppler ultrasonography also has the potential for judging the status (future ovulatory or anovulatory) of dominant follicles during the transitional period.39, 40 During the anovulatory transitional season, vascular changes in the follicle walls of both a future dominant anovulatory follicle and a future ovulatory follicle were studied from 25 mm until 7 days after the follicle was 30 mm.39 Blood flow area was already less for dominant-sized anovulatory follicles than for ovulatory follicles by the time blood flow determinations began at 25 mm (Figure 215.5). A hypothesis for anovulation that involves hormones and follicle angiogenesis during the transitional period has been discussed elsewhere.42 In this regard, preovulatory vascular changes have been compared between the first and later ovulations of the year in 40 pony mares for 6 days preceding ovulation.22 Although follicle blood flow area (Figure 215.6) increased towards the ovulation day in both groups, results demonstrated that follicle vascularization and the LH surge were attenuated preceding the first
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Figure 215.5 Means (± s.e.m.) for follicle diameters (a) and blood flow areas of the follicle wall (b) for mares with dominant (≥ 30 mm) ovulatory (n = 11) and anovulatory (n = 6) follicles. The asterisks indicate days when the difference between follicle groups was significant. An example of expansion of an anovulatory follicle to 38 mm, despite the greatly reduced follicle blood flow area is shown for a mare with double dominant follicles (c, d). These findings demonstrate that the expansion rate of dominant follicles is not an indicator of the status or health of the follicle. Adapted from Acosta et al.39
ovulation of the year, with no indication that estradiol was involved in the differences between the first and later ovulations. In regard to preovulatory follicle blood flow, recent studies have shown a daily increase in vascularity of the wall of the dominant follicle as it matures and approaches the day of ovulation10, 14, 22, 40 (Figure 215.7). However, on the day of ovulation, a few hours before evacuation an abrupt decrease in blood perfusion in the wall of the preovulatory follicle has been detected,10, 25 as described in detail in this chapter.
Signs of impending ovulation The above studies indicate that quantitative ultrasonic characteristics of the granulosa and anechoic band are useful for sequential assessment of the developmental progress of the preovulatory follicle. However, assessing progress requires judgment on relative changes either subjectively or by computerized pixel analyses. Discrete (non-quantitative) Bmode characteristics have been described for the pre-
ovulatory follicle and may be useful for predicting time of ovulation without the necessity for judging quantitative progress. These characteristics include the following: (i) decreased turgidity of the follicle under transducer pressure in mares43 and women,44 (ii) loss of spherical shape in mares31, 43, 45 and women,7, 44, 46 (iii) irregular inner surface of the granulosa termed crenation in women7, 33 and serration in mares11, 25 (see below for details), (iv) stigma or thin apical cone-shaped or nipple-like protrusion of the follicle (future ovulation site) in mares31, 43 and women,44 (v) apparent detached segments of granulosa or rent in the wall in mares43, 47 and women,48 and (vi) echoic spots floating in the antrum in mares43 and women.49, 50 Recently, we observed that both surfaces of the granulosa (interfaces with the antrum and with the theca interna) became irregular or with a notched appearance as ovulation approached and termed the phenomenon ‘serration of granulosa’11 (Figure 215.8). In this study, the following discrete end points were recorded as present or absent: (i) serration of granulosa, (ii) decreased turgidity, (iii) loss of spherical
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last 12-hour examination was detected concomitantly with serration, but was detected alone in 9–12% of previous examinations. Serration and decreased turgidity were present at each examination until ovulation 5 hours later. Loss of spherical shape initially occurred less frequently than decreased turgidity, but the incidence increased from 50% to 100% during 6 to 1 hours before ovulation. The incidence of an apical area reached 100% and echoic spots increased to 50% during 1 hour before ovulation. The results indicated that serration of granulosa and the other discrete characteristics were useful for predicting the time of ovulation within hours, but optimal efficiency would require examinations every few hours. In this regard, owing to the appearance of serration within the last 12 hours before ovulation and a decrease in follicle blood flow during the last hours before ovulation (e.g., 6 to 4 hours10, 25 ), a model for follicle maturity and impending ovulation has been proposed (Figure 215.10). The working hypothesis was that during a few hours before follicle evacuation, the blood flow becomes concentrated to the base of the follicle (Figure 215.11) and vessels protrude throughout the wall, causing the appearance of serration in the follicular wall opposite to the future site of follicle rupture. Although the presence or absence of an indicator of impending ovulation required judgment, the presence of discrete indicators would likely be more readily evaluated than the progression in thickness and echogenicity of the granulosa and prominence of an anechoic band described in other studies. Serration of granulosa was the most useful indicator of impending ovulation, considering its distinctive appearance and consistent development only during the late preovulatory period. Future work is needed to develop a system that utilizes serration for predicting the interval to ovulation in mares.
Prediction of ovulation: preliminary results (c) Figure 215.6 Color- (a, b) and power-Doppler (c) ultrasonograms from different mares illustrating various degrees of blood flow signals in the wall of preovulatory follicles. Images were captured 24 hours before the first (a; low blood flow) or later (b and c; high blood flow) ovulations of the season. Adapted from Gastal.1 (See color plate 99).
shape, (iv) apical area, and (v) echoic spots floating in the antrum (Figure 215.9). When records were examined for 24- and 12-hour intervals, serration was detected at the last examination before ovulation in 37% and 59%, respectively. Decreased turgidity at the
Prediction of ovulation in mares is a desirable goal for the equine industry and for research purposes. Currently, there are hormonal agents such as hCG, gonadotropin-releasing hormone (GnRH), and recombinant equine LH (reLH) that can induce ovulation in most (60–90%) mares within a predictable time after treatment (e.g., 24–48 hours); however, there is a proportion (10–40%) of mares that do not respond in a timely fashion to the administration of these hormones, and presumably ovulate on their own. Therefore, regardless of the use (or not) of hormonal treatment, there are several practical situations that need the prediction of impending ovulation within 24 hours or even within a few hours. These scenarios can
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(d)
(b)
(e)
(c)
(f)
Figure 215.7 Sequential color-Doppler ultrasonograms of preovulatory follicles of two mares at 60 (a), 48 (b), 32 (c), 48 (d), 40 (e), and 24 (f) hours before ovulation. Note the increase in color-Doppler signals (amount of color dots and intensity) as the follicle matured (top to bottom). Most of the color-Doppler signals are located within or extend beyond the width of the anechoic band of the follicle wall. Adapted from Gastal et al.10 (See color plate 100).
be exemplified by cases of horse breeder associations that allow only natural mating, clinics and farms that use cooled or frozen semen, limited semen quality or quantity as a result of stallion age or morbidity, assisted reproductive techniques, and research protocols. Therefore, the use of the main ultrasonographic signs of impending evacuation of the preovulatory
follicle (see previous section) can be helpful to predict the time of ovulation in mares. Few ultrasonographic studies of the preovulatory follicle have been done with a specific design to validate a methodology to predict the time of ovulation in mares. In a detailed study, echogenicity of the granulosa layer and prominence of an anechoic band
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(b)
(c)
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Figure 215.8 B-mode ultrasonograms from four different mares illustrating the presence of serration of granulosa. The images were taken at 20 (a), 1 (b, c), and 4 (d) hours before ovulation. As ovulation approaches, both surfaces of the granulosa (interfaces with the antrum and with the theca interna) become irregular. The farthest follicle from ovulation (a) does not contain serration; however, the follicles close to ovulation (b–d) present a distinct serrated granulosa (dashed line). Adapted from Gastal et al.10 and Ginther et al.25
(a)
(b) 100
90
Serration of granulosa
Percent of mares
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80
60
Decreased turgidity
50
Loss of spherical shape Apical area
40
*
70
*
Echoic spots in antrum
30
*
60 50
*
40
20
30
*
10
20
*
0
*
10 0
n = 22
48 to 37
26
36 to 25
26
24 to 13
27
12 to 1
n = 2
4
4
6
9
12
13
14
8
7
6
5
4
3
2
1
Number of hours before ovulation Figure 215.9 (a and b) Percentage of mares with discrete ultrasonographic indicators of impending ovulation at the given number of hours before ovulation. In (b) the percentage of mares with serration of granulosa and decreased turgidity are the same (100%) for each hour. An asterisk indicates a difference (P < 0.05) in percentage of mares between two indicators. A circle around a mean shows the beginning of an increase (P < 0.05) in percentage of mares for a given indicator of impending ovulation. Adapted from Gastal et al.11
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Maximum
Blood Flow
Minimum
Serration
Apex Ovulation
) Blood Flow (
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Blood Flow
Serration Apex ? Septated Evacuation, HAF, Atresia
-24
-12
-6
-5
-4
-3
-2
-1
Hours before expected evacuation Figure 215.10 Diagrammatic representation of a proposed working model to illustrate the relationship between preovulatory follicle blood flow and serration of granulosa for mares with a normal evacuation versus a prolonged septated evacuation. In normal evacuators, follicle blood flow reaches maximum 24 hours before ovulation and starts to decrease significantly over the 6 hours before evacuation. At the hours before evacuation, serration of granulosa appears in the wall and is associated with blood flow signals at the base of the follicle and opposite to a thin and avascular apical area. In septated evacuators, follicle blood flow continues to increase between 24 and 1 hours before the beginning of evacuation. Serration and color-flow signals are dispersed throughout the periphery of the follicle, including a potential apical area. (See color plate 101).
beneath the granulosa reached a maximum score in 70% of mares on the day before ovulation.9 The efficiency of these two echotexture characteristics was compared with follicle diameter as criteria for initiating breeding early in the ovulatory season. Results indicated that the ultrasonographic echotextural characteristics were superior to diameter in identifying the optimal breeding day in mares. Another study has concluded that the slopes of regression lines for the same previous characteristics were useful in predicting impending ovulation within 24–48 hours.32 Recently, we designed two studies to allow prediction of ovulation within different hours in hCG and non-hCG treated mares (Gastal EL, Silva LA, Rocha Filho AN, Chiquetto D, Reis CP, Gastal MO, unpublished observations). Prediction of ovulation in each experiment was carried out by two different operators without the knowledge of either the previous B-mode ultrasound scan record or day of the estrous cycle of each animal. In the first experiment (Table 215.1), ultrasonographic scanning was done
every 6 hours after a ≥35 mm follicle in four groups of mares. In the second experiment (Table 215.2), the frequency of scanning was every 24, 12, or 1 hours after a ≥32 mm follicle. Attempted predictions based on each scan within each group for both studies were classified as correct or incorrect. The mean percentage of correct predictions for both experiments was 92%. However, in about 36% of the mares prediction could not be attempted due to insufficient follicular wall characteristics. These results, although limited by the number of animals, demonstrate that the degree of certainty for correct diagnosis can be high for independent operators; however, there are several mares that do not show the combination of adequate ultrasonographic follicular wall signs to be judged as impending ovulation. Therefore, studies to predict impending ovulation with B-mode and color-Doppler ultrasonography using different combinations of follicular wall characteristics (e.g., serration of granulosa and decrease in blood flow) are warranted.
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(b)
9 12
6 3
(c) 12 11
100 1
Apical area
10
2
Preovulatory Follicle
9
3
Basal area
8
% of mares (n=21)
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Serration Blood-flow
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0 Serrated granulosa Vascularized theca
5
7 6
(d)
1
2
3
4
5
6
7
8
9
10 11 12
Clock-face position (e)
Figure 215.11 B-mode image (a) of a preovulatory follicle 1 hour before ovulation showing distinct serration of granulosa (dashed line) and a color-Doppler image (b) taken at approximately the same plane. The apex is to the right but is not apparent in this plane. The clock-face method for determining positions at the periphery of a preovulatory follicle is shown (c). The apex was designated as 12 o’clock. The mean clock-face location of the serrated and vascularized area of preovulatory follicle during the hour before ovulation in 21 mares with normal follicle evacuation (requiring < 1 hour) is presented (d). The most clockwise and counterclockwise clock-face positions were determined for each end point, and the mean (± s.e.m.) for the peripheries of the vascular and serrated beds are shown. The percentage of mares at each clock-face position with serration and percentage with blood flow signals in the wall of the follicle 1 hour before the beginning of normal evacuation is presented (e). Adapted from Ginther et al.25 (See color plate 102).
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Table 215.1 Prediction of impending ovulation within 6 hours using ultrasonographic follicle characteristics.a Experiment 1. Operator 1. Prediction attempted Groups
n
Correct
Incorrectb
Total
Prediction not attempted
2500 IU hCG 1500 IU hCG 500 IU hCG Control (saline)
29 20 23 17
23 (95.8%) 14 (93.3%) 13 (86.7%) 8 (80.0%)
1 1 2 2
24 (82.8%) 15 (75.0%) 15 (65.2%) 10 (58.8%)
5 5 8 7
TOTAL
89
58 (90.6%)
6
64 (71.9%)
25
a
Signs of impending ovulation were characterized by: thick and echogenic follicle wall; pronounced anechoic layer; irregular follicle shape; irregular internal follicle wall (“serration of granulosa”); echogenic specks floating in the follicular fluid; stigma formation; and rent on the follicle wall. b Hours when ovulation occurred after prediction: 24 hours (2500 IU group); 12 hours (1500 IU); 18 and 18 hours (500 IU); 12 and 18 hours (control). Gastal et al. (unpublished).
Types of preovulatory follicle outcomes Ovulation
Normal evacuation and infundibular fluid As ovulation approaches, a bulge at the apex of the follicle can be detected at the ovulation fossa by laparoscopy51 or by ultrasonographic imaging.10, 11, 43 This thin-walled and relatively avascular portion of the preovulatory follicle25 separates an infundibular fluid pocket and the follicular antrum. A preovulatory collection of fluid external to the ovary in the infundibular area has been detected by transrectal ultrasonography.47 In a recent detailed study, an accumulation of fluid in the infundibular area was discovered by transrectal ultrasonographic imaging (Figure 215.12) and was studied daily in both oviducts of 12 mares from day –10 to day 10
(day 0 = ovulation), and from day –6 to day 6 during 35 estrous cycles of young, intermediate, and old mares. The infundibulum was identified by processes (fimbriae) and folds in the pocket of fluid (5–20 mm in diameter). Frequency of detection of fluid in the infundibular area increased between day –10 (46% of oviducts) and day –3 (88%), and decreased between day –3 and day 7 (8%).23 The source of the fluid has not been determined but presumably originates from the oviduct, peritoneum, or both. The main findings of this study were that a substantial amount of fluid accumulated in the infundibular area in mares and that the amount of accumulation depended on the stage of the estrous cycle and the side of the preovulatory follicle or ovulation, but was independent of age. The changes in the amount of fluid accumulation in the infundibular area and endometrial edema (Figure 215.12) were similar and related to previously reported changes in systemic concentrations
Table 215.2 Prediction of ovulation considering different intervals between ultrasonographic examinations. Experiment 2. Operator 2.
Frequency of examination Every 24 h Every 12 h d
Every 1 h Total a
Predicted to ovulate within 24 h 24 h 12 h 1–3 h
Resultsa Correct
Incorrect
6 (86%) 10 (91%)c 9 (100%)c 9 (100%)
1 (14%) 1 (9%) 0 (0%) 0 (0%)
34 (94%)
2 (6%)
Prediction not attempted 11 (61%)b 12 (38%) 6 (40%) 29/65 (45%)
Percentage for predictions attempted. Percentage that were not attempted because of insufficient indicators of impending ovulation; it does not include 14 of 32 mares in which a prediction was made at other than the scheduled 24 hours. c Prediction made for either 24 hours or 12 hours in each mare. d Began at 36 hours after hCG treatment or at the examination with prediction of ovulation within 12 hours in non-hCG treated mares; includes only mares in which hourly ultrasound scan was done. Gastal et al. (unpublished). b
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40 Diameter (mm)
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inf
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ap
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pof
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x
15
(a)
Amount of fluid (score 0-3)
2.5
(d)
Infundibulum
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fim
z
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x w
0.5 0
(e)
(b) Endometrial echotexture (score 1-4)
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Uterus
3.5
z
3.0 2.5 2.0
fold
y
inf
x w
1.5 1.0 -10 -8
-6
-4 -2 0 2 4 Days from ovulation
6
8
10
(f)
(c) Figure 215.12 Means (± s.e.m.) for diameter of the largest follicle (a), amount of fluid in the infundibulum (b), and endometrial echotexture (c) centered to ovulation (n = 12 mares). There was an effect of day (P < 0.0001) for the diameter of the largest follicle, amount of fluid in the infundibulum, and score for endometrial echotexture. Means with different letters (wx = increase and yz = decrease) were different (P < 0.04). (d–f) B-mode ultrasonograms from transrectal imaging illustrate the accumulation of extraovarian fluid in the infundibular area (inf). The apex (ap) (d) of the preovulatory follicle (pof) protruding toward the infundibular area (d), fimbriae (fim) (e), and fold of the infundibulum (fold) (f) are shown. Images are from three mares 1 (d) and 3 (e, f) days before ovulation. Adapted from Gastal et al.23
of estradiol. During follicle evacuation at ovulation the follicular fluid enters the infundibular fluid accumulation. The first chance observations of continuous follicle evacuation by transrectal ultrasound were made in mares.52 In subsequent planned studies, two distinctive evacuation patterns (abrupt and gradual) were observed during continuous monitoring with B-mode ultrasound.47, 53 In about 50% of ovulations, evacuation of follicular fluid from the preovulatory
follicle is an abrupt process ranging from 5 to 90 seconds, with approximately 15% of the initial fluid remaining in the antrum. In the other 50%, release of follicular fluid is a slow and gradual process taking 6–7 minutes to evacuate about 90% of the initial volume. Complete loss of detectable follicular fluid from the antrum and the extraovarian space usually takes minutes or hours, but may last as long as 2 days.25, 47 On some occasions, residual antral fluid may not be lost before blood or transudate begins to collect
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(b)
(c)
(d)
(e)
(f)
Figure 215.13 Sequential color-Doppler ultrasonograms taken during an equine ovulatory process. (a) Preovulatory follicle approximately 60 minutes before the onset of evacuation; (b) evacuating follicle shortly after the onset of rupture of the follicle wall; and (c–f) evacuating follicle taken 2.1, 2.2, 2.4, and 3.2 minutes after image (b). The sequence of images illustrates the blood flow concentrated at the basal hemisphere of the follicle directly opposite the apex or rupture point during the evacuation process. (See color plate 103).
within the antral cavity. Evacuation can be suspected to be under way when the follicle is reduced in size and irregular in shape. Examination a few minutes later and sometimes a few seconds later, if early in the process, may indicate further evacuation and confirm that ovulation is under way or has occurred (Figure 215.13). Other reports25, 54 described evacuations that were considered atypical (see Septated ovulation).
Septated evacuation Follicle evacuations with atypical sites on the day of ovulation (estimated decrease in antrum) were compartmentalized with irregular echoic septa and contained apparent follicular fluid.54 Whether complete evacuation occurred was not determined, owing to a daily interval between examinations. Atypical sites occurred in approximately 7% of hCG-treated and non-treated mares and were less common in young (3–7 years; 2%, 1/64) than in intermediate (15–19 years; 8%, 4/48), and old mares (≥ 20 years; 16%, 6/38). Preliminary observations in our previous research projects, based on hourly ultrasonographic examinations before ovulation, suggested that septated evacuations (Figure 215.14) were prolonged and may
be related to the location of blood vessels at the periphery of the follicle. A recent study was performed using colorDoppler and B-mode film clips taken of the preovulatory follicle 1 hour or every hour during 12 hours preceding the beginning of evacuation in normal and septated evacuators.25 Locations of serrated granulosa and color-flow signals were determined by clock-face positions with the apex of the follicle (future ovulation site) at 12 o’clock (Figure 215.11). Mares were divided into a group with normal follicle evacuation (completion within 1 hour; n = 21 mares) and a group with septated evacuations (completion of evacuation in ≥ 3 hours and formation of echoic trabeculae in the antrum during evacuation; n = 5 mares; Figure 215.14). The percentage of follicle circumference with color-flow signals was greater 1 hour before the beginning of evacuation in the septated group (76%) than in the normal group (37%; Figure 215.15). For mares with hourly data available before evacuation (n = 8), there was a greater decrease in the percentage of follicle circumference with color-flow signals beginning 6 hours before ovulation in the normal group than in the septated group (Figure 215.15). In the normal-evacuation group, serration and blood flow signals were located
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Septated Evacuation
(a)
F
(b)
F
apex
(c)
(d)
Normal Evacuation
Septated Evacuation
(e)
(f)
Figure 215.14 Prolonged septated evacuation 1 hour before (a) and 1 hour after (b) the beginning of an evacuation. Color signals are dispersed throughout the periphery and in the septum. The septum is an apparent crease in the evacuating follicle. Ultrasonograms in B-mode (c) and color-Doppler (d) show a preovulatory follicle 1 hour after the beginning of a septated evacuation (requiring ≥ 3 hours) and two satellite follicles. The apex and the septa within the antrum contain color-flow signals. Corpora lutea 1 hour after completion of a normal evacuation (e) and 1 hour after the end of a prolonged septated evacuation (f) are shown. Color signals after a normal evacuation are limited to the base of the corpus luteum, but are dispersed throughout the corpus luteum after a septated evacuation (f). F, satellite follicle. Adapted from Ginther et al.25 (See color plate 104).
at the basal hemisphere of the follicle directly opposite the apex (Figures 215.15 and 215.16). The apical area was devoid of both serration and color-flow signals. In the septated evacuation group, color-flow signals and serration were detected at every clockface position in each mare at the hour before ovulation. Results supported the hypothesis that prolonged septated follicle evacuation is associated with vascularization and serration of a greater circumference of the follicle than for normal evacuation (Figure 215.16), and vascularization includes the apical
area (Figures 215.10 and 215.16). The results also indicated that detectable blood flow signals were lost from the apical pole over the few hours before ovulation in the normal evacuations but not in the septated evacuations. The mechanism involved in the loss of blood flow signals from the apical area in the normal evacuations is unknown, but may have been associated with the approach of a narrower apex toward the ovulation fossa. These findings suggest that, when a reduction in vascularization at a broad apex does not occur and ovulation occurs in the presence of apical
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Normal evacuation
apex apex
base
base
(a) 100
(b) Blood flow Septated evacuations
80 % of circumference
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Normal evacuations 60
40
20 -11
-9 -7 -5 -3 Hours from beginning of evacuation
-1
Figure 215.15 Color-Doppler ultrasonograms showing preovulatory follicles 1 hour before a normal evacuation (a) with color signals concentrated only at the base, and 1 hour before the beginning of a septated evacuation (b) with color signals dispersed throughout the periphery including the apical area. The graph shows the means (± s.e.m.) for percentage of follicle circumference with blood flow signals for normal evacuators (n = 6) and septated evacuators (n = 2) during 12 hours preceding the beginning of evacuation. The color-flow signals began to decrease 6 hours before ovulation in mares with normal evacuation but not in mares with prolonged septated evacuation. Adapted from Ginther et al.25 (See color plate 105).
vessels, the evacuation is prolonged and trabeculae form in the antrum during evacuation. Studies are necessary to evaluate whether the oocyte is released from the follicle during prolonged septated evacuation and to determine the fertility of this type of ovulation.
of the ovulation site usually do not have noticeable blood flow. However, during slow septated evacuations the blood flow and serration are spread around the follicle and future ovulation site (see above and Figures 215.14, 215.15, and 215.16).
Follicle blood flow during evacuation
Entry of follicular fluid into the abdomen
As mentioned above, preliminary results from these studies also indicate that, close to the beginning of follicle wall rupture and fluid evacuation, the blood flow is concentrated at the base of the structure and associated with serration of the granulosa layer (Figures 215.13 and 215.15). During the evacuation process in normal ovulators, the middle and apical parts
The mucosa of the infundibulum consists of complex primary and secondary folds, which are covered by distinct, irregular, finger-like processes called fimbriae.16, 55 An apparent combination of permanent and transient attachment of fimbriae around the ovulation fossa assures passage of the oocyte into the oviduct while allowing passage of the majority
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Septated evacuation
(a)
(b)
(c)
(d)
Figure 215.16 Schematic drawing showing the relationship between blood flow signals and serration of granulosa for normal evacuators and prolonged septated evacuators close to ovulation (a, b), and the dispersion of blood flow signals in the newly forming corpora lutea immediately after each type of evacuation (c, d). (See color plate 106).
of the discharged follicular fluid into the abdomen. In mares, the postovulatory follicular fluid is not essential for continued oocyte development, transport, and fertilization. This has been indicated by fertilization of equine oocytes surgically transferred from follicle to ampulla56, 57 and by ovulation and fertilization after all follicular fluid has been removed by aspiration.58, 59 The proportion of follicular fluid that enters the oviduct beyond the infundibulum is unknown for any species. Studies in mares have indicated that most of the follicular fluid passes through the fimbriae into the peritoneal cavity, leaving the oocyte and its granulosa investment for entry into the oviduct.47, 52, 53, 60, 61 One of these ultrasonographic studies47 has suggested that approximately 85% of the follicular fluid passes or is filtered between the surface of the ovulation fossa and the fimbriae of the infundibulum into the peritoneal cavity. In this regard, a spike in circulating concentrations of immunoreactive inhibin occurs on the day of ovulation,62 and results from the direct exposure of the peritoneum to the high concentrations of inhibin in the discharged follicular fluid.63, 64 During follicle evacuation, fluid in the oviduct was not detectable except for the fluid accumulation in the infundibular area that was present before follicle evacuation.47 The
fluid pocket in the infundibulum did not enlarge and the remaining oviduct did not become detectable by ultrasonographic imaging during and after follicular evacuation in mares, despite the large volume (e.g., 30–45 mL) of follicular fluid evacuated. In addition, we have not seen enlargement of any part of the oviduct during ultrasonographic monitoring of several preovulatory follicle evacuations in our research projects.
Early corpus luteum blood flow After ovulation, the basement membrane between the granulosa and thecal layers breaks down in association with invasion of blood vessels from the theca interna into the developing corpus luteum. The spatial relationship between the vascularity of the preovulatory follicle and developing corpus luteum has been recently studied in mares25 and for the first time in any species. Color-Doppler and B-mode film clips from 26 mares were taken of the preovulatory follicle 1 hour before the beginning of ovulation. The pattern of the progressive development of blood flow signals in various portions (basal third, middle third, and apical third) of the developing corpus luteum was evaluated every 12 hours until day 6. Color-flow
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signals were determined by clock-face positions with the apex of the follicle (future ovulation site) or corpus luteum at 12 o’clock (Figure 215.11). Mares with normal follicle evacuation and prolonged septated evacuation were considered. Results indicated that vascularization of the ovulation site in the normalevacuating mares and subsequent vascularization of the corpus luteum began at the base of the structure (Figure 215.16). This was shown by the greater percentage of blood flow signals in the basal third than in the middle and apical thirds of the newly forming corpus luteum. This process involved the incorporation of at least some of the vessels from the follicle base into the collapsed basal ovulation site. There was a progressive increase in blood flow signals from base to apex of the corpus luteum over the 6 days after ovulation. This was shown by the beginning of a successive increase in blood flow signals over the basal, middle, and apical areas. This finding is compatible with previous studies65, 66 that showed a progressive increase in plasma progesterone concentrations and percentage of corpus luteum with blood flow signals to near maximal levels during the first week after ovulation in normal evacuators. In the slow septated evacuation group, peripheral vessels seemed to contribute to the vessels of the newly forming corpus luteum uniformly in all parts (basal, middle, apical) of the structure (Figure 215.16). Hemorrhagic anovulatory follicle Extravasation of blood into follicles, ovulation sites, and corpora lutea is common in mares.16, 17 A hematoma that forms in the antrum instead of ovulation has been termed a hemorrhagic anovulatory follicle (HAF).52, 67 Similar structures have also been termed hemorrhagic follicles,17 anovulatory hemorrhagic follicles,68, 69 persistent anovulatory follicles,70 and autumn follicles.71 The economic importance of HAFs as a breeding-management problem in mares has been noted70, 72, 73 and reflects anovulation of a follicle after the mare has been bred. The extent of the problem is indicated by a 5% and 20% incidence during the early and late ovulatory season, respectively.74 Moreover, the syndrome seems to be more common in old mares and may occur repeatedly in an individual, sometimes encompassing much or all of the breeding season. The morphology and vascularity of HAFs in mares and the endocrinology immediately preceding HAF formation have been studied in control and HAF groups.75 The day of ovulation and the first day of HAF formation, as indicated by cloudiness of follicular fluid (Figure 215.17), have been defined as day 0. The frequency of discrete gray-scale ultrasonic
indicators of impending ovulation11 and follicle diameter on day –1 did not differ between future ovulating and HAF groups. On day –1, the circumference of the follicle wall of future HAFs had more colorDoppler signals of blood flow than in the control mares (Figure 215.17). In regard to hormones, higher estradiol concentrations occurred a few days before HAF formation, but systemic LH, follicle-stimulating hormone (FSH), and progesterone were not altered during conversion of a preovulatory follicle into an HAF. In a recent comparative study of induced waves using prostaglandin F2α (PGF) treatment on day 10 and ablation of follicles ≥ 6 mm and spontaneous ovulatory waves, an unexpected high incidence of HAFs (24%; n = 21) occurred in the induced waves and none in the spontaneous waves.76 The induced ovulatory waves differed considerably from spontaneous waves, including greater LH concentrations during much of the induced wave and greater growth rate of a smaller ovulatory follicle. In this regard, results of another recent study77 indicated that the high incidence of HAFs after PGF/ablation was associated with later follicle emergence and immediate and continuing greater LH concentration after PGF treatment, apparently augmented by an inherently high pretreatment LH concentration. Furthermore, a recent histological and immunohistochemical study was unable to determine conclusively the participation of several angiogenic factors or the LH receptor in HAF formation.78 Studies of HAF formation and the underlying mechanisms in mares may be of comparative importance for other animal species and women that have similar ultrasonographic structures. More details about HAFs can be found in Chapters 222 and 279. Atresia The process of atresia of a preovulatory follicle during a major ovulatory wave is not a common finding during the natural equine breeding season, but it occurs quite often during follicular waves in the spring and fall transitional seasons. A postulated hormonal mechanism for anovulation of a preovulatory follicle in mares has been proposed42 and its effects may possibly lead to changes in the follicular wall that could be appreciated through ultrasonography during a transitional season.
Vascularity of the preovulatory follicle versus fertility The degree of vascular perfusion of the ovary and follicles, assessed by color- or power-Doppler ultrasonography, has been used as a potential new technology for research and clinical studies of ovarian
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Figure 215.17 Color-Doppler ultrasonograms showing blood flow signals in the wall of the preovulatory follicle on the day before ovulation (a) and the day before anovulation or beginning of the formation of a hemorrhagic anovulatory follicle (HAF) (b). More blood flow signals can be seen in the wall of the future HAF. Gray-scale ultrasonograms (c–f) of the same mare illustrate the most common sequential events in formation of an HAF. Day 0 (equivalent to the day of ovulation), excessive floating specks in antrum. Day 1, floating specks and the beginning of echoic strands. Day 2, network of strands that quivered upon ballottement. (See color plate 107).
and follicle hemodynamics and to predict fertility in horses, cattle, and humans. Increased follicle blood flow, along with a rapid increase in LH at the terminal stage of follicle maturation, has been associated with meiosis resumption and completion of oocyte maturation. Greater vascularity of the preovulatory follicle has been associated with greater follicle diameter (women,79 mares,24 heifers41 ), re-
trieval rate of oocytes (women,79 mares,26 heifers80 ), retrieval rate of mature oocytes (mares26, 27, 81 ), in vitro fertilization rate (women,79 heifers80 ), pregnancy rate (women,79,82–86 mares,24 heifers41 ), and lower incidence of triploidy (women82 ). In addition, follicles with greater blood flow resulted in better embryos and more pregnancies after embryo transfer in women.79, 82, 83, 85, 87
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Figure 215.18 Means (± s.e.m.) for percentage of follicle circumference and prominence of color-Doppler signals of blood flow, and resistance index (RI) and pulsatility index (PI) for mares that became pregnant (n = 14) or non-pregnant (n = 7; left). An injection of 2500 IU of human chorionic gonadotropin (hCG) was given when the largest follicle was 34–37 mm (hour 0), and mares were bred 30 hours later (hour 30). The indices are inversely related to the extent of tissue vascular perfusion downstream from the point of assessment at the most prominent intraovarian color signal. Color-Doppler ultrasonograms (right) illustrate blood flow signals in the wall of preovulatory follicles of a mare that became pregnant (a) and a mare that did not become pregnant (b). Images were taken 30 hours after hCG treatment. Note the high (a) and low (b) percentages of follicular wall with blood flow signals. Adapted from Silva et al.24 (See color plate 108).
The relationships of oocyte maturity, 30 hours after hCG treatment, to blood flow (color-Doppler mode) and ultrasonographic characteristics of impending ovulation (B-mode) of the preovulatory follicle have been recently studied in mares.26 The vascularity of the preovulatory follicle tended to be greater for follicles that contained a mature oocyte. Follicles with an oocyte recovered had a significantly greater frequency of serration of granulosa and greater frequency of an apical area (indicators of impending ovulation). The frequency of serration of granulosa and decreased turgidity was greater in follicles that contained a mature oocyte. Results indicated that recovery of an oocyte and maturity of the recovered oocytes are both dependent upon the rate of follicle maturity in response to the hCG treatment. Recent studies in our laboratory tested the hypothesis that a higher pregnancy rate is associated with greater blood flow to the preovulatory follicle before breeding. The studies used color- and powerDoppler ultrasonography and indicated that follicle blood flow was greater in mares24 and heifers41 that became pregnant. In both of these studies, mares and heifers were previously treated with hCG and GnRH, respectively, to induce ovulation. Follicle blood flow
was evaluated at the time of treatment and natural breeding or artificial insemination. Although the results of both studies are preliminary, since they involved a limited number of animals, important statistical differences were found between animals that became pregnant versus non-pregnant in each study. In the mare study,24 B-mode echogenicity and thickness of the granulosa layer and prominence of the anechoic band beneath the granulosa increased similarly in both pregnant and non-pregnant groups. An increase in follicle diameter and percentage of follicle circumference with color-Doppler signals was greater between the time of hCG treatment (hour 0) and artificial insemination (hour 30) in the pregnant group than in the non-pregnant group (Figure 215.18). Spectral-Doppler measurements were made at the most prominent intraovarian color signal. Decreases in resistance and pulsatility indices were greater between hours 0 and 30 in the pregnant group than in the non-pregnant group, indicating increased vascular perfusion downstream from the spectral measurement in the pregnant group. Relative peak systolic velocity and time-averaged maximum velocity of blood flow at the point of spectral assessment was greater in the pregnant
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Ovulation: Part 2. Ultrasonographic Morphology of the Preovulatory Follicle group. Results supported the hypothesis that greater blood flow to the preovulatory follicle is associated with higher pregnancy rate. Similar results were found in the heifer study,41 demonstrating that highly vascularized preovulatory follicles are more likely to be associated with higher pregnancy rates, as previously seen for mares and women.
Influence of hCG antibodies on follicle and oocyte: preliminary results Treatment with hCG has been used frequently in mares when the preovulatory follicle is 32–35 mm to induce ovulation or to induce maturity of the oocyte for collection by aspiration of follicle contents 28–36 hours after treatment.16, 57 The effect of exposure to a previous hCG treatment or the presence of hCG antibodies on the reliability of ovulation induction by hCG administration has not been resolved adequately. Some reports have indicated a reduction in the efficacy of hCG for inducing ovulations when given during several estrous cycles,88–91 whereas others have indicated that refractoriness does not develop from repeated hCG treatments.92–96 Owing to high molecular weight (36.5 kDa) and glycoprotein properties of hCG, antibodies may develop following hCG treatment during repeated estrous cycles.27, 89, 93, 97 Most of the reports on the efficiency of hCG for induction of ovulation following previous hCG exposure did not have information on the presence of hCG antibodies. Apparently, the variable effectiveness of repeated hCG treatment is attributable to the production of antibodies only in some mares (e.g., 42%),27 and successful ovulation induction within 48 hours after hCG treatment in some mares (34%) despite high levels of antibodies.89 More elaborate studies, taking into account the effects of age and season, are needed to clarify many aspects on this topic. Studies are also needed to determine the maturity and quality of collected oocytes after an hCG treatment in the presence of circulating hCG antibodies. A recent study in mares27 investigated the effect of an ovulation-inducing dose of hCG in the presence versus absence of hCG antibodies on blood flow of the preovulatory follicle, follicular-fluid hormone concentrations, systemic estradiol and LH concentrations, and maturity and quality of recovered oocytes at 30 hours post-treatment. Mares were grouped as positive (n = 16) and negative (n = 44) for hCG antibodies before the experimental hCG treatment when the preovulatory follicle was ≥ 32 mm. The hCG antibodies were completely effective in neutralizing the hCG by 30 hours, as indicated by
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non-detectable hCG in the mares that were positive for antibodies. The percentage of the follicle wall with blood flow signals was less in the antibodypositive group than in the antibody-negative group. The concentrations of follicular-fluid estradiol and free insulin-like growth factor-1 (IGF1), and plasma estradiol were greater, and follicular-fluid progesterone and plasma LH were less in the antibodypositive group than in the antibody-negative group. The oocyte-recovery rate (62%, 37/60) between hCG antibody-positive (44%) and -negative mares (68%) tended to be different. The antibody-positive group had fewer mature (MII) oocytes than the antibodynegative group. The decrease in the percentage of Doppler signals of blood flow in the follicle wall of the antibody-positive group indicates that the antibodies neutralized the hCG so that hCG was not available for hastened development of the follicle and its oocyte. The highly effective neutralization of the hCG in the antibody-positive group indicates that a history of previous hCG treatment should be a consideration in using hCG for oocyte collection programs. These results suggest the need for the development of hCG protocols that would reduce the incidence of antibody formation ensuring follicle and oocyte maturation. Furthermore, studies with various doses of hCG and intervals from previous exposures, and a rapid and simple method for quantitating hCG antibodies in each mare, would have practical applications.
Acknowledgements I would like to thank the Eutheria Foundation, Cross Plains, WI, and the University of Wisconsin, Madison, WI, for financial support. Appreciation is expressed to Dr. Melba Gastal for helping with the figures of this chapter. Thanks are also due to Dr. O.J. Ginther for his scientific guidance during our studies and the following colleagues who have participated during the collection of data in some of the experiments: T.J. Acosta, M. Almamun, M.A. Beg, D. Cooper, F.X. Donadeu, M.O. Gastal, J.C. Jacob, A.P. Neves, B.L. Rodrigues, M.A. Siddiqui, L.A. Silva, and M.D. Utt.
References 1. Gastal EL. Recent advances and new concepts on follicle and endocrine dynamics during the equine periovulatory period. Anim Reprod 2009;6:144–58. 2. Ginther OJ, Beg MA, Gastal MO, Gastal EL. Follicle dynamics and selection in mares. Anim Reprod 2004;1:45–63. 3. Ginther OJ, Gastal EL, Gastal MO, Bergfelt DR, Baerwald AR, Pierson RA. Comparative study of the dynamics of follicular waves in mares and women. Biol Reprod 2004;71:1195–201.
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4. Ginther OJ, Beg MA, Gastal EL, Gastal MO, Baerwald AR, Pierson RA. Systemic concentrations of hormones during development of follicular waves in mares and women: a comparative study. Reproduction 2005;130:379–88. 5. Mihm M, Evans ACO. Mechanisms for dominant follicle selection in monovulatory species: comparison of morphological, endocrine and intraovarian events in cows, mares and women. Reprod Dom Anim 2008;43 (Suppl. 2): 48–56. 6. Baerwald AR. Human antral folliculogenesis: what we have learned from bovine and equine models. Anim Reprod 2009;6:20–9. 7. Martinuk SD, Chizen DR, Pierson RA. Ultrasonographic morphology of the human preovulatory follicle wall prior to ovulation. Clin Anat 1992;5:339–52. 8. Pierson RA, Chizen DR. Transvaginal ultrasonographic assessment of normal and aberrant ovulation. In: Jaffe R, Pierson RA, Abramowicz JS (eds) Imaging in Infertility and Reproductive Endocrinology. Philadelphia: J.B. Lippincott, 1994; pp. 129–42. 9. Gastal EL, Gastal MO, Ginther OJ. The suitability of echotexture characteristics of the follicular wall for identifying the optimal breeding day in mares. Theriogenology 1998;50:1025–38. 10. Gastal EL, Gastal MO, Ginther OJ. Relationships of changes in B-mode echotexture and colour-Doppler signals in the wall of the preovulatory follicle to changes in systemic oestradiol concentrations and the effects of human chorionic gonadotrophin in mares. Reproduction 2006;131:699–709. 11. Gastal EL, Gastal MO, Ginther OJ. Serrated granulosa and other discrete ultrasound indicators of impending ovulation in mares. J Equine Vet Sci 2006;26:67–73. 12. Carnevale EM. The mare model for follicular maturation and reproductive aging in the woman. Theriogenology 2008;69:23–30. 13. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Siddiqui MAR, Beg MA. Effects of age on follicle and hormone dynamics during the oestrous cycle in mares. Reprod Fertil Dev 2008;20:955–63. 14. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Beg MA. Agerelated dynamics of follicles and hormones during an induced ovulatory follicular wave in mares. Theriogenology 2009;71:780–8. 15. Hunter RHF. Physiology of the Graafian Follicle and Ovulation. Cambridge, UK: Cambridge University Press, 2003. 16. Ginther OJ. Reproductive Biology of the Mare, Basic and Applied Aspects. 2nd edn. Cross Plains, WI: Equiservices Publishing, 1992. 17. Ginther OJ. Ultrasonic Imaging and Animal Reproduction. Book 2. Horses. Cross Plains, WI: Equiservices Publishing, 1995; pp. 35–40. 18. Ginther OJ. Ultrasonic Imaging and Animal Reproduction. Book 4. Color-Doppler Ultrasonography. Cross Plains, WI: Equiservices Publishing, 2007. 19. Pierson RA. Folliculogenesis and ovulation. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 161–71. 20. Carnevale EM. Folliculogenesis and ovulation. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasound. Baltimore: Williams & Wilkins, 1998; pp. 201–11. 21. Bergfelt DR, Adams GP. Ovulation and corpus luteum development. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders Elsevier, 2007; pp. 1–13.
22. Gastal EL, Gastal MO, Donadeu FX, Acosta TJ, Beg MA, Ginther OJ. Temporal relationships among LH, estradiol, and follicle vascularization preceding the first compared with later ovulations during the year in mares. Anim Reprod Sci 2007;102:314–21. 23. Gastal EL, Jacob JCF, Gastal MO, Ginther OJ. Accumulation of fluid in the infundibulum during the estrous cycle in mares. J Equine Vet Sci 2007;27:251–9. 24. Silva LA, Gastal EL, Gastal MO, Beg MA, Ginther OJ. Relationship between vascularity of the preovulatory follicle and establishment of pregnancy in mares. Anim Reprod 2006;3:339–46. 25. Ginther OJ, Gastal EL, Gastal MO. Spatial relationships between serrated granulosa and vascularity of the preovulatory follicle and developing corpus luteum. J Equine Vet Sci 2007;27:20–7. 26. Ginther OJ, Gastal EL, Gastal MO, Siddiqui MAR, Beg MA. Relationships of follicle versus oocyte maturity to ultrasound morphology, blood flow, and hormone concentrations of the preovulatory follicle in mares. Biol Reprod 2007;77:202–8. 27. Siddiqui MAR, Gastal EL, Gastal MO, Beg MA, Ginther OJ. Effect of hCG in the presence of hCG antibodies on the follicle, hormone concentrations, and oocyte in mares. Reprod Dom Anim 2009;44:474–9. 28. Familiari G, Vizza E, Miani A, Motta PM. Ultrastructural and functional development of the theca interna. In: Familiari G, Makabe S, Motta PM (eds) Ultrastructure of the Ovary. Boston: Kluwer Academic Publishers, 1991; pp. 115–28. 29. Gastal EL, Donadeu FX, Gastal MO, Ginther OJ. Echotextural changes in the follicular wall during follicle deviation in mares. Theriogenology 1999;52:803–14. 30. Acosta TJ, Gastal EL, Gastal MO, Beg MA, Ginther OJ. Differential blood flow changes between the future dominant and subordinate follicles precede diameter changes during follicle selection in mares. Biol Reprod 2004;71:502–7. 31. Pierson RA, Ginther OJ. Ultrasonic evaluation of the preovulatory follicle in the mare. Theriogenology 1985;24:359–68. 32. Chan JPW, Tsung-Hsiang H, Chuang ST, Cheng FP, Chen CL, Mao CL. Quantitative echotexture analysis for prediction of ovulation in mares. J Equine Vet Sci 2003;23:397–402. 33. Picker RH, Smith DH, Tucker MH, Saunders DM. Ultrasonic signs of imminent ovulation. J Clin Ultrasound 1983;11:1–2. 34. Jaffe R, Ben-Aderet N. Ultrasonic screening in predicting the time of ovulation. Gynecol Obstet Invest 1984;18:303–5. 35. Carnevale EM, Checura CM, Coutinho da Silva MA, Adams GP, Pierson RA. Use of computer-assisted image analysis to determine the interval before and after ovulation. Proc Am Assoc Equine Pract 2002;48:48–50. 36. Ginther OJ, Utt M. Doppler ultrasound in equine reproduction: principles, techniques and potential. J Equine Vet Sci 2004;24:516–26. 37. Miyamoto A, Shirasuna K, Hayashi KG, Kamada D, Awashima C, Kaneko E, Acosta TJ, Matsui M. A potential use of color ultrasound as a tool for reproductive management: new observations using color ultrasound scanning that were not possible with imaging only in black and white. J Reprod Dev 2006;52:153–60. 38. Herzog K, Bollwein H. Application of Doppler ultrasonography in cattle reproduction. Reprod Dom Anim 2007;42:51–58. 39. Acosta TJ, Beg MA, Ginther OJ. Aberrant blood flow area and plasma gonadotropin concentrations during the
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40.
41.
42.
43.
44.
45.
46.
47.
48.
49.
50.
51. 52. 53.
54.
55. 56.
57. 58.
59.
development of dominant-sized transitional anovulatory follicles in mares. Biol Reprod 2004;71:637–42. Palmer E, Chavatte-Palmer P, Verdonck E. Field trial of Doppler ultrasonography of the preovulatory follicle in the mare. Anim Reprod Sci 2006;94:182–5. Siddiqui MA, Almamun M, Ginther OJ. Blood flow in the wall of the preovulatory follicle and its relationship to pregnancy establishment in heifers. Anim Reprod Sci 2009;113:287–92. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Seasonal influence on equine follicle dynamics. Anim Reprod 2004;1: 31–44. Carnevale EM, McKinnon AO, Squires EL, Voss JL. Ultrasonographic characteristics of the preovulatory follicle preceding and during ovulation in mares. J Equine Vet Sci 1988;8:428–31. Hanna MD, Chizen DR, Pierson RA. Characteristics of follicular evacuation during human ovulation. Ultrasound Obstet Gynecol 1994;4:488–93. Townson DH, Ginther OJ. Size and shape changes in the preovulatory follicle in mares based on digital analysis of ultrasonic images. Anim Reprod Sci 1989;21:63–71. Lenz S. Ultrasonic study of follicular maturation, ovulation and development of corpus luteum during normal menstrual cycles. Acta Obstet Gynecol Scand 1985;64:15– 19. Townson DH, Ginther OJ. Ultrasonic characterization of follicular evacuation during ovulation and the fate of the discharged follicular fluid in mares. Anim Reprod Sci 1989;20:131–41. Bourne TH, Jurkovic D, Waterstone J, Campbell S, Collins WP. Intrafollicular blood flow during human ovulation. Ultrasound Obstet Gynecol 1991;1:53–9. Mendelson EB, Friedman H, Neiman HL, Calenoff L, Vogelzang RL, Cohen MR. The role of imaging in infertility management. Am J Roentgenol 1985;144:415–20. Fleischer A, Kepple DM. Transvaginal Sonography: A Clinical Atlas, 2nd edn. Philadelphia: J.B. Lippincott, 1995; pp. 186–9. Witherspoon DM, Talbot RB. Ovulation site in the mare. J Am Vet Med Assoc 1970;157:1452–9. Ginther OJ, Pierson RA. Ultrasonic anatomy of equine ovaries. Theriogenology 1984;21:471–83. Townson DH, Ginther OJ. Duration and pattern of follicular evacuation during ovulation in the mare. Anim Reprod Sci 1987;17:155–63. Carnevale EM, Bergfelt DR, Ginther OJ. Follicular activity and concentrations of FSH and LH associated with senescence in mares. Anim Reprod Sci 1994;35:231–46. Sisson S, Grossman JD. The Anatomy of the Domestic Animals, 4th edn. Philadelphia: W.B. Saunders, 1953; pp. 607–9. McKinnon AO, Carnevale EM, Squires EL, Voss JL, Seidel GE. Heterogenous and xenogenous fertilization of in vivo matured equine oocytes. J Equine Vet Sci 1988;8:143–7. Carnevale EM. Oocyte transfer and gamete intrafallopian transfer in the mare. Anim Reprod Sci 2004;82–83:617–24. Palmer E, Duchamp G, Cribiu E, Mahla R, Boyazoglu S, B´ezard J. Follicular fluid is not a compulsory carrier of the oocyte at ovulation in mare. Equine Vet J 1997; (Suppl. 25): 22–4. Gastal EL, Gastal MO. Effects of intrafollicular injections of hCG, follicular puncture and follicular fluid aspiration to induce ovulation of preovulatory follicles in mares. In: Proceedings of the 14th International Congress on Animal Reproduction, vol. 1, 2000, p. 28.
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60. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Dynamics of the equine preovulatory follicle and periovulatory hormones: What’s new? J Equine Vet Sci 2008;28:454–60. 61. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Passage of postovulatory follicular fluid into the peritoneal cavity and the effect on concentrations of circulating hormones in mares. Anim Reprod Sci 2008;107:1–8. 62. Bergfelt DR, Mann BG, Schwartz NB, Ginther OJ. Circulating concentrations of immunoreactive inhibin and FSH during the estrous cycle of mares. J Equine Vet Sci 1991;11:319–22. 63. Nambo Y, Hasegawa T, Sato F, Oki H, Kusunose R, Nakai R, Nagata S, Watanabe G, Taya K. The release of follicular fluid into the peritoneal cavity during ovulation in mares. Theriogenology 2002;58:545–8. 64. Nambo Y, Hashimoto K, Nakai R, Asai Y, Watanabe G, Taya K. Biological significance of follicular fluid discharged into the peritoneal cavity at ovulation in mares. Anim Reprod Sci 2006;94:228–31. 65. Bollwein H, Mayer R, Weber F, Stolla R. Luteal blood flow during the estrous cycle in mares. Theriogenology 2002;57:2043–51. 66. Ginther OJ, Gastal EL, Gastal MO, Utt MD, Beg MA. Luteal blood flow and progesterone production in mares. Anim Reprod Sci 2006;99:213–20. 67. Ginther OJ, Pierson RA. Regular and irregular characteristics of ovulation and the interovulatory interval in mares. J Equine Vet Sci 1989;9:4–12. 68. Carnevale EM, Squires EL, McKinnon AO, Harrison LA. Effect of human chorionic gonadotropin on time to ovulation and luteal function in transitional mares. Equine Vet Sci 1989;9:27–9. 69. Lefranc A, Allen WR. Incidence and morphology of anovulatory haemorrhagic follicles in the mare. Pferdeheilkunde 2003;19:611–12. 70. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. 71. Knudsen O, Weiert V. Ovarian oestrogen levels in the nonpregnant mare: relationship to histological appearance of the uterus and to clinical status. J Reprod Fertil 1961;2:130–7. 72. McKinnon AO. Ovarian abnormalities. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Wilkins, 1998; pp. 233–51. 73. Pycock JF. Breeding management of the problem mare. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. Philadelphia: W.B. Saunders, 2000; pp. 195–228. 74. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Incidence, endocrinology, vascularity, and morphology of hemorrhagic anovulatory follicles in mares. J Equine Vet Sci 2007;27:130–9. 75. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Conversion of a viable preovulatory follicle into a hemorrhagic anovulatory follicle in mares. Anim Reprod 2006;3:29–40. 76. Ginther OJ, Jacob JC, Gastal MO, Gastal EL, Beg MA. Follicle and systemic hormone interrelationships during spontaneous and ablation-induced ovulatory waves in mares. Anim Reprod Sci 2008;106:181–7. 77. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Beg MA. Induction of haemorrhagic anovulatory follicles in mares. Reprod Fertil Dev 2008;20:947–54. 78. Ellenberger C, Muller K, Schoon HA, Wilsher S, Allen WR. Histological and immunohistochemical characterization of equine anovulatory haemorrhagic follicles (AHFs). Reprod Dom Anim 2009;44:395–405.
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79. Bhal PS, Pugh N, Chui D, Gregory L, Walker SM, Shaw RW. The use of transvaginal power Doppler ultrasonography to evaluate the relationship between perifollicular vascularity and outcome in in-vitro fertilization treatment cycles. Hum Reprod 1999;14:939–45. 80. Siddiqui MA, Gastal EL, Gastal MO, Almamun M, Beg MA, Ginther OJ. Relationship of vascular perfusion of the wall of the preovulatory follicle to in vitro fertilisation and embryo development in heifers. Reproduction 2009;137:689–97. 81. Siddiqui MAR, Gastal EL, Ju JC, Gastal MO, Beg MA, Ginther OJ. Nuclear configuration, spindle morphology and cytoskeletal organization of in vivo maturing horse oocytes. Reprod Dom Anim 2009;44:435–40. 82. Bhal PS, Pugh ND, Gregory L, O’Brien S, Shaw RW. Perifollicular vascularity as a potential variable affecting outcome in stimulated intrauterine insemination treatment cycles: a study using transvaginal power Doppler. Hum Reprod 2001;16:1682–9. 83. Coulam CB, Goodman C, Rinehart JS. Colour Doppler indices of follicular blood flow as predictors of pregnancy after in-vitro fertilization and embryo transfer. Hum Reprod 1999;14:1979–82. 84. Huey S, Abuhamad A, Barroso G, Hsu MI, Kolm P, Mayer J, Oehninger S. Perifollicular blood flow Doppler indices, but not follicular pO2, pCO2, or pH, predict oocyte developmental competence in in vitro fertilization. Fertil Steril 1999;72:707–12. 85. Du B, Takahashi K, Ishida GM, Nakahara K, Saito H, Kurachi H. Usefulness of intraovarian artery pulsatility and resistance indices measurement on the day of follicle aspiration for the assessment of oocyte quality. Fertil Steril 2006;85:366–70. 86. Shrestha SM, Costello MF, Sjoblom P, McNally G, Bennett M, Steigrad SJ, Hughes GJ. Power Doppler ultrasound assessment of follicular vascularity in the early follicular phase and its relationship with outcome of in vitro fertilization. J Assist Reprod Genet 2006;23:161–9. 87. Chui DKC, Pugh ND, Walker SM, Gregory L, Shaw RW. Follicular vascularity – the predictive value of transvaginal power Doppler ultrasonography in an in-vitro fer-
88.
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tilization programme: a preliminary study. Hum Reprod 1997;12:191–6. Sullivan JJ, Parker WG, Larson LL. Duration of estrus and ovulation time in nonlactating mares given human chorionic gonadotropin during three successive estrous periods. J Am Vet Med Assoc 1973;162:895–8. Wilson CG, Craig RD, Hughes PJ, Roser JF. Effects of repeated hCG injections on reproductive efficiency in mares. J Equine Vet Sci 1990;10:301–8. McCue PM, Hudson JJ, Bruemmer JE, Squires EL. Efficacy of hCG at inducing ovulation: a new look at an old issue. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2004, pp. 510–13. Green JM, Raz T, Epp T, Carley SD, Card CE. Relationships between utero-ovarian parameters and the ovulatory response to human chorionic gonadotrophin (hCG) in mares. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007, pp. 563–7. Loy RG, Hughes JP. The effects of human chorionic gonadotropin on length of estrus and fertility in the mare. Cornell Vet 1966;56:41–50. Roser JF, Kiefer BL, Evans JW, Neely DP, Pacheco CA. The development of antibodies to human chorionic gonadotropin following its repeated injection in the cyclic mare. J Reprod Fertil 1979; (Suppl. 27): 173–9. Michel TH, Rossdale PD, Cash RSG. Efficacy of human chorionic gonadotrophin and gonadotrophin releasing hormone for hastening ovulation in Thoroughbred mares. Equine Vet J 1986;18:438–42. Barbacini S, Zavaglia G, Gulden P, Marchi V, Necchi D. Retrospective study on the efficacy of hCG in an equine artificial insemination programme using frozen semen. Equine Vet Educ 2000;12:312–17. Gastal EL, Silva LA, Gastal MO, Evans MJ. Effect of different doses of hCG on diameter of the preovulatory follicle and interval to ovulation in mares. Anim Reprod Sci 2006;94:186–90. Duchamp G, Bour B, Combarnous Y, Palmer E. Alternative solutions to hCG induction of ovulation in the mare. J Reprod Fertil 1987;(Suppl. 35):221–8.
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Chapter 216
Luteal Development Don R. Bergfelt and Gregg P. Adams
Introduction The luteal gland (corpus luteum, CL) is a temporary endocrine gland within the ovary that forms in place of a previous dominant follicle after ovulation near the end of estrus (primary CL) or during diestrus or early pregnancy (secondary CL) in the mare as reviewed in numerous reference texts.1–9 A CL may also result from luteinization of a dominant anovulatory follicle, especially during early pregnancy (accessory CL). The primary CL is the principal source of progesterone that terminates estrous behavior and maintains the mare in a non-receptive state during diestrus. Moreover, the primary CL is the initial source of progesterone that assists in preparing the tubular genitalia, especially the uterus, to receive the blastocyst and support embryo and fetal development during early pregnancy. If the mare is not pregnant or spontaneously loses the embryo before approximately Day 16 after ovulation then the CL regresses in response to uterine synthesis and secretion of prostaglandin F2α (PGF2α ) and returns to estrus. If, however, the mare does respond to the ‘first luteal response to pregnancy’ (i.e., embryo suppression of the luteoytic mechanism) then the primary CL is maintained and continues to produce progesterone along with supplemental CL (combined secondary and accessory CL) that form after Day 30 until the placenta becomes the principal source of progesterone by Days 50 to 70 of gestation. Thus, the luteal gland plays a pivotal role during the estrous cycle and early pregnancy and, therefore, detection and evaluation of the CL is critical in providing structural and functional information to assist scientists, clinicians, and breeding farm managers in comprehending and controlling luteal gland development as well as mitigating potential luteal gland dysfunction. Transrectal palpation is relatively accurate for detecting ovulation in the mare,2 but it is a highly subjective and inaccurate approach for detecting and assessing the status of luteal gland development,
especially if the timing of ovulation is not known. In this regard, the unique anatomical arrangement of the equine ovary that prevents protrusion of the luteal gland from the ovarian surface lends itself to the power of ultrasonography. Ultrasonic imaging has proven to be an invaluable research and diagnostic tool in equine reproduction;2,10–16 it can provide immediate detection and a visual and objective evaluation of both structural and functional aspects of luteal gland development from maturation to regression. Although collection of a blood sample and assay of serum or plasma progesterone concentration is the ultimate direct test for detecting the functional status of a CL,1–9 it may be impractical when an immediate assessment of the luteal gland is required. In this regard, an ultrasound examination can provide indirect hormonal information regarding the functional status of luteal gland development. Based on the results from numerous studies as reviewed,1, 3, 8, 10,12–14 circulating concentrations of progesterone are highly correlated with ultrasonographic morphological changes (i.e., CL diameter and area, and luteal tissue echogenicity) and physiological changes (i.e., glandular blood flow) during maturation and regression in non-pregnant and pregnant mares. This chapter is intended to consolidate into a single source principal elements from previous2,10–15 and more recent8, 14, 16 publications on the ultrasonographic detection and evaluation of the equine CL. The approach of gathering information on the presence and status of the luteal gland with B-mode and color-Doppler ultrasonography will be discussed in the context of morphological, physiological, and endocrinological relationships primarily during the ovulatory season in non-pregnant mares but will include some aspects in pregnant mares. Most of the data presented herein are average values obtained from numerous mares under experimental conditions and may not necessarily reflect what is observed
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 216.1 Luteal gland terminology commonly used in association with the estrous cycle and early pregnancy in the mare Luteal structure
Definition
Primary corpora lutea
Result from single or multiple synchronous and asynchronous ovulations (≤ 2 days from primary ovulation) during the follicular phase at the end of estrus (estrogen dominance)
Secondary corpora lutea
Result from ovulations (> 2 days from primary ovulation) during the luteal phase of diestrus and early pregnancy. (progesterone dominance)
Corpora hemorrhagica
Primary or secondary corpora lutea that develop an intraluteal blood-filled cavity subsequent to ovulation
Accessory corpora lutea
Result from luteinization of anovulatory follicles, especially during early pregnancy
Supplemental corpora lutea
Include secondary and accessory corpora lutea that develop during diestrus and early pregnancy
Corpora albicantia
Regressed corpora lutea regardless of reproductive status
Adapted and revised from Ginther OJ. Prolonged luteal activity in mares – a semantic quagmire. Equine Vet J 1990;22:152–6.
under clinical or field conditions in individual animals. The mean results, which are often encompassed by an indicator of population variance (i.e., standard error of the mean, SEM), are intended to serve as a guide that can be used as a reference to estimate or predict a particular outcome in an individual mare.
Terminology Terminology associated with luteal gland development is primarily based on reproductive status (stage of the estrous cycle or pregnancy), hormonal status (estrogen or progesterone dominance) and pathological conditions associated with either the uterus or ovaries as discussed in the following section and summarized from the original source17 in Table 216.1.
especially during the period of eCG production associated with early pregnancy, results in accessory corpora lutea. The term supplemental corpora lutea includes both secondary and accessory CL. Regressed luteal glands, which typically occur during late diestrus (Figure 216.2a) or around the middle of pregnancy, are termed corpora albicantia whether or not they have remnants of an intraluteal cavity.
Luteal Gland Morphology a)
Corpus luteum without intraluteal cavity
b)
Corpus luteum with intraluteal cavity (corpus hemorrhagicum)
Luteal glands The primary corpus luteum results from ovulation of the dominant follicle of the primary follicular wave terminating the estrous cycle. Luteal glands that result from ovulation of dominant follicles of a secondary follicular wave during diestrus, or from large follicles present at the time of equine chorionic gonadotropin (eCG) production in early pregnancy are termed secondary corpora lutea. Notably, secondary CL result from ovulations occurring during periods of progesterone dominance (i.e., diestrous, early pregnancy) and need to be differentiated from primary CL, which develop from synchronous (same day) or asynchronous (1 to 2 days apart) double ovulations in association with estrus and estrogen dominance. Primary and secondary CL can develop with or without a blood-filled intraluteal cavity as depicted grossly and ultrasonographically in Figures 216.1 and 216.2. Luteal glands with an intraluteal cavity are termed corpora hemorrhagica (Figures 216.1b and 216.2a). Luteinization of anovulatory follicles,
Figure 216.1 Photographs and sonograms depicting development of (a) an early primary corpus luteum without an intraluteal cavity and (b) a corpus luteum with an intraluteal, bloodfilled cavity (corpus hemorrhagicum). Note and compare the demarcations between ovarian stroma and luteal tissue (a and b), and luteal tissue and blood clot (b) in the photographs with that in the sonograms. Measurement of cross-sectional diameter or area of the corpus luteum can be done with electronic calipers as illustrated by the yellow perpendicular lines or with the scale bars displayed on the left side of each sonogram. Gross morphology adapted from Ginther, 199233 and ultrasonic morphology adapted from Bergfelt et al.34
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Luteal Gland Development a)
B-Mode Ultrasonography
b)
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Figure 216.2 Sonograms taken with (a) B-mode and (b) color-Doppler ultrasonography depicting development of the primary corpus luteum without an intraluteal cavity and with an intraluteal blood-filled cavity (corpus hemorrhagicum, a) during various stages of the luteal phase. Note and compare changes in luteal gland size and luteal tissue echogenicity and blood flow from early to late diestrus. B-mode ultrasonography adapted from Bergfelt et al.34 and color-Doppler ultrasonography adapted from Ginther.35 (See color plate 109).
Luteal gland irregularities The term prolonged luteal function has been used previously to indicate luteal persistence without regard to etiology.1 However, with the advent of ultrasonography, specific conditions of uterine (i.e., pyometra, mucometra) and ovarian (i.e., secondary ovulations and hemorrhagic anovulatory follicles) origin have been shown to be related to prolongation of the luteal phase, as reviewed.1, 2, 5, 9 Briefly, the more contemporary phrase uteropathic persistence of the corpus luteum has been used to describe a prolonged luteal phase as a result of reduced synthesis and secretion of PGF2α associated with pyometra, which is a severe inflammatory condition of the uterus. Consequently, regression of the CL is delayed and the luteal phase or
estrous cycle is prolonged. In the absence of any apparent uterine or ovarian irregularity, the term idiopathic persistence of the corpus luteum has been suggested.17 Abbreviated luteal function has also been attributed to both uterine and ovarian irregularities, as reviewed1, 2, 5, 9 and as discussed in Chapters 222 and 223 in this text. Briefly, the phrase uteropathic abbreviation of the corpus luteum has been used to describe a shortened luteal phase as a result of premature synthesis and secretion of PGF2α associated with endometritis, which is a less severe inflammatory condition of the uterus. Consequently, regression of the CL is hastened and the luteal phase of the estrous cycle is shortened. The term luteal insufficiency has been used to describe abbreviated luteal function
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as a result of a dysfunctional CL in association with early embryonic loss. Notably, the prevalence of luteal insufficiency preceding embryonic loss during the ovulatory season is thought to be rare;18, 19 however, luteal gland dysfunction was demonstrated to precede embryonic loss following gonadotropin releasing hormone (GnRH)-induced ovulation during the anovulatory season, especially when diameter of the largest follicle was ≤ 15 mm at the start of treatment.20
Approach The techniques and methodologies involved in the use of B-mode and color-Doppler ultrasonography to detect and evaluate luteal gland development in the mare are founded on the basic principles of ultrasonography, which have been well documented in the horse.2,10–16 These references are invaluable for fully comprehending and interpreting the imaging processes involved in the various ultrasound techniques and methodologies discussed in this section that will allow for a safe and thorough examination of the ovaries so that detecting the presence of a CL is accurate and evaluation of its status is precise. Technique The latest makes and models of ultrasound instruments, imaging capabilities and incidental provisions vary widely. While some machines and transducers are specifically produced for veterinary use, most are produced for human use but can be readily adapted for use in animals. Transducers are available in various types (e.g., linear-array, convex-array) and frequencies (e.g., 3.5, 5.0, 7.5 MHz), and can be used intrarectally, intravaginally and transcutaneously. The transcutaneous route is not practical for detecting the luteal gland in the mare because of the distance between the transducer and ovary and consequent loss of resolution. Correspondingly, the transvaginal route may not be practical in field situations because of the extra effort to maintain sanitary conditions. However, in a clinical or research laboratory, the transvaginal route has been used as a relatively non-invasive and repeatable approach to reach the ovary and manipulate follicular and luteal development for basic and applied purposes.3, 8,12–16 Nonetheless, the transrectal route with a linear-array transducer and a frequency of 5 to 8 MHz is the most common ultrasound approach for scanning the ovary to detect and evaluate the CL from maturation to regression in all types of horses, including miniature horses/ponies21 and donkeys.22 In regard to the latter, some mares may be too small to allow both hand and transducer to enter and pass guardedly into the
rectum. However, by affixing a semi-rigid material (e.g., rubber-like tubing) to the transducer cord similar to that used for scanning sheep23 and prepubertal cattle,24 the transducer can be placed within the rectum and guided and maneuvered externally. Although this unguarded approach is considered safe and effective for scanning the reproductive tract, it is emphasized that the transducer is being manipulated externally in the rectum and extreme caution is recommended to protect against rectal injury. Location of either the left or right ovary by the transrectal approach using either the guarded or unguarded technique may be initially frustrating for individuals with minimal experience in one or both techniques, but is quickly grasped with a few examination attempts. Identification of the ovary may be facilitated by first viewing a cross-section of a uterine horn and, second, by slowly rotating the transducer laterally following the uterine horn cranially to the tip and beyond, when the ovary should come into view. In some instances, the ovary may be difficult to view or only partially visible or the image may not be clear because of intervening tissue or air (e.g., intestines) between the rectal wall and ovary. Correspondingly, the orientation or position of the ovary may change while an examination is in progress and the image may be partially or wholly obscured. In this regard, the ovary may have to be repositioned by transrectal palpation or, perhaps, by jogging the mare if palpation is not possible, to ensure that the entire ovary can be thoroughly scanned for accurate detection and precise assessment of luteal gland development. Methodology Ultrasonographic detection of the presence of a luteal gland within an ovary requires the capability to distinguish between ovarian stroma and luteal tissue. The intensity of the brightness and darkness assigned to the numerous pixels of a particular image on the ultrasound screen is referred to as tissue echogenicity,10–16 which is based on the ultrastructure of the luteal gland and surrounding tissue. Denser connective tissue of the ovarian stroma will reflect more of the ultrasound echoes from the transducer back to the transducer and be registered as light pixels (echoic), whereas less dense tissue of the luteal gland will reflect fewer of the ultrasound echoes and be registered as dark pixels (anechoic). In general, therefore, an ultrasound image of the CL and surrounding area will have various areas of lightness and darkness because of the different types of tissues and corresponding densities, which depend on the stage of development of the luteal gland. Hence, it is warranted to have a basic understanding of
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Luteal Development the principles of ultrasonography10–16 and ovarian macroscopic and microscopic anatomy, especially as it pertains to the luteal gland.1, 2 Morphological aspects of the tissue remodeling processes and the biochemical and physiological changes associated with luteogenesis in the mare have been reviewed.1, 2 In general, follicle theca cells begin degenerating just prior to ovulation and are nearly completely degenerated by 24 hours after ovulation as the follicular fluid is evacuated and the follicle wall collapses. Correspondingly, follicle granulosa cells are transformed from estrogensecreting cells to progesterone-secreting luteal cells. Initially, therefore, an immature CL will exhibit a relatively high degree of echogenicity during early diestrus (Figure 216.2a) as a result of the opposing walls of the follicle and denser luteal tissue associated with the early processes of luteinization. As the CL matures, luteal cells are characterized by an increase in diameter, vesiculated nuclei and fine vacuoles in the cytoplasm, all of which indicate an increase in cellular synthesis and secretion of progesterone. Concurrent with structural and functional changes associated with luteinization, neovascularization and genesis of blood and lymphatic vessels accompanies the invasion of folds of stromal tissue into the luteinizing tissue. Hence, a mature CL will exhibit a relatively lower degree of luteal tissue echogenicity during mid-diestrus (Figure 216.2a) as a result of sparser tissue associated with enhanced luteal cell hypertrophy and secretion, and increased glandular blood and lymph flow. Reportedly,1, 2 luteinization and vascularization are complete by 3–5 days after ovulation. As the CL regresses, luteolysis involves cellular degeneration and devascularization and, therefore, a regressed CL will exhibit a higher degree of luteal tissue echogenicity during late diestrus (Figure 216.2a). Regardless of the changes in luteal tissue echogenicity during maturation and regression, there is a distinct demarcation at the junction between the more vascularized and sparse luteal tissue of the CL and the less vascularized and dense connective tissue of the ovarian stroma (Figure 216.1a,b) due to a difference in impedance of the ultrasound echoes at the tissue interfaces. As a result, the CL is readily discernible from ovulation until late regression, when it is no longer distinguishable from ovarian stroma. The detection of a luteal gland subsequent to ovulation involves an understanding that development of a CL can proceed in a morphological manner that often involves the formation of an intraluteal blood-filled cavity.1–3,8, 10,12–14 The formation of an intraluteal cavity associated with corpora hemorrhagica (Table 216.1) occurs randomly from one estrous cycle to the next some 50–70% of the time.
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Even in mares with multiple ovulations during the estrous cycle or early pregnancy, formation of corpora hemorrhagica is considered an independent event. Based on the differences in tissue echogenicity, the intraluteal blood clot is readily distinguishable from the luteal tissue that encompasses the clot, as depicted in Figures 216.1b and 216.2a. Subsequent to ovulation, the ratio of non-luteal to luteal tissue of immature corpora hemorrhagica at early diestrus is relatively large (Figure 216.2a) and is first detectable in a majority of mares on the day of ovulation (Day 0; 28%) or Day 1 (62%). As corpora hemorrhagica reach maturity around mid-diestrus (Figure 216.2a), the luteal portion of the gland changes slightly compared with the intraluteal portion, which decreases in size in association with an increase in connective tissue and organization of the clot. In some instances, the intraluteal cavity can still be identified at late diestrus (Figure 216.2a). During early pregnancy, however, luteal glands that initially formed an intraluteal cavity may later become relatively uniform and resemble a CL that develops without a cavity. In regard to the latter, luteal glands develop without an intraluteal cavity 30–50% of the time and remain relatively uniform throughout their lifespan (Figures 216.1a and 216.2a). Ultrasonographic evaluation of luteal gland development in the mare has historically involved measuring CL diameter and area, luteal tissue echogenicity and, more recently, glandular blood flow.10,12–14 Quantitative data generated by these measurements have been used to characterize luteal gland development for basic and applied purposes during the estrous cycle and early pregnancy, as illustrated in Figures 216.3 and 216.4, respectively. Measurement of the cross-sectional diameter or area of the luteal gland can only be accomplished after the CL has been distinguished from the surrounding ovarian stroma as previously discussed. Although the CL may appear mushroom- or gourdlike in shape with an irregular boundary as ovulation is restricted to the fossa (Figure 216.1a,b), a relatively accurate assessment of size can be determined by taking the average of two lines of measurement using the electronic calipers of the ultrasound machine or by estimation using the calibrated grid that accompanies the ultrasound image, as shown in Figure 216.1a,b. After the CL has been frozen at its greatest cross-sectional size, the first measurement is made across the largest portion of the gland extending from the interface between stromal and luteal tissues on one side of the CL to the opposite side (Figure 216.1a,b). The second determination is made similarly but approximately perpendicular to the first. The first and second measurements are averaged to give an overall cross-sectional diameter.
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Days after ovulation Figure 216.3 Mean (± SEM) for (a) cross-sectional diameter and (b) area of the primary corpus luteum and (c) luteal tissue echogenicity and (d) blood flow in relation to circulating concentrations of progesterone (e,f) during various stages of the luteal phase. Adapted from Bergfelt and Adams.36
Diameter or radius of the CL can be converted to cross-sectional area using the formula for area of a circle (A = πr2 ). Notably, surface area (4πr 2 ) and volume (4/3πr 3 ) of the CL are representative of function and, therefore, may be better diagnostic measurements of the luteal gland for both clinical and research purposes as suggested.25, 26 Nonetheless, for a corpus hemorrhagicum, the cross-sectional diameter or area of intraluteal tis-
sue can be adjusted for the non-luteinized portion of the gland by subtracting the respective dimensions of the intraluteal cavity. Despite a larger overall diameter of a corpus hemorrhagicum, the absolute luteal tissue area is comparable between CL with and without an intraluteal cavity, which is supported by the similarity in daily profiles of progesterone concentrations.12 The mean daily profiles for diameter and area of the primary CL are depicted in Figure 216.3a,b
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Figure 216.4 Mean (± SEM) cross-sectional diameter of the primary corpus luteum (a), luteal tissue echogenicity (b), and circulating concentrations of progesterone (c) in non-pregnant mares (open circles) during the luteal phase and in pregnant mares (filled circles) during luteal maintenance (first luteal response to pregnancy) and resurgence (second luteal response to pregnancy). Note, mares with supplemental corpora lutea (third luteal response to pregnancy) were not included in the progesterone profile. Adapted from Bergfelt et al.34
during the estrous cycle and in Figure 216.4a during early pregnancy. Quantitating the degree of luteal tissue echogenicity is based on assigning a grayscale score to correspond with the degree of pixel lightness or darkness of an ultrasound image. Although most ultrasound machines will have as many as 256 gray shade levels, the human eye can only distinguish 18 to 20 levels. Nonetheless, the visual approach has involved assigning the luteal portion of the CL a grayscale score ranging from 0 (black) to 4 (white) according to a grayscale bar typically displayed on
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the ultrasound monitor. Alternatively, computerassisted image analysis may be a more objective and consistent approach to evaluate luteal tissue echogenicity, one that can utilize the full range of gray shades between black (score = 0) and white (score = 255). The availability of image analysis algorithms has been described.11, 27, 28 Regardless of visual or computer-assisted approaches to evaluate luteal tissue echogenicity, the gain settings (e.g., near, far, and overall fields) on the ultrasound scanner console need to be standardized from scan to scan within and among animals. It is emphasized that the operator is still responsible for selecting and delineating the areas of the luteal gland to be analyzed. In a review12 of a study using both the visual and computer-assisted approaches to evaluate luteal tissue echogenicity during the estrous cycle, hourly profiles in grayscale values were comparable. The mean daily profiles of luteal tissue echogenicity as determined using the visual approach are depicted in Figure 216.3c during the estrous cycle and in Figure 216.4b during early pregnancy. Considering the similarity in the results for luteal tissue echogenicity between the visual and computer-assisted approaches, the former approach seems more practical as a diagnostic tool for assessing luteal gland development. However, it should be recognized that more recent ultrasound machines are equipped with the capability to store numerous images and, perhaps, run the image analysis software. Alternatively, ultrasonic images can be downloaded to a computer at the clinic or laboratory, or to a server while still in the field using a wireless internet connection.27 Regardless, high-quality ultrasound images can be readily collected, archived, transported, and recalled for subsequent analysis or to confirm a previous diagnosis. Hence, the visual and computer-assisted image analysis approaches may be used alone or in combination to make an immediate or initial assessment of luteal gland echogenicity. However, for a more precise clinical diagnosis and for scientific hypothesis testing, the latter approach may be preferred. The feasibility of color-Doppler ultrasonography in equine reproduction was first documented13 in 1998, when it was demonstrated that color-flow images (indicative of blood flow) associated with the ovary (follicle and CL) and early pregnancy (embryo/fetus) could be generated and recorded. Since then, color-Doppler technology has advanced such that many portable ultrasound machines have been designed to include both B-mode and color-Doppler capabilities. The principles of B-mode/color-Doppler ultrasonography for use in domestic animal reproduction have recently been published by Ginther (2007),14 which includes results from numerous
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studies addressing basic and applied topics, especially in the mare. The CL is one of the most vascularized organs in the body and, in the mare, color-flow imaging can be readily applied to the relatively large luteal gland to evaluate glandular blood flow as an indicator of physiological integrity. In an earlier study,29 computer-assisted image analysis was used to estimate the number of colored pixels per luteal gland image during the estrous cycle. In a later study,14 a visual assessment was made of the percentage of the CL with color-Doppler signals. Both approaches provided an approximate quantitation of the extent of luteal tissue vascularity during the estrous cycle, as depicted qualitatively for the later approach14 in Figure 216.2b and quantitatively for the earlier approach29 in Figure 216.3d.
Structural and functional development Both earlier10–13 and more recent reviews8, 14 provide evidence that changes in the ultrasonographic morphology (e.g., diameter, area, echogenicity) of the CL are reflective of the ultrastructural changes associated with the physiology (e.g., blood flow) and endocrinology (e.g., progesterone) of the luteal gland during maturation and regression in the mare. The mean temporal relationships among these various aspects are summarized in Figure 216.3 during the estrous cycle and in Figure 216.4 during early pregnancy, to emphasize how the use of B-mode or color-flow ultrasonic imaging or a combination of both approaches can provide real-time structural and functional information for testing hypotheses and making diagnoses and prognoses concerning luteal gland development in the mare. Ultrasonographic characteristics of the CL in relation to progesterone during the estrous cycle
Early diestrus Beginning at ovulation on Day 0 until about Day 5, functional maturation of the primary CL is indicated by a progressive increase in circulating concentrations of progesterone (Figure 216.3e,f) that relate morphologically to an increase in CL diameter (Figure 216.3a) and area (Figure 216.3b), and an increase followed by a decrease in luteal tissue echogenicity (Figure 216.3c), and, physiologically, to an increase in glandular blood flow (Figure 216.3d). The transient change in echogenicity of the CL during early diestrus is attributed, in part, to overlap of the ovulation processes (e.g., evacuation of follicular fluid and
collapse of opposing follicle walls) with the initial processes involving luteinization and vascularization (e.g., invasion of folds of stromal tissue into the luteinizing tissue). Although changes in luteal tissue echogenicity during early diestrus may not be as readily observable as size and vascularity, luteal tissue echogenicity of the immature CL as determined by computer-assisted image analysis has been investigated as being predictive of the number of hours after ovulation that could potentially be used to assist in scheduling the timing of post-ovulatory insemination.30 Nonetheless, increase in CL diameter, area, and vascularity after ovulation are distinctly reflective of an increase in progesterone and, therefore, diagnostic of CL maturation during early diestrus.
Mid-diestrus Between Days 5 and 10, functional maturation of the primary CL is complete, as indicated by maximal progesterone concentrations (Figure 216.3e,f), which relate morphologically to a large CL diameter (Figure 216.3a) and area (Figure 216.3b), low luteal tissue echogenicity (Figure 216.3c) and, physiologically, to high glandular blood flow (Figure 216.3d). Notably, after Day 5, diameter, area, and vascularity of the CL begin to gradually decrease, which is not necessarily reflected in respective directional changes in luteal tissue echogenicity and progesterone concentrations. Although there may be some ambiguity in diagnosing a functionally mature CL at mid-diestrus based on some ultrasonographic characteristics, recording and integrating size, echogenicity and vascularity is still informative by eliminating an early or later stage of luteal gland development.
Late diestrus After Day 10, functional regression of the primary CL is indicated by a precipitous decrease in the concentration of progesterone (Figure 216.3e,f), which distinctly relates morphologically to a decrease in CL diameter (Figure 216.3a) and area (Figure 216.3b) and an increase in luteal tissue echogenicity (Figure 216.3c) and, physiologically, to a decrease in glandular blood flow (Figure 216.3d). Hence, the respective directional changes in size and luteal tissue echogenicity and vascularity are diagnostic of CL regression during late diestrus. Ultrasonographic characteristics of the CL in relation to progesterone during early pregnancy The use of ultrasonography during early pregnancy has been well documented;10–16 however, its use to study and monitor luteal gland development has
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Luteal Development received limited attention, which seems contrary to the important role that luteal gland function (i.e., progesterone production) plays in maintaining pregnancy for the first 50 to 70 days of gestation. Nonetheless, the temporal relationships between circulating concentrations of progesterone and CL diameter and luteal tissue echogenicity using B-mode ultrasonography are summarized and illustrated in Figure 216.4. Similar to the endocrinological and morphological relationships associated with the primary CL during the estrous cycle in non-pregnant mares (Figure 216.3), functional changes in circulating concentrations of progesterone (Figure 216.4c) relate to structural changes in cross-sectional diameter (Figure 216.4a) and luteal tissue echogenicity (Figure 216.4b) in pregnant mares until about Day 13 after ovulation. Thereafter, profiles of the functional and structural aspects differed as a result of regression of the primary CL in non-pregnant mares. In a field study,31 the cross-sectional area of the primary CL expressed as a volume ratio and progesterone concentrations were higher on Days 8 to 9 in pregnant mares compared to non-pregnant mares. It seems, therefore, that pregnancy status was indicated earlier during mid-diestrus (Days 8 to 9) using CL volume ratio and later during late diestrus (approximately Day 13) using CL diameter. Perhaps, quantitating the change in luteal tissue area as volume versus diameter is a more sensitive measure of luteal function, as suggested.26, 27 Regardless, separate studies that were done in pregnant mares document the practicality of ultrasonic imaging as an immediate aid in the diagnosis and prognosis of luteal gland development in association with the first luteal response to pregnancy.1 Moreover, morphological maintenance of size and echogenicity of the primary CL in pregnant mares is informative of the developmental status of the early embryo (i.e., intrauterine mobility) since the uterine luteolytic mechanism has been inactivated. Functional and structural resurgence of the primary CL in pregnant mares occurs between Days 30 and 35 after ovulation and represents the second luteal response to pregnancy as a result of formation of endometrial cups and synthesis and secretion of equine chorionic gonadotropin (eCG). Temporally, an increase in CL diameter precedes the increase in circulating concentrations of progesterone (Figure 216.4a,c). Apparently, luteal tissue echogenicity was less sensitive to detect resurgence of the CL, perhaps because values were already at minimal levels (Figure 216.4b). Nonetheless, ultrasonic imaging can be an immediate aid in the diagnosis and prognosis of luteal gland development in association with the second luteal response to pregnancy. Moreover, morphological resurgence of the primary CL is informative of the developmental status of the later embryo
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(i.e., embryo–uterine interaction in the formation of endometrial cups) since resurgence in diameter of the primary CL is not detected in mares with embryo loss before endometrial cup formation.32 Formation of supplemental luteal glands occurs during eCG production and represents the third luteal response to pregnancy.1 The initial formation of supplemental CL results from luteinization of dominant ovulatory follicles, termed secondary corpora lutea, and later from luteinization of unovulated dominant follicles, termed accessory corpora lutea (Table 216.1). The role of supplemental CL in maintaining pregnancy before a placental source of progesterone has been established has not been critically studied. For instance, it is not unusual to find mares at 60 to 160 days of pregnancy without supplemental CL, especially if they were bred late in the ovulatory season. Ultrasonographically, it appears that secondary CL develop in a morphologically similar fashion to the primary CL (Figure 216.1). That is, subsequent to ovulation, some secondary CL are uniform whereas others form an intraluteal blood-filled cavity (corpus hemorrhagicum). Since accessory corpora lutea develop from unovulated follicles, they inherently have a relatively uniform intraluteal blood-filled cavity and seem to follow a developmental pattern similar to that of hemorrhagic anovulatory follicles during diestrus or seasonal transitional periods discussed in Section C (Chapters 178, 180, 181, and 182). Notably, practitioners should be aware and differentiate accordingly between healthy ovaries that are enlarged due to increased follicular and luteal activity from 40 to 120 days of gestation, and neoplastic ovaries, which are pathological. Regardless, ultrasonic imaging can be an immediate aid in the diagnosis and prognosis of luteal gland development in association with the third luteal response to pregnancy.
Summary Ultrasonographic imaging allows immediate detection and evaluation of luteal gland development in mares. Technologies involving B-mode and colorDoppler ultrasonography used alone or together are invaluable as research, diagnostic, and prognostic tools in equine reproduction. Morphological (CL diameter and area and luteal tissue echogenicity) and physiological (glandular blood flow) characteristics of the luteal gland are highly correlated with the endocrinological (progesterone production) characteristics of maturation and regression during the estrous cycle and early pregnancy. Thus, ultrasonographic imaging is the most contemporary approach, for researchers, clinicians, and breeding farm managers to efficiently, effectively, and safely examine
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the ovaries for accurate detection and precise evaluation of stage of development of the equine luteal gland.
References 1. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices Publishing, 1992;642 pp. 2. McKinnon AO, Voss JL. Equine Reproduction, 2nd edn. Philadelphia: Lea & Febiger, 1993;1137 pp. 3. Samper JC. Equine Breeding Management and Artificial Insemination, 1st edn. Philadelphia: WB Saunders, 2000; 306 pp. 4. Davies Morel MCG. Equine Reproductive Physiology, Breeding and Stud Management. Oxford: CABI Publishing, 2003; 384 pp. 5. Frazer GS. Reproduction. In: Edward Robinson N (ed.) Current Therapy in Equine Medicine, 5th edn. Philadelphia: Saunders, 2003; pp. 1–930. 6. Knottenbelt DC, Pascoe RR, Leblanc M, Lopate C. Equine Stud Farm Medicine and Surgery. St Louis: Saunders/Elsevier, 2003;368 pp. 7. England G. Fertility and Obstetrics in the Horse. Oxford: Blackwell Publishing, 2005;307 pp. 8. Samper JC, Pycock JF, McKinnon AO. Current Therapy in Equine Reproduction. St Louis: Saunders/Elsevier, 2007; 492 pp. 9. Youngquist RS, Threlfall WR. Equine theriogenology. In: Current Therapy in Large Animal Theriogenology. St Louis: Saunders/Elsevier, 2007; pp. 3–218. 10. Ginther OJ. Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains, WI: Equiservices Publishing, 1986; 378 pp. 11. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Fundamentals. Book 1. Cross Plains, WI: Equiservices Publishing, 1995;225 pp. 12. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Horses. Book 2. Cross Plains, WI: Equiservices Publishing, 1995;394 pp. 13. McKinnon AO. Reproductive ultrasonography – The mare. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Philadelphia: Lea & Febiger, 1998; pp. 125–251. 14. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Color-Doppler Ultrasonography. Book 4. Cross Plains, WI: Equiservices Publishing, 2007; 258 pp. 15. Reef VB. Equine Diagnostic Ultrasound. St Louis: Saunders, 1998;424 pp. 16. Wolfgang K. Ultrasonography in the mare. In: Veterinary Reproductive Ultrasonography. Hannover: Schlutersche Verlagsgesellschaft, 2004; pp. 1–80. 17. Ginther OJ. Prolonged luteal activity in mares – a semantic quagmire. Equine Vet J 1990;22:152–6. 18. Irvine CHG, Sutton P, Turner JE, Mennick PE. Changes in plasma progesterone concentrations from Days 17 to 42 of gestation in mares maintaining or losing pregnancy. Equine Vet J 1990;22:104–6.
19. Bergfelt DR, Woods JA, Ginther OJ. Role of the embryonic vesicle and progesterone in embryonic loss in mares. J Reprod Fertil 1992;95:339–47. 20. Bergfelt DR, Ginther OJ. Embryo loss following GnRHinduced ovulation in anovulatory mares. Theriogenology 1992;38:33–43. 21. Gastal EL, Neves AP, Mattos RC, Petrucci BPL, Gastal MO, Ginther OJ. Miniature ponies: 1. Follicular, luteal and endometrial dynamics during the oestrous cycle. Reprod Fert Develop 2008;20:376–85. 22. Purdy SR. Reproduction in miniature donkeys. In: Matthews NS, Taylor TS (eds) Veterinary Care of Donkeys. Ithaca, NY: International Veterinary Information Service (IVIS) ( www.ivis.org), 2005; A2923.0405. 23. Ravindra JP, Rawlings NC, Evans ACO, Adams GP. Ultrasonographic study of ovarian follicular dynamics in the ewe. J Reprod Fertil 1994;101:501–9. 24. Adams GP, Evans ACO, Rawlings NC. Follicular waves and circulating gonadotrophins in 8-month-old prepubertal heifers. J Reprod Fertil 1993;100:27–33. 25. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Cattle. Book 3. Cross Plains, WI: Equiservices Publishing, 1998; 304 pp. 26. Lofstedt RM, Ireland WP. Measuring sphere-like structures using transrectal ultrasonography. Vet Radiol Ultrasound 2000;41:178–80. 27. Pierson RA, Adams GP. Remote assessment of ovarian response and follicular status using visual analysis of ultrasound images. Theriogenology 2003;51:47–57. 28. Singh J, Adams GP, Pierson RA. Promise of new imaging technologies for assessing ovarian function. Anim Reprod Sci 2003;78:371–99. 29. Bollwein H, Mayer R, Weber F, Stolla R. Luteal blood flow during the estrous cycle in mares. Theriogenology 2002;57:2043–51. 30. Carnevale EM, Checura CM, Coutinho da Silva MA, Adams GP, Pierson RA. Early luteogenesis: computerassisted image analysis of the corpus luteum in the first 24 hours postovulation. Theriogenology 2002;58:533–5. 31. Sevinga M, Schukken YH, Hesselink JW, Jonker FH. Relationship between ultrasonic characteristics of the corpus luteum, plasma progesterone concentration and early pregnancy diagnosis in Friesian mares. Theriogenology 1999;52:585–92. 32. Bergfelt DR, Woods JA, Ginther OJ. Role of the embryonic vesicle and progesterone in embryonic loss in mares. J Reprod Fertil 1992;95:339–47. 33. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices Publishing, 1992;642 pp. 34. Bergfelt DR, Pierson RA, Adams GP. Form and function of the corpus luteum. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Philadelphia: Lea & Febiger, 1998; pp. 221–232. 35. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Color-Doppler Ultrasonography. Book 4. Cross Plains, WI: Equiservices Publishing, 2007;258 pp. 36. Bergfelt DR, Adams GP. Ovulation and corpus luteum development. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders/Elsevier, 2007; pp. 1–13.
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Chapter 217
Pregnancy Don R. Bergfelt and Gregg P. Adams
Introduction Early pregnancy diagnosis and prognosis of embryo and fetal health throughout development leading to a vigorous foal at parturition is one of the most valuable and rewarding uses of reproductive ultrasonography in the mare. Advanced technology of B-mode and color-Doppler ultrasonography has provided a virtual window through which scientists, clinicians, and breeding farm managers can detect, monitor, and study morphological and physiological development of the conceptus and its interrelationships with the maternal reproductive system. This chapter is intended to consolidate principal elements from previous texts and review publications1–8 as well as multiple recent publications in scientific journals on the ultrasonographic detection and evaluation of the embryo and fetus into a single contemporary source of reference. The approach used for diagnosing and assessing the status of pregnancy with B-mode and color-Doppler ultrasonography will cover various aspects of pregnancy during the embryo stage (first detection of the embryonic vesicle to 40 days post-ovulation) and fetal stage (≥ 40 days post-ovulation). The term conceptus is used herein to encompass the embryo/fetus, extra-embryonic membranes, yolk, allantoic and amniotic sacs, and umbilical cord during the embryo or fetal stages of development, whereas the term embryonic vesicle is used more specifically in reference to the conceptus primarily during the yolk sac phase. The term embryo proper specifically refers to the early embryo that leads to the fetus apart from other aspects of the conceptus. The day of ovulation is designated day 0 for purposes of reference to the number of days and months of pregnancy and major developmental characteristics during the embryo and fetal stages. Notably, deliberate use of the terms embryo during days 0–40 and fetus, thereafter, should be considered apart from the text since they involve distinctly different develop-
mental aspects and interactive processes involving the conceptus with the uterus that may influence the manner in which an ultrasonographic examination is conducted.
Approach The basis for various routes of examination and techniques using B-mode and color-Doppler ultrasonography to detect and evaluate development of the conceptus during the embryo and fetal stages is in accordance with the fundamental principles of ultrasonography which have been well documented for the horse.1–3,5–8 These earlier sources of reference are invaluable for fully appreciating, comprehending, and interpreting the imaging processes involved in scanning the various aspects of the maternal reproductive tract and conceptus during the embryo and fetal stages. It is expected that information extracted from previous publications and summarized herein will facilitate a safe and thorough examination of the ovaries and uterus so that detecting the presence and morphological and physiological characteristics of the conceptus are precise and evaluation of the status of pregnancy is accurate. Routes of ultrasonographic examination Prior to an ultrasonographic examination, regardless of route, subdued lighting is preferred and the ultrasound machine should be positioned close to the operator and raised to eye level so controls (e.g., gains, brightness, contrast) can be reached readily for adjusting image quality and scrutinizing image detail, differentiating target from non-target tissues or organs, and interpreting the morphological and physiological attributes of target organs. A coupling medium is essential since liquid contact between the transducer face and mucosa or dermis is required for sound waves to be transmitted into the tissues; intervening bowel gas, fecal material, or air will partially
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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or wholly block the transmission of sound. Standard lubricants (e.g., methyl cellulose) used for transrectal palpation are satisfactory as a coupling medium. The latest makes and models of ultrasound machines, imaging capabilities, and incidental provisions vary widely. While some portable machines and transducers are specifically produced for veterinary use, many portable and larger base units are produced for human use but can be adapted readily for application in animals. Transducers are available in various types (e.g., sector, linear-array, convex-array) and frequencies (e.g., 2.5, 3.5, 5.0, 7.5 MHz), and can be used transvaginally, transrectally, and transcutaneously. Deciding on the type of equipment and route of imaging for research or clinical purposes will require professional judgment on practicality and logistics and complete knowledge of experimental designs or case histories.
Transvaginal imaging The transvaginal route is probably the least practical and conducive for diagnosing pregnancy since specialized equipment is involved and extra effort is necessary to maintain sanitary conditions, especially in field situations. Furthermore, additional effort is required to manipulate the uterus via the rectum so the uterine body and horns can be properly and thoroughly imaged when searching for an early embryonic vesicle. In regard to establishing and managing pregnancy, however, the transvaginal route has been used for artificial insemination, embryo transfer9 and the reduction of embryos or fetuses.10, 11
Transrectal imaging The transrectal route is the most conventional approach for examining the reproductive tract for breeding purposes and diagnosis of pregnancy in all types of mature horses, including miniature horses and ponies,1–3,5–8,12 as well as in donkeys.13 Typically, a 5.0- or 7.5-MHz or multiple frequency linear-array transducer is used transrectally until 70–80 days of pregnancy; thereafter, the fetus has grown such that the conceptus has usually descended into the abdominal cavity and out of range to be viewed in its entirety. In this regard, the transabdominal route may be used with a tranducer having more depth of penetration (e.g., 2.5–3.5 MHz) to get a more complete image of the entire fetus. If only portions of the conceptus or fetus need to be imaged then the transrectal route with a 5.0-MHz transducer should still be sufficient. Although the transrectal route of imaging the reproductive organs is not necessarily any easier or more difficult than transrectal palpation, a technical limitation of using this approach is that mares may
be too small to accommodate both hand and transducer into the rectum (e.g., ponies and miniature horses). Alternatively, a rigid or semi-rigid probe extension (e.g., rubber or plastic tubing) may be affixed to the transducer cord similar to that used for scanning small ruminants (e.g., calves and sheep14, 15 ), allowing the transducer to be placed within the rectum and maneuvered externally. Although this unguarded approach is considered safe and effective for scanning the reproductive tract, it is emphasized that the transducer is being manipulated externally in the rectum and extreme caution is recommended to protect against rectal injury, more so than the usual care needed during palpation.
Transcutaneous imaging A transcutaneous or transabdominal route can be used for imaging the conceptus or fetus wholly or partially in large and small horses (e.g., miniature) and ponies using a linear- or convex-array transducer with a frequency of 5.0–7.5 MHz for 70–150 days of pregnancy, or 2.5–3.5 MHz for > 150 days. In lieu of some technical and practical limitations of using the transrectal approach (e.g., small mares, large fetus), the transabdominal route has been used to image larger portions of the fetus to examine biophysical markers of fetal development,16 determine gender,17, 18 estimate day of pregnancy,19–22 and perform other more invasive procedures such as amnio- or allantocentesis, and fetal euthanasia.10, 11 In general, the transabdominal approach involves clipping the hair from the xiphoid to the cranial aspect of the mammary gland and sometimes laterally to the flank fold. After applying a coupling gel, the fetus is often visualized off to one side of midline. A useful landmark is location and identification of the fetal thorax with the beating heart and acoustic shadows from the fetal ribs. Regardless of whether the transrectal or transabdominal route is used, the ability to study sequential, sectional views of the ovaries, uterus, and conceptus in real-time requires a fundamental knowledge of gross23, 24 and ultrasonographic3–8 anatomy and physiology of the maternal reproductive tract and embryo/feto-placental unit. One must be able to direct the transducer to the structures of interest within the pelvic or abdominal cavities (e.g., ovaries, uterus, embryo, fetus, umbilicus) and be able to differentiate and interpret healthy from pathological characteristics. The following sections will summarize key developmental characteristics of the conceptus during the embryo and fetal stages that have been characterized primarily through the use of transrectal ultrasonographic imaging with B-mode3–7 and colorDoppler technology.8
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Pregnancy
Embryo stage Procedures for detection of the conceptus Anatomically, the uterus serves as a guide for the location and orientation of the transducer during a transrectal ultrasonographic examination. The orientation of a linear-array transducer to the uterus is shown in Figure 217.1. When examining the uterus for an early embryonic vesicle, slow, methodical movements covering the entire uterine body and both uterine horns will ensure detection. Rapid movements can result in a false-negative diagnosis since the ultrasound beam is very thin (i.e., 1–2 mm) and can quickly pass over a small vesicle (e.g., approximately 3 mm on day 9). Notably, it does not matter whether the cervix or ovaries are used as reference for beginning an examination as long as the approach covers both uterine horns and, especially, the uterine body. The following discussion describes the process of an examination beginning from the cervix. Beginning anterior to the internal cervical os, the uterine body is viewed longitudinally as shown in Figure 217.1. As the transducer is advanced cranially, it is rotated slightly side to side to view the entire
Ovary
Ovary RA
LA LM
RM LP RP
Right horn
Left horn
BA
Body
BM
BP
Cervix
Finish
Start
Figure 217.1 Diagrammatic representation of a dorsal view of the uterus. The rectangular shaped areas represent the face of the transducer over the uterine body and the left and right uterine horns; arrows represent the direction of movement of the transducer and letters represent major portions of the uterus (B, body; L, left horn; R, right horn) and segments of respective portions (A, anterior; M, middle; P, posterior). Adapted from Ginther.1
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width of the uterine body. At the corpus–cornual junction, the transducer is advanced and rotated slightly to the right to view the right uterine horn in cross-section. Scanning in this plane continues until the image of the anterior portion of the horn disappears and the right ovary comes into view. Routine imaging of the ovaries during early pregnancy evaluation is recommended not only to allow structural (e.g., diameter) and functional (e.g., echogenicity) assessment of the primary corpus luteum (see Chapter 216), but also to determine the presence of multiple corpora lutea. Detection of two corpora lutea would necessitate careful scrutiny of the uterus for potential twin embryonic vesicles (see Chapter 241). Diameters of the two or three largest follicles on each ovary may be recorded as a future reference if the mare is not pregnant and returns to estrus. After inspection of the ovary, the transducer is rotated in the opposite direction until the uterine horn again comes into view and is followed back towards the midline (Figure 217.1). At the corpus–cornual junction, the transducer is rotated laterally to the left to view a cross-section of the opposite uterine horn and ovary. Upon completion of the ultrasonographic examination, the entire uterus has been scanned twice, one or more ovulation sites (i.e., corpus luteum) have been detected, and the status of the primary corpus luteum and degree of follicular development have been assessed. In addition to establishing a systematic approach to an ultrasonographic examination of the uterus and ovaries, it is equally important to establish a consistent record-keeping system. One approach has been to divide the uterus into segments so that the location of the embryonic vesicle or any aberrant intrauterine structure or material (e.g., cyst, fluid) can be readily coded for future reference. As indicated in Figure 217.1, each major portion of the uterus (left horn, right horn, and body) can be subdivided into three segments (anterior, middle, and posterior). A segment is identified by moving the transducer from the structure of interest to each end of the respective segment of the uterine body or horn. Notably, it may be more appropriate to use cranial and caudal terminology instead of anterior and posterior to identify various segments of the uterus; however, the former terms cannot be differentiated readily when using abbreviations in the recording process as indicated in the following example. An example record entry might read: LAEV12 for left horn, anterior segment, embryonic vesicle of 12 mm. These recordings will be useful for updating the status of pregnancy at subsequent examinations. Notably, growth or loss of the embryonic vesicle can be clearly distinguished if earlier recordings indicated that the structure identified as a vesicle
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changed in location and/or size. Occasionally, uterine cysts mimic the expected size and appearance of an early embryonic vesicle.1, 2 Differentiation of a vesicle from a cyst can be made by documenting changes in size, location, or morphology of the structure within the uterine lumen. Early embryonic loss and uterine pathology are discussed in more detail in Chapters 221, 223, and 239. Yolk sac phase (days 11–20) Embryology of the equine conceptus has been thoroughly described in several reference texts3–5,23, 24 and should be studied to fully appreciate the dynamic interactions between the conceptus and uterus during critical periods for diagnosing and managing early pregnancy in mares. In general, the conceptus enters the uterus 6–8 days after ovulation as an early blastocyst with a temporary mucinous glycoprotein capsule, a single embryonic membrane, the ectoderm (trophoblast), and an inner cell mass (developing
Days after ovulation
embryonic disc). By approximately day 12, a second embryonic membrane, the endoderm, encircles the blastocoele transforming the blastocyst into a bilaminar yolk sac (Figure 217.2, days 10 and 12). By approximately day 14, a third embryonic membrane, the mesoderm, has emerged from the embryonic disc between the ectoderm and endoderm, transforming a portion of the two-walled yolk sac into a threewalled (trilaminar) yolk sac (Figure 217.2, days 14 and 16). Portions of ectoderm and mesoderm will eventually pass over the embryonic disc and together give rise to the amniotic sac encompassing the early embryo (Figure 217.2, day 18). From the hindgut of the embryo proper, the allantoic sac emerges into the exocelom and moves ventrally beneath the embryo (Figure 217.2, days 22 and 24). Other portions of the ectoderm and mesoderm (somatopleure) will give rise to the chorion (Figure 217.2, day 24) and, ultimately, the chorio-allantoic placenta (Figure 217.2, day 29) that is involved in endometrial cup formation (chorionic girdle) and implantation.
Sonogram of uterus and conceptus
Embryonic vesicle Blastoceole
Capsule
Inner cell mass
Ectoderm (trophoblast)
Day 10
Yolk sac
Day 12 Embryonic disc
Endoderm
Day 14 Mesoderm
Day 16
Figure 217.2 Diagrammatic representations of the embryology of the conceptus corresponding to respective ultrasonograms on designated days of pregnancy relative to ovulation. Adapted, in part, from Ginther.23 (See color plate 110).
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Pregnancy Days after Sonogram of uterus ovulation and conceptus
Day 18
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Embryonic vesicle Bilamina (ectoderm & endoderm) Trilamina (ectoderm, mesoderm & endoderm)
Amniotic sac
Day 22 Embryo proper
Day 24
Day 26
Allantoic sac
Chorion (SomatoPleure)
Chorionic girdle
Figure 217.2 (Continued)
Ultrasonographic characteristics of the embryonic vesicle, vesicle mobility, fixation, and orientation during the yolk sac phase followed by ascent and descent of the embryo/fetus and formation of the umbilical cord during the allantoic stage are presented as follows.
First detection of embryonic vesicle An embryonic vesicle can be detected as early as day 9; however, for practical purposes, pregnancy diagnosis is typically first done around day 12 when there is a high probability (98–100%) of detecting a larger vesicle and still early enough before fixation (days 15 or 16) to intervene if twin vesicles are detected (see Chapter 241). The early equine conceptus is spherical, fluidfilled, and produces a black (anechoic) image on the ultrasound screen (Figure 217.2, days 10–14). A bright
white (echoic) spot is frequently observed on the dorsal and ventral aspects of the blastocoele or yolk sac image. These image attributes are termed specular reflections and are not an image of the embryonic disc (Figure 217.2, day 12). Specular echoes can be a useful aid during the search for an early embryonic vesicle but should be differentiated from specular echoes that are also characteristic of some intrauterine cysts. In this regard, a vesicle may be distinguished from an intrauterine cyst by mobility and growth of the conceptus (see Chapter 223).
Mobility Intrauterine mobility of the embryonic vesicle is essential for eliciting the first luteal response to pregnancy (i.e., inhibition of the uterine luteolytic mechanism23 ) as reviewed in Chapter 227 and must be considered when scanning the entire uterus for a conceptus in this species. From days 9–11, small
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Sonogram of uterus and conceptus
Embryonic vesicle
Day 29 Allantochorionic placenta
Day 36
Fetus
Day 40
Umbilical chord
Day 45
Figure 217.2 (Continued)
embryonic vesicles (3–6 mm) are located about 60% of the time in the uterine body and, thereafter, with increasing frequency in either of the uterine horns. Maximum mobility (mean, 3.4 mm/min) of the vesicle is attained by about day 12 and is maintained until days 15 or 16, when fixation occurs. While longitudinal endometrial folds facilitate vesicle mobility, myometrial contractions move the vesicle, which can be observed using ultrasonographic imaging at all stages of the estrous cycle and early pregnancy.3 However, the greatest period of uterine contractility during early pregnancy occurs during the maximum mobility phase (days 12–15 or 16). The to-and-fro movements associated with contractions are thought to be influenced by continuous exposure to luteal gland progesterone in combination with follicular estrogen and, perhaps, estrogen and
prostaglandin E2 (PGE2 ) produced by the early conceptus.3, 4 In general, any structural or functional alteration of the early conceptus (e.g., undersized vesicle, decreased estrogen or PGE2 production) or uterus (e.g., large intrauterine cysts, endometrial inflammation) that may hinder vesicle mobility and its capability to block the luteolytic mechanism can result in PGF2a production and jeopardize the continuation of pregnancy, as discussed in Chapter 227.
Fixation Fixation of the early embryonic vesicle has been defined as the cessation of intrauterine mobility that occurs, on average, on day 15 in ponies and day 16 in
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Pregnancy horses at or near the corpus–corneal junction.3, 4 Concomitantly, as the size of the embryonic vesicle increases, the diameter of each uterine horn decreases and uterine tissue turgidity (i.e., tone of pregnancy) increases in response to production of luteal gland progesterone and follicular/conceptal estrogen. The continuation of uterine contractions force the vesicle along the endometrial folds until it reaches the sharp curvature or flexure of the uterine horn near the corpus–cornual junction, where impediment to movement is greatest as a result of increased vesicle size and decreased uterine diameter and increased tone. Thus, by days 15 or 16, the interaction between the early conceptus and uterus results in fixation of the embryonic vesicle at or near the caudal segment of a uterine horn. Notably, apparent cessation of embryonic mobility at a site other than the corpus–cornual junction is considered atypical and could lead to eventual loss of pregnancy (Chapters 221 and 239). Corresponding to the approximate day of fixation, a transient period of enhanced endometrial echo-heterogenicity has been described.3 The mild estrus-like appearance in the vicinity of where the embryonic vesicle is fixed is thought to be due, in part, to exposure of the uterus to estrogen emanating from the conceptus. It is emphasized that a local and transient increase in uterine echo-heterogenicity encompassing the time of fixation should not be confused with impending loss of the embryonic vesicle and subsequent return to estrus. Other ultrasonographic and palpable characteristics of the ovaries (e.g., decreased size or increased echogenicity of the corpus luteum), uterus (e.g., decreased turgidity or tone), and conceptus (e.g., mobility from site of fixation at or near the corpus–corneal junction) associated with loss of pregnancy should be considered (see Chapters 221 and 239). Color-Doppler ultrasonic imaging has been used recently to study endometrial vascular perfusion during mobility and fixation of the embryonic vesicle.8 Before fixation, subjective and objective (i.e., image analysis software) scoring of color-flow images indicated blood flow was greater in the uterine horn that temporarily contained the vesicle than in the opposite horn. After fixation, blood flow was greater in the fixed or gravid uterine horn. From a practical perspective, the results from these studies did not support the establishment of a vascular indicator to predict the future horn of fixation since endometrial vascular perfusion scores between uterine horns were not statistically different before fixation. Nevertheless, future studies are needed to determine whether the degree of endometrial perfusion at the site of embryo fixation may be used to assess health or impending loss of the early conceptus.
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Orientation Orientation refers to the rotation of the embryonic vesicle after fixation so that the three-walled portion of the vesicle (embryonic pole) assumes a ventral position, opposite the mesometrial attachment at the dorsal position. Reduced structural rigidity of the conceptus (dissipation of capsule), increased uterine turgidity (tone), massaging action of uterine contractions, and asymmetric encroachment of the dorsal uterine wall are integrated conceptal–maternal processes that force the thicker three-walled portion of the embryonic vesicle and embryonic pole ventrally leaving the thinner two-walled portion of the vesicle adjacent to the mesometrial area. Notably, a distinct guitar pick-shaped vesicle is often observed during an ultrasonographic examination around day 18, as shown in Figure 217.2. Orientation of the embryonic vesicle is thought to be necessary to accommodate species-specific development of the embryo/ fetus and formation of the umbilical cord and endometrial cups.
First detection of embryo proper Considering proper orientation of the embryonic vesicle, detection of the embryo proper is typically first observed as a small echoic structure in the ventral aspect of the yolk sac on approximately day 20 (Figure 217.2, day 22). Correspondingly, depending on imaging capability of the ultrasound machine and transducer, cardiac activity is first observed as a pulsation of the echoic image or as a color-Doppler signal (Figure 217.3, day 19). Although relatively rare, the first detection of the embryo proper may be observed in the dorsal aspect of the yolk sac, as documented in a recent study involving retrospective analyses of two different sets of data,25 as follows. Disorientation of the embryo proper is a consequence of disorientation of the embryonic vesicle such that the three-walled portion (embryonic pole) is oriented dorsally near the mesometrial area, as depicted in Figure 217.3, day 19.8, 25 In the first of two datasets, disorientation based on detection of the embryo proper in the dorsal aspect of the vesicle was observed in four out of 30 mares (13%) by day 19 and was associated with a flaccid uterus and a lack of encroachment of the dorsal endometrium in three mares. In the remaining mare, uterine tone and encroachment were apparently normal. Data were not available after day 19 to know the outcome of these pregnancies. In a second set of data, disorientation of the embryonic vesicle was detected in two mares with apparently normal uterine tone and encroachment. These pregnancies were subsequently monitored and compared with normally oriented vesicles,
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Orientation vs Dysorientation of Embryo Proper Ventral position of embryonic pole
Dorsal position of embryonic pole Ma
Ma
Ep Ep
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Ma
Ma
Ys
As
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Figure 217.3 A series of images taken with B-mode and color-Doppler ultrasonography showing orientation (left) and disorientation (right) of the embryo proper (Ep) from day 19 to day 45. Note the expanding allantoic sac (As), receding yolk sac (Ys), and formation of umbilical cord (Uc), such that the placental base of the cord lies dorsally in juxtaposition with the mesometrial attachment (Ma) despite the initial malposition of the embryonic vesicle. Adapted from Ginther and Silva.25 (See color plate 111).
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Pregnancy as depicted in a series of ultrasonograms in Figure 217.3. As the allantoic sac expanded and the yolk sac receded, the visibly echoic membranes of the future umbilical cord on either side of the disoriented embryo proper advanced simultaneously but differentially toward the mesometrial area (Figure 217.3, days 19 and 28). Distinctively, the umbilical cord membranes were malpositioned in a more vertical plane rather than a horizontal plane, which is normally observed (Figure 217.3, days 19 and 29). It appeared that, despite disorientation of the early conceptus, formation of the umbilical cord attachment in close proximity to the mesometrial attachment was conserved and that the conceptus was repositioned (Figure 217.3, day 44) in a manner that is normally observed (Figure 217.3, day 45). Hence, initial disorientation of the embryo proper upon first detection may not be detrimental to the conceptus provided uterine tone and encroachment are adequate for the respective stage of pregnancy. It has been postulated 8, 25 that the apparent conservation of umbilical cord formation dorsally in the vicinity of the mesometrial attachment is an adaptation to ensure that fetal placental blood vessels are in juxtaposition to maternal vessels during early pregnancy, and, perhaps, a mechanism to minimize or avoid entanglement and impingement by a large fetus later in pregnancy. Notably, disorientation of the early conceptus that is associated with a flaccid uterus may be indicative of impending embryo loss (see Chapters 221 and 239). While color-Doppler ultrasonic imaging may be used to further substantiate cardiac activity and viability of the embryo proper, recent investigation suggests that it may also be useful for predicting the future site or position of the embryo proper within the vesicle before the early embryo is first detected ultrasonographically.8 There was no statistically significant difference between the position of the early embryo as predicted by color-Doppler signals impinging upon the wall of the embryonic vesicle and the position observed by ultrasonography approximately 2.5 days later. Hence, the combination of B-mode and color-Doppler ultrasonography provides a powerful diagnostic and prognostic tool to predict or detect and verify the position and viability of the early embryo during a critical period of pregnancy.
Allantoic sac phase (days 21–40) Detection of the embryo proper with cardiac activity by approximately day 20 may be considered the beginning of the transition from the yolk sac phase to allantoic sac phase (Figure 217.2, day 22). As the allantoic sac expands and the yolk sac recedes, there is the
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distinct ultrasonographic characteristic of ascent of the embryo–amniotic unit (Figure 217.2, days 22–40). Thereafter, as the umbilical cord is elongated, there is descent of the fetal–amniotic unit (Figure 217.2, day 45). The unique characteristics of ascent and descent of the embryo/fetal–amniotic unit can be of value for estimating the day of gestation as well as determining the health of the pregnancy.3, 5
Embryo–amniotic ascent Development of the amniotic sac is complete and the allantoic sac begins to emerge from the hindgut of the embryo proper at approximately day 20 (Figure 217.2, day 22). By day 24, the allantoic sac has become a prominent anechoic area beneath the embryo proper, lifting the embryo–amniotic unit from the ventral floor of the vesicle as it expands and the yolk sac recedes (Figure 217.2, day 24). Opposing walls of the yolk sac membrane dorsally and the allantoic sac membrane ventrally appear as echoic lines on either side of the embryo proper that tend to maintain a horizontal position, except in situations involving disorienatation of the early conceptus8 and twinning,3, 5 as the early embryo is lifted toward the dorsal aspect of the vesicle toward the mesometrial area. Over days 24–33, the allantoic sac continues to expand as the yolk sac recedes (Figure 217.2, days 24–36) until the allantoic sac membrane engulfs the vestige of the yolk sac by day 40 (Figure 217.2). The convergence of the opposing walls of the allantois and their fusion around the urachus, blood vessels, and yolk sac results in the formation of the umbilical cord (Figure 217.2, day 45). Hence, the placental base of the umbilical cord is located dorsally in juxtaposition to the mesometrial area, and is encircled wholly or partially by endometrial cups that have colonized the dorsal endometrial wall.4, 23 Ultrasonographically, the late embryo/early fetus appears to uniquely dangle at the end of the umbilical cord in dorsal recumbency (Figure 217.2, day 45).
Embryo/fetal–amniotic descent Day 40 marks the transition from the embryo stage to the fetal stage; a distinctly different developmental period involving integrated movements of the fetus with periodic shifts in allantoic fluid as discussed in detail in a later section. At the beginning of the fetal stage, the umbilical cord elongates and the fetal–amniotic unit descends ventrally. At approximately day 45, descent is about 50% complete (Figure 217.2) and, by day 50, the fetal–amniotic unit has descended to the floor of the allantoic sac. These and other key developmental characteristics (e.g., size of
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the embryonic vesicle, crown–rump length of the embryo/fetus) have been compiled during the embryo and early fetal stages as an aid for estimating the day of pregnancy.1, 3, 5 Embryonic characteristics to estimate day of pregnancy Comparison of serial ultrasonographic images of the embryonic vesicle between horse and pony mares revealed no significant difference in size from day 9 to day 45.1, 3 Hence, the results have been combined to generate a growth profile of the embryonic vesicle as represented by three different regression lines illustrated in Figure 217.4: (i) linear from days 11–16 (growth rate, 3.4 mm/day), (ii) S-shaped from days 17–27, and (iii) linear from days 28–45 (growth rate, 1.7 mm/day). The apparent plateau in growth associated with the S-shaped portion of the growth profile is an illusion resulting from longitudinal expansion of the chorio-allantoic sac along the umbilical cord horn which is due, in part, to reduced rigidity of the embryonic vesicle and continuing increase in uterine tone associated with pregnancy.2, 23
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Confidence intervals were generated for the linear portions of the growth profile (Figure 217.4) so that day of pregnancy could be estimated with 95% accuracy for vesicles 6–23 mm (± 1.5 days) and 28–56 mm (± 4 days). The vertical diameter (height) of the embryonic vesicle does not change sufficiently during the S-shaped portion of the growth profile to accurately estimate day of pregnancy, but major developmental characteristics of the conceptus can aid in estimating the day of pregnancy and assessing viability during this period (Table 217.1). In regard to the latter, it may be more convenient and, perhaps, more accurate to estimate age morphologically by the changes in position of the embryo/fetus within the vesicle.2 Nevertheless, after day 40, maximal height of the embryonic vesicle is not a reliable indicator for estimating the day of pregnancy because of periodic shifts in allantoic fluid that begin to occur, as discussed in the next section. Alternatively, fetal body length (i.e., crown–rump) or other developmental aspects of the fetus (e.g., eye orbit and trunk) may be used to estimate the day of pregnancy,1, 3, 7,19–22 as presented in the final section of this chapter.
Fetal stage
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Figure 217.4 Graphical representation of a regression analysis (solid dots with regression lines) for estimating the day of pregnancy based on the observed size of the conceptus primarily during the embryo stage in horses and ponies. Note the Sshaped portion of the regression line between days 16 and 28 is not adequate for estimating day of pregnancy, but other key developmental characteristics can be used, as indicated in Table 217.1. Adapted from Ginther.1
Ultrasonographic imaging of the fetus and the fetoplacental unit from day 40 and beyond has not been studied as thoroughly as the embryo stage. Nonetheless, recent studies have characterized developmental aspects of the fetus, uterus, and associated membranes using transabdominal and transrectal Bmode16–18,21, 22 and color-Doppler ultrasonography.8 Characterization of uterine contractions and allantoic fluid shifts associated with fetal activity and mobility have led to a better understanding of how fetal–maternal interactions play a role in preparing the fetus for proper presentation prior to parturition during the latter months of pregnancy.4 In addition to B-mode and color-Doppler ultrasonographic examination of fetal cardiac activity and vascular blood flow of various fetal and maternal organs,8 observation of early motor activity may be an aid in assessing and monitoring fetal and maternal health during mid- to late gestation.16 To fully appreciate the events of fetal development and fetal–maternal interactions, the reader is directed to the original review article,4 which provides detailed documentation with ultrasonograms and illustrations of uterine constrictions, fetal activity, and rotational changes throughout the fetal stage and parturition. The following sections are a summary of fetal–maternal interactions as characterized by transrectal B-mode ultrasonography prior to parturition.
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Table 217.1 Characteristic developmental landmarks of the early equine conceptus (combined horses and ponies) that can be used in combination with the size of the embryonic vesicle illustrated in Figure 217.4 to estimate the stage of pregnancy when ovulation or breeding dates are not known. Adapted from Ginther.1 Estimated days of pregnancy
Developmental landmarks
9–11 12–14 15–16 17–19 20–22 23–24 25–27 28–30 31–33 34–36 37–45 46–50
Detection of embryonic vesicle as spherical, fluid-filled blastocoele Mobility of yolk sac vesicle between uterine horns Mobility cessation and fixation of yolk sac vesicle at posterior segment of a uterine horn Orientation and loss of sphericity of yolk sac Detection of embryo proper ventrally within yolk sac plus cardiac activity Embryo–amniotic unit ascends dorsally; yolk sac recedes/allantoic sac expands Yolk sac 75%; allantoic sac 25% Yolk sac 50%; allantoic sac 50% Yolk sac 25%; allantoic sac 75% Ascent complete; umbilical cord formation dorsally Umbilical cord elongates; fetal–amniotic unit descends ventrally 50% Descent complete; fetal–amniotic unit positioned ventrally within allantoic sac
Uterine constrictions and allantoic fluid shifts The fluid-filled chorio-allantoic sac gradually expands in association with placentation to fill the lumen of the uterus in a manner that involves alternating myometrial contractions and relaxation of segments of the uterus. The allantoic sac appears to first fill the uterine body, reaching the internal cervical os by approximately day 50, second, filling the umbilical cord horn and reaching the utero-tubal junction by day 60, and, third, the entire uterus is filled by day 65. Even after the allantoic sac has expanded to fill the entire uterus, transient contractions continue to constrict whole or partial segments of the uterus. Hence, at any given moment, a segment of the uterus may be constricted and temporarily inhibit fetal mobility at the site of closure, or become dilated allowing the fetal–amniotic unit to drift and move freely about the fluid-filled uterus or portions of it (Figure 217.5, day 100). The frequency of partial or complete constrictions of both the umbilical cord and non-cord horns begins to increase by approximately month 3 and reaches maximum during months 5–7 (Figure 217.6).
Activity and mobility Fetal activity and mobility are not mutually exclusive, but constitute a dynamic relationship resulting in movement of the fetus within the amniotic and allantoic sacs. Fetal activity refers to in-place movements of the head, limbs, and torso, whereas fetal mobility refers to whole-body changes in direction (presentation), recumbency, and locations within the uterus. In-place activity of the head is first observed
by approximately day 40, limb activity by day 45, and whole-body activity by day 50. Intensity and duration of fetal activity involving both in-place and whole body movements (i.e., presentation changes) increases and reaches maximum over months 2–4 (Figure 217.6). Concurrent with maximal changes in fetal presentation during months 2–4, there is an increase in length of the umbilical cord and ratio of allantoic fluid to fetal mass. The early fetus becomes highly mobile, changing location (between horns and body), recumbency (lateral, dorsal, sternal), and presentation (cranial and caudal aspects of fetus directed toward cervix) multiple times in concert with uterine constrictions and periodic allantoic fluid shifts. Over months 4–9 there is a decrease in the ratio of allantoic fluid to fetal mass and an increase in closure of the uterine horns such that fetal presentation changes decrease (Figure 217.6). Notably, however, in-place activity of the fetus is still prevalent.
Uterine horn closures and presentation Closure of the uterine horns increases after month 3, and, by month 6, the non-cord and cord horns, respectively, were completely closed such that allantoic fluid and the fetus were retained in the uterine body (Figure 217.5, day 190). During months 2–5, the fetus was as likely to be in a cranial presentation as a caudal presentation at any given time (Figure 217.6). After month 4, the percentage of presentation changes decreased, and by months 9–11 (after closure of the uterine horns), most fetal presentations were cranial (Figure 217.5, day 240, and Figure 217.6). For fetuses that may be in caudal presentation after uterine horn closures, positioning to a cranial presentation may
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80 60
Figure 217.5 Diagrammatic representation of the changes in fetal and uterine relationships. (a) Mobile fetus, periodic uterine constrictions, and shifts in allantoic fluid. (b) Fetus finds cranial presentation while confined to the uterine body by closed horns. (c) Hind limbs enclosed in uterine horn; fetus locked in a cranial presentation. Adapted from Ginther.4
be accomplished within the large fluid-filled uterine body (Figure 217.5, day 190). It has been hypothesized that the fetus recognizes a maternal cue (e.g., incline of ventral uterine body wall toward cervix) that prompts the fetus to present itself properly for parturition.4
Percentage of examinations or horn depth
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40 20 0 100
Depth of cord horn entered by hind limbs 80 60
Fetus entirely in uterine body
40
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20
Beginning around month 7, while the fetus is in a cranial position and in dorsal recumbency (Figure
Limbs permanently in cord horn (mean)
0 2
Hind limbs encased in cord horn
Almost all fetuses in cranial presentation
3
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5 6 7 8 Month of pregnancy
9
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Figure 217.6 Graphical summary of various aspects the fetus, uterus, and fetus–uterus interactions throughout pregnancy. Adapted from Ginther.4
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Figure 217.7 An ultrasonogram of the eye of an equine fetus measured at its maximum dimension from medial to lateral canthus using the electronic calipers. 1, Fetal skull; 2, bony orbit; 3, vitreous fluid; and 4, lens. Adapted from Turner et al.21, 22 with permission from Elsevier.
Table 217.2 Fetal eye measurements in light horse mares (400–550 kg) that can be used to estimate stage of pregnancy when ovulation or breeding dates are not known. Adapted from Turner et al.22 Days of pregnancy Confidence intervals Fetal eye (mm)
Estimate of mean
75%
Thoroughbreds, Standardbreds19 10 86 65–107 11 92 72–112 12 98 78–117 13 104 85–123 14 110 91–129 15 116 98–134 16 122 104–141 17 129 111–147 18 135 117–153 19 142 124–160 20 149 131–166 21 155 137–173 22 162 144–180 23 169 151–187 24 176 158–194 25 183 165–201 26 190 172–208 27 198 179–216 28 205 187–223 29 212 194–231 30 220 201–239 Thoroughbreds, Standardbreds, Quarter Horses, Arabians, and Warmbloods22 30 284 247–321 31 291 254–328 32 298 261–335 33 303 266–340 34 308 271–345 35 312 275–349 36 315 278–353 37 318 280–356
95%
50–123 57–127 64–131 71–136 78–142 85–148 91–154 98–160 104–166 111–173 118–179 124–186 131–193 138–200 145–207 152–214 159–221 166–229 173–237 180–245 187–253 220–348 227–355 234–361 239–367 244–372 248–376 251–380 252–384
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estimating the fetal stage of pregnancy.20 In mares with known days of pregnancy, the sums of the width plus length of the fetal eye as determined using transrectal ultrasonography with a 5-MHz transducer were highly correlated (r = 0.92) with gestational age. Regression analysis with 95% confidence intervals illustrating the relationship between the fetal eye and pregnancy from days 60 to 340 have been reviewed.5, 24 At approximately the same time, a comparable study by other investigators19 was done to examine the feasibility of using the fetal eye as well as other fetal characteristics (trunk, cranium, ribs plus intercostal space, and stomach) for estimating the day of pregnancy. In concurrence with the previous study,20 the fetal eye as well as the fetal trunk exhibited the highest correlation (r = 0.95) with the day of pregnancy as reviewed.2, 5 In more recent studies,21, 22 transrectal ultrasonography with either a 5.0- or 7.5-MHz transducer was used to measure the fetal eye during known days of pregnancy. The fetal eye was detected in > 90% of the examinations, and an image of the vitreous-filled orbit, including the lens, was frozen at its longest dimension (ostensibly from medial to lateral canthus),
measured, and recorded, as depicted in Figure 217.7. Three measurements of different images of the fetal eye were taken during each examination, and the mean maximum dimension was used to estimate the mean day of pregnancy in horse mares (Table 217.2). The gross weights of horse and pony fetuses diverge after approximately day 100.23 In recent studies,21, 22 involving ultrasonographic measurement of the fetal eye in horse and pony mares from mid- to late gestation, the fetal eye was larger in horses than in ponies beyond day 100. Hence, the need for separate horse and pony values for estimating day of pregnancy beyond day 100 was supported. Under simulated field conditions, a portable ultrasound machine with a 7.5-MHz linear array transrectal transducer was used to measure the fetal eye in a semiferal herd of ponies.21 The approach used to measure the fetal eye was similar to that previously described in horses22 (Figure 217.7). Although the fetal eye was detected in > 90% of the examinations, failure to detect the eye was most frequent during months 3 (35% of examinations) and 4 (19%), intermediate during months 5 (12%), 6 (12%), and 7 (13%), and least frequent during months 8 (2.5%), 9 (0%), 10 (8%), and 11 (0%). Notably, the period that the
Table 217.3 Fetal eye measurements in pony mares (100–250 kg) that can be used to estimate stage of pregnancy when ovulation or breeding dates are not known. Adapted from Turner et al.21 Days of pregnancy Confidence intervals Fetal eye (mm)
Estimate of mean
75%
95%
Shetland-type ponies21, 22 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35
209 203 197 190 183 175 167 158 150 140 131 121 110 99 88 77 65 52 39 26 13
184–251 177–245 170–238 153–230 155–223 147–214 138–206 129–197 120–187 110–177 100–167 89–156 78–145 66–133 54–121 42–109 29–96 16–83 2–69 0–55 0–41
160–275 153–269 146–262 140–254 131–238 123–238 115–230 106–220 96–211 86–201 76–191 65–180 54–169 43–157 31–145 18–133 5–120 0–107 0–93 0–79 0–64
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Pregnancy fetal eye was most difficult to detect during pregnancy corresponds with the period of maximal changes in presentation (Figure 217.6, months 2 and 4). Since the day of ovulation and breeding were not known, the days before parturition were used in a linear regression analysis to relate fetal growth and stage of pregnancy21, 22 (Table 217.3). It should be recognized that the means and ranges for estimating the day of pregnancy according to fetal eye measurements in horse and pony mares in Tables 217.2 and 217.3, respectively, are meant to serve as a guide for purposes of monitoring pregnancy, along with other physiological and morphological indicators of the fetus and dam.
Conclusion As ultrasound technology continues to advance and is diligently applied by scientists, clinicians, and breeding farm managers to diagnose, monitor, and study morphological and physiological development of the conceptus and its interrelationships with the maternal reproductive system, the nature of pregnancy will be further revealed. Historical as well as contemporary knowledge of the journey that the early embryo takes to become a late-term fetus provides a foundation for testing hypotheses associated with the many marvels of pregnancy and improving reproductive management practices that ultimately lead to a safe and successful delivery of a healthy foal and postpartum recovery of the dam.
References 1. Ginther OJ. Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains, WI: Equiservices Publishing, 1986. 2. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Fundamentals. Book 1. Cross Plains, WI: Equiservices Publishing, 1995. 3. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Horses. Book 2. Cross Plains, WI: Equiservices Publishing, 1995. 4. Ginther OJ. Equine pregnancy: physical interactions between the uterus and conceptus. In: Proceedings of the 44th Annual Convention of the American Association of Equine Practitioners, 1998, pp. 73–104. 5. McKinnon AO. Reproductive ultrasonography: the mare. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Philadelphia: Lea & Febiger, 1998; pp. 125–251. 6. Reef VB. Equine Diagnostic Ultrasound. St. Louis: Saunders, 1998. 7. Wolfgang K. Ultrasonography in the mare. In: Veterinary Reproductive Ultrasonography. Hannover: Schlutersche Verlagsgesellschaft, 2004. 8. Ginther OJ. Ultrasonic Imaging and Animal Reproduction: Color-Doppler Ultrasonography. Book 4. Cross Plains, WI: Equiservices Publishing, 2007.
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9. Silva LA, Gastal EL, Gastal MO, Jacob JCF, Reis CP, Ginther OJ. A new alternative for embryo transfer and artificial insemination in mares: ultrasound-guided intrauterine injection. J Equine Vet Sci 2004;24:324–32. 10. Rantanen NW, Kincaid B. Ultrasound guided fetal cardiac puncture: a method of twin reduction in the mare. In: Proceedings of the 34th Annual Convention of the American Association of Equine Practitioners, 1988, pp. 173–9. 11. Ragon AC. Management of twin pregnancy. In: Youngquist RS, Threlfall WR (eds) Current Therapy in Large Animal Theriogenology, 2nd edn. St. Louis: Saunders/Elsevier, 2007; pp. 114–17. 12. Gastal EL, Neves AP, Mattos RC, Petrucci BPL, Gastal MO, Ginther OJ. Miniature ponies. 1. Follicular, luteal and endometrial dynamics during the oestrous cycle. Reprod Fertil Dev 2008;20:376–85. 13. Purdy SR. Reproduction in miniature donkeys. In: Matthews NS, Taylor TS (eds) Veterinary Care of Donkeys. Ithaca, NY: International Veterinary Information Service (IVIS), 2005. 14. Adams GP, Evans ACO, Rawlings NC. Follicular waves and circulating gonadotrophins in 8-month-old prepubertal heifers. J Reprod Fertil 1993;100:27–33. 15. Ravindra JP, Rawlings NC, Evans ACO, Adams GP. Ultrasonographic study of ovarian follicular dynamics in the ewe. J Reprod Fertil 1994;101:501–9. 16. Reef VB. Late term pregnancy monitoring. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders/Elsevier, 2007; pp. 410–16. 17. Renaudin CD, Gillis CL, Tarantal AF. Transabdominal combined with transrectal ultrasonophic determination of equine fetal gender during midgestation. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997, pp. 252–5. 18. Holder RD. Fetal sex determination. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis: Saunders/Elsevier, 2007; pp. 343–56. 19. Kahn W, Leidl W. Ultrasonic biometry of horse fetuses in utero and sonographic representation of their organs [in German]. DTW Dtsch Tierarztl Wochenschr 1987;94: 509–15. 20. McKinnon AO, Squires EL, Pickett BW. Equine Reproductive Ultrasonography. Animal Reproduction Laboratory Bulletin No. 4. Fort Collins: Colorado State University, 1988. 21. Turner RM, McDonnell SM, Feit EM, Grogan EH, Foglia R. Real-time ultrasound measure of the fetal eye (vitreous body) for prediction of parturition date in small ponies. Theriogenology 2006;66:331–7. 22. Turner RM, McDonnell SM, Feit EM, Grogan EH, Foglia R. How to determine gestational age of the equine pregnancy in the field using transrectal ultrasonographic measurement of the fetal eye. In: Proceedings of the 52nd Annual Convention of the American Association of Equine Practitioners, 2006, pp. 250–5. 23. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices Publishing, 1992. 24. McKinnon AO, Voss JL. Equine Reproduction. Philadelphia: Lea & Febiger, 1993. 25. Ginther OJ, Silva LA. Incidence and nature of disorientation of the embryo proper and spontaneous correction in mares. J Equine Vet Sci 2006;26:249–56.
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Chapter 218
Equine Fetal Sex Determination Between 55 and 150 Days Richard D. Holder
In order to be successful, a considerable amount of time needs to be committed to learning how to perform a sex determination on an equine fetus. The veterinarian needs to decide if the area in which he is practicing warrants taking the time to become adept at performing sex determinations. If the area warrants it then it will be financially and professionally rewarding to those that persevere. It may take as many as 400 or 500 attempts at determinations before one is competent and confident. There are primarily three different stages of gestation at which fetal sex determinations can be made. The first stage is between 55 and 90 days. This stage involves finding the genital tubercle, a small, 2–3 mm hyperechoic bilobulated structure (Figure 218.6b and c) appearing like an equal sign. It is the precursor to the penis in the male and the clitoris in the female. The genital tubercle must be identified and located in relation to other fetal anatomical structures. The tubercle can be seen as early as 52 or 53 days of gestation and is located between the hind legs on the ventral midline. As the fetus ages, the tubercle appears to move toward the umbilical cord in the male and toward the tail in the female. Around day 55 from conception, the tubercle has usually migrated enough to make a determination. At this time the fetus is easily accessible by rectal palpation and an accurate determination (99+% accuracy) can be made a very high percentage of time on one examination. This examination takes 10 seconds to 2–3 minutes, depending on the experience of the examiner. The fetus continues to be easily accessible until around 70 days.
At this time, in some mares, the posterior part of the fetus can be difficult to access. Usually, it can be done until about day 80. At 80 days, the uterus begins to be pulled over the pelvic rim by the fluid of the pregnancy. The fetus is small and descends to the most ventral part of the uterus in the allantoic fluid. It remains difficult to reach until about day 90. At 90 days the fetus comes back into view as the uterus seems to elevate dorsally in the abdominal cavity and is easily accessible. The genital tubercle is less prominent now but the external genitalia (penis, prepuce, glans penis, and the gonads in the male and the mammary, teats, clitoris, and the gonads in the female) are just beginning to be evident. The second stage begins at 90 days and goes to about 150 days. Determinations are somewhat more subjective because of the need to evaluate the tissue densities between prepuce and muscle, or mammary and muscle, etc. Other fetal tissues frequently obscure the genitalia. It is usually not as clear-cut as the genital tubercle evaluations, and may take anywhere from 30 seconds to 5-10 minutes. A sex determination can be made 90% of the time at this stage, with 98+% accuracy, on one examination. After 150 days the fetus assumes an anterior presentation, making the posterior pelvic area of the fetus very difficult to access. In the author’s experience, determinations made transrectally between 150 and 200 days are fruitful only about 20% of the time. The third stage is from 150 days until the end of pregnancy. Sex determination must be done
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Equine Fetal Sex Determination Between 55 and 150 Days transabdominally to observe for external genitalia. This technique requires much more time, crawling under the mare’s abdomen, clipping or preparing the abdomen with alcohol, and a more powerful ultrasound machine. In the author’s hands, with transabdominal scanning after 160 days, a determination can be made only 0–20% of the time. Since this technique takes longer, is more dangerous, and yields a low percentage of diagnosis, the author strongly recommends that fetal sex determination is carried out at the earlier stages. When horse breeders became aware that the sex determination technique was extremely accurate, they readily incorporated the new procedure into their regimen as another tool for management. Figure 218.1 is a graph showing the increase in fetal sex determination requests from 1992 (when the service was initiated) until January 2007. Obviously the horse public has felt the need to know the information learned from a sex determination. If a practitioner is the only one in his or her area able to accurately perform a sex determination, the procedure can develop into a worthwhile profit center. Every year clients find new reasons for wanting to know the sex of the fetus. The information provides a valuable management tool to be used by the owner or the manager. Depending on the sire or the dam, the fetal gender may affect the value of the fetus. If the value of the fetus changes, then the value of the mare changes. This could affect values for appraisals, insurance coverage, sales reserves, collateral limits for loans, and so on. Some other considerations include cash flow predictions. For example, some individuals sell all the fillies and race all the colts, or vice versa. Knowledge of the gender would inform the owner how many he would be selling (income) and how many he would
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be paying training bills on (expenses), a year earlier. Possibly purchases would be made to alleviate a shortfall in one sex or the other in the foaling crop. Knowing the gender enables the owner to take advantage of restricted racing programs. For instance there is no reason to send a mare carrying a filly to New York to foal if the aim is to race New York bred colts. Perhaps an owner wants a particular sex by a certain stallion out of a specific mare. If the mating is successful, he can breed his mare to a different stallion the next year. If his mare is carrying the opposite sex of what he wanted, he could rebook his mare to the same stallion. Popular stallions fill their books early and one cannot wait to see what the sex is after birth and then choose the stallion. One client’s mare always had a dystocia when the mare carried a colt, but had no problems when she carried a filly. A decision was made to foal the mare at a clinic when it was determined that she was carrying a colt. The most common use of fetal sex determination is in keep-orsell decisions for the mare or her offspring depending on what she is carrying. Perhaps a mare has an offspring on the racetrack that is performing well. The owner may have an offer to purchase the mare or her offspring at her side. Suppose the mare had a weanling filly, and the owner was offered a lot of money for her but did not want to sell because he wanted to keep a filly out of the mare. The mare’s fetus could be sexed and, if she was carrying another filly, the weanling filly could be sold.
Materials needed for fetal sex determination
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Quality ultrasound machine with a 5 MHz linear array rectal transducer (Aloka 500 SSD or equivalent). Settings: If the overall gain, near gain, or far gain is set too high there is less contrast between
INCIDENCE OF INCREASING DEMAND 2,500 2,000
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Figure 218.1 Graph showing increase in requests for fetal sex determination between 1992 (when the service was initiated) and January 2007.
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the tubercle and its background, making it less distinct. Rectal palpation essentials: r Lube, sleeves; r Restraint: stocks, twitch, lip chain, etc.; r Tranquilizers: Depending on the situation, tranquilization is acceptable; however, this may cause the uterus to relax and descend further into the abdominal cavity away from the examiner. This may make the fetus more difficult to reach. The author uses 200 mg of xylazine and 10 mg of butorphanol tartrate mixed and given intravenously; r Propantheline bromide (30 mg i.v.) or buscopan (40 mg i.v.) may be used to prevent rectal straining. Ultrasound stand on wheels – for close, eye-level viewing. Subdued lighting – for good screen visualization. Hat – to remove surrounding glare. Fly spray (if needed) – to keep mare movement to a minimum. Videos or printers are helpful to verify the diagnosis and for record keeping.
Fetal sex not determined on first attempt: analysis 1811 mares were examined between 55 and 170 days’ gestation. In 61 mares (3.368%), no determination was made on first examination. In 31 of the 61 mares, determination was not possible because:
r r r r
In the remaining 30 mares (1.656%) no confident diagnosis was possible, or diagnosis was too difficult. 1546 mares were examined at less than 90 days in the United States. In Kentucky most mares were examined at 60–70 days. Of the 1546 mares:
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Fetal development
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55–60 days: The fetus is very accessible but it is small. The genital tubercle is difficult to see, and the tubercle may or may not be fully migrated. 60–70 days: This is the ideal time for the examination. The tubercle is distinct, fully migrated, and the fetus is easily accessible for viewing. 70–80 days: The fetus is slightly more difficult to reach and the tubercle is slightly less distinct. 80–90 days: It is more difficult to view the fetus, the tubercle is less distinct, the genitalia development is just beginning, and the fetus is frequently out of reach. However, maiden mares often can be examined at this time. 90–110 days: The fetus is usually accessible and the genitalia are gradually becoming more evident. 110–120 days: This is the ideal time for examination at the mid-gestation stage. The fetus is very accessible and the genitalia are very well developed. 120–140 days: The genitalia are well developed, but the posterior of the fetus may be difficult to access. 140–150 days: The fetus is large, with the posterior difficult to access. 150+ days: The fetus is large with an anterior presentation and the posterior is usually out of reach
18 mares were not pregnant 5 dead fetuses 4 mares were too fractious 4 mares were < 55 days and the tubercle had not migrated.
2 of 1298 – between 55 and 70 days had no determination; 10 of 239 – between 70 and 89 days had no determination; 4 of 9 – between 80 and 90 days had no determination. Clearly, this is the most difficult time for fetal sex determination.
The remaining 265 mares were examined between 55–170 days out of the United States. Outside of Kentucky, most mares were examined at the end of the breeding season at more random times. A total of 14 of 265 mares had no determination on the first examination, as follows:
r r
6 of 107 – less than 90 days or 5.6% had no determination; 8 of 158 – mares greater than 90 days or 5.06% had no determination.
Reason for difficulty at 80–90 days At about 80 days of gestation the fluid of the pregnancy pulls the uterus over the rim of the pelvis. The fetus is small and falls to the most ventral part of the uterus in the allantoic fluid (Figure 218.2a). However, the uterus does not continue to move more ventrally into the abdomen after 85 days. It actually elevates in the abdominal cavity as it becomes larger and the fetus is usually visible again at about 90 to 95 days of gestation. As the fetus grows it often
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Figure 218.2 Position of pregnant uterus in abdominal cavity (a) at 80 days’ gestation; (b) at 100 days’ gestation.
extends back into the pelvic cavity and is readily visible after around 100 days (Figure 218.2b). At this time the external genitalia are used to make a sex determination.
Precautions It is mandatory to keep good records because:
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Clerical errors will be made; It is necessary to have verification that the mare was sexed and a determination made; Occasionally identification mistakes are made at foaling.
Figure 218.3 Fetal sex determination certificate.
plane III (primarily useful for colt diagnosis; Figure 218.4d). Take-home hints
r Do not guess
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Recheck at a later date if unsure; Guessing leads to errors; Errors lead to loss of confidence in the procedure; Frequent equal signs can be seen around the umbilical area which are artifacts (Figure 218.19b).
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No guarantees Certificates are given (Figure 218.3) but not guarantees.
Ultrasonographic planes Four planes are commonly used in fetal sex determination: plane I (primarily useful for colts; Figure 218.4a): plane II female (ideal view for filly diagnosis; Figure 218.4b); plane II male (similar to female view but more anterior for colts; Figure 218.4c); and
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If the image is not clear, push down gently on uterus with the probe to bring the fetus closer to the probe. This changes the focal point for various areas of the fetus and creates movement of fluid in the allantoic fluid, causing the fetus to change position slightly. If the view of the fetus is not productive, turn the transducer 90◦ to the left. If the view is still not productive, turn the transducer 90◦ to the right. Somewhere in this 180◦ span is the proper scanning plane for viewing the tubercle. If the mare is straining, do not hesitate to use propantheline (30 mg i.v.) to lessen straining. This enables the examiner to move the probe to various positions to get different view angles of the fetus. A darker environment makes the details of the fetus easier to recognize. It may be necessary to place the probe in front of the hand to reach older fetuses. Evacuate fecal material as far anterior as possible.
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Figure 218.4 The four ultrasonagraphic planes used in fetal sex determination: (a) plane I (primarily useful for colts); (b) plane II female (ideal view for filly diagnosis); (c) plane II male (similar to female view but more anterior for colts); (d) plane III (primarily useful for colt diagnosis).
Plane I genital tubercle ribs front limbs hind limbs
(a) Figure 218.5 (a) Plane I diagram. (b) Plane I ultrasonographic view.
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Procedure at 55 days to 90 days Place proper restraint on the mare and evacuate feces thoroughly. If the mare is straining a smooth muscle relaxant may need to be given (propanthaline 30 mg i.v., or buscopan 40 mg i.v.) First determine the anterior–posterior positioning of the fetus. Find the skull (good anterior marker), the heart (good ventral marker), the umbilical entrance to the abdomen, a 6–7 mm dark hole just anterior to the genital tubercle (good ventral marker; Figure 218.12c,d), ribs coming off the vertebrae (good dorsal marker; Figures 218.11b, 218.19a), and the tail (good posterior marker). Using these markers for orientation, proceed to the posterior part of the fetus. Scan the area to see if an obvious genital tubercle is present. If not, try to get a transverse view of the abdomen where the plane of the ultrasound beam transects the fetus perpendicular to the axis of the fetal spine (plane III, Figures 218.11b, Figure 218.19a). It is necessary to look for markers (e.g., the ribs) that inform the examiner what is dorsal and what is ventral on the fetus. The spinal column exhibits as a hyperechoic equal sign and is a good dorsal marker (this looks slightly like a genital tubercle but is on the dorsal side rather than the ventral side (Figure 218.19a). The ribs coming off the vertebrae are a very good dorsal marker. The umbilical cord entering the abdominal cavity is a very good ventral marker. These markers give an idea of where to look for the genital tubercle. The genital tubercle for the male will be on the ventral midline just posterior to the umbilical entrance into the abdominal cavity (Figure 218.5). The genital tubercle for the female will be on the ventral midline just under the tail head (Figure 218.7b–e). If the ultrasound plane is perpendicular to the axis of the spine, continue with this plane until the view leaves the posterior part of the entire fetus. After there is no fetus in view, keeping this same plane, ease back toward the fetus. The first hyperechoic area to be seen on the dorsal side will be the tail head. It can be identified as the tail head because of the dorsal and ventral markers noted earlier. The next hyperechoic areas will be the genital tubercle just under the tail (in a female) and two hyperechoic areas (tibias) with no muscle mass around them about 1 cm from the tail, depending on the age of the fetus. These two hyperechoic areas are the tibias (Figure 218.7c,d). There now should be three or four hyperechoic areas in view. If it is a colt there will be three hyperechoic areas forming a tail-head–tibial triangle, with no hyperechoic area within this triangle. If it is a
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filly, a bilobulated hyperechoic equal sign will appear within the tail-head–tibial triangle, slightly closer to the tail head (Figure 218.7b–e). This view is an absolute must for identifying a filly. A colt tubercle can be visualized from many different angles including a frontal plane (plane I, Figure 218.5) that exhibits the front legs, ventral abdomen, and hind legs. The tubercle will appear on the ventral midline slightly anterior to a line drawn between the stifles and slightly posterior to the umbilical entrance into the abdominal cavity (Figure 218.6b,c). If there is a negative filly view (nothing within the tibial–tail-head triangle) then advance anteriorly on the fetus. The male tubercle will appear just at the ventral edge of the abdominal wall as a hyperechoic bilobulated equal sign (Figure 218.11c,d). Moving the transducer toward plane I, the umbilical entrance to the abdominal cavity appears as a small black hole anterior to the genital tubercle (Figure 218.11c,d). If the transducer is moved to plane II, a tubercle between the hind legs is visualized (Figure 218.6b,c). The tubercle should be anterior to a line drawn between the stifles to make sure it is a colt. Remember that at 52–53 days the tubercle is between the hind legs in both male and female. Not until at least 55 days of gestation can the direction in which the tubercle appears to be moving be determined. If the tubercle is anterior to a line drawn between the stifles it will be a colt. If the tubercle is posterior to a line between the stifles it will be a filly. If it is difficult to decide, wait 3 or 4 days and look again.
Procedure at 90 days to 160 days This procedure involves finding the external genitalia of the fetus. After 90 days the fetus begins to come back into view with the ultrasound. It is best to scan the entire fetus to get oriented anteriorly, posteriorly, dorsally, and ventrally. It is good to get a cross-sectional view of the abdomen and, keeping that plane, move off the fetus posteriorly. As you come back on to the fetus it is possible to get the same tail-head–tibial triangle with the clitoris showing up just under the tail head within the triangle (Figure 218.8). If nothing is under the tail head, notice the prominent ventral midline which appears as a white line (Figures 218.10a, 218.17). In a filly, following the prominent ventral midline posteriorly to the tail head, the clitoris, a small round hyperechoic mass (Figure 218.14a) will be encountered, or may appear as two hyperechoic dots lying on the ventral midline (Figure 218.14b). Following
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(c) Figure 218.6 (a) Plane II male diagram. (b,c) Plane II male, ultrasonographic view.
the ventral midline anteriorly, the mammary gland (Figure 218.9a–c) will be encountered. The mammary gland consists of three distinct parts: the bright small hyperechoic teats, on each lateral side of the gland (Figure 218.9a,c), and two dense halves of the mammary gland separated by a translucent line between the two halves (Figure 218.9a). Prepucial tissue is not as dense as a mammary gland, does not have a translucent line separating the halves, and obviously has no hyperechoic teats. In a male, following the ventral midline posteriorly using plane II, nothing will be encountered before reaching the tail head (Figure 218.17). Following the ventral midline anteriorly, the prepuce, a cone-shaped structure that comes to a point (Figure 218.10a,b), will be encountered. The penis can often readily be seen as a shaft-like structure with a bright hyperechoic mass on the end (Figure 218.10b– d). This bright hyperechoic tip is the glans penis. Using plane III view (a cross-section of the posterior ab-
domen) the prepuce will be seen as a cone-shaped structure off the ventral abdomen. Frequently the hyperechoic mass of the glans penis will be seen within the prepuce (Figure 218.12a–e). It is possible to get a plane III view of the glans penis without any of the prepuce in view (the glans penis frequently protrudes out the front of the prepuce). This will appear as a hyperechoic mass off the ventral abdominal wall (Figure 218.12a,d,e) just posterior to the urachus (umbilical entrance into the abdominal cavity, Figure 218.12c,d). The main marker to be utilized is the prominent ventral midline (Figures 218.10a, 218.16). The gonads can readily be seen but should used to support a diagnosis and not to make a diagnosis. The male gonads usually appear very homogeneous (Figure 218.18a) where as the female gonads have oval ring-like translucent areas (Figure 218.18b). These can be difficult determinations and are not reliable for a diagnosis.
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(e) Figure 218.7 (a) Plane II female diagram. (b–e) Plane II female, ultrasonographic view.
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Figure 218.8 Plane II female, ultrasonographic view.
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(c) Figure 218.9 (a–c) Plane II female, ultrasonographic view.
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Figure 218.10 (a–d) Plane II male, ultrasonographic view.
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Figure 218.11 (a) Plane III male diagram. (b–d) Plane III male, ultrasonographic view. (e) Cast of male fetus, 110 days.
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(e) Figure 218.12 (a–e) Plane III male, ultrasonographic view.
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Figure 218.13 Sagittal view.
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Figure 218.14 (a,b) Oblique view.
Figure 218.15 Oblique view.
Figure 218.16 Oblique view.
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Figure 218.17 Oblique view.
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Figure 218.18 Gonads. (a) Male gonads; (b) female gonads.
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Figure 218.19 (a,b) Artifacts.
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Further information Ginther OJ. Ultrasonic Imaging and Animal Reproduction. Horses Book 2. Crossplains, WI: Equiservices, 1995. Ginther OJ, Curran S, Ginther M. Fetal Gender Determination in Cattle and Horses. Crossplains, WI: Equiservices. Instructional video, available from Equiservices, 4343 Garfoot Road, Crossplains, WI 53528 (telephone 608-7984910). Holder RD. Sex determination in the mare between 55–150 days gestation. Proceedings of 46th Annual AAEP Convention, 2001; pp. 321–4.
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Holder RD. A Guide to Equine Fetal Sexing 55–150 days. Instructional video, available from: Jim Chryzan, Aloka Co., 10 Fairfield Blvd, Wallingford, CT 06492-7502 (telephone 7409276147); or Hagyard Davidson & McGee, 4250 Iron Works Pike, Lexington, KY 40511 (telephone 859-2558741). Holder RD. Chapter 53. In: Samper JC, Pycock J, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007. Holder RD. Chapter 5.22. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. Philadelphia, Saunders, 2003.
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Chapter 219
Fetal Gender Determination from Mid to Advanced Gestation Stefania Bucca
Equine fetal sex determination can be successfully accomplished by combining transrectal and transabdominal ultrasonography (US) techniques from 2 to 8 months’ gestation. The active nature of the equine fetus determines frequent changes in presentation up to 9 months’ gestation, making transrectal examination potentially diagnostic, even in advanced pregnancy. The ability to change presentation decreases as gestation advances.1, 2 Fetal size and encasement of the hindquarters within the fetal horn prevents further changes in presentation past 9 months’ gestation. After 9 months, fetal sex can still be determined in grossly undersized fetuses and when posterior or transverse presentations persist. Although the equine fetus can be successfully identified by transabdominal US as early as day 100 gestation, fetal sex diagnosis by this route is usually carried out from 4 to 8 months pregnancy and more commonly between days 150 and 260. In small ponies and miniature horses the acoustic window for diagnosis can be further stretched towards term. Convex and sector technologies are both more appropriate than linear ultrasonography for this diagnostic approach due to the better compliance of convex and sector transducers to the curvilinear contour of the mare’s abdomen. Probe frequencies ranging from 2 to 6 MHz are required to obtain a maximal scanning depth of 25–30 cm and successfully examine fetuses of gestations comprised between 5 and 8 months. Thorough patient preparation is rarely an option, as oftentimes mares examined for fetal sex determination are destined for the sales ring and clipping of the abdomen would not impact favorably on prospective buyers. Liquid paraffin provides an ex-
cellent alternative, but requires adequate transducer protection. During the summer months a fine coat will allow good transducer contact, just by spraying or sponging alcohol and applying US gel to the clean skin. In colder months and in mares with coarse, thick coats the quality of image, without clipping, will not be ideal, but may be improved greatly by brushing, washing, and scraping the mare’s ventral abdomen dry and then apply alcohol and a generous amount of US gel. A thick layer of subcutaneous fat will decrease US penetration through the mare’s abdominal wall and reduce image quality. Ideally, during the examination mares should be restrained in purposebuilt stocks, allowing easy access to the mare’s lower abdomen. It is advisable to commence the procedure by US per rectum, as diagnosis may be obtained by this route when the fetus lies in posterior presentation. Furthermore, by ascertaining presentation, clinicians may establish the extent of fetal engagement within the mare’s pelvis and predict which part of the abdomen will need to be examined. As a rule of thumb, with a fetus in posterior presentation, too far forward in the mare’s abdomen for transrectal sex determination, only the caudal half of the abdomen needs to be scanned in mid to advanced gestation, and diagnosis can be frequently carried out to term. In anterior presentation, fetal hindquarters are usually visualized in mid abdomen at 5–6 months’ gestation, mid to cranial up to 7 months and in the cranial third of the abdomen at 8 months. Mare’s parity and fetal activity may profoundly alter fetal position, from the outlined expected location. The likelihood of a rapid change of presentation is high, particularly in
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Fetal Gender Determination from Mid to Advanced Gestation pregnancies of 5–6 months. Transabdominal fetal sex determination provides the operator with unrestricted views and scanning time to obtain the most favorable access to fetal hindquarters, for accurate diagnosis. Fetal imaging and body orientation within the abdomen may not be as predictable as at an earlier gestational age or as it is perceived within the boundaries of the mare’s pelvis. Difficulties may be encountered in negotiating three-dimensional fetal orientation, when only two-dimensional imaging may be obtained by US. The fetal spinal cord often displays a pronounced curvature and frequently front and hindlimbs lie on opposite sides. The flexed pelvic limb may obscure the areas of diagnostic interest, with osseous components casting shadows over the pelvic organs. Umbilical cord segments and loops of maternal intestine with gaseous content may also interfere with proper imaging. Occasionally, lung parenchyma may block out US viewing in a cranially displaced fetus, resting against the mare’s diaphragm. Scanning planes of choice include: frontal plane to evaluate gonads, scrotal compartments, penis and penile urethra; cross-sectional planes to visualize the penis; frontal oblique and cross-sectional oblique planes to assess the perineum and ventral pelvis, which includes all primary sex organs. Active fetuses may swiftly change location within the uterus, and the mare’s excitement and anxiety strongly influence fetal activity. Carrying out the examination in quiet and familiar surroundings for the mare will usually facilitate good US imaging. Sedation of the mare profoundly curtails fetal activity and lowers the fetus towards the ventral uterine wall and abdominal floor, enhancing the quality of image. Fetuses undergoing dormant phases in awkward positions, whereby hindquarters cannot be easily investigated by US, may be aroused just by walking or trotting the mare out of the examination room. Transducer crystal configuration of sector and convex technologies produces a typical pie-shaped sonogram. Anatomical distortion, caused by a single oscillating crystal in sector technology, has been the reason for convex probes to become so rapidly popular with ultrasonographers, particularly when the abdominal content is to be evaluated. For ease of sonogram interpretation a standard technique should be adopted, as follows: r Both sides of the abdomen may need to be scanned. r To examine the left abdomen, the operator should stand close to the mare’s left flank, cranial to the stifle, facing the US unit screen, positioned towards the mare’s head; the transducer should be placed in her/his right hand and the US unit set at eye level.
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For the right abdomen, the operator should stand on the opposite side and hold the probe in her/his left hand. The craniocaudal orientation of the US probe is determined and US gel is liberally applied over its footprint. Scanning will commence over the ventral midline, in the area where fetal hindquarters are assumed to be resting, according to gestational age and results of transrectal examination; the skin had previously been prepared. When fetal parts are encountered, the rib cage and spinal cord are identified. The US beam is oriented parallel to the spinal cord and fetal craniocaudal orientation is determined, by scanning back and forth between cardiac area and stomach. Fetal presentation and position are determined. The US probe is then moved caudally within the fetus, to identify the hindquarters; ventrocaudal orientation is determined by scanning from the spine to the umbilical cord and/or fetal limbs. The resulting image on the screen will be the upside-down projection of the area examined.
From 5 to 8 months’ gestation, fetal sex determination is based on the visualization of primary sex organs. Ease of identification will depend on equipment, transducer contact, fetal cooperation, and operator’s ability to adjust probe orientation for optimal acoustic viewing. No structural changes are observed in the anatomy of fetal sex organs from 5 months’ gestation to term. The only substantial difference regards size and therefore accessibility, which determines quality of image, granted optimal US penetration and proximity of the fetus to the mare’s abdominal wall are available. In the male fetus the penis is usually visualized in great detail; hyperechoic dots are clearly visible on the tip of the penis encased within the prepuce.The penis may be clearly protruding from the sheath (see Figure 219.1); under those circumstances it becomes closely associated with the umbilical cord and displays rhythmic trembling, synchronous with the umbilical vessels’ pulsatile activity. The urethra (see Figure 219.2) may be visualized in its perineal or penile tracts, both in longitudinal and cross-sectional views. Cross-sectional imaging of the male urethra (see Figure 219.3) may display a typical circular image or appear ellipsoidal, when the sound beam is tangential to its course.The scrotum may be detected on a cross-sectional oblique view of the male caudal pelvis; a median rafe may sometimes be appreciated. On a frontal plane the scrotal boundaries are more difficult to define, but the scrotal compartments appear as a distinctive feature. The scrotal compartments (see Figure 219.4) appear as two symmetrical, hypoechoic, elliptical areas, obliquely oriented
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Figure 219.1 Frontal oblique scan of the pelvis of a 152-dayold, male fetus showing a longitudinal view of the tip of the penis, protruding from the sheath. Both testicular compartments can be identified. The umbilical cord is visualized cranial to the penis. Transabdominal scan, where left of sonogram is mare’s cranial.
Figure 219.2 Cross-sectional oblique scan of the perineum of a male fetus showing perineal urethra, scrotum and tip of penis, as indicated. Transabdominal scan of a mare 150 days pregnant. Left of sonogram, mare’s cranial.
Figure 219.3 Cross-sectional view of the caudal abdomen of a 218-day-old, male fetus: penis and urethra are visible, above the umbilical cord. Transabdominal scan, where left of sonogram is mare’s cranial.
Figure 219.4 Frontal oblique view of a 235-day-old, male fetus showing perineal urethra and testicular compartments. Transabdominal scan, where left of sonogram is mare’s cranial.
and located cranial to the perineal urethra and caudal to the base of the penis. Each one identifies the position of one gubernaculum testis. In crosssection, their location within the scrotum may be recognized. Fetal gonads are elliptically shaped and occupy the ventrocaudal abdomen, below each kidney, with an oblique orientation, divergent cranially. Caudally, they are closely associated to the bladder and urachus. Male fetal gonads (see Figure 219.5) appear uniformly echogenic, except for a central hyperechoic line running on the long axis, believed to correspond to the location of the testicular vein, in postnatal life. Color Doppler ultrasonography of the fetal gonad clearly identifies the course of the testicular vein in the male, while a strong cortical pattern is evident in the female fetus (see Figures 219.6 and 219.7). The epididymus may be visualized on the dorsolateral aspect of each gonad.
Figure 219.5 Frontal plane view, through mid abdomen in a 159-day-old, male fetus, showing both fetal gonads, cranial to the pelvis. The upper gonad displays a typical central hyperechoic line. Transabdominal scan, where left of sonogram is mare’s cranial.
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Figure 219.6 Color Doppler scan of a 182-day-old, fetal male gonad, displaying the course of the testicular vein. Transabdominal scan, where left of sonogram is mare’s cranial. (See color plate 112).
Figure 219.8 Cross-sectional oblique view of the perineum of a 242-day-old, female fetus, showing a ‘single line’ perineum and a triangular shaped mammary gland. Transabdominal scan, where left of sonogram is mare’s cranial.
The mammary gland (see Figure 219.7) is best visualized on a frontal oblique plane, where its shape may vary from triangular to trapezoidal. On this view, the cranial border appears slightly concave and the nipples (see Figure 219.8) are commonly identified as two hyperechoic protrusions of the cranial margin of the mammary gland, close to the inner thigh.The caudodorsal perineal location of the equine fetal vulva makes it difficult to visualise it on the same scanning plane as the perineum. Usually only the clitoris is identified on this view (see Figure 219.10), as a hyperechoic, caudal, ‘teardrop’ structure, bulging out, where the buttocks meet. The rest of the perineum is made out of a single echogenic line, extending from the clitoris to the mammary gland. In the equine filly fetus, gonads present a structural differentiation between cortex and medulla (see Figure 219.10), which translates into substantial echotex-
Figure 219.9 Cross-sectional oblique view of the perineum of a 237-day-old, female fetus, showing a triangular shaped mammary gland with a prominent nipple. Transabdominal scan, where left of sonogram is mare’s cranial.
Figure 219.7 Color Doppler scan of a 176-day-old, fetal female gonad, displaying a strong cortical pattern. Transabdominal scan, where left of sonogram is mare’s cranial. (See color plate 113).
Figure 219.10 Cross-sectional oblique view of the dorsal perineum of a 180-day-old, female fetus, showing a ‘single line’ perineum and the clitoris, to its upper end. Transabdominal scan, where left of sonogram is mare’s cranial.
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Figure 219.11 Frontal plane view, through mid abdomen in a 130-day-old, female fetus, showing both fetal gonads, at each side of the bladder, cranial to the pelvis. A faint line is separating cortex from medulla in the upper gonad. Transabdominal scan, where left of sonogram is mare’s cranial.
ture variation, between inner and outer layer. This sonographic distinction is very noticeable in mid pregnancy (see Figures 219.11 and 219.12), but becomes less evident as gestation advances. Fetal sex determination in mid to advanced gestation should be based on the identification of at least three different parameters. The size of the fully differentiated primary sex organs and the anatomical detail provided by good quality US equipment make diagnosis easy to accomplish, once proper fetal orientation is obtained. A systematic approach should be adopted when examining the equine fetus by US. Craniocaudal and dorsoventral orientation should be established first. US examination should then focus on the fetal hindquarters, scanning from the spinal cord to the umbilical cord and back. The area caudal to the root of the umbilical cord and cranial to convergence of the upper hindlimb should be carefully examined. Penis, scrotal lodges, and perineal urethra are identified in a craniocaudal scan of the male fetus. In the same area, the filly fetus displays the mammary gland and nipples, both characterized by a strong and uniform echogenicity. In addition, the female perineum shows a single line demarca-
Figure 219.12 Frontal plane view, through mid abdomen in a 120-day-old, female fetus, showing both fetal gonads, at each side of the bladder, in close contact with the umbilical arteries. Cortex and medulla are clearly identifiable. Transrectal scan, where left of sonogram is mare’s cranial.
tion between thighs (see Figure 219.8), as opposed to the well-defined double line (see Figure 219.2) in the male perineum, corresponding to the long, tubular male urethra. By following the perineum in a dorsal direction, the clitoris should be imaged, close to the root of the tail. A frontal plane scan, dorsal to the umbilicus and ventral to the kidneys, will finally identify the gonads and their distinctive sonographic appearance will confirm sex diagnosis.
References 1. Ginther OJ, Griffen PG. Equine fetal kinetics: Presentation and location. Theriogenology 1993;40:1–11. 2. Bucca S. Equine fetal gender determination from mid to advanced gestation by ultrasound. Theriogenology 2005;64:568–71. 3. Ginther OJ, Curran S, Ginther M. Fetal gender determination in cattle and horses. Video. Equiservices. 4343 Garfoot Rd, Crossplains, WI 53528. 4. Renaudin CD, Gillis CL, Tarantal AF, Coleman DA. Evaluation of equine fetal growth from day 100 of gestation to parturition by ultrasonography. J Reprod Fertil Suppl 2000;56:651–60.
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Chapter 220
Management of Twins Angus O. McKinnon
Introduction Twins are avoidable and should not persist on wellmanaged breeding farms. Clients understand this and demand the most sophisticated management programs available. Presented below are the management techniques in use at the Goulburn Valley Equine Hospital. The hospital is a major referral center and each year is presented with multiple referral cases of twin pregnancy that are diagnosed after cessation of the mobility phase. Historically, twins have been the single most important cause of abortion.1–5 With the advent of ultrasonography the incidence of abortion associated with twins has decreased dramatically to less than 6.1%,6 2.9%,7 or 1.7%8 of all abortions. Twins are often disastrous financially. Most twin pregnancies terminate in early fetal resorption or loss, late-term abortions, or the birth of small, growth-retarded foals. Mares aborting twins in late gestation frequently have foaling difficulties, damage their reproductive tracts, and are difficult to rebreed. If foals are born alive they are frequently small, demonstrate the effects of intrauterine growth retardation, and have a poor survival rate with many needing expensive sophisticated critical care. Origin, associations, and a general overview of twinning in the horse has been published elsewhere9, 10 (see Chapter 241). It is our responsibility to successfully manage early pregnancies such that no mare delivers or aborts twin foals. In consultation with farm managers, owners, and clients we must aim to utilize available equipment and technology commensurate with economic constraints and other owner/manager preferences to diagnose twin pregnancies as early as practically possible. It is also the responsibility of the veterinary profession to adequately inform owners/managers/clients of reasons why twins may be not diagnosed.
Twins are still occasionally missed despite multiple examinations. Reasons why twins may not be detected, despite repeated examination are: (i) difficulty distinguishing structures (this may be related to a poor examination environment, i.e., too much light, poor display characteristics of the ultrasonographic unit, mare movement and/or lack of restraint); (ii) variable growth patterns especially from asynchronous ovulations; (iii) inability to detect heartbeats of adjacent embryos; (iv) operator experience; (v) resolution of the equipment; and (vi) potential confusion of one or both pregnancies with uterine cysts. The most common cause of misdiagnosing the presence of twin pregnancies is examination prior to the time period that a second pregnancy (asynchronous ovulation) may be reasonably expected to be detected. Another reason is examining too quickly or terminating the examination once a single pregnancy has been detected in later (post-fixation) cases of bilaterally fixed (not unilateral) twin pregnancies. Also of note is the propensity of the uterus of early pregnant mares (days 20–35) to bulge forward at the corpus cornual junction, and it is possible with either a single pregnancy or twins to miss a pregnancy at the corpus cornual junction and yet scan the entire uterine horn to the body on both sides without the uterus ever leaving the screen.
Management techniques The mare is generally efficient at reducing unilateral twins to a single pregnancy. This is done by a competitive absorption of nutrients that is related to size and position of the early pregnancy and later to orientation of the embryo proper within the developing conceptus11 (see Chapter 241). However, any nonintervention decision needs to extrapolate the probability of successful reduction based on the age of identification, the orientation and fixation of the vesicles, and any disparity in size. It is not considered
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 220.1 (a and b) Asynchronous ovulations will result in vesicles of a different size. (c) Synchronous ovulations result in twins of similar size. (d) Gross disparity in size suggests either a large number of days between ovulations or abnormal embryonic development of the smaller vesicle.
good practice to leave twin pregnancies alone in the pre-fixation period unless one or both are too small to manipulate. Recognition ≤16 days from ovulation Embryo reduction on or before the day of fixation is not considered an important aspect of the natural correction of twins.12 The probability of a mare losing one or both vesicles of a set of twins from identification prior to fixation is minimal, and approximates that of early embryonic death for the same time period (per vesicle). The recognition of twin pregnancies prior to fixation day (day 16) is dependent on the day of examination relative to the day of ovulation. Asynchronous ovulations occasionally result in a gross disparity in vesicle size (Figure 220.1a,b), sometimes as much as 4–5 days, for example, identification of a day 11–12 and a day 16 vesicle concurrently. In instances such as this, examination one day earlier may have failed to detect the younger of the two pregnancies. Recognition that almost all twin pregnancies occur from multiple ovulation dictates
mandatory re-examination of all mares that have two corpora lutea and only a single vesicle detected prior to fixation (day 16). Recognition prior to fixation is also dependent on operator experience, resolution of the equipment (≥ 5 MHz preferred), monitor capabilities, restraint and other facilities (ability to darken the environment), the presence of uterine cysts, and the skill of the examiner. Twin or multiple pregnancies recognized prior to the day of fixation (Figure 220.1a–d) should be reduced to a singleton as soon as practically possible (see below). As smaller than expected single pregnancies have a higher rate of early embryonic death,9, 13 management techniques should consider the destruction of the smaller vesicle10 (Figure 220.1d) even if the body of evidence suggests minimal pre-fixation embryonic reduction. Recognition after fixation (after day 16) It should be clear that our philosophy is that non-intervention is only acceptable when twins are diagnosed as a unilateral occurrence between days
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Management of Twins 17 and 60 and then the decision depends on factors such as the value of the foal, the potential for rebreeding (endometrial cup formation, stallion availability, etc.), and the ability of the veterinarian to manually intervene. Intervention in twin pregnancies is strongly recommended in all other circumstances (see below). Intervention must occur before day 35 if endometrial cup formation is to be prevented and immediate rebreeding is contemplated. When bilateral twins are diagnosed after day 16 then destruction of one should be attempted as soon as possible. The incidence of loss of one of a set of bilateral fixed twins is similar to the probability of loss for a single pregnancy. For discussion on the outcome of nonintervention programs, please refer to Chapter 241.
When and how should we intervene? Recognition ≤16 days from ovulation The first technique for manual crush of the conceptus during the mobility phase utilized manual reduction with good results14 and was a variation from previously reported techniques for twin pregnancies.15, 16 The technique involved gentle manipulation of the embryonic vesicle to the tip of one uterine horn and manual rupture. When applied to single pregnancies it resulted in pseudopregnancy and when applied to twin pregnancies it resulted in a single pregnancy in seven of eight attempts.14 Utilizing the same embryo reduction techniques, later studies incorporated treatment of mares with single or multiple intramuscular injections of progestagens administration (hydroxyprogesterone caproate), an anti-prostaglandin (flunixin meglumine) plus progestagens, or given no treatment prior to manual embryonic rupture in the mobility phase.17, 18 Results were 10/10 (100%) mares maintaining pregnancy in the control group (no treatment, just manual rupture) and 37/40 (92.5%) for treated mares. The amount of PGF2α released was directly correlated with the pressure required to cause embryonic rupture. Flunixin meglumine inhibited PGF2α release after embryonic rupture. Treatment with progestagen plus flunixin meglumine or progestagen singly or multiply was not better than no treatment at all (although it was subsequently shown that the progestagen chosen had no ability to maintain pregnancy in ovariectomized mares and did not bind to progesterone receptors in the horse).19 Another report20 demonstrated that 60 of 66 mares (90.9%) maintained a single vesicle after manual reduction was attempted prior to fixation. Five of the six mares in which the procedure was not successful subsequently conceived. Since 198421 we
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have used a modification10 of the technique described originally.14 With this technique the ultrasound probe is used to manipulate the vesicles while keeping one or both vesicles in view during the manipulation and more importantly the crushing or rupture of the vesicle (Figure 220.2). Utilizing this technique it is possible to more accurately and quickly separate vesicles. It was originally proposed14 that when vesicles were in apposition mares be re-examined approximately 1 hour later. By utilizing the probe to manipulate vesicles, separation is achieved (prefixation) very quickly in most instances. Commonly the smaller vesicle is destroyed despite the lack of evidence to support pre-fixation reduction. On occasion, it is necessary to reattempt the embryo reduction technique 24–48 h after the original evaluation if the smaller of the two vesicles is less than 10 mm in diameter as sometimes these can be more difficult to rupture. Separation of vesicles should always be possible if the vesicles are still able to be identified as two spherical non-coalesced structures. Briefly, the technique involves separation of the vesicles using the probe. A finger is placed on either side of the probe to help stabilize the vesicle to be moved. Gentle sideto-side movement of the probe with pressure results in the two vesicles becoming separated (Figure 220.2). The separation is identified by lack of a vesicle under the probe (just the homogenous gray of the uterus). The vesicle can be crushed as close as 5 mm from the other but it is generally best to separate them at least 20 mm. This is in case the mare moves at the time of increasing pressure. The vesicle is crushed by gradually increasing the pressure using the probe. Occasionally, refractory cases may need a sudden increase in pressure much like a quick flick or snap at the end of the probe. This latter technique is quite useful for smaller (day 11–13) vesicles. When the vesicle is crushed it is not uncommon for its fluid to surround the other. This is not a problem at this stage of pregnancy. Later (≥ day 25) fluid surrounding the other vesicle is thought to be a potential problem. Generally, separation prior to fixation is not difficult and the crushing technique described above is appropriate for most vesicles between days 13 and 16. Smaller vesicles are often more difficult to crush and they should be managed with a rapid snap or crack with the tip of the probe (or left alone for 1 or 2 days). This technique can only safely be applied by placing a finger on either side of probe to restrict the movement of the vesicle while gentle pressure on the probe begins to distort it. At that time a sudden force (snap) is applied to the tip of the probe. Only people well experienced in rectal ultrasonography should attempt this technique as it can be dangerous for the mare.
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Figure 220.2 The technique for crush of a vesicle pre-fixation. The vesicles to be separated are identified. Frequent back and forth scanning may be necessary to continually confirm the location of each vesicle (a) and then gentle pressure is used to separate it from the other (b). As pressure is gradually increased the vesicle changes shape and starts to flatten (c). At the appropriate moment a little quick extra pressure completes ablation. It is quite common to see fluid immediately after the crush (d).
Specific variations that can result in vesicles being difficult to crush are when the vesicle is on the bladder (Figure 220.4a–c), when the mare has an excessively full bladder, or when the vesicle is immediately in front of the cervix (Figure 220.4d). When the vesicle is on a full bladder it will move away from the probe as soon as any pressure is applied, and it must be stabilized and managed by the snap technique. A mare is more likely to present with an excessively full bladder when mares have been held in yards for a few hours prior to being examined, and appear uncomfortable in their environment. These mares are best turned out into a yard by themselves after examination as most will urinate within 10 minutes due to stimulus from palpation. Alternatively the mare can be re-presented a day later (after being left not yarded). When the vesicle is immediately adjacent to the cervix it is difficult to apply the pressure due to
physical restriction to force created by the body of the cervix. In these cases it is best to move the vesicle away from the cervix before attempting to crush, or re-examine the mare in a few hours. Most twins in contact will have a horizontal orientation towards each other (Figs 220.2a and 220.3a); however, on occasion the twins may be vertically orientated (Figure 220.5a,b). Gentle pressure usually returns them to a horizontal orientation (Figure 220.5c). At the Goulburn Valley Equine Hospital, records were evaluated for 1716 Thoroughbred mare cycles and 1294 Standardbred mare cycles.22 Twins were diagnosed in 245 of 1716 cycles in Thoroughbred mares (14.3% of cycles) and 46 of 1294 of Standardbred cycles (3.5%). After twin reduction, mares are not routinely examined until the next scheduled examination, i.e., 21–25 days post-ovulation (detection
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(e) Figure 220.3 Separation of twins that are together is possible if they are able to assume a figure ∞ shape (a). Gentle pressure results in them moving apart (b, c and d). When they are separated enough to crush the vesicle has disappeared from the screen. When vesicles are in close proximity to each other at the time of the crush it is quite common for fluid from the crushed vesicle to surround the other (e).
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Figure 220.4 A vesicle immediately over the bladder. (b) When a vesicle is crushed over the bladder the ‘snap’ technique is recommended. (c) Fluid and yolk sac membranes from a vesicle crushed over the bladder. (d) A pregnancy (arrow) immediately adjacent to the cervix. The cervix is to the left of the screen and uterine cysts are to the right of the vesicle.
of the fetus). When mares were re-examined after pre-fixation embryonic reduction, 10/245 Thoroughbred mares (4%) had lost the remaining pregnancy and 8/46 Standardbred mares (17.4%) were not pregnant. The number of Thoroughbred mares losing the remaining pregnancy (4.0%) is similar to 3.7% (63/1716) which was the measured rate of early embryonic death (EED) on the same farms for mares with a single pregnancy diagnosed at day 13–15 and then subsequently found to be empty at the next scan. Interestingly, the number of Standardbred pregnancies lost was much higher after twin reduction (8/46, 17.4%) compared to the measured rate of EED (7.1%). This we believe was likely related to economics wherein clients with Standardbred mares were unwilling to examine for pregnancy early. Thus twins, when detected, are likely to be closer to final fixation or fixed already and harder to manage.
A further study from the Goulburn Valley Equine Hospital23 demonstrated that between detection and day 45 there were significantly fewer embryonic losses in mares diagnosed with multiple pregnancy (29/633, 4.6%) compared to singleton pregnancy (408/5414, 7.5%, P = 0.004). It is our contention that the procedure has developed to the stage that it is always expected that a single pregnancy will exist after pre-fixation embryo reduction is attempted. Unless a mistake occurs and the other vesicle is ruptured at the time of initial manipulation, we feel that any failure to survive the procedure is more likely a result of uterine inflammatory changes and infection or an embryonic defect rather than a result of the procedure. We believe that this is the most reliable technique available but feel it is important to highlight the experience of the personnel involved. From discussions with farm
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(c) Figure 220.5 On occasion the twins may be vertically orientated (a and b). Gentle pressure usually returns them to a horizontal orientation (c).
managers and other veterinarians, it is clear that only veterinarians involved with sophisticated reproductive management such as the routine use of ultrasonography can expect to achieve these types of results. Our strong recommendation to veterinarians and clients is that all mares are examined within 14–16 days of breeding. Expected time to ovulation after breeding will depend on frequency of examination and use of ovulation induction agents such as hCG or GnRH. Factors that may modify this decision are breed, mare value, ability of the stud master or owner to facilitate examination of the mare, and, on occasion, education of the owner.
Recognition after fixation (after day 16)
Days 17 to 20 In all cases of bilaterally fixed twins, one should be destroyed immediately. The mare has an extremely ef-
ficient biological embryo reduction mechanism that operates when twins are in apposition (unilaterally fixed).11, 24, 25 Reduction occurred in 100% of 22 mares with asynchronous ovulation (vesicles size difference greater than 4 mm in diameter) and 19/26 (73%) of mares with similar vesicle size (0–3 mm difference). Because the rate of embryo reduction between days 17 and 20 is so high for unilateral fixation, equine practitioners frequently elect to leave these developing pregnancies and determine their outcome later. Our philosophies are that if the two vesicles have coalesced into one larger vesicle with an ultrasonographic visible line in division (Figure 220.6a) they are left totally alone for 5–10 days, and more particularly so, if there is any unevenness in vesicle size. If the individual vesicles have still retained a spherical orientation or a spherical shape (like a figure ∞), then they can be separated gently with the probe and are crushed either in situ or after being manipulated apart (Figure 220.6b–h). Due to the nature of our
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Figure 220.6 (a–h) If the twin pregnancy has coalesced to a vertically orientated line it is difficult to separate them and they should be left alone (a). If the individual spherical relationship is still maintained (b) then they can be manipulated carefully (e–h). It is difficult to separate these vesicles and pressure is gently applied until one flattens and then ruptures (g and h).
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Figure 220.6 (Continued).
practice, few mares present with this configuration in the Thoroughbred population; however, it is not uncommon in the Standardbred population wherein economics dictate that pregnancy diagnosis is often delayed past the time the mobility phase has ended. Results from our practice with twins in this configuration are reduced compared to pre-fixation intervention procedures. Others have reported good results post-fixation. One group reported success in 49/50 cases post-fixation.18 The work of Bowman20 more closely parallels our experiences. With bilateral embryo fixation and intervention, Bowman reported almost no losses with 40/44 mares from day 16 to day 30 (90.9%) having a single pregnancy detected on day 45. With unilateral fixation the results were days 16 to 17 (16/18, 89%), days 18 to 19 (23/24, 95.8%), days 20 to 21 (8/13, 61.5%), days 22 to 24 (4/9, 47.4%), and days 25 to 30 (1/4, 25%). The high incidence of embryo reduction with unilateral fixation and the low incidence with bilateral fixation makes recommendations with twins in the days 17 to 20 period quite clear.
Days 30 to 35 During the period prior to the formation of endometrial cups, gentle pressure may be placed on one vesicle in cases of unilateral fixation. We do not attempt total ablation at this time as the resulting fluid from ruptured membranes sometimes surrounds the other fetus and effectively separates or prevents placental (chorionic girdle/trophoblast cells) attachments to the uterus. In these cases (total rupture of the vesicle), death of the remaining vesicle is very common. Between days 30 and 35 we attempt to pinch one vesicle with the probe and create a ‘snowflake’ effect which is the shedding of cells from the membranes. Demonstration of this effect almost always results in gradual loss of the affected conceptus. The pinching of the vesicle can be likened to membrane ‘slipping’ of bovine fetal membranes except that we use the probe to produce the effect. Occasionally, in mares with multiple cysts, twins may be missed and then identified at a later time. In general if they are unilateral it is best to leave them to the mare’s natural embryonic reduction method until day 30, when ablation can be attempted (Figure 220.9).
Days 21 to 30 All cases of bilaterally fixed twins of this age group are manipulated and one destroyed immediately (Figure 220.7). In most cases we do not attempt to manually destroy one vesicle with unilaterally fixed twins (Figure 220.8) of this age group until after day 30 and before day 35. At this age it is too easy to rupture both vesicles and the maximum success we believe we can expect is 50% (see previous section) which is less than or similar to the mare’s own biological reduction mechanism.
Days 36 to 60 Manual manipulation using ultrasonographic guidance From day 36 onwards it is a reasonable assumption that endometrial cup formation and subsequent eCG secretion will prevent many mares from returning to heat after early embryonic death due to varying susceptibility of the supplementary corpora lutea to prostaglandin. Abortion after 35 days is commonly associated with difficulties recycling the mare.26, 27 In
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Figure 220.7 Bilaterally fixed twins at ∼ day 24 (a and b). Firm pressure results in flattening of the vesicle (c) and ultimately rupture (d).
one study,28 when mares were aborted either between days 26 and 31 or between days 30 and 50, 8/11 became pregnant versus 2/7, respectively. This is similar to the work of Pascoe29 who concluded that the administration of a prostaglandin analog < 35 days of gestation was outstandingly successful as a method of treatment for twin pregnancy as mares could be rebred. Manual intervention at this time in our experience is very good in bilateral twin pregnancies (< 45 days) and approximately 65% successful in unilateral twin pregnancies. Success improves with use of more subtle pressure and damage to the chorioallantoic membrane rather than complete rupture in one attempt. Demonstration of the ‘snowflake effect’ without vesicle rupture consistently results in a gradual (48 h) stress of the fetus and ultimate loss of heartbeat. These pregnancies have the fetal fluids that become progressively more hyperechoic and reduce in size without interfering with the survival of the other
fetus (Figure 220.10). Early pregnancies tend to be resorbed without a major increase in echogenicity of the fetal fluids (Figure 220.11). It is important with these fetuses to always attempt to damage the same one. Multiple attempts, i.e. every day or every other day for 5 to 10 sessions may be necessary to elicit the desired response (Figure 220.12); however, quite frequently we are unable to create sufficient damage for fetal destruction without danger to the other pregnancy. In these cases, rather than creating major trauma (rupture of the vesicle), an alternative approach is sought after day 60. A combination of membrane slip and or oscillation of the fetus are used. If we are unsuccessful in establishing a response at this stage then the pregnancy is left until after day 100 (see below). Our anticipated success rates are 65% reduction and ∼ 30% of cases result in further examination after day 100. With careful manipulation the demise of both fetuses should occur less than 5% of the time.
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Figure 220.8 Twins at day 30 (a and b). The membrane between the twins can be clearly detected (c). Rupture of the vesicle is demonstrated by multiple membranes (d). It is not ideal to rupture membranes in unilateral pregnancies or bilateral pregnancies after ∼ day 35.
A variation for fetal destruction that is only possible between days 45 and 50 is oscillation and dislocation of the fetus from the umbilicus (Figure 220.13a–d). This technique uses the weight of the fetus on a specific length of umbilicus to create a whiplash effect. The frequency of the oscillation varies from 0.5 to 2 per second. Force is not the key, rather a technique that moves the fetus maximally. The oscillation technique works exceptionally well with bilaterally fixed 45–50-day pregnancies and quite well in unilaterally fixed pregnancies when the one fetus can be isolated. A variety of other methods have been reported as used to treat twins at this stage. Manual crushing was originally reported16 and results suggested that earlier crushing was better and, if possible, crushing should occur prior to day 31 because after day 35 sometimes manual rupture was not possible. The author quoted the following results between days
35 and 45: 60% resorption of both, 20% single foaling, and 20% survival of both. Pascoe29 demonstrated that needle puncture of one twin combined with nonsteroidal anti-inflammatory treatment (meclofenamic acid) resulted in no foals born. A more elaborate and invasive approach, such as intrafetal injection with saline via a laparotomy has been reported30 but is not really practical, or removal of one fetus via a video endoscope31 which also has been abandoned. An interesting report was the surgical technique for removal of one conceptus from mares with twin concepti more than 35 days of gestational age. 32, 33 Eight mares had bicornuate pregnancies and seven mares had unicornuate twin concepti. Five of six surviving mares with bicornuate twin concepti delivered a single viable foal, and none of the seven mares originally with unicornuate twin concepti produced a foal. The poor survival rate of unicornuate twin concepti was attributed to
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Figure 220.9 Twins identified with multiple cysts. (a) The fetus is visible in the lower left-hand corner. (b) The other twin is visible in the upper right-hand corner. (c) A single pregnancy is present 6 days after manipulation. (d) A single pregnancy is detected 18 days after manipulation.
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Figure 220.10 Identification of twin pregnancies at ∼ day 45 (a). Multiple traumas to the membranes/fetus have resulted in death of one fetus with increased echogenicity of the fetal fluids and lack of heartbeat (b).
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Figure 220.11 Normal 48-day pregnancy on the left and resorption of fluids and the fetus on the right.
disruption of the remaining chorioallantois during surgery. Transvaginal ultrasound-guided fetal puncture Transvaginal ultrasound-guided fetal puncture for destruction of one of a set of twin pregnancies has been reported.34 Fetal fluids from one fetus were aspirated while observing the relationships of the needle fetus yolk sac and/or allantochorion between days 20 and 45. Three of four bicornuate twin pregnancies resulted in a single pregnancy 10 days or greater after interference (similar to or less than our ability to manually destroy one conceptus in this configuration). Three of nine (33%) still had a viable single pregnancy after 10 days when twins were fixed together (between days 20 and 45). These results were disappointing; however, they may be improved with experience and/or antibiotic therapy at the time of intervention. Ultrasonographic-guided fluid withdrawal between days 40 and 5035 or days 50 and 65 has been studied in
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single pregnancies;36 however, these studies did not involve any twins. Macpherson35 found that pregnancy reduction was more effective following aspiration of allantoic fluid (83%, 10/12), as compared to simple allantoic puncture (50%, 6/12; P = 0.06). Our experiences with transvaginal ultrasound-guided fetal reduction are small (N = 5); however, between 45 and 60 days the fetus within the vesicle was difficult to position. We only have attempted to directly puncture the fetus, not to aspirate fluid, and are unlikely to persevere with this technique (fetal puncture at this age) due to the difficulties involved. All cases ended in loss of both fetuses, usually within 3 days of interference. A more recent report suggested around 20% success with transvaginal procedures.37 Another report38 demonstrated a reduction of unicornuate twins using transvaginal ultrasoundguided aspiration between 16 and 25 days of gestation with a success rate of 70.0% (14 viable foals from 20 twin pregnancies) but when reduction was performed after day 40 of gestation, all mares (one unicornuate, three bicornuate) lost both. The technique of ultrasonographic-guided transvaginal aspiration of fetal fluids has been excellently described, along with results from their work and others.37 The authors’ conclusion was that this technique should be used before day 35. It is our belief that, although this technique may well be suitable to use then, it is possible to have similar or better results with the ultrasonographic manipulations described above, which has the added benefit of giving the mare a chance to apply her own natural twin reduction methods.
Days 60 to 100 Between days 60 and 100 it becomes more difficult to damage the chorioallantois. In these cases we
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Figure 220.12 Increased debris and echogenicity of the fetal fluids (a) are a good sign of fetal demise. Occasionally an increase in size of the amnionic cavity (b) is observed with fetal demise.
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Figure 220.13 Oscillation and dislocation of the fetus from the umbilicus is a technique that uses the weight of the fetus on a specific length of umbilicus to create a whiplash effect. The oscillation technique works exceptionally well with bilaterally fixed 45–50-day pregnancies and quite well in unilaterally fixed pregnancies when the one fetus can be isolated. (a) The fetus before oscillation. (b) The debris immediately after dislocation is presumed to be blood from the umbilicus. (c) The umbilical stump without the fetus. (d) The fetus lying on the floor of the uterus after dislocation.
identify the most conveniently located (always the smallest) of the twins and repeatedly traumatize it by oscillation, or membrane slip, or attempt to damage the cranium with multiple attempts of single digit percussion. Approximately 50% succumb to this procedure; however, it is tedious and time consuming. A technique of cervical dislocation has been described as useful during this time period.39, 40 It appears well suited to pregnancies from days 70 to 90. Two techniques were described: first, dislocation of fetal head per rectum, and second, via a standing flank laparotomy. The authors currently favor flank laparotomy (K. Wolfsdorf, personal communication, 2008). The technique is straightforward using standard standing flank laparotomy procedures. Immediately prior to grasping the fetus the mare is administered propantheline as a uterine relaxant. The chosen fetus
is grasped and then the head is removed or dislocated from the neck by pinching it between the thumb and first finger (Figure 220.14a). A distinct popping sensation is evident. In all cases, the fetus that has been manipulated has been alive immediately after and for 24 hours or more. Ultrasonography is useful in demonstrating that the head is well separated from the neck after the surgery (Figure 220.14b,c). This technique can be utilized earlier in gestation than transabdominal needle injection (after 100 days, see later in this section), potentially reducing prostaglandin-mediated inflammation and possibly enhancing the capacity of the placenta to expand within the uterus and support the remaining fetus. After an initial success rate reported of two live foals from four mares, the authors currently report a success rate overall of 64%, from 44 cases with the
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(c) Figure 220.14 (a) Technique for head dislocation. (b) Demonstration of correct response to dislocation. A blind ending stump at the end of the neck is clearly visible (arrow). (c) The dislocated head (arrow) can be seen well away from the vicinity of the neck of the same foal depicted in (b).
most recent figures of 8 foals from the last 12 procedures (K. Wolfsdorf, personal communication, 2008). We can report one foal from five attempts. One mare aborted at 3–4 months after having a discharge and three had no foals. One mare was determined to be negative a few weeks after the procedure. The twin pregnancy was only 60 days old and the procedure was associated with great difficulty grasping the foal as the uterus had started to contract and resulted in some trauma to the uterus. The heads separated easily and all cases had two live foals around 3–4 days after decapitation with a clear separation of head and neck. None aborted live twins to our knowledge. It is not known why in our hands such a simple technique did not meet the same results as others reported. It is possible that a more traumatic neck separation might
result in a quicker death of the manipulated fetus. We plan on continuing to work with the procedure.
Day 100 onwards Probably the most common reason for being presented mares at this late stage of gestation with twins is failure of the aforementioned techniques. Less frequently, twins have been missed in earlier diagnostic attempts and there has been an increase in diagnosis of twins at this stage due to the widespread use of fetal sexing procedures (R. Holder, personal communication). Frequently mares have been identified with twins late in the breeding season and the owner has adopted a non-intervention approach. Because the possibility of fetal reduction after 100 days
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Figure 220.15 (a) Twins at ∼ 110 days (transrectal). The ribs, lungs, liver, aorta, and abdominal contents can be clearly identified. (b) Transabdominal scan of largest twin facing right-hand side of screen. (c) Transabdominal scan of smaller twin facing to the left. The puncture guide is visible in both (b) and (c). (d) The needle has been advanced to the body wall of the fetus. (e) Immediately after injection of penicillin into the chest. (f) The injected twin showing signs of advanced mummification.
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Figure 220.16 (a) Membranes containing mummified bones. (b) The client was kind enough to arrange the bones which were collected at birth.
is very low, and the probability of abortion or stillbirth is extremely high, an approach was developed to eliminate one pregnancy at a later stage of gestation.41 The technique involved transabdominal ultrasonographic identification of the twins and intracardiac injection of a lethal substance. The smaller twin was always identified. Initial results with saline and air were unsuccessful, but when the solution was replaced with potassium chloride, 7/18 mares (39.9%) had single live foals. We have been utilizing this technique since 1988 and can report similar experiences. Our initial success was not very promising (2/10 live foals) until the potassium chloride solution was replaced with 10–20 mL of procaine penicillin10 which has resulted in a live foal rate of 56%. The current procedure at the Goulburn Valley Equine Hospital is to tranquilize the mare with detomidine42 and to identify the smaller fetus or, in the case of evenly sized fetuses, the one with more potential for placental expansion. A 6–10 inch, 16 needle with a tip designed for ultrasonographic enhancement (Cook, Australia) is passed through the needle guide biopsy channel into either the heart, lungs or abdomen of the identified fetus (Figure 220.15a–f). Penicillin is injected and the fetus monitored for the next 5 to 10 minutes. If the needle delivers penicillin in the chest or abdomen the demise of the fetus still occurs; however, it just takes a little longer (up to 5 minutes). The apparent advantages of penicillin as we see them are: (i) it reduces iatrogenic bacterial contamination; (ii) it can be visualized ultrasonographically as it is injected; and (iii) fetal death can still be obtained without intracardiac needle placement. R We have attempted to place mares on Regumate and long-term oral antibiotics; however, we have found no difference in fetal survival rates with either treatment compared to those that have received
no treatment. At the time of needle puncture, mares are treated with systemic antibiotics for 3 days and intravenous phenylbutazone. There has been much discussion about the production of small dysmature foals from successful use of this technique. While this occasionally does occur, care in correct fetus identification (retaining the biggest fetus with the most potential for placental expansion) has lowered the occurrence of this outcome dramatically. This is more readily possible when there have been multiple early opportunities to observe fetal size, position with the pregnant horn(s), and uterine body and placental arrangements. After injection and death of the fetus it gradually becomes mummified. Commonly these fetal remnants can be detected within the placenta at birth (Figure 220.16a,b). Late in gestation, twins may be difficult to recognize using transrectal ultrasonography, except in those cases with observation of the twin membrane.43 Abdominal ultrasonography is useful to diagnose twins.41, 44 In the event of failure to diagnose twin pregnancies, occasionally abortion is heralded by lactation late in gestation (> 7 months). There are reports of successful maintenance of pregnancy despite premature lactation (> 1 month prior to foaling) in cases with twin gestation. Apparently the premature lactation is induced by fetal death and the beginning of mummification of one fetus, and thus is threatening to the remaining live fetus. Four mares with apparent impending twin abortion were able to deliver live single foals concurrent with a mummified twin after supplementation with progesterone was initiated upon recognition of inappropriate lactation late in gestation.45, 46 In these cases, although foals were born small, they survived and thrived normally. Further work will be necessary to determine which
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mares will respond best, or even at all, to supplementation with progesterone for initiation of premature lactation.
Induction of abortion Occasionally, owners elect a wait-and-see technique. This is often due to financial constraints. When twin pregnancies are allowed to proceed to foaling or abortion, there is a significant risk that the mare will develop foaling problems or have difficulty being rebred. After 100 days of gestation, natural fetal reduction is not reliable enough to let the mare continue the pregnancy. Multiple injections of prostaglandin should be used to induce abortion as soon after 100 days as possible (see Chapter 238). It is not considered advisable for any reason other than finances to induce abortion in earlier twin pregnancies. Many veterinarians at the time period of 30–35 days administer prostaglandins to abort the mare rather than allow endometrial cup formation and risk missing the opportunity to rebreed the mare in the same season. However, we encourage intervention through manual trauma to the placental membranes, because this can result in good outcomes, and manipulations can continue for a few days until the result is clear or destruction of both has occurred (see earlier section). From all of the preceding discussion it should be obvious to readers that in our opinion the best method of handling twins is early identification and destruction of one twin during the pre-fixation mobility phase (days 11 to 16).
References 1. Acland HM. Abortion in mares: diagnosis and prevention. Compend Cont Educ Pract Vet 1987;9:318–26. 2. Acland HM. Abortion in mares. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 554–62. 3. Jeffcott LB, Whitwell K. Twinning as a cause of foetal and neonatal loss in the Thoroughbred mare. J Comp Pathol 1973;83:91–106. 4. Kaur R, Sharma JK. Epidemiologic studies on abortions in Indian Thoroughbred mares. J Equine Vet Sci 1996;16:508–10. 5. Platt H. Aetiological aspects of abortion in the Thoroughbred mare. J Comp Pathol 1973;83:199–205. 6. Hong CB, Donahue JM, Giles RC, Petritesmurphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Equine abortion and stillbirth in Central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. 7. Ricketts SW, Barrelet A, Whitwell KE. A review of the causes of abortion in UK mares and means of diagnosis used in an equine studfarm practice in Newmarket. Pferdeheilkunde 2001;17(6):589–92.
8. Merkt H, Jochle W. Abortions and twin pregnancies in Thoroughbreds: Rate of occurence, treatment and prevention. J Equine Vet Sci 1993;13:690–4. 9. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects. Cross Plains, WI: Equiservices, 1992. 10. McKinnon AO, Rantanen NW. Twins. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Wilkins, 1998; pp. 141–56. 11. Ginther OJ. The nature of embryo reduction in mares with twin conceptuses: Deprivation hypothesis. Am J Vet Res 1989;50:45–53. 12. Ginther OJ. Twin embryos in mares. I: From ovulation to fixation. Equine Vet J 1989;21:166–70. 13. Carnevale EM, Ginther OJ. Relationships of age to uterine function and reproductive efficiency in mares. Theriogenology 1992;37:1101–15. 14. Ginther OJ. The twinning problem: From breeding to day 16. In: Proceedings of American Association of Equine Practitioners, 1983; pp. 11–26. 15. Pascoe RR. A possible new treatment for twin pregnancy in the mare. Equine Vet J 1979;15:40–2. 16. Roberts CJ. Termination of twin gestation by blastocyst crush in the broodmare. J Reprod Fertil Suppl 1982;32:447–9. 17. Pascoe DR, Pascoe RR, Hughes JP, Stabenfeldt GH, Kindahl H. Comparison of two techniques and three hormone therapies for management of twin conceptuses by manual embryonic reduction. J Reprod Fertil Suppl 1987;35:701–2. 18. Pascoe DR, Pascoe RR, Hughes JP, Stabenfeldt GH, Kindahl H. Management of twin conceptuses by manual embryonic reduction: comparison of two techniques and three hormone treatments. Am J Vet Res 1987;48:1594–9. Erratum: Am J Vet Res 1988;49(2):273. 19. McKinnon AO, Figueroa STD, Nobelius AM, Hyland JH, Vasey JR. Failure of hydroxyprogesterone caproate to maintain pregnancy in ovariectomized mares. Equine Vet J 1993;25:158–60. 20. Bowman T. Ultrasonic diagnosis and management of early twins in the mare. In: Proceedings of American Association of Equine Practitioners, 1986; pp. 35–43. 21. McKinnon AO, Squires EL, Pickett BW. Equine Diagnostic Ultrasonography. Animal Reproduction Laboratory, Colorado State University, 1988. 22. McKinnon AO. Reproductive efficiency of horses in Australia. http://www.gvequine.com.au/breeding efficiency.htm, 2000. 23. Nath LC, Anderson GA, McKinnon AO. Reproductive efficiency of Thoroughbred and Standardbred horses in northeast Victoria. Austr Vet J 2010;88:169–75. 24. Ginther OJ, Douglas RH, Woods GL. A biological embryoreduction mechanism for elimination of excess embryos in mares. Theriogenology 1982;18:475–85. 25. Ginther OJ. Postfixation embryo reduction in unilateral and bilateral twins in mares. Theriogenology 1984;22:213–23. 26. Baucus KL, Squires EL, Morris R, McKinnon AO. The effect of stage of gestation and frequency of prostaglandin injection on induction of abortion in mares. In: Proceedings of Equine Nutritional and Physiological Society, 1987; pp. 255–8. 27. Squires EL, Bosu WTK. Induction of abortion during early to mid gestation. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, London: Lea & Febiger, 1993; pp. 563–6. 28. Penzhorn BL, Bertschinger HJ, Coubrough RI. Reconception of mares following termination of pregnancy with prostaglandinF2 alpha before and after day 35 of pregnancy. Equine Vet J 1986;18:215–17.
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Management of Twins 29. Pascoe RR. Methods for the treatment of twin pregnancy in the mare. Equine Vet J 1983;15:40–2. 30. Hyland JH, Maclean AA, Robertson-Smith GR, Jeffcott LB, Stewart GA. Attempted conversion of twin to singleton pregnancy in two mares with associated changes in plasma oestrone sulphate concentrations. Austr Vet J 1985;62:406– 9. 31. Allen WR, Bracher V. Videoendoscopic evaluation of the mare’s uterus: III. Findings in thepregnant mare [see comments]. Equine Vet J 1992;24:285–91. 32. Pascoe DR, Stover SM. Surgical removal of one conceptus from fifteen mares with twin concepti. Vet Surg 1989;18:141–5. 33. Stover SM, Pascoe DR. Surgical removal of one conceptus from the mare with a twin pregnancy. Vet Surg 1987;16: 87. 34. Bracher V, Parlevliet JM, Pieterse MC, Vos PLAM, Wiemer P, Taverne MAM, Colenbrander B. Transvaginal Ultrasound-guided twin reduction in the mare. Vet Rec 1993;133:478–9. 35. Macpherson ML, Homco LD, Varner DD, Blanchard TL, Harms PG, Flanagan MN, Forrest DF. Transvaginal ultrasound-guided allocentesis for pregnancy elimination in the mare. Biol Reprod Mono 1995;1:215–23. 36. Squires EL, Tarr SF, Shideler RK, Cook NL. Use of transvaginal ultrasound-guided puncture for elimination of equine pregnancies during days 50 to 65. J Equine Vet Sci 1994;14:203–5. 37. Macpherson ML, Reimer JM. Twin reduction in the mare: current options. Anim Reprod Sci 2000;60:233–44.
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38. Mari G, Iacono E, Merlo B, Castagnetti C. Reduction of twin pregnancy in the mare by transvaginal ultrasound-guided aspiration. Reprod Domest Anim 2004;39:434–7. 39. Wolfsdorf K, Rodgerson DH, Holder R. How to manually reduce twins between 60 and 120 days gestation using cranio-cervical dislocation. Proceedings of the American Association of Equine Practitioners, 2005;51:284–7. 40. Wolfsdorf KE. Management of postfixation twins in mares. Vet Clin North Am Equine Pract 2006;22:713–25. 41. Rantanen NW, Kincaid B. Ultrasound guided fetal cardiac puncture: A method of twin reduction in the mare. Proceedings of the American Association of Equine Practitioners, 1988; pp. 173–9. 42. McKinnon AO, Carnevale EM, Squires EL, Jochle W. Clinical evaluation of detomidine hydrocholoride for equine reproductive surgery. Proceedings of the American Association of Equine Practitioners, 1988; pp. 563–8. 43. Ginther OJ, Griffin PG. Natural outcome and ultrasonic identification of equine fetal twins. Theriogenology 1994;41:1193–9. 44. Pipers FS, Adams-Brendemuehl CS. Techniques and applications of transabdominal ultrasonography in the pregnant mare. J Am Vet Med Assoc 1984;185:766–71. 45. Roberts SJ, Myhre G. A review of twinning in horses and the possible therapeutic value of supplemental progesterone to prevent abortion of equine twin fetuses the latter half of the gestation period. Cornell Vet 1983;73:257–64. 46. Shideler RK. Prenatal lactation associated with twin pregnancy in the mare: A case report. J Equine Vet Sci 1987;7:383–4.
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Chapter 221
Early Embryonic Loss Dirk K. Vanderwall
Early embryonic loss in the mare is generally defined as pregnancy failure that occurs up to day 40 of gestation, which corresponds to the time of transition from the embryo stage to the fetal stage of conceptus development.1 The diagnosis of early embryonic loss and recognition of factors contributing to its occurrence have been dramatically improved by the widespread use of transrectal ultrasonography for early pregnancy diagnosis. Under field conditions, transrectal ultrasonography is routinely used for pregnancy diagnosis as early as days 12–14 post ovulation, whereas under experimental conditions it may be used as early as days 10 or 11; therefore, ultrasonography allows direct (and repeatable) assessment of the conceptus during approximately three-quarters of the interval when early embryonic loss occurs. Before days 10 or 11, the conceptus is too small to be visualized with standard ultrasonographic equipment; therefore, other techniques have been used to study embryonic loss during that interval. Specifically, assisted reproductive techniques, such as embryo transfer, transvaginal ultrasoundguided follicle aspiration (TVA), and oocyte transfer, and experimental techniques, such as in vitro embryo culture and light/electron microscopy of oocytes/ embryos, have been used to study early embryonic development and embryonic loss prior to days 10 or 11. The incidence and etiological factors contributing to early embryonic loss in mares are discussed in Chapter 239. Briefly, based on serial examination with transrectal ultrasonography the incidence of embryonic loss is generally in the range of 5–15%; however, the highest embryonic loss rates (20–30% or higher) are detected in mares over 18 years of age.2 At those levels, early embryonic failure represents a considerable economic loss to the equine industry in the form of increased costs associated with additional breeding of mares and/or decreased foal production.
Diagnosis Early embryonic loss may or may not be accompanied by premonitory signs of impending failure of the pregnancy. Prior to development of the embryo proper, which usually becomes evident with transrectal ultrasonography by days 21 or 22 of gestation, pregnancy loss is generally characterized by the sudden disappearance of the embryonic vesicle between examinations. After development of the embryo proper, signs of impending pregnancy loss are more likely to be observed. Signs of impending embryonic loss detectable with transrectal ultrasonography include irregular shape of an embryonic vesicle, prolonged mobility of a vesicle (i.e., beyond day 16), excessive endometrial edema, an undersized vesicle (Figure 221.1), lack of development of the embryo proper (i.e., developing as a trophoblastic vesicle; Figure 221.2), loss of embryonic heartbeat (Figure 221.3), dislodgement of a vesicle with loss of fluid, increased echogenicity of fluid within the conceptus and abnormal development of the embryonic membranes (Figure 221.4).3 Clinical signs such as these have been observed in association with spontaneous pregnancy loss,4 experimentally induced pregnancy loss following ovariectomy or administration of prostaglandin F2α (PGF2α ),5 and more recently in pregnancy losses associated with newer assisted reproductive techniques such as cloning, which is characterized by an extremely high rate (> 80%) of early embryonic loss.6 Some pregnancies may exhibit more than one sign of impending failure; for example, conceptuses that were small for their gestational age were 50 times more likely to develop as a trophoblastic vesicle without an embryo proper than were normal-sized conceptuses.7 Another ultrasonographically detectable sign that may be indicative of an increased likelihood of pregnancy failure is the identification of an otherwise
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Figure 221.1 Transrectal ultrasonogram of a cloned mule conceptus on day 18 of gestation. The vesicle was 13 mm in diameter, which was small for its gestational age. The conceptus continued to develop until day 27 and was then spontaneously eliminated. Each division along the top and left margins equals 10 mm. Reprinted with permission from Vanderwall et al.6
normal-appearing conceptus that has become fixed in the uterine body instead of in the base of one of the uterine horns. In a recent retrospective analysis of clinical breeding records, the outcome of 30 ‘body pregnancies’ was determined.8 Nine of the 30 body pregnancies were in the cranial uterine body and the remaining 21 pregnancies were in the caudal 6 cm of the uterine body near the cervix (Figure 221.5). Of the nine cranial body pregnancies, seven were carried to term, one mare lost her pregnancy between days 35 and 42, and one mare aborted at 9 months of gestation. Of the 21 pregnancies located in the caudal body, 18 were monitored to follow the spontaneous outcome. Of those 18 caudal body pregnancies, only three (17%) were successfully carried to term; in contrast, 13 (72%) of the caudal body pregnancies were spontaneously lost by day 43, and one was lost at 4 months and another at 9 months of gestation. Although fixation in the cranial uterine body did not seem to alter early embryonic development [only one of nine (11%) conceptuses lost during early gestation], fixation in the caudal uterine body was associated with an extremely high rate of early embryonic loss as noted above. Because of the low probability
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Figure 221.2 Transrectal ultrasonogram of a cloned mule conceptus on day 30 of gestation. The vesicle was 28 mm in diameter (normal size), but the conceptus was apparently developing as a trophoblastic vesicle since there was no evidence of an embryo proper. The conceptus was not present when the mare was examined again 5 days later. Each division along the top and left margins equals 10 mm. Reprinted with permission from Vanderwall et al.6
of a successful outcome with the caudal body pregnancies, three of the caudal body pregnancies were terminated with administration of PGF2α .
Management In conjunction with recognizing signs of impending failure is the need to assess whether there are management changes or treatments that can be initiated to decrease the incidence of early embryonic loss and/or ‘rescue’ conceptuses showing signs indicative of impending failure. First and foremost, all broodmares should receive proper management to ensure adequate nutrition to maintain optimal body condition; appropriate vaccination/deworming and other routine healthcare such as dental prophylaxis; and, to the degree possible, efforts should be made to minimize stressful situations for mares, for example social stress due to introduction of new herd mates, competition for food sources, etc. Optimal management in conjunction with specific therapy aimed at resolving treatable reproductive abnormalities (e.g.,
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(a)
Figure 221.3 Transrectal ultrasonogram of a cloned mule conceptus on day 41 of gestation. The vesicle was 39 mm in diameter (measured in a different plane) and the embryo proper was evident; however, an embryonic heartbeat was not detected. The non-viable conceptus was recovered via transcervical lavage 7 days later for subsequent analysis. Each division along the top and left margins equals 10 mm.
poor perineal conformation, uterine infection) will increase the likelihood that mares will successfully maintain a pregnancy.9 Prophylactic administration of exogenous progesterone to pregnant mares in an effort to enhance maintenance of pregnancy is a widespread practice. Conclusive evidence that progesterone supplementation can help ‘rescue’ a conceptus that will otherwise fail is lacking; however, the use of exogenous progesterone may be warranted when faced with a conceptus that is ‘small-for-date’, since it is plausible that an undersized conceptus may be less likely to prevent luteolysis and maintain corpora luteal function. In addition, administration of exogenous progesterone is clearly indicated when there is clinical evidence, or suspicion, of systemic endotoxemia due to severe disease, since endotoxin has a deleterious effect on corpora luteal function (mediated through PGF2α release), which may lead to pregnancy failure due to insufficient progesterone levels.10 In pregnant mares, the fetal–placental unit begins secreting progesterone and related progestagens between days 80 and 100 of gestation, the levels of which become
(b)
Figure 221.4 Transrectal ultrasonograms of a cloned horse conceptus on day 14 of gestation (a) and the same conceptus on day 42 of gestation exhibiting disorganization of the conceptus membranes (b). Although an embryo proper was evident (b), cardiac activity was not detected. Remnants of the conceptus were retained in the uterine lumen until day 66 of gestation. Each division along the top and left margins equals 10 mm. Reprinted with permission from Vanderwall et al.6
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CAUDAL BODY
CERVIX
R08
R70 1
C3
2
3
4
Figure 221.5 Transrectal ultrasonogram of a conceptus on day 22 of gestation located in the caudal uterine body near the cervix. Reprinted with permission from Jobert et al.8
sufficient to maintain pregnancy after day 100; therefore, if progesterone therapy is instituted, it is generally discontinued between days 100 and 120 because there is no physiological basis for continued administration of exogenous progesterone after that time. Currently, there are no commercially available formulations of progesterone labeled for use in pregnant mares; therefore, the use of exogenous progesterone for maintenance of pregnancy is an ‘extra-label’ use. Two preparations of progesterone commonly administered to pregnant mares are daily oral administration of 0.044 mg/kg (1 mL per 110 pounds body weight) of the synthetic progestin altrenogest (ReguMate; Intervet, Inc., Millsboro, DE) or daily intramuscular administration of 0.2–0.3 mg/kg (100–150 mg total dose) progesterone in oil (available from compounding pharmacies). Although both treatments have been shown to be efficacious by their ability to maintain pregnancy in mares without an endogenous source of progesterone, the need for daily administration is a drawback of these formulations. In an attempt to eliminate the need for daily administration of exogenous progesterone preparations, there has been interest in the use of long-acting formulations of progesterone that can be administered at weekly (or longer) intervals. There are anecdotal
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reports of the clinical use of long-acting synthetic progestins labeled for use in women [e.g., medroxR ]) or 17-alphayprogesterone acetate (Depo-Provera hydroxyprogesterone hexanoate] for maintenance of pregnancy in mares; however, these formulations were unable to maintain pregnancy in mares without an endogenous source of progesterone.11, 12 It was recently reported that intramuscular administration of a compounded long-acting formulation containing 1.5 g progesterone every 7 days provided a sufficient level of progesterone to maintain pregnancy in mares that lacked an endogenous source of progesterone; therefore, this compounded formulation of progesterone appears to be an efficacious and suitable alternative to progesterone formulations that require daily administration.13 (See also Chapters 189 and 198). An important aspect of the clinical management of early embryonic loss is recognition that it will inevitably occur in some mares, particularly aged mares. An appropriate course of action is to diagnose its occurrence as early as possible in an effort to determine whether there were specific factors associated with the pregnancy loss and to provide an opportunity for rebreeding the mare as soon as possible, if the pregnancy loss occurs early enough in the breeding season. Early detection of embryonic loss can be accomplished by performing serial examinations with transrectal ultrasonography every 10 days to 2 weeks between days 14 and 40 of gestation. Clinically, serial examinations are routinely conducted through day 60 of gestation, at which time the frequency of examination is reduced, which is subsequently followed by a single ‘fall’ check. At each examination a complete assessment of the conceptus should be performed and recorded (i.e., size, location, presence of embryonic heartbeat, etc.); if the conceptus shows any signs of impending failure, the pregnancy can be monitored more closely, and a decision about the use of progesterone supplementation can be made as discussed above. Although in many cases it may be warranted to attempt to ‘rescue’ a conceptus displaying signs of impending pregnancy loss, there are times when an abnormal pregnancy may be selectively terminated. Two circumstances when it may be appropriate to actively terminate a pregnancy are when a conceptus is developing as a trophoblastic vesicle or when a conceptus becomes fixed in the caudal uterine body near the cervix as described previously. In the case of a trophoblastic vesicle, there is no potential for a viable outcome of the pregnancy, whereas in the situation of a caudal body pregnancy the likelihood of a viable outcome to the pregnancy is only 17%.8 Therefore, actively terminating the pregnancy in these situations may be indicated in order to facilitate rebreeding
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the mare in the same breeding season. Actively terminating an abnormal pregnancy is performed by administering a luteolytic dose of PGF2α alone or in combination with manual transrectal reduction of the conceptus. Although PGF2α treatment alone will reliably eliminate a conceptus during early gestation, on average complete loss of the conceptus takes more than 1 week when PGF2α is administered on day 30.5 By disrupting the embryonic vesicle, manual reduction may hasten complete elimination of the conceptus, although the length of time that conceptus fragments persist after manual reduction is unknown. Active termination is the option of choice if rebreeding is desired, and should be performed before day 35 of gestation to prevent the development of endometrial cups, which start to form and secrete equine chorionic gonadotropin (eCG) at that time.14 Terminating the pregnancy before the endometrial cups become functional is important, because, when a pregnancy is terminated after the cups become functional, many mares fail to resume normal cyclic activity, and frequently exhibit prolonged anestrus, anovulatory estrus, or ovulation unaccompanied by estrus.15
Summary An important aspect of the clinical management of early embryonic loss in the mare is to recognize that it will inevitably occur in some mares, and that the highest loss rates occur in aged mares. In general, the most appropriate course of action is to diagnose the occurrence of embryonic loss as early as possible in order to provide an opportunity for rebreeding the mare as soon as possible (if the loss occurs during the breeding season). Early detection of embryonic loss can best be accomplished by performing serial examinations with transrectal ultrasonography every 10 days to 2 weeks during early gestation; if a conceptus shows any signs of impending failure, the pregnancy can be monitored more closely. In some situations, active termination of a pregnancy demonstrating signs of impending failure may be warranted to facilitate rebreeding.
References 1. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992; pp. 345–6. 2. Vanderwall DK. Early embryonic loss in the mare. J Eq Vet Sci 2008;28:691–702. 3. Ball BA, Woods GL. Embryonic loss and early pregnancy loss in the mare. Compend Contin Educ Pract Vet 1987;9:459–70. 4. Ginther OJ, Bergfelt DR, Leith GS, Scraba ST. Embryonic loss in mares: incidence and ultrasonic morphology. Theriogenology 1985;24:73–86. 5. Ginther OJ. Embryonic loss in mares: nature of loss after experimental induction by ovariectomy or prostaglandin F2a . Theriogenology 1985;24:87–98. 6. Vanderwall DK, Woods GL, Roser JF, Schlafer DH, Sellon DC, Tester DF, White KL. Equine cloning: applications and outcomes. Reprod Fertil Dev 2006;18:91–8. 7. Vanderwall DK, Squires EL, Brinsko SP, McCue PM. Diagnosis and management of abnormal embryonic development characterized by formation of an embryonic vesicle without an embryo in mares. J Am Vet Med Assoc 2000;217:58–63. 8. Jobert ML, LeBlanc MM, Pierce SW. Pregnancy loss rate in equine uterine body pregnancies. Equine Vet Educ 2005;17:163–5. 9. Darenius K, Kindahl H, Madej A. Clinical and endocrine studies in mares with known history of repeated conceptus losses. Theriogenology 1988;29:1215–32. 10. Daels PF, Starr M, Kindahl H, Fredriksson G, Hughes JP, Stabenfeldt GH. Effect of Salmonella typhimurium endotoxin on PGF-2a release and fetal death in the mare. J Reprod Fertil Suppl 1987;35:485–92. 11. McKinnon AO, Tarrida Del Marmol Figueroa S, Nobelius AM, Hyland JH, Vasey JR. Failure of hydroxyprogesterone caproate to maintain pregnancy in ovariectomized mares. Equine Vet J 1993;25:158–60. 12. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32:83–5. 13. Vanderwall DK, Marquardt JL, Woods GL. Use of a compounded long-acting progesterone formulation for equine pregnancy maintenance. J Equine Vet Sci 2007;27:62–6. 14. Allen WR, Moor RM. The origin of the equine endometrial cups. I. Production of PMSG by fetal trophoblast cells. J Reprod Fertil 1972;29:313–16. 15. Squires EL, Hillman RB, Pickett BW, Nett TM. Induction of abortion in mares with Equimate: effect on secretion of progesterone, PMSG and reproductive performance. J Anim Sci 1980;50:490–5.
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Chapter 222
Ovarian Abnormalities Patrick M. McCue and Angus O. McKinnon
Ultrasonography of the ovary per rectum may be used in conjunction with behavioral observations, physical examination, and hormone analysis, along with ancillary procedures such as ovarian biopsy and karyotyping in the diagnosis and differentiation of ovarian abnormalities. A majority of ovarian abnormalities can be diagnosed on ultrasonographic examination alone, while some abnormalities may require a more extensive evaluation. Pathological conditions of the equine ovary can be divided into abnormalities of follicular development, ovulation, or luteal function, ovarian tumors, and miscellaneous other abnormalities.
Abnormalities of follicular development Alterations of follicular development may be associated with a variety of physiological and pathological conditions. A list of potential causes of abnormal follicular development is presented in Table 222.1. Ovarian senescence or age-related ovarian dysfunction Ovarian dysfunction has been identified as a cause of reduced fertility in mares approximately 20 years of age or older. Ovarian issues in older mares include a longer follicular phase,1 prolonged interovulatory period,2, 3 and delay in the first ovulation of the year. Complete failure of follicular development or ovarian senescence has been observed in aged mares and may be due to an insufficient number of primordial follicles. No effective treatments are available for promoting follicular growth in senescent ovaries.
opment may be limited to one or two small (< 10 mm) follicles on each ovary (Figure 222.1) or follicles may not be visible by ultrasonography. Subsequent examinations confirm the absence of follicular growth. Postpartum (‘lactational’) anestrus A majority of mares develop follicles and ovulate early in the postpartum period (i.e., foal heat ovulation) and continue to cycle if they are not bred or do not become pregnant. However, some mares may exhibit a temporary failure of follicular development or ovulation after foaling.4 Affected mares may remain anestrus or anovulatory for weeks or months before cyclic ovarian activity is initiated. Mares that foal early in the year are more prone to exhibit postpartum anestrus than mares foaling later in the spring or in the summer. ‘Lactational anestrus’ may be caused by the combined effects of lactation, body condition, and season. In addition, mares may present as failure to develop any follicular activity after foaling, failure to ovulate after some follicular development or failure to cycle after first developing, and ovulating a ‘foal heat’ follicle.
Ultrasonographic characteristics Examination of a mare in postpartum anestrus may reveal ovarian follicular activity ranging from inactive ovaries that resemble ovaries of a mare in deep seasonal anestrus (i.e., a few small follicles < 15 mm in diameter) to ovaries with moderate follicular activity that resemble ovaries of a mare in spring transition (i.e., multiple 20- to 30-mm follicles) (Figure 222.2).
Ultrasonographic characteristics
Chromosomal abnormalities
Palpation and ultrasonography of the ovaries per rectum reveal bilaterally small ovaries. Follicular devel-
A major sex chromosomal abnormality may be suspected in a mare of breeding age with primary
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Table 222.1 Potential causes of abnormal follicular development separated by conditions that are transient, physiological, and/or treatable versus conditions that are permanent, pathological, and/or non-treatable Transient or physiological
Permanent or pathological
Seasonal anestrus
Chromosomal abnormalities
Inadequate nutrition
Ovarian senescence (advanced age)
Postpartum or lactational anestrus Equine Cushing’s disease Exogenous hormone therapy GnRH vaccine administration Persistent endometrial cups Granulosa cell tumor on contralateral ovary
GnRH, gonadotropin-releasing hormone.
Figure 222.2 Ultrasonographic image of an ovary in an 11year-old Quarter Horse mare with postpartum anestrus. The mare developed a follicular wave in the early postpartum period that did not culminate in ovulation and subsequently became anestrus.
Ultrasonographic characteristics infertility and gonadal hypoplasia. The most commonly reported chromosomal abnormality of the horse is 63,X gonadal dysgenesis, in which only a single sex chromosome is present.5 Horses with gonadal dysgenesis develop as phenotypic females because of the absence of a Y chromosome. Affected horses are sometimes small in stature and have bilaterally small ovaries, a small flaccid uterus, and endometrial gland hypoplasia. Numerous other chromosomal abnormalities have also been reported in the mare,6 including translocations of non-sex chromosomes with normal cyclicity and poor fertility.7
Palpation per rectum reveals a pair of very small ovaries (i.e., 1.0–1.5 cm in length). Visible follicles are usually absent on ultrasonographic examination, but occasionally a small follicle < 10 mm in diameter may be observed (Figure 222.3). A presumptive diagnosis of chromosomal abnormality may be made only if all other causes of failure of follicular development have been ruled out. Definitive diagnosis of a chromosomal abnormality is based on chromosome analysis or karyotyping. Pituitary pars intermedia dysfunction (PPID) (equine Cushing’s syndrome) Mares with hypertrophy, hyperplasia, or adenoma formation in the pars intermedia of the pituitary
Figure 222.1 Ultrasonographic image of an ovary in a 28-yearold Arabian mare with ovarian senescence. Only one small follicle is visible.
Figure 222.3 Ultrasonographic image of an ovary in a mare with 63,X gonadal dysgenesis.
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Figure 222.4 A mare with equine Cushing’s disease.
(PPID) have been reported to have abnormal estrous cycles and reduced fertility.8, 9 A majority of horses diagnosed with PPID are older, with the average age being approximately 20 years. Clinical signs of PPID include hirsutism (Figure 222.4) and abnormal haircoat shedding patterns, polyuria, polydipsia, and hyperhidrosis.10 Potential causes of reproductive abnormalities in mares with PPID include an increased production of glucocorticoids and androgens from the adrenal cortex and destruction of gonadotrophs owing to compression of the anterior pituitary by the enlarged pars intermedia.11 It is presumed that a decrease in pituitary gonadotropin secretion is responsible for the reduction in ovarian follicular development.
Ultrasonographic characteristics Mares with PPID may have relatively normal estrous cycles and ultrasonographic examination may reveal significant follicular activity and/or a normal corpus luteum. However, in more advanced stages, follicular activity may be minimal or absent (Figure 222.5). Exogenous hormone treatment Administration of anabolic steroids, dexamethasone, estradiol esters, and a combination of progesterone plus estradiol have all been reported to suppress pituitary gonadotropin secretion and ultimately result in an inhibition of ovarian follicular activity.12–15 Potent agonists of GnRH, such as deslorelin acetate, have been demonstrated to cause downregulation of pituitary gonadotropin secretion and short-term alterations in ovarian function.16, 17
Ultrasonographic characteristics Mares may exhibit a suppression of follicular activity visible by ultrasonography characterized by an absence of follicles greater than 20 mm in diameter.
Figure 222.5 Ultrasonography of an ovary in a mare with equine Cushing’s disease.
GnRH vaccine administration Active immunization of mares against GnRH results in suppression of pituitary LH secretion18 and a subsequent reduction in ovarian follicular activity, suppression of estrus, and decreased fertility that is related to the magnitude and duration of antibody titer.18–23 In most instances, the effect is reversible over time as antibody titers wane and vaccinated mares eventually resume ovarian function and return to normal fertility.20, 22, 23 However, a recent report indicated that some mares immunized against GnRH have failed to have recrudescence to cyclicity even after several years.24
Ultrasonographic characteristics Mares have limited ovarian follicular activity during the period of high antibody titers following immunization against GnRH; however, in general, the duration of ovarian inactivity (> 6 months) persists well after the common time of antibody detection (< 90 days). The ultrasonographic image will be similar to that of a mare in seasonal anestrus. Persistent endometrial cups The presence of active, functional endometrial cups in a non-pregnant mare may result in complete
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Granulosa cell tumor on contralateral ovary
Figure 222.6 Ultrasonography of persistent endometrial cups in the uterus of a mare. Black arrow points to the endometrial cups.
suppression of ovarian follicular activity, or more commonly may result in follicular development and subsequent luteinization without ovulation (accessory corpora lutea like).25–27 Persistence of endometrial cups beyond their expected lifetime of 60–100 days has been reported in mares that sustained pregnancy loss as well as mares that carried to term and foaled normally.25, 27
Granulosa cell tumors (GCTs) produce inhibin and androgens that suppress pituitary FSH secretion and ultimately result in a small, inactive contralateral ovary.28 Surgical removal of the neoplastic ovary removes the source of pituitary suppression (Figure 222.7). The opposite ovary will usually resume follicular development and ovulation will often occur approximately 6–8 months after tumor removal, but the exact time will vary with individual mares and also the time of ovarian removal relative to seasonal effects on the ovary. In rare instances, a mare may not develop follicles for several years after removal of a GCT. In addition, mares with a GCT on one ovary and a functional contralateral ovary have been reported.29
Ultrasonographic characteristics Palpation and ultrasonographic examination of the ovary contralateral to the GCT will reveal a small ovary with limited to no follicular development (Figure 222.8).
Ultrasonographic characteristics
Idiopathic failure of follicular development
Ovarian ultrasonography may reveal inactive ovaries with minimal follicular activity (i.e. < 15-mm follicles), ovaries that appear normal, or, most commonly, mares with persistent endometrial cups repeatedly develop hemorrhagic anovulatory follicles (also called luteinized unruptured follicles) or accessory corpora lutea. Careful ultrasonographic examination of the uterus may reveal the persistent endometrial cups as one or more small echogenic areas at the base of a uterine horn (Figure 222.6). Confirmation is based on concentrations of equine chorionic
Occasionally, mares fail to exhibit significant follicular growth during the physiological breeding season for undetermined reasons. Idiopathic failure of follicular development is a default diagnosis made after all other potential causes are ruled out.
Ultrasonographic characteristics Inactive ovaries with follicles generally < 15 mm in diameter (Figure 222.9).
GTCT removal
1.2 1.0 0.8 Inhibin
c222
Inhibin
0.6 0.4 0.2 0 0
100
200
300
400
500
600
Time Figure 222.7 Inhibin levels in a mare following surgical removal of a granulosa–theca cell tumor.30
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Table 222.2 Abnormalities of the periovulatory period, separated into events that occur in the pre-ovulatory follicle and an event that occurs following ovulation Events in the pre-ovulatory follicle
Events after ovulation
Echogenic ovulatory follicle
Ovarian hematoma
Persistent anovulatory follicle Partially collapsed anovulatory follicle Hemorrhagic or luteinized anovulatory follicle
Figure 222.8 Ultrasonography of the ovary contralateral to a granulosa cell tumor.
pregnancy rate may be associated with degeneration of the oocyte if the echogenic follicle was atretic or entrapment of the oocyte within the follicle if ovulation was not normal.
Abnormalities of ovulation Abnormalities of the periovulatory period are relatively common in the mare (Table 222.2), with the most frequent condition being the hemorrhagic or luteinized anovulatory follicle. Echogenic ovulatory follicle Echogenic particles or spots were reported to occur within the follicular fluid of 6.2% of dominant follicles that ovulated in a retrospective study involving 721 mares and a total of 1694 estrous cycles.31 Pregnancy rate per cycle was higher following ovulation of a non-echogenic follicle (42.0%) than after ovulation of an echogenic follicle (29.4%). The echogenic particles may be granulosa cells shed from the follicular wall or red blood cells and/or fibrin associated with pre-ovulatory hemorrhage. The decrease in
Figure 222.9 Ultrasonographic image of the left ovary of a 13year-old American Quarter Horse mare with a 3-year history of idiopathic failure of follicular development.
Ultrasonographic characteristics Detection of a few small echogenic particles or spots within the follicular fluid of a follicle destined to ovulate (Figure 222.10) is more common than observation of multiple echogenic particles (Figure 222.11). It is usually not possible to predict whether a follicle containing multiple echogenic particles will ovulate when the particles are first detected. The clinical
Figure 222.10 Ultrasonographic image of a follicle with a few echogenic particles in the lumen prior to ovulation.
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Figure 222.12 Ultrasonographic image of a non-luteinized persistent anovulatory follicle.
respond to surgical drainage after a prolonged period of persistence (needle through paralumbar fossa or ultrasonography guided per vagina).
Figure 222.11 Ultrasonographic image of a follicle with multiple echogenic particles in the follicular lumen prior to ovulation.
significance is that follicles with multiple echogenic particles in the follicular lumen may not ovulate and ovulation may be associated with a reduction in fertility.
Persistent anovulatory follicle Persistent anovulatory follicles may be analogous to follicular cysts in cattle.30 These occur from failure of a dominant follicle to ovulate but are distinct from failure to ovulate associated with hemorrhage into the follicle (see below). They are often slow to regress and may remain for much of the breeding season. In addition, they may be associated with ovarian inactivity on both ovaries after the structure develops. Affected mares go out of heat and lose endometrial edema as estradiol levels decline. Nonluteinized anovulatory follicles eventually regress or undergo atresia and are eventually replaced by another dominant follicle. Ovulation failure occurs in approximately 8–12% of equine estrous cycles during the physiological breeding season.32 A majority of dominant follicles (85–90%) that fail to ovulate exhibit hemorrhage into the follicular lumen and eventually form luteal structures. The remaining 10–15% of large follicles that fail to ovulate during estrus remain as transient follicular structures, although, in some cases, they apparently
Ultrasonographic characteristics Persistent anovulatory follicles (non-luteinized) are recognized on ultrasonographic examination as large static follicular structures with few to no echogenic particles or strands within the lumen (Figure 222.12). Partially collapsed anovulatory follicle Occasionally, a partially collapsed follicle will be observed during an ultrasonographic evaluation. An initial impression might be that the follicle was examined in midovulation. However, in most instances this represents a follicle undergoing atresia and is another form of persistent anovulatory follicle and has been previously described as luteinized unruptured follicles (LUF).33 Evaluation of progesterone concentrations may indicate limited luteinization, with levels ranging from 1 to 3 ng/mL. One can usually easily differentiate a partially collapsed anovulatory follicle from a follicle undergoing true ovulation by a followup examination a few hours later or the next day. The clinical significance is that failure of true ovulation or release of the oocyte is incompatible with pregnancy.
Ultrasonographic characteristics The ultrasonographic image typically reveals a follicular structure with a variable amount of echogenic particles in the lumen and an irregular thick-walled border (Figure 222.13). Subsequent examinations often show little change over the next 3–4 days, followed by a slow regression in size.
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tory follicles are associated with hemorrhage into the follicular lumen followed by a progressive development of fibrin strands over the next 2–3 days and eventually complete infiltration of luteinized cells.32 Progesterone levels begin to increase within 1–2 days after initial detection of echogenic particles in the follicular lumen and commonly increase to greater than 8–10 ng/mL (Figure 222.14). Administration of a single dose of prostaglandins will usually result in complete regression of the luteinized structure, and appears to be best performed at least 9–10 days after initial detection of echogenic particles.
Ultrasonographic characteristics
Figure 222.13 Ultrasonographic image of a partially collapsed anovulatory follicle.
Hemorrhagic or luteinized anovulatory follicle The fate of a dominant follicle of a mare in estrus that fails to ovulate is usually formation of a transient luteal structure. Approximately 85–90% of anovula-
The initial feature characteristic of the early development of a luteinized anovulatory follicle is the presence of echogenic particles within the follicular lumen (Figure 222.15). These echogenic spots are likely clumps of red blood cells, fibrin, or clusters of granulosa cells. Significant hemorrhage into the uterine lumen is often visible the following day and is characterized by a diffuse echogenic fluid within the enlarged follicular lumen (Figure 222.16). Fibrin strands are subsequently identified as echogenic bands traversing the follicular lumen (Figure 222.17). Eventually the entire follicle is infiltrated with echogenic material as luteinization is completed (Figure 222.18). Administration of prostaglandins once luteinization has been completed results in regression of the structure into a
Figure 222.14 Progesterone concentrations in a mare with a luteinized anovulatory follicle. Day 1 refers to the day echogenic particles were first observed in the follicular lumen. The arrow indicates the day prostaglandins were administered.
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Figure 222.15 Diffuse pattern of echogenic spots within the lumen of a follicle that will eventually form a non-ovulatory luteal structure.
large corpus albicans (Figure 222.19) and a rapid decline of progesterone to less than 1 ng/mL. Occasionally, a large amount of unclotted blood is noted swirling around during manipulation of the anovulatory follicle (Figure 222.20). The delay in clotting is presumably due to the presence of anticoagulants in equine follicular fluid.34 Manipulation of the ovary will often elicit a swirling motion of the blood in a hemorrhagic anovulatory follicle.30 Ovarian hematoma The term ‘ovarian hematoma’ was originally coined in the period prior to ultrasonography and referred to a post-ovulation hemorrhagic event resulting in ovarian enlargement. Further, it was noted that the ovarian hematoma was the most common cause of unilateral ovarian enlargement.35 Ultrasonography has allowed us to differentiate between hemorrhage into
Figure 222.16 Increased concentration of echogenic spots and occasional echogenic strands within the lumen of a hemorrhagic follicle. This image was taken 1 day after Figure 222.15.
Figure 222.17 Echogenic fibrin strands criss-crossing the follicular lumen of a developing non-ovulatory luteal structure. This image was taken 1 day after Figure 222.16.
the lumen of an anovulatory follicle and hemorrhage that normally occurs after ovulation during development of a corpus hemorrhagicum. Excessive bleeding into the lumen of a collapsed follicle after ovulation may result in formation of a hemorrhagic structure that greatly exceeds the diameter of the pre-ovulatory follicle (i.e., the ovarian hematoma). The clinical need to differentiate between the two hemorrhagic conditions is that the ovarian hematoma may be associated with pregnancy by virtue of the fact that a true ovulation occurred.
Ultrasonographic characteristics An ovarian hematoma has ultrasonographic characteristics of an enlarged corpus hemorrhagicum.
Figure 222.18 Ultrasonographic photograph of a luteinized anovulatory follicle. This image was taken 9 days after the initial detection of echogenic spots within the lumen of the follicle. A single dose of cloprostenol (250 µg i.m.) was administered after this image was taken.
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Figure 222.19 Ultrasonographic image of a regressing luteinized anovulatory follicle taken 3 days after administration of a single dose of prostaglandins.
Figure 222.21 Ultrasonographic hematoma.
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Abnormalities of luteal function Fibrin strands traverse through a pool of congealing echogenic fluid as the blood clot begins to form (Figure 222.21). Subsequent examinations reveal increased echogenicity as the process of cellular infiltration and luteinization occurs.
Abnormalities of luteal function are associated with inadequate progesterone production, a shortened duration of progesterone secretion, or a prolonged duration of progesterone secretion (Table 222.3). Luteal insufficiency Primary luteal insufficiency implies a deficiency in progesterone production by the corpus luteum that forms after ovulation. Luteal insufficiency has been suggested to be a cause of subfertility in mares, although data are limited.36, 37 Clinically, luteal insufficiency may be noted most commonly 14–16 days after ovulation in pregnant mares with a small regressing corpus luteum. The cause of the luteal insufficiency is likely to be failure of maternal recognition of pregnancy. Supplementation with exogenous progestins can successfully maintain the pregnancy in the absence of endogenous progesterone.
Ultrasonographic characteristics A small corpus luteum (Figure 222.22) is associated with low progesterone levels that may be inadequate to maintain pregnancy. Size of the corpus luteum is positively correlated with progesterone levels.
Table 222.3 Abnormalities of luteal function in the mare Condition
Figure 222.20 Unclotted blood in the lumen of a hemorrhagic anovulatory follicle.
Luteal insufficiency Shortened luteal phase (premature luteolysis) Persistence of the corpus luteum (pseudopregnancy)
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Figure 222.22 A small corpus luteum in a pregnant mare.
Shortened luteal phase (premature luteolysis) Premature destruction of the corpus luteum (luteolysis) is associated with an early onset of estrus and a decrease in the interovulatory interval. The most common cause of premature luteolysis in the mare is endometritis. Inflammation of the endometrium can result in sufficient synthesis and release of prostaglandins to cause luteal regression.38
Ultrasonographic characteristics Luteal regression can be determined ultrasonographically by a decrease in size of the corpus luteum on sequential examinations.
Figure 222.23 Ultrasonographic image of a mature corpus luteum.
tent corpus luteum will have good tone in the uterus and cervix, a pale, tight and dry cervix on speculum examination, and a blood progesterone concentration above 1.0 ng/mL.
Ovarian tumors Persistent corpus luteum (pseudopregnancy) A corpus luteum that fails to regress within 14–16 days after ovulation in a non-pregnant mare is considered to be pathologically persistent.39 The most commonly recognized causes of a persistent corpus luteum are ovulations late in diestrus, embryonic loss after the time of maternal recognition of pregnancy, and chronic uterine infections (i.e., pyometra). Insertion of a sterile glass ball (i.e., marble) into the uterus,40 chronic administration of oxytocin during diestrus,41, 42 and ovulation during a course of altrenogest therapy43 can also cause persistence of the corpus luteum.
Ultrasonographic characteristics A persistent corpus luteum is indistinguishable on ultrasonographic examination from a normal mature corpus luteum (Figure 222.23). Mares with a persis-
Ovarian tumors may arise from the surface epithelium, ovarian stroma, or germ cells (Table 222.4). The most common ovarian tumor in the mare is the granulosa cell tumor (GCT).44 Cystadenomas, teratomas, and dysgerminomas are considered to be rare in the horse. Granulosa cell tumor Granulosa cell tumors are almost always unilateral, slow growing, and benign.28 Granulosa cell
Table 222.4 Ovarian tumors of the mare Ovarian tumor type
Tissue of origin
Granulosa cell tumor Cystadenoma Teratoma Dysgerminoma
Ovarian stroma Surface epithelium Germ cells Germ cells
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Figure 222.24 Ultrasonographic image of a granulosa cell tumor with multiple small cysts.
tumors are hormonally active and behavioral abnormalities such as prolonged anestrus, aggressive or stallion-like behavior, and persistent estrus or nymphomania may be expressed in affected mares. Palpation and ultrasonography are the principal techniques used in the diagnosis of a granulosa cell tumor.45, 46 Endocrine assays for detection or confirmation of a GCT include measurement of inhibin and testosterone.28,47–50
Ultrasonographic characteristics Examination of the affected ovary by transrectal ultrasonography often reveals a multicystic or honeycombed structure (Figures 222.24–222.26), but the tumor may also present as a solid mass or as a sin-
Figure 222.26 Ultrasonographic image of a granulosa cell tumor comprising several large cystic structures.
gle large cyst. Less commonly, the GCT may begin as a large single cyst with hemorrhage that can rupture and cause hemoperitoneum.51 The contralateral ovary is usually small and inactive, characteristics which are the hallmark of a clinical diagnosis of a GCT. Miscellaneous ovarian abnormalities
Figure 222.25 Image of the same granulosa cell tumor noted in Figure 222.24 after surgical removal.
Cysts within the region of the ovulation fossa or adjacent to the ovary may be observed during a routine ultrasonographic examination.30 Epithelial inclusion cysts (or ‘ovulation fossa cysts’) arise from the surface epithelium that lines the ovulation fossa that becomes embedded within the ovarian cortex following ovulation.52 Epithelial inclusion cysts may interfere with fertility if they obstruct the ovulation fossa and are more common in older mares.30 Parovarian (periovarian) cysts (or ‘fimbrial cysts’) arise from embryonic vestiges and cystic accessory structures. Small fimbrial cysts generally do not interfere with ovulation or oocyte transport, respectively, and are not usually associated with reduced fertility.33
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Figure 222.27 Ultrasonographic image of a cluster of fossa cysts in an older mare.
Ultrasonographic characteristics Epithelial inclusion cysts are recognized as a small cluster of cysts filled with clear (non-echogenic) fluid in the center of the ovary, in the region of the ovulation fossa (Figure 222.27). Parovarian cysts can be small (< 10 mm) to large (> 30 mm) fluid-filled structures adjacent to the ovary (Figures 222.28 and 222.29). Parovarian cysts may be inadvertently mistaken for an ovarian follicle. Differentiation between a parovarian cyst and an ovarian follicle may be made by manual palpation per rectum.
Figure 222.28 Ultrasonographic image of a parovarian cyst (left) and several small ovarian follicles (right).
Figure 222.29 Parovarian cyst (arrow) adjacent to the ovary of a mare.
References 1. Carnevale EM, Bergfelt DR, Ginther OJ. Follicular activity and concentrations of FSH and LH associated with senescence in mares. Anim Reprod Sci 1994;35:231–6. 2. Carnevale EM, Bergfelt DR, Ginther OJ. Aging effects on follicular activity and concentrations of FSH, LH, and progesterone in mares. Anim Reprod Sci 1993;31:287–99. 3. Vanderwall DK, Peyrot LM, Weber JA, Woods GL. Reproductive efficiency of the aged mare. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1989, pp. 153–6. 4. Nagy P, Huszenicza G, Juh´asz J, Kulcs´ar M, Solti L, Reiczigel J, Abav´ary K. Factors influencing ovarian activity and sexual behavior of postpartum mares under farm conditions. Theriogenology 1998;50:1109–19. 5. Hughes P, Benirschke K, Kennedy PC, Smith AT. Gonadal dysgenesis in the mare. J Reprod Fertil Suppl 1975;23:385–90. 6. Zhang TQ, Buoen LC, Weber AF, Ruth GR. Variety of cytogenetic anomalies diagnosed in 240 infertile equine. In: Proceedings of the 12th International Congress on Animal Reproduction and Artificial Insemination, 1992, pp. 1939– 41. 7. Lear TL, Lundquist J, Zent WW, Fishback Jr WD, Clark A. Three autosomal chromosome translocations associated with repeated early embryonic loss (REEL) in the domestic horse (Equus caballus). Cytogenet Genome Res 2008;120:117–22. 8. Beech J. Treatment of hypophysial adenomas. Compendium 1994;16:921–3. 9. Van Der Kolk JH. Equine Cushing’s disease. Equine Vet Educ 1997;9:209–14.
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Ovarian Abnormalities 10. Dybdal N. Pituitary pars intermedia dysfunction (equine Cushing’s-like disease). In: Robinson NE (ed.) Current Therapy in Equine Medicine, 4th edn. Philadelphia: Saunders, 1997; pp. 499–501. 11. McCue PM: Equine Cushing’s disease. Vet Clin North Am Equine Pract 2002;18:533. 12. Maher JM, Squires EL, Voss JL, Shideler RK. Effect of anabolic steroids on reproductive function of young mares. J Am Vet Med Assoc 1983;183:519–24. 13. Turner JE, Irvine CH. Effect of prolonged administration of anabolic and androgenic steroids on reproductive function in the mare. J Reprod Fertil Suppl 1982;32:213. 14. Asa CS, Ginther OJ. Glucocorticoid suppression of oestrus, follicles, LH and ovulation in the mare. J Reprod Fertil Suppl 1982;32:247. 15. Sharp DC, Grubaugh WR, Weithenauer J, Davis SD, Wilcox CJ. Effects of steroid administration on pituitary luteinizing hormone and follicle-stimulating hormone in ovariectomized pony mares in the early spring: pituitary responsiveness to gonadotropin-releasing hormone and pituitary gonadotropin content. Biol Reprod 1991;44:983–90. 16. Farquhar VJ, McCue PM, Nett TM, Squires EL. Effect of deslorelin acetate on gonadotropin secretion and subsequent follicular development in cycling mares. J Am Vet Med Assoc 2001;218:749–52. 17. Carnevale EM, Checura CH, Coutinho da Silva MA, Maclellan LJ, Squires EL. Use of deslorelin acetate to suppress follicular activity in mares used as recipients for oocyte transfer. Theriogenology 2001;55:358. 18. Safir JM, Loy RG, Fitzgerald BP. Inhibition of ovulation in the mare by active immunization against LHRH. J Reprod Fertil Suppl 1987;35:229–37. 19. Dowsett KF, Tshewang U, Knott LM, Jackson AE, Trigg TE. Immunocastration of colts and imunospeying of fillies. Immunol Cell Biol 1993;71:501–8. 20. Tshewang U, Dowsett KF, Knott LM, Trigg TE. Preliminary study of ovarian activity in fillies treated with a GnRH vaccine. Aust Vet J 1997;75:663–7. 21. Garza F, Thompson DL, French DD, Wiest JJ, St. George RL, Ashley KB, Jones LS, Mitchell PS, McNeill DR. Active immunization of intact mares against gonadotropin-releasing hormone: differential effects on secretion of luteinizing hormone and follicle-stimulating hormone. Biol Reprod 1986;35:347–52. 22. Elhay M, Newbold A, Britton A, Turley P, Dowsett K, Walker J. Suppression of behavioural and physiological oestrus in the mare by vaccination against GnRH. Aust Vet J 2007;85:39–45. 23. Card C, Raz T, LeHeiget R. GnRF immunization in mares: ovarian function, return to cycling, and fertility. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007, pp. 576–7. 24. Robinson SJ, McKinnon AO. Prolonged ovarian inactivity in broodmares temporally associated with administration of Equity. Aust Equine Vet 2006;25:85–7. ¨ 25. Allen WR, Kolling M, Wilsher S. An interesting case of early pregnancy loss in a mare with persistent endometrial cups. Equine Vet Educ 2007;19:539–44. 26. Willis LA, Riddle WT. Theriogenology question of the month. J Am Vet Med Assoc 2005;226:877–9. 27. Steiner J, Antzak DF, Wolfsdorf K, Saville K, Brooks S, Miller D, Bailey E, Zent W. Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. 28. McCue PM, Roser JF, Munro CJ, Liu IK, Lasley BL. Granulosa cell tumors of the equine ovary. Vet Clin North Am Equine Pract 2006;22:799–817.
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29. McCue PM, LeBlanc MM, Akita GY, Pascoe JR, Witherspoon DM, Stabenfeldt GH. Granulosa cell tumors in two cycling mares. J Equine Vet Sci 1991;11:281–2. 30. McKinnon AO. Ovarian abnormalities. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Wilkins, 1998; pp. 233–51. 31. McCue PM, Vanderwall DK, Squires EL. Embryo recovery rates in mares with echogenic pre-ovulatory follicles. Havemeyer Found Monogr Ser 2001;3:84–5. 32. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. 33. McKinnon AO, Voss JL, Squires EL, Carnevale EM. Diagnostic ultrasonography. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Baltimore: Williams & Wilkins, 1993; pp. 266–302. 34. Stangroom JE, de G Weevers R. Anticoagulant activity of equine follicular fluid. J Reprod Fertil 1962;3:269–82. 35. Bosu WTK, Van Camp SC, Miller RB, Owen R Ap R. Ovarian disorders: clinical and morphological observations in 30 mares. Can Vet J 1982;23:6–14. 36. Bergfelt DR, Woods JA, Ginther OJ. Role of the embryonic vesicle and progesterone in embryonic loss in mares. J Reprod Fertil 1992;95:339–47. 37. Allen WR. Luteal deficiency and embryo mortality in the mare. Reprod Dom Anim 2001;36:121–31. 38. Neely DP, Kindahl H, Stabenfeldt GH, Edqvist L-E, Hughes JP. Prostaglandin release patterns in the mare: physiological, pathophysiological, and therapeutic responses. J Reprod Fertil Suppl 1979;27:181–9. 39. Stabenfeldt GH, Hughes JP, Evans JW, Neely DP. Spontaneous prolongation of luteal activity in the mare. Equine Vet J 1974;6:158–63. 40. Nie GJ, Johnson KE, Braden TD, Wenzel JGW. Use of a glass ball to suppress behavioral estrus in mares. In: Proceedings of the 47th Annual Convention of the American Association of Equine Practitioners, 2001, pp. 246–8. 41. Stout TAE, Lamming GE, Allen WR. Oxytocin administration prolongs luteal function in cyclic mares. J Reprod Fertil 1999;116:315–20. 42. Vanderwall DK, Rasmussen DM, Woods GL. Use of exogenous oxytocin to block luteolysis in mares: a plausible method of long-term suppression of estrus. In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007, pp. 578–9. 43. Daels PF, McCue PM, DeMoraes M, Hughes JP. Persistence of the luteal phase following ovulation during altrenogest treatment in mares. Theriogenology 1996;46:799– 811. 44. Hughes JP, Kennedy PC, Stabenfeldt GH. Pathology of the ovary and ovarian disorders in the mare. In: Proceedings of the 9th International Congress on Animal Reproduction and Artificial Insemination, 1980, pp. 203–22. 45. Hinrichs K, Hunt PR. Ultrasonographic as an aid to diagnosis of granulosa cell tumour in the mare. Equine Vet J 1990;22:99–103. 46. White RA, Allen WR. Use of ultrasonographic echography for the differential diagnosis of granulosa cell tumour in a mare. Equine Vet J 1985;17:401–2. 47. Stabenfeldt GH, Hughes JP. Clinical aspects of reproductive endocrinology in the horse. Comp Contin Educ 1987;9:678–84. 48. McCue PM. Equine granulose cell tumors. In: Proceedings of the 38th Annual Convention of the American Association of Equine Practitioners, 1992, pp. 587–93. 49. Christman SA, Bailey MT, Wheaton JE, Troedsson MH, Ababneh MM, Santschi EM. Dimeric inhibin concentrations
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in mares with granulosa-theca cell tumors. Am J Vet Res 1999;60:1407–10. 50. Bailey MT, Troedsson MHT, Wheaton JE. Inhibin concentrations in mares with granulosa cell tumors. Theriogenology 2002;57:1885–95.
51. Alexander GR, Tweedie MA, Lescun TB, McKinnon AO. Haemoperitoneum secondary to granulosa cell tumour in two mares. Aust Vet J 2004;82:481–4. 52. McEntee K. Reproductive Pathology of Domestic Animals. San Diego: Academic Press, 1990; pp. 31–93.
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Chapter 223
Uterine Abnormalities Angus O. McKinnon and Patrick M. McCue
With ultrasonography the uterus can be examined non-invasively to evaluate normal reproductive anatomy, detect pathological changes, and monitor therapeutic regimen(s). The most common forms of uterine pathology detected by ultrasonography in the non-pregnant mare are accumulations of intrauterine fluid, inflammatory changes, uterine cysts, and air. Less commonly, foreign bodies, fetal remnants, debris, abscessation, persistent endometrial cups, and neoplastic conditions are observed. Applications of ultrasonography to periparturient conditions have demonstrated its usefulness in examining mares with problems such as placentitis and postpartum hemorrhage. Each year we see pregnant mares presented for either reproductive evaluation or breeding that have mistakenly been diagnosed as not pregnant during the previous breeding season. The reasons for this are varied, but commonly this occurs when the only pregnancy test is completed at day 13–15 postbreeding and the mare is not examined again. Ultrasonographic evaluation of the uterus will identify the pregnancy and will help avoid an inappropriate internal evaluation and iatrogenic abortion.
Inflammation of the uterus Inflammatory changes in the mare’s uterus are recognized by the presence of uterine fluid (see below) and/or markedly increased edema of the endometrium (endometrial folds). During the normal estrous cycle, endometrial edema is detected as a normal development and follows the pattern of estrogen production and thus, similar to estrogen, uterine edema peaks approximately 24 hours prior to ovulation.1, 2 Edema of the endometrium is graded from 1 to 3 as a normal development during the estrous cycle.2 See Chapter 179 for pictures of normal grades of uterine edema. Typically edema is minimal or absent close to ovulation and, when multiple frequent examinations are performed, a change from grade 3 to grade 1 or no edema is quite useful to pre-
dict impending ovulation.2, 3 When edema is detected in mares that are in diestrus or pregnant, or when edema is marked (> grade 3) it is a sign of inflammation (Figure 223.1a–d). Some uterine edema of the dorsal surface of the uterus in pregnant mares is normal between days 17 and 30 and is associated with increased tone, fixation, and local production of estrogens from the conceptus.4 Marked edema (grade 4) can mask the presence of smaller amounts of intrauterine fluid. On occasion, systemic administration of 20–30 mg of dexamethasone phosphate can result in a spectacular reduction of edema from grade 4 to grade 1 or even to non-detectable within an hour.
Uterine fluid Ultrasonography is extremely valuable for estimating quantity and quality of fluid in the uterine lumen. Palpation of the uterus per rectum is only accurate when quantity of intrauterine fluid is large (> 100 mL) and(or) when uterine tonicity changes. Confirmation of smaller volumes of intrauterine fluid, without invasive techniques such as lavage and cytological analysis, was difficult until direct, noninvasive visualization was made possible with ultrasonography. Volumes of fluid within the uterine lumen are estimated (graded as very small, small, medium, large, or excessive) with ultrasonography, and quality is graded from 1 to 4 according to degree of echogenicity5 (Figure 223.2a–d). Degree of echogenicity is related to amount of debris or white blood cell infiltration into the fluid. Grade 1 fluid has large numbers of neutrophils and grade 4 has very few neutrophils when echogenicity is related to inflammation. Some other forms of uterine fluid detected are urine, blood, mucus, biofilm, and a transudate associated with uterine edema (Figure 223.2e–l). Pyometra refers to an accumulation of purulent material within the lumen of the uterus and typically is identified by ultrasonography as a grossly enlarged
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 223.1 (a) Grade 4 edema (endometrial folds) of the uterine horn with a small amount of grade 4 fluid. (b) Grade 4 edema of the uterine body. (c) Edema of the uterus can be normal in pregnant mares between fixation and 30 days and is usually only visible dorsally. After this time the rapid expansion of the fluid compartments of the pregnancy are associated with progressive thinning of the uterus and loss of any folding. (d) Edema of this pregnant uterus is more pronounced than normal and involves both dorsal and ventral uterine surfaces and is likely an abnormality.
uterus, filled with echogenic fluid and a thickened uterine wall (Figure 223.2m). The degree of uterine thickening may be masked in instances of excessive fluid accumulation and distension but is appreciated when the uterus is drained. Large accumulations of fluid may be associated with physical obstruction or functional defects of the cervix. An intermittent vaginal discharge may be evident if the cervix is patent. Affected mares are usually not systemically ill and clinical signs such as septicemia, depression or anorexia are absent. Hematological changes are minimal, when present. Endometrial damage due to the chronic inflammation may result in failure of prostaglandin secretion and persistence of the corpus luteum. Diagnosis is based
on palpation and ultrasonography of the reproductive tract per rectum, and speculum examination, as well as culture and cytology if the cervix is patent. The prognosis for future fertility is poor and in most instances treatment is aimed at salvaging the mare for non-breeding purposes. Urine in the uterus may result from vesico-vaginal reflux and subsequent flow forward through an open cervix, and may also occur from contamination by a stallion that has urospermia. Urospermia results in an immediate decline in spermatozoal motility6 and thus impacts fertility. In addition, urine in the uterus (Figure 223.2e) creates a chemical endometritis and, if it persists, will interfere with maintenance
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Figure 223.2 (a) Grade 1 uterine fluid. (b) Grade 2 uterine fluid. (c) Grade 3 uterine fluid. (d) Grade 4 uterine fluid. (e) Urine in the uterus. (f) Blood in the uterus after a uterine biopsy. (g) Biofilm layer in uterine horn. (h) Biofilm layer in uterine body.
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Figure 223.2 (i) Biofilm layer in a pregnant uterus. (j) Mucus that contained very few segmented neutrophils on high power. (k) Pregnancy surrounded by grade 2 uterine fluid. (l) Pregnancy with a small amount of grade 4 uterine fluid surrounding it. (m) Pyometra can be detected by ultrasonography as a grossly enlarged uterus, filled with echogenic fluid, and a thickened uterine wall. (n) Catheter tip visible in fluid in the uterus.
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Uterine Abnormalities of pregnancy. Urine in the mare’s uterus is identified by hyperechoic accumulations, most commonly at the corpus corneal junction, which will not swirl easily when balloted with the probe. The echogenicity of the urine is thought to be from a combination of crystals and mucus. Urine in the uterus should be removed with voluminous lavage. Observations on quality and quantity of uterine fluid have been used to assess efficacy of various therapeutic procedures in individual animals treated for naturally occurring endometritis.7–10 Experiments5, 8, 9,11–22 have been conducted to determine the relationship of intrauterine fluid to fertility, and are discussed in more detail in Chapter 275. Some areas where recognition of intrauterine fluid may be of particular value are: (i) evaluation of the postpartum mare; (ii) diagnosis and treatment of persistent mating-induced endometritis; (iii) diagnosis and treatment of infectious endometritis; and (iv) effects on pregnancy rate and early embryonic death (EED). Ultrasonography can also be valuable as a diagnostic aid during a therapeutic uterine lavage procedure when fluid recovery is difficult (Fig 223.2n). A majority of cases involve older mares, in which the most dependent part of the uterus is located cranial and ventral to the pelvic brim. The location of the fluid pocket is noted, the cuff of the lavage catheter is deflated, and the catheter is advanced into the pool of accumulated fluid. In some instances it may be necessary to infuse approximately 500 mL of lavage fluid (i.e., lactated Ringer’s solution) to distend the uterus prior to advancing the catheter.
Ultrasonographic studies of the uterus after parturition In the equine industry, economic incentives influence breeders to attempt a foaling interval of 12 months or less. This commonly necessitates breeding of mares during the first postpartum estrus. However, fertility has been reported to be lower in mares bred during the first postpartum ovulatory period compared with mares bred during subsequent cycles,23–27 and EED has been reported higher for mares bred at this time.23, 24, 28, 29 This decreased fertility may be due to failure of elimination of microbes during uterine involution, introduction of microbes at breeding, or inadequate endometrial repair relative to when the embryo enters the uterus. In addition, presence of uterine fluid during estrus and diestrus has been shown to reduce fertility of mares.5, 11, 12
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A study5 was conducted to evaluate two hypotheses: (i) uterine involution and fluid accumulation could be effectively monitored with ultrasonography and used to predict fertility of mares bred during the first postpartum ovulatory cycle; and (ii) delaying ovulation with a progestin would result in improved pregnancy rates in mares bred during the first postpartum ovulatory period. The previously gravid horn was larger than the non-gravid horn for a mean of 21 days (range 15 to 25) after parturition. Uterine involution was most obvious at the corpus cornual junction. When the results of three ultrasonographic scans performed over a 5-day period were similar, the uterus was considered to be involuted. On the average, uterine involution was completed by day 23 (range 13 to 29). Quantity and quality of uterine fluid were not affected by progestin treatment. Number of mares with detectable uterine fluid decreased after day 5 postpartum. Uterine fluid generally decreased in quantity and improved in quality between days 3 and day 15. Fewer (P < 0.005) mares became pregnant when uterine fluid was present during the first postpartum ovulatory period (3 of 9, 33%), compared to when no fluid was detected (26 of 31, 84%). Mares with uterine fluid during breeding did not have appreciably larger uterine dimensions, compared with mares not having fluid. There was no relationship between uterine size on day of ovulation and pregnancy rate. Ovulation was delayed, and pregnancy rates improved in progestin-treated mares. More (P < 0.05) mares became pregnant (23/28, 82%) when they ovulated after day 15, in the first postpartum ovulatory period, than mares that ovulated before day 15 (6/12, 50%). This study demonstrated that ultrasonography was useful in detecting mares with postpartum uterine fluid and can be used to aid in determining whether a mare should be bred and/or treated during the first postpartum ovulatory period.5 Uterine fluid may be spermicidal during estrus and an excellent medium to support bacterial proliferation at any time. In addition, fluid may indicate poor uterine involution. When fluid is present during diestrus, it may be associated with infectious endometritis and with premature luteolysis or EED.12 Quantity of uterine fluid during the first postpartum ovulatory period appeared to be related to stage of uterine involution, and was reduced or eliminated by delaying the ovulatory period with progestins.5 Progestin treatment not only allowed time for elimination of uterine fluid before the first postpartum ovulation, but it also significantly delayed the first postpartum ovulation. Progestagen treatment has been demonstrated to delay onset of the first postpartum ovulatory period without affecting the rate of
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uterine involution.5,30–32 Long-term progestin administration to normal, cycling mares has not been shown to adversely affect fertility.33 However, treatment with progestins will affect uterine defense mechanisms34, 35 and thus care is recommended before prolonged progestin treatment is administered to postpartum mares or mares susceptible to infection. Since there were decreased pregnancy rates associated with uterine fluid, and increased pregnancy rates as ovulation was delayed, it was suggested both techniques could be used to manipulate breeding strategies and improve pregnancy rates from normal mares bred during the first postpartum ovulatory period.5 Further studies on postpartum intrauterine fluid has demonstrated that mares that show uterine fluid accumulation during foal heat also present a larger incidence of post-breeding fluid accumulation, and that treatments of mares with oxytocin did not decrease the incidence of post-breeding fluid accumulation or improve pregnancy rates. Uterine flushes performed 36–48 h after breeding in mares with uterine fluid after foal-heat ovulation also did not improve pregnancy rates.22 In another study, EED rates were significantly greater (P = 0.015) in mares with intrauterine fluid accumulation during foal heat (30.5%) than in mares without fluid accumulation (11.1%).17 A study from Australia clearly showed the generally held contention that fertility is poorer and embryonic death greater when mares are bred on foal heat compared to the next cycle that was induced by prostaglandin.24 First cycle pregnancy rates were 47.9% compared with 55.2% and overall pregnancy rates at the end of the breeding season were 83.3% and 89.7%, respectively. Pregnancy losses before day 45 were 10% and 3.9% and after day 45 were 9.3% and 3.6%, respectively.24 Results of clinical, microbiological, and hormonal examinations performed during the puerperal period provide useful information when deciding whether to use the foal heat for breeding or to initiate a therapy during this period of time.36 Interpretation of results of lavage or other postpartum treatments37, 38 should be viewed with caution when all mares are treated in different groups, because most mares do not need treatment in the immediate postpartum period and our aims should be to identify those that do need treatment.
Foal-heat breeding strategies Mares can be considered for a foal-heat breeding if they had a normal foaling (i.e., no dystocia), fetal membranes were not retained, and there was no prolonged discharge of fluid (lochia) from the uterus.
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All mares are examined at day 2–3 post-foaling. By that time, problems that may have occurred during foaling will be identified. We believe that this examination is critical if we are to prevent mares from becoming ‘problem broodmares.’ Each year we identify mares that, if inappropriately treated, could become long-term disasters. Approximately 5% of mares will have placental tags present (despite excellent staff management) and another 5% will have problems with metritis and uterine damage. Many (15%) will have delayed involution, suggested by increased size of the uterus and volume of fluids present. Such mares are treated aggressively with large-volume uterine lavage and ecbolics, antibiotics and anti-inflammatories, and monitored closely. Postpartum mares may be treated with an infusion of antibiotics in lactated Ringer’s solution (LRS) 2–3 days after foaling. This may help some individual mares and may prevent iatrogenic contamination. Mares with no abnormalities identified are scheduled to be re-examined on day 9–10 post-foaling. Mares will not be bred any earlier than day10. On day 9–10 if the mare has ovulated she is scheduled for PGF2α administration in 6 days. If no uterine fluid is detected and uterine tone is good, the mare may be scheduled to be bred. If there is a question, the mare is treated and or re-examined according to follicle size and presence or absence of uterine fluid. It would be rare for a foal-heat service to take precedence over a non-foal-heat service with a stallion with a busy book (> 150 mares per season with natural service). Breeding mares on foal heat with intraluminal fluid accumulations is avoided. These mares are treated and recycled. Lastly, there are two more guidelines that we adhere to. First, we only breed mares on foal heat that are young and reproductively healthy (i.e., < 12 years old and with their first, second, or third foal) and second, mares that have been confined without exercise (i.e., lunging) due to foal problems such as angular limb deformities, are not bred on foal heat, regardless.
Diagnosis of endometritis There are numerous techniques to diagnose endometritis. However, no technique is completely reliable. The common, currently accepted techniques are: (i) rectal palpation; (ii) vaginal speculum examination; (iii) bacterial culture of uterine contents; (iv) cytological examination of uterine contents; (v) endometrial biopsy; and (vi) ultrasonography.
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Uterine Abnormalities A study was conducted11 to examine the efficacy of individual diagnostic techniques to predict endometritis. The experimental model involved 60 intact mares that were treated with progesterone for 31 days, and 50 were inoculated with a broth of Streptococcus zooepidemicus. Reproductive evaluations were performed the day progesterone treatment began, 13 days after progesterone treatment began, and 2 and 7 days after various therapeutic regimens. Thus, for the 60 experimental mares, there were 240 individual examinations for endometritis. The following criteria were used to assess degree of endometritis: (i) ultrasonographic detection of intraluminal fluid accumulation; (ii) vaginal speculum examination: (iii) cytological examination of uterine contents; (iv) culture of uterine contents; and (v) acute and chronic inflammatory changes detected by endometrial biopsy. Each individual parameter was assigned a score from 0 to 3. The total index score of 0 to 18, calculated from summation of each component of reproductive evaluation, was used as a standard to determine if the mare had endometritis. To determine the efficacy of each individual diagnostic test, a predictive value for each test (for each score > 0) was calculated against a positive diagnosis of endometritis obtained from the total index score (at two different levels of diagnosis). Results from this experimental model indicated support for two conclusions: (i) bacterial culture was not as accurate in predicting endometritis as other diagnostic tests; and (ii) ultrasonographic detection of any uterine fluid accumulation was an accurate indicator of endometritis.11 This study had several limitations: (i) The model of endometritis was progesterone-dependent and may not accurately reflect the naturally occurring condition of endometritis. A progesterone-dependent model may result in increased bacterial proliferation, decreased neutrophil numbers and function, and decreased drainage of uterine contents. (ii) Without an independent standard to determine endometritis (i.e., used to compare the individual tests against a positive or negative diagnosis of endometritis), the predictive value in this study may have more accurately reflected each individual component’s influence on the total index score. Despite these drawbacks, the study demonstrated the usefulness of ultrasonography to diagnose endometritis. The application of ultrasonography becomes even more apparent when considering that it is non-invasive. All 10 progesterone-treated uninoculated control mares developed endometritis, most likely induced from repeated invasion into the uterus to collect data during the study. It can also be concluded that, despite accepted hygienic techniques, invasion of the uterus in progesterone-dominated
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mares resulted in endometritis. Other authors have made similar associations.39
Breeding the problem mare See also Chapter 272.
Pre-breeding assessment Ultrasonography is fundamental to any problem mare breeding program.7, 40, 41 The primary goal of the pre-breeding assessment period is to determine if a mare has a uterine environment capable of supporting spermatozoa long enough so they can reach the oviduct in a condition capable of initiating fertilization. It only takes a few hours for spermatozoa to pass through the utero-tubal junction and remain relatively free from toxic products in the uterus. However, fluid inflammatory products, when mixed with spermatozoa, cause an immediate decline in spermatozoal motility that is proportional to the amount of inflammatory products. Also, addition of uterine fluid of grades 1 to 2 to spermatozoa prior to breeding mares by artificial insemination (AI) resulted in a decreased embryo recovery (P < 0.05).42 Addition of an extender to the uterine fluid prior to addition of spermatozoa was reported to arrest the decline in spermatozoal motility ‘in vitro’.42 However, it appears that, for best fertility, removal of inflammatory products from the uterus prior to introduction of spermatozoa is the most logical approach. To meet these objectives, mares are examined in early estrus. Ultrasonographic identification of uterine fluid and culture and sensitivity results are used to determine whether uterine lavage, ecbolic agents such as oxytocin and/or cloprostenol,7–9,43–45 local antibiotics, or a combination of all of the above are necessary to obtain a uterus free from inflammatory products at the time of breeding. It is an inappropriate use of time and finances to breed mares destined to return to estrus; thus mares with abnormal uterine fluid accumulations detected at the time of breeding may have to be recycled as soon as PGF2α is capable of causing luteolysis. Lavage and retrieval of lactated Ringer’s solution from the uterus of mares immediately prior to breeding has been shown not to be detrimental,46 and should be considered in cases where considerable uterine fluid is detected prior to breeding, and recycling the mare is not an option.
Breeding management If the uterine environment is prepared properly at the time of breeding, then our goal is to prevent
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contamination at, or immediately after, breeding. Clearly, breeding contamination is more easily controlled with AI than with natural service. However, both techniques result in introduction of bacteria. Research in this area was originally reported by Kenney and colleagues in 1975.47 They demonstrated that hygienically collected semen from ‘non-infected’ stallions contained numerous types of aerobic bacteria and fungi. The total number of aerobic microorganisms in each of eight ejaculations from five stallions collected ranged from 0.09 to 36 million. However, addition of raw semen to non-fat dry milk seminal extenders containing either penicillinstreptomycin (1500 IU/mL and 1500 µg/mL, respectively) or gentamicin sulfate (1 mg/mL) resulted in no growth on any of the subcultures from treated samples, including zero time which was after about 5 minutes of exposure. There was heavy growth in all subcultures from raw semen.47 Aerobic bacteria commonly isolated from the urethra, semen, and prepuce of stallions are E. coli and other coliforms, Pseudomonas aeruginosa, beta-hemolytic and non-hemolytic streptococci (S. zooepidemicus), Klebsiella sp., hemolytic and non-hemolytic staphylococci, Proteus sp., and Corynebacterium. Further experimentation has demonstrated effective elimination of bacteria without affecting motility, using seminal extenders containing either penicillingentamicin or polymyxin B sulfate (1000 IU/mL). Thus, it appears that addition of raw semen to appropriate antibiotic-containing seminal extenders is one method of ensuring minimal contamination at the time of breeding.48–53 It is important to remember that some antibiotics affect spermatozoal motility at high concentrations and may adversely affect fertility.54, 55 Currently we favor the use of ticarcillin, timentin, or amikacin at 1 mg/mL in extender preparations.
Artificial insemination If possible, mares are inseminated without disturbing reproductive surgeries such as the Caslick procedure. The perineum is diligently cleaned and dried, as previously described, and 500 × 106 progressively motile spermatozoa (PMS) mixed with appropriate antibiotic-containing extender are inseminated. Proper technique to ensure cleanliness of the stallion and collection equipment is important. To ensure the antibiotics have had adequate time to eliminate bacterial growth, it is best to allow at least 15 minutes at 37◦ C prior to insemination. If a longer interval is required, extended semen may be cooled to 20◦ C and stored for up to 12 hours56 and often considerably longer at 4◦ C.57 To reduce contaminating
organisms to an absolute minimum, a method was devised47 to ‘wash’ spermatozoa by dilution with an antibiotic-containing extender, followed by centrifugation (300g) to produce a ‘soft’ pellet, decantation of the supernatant, and resuspension of the resulting pellet in fresh, warm extender. This technique has the added advantage of removing much of the seminal plasma which reduces motility after prolonged incubation, with minimal damage to spermatozoa;58 however, it is time-consuming and may not always be necessary.
Natural service Mares and stallions to be bred by natural service should be well cleaned. It is wise to avoid strong disinfectants that may cause overgrowth of potentially pathogenic bacteria after prolonged use. A technique of minimizing contamination by prebreeding infusion of 100 to 300 mL of antibiotic-enriched seminal extender has been described. This technique has advantages; however, caution should be advised. For maximum reproductive efficiency, 500 × 10 6 PMS should be deposited into the reproductive tract. However, lower spermatozoal numbers are quite effective in highly fertile stallions. This information was derived from AI with small volumes of semen or semen plus extender. Recent information has indicated that spermatozoal concentration or volume of inseminate may be important factors in fertility. When mares were bred with 250 × 106 PMS in 100 mL of extender, embryo recovery rate was significantly depressed (13.6%, P < 0.001) compared to mares bred with these same spermatozoal numbers from the same ejaculates in 10 mL of extender (70.6%).42 This finding becomes important when mares are bred by natural service to stallions that, due to frequent breedings, may have low spermatozoal numbers in normal ejaculate volumes (30 to 150 mL) and thus spermatozoal concentrations lower than the threshold for normal fertility. Our approach for mares bred by natural service is to use ultrasonographic detection of quality and quantity of fluid, combined with culture and sensitivity results, to determine optimum treatments. For instance, if mares have a large volume of fluid detected, then voluminous lavage of the uterus with a physiological solution such as lactated Ringer’s solution or Dulbecco’s phosphate buffered saline that is not expected to be detrimental to spermatozoal survival is instituted immediately prior to breeding. Increased temperature (41–45◦ C) of infused fluids or oxytocin may aid in evacuation of uterine contents. The aim is to clear the uterus of inflammatory products and enable spermatozoa to have a relatively safe
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Uterine Abnormalities passage into the oviduct before further inflammatory products are released. Lavage and retrieval of lactated Ringer’s solution from the uterus of mares immediately prior to breeding has been shown not to be detrimental.46 If small quantities of uterine fluid are detected, then intrauterine antibiotic solutions are infused before breeding (> 12 hours). If uterine fluid is detected at the time of scheduled breeding, depending on the type and volume of fluid, the mare is either recycled, a small volume of antibiotic containing extender (< 50 mL) is infused immediately prebreeding, or oxytocin is administered intravenously and the mare re-examined in 1–2 hours.59, 60 When organizing timing of breeding the problem mare, much effort is directed toward trying to breed only once, just prior to ovulation (< 12 hours). Induction of ovulation with human chorionic gonadotropin (hCG) or deslorelin acetate is routine (see Chapter 196). Mares are bred regardless of the number of pre-ovulatory follicles and multiple ovulations are actively encouraged. There are few effective, commercially available drugs to increase the number of ovulations per cycle. Recombinant folliclestimulating hormone (eFSH) (see Chapter 193), although effective, is very expensive for equine use and is no longer commercially available. Immunization against an inhibin subunit has been demonstrated to be effective in increasing ovulation rates in mares.61, 62 Conception rates are proportional to ovulation rates;63 thus treatment by immunization against inhibin should improve pregnancy rates in normal and subfertile mares and in mares bred to subfertile stallions. With intensive reproductive management, multiple pregnancies when diagnosed early present little difficulty in reduction to a singleton (see Chapter 220). Post-breeding management The aim of therapies in the immediate period after breeding is to: (i) reduce infection and inflammation to create a uterine environment capable of supporting pregnancy; and (ii) prevent further contamination. Spermatozoa are safely in the oviduct within 4 hours of breeding and are protected from inflammatory products in the uterus and/or uterine treatments by the utero-tubal junction. The embryo will not be transported from the oviduct through the utero-tubal junction until around 5.5 to 6 days after ovulation; however, because most intrauterine therapies have an attendant degree of inflammatory response, and to allow time for foreign material to be expelled or absorbed, no treatments are administered after day 4 post-ovulation. The cervix begins to exhibit increasing tone and improves as a barrier to
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infection within 2 days of ovulation, although it remains more relaxed when inflammation of the reproductive tract is present. In addition, the corpus luteum remains resistant to PGF2α released from local inflammatory responses until at least day 5 after ovulation. Approximately 12–24 hours after breeding, the uterine response to breeding and contamination is assessed by ultrasonography. If fluid is absent, then either no treatments are administered, or treatments are administered as indicated daily for 2 additional days. If small amounts of fluid are detected, the same treatment is applied after first lavaging the uterus with lactated Ringer’s solution. Saline is quite an irritant, especially with low pH.64 Usually 1–2 L are necessary to remove inflammatory products. When large amounts of fluid are detected, the fluid is recultured and removed by voluminous lavage until returning fluid is free from debris. Oxytocin may be added (20 to 40 IU/L) to flushing solutions and/or administered intravenously. Oxytocin increases myometrial contractions in estrogen-dominated reproductive tracts65, 66 and aids in expulsion of material. Administration of oxytocin causes uterine contractions for 30 to 45 minutes. Alternatively, cloprostenol may be administered to mares that do not evacuate their uterine fluid in response to oxytocin. Uterine contractions following cloprostenol administration last for 2 to 4 hours.41 From the preceding discussion it should be clear how fundamental ultrasonography is to good management decisions when breeding mares.
Effect of intrauterine fluid on pregnancy rate and early embryonic death Mares with intrauterine fluid detected prior to ovulation have reduced pregnancy rates. Mares with intrauterine fluid detected after ovulation have reduced pregnancy rates and increased EED. A study was designed to determine the influence of intrauterine fluid on pregnancy rate and EED.11 It was concluded from this study that: (i) presence of small quantities of intrauterine fluid during estrus in cycling mares did not affect pregnancy rates at either day 11 or 50; (ii) intrauterine fluid, detected 1 or 2 days after ovulation, did not affect day 11 pregnancy rates, but was associated with a significant increase in EED and reduced day 50 pregnancy rates; and (iii) presence of intrauterine fluid during diestrus was associated with a significant decrease in day 50 pregnancy rates. Another study revealed the incidence of intrauterine fluid during diestrus (12/43, 28%) was associated with the presence of an inflammatory process, as indicated by a high biopsy score, reduced
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progesterone concentrations, and a shorter interovulatory interval.12 Mares with intrauterine fluid during diestrus had a lower pregnancy rate at day 11 and a higher embryonic loss rate by day 20 than did mares without fluid. The progesterone profile and length of interovulatory interval for mares with uterine inflammation suggested that embryonic loss in this herd was due to uterine-induced luteolysis rather than primary luteal inadequacy.12 The effects on pregnancy rate of three different treatments to remove intrauterine fluid were assessed in 1267 mares.9 The mares were mated and allocated, in strict rotation, to four treatment groups: (i) untreated controls; (ii) intrauterine infusion of broadspectrum antibiotics; (iii) intravenous injection of oxytocin; (iv) intravenous injection of oxytocin followed by intrauterine antibiotics. The pregnancy status of the mares was determined 13 to 15 days and 27 to 30 days after ovulation by ultrasonography. The pregnancy rate of group 4 (72%) was higher than that of group 2 (64%, P < 0. 01) or group 3 (63%, P < 0. 01). The pregnancy rates of groups 2 and 3 were higher than that of group 1 (56%, P < 0.01). The treatment with antibiotics and oxytocin appeared to have an additive beneficial effect, which suggested two different modes of action of the combination treatment, namely, antibacterial activity and fluid drainage. In the untreated mares more fluid accumulated in the uterine lumen after mating, and this was the most likely reason for their lower pregnancy rate.9 In our breeding programs the amount and quality of fluid detected with ultrasonography is used to determine how to treat the mare. Large volumes of poor-quality (grade 1 or 2) fluid are treated with voluminous saline lavage, while small volumes of grade 4 fluid quite often are treated with local antibiotics and oxytocin and/or cloprostenol. When fluid is detected after ovulation then efforts are directed at reducing fluid and inflammation, such as intravenous non-steroidal anti-inflammatory agents, intrauterine antibiotics, immune stimulants,67 and systemic oxytocin. Cloprostenol is not used after ovulation due to the potential of modification of corpus luteum function.68–71
Uterine cysts Prior to ultrasonography, uterine cysts were most commonly diagnosed from postmortem examination and occasionally by rectal palpation72 (see Chapter 276). Subsequently they have been diagnosed by hysteroscopy and ultrasonography.73 Cysts in the uterus are fluid-filled and apparently have two origins. Endometrial cysts arise from endometrial glands, and are usually < 10 mm in diameter. The second form of
uterine cysts are lymphatic in origin (Figure 223.3a,b) and generally are larger than endometrial cysts. They are common in older mares,73–75 and have been associated with both normal and abnormal uterine biopsies.72 Size of uterine cysts may be indicative of origin. Little data has been reported on growth rate of uterine cysts. Despite the occasional large cysts, it is unlikely that they grow at a similar rate as the early embryonic vesicle (days 10 to 20). When visualized with ultrasonography, cysts are commonly rounded, sometimes with irregular borders, and occasionally are multiple or compartmentalized (lymphatic lacunae) (Figure 223.3c–f). Most cysts are luminal; however, occasionally transmural and subserosal cysts are detected. Movement of the early equine conceptus (days 10 to 16), presence of specular reflection, spherical appearance, and growth rate of the embryo should aid in its differentiation from uterine cysts. The relationship between infertility and uterine cysts is axiomatic as cysts increase in number as mares age.75 Cysts may impede movement of the early conceptus, restricting the reported ability of the vesicle to prevent luteolysis after day 10.76 Later in pregnancy, contact between the cyst wall and yolk sac or allantois may prevent absorption of nutrients (Figure 223.3f). This may be more important when considering the recognition that large uterine cysts are more commonly located at the junction of the uterine horn and body,75 which is the most common site of vesicle fixation.77 Finally, cysts are commonly indicative of uterine disease. They may reflect senility or be associated with endometrial fibrosis. It has been reported that there is an association between number of uterine cysts, age of mare,77 and endometrial biopsy score.12 The number of treatments proposed for uterine cysts probably reflects inability of any individual treatment to consistently be useful. Rupture of the fluid-filled structures has been attempted via uterine biopsy forceps, surgery, fineneedle aspiration and puncture via hysteroscopy. Electrocoagulative removal of cysts has also been described. Endometrial curettage and repeated lavage with warm saline (40 to 45◦ C) have also been advocated. Although there are no reports on respective efficiency of these treatments, endometrial curettage and saline lavage are frequently applied to treat the primary problem, which would appear to be lymphatic blockage. It was concluded from one study that: (i) uterine cysts, when detected by ultrasonography, were lymphatic in origin; (ii) uterine cysts did not change rapidly in size or shape, although they were more difficult to detect during estrus; (iii) treatment with infra-red radiation was not effective; (iv) there was no consistent location for uterine cysts; and (v) uterine cysts were commonly associated with chronic, infiltrative, lymphocytic endometritis.11
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Figure 223.3 (a) Uterine cyst on the surface of the endometrium. (b) Histology of a uterine lymphatic cyst (× 2). (c) Day 12 pregnancy in front of the cervix with two cysts of similar size immediately cranial. (d) Single cyst with inflammatory folds and fluid. (e) Multiple cysts associated with a pregnancy at the corpus corneal junction. (f) Larger number of cysts at the corpus corneal junction with a pregnancy. This number of cysts may interfere with absorption of nutrients although there are no definitive studies to prove this. (g) Pedunculated uterine cysts removed with a snare. (h) The suture material in the pipettes is used to saw off the attachment of the cyst while the pipettes protect the uterus from damage.
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Another study on 259 normal fertile Thoroughbred mares75 revealed the incidence of uterine cysts was 22.4%. Of the 95 cysts observed during the trial, 87.4% were located in the middle and posterior segments of both uterine horns. The size of all cysts ranged between 3 and 48 mm. When all mares were assigned to three age groups – A < 7 years (n = 116), B 7–14 years (n = 117), and C > 14 years (n = 26) – a significant (P < 0.01) increase in the number of cysts was observed with advancing age (4.3%, 29.1%, and 73.1%, respectively). The seasonal pregnancy rates at days 14 and 40 were significantly (P < 0.01) lower in mares with cysts (77.6% and 71.4%) compared to mares without cysts (91.5% and 88.0%). This suggested to the authors that the presence of uterine cysts plays an important role in the reduction of fertility of Thoroughbred mares.75 From our experience cysts are more commonly noted in older mares and are also more commonly recognized in the immediate postpartum period and at the corpus cornual junction. Uterine cysts are more commonly pedunculated and luminal; however, transmural and non-pedunculated cysts are also seen. Our preferred treatment for uterine cysts, when we believe treatment is necessary, has been to dilate the cervix while the mare is in heat and ablate the cyst manually. This is performed by placing the cyst in between two fingers and tearing or crushing the cyst off the uterine wall. Non-pedunculated cysts may be more difficult to destroy this way; however, they seldom appear to cause many problems. Recently, a simple improvised snare technique has been described to remove pedunculated uterine cysts.78 The snare consists of two insemination pipettes fixed together with a long portion of thick suture material as the snare. The suture material is passed over the cyst and fixed at the pedunculated base of the cyst. A sawing motion is used to transect the stalk (Figure 223.3g,h).
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Intrauterine air Air is recognized by the presence of multiple, hyperechogenic reflections (Figure 223.4a,b); occasionally a ventral reverberation artifact is present and it appears to be more prevalent slightly cranial to the cervix, although it can be present in the cranial body or uterine horns. When air is present < 24 h after artificial insemination, we consider it normal. However, we do not expect it to be detected in normal mares > 24 h after breeding. The observation of air in the uterus of mares that have not been cultured or bred recently is an indication of pneumouterus and reflects failure of the competency of the vaginal labia, vestibulovaginal sphincter, and(or) cervix.11 In our experience it has been of use to determine when some mares may need to undergo a Caslick procedure, particularly so when a clinical examination did not suggest the procedure. Occasionally air may be detected in a pregnant mare. In these cases the prognosis for pregnancy maintenance is poor.
Foreign bodies On occasion, strongly echogenic areas in the uterine lumen are observed with a concomitant echo shadow, seen with dense tissue such as fetal bone (Figure 223.5a). This might be expected after mummification. We have also identified a similar ultrasonographic image that was confirmed subsequently as the tip of a uterine culturette (Figure 223.5b). Another substance that creates a strong echo shadow is a chronically retained fetal membrane. These cases are usually recognized at the first postpartum examination; however, they may not be apparent until the echogenicity increases. In addition, we have seen a few interesting cases such as calcification on the uterine epithelial surface, suture material after a cesarean section, and
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Figure 223.4 (a) Air is detected as hyperechoic reflections in the uterine horn. (b) Air and fluid immediately cranial to the cervix.
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Figure 223.5 (a) Fetal remnants in a uterus with large amount of grade 2 fluid. (b) Echogenic mass that was confirmed to be the tip of a culturette. (c) Glass marble in the uterus. (d) Glass marble visualized by endoscopy.
post-treatment debris after multiple antibiotic suspension treatments into the uterus of a mare that had poor uterine clearance abilities. Marbles placed into the uterus to create a pseudo-pregnant state are easily visualized by ultrasonography (Figure 223.5c,d).
Retained endometrial cups In a normal pregnancy the endometrial cups are not detected; however, on occasion after EED and expulsion of fetal fluids, they may become visible. Mares can in unusual circumstances retain endometrial cups79, 80 in an active state for a long time (a year or more) despite either foaling, aborting or suffering from EED.81 These mares present as dry mares in the next breeding season and typically have irregular estrous cycles with abnormal ovarian function such as follicles that luteinize without ovulating. Presumably the abnormal ovarian function is associated with the high levels of equine chorionic gonadotropin (eCG)
that are secreted from the retained cups. Mares with retained cups may exhibit irregular or prolonged estrous or anestrous behavior. A diagnosis of retained endometrial cups can be suggested by ultrasonography and confirmed by endoscopy of the uterus or by measurement of serum eCG concentrations (Figure 223.6). Retained endometrial cups will eventually be removed by immunological rejection but this may take months. There is no technique currently available to consistently hasten the removal of aberrant persistent endometrial cups.81
Uterine adhesions Occasionally, after severe trauma to the reproductive tract or even after irritation from endometritis, transluminal uterine adhesions will form. Because of continuing secretions from the oviduct or endometritis in the horn proximal to the adhesions, fluid will accumulate in the tip of the affected horn. This fluid
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Mare: Diagnostic Ultrasonography traacetic acid–tromethamine (EDTA-tris;) (250 mL of 3.5 mM EDTA, 0.05 M tris, pH 8).41 (See Chapter 272).
Periparturient maternal disorders
Figure 223.6 Retained endometrial cups may on occasion be identified as hyperechoic plaques.
is well localized and should make the examiner suspicious of a complete adhesion. Diagnosis can be confirmed manually after first gently dilating the cervix or, if that is not possible, infusions of fluid (e.g., 1 L of lactated Ringer’s solution) into the uterus should demonstrate lack of fluid entering part of the uterus (typically a uterine horn). When adhesions exist it is not possible to dilate the tip of the affected horn because the fluid has restricted or denied entry. Adhesions may respond to manual breakdown and multiple infusions of an antibiotic extender; however, some appear to get worse with the irritation of treatment and may respond better to either an oil-based corticosteroid cream or rest from treatment for at least 3 weeks.
Less commonly recognized uterine pathology Uterine neoplasia, abscesses, and hematomas can be recognized with ultrasonography. Leiomyomas of the uterus are not uncommon, although careful scrutiny may be necessary to detect them as their detectable size varies from 10 mm up to as large as 60 mm. Leiomyomas are only slightly more echogenic than the uterus and they present as rounded discrete masses (Figure 223.7a–c). Ultrasonography may be useful in recognition of mucus layers and biofilm in the mare’s uterus (Figure 223.2g–i). Biofilm production is a protective mechanism of certain bacteria and is associated with increased difficulty to eliminate the infection with traditional treatment. Recent therapies showing promise for mucus and biofilm removal or modification are N-acetylcysteine (NAC) (30 mL of a 20% solution in 150 mL of saline)82 or ethylenediaminete-
A variety of injuries and disorders develop around the time of foaling and in the immediate postpartum period, that can have serious consequences for the mare. The age and breed of the dam have some influence on the incidence of these complications, but many are purely accidents associated with the rapid and often violent birth process. Frequently, diagnosis is difficult because the clinical signs of many postpartum problems are non-specific. Early recognition of peripartum disorders may improve the success rate of treatment. Some of these conditions are life-threatening, some will seriously compromise the future reproductive career of the mare, and some will cause considerable delay in the establishment of a healthy pregnancy, if not quickly recognized and treated. Ultrasonography has become an indispensable tool for the diagnosis and, in some cases, the monitoring of these conditions. Preparturient problems
Mummification and maceration See also Chapter 244. Mummification is not uncommon when one twin dies and the other continues to develop. Mummification of a single pregnancy is rare but has been reported.83–85 Occasionally lactation in a late pregnant mare may suggest fetal compromise, impending abortion, or mummification; however, commonly no clinical signs are recognized. Mummified fetuses are most commonly seen at birth or abortion. Maceration most commonly occurs when a single pregnancy dies and is not aborted. External signs of discharge may lead to the diagnosis. Treatment for both is gentle relaxation of the cervix and then delivery. We have seen naturally occurring cases of mummified fetuses with transabdominal ultrasonography but they can be difficult to detect (see Chapter 220). In all cases the mares were late in gestation and had started to lactate. The diagnosis of mummification was not easy, as all mares had an additional viable pregnancy present. In our limited experience it appears that identification of the fetal head or ribs as a separate entity from the live fetus are the easiest and most consistent fetal parts to find. As the mummification process proceeds it is harder to see fetal parts because they all assume a similar echogenicity. Only one case had a demonstrable ‘twin membrane,’ which probably was related to the more recent demise of that fetus. Early demise is associated with obvious echogenicity of fetal bones (Figure 223.8a,b).
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(c) Figure 223.7 (a) Small leiomyoma in the uterus. They typically lie in the submucosa or deeper and can on occasion be removed after gently dilating the cervix and tearing them off manually. (b) Large leiomyoma. (c) A large leiomyoma that has been highlighted by infusing lactated Ringer’s solution into the uterus.
Hydrops See also Chapter 243. Ultrasonography is useful for diagnosis of excessive accumulation of fluid in the uterus of a pregnant mare. Hydroallantois is a rare condition that occurs during the last trimester of pregnancy in primarily pluriparous mares after an otherwise uneventful gestation. Clinical signs include an enormous increase in abdominal size over 10–14 days, due to a rapid accumulation of allantoic fluid.86–88 Other clinical signs seen result from the pressure exerted by the excess amount of fluid. Mares may have anorexia, tachycardia, tachypnea, dyspnea, ventral edema, colic, and difficulty in defecating and walking. Horses with advanced cases may show evidence of ventral abdominal hernia or prepubic tendon rupture.
Evaluation of the genital tract per rectum reveals a huge, fluid-filled uterus and it may be difficult to palpate the fetus. Prognosis for obtaining a live foal is poor, and management of the case should be directed at saving the mare’s life.89 Parturition should be induced to relieve the pressure on the mare’s internal organs. Removal of the allantoic fluid slowly by rupture of the allantochorion should be attempted, as should siphoning of the fluid while supporting the mare with intravenous fluids and corticosteroids to prevent shock. Rather than fluid loss causing shock, we believe that the fluid loss removes the pressure on the abdominal vessels, and pooling and hypovolemic shock may follow when it occurs too quickly. Oxytocin may be administered to aid in evacuation of the uterine contents, although in mares with uterine inertia, the uterus may not respond to treatment.
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Mare: Diagnostic Ultrasonography monsters with gut abnormalities often have excessive fluid (8–10 times normal) in the amniotic cavity (hydrops amnii). Amniotic fluid is composed of saliva and secretions of the nasopharynx of the fetus. Although swallowing by the fetus may play a role in the maintenance of fetal fluid balance, other mechanisms may be important as there is no reported incidence of hydrops conditions with atresia or agangliosis of the digestive system. In cases of hydrops amnii, prognosis for the pregnancy is poor, because abortion and premature delivery are quite common. Fortunately, the prognosis for future fertility is fair to good, because the dam’s genital tract usually is normal after termination of the pregnancy.
Prepubic and abdominal wall rupture
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(b) Figure 223.8 (a) Mummification of a fetus that had recently died. (b) Fetal parts are visible in this image of a mummified/macerated fetus.
Hydrops amnii is an excessive accumulation of amniotic fluid in the amniotic cavity and it has a low incidence in all species.90 Hydrops amnii in the mare is reported less frequently than hydroallantois.91 The normal volume of amniotic fluid in the mare near term is between 3 and 7 L. Hydrops amnii tends to develop gradually over several weeks or months, during the latter half of gestation. The normal amniotic cavity has a direct communication with the digestive system by way of the mouth and anus. The fetus swallows fluid and may keep levels constant, as fetal
See also Chapter 250. A breakdown in the body wall of pregnant mares is common in older draft mares,89, 92 but has also been reported in other breeds. Conditions causing severe distention of the body wall, such as hydrops, twins, severe ventral edema or trauma, may result in rupture of the prepubic tendon, abdominal wall rupture or an abdominal hernia; however, many cases occur with no apparent reason. Mares usually present very close to foaling with severe, ventral edema running cranially from the udder and, in many cases, involving the udder itself. Unilateral edema may suggest partial rupture of the prepubic tendon or damage to the ventrolateral body wall. Affected mares have difficulty rising and are reluctant to move. Acute progression of either condition may result in severe distress, colic, tachypnea, tachycardia, sweating, internal hemorrhage, shock, and death. Prognosis for mare survival was better with non-intervention and when rupture was not associated with hydrops.93 Although the causes of these conditions are similar, the clinical appearance may help differentiate them. A mare with prepubic tendon rupture will have an elevated tail head and tuber ischii, due to the lack of ventral support of the pelvis, that results in lordosis and a sawhorse stance. The mammary gland may be shifted cranially from its normal position. Although a mare with an abdominal hernia or rupture may have an enlarged abdomen, her tail head and tuber ischii will be in a normal position. A diagnosis of prepubic tendon rupture or ventral body wall hernia may be difficult to confirm. In heavily pregnant mares, the presence of the fetus often makes palpation of the defect per rectum very difficult, and palpation transabdominally is usually impossible because of the amount of edema in the area. In our experience, ultrasonography can be useful in identifying body wall defects and herniation of intestine, but the extent of the damage may not
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Uterine Abnormalities be clearly defined until after parturition, when most of the edema is resolved. On occasion a diagnostic differential is severe edema in the prepartum mare. In cases of rupture the secretions in the mammary gland will commonly be bloodstained, while prepartum edema is not associated with blood in mammary secretions. This has important diagnostic significance as exercise is useful in the former condition. Mares suffering from abdominal wall ruptures are best left to provide foals by embryo transfer in the future; however, on occasion mares have successfully carried foals to term after a period of rest.
Uterine torsion Uterine torsion in the mare occurs late in gestation (> 8 months), more frequently in draft breeds than light horses, and is not usually associated with parturition (see Chapter 252), unlike in the cow. A low incidence of uterine torsion in mares is attributed to the sublumbar attachment of the ovaries and the dorsal insertion of the broad ligaments in the uterus. The causes of uterine torsion in the mare are not well defined, but include factors such as vigorous fetal movements, sudden falls, relative fetal oversize, diminished volume of fetal fluids, lack of tone in the pregnant uterus, long mesometrium, and the presence of a large, deep abdomen. Clinical signs are related to the severity of the torsion and include signs of abdominal pain such as restlessness, sweating, anorexia, frequent urination, sawhorse stance, looking at the flank, and kicking the abdomen. Signs may present for a variable duration (from few hours to 3 days or more) and they can mimic those of the early stages of parturition. The uterus may twist 90–540◦ on its long axis (Figure 223.9). In the mare, the twist typically involves the uterine body, not the cervix or the vagina, as occurs in the cow. If the torsion is less than 180◦ and
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blood flow to the uterus is not compromised, correction may not be required. The mare may show signs of shock if uterine rupture has occurred. Diagnosis is made by clinical signs and palpation per rectum of the gravid uterus; vaginal examination is only occasionally diagnostically useful. Palpation per rectum reveals tense broad ligaments, spiraling in the direction of the torsion. If the torsion is counterclockwise (as viewed from the rear), the left broad ligament will be taut and be felt to traverse under the uterus, while the right broad ligament will stretch dorsally over the distended uterus. The opposite will be present in a clockwise uterine torsion. The small colon undergoes a variable amount of constriction, due to the compressive forces of the displaced broad ligaments, and may impede the ability of the examiner to perform a complete rectal examination. In such cases, it is difficult to evaluate fetal viability or uterine integrity and it may even be impossible to determine which direction the uterus is rotated. Ultrasonography has not been useful for diagnosis of uterine torsion in our experience; however, it has been useful for determining if the fetus is still alive and also to evaluate placental attachment. In cases where the fetus has already died, management is different and frequently more aggressive. The prognosis for cases of equine uterine torsion depends on the degree of the circulatory disturbance resulting from involvement of the broad ligaments and uterine vessels. When the fetus is alive and the uterine wall is not severely congested and edematous, and treatment is prompt, the prognosis for maternal survival and the birth of a normal, live foal at term is good. A recent large retrospective study showed increased survival rates for both mares and foals when the uterine torsion occurred at less than 320 days’ gestation compared to a later occurrence.94 Overall mare survival was 84% (53/63). When uterine torsion occurred at < 320 days’ gestation, 97% (36/37) of mares survived compared to 65% (17/26) survival when uterine torsion occurred at ≥ 320 days’ gestation. Overall foal survival was 54% (29/54). When uterine torsion occurred at < 320 days’ gestation, 72% (21/29) foals survived compared to 32% (8/25) when uterine torsion occurred at ≥ 320 days’ gestation. Thirty mares were discharged from the hospital carrying a viable fetus following uterine torsion correction, and 25/30 (83%) of these mares delivered live foals that survived beyond the neonatal period.94
Preparturient uterine rupture
Figure 223.9 Uterine torsion at postmortem.
See also Chapter 253. Uterine rupture usually occurs in the peripartum period as a result of manual intervention or fetotomy
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or possibly because of violent intrapartum movement. However, in cases of uterine torsion95 or hydrops96 and in occasional idiopathic cases, the uterus can rupture before term. When the uterus ruptures before term, the mare may not show much pain, and if the fetus escapes into the abdomen and hemorrhage is not severe, uterine involution may occur without external signs. Diagnosis of uterine rupture may be facilitated by palpation and ultrasonography of the uterus per rectum, by fetal examination, abdominal centesis, and a history of moderate to severe abdominal pain during the last trimester of pregnancy. Once the uterus has ruptured and its contents have escaped, the uterine wall may feel thickened and corrugated, because the uterus contracts and begins to involute quite quickly. The site of rupture may be equally difficult to demonstrate by transrectal ultrasonography. However, transabdominal ultrasonography may reveal the presence of the fetus on the abdominal floor with excessive free abdominal fluid.
Placental separation and placentitis See also Chapters 242 and 254. Detachment of a significant portion of the allantochorion from the endometrium constitutes an emergency for the foal, owing to failure of adequate exchange of oxygen, carbon dioxide, and nutrients. Premature placental separation may occur in two forms in pregnant mares: acutely at parturition and more chronically during gestation, usually associated with placentitis. Chronic placentitis may be recognized by transabdominal ultrasonography; however, rectal ultrasonography is necessary to examine the cervix region in cases of ascending placentitis (Figure 223.10a–d). Chronic ascending placentitis or edema may result in separation and thickening of the placenta over the cervical region.
mares bred earlier in the season, Ginther suggested that mares may be able to make limited adjustments, so that their foals are born with an optimal chance for survival.98 The physiological control of this phenomenon appears to be related to photoperiod.99 Owners and farm managers are often concerned about pregnancies extending beyond the expected parturition date, and begin to exert pressure on the veterinarian to induce foaling. If no mammary development has occurred and fetal well-being has been assessed by ultrasonography, this practice should be strongly discouraged, even if the duration of gestation is longer than 1 year. We have seen numerous cases of foals from induction of parturition after a prolonged gestation where fetal immaturity resulted in an inability to survive for more than 24 hours. Patience and waiting for parturition to be initiated without intervention are indicated in most cases of prolonged gestation; however, fescue toxicosis also causes prolonged gestation and very large foals, and may well require specific intervention.100 Tall fescue (Lolium arundianceum) infected with the endophytic fungus Neotyphodium coenophialum (previously named Acremonium coenophialum), when ingested by pregnant mares, can have serious deleterious effects on the latter stages of equine pregnancy. Fescue toxicosis causes not only prolonged gestation, but also dystocia and hypogalactia. In affected foals, toxicosis causes dysmaturity and poor chances of survival (see Chapter 249). Placentas from affected mares are thick and edematous, and placental edema and premature separation of the chorioallantois have both been observed with ultrasonography.
Postparturient problems
Perivaginal hematomas and abcesses Prolonged gestation See also Chapter 253. Gestation usually lasts for 310–374 days, but normal pregnancies can be as long as 399 days.97 We have seen numerous cases of confirmed gestation longer than 400 days with one foal being born alive at 432 days. Foals born from such pregnancies are not usually oversize and do not predispose the mare to dystocia.97 Typical foals are small with a domed head and long tail and mane. The duration of gestation is partly controlled by nutrition and the genotype of the foal, but mares themselves may also control the duration of gestation. Based on the finding that average gestational durations were longer for
Intrapelvic perivaginal bleeding that results in perivaginal hematoma formation is not difficult to diagnose with ultrasonography. Large hematomas can be drained 2–3 days postpartum, after clot formation has occurred. If hematomas do not regress or do not become progressively firm, abscessation should be suspected. Occasionally perivaginal or abdominal, rather than retroperitoneal, abscesses may be confirmed by ultrasonography and by needle aspiration. The abscess should be incised and drained if not abdominal; however, on occasion we have drained abscesses into the uterus. Occasionally, large intrapelvic hematomas may leave permanent lumps of scar tissue that are palpable ventrally in the pelvic canal.
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Figure 223.10 (a) Abdominal ultrasonographic image of a mare with placentitis. (b) Transrectal ultrasonographic image demonstrating air at the cervix and in the fluid to the right-hand side. This fluid is outside the placenta. (c) Fluid outside the placenta with air interspersed. (d) Fluid outside the placenta, the chorioallantois, and fetus, from left to right.
Periparturient hemorrhage Hemorrhage from the middle uterine, utero-ovarian or, less frequently, external iliac arteries is a significant cause of peripartum colic syndrome and death, usually in older, multiparous brood mares (see Chapter 259). Hemorrhage from these vessels is often fatal, especially if the hemorrhage is into the abdominal cavity. The incidence of periparturient hemorrhage increases with age. Although it may occur before, during, or after parturition, it is most common and clinically significant during or within the first 24 h postpartum. Dystocia is not always associated with the hemorrhage and hemorrhage is often seen in mares after an apparently problem-free delivery. Fatal hemorrhage has also been reported after periparturient accidents such as uterine torsion, uterine tears or uterine prolapse. The clinical signs associated with periparturient hemorrhage depend on the
site and severity of the hemorrhage and on whether bleeding is contained within the broad ligaments or has escaped into the abdomen. Mares that bleed into the broad ligaments will exhibit variable signs of abdominal discomfort as a result of tension on the ligament and uterine serosa. Sometimes these signs may be overlooked as mares often show some abdominal discomfort associated with uterine contractions immediately postpartum. As blood loss and broad ligament distention increase, signs of abdominal pain, sweating, weakness, tachycardia, tachypnea, ataxia, and pale mucous membranes become more apparent. Rupture from the broad ligament or continued blood loss may lead to rapid onset of hemorrhagic shock. The course of clinical signs may be extremely rapid in cases of complete rupture and when hemorrhage occurs directly into the abdomen. Death often ensues within minutes to hours after the appearance of the first signs of discomfort. In other cases,
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Mare: Diagnostic Ultrasonography Retained placental membranes/ metritis-endotoxemia-laminitis syndrome
mares may have exhibited mild signs initially (associated with containment of hemorrhage within the broad ligament), only to collapse later. Mares suffering from uterine hemorrhage are sometimes just found dead. The time of onset of clinical signs appears to be related to prognosis, with mares that do not show signs in the first 12–24 hours having a better prognosis for long-term survival. In addition to the above, sometimes hemorrhages contained within the broad ligament may go undetected until a pre-breeding examination of the reproductive tract is performed. Mares that show persistent signs of abdominal pain or blood loss soon after parturition should be evaluated by ultrasonography and only gentle palpation of the uterus and broad ligaments. Both rectal and transabdominal ultrasonography are useful in the diagnosis and management of cases of periparturient hemorrhage in the mare (Figure 223.11a–c).
The fetal membranes are usually expelled 30 min to 3 hours after parturition. Horses in a natural environment may retain fetal membranes in the uterus for up to 24–48 hours without complications. These mares may produce a foal the same time the following year without treatment, which suggests that treatment is not always necessary. In an intensive management environment, mares may experience complications (endometritis, metritis, laminitis, and even death) that are related to retention of the fetal membranes 8 h or less. The percentage of postparturient mares with retained fetal membranes is believed to range from 2 to 10%101, 102 (see Chapter 260). The probability of retained placenta increases after dystocia, probably as result of trauma to the uterus, or myometrial
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(c) Figure 223.11 (a) Post-foaling middle uterine artery rupture the day of foaling (day 0). (b) Same mare as in (a), on day 26, demonstrating significant reduction in size. (c) Same mare as in (a) and (b), on day 58.
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Uterine Abnormalities exhaustion. Disturbance of the normal uterine contractions at parturition also might make retained placenta more likely. Retention is particularly likely if severe placentitis was present. Placental retention is more common in the non-gravid uterine horn, perhaps because of a progressive increase in the degree of placental folding and attachment from the gravid to the non-gravid horn. Partial placental retention is also more likely to occur in the non-gravid uterine horn)103 because the chorioallantoic membrane is thinner, resulting in easier tearing (Figure 223.12a,b). Ultrasonography can be used to visualize small pieces of retained fetal membranes in a postpartum mare, as they are typically surrounded by fluid. A variable portion of the placenta may be exposed through the vulvar opening. Occasionally, the veterinarian is alerted to the possibility of retained fetal membranes, when no placenta is found after foaling. Aseptic intrauterine examination may reveal the presence of the placenta in the uterine cavity. Alternatively, part of the placenta may remain in the uterus and continue to initiate mild straining or colic after most of the placenta has been removed. Not uncommonly, fetal membranes may be found days later when the mare has signs of discharge. Sequelae of retained placenta vary from none to the development of toxic metritis, septicemia or endotoxemia, laminitis, and death104 or uterine prolapse. Severe toxic metritis and laminitis after dystocia are believed to result from damage to uterine lining, that subsequently promotes absorption of endotoxins associated with the common bacterial proliferation in the postpartum mare. In patients with toxic metritis after dystocia and retained placenta, the uterine wall becomes thin and friable or even necrotic. Absorption of bacteria and bacterial toxins probably follows disruption of the endometrium and precipitates the peripheral vascular changes that lead to laminitis.104 On occasion fetal membranes may be detected with ultrasonography, days or even weeks after foaling. In addition, ultrasonographic examination will provide information regarding uterine involution and the presence of fluid within the uterine lumen (see above).
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Evaluation of the caudal reproductive tract
Figure 223.12 (a) A retained portion of placenta is visible using ultrasonography in this postpartum mare. (b) A piece of the non-pregnant horn was retained in this mare’s uterus.
Ultrasonography has not been particularly useful for diagnosis of cervical conditions; however, on a few occasions we have identified pathology such as cysts or scarring of the cervix. In general, manual or speculum examinations are more useful. Once the reproductive tract is fully involuted and the mare is in diestrus (when the cervix should be completely
closed), the cervix can be evaluated to determine whether it is capable of closing. Involution is also necessary to allow identification of the muscular and mucosal layers that must be debrided and sutured. We commonly find unidentified cervical tears when
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Figure 223.13 Ultrasonography was useful to visualize urine in the cranial vagina of this mare with vesico-vaginal reflux.
referred cases of problem mares. In addition, it is surprising how many mares with cervical defects will become pregnant. Ultrasonography can be useful in diagnosing cases of vaginal hematomas and abscesses (see above). Careful evaluation may even reveal urine or other fluid in the vagina or vestibule in mares with reproductive problems (Figure 223.13).
References 1. Hayes KEN, Pierson RA, Scraba ST, Ginther OJ. Effects of estrous cycle and season on ultrasonic uterine anatomy in mares. Theriogenology 1985;24:465–77. 2. McKinnon AO, Squires EL, Voss JL. Ultrasonic evaluation of the mare’s reproductive tract: Part I. Compend Cont Educ Pract Vet 1987;9:336–45. 3. Samper JC. Ultrasonograhic appearance and the pattern of uterine edema to time of ovulation in the mare. Proc Am Assoc Equine Pract 1997;43(189):191. 4. Griffin PG, Carnevale EM, Ginther OJ. Effects of the embryo on uterine morphology and function in mares. Anim Reprod Sci 1993;31:311–29. 5. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproductive tract of mares after parturition: Effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed first postpartum ovulatory cycles. J Am Vet Med Assoc 1988;192:350–353. 6. Griggers S, Paccamonti DL, Thompson RA, Eilts BE. The effects of ph, osmolarity and urine contamination on equine spermatozoal motility. Theriogenology 2001;56(4):613–22. 7. McKinnon AO, Voss JL. Breeding the problem mare. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, London: Lea & Febiger, 1993; pp. 368–78. 8. Pycock JF. Assessment of oxytocin and intrauterine antibiotics on intrauterine fluid and pregnancy rates in the mare. Proc Am Assoc Equine Pract 1994;40:19–20. 9. Pycock JF, Newcombe JR. Assessment of the effect of three treatments to remove intrauterine fluid on pregnancy rate in the mare. Vet Rec 1996;138(14):320–3.
10. Causey RC. Making sense of equine uterine infections: the many faces of physical clearance. Vet J 2006;172(3): 405–21. 11. McKinnon AO, Squires EL, Carnevale EM, Harrison LA, Frantz DD, McChesney AE Shideler RK. Diagnostic ultrasonography of uterine pathology in the mare. Proc Am Assoc Equine Pract 1987; 605–22. 12. Adams GP, Kastelic JP, Bergfelt DR, Ginther OJ. Effect of uterine inflammation and ultrasonically-detected uterine pathology on fertility in the mare. J Reprod Fertil Suppl 1987;35:445–54. 13. Ginther OJ, Garcia MC, Bergfelt DR, Leith GS, Scraba ST. Embryonic loss in mares: Pregnancy rate, length of interovulatory intervals, and progesterone concentrations associated with loss during days 11 to 15. Theriogenology 1985;24:409–18. 14. Knutti B, Pycock JF, Vanderweijden GC, Kupfer U. The influence of early postbreeding uterine lavage oil pregnancy rate in mares with intrauterine fluid accumulations after breeding. Equine Vet Educ 2000;12(5):267–70. 15. Malschitzky E, Schilela A, Mattos ALG, Garbade P, Gregory RM, Mattos RC. Effect of intra-uterine fluid accumulation during and after foal-heat and of different management techniques on the postpartum fertility of thoroughbred mares. Theriogenology 2002;58(2–4):495–8. 16. Malschitzky E, Schilela A, Mattos ALG, Garbade P, Gregory RM, Mattos RC. Pregnancy and embryo loss rates in non-lactating mares bred in the first or in other estrus cycles during the breeding season. Pferdeheilkunde 2003;19(6):641–5. 17. Malschitzky E, Schilela A, Mattos ALG, Garbade P, Gregory RM, Mattos RC. Intrauterine fluid accumulation during foal heat increases embryonic death. Pferdeheilkunde 2003;19(6):646–9. 18. Nikolakopoulos E, Watson ED. Uterine contractility is necessary for the clearance of intrauterine fluid but not bacteria after bacterial infusion in the mare. Theriogenology 1999;52(3):413–23. 19. Ozgen S, Schoon HA, Aupperle H, Sieme H, Klug E. Etiopathogenesis of equine intrauterine fluid accumulation. Pferdeheilkunde 2002;18(6):594–9. 20. Troedsson MT. Therapeutic considerations for matinginduced endometritis. Pferdeheilkunde 1997;13(5):516–20. 21. Watson ED, Barbacini S, Berrocal B, Sheerin O, Marchi V, Zavaglia G, Necchi D. Effect of insemination time of frozen semen on incidence of uterine fluid in mares. Theriogenology 2001;56(1):123–31. 22. Schilela A, Malschitzky E, Mattos AL, Garbade P, Gregory RM, Mattos RC. Effect of an intra-uterine fluid accumulation before and after the first postpartum ovulation on pregnancy rates in the mare. Pferdeheilkunde 2001;17(6):639–43. 23. Merkt H, Gunzel AR. A survey of early pregnancy losses in West German thoroughbred mares. Equine Vet J 1979;11:256–8. 24. Lowis TC, Hyland JH. Analysis of post-partum fertility in mares on a thoroughbred stud in southern Victoria. Aust Vet J 1991;68(9):304–6. 25. Woods GL, Baker CB, Baldwin JL, Ball BA, Bilinski J, Cooper WL, Ley WB, Mank EC, Erb HN. Early pregnancy loss in brood mares. J Reprod Fertil Suppl 1987;35:455–9. 26. Sullivan JJ, Turner PC, Self LC, Gutteridge HB, Bartlett DE. Survey of reproductive efficiency in the Quarter-horse and Thoroughbred. J Reprod Fertil Suppl 1975;(23):315–18.
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Uterine Abnormalities 27. Ishii M, Shimamura T, Utsumi A, Jitsukawa T, Endo M, Fukuda T, Yamanoi T. Reproductive performance and factors that decrease pregnancy rate in heavy draft horses bred at the foal heat. J Equine Vet Sci 2001;21(3): 131–6. 28. Lieux P. Comparative results of breeding on first and second post-foaling heat periods. Proc Am Assoc Equine Pract 1980; 129–32. 29. Platt H. Aetiological aspects of abortion in the Thoroughbred mare. J Comp Pathol 1973;83:199–205. 30. Loy RG, Evans MJ, Pemstein R, Taylor TB. Effects of injected ovarian steroids on reproductive patterns and performance in post-partum mares. J Reprod Fertil Suppl 1982;32:199–204. 31. Pope AM, Campbell DL, Davidson JP. Endometrial histology and post-partum mares treated with progesterone and synthetic GnRH (AY-24,031). J Reprod Fertil Suppl 1979;(27):587–91. 32. Sexton PE, Bristol FM. Uterine involution in mares treated with progesterone and estradiol-17-beta. J Am Vet Med Assoc 1985;186:252–6. 33. Squires EL, Heesemann CP, Webel SK, Shideler RK, Voss JL. Relationship of altrenogest to ovarian activity, hormone concentrations and fertility of mares. J Anim Sci 1983;56:901–10. 34. Evans MJ, Hamer JM, Gason LM, Graham CS, Asbury AC, Irvine CHG. Clearance of bacteria and non-antigenic markers following intra- uterine inoculation into maiden mares: Effect of steroid hormone environment. Theriogenology 1986;26:37–50. 35. Winter AJ. Microbial immunity in the reproductive tract. J Am Vet Med Assoc 1982;181:1069–73. 36. Glatzel PS, Belz JP. Fertility in mares after disturbed or undisturbed puerperal periods – significance of clinical, microbiological and hormonal examinations. Berl Mun Tierarztl Wochenschr 1995;108:367–72. 37. McCue PM, Hughes JP. The effect of postpartum uterine lavage on foal heat pregnancy rate. Theriogenology 1990;33(5):1121–9. 38. Blanchard TL, Varner DD, Brinsko SP, Meyers SA, Johnson L. Effects of postparturient uterine lavage on uterine involution in the mare. Theriogenology 1989;32(4): 527–35. 39. Hinrichs K, Spensley MS, McDonough PL. Evaluation of progesterone treatment to create a model for equine endometritis. Equine Vet J 1992;24:457–61. 40. LeBlanc MM. When to refer an infertile mare to a theriogenologist. Theriogenology 2008;70(3):421–9. 41. LeBlanc MM. The chronically infertile mare. Proc Am Assoc Equine Pract 2008;54:391–407. 42. Squires EL, Barnes CK, Rowley HS, McKinnon AO, Pickett BW, Shideler RK. Effect of uterine fluid and volume of extender on fertility. Proc Am Assoc Equine Pract 1989; 25–30. 43. Leblanc M, Neuwirth L, Mauragis D, Klapstein E, Tran T. Oxytocin enhances clearance of radiocolloid from the uterine lumen of reproductively normal mares and mares susceptible to endometritis. Equine Vet J 1994;26:279–82. 44. Combs CB, LeBlanc MM, Neuwirth L, Tran TW. Effects of prostaglandin-F2-alpha, cloprostenol and fenprostalene on uterine clearance of radiocolloid in the mare, Theriogenology 1996;45(8):1449–55. 45. LeBlanc MM. Effects of oxytocin, prostaglandin and phenylbutazone on uterine clearance of radiocolloid. Pferdeheilkunde 1997;13(5):483–5.
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46. Vanderwall DK, Woods GL. Effect on fertility of uterine lavage performed immediately prior to insemination in mares. J Am Vet Med Assoc 2003;222(8): 1108–10. 47. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: Technique and preliminary findings. Proc Am Assoc Equine Pract 1975; 327–35. 48. Danek J, Wisniewski E, Krumrych W, Dabrowska J. Prevalence and sensitivity to antibiotics of facultative pathogenic bacteria isolated from the semen of stallions. Med Weter 1994;50:385–7. 49. Vaillancourt D, Guay P, Higgins R. The effectiveness of gentamicin or polymyxin B for the control of bacterial growth in equine semen stored at 20 degrees C or 5 degrees C for up to forty-eight hours. Can J Vet Res 1993;57:277–80. 50. Padilla AW, Foote RH. Extender and centrifugation effects on the motility patterns of slow-cooled stallion spermatozoa. J Anim Sci 1991;69(8):3308–13. 51. Blanchard TL, Varner DD, Love CC, Hurtgen JP, Cummings MR, Kenney RM. Use of a semen extender containing antibiotic to improve the fertility of a stallion with seminal vesiculitis due to Pseudomonas aeruginosa. Theriogenology 1987;28(4):541–6. 52. Arriola J, Foote RH. Effects of amikacin sulfate on the motility of stallion and bull spermatozoa at different temperatures and intervals of storage. J Anim Sci 1982;54:1105–10. 53. Timoney PJ, O’Reilly PJ, Harrington AM, McCormack R, McArdle JF. Survival of Haemophilus equigenitalis in different antibiotic-containing semen extenders. J Reprod Fertil Suppl 1979;(27):377–81. 54. Pickett BW, Squires EL, McKinnon AO. Procedures for Collection, Evaluation and Utilization of Stallion Semen for Artificial Insemination, 2nd edn. Colorado State University: Animal Reproduction Laboratory, 1987. 55. Jasko DJ, Bedford SJ, Cook NL, Mumford EL, Squires EL, Pickett BW. Effect of antibiotics on motion characteristics of cooled stallion spermatozoa. Theriogenology 1993;40:885–93. 56. Francl AT, Amann RP, Squires EL, Pickett BW. Motility and fertility of equine spermatozoa in a milk extender after 12 or 24 hours at 20 degrees C. Theriogenology 1987;27(3):517–25. 57. Douglas-Hamilton DH, Osol R, Osol G, Driscoll D, Noble H. A field study of the fertility of transported equine semen. Theriogenology 1984;22:291–304. 58. Pickett BW, Sullivan JJ, Byers WW, Pace MM, Remmenga EE. Effect of centrifugation and seminal plasma on motility and fertility of stallion and bull spermatozoa. Fertil Steril 1975;26:167–74. 59. LeBlanc MM, Neuwirth L, Asbury AC, Tran T, Mauragis D, Klapstein E. Scintigraphic measurement of uterine clearance in normal mares and mares with recurrent endometritis. Equine Vet J 1994;26(2):109–13. 60. Neuwirth L, LeBlanc MM, Mauragis D, Klapstein E, Tran T. Scintigraphic measurement of uterine clearance in mares. Vet Radiol Ultrasound 1995;36:64–8. 61. McKinnon AO, Brown RW, Pashen RL, Greenwood PE, Vasey JR. Increased ovulation rates in mares after immunisation against recombinant bovine inhibin alphasubunit. Equine Vet J 1992;24:144–6. 62. McCue PM, Carney NJ, Hughes JP, Rivier J, Vale W, Lasley BL. Ovulation and embryo recovery rates
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following immunization of mares against an inhibin alpha-subunit fragment. Theriogenology 1992;38(5): 823–31. Squires EL, McKinnon AO, Carnevale EM, Morris R, Nett TM. Reproductive characteristics of spontaneous single and double ovulating mares and superovulated mares. J Reprod Fertil Suppl 1987;35:399–403. Pascoe DR, Stabenfeldt GH, Hughes JP, Kindahl H. Endogenous prostaglandin F-2-alpha release induced by physiologic saline solution infusion in utero in the mare: Effect of temperature, osmolarity, and pH. Am J Vet Res 1989;50:1080–3. Jones DM, Fielden ED, Carr DH. Some physiological and pharmacological factors affecting uterine motility as measured by electromyography in the mare. J Reprod Fertil Suppl 1991;44:357–68. Liu IKM, Troedsson MHT, Williams DC, Pascoe JR. Electromyography of uterine activity in the mare: comparison of single vs multiple recording sites. J Reprod Fertil Suppl 1991;44:744. Rohrbach BW, Sheerin PC, Cantrell CK, Matthews PM, Steiner JV, Dodds LE. Effect of adjunctive treatment with intravenously administered Propionibacterium acnes on reproductive performance in mares with persistent endometritis. J Am Vet Med Assoc 2007;231(1): 107–13. Brendemuehl JP. Effect of oxytocin and cloprostenol on luteal formation, function and pregnancy rates in mares. Theriogenology 2002;58(2–4):623–6. Nie GJ, Johnson KE, Wenzel JG, Braden TD. Luteal function in mares following administration of oxytocin, cloprostenol or saline on day 0, 1 or 2 post-ovulation. Theriogenology 2003;60(6):1119–25. Nie GJ, Johnson KE, Wenzel JG, Braden TD. Effect of administering oxytocin or cloprostenol in the periovulatory period on pregnancy outcome and luteal function in mares. Theriogenology 2003;60(6):1111–18. Gunthle LE, McCue PM, Farquhar VJ, Foglia RA. Effect of prostaglandin administration postovulation on corpus luteum development in the mare. Proceedings of Society for Theriogenology 2000; 139. Kenney RM, Ganjam VK. Selected pathological changes of the mare uterus and ovary. J Reprod Fertil Suppl 1975;23:335–9. McKinnon AO, Squires EL, Voss JL. Ultrasonic evaluation of the mare’s reproductive tract: Part II. Compend Cont Educ Pract Vet 1987;9:472–82. Stanton MB, Steiner JV, Pugh DG. Endometrial cysts in the mare. J Equine Vet Sci 2004;24(1):14–19. Tannus RJ, Thun R. Influence of endometrial cysts on conception rate of mares. J Vet Med A-Zbl Vet A-Physiol 1995;42:275–83. McDowell KJ, Sharp DC, Grubaugh W, Thatcher WW, Wilcox CJ. Restricted conceptus mobility results in failure of pregnancy maintenance in mares. Biol Reprod 1988;39:340–8. Ginther OJ. Fixation and orientation of the early equine conceptus. Theriogenology 1983;19:613–23. DeLuca CA, Gee EK, McCue PM. How to remove large endometrial cysts with an improvised snare: A simple technique for practitioners. Proc Am Assoc Equine Pract 2009;55:328–30. Silva MI, Nascimento EF, Cassali GD. Persistent endometrial cups in mare – case report. Arq Bras Med Vet Zootec 1995;47:31–6.
80. Willis LA, Riddle WT. Theriogenology question of the month. J Am Vet Med Assoc 2005;226(6):877–9. 81. Steiner J, Antczak DF, Wolfsdorf K, Saville K, Brooks S, Miller D, Bailey E, Zent W. Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. 82. Gores-Lindholm A, Causey R, Calderwood-Mays M, LeBlanc MM. Effect of intra-uterine infusion of diluted N-acetylcysteine on equine endometrium. Proc Am Assoc Equine Pract 2009;555:326. 83. Barber JA, Troedsson MH. Mummified fetus in a mare. J Am Vet Med Assoc 1996;208(9):1438–40. 84. McCue PM, Vanderwall DK, Squires EL. Fetal mummification in a mare. J Equine Vet Sci 1997;17(5):267–9. 85. Meyers PJ, Varner DD. Abortion of a mummified fetus associated with short uterine body in a mare. J Am Vet Med Assoc 1991;198(10):1768–70. 86. Waelchli RO, Ehrensperger F. Two related cases of cerebellar abnormality in equine fetuses associated with hydrops of fetal membranes. Vet Rec 1988;123: 513–14. 87. Blanchard TL, Varner DD, Buonanno AM, Palmer JE, Hansen TO, Divers TJ. Hydrallantois in two mares. J Equine Vet Sci 1987;7:222–5. 88. Allen WE. Two cases of abnormal equine pregnancy associated with excess foetal fluid. Equine Vet J 1986;18: 220–2. 89. Lofstedt RM. Miscellaneous diseases of pregnancy and parturition. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, London: Lea & Febiger, 1993; pp. 596–603. 90. Christensen BW, Troedsson MH, Murchie TA, Pozor MA, Macpherson ML, Estrada AH, Carillo NA, Mackay RJ, Roberts GD, Langlois J. Management of hydrops amnion in a mare resulting in birth of a live foal. J Am Vet Med Assoc 2006;228(8):1228–33. 91. Sertich PL, Reef VB, Oristaglioturner RM, Habecker PL, Maxson AD. Hydrops amnii in a mare. J Am Vet Med Assoc 1994;204:1481–2. 92. Jackson PG. Rupture of the prepubic tendon in a shire mare. Vet Rec 1982;111(2):38. 93. Ross J, Palmer JE, Wilkins PA. Body wall tears during late pregnancy in mares: 13 cases (1995–2006). J Am Vet Med Assoc 2008;232(2):257–61. 94. Chaney KP, Holcombe SJ, LeBlanc MM, Hauptman JG, Embertson RM, Mueller PO, Beard WL. The effect of uterine torsion on mare and foal survival: a retrospective study, 1985 – 2005. Equine Vet J 2007;39(1):33–6. 95. Wichtel JJ, Reinertson EL, Clark TL. Nonsurgical treatment of uterine torsion in seven mares. J Am Vet Med Assoc 1988;193:337–8. 96. Honnas CM, Spensley MS, Laverty S, Blanchard PC. Hydramnios causing uterine rupture in a mare. J Am Vet Med Assoc 1988;193:334–6. 97. Vandeplassche M. Obstetrician’s view of the physiology of equine parturition and dystocia. Equine Vet J 1980;12:45–9. 98. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992. 99. Hodge SL, Kreider JL, Potter GD, Harms PG, Fleeger JL. Influence of photoperiod on the pregnant and postpartum mare. Am J Vet Res 1982;43(10):1752–5. 100. Green EM, Loch WE, Messer NT. Maternal and fetal effects of endophyte fungus-infected fescue. Proc Am Assoc Equine Pract 1991; 29–44.
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Uterine Abnormalities 101. Vandeplassche M, Spincemaille J, Bouters R. Aetiology, pathogenesis and treatment of retained placenta in the mare. Equine Vet J 1971;3:144–7. 102. Threlfall WR. Retained placenta. In: McKinnon AO, Voss JL, (eds) Equine Reproduction. Philadelphia, London: Lea & Febiger, 1993; pp. 614–21.
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103. Perkins NR, Frazer GS. Reproductive emergencies in the mare. Vet Clin North Am Equine Pract 1994;10(3): 643–70. 104. Blanchard TL, Elmore RG, Varner DD, Garcia MC, Orsini JA, Youngquist RS. Dystocia, toxic metritis and laminitis in mares. Proc Am Assoc Equine Pract 1987;33:641–8.
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Chapter 224
Mammary Gland Bryan M. Waldridge
Introduction Mastitis, neoplasia, and other diseases of the mare’s udder are relatively uncommon. Ultrasonography is very useful for imaging and detecting abnormalities of the equine mammary gland. Suspected lesions are often discreet, and ultrasonography allows better determination of the size and extent of possible pathology. Ultrasound examination of the equine1 and bovine2, 3 mammary gland has been previously described. Ultrasonography in mares has been used to diagnose mammary gland adenocarcinoma, lymphangiectasia, and intramammary sarcoids.4 In cattle, ultrasonography aids in the diagnosis of abscesses, mastitis, obstructive lesions, teat malformations, fistulous tracts, and glandular fibrosis and atresia.2 Flock ¨ and Winter3 reported that in cows, Arcanobacterium pyogenes and Gramnegative pathogens caused more severe ultrasonographic changes than Gram-positive pathogens such as Streptococcus species. Ultrasound examinations may also be used to detect prohibited alterations to improve udder appearance in show cows.5, 6
Mammary gland anatomy The equine mammary gland is divided into right and left halves formed into laterally compressed cones. Each half of the mammary gland is usually also divided into separate cranial and caudal glandular systems. Each individual glandular system will have its own associated teat orifice (Figure 224.1). Most often there are two or rarely three teat orifices present for each side of the mammary gland. The teat orifice opens into a short lactiferous sinus (teat sinus) that receives milk from its system of lactiferous ducts.7
Ultrasound examination The mammary gland can be examined with any ultrasound probe of frequency 5 MHz or higher. Lesions
can be small or fairly inconspicuous and will be better imaged using higher frequency probes. Linear array probes are probably easiest to use, although this is dependent on the size and shape of the udder examined; other probes can also provide adequate diagnostic images. The udder should be cleaned of external debris; excellent image quality can be obtained by applying isopropyl alcohol to the udder skin to maintain probe contact. The most common difficulty encountered during ultrasound examination of the equine udder is excessive digital pressure with the probe that obliterates the contrast effect of mammary secretions. If the mare is lactating, it is very important to apply just enough contact pressure so as not to overly compress the fluid contents of the mammary gland. Standoff pads or probes with built-in fluid offset may improve ultrasound imaging of the udder, especially the teat.2, 4 Most mares are fairly tolerant of the procedure and can be examined without sedation. However, the examiner should be careful to position him or herself and the ultrasound machine in a safe location when examining maiden mares or mares with potentially very painful mastitis. Normal ultrasound examination Normal mammary gland parenchyma has a homogeneous hyperechoic appearance. In lactating mares, the large lactiferous ducts can be readily identified within the glandular tissue as they coalesce into progressively larger diameter lactiferous ducts and finally enter the lactiferous sinus in the teat. Lactating mares and mares in late gestation have an increased number and diameter of lactiferous ducts, compared to non-pregnant or non-lactating mares. Some mares may chronically lactate or normally have a small accumulation of mammary secretions in the lactiferous sinuses that cannot be easily followed proximally into the lactiferous ducts. The prominent hyperechoic and linear interglandular septum can be found
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 224.1 Plastinated cast of one side of a mammary gland from a non-lactating pony mare. The cranial and caudal glandular systems have been separated to better illustrate the lactiferous sinuses and lactiferous duct systems.
parallel to the long axis of the teat and dividing the individual glandular systems and lactiferous sinuses (Figures 224.2 and 224.3). Mammary gland blood vessels are more readily identified in lactating mares. Blood vessels appear thinner walled than lactiferous ducts and have a more hypoechoic lumen (Figure 224.4). However, some arteries may appear somewhat hyperechoic due to turbulent blood flow. Arteries may be seen to pulse with the heartbeat when the ultrasound probe is held in place. Color flow Doppler imaging gives the most useful information definitively to distinguish mammary gland blood vessels from lactiferous ducts.1
Figure 224.2 Lactiferous sinus and lactiferous ducts in a lactating mare. The lactiferous sinus can be seen in the upper right of the figure and two lactiferous ducts can be identified emptying into the lactiferous sinus. This image was obtained using a 7.5-MHz linear array probe. Ventral is to the right of the figure.
Figure 224.3 Lactiferous sinus and lactiferous ducts in a lactating mare. The lactiferous sinus can be identified in the upper right of the figure and several lactiferous ducts can be seen near and emptying into the lactiferous sinus. The prominent, linear interglandular septum is seen diagonally. This image was obtained using a 7.5-MHz linear array probe. Ventral is to the right of the figure.
Abnormal ultrasound findings In cases of mastitis, the milk will often have a hyperechoic, heterogeneous appearance (Figure 224.5). Both chronic intramammary infections and mastitis
Figure 224.4 Lactiferous sinus and mammary gland blood vessel in a non-lactating mare. Note that few lactiferous ducts can be identified, in contrast to lactating mares. The Y-shaped lactiferous duct has a more hyperechoic and prominent inner wall than the more rounded blood vessel. The interglandular septum can be seen near the top of the figure. This image was obtained using a 7.5-MHz linear array probe. Ventral is to the right of the figure.
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Figure 224.5 Mastitis in a lactating mare. The mammary secretions appear heterogeneous and more hyperechoic than normal milk. The mammary gland parenchyma also appears more hyperechoic. The lactiferous sinus is to the upper right of the figure and the interglandular septum can be seen to the lower right. This image was obtained using a 7.5-MHz linear array probe. Ventral is to the right of the figure.
Figure 224.6 Lactiferous sinuses in a lactating mare. The lactiferous sinuses are in the upper right of the figure. There is an adhesion within the upper lactiferous sinus and a proliferative lesion within the lower lactiferous sinus. This image was obtained using a 7.5-MHz linear array probe. Ventral is to the right of the figure.
may lead to cystic or cavitated areas in an enlarged or painful mammary gland. Glandular tissue may also appear more echodense and hyperechoic due to fibrosis or inflammatory cell infiltrate.4 Bovine colostrum may have an ultrasonographic appearance similar to mastitic milk, although colostrum initially has a more consistently uniform homogeneous echogenicity.2 Therefore, diagnosis of mastitis during early lactation by ultrasound examination alone may be misleading. Most cases of equine mastitis do not result in permanent glandular fibrosis or obstruction (Figure 224.6). Obstructive lesions are uncommon, but may be better observed with the infusion of sterile saline into the lactiferous sinus or ducts.2 Abscesses may sometimes occur with Streptococcus equi or other infections. Neoplasia is identified as focal, heterogeneous composite masses with marked loss of normalappearing glandular parenchyma.4
References 1. Waldridge BM, Ward TA. Ultrasound examination of the equine mammary gland. Equine Pract 1999;21:10–13. 2. Trostle SS, O’Brien RT. Ultrasonography of the bovine mammary gland. Comp Cont Educ Pract Vet 1998;20: S64–S74. ¨ 3. Flock M, Winter P. Diagnostic ultrasonography in cattle with diseases of the mammary gland. Vet J 2006;171:314–21. 4. Reef VB. Ultrasonographic evaluation of small parts. In: Equine Diagnostic Ultrasound. Philadelphia: WB Saunders, 1998; pp. 480–547. 5. O’Brien RT, Waller KR, Matheson JS. Ultrasonographic appearance of edema caused by injections in the mammary gland attachments of dairy cows. J Am Vet Med Assoc 2002; 221:408–10. 6. Trostle SS, O’Brien RT, Britt J, Waller KR. Ultrasonographic appearance of exogenous isobutane gas in the mammary glands of dairy cows. J Am Vet Med Assoc 1999;215:366–8. 7. Dyce KM, Sack WO, Wensing CJG. The pelvis and reproductive organs of the horse. In: Textbook of Veterinary Anatomy. Philadelphia: WB Saunders, 1987; pp. 522–41.
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Embryo Morphology, Growth and Development Keith J. Betteridge
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Fetal Membrane Differentiation, Implantation and Early Placentation W.R. (Twink) Allen, Susan Gower and Sandra Wilsher
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Maternal Recognition of Pregnancy Karen J. McDowell and Daniel C. Sharp
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Sex Determination and Differentiation Bruce W. Christensen and Vicki N. Meyers-Wallen
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Endocrinology of Pregnancy Jennifer C. Ousey
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Development and Morphology of the Placenta Sandra Wilsher and W.R. (Twink) Allen
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Pregnancy Examination Patrick M. McCue and Angus O. McKinnon
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Induction of Parturition Margo L. Macpherson and Dale L. Paccamonti
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Parturition Bruce W. Christensen
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Lactation Patrick M. McCue and Sofie Sitters
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Uterine Involution Mary E. Stanton
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Breeding Mares on Foal Heat Terry L. Blanchard and Margo L. Macpherson
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Inter and Extraspecies Equine Pregnancies W.R. (Twink) Allen, Julia H. Kydd, Roger V. Short and Douglas F. Antczak
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Induction of Abortion Claire E. Card
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Chapter 225
Embryo Morphology, Growth and Development Keith J. Betteridge
The reasons for regarding the first month of pregnancy as being of especial importance in equine reproduction all hinge on the concept that the embryo, from the beginning, participates in a ‘dialogue’ with the mare and in the establishment of an environment in which its gestation can continue. This concept underlies our understanding of how pregnancies are maintained or fail, and rationalizes the idea that the early uterine environment influences development with effects that extend into fetal, postnatal, and adult life and even into subsequent generations.1–4 It is a concept that applies to all mammals; however, the horse has played such an important role in the development of this concept over the past 50 years that it is appropriate to begin this chapter with a summary of how equine reproduction has contributed to today’s acceptance of the fact that embryos ‘talk’ and ‘listen’. There was a time that embryos, and indeed fetuses, were regarded more as passengers in the uterus than as active participants in pregnancy; they would be gestated and born under maternal control. Such a notion, which persisted until the early 1960s, is understandable in altricial species such as humans, in which total dependence on the mother continues for long after birth. Most newborn domestic animals, however, are relatively independent and watching their earliest activities makes it easy to understand that they must have been almost as physiologically active in utero at term, just a few hours earlier. That such is the case – and that the fetus at term plays an active role in determining when parturition should begin – became established after clinical observations of prolonged pregnancies in humans5 and domestic animals6 led to speculation that birth requires an intact hypothalamo–pituitary–adrenal axis in the fe-
tus.7 This contention was confirmed by classic endocrinological experimentation in sheep5, 8 and was not difficult to accept given the resemblance of the term fetus to the neonate. Gradually, however, it became apparent that the concept of ‘fetal autonomy’ applied to much less mature fetuses at progressively earlier stages of their gestation and could be extended to encompass a degree of ‘embryonic autonomy’ soon after fertilization. Figure 225.1 illustrates this progression for the horse.
Communication in the latter half of pregnancy After Zondek9 first reported that the urine of pregnant mares contains estrogens, much effort was devoted to establishing their source, chemistry, biological function, and pharmaceutical usefulness.10–15 Of cardinal importance is the relationship between the pattern of excretion of the estrogens (Figure 225.1) and the parallel enlargement and subsequent regression of the fetal gonads, first recognized by Cole et al.16 Once it became clear that the estrogens excreted in the mare’s urine had originated from placental metabolism of precursors synthesized in the fetal gonads,10 it was easy to accept that the fetus – with such an obviously developed endocrine and reproductive system – was playing an active role in its own gestation well before parturition. Similarly, research into the mechanisms of sexual differentiation, particularly by Alfred Jost, provided incontrovertible evidence that fetuses have autonomous endocrine systems at stages much earlier than term.17 Even at these earlier stages, however, the concept was relatively easy to accept, given that the fetal gonads and target organs involved are easily recognizable.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Months gestation Figure 225.1 Diagrammatic representation of how the concept of ‘fetal autonomy’ in the mare has extended to earlier and earlier stages of pregnancy following its initial demonstration at term. Key events of pregnancy are indicated in black; fetal and embryonic involvements are in gray. For details, see text.
Communication in the first half of pregnancy The era of urinary and serum gonadotropin discoveries in humans and horses during the 1920s and 1930s18, 19 is of special relevance to the growth of the concept of ‘fetal autonomy’. The realization that the gonadotropic activity of urine from pregnant women is due to a product of the conceptus, and the consequent naming of human chorionic gonadotropin (hCG), came early. The corresponding story in mares is more convoluted. The gonadotropic activity was found in the serum of pregnant mares rather than in their urine and was therefore named ‘pregnant mare serum gonadotropin’ or PMSG. Later, it was discovered that PMSG activity could be recovered from endometrial cups – the ulcerous structures that form opposite the chorionic girdle of the conceptus. Assays of PMSG, developed primarily as a means of diagnosing pregnancy, also showed that the levels found in serum between days 40 and 120 of pregnancy varied a great deal from mare to mare. Investigations of horse × donkey hybrid pregnancies in Poland first revealed that this variation is genetically controlled20 and the extensive studies of Allen and his colleagues in Cambridge beginning during the 1970s showed that the genetic variation in gonadotropin production is because the hormone is of fetal origin21 and should be called equine chorionic gonadotropin (eCG). Accepting this evidence of a ‘fetal’ contribution to the maintenance of pregnancy was made more difficult by the facts that (i) the embryo/fetus is much less well formed than at later stages when a fetal endocrine system is easy to envisage and (ii) the cells of the conceptus that provide the contribution are not in the embryo/fetus but in the extraembryonic membranes. Acceptance of the idea that conceptus tissues other than the embryo proper are involved in signaling to the mother resulted from embryo transfer work
in the 1960s (particularly by Rowson and Moor in Cambridge) that demonstrated the existence in ruminants of a ‘critical period’ for the ‘recognition of pregnancy’. The latter terms were coined at an important symposium on fetal autonomy.22 In my opinion, it is not appropriate to apply these terms to the establishment and maintenance of pregnancy in the mare in which these processes seem to be more of a continuum.23, 24 Nevertheless, the mare has provided especially valuable evidence of the autonomous activity of the embryo from the earliest cleavage stage; it was in horses that differential transport of fertilized and unfertilized eggs through the oviduct was first reported.24, 25 The gray question mark in Figure 225.1 is to indicate that much remains to be understood about exactly how a few embryonic cells can have such profound effects on the mother. Today, the idea of interaction with the maternal environment has been extended back to considerations of how embryos affect the oviduct,26 of how even a single cell – the follicular oocyte – influences ovulation rates in sheep,27 and how even the environment of that cell may influence the sex of an embryo resulting from its fertilization; in cattle, high levels of testosterone in follicular fluid increase the chances of its contained oocyte being fertilized by a Y-bearing spermatozoon in vitro.28 Discovery of how these earliest interactions are effected will require all the techniques of molecular and cell biology and it is useful to consider the horse as a model system in which to apply them. Such consideration requires knowledge of the roles of both partners in the conceptus–mare dialogue and so the balance of this chapter will summarize what we know – and, at least as important, what we do not know – about the partnership during the first month of pregnancy. Needless to say the choice of this relatively short period has been influenced by the author’s own research interests but it is not difficult to offer more objective reasons, too. First of all, the period covers the interval during which
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Embryo Morphology, Growth and Development embryo transfers and manipulations are possible in the horse. Second, it is a particularly important time for determining whether an established pregnancy will continue to term; for example, in their surveys of Thoroughbred studs in Newmarket, UK, Morris and Allen29 showed that 16–17% of pregnancies diagnosed by ultrasound at about day 15 are subsequently lost, 60% of them between days 15 and 35. Third, several aspects of equine pregnancy during the first month contrast with analogous events in other domestic species; understanding similarities and differences of reproductive strategies among species is fundamental to realizing the benefits of a comparative approach to reproductive biology. Fourth, it is now very clear that events during the earliest days of gestation are at the heart of the concept of ‘developmental origins of health and disease’ that is so impor-
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tant to human and veterinary medicine, as mentioned in the opening paragraph of this chapter. With these reasons in mind, interactions between the equine embryo and its environment during the first month of pregnancy will be discussed chronologically (Figure 225.2), starting from the time of ovulation and dividing the 4 weeks into six phases on physiological grounds, as in a recent review.30 The rapidity of early developmental changes makes it important to define the time of ovulation as accurately as possible and to recognize that it will always be subject to an error governed by the frequency of ovarian examination. With daily ultrasonography, for example, the first day on which a ruptured follicle is detected can be described as day 0.5 ± 0.5 because the rupture could have occurred at any time during the 24-hour interval between examinations.
0 2-cell (~24 h) 4-cell (24-48 h); Initial activation of embryonic genome 8-cell (72 h) Compaction (8- to 16-cell); principal activation of embryonic genome Cavitation (~120 h) Morula/Blastocyst enters the uterus
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Distinct embryonic shield; primitive streak First pairs of somites Fixation; loss of sialic acid from capsule Angiogenesis
Completion of amnion; blood in sinus terminalis
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Conceptus buoyant in saline
Intact capsule
Increasing osmolality and concentrations of fructose, oxytocin and arginine vasopressin in yolk-sac fluid
Emergence of allantois; Functional choriovitelline placenta Limb buds; heartbeat
Conceptus immobile at site of placentation
Allantochorionic placenta developing
28 Figure 225.2 Approximate timing of some key developmental events during the first 4 weeks of gestation of the equine conceptus. Adapted from Betteridge.24
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Phase 1: the oocyte and early cleavage stage embryos; day 0 (ovulation) to day 6
organelles and inclusions in the equine MII oocyte, before and after ovulation, and in the early zygote (Figure 225.3).30 The observations, which date to the 1980s, attracted little attention until the clear demonstration of polarity in mouse and human oocytes in the late 1990s renewed interest in the topic and led to speculation about how polarity might affect techniques such as ICSI, nucleus transfer, and cytoplasm transfer. Currently, there is considerable debate about the putative role of polarity in determining body axes, and of how oocyte polarity might be induced in the ovary. The interest is more than academic because polarity may be influenced by follicular cells communicating with the oocyte through transzonal processes – processes that could be disrupted by procedures such as oocyte maturation in vitro. Thus, the possibility that suboptimal maturation in vitro might adversely affect oocyte quality (and subsequent embryo development) through an effect on polarity cannot be ignored. It has also been suggested that cell surface charge polarity might be exploited in the mechanization of somatic cell nucleus transfer.36 However, it should be emphasized that putative relationships of oocyte polarity and first cleavages to body patterning remain extremely controversial, even in the mouse, and the inevitable limitations of experimental studies of early development in horses make it most unlikely that they will help resolve these controversies directly. Observational studies in horses, however, could be useful should sufficient early embryos become available. For example, initial cleavage of the polarized equine zygote often seems to give rise to ‘dark’ and ‘light’ blastomeres (Figure 225.3c,d) and it would be interesting to determine for how many cleavages these two categories are distinguishable in a population of freshly collected embryos. I have also suggested30 that a complementary approach to such a study could be to follow the cleavage of individual zygotes over time in an adequate culture system, perhaps in co-culture with oviduct cells. Because the early equine embryo is often
For almost all of this period the embryo remains in the oviduct and therefore inaccessible except by surgery31 or slaughter. This has severely limited the study of normal cleavage in vivo and may contribute to the fact that ‘normal’ in vitro fertilization (IVF) (i.e., excluding intracytoplasmic injection (ICSI) and somatic cell nucleus transfer (SCNT)) remains so lamentably unsuccessful in equids.32 The rates of fertilization following natural service are high and can exceed 90%.33 The most precise study of the progression of early cleavage in equine embryos is that of B´ezard et al.,34 who recovered them surgically after inducing ovulation with a pituitary extract and timing ovulation by ultrasonography at hourly intervals. At 6 hours after ovulation, oocytes were still enshrouded with cumulus cells and could not be examined for signs of fertilization. However, of a total of 18 oocytes or embryos recovered between 12 and 96 hours after ovulation, 16 (89%) were fertilized and cleaved in close synchrony (Table 225.1). The appearance of selected early cleavage stages is illustrated in Figure 225.3. Paternal and maternal pronuclei become visible by 12 hours postovulation34 and migrate into apposition within the subsequent 7 hours.35 Several features of embryonic development in horses during this time are of comparative interest and may provide insight into events in the oviduct that are essential to pregnancy and difficult to reproduce in vitro. Among these are: the pronounced polarity of the equine oocyte and early embryo; the coatings that are deposited on the outside of the zona pellucida; the manner in which the inner cell mass begins to become segregated; and the remarkable phenomenon of transit of embryos through the oviduct while unfertilized oocytes are retained there. Several investigators have been struck by the marked asymmetries in the distribution of cellular
Table 225.1 Numbers of equine embryos at various cleavage stages during the first 4 days after ovulation Number of cells per embryo Hours post ovulation 6 12 24 48 72 96 6–96
1 6 4 1
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Data from Bezard et al.34 ´
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Figure 225.3 Polarity in the equine oocyte and apparent segregation of zygote content during early cleavage. (a) An aspirated oocyte–cumulus complex. (b) A sectioned MII oocyte. (c) A three-cell embryo with one blastomere darker than the other two. (d) A four-cell embryo with two blastomeres darker than the other two. Note the ‘debris’ in the perivitelline space of (c) and (d), perhaps unusually prominent in these particular embryos. Scale bars: 100 µm for (a); 50 µm for (b–d). For sources, see Betteridge.30
ellipsoidal (Figure 225.3c,d) it would be particularly interesting to determine whether the physical restraint imposed on the zygote by this shape affects the plane of first cleavage, as it apparently does in mice. Oviductal equine embryos, if only they were more readily accessible, would be a valuable resource for studying the nature of early embryonic coats. These coats are more complex than the single term ‘zona pellucida’ implies but, until recently, they had been described in only rather general terms30 and undoubtedly deserve more attention, given their relevance to fertilization and early embryonic development in all farm species and their fascinating evolu-
tionary history37 and comparative differences.38 The relationship between embryonic coats and the socalled perivitelline space should also be borne in mind; the prominence of ‘debris’ around the blastomeres of some early cleaving horse embryos (Figure 225.3c,d) serves as a reminder that the perivitelline ‘space’ has, until recently, been neglected as an important structure. Knowing more about it could help our understanding of fertilization and embryogenesis because the ‘space’ is in fact filled with an extracellular matrix of unknown function. Segregation of the inner cell mass from the trophoblast at the time of blastocyst formation is much less rapid and distinct in horses than in ruminants
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Figure 225.4 An equine blastocyst recovered from the uterus 6.5 ± 0.5 days after ovulation, measuring 162 × 180 µm. (a) As seen with the dissecting microscope. Note the relatively indistinct inner cell mass inside the palisade of differentiated trophoblast. The beginnings of capsule deposition are seen lining the inside of the zona pellucida. (b) Semi-thin section of the same blastocyst, processed for electron microscopy, showing the inner cell mass occupying a large proportion of the blastocyst cavity. From study of Flood et al. 1982; see Betteridge.30
and pigs. Figure 225.4 exemplifies a pattern illustrated in early equine embryos by many investigators since the 1980s;30 the inner cells remain much more dispersed than in ruminants and pigs and fill much more of the space within the distinct palisade of trophoblast cells. This has the important practical effect of making it more difficult to distinguish between morulae and early blastocysts in horses than in the other farm species, a point to bear in mind when considering the stage of development of embryos passing into the uterus at about day 6. Investigations of the mechanisms whereby developing embryos, but not unfertilized oocytes, pass through the mare’s oviduct have not apparently advanced since the work of Woods and his colleagues in the early 1990s implicated embryonic production of prostaglandin (PG) E2 as an effector.17 It seems to me that perhaps the subject deserves more attention. In particular, the probability that oviductins contribute to the embryonic coats of the horse embryo during its stay in the oviduct needs to be studied. The protective, adhesive, and anti-adhesive functions of these secretory mucin-like molecules39 could, perhaps, also be important in the phenomenon. A further attraction of adding the horse to the species in which the oviductins are studied is the possibility that they could be used to improve sperm binding and fertilization in vitro, as in other species.40 M´en´ezo et al.41 – with human embryos – have re-
minded us of the need for rigor in any study of embryonic products presumed to play a role in communication with the mother. In rats, estrogen regulation of estrogen receptor β2 on ciliated oviductal epithelial cells may mediate gamete and embryo transport processes by processing protein–protein interactions.42 Furthermore, estrogens accelerate the passage of ova through the rat oviduct43 and affect the activity of tubal musculature through their effects on tight junctions.44 It has been reported that, by day 6, blastomeres of equine embryos contain the enzyme 5–3β-hydroxysteroid dehydrogenase and that, by day 8, measurable amounts of progesterone, androgens, and estrogens are to be found.45 Considering this, and the abundant production of estrogens and androgens by equine embryos at slightly later stages (see below), the possibility that they also have a role to play in differential transport of fertilized and unfertilized eggs in horses deserves to be investigated. Whatever the mechanism underlying the phenomenon, the selective transport of early embryos through the oviduct underlines that they need to be very metabolically active throughout even their first 6 days of existence. Cell division and differentiation, and concomitant changes in function, all require adenosine triphosphate (ATP), reducing equivalents, and precursors for synthesis of macromolecules; the total nuclear DNA increases 100-fold or more as the
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Embryo Morphology, Growth and Development equine zygote develops into a blastocyst. Unfortunately, the inaccessibility of embryos in the mare’s oviduct has, with the notable exception of a study ¨ and Hyland,46 precluded detailed study of by Bruck their energy metabolism (for example) before they enter the uterus (see below). However, studies in other species strongly suggest that the short- and long-term viability and health of the embryo, fetus, neonate, adult horse, and even the progeny of that adult horse have been irrevocably affected through metabolic interactions by the time the morula or early blastocyst is ready to pass into the uterus (Figure 225.5). Evidence for such a pronouncement is still accumulating,4, 47, 48 but the implications for research into the reproductive management of horses are likely to be important and long-lasting. The in-
PATERNAL GRANDAM
Oocyte
teractions, of course, are between the genotype of the embryo, established at fertilization, and the environment that it encounters in the oviduct. While the multiple epigenetic results of the interactions are beyond the scope of this chapter (for introductory references see Fleming et al.,1 Sinclair et al.,2 Gluckman et al.,3 Leese et al.,4 and Duranthon et al.49 ) and most have yet to be established in the horse specifically, one exception is activation of the embryonic genome which occurs during the fourth cell cycle as the embryo progresses from eight to 16 cells.50 It should also be noted that interbreed embryo transfers in horses have been important for demonstrating that upsetting normal fetal growth patterns in utero can influence postnatal cardiovascular function.51
MATERNAL GRANDAM
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Figure 225.5 Five principles underlying current and future experimental investigation of conceptus growth. 1, Growth of the diploid conceptus (and neonate), represented by the triangles, requires synchrony of developmental events in the conceptus and mother (represented by the interlocking waved borders between the two). 2, This synchrony depends on ‘dialogue’ between the mother and conceptus (double-ended arrows). 3, Maternal (as well as paternal and neonatal) environments are also influenced by the external factors, notably diet and management, which are more influential earlier (heavier arrows) than later in gestation. 4, Events leading to gametogenesis (broken lines) begin in utero and are therefore subject to parental and grandparental influences. 5, Pertubations of the environments affecting gametogenesis, embryogenesis, and synchronous developments can occur at any point along the broken lines and may be manifested immediately or after a delay. Manipulation of gametes and embryos in vitro is an obvious example of such pertubation. Others, however, notably dietary effects, are also important. Adapted from Betteridge.24
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Phase 2: the blastocyst before completion of the yolk sac; days 6–8 By the time the embryo passes into the mare’s uterus through the particularly distinct utero-tubal papilla about 6 days after ovulation,52 it is still within its unexpanded zona pellucida, usually with an overall diameter of about 180 µm and in transition between the morula and blastocyst stages (Figure 225.4). Instantly it becomes readily accessible by simple uterine lavage for both practical embryo transfer53 and research, both of which have taught us much more about embryos from this stage on than is known of earlier stages. As a result, it is known that critical changes, both morphological and physiological, rapidly ensue both inside and outside the differentiated trophoblast cells. Inside, the blastocyst cavity starts to enlarge, becomes better defined, demarcates the inner cell mass more clearly, and becomes lined with endoderm cells. Formation of the endodermal lining proceeds differently in horses than in ruminants and pigs. Rather than progress outwards from hypoblast cells of the inner cell mass (as in ruminants and pigs),30 equine endoderm cells first form separate ‘colonies’ sporadically distributed over the interior of the trophoblast. They then grow from these foci and coalesce into a continuous sheet.54 Outside, the capsule (an acellular mucin-like glycoprotein layer)30 is laid down between the trophoblast and the inner surface of the zona pellucida. The zona pellucida is then shed from the outside of the completed capsule during the first day or so in the uterus (Figure 225.6a). The processes involved in emergence of the encapsulated embryo from the zona pellucida have been considerably clarified recently; Stout et al.55 have observed that puncture of the zona pellucida by micromanipulation in vitro leads to ‘explosive’ shedding of the zona and emergence of the unfolding capsule through the breach. This is reminiscent of the effects of phase transition in biological gels.56 Nevertheless, many aspects of the formation of the capsule still merit attention in relation to our understanding of the embryonic coats encountered in the oviduct, and for the same comparative reasons. The capsule is structurally strong, essential to the continuation of pregnancy, and presumably of value in continuing the protective function of the zona pellucida. The trophoblastic cells certainly produce capsular material but, in vitro, the product is not transformed into the distinct acellular membrane form seen in vivo.57 This supports the notion that the material needs to be modified by cross-linking after secretion, as do mucins, and that conditions necessary for the transformation have not yet been produced in vitro.
(a)
(b) Figure 225.6 Conceptuses recovered from the uterus between days 6.5 and 12, illustrating key developmental events. (a) Loss of the zona pellucida (ZP) by peeling from around the newly formed capsule (Cp) surrounding the trophoblast (Tr) of a blastocyst (diameter 210 µm) with a distinct cavity and inner cell mass (ICM) on day 6.5 (M. Guillomot and K. J. Betteridge, unpublished observations). (b) At day 11.5, diameter 6 mm, the beginning of the phase of rapid expansion, with an intact capsule (Cp) and an embryonic disc showing no sign of neurulation but surrounded by the first signs of mesoderm formation.
It can be assumed that the blastocyst is highly mobile from the time it enters the uterus. There is no direct evidence that this is so until the conceptus first becomes detectable by transrectal ultrasonography, occasionally at day 9 but more reliably at about day 11, well after completion of the yolk sac (see below). However, strong indirect evidence of transuterine migration came from early embryo transfer studies in which flushing the entire uterus resulted in much higher blastocyst recovery rates than when the flush
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Assessing the metabolic activity of embryos at transferable stages could become of direct relevance to embryo transfer in horses if the promising signs (in cattle) that measuring oxygen uptake may be correlated with embryo viability63 are corroborated. For human embryos, two other approaches using ‘metabolism’ to predict viability show promise: amino acid turnover64 and metabolomic profiling.65 Metabolomic profiling involves spectrometric analysis of culture medium, and the profile is apparently related to oxidative metabolism, though exactly how is not known.
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Figure 225.7 Exponential increase in glucose uptake with diameter of equine expanded blastocysts (r = 0.903; P < 0.01). Redrawn from Lane et al.,61 with permission from John Wiley & Sons, Inc.
was confined to the horn adjacent to the ovulating ovary.58, 59 Initial formation of the blastocyst must be assumed to follow the patterns common to all mammals that have been investigated in any detail and to require large amounts of energy to support the pumping of fluid through the trophoblast. Experimentally, glucose metabolism augments 55-fold between days 4.5 and 7.5 (more than doubling when volume increases are taken into account)46 and up to 30-fold during the last day of that period.60 Lane et al.61 demonstrated that equine morulae take up equal amounts of pyruvate and glucose as energy sources with glucose consumption increasing slightly as the embryo cavitates. However, it is not until the blastocyst enters its rapid phase of expansion, considered in the next section, that the consumption of glucose increases exponentially (Figure 225.7). While there are hazards in extrapolating from experiments in vitro (in which substrates available to the embryo are controlled by the scientist) to making assumptions about what occurs in vivo (where the nature of substrates and their delivery is controlled by the mare and remains poorly understood), it is clear that the equine blastocyst is 5–10 times more active than bovine blastocysts of the same size in taking up and metabolizing glucose when it is available.60 It has ¨ et al.62 that progesteronealso been found by Bruck induced endometrial proteins can stimulate glucose metabolism in horse embryos, and they suggest that P19/uterocalin may be the protein responsible. The horse blastocyst differs from those of cattle and other species in metabolizing glucose carbon oxidatively through the Krebs cycle.60 Intriguingly, the predominant sugar within the yolk sac is fructose, not glucose (Figure 225.8).
Phase 3: completion of the yolk sac, pronounced mobility, and rapid expansion of the conceptus; days 8–17 As soon as the conceptus becomes detectable by ultrasonography it can be shown to be migrating throughout the uterine horns and body at a rate that accelerates with age up to about day 15.66 There is good evidence that the mobility of the conceptus is due to its own production of PGs.67–69 To watch, by ultrasonography, the uterine contractions that propel and deform the spherical conceptus is to appreciate the necessity for a strong supportive capsule. Equine embryos remain transferable at the beginning of this phase but success rates have generally been felt to drop markedly after day 8, partly because of the vulnerability of larger embryos to mechanical damage. Experimentally, however, good success rates have recently been obtained with day-10 embryos.70 Coalescence of the endoderm ‘colonies’ on the inside of the trophoblast at day 8 completes the formation of the yolk sac and initiates some important functional changes in the conceptus which result in its rapid growth and remarkable buoyancy. The pattern of increase in diameter of the spherical conceptus originally described by Ginther66 has been confirmed many times: growth is especially rapid between days 11 and 16, at which stage it enters a plateau. Figure 225.6b shows a conceptus at the beginning of this rapid expansion. Figure 225.8 a represents the rapid growth in terms of volume; it shows that the rate of fluid intake increases exponentially, reaching about 3 mL/day by day 16. As discussed elsewhere,30 what is remarkable about this phase of rapid expansion (Figure 225.9) is that the fluid within the yolk sac is markedly hypotonic (Figure 225.8b) and so the osmotic gradients generally assumed to underlie earlier blastocyst expansion in mammals cannot apply to equine yolk sac expansion. The hypo-osmolality of yolk sac fluid makes the conceptus buoyant in the
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Days after ovulation Figure 225.8 Changes in yolk sac fluid of the equine conceptus near the time of fixation (day 16 is indicated by the broken vertical line). (a) Daily volumes of 29 conceptuses calculated from diameters measured by ultrasonography (diamond symbols) and measured by aspiration at collection (round circles). (b) Osmolality (mOsm). (c) Fructose concentration (mM). (d) Protein concentration (µg/mL). Data from Betteridge.30
phosphate-buffered saline (280 mOsm) used to flush them from the uterus; the buoyancy begins at a diameter of 4–5 mm (days 10–11) and is maintained until sometime between days 18 and 20, at about the time the spherical shape of the conceptus is lost but while the capsule is still present (Figure 225.2; K. J. Betteridge and R.O. Waelchli, unpublished observations). Enders et al.54 have suggested that ‘third compartments’ (intercellular spaces between the trophoblast and the endoderm; Figure 225.10) are involved in fluid uptake by the conceptus. Our analyses of ion concentrations in these compartments and in the interior of the yolk sac by cryoscanning electron microscopy and energy dispersive X-ray microanalysis (SEM-EDX)71 partially support that suggestion. In conceptuses ≥ 6 mm in diameter, the concentrations of both sodium and potassium in the subtrophectodermal compartments were higher than those previously determined in uterine fluid, supporting the concept of osmotic intake of fluid from the uterine environment as far as the compartments. However, electrolyte concentrations in the compartments consistently exceeded those found in the yolk sac, making it likely that ‘uphill’ water transport, rather than a purely osmotic uptake, is involved in yolk sac fluid accumulation, probably mediated through the sodium pump. Budik et al.72 have shown that aquaporins are expressed in the yolk sac wall and are therefore likely to be important to fluid transport across the wall and to the expansion process. It
has also been suggested that capsule formation could contribute to conceptus expansion, and thereby to fluid intake, by mechanical means. The osmotic behavior of the equine conceptus from about day 8 has some very practical implications for cryopreservation. While morulae and early blastocysts collected immediately after entering the uterus (day 6) are readily cryopreserved by techniques used for ruminant embryos, success is rare with equine embryos after day 6. This is likely to be because the osmotic response of equine blastocysts to hypertonic cryoprotective agents changes markedly when the blastocysts reach a diameter of about 500 µm. Hochi et al.73, 74 have shown that smaller equine blastocysts respond like the blastocysts of other mammals by shrinking rapidly (within < 1 minute) to about 40–60% of their initial volume as water is withdrawn, then re-expanding to some 60–85% of their initial volume within 10 minutes as the cryoprotective agent permeates the embryo. In contrast, the largest blastocysts (498–980 µm in diameter) exposed to glycerol or ethylene glycol shrank only slowly and failed to re-expand within 20 minutes. This change in behavior, and the intolerance of cryopreservation, has been widely attributed to the appearance of the capsule but it should also be noted that it occurs at about the time and size that the endoderm is completed. Since the capsule is permeable to even large neutral molecules it is probably completion of the bilaminar omphalopleure that is
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Figure 225.10 The planed surface of a frozen horse conceptus collected at day 10 ± 1, 10 mm in diameter, and processed for cryoscanning electron microscopy and energy dispersive microanalysis, showing the prominent ‘third compartments’ (subtrophectodermal compartments; STC) between the trophectoderm and endoderm of the yolk-sac wall. From Crews et al.,71 with permission from CSIRO Publishing. (b)
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Figure 225.9 Conceptuses (A–C) and their embryonic discs (a–c) between days 14 and 18. The leading edge of the spreading mesoderm (i.e., the border between bi- and trilaminar yolk sac wall, or omphalopleure) at each stage is indicated by the vertical arrows. The conceptus at day 14.5, photographed from the side, is reflected in the surface of the collecting fluid in which it floats. The capsule is closely applied to the yolk sac in A and B but has become flaccid and free hanging in C. The discs show the sequence of developmental events leading to neurulation and somite formation, shown diagrammatically in Figure 225–14: (a) shows a dark crescent of cells at the posterior end of the disc (bottom of the photograph) which will form the primitive streak; (b) shows the neural plate (top), the neural folds, and the first four pairs of somites; (c) is photographed through the capsule (Cp) to show 11 pairs of somites and further closure of the neural tube with the head end to the right.
associated with the increased resistance to water withdrawal in a hypertonic environment. Conceptuses recovered on days 13 or 14 (12 mm in diameter) behave similarly to the largest ones described by Hochi et al.;73, 74 they largely retain their shape, turgidity, and hypo-osmolality when held for 90–110
minutes in NaCl solutions of 600 or 1500 mOsm.75 This resistance to water withdrawal is unlikely to be due to the capsule which, at least at later stages, does not prevent diffusion of ions; fluid collected from within an intact capsule after collapse of the yolk sac (due to trauma at collection) approaches isotonicity with its surrounding collection fluid within 1–2 hours (A. Butler, K. J. Betteridge and R. O. Waelchli, unpublished observations in six of seven such conceptuses; Figure 225.11). It is not surprising that the rapid expansion of the conceptus demands the ability to exponentially increase glucose uptake in vitro, as illustrated in Figure 225.7. The rapid growth of the conceptus between days 11 and 16 (Figure 225.9) is during the time that the conceptus exchanges essential signals with the mare by moving throughout the uterus rather than by elongating like its counterparts in ruminants and pigs.30 Also in contrast to ruminant embryos, the equine trophoblast does not produce interferon-τ as an effector of ‘recognition of pregnancy’. Interestingly, however, mRNA for interferon-δ is expressed on both sides of the conceptus–endometrial interface during this time.76 The functional significance of this remains to be established. Capsule production continues during this period of development and, since the first discovery of proteins bound to the capsule by Stewart et al.,77 much attention has been paid to various proteins in and around the early conceptus, particularly in uterine fluid, the capsule (Figure 225.12), and yolk sac fluid. Selective binding of some proteins to the capsule supports the concept that the capsule is like a ‘mailbox’
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Figure 225.12 Diagrammatic representation of the presence and prominence of proteins bound to the capsule before and after fixation (vertical broken line at day 16) of the equine conceptus. Adapted from Quinn et al.78 and Hayes et al.79
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(b) Figure 225.11 Twin conceptuses collected at 16.5 days after the first of two ovulations and measuring 27 mm and 21 mm in diameter by ultrasonography. At the time of collection (a), both floated in the phosphate-buffered saline (PBS) lavage; one yolk sac (A) was intact but the other (B) had ruptured inside an intact capsule (arrowhead). By 50 minutes after collection (b), the intact conceptus remained buoyant but the ruptured one had sunk in PBS. Fluid recovered from the intact conceptus 1.5 hours after collection from the mare was hypotonic (114 mOsm) whereas the fluid from conceptus B 1 hour after that had largely equilibrated with the PBS (273 mOsm). Note that the spherical form of the capsule is maintained despite the collapsed yolk sac wall and different osmotic gradients between the inside and outside at the times of the two photographs.
for delivery of various substances required by the embryo and endometrium in the conduct of their dialogue. For example, maternal p19/uterocalin binds to the capsule and is implicated mainly in delivery of lipids to the equine conceptus. It has also been shown that intact β 2 microglobulin (β 2 M) is a major protein associated with the embryonic capsule before fixation and is evidently not part of a complex with major histocompatibility complex (MHC) class I heavy α chain bands because these were undetectable in the capsule and uterine lavage.78, 79
Another probable component of systems for exchanging materials between the conceptus and the mare is found in the cellular yolk sac wall; before fixation these tissues express unusually large amounts of the GM2-activator protein (GM2AP), implicated in transport and catabolism of glycolipids.80 The yolk sac wall also produces insulin-like growth-factorbinding protein 3 that binds insulin-like growth factor (IGF-1; which is important for embryonic development), synthesizes and metabolizes steroids, and produces both PGF2α and PGE2 .30 The steroid metabolism differs between the bilaminar and trilaminar portions of the yolk sac wall, emphasizing the importance of defining functional differences between components of the conceptus during a period when rapid changes in proportions of these is occurring (Figure 225.9). Similar considerations apply to endodermal and trophectodermal layers of the wall. The equine endoderm deserves much more attention than it has received, given its multiple synthetic functions in other species, including humans81 and also some striking similarities between the environments in the equine yolk sac and the human exocoelomic cavity.82 The yolk sac fluid of the rapidly expanding conceptus contains relatively low concentrations of proteins (Figure 225.8d) but notable among them are p19/uterocalin and uteroglobin. Steroid production and metabolism by the early equine conceptus is very significant for the establishment and maintenance of pregnancy; estrogens, both free and as sulfoconjugates, accumulate in the yolk sac fluid as the pregnancy progresses through the attachment period (Figure 225.13).83 Furthermore, recent work has shown that quantities of the anabolic steroid norandrostenedione (NorA) in yolk sac fluid, mostly as a sulfate, increase to even higher levels;84 by day 16 the amount of this androgen exceeds that of estrone sulfate about threefold. The recent demonstration of receptors for estrogen and progesterone in the conceptus85 underlines the fact that more needs to be known about the roles that steroids play in the establishment and maintenance of pregnancy in horses.
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Figure 225.13 Total content of estrone sulfate in the equine conceptus before and after fixation (day 16 is indicated by the vertical broken line). Note the logarithmic scale. Data from Raeside et al.83
Estrogens influence vascularity and permeability and are presumed to be responsible for the localized endometrial edema around the conceptus once it becomes immobilized at about day 16. The edema can be recognized ultrasonographically66, 86 and quantified histologically a few days before localized endometrial hyperemia becomes obvious in the same location.87 Paradoxically, when estradiol is released from silastic spheres inserted into the uterus to emulate a conceptus, it enhances, rather than inhibits, endometrial release of luteolytic PGF2α in response to an oxytocin stimulus, so this particular estrogen seems unlikely to be involved directly in the maintenance of pregnancy.88 This is not to say that the conceptus plays no role in maintaining pregnancy, of course; it certainly inhibits PG release in response to oxytocin by days 11–13 of pregnancy.89 However, the search for specific ‘pregnancy recognition’ signals (analogous to interferon-τ in ruminants and estrogens in pigs, for example) has so far been fruitless,90 although there is some evidence for the involvement of a small molecule (3.5–6 kDa) produced by the conceptus. It has been suggested that the endometrium might metabolize progesterone to other more polar neutral steroids with roles to play in the maintenance of pregnancy.88, 91 Yolk sac fluid during this time contains very low concentrations of glucose but remarkably high concentrations of fructose, which increase with size (Figure 225.8c). Pairs of somites begin to appear in the embryo proper between days 15 and 16 in the horse (about 5 days earlier than in cattle)92 and accumulate rapidly, although with considerable variation from embryo to embryo (Figure 225.14). The mobility of the conceptus in the uterus is brought to an end at about days 16–17 through the process of ‘fixation’.66, 93
The prevailing view of the cause of ‘fixation’ is that it is predominantly mechanical; parallel increases in uterine tone and conceptus diameter coincide to prevent the conceptus from moving through the narrower uterine lumen at the flexure found at the base of each uterine horn.66 However, many changes in the conceptus seem to coincide with its immobilization at the time of fixation and may also be involved in the process; they include the physical properties of the capsule and the patterns of proteins that bind to it and pass through it into the yolk sac. Structurally, the capsule becomes flaccid (Figure 225.9c). This reflects the loss of sialic acid from what has recently been defined as a sialylated core type 1 O-linked glycan, Neu5 Ac-(2→3)-Gal-(1→3)GalNAc-(1→Ser/Thr).94 Measured dry weights of the capsule continue to rise until about day 18, but it is not known whether this is due to continued production or accumulation of bound proteins. Among the proteins bound to the capsule, p19/uteroglobin remains prominent but the β 2 M undergoes progressive degradation and a 17-kDa protein, secretory phospholipase A2, makes a transitory appearance (Figure 225.12). The osmolality of the yolk sac fluid begins to rise at about the time of fixation, as do its concentrations of proteins and fructose (Figure 225.8) and of arginine vasopressin and oxytocin.30 The rise in yolk sac fluid estrogen concentrations continues for some days and then reaches a plateau (Figure 225.13). In contrast to prefixation yolk sac fluid, uteroglobin and p19/uterocalin are no longer found to enter the sac.78 Immobilization of the conceptus at the time of fixation also coincides with its proper orientation (embryo proper ‘down’, antimesometrially). Silva and Ginther95 were able to detect the first signs of this process by color-Doppler ultrasonography. At the interface between the trophectoderm and the endometrium the capsule is likely to play a governing role in the fixation and orientation process and so mechanisms involved in anchoring the capsule are under investigation. The yolk sac can move to a limited extent within its capsule, at least in vitro after conceptus recovery, but how any such movement may relate to orientation is not known. The changes that occur in the embryo’s environment at the time of fixation and orientation are relatively acute, making them excellent end points to use in investigation of embryonic loss. They may be especially appropriate because of the concentration of losses between days 15 and 35 already referred to.29
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Figure 225.14 The rate (a) and manner (b) of somite formation in the equine embryonic disc. (a) Counts of somite pairs in individual conceptuses of ages known to within ± 0.5 days. (b) The embryonic disc seen from above displaying the development of the neuroectoderm and formation of somites in relation to the relative withdrawal of the primitive streak (courtesy of Drs M. Vejlsted and P. Maddox-Hyttel). (a) represents a stage very slightly after the disc shown in Figure 225–9a at 14.5 days; (c) represents the stage shown in Figure 225–9b at 16.5 days. 1, Primitive (Hensen’s) node; 2, primitive streak; 3, cut edge of chorion; 4, neural plate; 5, neural fold; 6, neural groove; 7, somite.
Experimental design for such investigations is difficult; which 16 of the first 100 day-15 conceptuses collected were destined to fail? This being an unanswerable question, an alternative approach is to examine conceptuses at given stages of pregnancies that have been ‘compromised’ (by luteolysis, for example) and compare them with control counterparts. Extraordinarily, the conceptus grows and develops quite normally for 3–4 days after removal of maternal progesterone; indeed, development after ovariectomy at day 12 can, on occasion, even proceed to the heartbeat stage at day 26.96 Luteolysis, however, does
affect desialylation of the capsule and the patterns of proteins that bind to it.30 After fixation the amniotic folds rise around the embryo proper, then meet and fuse to complete the amnion. This process is completed while the capsule is still present, which may well signify the importance of the capsule’s protective role because, until the amnion is completed, the somitic embryo is structurally vulnerable. Thus, when a conceptus is accidentally ruptured by the trauma of transcervical collection it most often splits neatly along the neural groove.
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Phase 5: loss of the capsule How and when the capsule is lost from around the trophectoderm has yet to be established precisely. It can be recovered intact at day 20 and in fragments for several days thereafter. It also appears to be complete in sections of uteri fixed by perfusion at day 2097 so day 21 is a reasonable estimate of its time of loss. Early work on possible enzymatic mechanisms of capsule removal98 should be followed up because disturbance of the normal removal of the capsule could be another factor affecting the survival or demise of the conceptus. It is also possible that continued yolk sac expansion in the absence of further capsule production causes mechanical rupture of the capsule. The conceptus loses its buoyancy in physiological saline between days 18 and 20, for reasons that remain entirely speculative.30
Phase 6: establishment of cell-to-cell contact with the endometrium, days 21–28 Beginning at about the time of loss of the capsule, the embryo proper comes into its own, with a completed amnion, emerging allantois, rapidly developing circulatory system, and recognizable body form (Figure 225.15). It was this stage of development that inspired the classic studies of Ewart.99 Building on his observations (which had received scant attention for 90 years), we have described structures that traverse the bilaminar omphalopleure even in the presence of an intact capsule and seem highly likely to be of functional significance (Figures 225.15 and 225.16).30 They deserve attention. By the end of the first month of gestation the degree of organogenesis in the equine embryo proper is striking (Figures 225.15c and 225.17) but no different than that seen in embryos of other domestic species. What makes the horse uniquely valuable, however, is the ease with which such large embryos, complete with their immediate environment contained within the extraembryonic membranes, can be recovered from docile donors repeatedly and without harm to either dam or conceptus. Given current concerns about the environment in general, and the effects of endocrine disruptors on sexual differentiation in particular,100 such ready access to the early stages of brain development and to yolk sac fluid reflecting its immediate steroidal environment suggests to me that the horse offers an unusual and useful model for study topics such as early sexual dimorphism. But horses did not evolve in order to provide research models; it is the manner in which this first month of pregnancy equips the embryo to navigate
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Figure 225.15 Conceptuses (a–c) and associated embryos proper between days 19 and 28. (a) Photographed collapsed in fixative, the embryo proper is seen at 6 o’clock and an acellular floating body (arrow; commonly seen) is within the yolk sac. The bilaminar yolk sac wall (top half of the conceptus) is decorated with trophoblastic discs (see Figure 225–16). Well-formed blood vessels extend to the sinus terminalis at the border of the bi- and trilaminar yolk sac wall. (b) The vascularized allantois (arrow) is seen emerging from the embryo proper; the sinus terminalis is more prominent; the bilaminar yolk sac wall is diminished relative to the trilaminar. The bracket indicates an area of trophoblastic discs (see Figure 225–16). (c) The allantois has become much more prominent and, by ultrasonography, would be seen to be displacing the embryo proper from the bottom of the embryonic vesicle towards the middle. (See color plate 114).
its ensuing 10 months in the uterus and then its entire postnatal life that is the marvel – and a puzzle fit for anyone with a yen for science and an appreciation of magnificent animals.
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(c) Figure 225.16 Trophoblastic discs of the type shown in Figures 225–15a,b as first illustrated by Ewart99 in 1915 (a), as next reported in 2002 in association with yolk sac tubercles and cords (b), and as seen by scanning electron microscopy (c). YS, yolk sac; Tr, trophoblast. From Betteridge et al.101
Concluding remarks There is perverse satisfaction to be taken from knowing that much that has been written in this chapter will be outdated by the time it is read; the potential for advancing our knowledge of reproduction in horses has increased immeasurably with the sequencing of the equine genome, and molecular biology, genomics, and proteomics have revolutionized the ways in which age-old questions can now be answered. Fortunately, there are many reasons for studying equine reproduction and improving its efficiency for its own sake; the equine industry is economically very significant in many countries, and poor reproductive performance is correspondingly costly. Fortunately, too, there is still room for good
observational studies upon which to base modern experimental approaches to answering the myriad questions to which observations in horses give rise. I hope that the observations recorded in this chapter will encourage the posing of yet more questions and stimulate some to take up the challenge of answering them.
Acknowledgements I am most grateful to all my colleagues (co-authors in cited papers), and particularly to Drs R.O. Waelchli, M.A. Hayes, J.I. Raeside and C. Tayade for their help with the work that has formed the backbone of this chapter. I am grateful to Dr. D. Rieger for helpful
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Figure 225.17 Longitudinal section of a 25-mm embryo (about day 39) from the collection of the late Dr. Arthur W. Marrable (courtesy of Dr. Peter Flood) illustrating the advanced stage of organogenesis reached at this stage. Remarkably, at least until day 37 (Antczak et al.102 ), embryos can be obtained intact by transcervical lavage without harm to the mare and are thus available for detailed study. This specimen lies a little beyond the first month of pregnancy but it has been included as a tribute to Dr. Marrable, who was my first mentor in embryology and among the early contributors to the renaissance in the study of equine development. 1, Tail; 2, genital tubercle; 3, descending colon; 4, loops of intestine; 5, spinal cord; 6, vertebral bodies; 7, liver; 8, stomach; 9, pancreas; 10, lung; 11, trachea; 12, esophagus; 13, ventricle; 14, aorta; 15, larynx; 16, tongue; 17, nasal cavity; 18, lateral ventricle; 19, fourth ventricle; 20, medulla oblongata.
discussions of embryo metabolism. Financial support for our own studies, from NSERC, OMAFRA, Equine Guelph, the E.P. Taylor Research Foundation, and the Grayson-Jockey Club Research Foundation Inc., is much appreciated.
References 1. Fleming TP, Kwong WY, Porter R, Ursell E, Fesenko I, Wilkins A, Miller DJ, Watkins AJ, Eckert JJ. The embryo and its future. Biol Reprod 2004;71:1046–54. 2. Sinclair KD, Lea RG, Rees WD, Young LE. The developmental origins of health and disease: current theories and epigenetic mechanisms. Soc Reprod Fertil Suppl 2007;64:425–43. 3. Gluckman PD, Hanson MA, Beedle AS. Non-genomic transgenerational inheritance of disease risk. Bioessays 2007;29:145–54. 4. Leese HJ, Hugentobler SA, Gray SM, Morris DG, Sturmey RG, Whitear S-L, Sreenan JM. Female reproductive tract fluids: composition, mechanism of formation and potential role in the developmental origins of health and disease. Reprod Fertil Dev 2008;20:1–8. 5. Longo LD. Classic pages in the American Journal of Obstetrics and Gynecology. Am J Obstet Gynecol 2000;182:473–4. 6. Holm LW. Prolonged pregnancy. Adv Vet Sci Comp Med 1967;11:159–205. 7. Holm LW, Short RV. Progesterone in the peripheral blood of Guernsey and Friesian cows during prolonged gestation. J Reprod Fertil 1962;4:137–41. 8. Liggins GC, Kennedy PC, Holm LW. Failure of initiation of parturition after electrocoagulation of the pituitary of the fetal lamb. Am J Obstet Gynecol 1967;98:1080–6.
9. Zondek B. Hormonale Schwangerschaftsreaction aus dem Harn beim Menschen und Tier. Klin Wochenschr 1930;9:2285–9. 10. Raeside JI, Liptrap RM, Milne FJ. Relationship of fetal gonads to urinary estrogen excretion by the pregnant mare. Am J Vet Res 1973;34:843–5. 11. Cox JE. Oestrone and equilin in the plasma of the pregnant mare. J Reprod Fertil Suppl 1975;23:463–8. 12. Raeside JI, Liptrap RM. Patterns of urinary oestrogen excretion in individual pregnant mares. J Reprod Fertil Suppl 1975;23:469–75. 13. Allen WR. Fetomaternal interactions and influences during equine pregnancy. Reproduction 2001;121:513–27. 14. Allen WR. Development and functions of the equine placenta. In: Powell DG, Furry D, Hale G (eds) Proceedings of a Workshop on the Equine Placenta, 2003. University of Kentucky, Lexington, 2004, pp. 20–33. 15. Allen WR, Stewart F. Equine placentation. Reprod Fertil Dev 2001;13:623–34. 16. Cole HH, Hart GH, Lyons WR, Catchpole HR. The development and hormonal content of fetal horse gonads. Anat Rec 1933;56:275–93. 17. Betteridge KJ. Comparative aspects of conceptus growth: a historical perspective. Reproduction 2001;122:11– 19. 18. Hamburger C. Historical introduction. In: Wolstenholme GEW, Knight J (eds) Gonadotropins: Physicochemical and Immunological Properties. London: J. and A. Churchill, 1965; pp. 2–10. 19. Greep RO. A vista of research on the mammalian gonadotropins. Biol Reprod 1973;8:2–10. 20. Bielanski W, Ewy Z, Pigoniowa H. [Preliminary investigations on differences in endocrine secretion in mares which are in foal to the stallion and to the ass]. Folia Biol 1955;3:19–30.
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21. Allen WR. Immunological aspects of the endometrial cup reaction and the effect of xenogeneic pregnancy in horses and donkeys. J Reprod Fertil Suppl 1982;31:57–94. 22. Wolstenholme GEW, O’Connor M (eds). Foetal Autonomy. London: J. and A. Churchill, 1969. 23. Sharp DC, McDowell KJ, Weithenauer J, Thatcher WW. The continuum of events leading to maternal recognition of pregnancy in mares. J Reprod Fertil Suppl 1989;37:101–7. 24. Betteridge KJ. Comparative aspects of equine embryonic development. Anim Reprod Sci 2000;60–61:691– 702. 25. Van Niekerk CH, Gernecke WH. Persistence and parthenogenic cleavage of tubal ova in the mare. Onderstepoort J Vet Res 1966;31:195–232. 26. Lee KF, Yao YQ, Kwok KL, Xu JS, Yeung WS. Early developing embryos affect the gene expression patterns in the mouse oviduct. Biochem Biophys Res Commun 2002;292:564–70. 27. McNatty KP, Moore LG, Hudson NL, Quirke LD, Lawrence SB, Reader K, Hanrahan JP, Smith P, Groome NP, Laitinen M, Ritvos O, Juengel JL. The oocyte and its role in regulating ovulation rate: a new paradigm in reproductive biology. Reproduction 2004;128:379–86. 28. Grant VJ, Irwin RJ, Standley NT, Shelling AN, Chamley LW. Sex of bovine embryos may be related to mothers’ preovulatory follicular testosterone. Biol Reprod 2008;78:812–15. 29. Morris LH, Allen WR. Reproductive efficiency of intensively managed Thoroughbred mares in Newmarket. Equine Vet J 2002;34:51–60. 30. Betteridge KJ. Equine embryology: an inventory of unanswered questions. Theriogenology 2007;68Suppl 1: S9–21. 31. Betteridge KJ, Mitchell D. A surgical technique applied to the study of tubal eggs in the mare. J Reprod Fertil Suppl 1975;23:519–24. 32. Galli C, Colleoni S, Duchi R, Lagutina I, Lazzari G. Developmental competence of equine oocytes and embryos obtained by in vitro procedures ranging from in vitro maturation and ICSI to embryo culture, cryopreservation and somatic cell nuclear transfer. Anim Reprod Sci 2007;98:39–55. 33. Ball BA, Little TV, Weber JA, Woods GL. Survival of Day4 embryos from young, normal mares and aged, subfertile mares after transfer to normal recipient mares. J Reprod Fertil 1989;85:187–94. 34. B´ezard J, Magistrini M, Duchamp G, Palmer E. Chronology of equine fertilization and embryonic development in vivo and in vitro. Equine Vet J Suppl 1989;8:105–10. 35. Grøndahl C, Grøndahl Nielsen C, Eriksen T, Greve T, Hyttel P. In vivo fertilization and initial embryogenesis in the mare. Equine Vet J Suppl 1993;15:79–83. 36. Heng BC, Cao T, Stojkovic M, Vajta G. Mammalian oocyte polarity can be exploited for the automation of somatic cell nuclear transfer—in the development of a ‘cloning biochip’. Med Hypotheses 2006;67:420–1. 37. Goudet G, Mugnier S, Callebaut I, Monget P. Phylogenetic analysis and identification of pseudogenes reveal a progressive loss of zona pellucida genes during evolution of vertebrates. Biol Reprod 2008;78:812–15. 38. Menkhorst E, Selwood L. Vertebrate extracellular preovulatory and postovulatory egg coats. Biol Reprod 2008;79: 790–7. 39. Malette B, Paquette Y, Merlen Y, Bleau G. Oviductins possess chitinase- and mucin-like domains: a lead in the search for the biological function of these oviduct-specific
40.
41.
42.
43.
44.
45.
46.
47.
48.
49.
50.
51.
52.
53. 54.
55.
ZP-associating glycoproteins. Mol Reprod Dev 1995;41: 384–97. Kan FW, Esperanzate PW. Surface mapping of binding of oviductin to the plasma membrane of golden hamster spermatozoa during in vitro capacitation and acrosome reaction. Mol Reprod Dev 2006;73:756–66. M´en´ezo Y, Elder K, Viville S. Soluble HLA-G release by the human embryo: an interesting artefact? Reprod Biomed Online 2006;13:763–4. Shao R, Weijdeg˚ard B, Fernandez-Rodriguez J, Egecioglu E, Zhu C, Andersson N, Thurin-Kjellberg A, Bergh C, Billig H. Ciliated epithelial-specific and regional-specific expression and regulation of the estrogen receptor-beta2 in the fallopian tubes of immature rats: a possible mechanism for estrogen-mediated transport process in vivo. Am J Physiol Endocrinol Metab 2007;293:E147–58. Croxatto HB. Physiology of gamete and embryo transport through the fallopian tube. Reprod Biomed Online 2002;4:160–9. ´ R´ıos M, Hermoso M, S´anchez TM, Croxatto HB, Villalon MJ. Effect of oestradiol and progesterone on the instant and directional velocity of microsphere movements in the rat oviduct: gap junctions mediate the kinetic effect of oestradiol. Reprod Fertil Dev 2007;19:634–40. Paulo E, Tischner M. Activity of delta(5)3betahydroxysteroid dehydrogenase and steroid hormones content in early preimplantation horse embryos. Folia Histochem Cytobiol 1985;23:81–4. ¨ I, Hyland JH. Measurements of glucose metabolism Bruck in single equine embryos during early development. J Reprod Fertil Suppl 1991;44:419–25. Leese HJ, Sturmey RG, Baumann CG, McEvoy T. Embryo viability and metabolism: obeying the quiet rules. Hum Reprod 2007;22:3047–50. Pantaleon M, Scott J, Kaye PL. Nutrient sensing by the early mouse embryo: hexosamine biosynthesis and glucose signaling during preimplantation development. Biol Reprod 2008;78:595–600. Duranthon V, Watson AJ, Lonergan P. Preimplantation embryo programming: transcription, epigenetics, and culture environment. Reproduction 2008;135:141–50. Grøndahl C, Hyttel P. Nucleologenesis and ribonucleic acid synthesis in preimplantation equine embryos. Biol Reprod 1996;55:769–74. Giussani DA, Forhead AJ, Gardner DS, Fletcher AJ, Allen WR, Fowden AL. Postnatal cardiovascular function after manipulation of fetal growth by embryo transfer in horses. J Physiol 2003;547 (Part 1): 67–76. Battut I, Grandchamp des Raux A, Nicaise JL, Fieni F, Tainturier D, Bruyas JF. When do equine embryos enter the uterine cavity? An attempt to answer. In: Katila T, Wade JF (eds) Proceedings of the 5th International Symposium on Equine Embryo Transfer. Havemeyer Foundation Monograph Series No. 3. Newmarket: R&W Publications, 2001; pp. 66–8. Stout TAE. Equine embryo transfer: review of developing potential. Equine Vet J 2006;38:467–78. Enders AC, Schlafke S, Lantz KC, Liu KM. Endoderm cells of the equine yolk sac from Day 7 until formation of the definitive yolk sac placenta. Equine Vet J Suppl 1993;15:3–9. Stout TAE, Meadows S, Allen WR. Stage-specific formation of the equine blastocyst capsule is instrumental to hatching and to embryonic survival in vivo. Anim Reprod Sci 2005;87:269–81.
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Embryo Morphology, Growth and Development 56. Pollack GH. Cells, Gels, and the Engines of Life: a New Unifying Approach to Cell Function. Seattle: Ebner and Sons, 2001, pp. 113–29. 57. Tremoleda JL, Stout TA, Lagutina I, Lazzari G, Bevers MM, Colenbrander B, Galli C. Effects of in vitro production on horse embryo morphology, cytoskeletal characteristics, and blastocyst capsule formation. Biol Reprod 2003;69:1895–906. 58. Oguri N, Tsutsumi Y. Non-surgical egg transfer in mares. J Reprod Fertil 1974;41:313–20. 59. Allen WR, Rowson LEA. Surgical and non-surgical egg transfer in horses. J Reprod Fertil Suppl 1975;23:525–30. 60. Rieger D. Embryo metabolism and implications for culture requirements. In: Morris LH-A, Foster L, Wade JF (eds) Proceedings of a Workshop Entitled ‘From Epididymis to Embryo’. Havemeyer Foundation Monograph Series No. 6. Newmarket: R&W Publications, 2001; pp. 59–61. 61. Lane M, O’Donovan MK, Squires EL, Seidel Jr GE, Gardner DK. Assessment of metabolism of equine morulae and blastocysts. Mol Reprod Dev 2001;59:33–7. ¨ 62. Bruck I, Anderson GA, Hyland JH. The influence of progesterone-induced proteins on glucose metabolism in early equine embryos. Theriogenology 1997;47:441–56. 63. Lopes AS, Madsen SE, Ramsing NB, Løvendahl P, Greve T, Callesen H. Investigation of respiration of individual bovine embryos produced in vivo and in vitro and correlation with viability following transfer. Hum Reprod 2007;22:558–66. 64. Brison DR, Houghton FD, Falconer D, Roberts SA, Hawkhead J, Humpherson PG, Lieberman BA, Leese HJ. Identification of viable embryos in IVF by noninvasive measurement of amino acid turnover. Hum Reprod 2004;19:2319–24. 65. Scott R, Seli E, Miller K, Sakkas D, Scott K, Burns DH. Noninvasive metabolomic profiling of human embryo culture media using Raman spectroscopy predicts embryonic reproductive potential: a prospective blinded pilot study. Fertil Steril 2008;90:77–83. 66. Ginther OJ. Equine pregnancy: physical interactions between the uterus and conceptus. In: Proccedings of the 44th Convention of the American Association of Equine Practitioners 1998, pp. 73–104. 67. Stout TAE, Allen WR. Role of prostaglandins in intrauterine migration of the equine conceptus. Reproduction 2001;121:771–5. 68. Stout TAE, Allen WR. Prostaglandin E2 and F2α production by equine conceptuses and concentrations in conceptus fluids and uterine flushings recovered from early pregnant and dioestrus mares. Reproduction 2002;123:261–8. 69. Aurich C, Budik S. Expression of enzymes involved in the synthesis of prostaglandins in early equine embryos. In: Stout TAE, Wade JF (eds) Proceedings of a Workshop on Maternal Recognition of Pregnancy in the Mare, III. Havemeyer Foundation Monograph Series No. 16. Newmarket: R&W Publications, 2004; pp. 31–2. 70. Wilsher S, Clutton-Brock A, Allen WR. Successful transfer of day 10 horse embryos: influence of donorrecipient asynchrony on embryo development. Reproduction 2010;193:575–85. 71. Crews LJ, Waelchli RO, Huang CX, Canny MJ, McCully ME, Betteridge KJ. Electrolyte distribution and yolk sac morphology in frozen hydrated equine conceptuses during the second week of pregnancy. Reprod Fertil Dev 2007;19:804–14.
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72. Budik S, Walter I, Tschulenk W, Helmreich M, Deichsel K, Pittner F, Aurich C. Significance of aquaporins and sodium potassium ATPase subunits for expansion of the early equine conceptus. Reproduction 2008;135:497–508. 73. Hochi S, Ogasawara M, Braun J, Oguri N. Influence of relative embryonic volumes during glycerol equilibration on the survival of frozen-thawed equine blastocysts. J Reprod Dev 1994;40:243–9. 74. Hochi S, Fujimoto T, Oguri N. Large equine blastocysts are damaged by vitrification procedures. Reprod Fertil Dev 1995;7:113–17. 75. Waelchli RO, Betteridge KJ. Osmolality of equine blastocyst fluid from day 11 to day 25 of pregnancy. Reprod Fertil Dev 1996;8:981–8. 76. Tayade C, Cnossen S, Wessels J, Linton N, Quinn BA, Waelchli R, Croy BA, Hayes MA, Betteridge KJ. IFN-δ, a type I interferon, is expressed by both the conceptus and endometrium during early equine pregnancy. Biol Reprod Special Issue 2008;Abstract 96. 77. Stewart F, Charleston B, Crossett B, Barker PJ, Allen WR. A novel uterine protein that associates with the embryonic capsule in equids. J Reprod Fertil 1995;105:65–70. 78. Quinn BA, Hayes MA, Waelchli RO, Kennedy MW, Betteridge KJ. Changes in major proteins in the embryonic capsule during immobilization (fixation) of the conceptus in the third week of pregnancy in the mare. Reproduction 2007;134:161–70. 79. Hayes MA, Quinn BA, Kierstead ND, Katavolos P, Waelchli RO, Betteridge KJ. Proteins associated with the early intrauterine equine conceptus. Reprod Dom Anim 2008;43Suppl. 2: 1–6. 80. Quinn B, Caswell DE, Lillie BN, Waelchli RO, Betteridge KJ, Hayes MA. The GM2-activator protein is a major protein expressed by the encapsulated equine trophoblast. Anim Reprod Sci 2006;94:391–4. 81. King BF, Enders AC. Comparative development of the mammalian yolk sac. In: Nogales FF (ed.) The Human Yolk Sac and Yolk Sac Tumors. Berlin: Springer-Verlag, 1993; pp. 1–32. 82. Jauniaux E, Poston L, Burton GJ. Placental-related diseases of pregnancy: Involvement of oxidative stress and implications in human evolution. Hum Reprod Update 2006;12:747–55. 83. Raeside JI, Christie HL, Renaud RL, Waelchli RO, Betteridge KJ. Estrogen metabolism in the equine conceptus and endometrium during early pregnancy in relation to estrogen concentrations in yolk-sac fluid. Biol Reprod 2004;71:1120–7. 84. Raeside JI, Christie HL. The presence of 19norandrostenedione and its sulphate form in yolk-sac fluid of the early equine conceptus. J Steroid Biochem Mol Biol 2007;108:149–54. 85. Rambags BPB, van Tol HTA, van den Eng MM, Colenbrander B, Stout TAE. Expression of progesterone and oestrogen receptors by early intrauterine equine conceptuses. Theriogenology 2008;69:366–75. 86. Griffin PG, Ginther OJ. Effects of the embryo on uterine morphology and function in mares. Anim Reprod Sci 1993;31:311–29. 87. Lefranc A-C, Allen WR. Endometrial gland surface density and hyperaemia of the endometrium during early pregnancy in the mare. Equine Vet J 2007;39:511–15. 88. Goff AK, Sirois J, Pontbriand D. Effect of oestradiol on oxytocin-stimulated prostaglandin F2α release in mares. J Reprod Fertil 1993;98:107–12.
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89. Goff AK, Pontbriand D, Sirois J. Oxytocin stimulation of plasma 15-keto-13,14-dihydroprostaglandin F-2α during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1987;35:253–60. 90. Stout TAE, Rambags BPB, van Tol HTA, Colenbrander B. Low molecular weight proteins secreted by the early equine conceptus. In: Stout TAE, Wade JF (eds) Proceedings of a Workshop on Maternal Recognition of Pregnancy in the Mare, III. Havemeyer Foundation Monograph Series No. 16. Newmarket: R&W Publications, 2004; pp. 50–2. 91. Raeside JI, Christie HL, Renaud RL, Waelchli RO, Betteridge KJ. Metabolism of neutral steroids by the equine conceptus in early pregnancy. In: Stout TAE, Wade JF (eds) Proceedings of a Workshop on Maternal Recognition of Pregnancy in the Mare, III. Havemeyer Foundation Monograph Series No. 16. Newmarket: R&W Publications, 2004; pp. 16–18. 92. Haldiman JT. Bovine somite development and vertebral anlagen establishment. Anat Histol Embryol 1981;10:289–309. 93. Ginther OJ. Fixation and orientation of the early equine conceptus. Theriogenology 1983;19:613–23. 94. Arar S, Chan KC, Quinn BA, Waelchli RO, Hayes MA, Betteridge KJ, Monteiro MA. Desialylation of core type 1 O-glycan in the equine embryonic capsule coincides with immobilization of the conceptus in the uterus. Carbohydrate Res 2007;342:1110–15. 95. Silva LA, Ginther OJ. An early endometrial vascular indicator of completed orientation of the embryo and the role
96.
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of dorsal endometrial encroachment in mares. Biol Reprod 2006;74:337–43. Kastelic JP, Adams GP, Ginther OJ. Role of progesterone in mobility, fixation, orientation and survival of the equine embryonic vesicle. Theriogenology 1987;27:655– 62. Enders AC, Liu IK. Lodgement of the equine blastocyst in the uterus from fixation through endometrial cup formation. J Reprod Fertil Suppl 1991;44:427–38. Denker H-W, Betteridge KJ, Sirois J. Shedding of the capsule and proteinase activity in the horse. J Reprod Fertil Suppl 1987;35:708. Ewart JC. Studies of the development of the horse. I. The development during the third week. Trans R Soc Edin 1915;51:287–329. Wilson CA, Davies DC. The control of sexual differentiation of the reproductive system and brain. Reproduction 2007;133:331–59. Betteridge KJ, Waelchli RO, Smith A, Hayes MA. Trophoblastic discs, yolk-sac tubercles and yolk-sac cords in the equine conceptus during the third and fourth weeks of pregnancy. In: Alvarenga M, Wade JF (eds) Proceedings of the 6th International Symposium on Equine Embryo Transfer. Havemeyer Foundation Monograph Series No. 14. Newmarket: R&W Publications, 2005; pp. 3–4. Antczak DF, Oriol JG, Donaldson WL, Poleman C, Stenzler L, Volsen SG, Allen WR. Differentiation molecules of the equine trophoblast. J Reprod Fertil Suppl 1987;35:371–8.
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Chapter 226
Fetal Membrane Differentiation, Implantation and Early Placentation W.R. (Twink) Allen, Susan Gower and Sandra Wilsher
Introduction By the time it reaches 20 days of age, the equine conceptus has survived some pretty unusual and dramatic events. Following its slow 6-day passage through the oviduct, where it causes its own propulsion by secreting prostaglandin E2 (PGE2 ) from day 4 to stimulate contractions and/or relaxation of the oviductal smooth muscle fibers,1–3 the embryo, now at the expanding morula or early blastocyst stage of development, finally passes through the very tight utero-tubal papilla and enters the uterus between 144 and 168 h after ovulation.4 There it remains mobile and moves constantly throughout the uterine domain over the next 10 days, driven by peristaltic contractions of the myometrium, again stimulated by the embryo itself through its rhythmical releases of prostaglandin F2α (PGF2α ) and PGE2 .5, 6 Within 12–24 h of entering the uterus, the outer trophectoderm cells of the still unexpanded blastocyst secrete appreciable quantities of a high-molecular-weight glycoprotein, which, trapped and thereby molded by the still-persisting zona pellucida, forms a very tough, elastic glycocalyx. This completely encapsulates the embryo for the next 20 or more days.7–9 Continued movement of the embryo throughout the uterine lumen for 10 days and the development and persistence of the blastocyst capsule well beyond hatching from the zona pellucida, appear to have at least three, and probably many more, vital functions. First, the capsule insures that the conceptus maintains its spherical shape and is therefore able to be propelled around the uterus.10 Second, it provides remarkable compressive and tensile strength to the
otherwise fragile embryo so enabling it to withstand the considerable myometrial forces that move it around the uterus; capsule-denuded day 7 embryos are unable to survive when transferred back to the uterus,11 although younger day 6 embryos bisected in vitro can re-form a capsule and survive in vivo when retransferred.12, 13 Third, the high negative electrostatic charge of the capsule makes it ‘sticky’ and therefore able to accumulate on its external surface considerable quantities of endometrial gland exocrine secretions (histotroph or ‘uterine milk’14 ). This is the sole source of nutrition for the still unattached and rapidly expanding conceptus and it is absorbed through the capsule and trophectoderm layer to reach the yolk sac fluid.15, 16 The intrauterine mobility of the conceptus also enables the embryo to either secrete an antiluteolytic hormone onto, or otherwise physically interact with, the endometrial epithelium over the majority of the latter’s surface area and thereby prevent it from secreting its cyclical luteolytic spike-like pulses of PGF2α ,17 and instead achieve luteostasis for maintenance of the ‘pregnancy’ state.18 Embryonic movement ceases abruptly on day 16–17 after ovulation19 when the increasing diameter of the conceptus physically prevents it from further passage through the narrowed uterine lumen.20 It now becomes ‘fixed’ at the base of one or other of the uterine horns where it is held firmly in place by a remarkable increase in myometrial tonicity.21, 22 Coincidentally, the vesicle rotates within the tightening uterine lumen, probably in response to a combination of the increased thickness of the endometrium in the mesemetrial (dorsal) segment of the uterus and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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the difference in thickness between the bi- and trilaminar portions of the embryonic membranes, so that the embryo proper comes to lie in the ventral (antimesometrial) quadrant of the uterine horn.20 Thus, by nature of the subsequent expansion of the allantois, the yolk sac, and eventually the attachment of the umbilical cord, occurs in the dorsal quadrant. Having stopped moving, the equine conceptus now begins embryogenesis proper, differentiation and development of the embryonic/fetal membranes, organogenesis, microvillous attachment of the trophoblast to the endometrial epithelium to achieve implantation and, finally, complex microcotyledonary interdigitation between fetal and maternal epithelial layers to create stable, nutritive placentation. This chapter summarizes the sequence of gross and microscopic anatomical changes associated with what may be termed the ‘placentation era’ of equine pregnancy, between 20 and 90 days of gestation. Material used for the description includes numerous conceptuses and biopsies of endometrium re-
covered opportunistically over 41 years at surgical hysterotomy and postmortem from many experimental mares at known stages of gestation,23 including 11 mares from which the gravid uterus was recovered intact at postmortem and perfused-fixed with the conceptus in situ.
Days 20–30 By day 20 after ovulation, the still-spherical conceptus is held firmly at the base of one uterine horn. The highly elastic blastocyst capsule still surrounds the conceptus although it is beginning to thin8 and it is easily dislodged from the surface of the trophoblast (Figure 226.1b). The embryo proper is beginning to take form at one pole of the conceptus and it now contains a primitive beating heart that pumps embryonic blood into the vitelline artery. This runs to the circumferential sinus terminalis at the advancing edge of the vascular mesoderm between the outer ectoderm (chorion) and
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Figure 226.1 (a) A 20-day conceptus exposed in situ in the gravid uterine horn of a perfused-fixed mare’s uterus viewed from the ventral aspect of the uterus. The vestigial embryo is seen in the ‘south’ (embryonic) pole and the sinus terminalis (arrowed) has advanced dorsally to only the ‘tropic of Capricorn’ region at this stage. Scale bar = 1 cm. (b) Photomicrograph of a section of the 20-day perfused-fixed pregnant uterus showing the tightly coiled and elastic blastocyst capsule wedged in a crevice in the floor of the endometrium following its sudden, explosive dislodgement from the embryonic membranes during dissection. Scale bar = 600 µm. (c) Non-vascularized choriovitelline membrane lying adjacent to the maternal endometrium at 20 days of gestation. Note the vestige of blastocyst capsule (arrowed) lying between the trophoblast layer and the endometrial epithelium. Scale bar = 45 µm. (d) Trilaminar (vascularized) choriovitelline membrane in association with the endometrium at 20 days of gestation. The large embryonic blood vessel contains nucleated red blood cells and undifferentiated vascular cells. A small quantity of exocrine secretion (histotroph; arrowed) is exuding from the mouth of an endometrial gland. Scale bar = 45 µm. (See color plate 115).
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Figure 226.1 (Continued) (e) Bicornuate twin conceptuses viewed in situ from the ventral aspect of a perfused-fixed uterus at 24 days of gestation. Each embryo is seen at the lateral border of the enlarging and now well-vascularized allantois. Scale bar = 0.5 cm. (f) A ‘thick’ histological section of both the gravid and non-gravid horns of a perfused-fixed horse uterus at 25 days of gestation. The embryo is seen in the antimesometrial (ventral) quadrant of the gravid horn and the arrows point to the position of the progenitor chorionic girdle. Note the thinning of the endometrium in this antimesometrial quadrant. Scale bar = 1 cm. (g) Unicornuate twin conceptuses viewed from the ventral aspect of the perfused-fixed uterus at 26 days of gestation. The conceptus on the right of the photograph is well vascularized and viable whereas that on the left is pale, bloodless, and clearly dead. Note how the absorptive choriovitelline membrane of the dead conceptus was abutted against the membranes of its viable co-twin rather than to the nutritive endometrium. Scale bar = 1 cm. (h) Embryonic blood vessels in the allantochorion of the viable unicornuate twin conceptus adjacent to the endometrium at 26 days of gestation. Note the accumulated histotroph, which is closely adhered to the trophoblast and endometrial epithelium. Scale bar = 45 µm. (See color plate 115).
inner endoderm (yolk sac) to give the trilaminar membrane, which at this stage occupies about onethird of the total surface area of the conceptus (Figure 226.1a). The allantois is just beginning to form as a sac-like outpouching of the embryonic hindgut. This will expand rapidly over the next 20 days and fuse with the outermost chorion and underlying vascularized mesoderm to form the allantochorion, which will eventually form the definitive placenta. The yolk sac, attached to the primitive embryonic gut through its umbilicus, does not expand further and instead regresses steadily as it becomes increasingly surrounded and replaced by the enlarging allantois. Eventually, between days 50 and 60 after ovulation, the now vestigial yolk sac remains as a remnant within the umbilical cord (see Figure 226.4b).
Histologically around day 20, the chorion is composed of cuboidal/low-columnar epithelioid trophoblast cells with pale-staining basally situated nuclei. It is backed in the trilaminar portion of the conceptus by elongated, fibroblast-like mesodermal cells (Figure 226.1d), scattered amongst which are primitive embryonic blood vessels containing undifferentiated primordial vascular cells and some nucleated red blood cells (Figure 226.1d). It sits snugly against, but is not in any way physically attached to, the luminal epithelium of the endometrium (Figure 226.1c). At this early stage, the endometrium surrounding the conceptus remains relatively thick and the stroma is packed to a good depth with tightly coiled secretory glands. These are lined by a columnar epithelium which is clearly hyperactive in terms
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of exocrine secretory activity. The apical regions or necks of the glands contain accumulated exocrine secretory material (histotroph) that is seen aggregating between the luminal epithelium and trophoblast and becomes closely attached to the latter (Figure 226.1d). The persisting blastocyst capsule at day 20 is apparently only loosely applied to the trophoblast layer and will readily ‘ping off’ the conceptus in toto if the latter is ruptured or otherwise disturbed (Figure 226.1b). Around day 25 of gestation the sinus terminalis and associated vascularized mesoderm have advanced ‘northwards’ from the ‘southern’ (ventral) embryonic pole to be in the equatorial region of the conceptus, and the allantois has expanded ‘southwards’ beneath the embryo thereby appearing to lift the latter ‘northwards’ to a level of about the ‘tropic of Capricorn’ in the ‘southern hemisphere’. The vitelline artery and its associated tracework of mesodermal blood vessels are now more prominent and the embryo proper, with its beating heart, is clearly visible encased within the primitive amnion (Figure 226.1g). In two perfused-fixed uteri, each carrying twin conceptuses, one with a conceptus situated in each uterine horn (bicornuate twins) at 24 days, the other with both vesicles abutted up to each other in the same uterine horn (unicornuate twins) at 26 days, the arrangement of the embryonic membranes is seen clearly after ‘opening the lid’ of the dorsal aspect of the gravid uterine horns (Figure 226.1e,g). In the bicornuate specimens at 24 days, the mesoderm has advanced and the yolk sac regressed although both membranes are now well vascularized (Figure 226.1e). In the unicornuate twins at 26 days, in contrast, one conceptus is well vascularized and is clearly viable while the membranes of its co-twin are pale, bloodless, and beginning to collapse away from the endometrium (Figure 226.1g). Very significantly, the dead twin is aligned sideways in the uterine lumen so that its highly absorptive bilaminar choriovitelline (yolk sac) membrane faces not onto the endometrium in the dorsal quadrant of the uterus as it should, but abuts directly onto the lateral surface of its co-twin, from which it can gain no sustenance (Figure 226.1g). This first-time clear display of the malpositioning of the dying unicornuate twin conceptus provides strong evidence to support the contention that it is simple starvation of essential histotroph occasioned in this manner that is the underlying cause of the very high rate (∼ 80%) of spontaneous conversion of unicornuate twin conceptuses to an ongoing singleton that occurs in Thoroughbred mares between 20 and 30 days of gestation.24, 25 It also emphasizes the fundamental importance of endometrial histotroph as the sole source of nutrition
for the unattached but rapidly growing equine conceptus during the first 40 days of gestation.26 Histologically at the embryo–maternal interface around day 25, the trophoblast layer is still closely applied, but not physically attached, to the luminal epithelium and histotroph is clearly evident exuding from the mouths of the endometrial glands and adhering to the surface of the overlying trophoblast cells for absorption into the allantoic blood vessels (Figure 226.1h). In both the 24- and 26-day perfusedfixed samples, remnants of the blastocyst capsule are still present between the trophoblast and luminal epithelium. Maternal neutrophils are seen passing through the luminal epithelium and appearing to phagocytose and generally clean up the debris of the disintegrating and autolyzing capsule. Furthermore, accumulation of lymphocytes and occasional macrophages in the endometrial stroma beneath areas of persisting and degenerating capsule gives the impression that, far from being a simple dissolution or autolysis of the capsule by the action of proteolytic enzymes,8 the process may include some degree of mild, maternal cell-mediated immune response possibly directed against tissue-specific trophoblast molecules that comprise the innermost layer of the capsule.27 At day 28–30 of gestation, the steadily expanding allantois now comprises about one-third of the total volume of the conceptus and it has pushed the embryo upward even closer to the dorsal or mesometrial quadrant of the uterus (Figure 226.2a). A pale, annulate band is now visible around the conceptus at the junction of the enlarging allantoic and regressing yolk sac membranes (Figure 226.2a). This is the commencing formation of the chorionic girdle,28 which will become thicker, paler, and more prominent during the next 10 days. Histologically, it is seen to be formed initially by a simple folding of the trophoblast layer in this discrete region, slightly akin to a crumpled rug on a shiny floor (Figure 226.2b). The allantoic and yolk sac vascular networks are now very prominent, and a small circle of nonvascularized bilaminar choriovitelline membrane persists beyond the sinus terminalis at what was originally the abembryonic pole of the conceptus (Figure 226.2a). This will become the center of attachment of the umbilical cord in due course. The allantochorionic and choriovitelline membranes are still closely applied, but not physically attached, to the luminal epithelium and histotroph is still prominent both within the apical portions of the endometrial glands and between the two epithelial layers (Figure 226.2c). Interestingly, portions of detached and partly degenerate capsule intermixed with neutrophils and cell debris are also still present between the maternal and fetal epithelial layers.
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Figure 226.2 (a) Intact conceptus at 28 days of gestation lying on its side. The persisting sinus terminalis surrounding the small circle of non-vascularized bilaminar choriovitelline membrane is at the ‘north’ (abembryonic) pole (asterisk). The enlarging allantois, fused with the outer chorion, is pushing the embryo ‘northwards’ towards the abembryonic pole. The developing chorionic girdle (arrowed) is just becoming visible as a narrow, pale, annulate band at the junction of the enlarging allantoic and regressing yolk sac membranes. (b) Section of a chorionic girdle at 30 days of gestation. The folds of trophoblast cells have elongated to create very simple gland-like structures between adjacent folds. Note the visible mitotic figures that illustrate the rapid hyperplasia of these specialized trophoblast cells. Scale bar = 45 µm. (c) Allantochorion ‘glued’ closely to the endometrial epithelium by the histotroph (‘uterine milk’) seen exuding from the mouths of two adjacent endometrial glands in a perfused-fixed uterus at 29 days of gestation. Scale bar = 45 µm. (d) Coils of blastocyst capsule, admixed with histotroph, still persisting as late as day 32 of gestation in a perfusedfixed uterus. The accumulation of lymphocytes in the adjacent endometrial stroma suggests a mild maternal cell-mediated immune response, perhaps against trophoblast-specific antigens in the capsular material. Scale bar = 45 µm. (See color plate 116).
Days 30–40 By day 32 after ovulation, the allantois has expanded to the point that the developing chorionic girdle, marking the junction between the expanding allantois and the regressing yolk sac, is now clearly visible as a pale white band of 1–1.5 cm width situated just ventral to the ‘equator’ of the conceptus. The circle of non-vascularized, bilaminar choriovitelline membrane still persists at the abembryonic pole. Histologically, the same apposition without firm attachment of the now more cuboidal trophoblast cells to the luminal epithelium of the endometrium is evident, with copious amounts of histotroph between the two layers (Figure 226.2c). A dense accumulation of capillaries is seen in the endometrial stroma immediately beneath the luminal epithelium and, surpris-
ingly, lengths of intact, but clearly degenerating, blastocyst capsule and associated neutrophils and cell debris can still be seen between the trophoblast and luminal epithelium. This clear persistence of capsular material well beyond the 21–23-day period of gestation originally proposed for its degeneration and disappearance by Oriol et al.6 raises the interesting question of how much its continued presence between maternal and fetal epithelial layers might interfere with the uptake of the all-important ‘uterine milk’ by the absorptive trophoblast cells of the latter. However, capsule persistence appears patchy and its close association with the endometrial epithelium and associated histotroph suggests the possibility that the degenerating capsular material, with its high content of uterocalin15 and no doubt other endometrial proteins intermixed within its glycocalyx structure, may
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Figure 226.2 (Continued) (e) The dorsal half of an intact horse conceptus at 34 days of gestation. The circle of bilaminar choriovitelline membrane persists at what was formerly the abembryonic pole (asterisk). The fetus, now enveloped in its amnion, lies immediately beneath, supported by the enlarging allantois. The pale, thickened annulate chorionic girdle (arrowed) is clearly seen at the junction of the well-vascularized allantois and the yellow degenerating yolk sac. Splits appearing in the chorionic girdle tissue reflect its maturity and readiness to dehisce from the fetal membranes and adhere to, and invade, the overlying maternal endometrium. Scale bar = 7 mm. (f) A mature chorionic girdle, backed by two large fetal blood vessels, sitting on the endometrium in a perfused-fixed uterus at 34 days of gestation. Scale bar = 600 µm. (g) High-power photomicrograph of the 34-day chorionic girdle seen in the previous figure. Note the even, epithelial-like arrangement of the tall columnar, already binucleated, trophoblast cells facing the much smaller cuboidal luminal epithelial cells of the endometrium and the convoluted glandlike arrangement of the trophoblast cells within the body of the girdle. Secretory material tentatively binds the surface of the girdle to the luminal epithelium of the endometrium. Scale bar = 45 µm. (h) The 34-day horse fetus dissected from the conceptus shown in Figure e. Organogenesis is essentially complete at this stage and both forelimb and hindlimb buds are protruding. It is therefore formally upgraded from the status of an embryo to a fetus.57 Scale bar = 2.5 mm. (See color plate 116).
constitute a further form of nutrients for the conceptus (Figure 226.2d). The chorionic girdle, progenitor tissue of the endometrial cups29 at day 30, is still composed of fairly simple folds of trophoblast cells (Figure 226.2b). However, the cells are now more columnar in shape and, as the folds become laterally compressed by uterine tone, they will begin to assume the appearance of simple gland-like structures (Figure 226.2g). Mitotic figures are seen in the rapidly multiplying trophoblast cells of the developing girdle at day 30 (Figure 226.2b) and in the mature, pre-invasive girdle at day 34 (Figure 226.2g). By day 34, the chorionic girdle, due to the continuing marked hyperplasia of the trophoblast cells, is now much thicker (Figure 226.2e) and the elongat-
ing folds of cells have become squashed together to form a line of simple excretory glands opening out towards the maternal surface (Figure 226.2f,g). Indeed, these glands do liberate an alcian blue-positive material which helps to bind fetal and maternal surfaces together and facilitate the separation of the entire mature girdle from the underlying basal layer of trophoblast to enable the girdle cells then to invade the endometrium via the mouths of the endometrial glands (Figure 226.2g).30, 31 On either side of the chorionic girdle the trophoblast of the allantochorionic and choriovitelline membranes remains cuboidal to low columnar and closely apposed to the luminal epithelium of the endometrium by the cementing properties of the endometrial gland histotroph. The embryo, having completed essential organogenesis and
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Fetal Membrane Differentiation, Implantation and Early Placentation developed visible limb buds, is now correctly termed a fetus (Figure 226.2h). At day 37 the fetus is still seen resting on the regressing yolk sac (Figure 226.3a). The chorionic girdle is in the middle of invading the endometrium to form the endometrial cups and in many areas the whole thickness of girdle is now well attached to the surface of the endometrium (Figure 226.3b. See also colour plate) with the leading edge girdle cells having destroyed and/or dislodged the luminal epithelium (Figure 226.3c). Virtually all the girdle cells have already become binucleate, as reported originally by Wooding et al.32 , although the two nuclei are still closely abutted against each other (Figure 226.3c). In other areas, although separated from the basal layer of trophoblast and firmly attached to the endometrium, the girdle cells have not yet damaged or invaded the maternal tissue to any great de-
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gree and, in some parts, the chorionic girdle still remains to be become attached to the endometrium. By day 38, however, the invasion process is well under way and the large flocculent, binucleate girdle cells are seen moving down the apical portions of the endometrial glands, dislodging the glandular epithelial cells from their basement membrane as they do so (Figure 226.3d). Interestingly, appreciable numbers of maternal lymphocytes have already accumulated around the attached, only partly invaded, chorionic girdle, presumably attracted by the known expression of paternally inherited class I major histocompatibility complex (MHC) antigens,33 and possibly also other trophoblast-specific foreign antigens, on the surface of the invading cells.27, 34 Yet despite the close physical presence of the two types of cell, antigenically foreign fetal trophoblast and maternal leukocytes, the
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Figure 226.3 (a) A 37-day horse conceptus exposed in situ. The fetus, in its amnion, is resting on the surface of the regressing yolk sac with the well-vascularized allantochorion nearer the camera. A band of pale, very young endometrial cups is seen at the edge of the yolk sac in the upper portion of the photograph (arrowed). Scale bar = 1.25 cm. (b) Chorionic girdle firmly attached to the endometrium and clearly separated from the underlying layer of trophoblast in a perfused-fixed uterus at 37 days of gestation. The girdle cells have disrupted the luminal epithelium but have not yet begun to pass down the endometrial glands. Scale bar = 170 µm. (c) High-power photomicrograph at the leading edge of the 37-day chorionic girdle attached to the luminal surface of the endometrium. Note the pseudostratified layer of very elongated, pale-staining binucleate girdle cells lined up on the apical surface to attack and destroy the luminal epithelial cells, darkly stained vestiges of which (arrowed) are still present. Scale bar = 45 µm. (d) Section from the uterus of a 38-day pregnant mare showing the chorionic girdle invading the endometrium. The palestaining epithelioid girdle cells have dislodged the luminal epithelium (darker staining cells on left of photograph) and are lifting the glandular epithelial cells off their basement membrane (arrowed) as they pass down the endometrial glands as the least line of resistance. Note the lymphocytes already accumulated in the endometrial stroma. Scale bar = 45 µm. (See color plate 117).
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Figure 226.3 (Continued) (e) Low-power section of the newly formed endometrial cup at day 40 of gestation. The apical portions of the endometrial glands have been obliterated and the cup is now a solid mass of enlarging trophoblast cells surrounded by a dark band of appreciable numbers of maternal leukocytes. Scale bar = 600 µm. (f) Horse conceptus at 42 days of gestation. The allantochorion has a pale, hazy appearance, perhaps due to commencing development of the chorionic villi. The chorionic girdle has invaded the endometrium to form the endometrial cups, leaving just a few vestigial tags of tissue at the junction of the well-vascularized allantochorion and the yellow, still regressing yolk sac. The circle of non-vascularized bilaminar choriovitelline membrane still persists at the previously abembryonic pole (asterisk). The fetus is enveloped by its amnion. Scale bar = 1.5 cm. (g) Perfused-fixed uterus at day 46 of gestation with the gravid horn opened ventrally to reveal the intact conceptus in situ. The vestigial yolk sac (asterisk) is seen below the fetus and the arrows point to both a small, isolated endometrial cup and a longer, unbroken line of cup tissue. Scale bar = 1.75 cm. (h) The endometrium–allantochorion interface in a perfused-fixed uterus at day 46 of gestation. Note the commencing interdigitation of simple chorionic villi with accommodating sulci in the surface of the endometrium. Also, the accumulated pink-staining histotroph in the mouths of some endometrial glands. Scale bar = 170 µm. (See color plate 117).
latter do not appear actively to attack the former (Figure 226.3e), as certainly occurs later in the lifespan of the endometrial cups. On either side of the developing cups, the trophoblast cells of the chorion are still only closely apposed, but not firmly attached, to the luminal epithelium which is beginning to show more pronounced undulations in its surface, perhaps as a prelude to impending trophoblast interdigitation. By this stage of gestation it is becoming noticeable that the endometrial glands in the stretched and thinning uterine wall around the expanding conceptus are spaced further apart and penetrate much less deeply into the endometrium than those elsewhere in the gravid horn and, especially, in the non-gravid horn. It must be supposed that the estrogens being secreted locally by the conceptus membranes35 and perhaps other locally produced growth factors,36 are
working hard to induce the endometrial glands to secrete increased quantities of vital histotroph to meet the fetal needs.
Days 41–60 From day 40 onwards, as myometrial tone declines, the conceptus starts to lose its spherical shape (Figure 226.3a,f,g) as continuing expansion of the allantois pushes the allantochorion further up the gravid uterine horn on one side and towards the uterine body on the other. This outermost membrane acquires a pale, hazy appearance to the naked eye, presumably due to the commencing development of blunt chorionic villi all over its surface (Figure 226.3f). Allantoic expansion, combined with yolk sac regression, has now
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Fetal Membrane Differentiation, Implantation and Early Placentation pushed the fetus ‘northwards’ or dorsally almost to the previously abembryonic pole where, by day 46, the vestigial yolk sac is contained by the encroaching allantoic membrane in what will form the umbilical cord (Figure 226.3g).37 Continued expansion and elongation means that the conceptus comes to occupy the remainder of the gravid horn and the whole of the uterine body by days 60–65 (Figure 226.4b), pushing on thereafter to finally fill the whole of the non-gravid horn by days 80–85 (Figure 226.4h). All the while the external or maternal surface of the allantochorion is becoming red and roughened in appearance due to the steady development of the microcotyledons (Figure 226.4h). Histologically, as seen in perfused-fixed uteri at days 43 and 46 of gestation, the still low-columnar
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trophoblast layer is at last firmly and stably adhered to the endometrium by a microvillous border on the apical surfaces of the two epithelial layers. Simple, blunt protrusions or villi of allantochorion interdigitate with corresponding upgrowths or sulci of endometrium, and capillary formation remains prominent in the subepithelial endometrial stroma (Figure 226.3h). The endometrial glands, although still sparsely arranged and relatively shallow in the stretched endometrium around the conceptus bulge, are continuing to produce copious amounts of histotroph, which is seen as a thin eosin-stained layer between the closely applied fetal and maternal surfaces and, by day 46, is already accumulating in ‘blobs’ or ‘piles’ above the mouths of endometrial glands (Figure 226.3h). The trophoblast overlying
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Figure 226.4 (a) Low-power section of an endometrial cup at day 46 of gestation. The cup is now a solid, densely packed mass of fetal cup cells interspersed amongst which are some partially distended endometrial glands. The apical surface of the cup is already concave due to overgrowth of the tissue at the lateral edges. The accumulated maternal leukocytes are already slightly reduced in numbers and they are restricted to the lateral edges and base of the cup tissue. Large lymph sinuses are forming in the stroma beneath the cup. Scale bar = 600 µm. (b) View of the 58-day perfused-fixed uterus with the conceptus in situ. The fetus lies on the ventral floor of the uterus close to the bifurcation of the uterine horns. Note the vestigial yolk sac (asterisk) forming part of the umbilical cord and the continuous line of pale, raised endometrial cups running away at an angle from the neck of the fetus. Scale bar = 2 cm. (c) Close-up view of the rather long, thin endometrial cups lying beneath the well-vascularized allantochorion in the perfused-fixed uterus of day 58 of gestation. Note the accumulation of yellow exocrine secretion (arrowed) in the concave surface of the cups in the upper part of the photograph. Scale bar = 5 mm. (d) High-power section of the now increasingly convoluted allantochorion–endometrium interface in the perfused-fixed uterus of day 58 of gestation. Note that most of the fetal red blood cells in the allantoic blood vessels have now lost their nuclei. Scale bar = 45 µm. (See color plate 118).
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Figure 226.4 (Continued) (e) High-power section of the placental interface at day 58 of gestation. Note the continuing prominent role of endometrial gland histotroph. Scale bar = 45 µm. (f) Section of the allantochorion–endometrium interface in the perfused-fixed uterus at day 68 of gestation. The interdigitation between maternal and fetal epithelial layers is becoming increasingly convoluted and the first signs of branching of the primary villi are evident. Scale bar = 170 µm. (g) High-power section of an areolus in the placental interface at day 68 of gestation. Note the specialized very tall columnar, highly absorptive trophoblast cells. Scale bar = 45 µm. (h) Intact conceptus recovered from a mare at day 93 of gestation. The reddened and thickening allantochorion has now expanded to occupy the whole uterus. The line of scars at the base of the right horn (arrowed) corresponds to the avillous areas that had overlain the endometrial cups. Scale bar = 8.5 cm. (See color plate 118).
these histotroph piles loses its microvillous attachment to the endometrial epithelium and the cells begin to become elongated, vacuolated, and pseudostratified as they take up, by pinocytosis, the accumulated histotroph for transport to the allantoic blood vessels (Figure 226.4g). By days 43–46 the endometrial cups are seen grossly as discrete, pale protuberances of 1–2 cm width projecting above the surface of the endometrium, and arranged in a circle or horseshoe in the area of endometrium abutting the yolk sac–allantois junction (Figure 226.3g). The form of the cups depends upon the configuration of the endometrium at the time of attachment and invasion of the chorionic girdle around days 35–37. The cups can be small, discrete structures of 2–6 cm length well separated from their neighbors (Figure 226.3g); they form in this manner due to the folding of the endometrium, which allows the chorionic girdle
to gain access only to the endometrium at the tips of adjacent folds. Alternatively, the cups may form long, unbroken ribbons of tissue as a consequence of the chorionic girdle being able to attach to, and penetrate, a more flattened, less convoluted endometrium (Figure 226.4g). Histologically around day 40, the enlarging binucleate cup cells, having progressed down the endometrial glands (Figure 226.3d) and now broken through their basement membranes, have progressed into the endometrial stroma (Figure 226.3e) where they still attract and come into close physical contact with appreciable numbers of maternal leukocytes (Figure 226.3d,e). The apical portions of most endometrial glands are obliterated during the invasion process but the fundic portions remain intact and become increasingly distended (Figure 226.4a), both as a result of accumulated exocrine secretion and as a consequence of some sort of hyperstimulation,
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Fetal Membrane Differentiation, Implantation and Early Placentation perhaps occasioned by the high concentrations of equine Chorionic Gonadotropin (eCG), the principal hormonal product of the fetal cup cells, to which the glands are exposed. These persisting basal glands can become grossly dilated during the 60–80-day lifespan of the endometrial cups and the trapped exocrine secretion they contain stains intensely with periodic acid–Schiff (PAS), thereby indicating a high content of glycogen.38 Indeed, this strong PAS staining induced Cole and Goss39 and Clegg et al.,40 mistakenly as it turned out, to suggest that the epithelium lining the dilated glands in the cups was probably the source of eCG. In reality, it is more likely that eCG simply diffuses from the cup cells into the gland luminae and accumulates in the trapped exocrine secretion produced by the hyperstimulated gland epithelium. By days 55–60 the fetus is now clearly horselike in appearance and is actively kicking and moving (Figure 226.4b).41 The umbilical cord is now well formed and is long enough to allow the fetus a considerable degree of movement within the proportionately large volume of allantoic fluid. The tracework of fetal blood vessels spreads out over the surface of the allantochorion from the base of the umbilical cord (Figure 226.4b). Histologically at the placental interface, fetal and maternal epithelial layers are now closely applied to each other by their microvillous union, and the simple undulations have deepened and lengthened appreciably to give the first intimation of what will eventually become the established microcotyledons (Figure 226.4d). However, very frequently between the short lengths of close interdigitation occur the previously mentioned areolae38 opposite the mouths of endometrial glands where secreted histotroph accumulates in a definite space between the luminal epithelium of the endometrium and the elongated and pseudostratified, absorptive trophoblast cells of the chorion (Figure 226.4e). The frequent occurrence of these areolae along the placental interface, and their persistence throughout gestation (Figure 226.4g)42 attests to the continuing importance of histotrophic nutrition throughout fetal life. The endometrial cups are well raised above the surface of the endometrium and are now showing the characteristic indentation in their luminal surface (Figure 226.4a), for which reason Schauder43 originally called them ‘endometrial cups’. This depression in the cup surface occurs as a result of the commencing degeneration and sloughing of large cup cells in the center of the apical surface of each cup – i.e., the cells farthest from adequate vasculature – combined with overgrowth of the cup cells and other tissues at the lateral edges of the cup (Figure 226.4a). Histologically, the cups now consist of a dense accumulation
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of the large, fetal endometrial cup cells interspersed with the dilated fundic portions of endometrial glands (Figure 226.4a). Interestingly, as highlighted previously by Grunig et al.,44 the density of maternal leukocytes accumulated around the periphery of the cup is notably reduced at this point in gestation compared to the numbers seen in earlier pregnancy. This reduction in the intensity of the maternal inflammatory response to the fetal cups is also evidenced when viewing the intact endometrial cups in situ in the uterus. For example, around days 40–45 a pronounced inflammatory reddening is visible in the stroma at the lateral edges of the cups whereas this hyperemia has disappeared completely by day 60 (Figure 226.4c).
Days 60–80 By day 68 the conceptus has expanded to fill the body and gravid uterine horn, with further progression to about one-third the length of the non-gravid horn. It extends further to reach the tip of the non-gravid horn by days 80–85 (Figure 226.4h). As mentioned, the allantochorion is becoming increasingly reddened and stippled in appearance due to the development of microcotyledons on its external surface, and the fetus still lies on its side on the ventral floor of the uterus close to the bifurcation of the uterine horns (Figure 226.4b,h. See also colour plate). The endometrial cups reach their maximum size and eCG-secreting potential by days 60–70 and, thereafter, increasing quantities of a sticky yellow exocrine amalgam accumulate in the central depression on the surface of each cup (Figure 226.4c. See also colour plate). This material, composed of a mixture of dead, sloughed cup cells and the inspissated exocrine secretion released from the now unblocked dilated endometrial glands, is extremely potent in eCG activity and may contain millions of units of gonadotropin per gram wet weight of tissue.45 Histologically, the cups are showing clear evidence of cell degeneration and sloughing in the central apical region, and the density of the accumulation of lymphocytes and other leukocytes in the stroma around the periphery of the cup is increasing again after the temporary decline between days 45 and 60. At the base of some cups, the accumulated lymphocytes can be seen attacking and killing the cup cells, thereby allowing the leukocytes to move further into the cup tissue and broaden the killing process. Eventually, between days 90 and 140, the entire necrotic cup and admixed inspissated exocrine cup secretion is sloughed off the surface of the endometrium to lie free in the uterine lumen between the endometrium and allantochorion. A scar devoid of
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microcotyledons persists on the surface of the allantochorion that had overlain the endometrial cups (Figure 226.4h) and, in the dorsal quadrant of the uterus, the lump of dehisced cup tissue and secretion may indent into the surface of the allantochorion to form pedunculated allantochorionic pouches, which hang into the allantoic cavity.40 At the fetal–maternal interface, the interdigitation, with folding and branching of the endometrium and attached allantochorion, is proceeding steadily to give the early stage microcotyledons (Figure 226.4f). Each multifolded and multibranched microcotyledon stems from a single blunt chorionic villus and accommodating endometrial sulcus first seen around days 40–43, and still the histotrophabsorbing pseudostratified areolae remain very evident along the allantochorion-endometrium border (Figure 226.4e,g).
Conclusions Thus, by days 80–85 of gestation in the mare and other female equids, life for the developing fetus has settled down to a stable period of steady growth following the dramatic changes, upheaval, and slow feto-maternal connection in early gestation. The allantochorion has advanced to occupy the whole of the uterine interior and is now busy establishing increasingly complex microcotyledonary attachment to the endometrium over its whole surface area for hemotrophic exchange, while still persisting with extensive histotrophic nutritional input in the form of the areolae, likewise spread across the entire chorionic surface. The eCG secreted by the fetal endometrial cups from days 37–38 has stimulated the development of a series of secondary or supplementary corpora lutea in the maternal ovaries to augment progesterone levels in maternal blood. Furthermore, the well-established allantochorion is now beginning to synthesize and secrete increasing quantities of 5αdihydroprogesterone and other biologically active 5α-reduced progestagens, utilizing maternal sources of cholesterol,46 to stimulate directly the continued secretion of histotroph by the endometrial glands and to maintain quiescence of the myometrium; the supplementary corpora lutea regress and the maternal ovaries become completely inactive from around mid-gestation.47 Also commencing around day 80, the multifunctional trophoblast begins to synthesize increasing quantities of phenolic and ring B unsaturated estrogens,48–50 utilizing C-19 precursor androgens secreted by the fetal gonads;51–54 these undergo tremendous enlargement and subsequent regression between days 80 and 320 of gestation.47, 55 Thus, the stage is finally set for the fetus to grow steadily dur-
ing the remainder of the 11-month gestation and, by nature of its dual hemotrophic and histotrophic forms of nutrition, to reach the remarkably precocious stage of development it must attain in order to survive birth and a rapid transition to an extrauterine existence.56
References 1. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 secretion by oviductal transport-stage equine embryos. Biol Reprod 1991;45:540–3. 2. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 hastens oviductal transport of equine embryos. Biol Reprod 1991;45:544–6. 3. Weber JA, Woods GL, Lichtenwalner AB. Relaxatory effect of prostaglandin E2 on circular smooth muscle isolated from the equine oviductal isthmus. Biol Reprod Monograph Series 1995;1:125–30. 4. Battut I, Colchen S, Fieni F, Tainturier D, Bruyas JF. Success rate when attempting to non surgically collect equine embryos at 144, 156 and 168 hours after ovulation. Equine Vet J Suppl 1997;25:60–2. 5. Stout TAE, Allen WR. Role of prostaglandins in intrauterine migration of the equine conceptus. Reproduction 2001;121:771–5. 6. Stout TAE, Allen WR. Prostaglandin E2 and F2α production of equine conceptuses and concentrations in conceptus fluids and uterine flushings recovered from early pregnant and dioestrous mares. Reproduction 2002;123:261–8. 7. Betteridge KJ. The structure and function of the equine capsule in relation to embryo manipulation and transfer. Equine Vet J Suppl 1989;8:92–100. 8. Oriol JG, Sharom FJ, Betteridge KJ. Developmentally regulated changes in the glycoproteins of the equine embryonic capsule. J Reprod Fertil 1993;99:653–64. 9. Oriol JG, Betteridge KJ, Clarke AJ, Sharom FJ. Mucin-like glycoproteins in the equine embryonic capsule. Mol Reprod Dev 1993;34:255–65. 10. Ginther OJ. Mobility of early equine conceptus. Theriogenology 1983;19:603–11. 11. Stout TAE, Meadows S, Allen WR. Stage-specific formation of the equine blastocyst capsule is instrumental to hatching and to embryonic survival in vivo. Anim Reprod Sci 2005;87:269–81. 12. McKinnon AO, Carnevale EM, Squires EL, Carney NJ, Seidel GE Jr. Bisection of equine embryos. Equine Vet J Suppl 1989;8:129–33. 13. Skidmore J, Boyle MS, Cran D, Allen WR. Micromanipulation of equine embryos to produce monozygotic twins. Equine Vet J Suppl 1989;8:126–8. 14. Amoroso EC. Placentation. In: Parkes AS (ed.) Marshall’s Physiology of Reproduction, 3rd edn. Vol. 2, London: Longmans Green, 1952; pp. 127–297. 15. Stewart F, Charleston B, Crossett B, Barker PJ, Allen WR. A novel uterine protein that associates with the blastocyst capsule in equids. J Reprod Fertil 1995;105:65–70. 16. Crossett B, Suire S, Herrler A, Allen WR, Stewart F. Transfer of uterine lipocalin from the endometrium of the mare to the developing equine conceptus. Biol Reprod 1998;59:483–90. 17. Neely DP, Kindahl H, Stabenfeldt GH, Edqvist LE, Hughes JP. Prostaglandin release patterns in the mare:
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18. 19. 20. 21. 22. 23. 24.
25.
26. 27.
28. 29.
30.
31.
32.
33.
34.
35.
36.
37.
38. 39.
physiological, pathophysiological and therapeutic responses. J Reprod Fertil Suppl 1979;27:181–9. Allen WR. Maternal recognition and maintenance of pregnancy in the mare. Anim Reprod 2006;2;4:209–23. Ginther OJ. Fixation and orientation of the early equine conceptus. Theriogenology 1983;19:613–23. Ginther OJ. Dynamic physical interactions between the equine embryo and uterus. Equine Vet J Suppl 1985;3:41–7. Bain AM. The ovaries of the mare during early pregnancy. Vet Rec 1957;80:229–31. Van Niekerk CH. I. Early clinical diagnosis of pregnancy in mares J S Afr Vet Med Ass 1965;36:55–8. Van Niekerk CH, Allen WR. Early embryonic development in the horse. J Reprod Fert Suppl 1975;23:495–8. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of Flatrace and National Hunt Thoroughbred mares and stallions in England. Equine Vet J 2007;39: 438–45. Morris LH-A, Allen WR. Reproductive efficiency of intensively managed Thoroughbred mares in Newmarket. Equine Vet J 2002;34:51–60. Allen WR. Fetomaternal interactions and influences during equine pregnancy. Reproduction 2001;121:513–27. Oriol JG, Poleman JC, Antczak DF, Allen WR. A monoclonal antibody specific for equine trophoblast. Equine Vet J Suppl 1989;8:14–18. Ewart JC. A critical period in the development of the horse. London: Adam and Charles Black, 1897. Allen WR, Moor RM. The origin of the equine endometrial cups. I. Production of PMSG by fetal trophoblast cells. J Reprod Fertil 1972;29:313–16. Allen WR, Hamilton DW, Moor RM. The origin of the equine endometrial cups. II. Invasion of the endometrium by trophoblast. Anat Rec 1973;177:485–502. Hamilton DW, Allen WR, Moor RM. The origin of the equine endometrial cups. III. Light and electron microscopic study of fully developed equine endometrial cups. Anat Rec 1973;177:503–18. Wooding FBP, Morgan G, Fowden AL, Allen WR. A structural and immunological study of chorionic gonadotrophin production by equine trophoblast and cup cells. Placenta 2001;22:749–67. Crump A, Donaldson WL, Miller J, Miller J, Kydd JH, Allen WR, Antczak DF. Expression of major histocompatibility complex (MHC) antigens on horse trophoblast. J Reprod Fert Suppl 1987;35:379–88. Antczak DF, Poleman JC, Stenzler L, Volsen SG, Allen WR. Differentiation molecules of the equine trophoblast. J Reprod Fert Suppl 1987;35:371–8. Heap RB, Hamon M, Allen WR. Studies on oestrogen synthesis by the preimplantation equine conceptus. J Reprod Fert Suppl 1982;32:343–52. Lennard SN, Stewart F, Allen WR, Heap RB. Growth factor production in the pregnant equine uterus. Biol Reprod Monograph Series 1995;1:161–70. Whitwell K, Jeffcott LB. Morphological studies on the fetal membranes of the normal singleton foal at term. Res Vet Sci 1975;19:44–55. Amoroso EC. Endocrinology of pregnancy. Br Med Bull 1955;11:117–25. Cole HH, Goss H. The source of equine gonadotrophin. In: Essays in Biology in Honour of Herbert M Evans. Berkeley: University of California Press, 1943; pp. 107–19.
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40. Clegg MT, Boda JM, Cole HH. The endometrial cups and allanto-chorionic pouches in the mare with emphasis on the source of equine gonadotrophin. Endocrinology 1954;54:448–63. 41. Allen WR, Bracher V. Videoendoscopic evaluation of the mare’s uterus; III. Findings in the pregnant mare. Equine Vet J 1992;24:285–91. 42. Allen WR, Gower S, Wilsher S. Immunohistochemical localisation of vascular endothelial growth factor (VEGF) and its two receptors (Flt-I and KDR) in the endometrium and placenta of the mare during the oestrous cycle and pregnancy. Reprod Dom Anim 2007;42:516–26. 43. Schauder W. Untersuchungen uber die eithaute und Embryotrophe des pferdes. Arch Anat Physiol 1912;259– 302. ¨ 44. Grunig G, Triplett L, Canady LK, Allen WR, Antczak DF. The maternal leucocyte response to the endometrial cups in horses is correlated with the developmental stages of the invasive trophoblast cells. Placenta 1995;16:539– 43. 45. Rowlands IW. Levels of gonadotrophin in tissues and fluids with emphasis on domestic animals. In: Cole HH (ed.) Gonadotrophins: Their Chemical and Biological Properties and Secretory Control. San Francisco: WH Freeman & Co., 1963; pp. 74–112. 46. Holtan DW, Houghton E, Silver M, Fowden AL, Ousey JC, Rossdale PD. Plasma progestagens in the mare and newborn foal. J Reprod Fert Suppl 1991;44:517–28. 47. Cole HH, Hart GH, Lyons WR, Catchpole HR. The development and hormonal content of fetal horse gonads. Anat Rec 1933;56:275–93. 48. Cox JE. Oestrone and equilin in the plasma of the pregnant mare. J Reprod Fert Suppl 1975;23:463–8. 49. Raeside JI, Liptrap RM. Patterns of urinary oestrogen excretion individual pregnant mares. J Reprod Fert Suppl 1975;23:469–75. 50. Urwin VE, Allen WR. Pituitary and chorionic gonadotrophin control of ovarian function during early pregnancy in equids. J Reprod Fert Suppl 1982;32:371–82. 51. Bhavnani BR, Short RV, Solomon S. Formation of oestrogens by the pregnant mare. I. Metabolism of 7–3Hdehydroisoandrosterone and 4–14C-androstenedione injected into the umbilical vein. Endocrinology 1969;85: 1172–9. 52. Bhavnani BR, Short RV, Solomon S. Formation of oestrogens by the pregnant mare. II. Metabolism of 14C-acetate and 3H-cholesterol injected into the fetal circulation. Endocrinology 1971;89:1152–7. 53. Pashen RL, Sheldrick EL, Allen WR, Flint APF. Dehydroepiandrosterone synthesis by the fetal foal and its importance as an oestrogen precursor. J Reprod Fert Suppl 1982;32:389–97. 54. Tait AD, Santikarn LC, Allen WR. Identification of 3b-hydroxy-5,7 pregnandien-20-one and 3b-hydroxy-5,7 androstadien-17-one as endogenous steroids in the foetal horse gonad. J Endocr 1983;99:87–92. 55. Hay MF, Allen WR. An ultrastructural and histochemical study of the interstitial cells in the gonads of the fetal horse. J Reprod Fert Suppl 1975;23:557–61. 56. Rossdale PD, Silver M. The concept of readiness for birth. J Reprod Fert Suppl 1982;32:507–10. 57. Working Party on Terminology. J Reprod Fert Suppl 1982;32:647–52.
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Chapter 227
Maternal Recognition of Pregnancy Karen J. McDowell and Daniel C. Sharp
The estrous cycle and the role of the uterus in luteal regression In farm animals each estrous cycle represents an opportunity for mating and for establishment of pregnancy. In the diestrus phase of the cycle the uterus is simultaneously prepared for two conflicting events: luteolysis and luteal maintenance for pregnancy. If there is no viable conceptus in the uterus approximately 2 weeks after ovulation, the corpus luteum (CL) regresses allowing for a return to estrus and another opportunity to establish pregnancy. If, however, there is a viable conceptus in the uterus, the conceptus must disrupt the luteolytic process in order to maintain progesterone production, which is requisite for a successful pregnancy. Thus successful pregnancy in farm animals requires a signal from the conceptus that results in maintenance of the corpus luteum, in a process known as ‘maternal recognition of pregnancy’.1 In mares an early form of pregnancy recognition takes place in the oviduct, which permits selective transport of the developing conceptus but does not permit transfer of unfertilized ova;2 this is likely mediated through conceptus production of prostaglandin E2 (PGE2 ).3 While transport of the developing conceptus through the oviduct to the uterus is clearly an obligatory step for a successful pregnancy, it does not guarantee successful accomplishment of the critical step of providing continued luteal support for that pregnancy. An essential consequence of maternal recognition of pregnancy and luteal maintenance is the continued secretion of uterine histotrophe (uterine milk), the nutrients required for conceptus growth and development. The uterus is necessary for regression of the CL in most mammalian species, with the human being a notable exception. Loeb4 first hysterectomized
guinea pigs and noted that the lifespan of the CL was extended far beyond normal. Ginther and First5 reported the effects of complete and partial hysterectomy in pony mares. Complete hysterectomy resulted in maintenance of the CL in all mares, and hemihysterectomy (one uterine horn and approximately one-half of the uterine body removed) resulted in maintenance of the CL in about half of the mares. In mares the side of uterine ablation relative to the CL was not important, indicating that the uterine signal for luteolysis likely reaches the CL via the systemic circulation. Thus the uterus, and more specifically the uterine endometrium, plays a critical role in normal luteal regression, and therefore must play a critical role in maternal recognition of pregnancy in mares.
Prostaglandin F2α , the natural uterine luteolysin, is altered in the presence of a conceptus It is now well established that prostaglandin F2α (PGF2α ) is the natural uterine luteolysin in farm animals, including mares.6–8 In cycling mares the onset of luteal regression at about 14 days after ovulation coincides with maximum PGF2α in the uterine vein and uterine lumen,9–11 maximum PGF2α content and production by the uterine endometrium,12 and maximum concentration of the PGF2α metabolites, 13,14dihydro-15-keto-PGF (PGFM) and 11-ketotetranor PGF, in the peripheral circulation.13–15 One of the critical requirements of maternal recognition of pregnancy is that the CL is maintained for continued secretion of progesterone. The conceptus could conceivably accomplish this goal through one or a combination of several ways:
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Maternal Recognition of Pregnancy 1. preventing the secretion of PGF2α (i.e., a luteostatic effect); 2. altering the distribution of PGF2α so that it does not reach the CL in an appropriate form (also a luteostatic effect); or 3. secreting a substance that counteracts the effects of PGF2α at the level of the CL (a luteotrophic or antiluteolytic effect).
In the mare there is abundant evidence to support the first of these mechanisms. Although the CL of pregnant mares are capable of binding PGF2α 16 and undergoing regression in response to exogenous PGF2α ,17 and the endometrium of pregnant mares is capable of producing PGF2α in vitro,12 there is considerable evidence that the capacity of the endometrium to release PGF2α is diminished in the presence of the conceptus. At the expected time of luteolysis, PGF2α concentration is lower in the uterine vein and uterine lumen,9, 18 and concentrations of PGFM and 11-ketotetranor metabolites are lower in the uterine lumen and peripheral circulation of pregnant compared to non-pregnant mares.7, 14, 15 In addition, co-incubation of endometrial explants with conceptus membranes in vitro reduced both PGF2α and PGFM concentrations over endometrium incubated alone.7, 8, 11 Co-incubating endometrial explant tissues with conceptus membranes contained in small bags of dialysis tubing suggested that the conceptus PGF2α inhibitory factor(s) may be of relatively low molecular weight.19 PGF2α production by endometrial explants was inhibited when conceptus membranes were contained within dialysis bags with molecular exclusion limits of 12 000–14 000, 6000–8000, and 3500, but not 1000 daltons. Thus the failure of the conceptus factors to suppress endometrial PGF2α production at the molecular exclusion limit less than 3500 suggests that the conceptus factor is larger than most common ions, steroids, or even other prostaglandins. The conceptuses of cattle and sheep secrete a protein hormone called interferon-tau, which dampens or suppresses PGF2α production by the uterine endometrium,20–23 and is considered to be the main signal for maternal recognition of pregnancy in those species. The proteins are acidic, with molecular weights of about 17 000 (sheep) and 21 000 (cows) daltons. Assessment of equine conceptus-produced proteins by two-dimensional polyacrylamide gel electrophoresis did not reveal similar proteins,24 no RNA transcripts for similar interferons were detected in equine conceptus tissues,25 and antibodies raised against the sheep interferons did not cross-react with conceptus-conditioned media or with uterine flushings from pregnant mares.19 Thus there is no evi-
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dence for maternal recognition of pregnancy in mares to be accomplished via a conceptus interferon-tau. Equine26–28 and porcine29, 30 conceptuses produce large amounts of estrogens, and estrogens appear to be the main conceptus signal for maternal recognition of pregnancy in pigs. Estrogens synthesized and secreted by the pig conceptus directly or indirectly alter the direction of PGF2α secretion by the uterine endometrium in pregnant compared to nonpregnant pigs.30–33 In non-pregnant gilts, at about 15 days after ovulation, PGF2α is secreted in an ‘endocrine’ direction, toward the myometrium, where it gains access to the vascular tree and is carried to the CL. In contrast, in pregnant gilts PGF2α is secreted in an ‘exocrine’ direction, toward the uterine lumen, where it is sequestered, thus protecting the CL.30 Evidence in support of this theory comes from a variety of studies, perhaps the most compelling of which involved the use of a two-chambered perfu¨ sion device called an Ussing chamber.34 In this device endometrial tissue can be placed between two chambers such that secretion of PGF2α toward the myometrial and luminal sides can be studied differentially. When such studies were conducted using endometrium from non-pregnant gilts the majority of PGF2α was secreted toward the myometrial side. However, when endometrium from pregnant gilts was used the majority of PGF2α was secreted toward the luminal side.35 When similar studies were conducted in horses, there was no sidedness to the direction of PGF2α secretion for either pregnant or nonpregnant endometrium.36 The majority of evidence in mares suggests an overall reduction in PGF2α synthesis and secretion by pregnant endometrium, as discussed above, not a redirection in its secretion. Administration of estrogens can increase the interovulatory intervals in mares;37, 38 the mechanism is via suppression of follicular development, not via maintenance of luteal function.38 If anything, conceptus-secreted estrogens could potentially jeopardize the pregnancy by enhancing prostaglandin secretion. Incubation of endometrium from nonpregnant mares in the presence of estradiol significantly increased in vitro prostaglandin F2α production in a dose-dependent fashion.12 Furthermore, in vivo systemic administration of estrogen followed by an oxytocin challenge resulted in a fivefold increase in PGFM compared to the control vehicle.39 In an attempt to create a more physiological test, Goff and co-workers employed Silastic spheres (1 cm) impregnated with estradiol, with a release rate similar to in vivo estrogen production by conceptuses, and inserted the spheres into the uterus of mares. When mares were challenged with oxytocin as before, there was a fourfold increase in PGFM compared with mares containing control spheres.39 Thus not only is
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there little compelling evidence that conceptus estrogen is the maternal recognition of pregnancy signal in mares, but also the enhancement of prostaglandin secretion by estrogens in non-pregnant mares suggests that whatever signal the conceptus does employ is very potent for it must override conditions that are highly favorable for prostaglandin secretion.
Conceptus estrogen secretion Although studies to date do not support a theory for estrogens being the primary signal in mares for maternal recognition of pregnancy, they do not preclude an important role for the conceptus estrogens in the establishment of pregnancy in this species. In fact, quite the opposite is likely true. Estradiol and estrone concentrations within the uterine lumen were markedly elevated in pregnant mares from days 10 to 20.40 Similarly, the uterine content of uteroferrin as well as other uterine-specific proteins increases markedly during this same time,24, 26 and the temporal associations between elevated estrogens in the uterine lumen and secretion of uterine-specific proteins are likely fundamentally connected. McDowell et al.41 reported that administration of progesterone to ovariectomized mares resulted in uterine secretory profile of proteins similar to those observed in uterine flushings of non-pregnant mares during the peak of the luteal phase. However, administration of progesterone + estradiol increased uterine-specific proteins, but not total protein, compared to treatment with just progesterone, an indication that conceptus estrogens may enhance uterine protein production or otherwise alter uterine function in pregnant mares. Thus at least certain uterine-specific proteins, thought to be important in establishment and maintenance of pregnancy, are enhanced by the actions of conceptus estrogen secretion in concert with progesterone secretion by the CL. Therefore it seems that pregnancy establishment in the horse requires one or more conceptus factors to ensure luteal maintenance and continued progesterone secretion, which in turn drives the production of uterine secretions essential for conceptus development. The secretion of these uterine factors could then be further amplified by estrogens from the conceptus. Interestingly, the early equine conceptus has been shown to express the mRNAs for the nuclear estrogen receptor β (ERβ) and the membrane-bound form of the progesterone receptor (PR).42 Protein for ERβ was localized to the same cells that secrete estrogens, the extraembryonic trophectoderm. No protein product for the nuclear PR was detected, and no immunostaining was performed for the membranebound form of PR.
Uterine protein secretion One of the consequences of progesterone production by the CL is the secretion of uterine proteins. Quantitative and qualitative changes in the protein content within the uterine lumen have been described.26, 43 When examined by two-dimensional polyacrylamide gel electrophoresis, the pattern of proteins synthesized and released by endometrium from cyclic mares was altered with advancing days after ovulation and CL formation, and proteins produced by the endometrium of pregnant mares at days 12 and 14 after ovulation were similar to those of diestrous mares.24 Major endometrial proteins identified in diestrous and early pregnant mares included the iron-binding protein uteroferrin, retinol binding protein,44 and a low molecular weight protein (Mr ∼ 17 000–18 000) named ‘U1’.45 ‘U1’ was later determined to be a member of the lipocalin family and found to associate with the acelluar capsule that surrounds the early equine conceptus.46 Other progesterone-dependent uterine proteins include uteroglobin,47 uterine Mx protein,48 transforming growth factor β1,49 and epidermal growth factor.50 These uterine proteins are likely important for embryo and/or placental development, but their roles in maternal recognition of pregnancy, if any, have not been established.
Conceptus mobility Early studies demonstrated an apparent paradox in that the endometrium of pregnant mares was capable of producing PGF2α , yet uterine luminal PGF2α concentrations were reduced in pregnancy.7, 12, 18 This paradox was resolved, as mentioned previously, when it was shown that co-incubations of endometrium from pregnant or non-pregnant mares with conceptus membranes reduced the PGF2α produced.8, 11, 19 These observations led to the realization that the inhibitory effects of conceptus products on endometrial production of PGF2α were likely shortlived or transient. However, the fact that the early equine conceptus remains spherical during the time of maternal recognition of pregnancy left the question of how can the small spherical conceptus in one uterine horn alter PGF2α production in the opposite horn. When Ginther published the interesting observation, determined by ultrasonography, that equine conceptuses are highly mobile from the time of first detection with ultrasonography until days 16 to 18 when they become ‘fixated’ at the junction of one uterine horn and the uterine body,51 the extensive mobility seemed likely to be an important mechanism to ensure PGF2α inhibition throughout the entire uterine lumen. To test this hypothesis,
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Maternal Recognition of Pregnancy McDowell and Sharp restricted conceptus mobility by ligating uterine horns at various locations, and the events leading to maternal recognition of pregnancy were studied.52, 53 Results of these studies revealed that a distinct relationship existed between the amount of uterus to which the conceptus had access and the success of pregnancy recognition. When conceptuses had access to most of the uterus (the tip of the uterine horn contralateral to the side of ovulation was ligated, i.e., a surgical control), all mares recognized pregnancy as determined by CL maintenance (monitored by ultrasound observations and elevated progesterone concentrations), failure to return to estrus, continued conceptus viability, and positive tests for equine chorionic gonadotropin (eCG) at day 40. When only about 50% of the uterus was accessible to the conceptus (i.e., the uterine horn contralateral to the side of ovulation was ligated at the bifurcation), 50% of the mares were unable to recognize pregnancy as determined by loss of the CL, return to estrus, and eventual loss of conceptus integrity. When conceptus mobility was restricted to one uterine horn (i.e., the uterine horn ipsilateral to the side of ovulation was ligated at the bifurcation) only about 12% of mares were able to recognize the pregnancy. Furthermore, in a second study with a different group of mares, uteri were ligated so as to maximally restrict conceptus movement, but the synthetic progestin altrenogest (Regumate; Hoescth-Roussel) was administered to provide a source of progestin. In this latter group, four of five mares maintained viable conceptuses and a positive ECG test at day 40, indicating that the major cause of conceptus loss in the groups with maximal conceptus restriction was loss of progestin.53 Thus conceptus mobility throughout the uterus clearly plays an important role in altering PGF2α production at the time of maternal recognition of pregnancy in horses. The extensive mobility may also permit conceptuses access to uterine histotrophe throughout the uterus. Conceptuses of other domestic species undergo extensive elongation at the time of maternal recognition of pregnancy, which enables maximal interaction of the trophoblast and uterine endometrium. Mobility of the spherical equine conceptus likely plays a similar role.19, 53 The equine conceptus is mobile within the uterine lumen from at least day 9 after ovulation until approximately day 15, at which time its mobility is restricted near the bifurcation of one uterine horn.54, 55 Stout and Allen56 reported that PGF2α concentrations in uterine flushings of pregnant mares increased on days 18 and 20 of pregnancy, after embryo fixation, to levels similar to those found in non-pregnant mares on day 14 (approximately 8 pg/mL at day 20 in pregnant mares compared to approximately 4 pg/mL on day 14 in non-pregnant mares). They proposed that
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Figure 227.1 Prostaglandin F (PGF) concentrations (ng/mL, ± SEM) in uterine flushings collected from non-pregnant mares (solid line) and from pregnant mares (dashed line). Adapted from Zavy et al.18
the conceptus simply delays the capability of the endometrium to produce PGF2α , and that after day 18 a mechanism alternative to the conceptus suppressing endometrial PGF2α production exists in order to avoid luteolysis. Likewise, in 1981 Vernon et al.12 had reported that endometrial production of PGF was greater at days 18 and 20 post-ovulation, and endometrial content of PGF was greater at 20 days postovulation in pregnant mares than at days 14 or 16 in non-pregnant mares. However, in contrast to Stout and Allen, Zavy and Sharp reported PGF2α concentrations in uterine flushings to be lower on days 12 through 20 in pregnant compared to non-pregnant mares (5.1 ng/mL at day 20 in pregnant mares compared to 26 ng/mL at day 14 in non-pregnant mares; Figure 227.1).7, 18 Zavy and Sharp concluded that although the endometrium of pregnant mares is capable of producing PGF2α , its capacity to do so is greatly inhibited by the presence of the conceptus, at least through day 20 of pregnancy (the latest day tested).
Changes in gene expression Complex changes in gene expression must occur at the proper time and in the appropriate tissues for pregnancy to be successful. Therefore research aimed at defining the regulation of gene expression in the conceptus and uterus around the time of maternal recognition of pregnancy is of critical importance. Differences in gene expression between day-12 and day-15 embryos were studied by suppression subtractive hybridization.57, 58 Some of the genes identified included pregnancy-associated glycoprotein (PAG), alpha-fetoprotein (AFP), calcyclin, phospholipase A2 (PLA2 ), fumarylacetoacetase, cytokeratins 8 and 18, and isocitrate and succinate
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dehydrases. Between day 12 and day 15 PAG and calcyclin transcript abundance increased approximately 30-fold, and AFP expression increased at least 1000-fold. Calcyclin is a calcium-binding protein of the S-100 family, and PLA2 is an enzyme that cleaves phospholipids to release fatty acids and is involved in prostaglandin, thromboxane, and leukotriene synthesis. Multiple forms of PLA2 appeared to be differentially regulated in the day-12 versus day-15 conceptuses.57 PAGs are placental antigens of the aspartic proteinase gene family that were initially characterized as pregnancy markers in the maternal circulation of domestic ruminants. They have since been reported for a variety of eutherian mammals. It has been suggested that porcine PAG may be involved in a luteoprotective mechanism by binding with luteal and uterine gonadotropin receptors.59 Equine PAG mRNA is restricted to the chorion both prior to implantation and in the term placenta,60 but a decisive function for equine PAG has not been eluciated. Two other proteins identified as produced by early equine conceptuses or found in yolk-sac fluid include ubiquitin (8500 daltons) and insulin (3500 daltons), and insulin was tested as a candidate protein for the equine maternal recognition of pregnancy signal. However, daily administration of insulin to non-pregnant mares did not result in luteal maintenance.61 Changes in gene expression around the time of maternal recognition of pregnancy remain poorly understood. The above studies with embryos examined individual genes60 or a small handful (50) of clones sequenced from an expression library.57, 58 Although there are studies examining differences in individual uterine proteins between cyclic and pregnant mares, as discussed above, to date there are no studies using suppression subtraction methods or transcriptome comparisons of pregnant versus nonpregnant equine endometrium. The recent release of the equine genome sequence by the Broad Institute (http://www.broadinstitute.org/mammals/horse) and upcoming equine genomic and expression microarray assays will greatly expand the tools with which large-scale comparisons of changes in gene expression by the equine embryo and uterus over time can be performed, as well as simplify comparisons among species for both genomic sequences and expression assays.
Relationship between oxytocin and prostaglandin F2α The corpora lutea of sheep and cattle produce high concentrations of oxytocin, which are secreted into the uterine vein at the time of luteolysis.62–64 Luteal oxytocin stimulates release of uterine PGF2α , and a
positive feedback loop between oxytocin and PGF2α plays a critical role in luteal regression in these species.65–67 In cycling pigs circulating oxytocin (from the pituitary and/or from the uterus) stimulates endocrine-directed PGF secretion to promote luteolysis. In pregnant pigs conceptus estrogen stimulates a large increase in uterine oxytocin, and the oxytocin within the uterus serves to reorient PGF secretion from endocrine to exocrine32, 33 (M.A. Mirnado, personal communication). Oxytocin likely plays a role in PGF2α release and luteolysis in mares as well, although its role is less well defined in horses than in ruminants or swine. The CL of horses produce small amounts of oxytocin,68 but there is no evidence that luteal oxytocin plays a significant role in regulating reproductive tract function in horses. As with other species, in the horse oxytocin is produced in the hypothalamus and released into the circulation from the neural lobe of the pituitary gland. When mares were fitted with catheters in the intercavernous sinus to measure pituitary effluent of oxytocin, high-magnitude luteolytic pulses of PGFM were associated with highmagnitude pulses of oxytocin.69 The high-oxytocin pulses appeared to coincide with or follow the onset of the PGFM pulses, and the authors concluded that uterine PGF2α may stimulate secretion of pituitary oxytocin release. In 1997 Behrendt et al.70 reported oxytocin production by the equine endometrium, and cloned and sequenced the oxytocin-neurophysin 1 cDNA (OT-NP1) from an equine endometrial cDNA library.70, 71 OT-NP1 mRNA levels were higher at estrus than at any other stage of the cycle and were upregulated by exogenous estradiol. The presence of a conceptus decreases message levels, as OT-NP1 mRNA levels were lower on day 15 of pregnancy than on day 15 of the cycle.71, 72 Likewise, immunohistochemical staining of the oxytocin peptide was greater at estrus than at diestrus, but was greater at day 14 of pregnancy than at any other stage examined.73 The low message levels but elevated protein levels at the time of maternal recognition of pregnancy indicate that there is differential regulation of transcription and translation, or that the protein localized by immunohistochemical staining was synthesized at a different location and transported to the endometrium for uptake. In examining the relationship between PGF2α and oxytocin in mares, there are apparent dichotomies between in vivo and in vitro studies or between different in vivo studies, depending on the dose and/or route of oxytocin administration. Several studies reported that exogenous PGF2α causes oxytocin release and that exogenous oxytocin causes PGF2α release. For example, when PGF2α was given to non-pregnant mares on days 8 or 13 after ovulation, oxytocin
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Maternal Recognition of Pregnancy concentrations in the peripheral circulation were elevated within 1 min of treatment.74 When oxytocin was given to cyclic mares on days 11 and 13 after ovulation there was a marked increase in peripheral PGFM. However, that increase in PGFM was eliminated when the mares were pregnant.75 Similarly, in in vitro studies, oxytocin causes PGF2α release from endometrial explants, and the magnitude of that response varies with day of the cycle in which the endometrium was obtained. In response to an oxytocin challenge significantly greater amounts of PGF2α were released from endometrial explants obtained at day 15 of the estrous cycle than from those obtained at estrus or on days 11 or 13 of the cycle.76, 77 The above studies would lead one to believe that there is a relatively simple positive feedback system between oxytocin and PGF2α , which is disrupted by presence of a conceptus. However, this story is complicated by studies demonstrating that exogenous oxytocin is capable of causing luteal maintenance in non-pregnant mares. When non-pregnant mares were injected with oxytocin daily from day 9 to day 14,75 or twice daily from day 8 until day 14 after ovulation,78 progesterone concentrations remained elevated and the mares did not return to estrus at the expected time. In addition when oxytocin was administered subcutaneously by osmotic minipumps on days 8 through 20 after ovulation, luteolysis was prevented in four of five mares.79 These in vivo studies suggest that exogenous oxytocin given intermittently may stimulate uterine PGF2α production, but when given at higher doses and/or on a daily basis or continuously via minipumps it may cause luteal maintenance. Thus in cycling mares exogenous PGF2α stimulates oxytocin release and exogenous oxytocin may either stimulate PGF2α release or result in luteal maintenance. It is clear that more work is needed to understand fully the precise mechanisms controlling PGF2α –oxytocin interactions at the time of luteolysis or maternal recognition of pregnancy in the mare. Both oxytocin and PGF2α are produced by equine endometrium and likely interact in a selfpropagating or positive-feedback system at the time of luteolysis in non-pregnant mares. However, static in vitro cultures typically performed in Petri dishes make it impossible to separate the interactions of the two hormones. Therefore an in vitro perifusion system was developed in which endometrial explants were placed in filter holders, and culture medium was passed through the filter holders by using syringe pumps80 (KJ McDowell and MM Ward, unpublished observations). Effluent was collected in culture tubes with the aid of a fraction collector. An example of PGF2α release from endometrial explants in response to oxytocin challenges is shown in Fig-
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ure 227.2. Oxytocin administration caused a significant increase in PGF2α release from endometrium of non-pregnant mares, but the effect was greatly diminished in endometrium of pregnant mares in the presence of a conceptus, indicating once again that the oxytocin–prostaglandin feedback system has become uncoupled in pregnancy. Endometrial prostaglandin release in mares is likely mediated through uterine endometrial oxytocin synthesis/release and uterine oxytocin receptors. In non-pregnant mares endometrial oxytocin receptor concentrations are low during estrus and increase steadily through day 12, then there is a significant increase in receptor concentrations between days 12 and 14. In pregnant mares, however, that dramatic increase in receptor concentration does not occur.81, 82 Thus inhibition of upregulation of oxytocin receptors may be an important part of maternal recognition of pregnancy in pregnant mares, and it has been suggested that the equine conceptus may produce substances that alter the interaction of the oxytocin with its receptor.81 There is also evidence that mares produce a conceptus-induced, endometrial-produced inhibitor of PGF2α (an endometrial prostaglandin synthesis inhibitor, or EPSI)83 (D.C. Sharp, unpublished observations), which produces a potent blockade to the conversion of arachidonic acid to PGF2α . These studies provide additional evidence that the conceptusdirected alteration of endometrial prostaglandin production is a critical step in maternal recognition of pregnancy in the horse. Equine conceptuses secrete both PGF2α and PGE2 .83, 84 Allen85 hypothesized that a relatively high proportion of pregnancy losses in the mare that occur between 12 and 30 days after ovulation may be attributed to the secretion of too much PGF2α by the conceptus. Although it seems paradoxical that conceptuses would produce these prostaglandins and simultaneously inhibit PGF2α in the uterine endometrium, it is plausible that the conceptusproduced prostaglandins, along with conceptus estrogens, are involved in stimulating local peristaltic uterine contractions, which are necessary for embryo mobility. Prostaglandins are produced by a variety of physiological stimuli that activate phospholipase A2 , which in turn cleaves arachidonic acid from intracellular phospholipid stores. Free arachidonic acid is rapidly oxidized in at least two separate pathways – one of which is catalyzed by cycloogenase (COX) enzymes leading to production of prostaglandins and thromboxanes;, while the other is catalyzed by lipoxygenase leading to the production of 5-hydroperoxyeicosatetraenoic acid (5-HPETE) and leukotrienes.86 There are two isoforms of COX, COX-1 and COX-2. In cows and sheep,
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Hours Perfused Figure 227.2 Prostaglandin F2α (PGF2α ) (ng/mg tissue) from endometrial explants collected from mares at day 14 of the estrous cycle (top panel) or at day 14 of pregnancy (bottom panel). Tissues were placed in perifusion chambers, with fractions collected each hour for 24 hours. In the perfusion system for this pregnant endometrium, membranes from a day 14 conceptus were placed in a separate filter holder immediately up-stream from the endometrium. Oxytocin (10−10 M or 10−6 M) was delivered over a 30-minute period, beginning at either 6 hours or 11 hours. PGF2α release in response to 10−10 M oxytocin was greater than to 10−0 M oxytocin, and the response was greatly reduced with pregnancy. From MM Ward and KJ McDowell, 2001, MS Thesis. University of Kentucky, unpublished.
endometrial expression of COX-2 was transiently induced during late diestrus, but COX-1 expression did not change (sheep) or was undetectable (cows). 87, 88 In horses, when endometrial biopsies were obtained on days 10, 13, and 15 of the estrous cycle and on day 15 of pregnancy, COX-2 was significantly increased at day 15 of the cycle but not on day 15 of pregnancy, and induction of COX-2 expression was localized to the surface epithelial cells in cycling animals only.89 In the same study, microsomal PGE2 and PGF2α synthases, localized mainly to the surface epithelium, did not change over the estrous cycle and were unaffected by pregnancy. These data are supported by the report of Eroh and Ealy that real-time quantitative analysis of the equine COX-2 mRNA in endometrium of non-pregnant mares was approximately 28-fold higher than in the endometrium of pregnant mares on day 14.90 Thus it appears that COX-1 is constitutively expressed in the uterine endometrium of farm species, and it is COX-2 that is
upregulated or downregulated at the time of luteolysis or maternal recognition of pregnancy. Bruemmer puts foward an intriguing hypothesis that maternal recognition of pregnancy in mares is regulated by the direction in which oxytocin is secreted from endometrial cells. When secreted into the lumen (apically) it acts in an autocrine manner stimulating prostaglandin production. When secreted basally into the myometrium it causes contractions, moving the spherical embryo toward another portion of the uterus (J. Bruemmer, personal communication). It is clear that the equine endometrium synthesizes and secretes both PGF2α and oxytocin. At the time of luteolysis in non-pregnamt mares these hormones (and perhaps pituitary oxytocin) interact in a selfpropagating or positive-feedback manner whereby pulses of PGF2α increase in frequency and magnitude ultimately causing the demise of the corpus luteum. During pregnancy the presence of the conceptus
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Maternal Recognition of Pregnancy disrupts or uncouples the self-propagating system and the corpus luteum is maintained. As discussed above, current studies indicate that during pregnancy the conceptus may downregulate (or inhibit upregulation) of endometrial oxytocin receptors, that the conceptus may induce an endometrial inhibitor of PGF2α , and that the conceptus may alter the direction in which oxytocin is secreted by the endometrium. Because maternal recognition of pregnancy is such an important physiological event there is likely redundancy in the system. Certainly over the next several years it will be interesting to see how these pieces of the puzzle fit together.
Uterine blood flow As the foregoing indicates, maternal recognition of pregnancy implies a complex dialog between conceptus and maternal unit. An interesting line of investigation has revealed that part of that dialog involves conceptus-induced changes in endometrial blood flow. Silva and Ginther91,92 used color Doppler ultrasonography to monitor blood perfusion in the endometrium of both uterine horns daily, beginning on day 1 (day of ovulation = day 0) until after fixation of the conceptus vesicle (day 16). After day 12, when the mobile conceptus occupied one uterine horn for more than 7 minutes, there was a significant increase in objective vascularity scores of the endometrium on that side. After fixation, the uterine horn containing the conceptus maintained a significantly higher blood perfusion, perhaps the result of conceptusproduced estrogens and prostaglandins. The same investigators also reported that thickening of the dorsal endometrium precedes fixation and likely contributes to orientation of the conceptus such that the embryo proper becomes positioned at 6.00 relative to the richly vascular portion of the mesometrium. Indeed, disorientation of the conceptus has been associated with early embryonic loss.93 Encroachment of the dorsal endometrium raises some fascinating questions. Grossly it has the appearance of a decidual reaction, but little is known about the changes in cellular make-up that result in differential enlargement of the dorsal endometrium but not the ventral endometrium. Furthermore, there is no apparent enlargement of the endometrium on the contralateral side, suggesting once again that the enlargement and encroachment is the result of conceptus–maternal communication.
Conclusions The evidence overwhelmingly supports the idea that luteolysis in non-pregnant mares is caused by a self-
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propagating system, whereby oxytocin stimulates release of PGF2α , and likewise PGF2α stimulates release of oxytocin, until elevated concentrations of PGF2α cause functional and structural regression of the corpus luteum. The initial signal that stimulates the oxytocin–PGF2α cascade is not clear, but it is clear that a conceptus-secreted substance disrupts the luteolytic cascade of events, suppressing PGF2α synthesis and release, with the end result being luteal maintenance. This sequence of events strongly indicates that the process is one of luteostasis. Furthermore, the dominant role of the conceptus in managing events of pregnancy establishment in mares is evident throughout the process. We tend to think in terms of the dam when speaking of maternal recognition of pregnancy because it is her physiological processes that are altered; however, it might be more instructive to think in terms of the conceptus and its need for nourishment. Indeed, maintenance of the corpus luteum may be thought of as a conceptuscontrolled mechanism to assure continuing histotrophe. One can only marvel at the simple elegance of the process initiated and maintained by the spherical equine conceptus as it moves throughout the uterine lumen distributing its luteostatic signal(s), utilizing available histotrophe, and directing angiogenic events to assure a hemotropic source of nutrition when metabolic needs and placental development make histotrophic nutrition less efficacious.
Acknowledgements This is publication No. 08-14-142 of the University of Kentucky Agricultural Experiment Station and is published with approval of the director.
References 1. Short RV. When a conceptus fails to become a pregnancy. In: Wolstenholme GEW, O’Conner M (eds) Foetal Autonomy. Ciba Foundation Symposium. London: J. and A. Churchill, 1969; pp. 2–26. 2. Betteridge KJ, Eaglesome MD, Flood PF. Embryo transport through the mare’s oviduct depends upon cleavage and is independent of the ipsilateral corpus luteum. J Reprod Fertil Suppl 1979;27:387–94. 3. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 hastens oviductal transport in equine embryos. Biol Reprod 1991;45:544–6. 4. Loeb L. The effect of extirpation of the uterus on the life and function of the corpus luteum in the guinea pig. Proc Soc Exp Biol Med 1923;20:441–64. 5. Ginther OJ, First NL. Maintenance of the corpus luteum in hysterectomized mares. Am J Vet Res 1971;32:1687–91. 6. Hershman L, Douglas RH. The critical period for the maternal recognition of pregnancy in pony mares. J Reprod Fertil Suppl 1979;27:395–401.
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7. Sharp DC, Zavy MT, Vernon MW, Bazer FW, Thatcher WW, Berglund LA. The role of prostaglandins in the maternal recognition of pregnancy in mares. Anim Reprod Sci 1984;7:269–82. 8. Sharp DC, McDowell KJ. Critical events surrounding the maternal recognition of pregnancy in mares. E Vet J Suppl 1985;3:19–22. 9. Douglas RH, Ginther OJ. Concentration of prostaglandins F in uterine venous plasma of anesthetized mares during the estrous cycle and early pregnancy. Prostaglandins 1976;11:251–260. 10. Zavy MT, Bazer FW, Sharp DC, Frank M, Thatcher WW. Uterine luminal prostaglandin F in cycling mares. Prostaglandins 1978;16:643–50. 11. Berglund LA, Sharp DC, Vernon MW, Thatcher WW. Effect of pregnancy and collection technique on prostaglandin F in the uterine lumen of pony mares. J Reprod Fertil Suppl 1982;32:335–41. 12. Vernon MW, Zavy MT, Asquith RL, Sharp DC. Prostaglandin F2alpha in the equine endometrium: steroid modulation and production capacities during the estrous cycle and early pregnancy. Biol Reprod 1981;25:581–9. 13. Neely DP, Kindahl H, Stabenfeldt GH, Edqvist LE, Hughes JP. Prostaglandin release patterns in the mare: physiological, patho-physiological and therapeutic responses. J Reprod Fertil Suppl 1979;27:181–9. 14. Kindahl H, Knudsen O, Madej A, Edqvist LE. Progesterone, prostaglandin F2a, PMSG and oestrone sulphate during early pregnancy in the mare. J Reprod Fertil Suppl 1982;32:353–9. 15. Kindahl H, Basu S, Fredriksson G, Goff AK, Kunavongkrit A, Edqvist LE. Levels of prostaglandin F2a metabolites in blood and urine during pregnancy. Anim Reprod Sci 1984;7:133–47. 16. Vernon MW, Strauss S, Simonelli M, Zavy MT, Sharp DC. Specific PGF-2 alpha binding by the corpus luteum of the pregnant and non-pregnant mare. J Reprod Fertil Suppl 1979;27:421–9. 17. Kooistra LH, Ginther OJ. Termination of pseudopregnancy by administration of prostaglandin F2a and termination of early pregnancy by administration of prostaglandin F2a or colchicine or by removal of embryo in mares. Am J Vet Res 1976;37:35–9. 18. Zavy MT, Vernon MW, Asquith RL, Bazer FW, Sharp DC. Effect of exogenous gonadal steroids and pregnancy on uterine luminal prostaglandin F in mares. Prostaglandins 1984;27:311–20. 19. Sharp DC, McDowell KJ, Weithenauer J, Thatcher WW. The continuum of events leading to maternal recognition of pregnancy in mares. J Reprod Fertil Suppl 1989;37:101–7. 20. Godkin JD, Bazer FW, Moffatt J, Sessions F, Roberts RM. Purification and properties of a major, low molecular weight protein released by the trophoblast of sheep blastocysts at day 13–21. J Reprod Fertil 1982;65:141–50. 21. Helmer SD, Hansen PJ, Anthony RV, Thatcher WW, Bazer FW, Roberts RM. Identification of bovine trophoblast protein-1, a secretory protein immunologically related to ovine trophoblast protein-1. J Reprod Fertil 1987;79: 83–91. 22. Roberts RM. Conceptus interferons and maternal recognition of pregnancy. Biol Reprod 1989;40:449–52. 23. Farin C E, Imakawa K, Hansen T R, McDonnell JJ, Murpy CN, Farin PW, Roberts RM. Expression of trophoblastic interferon genes in sheep and cattle. Biol Reprod 1990;43:210–18.
24. McDowell KJ, Sharp DC, Fazleabas AT, Roberts RM. Twodimensional polyacrylamide gel electrophoresis of proteins synthesized and released by conceptuses and endometria from pony mares. J Reprod Fertil 1990;89:107–15. 25. Baker CB, Adams MH, McDowell KJ. Lack of expression of alpha or omega interferons by the horse conceptus. J Reprod Fertil Suppl 1991;44:439–43. 26. Zavy MT, Mayer R, Vernon MW, Bazer FW, Sharp DC. An investigation of the uterine luminal environment of nonpregnant and pregnant pony mares. J Reprod Fertil Suppl 1979;27:403–11. 27. Flood PF, Betteridge KJ, Irvine DS. Oestrogens and androgens in blastocoelic fluid and cultures of cells from equine conceptuses of 10–22 days gestation. J Reprod Fertil Suppl 1979;27:413–20. 28. Heap RB, Hamon M, Allen WR. Studies on oestrogen synthesis by the preimplantation equine conceptus. J Reprod Fertil Suppl 1982;32:343–5. 29. Perry JS, Heap RB, Amoroso EC. Steroid hormone production by pig blastocysts. Nature 1973;245:45–7. 30. Bazer FW, Thatcher WW. Theory of maternal recognition of pregnancy in swine based on estrogen controlled endocrine versus exocrine secretion of prostaglandin F2alpha by the uterine endometrium. Prostaglandins 1977;14:397–400. 31. Bazer FW, Vallet JL, Harney JP, Gross TS, Thatcher WW. Comparative aspects of maternal recognition of pregnancy between sheep and pigs. J Reprod Fertil Suppl 1989;37:85–9. 32. Sample GL, Blackwell DM, Kubotsu SL, Mirando MA. Endocrine secretion of prostaglandin F2[alpha] in cyclic gilts is decreased by intrauterine administration of exogenous oxytocin. Anim Reprod Sci 2004;84:395–406. 33. Mirando MA, Hu J. Roles for oxytocin in luteolysis and pregnancy recognition in pigs. In: Proceedings of a Workshop on Maternal Recognition of Pregnancy and Early Embryonic Loss in the Mare, Havemeyer Foundation Monograph Series 16. Newmarket, UK: R&W Communications, 2005; pp 25–7. 34. Lacroix MC, Kann G. Discriminating analysis of “in vitro” prostaglandin release by myometrial and luminal sides of the ewe endometrium. Prostaglandins 1983;25:853–69. 35. Gross TS, Lacroix MC, Bazer FW, Thatcher WW, Harney JP. Prostaglandin secretion by perifused porcine endometrium: further evidence for an endocrine versus exocrine secretion of prostaglandins. Prostaglandins 1988;35:327–341. 36. Franklin KJ, Gross TS, Dubois DH, Sharp DC. In vitro prostaglandin secretion from luminal and myometrial sides of endometrium from cyclic and pregnant mares at day 14 post-estrus. Biol Reprod 1989;40(Suppl. 1):114. 37. Berg SL, Ginther OJ. Effect of estrogens on uterine tone and life span of the corpus luteum in mares. J Anim Sci 1978;47:203–8. 38. Woodley SL, Burns PD, Douglas RH, Oxender WD. Prolonged interovulatory interval after oestradiol treatment in mares. J Reprod Fertil Suppl 1979;27:205–9. 39. Goff AK, Sirois J, Pontbriand D. Effect of oestradiol on oxytocin-stimulated prostaglandin F2a release in mares. J Reprod Fertil 1993;98:107–12. 40. Zavy MT, Vernon MW, Sharp DC III, Bazer FW. Endocrine aspects of early pregnancy in pony mares: a comparison of uterine luminal and peripheral plasma levels of steroids during the estrous cycle and early pregnancy. Endocrinology 1984;115:214–19. 41. McDowell KJ, Sharp DC, Grubaugh W. Comparison of progesterone and progesterone + oestrogen on total and
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42.
43.
44.
45.
46.
47.
48.
49.
50.
51. 52.
53.
54.
55. 56.
57.
58.
specific uterine proteins in pony mares. J Reprod Fertil Suppl 1987;35:335–42. Rambags BPB, van Tol HTA, van den Eng MM, Colenbrander B, Stout TAE. Expression of progesterone and oestrogen receptors by early intrauterine equine conceptuses. Theriogenology 2008;69:366–75. Zavy MT, Bazer FW, Sharp DC, Wilcox CJ. Uterine luminal proteins in the cycling mare. Biol Reprod 1979;20: 689–98. McDowell KJ, Adams MH, Franklin KM, Baker CB. Changes in equine endometrial retinol-binding protein RNA during the estrous cycle and early pregnancy and with exogenous steroids. Biol Reprod 1995;52:438–43. Zavy MT, Sharp DC, Bazer FW, Fazleabas A, Sessions F, Roberts RM. Identification of stage-specific and hormonally induced polypeptides in the uterine protein secretions of the mare during the oestrous cycle and pregnancy. J Reprod Fertil 1982;64:199–207. Crossett B, Allen WR, Stewart F. A 19 kDA protein secreted by the endometrium of the mare is a novel member of the lipocalin family. Biochem J 1996;320:137–43 [abstr.]. Muller-Schottle F, Bogusz A, Grotzinger J, Herrler A, Krusche CA, Beier-Hellwig K, Beier HM. Full-length complementary DNA and the derived amino acid sequence of horse uteroglobin. Biol Reprod 2002;66:1723–8. Hicks BA, Etter SJ, Carnahan KG, Joyce MM, Assiri AA, Carling SJ, Kodali K, Johnson GA, Hansen TR, Mirando MA, Woods GL, Vanderwall DK, Ott TL. Expression of the uterine Mx protein in cyclic and pregnant cows, gilts, and mares. J Anim Sci 2003;81:1552–61. Lennard SN, Stewart F, Allen WR. Transforming growth factor beta 1 expression in the endometrium of the mare during placentation. Mol Reprod Dev 1995;42:131–40. Stewart F, Power CA, Lennard SN, Allen WR, Amet L, Edwards RM. Identification of the horse epidermal growth factor (EGF) coding sequence and its use in monitoring EGF gene expression in the endometrium of the pregnant mare. J Mol Endocrinol 1994;12:341–50. Ginther OJ. Mobility of the early equine conceptus. Theriogenology 1983;19:603–11. McDowell KJ, Sharp DC, Peck LS, Cheves LL. Effect of restricted conceptus mobility on maternal recognition of pregnancy in mares. E Vet . Suppl 1985;3:23–4. McDowell KJ, Sharp DC, Grubaugh W, Thatcher WW, Wilcox CJ. Restricted conceptus mobility results in failure of pregnancy maintenance in mares. Biol Reprod 1988;39:340–8. Leith GS, Ginther OJ. Characterization of intrauterine mobility of the early equine conceptus. Theriogenology 1984;22:401–8. Leith GS, Ginther OJ. Mobility of the conceptus and uterine contractions in the mare. Theriogenology 1985;24:701–11. Stout TA, Allen WR. Prostaglandin E(2) and F(2 alpha) production by equine conceptuses and concentrations in conceptus fluids and uterine flushings recovered from early pregnant and dioestrous mares. Reproduction 2002;123:261–8. Simpson KS, Adams MH, Behrendt-Adam CY, Baker CB, McDowell KJ. Identification and initial characterization of calcyclin and phospholipase A2 in equine conceptuses. Mol Reprod Dev 1999;53:179–87. Simpson KS, Adams MH, Behrendt-Adam CY, Baker CB, McDowell KJ. Differential gene expression in day 12 and day 15 equine conceptuses. J Reprod Fertil Suppl 2000;56:539–47.
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59. Panasiewicz G, Majewska M, Romanowska A, Dajnowiec J, Szafranska B. Radiocompetition of secretory pregnancyassociated glycoproteins as chorionic ligands with luteal and uterine gonadotrophin receptors of pregnant pigs. Anim Reprod Sci 2007;99:285–98. 60. Green JA, Xie S, Szafranska B, Gan X, Newman AG, McDowell K, Roberts RM. Identification of a new aspartic proteinase expressed by the outer chorionic cell layer of the equine placenta. Biol Reprod 1999;60:1069–77. 61. Stout TAE, Rambags PB, van Tol HTA, Colenbrander B. Proceedings of a Workshop on Maternal Recognition of Pregnancy in the Mare III. In: Havemeyer Foundation Monograph. Series No. 16, Stout, TAE, Wade, JF (eds). pp. 50–52. 62. Wathes DC, Swann RW. Is oxytocin an ovarian hormone ? Nature 1982;297:225–7. 63. Wathes DC, Swann RW, Birkett SD, Porter DG, Pickering BT. Characterization of oxytocin, vasopressin, and neurophysin from the bovine corpus luteum. Endocrinology 1983;113:693–8. 64. Ivell R, Richter D. The gene for the hypothalamic peptide hormone oxytocin is highly expressed in the bovine corpus luteum: biosynthesis, structure and sequence analysis. EMBO J 1984;3:2351–4. 65. Flint APF, Sheldrick EL. Evidence for a systemic role for ovarian oxytocin in luteal regression in sheep. J Reprod Fertil 1983;67:215–25. 66. Lafrance M, Goff AK. Effect of pregnancy on oxytocininduced release of prostaglandin F2 alpha in heifers. Biol Reprod 1985;33:1113–19. 67. Silvia WJ, Lewis GS, McCracken JA, Thatcher WW, Wilson L. Hormonal regulation of uterine secretion of prostaglandin F2a during luteolysis in ruminants. Biol Reprod 1991;45:655–63. 68. Stevenson KR, Parkinson TJ, Wathes DC. Measurement of oxytocin concentrations in plasma and ovarian extracts during the estrous cycle of mares. J Reprod Fertil 1991;93:437–41. 69. Vanderwall DK, Silvia WJ, Fitzgerald BP. Concentrations of oxytocin in the intercavernous sinus of mares during luteolysis: temporal relationship with concentrations of 13,14-dihydro-15-keto-prostaglandin F2a. J Reprod Fertil 1998;112:337–46. 70. Behrendt CY, Adams MH, Daniel KS, McDowell KJ. Oxytocin expression by equine endometrium. Biol Reprod 1997;56(Suppl. 1):134. 71. Behrendt-Adam CY, Adams MH, Simpson KS, McDowell KJ. Oxytocin-neurophysin I mRNA abundance in equine uterine endometrium. Domest Anim Endocrinol 1999;16:183–92. 72. Behrendt-Adam CY, Adams MH, Simpson KS, McDowell KJ. The effects of steroids on endometrial oxytocin mRNA production. J Reprod Fertil Suppl 2000;56:297– 304. 73. Watson ED, Bjorksten T, Buckingham J, Nikolakopoulos E. Immunolocalisation of oxytocin and neurophysin in the mare uterus. J Reprod Fertil Suppl 2000;56:289–96. 74. Utt MD, Acosta TJ, Wiltbank MC, Ginther OJ. Acute effects of prostaglandin F2[alpha] on systemic oxytocin and progesterone concentrations during the mid- or late-luteal phase in mares. Anim Reprod Sci 2007;97:63–73. 75. Goff AK, Pontbriand D, Sirois J. Oxytocin stimulation of plasma 15-keto-13,14-dihydro prostaglandin F-2a during the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1987;35:253–60.
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76. Smith KM. The relationship between oxytocin and prostaglandin F2a at the expected time of luteolysis in mares. MS thesis, University of Kentucky, Lexington, 2000. 77. Smith KM, Behrendt-Adam CY, Ward MM, Adams MH, McDowell KJ. Oxytocin causes prostaglandin F2a release from equine endometria in a day-, dose-, and time-dependent manner. Biol Reprod 2000;62(Suppl 1): 344. 78. Vanderwall DK, Rasmussen DM, Woods GL. Effect of repeated administration of oxytocin during diestrus on duration of function of corpora lutea in mares. J Am Vet Med Assoc 2007;231:1864–7. 79. Stout TAE, Lamming GE, Allen WR. Oxytocin administration prolongs luteal function in cyclic mares. J Reprod Fertil 1999;116:315–20. 80. Ward MM. Development of a perifusion culture system to examine oxytocin and prostaglandin F2alpha interaction in the mare. MS thesis, University of Kentucky, 2001. 81. Sharp DC, Thatcher M-J, Salute ME, Fuchs A-R. Relationship between endometrial oxytocin receptors and oxytocininduced prostaglandin F2a release during the oestrous cycle and early pregnancy in pony mares. J Reprod Fertil 1997;109:137–44. 82. Starbuck GR, Stout TAE, Lamming GE, Allen WR, Flint APF. Endometrial oxytocin receptor and uterine prostaglandin secretion in mares during the oestrous cycle and early pregnancy. J Reprod Fertil 1998;113:173–9. 83. Watson ED, Sertich PL. Prostaglandin production by horse embryos and the effect of co-culture with endometrium from pregnant mares. J Reprod Fertil 1989;87: 331–6. 84. Allen WR. Fetomaternal interactions and influences during equine pregnancy. Reproduction 2001;121:513–27.
85. Allen WR. The physiology of early pregnancy in the mare. In: Proceedings of the Annual Convention of the AAEP 2000, 46:338–54. 86. Arici A, Behrman HR, Keefe DL. Prostaglandins and prostaglandin-like products in reproduction: eicosanoids, peroxides, and oxygen radicals. In: Yen SS, Jaffe RB, Barbieri RL (eds) Reproductive Endocrinology. Philadelphia: W.B. Saunders, 1999; pp. 134–52. 87. Charpigny G, Reinoud P, Tamby J-P, Creminon C, Martal J, Maclouf J, Guillomot M. Expression of cyclooxygenase-1 and -2 in ovine endometrium during the estrous cycle and early pregnancy. Endocrinology 1997;138:2163–71. 88. Arosh JA, Parent J, Chapdelaine P, Sirois J, Fortier MA. Expression of cyclooxygenases 1 and 2 and prostaglandin E synthase in bovine endometrial tissue during the estrous cycle. Biol Reprod 2002;67:161–9. 89. Boerboom D, Brown KA, Vaillancourt D, Poitras P, Goff AK, Watanabe K, Dore M, Sirois J. Expression of key prostaglandin synthases in equine endometrium during late estrus and early pregnancy. Biol Reprod 2004;70: 391–9. 90. Eroh ML, Ealy ADS. Pregnancy status and steroid exposure impact the abundance of endometrial cyclooxygenase2 mRNA in mares. Biol Reprod 2007;77(Suppl. 1):113. 91. Silva LA, Ginther OJ. An early endometrial vascular indicator of completed orientation of the embryo and the role of dorsal endometrial encroachment in mares. Biol Reprod 2006;74:337–43. 92. Silva, L. A., E. L. Gastal, M. A. Beg, and O. J. Ginther. 2005. Changes in vascular perfusion of the endometrium in association with changes in location of the embryonic vesicle in mares. Biol. Reprod. 72:755–761. 93. Ginther OJ. Dynamic physical interactions between the equine embryo and uterus. E Vet J Suppl 1985;3:41–7.
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Chapter 228
Sex Determination and Differentiation Bruce W. Christensen and Vicki N. Meyers-Wallen
Normal sexual development Since Jost’s classical study in 1973, mammalian sexual development traditionally has been described as occurring in three chronological steps: establishment of chromosomal sex, gonadal sex, and phenotypic sex.1 Each step was described as being dependent on the previous one, i.e., chromosomal sex determines which gonad develops and the gonads determine the sexual phenotype. More recent studies indicate greater complexity in mammalian sexual development. Note that standard genetic nomenclature2 (http://www.genenames.org) is used throughout this chapter, as follows: Human gene symbols are in italics and capitalized (SOX9). Gene symbols for other vertebrates, such as the horse, are in italics and only the first letter is capitalized (Sox9). Protein symbols for vertebrates and humans are in plain text and capitalized (SOX9). Establishment of chromosomal sex At the end of meiosis each gamete normally contains only one sex chromosome, with oocytes bearing an X chromosome and sperm bearing either an X or a Y chromosome. Consequently, after fertilization a zygote has either an XX or an XY chromosomal constitution and normally develops as either a female or a male, respectively (see Figure 228.1). Early studies of abnormal mammals established that the presence of the Y chromosome was associated with testis development and a male sexual phenotype, irrespective of the number of X chromosomes present. This indicated that a gene on the Y chromosome was important for testis induction, leading to the search for a Y-linked testis determination factor. Mapping of the Y chromosome identified SRY (sex-determining region Y), a gene expressed in the XY gonad that
normally initiates the molecular pathway for testis formation in mammals (reviewed in DiNapoli and Capel, 2008).3 No counterpart to SRY has yet been identified as necessary for ovarian development. However, an XX chromosome constitution in oocytes is necessary for their survival in most mammals, which in turn is essential for the maintenance of normal follicles and later normal reproductive cyclicity.4 Errors in the number or structure of the sex chromosomes can therefore lead to abnormal male or female development. The sexually indifferent stage Between fertilization and development of the testis or ovary, XX and XY zygotes develop similarly and are sexually indifferent in that the reproductive system has not developed sufficiently to distinguish male from female by morphology alone. Embryogenesis of the mammalian urogenital tract during this period has been reviewed elsewhere.5 Briefly, the early embryonic reproductive tract is derived from endoderm, mesoderm, and ectoderm. The paired urogenital ridges (UGR) arise from mesoderm on the dorsal body wall, coursing parallel to the developing vertebral cord. The UGR contains the mesonephros and the wolffian (mesonephric) ducts and bipotential gonads, which arise lateral and medial to the mesonephros, respectively. The UGR coelomic epithelium lateral to the wolffian ducts invaginates to form the mullerian (paramesonephric) ducts. These arise in a cranial to caudal direction, coursing laterally then medially over the wolffian ducts and fusing caudally with the cranial aspect of the urogenital sinus. The latter originates from endoderm, arising ventral to the intestinal tract, and terminating caudally beneath the rectum (see Figure 228.2). Near the end of the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Pregnancy, Parturition and the Puerperal Period CHROMOSOMAL SEX:
XY
XX
?
INDIFFERENT EMBRYO
Sry
GONADAL SEX: OVARY
GONAD
TESTIS MIS
PHENOTYPIC SEX: Oviduct Uterus Carnial vagina
REGRESS
REMAINS OPEN: Caudal vagina Vestibule
Clitoris
MULLERIAN DUCTS
Regress TESTOSTERONE
WOLFFIAN DUCTS
Vas deferens Epididymis
UROGENITAL SINUS
DHT
GENITAL TUBERCLE
DHT
CLOSES: Urethra Prostate
Penis Insl3
REMAINS OPEN: Vulva
GENITAL SWELLINGS
DHT
CLOSES: Scrotum
Testis Descent
Figure 228.1 Overview of normal mammalian sexual development from fertilization to sexual differentiation. (From Meyers-Wallen 2008;37 used with permission.)
sexually indifferent stage, the foundations for both male and female duct systems are present and poised to receive signals that will determine their course of differentiation. Meanwhile the paired bipotential gonads have arisen medial to the mesonephros. While XX and XY gonads appear morphologically similar at this stage, the competition of gonadal sex determination, waged through gene transcriptional activation and suppression, has already begun (below). Morphological evidence of testis differentiation, such as the development of testis cords (below), marks the end of the sexually indifferent stage. The ovary retains sexually indifferent morphology longer than the testis, with development of the cortical sex cords being the first evidence of ovarian morphology. Gonadal sex determination and differentiation Early and recent experiments confirm that the ovary is not necessary for the development of a female phenotype in the fetus, but development of a functional testis is essential to normal male phenotypic development: When the bipotential gonads regress or are
removed from embryos at the sexually indifferent stage, both XX and XY animals develop a female phenotype. This indicates that the basic mammalian phenotypic plan is to develop as a female, but the plan is changed through signals dependent upon testis development. Because the presence or absence of a normal, functional testis is pivotal in determining sexual phenotype, the term sex determination in normal mammalian development has been equated with gonadal sex determination, the process by which the gonad becomes committed to ovarian or testis differentiation. In contrast, the term sex differentiation is the process whereby the embryonic reproductive tract develops into specific organs, such as the bipotential gonad differentiating into testis or ovary.
Cellular components of the differentiating testis and ovary The bipotential gonad contains three cell types that contribute to the testis and ovary: somatic cell precursors, primordial germ cells, and steroidogenic precursor cells. Somatic cell precursors, which can become either Sertoli or granulosa cells, arise from surface epithelial cells, control further differentiation
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Figure 228.2 Summary of mammalian reproductive tract differentiation. Note that both the wolffian and mullerian ducts are present at the indifferent gonad stage. (Reprinted with permission from Current Conceptions, Inc. Pathways to Pregnancy and Parturition, 2nd edition, by P. L. Senger).
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of the gonad, and support the germ cells. Near the end of the sexually indifferent stage, somatic cell precursors grow into the mesenchyme underlying the gonadal surface to form the sex cords. In males the sex cords penetrate the medulla to form the testis cords, precursors of seminiferous tubules. Conversely in females, the sex cords remain near the cortex to form cortical sex cords, precursors of ovarian follicles. The primordial germ cells migrate from the yolk sac via the hindgut mesentery into the bipotential gonad and become surrounded by somatic cell precursors that form seminiferous tubules or follicles. Mesenchymal cells in the interstitium develop into steroidogenic precursors that become either Leydig cells or thecal cells. The molecular steps controlling these cellular and organizational changes are the subject of intense investigation.
Molecular steps in the testis pathway It is now well accepted that the Sry gene on the Y chromosome encodes the testis-determining factor.6, 7 During the sexually indifferent stage, SRY protein is expressed in XY gonadal somatic cell precursors.8 While Sox9 is expressed in indifferent XX and XY gonads, it quickly ceases in XX gonads but is rapidly upregulated in XY gonads. It is likely that Sry expression indirectly upregulates gonadal Sox9 expression, which in turn induces the somatic cell precursors to differentiate as Sertoli cells;9, 10 see also these reviews.3, 8, 11, 12 Sertoli cell differentiation is the decisive cellular event defining testis differentiation, as it controls further cellular differentiation and organization. For example, Sertoli cells secrete the paracrine factor prostaglandin D2 (PGD2 ), which induces uncommitted somatic cell precursors to express Sox9 and become Sertoli cells.13, 14 Another important signaling molecule is Fgf9,15, 16 which acts in autocrine and paracrine fashion to promote Sox9 expression and induce cell proliferation and testis cord formation.3 When Fgf9 is knocked out in an XY individual, Sox9 expression is not maintained and Sertoli cells and testis cords do not develop; instead the gonad proceeds to follow the female developmental pathway.15, 16 Fgf9 expression acts in both an autocrine and paracrine fashion to maintain expression of Sox9 and recruit the differentiation of neighboring cells into Sertoli cells. Additionally, the upregulation of these genes blocks the Wnt4 pathway and ovarian development16 (see Figure 228.3).
Molecular steps in the ovarian pathway Evidence is mounting that the development of the ovary is not a passive process, as was once thought (see current review in Yao 2005).17 Transcriptional
balanced opposing signals Fgf9
Wnt4
Sry
Sox9
Wnt4
Fgf9
Testis
Wnt4
Sox9 Fgf9
Ovary
Figure 228.3 Model of balanced opposing signals between Fgf9 and Wnt4. In XY gonads, Sry upregulates Sox9 to establish a feed-forward loop that upregulates Fgf9 and silences Wnt4. In XX gonads, Wnt4 dominates and silences Fgf9 and Sox9. (From Kim and Capel 2006;35 used with permission from John Wiley & Sons, Inc.)
changes indicate that an ovarian pathway is active at the same time as the testis pathway during the sexually indifferent stage, before differentiation of cortical sex cords is evident. One hypothesis that attempts to explain the role of an active ovarian pathway is that an undiscovered gene Z suppresses Sox9 transcription during normal ovarian development.18.19 The Z gene itself would be subject to transcriptional suppression by Sry. While one gene that fits the description of Z has not been identified, there are examples of several competitive signals between the ovary and testis pathways at other steps. The final outcome of gonadal sex determination depends on an accumulation of signals that shifts the balance in favor of one pathway (see Figure 228.3). It is becoming increasingly evident that intracellular beta-catenin signaling is a critical step in the ovary determination pathway.20 While beta-catenin is a well-known downstream effector of the canonical Wnt pathway in other developing organs, in the ovary it is likely to be an effector of Rspo1 and Wnt4.21–24 Expression of these two genes promotes ovarian differentiation and blocks testis differentiation. Although Wnt4 is expressed in both XY and XX indifferent gonads, it quickly ceases in XY gonads but is rapidly upregulated in XX gonads at the end of the indifferent stage (see Figure 228.3). Wnt4 expression leads to beta-catenin signaling and expression of follistatin (Fst) and Bmp2, molecular markers of early ovarian differentiation. Mutations in Wnt4 cause similar effects on ovarian development: loss of Fst expression, failure of meiosis with loss of oogonia, and accumulation of ectopic adrenocortical cells that secrete androgens.25–28 Rspo1 is expressed in both XX and XY indifferent gonads, but by the end of this period is dramatically upregulated in XX gonads. This leads to beta-catenin signaling, either through Wnt4
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Sex Determination and Differentiation or a separate pathway, as recent studies indicate that Rspo1 acts upstream of Wnt4 (reviewed in Tevosian and Manuylov 2008).29 After beta-catenin accumulates in the cytoplasm and is stabilized, it enters the nucleus and activates gene transcription. The opportunity for competition between the ovarian and testis pathways probably occurs in the cytoplasm, as beta-catenin and SOX9 proteins can bind and inactivate each other.30, 31 In beta-catenin excess, stabilized beta-catenin enters the nucleus. Conversely in SOX9 excess, SOX9 protein enters the nucleus. Other mechanisms of beta-catenin and SOX9 interaction are also being investigated (reviewed in Tevosian and Manuylov 2008).29 Mice with Rspo1 deletions have delayed Wnt4 expression in the XX gonad, and exhibit abnormalities identified in Wnt4 null mice, but additionally they develop seminiferous tubules.23 In humans, RSPO1 deletions cause development of bilateral testes in XX individuals, indicating the importance of RSPO1 in blocking the testis pathway.32 Other genes that participate in ovarian differentiation include Pisrt1 and Foxl2.33–35 As research progresses, additional genes are likely to be found that will increase our understanding of the ovarian pathway. In summary, these complicated and emerging interactions between the testis and ovarian pathways provide a more complex explanation as to how the bipotential gonad becomes either testis or ovary. Currently the most likely scenario is competition between two opposing pathways (see Figure 228.3). If Sry is activated, the balance tips in favor of the male pathway, as this favors dramatic Sox9 expression that blocks the female pathway and leads to Sertoli cell differentiation. If Sry is not activated, the balance tips in favor of the female pathway, as this favors betacatenin stabilization that blocks the male pathway and leads to granulosa cell differentiation. Phenotypic sexual development The presence or absence of testis secretions influences the differentiation of internal and external genitalia. Sertoli cells produce anti-mullerian hormone (AMH), also known as mullerian inhibiting substance (MIS), which causes regression of the mullerian ducts through interaction with receptors in the target organs (see Figure 228.1). Leydig cells produce testosterone, which binds to androgen receptors in the target organs, prevents programmed cell death of the wolffian ducts, and stimulates their further differentiation. Wolffian ducts adjacent to the testis elongate and fold to form the epididymides. The middle portion of the wolffian ducts remains relatively straight and becomes the vasa deferentia, while the distal portion dilates to form the seminal vesicles.
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Another androgen, dihydrotestosterone (DHT), is derived from testosterone through the action of the 5α–reductase type 2, an enzyme expressed in DHT target tissues. Through binding with the androgen receptor, DHT induces the urogenital sinus to differentiate into the prostate and male urethra, the labioscrotal folds to fuse to form the scrotum, and the genital tubercle to form the prepuce and penis. Leydig cells also secrete insulin-like peptide 3 (INSL3), which binds its receptors in the gubernaculum, and together with testosterone and androgen receptors, controls testis descent into the scrotum (reviewed in Amann and Veeramachaneni 2007).36 In the absence of testicular secretions the target organs follow the female pathway. The wolffian ducts regress and the mullerian ducts differentiate into the oviducts, uterus, cervix, and cranial vagina. The urogenital sinus forms the caudal vagina and vestibule. The genital tubercle becomes the clitoris, and the labioscrotal folds remain open, forming the labia. The ovaries remain in an abdominal position.
Abnormalities of sexual development in the horse Terminology In the initial diagnostic process it remains useful to classify abnormalities of sexual development by the first step at which development differs from normal, e.g., at chromosomal, gonadal, or phenotypic sex. Realizing that the normal process at two of these steps can be disrupted by gene mutations, it is expected that many definitive diagnoses in future will be made at the molecular level. The terminology used to describe these patients is evolving, attempting to incorporate molecular discoveries into diagnostic algorithms containing older terminology that was based upon physical findings and histology. For example, intersex describes an animal with a sexually ambiguous phenotype, which does not indicate the cause, and is not a definitive diagnosis (reviewed in MeyersWallen, 2008).37 Similarly, the term true hermaphrodite only indicates that an animal has both testicular and ovarian tissue in one or both gonads, but is not a definitive diagnosis because there are several mechanisms by which this can occur. Finally, the term pseudohermaphrodite (male or female) indicates that the chromosomal and gonadal sex of the affected individual are in agreement but that the internal and/or external genitalia are ambiguous. Therefore this term only indicates that there is an error in phenotypic sex. Examples of these are discussed in context below in reviewing the equine literature. In consideration of the potential stigma and psychosocial consequences
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Monosomy X (XO syndrome)
(a)
(b)
(c) Figure 228.4 Perineal areas of three intersex equids showing similar phenotypes but each with a very different pathogenesis. Each has a hypoplastic penis within a vulva and increased anogenital distance; (a) a 2-year-old pony mosaic [with karyotype 63,X/64,XX/65,XX,del(Y)(q?); see section on mosaics; photo courtesy of M. Bugno]; (b) a 2-year-old draft cross XX true hermaphrodite (Sry negative, 64,XX with ovotestes; see section on sex reversals; photo courtesy of J. Brendemuehl) (photograph courtesy of J. Brendemuehi); and (c) a donkey with presumed testicular feminization (descended testes and normal male karyotype 62,XY; see section on pseudohermaphroditism; photograph courtesy of R. Torbeck).
for human patients, a new system has been proposed to replace such terms (intersex, true hermaphrodite, pseudohermaphrodite) with different nomenclature (disorders of sexual development). While the proposed system has yet to be universally adopted, it may more easily integrate molecular diagnoses in the future.38, 39 Presentation of an abnormal sexual phenotype will vary depending on the specific cause. Many of the abnormalities discussed below can manifest similarly, as variations of an ambiguous phenotype (see Figure 228.4). For that reason, it is not possible to make an etiological diagnosis based on physical appearance alone. For a definitive diagnosis of an intersex condition, minimally karyotyping and histopathological analysis of gonadal tissue are necessary. In some cases, evaluation of internal genitalia or further molecular diagnostics (e.g., PCR analysis for Sry) is needed. When gonadal tissue is not obtain-
Due to non-disjunction errors during meiosis, where paired sister chromatids fail to separate to opposite poles during anaphase, an ovum or sperm may not contain a sex chromosome. If such an abnormal gamete fuses with a normal coordinate gamete (e.g., an X-chromosome-bearing ovum fusing with a sperm lacking a sex chromosome, or an X-chromosomebearing sperm fertilizes an ovum lacking an X chromosome), the result will be a zygote with only one sex chromosome (63,XO; the 63,YO karyotype is lethal). While the resultant 63,XO zygote is often viable, it will not develop entirely normally. The body system to suffer the most is the reproductive system, but it is not alone. Monosomy X individuals (known as Turner syndrome in humans) have been reported in horses.40–43 The most common presenting complaint is an intended brood mare failing to cycle, or cycling with irregularity. These mares may tend to be smaller in stature and lack impressive athletic ability. Monosomy X mares may tease to the stallion as if in heat, but may not stand to be mounted. Examination of the reproductive tract usually shows the ovaries to be small and inactive. Oocytes are absent, as two X chromosomes are necessary to maintain oocyte viability, and as a result the follicles degenerate. Therefore follicular function is absent in XO mares, the exceptions being those that are actually chromosomal mosaics (discussed below). The hypoplastic uterus and cervix lack tone. Diagnosis is confirmed by karyotype (63,XO). Mature, maiden mares with inactive ovaries during the breeding season should have a karyotype evaluated if there is no history of GnRH vaccination.44
XXY syndrome Also due to meiotic, non-disjunctional errors, stallions with the XXY karyotype occur either when a Ybearing sperm fertilizes an ovum containing two X chromosomes, or when an XY-bearing sperm fertilizes a normal, X-bearing ovum. This is called Klinefelter syndrome in humans. One case of an XXXY intersex horse with Klinefelter-like syndrome has also been reported.45 XXY horses may have scrotal or cryptorchid testes,46 but all are sterile. Those with scrotal testes are usually assumed to be normal stallions and diagnoses are only made after puberty and unsuccessful
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Sex Determination and Differentiation attempts at breeding. Physical examination usually shows the testes to be soft and the penis is usually relatively small, in comparison with what is expected for that size stallion. Libido may be normal, but ejaculates will always be azoospermic. Histology of the testes will usually reveal decreased numbers of hypoplastic seminiferous tubules devoid of spermatogenesis. Cryptorchid Klinefelter stallions with an unknown history may be purchased as ‘geldings’ and then become suspect when they show stallion-like behaviors. Leydig cells in the testes produce testosterone concentrations comparable with (or even higher than) normal stallions,47 but the testes are unable to support spermatogenesis. Definitive diagnosis is by karyotype, identifying an extra X chromosome (karyotype 65,XXY). Although one case of a 65,XXY stallion has been reported,48 most cases reported were mosaics;46,49–52 see discussion of mosaics below.
Trisomy X Non-disjunction during meiosis can also result in trisomy of the X chromosome (65,XXX). These individuals usually appear as normal mares externally, but frequently show reduced fertility.53–56 In at least one reported case, a trisomy X mare seemed to have testicular formation and exhibit stallion-like behavior and physical appearance (crested neck and enlarged clitoris).54 An explanation for the appearance and behavior of this mare was not discovered and is not explained by the trisomy X alone.
Mosaics and chimeras A chimera occurs naturally when two littermates or twins share genetic material in utero, resulting in a genome that reflects, to some degree, the other individual. The fertilization of an oocyte by two separate sperm cells has also been implicated in some cases of chimerism. The classic example of chimerism in domestic animals is the freemartin syndrome (reviewed in Padula, 2005),57 most common in cattle. Chimerism is rare in horses due to the extremely low incidence of successful twinning and fundamental differences in timing of fetal development and type of placentation in the horse.58 Rare cases have been reported, as in the case of a Welsh pony with an 64,XX karyotype and bilateral ovotestes.59 In other cases, clinical consequences were lacking (i.e., the suspected mares born co-twin to a stallion went on to successfully breed and produce young).60 One documented case reports an 64,XX/64,XY chimerism in a Thoroughbred foal with an enlarged clitoris, increased anogenital distance, and palpable gonads
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within the inguinal region. These gonads were later removed and histologically shown to be testes.61 Mosaicism occurs during a mitotic nondisjunction event. When the non-disjunction involves the sex chromosomes, it can affect the progenitor cell lines and result in abnormal gonadal or phenotypic sexual development. Various combinations of mosaics are possible. Some are lethal, but others result in ambiguous sexuality. Phenotypes observed will vary by case and depend on the degree of non-disjunctional error. In the horse, various mosaic combinations have been reported: 63,XO/64,XX, 63,XO/64,XX, 63,XO/64,XY, 64,XX/65,XXY, 64,XX/65,XYY, 64,XX/65,XXX, 64,XX/65,XXX/6Y,XXY, 63,XO/64,XX/65,XXY, and 63,XO/64,XX/65,XX,del(Y)(q?). Abnormalities of gonadal sex determination: sex reversal In sex reversal the karyotype is normal, but the gonad does not agree with the sex chromosome constitution. For example, a 64,XX horse with testicular tissue would be a case of XX sex reversal. Typically these are either XX males having bilateral testes or XX true hermaphrodites having ovotestes. However, in XY sex reversal, if oocytes and follicles develop in the fetus, they most often degenerate such that ovarian architecture is usually unrecognizable at the time of presentation (gonadal dysgenesis). In the horse, both XX and XY sex reversal have been reported.51, 53, 55, 63,65–76 Fertility has been reported in some XY females.65, 70 However, these cases were reported prior to the advent of molecular diagnostic tests, which are needed to identify the reason that fertility could be present in such cases. The presence or absence of the Sry gene is a key part of the puzzle to understand the etiology behind these disorders. An XX sex-reversed horse that has an active Sry gene through a translocation event is easily explained; likewise an XY sex-reversed horse that is Sry-negative through translocational loss of the gene. In other species, XX sex reversals have been reported that are either Sry-negative77 or Sry-positive.78 To date, however, all XX sex reversal cases in the horse have tested negative for the Sry gene, indicating an error farther downstream.73, 74, 76, 79 Sry-negative XY sex reversal has been reported in mice and humans12,80–82 as well as in the horse.70, 75, 83 Sry-positive XY sex reversal has been reported in mice and humans, due to mutations in genes downstream of Sry.15,84–86 Sry-positive XY sex reversal in the horse, in the sense of an individual with an XY karyotype and ovarian development, has yet to be documented. However, it is rare to find ovaries in XY sex-reversed
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individuals, as most have gonadal dysgenesis, where neither functional testes nor ovaries develop. As neither testosterone nor AMH is produced in such individuals, they develop as phenotypic females that have a uterus. In the horse literature, the term ‘sex reversal’ has been used by some as a catch-all phrase to describe both genotypic and phenotypic abnormalities of sexual development (i.e., any 64,XY horse that did not develop into a normal male, regardless of whether the gonads were testes, ovaries or dysgenic gonads). In this vein, various authors have reported ‘sex reversal’ in XY horses, but many of these were cases of abnormal phenotypic sexual development (pseudohermaphrodites), specifically androgen insensitivity69, 70, 87 (see below). As discussed below, the latter have testes that produce testosterone and AMH, and the uterus is absent, features that are clinically distinguishable from the phenotype of XY sex reversal with gonadal dysgenesis. Diagnosis of sex reversal requires a karyotype identifying a normal (64,XX or 64,XY) karyotype and histopathological confirmation that the gonadal sex does not agree with the karyotype. In cases where gonadectomy is not performed, ultrasonographic evaluation of the gonads and internal reproductive system may provide sufficient information to reach a presumptive diagnosis. This is obviously not definitive, but can be beneficial to case management.63 Hormonal assays may also be performed and are discussed in the next section (see also Kent et al. 1988).69 Abnormalities of phenotypic sexual development: pseudohermaphroditism This section addresses cases in which the karyotype and gonad are in agreement but the phenotype is ambiguous. These are the pseudohermaphrodites, because the gonadal tissue is testicular or ovarian, but not both. Female pseudohermaphrodites include individuals with 64,XX karyotype, ovaries and masculinization of the external genitalia. These have yet to be reported in horses, but have been reported in other species.88 Male pseudohermaphrodites include individuals with 64,XY karyotype and testes, but retention of female internal and/or external genitalia. There are two major categories; either (i) mullerian derivatives persist (persistent mullerian duct syndrome, uterus masculinus) or (ii) failure of androgen dependent masculinization. In horses, the latter is most commonly reported.
Defects in androgen dependent masculinization: male pseudohermaphroditism In humans these have been linked to defects in testosterone biosynthesis, or defects in the 5α-reductase
type 2 enzyme.89–95 The term androgen insensitivity syndrome (AIS) refers only to the latter etiology, where both testosterone and DHT are present yet there is complete or partial failure of androgen dependent masculinization due to the androgen receptor defect. However, the uterus is absent as expected, as there is no defect in AMH or mullerian duct regression. As the androgen receptor gene is located on the X chromosome in all mammals, these mutations are inherited from the dam.91, 96 In human medicine, the term testicular feminization (Tfm) is reserved for male pseudohermaphroditism caused by complete androgen insensitivity syndrome (e.g., a non-functional androgen receptor). As there are several reported mutations that variably disrupt androgen receptor function, the associated phenotypes vary from infertile males to females that are sterile.95 A non-functional androgen receptor affects all androgen targets, internal and external. Phenotypic expression of other defects in androgen-dependent masculinization differs according to the type of defect. For example, 5α-reductase dysfunction affects only those target organs where DHT induces masculinization, which are the external genitalia (see Figure 228.1). However, this defect has not yet been identified in horses. Male pseudohermaphroditism has been reported in horses,63, 87, 97, 98 although definitive diagnosis is rare. Due to the lack of research to clarify the pathogenesis of the equine syndromes, and the lack of androgen receptor diagnostic tests, Tfm has been inappropriately used to describe many of these cases. Those likely to have dysfunctional androgen receptors have external female (or ambiguous) genitalia, although the body elsewhere may appear stallionlike (crested neck, well-muscled). The behavior is often reflective of stallions, with reports of herding mares, mounting or vocalizing in the presence of estrous mares, aggression towards other horses, and defecating in a pile in their enclosures.87, 98 Estrogens in stallions support good libido99 and may come from aromatization of circulating testosterone to estrogen in the brain tissue.100 Transrectal palpation of the internal reproductive tract will not reveal uterus or cervix, as the mullerian ducts regress as expected. Gonads will be smooth and ovoid, and may be located in the position of the ovaries, or anywhere between the caudal kidney and inguinal rings. Transrectal ultrasound evaluation will show these structures to be consistent with testes, not ovaries. Vaginal speculum examination will reveal a blind-ending, shortened vagina, which represents only the caudal vagina that is derived from the urogenital sinus. Demonstrating a 64,XY karyotype, testes, and female external genitalia and absence of the uterus is necessary for presumptive diagnosis of testicular feminization in the horse. Testicular tissue may be
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Sex Determination and Differentiation directly diagnosed by histopathology. Demonstration of a dysfunctional androgen receptor by in vitro binding studies or identification of an androgen receptor mutation establishes the definitive diagnosis. Besides providing diagnostic information, castration is useful in cases where the stallion-like behavior of the horse is undesirable. No correlation has yet been demonstrated in the horse between abdominal testes having an increased likelihood to develop seminomas or Sertoli cell tumors, as in the dog and human. In cases where surgical removal of the abdominal gonads is not a possibility or priority, presumptive diagnosis can be obtained from the ultrasonographic appearance or hormonal assays, as in other cases of cryptorchidism. Baseline serum testosterone concentrations are usually well above reference ranges for normal mares and reflect values more appropriate for normal stallions.69 If baseline concentrations of serum testosterone are not clearly diagnostic, a challenge test may be performed (see Chapters 105 and 158).
9.
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11.
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Summary Intersex conditions should be suspected in cases of infertility in horses, especially if any degree of phenotypic ambiguity is evident. Karyotype and gonadal histology are necessary for definitive diagnosis. In addition, to understand the etiology more specifically and provide genetic counseling, testing for specific gene mutations may be necessary.
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References 1. Jost A, Vigier B, Prepin J, Perchellet JP. Studies on sex differentiation in mammals. Recent Prog Horm Res 1973;29:1–41. 2. Wain HM, Bruford EA, Lovering RC, Lush MJ, Wright MW, Povey S. Guidelines for Human Gene Nomenclature. Genomics 2002;79(4): 464–70. 3. DiNapoli L, Capel B. SRY and the standoff in sex determination. Mol Endocrinol 2008;22:1–9. 4. Burgoyne PS, Baker TG. Meiotic pairing and gametogenic failure. Symp Soc Exp Biol 1984;38:349–62. 5. Staack A, Donjacour AA, Brody J, Cunha GR, Carroll P. Mouse urogenital development: a practical approach. Differentiation 2003;71:402–13. 6. Gubbay J, Collignon J, Koopman P, Capel B, Economou A, Munsterberg A, Vivian N, Goodfellow P, LovellBadge R. A gene mapping to the sex-determining region of the mouse Y chromosome is a member of a novel family of embryonically expressed genes. Nature 1990;346: 245–50. 7. Sinclair AH, Berta P, Palmer MS, Hawkins JR, Griffiths BL, Smith MJ, Foster JW, Frischauf A-M, LovellBadge R, Goodfellow PN. A gene from the human sexdetermining region encodes a protein with homology to a conserved DNA-binding motif. Nature 1990;346:240–4. 8. Sekido R, Bar I, Narv·ez V, Penny G, Lovell-Badge R. SOX9 is up-regulated by the transient expression of SRY
18. 19.
20.
21.
22.
23.
24.
2219
specifically in Sertoli cell precursors. Dev Biol 2004;274: 271–79. Kent J, Wheatley SC, Andrews JE, Sinclair AH, Koopman P. A male-specific role for SOX9 in vertebrate sex determination. Development 1996;122:2813–22. da Silva SM, Hacker A, Harley V, Goodfellow P, Swain A, Lovell-Badge R. Sox9 expression during gonadal development implies a conserved role for the gene in testis differentiation in mammals and birds. Nat Genet 1996; 14:62–8. Chaboissier M-C, Kobayashi A, Vidal VIP, Lutzkendorf S, van de Kant HJG, Wegner M, de Rooij DG, Behringer RR, Schedl A. Functional analysis of Sox8 and Sox9 during sex determination in the mouse. Development 2004;131:1891–901. Barrionuevo F, Bagheri-Fam S, Klattig J, Kist R, Taketo MM, Englert C, Scherer G. Homozygous Inactivation of Sox9 Causes Complete XY Sex Reversal in Mice. Biol Reprod 2006;74:195–201. Adams IR, McLaren A. Sexually dimorphic development of mouse primordial germ cells: switching from oogenesis to spermatogenesis. Development 2002;129:1155–64. Wilhelm D, Martinson F, Bradford S, Wilson MJ, Combes AN, Beverdam A, Bowles J, Mizusaki H, Koopman P. Sertoli cell differentiation is induced both cell-autonomously and through prostaglandin signaling during mammalian sex determination. Dev Biol 2005;287:111–24. Colvin JS, Green RP, Schmahl J, Capel B, Ornitz DM. Male-to-female sex reversal in mice lacking fibroblast growth factor 9. Cell 2001;104:875–89. Kim Y, Kobayashi A, Sekido R, DiNapoli L, Brennan J, Chaboissier M-C, Poulat F, Behringer RR, Lovell-Badge R, Capel B. Fgf9 and Wnt4 act as antagonistic signals to regulate mammalian sex determination. PLoS Biol 2006; 4:e187. Yao HH-C. The pathway to femaleness: current knowledge on embryonic development of the ovary. Mol Cell Endocrinol 2005;230:87–93. Goodfellow PN, Lovell-Badge R. SRY and sex determination in mammals. Annu Rev Genet 1993;27:71–92. McElreavey K, Vilain E, Cotinot C, Payen E, Fellous M. Control of sex determination in animals. Eur J Biochem 1993;218:769–83. Maatouk DM, DiNapoli L, Alvers A, Parker KL, Taketo MM, Capel B. Stabilization of beta-catenin in XY gonads causes male-to-female sex-reversal. Hum Mol Genet 2008;17:2949–55. Liu C-F, Bingham N, Parker K, Yao HHC. Sex-specific roles of beta-catenin in mouse gonadal development. Hum Mol Genet 2009;18:405–17. Kim KA, Zhao J, Andarmani S, Kakitani M, Oshima T, Binnerts ME, Abo A, Tomizuka K, Funk WD. R-spondin proteins: a novel link to β-catenin activation. Cell Cycle 2006;5:23–6. Chassot A-A, Ranc F, Gregoire EP, Roepers-Gajadien HL, Taketo MM, Camerino G, de Rooij DG, Schedl A, Chaboissier M-C. Activation of beta-catenin signaling by Rspo1 controls differentiation of the mammalian ovary. Hum Mol Genet 2008;17:1264–77. Tomizuka K, Horikoshi K, Kitada R, Sugawara Y, Iba Y, Kojima A, Yoshitome A, Yamawaki K, Amagai M, Inoue A, Oshima T, Kakitani M. R-spondin1 plays an essential role in ovarian development through positively regulating Wnt-4 signaling. Hum Mol Genet 2008;17:1278–91.
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25. Heikkila M, Prunskaite R, Naillat F, Itaranta P, Vuoristo J, Leppaluoto J, Peltoketo H, Vainio S. The partial female to male sex reversal in Wnt-4-deficient females involves induced expression of testosterone biosynthetic genes and testosterone production, and depends on androgen action. Endocrinology 2005;146:4016–23. 26. Vainio S, Heikkila M, Kispert A, Chin N, McMahon AP. Female development in mammals is regulated by Wnt-4 signalling. Nature 1999;397:405–9. 27. Jeays-Ward K, Hoyle C, Brennan J, Dandonneau M, Alldus G, Capel B, Swain A. Endothelial and steroidogenic cell migration are regulated by WNT4 in the developing mammalian gonad. Development 2003;130:3663–70. 28. Yao HH, Matzuk MM, Jorgez CJ, Menke DB, Page DC, Swain A, Capel B. Follistatin operates downstream of Wnt4 in mammalian ovary organogenesis. Dev Dyn 2004;230:210–15. 29. Tevosian SG, Manuylov NL. To beta or not to beta: Canonical beta-catenin signaling pathway and ovarian development. Dev Dyn 2008;237:3672–80. 30. Akiyama H, Lyons JP, Mori-Akiyama Y, Yang X, Zhang R, Zhang Z, Deng JM, Taketo MM, Nakamura T, Behringer RR, McCrea PD, de Crombrugghe B. Interactions between Sox9 and beta-catenin control chondrocyte differentiation. Genes Dev 2004;18:1072–87. 31. Hill TP, Spater D, Taketo MM, Birchmeier W, Hartmann C. Canonical Wnt/beta-catenin signaling prevents osteoblasts from differentiating into chondrocytes. Dev Cell 2005;8:727–38. 32. Parma P, Radi O, Vidal V, Chaboissier MC, Dellambra E, Valentini S, Guerra L, Schedl A, Camerino G. R-spondin1 is essential in sex determination, skin differentiation and malignancy. Nat Genet 2006;38:1304–9. 33. Pailhoux E, Vigier B, Chaffaux S, Servel N, Taourit S, Furet J-P, Fellous M, Grosclaude F, Cribiu EP, Cotinot C, Vaiman D. A 11.7-kb deletion triggers intersexuality and polledness in goats. Nat Genet 2001;29:453–8. 34. Blecher SR, Erickson RP. Genetics of sexual development: A new paradigm. Am J Med Genet A 2007;143A:3054–68. 35. Kim Y, Capel B. Balancing the bipotential gonad between alternative organ fates: A new perspective on an old problem. Dev Dyn 2006;235:2292–300. 36. Amann RP, Veeramachaneni DNR. Cryptorchidism in common eutherian mammals. Reproduction 2007;133:541– 61. 37. Meyers-Wallen VN. CVT Update: inherited disorders of the reproductive tract in dogs and cats. In: Bonagura J (ed.) Current Veterinary Therapy XIV. Philadelphia: Saunders, 2008. 38. Lee PA, Houk CP, Ahmed SF, Hughes IA, in collaboration with the participants in the International Consensus Conference on Intersex organized by the Lawson Wilkins Pediatric Endocrine S, the European Society for Paediatric E. Consensus Statement on Management of Intersex Disorders. Pediatrics 2006;118:e488–500. 39. Hughes IA. Disorders of sex development: a new definition and classification. Best Pract Res Clin Endocrinol Metab 2008;22:119–34. 40. Blue MG, Bruere AN, Dewes HF. The significance of the X0 syndrome in infertility of the mare. N Z Vet J 1978;26: 137–41. 41. Hughes JP, Kennedy PC. XO-gonadal dysgenesis in the mare (report of two cases). Eq Vet J 1975;7:109–12. 42. Kutzler MA. Theriogenology question of the month: X chromosome monosomy (XO syndrome) in the mare. J Am Vet Med Assoc 2001;219:751–2.
43. Makinen A, Suojala L, Niini T, Katila T, Tozaki T, Miyake YI, Hasegawa T. X chromosome detection in an XO mare using a human X paint probe, and PCR detection of SRY and amelogenin genes in 3 XY mares. Eq Vet J 2001;33: 527–30. 44. Robinson SJ, McKinnon AO. Prolonged ovarian inactivity in broodmares temporally associated with administration of Equity. Aust Equine Vet 2006;25:85–7. 45. Gluhovshci N, Bistriceanu M, Sucui A, Bratu M. A case of intersexuality in the horse with type 2 A + XXXY chromosome formula. Br Vet J 1970;126:522–5. 46. Makinen A, Katila T, Andersson M, Gustavsson I. Two sterile stallions with XXY-syndrome. Eq Vet J 2000;32: 358–60. 47. Arighi M, Bosu WTK. Comparison of hormonal methods for diagnosis of cryptorchidism in horses. Eq Vet Sci 1989;9:20–6. 48. Kubien EM, Pozor MA, Tischner M. Clinical, cytogenetic and endocrine evaluation of a horse with a 65,XXY karyotype. Eq Vet J 1993;25:333–5. 49. Basrur PK, Kanagawa H, Gilman JPW. An equine intersex with unilateral gonadal agenesis. Can J Comp Med 1969;33:297–306. 50. Bouters R, Vandeplassche M, De Moor A. An intersex (male pseudohermaphrodite) horse with 64XX/65XXY mosaicism. Eq Vet J 1972;4:150–3. 51. Dunn HO, Vaughan JT, McEntee K. Bilaterally cryptorchid stallion with female karyotype. Cornell Vet 1974; 64. 52. Fretz PB, Hare WCD. A male pseudohermaphrodite horse with 63XO?/64XX/65XXY mixoploidy. Eq Vet J 1976; 8. 53. Chandley AC, Fletcher J, Rossdale PD, Peace CK, Ricketts SW, McEnery RJ, Thorne JP, Short RV, Allen WR. Chromosome abnormalities as a cause of infertility in mares. J Reprod Fertil Suppl 1975;23:377–83. 54. Moreno-Millan M, Delgado Bermejo JV, Lopez Castillo G. An intersex horse with X chromosome trisomy. Vet Rec 1989;124:169–70. 55. Makinen A, Hasegawa T, Makila M, Katila T. Infertility in two mares with XY and XXX sex chromosomes. Eq Vet J 1999;31:346–9. 56. Bugno M, Slota E, Wieczorek M, Yang F, Buczynski J, Switonski M. Nonmosaic X trisomy, detected by chromosome painting, in an infertile mare. Eq Vet J 2003;35: 209–10. 57. Padula AM. The freemartin syndrome: an update. Anim Reprod Sci 2005;87:93–109. 58. Bouters R, Vandeplassche M. Twin gestation in the mare: the incidence of placental vascular anastomoses and their influence on the reproductive performance of heterosexual equine twins. J Reprod Fertil 1972;29:149. 59. Dunn H, Smiley D, Duncan J, McEntee K. Two equine true hermaphrodites with 64,XX/64,XY and 63,XO/64,XY chimerism. Cornell Vet 1981;71:123–35. 60. Marcum JB. The freemartin syndrome. Anim Breed Abstr 1974;42:227–242. 61. Power MM, Leadon DP. Diploid-triploid chimaerism (64, XX/96,XXY) in an intersex foal. Eq Vet J 1990;22:211– 14. 62. Hughes JP, Stabenfeldt GH, Kennedy PC. The estrous cycle and selected functional and pathologic ovarian abnormalities in the mare. Vet Clin N Am Large Anim Pract 1980;2:225–39. 63. Kuiper H, Distl O. Intersexuality in horses. Dtsch Tierarztl Wochenschr 2007;114:50–6.
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Sex Determination and Differentiation 64. Bugno M, Zabek T, Golonka P, Pienkowska-Schelling A, Schelling C, Slota E. A case of an intersex horse with 63,X/64,XX/65,XX,del(Y)(q?) karyotype. Cytogenet Genom Res 2008;120:123–6. 65. Sharp AJ, Wachtel SS, Benirschke K. H-Y antigen in a fertile XY female horse. J Reprod Fert 1980;58:157–60. 66. Power MM. XY sex reversal in a mare. Eq Vet J 1986; 18:233–6. 67. Trommershausen Bowling A, Millon L, Hughes JP. An update of chromosomal abnormalities in mares. J Reprod Fertil Suppl 1987;35:149–55. 68. Long SE. Chromosome anomalies and infertility in the mare. Eq Vet J 1988;20:89–93. 69. Kent MG, Schneller HE, Hegsted RL, Johnston SD, Wachtel SS. Concentration of serum testosterone in XY sex reversed horses. J Endocrin Invest 1988;11:609–13. 70. Kent MG, Shoffner RN, Hunter A, Elliston KO, Schroder W, Tolley E, Wachtel SS. XY sex reversal syndrome in the mare: clinical and behavioral studies, H-Y phenotype. Hum Genet 1988;79:321–8. 71. Constant SB, Larsen RE, Asbury AC, Buoen LC, Mayo M. XX male syndrome in a cryptorchid stallion. J Am Vet Med Assoc 1994;205:83–5. 72. Pailhoux E, Cribiu E, Parma P, Cotinot C. Molecular analysis of an XY mare with gonadal dysgenesis. Hereditas 1995;122:109–12. 73. Buoen LC, Zhang TQ, Weber AF, Ruth GR. SRY-negative, XX intersex horses: the need for pedigree studies to examine the mode of inheritance of the condition. Eq Vet J 2000;32:78–81. 74. Vaughan L, Schofield W, Ennis S. Sry-negative XX sex reversal in a pony: a case report. Theriogenology 2001;55: 1051–7. 75. Bugno M, Klukowska J, Slota E, Tischner M, Switonski M. A sporadic case of the sex-reversed mare (64,XY; SRY-negative): molecular and cytogenetic studies of the Y chromosome. Theriogenology 2003;59:1597–603. 76. Bannasch D, Rinaldo C, Millon L, Latson K, Spangler T, Hubberty S, Galuppo L, Lowenstine L. SRY negative 64,XX intersex phenotype in an American saddlebred horse. Vet J 2007;173:437–9. 77. Kocer A, Pinheiro I, Pannetier M, Renault L, Parma P, Radi O, Kim K-A, Camerino G, Pailhoux E. R-spondin1 and FOXL2 act into two distinct cellular types during goat ovarian differentiation. BMC Dev Biol 2008;8:36. 78. Reddy PP, Papenhausen PR, Suh Y-M, Riddick LM, Calvano CJ, Mandell J. XX sex reversal: molecular analysis of the SRY/ZFY regions. J Urol 1997; 158:1305–7. 79. Meyers-Wallen VN, Hurtgen J, Schlafer D, Tulleners E, Cleland WR, Ruth GR, Acland GM. Sry-negative XX true hermaphroditism in a Pasa Fino horse. Eq Vet J 1997;29:404–8. 80. Luo X, Ikeda Y, Parker KL, Renfree MB, McLaren A, Josso N. The cell-specific nuclear receptor steroidogenic factor 1 plays multiple roles in reproductive function [and Discussion]. Phil Trans R Soc B: Biol Sci 1995;350:279–83. 81. Achermann JC, Ito M, Ito M, Hindmarsh PC, Jameson JL. A mutation in the gene encoding steroidogenic factor-1 causes XY sex reversal and adrenal failure in humans. Nat Genet 1999;22:125–6. 82. Bouma GJ, Albrecht KH, Washburn LL, Recknagel AK, Churchill GA, Eicher EM. Gonadal sex reversal in mutant Dax1 XY mice: a failure to upregulate Sox9 in pre-Sertoli cells. Development 2005;132:045–54.
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83. Abe S, Miyake YI, Kageyama SI, Watanabe G, Taya K, Kawakura K. Deletion of the Sry region on the Y chromosome detected in a case of equine gonadal hypoplasia (XY female) with abnormal hormonal profiles. Eq Vet J 1999;31:336–8. 84. Bardoni B, Zanaria E, Guioli S, Floridia G, Worley KC, Tonini G, Ferrante E, Chiumello G, McCabe ERB, Fraccaro M, Zuffardi O, Camerino G. A dosage sensitive locus at chromosome Xp21 is involved in male to female sex reversal. Nat Genet 1994;7:497–501. 85. Zanaria E, Bardoni B, Dabovic B, Calvari V, Fraccaro M, Zuffardi O, Camerino G, Graves JAM, McLaren A. Xp Duplications and sex reversal [and Discussion]. Phil Trans R Soc B: Biol Sci 1995;350:291–6. 86. Le Caignec C, Delnatte C, Vermeesch JR, Boceno M, Joubert M, Lavenant F, David A, Rival JM. Complete sex reversal in a WAGR syndrome patient. Am J Med Genet A 2007;143A:2692–5. 87. Switonski M, Chmurzynska A, Szczerbal I, Lipczynski A, Yang F, Nowicka-Posluszna A. Sex reversal syndrome (64,XY; SRY-positive) in a mare demonstrating masculine behaviour. J Anim Breed Genet 2005;122:60–3. 88. Frimberger D, Gearhart JP. Ambiguous genitalia and intersex. Urol Int 2005;75:291–7. 89. Andersson S, Berman DM, Jenkins EP, Russell DW. Deletion of steroid 5[alpha]-reductase 2 gene in male pseudohermaphroditism. Nature 1991;354:159–61. 90. Imperato-McGinley J, Zhu YS. Androgens and male physiology: the syndrome of 5[alpha]-reductase-2 deficiency. Mol Cell Endocrinol 2002;198:51–9. 91. Migeon BR, Brown TR, Axelman J, Migeon CJ. Studies of the locus for androgen receptor: localization on the human X chromosome and evidence for homology with the Tfm locus in the mouse. Proc Natl Acad Sci USA 1981;78:6339–43. 92. McPhaul MJ, Griffin JE. Male pseudohermaphroditism caused by mutations of the human androgen receptor. J Clin Endocrinol Metab 1999;84:3435–41. 93. Sultan C, Paris F, Terouanne B, Balaguer P, Georget V, Poujol N, Jeandel C, Lumbroso S, Nicolas J-C. Disorders linked to insufficient androgen action in male children. Hum Reprod Update 2001;7:314–22. 94. Ahmed SF, Hughes IA. The genetics of male undermasculinization. Clin Endocrinol 2002;56:1–18. 95. Galani A, Kitsiou-Tzeli S, Sofokleous C, Kanavakis E, Kalpini-Mavrou A. Androgen insensitivity syndrome: clinical features and molecular defects. Hormones 2008; 7:217–29. 96. Lyon MF, Hawkes SG. X-linked gene for testicular feminization in mouse. Nature 1970;227:1217–19. 97. Crabbe BG, Freeman DA, Grant BD, Kennedy P, Whitlatch L, MacRae K. Testicular feminization syndrome in a mare. J Am Vet Med Assoc 1992;200:1689–91. 98. Howden KJ. Androgen insensitivity syndrome in a Thoroughbred mare (64,XY–testicular feminization). Can Vet J 2004;45:501–3. 99. Thompson DL Jr, Pickett BW, Squires EL, Nett TM. Sexual behavior, seminal pH and accessory sex gland weights in geldings administered testosterone and(or) estradiol-17B. J Anim Sci 1980;51:1358–66. 100. Lemazurier E, Sourdaine P, Nativelle C, Plainfoss`e B, S`eralini G-E. Aromatase gene expression in the stallion. Mol Cell Endocrinol 2001;178:133–9.
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Chapter 229
Endocrinology of Pregnancy Jennifer C. Ousey
Introduction The endocrine maintenance of pregnancy and initiation of parturition is a series of coordinated hormonal events involving interactions between the developing fetus, its major organ of support, the placenta, and the mare. In the early stages of pregnancy (< 45 days), hormones (progesterone, estrogens, gonadotropins) from the mare (ovaries, endometrium, brain) and the growing conceptus, play a major role in the endocrine recognition of pregnancy. As the allantochorion (placenta) attaches and starts to expand (40 to 120 days), it too assumes significant endocrine functions and secretory capabilities – equine chorionic gonadotropin (eCG), relaxin, oestrogens, and progestagens. The growing fetus develops in concert with the placenta and, from about 70 days, fetus and placenta use substrates obtained from the maternal circulation to synthesize the major endocrine products of pregnancy – progestagens and estrogens. This represents, quantitatively, the most significant endocrine activity for the majority of gestation and ensures the continued development of the fetus within a controlled uterine environment. From the end of the first trimester, the fetal endocrine organs begin to synthesize their own hormones, which facilitate normal fetal growth and development and, when necessary, allow the fetus to respond to fluctuations in the supply of substrates from the mare. Finally, in the periparturient period, there is upregulation of maternal and fetal endocrine activity in preparation for the processes of delivery and life ex utero, respectively. Much of the information about endocrinology during early pregnancy has been described in other chapters, and, therefore, only a brief synopsis will be given here. Instead the major endocrine developments of mid- and late pregnancy will be described, particularly after the establishment of the placenta, and detailed information given about the ontogeny of hormone profiles during late gestation and the periparturient period, in the healthy pregnant mare.
Progesterone and related compounds Progesterone (P4) and related progestagenic compounds are required to maintain pregnancy. High progestagen concentrations block prostaglandin synthesis, which otherwise would cause luteolysis of the primary corpus luteum (CL) during early pregnancy or, in later pregnancy, induce myometrial contractions. Studies have shown that repeated administration of prostaglandin F2α (PGF2α ) to either early or late pregnant mares will cause abortion/ delivery but can be prevented (in early pregnancy) if progesterone supplementation is given concurrently.1, 2 The primary CL secretes high concentrations of P4 between days 0 and 8 of pregnancy although levels decline between days 10 and 30 of pregnancy because this CL remains largely inactive without a luteotrophic stimulus.3, 4 After day 38–40, P4 levels increase through the luteinizing activity of eCG, secreted by the developing endometrial cups, which stimulates ovulation of secondary follicles (for references see Allen5 and Ginther6 ). Progesterone concentrations continue to rise and peak between 90 and 120 days of pregnancy following maximum eCG secretion. After this peak, P4 levels begin to decline as the development of supplementary CLs ceases in response to disappearance of eCG and sloughing of the endometrial cups. The maternal ovaries produce predominantly P4 in early pregnancy but this is metabolized into 5α-pregnan-3,20-dione (5α-DHP) and 3β-hydroxy-5α-pregnan-20-dione (3β-5P), and maternal plasma concentrations of these metabolites follow similar profiles to P4 during the first trimester.7 With the gradual regression of the CLs and decline in ovarian P4, production of 5α-DHP and other progestagens is sustained by the fetoplacental unit, which becomes the dominant source of progestagens for the remainder of pregnancy.8 The placenta develops as an active endocrine organ during its period of expansion and attachment
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Endocrinology of Pregnancy
is demonstrated by the studies of Holtan et al.15 These authors ovariectomized mares and showed that removal of the source of P4 before 70 days of pregnancy caused abortion in the majority of ovariectomized mares, whereas after 140 days there were no abortions because the pregnancies were maintained by feto-placental progestagen production.15 Although the young equine conceptus has 3βHSD activity and can produce progesterone and androgens (possibly as intermediates of estrogen production) from day 10, P4 concentrations in maternal plasma are essentially ovarian in origin.16, 17 However, as the fetus and placenta grow and their endocrine capacity increases, from day 60 a number of progestagens other than P4 start to appear in the maternal plasma.7 Much research has been devoted to identifying and quantifying the different progestagens present in fetal and maternal plasma, their ontogenic profiles, and how progestagen metabolism changes at parturition (for reviews see Chavatte et al. 18 and Ousey19 ). The main steroid precursor is cholesterol, which probably originates in the maternal circulation, crosses the placenta, and is converted into pregnenolone (P5) in the fetus by the cytochrome P450 side-chain cleavage enzyme (P450scc ), which is present in several fetal tissues including the adrenal glands, gonads, and trophoblast (Figure 229.1).18, 20
to the underlying maternal endometrium between 40 and 120 days of gestation. The trophoblast cells located between the regressing yolk sac and expanding allantochorion develop into the chorionic girdle, which invades the maternal endometrium and creates the endometrial cups, which secrete eCG until about 120 days of gestation.9 The non-invasive portion of the trophoblast, the allantochorion, gradually interdigitates with the underlying maternal endometrium to form discrete microcotyledons.10 By 90 days, these microcotyledons cover the whole surface of the maternal endometrium and their microvilli have begun to branch extensively in order to increase the efficiency of gas and nutrient exchange between the fetal and maternal circulations. The uterine blood flow increases substantially during the first trimester (from 70 to 4000 mL/min) concurrent with extensive growth in the height and branching of the microvilli.11–14 Consequently increasing quantities of steroid precursors are transported from the maternal to the fetal circulation and metabolized within the developing fetal and placental tissues to produce progestagens (and estrogens), which are then secreted back into the maternal circulation. Details of these metabolic pathways are described below. The ability of the feto-placental unit to support pregnancy through production of progestagens
FETUS
UTERO-PLACENTAL TISSUE
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P4 5α reductase
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5αDHP
5αDHP
20α reductase 20α5P
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Figure 229.1 Biosynthetic pathway of progestagens in the equine feto-placental unit: - - - cortisol pathway in late gestation fetus/foal; - - - alternative pathway to 5α-DHP (5α-pregnan-3,20-dione). P4, progesterone; P5, pregnenolone; P5ββ, pregnanediol; 17α-OH, 17α-hydroxylase; 3β-HSD, 3β-hydroxysteroid dehydrogenase; 20α5P, 20α-hydroxy-5alpha-pregnan-3-one. From Ousey.112
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Removal of the fetal gonads in mid-gestation does not alter the circulating progestagen concentrations whereas exogenous adrenocorticotropic hormone (ACTH) administration causes a rise in progestagens, suggesting that the adrenal glands are the major source of progestagen precursors.21, 22 P5 concentrations are much higher in the umbilical arteries compared to the umbilical vein, so P5 is directed from the fetus to the placenta–endometrium interface where most is converted to P4 and 5α-DHP by the enzymes 3β-HSD and 5α-reductase, located in the fetal trophoblast and endometrium, respectively (Figure 229.1).20, 23, 24 P5 is also metabolized directly into pregnanediol (P5ββ) and excreted into the maternal plasma. Some 5α-DHP is returned to the fetus via the umbilical vein for further metabolism in the fetal liver and kidneys whilst the rest is excreted into the uterine vein or converted within the placental-endometrial complex to other progestagens (Figure 229.1). The proportion of 5α-DHP returning to the fetus increases with increasing gestational age.25 The net uptake of progestagens by the feto-placental unit is greater than their net output. As a consequence of this substantial progestagen metabolism within the feto-placental tissues, circulating P4 concentrations are low. Instead high concentrations of P4 metabolites, particularly 20α-hydroxy-5-pregnan-3-one (20α-5P) and 5-pregnane-3β,20α-diol (βα-diol), and other progestagens, circulate in maternal plasma during the second half of pregnancy (Figure 229.1). Measured using quantitative gas chromatography-mass spectrometry, maternal plasma concentrations of various progestagens range from 50 to 1000 ng/mL whereas P4 concentrations are below the detection limit of the assay system, < 1 ng/mL.7, 8 For most of gestation, there is large uptake of P5 by the utero-placental (UP) tissues, minimal output of P4, and high output of 5α-DHP into the umbilical circulation, particularly towards term.26 Therefore, intracellular concentrations of P4 within the UP tissues are likely to be high. It is hypothesized that these high P4 concentrations may be important for uterine quiescence by regulating the enzyme prostaglandin dehydrogenase (PGDH) located in the maternal endometrium.20 In sheep, PGDH is upregulated by P4 while the activity of the enzyme responsible for prostaglandin synthesis (prostaglandin H synthase, PGHS) is downregulated by P4.27 If a similar mechanism exists in the pregnant mare, then high intracellular P4 concentrations will ensure that prostaglandins are maintained at low levels throughout pregnancy. At parturition, declining P4/progestagen concentrations due to blockade of 3β-HSD (see below) may allow upregulation of PGHS and prostaglandin synthesis, ultimately leading to myometrial contractility. At present, there is no
direct evidence to prove this; in fact, some evidence exists that 5α-DHP may be more important for regulation of myometrial activity. This is discussed in detail by Fowden et al.28 From the end of the first trimester of pregnancy, the fetal adrenal gland weight increases, the placental microvilli continue to grow and branch, and the umbilical and uterine blood flow increases.12, 29, 30 Consequently, fetal and maternal progestagen concentrations slowly rise over the second and third trimesters as greater quantities of steroid precursors are delivered to the uterus and metabolized within the feto-placental tissues.25 However, during the last few weeks of pregnancy, total progestagen concentrations increase rapidly, peaking at about 2–3 days before parturition, and then decline during the last few days/hours before parturition (Figure 229.2). The rise in total progestagen concentrations tends to be higher in Thoroughbred compared to pony breeds and may reflect greater adrenal P5 production in the Thoroughbred.31, 32 Although there is an exponential increase in uterine blood flow over the last month of gestation, the rise in progestagens is not simply a consequence of increased delivery of substrates. Also there is a large rise in fetal plasma ACTH, which drives synthesis of P5, a doubling of adrenal gland mass, and increased expression of the progestagenic enzymes in the adrenals and placenta.20, 33 This upregulation of progestagen synthesis towards term is important because there is also increased activity of uterotonic hormones, estradiol-17β,, oxytocin, and prostaglandins (see below). Consequently, increased progestagen suppression of myometrial activity is essential until the fetus is ready for delivery. Surprisingly, the upregulation of progestagen production may also contribute toward the signal for parturition. Chavatte et al.34 and others have demonstrated that most progestagens (except 5α-DHP) inhibit placental 3β-HSD in vitro at concentrations similar to those found in maternal plasma. Moreover, in vitro and in vivo administration of specific 3βHSD inhibitors (trilostane or epostane) blocks P4 production and transiently decreases progestagen concentrations in the pregnant mare whereas cortisol concentrations are increased.35, 36 This progestagen blockade of 3β-HSD activity may not only contribute toward diminished intracellular P4 concentrations and upregulation of uterine prostaglandin synthesis, but also divert P5 production away from P4 to 17α-hydroxyprogesterone and cortisol via induction of the adrenal enzyme, 17α-hydroxylase (17α-OH), which is upregulated in the periparturient period (Figure 229.1). The decline in progestagen concentrations occurs late in gestation and coincides with an increase in fetal cortisol concentrations.28, 33 The stimulus for induction of 17α-OH activity is
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Figure 229.2 Total plasma progestagen concentrations in Thoroughbred and pony mares during the last 2 months of pregnancy. Adapted from Ousey.112
unknown but may be due to increasing ACTH concentrations or removal of an as yet unknown inhibitory factor, for example dehydroepiandrosterone (DHEA).28 Maternal progestagen concentrations reach low levels soon after delivery. However, P5 concentrations in healthy foals remain high at birth and only gradually decline during the first 24 hours, confirming the fetus/foal as the source of P5.37
Estrogens Estrogens have been identified in the equine conceptus from as early as day 8 of pregnancy.38 Estradiol17β, estrone, and their sulfoconjugates have been measured in increasing concentrations in the blastocoelic/yolk sac fluid from days 10 to 26.16, 39 The concentration of estrogens produced by the conceptus rises with the growth in the diameter of the conceptus.40 Despite this, estrogen concentrations in the maternal plasma remain low during the first few weeks of pregnancy, indicating that the considerable amounts of estrogens produced by the conceptus are utilized within the uterus.41 It has been proposed that this significant estrogen production by the young conceptus/embryo may upregulate progesterone receptor numbers within the uterus for maintenance of P4 production, and promote feto-maternal nutrient and gaseous exchange at a time before the placenta is functional and when the growing embryo relies on histotrophic nutrition.39, 42, 43 Maternal concentrations of estrogens rise markedly at about 35–40 days
as a result of ovarian production, because if mares are ovariectomized or lose their functional CL, levels decline.44, 45 Estrogen concentrations change little until a second sustained rise starting at about day 80, which signifies the major period of estrogen production by the feto-placental unit. Estrogens are present in high concentrations (0.1–2 µg/mL in maternal plasma, urine, and allantoic fluid for most of pregnancy.46–49 The classical estrogens are estrone, estradiol-17β, and estradiol17α, and their sulfoconjugates (e.g., estrone sulfate), which predominate. The biosynthetic pathway of these estrogens is shown in Figure 229.3. In addition, the mare is unique in producing large quantities of ring B unsaturated estrogens, namely equilin and equilenin, and their hydroxyderivatives, which are shown in Figure 229.3b. Estrone and equilin are the principal steroids, and all the other estrogens are largely derived through them by metabolism in the mare. The main estrogen precursor, DHEA, is produced in large quantities by the enlarged fetal gonads, which have the necessary enzymes, P450scc and 17α-OH, to convert cholesterol to DHEA.50–52 However, the gonadal tissue lacks 3β-HSD and aromatase, the key enzymes in the estrogen biosynthetic pathway (Figure 229.2). Consequently, the fetal gonads cannot synthesize estrogens and instead excrete high concentrations of DHEA into the umbilical artery.50, 53 However, abundant aromatase and 3β-HSD activities in the placenta enable aromatization of DHEA via the conventional 4–5 pathway into the phenolic estrogens, estrone and estradiol-17α and -17β.24, 50
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FETAL GONADS
Cholesterol P450 scc P5 17αOH 17αOH Pregnenolone 17, 20 lyase DHEA
PLACENTA
DHEA 3β HSD Androstenedione
Testosterone
Aromatase
17β HSD Oestrone O
Oestradiol-17β OH
HO
HO 17β HSD
MARE
Oestradiol-17β sulphate
Oestrone Sulphate (a)
FETAL GONADS
Acetate
3β–hydroxy-5,7-androstadien-17-one [7-dehydro-DHEA]
PLACENTA
Equilin OH 17α-Dihydroequilin
17β-Dihydroequilin
HO
17α-Dihydroequilenin
Equilenin
17β-Dihydroequilenin
(b) Figure 229.3 Estrogen biosynthetic pathways in the equine feto-placental unit for (a) the classical phenolic estrogens and (b) the ring B unsaturated estrogens. P5, pregnenolone; 17α-OH, 17α-hydroxylase; 3β-HSD, 3β-hydroxysteroid dehydrogenase; 17β-HSD, 17β-hydroxysteroid dehydrogenase; DHEA, dehydroepiandrosterone; P450scc , P450 side-chain cleavage enzyme.
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Endocrinology of Pregnancy The ring B estrogens are produced by an alternative pathway from 5,7-androstadiene-3,17-diol and 7-dehydro-DHEA, also secreted by the fetal gonads and likewise aromatized by the placenta into 17βdihydroequilin and 17β–equilenin.47, 54, 55 Estrogen production does not differ between mares carrying male versus female fetuses, and the enzymes necessary for estrogen expression are expressed equally in the fetal ovaries and testes.24, 49 Total estrogen concentrations in maternal blood increase from about 80 days of gestation, and remain high until the last 2–3 months, when they gradually decline. This profile reflects the supply of estrogen precursors, which increases and decreases concurrent with the hypertrophy and subsequent regression of the fetal gonads.21, 54 Peaks in the concentrations of the phenolic and ring B unsaturated estrogens occur at different stages of gestation, with estrone concentrations peaking at about 5–6 months and equilin peaking at about 7–8 months; these differences may reflect the different pathways and precursor availability for their synthesis.21, 46, 56 Alternatively, Haluska and Currie57 reported that the rise and fall in plasma estradiol-17β concentrations occurred during a specific calendar period and was related to an average ambient photoperiod of 13.4 h light/day, rather than to the stage of gestation, which varied widely in their experimental mares. This interesting observation may help explain the variability in estrogen profiles between pregnant mares, and it requires clarification. Estrogens are uterotonic agents and generally oppose the action of progesterone. High estrogen concentrations are associated with increased blood flow, both during pregnancy and the estrous cycle in the mare.12, 58 In other species, estrogens promote myometrial activity by favoring synthesis of contractile proteins in myometrial cells, enhancing electrical coupling and expression of oxytocin receptors, and promoting prostaglandin release.59–61 It is likely that estrogens have a similar role in the mare, because fetal gonadectomy in mid-gestation causes an immediate decline in total estrogen concentrations and the mares deliver undernourished foals at term following labor with weak uterine contractions and low prostaglandin concentrations, compared to shamoperated control mares.21 These important findings demonstrate that estrogens are essential for uterine blood supply and delivery of nutrients to the developing fetus, in addition to playing a key role in the physical processes of labor. Until recently it was difficult to reconcile this latter role with the clear evidence of a prepartum decline in estogens beginning 2–3 months before birth and reaching low levels by the end of gestation. However, more detailed investigations during the last week of pregnancy have shown
-18 to -13 days Log n (variance) Oestradiol-17B
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Figure 229.4 Variance in maternal plasma estradiol-17β concentrations over 24 h in pony mares during the last 3 weeks of gestation. Adapted from O’Donnell et al.64
that, although total estrogen concentrations are declining, pulsatile releases of estradiol-17β are raised at night time, associated with elevated uterine myometrial activity (Figure 229.4).57,62–64 In non-human primates, estrogens mediate increases in plasma oxytocin (OX) concentrations and the switch from lowamplitude, irregular myometrial contractures, which occur throughout pregnancy, to the high-amplitude, regular contractions of labor.65 These findings indicate that the periparturient endocrine changes in the mare may, in fact, be more comparable to that of other domestic animals by demonstrating the expected decline in P5 (progestagens) and rise in estradiol17β (estrogens) albeit on an abbreviated time scale.
Relaxin Relaxin is a hormone of pregnancy being produced primarily by the placenta, specifically the placental trophoblast cells.66 Maternal concentrations increase from around 80 days of gestation as the placenta grows and tend to remain high thereafter.67 A further rise occurs during labor, which is probably initiated by rising OX concentrations because relaxin concentrations also increase following exogenous OX administration at term.67, 68 Postpartum relaxin concentrations diminish gradually with delivery of the placenta, but will remain elevated if the placenta is retained. Substantial differences in maternal plasma relaxin levels exist between breeds, which appear to be unrelated to placental size but may reflect differences in physiology between the breeds.69 Relaxin is thought to contribute to myometrial relaxation but, paradoxically, during active labor relaxin concentrations increase substantially. It has been suggested by Haluska et al. that these high levels
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may contribute to the brief (2–4 hours) period of myometrial quiescence that occurs before delivery, by increasing intracellular cAMP (cyclic adenosine monophosphate) concentrations in myometrial smooth muscle thereby lowering intracellular Ca2+ and myosin light chain kinase activity.70 These authors also proposed that the high concentrations of prostaglandins achieved during labor are necessary to overcome the myometrial relaxatory effect of relaxin. The other major hormone produced by the placenta in other species (e.g., primates, rodents, and ruminants) is placental lactogen. Its main physiological action during pregnancy is to stimulate growth and terminal differentiation of the mammary tissue for milk synthesis.71 One study investigated placental lactogen in the mare and concluded that the mare’s placenta does not synthesize placental lactogen.72
Prostaglandins Prostaglandins constitute a large family of hormones, mainly produced from the fatty acid precursor, arachidonic acid. They are not stored in cells but synthesized on demand when the necessary synthetic enzymes and substrates become available. Consequently they are labile and difficult to quantify in peripheral plasma. The major prostaglandins during pregnancy and labor in the mare, in common with other species, are prostaglandin F2α (PGF2α ) and prostaglandin E2 (PGE2 ). In early pregnancy PGF2α is luteolytic whereas in later pregnancy it stimulates myometrial contractions by increasing intracellular calcium concentrations in the myocytes.2, 27 PGE2 promotes cervical ripening and is used clinically for relaxation of the mare’s cervix.73 Delivery may be induced in the pregnant mare by small, repeated intramuscular (i.m.) doses of prostaglandins (5–10 mg PGF2α ) or its analog, chloprostenol (250–500 µg) throughout pregnancy.1, 2, 74 Both PGF2α and PGE2 are produced by the UP tissues but concentrations in peripheral plasma usually remain low (1–2 ng/mL) for the majority of gestation; levels increase slightly toward term (2–4 ng/mL) but then increase rapidly during first-stage labor (20 ng/mL) to reach high concentrations (100 ng/mL) during the expulsive phase of labor, when they stimulate strong uterine contractions.57, 75, 76
Oxytocin Oxytocin (OX) is secreted by the posterior pituitary gland. It is difficult to measure accurately in peripheral plasma because it is released in a pulsatile
manner and rapidly metabolized (half-life < 1 min). However, collection of blood from the intercavernous sinus (ICS) draining the pituitary, allows recovery of blood containing high concentrations of OX before the pituitary effluent is diluted in the peripheral circulation.77 In humans and other species, OX is produced by other tissues including luteal, chorion, amnion, and decidua, where it binds with local oxytocin receptors and acts in a paracrine manner.78 OX stimulates uterine contractions by increasing intracellular calcium release from intracellular stores in the myometrium.79 In the pregnant mare, plasma OX concentrations remain low (< 11 pg/mL) throughout gestation and only increase during labor after rupture of the allantochorion, when ICS concentrations may exceed 6000 pg/mL.57, 80, 81 Peak levels occur during the period of maximum contractions and fetal expulsion, when OX release is stimulated by stretching of the cervix and vagina (Ferguson reflex). In other species, the increase in OX at parturition is associated with increased uterine OX receptor numbers and formation of gap junctions to allow cell-to-cell communication; these changes are initiated by a decline in progestagens and an increase in estrogens; moreover, exogenous PGs can also stimulate a rise in OX receptor numbers.79, 82 Although uterine OX receptors have not yet been measured in the late pregnant mare, it is likely that they are, in fact, present in high concentrations as suggested by the increased sensitivity of the uterus to low doses of OX at this time. The decline in progestagens and increase in estradiol concentrations in the mare at term, favors the induction of OX receptors, synthesis of OX, and increased uterine electromyographic activity.63, 64
Glucocorticoids Whereas fetal cortisol concentrations increase rapidly in late gestation (see Chapter 12), maternal glucocorticoid concentrations change little during pregnancy. In pregnant women glucocorticoids increase about seven-fold over the course of pregnancy.83 However, in unstressed, pregnant mares cortisol concentrations are low, and uterine glucocorticoid receptor concentrations also are unchanged throughout pregnancy.84–87 Others have reported an increase in maternal cortisol 48 h before foaling.88, 89 It is unlikely that fetal cortisol concentrations are altered by fluctuations in maternal cortisol because the placental enzyme, 11β-hydroxysteroid dehydrogenase Type 2, prevents transfer by inactivating cortisol to cortisone.89 However, in stressed mares, high maternal cortisol levels are correlated with elevated fetal cortisol concentrations.28 It is not clear
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Endocrinology of Pregnancy whether this occurs by direct cortisol transfer across the placenta or indirectly by stimulation of other glucocorticoid-sensitive pathways.
Hormones involved in lactatogenesis, nutrition, and fetal growth Prolactin and growth hormone (and placental lactogen in other species) are essential for the initiation of lactation in most mammalian species and, unless one of these is present, the mammary gland will not grow or produce milk.71 A number of other hormones including progestagens, estrogens, insulin, and adrenal steroids also play a role. Prolactin, together with estrogens and progestagens, is important for lobuloalveolar growth of the mammary gland, but high progestagen concentrations are thought to inhibit milk production. Prolactin is secreted by the secretory cells in the anterior pituitary and is inhibited by dopamine and endogenous opioids.90 In the mare, plasma prolactin concentrations are relatively low until the last 2 of weeks of pregnancy, when levels increase substantially accompanied by development of the mare’s mammary gland and onset of lactogenesis.48, 91, 92 The importance of prolactin for lactogenesis is demonstrated by the agalactia and low prolactin concentrations observed in pregnant mares exposed to dopamine agonist drugs or ergot alkaloids.93, 94 However, prolactin is not essential for continued lactation because inhibition of prolactin by bromocriptine during peak lactation does not alter milk yield or mammary gland size.95 Growth hormone (GH), or somatotropin, is released from the pituitary and controls growth and lactation. It also stimulates insulin-like growth factor 1 (IGF1) release from the liver after birth and, therefore, is linked to other hormones of the somatotropic axis, including insulin and leptin. In primates GH is also produced by the placenta, and placental GH concentrations are correlated with fetal weight at birth.96 In the mare, peripheral concentrations of GH are low during pregnancy compared to concentrations in non-pregnant mares, and change little over the periparturient period.91 These results suggest that maternal GH does not play a direct role in fetal growth and lactation in the mare but may mediate its effects through IGF1 or by antagonizing the actions of insulin. Insulin-like growth factors and insulin are involved in growth and lactation, and high concentrations of IGF1 and insulin are found in mares’ colostrum at birth. IGFs are a large family of related protein hormones including IGF-binding pro-
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teins. IGF1 is produced mainly by the liver but also derives from the ovaries, hypothalamus, and uterus, and is involved in nutrient utilization, growth, and regulation of reproductive mechanisms by binding to receptors in reproductive tissues. IGF2 is found in higher concentrations than IGF1 in fetal plasma, and both are involved in feto-placental growth.98, 99 Plasma IGF1 concentrations rise in the late-pregnant mare, peak in both colostrum and plasma at parturition, before declining thereafter.91, 97, 100 The high concentrations at parturition promote lactogenesis and stimulate postpartum ovarian activity, whilst declining levels postpartum may be a response to the excessive metabolic demands of lactation causing energy deficits, as occurs in lactating dairy cows.91, 101, 102 Ontogenic profiles for IGF2 have not yet been reported for the late-pregnant mare. Insulin is released by the pancreatic β-cells in response to circulating glucose concentrations and plays a key role in the partitioning of nutrients between the mare and fetus. Basal insulin concentrations in the mare are higher in early than in later gestation, when concentrations decline.103 In early gestation, there is enhanced pancreatic β-cell sensitivity to glucose and hyperinsulinemia, which allows the glucose requirements of both the fetus and mare to be met even following periods of feed deprivation. However, in the third trimester, maternal basal insulin concentrations are lower and β-cell sensitivity to changes in glucose concentrations is less, which allows glucose to be preferentially directed toward the uterus and fetus, which is gaining weight rapidly at this time.104, 105 These changes in insulin secretion and sensitivity in late gestation can make the pregnant mare prone to hypoglycemia, even when in a wellfed condition, and indicate that the mare’s rate of glucose utilization is minimal at this time compared to that of the fetus and utero-placental tissues.103 Moreover, periods of short-term feed deprivation increase endogenous prostaglandin metabolite (PGFM) concentrations and may increase the risk of the mare delivering prematurely under such conditions.75 Leptin is another key hormone involved in nutrition and the regulation of appetite. It provides the brain with information about the body’s energy status, so high leptin concentrations lower voluntary feed intake. It is produced mainly by adipocytes but leptin receptors have also been identified in equine testes, ovaries, and other tissues, with high concentrations in obese animals.106, 107 In many species, plasma leptin concentrations are elevated throughout pregnancy as a result of upregulation of maternal adipose tissue synthesis, stimulated by estrogen concentrations, with leptin production by the fetus
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and placenta also contributing toward the hyperleptinemia.108 During early pregnancy leptin may be important for embryo implantation, and in later gestation, for fetal growth. Information on leptin profiles throughout pregnancy are lacking in the mare but during the last few weeks of pregnancy, plasma leptin concentrations are not significantly greater than in non-pregnant mares, with concentrations in both plasma and milk gradually declining before and after parturition.91, 100 It is not clear whether the relatively low leptin levels in the pregnant mare are due to an absence of placental leptin or whether they correlate directly with declining total estrogen concentrations. However, it seems more likely that the low levels are related to decreasing maternal adiposity in late gestation as the mare provides nutrients for the growing fetus at the expense of her own body mass.109
Fetal hormones A chapter on endocrinology of pregnancy would not be complete without mention of the endocrinology of the fetus. It is clear from the above and elsewhere in this book that the conceptus possesses steroidogenic enzymes and capacity to produce androgens and estrogens from as early as day 10.16 However, the steroids produced by the young conceptus/fetus are predominantly utilized within the uterine tissues and are not released in large volumes into the maternal circulation until the second and third trimesters, when the fetus excretes high concentrations of progestagen and estrogen precursors for metabolism by the placenta. This period coincides with hypertrophy of the steroidegenic tissue in the fetal adrenals and gonads.30, 110 As the fetal organs acquire endocrine cells and develop their secretory capacity, the organ weights increase, reaching a peak at different gestational ages depending on the period of major endocrine activity.105 The fetus also contributes to its growth and nutrition by maximizng its use of maternal nutrients through increasing activity of fetal somatotropic hormones. For example, fetal insulin secretion starts at about 100 days of gestation, which enables the fetus to utilize maternally derived glucose, particularly when there is post-prandial excess.103 Fetal triiodothyronine (T3 ) and thyroxine (T4 ) concentrations begin gradually to rise from 3 to 5 months of gestation as the fetal tissues become more metabolically active.111 In human fetuses, GH from the fetal pituitary is detected from 10 weeks of gestation, and numerous studies in other species have demonstrated the importance of fetal IGF1 and IGF2 in directing maternal nutrient transfer for fetal growth.96, 98 Ontogenic profiles for many of these fetal hormones remain to be elucidated in
the horse. There is major upregulation of fetal hormones toward parturition, particularly those of the hypothalamic–pituitary–adrenal (HPA) axis, associated with functional maturation of the fetal organs and preparation for life ex utero. The surge in equine fetal cortisol following the rise in fetal ACTH and some other prenatal hormone changes are described in other chapters (see Chapters 7 and 12). However, due to the difficulties in obtaining blood samples from the equine fetus during gestation, there is little information about the ontogeny of equine fetal endocrinology compared to that reported for other domestic and experimental animals. Hopefully this balance will be redressed in the near future.
References 1. Daels PF, Besognet B, Hansen B, Mohammed H, Odensvik K, Kindahl H. Effect of progesterone on prostaglandin F2 alpha secretion and outcome of pregnancy during cloprostenol-induced abortion in mares. Am J Vet Res 1996;57:1331–7. 2. Leadon DP, Rossdale PD, Jeffcott LB, Allen WR. A comparison of agents for inducing parturition in mares in the pre-viable and premature periods of gestation. J Reprod Fert Suppl 1982;32:597–602. 3. Holtan DW, Nett TM, Estergreen VL. Plasma progestins in pregnant, postpartum and cycling mares. J Anim Sci 1975;40:251–60. 4. Squires EL, Douglas RH, Steffenhagen WP, Ginther OJ. Ovarian changes during the estrous cycle and pregnancy in mares. J Anim Sci 1974;38:330–8. 5. Allen WR. Luteal deficiency and embryo mortality in the mare. Reprod Domest Anim 2001;36:121–31. 6. Ginther OJ. Reproductive Biology of the Mare, 2nd edn. Cross Plains, WI: Equiservices, 1992. 7. Holtan DW, Houghton E, Silver M, Fowden AL, Ousey J, Rossdale PD. Plasma progestagens in the mare, fetus and newborn foal. J Reprod Fert Suppl 1991;44:517– 28. 8. Short RV. Progesterone in blood: IV. Progesterone in the blood of mares. J Endocrinol 1959;19:207–10. 9. Allen WR, Hamilton DW, Moor RM. The origin of equine endometrial cups. II. Invasion of the endometrium by trophoblast. Anat Rec 1973;177:485–501. 10. Samuel CA, Allen WR, Steven DH. Studies on the equine placenta I. Development of the micro-cotyledons. J Reprod Fert 1974;41:441–5. 11. Abd-Elnaeim MM, Leiser R, Wilsher S, Allen WR. Structural and haemovascular aspects of placental growth throughout gestation in young and aged mares. Placenta 2006;27:1103–13. 12. Bollwein H, Weber F, Woschee I, Stolla R. Transrectal Doppler sonography of uterine and umbilical blood flow during pregnancy in mares. Theriogenology 2004;61:499–509. 13. Samuel CA, Allen WR, Steven DH. Ultrastructural development of the equine placenta. J Reprod Fert Suppl 1975;4:575–8. 14. Silver M. Some aspects of equine placental exchange and foetal physiology. Equine Vet J 1984;16:227– 33.
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Endocrinology of Pregnancy 15. Holtan DW, Squires EL, Lapin DR, Ginther OJ. Effect of ovariectomy on pregnancy in mares. J Reprod Fert Suppl 1979;27:457–63. 16. Flood PF, Betteridge KJ, Irvine DS. Oestrogens and androgens in blastocoelic fluid and cultures of cells from equine conceptuses of 10–22 days gestation. J Reprod Fert Suppl 1979;27:413–20. 17. Paulo, E, Tischner M. Activity of delta(5)3betahydroxysteroid dehydrogenase and steroid hormones content in early preimplantation horse embryos. Folia Histochem Cytobiol 1985;23:81–4. 18. Chavatte P, Holtan D, Ousey JC, Rossdale PD. Biosynthesis and possible biological roles of progestagens during equine pregnancy and in the newborn foal. Equine Vet J Suppl 1997;89–95. 19. Ousey JC. Peripartal endocrinology in the mare and foetus. Reprod Domest Anim 2004;39:222–31. 20. Han X, Rossdale PD, Ousey J, Holdstock N, Allen WR, Silver M, Fowden AL, McGladdery AJ, Labrie F, Belanger A, et al. Localisation of 15-hydroxy prostaglandin dehydrogenase (PGDH) and steroidogenic enzymes in the equine placenta. Equine Vet J 1995;27:334–9. 21. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J Reprod Fert Suppl 1979;27:499–509. 22. Rossdale PD, McGladdery AJ, Ousey JC, Holdstock N, Grainger L, Houghton E. Increase in plasma progestagen concentrations in the mare after foetal injection with CRH, ACTH or betamethasone in late gestation. Equine Vet J 1992;24:347–50. 23. Hamon M, Clarke SW, Houghton E, Fowden AL, Silver M, Rossdale PD, Ousey JC, Heap RB. Production of 5 alpha-dihydroprogesterone during late pregnancy in the mare. J Reprod Fert Suppl 1991;44:529–35. 24. Hasegawa T, Sato F, Nambo Y, Ishida N. Expression of steroidogenic enzyme genes in the equine feto-placental unit. J Equine Sci 2001;12:25–32. 25. Ousey JC, Forhead AJ, Rossdale PD, Grainger L, Houghton E, Fowden AL. Ontogeny of uteroplacental progestagen production in pregnant mares during the second half of gestation. Biol Reprod 2003;69:540–8. 26. Fowden AL, Ousey J, Forhead AJ, Rossdale PD, Grainger L, Houghton E. Uteroplacental production of 5αpregnane-3,20-dione (5α-DHP) in pregnant mares. Theriogenology 2002;58:821–4. 27. Challis JRG, Matthews SG, Gibb W, Lye SJ. Endocrine and paracrine regulation of birth at term and preterm. Endocr Rev 2000;21:514–50. 28. Fowden AL, Forhead AJ, Ousey JC. The endocrinology of equine parturition. Exp Clin Endocrinol Diabetes 2008;116:393–403. 29. Macdonald AA, Chavatte P, Fowden AL. Scanning electron microscopy of the microcotyledonary placenta of the horse (Equus caballus) in the latter half of gestation. Placenta 2000;21:565–74. 30. Yamauchi S. Histological development of the equine fetal adrenal gland. J Reprod Fert Suppl 1979;27:487–91. 31. Allen WR, Wilsher S, Stewart F, Stewart F, Ousey J, Ousey J, Fowden A. The influence of maternal size on placental, fetal and postnatal growth in the horse. II. Endocrinology of pregnancy. J Endocrinol 2002;172;237– 46. 32. Rossdale PD, Ousey JC, Cottrill CM, Chavatte P, Allen WR, McGladdery AJ. Effects of placental pathology on
33.
34.
35.
36.
37.
38.
39.
40.
41. 42.
43.
44.
45.
46. 47.
48.
49.
50.
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maternal plasma progestagen and mammary secretion calcium concentrations and on neonatal adrenocortical function in the horse. J Reprod Fert Suppl 1991;44:579–90. Fowden A, Silver M. Comparative development of the pituitary-adrenal axis in the fetal foal and lamb. Reprod Dom Anim 1995;30:170–7. Chavatte P, Rossdale PD, Tait AD. Modulation of 3β-hydroxysteroid dehydrogenase (3β-HSD) activity in the equine placenta by pregnenolone and progesterone metabolites. Equine Vet J 1995;27:342–7. Fowden AL, Silver M. Effect of inhibiting 3bhydroxysteroid dehydrogenase on plasma progesterone and other steroids in the pregnant mare near term. J Reprod Fert Suppl 1987;35:539–45. Schutzer WE, Kerby JL, Holtan DW. Differential effect of trilostane on the progestin milieu in the pregnant mare. J Reprod Fert 1996;107:241–8. Houghton E, Holtan D, Grainger L, Voller BE, Rossdale PD, Ousey JC. Plasma progestagen concentrations in the normal and dysmature newborn foal. J Reprod Fert Suppl 1991;44:609–17. Zavy JT, Mayer R, Vernon MW, Bazer FW, Sharp DC. An investigation of the uterine luminal environment of nonpregnant and pregnant pony mares. J Reprod Fert Suppl 1979;27:403–11. Raeside JI, Christie HL, Renaud RL, Waelchli RO, Betteridge KJ. Estrogen metabolism in the equine conceptus and endometrium during early pregnancy in relation to estrogen concentrations in yolk-sac fluid. Biol Reprod 2004;71:1120–7. Choi SJ, Anderson GB, Roser JF. Production of free estrogens and estrogen conjugates by the preimplantation equine embryo. Theriogenology 1997;47:457–66. Terqui M, Palmer E. Oestrogen pattern during early pregnancy in the mare. J Reprod Fert Suppl 1979;27:441–6. Heap RB, Hamon M, Allen WR. Studies on oestrogen synthesis by the preimplantation equine conceptus. J Reprod Fert Suppl 1982;32:343–52. Spencer TE, Bazer FW. Uterine and placental factors regulating conceptus growth in domestic animals. J Anim Sci 2004;82(E-Suppl.):E4–13. Daels PF, DeMoraes JJ, Stabenfeldt GH, Hughes JP, Lasley BL. The corpus luteum: source of oestrogen during early pregnancy in the mare. J Reprod Fert Suppl 1991;44:501– 8. Palmer E, Jousset B. Urinary oestrogen and plasma progesterone levels in non-pregnant mares. J Reprod Fert Suppl 1975;213–21. Cox JE. Oestrone and equilin in the plasma of the pregnant mare. J Reprod Fert Suppl 1975;23:463–8. Foster SJ, Marshall DE, Houghton E, Gower DB. Investigations into the biosynthetic pathways for classical and ring B-unsaturated oestrogens in equine placental preparations and allantochorionic tissues. J Steroid Biochem Mol Biol 2002;82:401–11. Nett TM, Holtan DW, Estergreen VL. Oestrogens, LH, PMSG, and prolactin in serum of pregnant mares. J Reprod Fert Suppl 1975;457–62. Raeside JI, Liptrap RM. Patterns of urinary oestrogen excretion in individual pregnant mares. J Reprod Fert Suppl 1975;23:649–75. Pashen RL, Sheldrick EL, Allen WR, Flint AP. Dehydroepiandrosterone synthesis by the fetal foal and its importance as an oestrogen precursor. J Reprod Fert Suppl 1982;23:32:389–97.
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51. Raeside JI, Gofton N, Liptrap RM, Milne FJ. Isolation and identification of steroids from gonadal vein blood of the fetal horse. J Reprod Fert Suppl 1982;32:383–7. 52. Raeside JI, Liptrap RM, McDonell WN, Milne FJ. A precursor role for DHA in a feto-placental unit for oestrogen formation in the mare. J Reprod Fert Suppl 1979;27:493–7. 53. Marshall DE, Gower DB, Silver M, Fowden A, Houghton E. Cannulation in situ of equine umbilicus. Identification by gas chromatography-mass spectrometry (GC-MS) of differences in steroid content between arterial and venous supplies to and from the placental surface. J Steroid Biochem Mol Biol 1999;68:219–28. 54. Raeside JI, Renaud RL, Christie HL. Postnatal decline in gonadal secretion of dehydroepiandrosterone and 3 betahydroxyandrosta-5,7-dien-17-one in the newborn foal. J Endocrinol 1997;155:277–82. 55. Tait AD, Santikarn S, Allen WR. Identification of 3 betahydroxy-5,7-pregnadien-20-one and 3 beta-hydroxy-5,7androstadien-17-one as endogenous steroids in the fetal horse gonad. J Endocrinol 1983;99:87–92. 56. Rance TA, Park BK. The measurement of oestrone, equilin and dehydroepiandrosterone in the peripheral plasma of pregnant pony mares by radioimmunoassay. J Steroid Biochem 1978;9:1065–9. 57. Haluska GJ, Currie WB. Variation in plasma concentrations of oestradiol-17 beta and their relationship to those of progesterone, 13,14-dihydro-15-keto-prostaglandin F-2 alpha and oxytocin across pregnancy and at parturition in pony mares. J Reprod Fert 1988;84:635–46. 58. Bollwein H, Mayer R, Weber F, Stolla R. Luteal blood flow during the estrous cycle in mares. Theriogenology 2002;57:2043–51. 59. Challis JR, Sloboda DM, Alfaidy N, Lye SJ, Gibb W, Patel FA, Whittle WL, Newnham JP. Prostaglandins and mechanisms of preterm birth. Reproduction 2002;124:1–17. 60. Egarter CH, Husslein P. Biochemistry of myometrial contractility. Bailli`ere’s Clin Obstet Gynaecol 1992;6:755–69. 61. Leung ST, Wathes DC, Young IR, Jenkin G. Effect of labor induction on the expression of oxytocin receptor, cytochrome P450 aromatase, and estradiol receptor in the reproductive tract of the late-pregnant ewe. Biol Reprod 1999;60:814–20. 62. Barnes RJ, Nathanielsz PW, Rossdale PD, Comline RS, Silver M. Plasma progestagens and oestrogens in fetus and mother in late pregnancy. J Reprod Fert Suppl 1975;23:617–23. 63. McGlothlin JA, Lester GD, Hansen PJ, Thomas M, Pablo L, Hawkins DL, LeBlanc MM. Alteration in uterine contractility in mares with experimentally induced placentitis. Reproduction 2004;127:57–66. 64. O’Donnell LJ, Sheerin BR, Hendry JM, Thatcher MJ, Thatcher WW, LeBlanc MM. 24-hour secretion patterns of plasma oestradiol 17beta in pony mares in late gestation. Reprod Domest Anim 2003;38:233–5. 65. Nathanielsz PW, Giussani DA, Mecenas CA, Wu W, Winter JA, Garcia-Villar R, Baguma-Nibasheka M, Honnebier MB, McDonald TJ. Regulation of the switch from myometrial contractures to contractions in late pregnancy: studies in the pregnant sheep and monkey. Reprod Fertil Dev 1995;7:595–602. 66. Klonisch T, Hombach-Klonisch S. Review: relaxin expression at the feto-maternal interface. Reprod Domest Anim 2000;35:149–52. 67. Stewart DR, Kindahl H, Stabenfeldt GH, Hughes JP. Concentrations of 15-keto-13,14-dihydro-prostaglandin F2 al-
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pha in the mare during spontaneous and oxytocin induced foaling. Equine Vet J 1984;16:270–4. Thorburn GD. A speculative review of parturition in the mare. Equine Vet J Suppl 1993;14:41–9. Stewart DR, Addiego LA, Pascoe DR, Haluska GJ, Pashen R. Breed differences in circulating equine relaxin. Biol Reprod 1992;46:648–52. Haluska GJ, Lowe JE, Currie WB. Electromyographic properties of the myometrium correlated with the endocrinology of the pre-partum and post-partum periods and parturition in pony mares. J Reprod Fert Suppl 1987;35:553–64. Cowie A, Forsyth I, Hart I. Hormonal Control of Lactation. New York: Springer-Verlag, 1980. Forsyth IA, Rossdale PD, Thomas CR. Studies on milk composition and lactogenic hormones in the mare. J Reprod Fert Suppl 1975;23:631–5. Rigby S, Love C, Carpenter K, Varner D, Blanchard T. Use of prostaglandin E2 to ripen the cervix of the mare prior to induction of parturition. Theriogenology 1998;50:897– 904. Ousey JC, Dudan FE, Rossdale PD, Silver M. Effects of fluprostenol administration in mares during late pregnancy. Equine Vet J 1984;16:264–9. Fowden AL, Ralph MM, Silver M. Nutritional regulation of uteroplacental prostaglandin production and metabolism in pregnant ewes and mares during late gestation. Exp Clin Endocrinol 1994;102:212–21. Silver M, Barnes RJ, Comline RS, Fowden AL, Clover L, Mitchell MD. Prostaglandins in maternal and fetal plasma and in allantoic fluid during the second half of gestation in the mare. J Reprod Fert Suppl 1979;27:531–9. Irvine CH, Alexander SL. A novel technique for measuring hypothalamic and pituitary hormone secretion rates from collection of pituitary venous effluent in the normal horse. J Endocrinol 1987;113:183–92. Mitchell BF, Fang X, Wong S. Oxytocin: a paracrine hormone in the regulation of parturition? Rev Reprod 1998;3:113–22. Fuchs AR, Fuchs F, Husslein P, Soloff MS, Fernstrom MJ. Oxytocin receptors and human parturition: a dual role for oxytocin in the initiation of labor. Science 1982;215: 1396–8. Vivrette SL, Kindahl H, Munro CJ, Roser J, Stabenfeldt GH. Effects of flunixin meglumine on pituitary effluent oxytocin, arginine vasopressin and 15-ketodihydroprostaglandin F2a concentrations and clinical parturient events during oxytocin-induced parturition in mares. Biol Reprod Monogr 1995;1:69–75. Vivrette SL, Kindahl H, Munro CJ, Roser JF, Stabenfeldt GH. Oxytocin release and its relationship to dihydro-15keto PGF2alpha and arginine vasopressin release during parturition and to suckling in postpartum mares. J Reprod Fert 2000;119:347–57. Garfield RE, Kannan MS, Daniel EE. Gap junction formation in myometrium: control by estrogens, progesterone, and prostaglandins. Am J Physiol 1980;238:C81–9. Dorr HG, Heller A, Versmold HT, Sippell WG, Herrmann M, Bidlingmaier F, Knorr, D. Longitudinal study of progestins, mineralocorticoids, and glucocorticoids throughout human pregnancy. J Clin Endocrinol Metab 1989; 68:863–8. Chavatte Palmer P, Duchamp G, Palmer E, Ousey JC, Rossdale PD, Lombes M. Progesterone, oestrogen and glucocorticoid receptors in the uterus and mammary
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glands of mares from mid- to late-gestation. J Reprod Fert Suppl 2000; 56:661–72. Hoffmann B, Gentz F, Failing K. Investigations into the course of progesterone-, oestrogen-and eCGconcentrations during normal and impaired pregnancy in the mare. Reprod Dom Anim 1996;31:717–23. Lovell JD, Stabenfeldt GH, Hughes JP, Evans JW. Endocrine patterns of the mare at term. J Reprod Fert Suppl 1975; 23:449–56. Silver M, Fowden AL. Prepartum adrenocortical maturation in the fetal foal: responses to ACTH. J Endocrinol 1994; 142:417–25. Fleeger JL, Harms PG, Dunn EL, Atkins DT. Levels of deoxycorticosterone and 21-hydroxy-5 alpha-pregnane3,20-dione in the peripheral circulation of the prepartum and postpartum mare. Biol Reprod 1979;21:433–7. Nathanielsz PW, Rossdale PD, Silver M, Comline RS. Studies on fetal, neonatal and maternal cortisol metabolism in the mare. J Reprod Fert Suppl 1975; 23:625–30. Aurich C, Gerlach T, Aurich JE, Hoppen HO, Lange J, Parvizi N. Dopaminergic and opioidergic regulation of gonadotropin and prolactin release in stallions. Reprod Domest Anim 2002;37:335–40. Heidler B, Parvizi N, Sauerwein H, Bruckmaier RM, Heintges U, Aurich JE, Aurich C. Effects of lactation on metabolic and reproductive hormones in Lipizzaner mares. Domest Anim Endocrinol 2003;25:47–59. Worthy K, Escreet R, Renton JP, Eckersall PD, Douglas TA, Flint DJ. Plasma prolactin concentrations and cyclic activity in pony mares during parturition and early lactation. J Reprod Fert 1986;77:569–74. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995;73:899–908. Ireland FA, Loch WE, Worthy K, Anthony RV. Effects of bromocriptine and perphenazine on prolactin and progesterone concentrations in pregnant pony mares during late gestation. J Reprod Fert 1991;92:179–86. Neuschaefer A, Bracher V, Allen WR. Prolactin secretion in lactating mares before and after treatment with bromocriptine. J Reprod Fert Suppl 1991;44:551–9. Mullis PE, Tonella P. Regulation of fetal growth: consequences and impact of being born small. Best Pract Res Clin Endocrinol Metab 2008;22:173–90. Hess-Dudan F, Vacher PY, Bruckmaier RM, Weishaupt MA, Burger D, Blum JW. Immunoreactive insulin-like growth factor I and insulin in blood plasma and milk of mares and in blood plasma of foals. Equine Vet J 1994;26:134–9.
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98. Fowden AL. The insulin-like growth factors and fetoplacental growth. Placenta 2003;24:803–12. 99. Velazquez MA, Spicer LJ, Wathes DC. The role of endocrine insulin-like growth factor-I (IGF-I) in female bovine reproduction. Domest Anim Endocrinol 2008;35: 325–42. 100. Berg EL, McNamara DL, Keisler DH. Endocrine profiles of periparturient mares and their foals. J Anim Sci 2007;85:1660–8. 101. Lucy MC. Functional differences in the growth hormone and insulin-like growth factor axis in cattle and pigs: implications for post-partum nutrition and reproduction. Reprod Domest Anim 2008;43(Suppl. 2): 31–9. 102. Sticker LS, Thompson DL Jr, Fernandez JM, Bunting LD, DePew CL. Dietary protein and(or) energy restriction in mares: plasma growth hormone, IGF-I, prolactin, cortisol, and thyroid hormone responses to feeding, glucose, and epinephrine. J Anim Sci 1995;73:1424–32. 103. Fowden AL, Comline RS, Silver M. Insulin secretion and carbohydrate metabolism during pregnancy in the mare. Equine Vet J 1984;16:239–46. 104. Fowden AL. Comparative aspects of fetal carbohydrate metabolism. Equine Vet J Suppl 1997;24:19–25. 105. Platt H. Growth of the equine foetus. Equine Vet J 1984;16(4):247–52. 106. Buff PR, Dodds AC, Morrison CD, Whitley NC, McFadin EL, Daniel JA, Djiane J, Keisler DH. Leptin in horses: tissue localization and relationship between peripheral concentrations of leptin and body condition. J Anim Sci 2002;80:2942–8. 107. Frank N, Elliott SB, Brandt LE, Keisler DH. Physical characteristics, blood hormone concentrations, and plasma lipid concentrations in obese horses with insulin resistance. J Am Vet Med Assoc 2006;228:1383–90. 108. Henson MC, Castracane VD. Leptin in pregnancy: an update. Biol Reprod 2006;74:218–29. 109. Powell D, Lawrence L, Parrett D, DiPietro J. Body composition changes in broodmares. In: Proceedings of the 11th Equine Nutrition and Physiology Symposium, 1989, pp. 91–4. 110. Cole H, Hart G, Lyons W, Catchpole H. The development and hormonal content of fetal horse gonads. Anat Rec 1933;56:275–93. 111. Allen AL, Doige CE, Haas SD, Card CE, Fretz PB. Concentrations of triiodothyronine and thyroxine in equine fetal serum during gestation. Biol Reprod Monogr 1995;1: 49–52. 112. Ousey JC. Veterinary Clinics of North America. Advances in Reproduction 2006;22(3):727–47.
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Chapter 230
Development and Morphology of the Placenta Sandra Wilsher and W.R. (Twink) Allen
Introduction By any standards, embryogenesis and placentation in the horse is a slow and steady process that involves a number of interesting and biologically puzzling features. At the very outset, only the embryo, but not an unfertilized oocyte,1 passes all the way down the oviduct to reach the uterus as late as 6–6.5 days after ovulation.2 The embryo achieves this selective passage by secreting, from as early as day 4, appreciable quantities of prostaglandin E2 (PGE2 ) onto the oviductal mucosa;3 the PGE2 binds strongly to and relaxes the circular smooth muscle fibers of the oviduct wall,5 thereby allowing the embryo and any unfertilized oocytes from previous ovulations to pass on down and into the uterus.6, 7 Soon after entering the uterus on day 6–6.5, and before the embryo hatches from its zona pellucida, the trophectoderm cells begin releasing appreciable quantities of a glycoprotein-rich secretion, which, due to the continuing presence of the zona pellucida, is molded into a transparent, thin, very elastic glycocalyx, which completely envelops the embryo from then until around day 23 after ovulation.8–11 This uniquely equine blastocyst capsule has two clear functions and probably many more that are not yet fully appreciated. First, its tough and elastic physical properties maintain the spherical shape of the equine conceptus and prevent elongation and remodeling of the trophoblast, which commences around days 10–12 after ovulation in the conceptuses of pigs and ruminants.12 In addition, the elasticity of the capsule gives the conceptus the strength to withstand the strong myometrial contractions that propel it constantly throughout the uterine lumen between days 7 and 16 after ovulation,13, 14 presumably to release its antiluteolytic maternal recognition of pregnancy
signal onto sufficient area of endometrium to achieve the necessary luteostasis of pregnancy.15, 16 Second, the strongly negative electrostatic charge exhibited by the capsule makes it ‘sticky’ and hence able to attract and adhere the positively charged protein-rich endometrial gland secretions (‘uterine milk’) upon which the free-living equine conceptus must survive exclusively during the first 40 days of its existence.12 The mechanisms involved in the passage of histotrophe through the capsule and onto the underlying trophectoderm for imbibition by the embryo are only partly elucidated,17–20 but it is already clear that the capsule is essential to the survival of the conceptus during this early mobile phase of its development. Physical removal of the capsule from day-8 embryos results in their death when transferred back to recipient mares, whereas equivalent mechanical ‘mishandling’ of similar age embryos in vitro before re-transfer to recipients results in normal pregnancy rates.21, 22 From day 17 of gestation the still spherical conceptus is held stationary at the base of one or other of the uterine horns by the striking increase in myometrial tonicity that occurs from around this time.23, 24 The blastocyst capsule begins to fragment and disappear from days 21–23,11 presumably as a consequence of the actions of glycolytic enzymes secreted by the endometrial luminal and/or glandular epithelia or even the trophoblast itself.9 From day 21 or thereabouts the allantois appears as an outpouching of the embryonic hindgut, and it grows rapidly over the next 20 days, fusing with the outermost chorion to form the definitive allantochorion and progressively regressing the yolk sac.25 Between days 25 and 35 of gestation a discrete annulate portion of the trophoblast at the junction of the enlarging allantois and regressing yolk sac membranes forms a series of simple folds.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Development and Morphology of the Placenta The trophoblast cells at the tips of each fold multiply extremely rapidly to form simple glandlike structures, which then become increasingly compressed by the typical pressure created by the continuation of myometrial tone and conceptus expansion to form a pale, thickened band of hyperplastic specialized trophoblast cells known as the chorionic girdle.23, 26, 27 Then, around day 35–38 of gestation the entire chorionic girdle simply peels off the fetal membranes and becomes attached to the underlying luminal epithelium of the endometrium by an exocrine secretory material secreted by the simple glandlike structures that comprise the girdle.28 This enables the now binucleated29 and highly invasive30 girdle cells to penetrate the luminal epithelium and progress down the lumens of the endometrial glands to penetrate the basement membrane and invade the endometrial stroma.28 Here, the girdle cells soon become sessile, round up, and begin to secrete equine chorionic gonadotropin (eCG); the girdle cells enlarge greatly, thereby becoming tightly packed in the stroma to form the endometrial protuberances known as endometrial cups.31–33 The structure and biological properties of their curious gonadotropic product, eCG, are described in more detail in Chapter 172. Until day 40 after ovulation, the non-invasive trophoblast of the equine allantochorion lies in simple apposition but not intimately associated with the luminal epithelium of the endometrium, but after this time it at last forms a microvillous attachment between the maternal and fetal epithelial layers and true allantochorionic placentation finally commences.34, 35
Development of the microcotyledons Around day 40 the chorion establishes a microvillous attachment to the luminal epithelium of the endometrium, and blunt primary villi of allantochorion begin to interdigitate with corresponding endometrial upgrowths (sulci) from the surface of the endometrium. As gestation advances several adjacent primary villi may coalesce to form one microcotyledon with a common stem,34 or a single primary villus may become multibranched to achieve the same result.12, 36 Continued secondary and tertiary branching of the chorionic villi and the corresponding endometrial sulci leads to the increasingly complex structure of the mature microcotyledon.34, 35 The microcotyledonary architecture maximizes the surface area contact between maternal and fetal epithelial layers without a breakdown of either tissue, and the microcotyledons development is essentially complete by mid-gestation when they exhibit a diameter of 1 to 2 mm. Nevertheless, further remodeling of each mi-
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Figure 230.1 A 250-day fetus enveloped in the allantochorion. By this stage of gestation the allantochorion occupies the entire uterus. Development of the microcotyledons, although essentially complete at this stage, will continue until term. Note the areas devoid of microcotyledons, i.e., at the cervical star (overlying the fetal forehead) and the scarlike areas that overlie where the endometrial cups were formed (overlying the fetal hindquarters).
crocotyledon continues until term, with a progressive lengthening and sub-branching of the villi in the last 2 months of gestation.37 In the healthy allantochorion the only areas devoid of microcotyledons are the avillous scarlike areas that overlie the endometrial cups,32 the cervical star adjacent to the internal os of the cervix,12 and the small circular area apposed to the utero-tubal papilla at the tip of each uterine horn (Figure 230.1). This indicates that, for microcotyledon formation to occur, locally acting stimulatory factors are required from the opposing glandular endometrial surface.38, 39 Indeed, throughout gestation, the endometrial glands remain open to the surface between adjacent microcotyledons.34 They continue to exude their protein-rich histotroph, which is taken up by groups of specialized tall, columnar, pseudostratified trophoblast cells known as areolae12, 40 (Figure 230.2). In addition to the gross changes occurring in the enlarging microcotyledons, other microscopic modifications increase the area of placental exchange and bring the fetal and maternal circulations into the closest possible proximity. For example, by day 300, the maternal luminal epithelium has thinned to only onethird of its original height, concomitantly with progressive indentation of the trophoblast layer by fetal capillaries to reduce the diffusion distance between maternal and fetal circulations to as little as 3 µm.35
Cellular proliferation of the equine placenta While the precise nature of the stimuli that drive the astounding growth and architectural remodeling of the allantochorion and the endometrium throughout
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Figure 230.2 A scanning electron micrograph of the surface of the endometrium at 309 days of gestation. The allantochorion has been peeled away leaving the maternal crypts visible, into which the microcotyledons were held. Between these raised crypts the mouths of the endometrial glands open, releasing their histotrophic nutrition throughout gestation.
gestation remains unknown, steroid hormones and/ or locally produced growth factors are the most likely candidates for this mitogenic impetus. Immunohistochemical localization of the proliferation marker Ki67 in placental and endometrial samples collected from mares between days 18 and 250 of gestation showed that, even as early as day 18, proliferation is apparent in the superficial strata of the endometrium and this effect can be mimicked by supplementing non-pregnant mares with estradiol during the last 6 days of a prolonged period of exogenous progesterone administration.41 Furthermore, the specialized trophoblast cells of the chorionic girdle proliferate vigorously during days 30–34 of gestation but cease to divide when they invade the endometrium to form the endometrial cups,28 while the non-invasive trophoblast cells of the allantochorion show an increase in mitotic activity after day 38. Interestingly, the luminal epithelium of the endometrium starts to proliferate only after the primary allantochorionic villi have formed, and this proliferation occurs only in the uterine horn into which the placenta is expanding.41 This localized effect implies that the presence of the allantochorion is essential for the remodeling of the underlying endometrium. During the second half of gestation proliferation becomes confined to the periphery of the microcotyledons as they continue to grow to meet the increasing nutritional demands of fetal growth.42
Growth factors associated with placentation Epidermal growth factor (EGF) was one of the first growth factors identified to be associated with pla-
centation in the mare, when Stewart et al.43 demonstrated a dramatic upregulation in EGF mRNA in the epithelium of the apical portions of the endometrial glands around day 35 of gestation. Further studies from the same laboratory then demonstrated that prolonged exposure of anestrous or ovariectomized mares to high circulating levels of progesterone for 35 days produced the same upregulation of endometrial EGF mRNA, with little or no synergistic effect of concomitant estrogens.44 Three other growth factors have been investigated during early pregnancy in the horse, namely, insulinlike growth factor 2 (IGF2), hepatocyte growth factorscatter factor (HGF-SF), and transforming growth factor β1 (TGFβ1). The first of these, IGF2, is known, from work in other species, to play an important role in feto-placental growth through its metabolic, mitogenic, and differentiative actions.45 In the horse, IGF2 mRNA becomes detectable in the trophoblast layer of the allantochorion soon after day 33 and persists at high concentrations in the allantochorion and other fetal tissues throughout the development of the placenta, while remaining undetectable in the maternal endometrium.46, 47 HGF-SF mRNA expression is limited to just the allantoic mesoderm while its receptor, c-met, is expressed in the trophoblast, with particularly high-intensity expression levels in the specialized invasive trophoblast cells of the chorionic girdle.48, 49 TGFβ1 mRNA appears in the luminal and glandular epithelia of the endometrium from around day 30 of gestation,50 with the suggestion that it may control the differentiation and/or invasiveness of the chorionic girdle cells, and play a significant role in remodeling both fetal and maternal tissues during placentation. More recently, Arai et al.51 described the expression of IGF-1 and activin in the equine placenta during the latter two-thirds of pregnancy and they noted the possible role of both factors in reg¨ ulating placental development. Finally, Grunig and Antczak52 proposed that cytokines, such as tumor necrosis factor α (TNFα) may also play an important role in regulating placental growth.
Measurement of feto-maternal contact and placental efficiency The gross area of the allantochorion and its macroscopic and microscopic architecture have been positively correlated with foal birthweight.38,53–55 At the microscopic level, stereology56 can be employed to further assess such relationships, because the technique allows derivation of three-dimensional parameters from two-dimensional histological sections providing strict criteria have been applied to the collection of the samples. For the equine placenta
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Figure 230.3 A scanning electron micrograph of the microcotyledonary villi overlain with a diagrammatic representation of surface density. Calculations are performed on twodimensional histology slides and result in a three-dimensional parameter, i.e., the surface area of the microcotyledons per unit volume of the chorion. Reprinted with permission from Wilsher and Allen.60
stereological measurements allow calculation of the surface area of contact within the microcotyledons, termed surface density (Figure 230.3) and, by further calculation, the total microscopic area of fetomaternal contact at the placental interface.55 Hence, the method can be employed to enumerate changes both in the surface area of the microcotyledons and in the total feto-maternal contact across the placental interface under different conditions. In addition, calculation of the weight of the foal at birth per square meter of microscopic feto-maternal contact gives an insight into placental efficiency and allows investigation of the different strategies the placenta may use to adapt itself to adverse or advantageous circumstances.
Influences of maternal size and genotype Maternal size, and hence uterine size, profoundly affects the birthweight of the foal, as demonstrated originally by Walton and Hammond57 in their classic between-breed crossing of small Shetland ponies with large Shire horses. Tischner and Klimczak58 observed a similar effect when they transferred Pony embryos into the uteri of larger draft-type recipient mares and compared birth size and subsequent development of the resulting foals with sex-matched siblings born from the genetic Pony mothers. Although both experiments demonstrated clearly that uterine size profoundly influences prenatal and postnatal growth of the foal, neither study investigated placental parameters in any detail or any adaptive changes the placental microcotyledons
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may have undergone. Accordingly, Allen et al.55 similarly utilized between-breed embryo transfer to create Thoroughbred-in-Pony (Tb-in-P) pregnancies in which the genetically larger Thoroughbred fetus endured cramping and nutritional deprivation in utero, and reciprocal Pony-in-Thoroughbred (P-inTb) pregnancies in which the smaller pony fetus was exposed to nutritional and spatial excesses in utero. Control Thoroughbred-in-Thoroughbred (Tb-in-Tb) and Pony-in-Pony (P-in-P) pregnancies were created using artificial insemination. The study shed considerable new light on both the gross and microscopic development of the placenta in both control and experimental pregnancies. The mass, gross area, and volume of the allantochorion were significantly greater in the Tb-in-Tb than in the P-in-P pregnancies, whereas intermediate values were obtained for the experimental P-inTb and Tb-in-P groups. Analysis of the data showed that both maternal and fetal genotypes played significant roles in determining these parameters. For example, stereological assessment of the surface area per unit volume (Sv ) of the microcotyledons showed that Thoroughbred mares exhibited a significantly higher mean Sv than Pony mares, regardless of the genotype of the fetus they were carrying. Hence, it could be concluded that the plexiform nature of the microcotyledons is under maternal control. On the other hand, in the ‘deprived’ Tb-in-P pregnancies, the microcotyledonary villi were longer than those in the control P-in-P pregnancies, although the Sv remained unchanged; presumably this had occurred as a compensatory measure to increase the area of fetomaternal contact. However, despite this attempt to redress the shortfall, a reduction in the total area of fetomaternal contact across the placental interface was still apparent; namely, 42.0 (± 4.4) m2 in the Tb-in-Tb control pregnancies to 29.4 (± 2.6) m2 in the Tb-in-P pregnancies. This was reflected in the significantly lower birthweight of the Tb-in-P foals compared to their Tb-in-Tb controls. When calculating kilograms of foal birthweight per square meter of feto-maternal contact as a measure of placental efficiency, an overall mean figure of 1.23 kg/m2 was observed, with no major differences between the four groups. In conclusion, maternal size and genotype, of both the dam and the fetus, can act to restrict or enhance growth of the allantochorion and, hence, the available area for fetomaternal exchange.
Influence of age and parity Age-related degenerative changes in the endometrium may also limit placentation by reducing the effective area for feto-maternal exchange and
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thereby lead to intrauterine growth retardation (IUGR) of the fetus.59 Fewer and shorter chorionic villi develop on the areas of allantochorion apposed to degenerate endometrium in aged mares, which causes delayed and abnormal development of the microcotyledons and a resulting reduction in weight gain in the fetus.38 Therefore, it was not surprising to find that stereological assessment of the placental microcotyledons of 84 Thoroughbred mares, separated into four groups on the bases of age and parity – primiparous mares aged ≤ 6 years (n = 24) and multiparous mares aged 5–9 (n = 41), 10–15 (n = 10) and ≥ 16 (n = 9) years – showed significantly lower Sv values in the oldest animals. However, the maiden primiparous mares also showed significantly lower Sv values than their secundiparous and younger multiparous counterparts, despite the healthy, virginal endometrium in the maiden animals. Similarly, 11 young Thoroughbred mares followed in their first and second successive pregnancies showed significantly lower Sv values in the first parity.60 Likewise, pregnant fillies aged 1 and 2 years from which their conceptuses were removed surgically on day 70 of gestation, also showed a significant increase in Sv values between the first and second pregnancy, despite the confounding factors of immaturity of both the placenta and the dam.39 It would appear, therefore, that the equine uterus needs to be ‘primed’ in some way by a first pregnancy for microcotyledon development to reach its full potential. In the pregnant mare, large quantities of mitogenic growth factors such as epidermal growth factor (EGF) are secreted by the endometrial glands,43 and differences in the quantities of such factors may play a significant role in bringing about the observed reduction in microcotyledon Sv in older animals. However, although Gerstenberg39 reported a reduction in EGF production by damaged glands in older mares from around day 35 post-ovulation, an ultrastructural study of the secretory endometrium by Tunon et al.61 failed to show differences between nulli-, primi-, and secundiparous mares during estrus. It is interesting to speculate that the remodeling of the endometrium that occurs during a first pregnancy may not completely resolve following parturition so that pregnancy results in permanent anatomical changes. This theory has been proposed as the basis for the increases in birthweight measured in human infants born from second and subsequent gestations.62
Placental efficiency Assessment of placental efficiency in terms of kilograms of foal birthweight per square meter of micro-
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Microcotyledonary surface density (µm−1) Figure 230.4 The relationship between placental efficiency (kilograms of foal birthweight per m2 of microscopic surface area of the microcotyledons) and the surface density of the microcotyledons (y = – 33.7x + 2.4; r = 0.6; n = 84; p < 0.001).
scopic feto-maternal contact showed that the allantochorion of both primiparous and multiparous mares aged ≥ 16 years appeared to be more ‘productive’ than those of the other groups of multiparous mares. Indeed, regardless of age and parity of the mare, any increase in microscopic surface area was correlated with decreases in placental efficiency. The same correlation was also evident when plotting the Sv of the microcotyledons against placental efficiency (Figure 230.4). Such findings indicate that smaller placentas, or less well-developed microcotyledons, become markedly more efficient in extracting nutrients for the growing fetus. It is important to remember that placental efficiency does not relate to foal birthweight, and large and small foals can develop from both efficient and inefficient placentas. These same correlations matching increases in placental weight with decreases in placental efficiency have been reported in the pig,63, 64 in which placental efficiency was correlated strongly with both placental and endometrial vascularity.65 Additionally, differences in placental efficiency and vascularity have been reported between different breeds of pig. For example, the prolific Meishan breed achieves an increase in litter size over its equals by virtue of smaller, but more vascular, placentas.66 This example also highlights the concept of placental efficiency whereby the Meishan placenta, although smaller, has the capacity to produce, weight-for-weight, more piglet per placenta, as a result of enhanced placental and endometrial vascularity. It remains to be determined if the differences observed in placental efficiency between horses may be related to placental vascularity. Alterations in endometrial vascularity and maternal uterine blood flow have been observed in mares of varying age and parity.67, 68 Such differences may influence the ability
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Development and Morphology of the Placenta of the placenta to become more or less efficient. Furthermore, experimental reduction of placental size in the ewe has been shown to enhance placental transport of antipyrine and facilitate diffusion of glucose.69
Nutritional insults during gestation It is clear from work in other species that placental development can be enhanced or compromised by a range of external influences. For example, a critical period of sensitivity occurs between approximately 40 and 80 days of gestation in the ewe, which coincides with the time of rapid proliferative growth of the placenta.70 During this period placental growth may be modulated nutritionally by the size,71, 72 body condition,73 and degree of maturity of the ewe.74, 75 There is a paucity of experimental data relating maternal nutrition with placental development and function in the horse. During an experiment undertaken to examine the effects of moderate versus excessive maternal nutrition during pregnancy on fetal and placental development in maiden primigravid Thoroughbred mares, both groups of fillies became severely infected with Streptococcus equi (‘strangles’) when they were between 90 and 150 days of gestation and hence still in the proliferative growth phase of placentation. The disease resulted in a dramatic weight loss in all the fillies and it was subsequently possible to show that their placentas had modified and adapted their function to cope with the adverse circumstances. Maternal weight loss during the infection taken as a percentage of pre-infection bodyweight was used to indicate the degree of nutritional insult the fillies had suffered. This correlated negatively with the weight and volume of the placenta at term, whereas placental efficiency was enhanced when both maternal weight and placental weight fell.76 Evidence has emerged that pigs selected for postnatal survival exhibit an increase in placental efficiency77 so that the above changes in placental efficiency observed in the nutritionally stressed pregnant mares may indicate an adaptive strategy to increase the provision of nutrients to the fetus at critical times during gestation – for example, to meet the challenge of increasing fetal demands in late gestation or to make up for reduced feto-maternal contact across the placental interface. The plexiform nature of the microcotyledons in the maiden fillies that suffered strangles, as indicated by their mean Sv values, was not modified by the nutritional insult and was equivalent to the mean obtained in healthy primiparous Thoroughbreds.60 In women the Sv of the placental villi can be modified by many factors, including intrauterine growth restric-
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tion,78 pre-eclampsia,79–81 hypoxia at high altitude,82 and exercise.83 By contrast, the only factors that appear able to influence the Sv of the villi of the microcotyledons in the mare are maternal age, parity, and genotype,55, 60, 76 three factors that also influence the architecture and health of the maternal endometrium.59, 84 Thus the maternal endometrium seems to exert the overriding influence on microcotyledon development in the mare. It is likely that the epitheliochorial structure of the mare’s placenta with the intimate contact of the placental villi and maternal endometrial crypts, in contrast to the hemochorial structure of the human placenta with its associated breakdown of maternal tissues,85 probably influences the ability of the chorionic villi to modify themselves under different circumstances.
Placental vascularity As in all mammalian placentas, extensive development of maternal and fetal capillary networks in the mare parallels the cellular development at the feto-maternal interface so as to maximize the extent and efficiency of hemotrophic exchange.42 In normal human pregnancy, development of the placental capillary networks begins with vasculogenesis, in which vascularization of the first rudimentary villi is the result of local de novo formation of capillaries from blood islands, rather than the extension of embryonic vessels into the placenta. Feto-placental angiogenesis then shapes development of the villi, so that an initial phase of branching angiogenesis, resulting in the formation of tightly looped capillaries, followed by one of increased non-branching angiogenesis leading to the formation of longer capillaries, is the norm.86, 87 This essentially biphasic development of placental vascularity also occurs in the equine placenta, commencing with complex interbranched capillary networks within the developing chorionic villi in early gestation, which change increasingly in the second half of pregnancy to elongated non-branching capillaries that run parallel to one another within the villi.42 Near term in the mare, vascular casts show that the extensive allantochorion, which now has a total microscopic area of feto-maternal contact at the placental interface averaging 40 m2 in a Thoroughbred mare,60 consists of no more than a single thin layer of low columnar-to-cuboidal trophoblast cells, which overlies and provides the essential structural framework for an incredibly densely packed mass of fetal capillaries supported in minimal amounts of allantoic mesoderm (230.5). In women, the relationship between the placental villi and their capillary networks suggests that the villous trophoblast is a plastic layer that adapts in parallel, or in response to,
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Figure 230.5 A scanning electron micrograph of the equine microcotyledonary villi at 309 days of gestation. The trophoblast cells covering the villi have been removed and the complex fetal capillary system can be seen. It is evident from this micrograph how the fetal villi are composed almost entirely of the capillary network.
the changing structure of the underlying vasculature. This idea implies that the trophoblast does not sculpt the vasculature and that angiogenesis is responsible for villous development and differentiation.86 To what extent this scheme might hold true in the horse, with enrapment of the villi by the maternal epithelium, remains to be determined. The major driving force behind the development of the capillary networks is vascular endothelial growth factor (VEGF), a homodimeric glycoprotein that exists in five alternatively spliced forms containing 121–206 amino acids in each of the monomers.88, 89 It is a potent mitogen, morphogen, and chemoattractant for endothelial cells and is widely recognized as the most potent stimulator of vasculogenesis and angiogenesis.87 VEGF, and its two main receptor molecules, VEGF-I (Flt) and VEGF-II (KDR), have been shown to be expressed in the endometrium, decidual tissue, and trophoblast of the human90–98 and non-human primates,99 and also in the syndesmochorial placenta of the sheep100 and the epitheliochorial placenta of the pig.101 Hypoxia is known to be a potent stimulator of VEGF synthesis,102, 103 and estrogens also stimulate VEGF production in the ewe,104 in pregnant rats,105 in women,93 and in sows.106 Localization of VEGF and its two receptors, Flt-I and KDR, in the endometrium and placenta of the mare during the estrous cycle and pregnancy has also been demonstrated immunohistochemically.107 As in the pig with its similar non-invasive epitheliochorial placenta,87, 108, 109 the two tissues in the pregnant mare’s uterus that labeled strongly and consistently throughout gestation were the maternal and fetal epithelial layers – the luminal and glandular epithelia in
the endometrium on the maternal side and the outermost trophoblast layer of the allantochorion on the fetal side. In addition to their shared epitheliochorial placentation, porcine and equine embryos both secrete appreciable quantities of estrogens from as early as day 10 after ovulation.110, 111 Charnock-Jones et al.89 postulated that the increase in angiogenesis observed in the sow at the mesometrial side of the uterus where the embryonic disk first establishes contact with the endometrium112, 113 is likely to be stimulated by the paracrine action of blastocyst estrogens stimulating VEGF release in the endometrium. Since estrogens have been shown to stimulate VEGF production in other species93, 104, 105 there is a likely angiogenic action of blastocyst estrogens in the horse. Indeed, strong staining for KDR is apparent in the subepithelial stroma of the endometrium directly beneath the conceptus as early as day 18 after ovulation,107 and Doppler ultrasonography shows a pronounced increase in endometrial blood flow in the gravid versus the non-gravid uterine horn around the same stage.114
Structure and function Numerous factors dictate the rate of development and architecture of the equine placenta, only some of which are understood. Although this chapter has concentrated upon the structure of the allantochorionic portion of the placenta, it must be remembered that all parts of this unique organ will govern its development and function. For example, what role might the morphology of the umbilical cord play in influencing placental structure? Furthermore, since structure is closely correlated to function, any deficiencies in the former may adversely affect the latter to the extent of causing related deficits of fetal growth and maturity. In addition, long-term changes in the physiology and metabolism of the offspring right through to adulthood can result from an inappropriate intrauterine existence occasioned by an inadequately functioning placenta.115, 116
References 1. Van Niekerk CH, Allen WR. Early embryonic development in the horse. J Reprod Fertil Suppl 1975;23:495–8. 2. Battut I, Colchen S, Fi´eni F, Tainturier D, Bruyas JF. Success rates when attempting to non surgically collect equine embryos at 144, 156, and 168 hours after ovulation. Equine Vet J Suppl 1997;25:60–62. 3. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 secretion by oviductal transport stage equine embryos. Biol Reprod 1991;45:540–3.
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24. Ginther OJ. Fixation and orientation of the early equine conceptus. Theriogenology 1983;19:613–23. 25. Ewart JC. Studies on the development of the horse I. The development during the third week. Trans R Soc Edin 1915;51:287–329. 26. Ewart JC. A Critical Period in the Development of the Horse. London: Adam and Charles Black, 1897. 27. Allen WR, Moor RM. The origin of the equine endometrial cups. I. Production of PMSG by the fetal trophoblast cells. J Reprod Fertil 1972;29:313–16. 28. Allen WR, Hamilton DW, Moor RM. The origin of the equine endometrial cups. II. Invasion of the endometrium by trophoblast. Anat Rec 1973;177:485–502. 29. Wooding FBP, Morgan G, Fowden AL, Allen WR. A structural and immunological study of chorionic gonadotrophin production by equine trophoblast girdle and cup cells. Placenta 2001;22:749–67. 30. Moor RM, Allen WR, Hamilton DW. Origin and histogenesis of equine endometrial cups. J Reprod Fertil Suppl 1975;23:391–6. 31. Cole HH, Goss H. The source of equine gonadotrophin. In: Essays in Biology in Honor of Herbert M Evans. Berkeley: University of California Press, 1943; pp. 107–19. 32. Clegg MT, Boda JM, Cole HH. The endometrial cups and allantochorionic pouches in the mare with emphasis on the source of equine gonadotrophin. Endocrinology 1954; 54:448–63. 33. Hamilton DW, Allen WR, Moor RM. The origin of the equine endometrial cups. III. Light and electron microscopic study of fully developed equine endometrial cups. Anat Rec 1973;177:503–18. 34. Samuel CA, Allen WR, Steven DH. Studies on the equine placenta. I. Development of the microcotyledons. J Reprod Fertil 1974;41:441–5. 35. Samuel CA, Allen WR, Steven DH. Ultrastructural development of the equine placenta. J Reprod Fertil Suppl 1975;23:575–8. 36. Leiser R, Pfarrer C, Adb-Elnaeim M, Dantzer V. Fetomaternal exchange in epitheliochorial placental types studied by histology and microvascular corrosion casts. Troph Res 1998;12:21–39. 37. MacDonald AA, Chavatte P, Fowden AL. Scanning electron microscopy of the microcotyledonary placenta of the horse (Equus caballas) in the latter half of gestation. Placenta 2000;21:565–74. 38. Bracher V, Mathias S, Allen WR. Influence of chronic degenerative endometritis (endometrosis) on placental development in the mare. Equine Vet J 1996;28:180– 8. 39. Gerstenberg C. Factors controlling epitheliochorial placentation in the mare. University of Cambridge, PhD thesis, 1999. 40. Steven DH. Placentation in the mare. J Reprod Fertil Suppl 1982;31:41–55. 41. Gerstenberg C, Allen WR, Stewart F. Cell proliferation patterns during development of the equine placenta. J Reprod Fertil 1999;117:143–52. 42. Abd-Elnaeim MMM, Leiser R, Wilsher S, Allen WR. Structural and haemovascular aspects of placental growth throughout gestation in young and aged mares. Placenta 2006;27:1103–13. 43. Stewart F, Power CA, Lennard SN, Allen WR, Amet L, Edwards RM. Identification of the horse epidermal growth factor (EGF) coding sequence and its use in monitoring
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EGF gene expression in the endometrium of the pregnant mare. J Mol Endocrinol 1994;12:341–50. Gerstenberg C, Allen WR, Stewart F. Factors controlling epidermal growth factor (EGF) gene expression in the endometrium of the mare. Mol Reprod Dev 1999;53:255–65. Jones JI, Clemmons DR. Insulin-like growth factors and their binding proteins: biological actions. Endocr Rev 1995; 16:3–35. ¨ K, EnJoujou-Sisic K, Gran´erus M, Wetterling H, Wikstrom ¨ W, Jeffcott L, Schofield PN, Welin A. Developmengstrom tal regulation of insulin-like growth factor II expression in the horse. Cell Biol Int 1993;17:603–7. Lennard SN, Stewart F, Allen WR. Insulin-like growth factor II expression in the fetus and placenta of the horse during the first half of gestation. J Reprod Fertil 1995;103: 169–79. Stewart F, Lennard SN, Allen WR. Mechanisms controlling formation of the equine chorionic girdle. Biol Reprod Monogr 1995;1:151–9. Stewart F. Roles of mesenchymal-epithelial interactions and hepatocyte growth factor-scatter factor (HGF-SF) in placental development. Rev Reprod 1996;1:144–8. Lennard SN, Stewart F, Allen WR. Transforming growth factor β1 (TGF β1) expression in the endometrium of the mare during placentation. Mol Reprod Dev 1995;42: 131–40. Arai KY, Tanaka Y, Taniyama H, Tsunoda N, Nambo Y, Nagamine N, Watanabe G, Taya K. Expression of inhibins, activins, insulin-like growth factor-I and steroidogenic enzymes in the equine placenta. Domest Anim Endocrinol 2006;31:19–34. ¨ Grunig G, Antczak DF. Horse trophoblasts produce tumour necrosis factor α but not interleukin 2, interleukin 4, or interferon γ . Biol Reprod 1995;52:531–9. Rossdale PD. A clinical assessment of the health status of the newborn Thoroughbred foal. Royal College of Veterinary Surgeons, Fellowship thesis, 1966. Cotrill CM, Jeffers LU, Ousey JC, McGladdery AJ, Rickett SW, Silver M, Rossdale PD. The placenta as a determinant of fetal well-being in normal and abnormal equine pregnancies. J Reprod Fertil Suppl 1991;44:591–601. Allen WR, Wilsher S, Turnbull C, Stewart F, Ousey J, Rossdale PD, Fowden AL. Influence of maternal size on placental, fetal and postnatal growth in the horse. I. Development in utero. Reproduction 2002;123:445–53. Brown MA, Howard CV, Jolleys GD. Principles of stereology. In: Wootton R, Springhall DR, Polak JM (eds) Image Analysis in Histology: Conventional and Confocal Microscopy. Cambridge, UK: Cambridge University Press, 1995, 96– 120. Walton A, Hammond J. The maternal effects on growth and conformation in Shire horse-Shetland pony crosses. Proc R Soc Lond B Biol Sci 1938;125:311–35. Tischner M, Klimczak M. The development of Polish ponies after embryo transfer to larger recipients. Equine Vet J Suppl 1989;8:62–3. Kenny RM. Proceedings of the John P. Hughes International Workshop on Equine Endometritis. Equine Vet J 1993;25:184–7. Wilsher S, Allen WR. The effects of maternal age and parity on placental and fetal development in the mare. Equine Vet J 2003;35:476–83. Tunon AM, Rodriguez-Martinez H, Haglund A, Al-bihn A, Magnusson U, Einarsson S. Ultrastructure of the secretory
62.
63.
64. 65.
66.
67.
68.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
endometrium during oestrus in young maiden and foaled mares. Equine Vet J 1995;27:382–88. Khong TY, Adema ED, Erwich JJ. On an anatomical basis for the increase in birthweight of second and subsequent born children. Placenta 2003;24:348–53. Wilson ME, Biensen NJ, Ford SP. Novel insight into the control of litter size in pigs, using placental efficiency as a selection tool. J Anim Sci 1999;77:1654–8. Wilson ME, Ford SP. Comparative aspects of placental efficiency. Reproduction (Suppl.) 2001;58:223–32. Vonnahme KA, Wilson ME, Ford SP. Relationship between placental vascular endothelial growth factor expression and placental/endometrial vascularity in the pig. Biol Reprod 2001;64:1821–5. Ford SP. Embryonic and fetal development in different genotypes in pigs. J Reprod Fertil Suppl 1997;52:165– 76. Stolla R, Bollwein H. Colour Doppler sonography of the uterine artery in subfertile mares. J Reprod Fertil 1997;13:547 [abstr.]. Bollwein H, Woschee I, Stolla R. Uterine durchblutung wahrend der graviditat der stute. Pferdheilkunde 1999; 15:595–8. Owens JA, Falconer J, Robinson JS. Restriction of placental size in sheep enhances efficiency of placental transfer of antipyrine, 3-O-methyl-D-glucose but not urea. J Dev Physiol 1987;9:457–64. Ehrhardt RA, Bell AW. Growth and metabolism of the ovine placenta during mid-gestation. Placenta 1995;16: 727–41. Russell AJF, Food JZ, White IR, Davies GJ. The effect of weight at mating and of nutrition during mid-pregnancy on the birth weight of lambs from primiparous ewes. J Agri Sci 1981;97:723–9. McCrabb GJ, Egan AR, Hosking BJ. Maternal undernutrition during mid-pregnancy in sheep: variable effects on placental growth. J Agric Sci 1992;118:127–32. Clarke L, Heasman L, Juniper DT, Symonds ME. Maternal nutrition in early-mid gestation and placental size in sheep. Brit J Nutr 1998;79:349–64. Wallace JM, Aitkin RP, Cheyne MA. Nutrition partitioning and fetal growth in rapidly growing adolescent ewes. J Reprod Fertil 1996;107:183–90. Wallace JM, Bourke DA, Aitkin RP, Cruikshank MA. Effect of switching maternal nutrient intake at the end of the first trimester on placental and fetal growth in adolescent sheep carrying singleton fetuses. J Reprod Fertil 1997;19:14–15 [abstr.]. Wilsher S, Allen WR. Effects of a Streptococcus equi infection-mediated nutritional insult during mid-gestation in primiparous Thoroughbred fillies. Part 1. Placental and fetal development. Equine Vet J 2006;38:549–57. Wilson ME, Ford SP, Vonnahme KA. Factors influencing placental growth and efficiency in pigs. In: Wilsher S, Wade JF (eds) Proceedings of a Workshop on Embryonic and Fetal Nutrition. Havemeyer Foundation Monograph Series No. 10, 2003; pp. 33–5. Mayhew TM, Ohadike C, Baker PN, Crocker IP, Mitchell C, Ong SS. Stereological investigation of placental morphology in pregnancies complicated by pre-eclampsia with and without intrauterine growth restriction. Placenta 2003;24:219–26. Boyd PA, Scott A. Quantitative structural studies on human placentas associated with pre-eclampsia, essential
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86.
87.
88.
89.
90.
91.
92.
93.
94.
95.
hypertension and intrauterine growth retardation. Br J Obstet Gynec 1985;184:146–52. Teasdale F. Histomorphometry of the human placenta in maternal pre-eclampsia. Am J Obstet Gynecol 1985;152: 25–31. Burton GJ, Reshetnikova OS, Milovanov AP, Teleshova OV. Stereological evaluation of vascular adaptations in human placental villi to differing forms of hypoxic stress. Placenta 1996;17:49–55. Mayhew TM. Changes in fetal capillaries during preplacental hypoxia: growth, shape remodelling and villous capillarization in placentae from high-altitude pregnancies. Placenta 2003;24:191–8. Jackson MR, Gott P, Lye SY, Knox Ritchie JW, Clapp JF III. The effects of maternal aerobic exercise on human placental development: Placental volumetric composition and surface areas. Placenta 1995;16:179–91. Lefranc AC, Allen WR. Influence of breed and oestrous cycle on endometrial gland surface density in the mare. Equine Vet J 2007;39:506–10. Steven DH. Anatomy of the placental barrier. In: Steven DH (ed.) Comparative Placentation. Essays in Structure and Function. New York: Academic Press, 1975; pp. 25–57. Kaufmann P, Mayhew TM, Charnock-Jones DS. Aspects of human fetoplacental vasculogenesis and angiogenesis. II Changes during normal pregnancy. Placenta 2004;25: 114–26. Charnock-Jones DS, Kaufmann P, Mayhew TM. Aspects of human fetoplacental vasculogenesis and angiogenesis. I. Molecular regulation. Placenta 2004;25:103–13. Houcke KA, Ferrara N, Winer, J, Cachianes G, Li B, Leung DW. The vascular endothelial growth factor family: identification of a fourth molecular species and characterisation of alternative splicing of RNA. Mol Endocrinol 1991;5:1806–14. Charnock-Jones DS, Sharkey AM, Rajput-Williams J, Burch D, Schofield JP, Fountain SA, Boocock CA, Smith SK. Identification and localization of alternately spliced mRNAs for vascular endothelial growth factor in human uterus and estradiol regulation in endometrial carcinoma cell lines. Biol Reprod 1993;48:1120–8. Sharkey AM, Charnock-Jones DS, Boocock CA, Brown KD, Smith SK. Expression of mRNA for vascular endothelial growth factor in human placenta. J Reprod Fertil 1993;99:609–15. Charnock-Jones DS, Sharkey AM, Boocock CA, Ahmed A, Plevin R, Ferrara N, Smith SK. Vascular endothelial growth factor receptor localization and activation in human trophoblast and choriocarcinoma cells. Biol Reprod 1994;51: 524–30. Jackson MR, Carney EW, Lye SJ, Ritchie JW. Localization of two angiogenic growth factors (PDECGF and VEGF) in human placenta throughout gestation. Placenta 1994;15:341–53. Ahmed A, Li XF, Dunk C, Whittle MJ, Rushton DI, Rollason T. Colocalisation of vascular endothelial growth factor and its Flt-1 receptor in human placenta. Growth Factors 1995;12:135–43. Clark DE, Smith SK, Sharkey AM, Sowter HM, CharnockJones DS. Hepatocyte growth factor-scatter factor and its receptor c-met: localization and expression in the human placenta throughout pregnancy. J Endocrinol 1996;151: 459–67. Sharaishi S, Nakagawa K, Kinukawa N, Nakano H, Sueishi K. Immunohistochemical localization of vascular
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endothelial growth factor in the human placenta. Placenta 1996;17:111–21. 96. Torry DS, Holt VJ, Keenan JA, Harris G, Caudle MR, Torry RJ. Vascular endothelial growth factor expression in cycling human endometrium. Fertil Steril 1996;66:72–80. 97. Vuckovic M, Ponting J, Terman BI, Niketic V, Seif MW, Kumar S. Expression of the vascular endothelial growth factor receptor, KDR, in human placenta. J Anat 1996;188:361–6. 98. Shore VH, Wang T-H, Wang C-L, Torry RJ, Caudle MR, Torry DS. Vascular endothelial growth factor, placenta growth factor and their receptors in isolated human trophoblast. Placenta 1997;18:657–65. 99. Wulff C, Wilson H, Dickson SE, Wiegand SJ, Fraser HM. Hemochorial placentation in the primate: expression of vascular endothelial growth factor, angiopoietins, and their receptors throughout pregnancy. Biol Reprod 2002;66:802–12. 100. Cheung CY, Singh M, Ebaugh MJ, Brace RA. Vascular endothelial growth factor gene expression in ovine placenta and fetal membranes. Am J Obstet Gynecol 1995;173:753–9. 101. Charnock-Jones DS, Clark DE, Licence D, Day K, Wooding FBP, Smith SK. Distribution of vascular endothelial growth factor (VEGF) and its binding sites at the maternal-fetal interface during gestation in pigs. Reproduction 2001;122:753–60. 102. Shweiki D, Itin A, Neufeld G, Gitay-Goren H, Keshet E. Patterns of expression of vascular endothelial growth factor (VEGF) and VEGF receptors in mice suggest a role in hormonally regulated angiogenesis. J Clin Invest 1993; 91:2235–43. 103. Goldberg MA, Schneider TJ. Similarities between the oxygen-sensing mechanisms regulating the expression of vascular endothelial growth factor and erythropoietin. J Biol Chem 1994;269:4355–9. 104. Reynolds LP, Kirsch JD, Kraft KC, Redmer DA. Time course of the uterine response to estradiol-17β in ovariectomised ewes: expression of angiogenic factors. Biol Reprod 1998;59:613–20. 105. Cullinan-Bove K, Koos RD. Vascular endothelial growth factor/vascular permeability factor expression in the rat uterus: rapid stimulation by estrogen correlates with estrogen-induced increases in uterine capillary permeability and growth. Endocrinology 1993;133:829–37. 106. Anderson LH, Christenson LK, Ford SP. Estrogen associated changes in uterine blood flow and maternal recognition of pregnancy in Chinese (Meishan) pigs. Anim Reprod Sci 1993;36:135–43. 107. Allen WR, Gower S, Wilsher S. Immunohistochemical localization of vascular endothelial growth factor (VEGF) and its two receptors (Flt-I and KDR) in the endometrium and placenta of the mare during the oestrous cycle and pregnancy. Reprod Domest Anim 2007;42:516–26. 108. Winther H, Ahmed A, Dantzer V. Immunohistochemical localization of vascular endothelial growth factor (VEGF) and its two specific receptors, Flt-1 and KDR, in the porcine placenta and non-pregnant uterus. Placenta 1999;20:35– 43. 109. Dantzer V, Winther H. Histological and immunohistochemical events during placentation in pigs. Reproduction (Suppl.) 2001;58:209–22. 110. Bazer FW, Thatcher WW. Theory of maternal recognition of pregnancy in swine based on oestrogen controlled endocrine versus exocrine secretion of prostaglandin F2α by the uterine endometrium. Prostaglandins 1977;14:397– 401.
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111. Heap RB, Hamon M, Allen WR. Studies on oestrogen synthesis by the preimplantation equine conceptus. J Reprod Fertil Suppl 1982;32:343–52. 112. Keys JL, King GJ. Microscopic examination of porcine conceptus-maternal interface between 10 and 19 days of pregnancy. Am J Anat 1988;188:221–38. 113. Dantzer V, Leiser R. Initial vascularisation in the pig placenta: demonstration of nonglandular areas by histology and corrosion casts. Anat Rec 1994;238:177–90.
114. Silva LA, Gastal EL, Beg MA, Ginther OJ. Changes in vascular perfusion of the endometrium in association with changes in location of the embryonic vesicle. Biol Reprod 2005;72:755–61. 115. Barker DJ. Intrauterine programming of coronary heart disease and stroke. Acta Paediatr (Suppl.) 1997;423:178–82. 116. Barker DJ. Chronic disease in humans originates in the intra-uterine environment. Anim Reprod Sci 2006;94: 343–54.
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Chapter 231
Pregnancy Examination Patrick M. McCue and Angus O. McKinnon
Economic pressures in the equine breeding industry create a demand for accurate early pregnancy diagnosis and detection of pregnancy loss. Techniques used for pregnancy diagnosis and the timing of specific pregnancy tests are based on normal physiological events in the pregnant mare (Table 231.1). Accurate breeding dates are consequently helpful when determining when to test for pregnancy. If breeding dates are not available or unknown (i.e. a pasture-bred mare), the clinician should be made aware of the first and last potential breeding dates. All methods of pregnancy examination have limitations and some may be associated with a false positive or false negative diagnosis. An understanding of reproductive physiology of the pregnant mare (Figure 231.1) as well as the advantages and disadvantages of the various procedures for pregnancy diagnosis is important to maximize the odds of making an accurate diagnosis. Techniques for detection of pregnancy in the mare include evaluation of behavioral characteristics, manual palpation or ultrasonography of the reproductive tract, hormone analysis and miscellaneous other tests (Table 231.2). The type of test selected may depend on: (i) experience of the clinician; (ii) equipment and facilities available; (iii) stage of gestation; (iv) size and behavior of the mare; (v) medical complications (previous rectal tear, impending abortion, suspected pregnancy loss, etc.), or other factors. The primary method for initial pregnancy examination currently recommended is transrectal ultrasonography performed 14 to 16 days after ovulation. This protocol allows for accurate diagnosis of pregnancy status, early detection and management of twins, evaluation of the corpus luteum, and assessment of the presence or absence of uterine edema. Subsequent re-evaluation of pregnancy status at key stages of gestation is also recommended. For example, knowledge of the pregnancy status of a mare at
the time of endometrial cup formation is critical, as mares that lose their pregnancy after cup formation are unlikely to cycle or become pregnant again that season. In addition, some breeding contracts and insurance companies underwriting mares against pregnancy loss provide specific guidelines as to acceptable techniques and gestational age for pregnancy examinations (i.e. often 42 to 45 days of pregnancy).
Behavior A majority of non-pregnant mares exhibit a regular and predictable pattern of behavioral estrus. Mares that have been bred and return to estrus within 18 days after ovulation can usually be presumed to be not pregnant.2 However, early pregnant mares may show behavioral estrus and stand for a stallion.3 Asa and colleagues4 estimated that 5–10% of pregnant mares exhibit behavioral estrus. Hayes and Ginther5 noted that pregnant mares carrying a female conceptus had a higher incidence of behavioral estrus than mares carrying a male conceptus. Conversely, the cyclic pattern of estrus ceases in a majority of mares that become pregnant (i.e. they do not come back into heat). A misconception by horse owners is that all mares that have been bred and do not return to estrus are pregnant. Non-pregnant mares that fail to return to estrus may have a ‘silent heat’, persistent corpus luteum, early embryonic loss, luteinized anovulatory follicle, failure of follicular development, variation in estrous cycle length, or other factors affecting their behavior. An early accurate pregnancy diagnosis in mares that fail to return to estrus can provide an owner with critical information with which to make management decisions.
Visual assessment and ballottement of the abdomen External visual examination of a pregnant mare during the first half of pregnancy, or even for a long time
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 231.1 Physiological benchmarks during the first 3 months of pregnancy in mares Gestational age
Physiological event
0 1–3 days 1–5+ days 5–6 days
Ovulation and fertilization of the oocyte Onset of early pregnancy factor production Formation of the primary corpus luteum Passage of the embryo through the isthmus of the oviduct and into the uterus, facilitated by secretion of PGE2 by the embryo
6–16 days 12–15 days 16–17 days 24–25 days 35–37 days 35–40 days 45–60 days 70–90 days 90–100 days
Migration of embryo throughout uterus Maternal recognition of pregnancy ‘Fixation’ of embryo in uterus Embryonic heartbeat first routinely evident with ultrasonography Endometrial cups begin to form; secretion of eCG begins Increase in estrogen conjugate secretion by the corpus luteum in response to eCG Secondary corpora lutea begin to develop Placental progestins begin to be produced Estrogens and androgen production begins from feto-placental unit
eCG, equine chorionic gonadotropin; PGE2 , prostaglandin E2 .
thereafter, may fail to reveal her to be in foal. Typically, though, by the fifth to sixth month of pregnancy the abdomen may take on a ‘pear-shape’, with the greatest width in the ventral abdomen.6 In some mares a distinct change in abdominal shape may be noted as the fetus ‘drops’ into a more ventral position in late gestation. Direct ballottement of a fetus from the outside may be possible in a late-pregnant mare, but is often difficult due to the thickness and muscular tension of the abdominal wall. Movement of the fetus may occasionally be observed through the abdominal wall during late pregnancy.
Speculum examination Changes in cervical characteristics may be suggestive of pregnancy, but are not totally diagnostic. The cervix of a normal pregnant mare should appear dry, pale-white, tightly closed, and located high on the wall of the cranial vaginal vault by day 30. A sticky vaginal secretion is often evident.
Palpation of the reproductive tract Palpation of the reproductive tract per rectum was historically the most common technique utilized
Primary C.L. Accessory C.L. Endometrial cups Foetal gonads
Progestagens 20
10
400
150 Oestrogens
300
100 Parturition
200
eCG 50 100 FSH
0
0
0
40
80
120
100 200 Days of pregnancy
240
280
320
Plasma conjugated oestrogens (ng/ml)
30 Plasma progestagens (ng/ml)
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Serum eCG (i.u./ml)
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Figure 231.1 Reproductive physiology of the pregnant mare. C. L., corpus luteum (or corpora lutea); eCG, equine chorionic gonadotropin; FSH, follicle-stimulating hormone. From Allen et al.1
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Pregnancy Examination Table 231.2 Options for pregnancy examination in the mare Day of gestation
Pregnancy test
1–120 11–term 18–term 18–90 40–120 80–term 90–term
Early pregnancy factor (EPF)a Ultrasonography Palpation Progesteroneb Equine chorionic gonadotropin (eCG)c Relaxin Estrogens (conjugated or total)
a
Not commercially available. Not a direct indicator of pregnancy. c False positive and false negative results are possible. b
for pregnancy determination prior to the advent of ultrasonography. A diagnosis of pregnancy by palpation is based on detection of a bulge in the ventral aspect of the uterus, typically at the base of one uterine horn, and the presence of increased uterine and cervical tone. It has been suggested by various authors that a positive diagnosis of pregnancy should be possible by manual palpation of the reproductive tract by day 17 to 20,3 day 25 to 30,7 or day 30 to 35 after ovulation.8 Several factors combine to determine when a positive manual diagnosis of pregnancy can be made, including the experience and skill of the clinician and the type of mare being examined. An accurate early pregnancy diagnosis by transrectal palpation is often easier in maiden, open, or barren mares than in postfoaling mares. An increase in uterine tone and uterine wall thickness may be noted in pregnant mares between 12 and 18 days after ovulation.3 The tight, tubular nature of the uterus is greater in pregnant mares than in mares
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in diestrus or during prolongation of the corpus luteum. In addition, the cervix of a pregnant mare becomes elongated and very firm, taking on a ‘rod-like’ or ‘pencil-like’ consistency.9 The embryonic vesicle itself may be detected by day 20 with careful palpation by a skilled practitioner. Gentle palpation of the reproductive tract by an experienced individual does not harm the pregnancy.7, 8 Cupping the uterus in a hand and feeling along the ventral aspect of the uterus with the fingers is the optimal technique for manual detection of an embryonic bulge. The embryonic bulge will typically be located on the ventral aspect of the uterus at the base of one uterine horn. The size of the embryonic/fetal bulge, uterine characteristics, and ability to ballot the fetus will depend on the stage of gestation (Table 231.3; Figure 231.2). Manual palpation is not an adequate technique for the early detection and management of twins, detection and characterization of uterine fluid, or evaluation of the corpus luteum. As a consequence, ultrasonography has become a routine technique for detecting early pregnancy and monitoring ovarian and uterine function.
Ultrasonographic examination The most reliable test for early pregnancy evaluation is transrectal ultrasonography. An embryonic vesicle may be detected as early as day 10 or 11 after ovulation. Routine initial pregnancy examinations are typically performed 12 to 16 days post-ovulation. An ultrasonographic examination performed on or before day 16 is also beneficial for the identification and management of twin pregnancies, scheduling rebreeding in open mares, and early detection of
Table 231.3 Characteristics of the equine pregnancy noted during manual palpation by day of gestation Day of gestation
Characteristics noted on palpation
20
Embryonic vesicle is 30–40 mm in diameter; good uterine tone overall; slight decrease in tone of the uterine wall over the vesicular bulge
30
Embryonic vesicle is 40–50 mm in diameter; noted as a discrete fluid swelling in the ventral aspect of the uterus at the base of one horn (corpus-cornual junction); decreased tone over bulge more obvious
40
Fetal bulge is approximately 65 mm in diameter; decreased tone at site of bulge; tone of pregnant horn has decreased; cervical tone is excellent
50
Fetal bulge is at least 8 cm in diameter; fluid-filled vesicle extends from base of uterine horn into uterine body
60
Fetal bulge is 10–13 cm in diameter; loss of tone at bulge is more obvious; uterus begins to be pulled cranially and ventrally over the brim of the pelvis; ovaries pulled closer together; urinary bladder may be mistaken for fetal-fluid vesicle
60–100
Uterus increased in size, located farther ventral in the abdomen; ovaries are closer together and located in a more ventral position; direct palpation or ballottement of the fetus may be difficult; pyometra may be mistaken for pregnant uterus
90–120
Uterus in abdominal position; fetus can usually be easily palpated
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(a)
(b)
(c)
(d)
Figure 231.2 Drawing of uterus at various stages of early pregnancy. (a) 20 days pregnant; (b) 30 days pregnant; (c) 40 days pregnant; (d) 60 days pregnant. From McKinnon.10
problems associated with pregnancy. A thorough, systematic approach to ultrasonographic pregnancy diagnosis is imperative in order to correctly diagnose pregnancy status and detect twins (Figure 231.3). It is recommended that the entire uterus be scanned in
Figure 231.3 Ultrasonographic image of twin pregnancies, with embryonic vesicles separated slightly.
one continuous motion in order to avoid missing a section of the uterus. The most common locations for a single embryo or a twin embryo to be missed are the caudal uterine body adjacent to the cervix or the tip of a uterine horn. Determination of the number of ovulations and a time-course of ovulations is helpful in the subsequent detection and management of twins. Asynchronous ovulations that occur 1 to 2 days or more apart may result in twin pregnancies that are initially difficult to detect. A 12-day pregnancy examination based on the date of the first ovulation may reveal a day-12-sized embryo, but the second embryo from the latter ovulation may be too small to be detected. A follow-up ultrasonographic examination would therefore be indicated. Twin embryos that are adjacent to each other may be difficult to identify initially, but on careful examination a thin echogenic line can be noted separating the two embryos (Figure 231.4). It is of great benefit to identify, measure, and map the location of uterine cysts prior to breeding a mare. In some instances a uterine cyst may be confused with an embryonic vesicle. This may be crucial
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Figure 231.4 Ultrasonographic image of twin pregnancies, with embryos touching each other.
and potentially frustrating during an examination for twins. Re-evaluation of the mare after an initial ultrasonograhic examination for pregnancy may be necessary. The fact that ‘cysts do not grow (rapidly) and do not move’ within the uterus may be beneficial in differentiation of cysts from an embryonic vesicle. The equine embryo migrates throughout the uterus until day 16 or 17 (stage of fixation) and generally increases in diameter at a rate of approximately 3 to 5 mm per day. Rarely, a final diagnosis of pregnancy status may have to wait until after day 25 to determine if an embryonic vesicle, with an embryo proper and heartbeat, is present amongst a cluster of cysts (Figure 231.5). Examination of the ovaries for the presence and characteristics of a corpus luteum, and of the uterus for evidence of edema, is valuable at the initial pregnancy examination. Pregnancies may be saved by initiating progesterone supplementation in cases of corpus luteum regression. Measurement of blood
Figure 231.5 Ultrasonographic image of a 25-day pregnancy and adjacent cysts (note the abnormal orientation of the fetus).
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progesterone level may be indicated to confirm the functional status of the corpus luteum. The ultrasonographic appearance of the equine embryo changes from a round shape (day 12 to 16) to a more triangular (‘guitar pick’) shape on day 17 to 21. Growth rate of the vesicle plateaus between 17 and 24 days, after which growth resumes at a slightly slower rate11 (Figure 231.6). The developing embryonic vesicle may have a somewhat irregular shape from day 25 to 35. Ultrasonographic images of early equine pregnancies are presented in Figures 231.7 to 231.10. A routine follow-up examination by ultrasonography is generally recommended between 25 and 35 days post-ovulation to confirm that the pregnancy is ongoing and that a heartbeat is present. It is also a useful time to reconfirm that twins are not present, as twin pregnancies or abortions have been reported in mares in which a single embryonic vesicle was observed during pregnancy examination at 14 days13–15 (Figure 231.11). The embryo proper may become visible 20 to 23 days after ovulation, and a heartbeat can usually be observed by day 24 to 25.16 Absence of a heartbeat should be confirmed by a subsequent examination at least 24 to 48 hours later, before a final diagnosis of embryonic loss is made and prostaglandins are administered. Empty trophoblastic vesicles (ETV), defined as the presence of an embryonic (trophoblastic) vesicle without an embryo proper, have been identified in mares.17 An embryo destined to form an ETV often exhibits normal early growth (i.e. day 11 to 16). Ultrasonographically, an ETV is recognized as a static oval or irregular embryonic structure without an embryo proper after day 24 to 25 of gestation (Figure 231.12). Empty trophoblastic vesicles do not form endometrial cups, even if the vesicle is still present in the uterus after day 35 (PM McCue, unpublished observations). Mares determined to be empty prior to day 35 can usually be rebred that season as endometrial cups have not yet developed. Administration of prostaglandins may be required to bring these mares back into heat due to persistence of the corpus luteum. Mares that lose their pregnancy after day 35 will usually not cycle again that season or are very difficult to get in foal due to the presence of endometrial cups, the production of equine chorionic gonadotropin and subsequent supplementary corpus luteum (CL) development. Supplementary CLs are either secondary or accessory CLs, and may be identified in pregnant mares after 45 to 60 days of gestation. Secondary CLs form from ovulation and normal luteinization. Accessory CLs form following equine chorionic gonadotropin
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Mare: Pregnancy, Parturition and the Puerperal Period 62 60 58 56 54 52
Mean of height and width (mm) of ultrasound image
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50 48 46 44 42 40 38 36 34 32
Regression lines Horses Ponies Actual means Horses Ponies
Days 11–16 Days 16–28
30 28
Days 28–45
26 24 22 20 18 16 14 12 10 8 6 4 2 0 0 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 Day of pregnancy
Figure 231.6 Growth rate of early equine pregnancies. Adapted from Ginther,12 courtesy of Equiservices.
(eCG)-induced luteinization of follicles stimulated by endogenous follicle-stimulating hormone (FSH) that do not ovulate.1 Ultrasonographically, the progression of accessory CL formation includes development of a large follicle, identification of echogenic particles or strands within the follicular lumen, followed by infiltration of echogenic material within the follicular lumen. The mature secondary CL has similar ultrasonographic characteristics to the primary CL. Multiple supplementary CLs may be present on each ovary and the size of the ovary is subsequently increased (Figure 231.13). It should be noted that not every mare develops supplementary CLs. Observation of supplementary CLs may be important for management decisions in mares being supplemented with exogenous progesterone or progestins. Determination of fetal sex can initially be made between days 55 and 90 of gestation by evaluating the location of the genital tubercle relative to other fetal structures.18 Accurate fetal sex determination requires both quality ultrasonographic equipment and significant clinical experience. Transabdominal ultrasonography, possibly in conjunction with endocrine testing, is sometimes necessary later in gestation for determination of pregnancy status in mares that cannot be examined by a transrectal approach, such as miniature mares, fractious
mares, or mares with a history of a rectal tear, or to monitor fetal viability or placental health. A schedule of potential ultrasonographic examination dates is presented in Table 231.4. Frequency and schedule of initial and subsequent examinations are often dictated by factors such as availability of the mare or clinician, potential for twins, results of previous pregnancy examinations, a history of pregnancy loss in a particular mare, and economics.
Biological, chemical and immunological tests for pregnancy A variety of tests, including biological, chemical, and immunological tests, have been described for the diagnosis of pregnancy in mares. Tests have been performed on serum, plasma, milk, and feces. A majority have estimated the concentration of either eCG in serum or estrogens in blood or urine. Prior to the advent of immunological tests, biological tests with laboratory animals, birds, amphibians, and even plants were used to detect the presence of eCG, estrogens, or other factors in pregnant mare’s blood or urine. For a historical perspective, a list of some of the original diagnostic tests is presented in Table 231.5. An overview of each parameter is presented later in this chapter.
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Figure 231.7 Ultrasonographic image of a 12-day pregnancy. Figure 231.9 Ultrasonographic image of a 26-day pregnancy.
Hormonal tests for pregnancy
Hormones of the pregnant mare
Radioimmunoassays (RIA) and enzyme-linked immunosorbent assays (ELISA) are currently used for measurement of hormone levels in blood to determine pregnancy status and monitor fetal-placental health (Table 231.6). In many instances, an accurate diagnosis of true pregnancy status may best be determined by evaluation of a combination of endocrine parameters. Individually, all endocrine tests have limitations associated with a false positive or a false negative diagnosis. A hormone or other factor may be considered a direct indicator of pregnancy if the source is limited to the fetus and/or placenta.
Progesterone Measurement of blood progesterone levels has been evaluated as an aid in differentiating pregnant mares from non-pregnant mares that fail to return to estrus following breeding (i.e. persistent corpus luteum, ‘silent heat’ mares, or mares with a prolonged absence of follicular activity).2 Pregnant mares should theoretically have continued secretion of progesterone from the ovarian corpus luteum beyond the period of maternal recognition of pregnancy (MRP) (day 12 to 14), whereas non-pregnant mares should
Figure 231.8 Ultrasonographic image of a 16-day pregnancy.
Figure 231.10 Ultrasonographic image of a 50-day pregnancy.
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Figure 231.13 Ultrasonographic image of multiple supplementary (accessory) corpora lutea in a pregnant mare.
Figure 231.11 Ultrasonographic image of a 25-day pregnancy examination showing two embryos proper, each of which had a distinct heartbeat, in a recipient mare following transfer of a single embryo. A single embryonic vesicle was observed during a routine 14-day pregnancy examination.
experience a rapid decline in progesterone following the release of prostaglandins from the endometrium. Unfortunately, some non-pregnant mares will have elevated progesterone beyond the time of MRP (i.e. mares with a persistent corpus luteum, luteinized anovulatory follicle, or a diestrus ovulation).51–53 Consequently, measurement of progesterone in the blood of mares is of limited value for an accurate pregnancy diagnosis. Detection of a progesterone concentration of less than 1 ng/mL usually indicates absence of a pregnancy,2 unless the mare is being supplemented with a synthetic progestin. However, occasionally a 16-day ultrasound pregnancy examina-
Figure 231.12 Ultrasonographic image of an empty trophoblastic vesicle (28 days after ovulation).
tion will reveal an embryo in the presence of mild uterine edema and a regressing corpus luteum (Figures 231.14 and 231.15). Low plasma progesterone in these cases suggests failure of maternal recognition of pregnancy and progesterone supplementation would be indicated. Progesterone measurement is valuable in determining if a pregnant mare is producing sufficient progesterone to maintain her pregnancy. Generally, a serum concentration of ≥ 4 ng/mL is considered adequate to maintain pregnancy when measured during the first 2 to 3 months of gestation, when the ovary is the primary source of progesterone. Progesterone levels may rise substantially by day 60 in mares that form supplementary CLs. Evaluation and interpretation of progesterone levels in samples collected after day 100 are more complicated. Most progesterone assays utilize an antibody specific for natural progesterone (P4 ), with some potential cross-reactivity with other progestins. After day 100, the equine placenta produces a variety of progestin compounds, which are often not detected by a progesterone assay or have limited cross-reactivity in the assay.54 Consequently, it is not unusual to detect a ‘progesterone’ level of 2.5 to 4.0 ng/mL in a normal pregnant mare in the second half of gestation. Interpretation of progesterone values must be based on the stage of pregnancy. Equine chorionic gonadotropin (eCG) Detection of elevated levels of equine chorionic gonadotropin (eCG), formerly called pregnant mare’s serum gonadotropin (PMSG), in the blood of mares may be used as a direct positive indicator of pregnancy. Equine chorionic gonadotropin is produced by specialized placental cells called endometrial cups, which begin to form and invade the maternal
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Table 231.4 Schedule of potential ultrasonographic examinations in the early pregnant mare. Many modifications of this schedule are used in clinical practice Gestation day post-ovulation
Examination procedure
12–15
Initial pregnancy examination; identification and management of twins; evaluation of the corpus luteum, follicle status and presence of endometrial edema
24–27 ∼35 ∼45
Examination for an embryo proper; detection of a heartbeat; reconfirm absence of twins Confirmation of pregnancy status at the stage of endometrial cup formation This is historical representing ∼2 cycles wherein the mare does not show heat and also is often the time service fees are payable for pregnant mares
∼60 55–90 ∼150
Evaluation of pregnancy status and supplementary corpora lutea development Fetal sex determination Confirmation of pregnancy and administration of first EHV-1 vaccine
EHV-1, equine herpesvirus type 1.
endometrium at approximately day 35 of gestation.1 An increase in blood eCG levels can be detected by day 35 to 42, with maximum concentrations achieved by day 55 to 70.1, 37 Complete regression of the cups and an absence of eCG occurs by day 100 to 140. 22, 37 A mare carrying a twin pregnancy may have a
markedly elevated eCG level,55 whereas a mare carrying a mule fetus may have very low or no detectable eCG due to early immune rejection of the endometrial cups.56 The presence of a gonadotropic hormone in pregnant mares was first reported by Cole and Hart in
Table 231.5 Historical biological, chemical, and immunological tests for pregnancy determination in the mare Type of Test
Hormone
Method
Reference
Biological
Equine chorionic gonadotropin (eCG) (serum)
Mouse test (Aschheim–Zondek test)
Hart and Cole (1932);19 Jeffcott et al. (1969);20 Santamarina and Joven (1959)21
Rat test
Cole and Hart (1930);22 Zondek and Sulman (1945);23 Valle (1947)24
Friedman (rabbit) test Male toad (frog) test Cockerel test Mouse/Rat test Capon test White lupine test
Morales and Damonte (1950)25 Neto (1949);26 Sakuma (1952)27 Bratescu et al. (1955)28 Miller (1934);29 Mayer (1944)30 Greenwood and Blyth (1934)31 Novazzi (1949)32
Estrogen (urine) Unknown (serum) Chemical
Estrogen (urine)
Cuboni test Lunaas test Phenolsuphonic acid test
Cuboni (1934)33 Lunaas (1962)34 Mayer (1944)30
Immunological
eCG (serum)
Hemagglutination inhibition
Wide and Wide (1963);35 Chak and Bruss (1968);36 Allen (1969);37 Jeffcott et al. (1969);20 Mitchell (1971)38 Richards (1967);39 Wormstrand (1969)40 McCarthy and Pennington (1966)41 de Coster et al. (1980)42 Stewart et al. (1976);43 Fay and Douglas (1982)44 Mia et al. (1981);45 Squires et al. (1983)46
Agar gel diffusion (AGD) Immunoelectrophoresis Latex agglutination test Radioreceptor assay Enzyme-linked immunosorbent assay (ELISA) Radioimmunoassay (RIA)
PMSG, pregnant mare’s serum gonadotropin.
Nett et al. (1975; estrogen);47 Terqui and Palmer (1979; estrogen);48 Menzer and Schams (1979; PMSG);49 Kindahl et al. (1982; both)50
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Table 231.6 Endocrine tests for pregnancy diagnosis in the mare Hormone
Source
Time course
Comments
Progesterone (P4 )
Corpus luteum and/or placenta
Initial elevation after primary ovulation; additional rise after formation of secondary corpora lutea begins on day 45 to 60
Not a direct indicator of pregnancy; levels > 1.0 ng/mL associated with presence of luteal tissue in a non-pregnant or early pregnant mare; placenta begins to produce progestins by day 70, which may be detected in a progesterone assay; levels < 1.0 ng/mL are generally incompatible with pregnancy, in the absence of additional progestin support
Equine chorionic gonadotropin (eCG)
Endometrial cups
Cup formation begins at approximately day 35 of pregnancy; eCG may be detected in blood by day 35 to 42; maximum concentrations are reached by day 55 to 70; cups regress by day 100 to 140; eCG production and duration of secretion is highly variable between mares
Direct indicator of pregnancy; detection of eCG indicates presence of endometrial cups; false negatives may occur if testing before cup formation or after cup regression in pregnant mare; false positive may occur following pregnancy loss after cup formation
Estrone sulfate
Ovary (corpus luteum, CL) and feto-placental unit
Initial rise at day 35–45 of ovarian CL origin; substantial increase at day 90 of placental origin
Direct indicator of pregnancy after day 90; also useful in monitoring health status of feto-placental unit
Relaxin
Placenta
Increased blood levels by day 80; levels remain elevated until foaling
Direct indicator of pregnancy in the mare; may be useful as a marker for placental health
1930.22 A wide variety of biological tests were initially employed to evaluate the level of gonadotropic hormone in the urine or blood of pregnant mares. The pioneering test was the ‘mouse test’, which involved injection of mare’s urine into immature female mice approximately 21 days old. Ovarian follicular development, blood spots or enlargement of the
uterus were indicative of a positive test.19, 20 Some of the other interesting historic biological tests for eCG utilized rabbits, male toads, and cockerels.25–28 The Friedman test relied on the induction of ovulation in rabbits for a positive test.25 Male toads injected with mare’s serum were reported to excrete spermatozoa into their cloaca if eCG was present.26, 27 Injection of
Figure 231.14 Ultrasonographic images of an embryonic vesicle (left) and ovarian corpus luteum (right) on a 12-day pregnancy examination. Serum progesterone level on day 12 was 7.95 ng/mL.
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Figure 231.15 Ultrasonographic images of an embryonic vesicle (left) and ovarian corpus luteum (right) on a 16-day pregnancy examination in the same mare as above. Serum progesterone level on day 16 was 0.7 ng/mL. The pregnancy was maintained successfully with altrenogest supplementation.
pregnant mare’s serum into cockerels resulted in enhanced comb growth.28 Biological tests for the presence of eCG have been replaced by enzyme-linked immunosorbent assays and radioimmunoassays. Two major problems exist with using eCG levels for pregnancy diagnosis. First, a false negative diagnosis (i.e. no eCG detected in a mare that is truly pregnant) can be made if a blood sample is collected prior to day 35, before endometrial cups form, or after day 120 of pregnancy after regression of the cups. Second, a false positive pregnancy diagnosis (i.e. elevated eCG detected in a mare that is truly not pregnant) can be made if a blood sample is collected from a mare that lost her pregnancy after endometrial cup formation. Mares that experience pregnancy loss after day 40 of gestation will not return to estrus until cup degeneration is complete and after all luteal tissue has regressed. Therefore, detection of elevated eCG in the blood of a mare will only confirm that endometrial cups are present and do not indicate current pregnancy status or fetal health. Persistence of endometrial cups with prolonged secretion of eCG has also been reported following abortion or the next season after a full-term pregnancy.52,57–59 Estrogens Conjugated estrogens (i.e. estrone sulfate) and total estrogens (conjugated plus unconjugated estrogens)
in blood, milk, urine, or feces have been used for diagnosis of pregnancy and as an indicator of fetal viability in domestic and feral horses.60–64 Initial equine pregnancy tests based on estrogen levels were chemical tests performed on urine samples.65 Acid hydrolysis was used to free estrogens from their conjugated form. The estrogens were then extracted from the sample and a fluorescence reaction subsequently evaluated. Cuboni (1934)33 and Kober (1935)66 tests for estrogens in urine were able to detect pregnancy after 100 to 110 days of gestation in mares. Biological tests for pregnancy in mares based on elevated estrogens relied on cornification of vaginal smears in ovariectomized mice or rats29, 30 or changes in plumage in capons31 injected with mare’s urine. Measurements of total estrogens or conjugated estrogens are now commonly performed by radioimmunoassay or enzyme immunoassay. Total estrogen and conjugated estrogen levels initially increase between days 35 and 40 of pregnancy. 48, 67, 68 This first increase in estrogens is due to production by the ovaries, since the increase is prevented by ovariectomy.48, 68 Daels and co-workers68 reported that the rise of estrogens was also eliminated in the absence of a functional corpus luteum (Figure 231.16). Equine chorionic gonadotropin produced by the endometrial cups is responsible for stimulation of estrogen production by the corpus luteum.68 This
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Group I Group II Group III
1.2 Urinary oestrogen conjugate (ng × 103 mg Cr)
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1
0.8
0.6
0.4
0.2
0 20
25
30
35
50 45 40 Day of pregnancy
55
60
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Figure 231.16 Concentrations of oestrogen conjugate in urine during early pregnancy. Group 1 consists of pregnant mares ovariectomized on day 18 to 19 of pregnancy and treated with altrenogest; Group 2 is pregnant mares and Group 3 is pregnant mares treated with altrenogest. From Daels et al.68
early rise in estrogens cannot be used as a direct indicator of pregnancy in mares. A second larger increase in estrogens begins after day 55 of pregnancy (Figure 231.17). The source of estrogen production at this stage of gestation is the feto-placental unit.69–71 The pattern of estrogen secretion during gestation mirrors the pattern of fetal gonad development. Estrogen concentrations increase by the end of the first trimester, reach peak concentrations by day 210–240, and then gradually decline over the remainder of gestation.69 Consequently, an increase in total estrogens or conjugated estrogens after day 70 to 80 of gestation can be used as a direct indicator of pregnancy. Measurement of estrogen levels after the third month of pregnancy can also be used to evaluate fetal viability. Fetal death leads to a rapid decline in total estrogen and conjugated estrogen levels.72 Determination of estrone sulfate levels in urine or serum is the basis of several stall-side pregnancy tests.73
Relaxin Relaxin is a protein hormone produced primarily by the placenta in the mare. Production of relaxin by the placenta begins at approximately 80 days of gestation (Figure 231.18). Concentrations peak between days 176 and 200 and remain elevated to term.74 Relaxin concentrations decrease rapidly following foaling and expulsion of the placenta. Measurement of relaxin has been used as an indicator of pregnancy75
and as a biomarker for placental function in the mare.76, 77 Other tests for pregnancy
Early pregnancy factor An immunosuppressive glycoprotein called ‘early pregnancy factor’ (EPF) was first described in the mouse78 and has since been reported in multiple mammalian species. The first report of EPF in mares was by Branco and colleagues in 1989.79 In early reports, EPF was detected in maternal blood using the rosette inhibition test (RIT) 7 to 10 days after ovulation.79, 80 Takagi and co-workers81 evaluated the RIT in mares before and after mating, in embryo donor mares prior to and after flushing, and in recipient mares after transfer. Both Takagi et al.81 1998 and Ohnuma et al.82 noted an increase in EPF to a level consistent with pregnancy on day 2 after ovulation in mated mares (Figure 231.19). The EPF activity in donor mares decreased to a non-pregnant level 2 days after an embryo flush procedure. Levels of EPF increased 2 to 3 days after receiving an embryo in recipient mares subsequently confirmed to be pregnant by ultrasonography. It was noted that two patterns of EPF activity were evident in recipient mares that did not become pregnant. In some mares, EPF activity remained low after transfer, suggesting that the embryo was not viable after transfer. In other mares, EPF activity increased 2 days after transfer, but subsequently decreased to non-pregnant levels by 5 days
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Figure 231.17 Concentrations of E1 (estrone-euilin-equilenin) and E2 (estradiol-17β and estradiol-17α) in the plasma of mares throughout gestation. From Nett et al.69
after transfer, suggesting that the embryos were initially viable after transfer and then died. Early pregnancy factor has been detected as early as 24–72 hours after mating, and was reported to re-
main elevated until the second trimester.82 Collectively, these studies suggest that early pregnancy factor is initially produced by the early developing embryo while still in the oviduct and that EPF may
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Figure 231.18 Plasma relaxin concentrations in Standardbred mares (±SEM) segregated by those carrying colt (open circles, n = 9) vs. filly (filled circles, n = 12) offspring. From Stewart et al.76
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(b) Figure 231.19 Concentration of early pregnancy factor in non-pregnant and pregnant mares. RIT, rosette inhibition test. From Ohnuma et al.82 Courtesy of the American Journal of Reproductive Immunology.
be a useful marker for pregnancy and embryo viability. Unfortunately, evaluations of a commercial lateral flow assay for ‘early conceptus factor’ have indicated that the test is unreliable for determination of the pregnancy status of a mare.83–85
Pregnancy specific proteins Mare pregnancy protein-1 (MMP-1)85 and β 2 pregnancy protein87 were reported to be present in pregnant mares and absent in non-pregnant mares. It was suggested that these proteins may be of maternal (uterine) origin and produced in response to a signal from the conceptus.87
Mucin (Kurasawa) test Changes in cervical-vaginal mucus have been used to differentiate pregnant from non-pregnant mares.88
Stained smears from pregnant mares are thick and dark, with globules of mucus and epithelial cells. Smears from non-pregnant mares are thin and pale and do not contain globules of mucus. However, it is currently difficult to see a use for this test as it carries the risk of introduction of infection to the vagina/cervix.
Summary Ultrasonography is the most useful tool for early pregnancy diagnosis, identification of twins, and detection of uterine or ovarian abnormalities. Endocrine tests may be helpful for pregnancy diagnosis if transrectal palpation cannot be safely performed or if the stage of pregnancy is unknown. A combination of endocrine tests, including progesterone, eCG and estrone sulfate, may be preferred over a single
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References 1. Allen WR, Wilsher S, Stewart F, Ousey J, Fowden A. The influence of maternal size on placental, fetal and postnatal growth in the horse. II. Endocrinology of pregnancy. J Endocrinol 2002;172:237–46. 2. Palmer E, Thimonier J, Lemon M. Early pregnancy diagnosis in the mare by estimation of the level of progesterone in the peripheral blood. Livest Prod Sci 1974;1:197–206. 3. van Niekerk CH. The early diagnosis of pregnancy, the development of foetal membranes and nidation in the mare. J S Afr Vet Med Assoc 1965;36:483–8. 4. Asa CS, Goldfoot DA, Ginther OJ. Assessment of the sexual behavior of pregnant mares. Horm Behav 1983;17: 405–13. 5. Hayes KEN, Ginther OJ. Relationship between estrous behavior in pregnant mares and the presence of a female conceptus. J Equine Vet Sci 1989;9:316–18. 6. Roberts SJ. Gestation and pregnancy diagnosis in the mare. Curr Ther Theriogenol 1986;2:670–8. 7. Bain AM. The manual diagnosis of pregnancy in the thoroughbred mare. New Zeal Vet J 1967;15:227–30. 8. Zemjanis R. Pregnancy diagnosis in the mare. J Am Vet Med Assoc 1961;139:543–7. 9. Solomon WJ. Rectal examination of the cervix and its significance in early pregnancy evaluation of the mare. P Annu Conv Am Equine 1971;17:73–80. 10. McKinnon AO. Diagnosis of pregnancy. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 501–8. 11. Ginther OJ. Intrauterine movement of the early conceptus in barren and postpartum mares. Theriogenology 1984;21:633–44. 12. Ginther OJ. Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains, WI: Equiservices, 1986. 13. Govaere J, Hoogewijs M, De Schauwer C, Van Zeveren A, Smits K, Cornillie P, de Kruif A. An abortion of monozygotic twins in a warmblood mare. Reprod Domest Anim 2009;44:852–4. 14. McCue PM, Thayer J, Squires EL, Brinsko SP, Vanderwall DK. Twin pregnancies following transfer of single embryos in three mares: a case report. J Equine Vet Sci 1998;18:832– 4. 15. Vullers A, Vullers F, Govaere G, Van Poucke M, Van Zeveren A, de Kruif A. Monozygotic twins after transfer of a single embryo. Reprod Domest Anim 2009;44:131. 16. Allen WR, Goddard PJ. Serial investigations of early pregnancy in pony mares using real time ultrasound scanning. Equine Vet J 1984;16:509–14. 17. Vanderwall DK, Squires EL, Brinsko SP, McCue PM. Diagnosis and management of abnormal embryonic development characterized by formation of an embryonic vesicle without and embryo in mares. J Am Vet Med Assoc 2000;217:58–63. 18. Holder RD. Fetal sex determination. In: Samper J, Pycock J, McKinnon AO (eds) Current Therapy in Equine Reproduction. Philadelphia: W.B. Saunders, 2007; pp. 343–56. 19. Hart G, Cole H. A practical method for diagnosis of pregnancy in the mare. J Am Vet Med Assoc 1932;80:604–14.
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20. Jeffcott LB, Atherton JG, Mingay J. Equine pregnancy diagnosis: a comparison of two methods for the detection of gonadotrophin in serum. Vet Rec 1969;84:80–3. 21. Santamarina E, Joven LL. Evaluation of reliability of a diagnosis test for pregnancy in mares based on the presence of gonadotrophic hormones. J Am Vet Med Assoc 1959;135:383–7. 22. Cole HH, Hart GH. Properties of gonad-stimulating principle of mares’ serum. Am J Physiol 1930;93:57. 23. Zondek H, Sulman F. A twenty-four hour pregnancy test for equines. Nature 1945;155:302–3. 24. Valle JR. Ovarian hyperemia in the immature rat as a pregnancy test in equids. Proc Soc Exptl Biol Med 1947;66: 352–4. 25. Morales CC, Damonte FR. The Friedman test in horses. Rev Fac Agron 1950;12:221–52. 26. Neto JFT. The reaction of the male toad to pregnant mare’s serum and its comparative study with the Cole–Hart Test. Am J Vet Res 1949;10:74–6. 27. Sakuma Y. The semen excretion test of male batrachia for the diagnosis of early pregnancy in mare. Tohoku J Agric Res 1952;3:69–81. 28. Bratescu I, Filimon St, Otel V. Early pregnancy diagnosis in the mare by the biological test using cockerels. Anal Inst Cerc Zooteh 1955;13:317–23. 29. Miller WC. Biological methods of diagnosing equine pregnancy. I. The mouse test. P R Soc London 1934;116: 237–47. 30. Mayer DT. A comparative study of two biologic and two chemical techniques of pregnancy diagnosis in the mare. Am J Vet Res 1944;5:209–14. 31. Greenwood AW, Blyth JSS. Biological methods of diagnosing equine pregnancy. II. The Capon Test. P R Soc London 1934;116:247–58. 32. Novazzi G. Pregnancy diagnosis in mares by the white lupin test. Profilassi 1949;22:96–100. 33. Cuboni E. A simple and rapid chemical hormonic pregnancy piagnosis. Klin Wochenschr 1934;13:302–3. 34. Lunaas T. A rapid method for the quantitative estimation of urinary oestrogens in the pregnant mare. Acta Chem Scand 1962;16:2064–5. 35. Wide M, Wide, L. Diagnosis of pregnancy in mares by an immunological method. Nature 1963;198:1017–18. 36. Chak R, Bruss M. The MIP test for diagnosis of pregnancy in mares. P Annu Conv Am Equine 1968, pp. 53–55. 37. Allen WR. Factors influencing pregnant mare serum gonadotropin production. Nature 1969;223:64–6. 38. Mitchell D. Early fetal death and serum gonotrophin test for pregnancy in the mare. Canadian Vet J 1971;12:41–4. 39. Richards CB. Simple immunological method for the diagnosis of pregnancy in mares. Nature 1967;215:1280–1. 40. Wormstrand A. Immunological pregnancy diagnosis in the mare. Acta Vet Scand 1969;10:299–308. 41. McCarthy C, Pennington G. A preliminary study of the immunoelectrophoretic properties of pregnant mares serum (PMS) together with its application to the diagnosis of pregnancy in the mare. Cell Mol Life Sci 1966;22:33–5. 42. de Coster R, Cambiaso CL, Masson PL. Immunological diagnosis of pregnancy in the mare by agglutination of latex particles. Theriogenology 1980;16:433–6. 43. Stewart F, Allen WR, Moor RM. Pregnant mare serum gonadotrophin: Ratio of follicle stimulating hormone and luteinizing hormone activities measured by radioreceptor assay. J Endocrinol 1976;71:371–82.
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44. Fay JE, Douglas RH. The use of radioreceptor assay for the detection of pregnancy in the mare. Theriogenology 1982;18:431–44. 45. Mia AS, Tierney M, Rohovsky MW. A rapid micro ELISA test for detection of equine pregnancy. Proceedings of the Conference on Research Workers in Animal Diseases 1981;62:18. 46. Squires EL, Voss JL, Villahoz MD. Immunological methods for pregnancy detection in mares. P Annu Conv Am Equine 1983;29:45–51. 47. Nett TM, Holtan DW, Estergreen VL. Oestrogen, LH, PMSG and prolactin in serum of pregnant mares. J Reprod Fertil 1975;23(Suppl.):457–62. 48. Terqui M, Palmer E. Oestrogen pattern during early pregnancy in the mare. J Reprod Fertil 1979;27(Suppl.): 441–6. 49. Menzer C, Schams D. Radioimmunoassay for PMSG and its application to in vivo studies. J Reprod Fertil 1979;55:339–45. 50. Kindahl H, Knudsen O, Madej A, Edqvist LE. Progesterone, prostaglandin F-2α, PMSG and oestrone sulphate during early pregnancy in the mare. J Reprod Fertil 1982;32(Suppl.):353–9. 51. Stabenfeldt GH, Hughes JP, Evand JW, Neely DP. Spontaneous prolongation of luteal activity in the mare. Equine Vet J 1974;6:158–63. 52. Hoffmann B, Gentz F, Failing K. Investigations into the course of progesterone, oestrogen and eCG concentrations during normal and impaired pregnancy in the mare. Reprod Domest Anim 1996;31:717–23. 53. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. 54. Holtan DW, Nett TM, Estergreen VL. Plasma progestins in pregnant, postpartum and cycling mares. J Anim Sci 1975;40:251–60. 55. Rowlands I. Serum gonadotrophin and ovarian activity in the pregnant mare. J Endocrinol 1949;6:184–91. 56. Taylor TS, Honey PG. Use of immunological pregnancy testing in mares carrying mule fetuses. Equine Pract 1980;2:25–6. 57. Allen WR, Kolling M, Wilsher S. An interesting case of early pregnancy loss in a mare with persistent endometrial cups. Equine Vet Educ 2007;19:539–44. 58. Willis LA, Riddle WT. Endometrial cups. J Am Vet Med Assoc 2004;226:877–9. 59. Steiner J, Antczak DF, Wolfsdorf K, Saville K, Brooks S, Miller D, Bailey E, Zent W. Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. 60. Bamberg E, Choi HS, Moestl E, Wurm W, Lorin D, Arbeiter K. Enzymatic determination of unconjugated oestrogens in faeces for pregnancy diagnosis in mares. Equine Vet J 1984;16:537–9. 61. Sist MD, Williams JF, Geary AM. Pregnancy diagnosis in the mare by immunoassay of estrone sulfate in serum and milk. J Equine Vet Sci 1987;7:20–3. 62. Sist MD, Youngblood MA, Williams JF. Using fecal estrone sulfate concentrations to detect pregnancies. Vet Med 1987;82:1036–8. 63. Kirkpatrick J, Kasman L, Lasley B, Turner J Jr. Pregnancy determination in uncaptured feral horses. J Wildlife Manage 1988;52:305–8. 64. Lasley B, Ammon D, Daels P, Hughes J, Munro C, Stabenfeldt G. Estrogen conjugate concentrations in plasma and urine reflect estrogen secretion in the nonpregnant and pregnant mare: a review. Equine Vet Sci 1990;10: 444–8.
65. Cox JE, Galina CS. A comparison of the chemical tests for oestrogens used in equine pregnancy diagnosis. Vet Rec 1970;86:97–100. 66. Kober S. Uber sexualhormone bei den haustieren. Klin Wochenschr, 1935;14:381. 67. Hyland JH, Wright PJ, Manning SJ. An investigation of the use of plasma oestrone sulphate concentrations for the diagnosis of pregnancy in mares. Aust Vet J 1984;61:123. 68. Daels PF, Shideler S, Lasley BLl, Hughes JP, Stabenfeldt GH. Source of oestrogen in early pregnancy in the mare. J Reprod Fertil 1990;90:55–61. 69. Nett TM, Holtan DW, Estergreen VL. Plasma estrogens in pregnant and postpartum mares. J Anim Sci 1973;37: 962–70. 70. Raeside JI, Liptrap RM, Milne FJ. Relationship of fetal gonads to urinary oestrogen excretion by the pregnant mare. Am J Vet Res 1973;34:843–5. 71. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J Reprod Fertil 1979;27(Suppl.):499–509. 72. Bosu WTK, Turner L, Franks T. Estrone sulphate and progesterone concentrations in the peripheral blood of pregnant mares: Clinical implications. Proceedings of the International Congress on Animal Reproduction and Artificial Insemination, 1984;2:78–82. 73. Henderson K, Stewart J. A dipstick immunoassay to rapidly measure serum oestrone sulfate concentrations in horses. Reprod Fertil Dev 2000;12:138–89. 74. Stewart DR, Stabenfeldt G. Relaxin activity in the pregnant mare. Biol Reprod 1981;25:281–9. 75. Ponthier J, van de Weerdt M, Deleuze S. Pregnancy diagnosis in the mare by semi quantitative relaxin quick assay kit. Reprod Domest Anim 2008;43:11. 76. Stewart DR, Stabenfeldt GH, Hughes JP, Meagher DM. Determination of the source of equine relaxin. Biol Reprod 1992;27:17–24. 77. Ryan P, Vaala W, Bagnell C. Evidence that equine relaxin is a good indicator of placental insufficiency in the mare. P Annu Conv Am Equine 1998;44:62–3. 78. Morton H, Hegh V, Clunie GJA. Immunosuppression detected in pregnant mice by rosette inhibition test. Nature 1974;249:459–60. 79. Branco MDL, Kuchembuck MRG, Papa FO, Campos Filho EP. Detection of early pregnancy in mares by the rosette inhibition test and measurement of serum progesterone. Equine Vet J 1989;8:19–20. 80. Ohnuma K, Ito K, Miyake Y–I, Takahashi J, Yasuda Y. Detection of early pregnancy factor (EPF) in mare sera. J Reprod Dev 1996;42:23–8. 81. Takagi M, Nishimura K, Oguri N, Ohnuma K, Ito K, Takahashi J, Yasuda Y, Miyazawa K, Sato K. Measurement of early pregnancy factor activity for monitoring the viability of the equine embryo. Theriogenology 1998;50:255–62. 82. Ohnuma K, Yokoo M, Kazuei I, Nambo Y, Miyake Y–I, Komatsu M, Takahashi J. Study of early pregnancy factor (EPF) in equine. Am J Reprod Immunol 2000;43: 174–9. 83. Metcalf ES, McCue PM, Jasko DJ. Evaluation of a test for equine early conception factor. P Annu Conv Am Equine 2004;50:518–20. 84. Horteloup MP, Threlfall WR, Funk JA. The early conception factor (ECFTM ) lateral flow assay for non-pregnancy
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Pregnancy Examination determination in the mare. Theriogenology 2005,64: 1061–71. 85. Parker E, Tibary A, Vanderwall DK. Evaluation of a new early pregnancy test in mares. J Equine Vet Sci 2005;25:66–9. 86. Gidley-Baird AA, Teisner B, Hau J, Grudzinskas JG. Immunochemical demonstration of a new pregnancy protein in the mare. J Reprod Fertil 1983;67:129–32.
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87. Lea RG, Bolton AE. An immunochemical demonstration of a pregnancy-specific protein in the horse and its use in the serological detection of early pregnancy. J Reprod Fertil 1988;84:431–6. 88. Day F, Miller WC. A comparison of the efficiency of methods of diagnosing equine pregnancy, with special reference to the mucin test. J Brit Vet Rec 1940;52:711–16.
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Chapter 232
Induction of Parturition Margo L. Macpherson and Dale L. Paccamonti
Introduction Parturition is an efficient, explosive event in the mare. Stage I of parturition, the phase important for initial cervical dilation and fetal positioning, generally takes a few hours. Stage II, marked by rupture of the chorioallantois and delivery of the foal, often occurs in less than 30 minutes. These highly coordinated events are crucial to the well-being of both the dam and the fetus. Altering the component parts of parturition, through medical intervention, can significantly impact maternal and fetal health. However, some conditions necessitate controlling parturition so that trained veterinary personnel are present during the delivery period. For instance, normal parturition is compromised in mares with body wall defects, ruptured prepubic tendons, or hydropic conditions. Poor abdominal press, dystocia, and/or hypovolemic shock can be consequences of these conditions. Inducing parturition can allow for prompt intervention, which can be life saving for both the mare and the foal. However, inducing parturition can also have serious consequences. Dystocia, premature placental separation, fetal hypoxia, and dysmaturity can all occur as a result of induced parturition. The risks and benefits of this procedure should be carefully reviewed so that all involved parties are prepared for the potential outcomes. A first step is to review the specific criteria that have been defined to aid in the decision-making process for inducing parturition in the mare.
Criteria for induction of parturition A prerequisite to inducing labor in a mare is determining whether the fetus is capable of surviving extrauterine life. Several physiological processes occur within the fetus prior to delivery, ensuring that it will survive after birth. The normal fetus must have appropriate energy reserves, functional lungs and gut,
and the ability to suck, swallow, and maintain body temperature after delivery.1 In most domestic species, fetal maturation is associated with a prenatal increase in adrenocortical activity a few weeks prior to birth.2 The equine fetus is unique in that there is little adrenocortical activity until 24–48 hours before birth. Final maturation of the equine fetus occurs during this period. Consequently, the equine fetus is at substantially greater risk of dysmaturity/prematurity if delivered at an inappropriate time. Several indicators have been identified that suggest fetal and maternal ‘readiness for birth’. Purvis3 described three criteria considered essential prior to inducing parturition in the mare. These criteria include adequate gestational length, mammary development and milk/colostrum production, and cervical softening. Gestational length Gestational length is often cited as a criterion prior to inducing parturition in mares. Early studies suggested that mares with a minimum gestational length of 330 days would consistently have mature foals. These mares were considered good candidates for induction.3 However, gestational length is highly variable between mares, ranging from 320 to 362 days in light horse mares.4, 5 Considering the late maturational phase of the equine fetus, not all fetuses are mature 330 days from the last breeding. Gestational length can be useful when making a decision to induce parturition if the normal gestation for a given mare is known. Most mares have a similar gestational length from year to year, so historical information can be very useful when estimating expected foaling dates and fetal maturity. However, one must consider that day length can affect gestational length.4, 6 Mares foaling during short days typically have a longer gestation, while mares foaling during long days have a shorter gestation. Given these
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Induction of Parturition several points, gestational length is an insensitive parameter for determining a mare’s readiness to give birth. This end point should only be used in conjunction with other signs when deciding to induce parturition. Mammary development and secretions Mammary development and colostrum production in the mare are considered to be the most reliable indicators of imminent parturition as well as fetal maturity.7, 8 Peaker et al.8 measured components of mammary secretions prepartum and found that calcium rose to > 10 mol/L 1 to 6 days before foaling. This was followed by reports supporting the finding of a rapid prepartum rise in calcium in mammary secretions.7, 9 Interestingly, a similar abrupt rise in calcium occurs in allantoic fluid shortly before parturition.10 Not only does milk calcium rise rapidly, but the relative concentrations of sodium and potassium invert, with potassium concentrations becoming greater than sodium in the last few days of gestation. Based on these characteristic changes, mammary secretions were deemed to be a very good way to assess fetal maturity or ‘readiness for birth’.7, 9 Foals born from mares induced to foal when calcium was < 10 mmol/L had a poorer chance for survival compared with foals induced when calcium was > 10 mmol/L.7, 9 Precise measurement of mammary secretion electrolyte concentrations requires a flame spectrophotometer or a laboratory chemistry analyzer. Using these systems, elevation of mammary secretion calcium above 40 mg/dL and potassium concentration greater than sodium generally indicates fetal maturity in the normal equine pregnancy.7–9 However, changes in mammary secretion electrolytes most commonly occur at night, coincident with the period when the majority of mares foal. Laboratory evaluation of mammary secretions is impractical during this time period for the field practitioner. Several stall-side tests are available to measure calcium (Ca2+ ) concentration in mammary secretions. Stall-side test kits are modified water hardness tests, which often measure both divalent cations, magnesium and calcium, from a single milk sample. 11–13 Measurement of both cations complicates interpretation of results as correlation with fetal maturity is based on concentrations of calcium, and not magnesium, in mammary secretions. Furthermore, the concentration of magnesium in mammary secretions increases earlier than calcium and the rise is more gradual. The peak concentration of magnesium is reached earlier than that of calcium, and magnesium often declines at parturition.13, 14 Therefore, if a test measuring both cations is used to
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decide when to induce parturition, high magnesium measured by the test might prematurely indicate that the fetus was ‘ready’ for birth. The calcium concentration in the secretions might still be low, and calcium is a more accurate indicator of fetal maturity. As such, if mammary secretion electrolytes are used to decide when to induce parturition, it is critical that only calcium is measured – whether using a laboratory assay or a water hardness kit that measures only calcium. When stall-side tests for measuring mammary secretion electrolytes are the only option, accurate interpretation of the calcium values must be considered. Specifically, 10 mmol/L (400 ppm) calcium in mammary secretions has been well established as a good indicator of fetal readiness for birth.7–9,14 Some studies have referred to 200 or 250 ppm calcium carbonate, measured using a stall-side test, as benchmarks for readiness for birth.15, 16 However, these values carry potential danger if used in assessing readiness for induction of parturition. Calcium in milk is not in the form of calcium carbonate. Results from tests that measure calcium carbonate in a solution should be converted to calcium for correct interpretation. To convert ppm calcium carbonate to ppm calcium, results should be divided by 2.5 because the molecular weight of calcium carbonate is 100 and the molecular weight of calcium is 40 (100/2.5 = 40) (ppm is derived by multiplying mmol × molecular weight). Therefore, 200 and 250 ppm calcium carbonate are equivalent to only 80 and 100 ppm calcium, respectively, or 2 and 2.5 mmol calcium carbonate or calcium (mmol are not dependent on molecular weight). If these values were used (calcium carbonate concentration) to elect to induce parturition, it is very likely that the fetus would be dysmature after delivery. An additional factor to consider is the dilution of mammary secretions prior to using a water hardness test for measuring calcium (carbonate). For example, when using a commercially available test in the USA (Titrets; Chemetrics, Calverton, VA), a convenient dilution ratio to use is one part milk to four parts water. The test result should then be multiplied by a factor of five to correct for the dilution effect. If both corrections for conversion of calcium carbonate to calcium and the dilution ratio are considered, the overall correction factor is two. Here is an example calculation: (Test result × 5) to correct for dilution effect (Test result/2.5) to convert calcium carbonate to calcium (Test result × 5/2.5 = test result × 2)
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The dilution value may be different for different tests; therefore, close attention should be paid to this point. For instance, another commercially available test in the USA (FoalWatch; Chemetrics, Calverton, VA) employs a 1:7 dilution (milk:diluent). In this case, if one measures 200 ppm calcium carbonate (uncorrected results), which was cited as a predictor of foaling in one study,16 the actual calcium concentration is 560 ppm calcium, equivalent to 14 mmol/L. These values were in agreement with the earlier reports of adequate calcium concentrations as indicators of fetal readiness for birth.7, 9, 13 In some cases, mammary secretion electrolyte concentrations may not reliably indicate impending parturition. Samples taken early in the day may be low and not reflect electrolyte changes that occur in the evening or at night, shortly before parturition (when calcium concentrations often increase). Sampling should occur as late in the day as possible, particularly when induction is considered. If induction must be scheduled during the daytime (to ensure adequate personnel are present), samples can be taken late in the day prior to potential induction and again first thing in the morning. Comparing values can be useful prior to making a final decision for induction. Parity can also affect changes in mammary secretion electrolytes. Multiparous mares show more consistent changes in electrolyte values prior to foaling. Primiparous or ‘maiden’ mares often have slow mammary gland development, and mammary secretion electrolytes may not change until immediately prior to parturition. Vigilant monitoring of mammary secretion electrolytes is necessary to predict impending parturition in maiden mares. Maiden mares can be especially challenging when considering induced parturition. Also challenging are mares with placental pathology, such as placentitis. Mares with abnormal placentas show precocious mammary gland development and secretion electrolyte changes, most commonly a precocious rise in calcium.17 Rossdale and coworkers17 monitored mammary secretion electrolytes in 25 mares with abnormal pregnancies. Seventeen mares (17/25, 68%) had elevation of mammary secretion calcium greater than 10 mmol/L before 310 days of gestation. Placental pathology was identified in 16 (16/17, 94%) mares with prematurely elevated mammary secretion calcium. Ten of the 16 foals (63%) were either delivered dead or died shortly after birth. Results from these studies indicate that changes in mammary secretion calcium from mares with abnormal pregnancies should be interpreted cautiously. When placental pathology is present, sodium and potassium values can sometimes provide additional information that indicates impending parturition.
Cervical softening Cervical softening is the final criterion cited by Purvis3 as critical to the success of induced parturition. The importance of cervical relaxation prior to induction has been a controversial point. Studies in humans18–20 have associated poor cervical relaxation with failed induction, prolonged labor, and a high cesarean birth rate. Early reports examining the role of cervical dilation prior to induction in mares21, 22 suggested that outcome was successful in mares having a tightly closed, mucus-covered cervix. More recent studies in mares23, 24 showed a positive effect of cervical dilation on neonatal viability after induced parturition. Mares with a spontaneously dilated cervix (determined by digital examination per vagina) prior to induction delivered their foals more quickly than those mares with a closed cervix.23 Foals delivered after a shorter interval from initial administration of oxytocin were better adapted after birth. Foals delivered from mares with a dilated cervix stood and nursed more quickly and had fewer signs of intrapartum asphyxia (hypercapnia, maladjustment) than foals from mares with a closed cervix. Mares suffering intrapartum complications (premature placental separation, dystocia) all had a non-dilated cervix prior to induction. Rigby and co-workers24 further investigated the role of pre-induction cervical dilation on foal outcome by using prostaglandin E2 (PGE2 ) to initiate cervical relaxation prior to induction. Prostaglandin E2 has been administered to women to promote cervical softening and dilation before inducing labor.25–27 In women, PGE2 treatment significantly enhanced cervical dilation and effacement, shortened the induction–delivery interval, reduced induction failures, and lowered cesarean section rate in some studies.28, 29 Rigby et al.24 treated seven mares with 2.5 mg intracervical PGE2 and four mares with intracervical saline. Pretreatment cervical dilation was assessed in all mares by determining how many fingers would easily penetrate the cervix. Differences were not seen between PGE2 -treated and salinetreated mares for number of oxytocin treatments required to induce chorioallantois rupture. Mean intervals from initial injection of oxytocin to rupture of the chorioallantois or delivery of the foal also did not differ between treatment groups. Mares treated with PGE2 tended to have greater cervical dilation at the time of induction than saline-treated mares. Foals delivered from PGE2 -treated mares suckled more quickly than foals from saline-treated mares. While treatment groups did not differ statistically in this study, the authors concluded that cervical softening prior to induction of parturition favored a shorter delivery period and positively impacted neonatal
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Induction of Parturition adaptability. Additionally, the authors acknowledged that PGE2 treatment enhanced cervical softening, but they speculated whether cervical manipulation alone may have contributed to cervical changes seen prior to induction in this study. In summary, several factors can contribute to ensuring the success of induced parturition in the mare. Adequate udder development, changes in mammary secretion electrolytes, and cervical softening are all important considerations prior to induction.
Methods of induction A variety of agents and methods have been used to induce parturition in the mare, including administration of glucocorticoids, prostaglandins, and oxytocin. Glucocorticoids have shown limited efficacy for inducing parturition in the mare,30–32 given the need for high doses over multiple days to initiate parturition. This method is not practical for the field practitioner. However, interesting work by Ousey et al.33 showed that treating normal mares with 100 mg dexamethasone for 3 days (similar to the protocol for induction), starting on day 315 of gestation, would not only shorten gestation but also result in delivery of a mature foal. While the use of glucocorticoids to induce parturition in most mares would not be practical, this tool might be useful for salvaging a high-risk pregnancy by initiating early fetal maturation and delivery. An important clinical finding from this study was that supplementation with colostrum was necessary in several of the foals from mares induced to foal after dexamethosone treatment. Exogenous prostaglandins have been successful in initiating parturition in the mare. Both natural and synthetic prostaglandins produce potent myometrial activity in the mare.34, 35 However, natural prostaglandin (PGF2α ) has not proven to be a reliable induction agent for the mare due to a higher number of peripartum abnormalities (premature placental separation, dystocia, cracked ribs in foals) and prolonged time from drug administration to delivery of a foal.36 Synthetic prostaglandins (fluprostenol, fenprostalene, prostalene) have been more reliable for initiating parturition in the mare,34, 36, 37 but these products are no longer commercially available. Cloprostenol, a synthetic prostaglandin that is readily available for use in horses, has not been critically tested as an induction agent. Oxytocin has been widely used for inducing parturition in the mare.3, 21, 23,38–40 Oxytocin has a rapid effect resulting in delivery 15–90 minutes following administration. Various methods and doses of oxytocin induction have been described. Single-dose bolus injection (2.5–120 IU oxytocin, i.v.,
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i.m. or subcutaneously),3, 21, 23, 38, 39 repeated injections (2.5–20 IU oxytocin, i.v., i.m., or subcutaneously) at 15–20-minute intervals,23, 40 or intravenous drip (60–120 IU oxytocin in 1 liter saline delivered at a rate of 1 IU/min)23 are all proven methods for inducing parturition. Workers in Texas23 compared different methods of oxytocin administration for inducing parturition in mares to determine if method of administration had untoward effects on the foal. Three treatments were used to induce parturition: (i) a bolus intramuscular injection of 75 IU oxytocin; (ii) intramuscular injection of 15 IU oxytocin every 15 minutes with a maximum dose of 75 IU; or (iii) intravenous administration of oxytocin in 1 L 0.9% NaCl at a rate of 1 IU/min for a maximum dose of 75 IU. Delivery time intervals and foal vitality parameters were measured. The method of oxytocin administration did not affect the overall time interval from initial oxytocin administration to delivery of the foal. Furthermore, neonatal blood gas values, vitality assessments, and plasma cortisol concentrations were similar between foals from different treatment groups. Across treatment groups, longer delivery times resulted in reduced foal vigor (standing, sucking) and vitality. Mares that had a softened cervix prior to induction had shorter delivery times and more vigorous foals. The authors concluded that method of induction had little impact on foal viability, even with the high doses of oxytocin used in some mares. Case selection and adherence to criteria for induction were cited as critical factors for a successful induction. Dose of oxytocin administered to induce parturition has also been examined. Administration of very low doses of oxytocin has been advocated as a safer means for inducing parturition in the mare. Pashen 40 first investigated administration of 2.5–10 IU oxytocin i.v., to induce parturition in four pony mares and one jenny at 323–334 days of gestation. Mares foaled following administration of 2.5–10 IU i.v. in total. Additionally, plasma prostaglandin metabolite (PGFM) profiles were similar to those seen in spontaneously foaling mares.41 Pashen concluded that low doses of oxytocin were physiological and efficacious for inducing parturition in mares. Camillo and coworkers11, 38 repeated this work with a slight modification in protocol. Mammary secretion calcium concentrations were determined in 24 Haflinger pony mares at 320 days gestation. Mares having 8 mmol/L Ca2+ were considered near to foaling and one dose of 2.5 IU oxytocin was administered intravenously. Mares that did not foal within 1 hour of oxytocin administration were judged not physiologically ready for parturition. The same mares received a second dose of oxytocin (2.5 IU, i.v.) the following evening, and then at a similar time, once daily, until foaling. Fourteen of 17 mares (14/17, 82%) foaled after the
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first injection of oxytocin, one mare foaled after oxytocin administration on day 2, and two mares foaled after oxytocin on day 3. The group concluded that a single, low-dose injection of oxytocin was efficacious for inducing parturition in mares. Furthermore, the workers suggested that this induction scheme would work only in mares that had a mature fetus that was physiologically ready for delivery. They felt that mares foaling on days subsequent to the initial injection did not respond to oxytocin because the fetus was in the final maturational phases prior to delivery. Workers from France42 further investigated the efficacy of low-dose oxytocin for inducing parturition in the mare. The objectives of the work were to test low-dose oxytocin in larger breed mares (light breed and draft horses), and to test a range of doses (0.1–2.0 IU i.v., once daily). Mares were administered oxytocin once mammary secretion calcium measured 13 mmol/L. Treatment was initiated at 17.00 hours. More mares foaled after administration of 2.0 IU oxytocin than with lower doses; however, only 60% of mares responded after the first injection (day 1). Mares administered 2 IU oxytocin retained fetal membranes more frequently than untreated control mares. The authors concluded that low-dose oxytocin was not a reliable tool for inducing parturition in mares, resulted in a higher incidence of retained fetal membranes, and that foals had lower cortisol levels at birth (indicating prematurity). The authors speculated that higher doses of oxytocin (3–5 IU) might have provided a more reliable induction response in larger breed horses tested. A recent study tested low-dose oxytocin (2.5 IU i.v.) for inducing parturition in a large, field population of mares.43 The goals of this study were to support or refute results from previous low-dose oxytocin studies by applying the method to a large population of mares (n = 350). One hundred and seventysix mares were allowed to foal spontaneously and 174 mares were subjected to treatment with oxytocin. Mares were enrolled in the study when results from a commercially available test for mammary secretion electrolytes (Foalwatch; Chemetrics, Calverton, VA) reached a minimum of 200 ppm calcium carbonate, which is equivalent to 560 ppm calcium given the dilution ratio used (one part milk to seven parts diluents). Mares were administered 3.5 IU oxytocin (i.v.) once daily, and monitored for foaling. The majority (69%) of mares foaled within 120 minutes of an injection; however, only 51% of mares foaled on the first day of treatment. Roughly one-third of mares foaled greater than 2 hours post-injection. All foals were normal after delivery and few mares experienced dystocia (treatment n = 2; control n = 3). The authors concluded that the method was moderately efficacious for inducing parturition. Important points
to consider if using this technique would include continued surveillance of mares after administration of oxytocin to ensure assistance during delivery for mares foaling after the expected time interval (2 hours).
Conclusions In summary, several factors impact the success of induced parturition in the mare. Fetal readiness for birth is paramount to survival of the foal after birth. Changes in mammary secretion electrolytes may be used to predict fetal maturity prior to induced parturition. Cervical relaxation can facilitate expedient delivery and improve postpartum neonatal vigor. Oxytocin is the current agent of choice for induction of parturition in the mare. Method of oxytocin administration does not impact neonatal adaptability after induced birth. Low-dose oxytocin is effective for inducing parturition in the mare, particularly in mares that are close to natural foaling.
References 1. Silver M, Fowden A. Induction and labour in domestic animals: endocrine changes and neonatal viability. In: Kunzel W, Jobim MIM (eds) The Endocrine Control of the Fetus. Berlin: Springer-Verlag, 1988; pp. 401–10. 2. Liggins GC. Adrenocortical-related maturational events in the fetus. Am J Obstet Gynecol 1976;126:931–9. 3. Purvis AD. The induction of labor in mares as a routine breeding farm procedure. Proceedings of the 23rd Annual Conference of the American Association of Equine Practitioners 1977;23:145–60. 4. Howell C, Rollins W. Environmental sources of gestation length in the mare. J Anim Sci 1951;10:805. 5. Laing JA, Leech FB. The frequency of infertility in Thoroughbred mares. J Reprod Fert Suppl 1975;23: 307–10. 6. Hodge SL, Kreider JL, Potter GD, Harms PG, Fleeger JL. Influence of photoperiod on the pregnant and post partum mare. Am J Vet Res 1982;10:1752–5 7. Ousey JC, Dudan F, Rossdale PD. Preliminary studies of mammary secretions in the mare to assess fetal readiness for birth. Equine Vet J 1984;16:259–63. 8. Peaker M, Rossdale PD, Forsyth IA, Falk M. Changes in mammary development and composition of secretion during late pregnancy in the mare. J Reprod Fertil Suppl 1979;(8)555–61. 9. Leadon DP, Jeffcott LB, Rossdale PD. Mammary secretions in normal spontaneous and induced premature parturition in the mare. Equine Vet J 1984;16:256–9. 10. Paccamonti D, Swiderski C, Marx B, Gaunt S, Blouin D. Electrolytes and biochemical enzymes in amniotic and allantoic fluid of the equine fetus during late gestation. Biol Reprod Monogr 1995;1:39–48. 11. Camillo F, Cela M, Romagnoli S. Day-time management of the foaling mare: use of a rapid mammary Ca++ determination followed by a low dose of oxytocin. 1992;2:883. 12th International Congress on Animal Reproduction. The Hague, Netherlands 1992:883–85.
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Induction of Parturition 12. Cash RSG, Ousey JC, Rossdale PD. Rapid strip test method to assist management of foaling mares. Equine Vet J 1985;17:61–2. 13. Ousey JC, Delclaux M, Rossdale PD. Evaluation of 3 strip tests for measuring electrolytes in mares pre-partum mammary secretions and for predicting parturition. Equine Vet J 1989;21:196–200. 14. Rook JS, Braselton WE, Nachreiner RF, Lloyd JW, Shea ME, Shelle JE, Hitzler PR. Multi-element assay of mammary secretions and sera from periparturient mares by inductively coupled argon plasma emission spectroscopy. Am J Vet Res 1997;58:376–8. 15. Ley WB, Hoffman JL, Meacham TN, Sullivan TL, Kiracofe RL, Wilson ML (1989b), Daytime Management of the Mare.1. Pre-Foaling Mammary Secretions Testing, Journal of Equine Veterinary Science 9:88–94 16. Ley WB, Bowen JM, Purswell BJ, Irby M, Greivecrandell K. The sensitivity, specificity and predictive value of measuring calcium-carbonate in mares prepartum mammary secretions. Theriogenology 1993;40:189–98. 17. Rossdale PD, Ousey JC, Cottrill CM, Chavatte P, Allen WR, McGladdery AJ. Effects of placental pathology on maternal plasma progestagen and mammary secretion calcium concentrations and on neonatal adrenocortical function in the horse. J Reprod Fertil 1991;44: 579–90. 18. Brindley BA, Sokol RJ. Induction and augmentation of labor: basis and methods for current practice. Obstet Gynecol Surv 1988;43:731–40. 19. Clark SL, Miller DD, Belfort MA, Dildy GA, Frye DK, Meyers JA. Neonatal and maternal outcomes associated with elective term delivery. Am J Obstet Gynecol 2009;200: 156. 20. Schlembach D, MacKay L, Shi L, Maner WL, Garfield RE, Maul H. Cervical ripening and insufficiency: From biochemical and molecular studies to in vivo clinical examination. Eur J Gynecol Reprod Biol. 2009; 144 Supple 1: S70–6. 21. Jeffcott LB, Rossdale P. A critical review of current methods for induction of parturition in the mare. Equine Vet J 1977;9:208–15. 22. Meyers SA, Leblanc MM. Induction of parturition – clinical considerations for successful foalings. Vet Med 1991;86:1117–21. 23. Macpherson ML, Chaffin MK, Carroll GL, Jorgensen J, Arrott C, Varner DD, Blanchard TL. Three methods of oxytocin-induced parturition and their effects on foals. J Am Vet Med Assoc 1997;210:799–803. 24. Rigby S, Love C, Carpenter K, Varner D, Blanchard T. Use of prostaglandin E-2 to ripen the cervix of the mare prior to induction of parturition, Theriogenology 1998;50:897–904. 25. Ferguson JE, Ueland FR, Stevenson DK, Ueland K. Oxytocin-induced labor characteristics and uterine activity after preinduction cervical priming with prostaglandin-E2 intracervical gel. Obstet Gynecol 1988;72:739–45. 26. Hales KA, Rayburn WF, Turnbull GL, Christensen HD, Patatanian E. Double-blind comparison of intracervical and
27.
28.
29.
30. 31.
32. 33.
34.
35.
36.
37.
38.
39. 40. 41.
42.
43.
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intravaginal prostaglandin E(2) for cervical ripening and induction of labor. Am J Obstet Gynecol 1994;171:1087–91. Rayburn WF. Prostaglandin-E2 gel for cervical ripening and induction of labor – a critical analysis. Am J Obstet Gynecol 1989;160:529–34. Smith CV, Rayburn WF, Connor RE, Fredstrom GR, Phillips CB. Double-blind comparison of intravaginal prostaglandin-E2 gel and chip for preinduction cervical ripening. Am J Obstet Gynecol 1990;163:845–7. Stewart JD, Rayburn WF, Farmer KC, Liles EM, Schipul AH, Stanley JR. Effectiveness of prostaglandin E-2 intracervical gel (Prepidil), with immediate oxytocin, versus vaginal insert (Cervidil) for induction of labor. Am J Obstet Gynecol 1998;179:1175–80. Alm CC, Sullivan JJ, First NL. Dexamethasone induced parturition in mare. J Reprod Fertil Suppl. 1975;23:637–40. Alm CC, Sullivan JJ, First NL. Induction of premature parturition by parenteral administration of dexamethasone in mare. J Am Vet Med Assoc 1974;165:721–2. First NL, Alm CC. Dexamethasone-induced parturition in pony mares. J Anim Sci 1977;44:1072–5. Ousey JC, Kolling M, Allen WR. The effects of maternal dexamethasone treatment on gestation length and foal maturation in Thoroughbred mares. Anim Reprod Sci 2006;94:436–8. Ley WB, Hoffman JL, Crisman MV, Meacham TN, Kiracofe RL, Sullivan TL. Daytime foaling management of the mare. 2. Induction of parturition. J Equine Vet Sci 1989;9:95–9. Rossdale PD, Jeffcott LB, Allen WR. Foaling induced by a synthetic prostaglandin analog (fluprostenol). Vet Rec 1976;99:26–8. Alm CC, Sullivan JJ, First NL. Effect of a corticosteroid (dexamethasone), progesterone, estrogen and prostaglandinF2alpha on gestation length in normal and ovariectomized mares. J Reprod Fertil 1975; 637–40. Rossdale PD, Pashen RL, Jeffcott LB. The use of synthetic prostaglandin analogue (fluprostenol) to induce foaling. J Reprod Fertil Suppl 1979;27:521–9. Camillo F, Marmorini P, Romagnoli S, Cela M, Duchamp G, Palmer E. Clinical studies on daily low dose oxytocin in mares at term. Equine Vet J 2000;32:307–10. Hillman RB. Induction of parturition in mares. J Reprod Fertil Suppl 1975;23:641–4. Pashen RL. Low-doses of oxytocin can induce foaling at term. Equine Vet J 1980;12:85–7. Pashen RL, Allen WR. Endocrine changes after fetal gonadectomy and during normal and induced parturition in the mare. Anim Reprod Sci 1979;2:271–88. Chavatte-Palmer P, Arnaud G, Duvaux-Ponter C, Zanazi C, Gerard M, Ponter A, Kindahl H, Clement F. The use of microdoses of oxytocin in mares to induce parturition. Theriogenology 2002;58:837–40. Villani M, Romano G. Induction of parturition with daily low-dose oxytocin injections in pregnant mares at term: Clinical applications and limitations. Reprod Domest Anim 2008;43:481–3.
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Chapter 233
Parturition Bruce W. Christensen
Dystocia is not a common occurrence in the mare, but when it does occur it rapidly becomes a critical situation with regard to the life of the foal and potentially the mare. For this reason, there is great interest in observing each foaling event in order to intervene early if a problem arises. As with other mammalian species, labor in the mare can be divided into three stages. Stage one involves preparatory changes leading up to stage two, which is the expulsion of the fetus. Stage three is the expulsion of the fetal membranes.
stage of parturition compared with mares not supplemented with progestagens (specifically altrenogest5 ). Foals born to mares supplemented with altrenogest showed maladaptive changes compared with control foals, including lower initial respiratory rates (52 ± 4 vs 88 ± 10 bpm), higher pH (7.41 ± 0.01 vs 7.37 ± 0.01), and higher base excess. Base cortisol concentrations were also higher in foals born to treated versus untreated mares.5
Estrogens
Endocrinology Progestagens Maternal concentrations of progesterone (P4), which are high in early pregnancy, fall during midpregnancy and remain low thereafter1 until a rise at the very end of gestation. In addition to P4, various related 5α-pregnanes produced by the feto-placental unit rise during the final months of gestation.2 It has been proposed that the active progestagen in late pregnancy in the mare, as in the sheep, may be 3β-hydroxy-5-pregnen-20-one (P5), a precursor to P4 in the steroidogenic pathway, and that a significant amount of P5 could be produced by the fetal adrenal glands, which would then aid the fetoplacental unit in pregnancy maintenance during the second half of pregnancy.3 The specific effects of these progestagens have yet to be determined, especially their ability to induce endometrial quiescence in the mare. Though overall progestagen concentrations are decreasing prior to parturition, they do not reach baseline concentrations until after parturition. Mares foal in the face of these relatively elevated progestagen concentrations.4 Mares supplemented with exogenous progestagens will foal despite continued supplementation, but spend more time in the second
Maternal estrogen concentrations are elevated during the second half of pregnancy, highest during the seventh and eighth months, falling gradually until term, and then rapidly returning to baseline after foaling.6 Estrogens play an important role at parturition in supporting myometrial contractility by stimulating prostaglandin production, and increasing oxytocin receptors and myometrial gap junctions.7, 8 The major source of these estrogens is the fetal gonads. The exception to the general, gradual decrease in estrogens is the notable rise in estradiol-17β (E2), which is known in other species to induce prostaglandin F2α (PGF2α ) synthesis.9, 10 The rise in E2 may also account for the presence of oxytocin receptors in the pregnant mare endometrium throughout late gestation,11 the release of oxytocin from the posterior pituitary, the release of PGF2α from the endometrium, and the formation of myometrial gap junctions.12 A study in the mare demonstrated that removing fetal gonads resulted in a drop in maternal plasma total estrogen concentrations, lower 13,14-dihydro-15oxo-PGF-2α (PGFM) concentrations, and weaker myometrial contractions compared with sham-operated mares.13 In non-human primates, increased estrogen concentrations are associated with a change in the amplitude and frequency of oxytocin pulses.14 Earlier in pregnancy, myometrial contractures are low
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Parturition amplitude and irregular waves, resulting in a tonic uterus. At term, higher estrogen levels mediate increased oxytocin pulses, resulting in the strong, progressive uterine contractions that are a part of labor. This illustrates that even though overall estrogens are decreasing late in pregnancy, local estrogen effects may be increasing. Relaxin Relaxin concentrations are elevated during the second half of pregnancy and are thought to originate from the placenta. During the second stage of labor, relaxin spikes in the maternal serum concentration, and then falls to baseline after stage three parturition is complete.15, 16 Oxytocin has been indicated as the hormone that initiates relaxin release from the placenta.16 Whether or not relaxin inhibits uterine activity in the mare has yet to be determined. Studies in the rat have shown that relaxin causes cervical softening,17, 18 and the same may be true in the mare. Cortisol A rise in fetal cortisol occurs in the day or two prior to parturition. This rise is not due to a rise in adrenocorticotropic hormone (ACTH), which may be high previous to fetal readiness for birth, but more importantly is the result of maturation of the fetal adrenal glands, with receptors able to respond to the ACTH and trigger cortisol secretion.19 In other species, the rise in cortisol has been shown to be important in the maturation of multiple fetal organ systems, causing surfactant release into fetal lungs and activating enzyme systems in the gastrointestinal system, liver, and placenta.20, 21 In the placenta, cortisol stimulates the steroidogenic pathways that convert cholesterol to P4 further towards estrogen production. The resultant drop in P4 and rise in estrogens causes a relaxation of the cervix and a cessation of the P4 block on uterine contractility. It has been suggested that this drop in P4 also allows the release of oxytocin from the pituitary, further relaxing the cervix and inducing myometrial contractions.3 The rapid rise in estrogen concentrations leads to PGF2α production, which induces myometrial contractions and signals the beginning of active parturition.22 Prostaglandins In other species, prostaglandin E2 (PGE2 ) is secreted in greater concentrations in the last portion of gestation and presumably acts synergistically with relaxin to help soften the cervix.23 For up to 3 hours prior to stage two, prostaglandins rise gradually, enhancing myometrial contractility and sensitizing the endometrium to
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further prostaglandin release. Following this period, and corresponding with the strong uterine contractions of second-stage labor, prostaglandins rise quickly and precipitously.13 Oxytocin The entry of the fetus into the birth canal causes physical stretching of the cervix and cranial vagina, eliciting a neuroendocrine reflex release of oxytocin from the posterior pituitary (the Ferguson reflex). Oxytocin, as its name suggests, has a powerful oxytocic effect on the pregnant uterus, causing marked contractions. Exogenous oxytocin causes induction of parturition, in part by inducing a rise in endogenous prostaglandin concentrations.24 It is likely that endogenous oxytocin has a similar effect. Oxytocin remains basal throughout pregnancy until stage-two labor,11 when highest concentrations occur between rupture of the chorioallantois and completion of delivery of the foal. Peak concentrations are noted with the appearance of the fetal head (corresponding, perhaps, with engagement of the fetal shoulders within the birth canal). Concentrations of maternal oxytocin do not return to baseline until expulsion of the fetal membranes.
Predicting parturition The gestational length for the mare is highly variable, with normal pregnancies usually ranging from around 330 to 360 days (315 to 350 for pony breeds). Mares with placentitis or other placental pathology may foal earlier than 320 days and still deliver a viable, though small, foal.25 The mechanism of this precocious maturation is not yet known. With support, other foals born between 300 and 320 days can survive, though the prognosis is guarded and based on the individual degree of physiological maturity of the foal. See Chapter 233 on prematurity and dysmaturity for more information. Most mares foal during night-time hours (usually between 9 p.m. and 11 p.m. or 2 a.m. and 6 a.m.26 ); it has been said that the fetus determines the day, but the mare determines the hour. This high degree of control by the mare has been attributed to central nervous system control over endogenous oxytocin release.11 Because the day of parturition is highly variable and unpredictable in relation to ovulation or breeding dates, and the fact that most mares will foal at a time of the day when workers are not readily available, there is great interest in predicting the night when workers will most likely need to lose sleep, rather than having workers available every night. Some specific methods of determining fetal maturity have been addressed in Chapter 233 in
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reference to determining a date when induction of parturition may be safely attempted. Some of these methods have value in predicting the date of natural parturition and will be discussed again briefly here. Physical signs Physical changes accompany the prepartum mare as she gets close to her anticipated foaling date. Sacrosciatic ligaments soften, the cervix relaxes, and the vulva relaxes and lengthens. Mammary glands develop and begin to fill with colostrum, which collects on the tip of each teat and forms a waxy residue (known appropriately as ‘waxing’). Some mares may drip, or even stream milk in the days or hours approaching foaling, which may decrease the colostral quantity and quality.27 Using these physical signs to predict the day of parturition will frustrate any horse worker as they are all highly variable and unpredictably correlated with the actual foaling date, especially in maiden mares. The evaluation of ligament softening and vulvar changes is subjective and these changes may occur any number of days prior to parturition. Ligaments may relax, tighten, and relax again a number of times prior to the expected date. Some mares, especially maidens, are notorious for not displaying any appreciable mammary development until hours before, or even a day or two after, parturition.
ways secrete calcium at the same rate as multiparous mares.30 Most mares (99%) with mammary secretion calcium concentrations above 200 ppm foaled within 72 hours of the test. Those mares (1%) going longer were multiparous mares known for heavy milk production in subsequent lactations.30 In another study, calcium was found to remain constant at a mean concentration of 5 mmol/L until 3 days prior to foaling, and then increase to greater than 10 mmol/L on the day of foaling.31
Sodium and potassium Workers have evaluated other electrolytes in mammary secretions in addition to calcium. In particular, sodium and potassium may give valuable information about approaching parturition. Sodium decreases and potassium increases significantly immediately prior to parturition.31 Potassium concentrations increased gradually to a mean value of 30 mmol/L 3 days prior to parturition and then increased rapidly to a mean value of 47 mmol/L on the day prior to parturition. Sodium concentrations decreased from approximately 87 mmol/L 3 days prior to parturition to 37 mmol/L on the day prior to parturition.31 From the period of inversion, i.e., when the potassium value becomes greater than the sodium value, mares tend to foal 24–36 hours later.31–33 These electrolyte correlations are not reliable in mares with pathologies, such as placentitis.
Mammary secretion electrolytes
Calcium
Jennies
Given the unreliability of physical changes and the wide variation of possible individual gestational lengths, other parameters have been investigated to predict better the day of natural foaling in the mare. Measurements of mammary secretion electrolytes may help. Water hardness test strips that evaluate calcium have not proven helpful in determining on which day the mare will foal, but have been helpful in determining on which day she will not foal.28 This is due to the fact that calcium concentrations may remain relatively elevated (which is the extent of the analysis for a qualitative test) for one to many days prior to foaling, but the mare is less likely to foal if calcium concentrations are low. As before, maiden mares may be the exception to this. A quantitative colorimetric test (FoalWatchTM ; CHEMetrics, Inc., Calverton, VA) has proven more predictive.29 This test gives a quantitative assessment of mammary secretion calcium concentrations in parts per million (ppm). Most mares (98%) with mammary secretion calcium concentrations less than 200 ppm will not foal within 24 hours of the test. The 2% false negative results are once again reportedly from maiden mares, which apparently do not al-
Similar changes have been noted in the jenny.34 Mammary secretion calcium and potassium concentrations progressively rose, and sodium concentrations fell, in the days prior to parturition. Calcium concentrations reached 10.3 mmol/L (± 0.65 mmol/L) on the day of foaling, no different from the mare. Much like the mare, in some jennies this value was reached as many as 4 days prior to foaling, and so is more useful as a predictor of when the jenny will likely not foal (if < 10 mmol/L) rather than when she will foal. Sodium and potassium concentrations decreased and increased, respectively, at different rates than in the mare. In the jenny, sodium decreased more gradually and potassium showed two periods of significant increase (between days 10 and 8 and again between days 6 and 4 prior to parturition). Sodium and potassium concentration showed an inversion in the jenny 24–48 hours prior to foaling.
Stage one of parturition The first stage of parturition in the mare includes both physiological changes and external signals. It is during this stage that the fetus changes from a
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Parturition dorsopubic to a dorsosacral position.35 The cervix and pelvic ligaments soften. Endometrial contractions start, but are not visible externally as abdominal contractions. The mare may act colicky, observing her abdomen, circle, and alternate between lateral recumbency, standing, and rolling. The mare may exhibit pollakiuria. Sweating may be noticed first under the tail and mane, but eventually spreads to cover the entire body. Usually stage one parturition lasts 30 minutes to 4 hours, but can in rare instances be interrupted and last for a few days. The transition from stage one to stage two is marked by the rupture of the chorioallantoic membrane at the cervix and the passage of the allantoic fluid (‘breaking of the water’).
Stage two of parturition Stage two commences with the rupture of the chorioallantois and ends with the complete passage of the fetus. The allantoic fluid, expelled at the rupture of the chorioallantois, is the urine of the fetus and is yellow in color and generally is between 8 and 15 liters in volume.36 The first visible part of the fetus is one of the front hooves, usually covered still by the amnion, followed a few centimeters behind by the second front hoof. The second hoof will appear usually in line with the cannon bone of the leading leg. The nose of the fetus follows, usually falling somewhat in line with the cannon bone of the trailing leg. The appearance of both forelimbs ahead of the nose (rather than all three appearing at the same time) verifies that all joints of the forelimbs (particularly the elbows) are extended. The staggered appearance of the leading leg and trailing leg verifies that one of the shoulders is advanced in front of the other one. This is important because the widest part of the fetus is the shoulders. When the nose is making its appearance past the vulva, the shoulders are approximately engaging the narrowest part of the pelvic canal, hence this represents the most difficult and lengthy part of delivery for the mare. If the fetus were to try to pass through the pelvic canal without advancing one shoulder in front of the other it would be akin to a broad-shouldered man trying to walk straight-on through a very narrow gate, as opposed to putting one arm and shoulder through first and letting the other one follow. The mare will typically exhibit strong contractions at this stage for quite a few minutes, making just a little progress each time, before successfully clearing the fetal shoulders through the pelvic canal. The amnion usually breaks during the passage of the shoulders, and the neonate breathes air for the first time, even though the umbilicus is still attached to the placenta and functioning. Clear fluid may be seen to drain from the mouth and nostrils of the neonate; this originates from the
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respiratory and gastrointestinal tract. Once the shoulders are clear, with the entire fetal head visible past the vulva, the trunk and hips of the fetus pass comparatively quickly – this has been described as ‘explosive’. The mare will often rest at this point, with the hindlimbs of the fetus still inside her reproductive tract, for quite a few minutes. Blood is still pulsing through the umbilical vessels and it is thought that the neonate recovers some of this ‘reserve’ blood during this resting period. Stage two usually lasts between 20 and 30 minutes; if stage two lasts longer than 20 minutes, especially without evidence of the fetus, the mare should be evaluated for dystocia. After some time resting, the fetus may either kick out of the mare completely, or the mare will stand up. The physical distance created by this movement will break the umbilical cord at a natural stricture in the cord. The stretch and breakage of the vasculature in this fashion results in a constriction of the vessels and almost no blood loss. The mare should turn and introduce herself to the neonate with licking, nudging, and (if it has not started already) vocalizations. The neonate will often respond in kind with vocalizations. The neonate should be in sternal position within 30 minutes of birth and be standing and suckling on its own within 3 hours. To avoid failure of passive transfer, a foal should ingest 1–1.5 L of good-quality colostrum in the first 6 hours of life. 37, 38 It is a good idea to evaluate a colostral sample from the mare at this stage, especially in cases of suspicion of colostral insufficiency. Using a colostrometer a specific gravity > 1.060 correlates with colostral IgG concentrations > 3000 mg/dL and foals having serum IgG concentrations > 520 mg/dL 24 hours postpartum.39 Using a wine refractometer, readings of > 23% sugar and > 16◦ alcohol correlate with colostral IgG concentrations > 6000 mg/dL and foal serum IgG concentrations > 670 mg/dL at 24 hours postpartum.40 See Chapter 234 on Colostrum Assessment of Quality and Artificial Supplementation for more information. Foals from mares with inadequate colostrum should be supplemented with 200–400 mL high-quality colostrum (e.g., colostral specific gravity > 1.080) within 12 hours of birth.38 See Chapter 234 on Assessment of Passive Transfer for more information.
Stage three of parturition Stage three is the passage of the fetal membranes and usually follows between 15 minutes and 4 hours after completion of stage two. If stage three lasts longer than 6 hours, treatment should be initiated for retained fetal membranes; such treatment includes antibiotics, anti-inflammatory drugs, oxytocin, and uterine lavage (see Chapter 260). When the membranes pass they should be examined to
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(a)
(b)
Figure 233.1 The allantoic side of the chorioallantois. (a) A mare’s chorioallantois arranged in an ‘F’ formation for assessment. The top arm of the ‘F’ is the gravid horn (white arrow) and the lower arm of the ‘F’ is the non-gravid horn (black arrow). The body of the uterus makes up the back of the ‘F’; the extra arm is the umbilicus attached to the amnion (open arrow). Note the prominent blood vessels visible on this side of the membranes. The hippomane is in the upper left corner. (b) The chorionic side of the chorioallantois. The chorion in the mare is diffusely covered with microcotyledons, giving a freshly expelled chorion a red, velvety appearance. The tip of the gravid horn in this example is hyperemic, pathogenesis unknown.
determine their completeness and to detect any gross pathologies. The membranes will often be expelled inside out with the allantoic side of the chorioallantois facing out and the chorionic side inward (Figure 233.1a). Placental evaluation Examining the fetal membranes after every foaling event is a responsible practice that will help the clinician diagnose pathologies such as placentitis and dystocia after the fact, and partial retention of fetal membranes before clinical signs occur in the mare. In situations where foaling occurs unattended, especially outside, it may not be possible to find the fetal membranes. If they are found they may be partially destroyed. In all cases when foaling is observed, the fetal membranes should be saved and evaluated within the next few hours or day. Turning the membranes right side out (with the chorionic side external), you can arrange the membranes on the ground in an ‘F’ shape (Figure 233.1b). Stretching the membranes out thus allows the assessment of the completeness of the fetal membranes. The
‘gravid’ horn (i.e., the one that contained the fetal hindlimbs) will be positioned on the top portion of the ‘F’ while the stem of the ‘F’ comprises the body of the fetal membranes and the lower arm of the ‘F’ is the ‘non-gravid’ horn (i.e., the horn that did not contain any of the fetus). When a portion of the fetal membranes is retained in the mare due to a tear during stage three, it should be detected during examination of the membranes in this fashion. The most common site for a tear to occur is at the tip of the non-gravid horn. This horn will be noticeably thinner than the gravid horn, which may contribute to its tendency to tear. Some have advocated filling the fetal membranes with water to detect tears. This has limited use since often tears occur that do not represent actual retained portions, but simply a traumatic tear in the membranes that occurred as a result of normal parturition, without a portion of the membranes remaining in the uterus. The task of the person evaluating the membranes is to try to discern if holes in the membranes represent retained pieces, or simple tears. A missing tip of a horn is easier to assess than a tear in the body (Figure 233.2).
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determine if the vessels seem to meet, or if a portion is entirely missing. When in doubt, it is best to err on the side of caution, assume that a piece may be retained, and treat the mare accordingly with antibiotics, anti-inflammatory agents, oxytocin, and uterine lavage until a retained portion of the fetal membranes can be recovered or ruled out. In addition to evaluating the intactness of the fetal membranes, the chorion should be evaluated for pathologies indicative of intrauterine disease. Placentitis will often leave gross lesions on the chorion, most commonly a purulent exudate and avillous areas. This latter lesion should be differentiated from normal avillous areas of the chorion. The mare has diffuse placentation, resulting in a chorion that is covered by microcotyledonary villi everywhere the chorion had contact with the endometrium. Areas lacking villi naturally will be found: Figure 233.2 Chorioallantois of a mare with the chorionic surface facing out. Note the missing tip of the non-gravid horn (arrow).
Turning the membranes back to where the allantoic portion is external may be advantageous in situations where there is doubt about whether a tear represents a retained piece or not. The allantoic portion of the membranes contains the vasculature of the membranes and one can often put pieces back together to
(a)
1. at the tip of each horn at the utero-tubular junction where the oviductal papilla is found; 2. at the base of the horns opposite the attachment of the umbilicus; 3. at the end of the body where the cervix lay adjacent to the placenta (resulting in the ‘cervical star’); and 4. anywhere along the membranes where the chorion folded on itself and formed a crease where it touched itself instead of the endometrium.
(b)
Figure 233.3 Chorioallantoides of mares with the chorionic surface facing out. (a) The avillous lesion near the cervical area of the body of the chorion (arrow) indicates an ascending placentitis and consequent detachment of the fetal membranes from the endometrium with loss of microvilli. (b) The discrete avillous lesion on the body of the chorion, without contact with the cervical area, indicates a separation of the placenta not due to an ascending pathogen; this type of lesion is typical of a nocardioform placentitis.
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Figure 233.5 Allantoic surface of fetal membranes showing raised, wartlike lesions, allantoic adenomatous hyperplasia (arrows), which were located opposite chorionic lesions indicative of placentitis.
Figure 233.4 Chorioallantois of a mare with the chorionic surface facing out. Note the intact cervical star (arrow) indicating a ‘red bag’ dystocia and likely dummy foal. Photo courtesy of Malgorzata Pozor.
Other than these common areas, the rest of the chorion should be covered uniformly in a red, velvety layer of villi. An avillous lesion near the cervical star indicates an ascending, transcervical placentitis,
(a)
whereas an avillous lesion near the base of the horns may be indicative of a nocardioform placentitis (Figure 233.3). Nocardia placentitis more typically is associated with a thick, often brown mucoid exudate at the site of placental separation at the base of the uterine horn or body. Diffuse avillous lesions indicate a hematogenous pathogenesis. Definitive diagnosis of placentitis and treatment are discussed in Chapter 242. Other pathologies will occasionally be noted. An intact cervical star indicates premature separation of the chorioallantois (i.e., ‘red bag’ delivery; Figure 233.4), and may result in hypoxia and a ‘dummy foal’. Twins will result in two separate sets of fetal membranes, each with only one horn, or portion thereof, and a large avillous area at the base where they lay adjacent to each other inside the uterus.
(b)
Figure 233.6 Amniotic portions of fetal membranes from two foals. Note the translucent nature of the normal amnion (a) compared to the thicker, opaque amnion (b) that was a result of inflammation in utero.
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Parturition Occasionally on the allantois, opposite an avillous chorionic lesion, raised wartlike lesions called chorionic adenomatous hyperplasia may be detected (Figure 233.5). These lesions are associated with infectious placentitis, either bacterial or fungal,41, 42 but the specific pathogenesis or significance of the adenomatous hyperplasia is unknown. A similar gross lesion in a mare was diagnosed as a placental teratoma.43 Evaluation of the amnion should involve assessment of its color and thickness. A normal amnion should be translucent, allowing easy visualization of your hand through the tissue (Figure 233.6a). A thickened amnion will be opaque, and the thickening may indicate an inflammatory response in utero (Figure 233.6b). The normal color of the amnion is gray to whitish. A brownish or greenish stain on the amnion usually indicates that the meconium was expelled in utero, an indication of fetal stress, as may be the case in a dystocia.
References 1. Holtan DW, Houghton E, Silver M, Fowden AL, Ousey J, Rossdale PD. Plasma progestagens in the mare, fetus, and newborn foal. J Reprod Fert Suppl 1991;44:517–28. 2. Seamans KW, Harms PG, Atkins DT, Fleeger JL. Serum levels of progesterone, 5α-dihydro-progesterone and hydroxy-5α-pregnones in the pre-partum and post-partum equine. Steroids 1979;33:55–63. 3. Thorburn GD. A speculative review of parturition in the mare. Equine Vet J 1993;14:41–9. 4. Lovell JD, Stabenfeldt GH, Hughes JP, Evans JW. Endocrine patterns of the mare at term. J Reprod Fertil Suppl 1975;23:449–56. 5. Palm F, Neuhauser S, Ambuehl F, Moestl E, Aurich C. Effect of altrenogest-treatment in late pregnant-mares on parturition, neonatal adaptation and adrenocortical function in new-born foals. Reprod Domest Anim 2008;43(Suppl. 3): 109. 6. Pashen RL. Maternal and foetal endocrinology during late pregnancy and parturition in the mare. Equine Vet J 1984;16:233–8. 7. Silver M. Prenatal maturation, the timing of birth and how it may be regulated in domestic animals. Exp Physiol 1990;75:285–307. 8. Egarter CH, Husslein P. Biochemistry of myometrial contractility. In: Elder MG (ed.) Bailli`ere’s Clinical Obstetrics and Gynaecology – Prostaglandins. London: Bailli`ere Tindall, 1992; pp. 755–69. 9. Currie WB, Wong MSF, Cox RF, Thorburn GD. Spontaneous or dexamethasone induced parturition in the sheep and goat; changes in plasma concentrations of maternal prostaglandin F and foetal oestrogen sulphate. Mem Soc Endocr 1973;20:95–118. 10. Flint AP, Anderson BM, Patten PT, Turnbull AC. Proceedings: Uterine venous prostaglandin F and oestrogen levels during ovine parturition: acute effects of cervical stimulation. J Endocrinol 1974;61(1):XXXV–XXXVI. 11. Haluska GJ, Currie WB. Variation in plasma concentrations of oestadiol-17β and their relationship to those of progesterone, 13,14-dihydro-15-keto-prostaglandin F-2α and oxy-
12.
13.
14.
15. 16. 17.
18.
19.
20.
21.
22.
23. 24.
25. 26.
27.
28.
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tocin across pregnancy and at parturition in pony mares. J Reprod Fertil 1988;84:635–46. Garfield RE, Karmen MS, Daniel EE. Gap junction formation in myometrium: control by estrogen, progesterone and prostaglandins. Am J Physiol 1980;237:C81–C89. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy, and parturition in the mare. J Reprod Fertil Suppl 1979:27;499–509. Nathanielsz PW, Giussani DA, Mecenas CA, Wu W, Winter JA, Garcia-Villar R, Baguma-Nibasheka M, Honnebier BOM, McDonald TJ. Regulation of the switch from myometrial contractures to contractions in late pregnancy: studies in the pregnant sheep and monkey. Reprod Fertil Dev 1995;7:293–300. Stewart DR, Stabenfeldt GH. Relaxin activity in the pregnant mare. Biol Reprod 1981;25:281–9. Stewart DR, Stabenfeldt GH, Hughes JP. Relaxin activity in foaling mares. J Reprod Fertil Suppl 1982;32:603–9. Hwang JJ, Sherwood OD. Monoclonal antibodies specific for rat relaxin III. Passive immunization with monoclonal antibodies throughout the second half of pregnancy reduces cervical growth and extensibility in intact rats. Endocrinology 1988;123:2486–90. Hwang JJ, Shanks RD, Sherwood OD. Monoclonal antibodies specific for rat relaxin IV. Passive immunization with monoclonal antibodies during the antepartum period reduces cervical growth and extensibility, disrupts birth, and reduces pup survival in intact rats. Endocrinology 1989;125:260–6. Silver M, Ousey JC, Dudan FE, Fowden AL, Knox, J, Cash RSG, Rossdale PD. Studies on equine prematurity 2: post natal adrenocortical activity in relation to plasma adrenocorticotrophic hormone and catecholamine levels in term and premature foals. Equine Vet J 1984;16:278– 86. Moog F. Enzyme development and functional differentiation in the fetus. In: Waisman HA, Kerr GR (eds) Foetal Growth and Development. New York: McGraw Hill, 1970; pp. 29–48. Liggins GC, Kitterman JA. Development of the fetal lung. In: Whelan J (ed.) The Fetus and Independent Life. CIBA Foundation Symposium No. 86. London: Pitman, 1981; pp. 308–30. Challis JRG, Harrison FA, Heap RB, Horton EW, Poyser NL. A possible role of oestrogens in the stimulation of PGF2α output at the time of parturition in a sheep. J Reprod Fertil 1972;30:485–8. Thorburn GD. The placenta, PGE2, and parturition. Early Hum Dev 1992;29:63–73. Barnes RJ, Comeline RS, Jeffcott LB, Mitchell MD, Rossdale PD, Silver M. Foetal and maternal plasma concentrations of 13,14 dihydro-15-oxo-prostaglandin F in the mare during late pregnancy and at parturition. J Endocrinol 1978;78:201–15. Rossdale PD. Clinical view of disturbances in equine foetal maturation. Equine Vet J Suppl 1993;14:3–7. Bain AM, Howey WP. Observations on the time of foaling in Thoroughbred mares in Australia. J Reprod Fertil Suppl 1975;23:545. Morris DD, Meirs DA, Merryman GS. Passive transfer failure in horses: incidence and causative factors on a breeding farm. Am J Vet Res 1985;46:2294–9. Ousey JC, Delclaux M, Rossdale PD. Evaluation of three strip tests for measuring electrolytes in mares’ pre-partum
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mammary secretions and for prediction of parturition. Equine Vet J 1989;21:196–200. Ley WB. Management of the foaling mare: prediction readiness for birth and inducing foaling. Vet Med 1994;89:570–7. Ley WB, Bowen JM, Purswell BJ, Irby M, Greive-Crandell K. The sensitivity, specificity and predictive value of measuring calcium carbonate in mares’ prepartum mammary secretions. Theriogenology 1993;40:189–98. Rook JS, Braselton WE, Nechreiner RF, Lloyd JW, Shea ME, Shelle JE, Hitzler PR. Multi-element assay of mammary secretions and sera from periparturient mares by inductively coupled argon plasma emission spectroscopy. Am J Vet Res 1997;58:376–8. Ousey JC, Dudan F, Rossdale PD. Preliminary studies of mammary secretions in the mare to assess foetal readiness for birth. Equine Vet J 1984;16:259–63. Bennett DG. Confronto tra parto indotto artificialmente e parto spontaneo nella cavella. Ippologia 1990;3:11–18. Carluccio A, De Amicis I, Panzani S, Tosi U, Faustini M, Veronesi MC. Electrolytes changes in mammary secretions before foaling in jennies. Reprod Domest Anim 2008;43:162–5. Jeffcott LB, Rossdale PD. A radiographic study of the fetus in late pregnancy and during foaling. J Reprod Fertil Suppl 1979;27:563–9. Arthur GH. The development of the conceptus. In: Veterinary Reproduction and Obstetrics, 4th edn. London: Bailli`ere Tindall, 1975; p. 4142.
37. Massey R, LeBlanc M, Klapstein E. Colostrum feeding of foals and colostrum banking. In: Proceedings of the 37th Annual Conference of the American Association of Equine Practitioners, 1991, pp. 1–8. 38. LeBlanc M, Tran T, Baldwin J, Pritchard E. Factors that influence passive transfer of immunoglobulins in foals. J Am Vet Med Assoc 1992;200:179–83. 39. LeBlanc M, McLaurin B, Boswell R. Relationships among serum immunoglobulin concentration in foals, colostral specific gravity, and colostral immunoglobulin concentration. J Am Vet Med Assoc 1986;189:57–60. 40. Chavatte P, Clement F, Cash R, Gronget J. Field determination of colostrum quality by using a novel, practical method. In: Proceedings of the 44th Annual Convention of the Amercian Association of Equine Practitioners, 1998, pp. 206–9. 41. Hong CB, Donahue JM, Giles RC Jr, Petrites-Murphy MB, Poonacha KB, Tramontin RR, Tuttle PA, Swerczek TW. Adenomatous hyperplasia of equine allantoic epithelium. Vet Pathol 1993;30:171–5. 42. Shivaprasad HL, Sundberg JP, McEntee K, Gordon L, Johnstone AC, Lombardo de Barros CS, Hoffman RL. Cystic adenomatous hyperplasia of the equine allantois: a report of eight cases. J Vet Diagn Invest 1994;6:107–10. 43. Gurfield N, Benirschke K. Equine placental teratoma. Vet Pathol 2003;40:586–8.
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Chapter 234
Lactation Patrick M. McCue and Sofie Sitters
Appropriate and timely development of the mammary gland, colostrum production, and subsequent lactation play key roles in the survival and health of a neonatal foal. The mammary gland is responsible for providing the neonate with immunoglobulins critical to survival and supplies nutrition to the foal during the first few months of life.
Anatomy of the mammary gland The mammary gland of the mare consists of paired mammae, separated by a fascial septum, each with a glandular body and teat.1 The glandular portion of each half is divided by fibroelastic capsules into two, or occasionally three, lobes. The mammary gland is located high in the inguinal region of the mare. Milk is secreted by specialized epithelial cells into the lumen of alveoli and small ducts. Myoepithelial cells surround the alveoli and are important in the ejection of milk following a sucking stimulus.2 Milk passes through progressively larger lactiferous ducts and then into the teat cistern before exiting at the teat orifice. Each of the mare’s teats serves a cranial and caudal lobe of mammary tissue and has two corresponding teat orifices (Figure 234.1). The anatomy of the equine mammary gland and relationship between teat orifices and lobes is important for understanding diseases (i.e., mastitis) and treatments.
Physiology of lactation Development of the mammary gland and onset of lactation are complex processes under control of the hypothalamic–pituitary axis, ovaries, and placenta. Mammary growth and galactopoiesis are influenced by ovarian and adrenal steroids, prolactin, oxytocin, growth hormone, insulin, thyroid hormones, and other factors. Estrogen induces development of the mammary ducts. Progesterone stimulates lobuloalveolar growth, while at the same time inhibiting lac-
togenesis.2 The trigger that initiates lactogenesis is thought to be the decline in progesterone and increase in prolactin that occurs late in gestation.2 Plasma prolactin concentration in the mare increases during the last week of gestation and remains elevated during early lactation before declining to baseline by 1–2 months postpartum.3 Maximum concentrations of prolactin are reached 2–3 days after foaling.4 Production of prolactin by lactotrophs in the anterior pituitary is regulated by hypothalamic secretion of dopamine. The presence of a placental lactogen as exists in primates, rodents, and ruminants has not been demonstrated in the mare.5 Oxytocin is synthesized in the supraoptic and paraventricular nuclei of the hypothalamus. Nursing by the foal stimulates the release of oxytocin from the neurohypophysis.6 Oxytocin release induces contraction of myoepithelial cells surrounding the alveoli, causing expulsion of milk from the alveoli and small ducts into the larger lactiferous ducts and teat cisterns (i.e., ‘milk let-down’). Milk let-down or ejection in the mare can also occur in the absence of tactile stimulation or a measurable oxytocin release.7
Components of colostrum and milk The epitheliochorial placenta of the mare prevents transfer of maternal antibodies to the fetus in utero. The foal depends entirely on passive transfer of antibodies in colostrum for protection against infection during the early neonatal period. Antibodies (primarily immunoglobulin IgC with some IgM class antibodies) from the mare’s blood are selectively concentrated in the colostrum during the last 2 weeks of pregnancy (Figure 234.2).8–11 The average IgG concentration of equine colostrum at foaling is approximately 7000 mg/dL (70 g/L), with a normal range of approximately 3000–12 000 mg/dL (30–120 g/L) (Table 234.1).11–24 Immunization of mares 4–6 weeks prior to foaling will increase
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Pregnancy, Parturition and the Puerperal Period Table 234.1 Antibody (IgG) concentration in colostrum of mares at the time of parturition
Figure 234.1 Mammary gland of the mare showing two teat orifices on the right teat.
the concentration of IgGs against pathogens in colostrum.25 Colostrum is also an important source of nutrients, vitamins, complement, and lactoferrin. In addition, colostrum promotes cellular immunity and may have a variety of other physiological activities such as the activation of granulocytes and the provision of a local protective effect in the gastrointestinal tract.8,10–12 In multiparous mares, the average rate of colostrum secretion is approximately 300 mL/ h and the total volume of colostrum produced is 2–5 liters.11, 12, 16, 18 Premature lactation can significantly decrease the amount of immunoglobulins available to the neonate and may result in failure of passive transfer. IgG concentration in the colostrum of a dam is highly correlated with the serum IgG of her neonatal foal after suckling.13–16,19, 28 After a neonatal foal has ingested colostrum, specialized enterocytes in the small intestine nonselectively absorb the colostral immunoglobulins by pinocytosis. The enterocytes engulf droplets of colostrum from the intestinal lumen, transfer the droplets in small vacuoles across the cell, and then 35
12 000
30
10 000
25
8000
20
6000
15
4000
10
IgG (mg/dL)
Sample size
Various breeds Various breeds Not specified Crossbred Thoroughbred Thoroughbred Thoroughbred Arabian Arabian Arabian Quarter Horse Quarter Horse
5450 11 324 16 583 8867 5036 2354 4608 9691 10 847 6394 5843 1080
139 18 6 10 39 32 22 14 12 25 21 1
3932 5128 8912 4600 6438
15 136 36 9 11
Standardbred Standardbred Standardbred Shetland pony Heavy draft mares
Reference Erhard et al.14 Massey et al.18 Lavoie et al.16 Curadi et al.13 LeBlanc et al.17 McGuire et al.26 Pearson et al.20 Pearson et al.20 Perryman24 LeBlanc et al.17 LeBlanc et al.17 McGuire and Crawford27 LeBlanc et al.17 Morris et al.19 Kohn et al.15 Rouse and Ingram21 Townsend et al.23
discharge the contents into lymphatic vessels. The antibodies are subsequently transported via the lymphatic system to the bloodstream.8,10–12 The absorption of colostral antibodies is greatest during the first 6–8 hours after birth, and has essentially stopped by 24–36 hours of age because the specialized enterocytes have been replaced by epithelial cells not capable of pinocytosis.8,10–12,20, 29, 30 To acquire sufficient antibodies to offer reasonable protection against infectious disease, the average foal needs to ingest approximately 2–4 liters of good quality colostrum within the first few hours of life.8,10–12,16–18,24 Concentrations of IgG in the mammary secretions decline rapidly during the first 24 hours after foaling as the foal suckles (Figure 234.3).15, 16, 20, 21, 31 35
10 000
30 8000 25 6000
20
4000
15
Brix score (%)
14 000
Breed
IgG (mg/dL)
IgG (mg/dL)
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5
0
0 1
3
5
7
9
11
13
15
17
19
21
23
Time (days), day 23=day of foaling BRIX
RID
Figure 234.2 Antibody (immunoglobulin G; IgG) levels in the mammary secretion of a mare in the days prior to foaling measured by single radial immunodiffusion (RID) and estimated using a Brix (sugar) refractometer.
5 0
0 0
2
4
6
8
10
12
14
16
18
20
22
Time (hours) RID
BRIX
Figure 234.3 Antibody (IgG) levels in the mammary secretion of a mare in the hours after foaling and during suckling of the newborn foal as measured by single radial immunodiffusion (RID) and estimated using a Brix (sugar) refractometer.
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Table 234.2 Relationship between immunoglobulin (IgG) concentrations as measured by single radial immunodiffusion and estimation of colostrum quality using Brix refractometry and specific gravity
Colostrum quality Poor Fair Good Very good
IgG concentration (g/L)
Brix (%)
Specific gravity
0–28 28–50 50–80 > 80
< 15 15–20 20–30 > 30
< 1.06 1.06–1.07 1.08–1.09 ≥ 1.10
This has important ramifications for collection of colostrum for banking. Colostrum should be harvested for frozen storage either immediately after the mare foals or within the first 2 hours. The quality of colostrum collected subsequently will be lower (i.e., have a lower IgG concentration). The concentration of colostral immunoglobulins and therefore relative quality can be quantified using radial immunodiffusion or estimated using a Brix (sugar) refractometer or by measurement of specific gravity using a commercial colostrometer (Table 234.2).28, 32 Other less widely used techniques include gluteraldehyde precipitation,8, 11, 12 latex agglutination,11, 12, 33 and an enzyme-linked immunosorbent assay (ELISA).14 Direct quantification of IgG levels in colostrum can be performed using the single radial immunodiffusion (RID) assay.11, 12, 34, 35 Unfortunately, the RID cannot be performed easily in a field situation and it takes approximately 24 hours to obtain a result. The colostrometer measures the density or specific gravity of colostrum (Figure 234.4).11, 12, 18, 28, 33 Colostrum with high IgG levels has a higher density and, therefore, specific gravity. However, determination of the specific gravity of equine colostrum with a colostrometer depends on accurate measurement of an exact volume (15 mL) of colostrum; even small errors in volume measurement can lead to significant inaccuracy in specific gravity determination.36 Refractometry measures the concentration of dissolved solids in a solution. In the case of a Brix refractometer, a small amount of colostrum is placed on the prism and the light plate is closed so that colostrum is spread evenly across the prism (Figure 234.5). The refractometer is then pointed at a light source and the deviation or refraction of light is evaluated as a percentage score (Figure 234.6). Colostrum with a low concentration of dissolved solids (i.e., low IgG level) will scatter less light and therefore have a lower percentage score. Colostrum with high amounts of dissolved solids (i.e., high IgG levels) will cause more light scatter and thereby achieve a higher percent-
Figure 234.4 Evaluation of colostrum specific gravity using a colostrometer. The glass colostrometer containing 15 mL of colostrum is allowed to float in a graduated cylinder filled with distilled water. The specific gravity of the colostrum is evaluated using a scale on the colostrometer
age score. Brix refractometer evaluation of equine colostrum quality has been shown to be both repeatable (r = 0.98) and highly correlated with foal plasma IgG levels (r = 0.85), as measured by the radial immunodiffusion assay (Figure 234.7).32, 36
Figure 234.5 Placing a drop of colostrum onto the prism of a Brix refractometer.
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100 Colostrum IgG (g/L)
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80 60 40 20 0 0
10
30
20
40
Refractometer % reading Figure 234.7 Correlation between Brix refractometer score and immunoglobulin G (IgG) concentration of colostrum measured by single radial immunodiffusion (SRID).32
are available to estimate calcium concentrations in prepartum mammary secretions and therefore predict the interval to spontaneous parturition. Lactation in the mare peaks at 1–2 months postpartum. Average daily milk production in Quarter Horse mares ranges from 11.8 kg on day 30 of lactation to 9.8 kg on day 150 of lactation.38 The average daily milk production in a Thoroughbred mare is approximately 10–12 kg/day.48 Daily milk production in the mare is equivalent to 2.1–3.4% of bodyweight.37
Figure 234.6 View through the refractometer showing Brix % and the quality score. Note that, in this instance, the interface of the dark field (above) is approximately 21.5%, which would be on the low end of the scale for ‘good quality’ colostrum.
Concentrations of total solids, protein, and amount of gross energy decrease abruptly during the first 12 hours following foaling and continue to decline slowly throughout lactation (Table 234.3).37, 38 Conversely, lactose concentrations tend to rise slowly throughout lactation.38 The mare has milk that is low in total solids, fat, and protein but high in lactose, which is somewhat rare in mammals.39, 40 Mineral composition of mare’s milk also varies with stage of lactation. Concentrations of total solids, ash, calcium, phosphorus, magnesium, copper, and zinc all decrease as lactation progresses (Table 234.4).42–44 Composition of milk from nondomestic equids is similar to that of the domestic horse.40 Mineral and electrolyte concentrations in prepartum mammary secretions can be used as an indicator of impending parturition and to evaluate fetal readiness for birth.45–47 Commercial kits
Milk replacers It may become necessary to provide a foal with an alternative source of milk because of death or injury to the dam, rejection of the foal by the mare, or inadequate lactation. Orphaned neonates must first be provided an adequate quantity of colostrum with high immunoglobulin content. Foals should receive 250 mL of colostrum (Brix > 25%) every hour for the first 6 hours of life.49 This may be accomplished via a bottle or passage of a small nasogastric tube. Table 234.3 Composition of equine milk (adapted from National Research Council: Nutrient Requirements of Horses41 )
Weeks of lactation 1–4 5–8 9–21
Energy (kcal/100 g)
Protein (%)
Fat (%)
Lactose (%)
Total solids (%)
58 53 50
2.7 2.2 1.8
1.8 1.7 1.4
6.2 6.4 6.5
10.7 10.5 10.0
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Table 234.4 Mineral constituents of equine milk (adapted from National Research Council41 ) Concentration of minerals (mg/L) [mmol/L] in equine milk Weeks of Lactation 1–4 5–8 9–21
Ca
P
Mg
K
Na
1200 [30] 1000 [25] 800 [20]
725 [23.4] 600 [19.4] 500 [16.1]
90 [3.9] 60 [2.6] 45 [2.0]
700 [18.0] 500 [12.8] 400 [10.3]
225 [9.8] 190 [8.3] 150 [6.5]
Once the foal has received colostrum, the optimal approach would be to foster the foal onto a nurse mare if one is available.50, 51 Alternatively, orphaned foals have been fostered successfully onto cows or goats.51 If the foal cannot be fostered onto a nurse mare, it can be hand reared on a milk substitute such as fortified cow’s milk or a commercial milk replacer (Table 234.5). Cow’s milk (whole) has almost twice the fat content and only two-thirds the sugar content of mare’s milk. Low-fat cow’s milk (2% milk fat) can be used as a substitute for mare’s milk if 20 g dextrose is added per liter. Fortified cow’s milk should be fed to a total volume of 10% of the foal’s bodyweight at 1 day of age.50 The amount fed should be gradually increased to 25% of bodyweight per day from day 10 until weaning. Many mare milk replacer powders are commercially available (Table 234.6). Milk replacers should contain about 15% fat and 22% crude protein and have a fiber content of less than 0.5%. Manufacturers of milk replacers often recommend feeding a solution more concentrated than normal mare’s milk. However, Naylor and Bell49 recommend that foals be fed a more dilute (12.5%) solution when using commercial milk replacers to avoid potential problems with constipation and dehydration. Therefore, a greater volume of diluted milk replacer solution should be fed to provide the recommended total dry matter intake.
Table 234.5 Comparison of the composition of mare’s milk with various substitutes (adapted from Naylor and Bell50 ) Mare’s milk
Milk replacera
Cow’s milk
Goat’s milk
Fat, % of dry matter (DM)
15
14.4
29
34
Protein, % of DM Sugar, % of DM Fiber, % of DM Total solids
22.8 58.8 0 11.6
20.2 52.6 0.2 20
27 38 0 12.3
25 31 0 13.2
Item
a
Foal-Lac, Borden, Inc., Hampshire, IL.
Foals can be artificially reared using a bottle, or they can be taught to drink from a bucket. Foals nurse from their dams more frequently during the first week of life than at any other age.52 Consequently, recommended frequency of feeding orphan foals gradually decreases from every 1–2 hours during the first few days after birth to every 4–6 hours after 2 weeks of age.49 Overall weight gains in foals reared on an artificial diet have been reported to be similar to gains in foals allowed to nurse from their dams.53, 54 In general, however, it is recommended that orphan foals be provided a nurse mare or an equine companion to facilitate development of normal equine social skills and to avoid inappropriate future horse–human behavioral problems.
Dietary requirements for lactation Foaling mares should be fed good-quality roughage at a rate of at least 1–2% of bodyweight of dry matter per day. Supplementation with concentrates should be based on quality of the roughage, environmental conditions, nutritional status of the mare, and stage of lactation. Nutrient requirements of mares during early lactation are substantially greater than those of the pregnant or open mare. Energy requirements during early and late lactation are 1.7 and 1.5 times, respectively, greater than necessary for maintenance in an open mare.41 The National Research Council indicated that the percentages of dietary protein, calcium, and phosphorus Table 234.6 Commercial milk replacers for artificial rearing of foals Name
Manufacturer
Foal-Lac FoalGro Grow-N-Glow Mare’s Match Foals First Mare Replacer Powder Mares Milk Plus
Borden, Inc., Hampshire, IL Grober Nutrition, Cambridge, Ontario, Canada Merrick’s Inc., Middleton, WI Land O’Lakes Purina Feed, LLC, Gray Summit, MO Progressive Nutrition, LLC, Harlan, IA Start To Finish Products, Carpentersville, IL Buckeye Nutrition, Dalton, OH
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during early lactation are approximately 1.7, 2.2, and 2.0 times that required for maintenance. A 450-kg mare in early lactation, for example, would require a diet that contained 25.6 Mcal digestible energy, 12% protein, 0.47% calcium, and 0.30% phosphorus. Those requirements could be provided by feeding a ration of 6 kg of alfalfa hay and 4.1 kg of grain per day. All horses should have free access to a clean water supply and a salt–mineral supplement. Water requirements of a lactating mare may be 50–70% above that required for a non-pregnant mare.41
related to the degree of hypocalcemia. Without treatment, tetanic convulsions begin within 24 hours and death occurs within 48 hours. Treatment with a 20% calcium gluconate solution (250–500 mL per 450 kg, diluted 1:4 in saline or dextrose) results in rapid recovery. Calcium solutions should be administered slowly intravenously while monitoring cardiovascular function. Retreatment may be required in some cases.
Agalactia Weaning Foals are commonly weaned at 4–6 months of age. Weaning of individual foals may be accomplished by removing the dam from the box stall of the foal to another area of the farm, which is out of visual and auditory contact with the foal. After a few days in the stall, the foal may be placed into a small paddock with other weanlings. Alternatively, groups of foals housed with their dams in large paddocks may be weaned together by removing one or two mares every other day over a period of 1–2 weeks. An old, gentle mare may be left with the weaned foals to have a quieting effect.51 Alveolar cells continue to secrete milk as long as milk is removed regularly from the mammary gland. After nursing ceases, mammary glands may become mildly to moderately distended with milk. Accumulation of milk initiates involutional changes in the mammary gland, resulting in decreased milk synthesis and glandular regression. Glandular distension will gradually subside without treatment. It is not recommended to ‘milk-out’ a mare with a distended mammary gland after weaning as this may lead to a delay in glandular involution. Mastitis occasionally occurs in the mare after weaning of the foal. Therefore, the mammary gland should be routinely monitored for development of mastitis for several weeks after weaning.
Eclampsia Lactation tetany, or eclampsia, is an uncommon disease that may occur in lactating mares in the early postpartum period, stressed lactating mares, or after weaning.55, 56 Mares producing large quantities of milk that are subjected to hard physical work or prolonged transport are most commonly affected. The syndrome is characterized by low serum calcium levels (4–5 mg/dL) and occasionally low magnesium levels. Severity of clinical signs – which may include a stiff gait, tachypnea, synchronous diaphragmatic flutter, profuse sweating, and muscle tremors – is
Agalactia is the failure of the mammary gland to secrete colostrum or milk following parturition. Clinical consequences of agalactia or hypogalactia in the periparturient mare are failure of passive transfer of immunoglobulins and inadequate milk supply to the neonate. Prepartum failure of mammary development may occur spontaneously in some (usually primiparous) mares, as a consequence of ingestion of an ergot alkaloid, administration of a dopamine agonist (e.g., pergolide or bromocryptine), inadequate nutrition, or other factors. In the USA, agalactia is most commonly caused by ingestion of tall fescue grass (Festuca arundinacea) infected with the ergot alkaloid-producing fungus Neotyphodium coenophialum (formerly Acremonium coenophialum) leading to a condition commonly referred to as ‘fescue toxicosis’.57–60 In Brazil, ergotalkaloid-induced agalactia has been reported to follow ingestion of black oats (Avena strigosa) infected with the fungus Claviceps purpurea.61, 62 A recent study in Argentina suggested that ingestion of rye grass infested with the endophyte Neotyphodium lolii may cause clinical signs similar to ‘fescue toxicosis’ in pregnant mares.63 Agalactic mares often have serum prolactin concentrations that are lower than concentrations in normal lactating mares.59, 64 Ergot alkaloids depress prolactin secretion by their action as dopamine DA2 receptor agonists and serotonin antagonists.65 Clinical signs of ‘fescue toxicosis’ can be reproduced by administration of bromocryptine, a semisynthetic ergopeptine alkaloid.60, 66 Pregnant mares maintained on pastures contaminated with a fungus producing an ergot alkaloid may exhibit a lack of udder development and agalactia, thickened fetal membranes that may be retained following parturition, prolonged gestation, abortion, uterine rupture, dystocia, rebreeding problems, and delayed ovarian activity.59,61–63 Administration of thyrotropin-releasing hormone (TRH) has been shown to stimulate prolactin secretion in the mare.64, 67, 68 Other agents that cause an increase in prolactin by blocking the inhibitory action of dopamine include reserpine, phenothiazine
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Table 234.7 Medications used to stimulate lactogenesis in agalactic mares Drug
Dose
Route
Reference
Thyrotropin-releasing hormone (TRH) Reserpine Perphenazine Domperidone Metoclopramide Sulpiride
2.0 mg 0.5–2.0 mg 0.3–0.5 mg/kg BW 1.1 mg/kg BW 25 mg 50 mg/100 kg BW
s.c., q.12 h i.m., q.48 h p.o., q.12 h p.o., q.24 h i.m., q.12 h i.m., q.12 h
Caudle et al.67 Cowles57 Green and Loch57 Redmond et al.71 Singh and Chanel78 Chavatte-Palmer et al.74
BW, bodyweight; i.m., intramuscular; p.o., per os; s.c., subcutaneous.
tranquilizers, butyrophenones, metoclopramide, domperidone, and sulpiride.57,68–71 Treatment of mares with agalactia consists of removal of any alkaloid-containing feed source and stimulation of pituitary prolactin synthesis. Pregnant mares being administered the dopamine agonist pergolide mesylate for treatment of equine Cushing’s disease (ECD) should have the medication discontinued 2 weeks prior to the expected foaling date to allow for proper mammary development.72 Treatment of ‘fescue toxicosis’, stimulation of endogenous prolactin levels, and/or induction of galactopoiesis in agalactic mares has been reported after administration of thyrotropin-releasing hormone,67 reserpine,57 domperidone,71, 73 sulpiride,74, 75 metoclopramide,63, 68 perphenazine,58 fluphenazine decanoate,76, 77 or the serotonin 5HT1A receptor agonists buspirone and dichlorophenyl piperazine63 (Table 234.7). In one report administration of both metoclopramide and domperidone to ergot-infected mares induced milk production, but metoclopramide improved ovarian activity earlier.63 In a comparative trial, domperidone was more effective at resolving both pre- and postpartum agalactia, and prolonged gestation than reserpine.60 Reserpine, administered orally at a dose of 0.01 mg/kg q.24 h, resolved postpartum agalactia, but was also associated with moderate sedation and diarrhea. In contrast, domperidone does not cross the blood–brain barrier. Administration of domperidone should begin 30 days prior to the expected due date if mares are to remain on infested pastures. Domperidone therapy for mares removed from infested pastures, but not exhibiting proper udder development, should begin 10 days prior to the expected foaling date. Administration of domperidone to late-term pregnant mares is associated with a premature rise in milk calcium levels which may interfere with the use of milk calcium levels to predict foaling (P.M. McCue, personal observation). Management recommendations for the prevention of fescue-associated reproductive problems include reseeding fescue pastures with legumes, removal of
pregnant mares from fescue pastures 1–3 months before foaling, or supplementation with a non-fescue hay, such as alfalfa.57, 59 Causes of lactation failure in the mare other than ergot alkaloid ingestion are not well documented. Factors that may lead to hypogalactia include inadequate nutrition, selenium deficiency, and stress.57
Induction of lactation Stimulation of milk production in non-pregnant mares has been accomplished by a combination of exogenous hormones and milking. The primary purpose of induction of lactation in a non-pregnant mare is creation of a nurse mare to provide nutrition and an equine companion for an orphan foal. In most cases, mares do not produce a significant amount of IgG in an induced lactation.74, 75 The following protocol (Box 234.1) has been adapted from several studies.74, 75, 79, 80 A recent report indicated that pretreatment of mares with 11 mg of estradiol benzoate prior to the onset of dopamine antagonist therapy and every other day during dopamine antagonist therapy significantly increased the prolactin response.81 Consequently, it may be beneficial to continue estradiol therapy during induction of lactation for more than a single day as originally described.
Galactorrhea Premature udder development Pregnant mares may exhibit mammary development and begin lactation weeks or months before the expected foaling date. The causes of premature udder development and lactation in pregnant mares include impending abortion, in utero death of a fetus or one of twin fetuses (Figure 234.8), placental separation, and placentitis.31, 82, 83 Occasionally, a pregnant mare will, for unknown reasons, show mammary enlargement during mid- to late gestation, which subsequently spontaneously regresses. Mares with premature udder development should be examined for
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Box 234.1 Procedure for induction of lactation in a non-pregnant mare
r r r r
r r r
Select a calm, gentle, preferably cycling mare that has given birth and lactated previously. Administer estradiol benzoate (50 mg/500-kg mare), intramuscularly once. One may consider continuing estradiol therapy every other day during the dopamine antagonist treatment period as noted above. Begin daily administration of altrenogest (22 mg/500-kg mare), orally once per day and a dopamine antagonist; choices include: – sulpiride (1 mg/kg, intramuscularly, twice daily) – domperidone (1.1 mg/kg, orally, twice daily, q.12 h). Start milking six times daily on the 4th to 7th day of treatment, when the mammary gland has increased in size: – hand milking initially until milk production increases – machine milking (goat-type milking machine) – 120 aspirations/min, vacuum 22–37 cmHg, set at an alternation of suction relaxation ratio of 50:50. Oxytocin (1–5 IU, i.m. or i.v.) may be administered 1–2 minutes prior to milking to promote milk let-down. Introduce foal after 3–4 days of milking, when production has reached 3–5 L/day day in a 500-kg mare. Discontinue altrenogest on day 7 of treatment and discontinue dopamine antagonist treatment several days after adoption has been completed.
evidence of placentitis by transrectal and possibly transabdominal ultrasonography and measurement of the combined thickness of the uterus and placenta (CTUP) as well as evaluated for separation of the placenta from the uterus.84 Mares identified with placentitis should be treated with a combination of antibiotics (i.e., trimethoprim–sulfamethoxazole, 15–30 mg/kg, p.o., q.12 h), supplemental progesterone (or
Figure 234.8 Mummified twin fetuses removed from a mare approximately 340 days after mating. The mare was noted to have significant udder development and ‘came into milk’ at approximately 6 months of gestation. The mammary development regressed and the mare presented with a vaginal discharge near her original due date. It is likely that the mare started lactating following the death of one or both of the twin fetuses.
progestin i.e., altrenogest, 0.088 mg/kg, p.o. q.24 h), and possibly pentoxifylline (8.5 mg/kg, p.o., q.12 h) until the mare foals.85 Premature lactation Premature lactation is one of the most important causes of failure of passive transfer caused by the loss of colostrum.31 Lactation that occurs even a few hours before foaling may significantly reduce the quantity of immunoglobulins available to the neonate. Immunoglobulin content of presuckle colostrum of mares that have run milk before foaling should be estimated by use of a Brix (sugar) refractometer or by determination of colostral specific gravity. A general guideline for owners and foaling personnel is that mares that begin to run milk (i.e., in a continuous stream) in the hours prior to foaling should be milked out and the colostrum stored at 4◦ C for the foal (Figure 234.9).86 In contrast, mares that drip prior to foaling may be left alone, as they do not experience as much colostrum loss. When in doubt, it is certainly better to collect the colostrum instead of letting it run onto the ground. Banked colostrum can be administered to the foal by bottle or by stomach tube. Foals born to mares that lost significant colostrum before foaling, had poor quality colostrum at foaling, or had an inadequate volume of colostrum should be supplemented with frozen–thawed colostrum, a lyophilized immunoglobulin preparation, a concentrated serum product, or plasma.87 The plasma immunoglobulin concentration of foals may be assessed
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Figure 234.9 Mare running milk prior to foaling.
at 12 hours of age to evaluate passive transfer when oral administration may still be effective or at ≥ 24 hours of age after the period of passive transfer is complete. No specific treatment for premature lactation exists because it is usually a symptom of another underlying disease. The efficacy of progestagen therapy for mares that have premature udder enlargement and lactation during midgestation is not known.
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of galactorrhea in these animals are not well defined. The most commonly identified cause of inappropriate lactation in mares is an elevation in prolactin secondary to ECD.90 Consequently, ECD should be ruled out before other potential causes of inappropriate lactation are considered. In many instances, udder development is incorrectly attributed to mastitis. A brief manual examination of the udder and evaluation of the character of the secretion is often sufficient to rule out mastitis, although collection of samples for culture and cytology may be warranted. Treatment strategies for inappropriate lactation in mares should be based on the cause. Mares with ECD should be treated with medications such as pergolide mesylate (1–2 mg, q.24 h) or cyproheptadine (0.5 mg/kg q.12 h). Mares exhibiting udder development and lactation not associated with ECD may be managed with a combination of pergolide therapy and dietary management. A reduction in dietary intake of energy and protein may be beneficial to reduce milk production. This should include a discontinuation of any grain-based supplements and a change in forage from alfalfa to grass hay.
Mastitis Inappropriate lactation Neonates occasionally exhibit precocious mammary development with associated secretory activity, referred to as ‘witch’s milk’. This phenomenon, which has been observed in most domestic large animal species, has been attributed to elevations of lactogenic hormones of maternal origin in fetal circulation. 88 The incidence of galactorrhea in human infants has been reported to be approximately 6%.89 Transient glandular development and lactation are also occasionally observed in weanling foals, yearlings, and adult mares with no previous or recent history of pregnancy or lactation (Figure 234.10). Causes
Figure 234.10 Udder development with milk production in a mare that had not been pregnant for several years.
Incidence of mastitis, or inflammation of the mammary gland, is much lower in mares than in dairy cattle. Clinical signs associated with mastitis include a warm, swollen, and painful udder (Figure 234.11); ventral edema; and fever.91, 92 Occasionally, systemic signs of illness, such as depression and anorexia, or a mild degree of lameness in the hindlimb adjacent to the affected mammary gland, will be exhibited. Mastitis usually occurs in adult horses, but has been reported in a 15-week-old filly.93 A seasonal predilection for mastitis has been reported, with most mares affected during the summer.91 Mastitis occurs most
Figure 234.11 Enlarged right hind quarter of a mare with clinical mastitis.
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Table 234.8 Organisms cultured from milk or mammary exudate of mares with clinical mastitis (from McCue and Wilson91 )
Box 234.2 Causes of mammary gland enlargement in mares (adapted from Brendemuehl103 )
Organism Streptococcus zooepidemicus S. viridans S. agalactiae Staphylococcus sp. Actinobacillus suis sp. Pasteurella ureae Enterobacter aerogenes
Number of isolates
Percent of all isolates
7 1 1 2 2 1 1
36.8 5.3 5.3 10.5 10.5 5.3 5.3
commonly during lactation or within 8 weeks after weaning a foal.91, 94, 95 However, mastitis can occur in mares that have never been pregnant or have not lactated in over a year.88, 91,96–98 Diagnosis of mastitis is usually made on the basis of clinical signs. Cytological evaluation and culture of milk or mammary exudate may be used to confirm presence of mastitis. Cytological evaluation of milk samples from mares with clinical and subclinical mastitis often shows a large number of neutrophils.91, 99, 100 The most common bacterial organism obtained from aerobic culture of aseptically collected samples is Streptococcus zooepidemicus (Table 234.8).91, 95, 99101 Rarely, fungal organisms may be the cause of mastitis in a mare.102 A combination of frequent milking, hotpacks or hydrotherapy, systemic antibiotics (i.e., trimethoprim–sulfamethoxazole), and possibly infusion of a bovine intramammary antibiotic preparation is recommended for the treatment of equine mastitis. Non-steroidal anti-inflammatory drugs such as flunixin meglumine or phenylbutazone may be indicated to relieve patient discomfort, decrease inflammation, and stabilize an elevated body temperature. Clinical signs are usually abated within 2–3 days after medical treatment and the udder is almost normal within 1 week.
r r r r r r r r r r r r r
Aberrant larval migrans Abscessation of the mammary gland Avocado toxicity Cutaneous histoplasmosis Failure of a foal to suckle Galactorrhea Inappropriate lactation Mastitis Periparturient udder edema Premammary adipose tissue Trauma Tumors of the mammary gland Weaning of a foal
In some mares, a large plaque of edema may extend from the udder cranially to the chest (Figure 234.12). The enlargement may be identified as edema as it ‘pits’ upon digital pressure. The added weight of edema may predispose mares to rupture of abdominal musculature or the prepubic tendon. Causes of pre-foaling ventral edema are considered to be reduced efficacy of lymphatic circulation owing to the weight of the gravid uterus and a general lack of exercise. Physiological udder edema is most common in primiparous mares. Treatment, which is usually not necessary, consists of light controlled exercise, hydrotherapy, and possibly diuretics. Mares exhibiting significant discomfort may benefit from administration of a non-steroidal anti-inflammatory drug, such as phenylbutazone or flunixin meglumine. Ventral edema generally dissipates within 2–3 days after foaling.
Miscellaneous causes of mammary gland enlargement Additional causes of mammary gland enlargement in mares are presented in Box 2. Udder edema Accumulations of ventral edema during the last weeks of gestation and the periparturient period may result in mammary gland enlargement. Affected mares may exhibit mild discomfort, a reluctance to move, and may refuse to allow their foal to nurse.104
Figure 234.12 Mare with ventral edema prior to foaling.
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Lactation Abscessation of the mammary gland Abscessation of the mammary gland or adjacent lymph nodes in California is most commonly associated with Corynebacterium pseudotuberculosis infections. Other bacteria cultured from mammary abscesses include Streptococcus equi ssp. zooepidemicus, S. equi ssp. equi, Staphylococcus sp., Peptostreptococcus anaerobius, and Actinomyces naeslundii. Clinical signs include a unilaterally swollen, painful mammary gland, ventral edema, and often lameness. Abscessation is usually not associated with lactation. Diagnosis is based on manual palpation of the mammary gland, ultrasonography, as well as cytology and culture of mammary secretion or needle aspirate. Treatment includes the establishment of ventral drainage, lavage, hotpacks or hydrotherapy, antimicrobial agents, and non-steroidal anti-inflammatory drugs.
Tumors of the mammary gland Tumors of the equine mammary gland are rare, especially compared with the incidence in certain other domestic animals such as the bitch.105–107 Feldman108 reported a 0.11% rate of mammary tumors in a retrospective study of horses sent to slaughter in France. In another study, a total of 45 cases of mammary tumors were found during examinations of approximately 20 000 mares in a Paris abattoir.109 More recently, Laugier and colleagues110 noted only one case of mammary carcinoma in 1771 horses necropsied in Normandy between 1986 and 2003. A variety of case reports of individual horses with adenocarcinomas or other invasive tumors have been published.105111–116 Recently, a mammary gland adenoma without metastasis was also reported.117 Skin tumors affecting the mammary gland include sarcoids and, in gray mares, melanomas.
References 1. Getty R. Sisson and Grossman’s the Anatomy of the Domestic Animals. Philadelphia: W.B. Saunders, 1975; pp. 296–7. 2. Mepham TB. Physiology of Lactation. Milton Keynes: Open University Press, 1987; pp. 109–27. 3. Worthy K, Escreet JP, Renton JP, Eckersall PD, Douglas TA, Flint DJ. Plasma prolactin concentrations and cyclic activity in pony mares during parturition and early lactation. J Reprod Fertil 1986;77:569–74. 4. Heidler B, Parvizi N, Sauerwein H, Bruckmaier RM, Heintges U, Aurich JE, Aurich C. Effects of lactation on metabolic and reproductive hormones in Lipizzaner mares. Domest Anim Endocrinol 2003;25:47–59. 5. Forsyth LA, Rossdale PD, Thomas CR. Studies on milk composition and lactogenic hormones in the mare. J Reprod Fertil Suppl 1975;23:631–635. 6. Sharma OP. Release of oxytocin elicited by suckling stimulus in mares. J Reprod Fertil 1974;37:421–3.
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7. Ellendorff F, Schams D. Characteristics of milk ejection, associated intramammary pressure changes and oxytocin release in the mare. J Endocrinol 1988;119:219–27. 8. Blackmer JM, Sellon DC, Hines MT. Failure of passive transfer. In: Smith BP (ed.) Large Animal Internal Medicine, 3rd edn. St. Louis: Mosby, 2002; pp. 1592–5. 9. Genco RJ, Yecies L, Karush F. The immunoglobulins of equine colostrum and parotid fluid. J Immunol 1969;103;437–44. 10. Hines MT. Immunodeficiencies of foals. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 5th edn. Philadelphia: Saunders, 2003; pp. 692–7. 11. Knottenbelt DC, Holdstock N. Methods of assessing colostrum quality. In: Knottenbelt DC (ed.) Equine Neonatology Medicine and Surgery. Edinburgh: Saunders, 2004; pp. 393–4. 12. Knottenbelt DC, Holdstock N. The role of colostrum in immunity. In: Knottenbelt DC (ed.) Equine Neonatology Medicine and Surgery. Edinburgh: Saunders, 2004; pp. 15–18. 13. Curadi MC, Minori D, Demi S, Orlandi M. Immunotransfer in the foal. Ann Facolta Med Vet Pisa 2001;54:127–40. 14. Erhard MH, Luft C, Remler HP, Stangassinger M. Assessment of colostral transfer and systemic availability of immunoglobulin G in newborn foals using a newly developed enzyme-linked immunosorbent assay (ELISA) system. J Anim Physiol Nutr (Berl) 2001;85:164–73. 15. Kohn CW, Knight D, Hueston W, Jacobs R, Reed SM. Colostral and serum IgG, IgA, and IgM concentrations in Standardbred mares and their foals at parturition. J Am Vet Med Assoc 1989;195:64–8. 16. Lavoie JP, Spensley MS, Smith BP, Mihalyi J. Colostral volume and immunoglobulin G and M determinations in mares. Am J Vet Res 1989;50:466–70. 17. LeBlanc MM, Tran T, Baldwin JL, Pritchard EL. Factors that influence passive transfer of immunoglobulins in foals. J Am Vet Med Assoc 1992;200:179–83. 18. Massey RE, LeBlanc MM, Klapstein EF. Colostrum feeding of foals and colostrum banking. In: Proceedings of the 37th Annual Convention of the American Association of Equine Practitioners, 1991, pp. 1–8. 19. Morris DD, Meirs DA, Merryman GS. Passive transfer failure in horses: incidence and causative factors on a breeding farm. Am J Vet Res 1985;46:2294–9. 20. Pearson RC, Hallowell AL, Bayly WM, Torbec RL, Perryman LE. Times of appearance and disappearance of colostral IgG in the mare. Am J Vet Res 1984;45:186–90. 21. Rouse BT, Ingram DG. The total protein and immunoglobulin profile of equine colostrum and milk. Immunology 1970;19:901–7. 22. Shideler RK, Squires EL, Osborne M. Immunoglobulin concentration and blood chemistry parameters in the newborn foal: relationship to sex, type of dam, gestation length and mare serum and colostrums concentrations. In: Proceedings of the 32nd Annual Convention of the American Association of Equine Practitioners, 1986, pp. 145–51. 23. Townsend HG, Tabel H, Bristol FM. Induction of parturition in mares: effect on passive transfer of immunity to foals. J Am Vet Med Assoc 1983;182:255–7. 24. Perryman LE, McGuire TC. Evaluation for immune system failures in horses and ponies. J Am Vet Med Assoc 1980;176:1374–7. 25. Sheoran AS, Karzenski SS, Whalen JW, Chrisman MV, Powell DG, Timoney JF. Prepartum equine rotavirus
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vaccination including strong specific IgG in mammary secretions. Vet Rec 2000;146:672–3. McGuire TC, Crawford TB, Hallowell AL, Macomber LE. Failure of colostral immunoglobulin transfer as an explanation for most infections and deaths of neonatal foals. J Am Vet Med Assoc 1977;170:1302–4. McGuire TC, Crawford TB. Passive immunity in the foal: measurement of immunoglobulin classes and of specific antibody. Am J Vet Res 1973;34:1299–303. LeBlanc MM, McLaurin BI, Boswell R. Relationships among serum immunoglobulin concentration in foals, colostral specific gravity, and colostral immunoglobulin concentration. J Am Vet Med Assoc 1986;189:57–60. Jeffcott LB. Duration of permeability of the intestine to macromolecules in the newly-born foal. Vet Rec 1971;88:340–1. Jeffcott LB. Studies on passive immunity in the foal. J Comp Pathol 1974;84:93–101. Jeffcott LB. Passive transfer of immunity to foals. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia: Saunders, 1987; pp. 210–15. Cash RSG. Colostral quality determined by refractometry. Equine Vet Educ 1999;11:36–8. Waelchli RO, Hassig M, Eggenberger E, Nussbaumer M. Relationships of total protein, specific gravity, viscosity, refractive index and latex agglutination to immunoglobulin G concentration in mare colostrums. Equine Vet J 1990;22:39–42. Fleenor WA, Stott GH. Single radial immunodiffusion analysis for quantitation of colostral immunoglobulin concentration. J Dairy Sci 1981;64:740–7. Mancini G, Carbonara AO, Heremans JF. Immunochemical quantitation of antigens by single radial immunodiffusion. Immunochemistry 1965;2:235–54. Chavatte P, Cl´ement F, Cash R, Grongnet JF. Field determination of colostrum quality by using a novel, practical method. In: Proceedings of the 44th Annual Convention of the American Association of Equine Practitioners, 1998, pp. 206–9. Gibbs PG, Potter GD, Blake RW, McMullan WC. Milk production of Quarter Horse mares during 150 days of lactation. J Anim Sci 1982;54:496–9. Ullrey DE, Struthers RD, Hendricks DG, Brent BE. Composition of mare’s milk. J Anim Sci 1966;25:217–22. Lukas VK, Albert WW, Owens FN, Peters A. Lactation of Shetland mares. J Anim Sci 1972;34:350. Oftedal OT, Jenness R. Interspecies variation in milk composition among horses, zebras and asses (Perisodactyla: Equidae). J Dairy Res 1988;55:57–66. National Research Council. Nutrient Requirements of Horses, 5th edn. Washington DC: National Academy Press, 1989. Linton RG. The composition of mare’s milk. II. The variation in composition during lactation. J Dairy Res 1937;8:143–72. Schryver HF, Oftedal OT, Williams J, Cymbaluk NF, Antczak D, Hintz HF. A comparison of the mineral composition of milk of domestic and captive wild equids (Equus Przewalski, E. zebra, E. burchelli, E. caballus, E. assinus). Comp Biochem Physiol 1986;85A:233–5. Schryver HF, Oftedal OT, Williams J, Soderholm LV, Hintz HF. Lactation in the horse: the mineral composition of mare milk. J Nutr 1986;116:2142–7. Peaker M, Rossdale PD, Forsythe LA, Falk M. Changes in mammary development and the composition of secretion
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50. 51. 52.
53.
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during late pregnancy in the mare. J Reprod Fertil Suppl 1979;27:555–61. Leadon DP, Jeffcott LB, Rossdale PD. Mammary secretions in normal spontaneous and induced premature parturition in the mare. Equine Vet J 1984;16: 256–9. Ousey JC, Dudan F, Rossdale PD. Preliminary studies of mammary secretions in the mare to assess foetal readiness for birth. Equine Vet J 1984;16:259–63. Knottenbelt DC. The mammary gland. In: Equine Stud Farm Medicine and Surgery. Edinburgh: Saunders, 2003; pp. 343–52. Naylor JM, Bell RJ. Feeding the sick or orphaned foal. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia: Saunders, 1987; pp. 205–9. Naylor JM, Bell R. Raising the orphan foal. Vet Clin North Am Equine Pract 1985;1:169–78. Rossdale PD, Ricketts SW. Equine and Stud Farm Medicine, 2nd edn. Philadelphia: Lea & Febiger, 1980. Carson K, Woods-Gush DGM. Behaviour of Thoroughbred foals during nursing. Equine Vet J 1983;15: 257–62. King SS, Nequin LG. An artificial rearing method to produce optimum growth in orphaned foals. J Equine Vet Sci 1989;9:319–22. Knight DA, Tyznik WJ. The effect of artificial rearing on the growth of foals. J Anim Sci 1985;60:1–5. Blood DC, Radostits OM, Henderson JA. Veterinary Medicine, 6th edn. London: Bailliere Tindall, 1983. Hintz HF. Eclampsia. In: Robinson NE (ed.) Current Therapy in Equine Medicine. Philadelphia: Saunders, 1983; p. 111. Cowles RR. Lactation failure in the mare. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1983, pp. 222–33. Green EM, Loch WE. Fescue-induced agalactia in mares: management and treatment. In: Proceedings of the American College Veterinary Internal Medicine Forum, 1990, pp. 551–3. Henton JE, Lothrop CD, Dean D, Waldrop V. Agalactia in the mare: a review and some new insights. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1983, 203–21. Evans TJ, Youngquist RS, Loch WE, Cross DL. A comparison of the relative efficacies of domperidone and reserpine in treating equine ‘fescue toxicosis.’ In: Proceedings of the 45th Annual Convention of the American Association of Equine Practitioners, 1999, pp. 207–9. Copetti MV, Santurio JM, Boeck AAP, Silva RB, Bergermaier LA, Lubeck I, Leal ABM, Leal AT, Alves SH, Ferreiro L. Agalactia in mares fed with grain contaminated with Claviceps purpurea. Mycopahologia 2001;154:199–200. Riet-Correa F, Mendez MC, Schild AL, Bergamo PN, Flores WN. Agalactica, reproductive problems and neonatal mortality in horses associated with the ingestion of Claviceps purpurea. Aust Vet J 1988;65:192–3. Lezica RP, Filip R, Gorzalczany S, Ferraro G, de Erausquin GA, Rivas C, Ladaga GJB. Prevalence of ergot derivatives in natural ryegrass pastures: detection and pathogenicity in the horse. Theriogenology 2009;71:422–31. Lothrop CD, Henton JE, Cole BB, Nolan HL. Prolactin response to thyrotrophin-releasing hormone stimulation in normal and agalactic mares. J Reprod Fertil Suppl 1987;35:277–80.
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Lactation 65. Floss HG, Cassady JM, Robbers JE. Influence of ergot alkaloids on pituitary prolactin and prolactin-dependent processes. J Pharm Sci 1973;62:699–715. 66. Ireland FW, Loch WE, Worthy K, Anthony RV. Effects of bromocriptine and perphenazine on prolactin and progesterone concentrations in pregnant pony mares during late gestation. J Reprod Fertil 1991;92:179–86. 67. Caudle AB, Thompson FN, Kemppainer RJ, Nett TM, Brown J, Williams DJ. Effects of thyrotropin releasing hormone in agalactic mares. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1984, pp. 235. 68. Johnson AL, Becker SE. Effects of physiologic and pharmacologic agents on serum prolactin concentrations in the non-pregnant mare. J Anim Sci 1987;65:1292–7. 69. Brander GC, Pugh DM, Bywater RJ. Veterinary Applied Pharmacology & Therapeutics, 4th edn. London: Bailliere Tindall, 1982. 70. Loch W, Worthy K, Ireland F. The effect of phenothiazine on plasma prolactin levels in non-pregnant mares. Equine Vet J 1990;22:30–2. 71. Redmond LM, Cross DL, Strickland JR, Kennedy SW. Efficacy of domperidone and sulpiride as treatments for fescue toxicosis in horses. Am J Vet Res 1994;55: 722–9. 72. Divers TJ. Pergolide and cyproheptadine: which medications to chose for treatment of equine Cushing’s disease. J Equine Vet Sci 2008;28:370–1. 73. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995;73:899–908. 74. Chavatte-Palmer P, Arnaud G, Duvaux-Ponter C, Brosse L, Bougel S, Daels P, Guillaume D, Clement F, Palmer E. Quantitative and qualitative assessment of milk production after pharmaceutical induction of lactation in the mare. J Vet Intern Med 2002;16:472–7. 75. Daels PF, Duchamp G, Porter D. Induction of lactation and adoption of foals by non-parturient mares. In: Proceedings of the 48th Annual Convention of the American Association of Equine Practitioners, 2002, pp. 68–71. 76. Bennett-Wimbush K, Loch W. A preliminary study of the efficacy of fluphenazine as a treatment for fescue toxicosis in gravid pony mares. J Equine Vet Sci 1998;18:169–73. 77. Ryan PL, Bennett-Wimbush K, Vaala WE, Bagnell CA. Systemic relaxin in pregnant pony mares grazed on endophyte-infected fescue: effects of fluphenazine treatment. Theriogenology 2001;56:471–83. 78. Singh MP, Chanel KS. Treatment of agalactia in two mares. Equine Vet Educ 1997;9:60–1. 79. Steiner JV. How to induce lactation in non-pregnant mares. In: Proceedings of the 52nd Annual Convention of the American Association of Equine Practitioners, 2006, pp. 259–60. 80. Daels PF. Induction of lactation and adoption of the orphan foal. In: Proceedings of the 8th American Association of Equine Practitioners Resort Symposium, 2006, pp. 1–6. 81. Kelley KK, Thompson Jr DL, Storer WA, Mitcham PB, Gilley RM, Burns PJ. Estradiol interactions with dopamine antagonists in mares: prolactin secretion and reproductive traits. J Equine Vet Sci 2006;26:517–28. 82. Shideler RK. Prenatal lactation associated with twin pregnancy in the mare: a case report. J Equine Vet Sci 1987;7:383–5. 83. Whitwell KE. Fetal membrane abnormalities. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia: Saunders, 1987; pp. 528–31.
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84. Renaudin CD, Troedsson MHT, Gillis CL, King VL, Bodena A. Ultrasonographic evaluation of the equine placenta by transrectal and transabdominal approach in the normal pregnant mare. Theriogenology 1997;47:559–73. 85. Macpherson ML, Bailey CS. Treating the mare with placentitis: a clinical approach. J Equine Vet Sci 2008;20:703–7. 86. Chavatte P. Lactation in the mare. Equine Vet Educ 1997;9:62–67. 87. Vivrette S. Colostrum and oral immunoglobulin therapy in newborn foals. Compendium 2001;23:286–91. 88. Roberts MC. Pseudomonas aeruginosa mastitis in a dry non-pregnant pony mare. Equine Vet J 1986;18:146–7. 89. Madlon-Kay DJ. Witch’s milk. Am J Dis Child 1986;140:252–3. 90. McCue PM. Equine Cushing’s disease. North Am Vet Clin Equine 2002;18:533–43. 91. McCue PM, Wilson WD. Equine mastitis: a review of 27 cases. Equine Vet J 1989;21:351–3. 92. Perkins NR, Threlefall WR. Mastitis in the mare. Equine Vet Educ 1993;5:192–5. 93. Pugh DG, Magnusson RA, Modransky PD, Darnton KR. A case of mastitis in a young filly. J Equine Vet Sci 1985;5:132–4. 94. Addo PB, Wilcox GE, Taussig R. Mastitis in a mare caused by C. ovis. Vet Rec 1974;95:193. 95. Al-Graibawi MAA, Sharma VK, Ali SK. Mastitis in a mare. Vet Rec 1984;115:383. 96. O’Reilly LM. The isolation of Bazeley and Battle’s type 5 Streptococcus from a case of equine mastitis. Irish Vet J 1965;19:198–202. 97. Prentice MWM. Mastitis in the mare. Vet Rec 1974;94:380. 98. Welsh RD. The significance of Streptococcus zooepidemicus in the horse. Equine Pract 1984;6:6–16. 99. Freeman KP, Roszel JF, Slusher SH, Young D. Cytologic features of equine mammary fluids: normal and abnormal. Comp Cont Educ Pract Vet 1988;10:1090–9. 100. Freeman KP. Cytological evaluation of the equine mammary gland. Equine Vet Educ 1993;5:212–13. 101. Strong MG. Mastitis in the mare. Vet Rec 1974;94:526. 102. Walker RL, Johnson BJ, Jones KL, Pappagianis D, Carlson GP. Coccidioides immitis mastitis in a mare. J Vet Diagn Invest 1993;5:446–8. 103. Brendemuehl JP. Mammary gland enlargement in the mare. Equine Vet Educ 2008;20:8–9. 104. Spensley MS. Alterations in lactation. In: Smith BP (ed.) Large Animal Internal Medicine. St. Louis: C.V. Mosby, 1990; pp. 258–62. 105. Acland HM, Gillette DM. Mammary carcinoma in a mare. Vet Pathol 1982;19:93–5. 106. Moulton JE. Tumors in Domestic Animals. Berkeley: University of California Press, 1961. 107. Munson L, Moresco A. Comparative pathology of mammary gland cancers in domestic and wild animals. Breast Dis 2007;28:7–21. 108. Feldman WH. Malignant growths in domestic animals. J Am Vet Med Assoc 1929;75:192–200. 109. Martel H. Cancer in horses. Am Vet Rev 1913–14; 44:299–300. 110. Laugier C, Tapprest J, Foucher N, Doux N, Geroge C, Longeart L, le Net JL. Prevalence of equine tumours in 1771 horses examined post-mortem. Pract Vet Equine 2004;36:21–35. 111. Munson L. Carcinoma of the mammary gland in a mare. J Am Vet Med Assoc 1987;191:71–2.
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112. Schmahl VW. Solides karzinom der mamma bei einem pferd. Berl Munch Tierarztl Wochenschr 1972;85: 141–2. 113. Foreman JH, Weidner JP, Parry BW, Hargis A. Pleural effusion secondary to thoracic metastatic mammary adenocarcinoma in a mare. J Am Vet Med Assoc 1990;197: 1193–5. 114. Seahorn TL, Hall G, Brumbaugh GW, Honnas CM, Lovering SL, Snyder JR. Mammary adenocarcinoma in four mares. J Am Vet Med Assoc 1992;200:1675–7.
115. Reppas GP, McClintock SA, Canfield PJ, Watson GF. Papillary ductal adenocarcinoma in the mammary glands of two horses. Vet Rec 1996;138:518–19. 116. Hirayana K, Honda Y, Sako T,. Okamoto M, Tsunoda N, Tagami M, Taniyama H. Invasive ductal carcinoma of the mammary gland in the mare. Vet Pathol 2003;40:86–91. 117. Spadari A, Valentini S, Sarli G, Spinella G, Millanta F. Mammary adenoma in a mare: clinical, histopathological and immunohistochemical findings. Equine Vet Educ 2008;20:4–7.
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Chapter 235
Uterine Involution Mary E. Stanton
Uterine involution in the mare is a very efficient process. It is remarkable that a mare can produce a 50 kg foal and be ready to conceive within 2 weeks. Mare gestation length averages 340 days, and unless mares conceive during foal heat their subsequent foaling will occur approximately 10 days later in each successive year.1, 2 In breeds with a finite reproductive season, late-foaling mares may have an open year resulting in a loss of revenue from the lack of a foal and the costs associated with supporting a barren mare.1 Therefore, from a management perspective it is imperative that mares average 25 days from foaling to conception in order to maintain a sustainable yearly foaling schedule.3 The success of this endeavor depends on the process of uterine involution and appropriate timing of foal heat breeding. The equine placenta is favorably structured to allow for a rapid return to fertility following parturition. The combination of a microcotyledonary placenta and a potentially fertile postpartum estrus makes it possible for mares to maintain the goal of foaling annually. Normal foaling results in little damage to fetal or maternal tissues4 and fetal membranes are typically expelled approximately 60 minutes after foaling.5 The normal maternal contact surface of the allantochorionic membrane has a uniform distribution of tufts of microvilli that appear similar to a red velvet surface. The microvilli have a central vascular connective tissue core covered by a single layer of chorionic epithelium.4 In stillborn births and cases of placentitis there is often a degenerative appearance of the chorionic epithelium with an absence of microvilli and fragmentation of the plasma membrane.4 The microvilli of the chorion form microcotyledons that attach to maternal crypts. The crypts or microcaruncles of the mare are located within the endometrium, unlike the cow, in which the caruncles are outgrowths that must be sloughed as a part of involution.6 The fetal–maternal surface is interfaced by an electron dense material that connects the fetal
and maternal cell membranes. Normal placental separation involves dissolution of this material allowing disengagement of the microvilli from the maternal crypts.4 Placental separation generally leaves the uterine epithelium undamaged. In mares that experience uncomplicated pregnancies and parturition the uterus is grossly normal in terms of size and fluid content by 7 days post-foaling. Histological evaluation, however, indicates that uterine involution occurs over an average of 14 days. Uterine biopsy samples taken within 48 hours after placental expulsion show stratified uterine epithelium. The cells vary between cuboidal to low columnar with the presence of edema.4, 6 Maternal septae are present which extend from the luminal surface into the stratum spongiosum. The septae are formed by clusters of primary and secondary epithelial cell lined crypts.6 The maternal crypts are complex, described as a spongy labyrinth joined by junctional complexes associated with a fine fibrillar material. The lateral membranes of the cells are highly convoluted and the concave surfaces show little disruption. Some of the cells containing myelin figures appear compressed. The number of compressed cells increases as the time from placental expulsion elapses.6 The crypts appear shallower by 80 hours after the placenta is passed.4 The majority of the lumina are empty with a few polymorphonuclear cells (PMNs). There is evidence of karyolysis at the level of the microcaruncle and a diffuse mild infiltrate of inflammatory cells (PMNs) in the stratum compactum below the luminal epithelium by day 1 postpartum.7 The stratum spongiosum contains many dilated glands with various amounts of cellular debris. Epithelial cells lining the glands are columnar and vacuolated.7 In contrast, biopsies taken from mares who delivered stillborn foals show significant uterine epithelial damage. Large numbers of PMNs were found in the crypt lumens and between epithelial cells. The crypts also contained a large amount of cellular
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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material thought to be of fetal origin. Manual removal of a placenta resulted in similar histological changes. In contrast, two mares who presented with dystocia related to malpresentation were also biopsied within 14 hours of parturition and did not show significant uterine epithelial damage.4 In the mare microcaruncles are reduced by karyorrhexis and cytoplasmic vacuolization with subsequent cell lysis and phagocytosis, or shrinking of the maternal epithelial cells, condensation of their contents and collapse of the crypt lumen.6 The process is rapid, but occurs more slowly in mares with induced parturition.7 The connective tissue capsule surrounding the microcotyledon at 12 hours postpartum is not visible by day 3 and 4 postpartum, and lumina are not seen at day 3.4, 7 By day 5 the microcaruncles consist of lymphocytes, macrophages, and degraded epithelial cells.6 The level of inflammation seen is variable.6 By day 6 or 7 postpartum the microcaruncles are either no longer present or consist of focal areas of crypts lined with contracted or vacuolated epithelial cells.6 Siderocytes are numerous in the stratum compactum by day 8 with a sparser distribution through the stratum spongiosum.7 At day 11, a common time for foal heat ovulation, cystic distension of uterine glands has resolved in areas without periglandular fibrosis. The endometrium appears normal by day 14 with the presence of siderocytes and sporadic inflammatory changes. Inflammation was noted inconsistently throughout a 32-day observation period.6 The physiological activity of reproductive hormones also plays a role in uterine involution. Progesterone blocks uterine contractions, but decreases in late pregnancy and remains low until the postpartum ovulation.8–10 Plasma estrogen is low at the time of parturition, but increases corresponding to the recruitment of follicles and the onset of postpartum estrous.8, 10 Estrogen has the beneficial effect of facilitating uterine contraction to return the uterus to its pre-gravid size. Additionally, estrogen influences the recruitment of neutrophils to remove bacteria and debris, improving the inflammatory status of the uterine environment. The uterus is generally contaminated by bacteria during stages two and three of labor, particularly during assisted foaling in which an attendant may inadvertently introduce bacteria while assessing fetal position or attempting to assist delivery.11 The uterus is designed to clear bacteria by passing the placenta and the associated contractions that move debris caudally through the cervix. Lochia is initially seen as a red blood-tinged discharge that turns brown and mucoid. The quantity decreases through the first postpartum estrus and should resolve by day 15 post-foaling.12 Species of bacteria reported
on cultures from postpartum mares include Streptococcus equi zooepidemicus, Escherichia coli, Enterobacter spp, Klebsiella spp, Pseudomonas spp, Staphylococcus aureus, Acinetobacter calcoaceticus, Corynebacerium spp, Pasteurella spp, Bacillus spp, and Proteus spp.6, 7,11–14 The most common organism isolated is Streptococcus equi zooepidemicus though mixed infections are common.11 Positive uterine cultures from mares to be bred on foal heat are not unusual. They may accompany cytology results that are positive for inflammation or be an indication of contamination without actual pathology. Many techniques have been explored to promote uterine involution and foal heat breeding. One of the simplest methods is to ensure that the mare is allowed to exercise to help with evacuation of uterine fluid. Mares that are stalled because of a sick foal should be evaluated in case intervention is needed to prevent metritis.15 Uterine lavage using warm physiological saline has been evaluated in normal mares and found to have no benefit in improving pregnancy rates.1 Lavage does not affect uterine body or horn diameter, the presence of fluid in the lumen, histological characteristics of the involuting uterus, or the number of mares which ovulate by day 11 postpartum.1 There is potential to introduce bacteria through uterine lavage techniques. Ecbolics have been evaluated in several different regimens. Oxytocin administered at a dose of 20 units i.v. or fluprostenol given at 500 µg i.m. twice daily for 10 days postpartum were proven ineffective for increasing the rate of involution. Uterine size and fluid expulsion were not appreciably altered, potentially due to duration of action of the ecbolics or the dosing regimen. It is possible that physiological doses given more frequently may be efficacious.14 Oxytocin 30 units i.m. and cloprostenol 250 µg im given 12 hours after parturition and continuing every 12 hours for 3 days had no effect on decreasing uterine diameter, time of foal heat ovulation, pregnancy rates, or early embryonic death.16 However, prostalene has successfully been used in postpartum mares beginning on the day of foaling and continuing for 10 days or until the mare was first bred at foal heat. In this study,17 by comparison, 76.9% of treated mares versus 44.4% of control mares became pregnant. Successful treatment of mares to enhance foal heat fertility may focus on timing ovulation to coincide with complete histological involution. Progesterone 150 mg and estradiol-17β 10 mg i.m. are used to delay postpartum estrus. Treatment should begin within 18 hours of foaling. This can effectively delay the initial wave of follicular recruitment to allow more time for involution. Histologically, treated mares showed greater uterine gland proliferation and density, and the proportion of ciliated to secretory
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Uterine Involution epithelial cells lining the endometrium was greater.13 Progestin alone did not affect uterine size or fluid accumulation during involution as detected by ultrasonography.18 However, using altrenogest to delay ovulation beyond day 8 postpartum enhanced pregnancy rates.18 A recent retrospective study of foal heat breeding shows pregnancy rates of 72% as compared to 76% for later postpartum estrous cycles.19 Foal heat pregnancy rates were negatively affected by increasing age of the mare and mating by natural service. The presence of intraluminal fluid which required treatment decreased postpartum estrus pregnancy rates18, 19 and increased early embryonic death loss rates.19 This contrasts with earlier reports of mean foal heat pregnancy rates of 44.1% to 60% for mares bred naturally and artificially inseminated.7, 20 However, this does not take into account potential differences among breeds and among methods of mare management. Uterine involution is a combination of reduction of uterine size, removal of debris, glandular contraction, and alteration of the endometrial surface to create an environment that is hospitable to a new conceptus. It is a rapid process in the horse, functionally complete by day 7 and histologically complete by day 14.12 Stillbirths, dystocia, and placentitis delay involution through damage to the placenta resulting in disruption of normal placental separation and retention of fetal debris.4 In normal mares uterine lavage and administration of ecbolic agents has not enhanced uterine involution.1, 14, 16 Mares that give birth without complications and subsequently have an uneventful involution period have foal heat pregnancy rates that are comparable to breeding on subsequent cycles.19
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References 1. Blanchard TL, Varner DD, Brinsko SP, Meyers SA, Johnson L. Effects of postparturient uterine lavage on uterine involution in the mare. Theriogenology 1989;32:527–35. 2. Lenz TR. One practitioner’s approach to foal heat breeding. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 1986; pp. 111–19. 3. Zent W. The postpartum breeding mare. In: Samper JC, Pycock J. McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; p. 455. 4. Steven DH, Jeffcoat LB, Mallon KA, Ricketts SW, Rossdale PD, Samuel CA. Ultrastructural studies of the equine
19.
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uterus and placenta following parturition. J Reprod Fert Suppl 1979;27:579–86. Rossdale PD, Ricketts SW. The Practice of Equine Stud Medicine. London: Bailli`ere Tindall, 1974; pp. 165–7. Gygax AP, Ganjam VK, Kenney RM. Clinical, microbiological and histological changes associated with uterine involution in the mare. J Reprod Fert Suppl 1979;27:571–8. Bailey JV, Bristol FM. Uterine involution in the mare after induced parturition. Am J Vet Res 1983;44:793–7. Hillman RB, Loy RG. Estrogen secretion in mares in relation to various reproductive states. In: Proceedings of the American Association of Equine Practitioners, 1969; 111–19. McDonald LW. Veterinary Endocrinology and Reproduction, 3rd edn. Philadelphia: Lea & Febiger, 1980. Nett TM, Holtan DW, Estergreen VL. Plasma estrogens in pregnant and postpartum mares. J Anim Sci 1973;37: 962–70. Card C, Lopate C. Infectious diseases of the puerperal period. In: Youngquist RS, Threlfall WR (eds) Current Therapy in Large Animal Theriogenology, 2nd edn. St Louis: Saunders, Elsevier, 2007; pp. 138–43. Blanchard TL, Varner DD. Uterine involution and postpartum breeding. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 622–5. Sexton PE, Bristol FM. Uterine involution in mares treated with progesterone and estradiol-17β. J Am Vet Med Assoc 1985;186:252–6. Blanchard TL, Varner DD, Brinsko SP, Quirk K, Rugila JN, Boehnke . Effects of ecbolic agents on measurements of uterine involution in the mare. Theriogenology 1991;36:559–71. LeBlanc MM. Immediate care of the postpartum mare and foal. In: Youngquist RS, Threlfall WR (eds) Current Therapy in Large Animal Theriogenology, 2nd edn. St Louis: Saunders, Elsevier, 2007; pp. 134–8. Gunduz MC, Kasickci G, Kaya HH. The effect of oxytocin and PGF2α on the uterine involution and pregnancy rates in postpartum Arabian mares. Anim Reprod Sci 2008;104:257–63. Ley WB, Purswell BJ, Bowen JM. The effects of prostalene and alfaprostol as uterine myotonics, and the effect on postpartum pregnancy rate in the mare following daily treatment with prostalene. Theriogenology 1988;29:1113–22. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproductive tract of mares after parturition: Effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed first postpartum ovulatory cycles. J Am Vet Med Assoc 1988;192:350–3. Blanchard TL, Thompson JA, Brinsko SP, Stich KL, Wendt KM, Rigby SL. Mating mares on foal heat: A Five-Year Retrospective Study. In: Proceedings of the American Association of Equine Practitioners, 2004; pp. 525–30. Kattila T, Koskinen E, Oijala M, Parviainen P. Evaluation of the postpartum mare in relation to foal heat breeding. II. Uterus swabbing and biopsies. J Am Vet Med Assoc 1988;35:331–9.
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Chapter 236
Breeding Mares on Foal Heat Terry L. Blanchard and Margo L. Macpherson
Introduction Maintaining yearly foal production is important for economic reasons. Income is usually generated from selling offspring, which offsets maintenance and breeding expenses incurred by the mare owner. Since mares have an average gestational length of 333 to 345 days,1 they must become pregnant within 1 month postpartum to continue producing foals each year. Mating mares on the first postpartum estrus (foal heat, which begins 5–12 days after foaling2 with most mares ovulating within 20 days postpartum3 ) is one method used to improve the chance of maintaining yearly foal production. Advocates for mating on foal heat cite the following advantages: (i) lowering the odds that mares foaling early in the year will re-enter anestrus after the foal heat ovulation,3–6 which would increase the parturition-to-conception interval; (ii) avoiding a delay (3 weeks or more) in the foaling date the next year compared to mares mated for the first time on their second postpartum estrus;7, 8 and (iii) cumulative season pregnancy rates are often similar between mares mated for the first time on their foal heat and mares mated for the first time on later postpartum estrous periods.3, 9 Those that routinely avoid mating on the first postpartum estrus cite the following disadvantages: (i) lower (perhaps averaging 20%) pregnancy rates achieved for mares mated on foal heat compared to mares mated for the first time on later postpartum estrous periods;2, 3, 10, 11 and (ii) potential for greater pregnancy losses may occur in mares conceiving on foal heat matings compared to mares conceiving on later postpartum estrous periods.12 The mare is unique among domestic large animal species in regard to the early return to a fertile estrus after parturition. Major factors thought to be related to fertility achieved on foal heat breeding include an uncomplicated birth free from genital tract trauma, prompt passage of the fetal membranes, rapid uterine
involution including expulsion of lochia, and early return to regular estrous cycles.
Uterine involution Uterine involution after parturition is a complex set of events that restores the readiness of the uterus for re-establishing pregnancy. Involution occurs rapidly after a normal parturition. Factors such as tissue remodeling, uterine contractility, and reduction in uterine size and fluid content are linked to successful uterine involution. In normal foaling mares, the villi of the diffuse epitheliochorial placenta separate neatly at the maternal–fetal interface requiring minimal endometrial remodeling after placental expulsion.13 The endometrial surface is intact (as viewed by scanning electron microscopy) and contains prociliated and secretory cells on the first day after parturition.14 Cilia become longer as prociliated cells become ciliated, conspicuously appearing around gland orifices. Secretory products are visible, although not as prominently as in fully involuted endometria. Histologically, the microcaruncles are completely resorbed by day 7 postpartum.15, 16 Glandular distention, which is prominent in pregnancy, may resolve as early as day 4 postpartum.16 Glandular density, depth of gland penetration into the lamina propria, and luminal epithelial cell height progressively increase in the first 1–2 weeks postpartum.17–19 Further evidence for an increased glandular activity is the observed increase in mitotic activity of the glandular epithelial cells through day 12 postpartum.16 An influx of neutrophils into the endometriuim is common up to day 5 postpartum, followed by a steady decline in numbers through the first postpartum estrus.15 Lymphocytes become the predominant inflammatory cell type after day 5 postpartum. Siderophages are present in large numbers in foaling mares, and often persist (albeit in decreasing numbers with less prominent stain uptake) for up to 7 months or longer. The
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Breeding Mares on Foal Heat endometrium of most mares appears histologically indistinguishable from its fully involuted pre-gravid state by day 14 postpartum.15 Several gross uterine changes occur during involution. Presumably, increased uterine contractility contributes to reduction of uterine size and evacuation of contents, but how important contractile activity is in the reduction of uterine size is not known. Little published data is available regarding contractility patterns of the postpartum mare. Using electromyographic measurements, Haluska and coworkers20 demonstrated that contractile activity increased abruptly after placental passage. This activity was replaced by low-level, erratic bursts in the early days following parturition. Griffin and Ginther21 used ultrasonography to measure uterine contractility relative to other morphological assessments in postpartum mares. Subjective measures of contractile activity were made daily with longitudinal scans of the uterine body completed at 1-minute intervals. They detected minimal contractile activity in the immediate postpartum period and after the first ovulation. From their findings, the workers concluded that uterine contractility plays a minimal role in uterine involution and evacuation of uterine contents. The authors conceded that the measurement times were short, and that episodic bursts of uterine contractility may have been missed. To date, uterine contractility in the postpartum mare has been inadequately studied, and the role of resorption or mechanical evacuation of uterine contents in involution is unclear. Certainly, rapid reduction in uterine length and diameter occurs in the early postpartum period of the mare. Using ultrasonography to measure changes, uterine diameter was found to reach a pre-gravid state in a mean of 23 days postpartum.22 Discrepancies between the previously non-gravid and gravid horns can be expected to resolve as early as day 15 postpartum,23 or as late as day 35 postpartum in some mares.21 Exercise probably may also play a role in gross involution of the uterus, as ultrasonographic measurements taken at 5-day intervals in the postpartum period revealed the mean diameter of the previously gravid uterine horn reduced more rapidly in mares maintained on pasture compared to mares confined to stalls.24 Luminal contents (lochia) are discharged from the uterus via the cervix during uterine involution. Complete cervical closure is dependent on progesterone, which is lacking from the circulation until corpus luteum formation occurs following the first postpartum ovulation.15 Lochial discharge is typically seen at the vulva as an exudate between days 3 and 6 postpartum.25 Color of lochia varies from red to reddish brown, but sometimes can be yellowish when mucopurulent. Normal lochia is not malodorous; it is
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easily distinguished from the fetid uterine discharge of mares with acute early postpartum metritis. Ultrasonographically, intrauterine fluid is frequently seen in the first few days after foaling.21, 22 The echogenic character of postpartum fluid probably varies with cellularity and mucus content. McKinnon et al.22 reported that fluid echogenicity decreased as normal involution progressed, while others17, 18, 21 reported that intrauterine fluid of involuting mares was minimally echogenic with little change in ultrasonographic character occurring over time. Uterine contents should begin to decrease around day 5 in normal mares, and fluid should be minimal by day 10 and undetectable by day 15 postpartum.17, 18, 21, 22 However, mares confined during the postpartum period may retain intrauterine fluid longer than exercised mares (Macpherson, personal observation). McKinnon and co-workers22 examined the effect of intrauterine fluid accumulation on pregnancy rates of mares bred on the first postpartum estrus. Mares with ultrasonographically detectable uterine fluid accumulations during the ovulatory period had significantly lower pregnancy rates than mares without fluid. Pycock and Newcombe26 demonstrated that the presence of any intrauterine fluid at the time of breeding (even as little as 1–20 mm distension of the uterine lumen) in both lactating and non-lactating mares resulted in reduced pregnancy rates compared to mares without intrauterine fluid present at the time of breeding. Blanchard et al.9 reported that even when foaling mares bred on foal heat were treated to remove intrauterine fluid, pregnancy rates were lower and subsequent pregnancy losses were higher than in mares bred on foal heat when no intrauterine fluid was present. Results from these studies suggest that the presence of intrauterine fluid on the first postpartum estrus may be a good clinical indicator that involution is not complete and the uterus is therefore not ready to establish a viable pregnancy, and confirm that breeding should be delayed until intrauterine fluid no longer remains.
Treatment effects on uterine involution A number of treatment protocols have been tried in an effort to hasten uterine involution and thus improve postpartum pregnancy rates in normal foaling mares.17–19,27–30 Treatments have included repeated ecbolic (oxytocin, prostaglandin) administration, performance of single or repeated large volume uterine lavage(s), and administration of steroid hormones (altrenogest or progesterone, estradiol, or progesterone plus estradiol) for a varying number of days in the early postpartum period. Assessment
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of gross and histological involution (including intrauterine fluid retention, and recovery of bacteria from endometrial swabbing in some studies) failed to demonstrate an improvement in any of these parameters in the normal foaling early postpartum mare. Only one study reported an improvement in pregnancy rates.28 In conclusion, the preponderance of current evidence would suggest that treatments aimed at hastening uterine involution to improve foal heat fertility in normal foaling mares is likely to be unrewarding. In the normal foaling mare, histological involution and fluid expulsion should be complete by 14–15 days after parturition, and gross involution should be complete by 25–32 days after foaling. Using this framework, the practitioner can judge whether postpartum involution is proceeding normally, and can then use this conclusion to support a decision about whether to breed a given mare on foal heat.
Periparturient problems Periparturient problems, including dystocia or retained placenta, have been shown to delay uterine involution.31 Mares experiencing a dystocia frequently suffer trauma to the caudal genital tract that will impair restoration of fertility, and may require several weeks to resolve. Incomplete placental detachment, uterine trauma, and uterine infection retard involution rate in mares with retained placenta.25 Mares with ‘disturbed puerperium’ (including uncomplicated retained placenta) have been shown to have an increased influx of neutrophils in the endometrium, and to require a longer period of time for return of luminal and glandular epithelium to a pre-gravid state (and lower pregnancy rates
when bred) compared to normal foaling mares.32 Steiger et al.31 postulated that uterine hypoxia was caused by dystocia, perhaps due to myometrial exhaustion, that prevents normal desquamation of epithelial cells into the uterine lumen, thus leading to ‘disturbed’ endometrial involution. They also postulated dystocia and retained placenta resulted in an unphysiological delay in glandular re-differentiation that culminated in an endometrium poorly suited for establishing a new pregnancy. Clearly, mares having a difficult birth and/or retained placenta are not good candidates for foal heat breeding.
Characteristics of the first postpartum estrus, and the occurrence of anestrus The first postpartum estrus generally begins 5–12 days after foaling;2 hence, the terminology ‘foal–heat’ or ‘9-day heat’. In a study involving Thoroughbred mares that was performed in central Kentucky, ovulation during foal heat was reported on average at 10 days postpartum, with most mares ovulating within 20 days postpartum.3 The influence of artificial lighting was not critically evaluated in that study. In a study involving Quarterhorse-type mares maintained on pasture in southeast Texas (i.e., nonlighted), the median interval to foal-heat ovulation was 13 days, with 50% of mares ovulating 11–16 days postpartum9 (see Figure 236.1). Loy3 detected an underlying effect of season on the interval from parturition to first ovulation, with a greater incidence of first postpartum ovulations occurring by 10 days postpartum in mares foaling later in the spring than in mares foaling earlier in the winter. He likewise reported that most mares having a prolonged period of postpartum anestrus (> 30 days to first ovulation
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Day of First Postpartum Ovulation
Figure 236.1 Day of first postpartum ovulation for 399 Quarterhorse type mares maintained on pasture in southeast Texas. Mares foaled between late December and early May, and were not exposed to artificial lighting supplementation. Data from mares experiencing postpartum anestrus was excluded.
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Figure 236.2 Incidence of postpartum anestrus by month of foaling in 557 Quarterhorse type mares foaled on pasture in southeast Texas. Mares were not exposed to artificial lighting supplementation.
postpartum) were early foaling mares (i.e., foaling dates of January–March). He also noted that many mares with postpartum anestrus developed follicles at the expected time in the early postpartum period, but failed to ovulate and then entered a variable period of ovarian inactivity. By contrast, in the 5-year study of foaling mares maintained on pasture (unlighted) in southeast Texas, early foaling mares (January–March) that had prolonged intervals to their first postpartum ovulation did not seem to develop large (> 25 mm diameter) follicles and then fail to ovulate; instead, ovaries almost invariably remained small with follicles < 15–20 mm diameter for prolonged periods of time prior to experiencing their first postpartum estrus and ovulation.9 The incidence of mares experiencing postpartum anestrus in that study (see Figure 236.2) decreased from 27% for mares foaling in December–January, to 23% for mares foaling in February, 14% for mares foaling in March, and < 3% for mares foaling in April. No mares foaling in May experienced anestrus. Median duration of postpartum anestrus likewise appeared longer for mares foaling in December–January (67 days) than those foaling in February (52 days), March (45 days), and April (43 days). While not proven, it is suspected that the postpartum anestrus seen in many foaling mares is thus related to season (short daylength) when parturition occurs. In this regard, supplementing artificial light to late pregnant mares has been shown to reduce the incidence of postpartum anestrus.5
Fertility associated with breeding on foal heat Pregnancy rates from mares bred on the first postpartum estrus averaged 20% lower than mares bred in subsequent heat cycles.3, 10 Using multiple logis-
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tic regression to adjust for factors affecting fertility of Thoroughbreds bred by natural service in central Kentucky, mating on foal heat was found to significantly lower the odds of establishing pregnancy (odds ratio of 0.64 compared to foaling mares not bred on foal heat).11 However, not all studies report lowered pregnancy rates in mares bred on foal heat, and whether method of mating affects outcome from foal heat breeding remains undetermined. Camillo et al.33 bred 401 mares using artificial insemination. Mares bred during the first postpartum estrus obtained a 72% (182/253) pregnancy rate. Mares bred for the first time on the second postpartum estrus obtained an 85% (22/26) pregnancy rate. In their study, pregnancy rates for postpartum mares were comparable to those of maiden, barren, and non-lactating mares (78% and 76%) on first and second cycles, respectively. In a 5-year study performed in southeast Texas, Blanchard et al.9 likewise reported comparable first-cycle pregnancy rates in foaling mares mated for the first time on foal heat (72%; 288/399) or for the first time on later postpartum estrous periods (75%; 120/158). In that study, multiple logistic regression was used to adjust for factors found to significantly affect fertility (e.g., year, stallion used for mating, age of mare, method of mating, and whether or not the mare was treated for fluid accumulation in that cycle), and adjusted pregnancy rates for foal heat matings were lower in mares bred by natural cover (66%) than for mares bred by artificial insemination (83%). By contrast, Pycock (personal communication) reviewed 5 years of breeding records and found a 60% per cycle pregnancy rate for mares bred by natural service during the first postpartum estrus, and a 73% pregnancy rate for mares bred by natural service on subsequent cycles. Using stored (chilled) semen for artificial insemination, he found a 50% per cycle pregnancy rate when breeding on the first postpartum estrus in contrast to a 65% per cycle pregnancy rate from subsequent cycles. Obviously, further studies directly comparing different methods of breeding are warranted to determine their effects on fertility achieved in mares bred on foal heat. However, these recent reports emphasize pregnancy rates can be high enough to warrant breeding on the first postpartum estrus.
Influence of the day of postpartum ovulation on fertility Loy3 reported higher pregnancy rates in mares bred on the first postpartum estrus if ovulation occurred after day 10. The timing of several events occurring during the postovulatory period supports this concept. If ovulation and fertilization occur on day 10 postpartum, the embryo will move through
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Table 236.1 Pregnancy rates and losses in Quarterhorse-type mares maintained on pasture and bred on foal heat in southeast Texas by day of postpartum ovulation (DPPOV) DPPOV 7–10 11–15 16–20 >20
n 63 220 76 40
Pregnancy rate
Pregnancy loss rate
67% (42/63) 72% (158/220) 69% (50/72) 80% (32/40)
14% (6/42) 9% (14/158) 6% (3/50) 9% (3/32)
48% of all losses occurred by 42 days, 19% of all losses occurred between 42 days and fall pregnancy examination, and 33% of all losses occurred between fall pregnancy examination and term. Note: While pregnancy rates and pregnancy loss rates did not differ between foaling mares bred for the first time on the first or later postpartum estrus periods, the pregnancy loss rate appears greater for mares bred on foal heat that ovulated in the first 10 days postpartum. Source: Blanchard et al. 20049
the oviduct and into the uterus 5–6 days after fertilization,34, 35 and arrive in the uterine lumen by day 15 or 16 at a time when histological uterine involution is complete15 ) and the uterus is well prepared to accept an embryo. By contrast, mares ovulating prior to day 10 postpartum will experience embryo entry into the uterus at a time when the endometrium may not be histologically normal, and thus may not be prepared to nurture the embryo. Thus, mares ovulating after 10 days postpartum are purported to have better pregnancy rates. Whether this phenomenon holds true for all mares on all farms remains in question. Loy3 also reported foal heat pregnancy rates were farm dependent (i.e., management-related in that some farms had high foal heat pregnancy rates while others had low foal heat pregnancy rates). On farms experiencing low foal heat pregnancy rates, it is possible that only breeding mares that ovulate after day 10 postpartum may still not improve fertility. By contrast, on farms where foal heat breedings result in high pregnancy rates, there may not be a significant advantage to restricting first postpartum estrus breedings only to those mares that ovulate after day 10 postpartum. For example, Texas workers9 recently reported no difference in foal heat pregnancy rates between Quarterhorse-type mares ovulating either ≤ 10 or > 10 days postpartum (see Table 236.1). It may be important to note that the mares in that study were relatively young and maintained on pasture, allowing plenty of exercise to aid in uterine evacuation. Age was found to significantly affect pregnancy rate achieved on foal heat breeding in that study (i.e., for each year increase in age, the odds ratio for establishing pregnancy on foal heat mating was 0.937). Because of Loy’s (1980)3 findings that mares ovulating after 10 days postpartum have higher foal heat pregnancy rates, researchers have attempted to delay the first postpartum ovulation with exogenous steroid regimens in an effort to improve pregnancy
rates in mares bred on the first postpartum estrus. Loy et al.36 administered progesterone, intramuscularly, to postpartum mares in an effort to delay the first ovulation after foaling. Six mares were treated with 100 mg progesterone for 10 days after foaling. Four mares ovulated on days 15 to 17 postpartum and two mares ovulated on days 23 and 25 postpartum. Five of the six mares were bred and became pregnant. In a similar study, McKinnon et al.22 treated postpartum mares with altrenogest (0.044 mg/kg, orally) to delay the first postpartum ovulation. Mares received: (i) daily treatment with altrenogest for 8 days, beginning the day after parturition, and were treated once with prostaglandin on day 9; (ii) daily treatment with altrenogest for 15 days, beginning on the day after parturition; or (iii) no treatment (controls). Ovulation was delayed in treated mares and pregnancy rates were higher in mares that ovulated after day 15 (23/25, 82%) postpartum compared to mares that ovulated before day 15 (6/12, 50%) postpartum. Mares treated with altrenogest for 8 days, with prostaglandin administration on day 9, had higher pregnancy rates (11/12, 92%) than control mares (9/15, 60%) or mares treated with altrenogest for 15 days without prostaglandin treatment (9/13, 69%). Mares with delayed uterine involution, regardless of cause, might be poor candidates for progestogen therapy to delay the first postpartum ovulation as it might predispose them to uterine infection. The effects of exogenous estradiol-17β administration alone on duration of postpartum estrus and interval to ovulation were examined by Texas workers.19 Treatment using a timed-release (12 to 15 days) estradiol-17β microsphere preparation did not alter intervals to the first postpartum estrus or ovulation, and did not alter foal heat pregnancy rates. A combination of progesterone and estradiol has been used successfully to delay the first postpartum ovulation. Follicular development and ovulation are suppressed through the negative feedback effects of the steroid combination. Loy et al.37 injected mares with 150 mg progesterone and 10 mg estradiol-17β, i.m., beginning 12 hours after foaling, once daily for 6 days. Ovulation was delayed approximately 5 days using this therapy, but pregnancy rates were not improved in treated mares. Bristol et al.38 also used this regimen to synchronize the first postpartum estrus in foaling mares in a PMU herd of mares used to collect urine. Mares were treated from 1–10 days with 150 mg progesterone and 10 mg estradiol17β. Mares treated for 1–5 days were in estrus 9.3 days (range 7–14 days) post-treatment and ovulated 13.7 days (range 11–16 days) post-treatment. Mares treated for 6–10 days postpartum were in estrus 9.4 days (range 8–13 days) post-treatment and ovulated 12.6 days (range 10–16 days) post-treatment. Mares
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Breeding Mares on Foal Heat were turned into pastures with stallions 7 days after the end of treatment, which resulted in an overall first postpartum estrus pregnancy rate of 81%. Because steroid treatments administered for several days may not result in significant time savings in interval from foaling to conception compared to non-steroid treated mares, Texas workers investigated shorter steroid treatment regimes. Bruemmer et al.30 compared the effects of administering 150 mg progesterone plus 10 mg estradiol-17β only twice (at 1 and 24 hours after foaling), or a single injection of 300 mg progesterone plus 20 mg estradiol17β given within 12 hours after foaling, to salinetreated (control) first postpartum estrus mares. No differences in interval to first postpartum ovulation occurred between steroid-treated and control mares. However, no progesterone plus estradiol-17β-treated mares (0/20) ovulated prior to day 10 postpartum, whereas six of 30 control mares did. A limitation of this study was that it was performed on mares foaling early in the year, when few of them ovulate prior to day 10 postpartum (i.e., due to seasonal effects on interval to first postpartum estrus and ovulation). The Texas workers proposed one or two days of steroid treatment in mares foaling later in the year would be more likely to be advantageous for foal heat breeding, as late foaling mares are more prone to ovulate prior to day 10 postpartum.
Strategies for breeding mares on the first postpartum estrus The authors recognize that no one protocol will work for all mares when breeding during the postpartum period. However, the following approach is recommended as one that is justifiable, based on available published data. First, all mares should be examined in the postpartum period, beginning no later than 6–8 days after foaling. A visual examination of the caudal reproductive tract will reveal the presence of urine pooling, pneumovagina, or unresolved trauma to the vagina, vestibule or vulva. Although these conditions usually will improve over time, injured mares are not good candidates for breeding on the first postpartum estrus. Ultrasonographic examination of the reproductive tract should be performed to reveal the presence of intrauterine fluid accumulation and size of any developing follicle(s). Depth and character of uterine fluid should be monitored by repeating examinations at 1- to 2-day intervals. If intrauterine fluid accumulation remains at the time when foal heat breeding is anticipated, it is better to treat the mare to remove the fluid than to breed the mare. Mares judged to be involuting normally, with no significant intrauterine fluid accumulation, are good candidates
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for foal heat breeding. If the mare ovulates prior to day 10 postpartum, breeding on foal heat can be bypassed and prostaglandin can be administered 5–6 days after ovulation to induce an earlier return to estrus for breeding. It is thought that this protocol allows the uterine environment more time for involution, as well as providing a shorter interval (1–2 weeks) to breeding than that achieved by waiting for the second postpartum estrus to spontaneously occur (typically around day 30). The exception to this first postpartum estrus breeding strategy would be that older multiparous mares are probably not good candidates for breeding on foal heat. In fact Ginther2 states that it is not known whether depressed pregnancy and foaling rates in mares mated on foal heat are caused by age or multiparity. Lowis and Hyland (1991)39 compared pregnancy rates in mares bred on foal heat to that of mares bred on a prostaglandin-induced second postpartum estrus. Mares bred on the prostaglandin-induced estrus had more favorable rates in all parameters (first cycle pregnancy rate of 55% vs 48%; seasonal pregnancy rate of 90% vs 83%; cycles per pregnancy of 1.4 vs 1.7; pregnancy loss 4% and 9% in prostaglandininduced second postpartum estrus mares vs foal heat mares) except for a slight delay in foaling to conception interval (9.4 day advantage for mares bred on foal heat). Despite attempting to avoid breeding of first postpartum estrus mares with intrauterine fluid accumulations, some mares will accumulate fluid after the foal heat breeding. Such mares should be treated to remove any fluid accumulation and resolve any infection that may have been established. Performing uterine lavage (3 L lactated Ringer’s or saline solution) in these mares is important, and is thought not to interfere with fertility if the lavage is performed at least 4 hours after breeding.40 Brazilian workers41 did find that uterine lavage to remove fluid accumulation after breeding of first postpartum estrus mares improved 42-day pregnancy rates compared to non-lavaged mares that had accumulated fluid after foal heat breeding. Administration of uterine ecbolics (e.g., oxytocin) as the sole treatment, or in conjunction with uterine lavage, has also been shown to have value as post-breeding therapy to improve pregnancy rates.42–44 While the inclusion of antibiotic infusions after breeding of postpartum mares remains controversial, Pycock45 reported that postpartum mares treated with antibiotics plus oxytocin had higher pregnancy rates than either mares treated only with oxytocin or mares that were left untreated. Pascoe46 also reported that an improved foal heat pregnancy rate was achieved in mares infused with antibiotics after breeding compared to mares that were not infused with antibiotics. If antibiotic infusions are
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chosen as a complement to the therapy administered to mares that accumulate fluid after foal heat breeding, the authors choose to use antimicrobials known to be effective against the most common bacteria that are typically recovered from the uterus of postpartum mares (i.e., beta-hemolytic Streptococcus and coliforms such as Escherichia coli and Enterobacter spp). In conclusion, not all mares are suitable candidates for breeding during the first postpartum estrus. However, using careful selection of mares, breeding during foal heat can result in favorable pregnancy rates in a highly efficient manner, and can reduce parturition to conception interval to help to maintain yearly foal production in mares.
References 1. Rossdale PD, Ricketts SW. 1980. Equine Stud Farm Medicine, 2nd edn. Philadelphia: Lea & Febiger, 1980; pp. 260–76. 2. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992; pp. 504–6. 3. Loy RG. Characteristics of post partum reproduction in the mare. Vet Clin N Amer: Large Anim Prac 1980;2:345–59. 4. Hodge SL, Kreider JL, Potter GD, Harms PG, Fleeger JL. Influence of photoperiod on the pregnant and post partum mare. Am J Vet Res 1982;10:1752–55. 5. Palmer E, Driancourt MA. Some interactions of season of foaling, photoperiod and ovarian activity in the equine. Livest Prod Sci 1983;110:197–210. 6. Nagy P, Huszenicza G, Juhasz J, Kulcsar M, Solti L, Reiczigel J, Abavary K. Factors influencing ovarian activity and sexual behavior of post partum mares under farm conditions. Theriogenology 1998;50:1109–19. 7. Lieux P. Comparative results of breeding on first and second post-foaling heat periods. Proceedings of the 26th Annual Meeting of the Association of Equine Practitioners, 1980; pp. 129–32. 8. Lenz TR. One practitioner’s approach to foal heat breeding. Proceedings of the Society for Theriogenology, 1986; pp. 111–19. 9. Blanchard TL, Thompson JA, Brinsko SP, Stich KL, Wendt KM, Varner DD, Rigby SL. Mating mares on foal heat: a five-year retrospective study. Proceedings of the American Association of Equine Practitioners 2004;50:525–30. 10. Koskinen E, Katila T. Uterine involution, ovarian activity and fertility in the post-partum mare. J Reprod Fertil 1987;Suppl 35:733–4. 11. Blanchard TL, Love CL, Thompson JA, Ramsey J, O’Meara A. Role of reinforcement breeding in a natural service mating program. Proceedings of the American Association of Equine Practitioners 2006;52:384–6. 12. Meyers PJ, Bonnett BN, McKee SL. Quantitating the occurrence of early embryonic mortality on three equine breeding farms. Can Vet J 1991;32:665–72. 13. Steven DH. Placentation in the mare. J Reprod Fertil 1982;Suppl 31:41–55. 14. Blanchard TL, Elmore RG, Kinden DA, Berg JN, Mollett TA, Garcia MC. Effect of intrauterine infusion of Escherichia coli endotoxin in postpartum pony mares. Am J Vet Res 2985;46(5):2157–62.
15. Gygax AP, Ganjam VK, Kenney RM. Clinical, microbiological and histological changes associated with uterine involution in the mare. J Reprod Fert 1979;Suppl 27: 571–8. 16. Bailey JV, Bristol FM. Uterine involution in the mare after induced parturition. Am J Vet Res 1983;44(5):793–7. 17. Blanchard TL, Varner DD, Brinsko SP, Meyers SA, Johnson L. Effects of postparturient uterine lavage on involution in the mare. Theriogenology 1989;32(4):527–35. 18. Blanchard TL, Varner DD, Brinsko SP, Quirk K, Rugila JN, Boehnke L. Effects of ecbolic agents on measurements of uterine involution in the mare. Theriogenology 1991;36(4):559–71. 19. Arrott C, Macpherson M, Blanchard T, Varner D, Thompson J, Simpson B, Bruemmer J, Vogelsang S, Fernandez M, Fleet T, Burns P. Biodegradable estradiol microspheres do not affect uterine involution or characteristics of post partum estrus in mares. Theriogenology 1994;42:371–84. 20. Haluska GJ, Lowe JE, Currie WB. Electromyographic properties of the myometrium correlated with the endocrinology of the pre-partum and post-partum periods and parturition in pony mares. J Reprod Fertil 1987;Suppl 35:553–64. 21. Griffin PG, Ginther OJ. Uterine morphology and function in post partum mares. J Eq Vet Sci 1991;11(6): 330–9. 22. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproductive tract of mares after parturition: effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed post partum ovulatory cycles. J Am Vet Med Assoc 1988;192(3) 350–3. 23. Sertich PL, Watson ED Plasma concentrations of 13,14dihydro-15-ketoprostaglandin F2 alpha in mares during uterine involution. J Am Vet Med Assoc 1992;210(3):434–7. 24. Hooper RN, Blanchard TL, Taylor TS, Schumacher J, Varner DD. Identifying and treating uterine prolapse and invagination of the uterine horn. Vet Med 1993;88:60–5. 25. Vandeplassche M, Bouters R, Spincemaille J, Bonte P, Coryn M. Observations on involution and puerperal endometritis in the mare. Irish Vet J 1983;37:126–32. 26. Pycock JF, Newcombe JR. The relationship between intraluminal uterine fluid, endometritis, and pregnancy rate in the mare. Eq Pract 1996;18(6):19–22. 27. Shideler RK, McChesney AE, Squires EL, Osborne M. Effect of uterine lavage on clinical and laboratory parameters in post partum mares. Eq Prac 1987;7(9):20–6. 28. Ley WB, Purswell BJ, Bowen JM. Effect of PGF2-alphaanalogue administered during the post partum period upon pregnancy rate. J Eq Vet Sci 1988;8(2):141–3. 29. McCue PM, Hughes JP. The effect of post partum lavage on foal heat pregnancy rate. Theriogenology 1990;33(5):1121–9. 30. Bruemmer JE, Brady HA, Blanchard TL. Uterine involution, day and variance of first post partum ovulation in mares treated with progesterone and estradiol-17β for 1 or 2 days post partum. Theriogenology 2002;57:989–95. 31. Steiger K, Kersten F, Aupperle H, Schoon D, Schoon HA. Puerperal involution in the mare – morphological studies in correlation with the course of birth. Theriogenology 2002;58:783–6. 32. Belz JP, Glatzel PS. Fertility in mares after a disturbed as well as an undisturbed puerperium. Significance of histological and cytological examinations of the uterus. Tierarztl Prax 1995;23(3):267–72. 33. Camillo F, Marmorini P, Romagnoli S, Vannonzzi I, Bagliacca M. Fertility at the first post partum estrous compared with fertility at the following estrous cycles in
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35.
36.
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foaling mares and with fertility of nonfoaling mares. J Eq Vet Sci 1997;17(11):612–16. Oguri N, Tsutusumi Y. Non-surgical recovery of equine eggs, and an attempt at non-surgical egg transfer in horses. J Reprod Fertil 1972;31:187–95. Hinrichs K, Riera FR. Effect of administration of prostaglandin F2 α on embryo recovery from the uterus on day 5 after ovulation in mares. Am J Vet Res 1990;51:451–3. Loy RG, Hughes JP, Richards WPC, Swan SM. Effects of progesterone on reproductive function in mares after parturition. J Reprod Fertil 1975;Suppl 23:291–5. Loy RG, Evans MJ, Pemstein R, Taylor TB. Effects of injected ovarian steroid on reproductive patterns and performance in post-partum mares. J Reprod Fertil 1982;Suppl 32:199–204. Bristol F, Jacobs KA, Pawyshyn V. Synchronization of estrus in post-partum mares with progesterone and estradiol17β. Theriogenology 1983;19:779–85. Lowis TC, Hyland JH. Analysis of post-partum fertility in mares on a thoroughbred farm in southern Victoria. Aust Vet J 1991;68(9):304–6. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post insemination on pregnancy rates in mares. Theriogenology 1991;35:1111–19.
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41. Malschitzky E, Schilela A, Mattos ALG, Garbade P, Gregory RM, Mattos RC. Effect of intra-uterine fluid accumulation during and after foal-heat and of different management techniques on the post partum fertility of thoroughbred mares. Theriogenology 2002;58: 495–8. 42. Pycock JF. A new approach to treatment of endometritis. Eq Vet Ed 1994;6:36–8. 43. LeBlanc MM, Neuwirth L, Mauragis D, Klapstein E, Tran T. Oxytocin enhances clearance of radiocolloid form the uterine lumen of reproductively normal mares and mares with recurrent endometritis. Eq Vet J 1994;29: 279–82. 44. Rasch K, Schoon HA, Sieme H, Klug E. Histomorphological endometrial status and influence of oxytocin on the uterine drainage and pregnancy rate in mares. Eq Vet J 1996;28:455–60. 45. Pycock J. Assessment of oxytocin and intrauterine antibiotics on intrauterine fluid and pregnancy rates in mares. Proceedings of the American Association of Equine Practitioners 1994;40:19–20. 46. Pascoe D.R. Effect of adding autologous plasma to an intrauterine antibiotic therapy after breeding on pregnancy rates in mares. Biol Reprod 1995;Mono 1:539–43.
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Chapter 237
Inter and Extraspecies Equine Pregnancies W.R. (Twink) Allen, Julia H. Kydd, Roger V. Short and Douglas F. Antczak
The genus Equus is unusual, but not unique, within the mammalian class for the ability of its phenotypically and karyotypically diverse member species to interbreed freely and produce live, though usually infertile, offspring. Species range from the ancient horse of Mongolia (Equus przewalskii), with the highest diploid chromosome number of 66, through the many breeds and types of domestic horse (E. caballus, 2n = 64), to the European ass or domestic donkey (E. asinus, 2n = 62), to the many African and Asiatic species of wild ass (e.g., E. hemionus, 2n = 54), to the Gr´evy’s zebra of Somalia and northern Kenya (E. grevyi, 2n = 46), the many subspecies of common zebra in Central and East Africa (E. burchelli, 2n = 44), and the now rare mountain zebra of southwest Africa (E. zebra, 2n = 32). Cataloged evidence concerning the range of hybrid foals born during the past 100 years makes it safe to assume that an adult cycling female from any of the species inseminated with fertile semen from a male of another species will conceive and give birth to a viable hybrid foal.1–4 The mule (2n = 63), the result of the mating of a horse mare (E. caballus) to a domestic jack donkey (E. asinus), and the reciprocal cross, the hinny (2n = 63), are by far the most common equine hybrids, simply because their parents are the only two species of equids to have been domesticated in significant numbers. The mule was the first and only commercially viable interspecific hybrid to be bred by humans, and many millions of them have been produced since long before the time of Christ to the present day, for their perfect blend of the physical and mental attributes of the two parental species. But, in addition to its physical prowess, the mule has intrigued and puzzled biologists and geneticists since the time of Aristotle. As described so eloquently by Short,4 ‘the clear and equal mixture in the mule of the phenotypic
characteristics of both parents ran counter to Aristotle’s ‘seed and soil’ view of reproduction in which the male provided the essential “seed” for the new offspring while the female played an entirely passive role by supplying merely the “soil” in which the male seed would grow.’ Much later, as Short quoted, ‘the Danish anatomist Stensen (1638–1686) and the Dutch biologist De Graaf (1641–1673) used the mule as an example in their proposals that the females of all mammalian species also contribute “seed” toward the generation of new individuals. Stensen noted the presence of follicles in the ovaries of one of two female mules he was able to dissect, just as he had seen these fluid-filled “eggs” in the ovaries of women, cattle, sheep, hares, and many other species.5 And, as quoted by Jocelyn and Setchell,6 De Graaf noted: ‘if a mare receives the semen of an ass, the fetus has the features not of the father alone, but those of both parents mixed together. . . . Mothers, therefore, as well as fathers, have child-producing semen.’ In more recent times the infertility of mules and hinnies has similarly aroused the interest of geneticists. Wodsedalek7 was first to conclude that spermatozoa are not produced in the testes of male mules because of an incompatibility between the paternal and maternal sets of chromosomes leading to a block in meiosis. More than 50 years later, Taylor and Short8 demonstrated that the same chromosomal incompatibility leads to partial meiotic arrest in female mules and hinnies, which results in a severely depleted stock of oocytes at birth in these hybrids. Thus equine hybrids, especially the mule, have proved to be of scientific interest as well as commercial value for many centuries past. But perhaps of equal and more modern scientific interest are the immunological and developmental questions posed by the discovery that at least the horse, donkey, and mule within the equid family will accept, carry to
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Inter and Extraspecies Equine Pregnancies term, give birth to, and rear successfully true xenogeneic extraspecific foals following between-species embryo transfer.9 In this chapter we attempt to review some of the more puzzling immunological and endocrinological aspects of the establishment of pregnancy and the survival of the xenogeneic fetus in mares and other equids when they are carrying hybrid interspecific and transferred extraspecific foals.
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Table 237.1 Conception rates achieved in 51 jenny donkeys mated to horse stallions or inseminated with fresh stallion semen in repeated estrous cycles
Interspecies pregnancies Totals
Number of donkeys mated/ inseminated
Number of hinny embryos recovered at days 3–8 post-ovulation or number of confirmed hinny pregnancies
Conception rate (%)
125 34 159
18 5 23
14.4 14.7 14.5
The mule and hinny There seems little doubt that the interspecies mating of the jack donkey and the horse mare is just as fertile, in terms of conception rates, as the straightforward intraspecies mating between either of the parental species. Many millions of mules have been produced over the centuries in the Asian and Indian subcontinents, in southern Europe and north Africa, and, in more recent times, in North and South America. These pregnancies have been established both by natural mating and by artificial insemination, and although strict comparisons of conception rates achieved per insemination or per estrous cycle have not been reported in the scientific literature, it is a situation for which common knowledge and practical horsemanship dictates that the interspecies mating for mule production is of normal fertility. This high fertility of the mating to produce the mule is certainly not matched when attempting to produce the reciprocal hybrid, the hinny, by mating jenny donkeys to horse stallions.10, 11 Once again, the situation does not appear to have been researched thoroughly or recorded in any detail in the literature. However, few confirmed hinnies exist in countries like Spain, Portugal, Greece, and India, where enormous numbers of mules are still produced and used routinely by smallholders and peasant farmers. Enquiry of such farmers always yields the same response, namely that the horse stallion is much less aroused by the estrous display of the jenny donkey than vice versa and, hence, he is less than keen to mate with her. But even if mating is achieved, only rarely does conception occur. This relative infertility of the hinny mating is also the experience of the authors. During a 7-year period in two laboratories (Cambridge, UK, and Cornell University, USA), a total of 159 attempts were made in successive breeding seasons to establish hinny pregnancies in 51 jenny donkeys. These jennies appeared to be of normal fertility and were cycling normally and ovulating regularly. A total of six pony and horse stallions, all of proven high fertility, were
used with a mixture of natural mating and artificial insemination on alternate days during estrus. In 125 instances, the mated/inseminated jenny donkey was left untouched in the hope of establishing a normal ongoing hinny pregnancy; this occurred in 18 cases to give a pregnancy rate of only 14.4%. In the other 34 instances, the uterus of the donkey was flushed non-surgically on day 7 or 8 after ovulation (n = 29) or the ipsilateral oviduct was flushed on day 2 or 3 (n = 5) as described previously12 in an attempt to recover a hinny embryo for transfer to another recipient animal; this was successful in five cases to give an embryo recovery rate of only 14.7% – similar to the pregnancy rate and much lower than the recovery rates of 60–70% obtained when flushing mares or jenny donkeys following normal intraspecific matings or inseminations12 (Table 237.1). This striking disparity in fertilization and/or pregnancy rates when carrying out reciprocal hybrid matings has parallels in other species. In sheep × goat crosses, for example, Hancock and colleagues reported high fertilization rates when inseminating female goats with ram semen, but many fewer conceptions (± 10%) when reciprocally inseminating ewes with goat semen.13–15 Similarly, hybrid matings between the hare (Lepus americanus) and the rabbit (Oryctolagus cuniculus) produce high rates of fertilization when rabbits are inseminated with hare semen whereas < 10% of hare oocytes are fertilized by rabbit semen.16 The mechanisms responsible for these large differences in fertilization rates in interspecies matings have yet to be determined, although the phenomenon of gene imprinting could possibly play a role.17 Other equine hybrids As mentioned previously, viable hybrid offspring may be created by crossing of any two of the considerable array of species of equids. Hybrids that have been produced have been well cataloged,3 and they
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Mare: Pregnancy, Parturition and the Puerperal Period duced conception rate when attempting to produce hinnies compared with mules. To speculate that fertilization may occur readily in any cross where the sire has a lower diploid chromosome complement than the dam is tempting – e.g. a domestic horse mare (E. caballus, 2n = 64) mated to a domestic jack donkey (E. asinus, 2n = 62) or to a male Grant’s zebra (E. burchelli, 2n = 44). Conversely, conception rate is greatly reduced in any combination in which the sire has a higher diploid number of chromosomes than the dam (e.g. E. caballus ♂ × E. asinus ♀ or E. asinus ♂ × E. burchelli ♀).
Figure 237.1 Fl -interspecies hybrid ‘zebronkey’, which resulted from the mating of a male Burchell’s zebra (Equus burchelli b¨ohmii, 2n = 44) to a European jenny donkey (E. asinus, 2n = 62). From King.64
include such unlikely combinations as the progeny of the domestic donkey (E. asinus) mated to: (i) the Somali, African, and Asiatic wild asses; (ii) Przewalski’s and domestic horses; and (iii) Burchell’s, Gr´evy’s, and mountain zebras (Figure 237.1). Similarly, a wide range of hybrids has been produced by mating domestic horses (E. caballus) to all the main zebra species, including Burchell’s, Chapman’s, Grant’s, Hartmann’s, and Gr´evy’s (Figure 237.2), and to a wide range of the wild asses, including Abyssinian, Asiatic, Tibetan, Indian, and Persian subspecies. In all these cases of successful birth, the female of the partnership was the horse, in a manner similar to the production of mules. In any instances in which reciprocal crosses were attempted by mating a male domestic horse to a female wild ass or zebra, conceptions failed to occur.3 This seems to mimic the previously described situation of a greatly re-
Figure 237.2 Fl -interspecies hybrid ‘zebrorse’ that was produced by the mating of a male Grevy’s zebra (Equus grevyi, 2n = 46) to a domestic horse mare (E. caballus, 2n = 64). From King.64
Fertility of equine hybrids The infertility of mules has perplexed and fascinated horse breeders and scientists through the ages. Wodsedalek7 was first to provide the real explanation of the situation when, after studying the testes of three male mules histologically, he concluded that spermatozoa could not be produced because of the incompatibility between the paternal (donkey) and maternal (horse) sets of chromosomes resulting in a block in meiosis. This breakdown of spermatogenesis at the pachytene stage of meiotic prophase was confirmed in male mules and hinnies,18 and the same sort of chromosomal incompatibility, causing partial meiotic arrest in oogenesis during fetal life, was described in female mules and hinnies.8 A number of isolated and unverified reports of individual fertile female mules were made during the early part of the twentieth century, the most notable of which was an animal named ‘Old Bec’ that was owned by Texas A&M College in the 1920s. She had all the morphological characteristics of a mule and was reported to have given birth to two foals in her lifetime. One of these, sired by a jack donkey, was a female, was typically mule-like in appearance, and was infertile. The other, a colt sired by a horse, was completely horse-like in appearance and was apparently fully fertile; he sired many male and female foals that were all horse-like in appearance and were fertile.19 However, some years later another allegedly fertile female mule was karyotyped and it was discovered to be a normal donkey.20 This revelation, and the finding of few spermatozoa in the centrifuged ejaculates of male mules and hinnies,18 led Short to declare firmly that all mules and hinnies, without exception, should be considered infertile unless conclusive proof to the contrary was provided.5 Curiously, such proof was soon forthcoming and, during the next two decades, no fewer than five female mules, one in China,21 two in North America,22, 23 one in Brazil,24 and one in Morocco25 have all been karyotyped as mules and shown conclusively to produce one or more foals.
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Figure 237.3 F2 -interspecies foals produced by mating a male Przewalski’s horse × domestic horse Fl hybrid to two domestic horse mares. (a) The F2 -filly foal had a karyotype of 2n = 65 and was similar to Przewalski’s horse in appearance, whereas the F2 -colt foal (b) had a karyotype of 2n = 64 and, apart from a prominent dorsal stripe, was indistinguishable from a normal horse foal.
Similarly, a female hinny in China has been karyotyped correctly as a hybrid and has produced a donkey-like foal after having been mated to a jack donkey.26 Chandley27 discussed at some length the mechanisms that might function to enable these odd female mules and hinnies to be fertile. She favored the earlier ‘affinity’ hypothesis of Michie,28 whereby chromosomally balanced haploid gametes might occasionally be produced by the movement of centromeres of similar ancestry to opposite poles at anaphase I of meiosis. This could leave the whole set of female parental chromosomes remaining in the oocyte with subsequent elimination to the polar body of the paternal set. Elimination of the donkey paternal set of chromosomes from the mule oocyte would leave only the maternal horse set remaining so that another mule would be produced if the animal was mated to a jack donkey, whereas a pure horse would result if the mating were to a horse stallion. This is exactly what appears to have happened in the case of ‘Old Bec’s’ mule and horse foals in the 1920s, the mule foal born from the mule in Nebraska in 1985,19 and the three progeny produced by the fertile female mule in Brazil.24 However, when the offspring of the fertile mule and the hinny in China were investigated by Chandley and her colleagues,21, 26, 29 they showed a mixture of horse and donkey chromosomes and, therefore, demonstrated inheritance of both paternal and maternal chromosomes in the oocyte that was fertilized. The more recent application of cross-species chromosome painting to equids has suggested how such departures from the ‘affinity hypothesis’ might occur, and how certain combinations of meiotic pairings of horse and donkey chromosomes can lead to the production of true backcross animals.30, 31 With the
wider appreciation of the scientific value of these rare cases of fertility in mules, it is likely that increasing numbers of such equids will be characterized in the future. Not all the possible equid hybrids seem to be as infertile as the mule and hinny. Both the male and female offspring (2n = 65) of the cross between Przewalski’s horse (E. przewalskii 2n = 66) and the domestic horse (E. caballus, 2n = 64) are fully fertile32–34 (Figure 237.3). Furthermore, Gray3 lists a number of examples where hybrids between the domestic donkey (E. asinus, 2n = 62) and both Burchell’s zebra (E. burchelli, 2n = 44) and Chapman’s zebra (E. burchelli antiquorum, 2n = 44) were reported to be fertile, although none of these cases was verified. However, examples of hybrids between the domestic donkey and other members of the wild ass family, including the Persian wild ass (E. hemionus onager) and the Somali wild ass (E. asinus africanus) have been reported as being sterile, as have hybrids between the various species of zebra and the domestic horse.
Extraspecies pregnancies The use of between-species embryo transfer to establish xenogeneic equine pregnancies has highlighted some unexpected and intriguing interacting influences of fetal genotype and maternal uterine environment on placental development and implantation. Horse embryos were first successfully transferred to donkey recipients and, vice versa, donkey embryos were transferred to horses, in the authors’ laboratories in 1979, and further transfers of more exotic equine embryos to domestic horses or donkeys were undertaken in collaboration with other laboratories after that. In all instances, the techniques used
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for synchronizing and diagnosing ovulation in donor and recipient animals, and the methods employed for the non-surgical recovery of embryos from donors and their surgical transfer to recipients, were similar to those described previously.12 Embryo recovery and transfer The great majority of embryos were recovered on days 7 or 8 after ovulation and were, therefore, at the early blastocyst or expanded blastocyst stages when transferred. Similarly, most of the recipients had ovulated synchronously with, or 1 to 2 days after, the donor and only on a few occasions were recipients used that had ovulated 1 day before, or 3 days after, the donor. Pregnancy in the recipients was diagnosed by a combination of palpation and real-time ultrasound scanning of the uterus per rectum,35 commencing 8 to 10 days after transfer, with repeated examinations at intervals of 10 to 14 days until around day 100 of gestation. Jugular vein blood samples were recovered three times a week throughout gestation from all pregnant recipients to monitor routinely profiles of equine chorionic gonadotropin (eCG),36 progesterone,37 and occasionally the onset, specificity, and titer of maternal antibodies to paternal lymphocyte alloantigens.38 Most pregnancies were left untouched to proceed to term or to abort spontaneously, whichever was destined to be the outcome. In other pregnancies, especially in horse mares carrying donkey fetuses and, vice versa, in jenny donkeys carrying horse fetuses, the conceptus was electively removed from the uterus via midline hysterotomy under general anesthesia for gross and histological examinations of many parameters, including fetal growth, endometrial cup development and/or degeneration, and the rate of growth, degree of interdigitation, and general histological appearance of the non-invasive allantochorion–endometrium interface.9 Horse-in-donkey pregnancy Six extraspecies horse-in-donkey pregnancies were created over a 3-year period. These arose from nine transfers (67%) of single horse embryos recovered on days 7 or 8 after ovulation; the nine embryos were obtained from 13 recovery attempts (77%). On one occasion the horse embryo was transferred to a jenny donkey that had been mated to a jack donkey in the previous estrus. This animal successfully implanted the two conceptuses – her own intraspecific donkey embryo conceived normally and the extraspecific horse conceptus transferred to her subsequently. Of these six horse-in-donkey pregnancies, two were allowed to continue uninterrupted to term and
the jennies gave birth to healthy horse foals on, respectively, days 336 and 363 of gestation.9 Both foals were slightly smaller at birth than expected (Figure 237.4), probably because of the restrictive effects of the physically smaller uteri of the donkey surrogate mothers, compared with the uterine and general body size and ‘roominess’ of the true genetic horse mothers. Nevertheless, both foals were robust and they stood and sucked unaided. Two other singleton horse-in-donkey conceptuses were removed by surgical hysterotomy on days 59 and 62, and the donkey carrying the intraspecific donkey and extraspecific horse twin conceptuses was killed on day 63 for recovery of the gravid uterus. In all three of these recipient donkeys, a horseshoe or semicircle of large endometrial cups was associated with the horse conceptus. These cups were relatively broad and they protruded prominently from the luminal surface of the endometrium, just as in donkeys carrying interspecific hinny conceptuses.10 In the donkey carrying the twins, however, the endometrial cups associated with the intraspecies donkey conceptus were, typically, much narrower and generally smaller than those associated with the horse conceptus11 (Figure 237.5). This striking difference in the size and general development of the endometrial cups between the horse and donkey conceptuses in the same animal reflected the pronounced influence of fetal genotype on the development of the placental progenitor of the cups, the chorionic girdle. In another parallel to hinny pregnancy, the ovaries of the recipient donkeys carrying horse fetuses were greatly enlarged because of the development of multiple secondary corpora lutea and luteinized follicles (Figure 237.5). This, in turn, led to extremely high concentrations of progesterone in maternal peripheral plasma39 (Figure 237.6) and was almost certainly the result of the higher ratio of follicle-stimulating hormone-like to luteinizing hormone-like biological activities in eCG, causing hyperstimulation of the surrogate donkey ovaries (see Figure 237.5), as described previously in donkeys carrying interspecies hinny conceptuses.40 Histological examination of the placental interface in these three donkeys carrying extraspecific horse conceptuses revealed that implantation and placentation were proceeding normally. The horse allantochorion was well interdigitated with the donkey endometrium, and a firm microvillous attachment existed between the trophoblast and the endometrial epithelium. No sign of any leukocyte aggregations were evident at this junction. The horse endometrial cups were large and the majority of the binculeate eCG-secreting cup cells appeared viable and active. Large numbers of lymphocytes were clustered around each cup, and in a manner similar to
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Figure 237.4 Donkey and horse extraspecific foals with their horse and donkey surrogate mothers. Note how big and robust the donkey foal is compared with the horse foal, which developed in the smaller uterus of its donkey surrogate mother.
interspecies hinny pregnancy, but in contrast to the situation in interspecies mule pregnancy, the accumulated cells remained in the surrounding endometrial stroma, with little invasion and destruction of the cup tissue itself. Thus, it was apparent that endometrial cups had developed as expected and implantation and placentation were occurring normally in these three recipient donkeys, as clearly it had done in the two that carried their foals to term. However, the remaining
Figure 237.5 Opened uterus of a jenny donkey carrying its own intraspecific donkey fetus (left horn) and a transferred extraspecific horse fetus (right horn) at day 63 of gestation. Note the small, narrow donkey endometrial cups compared to the much larger and more active horse endometrial cups. Note also the hyperstimulated maternal ovaries due to the much higher follicle-stimulating hormone (FSH):luteinizing hormone (LH) ratio of the horse CG (chorionic gonadotropin).
donkey carrying a horse conceptus aborted spontaneously on day 80, following an atypical steep decline in maternal serum eCG concentrations from day 64 onward (Figure 237.7). Embryo transfer in mules To compare the humoral and cell-mediated maternal immune responses of the F1 interspecies mule to antigens expressed by the fetuses of its component parental species, five horse and six donkey embryos were transferred surgically to cycling female mules. Four other horse embryos and a donkey embryo were transferred to non-cycling mules with streak gonads; these animals were given a daily oral dose of 27.5 mg of the synthetic progestagen, altrenogest (Regumate; Hoechst Animal Health, Milton Keynes, UK), beginning 5 days before transfer, to simulate diestrus. The results of this experiment have been described previously.41–43 One donkey and three horse pregnancies became established in the cycling recipient mules, and one horse pregnancy got underway in a progestagentreated anestrous mule (Table 237.2). One donkey and two of the horse foals were carried safely to term and were born healthy and robust between days 344 and 366 (Figure 237.7); all three of the mule surrogate mothers lactated normally and showed strong maternal behavior toward their foals.44 One horse-in-mule conceptus was removed by surgical hysterotomy at day 73 for gross and histological examinations of the
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Figure 237.6 Peripheral plasma progesterone (solid circles) and serum eCG (open circles; hatched area) concentrations measured during early pregnancy in two surrogate jenny donkeys carrying transferred extraspecies horse conceptuses. Note the high plasma progesterone concentrations in both animals. The donkey in the top graph was killed on day 59, whereas the donkey in the lower graph aborted spontaneously on day 80 when serum equine chorionic gonadotropin (eCG) concentrations had fallen rapidly and prematurely to < 5 IU/mL. An endometrial biopsy recovered non-surgically on the day after abortion showed an intense inflammatory reaction of neutrophils and lymphocytes.
endometrial cups and non-invasive placenta. In the progestagen-treated acyclic pregnant mule recipient, the fetus died spontaneously around day 55 and the resorbing conceptus was expelled a few days later. This animal was also subjected to hysterotomy on day 84 to recover persisting endometrial cup tissues for histological evaluation. Both the amounts of eCG secreted and the duration of eCG secretion differed markedly between the two pregnancy genotypes in
these hybrid surrogate mothers.42 For example, peak serum eCG concentrations in the four mules carrying horse fetuses ranged from 38 to 352 IU/mL, and eCG remained detectable until days 92 to 119 of gestation (Figure 237.8). Thus, the profiles were essentially similar to those found in mares carrying normal intraspecies horse pregnancies.45 In the mule carrying the donkey fetus, however, serum eGG concentrations peaked at only 10 IU/mL and the hormone had
Figure 237.7 Surrogate Fl mule mothers with their horse (a) and donkey (b) foals following embryo transfer. Note the vitality and forward development of both foals following their gestation in the uteri of the large recipient mules.
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Table 237.2 Recovery and transfer of horse (Equus caballus) and donkey (E. asinus) embryos to F1 hybrid cycling and anestrous progestagen-treated female mules
Type of transfer
Number of embryos transferred
Number of recipients pregnant (%)
Horse to cycling mule
5
3 (60)
Donkey to cycling mule Horse to anestrous mule Donkey to anestrous mule
3 4 1
1 (33) 1 (25) 0
Outcome of pregnancies 2 foaled (days 344 and 366) 1 removed (day 73) Foaled (day 357) Resorbed (day 55) –
Data from Antczak et al.42 and Shaw and Haupt.44
become undetectable by as early as day 84 of gestation42 (Figure 237.7). Thus, in this animal the secretion pattern was much more like that described in mares carrying interspecies mule fetuses.46 Striking differences existed also in the gross and histological appearances of the endometrial cups recovered from two of the mules carrying horse fetuses. In the animal that underwent hysterotomy at day 73 while still pregnant, the cups were already cheesy and yellowish in appearance as a result of a significant degree of cell degeneration and a copious amount of eCG-rich exocrine secretion accumulated on their luminal surfaces. Histologically, most of the cup tissue was already necrotic and had become mixed with the liberated gland secretions to give the honey-colored pabulum trapped between the luminal surface of the cup and the overlying allantochorion. An extremely dense band of maternal lymphocytes and other leukocytes surrounded the cup, completely separating it from the adjacent maternal tissues (Figure 237.9a). In marked contrast, the endometrial cups recovered on day 84 from the altrenogest-treated mule that had undergone spon-
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Figure 237.8 Serum equine chorionic gonadotropin (eCG) profiles measured between 40 and 140 days of gestation in four pregnant female mules, three of which were carrying a horse conceptus and the other a donkey conceptus following embryo transfer. From Antczak et al.42
taneous fetal death some 30 days previously were pale, convex rather than saucer-shaped in outline, and still clearly viable and active. Histologically, the endometrium of this acyclic animal was immature and prepubertal in general appearance, with only a few endometrial glands that were straight and non-branched. The persisting endometrial cups were composed, typically, of the mass of large, binucleated eCG-secreting endometrial cup cells of fetal origin. However, as a consequence of the loose and immature architecture of the endometrial stroma, the cup cells were much less densely packed together than normal, so the cup as a whole had an edematous appearance (Figure 237.9b). But most striking of all, few lymphocytes had accumulated in the stroma around the cup, thereby giving a picture more like the histological appearance of the endometrial cups in intraspecies donkey pregnancy,10 in complete contrast to the dense aggregation of lymphocytes around the cups in the other mule still carrying a viable horse fetus at day 73. This big difference in the intensity of the leukocyte accumulations around the endometrial cups in the two horse-in-mule pregnancies was almost certainly caused by the absence of the fetus and noninvasive placenta from the second animal. A similar marked decline in the intensity of maternal leukocyte response to the endometrial cups, and consequently a longer persistence of viable cups in the endometrium, has been observed on a number of occasions in mares that have spontaneously aborted their normal, intraspecies horse conceptus soon after development of the endometrial cups at day 40. The equally large differences in eCG production and endometrial cup lifespan between the horse-in-mule and the donkey-in-mule pregnancies further highlighted the profound and interactive influences of fetal genotype and maternal uterine environment on the development and invasiveness of the chorionic girdle and on the expression of, and maternal cellmediated responses to, fetal antigens on these specialized trophoblast cells.
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Figure 237.9 Histological sections of endometrial cups recovered from a female mule carrying a viable transferred horse fetus at day 73 of gestation (a, × 10; b, × 50) and from a non-cycling progestagen-treated female mule that had aborted a transferred horse fetus 30 days previously at day 54 of gestation (c, × 10; d, × 50). Note the pronounced contrast between these two animals in the viability of the fetal cup cells and in the intensity of the maternal leukocyte accumulation around the cups. Only a small patch of lymphocytes was observed around the cups from the mule that aborted its conceptus (arrow in c). Sections were fixed in Bouin’s solution and stained with hematoxylin and eosin.
Mule embryos in donkeys The early studies of one author (Allen)10, 36 showed dramatic differences in the size, productivity, and lifespan of the endometrial cups in mares carrying intraspecies horse or interspecies mule conceptuses versus donkeys carrying intraspecies donkey or reciprocal interspecies hinny conceptuses. These findings appeared to reflect genomic imprinting influencing placental development, as highlighted by the classical studies of Barton et al.17 when comparing fetal and placental development in gynogenetic and androgenetic mouse embryos constructed by micromanipulation and polar body injection; they concluded that the maternal genome contributes preferentially to the makeup of the embryo while the paternal genome contributes likewise to the placenta. However, one experiment involving micromanipulation and between-species embryo transfer appeared to refute this notion.
A mule morula recovered on day 6 after ovulation from the donor mare was bisected using the micromanipulator,47 and one of the two resulting demiembryos was transferred back to a mare while the other was transferred to an unmated recipient donkey. Two other intact mule blastocysts recovered on day 7 were also transferred to unmated donkey recipients. All four recipients became pregnant and were bled thrice weekly from day 30 after ovulation to measure eCG profiles. Their uteri and ovaries were recovered postmortem between days 60 and 63 for gross and microscopic examination of the endometrial cups and secondary luteal development in the ovaries. In the surrogate mare carrying one demi-mule embryo, small, narrow, and prematurely necrotic endometrial cups were present in the uterus on day 63 and she showed a peak serum eCG concentration of only 11 IU/mL on day 56. Thus, the situation was
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whose binding affinity has been shown to be very species-specific, would seem the most likely candidate for this role in equine pregnancy. Embryo transfer in exotic equids Over a 2-year period attempts were made at the Zoological Society of London to recover embryos non-surgically from two Przewalski’s horse mares (Equus przewalskii, 2n = 66) and two Grant’s zebra mares (E. burchelli, 2n = 44) for transfer to domestic pony mares (E. caballus, 2n = 64) and jenny donkeys (E. asinus, 2n = 62).43, 55 Estrus in the donor animals was induced by an intramuscular (i.m.) injection, given between days 6 and 14 of diestrus, of 250 µg of the prostaglandin analog, cloprostenol (Estrumate; Coopers Animal Health, Berkhamstead, UK), which was delivered by a compressed air dart from a blowgun. The treated animals were placed in a pen with the appropriate male for 1 hour each day and observed for signs of estrus and successful mating. Ovulation was assumed to have occurred during the 24-hour period before the cessation of estrous behavior and embryo recovery attempts were carried out between 6 and 10 days later, mostly on days 7 or 8. Briefly, the zebra or Przewalski’s horse donor mare was tranquilized by an i.m. injection of 1.1–2.9 mg etorphinehydrochloride and 4–12 mg acepromazine maleate (0.9–1.2 mL; Large Animal Immobilon; C-Vet Ltd, Suffolk, UK). The sedated animal was then blindfolded and restrained standing, or in sternal recumbency. A flexible 24-Fr gauge Gibbon balloon flushing catheter was passed through the cervix and the uterus was then irrigated three times, each with 700–1000 mL of flushing medium, as described previously. When located, the embryo was transferred to 5 mL of fresh transfer medium in a conical sterile plastic tube. This was placed in a shirt breast pocket and driven 72 miles by car to Cambridge (1.5–2 h transit time) for surgical or non-surgical transfer to the selected, synchronized recipient pony mare or jenny donkey. The embryo recovery and pregnancy rates achieved in these experiments are summarized in Table 237.3. Eleven Przewalski’s horse embryos were obtained from 18 recovery attempts performed on the two donor mares (61%). Of these embryos, nine were transferred surgically and two were transferred non-surgically to recipient pony mares. This resulted in seven pregnancies initially (64%), as diagnosed by ultrasonographic scanning of the uterus between days 15 and 18 after ovulation.35 However, an embryo proper did not appear after day 21 in two of these vesicles, which then diminished steadily in size and finally disappeared during the next 12
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Table 237.3 Recovery and transfer of Przewalski’s horse (Equus przewalskii) and Grant’s zebra (E. burchelli) embryos to domestic horse (E. caballus) and donkey (E. asinus) recipients Type of transfer recipients
Embryo recovery attempts
No. embryos recovered (%)
No. embryos transferred/ no. pregnant (%)
Przewalski’s horse-to-horse
18
11 (61)
11/7 (64)
2 resorbed (days 20–40) 1 resorbed (days 85–101) 1 stillborn at term 3 live foals
Zebra-to-horse
25
14 (56)
5/3 (60)
1 resorbed (days 59–64) 1 stillborn (day 350) 1 live foal
8/2 (25)
1 resorbed (days 53–64) 1 aborted (day 292)
Zebra-to-donkey
Outcome of pregnancies
Data from Kydd et al.43 and Summers et al.55
days. A third conceptus was aborted spontaneously between days 85 and 101. The other four pregnancies proceeded to term and three live and healthy foals were born (Figure 237.10). The fourth mare foaled unobserved in the paddock and her foal was unfortunately suffocated by the amnion. Fourteen zebra embryos were obtained from a total of 25 embryo recovery attempts (56%) performed on the two donor zebras. Of these, five embryos were transferred surgically to recipient pony mares, which resulted in three established pregnancies (60%). However, one fetus died and the conceptus was resorbed between days 59 and 66, and a second foal was stillborn by cesarean section performed on
day 350. This mare had shown increasingly severe polyarthritis during the previous 3 weeks. The condition was considered likely to be the manifestation of some form of immunologically based pregnancy toxemia, because the joint swelling and soreness disappeared completely within a few hours after removing the conceptus. The third foal was born alive and robust at day 367 and it was reared successfully by its surrogate horse mother (Figure 237.11a). Of the original zebra embryos, eight were transferred surgically to synchronized recipient jenny donkeys to yield only two pregnancies (25%). One of these was resorbed between days 53 and 64, whereas the other foal was born prematurely at day 292 and lived for only
Figure 237.10 Surrogate domestic horse mare (E. caballus, 2n = 64) with her newborn Przewalski’s horse foal (2n = 66), resulting from embryo transfer on day 7 after ovulation. From Kydd et al.43
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(b) Figure 237.11 (a) A 3-day-old Grant’s zebra foal (Equus burchelli, 2n = 44) with its surrogate domestic horse mother (E. caballus, 2n = 64). (b) A well-formed Grant’s zebra fetus aborted by its surrogate jenny donkey mother (E. asinus, 2n = 62) on day 292 of gestation.
3 hours before succumbing to respiratory incompetence (Figure 237.11b). This surrogate donkey mother also showed clinical signs of severe polyarthritis for 5 days before the premature birth (Table 237.3). Thus, from the total of seven Przewalski’s horse and five Grant’s zebra pregnancies established initially in the recipient mares and jenny donkeys, only
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three Przewalski’s horse and one zebra foal were born alive. The two Przewalski’s horse conceptuses that were resorbed before day 40 were most likely to have failed as a result of lethal damage to the inner cell mass (ICM) sustained during the recovery, transport, and transfer procedures, which then led to the development of anembryonic trophoblast vesicles. The one Przewalski’s horse and two Grant’s zebra conceptuses that failed between days 53 and 101 may not have implanted properly for reasons that are not understood. One possibility, however, is that the xenogeneic conceptus may have elicited a generalized cell-mediated rejection response from the maternal immune system. The donkey and the horse recipients that aborted well-developed zebra foals near term both showed clinical signs of the severe non-infective polyarthritis described previously before the abortion, and both exhibited high concentrations of antigen-antibody complexes in their peripheral blood at this time.55 Such untoward immunological signs were not exhibited by the other pony mare that gave birth to the live zebra foal, nor by an aged Quarterhorse mare in Kentucky that also carried an extraspecific zebra foal to term after non-surgical transfer.56 In endocrinological terms, the Przewalski’s horsein-horse pregnancies were characterized by serum eGG profiles that were essentially similar to those measured in normal intraspecies horse-in-horse pregnancies.45 Considerable variation existed, however, among the four animals involved, both in the peak concentrations (5 to 135 IU/mL) and in the stage of gestation when eCG disappeared from the blood (55 to 95 days). In contrast, minimal amounts of eCG (0.5 to 1.5 IU/mL) were detected for only a few days of gestation between days 40 and 56 in the three mares carrying zebra conceptuses, whereas the peak concentrations were higher (7–22 IU/mL) and eCG secretion was more prolonged (days 39 to 83) in both the donkeys carrying zebra fetuses.55 These big differences in serum eCG concentrations between the different pregnancy genotypes probably reflected differences in the breadth and overall development of the progenitor chorionic girdle. Similarly, the equally large variation in the duration of eCG secretion no doubt mirrored differences in the surrogate maternal cell-mediated responses to xenogeneic fetal antigens expressed at the fetomaternal interface. Certainly, the most striking examples of both these aspects were seen in the three mares carrying zebra conceptuses. The low concentrations of eCG indicated a greatly reduced development of the zebra chorionic girdle in the horse uterus, compared with the greater degree of development in the zebra uterus, as evidenced by the much higher concentrations of eCG measured in the blood of the two donor Grant’s zebra mares when
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carrying their own interspecies zebra conceptuses at the conclusion of the embryo transfer experiment.55 Similarly, the brief period when eCG was present in maternal blood indicated an even more rapid cellmediated destruction of the small zebra-in-horse endometrial cups that occurred in mares carrying interspecies mule conceptuses.36 The donkey-in-horse model of pregnancy failure The one example of equine extraspecific pregnancy that seems to differ markedly from the other types examined to date is that created when a donkey embryo (E. asinus, 2n = 62) is transferred to the uterus of a horse recipient (E. caballus, 2n = 64). In this situation, the donkey chorionic girdle develops as an extremely narrow and scanty band of trophoblast tissue compared with its counterpart in intraspecies horse pregnancy, and it fails completely to invade the surrogate horse endometrium at days 36 to 38 to form endometrial cups. This results in total absence of eCG in maternal blood at any time during gestation.9 Nevertheless, the donkey fetus and its membranes continue to grow and develop until around days 60 to 65, but in most cases, this occurs without the normal interdigitation of allantochorionic villi with corresponding crypts in the endometrium that commences around day 42 in conventional intraspecies equine pregnancy.9, 57 In around 70% of donkey-in-horse pregnancies, implantation does not occur at all, with the result that the fetus becomes increasingly starved and stressed until it finally dies and is aborted between days 80 and 100 (Figure 237.12). Furthermore, this process of fetal degeneration is accompanied by a generalized infiltration of lymphocytes and other leukocytes into the area of endometrium that is in contact with the poorly attached donkey trophoblast; this response has the appearance of a cell-mediated rejection reaction, mounted in this case in response to maternal recognition of xenogeneic donkey antigens at the poorly developed placental interface (Figure 237.13). In the other 30% of donkey-in-horse pregnancies, however, the rate of attachment and interdigitation of the allantochorion and endometrium is somewhat slower than it is in either intraspecies horse or donkey pregnancy. It eventually occurs, however, and these ‘successful implanters’ then seem to develop relatively normally until at or near term, at which time a second clear division occurs. In about half the cases, a live and particularly large and robust donkey foal is born, having clearly benefited in utero from the larger area of placental exchange in the bigger uterus of the recipient mare (see Figure 237.7). But in the other half, the donkey foal is either aborted dur-
Figure 237.12 A degenerating donkey conceptus (Equus asinus, 2n = 62) aborted by its surrogate horse mother (E. caballus, 2n = 64) on day 83 of gestation. Note the pale and bloodless allantochorion and the cachectic fetus within the heavily bloodstained amniotic fluid.
ing the last 4–8 weeks of pregnancy, or is born at or near term in a live but weak, dysmature, and nutritionally deprived state. This latter group shows a placenta that is much smaller and lighter than normal because of the combination of fewer and less welldeveloped microcotyledons per area of tissue. This, in turn, is clearly related to the slower rate of differentiation and interdigitation of the allantochorion much earlier in pregnancy. Thus, a complete gradation appears to exist within this one type of extraspecific donkey-in-horse pregnancy in terms of implantation, placentation, and fetal survival. This ranges from apparently normal in those few mares that give birth to big and robust donkey foals, to another small ‘just made it’ group that give birth to small but dysmature foals at or beyond term, to another small ‘almost made it’ group that abort severely undernourished foals within a few weeks before term, to the majority of ‘didn’t have a chance’ animals that undergo the now well-characterized pattern of failed implantation and inadequate placentation leading to fetal starvation, death, and abortion before day 100. During the past decade the authors have carried out a variety of experiments aimed at elucidating
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Figure 237.13 Histological sections of the allantochorion–endometrium interface in (a) a mare carrying a viable intraspecies horse conceptus at day 60 of gestation, and (b) a mare carrying a transferred extraspecies donkey conceptus at day 71. The typical branched chorionic villi interdigitating with endometrial crypts of the implanting horse-in-horse pregnancy are absent in the failing donkeyin-horse pregnancy. Instead, appreciable quantities of exocrine secretion tend to separate the trophoblast from the endometrial epithelium while large numbers of lymphocytes and other leukocytes accumulate in the endometrial stroma (× 100).
the underlying causes of the high rate of pregnancy failure in the donkey-in-horse model, which differs so strikingly from its counterparts among the range of equine extraspecies pregnancies and also from early embryonic death observed commonly in normal horse-in-horse pregnancies. Initially, hormone deficiency seemed likely to play an important part in the syndrome, because the failure of endometrial cup development meant that (i) eCG was completely absent from maternal blood during the first half of pregnancy; (ii) the normal rises in plasma progesterone concentrations after day 40–50 did not occur in most animals because of an absence of secondary luteal development;9 and (iii) the normal sharp rise in plasma conjugated estrogen concentrations between days 36 and 40 resulting from eCG stimulation of the maternal ovaries58–60 also failed to occur. Moreover, plasma progesterone concentrations declined steadily over a 15- to 20-day period in mares destined to abort their donkey conceptuses during the first 100 days of pregnancy, and the levels reached baseline coincidentally with expulsion of the degenerating fetus and membranes (Figure 237.14a). However, supplementation of four mares carrying donkey conceptuses with high daily doses of the orally active synthetic progestagen altrenogest (Regumate; Hoechst Animal Health, Milton Keynes, UK) failed to prevent fetal death during the characteristic period in all four animals. Furthermore, the exogenous progestagen hastened the demise of the primary corpus luteum in the treated mares, as evidenced by a more rapid decline of plasma progesterone concentrations to basal values compared with the situation in the untreated mares (Figure 237.14b). Similarly, fetal death and abortion occurred during
the characteristic period (days 83 to 105) in six of seven mares carrying donkey conceptuses that were given high doses (20 000 to 300 000 IU) of partially purified eCG on one or more occasions between days 38 and 60 of gestation and which showed measurable eCG concentrations in their serum after this exogenous treatment (Table 237.4). Two types of immunological therapy were applied to the model and both seemed to increase the rate of fetal survival and the chance of successful pregnancy in the treated animals.9 The first of these was a type of passive immunization whereby six mares carrying donkey fetuses were given repeated intravenous infusions, every 5 days between days 40 and 80 of gestation, of large volumes (0.75–1.2 L) of serum recovered from mares carrying normal intraspecific horse conceptuses at equivalent stages of gestation. Three of these mares (50%) then carried their donkey foals to term whereas all three of the control mares similarly infused with non-pregnant mares’ serum aborted at the usual time (Table 237.5). The second type of immunological treatment involved active immunization, following reports in the literature concerning the successful treatment of women with histories of repeated spontaneous abortion by immunizing them with paternal or thirdparty lymphocytes.61 Nine mares carrying donkey pregnancies were immunized, on four occasions between days 28 and 52 after ovulation, with approximately 200 × 106 mixed washed peripheral blood lymphocytes recovered either from the donkey sire and dam of the particular donkey embryo being carried, or from a panel of four unrelated male and female donkeys. Six of the nine lymphocyte-treated mares remained pregnant to > 150 days of gestation
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Mare: Pregnancy, Parturition and the Puerperal Period DONKEY X DONKEY
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12
MARE TO
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MARE HT
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(a)
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Days of pregnancy DONKEY X DONKEY
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MARE WJ
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MARE OA
6
20
(b)
40
60
80
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120
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DAYS OF PREGNANCY
Figure 237.14 Peripheral plasma progesterone concentrations measured in mares carrying transferred extraspecies donkey conceptuses that were (a) not given any exogenous hormonal therapy, and (b) given 30 mg altrenogest daily from around day 35 of gestation onward (hatched horizontal bar). The vertical arrows in (a) show the days on which three of the untreated mares aborted their degenerating conceptus. The curved arrows in (b) indicate the period of gestation when the fetus died and resorption of the conceptus began in all four of the altrenogest-treated mares. Note the premature decline in endogenous plasma progesterone concentrations in the group of treated mares. From Allen.9
to give an improved fetal survival rate of 67% (Table 237.5). None of the immunized mares showed any detectable eCG in their blood at any stage and, as judged by the relatively flat peripheral plasma pro-
gesterone profiles exhibited between days 40 and 100, none of them had any secondary luteal development. Nevertheless, on the basis of plasma progesterone assays and other clinical parameters, it was impossible to predict prospectively, or to determine retrospectively, why the three mares still aborted their donkey conceptuses despite having been immunized (Figure 237.15). These striking and inexplicable differences in the responses of individual mares carrying donkey conceptuses to immunological and other forms of treatment attempted to date leave the question as to whether the whole phenomenon of early pregnancy failure in this model is mediated immunologically, or whether it has a genetic basis. Unfortunately, the difficulty and cost of establishing such xenogeneic pregnancies makes it virtually impossible to carry out controlled experiments with sufficient numbers of replicates to provide statistically valid results and comparisons. Nevertheless, one striking observation that has emerged from the results of the experiments undertaken so far points strongly to the involvement of both genetic and immunological factors in the process. Namely, the high correlation that exists between the outcome of successive pregnancies in individual mares to which donkey conceptuses were transferred in successive breeding seasons. For example, of 10 mares that carried their first donkey pregnancies to term, no fewer than nine carried a second donkey conceptus successfully; two of these same mares were given a third successive donkey pregnancy and these were also carried successfully without any form of exogenous therapy.62 Conversely, all of seven mares that aborted their first donkey pregnancies aborted second donkey conceptuses in the following breeding season; six of these seven second abortions occurred significantly earlier in gestation and both of the third successive abortions occurred even earlier (Figure 237.16). This positive correlation between the outcome of the first and subsequent donkey pregnancies points strongly toward a genetic basis for the phenomenon (Figure 237.17). However, the pronounced decrease in the interval from embryo
Table 237.4 The extraspecies donkey-in-horse pregnancy model of pregnancy failure: endocrinological treatments
Treatment
Number of mares
Number aborted before day 120
None (control)
Fetal survival (%)
12
11
8
Progestagen (altrenogest) from day 36
4
4
0
Equine chorionic gonadotropin (20 000 to 300 000 IU repeatedly from day 38)
7
6
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Table 237.5 The extraspecies donkey-in-horse pregnancy model of pregnancy failure: immunological treatments
Treatment
Number of mares
Number that aborted before day 120
Fetal survival (%)
None (control) Infusions of pregnant mares serum Immunized with donkey lymphocytes
24 6 9
19 3 3
21 50 67
transfer to abortion in second and later donkey-inhorse pregnancies is reminiscent of immunological memory, one of the hallmarks of the immune system. In summary, strong suggestive evidence exists that the characteristic degeneration leading to death and expulsion of the fetus and membranes that occurs between days 80 and 100 of gestation in around 70% of the extraspecific donkey-in-horse pregnancies involves a vigorous maternal cell-mediated rejection response to the mare’s recognition of foreign fetal antigens. Enormous numbers of lymphocytes accumulate in the endometrium in contact with the degenerating allantochorion (see Figure 237.13b); and these have been observed tracking through the endometrial epithelium, apparently en route to attack the xenogeneic trophoblast cells.9, 62 Furthermore, the severe hemolysis of the fetal red blood cells that occurs before the death of the donkey fetus is also a characteristic feature of another murine model of early pregnancy failure, namely, that occurring when rare Mus caroli (mouse) embryos are transferred to the uteri of the common laboratory mouse
DONKEY
HORSE (IMMUNIZED PARENTAL DONKEY LYMPHOCYTES)
1984
L∅
16
‘‘QL’’
(M. musculis).63 But whether these immunological changes are actually the cause of fetal death leading to abortion, or whether they occur merely as a consequence of developmental placental failure enabling maternal recognition of fetal antigens that would otherwise remain masked, remains unclear.
Abortion of extraspecies donkey - in - horse pregnancies rejective - type memory in successive gestations 100
80
60
40
20
0 DN
GW
RC
DS
SL
RAA
SR
Mare
Figure 237.16 The successively earlier stages of gestation when abortion occurred in seven pony mares carrying their first (filled bar), second (open bar), and third (hatched bar) successive transferred donkey-in-horse pregnancies.
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40 60 DAYS OF PREGNANCY
80
100
Figure 237.15 Peripheral plasma progesterone concentrations measured in four mares carrying transferred extraspecies donkey conceptuses that were each immunized four times between days 28 and 52 of gestation with washed parental donkey lymphocytes. In the two mares that aborted (arrows), plasma progesterone concentrations were equal to, or higher than, those in the two mares that remained pregnant.
Figure 237.17 Genetic drift in the equine nursery. Surrogate domestic horse (Equus caballus, 2n = 64) mothers with their transferred extraspecific foals; a Przewalski’s horse (E. przewalskii, 2n = 66), a domestic donkey (E. asinus, 2n = 62), and a Grant’s zebra (E. burchelli, 2n = 44).
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References 1. Tegetmeier WB, Sutherland CL. Horses, Asses, Zebras, Mules and Mule Breeding. London: Horace Cox, 1895. 2. Ewart JC. On zebra-ass hybrids; with observations on the relationships of the zebras. Veterinarian 1898;71:185–202. 3. Gray AP. Mammalian Hybrids. Slough, UK: Commonwealth Agricultural Bureaux, 1971. 4. Short RV. The contribution of the mule to scientific thought. J Reprod Fert Suppl 1975;23:359–64. 5. Short RV. The evolution of the horse. J Reprod Fert Suppl 1975;23:1–6. 6. Jocelyn HD, Setchell BP. Regnier de Graaf on the human reproductive organs. J Reprod Fert Suppl 1972;17 [trans.]. 7. Wodsedalek JE. Causes of sterility in the mule. Biol Bull Mar Biol Lab Woods Hole 1916;30:1–56. 8. Taylor MJ, Short RV. Development of the germ cells in the ovary of the mule and hinny. J Reprod Fert 1973;32: 441–5. 9. Allen WR. Immunological aspects of the equine endometrial cup reaction and the effect of xenogeneic extraspecies pregnancy in horses and donkey. J Reprod Fert Suppl 1982;31:57–94. 10. Allen WR. The influence of fetal genotype upon endometrial cup development and PMSG and progestagen production in equids. J Reprod Fert Suppl 1975;23:405–13 11. Allen WR. Maternal recognition of pregnancy and immunological implications of trophoblast-endometrium interactions in equids. In: Maternal Recognition of Pregnancy. CIBA Foundation Symposium No. 64. Amsterdam: Excerpta Medica, 1979; pp. 323–52. 12. Allen WR. Embryo transfer in the horse. In: Adams CE (ed.) Mammalian Egg Transfer. Boca Raton: CRC Press, 1982; pp. 126–54. 13. Hancock JL, McGovern PT. The transport of sheep and goat spermatozoa in the ewe. J Reprod Fert 1964;15:283–7. 14. Hancock JL. Animal hybrids: Causes of death before birth. New Scientist 1967;35:504. 15. Hancock JL, McGovern PT, Stamp JT. Failure of gestation of goat X sheep hybrids in goats and sheep. J Reprod Fert Suppl 1968;3:29–36. 16. Chang MC, Marston JH, Hunt DM. Reciprocal fertilization between the domesticated rabbit and the snowshoe hare with special reference to insemination of rabbits with an equal number of hare and rabbit spermatozoa. J Exp Zool 1964;155:437–45. 17. Barton SC, Surani MAH, Norris ML. Role of paternal and maternal genomes in mouse development. Nature 1984;311:374–6. 18. Chandley AC, Jones RC, Dott HM, Allen WR, Short RV. Meiosis in interspecific equine hybrids. I. The male mule (Equus asinus × E. caballus) and hinny (E. caballus × E. asinus). Cytogenet Cell Genet 1974;13:330–41. 19. Anderson WS. Fertile mare mules. J Hered 1939;30:549–51. 20. Benirschke K, Low RJ, Sullivan MM, Carter RM. Chromosome study of an alleged fertile mare mule. J Hered 1964;55:31–8. 21. Chandley AC, Clarke CA. Gum mula peperit. J R Soc Med 1985;78:800–1. 22. Ryder OA, Chemnick LG, Bowling AT, Benirschke K. Male mule qualifies as the offspring of a female mule and Jack donkey. J Hered 1985;76:379–81. 23. Chandezi M. August/September update on the miracle mule baby. Mules and More 2007, July.
24. Henry M, Gastal EL, Pissheiro LEL, Guimaraes SEF. Mating patterns and chromosome analysis of a mule and her offspring. Biol Reprod Monogr Ser 1995;1:273–80. 25. Kay G. A foal from a mule in Morocco. Vet Rec 2003;152:92. 26. Rong R, Chandley AC, Song J, McBeath S, Tan PP, Bai Q, Speed RM. A fertile mule and hinny in China. Cytogenet Cell Genet 1988;47:134–9. 27. Chandley AC. Fertile mules. J R Soc Med 1988;81:2. 28. Michie D. Affinity: a new genetic phenomenon in the house mouse, evidence from distant crosses. Nature 1953;171:26–7. 29. Rong R, Cai H, Wei J. Fertile mule in China and her unusual foal. J R Soc Med 1985;78:821–5. 30. Zong E, Fan G. The variety of sterility and gradual progression to fertility in hybrids of the horse and donkey. J Hered 1989;62:393–406. 31. Yang F, Fu B, O’Brien PC, Nie W, Ryder OA, FergusonSmith MA. Refined genome-wide comparative map of the domestic horse, donkey and human based on cross-species chromosome painting: insight into the occasional fertility of mules. Chromosome Res 2004;12:65–76. 32. Koulischer L, Frechkof S. Chromosome complement: A fertile hybrid between Equus przewalskii and Equus caballus. Science 1966;151:93–5. 33. Short RV, Chandley AC, Jones RC, Allen WR. Meiosis in interspecific equine hybrids. II. The Przewalski horse/domestic horse hybrid (Equus przewalskii × E. caballus). Cytogenet Cell Genet 1974;13:465–78. 34. Chandley AC, Short RV, Allen WR. Cytogenetic studies of three equine hybrids. J Reprod Fert Suppl 1975;23:365–70. 35. Simpson DJ, Greenwood RES, Ricketts SW, Rossdale PD, Sanderson M, Allen WR. Use of ultrasound echography for early diagnosis of single and twin-pregnancy in the mare. J Reprod Fert Suppl 1982;32:431–9. 36. Allen WR. A quantitative immunological assay for pregnant mare serum gonadotrophin. J Endocrinol 1969;43:581–91. 37. Allen WR, Sanderson MW. The value of a rapid progesterone assay (AELIA) in equine stud veterinary medicine and management. In: Bain-Fallon Memorial Lectures. Sydney. Australian Equine Veterinarian Association, 1987, pp. 75–82. 38. Kydd J, Miller J, Antczak DF, Allen WR. Maternal anti-fetal cytotoxic antibody responses of equids during pregnancy. J Reprod Fert Suppl 1982;32:361–9. 39. Sheldrick EL, Wright PJ, Allen WR, Heap RB. Metabolic clearance rate, production rate and source of progesterone in donkeys with fetuses of different genotypes. J Reprod Fert 1977;51:473–6. 40. Stewart F, Allen WR, Moor RM. Influence of foetal genotype on the follicle-stimulating hormone luteinizing hormone ratio of pregnant mare serum gonadotrophin. J Endocrinol 1977;73:419–25. 41. Davies CJ, Antczak DF, Allen WR. Reproduction in mules: Embryo transfer using sterile recipients. Equine Vet J 1985;Suppl. 3:63–7. 42. Antczak DF, Davies CJ, Kydd J, Allen WR. Immunological aspects of pregnancy in mules. Equine Vet J 1985;Suppl. 3:68–72. 43. Kydd J, Allen WR, Boyle MS, Shephard A, Summer P. Transfer of exotic equine embryos to domestic horses and donkeys. Equine Vet J 1985;Suppl. 3:80–4. 44. Shaw E, Haupt KA. Pre- and post partum behaviour in mules impregnated by embryo transfer. Equine Vet J 1985;Suppl. 3:73.
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Inter and Extraspecies Equine Pregnancies 45. Allen WR. The immunological measurement of pregnant mare serum gonadotrophin. J Endocrinol 1969;43:593–8. 46. Allen WR. Factors influencing pregnant mare serum gonadotrophin production. Nature 1969;223:64–6. 47. Skidmore J, Boyle MS, Cran D, Allen WR. Micromanipulation of equine embryos to produce monozygotic twins. Equine Vet J 1989;Suppl. 8:126–8. 48. Lennard SN, Stewart F, Allen WR. Insulin-like growth factor II gene expression in the fetus and placenta of the horse during the first half of gestation. J Reprod Fert 1995;103:169–79. 49. Lennard SN, Stewart F, Allen WR. Transforming growth factor β1 (TGFβ1) expression in the endometrium of the mare during placentation. Mol Reprod Develop 1995;42:131–40. 50. Lennard SN, Stewart F, Allen WR, Heap RB. Growth factor production in the pregnant uterus. Biol Reprod Monogr Ser 1995;1:161–70. 51. Stewart F, Lennard SN, Allen WR. Mechanisms controlling formation of the equine chorionic girdle. Biol Reprod Monogr Ser 1995;1:151–9. 52. Dechiara TM, Efstratiadis A, Robertson EJ. A growth deficiency phenotype in heterozygous mice carrying an insulin-like growth factor II gene disrupted by targeting. Nature 1990;345:78–80. ¨ 53. Barlow DF, Stoger R, Herrmann BG, Saito K, Schweifer N. The mouse insulin-like growth factor type-2 gene is imprinted and closely linked to the Tme locus. Nature 1991;349:84–7. 54. Allen WR, Skidmore JA, Stewart F, Antczak DF. Effects of fetal genotype and uterine environment on placental development in equids. J Reprod Fert 1993;47:55–60.
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55. Summers PM, Shephard AA, Hodges JK, Kydd JH, Boyle MS, Allen WR. Successful transfer of the embryos of Przewalski’s horse (Equus przewalskii) and Grant’s zebra (E. burchelli) to domestic mares (E. caballus). J Reprod Fert 1987;80:13–20. 56. Bennett SD, Foster WR. Successful transfer of a zebra embryo to a domestic horse. Equine Vet J 1985;Suppl. 3: 78–9. 57. Samuel CA, Allen WR, Steven DH. Ultrastructural development of the equine placenta. J Reprod Fertil Suppl 1975;23:575–8. 58. Squires EL, Ginther OJ. Follicular and luteal development in pregnant mares. J Reprod Fert Suppl 1975;23:429–33. 59. Terqui M, Palmer E. Oestrogen pattern during early pregnancy in the mare. J Reprod Fert Suppl 1979;27: 441–6. 60. Kindahl H, Knudsen O, Madej A, Edqvist LE. Progesterone, prostaglandin F2a, PMSG and oestrone sulphate during early pregnancy in the mare. J Reprod Fert Suppl 1982;32:353–9. 61. Mowbray JF, Gibbins CR, Liddell H, Reginald PW, Underwood JL, Beard RW. Controlled trial of treatment of recurrent spontaneous abortion by immunisation with paternal cells. Lancet 1985;i:941–3. 62. Allen WR, Kydd JH, Boyle MS, Antczak DF. Extra-specific donkey-in-horse pregnancy as a model of early fetal death. J Reprod Fert Suppl 1987;35:197–209. 63. Croy AB. Use of experimental embryo transfer to study the role of the immune system in embryonic death. Equine Vet J 1985;Suppl. 3:49–52. 64. King JM. Comparative aspects of reproduction in Equidae. PhD thesis, University of Cambridge, 1965.
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Chapter 238
Induction of Abortion Claire E. Card
Review of the physiology of pregnancy Before day 35 of gestation the mare is dependent upon luteal progesterone secretion for the maintenance of pregnancy. Early studies showed that ovariectomy between days 50 and 70 of gestation resulted in 45% of mares aborting within 10–15 days.1 This was direct evidence of the mare’s dependence on luteal progesterone for pregnancy maintenance. The maintenance of pregnancy gradually loses its dependency on luteal progesterone, and is replaced by progestagen secretion from the placenta in the mare.1, 2 Endometrial cup formation begins at 35 days and continues through about 150 days of pregnancy. This coincides with the secretion of equine chorionic gonadotropin (eCG). Pregnancy loss or termination during this stage of pregnancy does not intercept eCG secretion.3 Indeed repeated daily prostaglandin treatments beginning at 42 days of pregnancy resulted in luteolysis and pregnancy loss, but did not interfere with eCG secretion.4 The biological activity of eCG is luteotrophic. Pregnant mares continue to have follicular wave activity in early pregnancy. The concurrent secretion of eCG with follicular waves in the mare results in the luteinization of large follicles, which form supplementary (accessory) corpora lutea (CLs). Supplementary CL formation driven by eCG secretion prevents the mare from returning to estrus even in the event of pregnancy loss.3 The subsequent demise of the cup tissue at around 120 days of pregnancy results in falling eCG secretion and the CLs eventually regress.
Pharmacological agents used to induce abortion The ability of pharmacological agents to terminate pregnancy in the mare is influenced by the gestation age of the mare, and the nature and dose of the
agent. A variety of pharmacological agents and solutions, alone or in combination, have been used for induction of abortion in mares, either through systemic administration, intrauterine infusion, or cervical application. The pharmacological agents include: prostaglandin, prostaglandin analogs, human chorionic gonadotropin (hCG), oxytocin, prostaglandin E, procaine penicillin, cholchicine, dexamethasone, saline, hypertonic saline, and dilute iodine solutions. Prostaglandin is reported to be the most common product and is highly efficacious.
Treatment to induce abortion Abortion is defined as pregnancy loss in the period from the completion of fetal organogenesis through late gestation when the fetus is not yet viable (50–300 days). Considerations for treatment include the stage of pregnancy, products available, desired time frame from treatment to abortion, cost, the likelihood of complications, and welfare of the mare. Historically prostaglandin is known for both its luteolytic and abortifacient effects.5 Thus treatment of mares in early pregnancy with prostaglandin mimics ovariectomy and induces early pregnancy loss.1, 6 Later in pregnancy, when luteal dependency is lost, prostaglandin appears to induce abortion through direct action on the myometrium of the mare.7 Early studies revealed the relative insensitivity of the pregnant mare to the induction of abortion using natural and synthetic prostaglandin analogs. Fenprostalene administered to mares during midpregnancy from 99 to 153 days, at luteolytic and twice the luteolytic doses, two injections 12 hours apart, was not effective in inducing abortion.8 Similarly, single low-dose injections of 1.25 mg or 2.5 mg of prostaglandin F2α (PGF2α ) resulted in abortion in only about half of mares during 40–150 days of pregnancy.6, 9 In general there is a paucity of controlled
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Induction of Abortion
Figure 238.1 A 7.5-month fetus following abortion induced with prostaglandin.
studies comparing prostaglandin treatment with intrauterine treatments for the induction of abortion. There are a variety of effective protocols available to induce abortion. Systemic administration of prostaglandin/prostaglandin analogs is presently one of the preferred methods for inducing abortion,5–7,10–15 and may be used throughout gestation. Prostaglandin/prostaglandin analogs are commonly recommended for pregnancy termination in the first third of pregnancy.12, 16 Systemic administration of a prostaglandin/prostaglandin analog is less invasive than intrauterine treatment and is highly effective. The most common treatment to induce pregnancy loss before 35 days is PGF2α or its analogs.6 From days 35 to 300 of pregnancy, repeated daily to twice daily intramuscular treatments with 10 mg of PGF2α or 250 µg cloprostenol are usually administered (Figure 238.1). Typically, two to four treatments of prostaglandin are required to induce abortion.4, 7, 13 Douglas and Ginther reported that 13 mares treated with 2.5 mg of PGF2α at 12-hour intervals aborted following an average of 3.7 injections (range of 1–7), with no anorexia, diarrhea, colic, or sweating.9 The cervix of the pregnant mare is long and tight, and must be dilated in order to introduce a catheter or infusion pipette. Prostaglandin E1 (misoprostol 200 micrograms) and E2 (various formulations) are reported to induce cervical dilation within 90–120 minutes in mares from 80 to 300 days of gestation. Subsequent treatment of the PGE-treated mares with oxytocin failed to induce abortion, and manual removal of the fetus was performed. Retention of fetal membranes for up to 40 hours was reported, but did not have adverse effects on the health and breeding potential of the mares.14 The addition of estradiol benzoate (5 mg two times, 12 hours apart) and oxytocin (50 IU 12 hours later) to the prostaglandin treatment decreased the
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time to abortion in two mares to 13 and 25 hours, compared with two other mares that aborted at 37 and 61 hours following 10 mg PGF2α twice daily, 12 hours apart.2 Oxytocin treatment resulted in additional secretion of PGF2α .2 If abortion has not occurred by the third day of treatment, 20 IU of oxytocin may be administered intramuscularly 30 minutes after the administration of prostaglandin PGF2α . Oxytocin administration is associated with a more sustained release of PGF2α .2 Following PGF2α or cloprostenol administration, mares will frequently show transient side effects within 20 minutes of administration; these include: profuse sweating, abdominal discomfort/colic-like signs, flehman response, ataxia, and increased defecation. In addition treated mares may have episodes of discomfort between the administrations of prostaglandin before abortion occurs. Presumably this relates to endogenous prostaglandin release, such as has been reported following treatment with cloprostenol.7 Interestingly, treatment with flunixin meglumine (500 mg i.v.) at 8-hour intervals did not delay the time to abortion in mares treated with oncedaily cloprostenol, suggesting flunixin meglumine did not modulate endogenous prostaglandin release.17 Progesterone (300 mg/day i.m.) or oral altrenogest (44 mg/day) administration to mares that have been treated with prostaglandin to induce abortion appears to have a protective effect by inhibiting endogenous prostaglandin secretion.18 The nonsteroidal anti-inflammatory drugs (NSAIDs), such as flunixin meglumine, fail to inhibit endogenous prostaglandin release during induced abortion.17 While the mechanism has not been identified, it may be that progesterone and altrenogest alter cytokine secretion. The fetus may be delivered with the fetal membranes intact or ruptured, and it may be delivered non-viable but alive. Sensitivity to owners’ concerns about animal welfare are important in this regard. Iatrogenic disruption of the fetal membranes, such as with biopsy forceps, or transcervical passage of a pipette to deliver iodine, procaine penicillin G, or hypertonic saline into the uterus will also disrupt the pregnancy, but are considered more invasive and carry the risk of bacterial contamination.8 Intra-allantoic infusion of 1 liter of saline containing 0.5% povidone-iodine in 6/6 mares at day 75 of pregnancy induced abortion more rapidly than daily PGF2α. These mares were reported to have normal uterine involution.19 Single or repeated infusions of sterile saline, sometimes followed by recovery of the fluid, have been described to terminate pregnancy, with high efficacy reported prior to placental attachment at about 80 days.12, 16 A variety of other
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intrauterine treatments have been used to induce pregnancy loss including: 100–200 mL of saline, hypertonic saline, dilute povidone or Lugol’s iodine, procaine penicillin G, and dexamethasone. Miscellaneous drugs include the use of human chorionic gonadotropin (hCG; 2000 IU every other day beginning on day 39 for three treatments), estrogen, and fetal injection with colchicine or potassium chloride.16, 20 Injection of hCG after day 40, presumably related to eCG secretion, was ineffective.20 The pharmacological agents used in intrauterine infusions function by being directly embryocidal; disturbing the function of the endometrium; or disrupting the function of the placenta. Physical disruption of the placental membranes in the horse triggers pregnancy loss, by inducing physical damage to the placenta and presumably activating cytokine production and initiating an inflammatory cascade.
been reported (F. Bristol, personal communication) to be normal after prostaglandin-induced parturition in the mare.25 Presently hormonal methods of lactation induction in pluriparous mares provide an additional management option to support orphan foals, one that does not require the induction of abortion.26 Fetal anomalies can include abembryonic vesicles or fetal malformation.27 Abnormal embryonic development is often self-correcting in that spontaneous pregnancy loss occurs. When a fetal anomaly does not result in timely pregnancy loss, induction of abortion may be indicated to allow the mare to be rebred earlier. Maternal compromise from pregnancy usually is related to the stress of carrying the additional weight of the fetus, fluids, and placenta. Problems related to pregnancy include: ventrolateral abdominal muscle disease, hydropic conditions, and laminitis.
Indications for induction of abortion
Ventrolateral abdominal muscle disease and prepubic tendon rupture
The common indications for induction of abortion include: mismating, twin pregnancy, nurse mare, fetal anomaly, or maternal compromise from pregnancy.6 Mismating in the mare is best treated early in gestation, when ultrasound may be used to detect an embryo and/or prostaglandin can be administered to induce luteolysis. From 5 days post-ovulation to day 35 of gestation the mare will quickly return to estrus. Therefore management steps must be taken, such as fence repairs, to prevent re-exposure of the mare to the unwanted sire. In addition mares treated with PGF2α beginning at day 42 of pregnancy with 5 mg PGF2α daily, aborted within 3–4 days of the first injection and displayed estrus 2–6 days later, and were reported to ovulate.4 However, due to endometrial cup formation termination of pregnancy is recommended before 35 days, if the mare is to be rebred that season. As noted previously as pregnancy continues multiple days of prostaglandin treatment or other interventions are needed to induce abortion. Twin pregnancies are sometimes left undetected in mares. Prostaglandin is an affordable and convenient option for termination of a twin termination pregnancy. Consideration should be given to the options available for the reduction of a twin pregnancy to one viable fetus through: transrectal crushing of the embryonic vesicle; allantocentesis; fetal craniocervical dislocation; or transabdominal fetal cardiothoracic puncture.21–24 These techniques require specialized equipment and/or training. Induced abortion has sometimes been used to provide a nurse mare for another foal. Preterm induction with prostaglandin is more likely to result in lactation compared to oxytocin induction, and lactation has
Ventrolateral abdominal muscle disease, also referred to as body wall tears, is an uncommon problem in late pregnancy.28 Compromise or rupture of the prepubic tendon is an extremely severe form of this condition. The history often suggests that the mare was kicked, or slipped; however, in the majority of cases this is an assumption by the mare’s owner, and the actual injury to the mare was not observed. Mares from midto late pregnancy are affected. The clinical presentation includes excessive, hot, pathological ventral edema. Mares are reluctant to move and are painful. Figure 238.2 shows a mare with severe bilateral ventrolateral abdominal hernias with prepubic tendon involvement. The skin of the ventral abdomen
Figure 238.2 Mare with bilateral ventrolateral abdominal muscle disease and prepubic tendon rupture; note discoloration of the skin. White arrow points to the teat on the cranially displaced udder.
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Figure 238.3 Mare with bilateral ventrolateral abdominal muscle disease and prepubic tendon rupture.
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In some mares, both allantoic and amnionic fluid compartments appear to be involved in the hydropic condition. Pressure from the enlarged pregnant uterus is believed to cause excessive physical loading of the abdominal muscles, and problems with vascular perfusion. Diagnosis of a hydropic condition is made based on transrectal palpation/ultrasonography and transabdominal ultrasonography, and following diagnosis careful monitoring and pregnancy termination are usually recommended.33 Treatment is directed toward terminating pregnancy in the mare in a manner to avoid risk of complications from physical stress and shock from fluid imbalances. Laminitis
is discolored and weeping (Figure 238.3). The majority of ventrolateral abdominal muscle disease cases are idiopathic. If recheck examinations at 24-hour intervals shows ultrasonographic evidence of continued abdominal muscle deterioration, and serum biochemistry shows persistently elevated or significantly increased muscle enzymes, this suggests ongoing myonecrosis and damage, and is an indication to induce abortion. Bloody mammary secretion indicates that the rectus abdominus is involved, and this is considered pathognomonic for the condition. Udder displacement suggests tearing or rupture of the prepubic tendon. Mares with prepubic tendon involvement are at risk of full abdominal muscular rupture, eventration, herniation, ileus, colic, and death. It is important in near-term cases to ascertain if the survival of the fetus or mare is more important. Conservative management to allow time for natural fetal maturation or induced fetal maturation using maternal systemic administration of dexamethasone is sometimes attempted.28 Hydropic conditions Hydropic conditions in the mare include hydramnios and hydrallantois.29–31 Hydramnios (hydrops amnii) in the mare is a rare occurrence, of unknown etiology. It is sometimes associated with craniofacial deformities of the foal. A lack of fetal deglutition may have a role in the pathogenesis, although it cannot be considered the only cause, as some foals are normal. Hydrallantois is an uncommon condition in mares. The etiology of hydrallantois remains obscure. However, congenital abnormalities of the fetus, placentitis, and twin pregnancies have been associated with the condition.32 The mare undergoes a rapid increase in abdominal size over a 6- to 14-day period.
Laminitis is sometimes reported in mares in late pregnancy without other inciting causes. Excessive recumbency, saw horse stance, and reluctance to move are usually present. Hormonal and mechanical factors may be involved in the pathogenesis of laminitis in these mares. Therapeutic farrier care, attention to diet, and NSAIDs may all benefit the laminitic mare. Radiographs of the third phalanges will aid in prognosis. Induction of abortion or parturition may be beneficial in these cases. Abortion is therefore medically indicated in cases:
r r r r r
where the foal is non-viable; after mismating; when twins are present and fetal reduction is not possible; when the pregnancy is compromising the health and welfare of the mare; where medical management is not practical or non-affordable.
Complications related to induction of abortion Following induced abortion the retention of fetal membranes, reproductive tract trauma, dystocia, endometritis, and endotoxemia are all possible complications. It is recommended that the fetal membranes are evaluated for completeness. Near-term abortions may be more likely to result in dystocia due to fetal size. The large size of the fetus after 8 months of pregnancy means that there is a higher likelihood for dystocia, and it may be therefore advisable to let the mare carry to term. Generally the mare’s temperature, appetite and demeanor should be monitored for a few days following an induced abortion. Excess vaginal discharge or fever are indications for further evaluation of the mare, including a detailed examination of the reproductive tract.33
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References 1. Holtan DW, Squires EL, Ginther OJ. Effect of ovariectomy on pregnancy in mares. J Anim Sci 1975;41:359–60. 2. Madej A, Kindahl H, Nydahl C, Edqvist LE, Stewart DR. Hormonal changes associated with induced late abortions in the mare. J Reprod Fertil Suppl 1987;35:479–84. 3. Allen WR. Maternal recognition and maintenance of pregnancy in the mare. Anim Reprod (Belo Horizonte) 2005;2:209–23. 4. Rathwell AC, Asbury AC, Hansen PJ, Archbald LF. Reproductive function of mares given daily injections of prostaglandin F2alpha beginning at day 42 of pregnancy. Theriogenology 1987;27:621–30. 5. Christiansen IJ. Prostaglandins for termination of pregnancy. Arsberetning, Institut for Sterilitetsforskning 1980;23:96–108. 6. Douglas RH, Squires EL, Ginther OJ. Induction of abortion in mares with prostaglandin F2alpha. J Anim Sci 1974;39:404–7. 7. Daels PF, Mohammed H, Montavon SME, Stabenfeldt GH, Hughes JP, Odensvik K, Kindahl H. Endogenous prostaglandin secretion during cloprostenol-induced abortion in mares. Anim Reprod Sci 1995;40:305–21. 8. Bosu WTK, McKinnon AO. Induction of abortion during midgestation in mares. Can Vet J 1982;23:358–60. 9. Douglas RH, Ginther OJ. Effects of prostaglandin F2alpha on the oestrous cycle and pregnancy in mares. J Reprod Fertil Suppl 1975;23:257–61. 10. Lange W, Kretschmer HJ, Schutzler H, Schubert E, Kley R. Use of prostaglandin analogues to induce abortion in PMSG donor mares. Monatsh Veterinarmed 1989;44:605–8. 11. Squires EL, Hillman RB, Pickett BW, Nett TM. Use of Equimate for induction of abortion: effect on secretion of progesterone, pregnant mare serum gonadotropin and reproductive performance. J Anim Sci 1979;49(Suppl. 1):338. 12. Paccamonti DL. Elective termination of pregnancy in mares. J Am Vet Med Assoc 1991;198:683–9. 13. Vanleeuwen W, Noden PA, Dieleman SJ. Induced-abortion with 2 prostaglandin-F2-alpha analogs in mares – plasma progesterone changes. Vet Quart 1983;5:97–100. 14. Volkmann DH, Cramer KGMd. Prostaglandin E2 as an adjunct to the induction of abortion in mares. In: Equine Reproduction V: Proceedings of the Fifth International Symposium on Equine Reproduction, 1991. 15. Wichtel JJ, Evans LE, Clark TL. Termination of midterm pregnancy in mares using intra-allanotic dexamethasone. Theriogenology 1988;29:1261–7. 16. Loftsted R. Termination of unwanted pregnancy in the mare. In: Morrow D (ed.) Current Therapy in Theriogenology. Philadelphia: WB Saunders, 1986; p. 715. 17. Daels PF, Mohammed HO, Odensvik K, Kindahl H. Effect of flunixin meglumine on endogenous prostaglandin
18.
19.
20. 21.
22.
23. 24. 25. 26.
27.
28.
29.
30. 31. 32. 33.
F2alpha secretion during cloprostenol-induced abortion in mares. Am J Vet Res 1995;56:1603–10. Daels PF, Besognet B, Hansen B, Mohammed HO, Odensvik K, Kindahl H. Effect of progesterone on prostaglandin F-2 alpha secretion and outcome of pregnancy during cloprostenol-induced abortion in mares. Am J Vet Res 1996;57:1331–7. Varner DD, Meyers PJ, Evans JW, Wiest JJ, Kloppe LH, Elmore RG. Effect of equine abortifacients on fetal viability and post-abortion reproductive performance. In: Proceedings of the 11th International Congress on Animal Reproduction and Artificial Insemination, Dublin, Ireland, 26–30 June, 1988; Vol. 2. Allen WE. Pregnancy failure induced by human chorionic gonadotrophin in pony mares. Vet Rec 1975;96:88–90. Morar I, Groza I, Bogdan L, Ciupe S, Pop R. Twin pregnancy in mares, diagnostic and management. Lucrai Stiinifice – Medicina Veterinara, Universitatea de Stiinte Agricole si Medicina Veterinara “Ion Ionescu de la Brad” Iasi 2004;47:539–42. Meshram BN, Kamble MV, Dalvi RS. Detection and management of twin pregnancies in Thoroughbred mares. Intas Polivet 2004;5:311–12. Wolfsdorf KE. Management of postfixation twins in mares. Vet Clin N Am-Equine P 2006;22:713–25. Macpherson ML, Reimer JM. Twin reduction in the mare: current options. Anim Reprod Sci 2000;60:233–44. Bristol F. Induction of parturition in near-term mares by prostaglandin F-2 alpha. J Reprod Fertil Suppl 1982;32:644. Daels PF, Duchamp G, Porter D. Induction of lactation and adoption of foals by non-parturient mares. In: Proceedings of the 48th Annual Convention of the American Association of Equine Practitioners, Orlando, Florida, USA, 4–8 December 2002, pp. 68–71. Vanderwall DK, Squires EL, Brinsko SP, McCue PM. Diagnosis and management of abnormal embryonic development characterized by formation of an embryonic vesicle without an embryo in mares. J Am Vet Med Assoc 2000: 217(1); 58–63. Ross J, Palmer JE, Wilkins PA. Body wall tears during late pregnancy in mares: 13 cases (1995–2006). J Am Vet Med Assoc 2008;232:257–61. Henry MM, Morris DD, Pugh DG. Hydrallantois associated with twin pregnancy in a mare. Equine Pract 1991; 13:20–3. Bucca S. Diagnosis of the compromised equine pregnancy. Vet Clin N Am-Equine P 2006;22:749–61. Bucca S, Romano G. Case of equine hydramnios in late pregnancy. Ippologia 2000;11:35–8. Blanchard TL. Managing the mare with hydrallantois. Vet Med 1989;84:790–2. Frazer GS, Embertson R, Perkins NR. Complications of late gestation in the mare. Equine Vet Educ 1997;9:306–11.
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239.
Embryonic Loss Barry A. Ball
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Abortions and Stillbirths: A Pathologists Overview Katherine E. Whitwell
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Origin and Outcome of Twin Pregnancies Angus O. McKinnon
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Placentitis Mats H.T. Troedsson and Margo L. Macpherson
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Hydrops Rudolf O. Waelchli
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Fetal Maceration and Mummification Claire E. Card
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Equine Herpesvirus Peter J. Timoney
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Equine Viral Arteritis Peter J. Timoney
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Contagious Equine Metritis Peter J. Timoney
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Mare Reproductive Loss Syndrome David G. Powell
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Fescue Toxicosis Dee L. Cross
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Prepubic and Abdominal Wall Rupture Dwayne H. Rodgerson
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Uterine Prolapse Michael A. Spirito and Kim A. Sprayberry
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Uterine Torsion James R. Vasey and Tom Russell
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Abnormalities of Pregnancy ¨ Robert M. Lofstedt
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Maintenance of Pregnancy Angus O. McKinnon and Jonathan F. Pycock
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Dystocia Management Grant Frazer
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Fetotomy Grant Frazer
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Cesarian Section David E. Freeman
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Referral Dystocias Rolf M. Embertson
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Periparturient Hemorrhage T. Douglas Byars and Thomas J. Divers
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Retained Fetal Membranes Walter R. Threlfall
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Postpartum Metritis Terry L. Blanchard
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Chapter 239
Embryonic Loss Barry A. Ball
Introduction In contrast to other domestic livestock, the horse is maintained as part of the breeding population for a relatively longer period of time. In many situations, older mares may be more valuable because of their proven produce record, and it is common to find mares that are bred at greater than 20 years of age. For these reasons, age-related reproductive failure has received considerable study. This chapter will summarize our current understanding of reproductive failure in mares, particularly as it relates to embryonic loss. Foaling rates decline with maternal age after 14 to 16 years.1–3 Although pregnancy losses during late gestation increase in older mares, it appears that failures of early pregnancy account for the majority of the reduced fertility in aged mares. The widespread application of transrectal ultrasonography for pregnancy detection in mares at Days 10 to 14 after ovulation led to evaluation of the incidence of embryonic loss in mares between Days 14 and 40, and these studies indicate that overall pregnancy rates declined and that detected embryonic loss rates increased with increasing mare age.1, 4 A recent review evaluated the overall incidence of detected embryonic loss in mares as approximately 7.7%.5 Data from field studies such as these provided a strong impetus to examine very early embryonic mortality in mares.
aged mares survive after embryo transfer to healthy recipient mares.7 Because the equine embryo enters the uterus at Day 6 to 6.5, it was unclear from these observations whether the decline in number and apparent viability of blastocysts from aged mares was related to an abnormal uterine environment, abnormalities of the oviductal environment, or abnormalities inherent in the embryo. Several studies have characterized the early equine embryo during oviductal passage in mares of different ages and fertility status. As in other species, the overall fertilization rate in young mares appears high (> 90%) based upon the cleavage rate of ova recovered 2 days after fertilization whereas the fertilization rate in aged mares was 80 to 90%. Estimates of the embryonic loss rate between fertilization and Day 10 were 9% for young mares compared to 60 to 70% for aged mares.8–10 Although pregnancy rates at Day 2 were similar in young and aged mares, by 4 days after fertilization, there was a significant reduction in pregnancy rates in aged mares. This finding suggested that the interval between Day 2 and 4 might represent a critical period in pregnancy failure in aged mares. These studies confirmed the importance of very early embryonic loss in the reduced fertility of aged mares; however, such studies do not provide information as to the causation of these losses.
Factors in embryonic loss Incidence of early embryonic loss Prior to Day 10, ultrasonographic evaluation cannot accurately detect the early equine conceptus; however, several reports based upon embryo collection and transfer provide information regarding very early embryonic loss in mares. From these studies, it appears that there is a sharp decline in the recovery rate of blastocysts from the uterus of aged mares.6, 7 In addition, blastocysts from aged mares have more morphological defects,6 and fewer blastocysts from
Pre-uterine events associated with embryonic loss in mares Events that occur prior to Day 6 to 6.5 of pregnancy in the mare within the oviduct have received limited study due to the limited accessibility to the oviduct. However, as noted above, a relatively high rate of embryonic loss appears to occur prior to entry of the embryo into the uterus, and therefore these pre-uterine events are of particular importance in understanding
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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early embryonic loss in mares. The role of pre-uterine events in embryonic wastage in aged mares was examined by collection of embryos from the uterine tube of aged and young mares at 4 days after fertilization.9 The viability of these embryos was assessed after transfer to the uterus of young recipient mares. The survival of embryos from aged mares was significantly lower than those from young mares after placement into a normal uterine environment, which indicates a role for either the oviductal environment or defects inherent in the early embryo for the high rate of early losses in aged mares. The equine embryo resides in the oviduct for a relatively long time (6 days) and reaches a late stage of development (morula or early blastocyst) before entering the uterus. There are, consequently, a number of important events that occur during oviductal transit that could affect embryonic survival. During this time, the embryonic genome is activated, and the transition from maternal to embryonic control of development has been proposed as a critical juncture in embryonic development.11 Early blastomere cleavage progresses to further differentiation including compaction, blastulation, and initial formation of the inner-cell mass and trophoblast. In order to investigate developmental capacity of early embryos, we examined the in vitro development of early, Day-4 embryos from both aged and young mares during co-culture with oviductal epithelial cells. There was no catastrophic failure of development of embryos from aged mares during in vitro culture as the developmental rate of embryos from young and aged mares to blastocysts was similar. However, embryos from aged mares had fewer blastomeres and poorer quality scores than did embryos from young mares.10 This study supported a reduction in embryo quality in embryos from aged mares during the preblastocyst period. Oviductal transit of the embryo in the mare is unique among the domestic animals because the early embryo actively regulates transit from the oviduct into the uterus.12, 13 Regulation of embryo transit through the isthmic portion of the oviduct is dependent upon normal development of the embryo14 and is regulated through secretion of prostaglandin E2 in part by the early conceptus.15 Therefore, if embryonic development is abnormal during the early cleavage stages, the equine blastocyst will not passage through the oviduct into the uterus as expected at Day 6. Anecdotally, embryo transit into the uterus of aged mares appears to be delayed and many embryo transfer practitioners delay embryo recovery attempts from such mares until Day 8 after ovulation. Although the relationship between delayed rates of embryonic development and embryo transit through the oviduct of older
mares has not been established, it is possible that delayed embryonic development in these mares is also associated with delayed or abnormal transit through the oviduct. Oviductal (uterine tubal) factors Because the period of oviductal transit of the embryo appears to be critical to early embryonic development and loss, two studies were conducted to evaluate the role of the uterine tubal environment on early embryonic loss in aged mares. In the first study, histopathological evaluation of both the ampulla and isthmic regions of the uterine tube was conducted on aged and young mares. In contrast to the findings associated with the endometrium, there was no association between mare age and the incidence of salpingitis.16 In a second study, differences in proteins synthesized and secreted in vitro by explanted oviductal tissue was examined by incorporation of 35 S-methionine, two-dimensional polyacrylamide gel electrophoresis, and fluorography. Both qualitative and quantitative differences in the patterns of proteins from oviductal epithelium were detected between young and aged mares; however, the importance of these differences relative to embryonic development remains to be determined.17 Another, unique, feature of the equine oviduct is the presence of oviductal masses (Figure 239.1) or ‘plugs’ that have been characterized by several authors.12, 18, 19 The exact origin of these masses has been debated, but it appears likely that these aggregates of collagen and fibroblasts originate from connective tissue debris that is carried into the oviduct at the time of ovulation.18 Irrespective to their origin, these oviductal plugs are observed with considerable frequency in mares, and it has been proposed that they may act as physiological ‘ball’ valves which allow the passage of sperm up the oviduct but prevent the larger embryo from passing through the
Figure 239.1 Low-magnification micrograph demonstrating an equine oviductal ‘plug’ recovered after lavage of the oviduct of a mare.
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Embryonic Loss oviduct and into the uterus.19 In support of this hypothesis, Zent et al. demonstrated that lavage of the oviduct to remove these oviductal plugs improved fertility in three of five mares with long-standing subfertility.20 More recently, Allen and co-workers19 used the novel approach of application of PGE2 gel (Prepodil) to the serosal surface of the oviduct in 15 mares with long-standing subfertility. Laparoscopic application of PGE2 to the oviduct was theorized to induce relaxation of the oviductal smooth muscle with subsequent passage of these oviductal plugs into the uterus, thereby relieving the oviductal obstruction. Although the study was not controlled, 14 of these 15 mares established detectable pregnancies subsequent to this treatment.19 Oocyte abnormalities Deterioration of oocytes associated with increased maternal age has been well described in other species.21 Unfortunately, few studies have addressed this issue in the horse. In one study, we examined the in vitro maturation of oocytes from young and aged mares as well as the rate of aneuploidy in oocytes from each group. Unfortunately, inadequate numbers of chromosomal spreads were available to assess the rate of aneuploidy in oocytes from these two groups of mares, and the relative incidence of aneuploidy in oocytes from aged mares remains to be defined.22 However, oocytes from aged mares reached metaphase II at a much lower rate than did oocytes from younger mares, with more oocytes from older mares arresting at metaphase I, which suggests that meiotic division of oocytes from older mares is more likely to be abnormal.22 In another report, Carnevale and Ginther reported the results of gamete intrafallopian transfer studies in which oocytes were collected from both young and aged mares and transferred into the uterine tube of young recipient mares.23 Oocytes from aged mares resulted in significantly fewer pregnancies than those from young mares after transfer to the uterine tube of a young recipient mare. This study provides the most convincing evidence to date of an age-related decline in oocyte quality in mares as a major factor in the reduced fertility of these mares. It appears likely that this reduction is secondary to abnormalities during meiosis in oocytes from aged mares possibly resulting in an increased incidence of aneuploidy; however, this remains to be established for the horse. Although the mechanism responsible for reduced oocyte quality from aged mares remains undefined, a recent study indicated that there is a quantitative reduction in mitochondria in oocytes from aged compared to young mares.24 Mitochondria from in vitro matured oocytes from aged mares also demon-
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strated more ultrastructural abnormalities. Because the events of oocyte maturation and fertilization are highly energy-dependent processes, and because there are major changes in distribution of mitochondria during oocyte maturation in the horse,25 these findings suggest that alterations in mitochondria in oocytes from aged mares may be an important underlying factor associated with reduced oocyte quality in these mares. Delayed ovulation and preovulatory aging of oocytes has been proposed as another factor associated with abnormal oocytes in aged females. Aged mares appear to undergo a reproductive senescence characterized by lengthening of the follicular phase, irregular ovulations, and eventually a cessation of follicular activity.23, 26, 27 The onset of these changes appears to occur over a relatively broad age range but appears most commonly in mares > 20 years of age. Prolongation of the follicular phase appears to be associated with an elevation of both FSH and LH in aged mares.27 The relationship of declining follicular populations, altered gonadotropin levels, and prolonged follicular development in aged mares with an increased incidence of abnormal oocytes remains to be explored as an explanation for age-related infertility in mares. In cattle, a decreased concentration of circulating progesterone in diestrus prior to the subsequent estrus influences LH pulse frequency and may lead to prolonged follicular development and premature resumption of meiosis.28 Oocytes derived from these large persistent follicles are able to initiate fertilization but have a high rate of embryonic death that occurs by the 2- to 16-cell stage.28 Although similar data are lacking for the mare, it appears likely that prolonged follicular development in older mares may contribute to increased abnormalities in the oocyte with a subsequent increase in embryonic loss. Aging of the oocyte after ovulation with subsequent fertilization may also result in abnormalities of oocyte function. In mares, aging of the oocyte subsequent to ovulation results in a progressive decline in pregnancy rates,29, 30 a delayed embryonic development, and possibly an increased rate of subsequent embryonic loss. Although the mechanism for the reduced developmental capacity of embryos derived from oocytes that have undergone post-ovulatory aging in mares is not known, information from other species suggests that breakdown of the meiotic spindle or reduced efficacy of the cortical reaction with subsequent polyspermic fertilization are likely factors to consider. Genetic factors Although infrequently reported, autosomal translocations have been described in phenotypically
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normal mares that were associated with reduced fertility.31, 32 More recently, a high rate of recurrent early embryonic loss was reported in three Thoroughbred mares that had reciprocal autosomal translocations detected by karyotyping.33 These findings suggest that mares with repeated early embryonic losses may be considered as candidates for karyotypic investigation in the absence of other abnormal clinical findings.
Uterine period Endocrine factors Progesterone is critical for the continued maintenance of pregnancy in mares throughout gestation and applications for progestin administration in pregnant mares have been previously reviewed.34, 35 During the embryonic period (through Day 40) progesterone is produced solely by the primary corpus luteum that is formed at the time of the initial ovulation. Lysis of the corpus luteum associated with failure of maternal recognition of pregnancy36–38 can lead to early embryonic death. Failure of the embryo to block luteolysis has been identified in mares with embryonic loss prior to day 20 and was characterized by the presence of embryonic vesicles that were too small for days of age and by failure of fixation.39 In these mares, serum progesterone concentrations were lower at days 12, 15, and 18 as was the diameter of the corpus luteum. Although the role of abnormal luteal function and deficiency of progesterone secretion during early pregnancy is cited as a cause of early embryonic loss in mares,39 there is little, if any, scientific evidence to support such a cause of embryonic loss in mares.40–42 However, in cattle, circulating progesterone concentrations in the late embryonic period (5 weeks) were positively associated with embryo survival over the next several weeks. Cows with low progesterone at 5 weeks had a higher rate of late embryonic death.28 The frequent use of exogenous progestins to prevent early embryonic loss in mares is not well justified based upon available scientific evidence. This said, the veterinary practitioner is frequently faced with clients who demand the use of exogenous progestins in pregnant mares. The practitioner is trapped in a ‘Catch 22’ in such situations and frequently resorts to the use of such therapy as a means to satisfy client demands. Interpretation of circulating progesterone concentrations in the early pregnant mare can be useful to assess luteal function in situations where inadequate progestational support is suspected clinically. Mares that have poor uterine or cervical tone, increased uterine edema, failure of fixation of the conceptus at Day 16 post-ovulation, or do not have a detectable
corpus luteum with ultrasonography of the ovaries might all represent situations in which determining serum progesterone concentrations would be useful. Minimal accepted concentrations of serum progesterone vary according to the laboratory or study reported; however, values between 2.0 and 4.0 ng/mL are typically used as the minimal serum progesterone concentration during early gestation.34, 43 Because serum progesterone concentrations vary over time, it is typically recommended that a minimum of two daily samples be evaluated to more accurately assess the progestational status of the pregnant mare. It has become common for practitioners to determine serum progesterone concentrations in embryorecipient mares at the time of embryo transfer and at the first pregnancy determination.44 In mares used for embryo recipients, there appears to be a relationship between circulating progesterone concentrations at 7 days after transfer and subsequent survival of transferred embryos; however, there was no relationship between progesterone concentrations at the time of transfer and subsequent embryo survival.44 These results suggest that non-surgical embryo transfer techniques may reduce serum progesterone concentrations, possibly associated with a partial luteolysis initiated at the time of non-surgical embryo transfer. Hypothyroidism has also been cited as a potential factor contributing to reduced fertility or early embryonic loss in mares. Two recent reports have examined the relationship between thyroid status and early pregnancy in mares. There was no association between basal serum T4 concentrations or thyroid supplementation and early pregnancy rates in mares.45 There was also no association between thyroid function as determined by TRH stimulation testing.46 Together these studies suggest that thyroid supplementation is unlikely to affect early embryonic loss rates in mares. Endometrial factors Clinically, abnormalities of the endometrium have been considered an important factor in the reduced fertility of aged mares. A number of studies have characterized degenerative changes in the equine endometrium as a function of increased mare age, and the decline in mare fertility with increased age has been frequently attributed to these changes. These changes include endometrosis, and endometritis as well as vascular changes. We examined the impact of uterine environment on embryo survival in young and aged mares in which endometrial histopathology was evaluated.47 As expected, aged mares had a higher incidence of degenerative and inflammatory changes in the endometrium. When morphologically
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Embryonic Loss normal, Day-7 or -8 blastocysts were transferred into the uterus of young and aged recipient mares, embryo survival rates at Days 12 and 28 were not different between young and aged mares (55% vs 45% at Day 12). In another report comparing the effect of recipient mare age on subsequent embryonic survival and embryonic loss rates, there were no differences in survival of equine embryos transferred into mares that were 2 to 9 years of age compared to 10 to 18 years of age based upon embryo survival rates at Day 12.48 There tended, however, to be more embryonic losses between Day 12 and Day 50 in the older mares that received embryos. These two studies suggest that uterine effects did not play a major role in the high incidence of early (prior to Day 12) embryonic loss that had been previously detected in aged mares. The impact of uterine abnormalities may be more likely during later periods in gestation in these mares and this conclusion is supported by the higher rate of embryonic loss in older mares between Days 12 and 50.48 The uterine environment clearly plays a role in embryonic losses detected by ultrasound in subfertile mares and many of these losses are detected in the period between the first detection of the embryonic vesicle with ultrasound at Day 11 and Day 15.37 Beyond Day 20, embryo loss rates in subfertile mares appeared no higher than those detected in normal populations of mares. Endometritis has long been held as an important factor in embryo losses in subfertile mares, and this is supported by the finding that the rate of embryo loss in mares with intrauterine collections of fluid during early pregnancy are associated with higher rates of embryo loss, lower progesterone concentrations, and more evidence of inflammation based upon endometrial histopathology.37, 49 A negative impact of foal-heat breeding on embryonic loss has been reported3 but has not been a consistent finding across studies.1 In some cases it is difficult to differentiate the effect of the postpartum uterine environment and other effects such as lactation or nutritional status.50 Interestingly, there appears to be an effect of side of fixation of the conceptus relative to the prior pregnancy in postpartum mares. The rate of pregnancy loss (as determined by palpation per rectum) was higher when the conceptus was located in the previously gravid uterine horn. 51 These observations await confirming studies based upon transrectal ultrasonography, but suggest that the local uterine environment may indeed play a role in survival of the conceptus in the postpartum mare. Lymphatic cysts Among the most common pathology identified by hysteroscopy in mares is the presence of uterine cysts.
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Two general categories of uterine cysts have been described including either glandular cysts or lymphatic cysts.52 However, most macroscopically evident uterine cysts are lymphatic in origin because glandular cysts tend to remain microscopic in size.53 Lymphatic cysts occur with increasing frequency in mares after 10 years of age.37, 54 Although the pathogenesis of lymphatic cysts remains unclear, it is likely that these distended lymphatics originate as a consequence of vascular degenerative changes and fibrosis in the endometrium that occur with increased age and parity in mares.55 The effect of lymphatic cysts on fertility in mares remains controversial. Many studies which have examined the relationship between the presence of lymphatic cysts and fertility in mares are confounded by mare age because the presence of cysts is strongly age-associated and because fertility is also age-dependent. The frequency of lymphatic cysts in mares over 10 years of age varies between 13% and 27% of mares examined54,56–58 with most cysts being located near the base of the uterine horn (near the site of embryo fixation). Eilts et al.56 found no relationship between the presence or absence of lymphatic cysts on either pregnancy rates or pregnancy loss rates in 274 Thoroughbred mares. In contrast, several other studies suggest that the presence of lymphatic cysts had a negative impact on fertility54, 57 although other effects such as mare age were not considered in these studies. Irrespective of the effect of lymphatic cysts on fertility, the presence of single or multiple lymphatic cysts can make early pregnancy diagnosis and twin pregnancy diagnosis based upon transrectal ultrasonography in mares much more difficult. Although cysts can often be distinguished from the embryonic vesicle based upon their appearance (lymphatic cysts are often irregular and multiloculated), solitary cysts can be confused with the early conceptus and the presence of multiple cysts can greatly hamper early pregnancy diagnosis. Therefore, removal of lymphatic cysts has become a more common procedure in barren mares. Ablation of lymphatic cysts can be conducted using either laser photoablation or loop electrocautery depending upon the preference of the veterinarian and the availability of equipment. Photoablation is based upon neodymium : yttrium (Nd:YAG) laser irridation.59–61 Loop electrocautery is an alternative for ablation of lymphatic cysts that has advantages of reduced equipment costs and reduced thermal trauma to the endometrium.62 There are no controlled studies which have examined the effect of lymphatic cyst ablation on subsequent mare fertility. Several clinical reports, however, suggest that subfertile mares that underwent cyst
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ablation were able to establish pregnancy after the procedure. Griffin and Bennett60 presented followup information on 39 mares that had undergone laser cyst ablation. These mares were barren with a mean age of 18 years. Of these mares, 26 of 39 (66%) conceived the following season. Unfortunately, this study was not controlled and no further information (such as endometrial biopsy) was available to characterize these mares. Other clinical reports have also suggested that ablation of lymphatic cysts can improve mare fertility59, 63 but appropriately controlled studies are needed to accurately assess the benefits of cyst ablation on fertility of mares. Effect of sire on embryonic loss Some3, 4, 64, 65 but not all66 field studies have described an effect of sire on the detected incidence of early pregnancy loss in mares. In two studies in Thoroughbreds, there appeared to be a subpopulation of stallions with increased rates of early pregnancy loss rates in mares.3, 4 In the group of 49 Thoroughbred stallions which had mated > 30 mares, 10 stallions had a higher rate of early embryonic loss (13–23%).4 Interestingly, increasing sire age was not associated with the rate of early pregnancy loss in mares bred to those sires in this study,4 contrary to observations from other species. The influence of stallion on embryonic loss rate in mares could be related to genetic factors, infectious diseases, or damaged sperm chromatin among others. There is relatively little information concerning possible genetic influence on embryonic loss in horses. In cattle, a 10% inbreeding coefficient of the embryo was associated with a 1% decline in fertility rates.67 Similar studies have not been conducted for the horse; however, increased inbreeding is likely to increase the incidence of early pregnancy loss associated with recessive lethal traits. Early pregnancy loss can also be associated with structural chromosomal abnormalities in cattle such as the 1/29 Robertsonian translocation.68 An increased rate of early pregnancy loss was present in mares bred to a Thoroughbred stallion with an autosomal translocation.69 These studies suggest that genetic factors should be considered as a possible cause of embryonic loss in mares bred to individual stallions with unexplained high rates of embryonic loss in their mares. Damage to sperm chromatin is another potentially important stallion factor that may influence early embryonic loss rates in mares. Sperm chromatin damage may increase during sperm storage70 or after cryopreservation.71 Spermatozoa have no ability to repair this damage although some DNA repair may occur in the oocyte after fertilization. If damage to sperm DNA is extensive, fertilization may occur; however,
the resulting embryo undergoes embryonic loss at a much higher rate as demonstrated by experimental studies in mice and in cattle.72, 73 Immune-mediated effects on pregnancy loss Although stallion ‘incompatibilities’ have been suggested to cause pregnancy loss in mares (see above), there is little documented evidence for such causes. In horses, immune recognition of paternal major histocompatability antigens appears to occur early in pregnancy,74 and these antigens are expressed by the invasive trophoblast of the chorionic girdle as soon as 32 to 36 days of gestation.74 In extraspecific pregnancies in equids (donkey embryos transferred to horse recipient mares) a high proportion of donkey embryos are lost by Days 85 to 90 of gestation, and these losses are characterized by a failure of the invasive trophoblast of the donkey conceptus to form endometrial cups within the uterus.76 These pregnancy losses are associated with little to no eCG present in the recipient mare and a failure of formation of normal allantochorionic placenta.77 Although the basis for the high rate of early fetal loss in these extraspecific pregnancies remains to be determined, it has been postulated that these early fetal losses have an immunological basis.77 This postulate was supported by the observation that immunization of pregnant mares with donkey peripheral lymphocytes during early gestation (between Days 20–50) increased the survival of donkey embryos in horse recipient mares. However, subsequent attempts to reduce the incidence of recurrent early pregnancy loss in mares based upon immunization with paternal lymphocytes had no effect on the incidence of early pregnancy loss.78 Effect of heat stress on embryonic loss Heat stress has long been defined as an important factor in early embryonic loss in ruminants; however, there has been little information available concerning the effect of heat stress on early pregnancy in mares. In a recent abstract, exercise-induced hyperthermia in mares was associated with a reduced embryo recovery rate at Day 7, possibly implying a similar effect of heat stress on early pregnancy in mares.79 Future studies should examine this possibility in the horse and address management issues to control heat stress in mares in hot climates. Effect of nutrition on embryonic loss Adverse affects of poor nutrition and low body condition scores on fertility in mares have been described,80, 81 and mares that experience increased
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nutritional demands during lactation appear to be at risk of higher early pregnancy loss rates if underfed.50, 80 In another trial, mares fed a low-protein diet had a higher rate of early pregnancy loss than did mares fed a high-quality diet (36% vs 7%, respectively).82 Lactating mares that experienced embryo loss had weight loss (25 kg) which was not experienced by mares fed a higher-quality ration. Further observations on this group of mares suggested that there were changes in serum progestagen concentrations associated with dietary restriction although mares on the low-protein diet had higher serum progesterone concentrations than did mares fed normally.83
Ultrasonographic indicators of impending embryonic loss A number of ultrasonographic findings during early pregnancy in mares may suggest impending embryonic loss.84 Embryos detected by transrectal ultrasonography that are significantly retarded in development at Days 14 to 21 have a higher rate of subsequent embryonic loss than do embryos that are normally sized for days of age.41,85–87 Although the causal relationship between delayed embryonic development and subsequent embryo loss is not known, two possible mechanisms may account for these failures. If the early embryo growth is retarded due to an embryonic defect, the embryo may not signal appropriately to block luteolysis in the mare, with subsequent embryonic loss. Similarly, the early embryo with retarded development may be inherently defective with subsequent failure associated with those defects. Such embryos have been described in some cases as trophoblastic vesicles in which the embryonic trophoblast continues to develop without subsequent development of the inner-cell mass and embryo proper. These pregnancies are typically lost by approximately Day 30 of gestation.88 The presence of intraluminal fluid accumulations during Days 11 to 15 is also associated with a high rate of embryonic loss, and many of these losses appear related to endometritis.84 There is also a higher rate of embryonic loss if the embryonic vesicle fails to undergo fixation at the base of the uterus horn around Day 16 or if an embryonic vesicle that was previously fixed moves caudally into the uterine body.84 After the appearance of the embryo proper, detection of embryonic cardiac motion beginning around Day 20–22 of gestation is a useful indicator of embryonic viability (Figures 239.2–239.5). The more widespread availability of color Doppler ultrasonography provides a useful adjunct for determination of embryonic blood flow which may be de-
Figure 239.2 Photomicrograph of an equine oocyte demonstrating large number of lipid droplets (L) along with polar distribution of mitochondria (arrows).
tected as early as Days 17 to 20 under experimental conditions.89
Treatment/management of embryonic loss Embryonic loss, particularly when it occurs over multiple cycles in a mare, is a particularly challenging and frustrating reproductive problem to manage. In many cases, diagnostic approaches may be limited to ultrasonographic imaging or measurement of circulating progesterone, and these may be undertaken
Figure 239.3 Power Doppler ultrasound scan of a Day 28 equine conceptus demonstrating blood flow within the conceptus. (See color plate 119).
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Figure 239.4 Ultrasonographic image of a Day 32 conceptus undergoing embryonic death. The conceptus demonstrates general disorganization, excess embryonic membranes, and loss of fetal heartbeat.
after the loss has occurred. Nonetheless, there are several approaches which may improve the chance of successfully re-establishing pregnancy in mares that have undergone embryonic loss. In older, problem-breeding mares, it will be useful to place these animals under artificial photoperiod in
Figure 239.5 Color Doppler ultrasonographic image of a Day 45 equine conceptus with displacement of the fetus toward the cervix and loss of cardiac motion. Blood flow is detectable in the caudal vaginal artery but not within the conceptus. (See color plate 120).
order to begin the breeding season as early as possible. An early start to the breeding season will provide as many cycles as possible to establish pregnancy. Likewise, frequent pregnancy examination with ultrasound will allow the early detection of embryonic loss and allow the mare to be re-bred as soon as possible, thus allowing more attempts per season to establish pregnancy. In older mares, it is also prudent to utilize ovulation induction with preparations such as human chorionic gonadotropin (hCG) or GnRH analogs in order to limit the duration of the follicular phase and thereby reduce the prospects of oocyte aging that may contribute to abnormalities of the oocyte in these animals. Because this group of mares is also more prone to post-mating endometritis, aggressive breeding management utilizing uterine lavage and administration of uterine ecbolics should help avoid problems attributed to uterine inflammation that persists into diestrus. Insemination or breeding once per cycle is beneficial in older mares and will further minimize post-mating endometritis. As noted above, supplementation with exogenous progesterone has been commonly used in attempts to prevent embryonic loss although the efficacy of such treatment has yet to be established. In situations where administration of exogenous progesterone is selected as a means to treat or prevent embryonic loss in mares, it seems prudent to choose a progestin and a treatment protocol that has been demonstrated to maintain pregnancy in the absence of any exogenous progesterone. A number of progestins have been shown to be ineffective in maintaining early pregnancy in the absence of endogenous progesterone including medroxyprogesterone acetate, hydroxyprogesterone caproate or hexanoate, norgestomet, and megesterol acetate.90, 91 Preparations which have been shown to be effective in maintaining pregnancy include daily altrenogest (0.44 mg/kg; orally), daily injectable progesterone in oil (150–300 mg, i.m.) or various preparations of controlled-release progesterone which allow longer intervals between administration.92–95 Although the recommended duration of progesterone supplementation varies, in most cases it seems prudent to continue administration of supplemental progestins through 100–120 days of gestation.34 This includes the period of time in which placental production of progestins is adequate to maintain pregnancy as well as the period of time in which much of placental formation is complete in mares. If progestin therapy is discontinued prior to 60 days of gestation, it may be prudent to assay serum progesterone concentrations in the mare before discontinuing progesterone administration. This is possible in those mares being supplemented with altrenogest which does not crossreact in most progesterone assays.
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Embryonic Loss Although no long-term deleterious effects on the conceptus have been established for the administration of supplemental progestins in mares, care should be used in supplementation with progestins. Mares should be monitored regularly for continued development of normal pregnancy because death and retention of the conceptus may occur in later gestation in mares receiving progesterone.96 Mares that are given supplemental progestins soon after mating and ovulation should be monitored closely for development of post-mating endometritis, which may be exacerbated by exogenous progestin administration. In a series of field trials, mares were given the GnRH analog, buserelin (20 or 40 µg) between days 8 to 12 after mating and ovulation in mares.97 Overall pregnancy rates were significantly increased by approximately 10% in buserelin-treated mares compared to controls. The authors of this study speculated that differences in early pregnancy rates in their study may have been due to a luteotrophic effect of administration of GnRH possibly through the induction of additional corpora lutea formation;97 however, a subsequent study was unable to demonstrate either luteinization or ovulation of diestrous follicles in mares after administration of another GnRH analog, deslorelin, at days 8 or 12.98 More recently, mares that received 40 µg of buserelin at day 10 after artificial insemination and ovulation had an increase in circulating LH concentrations but no change in circulating progesterone concentrations.99 Pregnancy rates, although numerically higher in buserelintreated mares, were not significantly different from controls in this particular study.99 Overall, these studies suggest that administration of potent GnRH analogs during early diestrus in mares may positively influence pregnancy rates although the mechanism of action of such treatment remains to be determined. In other species, multiple studies suggest a positive effect of administration of GnRH analogs during early diestrous on fertility100 although this result has not been supported in other reports.101 In addition to administration of exogenous progestins to mares in attempts to prevent embryonic loss, a number of studies have examined the administration of corticosteroids, non-steroidal antiinflammatory drugs (NSAIDs), and immunomodulators to the mare to improve early pregnancy rates or prevent embryonic loss. The administration of immunomodulators or immunostimulants have been used in attempts to improve fertility in mares with persistent endometritis.102, 103 In a recent report, fertility of mares with persistent endometritis was improved after administration of Propionibacterium acnes (EqStimTM ) compared to control mares, and mares treated with P. acnes tended to have a lower rate of pregnancy loss.102 Although the mechanism of action
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is unclear, these studies suggest that in some mares with persistent endometritis, the administration of immunostimulants should be considered as a part of the therapeutic regimen. Likewise, investigators have suggested that administration of either corticosteroids or NSAIDs may also improve fertility or reduce early pregnancy loss in mares. Acute administration of flunixin meglumine at the time of nonsurgical embryo transfer has been suggested as a means to improve pregnancy rates44 although controlled studies are lacking to support a positive effect of NSAID administration on embryo survival. Another study demonstrated an increased early pregnancy rate in barren mares that received prednisolone (0.1 mg/kg, twice daily) for 4 days before and at the time of artificial insemination with frozen semen compared to control mares.104 Wilsher et al. demonstrated that administration of NSAIDs (meclofenamic acid) increased the range of synchrony between donor and recipient mares that resulted in successful embryo transfer.105 Administration of meclofenamic acid to recipient mares beginning at 9 days after ovulation supported the establishment of pregnancy in recipient mares that ovulated up to 3 days prior to the donor mare although the mechanism of action did not appear to act via suppression of prostaglandin F2α -mediated luteolysis.105 Together, these studies suggest that the immune response around the time of mating may alter the subsequent uterine environment and embryo survival, and future studies should be directed towards a better understanding of the mechanisms involved as they relate to embryonic loss in mares. Because of the potential adverse effect of chronic or persistent endometritis on survival of the early embryo in mares, some authors have advocated administration of systemic antibiotics to mares with recurrent early embryonic loss.35 In situations where chronic endometritis has been documented or is suspected, administration of broad-spectrum antibiotics during early gestation may be indicated. No controlled studies exist to support this application; however, anecdotal information suggests that treatment with systemic antibiotics may be helpful in selected cases. Assisted reproductive techniques may offer some benefit to improve the prospects of establishing pregnancy from older, subfertile mares. Unfortunately, embryo recovery from uterine flushes is often reduced, and survival of embryos from such mares is lower after embryo transfer. Likewise, the ability to establish pregnancies from older, subfertile mares also appears to be somewhat reduced after procedures such as oocyte transfer, and more attempts will often be necessary to successfully establish
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pregnancy from these mares. Carnevale et al.106 reported that Day 50 pregnancy rates in recipient mares that received oocytes tended to be lower from donor mares over 20 years of age than from donor mares less than 20 years of age. When mares at the extreme of age ranges were compared, mares less than 15 years of age established pregnancies from 50% of oocytes compared to 16% of oocytes from mares over 23 years of age.106
References 1. Woods GL, Baker CB, Baldwin JO, Ball BA, Bilinski JL, Cooper WL, Ley WB, Mank EC, Erb HN. Early pregnancy loss in broodmares. J Reprod Fertil Suppl 1987;35: 455–9. 2. McDowell KJ, Powell DG, Baker CB. Effect of book size and age of mare and stallion on foaling rate in Thoroughbred horses. J Equine Vet Sci 1992;12:364–7. 3. Morris LHA, Allen WR. Reproductive efficiency of intensively managed Thoroughbred mares in Newmarket. Equine Vet J 2002;34:51–60. 4. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of Flatrace and National Hunt Thoroughbred mares and stallions in England. Equine Vet J 2007; 39:438–45. 5. Vanderwall DK, Newcombe JR. Early embryonic loss. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 374–83. 6. Woods GL, Hillman RB, Schlafer DH. Recovery and evaluation of embryos from normal and infertile mares. Cornell Vet 1986;76:386–94. 7. Vogelsang SG, Vogelsang MM. Influence of donor parity and age on the success of commercial equine embryo transfer. Equine Vet J Suppl 1989;8:71–2. 8. Ball BA, Little TV, Hillman RB, Woods GL. Pregnancy rates at Days 2 and 14 and estimated embryonic loss rates prior to Day 14 in normal and subfertile mares. Theriogenology 1986;26:611–19. 9. Ball BA, Little TV, Weber JA, Woods GL. Viability of Day4 embryos from young, normal mares and aged, subfertile mares after transfer to normal recipient mares. J Reprod Fertil 1989;85:187–94. 10. Brinsko SP, Ball BA, Miller PG, Thomas PGA, Ellington JE. In vitro development of day two embryos obtained from young, fertile mares and aged, subfertile mares. J Reprod Fertil 1994;102:371–8. 11. Brinsko SP, Ball BA, Ignotz GG, Thomas PGA, Currie WB, Ellington JE. Initiation of transcription and nucleologenesis in equine embryos. Mol Reprod Dev 1995;42: 298–302. 12. Van Niekerk CH, Gerneke WH. Persistence and parthenogenetic cleavage of tubal ova in the mare. Onderstepoort J Vet Res 1966;31:195–232. 13. Betteridge KJ, Mitchell D. Direct evidence of retention of unfertilized ova in the oviduct of the mare. J Reprod Fertil 1974;39:145–8. 14. Betteridge KJ, Eaglesome MD, Flood PF. Embryo transport through the mare’s oviduct depends on cleavage and is independent of the ipsilateral corpus luteum. J Reprod Fertil Suppl 1979;27:387–94.
15. Weber JA, Freeman DA, Vanderwall DK, Woods GL. Prostaglandin E2 hastens oviductal transport of equine embryos. Biol Reprod 1991;45:544–6. 16. Ball BA, Brinsko SP, Schlafer DH. Histopathologic examination of the oviduct and endometrium of fertile and subfertile mares. Pferdeheilkunde 1997;13:548–9. 17. Brinsko SP, Ignotz GG, Ball BA, Thomas PGA, Currie WB, Ellington JE. Characterization of polypeptides synthesized and secreted by oviductal epithelial cell explants obtained from young, fertile mares and aged, subfertile mares. Am J Vet Res 1996;57:1346–58. 18. Lantz K, Enders A, Liu I. Possible significance of cells within intraluminal collagen masses in equine oviducts. Anat Rec 1998;252:568–79. 19. Allen WR, Wilsher S, Morris L, Crowhurst JS, Hillyer MH, Neal HN. Laparoscopic application of PGE2 to reestablish oviducal patency and fertility in infertile mares: a preliminary study. Equine Vet J 2006;38:454–9. 20. Zent WW, Liu IKM, Spirito MA. Oviduct flushing as a treatment for infertility in the mare. Equine Vet J Suppl 1993;15:47–8. 21. Armstrong DT. Effects of maternal age on oocyte developmental competence. Theriogenology 2001;55:1303–22. 22. Brinsko SP, Ball BA, Ellington JE. In vitro maturation of equine oocytes obtained from different age groups of sexually mature mares. Theriogenology 1995;44:461–9. 23. Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod 1995; Mono. 1: 209–14. 24. Rambags BPB, van Boxtel DCJ, Tharasanit T, Lenstra JA, Colenbrander B, Stout TAE. Maturation in vitro leads to mitochondrial degeneration in oocytes recovered from aged but not young mares. Anim Reprod Sci 2006; 94:359–61. 25. Torner H, Alm H, Kanitz W, Goellnitz K, Becker F, Poehland R, Bruessow KP, Tuchscherer A. Effect of initial cumulus morphology on meiotic dynamic and status of mitochondria in horse oocytes during IVM. Reprod Dom Anim 2007;42:176–83. 26. Carnevale EM, Bergfelt DR, Ginther OJ. Aging effects on follicular activity and concentrations of FSH,LH, and progesterone in mares. Anim Reprod Sci 1993;31:287–99. 27. Carnevale EM, Bergfelt DR, Ginther OJ. Follicular activity and concentrations of FSH and LH associated with senescence in mares. Anim Reprod Sci 1994;35:231–46. 28. Inskeep EK. Preovulatory, postovulatory, and postmaternal recognition effects of concentrations of progesterone on embryonic survival in the cow. J Anim Sci 2004; 82:E24–39. 29. Woods JA, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic loss rate in mares. Equine Vet J 1990;22:410–15. 30. Huhtinen M, Koskinen E, Skidmore JA, Allen WR. Recovery rate and quality of embryos from mares inseminated after ovulation. Theriogenology 1996;45:719–26. 31. Power MM. The first description of a balanced reciprocal translocation [t(1q;3q)] and its clinical effects in a mare. Equine Vet J 1991;23:146–9. 32. Lear TL, Layton G. Use of zoo-FISH to characterise a reciprocal translocation in a thoroughbred mare: t(1;1 6)(q16;q21.3). Equine Vet J 2002;34:207–9. 33. Lear TL, Lundquist J, Zent WW, Fishback WD, Clark A. Three autosomal chromosome translocations associated with repeated early embryonic loss in the domestic horse (Equus caballus). Cytogenet Genome Res 2008;120:117–22.
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Maximum
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% of mares (n=21)
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0 Serrated granulosa Vascularized theca
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Normal evacuation
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base
base
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100
Septated evacuations 80 % of circumference
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40
20 -11
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80 Pregnant 70 60 Nonpregnant
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Prominence (score)
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0 30 Hours after hCG treatment
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Luteal Gland Development a)
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b)
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Plate 109 (Figure 216.2a-b – Volume 2)
Days after ovulation
Sonogram of uterus and conceptus
Days after Sonogram of uterus ovulation and conceptus
Embryonic vesicle Blastoceole
Capsule
Day 10
Day 18 Inner cell mass
Embryonic vesicle
Ectoderm (trophoblast)
Bilamina (ectoderm & endoderm) Trilamina (ectoderm, mesoderm & endoderm)
Yolk sac
Amniotic sac
Day 12 Embryonic disc
Endoderm
Day 22 Embryo proper
Day 14 Mesoderm
Day 24
Chorion (SomatoPleure)
Day 16 Day 26
Plate 110 (Figure 217.2 – Volume 2)
Chorionic girdle
Allantoic sac
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Day 36
Fetus
Day 40
Umbilical chord
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Plate 110 (Figure 217.2 – Volume 2, continued) Orientation vs Dysorientation of Embryo Proper Ventral position of embryonic pole
Dorsal position of embryonic pole Ma
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Ep Ep
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Day 19
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Ep
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Day 28
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Ma
Uc
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Fetus Fetus Day 45
Plate 111 (Figure 217.3 – Volume 2)
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Plate 119 (Figure 239.3 – Volume 2)
Plate 122 (Figure 268.1 – Volume 2) Plate 121 (Figure 248.4 – Volume 2) C C
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B
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Plate 123 (Figure 268.2 – Volume 2)
Plate 125 (Figure 271.6 – Volume 2)
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A
Plate 126 (Figure 271.7 – Volume 2)
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Plate 134 (Figure 278.5 – Volume 2)
Plate 135 (Figure 278.6 – Volume 2)
Plate 136 (Figure 278.7 – Volume 2)
Plate 137 (Figure 278.8 – Volume 2)
Plate 138 (Figure 278.9 – Volume 2)
Plate 139 (Figure 278.10 – Volume 2)
Plate 140 (Figure 281.2 – Volume 2)
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Embryonic Loss 34. Ball BA, Daels PF. Early pregnancy loss in mares: applications for progestin therapy. In: Current Therapy in Equine Medicine IV. 1997; pp. 531–4. 35. McKinnon AO, Pycock JF. Maintenance of pregnancy. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 389–409. 36. McDowell KJ, Sharp DC, Grubaugh W, Thatcher WW, Wilcox CJ. Restricted conceptus mobility results in failure of pregnancy maintenance in mares. Biol Reprod 1988;39:340–8. 37. Adams GP, Kastelic JP, Bergfelt DR, Ginther OJ. Effect of uterine inflammation and ultrasonically-detected uterine pathology on fertility in the mare. J Reprod Fertil Suppl 1987;35:445–54. 38. Daels PF, Stabenfeldt GH, Hughes JP, Odensvik K, Kindahl H. Effects of flunixin meglumine on endotoxininduced prostaglandin F2a secretion during early pregnancy in mares. Am J Vet Res 1991;52:276–81. 39. Bergfelt DR, Woods JA, Ginther OJ. Role of the embryonic vesicle and progesterone in embryonic loss in mares. J Reprod Fertil 1992;95:339–47. 40. Irvine CHG, Sutton P, Turner JE, Mennick PE. Changes in plasma progesterone concentrations from Days 17 to 42 of gestation in mares maintaining or losing pregnancy. Equine Vet J 1990;22:104–6. 41. Papa FO, Lopes MD, Alvarenga MA, Meira C, Luvizotto MCR, Langoni H, Ribeiro EF, Azedo AE, Bomfim ACM. Early embryonic death in mares: clinical and hormonal aspects. Braz J Vet Res Anim Sci 1998;35:170–3. 42. Allen WR. Luteal deficiency and embryo mortality in the mare. Reprod Domest Anim 2001;36:121–31. 43. McKinnon AO, Squires EL, Carnevale EM, Hermenet MJ. Ovariectomized steroid-treated mares as embryo transfer recipients and as a model to study the role of progestins in pregnancy maintenance. Theriogenology 1988;29(5): 1055–63. 44. Foss R, Crane A. Serum progesterone changes in embryo transfer recipients. Proc Am Assoc Equine Prac 2004; 50:521–4. 45. Gutierrez CV, Riddle WT, Bramlage LR. Serum thyroxine concentrations and pregnancy rates 15 to 16 days after ovulation in broodmares. J Am Vet Med Assoc 2002; 220:64–6. 46. Meredith TB, Dobrinski I. Thyroid function and pregnancy status in broodmares. J Am Vet Med Assoc 2004;224:892–4. 47. Ball BA, Hillman RB, Woods GL. Survival of equine embryos transferred to normal and subfertile mares. Theriogenology 1987;28:167–74. 48. Carnevale EM, Ramirez RJ, Squires EL, Alvarenga MA, Vanderwall DK, McCue PM. Factors affecting pregnancy rates and early embryonic death after equine embryo transfer. Theriogenology 2000;54:965–79. 49. Carnevale EM, Ginther OJ. Relationships of age to uterine function and reproductive efficiency in mares. Theriogenology 1992;37:1101–15. 50. Potter JT, Kreider JL, Potter GD, Forrest DW, Jenkins WL, Evans JW. Embryo survival during early gestation in energy deprived mares. J Reprod Fertil Suppl 1987;35: 715–16. 51. Gilbert RO, Marlow CH. A field study of patterns of unobserved foetal loss as determined by rectal palpation in foaling, barren and maiden thoroughbred mares. Equine Vet J 1992;24:184–6.
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52. Kenney RM, Ganjam VK. Selected pathological changes of the mare uterus and ovary. J Reprod Fertil Suppl 1975; 23:335–9. 53. Kenney RM. Cyclic and pathologic changes of the mare endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1987;172:241–62. 54. Leidl W, Kaspar B, Kahn W. [Endometrial cysts in the mare. 2. Clinical studies: occurrence and significance]. Tierarztl Prax 1987;15:281–9. 55. Grueninger B, Schoon HA, Schoon D, Menger S, Klug E. Incidence and morphology of endometrial angiopathies in mares in relationship to age and parity. J Comp Path 1998;119:293–309. 56. Eilts BE, Scholl DT, Paccamonti DL, Causey RC, Klimczak JC, Corley JR. Prevalance of endometrial cysts and their effect on fertility. Biol Reprod 1995;Mono 1:527–32. 57. Tannus RJ, Thun R. Influence of endometrial cysts on conception rate of mares. Zentralbl Veterinarmed A 1995; 42:275–83. 58. Kaspar B, Kahn W, Laging C, Leidl W. [Endometrial cysts in the mare. 1. Post-mortem studies: occurrence and morphology]. Tierarztl Prax 1987;15:161–6. 59. Blikslager AT, Tate LP Jr, Weinstock D. Effects of neodymium:yttrium aluminum garnet laser irradiation on endometrium and on endometrial cysts in six mares. Vet Surg 1993;22:351–6. 60. Griffin RL, Bennett SD. Nd:YAG Laser photoablation of endometrial cysts: A review of 55 cases (2000–2001). Proceedings of the Annual Convention of the American Association of Equine Practitioners 2002;48:58–60. 61. Steiner JV. Mare reproductive tract. In: Slovis NM (ed.) Atlas of Equine Endoscopy. St Louis: Mosby, 2004; pp. 183–206. 62. Bartmann CP, Stief B, Schoon HA. [Thermal injury and wound healing of the endometrium subsequent to minimally invasive transendoscopic use of Nd:YAG-laserand electrosurgery in horses]. Dtsch Tierarztl Wochenschr 2003;110:271–80. 63. Stanton MB, Steiner JV, Pugh DG. Endometrial cysts in the mare. J Equine Vet Sci 2004;24:14–19. 64. Platt H. Aetiological aspects of abortion in the Thoroughbred mare. J Comp Pathol 1973;83:199–205. 65. Moberg R. The occurrence of early embryonic death in the mare in relation to natural service and artificial insemination with fresh or deep-frozen semen. J Reprod Fertil Suppl 1975;23:537–9. 66. Chevalier-Clement F. Pregnancy loss in the mare. Anim Reprod Sci 1989;20:231–44. 67. VanRaden PM, Miller RH. Effects of nonadditive genetic interactions, inbreeding, and recessive defects on embryo and fetal loss by seventy days. J Dairy Sci 2006;89: 2716–21. 68. King WA, Linares T, Gustavsson I. Cytogenetics of preimplantation embryos sired by bulls heterozygous for the 1/29 translocation. Hereditas 1981;94:219–24. 69. Long SE. Tandem 1;30 translocation: a new structural abnormality in the horse (Equus caballus). Cytogenet Cell Genet 1996;72:162–3. 70. Love CC, Thompson JA, Lowry VK, Varner DD. Effect of storage time and temperature on stallion sperm DNA and fertility. Theriogenology 2002;57:1135–42. 71. Baumber J, Ball BA, Linfor JJ, Meyers SA. Reactive oxygen species and cryopreservation promote DNA fragmentation in equine spermatozoa. J Androl 2003;24: 621–8.
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Mare: Problems of Pregnancy
72. Sakkas D, Urner F, Bizzaro D, Manicardi G, Bianchi PG, Shoukir Y, Campana A. Sperm nuclear DNA damage and altered chromatin structure: effect on fertilization and embryo development. Hum Reprod 1998;13:Suppl 4:11–19. 73. Fatehi AN, Bevers MM, Schoevers E, Roelen BAJ, Colenbrander B, Gadella BM. DNA damage in bovine sperm does not block fertilization and early embryonic development but induces apoptosis after the first cleavages. J Androl 2006;27:176–88. 74. Antczak DF, Miller J, Remick LH. Lymphocyte antigens of the horse II. Antibodies to ELA antigens produced during equine pregnancy. J Reprod Immunol 1984;6: 283–97. 75. Bacon SJ, Ellis SA, Antczak DF. Control of expression of major histocompatibility complex genes in horse trophoblast. Biol Reprod 2002;66:1612–20. 76. Antczak DF, Allen WR. Maternal immunologic recognition of pregnancy in equids. J Reprod Fertil Suppl 1989; 37:69–78. 77. Allen WR, Short RV. Interspecific and extraspecific pregnancies in equids: anything goes. J Hered 1997;88:384–92. 78. Mathias S, Allen WR. Immunization with stallion lymphocytes for treatment of recurrent spontaneous abortion in Thoroughbred mares. J Reprod Fertil Suppl 2000; 56:645–50. 79. Mortensen C, Choi YH, Hinrichs K, Ing N, Kraemer D, Vogelsang S, Vogelsang M. Effects of exercise on embryo recovery rates and embryo quality in the horse. Anim Reprod Sci 2006;94:395–7. 80. Henneke DR, Potter GD, Kreider JL. Body condition during pregnancy and lactation and reproductive efficiency of mares. Theriogenology 1984;21:897–909. 81. Newcombe JR, Wilson MC. Age, body weight, and pregnancy loss. J Equine Vet Sci 2005;25:188–94. 82. van Niekerk FE, Van Niekerk CH. The effect of dietary protein on reproduction in the mare. VII. Embryonic development, early embryonic death, foetal losses and their relationship with serum progestagen. J S Afr Vet Assoc 1998;69:150–5. 83. van Niekerk FE, Van Niekerk CH. The effect of dietary protein on reproduction in the mare. VI. Serum progestagen concentrations during pregnancy. J S Afr Vet Assoc 1998;69:143–9. 84. Ginther OJ. Embryonic loss. In: Ultrasonic Imaging and Animal Reproduction; Book 2 Horse. Cross Plains, WI: Equiservices, 1995; pp. 175–83. 85. Ginther OJ. Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains, WI: Equiservices, 1986; pp. 542–3. 86. Newcombe JR. Embryonic loss and abnormalities of early pregnancy. Equine Vet Educ 2000;12:88–101. 87. Newcombe JR. The relationship between the number, diameter, and survival of early embryonic vesicles. Pferdeheilkunde 2004;20:214–20. 88. Vanderwall DK, Squires EL, Brinsko SP, McCue PM. Diagnosis and management of abnormal embryonic development characterized by formation of an embryonic vesicle without an embryo in mares. J Am Vet Med Assoc 2000;217:58–63. 89. Silva LA, Ginther OJ. An early endometrial vascular indicator of completed orientation of the embryo and the role of dorsal endometrial encroachment in mares. Biol Reprod 2006;74:337–43. 90. McKinnon AO, Tarrida del Marmol Figueroa S, Nobelius AM, Hyland JH, Vasey JR. Failure of hydroxyproges-
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terone caproate to maintain pregnancy in ovariectomised mares. Equine Vet J 1993;25:158–60. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32: 83–5. Shideler RK, Squires EL, Voss JL, Eikenberry DJ, Pickett BW. Progestagen therapy of ovariectomized pregnant mares. J Reprod Fertil Suppl. 1982;32:459–64. Ball BA, Wilker C, Daels PF, Burns PJ. Use of progesterone in microspheres for maintenance of pregnancy in mares. Am J Vet Res 1992;53:1294–7. Knowles J, Squires EL, Shideler RK, Tarr SF, Nett TM. Relationship of progesterone to early pregnancy loss in mares. J Equine Vet Sci 1993;13:528–33. Vanderwall DK, Marquardt, JL, Woods GL. Use of a compounded long-acting progesterone formulation for equine pregnancy maintenance. J Equine Vet Sci 2007;27: 62–6. Barber JA, Troedsson MH. Mummified fetus in a mare. J Am Vet Med Assoc 1996;208:1438–40. Newcombe JR, Martinez TA, Peters AR. The effect of the gonadotropin-releasing hormone analog, buserelin, on pregnancy rates in horse and pony mares. Theriogenology 2001;55:1619–31. Glazar BS, McCue PM, Bruemmer JE, Squires EL. Deslorelin on Day 8 or 12 postovulation does not luteinize follicles during an artificially maintained diestrous phase in the mare. Theriogenology 2004;62:57– 64. Kanitz W, Schneider F, Hoppen HO, Unger C, Nurnberg G, Becker F. Pregnancy rates, LH and progesterone concentrations in mares treated with a GnRH agonist. Anim Reprod Sci 2007;97:55–62. Peters AR. Veterinary clinical application of GnRH questions of efficacy. Anim Reprod Sci 2005;88:155–67. Bartolome JA, Melendez P, Kelbert D, Swift K, McHale J, Hernandez J, Silvestre F, Risco CA, Arteche ACM, Thatcher WW. Strategic use of gonadotrophin-releasing hormone (GnRH) to increase pregnancy rate and reduce pregnancy loss in lactating dairy cows subjected to synchronization of ovulation and timed insemination. Theriogenology 2005;63:1026–37. Rohrbach BW, Sheerin PC, Cantrell CK, Matthews PM, Steiner JV, Dodds LE. Effect of adjunctive treatment with intravenously administered Propionibacterium acnes on reproductive performance in mares with persistent endometritis. J Am Vet Med Assoc 2007;231:107–13. Rogan D, Fumoso E, Rodriguez E, Wade J, Sanchez Bruni SF. Use of a mycobacterial cell wall extract (MCWE) in susceptible mares to clear experimentally induced endometritis with Streptococcus zooepidemicus. J Equine Vet Sci 2007;27:112–17. Dell’Aqua JA, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Melo CM. Modulation of acute uterine inflammatory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94:270– 3. Wilsher S, Kolling M, Allen WR. Meclofenamic acid extends donor-recipient asynchrony in equine embryo transfer. Equine Vet J 2006;38:428–32. Carnevale EM, Coutinho da Silva MA, Panzani D, Stokes JE, Squires EL. Factors affecting the success of oocyte transfer in a clinical program for subfertile mares. Theriogenology 2005;64:519–27.
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Abortions and Stillbirths: A Pathologists Overview Katherine E. Whitwell
For an owner, a pregnancy failure in a broodmare represents a series of losses – of expectations, of finances (no return from a year’s management, veterinary fees and stud fees) and of potential genetic improvement. In addition it can jeopardize the mare’s immediate and future health and, depending on the nature of the loss, may become a real or potential source of pregnancy loss in cohort mares. It is therefore vitally important that all losses are assessed so appropriate treatment can be given, and other current and future losses prevented. Even good surveillance of both the pregnant mare and her environment is not always able to identify pregnancy loss. With early fetal losses there may well be no aborted material found, either within the mare or after expulsion into her environment. Investigations to evaluate possible causes therefore may need to rely entirely on assessment of data acquired direct from clinical examinations and screening of the mare herself. After 4 months of gestation, aborted material is more likely to be found, particularly if stud staff are vigilant in observing the mare (her perivaginal and mammary regions). In stable bedding small, usually autolysed, conceptual remnants are easily missed. In paddocks too, particularly large ones, aborted material may go unnoticed and become totally or partially predated and weathered so as to be unidentifiable. Whenever aborted material is found, no matter how incomplete or decomposed, it is essential that a detailed examination be performed. This is preferably done at a lab that deals regularly with equine material and has facilities to carry out gross fetal and placental examinations, histology, and microbiology (including virology and bacteriology). However if the clinician prefers to conduct the necropsy then the following aims and routines are recommended.
Aims of an abortion investigation The main aim is to arrive at an exact and positive etiological diagnosis. If this proves to be impossible – either because the aborted material is of poor quality or incomplete, or because no diagnostic lesions are identified, then it is useful if at least the investigation can exclude certain conditions known to cause pregnancy loss (e.g. equine herpesvirus, other infective fetal conditions considered relevant, placental infections, umbilical cord-related problems, twinning, and severe developmental disease.)
Suggested routine for conducting an abortion investigation Precautions and procedures Different countries have differing legal requirements when dealing with certain infectious conditions. Each year a ‘Codes of Practice’ booklet1 is published by the UK’s Horserace Betting Levy Board (HBLB), which provides an annual update on voluntary recommendations for owners and veterinarians to assist in the control of certain infectious diseases, including several that can compromise pregnancy and/or broodmare reproductive health, including contagious equine metritis (CEM), Klebsiella pneumoniae, Pseudomonas aeruginosa, equine herpesvirus (EHV), and equine viral arteritis (EVA). The Codes are common to the UK, Ireland, France, Germany and Italy: it is claimed that their implementation has resulted in a significant decrease in disease outbreaks. In general when a mare aborts or a neonate dies there is a need always to:
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wear disposable gloves when handling a mare’s genitalia and aborted material;
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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use leakproof containers for moving abortuses/foals; disinfect potentially contaminated sites, equipment, and vehicles; isolate the mare and regulate horse movements on and off the site pending the results of the investigations.
the use of microbiology and histology. Routine investigations should never be curtailed just because an abortion appears grossly to have one condition. Some aborted fetuses have been known to suffer from two conditions, for example EHV abortion in a twin pregnancy. A recommended necropsy routine
Specific guidelines are given in the Codes of Practice for control of these conditions. For more detailed recommendations the reader should consult the booklet. Obtain a clinical history This is preferably acquired prior to a postmortem being carried out. In cases of abortion or stillbirth a clinical history can be of considerable help in evaluating the significance of the postmortem findings. Details concerning the gestation length, the mare’s past and present general and reproductive health, her vaccination history and any treatments recently given, her diet and management, travels abroad, the parturition, details of the environment and the presence of other pregnancy losses on the premises are all useful. Details of any recent stressful or traumatic events may be useful. General approach
Early pregnancies These may be retained in utero for a period after fetal death resulting in the tissues becoming very autolysed or decomposed, and often unsuitable for histological evaluation. However, even in severely predated or decomposed abortuses samples of some fetal organs (thymus, lung, liver, spleen) and often allantochorion may be located and screened for EHV infection using polymerase chain reaction (PCR). An apparently useless, fetid, shapeless lump of allantochorion can sometimes be expanded by the slow instillation of water or air into the membrane (via water hose or enema pump) so that morphological features of two horns and the body become identifiable.
Later pregnancies To optimize the chance of making a positive diagnosis both the fetus and the whole of the placenta should be examined conscientiously at necropsy, taking objective measurements from both. All observations and measurements are preferably noted directly onto forms at the time – so as to be sure they are not overlooked or confused with other investigations. Digital photos are an excellent format to record both fetal and placental features. Twinning and most cases of cervical placentitis are usually identified grossly without difficulty but all investigations benefit from
Fetal external examination If the fetus is still attached to the placenta, cut the cord, weigh the fetus, and measure the crown–rump length. Spend time evaluating the fetus externally – the degree of rigor mortis, the amount of autolysis, the coat length, and color of the mucosae. Check for congenital abnormalities (deformations and malformations) including active palpation of the navel for umbilical hernia; also check the perineum for meconium staining, and the nares for froth after breathing. Forcibly extend bent limbs to determine whether there is a true contracture or just rigor mortis. Even a small amount of knee contracture may cause stillbirth due to prolongation of second-stage labor, placental detachment, and asphyxiation.
Fetal internal examination A blood sample may be taken. The best view of the viscera (with spleen uppermost) is obtained when the fetus is laid on its right side, the left limbs are reflected over the back and the left abdominal and thoracic walls are opened and reflected. If the left abdominal ventral incision is parasagittal it enables the state of the intact umbilical vein, the navel and bladder apex to be assessed – in particular in stillbirths to determine whether the vein has contracted (indicating that the fetus was alive for a period during second-stage labor.) The volume and color of serosal fluids are established immediately, before any iatrogenic bleeding is caused. To avoid contaminating the organs it is preferable to carry out the fetal microbiological sampling before handling or removing viscera. For EHV PCR, disposable forceps and scalpel are used to place samples of liver, spleen, lung and thymus into virus transport medium (VTM). For Leptospira, PCR samples of fetal kidney (cortex and medulla) and vitreous humor are placed in transport medium. Two bacteriology swabs (Amies charcoal) are taken aseptically from fetal liver or heart blood, and lung or stomach. After viewing them in situ, first the abdominal and then the thoracic viscera are removed individually, inspected and sliced into thin slices, one or more being placed in 10% formalin fixative for subsequent histological examination. Gross examination should include inspection of the spleen, liver, kidneys, navel, bladder, gastric contents, small and large
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bowel (checking for asphyxial intussusceptions in stillbirths), opening the lower airways to check for air inhalation, and checking the heart for anomalies and recent bruising. The distal ends of the ribs inside and outside the thorax should be checked for evidence of recent bruising or fracture due to birth trauma. After splitting the skull, the pharynx and brain are examined. The fetal organs harvested for histology should include liver, spleen, kidney, adrenal, lung, thymus, thyroid, and sometimes brain. Finally the main limb joints, particularly those of the upper limbs, are opened to check for hemarthrosis. To enable optimal control measures for infections such as EHV a rapid turnaround for the results of screening is advisable (PCR, histology, immunostaining).
Placental examination This is a vital part of the diagnostic investigation and is best conducted on a large, elevated, well-lit, flat board. Four pieces of chorion including a cervical fold, and a sample from amnion and cord surface, are placed in fixative for histology. The chorion is spread out into an F shape for inspection of shape and integrity: both surfaces should be examined, starting with the side outermost. Using sterile instruments at least four samples of chorion should be sampled into VTM for EHV PCR (if Leptospira screening is required, samples of chorion and amniotic cord surface are placed into VTM for PCR). On the chorion’s allantoic side the cord attachment site is identified (normally at the base of the pregnant or non-pregnant horn) and a note made of the vascular pattern.
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In type I the cord horn is the pregnant horn at term: one artery goes to the pregnant horn only and the other supplies the body and the non-pregnant horn. In type II the cord horn is not the pregnant horn at term: one artery supplies the non-pregnant horn and the other supplies the pregnant horn and body. In type III one artery supplies both horns and the body, the other much smaller artery supplies only a small area on the convexity at the base of one horn (K.E. Whitwell, unpublished, cited in ref. 2).
If an umbilical cord problem is suspected the vascular examination can include the opening of some vessels with fine scissors, to identify visible intravascular thrombi. Inspection of the villous side of the chorion commences with bacteriological swabbing of cervical fold surfaces (and any site suspected of placentitis). Histological samples should include a cervical star fold. The whole villous surface should be scrutinized. Left or right sidedness of the pregnant horn can be determined by horn tip elevation and
Figure 240.1 Pregnant horn determination. With the villous side outermost and the body towards the examiner, the horn tips are elevated. The larger convexity is at the base of the pregnant horn (the left horn in this placenta.)
shape assessment with the convexity at the base of the pregnant horn being larger and hanging downwards with a more distinct ventral bulge (Figure 240.1). It is the tip of the non-pregnant horn that gets retained in utero: if missing the clinician should be notified in case it is retained. The amnion is checked for old tears, focal lesions or opacity, and for meconium staining. Umbilical cord: The total and amniotic cord lengths are measured after untwisting any twists. In stillbirths the presence or absence of the two (normal) umbilical artery worm-like projections from the cord end is noted. (Their presence suggests that the fetus was alive when the cord ruptured.) The amniotic cord surface, stroma, urachus, and vessels are inspected, particularly at any indented twist sites. In the allantoic cord similar checks for vascular integrity are made and the yolk sac identified after incising into the infundibulum.
Diagnostic trends in abortion statistics Many individual causes of pregnancy loss are discussed in detail by other authors in this textbook. Retrospective surveys and overviews of the causes of abortion and stillbirth are periodically published by pathologists at diagnostic labs in many countries. These highlight how the diagnoses made have changed over the decades, the causal trends being influenced by a number of disparate factors, including:
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Technological developments in clinical medicine. For example, the increasing clinical use of ultrasonographic scanning of the pregnant uterus
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to facilitate twin reduction has been directly responsible for the incidence of twin abortions falling dramatically from being the commonest diagnosis made, to being 6% of the diagnoses (see Chapter 241). Increased use of vaccines for control of EHV. This is believed to have lowered the incidence of EHV abortions. Developments in laboratory diagnostic methods. PCR techniques and immunostaining have revolutionized the speed and accuracy of diagnosis. For example, placenta-positive/fetus-negative EHV cases have now been confirmed3 – perhaps solving the riddle of some EHV fetus-negative abortions seen at the start of EHV outbreaks. Strong opinions held by eminent pathologists. For example, neither Leo Mahaffey in Newmarket, UK,4 or Jim Rooney in Lexington5 believed umbilical cord torsion was a cause of fetal death. In subsequent years mounting gross and histological evidence confirmed that a diagnosis of fetal death from vascular compromise and thrombosis associated with long and twisted cords was a valid diagnosis. With supportive comparative input from human pediatric pathology the diagnosis became increasingly accepted, so that recent UK surveys
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reveal it is now the commonest cause of diagnosed loss (see below). Environmental changes influencing insect populations. A massive caterpillar multiplication heralded the 2001 Lexington outbreak of mare reproductive loss syndrome (MRLS). Research into MRLS and also amnionitis/funisitis cases in Australia revealed that ingestion of caterpillar spines from a contaminated environment, and development of intestinal granulomata, probably underlie the abortifacient nature of the insult (see Chapter 248).
Summary list of causes of abortion and stillbirth As outlined above, geographical location, clinical and laboratory technologies used, and the current perceptions and deductions of investigators all strongly influence published data. Table 240.1 summarizes many of the infective and non-infective published causes of loss over the past few decades. What today may be a new cause of concern and loss may assume very different importance in another geographical area, in another breeding season. The long list of infective agents highlights the fact that most of the organisms cultured from the mare’s non-pregnant
Table 240.1 Summary of recorded causes of equine abortion and stillbirth A. Infective. Grouped alphabetically as a) Common in some geographic areas, and b) Occasional or Rare. Viral a) Equine herpesvirus EHV1 Equine viral arteritis EVA b) Equine herpesvirus EHV4 Equine Infectious Anaemia Bacterial
a) Streptococcus zooepidemicus Escherichia coli Pseudomonas aeruginosa Leptospira sp Nocardioform actinomycetes (egs Nocardia sp; Amycolatopsis sp; Crosiella equi; Rhodococcus rubropertinctus Cellulosimicrobium cellulans) b): Actinobacillus equuli Aeromonas hydrophila Brucella sp. Cellulosimicrobium cellulans Chlamydia psittaci Citrobacter sp Corynebacterium equi Corynbacterium pseudotuberculosis Enterobacter aerogenes Enterobacter agglomerans Erwinia herbicola Flavobacterium sp Gemella haemolysans Klebsiella pneumoniae Mycoplasma sp (Continued)
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Table 240.1 (Continued) Proteus sp Pseudomonas sp Salmonella abortus-equi Salmonella typhimurium Staphylococcus aureus Streptococcus equisimilis Streptococcus faecalis Taylorella equigenitalis Fungal
a) Aspergillus fumigatus Mucor sp b): Absidia sp (eg A.corymbifera; A.ramose) Acholelplasma sp Allescheria boydii Aspergillus sp (eg A.terreus; A.nidulans; A.ustus) Bipolaris specifera Candida sp (incl C.rugosa) Coccidiodes immitis Cryptococcus neoformans Histoplasma capsulatum Mycoplasma sp Neorickettsia (previously Ehrlichia) risticii (Potomac horse fever) Rhizomucor pusillus Thermomyces lanugino
Protozoal
b) Encephalitozoon cuniculi Piroplasmosis (Theileria equi and Babesia caballi)
B. Non-infectiive conditions which can lead to pregnancy loss Multiple births Twins and, rarely, triplets Umbilical cord problems Vascular compromise. Excessive twisting in a long cord Tight wrapping of the cord around the fetus or a limb Cord strangulation by an excessively long allantoic pouch Cord strangulation by a large yolk sac remnant Very short cord. Can lead to intrapartum stillbirth Neoplasia – cord angioma Placental problems Clearly defined sites of ischaemic necrosis, usually at the cervical pole. Body pregnancy Bicornual pregnancy Hydrops – amniotic or allantoic Ruptured amnion Maternal problems
Severe general illness (colic; pyrexia, starvation; neoplasia) Uterine pathology Congenitally malformed uterus Cervical incompetence Endometrial infarction Endometrial scarring, degeneration, ageing change (poor placental support) Trans-uterine adhesions Severe trauma to the pregnant uterus (fractured fetal limbs; ruptured amnion) Parturition problems leading to stillbirth (fractured pelvis; ruptured uterus; premature placental separation)
Fetal problems
Chromosomal abnormality Severe congenital malformations (of head, spine, body wall, heart, intestinal tract, limbs) Severe congenital deformations (of limbs or spine leading to intrapartum stillbirths) Abnormal presentation leading to stillbirth Thyroid disease Congenital fetal neoplasia
Environmental problems
Fescue toxicosis Spiny caterpillar-based losses (MRLS in Lexington, USA; Amnionitis EAFL in Australia)
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reproductive tract, as well as many environmental organisms have the capacity to become opportunistic pathogens, particularly if the host is compromised.
Some specific causes of loss Important infective and non-infective causes of abortion or stillbirth are also discussed in chapters that deal with the agent or cause specifically (see Chapters 241, 242, 245, 246, 247, 248, 249 and 250). Umbilical cord problems Evaluation of umbilical cords from a population of Thoroughbred (TB) mares, selected for the normality of the foaling and of the foal born, provided the standards by which to judge cords from other notso-normal foalings and from abortions.6 Variations in total cord length (95% lay between 36 and 83 cm), twisting and urachal integrity were recorded. This knowledge was subsequently used to assist in the assessment of cord abnormalities in abortions and stillbirths.
Figure 240.2 Amniotic umbilical cord that has been tightly twisted. Untwisted it shows stromal hemorrhage, edema, pallor at a tight twist, and stretch marks in the surface covering.
Vascular occlusion Fetuses dying in utero due to vascular obstruction in the umbilical cord are not usually aborted immediately, so the tissues display variable degrees of (sterile) autolysis when aborted. They tend to have non-specific changes of distended abdomen, bloody fluid in serosal cavities and sometimes edema around the navel. The gross evidence to support the diagnosis lies in the cord and chorion. Cord lesions associated with tight twists include local intimal or medial vascular wall tears and associated intramural and stromal hemorrhages, local edema even sufficient to cause ‘stretch tears’ at the amniotic surface, and small aneurysmal dilatations (Figure 240.2). Fetuses often have elongated navel stalks – possibly from chronic cord tension (Figure 240.3). Occasionally thrombi can be seen macroscopically within chorioallantoic vessels opened with scissors (Figure 240.4). Histological evidence supports a diagnosis of poor/failing circulation in the chorion: sections show degenerative changes in some stromal blood vessel walls (intimal and smooth muscle degenerative change), microthrombosis (luminal fragments as well as obstruction and mineralization), and characteristically villus core necrosis and patchy mineralization, but with an intact villous epithelium (presumably maintained by perfusion across the still in-contact feto-maternal junction). Statistical reviews from both diagnostic labs in Newmarket, UK (1988–19977 and 1996–20018 ), showed that umbilical cord torsions
Figure 240.3 Aborted fetus displaying an elongated navel stalk, possibly resulting from cord tension.
Figure 240.4 Large thrombi obstructing chorioallantoic blood vessels in a twisted cord abortion.
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Ischemic necrosis of defined sites of allantochorion
Figure 240.5 Allantoic surface of caudal body of chorion. Clear line demarcates border of ischemic necrotic tissue.
resulting in vascular compromise and fetal death comprised the most frequent diagnosis (35.7% of abortion and neonatal deaths, and 46.2% of pre-term abortions, respectively). In countries with a high incidence of other conditions causing pregnancy failure, cord problems have not ranked so highly. Ultrasonographic scanning indicates that up until the time of closure of both uterine horns at month 8 (when fetal hindlimbs become lodged up what then becomes known as the ‘pregnant horn’), the fetus can move within the uterus, both within the amniotic sac and, in the earlier stages as a fetal-amniotic unit within the allantochorion.9 Fetuses are remarkably active and their rotational movements can result in 360-degree twisting of the amniotic and/or the allantoic parts of the cord. What governs cord lengths has not been definitively established although a statistical analysis of data from normal foals showed both genetic (male foals had longer cords than females, and there was a sire effect) and environmental influences (cord length increased with age of mare). 10 Cord lengths were statistically longer in cases aborted due to vascular compromise than in abortions associated with infective conditions. The reason for excessive twisting has not been fully explained. Medical pathologists have highlighted similar vascular problems associated with long-cordedness and cord twisting in human pregnancy losses.11 Their studies indicate that the associated vascular compromise does not always have a fatal outcome, some live births having evidence of long-term tissue damage. In equine fetuses, sudden and fatal vascular compression/cord torsions tended to occur before signs of urinary retention (urachal dilatation) developed at twist sites (51/2 to 71/2 months). Where urachal dilatations had developed the pregnancies tended to continue longer (up to 6–10 months).10 It is quite feasible that some equine live births may have subclinical tissue defects arising from degrees of poor placental perfusion due to cord twist vascular compromise.
Clearly defined areas of full-thickness necrosis, mainly involving the caudal body/cervical pole (Figure 240.5), but occasionally also involving other sites and even amnion, are also seen in abortions with excessively long cords. The abortions occur throughout gestation. It has also been noted in a twin. The reason for the lesions is not apparent. The necrotic site may eventually leak fetal fluids and has been known to become secondarily infected via the cervix. Loss of functional chorion eventually leads to fetal growth retardation.
Body pregnancy This is a rare and little understood chronic condition usually causing abortion of a growth-retarded fetus at 8 to 10 months. In normal term TB mares, 95% of chorions have a maximum mid-body width of 52 cm.6 In body pregnancy the horns collapse and shrink and their lumen is reduced or obliterated so they cannot contain the fetal hindlimbs (Figure 240.6). The fetus occupies the uterine body only, which is wider than normal (up to 65 cm) to accommodate it. Accompanying defects may include a very long cord, areas of cervical necrosis, thickened
Figure 240.6 Chorion from a body pregnancy abortion with narrow horns and a wide body.
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Figure 240.7 Two urachal dilatations near the end of an umbilical cord.
amnion and allantoitis. The cord attachment site is usually normal.
equine twins are regarded as non-identical and dizygotic. Before routine use of ultrasonography, abortion diagnostic submissions from TB breeding areas were dominated by twinning. Platt, based at a lab in Newmarket, UK, reported in 1973 that 29% of the 1969–1971 abortion submissions resulted from twinning.13 Without the incursion of any new emerging abortigenic condition, when the same lab published a 1988–1997 review, twinning had fallen dramatically to 6%.7 The ready availability of so many twin abortions provided the opportunity in the early 1970s to study 62 sets of twins and their membranes.14 The potential for twinning is high in TBs compared to some other breeds but their ability to produce live and viable twin foals is poor. Both foals survived to 2 weeks in only 14.5% of the 62 pregnancies, and one foal survived to 2 weeks in 21%.14 Draft breeds, with a larger uterine capacity, are reputedly more successful.
Placental arrangements in aborted twins Urachal obstruction The urachus conveys urine from the bladder to the allantoic cavity. As it has no smooth muscle coat it is vulnerable to local dilatation by urine at sites of obstruction, for example as caused by twisting (Figure 240.7). Many normal foals are born with one or several urachal dilatations along the amniotic cord but if an obstruction is marked, urachal dilatation can be very large and even rupture into the amniotic cavity. A proximal urachal obstruction can lead to a large dilated fetal bladder and contribute to pervious urachus in the foal, and potentially to ruptured bladder.
Short cords These are rare and said to be associated with fetal inactivity. If a mare has a long uterine body the tension on a short cord may cause it to rupture prematurely during parturition, with a risk of stillbirth.
Abnormal cord attachment site This reflects an abnormal uterine implantation site of the conceptus: the cause is not known. Instead of the normal site at the base of one horn, the site can be up one horn, in the rostral ventral aspect of the uterine body, or more caudally in the uterine body. The subject has recently been reviewed.12 Some pregnancies are unaffected but in others there are associated problems, some related to excessive cord length (e.g. of 170 cm). Twinning Based on double ovulations, differences in fetal sex, markings and possession of separate placentae,
The twin fetal size achieved is proportional to its gestational age and its surface area of viable, villous chorion. The twin placenta study14 defined three main placental patterns: in type A (79%) the chorion of larger twin 1 occupies the uterine body and one horn. In the less common type B (11%) the twin placentae are of similar size, each occupying one horn and one (longitudinal) half of the uterine body. Type C (10%) is similar to type A but the larger twin 1’s placenta also extends up into the second horn leaving twin 2 only about 15% of the total surface area. In type C when the small twin 2 dies it becomes dessicated and mummifies, and deeply invaginated into twin 1’s placenta. For a given gestation, type A twin 1 fetuses are the largest, and type C twin 2 fetuses the smallest (see Chapter 241). A report in 198015 identified a type C variant: instead of twin 2 conceptual remnants invaginating into twin 1’s chorion, they tracked external to it, presumably under gravity, from the horn tip down to the horn base. Three other ‘atypical’ twin placentae were also recorded.15 Twin 1’s chorion filled both horns and the body whilst remnants of the twin 2 s occupied small sites between the uterine wall and twin 1’s chorion in the rostral, mid and caudal body, respectively, the latter close to the cervical pole. They could easily have been overlooked. In these cases and in type C the twin 1 foals were able to achieve single foal size.
The in-contact avillous area between twin 1 and twin 2 chorions There is much variation in the gross and histological appearances at this site. The two chorions abut, and in types A and C variable degrees of (intact) chorionic invagination occur (generally twin 2’s into twin 1’s).
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Figure 240.8 Separated type A twin placentae showing an extreme form of pseudo-anastomosis. Fresh erythrocytes from the (lower) chorion 1 are reperfusing a band of the mummifying chorion 2 adjacent to the in-contact area.
On inspection, some twin chorions fall or pull apart with ease and show a smooth, opaque/translucent surface with or without a little local hemorrhage. Others cannot be separated without tearing, due to bridging adhesions, and sometimes cystic areas and calcification. Blood vessels can sometimes be seen crossing the site. Indian ink injection confirmed crossover of blood from one side to the other, but it was not clear whether these were anastomoses or only pseudo-anastomoses.14 In Type C, after death and mummification of twin 2 the apparent extension of twin 1’s chorion further up into twin 2’s horn space might be construed as a form of survival strategy for twin 1. Another phenomenon/survival strategy occasionally noted is when one twin has been dead for some time and the other is relatively fresh. At the in-contact site blood vessels can be seen to have crossed from twin 1’s chorion into the dead twin 2’s in-contact (autolysed) chorion: the vascularized tissue appears as viable islands amongst very autolysed tissue. In some twins this neoperfusion extends to the edges of the in-contact area and beyond so that a narrow band of twin 2’s villous chorion appears viable. Figure 240.8 shows an extreme form of this pseudoanastomosis. Histological evidence in some twins of periportal inflammatory infiltration in the liver, plasma cells in infiltrates at the chorion in-contact interface, and small quantities of gamma-globulin in twin fetal serum support the notion that an adverse inter-twin immunological response may contribute to some twin deaths.
Anomalous vascular patterns in term single placentae The normal umbilical cord attachment site is at a horn base (the cord horn) and defines where the
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embryonic vesicle becomes fixed at ∼ day 16. At term the attachment site is sometimes not in the same horn as the horn containing the fetal hindlegs (the pregnant horn). The vascular patterns of the vessels extending over the chorionic allantoic surface to and from the umbilical cord vessels have been described earlier in this chapter. In the uncommon third anomalous pattern, type III, it was noted that scanning records showed that a twin reduction had been performed in many of them. More structured studies are required to determine if the deprivation hypothesis16 of events around implantation time can account for the distinctive type III pattern developing.
Placentitis This term usually refers to inflammation of the chorioallantois caused by an infective agent, but it often also involves the amnion and umbilical cord. The inflammatory response can be slight – inflammatory cells entering the villi, stromal edema, separation from the endometrial crypts – to moderate – more extensive villitis, extension of inflammatory material into the extraembryonic coelom (EEC), build-up of endometrial exudates and secretions at the surface, and reactive change in the stroma and allantoic surface – to severe – with loss of villi, ischemia, fibrosis, and degrees of adenomatous hyperplasia on the allantoic surface. The amnion also can be variably affected by inflammatory change, and have areas of ischemic necrosis and calcification. The surface of the umbilical cord may show variable degrees of inflammation (funisitis) with inflammatory debris at the surface. A large number of infective agents have been implicated (see Table 240.1), many being opportunistic or environmental invaders. Within and between geographical centers there are annual variations in numbers and types of placentitis. A pregnant mare’s clinical history – previous abortions, vaginal discharge, inappropriate mammary development, dripping of milk, perineal pruritus, abnormal transrectal or transabdominal ultrasonographic findings (e.g. exudate build-up over separated chorion), estrous, pyrexia, abdominal discomfort, malaise – may alert the investigator to search particularly carefully for some form of placentitis. As the mare’s present and future general and reproductive health may be compromised by a uterine infection the search should not stop at a gross inspection, but be detailed and include histological and microbiological assessment (see earlier). In general the location of the placentitis reflects the route by which the infection entered the uterus (three main focal patterns and one diffuse are recognized).
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Cervical pole placentitis
More diffuse, often microscopic placentitis
In some countries (including the UK) this is the commonest type. By 60 days of gestation the chorion has extended to the cervix.16 If there are bacteria and/or fungi on the cervix, and if the cervical seal is compromised, they can enter the uterus and cause placentitis. Anatomical, hormonal or even neurological factors contributing to cervical incompetence and/or increased perineal or vaginal contamination will increase the risk of cervical placentitis. As well as placentitis many infections cause fetal inflammatory lesions (e.g. in liver and/or lung). A fetal septicemia tends to provoke abortion rapidly, often before placentitis has become grossly evident. In one series of 200 placentitis cases,17 12% were not visible grossly. The more chronic the infection the more extensive or pronounced the lesions of chorionic thickening and fetal growth retardation are likely to be. Some infections can spread rostrally to involve most of the chorionic body. Thickening at the cervical star can prevent it rupturing at birth or abortion, so that the chorion tears across the rostral body. Some fetuses with fungal infections are born alive but premature and compromised. Although most infections continue to invade the chorion, Taylorella equigenitalis is of low invasiveness and can become isolated into small invaginating allantochorionic pouches at the cervical pole, each containing desiccated inflammatory debris (of a similar appearance to invaginated endometrial cups).
In these cases the infections causing placentitis are believed to be blood-borne. In some rare cases mares may show pyrexia and signs of septicemia before aborting. Multiple small placentitis foci diffusely spread within the chorion are present. In several countries (USA, Canada, South America, New Zealand, Australia and some European countries including the UK) an increasingly recognized example of abortion and stillbirth due to a diffuse placental villitis is that caused by a number of different leptospiral serovars. Its demonstration requires specialized diagnostic methods, not routinely used at all diagnostic centers – serology on mares and fetuses, microbiological tissue testing for leptospires including PCR and culturing, and microscopy (silver stains, immunofluorescence and immunostaining). Labs using specific methods, for example in Northern Ireland and Lexington, USA, have identified the importance of leptospiral pregnancy losses for many years.20, 21 Poonacha et al.21 provide helpful descriptions of the gross and microscopic changes in fetal and placental tissues. The allantochorion of 56% of placentae had rather non-specific gross lesions. However, 95% of chorions, 43% of amnions and 51% of umbilical cords showed histopathological lesions. Diffuse villitis of the chorion and funisitis of the cord were two of the more characteristic placental lesions. For specific PCR testing using leptospiral transport medium, the tissues recommended for sampling include fetal liver, kidney, and adrenal gland, and chorion, cord surface, and amnion.
Horn base with/without rostral body placentitis A less common type of chronic-active placentitis occurs at the base of one or both uterine horns or rostral body. Gram-positive ‘nocardioform’ filamentous branching microorganisms have been associated with the distinctive lesions, where sites with villus loss are often coated in a brownish muddy mucoid exudate. Allantoic surfaces of the lesions may show adenomatous hyperplasia. The infection causes late abortions, stillbirths or premature births. Just how the bacteria invade the uterus has not been determined: it has been postulated that they enter at covering and remain at the most ventral aspect of the uterine floor or horn bases throughout pregnancy.18 Chopin et al.19 in Australia recently suggested that caterpillars may play a part in the pathogenesis of nocardioform placentitis.
Recurrent pregnancy loss Certain types of abortion have a risk of recurrence in a subsequent pregnancy: twinning and cervical placentitis are well-known examples. Thankfully steps can be taken clinically to reduce the risk. One worrying recurrence that is not possible to control is the risk factor of long-cordedness. Cord-related losses are numerically important in UK broodmares. Known influences on cord length only have minor effects. Only a better understanding of the major influences will enable a reduction in the losses. The medical profession is studying the phenomenon of long-cordedness and so it behoves veterinarians to keep up to date with any new comparative data that may be forthcoming. Diagnostic uncertainties
Focal single sites elsewhere Very rarely focal placentitis is believed to relate to a single site where there is a focal uterine infection or abscess.
The percentage of investigated abortions with no diagnosis has steadily fallen. Although there has been progress in many areas of etiological abortion diagnosis there remains much that is only
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Abortions and Stillbirths: A Pathologists Overview descriptive. With conscientious observation and recording, pathologists have defined and named many fetoplacental lesions, but their etiology and/or exact pathogenesis often remains speculative. Some examples include the phenomena of body pregnancy, ‘contracted’ fetuses, chronic fetal diarrhea, chorionic villus hypoplasia/atrophy, premature placental separation and even nocardioform placentitis and MRLS. Improved understanding of these may require a more holistic approach. Investigations may need to broaden, to consider uterine and/or maternal general health and physiology, rather than be restricted to assessing the products of the abortion. When for some reason a pregnant or aborting mare dies, there is a unique window of opportunity to examine the maternal side of the pregnancy by extending the investigation to include a detailed maternal postmortem examination. It might provide answers to some of the unanswered riddles. Hence, in the unwelcome event of a future catastrophic outbreak of abortifacient disease in a broodmare population, judicious culling of one or two affected mares for detailed necropsy could bring a much broader understanding of the underlying disease process.
Acknowledgement K.W. wishes to thank the Animal Health Trust and Beaufort Cottage Laboratories, Newmarket, UK, and their staff, for the opportunity over several decades to conduct the diagnostic and research work that forms the basis for this chapter.
References 1. Anon. HBLB Codes of Practice. London: Horserace Betting Levy Board, 2010 (http://www.hblb.org.uk/ document.php?id = 43). 2. Rossdale PD, Ricketts SW. Evaluation of fetal membranes at foaling. In: Rossdale PD, Mair TS, Green RE (eds) Reproduction: Foaling Part 1: Maternal Aspects. Newmarket: Equine Veterinary Journal Ltd, 2002; pp. 78–82. 3. Smith KC, Whitwell KE, Blunden AS, Bestbier ME, Scase RJ, Geraghty RJ, Nugent J, Davis-Poynter NJ, Cardwell JM. Equine herpesvirus-1 abortion: atypical cases with lesions largely or wholly restricted to the placenta. Equine Vet J 2004;36: 79–82. 4. Mahaffey LW. Abortion in mares. Vet Rec 1968;82:681–6.
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5. Rooney JR, Robertson JL. Equine Pathology. Ames: Iowa State University Press, 1996. 6. Whitwell KE, Jeffcott LB. Morphological studies on the foetal membranes of the normal singleton foal at term. Res Vet Sci 1975;14:44–55. 7. Smith KC, Blunden AS, Whitwell KE, Dunn KA, Wales AD. A survey of equine abortion, stillbirth and neonatal death in the UK from 1988 to 1997. Equine Vet J 2003;35:496– 501. 8. Ricketts SW, Barralet A, Whitwell KE. A review of the causes of abortion in UK mares and means of diagnosis used in an equine studfarm practice in Newmarket. Equine Vet Educ Manual 2003;6:18–21. 9. Ginther OJ. Equine pregnancy: Physical interactions between the uterus and conceptus. AAEP Proc 1998;44: 73–104. 10. Whitwell KE, Wood J. The length of the umbilical cord: association with abortion in Thoroughbreds and investigations into factors influencing length. In: Proceedings of 31st BEVA Congress, Equine Veterinary Journal, Newmarket, UK; 1992; pp. 53–6. 11. Baergen RN. Manual of Benirschke and Kaufmann’s Pathology of the Human Placenta. Springer, 2005. 12. Wilsher S, Ousey J, Allen WR. Abnormal umbilical cord attachment sites in the mare: a review illustrated by three case reports. Equine Vet J 2009;41:930–939. 13. Platt H. Aetiological aspects of abortion in the Thoroughbred mare. J Comp Pathol 1973;83:199–205. 14. Jeffcott LB, Whitwell KE. Twinning as a cause of foetal and neonatal loss in the Thoroughbred mare. J Comp Pathol 1973;83:91–106. 15. Whitwell KE. Investigations into fetal and neonatal losses in the horse. Vet Clin N Am-Large 1980;2 (2): Nov. 16. Ginther O.J. In: Reproductive Biology of the Mare, 2nd edn. Equiservices, 1992; p. 553. 17. Whitwell KE. Infective placentitis in the mare. In: Powell DG (ed.) Equine Infectious Diseases V. Lexington, KY: University Press of Kentucky, 1988; pp. 172–80. 18. Christensen WC, Roberts JE, Pozor MA, Giguere S, Sells SF, Donahue JM. Nocardioform placentitis with isolation of Amycolatopsis spp in a Florida-bred mare. J Am Vet Med Assoc 2006;228:1234–9. 19. Chopin JB, Muscatello G, Goswami P, Begg AP. Nocardioform placentitis in Australia with implications for equine amnionitis and foetal loss (EAFL) and mare reproductive loss syndrome (MRLS). Proc. 10th International Symposium on Equine Reproduction. Suppl. Anim. Reprod. Sci. (2010) S347–S348. Ed MJ Evans. 20. Ellis WA, Bryson DG, O’Brien JJ, Neill SD. Leptospiral infection in aborted equine foetuses. Equine Vet J 1983;15:321–4. 21. Poonacha KB, Donahue JM, Giles RC, Hong CB, PetritesMurphy MB, Smith BJ, Swerczek TW, Tramontin RR, Tuttle PA. Leptospirosis in equine fetuses, stillborn foals and placentas. Vet Pathol 1993;30:362–9.
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Chapter 241
Origin and Outcome of Twin Pregnancies Angus O. McKinnon
Introduction Three things about twins have been known for a long time. First, twins are repeatable within the same mare; second, twinning rate varies according to breed; and third, the more fertile the stallion, the more twins he is expected to achieve. Historically, twins have been the single most important cause of abortion.1–5 With the advent of ultrasonography, the incidence of abortion associated with twins has decreased dramatically to less than 6.1%,6 2.9%,7 or 1.7%8 of all abortions. Twins are often disastrous financially. Most twin pregnancies terminate in early fetal resorption or loss, late term abortions, or the birth of small growthretarded foals. Mares aborting twins in late gestation frequently have foaling difficulties, damage their reproductive tracts, and are difficult to rebreed. If foals born alive they are frequently small, demonstrate the effects of intrauterine growth retardation, and have a poor survival rate, with many needing expensive sophisticated critical care.
Origin of twins The origin of equine twins is almost exclusively dizygotic. Zygosity refers to the origin of the twin. Dizygotic twins originate from two separate fertilized oocytes with two different spermatozoa, and monozygotic refers to identical twins that arose from one fertilized oocyte. Monozygotic twins Twins in the horse are unlikely to be identical. Almost all cases of twin pregnancy are expected to be dizygotic and thus related to multiple ovulations, although there are cases of monozygotic twin pregnan-
cies that have been suspected9–11 or confirmed in the horse.12–14 Exactly how monozygotic twin formation occurs is not known, although pinching by the zona pellucida or damage to the inner cell mass are popular theories.15 Time of development probably influences placental arrangements. Four types of placental arrangements are recognized in human monozygotic twins:16
r r
r r
Dichorionic/Diamniotic; twins are totally separate and share only a common contact area of the chorion. These occur when the embryo divides before blastocyst formation. Monochorionic/Diamniotic; twins share a common chorion but have separate amniotic cavities. These occur when the embryo divides after blastocyst formation and development of the inner cell mass and the trophectoderm. Monochorionic/Monoamnionic; twins separate after formation of the ectoplacental cone and share a common chorion and amnion. Conjoined twins; twins incompletely separate.
In the horse, one apparent set of dichorionic/diamniotic monozygotic twins has been reported,9 while all other suspected or confirmed cases of monozygotic twins have been monochorionic/diamniotic.9,11–13 It is of interest that the three cases involving embryo transfer9 were all from transfer of morulas, which by definition have not separated into the inner cell mass and the trophectoderm,17 and that the two monochorioinc/diamniotic twins were from morulas that had been cooled and transported before transfer. It is possible that embryo transfer pregnancies are overrepresented in the reports of monozygotic twins.9, 14 It is not expected
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Origin and Outcome of Twin Pregnancies to be possible to determine the true incidence of dichorionic/diamniotic monozygotic twins as both should be present at the initial ultrasonographic pregnancy diagnosis on or around day 13 after ovulation, and would be indistinguishable from dizygotic twins apart from the number of corpora lutea present. In this regard, a study reporting a probable incidence of monozygotic multiple pregnancies of 2.6%10 should be interpreted with caution, as only dichorionic/diamniotic monozygotic pregnancies would be detected at the first pregnancy examination and the identification was based on identification of one less CL than the number of vesicles detected10 which is not always accurate, even in experienced hands. The reason for lack of monozygotic twin formation in the horse is probably related to the capsule.18 The capsule forms in equine embryos aged around 6 days, shortly after their entry into the uterus.19, 20 When embryos were cultured prior to the formation of the capsule, hatching occurred similar to the bovine;20 however, when embryos were cultured after formation of the capsule, the zona pellucida continued to become progressively thinner and finally fell away from the developing conceptus. Other species that do not have a capsule, such as sheep, cattle, and humans, all have the ability to give birth to monozygotic twins which may form from pinching of a hatching embryo by the zona pellucida. Identification of dichorionic/diamniotic monozygotic pregnancies must rely on the use of DNA microsatellite loci as placental arrangements are indistinguishable from dizygotic twins. Monochorioinc/diamniotic twins can almost always be assumed to be identical.
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241.1).22–24 Twins are more common in Thoroughbreds and Draught type horses than Quarter Horses, Ponies and Arabians.25 Multiple ovulation rate is correlated with twinning rates. Multiple ovulation rates have been reported to be 15–25% in Thoroughbreds, 13–15% in Standardbreds, 24% in Draught Horses, 8–10% in Quarter Horses and Appaloosas, 2–3% in domestic ponies, and 0% in Korean wild ponies.25
Age Multiple ovulation has been reported from a slaughterhouse study to be more common in older mares (18% in mares aged ≥ 6 years) than younger mares (14% for mares ≤ 5 years).26 Twinning rate has been reported to be 2.8%, 3.4%, 3.6%, and 6.8% for mares aged 4–7, 8–11, 12–15, and 16–20 years, respectively.27
Reproductive status Fewer multiple ovulations are expected on the first postpartum estrus compared to other cycles.25 The highest number of twin pregnancies are recorded in barren mares, followed by maiden mares, and then lactating mares.25, 27, 28
Season Although various reports have been published indicating that twins were more common either at the beginning or the end of the breeding season, many are confounded and not supported by large numbers.29 It is most likely that season does not play a major part in the probability of twins.25
Dizygotic twins Twin pregnancies originating from multiple ovulations are dizygotic. A strong association between twinning rate and breed has been reported. Other associations that have been reported or suggested are age, reproductive status, season, and use of drugs to control ovulation. Twinning appears to have a high degree of repeatability and heritability.21
Breed More twins are diagnosed per cycle in Thoroughbreds compared with Standardbreds (Table
Ovulation induction Multiple pregnancies are more commonly detected when ovulation induction is used.23, 30, 31 It is not clear if a higher probability of twins is a result of more ovulations or a better ability to detect twins due to a closer synchrony of ovulation. Twin pregnancy rates were higher when either deslorelin acetate (Ovuplant) or hCG were administered: 19.7% (29/147) and 14.8% (154/1042), respectively, compared to control, non-treated mares (11.8%; 62/527).23 Overall pregnancy rates were also improved: 72.0% (147/204),
Table 241.1 Identification of twin pregnancies in populations of Thoroughbred and Standardbred mares
Study 24 23 22
Twins per mare Twins per cycle Twins per cycle
Thoroughbred (n)
Thoroughbred (%)
Standardbred (n)
Standardbred (%)
97/629 236/2436 510/6123
15.4 9.7 8.3
39/635 46/2086 123/2690
6.1 2.2 4.6
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72.0% (1042/1447), and 64.5% (527/817), respectively, for deslorelin acetate, hCG, and control groups.23 Despite clear associations in some studies,23, 30, 31 other reports indicate no effect of ovulation induction on the probability of twin detection.32, 33 Reasons for differences may be related to techniques used to determine when to use drugs to hasten ovulation, dosage of drugs or farm management practices, or unrecognized confounding variables. It has been shown that twins in mares are as likely to result from synchronous versus asynchronous ovulation.34 and that pregnancy rate per follicle was identical for double ovulations on opposite ovaries as compared to that obtained from single ovulations per cycle. A point of interest in relation to the origin of twin pregnancies is the recognition that twin pregnancies were more likely to occur (P < 0.05) if ovulations were bilateral versus unilateral (77% bilateral versus 60% unilateral ovulations)35 and a similar effect was noted with embryo recovery.36 The probable mechanism for the decreased frequency of twins in unilateral ovulations may be related to the oocyte capture by the fimbria being disturbed somewhat in cases of multiple large follicles on one ovary. Apart from the rare occurrence37 of trophoblastic vesicles (an absence of the embryo/fetus to develop inside the vesicle/conceptus) there is no evidence of pre-
fixation embryo reduction35 and thus disparity in size between vesicles; pre-fixation is almost always due to asynchronous ovulations.
Outcome of twin pregnancies Understanding the outcome, if twin pregnancies are left to develop, is of great importance to veterinarians, farm managers, and owners. Also important is when to intervene and what probability of success such interventions may be expected to achieve (see Chapter 220). The mare is very efficient at reducing twins to a single pregnancy. This is achieved by a competitive absorption of nutrients that is related to size and position of the early pregnancy and later to orientation of the embryo proper within the developing conceptus25, 38 (Figure 241.1). However, the implementation of any non-intervention program for resolution of twins depends on the age at identification, the orientation of the vesicles, and any disparity in size. Pre-fixation Embryo reduction before or on the day of fixation is not considered an important aspect of the natural correction of twins.39 The probability of a mare losing one or both vesicles of a set of twins from
Figure 241.1 Deprivation hypothesis redrawn from Ginther (1992).25 The deprivation hypothesis proposed by Ginther25 suggests that embryonic reduction is associated with a competitive absorption of nutrients that is related to size and position of the early pregnancy and later to orientation of the embryo proper within the developing conceptus. The bi-walled portion of the embryo (clear area) consists of the endoderm and ectoderm and the tri-walled portion (gray area) the addition of the mesoderm. Endometrial contact (surface area) of the vascularized tri-walled portion influences absorption of nutrients and is determined by the orientation process. When the tri-walled membranes have little contact surface area with the endometrium (column A day 17) the competitive advantage of the other embryo with better surface area contact results in rapid destruction of the disadvantaged embryo. The embryonic reduction process is less rapid as more of the tri-walled membrane has surface area contact with the endometrium (column B and C) and almost non-existent when even surface area contact occurs (column D). The more surface area contact of the disadvantaged embryo the more chance there will be of the development of the allantois in the smaller or disadvantaged embryo. The orientation of the embryos is influenced by their relative size. Dissimilar embryos are more likely to assume the orientation in column A and similar-sized embryos column B-D). Adapted from Ginther.25
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Figure 241.2 Day 11 and 14 pregnancy.
identification prior to fixation is minimal, and approximates that of early embryonic death for the same time period (per vesicle). The recognition of twin pregnancies prior to fixation day (day 16) is dependant on the day of examination relative to the day of ovulation. Asynchronous ovulations occasionally result in a gross disparity in vesicle size, sometimes as much as 4–5 days, i.e., identification of a day 11–12 and a day 15–16 vesicle concurrently (Figure 241.2). In instances such as this, examination one day earlier may have failed to detect the younger of the two pregnancies. Recognition that almost all twin pregnancies occur from multiple ovulation dictates mandatory re-examination of all mares that have two CLs and only a single vesicle detected prior to fixation (day 16). Recognition prior to fixation is also dependent on operator experience, resolution of the equipment (≥ 5 MHz preferred), monitor capabilities, restraint and other facilities (ability to darken the environment), the presence of uterine cysts, and the skill of the examiner. As smaller-than-expected single pregnancies have a higher rate of early embryonic death,25, 40 management techniques should consider the destruction of the smaller vesicle41 even if the body of evidence suggests minimal pre-fixation embryonic reduction. Recognition after fixation (after day 16) The recognition of unilaterally fixed twins from day 17 through to 21 (prior to clear recognition of the developing fetus within the vesicle) may be the most difficult time to determine if there are twins present. Ultrasonographically, all that is detectable is a thin line (the apposition of the two yolk sacs) running vertically approximately in the middle of what appears to be a slightly oversized vesicle (Figure 241.3a–c). Recognition of the fetus(es) within the vesicle a few days later makes differentiation eas-
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ier(Figure 241.3d). Occasionally, an inexperienced operator may confuse an abnormally orientated 28- to 30-day single pregnancy with 17- to 20-day unilaterally fixed twins. From days 22 to 60 the presence of multiple fetuses, umbilical cords, and general excess in the number of visible membranes should alert the practitioner to the likelihood of more than one pregnancy (Figure 241.4a–c). The junction between two developing fetuses (after 30 days) results in an ultrasonographically visible common membrane representing the area of apposition between the two allantochorions. This common membrane has been referred to as the twin membrane42 and has diagnostic potential, particularly later in pregnancy when it might not be possible to view both fetuses transrectally (> 100 days) (Figure 241.5). After 100 days, careful transabdominal ultrasonography may be necessary to determine the presence of twins.
Days 17 to 40 The outcome of pregnancies post-fixation is dependent upon their size (diameter) and the nature of their fixation. Unilateral (both fixed together at the same corpus cornual junction) fixation reduction is much higher than bilateral (one on each side) fixation reduction. Fortunately, unilateral fixation is much higher (approximately 70%) compared to bilateral (30%).38 In 28 mares with known ovulatory patterns, synchronous ovulations did not affect the type of fixation (9/17 unilateral, 8/17 bilateral). However, for asynchronous ovulation the frequency of unilateral fixation (10/11) was greater (P < 0.01) than the frequency of bilateral fixation (1/11). The incidence of embryo reduction was greater (P < 0.01) for unilateral fixation (14/19) than for bilateral fixation (0/9) and was greater (P < 0.05) for asynchronous ovulation (9/11) than for synchronous ovulation (5/17).38 Practically speaking, this means that if asynchronous ovulations have resulted in significant age differences between vesicles (i.e., > 3 mm diameter at day 15) then rate of natural embryonic reduction is very high.38 In cases of unilateral fixation, 22 of 22 mares with vesicles of dissimilar size had reduction compared to 19 of 26 (73%) with vesicles of similar size.43 As a result of work studying reduction of unilateral versus bilateral twin pregnancies in mares from days 17 to 40, Ginther proposed that the nutrient intake from the larger vesicle (before the fetus was present) prevented adequate nutrition of the smaller vesicle. Later the position of the fetus proper and its emerging allantoic sac seemed to determine whether a given conceptus survived or underwent late reduction. The fetus, the vascularized wall of the yolk sac adjacent to the fetus, and the emerging allantoic sac were exposed to the endometrium (uterine lumen) in the surviving
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(a)
(b)
(c)
(d)
Figure 241.3 (a) On day 20 all that is detectable ultrasonographically is a thin line (the apposition of the two yolk sacs) running approximately in the middle of what appears to be a slightly oversized vesicle. These may on occasion be difficult to detect. (b) Coalescing twin vesicles on day 16. (c) Occasionally the line between twin vesicles may be horizontal rather than vertical. (d) Twins old enough to have fetuses present makes recognition easier (day 23 fetus visible with the yolk sac on right side of the vesicle).
vesicles. In the vesicles that underwent reduction, much of the corresponding area of the vesicle wall was covered by the wall of the adjacent survivor and is thus deprived of adequate embryonal–maternal exchange and therefore regresses (Figure 241.1). In summary, dissimilarity in diameter increases the likelihood of unilateral fixation, increases the incidence of reduction for unilateral fixed vesicles, hastens the day of occurrence of reduction and shortens the interval from initiation to completion of reduction. The incidence of reduction for bilaterally fixed vesicles is negligible and approximates that of standard early embryonic death in this period. Of the 85% of reductions by day 40 in cases of unilateral fixed twin pregnancies, 59% of reductions had occurred between day 17 and 20, 27% between day 21 and day 30, and 14% between days 31 to 38. The ma-
jority of early reductions occurred spontaneously (by day 20) as compared to reductions after day 20 that were preceded by a gradual decrease in size of the eliminated vesicle. In addition, when twins were dissimilar in diameter (4 mm or more) they were more likely to undergo reduction by day 20.43 Other studies have demonstrated similar results. The hypothesis of an early embryonic reduction mechanism for elimination of excess embryo in mares was not new and had been suggested as early as 1982.44 However, ultrasonography was necessary to adequately document the occurrence and nature of the reduction.
Day 40 onwards Ginther and Griffin42 examined the natural outcome of bilateral twins (one in each horn) that were viable on day 40 in 15 pony mares. Readers should be
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(b)
(c) Figure 241.4 (a–c) From days 22 onwards the presence of multiple fetuses, umbilical cords and general excess in the number of visible membranes should alert the practitioner to the likelihood of more than one pregnancy.
Figure 241.5 A common membrane that has been referred to as the twin membrane (junction between the two chorioallantoic membranes) has diagnostic potential for twin identification, particularly later in pregnancy. Note also in this picture the thinner amniotic membrane.
aware that pony mares are not necessarily a good model for larger breeds, as the incidence of twins is low and evidence is suggestive that the larger the breed (Draught, Thoroughbred and Warmblood), the higher the probability of maintaining twins. Fifteen pony mares were monitored by ultrasonography until the outcome of the pregnancy was determined. In all, 66% (10/15) of the pregnancies suffered from either death of both (80% 8/10) or death of one fetus (20% 2/10) during months 2 or 3. For the remaining five pregnancies (5/15), nothing occurred from then until month 8. Between months 8–11, two mares lost one fetus (fetal death was associated with mummification) and two mares lost both. The two mares that lost one pregnancy both delivered undersized weak foals at birth. One mare (7%) delivered live twins at term, and two normal foals were born from mares losing the one pregnancy (absorption of the fetus rather than mummification) in month 2. In this study, six live foals were born (two of normal size),
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Figure 241.6 (a) Normal, singleton placentation. (b) Type A placentation was seen in 79% of cases (48 sets of twins). In this group 31/48 mares lost their pregnancies between 3 and 9 months (64.5%). (c) Type B placentation occurred in 11% of cases (seven sets) and both foals were usually similar in size and were usually born alive. Nine foals survived to 2 weeks from six sets of twins that made it to term. (d) Type C placentation was seen in 10% of cases (six sets of twins). In this group three foals were born alive from six pregnancies at term or resulted in mummification and subsequent abortion of one mummified and one freshly dead fetus.5 Redrawn from Jeffcott and Whitwell (1973).5 (e) Type B placentation that ended in abortion. Note equal size of each fetus, similar surface areas of the chorioallantois and large avillous contact area between the two membranes. (f) Type C placentation has resulted in mummification and subsequent abortion of one mummified and one freshly dead fetus.
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Origin and Outcome of Twin Pregnancies from a total of 15 mares and 30 fetuses. This incidence is similar to previous reports wherein of 130 pregnant mares with twins, only 17 live foals (13%) were produced.45 An interesting observation from the later report was that, from the 102 mares that delivered live or dead twins in the previous year, only 37 produced live foals the next season, and thus over two seasons there was an average of 23% producing live foals.45 An earlier study5 was extremely useful in categorizing outcome of twins that managed to survive to later pregnancy. Twinning accounted for 22% of the cases of abortion and stillbirth between 1967 and 1970. Sixty-two sets of twins and their placentas were examined from Thoroughbred mares. All were considered to be dizygous. Abortion or stillbirth of both twins from 3 months of gestation to term occurred in 64.5% of mares, although most (72.6%) slipped from 8 months to term. In the remaining cases one twin (21%) or both twins (14.5%) were born alive. Most foals at term were stunted and emaciated and of the 31 alive at birth only 18 had survived to 2 weeks of age.5 In this study, twin placentation was divided into three morphological groups according to the disposition of the chorionic sacs within the uterus. Type A placentation (Figure 241.6b) was seen in 79% of cases (48 sets of twins). One fetus occupied one horn and most of the body (mean 68% of the total functional surface area) while the other twin occupied only one horn and usually only a small part of the adjacent body. Where the chorions abutted there was a variable degree of invagination of the smaller chorion into the allantoic cavity of the larger twin. These pregnancies frequently ended in abortion or stillbirth of one or both twins. In this group 31/48 lost their pregnancies between 3 and 9 months (64.5%). The gestation length in this group was frequently shorter, and at birth the larger twin had a much greater chance of survival than the smaller one. In this group only six foals of 48 sets were born alive. Of the six fetuses born alive, five were the larger twin. Type B placentation (Figures 241.6c,e) occurred in 11% of cases (7 sets) and the placentas were orientated such that the villous surface areas were more or less equally divided and each fetus occupied one horn and half of the body. Both foals were usually similar in size and were usually born alive. Nine foals survived to 2 weeks from six sets of twins that made it to term. In this group (7 total) one aborted at 7 months. Type C placentation (Figures 241.6d,f) was seen in 10% of cases (six sets of twins). In this group there was a greater disparity between the surface area of the two chorions (85% versus 15%). The smaller twin, occupying only part of one horn, died earlier on and became mummified. The larger twin was usually born alive with a fair chance of survival. In this group three foals were
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born alive from six pregnancies at term.5 The authors attributed the loss of twin fetuses and poor survival rates to placental insufficiency. Readers are encouraged to obtain the article of Jeffcott and Whitwell as, with the current use of ultrasonography, it is unlikely that a population of this magnitude of twin pregnancies progressing to an inevitable conclusion will ever be available again. When one or both of opposite sex twins are born alive and survive, their reproductive potential should not be affected. Cattle that have opposite sex pregnancies commonly develop the ‘freemartin condition’ and this is related to vascular connections between both allantochorions; XX/XY chimerism develops, hormones are interchanged, and ultimately there is masculinization of the female tubular reproductive tract to varying degrees.46 In horse twin pregnancies, interplacental vascular anastomoses were macroscopically visible in ∼ 25% of the cases and resulted in blood chimerism without interfering with fertility, and no freemartins were observed despite evidence of chimerism.47 This was explained by the relatively late establishment of vascular anastomoses of the allantochorion in the horse (after gonad differentiation has occurred) compared to the cow, where blood vessel anastomoses begin as early as 50 days.47 It should be clear that our philosophy is that nonintervention is only acceptable when twins are diagnosed as a unilateral occurrence between days 17 and day 40, and then the decision depends on factors such as the value of the foal, the potential for rebreeding, and the ability of the veterinarian to manually intervene. Intervention in twin pregnancies is strongly recommended in all other circumstances (see Chapter 220).
References 1. Acland HM. Abortion in mares. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993: pp. 554–62. 2. Acland HM. Abortion in mares: diagnosis and prevention. Compend Cont Educ Pract Vet 1987;9:318–26. 3. Kaur R, Sharma JK. Epidemiologic studies on abortions in Indian thoroughbred mares. J Equine Vet Sci 1996;16(11):508–10. 4. Platt H. Aetiological aspects of abortion in the Thoroughbred mare. J Comp Pathol 1973;83:199–205. 5. Jeffcott LB, Whitwell K. Twinning as a cause of foetal and neonatal loss in the Thoroughbred mare. J Comp Pathol 1973;83:91–106. 6. Hong CB, Donahue JM, Giles RC, Petritesmurphy MB, Poonacha KB, Roberts AW et al. Equine abortion and stillbirth in Central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. 7. Ricketts SW, Barrelet A, Whitwell KE. A review of the causes of abortion in UK mares and means of diagnosis
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9.
10. 11.
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21. 22.
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25. 26. 27. 28.
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used in an equine studfarm practice in Newmarket. Pferdeheilkunde 2001;17(6):589–92. Merkt H, Jochle W. Abortions and twin pregnancies in Thoroughbreds: Rate of occurence, treatment and prevention. J Equine Vet Sci 1993;13(12):690–4. McCue PM, Thayer J, Squires EL, Brinsko SP, Vanderwall DK. Twin pregnancies following transfer of single embryos in three mares: a case report. J Equine Vet Sci 1998;18:832–4. Newcombe JR. The probable identification of monozygous twin embryos in mares. J Equine Vet Sci 2000;20(4):269–74. Rooney JR. The fetus and foal. In: Rooney JR (ed.) Autopsy of the Horse. Philadelphia: Williams & Wilkins, 1970; pp. 121–33. Meadows SJ, Binns MM, Newcombe JR, Thompson CJ, Rossdale PD. Identical triplets in a Thoroughbred mare. Equine Vet J 1995;27:394–7. Govaere J, Hoogewijs M, De Schauwer C, Van Zeveren A, Smits K, Cornillie P et al. An abortion of monozygotic twins in a warmblood mare. Reprod Domest Anim 2009;44(5):852–4. Mansill, SS, R J Arnott, C C Love, K Hinrichs. Monozygotic equine twins after embryo trasnfer verified by DNA typing. Proc Am Assoc Equine Pract 2010;54:278. Hall JG. Twinning. The Lancet 2003;362:735–743. Scott L. The origin of monozygotic twinning. Reproductive BioMedicine Online 2002;5(3):276–84. McKinnon AO, Squires EL. Morphologic assessment of the equine embryo. J Am Vet Med Assoc 1988;192:401–6. McKinnon AO, Voss JL, Squires EL, Carnevale EM. Diagnostic ultrasonography. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, London: Lea & Febiger, 1993: pp. 266–302. Betteridge KJ, Eaglesome MD, Mitchell D, Flood PF, Beriault R. Development of horse embryos up to twenty two days after ovulation:observations on fresh specimens. J Anat 1982;135:191–209. McKinnon AO, Carnevale EM, Squires EL, Carney NJ, Seidel GE, Jr. Bisection of equine embryos. Equine Vet J (Suppl) 1989;8:129–33. Ginther OJ. Twinning in mares: a review of recent studies. J Equine Vet Sci 1982;2:127–35. Nath LC, Anderson GA, McKinnon AO. Reproductive efficiency of Thoroughbred and Standardbred horses in northeast Victoria. Aust Vet J 2010;88:169–175. McKinnon AO. Reproductive efficiency of horses in Australia. http://www.gvequine.com.au/breeding efficiency. htm. 2000. Bowman T. Ultrasonic diagnosis and management of early twins in the mare. Proceedings of American Association of Equine Practitioners, 1986; pp. 35–43. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992. Henry M, Coryn M, Vandeplassche M. Multiple ovulation in the mare. Zentralbl Veterinarmed A 1982;29:170–84. Deskur S. Twinning in Thoroughbred mares in Poland. Theriogenology 1985;23:711–18. Ginther OJ. Effect of reproductive status on twinning and on side of ovulation and embryo attachment in mares. Theriogenology 1983;20(4):383–95.
29. Merkt H, Jochle W. More twin pregnancies as season progresses. J Equine Vet Sci 1999;19(9):536. 30. Perkins NR, Grimmett JB. Pregnancy and twinning rates in Thoroughbred mares following the administration of human chorionic gonadotropin (HCG). N Z Vet J 2001;49(3):94–100. 31. Veronesi MC, Battocchio M, Faustini M, Gandini M, Cairoli F. Relationship between pharmacological induction of estrous and/or ovulation and twin pregnancy in the Thoroughbred mares. Domest Anim Endocrinol 2003;25(1):133– 40. 32. Davies Morel MC, Newcombe JR. The efficacy of different hCG dose rates and the effect of hCG treatment on ovarian activity: Ovulation, multiple ovulation, pregnancy, multiple pregnancy, synchrony of ovulation in the mare. Anim Reprod Sci 2008;109:189–99. ¨ 33. Jochle W, Trigg TE. Control of ovulation in the mare with OvuplantTM : A short-term release implant (Sti) containing the GnRH analog deslorelin acetate – studies from 1990 to 1994. J Equine Vet Sci 1994;14:632–44. 34. Ginther OJ. Relationships among number of days between multiple ovulations, number of embryos, and type of embryo fixation in mares. J Equine Vet Sci 1987;7:82–8. 35. Ginther OJ, Bergfelt DR. Embryo reduction before day 11 in mares with twin conceptuses. J Anim Sci 1988;66:1727–31. 36. Squires EL, McClain MG, Ginther OJ, McKinnon AO. Spontaneous multiple ovulation in the mare and its effect on the incidence of twin embryo collections. Theriogenology 1987;28:609–13. 37. Vanderwall DK, Squires EL, Brinsko SP, McCue PM. Diagnosis and management of abnormal embryonic development characterized by formation of an embryonic vesicle without an embryo in mares. J Am Vet Med Assoc 2000;217(1):58–63. 38. Ginther OJ. The nature of embryo reduction in mares with twin conceptuses: Deprivation hypothesis. Am J Vet Res 1989;50:45–53. 39. Ginther OJ. Twin embryos in mares. I: From ovulation to fixation. Equine Vet J 1989;21:166–70. 40. Carnevale EM, Ginther OJ. Relationships of age to uterine function and reproductive efficiency in mares. Theriogenology 1992;37(5):1101–15. 41. McKinnon AO, Rantanen NW. Twins. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. [place]: Williams & Wilkins, 1998: pp. 141–56. 42. Ginther OJ, Griffin PG. Natural outcome and ultrasonic identification of equine fetal twins. Theriogenology 1994;41:1193–9. 43. Ginther OJ. Twin embryos in mares. II: Post fixation embryo reduction. Equine Vet J 1989;21:171–4. 44. Ginther OJ, Douglas RH, Woods GL. A biological embryoreduction mechanism for elimination of excess embryos in mares. Theriogenology 1982;18(4):475–85. 45. Pascoe RR. Methods for the treatment of twin pregnancy in the mare. Equine Vet J 1983;15(1):40–2. 46. Padula AM. The freemartin syndrome: an update. Anim Reprod Sci 2005;87(1–2):93–109. 47. Vandeplassche MM. Special features of twin gestation in the mare. Vlaams Diergeneesk Tijdschr 1993;62:151–4.
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Chapter 242
Placentitis Mats H.T. Troedsson and Margo L. Macpherson
Introduction Placentitis in mares poses a significant threat to fetal and neonatal viability.1 It is commonly caused by bacteria ascending through the vagina, but can also enter the uterus by a hematogenous route.2–4 The most frequent bacterial pathogens implicated in equine placentitis are Streptococcus equi subsp. zooepidemicus, Escherichia coli, Klebsiella pneumoniae, Pseudomonas aeruginosa, and Leptospira species.5 Nocardioform actinomycetes have been reported in parts of the United States. The route of infection for this organism is poorly understood. Aspergillus fumigatus and other mucor species are the most common diagnosed causes of mycotic abortion in mares. While bacterial infection initiates fetal disease, recent work from an experimental model of ascending placentitis in pony mares showed that premature delivery may occur secondary to inflammation of the chorion rather than as a consequence of fetal infection.6, 7 These inflammatory processes result in prostaglandin production (PGE2 and PGF2α ) and stimulation of myometrial contractility, resulting in preterm delivery.8, 9 However, in some chronic cases of placentitis, foals will experience accelerated fetal maturation. These foals will be delivered prematurely, but will be mature enough to survive in the extrauterine environment. In humans, it is thought that indirect stimulation of the fetal hypothalamic-pituitary-adrenal axis by proinflammatory cytokines is responsible for precocious fetal maturation.10, 11 If this phenomenon is also true for equine fetuses, then delaying premature labor long enough to allow accelerated fetal maturation to occur may improve the chance of foal survival. To achieve this goal, it is necessary to promptly diagnose and effectively treat the disease.
Diagnosis of placentitis The most common clinical signs of placentitis in mares are premature udder development (with or
without streaming of milk) and vulvar discharge. Transrectal and transabdominal ultrasonography, possibly in combination with endocrinological assays, provide additional tools for diagnosing and monitoring progression of placentitis in mares.
Ultrasonographic monitoring of mares with placentitis Ultrasonography is an excellent tool for monitoring fetal and placental changes in mares affected by placentitis. Both transabdominal and transrectal evaluations can provide meaningful, and somewhat different, information. Mares often develop subclinical placentitis without overt clinical signs. Since serial ultrasonographic evaluations are not commonly employed in late gestation mares, diagnosis in subclinically affected mares can be missed. Subclinical disease may also result in subtle ultrasonographic changes that are not easily distinguished from normal findings. In spite of these hurdles, ultrasound remains one of the best tools available for diagnosing equine placental infections. Transrectal ultrasonography Transrectal ultrasonography of the caudal allantochorion in late gestational mares provides an excellent image of the placenta close to the cervical star12 (Figure 242.1). The cervical star region is most commonly affected in mares with ascending placentitis. Monitoring changes in this area can give an indication of subclinical and clinical disease. The area imaged in this region is the combined unit of the uterus and the chorioallantoic membrane from the placenta. This unit has been termed the combined thickness of the uterus and placenta (CTUP). 12 To perform the procedure, a 5–7.5 MHz linear transducer should be positioned 1–2 inches cranial of the cervical-placental junction, and then moved laterally until a major uterine vessel is visible at the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 242.1 Transrectal ultrasonography of the caudal part of the placenta in a late gestational mare. A, amniotic membrane; x – -x, CTUP.
ventral aspect of the uterine body.12 The CTUP should then be measured between the vessel and the allantoic fluid (Figure 242.1). It is important to obtain all CTUP measurements from the ventral aspect of the uterine body, since physiological edema of the dorsal aspect of the allantochorion has been noted in normal pregnant mares during the last month of gestation.12 In addition, care should be exercised to be certain that the amniotic membrane is not adjacent to the allantochorion, since this may result in a false increased CTUP. When possible, three measurements are taken and averaged. Normal values for CTUP have been established (Table 242.1).12, 13 Increases in CTUP > 8 mm between day 271 and 300, > 10 mm between day 301 and 330, and > 12 mm after day 330 have been associated with placental failure and pending abortion.13, 14 In some cases of placentitis, hyperechoic fluid (purulent material) can be noticed separating the uterus and the placenta (Figure 242.2). Measurements in these cases are meaningless as one is no longer measuring the combined unit. Additional parameters that can be evaluated using transrectal ultrasonography include changes in the amniotic membrane and fluid character. Amniotic thickening, which occurs in some cases of placentitis, can be identified using transrectal ultrasonogra-
Figure 242.2 Placental separation and accumulation of purulent material at the cervical star area in a mare with ascending placentitis.
phy (Figure 242.3). Changes in allantoic and amniotic fluid character can also be identified using transrectal ultrasonography. In normal mares, allantoic fluid is commonly hypoechoic with some specular material, while amniotic fluid is frequently a shade more hyperechoic (gray) than allantoic fluid. Marked changes in these fluid characteristics can also confirm presence of a placental infection or stress to the fetus.
AMNION THICK
Table 242.1 Normal upper limits for the combined thickness of the uterus and the placenta (CTUP) during late gestation12, 13 Gestation length
Normal CTUP
151–270 days 271–300 days 301–330 days 331–
<7 mm <8 mm <10 mm <12 mm
Figure 242.3 Thickening of the amniotic membrane in a mare with ascending placentitis.
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Placentitis
Figure 242.4 Placental lesions consistent with a nocardioform type of placentitis. Characteristics of this placentitis include a non-villous area covered with a thick, mucoid exudate located at the juncture of the uterine body and pregnant horn.
Transabdominal ultrasonography Transabdominal ultrasonography can also be a useful tool for placental evaluation in mares with suspected placentitis.12,15–17 Using transabdominal ultrasonography, the caudal portion of the allantochorion (cervical star area) cannot be imaged, which prevents the clinician from diagnosing ascending placentitis in its early stages. However, placental thickening and partial separation of the allantochorion from the endometrium can be detected using transabdominal ultrasonography in mares with placentitis originating from a hematogenous infection. In addition, mares infected with nocardioform bacteria will often have placental separation and purulent material at the ventral aspect of the base of the gravid horn and the junction of the uterine body (Figure 242.4). This finding is only identified using transabdominal ultrasonography. To examine the placenta from a transabdominal approach, a 3.5 MHz sector transducer is most frequently used. A 5 MHz linear transducer can be used for transabdominal evaluation of the placenta; however, depth penetration can limit evaluation of the fetus. All four quadrants of the placenta should be examined; right cranial, right caudal, left cranial, and left caudal. Measurements of the CTUP can be made with this technique if the fetus is not in close apposition with the uterine wall. Mares with normal pregnancies should have a minimum combined thickness of the uterus and the placenta (CTUP) of 7.1 ± 1.6 mm, and a maximal CTUP of 11.5 ± 2.4 mm.16 Pregnancies with an increased CTUP have been associated with the delivery of abnormal foals.16 Mares grazing endophyte-infected fescue often experience premature separation of the allantochorion, increased allantochorion weight and thickness, and re-
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tained placenta. A significant increase in uteroplacental thickness and premature separation of the allantochorion has been demonstrated on transabdominal ultrasonographic examination of endophyte-infected mares. However, the thickness was not observed until an average of 8 hours before the onset of labor.18 In addition to diagnosing placental disease, transabdominal ultrasonography is very useful for monitoring fetal health. Transabdominal ultrasonography of the equine fetus can be performed reliably after 90 days of gestation. At this time, the uterus drops over the pelvic brim and is visible from the ventral abdomen. The assessment of fetal well-being is obtained through measurement of heart rate, size, movement, and tone of the fetus. The thickness of the fetal membranes, echogenicity and quantity of the allantoic and amniotic fluids, and the number of fetuses provide information to evaluate the fetus. The fetus is visible in the inguinal area and between the mammary glands in early gestation. As pregnancy progresses, the fetus is found progressively more cranially. It is necessary to apply alcohol or to clip the hair on the abdomen to obtain a diagnostic image of the fetus. In late gestation, the mare should be examined from the mammary glands to the xyphoid extending to the level of the stifles on both sides of the abdomen. Variable gestational length, size and body type of the mare, and position of the fetus will affect the choice of transducer. The highest frequency transducer that will penetrate to the desired depth should be chosen. Generally, a 2.5 or 3.5 MHz transducer is required to image the fetal heart in late gestation since a depth of 30 cm is often required.19, 20 Either a curvilinear or sector scanner is preferred because they produce a pie-shaped image that allows an increasingly larger field of view in the deeper section of the image. Sedation of the mare will affect the heart rate, tone and movement of the fetus and should be avoided if possible. The ventral abdomen of the mare should be scanned in the sagittal and transverse plane. The entire uterus should be scanned to determine the number of fetuses and the position of the fetus. To determine fetal orientation, the uterus should be scanned in a sagittal section.19, 20 In late gestation the head of the fetus is near the brim of the pelvis. Orbital diameter can usually only be obtained by transrectal scanning.21, 22 The fetus is in dorsal recumbency with the vertebrae closest to the ventral abdominal wall.20 The thorax of the fetus can be localized by the recognizable striped pattern (Figure 242.5). The heart is found in the cranial aspect of the thorax. The fetal heart rate has been used as an indicator of fetal well-being.23 A poor outcome of parturition was associated with bradycardia or tachycardia in the fetus.
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Figure 242.5 Transabdominal ultrasound image of the fetal thorax obtained with a 2.5 MHz sector scanner transducer. The shadows are caused by the vertebrae and ribs. Ventral is the top of the image and dorsal is at the bottom.
Heart rate of the fetus peaks at 3 months of gestation to a mean of 196 beats per minute (bpm) and then gradually decreases throughout pregnancy.20 The decrease in fetal heart rate is a result of increasing parasympathetic tone to the heart.23 The average fetal heart rate in a fetus greater than 300 days gestation is 75 ± 7 bpm.23 Fetal heart rate slows by approximately 10 bpm at greater than 330 days gestation. Fetal heart rates vary with activity level, which should be considered when examining a mare. Consistently low or high fetal heart rates are associated with fetal stress. Fetal heart rate can be obtained either by using a stopwatch while monitoring the B-mode image or utilizing M-mode. M-mode displays movement at a fixed position of the transducer. The M-mode cursor is moved so that it intersects the heart. The M-mode image is activated. The image displayed will show movement of the heart over time (Figure 242.6). The heart rate is automatically calculated by measuring the time between two cardiac cycles. M-mode analysis is more accurate in assessing the fetal heart rate than the stopwatch method. Serial examinations should be performed to verify fetal well-being or distress. Mares considered ‘at risk’ for pregnancy loss are often examined on a daily basis. Fetuses experiencing distress are often evaluated several times a day. This is particularly true when determining if fetal distress is significant enough to prompt intervention such as induction of parturition.
Endocrine monitoring of the placenta
Figure 242.6 Transabdominal ultrasound image of the fetal heart obtained with a 2.5 MHz curvilinear scanner transducer. The B-mode image is at the left of the picture. The M-mode image is at the right of the picture. The cursor marks delineate two cardiac cycles. The calculated heart rate is 100 bpm.
metabolizes progestagens.24 This endocrine function of the placenta is important for maintenance of pregnancy after the endometrial cups and the secondary corpora lutea disappear around day 160–180 of gestation. Fetal-placental progesterone is rapidly metabolized to 5α-pregnanes. Mares with placental pathology may have increased plasma concentrations of progestagens as a result of stress to the fetal placental unit.25, 26 Unfortunately, 5α-pregnanes are not readily assayed in a commercial setting, so diagnosis of placental disease using 5α-pregnane concentrations is not possible. There is cross-reactivity between 5α-pregnanes and progesterone using some commercial radio-immunoassays for progesterone. In recent studies27 using an experimental model to induce placentitis, it was found that mares that develop a chronic form of placentitis responded with increased plasma progestin concentrations. Conversely, mares that developed acute placentitis and abortion soon after infection experienced a rapid drop in plasma progestin concentrations. It was suggested that measurement of repeated samples of plasma progestin concentrations in mares with placentitis might be a useful method to identify mares that may abort or deliver prematurely.26 Furthermore, sensitivity can be improved when progestin assays are combined with evidence of placental thickening, as detected using transrectal ultrasonography.27 Estrogen
Progesterone The equine placenta is part of an endocrine fetal–placental interaction, which synthesizes and
Serum estrogen concentrations are elevated in pregnant mares between 150–310 days of gestation.29 The predominant estrogens in pregnant mares are
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Placentitis estrone, equilin, equilenin, estradiol-17ß, and estradiol-17α. Determination of serum concentrations of estrone sulfate has been suggested to be useful in determining fetal well-being, and more recently, to monitor placental insult.30 Douglas29 observed that mares aborting from placentitis had serum estrogen concentrations below those normally detected in pregnant mares, and the author suggested that supplementation with estradiol cyprionate (ECP) or estradiol-17ß may prevent abortion in mares with placentitis.29 Relaxin Relaxin is produced by the equine placenta, and can be detected in peripheral blood plasma from day 80 of gestation and throughout the pregnancy.28 The role of relaxin during pregnancy is not fully understood, but there is evidence that placental relaxin production is compromised in mares at risk of aborting their fetuses.31 In a recent study, low circulating relaxin concentrations were associated with poor pregnancy outcome in mares with compromised placental function.32 However, relaxin concentrations were not useful in evaluating therapeutic efficacy in mares with placentitis.
Treatment of equine placentitis Treatment strategies for mares with ascending placentitis are currently ill-defined. Many treatment regimens have been extrapolated from other species, such as humans. Treatment efforts are directed at several factors including combating infection, reducing inflammation and controlling myometrial activity. Antimicrobial and anti-inflammatory therapy The majority of placental infections are caused by opportunistic bacteria migrating into the uterus from the caudal reproductive tract. The most common bacteria isolated in equine placentitis/abortion include Streptococcus equi subsp. zooepidemicus, Escherichia coli, Pseudomonas aeruginosa, Klebsiella pneumoniae, and nocardioform species.1 If a sample from vaginal discharge cannot be obtained to identify a specific bacterium, broad-spectrum antibiotic therapy is an essential first step in treating placentitis. Several studies in humans and non-human primates suggest that proinflammatory cytokines play a key role in the pathogenesis of infection-associated preterm delivery.8 Bacteria or bacterial products in the fetal membranes stimulate cell-mediated immune mechanisms with subsequent release of proinflammatory cytokines from macrophages and the decidua. In turn, proinflammatory cytokines stimulate release of
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prostaglandins E2 (PGE2 ) and F2α (PGF2α ) from the endometrium. Prostaglandins then initiate uterine contractions. As a consequence, treatment of placentitis in women and non-human primates has been varied.33–35 Antibiotic efficacy for reducing the incidence of preterm labor, when used alone, has been limited.33–35 Anti-inflammatory and immunomodulatory therapies have been shown to suppress inflammatory components of preterm labor in experimental models.36, 37 Recent work in non-human primates38 showed, not surprisingly, that antibiotic therapy (ampicillin) in combination with dexamethasone (immunomodulatory) and indomethacin (cyclooxygenase inhibitor) was more effective in suppressing bacterial infection, amniotic fluid cytokines, prostaglandins and uterine contractions than antibiotic therapy alone. Studies in these species indicate that combined therapy is necessary to stem the bacterial infection as well as to suppress the subsequent inflammatory response. Treatment approaches for equine placentitis have been largely anecdotal. Early studies evaluated the ability of commonly used antibiotics to penetrate the placenta in normal pregnant mares.39, 40 Sertich and Vaala40 administered potassium penicillin G and gentamicin sulfate, both alone and in combination or trimethoprim sulfadiazine to 11 normal, late pregnant mares. Samples were obtained from allantoic and amniotic fluid, when possible, and serum. Penicillin and gentamicin were detected in the mares’ serum at normal concentrations but not in amniotic fluid or foal serum. Penicillin was detected in the allantoic fluid of one mare. Trimethoprim sulfadiazine was recovered from both amniotic (n = 2) and allantoic (n = 4) fluid and was detectable in neonatal serum of two foals. The authors concluded that trimethoprim sulfadiazine penetrated fetal membranes sufficiently to combat bacteria that were sensitive to the drug. Santschi and Papich39 monitored the pharmacokinetics of gentamicin in late pregnancy and early lactation. Seven mares were treated with one dose of gentamicin sulfate prior to parturition, again after parturition, and pharmacokinetic analysis was performed on serum samples. Gentamicin was detected in mare serum samples at expected concentrations, but not detected in foal serum or in amniotic fluid samples. The authors concluded that lack of gentamicin in foal serum or amniotic fluid either indicated that gentamicin did not readily pass the equine placenta, or that the time between drug administration and sample collection may have been too short for sufficient drug distribution. In vivo microdialysis has recently been used to detect concentrations of commonly used drugs in allantoic fluid of pregnant pony mares.41 In the first study,
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five reproductively normal mares (269–271 days of gestation) had microdialysis probes placed in the allantoic cavity and jugular vein. Mares were treated intravenously with potassium penicillin G (22 000 IU/kg, q 6 h), and gentamicin (6.6 mg/kg, q 24 h). Allantoic fluid and blood were collected for 24 hours. Both antibiotics were present in allantoic fluid, albeit at lower concentrations than present in serum. Concentrations of penicillin in allantoic fluid achieved the minimum inhibitory concentration (MIC) against Streptococcus equi subsp. zooepidemicus. Gentamicin concentrations in allantoic fluid were adequate to be effective against Escherichia coli or Klebsiella pneumoniae. Also using microdialysis, the pharmacokinetics of trimethoprim sulfamethoxazole and pentoxifylline was studied in allantoic fluid of 10 pregnant pony mares.42 The drugs were chosen due to oral bioavailability and good uterine penetration. Pentoxifylline was evaluated because it appears to downregulate proinflammatory cytokines.43, 44 Five mares were inoculated, intracervically, with Streptococcus equi subsp. zooepidemicus 5 days before drug administration and measurement of drug concentrations by microdialysis. Five mares served as uninfected controls. All mares were treated with trimethoprim sulfa (30 mg/kg, b.i.d.) and pentoxifylline (8.5 mg/kg, b.i.d.), orally, for 14 days. Both drugs penetrated fetal membranes and were detected in allantoic fluid of treated and control mares. Four of the five infected mares aborted. Three mares aborted after termination of drug therapy (10, 17, and 19 days after the last day of treatment), one mare aborted on the 13th day of drug therapy, and one mare carried a normal foal to term (40 days after cessation of drug therapy). All control mares carried pregnancies to term and delivered healthy foals. These data suggested that trimethoprim sulfa and pentoxifylline might delay preterm delivery in mares with placentitis, but that treatment time was not sufficient to consistently improve neonatal outcome. The effects of long-term treatment with trimethoprim sulfamethoxazole and pentoxifylline on pregnancy outcome in mares with experimentally induced placentitis was investigated more recently.45 Mares (n = 11) were inoculated, intracervically, with Streptococcus equi subsp. zooepidemicus. Six mares were treated with the drug regimen described above from the onset of clinical signs until delivery or abortion. Five mares served as infected, untreated controls. Five of six treated mares aborted up to 27 days after bacterial inoculation. One mare delivered a healthy, live foal. Four of five control mares aborted after bacterial inoculation. One mare delivered a healthy, live foal. The treatment regimen did not improve fetal viability in this model, but treated
mares tended to carry pregnancies longer than untreated mares after inoculation. Furthermore, fetal and placental tissues were analyzed for presence of trimethoprim sulfamethoxazole and pentoxifylline. Drugs were detected in all fetal and placental tissues analyzed. Progestins Progestin therapy is currently being implemented in humans to halt preterm labor. A recent study 46 showed a beneficial effect when women with a documented history of spontaneous preterm delivery were treated with progesterone. The incidence of recurring spontaneous preterm delivery was significantly lower in women treated with 17α-hydroxyprogesterone, than in untreated women (36.3% vs 54.9%, respectively). Presumably, the antiprostaglandin effect of progestins would contribute to reduced myometrial activity by interfering with upregulation of prostaglandin and oxytocin receptors.47 Without receptor formation, gap junction formation would be inhibited and coordinated uterine contractility prevented. Daels and co-workers48 tested the effects of progesterone and altrenogest, a synthetic progestin, on pregnancy maintenance in mares treated with the prostaglandin analog, cloprostenol. Sixteen mares with pregnancies ranging from 93 to 153 days of gestation were included in the study. Cloprostenol (250 µg) was administered to all mares, intramuscularly, for 5 consecutive days. Progesterone (300 mg, i.m., s.i.d.) was administered to eight mares, and altrenogest (44 mg, p.o., s.i.d.) was administered to eight mares. Cloprostenol-treated control mares were extrapolated from a previous study. Five of eight mares (63%) in the progesterone-treated group maintained pregnancies after cloprostenol treatment, while three mares aborted during treatment. All eight mares (100%) treated with altrenogest maintained pregnancies. None of the control mares (0%) maintained pregnancies after cloprostenol treatment. Administration of exogenous progestins to mares treated with cloprostenol was associated with an endogenous decrease in prostaglandin metabolite concentrations. This important study demonstrated that progestin supplementation was able to prevent prostaglandininduced abortion in most cases. Progestin supplementation does not appear to prevent the initiation of normal parturition. However, a recent report suggested that progestin supplementation may slow down parturition, putting the health of the newborn foal at risk.49 These data deserve further study since clinical experiences do not always reflect this finding. Regardless of the effect of progestin supplementation on parturition, parturition of high-risk mares should
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Placentitis always be attended for assistance if needed and immediate attention to the newborn. Tocolytics The goal of tocolytic therapy is to prevent, or disrupt, uterine contractions and premature labor. A variety of agents have been used in women with preterm labor including: magnesium sulfate, betasympathomimetic agents (ritodrine, terbutaline), prostaglandin synthesis inhibitors (indomethacin, suldinac, ibuprofen, aspirin), calcium channel blockers (nifedipine), and oxytocin antagonists (atosiban). 35 Tocolytic agents have not been shown to significantly prolong pregnancy or improve neonatal outcome when used alone. Historically, tocolytics prolong pregnancy for up to 48 hours during which time glucocorticoids can be administered to the mother in an effort to expedite fetal maturation. Side effects from tocolytic agents can be significant.50 Clenbuterol, a beta-sympathomimetic agent, has been used in clinical equine practice. The effects of clenbuterol administration on uterine tone and maternal and fetal heart rates were examined by Card and Wood.51 Clenbuterol was administered intravenously (300 µg) to four pregnant mares at 30, 40, 50, 60 days of gestation and monthly thereafter until parturition. The final dose was administered when the mare was thought to be close to parturition as indicated by concentration of calcium and magnesium (120 ppm) measured using water hardness test strips. Fetal heart rate, maternal heart rate, and uterine tone (measured by palpation) were recorded after drug administration. Mares and fetuses experienced transient tachycardia after drug administration at all time points. Resting uterine tone changed significantly after clenbuterol administration to mares early in gestation. Uterine relaxation after clenbuterol administration was less profound later in gestation when uterine tone was decreasing naturally. Uterine relaxation occurred within 3 minutes of drug administration and persisted up to 120 minutes. The authors concluded that clenbuterol was effective in causing uterine relaxation throughout gestation, and that the side effects were minimal and transient. A more recent study reported the effects of clenbuterol when administered to 29 mares late in gestation.52 Beginning day 320 of gestation, mammary secretion electrolyte changes were monitored using a calcium strip test. Treatment started when calcium levels reached a maximum level of 13 mM. Mares were treated with varying doses of clenbuterol, i.v. – 0.6 mg (n = 6); 1 mg (n = 5), and 1.5 mg (n = 4). Fifteen control mares were treated with saline. All mares were treated once daily, at 10:00 pm, until parturition. No differences were detected between groups
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for length of gestation, number of treatments, or time to foaling after the last dose was administered. Treatment dose did not affect outcomes. Mares in the lowdose treatment groups (0.6 mg and 1 mg) showed no side effects, while mares treated with 1.5 mg showed transient signs of abdominal distress and sweating. All foals were clinically normal except one foal from the treatment group that died after dystocia. The authors concluded that clenbuterol was not effective in preventing the onset of myometrial contractions in normal foaling mares at term. Treated mares in this study actually foaled earlier in the evening than untreated mares. The authors speculated that the relaxant effects of clenbuterol may have promoted cervical relaxation and subsequent parturition. Based on side effects detected when clenbuterol is administered to pregnant mares,51, 52 and lack of effect for delaying normal parturition, the authors suggest that this agent has limited usefulness in horses. Combined therapy Work from a large-scale clinical trial examined the efficacy of multi-pronged, long-term therapy for treating equine placentitis.53 Investigators examined records of 477 mares over 6 years. Fifteen mares were diagnosed with placentitis. Criteria for treatment included increased thickness of the uteroplacental unit using transrectal ultrasound, placental separation and/or vulvar discharge and udder development. The average gestational age at diagnosis was 8.6 months. Mares were treated with a combination of systemic antibiotics (trimethoprim sulfa, ceftiofur or penicillin and gentimicin), pentoxifylline, altrenogest, and non-steroidal anti-inflammatory agents. Mares were treated until abortion or delivery of a foal. Of the treated mares, 84% carried their foals to term and 73% delivered live foals. Birth weights of surviving foals from mares treated for placentitis were similar to foals from non-affected mares. Data from these studies suggest that longterm antibiotic and anti-inflammatory treatment may positively impact pregnancy outcome in mares with placentitis. Recent work from the University of Florida54 supports treatment with combined therapy. Treatment with trimethoprim sulfamethoxazole (30 mg/kg, p.o., b.i.d.), pentoxifylline (8.5 mg/kg, p.o., s.i.d.) and altrenogest (RegumateTM , 0.088 mg/ kg, p.o., s.i.d.) was evaluated in 12 mares with experimentally induced placentitis. Five mares served as infected, untreated controls. Ten of twelve treated mares delivered live foals after treatment according to the protocol. All five of the untreated mares aborted or delivered non-viable foals. The combined effects of the drugs may have provided the necessary intervention for preterm delivery by addressing
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microbial infection, inflammation, and uterine contractility. Furthermore, early diagnosis of disease and initiation of treatment likely contributed to the positive outcome in this study. Results from this study also supported long-term therapy in affected mares (from the onset of clinical signs to delivery of the foal).
Conclusions Placentitis in the mare remains a clinical enigma. Efficient diagnosis of the disease relies upon vigilant monitoring of late pregnant mares using both gross and laboratory findings. Treatment strategies are better directed with results from recent studies. However, the treatment approaches available are far from perfect and treatment outcome is hard to predict. Early intervention with placental infections is likely the key to treatment success. A multifaceted treatment approach is necessary to resolve this complex disease. While success is hard won when treating placentitis in mares, delivery of a live foal after weeks or months of treatment is satisfying to all involved.
References 1. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3527 cases (1986–1991). J Am Vet Med Assoc 1993;203:1170–5. 2. Hong CB, Donahue JM, Giles RCJ, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Etiology and pathology of equine placentitis. J Vet Diag Invest 1993; 5:56–63. 3. Platt H. Infection of the horse fetus. J Reprod Fertil Suppl 1975;605–10. 4. Whitwell K. Infective placentitis in the mare. In: Powell DG (ed.) Equine Infectious Diseases V. Lexington, KY: University Press of Kentucky, 1988; pp. 172–80. 5. Acland HM. Abortion in mares. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 554–62. 6. LeBlanc MM, Giguere S, Brauer K, Paccamonti DL, Horohov DW, Lester GD, O’Donnell LJ, Sheerin BR, Pablo L, Rodgerson DH. Premature delivery in ascending placentitis is associated with increased expression of placental cytokines and allantoic fluid prostaglandins E-2 and F-2 alpha. Theriogenology 2002;58:841–4. 7. Mays MBC, LeBlanc MM, Paccamonti D. Route of fetal infection in a model of ascending placentitis. Theriogenology 2002;58:791–2. 8. Dudley DJ, Trautman MS. Infection, inflammation, and contractions – the role of cytokines in the pathophysiology of preterm labor. Semin Reprod Endocrinol 1994;12:263–72. 9. Pollard JK, Mitchell MD. Intrauterine infection and the effects of inflammatory mediators on prostaglandin production by myometrial cells from pregnant women. Am J Obst Gynec 1996;174:682–6.
10. Besedovsky HO, del Rey A. Immune-neuro-endocrine interactions: facts and hypotheses. Endocr Rev 1996;17:64–102. 11. Gravett MG, Hitti J, Hess DL, Eschenbach DA. Intrauterine infection and preterm delivery: evidence for activation of the fetal hypothalamic-pituitary-adrenal axis. Am J Obst Gynec 2000;182:1404–1413. 12. Renaudin CD, Troedsson MH, Gillis CL, King VL, Bodena A. Ultrasonographic evaluation of the equine placenta by transrectal and transabdominal approach in the normal pregnant mare. Theriogenology 1997;47:559–73. 13. Troedsson MHT, Renaudin CD, Zent WW, Steiner JV. Transrectal ultrasonography of the placenta in normal mares and mares with pending abortion: a field study. In: Proceedings of the American Association of Equine Practitioners, 1997; pp. 256–8. 14. Renaudin CD, Liu IKM, Troedsson MHT, Schrenzel MD. Transrectal ultrasonographic diagnosis of ascending placentitis in the mare: a report of two cases. Equine Vet Educ 1999;11:69–74. 15. Adams-Brendemuehl C, Pipers FS. Antepartum evaluations of the equine fetus. J Reprod Fertil Suppl 1987;35:565–73. 16. Reef BV, Vaala WE, Worth LT, Sertich PL, Spencer PA. Ultrasonographic assessment of fetal well-being during late gestation: development of an equine biophysical profile. Equine Vet J 1996;28:200–8. 17. Vaala WE, Sertich PL. Management strategies for mares at risk for periparturient complications. Vet Clin N Am Equine Pract 1994;10:237–65. 18. Boosinger TR, Brendemuehl JP, Schumacher J, Bransby DI, Kee D, Shelby RA. Effects of short-term exposure to and removal from the fescue endophyte Acremonium coenophialum on pregnant mares and foal viability. In: Equine Reproduction VI. Madison, WI: Society for the Study of Reproduction, 1995; pp. 61–7. 19. Pipers FS, Adams-Brendemuehl CS. Techniques and applications of transabdominal ultrasonography in the pregnant mare. J Am Vet Med Assoc 1984;185:766–71. 20. Reef VB, Vaala WE, Worth LT, Spencer PA, Hammett B. Ultrasonographic evaluation of the fetus and intrauterine environment in healthy mares during late gestation. Vet Radiol Ultrasound 1995;36:533–41. 21. Turner RM, McDonnell SM, Feit EM, Grogan EH, Foglia R. How to determine gestational age of an equine pregnancy in the field using transrectal ultrasonographic measurement of the fetal eye. In: Proceedings of the American Association of Equine Practitioners, 2006; pp. 250–5. 22. Turner RM, McDonnell SM, Feit EM, Grogan EH, Foglia R. Real-time ultrasound measure of the fetal eye (vitreous body) for prediction of parturition date in small ponies. Theriogenology 2006;66:331–7. 23. Curran S, Ginther OJ. M-mode ultrasonic assessment of equine fetal heart rate. Theriogenology 1995;44:609–17. 24. Pashen RL. Maternal and foetal endocrinology during late pregnancy and parturition in the mare. Equine Vet J 1984;16:233–8. 25. Rossdale PD, Ousey JC, Cottrill CM, Chavatte P, Allen WR, McGladdery AJ. Effects of placental pathology on maternal plasma progestagen and mammary secretion calcium concentrations and on neonatal adrenocortical function in the horse. J Reprod Fertil Suppl 1991;44:579–90. 26. Stawicki RJ, Ruebel H, Hansen PJ, Sheerin BR, O’Donnell LJ, Gester GD, Paccamonti DL, LeBlanc MM. Endocrinological findings in an experimental model of ascending placentitis in the mare. Theriogenology 2002;58:849–52.
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Placentitis 27. Sheerin PC, Morris S, Kelleman AA, Stawicki R, Sheerin BR, LeBlanc MM. Diagnostic efficiency of transrectal ultrasonography and plasma progestin profiles in identifying mares at risk for premature delivery. In: Proceedings of the American Association of Equine Practitioners, Focus in Equine Reproduction, 2003; pp. 22–3. 28. Stewart DR, Stabenfeldt GH, Hughes JP. Relaxin activity in foaling mares. J Reprod Fertil Suppl 1982;32:603–9. 29. Douglas RH. Endocrine and estrogens. Proceedings of the Society for Theriogenology, 2004; pp. 106–15. 30. Kashman LH, Hughes JP, Stabenfeldt GH, Starr MD, Lasley BL. Estrone sulfate concentrations as an indicator of fetal demise in horses. Am J Vet Res 1988;49:184–7. 31. Ryan P, Bennet-Wimbush K, Vaala WE, Bagnell CA. Relaxin as a biochemical marker of placental insufficiency in the horse: a review. Pferdeheilkunde 1999;15:622–6. 32. Ryan PL, Christiansen DL, Hopper RM, Bagnell CA, Vaala WE, Leblanc MM. Evaluation of systemic relaxin blood profiles in horses as a means of assessing placental function in high-risk pregnancies and responsiveness to therapeutic strategies. Ann NY Acad Sci 2009;1160:169–78. 33. Lamont RF. Infection in the prediction and antibiotics in the prevention of spontaneous preterm labour and preterm birth. BJOG: Int J Obst Gynaec 2003;110:71–5. 34. Lamont RF. Can antibiotics prevent preterm birth – the pro and con debate. BJOG: Int J Obst Gynaec 2005;112:67–73. 35. Ramsey PS, Rouse DJ. Therapies administered to mothers at risk for preterm birth and neurodevelopmental outcome in their infants. Clin Perinatol 2002;29:725–43. 36. Sadowsky DW, Haluska GJ, Gravett MG, Witkin SS, Novy MJ. Indomethacin blocks interleukin 1-beta-induced myometrial contractions in pregnant rhesus monkeys. Am J Obst Gynec 2000;183:173–80. 37. Sadowsky DW, Novy MJ, Witkin SS, Gravett MG. Dexamethasone or interleukin-10 blocks interleukin-1-betainduced uterine contractions in pregnant rhesus monkeys. Am J Obst Gynec 2003;188:252–63. 38. Gravett M, Sadowsky D, Witkin S, Novy M. Immunomodulators plus antibiotics to prevent preterm delivery in experimental intra-amniotic infection (IAI). Am J Obst Gynec 2003;189:S56. 39. Santschi EM, Papich MG. Pharmacokinetics of gentamicin in mares in late pregnancy and early lactation. J Vet Pharmacol Ther 2000;23:359–63. 40. Sertich PL, Vaala WE. Concentrations of antibiotics in mares, foals and fetal fluids after antibiotic administration in late pregnancy. In: Proceedings of the American Association of Equine Practitioners, 1992; pp. 727–36. 41. Murchie TA, Macpherson ML, LeBlanc MM, Luznar SL, Vickroy TW. A microdialysis model to detect drugs in the allantoic fluid of pregnant pony mares. Proceedings of the American Association of Equine Practitioners, 2003; pp. 118–21.
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42. Rebello S, Macpherson M, Murchie T, LeBlanc M, Vickroy T. The detection of placental drug transfer in equine allantoic fluid. Theriogenology 2005;64:776–77. 43. Lauterbach R, Zembala M. Pentoxifylline reduces plasma tumour necrosis factor-alpha concentration in premature infants with sepsis. Eur J Pediatr 1996;155:404–9. 44. Lauterbach R, Pawlik D, Zembala M, Szymura-Oleksiak J, Lisowska-Miszczyk I, Kowalczyk D, Bury J. Pentoxyfylline in and prevention and treatment of chronic lung disease. Acta Paediatrica Suppl 2004;93:20–22. 45. Graczyk J, Macpherson ML, Pozor MA, Troedsson MHT, Eichelberger AC, LeBlanc MM, Vickroy TW. Treatment efficacy of trimethoprim sulfamethoxazole and pentoxifylline in equine placentitis. Anim Reprod Sci 2006;94:434–5. 46. Meis PJ, Klebanoff M, Thom E, Dombrowski MP, Sibai B, Moawad AH, Spong CY, Hauth JC, Miodovnik M, Varner MW, Leveno KJ, Caritis SN, Iams JD, Wapner RJ, Conway D, O’Sullivan MJ, Carpenter M, Mercer B, Ramin SM, Thorp JM, Peaceman AM. Prevention of recurrent preterm delivery by 17 alpha-hydroxyprogesterone caproate. N Eng J Med 2003;348:2379–85. 47. Garfield RE, Kannan MS, Daniel EE. Gap junction formation in myometrium: control by estrogens, progesterone, and prostaglandins. Am J Physiol 1980;238:C81–9. 48. Daels PF, Besognet B, Hansen B, Mohammed H, Odensvik K, Kindahl H. Effect of progesterone on prostaglandin F2 alpha secretion and outcome of pregnancy during cloprostenol-induced abortion in mares. Am J Vet Res 1996;57:1331–7. 49. Palm F, Neuhauser S, Ambuehl F, Moestl E, Aurich C. Effect of altrenogest-treatment in late pregnant mares on parturition, neonatal adoption and arenocortical function in newborn foals. Proceedings of the 16th International Congress on Animal Reproduction, 2008; pp. 109. 50. Goldenberg RL. The management of preterm labor. Obst Gynec 2002;100:1020–37. 51. Card CE, Wood MR. Effects of acute administration of clenbuterol on uterine tone and equine fetal and maternal heart rates. In: Equine Reproduction VI. Madison, WI: Society for the Study of Reproduction, 1995; pp. 7–11. 52. Palmer E, Chavatte-Palmer P, Duchamp G, Levy I. Lack of effect of clenbuterol for delaying parturition in late pregnant mares. Theriogenology 2002;58:797–9. 53. Troedsson MHT, Zent WW. Clinical ultrasonographic evaluation of the equine placenta as a method to successfully identify and treat mares with placentitis. In: Proceedings of a Workshop on the Equine Placenta, 2004; pp. 66–7. 54. Bailey CS, Macpherson ML, Graczyk J, Pozor MA, Troedsson MHT, LeBlanc MM, Vickroy TW. Treatment efficacy of trimethoprim sulfamethoxazole, pentoxifylline, and altrenogest in equine placentitis. Theriogenology 2007;68:516–17.
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Chapter 243
Hydrops Rudolf O. Waelchli
Introduction Hydrops allantois (hydrallantois) and hydrops amnion (hydramnion) are rare conditions in the mare but, because of their dramatic appearance, are universally recognized. Hydrallantois can occur at any time from 6 to 7 months of gestation to term and is characterized by a sudden increase in allantoic fluid leading to abdominal distension over a few days. Hydramnion, an excess accumulation of amniotic fluid, is less common in mares, but also causes problems in the latter half of gestation. Normally, there are no inflammatory changes in the placenta, but there are exceptions.1 A mare with hydrops may be dyspneic and her well-being is further affected in several other ways. The etiology of the condition remains obscure.2 Most published reports on hydrops conditions involve single cases or small numbers of mares with a variety of breeds involved. Roberts3 included hydramnios, hydrallantois, edema of the allantochorion, fetal anasarca, fetal edema with ascites and hydrothorax in the condition collectively termed dropsy. Many affected mares abort spontaneously. The terminology used to describe the dropsical conditions in farm animals varies and terms used include hydrops amnii,3–6 hydrops amnion,7 hydrops of the amnion,3 hydramnios,3,5–10 hydrops of the fetal sacs,11 dropsy of the fetal sacs and dropsy of fetal membranes,2 hydrallantois,3, 5, 10, 12, 13 hydroallantois,4 hydrops allantois,1, 3, 7 hydrops of fetal membranes,10, 14 placental hydrops,15 hydrops uteri16 or, simply, excess fetal fluid.4 Another accepted term is hydramnion. In human medicine, the term polyhydramnios is used to describe an increase in the amount of amniotic fluid (to more than 2 L), which is commonly associated with fetal abnormalities such as fetal or neonatal hydrops with anasarca, ascites, pleural or pericardial effusion and skeletal malformations. In the human embryo, the allantois does not form a vesicle outside the fetus and, therefore, hydrallantois does not occur in women. Hydrallantois occurs sporadically in cattle –
commonly associated with multiple fetuses – and is seen 10 to 15 times more frequently than hydramnion. Hydramnion occurs most commonly in cattle and occasionally in sheep, but is rare in pigs, carnivores, and horses.3 The normal amounts of fetal fluids during pregnancy in the mare were reported by Arthur.11 The allantoic fluid gradually increases to about 8 to 15 L at term. Amniotic fluid, on the other hand, is very low in volume during the first 3 months, increases rapidly thereafter and is similar in amount to allantoic fluid at mid-pregnancy. The amount then decreases to about 3 to 5 L at term. The origin of fetal fluids has been the subject of many investigations. Generally, the biochemical composition of allantoic fluid resembles urine and that of amniotic fluid resembles a transudate. Both the allantoic and amniotic fluid cavities are connected to the fetal kidneys via the urachus and the urethra, respectively, and it appears that fetal urine is an important part of both fluids.5 However, the biochemical compositions of both fluids change during the different stages of pregnancy.17–21 Allantoic fluid reflects a transudate in the very early stages but, after about 3 months (and after the maturation of the fetal kidneys), the composition resembles modified urine and is notably characterized by an exponential increase in creatinine. Active transport across the chorioallantoic membranes also contributes to the allantoic fluid.17 The mechanisms involving the production of amniotic fluid are not clear, although the fluid is considered a secretory product of the amnion. In addition, secretions of the oropharynx, saliva and, towards the end of gestation, fetal urine from the urethra, all contribute to the fluid.19, 22 Towards the end of gestation, normal allantoic fluid of the mare varies from watery and clear to opaque brown; amniotic fluid is slightly viscous, transparent to slightly opaque, and white to yellowish. The gross appearance of fetal fluids can vary among individual mares.20, 22
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Hydrops In one report on hydrops in mares, the amount of allantoic fluid ranged from 120 to 220 L; one mare had concurrent hydramnion with 15 L of amniotic fluid.2 In a mare in which hydramnion had caused uterine rupture, the amniotic fluid was estimated at 96 L.9 In the mare, hydrallantois1, 2, 4, 10,12–14 occurs more frequently than hydramnion2, 4, 6, 7, 9 and is the most common dropsical condition.
Clinical signs Hydrallantois typically occurs in mid- to late pregnancy and is characterized by dramatic abdominal enlargement over a period of a few days to up to 2 weeks. Before this happens, the gestation of affected mares appears uneventful.2 Affected mares may be primiparous10 or, more commonly, multiparous,2 and of any age. Abdominal enlargement is accompanied by clinical signs that occur as a consequence of uterine distension. These include anorexia, decreased fecal output, depression, reluctance to move or lie down, ventral pitting edema, and dyspnea when lying down. Lymphatic obstruction by the enlarged uterus is believed to be responsible for ventral edema, which is often seen.12 Affected mares are uncomfortable, may shift weight, or even be seriously ill.2 Hydrallantois causes the abdomen to be barrel-shaped when viewed from behind, whereas, in hydramnios, the abdomen may be pear-shaped and softer.6 In cattle, hydramnion causes a more gradual enlargement of the abdomen than does hydrallantois.3 Spontaneous abortion is a common sequel to hydrallantois and is often complicated by uterine inertia because of overextension of the uterine musculature.
Diagnosis History (including pregnancy status and speed of onset of signs), inspection of the mare and clinical signs allow a tentative diagnosis of a hydrops condition, particularly hydrallantois. The abdominal enlargement is greater than that expected for the given stage of gestation.10 Transrectal palpation of the uterus is the first and easiest procedure to confirm the diagnosis. A grossly extended uterus is readily palpated. There may be restricted space for the examining arm and generally no fetus can be felt,1, 2, 4, 14 although there are exceptions.12 This is in contrast to a normal late pregnancy in which a fetus can usually be palpated.12 With hydrops, the uterus bulges dorsally above the pelvic brim or even into the pelvic cavity making transrectal examination difficult. Deep palpation is difficult because of the tense and distended abdomen.2 Transrectal or transabdominal ultrasonography has been used for the diagnosis of hydrops. Differ-
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ent variables have been assessed as end points, including fetal visibility and the presence of twins,7, 12 fetal viability,4, 6, 9 fetal heart rate,6, 9, 12 edema of the amnion,6, 7 combined thickness of the uterus and placenta,7 echogenicity4, 6 and amount of fetal fluid.6, 7, 10 Ultrasonography does not always allow the differentiation of amniotic and allantoic fluid, especially when the allantoamnion is not visualized. If two fluid compartments can be visualized with a septum (the allantoamnion) between them, the compartment closer to the body wall of the mare is likely to be the allantoic cavity.7 Because the amount of amniotic fluid in a normal pregnancy is relatively small, the amnion appears closely associated with the fetus and only small amniotic fluid pockets are visible. Ultrasonography has been used to support a diagnosis of hydrops amnii in a mare based on the observation that the allantoamnion was not intimately associated with the fetus and the latter was surrounded by a large amount of fluid.6 In accordance with human obstetrics, an equine biophysical profile has been established for the ultrasonographic assessment of fetal well-being during late gestation.23 One can also attempt to diagnose a hydrops condition (e.g., hydramnion versus hydrallantois) on the basis of the appearance and composition of the fetal fluids, procured either by ultrasound-guided transcutaneous aspiration in the pregnant mare7,19–21,24 or at delivery. The compositions of allantoic and amniotic fluid throughout gestation,18 during the last few months21 or close to or at term19, 20, 25 have been investigated. However, while biochemical analysis of the fluid is helpful for the differentiation of normal amniotic and allantoic fluid, its usefulness for the differentiation of hydramnion and hydrallantois in the mare is less clear.10 Because of the rarity of the condition in the mare, there have been no large systematic studies on the composition of allantoic and amniotic fluid in mares with hydrops. Such a study in cows revealed several interesting and diagnostically useful observations:17 in the beginning of the normal bovine pregnancy, the concentrations of sodium, potassium, and chloride in the allantoic fluid resemble those found in extracellular fluid, whereas after about 3 months of pregnancy, the concentrations of sodium and chloride decrease and that of potassium increases. In contrast, amniotic fluid resembles extracellular fluid throughout pregnancy with respect to the concentrations of these three ions. In hydrallantois, the electrolyte composition of the allantoic fluid differs from the corresponding normal allantoic fluid and closely resembles that of extracellular fluid. The amniotic fluid, in hydrallantois, deviates little from normal except that it seems to be mildly contaminated by urine. Normal allantoic fluid is characterized partly by fetal urine, partly by transport
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activity across the chorioallantoic membrane. It is not known whether all these observations also apply to the mare but some reports suggest that at least some do. Concentrations of calcium, sodium, potassium, and magnesium2 and protein and creatinine10 were serum-like in allantoic fluid of mares with hydrallantois. Likewise, in three cases of hydrallantois seen by this author, concentrations of sodium, potassium, calcium, magnesium, phosphorus, and chloride closely resembled those of serum, whereas creatinine concentrations tended to be higher than serum creatinine concentrations.14 It is not known whether the composition of amniotic fluid changes in mares with hydramnion. When fluid with a serum-like electrolyte concentration is collected during the delivery of a dropsical fetus, it is difficult to make a definitive diagnosis (hydrallantois versus hydramnion) based on electrolyte concentrations alone without proper clinical and obstetrical examinations because the fluid could reflect hydrallantois, normal amniotic fluid or, possibly, hydramnion. In two reports, the dropsical fluid had a serumlike electrolyte concentration and was identified as amniotic fluid; one case in association with obstetrical signs of hydramnion4 and the other on the basis of ultrasonographic identification of the amniotic cavity and concurrent analysis of fluid from a second ultrasonographically defined compartment identified as normal allantoic fluid.7 These two studies indicate that in hydramnion the electrolyte concentrations of amniotic fluid may not deviate significantly from normal. Other than hydrops, the differential diagnosis of sudden abdominal enlargement primarily includes, other than hydrops, twin pregnancy, and ascites caused by congestive heart failure or chronic liver failure.10 It should be remembered that dropsical conditions may occur in association with twin or multiple pregnancies.4, 10, 12
Possible causes The pathogenesis of hydrallantois seems to be associated with structural or functional changes in the chorioallantoic membranes such as dysfunction of chorionic ion pumps.1 The placenta may be macroscopically and microscopically normal or it may be edematous and cystic.2 Concurrent dysfunction of the fetal renal tubules cannot be excluded;17 in cattle, but not in horses, a high incidence of fetal hydronephrosis associated with hydrallantois has been noted,11 although these changes are probably secondary to hydrops.3 The rather sudden disturbance in the fluid exchange between uterus and fetus in hydrops is striking and its etiology remains unknown; there could possibly be increased secretion into, or
reduced resorption from, the allantoic sac, or both.2 One group views the fetal fluid as the result of equilibrium between its production (primarily in the fetal lung and kidney) and removal (by fetal swallowing).23 The cause of hydramnion is also unclear; as in hydrallantois, there may be increased secretion into, or decreased resorption from, the amnion, or possibly both.2 The presence of meconium implies swallowing of fluid by the fetus in utero and it would not seem far-fetched to propose that the fetus itself actively regulates the amount of amniotic fluid, and that, consequently, a lack of swallowing would be involved in the development of hydramnion. This hypothesis was tested, and subsequently rejected, by Wintour et al.8 who experimentally induced esophageal atresia in ovine fetuses and found that it failed to cause hydramnion. The conclusion from that experiment was that although fetal swallowing does normally provide a significant means of disposal of sodium from the amniotic fluid, other undefined mechanisms regulate the volume and composition of amniotic fluid. Although there is no firm evidence that hydrops has a genetic component, it is interesting to note that in one report, six of eight cases seen during a 19year period were in Belgian draught horses, leading to a suggestion of a breed disposition,2 and that in another report, at least two cases of hydrops in Haflinger mares involved the same stallion.14
Complications and prognosis Hydrops is fraught with potential complications including rupture of the prepubic tendon, herniation of the abdominal wall, uterine rupture, abortion, dystocia, and retained placenta. Complications associated with delivery are very common and include uterine inertia, delayed involution, puerperal fluid pooling, laminitis, and shock.2, 10 Uterine inertia during abortion or parturition is believed to be the result of overstretching of the abdominal and uterine musculature. Shock is believed to occur as a result of pooling of fluid in splanchnic vessels when the abdominal pressure is released.2, 12 The prognosis for hydrops allantois and hydrops amnion is guarded to poor with respect to a favorable pregnancy outcome or even a full-term gestation. Close monitoring of the mare and fetus and intensive treatment of both after delivery can result in a viable foal after hydrops allantois1 or hydrops amnion.7 Prognosis for survival of the mare after hydrallantois is good if measures are taken to prevent or treat shock, retained placenta, delayed uterine involution and other complications such as laminitis.2, 9 The prognosis for future use of the patient as a brood mare is favorable; although hydrops may occur in subsequent pregnancies,14 this is
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Hydrops uncommon and affected mares can be rebred to produce normal foals.1, 2 It is difficult to formulate a prognosis for hydramnion because of the small number of equine cases that are on record.
Treatment Once a hydrops condition has been confirmed, a logical treatment is the induction of abortion or labor to relieve pressure on internal organs and to avoid further problems, such as rupture of the prepubic tendon, especially if the uterus is severely distended to a level above the pelvic brim, and obstructs the pelvic canal. The decision to end the pregnancy is easy when the mare has signs of discomfort, is anorexic, and has respiratory distress when lying down. Various protocols have been used to induce abortion in mares with hydrops. The easiest method is to perforate the allantochorion after manual dilation of the cervix.4, 12, 13 If the cervix is tightly closed, the administration of an estrogenic drug10 or a prostaglandin14 the day before may facilitate cervical dilation. Oxytocin given intravenously in small repeated amounts of 2.5 to 5 units, or 50 units administered intramuscularly, or intravenously over 1 hour, can be attempted, although the latter regimens did not cause abortion in all mares with hydrops.2 Perforation of the chorioallantoic membrane is usually achieved digitally but may require an instrument such as a teat cannula or blunt scissors. Surprisingly, the chorioallantois does not bulge through the dilated cervix, nor does the fetal fluid always escape under great pressure after puncturing the allantochorion.2, 14 This is because of uterine inertia. The goal is a slow and gradual release of the fetal fluid so that the vascular system of the mare can adapt and shock be avoided. If fluid does not escape spontaneously, it may be removed by siphoning, and a stalled delivery restarted by the administration of oxytocin. In an attempt to prevent or combat hypovolemic shock during or following the release of fetal fluid, pretreatment of the mare with corticosteroids and/or placement of one or two large-bore jugular vein catheters may be indicated; fluid can then be administered quickly should this become necessary. Once a certain amount of fetal fluid has escaped from the uterus, the fetus or the fetuses can be identified within the uterus and extracted. The fetus is normally alive at delivery but is not expected to survive.2 Complications may occur with the presence of skeletal malformations of the fetus such as limb deformities, torticollis or ascites.2 The mare is carefully monitored for placental expulsion; retained placenta is common after hydrops because of uterine inertia and, possibly, placental immaturity or changes such as thickening and edema.2 The normal placenta of the mare weighs
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11% of the fetal weight26 but in two mares with hydrallantois, it weighed 18 and 33%14 and, in a mare with hydramnion, 24% of the fetal weight.7 The reasons for this are edema or cystic changes of the placenta.2 Alternatively, when the condition of the mare is not as severe and not life-threatening, or if the potential value of the foal precludes the induction of abortion, the mare and fetus can be monitored and the mare treated supportively by applying abdominal and limb support wraps and frog pads. Nonsteroidal anti-inflammatory drugs and progesterone may also be given. The calcium concentration of the mammary secretion must be monitored to detect fetal readiness for birth,27 at which time treatment with progesterone is discontinued. The monitoring of the fetus is done via ultrasonography and is based on various factors that are related to fetal well-being and pregnancy outcome. These factors make up an equine biophysical profile and include fetal heart rate, aortic diameter, maximum fetal fluid depth, uteroplacental thickness and contact, and fetal activity.23, 28, 29 These factors have been used to monitor a late-term fetus of a hydramnion pregnancy that went to term and produced a viable foal.7
The fetus Hydrallantois has occurred in mares in association with abnormal fetal positions, placental abnormalities, twins and deformed or non-viable foals;2, 10, 12 in one report of eight cases of hydrops, four fetuses were abnormal.2 Abnormalities seen in hydrops conditions include limb deformities and torticollis,2 hydrocephalus,2, 14 scoliosis of the vertebral column, overextended fetlocks, cerebellar and cerebral hypoplasia, internal hydrocephalus, hydranencephaly, and cerebellar aplasia,14 brachygnatia,6 and visceral herniation.4 An 8-month-old fetus aborted by a mare with hydrops allantois, seen by the author, had severe brachygnathia superior, a short lumbar region suggesting perosomus elumbis, and arthrogryphosis of the hind legs with fixation of the fetlocks. A cause–effect relationship between the observed anomalies and the dropsy condition cannot be established. In cattle, defective and anomalous fetuses are typically seen in association with hydramnion.3 Congenital anomalies, particularly those involving the brain or other parts of the head, may occur in horse fetuses associated with hydramnion6 or hydrallantois.2, 14
References 1. Koterba AM, Haibel GK, Grimmet JB. Respiratory distress in a premature foal secondary to hydrops allantois and placentitis. Comp Contin Educ Pract Vet 1983;5(3):S121–5.
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2. Vandeplassche M, Bouters R, Spincemaille J, Bonte P. Dropsy of the fetal sacs in mares: induced and spontaneous abortion. Vet Rec 1976;99:67–9. 3. Roberts SJ. Veterinary Obstetrics and Genital Diseases, 3rd edn. Woodstock, VT: Stephen J Roberts, 1986. 4. Allen WE. Two cases of abnormal equine pregnancy associated with excess foetal fluid. Equine Vet J 1986;18(3):220–2. 5. Wintour EM, Laurence BM, Lingwood BE. Anatomy, physiology and pathology of the amniotic and allantoic compartments in the sheep and cow. Aust Vet J 1986;63(7):216–21. 6. Sertich PL, Reef VB, Oristaglio-Turner RM, Habecker PL, Maxson AD. Hydrops amnii in a mare. J Am Vet Med Assoc 1994;204(9):1481–2. 7. Christensen BW, Troedsson MHT, Murchie TA, Pozor MA, Macpherson ML, Estrada AH, Carrillo NA, Mackay RJ, Roberts GD, Langlois J. Management of hydrops amnion in a mare resulting in birth of a live foal. J Am Vet Med Assoc 2006;228(8):1228–33. 8. Wintour EM, Barnes A, Brown EJ, Hardy KJ, Horacek I, McDougall JM, Scoggins BA. The role of deglutition in the production of hydramnios. Theriogenology 1977;8(4):160. 9. Honnas CM, Spensley MS, Laverty S, Blanchard PC. Hydramnios causing uterine rupture in a mare. J Am Vet Med Assoc 1988;193(3):334–6. 10. Henry MM, Morris DD, Pugh DG. Hydrallantois associated with twin pregnancy in a mare. Equine Pract 1991;13(7):20–3. 11. Arthur GH. The fetal fluids of domestic animals. J Reprod Fertil Suppl. 1969;9:45–52. 12. Blanchard TL, Varner DD, Buonanno AM, Palmer JE, Hansen TO, Divers TJ. Hydrallantois in two mares. J Equine Vet Sci 1987;7(4):222–5. 13. Blanchard TL. Managing the mare with hydrallantois. Equine Pract 1989;84:790–2. 14. Waelchli RO, Ehrensperger F. Two related cases of cerebellar abnormality in equine fetuses associated with hydrops of fetal membranes. Vet Rec 1988;123:513–14. 15. Perkins NR, Frazer GS. Reproductive emergencies in the mare. Vet Clin N Am Equine Pract 1994;10(3):643–70. 16. Jones SL, Fecteau G. Hydrops uteri in a caprine doe pregnant with goat-sheep hybrid fetuses. J Am Vet Med Assoc 1995;206(12):1920–2. 17. Skydsgaard JM. The pathogenesis of hydrallantois bovis. Acta Vet Scand 1965;6:193–207.
18. Dickerson JWT, Southgate DAT, King JM. The origin and development of the hippomanes in the horse and zebra. J Anat 1967;101(2):285–93. 19. Williams MA, Schmidt AR, Carleton CL, Darien BJ, Goyert GL, Sokol RJ, Derksen FJ. Amniotic fluid analysis for ante-partum foetal assessment in the horse. Equine Vet J 1992;24(1):236–8. 20. Williams MA, Wallace SS, Tyler JW, McCall CA, Gutierrez A, Spano JS. Biochemical characteristics of amniotic and allantoic fluid in late gestational mares. Theriogenology 1993;40:1251–7. 21. Paccamonti D, Swiderski C, Marx B, Gaunt S, Blouin D. Electrolytes and biochemical enzymes in amniotic and allantoic fluid of the equine fetus during late gestation. Biol Reprod 1995;Monograph 1:39–48. 22. Tillmann H. Die Fruchtw¨asser. In: Rosenberger G, Tillman H (eds) Tiergeburtshilfe. Berlin: Paul Parey, 1978; pp. 50–2. 23. Reef VB, Vaala WE, Worth LT, Sertich PL, Spencer PA. Ultrasonographic assessment of fetal well-being during late gestation: development of an equine biophysical profile. Equine Vet J 1996;28(3):200–8. 24. Schmidt AR, Williams MA, Carleton CL, Darien BJ, Derksen FJ. Evaluation of transabdominal ultrasound-guided amniocentesis in the late gestational mare. Equine Vet J 1991;23(4):261–5. 25. Schott HC, Mansmann RA. Biochemical profiles of normal equine amniotic fluid at parturition. Equine Vet J Suppl. 1988;5:52. 26. Schlafer DH. The equine placenta before and after birth: recognition of normal features and appreciation of lesions of clinical importance. In: Proceedings, Mare Reproduction Symposium, Society for Theriogenology, 1996; pp. 122–37. 27. Ousey JC, Dudan F, Rossdale PD. Preliminary studies of mammary secretions in the mare to assess foetal readiness for birth. Equine Vet J 1984;16:259–63. 28. Adams-Brendemuehl C. Fetal assessment. In: Koterba AM, Drummond WH, Kosch PC (eds) Equine Clinical Neonatology. Philadelphia: Lea & Febiger, 1990; pp. 16–33. 29. Reef VB, Vaala WE, Worth LT, Spencer PA, Hammett B. Ultrasonographic evaluation of the fetus and intrauterine environment in healthy mares during late gestation. Vet Radiol Ultrasound 1995;36(6):533–41.
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Chapter 244
Fetal Maceration and Mummification Claire E. Card
Definition of fetal maceration Maceration of a fetus may develop when a fetus dies in utero but is not expelled because of uterine inertia, fetal malpositioning, inadequate cervical dilation, or other pathological process.1 Subsequent bacterial contamination results in fetal autolysis, but in fetuses that are greater than 3 months of gestation, ossification prevents skeletal resorption. In mares, uterine retention of a macerated fetus is uncommonly associated with systemic illness despite the presence of bacteria and purulent material. Bacterial contamination with Gram-negative organisms would be expected to be associated with the production of endotoxins during fetal degeneration. It is probable that the endometrium heals quickly after placental separation, and the intact endometrium prevents systemic absorption. Most macerated fetuses arise from midgestational losses.2
Diagnosis and predisposing causes of fetal maceration The diagnosis of fetal maceration is based on clinical findings following examination using: rectal palpation, ultrasonography, endometrial cytology/culture, and hysteroscopy. A persistent or bloody vaginal discharge has been reported along with a crepitant uterine horn with dense hyperechoic fluid and linear hyperechoic structures in a mare with a macerated fetus. Fetal bones may be identified using ultrasonography or hysteroscopy. In one case the fetal bones were adhered to the endometrium with inflammatory debris.2 The cause of fetal death is seldom identified but may include: cervical abnormalities, ascending placentitis, incomplete cervical dilation preventing ex-
pulsion of all fetal parts, or dystocia resulting from abortion of twins or an anomalous fetus. In cattle, fetal maceration and pyometra are associated with anestrus, cervical closure, and persistence of a corpus luteum. There are few reports of ovarian status during retention of a macerated fetus in a mare; however, one mare was reported to show estrus, was bred, and had evidence of a recent ovulation. It has been speculated that a mare with a macerated fetus may show anestrus, or perhaps have behavioral signs similar to those seen with pyometra, where estrus intervals can be absent, shortened, lengthened, or normal.
Definition of fetal mummification Fetal mummification follows the death of a mid- to late-gestational fetus in a sterile environment. The fetus undergoes autolysis and becomes dehydrated, compact, and has a leathery skin. The placenta may also be mummified. Both the fetus and placenta are a dark-brown color. The fetal fluids are reabsorbed in the process of autolysis.3
Causes of fetal mummification The most common cause of fetal mummification is a twin pregnancy. The majority of surviving twin pregnancies abort between 9 and 11 months of gestation, but in rare cases one twin may mummify and the pregnancy continues. There are a few reports of single mummified fetuses in mares that had been treated with progesterone or altrenogest.3–5 Fetal mummification in twin pregnancy may be due to the limitations of placental and fetal development. Eventually the inadequate placental surface area results in compromise and subsequent death of a fetus (Figure 244.1). In these cases where mummification occurs it has been reported that some mares
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Problems of Pregnancy tion.6, 7 In these cases the bones are walled off into an inverted sac, which is attached to the placenta on the allantoic side (Figure 244.2a–c). Hormone supplementation has rarely been associated with fetal mummification; presumably its actions somehow interfere with fetal expulsion. Exogenous progestagens do not inhibit luteolysis in the mare. The process or products of mummification may have effects on the endometrium, such as inhibition of prostaglandin or proinflammatory cytokine production, which in turn affect uterine and cervical function.3
Figure 244.1 Image of an aborted mummified twin fetus.
Treatment
show premonitory signs of abortion, such as udder development, which subsequently abates, and the mare aborts or delivers the remaining fetus and a mummified fetus at a late date. Ultrasound-guided fetal thoracic or cardiac injection also has the ability to result in fetal mummification of the dead twin, as does craniocervical disloca-
Therapy is aimed at removing the autolyzed fetus from the uterus. Manual removal may be attempted if the cervix can be dilated. If the mare has elevated progesterone levels associated with a corpus luteum, prostaglandin is indicated before manual removal. Treatment with estradiol 17-β intramuscularly (i.m.)
(a)
(b)
(c)
Figure 244.2 Term placenta from a twin pregnancy where a fetal reduction by transabdominal fetal cardiac puncture was performed at around 100 days of gestation. (a) The allantoic surface of the placenta with the mummified fetal remnants in a sac in the upper left corner of the image. (b) Close-up of the fetal remnant sac. (c) Radiograph of the fetal remant sac with ossified bones.
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Fetal Maceration and Mummification the day before and/or prostaglandin E1 (misoprostol 200 micrograms), or prostaglandin E2 (various preparations) administered at least 90–120 minutes before planned removal, locally on and in the cervical canal, may also assist with cervical relaxation.8, 9 Antibiotic administration based on culture and sensitivity if there are bacteria present, or uterine lavage to remove debris, are also indicated. An endometrial biopsy is indicated in cases where the retention is longstanding or the inflammation is severe, to determine the prognosis for breeding.
References 1. Ve’pzina J, Marcoux M, Phaneuf JB. Maceration of the foetus in a mare. Can Vet J 1975;16:20–1. 2. Burns TE, Card CE. Fetal maceration and retention of fetal bones in a mare. J Am Vet Med Assoc 2000;217:878–80.
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3. McCue PM, Vanderwall DK, Squires EL. Fetal mummification in a mare. J Equine Vet Sci 1997;17:267–9. 4. Meyers PJ, Varner DD. Abortion of a mummified fetus associated with short uterine body in a mare. J Am Vet Med Assoc 1991;198:1768–70. 5. Newcombe JR, Wilson MC. Mummified singleton foetus in a mare. Aust Equine Vet 1998;16:33–4. 6. Wolfsdorf KE. Management of postfixation twins in mares. Vet Clin N Am-Equine 2006;22:713–25. 7. Ball BA, Schlafer DH, Card CE, Yeager AE. Partial reestablishment of villous placentation after reduction of an equine co-twin by foetal cardiac puncture. Equine Vet J 1993;25:336–8. 8. Volkmann DH, de Cramer KGM. Prostaglandin E2 as an adjunct to the induction of abortion in mares. In: Equine Reproduction V: Proceedings of the Fifth International Symposium on Equine Reproduction, 1991, pp. 722–3. 9. Rigby S, Love C, Carpenter K, Varner D, Blanchard T. Use of prostaglandin E2 to ripen the cervix of the mare prior to induction of parturition. Therio 1998;50:897–904.
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Chapter 245
Equine Herpesviruses Peter J. Timoney
Introduction Nine equid herpesviruses have thus far been identified, of which five, equid herpesviruses 1 to 5, are known to infect the horse.1 Of these, equid herpesviruses 1 and 4 (EHV-1 and -4) are the most important for horse industries worldwide based on their veterinary medical and economic significance.2–4 Both viruses are endemic in domesticated equid populations in the vast majority of countries.5, 6 Equine herpesvirus 1 is the more important of the two viruses, being the major cause of a multisyndromic disease, equine rhinopneumonitis. The term encompasses a range of syndromes: respiratory illness,5, 7 abortion,8, 9 a very frequently fatal pneumonitis in neonatal foals,10, 11 myeloencephalopathy,2, 4,12–15 and very rarely, a fatal, multisystemic, non-neurological syndrome.16 It is believed that EHV-1 and EHV-4 have co-evolved with horses over millions of years, in the course of which they have acquired the ability to establish a lifelong carrier state in their natural host species.2, 6 Of the remaining three herpesviruses known to infect the horse, none have the clinical impact of EHV-1 or EHV-4. Equid herpesvirus 3 is the cause of equine coital exanthema, a highly contagious though relatively benign, non-systemic, venereally transmissible disease affecting the external genitalia of the stallion and the mare.17–20 There is no evidence implicating the virus in infertility or abortion in the mare.21–23 Equid herpesviruses 2 and 5 are gammaherpesviruses that have been associated with a number of clinical entities in the horse. Upper and lower respiratory disease,24–26 keratoconjunctivitis,27, 28 chronic follicular pharyngitis, and poor performance have been linked to infection with equid herpesvirus 2.29 Recent evidence would suggest that a newly recognized lung disease, equine multinodular pulmonary fibrosis, develops in association with infection with equid herpesvirus 5.30, 31 Equid herpesviruses 2, 3, and 5 will not be given
further consideration here because they appear to lack the potential to adversely affect pregnancy and cause abortion in the mare.
Causal agents Equid herpesviruses 1 and 4 are members of the genus Varicellovirus, subfamily Alphaherpesvirusvirinae, family Herpesviridae.32 Originally considered subtypes of the one virus, EHV-1,33 they have since been shown to be closely related but distinct alphaherpesviruses, with nucleotide sequence identity within individual homologous genes ranging from 55% to 84% and amino acid identity from 55% to 96%.34, 35 Whereas the natural host range of EHV-4 is restricted to horses, EHV-1 can occasionally infect cattle, cervids, captive camelids, and laboratory mice;36–39 it has also been adapted to replicate in hamsters.40 Both EHV-1 and EHV-4 are genetically stable viruses in their natural host species, the horse.2,41–43 However, considerable sequence variation has been demonstrated in a region of open reading frame (ORF) 68 of field strains of EHV-1.44 This polymorphism has been used in molecular epidemiological studies as a marker system for assigning isolates to one of six common strain groups. Most of the glycoproteins identified with EHV-1 and EHV-4 possess epitopes that induce both type-specific and typecommon serological responses.45, 46 The exception is glycoprotein G, which elicits only a type-specific serological response in the horse.47 There is no evidence of major antigenic variation among strains of either virus, although minor antigenic differences can be detected by detailed monoclonal antibody analysis.2 In contrast to the reported lack of significant antigenic variability, individual isolates/strains of EHV-1 and EHV-4 can differ markedly in their pathogenicity for the horse.48, 49 Historical evidence has shown that electropherotypes 1B and 1P of EHV-1 can be of major significance in causing outbreaks of abortion.2, 50
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Equine Herpesviruses Equid herpesviruses 1 and 4 are readily inactivated outside the body, by detergents, disinfectants, lipid solvents, and heat. There is some evidence to indicate that either virus can survive for up to a week under cool, damp, ambient conditions.51 Anecdotally, it has been suggested that the period of viability of EHV-1 on contaminated fodder may be considerably longer, perhaps several weeks, given conditions conducive to survival of the virus.10 Differentiation of EHV-1 from EHV-4 has helped greatly in clarifying their respective roles in the various clinico-pathological syndromes comprising equine herpesviral disease or equine rhinopneumonitis. Whereas both EHV-1 and EHV-4 can cause acute febrile respiratory illness on primary infection, the majority of such cases are due to EHV-4, especially in young foals and in some countries, also in 2- and 3-year-old horses in training.3, 5, 6, 52 A frequently encountered sequel to EHV-1 and, less commonly, EHV-4 infection is abortion in the pregnant mare.3, 8, 9 Equid herpesvirus 1 is regarded as the principal cause of contagious abortion in mares in many countries.2, 6, 53 Economic losses can be very significant, especially where the virus is introduced into unprotected populations of pregnant mares. Although less frequently implicated, EHV-4 can cause sporadic cases of abortion.50, 54 Inadequately protected mares exposed very late in pregnancy to either virus, especially EHV-1, may not abort, however, but give birth to a congenitally infected live foal.10, 11, 55, 56 The third major clinical outcome primarily of EHV-1 infection is a neurological syndrome, specifically a myeloencephalopathy.12, 13, 15,57–60 It is frequently, but not invariably, caused by neuropathogenic strains of EHV-1. These are characterized by a unique point mutation in the catalytic subunit of the polymerase gene, which is located in ORF 30 of the virus.44, 61 This mutation results in a single amino acid change: asparagine in the wild-type virus (ORF30 A2254 ) becomes aspartic acid in the mutant virus (ORF30 A2254 →G). It should be noted, however, that a small percentage of outbreaks of herpesvirus myeloencephalopathy are caused by wild-type virus lacking this mutation.62 Equid herpesvirus 1 has also been linked to a fatal non-neurological syndrome characterized by a multisystemic vasculitis, severe pulmonary edema, and a mild enterotyphocolitis.16 Only a few cases of this manifestation of EHV-1 infection have been recorded to date.
Transmission Transmission and dissemination of EHV-1 and EHV-4 takes place primarily by the respiratory route through direct or indirect contact with infective nasal
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or conjunctival secretions, aborted fetuses, placental membranes and fluids, and various secretions and excretions of congenitally infected foals.3,63–65 Exposure occurs through inhalation of infective droplets or by ingestion of virus-contaminated food or water. Both EHV-1 and EHV-4 can readily spread among susceptible horses, with transmission rates approaching 100%. Clinical cases are most contagious during the first week after infection, when up to 106 plaque-forming units of virus can be isolated from swabs of nasal secretion. Viral shedding by the respiratory route occurs for 7 to 15 days in naive individuals.66 The magnitude and duration of shedding is reduced in previously infected horses or in those in which reactivation of latent virus has taken place.67 Initial infection of foals with EHV-1 or EHV-4 most often occurs either before or after weaning and is not always associated with observable signs of illness.2, 5, 6 Up to 30–60% of young horses have been infected with one or both viruses by 1 year of age. Foals most probably acquire infection from contact with a mare or mares that, it is believed, experiences reactivation of a latent infection. Transplacental transmission occurs readily in the case of EHV-1 and infrequently with respect to EHV4. This usually but not invariably results in abortion during the final trimester of pregnancy. Failure of either virus to induce abortion in mares prior to the fifth month of pregnancy has been attributed to the fact that endometrial vascular changes and viral antigen expression in endothelial cells in these individuals are significantly less than those observed in mares infected later in gestation.68 Thrombosis was rare and thrombo-ischemic lesions were absent in mares experimentally infected with EHV-1 in early pregnancy. These findings contrast sharply with those observed in mares in the final trimester of pregnancy. Abortion in such cases is associated with multifocal thrombosis and cotyledonary infarction, which presumably results in placental separation and fetal death.69, 70 It has been postulated that resistance of the mare in early pregnancy to EHV-1 abortion may be related to hormonal stability of the placenta in response to vascular injury.71 Smith72 proposed that the equine endometrium is less susceptible to such damage and to the resultant prostaglandin release in early as opposed to late gestation. A pregnant mare exposed to infection very late in gestation may not abort but instead give birth to a live foal affected with a very frequently fatal pneumonitis, presumably the result of congenital infection with the virus.2, 4, 53 Horses that are carriers of EHV-1 and/or EHV-4 can, upon reactivation of the latent virus, serve as a source of infection for inadequately protected, incontact horses. Virus replication and shedding into
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the respiratory tract takes place in such individuals for a limited period of time, usually no longer than 36 to 48 hours.67 Reactivation of latency, principally of EHV-1, in the pregnant mare may result in abortion in that individual if its pre-existing level of protective immunity is insufficient.4, 53 Should this occur, the aborted fetus and placenta can be highly productive sources of virus as previously mentioned. It should be made clear that where reactivation of latent EHV-1 occurs in a pregnant mare, aside from the potential for abortion to supervene in the individual animal, infectious virus is also shed into the nasopharynx for 36–48 hours and can serve as a source of infection for any susceptible in-contact mares. Aside from the traditionally accepted modes of transmission of EHV-1 and EHV-4, there have been a few isolated cases recorded where infectious EHV1 or viral DNA has been demonstrated in stallion semen and in an embryo from a clinically healthy donor mare.73–75 In other studies, however, viral nucleic acid could not be detected in the semen of EHV-1 seropositive stallions.76 The significance of these findings and the potential for venereal transmission of EHV-1 requires further investigation.
Pathogenesis There are significant differences between the pathogenesis of EHV-1 and that of most strains of EHV4.2–4 Whereas both viruses infect the respiratory tract epithelium and associated lymph glands, strains of EHV-1 also have a predilection for vascular epithelium, particularly in the nasal mucosa, lung, adrenal, thyroid, and uterus as well as the central nervous system (CNS), especially but not exclusively in the case of infection with neuropathogenic strains of the virus.62 In contrast, the majority of strains of EHV4 are restricted to the respiratory tract and associated lymph glands. Whereas a leukocyte-associated viremia is characteristic of EHV-1 infection, it occurs with only a limited number of strains of EHV-4.2, 4, 49
Respiratory disease Upon respiratory exposure, both EHV-1 and EHV-4 infect and multiply in the nasal and nasopharyngeal epithelium of inadequately protected individuals.77, 78 Over the following 4 to 7 days, considerable amounts of either virus are shed into the respiratory tract. Within 12 to 24 hours of infection, virus is transported via infected leukocytes to the lymph glands associated with the respiratory tract, where it undergoes further replication.78 Migration of EHV-1 and certain strains of EHV-4 ensues, with virus being transported from the lymph glands
into the bloodstream and lymphatic circulation via infected leukocytes. The resultant cell-associated viremia involves monocytes and CD5+ and CD8+ T-lymphocytes.79–81 Widespread virus dissemination follows, with additional replication taking place in the vascular epithelium of the target sites of EHV-1; these sites may also be subject to infection by a minority of strains of EHV-4.82 In addition, there is evidence that cell-to-cell infection can occur in infected blood vessels.53 A very important feature of the pathogenesis of EHV-1 and EHV-4 is the propensity of each virus to gain entry to the neuronal fibers of the trigeminal nerve innervating the nasopharyngeal mucosa and conjunctiva, and migrate centripetally to the trigeminal ganglia where each can establish latency.83–86 This may precede or coincide with onset of the cellassociated viremia. Abortion The cell-associated viremia is a key event in the pathogenesis of abortion/neonatal disease due to EHV-1 and some strains of EHV-4.2, 48 Once virus reaches the pregnant uterus, it infects the endothelial cells of the uterine vasculature and sets up an endometrial vasculitis, damaging both the endometrium and the chorioallantois.53, 71 In turn, this results in thrombosis in affected vessels and areas of infarction in the microcotyledons of the placenta.70 The extent and severity of thrombosis that occurs will determine whether abortion will supervene or not. Where thrombotic lesions are focal and not generalized, virus spreads to the fetus by infected leukocytes, causing widespread necrotic lesions in many tissues. Concurrently, local edema develops at the feto-maternal interface, resulting in premature separation of the chorioallantois from the endometrium. The outcome is abortion with fetus and placenta positive for virus. However, in cases of extensive thrombotic lesions, placental ischemia rapidly ensues with premature placental separation and abortion.69 Under such circumstances, the virus has not had sufficient time to invade the fetus, and consequently the placenta but not the fetus is positive for virus. Neonatal disease Where pregnant mares are exposed to infection with EHV-1 and certain strains of EHV-4 very close to term, they may not abort but give birth to a diseased foal, presumably the result of congenitally acquired infection.10, 11, 55 Such foals are either born weak and very frequently succumb to a viral pneumonitis within the first 24 hours of life, or are born apparently normal but develop severe respiratory distress
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Equine Herpesviruses within 18 to 24 hours and die 12 to 72 hours later. There has been one report linking mortality in foals in the first few weeks of life with EHV-1 infection.87 Although normal at birth, such foals succumbed to a variety of secondary bacterial infections within 2 to 4 weeks.
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lings.5 Older animals and those that have been previously exposed to or vaccinated against EHV-1 or EHV-4 generally display minimal if any evidence of disease.77 Younger horses develop an acute febrile respiratory illness following an incubation period ranging from 2 to 10 days (usually 1–3 days).66 Affected individuals may develop any combination or all of the following:
Myeloencephalopathy Equine herpesvirus myeloencephalopathy is usually a sequel to a primary febrile, respiratory infection or a case of herpesvirus abortion.10, 15, 60, 88, 89 Of critical significance in the development of this syndrome is the cell-associated viremia that follows EHV-1 infection2 and, very uncommonly, EHV-4 infection.90–92 This enables dissemination of virus to the vasculature of the CNS as well as other sites in the body. An important factor in determining whether neurological disease develops from this point is the property of endotheliotropism, which is inherent in certain strains of EHV-1, namely neuropathogenic or mutant strains.44, 61, 62, 93 Such strains give rise to a cell-associated viremia of earlier onset, greater magnitude, and longer duration than that observed with non-neuropathogenic or wild-type virus.94 Under such circumstances, the CNS vasculature is exposed to a considerable concentration of virus-infected leukocytes with a consequent greater risk of infection of the vascular endothelium and development of neurological disease. Thrombosis, hemorrhage, and areas of infarction ensue, changes that in turn result in secondary ischemia of affected regions of the CNS. Thrombo-ischemic damage to the neural parenchyma leads to neuronal degeneration, axonal swelling, and, in time, foci of malacia.4, 60 Vascular lesions in the spinal cord and/or brain of affected horses are suggestive of an immune-mediated vasculitis.60 Some have proposed that an anamnestic immune reaction is a prerequisite to generation of an immune-mediated vasculitis that may be involved in the pathogenesis of equine herpesviral neurological disease.53, 60 A number of virus strain-related or hostrelated factors can play a role in determining the extent and severity of the lesions observed in cases of the disease.
Clinical features Respiratory disease Respiratory disease caused by EHV-1 or EHV-4 is clinically indistinguishable.2, 3, 5, 6 Primarily an upper respiratory tract infection, it is characterized by a rhinopharyngitis and a tracheobronchitis, with signs of illness seen chiefly in weanling foals and year-
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fever (39–41◦ C), which may be biphasic; leukopenia involving both a neutropenia and a lymphopenia; serous to mucopurulent nasal discharge; conjunctivitis; anorexia; depression; occasional coughing; not infrequently, a degree of enlargement of the mandibular and even the retropharyngeal lymph glands.5
While the duration of the febrile response is usually 2 to 5 days, the nasal discharge and cough can last for 1 to 3 weeks as a result of secondary bacterial infection. Severity of the disease varies with age of the horse and level of pre-existing immunity to either EHV-1 or -4. Equine rhinopneumonitis is not lifethreatening per se, and clinical recovery from uncomplicated cases of infection is usually complete within a few weeks, although some 2–3-year-old horses may subsequently develop the ‘poor performance syndrome’.95 This has been associated with bronchial hypersensitivity and impaired ability to perform to the individual’s considered athletic potential. Abortion Abortion caused by EHV-1 and, uncommonly, EHV4, is very seldom preceded by any clinical signs of illness in the mare.3, 9, 63 Although abortion rates as high as 75% can occur,96 under conventional systems of good management such occurrences, or ‘abortion storms’, are less commonly encountered than in the past. Most cases of equine herpesvirus abortion are isolated or sporadic in nature and frequently involve maiden mares.7 The incubation period can range from 1 week to a few months, with the majority of abortions occurring within 8–9 weeks of infection, usually 15–30 days. While herpesvirus abortion can take place as early as the fifth month of pregnancy, most occur during the final trimester between the seventh and eleventh months of gestation for the reasons stated previously. The fetus is expelled fresh and is frequently enveloped in the amniotic membrane.97, 98 Some foals are still alive at time of
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expulsion but die shortly afterwards from the extensive viral-induced damage to their lungs. Abortion due to EHV-1 or EHV-4 has no adverse effect on the subsequent fertility of the mare.63 Viral antigen cannot be detected in the mare’s uterus beyond 48 hours after abortion of an infected fetus. Mares rarely abort from either virus in successive years but may abort later in their reproductive life if reinfected during pregnancy, when they may be vulnerable to the abortigenic effects of the virus.99 Neonatal disease Occasionally, foals are born that are either ill at birth or become ill within the first 1 to 2 days of life.10, 11, 55 These foals are infected with EHV-1 or, very infrequently, EHV-4. Affected foals fail to nurse, become lethargic, pyretic, leukopenic, hypoxic, and dyspneic, and develop intractable diarrhea. The clinical condition of such foals rapidly deteriorates and they usually succumb to a fulminant viral pneumonia in a few days, even if provided with intensive supportive treatment. There has been one report of herpesvirusinfected foals surviving for a few weeks before dying from a variety of secondary bacterial infections.87 Although not conclusively proven, it is believed that cases of neonatal disease are the result of mares being exposed to EHV-1 or possibly EHV-4 very late in gestation, not aborting, but giving birth to a congenitally infected foal affected with a viral pneumonitis, which is very frequently fatal.10, 11, 55, 56 Myeloencephalopathy Neurological disease is a sporadic, though no longer an uncommon outcome of infection with EHV-1 and, rarely, some strains of EHV-4.12–15,59, 91, 92 Recognized clinically for many years, the disease can be associated with a high morbidity and case-fatality rate. 44, 62, 93 Frequently but not invariably, cases of equine herpesvirus myeloencephalopathy are a sequel to an episode of respiratory disease, abortion, or febrile illness.10, 15, 59, 88, 89 During an extensive outbreak of the disease at a university equestrian center in 2003, the onset of neurological signs ranged from 1 to 4 days after the end of the febrile period.59 Equine herpesvirus myeloencephalopathy can supervene in mares within a week of their aborting due to EHV-1 infection. Individual cases or disease outbreaks have also been reported in horses that displayed no premonitory signs of herpesvirus-related infection; however, some such instances have been associated with clinical evidence of herpesvirus disease in one or more cohort animals. The disease can occur in horses of either gender and any age, and it is most often observed in late winter, spring, or early summer.2, 58 It is more common
in horses previously infected with EHV-1. Viral reactivation and concomitant nasal shedding of neuropathogenic strains by latently infected carrier animals is believed responsible for initiating some outbreaks of neurological disease.100 After an incubation period of 1 to 10 days, illness is rapid in onset and reaches maximal severity within 2 to 3 days.53 Clinical signs are very variable, depending on the distribution and extent of the lesions in the CNS. Fever of short-term duration may or may not precede the onset of neurological signs. The degree of neurological dysfunction ranges from a mild, temporary ataxia, frequently of the hindlimbs, to impairment of proprioceptive reflexes, paresis, toe-dragging of the hindlimbs, bladder atony with urinary retention or incontinence, fecal retention, sensory deficits of the perineal area, which may extend to the inguinal region and part of the hindlimbs, decreased tail tone, and limb edema, to very severe cases where there is posterior paraplegia or even quadriplegia, recumbency, and death. Neurological deficits may be symmetric or asymmetric. Some affected individuals may exhibit cranial nerve deficits, developing a head tilt, tongue weakness, nystagmus, and even blindness. While the prognosis for non-recumbent horses is usually very good, it is guarded to poor for those that remain recumbent for 2 to 3 days or longer. Even if such severely affected animals survive with intensive supportive care, they may be left with permanent residual deficits. The period required to achieve full clinical recovery can range from days to weeks to many months. Case-fatality rates in outbreaks of equine herpesvirus myeloencephalopathy can range from 1% to as high as 50%. Pulmonary vasculotropic infection The term refers to a rarely encountered multisystemic, non-neurological syndrome in young adult horses that has been associated with infection with EHV-1.16 It is a peracute disease with a fatal outcome, and is characterized by a high fever, anorexia, severe depression, respiratory disease, and a very short clinical course. Death may supervene without prior detection of clinical signs. A multisystemic vasculitis, especially of the smaller vessels in the lungs, is the primary pathological feature of the disease. Other clinical outcomes of EHV-1 infection Equine herpesvirus-1 infection of incompletely protected or naive stallions, may have a temporary adverse effect on their fertility. Scrotal edema together with prolonged pyrexia can result in reduced libido and a decrease in the percentage of normal sperm.58.101
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Equine Herpesviruses An uncommon outcome of EHV-1 infection can be ocular disease involving uveitis and a chorioretinitis.101 Observed in foals, the chorioretinal lesions may develop up to 5 weeks after respiratory infection.102 Focal and diffuse lesions have been described, which if severe may lead to permanent impairment of vision and even blindness.
Epidemiology Equine herpesvirus 1 and 4 infections are commonplace in horse populations virtually worldwide.5, 6 The two viruses are highly contagious, with the result that equine herpesvirus respiratory disease occurs annually wherever horses are congregated. Although both viruses can infect horses of all ages, signs of respiratory illness are observed primarily in weanlings and yearlings.2, 5, 52, 103 Since immunity to either infection is short term, horses experience reinfections repeatedly throughout their life. Significant factors known to be involved in the epidemiology of EHV1 and EHV-4 include: virus characteristics, modes of transmission, carrier state, immune status of at-risk populations of equids, pregnancy status of mares, and a variety of management practices that can influence the incidence and severity of the disease syndromes caused by either of these viruses. Virus characteristics Differences in the biological properties of EHV-1 and EHV-4 are responsible for dissimilarities not only in the pathogenesis but also in the epidemiology of the respective infections.2 The cell-associated viremia that is characteristic of EHV-1 but only infrequently encountered with strains of EHV-4 is the major determinant of the role each virus plays in causing abortion. Historically, abortion was believed to be the result of contact of inadequately protected pregnant mares with cases of equine rhinopneumonitis in preweaning or weanling foals.7 This view has since been modified based on the finding that the vast majority of herpesvirus infections in foals are caused by EHV-4, which is implicated only infrequently in sporadic cases of abortion. There is evidence that certain strains of EHV-1 and, much less frequently, EHV-4, vary in their pathogenic potential to cause abortion, neonatal disease, or myeloencephalopathy.48, 49 Electropherotypes 1B and 1P of EHV-1 have been especially significant in outbreaks of abortion over the years.2, 50 Of major importance is the more recent discovery of a single mutation in the viral polymerase gene of particular strains of EHV-1 that results in a mutant virus with enhanced neuropathogenic potential.44, 61 As previously stated, such strains possess
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enhanced replicative properties resulting in tenfold higher virus levels than the wild-type virus in the upper respiratory tract, blood leukocytes, and vascular endothelium.93, 94 They are capable of causing a cellassociated viremia that is earlier in onset, of greater magnitude, and longer in duration than that of nonneuropathogenic strains or wild-type virus. The findings of a retrospective study of outbreaks of equine herpesvirus myeloencephalopathy recorded since 1970, confirmed that the frequency of the disease has increased considerably in North America since 2000, with the vast majority of occurrences (24/26) caused by strains of the mutant virus belonging to a genetic substrain of EHV-1.44, 62 Outbreaks have been characterized by a high clinical attack rate and a significant case-fatality rate. The dramatic resurgence of large-scale occurrences of EHV-1 neurological disease and the apparent evolving increase in pathogenicity of the causal agent have led to the US Department of Agriculture designating the hypervirulent, mutant type of EHV-1 as a potentially emerging equine pathogen.104 There is every reason to believe that this trend is likely to continue as neuropathogenic strains of EHV-1 become more widely established in equine populations in different countries related to the international movement of horses. Of additional significance in explaining the increase in frequency of outbreaks of EHV-1 myeloencephalopathy is the failure of currently available vaccines to protect against this clinical manifestation of herpesvirus infection.62 By comparison, herpesviral abortion has been effectively controlled for many years in well-managed populations of broodmares through a combination of sound management practices and prophylactic vaccination against the disease. Transmission The principal modes of transmission of EHV-1 and EHV-4 have already been described.3, 64, 65 As previously indicated, both viruses are highly contagious and can spread rapidly among susceptible horses, primarily through direct or indirect contact with the respiratory secretions of virus-shedding individuals. These may be horses acutely infected with either virus or those that may have experienced reactivation of a latent infection. Carrier state The primary natural reservoir of EHV-1 and EHV-4 is the carrier horse that is latently infected with one or both viruses.86,105–107 After the initial lytic phase of infection in the respiratory tract, EHV-1 and EHV-4 enter a latent state in CD8+ T-lymphocytes, both in
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Life Cycle of EHV-1 (Central Role of Latency)
Latently Infected Carrier Horse Latency Establishment
Latency Reactivation
Recruitment of New Hosts into Cycle
Nasal Shedding Pregnant Mares Infection of Young Horses
Abortion Figure 245.1 The life cycle of equid herpesvirus 1 (EHV-1). Figure devised by Dr G.P. Allen.
circulation and in the lymph nodes draining the tract. 108 Additionally, both viruses can become latent in the sensory nerve cell bodies in the trigeminal ganglia.85 The carrier state is believed to be lifelong. A number of surveys have shown that the carrier rate in adult horses can be as high as 40–60%, with up to one-third of persistently infected horses harboring both EHV-1 and EHV-4 in a latent state.85 In a study of 132 necropsied mares in central Kentucky, latency of the wild type of EHV-1 was confirmed in 46% of the animals tested compared to 8% for the mutant or neuropathogenic strains of the virus.109 The natural reservoir of latently infected horses is a major source of infection for young or naive horses early in life2, 5 (Figure 245.1). Horses harboring latent EHV-1 and EHV-4 experience periodic episodes when infectious virus is reactivated from its quiescent state and is shed in the respiratory secretions, thus serving as a potential source of infection for any in-contact susceptible horses.83, 84, 105 Reactivation of latency can be triggered by a variety of environmental factors including but not exclusive of transportation, handling, change in accommodation, overcrowding, administration of large doses of corticosteroids, and the stress of training and performance. Horses that experience reactivation are invariably asymptomatic and are undetectable under conventional conditions of management. Reactivation of latency in a carrier mare that is pregnant may be a source of virus that results in abortion in that individual or in other in-contact mares, as previously mentioned.53 Latency and reactivation of latent infection are considered to play very important roles in the epidemiology of EHV-1 and EHV-4.
Immunity As protective immunity following infection with either EHV-1 or EHV-4 is short-lived, reinfection can occur every 3 to 6 months during a horse’s life.2, 4, 99, 110 Although very little cross-immunity develops between the two viruses after primary infection of the respiratory tract, significant crossprotection against EHV-4 can evolve following repeated infections with EHV-1.5,111–113 Mucosal immunity at the site of entry of EHV1 and EHV-4 into the body, namely the respiratory epithelial surface, is of primary importance in preventing the disease syndromes comprising equine rhinopneumonitis, especially those involving a blood leukocyte-associated viremia. Mucosal humoral and cellular immunity provide the first line of defense in affording protection against equine herpesvirus abortion and myeloencephalopathy.53 Cell-mediated immunity is of considerable importance in resistance to infection with EHV-1 or EHV-4.114 It has been shown that the levels of pre-existing EHV-1-specific cytotoxic T-lymphocyte precursor cells are critical in determining whether horses are protected against EHV-1-related diseases.62 There is evidence that the presence of functional cytotoxic T-lymphocytes with specific cytolytic activity against EHV-1 can prevent the development of abortion or neurological disease by controlling the magnitude of the post-infection leukocyte-associated viremia. While reinfections with EHV-1 or EHV-4 are invariably subclinical, they do give rise to limited viral replication, respiratory shedding, and, in the case of EHV-1 and infrequently EHV-4, a leukocyte-associated viremia and the
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Equine Herpesviruses potential for abortion, neonatal disease, or neurological disease to ensue.4, 53, 67 Pregnancy There is little doubt that the uterus of the pregnant mare infected with EHV-1 or EHV-4 between 7 and 11 months of gestation functions as a very efficient amplifier of either virus. The fetus, placenta, and reproductive tract secretions of the mare immediately following abortion due to EHV-1 are very important sources of infection.64 Direct or indirect contact with these materials is often implicated in viral spread on an affected farm or to adjacent premises. Management practices A range of management-related practices can influence the epidemiology of EHV-1 or EHV-4 infections. 5, 115, 116 Included among the more significant are:
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the control measures taken to restrict the introduction of infection onto premises; how pregnant mares are managed, especially after the first trimester of pregnancy; whether steps are taken to minimize the risk of management-related stress and the potential for reactivation of latency in the carrier animal; whether at-risk horses are kept on a program of regular immunization against both infections.63
Overcrowding and addition of mares during the final months of the foaling season favor reactivation of latency, widespread dissemination of EHV-1, and the likelihood of multiple cases of abortion. The incidence of abortion is higher in mares transported during late pregnancy or in those that have recently gone through a sale.2, 53
Economic significance Infection with EHV-1 and, to a lesser extent, EHV4 can result in significant economic loss.3, 117 Outbreaks of abortion, death in neonatal foals, and illness and deaths associated with outbreaks of equine herpesvirus myeloencephalopathy are the principal sources of financial loss experienced by the equine industry. Not to be overlooked in the case of outbreaks of herpesvirus neurological disease at performance venues such as racetracks, are the additional losses incurred due to cancellation of event entries, disruption of training, and restrictions on movement from affected premises.104
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Diagnosis Viral rhinopneumonitis caused by EHV-1 or EHV-4, no matter how suggestive the clinical features and the age group involved, cannot be differentiated on clinical grounds alone from equine influenza, equine viral arteritis, or certain other respiratory infections of the horse.5, 6 A diagnosis can be confirmed by virus isolation from nasal or nasopharyngeal swabs, and from the buffy coat of unclotted blood samples in the case of EHV-1 and specific strains of EHV-4, or by detection of viral nucleic acid by the polymerase chain reaction (PCR) assay.6 In view of significant differences in the pathogenesis and epidemiology of EHV1 and EHV-4, it is very important to determine which virus type is involved in outbreaks of respiratory disease.2, 5 Testing of paired (acute and convalescent) sera collected 2 to 4 weeks apart by the camflement fixation test can provide serological confirmation of recent equine herpesvirus infection.6 Determination of whether EHV-1 or EHV-4 is involved is possible using type-specific enzyme-linked immunosorbent assays (ELISAs). Diagnosis of equine herpesvirus abortion is based upon the presence of characteristic pathological lesions in the fetus and/or the placenta,7 virus detection by isolation in cell culture and/or by PCR, and demonstration of viral antigen in frozen or formalinfixed placental and fetal tissues by immunofluorescence or indirect immunoperoxidase staining.6, 53 Appropriate specimens for virus isolation, virus detection by PCR, and histopathological or immunohistochemical examination include lung, liver, adrenal, thymus or spleen, and chorioallantois. Whereas serological examination of sera from mares that have aborted is of little diagnostic value, the converse is the case with respect to serum from aborted fetuses or stillborn foals, or pre-colostral samples from cases of equine herpesvirus neonatal disease, which may have significant levels of virus-neutralizing antibodies.118 Of diagnostic assistance in the field in helping to differentiate herpesvirus abortion from that caused by equine arteritis virus is the fact that in the latter infection, the mare may have exhibited clinical signs of equine viral arteritis prior to aborting.119 Furthermore, fetuses infected with equine arteritis virus very seldom display any gross lesions, in contrast to those infected with EHV-1 or EHV-4, which frequently exhibit a range of lesions strongly indicative of this infection. Antemortem diagnosis of equine myeloencephalopathy is based upon presence of clinical signs consistent with the disease together with detection of EHV-1 DNA by PCR in nasal or nasopharyngeal swabs or washings, circulating blood leukocytes, or cerebrospinal fluid.62 Virus isolation and serological
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examination of paired sera and/or cerebrospinal fluid are of limited diagnostic value. It is highly advantageous for guiding the planning and implementation of appropriate strategies for controlling outbreaks of equine myeloencephalopathy, to employ a real-time PCR assay that can not only confirm the presence of EHV-1 DNA but also determine whether the horses are infected with the wild-type or mutant (neuropathogenic) strains of the virus.120 On postmortem examination, confirmation of a diagnosis of the disease is based upon demonstration of the characteristic lesions of vasculitis in the white matter of the spinal cord or brain with resultant hemorrhage, thrombosis, and secondary ischemic degeneration.90, 121, 122 The presence of equine herpesviral antigen in vascular endothelium of affected small arteries and veins is additional confirmatory evidence of equine myeloencephalopathy.123 Among the range of diseases that need to be considered in the context of a differential diagnosis are rabies, equine protozoal myeloencephalitis, upper motor neuron disease, Borna disease, botulism, any of the equine viral encephalomyelitides, mechanical injuries, and the wobbler syndrome.124 Specimens from a suspected case of equine herpesvirus disease should be transported to an appropriate laboratory with minimum delay. Samples should be kept refrigerated, not frozen, to optimize the chances of virus isolation. Detection of latent EHV-1 or EHV-4 is not possible using classical diagnostic methods for these infections. In the live animal, detection of latent EHV-1 in peripheral blood leukocytes can be attempted using PCR detection of latency associated transcript (LAT) sequences.108 While nested PCR and co-cultivation procedures have been employed with some success in demonstrating latent infection in lymphatic gland or trigeminal ganglionic tissues removed at necropsy examination,106, 107, 125, 126 recent reports have shown that detection of latency in antemortem or postmortem tissues can be greatly enhanced using a magnetic bead, sequence-capture nested PCR assay.100, 109
Treatment There is no specific treatment currently available for equine herpesvirus respiratory disease.5 Nonsteroidal anti-inflammatory drugs (NSAIDs) can be administered to control inflammation and fever. Antibiotic treatment is indicated in horses with a mucopurulent nasal discharge or pulmonary involvement. Rest during the acute febrile phase of the infection and for a short period thereafter is recommended to minimize secondary bacterial complications. There is no treatment to prevent abortion in pregnant mares
infected with EHV-1 or EHV-4. Neonatal foals affected with herpesvirus pneumonitis usually succumb despite intensive symptomatic treatment. The efficacy of antiviral therapy with the analogs of aciclovir (acyclovir) has yet to be fully evaluated and proven. A range of drugs including but not exclusive of corticosteroids, dimethyl sulfoxide, and NSAIDs are used to control and promote resolution of the inflammatory response in the CNS of cases of equine herpesvirus myeloencephalopathy.60 Additionally, electrolyte and fluid replacement therapy and broadspectrum antimicrobials are indicated in recumbent horses due to the potential for dehydration, pneumonia, and cystitis. Intensive supportive care is required if horses that are recumbent for 2 to 3 days or longer are to stand a chance of surviving. Full clinical recovery can take an extended period of time in such cases.
Prevention and control To reduce the veterinary medical and economic consequences of equine herpesvirus-related diseases, current control programs are focused on limiting the incidence and clinical severity of the various clinical syndromes caused by EHV-1 or EHV-4 and restricting the spread of infection during outbreaks of disease.5, 63, 115, 116 Prevention and control can best be achieved through implementation of sound management practices integrated with a regular program of prophylactic vaccination against either virus. It is important to emphasize that vaccination per se is not a substitute for employing good management practices.2, 4, 53 Management practices Critical to prevention of outbreaks of equine herpesvirus diseases, namely respiratory disease, abortion, neonatal pneumonitis, and myeloencephalopathy, is reducing the risk of introduction of either EHV-1 or EHV-4 infection onto a farm or other type of facility. A special source of risk in the case of breeding farms are mares in late pregnancy coming from a saleyard or from abroad. Risk reduction can only be achieved by requiring a period of temporary isolation of all new arrivals onto the premises, during which they are kept under observation. The recommended period of isolation with respect to breeding farms is 3 to 4 weeks. Horses on breeding farms should, where feasible, be maintained in smaller, physically isolated groups to reduce the risk of introduction of infection and also to minimize the extent of an outbreak if one occurs.96 In the case of pregnant mares, it is especially important that they be segregated from weanlings
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Equine Herpesviruses and other horses, divided into groups by stage of gestation, and remain so until all the mares in a group have foaled.2, 63 If pregnant mares are removed from their group and subsequently return to the farm, they should not be introduced into their original group. Subject to the availability of facilities, pregnant maiden mares should not be mixed with older in-foal mares. Any foster mares introduced onto premises should not be allowed contact with pregnant mares. Additional to the foregoing, it is important to minimize management-related practices that are stressful to sizeable groups of horses.2, 4, 63 These include: overcrowding, disruption of established social structures, transportation over long distances, intercurrent disease, relocation, comingling horses from different sources at sales, training stables, vaccination frequency, and abrupt en masse weaning. Doing so reduces the possibility of stress-related reactivation of latent infection in carrier animals. Under the conditions that prevail at racetracks, show events, training centers, and boarding/riding establishments, it is much more difficult to mitigate the risk of introduction and to control the spread of EHV-1, specifically the neuropathogenic strains of the virus, than it is on breeding farms.93 The densely populated, pressurized, highly stressful environment that is commonplace at performance events and training facilities runs counter to being able to effectively reduce the risk of occurrences of herpesvirus myeloencephalopathy. At the very least, the management of such facilities/events should have an established plan setting out the entry requirements for horses, including the ability to provide temporary isolation and observation of all new arrivals, together with a rapid response plan were an outbreak of the disease to occur. In the event of a suspected outbreak of herpesvirus, abortion, or myeloencephalopathy, all affected animals and in-contacts should be promptly isolated.2, 4, 115 The importance of early recognition of a disease problem cannot be overemphasized, if spread of infection is to be rapidly curtailed. In the case of abortion or pneumonitis in a neonatal foal, appropriate sanitary measures must be taken to contain the infection. The foaling stall or site where the abortion occurred should be thoroughly disinfected and any bedding preferably burnt. The fetus and placenta should be transferred to a plastic bag for submission to a diagnostic laboratory. The hindquarters and perineal region of the aborting mare should be washed down with an appropriate disinfectant and the mare isolated for a recommended period of 4 weeks. If feasible, all in-contact pregnant mares should be divided into smaller groups and kept isolated until they foal. Considerable care should be observed when han-
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dling potentially infectious material to avoid the possibility of indirect transfer of infection via contaminated clothing, equipment, and utensils. Movement of at-risk horses off a premises on which a case of herpesvirus abortion has been confirmed should be restricted for one month. Vaccination Prevention and control of equine herpesvirus respiratory disease and abortion is based on a program of prophylactic vaccination that is focused on induction of herd immunity against EHV-1 and EHV-4 in conjunction with sound, time-tested management practices.5, 63, 115, 127 To be fully successful in conferring protection against any of the disease syndromes caused by either virus, a vaccine must be capable of: (i) controlling viral replication in and dissemination from the respiratory tract very early in the course of infection; (ii) effectively neutralizing any progeny virus that is released into the respiratory secretions, interstitial spaces, and blood or lymphatic circulations; and (iii) suppressing the leukocyte-associated viremia that plays a critical role in the pathogenesis of abortion and myeloencephalopathy.53 None of the currently available equine herpesvirus vaccines comprising modified-live and inactivated products meets all of these criteria. As a consequence, they have limited effectiveness in conferring protection against respiratory disease and abortion. Presently, there is no licensed product that has proven efficacy in preventing equine herpesvirus myeloencephalopathy.62, 93 Adult horses respond to immunization with most available herpesvirus vaccines by developing high serum antibody titers against EHV-1/EHV-4.127–132 Responses of lesser magnitude are observed in young immunologically naive foals.133, 134 Cellular immune responses, both proliferative and cytotoxic, have been reported for a limited number of vaccines.131, 134 Although none of the approved vaccines is currently fully protective against herpesvirus respiratory disease and abortion, vaccination does reduce the incidence of these diseases, their clinical severity, and the duration and quantity of virus shed.33 In spite of their relative efficacy, vaccines are an integral and important component of prevention and control programs against EHV-1- and EHV-4-related respiratory disease and abortion.5, 53, 115, 127 Frequent revaccination is necessary to maintain adequate levels of immunity.3, 10 Broodmares should be vaccinated during the latter two trimesters of pregnancy or at 5, 7, and 9 months of gestation, to provide protection during the period when the unborn foal is most vulnerable to infection with the virus. It is deemed advisable to vaccinate foals with modified live or inactivated
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vaccines initially at 4–6 months of age followed by a second injection 4–6 weeks later and a third dose at 10–12 months of age.135 Revaccination is recommended at 6-monthly intervals. Irrespective of the vaccine selected, all categories of horses on a premises, especially pregnant mares, should be included in any program of regular vaccination against equine rhinopneumonitis as advised by the farm veterinarian. Efforts are continuing to develop more effective vaccines against the various clinical syndromes caused by EHV-1 or EHV-4.136 This is especially important with respect to herpesvirus myeloencephalopathy, for which presently there is no product with a manufacturer’s claim of efficacy in preventing this disease.62, 93 Aside from safety, it is hoped that future equine herpesvirus vaccines will provide a high level of protection against clinical disease, stimulate a long-lasting immunity, prevent establishment of viral latency, and are easy to administer. At the present time, maximal protection against EHV1/EHV-4-related disease is best achieved through observance of sound management practices implemented in conjunction with a regular program of immunization against both infections.3–5, 53
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Acknowledgements This chapter is dedicated in memory of the late Dr George Allen, former Office International des Epizooties (OIE) designated international expert on equine rhinopneumonitis, long-time friend, and wonderful colleague, whose research furnished much of the information on equine herpesvirus 1 and 4 infections on which this chapter is based, including the schematic representation of the life cycle of equine herpesvirus 1 in Figure 245.1.
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References 22. 1. Roizman B, Desrosiers RC, Fleckenstein B, Lopez C, Minson AC, Studdert MJ. Herpesviridae. In: Murray FA, Faucet CM, Bishop DHL, Ghabrial SA, Jarvis AW, Matelli GP, Mayo MA, Summers MD (eds) Virus Taxonomy. Sixth Report of the International Committee on Taxonomy of Viruses. New York: Springer-Verlag, 1995; pp. 114–27. 2. Allen GP, Bryans JT. Molecular epizootiology, pathogenesis, and prophylaxis of equine herpesvirus-1 infections. Prog Vet Microbiol Immunol 1986;2:78–144. 3. Bryans JT, Allen GP. Equine viral rhinopneumonitis. Rev Sci Tech OIE 1986;5:837–47. 4. Bryans JT, Allen GP. Herpesviral diseases of the horse. In: Wittmann G (ed.) Herpesvirus Diseases of Cattle, Horses and Pigs. Boston: Kluwer, 1989; pp. 176–229. 5. Allen GP. Respiratory infections by equine herpesvirus types 1 and 4. In: Lekeux P (ed) Equine Respiratory Diseases. Ithaca: International Veterinary Informa-
23.
24.
25.
26.
27.
tion Service, 2002 (online publication); http://www.ivis. org/special books/Lekeux/allen/chapter frm.asp?LA=l. Allen GP. Equine rhinopneumonitis. In: OIE Biological Standards Commission (ed.) OIE Manual of Standards for Diagnostic Tests and Vaccines, 6th edn. Paris: Office International des Epizooties,. 2008; pp. 894–903. Doll ER, Bryans JT. Epizootiology of equine viral rhinopneumonitis. J Am Vet Med Assoc 1963;142:31–7. Dimock WW. The diagnosis of virus abortion in mares. J Am Vet Med Assoc 1940;96:665–6. Dimock WW, Edwards PR, Bruner DW. Equine virus abortion. Kentucky Agricultural Experiment Station Bulletin 1942;426:1–20. Campbell TM, Studdert MJ. Equine herpesvirus type 1. Vet Bull 1983;53:135–46. Dixon RJ, Hartley WJ, Hutchins DR, Lepherd EE, Feilen C, Jones RF, Love DN, Sabine M, Wells AL. Perinatal foal mortality associated with a herpesvirus. Aust Vet J 1978;54:103–5. Bitsch V, Dam A. Nervous disturbances in horses in relation to infection with equine rhinopneumonitis virus. Acta Vet Scand 1971;12:134–6. Carroll CL, Westbury HA. Isolation of equine herpesvirus 1 from the brain of a horse affected with paresis. Aust Vet J 1985;62:345–6. Charlton KM, Mitchell D, Girard A, Corner AH. Meningoencephalomyelitis in horses associated with equine herpesvirus 1 infection. Vet Pathol 1976;13:59–68. Saaxegard F. Isolation and identification of equine rhinopneumonitis virus (equine abortion virus) from cases of abortion and paralysis. Nord Vet Med 1966;18: 504–12. Del Piero F, Wilkins PA. Pulmonary vasculotropic EHV-1 infection in equids. Vet Pathol 2001;38:474–5. Bitsch V. Cases of equine coital exanthema in Denmark. Acta Vetm Scand 1972;13:281–3. Blanchard TL, Kenney RM, Timoney PJ. Venereal disease. Vet Clin N Am-Equine 1992;8:191–203. Bryans JT. Herpesviral diseases affecting reproduction in the horse. Vet Clin N Am-Large 1980;2:303–12. Craig JF, Kehoe D. Horse pox and coital exanthema. J Comp Pathol Therap 1921;34:126–9. Bryans JT, Allen GP. In vitro and in vivo studies of equine ‘coital’ exanthema. In: Bryans J, Gerber H (eds) Proceedings of the Third International Conference on Equine Infectious Diseases, Paris, 1972. Karger, 1973; pp. 322–42. Gibbs EP, Roberts MC, Morris JM. Equine coital exanthema in the United Kingdom. Equine Vet J 1972;4:74–80. Pascoe RR. The effect of equine coital exanthema on the fertility of mares covered by stallions exhibiting the clinical disease. Aust Vet J 1981;57:111–14. Ames TR, O’Leary TP, Johnston GR. Isolation of equine herpesvirus type 2 from foals with respiratory disease. Comp Cont Educ Pract Vet 1986;8:664–70. Blakeslee J, Olsen RG, McAllister ES, Fassbender J, Dennis R. Evidence of respiratory tract infection induced by equine herpesvirus, type 2, in the horse. Can J Microbiol 1975;21:1940–6. Burrell MH, Whitwell KE, Wood JL, Mumford JA. Pyrexia associated with respiratory disease in young thoroughbred horses. Vet Rec 1994;134:219–20. Collinson PN, O’Rielly JL, Ficorilli N, Studdert MJ. Isolation of equine herpesvirus type 2 (equine gammaherpesvirus 2) from foals with keratoconjunctivitis. J Am Vet Med Assoc 1994;205:329–31.
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Equine Herpesviruses 28. Dwyer RM, Vickers ML. Ophthalmic disease outbreak. Equine Dis Quarterly 1997;5:4–5. 29. Studdert MJ. Equine herpesvirus 2 and disease [editorial]. Equine Vet J 1996;28:426–8. 30. Williams KJ, Maes R, Del Piero F, Lim A, Wise A, Bolin DC, Caswell J, Jackson C, Robinson NE, Derksen F, Scott MA, Uhal BD, Li X, Youssef SA, Bolin SR. Equine multinodular pulmonary fibrosis: A newly recognized herpesvirus-associated fibrotic lung disease. Vet Pathol 2007;44:849–62. 31. Wong DM, Belgrave RL, Williams KJ, Del Piero F, Alcott CJ, Bolin SR, Marr CM, Nolen-Walston R, Myers RK, Wilkins PA. Multinodular pulmonary fibrosis in five horses. J Am Vet Med Assoc 2008;232:898–905. 32. Davison A, Eberle R, Desrosiers RC, Fleckenstein R, McGeoch DJ, Pellet PE, Roizman B, Studdert MJ. Herpesviridae. In: Van Regenmortel MHV, Fauquet CM, Bishop DHL, Carstens EB, Estes MK, Lemon SM, Maniloff J, Mayo MA, McGeoch DJ, Pringle CR, Wickner RB (eds) Virus Taxonomy. Seventh Report of the International Committee on Taxonomy of Viruses. Orlando: Academic Press, 2000; pp. 114–27. 33. Ostlund EN. The equine herpesviruses. Vet Clin N AmEquine 1993;9:283–94. 34. Telford EA, Watson MS, McBride K, Davison AJ. The DNA sequence of equine herpesvirus-1. Virology 1992;189:304–16. 35. Telford EA, Watson MS, Perry J, Cullinane AA, Davison AJ. The DNA sequence of equine herpesvirus-4. J Gen Virol 1998;79:1197–203. 36. Awan AR, Chong Y-C, Field HJ. The pathogenesis of equine herpesvirus type 1 in the mouse: A new model for studying host responses to the infection. J Gen Virol 1990;71:1131–40. 37. Chowdhury SI, Ludwig H, Buhk HJ. Molecular biological characterization of equine herpesvirus type 1 (EHV-1) isolates from ruminant hosts. Virus Res 1988;11:127–39. 38. Crandell RA, Drysdale S, Stein TL. A comparative study of bovine herpesvirus 1247 and equine herpesvirus 1 in ponies. Can J Comp Med 1979;43:94–7. 39. Rebhun WC, Jenkins DH, Riis RC, Dill SG, Dubovi EJ, Torres A. An epizootic of blindness and encephalitis associated with a herpesvirus indistinguishable from equine herpesvirus I in a herd of alpacas and llamas. J Am Vet Med Assoc 1988;192:953–6. 40. Doll ER, Bryans JT, McCollum WH, Crowe EW. Propagation of equine abortion virus in Syrian hamsters. Cornell Vet 1956;46:68–82. 41. Allen GP, Yeargan MR, Bryans JT. Alterations in the equine herpesvirus 1 genome after in vitro and in vivo virus passage. Infect Immun 1983;40:436–9. 42. Studdert MJ. Restriction endonuclease DNA fingerprinting of respiratory, foetal and perinatal foal isolates of equine herpesvirus type 1. Arch Virol 1983;77:249–58. 43. Studdert MJ, Crabb BS, Ficorilli N. The molecular epidemiology of equine herpesvirus 1 (equine abortion virus) in Australasia 1975 to 1989. Aust Vet J 1992;69:104–11. 44. Nugent J, Birch-Machin I, Smith KC, Mumford JA, Swann Z, Newton JR, Bowden RJ, Allen GP, DavisPoynter N. Analysis of equid herpesvirus 1 strain variation reveals a point mutation of the DNA polymerase strongly associated with neuropathogenic versus nonneuropathogenic disease outbreaks. J Virol 2006;80:4047–60.
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45. Crabb BS, Allen GP, Studdert MJ. Characterization of the major glycoproteins of equine herpesviruses 4 and 1 and asinine herpesvirus 3 using monoclonal antibodies. J Gen Virol 1991;72:2075–82. 46. Yeargan MR, Allen GP, Bryans JT. Rapid subtyping of equine herpesvirus 1 with monoclonal antibodies. J Clin Microbiol 1985;21:694–7. 47. Crabb BS, Nagesha HS, Studdert MJ. Identification of equine herpesvirus 4 glycoprotein G: A type-specific, secreted glycoprotein. Virology 1992;190:143–54. 48. Mumford JA, Hannant D, Jessett DM, O’Neill T, Smith KC, Ostlund EN. Abortigenic and neurological disease caused by experimental infection with equid herpesvirus-1. In: Plowright W, Nakajima H (eds) Equine Infectious Diseases VII. Newmarket (Suffolk): R & W Publications, 1994; pp. 261–75. 49. Patel JR, Edington N, Mumford JA. Variation in cellular tropism between isolates of equine herpesvirus-1 in foals. Arch Virol 1982;74:41–51. 50. Allen GP, Yeargan MR, Turtinen LW, Bryans JT, McCollum WH. Molecular epizootiologic studies of equine herpesvirus-1 infections by restriction endonuclease fingerprinting of viral DNA. Am J Vet Res 1983;44:263– 71. 51. Doll ER, McCollum WH, Bryans JT, Crowe ME. Effect of physical and chemical environment on the viability of equine rhinopneumonitis virus propagated in hamsters. Cornell Vet 1959;49:75–81. 52. Gilkerson JR, Whalley JM, Drummer HE, Studdert MJ, Love DN. Epidemiology of EHV-1 and EHV-4 in the mare and foal populations on a Hunter Valley stud farm: Are mares the source of EHV-1 for unweaned foals. Vet Microbiol 1999;68:27–34. 53. Allen GP, Kydd IK, Slater ID, Smith KC. Advances in understanding of the epidemiology, pathogenesis and immunological control of equid herpesvirus-1 abortion. In: Wernery U, Wade J, Mumford J, Kaaden O-R (eds) Equine Infectious Diseases VIII. Newmarket (Suffolk): R & W Publications, 1999; pp. 129–46. 54. Studdert MJ, Blackney MH. Equine herpesviruses: on the differentiation of respiratory from foetal strains of type 1. Aust Vet J 1979;55:488–92. 55. Blunden AS, Smith KC, Binns MM, Zhang L, Gower SM, Mumford JA. Replication of equid herpesvirus 4 in endothelial cells and synovia of a field case of viral pneumonia and synovitis in a foal. J Comp Pathol 1995; 112:133–40. 56. Murray MJ, Del Piero F, Jeffrey SC, Davis MS, Furr MO, Dubovi EJ, Mayo JA. Neonatal equine herpesvirus type 1 infection on a thoroughbred breeding farm. J Vet Intern Med 1998;12:36–41. 57. Chowdhury SI, Kubin G, Ludwig H. Equine herpesvirus type 1 (EHV-1) induced abortions and paralysis in a Lipizzaner stud: A contribution to the classification of equine herpesviruses. Arch Virol 1986;90:273–88. 58. Greenwood RE, Simson AR. Clinical report of a paralytic syndrome affecting stallions, mares and foals on a thoroughbred studfarm. Equine Vet J 1980;12:113– 17. 59. Henninger RW, Reed SM, Saville WJ, Allen GP, Hass GF, Kohn CW, Sofaly C. Outbreak of neurologic disease caused by equine herpesvirus-1 at a university equestrian center. J Vet Intern Med 2007;21:157–65. 60. Wilson WD. Equine herpesvirus 1 myeloencephalopathy. Vet Clin N Am-Equine 1997;13:53–72.
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61. Goodman LB, Loregian A, Perkins GA, Nugent J, Buckles EL, Mercorelli B, Kydd JH, Palu G, Smith KC, Osterrieder N, Davis-Poynter N. A point mutation in a herpesvirus polymerase determines neuropathogenicity. PLoS Pathogens 2007;3:1583–92. 62. Allen GP, Timoney PJ. Recent advances in our understanding of equine herpesvirus-1 (EHV-1) myeloencephalopathy. In: Proceedings of the 111th Annual Meeting of the United States Animal Health Association, 2008, pp. 373–80. 63. Bryans JT, Allen GP. Control of abortigenic herpesviral infections. Curr Ther Theriogenol 1986;2:711–14. 64. Burrows M. The general virology of the herpesvirus group. In: Bryans JT, Gerber H (eds) Equine Infectious Diseases II. Basel: Karger, 1970; pp. 1–12. 65. Burrows R, Goodridge D. In vivo and in vitro studies of equine rhinopneumonitis strains. In: Bryans JT, Gerber H (eds) Equine Infectious Diseases III. Basel: Karger, 1972; pp. 306–21. 66. Gibson JS, Slater JD, Awan AR, Field HJ. Pathogenesis of equine herpesvirus-1 in specific pathogen-free foals: Primary and secondary infections and reactivation. Arch Virol 1992;123:351–66. 67. Burrows R, Goodridge D. Experimental studies on equine herpesvirus type 1 infections. J Reprod Fertil Suppl 1975;23:611–15. 68. Smith KC, Mumford JA, Lakhani K. A comparison of equid herpesvirus-1 (EHV-1) vascular lesions in the early versus late pregnant equine uterus. J Comp Pathol 1996;114:231–47. 69. Smith KC, Whitwell KE, Binns MM, Dolby CA, Hannant D, Mumford JA. Abortion of virologically negative foetuses following experimental challenge of pregnant pony mares with equid herpesvirus 1. Equine Vet J 1992;24:256–9. 70. Smith KC, Whitwell KE, Mumford JA, Gower SM, Hannant D, Tearle JP. An immunohistological study of the uterus of mares following experimental infection by equid herpesvirus 1. Equine Vet J 1993;25:36–40. 71. Edington N, Smyth B, Griffiths L. The role of endothelial cell infection in the endometrium, placenta and foetus of equid herpesvirus 1 (EHV-1) abortions. J Comp Pathol 1991;104:379–87. 72. Smith KC. Studies on the pathogenesis of abortigenic infection by equid herpesvirus-1 in pregnant pony mares. PhD thesis, University of London, 1994. 73. Carvalho R, Passos LM, Oliveira AM, Henry M, Martins AS. Detection of equine herpesvirus 1 DNA in a single embryo and in horse semen by polymerase chain reaction. Arq Bras Med Vet Zoo 2000;47:52–4. 74. Hebia I, Fieni F, Duchamp G, Destrumelle S, Pellerin JL, Zientara S, Vautherot JF, Bruyas JF. Potential risk of equine herpes virus 1 (EHV-1) transmission by equine embryo transfer. Theriogenology 2007;67:1485–91. 75. Tearle JP, Smith KC, Boyle MS, Binns MM, Livesay GJ, Mumford JA. Replication of equid herpesvirus-1 (EHV-1) in the testes and epididymides of ponies and venereal shedding of infectious virus. J Comp Pathol 1996;115:385–97. 76. Brown JA, Mapes S, Ball BA, Hodder AD, Liu IK, Pusterla N. Prevalence of equine herpesvirus-1 infection among Thoroughbreds residing on a farm on which the virus was endemic. J Am Vet Med Assoc 2007;231:577–80. 77. Kydd JH, Smith KC, Hannant D, Livesay GJ, Mumford JA. Distribution of equid herpesvirus-1 (EHV-1) in the
78.
79.
80.
81.
82.
83.
84.
85.
86.
87.
88.
89.
90.
91.
92.
93. 94.
respiratory tract of ponies: Implications for vaccination strategies. Equine Vet J 1994;26:466–9. Kydd JH, Smith KC, Hannant D, Livesay GJ, Mumford JA. Distribution of equid herpesvirus-1 (EHV-1) in respiratory tract associated lymphoid tissue: implications for cellular immunity, Equine Vet J 1994;26:470–3. Lunn DP, Holmes MA, Gibson JS, Field HJ, Kydd JH, Duffus WPH. Haematological changes and equine lymphocyte subpopulation kinetics during primary infection and attempted re-infection of specific pathogen free foals with EHV-1. Equine Vet J Suppl 1991;12:35–40. McColloch J, Williamson SA, Powis SJ, Edington N. The effect of EHV-1 infection upon circulating leucocyte populations in the natural equine host. Vet Microbiol 1993;37:147–61. Smith DJ, Iqbal J, Purewal A, Hamblin AS, Edington N. In vitro reactivation of latent equid herpesvirus-1 from CD5+/CD8+ leukocytes indirectly by IL-2 or chorionic gonadotrophin. J Gen Virol 1998;79:2997–3004. Matsumura T, Sugiura T, Imagawa H, Fukunaga Y, Kamada M. Epizootiological aspects of type 1 and type 4 equine herpesvirus infections among horse populations. J Vet Med Sci 1992;54:207–11. Browning GF, Bulach DM, Ficorilli N, Roy EA, Thorp BH, Studdert MJ. Latency of equine herpesvirus 4 (equine rhinopneumonitis virus). Vet Rec 1988;123:518–19. Burrows R, Goodridge D. Studies of persistent and latent equid herpesvirus-1 and herpesvirus-3 infections in the Pirbright pony herd. In: Wittmann G, Gaskell RM, Rziha H-J (eds) Latent Herpesvirus Infections in Veterinary Medicine. The Hague: Martinus Nijhoff, 1984; pp. 307–19. Edington N, Welch HM, Griffiths L. The prevalence of latent equid herpesviruses in the tissues of 40 abattoir horses. Equine Vet J 1994;26:140–2. Slater JD, Borchers K, Thackray AM, Field HJ. The trigeminal ganglion is a location for equine herpesvirus 1 latency and reactivation in the horse. J Gen Virol 1994;75:2007–16. Bryans JT, Swerczek TW, Darlington RW, Crowe MW. Neonatal foal disease associated with perinatal infection by equine herpesvirus-1. J Equine Med Surg 1977;1:20–6. Crowhurst FA, Dickinson G, Burrows R. An outbreak of paresis in mares and geldings associated with equid herpesvirus 1. Vet Rec 1981;109:527–8. Jackson TA, Kendrick LW. Paralysis of horses associated with equine herpesvirus 1 infection. J Am Vet Med Assoc 1971;158:1351–7. Edington N, Bridges CG, Patel JR. Endothelial cell infection and thrombosis in paralysis caused by equid herpesvirus-1: Equine stroke. Arch Virol 1986;90:111–24. Meyer H, Thein P, Hubert P. Characterization of two equine herpesvirus (EHV) isolates associated with neurological disorders in horses. Zbl Vet Med B 1987;34:545–8. Van Maanen C, Vreeswijk J, Moonen P, Brinkhof J, De Boer-Luijtze E, Terpstra C. Differentiation and genomic and antigenic variation among fetal, respiratory, and neurological isolates from EHV1 and EHV4 infections in The Netherlands. Vet Quart 2000;22:88–93. Allen GP. Demystifying neurologic herpes. Equine Dis Quart 2007;16:3–4. Allen GP, Breathnach CC. Quantification by real-time PCR of the magnitude and duration of leucocyteassociated viraemia in horses infected with neuropathogenic vs. non-neuropathogenic strains of equine herpesvirus-1. Equine Vet J 2006;38:252–7.
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Equine Herpesviruses 95. Mumford JA, Rossdale PD. Virus and its relationship to the “poor performance” syndrome. Equine Vet J 1980;12:3–9. 96. Mumford JA, Rossdale PD, Jessett DM, Gann SJ, Ousey J, Cook RF. Serological and virological investigations of an equid herpesvirus 1 (EHV-1) abortion storm on a stud farm in 1985. J Reprod Fertil Suppl 1987;35:509–18. 97. Gleeson LJ, Coggins L. Response of pregnant mares to equine herpesvirus 1 (EHV1). Cornell Vet 1980;70:391–400. 98. Prickett ME. The pathology of disease caused by equine herpesvirus 1. In: Bryans JT, Gerber H (eds) Equine Infectious Diseases II. Basel: S. Karger, 1970; pp. 24–33. 99. Bryans JT. On immunity to disease caused by equine herpesvirus 1. J Am Vet Med Assoc 1969;155:294–300. 100. Allen GP. Antemortem detection of latent infection with neuropathogenic strains of equine herpesvirus-1 in horses. Am J Vet Res 2006;67:1401–05. 101. McCartan CG, Russell MM, Wood JL, Mumford JA. Clinical, serological and virological characteristics of an outbreak of paresis and neonatal foal disease due to equine herpesvirus-1 on a stud farm. Vet Rec 1995;136:7–12. 102. Slater JD, Gibson JS, Barnett KC, Field HJ. Chorioretinopathy associated with neuropathology following infection with equine herpesvirus-1. Vet Rec 1992;131: 237–9. 103. Gilkerson J, Jorm LR, Love DN, Lawrence GL, Whalley JM. Epidemiological investigation of equid herpesvirus4 (EHV-4) excretion assessed by nasal swabs taken from thoroughbred foals. Vet Microbiol 1994;39:275–83. 104. United States Department of Agriculture. Equine herpesvirus myeloencephalopathy: A potentially emerging disease. Animal Plant Health Inspection Service, Veterinary Services, Centers for Epidemiology and Animal Health, 2007; pp. 40–43. 105. Edington N, Bridges CG, Huckle A. Experimental reactivation of equid herpesvirus 1 (EHV 1) following the administration of corticosteroids. Equine Vet J 1985;17:369–72. 106. Edington N, Chesters P, Azam H, Welch H, McGladdery A, Purewal A. Profiles of alphaherpesviruses in circulating leucocytes from Thoroughbred mares and foals using PCR and co-cultivation. In: Nakajima H, Plowright W (eds) Equine Infectious Diseases VII. Newmarket (Suffolk): R & W Publications, 1994; pp. 251–4. 107. Welch HM, Bridges CG, Lyon AM, Griffiths L, Edington N. Latent equid herpesviruses 1 and 4: detection and distinction using the polymerase chain reaction and co-cultivation from lymphoid tissues. J Gen Virol 1992;73:261–8. 108. Chesters PM, Allsop R, Purewal A, Edington N. Detection of latency-associated transcripts of equid herpesvirus 1 in equine leukocytes but not in trigeminal ganglia. J Virol 1997;71:3437–43. 109. Allen GP, Bolin DC, Bryant U, Carter CN, Giles RC, Harrison LR, Hong CB, Jackson CB, Poonacha K, Wharton R, Williams NM. Prevalence of latent, neuropathogenic equine herpesvirus-1 in the Thoroughbred broodmare population of central Kentucky. Equine Vet J 2008;40:105–10. 110. Doll ER, Crowe ME, Bryans JT, McCollum WH. Infection immunity in equine virus abortion. Cornell Vet 1955;45:387–410. 111. Edington N, Bridges CG. One way protection between equid herpesvirus 1 and 4 in vivo. Res Vet Sci 1990;48:235–9.
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112. Fitzpatrick DR, Studdert MJ. Immunologic relationships between equine herpesvirus type 1 (equine abortion virus) and type 4 (equine rhinopneumonitis virus). Am J Vet Res 1984;45:1947–52. 113. Stokes A, Corteyn AH, Murray PK. Clinical signs and humoral immune response in horses following equine herpesvirus type-1 infection and their susceptibility to equine herpesvirus type-4 challenge. Res Vet Sci 1991;51:141–8. 114. Allen GP, Yeargan MR, Costa LR, Cross R. Major histocompatibility complex class I-restricted cytotoxic Tlymphocyte responses in horses infected with equine herpesvirus 1. J Virol 1995;69:606–12. 115. Bryans JT. Application of management procedures and prophylactic immunization to the control of equine rhinopneumonitis. In: Proceedings of the 26th Annual Convention of the American Association of Equine Practitioners, Anaheim, 1981, pp. 259–72. 116. Powell, D. Prevention, treatment of equine herpesvirus. Thoroughbred Times 31January, 1992, p. 9. 117. Crabb BS, Studdert MJ. Equine herpesviruses 4 (equine rhinopneumonitis virus) and 1 (equine abortion virus). Adv Virus Res 1995;45:153–90. 118. Crandell RA, Angulo AB. Equine herpesvirus-1 antibodies in stillborn foals and weak neonates. Vet Med 1985;80:73–74, 76. 119. Timoney PJ, McCollum WH. Equine viral arteritis. In: Coetzer JAW, Tustin RC (eds) Infectious Diseases of Livestock. Vol. 2, 2nd edn. Oxford University Press Southern Africa, 2004; pp. 924–32. 120. Allen GP. Development of a real-time PCR assay for rapid diagnosis of neuropathogenic strains of equine herpesvirus-1. J Vet Diagn Invest 2007;19:69–72. 121. Jackson TA, Osburn BI, Cordy DR, Kendrick JW. Equine herpesvirus 1 infection of horses: studies on the experimentally induced neurologic disease. Am J Vet Res 1977;38:709–19. 122. Platt H, Singh H, Whitwell KE. Pathological observations on an outbreak of paralysis in broodmares. Equine Vet J 1980;12; 118–26. 123. Whitwell KE, Gower SM, Smith KC. An immunoperoxidase method applied to the diagnosis of equine herpesvirus abortion, using conventional and rapid microwave techniques. Equine Vet J 1992;24:10–12. 124. De Lahunta A. Diagnosis of equine neurologic problems. Cornell Vet 1978;68(Suppl. 7): 122–32. 125. Borchers K, Slater J. A nested PCR for the detection and differentiation of EHV-1 and EHV-4. J Virol Methods 1993;45:331–6. 126. Borchers K, Wolfinger U, Lawrenz B, Schellenbach A, Ludwig H. Equine herpesvirus 4 DNA in trigeminal ganglia of naturally infected horses detected by direct in situ PCR. J Gen Virol 1997;78:1109–14. 127. Bryans JT, Allen GP. Application of a chemically inactivated, adjuvanted vaccine to control abortigenic infection of mares by equine herpesvirus 1. Dev Biol Stand 1982;52:493–8. 128. Bryans JT. Serologic responses of pregnant thoroughbred mares to vaccination with an inactivated equine herpesvirus 1 vaccine. Am J Vet Res 1980;41:1743–6. 129. Burrows R, Goodridge D, Denyer MS. Trials of an inactivated equid herpesvirus 1 vaccine: Challenge with a subtype 1 virus. Vet Rec 1984;114:369–74. 130. Crandell RA, Mock RE, Lock TF. Vaccination of pregnant ponies against equine rhinopneumonitis. Am J Vet Res 1980;41:994–6.
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131. Ellis JA, Bogdan JR, Kanara EW, Morley PS, Haines DM. Cellular and antibody responses to equine herpesviruses 1 and 4 following vaccination of horses with modified-live and inactivated viruses. J Am Vet Med Assoc 1995;206:823–32. 132. Purdy CW, Ford SJ, Porter RC. Equine rhinopneumonitis vaccine: Immunogenicity and safety in adult horses, including pregnant mares. Am J Vet Res 1978;39:377– 83. 133. Breathnach CC, Allen GP, Holland RE, Conboy HS, Berry DB, Chambers TM. Problems associated with vaccination of foals against equine herpesvirus-4 (EHV4) and the role of anti-EHV-4 maternal antibodies. In:
Wernery U, Wade JF, Mumford JA, Kaaden O-R (eds) Equine Infectious Diseases VIII. Newmarket (Suffolk): R&W Publications, 1999; pp. 426–7. 134. Ellis JA, Steeves E, Wright AK, Bogdan JR, Davis WC, Kanara EW, Haines DM. Cell-mediated cytolysis of equine herpesvirus-infected cells by leukocytes from young vaccinated horses. Vet Immunol Immunopathol 1997;57:201–14. 135. American Association of Equine Practitioners. Guidelines for Vaccination of Horses. 2008; http://www. aaep.org/vaccination guidelines.htm. 136. Minke JM, Audonnet JC, Fischer L. Equine viral vaccines: The past, present and future. Vet Res 2004;35:425–43.
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Chapter 246
Equine Viral Arteritis Peter J. Timoney
Introduction Equine viral arteritis (EVA) is an acute contagious disease of equids, caused by equine arteritis virus (EAV).1, 2 It is principally characterized by fever, depression, leukopenia, dependent edema, and respiratory signs. Although not typically life-threatening to otherwise healthy adult horses, the disease is of primary concern because of the risk of abortion in the pregnant mare,3 illness and death in young foals,4–6 and the predilection of the causal virus to establish the carrier state in the stallion.7–9 Although descriptions of a clinically similar disease were recorded in the veterinary literature for many years,10–12 it was not until 1953, following an extensive outbreak of respiratory disease and abortion on a Standardbred farm near Bucyrus, Ohio, that EVA was unequivocally defined as a separate equine viral disease,1 its pathological features characterized,13 and the etiological agent isolated.1 Equine viral arteritis garnered little international attention until the occurrence of the disease on numerous Thoroughbred breeding farms in Kentucky in 1984.14 The event caused major concern over the risk of spread of EAV through movement of horses and the potential for major outbreaks of abortion in unprotected populations of pregnant mares, as well as establishment of the carrier state in stallions. This led to a significant reassessment of the importance of EVA by horse breeding and racing countries worldwide and the imposition of stringent restrictions on the movement of horses that were positive for serum antibodies to the virus.2 While many countries have since eased their respective import requirements, the disease continues to have a major financial impact on the international trade in horses and equine semen.15
worldwide.2, 16 Presence of the virus has been confirmed in countries in North and South America,16–18 Europe,19–24 Asia,25, 26 Africa,24, 27 and Australia and New Zealand.28 The prevalence of infection has been shown to vary both between countries and between different horse breeds in the same country.2, 16 Historically, the highest rates of infection have been reported in Standardbreds,16, 29 although there is more recent evidence to suggest that the seropositivity rate in various Warmblood breeds is on the increase.30 Notwithstanding the widespread global distribution of EAV,2 confirmed outbreaks of EVA have historically been relatively infrequent and only recorded in Europe and North America. There has been an increase in the frequency of occurrence of the disease in recent years, especially in countries with a significant import trade in horses and equine semen.31 While this trend may be due in part to greater awareness of EVA, and to improved diagnostic capability, it is much more likely a direct consequence of the increased volume of horse movements, especially stallions, and the rapidly growing trade in cryopreserved semen.2, 31 There have been numerous instances where imported carrier stallions or infective semen have been responsible for significant outbreaks of EVA, including abortion in mares and mortality in young foals.32–34 Equine arteritis virus has also been widely disseminated within countries through the shipment of fresh-cooled or cryopreserved semen, as was well exemplified by the 2006 multistate occurrence of EVA in the USA,35 and in 2007 by the spread of the disease in France associated with shipment of semen from a draft stallion.36
Causal agent Distribution Virological and/or serological evidence of EAV has been found in equine populations in many countries
Equine arteritis virus is an enveloped, singlestranded, positive-sense RNA virus with quasispecies structure; it is the prototype member of the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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genus Arterivirus, family Arteriviridae, order Nidovirales.37, 38 Other taxonomically related viruses belonging to the same genus include porcine reproductive and respiratory syndrome virus, lactic dehydrogenase elevating virus of mice, and simian hemorrhagic fever virus.37, 39 Evidence to date would indicate that EAV is a natural pathogen of equids, especially horses.2 While there is one report of the detection of EAV nucleic acid in the tissues of an aborted alpaca fetus,40 the significance of this finding has yet to be established. All of the strains of EAV examined so far are antigenically related to one another and to the original prototype (Bucyrus) strain of the virus.2, 41 Although only one major serotype of EAV has been identified at this time, differences in antigenicity and pathogenicity have been observed among geographically and temporally distinct isolates of the virus.42–44 Some strains are capable of causing the range of clinical signs collectively referred to as EVA, whereas others elicit only a mild to moderate febrile response.2, 45 Equine arteritis virus is not a particularly resistant virus outside the body. It is readily inactivated under a range of physico-chemical conditions. However, the virus is remarkably stable at freezing temperatures, remaining infective in cryopreserved semen for many years.2
Transmission The most important modes of transmission of EAV are via the respiratory route by the acutely infected equine,1, 2, 46 and via the venereal route by the acutely or chronically infected stallion.2, 14, 47 Although shed in various secretions and excretions during the acute phase of the infection,46, 48, 49 the virus is present in greatest concentration in the respiratory tract.46 Spread of EAV occurs through direct contact with infective respiratory secretions, which can be positive for virus for 2 to 16 days. This is the principal means of viral dissemination on breeding farms, racetracks, and other performance venues, and wherever horses are congregated.2, 17, 50, 51 Equine arteritis virus can also be spread very effectively by the acutely infected stallion, which can shed high concentrations of virus in its semen.14 Venereal transmission is the only means of spread of EAV by the persistently infected or carrier stallion.2, 8 Equine arteritis virus can also be transmitted across the placenta from an infected mare to her unborn foal. Depending upon the stage in pregnancy at which viral exposure occurs, infection may result in abortion or the birth of a congenitally infected foal.5, 6, 52 Infection can also be transmitted by embryo transfer where the donor mare is bred with virus-infective semen (C. Broaddus, personal communication).
An additional means of spread is by the indirect transfer of EAV through a variety of fomites contaminated with infective secretions, semen, or the tissues and fluids of an aborted fetus or young foal.2, 14
Clinical response The clinical outcome following exposure to EAV is influenced by several factors – related to the virus, host, and environment.2 The majority of primary cases of infection are asymptomatic.2, 45 The clinical signs of EVA, which can vary in range and severity, develop after an incubation period of 2 to 13 days, usually 6 to 8 days after venereal exposure.2 Typical cases can present with all or any combination of the following: fever, anorexia, depression, leukopenia, dependent edema, especially of the lower limbs, scrotum, and prepuce in the male and mammary glands in the female, supraorbital or periorbital edema, nasal and/or ocular discharge, conjunctivitis, photophobia, and a skin rash, which may be localized to the sides of the face, neck, or pectoral region or may be generalized over the surface of the body. A range of other clinical signs including coughing, respiratory distress, submaxillary lymphadenopathy, posterior paresis or ataxia, adventitious swelling in the intermandibular space, beneath the sternum, the shoulder region or other parts of the body, gingival and buccal erosions, and diarrhea have been infrequently observed in some cases of the disease.1, 2, 14, 53 The duration of clinical illness can range from several days up to 2 weeks. In general, the severity of EVA tends to be greater in very young or aged horses, in debilitated individuals and in horses subjected to physical stress.2, 17 With few exceptions, horses affected with EVA, even those exhibiting severe clinical signs, make uneventful recoveries. Horses in training may experience a period of impaired performance during the acute and early convalescent stages of the infection.17, 51 While mortality is a very rare sequel to EAV infection in otherwise healthy adult horses,54 it has been recorded in foals congenitally infected with the virus as well as in those a few weeks to months of age.4–6,33 Of special importance is the potential of many strains of EAV to cause abortion.1, 2, 32, 33, 52 Abortion rates can range from less than 10% to greater than 70% in unprotected mares.2, 49, 55 Abortion can be a sequel to clinical or asymptomatic infection and supervenes late in the acute phase or early in the convalescent phase of the infection. Natural or experimental cases of EAV-related abortion can occur in mares from 3 to over 10 months of gestation.2 Viral exposure of the unprotected mare very late in pregnancy may not result in abortion but rather the birth of a congenitally infected foal.6, 33 Whether dealing with a case of EVA-related abortion or a congenitally
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Equine Viral Arteritis infected foal, the placenta and placental fluids, fetus, or affected newborn foal are very productive sources of EAV for any unprotected in-contact animals. A source of confusion among some breeders, owners, and even veterinarians is whether an unprotected mare bred with EAV-infective semen can harbor the virus following conception and abort due to the infection weeks or months later. There is no evidence that this occurs. Based on investigation of naturally occurring outbreaks of EVA and experimentally induced infection, abortion is a sequel to the pregnant mare being exposed invariably by the respiratory route and not the venereal route. The usual scenario is that the unprotected pregnant mare comes into direct contact with another horse(s) acutely infected with EAV, very frequently the result of recent insemination with virus-infective semen. There is considerable evidence that mares that abort due to this infection do not experience any virus-related impairment of fertility in subsequent breeding seasons.56 Stallions affected with EVA that develop a significant febrile response and severe edema of the scrotum and prepuce can experience a period of temporary subfertility unless treated promptly to mitigate the severity of these clinical signs.57 Detectable changes include a marked decrease in the percentage of normal sperm; this commences approximately 1 week after infection, reaches a nadir about week 7, before returning to pre-infection levels by week 16. There are concomitant changes in the percentage of motile sperm and sperm concentration, both of which follow a similar response pattern. The temporary subfertility observed in acutely affected stallions is not considered to be directly virus-related. Such stallions do not appear to experience any resultant long-term adverse effects in their fertility.
Pathogenesis Based on studies of experimentally induced infection,4, 46, 58 following exposure EAV rapidly invades the respiratory epithelium, multiplying in both bronchial and alveolar macrophages before migrating to the regional lymph nodes. Within 3 days, a leukocyte-associated viremia develops and the virus becomes distributed in the tissues and fluids throughout the body. By days 6 to 8, EAV has localized in the vascular endothelium and medial myocytes of blood vessels with development of the vasculitis characteristic of the disease.13, 59, 60 These lesions give rise to edema and hemorrhage in various tissues and organs throughout the body. There is evidence that replication of EAV in macrophages and vascular endothelium leads to activation and increased transcription of genes that encode proinflammatory mediators, and these likely play a
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significant role in the pathogenesis of EVA.61 Maximal vascular injury is evident about 10 days into the infection, after which the lesions start to resolve.58 The pathogenesis of abortion resulting from EAV infection remains to be determined. While myometritis has been reported in mares experimentally infected with the highly virulent Bucyrus strain of EAV,62, 63 it is debatable whether this is a feature of naturally occurring cases of EVA-related abortion. Pathological evidence would indicate that fetal death occurs prior to parturition. Equine viral arteritisrelated cases of abortion are frequently associated with minimal evidence of vascular or other tissue damage involving the placenta and/or the fetus.55, 62
Epidemiology A range of virus-, host-, and environment-related factors are considered to be involved in the epidemiology of EVA.2 Among those identified of importance are:
r r r r r
variation in pathogenicity and other phenotypic characteristics among naturally occurring strains of EAV; modes of transmission during acute and chronic phases of the infection; occurrence of the carrier state in the stallion; nature and duration of acquired immunity to infection; changing trends in the horse industry.
Viral characteristics It has been known for many years that field strains of EAV can differ significantly from one another based on their inherent ability to cause disease, including abortion. Limited experimental studies have confirmed the existence of lento-, meso-, and velogenic strains of the virus.2, 43 The carrier stallion is believed to be the primary source of genetic and phenotypic diversity among naturally occurring strains of EAV.64 Modes of transmission The principal routes of transmission of EAV by the acutely infected equid or by the persistently infected (carrier) stallion have already been described.2, 8, 14 Knowledge not only of the different routes of transmission but also of the duration of viral shedding is of critical importance when attempting to develop effective prevention and control programs for this disease. Carrier state Long-term persistence of EAV, or the carrier state, has only been identified in the stallion and not in the mare, gelding, or sexually immature colt.2, 65, 66
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The carrier stallion is widely considered the natural reservoir of EAV and of major epidemiological importance in dissemination and perpetuation of the virus.2, 45 Frequency of the carrier state can vary from less than 10% to greater than 70%.7, 8 Duration of viral persistence in the stallion may be several weeks, months, or many years. Not all long-term carrier stallions remain persistently infected for life. A percentage will spontaneously clear the carrier state after a variable interval, with no evidence of subsequent reversion to a virus-shedding state.7 Establishment and maintenance of the carrier state has been shown to be androgen-dependent,67 with EAV harbored in certain of the accessory sex glands, especially the ampulla of the vas deferens. Carrier stallions are constant semen shedders of the virus, which is present in the spermrich fraction of the ejaculate.8 There is no evidence that carrier stallions can transmit EAV other than by the venereal route. Efficiency of virus transmission, whether by natural service or artificial insemination, can be as high as 85–100%. Carrier stallions are invariably seropositive for antibodies to EAV, are clinically normal, and appear to experience no adverse effect on their fertility.7
Immunity Compared to most other equine infectious diseases, natural infection with EAV stimulates a strongly protective, long-lasting immunity against the disease.68, 69 Foals born of seropositive mares are passively protected against EVA for the first 3 to 5 months of life.70 Vaccination can also stimulate a level of immunity protective against development of clinical disease and establishment of the carrier state in the stallion.56 First-time vaccinates may still undergo a limited reinfection cycle upon natural exposure to the virus.71
Industry trends Of various developments that have occurred in the horse industry worldwide over the past 30 to 40 years, two, namely continued growth in the volume of international movement of horses and increased trade in cryopreserved semen, have impacted significantly on the global distribution of EAV.31
Economic significance Sources of economic losses attributable to EAV infection include: (i) outbreaks of abortion and death in young foals; (ii) decreased commercial value of carrier stallions and the demand to breed to them; (iii) loss of export markets for carrier stallions, virusinfective semen, and, in the case of some countries,
any horse that is positive for antibodies to the virus or comes from premises on which there is evidence of EAV infection; and (iv) cancellation of race entries and disruption of training schedules during outbreaks at racetracks.2, 72
Diagnosis No matter how suggestive the clinical features of a suspect case or outbreak of EVA may be, they are not of themselves diagnostic of the disease.2 There are a wide range of other infectious as well as noninfectious diseases that closely resemble the symptomatology of EVA. In view of the mimicry of clinical signs, laboratory confirmation of a provisional diagnosis of EVA should be obtained without delay. Where acute EAV infection is suspected, laboratory confirmation can be accomplished by virus isolation, detection of viral nucleic acid by reverse transcription-polymerase chain reaction (RT-PCR) assay or visualization of viral antigen, and/or demonstration of a specific antibody response by testing paired (acute and convalescent) sera collected 3 to 4 weeks apart.73 Appropriate specimens to collect from suspected cases of infection include nasopharyngeal swabs or washings, and citrated or EDTA (ethylene diamine tetra-acetic acid) as well as clotted blood samples.2, 73 Collection of specimens for EAV detection should be instigated as soon as possible after the onset of fever or other clinical signs to optimize the chances of confirmation of a diagnosis. In suspect cases of EVA abortion, virus isolation/detection should be attempted from a range of placental and fetal tissues and fluids, especially fetal lung, liver, and lymphoreticular tissues.2, 73 Specimens should also be taken for histological and immunohistochemical examination for the vascular lesions characteristic of EAV infection. Where EVA is suspected in cases of mortality in young foals or older horses, a wide range of tissues and fluids should be collected and subjected to the same laboratory screening as specimens from cases of abortion. In detection of the carrier state, the serological status of the stallion being evaluated must firstly be established.2, 73, 74 Only stallions with a neutralizing antibody titer of ≥ 1:4 and without a certified history of vaccination against EVA should be considered potential carriers of EAV. The carrier state is confirmed by demonstration of the virus in semen containing the sperm-rich fraction of the ejaculate by virus isolation or detection by RT-PCR assay. A more costly, less timely, but equally reliable means of detection of the carrier state is to test breed a stallion to two seronegative mares and check these for seroconversion to the virus after 28 days.74
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Equine Viral Arteritis
Treatment Treatment of horses with EVA is currently limited to controlling the severity of clinical signs of disease.2 Particular attention should be paid to mitigating the fever and dependent edema in affected stallions. Currently, there is no effective treatment for the viral-related pneumonia or pneumoenteritis in young foals, which invariably has a fatal outcome. Non-surgical interventions for facilitating elimination of the carrier state in the stallion have not as yet proven uniformly effective. Attempts at hyperimmunization of a limited number of carrier stallions with R ; see the modified live virus (MLV) vaccine (Arvac below) with a view to promoting viral clearance from the reproductive tract have been unsuccessful. Temporary downregulation of testosterone production through the use of gonadotropin-releasing hormone (GnRH) antagonists75 or GnRH immunization76 (immunocastration) has been associated with elimination of EAV in some but not all treated stallions. Additional studies are needed to confirm that the induced downregulation of GnRH production does not have any long-term adverse effect on the fertility of treated individuals. Recent work has shown that it is possible to reduce but not eliminate the viral content in EAV-infective semen through the use of density gradient centrifugation and a ‘swim-up’ protocol for processing the semen.77 However, a foolproof method of decontaminating semen without compromising its fertility remains to be developed.
Prevention and control Sufficient is known about EAV and the epidemiology of the infection on which to base effective measures for the prevention and control of EVA.2 Current prophylactic programs are targeted at breeding populations, which experience the greatest health-related consequences and economic impact from EVA. These are focused on minimizing the risk of introduction and dissemination of EAV on farms to prevent abortions, death in young foals, and establishment of the carrier state in stallions. Although the virus can cause extensive outbreaks of disease at racetracks, shows and other equine performance events where horses are closely congregated, currently the risk is not perceived great enough to warrant implementing an EVA control program to prevent such contingencies.51 Two vaccines are commercially available against R EVA: a modified live virus (MLV) vaccine78 (Arvac ; Fort Dodge Animal Health), which is approved for use in Canada and the USA, and an inactivated vacR ; Fort Dodge Animal Health), which cine (Artervac is conditionally licensed in a limited number of European countries.2, 73 Use of the MLV vaccine since 1985
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has confirmed its safety and immunogenicity for stallions,79, 80 non-pregnant mares, fillies, and colts. It is not recommended for use in pregnant mares, especially in the final 2 months of gestation, and in foals less than 6 weeks of age unless in situations of high risk of natural exposure to EAV.2 The inactivated vaccine, though safe for pregnant mares, is not as immunogenic as the MLV vaccine.73 There is no evidence that the MLV vaccine can establish the carrier state in the stallion.2, 7 Primary vaccination provides protection against clinical disease for several years.47, 69 Following natural infection with EAV, firsttime vaccinated horses may undergo a limited reinfection cycle with the possibility of short-term respiratory shedding of very small amounts of virus.71 Revaccination with the MLV vaccine induces a substantial anamnestic antibody response, which evidence would indicate can persist for at least several years.2 Experience has shown the importance of combining sound management practices with a program of targeted vaccination in mitigating the risk of outbreaks of EVA.2 Critical to the success of such programs is minimizing or eliminating direct or indirect contact of EAV-naive (i.e. unprotected) horses with the secretions/excretions of infected individuals. To reduce the possibility of this occurring, all horses arriving from other premises (sales, racetracks, shows, veterinary clinics, farms) should be isolated from resident horses for 4 weeks. Teaser stallions and nurse mares should not be overlooked in this regard. Furthermore, all pregnant mares should be segregated and maintained in small groups until they have foaled. Of major importance in achieving control over the dissemination of EAV in breeding populations is containment of the number of carrier stallions.2 Reduction of the natural reservoir of the virus can be accomplished through vaccination of seronegative breeding stallions, teaser stallions, and colts above 6 months and less than 12 months of age – all highly recommended measures.56 An essential part of any control program against EVA is the need to screen existing stallion populations for presence of the carrier state. Individual carriers should be physically isolated and appropriate sanitary measures taken when breeding or collecting them to prevent inadvertent spread of infection to other horses on the premises. Such stallions should preferably be bred to naturally seropositive mares or mares previously vaccinated against EVA. In the case of farms using artificial insemination, all shipped semen should be screened for the presence of EAV, especially if the semen originated from abroad.34, 56 Mares inseminated with infective semen should be managed similarly to those bred by natural service to a carrier stallion.2 The importance
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of stallion owners informing mare owners of the carrier status of stallions under their care cannot be overemphasized. Similarly, those importing and retailing stallion semen that is known to be infective should communicate that fact to mare owners intending to use such semen so that they can take the necessary precautions to avoid unintentional spread of the virus and the risk of an outbreak of EVA. In the event of a suspected outbreak of EVA, regardless of circumstances under which it occurs, the appropriate animal health regulatory authorities should be notified without delay. Affected and in-contact horses should be isolated, and restrictions placed on the movement of horses onto and off the affected premises. It is important to obtain laboratory confirmation of a provisional clinical diagnosis of EVA as soon as possible. If a breeding farm is involved, all breeding activity should be suspended until further notice. Stalls and equipment associated with affected horses should be thoroughly cleaned and disinfected. In such situations, early vaccination of the at-risk equine population on a premises has proven effective in helping to curtail further spread of EAV and limit the morbidity rate during outbreaks of EVA in the USA.50 Restrictions on movement should be maintained for at least 3 weeks after the last clinical case or confirmed asymptomatic case of EAV infection. From an international viewpoint, countries, and specifically their respective equine industries, differ considerably in their stance over the significance of EVA.81 Whereas some adopt a ‘zero tolerance’ approach to the disease and seek to maintain freedom from it, other countries/industries are prepared to ‘live with’ the infection, based on its perceived veterinary medical and economic importance. Regardless of which approach is pursued, experience gained over the past 25 years has repeatedly confirmed that EVA is a very manageable and controllable disease.2, 56
References 1. Doll ER, Bryans JT, McCollum WH, Crowe MEW. Isolation of a filterable agent causing arteritis of horses and abortion by mares. Its differentiation from the equine abortion (influenza) virus. Cornell Vet 1957;47:3–40. 2. Timoney PJ, McCollum WH. Equine viral arteritis. Vet Clin N Am-Equine 1993;9:295–309. 3. Doll ER, Knappenberger RE, Bryans JT. An outbreak of abortion caused by the equine arteritis virus. Cornell Vet 1957;47:69–75. 4. Del Piero F, Wilkins PA, Lopez JW, Glaser AL, Dubovi EJ, Schlafer DH, Lein DH. Equine viral arteritis in newborn foals: clinical, pathological, serological, microbiological and immunohistochemical observations. Equine Vet J 1997;29:178–85.
5. Golnik W, Michalska Z, Michalak T. Natural equine viral arteritis in foals. Schweiz Arch Tierh 1981;123:523–33. 6. Vaala WE, Hamir AN, Dubovi EJ, Timoney PJ, Ruiz B. Fatal, congenitally acquired infection with equine arteritis virus in a neonatal Thoroughbred. Equine Vet J 1992;24: 155–8. 7. Timoney PJ, McCollum WH. Equine viral arteritis: further characterization of the carrier state in stallions. J Reprod Fertil Suppl 2000;56:3–11. 8. Timoney PJ, McCollum WH, Murphy TW, Roberts AW, Willard JG, Carswell GD. The carrier state in equine arteritis virus infection in the stallion with specific emphasis on the venereal mode of virus transmission. J Reprod Fertil 1987;35:95–102. 9. Timoney PJ, McCollum WH, Roberts AW, Murphy TW. Demonstration of the carrier state in naturally acquired equine arteritis virus infection in the stallion. Res Vet Sci 1986;41:279–80. 10. Clark J. Transmission of pink-eye from apparently healthy stallions to mares. J Comp Pathol Theriogenol 1892;5: 261–4. 11. Pottie A. The propagation of influenza from stallions to mares. J Comp Pathol Therap 1888;1:37–8. 12. Reeks HC. The transmission of pink-eye from apparently healthy stallions to mares. J Comp Pathol Therap 1902;18:97–102. 13. Jones TC, Doll ER, Bryans JT. The lesions of equine viral arteritis. Cornell Vet 1957;47:52–68. 14. Timoney PJ. Clinical, virological and epidemiological features of the 1984 outbreak of equine viral arteritis in the Thoroughbred population in Kentucky, USA. In: Proceedings of the Grayson Foundation International Conference of Thoroughbred Breeders’ Organizations on Equine Viral Arteritis, 1984. Grayson Jockey Club Equine Research Foundation, 1984, pp. 24–33. 15. Timoney PJ, McCollum WH. Equine viral arteritis in perspective in relation to international trade. J Equine Vet Sci 1993;13:50–2. 16. McCollum WH, Bryans JT. Serological identification of infection by equine arteritis virus in horses of several countries. In: Proceedings of the 3rd International Conference on Equine Infectious Diseases. Basel: S. Karger, 1973; pp. 256–63. 17. McCollum WH, Swerczek TW. Studies of an epizootic of equine viral arteritis in racehorses. J Equine Med Surg 1978;2:293–9. 18. Nosetto PEO, Etcheverrigaray ME, Oliva GA, Gonzalez ET, Samus SA. Arteritis viral equina: Deteccion de anticuerpos en equinos de la republica Argentina. Zbl Vet Med B 1984;31:526–529. 19. Camm IS, Thursby-Pelham C. Equine viral arteritis in Britain. Vet Rec 1993;132:615. 20. de Boer GF, Osterhaus ADME, van Oirschot JT, Wemmenhove R. Prevalence of antibodies to equine viruses in the Netherlands. Vet Quart 1979;1:65–74. 21. Eichhorn W, Heilmann M, Kaaden OR. Equine viral arteritis with abortions: serological and virological evidence in Germany. J Vet Med B 1995;42:573–6. 22. Jaksch VW, Sibalin M, Taussig E, Pichler L, Burki F. Naturliche falle und experimentelle ubertragungen equiner virus-arteritis in Osterreich. Deut Tierarztl Woch 1973;80:317–40. 23. Monreal L, Villatoro AJ, Espada Y, Vinas L. Clinical signs of 1992 EVA outbreak in Barcelona. The first outbreak diagnosed in Spain. Swiss Vet 1993;11-S.
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Equine Viral Arteritis 24. Moraillon A, Moraillon R. Results of a serological survey of viral arteritis in France and in several European and African countries. In: Proceedings of the 4th International Conference on Equine Infectious Diseases. New Jersey: Veterinary Publications Inc., 1978; pp. 467–73. 25. Konishi S, Akashi H, Sentsui H, Ogata M. Studies on equine viral arteritis. I. Characterization of the virus and trial survey on antibody with vero cell cultures. Jpn J Vet Sci 1975;37:259–67. 26. Matumoto M, Shimizu TT, Ishizaki R. Constation d’anticorps contre le virus de l’arterite equine dans le serum de juments indiennes. In: Proceedings of the Societ´e Franco-Japonaise de Biologie. Societ´e Franco-Japanaise De Biologie, 1965; pp. 1262–4. 27. Paweska JT, Barnard BJH. Serological evidence of equine arteritis virus in donkeys in South Africa. Onderstepoort J Vet Res 1993;60:155–8. 28. Huntington PJ, Forman AJ, Ellis PM. The occurrence of equine arteritis virus in Australia. Aust Vet J 1990;67: 432–5. 29. van Maanen C, Heldens J, Cullinane AA, van den Hoven R, Westrate M. The prevalence of antibodies against equine influenza virus, equine herpesvirus 1 and 4, equine arteritis virus and equine rhinovirus 1 and 2 in Dutch standardbred horses. Vlaams Diergen Tijds 2005;74:140–5. 30. Hullinger PJ, Gardner IA, Hietala SK, Ferraro GL, MacLachlan NJ. Seroprevalence of antibodies against equine arteritis virus in horses residing in the United States and imported horses. J Am Vet Med Assoc 2001;219:946–9. 31. Timoney PJ. Factors influencing the international spread of equine diseases. Vet Clin N Am-Equine 2000;16:537–51. 32. Balasuriya UBR, Evermann JF, Hedges JF, McKeirnan AJ, Mitten JQ, Beyer JC, McCollum WH, Timoney PJ, MacLachlan NJ. Serologic and molecular characterization of an abortigenic strain of equine arteritis virus isolated from infective frozen semen and an aborted equine fetus. J Am Vet Med Assoc 1998;213:1586–9. 33. McCollum WH, Timoney PJ, Lee JW Jr, Habacker PL, Balasuriya UBR, MacLachlan NJ. Features of an outbreak of equine viral arteritis on a breeding farm associated with abortion and fatal interstitial pneumonia in neonatal foals. In: Wernery U, Wade JF, Mumford JA, Kaaden O-R (eds) Proceedings of the 8th International Conference on Equine Infectious Diseases. Newmarket: R&W Publications, 1999; pp. 559–60. 34. Timoney PJ. Equine viral arteritis. AAEP Report 2000; 7. 12. 35. Timoney PJ, Creekmore L, Meade B, Fly D, Rogers E, King B. 2006 Multi-state occurrence of EVA. In: Proceedings of the 110th Annual Meeting of the United States Animal Health Association, 2007, pp. 354–62. 36. Pitel PH, Legrand L, Marcillaud-Pitel C, Miszczack F, Guix E, Pronost S, Fortier G. L’epizootie d’arterite virale equine de 2007: diagnostic et epidemiosurveillance. Prat Vet Equine 2007;39:19–27. 37. Cavanagh D. Nidovirales: a new order comprising Coronaviridae and Arteriviridae. Arch Virol 1997;142:629–33. 38. Snijder EJ, Meulenberg JJM. The molecular biology of arteriviruses. J Gen Virol 1998;79:961–79. 39. Plagemann PGW, Moennig V. Lactate dehydrogenaseelevating virus, equine arteritis virus, and simian hemorrhagic fever virus: A new group of positive-strand RNA viruses. Adv Virus Res 1992;4:99–192. 40. Weber H, Beckmann K, Haas L. Equine arteritis virus (EAV)-induced abortion in alpacas. Deut Tierarztl Woch 2006;113:162–3.
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41. Fukunaga Y, McCollum WH. Complement-fixation reactions in equine viral arteritis. Am J Vet Res 1977;38:2043–6. 42. Balasuriya UBR, Patton JF, Rossitto PV, Timoney PJ, McCollum WH, MacLachlan NJ. Neutralization determinants of laboratory strains and field isolates of equine arteritis virus: Identification of four neutralization sites in the amino-terminal ectodomain of the GL envelope glycoprotein. Virology 1997;232:114–28. 43. McCollum WH, Timoney PJ. Experimental observation on the virulence of isolates of equine arteritis virus. In: Wernery U, Wade JF, Mumford JA, Kaaden O-R (eds) Proceedings of the 8th International Conference on Equine Infectious Diseases. Newmarket: R&W Publications, 1999; pp. 558–9. 44. Murphy TW, McCollum WH, Timoney PJ, Klingeborn B, Hyllseth B, Golnik W, Erasmus B. Genomic variability among globally distributed isolates of equine arteritis virus. Vet Microbiol 1992;32:101–15. 45. Timoney PJ, McCollum WH. The epidemiology of equine viral arteritis. In: Proceedings of the 32nd Annual Convention of the American Association of Equine Practitioners, 1986, pp. 545–51. 46. McCollum WH, Prickett ME, Bryans JT. Temporal distribution of equine arteritis virus in respiratory mucosa, tissues and body fluids of horses infected by inhalation. Res Vet Sci 1971;2:459–64. 47. McCollum WH. Responses of horses vaccinated with avirulent modified-live equine arteritis virus propagated in the E. Derm (NBL-6) cell line to nasal inoculation with virulent virus. Am J Vet Res 1986;47:1931–4. 48. Fukunaga Y, Imagawa H, Tabuchi E, Akiyama Y. Clinical and virological findings on experimental equine viral arteritis in horses. Bull Equine Res Inst 1981;18:110–18. 49. McCollum WH, Timoney PJ. The pathogenic qualities of the 1984 strain of equine arteritis virus. In: Proceedings of the Grayson Foundation International Conference of Thoroughbreed Breeders Organizations, 1984, pp. 34–47. 50. Scollay MC, Foreman JH. An overview of the 1993 equine viral arteritis outbreak at Arlington International Racecourse. In: Proceedings of the 39th Annual Convention of the American Association of Equine Practitioners, 1993, pp. 255–6. 51. Timoney PJ. Equine viral arteritis: how significant a threat to the horse? In: Albert PH, Morton T, Wade JF (eds) Proceedings of the 15th International Conference of Racing Analysts and Veterinarians. Newmarket: R & W Publications, 2005; pp. 238–44. 52. Golnik W. Viruses isolated from aborted foetuses and stillborn foals. In: Plowright W, Rossdale PD, Wade JF (eds) Equine Infectious Diseases VI: Proceedings of the 6th International Conference. Newmarket: R&W Publications, 1992; pp. 314. 53. Monreal L, Villatoro AJ, Hooghuis H, Ros I, Timoney PJ. Clinical features of the 1992 outbreak of equine viral arteritis in Spain. Equine Vet J 1995;27:301–4. 54. O’Connor B. Equine viral arteritis in a Thoroughbred filly. Can Vet J 1993;34:506–7. 55. Cole JR, Hall RF, Gosser HS, Hendricks JB, Pursell AR, Senne DA, Pearson JE, Gipson CA. Transmissibility and abortigenic effect of equine viral arteritis in mares. J Am Vet Med Assoc 1986;189:769–71. 56. Timoney PJ, McCollum WH. Equine viral arteritis. In: Coetzer JAW, Tustin RC (eds) Infectious Diseases of Livestock, Vol. 2, 2nd edn. Cape Town: Oxford University Press Southern Africa, 2004; pp. 924–32.
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57. Neu SM, Timoney PJ, Lowry SR. Changes in semen quality in the stallion following experimental infection with equine arteritis virus. Theriogenology 1992;37:407–31. 58. Crawford TB, Henson JB. Immunofluorescent, light microscopic and immunologic studies of equine viral arteritis. In: Proceedings of the 3rd International Conference of Equine Infectious Diseases. Basel: S. Karger, 1973; pp. 282–302. 59. Jones TC. Clinical and pathologic features of equine viral arteritis. J Am Vet Med Assoc 1969;155:315–17. 60. Prickett ME, McCollum WH, Bryans JT. The gross and microscopic pathology observed in horses experimentally infected with the equine arteritis virus. In: Bryans, Gerber H (eds) Equine Infectious Diseases III: Proceedings of the Third International Conference. Basel: S. Karger, 1973; pp. 265–72. 61. Moore BD, Balasuriya UB, Watson JL, Bosio CM, MacKay RJ, MacLachlan NJ. Virulent and avirulent strains of equine arteritis virus induce different quantities of TNF-alpha and other proinflammatory cytokines in alveolar and bloodderived equine macrophages. Virology 2003;314:662–70. 62. Coignoul FL, Cheville NF. Pathology of maternal genital tract, placenta, and fetus in equine viral arteritis. Vet Pathol 1984;21:333–40. 63. Del Piero F. Equine viral arteritis. Vet Pathol 2000;37:287–96. 64. Hedges JF, Balasuriya UBR, Timoney PJ, McCollum WH, MacLachlan NJ. Genetic divergence with emergence of novel phenotypic variants of equine arteritis virus during persistent infection of stallions. J Virol 1999;73:3672–81. 65. Holyoak GR, Little TV, McCollum WH, Timoney PJ. Relationship between onset of puberty and establishment of persistent infection with equine arteritis virus in the experimentally infected colt. J Comp Pathol 1993;109:29–46. 66. McCollum WH, Little TV, Timoney PJ, Swerczek TW. Resistance of castrated male horses to attempted establishment of the carrier state with equine arteritis virus. J Comp Pathol 1994;111:383–8. 67. Little TV, Holyoak GR, McCollum WH, Timoney PJ. Output of equine arteritis virus from persistently infected stallions is testosterone-dependent. In: Plowright W, Rossdale PD, Wade JF (eds) Equine Infectious Diseases VI: Proceedings of the 6th International Conference. Newmarket: R&W Publications, 1992; pp. 225–9. 68. Doll ER, Bryans JT, Wilson JC, McCollum WH. Immunization against equine viral arteritis using modified live virus propagated in cell cultures of rabbit kidney. Cornell Vet 1968;58:497–524. 69. McCollum WH. Vaccination for equine viral arteritis. In: Equine Infectious Diseases II: Proceedings of the
70. 71.
72.
73.
74.
75.
76.
77. 78.
79.
80.
81.
2nd International Conference. Basel: S. Karger, 1970; pp. 143–51. McCollum WH. Studies of passive immunity in foals to equine viral arteritis. Vet Microbiol 1976;1:45–54. McCollum WH, Timoney PJ, Roberts AW, Williard JE, Carswell GD. Response of vaccinated and non-vaccinated mares to artificial insemination with semen from stallions persistently infected with equine arteritis virus. In: Powell DG (ed.) Proceedings of the 5th International Conference on Equine Infectious Diseases. Lexington: The University Press of Kentucky, 1988; pp. 13–18. Timoney PJ, McCollum WH. Equine viral arteritis: Current clinical and economic significance. In: Proceedings of the 37th Annual Convention of the American Association of Equine Practitioners, 1991, pp. 403–9. Timoney PJ. Equine viral arteritis. In: OIE Manual of Standards for Diagnostic Tests and Vaccines, 6th edn. Paris: Office International des Epizooties, 2008; pp. 904–18. Timoney PJ, McCollum WH, Roberts AW. Detection of the carrier state in stallions persistently infected with equine arteritis virus. In: Proceedings of the Annual Meeting of the American Association of Equine Practitioners, 1987, pp. 57–65. Fortier G, Vidament M, deCraene F, Ferry B, Daels PF. The effect of GnRH antagonist on testosterone secretion, spermatogenesis and viral excretion in EVA-virus excreting stallions. Theriogenology 2002;58:425–7. Burger D, Janett F, Vidament M, Stump R, Fortier G, Imboden I, Thun R. Immunization against GnRH in adult stallions: Effects on semen characteristics, behaviour and shedding of equine arteritis virus. Anim Reprod Sci 2006;94:107–11. Morrell JM, Geraghty RM. Effective removal of equine arteritis virus from stallion semen. Equine Vet J 2006;38:224–9. McCollum WH. Development of a modified virus strain and vaccine for equine viral arteritis. J Am Vet Med Assoc 1969;155:318–22. McKinnon AO, Colbern GT, Collins JK, Bowen RA, Voss JL, Umphenour NW. Vaccination of stallions with a modified live equine viral arteritis virus. J Equine Vet Sci 1986;6:66–9. Timoney PJ, Umphenour NW, McCollum WH. Safety evaluation of a commercial modified live equine arteritis virus vaccine for use in stallions. In: Powell DG (ed.) Equine Infectious Diseases V: Proceedings of the Fifth International Conference. Lexington: The University Press of Kentucky, 1988; pp. 19–27. Newton JR. Controlling EVA in the 21st century: ‘zero tolerance’ or ‘live and let live’? Equine Vet Educ 2007;19:612–16.
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Chapter 247
Contagious Equine Metritis Peter J. Timoney
Introduction Contagious equine metritis (CEM) is a bacterial disease of the reproductive tract of equids that evidence would indicate is primarily transmitted by the venereal route.1–4 It was first recognized as an emergent equine disease in 1977 following extensive outbreaks of a contagious venereal infection on Thoroughbred breeding farms in the Newmarket area, United Kingdom (UK), and also in Ireland.5 There is strong circumstantial evidence to suggest, however, that a disease clinically resembling CEM had occurred on certain breeding farms in Ireland in 1975 and 1976 that was associated with the importation of a Thoroughbred stallion from continental Europe.5–7 Contagious equine metritis is a non-systemic reproductive disease, clinical signs of which are only manifest in the mare, not in the stallion.5, 8 Infection in the mare is principally characterized by an acute endometritis, cervicitis, and vaginitis of variable severity and a mucopurulent vaginal discharge, ranging from copious to minimal in amount.1–4 In one experimental study, salpingitis was also demonstrated in a significant percentage of cases.9 Abortion is a rarely reported sequel to infection.10 The etiological agent of this previously unrecognized disease is Taylorella equigenitalis (T. equigenitalis), a bacterium not known to exist prior to its initial isolation from typically affected mares and contaminated stallions in 1977.2–4,11, 12 T. equigenitalis can establish a carrier state of variable duration in both the stallion and the mare, which may last for years in the case of certain individuals.7, 13, 14 Contagious equine metritis has given rise to significant international concern since it was first identified as a novel venereal disease of equids in 1977.7, 11, 14 This was fueled by initial reports of the dramatic clinical features of the disease and how readily it could spread in breeding populations. International concerns focused on the economic impor-
tance of T. equigenitalis as a cause of short-term infertility in the mare, frequency of the carrier state in the stallion and the mare, and lastly, the contagiousness of the causal bacterium by direct or indirect venereal contact. As a consequence of international opinion, CEM has become one of the most regulated diseases with respect to the global movement of horses of breeding age and the trade in equine germplasm.7, 15 Since CEM was first discovered in 1977, the disease has been confirmed in equine populations in some 29 countries throughout the world including but not exclusive of those in many European countries, Japan, Australia, and North and South America.16 In countries in which CEM is currently known or believed to occur, the infection is endemic, largely in non-Thoroughbred breeds. On numerous occasions, the disease has been spread through the importation of a carrier stallion or mare.15 The risks of international dissemination of T. equigenitalis have been enhanced by changing commercial trends in the equine industry worldwide. These include: shuttling of stallions from northern to southern hemispheres or vice versa; shipment of mares between countries or hemispheres for breeding purposes; and finally, acceptance of the use of cryopreserved semen by the majority of horse breed registries.15 Over the years, experience has shown that the risks of introduction of CEM are greater when importing non-Thoroughbred stallions or mares from countries in which the disease is known or strongly suspected to be endemic, and a Code of Practice setting out the measures needed to prevent and control the disease is either lacking, or if available, not widely implemented.17 The 2008/9 disease event in the USA highlighted an important and frequently overlooked feature of acute CEM infection in the mare. It is an established fact that many such cases fail to develop overt signs of reproductive tract disease.11, 14 Widespread asymptomatic infection has been characteristic of numerous
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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outbreaks of CEM over the years. Under such circumstances, there is a significant risk of T. equigenitalis remaining undetected in a breeding horse population for months, or even years. This is believed to have been the case with respect to the most recent rediscovery of CEM in the USA in December 2008. Epidemiological evidence would indicate that the causal bacterium very likely had been in the country since 2000. Owners, breeders, and veterinarians must be alert to the oftentimes insidious nature of the infection and the need to include CEM in any differential diagnosis of short-term infertility in the mare, even in the absence of inflammatory changes in the proximal and distant reproductive tract and/or a vaginal discharge. Its presence can so easily be missed unless potential cases of infection are diagnostically evaluated for T. equigenitalis.
Causal agent The causal agent of CEM is T. equigenitalis, a Gramnegative, non-acid-fast, non-motile bacterium with fastidious growth requirements.2–4,11, 18 In terms of morphology, it frequently presents as a bipolar staining cocco-bacillus, especially in smears of vaginal exudate from an acutely affected mare. In longstanding cases of infection, the organism is more often pleomorphic, frequently bacillary in appearance.19 Two biotypes of T. equigenitalis exist, one that is streptomycin-sensitive and the other that is streptomycin-resistant.2, 4, 14, 20 In vitro assessment of the activity of strains of T. equigenitalis against different antimicrobials has repeatedly shown that they are sensitive to a wide range of antibiotics and disinfectants.11, 12, 18 Sensitivity/resistance patterns of the bacterium have remained virtually unchanged since its initial discovery in 1977. T. equigenitalis is not a particularly resistant organism outside the body, surviving in swabs coated with infective vaginal discharge for 48 hours at 20◦ C to 22◦ C and for up to 78 hours on swabs held at 4◦ C.21 It is rapidly killed by a range of commonly used disinfectants. Fomites such as artificial vaginas or a phantom mare can be readily contaminated if used with a carrier stallion, and serve as a source of T. equigenitalis for other stallions on the same premises unless appropriately decontaminated between semen collections.16 Investigative studies into the host range of T. equigenitalis have shown that, while CEM has been experimentally reproduced in donkeys,22 attempts to achieve transmission to cattle, sheep, swine,23 and cats have been unsuccessful.24 However, certain species of rodents, specifically congenic strains of mice, are susceptible to experimental infection when exposed by the intrauterine route.25 There is no evidence that T. equigenitalis is transmissible to humans.
Cultivation of T. equigenitalis requires enriched media such as Eugon chocolate agar containing a selection of antibiotics that control the majority of the bacterial contaminants that frequent the sites of persistence of this bacterium in the stallion and the mare. 26, 27 Primary isolation of the bacterium, which can be slow growing, requires that inoculated culture plates are incubated under microaerophilic conditions, i.e., in the presence of 5 to 10% carbon dioxide in air. An incubation time of at least 7 days is advisable before certifying cultures as negative for T. equigenitalis. While colonies of many strains of this bacterium can be visualized after 72 hours of incubation, there is evidence that the very occasional strain may not be detectable before 10 to 14 days.28 T. equigenitalis is a relatively unreactive bacterium, non-fermentative and non-proteolytic but positive for cytochrome oxidase, catalase, and alkaline phosphatase and on fluorescent antibody and latex agglutination testing.11, 26 It has been known for a significant number of years that strains of T. equigenitalis vary in pathogenicity for the mare.19, 29 In earlier experimental studies, mares challenged with larger colonial variants of the bacterium developed clinical signs of CEM, whereas those exposed to small colonial variants developed an inapparent infection.30 Aside from variation in pathogenicity, molecular genotyping of strains of T. equigenitalis using pulsedfield gel electrophoresis or crossed-field gel electrophoresis and restriction enzymes Apa 1, Not 1, and Nae 1 has confirmed the existence of a diversity of genotypes among countries in which CEM is known to occur.31 Strains from some countries are represented by a single genotype,32 whereas those from others comprise several different genotypes.33–35 At this point, no correlation has been established between a particular genotype(s) and its pathogenicity for the mare. For 20 years following its initial isolation in 1977, T. equigenitalis was the only species of Taylorella known to exist. Within a very short interval between late 1997 and early 1998, however, another species, designated T. asinigenitalis, was isolated from donkey jacks, a stallion, and several mares in California and Kentucky.36 The newly described organism is a very close ‘look-alike’ that shares many morphological, cultural, and biochemical characteristics with T. equigenitalis. It is genomically distinct enough, however, to be taxonomically classified as a separate species.36 Unlike T. equigenitalis, T. asinigenitalis has so far not been associated with naturally occurring disease in equids.
Transmission Transmission of T. equigenitalis occurs primarily by the venereal route.1–4,14 This very frequently takes
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Contagious Equine Metritis place through direct genital contact between an infected mare and a stallion, or vice versa. A mare can also be exposed to infection by artificial insemination with fresh-cooled or extended semen from a carrier stallion. By comparison with natural service, however, the risk of transmission by this means is very considerably less, provided that the extender used contains an appropriate antibiotic or antibiotics to which T. equigenitalis is sensitive (Klein et al., unpublished data). The potential risk of transmission of CEM through the use of cryopreserved semen, although considered minimal, has not yet been established. Transmission of T. equigenitalis can also be readily accomplished through indirect genital contact with fomites contaminated with the organism, such as vaginal specula, forceps, obstetrical sleeves, tail bandages, etc., used in examination of the reproductive tract of the mare, or by personnel failing to observe adequate hygienic precautions in handling mares or stallions at time of breeding.7, 14 Spread by this means was responsible for much of the dissemination of CEM on, and between, breeding farms in the UK and Ireland in 1977. Failure to observe appropriate sanitary measures when handling the external genitalia of stallions has also been implicated in the indirect transmission of the disease. Among the items considered of major importance as potential sources of contagion in stallion semen-collection centers is the external surface of the phantom, artificial vaginas, and wash buckets. There is strong circumstantial evidence implicating transmission of T. equigenitalis between stallions by this means at several of the stallion semen-collection centers involved in the 2008/9 CEM disease event in the USA. Although of lesser epidemiological significance, there is proof that some pregnant mares can continue to harbor T. equigenitalis in the uterus throughout pregnancy.37, 38 This may give rise to congenital infection of the fetus which, on rare occasions, can result in abortion.10 Alternatively, the fetus can survive the infection and be born normal but culture positive for T. equigenitalis.38 It is believed some foals out of carrier mares can also be exposed to T. equigenitalis through contact of their external genitalia with a CEM-positive placenta and/or contaminated clitoral area of the mare at time of foaling.39 Furthermore, it is conceivable that exposure of the postpartum foal can occur through contact of its external genitalia with bedding or pasture contaminated with infective placental fluids or vaginal discharge. A frequently overlooked yet potentially important mode of indirect venereal transmission of T. equigenitalis is through the use of contaminated wash cloths or sponges between colts or stallions.39 This has been implicated in the past39 and also, more recently, as the only explicable way transmission of T. equigenitalis could have oc-
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curred in colts and stallions under specific conditions of management.
Clinical response Contagious equine metritis is an acute, venereally transmissible disease that is restricted to the reproductive tract in the mare.1–4,11, 14, 40 It is a non-lifethreatening infection characterized by short-term infertility and, very rarely, abortion around 7 months’ gestation.10 In contrast, T. equigenitalis does not elicit a clinical response in the stallion, existing as a commensal on the external genitalia, causing neither a local nor a systemic inflammatory reaction.11, 14, 40 There has been one report, however, describing the isolation of T. equigenitalis from the testis, epididymis, seminal vesicles, and urethra of a stallion upon necropsy examination.41 This finding would indicate that, perhaps on rare occasions, exposure of stallions to the bacterium may lead to an ascending infection of the reproductive tract. The outcome of primary exposure of mares to T. equigenitalis can vary from overt disease to asymptomatic infection.11, 14, 29, 42 Mares, but not stallions, develop a humoral antibody response to T. equigenitalis that is usually short-lived.43–45 To date, there is no firm evidence of variation among horses or pony breeds with respect to susceptibility to T. equigenitalis infection. After an incubation period of 13 days, typically affected mares present with an odorless, grayish white, mucopurulent vaginal discharge of uterine origin.13, 19 This can vary considerably in amount, color, and consistency and can persist for 2 weeks or longer in untreated cases. In severe cases, the discharge may be so copious as to run down the hindquarters, wetting the entire perineal area and inside of the thighs, and matting the tail hairs.11, 14 The discharge is accompanied by an endometritis, cervicitis, and vaginitis of variable severity.1–4,14, 19, 40 Intrauterine fluid accumulation can often be detected on ultrasonographic examination in such cases. On vaginoscopic examination, discharge can be observed seeping between the folds of the external os of the cervix and accumulating on the floor of the vagina.19, 40 Irrespective of presence or absence of clinical signs of infection, most mares fail to conceive upon primary exposure to T. equigenitalis and return to estrus after a shortened diestrous period.7, 11, 14 Resultant infertility is short-term, however, and no long-term adverse effects have been observed in infected mares.11 Experimentally, re-exposure of mares previously affected with CEM was associated with minimal or zero clinical evidence of the disease.46–48 Persistence of T. equigenitalis in the reproductive tract of the mare need not compromise maintenance of a
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normal pregnancy and the birth of a healthy foal.37, 38 Some chronically infected pregnant mares may develop an intermittent vaginal discharge that can be indicative of a localized placentitis.3, 38
their ability to replicate in these cells.50 They found a positive association between ability to replicate in endothelial cells under in vitro conditions and the severity of clinical signs observed in infected mares.
Pathogenesis Several studies have been carried out on experimentally induced CEM in the mare.3, 9, 13, 47, 49 Within 3 days following intrauterine inoculation of T. equigenitalis, acutely infected mares develop a severe, diffuse endometritis and cervicitis, with accumulation of a variable quantity of a gray, watery exudate between the endometrial folds which become edematous and congested. The exudate increases in quantity over the following 7 to 12 days before starting to decrease in amount but increase in viscosity.9, 49 Mild multifocal salpingitis was a frequently observed feature in one experimental study.9 The endometrial changes were accompanied by edema and congestion of the mucous membranes of the cervix and vagina.49 Histologically, there was evidence of a severe, acute inflammation of the uterus and cervix within 2 days of experimental challenge. This was associated with edema of the lamina propria and neutrophil infiltration and migration through the endometrial surface. A decrease in the edema followed which was succeeded by an influx of lymphocytes, macrophages, and plasma cells. Cellular infiltration of the endometrium became primarily plasmacytic by day 14.9 The diffuse endometritis and cervicitis subsequently decreased in severity. However, a mild diffuse or multifocal lymphocytic endometritis and cervicitis persisted in mares in one experimental study that was terminated 116 days after challenge.9 In earlier studies by Platt et al.13 and Ricketts et al.,3 the authors described widespread destruction and denuding of the luminal epithelium in the acutely infected mare, leading to pseudo-stratification, regenerative activity in the uterine glands, and intercellular basal vacuolation of luminal epithelium. In the study by Wada et al.,49 T. equigenitalis organisms were observed multiplying on the surface of the endometrium and in the lumen of the uterine glands. There was no evidence, however, of bacterial invasion of the surface epithelium nor of destructive lesions involving the endometrium as described in the studies by Platt et al.13 and Ricketts et al.3 Differences in pathogenicity between the strains used in the respective studies and in the challenge dose of T. equigenitalis may have been responsible for the variation in pathological changes observed in the different groups of experimentally infected mares.9, 49 In a study using cultured endometrial cells, Nancy et al. showed that strains of T. equigenitalis differed in
Epidemiology A range of agent-, host-, and environment-related factors are involved in the epidemiology of CEM. These include, but are not necessarily exclusive of variation in, pathogenicity and other phenotypic characteristics among strains of T. equigenitalis,19, 29 modes of transmission,11, 14 occurrence of the carrier state in the stallion and the mare,40,51–53 industry trends influencing international/national movement of stallions and mares for breeding including the shipment of semen,14, 15 and finally, management practices in vogue when breeding stallions by natural service or collecting semen for artificial insemination (AI). For many years, it has been known that strains of T. equigenitalis can differ significantly from one another with respect to their inherent ability to cause a clinical response in previously unexposed mares. At the time CEM was first recognized as a highly contagious equine venereal infection, many outbreaks of the disease were characterized by a high clinical attack rate in naive mares.1–3 Within a year, however, the clinical features of the disease changed dramatically, with the majority of first-time exposed mares developing asymptomatic infections, and very few individuals exhibiting overt signs of reproductive disease.29, 54 That trend continues today, especially in countries and in breeds in which the disease is endemic. At this point, there is no specific explanation for the changing clinical pattern of CEM and no information on whether the long-term carrier stallion or mare is the source of genomic and phenotypic diversity, including alteration in pathogenicity, among naturally occurring strains of T. equigenitalis.
Modes of transmission The principal modes of transmission of T. equigenitalis, both direct and indirect, have already been described. Formulation of effective measures for the prevention and control of CEM is predicated, in large measure, not only on the different routes of transmission but also on the duration of bacterial shedding by the acutely infected mare.19 Determination of the pattern of shedding can present significant difficulties, however, when dealing with a chronically infected mare or carrier stallion.37, 55
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Contagious Equine Metritis Carrier state It is widely accepted that the single most important factor in the epidemiology of CEM is occurrence of the carrier state in the stallion and the mare.40, 55 On repeated occasions, the international spread of CEM has been identified with the importation of a carrier stallion and, less frequently, a carrier mare, into a country whose resident equine population is free from the disease.14, 15 Such introductions have, on occasion, had a significant economic impact on the equine industry in the importing country, as was attested to following the widespread occurrence of CEM in Kentucky in 1978.56 A variable percentage of stallions can become inapparent carriers of T. equigenitalis.11, 16, 40 The sites of persistence favored by the bacterium include the urethral fossa and urethral sinus or diverticulum, the terminal urethra, the external surface of the penis and the prepuce, and, less frequently, the pre-ejaculatory fluid.14, 26, 40, 55 Although duration of the carrier state in the stallion can be variable, T. equigenitalis can persist as a surface contaminant on the external genitalia of some stallions for an extended number of years.14, 15 The transmission rate of T. equigenitalis to naive mares bred by natural service can vary between carrier stallions.51 This is presumably influenced by concentration of the bacterium on the external genitalia and the level of innate resistance/susceptibility to infection of the individual mares being bred to a particular stallion. The risk of transmission is greatly reduced, however, if AI is used and semen is extended in a commercial extender containing antibiotics such as amikacin or ticarcillin, to which T. equigenitalis is sensitive (Klein et al., unpublished data). This was supported by the findings of the 2008/9 CEM disease event in the USA, when only 3 out of over 700 exposed mares, the vast majority bred with extended semen from carrier stallions, were confirmed to be long-term carriers.57 The transmission rate of T. equigenitalis may in fact have been higher than this statistic would indicate, had the exposed population of mares been sampled within the first 2 to 6 months after breeding. Early experimental studies have shown that the causal organism of CEM can persist for a period of several weeks to a number of months following the acute phase of the infection in a varying percentage of mares.13, 19 Because of the extended interval between breeding and eventual sampling of the mares traced in connection with the 2008/9 CEM event, any such short-term carrier mares clearly would not have been detected. The risk of spread of CEM using frozen semen from a carrier stallion has not been determined under field conditions. While it may be equivalent to
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what it might be using fresh-cooled semen, it could also be less. The additional steps of centrifugation and removal of most of the extended seminal plasma involved in processing semen for cryopreservation may reduce the concentration of T. equigenitalis below the threshold level needed to establish infection in most na¨ıve mares. An experimental study carried out by Klein et al. (unpublished data) in a limited number of mares failed to establish transmission of T. equigenitalis using frozen semen from a carrier stallion. Notwithstanding the findings of this study, the risk of transmission of CEM through the use of cryopreserved semen cannot be excluded. Additional to the stallion, the carrier state can also be established in a variable percentage of mares after primary infection with T. equigenitalis, irrespective of presence or absence of prior clinical signs of CEM. Frequency of the carrier state can be as high as 20 to 25%.58 While most persistently infected mares are asymptomatic carriers, a very small percentage may have an intermittent vaginal discharge.3, 38 Duration of the carrier state can be for weeks, months or years. Based on the site(s) of sequestration of T. equigenitalis, two types of carriers are recognized, clitoral and uterine.26, 55 In mares that are clitoral carriers (and these are in the majority), the bacterium is harbored in the clitoral sinuses (medial and lateral if present) and in the clitoral fossa. Uterine carriers are those in which infection persists in the proximal rather than the distal reproductive tract, namely, the uterus.7 The shedding pattern of T. equigenitalis by carrier mares can vary considerably, with significant fluctuations in the numbers of detectable organisms being observed in some animals over time.19, 37 Aside from the foregoing, T. equigenitalis can persist in the uterus during pregnancy and may be isolated from the placenta at foaling,38 and also, infrequently, from the external genitalia of the newborn foal, especially colt foals. If undetected, such individuals may continue to harbor the organism for an extended period, even after they have reached sexual maturity and can serve as a potential source of infection for future outbreaks of CEM.39 As already indicated, transmission of T. equigenitalis occurs through direct or indirect venereal contact with an infected mare or carrier stallion. A carrier teaser stallion should also not be overlooked as a potential source of transmitting CEM.7, 59 Apart from the acknowledged efficiency of natural breeding as a means of transmitting infection between an infected mare and a stallion, and vice versa, indirect contact through the use of contaminated fomites can play and has played a major role in spread of the disease. This was supported by the findings of the 2008/9 CEM event in the USA, which provided very strong circumstantial evidence of
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indirect stallion-to-stallion transmission of T. equigenitalis at several stallion breeding and semencollection centers in three of the states involved in that disease occurrence.
Immunity There is evidence to indicate that mares develop some level of local immunity following infection with T. equigenitalis.47 This would appear to provide a degree of clinical protection against disease, the duration of which has not been established. It is evident from early field studies on CEM that, while infection in the mare is characterized by short-term infertility, repeat live breeding to a carrier stallion does not necessarily prevent conception and establishment of pregnancy in all cases.
Industry trends As indicated previously, some of the changes that have taken place in the horse industry worldwide over the past 40 to 50 years have contributed significantly to the international dissemination of CEM. These include ‘shuttling’ of stallions between northern and southern hemispheres for breeding, and national and international movement of mares for breeding to specific sires.15 While AI with extended semen from a carrier stallion represents a much lesser risk of spread of the disease, the potential for transmission of T. equigenitalis using cryopreserved semen from a similar source remains to be established at this point.
Economic significance The occurrence of CEM in the Newmarket area of the UK and in Ireland in 1977 resulted in major disruption of that year’s Thoroughbred breeding season in both countries, and a significant decline in stallion income and live foal percentages for infected mares. The economic impact of the disease event in the UK alone was estimated at the time to be in the region of $20–30 million. The first known occurrence of CEM in Kentucky, USA, in the following year, even though it was relatively short-term in duration, had a similar disruptive effect on the state’s Thoroughbred breeding industry. Lost income from stallion breeding fees, failure to get mares in foal, transportation bans and associated losses were estimated at $1 million for every day that mares were not bred and movement was restricted, until CEM was brought under control several weeks into the event.60 It took an estimated $13.5 million to eradicate the disease. The most recent finding of CEM in the USA in late 2008
has come at considerable cost to the USDA and to the horse industry. Rediscovery of the disease came to light with the identification of the carrier state in a Quarterhorse stallion that was being routinely screened for T. equigenitalis prior to approval of his semen for export.57 A total of 23 stallions, including one that is currently a gelding, have been confirmed positive for T. equigenitalis over the course of very extensive federal and state field investigations that have been carried out to date. Five mares have also been cultured positive for the organism. None of the aforementioned carriers were considered the source of this disease event. The 23 positive stallions were found in seven states: Georgia, Illinois, Indiana, Iowa, Kentucky, Texas, and Wisconsin. The five positive mares were located in California, Illinois, and Wisconsin. In addition to the CEM-positive stallions and mares, a further 977 horses were considered exposed to T. equigenitalis, either directly or indirectly. A total of 1005 positive/exposed horses have been traced to all but the states of Hawaii and Rhode Island. Expenses incurred to this point in undertaking the very extensive field investigations and tracing of animals involved, the testing and treatment of culture positive stallions and mares and related expenses, have already totaled several million dollars. A more detailed description of the 2008/9 occurrence is available from USDA, APHIS.57
Diagnosis Contagious equine metritis cannot be diagnosed on clinical grounds alone because of its similarity to other, more commonly encountered, bacterial infections of the reproductive tract of the mare.14 These can be caused by: Streptococcus zooepidemicus, Pseudomonas aeruginosa, certain capsule types (1, 2, 5, and possibly 7 and 39) of Klebsiella pneumoniae, Escherichia coli, and a miscellany of other bacteria.26 Bacteriological culture is still widely accepted as the most definitive means of confirming a diagnosis of CEM in the mare and the stallion.16 Notwithstanding problems inherent in attempting isolation of a slow-growing, highly fastidious organism from heavily contaminated sites in the horse, culture of T. equigenitalis has the advantage of providing the opportunity to determine a strain’s cultural, antimicrobial sensitivity, pathogenic and antigenic characteristics. Contagious equine metritis should always be considered in a differential diagnosis when encountering mares that develop a mucopurulent vaginal discharge 1 to 13 days after being bred or returning prematurely in estrus after a shortened diestrous period.14 While the polymerase chain reaction (PCR) assay offers a more rapid and cost-effective test than culture for the detection of
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Contagious Equine Metritis T. equigenitalis on swabs from mares and stallions,61–66 it has yet to be fully validated for detection of the bacterium in fresh-cooled and cryopreserved semen as well as in equine embryos. The assay is especially useful, however, when attempting detection of T. equigenitalis on swabs from the heavily contaminated sites of persistence in the mare and the stallion and from mares and stallions recently treated with antibiotics. When encountering cases of mares with a vaginal discharge, the possibility of CEM should not be overlooked. Smears should be made of the discharge, stained, and examined for evidence of bacteria morphologically resembling T. equigenitalis either extracellularly or phagocytosed by neutrophils.4, 14, 19, 67 If present, the evidence is suggestive, but of itself, not confirmatory of a diagnosis of CEM. The sites of choice for detection of T. equigenitalis in the barren, maiden or post-parturient mare include the uterus and, if unable to obtain an endometrial swab, the cervix, together with the clitoral sinuses and clitoral fossa.14, 26 It is important to use swabs of smaller diameter, pre-moistened with sterile water, to ensure effective sampling of the clitoral sinuses.55 There is some evidence to indicate that swabbing the uterus/cervix in early estrus optimizes the chances of detection of T. equigenitalis.14 The clitoral fossa and clitoral sinuses and any vulvar discharge that may be present are the only sites/specimens to be swabbed in the pregnant mare.16 In the case of the stallion, the preferential sites of persistence of T. equigenitalis are the urethral fossa and urethral sinus or diverticulum, the terminal urethra, and the external surface of the penis and the prepuce.11, 14, 40 Although pre-ejaculatory fluid has historically been included among the specimens collected from a putative carrier stallion, experience has shown it is not as diagnostically reliable as the other aforementioned sites for the detection of T. equigenitalis.39 Swabbing of the sites of persistence in the stallion is optimized when the penis is fully erect at time of sampling.14 Additional to bacteriological culture and PCR assay, the carrier state in the stallion can also be detected by test breeding a stallion by natural service to two T. equigenitalis-cultured and serologically negative mares.26 After breeding, the mares are checked for evidence of infection by repeated culture or PCR assay and also for seroconversion by testing blood collected between days 21 and 45 for the development of complement-fixing antibodies to the bacterium. Swabs collected for the diagnosis of CEM should be obtained by a veterinarian to ensure the appropriate sites are reliably sampled. To enhance the chances of isolation of T. equigenitalis from mares or stallions,
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three sets of swabs should be collected at intervals of 3–7 days. Mares or stallions that have received systemically or locally administered antibiotics should not be sampled for 7 days or 21 days, respectively, after the completion of treatment. In handling swabs collected for CEM diagnosis, every effort must be made to enhance survival of T. equigenitalis if present, and to control replication of other bacterial species that colonize the sampling sites. This can best be achieved by inserting swabs into individual tubes of antibiotic-free Amies transport medium with charcoal immediately after collection68 and transporting the specimens, preferably under conditions of refrigeration, to a laboratory officially approved for the purpose of testing for CEM. 14, 26 Swabs must be received in satisfactory condition and plated out within 48 hours of collection. Serological testing for the diagnosis of T. equigenitalis infection is only applicable in the mare and not in the stallion, which does not develop a detectable serum antibody response to the bacterium.14 Even in the mare, serological testing is only of value in detecting recent exposure to T. equigenitalis.14, 19, 44 As the humoral antibody response is, in the majority of cases, relatively short-lived, serology has no reliability as a means of identification of the carrier state in the mare.54
Treatment From a clinical perspective, treatment applies only to a mare or mares acutely affected with CEM for the purpose of expediting resolution of clinical signs. Mares with acute endometritis can be effectively treated by the intrauterine infusion of ampicillin, 2 g in 60 mL of normal saline or gentamicin (buffered) for 3 to 5 days.69 This may be combined with sulfamethoxazole trimethoprim 30 mg/kg administered orally for 5 days. It has been suggested that treatment of the acutely affected mare may be contraindicated in that it may enhance the chances of persistence of T. equigenitalis in the clitoral area after the organism has been cleared from the proximal reproductive tract.7 As yet, however, there is no firm evidence in support of such speculation. By far the more important application of treatment is to eliminate T. equigenitalis from its sites of persistence in the mare and the stallion. Treatment of clitoral carrier mares must be preceded by removal of any smegma-like material from the clitoral sinuses by flushing them with a cerumenolytic agent (a compound that softens and facilitates removal of cerumen), followed by irrigation of the sinuses and clitoral fossa with a 4% solution of chlorhexidine gluconate.69 Sinuses and fossa are
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packed with 0.2% nitrofurazone or 1% silver sulfadiazine ointment. Nitrofurazone ointment should not be used in mares or stallions that eventually could be processed for human consumption. The clitoral fossa and sinuses are scrubbed for 4 more days with 4% chlorhexidine solution and packed on each occasion with 0.2% nitrofurazone or 1% silver sulfadiazine ointment.16 Mares that are intrauterine carriers of T. equigenitalis need to be treated by the intrauterine infusion of ampicillin, ticarcillin, or gentamicin together with systemic administration of sulfamethoxazole trimethoprim, as already described. It is advisable to combine the foregoing with topical treatment of the distal reproductive tract to eliminate the risk of localization of T. equigenitalis in the clitoral sinuses and fossa. Some mares may require several courses of treatment before a successful outcome is obtained.16 Very infrequently, surgical ablation of the clitoral sinuses may be necessary to eliminate the carrier state.70 The first step in treatment of the carrier stallion is, with the penis fully extruded, to remove all smegma from the penis and prepuce, urethral fossa, and urethral sinus or diverticulum by thorough washing of the external genitalia with a 4% solution of chlorhexidine gluconate.69 After drying of the penis and prepuce, the latter together with the urethral fossa and urethral sinus or diverticulum are liberally anointed and packed with 0.2% nitrofurazone or 1% silver sulfadiazine ointment. This regimen is repeated daily for 5 days. While in many cases one course of treatment is successful in eliminating the carrier state, the occasional stallion may require repeat treatment(s) before it is finally cleared of T. equigenitalis. In such cases, it is advisable to consider supplementing local treatment with a 5- to 10-day course of parenterally administered antibiotics, as described for the mare. Treated stallions and mares must be retested for presence of CEM no less than 21 days after completion of treatment, to confirm their freedom from T. equigenitalis.69 Efforts to treat extended or fresh-cooled semen from a carrier stallion to render it completely noninfective have met with mixed success. In a very early experimental study of the survival of T. equigenitalis in ‘spiked’ semen, of three antibiotic-containing extenders evaluated, one supplemented with gentamicin proved most effective when extended semen was held at 22 ± 2◦ C rather than 4◦ C or 37◦ C.71 However, such treatment did not entirely eliminate infectivity under the conditions of the study. This is supported by the findings of a recent study in which, using an extender containing amikacin and penicillin or ticarcillin, infectivity levels of T. equigenitalis in extended semen from a carrier stallion were greatly reduced, but not entirely eliminated (Klein et al., unpublished data). The procedures used in processing semen
for cryopreservation may reduce infectivity levels even lower than those in fresh-cooled or extended semen.
Prevention and control Contagious equine metritis is a preventable and controllable disease. Of singular importance in any national prevention program is the need to make the disease notifiable or, if non-notifiable, reportable.7, 11, 72 Furthermore, broadly based horse industry support for any such program is crucial to ensuring its success. The availability of animal health laboratories with the requisite expertise and experience in the diagnosis of CEM is an essential component of any prevention and control initiative.7 It should be noted that currently there is no vaccine to immunize against CEM.14, 15 Consideration of how best to prevent and control CEM is largely influenced by whether a country is free of the disease or whether the disease is endemic in its equine population. In the former instance, emphasis has to be placed on seeking pre-entry certification of freedom of infection when importing stallions and mares from known CEM-affected countries.14, 15 This must be complemented by officially requiring that all such imports also be subjected to post-entry quarantine and testing, to confirm their negative carrier status for T. equigenitalis. The nature and extent of diagnostic testing for this bacterium in the stallion and the mare should be in full accordance with what has already been described. In the case of imported stallions, it is imperative that post-entry screening include not just bacteriological culture and PCR assay but also test breeding. It is worth emphasizing that close to 70% (16/23) of the stallions confirmed carriers of T. equigenitalis on post-entry testing in the US between 1997 and 2007 were detected by test breeding alone and not by bacteriological culture (Timoney, unpublished data). This finding may be biased in that it has been rumored that many of the imported stallions had been extensively treated for CEM in the country of origin shortly before being subjected to pre-export testing for CEM. Notwithstanding the rigor and expense of the post-entry testing requirements in the US, there is always the risk of a carrier mare, and especially a stallion, escaping detection and being released from quarantine. This very scenario is the most likely explanation to account for the reintroduction of CEM into the US, perhaps as far back as 2000, but whose presence was only discovered by chance in late 2008. Certain countries that have experienced the introduction of CEM into their respective equine populations or in which the disease is endemic have developed voluntary Codes of Practice.72 Such Codes
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Contagious Equine Metritis provide practical guidelines for veterinarians, owners, and breeders alike on how to prevent and control CEM and other venereally transmissible diseases. Control of CEM is based on comprehensive bacteriological screening of breeding stallion and mare populations, to identify asymptomatic carrier animals and possible clinical cases of infection. Mares and stallions positive for T. equigenitalis must be isolated and not bred until treated and confirmed free from the bacterium. The Codes emphasize the importance of improved standards of hygiene on breeding farms and the benefits of using disposable or sterile materials and equipment. Failure to uphold adequate biosecurity and sanitary measures at various stallion breeding and semen-collection centers would appear to have been important for indirect stallionto-stallion transfer of T. equigenitalis on premises involved in the 2008/9 CEM event in the USA. The potential risks of transmission of CEM by AI with extended semen must not be overlooked, even if they are considerably less than if breeding by natural service. Experience has shown that where Codes of Practice have been widely implemented in countries, it has been possible to bring CEM under rigorous control, and in some instances even to successfully eradicate the disease. Contagious equine metritis remains a disease of international importance, with the potential to cause considerable economic loss in previously unexposed breeding populations. The ease with which CEM can be spread internationally through the movement of carrier stallions and mares represents a continuing threat to countries with established freedom from the disease. The re-emergence of CEM in the USA in late 2008 underscores the importance of having a national monitoring, surveillance, and reporting program for venereal diseases such as CEM that, once introduced, can so readily be disseminated within a country through movement of stallions, mares or through shipment of infective semen. All sectors of the equine industry must be ever vigilant if such disease incursions are to be detected at the earliest opportunity, and the risk of any of them becoming endemic is to be averted. The responsibility lies primarily with the horse industry to ensure that the appropriate precautions are taken to safeguard it from the potential economic consequences of CEM and analogous diseases.
References 1. Crowhurst RC. Genital infection in mares. Vet Rec 1977;100:476. 2. Platt H, Atherton JG, Simpson DJ, Taylor CE, Rosenthal RO, Brown DFJ, Wreghitt TG. Genital infection in mares. Vet Rec 1977;101:20.
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3. Ricketts SW, Rossdale PD, Wingfield-Digby NJ, Falk MM, Hopes R, Hunt MDN, Peace CK. Genital infection in mares. Vet Rec 1977;101:65. 4. Timoney PJ, Ward J, Kelly P. A contagious genital infection of mares. Vet Rec 1977;101:103. 5. Powell DG. Contagious equine metritis. Equine Vet J 1978;10:1–4. 6. Timoney PJ. Contagious equine metritis in Ireland (1977–1978). Irish Vet J 1979;33;14–15. 7. Timoney PJ, Powell DG. Contagious equine metritis: epidemiology and control. J Equine Vet Sci 1988;8:42–6. 8. Platt H, Atherton JG. Contagious equine metritis. Vet Rec 1977;101:434. 9. Acland HM, Kenney RM. Lesions of contagious equine metritis in mares. Vet Path 1983;20:330–41. 10. Nakashiro H, Naruse M, Sugimoto C, Isayama Y, Kuniyaou C. Isolation of Haemophilus equigenitalis from an aborted equine fetus. Nat Inst Anim Hlth Q (Tokyo) 1981;21:184–5. 11. Platt H, Taylor CE. Contagious equine metritis. Med Microbiol 1982;1:49–96. 12. Sugimoto C, Isayama Y, Sakazaki R, Kuramochi S. Transfer of Haemophilus equigenitalis Taylor et al. 1978 to the genus Taylorella gen. nov. as Taylorella equigenitalis comb. nov. Curr Microbiol 1983;9:155–62. 13. Platt H, Atherton JG, Simpson DJ. The experimental infection of ponies with contagious equine metritis. Equine Vet J 1978;10:153–9. 14. Powell DG. Contagious equine metritis. Adv Vet Sci Comp Med 1981;25:161–84. 15. Timoney PJ. Factors influencing the international spread of equine diseases. In: Timoney PJ (ed.) Veterinary Clinics of North America: Equine Practice. Philadelphia: Saunders, 2000; pp. 537–51. 16. Timoney PJ. Contagious equine metritis. Comp Immun Microbiol Infect Dis 1996;19:199–204. 17. Timoney PJ. Infectious diseases and international movement of horses. In: Sellon D, Long M (eds) Infectious Diseases of Horses, 1st edn. St Louis, MO: Saunders Elsevier, 2007; pp. 549–56. 18. Taylor CE, Rosenthal RO, Brown DF, Lapage SP, Hill LR, Legros RM. The causative organism of contagious equine metritis 1977: proposal for a new species to be known as Haemophilus equigenitalis. Equine Vet J 1978;10:136–44. 19. Timoney PJ, McArdle JF, O’Reilly, PJ, Ward J. Infection patterns in pony mares challenged with the agent of contagious equine metritis 1977. Equine Vet J 1978;10:148–52. 20. Swerczek TW. Contagious equine metritis in the USA. Vet Rec 1978;102:512–13. 21. Timoney PJ, Harrington AM, McArdle JF, O’Reilly PJ. Survival properties of the causal agent of contagious equine metritis 1977. Vet Rec 1978;102:152. 22. Timoney PJ, O’Reilly PJ, McArdle JF, Ward J, Harrington AM. Contagious equine metritis: experimental infection in the donkey. Vet Microbiol 1985;10:259–68. 23. Timoney PJ, O’Reilly PJ, McArdle JF, Ward J. Attempted transmission of contagious equine metritis 1977 to other domestic animal species. Vet Rec 1978;102:152. 24. Timoney PJ, Shin SJ, Lein DH, Jacobson RH. Transmissibility of the contagious equine metritis organism for the cat. Comp Immunol Microbiol Inf Dis 1984;7:131–40. 25. Timoney PJ, Shin SJ, Jacobson RH. Variable persistence of the contagious equine metritis organism in the genital tract of CBA/J, CBA/N, LAF1/J, BALB/c and congenitally thymus-deficient (nude) mice. J Comp Pathol 1985;95:137–49.
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26. Heath P, Timoney P. Contagious equine metritis. In: O.I.E. Manual for Diagnostic Tests and Vaccines for Terrestrial Animals, 6th edn. Paris: Office International des Epizooties, 2008; pp. 838–44. 27. Timoney PJ, Shin SJ, Jacobson RH. Improved selective medium for isolation of the contagious equine metritis organism. Vet Rec 1982;111:107–8. 28. Ward J, Hourigan M, McGuirk J, Gogarty A. Incubation times for primary isolation of the contagious equine metritis organism. Vet Rec 1984;114:298. 29. Timoney PJ. CEM in Ireland. Vet Rec 1978;103:475–6. 30. Kanemaru T, Kamada M, Anzai T, Kumanomido T. Contagious equine metritis: the pathogenicity for mares of small and large colonial variants of Taylorella equigenitalis isolated from a laboratory strain. In: Powell DG (ed.) Equine Infectious Diseases V: Proceedings of the Fifth International Conference. Lexington, KY: University Press of Kentucky, 1988; pp. 155–63. 31. Matsuda M, Moore JE. Recent advances in molecular epidemiology and detection of Taylorella equigenitalis associated with contagious equine metritis (CEM). Vet Microbiol 2003;97:111–22. 32. Thoresen SI, Jenkins A, Ask E. Genetic homogeneity of Taylorella equigenitalis from Norwegian trotting horses revealed by chromosomal DNA fingerprinting. J Clin Microbiol 1995;33:233–4. 33. Kagawa S, Klein F, Corboz L, Moore JE, Murayama O, Matsuda M. Demonstration of heterogeneous genotypes of Taylorella equigenitalis isolated from horses in six European countries by pulsed-field gel electrophoresis. Vet Res Commun 2001;25:565–75. 34. Matsuda M, Miyazawa T, Moore JE, Buckley TC, Thomas LA. Molecular genotyping by pulsed-field gel electrophoresis of restricted genomic DNA of strains of Taylorella equigenitalis isolated in Ireland and in the United States. Vet Res Comm 1998;22:217–24. 35. Matsuda M, Kagawa S, Sakamoto Y, Miyajima M, Barton M, Moore JE. Detection of heterogeneous genotypes among Australian strains of Taylorella equigenitalis. Austr Vet J 2000;78:56–7. 36. Jang SS, Donahue JM, Arata AB, Goris J, Hansen LM, Earley DL, Vandamme PAR, Timoney PJ, Hirsh DC. Taylorella asinigenitalis sp. nov., a bacterium isolated from the genital tract of male donkeys (Equus asinus). Int J Syst Evol Micr 2001;51:971–6. 37. Timoney PJ, Ward J, McArdle JF. CEM and the foaling mare. Vet Rec 1978;102:246–7. 38. Powell DG, Whitwell K. The epidemiology of contagious equine metritis (CEM) in England 1977–1978). J Reprod Fertil Suppl 1979;27:331–5. 39. Timoney PJ, Powell DG. Isolation of the contagious equine metritis organism from colts and fillies in the United Kingdom and Ireland. Vet Rec 1982;111:478–82. 40. Platt H, Atherton JG, Dawson FL, Durrant DS. Developments in contagious equine metritis. Vet Rec 1978;102:19. 41. Schluter H, Kuller HJ, Friedrich U, Selbitz HJ, Marwitz T, Beyer C, Ullrich E. Epizootiology and treatment of contagious equine metritis (CEM), with particular reference to the treatment of infected stallions. Praktische Tierarztliche 1991;72:503–11. 42. Timoney PJ. Contagious equine metritis in Ireland. Vet Rec 1979;105:172–3. 43. Benson JA, Dawson FLM, Durrant DS, Edwards PT, Powell DG. Serological response in mares affected by contagious equine metritis 1977. Vet Rec 1978;102:277–80.
44. Bryans JT, Darlington RW, Smith B, Brooks RR. Development of a complement fixation test and its application to diagnosis of contagious equine metritis. J Equine Med Surg 1979;3:467–72. 45. Croxton-Smith P, Benson JA, Dawson FLM, Powell DG. A complement fixation test for antibody to the contagious equine metritis organism. Vet Rec 1978;103:275–8. 46. Fernie DS, Batty I, Walker PD, Platt H, Mackintosh ME, Simpson DJ. Observations on vaccine and post-infection immunity in contagious equine metritis. Res Vet Sci 1980;28:362–7. 47. Sahu SP, Pierson RE, Dardiri AH. Contagious equine metritis: effect of intrauterine inoculation of contagious equine metritis agent in pony mares. Am J Vet Res 1980;41:5–9. 48. Timoney PJ, O’Reilly PJ, McArdle JF, Ward J, Harrington AM. Responses of mares to rechallenge with the organism of contagious equine metritis 1977. Vet Rec 1979;104:264. 49. Wada R, Kamada M, Fukunaga Y, Kumanomido T. Studies on contagious equine metritis. IV. Pathology in horses experimentally infected with Haemophilus equigenitalis. Bull Equine Res Inst 1983;20:133–43. 50. Nancy M, Bleumink-Pluym C, Laak EA, Houwers D, van der Zeist BA. Differences between Taylorella equigenitalis strains in their invasion of and replication in cultured cells. Clin Diag Lab Immunol 1996;3:47–50. 51. Bryans JT, Hendricks JB. Epidemiological observations on contagious equine metritis in Kentucky, 1978. J Reprod Fertil Suppl 1979;27:343–9. 52. Crowhurst RC, Simpson DJ, Greenwood RE, Ellis DR. Contagious equine metritis. Vet Rec 1979;104:465. 53. Swerczek TW. Contagious equine metritis – outbreak of the disease in Kentucky and laboratory methods for diagnosing the disease. J Reprod Fertil Suppl 1979;27:361–5. 54. Day FT, Crowhurst RC, Simpson DJ, Greenwood RE, Ellis DR, Eaton Evans W. An outbreak of contagious equine metritis in 1977 and its effect the following season. J Reprod Fertil Suppl 1979;27:351–4. 55. Simpson DJ, Eaton Evans WE. Isolation of the CEM organism from the clitoris of the mare. Vet Rec 1978;102:19–20. 56. Holden C. Outbreak of equine VD stirs fear in Kentucky. Science 1978;200:181–5. 57. USDA, APHIS 2010, Animal Health. (Contagious Equine Metritis. http://www.aphis.usda.gov/newsroom/ hot issues/cem/index.shtml Accessed May 24, 2010). 58. Wood JLN, Cardwell J, Castillo-Olivares J, Irwin V. Transmission of diseases through semen. In: Samper JC, Pycock J, McKinnon AO (eds) Current Therapy in Equine Reproduction. Philadelphia: Saunders, 2007; pp. 266–74. 59. Dingle PJ. Contagious metritis in mares. Vet Rec 1977;101:214. 60. Kristula MA, Smith BI. Diagnosis and treatment of four stallions, carriers of the contagious metritis organism – case report. Theriogenology 2004;61:595–601. 61. Arata AB, Cooke CL, Jang SS, Hirsh DC. Multiplex polymerase chain reaction for distinguishing Taylorella equigenitalis from Taylorella equigenitalis-like organisms. J Vet Diagn Invest 2001;13:263–4. 62. Anzai T, Eguchi M, Sekizaki T, Kamada M, Yamamoto K, Okuda T. Development of a PCR test for rapid diagnosis of contagious equine metritis. J Vet Med Sci 1999;61:1287–92. 63. Chanter N, Vigano F, Collin NC, Mumford JA. Use of a PCR assay for Taylorella equigenitalis applied to samples from the United Kingdom. Vet Rec 1998;143:225–7. 64. Miserez R, Frey J, Krawinkler M, Nicolet J. Identification and diagnosis of Taylorella equigenitalis by a DNA
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Contagious Equine Metritis amplification method (PCR). Schweizer Archiv fur Tierheilkunde 1996;138:115–20. 65. Premanandh J, George LV, Wernery U, Sasse J. Evaluation of a newly developed real-time PCR for the detection of Taylorella equigenitalis and discrimination from T. asinigenitalis. Vet Microbiol 2003;95:229–37. 66. Wakeley PR, Errington J, Hannon S, Roest HI, Carson T, Hunt B, Sawyer H, Heath P. Development of a real-time PCR for the detection of Taylorella equigenitalis directly from genital swabs and discrimination from Taylorella asinigenitalis. Vet Microbiol 2006;118:247–54. 67. Wingfield Digby NJ. The technique and clinical interpretation of endometrial cytology in mares. Equine Vet J 1978;10:167–70.
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68. Atherton JG. Isolation of CEM [contagious equine metritis] organism. Vet Rec 1978;102:67. 69. Kristula M. Contagious equine metritis. In: Sellon DC, Long MT (eds) Equine Infectious Diseases. St Louis, MO: Saunders Elsevier, 2007; pp. 351–3. 70. Swerczek TW. Elimination of CEM organism from mares by excision of clitoral sinuses. Vet Rec 1979;105:131–2. 71. Timoney PJ, O’Reilly PJ, Harrington AM, McCormack R, McArdle JF. Survival of Haemophilus equigenitalis in different antibiotic-containing semen extenders. J Reprod Fertil Suppl 1979;27:377–81. 72. Powell DG, David JS, Frank CJ. Contagious equine metritis: the present situation reviewed and a revised code of practice for its control. Vet Rec 1978;103:399–402.
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Chapter 248
Mare Reproductive Loss Syndrome David G. Powell
Introduction During April and May of 2001 the pregnant mare population of central Kentucky and southern Ohio, comprising multiple breeds of horses, experienced a high level of fetal loss. They included early (35–100day) fetal losses (EFL) plus late-term abortions and the birth of weak foals, many of which died, referred to as late fetal losses (LFL). With no immediate explanation for the cause it was identified as mare reproductive loss syndrome (MRLS). Also during May equine clinicians diagnosed an increased incidence of unilateral endophthalmitis and pericarditis among horses of all breeds, ages, and sexes. In the following year toward the end of April 2002, cases of MRLS were again reported on several farms in central Kentucky albeit at a reduced level compared to 2001. During subsequent years reports of MRLS have been extremely sporadic, with cases reported in Florida, Kentucky, and New Jersey. As a result of the outbreaks that occurred during 2001 and 2002 a case definition for MRLS was developed1 as follows. Early fetal loss (EFL) occurred within 35 to 100 days after mares were bred. In mares that underwent ultrasonographic examination during this period, fetal death followed by expulsion of the fetus was associated with the presence of abnormal echogenic fluid (cloudy and flocculent) around the fetus. Late fetal loss (LFL) occurred as abortion during the last trimester of pregnancy. LFL was associated in many but not all cases with a swollen and engorged placenta (premature placental separation or ‘red bag’ syndrome). Foals born alive were often weak and required intensive veterinary care, but many died. In addition to reproductive losses, a number of cases of pericarditis (inflammation surrounding the heart) and severe unilateral uveitis (inflammation
of one eye) were associated with the occurrence of MRLS. A small number of cases of suppurative encephalitis were also reported in adult horses during the outbreak of MRLS in 2001.2 A study undertaken by the Department of Equine Business at the University of Louisville beginning in the summer of 2001 estimated the economic loss to the equine industry in Kentucky over the 4-year period 2000–2003 at $336 million.3 Estimated figures for subsequent losses in 2002 raised that figure to approximately $500 million. Efforts to identify the origin and cause of MRLS have concentrated on epidemiological investigations followed by experimental challenge studies. The findings focused on the role of the eastern tent caterpillar (ETC) (Malacosoma americanum) as the source of the outbreak although the exact cause has still to be identified.
Clinical description Veterinarians specializing in equine reproductive practice were examining pregnant mares bred during February through March during the last week of April 2001 utilizing ultrasonography to determine the sex of the fetus (Figure 248.1). They reported an unusually high number of non-viable fetuses, lacking a heartbeat, plus the presence of cloudy, flocculent material in the allantoic and amniotic fluids. Subsequent examination confirmed the fetus was no longer present and cases were referred to as EFL. Mares typically showed no clinical signs and rarely were the expelled fetuses identified within the fetal membranes and fluids. Occasionally a mare showed a serosanguineous or purulent discharge from the vulva. Some mares showed the fetus and membranes protruding from the vulva (Figure 248.2) or the fetus located either in the vagina or uterus. A small
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Fetal Sexing Ultrasound Examination 60-70 days Normal
Early Fetal Loss
allantois fetus amnion genital tubercle
fetus
Figure 248.1 Fetal sexing, ultrasound examination at 60–70 days gestation. (a) Normal fetus, and (b) early fetal loss. Reproduced courtesy of Dr Walter Zent, Hagyard Equine Medical Institute.
Figure 248.2 Expelled early fetal loss (EFL). Reproduced courtesy of Dr Stuart E. Brown II, Hagyard Equine Medical Institute.
number (< 5%) exhibited mild signs of colic, abdominal straining, or low-grade fever 1 to 3 days before EFL occurred. During the same period the Livestock Disease Diagnostic Center (LDDC) in Lexington reported a significant increase in the number of late-term abortions submitted for postmortem examination. Clinical signs in mares described by Byars and Seahorn4 were inconsistent and varied from normal gestational term to within 30 days of foaling. The majority displayed no signs of impending parturition, although others exhibited restlessness, discomfort, sweating, mild signs of colic, muscular tremors, and stiffness resembling acute laminitis. In many cases delivery was described as explosive, with the absence of stage 1 parturition and with stage 2 and 3 occurring simultaneously. Premature placental separation was common, with the appearance of the engorged chorioallantoic membrane, or ‘red bag presentation’, indicating that the allantochorion was passed with the fetus. Due to abnormal positioning of the fetus, dystocia was frequent, with the mare reluctant to foal in the recumbent position. The majority of mares lacked mammary development If delivered alive, neonates were weak, dehydrated and dyspneic, requiring immediate intensive veterinary assistance. Clinical signs were associated with neonatal asphyxia and consistent with systemic inflammatory response syndrome (SIRS). Clinical pathology revealed dehydration, neutropenia, hypoglycemia, and inadequate colostral transfer. The placenta was heavy and edematous, weighing up to
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Figure 248.4 Unilateral endophthalmitis. Reproduced courtesy of Dr Claire Latimer, Rood & Riddle Equine Hospital. (See color plate 121).
Figure 248.3 Draining fluid from the chest cavity of a case of pericarditis. Reproduced courtesy of Dr Stuart E. Brown II, Hagyard Equine Medical Institute.
18 kg (40 lb), with hemorrhages on the surface of the umbilical cord.4 During April and May of 2001 and 2002 an increase in the number of cases of fibrinous pericarditis was also reported among the equine population of central Kentucky.5 The presenting signs included fever, azotemia, colic, weight loss, anorexia, lethargy, tachypnea, and tachycardia. Echocardiograms indicated an effusive pericarditis with evidence of right-sided heart failure with pleural effusion and/or ascites. Fluid evacuated from the pericardial space ranged from 1 to 22 liters (Figure 248.3). In two-thirds of the cases examination of the fluid indicated that it was sterile, with a variety of bacteria isolated from the remaining cases. The white cell count ranged from 900 to 28 000 cells/cmm and total protein from 3.3 to 5.4 g/dL. Approximately 50% of cases died or were euthanized and submitted to the LDDC for postmortem examination. Also during the same period an increased incidence of an acute-onset unilateral, exudative endophthalmitis (Figure 248.4) was observed.6 Cases exhibited corneal edema and exudates in the anterior
and posterior segments with variable degrees of pain, periocular swelling, discharge, and hyphema. Culture and cytology of aqueous and vitreous samples were unproductive. The overall number of cases due to the various manifestations of MRLS occurring during 2001 and 2002 is shown in Table 248.1. Figures derived from the Fact Book 20047 comparing numbers of registered Thoroughbred foals born in Kentucky during 2001 and 2002 indicate a 17% fall from 9837 to 8166. This drop was attributable primarily to EFL.
Pathological findings Lesions associated with LFL have been described by Williams et al.8 They included inflammation of the umbilical cord (funisitis) and amnion (amnionitis), pneumonia, fetal bacteremia, and sometimes placentitis. Findings were suggestive of in utero fetal illness and distress but lacked specificity to permit diagnosis of the cause. Fetuses were of normal size and weight for gestational age and in a good state of postmortem preservation. The most striking changes were observed on the umbilical cord especially the amniotic segment. They included roughening of the surface, a gray-yellow discoloration, enlargement due to stromal edema and surface hemorrhages (Figure 248.5). Histopathological findings revealed the lungs of LFL cases to contain desquamated cells of amnionic fluid origin and small numbers of neutrophils and macrophages in the alveolar spaces. Bacteria
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Table 248.1 Cases of mare reproductive loss syndrome, 2001–02 Year
Early fetal loss (EFL)
Late fetal loss (LFL)
Uveitis
Pericarditis
2001 2002
2000–3000 Approx. 500
550 165
Approx. 30 6
50–60 9
were often present in the alveoli. Bacteria were observed on the surface of umbilical cords with loss of epithelium and light to heavy infiltrates of neutrophils and macrophages on the surface. Bacterial cultures from umbilical cords, fetal lungs, and placentas yielded alpha-hemolytic Streptococcus spp. and/or Actinobacillus spp. in approximately 80% of the cases.9 They represent two groups of bacteria not normally considered important causes of equine abortions. Actinobacillus spp. isolates recovered from fetuses and pericardial fluid samples were identical to Actinobacillus spp. found in the oral cavity and alimentary tracts of healthy horses.10 The macroscopic and microscopic findings associated with cases of pericarditis have been reported by Bolin et al.11 The most obvious lesion was a layer of fibrin covering the epicardium in the form of villonodular deposits with gross thickening of the pericardium (Figure 248.6). Bacteria isolated from pericardial fluid and/or the myocardium included Actinobacillus and Streptococcus spp.
an abrupt onset and dramatic increase in number of reported cases of both EFL and LFL with the possible addition of cases of pericarditis and unilateral endophthalmitis in the population at risk. Cases identified during the spring involving horses grazing pastures containing or in close proximity to trees of the family Roseaceae, particularly black cherry containing ‘tents’ of the ETC, are an important consideration. The differential diagnosis includes fescue toxicity, abortion due to a variety of infectious agents including equine herpesvirus and equine arteritis virus, chemical and plant toxins, and mycotoxins.
Treatment
Presenting features of MRLS include EFL at 35 days and beyond, LFL, birth of weak foals with acute respiratory distress, unilateral endophthalmitis, and pericarditis, although not all may be observed during an outbreak. The diagnosis of MRLS is dependent on an appreciation of epidemiological parameters including
Little if any success was achieved with treating mares showing early signs of EFL based on ultrasonographic examination showing the presence of echogenic fluid and a viable fetal heartbeat. The prognosis for survival of neonatal foals exposed to MRLS was dependent on improved blood values and developing a normal respiratory pattern.4 Treatment was empirical utilizing antibiotics, plasma, fluid therapy, enteral and/or parenteral nutrition, vitamin E, dimethyl sulfoxide (DMSO) or mannitol, anti-inflammatory medication, oxygen supplementation, and intensive nursing care. Treatment of cases of pericarditis involved pericardiocentesis, often on more than one occasion. Supportive treatment included intrapericardial and systemic administration of corticosteroids and
Figure 248.5 Funisitis of amniotic segment of umbilical cord from aborted fetus. Reproduced courtesy of Dr Neil Williams, Livestock Disease Diagnostic Center, University of Kentucky.
Figure 248.6 Pericarditis with nodules of fibrin covering the epicardium. Reproduced courtesy of Dr David Bolin, Livestock Disease Diagnostic Center, University of Kentucky.
Diagnosis
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antibiotics.5 Cases of endophthalmitis did not respond to treatment.6 In 2001 very few mares that lost their fetuses and were bred back the same season became pregnant. The primary reason was their inability to cycle normally due to the presence of eCG (equine chorionic gonadotropin) in the blood circulation resulting from persistence of endometrial cups produced before the pregnancy was lost. These mares experienced no problems in becoming pregnant when they were bred in 2002.
Epidemiology Preliminary observations during the spring and summer of 2001 confirmed that mare reproductive losses occurred simultaneously on many farms over a widespread area of central Kentucky, commencing during the latter part of April and peaking during the early part of May. This observation was more indicative of an environmental toxin as distinct from an infectious agent. The period coincided with a massive invasion of ETC that completely denuded the vast numbers of wild black cherry trees that had grown up on pastures and fence lines on the majority of horse farms. On falling to the ground caterpillars created a dense mat on pastures, roadways, fences, and buildings, filling equipment such as water troughs and feed buckets. An extensive epidemiological study undertaken in the summer of 2001 on 133 farms in central Kentucky identified high to medium levels of ETC and wild cherry trees as strongly associated with a high incidence of EFL in and around impacted pastures.12 Case–control studies examining specific clinical aspects including EFL,13 LFL,14 and pericarditis15 identified environmental factors as predisposing to MRLS, including the presence of ETC. The biology of the ETC as it relates to MRLS has been described by Fitzgerald.16 ETC is largely restricted to populating trees of the family Roseaceae, preferring the black cherry throughout the eastern half of the USA. Overwintered eggs hatch in the spring just as buds break and leaves begin to unfold, around mid-March in central Kentucky. The caterpillars gather at a branch fork and build a communal silk shelter (tent) and feed on surrounding young leaves. Whole trees become festooned with silken tents and are completely denuded of leaves. When the larvae are fully grown, usually by mid-May, they abandon their host tree, and fall to the ground in search of pupation sites, often traveling long distances. On finding a suitable site, the caterpillars spin cocoons, spending 2–3 weeks in the pupal phase before emerging as moths between June and early July, whereupon they mate and lay large numbers of eggs.
Figure 248.7 Caterpillar ‘tent’ on branch of cherry tree. Reproduced courtesy of Mr Henry Q. Murphy.
The ETC has irruptive population dynamics characterized by periodic region-wide outbreaks followed by abrupt declines that are related to diseases, natural enemies, and environmental factors including temperature. Inclement weather causes a high mortality as the ETC is unable to digest food below 15◦ C. During late April to early May in central Kentucky ETC were at the peak of their population cycle and warm temperatures were conducive to widespread dispersal (Figures 248.7 to 248.10). Epidemiological studies revealed a strong temporal and spatial association between MRLS and the presence of ETC. Other studies to identify possible causes, including mycotoxins, fescue toxicosis, several plant toxins, cyanide, and infectious disease agents, produced negative results. A summary of these findings may be found in the Proceedings of the First Workshop on MRLS1 (http://www.ca.uky. edu/agc/pubs/sr/sr2003–1/sr2003–1.htm). Consequently several experimental studies were undertaken to determine the role of ETC. Three challenge studies undertaken by Webb et al.17 provided evidence that ingestion of ETC induces MRLS-type
Figure 248.8 Buckets filled with caterpillars, spring 2001. Reproduced courtesy of Mr Henry Q. Murphy.
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Figure 248.9 Eastern tent caterpillar on leaf of cherry tree. Reproduced courtesy of Mr Henry Q. Murphy.
abortions in mares; these were supported by a fourth study undertaken by Bernard et al.18 The studies provided evidence that abortigenic activity was associated with the exoskeleton or cuticle of the caterpillar, and that this abortigenic activity was inactivated by autoclaving and not easily removed by extraction with aqueous or organic solutions. A study in pigs19 demonstrated that ingested ETC caused abortion in a non-equid species and revealed that the caterpillars’ setae (hairs) became embedded in the deep submucosa of the pig’s alimentary tract. Similar lesions were observed in mares fed ETC. Whilst these studies provide evidence of source they do not provide clarification of the pathogenesis of MRLS. Tobin et al.20 proposed the ‘septic pene-
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trating setal hypothesis’ whereby as a result of ingestion, intestinal penetration by barbed setal fragments of ETC with associated bacterial ‘hitchhikers’ is followed by hematogenous distribution of septic materials to susceptible tissue sites giving rise to the various manifestations of MRLS. No species of caterpillar is presently known to serve as a vector of a vertebrate pathogen or to induce abortion in mammals. The bacterium Serratia marcesens has been isolated from the gut of caterpillars and is recognized as a cause of disease in horses;16 it was isolated from fetuses in the experimental challenge studies but not from field cases. The role of bacteria as primary or secondary pathogens in the etiology of MRLS remains unresolved. The abnormal weather pattern through April 2001 in central Kentucky, comprising low rainfall and high temperatures, punctuated by severe frosts in mid-April, stimulated an investigation as to the possible role of weather acting as a trigger factor for MRLS.21 Examination of weather patterns from previous years identified a similar pattern in 1981. During 1980 and 1981 EFL was identified on a number of farms in central Kentucky in a report prepared by Dr J.T. Bryans (unpublished, 1981). Whilst the etiology was not identified and there was no information on the presence or absence of ETC, there is a report of the isolation of alpha-hemolytic Streptococcus and Actinobacillus spp. (P.J. Timoney, personal communication) from EFL. These observations provide circumstantial evidence that MRLS had occurred in central Kentucky prior to 2001. In 2004 a series of abortions occurred among pregnant mares on farms in New South Wales. The term ‘equine amnionitis and fetal loss’ (EAFL) was coined to describe the condition, which has been compared to MRLS.22 Whilst the pathology of fetal pneumonia, placentitis, and funisitis is similar to MRLS, the two conditions differ in other aspects. A possible etiology of EAFL has been suggested resulting from exposure to processionary caterpillars (Ochragaster lunifer).23
Prevention
Figure 248.10 Egg mass of overwintering eastern tent caterpillar. Reproduced courtesy of Dr Lee H. Townsend, Department of Entomology, University of Kentucky.
The evidence linking MRLS to exposure of pregnant mares to ETC led to studies to manage ETC infestations.24 Investigators monitored the emergence from egg masses to guide timing of control actions and evaluated reduced-risk insecticides and treatment strategies including foliar sprays, trunk injections, winter egg mass treatments, and barrier sprays to intercept larvae entering pastures. Spraying black cherry trees is time-consuming and costly. Many farms share tree lines with residential areas or with other farms so spray drift and liability are a concern. Few insecticides in the USA are registered for use on
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pastures, and others that are effective against treefeeding caterpillars have grazing restrictions on their label. Winter treatment of egg masses with two pyrethroid insecticides, bifenthrin (OnyxTM ; FMC Corp., Philadelphia, PA) or permethrin (AstroTM , FMC Corp., Philadelphia, PA), prevented emergence. However, egg masses are difficult to identify from the ground so treating whole trees in winter is less efficient than targeting nests with young larvae in early spring. Treatment may be justified in years when ETC populations are at a level commensurate with a high risk of MRLS. Bifenthrin and spinosad (SpinTorTM ; Dow AgroSciences, Indianapolis, IN), a metabolite of the actinomycete bacterium Saccharopolyspora spinosa Mertz & Yao, were effective as foliage sprays, and their fieldweathered residues remained active for at least 7 days. Bacillus thuringiensis Berliner var. kurstaki (Btk) (DiPel DFTM ; Valent, Richardson, TX), a microbial insecticide active against many caterpillar species, was less active with shorter residual activity compared to bifenthrin and spinosad. Bifenthrin proved to be the fastest-acting, most effective, longest-lasting foliage insecticide tested. Applied 2–3 weeks after the first eggs hatch, it provided tree-wide control with a single application. It is labeled in the USA as TalstarOne (FMC, Philadelphia, PA), which lists ETC on its label. Microinjection of trees is well suited to use on horse farms as it eliminates spray drift and can be undertaken in most weather conditions. Dicrotophos (Bidrin) formulated in 2-mL capsules (Inject-A-Cide B) using the Mauget MicroInjection System (Mauget, Arcadia, CA), is labeled for systemic control of ETC. Dicrotophos trunk injections controlled ETC from shortly before egg hatch until late April when caterpillars were still feeding in trees. It is a Restricted Use Pesticide in the USA due to high acute mammalian toxicity but microinjection allows certified applicators to use it with relatively low hazard risk. The effectiveness of barrier treatments around trees, paddocks, and fence areas infested with ETC was also evaluated. Application of malathion and carbaryl proved ineffective but permethrin (AstroTM ; FMC, Philadelphia, PA) was effective in providing residual control over a 7-day period. Whilst perimeter treatment may be warranted it is recommended that the main defense should target larvae before they abandon their host trees. There remain several important unanswered questions with respect to MRLS, including the pathogenesis of the various clinical manifestations and its precise etiology. In the interim, strict attention by management on horse farms in central Kentucky to prevent contact between pregnant mares and ETC
has resulted in a significant reduction in the incidence of MRLS since 2002. To what extent this has been influenced by reduction in ETC through natural changes in population density, extensive elimination of cherry trees, and pesticide use on horse farms remains unanswered. The possibility that factors yet to be identified contribute to MRLS remains to be considered.
References 1. Powell DG, Troppman A, Tobin T. (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003 (http://www.ca.uky.edu/agc/pubs/sr/ sr2003–1/sr2003–1.htm) 2. Sebastian M, Giles R, Donahue J, Sells S, Tramontin R, Poonacha KB, Roberts J, Harrison L, Seahorn T, Sprayberry K, Bernard W. Encephalitis due to Actinobacillus species in three adult horses [abstract]. Vet Pathol 2002;35:630. 3. Thalheimer R, Lawrence RG. The economic loss to the Kentucky breeding industry from Mare Reproductive Loss Syndrome (MRLS) of 2001. URL: http://cobweb2.louisville. edu/eip/Newsletters/research/MRLS.asp 4. Byars TD, Seahorn TL. Clinical observations of Mare Reproductive Loss Syndrome in critical care mares and foals. In: Powell D, Troppman A, Tobin T (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003; pp. 15–6. 5. Slovis, N. Clinical observations of the pericarditis syndrome. In: Powell D, Troppman A, Tobin T (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003; pp. 18–20. 6. Latimer, C. Endophthalmitis: a syndrome associated with Mare Reproductive Loss Syndrome? In: Powell D, Troppman A, Tobin T (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003; pp. 17–18. 7. Jockey Club. Fact Book 2004: a Guide to the Thoroughbred Industry in North America. Lexington, KY: Jockey Club, 2005. 8. Williams NM, Bolin DC, Donahue JM, Giles RC, Harrison LR, Hong CB, Poonacha KB, Roberts JF, Sebastian MM, Smith BJ, Smith RA, Swerczek TW, Tramontin RR, Vickers ML. Gross and histopathological correlates of Mare Reproductive Loss Syndrome. In: Powell D, Troppman A, Tobin T (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003; pp. 24–5. 9. Donahue JM, Sells SF, Harrison LR, Hong CB, Poonacha KB, Roberts JF, Sebastian M, Smith R, Swerczek T, Tramontin R, Vickers ML, Williams N. Bacteria associated with Mare Reproductive Loss Syndrome: late fetal losses. In: Powell D, Troppman A, Tobin T (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003; pp. 27–9. 10. Donahue JM, Sells SF, Bolin DC. Classification of Actinobacillus spp. isolates from horses involved in mare reproductive loss syndrome. Am J Vet Res 2006;67:1426–32. 11. Bolin DC, Donahue JM, Vickers ML, Harrison L, Sells S, Giles RC, Hong CB, Poonacha KB, Roberts J, Sebastian MM,
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Swerczek TW, Tramontin R, Williams NM. Microbiologic and pathologic findings in an epidemic of equine pericarditis. J Vet Diagn Invest 2005;17:38–44. Dwyer RM, Garber LP, Traub Dargatz JL, Meade BJ, Powell D, Pavlick MP, Kane AJ. Case-control study of factors associated with excessive proportions of early fetal losses associated with mare reproductive loss syndrome in central Kentucky during 2001. J Am Vet Med Assoc 2003;222:613–19. Cohen ND, Donahue JG, Carey VJ, Seahorn JL, Piercy D, Donahoe JK, Williams DM, Brown SE, Riddle TW. Casecontrol study of early-term abortions (early fetal losses) associated with mare reproductive loss syndrome in central Kentucky. J Am Vet Med Assoc 2003;222:210–17. Cohen ND, Carey VJ, Donahue JG, Seahorn JL, Donahoe JK, Williams DM, Harrison LR. Case-control study of late-term abortions associated with mare reproductive loss syndrome in central Kentucky. J Am Vet Med Assoc 2003;222:199–209. Seahorn JL, Slovis NM, Reimer JH, Carey VJ, Dohanue JG, Cohen ND. Case-control study of factors associated with fibrinous pericarditis among horses in central Kentucky during Spring 2001. J Am Vet Med Assoc 2003;223:832–8. Fitzgerald T. The biology of the tent caterpillar as it relates to Mare Reproductive Loss Syndrome. In: Powell D, Troppman A, Tobin T (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003; pp. 84–7. Webb BA, Barney WE, Dahlman DL, DeBorde SN, Weer C, Williams NM, Donahue JM, McDowell KJ. Eastern tent caterpillars (Malacosoma americanum) cause mare reproductive loss syndrome. J Insect Physiol 2004;50:185–93. Bernard WV, LeBlanc MM, Webb BA, Stromberg AJ. Evaluation of early fetal loss induced by gavage with eastern
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tent caterpillars in pregnant mares. J Am Vet Med Assoc 2004;225:717–21. McDowell KJ, Williams NM, Donahue JM, Poole L, Barney WE, Coe B, DeBorde SN, Ennis L, Newman KE, Linemann M, Lynn B, Webb BA. Deductive investigations of the role of eastern tent caterpillars in Mare Reproductive Loss Syndrome. In: Powell DG, Furry D, Hale G (eds) Proceedings of a Workshop on the Equine Placenta. Lexington, KY: College of Agriculture, University of Kentucky, 2004; pp. 99–103. Tobin T, Harkins JD, Roberts JF, VanMeter PW, Fuller TA. The Mare Reproductive Loss Syndrome and the Eastern Tent Caterpillar II: a toxicokinetic/clinical evaluation and a proposed pathogenesis: septic penetrating setae. Int J Appl Res Vet Med 2004;2:142–58. Priddy T, Pieper C, Wang W. Climatic correlations of the 2001 and 2002 episodes of mare reproductive loss syndrome. In: Powell D, Troppman A, Tobin T (eds) Proceedings of the First Workshop on Mare Reproductive Loss Syndrome. Lexington, KY: College of Agriculture, University of Kentucky, 2003; pp. 37–40. Todhunter KH, Perkins NR, Wylie RM, Chicken C, Blishen AJ, Racklyeft DJ, Muscatello G, Wilson MC, Adams PL, Gilkerson JR, Bryden WL, Begg AP. Equine amnionitis and fetal loss: The case definition for an unrecognized cause of abortion in mares. Aust Vet J 2009;87:35–8. Cawdell-Smith AJ, Todhunter KH, Perkins NR, Bryden WL. Processionary caterpillars are an abortifacient in mares. Proc Aust Soc Anim Prod 2008;27:75. Potter DA, Foss L, Baumler RE, Held DW. Managing Eastern tent caterpillars Malacosoma americanum (F) on horse farms to reduce risk of mare reproductive loss syndrome. Pest Manag Sci 2005;61:3–15.
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Chapter 249
Fescue Toxicosis Dee L. Cross
Tall fescue (Lolium arundianceum) infected with the endophytic fungus Neotyphodium coenophialum (E+) negatively affects herbivores that consume it. Horses (Equus caballus) are more sensitive to the alkaloids in E+ tall fescue than cattle (Bos spp.). During the Middle Ages, there were numerous incidences of human toxicosis involving hallucinations, convulsions, delusions, and gangrene. The syndrome became known as St Anthony’s fire. In some instances, people actually jumped from buildings during hallucinogenic states. Barger (1931)1 determined that ergot alkaloids from the Claviceps purpurea fungus were the causative agent. Ergot alkaloids are characterized by an indole group which is a component of a tetracyclic ring known as an ergoline ring.2 Most of the sclerotia, containing the ergot bodies, can be removed during the grain cleaning process. However, small amounts have been detected in a variety of grains destined for human consumption.3
Interaction of ergot alkaloids with receptors Ergot alkaloids exhibit a high affinity for αadrenoreceptors, D2 dopamine receptors, and several subtypes of 5-HT receptors.4 The various alkaloids may act as receptor agonists or antagonists, depending on the receptor and the particular alkaloid. Ergovaline was shown to activate 5-HT2A receptors and thus induce contractions of blood vessels.5 Thus, blood flow to the vascular beds was reduced. Further research by Schoning et al.6 confirmed the interaction of ergovaline with 5-HTIB/ID , and αadrenoreceptors. Many of the toxic effects of ergot alkaloids can be explained by changes in vascular blood flow as proposed by Cross.7 Strickland et al.8 reported that ergovaline was a D2 dopamine receptor agonist in rat pituitary cells. In this study, ergovaline
blocked the release of prolactin by cultured pituitary cells. The D2 dopamine receptor antagonist, domperidone, prevented the prolactin-lowering effect of ergovaline. Therefore, the effects of ergot alkaloids in milk production and agalactia can probably be explained by their effect on D2 dopamine receptors. Domperidone’s effects on placental health, gestation length, dystocia, foal vigor, mare serum estrogen and progestogens, and subsequent mare fertility are apparently due to its 5-HT and α−adrenoreceptor antagonist activity.
Effect of ergot alkaloids on the immune system Immunologists generally agree that recent evidence suggests a balance between prolactin (PRL) and glucocorticoids is important in regulating the immune response. Numerous studies have reported a reduction in PRL levels when ergot alkaloids are consumed (reviewed by Cross et al.).9 Natural killer cell activity is suppressed by several dopamine receptor antagonists (huxthine, fluphenazine, pimocide, and haloperidol, in a PRL-dependent manner.10 They theorized interaction between ergot alkaloids and lymphoid cells and tumor cells based on indirect findings: (i) expression of dopamine, serotonin or α-adrenoreceptors or lymphocytes that can be involved in the action of ergopeptines; (ii) inhibition of signaling pathways through intracellular enzymes (serine/threonine kinases, Ras-MAPK), leading to mitogenesis by activation of the adenylate cyclase system (D2 agonistic, PRL inhibitory eeroglines); (iii) interaction with the nuclear structures or direct binding with DNA, including derivatives of agroclavine, festuclavine, and several ergopeptines (LSD, ergosine, ergosinine, and dihydroergosine).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Dee L. Cross
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Toxicity of the ergot alkaloids of Neotyphodium coenophialum Since the release of ‘Kentucky-31’ tall fescue seed in 1942, it has been widely used by livestock producers and the turf grass industry. It is hardy in a wide range of soil types, resistant against heavy traffic and overgrazing, and relatively drought and pest resistant when compared to other pasture grasses.11 After its release, it was observed that cattle consuming the grass did not perform as well as the nutritional analysis of the forage indicated that they should have performed. Instead, tall fescue began to gain a reputation for causing reproductive problems and poor pre- and post-weaning calf performance. Cattle consuming tall fescue during hot summer months appeared to be more stressed than normal and frequently stood in water or created mud wallows. Occasionally, lameness was detected and in extreme cases a foot would become detached, especially after cold winter weather. Horse owners also began to notice agalactia, prolonged gestations, foaling difficulties, and sometimes mare and foal deaths due to dystocia in pregnant mares grazing tall fescue pastures.12 Bacon et al.13 reported the first conclusive evidence of the presence of an endophytic fungus in tall fescue that was later identified as Neotyphodium coenophialum.14
Toxicosis in horses For many years, veterinarians and horse owners reported reproductive problems in mares that con125
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sumed tall fescue.12, 15, 16 Monroe et al.17 determined that the endophyte of tall fescue is the causative agent for reproductive abnormalities in gravid mares (Figure 249.1). Monroe reported that increased gestation lengths, agalactia, foal and mare mortality, tough and thickened placentas, weak and dysmature (largeframed emaciated looking) foals, and reduced serum prolactin and progestagen levels occurred in mares consuming E+ pasture, whereas gravid mares on endophyte-free (E–) pasture appeared normal. Gravid mares
Increased length of gestation Gestation length of mares increased 27 d when consuming E+ fescue, compared to mares consuming E– fescue.17 Putnam et al.,18 Earle et al.,19 and Redmond et al.20 observed similar results. Severe dystocia is a frequent observation of mares that try to foal after an extended gestation period. Supplementing E– and E+ mares on pastures with 50% of their National Research Council (NRC) requirements for energy, using whole shelled corn, provided no beneficial effects on length of gestation or dystocia. In the Earle et al.19 study, 66% of the mares grazing E+ with energy supplementation exhibited prolonged length of gestation and died due to dystocia, whereas the same signs and effects were noted for 50% of the mares grazing E+ without supplementation. Putnam et al.18 reported 10 of 11 mares grazing E+ fescue experienced obvious clinical dystocia, and four of them died during parturition, with only one foal surviving the natal period.
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Figure 249.1 Comparison of the effects of E+ and E– tall fescues on gestation length, foal mortality, agalactia, incidence of placental retention, and rebreeding response in mares. Stars indicate significant difference between treatments (P < 0.05). Adapted from Monroe et al., 1988.17
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The dystocia appears to be a result of inadequate preparation of the reproductive tract for foaling, prolonged gestation, and fetal malpresentation. Due to prolonged gestation, foals usually have larger than normal skeletal frames, increasing the difficulty of expelling the fetus through an unprepared tract.17, 18, 21 Additionally, foals are often rotated 90–180◦ from the normal position for delivery.17, 20, 22 The failure of events that initiate preparation for and cause normal parturition results in the subsequent catastrophic events of dystocia, as well as, in some instances, mare and foal mortality when mares graze E+ fescue.
Agalactia The effects of endophyte consumption on milk production vary among species. Cattle,23–26 sheep,27 and mice28, 29 have been shown to have reduced milk yields, whereas horses17 (Figure 249.2) and rabbits (Oryctolagus spp.)30 exhibit reduced milk yields or complete agalactia. The connection between tall fescue toxicosis and lactogenesis seems to be through the effects of the ergot alkaloids on lactogenic hormones. Cattle, sheep, and mice have both placental lactogen and prolactin.31 In contrast, horses and rabbits rely on prolactin to stimulate prepartum lactogenesis (milk synthesis).31 The depressive effects of the
ergot alkaloids on prolactin secretion may suppress the effect of prolactin on lactogenesis in cattle, sheep, and mice, but have little or no effect on placental lactogen (a lactogenic hormone). Consequently, the placental lactogen and the small level of pituitary prolactin, a hormone produced by the anterior pituitary, may be sufficient to initiate prepartum (prior to parturition) lactogenesis in these species and allow lactation to begin after parturition. In the horse, it seems that the reduced prolactin secretion from the pituitary lactotrophic cells results in agalactia. The alkaloids in the tall fescue/fungal endophyte symbiont are serving as D2 dopamine receptor agonists at the pituitary level.32 In addition, the horse, unlike ruminant herbivores, does not benefit from pre-gastric metabolism of alkaloids and would be subject to absorption of larger amounts of the alkaloids from E+ tall fescue.33 .’ Eighty-eight percent of mares were agalactic when maintained on E+ fescue up to foaling17 (Figure 249.1). The milk of agalactic mares often appears as a brown or straw-colored oily-looking fluid, rather than the white milk of normal mares. This fluid has little nutritional value, and foals invariably die unless bottle-fed. Another frequent complication affecting foal viability is the lack of normal immunoglobulins in foals from mares that have the straw-colored fluid rather than white milk.34
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Endophyte-Infected Control Figure 249.2 Effects of varying levels of domperidone (1.1, 1.65, or 2.2 mg/kg, p.o., s.i.d.) on mammary gland development in periparturient (near to parturition date) mares grazing E+ tall fescue pastures. Day –31 represents the average number of days prior to the calculated date of parturition when pretreatment samples were obtained (pretreatment samples obtained 1.2 d before initiation of drug treatment). Stars indicate first detectable difference (P < 0.05) from pretreatment values within treatment. Numbers indicate number of animals remaining in treatment group when sample was obtained (from Redmond, 1994).59
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Fescue Toxicosis Placentas The placentas of mares grazing E+ tall fescue are thickened, reddish colored, and heavier with an increased rate of retention than for E– mares.17 Frequently, the foal is presented normally but encased in a tough, thickened chorioallantois (a membrane surrounding the foal in the placenta) membrane, which it cannot break through; consequently, the foal suffocates unless an attendant is present to cut the chorioallantois immediately. Using an Ingstrom meter to measure stress and strain, these E+ placentas appeared to be more resistant to forces that would tear them, partially explaining why some foals are unable to break through the thickened placentas.17 Taylor et al.22 reported heavier and thicker placentas from many mares consuming E+ seed than from mares consuming E– seed; DNA, RNA, and collagen content were greater in the placentas of mares consuming E+ seed. Boosinger et al.35 reported on mares that grazed E+ fescue either continuously, from 300 d of gestation to foaling, or from gestation days 60 to 300, or had no exposure to fescue. They observed an increase in weight and width of combined chorioallantois from mares exposed to E+ fescue continuously or from day 300 to foaling. The same authors reported increased placental thickness in E+ mares immediately before parturition.35 Using 12 E+ and 12 E– mares, increased placental thickness was observed in 10 of the E+ mares a mean of 6.5 h prior to the onset of parturition, while another E+ mare demonstrated an increase in placental thickness 32 h prior to parturition. An elective cesarean section was performed in one E+ mare at 358 d of gestation and within 2 h of noting an increase in placental thickening. At surgery, the placenta was reported to be thickened in a plaque-like fashion in the ventral portion of the gravid horn (the uterine horn containing the fetus). The thickened portion of the chorioallantois was noted to be separated from the uterus. Placental separations are common during the last trimester in mares grazing E+ fescue and are commonly referred to as red-bag. Frequently these mares develop udders prematurely and may leak milk. This is the only visual sign that a possible placental separation is eminent. Clinicians have reported that, in some cases, where the udder develops early also, there is some placental separation, visible by ultrasound, just caudal to the cervical star. In some case where placental separation occurs according to the preceding indicators, domperidone is administered twice daily at the 1.1 mg/kg bodyweight dose. This is continued for approximately 2 weeks. Many times the udder will return to normal and leakage of milk cease. About 2 weeks before foaling, domperidone
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dosing is reinstated at the 1.1 mg/kg level until foaling. In human medicine, physicians frequently use ergot alkaloids to stop post-parturition uterine bleeding. The ergot alkaloids reduce blood flow by activating α-adrenoreceptors in the uterus (αadrenoreceptor agonists). Apparently, during late gestation the uterus is suffering from reduced blood flow in mares grazing E+ fescue. This is similar to the effects that ergot alkaloids have on increasing body temperature in cattle during warm weather (summer syndrome). In summer syndrome, the ergot alkaloids activate the peripheral α-adrenoreceptors and shunt blood away from the peripheral tissues, where cooling can occur, and into body core, thus raising body temperature. In the gravid mare, domperidone blocks the α-adrenoreceptors (α-adrenoreceptor antagonists) and allows near normal blood flow to return, thus preventing the cascade of events resulting in a premature placental separation. The reason for udder development and leaking of milk that occurs in this malady is not known; however, the uterus may be signaling the udder that parturition is imminent. Therefore, it is important that the attending physician begin ultrasound of the uterus should udder development and/or leaking of milk start in any mare that has been exposed to E+ pasture or hay during late gestation.
Foal vigor and viability Monroe et al.17 observed large-framed, dysmature and emaciated looking (poor muscle mass) foals with overgrown hooves from mares grazing E+ fescue, when those foals survived the birthing process, and which had an average gestation length 27 d past the expected foaling date (Figure 249.1). These foals appeared weak and many times exhibited a ‘dummy-like’ behavior. Later, with proper care, the foals appeared normal.17, 19 Septicemia (bacterial infection at or near birth) is a frequent problem and is likely a result of the low level of passive immunity. Putnam et al.18 reported that of 11 mares grazing E+ fescue, only three foals were alive at birth, and only one of the three survived the first month of life. In this study, dysmaturity or neonatal death of foals was not observed in 11 mares grazing E– pastures. Taylor et al.22 and Kosanke et al.36 observed lack of lung maturation in stillborn foals born to E+ mares. Amniotic fluid from E+ mares lacked pulmonary phospholipids and phosphatidylethanolamine was present in only 12% of those E+ mares.37 These data suggest that lack of lung maturation may be a contributing factor to the high rate of foal death observed from mares on E+ fescue. Boosinger et al.35 examined
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several organs and tissues from foals of E+ and E– mares. Histological studies of thyroid glands from foals exposed to E+ continuously or after gestation day 300 revealed numerous distended colloid-filled thyroid follicles lined by flattened cuboidal epithelial cells. Mean plasma T3 concentrations were reduced in these foals. Foals from mares exposed to E+ continuously or from day 300 to foaling demonstrated a response to thyroid-stimulating hormone (TSH) by showing improved mental alertness, desire to stand, and good suckle reflex. Brendemuehl38 collected colostrum from normal mares and tested it for IgG concentration. Foals from mares exposed to E+ fescue continuously or from gestation day 300 were administered 1 L of the pooled, collected colostrum collections by nasogastric tube within 1 h of birth. Compared to control foals not from E+ or E– fescue mares, these foals had decreased serum IgG concentrations. These data, as well as other data, suggest that foals from E+ mares receive less IgGs from the milk produced by the mares, and their absorption rate is lower even if the milk IgG levels are at or near normal levels. These factors, combined with the lower level of colostrum production in E+ mares, explain why many foals from E+ mares quickly become septic. The lower colostrum and nutrient intake from milk, especially IgG, probably accounts for many foal deaths from E+ mares which have live foals at birth. Brendemuehl et al.39 observed lower serum T3 , ACTH, cortisol, and total progestagen levels in foals from E+ mares compared to foals from E– mares.
Body temperature, blood flow, and laminitis In cattle and sheep, blood flow to the peripheral tissues decreased and body temperature increased when tall fescue seed was included in the diet.40 The reduction in blood flow to the peripheral tissues is probably related to increased body temperature because the animal is less efficient in cooling itself. Unlike cattle and sheep, pregnant mares exhibit no increase in body temperature when exposed to the endophytic toxins.17, 18 However, horses sweat more freely than cattle and are more capable of cooling themselves. Putnam et al.18 observed increased sweating in gravid mares grazing E+ tall fescue. In cattle, it seems that peripheral vasoconstriction (reduction in size of the blood vessel) caused by the alkaloids of E+ tall fescue results in ‘fescue foot’.41 Rohrback et al.42 reviewed data from 185 781 horses of which 5536 had a diagnosis of laminitis. Although these data were preliminary, they concluded that there appeared to be a relationship between laminitis in horses and consumption of E+ tall fescue.
Abney et al.43 observed a vasoconstrictive effect of ergot alkaloids on equine vessels in vitro. Carbohydrate overload and aqueous extract of black walnut44 are associated with the development of laminitis in horses. The aqueous extract of black walnut caused post-capillary venoconstriction (reduction in size of the vein), increased capillary hydrostatic pressure, and transvascular fluid movement (movement through the capillary wall), resulting in increased tissue pressure, edema, vascular collapse, and ischemia (reduced blood perfusion) in the equine digit.45 It is possible that through interaction of the ergot alkaloids with the adrenergic receptors of the sympathetic nervous system, similar responses may be occurring in horses consuming E+ fescue. Direct evidence for this theory does not exist.
Mare abortions and fertility Abortions in mares occur after rapid separation of the placenta from the endometrium, a layer of the uterus. This can occur any time during the last 6 months of gestation but is most often seen during the last 45 days and most frequently during the last 2 weeks. Of 1211 abortion/stillbirths presented to a diagnostic laboratory in Kentucky, placentitis and dystocia were the commonly diagnosed causes (about 19.5%, each), with congenital abnormalities (8%), twins (6%), umbilical cord torsion, and premature placental edema (4%, each) and equine herpesvirus and other bacterial infections (3%, each) being the other diagnostic causes.46 Pregnancy rates can be negatively affected by E+ fescue. Brendemuehl et al.47 observed the effects of E+ fescue on mare cyclicity, pregnancy rates, and embryonic death rates. Mares grazing E+ pastures had prolonged luteal functions, decreased per cycle pregnancy rates, and increased early embryonic death rates, when compared to those grazing E– pastures. Growing horses
Effects on growth rate and digestibility Consumption of E+ tall fescue, or treatment with tall fescue extract, causes a reduction in rate of gain and feed intake in cattle,48–50 rats (Rattus spp.),51 and rabbits.30 No reduction in growth rate was observed in yearling horses when corn-based concentrates were used to supplement E+ or E– hay.52 Pendergraft and Arns53 observed similar gains in yearling horses consuming E+ or E– hay with concentrate supplementation to meet NRC requirements for growth. However, average daily gains were reduced by 57% (0.24 and 0.56 kg for high- and low-endophyte treatments, respectively) in yearling horses grazing E+ pasture without supplementation, with a similar reduction in gain for steers in the same study.54
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Fescue Toxicosis Redmond et al.20 and McCann et al.52 observed lower intake and digestibility for E+ hay fed to mature geldings and yearling horses, respectively. McCann et al.55 and Pendergraft and Arns53 found no differences in digestibility due to the presence of the endophyte in hay when yearling horses were fed concentrate with the hay. Concentrate supplementation was used in both studies to meet NRC requirements for growth for yearling horses. These results suggest that the effects of endophyte consumption on digestibility and growth rate may be lessened by the inclusion of concentrates in the diet. In contrast, energy supplementation has no beneficial effects for alleviating lactation and reproductive problems in gravid mares that graze E+ tall fescue pastures.19 Performance horses Vivrette et al.56 evaluated the effect of E+ fescue on cardiorespiratory and thermoregulatory aspects of performance horses. Fourteen Quarter Horses grazed either E– or E+ tall fescue pastures. Subsequently, the horses were exercised at 4 m/s (9 mile/h), a brisk trot, for 0.5 h and a distance of 7.2 km (4.5 mile) over rolling terrain. Respiratory rates were higher in horses grazing E+ pasture during the 3.5 h postexercise period. In horses that had grazed E+, heart rates and skin temperatures remained above preexercise levels for 2.5 and 3.0 h, respectively. In E– horses, heart rates and skin temperatures of horses from E– pastures returned to pre-exercise level 1.5 h after exercise. Water consumption following exercise for E– and E+ horses was 21.2 and 35.9 L (5.6 and 9.5 gal), respectively. These data suggest a negative effect of E+ fescue on performance when horses have been grazing E+ pasture immediately prior to exercise.
Management and treatment of N. coenophialum toxicosis Pasture management Personal interviews of horse owners and veterinarians and clinical research have revealed that many horses exhibit most of the signs of tall fescue toxicosis while consuming even small quantities in hay, small patches of E+ fescue in paddocks, or even by grazing a small quantity of E+ fescue under paddock fences.57 Therefore, to prevent toxicosis in horses, E+ tall fescue must be completely eliminated in pastures. Personal experience and interviews with livestock owners throughout the United States attest to the extreme difficulty of eliminating E+ fescue from pastures. Experience has shown that, unless pastures are completely devoid of E+ plants and viable seed,
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the E+ plants begin to thrive and become significant problems within 1–3 y after replanting pastures. Best success with pasture reseeding has come through the use of chemical destruction of the fescue or an intense fallow followed by aggressive choke (crops that have a dense canopy) crops for 2 y before reseeding to another pasture forage. Establishment of clover or other forage mixes seems to be a reasonable alternative for cattle, but not for horses. Grazing behavior The horse is a notoriously selective grazer and will select many alternative forage species before consuming E+ tall fescue. Under low grazing pressures, many mares will spot graze other species of forage and never exhibit any signs of fescue toxicosis. Changes in grazing pressure or lack of availability of alternative forages can quickly force E+ fescue consumption and the classical signs of fescue toxicosis. This partially explains why some horse owners appear to have little or no fescue toxicosis when a few mares are grazing a large acreage of mostly E+ fescue, and other horse owners routinely have problems with mares grazing E+ fescue. Removal of mares from E+ pastures There is evidence to suggest that the effects of E+ fescue are greatly reduced if mares are withdrawn from E+ pastures at least 30–40 d prior to the date of expected foaling.35, 58 However, most veterinarians recommend removal of mares from E+ pastures from 30–40 d and up to 90 d prior to expected foaling. With this management approach, red-bag and the other signs of fescue toxicosis are greatly minimized. Therapeutic treatment
Dilution of toxin intake Gravid mares were fed 50% of the NRC requirement for energy as cracked corn for the last 90 d of gestation;19 this practice, while the mares are grazing an E+ fescue pasture had no effect on eliminating the negative results of consuming E+ fescue. Foal mortality was 66% and 100% for the energy and no energy supplement treatments, respectively. This study confirmed the severity of the problem under the typical grazing conditions found in the southeastern United States.
Dopamine antagonists (receptor blockers) Strickland et al.8, 32 studied the effects of ergot and loline alkaloids of E+ tall fescue on prolactin release by isolated and perfused (circulation of media
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Figure 249.3 Effect of E+ fescue and domperidone (1.1, 1.65, or 2.2 mg/kg, p.o., s.i.d.) treatment on serum prolactin levels in gravid mares. First detectable differences (P < 0.05) from pretreatment levels are indicated by stars. Unless otherwise indicated, data points represent four mares/treatment. Mares which were not showing signs of impending parturition 7 d after the calculated date of parturition, as determined by veterinary examination, were relocated to an E– pasture (from Redmond, 1994).59
through the cells) rat pituitary cells. The ergot alkaloids lowered prolactin concentrations and mimicked dopamine action. The use of a D2 dopamine receptor antagonist (domperidone) blocked the effect of the ergot alkaloids and prevented their prolactin lowering effect. Domperidone is a D2 dopamine receptor blocker that does not cross the blood brain barrier and elicit neuroleptic (central nervous system effects) side effects. Domperidone was administered orally (1.1 mg/kg bodyweight) to gravid mares grazing E+ tall fescue.20 It resulted in an increase in serum prolactin and progestagens (progesterone and its derivatives) and provided what seemed to be nearly complete recovery of gravid mares from tall fescue toxicosis with no observed neuroleptic side effects. Treated mares produced milk, had live and healthy foals, and their gestation length was similar to the calculated gestation length. Subsequently, a dose titration study (Figure 249.2–249.4) was conducted to determine the minimum effective dose of domperidone for treating tall fescue toxicosis.59 Again, domperidone provided recovery from tall fescue toxicosis in gravid mares and the minimum effective oral dose was 1.1 mg/kg bodyweight when administered daily for 30 d before foaling. In a field study,57 domperidone was administered to 1423 periparturient mares grazing tall fescue in several US states. Veterinarians and/or horse owners reported the drug to be 95% effective in preven-
tion of the signs of fescue toxicosis. Mares consuming the drug in a preventive mode did not experience increased gestation, dystocia, agalactia or lower than normal milk production, retained placentas, premature placental separations, dead, weak, or dysmature foals, as had been observed in non-treated control mares.
Mechanism of action of domperidone for treating equine fescue toxicosis The action of domperidone as a D2 dopamine receptor blocker prevents the ergot alkaloids from mimicking dopamine actions. The most apparent action of dopamine and the ergot alkaloids in the fescue/ endophyte symbiont is their prolactin lowering effect. With administration of domperidone to mares consuming E+ tall fescue, prolactin returned to normal levels and even increased above normal levels in most instances.20 Certainly prolactin is a major factor in equine fescue toxicosis; however, as demonstrated in the preceding review of endocrine effects of E+ fescue, prolactin is only one of many hormones which are influenced. Prolactin, progestagens, and estrogen are certainly major factors in the milk production maladies observed in E+ mares – and administration of domperidone returns these hormones to near normal levels and functions. In addition, the effect of domperidone
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on peripheral circulation as an alpha-1 receptor antagonist and 5-HT receptor antagonist provides for its remediation of the negative microcirculatory effects of the alkaloids. Since the hypothalamic–pituitary–adrenal (HPA) axis of the fetus in mares consuming E+ tall fescue appears to be compromised, resulting in prolonged gestation lengths and the associated problems, domperidone may be having some effect on the HPA system. This hypothesis is supported by the fact that mares that receive domperidone while grazing E+ tall fescue foal at or near their expected foaling date with normal, healthy foals. Adrenocorticotropic hormone (ACTH) levels in foals from mares consuming E+ fescue are low. Since ACTH is the stimulus for adrenal cortisol release, and since normal fetal adrenal cortisol levels appear to be necessary to trigger parturition, it can be speculated that domperidone may be affecting this system. Zerbe et al.60 administered domperidone to dogs (Canis familiaris) and observed an enhanced ACTH response to corticotropic releasing hormone (CRH) injections. Thus, domperidone could be reversing the effects of E+ tall fescue on gestation length by causing an increase in adrenal cortisol through CRH stimulated release of ACTH. Direct evidence to confirm this theory does not exist.
Acknowledgement The author wishes to express appreciation to Dr A.F. Parlow of the National Hormone and Pituitary Program, UCLA Medical Center, Torrance, CA, for supplying the antigen and antisera for the equine prolactin assays conducted at Clemson University, and to the many graduate student co-authors working with the author while a professor at Clemson University.
References 1. Barger G. Ergot and Ergotism. London: Garney & Jackson, 1931. 2. Lorenz K. Ergot on cereal grains. Crit Rev Food Sci Nutr 1979;11:311–50. 3. Scott PM, Lambaert GA, Pellaers P, Bacler S, Lappi J. Ergot alkaloids in grain foods sold in Canada. J AOAC Int 1992;75:773–9. 4. Pertz H, Eich E. Ergot alkaloids and their derivatives as ligands for serotonergic dopaminergic and adrenergic receptors. In: Kren V, Cvak L (eds) Ergot: The Genus Claviceps. Amsterdam: Harwood, 1999; pp. 411–40. 5. Dyer DC. Evidence that ergovaline acts on serotonin receptors. Life Sci 1993;53:223–8. 6. Schoning C, Flieger M, Pertz HH. Complex interaction of ergovaline with 5-HT2A. 5-HT1B/1D, and alpha-receptors in isolated arteries of rat and guinea pig. J Animal Sci 2001;79:2202–9.
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7. Cross DL. Fescue toxicosis in horses. In: Bacon CW, Hill NS (ed.) Neotyphodium/Grass Interactions. New York, London: Plenum Press, 1997; pp. 289–309. 8. Strickland JR, Cross DL, Birrenkott GP, Grimes LW. Effect of ergovaline, loline, and dopamine antagonists on rat pituitary cell prolactin release in vitro. Am J Vet Res 1994;55:716–21. 9. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: Signs and solutions. J Anim Sci 1995;73:899–908. 10. Fiserova A, Pospisil M. Role of ergot alkaloids in the immune system. In: Kren V, Clark L (eds) Ergot: The Genus Claviceps. Amsterdam: Harwood, 1999; pp. 451–67. 11. Siegel MR, Johnson MC, Varney DR, Nesmith WC, Buckner RC, Bush LP, Burrus PB II, Jones TA, Boling JA. A fungal endophyte in tall fescue: Incidence and dissemination. Phytopathology 1984;74:932–40. 12. Garrett LM, Heimann ED, Wilson LL, Pfander WH. Reproductive problems of pregnant mares grazing fescue pastures. J Anim Sci 1980;50:322 (Abstr). 13. Bacon CW, Porter JK, Robbins JD, Luttrell ES. Epichloe typhina from toxic tall fescue grasses. Appl Environ Microbiol 1977;34:576–81. 14. Glen AE, Bacon CW, Price R, Hanlin RT. Molecular phylogeny of Acremonium and its taxonomic implications. Mycologia 1996;88:369–83. 15. Villahoz MD, Moras EV, Barboni AM, Scharf V, Menchaca ES, de Guglielmone R. Reproductive problems of pregnant mares grazing fescue pastures in Argentina. 10th International Congress on Animal Reproduction and Artificial Insemination. 10–14 June 1984. Urbana-Champaign, IL; 1984; pp. 2:100–1. 16. Poppenga RH, Mostrum MS, Haschek WM, Lock TF, Buck WB, Beasley VR. Mare agalactia, placental thickening, and high foal mortality associated with the grazing of tall fescue: a case report. Proceedings of American Association of Veterinary Laboratory Diagnosis, 21 October 1984, Fort Worth, TX, 1984; pp. 325–6. 17. Monroe JL, Cross DL, Hudson LW, Henricks DM, Kennedy SW, Bridges WC Jr. Effect of selenium and endophytecontaminated fescue on performance and reproduction in mares. J Equine Vet Sci 1988;8:148–53. 18. Putnam MR, Fransby DL, Schumacher J, Boosinger TR, Bush L, Shelby RA, Vaughan JT, Ball D, Brendemuehl JP. Effects of the fungal endophyte Acremonium coenophialum in fescue on pregnant mares and foal viability. Am J Vet Res 1991;52:2071–4. 19. Earle WF, Cross DL, Hudson LW, Redmond LM, Kennedy SW. Effect of energy supplementation on gravid mares grazing endophyte-infected fescue. J Equine Vet Sci 1990;10:126–30. 20. Redmond LM, Cross DL, Strickland JR, Kennedy SW. Efficacy of domperidone and sulpiride for fescue toxicosis in horses. Am J Vet Res 1994;55:722–9. 21. Redmond LM, Cross DL, Jenkins TC, Kennedy SW. The effect of Acremonium coenophialum on intake and digestibility of tall fescue hay in horses. J Equine Vet Sci 1991;11: 215–19. 22. Taylor MC, Loch WE, Ellersieck M. Toxicity in pregnant pony mares grazing Kentucky-31 fescue pastures. Nutr Rep Int 1985;31:787–95. 23. Strahan SR, Hemken RW, Jackson JA Jr, Buckner RC, Bush LP, Siegel MR. Performance of lactating dairy cows fed tall fescue forage. J Dairy Sci 1987;70:1228–34. 24. Porter JK, Thompson FN Jr. Effects of fescue toxicosis on reproduction in livestock. J Anim Sci 1992;70:1594–603.
25. Schmidt SP, Osborn TG. Effects of endophyte-infected tall fescue on animal performance. Agric Ecosystems Environ 1993;44:233–62. 26. Irwin JW. The effect of a D2 dopamine receptor antagonist on fescue toxicosis in beef cattle. PhD dissertation. Clemson University, Clemson, SC, 1997. 27. Stidham WD, Brown CJ, Daniels LB, Piper EL, Featherstone HE. Toxic fescue linked to reduced milk output in ewes. Arkansas Farm Res 1982;31:9. 28. Zavos PM, Varney DR, Jackson JA Jr, Hemken RW, Siegel MR, Bush LP. Lactation in mice fed endophyte-infected tall fescue seed. Theriogenology 1988;30:865–75. 29. Siegel MR, Bush LP. Lactation in mice fed endophyteinfected tall fescue seed. Theriogenology 1988;30:865–87. 30. Daniels LB, Ahmed A, Nelson TS, Piper EL, Beasley JN. Physiological responses in pregnant white rabbits given a chemical extract of toxic tall fescue. Nutr Rep Int 1984;29:505–10. 31. Forsyth IA. Variation among species in the endocrine control of mammary growth and function: the roles of prolactin, growth hormone and placental lactogen. J Dairy Sci 1986;69:886–903. 32. Strickland JR, Cross DL, Jenkins TC, Petroski RJ, Powell RG. The effect of alkaloids and seed extracts of endophyte-infected tall fescue on prolactin secretion in an in vitro rat pituitary perfusion system. J Anim Sci 1992;70: 2779–86. 33. Wachenheim DE, Blythe LL, Craig AM. Characterization of rumen bacteria pyrrolizidine alkaloid biotransformation in ruminants of various species. Vet Hum Toxicol 1992;34:513–16. 34. Kouba JM. Effect of endophyte-infected tall fescues and pre- and postpartum domperidone treatment on gravid mares. MS Thesis. Clemson University, Clemson, SC, 1995. 35. Boosinger TR, Brendemuehl JP, Schumacher J, Bransky DI, Lee D, Shelly RA. Effects of short-term exposure to and removal from the fescue endophyte Acremonium coenophialum on pregnant mares and foal viability. Biol Reprod Mono 1995;1:61–7. 36. Kosanke JL, Loch WE, Worth K, Ellersieck MR. Effect of toxic tall fescue on plasma prolactin and progesterone in pregnant pony mares. Proceedings of Equine Nutrition and Physiological Society Symposium, 1989; p. 663. 37. Clare KA, Green EM, Strickland JR, Oliver JW, Andrews FM. Effect of endophyte infected tall fescue on equine fetal pulmonary maturity. Proceedings of American College of Veterinary Internal Medicine, 12th Veterinary Medical Forum, 1994. 38. Brendemuehl JP. Fescue toxicity in the broodmare – from conception to foaling. Proceedings of Auburn University Horse Course, Auburn, AL: 1995; 17 pp. 17. 39. Brendemuehl JP, Williams MA, Boosinger TR, Ruffin DG. Plasma progestogen, tri-iodothyronine and cortisol concentrations in postdate gestation foals exposed in utero to the fescue endophyte Acremonium coenophialum. Biol Reprod Mono 1995;1:53–9. 40. Rhodes NT, Paterson JA, Kerley MS, Garner HE, Laughlin MH. Reduced blood flow to peripheral and core body tissues in sheep and cattle induced by endophyte-infected tall fescue. J Anim Sci 1991;69:2033–26. 41. Solomons RN, Oliver JW, Linnabary RD. Reactivity of dorsal pedal vein of cattle to selected alkaloids associated with Acremonium coenophialum infected fescue grass. Am J Vet Res 1989;45:942.
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Fescue Toxicosis 42. Rohrbach BW, Green EM, Oliver JW, Schneider JF. Aggregate risk study of exposure to endophyte-infested (Acremonium coenophialum) tall fescue as a risk factor for laminitis in horses. Am J Vet Res 1995;56:22–6. 43. Abney LK, Oliver JW, Reinemeyer CR. Vasoconstrictive effects of tall fescue alkaloids on equine vasculature. J Equine Vet Sci 1993;13:334–40. 44. Galey FD, Beasley VR, Schaeffer D. Effect of an aqueous extract of black walnut (Juglans nigra) on isolated equine digital vessels. Am J Vet Res 1990;51:83–8. 45. Eaton SA, Allen D, Eades SC. Digital Starling forces aid hemodynamics during early laminitis induced by an aqueous extract of black walnut (Juglans nigra) in horses. Am J Vet Res 1995;56:1338–43. 46. Hong CB, Donahue JM, Giles RC Jr, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Equine abortion and stillbirth in Central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. 47. Brendemuehl JP, Boosinger TR, Pugh DG, Shelby RA. Influence of endophyte-infected tall fescue on cyclicity, pregnancy rate, and early embryonic loss in the mare. Theriogenology 1994;42:489. 48. Schmidt SP, Hoveland CS, Clark EM, Davis ND, Smith LA, Grimes HW, Holliman JL. Association of an endophytic fungus with fescue toxicity in steers fed Kentucky 31 tall fescue seed or hay. J Anim Sci 1982;55:1259– 63. 49. Hoveland CS, Schmidt SP, King, CC Jr, Odom JW, Clark EM, McGuire JA, Smith LA, Grimes HW, Holliman JL. Steer performance and association of Acremonium coenophialum fungal endophyte on tall fescue pasture. Agron J 1983;75:821–4. 50. Bond J, Bolt DJ. Growth plasma prolactin and ovarian activity in heifers grazing fungus-infected tall fescue. Nutr Rep Int 1986;34:93–102.
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51. Neal WD, Schmidt SP. Effects of feeding Kentucky 31 tall fescue seed infected with Acremonium coenophialum to laboratory rats. J Anim Sci 1985;61:603–11. 52. McCann JS, Heusner GL, Amos HE, Thompson, DL Jr. Growth rate, diet digestibility, and serum prolactin of yearling horses fed non-infected and infected tall fescue hay. J Equine Vet Sci 1992;12:240–3. 53. Pendergraft J, Arns MJ. Tall fescue utilization in exercised yearling horses. Proceedings of 13th Equine Nutrition and Physiological Symposium, 21–23 January, Gainesville, FL: 1993; p. 106. 54. Aiken GE, Bransby DI, McCall CA. Growth of yearling horses compared to steers on high- and low-endophyte infected tall fescue. J Equine Vet Sci 1993;13:26–8. 55. McCann JS, Heusner GL, Amos HE, Thompson DL Jr. Effects of 0 and 94% endophyte-infected tall fescue hay on growth rate, diet digestibility, and serum prolactin levels in yearling horses. Prof Anim Sci 1993;9:39–44. 56. Vivrette S, Stebbins ME, Martin O, Dooley K, Cross DL. Cardiorespiratory and thermoregulatory effects of endophyte-infected fescue in exercising horses. J Equine Vet Sci 2001;21:65–7. 57. Cross DL, Anas K, Bridges WC, Chappell JH. Clinical effects of domperidone on fescue toxicosis in pregnant mares. Proc Am Assoc Equine Pract 1999;45:203–6. 58. Taylor W. Effect of removal time on reproductive performance in pregnant mares grazing endophyte-infected fescue. Master’s Thesis, Clemson University, Clemson, SC, 1993. 59. Redmond LM. Efficacy of dopamine receptor antagonists as treatments for equine fescue toxicosis. PhD dissertation, Clemson University, Clemson, SC, 1994. 60. Zerbe CA, Clark TP, Sartin JL, Kemppainen RJ. Domperidone treatment enhances corticotrophin-releasing hormone stimulated adrenocortocotropic hormone release from dog pituitary. Neuroendocrinology 1993;57:282–8.
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Chapter 250
Prepubic and Abdominal Wall Rupture Dwayne H. Rodgerson
Body wall ruptures in prepartum mares occur uncommonly, but do present a unique challenge to practitioners (Figure 250.1). Case reports and small case series have been described to provide a better knowledge of the condition.1–5 Mares developing a change in the appearance of the ventral abdomen or ventral edema could be developing soft tissue tearing involving the abdominal musculature (Figure 250.2). The structures most commonly involved include the prepubic tendon or rectus abdominus muscle. The anatomy of the prepubic tendon is well described and complex in the horse.6 Body wall defects involving the external abdominal oblique, internal abdominal oblique, and transverse abdominal muscles can also be involved separately or in conjunction with tears in the prepubic tendon. An understanding of the predisposing factors and early clinical signs can potentially help veterinarians improve case outcome. The exact cause of abdominal wall herniation or prepubic tendon rupture is often unknown. Trauma to the abdominal wall in late gestation can result in abdominal wall herniation. Predisposing factors described include hydrops allantois, hydrops amnion, twins, and fetal giants.1, 3, 5 These factors all increase the amount of weight the ventral abdomen supports. Mares with abdominal wall defects will develop ventral edema that can extend from the mammary glands to the xiphoid. Mares may show signs of abdominal pain (looking at flank, sweating, and pawing), and this can vary based on the location and size of the defect. An elevation in heart rate and respiratory rate may be noted due to increased degree of pain and discomfort. Mares with prepubic tendon ruptures may be reluctant to move, show a sawhorse stance (elevated tail head and ischial tuberosity), and the mammary glands may be displaced cranially or flattened. In some cases, the secretions from the mam-
mary glands may be bloody. Palpation of the ventral abdomen often elicits a painful response. In many cases a large hematoma will develop subsequent to the muscle tearing. In some cases, abscess formation can also occur. Ultrasonography is used to assess the location and size of the body wall defect (Figure 250.3). The body wall defect is often bilateral with one side having a larger defect. Assessment of the fetal viability can also be performed. The technique also allows determining if a hydrops allantois or hydrops amnion is present. The condition of the placenta can be assessed and, in some cases, placental separation may be noted. The bowel wall close to the body wall defect should be assessed to determine if there is evidence of edema and possible strangulation. Adhesions within the abdominal defect have also been noted in some cases. Conservative management strategies that avoid induction of parturition or elective cesarean section and allow for natural parturition have recently been proposed as the treatment of choice in a small group of mares with body wall tears or prepubic tendon rupture.5 In acute cases of body wall ruptures, conservative management should be the treatment of choice initially. Treatment should be aimed at trying to prevent any further tearing to the abdominal musculature. Mares should be confined to a stall to restrict movement. Abdominal support wraps should be utilized to help take some of the stress off the abdominal musculature. The wraps should be well padded to prevent pressure sores from developing. Non-steroidal anti-inflammatory agents and analgesic therapy should be added to the treatment regimen to minimize inflammation and pain. Systemic antibiotics should also be considered to prevent abscess formation. The fetus should be continually
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 250.1 (a) A mare with a body wall defect on the right body wall just cranial to the right stifle. (b) The same mare with a body wall defect on the right body wall just cranial to the right stifle. This figure better demonstrates the location and size of the body wall defect.
assessed using fetal electrocardiogram monitoring to ensure the viability of the fetus. If the fetus is starting to show signs of stress or compromise then interventional management may be warranted. This would entail either induction of parturition or an elective cesarean section to deliver the foal. However, the gestational age should be taken into consideration in the decision to use interventional management. In late term mares, the induction of parturition can be initiated over 5 to 7 days by administering 100 mg/day of dexamethasone.7 This can potentially help improve the overall viability and maturation of the neonate. However, the size of the body wall defect
and progression of further muscle tearing may limit this option. Though conservative management has recently been recommended over interventional management, the exact process of making decisions may vary from case to case based on the value of the mare or foal, age of the mare, severity of the defect, foal’s or mare’s potential viability, or economical concerns of the owner. Therefore, the choice of therapy for pregnant mares with body wall tearing or prepubic tendon rupture will vary. Surgical repair of body wall defects in mares can be performed in select cases. The decision to
Figure 250.2 A pregnant mare with a prepubic tendon disruption.
Figure 250.3 A pregnant mare with a prepubic tendon disruption. The ventral abdomen is examined by ultrasound to determine the extent of the disruption and fetal viability.
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surgically repair a body wall defect will often depend on the location and size of the defect. Surgical repair is often recommended to be carried out 12 to 16 weeks after foaling. In acute cases, secure closure of defects is not possible due to the fragility of the tissue. Edematous tissue will not support a closure with significant tension. Surgical repair should be performed once the border of the defect has formed adequate scar tissue to secure suture material and/or synthetic mesh. The method of repair is generally based on the size of the defect. Surgical repair may not be warranted in all cases and is generally not necessary for very small body wall defects. For larger defects, mares should be taken off feed for approximately 12 to 24 hours to minimize gastrointestinal distension. Broad-spectrum antibiotics and non-steroidal anti-inflammatory agents should be administered immediately prior to surgery and continued postoperatively. Mares should be positioned in dorsal recumbency with slight rotation to the side to provide adequate exposure to the body wall defect. The ventral abdomen is clipped and aseptically prepared. A curvilinear incision is made around the defect and the skin is elevated over the defect. If the defect can be apposed, the herniated peritoneum can be resected or left in situ. The peritoneum should be resected carefully to ensure there are no adhesions present on the peritoneal surface. Interrupted stay sutures are placed through the edges of the defect using an appositional or overlapping pattern (Figure 250.4). Absorbable and non-absorbable suture material can be used. If the hernia edges cannot be apposed due to tension, then closure may require the placement of a synthetic mesh. If possible, the mesh should be placed on the outside of the peritoneum and body wall closure.8, 9 Placement of the mesh
Figure 250.4 Closure of a body wall defect. The interrupted sutures have been pre-placed using a vest-over-pants suture pattern (also know as a modified horizontal mattress or Mayo overlap technique).
within the abdomen can result in adhesion formation between the mesh and bowel. The mesh is secured to the defect edges and the skin is closed over the mesh. Proper aseptic technique is extremely important when placing non-absorbable sutures or mesh to prevent chronic implant infections. Postoperatively, all repairs should be supported using abdominal support such as abdominal wraps. The support wraps should be changed daily or every 3 to 5 days depending on the method of application. Velcro wraps can be changed and reset daily. Abdominal wraps placed using elastic tape can be changed every 3 to 5 days and only the ventral portion of the wrap should be removed. The prognosis for the survival of the foal in mares with body wall ruptures is reported to be good when the mare is initially treated conservatively. Parturition should be allowed to occur naturally or be induced if the gestation age of the foal is known. The prognosis for mares with body wall ruptures depends on the size and location of the defect. Small defects generally have a better prognosis than larger defects. Surgical repair of larger defects is also more difficult and predisposed to failure. It is discouraged to breed mares again with prepubic tendon ruptures. Mares with small body wall defects can be bred again; however, close monitoring of the defect should be performed during the last trimester. Future fertility of mares surviving with large body wall defects may be achieved using embryo transfer.
References 1. Adams S. Rupture of the prepubic tendon in a horse. Equine Pract 1979;1:17–19. 2. Hanson R, Todhunter R. Herniation of the abdominal wall in pregnant mares. J Am Vet Med Assoc 1986;189:790–3. 3. Jackson P. Rupture of the prepubic tendon in a shire mare. Vet Rec 1982;111:38. 4. Mirza MH, Paccamonti D, Martin GS, Ramirez S, Pinto C. Theriogenology question of the month. Rupture of the prepubic tendon with additional tearing of the abdominal tunic. J Am Vet Med Assoc 1997;211:1237–38. 5. Ross J, Palmer JE, Wilkins PA. Body wall tears during late pregnancy in mares: 13 cases (1995–2006). J Am Vet Med Assoc 2008;232:257–61. 6. Habel RE, Budras KD. Anatomy of the prepubic tendon in the horse, cow, sheep, goat, and dog. Am J Vet Res 1992;53:2183–95. 7. Ousey JC, Kolling M, Allen W. The effects of maternal dexamethasone treatment on gestation length and foal maturation in Thoroughbred mares. An Reprod Sci 2006;94:436–8. 8. Kelmer, G, Schumacher J. Repair of abdominal wall hernias in horses using primary closure and subcutaneous implantation of mesh. Vet Rec 2009;163:677–9. 9. van der Velden MA, Klein WR. A modified technique for implantation of polypropylene mesh for the repair of external abdominal hernias in horses: a review of 21 cases. Vet Q 1994;16(Suppl 2): S108–10.
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Chapter 251
Uterine Prolapse Michael A. Spirito and Kim A. Sprayberry
Uterine prolapse is an uncommonly encountered problem in the mare, but must be dealt with as a life threatening emergency. Uterine prolapse is defined as the falling or sliding of the uterus into and beyond the vaginal vault. In the mare, the reproductive tract extends from the level of the 4th or 5th lumbar vertebra to the vulva. The uterus is suspended in the abdominal cavity by the broad ligament, which derives its attachment from the lateral sublumbar and pelvic walls. The arteries, veins, lymphatic vessels, loose collagenous and adipose tissue, and sheets of smooth muscle continuous with the outer layer of the uterine tube and uterus are contained between the 2 layers of the broad ligament, which becomes continuous with the lateral ligaments of the bladder.1 The broad ligament attaches to the dorsal aspect of the uterus and cervix. In the pregnant mare, the uterus and supporting structures become remarkably developed, concomitant with development of the fetus. The importance of these adaptations to pregnancy should not be underestimated by the practitioner confronted with a mare with uterine prolapse. In the foaling mare, the anatomy of this area becomes noticeably altered in the days and hours preceding parturition. These changes include a general relaxation of the supporting structures of the uterus and all of the structures in the pelvic canal, including the bladder, urinary sphincter, rectum, vulva, and muscles of the pelvic limbs. In addition, marked swelling and congestion of this area develop prior to and in preparation for parturition. The uterine arteries, which, in nongravid mares are several millimeters in diameter, attain a diameter of up to 10mm in the late stages of pregnancy. The broad ligament is substantially lengthened to accommodate the degree of enlargement in the uterus at the time of birth. The causes of uterine prolapse are multiple. The most common cause seen in the authors’ practice is
dystocia, followed by retained placenta, and lastly, as a sequela of abortion. It is not uncommon for some degree of prolapse to occur after abortion even in a mare that is only in the 8th month of gestation. Other less-common causes of uterine prolapse are straining and tenesmus (often secondary to undiagnosed pain in the pelvic region), colic, and cribbing.2 In addition, positioning a periparturient mare in dorsal recumbency for a surgical procedure substantially enhances the risk of uterine prolapse. At the authors’ practice, the vulva is held closed with towel clamps to decrease this risk in periparturient mares undergoing surgery. With all instances of dystocia, the uterus should be digitally examined after delivery of the foal. It is very common in these situations to detect invagination of the ovarian pole of the uterus, which can lead to complete uterine prolapse if not corrected. On rare occasions, the uterus may prolapse concurrently with the foal’s exit as it is pulled from the birth canal. Prolapse of the uterus can arise in primiparous mares as well as in multiparous mares, and prolapse has been reported in a nulliparous yearling.3 Relaxation of the supporting structures of the uterus, increase in size of the uterus, and intensity of uterine contractions leading up to and following foaling predispose the foaling mare to uterine prolapse. Elongation of the mesometrial attachments, combined with a flaccid, atonic, or relaxed uterus, are fundamental ingredients in the etiology of uterine prolapse. It is believed that older multiparous mares are more disposed to uterine prolapse. Diagnosis of uterine prolapse is generally straightforward. Complete uterine prolapse is recognizable by viewing of the engorged, edematous, congested uterus protruding from the vulva, and in some instances extending to the ground. Affected mares may be recumbent or standing. The mare may be in distress and have clinical signs of shock, or may appear stable at the time of initial evaluation.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 251.1 Photograph of a postpartum mare with uterine prolapse.
The condition of the uterus, and consequently the mare, is largely dependent on the length of time the prolapse has been in progress. The shorter the duration of the prolapse, the less edema and enlargement will be encountered. The more quickly the problem is diagnosed and resolved, the more likely the mare will recover uneventfully. The smaller the quantity of the uterus that is prolapsed, the less likelihood there will be for rupture of a uterine artery. Mares in which a uterine artery ruptures have a poorer prognosis and can be difficult to save. It can occasionally be challenging to differentiate uterine prolapse from rectal prolapse or evisceration of bowel segments through a rent in the uterus or vagina. Prolapse of the bladder should also be considered. If at all possible, uterine prolapse should be managed on the farm by an experienced practitioner. If that is not possible and the exact diagnosis is unclear, the mare should be transported to a clinic as quickly as possible. Keeping the uterus positioned as high as the pelvic brim as much as possible reduces the strain on the uterine arteries and supporting soft tissues, and the uterus will become less congested and edematous. The uterus should be gently
cleansed with water and a plastic bag or sheet should be used to support and suspend the uterus. Occasionally, a board may also be helpful to elevate the uterus. When the uterus is suspended, the mare will instantly become more comfortable and may strain less. Manual replacement can be accomplished by positioning the mare so that the hindquarters are higher than the front end. This can be achieved by placing the mare on an incline or a small hill, or by standing her front end in a pit or depression. Prior to replacement, the uterus should be cleansed with a dilute povidone solution or a warm-water shower with a hose. The goal is to remove as much gross contamination as possible while delaying reduction of the prolapse as little as possible. Placental attachments still adhering to the uterus should be removed. However, rapid resolution of the problem is more important than scrupulous cleansing of the uterus. It is very important to evaluate the uterus for tears prior to reducing the prolapse. If a tear is found, it should be repaired by placing sutures in an inverting pattern in two layers. The uterus is manipulated with the flat of the hand, with every effort made to keep pressure evenly distributed and avoid rupture or tearing of the potentially friable organ. The concern of many is to reduce some of the uterine edema prior to replacement, but it is more important to restore the uterus to its normal anatomic position as the edema will rapidly subside once this is accomplished. To replace the uterus, identify the everted tip of the uterus and apply steady pressure against it while pushing it into the vagina and through the cervical ring. The rest of the uterus will follow. In mares that are particularly fractious and painful, an epidural may assist in replacement of the uterus. In the authors’ clinic, it is not unusual to anesthetize and hoist the mare by the hind limbs; with this technique, gravity helps return the uterus to normal anatomic position. It is important to verify by manual palpation the position of both the gravid and nongravid horns after reduction of the prolapse. This will help ensure that there is no longer any invagination or telescoping of the horns. If an invaginated horn is not completely reduced, colic, straining, and a higher probability of reccurrence will result.4 If it is difficult to position the uterus correctly, a rectal sleeve filled with water and tied off and placed inside of another sleeve will provide an excellent probe with which to aid in repositioning of the inverted horn. This is as effective as filling the uterus with water and more easily controlled. The mare may be treated by means of intrauterine antimicrobial boluses or an antimicrobial infusion after reduction of the prolapse. It is advisable to place retaining sutures in the vulva with umbilical tape or heavy suture material after manipulation. Sutures may be removed in 24 hours.
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Uterine Prolapse A complicating factor occasionally encountered after reduction of a prolapsed uterus is continued tenesmus. This complication can be difficult to manage. In these situations mares may continue to strain despite a normally positioned organ and an absence of other apparent problems. Management of these mares involves placement of a catheter in the epidural space and pain management with with a combination of butorphanol and xylazine. If uterine artery rupture has occurred as a sequel to prolapse, signs of shock, and, in many cases, terminal hemorrhage is detected. Once the uterus has been replaced and repositioned in situ, the mare should undergo close evaluation for medical problems that may develop as complications of compromised vascular supply, especially endotoxemia. Injury to ovarian or uterine vasculature may manifest acutely, as arterial rupture and internal hemorrhage and shock, or may manifest more gradually in the following days as the effects of disruption of blood supply to a portion of the uterus become apparent. Mares with the latter condition may have malaise, fever, alterations in the hemogram, or colic. During the week following reduction of uterine prolapse, the mare should be monitored with physical examination and blood work, and possibly also with ultrasound imaging and manual examination of the uterine lumen to ensure ongoing return to normal with regard to tone and viability. Mares with signs of compromise at admission or initial examination should be stabilized with intravenous fluids and broad-spectrum antimicrobials. Pain should be managed with analgesics, which can be used according to clinician preference. In mares managed in the field, installation of an epidural catheter may not be feasible or possible, and placement can be technically difficult. In those mares, a multimodal approach to analgesia may yield favorable results and may include administration of some combination of flunixin meglumine, butorphanol, and intravenous lidocaine infusion. Detection of leukopenia secondary to neutropenia, a neutrophilic left shift, sustained high fibrinogen concentration, or other changes should prompt attention to prophylaxis for complications of endotoxemia, such as laminitis. Development of such hematologic changes several days after reduction of the prolapse, in association with fever or signs of malaise, should prompt suspicion of a compromised segment of uterine wall or full-thickness tear. Diagnostic techniques used to assess affected mares for this complication would include some combination of manual exploration of the uterine lumen, abdominal palpation per rectum, transabdominal ultrasound, transrectal ultrasound, abdominocentesis, en-
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doscopic imaging of the uterine lumen, laparoscopy, or celiotomy. Signs of acute rupture of the ovarian or middle uterine artery are straightforward and are those of acute hemorrhagic shock, including tachycardia, sweating, trembling, pallorous (rather than toxic or cyanotic) mucous membranes, and cold extremities. These signs are referable to acute hypovolemia, not anemia, and many mares can be stabilized with partial-volume resuscitation with crystalloid fluids, with or without transfusion of whole blood or a synthetic oxygen-carrying solution. Artery rupture can be diagnosed by transabdominal ultrasound or via gentle rectal palpation and recognition of asymmetric bulging in a broad ligament; containment of the extravasated blood in the broad ligament is associated with a better prognosis for survival than that in mares that hemorrhage freely into the peritoneal cavity. However, mares in the latter situation often also respond favorably to management with permission hypotension and low-volume isotonic fluids, although in some instances transfusion with fresh whole blood or an oxygen-carrying substitute may be necessary. Diagnosis of acute intraabdominal arterial rupture can be made most promptly via transabdominal ultrasound imaging or abdominocentesis; hematologic assessment during the acute phase of hemorrhage is unhelpful, as decreases in hematocrit and serum protein concentration will not be detected unless the mare survives long enough to volume replete through drinking or is supported with IV fluids. Changes in the hemogram (especially in protein concentration) typically become apparent 6 to 12 hours after the hemorrhagic event. The magnitude of decrease in serum protein concentration is a better indicator of the severity of circulating blood loss than is the decrease in hematocrit. The prognosis for mares with uterine prolapse is good if there are no tears in the uterus and the uterine arteries have not been damaged.5, 6 Most mares that survive the initial uterine prolapse and subsequent reduction of the uterus have a normal recovery. Mares that die from uterine prolapse are almost exclusively those that have tearing of the uterine arteries. At the Hagyard Equine Medical clinic, of 18 mares admitted for uterine prolapse from 2001 through 2007, 15 survived to discharge from the hospital. Many mares with uterine prolapse are managed in the field and are not referred to a referral center. It is nearly always preferable to reposition the uterus in the field rather than expose the mare to the additional time required for transport to a clinic. It is generally accepted that the uterus will normalize in a short period of time after correction of the prolapse. The edema will dissipate from the uterus fairly quickly, and by several days after the event, the
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uterus will scan and palpate as would be expected in a normal postpartum mare. The prognosis for reproductive soundness is very good as well. There should be no long term negative effects expected as a result of uterine prolapse. There is no tendency for uterine prolapse to recur at subsequent parturitions.7–9 Mares that have experienced uterine prolapse will cycle and conceive and can have normal reproductive lives, although they may be difficult breeders if rebred in the same season as the prolapse.
References 1. Kainer RA Reproductive organs of the mare. In McKinnon AO, Voss JL. Equine Reproduction. Malvern, Pennsylvania: Lea & Febiger, 1993:5–19.
2. Slack A. 1973 Uterine prolapse in a mare. J Am Vet Med Assoc. 162:780. 3. Schambourg MA, Idiopathic prolapse of 1 uterine horn in a yearling filly. 2004 Can Vet J; 45:602–604. 4. Pascoe J, Pascoe R: 1988 Displacements, malpositions and miscellaneous injuries of the mares urogenital tract, Vet Clin North Am Equine Pract 4:439–450. 5. Correspondence: 1975 Uterine prolapse in the mare. Vet Rec March [Vol 96, Issue 9, Pages 207–8. 6. Letter: 1975 Uterine Prolapse in the mare Nisbet, A. Vet Rec Apr [Vol 96, Issue 14, Page 324. 7. Burgess J. 1975 Uterine prolapse in the mare. Vet Rec, 96, 513. 8. Cran H.R. 1975 Uterine prolapse in the mare. Vet Rec 97, 19. 9. Gray J.D. 1975 Complete uterine prolapse in the mare. Vet Rec, 96,98.
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Chapter 252
Uterine Torsion James R. Vasey and Tom Russell
Uterine torsion is an infrequent but serious complication of pregnancy in mares. It may account for 5–10% of serious dystocia in the equine.1 The factors which initiate uterine torsion are unclear but may include fetal activity and/or rolling of the mare. The degree of torsion can vary from 180◦ to 540◦ , in either a clockwise or counterclockwise direction as viewed from the rear. The broad ligament, or mesometrium, which suspends the uterus, has an extensive base in the sublumbar region, which limits the potential for uterine torsion in the mare compared with the cow. No breed or age predilection has been demonstrated in the horse.
Diagnosis The condition generally occurs in the last trimester of gestation and should be a major differential diagnosis in mares which present at 8 months of gestation or later with signs of colic. Cases have been reported from as early as 5 months of gestation. Most cases present soon after onset, depending on frequency of observation. The more rarely described chronic presentation will be dealt with later in the chapter. Presenting signs associated with uterine torsion include depression, periodic pawing, looking at the flank, kicking the abdomen, and rolling. The severity of pain and level of elevation of heart rate are related to the degree of torsion and any vascular compromise and/or concurrent involvement of the gastrointestinal tract. Rectal temperature and respiratory rate are within the normal limits or only slightly elevated. Abdominal auscultation will reveal normal or reduced gastrointestinal sounds. Dystocia may be the presenting sign if torsion of the uterus occurs near term. Abdominal contractions of foaling are absent as the torsion prevents the fetus from being delivered into the pelvic canal. Usually the mild, intermittent signs of colic respond temporarily to analgesics.
Unlike in the cow, torsion generally occurs cranial to the cervix in the horse; consequently vaginal examination is not usually diagnostic for the condition, but should be carried out to assess cervical tone and state of dilation, and to identify any abnormal discharge. A definitive diagnosis is based on careful rectal examination. In late pregnancy, the broad ligaments are pulled tightly downward in front of the brim of the pelvis by the gravid uterus. Identification of the ovarian pedicle and uterine ligaments should suggest whether the rotation is clockwise or counterclockwise. Both occur with equal frequency.2 The uterine ligament on the side to which the rotation has occurred is pulled directly downwards and under the uterine body. The other uterine ligament is displaced medially and runs over the uterine body towards the side of rotation and then downwards. The greatest tension is on the ligament that the rotation is turned towards, and the severity of the torsion may be inferred by the amount of tension on the ligaments. Occasionally the direction of torsion is difficult to diagnose per rectum. The position and orientation of the rectum may occasionally be useful in this regard. Some constriction of the small colon may exist, depending on stage of gestation and degree and site of the torsion, which may restrict abdominal exploration per rectum. Tympany of small or large colon may also make rectal evaluation difficult. Palpation of the body of the uterus may indicate that it has become involved in the torsion, in which case the fetus cannot be readily palpated because of cranial displacement in the abdomen. All palpable organs within the abdominal cavity should be examined to rule out a concurrent or associated gastrointestinal condition. Any tense band in the pregnant mare needs to be carefully examined to distinguish between the taeniae coli and the broad ligaments of the uterus. The uterine wall should be carefully
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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palpated to assess degree of congestion, necrosis, or possible rupture. Rupture of the uterus is a not uncommon sequel to prolonged uterine torsion. The position, extent, and duration of the tear will influence surgical access and resultant prognosis. Transabdominal ultrasonographic examination can provide crucial information with regard to management of the torsion, and is best performed with a low-frequency (2.5–4 MHz) probe. Clipping is helpful in obtaining the best diagnostic image, but wetting the hair with alcohol will usually suffice in fine-coated animals. A full ultrasonographic abdominal exploration should be undertaken. Concurrent surgical gastrointestinal conditions may be present which will influence choice of treatment. A careful search should be made for distended or edematous viscera. The foal should be assessed for signs of viability and/or distress. Fetal heart rate is a useful indicator of fetal stress levels. The uterine wall should be assessed for signs of edema, and colorDoppler may give some indication of myometrial blood flow. Copious free fluid in the abdomen should alert the clinician to the possibility of uterine rupture, although peritonitis and hemorrhage should also be considered and evaluated by abdominocentesis. With uterine rupture the foal can usually be clearly seen to be floating free in the abdomen. Abdominal paracentesis is indicated, particularly if rectal examination does not confirm a diagnosis of uterine torsion, or when a concurrent gastrointestinal lesion is suspected.3 Peritoneal fluid may be difficult to obtain from a mare in late pregnancy because the uterus occupies a large amount of the ventral abdomen. Ultrasonographic location of peritoneal fluid pockets may be helpful; nevertheless retrieval of fluid may not be possible. The cytology of peritoneal fluid retrieved from mares prepartum and postpartum does not vary significantly from normal peritoneal fluid. Sero-sanguinous fluid may indicate devitalization of intestine or the uterus, and usually worsens the prognosis as compared to an uncomplicated uterine torsion. Peritoneal lactate may also provide useful information in this regard.
Treatment Non-surgical correction
Manual rotation through the cervix When uterine torsion has been diagnosed at term, an attempt may be made to correct the torsion manually per vaginum. Successful delivery depends on passage through the cervical canal of the clinician’s well-lubricated hand and arm. This will usually only be possible if the torsion is less than 270◦ . If the fe-
tal membranes are intact, they should be ruptured to release fluids and reduce the size and weight of the uterus and its contents. It is essential that the mare remains standing throughout the procedure; hence sedatives should be avoided but epidural anesthesia may help eliminate abdominal straining. Where possible, elevation of the mare’s hindquarters on an inclined plane will provide more space in the caudal abdomen, as the large and small bowel will move forward. The clinician’s hand and arm should be inserted as deeply as possible into the uterus and a substantial part of the fetus (upper forelimb or body) should be grasped. The fetus and uterus are then rocked back and forth through a small arc (25 to 30 cm) until, with an extra effort in a semicircular motion, opposite to the direction of the torsion, detorsion is accomplished. A second effort may be necessary for complete detorsion. Complete detorsion is recognized by normal dorsopubic delivery of the fetus, lack of twisting of the cervix or uterine body, and normal position of the uterine ligaments detected by rectal palpation. Following correction of the torsion the mare should spontaneously commence second-stage labor, although this may be delayed because vascular congestion and edema diminish uterine contractility. If the mare does not spontaneously deliver the foal following full dilation of the cervix, either the mare can be induced to foal with 10–20 IU of intramuscular oxytocin, or the foal can be delivered with manual assistance. Over 80% of uterine torsion seen at parturition can be corrected by this technique.1 Mares in advanced pregnancy with a partial uterine torsion in which the cervix is sufficiently dilated to deliver the foal need to be carefully evaluated. Traction applied to the foal in this case may cause uterine rupture, resulting in fatal hemorrhage or peritonitis. Before attempting delivery, a rectal examination should establish if a uterine torsion exists and then appropriate corrective procedures should be undertaken as required. Some uterine torsions of less than 180◦ may correct spontaneously.
Rolling Rolling the anesthetized mare can be used to correct uterine torsion but should not be used near term because of the increased risk of uterine rupture.4 The mare is anaesthetized and placed in lateral recumbency on the side toward which the torsion is directed. The mare is then turned quickly in the direction of the twist, with the objective of turning the body around the uterine axis. For example, if a clockwise torsion (when viewed from behind the horse) is diagnosed, the mare is positioned in right lateral recumbency and the mare rotated in a clockwise direction. This technique depends on fetal weight
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Figure 252.1 Technique for non-surgical correction of a uterine torsion. The mare is placed in lateral recumbency, the side the uterus has turned toward is down. In this case, the mare has a clockwise uterine torsion. The mare is slowly rolled over to the opposite side while a board and weight are used to keep the uterus from rotating with the mare. (Adapted from Bowen JM, Gaboury C, Bousquet D. Non-surgical correction of a uterine torsion in the mare. Vet Rec 1976;99:495–6.)5
maintaining a constant uterine position while the maternal position changes. Often, rolling needs to be repeated or the mare rocked back and forth while positioned in dorsal recumbency to help correct the uterine torsion. A modification of this technique involves the use of a plank of wood to help stabilize the fetus while the mare is rotated (see Figure 252.1).5 A person kneels on a board (2 to 3 m long and 20 to 30 cm wide) that has one end across the recumbent mare’s upper paralumbar region and the other end on the floor near the mare’s feet. The mare is then rolled as described above, although more slowly, to reduce risk of uterine rupture. In cases where the fetus is ballotable through the abdominal wall, the plank is repositioned during the rolling procedure as necessary to
maintain maximal leverage on the fetus. To achieve correction, this procedure may need to be repeated several times. Progress is assessed by rectal palpation, although this may be difficult while the mare is recumbent. Although difficult, and carrying the risk of rectal penetration, on occasion the fetus can be stabilized per rectum while the mare is carefully rolled.
Surgical correction
Standing flank laparotomy In tractable mares, without evidence of uterine rupture, uterine torsion may be corrected by standing flank laparotomy. The flank incision is made on the
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side toward which the torsion is rotated. For example, in the case of a clockwise torsion, the incision is made in the right flank. It is easier to detorse the uterus by lifting and then pushing, than by pulling, which may predispose to tearing of the uterine wall and subsequent rupture. If the direction of torsion is not clear, then the incision should be made in the left flank to avoid the cecum. The mare is placed in stocks and sedated only enough to ensure that she remains standing during the procedure. The appropriate paralumbar fossa is clipped and prepared for aseptic surgery. Local anesthetic (40–80 mL) is infused in a vertical line at, or slightly cranial to, the anticipated incision in the middle of the paralumbar fossa. A modified grid approach is used. The skin and external abdominal oblique muscle are incised vertically, and then the internal abdominal oblique and transverse abdominal muscles are separated by blunt dissection along the direction of their fibers. The peritoneum is punctured with a thrusting single finger to avoid visceral penetration. The direction of the torsion is confirmed by palpating the direction of displacement of the broad ligaments of the uterus and palpating the dorsal surface of the uterine body forward from the cervix. The surgeon’s arm is passed ventrally along the body wall and a prominent part of the foal within the uterus is grasped and used to gently rock and lift the uterus towards the surgeon. Resolution of the torsion is usually immediately obvious, and is confirmed by palpation of the broad ligaments and the dorsal surface of the uterus beginning at the cervix. On occasion, lengthening the incision to allow two hands to enter the abdomen or a bilateral flank incision with two surgeons is sometimes necessary, especially in late gestation. Closure of the incision is routine. The external and internal abdominal oblique muscles and subcutaneous tissues are apposed by a simple continuous suture pattern with a No. 1 or 2 absorbable material. The skin is closed with a continuous simple or interlocking pattern of non-absorbable material.
Ventral midline laparotomy Following induction of general anesthesia, the mare is positioned in dorsal recumbency and a 15–25 cm long ventral midline incision is made immediately cranial to the umbilicus. Consideration should be given to minimizing the length of the incision, such that less strain is placed on the wound at parturition. This approach allows better access to the abdomen than with a standing flank approach, and enables both arms to be inserted through the incision to correct the torsion. This approach is preferred where hysterotomy or repair of a uterine tear is necessary. If a hysterotomy is indicated, this can be performed prior to detorsion, which allows easier correction of
the problem. Flooding the abdominal cavity with up to 20 litres of warm saline has been advocated to allow the uterus to be ‘floated’ into its normal orientation.6 The ventral midline approach allows treatment of concomitant gastrointestinal problems, and visualization of uterine viability. Disadvantages include the stress of general anesthesia on the mare and foal, the requirement for surgical facilities, and the potential for catastrophic dehiscence if the mare goes into labor soon after surgery. This latter complication may be managed by anesthetizing the mare at the onset of stage II parturition and delivering the foal using controlled vaginal delivery. Before surgery, the mare is given broad-spectrum antibiotics and a non-steroidal anti-inflammatory drug, such as flunixin meglumine (0.5 to 1.1 mg/kg i.v.), to counteract the risk of endotoxemia associated with abdominal surgery. A foal care team may be needed if cesarean section is carried out. Following surgery, antibiotics, intravenous fluids, and nonsteroidal anti-inflammatory drugs are given as necessary. Tocolytic agents such as clenbuterol may be used for the first several days after surgery in order to minimize the chance of premature chorioallantoic separation.6 Supplementation with altrenogest until parturition has also been advocated, although one study assessing its use in mares with medical and surgical colic did not detect a difference in foaling rate.7 Following surgery, the mare is confined to a stall for 3 to 4 weeks. The mare should be observed for any signs of abortion such as loss of cervical mucus plug, udder development, vulval discharge, or early signs of parturition. Non-surgical or surgical correction of uterine torsion does not adversely affect mares’ subsequent reproductive performance unless either uterine rupture has occurred or a cesarean section was performed.
Choice of treatment The optimum treatment for a mare with uterine torsion will depend on (i) economic considerations, (ii) facilities available, (iii) stage of pregnancy, (iv) duration of clinical signs, (v) viability of the foal and viability of the uterus, and (vi) whether signs of abortion or parturition are present. For a mare at term, when the cervix has dilated, manual correction via the cervix should be attempted. When lack of facilities, or economical considerations, preclude surgery, then rolling the mare may be indicated. This is most likely to be successful in those mares in earlier gestation. Early reports using this non-surgical method were associated with a high mortality rate of both mare and foal, possibly because of separation of the chorioallantois from the
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Uterine Torsion endometrium with subsequent abortion, premature birth, death of the fetus, or uterine rupture. However, more recent reports have given much better results, possibly because of improved rolling technique, selection of non-term mares, and the lack of interference with the cervix during corrective procedures.8 The potential for uterine rupture, or premature placental separation and subsequent abortion, should be carefully explained to the owner prior to using this technique. A quiet, mare, 6 weeks or more from term, with an uncomplicated uterine torsion may be treated using a standing flank laparotomy. However, sudden recumbency of the mare during manipulation may occur, leading to potential visceral contamination or even evisceration, and the procedure can be physically demanding, and potentially dangerous to the surgeon. It cannot be used in mares in intractable pain. In most situations ventral midline celiotomy represents the most versatile method of correction, and is the method of choice in most recent reports for mares in late gestation, or where concurrent surgical gastrointestinal lesions have been diagnosed or suspected. It is also usually the most expedient method of dealing with a dead fetus, where concurrent detorsion and hysterotomy may be performed. Fetal death subsequent to surgery has been linked to increasing surgery time and degree of hypotension during surgery.6 Particular attention should be paid to these factors when this technique is chosen.
Prognosis One report4 documents the birth of six live foals from seven mares on which rolling was performed. One mare was euthanized due to uterine rupture. An older report regarding the survival of mares with uterine torsion treated surgically, predominantly by flank laparotomy, had a success rate of 70% foals born alive, of those determined to be alive at surgery.2 One-fifth of the foals were presented dead at the initial surgery. A large retrospective study of 63 mares gives some useful guidance regarding prognosis and management of the uterine torsion.8 In total, 84% of presented mares survived. The stage of gestation in which torsion occurred was significantly associated with mare survival. Mares that developed torsion at ≥ 320 days of gestation were less likely to survive (65%) compared to mares with uterine torsion at < 320 days (97%). The method of correction of uterine torsion was not associated with mare survival but was associated with foal survival when the torsion occurred at < 320 days’ gestation. Survival in these foals was significantly better when uterine torsion was corrected via standing flank laparotomy as compared to ventral midline celiotomy. Overall foal survival was
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56%. However, only 32% of foals survived when presented after 320 days of gestation, as compared to a 72% survival rate when presented before 320 days of gestation. This statistic should be considered when confronted with mares in late-stage gestation, and consideration should be given to whether saving the mare or the foal is paramount. Foals left in situ in late-stage gestation have a high probability of dying, and often present with dystocia at parturition. This may be difficult to correct, possibly requiring further surgery, and is likely to put severe strain on a ventral midline laparotomy incision. Mean heart rate at admission was significantly lower in surviving mares than for mares that died. Most mares with an admission heart rate below 60 b.p.m. survived, while most with an admission heart rate above 70 b.p.m. died.
Chronic uterine torsion This condition has been described in detail in two mares.9 Presenting signs were vague and included inappetence and chronic depression of several weeks’ duration. One mare was febrile; the other had diarrhea and mild laminitis. Rectal examination showed an abnormal mass to be present in the caudal abdomen, and definitive signs of torsion were absent. At surgery, extensive adhesions were found between the uterus and the peritoneum and other viscera. The uterus was hemorrhagic and friable. Removal of the fetal fluids was accomplished creating a 1 cm incision in the middle of a pursestring suture, through which a suction tip was placed. Removal of the dead fetus was then undertaken before detorsion and removal of the uterus by ovariohysterectomy. A caudal ventral midline approach was necessary to gain access to the cervix. The ovarian vessels were cut and crushed with an emasculator and ligated using 1 polydioxanone. A surgical stapler was applied cranial to the cervix and used to retract the uterus cranially. The uterus was transected caudal to the stapler, and the uterine stump was oversewn with 1 polydioxanone in a Cushing pattern in two layers. The broad ligaments could not be identified and thus ligation of the associated vessels was not necessary. Obviously this procedure ends the animal’s reproductive usefulness and this should be made clear to the owner before the decision is made to proceed with this surgery.
References 1. Vandeplassche M, Spincemaille J, Bouters R, Bonte P. Some aspects of equine obstetrics. Equine Vet J 1972; 4: 105–13. 2. Pascoe JR, Meagher DM, Wheat JD. Surgical management of uterine torsion in the mare: a review of 26 cases. J Am Vet Med Assoc 1981;179(4):351–4.
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3. Pascoe JR, Pascoe RR. Displacement, malpositions and miscellaneous injuries. Vet Clin N Am Equine Prac 1988;4: 439–50. 4. Witchell JJ, Reinertson EL, Clark TL. Non-surgical treatment of uterine torsion in seven mares. J Am Vet Med Assoc 1988;193(3):337–8. 5. Bowen JM, Gaboury C, Bousquet D. Non-surgical correction of a uterine torsion in the mare. Vet Rec 1976;99: 495–6. 6. Jung C, Hospes R, Bostedt H, Litzke LF. Surgical treatment of uterine torsion using a ventral midline laparotomy in 19 mares. Aust Vet J 2008;86(7):272–6.
7. Tracey SC, Whitehead AE. Foaling rates and risk factors for abortion in pregnant mares presented for medical or surgical treatment of colic: 153 cases (1993–2005). Can Vet J 2009;50(5):481–5. 8. Chaney KP, Holcombe SJ, LeBlanc MM, Hauptman JG, Embertson RM, Mueller PO, Beard WL. The effect of uterine torsion on mare and foal survival: a retrospective study, 1985–2005. Equine Vet J 2007;39(1):33–6. 9. Doyle AJ, Freeman DE, Sauberli DS, Hammock PD, ¨ Lock TF, Rotting AK. Clinical signs and treatment of chronic uterine torsion in two mares. J Am Vet Med Assoc 2002;220(3):349–53.
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Chapter 253
Abnormalities of Pregnancy ¨ Robert M. Lofstedt
Fetal malformation due to transverse and posterior presentation Although this is of no significance in other species, the position of the fetus within the uterus appears to affect the incidence of fetal malformation in horses. If a pregnancy is examined during the first half of gestation, one need not become concerned if the fetus is found to be in posterior or transverse presentation because it may change its presentation until late in gestation (see below). This is due to the fact that the equine fetus is more mobile than the ruminant fetus, because the amnion is separate from the chorion. This allows the fetus to rotate within the amnion and the amnion to rotate within the chorioallantois (allantochorion). Eventually however, fetal presentation dose become fixed, usually by 8 months of gestation and, in almost every case, in anterior longitudinal presentation.1 There is no doubt that posterior longitudinal gestation is abnormal in horses. Before making a diagnosis of posterior presentation in late gestation and discussing the possibility of foaling complications and fetal malformation with an owner, one should consider the normal position of the fetus in the abdomen during late gestation. When a rectal examination is performed at this time, for example, to examine placental thickness in cases of threatened abortion, the hind legs of the fetus can be palpated close to the pelvic inlet. This does not mean that the fetus is in posterior presentation. It is merely because the fetus lies in the uterus in a tight ‘C’ position, with its head and forelimbs close to the cervix and its hind legs suspended above it, adjacent to the ovaries and often within easy reach of the operator. In 601 cases of dystocia,2 it was found that fetal anomalies were disproportionally common when the fetus had developed in posterior or transverse presentation. When fetuses were in posterior presentation, the proportion of fetal malformation was almost
20 times higher than expected and, when they were in transverse presentation, about 200 times higher. This suggests that that normal fetal growth and movement cannot be accommodated by the uterus if a fetus is lying in an abnormal presentation. This may also apply when the posture of the fetus is abnormal. When the head of the foal is retained (i.e., there is neck flexion), uterine compliance appears to be so limited in late gestation that the neck vertebrae are forced to ossify within in a flexed posture, causing a common malformation known as ‘wry-neck’. Although these assumptions are logical, they are, nevertheless, surprising in terms of the fact that the uterine body is full of fluid in late gestation and the head and neck of the fetus are so mobile at that time.3 When fetal anomalies are severe enough to preclude normal birth and cesarean section is required, foals may be born with severe malformations such as scoliosis, torticollis, and limb deformities. The prognosis for these foals may appear to be dismal, but apparently spontaneous recovery is possible.2 Therefore, decisions for euthanasia should be delayed until it is obvious that the prognosis is poor. When the entire conceptus develops in the uterine body with minimal involvement of the uterine horns, the condition is known as a ‘body pregnancy’.4 After examining large numbers of placentas over a period of 25 years, the author has no doubt that this is abnormal, but a search of the literature and personal experience does not suggest a link between body pregnancies and fetal anomalies.
Uterine torsion Uterine torsion is usually a cause of colic, especially during the latter third of gestation. This is very different to the situation in cows, where torsion is essentially a problem associated with calving. It is essential to be aware of this difference because, once uterine torsion has been relieved in a mare, no
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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consideration is given to delivering the foal as it is usually too immature to survive. In some cases, however, torsion can occur at term and then delivery of the foal is feasible. Uterine torsion should be suspected in any mare that is pregnant during the last one-third of gestation and is presented with colic. The diagnosis is facilitated by the mesometrial ligament being pulled tightly to one side and may often be clearly palpable per rectum. The basic principle for non-surgical correction of uterine torsion in mares is the same as for cows, i.e., one attempts to ‘roll the dam around her uterus, leaving her uterus behind’. If the direction of torsion is not absolutely obvious on rectal examination, torsion should be corrected by surgery.5 Although a minimum of restraint is required in cows, general anesthesia must be used to roll a mare. After induction, it is helpful if a plank is placed across the abdomen to stabilize the uterus while the mare is rotated on her longitudinal axis (Figure 253.1). Even if a plank is not available, one should still attempt to correct the torsion non-surgically, because the value of placing a plank across the abdomen has not been established in controlled studies. Common practice suggests that, if non-surgical correction fails, one should resort to a laparotomy to correct the torsion. Although this may be sound practice in a farm setting, retrospective analysis6 has suggested that surgical correction will result in a more favorable outcome than rolling if it is used as the initial approach to the problem. Surgical correction has been associated with a high mare and foal survival rate, especially when tor-
sion occurs early in gestation.6 Surgery can be performed by either a standing flank approach or via midline laparotomy. Correction via a flank approach is facilitated by starting a rocking motion, then flipping the uterus over when sufficient momentum has been achieved.5 However, because of the possibility of evisceration if a mare collapses during a standing flank approach, a ventral midline approach under general anesthesia may be the safest surgical approach in some cases.6 With respect to the foal, an average survival rate of approximately 80% can be expected, with survival rates > 90% when torsion occurs at less than 10 months of gestation. The survival of the mare does not appear to be significantly different between the various approaches to correcting torsion.6 Occasionally the foal dies as a result of blood vascular embarrassment due to torsion, and may be aborted soon after the torsion is corrected. With this in mind, in all cases of uterine torsion the viability of the fetus should be determined by transabdominal ultrasonography7 or identification of movement per rectum. In that context, one should be aware that fetal movement is normally frequent and vigorous during the last half of gestation.3 Therefore an absence of fetal movement for several minutes should be regarded with suspicion. To exclude the possibility of fetal death in the absence of ultrasonography, one can measure estrone sulfate concentrations, which are elevated if the foal stays alive.
Pre-term uterine rupture As mentioned above, mares with uterine torsion are usually presented with colic, but rupture of the
Figure 253.1 The basic principle for non-surgical correction of uterine torsion is to twist the animal around its own uterus. A plank such as that shown here can be used to stabilize the uterus while the mare is rotated on her longitudinal axis, but this is not essential. Image used with the permission of Dr G.F. Richardson, Department of Health Management, Atlantic Veterinary College, University of Prince Edward Island, Canada.
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Abnormalities of Pregnancy twisted uterus reportedly also occurs without observations of severe pain. It is difficult to imagine that the mare does not experience pain when this occurs, so it is likely that this often goes unnoticed by the owner. Prior to rupture, it is likely that the uterus becomes friable due to vascular strangulation but, obviously, there is a risk that uterine rupture can also occur during attempts at correcting the torsion, either surgically or non-surgically. Exsanguination and death after a uterus ruptures appears to be unusual even though the uterus has an excellent blood supply.8, 9 Nevertheless, exsanguination has been reported.10 Uterine rupture may be diagnosed using transabdominal ultrasonography where the fetus may be seen on the floor of the abdomen, lacking fetal fluids, movement and a heartbeat Electrocardiography of the fetus is often difficult but it may assist in making a diagnosis when ultrasonography is not available.9, 11 Presented in Figure 253.2 is a case of a mare with uterine rupture. A rectal examination was performed and it was determined that a normal, nonpregnant uterus was present. The mare was in estrus at that time and was artificially inseminated. Within 48 hours she had died from per-acute peritonitis due to semen having escaped from the uterus, into the peritoneal cavity. At postmortem, a dead fetus, close to maturity, was pulled from the abdomen of the mare and as shown in Figure 253.2, a rupture site was discovered
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in the ventral portion of the uterus. It was from this hole that the foal had escaped into the peritoneal cavity. Amazingly, the rupture site had healed remarkably well, leaving only a small hole in the wall of the uterus. In the case of uterine rupture illustrated here, the uterus had involuted well and the site of rupture had closed sufficiently so that it was not obvious on palpation. Therefore, in some cases, if the uterine environment is sterile, the uterus may rupture, torsion may correct itself, and involution will follow. The fetus may never be discovered in the abdomen. In most cases, laparotomy, fetal delivery and hysteropexy may be performed to recover the fetus after the uterus has ruptured.8, 10,12–15 The uterus itself is usually repaired through a ventral midline incision.8 The uterus can also be repaired by prolapsing it through the vagina, then suturing the tear16 but, obviously, this approach can only be used when a uterus ruptures close to the end of gestation.
Entrapment of the uterus within the mesentery Entrapment of the uterus within the mesentery is an extremely rare cause of fetal distress or dystocia. Indeed, the case described here may be unique. This case, presented in Figure 253.3, occurred in a Belgium draft mare referred to our clinic several hours after starting to foal. The mare had been experiencing severe colic and dystocia. Attempts by the referring veterinarian to deliver the foal were
Figure 253.2 Uterine rupture occurred in this mare, presumably as a result of uterine torsion. The mare was kept in a pasture so the symptoms of rupture would not have been noticed. Because of abdominal enlargement, however, the owner submitted this mare for a pregnancy diagnosis. Used with permission from Dr Lawrence Evans, Chairperson, Department of Theriogenology, Iowa State University, Ames, IA, USA.
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Figure 253.3 Entrapment of the uterus by a rent in the mesentery. This caused severe colic and dystocia.
unsuccessful. During a rectal examination to ascertain the presence of possible uterine torsion, the foal was not palpable, yet it was clearly palpable during per vagina examination. The owners elected to euthanize the mare. On postmortem examination a large rent was discovered in the mesentery. It appeared that this rent had occurred before pregnancy began, or perhaps during early pregnancy. At that time, the uterus must have passed through the hole in the mesentery. The foal had developed to term, with the uterus remaining on the side of the mesentery that was distal to the vagina. This explained why the foal could not be palpated per rectum yet was easily palpable per vagina. Essentially, the hand of the operator passed down the lumen of the genital tract, through the hole in the mesentery.
Hydrops of the fetal membranes Excess fluid can accumulate in the amnion (hydramnion, hydramnios or hydrops amnion)9, 17 or the allantois (hydrallantois or hydrops allantois)9,17–21 in mares during late gestation. Although hydrops allantois is more commonly reported, both conditions are rare. It is also possible that the two conditions have occasionally been confused with one another because it is not always obvious which of the two fluid compartments is affected. Mares with hydrops allantois always present with abdominal enlargement and may show abdominal discomfort and remarkable ventral edema. Also, as though ventral abdominal rupture is impending; they walk with difficulty and may even become recumbent. In fact, some of these mares will develop ventral ruptures.22 Affected mares also show labored breathing because of fluid pressure on their diaphragms. They may develop inguinal hernias and occasionally the uterus may rupture. Spontaneous abortion may also occur.18, 20
The incidence of hydrops amnion is too low to describe its general characteristics accurately, but case reports suggest that they are similar to those of hydrops allantois.9, 17, 18 In cases of hydrops, massive amounts of uterine fluid will be seen on transabdominal and transrectal scanning. However, the use of ultrasonography to verify the type of hydrops is open to criticism because both fluid compartments can lie close to the fetus and they may have similar echogenicity, especially in late gestation.7 Amniotic and allantoic fluid can, however, be distinguished from one another according to their biochemistry. The composition of allantoic fluid closely resembles urine and therefore has a higher specific gravity (∼ 1.026) than amniotic fluid (∼ 1.009) which closely resembles a plasma transudate. There are many other differences between these fluids but perhaps most useful are the high sodium and chloride concentrations in amniotic fluid (∼ 130 and ∼ 120 mmol/L respectively): about two to three times higher than the level in allantoic fluid.23 Finally, the amniotic fluid contains structures called lamellar bodies. These are membranous, platelet-sized structures that arise from type 2 pneumocytes in increasing numbers as pregnancy progresses.24 Although these data are potentially useful, one is generally not inclined to sample fetal fluid because amniocentesis itself can precipitate abortion.25 Also, the effect of hydrops on the composition of fetal fluids is not known. Nevertheless, if there is uncertainty as to the nature of the fluid liberated at the time of foaling, it may be valuable to collect a sample and measure its specific gravity and sodium or chloride concentration. To ascertain a diagnosis of hydrops of either type, ultrasonography should be used and weight gain should be monitored. The type of hydrops is based on the pre-foaling weight of the mare and, if possible, capture of fluid expelled at the onset of foaling.9,17–21 However, this is difficult, somewhat impractical, and certainly inaccurate. The total amount of fetal fluid at term can be calculated by subtracting fetal and placental weights from the difference between pre- and post-foaling weights of the mare. The placenta should also be examined to ascertain which fluid compartment was affected. In some cases, so much fluid is released when the fluid compartments rupture during foaling, that these comments are redundant and there is little uncertainty as to which placental compartment was affected. At term, one should expect about 30 liters (∼ 8 gal) of fetal fluid in total. The normal volume of allantoic fluid in mares varies from 8 to 18 L (∼ 2 to 5 gal) and the volume of amniotic fluid from 3 to 7 L (∼ 1 to 2 gal).10 Also, the normal weight of the placenta varies between 2.5 and 6.4 kg (∼ 5 and 14 lb) in various breeds.18, 26 When the weight of the foal and its
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Figure 253.4 A 17-year-old Standardbred mare presented with signs of impending rupture of ventral abdominal structures, two weeks before the expected date of foaling.
placenta are subtracted from the pre-foaling weight of the mare, one has a good estimate of the weight of the fetal fluids. Knowing that 1 kg of fetal fluid is approximately equal to 1 liter of the same, the total volume of both compartments of fetal fluid can easily be calculated. An example of where this calculation was used is shown in Figure 253.4. In that case, the referring veterinarian reported rapid abdominal enlargement over a 2-week period before presentation. Although the uterus was distended with fluid, a live fetus could be detected by ballottement per rectum. In a single day after admission, this mare gained 10 kg in weight, strongly suggesting some form of hydrops. Because of an impending ventral rupture, foaling was introduced using oxytocin and a 30 kg live filly foal was delivered in posterior presentation. Normal Standardbred foals usually weigh between 45 and 60 kg at birth, therefore this foal was underweight. The mare weighed 710 kg on admission and 570 kg immediately after foaling. Using the arithmetic method described above, the actual fluid volume lost during foaling was determined to be about 90 liters. An estimate of the fluid volume that escaped when the chorioallantois ruptured (before the amnion ruptured) was ‘approximately 100 L’. Therefore there was little doubt that most of the fluid originated from the allantoic cavity, i.e., this was a case of hydrops allantois, not hydrops amnion.
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Shortly after foaling, this mare went into severe hypovolemic shock, requiring aggressive fluid therapy. Due to the mare’s severe pain and damage of ventral abdominal structures, the owners elected euthanasia. Treatment of hydrops is directed at induction of abortion and medical support of the mare. In some cases of hydrops, induction of abortion with oxytocin is not effective18, 27 because the myometrium is so distended and atonic. If oxytocin is not effective, abortion can be induced by gradual manual dilation of the cervix over a period of 15 to 20 minutes, followed by slow drainage of the fetal fluid and forced extraction of the fetus. Upon rupture of the chorioallantois, the normally expected rush of fetal fluid may be absent17, 18 because of uterine atony. There is also a tendency for blood to pool in the abdomen after the foal has been delivered, probably because of a sudden decrease in the pressure that has been chronically exerted on the abdominal vessels. Therefore severe shock and death may ensue if intravenous fluid is not administered to maintain blood pressure. After the danger of hypovolemic shock has passed, oral electrolyte mixtures can be used to decrease the cost of treatment. Live foals have been born to mares with hydrops allantois but in some reports they were too premature to survive, while others had abnormalities such as ventral herniation, growth retardation, wry neck, limb ankylosis and deformity, brachygnathia, hydrocephalus, multiple mesotheliomas, scoliosis, and hydrencephaly.17, 18, 22 Although hydrops causes uterine inertia, once the fluid has been removed, uterine involution may progress normally. There is a lack of information on the fertility of mares that have been treated for hydrops. However, mares have been reported to conceive and foal again after experiencing hydrops allantois.18, 19 The etiology of hydrops remains unknown, but blood vascular embarrassment secondary to torsion of the amnion and umbilicus has been implicated.9 Hydrops has also been related to placentitis19, 22 and genetic predispositions have also been suggested.21 In cattle, hydrops allantois is often seen in IVF pregnancies and is almost reproducible by that technology. However, even in affected cattle, the etiology of hydrops is not understood.
Fetal mummification and maceration When a fetus dies but is not aborted, its fetal fluids and the body fluids are absorbed by the endometrium resulting in a shriveled up (dehydrated) conceptus that resembles a mummy. It is not
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known why most dead fetuses are aborted but others remain in the uterus and become mummified. If the cervix opens and the uterine environment is no longer sterile, a mummified fetus becomes infected with bacteria and maceration takes place, converting the fetus and its membranes into a mass of bones within a soupy fluid. In the case of twin pregnancies, one of the co-twins often dies spontaneously or after it is destroyed by a veterinarian. When this happens, it is often stated that it has been ‘re-absorbed’ or ‘resorbed’, implying that the uterus has some sort of digestive function. As far as this author is aware, the endometrium has no digestive capacity at all, although it is certainly able to absorb various fluids and drugs. Therefore, many embryos that die are not autolyzed completely but remain attached to their co-twins until term. It is logical that this would depend on the age of the fetus when death occurs. When fetal membranes are examined after foaling, small, mummified co-twins are occasionally seen on the chorion of the foal, barely recognizable as fetuses. Figure 253.5 shows a fetus that was recently destroyed by crushing. It had become incorporated into the chorion of its co-twin. In general, little attention is paid to the co-twin that is crushed because it and its fluids disappear almost instantly from ultrasonographic view after a crush has been performed. Figure 253.5 demonstrates that a crushed embryo may persist for some time alongside its normal co-twin, possibly to term. One can only speculate how mummies may occur, but presumably some of the numerous causes of abortion that have been described in horses may also cause mummification of fetuses. Specific infectious or genetic causes of mummification have not been identified in horses. Interestingly, conventional equine reproductive physiology suggests that a single mummy should not persist in mares because a normal placenta must
Figure 253.5 A degenerating pregnancy seen on the chorion of its co-twin 21 days after it had been destroyed by transrectal palpation.
Figure 253.6 A mummified equine fetus that was not accompanied by a living co-twin. The fetus appeared to have died at approximately 5 months of gestation and was discovered when the mare had passed her foaling date. Used with permission from Dr Lawrence Evans, Chairperson, Department of Theriogenology, Iowa State University, Ames, IA, USA.
be present to produce progestogens for the maintenance of pregnancy. Indeed, it is not surprising to find that single mummified fetuses can occur in mares that have been fed altrenogest.28 However, it is remarkable that this author and many others (J.P. Hughes, University of California at Davis, personal communication; T.L. Clark, Iowa State University, personal communication) 29–33 have encountered singleton pregnancies either undergoing mummification, or in a state of complete mummification (see Figure 253.6) or maceration, similar to the situation in cattle. In cattle, this is thought to be due to the suppression of luteolysis, causing a persistent luteal phase, but obviously this cannot be the situation in mares where pregnancy is completely independent of luteal tissue in the latter half of gestation. When fetuses die before becoming mummified, affected mares may show signs of mammary enlargement and lactation, as is the case for conventional abortions.29, 34, 35 However, these signs may be so subtle that they go unnoticed by owners. As a result, mummies are usually detected only after mares have passed their expected foaling dates. However, an unusual case has been described where a mummified fetus was discovered because it had caused a hemorrhagic vaginal discharge.36 A diagnosis of mummification is quite simple. There is a palpable absence of fetal fluid and the uterus is contracted around the fetus. If the fetus has been dead for some time and has become dehydrated, the angularity of its skeleton will be obvious. A highly echogenic mass is seen during transrectal ultrasonography. If a mummified fetus becomes macerated, the presentation is essentially the same except that a vaginal discharge may be present. This author has only seen one case of fetal maceration in 30 years and interestingly that mare was systemically normal, similar to the situation with pyometra. That
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After the uterine contents have been extracted, the uterus should be flushed with saline and antibiotics. The reproductive potential of mares after treatment for fetal mummification or maceration is unknown.
Ventral ruptures: rupture of the prepubic tendon and the abdominal muscles
Figure 253.7 Maceration of an advanced age fetus in a mare. The fetus was discovered during a rectal examination after a complaint of prolonged gestation. As in cases of pyometra in mares, this mare showed no systemic signs of disease. Treatment included tranquilization followed by slow, manual dilation of the cervix, over a period of approximately 20 minutes. When the cervix was dilated enough to admit a hand, the bones and debris were extracted and the uterus flushed with saline and antibiotics. Oxytocin treatment could follow flushing if required.
pregnancy also consisted of a single fetus and, judging by the size of its bones, it had obviously died in late gestation. Again, this demonstrates that single dead pregnancies can persist in advanced gestation in mares (Figure 253.7). It is not known if mares with macerated or mummified fetuses are consistently anestrous as is the case in cattle, or if they experience variable cyclicity like mares with pyometra.37 Experience with these cases and the equine cervix in pregnant and non-pregnant mares, suggests to this author that (in contrast to the situation in cows) macerated or mummified fetuses should be easily removable from the fetus in most cases. The technique used is the same as that used for removing (aborting) fetuses manually in the second half of gestation, when prostaglandins are not consistently abortifacient. With patience, lubrication, and a gentle approach, the cervix can be dilated safety and a live fetus or fetal debris can be removed. Pretreatment with 10 to 15 mg of estradiol cypionate 24 hours before attempting extraction has not been of value in the hands of this author. Although controlled studies have not yet shown it to be effective, 500 to 1000 µg of misoprostol (PGE1) may be valuable when applied to the cervix several hours before attempted removal. Cream based preparations can be obtained from compounding pharmacies or oral forms of misoprostol used for companion animals can be crushed and mixed with methylcellulose. A caesarean section should not be required.
Ventral herniation and rupture of the prepubic tendon are often discussed under separate headings. However, this author prefers to discuss ventral herniation of the abdominal wall, rupture of the prepubic tendon, and rupture of the rectus abdominis muscle under the collective heading of ventral ruptures. These conditions can occur together22 and appear to have the same predisposing causes. They also have similar or identical clinical presentations and are usually impossible to distinguish from one another in living mares. Although the margins of a rupture can be delineated after foaling, this author believes that postmortem examination is the only way to make definitive statements on which structures have been damaged. Ventral ruptures are most common in older mares of any breed, but it has been suggested that draft mares may be predisposed to ruptures. Extraordinary uterine weight due to twins or hydrops are additional predisposing causes.10, 22, 38, 39 In most cases, however, there is no obvious predisposing cause for ventral ruptures. Mares with ruptured ventral abdominal structures (Figure 253.8) are typically close to foaling and are presented with abdominal pain and reluctance to walk. The abdomen is usually dependant or may bulge ventrolaterally, but complete eventration (protrusion of the bowels from the abdomen into a subcutaneous sac) is rare, although this can be seen after the foal is removed and the edema has resolved. Affected mares invariably show a thick plaque of ventral edema; far more obvious than the mild ventral edema commonly seen at term. Although it may sometimes be difficult to distinguish between the two conditions, the pain that accompanies ventral ruptures usually distinguishes it from normal prepartum edema. The edema may arise because the uterus presses on the caudal epigastric and caudal superficial epigastric veins, the chief vessels that drain this area. Edema may also be caused by trauma to the abdominal muscles. Sometimes edema around the mammary gland may cause the teats to become indistinct and the neonate may even have difficulty suckling. Fortunately, edema subsides rapidly after foaling and the foal can usually suckle normally after a few days. Occasionally blood may appear in the milk because
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Mare: Problems of Pregnancy Repair of ventral abdomen defects after foaling has been described,39 but the defect is often invisible after several weeks, obviating the need for surgery in most mares. If there is a need to repair ventral ruptures, it has been suggested that surgery be postponed for at least 2 months after foaling so that edema subsides completely and the tissue margins of the hernia become stronger and better defined.42 Ventral ruptures do not usually affect the life of the mare unless she is bred again, a significant risk in the opinion of this author. Structures that have failed under the stress of normal pregnancy are likely to fail again. If additional pregnancies are planned and breed regulations allow it, embryo transfer should be considered.
Prolonged gestation
Figure 253.8 Impending rupture of ventral abdominal structures during late gestation in two mares. Ventral plaque of edema (more obvious than the usual ventral edema of late gestation) and a painful gait are indicators of this condition. Ventral edema can be so severe that the architecture of the mammary gland is almost obscured (inset). The margin between the plaque of edema and the ventral abdomen is indicated by several arrows.
tension on the mammary gland can tear some of its blood vessels. If a mare with a ventral abdominal defect has a gestation length of more than 330 days, induction of foaling should be considered40 because rupture of uterine blood vessels can occur10 and abdominal discomfort and weakness can progress to a point where the mare is no longer ambulatory. Before induction is attempted, the status of electrolytes in the mare’s milk should be considered. Although spontaneous foaling is possible (Jackson38 and unpublished observations by the author), traction may be required because the mare is not always able to exert an abdominal press. Passive immunity in the foal should also be monitored because it may be incomplete when foaling is induced.41 If pregnancy is not sufficiently advanced to allow induction, abdominal support (in the form of a special abdomen bandage or wrapping with wide adhesive tape) should be supplied until induction is feasible. A laxative diet should also be fed to decrease tenesmus during defecation.
Pathologically prolonged gestation due to abnormalities of the hypothalamic–hypophysial–adrenal axis is well known in ruminants10 but this syndrome does not seem to occur in horses. Gestation in mares usually lasts for 310 to 374 days, but normal pregnancies can be as long as 412 days.43–45 Interestingly, foals born from normal, prolonged pregnancies are not usually oversized, as they are in cattle, and do not predispose the mare to dystocia.46 In one unusual case a foal was born at 432 days of gestation but was somewhat abnormal, having a domed head and a long hair coat (AO McKinnon, personal communication, 2009). Little is known about the physiological control of the length of gestation, but its duration is known to be several days longer on average in colt than filly foals or twins.46 Studies relating gestation lengths to season have produced conflicting result, i.e., some showed that long, sustained daylength resulted in shorter gestation length and, conversely, that mares foaling in summer had comparatively short gestation lengths.45, 47 However, one study in Thoroughbreds46 showed that foals born in winter had shorter gestation lengths than those born later in the year. The reason for this disparity is unknown. The duration of gestation in horses is also partly controlled by nutrition and the genotype of the foal43, 48 but mares themselves may also control the duration of gestation to a limited extent so that their foals are born with an optimal chance for survival.48 Finally, it is possible that horses may also experience embryonal diapause, i.e., delayed embryonic development in a state of suspended animation. Some readers may be surprised with this speculation but it is not unreasonable. Embryonal diapause is well known in many other mammals.49 In all cases, embryonal diapause is referred to as occurring in preimplantation embryos, but the age of the embryo in
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Abnormalities of Pregnancy diapause is not well defined so it is possible that this may also occur in mares. In summary, there is clearly a huge range in gestation length. Therefore it is not easy to decide when gestation is pathologically prolonged. In the case of pathologically prolonged gestation due to the ingestion of ergot alkaloids (see below), this may be an easier decision because of the grazing history of the mare and an absence of mammary gland development. Many clients become concerned when their mares have not foaled after a year of gestation and begin to exert pressure on the veterinarian to induce foaling. If there is no evidence of fescue pastures, fetal movement is palpable but there is no mammary development, it is important that the clinician should not yield to this pressure, even if the duration of gestation is longer than 1 year. Instead, the owner should be reassured that this is probably a normal phenomenon and that waiting will result in a normal foal, with adequate colostrum and normal passive immunity. Prolonged gestation due to fescue toxicosis has been characterized. In these cases, gestation in affected mares is significantly longer on average than it is in mares not grazing on infected pastures. When pastures are infected with the endophyte Neotyphodium coenophialum or when they are fed ergot alkaloids such as bromocryptine to mimic fescue ingestion, gestation lengths may be 350 days or longer.50 As mentioned, mares that have ingested Neotyphodium coenophialum are also affected by agalactia. In addition, they have thickened placental membranes that may cause asphyxia of the foal during birth. The endocrinology of the syndrome is obscure, but from the known action of ergot alkaloids, it is apparent that prolactin as well as dopamine and other catecholamines are involved in the initiation of foaling. The dopamine receptor site antagonist domperidone is especially effective in treating mares (more so than phenothiazine derivatives or reserpine) that may be affected by fescue toxicosis. To prevent fescue toxicosis, doperidone can be fed from about 320 days of gestation until foaling, at a dose of 1.1 mg/kg.50, 51 Better still, mares should be removed from infected pastures at least 6 weeks before foaling.51
Hemorrhage associated with pregnancy and parturition Hemorrhage from the uterus during pregnancy and foaling Fatal extrauterine hemorrhage can occur during pregnancy when major uterine vessels rupture. This may be caused by uterine torsion, uterine rupture, or trauma experienced during violent foaling (par-
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ticularly assisted foaling), exercise, or transport10 although its cause is not obvious in most cases. Most cases of hemorrhage occur at foaling. Rupture can involve the uterine (middle uterine) arteries, utero-ovarian arteries, or external iliac arteries.52 This author has also seen a fatal hemorrhage from a mesenteric vessel that occurred during foaling. Severe intrauterine hemorrhage can also be seen when there is foaling trauma or when retained placentas are forcibly removed. There seems to be little doubt that older mares are especially predisposed to rupture of major vessels during pregnancy or foaling. In three reports53–55 that included a total of 30 mares, the average age of 25 mares that died from hemorrhage was 18.5 years. The youngest mare was 12 years old. A subgroup of 5 mares54 that only had hematomas of the broad ligament appeared to be slightly younger, with a mean age of about 12 years. Therefore, rupture of major vessels appears to be associated with aging. A causal link has been suggested between low serum copper concentrations, aging, and vascular degeneration.53 It was shown that aged mares had significantly lower serum copper concentrations than young mares, and serum copper concentrations were lowest in aged mares that suffered fatal vascular ruptures. Aging and repeated pregnancies appear to cause increased pressure within arteries because (counter-intuitively) intra-arterial pressure actually increases as blood vessels dilate (Laplace’s law). Together with excessive tension on the arteries, increased internal pressure contributes to their rupture. It has been suggested that the birth of a foal may exert pressure on the external iliac arteries, causing local vascular hypoxia, damage to the arterial walls, and subsequent rupture.52 This is a reasonable suggestion because there is no tension on these arteries during pregnancy. It is interesting that the majority of ruptured uterine blood vessels occurred on the right-hand side in one study. This was associated with the presence of the cecum in the right flank, causing displacement of the uterus to the left with a subsequent increase in tension on the right uterine artery.56 When ruptured uterine and external iliac arteries were examined by culture and biopsy, no infectious organisms or parasites were observed.52 When a mesometrial blood vessel ruptures, hemorrhage is often contained within the mesometrium but, in some cases, an artery may rupture directly into the abdomen. Alternatively, bleeding can dissect the mesometrium until it ruptures some distance away from the ruptured vessels. Acute exsanguination may follow.52
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When a large artery ruptures during foaling, it may not be noticed immediately because the discomfort of foaling and placental expulsion can mask the pain of hemorrhage. If the hemorrhage is severe, however, signs of vascular rupture will soon be recognized, including colic, sweating, rapid pulse, anemia, and death.57 In some cases, death may be immediate; in others a hematoma may rupture, causing sudden death several weeks after foaling.54 In many cases of peripartum hemorrhage, evidence of hemorrhage may only be detected when a hematoma is palpated as a large firm mass in the mesometrial ligament, often at the time of foal-heat breeding (Figure 253.9a). These hematomas have a solid to semi-solid appearance on ultrasound, the bulk of which eventually disappears, in most cases by the following breeding season. When there is severe intrauterine hemorrhage, for example, after dystocia or when a placenta has been removed forcibly, a form of tamponade may occur, filling the uterus with blood. However, hemorrhage is not usually fatal. In such cases, broad-spectrum antibiotic coverage should be applied and the clotted
blood should be left in the uterus so that it is expelled as part of the lochia. Removing the clotted blood may re-start hemorrhage. To prevent further hemorrhage, there is general agreement that mares should be in a quiet and comfortable place to rest, to prevent elevation of blood pressure. If possible, the foal should not be removed from the stall.5 Analgesic treatments such as xylazine, detomidone, flunixin meglumine, and butorphanol may also be beneficial. Although the administration of oxytocin will have no value when the external iliac or major uterine arteries have ruptured, it may decrease hemorrhage when bleeding is myometrial in origin. Therefore, doses of 10 to 20 IU every 30 minutes should be considered in all cases of peripartum hemorrhage. Additional treatments include tetanus prophylaxis, antibiotics and antifibrinolytic therapy (aminocaproic acid). Current literature does not support the use of formalin as a coagulant. Treatment of severe hemorrhage from ruptured arteries can be surgical or non-surgical, but a high death rate can be expected in both cases. There may be little success with blood transfusion but it is an attractive
Figure 253.9 (a) A hematoma seen in the mesometrium (ME) at the time of foal-heat breeding. This mare showed no symptoms at the time of foaling. The structure was clearly palpable per rectum, hence the ultrasonograph. (b) A similar hematoma (ME) to that described in (a) but more consolidated and echogenic. (c) Intraluminal hemorrhage (L UI) in a mature mare. For the mare with intraluminal hemorrhage, a foul-smelling vaginal discharge was noted at 35 days post-foaling. The mare was systemically normal but ultrasound and a gloved hand examination of the uterine contents revealed masses of clotted blood. This was presumed to be due to a severe partial thickness endometrial tear during foaling.
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Abnormalities of Pregnancy alternative to laparotomy because the surgical risk is significant in these cases. If transabdominal ultrasonography shows that there is free blood in the abdomen (often hyperechogenic because of fibrin formation) and anemia is severe, transfusion should begin as soon as possible and plasma expansion therapy should also be used. If the packed cell volume is less than 20%, whole blood is required, somewhere in the range of 15 to 20 liters. Up to 8 L (∼ 2 US gal) of blood can be removed from a healthy horse without adverse effect. Obviously this is not enough blood to raise the PCV to a normal level, however it should raise the PCV of the recipient to above critical levels. One liter of blood (∼ 2 pints) can be administered every 10 minutes58 but when exsanguination is occurring, the rate of administration can be faster. Movement of the mare should be restricted to decrease the possibility of dislodging blood clots and re-initiating bleeding. If one has time to compute exact amounts, the precise volume of blood required can be calculated by multiplying 8% (0.08) of the bodyweight of the mare by the missing fraction of the normal packed cell volume (PCV). For example, if a mare weighs 450 kg and her PCV is 18%, about 55% of her PCV has been lost. Therefore the volume of whole blood required is 0.08 × 450 × 0.55 = 19.8 L (∼ 5 gal). It is important to realize that the PCV at the time of arterial rupture will not necessarily reflect the true blood loss because of the compensatory effects of arterial constriction and splenic contraction. Often the packed cell volume and total protein are within normal limits in the first hours of blood loss (computations of the blood required should take this into account). Blood should be collected in a 4% solution of sodium citrate, if possible, from a gelding. This is because geldings cannot have preformed antibodies against the A and Q blood types. If the mare survives, her blood should be typed to assess the possibility of neonatal isoerthrolysis in future pregnancies. Hemorrhage due to vaginal and vestibular trauma during foaling Occasionally blood vessels in the vagina, vestibule, and perineal area are damaged during foaling (Figure 253.10). Some cases of hemorrhage can be fatal but, fortunately, most are not serious and require no treatment except tetanus prophylaxis and perhaps antibiotics. Even third-degree lacerations of the perineum are seldom life threatening. However, if vaginal hemorrhage is profuse and vessels cannot be ligated, a large tampon made of rolled cotton and umbilical tape may
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Figure 253.10 Severe extrauterine hemorrhage after foaling in a 9-year-old Standardbred mare that had an assisted foaling because of an apparent ‘hip lock’. The mare appeared to be partially paralyzed in her hindquarters and was unable to rise. Her heart rate was elevated at 80 beats per minute and later rose to 130 beats per minute despite appropriate treatment. Hemorrhage from the vulva lips was obvious, soaking the adjacent pasture. The owner elected to euthanize the mare. The lower image shows some of the postmortem findings: extensive muscular and fibrous tissue necrosis within the vaginal wall. There was also subluxation of the right sacroiliac joint. Severe coalescing vaginal and vulvar hematomas were also present.
be used as a pressure bandage. It should be covered with petroleum jelly and an oily antibiotic preparation such as a mastitis preparation, to facilitate its removal. When vaginal vessels are damaged, vaginal hematomas may occur (Figure 253.11). Large hematomas may cause difficulty in defecation, so laxative diets should be fed until hematomas becomes reduced in size. The use of antibiotics and tetanus prophylaxis should be contemplated as well.
Hemorrhage present at the external genitalia during pregnancy Although the placenta is well vascularized, placental detachment and impending abortion are not associated with hemorrhage from the cervix as is the
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Figure 253.12 Vaginal varicose veins in an aged mare. The vagina of this postmortem specimen has been everted to demonstrate the affected vessels.
Figure 253.11 A vaginal hematoma caused by foaling trauma. Used with permission from Dr Lawrence Evans, Chairperson, Department of Theriogenology, Iowa State University, Ames, IA, USA.
case in humans. Therefore owners can be assured that blood seen at the vulvar lips is usually unimportant and most likely originates from vaginal varicosities or, occasionally, vaginal trauma. An examination with a vaginal speculum will confirm these diagnoses. Of course, blood can also originate from the urinary tract, therefore exclusion of urinary tract pathology is important. Varicose veins occasionally develop in the vagina during pregnancy and in estrus, especially in older mares (Figure 253.12). The etiology of vaginal varicose veins is unknown59 and the reason for their presence is puzzling because there is no effect of gravity on the blood in these veins. Perhaps these veins just weaken with age. Increased blood flow to the genitalia under the influence of estrogens during pregnancy or estrus may contribute to their distention as well. Impeded drainage during pregnancy due to abdominal filling may also be contributory. Occasionally these veins may rupture and it is on these occasions that blood is seen at the vulva lips. Hemorrhage is seldom severe but it can attract flies and provokes suspicions of impending abortion. Vaginal varicose veins can be up to 4 or 5 mm in diameter and are usually most obvious in the dorsal vaginal wall, just cranial to the urethral opening on the vestibular-vaginal seal, i.e. the remnant of the hymen. They can easily be missed during a speculum examination if the speculum is withdrawn quickly
and the caudo-dorsal part of the vagina is not examined carefully. These vessels can be ligated conventionally or the ‘Ligasure’ system (www.ligasure.com) can be used (A.O. McKinnon, personal communication). Endovascular laser treatment, conventional laser treatment, or cryotherapy have also been described. Anecdotal reports suggest that ‘Preparation H’, a product used for hemorrhoids in humans, may be valuable for treating vaginal varicose veins. However, the composition of this product varies from country to country, some preparations containing phenylephrine, others not. This may explain its variable efficacy in these reports.
Colic during pregnancy When evaluating mares for colic in late gestation, one should note that the resting heart rate for mares in late gestation is usually higher than normal (40 to 60 bpm vs 28 to 40 bpm). Also, rectal examination may be unrewarding because the enlarged uterus my obscure problems in the intestine. Abdominocentesis is often performed in pregnant mares, but ultrasonography is required to prevent uterine penetration.60 Large colon torsion is the most common nongenital cause of colic in mares and it recurs more frequently in broodmares than other horses.61 Cecal rupture has also been associated with foaling.62 Therefore it is not uncommon to have to perform colic surgery on pregnant mares. As reviewed by Steel and Gibson,42 abortion can be a sequel to medical or surgical colic, especially if signs of endotoxemia (depression, anorexia, fever, tachycardia, leukopenia, and dehydration) are present. Cyclooxygenase inhibitors such as flunixin meglumine may be valuable in these cases.
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Abnormalities of Pregnancy If colic occurs during early gestation it is advisable to administer non-steroidal anti-inflammatory drugs (NSAIDs) and altrenogest (0.044 mg/kg) to maintain pregnancy in the event of luteolysis. Altrenogest has an advantage over native progesterone treatment because endogenous serum progesterone concentrations can be measured while altrenogest is given. This is because altrenogest does not cross-react with most progesterone assays. Also, it has the convenience of being an orally administered hormone. Antibiotics and fluids should be given as is appropriate. When mares are not severely debilitated, the incidence of post-colic abortions and the prognosis for fetal survival after intestinal colic appears to be only slightly higher (16 to 19%)63, 64 than the general abortion rate in mares. However, a high incidence of postcolic abortion was associated with severe intestinal debility where more than half of all severely affected mares aborted their pregnancies.65 Also, perceived risk factors such as the stage of gestation when colic occurred, the duration of anesthesia, and starvation for longer than 30 hours were not risk factors for abortion.63, 64
References 1. Bucca S, Fogarty U, Collins A, Small V. Assessment of feto-placental well being in the mare from mid- gestation to term: Transrectal and transabdominal ultrasonographic features. Theriogenology 2005;64:542–57. 2. Vandeplassche MM. The pathogenesis of dystocia and fetal malformation in the horse. J Reprod Fert Suppl 1987;35:547–52. 3. Ginther OJ. Equine pregnancy: physical interactions between the uterus and conceptus. Milne lecture. Proc Am Assoc Equine Pract 1998;44:73–104. 4. Whitwell K. The nature of adverse placental features in equine pregnancies in the UK and their relative danger to the fetus. Workshop on comparitive placentology. Victoria, Canada. Dorothy Havemeyer Foundation Monograph Series 2005;No. 17:25–7. 5. Le Blanc MM. Common peripartum problems in the mare. Equine Vet Sci 2008;28:709–15. 6. Chaney KP, Holcombe SJ, LeBlanc MM, Hauptman JG, Embertson RM, Mueller POE, Beeard WL. The effect of uterine torsion on mare and foal survival: a retrospective study 1985–2005. Equine Vet J 2007;39:33–6. 7. Reef VB Vaala WE, Worth LT, Spencers PA, Hammett B. Ultrasonic evaluation of the fetus and intrauterine environment in healthy mares during late gestation. Vet Radiol Ultrasound 1995;36:533–41. 8. Pascoe JR, Meagher DM, Wheat JD. Surgical management of uterine torsion in the mare: a review of 26 cases. J Am Vet Med Assoc 1981;179:351–4. 9. Honnas CH, Spensley MS, Laverty S, Blanchard PC. Hydramnios causing uterine rupture in a mare. J Am Vet Med Assoc 1988;193:332–6. 10. Roberts SJ. Torsion of the uterus during pregnancy. In: Veterinary Obstetrics and Genital Diseases. Theriogenology. 3rd edn. Roberts, 1986.
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11. Buss DD, Asbury AC, Chevalier L. Limitations of equine fetal electrocardiography. J Am Vet Med Assoc 1980;177:174–6. 12. Wheat JD, Meager DM. Uterine torsion and rupture in mares. J Am Vet Med Assoc 1972;160:881–4. 13. Jones RD. Diagnosis of uterine torsion in mare and correction by standing flank laparotomy. Can Vet J 1976;17:111–13. 14. Maxwell JAL. The correction of uterine torsion in a mare by caesarian section. Austr Vet J 1979;55:33–4. 15. Wichtel JJ, Reinertson EL, Clark TL. Nonsurgical treatment of uterine torsion in seven mares. J Am Vet Med Assoc 1988;193:337–8. 16. Fischer AT, Phillips TN. Surgical repair of a ruptured uterus in five mares. Equine Vet J 1986;18:153–5. 17. Allen WE. Two cases of abnormal equine pregnancy associated with excess foetal fluid. Equine Vet J 1986;18:220–2. 18. Vandeplassche M, Bouters R, Spincemaille J, Bonte P. Dropsy of the fetal sacs in mares: Induced and spontaneous abortion. Vet Rec 1976;99:67–9. 19. Koterba AM, Haibel GK, Grimmet JB. Respiratory distress in a premature foal secondary to hydrops allantois and placentitis. Comp Contin Educ 1983;7:S121–5. 20. Blanchard TL, Varner DD, Buonanno AM, Palmer JE, Hansen TO, Divers JT. Hydrallantois in two mares. J Equine Vet Sci 1987;7:222–5. 21. Waelchli RO, Ehrensperger F. Two related cases of cerebellar abnormality in equine fetuses associated with hydrops of fetal membranes. Vet Rec 1988;123:513–14. 22. Hanson RR, Todhunter RJ. Herniation of the abdominal wall in pregnant mares. J Am Vet Med Assoc 1986;189:790–3. 23. Williams MA, Wallace SS, Tyler JW, McCall CA, Gutierrez A, Spano JS. Biochemical characteristics of amniotic and allantoic fluid in late gestational mares. Theriogenology 1993;40:1251–7. 24. Castagnetti C, Mariella J, Serrazanetti GP, Grandis A, Medrlo B, Fabbri M, Mari G. Evaluation of lung maturity by amniotic fluid analysis in equine neonate. Theriogenology 2007;67:1455–62. 25. Williams MA, Goyert NA, Goyert GL, Sokol RJ. Preliminary report of transabdominal amniocentesis for the determination of pulmonary maturity in an equine population. Equine Vet J 1988;20:457–8. 26. Jennings W. Some common problems in horse breeding. Cornell Vet 1941;31:197–216. 27. Lofstedt R. Termination of unwanted pregnancy. In: Morrow DA (ed.) Current Therapy in Theriogenology, 2nd edn. Philadephia: Saunders, 1986; pp. 715–18. 28. Threlfall WR. Equine fetal mummification as a result of progestagen therapy. Equine Vet Educ 2005;17:238–9. 29. Gilbert RO, Bosu WTK, Levine SS, Smith DF. Intrauterine death and onset of mummification of a single equine foetus. Equine Vet J 1989;21:301–2. 30. Meyers PJ, Varner DD. Abortion of a mummified fetus associated with short uterine body in a mare. J Am Vet Med Assoc 1991;198:1768–70. 31. Barber JA, Troedsson MH. Mummified fetus in a mare. J Am Vet Med Assoc 1996;208(9):1438–40. 32. McCue PM., Vanderwall DK, Squires EL. Fetal mummification in a mare. J Equine Vet Sci 1997;17:267–9. 33. Threlfall WR. Case Report: Singleton mummified fetus in a Standardbred mare. Equine Vet Educ 2005;17: 235–7. 34. Roberts SJ, Myhre G. A review of twinning in horses and the possible therapeutic value of supplemental progesterone to prevent abortion of equine twin fetuses in the latter half of the gestation period. Cornell.Vet 1983;73:257–64.
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35. Shideler RK. Prenatal lactation associated with twin pregnancy in the mare: A case report. J Equine Vet Sci 1987;7:383–4. 36. Burns TE, Card CE. Fetal maceration and retention of fetal bones in a mare. J Am Vet Med Assoc 2000;217:878–80. 37. Hughes JP, Stabenfeldt GH, Kindahl H, Kennedy PC, Edqvist LE, Neely DP, Schalm OW. Pyometra in the mare. J Reprod Fert Suppl 1979;27:321–9. 38. Jackson PGG. Rupture of the prepubic tendon in a Shire mare. Vet Rec 1982;111:38. 39. Tulleners EP, Fretz P.B. Prosthetic repair of large abdominal wall defects in horses and food animals. J Am Vet Med Assoc 1983;182:258–62. 40. Carleton CL, Threlfall WR. Induction of parturition in the mare. In: Morrow DA (ed.) Current Therapy in Theriogenology, 2nd edn. Philadelphia: Saunders, 1986; pp. 689–92. 41. Townsend HGG, Tabel H, Bristol FM. Induction of parturition in mares: Effect of passive transfer on immunity of foals. J Am Vet Med Assoc 1983;183:255–7. 42. Steel CM, Gibson KT. Colic in the pregnant and periparturient mare. Equine Vet Educ Man 2002;5:5–15. 43. Rossdale PD, Ricketts SW. The Practice of Equine Stud Medicine. Baltimore: Williams & Wilkins, 1974. 44. Vandeplassche M. Obstetrician’s view of the physiology of equine parturition and dystocia. Equine Vet J 1980;12: 45–9. 45. Hura V, Hajurka J, Kacmarik J, Csicsai G, Valocky I. HTML report from UVL, 04101 Kosice, Komensk´eho 73, Slovak Republic, 1997. 46. Davies Morel MC, Newcombe JR, Holland SJ. Factors affecting gestation length in the Thoroughbred mare. Anim Reprod Sci 2002;16:175–85. 47. Hodge S, Kreider J, Potter G, Harms P, Fleeger J. Influence of photoperiod on the pregnant and post partum mare. Am J Vet Res 1982;43:1752–5. 48. Ginther O.J. 1979. Reproductive Biology of the Mare: Basic and Applied Aspects. Cross Plains, WI: Ginther, 1979. 49. Rowlands IW, Weir BJ. Mammals: non primate eutherian. In: Lamming GE (ed.) Marshall’s Physiology of Reproduction, vol. 1. Edinburgh: Churchill Livingstone, 1984; pp. 455– 658.
50. Evans T, Youngquist RS, Loch WE. A comparison of the relative efficacies of domperidone and reserpine in treating equine ‘fescue toxicosis’. Proc Am Assoc Equine Pract 1999;45:207–9. 51. Cross DL, Redmond LM, Strickland JR. Equine fescue toxicosis: signs and solutions. J Anim Sci 1995;73:899–908. 52. Rooney JR. Internal hemorrhage related to gestation in the mare. Cornell Vet 1964;54:11–17. 53. Stowe HD. Effects of age and impending parturition upon serum copper of Thoroughbred mares. J Nutr 1968;95:179–83. 54. Pascoe RR. Rupture of the utero-ovarian or middle uterine artery in the mare at or near parturition. Vet Rec 1979;104:77. 55. Wenzel JGW, Caudle AB, White NA. Treating for uterine intramural hematoma in a horse. Vet Med 1985;80:66–9. 56. Asbury AC The reproductive system. In: Mannsman RA, McAllister ES (eds) Equine Medicine and Surgery, 3rd edn. Santa Barbara, CA: American Veterinary Publications, 1982; pp. 1305–67. 57. Walker DF, Vaughan JT. Bovine and Equine Urogenital Surgery. Philadelphia: Lea & Febiger, 1980. 58. Becht JL, Gordon BJ. Blood and plasma therapy. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia: Saunders, 1987; pp. 317–22. 59. Frazer GS. Differential diagnosis for vaginal haemorrhage in the mare. Equine Vet Educ 2005;17:153–5. 60. Dolente BA. Critical peripartum disease in the mare. Vet Clin N Am Equine Pract 2004;20:151–65. 61. Hance SR, Embertson RM. Colopexy in broodmares. J Am Vet Med Assoc 1992;201:782–7. 62. Rossdale PD. Differential diagnosis of post parturient haemorrhage in the mare. Equine Vet Educ 1994;6:135–6. 63. Santschi EM, Slone DE, Gronwall R, Juzwiak JS, Moll HD. Types of colic and incidence of post colic abortion in pregnant mares: 105 cases (1984–1988). J Am Vet Med Assoc 1991;199:374–7. 64. Boening KJ, Leendertse IP. Review of 115 cases of colic in the pregnant mare. Equine Vet J 1993;25:518–21. 65. Slone DE. Treatment of pregnant mares with colic: practical considerations and concerns. Comp Cont Educ Pract Vet 1993;15:117–20.
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Chapter 254
Maintenance of Pregnancy Angus O. McKinnon and Jonathan F. Pycock
There is little doubt that improved knowledge and techniques such as ultrasonography and hormonal manipulations have improved pregnancy rates; however, there is no clear corresponding drop in pregnancy losses. Part of the reason for lack of progress on pregnancy loss may be management related. The long gestation of the mare (∼ 340 days) hardly gives her time to prepare for the next pregnancy and there is very little selection for fertility at all in horses. The better the performance, the more likely clients are to want pregnancies. In addition, the increasing pregnancy rate per cycle and increased total number of pregnant mares at the end of the breeding season1–4 may mean we obtain pregnancies from mares that have persistent and not appropriately controlled inflammatory conditions of the uterus. If the oocyte is defective or if the embryo has a major chromosome aberration, it is unlikely we can influence the outcome; however, we can modify a poor oviductal or uterine environment with assisted reproductive techniques such as intracytoplasmic sperm injection, or oocyte or embryo transfer. Mares destined to carry their own pregnancies with potential for embryonic or later loss may be managed with some of the techniques presented below. In order to properly evaluate possible therapies to help mares maintain pregnancy, we first need to understand some of the reasons why mares suffer from either early embryonic death (EED) or later pregnancy losses. In general EED is defined as pregnancy loss before the fetal stage (day 40). Pregnancy losses after this period are more difficult to compare between studies as they may be listed as fetal loss, resorption, stillbirth, or abortion.
Early embryonic death Improvement in ultrasonographic equipment permitted initial investigation of early embryonic losses
between days 10 and 20 of gestation.5 This technique, combined with embryo recovery, permits investigation of embryo losses between day 6, which is the first time an embryo can be routinely recovered from the uterus, and day 10–11, which is the first time the vesicle can be consistently detected by ultrasonography. The real incidence of EED prior to day 6 is unknown (see Chapter 239). However, it was suggested that a major proportion of EED in infertile mares occurred in the oviduct.6 In aged subfertile mares the primary problem may actually be the oocyte.7 Increasing age has been associated with poorer pregnancy rates and increased pregnancy loss2 as detected with ultrasonography, and increased losses of oviductal embryos.6, 8 The incidence of EED has been reported to be between 5% and 30% of established pregnancies. 9, 10 In early studies, utilizing ultrasonography, it appeared that EED occurred in mares much earlier than previously reported.5, 11 In a recent large study Allen et al. reported pregnancy losses between day 15 and 42 of pregnancy between 7–8%, and only 6–8% to term.3 Various causes and factors responsible for EED in mares, apart from presence of twins, have been suggested, which included nutrition,12, 13 plant estrogens and photoperiod,14 endophyte-infected tall fescue,15 seminal treatments,16 lactation stress, and foal-heat breeding,17 genital infections,9, 10, 17 chromosomal abnormalities, hormonal deficiencies,13 anabolic steroids,5 stress,13 failure of maternal recognition, and deficiency of pregnant mare serum gonadotropin (eCG or PMSG) production,18 immunological factors,19, 20 and even a higher incidence from some individual stallions.21 Lactating mares and mares bred during foal heat have been reported to have a higher incidence of EED than non-lactating mares.17, 22, 23 However, in a survey of 2562 pregnancies in lactating and non-lactating mares, the incidence of EED was similar.24 Prior to the advent of ultrasonography, recognition and timing of EED was difficult. From data
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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collected over two breeding seasons5 involving 356 mares diagnosed pregnant utilizing ultrasonography, the overall incidence of EED through day 50 post-ovulation was 17.3%. The majority (77.1%) of EED occurred prior to day 35 post-ovulation. During the period 15 to 35 days post-ovulation, a greater (P < 0.05) incidence of EED occurred between days 15 to 20 (26.2%) and 30 to 35 (29.5%) post-ovulation compared to other time periods. Maternal recognition of pregnancy has been reported to occur between 14 and 16 days post-ovulation.25 Embryo development and morphology was useful to predict EED in an embryo transfer program.26 ‘638 embryo transfers conducted over 3 yr were retrospectively examined to determine which factors (recipient, embryo and transfer) significantly influenced pregnancy and embryo loss rates and to determine how rates could be improved. On Day 7 or 8 after ovulation, embryos (fresh or cooled/transported) were transferred by surgical or nonsurgical techniques into recipients ovulating from 5 to 9 d before transfer. At 12 and 50 d of gestation (Day 0 = day of ovulation), pregnancy rates were 65.7% (419 of 638) and 55.5% (354 of 638). Pregnancy rates on Day 50 were significantly higher for recipients that had excellent to good uterine tone or were graded as “acceptable” during a pretransfer examination, usually performed 5 d after ovulation, versus recipients that had fair to poor uterine tone or were graded “marginally acceptable”. Embryonic factors that significantly affected pregnancy rates were morphology grade, diameter and stage of development. The incidence of early embryonic death was 15.5% (65 of 419) from Days 12 to 50. Embryo loss rates were significantly higher in recipients used 7 or 9 d vs 5 or 6 d after ovulation. Embryos with minor morphological changes (Grade 2) resulted in more (P < 0.05) embryo death than embryos with no morphological abnormalities (Grade 1). Between Days 12 and 50, the highest incidence of embryo death occurred during the interval from Days 17 to 25 of gestation. Embryonic vesicles that were imaged with ultrasound during the first pregnancy exam (5 d after transfer) resulted in significantly fewer embryonic deaths than vesicles not imaged until subsequent exams. In the present study, embryo morphology was predictive of the potential for an embryo to result in a viable pregnancy. Delayed development of the embryo upon collection from the donor or delayed development of the embryonic vesicle within the recipient’s uterus was associated with a higher incidence of pregnancy failure. Recipient selection (age, day after ovulation, quality on Day 5) significantly affected pregnancy and embryo loss rates.’26 In addition a higher rate of EED has been reported when insemination is delayed greater than 12 hours27 or greater than 18 hours post ovulation.28 It should be noted that formation of endometrial cups occurs on approximately day 35. EED is diag-
nosed when an embryonic vesicle seen previously is not observed on two consecutive ultrasonographic scans and/or when only remnants of a vesicle are observed. It is strongly recommended that all suspected cases of EED should have two examinations as even experienced veterinarians have made mistakes when the uterus of early pregnant mares (day 20–35) bulges forward at the corpus cornual junction and a pregnancy is missed at the corpus cornual junction even though the entire uterine horn to the body on both sides has been examined without the uterus ever leaving the screen. Ultrasonographic criteria for impending EED are obviously small for gestation length vesicles,29 uterine edema, fluid in the uterine lumen and vesicular fluid that contains echogenic spots (Figure 254.1a–c). EED is suspected, particularly after day 30, when no fetal heartbeat is observed, there is poor definition of fetal structure, fetal fluids are very echogenic, or the largest diameter of the fetal vesicle is two standard deviations smaller than the mean established for that specific day of age. Vesicles increasing in size more slowly than normal may also be characteristic of EED.11 Indications obtained by ultrasonographic scanning of impending loss at later stages include: failure of fixation, an echogenic ring within the vesicle, a mass floating in a collection of fluid and a gradual decrease in volume of placental fluid with disorganization of placental membranes (Figure 254.1d). On occasion EED may be anticipated by identification of endometrial folds in an early pregnant uterus with or without visible fluid. Ultrasonographic scanning, during early pregnancy, is an extremely useful management tool for pregnancy detection and determination of EED. However, if pregnancy rates are not reported until day 50, the discrepancy between pregnancy and foaling rates decreases. In one study the risk of pregnancy loss decreased as gestation increased. ‘3740 mares were diagnosed pregnant by ultrasound around day 22 (± 5) (day 0 being the day of last service) and checked again around day 44 (± 12). Parturition or abortion was recorded for all mares which were still pregnant at the second test. Pregnancy loss between the two early pregnancy diagnoses was 8.9% (n = 3740). This rate was 5.5% (n = 2984) when one normal single pregnancy was diagnosed on the first test. It was increased when uterine endometrial cysts were present (24.4%; n = 86) or when the conceptus looked abnormal (34.8%; n = 279). Following 278 twin pregnancy tests, 9.7% of total resorptions of both twins and 61.5% of unilateral resorption of one of them were observed. Following 113 crushings of one twin conceptus, 20.3% of resorption of the remaining twin conceptus was noted. Abortion rate between day 44 and day 310 was 9.1% (n = 2988). Abortions were more frequent in the case of
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Figure 254.1 (a) Small amount of grade 4 fluid seen around a day 14 pregnancy. This pregnancy developed normally. (b) Large amount of extremely echogenic (grade 1) fluid seen around a day 12 pregnancy. This pregnancy did not survive. (c) A 30-day pregnancy which is a trophoblastic vesicle. No fetus can be detected in this slower growing than normal pregnancy. (d) Disruption of membranes and increased echogenicity is a common sign of EED after day 30.
twins pregnancies (52.8%; n = 53). A higher abortion rate of the remaining twin conceptus was observed after crushing (23.8%; n = 80) or after natural resorption of one of the twins (13.5; n = 148). Early and late pregnancy loss increased with the age of the mare. Resorption rate, but not abortion rate, increased with parity. Breed had no effect on pregnancy loss. Reproductive status was not linked with embryonic death frequencies, but abortion rate increased if the mare became pregnant during foal heat. Similar resorption rates were observed in 261 different stallion “harems” (groups of mares served by the same stallion). Similarly, the rate of abortion was not linked to the stallion (192 groups of mares). The day by day risk of pregnancy loss decreased steadily as pregnancy progressed.’30 In this later study the higher than expected pregnancy loss from mares with twin pregnancies that either were assisted in or spontaneously eliminated one pregnancy was surprising and does not parallel our experiences.
Abortion Abortion can be infectious or non-infectious and is discussed under specific causes in various chapters in the text; there is also an overview of abortion (see Chapter 240). Many studies have been reported which implicate the relative importance of different causes. Unfortunately there are still many gaps in our knowledge, and one of the common pathological report is ‘unable to determine the cause’. This may on occasion be the result of not providing adequate or suitable material in a sterile or even timely manner to the laboratory. ‘A total of 309 miscarried equine fetuses and foals were submitted for necropsy at the Institut de pathologie du cheval between May 1st, 1986 and April 30th, 1990. An infectious origin could be established in 34.6% of the cases, of which 79.0% were of bacterial aetiology and 21.0% of viral aetiology. Of the bacteria, 26.1% of the isolates were streptococci and 19.3% were Escherichia
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coli. The only virus involved was the equine rhinopneumonitis virus.’31 ‘Pathology case records of 3,514 aborted fetuses, stillborn foals, or foals that died < 24 hours after birth and of 13 placentas from mares whose foals were weak or unthrifty at birth were reviewed to determine the cause of abortion, death, or illness. Fetoplacental infection caused by bacteria (n = 628), equine herpesvirus (143), fungi (61), or placentitis (351), in which an etiologic agent could not be defined, was the most common diagnosis. Complications of birth, including neonatal asphyxia, dystocia, or trauma, were the second most common cause of mortality and were diagnosed in 19% of the cases (679). Other common diagnoses were placental edema or premature separation of placenta (249), development of twins (221), contracted foal syndrome (188), other congenital anomalies (160), and umbilical cord abnormalities (121). Less common conditions were placental villous atrophy or body pregnancy (81), fetal diarrhea syndrome (34), and neoplasms or miscellaneous conditions (26). A diagnosis was not established in 16% of the cases seen (585). The study revealed that leptospirosis (78) was an important cause of bacterial abortion in mares, and that infection by a nocardioform actinomycete (45) was an important cause of chronic placentitis.’32 ‘Pathologic and microbiologic examinations were performed on 1,211 aborted equine fetuses, stillborn foals, and placentas from premature foals in central Kentucky during the 1988 and 1989 foaling seasons to determine the causes of reproductive loss in the mare. Placentitis (19.4%) and dystocia-perinatal asphyxia (19.5%) were the 2 most important causes of equine reproductive loss. The other causes (in decreasing order) were contracted foal syndrome and other congenital anomalies (8.5%), twinning (6.1%), improper separation of placenta (4.7%), torsion of umbilical cord (4.5%), placental edema (4.3%), equine herpesvirus abortion (3.3%), bacteremia (3.2%), fetal diarrhea (2.7%), other placental disorders (total of 6.0%), and miscellaneous causes (1.6%). A definitive diagnosis was not established in 16.9% of the cases submitted. Streptococcus zooepidemicus, Escherichia coli, Leptospira spp., and a nocardioform actinomycete were organisms most frequently associated with bacterial placentitis, and Aspergillus spp. was the fungus most often noted in mycotic placentitis. No viral placentitis was noticed in this series. Dystocia-perinatal asphyxia was mostly associated with large foals, maiden mares, unattended deliveries, and malpresentations. The results of this study indicate that in central Kentucky, the non infectious causes of equine reproductive loss outnumber the infectious causes by an approximate ratio of 2:1, placental disorders are slightly more prevalent than non-placental disorders, Leptospira spp. and a nocardioform actinomycete are 2 new important abortifacient bacteria in the mare, the occurrence of contracted foal syndrome is unusually frequent, the incidence of twin abortion has
sharply declined, and torsion of the umbilical cord is an important cause of abortion in the mare.’33 A survey of equine abortion, stillbirth, and neonatal death in the UK from 1988 until 1997 was reported by Smith:34 ‘A detailed review of laboratory records for equine abortion is fundamental in establishing current disease trends and suggesting problems important for further research. OBJECTIVES: To review the causes of abortion and neonatal death in equine diagnostic submissions to the Animal Health Trust over a 10 year period. METHODS: The diagnoses in 1252 equine fetuses and neonatal foals were reviewed and analysed into categories. RESULTS: Problems associated with the umbilical cord, comprising umbilical cord torsion and the long cord/cervical pole ischaemia disorder, were the most common diagnoses (38.8%: 35.7% umbilical cord torsion and 3.1% long cord/cervical pole ischaemia disorder). Other noninfective causes of abortion or neonatal death included twinning (6.0%), intrapartum stillbirth (13.7%) and placentitis, associated with infection (9.8%). E. coli and Streptococcus zooepidemicus were the most common bacteria isolated. Neonatal infections not associated with placentitis accounted for 3.2% of incidents; and infections with EHV-1 or EHV4 for 6.5%. CONCLUSIONS: Definitive diagnosis of equine abortion is possible in the majority of cases where the whole fetus and placenta are submitted for examination. Potential relevance: Given the high incidence of umbilical cord torsion and related problems as causes of abortion in UK broodmares, more research on factors determining umbilical cord length and risk of torsion is essential.’34 It is interesting to note the bias of region on diagnosis. In the UK, umbilical torsion appears to be a major cause of pregnancy loss, whilst in the USA, nocardioform placentitis appears to be increasing in frequency. ‘Observations on 2,000 pregnancies revealed that 175 (8.7%) equine abortions occurred over a 4-year period. The peaks of abortion were at mid-gestation (20–32 weeks) and at late gestation (37–44 weeks). Abortion was higher in mares bred to donkey (10.4%) than in those bred to horse stallion (7.8%). Maximum abortions (80%) occurred in winter. Out of 175 equine abortions, 102 (58.3%) were due to infectious (bacterial, viral, fungal) causes, while 41.7% were due to non infectious causes, viz. placentitis, malpresentation, twin pregnancy, foetal abnormality, toxaemia and trauma. Sixty-one abortions were grouped as due to unspecified non infectious causes.’35 Severe stress, such as colic, would appear to play a major part in late term pregnancy loss.
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Maintenance of Pregnancy ‘The records of 105 pregnant mares and 105 nonpregnant horses with colic admitted to an equine hospital were reviewed. The 2 groups had similar types of colic and short-term survivability. Of the 105 pregnant mares, 31 were treated medically and 74 required surgical intervention. Thirty-three of the 105 mares died or were euthanatized. Thirteen (18%) of the 72 remaining mares aborted. Of 4 mares with severe medical cases, 2 died, 1 aborted, and 1 aborted and died. Of 27 horses with medical cases that required less intensive treatment, none died and 2 aborted. Of the 74 horses that required surgery, 45 survived to termination of pregnancy (foaling or abortion); 36 of these mares (80%) had a live foal. The type of surgical lesion had no effect on pregnancy outcome. Stage of gestation at initial examination, duration of anesthesia, or intraoperative hypoxia or hypotension had no effect on pregnancy outcome. However, when hypoxia occurred during colic surgery in the last 60 days of pregnancy, the mares either aborted or delivered severely compromised foals that did not survive.’36 Stresses early in pregnancy associated with transport37 or rectal palpation for pregnancy diagnosis38 apparently have little or no effect on fetal loss. ‘Incidence of early embryonic death (EED) and associated changes in serum cortisol, progesterone and plasma ascorbic acid (AA) in transported mares were investigated. Mares were transported for 472 km (9 h) during either d 16 to 22 (T-3 wk, n = 15) or d 32 to 38 (T-5 wk, n = 15) of gestation. Blood samples were drawn from control, non-transported mares (NT-3 wk, NT5wk, n = 24) and transported mares pre-trip, midtrip, and at 0, 12, 24, 48 and 72 h post-transport and daily for the next 2 wk. Incidence of EED between transported and non-transported mares was not different (P > .05). Serum cortisol in all transported mares increased (P < .05) relative to pre-trip values at midtrip and 0 h post-transport. Relative to NT mares, serum cortisol was higher (P < .05) at midtrip in T-3 wk mares and 0 h post- transport in T-5 wk mares. Serum progesterone in all T mares increased (P less than.05) at midtrip relative to pre-trip values and was higher (P < .05) in T-3 wk mares than in NT-3 wk mares at midtrip and 0 h post-transport. Post-transport decreases (P < .05) in concentrations of progesterone were observed in mares that aborted. Plasma AA in transported mares increased (P < .05) at midtrip in T-5 wk mares and decreased (P < .05) relative to pre-trip values at 24 and 48 h post-transport (T-3 wk and T-5 wk mares, respectively.’37 Placentitis There is little doubt that placentitis is a disease more commonly diagnosed in intensively managed broodmare farms39 (see Chapter 242). It is possible
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that our sophisticated management procedures have led to more mares becoming pregnant that, without intense management, might have struggled to become pregnant, thus creating a group of mares either susceptible to, or more likely to be affected by, placentitis. Placentitis may be recognized by vulval discharge, premature lactation, or spontaneous abortion. Both transrectal and transabdominal ultrasonography have been useful in examining the placenta at the region of the cervical star (Figure 254.2a–c). Thickness of the uterus and placenta (CTUP) can be measured, with values > 12 mm considered abnormal.40–44 Other ultrasonographic findings are separation of the placenta, particularly at the cervical star, by pus, mucus or blood and, on occasion, air. ‘Feto-placental infection caused by microorganisms was the most common cause of death of fetuses, stillborn foals, and foals that died within 24 hrs after birth in central Kentucky. Pathologically, three types of placentitis were seen: ascending, diffuse, and focal mucoid. The pathogenesis in each form is believed to be different and each is associated with certain types of causative bacteria or fungi. Ascending placentitis was the most common type of placentitis prior to the 1998 foaling season in Kentucky. This form is thought to be the result of microorganisms gaining access to the cervical portion of the placenta during gestation by spread from the lower reproductive tract through the cervix. Streptococci and E. coli were the most commonly isolated bacteria. Diffuse or multifocal placentitis was less commonly diagnosed and is associated with hematogenous spread of microorganisms to the uterus of the mare with subsequent infection of the placenta. This form was associated with infection by bacteria in the genera Leptospira, Salmonella, Histoplasma, and Candida. Focal mucoid placentitis is also known as nocardioform placentitis. Nocardioform placentitis has emerged as the most commonly diagnosed type of placentitis over the last two foaling seasons and is characterized by unique pathology and distinct bacteria. At present the pathogenesis of this form is unknown. Diagnosis of placentitis during gestation is often difficult. Most mares show no outward signs of infection. Some mares will undergo premature mammary development with lactation and, occasionally, a vaginal discharge is present. Transrectal and transabdominal ultrasound examinations are useful in arriving at a diagnosis if placentitis is suspected. Various treatment modalities have been utilized in an attempt to maintain gestation for as long as possible to enhance foal viability. Placentitis results in several outcomes. In addition to abortions and stillbirths, the mare may produce small weak foals or normal foals. The small, weak neonates represent a special management and medical challenge. These individuals have an increased risk of sepsis and orthopedic problems requiring extensive care.’45
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(c) Figure 254.2 (a) Note extreme size of the combined thickness of the placenta and uterus (CTUP) between arrows in this transrectal ultrasonographic image. (b) Note extreme size of the CTUP in this transabdominal ultrasonographic image. (c) Air has penetrated through the cervix and around the placenta, making identification of the CTUP difficult.
‘Placentitis is a common cause of equine abortions. In a majority of cases, the route of infection is believed to be ascending through the cervix, and an area of the chorion adjacent to the cervical star shows characteristic pathology in aborting mares. This area is depleted of chorionic villi, thickened, discolored, and covered by fibronecrotic exudate. Mares with placentitis due to a hematogenous infection show multifocal pathology of the chorionic surface of the placenta. Clinical signs include udder development, premature lactation, cervical softening, and vaginal discharge. Treatment is often unsuccessful once the mare has developed clinical signs. Ultrasonographic evaluation of the placenta in late gestational mares allows the clinician to detect preclinical signs of placentitis and premature separation, which then could be treated in its early stages. Transabdominal ultrasonography: Focal areas of uteroplacental thickening and partial separation of the allantochorion from the endometrium in the uterine body and horns, can be observed by this approach. The com-
bined thickness of the uterus and the placenta (CTUP) should not be more than 12 mm at any site. In addition to evaluating the placenta a biophysical profile of the fetus can be performed by a transabdominal approach. Transrectal ultrasonography: This approach provides an excellent image of the caudal portion of the allantochorion adjacent to the cervical star. Using transrectal ultrasonographic evaluation of the placenta, we have observed abnormal thickness and partial separation of the allantochorion from the endometrium in mares with clinical signs of placentitis. In advanced stages, the space between the uterus and the placenta is filled with hyperechoic fluid. Normal values of the CTUP in an area immediately cranially and ventrally of the cervix was determined to be: < 8 mm between day 271 and 300; < 10 mm between day 301 and 330; and < 12 mm after day 330. Increased CTUP suggests placental failure and pending abortion. Treatment of placentitis should be aimed toward elimination of the infectious agents, reduction of the inflammatory
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Maintenance of Pregnancy response, and reduction of the increased myometrial contractility in response to the ongoing inflammation. Broad spectrum antibiotics, anti- inflammatories (flunixin meglumine, 1.1 mg/kg BID; phenylbutazone, 4 mg/kg BID) and tocolytics (Altrenogest; Clenbuterol), are recommended for treatment of placentitis. Pentoxyfylline (7.5 mg/kg p.o. BID) has been suggested to increase oxygenation of the placenta through an increased deformability of red blood cells.’46 The approach of treating mares with placentitis at our practices is to begin with mares on broad-spectrum antibiotics at first recognition. Commonly we use trimethoprim sulpha combinations or penicillin/gentamicin. Mares are supplemented with long-acting altrenogest or progesterone preparations (compounded) as well as non-steroidal antiinflammatory agents. Other treatments that may be considered are tocolytics such as clenbuterol or use of pentoxyfylline to increase oxygenation of the placenta.4 Duration and intensity of treatment are dictated by clinical signs (lactation, fetal heart rate, and placental thickness measured at the cervix and midventral uterine body). Identification of a fetal heart rate under 65 within two months of foaling is usually associated with a very poor prognosis, and owners’ consent to continuing treatment is requested.
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many aspects of the epidemiology, clinical presentations, and manifestations of MRLS. It allows quantitative interpretation of experimental and field MRLS data and has implications for the basic mechanisms underlying MRLS. The results support suggestions that MRLS is caused by exposure to or ingestion of ETCs. The results also show that high levels of ETC exposure produce intense, focused outbreaks of MRLS, closely linked in time and place to dispersing ETCs, as occurred in central Kentucky in 2001. With less intense exposure, lag time is longer and abortions tend to spread out over time and may occur out of phase with ETC exposure, obscuring both diagnosis of this syndrome and the role of the caterpillars.’47 Equine amnionitis and fetal loss (EAFL) appears to be a similar syndrome reported in Australia.48 From the preceding abstract quotations it appears that infectious causes of abortion, twins, umbilical cord torsion, and stress constitute a major proportion of reported fetal demise late in pregnancy, and many poorly defined factors such as uterine environment and the hormonal milieu are involved earlier in pregnancy. A background on endocrinology of the mare during pregnancy may be useful in providing rational approaches to therapy (see Chapter 229).
Endocrinology of pregnancy Mare reproductive loss syndrome A description of mare reproductive loss syndrome (MRLS) is contained in Chapter 248, and the steps taken to elucidate the etiology of MRLS are described below. ‘During 2001, central Kentucky experienced acute transient epidemics of early and late fetal losses, pericarditis, and unilateral endophthalmitis, collectively referred to as mare reproductive loss syndrome (MRLS). A toxicokinetic/statistical analysis of experimental and field MRLS data was conducted using accelerated failure time (AFT) analysis of abortions following administration of Eastern tent caterpillars (ETCs; 100 or 50 g/day or 100 g of irradiated caterpillars/day) to late-term pregnant mares. In addition, 2001 late-term fetal loss field data were used in the analysis. Experimental data were fitted by AFT analysis at a high (P < .0001) significance. Times to first abortion (“lag time”) and abortion rates were dose dependent. Lag times decreased and abortion rates increased exponentially with dose. Calculated dose X response data curves allow interpretation of abortion data in terms of “intubated ETC equivalents.” Analysis suggested that field exposure to ETCs in 2001 in central Kentucky commenced on approximately April 27, was initially equivalent to approximately 5 g of intubated ETCs/day, and increased to approximately 30 g/day at the outbreak peak. This analysis accounts for
In mares, maternal recognition of pregnancy associated with production of an immunosuppressive agent, a pregnancy-specific protein called early pregnancy factor, may be as early as 48 hours after conception. Early pregnancy factor has been detected in mice, sheep, human beings, and mares (see Chapter 231) and may, in the future, aid in detection of pregnancy and EED. The second time for maternal recognition of pregnancy probably arises on or before day 5 or 6 after ovulation, as fertilized ova are transported from the oviduct through the utero-tubal junction into the uterus by 5 or 6 days after ovulation, but unfertilized oocytes generally are retained in the oviducts.49–52 The third recognizable time for maternal recognition (or reinforcement) of pregnancy occurs between 12 and 16 days. During this time, the conceptus is extremely mobile and probably secretes an antiluteolytic substance that prevents release of PGF2α , thus maintaining the corpus luteum (CL) and preventing the mare from returning to estrus. ‘Two experiments were performed to determine the critical time at which the equine blastocyst must be present within the uterus of the mare to prevent regression of the corpus luteum, and thus establish the critical time for the maternal recognition of pregnancy. A non-surgical blastocyst collection technique was developed to study
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this relationship between the blastocyst and the maternal ovary. Results from these experiments demonstrated that the cyclic life-span of the corpus luteum is not affected by the presence of the blastocyst within the mare’s uterus until after Day 14 after ovulation. Luteal function was prolonged when blastocysts were removed on Day 15 or later. The critical period for the maternal recognition of pregnancy in the Pony mare appears to be confined to the period between Days 14 and 16 after ovulation.’53 ‘Two experiments were conducted to assess the effect of exogenous hormone treatment on uterine luminal prostaglandin F (PGF). In the first experiment ovariectomized pony mares received either corn oil (21 days, n = 3), estradiol valerate (21 days, n = 3), progesterone (21 days, n = 3) or estradiol valerate (7 days) followed by progesterone (14 days, n = 4). Progesterone treated mares had higher (P < .01) uterine luminal PGF compared with all other groups, and no differences were detected between other treatment comparisons. In Experiment II, uterine fluid was collected from 4 ovariectomized horse mares before and after treatment with estradiol valerate (7 days) followed by progesterone (50 days). Pre-treatment uterine luminal PGF levels were lower (P < .001) than post-treatment levels (.03 vs 76.80 ng/ml). In a third experiment PGF was measured in uterine fluid of pony mares on days 8, 12, 14, 16, 18 and 20 of the estrous cycle and pregnancy. In non pregnant mares a day effect (P less than.03) was observed in which uterine fluid PGF increased during the late luteal phase and declined thereafter. In contrast, no day effect was observed in pregnant animals and uterine luminal PGF was lower (P < .001) than in cycling animals. These studies indicate that exogenous progesterone administration results in a large increase in uterine luminal PGF, whereas, pregnancy results in suppression. Taken collectively with previous work from our laboratory, these results suggest that while the endometrium of pregnant mares is capable of producing large amounts of PGF, the presence of a conceptus impedes its synthesis and/or release which allows for luteal maintenance.’54 ‘Comparisons of estrone, 17 beta-estradiol, and plasma progestin concentrations were made in uterine fluid and peripheral blood of non pregnant and pregnant pony mares. Concentrations of these steroids were also measured within yolk sac fluid from blastocysts on days 12, 14, 16, and 18 of pregnancy to obtain more complete analyses of the uterine environment (uterine fluid plus yolk sac fluid) of early pregnancy. Thirty mares were randomly assigned to six treatment groups (n = 5/group), and uterine fluid and peripheral blood samples were obtained on days 8, 12, 14, 16, 18, and 20 postovulation. After a recovery period of one estrous cycle, mares were bred at their next estrus. Animals were hysterectomized on the same treatment day to which they had previously been assigned in the non pregnant phase of this study. Using this design, uterine fluid and pe-
ripheral blood samples were collected from each mare on equivalent days of the estrous cycle and pregnancy. Significant differences in day trends were found between non pregnant and pregnant animals for estrogens and progestins in both uterine fluid and peripheral plasma. Furthermore, these data demonstrate that large increases in estrogens occur after day 12 of pregnancy in uterine and yolk sac fluids, with estrone becoming the predominant estrogen by days 18 and 20 in yolk sac and uterine fluids, respectively. These changes were not detected in peripheral plasma, which indicates that changes occurring within the uterine environment are not discernible in the systemic circulation during early pregnancy. These results indicate that the large amounts of estrogens appearing in uterine fluids during early pregnancy are of conceptus origin and may be an important factor in regulating the environment in which the conceptus develops.’55 The next piece of the puzzle was also provided by researchers in Florida:25 ‘Cycling pony mares were bred and used to test the effect of restricted conceptus mobility on luteal maintenance (i.e. maternal recognition of pregnancy). In Experiment 1, uterine horns were ligated to restrict conceptus mobility to one uterine horn, Group 1; one horn plus the uterine body, Group 2; or one horn, the body and approximately 80% of the second horn, Group 3. Pregnancies were monitored with real-time ultrasonography. Four of five mares in Group 1 and two of four mares in Group 2 returned to estrus (Day 16.0 ± 1.9 and 14.5 ± 0.7, respectively) and subsequently lost the embryonic vesicles (Day 17.2 ± 1.2 and 15.7 ± 0.7, respectively). None of the four mares in Group 3 lost the vesicles. There was a significant effect of the interaction of treatment (amount of uterus available to the conceptus) and day on plasma progesterone (P) concentration (P < 0.005). In Experiment 2, conceptus mobility was restricted to one uterine horn in two groups of mares, of which the second was treated with the synthetic progestin, Regumate (allyl trenbolone). In the first group, each of three mares lost the vesicle (Day 17.3 ± 4.3). In the second group, four of five mares maintained the pregnancies, indicating that pregnancy failure was due to the effects of declining P. These data indicate that restricted conceptus mobility results in luteolysis in the mare, and that the subsequent decline in P leads to embryonic death. This supports the notion that unrestricted mobility of the equine conceptus, allowing it to interact with most the uterine endometrium, is necessary for luteal maintenance and conceptus survival.’25 These studies have important ramifications on effects of different uterine environment on ability of mares to maintain pregnancy, for example, in cases of uterine cysts that impair mobility, and the possible therapeutic implications of progestagens.
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Maintenance of Pregnancy Estrogens A minor elevation of maternal (ovarian) production of estrogen occurs between days 35 and 60 but has little diagnostic significance. A major increase in plasma estrogen concentration occurs after day 60.56 This increase comes from the fetal placental unit and, thus, is not affected by ovariectomy. ‘Plasma total (conjugated + unconjugated) oestrogens were measured from Day 0 to 100 of pregnancy and compared with the levels found during the oestrous cycle. From Day 0 to 35 of gestation, the concentrations were similar to those during dioestrus. An increase in total oestrogens between Days 35 and 40 was followed by a plateau of 3 ng/ml between Days 40 and 60 which was slightly higher than preovulatory concentrations. This first increase in total oestrogen level was produced by the ovaries since values were suppressed after ovariectomy; stimulation may be due indirectly to PMSG causing follicular growth. After Day 60, a second increase was detected and considered to be of feto-placental origin as the levels at this time were not suppressed after ovariectomy. By Day 85, oestrogen concentrations exceeded those detected in non-pregnant mares so that a direct measurement of total oestrogens in plasma by a simplified radioimmunoassay after Day 85 of gestation offers a reliable method of pregnancy diagnosis.’56 Concentrations of estrogen in urine and serum peak at about day 210 of gestation57 and are thought to come from fetal gonads, which are enlarged at this time. If fetal gonads are removed, plasma estrogen concentrations drop immediately to very low levels.58 In general, estrogen detection in serum or urine has only limited value for pregnancy diagnosis since estrogen measurements become reliable only relatively late in pregnancy. Earlier it was not recommended to test for estrogens before 150 days of pregnancy, but more recently, measurement of estrone sulfate (conjugated) for diagnosis of pregnancy and evidence of a viable conceptus has been advocated after day 60 of pregnancy.59 Fetal death results in an immediate decrease in estrone sulfate concentration.59 If a mare in the second or third trimester of pregnancy begins to lactate, measurement of estrone sulfate may provide an index of fetal viability. This may be particularly useful when other tests of fetal viability such as ultrasonography and/or electrocardiographic monitoring are not available. Biological, chemical, and immunological tests are available for estrogen determination. Biological tests such as increased nipple length in guinea pigs or induction of the vaginal cytological findings typical of estrus in immature or ovariectomized mice rarely are used. They have been replaced by chemical tests, largely because of cost and time. Chemical
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tests for urinary estrogen rely on color changes and fluorescence in the presence of warm, concentrated sulfuric acid. Radioimmunoassay can be used for determination of plasma total estrogen (conjugated plus unconjugated)56 concentration or urinary estrone conjugates. The method described for plasma analysis56 requires enzymatic hydrolysis and extraction, and may not reveal all estrone conjugates because the concentration of estrone sulfate is 1000 times higher in the urine. Direct conjugated estrogen measurement on serum or milk has been described,60 and differentiation between pregnancy and estrus was possible after approximately 50 days. In mares, an enzymatic determination of unconjugated estrogens in the feces for pregnancy diagnosis has been described.61 It was possible to confirm pregnancy after 120 days of gestation, and because this technique may be totally noninvasive to the mare, some benefits to the equine reproductive industry may be realized. Recently ‘Immunoreactive urinary oestrogen conjugates were assessed in daily urine samples (∼ 5 samples/week) collected from 8 Przewalski’s mares maintained under semi free-ranging pasture conditions. The relative percentage contributions of immunoreactive urinary oestrogens during different reproductive states (oestrus, luteal phase, early, mid- and late gestation) were determined using high-pressure liquid chromatography. In general, conjugated forms of oestrone (oestrone sulphate and oestrone glucuronide) were the major excreted immunoreactive oestrogens in non-pregnant and pregnant Przewalski’s mares. Variations in urinary oestrogen conjugates indicated that the onset of oestrous cyclicity coincided with increasing daylengths, and the non-conception oestrous cycle was 24.1 ± 0.7 days (n = 17) in duration. Most copulations (29/35, 82.9%) were observed between Day −4 and Day +1 from the preovulatory oestrogen conjugates peak (Day 0). Based on known copulation dates, the mean gestation length was 48.6 ± 0.4 weeks (range 47.3–50.3 weeks). During pregnancy, urinary excretion of oestrogen conjugates increased apprx 300-fold over levels in non-pregnant mares, reaching peak concentrations by Week +24 (51% of gestation). These results demonstrate that longitudinal reproductive events, including oestrous cyclicity and pregnancy, can be monitored precisely by evaluating urinary oestrogen conjugates in samples from Przewalski’s mares maintained under semi-free-ranging conditions.’62 Further studies on uses of estrogen analysis are presented below. ‘Serum and urinary estrone sulfate concentrations were determined in 7 pregnant mares before and after prostaglandin-induced abortion (n = 4) or surgical removal of the fetus (n = 3) to determine the source of
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estrogen during early pregnancy (gestation days [GD] 44 to 89). Estrone sulfate concentrations also were determined in serum samples (stored frozen for 2 years) from 3 mares that had been ovariectomized between GD 51 and 58. Estrone sulfate concentrations decreased in serum and urine after expulsion or removal of the fetus (urinary patterns were more definitive than were patterns for serum), whereas a transient decrease in serum estrone sulfate concentration was observed after ovariectomy. Seemingly, products of conception are the major source of estrone sulfate during early pregnancy, although there appears to be some ovarian contribution. Serum or urinary estrone sulfate measurements provide a simple and accurate test for fetal viability after GD 44 in the mare.’63 ‘The removal of one of twin embryos was attempted by infusion of 24% (w/v) saline into the gestation sac in 2 mares by laparotomy. The treatment was successful in one mare (Case 1) and the untreated embryo remained viable. However, neither foetus survived in the second mare (Case 2). Plasma oestrone sulphate (E1S) concentrations fell immediately after treatment in both mares but recovered to approximately 50% of pre-treatment levels in Case 1. In Case 2 plasma E1S concentrations declined steadily and were< 1 ng/ml within 6 days of treatment. These preliminary results suggest that the method may be useful for selective removal of one of twin embryos in mares. Furthermore, plasma E1S concentrations may be a useful indicator of embryonic viability.’64 ‘Two mares received PGF-2 alpha twice daily until abortion and 2 mares received a combined treatment with oestradiol benzoate and oxytocin. The mares were about 150 days pregnant. The PG-treated animals aborted after 37 and 61 h, respectively, and the fetuses were expelled in intact fetal membranes. The other 2 mares aborted 13 and 27 h after the first oxytocin injection, respectively, and showed strong uterine contractions and expelled the fetuses in disrupted fetal membranes. Concentrations of 15-ketodihydro-PGF-2 alpha increased both after PG and oxytocin injections and in association with the abortion, but after the PG-induced abortion there was an immediate return to basal levels and after the oxytocin-induced abortion there was a large increase in the concentrations, indicating damage of the uterus. Progesterone and relaxin concentrations followed the placental function and decreased in association with the abortions. Oestrone sulphate values differed in the two groups; the oxytocin-treated animals showed a rapid decrease while the mares treated with PG showed first a marked increase and then a decrease. Concentrations of PMSG appeared to be unaffected by the abortions.’65 ‘We evaluated 2 rapid non-instrumented field tests for pregnancy on feral horses (Equus caballus) because previous techniques required sophisticated and expensive instrumentation that limited their usefulness to field researchers. The measurement of urinary estrone conjugates (E-1C) by an enzyme immunoassay and
based on observable color changes was 100% accurate when compared with instrumented spectrophotometrically measured tests for E-1C. A non-instrumented “dipstick” enzyme immunoassay for equine chorionic gonadotropin-like (eCG) molecules was 83% accurate in diagnosing mare pregnancies when compared with the results of the instrumented test for E-1C. The decrease in accuracy using the non-instrumented eCG test resulted from a time period of 40–140 days when eCG was measurable, whereas E-1C elevation were measurable after day 35 of a mare’s pregnancy. Our results indicate that it is possible to detect pregnancy in free-roaming horses under field conditions and without instrumented assays; such field tests provide opportunities to study fecundity and fetal loss in a variety of free-roaming animals.’66 ‘Plasma cortisol, oestrone sulphate and progestagens were measured in 22 stressed pregnant mares (gestation length 17–336 days) as indicators of fetal viability. Mares were bled every 12 h from time of admission, and plasma was stored at –70 degrees C until assayed. Four normal mares were bled twice weekly from Day 270 to parturition to provide baseline endocrine data. Cortisol and progestagen concentrations were measured by radioimmunoassay and oestrone sulphate was measured by enzyme immunoassay. Mares were grouped according to clinical diagnosis: surgical colic (Group 1, n = 11), medical colic (Group 2, n = 7), and uterine torsion (Group 3, n = 4). Of the 16 mares in Groups 1 and 2 that survived to discharge, 12 mares foaled normally and 4 aborted, 3 during hospitalization. Following surgical treatment of uterine torsion, 2 mares aborted and 2 mares carried foals to term. Plasma cortisol was greater than 30 ng/ml in 19 of the 22 stressed mares at presentation and was less than 30 ng/ml in normal mares at all collections. Cortisol concentrations remained elevated in mares during post admission complications. The mean cortisol concentration of mares with colic that subsequently aborted was higher at presentation, but not statistically different, than levels of mares that did not abort (135 ± 35 ng/ml and 83 ± 19 ng/ml, respectively; mean ± s.e.m.). Progestagen concentrations in normal mares ranged from 2–25 ng/ml. Progesterone concentrations in subject mares that subsequently foaled did not differ from normal mares, whereas concentrations in aborting mares (n = 5) either dropped rapidly (n = 3) or were low at the time of admission and decreased to < 2 ng/ml (n = 2). Oestrone sulphate concentrations that ranged from 7–5000 ng/ml, decreased after foaling or abortion. However, the concentrations remained above 38 ng/ml (range, 38–480 ng/ml) in those mares (n = 7) sampled post foaling or post abortion. Our results show that mares with colic that subsequently aborted had higher mean plasma cortisol concentrations at admission than mares that carried foals until term. No single sample of the 3 hormones measured was a reliable indicator of abortion. Oestrone sulphate was not an accurate indicator of foetal viability during late pregnancy.’67
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Maintenance of Pregnancy Equine chorionic gonadotropin Equine chorionic gonadotropin (eCG; also referred to as pregnant mare serum gonadotropin – PMSG) is produced by fetal trophoblast cells that invade the maternal endometrium on days 35 through 38.68 These cells form ‘endometrial cups’ that are white, raised plaques that increase in size until day 60 of gestation. A maternal immunological rejection against the endometrial cups results in a decrease in their size beginning around day 70 and continuing through day 120 when they are no longer functional. Plasma concentrations of eCG parallel the growth and regression of endometrial cups and are first detected from days 35 through 40, peak between days 60 and 65, and decrease to low or non-detectable values between 120 and 150 days of gestation. The exact function of eCG is undetermined and is the source of much controversy. Current popular theory is that eCG is responsible for a luteinizing hormone-like activity that causes either ovulation and(or) luteinization of follicles resulting in formation of secondary or accessory CL formation respectively. Serum analysis for eCG should be performed between days 40 and 120 of gestation. When abortion occurs after day 35, endometrial cups may persist for a time, which means that: (i) eCG is still detectable and may be responsible for a false-positive pregnancy diagnosis, and (ii) these mares commonly fail to return to estrus. In addition, when mares are carrying a mule fetus, the immunological rejection of the endometrial cups occurs much earlier, and extremely low or nondetectable concentrations of ECG exist despite the persistence of pregnancy. An interesting observation is the apparent abortigenic ability of hCG when given to early (< 35 day) pregnancies.69 It is interesting to speculate on the interactions of these gonadotropins, especially in light of evidence that suggests that not only are the secondary corpora lutea forming in association with eCG, but that the primary CL of pregnancy has a resurgence that is temporally associated with the beginning of eCG production.70 ‘The temporal association between the first detectable increase in concentrations of circulating progesterone and equine chorionic gonadotropin (eCG) was studied in pregnant mares. Blood samples were collected on days 15, 20 and 25–40 from pony mares (experiment 1; n = 11) and days 11–40 from horse mares (experiment 2; n = 10). Mares were ultrasonically examined daily to evaluate the primary corpus luteum (experiment 3; n = 2–6) and to detect the formation of secondary corpora lutea (experiments 1 and 2). Formation of secondary corpora lutea was not detected in any mare prior to day 40 in experiment 1, but was detected in one mare on day 38 and in two mares on day 39 in experiment 2. In both experiments 1 and 2, there was a significant main effect of day
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for circulating concentrations of progesterone and was attributable to decreased (P < 0.05) concentration between days 12 and 15 (experiment 2) and increased concentrations after day 32 (experiment 1) or day 34 (experiment 2). Progesterone concentrations were higher (P < 0.05) on days 38, 39, and 40 than on days 29, 31, and 32 in experiment 1 and higher (P < 0.05) on day 40 than on days 32, 33 and 34 in experiment 2. When progesterone concentration in experiment 2 were normalized to the day that eCG was first detected (mean, day 35), mean concentrations were lower (P < 0.05) before detection of eCG than on the second day after detection. The cross-sectional area of the ultrasonic image of the primary corpus luteum (experiment 3) was greater (P < 0.05) on days 39, 40, 41, and 42 than on days 30 and 33. The cross-sectional increases in area of the primary corpus luteum in experiment 3, closely paralleled the increases in progesterone concentration in experiments 1 and 2. Combined results of the 3 experiments supported the hypothesis that resurgence of primary corpus luteum occurs in pregnant mares between days 30 and 40 and that the resurgence is a response to release of eCG into the circulation. More specifically, the results indicated that, on the average, the resurgence of the primary corpus luteum in the mare began on day 35.’70 Biological tests rely on the effects of eCG obtained from mare serum, upon the reproductive tract of laboratory animals, for example, stimulation of rabbit ovaries (Friedman test), rat uterus, mouse uterus, and production of spermatozoa from male toads. A practical, inexpensive bioassay involves the injection of 0.5 to 1 mL of mare serum, subcutaneously into immature, 21-day-old female mice. Saline-injected mice serve as controls. The presence of eCG will cause enlargement and congestion of the mouse uterus compared with controls when examined at 72 hours. The sensitivity of the biological assays is, in general, good. However, there is little diagnostic application for these assays because of the expense and time involved and the excellent immunological testing methods now available. Immunological methods of pregnancy diagnosis generally rely on an antigen–antibody reaction. The antibody is produced by hyper-immunization of another animal such as a rabbit, with eCG as the antigen. Some reported immunological assay systems are the hemagglutination inhibition test (HI), mare immunological pregnancy (MIP) test, RIA, direct latex agglutination (DLA), and ELISA. In addition, a radioreceptor assay has been described. Serum from pregnant mares (diagnosed by ultrasonography) has been used to compare the efficacy of the MIP test, DLA assay, and ELISA.71 None of the seven samples from pregnant mares obtained on day 35 resulted in positive reactions with the HI or DLA assay, but three out of seven (43%) were diagnosed positive, using ELISA. On day 40,
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a greater (P < 0.01) number of mares were detected pregnant with the ELISA (16/16, 100%) compared to the HI (5/16, 31%) or DLA assay (6/16, 37%). Between days 45 and 100, there were no differences in efficacy among the three tests. At 50 days and later, efficacy of the three tests ranged from 90% to 100%. Sera collected from 10 non-pregnant mares were used as control samples. None of these samples resulted in positive reaction. The authors concluded that all three tests were simple, quick methods that could be used for detection of pregnancy in mares.71 Although the DLA assay was the easiest to conduct, all three assay procedures could be performed in < 2 hours, and the ELISA, because of increased sensitivity, could be used to detect pregnancy slightly earlier than the other two methods. Progesterone Progesterone is produced by the CL and functions to maintain pregnancy. The CL forms from luteinization of granulosa cells and theca interna cells from the wall of an ovulated follicle. The primary CL is formed from the follicle that released the oocyte that resulted in pregnancy. After fertilization and recognition of pregnancy, the primary CL is maintained beyond day 15 until at least days 140 through 180 and probably longer. Progesterone concentrations peak at approximately day 20, then decline slightly until days 40 through 45, at which time supplementary CL formation occurs, resulting in an increase in progesterone measurements until days 80 to 90 of pregnancy. It is speculated that supplementary CLs form from follicles that are stimulated by 10-day cyclic pulses of follicle-stimulating hormone (FSH) and that either ovulation (secondary CL) and/or luteinization (accessory CL) is initiated by eCG. Ovariectomy commonly will result in abortion before day 60, but not after day 80.72, 73 Pregnancy is maintained by progestins (progesterone-like compounds) produced by the placenta beginning around days 60 through 90, with increasing concentrations until parturition. ‘Pony mares were bilaterally ovariectomized at different stages of pregnancy between Days 25 and 210. Abortion or fetal resorption occurred within 2 to 6 days after operations in all 14 mares ovariectomized between Days 25 and 45 and after an interval of 10 to 15 days in 9 of 20 other ovariectomized between 50 and 70 days. All 12 mares ovariectomized on either 140 or 210 days carried their foals to normal term. The termination of early pregnancy was preceded by a loss of uterine tone and of a palpable uterine bulge. The mean length of gestation in all mares in which pregnancy was not interrupted by ovariectomy was not significantly different from that
in a group of contemporary control mares. Plasma progestagen concentrations dropped to less than 2 ng/ml after ovariectomy, whether or not pregnancy was maintained. Mares ovariectomized on Day 25 and injected with 100 mg progesterone daily for 10 or 20 days remained pregnant during treatment but showed a loss of uterine tone and the fetal bulge disappeared within 4 to 6 days after the end of treatment. Non-pregnant ovariectomized or intact seasonally anoestrous mares injected i.m. with 50 or 100 mg progesterone daily for 8 weeks showed changes in uterine tone, length and thickness similar to those occurring in mares during early pregnancy.’72 When used to assess ovarian activity in mares, measurement of plasma progesterone concentration is more reliable than observation of estrual behavioral patterns. Progesterone concentrations between 4 and 9 ng/mL are found 5 to 10 days after ovulation. If pregnancy ensues, higher concentrations are observed during days 16 through 20, compared to mares destined to return to estrus. In one study, measurement of plasma progesterone concentration in non-pregnant and pregnant mares at two stages, post-ovulation days 13 through 17 and 18 through 22, was 3.08 and 0.77 ng/mL in non-pregnant mares and 7.72 and 6.34 ng/mL in pregnant mares, respectively. Regardless, some pregnant mares will have low progesterone measurements and some diestrous mares (with a retained CL) will have high progesterone measurements similar to those in pregnant mares. There are numerous progestins. If the assay for progesterone is not highly specific, many other progestins including progesterone metabolites will be measured in the serum and milk, particularly after day 60 to 90 of gestation when progestins from the fetal placental unit are produced. In general, the RIA and ELISA are highly specific for progesterone, whereas competitive protein-binding assay will detect progestins as well. Since progesterone has been shown to play a key role in pregnancy maintenance, it is not surprising that progesterone administration to habitually aborting mares has become a fairly common practice. Generally, most of the pregnancy losses in mares occur during the first two months of gestation. In one study74 various doses and regimes of administering progesterone were evaluated for their efficacy in maintaining pregnancy in ovariectomized pregnant mares. Forty-eight light-horse mares were confirmed pregnant on day 29 or 30 and randomly assigned to one of six treatments: (i) non-injected controls, (ii) subcutaneous injection of 250 mg of progesterone in oil every other day, (iii) and (iv) intramuscular injection of 500 mg or 1000 mg of repositol progesterone every four days, respectively, or (v) and (vi) 22 and 44 mg of altrenogest daily, respectively.
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Maintenance of Pregnancy Hormonal treatments began on day 29 or 30 and ended on day 100 of gestation. As expected, none of the control mares maintained pregnancy after ovariectomy. Progesterone in oil (250 mg) administered every other day was only marginal in maintaining pregnancy (5 of 8). Administration of repositol progesterone (500 mg every four days) was ineffective in maintaining pregnancy (1 of 8). In contrast, all mares administered 1000 mg repositol progesterone every four days maintained their pregnancy. There appeared to be no difference between the two doses of altrenogest in maintenance of pregnancy; 7 or 8 mares maintained pregnancy in each group. Occasionally, mares experienced low levels of progesterone (less than 1 ng per mL) and still maintained pregnancy. Therefore, progesterone insufficiency should not be based on a single blood sample, but should be based on two or three samples collected during a given week. In this study, mares that had > 4 ng of progesterone per ml of blood were less likely to abort than those mares with levels less than 4 ng per mL. The level of progesterone achieved by exogenous treatment is dependent upon the type of progesterone used, the release profile of the progestagen, and thus the frequency of administration. Aqueous progesterone is cleared extremely rapidly from the blood and, therefore, should not be recommended for pregnancy maintenance. Progesterone suspended in an oil base (sesame, safflower, corn oil, etc.) needs to be administered on a daily basis. Repositol progesterone was the most common form of progesterone available to the practitioner. This progesterone is suspended in a propylene glycol base and is cleared relatively slowly from the blood of the mare. Injections of repositol progesterone were given every 4 to 7 days. Recent technology (see Chapter 198) has resulted in delivery vehicles that allow for administration once a week of either progesterone or altrenogest intramuscularly. Daily administration of oral altrenogest is recommended. Based on current information, it appears that if progesterone in oil is used, then 200 to 300 mg should be given daily. Altrenogest at a level of 0.044 mg per kg of bodyweight would appear to be effective in maintaining pregnancy. Readers are referred to Chapter 198 for information on microsphere, SABER, and the new biorelease compounded progestagen or progesterone products. Embryos were transferred into ovariectomized mares administered either 300 mg of progesterone in oil daily or 22 mg of altrenogest administered orally on a daily basis.75 Of 20 embryos transferred into ovariectomized mares treated with injectable progesterone, 15 resulted in a pregnancy, compared to 14 of 20 embryos transferred into ovariectomized mares administered altrenogest.
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‘Embryo transfer into ovariectomized steroid-treated mares was used as a model to evaluate various progestin/estradiol treatments and to determine the level of progesterone necessary for the maintenance of pregnancy in mares. Once a donor mare was in estrus and had a > 35 mm follicle, an ovariectomized recipient was selected and assigned to one of three groups: 1) 1 mg estradiol (E-2) was injected subcutaneously daily until the donor mare ovulated; on the day of the donor mare’s ovulation, daily intramuscular injections of 300 mg progesterone (P4) were commenced and continued until the end of the experiment (Day 35); 2) E-2 and P4 treatments were identical except E-2 was continued daily until Day 20; and 3) The same E-2 treatment as Group 1, 0.044 mg altrenogest per kilogram body weight were administered daily until Day 35. Embryos were recovered 7 d after the donor mare’s ovulation and were transferred via surgical flank incision. Twenty additional embryos (controls) were transferred into intact recipients that ovulated 1 d before to 3 d after the donor. Pregnancy rates did not differ (P > 0.05) among groups at Days 14 or 35. Pregnancy rates at Day 35 for mares administered injectable P4 (70%) were identical to those given altrenogest. Overall, pregnancy rates for ovariectomizedprogestin treated recipients (28 of 40, 70%) were similar (> 0.05) to that of intact mares (16 of 20, 80%). Dose of P4 was decreased in Groups 1 and 2 to 200 mg (Days 35 to 39), 100 mg (Days 40 to 44), 50 mg (Days 45 to 49) and 0 mg (> Day 50). Blood samples were collected once on Days 34, 35, 39, 40, 44, 45, 49 and 50 and assayed for P4. Dose of Altrenogest was decreased to 0.022, 0.011, 0.0055 and 0 mg per kilogram body weight at Days 35 to 39, 40 to 44, 45 to 49 and > 50. Number of mares in Groups 1 and 2 that lost their pregnancy while given 200, 100, 50 or 0 mg P4 was 0, 2, 8 and 4, respectively. Doses of 0.022, 0.011, 0.0055 and 0 mg altrenogest per kilogram body weight resulted in 0, 6, 4 and 3 mares aborting. Fetal death did not occur until concentrations of P4 decreased below 2.56 ng/ml 24 h after injection.’75 Apart from occasional enlargement of the clitoris seen in neonatal fillies,76 prolonged administration of altrenogest has not resulted in any deleterious effects.
Therapeutic considerations Management
General There are few, if any, treatments to consistently decrease incidence of EED, but artificial insemination can limit bacterial challenge to a mare’s uterus,77 thus reducing potential losses from endometritis. In addition, any new information on causes and treatment of endometritis should result in increased breeding efficiency. Strict adherence to repair of external conformation defects and thus a reduction in caudal
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reproductive tract contamination is important, as is efficient handling of twins, better management of placentitis, recognition of exposure factors for MRLS, and prevention of devastating outbreaks of viral abortion78 through vaccination, will all help reduce pregnancy losses. The transfer of embryos from mares with poor uterine biopsy grades into normal recipient mares is recommended to provide an environment more conducive to pregnancy maintenance. Unfortunately, recovery of embryos from subfertile mares is low. Perhaps the approach most likely to result in a decrease in incidence of EED is improving management factors related to uterine environment, nutrition, environmental temperature, infectious diseases, and other stresses. It seems that most problems of the horse are management related. There are few diseases or problems that our imposed management practices have not influenced or created. Take the case of the habitually aborting mare. It is most likely she has persistent and chronic uterine inflammation or fibrosis. We probably created it through poor hygiene, lack of exercise, amalgamation of too many mares in a stressful environment, and lack of appropriate therapy at the time of breeding or later. The pregnancy rates and foaling rates are generally better on well-managed breeding farms, despite the fact that there is absolutely no selection for reproductive efficiency, just performance-related parameters. Our belief is that persistent, unidentified or poorly treated chronic inflammatory conditions of the uterus are responsible for the majority of problems of EED and many cases of abortion.
Foal heat breeding One of the most important management issues still neglected somewhat is the practice of foal heat breeding. Many of our clients wish to breed mares on foal heat. The reason is generally obvious. The gestation of the mare averages 340 days, but a significant number are longer. Most of us involved with commercial breeding farms recognize a financial penalty for foals born later than the normal accepted times. Without breeding at foal heat it may be hard to maintain a foaling interval of 365 days in some cases. In addition, clients with brood mares on farms sometimes are paying high agistment costs and place pressure on managers to get their mares pregnant and send them home as soon as possible. Sometimes farm mangers cannot resist the temptation to breed at foal heat despite the potential problems, just because the mare is showing heat and their knowledge that foalheat mares ovulate quite quickly. However, fertility has been reported lower in mares bred during the first postpartum ovulatory pe-
riod compared with mares bred during subsequent cycles,17, 79 and EED has been reported higher for mares bred at this time.17, 21, 23, 79 This decreased fertility may be due to failure of elimination of microbes during uterine involution17, 21 or their introduction at breeding. In addition, presence of uterine fluid during estrus80 and diestrus81, 82 has been shown to reduce fertility of mares. A study from Australia clearly showed the generally held contention that fertility is poorer and embryonic death greater when mares are bred on foal heat, compared to the next cycle that was induced by prostaglandin.79 First-cycle pregnancy rates were 47.9% compared with 55.2%, and second-cycle pregnancy rates were 46% and 57.7%, respectively. Overall pregnancy rates at the end of the breeding season were 83.3% and 89.7%, respectively. Pregnancy losses before day 45 were 10% and 3.9% and after day 45 were 9.3% and 3.6%, respectively.79 A study80 was conducted to evaluate two hypotheses: (i) uterine involution and fluid accumulation could be effectively monitored with ultrasonography and used to predict fertility of mares bred during the first postpartum ovulatory cycle, and (ii) delaying ovulation with a progestin would result in improved pregnancy rates in mares bred during the first postpartum ovulatory period. The previously gravid horn was larger than the non-gravid horn for a mean of 21 days (range 15 to 25) after parturition. Uterine involution was most obvious at the corpus cornual junction. When the results of three ultrasonographic scans were similar, over a 5-day period, the uterus was considered to be involuted. On the average, uterine involution was completed by day 23 (range 13 to 29). Quantity and quality of uterine fluid were not affected by progestin treatment. Number of mares with detectable uterine fluid decreased after day 5 postpartum. Uterine fluid generally decreased in quantity and improved in quality between days 3 and day 15. Fewer (P < 0.005) mares became pregnant when uterine fluid was present during the first postpartum ovulatory period (3 of 9, 33%), compared to when no fluid was detected (26 of 31, 84%). Mares with uterine fluid during breeding did not have appreciably larger uterine dimensions, compared with those mares not having fluid. There was no relationship between uterine size on day of ovulation and pregnancy rate. Ovulations were delayed, and pregnancy rates improved in progestin-treated mares. More (P < 0.05) mares became pregnant (23/28, 82%) when they ovulated after day 15, in the first postpartum ovulatory period, than mares that ovulated before day 15 (6/12, 50%). Ultrasonography has been proven useful in detecting mares with postpartum uterine fluid. Further, it could be used to aid in determining whether a mare should be bred, treated, or not bred during the first postpartum ovulatory
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Maintenance of Pregnancy period. During estrus, uterine fluid may be spermicidal and/or an excellent medium to support bacterial proliferation. When fluid is present during diestrus, it may cause premature luteolysis or EED.82 Quantity of uterine fluid during the first postpartum ovulatory period appeared to be related to stage of uterine involution, and was reduced or eliminated by delaying the ovulatory period with progestins. Progestin treatment not only allowed time for elimination of uterine fluid before the first postpartum ovulation, but it also significantly delayed the first postpartum ovulation. Results of this study concurred with those of others in which it was concluded that progestin treatment delayed onset of the first postpartum ovulatory period, but did not affect rate of uterine involution.83–85 Long-term progestin administration to normal, cycling mares has not been shown to adversely affect fertility.86 However, treatment with progestins will affect uterine defense mechanisms87, 88 and thus care is recommended before prolonged progestin treatment is administered to postpartum mares or mares susceptible to infection. Since there were decreased pregnancy rates associated with uterine fluid, and increased pregnancy rates as ovulation was delayed, it was suggested both techniques could be used to manipulate breeding strategies and improve pregnancy rates from normal mares bred during the first postpartum ovulatory period. The approach used at our practices for management of foal heat breeding is listed below.
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All mares are examined at day 2–3 post foaling. By that time, if problems have occurred during foaling then they will be identified. We believe that this examination is critical if we are to prevent mares from becoming ‘problem broodmares’. Each year we identify mares that, if inappropriately treated, could become long-term disasters. Approximately 5% of mares will have placental tags present (despite excellent staff management) and another 5% problems with metritis and uterine damage. Many (15%) will have delayed involution, suggested by increased size of the uterus and volume of fluids present. Such mares are treated aggressively with large-volume uterine lavage and ecbolics, antibiotics and anti-inflammatories, and monitored closely. All mares are treated with an infusion of lactated ringer’s saline with antibiotics. We believe that this helps only the occasional mare but, more importantly, we have invaded the mares’ reproductive tract and the treatment is aimed at preventing iatrogenic contamination. Mares with no abnormalities identified are scheduled to be re-examined on day 9–10 post foaling. Mares will not be bred any earlier than day10. On
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day 9–10, if the mare has ovulated, she is scheduled for PGF2α administration in 6 days. If no uterine fluid is detected and uterine tone is good, the mare may be scheduled to be bred. If there is a question, the mare is treated and or re-examined according to follicle size and presence or absence of uterine fluid. It would be rare for a foal-heat breed to take precedence over a non-foal-heat breed with a stallion with a busy book (> 150 mares per season with natural service). Breeding mares on foal with intraluminal fluid accumulations is avoided. These mares are treated and recycled. Lastly there are two more guidelines that we adhere to. First, we only breed mares on foal heat that are young and reproductively healthy (i.e., < 12 years old and with their first, second or third foal); and second, mares that have been confined without exercise (i.e., lunging) due to foal problems such as angular limb deformities, are not bred on foal heat regardless.
Hormones
Progestagens By far the most popular treatment to aid in pregnancy maintenance is the supplementation of progesterone or progestagens. This occurs despite advice that there is no scientific evidence to support the use of such compounds.18, 89, 90 Most studies suggest that primary luteal inadequacy is not likely a cause of decreased progesterone levels.91 ‘Plasma progesterone concentrations were measured in 179 mares bled on alternate days commencing with a positive pregnancy diagnosis on Days 17 to 18 after ovulation and concluding on Days 42 to 45. During this period 17 mares (10 per cent) lost their pregnancies, 11 before Day 25. In 15 mares the timing of the pregnancy loss could be determined with adequate accuracy; in only one did a decline in progesterone precede the loss. Thus pregnancy loss between Days 17 and 42 was rarely caused by a fall in plasma progesterone.’91 The exception is failure of mares to maintain either a CL or pregnancy, when induced to ovulate from either the early transition or anestrus.92, 93 ‘Two experiments were conducted using a 21-day GnRH analogue treatment regimen to induce ovulation in seasonally anovulatory mares. In Experiment 1, non treated (n = 20) and treated (n = 83) mares were defined as having inactive ovaries (largest follicle < 15 mm) at the start of treatment. In Experiment 2, non treated (n = 7) and treated (n = 10) mares were defined as having active ovaries (largest follicle 21 to
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29 mm) at the start of treatment. Less mares ovulated that had inactive ovaries at the start of treatment compared to mares that had active ovaries (26% versus 70%). In mares that had inactive ovaries at the start of treatment, the pregnancy rate on Day 11 (ovulation = Day 0) was not different between treated (14/22; 64%) and non-treated (14/20; 70%) mares. However, the embryo-loss rate between Days 11 and 40 was significantly higher in treated (9/14; 64%) than in nontreated (1/14; 7%) mares. Losses tended to occur more frequently between Days 15 and 25. A significant decrease in progesterone on Day 15 in treated mares with embryo loss corresponded to the first day that embryo loss was detected. In mares that had active ovaries at the start of treatment, pregnancy rate, embryo-loss rate, and ovarian and hormonal end points were not significantly different between the treated and non-treated mares. The results indicated that hormonal capacity was reduced and embryo-loss rate was high following GnRH-induced ovulation in mares that had inactive ovaries at the start of treatment.’92 Another report suggests that, although the incidence is apparently low, we should continue to watch out for mares with low progesterone that occurs before obvious fetal demise.94 ‘Characteristics of spontaneous embryonic loss in 21 mares were compared with those of 52 contemporary mares that maintained pregnancy. Embryonic losses were, in retrospect, grouped according to day of loss and length of the interovulatory interval, respectively, as follows: group 1, ≤ day 20 and < 30 days (n = 10); group 2, ≤ day 20 and > 30 days (n = 3); and group 3, > day 20 and > 30 days (n = 8); ovulation was day 0. Mean diameter of the embryonic vesicle in group 1 was smaller (P < 0.05) on days 12–18 than in the pregnancy-maintained group, but among the pregnancy-maintained group and the embryonicloss groups, the mean individual growth rates of vesicles was similar (no significant difference). A more frequent (P < 0.05) location of the vesicles in the uterine body on day 13 in group 1 was due to a greater proportion of small vesicles and for day 18 was due to a greater incidence of fixation failure. Luteal regression occurred at the expected time in 77% of the mares with loss sooner than day 20. Low concentration of progesterone on days 12, 15 and 18, a detected decrease in diameter of the corpus luteum on days 15 and 18, and an interovulatory interval of ≤ 30 days indicated that luteolysis was not prevented by the embryonic vesicle in group 1. However, in mares in group 2, the corpus luteum was maintained as indicated by luteal diameters, progesterone concentrations, and prolonged interovulatory intervals. Thus, luteal maintenance occurred in 23% of mares with embryonic loss at ≤ day 20. In group 3 (loss > day 20), circulating concentrations of progesterone were either maintained (four mares) after embryonic death (cessation of embryonic heartbeat) or appeared to decrease
before death (two mares) or on the day of death (one mare). The decrease in progesterone on the day of death seemed to be associated with acute endometritis. In another mare, embryonic loss was associated with continuing low concentrations of progesterone, a flaccid uterus, an oversized embryonic vesicle, failure of development of an embryo proper and collapse of the large vesicle on day 42. The apparent drop in progesterone before embryonic death in two of eight mares with loss after day 20 could have been associated with primary luteal insufficiency.’94 Thus, although in many cases pregnant mares are probably unnecessarily treated with progestins, such treament would appear to do no harm. An exception is mares that are chronically infected. Administration of either altrenogest or natural progesterone to a chronically infected mare will enhance the infection process. The mare should first be examined using ultrasonography, cultured and a uterine biopsy obtained in order to confirm that she is not harbouring a uterine infection. Another exception is the use of compounds that are ineffective for the intended purpose. Studies at the Goulburn Valley Equine Hospital were able to demonstrate failure of hydroxyprogesterone caproate to maintain pregnancy in ovariectomized mares.95 Hydroxyprogesterone caproate is commonly used in many countries and apparently it has not been critically studied. Despite lack of definitive studies on frequency of a primary luteal defect on EED or the relative contributions of stress, PGF2α release associated with uterine inflammation, genetic abnormalities or uterine infection, many veterinarians, breeding farm managers, and mare owners are convinced of the beneficial effect of supplemental progestagens. Perhaps inappropriate administration of therapeutic substances that lack scientific documentation of efficacy is an even bigger form of reproductive wastage due to cost and lack of specified effect. Hydroxyprogesterone caproate has been specifically advocated for use in pregnancy maintenance of mares (manufacturers’ recommendations). Despite lack of research data supporting the drug for this purpose, its use appears to be widespread. The results of this experiment suggest that pregnancy in mares with inadequate luteal function cannot be maintained with hydroxyprogesterone caproate at 4–15 times the recommended dose rate. Pregnancy can, however, be maintained by altrenogest supplementation together with the contribution of endogenous progesterone from supplementary CL development which will occur despite exogenous therapy.95 A further study was useful in demonstrating just how little effect many of the progestagens either recommended or used actually have.96 In this study, 25 mares that were diagnosed pregnant on day 14 after ovulation were divided into five groups of five and
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Maintenance of Pregnancy treated as outlined below; all mares were injected i.m. with 5 mg PGF2α to induce luteolysis of the primary CL on day 18 of pregnancy. All supplemental treatments were begun on day 16. Group 1: 1000 mg medroxyprogesterone acetate administered i.m. every 7 days; Group 2: 500 mg hydroxyprogesterone hexanoate administered IM every 4 days; Group 3: 0.044 mg/kg oral altrenogest daily; Group 4: 15 mg (five implants) norgestomet subcutaneously; Group 5: 500 mg megesterol acetate orally daily. Serum was analysed for progesterone using an amplified enzyme-linked immunoassay (AELIA), developed and validated for horse serum,97 until either abortion occurred or mares were ≥ 24 days pregnant with a fetus and detectable heartbeat. Ultrasonography was performed daily to monitor pregnancy, presence of endometrial folds, time to the next ovulation and time to either pregnancy loss or recognition of the fetus within the vesicle and normal growth. All five mares in each of groups 1, 2, 4, and 5 aborted between 2 and 8 days after PGF2α administration, whereas none of the mares in group 3 given altrenogest daily aborted. Serum progesterone concentrations in all mares had fallen to less than 2 ng/mL the day following PGF2α administration and they remained very low or undetectable on the following days. All the mares in Groups 1, 2, 4, and 5 returned to estrus and ovulated within 11 days after PGF2α administration. The results of this experiment demonstrated convincingly that medroxyprogesterone acetate, hydroxyprogesterone hexanoate, norgestomet and megesterol acetate were unable to maintain pregnancy in mares in the absence of endogenous progesterone secreted by a viable CL. It is reasonable to conclude from these findings that the progestagens used in this study were not likely to be of any more use when administered at a later time in pregnancy. Pregnancy in the mare can be maintained solely by placental sources of progestagens from around day 80–100 of gestation,72, 74, 98, 99 but prior to this time ovarian corpora lutea, both primary and accessory18, 74 are the only endogenous source of this hormone. The progestagens chosen in this study are those commonly administered by equine veterinary clinicians either to prevent abortion or to modify undesirable behavior in performance horses, and the dose rates of progestagen administration were from manufacturer recommendations. Altrenogest was used as a positive control since it had been shown in a previous study to maintain pregnancy in ovariectomized mares.95
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We are now at least cognisant of a few progestagens that we can use: progesterone at around 150 mg/day, altrenogest at 22 mg/day, and some of the newer compounded formulations. Recent data on long-acting administration with bio-release compounds is presented in Chapter 198). Little data exist on supplementation later in pregnancy when the placenta is the only source of progestins. A study examined progestins at this time and effect of supplementation.99 ‘Two experiments were conducted to determine when a placental source of progestin was sufficient for maintaining pregnancy in the mare. In the first study, embryos were transferred into ovariectomized mares and pregnancy was maintained with altrenogest. Altrenogest treatment was terminated at either day 100 (n = 6) or day 150 (n = 6). Twelve ovarian-intact mares were assigned to a second experiment on day 100 of gestation. On day 160 of gestation, these mares were assigned to one of three treatments: 1) ovariectomy on day 160 and given altrenogest to day 200 (n = 4); 2) ovariectomy on day 180 and given Altrenogest to day 250 (n = 4); or 3) ovariectomy on day 200 and given altrenogest to day 300 (n = 4). Blood samples were collected every 2 weeks from all mares in both experiments from day 100 to parturition and assayed for concentrations of progestins. Pregnancy loss from day 100 to parturition was not different among groups in either experiment. Serum concentrations of progestins in ovary-intact mares were greater (P < 0.05) than those in ovariectomized pregnant mares until day 130, after which they were similar. Serum concentrations of progestins in the ovariectomized pregnant mares rose gradually from day 100 until near parturition. Serum concentrations of progestins in the ovary-intact pregnant mares declined from day 100 to day 157, did not vary significantly from day 157 to day 245, then rose until near parturition. Serum concentrations of progestins tended to decrease the sixth day prior to parturition. From these data, it was concluded that even though the ovary is a significant source of circulating progestins until day 130 of pregnancy in the mare, the feto-placental unit produces sufficient progestins to maintain pregnancy by day 100 of gestation.’99 Unfortunately this and other studies have not been able to address the question of whether or not supplementation of the feto-placental unit would ever be likely to be required. The other question not adequately addressed is when and how to withdraw progesterone supplementation. Generally speaking in our practices progesterone is withdrawn if supplementary CL development can be demonstrated before 60 days, accompanied by a progesterone measurement of greater than 4 ng/mL. After 100 days as placenta
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progestagens are maintaining pregnancy supplementation may be withdrawn. Typically we recommend a tapered withdrawal of progestagens such as a halfdose for a week before total withdrawal.
Supplementation of estrogens Supplementation of estrogens has been recommended100 or suggested for further examination.101 Nishikawa suggested the rationale behind supplementation was to prolong the life of the CL. He administered stilbestrol in either tablet or injectable form at a dose of 60–90 mg (injectable) or 144–200 mg (orally) over a 2-month period starting at the third or fourth month of gestation. Fifty-seven percent of supplemented animals had miscarried one to five times prior to the years in which the experiment was performed (1949–54). Significantly fewer (P < 0.0001) treated animals aborted (19/576; 3.3%) compared to control animals (474/3734; 12.7%). Nishikawa states ‘that the treatment with estrogen is definitely effective for preventing abortion in mares’, and ‘the treatment was highly praised by farmers’.100 Despite this convincing evidence, further studies have failed to find any effect of diethylstilbestrol on progesterone secretion.102 The work of Pashen103 suggests that we need to look further at the role of the fetal gonads in pregnancy maintenance and parturition. ‘The effects of fetal gonadectomy on steroid production and the maintenance of pregnancy in the mare were studied. Removal of the fetal gonads resulted in an immediate fall in maternal plasma concentrations of conjugated and unconjugated oestrogens whereas progestagen levels remained unchanged. Hormone profiles in mares carrying sham-operated fetuses remained similar to those in un-operated control mares. Plasma levels of 13,14-dihydro-15-oxo-PGF-2 alpha (PGFM) were much lower, and uterine contractions weaker, during labour in mares carrying gonadectomized foals than in control mares. Pregnancy was maintained until parturition at term in the mares carrying gonadectomized fetuses. However, 3 of the 4 gonadectomized foals were dysmature and died during or soon after birth. The biosynthetic pathways involved in the production of oestrogens by the feto-placental unit and the possible role of oestrogens in fetal development in the pregnant mare are discussed.’103
Supplementation with GnRH A slight increase in pregnancy rate per cycle and, more importantly, a decrease in EED per cycle has been demonstrated with the GnRH agonist buserelin.104 ‘Two trials involving 578 mares were performed to investigate the effect of a single intramuscular treatment
of 40 µg buserelin, an analog of gonadotrophin releasing hormone, on pregnancy rate in mares. All mares were bred by natural mating and were allocated into pairs. One mare in each pair was injected with buserelin either on Day 10 or 11 (Trial 1) or on Days 8 to 10 (Trial 2) after ovulation. Pregnancy status of mares was determined by transrectal ultrasonographic examination on Day 14 or 15 after the day of ovulation and was repeated between Days 28 and 30 of pregnancy. In Trial 1, buserelin treatment increased the pregnancy rate at Days 14 and 15 (72.5 vs 66.6%; P < 0.01). At the second pregnancy examination, pregnancy losses were lower in the treated group of mares (4.1 vs 7.4%; P < 0.05). In Trial 2, buserelin also improved the pregnancy rate (57.2 vs 53.5%; P < 0.05) at Days 14 and 15. Pregnancy losses between the first and second examinations were lower in the treated group of mares (6.5 vs 12.0%; P < 0.05). Buserelin increased pregnancy rates after breeding at the first estrus in both trials. In addition, buserelin treatment increased the pregnancy maintenance rate at Days 28 to 30.’104 This finding has potentially interesting ramifications, but needs to be replicated by others. Further work by Newcombe and others105 has strengthened the data: ‘A series of trials over a four-year period on a total of 2,346 mares was conducted, to determine the effect of a single dose of the GnRH analog buserelin (20 to 40 µg im or sc) on pregnancy rates when given between 8 and 12 days after service. Although there were some statistically significant improvements in pregnancy rates in individual trials, meta-analysis of the data overall showed significant improvements at all times examined, i.e. 13 to 16, 19 to 23, 28 to 31 and 38 to 42 days after service. These results indicate that treatment of mares with 20 to 40 µg buserelin between Days 8 and 12 significantly increases pregnancy rates by approximately 10 percentage points.’105 In addition, another study has supported the hypothesis (although the pregnancy rates were quite low).106 It is not currently understood why this therapy is beneficial. One possibility is that the GnRH may be luteotropic and thus stimulate the primary CL of pregnancy. Other considerations
Antibiotics Our belief is that persistent, unidentified or poorly treated chronic inflammatory conditions of the uterus are responsible for the majority of problems of EED and abortion. It is common to visualize with ultrasonography significant uterine inflammation (inappropriate uterine edema) and excessive fluid in mares
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Maintenance of Pregnancy that have undergone early pregnancy loss and then are given prostaglandins to return them to estrus. Therapies aimed at reducing this inflammation, infection, and irritation are a primary goal in our practice. To this end, mares with a history of recurrent pregnancy loss are biopsied wherever possible and, if the primary problem is uterine inflammation, they are placed on broad-spectrum antibiotics at intervals throughout pregnancy.107 The results from our practices appear very encouraging. When pregnancy continues, antibiotics are administered for 5 consecutive days each and every month. As with many treatments of endometritis, there is no experimental data available to document the efficacy of such treatments. However, at the Goulburn Valley Equine Hospital we have been using this approach since before 1993 and have little doubt that the number of mares maintaining pregnancy is much higher than those mares not treated. A study was reported wherein potential cases of chronic uterine infection/inflammation were treated with antibiotics, monthly, during all of the pregnancy. Because not all EED and abortion is caused by bacterial infection/inflammation, it was not expected that all mares would maintain pregnancy. The mares selected for treatment had all undergone either EED or abortion at least once in the last season prior to entering the trial.108, 109 In this study 43 mares had entered the program, they had a mean age of 14.2 years, and had EED or abortion an average of 2.2 times in the preceding 3–4 years. At the conclusion of the study 36 mares had become pregnant (84%). Of the 36 pregnant mares, 3 underwent EED (8.3%), 4 aborted (11.1%), and 29 had foals (80.6%).109 It needs to be recognized that it may not be possible to obtain valid significant scientific data from these studies. In the study presented above there were no controls, and caution is advised in interpreting studies without contemporary controls. A publication from South Africa highlights well the effect of post-foaling irritation and involution.110 ‘Records of 1,009 pregnancies in 574 foaling, barren and maiden Thoroughbred mares on a single stud farm, over a period of 12 years were examined. The farm is situated in the Eastern Cape Province of South Africa, at an elevation of 1800 m, and in an area of climatic extremes. Records of 604 pregnancies in 249 foaling Thoroughbred mares were examined. For these purposes, those pregnancies in which a mare conceived in the same breeding season during which she had foaled were considered as pregnancies in foaling mares. Pregnancy was confirmed by rectal palpation by a single experienced practitioner. Of the 604 pregnancies examined, conceptus attachment occurred in the horn opposite the previously gravid horn in 345 cases (57%), and in the previously gravid
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horn in 259 cases (43%; P < 0.005). Unobserved foetal loss after pregnancy diagnosis amounted to 30 (9%) in the former group, while in the latter group (pregnancy established in the postgravid horn) 46 pregnancies were lost (18%; P < 0.005). This study confirmed that conceptus attachment tends to occur in the uterine horn opposite the previously gravid horn in foaling Thoroughbred mares conceiving during the same season. A significantly higher incidence of foetal loss accompanied conceptus attachment in the postgravid horn. Of 242 pregnancies in 162 previously barren mares, 95 (39%) occurred in the left uterine horn and 147 (61%) in the right horn (P < 0.005). The incidence of pregnancy failure in this group was 7%. The side of attachment did not affect the rate of loss. Evaluation of the records of 163 maiden mares revealed that conceptus attachment occurred in the left uterine horn in 58 (36%) pregnancies and in the right horn in 105 (64%) pregnancies (P < 0.005), which is consistent with previously reported observations. Pregnancy failure was recorded in 4% of maiden mares. Side of attachment did not influence rate of loss in this group.’110 An identical study has recently been published.111 These authors reported that in 1383 Thoroughbred mares monitored by ultrasonography during estrus and early pregnancy, significantly (P < 0.01) more pregnancies were located in the previously non-gravid horn (79.2%) than the previously gravid horn (20.8%). In addition, the number of pregnancies in the previously gravid horn significantly increased with mare age (P < 0.01) and foaling to conception interval (P < 0.05). Similar to the South African study, more pregnancies located in the previously gravid horn were lost (16.5%) compared to losses of those that were located in the previously non-gravid horn (4.6%) (P < 0.001).111 Lastly, we would serve our clients best by trying to identify which mares are in a high-risk category for abortion or EED, rather than trying to diagnose what has made the mare lose the pregnancy. ‘Equine high-risk pregnancy can be caused by conditions of the fetus or placenta. Placental abnormalities are the most common cause of equine abortion. Conditions that lead to equine high-risk pregnancy can be caused by infectious agents, physical abnormalities, or the ingestion of toxins. The diagnosis of fetal and placental conditions can be very challenging, as clinical signs are often absent; the most common premonitory sign of fetal or placental disease is premature lactation. Most congenital abnormalities (arthrogryposis, hydrocephalus, and hydrops, etc) have no treatment. Therapy in these cases is directed at facilitating parturition to salvage the future breeding potential of the mare. Patients with some infectious causes of high-risk pregnancy, such as bacterial placentitis, can be successfully treated. Early
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recognition of the infection and aggressive antimicrobial therapy, in some cases, control the infection and prolong gestation until a viable fetus is born. Additional therapy for placentitis can include the use of non-steroidal antiinflammatory drugs and supplemental progestins.’112 In this regard fetal monitoring has been useful in identification of placental problems and to aid in decision processes based on probability of successful treatments.
Immune stimulation The earliest report on using non-specific immune stimulation suggested a decrease in the numbers of neutrophils after intramuscular administration during estrus.113, 114 These reports of immune stimulation having an apparent benefit on the mare’s reproductive tract resulted in us utilizing the therapy R (EqStim ; Neogen Corporation) during estrus and in early pregnancy. The aim has been to reduce acute inflammation. ‘Thirty-seven mares, barren for at least one year, were diagnosed to have endometrial inflammation by cytological examination and/or bacterial culture. Sixteen of these mares (the control group) were treated with prostaglandin and reevaluated cytologically within 10 days of behavioral estrus. Fifteen out of sixteen (15/16) (93.8%) remained positive for endometrial inflammation. Twenty-one mares (immunologically treated group) were treated with Propionibacterium acnes by injecting 1 ml per each 113.3 kilograms body weight intravenously on days 1, 3, and 7 following the diagnosis of endometrial inflammation. They also received prostaglandin. Twelve of the immunologically treated mates were examined cytologically as were the control mares. One of twelve examined (1/12) (8.3%) was positive cytologically for endometrial inflammation when examined within 10 days after behavioral estrus. The other nine mares in the treated group were bred on the first behavioral estrus following treatment. Nine of nine (9/9) were determined to be pregnant to 60 days gestation. The other treated mares were bred following the cytological evaluation. Eighteen of the twenty-one treated mares(18/21) (85.7%) were confirmed pregnant to 60 days gestation.’114 More recent publications have examined the effects of either one of two immune modulators. The R , mentioned first, Propionibacterium acnes (EqStim 115 above) was reported on recently, and the second, TM a cell wall extract of Mycobacterium phlei (Settle ; Bioniche Animal Health) has been the subject of numerous recent publications. Propionibacterium acnes, when administered intravenously to mares with a cytological diagnosis of per-
sistent endometritis, was associated with increased pregnancy and foaling rates. The effect was best when the immune stimulant was administered from 8 days before up to 2 days after breeding.117 ‘Mares were treated with P acnes or placebo (both administered IV) on days 0, 2, and 6. No attempt was made to alter additional treatments administered by attending veterinarians. Information on breeding history, physical examination findings, results of cytologic examination and microbial culture of uterine samples, additional treatments administered, breeding dates, results of pregnancy examinations, whether a live foal was produced, and reactions to treatment was recorded. . . . In multivariate logistic regression models, mare age, year of entry into the study, and first breeding within 8 days after first treatment with P acnes or placebo were significantly associated with pregnancy. Fewer number of cycles bred and younger age were significantly associated with delivery of a live foal in a separate multivariate analysis. Results of multivariate logistic regression modeling indicated that mares treated with P acnes were more likely to become pregnant and to deliver a live foal, compared with placebo-treated controls. CONCLUSIONS AND CLINICAL RELEVANCE: IV administration of P acnes as an adjunct to conventional treatments in mares with a cytologic diagnosis of persistent endometritis improved pregnancy and live foal rates. The optimal effect was detected in mares bred during the interval extending from 2 days before to 8 days after first treatment with P acnes.’115 The studies by Fumuso116, 117 on Mycobacterium phlei are similarly convincing. ‘Our objective was to characterize immune parameters in susceptible (SM) and resistant (RM) mares, with and without artificial insemination (AI) and immunomodulation. Eight RM and eight SM were selected based on their reproductive history and functional tests. Both groups of mares were evaluated during three consecutive cycles: Cycle 1, untreated cycle (control); Cycle 2, AI with dead semen; Cycle 3, AI with dead semen and immunomodulation. Endometrial biopsies were taken during the three cycles as follows: Cycle 1 – at estrus, when follicles > or = 35 mm and at diestrus (7 ± 1 days after ovulation); Cycle 2 – at estrus 24 h post-AI, and at diestrus; Cycle 3 – at estrus 24 h after treatment with a Mycobacterium phlei cell-wall extract (MCWE) and AI, and at diestrus. The mRNA transcription (mRNAT) of IL-8 and IL-10 were determined by real-time PCR. Image analysis of immunohistochemistry slides was performed using digital software (Image-Pro Plus v 5.0; Media Cybernetics); the percentage of stained area was determined for Major Histocompatibility Complex II (MHC-II), polymorphonuclear leukocytes (PMN) and T lymphocytes (TL) on each tissue section. In Cycle 1, SM had significantly higher MHC-II, TL, PMN and
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Maintenance of Pregnancy IL-8 than RM during estrus (P < 0.006, P < 0.0005, P < 0.05, respectively), while transcription of IL-10 was significantly lower than in RM (P < 0.0001). During diestrus, SM had higher levels of TL, PMN and IL-8 than RM (P < 0.0001). After AI (Cycle 2), SM had higher levels of IL-8 and lower levels of IL-10 than RM at estrus and no differences were detected for MHC-II, TL and PMN positive cells. During diestrus in the same cycle, all the immune parameters were higher in SM mares (P < 0.005, P < 0.0004, P < 0.0001, P < 0.02, respectively). When MCWE was applied at the time of AI (Cycle 3), SM expressed significant higher levels of IL-10 24 h after treatment (P < 0.005), which were also higher than in the control Cycle 2 or after AI (Cycle 2). However, no significant differences were detected for MHC-II, lymphocytes-PMN or IL-8 between SM and RM during diestrus in Cycle 3. This study showed that SM had higher levels of all immune parameters except IL-10 than RM during Cycle 1. After AI (Cycle 2), the inflammatory condition persisted in SM but not RM mares until day 7 post-ovulation. Following treatment with MCWE at the time of AI (Cycle 3) uterine immunological changes in SM resulted in an endometrial immune environment similar to that found in normal RM.’117
Taken together, these indicate that immune stimulation can be expected to reduce neutrophil numbers in mares with persistent mating-induced endometritis, with a corresponding increase in pregnancy and foaling rates. When a mare has a history of habitually aborting or experiencing EED, the decision of whether to use immune stimulation, supplement or not with progestagens, antibiotics or other treatments is largely at the discretion of the owners and managers. Many owners and mare or farm managers, having invested so much time and money in getting the mare pregnant, will feel compelled to supplement or treat regardless of lack of documented efficacy and our advice.
References 1. Nath LC, Anderson GA, McKinnon AO. Reproductive efficiency of Thoroughbred and Standardbred horses in north-east Victoria. Aust Vet J 2010;88:169–75. 2. Morris LH, Allen WR. Reproductive efficiency of intensively managed Thoroughbred mares in Newmarket. Equine Vet J 2002;34(1):51–60. 3. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of Flatrace and National Hunt Thoroughbred mares and stallions in England. Equine Vet J 2007;39(5):438–45. 4. Ricketts S, Troedsson MHT. Fertility expectations and management for optimal fertility. In: Sampieri F, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007: pp. 53–69.
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5. Villahoz MD, Squires EL, Voss JL, Shideler RK. Some observations on early embryonic death in mares. Theriogenology 1985;23:915–24. 6. Ball BA, Little TV, Hillman RB, Woods GL. Pregnancy rates at days 2 and 14 and estimated embryonic loss rates prior to day 14 in normal and subfertile mares. Theriogenology 1986;26:611–20. 7. Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod Mono 1995;1:209–14. 8. Brinsko SP, Ball BA, Miller PG, Thomas PGA, Ellington JE. In-vitro development of day-2 embryos obtained from young, fertile mares and aged, subfertile mares. J Reprod Fertil 1994;102:371–8. 9. Day FT. Clinical and experimental observations on reproduction in the mare. J Agri Sci 1940;30:244–61. 10. Kenney RM. Cyclic and pathologic changes of the mare endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1978;172:241–62. 11. Ginther OJ, Bergfelt DR, Leith GS, Scraba ST. Embryonic loss in mares: Incidence and ultrasonic morphology. Theriogenology 1985;24:73–86. 12. Belonje PC, van Niekerk CH. A review of the influence of nutrition upon the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1975;( 23): 167–9. 13. van Niekerk CH, Morgenthal JC. Fetal loss and the effect of stress on plasma progestagen levels in pregnant Thoroughbred mares. J Reprod Fertil Suppl 1982;32:453–7. 14. Swerczek TW. Early fetal death and infectious placental disease in the mare. Proc Am Assoc Equine Pract 1980;173–9. 15. Brendemuehl JP, Boosinger TR, Pugh DG, Shelby RA. Influence of endophyte-infected tall fescue on cyclicity, pregnancy rate and early embryonic loss in the mare. Theriogenology 1994;42:489–500. 16. Moberg R. The occurrence of early embryonic death in the mare in relation to natural service and artificial insemination with fresh or deep-frozen semen. J Reprod Fertil Suppl 1975;(23):537–9. 17. Merkt H, Gunzel AR. A survey of early pregnancy losses in West German thoroughbred mares. Equine Vet J 1979;11:256–8. 18. Allen WR. Is your progesterone therapy really necessary? Equine Vet J 1984;16:496–8. 19. Liu IK, Shivers CA. Antibodies to the zona pellucida in mares. J Reprod Fertil Suppl 1982;32:309–13. 20. Shivers CA, Liu IK. Inhibition of sperm binding to porcine ova by antibodies to equine zonae pellucidae. J Reprod Fertil Suppl 1982;32:315–18. 21. Platt H. Aetiological aspects of abortion in the Thoroughbred mare. J Comp Pathol 1973;83:199–205. 22. Fiolka G, Kuller HJ, Lender S. Embryonic mortality of horse. Monatshefte Fuer Veterinaermedizin 1985;40:835–8. 23. Lieux P. Comparative results of breeding on first and second post-foaling heat periods. Proc Am Assoc Equine Pract 1980;129–32. 24. Bain AM. Foetal losses during pregnancy in the thoroughbred mare: a record of 2,562 pregnancies. N Z Vet J 1969;17:155–8. 25. McDowell KJ, Sharp DC, Grubaugh W, Thatcher WW, Wilcox CJ. Restricted conceptus mobility results in failure of pregnancy maintenance in mares. Biol Reprod 1988;39:340–8. 26. Carnevale EM, Ramirez RJ, Squires EL, Alvarenga MA, Vanderwall DK, McCue PM. Factors affecting pregnancy
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rates and early embryonic death after equine embryo transfer. Theriogenology 2000;54(6):965–79. Woods J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic loss rate in mares. Equine Vet J 1990;22: 410–15. Koskinen E, Lindeberg H, Kuntsi H, Ruotsalainen L, Katila T. Fertility of mares after postovulatory insemination. Zentralbl Veterinarmed [A] 1990;37:77–80. Vanderwall DK, Squires EL, Brinsko SP, McCue PM. Diagnosis and management of abnormal embryonic development characterized by formation of an embryonic vesicle without an embryo in mares. J Am Vet Med Assoc 2000;217(1):58–63. Chevalier Clement F. Pregnancy loss in the mare. Anim Reprod Sci 1989;20:231–44. Fontaine M, Collobertlaugier C, Tariel G. Abortion and stillbirth of infectious origin in the mare – a study of necropsy of aborted fetuses, premature and stillborn foals. Point Vet 1993;25:79–84. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases(1986–1991). J Am Vet Med Assoc 1993;203:1170–5. Hong CB, Donahue JM, Giles RC, Petritesmurphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Equine abortion and stillbirth in Central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. Smith KC, Blunden AS, Whitwell KE, Dunn KA, Wales AD. A survey of equine abortion, stillbirth and neonatal death in the UK from 1988 to 1997. Equine Vet J 2003;35(5):496–501. Garg DN, Manchanda VP. Prevalence and etiology of equine abortion. Indian J Anim Sci 1986;56:730–5. Santschi EM, Slone DE, Gronwall R, Juzwiak JS, Moll HD. Types of colic and frequency of postcolic abortion in pregnant mares:105 cases (1984–1988). J Am Vet Med Assoc 1991;199:374–7. Baucus KL, Ralston SL, Nockels CF, McKinnon AO, Squires EL. Effects of transportation on early embryonic death in mares. J Anim Sci 1990;68:345–51. Irwin CFP. Early pregnancy testing and its relationship to abortion. J Reprod Fertil Suppl 1975;23:485–488. Donahue JM, Williams NM. Emergent causes of placentitis and abortion. Vet Clin North Am Equine Pract 2001;16(3):443. Reef VB, Vaala WE, Worth LT, Sertich PL, Spencer PA. Ultrasonographic assessment of fetal well-being during late- gestation – development of an equine biophysical profile. Equine Vet J 1996;28:200–8. Renaudin CD, Troedsson MH, Gillis CL, King VL, Bodena A. Ultrasonographic evaluation of the equine placenta by transrectal and transabdominal approach in the normal pregnant mare. Theriogenology 1997;47(2):559–73. Renaudin CD, Liu IM, Troedsson MT, Schrenzel MD. Transrectal ultrasonographic diagnosis of ascending placentitis in the mare: a report of two cases. Equine Vet Educ 1999;11(2):69–74. Renaudin CD, Troedsson MT, Gillis CL. Transrectal ultrasonographic evaluation of the normal equine placenta. Equine Vet Educ 1999;11(2):75–6. Troedsson MHT, Renaudin CD, Zent WW, Steiner JV. Transrectal ultrasonography of the placenta in normal
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mares and mares with pending abortion. Proc Am Assoc Equine Pract 1997;43:256–8. Zent WW, Williams NM, Donahue JM. Placentitis in Central Kentucky broodmares. Pferdeheilkunde 1999;15(6):630–2. Troedsson MH. Ultrasonographic evaluation of the equine placenta. Pferdeheilkunde 2000;17(6):583. Sebastian M, Gantz MG, Tobin T, Harkins JD, Bosken JM, Hughes C, Harrison LR, Bernard WV, Richter DL, Fitzgerald TDl. The mare reproductive loss syndrome and the eastern tent caterpillar: A toxicokinetic/statistical analysis with clinical, epidemiologic, and mechanistic implications. Vet Ther 2003;4(4): 324–39. Todhunter KH, Perkins NR, Wylie RM, Chicken C, Blishen AJ, Racklyeft DJ, Muscatello G, Wilson MC, Adams PL, Gilkerson JR, Bruden WL, Bagg AP. Equine amnionitis and fetal loss: the case definition for an unrecognised cause of abortion in mares. Aust Vet J 2009;87(1):35–8. Betteridge KJ, Mitchell D. Direct evidence of retention of unfertilized ova in the oviduct of the mare. J Reprod Fertil 1974;39:145–8. Betteridge KJ, Eaglesome MD, Flood PF. Embryo transport through the mare’s oviduct depends upon cleavage and is independent of the ipsilateral corpus luteum. J Reprod Fertil Suppl 1979;(27):387–94. Flood PF, Jong A, Betteridge KJ. The location of eggs retained in the oviducts of mares. J Reprod Fertil 1979;57:291–4. van Niekerk CH, Gerneke WH. Persistence and parthenogenic cleavage of tubal ova in the mare. Onderstepoort J Vet Res 1966;31:195–232. Hershman L, Douglas RH. The critical period for the maternal recognition of pregnancy in pony mares. J Reprod Fertil Suppl 1979;(27):395–401. Zavy MT, Vernon MW, Asquith RL, Bazer FW, Sharp DC. Effect of exogenous gonadal steroids and pregnancy on uterine luminal prostaglandin F in mares. Prostaglandins 1984;27:311–20. Zavy MT, Vernon MW, Sharp DC3, Bazer FW. Endocrine aspects of early pregnancy in pony mares: a comparison of uterine luminal and peripheral plasma levels of steroids during the estrous cycle and early pregnancy. Biomed Tech (Berlin) 1984;115:214–19. Terqui M, Palmer E. Oestrogen pattern during early pregnancy in the mare. J Reprod Fertil Suppl 1979;27:441–6. Nett TM, Holtan DW, Estergreen VL. Oestrogens, LH, PMSG and prolactin in serum of pregnant mares. J Reprod Fertil Suppl 1975;23:457–62. Raeside JI, Liptrap RM, Milne FJ. Relationship of fetal gonads to urinary estrogen excretion by the pregnant mare. Am J Vet Res 1973;34:843–5. Jeffcott LB, Hyland JH, Maclean AA, Dyke T, RobertsonSmith G. Changes in maternal hormone concentrations associated with induction of fetal death at day 45 of gestation in mares. J Reprod Fertil Suppl 1987;35:461–7. Hyland JH, Wright PJ, Manning SJ. An investigation of the use of plasma oestrone sulphate concentrations for the diagnosis of pregnancy in mares. Aust Vet J 1984;61: 123. Bamberg E, Choi HS, Mostl E, Wurm W, Lorin D, Arbeiter K. Enzymatic determination of unconjugated oestrogens in faeces for pregnancy diagnosis in mares. Equine Vet J 1984;16:537–9.
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Maintenance of Pregnancy 62. Monfort SL, Arthur NP, Wildt DE. Monitoring ovarian function and pregnancy by evaluating excretion of urinary estrogen conjugates in semi-free-ranging Przewalski’s horses (Equus przewalskii). J Reprod Fertil 1991;91:155–64. 63. Kasman LH, Hughes JP, Stabenfeldt GH, Starr MD, Lasley BL. Estrone sulfate concentrations as an indicator of fetal demise in horses. Am J Vet Res 1988;49:184–7. 64. Hyland JH, Maclean AA, Robertson-Smith GR, Jeffcott LB, Stewart GA. Attempted conversion of twin to singleton pregnancy in two mares with associated changes in plasma oestrone sulphate concentrations. Aust Vet J 1985;62:406–9. 65. Madej A, Kindahl H, Nydahl C, Edqvist LE, Stewart DR. Hormonal changes associated with induced late abortions in the mare. J Reprod Fertil Suppl 1987;35:479–84. 66. Kirkpatrick JF, Lasley BL, Shideler SE, Roser JF, Turner JW. Non-instrumented immunoassay field tests for pregnancy detection in free-roaming feral horses. J Wildlife Management 1993;57:168–73. 67. Santschi EM, LeBlanc MM, Weston PG. Progestagen, oestrone sulphate and cortisol concentrations in pregnantmares during medical and surgical disease. J Reprod Fertil Suppl 1991;44:627–34. 68. Allen WR. Hormonal control of early pregnancy in the mare. Vet Clin North Am Large Anim Pract 1980;2:291–302. 69. Allen WE. Pregnancy failure induced by human chorionic gonadotrophin in pony mares. Vet Rec 1975;96:88–90. 70. Bergfelt DR, Pierson RA, Ginther OJ. Resurgence of the primary corpus luteum during pregnancy in the mare. Anim Reprod Sci 1989;21:261–70. 71. Squires EL, Voss JL, Villahoz MD. Immunological methods for pregnancy detection in mares. Proc Am Assoc Equine Pract 1983;45–51. 72. Holtan DW, Squires EL, Lapin DR, Ginther OJ. Effect of ovariectomy on pregnancy in mares. J Reprod Fertil Suppl 1979;(27):457–63. 73. Shideler RK, Squires EL, Voss JL, Eikenberry DJ. Exogenous progestin therapy for maintenance of pregnancy in ovariectomized mares. Proc Am Assoc Equine Pract 1981;27:211–19. 74. Shideler RK, Squires EL, Voss JL, Eikenberry DJ, Pickett BW. Progestagen therapy of ovariectomized pregnant mares. J Reprod Fertil Suppl 1982;32:459–64. 75. McKinnon AO, Squires EL, Carnevale EM, Hermenet MJ. Ovariectomized steroid-treated mares as embryo transfer recipients and as a model to study the role of progestins in pregnancy maintenance. Theriogenology 1988;29:1055–64. 76. Naden J, Squires EL, Nett TM. Effect of maternal treatment with altrenogest on age at puberty, hormone concentrations, pituitary response to exogenous GnRH, estrous cycle characteristics and fertility of fillies. J Reprod Fertil 1990;88:185–96. 77. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares:Technique and preliminary findings. Proc Am Assoc Equine Pract 1975;327–35. 78. Studdert MJ. Vaccination of foals and pregnant mares with Duvaxyn EHV1, 4 vaccine. Vaccine 2002;20(7–8):992. 79. Lowis TC, Hyland JH. Analysis of post-partum fertility in mares on a Thoroughbred stud in southern Victoria. Aust Vet J 1991;68:304–6. 80. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproduc-
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tive tract of mares after parturition: effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed first postpartum ovulatory cycles. J Am Vet Med Assoc 1988;192:350–3. McKinnon AO, Squires EL, Carnevale EM, Harrison LA, Frantz DD, McChesney AE, Shideler RK. Diagnostic ultrasonography of uterine pathology in the mare. Proc Am Assoc Equine Pract 1987;605–22. Adams GP, Kastelic JP, Bergfelt DR, Ginther OJ. Effect of uterine inflammation and ultrasonically-detected uterine pathology on fertility in the mare. J Reprod Fertil Suppl 1987;35:445–54. Loy RG, Evans MJ, Pemstein R, Taylor TB. Effects of injected ovarian steroids on reproductive patterns and performance in post-partum mares. J Reprod Fertil Suppl 1982;32:199–204. Pope AM, Campbell DL, Davidson JP. Endometrial histology and post-partum mares treated with progesterone and synthetic GnRH (AY-24,031). J Reprod Fertil Suppl 1979;(27):587–91. Sexton PE, Bristol FM. Uterine involution in mares treated with progesterone and estradiol-17beta. J Am Vet Med Assoc 1985;186:252–6. Squires EL, Heesemann CP, Webel SK, Shideler RK, Voss JL. Relationship of altrenogest to ovarian activity, hormone concentrations and fertility of mares. J Anim Sci 1983;56:901–10. Evans MJ, Hamer JM, Gason LM, Graham CS, Asbury AC, Irvine CHG. Clearance of bacteria and non-antigenic markers following intra- uterine inoculation into maiden mares: Effect of steroid hormone environment. Theriogenology 1986;26:37–50. Winter AJ. Microbial immunity in the reproductive tract. J Am Vet Med Assoc 1982;181:1069–73. Allen WR. Progesterone and the pregnant mare: unanswered chestnuts [editorial;comment]. Equine Vet J 1993;25:90–1. Wilker C, Daels PF, Burns PJ, Ball BA. Progesterone therapy during early pregnancy in the mare. Proc Am Assoc Equine Pract 1991;161–72. Irvine CHG, Sutton P, Turner JE, Mennick PE. Changes in plasma progesterone concentrations from days 17 to 42 of gestation in mares maintaining or losing pregnancy. Equine Vet J 1990;22:104–6. Bergfelt DR, Ginther OJ. Embryo loss following GnRHinduced ovulation in anovulatory mares. Theriogenology 1992;38:33–43. McCue PM, Troedsson MH, Liu IK, Stabenfeldt GH, Hughes JP, Lasley BL. Follicular and endocrine responses of anoestrous mares to administration ofnative GnRH or a GnRH agonist. J Reprod Fertil Suppl 1991;44: 227–33. Bergfelt DR, Woods JA, Ginther OJ. Role of the embryonic vesicle and progesterone in embryonic loss in mares. J Reprod Fertil 1992;95:339–47. McKinnon AO, Tarrida del Marmol Figueroa S, Nobelius AM, Hyland JH, Vasey JR. Failure of hydroxyprogesterone caproate to maintain pregnancy in ovariectomized mares. Equine Vet J 1993;25:158–60. McKinnon AO, Lescun TB, Walker JH, Vasey JR, Allen WR. The inability of some synthetic progestagens to maintain pregnancy in the mare. Equine Vet J 2000;32(1): 83–5. Allen WR, Sanderson MW. The value of a rapid progesterone assay (AELIA) in equine stud veterinary medicine
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and management. Proc Bain-Fallon Mem Lect, Sydney 1987;76. Holtan DW, Squires EL, Ginther OJ. Effect of ovariectomy on pregnancy in mares. J Anim Sci 1975;41:359. Knowles JE, Squires EL, Shideler RK, Tarr SF, Nett TM. Progestins in mid-pregnant to late-pregnant mares. J Equine Vet Sci 1994;14:659–63. Nishikawa Y. Studies on reproduction in horses: Singularity and artificial control in reproductive phenomena. Tokyo: Japan Racing Association, 1959. Varner DD, Blanchard TL, Brinsko SP. Estrogens, osytocin and ergot alkaloids – uses in reproductive management of mares. Proc Am Assoc Equine Pract 1988;219–41. Nett TM, Pickett BW. Effect of diethylstilboestrol on the relationship between LH, PMSG and progesterone during pregnancy in the mare. J Reprod Fertil Suppl 1979;(27):465–70. Pashen RL, Allen WR. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J Reprod Fertil Suppl 1979;(27):499–509. Pycock JF, Newcombe JR. The effect of the gonadotropinreleasing-hormone analog, buserelin, administered in diestrus on pregnancy rates and pregnancy failure in mares. Theriogenology 1996;46:1097–101. Newcombe JR, Martinez TA, Peters AR. The effect of the gonadotropin-releasing hormone analog, buserelin, on pregnancy rates in horse and pony mares. Theriogenology 2001;55(8):1619–31. Unger C, Schneider F, Hoppen HO, Becker F, Nurnberg G, Kanitz W. Pregnancy rates in mares after application of gonadotropin releasing hormone analogue, Buserelin, in warmblood mares. Theriogenology 2002;58(2–4):635–8. McKinnon AO, Voss JL. Breeding the problem mare. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, London: Lea & Febiger, 1993; pp. 368–78. McKinnon AO. Reproductive efficiency of horses in Australia. http://www.gvequine.com.au/breeding efficiency.htm. 2000.
109. McKinnon AO. Why is my mare loosing her pregnancy? Equine Research Seminar,119–144. 1998. University of Sydney, Post Graduate Foundation in Veterinary Science, Sydney. Equine Research Seminar 98. 110. Gilbert RO, Marlow CHB. A field study of patterns of unobserved foetal loss as determined by rectal palpation in foaling, barren and maiden thoroughbred mares. Equine Vet J 1992;24:184–6. 111. Davies Morel MC, Newcombe JR, Hinchliffe J. The relationship between consecutive pregnancies in Thoroughbred mares. Does the location of one pregnancy affect the location of the next, is this affected by mare age and foal heat to conception interval or related to pregnancy success. Theriogenology 2009;71(7):1072–8. 112. Santschi EM, LeBlanc MM. Fetal and placental conditions that cause high-risk pregnancy in mares. Compend Cont Educ Pract Vet 1995;17:710–21. 113. Zingher AC. Evaluation of a Propionobacterium acnes immunostimulant in the treatment of equine endometritis. Proc Am Assoc Equine Pract 1993;171–2. 114. Zingher AC. Effects of immunostimulation with PropioniR ) in mares cytologically positive bacterium acnes (Eqstim for endometritis. J Equine Vet Sci 1996;16:100–3. 115. Rohrbach BW, Sheerin PC, Cantrell CK, Matthews PM, Steiner JV, Dodds LE. Effect of adjunctive treatment with intravenously administered Propionibacterium acnes on reproductive performance in mares with persistent endometritis. J Am Vet Med Assoc 2007;231(1): 107–13. 116. Fumuso E, Gigure S, Wade J, Rogan D, VidelaDorna I, Bowden RA. Endometrial IL-1 beta, IL-6 and TNF-alpha, mRNA expression in mares resistant or susceptible to post-breeding endometritis – Effects of estrous cycle, artificial insemination and immunomodulation. Vet Immunol Immunopathol 2003;96(1–2):31–41. 117. Fumuso EA, Aguilar J, Giguere S, Rivulgo M, Wade J, Rogan D. Immune parameters in mares resistant and susceptible to persistent post-breeding endometritis: effects of immunomodulation. Vet Immunol Immunopathol 2007;118(1–2):30–9.
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Chapter 255
Dystocia Management Grant Frazer
Introduction In any informed discussion of obstetrics it is essential that there be a uniform appreciation of the terminology. The term ‘breech presentation’ is often used as if it were synonymous with ‘posterior presentation’. It actually refers to a postural abnormality in which the posterior presentation is complicated by bilateral hip flexion with retention of both hindlimbs. Likewise, the term ‘retained placenta’ is part of the common vernacular, but it is actually a misnomer. The correct term for prolonged stage III of labor is ‘retention of the fetal membranes’ since if the endometrial portion of the placenta were not retained there would be a uterine prolapse. The terms malpresented, or malpositioned, fetus are often used erroneously when the intent is to convey that the fetal head and/or neck – or a limb(s) – was not correctly aligned within the birth canal. While the words anterior and posterior are still widely used in the veterinary literature, the correct Nomina Anatomica Veterinaria (NAV) terms are cranial and caudal.1 The word dystocia implies that there is some impediment to the normal parturient process – be it maternal (e.g., uterine torsion, abdominal hernia) or fetal (size, presentation, position, posture) in origin.2, 3 Vandeplassche suggested that it may be preferable to note that the fetus is maldisposed (fetal maldisposition) since this is an all-encompassing term that implies any combination of abnormalities – be it in presentation, position and/or posture.4
r r
Presentation – the orientation of the fetal spinal axis to that of the mare, and also the portion of the fetus that enters the vaginal canal first (cranial or caudal longitudinal; ventro- or dorso-transverse). Position – the relationship of the fetal dorsum (longitudinal presentation) or head (transverse presentation) to the quadrants of the mare’s pelvis (dorsosacral, right/left dorsoilial, dorsopubic; right/left cephalo-ilial).
r
r r r
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Posture – the relationship of the fetal extremities (head, neck, limbs) to its body. Postural abnormalities are the most common cause of equine dystocia, and can be difficult to correct due to the remarkably long head, neck and limbs, of the foal. Repulsion – the procedure whereby the fetus is repelled from the pelvic inlet such that the increased space created in the uterus can be utilized to correct the maldisposition.5 Mutation – manipulation of the fetal extremities, together with correction of any positional abnormalities, to assist vaginal delivery. Version – manual changing of the polarity of the fetus with reference to the mother. It is the manipulation that is employed when attempting to correct a transverse presentation. The head and forelimbs are repelled into the uterus while the hindquarters are pulled into the pelvic canal.5 Tocolysis – the administration of agents (clenbuterol, isoxsuprine) that inhibit myometrial contractions and thus relax the uterus. In many countries these tocolytic agents are administered by injection to provide more space for obstetrical manipulations.5, 6
In-utero fetal kinetics Fetal rotation within the amniotic cavity, and amniotic sac rotation within the allantoic cavity, results in the characteristic twisting of the equine umbilical cord.7–9 Between 5–7 months of gestation it appears that contraction of the horns serves to confine the allantoic fluid and fetus to the uterine body.8,10–13 It has been theorized that neurological signals within the maturing inner ear may orient the dorsum of the fetus with the concave slope of the ventral uterine wall, and prompt it to lie with the cranial aspect elevated towards the cervix.10–14 The acute angle between the horn and body by 7 months of gestation means that the fetus must be in dorsal recumbency before the hindlimbs can gain entry. After 9 months
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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the hindlimbs remain enclosed within the uterine horn, and the hooves reach to the tip by the tenth month.8,10–13 After this time the fetus is locked into a cranial presentation and dorsopubic position and this likely explains how 98.9% of foals are delivered in cranial presentation, with 1.0% being caudal, and only 0.1% being transverse.4, 15 A classic radiographic study demonstrated that the full-term equine fetus is lying in a dorsopubic position with the head, neck, and forelimbs flexed.16 It is still possible for the entire pregnancy (uterus and fetus) to rotate approximately 90◦ on the lower maternal abdominal wall. This occurs because any rotational movement of the caudal half of the fetus (pelvis and hindlimbs) by necessity will involve the close-fitting uterus. In extreme cases this rotation may progress into a clinical uterine torsion.1, 2, 13, 17 Close to term the horn containing the hindlimbs comes to rest on the dorsal surface of the uterine body, with the tip of the horn directed towards the cervix.11, 13 Thus, the uterine body and gravid horn form what can be described as a U-shaped tube that tapers towards the tip. In some cases it is possible for the hindlimb hooves in the horn tip to be pushed so far caudally that they actually come to lie over the fetal head. Thus, when performing a per rectum evaluation of the pregnancy close to term, the fetal hooves that are palpable may sometimes belong to the hindlimbs (see Figure 255.1). Vigorous piston-like thrusts of the hindlimbs, in association with elevation of the fetal rump, can result in the hooves being pushed well past the cervix into the rectogenital
Figure 255.1 The horn containing the hindlimbs comes to rest on the dorsal surface of the uterine body in late gestation, such that the tip of the horn is directed towards the cervix. As the fetus moves, the hindlimbs may push the horn tip caudally such that the hindlimb hooves come to lie over the fetal head. Although the forelimbs and neck are usually flexed, on some occasions the head or forelimbs may be extended such that the limbs are readily palpable above the fetal head.
pouch.11–13 This observation may explain the acute colic episodes that have been previously attributed to uterine dorso-retroflexion.6 Although the gel-like pads on the fetal hooves may serve to protect the placenta and uterine wall from the vigorous activity of the hindlimbs, tears at the tip of the gravid horn can still occur during an unassisted foaling.1, 13,18–20
Signs of impending parturition Some mares will develop a ventral, edematous plaque of variable thickness (2 to 4 inches or 50–100 mm) in late gestation. This is a physiological phenomenon that can result from the increased weight of the gravid uterus compressing lymphatic drainage and venous return from the ventral abdomen. The swelling must be differentiated from the painful inflammatory edema that is associated with impending – or actual – ventral body wall rupture (prepubic tendon, ventral hernia). In cases of rupture the mare may be reluctant to move, the mammary secretions will often contain blood, and ultrasonography may demonstrate tearing and edema within the musculature. If the prepubic tendon has ruptured the mare will develop a lordosis due to tilting of the pelvic floor. A canvas abdominal sling may provide sufficient support to permit some of these mares to carry the foal to term.1, 2 While changes in the hormonal milieu during the last weeks and days of gestation result in physical changes in the mare, the actual signs of impending parturition are quite variable between mares. The pregnant mare’s udder may start to increase in size during the last 4 to 6 weeks of gestation, and it generally becomes prominent within 2 weeks of foaling. The size of the udder is influenced by the parity of the mare, and some maiden mares will foal with minimal mammary changes. Although it may not be readily apparent in a well-muscled mare, relaxation of the sacrosciatic ligaments can cause a discernable hollow to develop on either side of the tail head. While this change may be easier to discern in older mares, a palpable softening of the ligaments can usually be detected. Often any elongation and relaxation of the vulva may not be noticeable until foaling is imminent.1 Even though the most notable prepartum change is usually in the mammary gland, the onset of secretory activity is quite variable. Some mares may be observed to develop wax-like material on the teat orifices (‘wax up’) several days before foaling, while others may never develop these honey-colored beads of dried secretion on the teat orifices. The teats usually fill with colostrum 24 to 48 hours before foaling, but some mares may drip or ‘run’ milk for
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Dystocia Management several days prior to the onset of parturition. In these cases a substantial volume of colostrum may be lost, and plans should be made to supplement the foal’s immunoglobulin intake. Thus, while the onset of secretory activity can be a useful guide it cannot be used as a reliable indicator of impending parturition. However, electrolyte changes in mammary secretions can be used to predict ‘readiness for birth’.21–23 The clinician must remember that these electrolyte changes are not a reliable indicator of fetal maturity if a placental anomaly is present (e.g., placentitis, twins), and premature mammary development may be the first sign of an impending abortion.1
The normal parturient process Although the caudal aspect of the fetus is in a dorsopubic position and intimately associated with the ventral uterine wall, the cranial portion has room to rotate within the uterine body itself. Recent ultrasonographic studies have shown that in a mare close to term (> 330 days’ gestation) the cranial aspect of the fetus may be in dorsopubic position approximately 60% of the time, and in dorsoilial position during about 40% of observations. Although the forelimbs and neck are usually flexed (about 80%), on some occasions the head or limbs may be extended.12, 13 Thus, palpation of the fetus cannot be used to detect impending dystocia (postural or positional complications) unless labor is already well advanced. However, if a transverse or caudal presentation is suspected then a detailed transabdominal ultrasonographic evaluation is indicated to confirm the location of the fetal thorax and head. In both cases planned intervention may be warranted. In some cases transabdominal ultrasonography may suggest that a fetus is transversely presented, yet the foal will be delivered normally. The so-called ‘body pregnancy’ tends to cause abortion in the latter stages of gestation when the nutritional demands of the growing fetus are no longer being met. In these cases the placenta is underdeveloped, and is characterized by the presence of short horns. It is this lack of development of a gravid horn that causes intrauterine fetal growth retardation and abortion due to placental insufficiency.1, 24, 25 In order to quickly recognize that something is abnormal, owners and farm managers must be fully aware of all the characteristics of the normal parturient process. The vast majority of mares deliver their foals during the evening hours. Mares vary in their display of symptoms that are consistent with the onset of first-stage labor. The first stage of parturition is characterized by the development of coordinated uterine contractions that create increased intrauterine pressure. In some, the increasing intensity of the
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uterine contractions may be manifest by restless behavior similar to that of mild colic. Patchy sweating often develops, and some mares expel colostrum. The mare may look at her flank, frequently lie down and get up, stretch as if to urinate, and pass small amounts of feces.5 In some mares a normal delivery may be preceded by one or more episodes of false labor. Although the fetus probably determines the day, the mare appears to be able to choose the hour for delivery. It is not uncommon for labor signs to abate if the mare is unduly disturbed by the external environment. Thus, it is important that supervision of foaling mares be unobtrusive. Any untoward activity may lead to a stress response and suppression of uterine contractions – especially in nervous mares.1 Some clinicians report that on farms with significant night-time activity there may be an increase in the number of daytime deliveries, often soon after the mares are returned to pasture from the foaling yards. The increasing uterine tone during stage I of parturition pushes the ‘cervical star’ region of the chorioallantoic sac against the hormonally relaxed cervix such that gradual dilation occurs. It has been proposed that the uterine contractions somehow stimulate the fetus to play an active role in positioning itself for delivery, particularly the extension of forelimbs. The characteristic side-to-side rolling each time the mare assumes lateral recumbency is also thought to assist in positioning the foal correctly. It makes purposeful movements such that the cranial aspect rotates through a dorsoilial position as the head and forelimbs are extended up into the birth canal13, 16 (see Figure 255.2). Dystocia cases that involve a dorsopubic position with flexed extremities suggest that the fetus was compromised prior to, or very early in, the parturient process.5, 13, 16, 26 In some
Figure 255.2 During a dystocia a dead fetus may be found in dorsopubic position with flexed neck and limbs. The fetus must be healthy and viable to be able to make purposeful movements that rotate the cranial aspect through a dorsoilial position while the head and forelimbs are extended up into the birth canal.
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cases this may be indicative of advanced placentitis or fetal infection, and thus submission of tissue from a stillborn fetus and attendant membranes is recommended.24 The neck and forelimbs will not normally be returned to a flexed posture once a viable fetus in stage I of delivery has actively extended them. However, fetlock flexion can develop if the hoof catches on the pelvic brim or on a fold of vaginal mucosa. Continued expulsive efforts from the mare can force the carpus to flex as well.1, 5 Rupture of the chorioallantois and discharge of the watery allantoic fluid marks the onset of stage II of labor. This occurs when the fetlocks – and sometimes the knees – are at the level of the external cervical opening.13 The second stage of labor is characterized by strong abdominal contractions that provide the expulsive force necessary to expel the fetus. Second-stage labor in the mare is rapid, with most foals being delivered within 20–30 minutes after the chorioallantoic membrane ruptures. Passage of the foal into the pelvic inlet initiates a reflex release of oxytocin from the posterior pituitary, thereby enhancing uterine contractility.5 Most mares assume lateral recumbency once active straining commences, although it is not uncommon for the mare to get up once or twice during second-stage labor. The most forceful contractions occur as the fetal thorax passes through the pelvic cavity. Pre-existing defects in the abdominal musculature (prepubic tendon rupture; ventral hernia) adversely impact on the mare’s ability to mount sufficient intra-abdominal pressure to expel the foal without assistance. These mares warrant close supervision since traction may need to be applied to the foal’s forelimbs once stage II of labor has commenced.1, 2 Appearance of one hoof within the translucent fluid-filled amnion at the vulvar lips can be expected to occur within 5 minutes of rupture of the chorioallantois.27 A dark yellowish-green discoloration of the amnion and fluid is indicative of passage of meconium due to fetal stress. One forelimb should precede the other by several inches, such that the shoulders enter the pelvis successively. The measurement across the shoulders represents the widest portion of the foal. Thus, the tendency for one limb to advance ahead of the other ensures that the cross-sectional diameter of the fetus at the shoulders is reduced. The soles of the hooves should be directed downward and the foal’s head should be resting between the carpi, with the muzzle reaching the proximal metacarpus. By the time the nose reaches the vulva the cranial half of the torso has already rotated from a dorsopubic to a dorsoilial position. The foal’s withers continue to rotate into a dorsosacral position as the head appears through the vulvar lips.13 At this time the foal’s shoulders have entered the pelvic canal.1
While the amniotic sac may rupture during the mare’s expulsive efforts, the foal can be delivered with a portion of the sac wrapped around its head.27 Although a viable foal should be able to free itself of the fetal membranes, foaling attendants should be instructed to free the foal’s head promptly to avoid the possibility of suffocation. While the cranial aspect of the foal is passing through the birth canal the hindlimbs remain locked within the uterine horn. The fetal pelvis rotates through a dorsoilial position into a dorsosacral position. Once the fetal abdomen begins to pass through the vulvar lips the stifles engage the pelvic brim and this causes the hindlimbs to extend. The foal’s rump remains closely apposed to the cranial dome of the uterine body as the contracting uterus and abdominal press combine to expel the foal. Forceful straining ceases once the foal’s hips are delivered, and most mares will rest in lateral recumbency for several minutes. An active foal will extract its hindlimbs as it struggles to stand. At the time of the foal’s delivery the cranial aspect of the contracting uterine body is only about 12 inches from the cervix.1, 13
Detecting foaling problems The incidence of dystocia varies among breeds, with estimates ranging from 4% in Thoroughbreds to as much as 1 in 10 foalings in the heavily muscled Draft breeds. Hydrocephalus can occur in all breeds. Ponies are more likely to experience difficulties due to a large fetal skull that may resemble hydrocephaly in some cases.4, 28, 29 Primiparous dams generally require a longer time to expel the fetus than multiparous dams.5, 27 Delayed delivery (prolonged foaling time) increases the likelihood of fetal asphyxia, or neonatal problems associated with hypoxia due to placental separation.24, 30 Often these losses occur in mares that were unattended at the time of foaling. The short, explosive birth process, and the mare’s tendency to foal at night, increases the probability of unattended delivery.30 Prepartum mares warrant close attention since careful monitoring of parturition may help to reduce the incidence of deaths attributable to neonatal asphyxia. A number of foaling monitors and video camera systems are available to alert personnel that foaling is imminent.1, 31 Attendants should suspect that the mare is experiencing obstetrical problems if either the first or second stage of parturition is prolonged, or not progressive. Occasionally, the chorioallantois fails to rupture and the velvety, red membrane (‘red bag’) appears at the vulvar lips. This is a common complication of induced parturition, and placental edema is a major problem in mares that have been exposed to endophyte-infected fescue hay or pasture.32, 33 In
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Dystocia Management the latter scenario the pregnancy may be prolonged, and the mare is also likely to be agalactic. Premature separation of the placenta is an emergency situation because the foal is deprived of its maternal oxygen supply. Foaling attendants should be instructed to break the membrane, and to provide gentle traction in unison with the mare’s expulsive efforts. Although the foal should be delivered as quickly as possible, injudicious traction at this time can create a laceration or fetal rib fractures in an incompletely dilated cervix.1, 3 Time is the critical factor when managing an equine obstetrical case.34 Experienced personnel can save many foals by early recognition of dystocias, and by prompt intervention before serious complications arise. If the cause can be easily, quickly, and atraumatically corrected by the attendant then this should be done without delay. The secret to foal survival is a combination of experience and expediency. If farm personnel are experienced, and referring veterinarians are well-trained in dystocia management, prompt referral to a hospital in close proximity may permit delivery of a viable foal – even after an hour has elapsed since the chorioallantois ruptured.34, 35 It is probable that limited intervention causes less straining and thus less disruption of the fetal membranes and thus permits longer placental support of the fetus. Signs that a mare may be experiencing difficulties include failure of any fetal parts, or of the amniotic membrane, to appear at the vulval lips within 5 minutes after rupture of the chorioallantois. If there is no evidence of strong contractions, and/or no progression in the delivery process within 10 minutes of rupture of the chorioallantois, then a vaginal examination is warranted. Other causes for concern are hooves upside-down, or any abnormal combination of extremities appearing at the vulva (only one limb; hooves and nose in abnormal relationship; nose but not hooves; both forelimbs protruding but no head visible at the level of the carpi).27 The most common impediments to delivery are malpostures of the long fetal extremities (head and neck, or limbs).4, 5, 26 Failure to quickly identify a dystocia often results in the fetus becoming impacted in the birth canal by the mare’s uterine contractions and strong abdominal straining.1
Clinical examination of the parturient mare It is prudent for an equine practitioner to have a well-organized dystocia kit readily accessible during the foaling season. Supplies necessary to provide cardiovascular and respiratory support to a compromised neonate should also be readily available. The
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obstetrical equipment should be clean and preferably sterile. A nasogastric tube, pump, and bucket should be kept exclusively for delivering volume-expanding lubricant (e.g., carboxymethylcellulose) into the uterus. This is especially important in those countries (e.g., the United States) where injectable tocolytic agents (clenbuterol, isoxsuprine) are not available for obstetrical use.5 Obstetrical chains, nylon ropes, and handles are the basic requirements, and blunt eye hooks may occasionally be indicated.3, 5, 36 A detorsion rod, an obstetrical snare, and a Kuhn’s crutch repeller (with long nylon rope) can be useful in some instances, but require additional expertise.5, 37 It goes without saying that these metal instruments can easily traumatize the reproductive tract and should be used with caution. The mechanical extractive devices that are often used in bovine practice can exert inordinate force and in the author’s opinion they do not have a place in equine obstetrics. However, prior experience in resolving bovine dystocias will be invaluable when a veterinarian is faced with the long extremities and forceful straining that inevitably complicate the resolution of equine obstetrical cases. A minimum of two people should be available to assist the obstetrician: one to hold the mare’s head and at least one other individual to hold instruments, and to assist with fetal delivery.1 Two experienced veterinarians should be available for induced foalings. Dystocia in a mare causes stress for both the owner and the veterinarian. These cases should always be regarded as an emergency, especially if there is a possibility that the foal is still alive. The veterinarian must simultaneously consider the best interests of two patients and a sometimes emotional owner. Apart from potential loss of the foal, the mare’s future fertility can be easily compromised, and in some cases the mare may even suffer fatal complications. Initially the veterinarian should make a cursory assessment of the mare’s general physical condition noting, in particular, mucous membrane color and refill time (hemorrhage, shock).3 The perineal area should be inspected for the presence and nature of any vulvar discharge, the presence of fetal membranes, and to identify any fetal extremities. Excessive hemorrhage is indicative of soft tissue trauma. Moist fetal extremities may help to confirm that assistance was summoned promptly, whereas dry and swollen tissues are typical of more protracted cases. A malodorous discharge strongly suggests the presence of an emphysematous fetus – an uncommon occurrence that may be encountered in neglected cases.5 Occasionally a foaling mare may present with a rectal prolapse, an everted bladder, or with loops of bowel (small intestine, colon) protruding from the vulvar lips.3 While performing this external examination of the mare the following questions
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should be posed to the owner – age and parity number; expected foaling date; duration and intensity of labor signs; time since rupture of the chorioallantois; appearance of fetal membranes, limbs, or muzzle at vulva; and any attempts at manual intervention? It is not uncommon to be informed that the muzzle was present when an attempt was made to correct a simple limb malposture. Unfortunately these cases have often been complicated by a flexed head and neck posture before veterinary assistance is requested. Reports of fetal movement observed in the mare’s flanks are often unreliable since the mare’s expulsive efforts can easily cause deceptive ‘movement’ of exposed extremities in what is already a dead fetus.1 This author believes that the use of rigid, closedend stocks is contraindicated because of the propensity for parturient mares to become recumbent. However, adequate restraint is essential because the behavior of a mare in the second stage of labor is so unpredictable – and it can be violent. If used, the stocks should be open-ended with movable sides. The internal examination should be performed in a clean area with good footing, and with sufficient room to ensure safety for all personnel involved. Whenever possible the author prefers to perform the initial examination on a standing mare in a large, well-bedded stall – with nothing more than a twitch or lip chain for restraint. The person handling the mare should stand on the same side as the obstetrician, and be instructed to apply intermittent tension to the twitch or lip chain when asked to do so. If the mare attempts to kick, the handler should know to pull the mare’s head towards them such that the hindquarters will move away from the obstetrician. Although not essential, an initial rectal examination may rule out the presence of a term uterine torsion, determine the condition of the uterine wall (tears, spasm), and may provide useful information regarding the disposition of the fetus.3, 17 Prior to any vaginal examination the mare’s tail should be wrapped and the perineal area thoroughly cleansed. The clinician’s arms and hands should be scrubbed with disinfectant soap. The use of sterile sleeves is optional. The author routinely uses shoulder-length rubber sleeves with surgical gloves attached. Although disposable plastic sleeves decrease tactile sensation, experienced obstetricians believe that the use of a sleeve reduces the amount of trauma inflicted upon the vaginal mucous membranes. Liberal application of lubricant is essential since the mare’s vagina and cervix are easily traumatized, and future fertility can be jeopardized by the resultant adhesions and fibrosis. White petroleum jelly (Vaseline) is useful when performing the initial exploratory examination. If there is evidence of prior vaginal manipulations it may be prudent to perform an abdominocentesis
to ensure uterine integrity before instilling large volumes of liquid obstetrical lubricant.1, 20, 38 A rapid, thorough check should be made for lesions, lacerations, and contusions of the genital tract. The presence of any pelvic abnormalities that may impede passage of the fetus should be noted, the degree of cervical relaxation determined, and an assessment made as to whether or not the uterus is tightly contracted around the fetus. Although fetopelvic disproportion is uncommon in the mare, it can be a factor in some equine dystocias, especially those in primiparous mares.24, 30, 39, 40 The disposition of the fetus should be noted (presentation, position, and posture), and fetal viability determined.3 Care should be exercised because an active fetal response to manipulations can easily complicate what was initially a simple dystocia. This is often the cause of the more extreme head and neck malpostures.3, 26, 36 Placement of a rope snare behind the ears and into the mouth will ensure that the clinician always has control of the head. This will facilitate easy correction of a potentially life-threatening development such as lateral deviation of the head and neck if the fetus pulls away from the clinician’s arm. When obvious fetal movement is absent, limb withdrawal may be initiated in response to pinching of the coronary band. Slight digital pressure through the eyelid may arouse a response, as may stimulation of the tongue (swallowing). If the thorax can be reached, fetal heartbeat is definitive. In caudally presented cases, the digital and anal reflexes are useful as indicators of fetal viability.3, 5 Failure to elicit a response is not always an absolute confirmation of fetal death, and when there is doubt about fetal viability an ultrasonographic assessment of the fetal heart may be made with a 3.5 MHz transabdominal probe.1 Once the cause of the dystocia has been determined, the obstetrician should ensure that the owner understands the various treatment options and costs, and the inherent risks that may be associated with each approach. The prognosis for survival of both the mare and foal, and the impact of the treatment options on the mare’s future fertility, should also be discussed. Sometimes the owner will place more value on the mare’s non-reproductive attributes, while in other cases the mare’s future fertility is of paramount importance. The economics of the case, the expertise of the clinician, and the proximity of hospital facilities are all factors to be considered when planning the best course of action to resolve a dystocia case.1, 3, 34, 36, 41–43 There is no place for heroics in equine obstetrics. The clinician should recognize their limitations, and be prepared to refer the case, for the mare’s sake.35 The clinician must be decisive, and if a brief manipulation is ineffective then an alternate approach should be considered. If a well-equipped veterinary
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Dystocia Management hospital is close by then immediate referral for general anesthesia, controlled vaginal delivery, or cesarean section may offer the best prospect for foal survival and the mare’s subsequent fertility.34, 42 Prolonged vaginal manipulations are contraindicated if future fertility is an objective. The mucous membranes are easily abraded and will inevitably heal with transluminal adhesions and fibrosis. Repeated insertion and removal of the obstetrician’s arm not only traumatizes the vaginal canal and cervix, but also increases the bacterial load that is carried into the uterus. If the fetus is dead, many malposture cases can be rapidly resolved by one or two fetotomy cuts – provided that the clinician has the appropriate skills and equipment.41, 44, 45 Unfortunately fetotomy is often used as a last resort when prolonged manipulations have failed to correct the problem, and extensive soft tissue trauma has already been inflicted. Poor technique and inappropriate fetotomy cuts often lead to infertility. The alternative is cesarean section.1, 34, 42, 46
Chemical restraint Any tranquilization of a foaling mare should be performed with due consideration for cardiopulmonary function and fetal oxygenation. The ultimate choice of restraint will vary with the obstetrician, and will be determined by such factors as the mare’s demeanor, the complexity of the dystocia, and the expectation of fetal viability. If delivery of a live foal is anticipated, the clinician should consider the potential for altered placental perfusion and fetal cardiovascular compromise before administering any tranquilizers to the mare. Light sedation with acetylpromazine should have minimal effect on the foal. If extra restraint is required, xylazine is preferable to detomidine if the fetus is viable because its depressant effects are of much shorter duration. Xylazine or detomidine should be used on its own to sedate a dystocia case with some caution since some mares sedated with these compounds have been noted to become hypersensitive over the hindquarters. A xylazine-acepromazine combination will provide good sedation in a quiet mare, but a xylazine-butorphanol combination may be preferable if extra sedation and analgesia is necessary. Reduced dosages should be used initially, especially in Draft breeds and Miniatures that appear to respond more intensely to tranquilizers and sedative-hypnotic drugs than light breeds. Since opioid analgesics can cause some gastrointestinal stasis and impaction, it is especially important to administer mineral oil once the dystocia has been resolved if these agents have been used.1, 47, 48 Short-term general anesthesia may be indicated when minor postural abnormalities are present, yet
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maternal expulsive efforts make correction difficult – or the attempts at correction have already exceeded 10 minutes. Although the uterus in an anesthetized mare will be relaxed, further relaxation can still be achieved with administration of tocolytics. Intrauterine manipulations of anesthetized mares in lateral recumbency can still be difficult due to the increased abdominal pressure. The chances for successful resolution are enhanced if a hoist is available since the effects of gravity will greatly assist with fetal repulsion and manipulation. Hobbles should be placed on the hind pasterns, and the mare’s hindquarters briefly elevated some 2 to 3 feet from the floor. A xylazine-ketamine combination provides a smooth, short (10–15 minute) duration general anesthetic, but the addition of guaifenesin can provide an additional 10–20 minutes for fetal manipulation.47 However, the mare should not remain in an elevated, dorsally recumbent position for very long. Apart from respiratory and cardiovascular compromise, hindlimb paresis can develop after prolonged suspension. Some of the mare’s weight can be taken off the limbs by placing supportive pads under the hindquarters. If attempts at correction are successful, the mare should be lowered into lateral recumbency so that the foal can be extracted. In specialist equine hospitals that are located close to well-managed broodmare farms, the fetus is often still alive when the mare arrives. It is common practice to immediately anesthetise these mares, and to maintain them with controlled ventilation and intravenous fluids.34, 49 By using the hind-end elevation technique, almost three-quarters of such cases can be resolved by controlled vaginal delivery. However, if the fetus is still alive and has not been delivered within 15 minutes, a cesarean section is performed. Under these optimal circumstances a 30% foal survival rate has been reported.1, 34, 46, 50 The time involved in administering effective epidural anesthesia often makes this form of restraint impractical if a live foal is to be delivered. The majority of fetotomy procedures (one or two cuts) can be made on a standing mare, and in these cases caudal epidural anesthesia may be used at the clinician’s discretion. Although an epidural does not prevent the mare’s myometrial contractions or the abdominal press, it will reduce vaginal sensitivity, and thus suppresses the Ferguson reflex associated with vaginal manipulations.5, 47 If an epidural is utilized, a xylazine-lidocaine combination is recommended. A low dose is essential to avoid the complication of ataxia – especially if referral to a hospital is a possibility.51 Myometrial contractions (uterine spasm) can be controlled by injectable tocolytic agents (isoxsuprine, clenbuterol) if they are available for veterinary use.6 Although intravenous or intramuscular administration will cause rapid
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uterine relaxation, it is not known whether the oral formulations will achieve sufficient blood levels to provide the same effect on the parturient uterus.1, 52
Fetal manipulation (mutation) Often space limitations within the pelvic canal dictate that the fetus must be repelled deeply into the uterus before it is possible to manipulate the fetal body, and especially the long extremities. However, overzealous intervention is a major cause of uterine rupture and it may not be possible to achieve meaningful – yet safe – repulsion if the uterus is contracted down around the fetus.5, 20, 38,53–59 Tocolytic agents are effective in relaxing a contracted uterus, but injectable formulations are not available in all countries.5, 6 Pumping warm obstetrical lubricant around the fetus will induce some uterine relaxation in most cases, and may provide the extra space that is required to safely mutate the malposture.3, 36 However, it is imperative that the obstetrician first verifies that the uterus is intact, especially if there has been prior intervention by inexperienced personnel.38 Most iatrogenic lacerations occur in the uterine body, whereas a rupture at the tip of the gravid horn is more likely to be caused by the thrusting activity of the fetal hindlimbs.1, 13, 20 If a live fetus is found to be in an abnormal position (dorsopubic or dorsoilial) it may be possible to rotate it by applying lateral pressure over the shoulder. An active fetal response will often facilitate correction into a dorsosacral position. If the foal is dead, and the forelimbs are extended, it may be possible to mimic the normal rotational movements by combining torque with gentle traction on the obstetrical chains. Judicious use of a bovine detorsion rod can also facilitate this maneuver.5 Correction of a malposture of the fetal extremities is the most common reason for intervention. While attempts to extract a fetus with an uncorrected limb malposture pose a risk for both the fetus and mare, it is sometimes possible to be successful if the mare has a large pelvis and the fetus is very small or premature.5 However, the cervix and vagina can easily be lacerated, and extreme pressures created by extraction attempts with an uncorrected malposture can cause fetal rib fractures and severe internal trauma. It should be noted that rib fractures and contusions can also occur during an unassisted delivery.1, 39, 40 Once any malpositions and/or malpostures have been corrected, any traction must be applied with careful regard for both maternal and fetal wellbeing.36 Often traction applied entirely by hand is all that is necessary, although obstetrical straps or chains may provide a better grip. If the chains are applied with a loop above the fetlock and a half hitch below the joint, the extractive force will be spread over a
larger surface and this may decrease the possibility of a crush fracture. The eye of the chain or strap should be positioned on the dorsal aspect of the limbs.5 Any traction that is applied to the head should be merely to correct a malposture and to direct passage through the vaginal canal. A rope head snare (behind ears and into mouth) may be preferable to the use of eye hooks in foals since they are more susceptible to trauma than are calves. This is obviously not an issue if the foal is dead. Although tension on the head snare can force the mouth open, it is unlikely that the incisors will have broken the gum sufficiently to be of any concern. Nevertheless, care should be taken to protect the uterus if retrieval of the head becomes necessary. A mandibular snare is sometimes useful on a viable foal, but it should be used merely to direct the head and not as a point of traction. Mandibular fractures can easily occur if excessive force is applied.1, 5 If the mare is not anesthetized, any traction on the limbs should be applied as an adjunct to her expulsive efforts – assisted vaginal delivery. Traction should cease when the dam stops straining, thereby permitting rest and recovery. This approach is essential to ensure complete cervical dilation, and to permit adequate dilation of the caudal reproductive tract, especially in primiparous mares.36 Copious lubrication and slow traction, with continuous monitoring of cervical dilation, are essential when controlled vaginal delivery is performed on an anesthetized mare.46 No more than two or three people (depending on size and strength) should apply traction to the fetus.36 Excessive use of force may injure a viable foal and inflict maternal soft tissue trauma. Serious birth-related injuries in a foal may include fractured ribs, bruised costochondral junction of ribs, hemarthrosis of shoulder joints, as well as internal hemorrhage and/or a ruptured viscus.1, 24, 30, 39, 40
Cranial (anterior) presentation Fetal compromise and/or death in stage I or early stage II will preclude the normal positional and/or postural changes. Primary uterine inertia due to subclinical hypocalcemia may affect the normal fetal righting reflexes. It could also be that the foal has not been assisted in its positional changes by the normal rolling that accompanies the early phase of parturition in the mare. If an inexperienced foaling attendant has prematurely summoned veterinary assistance the foal may not be in a normal dorsosacral position. When a live fetus is found to be in an abnormal position (dorsopubic or dorsoilial) it may be possible to rotate it by applying lateral pressure over the shoulder. Another alternative is to let the mare roll and re-examine the position. An active fetal response will often facilitate correction into
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Dystocia Management a dorsosacral position. If the foal is dead, and the forelimbs are extended, it may be possible to mimic the normal rotational movements by combining torque with gentle traction on the obstetrical chains.1 A cranially presented fetus, with forequarters in a dorsosacral position and with head and forelimbs extended, should require minimal traction to complete the delivery. It is important that one limb is advanced slightly ahead of the other. This ensures that the width of the fetus across the shoulders is reduced. Provided that there is adequate lubrication, most foals can be successfully delivered. However, if progress is not being made then all traction should cease. The vaginal canal must be fully explored since there are three possible explanations that can be readily identified.1, 36 1. Incomplete extension of a forelimb (elbow lock): This malposture should be immediately suspected if the fetal muzzle lies at the same level as the fetlocks instead of over the proximal metacarpal bones.5, 36 The flexed shoulder and elbow joint causes increased depth and width of the fetus within the maternal pelvic inlet (see Figure 255.3). If both forelimbs are equally advanced into the vaginal canal there is a high risk of impaction of the fetal elbows on the pelvic brim. Palpation will reveal that the fetal olecranon is caught against the pubis. Correction involves repelling the fetal trunk while dorsomedial traction is applied to the affected forelimb. This will raise the elbow up over the floor of the pelvic inlet and permit full extension.1, 57 2. Absolute feto-pelvic disproportion (fetal oversize): The shape of the mare’s pelvis and the more elongated equine fetus make this condition less
Figure 255.3 Incomplete extension of a forelimb (elbow lock) should be suspected if labor is not progressing and the fetal muzzle lies at the same level as the fetlocks. The normal posture for successful delivery has one forelimb preceding the other such that one shoulder enters the pelvis ahead of the other. The foal’s head should be resting between the carpi.
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common in horses than in cattle.5, 26, 34 The size of the uterus (breed variation) plays a much greater role in determining ultimate fetal size than does the stature of the stallion.60, 61 Despite this, owners will often express concern about fetal oversize if the foal has been carried several weeks past the expected due date. Although this is a major concern with prolonged gestation in cattle, it is seldom an issue in overdue mares.5 In fact, some cases of prolonged gestation in mares may involve a smaller than normal, dysmature fetus.31 However, relative feto-pelvic disproportion may be a significant factor in primiparous mares, and obstetrical assistance (traction) is required much more often in these younger animals.4, 24, 28, 30, 39 Foals from primiparous mares are more likely to sustain thoracic trauma compared to foals from pluriparous mares.39 Approximately 30% of referral hospital dystocia cases are in primiparous mares, and these younger mares were disproportionately represented in a report on dystocia and neonatal asphyxia from the central Kentucky area.15, 24, 26, 27, 30, 39 Apart from fetal issues (malposition, malposture), dystocia in primiparous mares may be further complicated by a tight vaginovestibular sphincter. This will predispose such cases to lacerations and rectovaginal tears.39 If copious lubrication and gentle traction do not facilitate delivery then cesarean section or partial fetotomy are the only alternatives.1 3. Ventro-vertical presentation with hip flexion and impaction of one (‘hurdling’ posture) or both hindlimbs (‘dog-sitting’ posture) (see Figure 255.4): The unilateral posture is more common.26
Figure 255.4 If the forelimbs, head, and thorax are all correctly aligned but no progress is being made, all traction should cease until the pelvic floor has been examined. If either bilateral (‘dog-sitting’ posture) or unilateral (‘hurdling’ posture) hip flexion are present the fetal hindlimbs (hooves) will push against the pelvic brim whenever traction is applied to the forelimbs.
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The dystocia is characterized by partial delivery of the anterior aspect of the fetus, followed by unproductive straining whereby the delivery fails to progress despite the mare’s vigorous expulsive efforts. It has been suggested that this condition could be a variant of a ventro-transverse presentation where the cranial aspect of the fetus has become dislodged and entered the vaginal canal.5 The mechanics of this malposture cause the fetal hindlimbs (hooves) to push against the pelvic brim whenever traction is applied to the forelimbs.4, 5, 36 The cervix is easily damaged by the impacted limb(s) during the mare’s expulsive efforts, and especially due to inappropriate traction from inexperienced personnel. It is essential that all traction is stopped if progress is not being made even though the forelimbs, head, and thorax are all correctly aligned.1 Correction of this malposture is contingent upon the fetus being repelled enough to permit the obstetrician’s fingers to sweep the floor of the pelvic inlet. This is not always possible, especially if the fetus has been forcefully impacted in the birth canal. In extreme cases the hindlimb may actually extend under the fetus and up into the vagina.26 Although the hindlimb may be successfully repelled if the fetus is alive, it is a difficult procedure and is associated with some risk of uterine laceration. Limitations of arm length often preclude grasping the hindlimb. Sometimes merely rotating the fetus will dislodge the trapped hoof. One technique that is not without risk is to use a snare or threaded crutch repeller to loop around the pastern.5 A long cord with a knot in one end is pulled through one eye of a Kuhn’s crutch. The free end is passed through the other eye and the resulting loop placed over the trapped hoof. Tension is maintained on the long cord as the crutch repeller is advanced up the limb and seated against the pastern/fetlock. If used judiciously the instrument may serve to repel the hoof away from the pelvic brim.37, 62 Although it is not an ideal ‘obstetrical’ instrument, some practitioners have used a twitch to facilitate limb repulsion and delivery of a viable foal. When the fetus is dead, repulsion attempts should not be performed on a standing mare since in these cases the repelled fetal hindlimb often will remain flexed instead of returning to its normal position. This then poses an extreme risk that the hoof will puncture the ventral uterine wall as the foal is being extracted.41, 45 It is recommended that general anesthesia and hoisting of the hindquarters be employed to reduce the risk of ventral uterine rupture.63 A surgical alternative to a complete cesarean section involves manipulation of the hindlimb through a ventral midline incision. An assistant may be able to ex-
Figure 255.5 The ‘foot-nape’ posture can impede delivery by increasing the diameter across the fetal chest, and the elbow may become lodged against the pubis. Unless it is prompted corrected, this type of dystocia can lead to a laceration of the vaginal roof as the mare’s straining can force the fetal hoof against the vaginal wall, and in extreme cases it may perforate completely through into the rectum (‘rectovaginal fistula’). If the mare continues to strain with the distal limb trapped in the rectum a tear can develop through the caudal rectovaginal shelf and anal sphincter (‘third-degree perineal laceration’).
tract the fetus through the vagina once the hindlimb has been grasped from within the abdominal cavity. If successful, this technique will reduce the potential for peritoneal contamination that can be associated with cesarean section.62 Since this type of dystocia is especially likely to traumatize the tissues on the pelvic floor, the integrity of the ventral aspect of the cervix should be ascertained prior to rebreeding.1 Foot-nape posture One or both of the forelimbs is displaced over the foal’s head and pushed against the vaginal roof (see Figure 255.5). The abnormal posture increases the diameter across the fetal chest, and the elbows may be lodged against the pubis.5 This malposture can cause significant trauma to the vagina and perineum if it is not corrected immediately. Incorrect placement of the fetal hooves can cause severe trauma to the distal aspect of the birth canal. This is especially likely in maiden mares that do not have sufficient laxity in their vaginal and vestibular tissues to permit passage of a misplaced limb. The mare’s straining can cause the fetal hoof to lacerate the vaginal roof. In extreme cases a rectovaginal fistula is created by the hoof tearing through into the rectum. This may be all that occurs if the foal then withdraws its hoof from the rectum prior to delivery. However, if the mare’s strong expulsive efforts force the trapped limb caudally it will dissect through the rectovaginal shelf – and may even rupture through the anal sphincter. The
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Dystocia Management resulting cloaca-like opening is known as a thirddegree perineal laceration.64–66 The foot-nape malposture is corrected by repelling the fetus into the uterus, lifting the limbs off the neck, and placing them in a lateral position. The head is then raised while at the same time obstetrical chains are used to apply ventromedial traction to the limbs until they come to lie under the head. Once the malposture has been corrected, fetal extraction can usually proceed quite uneventfully.5, 36 If a live foal is found to have a limb lodged in the rectum (fistula) and it cannot be repelled into the vagina, it may be necessary to incise the perineum and surgically create a third-degree perineal laceration. This will facilitate expeditious delivery of a viable neonate. Appropriate postpartum care should permit successful reconstruction of the perineal area within 4 to 6 weeks.67 If the foal is dead it may be possible to amputate the trapped limb with a fetatome, but care must be taken to ensure that the sharp bony stump does not create further trauma.1 Carpal flexion If one or both limbs are not visible at the vulvar lips then either a uni- or a bilateral flexion of the carpus may be present. The affected limb is often lodged in the pelvic inlet.36 If head/neck flexion is also present then this malposture should be corrected before the carpal flexion is resolved (see later discussion). Repulsion of the fetus back into the uterus will permit the flexed limb to be grasped just proximal to the fetlock. Many times it is necessary to grasp the carpus or mid-to-proximal metacarpus in order to achieve sufficient repulsion to create enough room to properly mutate. This is particularly true in cases of carpal contraction. By rotating the wrist, the carpus can be rotated dorsolaterally while the flexed fetlock is brought medially and caudally into the birth canal.5 This maneuver allows maximal use of available space by obliquing the extremity through the pelvic inlet. It is important that the clinician’s hand is cupped over the bottom of the fetal hoof at all times while attempts are made to straighten the limb. Failure to do so may result in injury to the reproductive tract. Application of an obstetrical chain or rope snare to the pastern may be a useful aid since controlled traction can be applied externally while the internal hand manipulates the distal limb, and then covers and guides the hoof.1, 5, 36 Since most carpal flexions are relatively easy to correct manually, the obstetrician should be cognizant of the possibility of contracted tendons when difficulties are encountered. Flexural deformities are considered to be the most common congenital anomaly of foals.15, 24, 29, 30, 41, 44, 45 Limb contractures are generally bilateral, with the forelimbs being
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affected more often than the hindlimbs, although it is not uncommon for all four limbs to be involved.15, 24, 29, 30, 34 Severely affected limbs cannot be straightened, and needless trauma can be inflicted on the genital tract by prolonged attempts to manually correct this malposture.1 If the mare has been in dystocia for some time, the amount of uterine contraction may prevent meaningful repulsion of the fetus. The fetus is invariably dead in such cases, and a relatively simple fetotomy cut at the level of the distal row of carpal bones can permit atraumatic extraction within minutes.1, 4, 44, 45 Head and neck malpostures These malpostures can occur primarily, or on occasion secondarily, when a viable foal pulls back from vaginal manipulations that are intended to correct a minor postural problem. If the muzzle engages the uterine wall, or floor of the pelvic inlet, the mare’s forceful expulsive efforts can drive the head and neck ventrally, or laterally along the thorax, while the forelimbs are pushed further into the vaginal canal (see Figure 255.6). A rope head snare (behind ears and into mouth) that is applied before manipulating the limbs can facilitate head retrieval if a malposture does develop. The clinician should remember that any tension on the head snare can force the mouth open, and thus care must be taken to protect the uterus if retrieval becomes necessary. Head and neck malpostures are very difficult to correct because the long neck often makes it impossible to reach the foal’s head once this type of dystocia has occurred. The long nose and muzzle are an added complication.5, 36 In some especially difficult cases the displaced head will be rotated axially at the atlanto-occipital joint such that the fetal mandibles are uppermost.5 Inexperienced clinicians should consider referral as soon as a head and neck malposture is diagnosed.36 In
Figure 255.6 The most common reason for referring a dystocia case to a veterinary hospital are head and neck malpostures. They can be extremely difficult to correct because the long fetal neck often makes it impossible to reach the head. The length of the head and muzzle are an added complication.
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fact, reflection of the fetal head and neck is the most common reason for referring a dystocia case to a veterinary hospital.4, 15, 26, 34 Prolonged, unrewarding manipulations can easily jeopardize the mare’s future fertility. As with contracted tendons, it is essential that the practitioner considers the possibility of a ‘wry neck’ when attempting to correct a displaced head and neck.15, 24, 30 This congenital curvature of the cervical vertebrae is not amenable to correction by mutation, and needless trauma can be inflicted on the genital tract by unrewarding attempts to achieve the impossible.1 Even when the head/neck is apparently straightened it may still have a kink, and simple traction on the snare only serves to impact the foal within the birth canal. In these cases repulsion of the foal is necessary to provide sufficient space to fully explore the condition of the neck.
Lateral reflexion of the head and neck If the fingers can reach the foal’s muzzle it may be possible to exert traction on the corner of the mouth while moving the wrist to apply opposing pressure against the side of the head. Some obstetricians suggest applying a clamp (with a retrieving cord attached) to an ear so that the head can be retracted enough to permit placement of a mandibular snare. While application of a snare can be a useful method for correcting a lateral head and neck malposture, minimal tension should be applied to the mandible since it is relatively easy to cause a fracture. Gentle traction may be applied to the snare while the internal hand repels the rest of the head and neck. Attempts to grasp the orbital grooves should be made with caution if the foal is alive since pressure on the eyeball may provoke evasive movements that exacerbate the malposture. On occasion a reactive foal may be made to right its head straight by pinching it behind the ears, but this may also be counterproductive if the head is moved further away. Eye hooks can easily cause damage to the foal’s eye, and thus should be used with caution on a live fetus.5, 15 If a portion of the head cannot be grasped, it may be possible to use an obstetrical chain looped around the neck to pull the head within reach. This should be done with caution so that the muzzle is not forced against the uterine wall with such pressure that a rupture results.1 If the fetus is dead a pair of forceps (with retrieving cord attached) can be attached to the skin of the neck in an attempt to pull the head caudally. Another technique that requires extreme caution is the use of a Kuhn’s crutch.5, 37 In a manner similar to that previously described for correction of a ‘dogsitter’ the long cord is fixed to one eye of the crutch. The free end is then attached to a curved introducer that is passed over the neck and retrieved ventrally.
The cord is withdrawn, untied from the introducer, and then passed through the other eye of the Kuhn’s crutch. The free end of the long cord must be held firmly with the crutch handle. Moderate tension is maintained on the cord as the crutch is advanced along the lateral aspect of the neck towards the head.5, 37 If applied correctly the loop around the foal’s neck will prevent the Kuhn’s crutch from slipping off the neck and possibly rupturing the uterus. The advantage of the crutch is that there should be sufficient space for an arm to repel the fetal body while gentle traction on the crutch is used to draw the long head caudally. Once the hand can grasp the muzzle it should protect the uterine wall while guiding the head into a normal posture for delivery. Experienced clinicians may place the ‘U’ of the Kuhn’s crutch in the carina (where the trachea and jugular enter the chest cavity) to repel the fetal chest. However, extreme care must be exercised since if the crutch dislodges it can easily lacerate the uterine wall. Irrespective of the method chosen, correction of a lateral neck flexion can be extremely difficult. Factors influencing the likelihood of success include uterine tonicity, the clinician’s arm length and skill, and the presence or absence of torticollis and facial scoliosis.1, 5, 15, 24, 30, 36
Ventral deviation of the head and neck This is relatively easy to correct if the long fetal nose is just below the brim of the pelvis (vertex or poll posture). The head should be rotated laterally in an arc before attempting to bring the muzzle up over the pelvic brim. In more severe cases the neck is tucked down between the forelimbs (nape posture), and in extreme cases the mandible rests against the sternum (see Figure 255.7). In the less extreme
Figure 255.7 ‘Vertex’ or ‘poll’ posture is a simple ventral deviation of the head (long fetal nose is just below the pelvic brim) and is relatively easy to correct. ‘Nape’ posture is a more difficult dystocia since the neck is tucked down between the forelimbs. In extreme cases the mandible rests against the sternum.
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Dystocia Management deviations a mandibular snare may be useful to apply gentle traction to the head while the poll is repelled. Excessive tension on the snare can easily fracture the mandible. The fetus may have to be repelled deeply into the uterus to provide sufficient space to correct an extreme ventral malposture. This may not be possible unless one or both forelimbs are placed in a carpal flexion, and pushed back into the uterus. A retrieving rope loop or obstetrical chain should be attached to the pastern to facilitate replacement of the limb once the neck flexion has been corrected. It is generally much easier to manipulate the head and neck when the forelimbs are not engaged in the pelvic canal. In a dystocia where there is a head and neck flexion occurring concurrently with bilateral carpal flexion, it is imperative that the neck flexion be corrected before the limbs are extended into the vaginal canal. This type of dystocia typically occurs if the fetus has died prior to, or early in stage I of labor. Cesarean section or fetotomy are indicated if attempts to reposition the head and neck in a timely manner are unsuccessful.1, 5, 41,44–46 Shoulder flexion This malposture may be unilateral (‘freestyle swimming’ posture) (see Figure 255.8) with head and one forelimb presented, or bilateral where only the fetal head protrudes through the vulvar lips.5, 15, 26, 36 An immediate cesarean section may be preferable if the foal is alive since the long forelimbs make manual correction of these malpostures difficult and timeconsuming. The head and neck must be repelled back into the uterus to gain access to the retained limb(s), and this is seldom easy to achieve.5 A soft rope snare should be placed on the fetal head so that
Figure 255.8 The long fetal extremities can make correction of shoulder flexion malpostures difficult and time-consuming. It is usually extremely difficult to repel the head and neck back into the uterus to gain access to the retained limb.
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it can be readily retrieved once the shoulder flexion has been corrected. If the limb(s) can be reached, the mutation is performed in two stages. A carpal flexion is created by locating the retained humerus, and working down to the distal radius. The limb is then pulled caudally and medially as the fetal body is repelled. If a rope loop can be passed around the radius the external hand can apply traction on the distal extremity while the internal hand repels the fetus by applying pressure to the shoulder. Once the flexed carpus has been hooked over the brim of the pelvis it can be extended as described previously. It must be remembered that it is not always possible to repel the head sufficiently to gain access to the retained forelimb. In these cases cesarean section may be the only option for delivery of a live foal. While attempts to extract a fetus with an uncorrected limb malposture pose a risk for both fetus and mare, it is sometimes possible to be successful if the mare has a large pelvis and when the fetus is very small or premature.5 While some clinicians may consider delivering a foal with this malposture, they are unlikely to attempt such a delivery in a maiden mare. The cervix and vagina can easily be lacerated, and extreme pressures created by extraction attempts with an uncorrected limb malposture can cause fetal rib fractures and severe internal trauma. Before attempting an uncorrected delivery the clinician must ensure that a carpal flexion is not present, that fetal oversize is not an issue, and that copious lubrication has been applied.1
Caudal (posterior) presentation If the soles of the hooves are facing upwards then one of two abnormalities should be suspected. Either the foal is in caudal presentation, or a fetus in cranial presentation has failed to rotate as it extended the forelimbs. Palpation of the hocks somewhere in the vaginal canal will confirm the diagnosis. There is only one joint (fetlock) between the hoof and the hock, whereas a forelimb has two joints (fetlock, carpus) between the hoof and the elbow.5 In a caudal presentation there is an increased risk of fetal hypoxia due to compression of the umbilical cord under the fetal thorax (see Figure 255.9). Likewise, premature rupture of the umbilical cord while the fetal head is still retained within the uterine lumen can also cause fetal death.36 In uncomplicated posterior presentations all that may be needed to facilitate delivery is gentle traction on the hindlimbs in conjunction with the mare’s expulsive efforts. However, caudal presentation often predisposes the mare to dystocia because the synchronized rotation of the fetal body – and extension of the extremities – that has been described for cranial presentation
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Figure 255.9 Caudal presentation can easily result in fetal death since there is an increased risk of fetal hypoxia due to compression of the umbilical cord under the fetal thorax. Premature rupture of the umbilical cord while the head is still retained within the uterine lumen can also cause fetal death.
Straightening a flexed hock entails considerable risk since the hock is invariably forced against the dorsal uterine wall. In cases where the uterus is contracted there is a real possibility of creating a laceration or perforation. The metatarsus is grasped and the fetus repelled into the uterus while the hock is pushed dorsolaterally. The procedure for medially obliquing the extremity into the birth canal is similar to that previously described for correction of a carpal flexion.5 An obstetrical chain or strap can be used to apply medial traction to the distal limb. It is important that the hoof is cupped in the hand as it is guided into the pelvic canal. If the foal is dead, and the obstetrician is experienced, it may be safer to correct a flexed hock dystocia by fetotomy.1, 5, 41,44–46 Hip flexion
13, 16
does not occur. In many cases the abnormally presented fetus will be in a dorsoilial position.1, 15, 26 Although only about 1% of foals are presented caudally, this malpresentation may account for 14–16% of referral hospital dystocia cases since any postural abnormalities of the long limbs create a major complication.4, 15, 26, 34 Typically both hindlimbs are involved in the malposture, suggesting that the etiology is associated with a failure of the normal extension mechanism.4, 15, 26 Both bilateral hock and hip flexion cases are extremely difficult to correct under field conditions due to space limitations within the birth canal.1
Approximately half of referred caudal presentation cases are breech (bilateral hip flexion posture). Even under ideal hospital conditions the manipulations involved in attempting to correct a bilateral hip flexion (breech) posture are especially time-consuming since the limbs are displaced under the fetal body (see Figure 255.10). Cesarean section will usually provide the best prognosis for fetal viability and future fertility in these cases.4, 15, 26, 45 The comments for managing a hock flexion apply because, if mutation is attempted, the bilateral hip flexion must first be converted into a bilateral flexed hock posture.5, 36 The distal tibia is grasped and pulled caudo-dorsally until the hock is in a flexed posture. A rope or
Hock flexion This malposture accounts for about one-quarter of referred caudal presentation cases.4, 15, 26 The flexed hocks are palpable either at the pelvic inlet, or impacted more caudally in the vaginal canal. These, and bilateral hip flexion (‘breech’) malpostures, can be extremely difficult to correct because it may not be possible to safely repel the fetus far enough cranially to permit manipulation of the retained limbs. Tocolytic drugs are extremely useful in these cases. Expansion of the uterus with large volumes of lubricant may also facilitate manipulations. Although a Kuhn’s crutch can be used to repel the fetal rump, the instrument can easily perforate the uterus if it slips off the fetus. Anchoring one of the forks in the fetal anus/rectum may reduce the likelihood of this mishap. While an assistant’s arm may also suffice, there are safety issues when an assistant and clinician are working this closely in a standing mare. Certainly a well-lubricated assistant’s arm can provide meaningful fetal repulsion in an anesthetized mare that has the hindquarters elevated. The obstetrician is then able to focus on manipulating the retained limbs.1
Figure 255.10 Cesarean section will often provide the best prognosis for fetal survival and future fertility with this type of dystocia. Bilateral hip flexion (‘breech’ posture) can be extremely difficult to correct because the limbs are displaced under the fetal body. The clinician must first create a bilateral hock flexion before the limbs can be extended. Straightening the limb can force the hock against the dorsal uterine wall such that there is potential for uterine rupture.
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Dystocia Management obstetrical chain looped around the limb can help to retrieve the hock.5 If successful, it is important to remember to flex both hocks before attempting to straighten either limb. If one limb is extended into the vaginal canal while the other hip remains flexed, the fetal body will move back into the pelvic canal, and this will make it extremely difficult to access the retained limb.31 If the fetus is dead – and cesarean section is not an option – this author recommends attempting to convert the bilateral hip flexion into a hock flexion posture, followed by correction with two fetotomy cuts through the distal row of tarsal bones. Provided that the clinician is experienced in the use of a fetatome, this may be safer and cause less trauma than attempting to straighten the limbs.1
Transverse presentation Only about 1 in 1000 foals will be presented transversely. This is the culmination of a bicornual gestation in which the fetus was tightly packed into the uterine body and variable portions of both horns. Neither horn is as fully expanded as would be expected in a normal pregnancy.4, 13 However, the net effect of this abnormal gestation is that overall placental surface area may be increased, and thus the foals are often larger than normal – a factor to be considered if an attempt at vaginal delivery is contemplated. Under less than ideal management conditions these mares may not be recognized as being in labor. The normal vaginal pressure stimuli are not present and only weak – if any – abdominal straining occurs.5, 15 Successful resolution of these dystocia cases requires a significant amount of obstetrical experience. This explains why these extremely rare presentations account for 10–16% of referral hospital dystocia cases.15, 26 If the fetus is alive the delivery method of choice is cesarean section.46
Ventro-transverse The majority of transverse presentations are ventrotransverse (see Figure 255.11) with the abdomen and limbs of the fetus presented towards the birth canal.4, 15, 26 Although the widespread adoption of ultrasonography has markedly reduced the likelihood of a twin birth, this possibility must always be explored when more than two limbs are present in the birth canal.24, 30 In some instances it may be possible to repel the head and forequarters of the fetus, while extending the two hindlimbs into the pelvic canal (version). If the manipulations are successful, the transverse presentation is thus converted into a caudal presentation for vaginal delivery.4 The likelihood of successfully resolving one of these cases is
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Figure 255.11 Transverse presentations are rare, with the majority being ventro-transverse. In these cases the clinician will often palpate multiple limbs within the birth canal, and time should be taken to determine if twins are present. If the fetus is alive, the delivery method of choice for transverse presentations is by cesarean section since successful mutation of these cases requires a significant amount of obstetrical experience. In some instances it may be possible to repel the head and forequarters of the fetus, while extending the hindlimbs into the pelvic canal. Fetotomy is seldom indicated due to the necessity for several cuts.
improved if the mare has been anesthetized and the hindquarters elevated.36 However, extreme caution should be used during any manipulations since transverse fetal presentations are associated with a markedly increased incidence of fetal malformations. This is likely to be a consequence of restricted fetal movement in utero.5 In approximately one-third of cases attempts at correction may be complicated by the presence of flexural limb deformities, angular limb deformity, and vertebral deformity such as wry neck.15 Dorso-transverse In these extremely rare presentations the spinal column of the fetus is presented towards the birth canal. These cases warrant an immediate referral for cesarean section, even if the foal is dead.4, 15, 26 Although an experienced obstetrician may be able to deliver a transversely presented fetus by multiple fetotomy cuts, the owner should be advised that this will be a difficult and time-consuming procedure. There is a high risk of trauma that most likely will impede the mare’s future fertility.1, 36, 41, 44, 45
Fetal anomalies Apart from the ‘contracted foal syndrome’ (limb contractures, scoliosis and/or torticollis), fetal anomalies that have been reported in the horse include hydrocephalus, anencephaly, meningoencephalocele,
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microophthalmia, craniofacial malformations (wry nose), cleft palate, umbilical hernia, diaphragmatic hernia, cardiac anomalies, epitheliogenesis imperfecta, dystrophic chondropathy, scoliosis, and hydronephrosis.24, 29 In most cases these congenital anomalies are not related to hereditary factors and do not cause dystocia.29 A very large umbilical hernia will usually cause evisceration of the fetus during parturition (this is more common with assisted deliveries). Although these cases will present like a schistosomus reflexus birth in cattle (viscera protruding through the vulvar lips), the equine fetus does not appear to develop the dorso-retroflexion of the vertebral column that characterizes the ruminant condition.3, 5, 24, 29, 68, 69 Hydrocephalus is an occasional cause of equine dystocia, especially in pony breeds.4, 15, 29 The condition occurs when increased intracranial pressure causes the bones of the skull to enlarge, sometimes almost doubling the size of the head.15 The skull is often very thin, and many affected foals can be delivered after incising the soft portion of the skull with a finger knife, allowing the skull to collapse.5 The trunk of the hydrocephalic fetus is generally smaller than normal and seldom interferes with delivery.36 If the enlarged cranium is densely ossified then a fetotomy cut may be necessary to reduce the size of the head.1, 5, 41, 44, 45
Conclusion The third stage of parturition involves expulsion of the fetal membranes, and is regarded as being normal if completed within 30 minutes to 3 hours.5, 67 Owners should be advised to seek veterinary assistance if passage of the membranes is delayed. Metritis and laminitis are common sequelae of membrane retention. In an attempt to reduce the incidence of membrane retention this author routinely employs the Burn’s technique after resolving a dystocia case.18, 70 This involves distending the chorioallantois with saline, and then tying off the opening. A low dose of oxytocin (10–20 IU) is then administered and most mares will pass the entire chorioallantoic sac within 30 minutes.67 Although many of these cases may well pass the membranes unaided, it is recognized that obstetrical procedures predispose the mare to membrane retention.71 The advantage of encouraging rapid passage of the entire chorioallantoic sac, with any introduced contaminants contained within, warrants the minimal effort involved in employing this technique. The fetal membranes should be examined as a matter of routine to ensure that they have been passed intact, and to check for any placental anomalies that may forewarn of impending problems in the neonate. Abdominal discomfort in
the postpartum mare may be due to uterine contractions, especially if the mare has been treated with oxytocin to promote passage of the fetal membranes. Other causes of abdominal pain should not be discounted, including rupture of the uterine artery. An abdominocentesis can provide useful prognostic information in the immediate postpartum period.18, 38 Variable amounts of perineal swelling are not uncommon after a dystocia, and the associated discomfort may contribute to fecal impaction. Routine administration of mineral oil can ease fecal passage in mares that have experienced a dystocia.1
References 1. Frazer G. Dystocia and fetotomy. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 417–34. 2. Frazer GS, Emberstson R, Perkins NR. Complications of late gestation in the mare. Equine Vet Educ 1997;9:306–11. 3. Frazer GS, Perkins NR, Embertson RM. Normal parturition and evaluation of the mare in dystocia. Equine Vet Educ 1999;11:41–6. 4. Vandeplassche M. Dystocia. In: McKinnon AO, Voss J (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 578–87. 5. Roberts SJ. Veterinary Obstetrics and Genital Diseases (Theriogenology), 3rd edn. Woodstock, VT, 1986. 6. Vandeplassche M. Prepartum complications and dystocia. In: Robinson N (ed.) Current Therapy in Equine Medicine 2. Philadelphia: Saunders, 1987; pp. 537–42. 7. Vandeplassche M, Lauwers H. The twisted umbilical cord: an expression of kinesis of the equine fetus? Anim Reprod Sci 1986;10:163–75. 8. Ginther OJ. Equine fetal kinetics: allantoic-fluid shifts and uterine-horn closures. Theriogenology 1993;40:241–56. 9. Williams NM. Umbilical cord torsion. Eq Dis Q 2002;10:3–4. 10. Ginther OJ, Griffin PG. Equine fetal kinetics: presentation and location. Theriogenology 1993;40:1–11. 11. Ginther OJ, Williams D, Curren S. Equine fetal kinetics: entry and retention of fetal hind limbs in a uterine horn. Theriogenology 1994;41:795–807. 12. Ginther OJ. Equine physical utero-fetal interactions: a challenge and wonder for the practitioner. J Equine Vet Sci 1994;14:313–18. 13. Ginther OJ. Equine pregnancy: physical interactions between the uterus and conceptus. Am Assoc Equine Pract 1998;44:73–104. 14. Curren S, Ginther OJ. Ultrasonic fetal gender diagnosis during months 5 to 11 in mares. Theriogenology 1993;40:1127–35. 15. Vandeplassche M. The pathogenesis of dystocia and fetal malformation in the horse. J Reprod Fert 1987; Suppl. 35:547–52. 16. Jeffcott LB, Rossdale P. A radiographic study of the fetus in late pregnancy and during foaling. J Reprod Fert 1979; Suppl. 27:563–9. 17. Frazer GS. Uterine torsion. In: Robinson N (ed.) Current Therapy in Equine Medicine 5. Philadelphia: Saunders, 2002; pp. 311–15. 18. Frazer GS. Postpartum complications in the mare. Part 1: Conditions affecting the uterus. Equine Vet Educ 2003;5:45–54.
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Dystocia Management 19. Dascanio JJ, Ball BA, Hendrickson DA. Uterine tear without a corresponding placental lesion in a mare. J Am Vet Med Assoc 1993;202:419–20. 20. Sutter WW, Hopper S, Embertson RM, Frazer GS. Diagnosis and surgical treatment of uterine lacerations in mares (33 cases). Am Assoc Equine Pract 2003;357–59. 21. Ley WB, Bowen JM, Purswell BJ. The sensitivity, specificity and predictive value of measuring calcium carbonate in mare’s prepartum mammary secretion. Theriogenology 1993;40:189–98. 22. Ousey JC, Delclaux M, Rossdale PD. Evaluation of three strip tests for measuring electrolytes in mares’ prepartum mammary secretions and for predicting parturition. Equine Vet J 1989;21:196–200. 23. Paccamonti DL. Milk electrolytes and induction of parturition. Pferdeheilkunde 2001;17:616–18. 24. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith BJ, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203:1170–5. 25. Whitwell KE. Investigations into fetal and neonatal losses in the horse. Vet Clin N Am 1980;2:313–31. 26. Frazer GS, Perkins NR, Blanchard TL, Orsini J, Threlfall WR. Prevalence of fetal maldispositions in equine referral hospital dystocias. Equine Vet J 1997;29:111–16. 27. Ginther OJ, Williams D. On-the-farm incidence and nature of equine dystocias. J Equine Vet Sci 1996;16:159–64. 28. Vandeplassche M. Selected topics in equine obstetrics. Am Assoc Equine Pract 1992;623–8. 29. Crowe MW, Swerczek TW. Equine congenital defects. Am J Vet Res 1985;46:353–8. 30. Hong CB, Donahoe JM, Giles RC, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Equine abortion and stillbirth in central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. 31. Hillman RB. Dystocia management at the farm. In: Periparturient Mare and Neonate Symposium. San Antonio, TX: Society for Theriogenology, 2000; pp. 65–71. 32. Macpherson ML, Chaffin MK, Carroll GL, Jorgensen J, Arrott C, Varner DD, Blanchard TL. Three methods of oxytocin-induced parturition and their effects of foals. J Am Vet Med Assoc 1997;210:799–803. 33. Brendemeuhl JP. Fescue and Agalactia – Pathophysiology, Diagnosis and Management. In: Periparturient Mare and Neonate Symposium. San Antonio, TX: Society for Theriogenology, 2000; pp. 25–32. 34. Byron C, Embertson RM, Bernard WV, Hance S, Bramlage L, Hopper S. Dystocia in a referral hospital setting: approach and results. Equine Vet J 2002;35:82–5. 35. Norton JL, Dallap BL, Johnston JK. Retrospective study of dystocia in mares at a referral hospital. Equine Vet J 2007;39:37–41. 36. Frazer GS, Perkins NR, Embertson RM. Correction of equine dystocia. Equine Vet Educ 1999;11:48–53. 37. Arthur G, Noakes D, Pearson H, Parkinson TJ. Veterinary Reproduction and Obstetrics, 7th edn. London: Saunders, 1996. 38. Frazer GS, Burba D, Paccamonti D, LeBlanc MM, Embertson R, Hance S, Blouin D. The effects of parturition and peripartum complications on the peritoneal fluid composition of mares. Theriogenology 1997;48:919–31. 39. Jean D, Laverty S, Halley J, Hanningan D, Leveille R. Thoracic trauma in newborn foals. Equine Vet J 1999;31:149–52.
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40. Schambourg M, Laverty S, Mullim S, Fogarty UM, Halley J. Thoracic trauma in foals: post mortem findings. Equine Vet J 2003;35:78–81. 41. Frazer GS. Fetotomy technique in the mare. Equine Vet Educ 2001;13:195–203. 42. Freeman DE, Hungerford LL, Schaeffer D, Lock TF, Sertich PL, Baker GJ, Vaala WE, Johnston JK. Caesarean section and other methods for assisted delivery: comparison of effects on mare mortality and complications. [see comments]. Equine Vet J 1999;31:203–7. 43. Embertson R. Dystocia and caesarean sections: the importance of duration and good judgement. Equine Vet J 1999;31:179–80. 44. Frazer GS. Review of the use of fetotomy to resolve dystocia in the mare. Am Assoc Equine Pract 1997;262–8. 45. Bierschwal CJ, deBois C. The Technique of Fetotomy in Large Animals. Bonner Springs, KS: VM Publishing, 1972. 46. Embertson RM. The indications and surgical techniques for cesarean section in the mare. Equine Vet Educ 1992;4:31–6. 47. LeBlanc MM. Sedation and anesthesia of the parturient mare. In: Periparturient Mare and Neonate Symposium. San Antonio, TX: Society for Theriogenology, 2000; pp. 59–64. 48. Luukkanen L, Katila T, Koskinen E. Some effects of multiple administration of detomidine during the last trimester of equine pregnancy. Equine Vet J 1997;29:400–3. 49. Taylor PM. Anaesthesia for pregnant animals. Equine Vet J 1997; Suppl. 24:1–6. 50. Embertson RM, Bernard WV, Hance SR, Smith S. Hospital approach to dystocia in the mare. Am Assoc Equine Pract 1995;13–14. 51. Grubb TL, Reibold TW, Huber MJ. Comparison of lidocaine, xylazine, and xylazine/lidocaine for caudal epidural analgesia in horses. J Am Vet Med Assoc 1992;201: 1187–90. 52. Card CE, Wood MR. Effects of acute administration of clenbuterol on uterine tone and equine fetal and maternal heart rates. Biol Reprod 1995; Mono 1:7–11. 53. Zent WW. Postpartum complications. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia: Saunders, 1987; pp. 544–7. 54. Pascoe J, Pascoe R. Displacements, malpositions and miscellaneous injuries of the mare’s urogenital tract. Vet Clin N Am 1988;4:439. 55. Fisher AT, Phillips TN. Surgical repair of a ruptured uterus in five mares. Equine Vet J 1986;18:153–5. 56. Rossdale PD. Differential diagnosis of postparturient hemorrhage in the mare. Equine Vet Educ 1994;6:135–6. 57. Perkins NR, Frazer GS. Reproductive emergencies in the mare. Vet Clin N Am – Equine Pract 1994;10:643–70. 58. Hooper RN, Blanchard TL, Taylor TS. Identifying and treating uterine prolapse and invagination of the uterine horn. Vet Med 1991;88:60. 59. Brooks DE, McCoy DJ, Martin GS. Uterine rupture as a postpartum complication in two mares. J Am Vet Med Assoc 1985;187:1377–9. 60. Tischner M, Klimczak M. The development of Polish ponies born after embryo transfer to large recipients. Equine Vet J 1989; Suppl. 8:62–3. 61. Wilsher S, Allen WR. The influence of maternal size, age and parity on placental and fetal development in the horse. In: Katila T, Wade J (eds) Havemeyer Monograph Vol 3. New York: D.R. Havemeyer Foundation, 2000. 62. Hunt RJ. Obstetrics. In: Robinson NE (ed.) Current Therapy in Equine Medicine 5. Philadelphia: Saunders, 2003; pp. 319–22.
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63. Baldwin JL, Cooper WL, Vanderwall DK. Dystocia due to anterior presentation with unilateral or bilateral hip flexion posture (‘dog-sitting’ presentation) in the mare: incidence, management, and outcomes. Am Assoc Equine Pract 1991;229–240. 64. Aanes WW. Surgical management of foaling injuries. Vet Clin N Am 1988;4:417. 65. Trotter G. The vulva, vestibule, vagina, and cervix. In: Auer JA, Stick JA (eds) Equine Surgery, 2nd edn. Philadelphia: Saunders, 1999. 66. Freeman D. Rectum and anus. In: Auer JA, Stick JA (eds) Equine Surgery, 2nd edn. Philadelphia: Saunders, 1999; pp. 286–93.
67. Frazer GS. Postpartum complications in the mare. Part 2: Fetal membrane retention and conditions of the gastrointestinal tract, bladder and vagina. Equine Vet Educ 2003;5:118–28. 68. Bezeck DM, Frazer GS. Schistosomus reflexus in large animals. Compend Contin Educ Pract Vet 1994;16:1393–9. 69. Whitwell KE. Morphology and pathology of the equine umbilical cord. J Reprod Fert 1975; Suppl. 23:599–603. 70. Burns SJ, Judge NG, Martin FE, Adams LG. Management of retained placenta in mares. Am Assoc Equine Pract 1977; 381. 71. Vandeplassche M. Aetiology, pathogenesis and treatment of retained placenta in the mare. Equine Vet Educ 1971;3:144.
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Chapter 256
Fetotomy Grant Frazer
Most equine dystocias are readily corrected by assisted vaginal delivery. More complicated cases can often be resolved following general anesthesia and hoisting of the hindquarters – termed controlled vaginal delivery. General anesthesia provides excellent muscle relaxation; and hoisting of the hindquarters provides additional free space within the uterus such that the fetus may be repelled and obstetrical manipulations performed. It should be remembered, however, that prolonged intervention can generally be expected to result in severe trauma to the genital tract, even if performed by a skillful and experienced operator.1–5 If the fetus is dead, many malposture cases can be rapidly resolved by fetotomy in the standing mare – provided that the clinician has the appropriate skills and equipment.4–7 Caudal epidural anesthesia and/or sedation may be used at the clinician’s discretion.8, 9 This may eliminate the need for general anesthesia. When applied correctly, a fetotomy procedure permits the clinician to avoid the additional cost and risk of cesarean section to remove a dead foal.5,9–13 Partial fetotomy (one, two or, exceptionally, three cuts) permits repositioning of the fetus such that controlled vaginal delivery is possible.14 A series of 132 severe dystocias that were resolved by partial fetotomy included reflection of the head and neck (54.5%), breech presentation with deformity or ankylosis of the hindlimbs (12.9%), partial transverse presentation (18.9%), deformity, ankylosis or reflection of the forelimbs (9.0%), and hydrocephalus/dicephalicus (4.5%). On the basis of these 132 cases Vandeplassche stated that partial fetotomy was the method of choice to rapidly and safely resolve over 80% of cases that were not amenable to mutation alone.15 The only alternative in such cases is a cesarean section. In another dystocia study7 almost half of the cases were resolved by fetotomy, and 73% (51/70) of these were performed on a standing tranquilized mare.7 A one- to three-cut fetotomy was
performed in cases where the head and neck, and/or limbs were abnormally placed. One or two cuts were sufficient to correct 57% (40/70) of the cases, and another 15 required three cuts. More than three cuts (15/70) were performed when economics of the case precluded the expense of cesarean section or where, in the opinion of the clinician, the autolyzed state of the fetus presented too great a surgical risk. Of the cases not resolved by mutation alone, 66% were corrected by fetotomy and 34% by cesarean section.7 While one or two fetotomy cuts can permit rapid, atraumatic extraction of a dead fetus, the procedure is often used as a last resort when prolonged manipulations have failed to correct the problem, and extensive soft tissue trauma has already been inflicted.5 Repeated in-and-out arm movements are contraindicated as the mucous membranes of the mare’s vagina and cervix are easily abraded. If the surface epithelium of the vagina is destroyed then exposure of the underlying tissue can result in adhesions that may adversely affect fertility. Cervical and vaginal scarring and adhesions are almost inevitable after prolonged intervention. An irritated equine vagina takes on the texture of sandpaper, and the scar tissue laid down during the healing process results in what can best be described as a ‘stovepipe vagina’.4 While unnecessary delay, poor technique, and inappropriate fetotomy cuts often lead to infertility, recent studies have confirmed that if a one- or twocut fetotomy is performed by an experienced obstetrician then the mare’s reproductive future is not necessarily impeded.8, 9 The economic value of the mare, expertise, equipment and facilities available to the veterinarian, and owner’s preference ultimately determine which method is chosen for those cases where attempts at manual correction of malpositions and malpostures have failed. Economic considerations include the future use of the mare (breeding vs performance value), veterinary fees (anesthesia and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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surgery vs fetotomy) and the extended hospitalization required for postsurgical management.5
using atraumatic technique can minimize complications and maintain future fertility.
Fertility concerns
Equipment and general procedures
Fetotomy performed by a skilled veterinarian should be considered as a viable option for quick and safe correction of dystocia. One or two well-placed fetotomy cuts can dramatically shorten the intervention time and permit atraumatic delivery of a non-viable fetus. However, if the obstetrician is not familiar with correct fetotomy technique then the best option for the mare’s reproductive future may well be a cesarean section.5 Complete perforation of the uterine wall is not necessary for peritonitis to develop if traumatic obstetrical manipulations have caused an acute hemorrhagic necrosis of the endometrium.16 Peritonitis can develop subsequent to severe necrotizing endometritis if the uterine wall is traumatized, and areas of transmural necrosis extending through the myometrium to the uterine serosa can be seen at necropsy in such cases.17 The normal parturient process in the mare does not entail epithelial loss from the endometrium and it appears that an intact endometrium is able to prevent absorption of endotoxin and bacteria, whereas devitalization permits diapedesis and peritoneal contamination.17 The effectiveness of the barrier provided by a healthy uterine wall has been clearly demonstrated by a study that examined changes in peritoneal fluid content after protracted dystocias.18 That work demonstrated that a correctly performed fetotomy does not cause changes in the composition of the peritoneal fluid in post-dystocia mares. Italian researchers examined the records of 72 dystocia cases in which fetotomy was performed in order to determine survival rate, complications, and short-term fertility rates.8 Survival rate after fetotomy was 95.8%, with vaginal and/or cervical lacerations occurring in 2.8% of cases. This survival rate was considered to be high when compared with rates reported after cesarean section.19–21 Mares (n = 68) were bred 2 to 3 months after fetotomy and had good short-term fertility, with a mean pregnancy rate of 79.4% at 45 days after breeding.8 A smaller study of 20 dystocia cases that were resolved by fetotomy also reported that partial fetotomy does not necessarily impede the future fertility of the mare.9 In this study a breeding season was defined as a year in which each mare was bred regardless of pregnancy. There were 24 breeding seasons for which 20 mares were confirmed to be pregnant (83%), and live foaling rate had been 100% for those not currently pregnant at the time the paper was published. Expedient decision-making by experienced clinicians
The author prefers to perform partial fetotomy procedures on a tranquilized, standing mare in a large stall bedded with non-slip material, preferably clean straw. Parturient mares are unpredictable and the clinician should take whatever precautions are necessary to permit a safe examination – both for the mare and the veterinarian. The presence of a dead fetus means that only the mare need be considered when administering chemical restraint. The author routinely uses low doses of xylazine (0.3 to 0.5 mg/kg i.v.) and butorphanol (0.01 to 0.02 mg/kg i.v.) in conjunction with judicious application of a twitch or lip chain. The twitch should be tightened as needed to distract the mare from the obstetrician’s activities. It should be loosened when not needed so that it does not lose its effectiveness. Epidural anesthesia may not eliminate the abdominal contractions of mares, but it does provide analgesia to the perineal region and it can be useful to reduce the reflex straining initiated by manipulations within the birth canal. In those mares subjected to a prolonged fetotomy procedure the use of general anesthesia and hoisting of the hindquarters is recommended. It should be remembered, however, that complete fetotomy can generally be expected to result in severe trauma to the genital tract, even if performed by a skillful and experienced operator.4 It is essential that the correct equipment is available5, 6 (Figure 256.1):
r
Fetatome and threader: double-barreled instrument with a hand grip and notched oval plate for anchoring obstetrical chains;
Figure 256.1 Essential equipment – a fetatome and threader (with cleaning brush), saw wire with handles, wire cutter, curved wire introducer, fetotomy (palm) knife, Krey hook, and obstetrical chains with handles.
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Fetotomy
r r r r r r r
Fetotome saw wire: available in spools (approximately 4.6 m [15 feet] are required to double thread the fetatome, pass around the fetus, and still provide the assistant with sufficient length to saw); Sterile wire cutters should be kept in the obstetrical kit; Wire saw handles that permit the wire to be quickly attached and securely held; Wire introducer: curved instrument for passing the wire over or around a fetal part; Krey hook: this expandable, two-armed hook comes with a stop that prevents over-closure of the instrument should it become disengaged from the fetus; Fetotomy (palm) knife: used to seat the wire for some cuts and to remove bone fragments prior to fetal extraction; A complete obstetrical kit (chains, etc.) should include bucket liners, lubricant, and a sterile stomach tube with pump.
Careful thought should precede all manipulations. Repeated vaginal entry and internal maneuvers only serve to traumatize the birth canal, and increase the level of bacterial contamination. Application of copious volumes of lubricant is essential because the mare’s genital tract is very sensitive to trauma, and the uterus is easily ruptured. A clean stomach tube and pump are used to gently instill an obstetrical lubricant mixture into the uterine lumen as often as necessary during the procedure to keep the tract coated with lubricant. If the uterus is contracted, the lubricant tends to induce some uterine relaxation and thereby create additional space between the uterus and fetus. In countries where they are available, systemic spasmolytic (tocolytic) agents (isoxsuprine; clenbuterol) relax the uterine smooth muscle and therefore provide more space for manipulations.5 Once the wire has been threaded around the fetal part to be amputated, sufficient tension is applied to the wire to ensure that it is not crossed or kinked. The veterinarian must ensure that the head of the fetatome is in the correct position and then cover it with a hand, while using the other arm to hold the fetatome securely during the cutting procedure. An assistant is instructed to start the cut by slow, short, to-and-fro arm movements. Once the wire is seated, the individual performing the fetotomy cut should be instructed to use long rather than short strokes. The length of the arm movements is increased, as is the amount of pressure. Longer movements decrease the amount of heat generated and spread the wear on the wire, making breakages less likely. However, excessive tension on the wire may cause it to snap. The obstetrician should ensure that the head of the fetatome remains positioned correctly at all times.4
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Although a correctly performed cut can be completed very quickly, the obstetrician should not then rapidly extract the fetus. Since the normal dilation processes of stages I and II have been compromised, it is important to apply copious lubrication, and then to proceed slowly so that the cervical and vaginal tissues can stretch without tearing.5
Specific cuts Anterior presentation
Carpal flexion If the mare has been in dystocia for some time, the amount of uterine contraction may prevent meaningful repulsion of the fetus. The fetus is invariably dead in such cases, and a relatively simple fetotomy cut at the level of the distal row of carpal bones can permit atraumatic extraction within minutes.2, 4, 6 Although most carpal flexions are relatively easy to correct manually, the clinician should always consider the possibility of contracted tendons when difficulties arise.2, 7, 14, 22 If the fetus is dead, a fetotomy cut through the distal row of carpal bones is preferable to prolonged attempts at manual correction which, in the case of contracted tendons, may be impossible.2, 6, 7 In a survey of 668 foals that died due to complications of birth the contracted foal syndrome was the most common congenital anomaly diagnosed. 23, 24 This rigid deformity generally means that the fetus must be extracted by cesarean section. Less severe cases may be managed by fetotomy. Limb contractures are generally bilateral and more common in the forelimbs (87%) than the hindlimbs (74.5%).24 Contracted fetlocks are more common than contractures of either the carpus or tarsus. To perform this cut, one channel of the fetatome is threaded and the free end attached to a curved wire introducer. This is passed around the carpus of the flexed limb and the wire returned to the outside. The second channel of the fetatome is then threaded and the head of the instrument held firmly against the distal carpal joint (Figure 256.2). Cutting through the intercarpal joint ensures that the increased diameter of the distal radius in the area of the growth plate remains. This then acts as an anchor point for the obstetrical chain, thereby preventing it from slipping off as traction is applied. A common error is to make the cut above or below the carpal joint.5 Apart from loss of an anchor point (too high), an incorrectly placed cut (above or below the joint) will result in a sharp stump of bone that can easily lacerate the uterus and/or vagina.4 The amputated distal limb should always be removed from the uterus before the fetus is extracted.6 Care should be taken to
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Figure 256.2 When removing a flexed carpus, the head of the fetotome should be positioned such that the wire will make a cut at the level of the distal row of carpal bones. See text for details.
Figure 256.3 A relatively simple fetotomy cut can permit atraumatic extraction of a fetus with head and neck malposture and this may be the preferred option in experienced hands if the fetus is dead. See text for details.
extend the proximal limb fully before applying traction. This will ensure that an elbow lock does not prevent fetal extraction.
Neck amputation One channel of the fetatome is threaded and the wire introducer attached to the free end is passed between the neck and chest wall. If the fetus is tightly impacted in the birth canal it may be difficult to pass the wire introducer between the flexed head and neck. The head of the fetatome is held as close to the thoracic cavity as possible so that amputation is near the base of the neck (Figure 256.3). A Krey hook may be attached to the exposed vertebra to facilitate extraction of the severed head and neck. The hook is then anchored to the vertebral stump of the fetus and, together with traction on leg chains, extraction is readily achieved.4, 6 It is essential that a hand is placed over the neck stump while traction is applied to the Krey hook chain and forelimbs. This ensures that the uterine wall, cervix, and vagina are not traumatized by bone fragments. If a wry neck is suspected, it may be preferable to make the first cut through the greater curvature, and then a second at the thoracic inlet after the head and proximal neck stump have been removed.
Deviation of the head and neck Deviation of the head and neck, either laterally alongside the thorax, or ventrally between the forelimbs, can be very difficult to correct. In fact, it is the major reason for referral of equine dystocias to specialist hospitals.7, 14 It is important to be able to differentiate a wry neck from a mere lateral deviation. A wry neck cannot be straightened, and prolonged unproductive manipulations serve only to traumatize the genital tract.22, 25 If a fetus with head and neck malposture is dead when the veterinarian arrives then a relatively simple fetotomy cut can resolve these cases quite atraumatically.14 The author believes that this approach is preferable to prolonged – and often unsuccessful – attempts at manual correction of this extremely difficult malposture.5 Two options are possible: either direct amputation of the flexed head and neck, or amputation of the opposite forelimb. Removing the entire forelimb may permit the obstetrician to advance a forearm deeper into the uterus such that the head can be reached. Alternatively, the extra space in the pelvic canal may now permit the fetus to be extracted several inches further such that the retained head can be reached. If the malposture cannot be corrected, then the extra space facilitates passage of the curved wire introducer and a second cut can then readily remove the retained head and neck.3, 4, 6, 14, 25 In Vandeplassche’s dystocia population, 89% (154/173) of the anteriorly presented cases (head and neck reflection) that were not amenable to manual correction were subsequently resolved by predominately one-cut (40%) or two-cut (30%) fetotomies.1, 2, 14
Removal of an extended forelimb When the flexed neck cannot be reached, or it is too tightly flexed to pass the wire introducer, it may be possible to amputate the opposite forelimb. The head of the pre-threaded fetatome is guided up the lateral aspect of the forelimb and the wire loop is seated in the axilla. The fetatome is then advanced further into the uterus such that the head is positioned dorsocaudal to the fetal scapula. It is essential that the wire is not dragged over the vaginal and cervical mucous membranes. Once tension is applied to the wires, the loop should be seated in the axilla such that the ventral wire passes between the fetal elbow joint and chest, and the dorsal wire rests medial to the humeroscapular joint.4
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Figure 256.4 When amputating an extended forelimb, an obstetrical chain attaches the fetlock to the fetatome, and holds the forelimb in extension. See text for details.
An obstetrical chain attaches the fetlock to the fetatome, and holds the forelimb in extension. The veterinarian must ensure that the head of the fetatome remains deep in the uterus (dorsocaudal to the scapula) during the sawing process. A correctly placed cut will remove the entire forelimb by dissecting up through the muscular attachments between the scapula and chest wall (Figure 256.4). The obstetrician should endeavor to remove the scapula in its entirety, but the length of the fetal limb means that it is common to have a dorsal remnant of the scapula attached to the fetal trunk. It is critical that this portion be removed by hand (using a palm knife) before fetal extraction is attempted, since sharp edges may lacerate the already friable tissues of the cervix and vagina.4, 6 A dangerous complication occurs if the fetatome is permitted to move caudally during the cutting procedure. The misplaced cut will most likely occur through the proximal humerus, creating an extremely sharp bone stump, and leaving the width of the shoulder joint intact. If the procedure is performed correctly the extra space in the pelvic canal permits the fetus to be pulled further into the birth canal. Often this then permits the retained head to be reached, and manipulated into correct alignment for delivery. Failing this, a second cut should be made through the now accessible flexed neck.
Retained forelimb (shoulder flexion) Uni- or bilateral shoulder flexion inevitably causes dystocia.2, 4 Manual correction alone is possible but may take some time to achieve due to the long limbs of the equine fetus.2, 22, 26 If the fetus is dead, a fetotomy cut to remove the head with as much neck as can be reached may provide sufficient space to correct the malposture.3–6 This approach is definitely indicated if the mare is presented with only
Figure 256.5 Removal of the head with as much neck as possible often provides sufficient space for an arm to correct the malposture if a fetus with uni- or bilateral shoulder flexion is determined to be dead. If this is unsuccessful, an experienced obstetrician may be able to perform a fetotomy to permit atraumatic extraction. See text for details.
the dead foal’s head protruding through the vulvar lips. If the head and neck are removed and the shoulder flexion still cannot be manually corrected, an experienced obstetrician may be able to remove the retained limb in utero by performing a second fetotomy cut. A palm knife is used to make a deep incision medial to the dorsal border of the scapula prior to introduction of the saw wire. It is essential that a long cord is attached to the palm knife so that it can be easily retrieved if the finger ring slips off. The curved wire introducer is passed down between the retained limb and chest wall, retrieved ventrally, then withdrawn under the humeroscapular joint. Once the second channel of the fetatome is threaded it is positioned medial to the shoulder joint, and the wire loop is seated in the dorsal incision. In this instance the direction of the fetotomy cut will be down through the muscular attachments that hold the scapula to the thorax (Figure 256.5).3, 5, 6 Correct positioning of the head of the fetatome and the wire loop is essential. If the wire slips out of the dorsal incision an erroneous cut most likely will be made through the proximal portion of the retained humerus, creating an extremely sharp bone stump, and leaving the width of the shoulder joint intact.
Feto-pelvic disproportion This condition is uncommon in the mare. In experienced hands a transverse section of the lumbar vertebrae, followed by bisection of the pelvis, will usually permit extraction of a large fetus.27
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Ventro-vertical presentation with uni(hurdling) or bilateral hip (dog sitting) flexion In experienced hands partial fetotomy may be a viable alternative to cesarean section if the foal is dead.3–6,28 However, the owner must accept that correction of this type of dystocia by fetotomy is not without risk to the mare.29 This malposture is often not detected until the head and chest have passed through the birth canal, and thus fetal repulsion may not be possible. An initial cut transects the fetal trunk. If the hindquarters are repelled so that the hindlimbs can be retrieved to create a posterior presentation, there is potential for a uterine laceration from the exposed vertebral stump. Extreme caution is indicated.3, 28 Another approach entails removal of the anterior aspect, evisceration of the retained portion, then undermining the dorsal skin sufficiently to permit removal of the lumbar vertebrae as close to the pelvis as possible. The skin flaps are then tied together to cover the vertebral stump. The pelvis is repelled into the uterus and chains are attached to the hindlimbs such that the caudal portion of the fetus is delivered in posterior presentation.6, 14, 28 A safer approach may be to bisect the pelvis, provided that the wire can be passed over the tailhead and threaded down between the hindlimbs.3, 6, 29 An alternate approach in the anesthetized mare has been described in which a forelimb is removed, the costochondral junctions incised, and the fetus eviscerated. The curved fetotomy wire introducer is then passed far back over the fetal tailhead and retrieved by passing an arm underneath into the inguinal region of the fetus. The malpostured hindlimb is sectioned in the coxofemoral area and the fetus delivered by applying traction on the remaining forelimb.28 This method eliminates the possibility of uterine rupture if the offending hindlimb remains flexed within the uterus after manual repulsion. If a cesarean section is to be performed it is common practice to cut through the foal’s vertebral column, and remove the fetal portion that is protruding through the vulvar lips. Thus, only the retained hindquarters and trunk are subsequently removed through the surgical site.
Hydrocephalus This condition is not uncommon in equine fetuses, especially those of the pony breeds.2 In the author’s referral hospital caseload, approximately 5% of dystocias involve a hydrocephalic fetus.7 In many cases, the extremely thin skull bones can be crushed manually to facilitate delivery. If the enlarged cranium is densely ossified then a fetotomy cut may be neces-
sary to reduce the size of the head.3–6 A single fetotomy cut can be performed by looping the saw wire behind the ears and holding the head of the fetatome between the eyes, thereby permitting removal of the dorsal half of the fetal head, and then assisted vaginal delivery of the fetus.3, 4, 6 Alternately, if the head is turned back, the wire loop may be placed in the mouth and the head of the fetatome held at the base of the skull. In both scenarios it is important to place a hand over the cut surface to protect the vaginal canal during extraction. The trunk of a hydrocephalic fetus is generally smaller than normal and therefore seldom interferes with delivery.25 Posterior presentation In Vandeplassche’s dystocia population, 78.6% (44/56) of posterior presentations not amenable to manual correction were subsequently resolved by fetotomy (mean 2.8 cuts).1, 2, 14
Hip flexion In referral hospital populations, approximately 50% of posteriorly presented cases have bilateral hip flexion (breech presentation).7, 14 These cases are not readily amenable to fetotomy, since introduction of the wire is not easily performed. Caesarean section is the best option if the fetus is alive, and often even when the fetus is dead.3, 4, 6, 14, 22, 26 It may not be possible to grasp the wire from below even if it can be passed over the fetal rump and down between the retained limb and body wall. If the wire can be retrieved, it is threaded down the other fetatome channel and the head of the instrument is placed firmly against the perineum on the side of the tail opposite the limb being amputated. The goal is to make a diagonal cut through the pelvis. This removes the entire hindlimb and the caudal aspect of the pelvis that includes the acetabulum.3 Once the amputated limb has been removed from the pelvic cavity there may be sufficient space to correct the remaining malposture and extract the fetus. If the fetus is dead – and cesarean section is not an option – this author recommends attempting to convert the bilateral hip flexion into a hock flexion posture, followed by correction with two fetotomy cuts through the distal row of tarsal bones. Anesthetizing the mare, hoisting the hindquarters, then instilling copious amounts of lubricant will eliminate straining and increase the abdominal space.14, 22 Even if the hip flexion is corrected, the uterine body can still easily be ruptured at its dorsal aspect as the hock is then straightened. Provided that the clinician is experienced in the use of a fetatome, cuts through the distal row of tarsal bones may be safer and cause less trauma than attempting to straighten the limbs.
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Conclusions
Figure 256.6 There is considerable risk of uterine rupture when attempting to straighten a ‘flexed hock’ malposture since the hock is invariably forced against the dorsal uterine wall. Thus, it may be safer to correct this dystocia by fetotomy if the fetus is dead. See text for details.
Hock flexion Approximately 25% of posteriorly presented cases referred to equine hospitals involve bilateral hock flexion.7, 14 If the foal is dead and the obstetrician is experienced, it may be safer to correct a flexed hock dystocia by fetotomy than by attempting to manually correct the malposture.2–6,10, 14 The fetotomy technique is essentially the same as that described for amputation of a flexed carpus, with the cut being made through the lower row of tarsal bones. One channel of the fetatome is pre-threaded, a curved wire introducer is passed around the hock joint, and the second channel threaded. The head of the fetatome is then positioned such that the limb is sectioned through the distal row of tarsal bones (Figure 256.6). The comments about adverse effects of high or low cuts at the carpus also apply to hock joint fetotomy cuts. Once the amputated portion has been removed, an obstetrical rope or chain can then be fixed above the os calcis and use it as a point of traction.3, 4, 6 It may be necessary to repel the fetus to extend the stifle joints up into the pelvic canal. Transverse presentation The author has extracted several transversely presented fetuses by means of fetotomy, but these are extremely difficult cases, and the procedure in this instance should be viewed as a last resort to save the mare’s life when cesarean section is not an option. In Vandeplassche’s dystocia population, transverse presentation of the fetus was the predominant reason for cesarean section.1, 2, 14 Those cases that were resolved by fetotomy required an excessive number of cuts (mean 3.3).
The aim of a fetotomy is rapidly to decrease the size of a fetus such that safe extraction can proceed. This avoids the stress and injury that follows prolonged manipulations and excessive traction, and also the additional expense and risks inherent in performing a caesarean section.4, 6, 14 The results from fetotomy can vary tremendously, depending to a large extent upon the level of expertise offered by the obstetrician and facilities available.2, 16 Certainly, injudicious use of a fetatome can jeopardize the fertility and, possibly, the life of a mare.16, 20 A common fault is to choose fetotomy only after the birth canal has already been traumatized by unproductive attempts at manual correction.4, 6 A recent study demonstrated that when a fetotomy is performed correctly there should be no damage to the uterus, as indicated by changes in the protein composition and cellularity of the peritoneal fluid.30 Likewise, subsequent fertility is not necessarily compromised if the procedure is performed by an experienced clinician.4, 8, 9
References 1. Vandeplassche M. Selected topics in equine obstetrics. Proceedings of the American Association of Equine Practitioners, 1992. 2. Vandeplassche M. Dystocia. In: McKinnon AO, Voss J (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 578–87. 3. Roberts SJ. Veterinary Obstetrics and Genital Diseases (Theriogenology), 3rd edn. Woodstock, VT: 1986. 4. Frazer GS. Review of the use of fetotomy to resolve dystocia in the mare. Proceedings of the American Association of Equine Practitioners, 1997. Phoenix, AZ. 5. Frazer GS. Fetotomy technique in the mare. Equine Vet Educ 2001;13:195–203. 6. Bierschwal CJ, deBois C. The Technique of Fetotomy in Large Animals. Bonner Springs, KS: VM Publishing, 1972; pp. 6–50. 7. Frazer GS, Perkins NR, Blanchard TL, Orsini J, Threlfall WR. Prevalence of fetal maldispositions in equine referral hospital dystocias. Equine Vet J 1997;29(2):111–16. 8. Carluccio A. Survival rate and short-term fertility rate associated with the use of fetotomy for resolution of dystocia in mares: 72 cases (1991–2005). J Am Vet Med Assoc 2007;230(10):1502–5. 9. Nimmo MR, Slone DE, Hughes FE, Lynch TM, Clark CK. Fertility and complications after fetotomy in 20 brood mares (2001–2006). Vet Surg 2007;36(8):771–4. 10. Embertson RM. The indications and surgical techniques for cesarean section in the mare. Equine Vet Educ 1992;4: 31–6. 11. Freeman DE, Hungerford LL, Schaffer D, Lock TF, Sertich PL, Baker GJ, Vaala WE, Johnston JK. Caesarean section and other methods for assisted delivery: comparison of effects on mare mortality and complications. [see comments]. Equine Vet J 1999;31(3):203–7.
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12. Byron C, Embertson RM, Bernard WV, Hance S, Bramlage L, Hopper S. Dystocia in a referral hospital setting: approach and results. Equine Vet J 2002;35(1):82–5. 13. Norton JL. Retrospective study of dystocia in mares at a referral hospital. Equine Vet J 2007;39(1):37–41. 14. Vandeplassche M. The pathogenesis of dystocia and fetal malformation in the horse. J Reprod Fert 1987;Suppl. 35:547–52. 15. Vandeplassche M. Obstetrician’s view of the physiology of equine parturition and dystocia. Equine Vet J 1980;12: 45–9. 16. Blanchard TL, Bierschwal CJ, Youngquist RS, Elmore RG. Sequelae to percutaneous fetotomy in the mare. J Am Vet Med Assoc 1983;182:1127. 17. Blanchard TL, Vaala WE, Straughn AJ, Acland HM, Kenny RM. Septic/Toxic metritis and laminitis in a postpartum mare: case report. J Equine Vet Sci 1987;7:32–4. 18. Frazer GS, Burba D, Paccamonti D, LeBlanc MM, Embertson R, Hance S. Blouin D. The effects of parturition and peripartum complications on the peritoneal fluid composition of mares. Theriogenology 1997;48:919–31. 19. Lynch JL. Retropsective study of dystocia in mares at a referral hospital. Equine Vet J 2007;39(1):37–41. 20. Freeman DE, Hungerford LL, Schaffer D, Lock TF, Sertich PL, Baker GJ, Vaala WE, Johnston JK. Caesarean section and other methods for assisted delivery: comparison of effects on mare mortality and complications. Equine Vet J 1999;31(3):203–7. 21. Byron CR, Embertson RM, Bernard WV, Hance S, Bramlage L, Hopper S. Dystocia in a referral hospital setting: approach and results. Equine Vet J 2003;35(1):82–5.
22. Frazer GS, Perkins NR, Embertson RM. Correction of equine dystocia. Equine Vet Educ 1999;11(1):48–53. 23. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith BJ, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203(8):1170–5. 24. Hong CB, Donahue JM, Giles RC. Equine abortion and stillbirth in central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5(4):560–6. 25. Youngquist RS. Equine obstetrics. In: Morrow D (ed.) Current Therapy in Theriogenology. Philadelphia: Saunders, 1986;. pp. 693–9. 26. Frazer GS, Perkins NR, Embertson RM. Normal parturition and evaluation of the mare in dystocia. Equine Vet Educ 1999;11(1):41–6. 27. Vandeplassche M. Prepartum complications and dystocia. In: Robinson N (ed.) Current Therapy in Equine Medicine 2. Philadelphia: Saunders, 1987; pp. 537–42. 28. Baldwin JL, Cooper WL, Vanderwall DK. Dystocia due to anterior presentation with unilateral or bilateral hip flexion posture (‘dog-sitting’ presentation) in the mare: incidence, management, and outcomes. Proceedings of the American Association of Equine Practitioners, 1991. 29. Hunt RJ. Obstetrics. In: Robinson NE (ed.) Current Therapy in Equine Medicine 5. Philadelphia: Saunders, 2003; pp. 319–22. 30. Frazer G, Burba D, Paccamonti D, LeBlanc MM, Embertson R, Hance S, Blouin D. The effects of parturition and peripartum complications on the peritoneal fluid composition of mares. Theriogenology 1997;48(6):919–31.
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Chapter 257
Cesarean Section David E. Freeman
Cesarean section is one of four options for treating dystocia in mares, and although it is the most effective procedure for this purpose, it is not always necessary. Guidelines for correction of dystocia in mares1–4 can be influenced by several factors, including operator experience and expertise with all available treatment options, foal presentation and position, mare’s temperament, foal viability, duration of dystocia, and severity of vaginal trauma.5–7 Cost factors must also be considered, because cesarean section is generally more expensive than other treatments. Details about other treatments and selection factors can be found in Chapters 255, 256 and 258.
in that study was inappropriate case selection for this procedure, poor technique for CVD, or financial constraints that precluded cesarean section.7 Alternatively, clinicians might have persisted for too long with CVD when the foal was dead, because these mares did not require the rapid delivery needed for a live foal.7 In a more recent study, a 94% survival rate to discharge for mares after CVD (166/177) was attributed to shorter duration of dystocia before CVD at the hospital,8 so drying and swelling of the reproductive tract were probably less severe than after more protracted dystocias.7 Based on these findings, it was concluded that more prompt application of CVD could improve mare survival after this procedure.
Cesarean section versus vaginal delivery
Cesarean section: a team approach
Assisted vaginal delivery (AVD) can be used at the farm to resolve most dystocias, but when it fails, the next most likely step is to anesthetize the mare for controlled vaginal delivery (CVD), usually at a referral hospital. CVD eliminates the mare’s resistance to repositioning the foal and allows her hindquarters to be elevated so that gravity can be used to repel the foal into the uterus. It can be used to deliver live and dead foals, with or without fetotomy for the latter.5,8–10 Although cesarean section is a more invasive procedure than CVD for correction of dystocia, it had a significantly lower short-term mortality (15%) than CVD (29%) in one study.7 The tendency for multiple complications, the nature of those complications (predominantly metritis), and the high mortality after CVD suggest that mares treated by this method experienced more reproductive tract trauma than those treated by cesarean section in that study.7 Trauma could be the product of severity or duration of dystocia, the nature of previous attempts at delivery, and experience of the operator with vaginal delivery. Most likely, the high mortality for mares after CVD
Ideally, the goals for dystocia correction should be to save the lives of the mare and foal and to preserve the reproductive efficiency of the mare. These goals can only be attained with a team approach that involves farm personnel and referring veterinarian at the point of referral, and theriogenologists, surgeons, veterinary technicians, anesthesiologists, and neonatologists at the point of definitive treatment. The following comprehensive hospital approach to cesarean section was successful in one study for mares with dystocia and live foals at the time of presentation.8 When notified of a dystocia referral, the anesthetist, surgeon, neonatal team, and their assistants assemble quickly. Immediately after arrival, the mare is walked directly into an induction stall where a very brief physical examination is performed.8 Assisted vaginal delivery is rarely attempted at the hospital if this was unsuccessful at the farm. Sedation with xylazine (0.8 mg/kg bodyweight [bwt] intravenously [i.v.]) is followed by anesthesia with diazepam (0.08 mg/kg bwt) and ketamine (2.2 mg/kg bwt i.v.), maintained with isoflurane or halothane in oxygen. The hobbled hind pasterns of the anesthetized
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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mare are hoisted to raise the pelvis about 1 meter (3 feet) above the floor, and the foal is repositioned as needed to allow vaginal delivery.8 During CVD, the ventral abdomen is prepared for a cesarean section by clipping and scrubbing for midline celiotomy, and time is closely observed through this and subsequent steps. As soon as it becomes evident that a live foal cannot be delivered within 15 minutes of CVD, a cesarean section is performed. If the foal is alive, it should be delivered within 20 minutes from induction of anesthesia, or 20 minutes from the time of deciding to perform the cesarean section if the mare is already anesthetized. If at any point during CVD it is determined that the foal is dead, the surgeon can then decide to continue with CVD, or use fetotomy or cesarean section. If CVD consumes considerable time or traumatizes the reproductive tract, or the fetotomy requires many cuts, the surgeon must terminate them in favor of cesarean section to avoid the risk of further damage to the mare.7
Surgery For cesarean section after failed CVD and with a live foal, the mare is transferred quickly to a surgery table in dorsal recumbency and the ventral abdomen receives a final surgical preparation. In the case of a dead foal, cesarean section is approached with a lesser sense of urgency. Cesarean sections are generally performed through a caudal ventral midline celiotomy7, 8,11–13, 17 , with the mare in dorsal or dorsolateral recumbency with the ventral midline tilted toward the primary surgeon. The mare is draped as for an exploratory celiotomy, and a 35–40-cm ventral midline incision is directed cranially from a point caudal to the umbilicus for a dystocia and cranial to the umbilicus for an elective cesarean section. The uterine horn containing the hindlimbs (or forelimbs) is exteriorized and large stay sutures or Babcock forceps are placed, one near the tip of the uterine horn, close to the fetal feet, and one toward the body of the uterus, close to the point of the hocks. The assistant surgeon uses these to elevate the uterine horn as far above and away from the abdominal incision as possible. Drapes or large laparotomy sponges are used to isolate the exteriorized uterine horn from the rest of the surgical field and prevent contamination from uterine contents. The uterine wall and chorioallantois are incised from the point of the fetal hock to its fetlock, which is sufficient to accommodate most foals without the risk of uterine tearing from one end of the incision. The amniotic membrane is incised and the hindlimbs are grasped and pulled out of the uterus, and the foal is transferred to a third assistant. As the foal is delivered, the uterus is drawn with it out of the
incision so the cut edges are exteriorized as much as possible. In some cases, sterile obstetrical chains or straps are applied above the fetlock and used to apply careful traction by non-sterile assistants. If a large draft-breed foal is to be delivered by cesarean section, the surgery table should be placed under or close to a hoist so that this can be used to pull the foal from the uterus, if necessary. The umbilical cord is torn near the body of the foal and the ends are clamped temporarily. Strict adherence to the traditional 3–4minute delay before separation of the umbilical cord from the foal is of questionable benefit.3 The foal is quickly transferred to the neonatology room for evaluation and resuscitation if necessary. The uterus is checked for tears and a second foal, the uterine incision is closed (see below), and the abdominal incision is closed as for exploratory celiotomy in a horse.14
Hysterotomy closure Closure of the uterine incision in the mare presents a unique challenge, because the cut edges are prone to hemorrhage, which can be of sufficient magnitude to cause postoperative anemia in many mares and hemorrhagic shock and death in some.3, 5, 7 Therefore, the closure technique must incorporate steps directed at achieving hemostasis along the edges of the hysterotomy. If the chorioallantois is easily separated from the uterus, it can be removed, although complete removal is usually difficult and unsafe during surgery. Regardless, the chorioallantois is peeled back from the cut edges for 10 cm to prevent accidental inclusion in the suture line, and all obvious large bleeders are individually clamped and ligated. Uterine hemorrhage is also controlled by a hemostatic suture line or whipstitch.3, 5 For this, a continuous suture pattern of No. 1 absorbable suture is placed along each edge of the uterine incision in a Ford interlocking pattern (Figure 257.1), with continuous tension on the suture to compress the large plexus of veins between the myometrium and endometrium.5, 15 The uterus is generally closed in two layers with continuous inverting patterns (see below). The exteriorized uterine horn is lavaged with sterile physiological solution and the contaminated drapes are removed from the surgery field. Oxytocin (40 IU i.v.) is then administered to stimulate immediate uterine contractions and aid in expulsion of the placenta. The abdominal cavity can be lavaged with sterile saline or other physiological solution, and the fluid is removed by suction. Mares are monitored for retained placenta after delivery, and if the placenta is not delivered within 3–6 h of recovery from anesthesia, oxytocin in physiological solution is given intravenously intermittently until placental passage. Potassium penicillin (40 000 units/kg bwt i.v., q 6 h),
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(a)
Figure 257.1 Method of using a hemostatic suture or ‘whipstitch’ in a Ford interlocking pattern. Notice that the chorioallantois is peeled back from the cut edges for 10 cm to prevent accidental inclusion in the suture line. From Embertson,3 with permission.
gentamicin (6.6 mg/kg bwt i.v., q 24 h), and flunixin meglumine (1 mg/kg bwt i.v., q 12 h) are administered routinely for 1–3 days or more as required. Most mares recover rapidly after cesarean section and do not require intensive care or fluid therapy.
Hemostatic suture: to use or not? Because the major cause of mortality in mares related directly to cesarean section is hemorrhage from the hysterotomy5 , a hemostatic suture or ‘whipstitch’ (Figure 257.1) is used by many surgeons,3, 11, 12,16–18 and credited with an improved survival rate.15 Although some authors consider the hemostatic suture to be mandatory,3, 13 Cox19 proposed that a sufficient hemostatic effect might be achieved with intraoperative and postoperative oxytocin. Freeman et al.20 described an alternative approach that omits the hemostatic suture and uses the first closure row to achieve the same effect. Ligation of individual vessels alone, although recommended for obvious large bleeders, is unsatisfactory because it is time-consuming and unreliable. The most popular order of patterns for hysterotomy closure in mares is the Cushing pattern followed by Lembert,3, 11, 12,16–18 which could explain the reported high prevalence of uterine edge hemorrhage in mares. The Cushing or Connell patterns would seem to be the least hemostatic of available patterns because they appose alternating segments of incision edges, thereby excluding intervening
(b)
(c)
Figure 257.2 Failure of the Cushing pattern to exert a hemostatic effect on incised edges of a hollow viscus. (a) Method of suture placement so that alternating bites appose one edge against another edge that is not included in the suture. (b) In this view, each bite is illustrated by broken lines to show that it is opposite a cut edge that is not included in the suture pattern. (c) To tighten the suture, it is pulled to create tension lines in the direction of the short arrows. This pulls the edge included in each bite (curved arrow) against an edge without a bite, so that there is minimal compression of tissues along the cut edges, including the intramural vessels.
edges in the suture (Figure 257.2). The alternating segments within each bite are not compressed across the incision to the apposing edge, and this might not be adequate to occlude critical vessels in the uterine wall (Figure 257.2). Hence, the hemostatic suture would be necessary for surgeons that use these patterns in the first row. A Lembert, full-thickness appositional pattern, or a blend of both patterns, could be used in the first row instead of the hemostatic pattern. These patterns encircle apposing edges of the incision and compress them directly against each other along the line of tension, thereby ‘strangulating’ large vessels in the uterine wall (Figure 257.3). One recommended approach is to place the Lembert suture pattern so that it takes a thick section of the uterine wall with a heavy suture material, such as size 2 or 3 synthetic absorbable material (preferably polyglactin 910), and includes at least 2 cm of tissue 0.25 cm from the cut edge. This suture could penetrate the mucosa and thereby increase the chances of encircling all submucosal bleeders. As each bite is placed, the suture is pulled tightly by an assistant to suspend the uterus and maintain tension by weight of the organ, thereby compressing the tissue edges along the same lines of tension as in a whipstitch. A heavy suture material is required to accept the amount of tension needed to achieve this effect. A Cushing pattern can then be used to support and bury the Lembert pattern, because it exposes a smaller amount of suture material to the peritoneal cavity and therefore should reduce the risk of postoperative adhesions.21 A Lembert pattern as the second row over a Cushing would be unlikely to accomplish the same effect because the Lembert bites would be too far back
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Mare: Problems of Pregnancy could be time-consuming in draft-breed mares, because a large hysterotomy is required to deliver a draft-breed foal. The added time could increase the existing high risk of post-anesthetic myopathy in these mares.20
Lembert
Elective cesarean section DEF
(a)
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Figure 257.3 Method by which the Lembert pattern exerts a hemostatic effect on incised edges of a hollow viscus. (a) Method of suture placement so that alternating bites appose one edge against another edge and both are included in the suture. (b) This view shows how the bites (one represented by broken lines) include apposing tissue edges in the suture pattern. (c) As the suture is tightened, it pulls the apposing edges included in each bite (curved arrows) against each other, so that the enclosed tissues, including the intramural vessels, are compressed in the directions of the short arrows.
from the cut edges and hence from the major bleeding vessels. In a study that allowed comparison between mares that did have the hemostatic suture (n = 31) and a comparable group of mares that did not (n = 35), the hemostatic suture did not reduce the risk of anemia.20 Also, the percentage of mares that developed anemia (packed cell volume [PCV] < 30%) after cesarean section in this study (22%) was lower than the 59% prevalence of anemia in 17 mares reported by Juzwiak et al.,17 all of which had a hemostatic suture. The prevalence of severe hemorrhage in this study was 6% in mares without the hemostatic suture and 10% in those that did, which is lower than the 26% reported by Vandeplassche et al.5 in 19 mares. Although the hemostatic suture did not prevent anemia or severe uterine hemorrhage in this study, the only deaths from uterine hemorrhage were two mares in which this suture was not used.20 Although the higher mortality without the hemostatic suture would support its use, the numbers were too small to allow statistical comparison, and differences in treatment for hemorrhagic shock between the two groups could also be a confounding variable. One of the two mares that died from uterine hemorrhage and that did not have the hemostatic suture, had a Cushing pattern as the first layer.20 The other mare had a Lembert pattern; however, this mare underwent resection of 6 meters (20 feet) of strangulated small intestine in the same anesthetic procedure as the cesarean section. It is possible that postoperative cardiovascular complications and clotting disorders from the gastrointestinal disease contributed to her demise or the Lembert pattern was not applied as recommended.20 The disadvantage of the hemostatic suture is the added surgery time, and although this is short, it
Because shortening the duration of stage II of parturition (chorioallantoic rupture to delivery of foal) is the most critical factor for delivering a live foal,7, 8, 22 mares with a history of prior dystocia or with a known physical obstruction to rapid vaginal delivery (healed pelvic fractures and pelvic masses) should be moved to the hospital for foal watch and rapid resolution of dystocia if needed. It is not unusual for such mares to require cesarean section in successive years,7 with three being the highest number reported in one study.12 The prognosis for mare and foal is excellent in such cases,7, 12 but these mares require the same level of prepartum monitoring as any mare with a high-risk pregnancy to accurately predict time of intervention.23 They also benefit from a team approach and facilities that involve theriogenologists, surgeons, and neonatologists.
Combined colic surgery and cesarean section Despite the temptation to do a cesarean section to facilitate surgical correction of a colic, this is an error that is difficult to defend and is rarely if ever truly required for that purpose. Even when foals delivered by cesarean section during colic surgery are close to or at term, they have a low survival rate, even after aggressive intensive care, which demonstrates that an estimate of term is a poor predictor of foal survival.7 In a mare with colic, especially if the mare does not become hypotensive under anesthesia and the prognosis is reasonable, it would be more prudent to leave the foal in place for natural delivery than to deliver a premature foal.7, 24 Also the risk of abortion after colic surgery is low, except in the mare with severe endotoxemia.25 There is no evidence that removal of the foal improves the mare’s chances of survival from colic surgery, and the simultaneous combination of two major procedures could jeopardize the mare’s life.7
Complications Many complications after cesarean section, such as retained placenta, metritis, laminitis, uterine hemorrhage, and abdominal incisional complications have been described in draft-breed mares and with
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Cesarean Section abdominal incisions other than the ventral midline.1, 5, 6, 18 Post-anesthetic myopathy and abdominal wound dehiscence might also be more likely complications after cesarean section in draft mares. If protracted CVD fails in draft mares, the additional time for cesarean section could prolong anesthesia sufficiently to cause post-anesthetic myopathy. Peritonitis is rare after cesarean section,6 and might be more likely after protracted vaginal delivery than after cesarean section.7, 26 Any mare that has had an epidural shortly before cesarean section, such as to facilitate AVD, is at risk of difficulty rising in the recovery stall and might need to be assisted to her feet. Assisted recovery should be considered for older mares (≥ 10 years), especially Thoroughbreds, because these mares are at risk of long-bone fracture during anesthetic recovery. The risk of retained placenta increases after elective cesarean section, which has been attributed to removal of the foal before onset of first-stage labor.12, 16 Retained placenta is also more likely after delivery of a live than a dead foal.27 Contrary to data from older reports, the placenta can be retained for many days in light-breed mares without risk of laminitis,7, 27 and this risk might be reduced by post-operative treatment with broad-spectrum antibiotics and flunixin meglumine.7
Outcome: mare Reported survival rates for mares after cesarean section for dystocia exceed 80%, and can be 100% after elective cesarean section (Table 257.1). When case selection for method of dystocia resolution is
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appropriate, other methods, such as fetotomy, AVD, and CVD can yield superior survival rates, although the differences are small.7–10 In one study on mares with dystocia in Kentucky, average pre-dystocia live foaling rate for all mares with available records was 84%, and post-dystocia live foaling rate for mares that had cesarean section was 72% over the full study period.8 Same-season breeding appears to be rewarding for select dystocia cases,9 including cesarean section, although mares used for producing Thoroughbred racehorses might require at least 6 weeks to allow the abdominal incision to handle natural cover.8 Depending on the surgery date, this could put them close to the end of the designated breeding season; however, this should not factor into the decision-making process, if a cesarean section gives the best chance of delivering a live foal.8
Outcome: foal Because duration of dystocia has an enormous effect on foal survival,3 survival data can be highly variable among different reports, and often disappointing (Table 257.1). In a study on 247 dystocias in mares, the period from hospital arrival to delivery for foals alive at discharge (23.0 ± 14.1 min) was not significantly different than that for non-surviving foals (24.8 ± 10.6 min); however, time from chorioallantoic rupture to delivery of foals alive at discharge (stage II) was significantly less than for foals that did not survive (71.7 ± 34.3 min vs 85.3 ± 37.4 min). Therefore, when the foal is alive, the methods selected for correction of dystocia should be conducted in a
Table 257.1 Mare and foal survival rates after cesarean section
Cesarean section type Emergency
Elective
e
Elective and emergency With colic surgery
Mare survival n (%)
Foal survival – birth n (%)
Foal survival – discharge n (%)
51/63 (80) 41/48 (85) 54/61 (89) (81 to 87)c 10/10 (100)
22/63 (35) NRa NRb
18/63 (29) NRa 19/61 (31)
10/10 (100) (88 to 100)c 8/8 (100) 6/19 (32) 8/8 (100)
9/10 (90) (83 to 86)c 7/8 (88) 2/19 (11) 3/8 (38)
5/5 (100)d 17/19 (89) 5/8 (63)
Authors (year) Vandeplassche5 Freeman et al.7 Byron et al.8 Norton et al.22 Freeman et al.7 Norton et al.22 Watkins et al.12 Juzwiak et al.17 Freeman et al.7
NR, not recorded. There was no breakdown for foal survival by treatment for dystocia in this study, but 11 of 102 foals (11%) were alive at delivery and 5/102 (5%) survived to discharge after correction of dystocia. b There was no breakdown for foal survival by treatment for this category, but 104 of 247 foals (42%) were alive at delivery and 73 (30%) survived to discharge. c Figures given are percentages before and after implementation of a coordinated dystocia management protocol. d One of five mares had three cesarean sections, another had two, and the remainder had one each. e Two mares were elective cesarean sections and 17 were dystocias. a
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sequence that minimizes total delivery time and favors survival of the foal. In one study, foals that survived dystocia were delivered within 90 minutes from start of labor or from mares with term uterine torsion.7 In another study,8 154 minutes was the longest time recorded for delivery of a live foal (method not specified). However, even longer intervals can be expected in rare cases, if the foal can be supported by ex utero intrapartum treatment with oxygen or if the type of dystocia is not associated with cord compression or placental separation.22 Nonetheless, for each 10-minute increase in stage II beyond 30 minutes, the risk of foal mortality at delivery increases by 10% and mortality by discharge increases by 16%.22
Conclusions Cesarean section should be regarded as one option in a continuum of steps required to save the lives of a mare and foal and to restore the mare to normal reproductive activity. It is indicated for correction of dystocias that cannot be corrected by vaginal delivery and for those cases in which other methods would only delay delivery of a live foal. The ideal approach to a typical equine dystocia would involve all individuals, from farm personnel to neonatologists, with prompt selection and application of the appropriate procedure for each case, and with each delivery procedure conducted by an experienced individual. When a cesarean section is performed, it should not be regarded as a last resort, but as the step most likely to produce a satisfactory outcome.
References 1. Vandeplassche MM. Cesarean section in horses. In: Grunsell CSG, Hill FWG (eds) The Veterinary Annual, 14th edn. Bristol: John Wright, 1973; pp. 73–8. 2. Blanchard TL, Varner DD, Elmore RG, Martin MT, Scrutchfield WL, Taylor TS. Management of dystocias in mares: examination, obstetrical equipment and vaginal delivery. Comp Cont Educ Pract Vet 1989;11:745–53. 3. Embertson RM. The indications and surgical techniques for Cesarean section in the mare. Equine Vet Educ 1992;4:31–6. 4. Embertson RM, Bernard WV, Hance SR, Smith S. Hospital approach to dystocia in the mare. P Annu Conv Am Assoc Equine P 1995;41:13–14. 5. Vandeplassche M, Spincemaille J, Bouters R, Bontc P. Some aspects of equine obstetrics. Equine Vet J 1972;4:105–9. 6. Vandeplassche M. Obstetrician’s view of the physiology of equine parturition and dystocia. Equine Vet J 1980;12: 45–9. 7. Freeman DE, Hungerford EE, Lock TE, Sertich PE, Vaala WE, Baker GJ, Johnston JK. Cesarean section and other methods for assisted delivery: comparison of effects on mare mortality and complications. Equine Vet J 1999;31:203–7.
8. Byron CR, Embertson RM, Bernard WV, Hance SR, Bramlage LR, Hopper SA. Dystocia in a referral hospital setting: approach and results. Equine Vet J 2002;35:82–5. 9. Carluccio A, Contri A, Tosi U, De Amicis I, De Fanti C. Survival rate and short-term fertility rate associated with the use of fetotomy for resolution of dystocia in mares: 72 cases (1991–2005). J Am Vet Med Assoc 2007;230: 1502–5. 10. Nimmo MR, Slone DE Jr, Hughes FE, Lynch TM, Clark CK. Fertility and complications after fetotomy in 20 brood mares (2001–2006). Vet Surg 2007;36:771–4. 11. Taylor TS, Blanchard TI, Varner DD, Scrutchfield WI, Martin MT, Elmore RG. Management of dystocia in mares: uterine torsion and cesarian section. Comp Cont Educ Pract Vet 1989;11:1265–73. 12. Watkins JP, Taylor TS, Day WC, Varner DD, Schumacher J, Baird AN, Welch RD. Elective cesarian section in mares: eight cases (1980–1989). J Am Vet Med Assoc 1990;197:1639–45. 13. Trotter GW, Embertson RM. Surgical diseases of the cranial reproductive tract. In: Auer J (ed.) Equine Surgery. Philadelphia: W.B. Saunders Co., 1992; pp. 750–61. ¨ 14. Freeman DE, Rotting A, Inoue OJ. Abdominal closure and complications. Clin Tech Equine P 2002;1:174–87. 15. Vandeplassche M, Bouters R, Spincemaille J, Bonte P. Cesarean section in the mare. P Annu Conv Am Assoc Equine P 1977;23:75–80. 16. Edwards GB, Newcomhe JR. Elective cesarean section in the mare for the production of gnotobiotic foals. Equine Vet J 1974;6:122–6. 17. Juzwiak JS, Slone DE, Santschi EM, Moll HD. Cesarean section in 19 mares: results and postoperative fertility. Vet Surg 1990;19:50–2. 18. Stashak TS, Vandeplassche M. Cesarian section. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea and Febiger, 1993; pp. 437–43. 19. Cox JE. Some aspects of equine obstetrics. In: Grunsell CSG, Hill FWG (eds) The Veterinary Annual, 22nd edn. Bristol: John Wright and Sons, 1982; pp. 153–8. 20. Freeman DE, Johnston JK, Baker GJ, Hungerford LL, Lock TF. An evaluation of the hemostatic suture in hysterotomy closure in the mare. Equine Vet J 1999;31:208–11. 21. Elkins TE, Stovall TG, Warren J, Ling EW, Meyer NE. A histologic evaluation of peritoneal injury and repair: implications for adhesion formation. Obstet Gynecol 1987;70: 225–8. 22. Norton JL, Dallap BL, Johnston JK, Palmer JE, Sertich PL, Boston R, Wilkins PA. Retrospective study of dystocia in mares at a referral hospital. Equine Vet J 2007;39:37–41. 23. Vaala WE, Sertich PI. Management strategies for mares at risk for periparturient complications. Vet Clin N Am Equine Pract 1994;10:237–65. 24. Slone DE. Treatment of pregnant mares with colic: practical considerations and concerns. Comp Cont Educ Pract Vet 1993;15:117–20. 25. Santschi EM, Slone DE, Gronwall R, Juzwiak JS, Moll HD. Types of colic and frequency of postcolic abortion in pregnant mares: 105 cases (1984–1988). J Am Vet Med Assoc 1991;199:374–7. 26. Blanchard TL, Bierschwal CJ, Youngquist RS, Elmore RG. Sequelae to percutaneous fetotomy in the mare. J Am Vet Med Assoc 1983;182:1127. 27. Threlfall WR. Retained placenta. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea and Febiger, 1993; pp. 614–21.
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Chapter 258
Referral Dystocias Rolf M. Embertson
Introduction Natural birthing occurs rapidly in the horse. From the onset of stage II of labor, when the chorioallantois ruptures, the forefeet and muzzle of the foal are usually visible at the vulvar lips within 5 to 10 minutes and the foal usually delivered within 15 to 25 minutes. A slower rate of progression of stage II, or recognition of other abnormalities (e.g., premature placental separation, or a foot or muzzle protruding through the anus), warrant concern and some degree of assistance. Dystocia is one of the few true emergencies encountered in the horse, where delays of 10 to 15 minutes may have a significant effect on survival of the foal. The survival rate of foals decreases rapidly if delivered after 45–60 minutes following chorioallantois rupture, but survival is still possible up to 160 minutes.1, 2 Duration not only affects the foal, but plays an important role in trauma to the mare’s reproductive tract, long-term fertility, and potentially her life. The goal when facing dystocia isdelivery of a live healthy foal in a manner that yields a reproductively sound mare. Dystocias are usually (and ideally) resolved at the farm, with the mare in a familiar environment. Experienced attendants, sometimes with the help of an experienced veterinarian, resolve most dystocias. Occasionally, a dystocia requires more than minor repositioning of the fetus to enable delivery. More difficult dystocias require equipment, facilities, and expertise not usually available at the farm. Duration quickly becomes an important factor. The outcome depends on rapid recognition, intervention, and if necessary, referral. Referring the mare to a nearby hospital is often the best choice for difficult dystocias. Referral hospitals should have the equipment and personnel available to resolve any dystocia. Hospitals that accept the responsibility for treating mares in dystocia should
have personnel that are willing to live this lifestyle during foaling season. Prompt treatment of the mare upon arrival is necessary to achieve the best results.
Procedures used to resolve dystocia There are essentially four procedures used to resolve dystocia in the mare.3 Assisted vaginal delivery (AVD) refers to when the mare is conscious, although perhaps sedated, and is assisted to some degree in vaginal delivery of an intact foal. Controlled vaginal delivery (CVD) refers to when the mare is anesthetized and the clinician is in complete control of vaginal delivery of an intact foal. Fetotomy refers to the procedure where the dead foal is reduced to more than one part to remove the foal vaginally from the mare (see Chapter 256). The mare can either be sedated or anesthetized. Caesarean section (C-section; see Chapter 257) refers to delivery of a foal through an incision in the uterus. Except for fetotomy, these procedures can be used on a live or dead foal. The above procedures are employed as needed to resolve the problem encountered. There is not one best procedure for every situation. It is important to recognize when one procedure will work better than another. Regardless of the procedure, resolution of a dystocia is best performed more by finesse than force. Euthanasia of the mare is another option, although not actually a procedure to resolve dystocia.
Preparation of the hospital Accepting the responsibility of managing dystocias requires readiness of personnel and facilities. At the beginning of each foaling season the approach to dystocias is reviewed with the experienced and new staff members. Potential improvements are discussed. The primary dystocia clinician should be someone who is comfortable performing CVD, C-section, or fetotomy, as there should be no delays
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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in proceeding with whatever method of resolution is needed. Ideally, an intern or seasoned staff member who is competent at anesthesia stays at the hospital during foaling season. This avoids delays in inducing anesthesia of mares with dystocia that arrive with minimal warning. The emergency schedule and phone numbers should be readily available to the person answering the emergency calls. Everyone involved should be aware of their responsibilities before the foaling season commences. During foaling season, the on-call members of the dystocia and neonate teams should be able to arrive at the hospital at the same time as, or before, the mare with dystocia. Controlled vaginal delivery is best performed in a padded surgical induction/recovery stall. This room should be close to the anesthetic equipment, the surgery table, preparation area, and surgery room. A dystocia cart, containing all the equipment and items potentially needed for CVD or fetotomy, should be positioned just outside the induction stall. The foal bed, onto which the newborn is immediately placed for resuscitation, should be at a comfortable height for the clinician and staff. The bed should be on sturdy wheels, so it can be easily moved to just outside the surgery room or to the foal’s stall in the ICU facility. The equipment potentially needed to resuscitate the neonate should be readily available on or near the cart, for example, endotracheal tubes, oxygen supply, intravenous catheters, and intravenous fluids.
Hospital procedure for dystocia The protocol for managing mares with dystocia in different hospitals varies with personnel, facility, referring veterinarians, and proximity to client farms. The approach to dystocia currently employed at our hospital has evolved to the following. During the emergency call, only pertinent information is asked for. Immediately following the call, the on-call dystocia and neonate teams are quickly notified and assemble at the hospital within 15 to 20 minutes. Upon arrival of the mare, debris is rinsed from her mouth, and she is walked into the induction stall. As this occurs, her status is quickly assessed and more information obtained from farm personnel regarding the time of chorioallantois rupture, the perceived reason for dystocia, what has been done, and what drugs have been given. Controlled vaginal delivery is planned unless circumstances dictate otherwise. Xylazine is administered and while waiting for sedation an intravenous catheter is placed, the mare’s tail is wrapped, and the perineal area cleansed. Anesthesia is induced with diazepam and ketamine. An endotracheal tube is passed. Hobbles are placed on the limbs; the mare is hoisted and positioned to allow room for the clinician to work. The forelimbs are
Figure 258.1 Fetal manipulation during controlled vaginal delivery. Note the preparation of the abdomen for a C-section during the procedure.
disconnected from the hoist and the hindlimbs are elevated to a level where the pelvis is approximately 3 feet off the floor. Isoflurane and oxygen are used to maintain anesthesia and mechanical ventilation used to overcome some of the effects of the weight of the abdominal organs on the diaphragm. The fetus is examined and repositioned in an attempt to enable vaginal delivery (Figure 258.1). During manipulation of the fetus, the mare’s ventral abdomen is clipped and prepared for a C-section. When the head and distal forelimbs are straightened and exteriorized, straps are placed over the foal’s fetlocks. A large amount of lubricant is placed in the uterus and vagina and the mare is lowered into lateral recumbency. Traction is applied and the foal is pulled out of the mare. The umbilical cord is quickly clamped and transected and the foal is lifted onto the adjacent foal bed. The foal is placed in sternal recumbency with its head hanging down to allow drainage of fluids. A nasotracheal tube is passed and oxygen started, heart and respiratory rates are determined, blood is drawn for analysis, and an intravenous catheter placed. The clinician’s assessment of the foal determines further treatment and when the foal is taken to its stall. Following delivery of the foal, an attempt is made to remove the placenta. Excess traction should be avoided to prevent tearing and incomplete removal or inadvertent inversion of a uterine horn. If removed, the chorioallantois is examined for any missing tissue, which may be in the uterus. If not removed, it is tied up to provide weight and traction on the retained membranes. The mare is placed on a thick foam mat for recovery. The recovery stall floor should be cleaned of blood and lubricant and then dried to provide stable footing and facilitate an uncomplicated recovery. If the CVD is unsuccessful within 15 minutes, the anesthetist informs the clinician of this time
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Referral Dystocias period. A decision is made at that time whether to continue CVD, perform a C-section, or, if the fetus is known to be dead, a fetotomy. Occasionally, it is quickly determined that CVD cannot be safely performed (e.g., transverse presentation, dog-sitting posture), at which time a C-section or fetotomy is indicated. A decision to perform a C-section should result in delivery of a foal within 20 minutes. The mare is hoisted and positioned on the surgery table with her ventral midline angled slightly towards the surgeon. The final surgical preparation is performed and the mare rolled into the surgery room. During the final preparation of the mare, the surgeon and assistant surgeon also prepare for surgery. Draping begins as soon as the mare is positioned in the surgery room. A caudal ventral midline incision is made, the uterine horn containing the hind feet is exteriorized and isolated with wet laparotomy sponges, and stay sutures are placed to stabilize the horn. An incision is made through the uterine wall and chorioallantois. The amnion is usually opened separately. The hindlimbs of the fetus are grasped, the fetus pulled up and partially out of the uterus. The hindquarters are handed off to an assistant wearing a sterile gown and gloves while the umbilical cord is clamped and transected. The foal is delivered and carried to the nearby foal bed for resuscitation. Occasionally, the hindlimbs are not present in a uterine horn, forcing the uterotomy incision to be made in the body of the uterus with the uterus still in the abdomen. This results in increased contamination of the mare’s abdomen during delivery of the foal from the uterus. The chorioallantois is usually too tightly attached to remove from the uterus at this point. It is separated from the endometrium along the margins of the uterine incision and the incision closed. In our hospital, the incised uterine edges are sutured in a continuous pattern, and prominent bleeding vessels individually ligated, to minimize hemorrhage, prior to closure. The uterus is closed in two layers, using an inverting pattern in either the second or both layers. Intravenous oxytocin is then administered to help contract the uterus. The abdomen is examined for any other significant abnormalities, then lavaged. The volume of lavage fluid depends on the degree of perceived contamination and experience of the surgeon. Occasionally, a catheter/drain is placed to allow for postoperative abdominal lavage. The abdomen is closed and the mare allowed to recover in routine fashion. If the fetus is dead, a fetotomy may be more appropriate than a C-section to remove the foal in a manner least traumatic for the mare. This decision is influenced by the reason for dystocia, experience of the surgeon, and financial constraints. A fetotomy should not involve more than two cuts.
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Post-dystocia management of the mare The mare, and hopefully the foal, is discharged to the farm as soon as appropriate. Regardless of the method used to resolve the dystocia, the uterus and vagina have been contaminated and traumatized. Antibiotics and non-steroidal antiinflammatory drugs are indicated postpartum. The uterus should be lavaged once or twice daily for a few days and the mare should receive some degree of exercise. Thorough lavage of the uterus provides the opportunity to palpate the caudal uterus, cervix, and vagina for any significant abnormalities. It is not unusual for a mare to retain her placenta beyond the accepted norm of 3 hours. There are several approaches to treatment of retained fetal membranes.4 Treatment with oxytocin is usually effective. The author generally administers 40 IU in 1 liter of lactated Ringer’s solution, administered intravenously over 45–60 minutes within 3–4 hours postpartum. If the mare becomes too uncomfortable the rate is decreased. One must be aware that this may infrequently contribute to uterine eversion and, rarely, hemorrhage in the broad ligament. If not effective, the oxytocin dose is doubled and repeated as needed at 4- to 6-hour intervals until the placenta is passed. Manual traction applied to the placenta during uterine lavage can be helpful. It is much preferable to aggressively treat the retained placenta, than the metritis and laminitis that may result from a prolonged retained placenta.
Considerations regarding resolution of dystocia The effects of duration of dystocia on the fetus and the mare cannot be overemphasized. The time period from when the chorioallantois ruptures until dystocia is recognized (> 10 minutes), the mare is loaded in a trailer and transported to a hospital (> 20 minutes), and then unloaded, walked into the hospital, anesthetized and hoisted (∼ 10 minutes) combines to at least 40 minutes duration and usually more under the best circumstances. This should provide enough motivation for the clinician to work as quickly and efficiently as possible. Progress should be critically assessed after 15 minutes of manipulation. Assisted vaginal delivery is rarely attempted on referral dystocias. This has usually already been tried at the farm by experienced staff and further attempts are generally a waste of valuable time. Controlled vaginal delivery has significant advantages over continued AVD. Anesthesia and elevation of the hindquarters facilitates manipulation of the fetus without the mare straining. The clinician can work
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efficiently, in a relatively comfortable position, with minimal concern of injury to them or others. During induction and positioning of the mare, her perineal region is cleansed and the primary clinician scrubs their hands and arms. Further contamination of the uterus beyond what is already present should be minimized. Sterile gloves and sleeves are generally a hindrance, and not used. The position, posture, and presentation of the fetus can usually be quickly determined. However, viability of the fetus can be difficult to establish if the ventral thorax cannot be reached. The most accurate determination of a live fetus is palpation of a heartbeat through the thoracic wall. If uncertain, assume the foal is viable. Generally, foals that have not engaged in the pelvic canal remain viable for a longer period than those that have. Experience and judgment play a large role in determining how best to reposition the fetus for delivery and if and when CVD should be abandoned and C-section or fetotomy performed. It is very important to minimize trauma to the mare’s reproductive tract and to the foal during delivery. Initially, the mare has enough natural lubrication with the fluids and tissues involved. With increasing duration, the tissues of the placenta and hair coat of the foal lose this lubrication. In addition, the walls of the uterus, cervix, and vagina become edematous, less slippery, and more susceptible to trauma. Supplementing lubricant to the uterus prior to manipulation may allow easier repositioning of fetal limbs and head. However, excessive lubrication may make grasping and repositioning a lower limb or muzzle more difficult. Once the fetus is manipulated into the desired positioned for delivery, with straps in place on the distal forelimbs, a copious amount of lubricant should be added to the uterus and vagina to reduce as much friction, and thus trauma, as possible as the foal is being pulled out of the mare. Different lubricant products are available. The author uses a commercially available non-sterile product containing sodium carboxymethylcellulose. Approximately 2 to 4 liters are used to facilitate CVD. Excessive force may result in uterine, cervical, and vaginal tears in the mare and fractured ribs and torn abdominal muscles in the foal. A brief explanation follows of manipulations that may be useful during CVD to resolve some causes of dystocia. Fetal manipulation is usually performed with the mare’s hind end hoisted, but her forelimbs lying in left lateral recumbency. Following correction of the abnormality, the mare is lowered into left lateral recumbency and the foal is pulled out. If the fetus is rotated abnormally within the mare, following correction of any malposture, the mare may be rotated around the fetus into right lateral recumbency
to achieve a more dorsosacral position prior to applying traction to deliver the foal. In most dystocias with a cranial presentation, positioning the head over the forelimbs, within the pelvic canal, enables a vaginal delivery. Ventral deviation of the head (flexed) can be corrected by repelling the fetus, then cupping a hand around the muzzle to draw it into the pelvic canal. This may require both hands. If the head is turned back to either side, a finger is introduced into the corner of the mouth to pull the head straight until the muzzle can be cupped with a hand as it is directed into the pelvic canal. It can be difficult to reach the corner of the mouth of the fetus. A snare placed in the mouth and around the poll of the head may be needed to pull the head into the pelvic canal. The most common malposture encountered is unilateral or bilateral flexed forelimbs. Some of these are actually congenital flexural deformities. Usually the fetus can be repelled and a cannon bone grasped. A hand is slid down the cannon bone to the fetlock as the leg is extended, and then the foot is cupped and drawn up into the pelvic canal. The same maneuver is performed on the other limb if needed. Occasionally, the pelvic entrance does not provide enough room to allow both elbows through at the same time, requiring one leg to be advanced slightly ahead of the other. During delivery, if the foal becomes stuck at the pelvis or stifles, repelling and slightly rotating the foal, followed by pulling the foal’s head and forelimbs down towards the mare’s hind feet, will resolve the obstruction. Caudal presentation of the fetus is very unusual. If both hind feet can be manipulated into the pelvic canal, vaginal delivery is usually accomplished. If this is not feasible, a C-section is indicated. Cranial presentation, but with three or four feet palpable, is usually safest resolved by C-section. Unusual circumstances may alter the normal hospital approach to referral dystocia. Rarely, a mare may arrive down in a trailer and unable to stand. Usually these mares have encountered a prolonged attempt at resolution at the farm. Ideally, the trailer is positioned under a hoist that extends beyond a recovery stall to the outside. The mare is anesthetized, pulled to the end of the trailer, hoisted, and moved by rail or mobile table to the surgery area. Depending on the circumstances, the mare is then prepared for CVD, C-section, or fetotomy. Also rarely, a mare may arrive that has partially eviscerated bowel during foaling through a tear in the caudal uterus and/or cranial vagina. If the farm personnel have minimized bowel trauma and contamination and prevented further straining by the mare, it is still possible to end with a live foal and mare. The affected bowel is lavaged and returned to the abdomen. The vulva is sutured closed and a C-section performed. Following
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Referral Dystocias delivery of the foal and uterine closure, the affected bowel is treated as needed and the tear assessed and closed, if possible. These tears are usually too far caudal to address abdominally, requiring repair from a vaginal approach subsequent to recovery. The abdomen should be lavaged, a catheter placed for subsequent lavage, and then closed. Postoperative uterine adhesions are of concern for these mares.
Procedures for resolution of dystocia As stated previously, the decision regarding which procedure to use to resolve a dystocia depends on the situation encountered. Many factors play an important role in this decision, including cause and duration of dystocia, viability of the fetus, equipment and facilities available, financial perspective of the owner, and experience and judgment of the clinician, among others. Reviewing and understanding the results of different reports regarding dystocia resolution should provide the clinician with the most appropriate approach to the dystocia encountered.1, 2,4–7 There are few reports evaluating a case series of dystocias in mares where resolved in an equine hospital.1, 2, 5 In one report from a private equine hospital, the percentage of dystocias (n = 247) resolved, by procedure,was as follows: AVD 0%, CVD 71%, C-section 25%, and fetotomy 4%.1 The mean duration from chorioallantois rupture to delivery in that report for CVD was 101 minutes and for C-section was 129 minutes. The median duration to resolution for all dystocias was not reported. The percentage of referral dystocias (n = 47) resolved, by procedure, at a university hospital using a similar coordinated dystocia management protocol was as follows: AVD 11%, CVD 23%, C-section 43%, fetotomy 11%, and euthanasia 2%.2 The median duration from chorioallantois rupture to dystocia resolution in that report was 250 minutes. It is difficult to compare and contrast different studies as different factors play a role in how a dystocia is approached. In an environment where the broodmare farms are near a hospital and the duration of dystocia until definitive treatment is relatively short, CVD can be an effective approach to resolution of dystocia. Increasing duration of dystocia, due to distance to a hospital and time for recognition or decisions, in addition to other factors, may make other approaches more appropriate than CVD.
Survival of the mare and foal Survival of the mare in dystocia depends on several factors, but probably the most important is the avoidance of prolonged and inappropriate attempts to deliver the foal. Minimizing the duration of manipula-
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tion and trauma to the tissues in and around the reproductive tract will yield better results for survival and fertility. The causes of death related to dystocia in mares would include, among others, tears to the reproductive tract with evisceration, hemorrhage, or infection, endometritis with laminitis, musculoskeletal injuries, trauma to the intestinal tract, hemorrhage from uterine arteries, and financial constraints. It is difficult to compare short-term survival rates of mares with dystocia between different studies. In studies reporting all the procedures used to resolve dystocia, mare survival rates ranged from 82% to 91%.1, 2, 5 Studies focusing on the results following fetotomy, performed in a referral hospital or on a farm, yielded mare survival rates of 90% to 96%.4, 6, 7 The mare survival rate following CVD, as defined in this chapter, was 94% in one report.1 Mare survival rates following C-section to resolve dystocia have ranged from 82% to 89%.1, 2, 7, 8 Foal survival rates from mares in dystocia resolved in a referral hospital are affected by duration of dystocia (see below). In studies where all the procedures used to resolve dystocia were examined, the percentage of foals born alive ranged from 11% to 44%.1, 2, 5 The percentage of foals that were discharged from the hospital in those studies ranged from 5% to 30%. The percentage of foals discharged from the hospital following CVD was 32% in one report.1 The percentage of foals discharged from the hospital following C-section to resolve the dystocia has ranged from 0% to 31%.1, 5, 8 Interestingly, an older study examining foal survival following C-section for dystocia at farms yielded a rate of 30%.7 Following live birth, the most common reasons for euthanasia or foal death were malformation of the foal or neonatal maladjustment syndrome.
Duration of dystocia Duration of dystocia directly affects foal survival. Two studies have shown significant differences in duration of dystocia (from chorioallantois rupture to delivery) between foals surviving and not surviving to discharge from the hospital.1, 2 In one study, the group of surviving foals had a dystocia duration mean of 71.7 minutes, median of 60 minutes, and range of 32 to 154 minutes.1 The non-survivor foals had a dystocia duration mean of 85.3 minutes, median of 79 minutes, and range of 31 to 254 minutes. The duration means were statistically significant. In the other study, the group of surviving foals had a median dystocia duration of 71 minutes and range of 30 to 162 minutes and the non-survivor foals had a median dystocia duration of 249 minutes and range of 30 to 783 minutes.2 The median dystocia duration was significantly different. In these studies, no foal
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survived to discharge following dystocia duration of more than 162 minutes. Survival of the mare is also affected by duration of dystocia, as increasing duration usually results in increasing trauma to the reproductive tract and adjacent tissues. However, some of the causes of mare death are acute injuries and not related to the period of time in dystocia. Fertility is directly affected by trauma to the reproductive tract.
The above information shows evidence that increasing duration of dystocia prior to resolution decreases conception rates and live foaling rates. Live foaling rates are adversely affected by dystocia, but can approach normal. It is difficult to compare these papers because of the different factors involved. However, these papers provide strong evidence that minimizing trauma to the mare’s reproductive tract during resolution of dystocia will improve postdystocia fertility.
Fertility Fertility of the mare following dystocia has been evaluated in several studies.1, 4, 6, 8, 9 Byron’s study evaluated fertility following all procedures used to resolve dystocia. The other studies examined fertility following only fetotomy or only C-section. In Byron’s paper, cumulative pre-dystocia live foaling rates for the Thoroughbred mares in the study were 84%.1 Cumulative post-dystocia live foaling rates were 67% (72% for C-section and 66% for CVD). In Abernathy’s study of C-section mares, most of which were also in Byron’s study, cumulative predystocia live foaling rates were 79%.8 . Cumulative post-dystocia live foaling rates over a 3-year period were 68% and 52% for mares in dystocia < 90 minutes and > 90 minutes, respectively. Cumulative post-dystocia pregnancy rates (pregnant at 45 days) over a 3-year period were 79% and 64% for mares in dystocia < 90 minutes and > 90 minutes, respectively. Carluccio evaluated the 45-day pregnancy rate the same year following fetotomy to resolve dystocia.4 This study showed a 79% pregnancy rate (at 45 days) in mares who had undergone fetotomy compared to an 82% rate in control mares (normal parturition). Nimmo’s study evaluated pregnancy and live foal rates for mares following fetotomy to resolve dystocia.6 The cumulative pregnancy rate was 86% and the foaling rate was 86%.
References 1. Byron CR, Embertson RM, Bernard WV, Hance SR, Bramlage LR, Hopper SA. Dystocia in a referral hospital setting: approach and results. Equine Vet J 2002;35:82–5. 2. Norton JL, Dallap BL, Johnston, Palmer JE, Sertich PL, Boston R, Wilkins PA. Retrospective study of dystocia in mares at a referral hospital. Equine Vet J 2007;39:37–41. 3. Embertson RM. Editorials – Dystocia and caesarean sections: the importance of duration and good judgment. Equine Vet J 1999;31:179–80. 4. Carluccio A, Contri A, Tosi U, De Amicis I, De Fanti C. Survival rate and short-term fertility rate associated with the use of fetotomy for resolution of dystocia in mares: 72 cases (1991–2005). J Am Vet Med Assoc 2007;230:1502–5. 5. Freeman DE, Hungerford EE, Lock TE, Sertich PE, Vaala WE, Baker GJ, Johnston JK. Cesarean section and other methods for assisted delivery: comparison of effects on mare mortality and complications. Equine Vet J 1999;31:203–7. 6. Nimmo MR, Slone DE, Hughes FE, Lynch TM, Clark CK. Fertility and complications after fetotomy in 20 Brood Mares ( 2001–2006). Vet Surg 2007;36:771–4. 7. Vandeplassche M.. Obstetrician’s view of the physiology of equine parturition and dystocia. Equine Vet J 1980;12: 45–9. 8. Abernathy KK, LeBlanc MM, Embertson RM, Pierce SW, Stromberg A. Survival and fertility rates after cesarean section. Proc Am Assoc Equine Pract 2009;55:268. 9. Juzwiak JS, Slone DE, Santschi EM, Moll HD. Cesarean section in 19 mares: results and postoperative fertility. Vet Surg 1990;19:50–2.
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Chapter 259
Periparturient Hemorrhage T. Douglas Byars and Thomas J. Divers
Periparturient hemorrhage in the broodmare includes the abdominal blood loss events associated with pregnancy. The hemorrhagic event can occur prior to parturition or during or post-foaling. The clinical presentations can be subtle or profound, with hemorrhagic peracute blood loss into the abdomen, broad ligament, or uterus. The clinical presentation may initiate as hemorrhagic shock, abdominal pain, and weakness with compartmental syndrome, i.e., broad ligament containment, or as mural to the uterine wall followed by the vascular and uterine wall failures causing blood loss into the uterus. Hemorrhagic discharge from the uterus may be dramatic in a few cases.
Pathogenesis and epidemiology The utero-ovarian artery arises from the external iliac artery and is short and broadened to the ovary. The artery then traverses along the cornu of the uterus within the broad ligament and eventually gives rise to the middle uterine artery along the uterine wall. The anatomical site of blood loss can directly relate to the prognosis for survival by virtue of the opportunity for eventual tamponade such as is provided within the broad ligament. If the uterine artery rupture occurs proximal to the ovary, arterial hemorrhage within the larger abdominal cavity has less tendency for compressive containment of the vascular lesion. Whenever rupture occurs clot formation is initiated by platelet contact activation with the vascular endothelium, the extrinsic and intrinsic clotting proteases, and vasospasm. If the vascular lesion represents a large uterine artery tear the coagulation system is overwhelmed and unable to provide a stabilized (polymerized) clot to slow or stop the hemorrhage. Hemorrhage usually occurs in multiparous broodmares of middle to older ages. Although a mare may have bled in association with a previous foal-
ing, the tendency to bleed with subsequent foalings is not consistent. Recognition of aging in broodmares with periparturient hemorrhage has been associated with decreased serum copper levels. Previous pregnancies and foaling are likewise not consistent, and rarely maiden mares may also experience periparturient rupture of the uterine artery. In some mares hemorrhage is subclinical and a hematoma is discovered at a later date during a routine reproductive examination. Each hemorrhagic event can cause fibrosis and a loss of arterial elasticity.
Diagnosis Diagnosis of suspected uterine artery hemorrhage is determined by the clinical signs of hemorrhagic shock, palpation of a painful enlarged broad ligament hematoma, or the presence of a painful hematoma of the uterine wall. Ultrasound diagnostics are pivotal to assessing the hemorrhage. A 5-MHz reproductive probe is adequate and when placed onto theventral abdomen free blood within the abdominal cavity can be observed as a ‘ground-glass’ echodense fluid that swirls with the motion of abdominal excursions or visceral movement (Figure 259.1). If abdominal clots are also present these are observed as amorphous clumps outside of the bowel or uterus within the abdominal cavity. If particulate fluid is noted the particles could represent bowel rupture (usually cecal) with peritonitis. Another source of both blood and particulate material may be observed with a tear or hole in the uterus. A peritoneal centesis with microscopic examination of the cellular or particulate content of the fluid can assist in determining if bowel rupture or uterine induced peritonitis is present. In rare cases, bladder rupture with uroperitoneum may be present. Intrauterine hemorrhage will usually result in significant amounts of blood noted from the vagina with excessive fibrinolytic activity. Rectal examination may reveal an enlarged brood ligament,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 259.1 Hemorrhage that is free in the abdominal cavity can be identified by swirling and the “ground glass” appearance of the fluid (arrow) often with viscera in the background (arrowhead).
and vaginal examination is often broad with the exception of obvious tears. Tears to the uterus palpate as painful by either rectal or vaginal examination.
Differential diagnosis The most common differential ruleouts to suspected periparturient hemorrhage are a tear in the uterus or visceral rupture. Although infrequent, the presence of hemorrhage in both the thoracic and abdominal cavities in association with parturition is most consistent with a diaphragmatic hernia.
Clinical signs Mares may be moderately depressed with pale mucous membranes. The heart rate is usually elevated and fever is similarly not present early in the course of the blood loss. Mares with significant hemorrhagic shock may be shaking with muscle fasciculations, sweating, pale oral and vulvar mucous membranes, cold ears and distal limbs, and elevated heart rates near 80 bpm or greater. Some mares will be weak and appear ataxic. A shrill whinny may occasionally be noted. Mares with vaginal blood loss should not be confused with local blood loss due to vaginal varicosities.
Treatment If the foal is in danger it should be separated from the mare. If the mare appears stable it is best for the mare not to remove the foal as anxiety from removal of the foal, repeated rectals, twitching, etc. may increase blood pressure and decrease clot formation. Mares
with colic should receive flunixin meglumine and additive analgesics if needed. In mares with mild clinical signs, stall rest and symptomatic care may be the limits of treatment. In mares with obvious or impending hemorrhagic shock an intravenous catheter should be placed and 5 mL/kg of hypertonic saline may be administered followed by crystalloids containing 20 g of antifibrinolytic aminocaproic acid (loading) followed by 10-g doses three to four times daily in fluids. Although hypertonic saline is indicated for mares with hypotensive shock it should not be used in mares that are stable since a rapid increase in blood pressure may displace an already formed clot. If colloids are administered the choices include plasma, whole blood, or synthetic hemoglobin (Oxyglobin; OPK Biotech). Hetastarch is currently contraindicated as being potentially fibrinolytic with dilution of clotting proteins. Whole blood can be provided from a suitable donor horse or as an autotransfusion. The autotransfusion is performed in mares with intra-abdominal hemorrhage. A Foley catheter or 24-Fr thoracic tube is placed into the ventral abdomen and the blood is collected into sterile acid-citrate-dextrose solution (ACD) bags or bottles and transfused back into the mare. Approximately one-third of the ACD can be removed as many clotting factors are no longer present/active in the abdominal blood. Oxygen may be considered as an adjunct treatment. Corticosteroids (anti-shock dose of 1–2 mg/kg i.v.) and naloxone (0.02–0.03 mg/kg i.v.) should be administered to stabilize endothelial membranes and neutralize vasodilatory endorphins, respectively. Antibiotics are considered supportive or indicated as a precaution against secondary bacterial translocation from hypoperfused bowel. Hypotensive resuscitation can be provided by the intramuscular administration of 1 mL of acepromazine three to four times daily. The rationale for decreasing the blood pressure is to decrease blood flow and pressure at the site of the arterial lesion. Likewise, modest fluid therapy to maintain low normal blood pressure can help perfuse organs yet not dislodge clots. Ideally systolic blood pressure should be 60–80 mmHg, and urination should be observed to help insure that adequate perfusion pressure is present. Abdominal compression by wrapping the abdomen may assist in tamponade. Abdominal compression is utilized in humans and small animals, and may be provided to the mare by abdominal bandaging with Elastocon or the application of a commercial abdominal bandage (BOA; Wire2Wire, Paris, KY). The BOA bandage is adjustable and secured using Velcro. Surgery to tie off the uterine artery is difficult and requires the dexterity of a skilled surgeon capable of providing manual ligation. Ligatures have
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Periparturient Hemorrhage been known to ‘slip’ post-operatively although some mares have survived due to heroic surgical intervention
Prognosis The prognosis for survival is based upon the site of the hemorrhage, the response to treatment, and sufficient stall rest. A fully polymerized clot will occur in 5–7 days but the mare should not be moved for an additional 5–7 days. A 30-day post-hemorrhage reproductive examination may determine the suitability of breeding the mare back within the same year. Breeding with ovulation from the non-hemorrhagic contralateral ovary is preferable.
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Bibliography 1. Britt B. Postpartum hemorrhage In: Robinson NE (ed.) Current Therapy 5. Philadelphia: W.B. Saunders/Elsevier, 2003; pp. 327–30. 2. Lofstedt RM. Hemorrhage associated with pregnancy and parturition. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea and Febiger, 2003; pp. 600–2. 3. McKinnon AO. Reproductive ultrasonography Part III. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Wilkins, 1998; pp. 196–7. 4. Sisson S, Grossman JD. Anatomy of Domestic Animals, 4th edn. Philadephia W.B. Saunders, 1969; 680 pp. 5. Sprayberry KA. Hemorrhage and hemorrhagic shock. In: Bluegrass Equine and Critical Care Symposium, 1999, pp. 1–16. 6. Steiner JV, Hillman RB, Orsini JA, Divers TJ, Schlafer DH. Reproductive system. In: Orsini JA, Divers TJ (eds) Equine Emergencies. St Louis: W.B. Saunders/Elsevier, 2008; p. 427.
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Chapter 260
Retained Fetal Membranes Walter R. Threlfall
Introduction Retained fetal membranes, also referred to as retained afterbirth and retained placenta, have had a variety of literary definitions. All authors consider a retained placenta to be the failure of passage of part or all of the allantochorionic membrane with or without the amnionic membrane within a specific time limit. This length of time postpartum, however, varies and has included 30 minutes,1 1 hour,2 1.5 hours,1 2 hours,3 3 hours,4 and 6–12 hours.5 Retained placenta is reported to be the most common postpartum problem in the mare.6 The incidence of this condition is difficult to assess owing to the lack of a specific postpartum time interval in the definition. Its occurrence is, however, reported to be between 2% and 10.5% of foalings.7 There is a reported higher incidence in draft mares,8, 9 and after dystocia,7 prolonged gestation,10 hydrops,11 and cesarean.7 Although retained placentas have been reported to occur at a higher incidence in association with abortion, stillbirth, and twinning in lightweight horses, one report12 indicated no increase with these conditions if they occurred without accompanying dystocia. Furthermore, the reported occurrence of retained placenta did not differ between the birth of a weak or diseased foal and that of a healthy foal. The sex of the foal did not influence the incidence of retained placenta. The age of the mare may be related to the incidence of retained placenta on some farms; the reason for this farm effect has not been reported. Mares over 15 years old may have a significantly higher incidence of retained placenta than younger mares. Time of year appeared to influence the incidence of retained placenta only on certain farms; when a difference existed there was a higher incidence after March 31(northern hemisphere). Mares that were barren or maiden or had foaled the previous year had no difference in the incidence of retained placenta after foaling. There was
no reported difference in placental passage in mares artificially bred at foal heat versus mares artificially bred at other heats. Mares bred naturally at foal heat versus by artificial insemination reportedly experienced an increased incidence of retained placentas.13 In uncomplicated placental membrane retention cases, pregnancy rates after first breeding, at the end of the breeding season, at midgestation, and at term may6, 14 or may not12 be affected by foal heat breeding. Mares that have had retained placentas before demonstrated a threefold greater probability of having retained placenta after foaling than mares without this history.12 A possible reason for this increase is that formation of pathological adherences between the endometrium and chorion during the first retention of the placenta may recur at subsequent pregnancies. Mares that develop uterine or systemic infections before or during pregnancy have an increased occurrence of retained placentas. Any debilitating condition such as senility, excessive fatigue, poor condition, or poor environment may increase the incidence of retained placenta and puerperal infections. Retained placenta occurrence is reportedly 28% after fetotomy and averages 50% following cesarean section, with the percentage occurring with a live fetus at the beginning of surgery, twice that with a dead fetus.15 In the mare, normal separation of trophoblastic cells from the uterine epithelium and expulsion of placenta through the cervix and vagina occur in the third stage of labor (Figure 260.1).16 It is assumed that rupture of the umbilical cord causes collapse of fetal placental vessels and shrinkage of chorionic villi (Figure 260.2).1 Postpartum uterine contractions reduce uterine size and probably the amount of blood circulating within the endometrium; thus the maternal crypts relax. Uterine contractions, originating at the apices of the horns and progressing towards the cervix, evert the apical pole of the allantochorion, squeezing the base of endometrial crypts and dilating
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Gravid uterine horn
Ovary Oviduct
Non gravid uterine horn
Uterine body
Cervix
Figure 260.1 Postpartum uterus with placenta attached to endometrial surface.
them. Moving towards the cervix, the everted allantochorion pulls the chorionic villi from endometrial crypts (Figure 260.3). Presence of the allantochorion at the cervix stimulates oxytocin release and additional uterine contractions, which are accompanied by further abdominal expulsive efforts. Portions of the freed placenta pass through the vagina and vulva adding the weight of the placenta to the expulsive efforts of the abdomen and uterus. Oxytocin seems to play a role in postpartum uterine contractions since there is an increased concentration of this hormone in the blood during the third stage of labor.17
Causes The cause for failure of the placenta to be passed within the first 3 hours of stage 3 of parturition is unclear. Attachment of the microcotyledons (mi-
crovilli) are tightly adhered by the seventh month of gestation.7 This attachment makes complete separation difficult to impossible unless the microvilli are released. It is reported that either the gravid horn18, 19 or the nongravid horn allantochorion of a retained placenta is attached to the endometrium, thus preventing expulsion. A third report20 stated that it is neither horn but actually the area between the two horns that remains attached. The generalized opinion today is that areas of allantochorion near the tip of the nonpregnant horn have failed to separate. Retention of a portion of the placenta is more likely to occur in the nongravid horn than in the gravid one. This is thought to be due to the actual thickness of the chorioallantoic membrane being thinner in this area before it becomes edematous from retention.21 This could allow it to tear more easily when the weight of the already passed placenta is pulling on it. Clinical and pathological observations suggest
Microvilli Detachment of placenta from gravid horn
Figure 260.2 Separation of microvilli from endometrial crypts beginning within gravid horn.
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Figure 260.3 Continued separation with eversion of placenta.
mechanical interference and hormonal imbalance as possible causes. In addition, allantochorion thickness, length of villi, and degree of attachment of the placenta increase from gravid to nongravid horn. An explanation15 for these characteristics includes the presence of more developed microvilli in uterine horns than in the body; also, microvilli are more branched and larger in the nonpregnant horn. Furthermore, folding of the placenta and endometrium within the nongravid horn is more pronounced.22 Lastly, uterine involution occurs at a slower rate in the nongravid than in the gravid horn. These events, working in combination or alone, help to explain the slower release of the placenta from the nongravid horn. Placental edema is reported to be partially responsible for retention.23 Presence of edema and increased weight supposedly lead to weighing of placentas. The placenta should reportedly weigh 6.4 kg. However, the weights of 200 normal Thoroughbred placentas ranged from 4 to 8 kg, so this practice is no longer recommended. Furthermore, edema was apparently more prevalent in the gravid than in the nongravid horn. The failure of sufficient concentrations of oxytocin to be released at the appropriate time could also add to the occurrence of retained placenta. The increased occurrence of retained placentas following a dystocia is probably related to the trauma of the endometrial tissue and fatigue of the myometrium. Bloodborne or ascending infection after cervical relaxation and endometritis during pregnancy could induce inflammatory choriouterine adherences and delay separation of the placenta.1 This type of lesion, along with placentome sclerosis, has also been observed in the cow with retained placenta and after experimental infections with Brucella abortus and Aspergillus species. It has been suggested1 that contaminated air entering the uterus at the time of parturition could carry in bacteria that could reduce
speed of placental passage. It is difficult to imagine how these contaminating bacteria could have such an effect within 30–60 minutes. It does seem more plausible that bacteria entering at the time of breeding or foaling owing to poor hygiene could have an effect on the next pregnancy. Mares that have uterine infections before breeding, that become infected at breeding, or that are bred to an infected stallion may have retarded placental separation at the next parturition.1 Increases in fetal membrane retention are more likely if the mare had severe placentitis prepartum.21 Infection as a cause of retained placenta may not be as important as was once thought since infections between endometrium and placenta are usually present near the internal os and surrounding area.1 These infections have not led to placental retention and do not account for the last attachment freed being located near the tip of the uterine horn. Many authors believe that retained fetal membranes are primarily due to uterine inertia and hormonal imbalance because mares affected with retained placentas do not exhibit the mild signs of colic associated with postpartum uterine contractions.24 This hormonal imbalance could result in abnormal myometrial activity, not necessarily decreased movement or contractions. Excellent results obtained with treatment of retained placenta with oxytocin suggest a possible deficiency or lack of this hormone in the circulation.7, 21, 25 One report on a limited number of pony mares revealed that oxytocin increased in the serum during the second and third stages of a normal labor.17 Furthermore, when oxytocin was used to induce parturition, increasing doses shortened the time to the second and third stages of labor.26 Although some authors believe uterine inertia can cause retained placenta in the cow, it is probably not the sole cause in the mare since, reportedly,7 uterine tone frequently increases after dystocia with placental retention. It is widely
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Retained Fetal Membranes accepted that uterine inertia can be caused by low blood calcium, overstretching of the myometrium as in hydrops allantois, twinning and oversized fetus, myometrial degeneration resulting from bacterial infection, and myometrial exhaustion after dystocia in poorly conditioned animals.1 Uterine inertia was also incriminated in premature delivery in which the normal sequence of hormonal changes had not occurred. Low concentrations of circulating cortisol at calving have been associated with retained placenta but this has not been reported in the mare. Decrease in plasma progesterone before calving was slower in cows that retained the placenta. High blood concentrations of progesterone at calving interfered with uterine contractions and with detachment and expulsion of the placenta. A report on one mare with a retained placenta indicated serum progesterone remained elevated at prepartum concentrations instead of decreasing as occurred in all mares that passed the placenta within 30 minutes.27 Another report indicated that relaxin activity remained elevated in a mare with a retained placenta.28 In both the above reports the presence of the placenta within the uterus may have acted as a source of progesterone or relaxin. The role of prostaglandins (PGs) in passage of the placenta remains unknown. A report29 on retained placentas in cows indicated that prostaglandin F2α (PGF2α ) decreased and prostaglandin E2 (PGE2 ) increased in association with retained placenta. It has been reported that Highland ponies on one breeding farm all (four out of four) retained their fetal membranes during one breeding season.30 The cause of the retention was not determined except in one mare, which had a lowered calcium concentration in the blood. This number may have been increased if the calcium had been tested for the biologically active form and not total serum calcium.30 Another possible method to determine whether calcium was the cause of the retained fetal membranes would be to take serial samples for calcium testing and observe whether a decrease was occurring.31 Fescue grass (Festuca arundinacea) infected with the endophyte Acremonium coenophialum has reportedly caused retained fetal membranes.32 Although cows that have become too obese through being fed high-nutritive diets have an increased incidence of retained fetal membranes, this has not been reported to be the case in a horse.33 The Friesian breed of horses has a higher incidence of retained fetal membranes than other breeds of horses.34 In a study based on 436 Friesian mares the incidence of retained placentas (longer than 3 hour retention) was 54%. This relationship may be at least partly due to inbreeding.35 Mares bred earlier in the breeding season had a lower percentage (52.7%) of placental retention than did mares bred later in the
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season (59.5%). This study did not find a correlation between sex of the foal and retained membranes, as was shown in another report.12 There was no relationship between month of breeding or gestational length in regard to retained placentas. Age of the broodmare may influence the occurrence of retained placentas.
Examination of the placenta The examination of the placenta, when passed, is essential to the determination of whether all of the membrane has been passed. Findings to be considered normal are twisting of the umbilical cord as long as the cord appears normal and not edematous; plaques of epithelial cells on the cord and on the amnion; small dilations of the urachus in the cord; velvet-like appearance to the entire attachment side of the chorioallantoic membrane; and small sacculations representing the previous endometrial cup structures. When a retained placenta is finally passed and has been shredded during its retention, it is difficult to state with certainty that all of the membrane has passed. Attempts to line up vessels within the membrane should be made, and observation of missing pieces of tissue should be noted before progressing to any form of therapy. This may not always be possible but should be attempted. The areas of chorionic tissue should always be examined to determine areas without endometrial attachment and biopsies taken if indicated.
Signs The most obvious sign of a mare with retained fetal membranes is the appearance of tissue protruding from the vulva. It may range from slightly visible to dragging on the ground. A retained placenta with a portion of it protruding for a relatively long period of time may make examination for completeness impossible. A retained placenta present with no placenta visible is usually due to a portion of the placenta remaining attached to the endometrium rather than the entire placenta being present within the uterus, although the latter is possible. Mares that normally pass the placenta during stage 3 of parturition show slight to mild colic due to uterine contractions.1 Mares with retained placentas infrequently demonstrate this activity and, when present, it is diminished. A retained placenta protruding from the vulva and touching the hocks should be tied up away from the hocks below the vulva (Figure 260.4).36 This procedure will reduce the probability that the mare will kick at the placenta and in doing so possibly injure the foal.1 The mare will also be less likely to step on the placenta, causing it
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Figure 260.4 Placenta tied in knot to avoid hocks.
to tear or causing the tip of the uterine horn to invaginate within itself or to evert through the vulva.
Complications of retained fetal membranes Sequelae of retained fetal membranes in the mare may vary from none to metritis, laminitis, septicemia, or death.37 Acute metritis and laminitis, as described in heavy breeds, are not as commonly seen in Thoroughbreds and other light breeds. Older literature suggests the longer the placenta is present the greater the risk of uterine infection and the greater the delay in uterine involution.38 A major concern regarding a retained placenta to most veterinarians and horse owners is that the mare may develop laminitis.39 It is reported that laminitis is more common in draft horses, and it occurs especially with a profuse uterine discharge. The suggested mechanism of action was that delayed uterine involution resulted in autolysis of the placenta within the uterus. This condition provided the environment in which bacteria could rapidly multiply, resulting in local inflammation and absorption of uterine contents that created a septicemia or toxemia. Although Streptococcus zooepidemicus is the predominant infective organism within the uterus at the time of a retained placenta,1 other Gram-negative bacteria capable of endotoxin and histamine production may be more important. Furthermore, laminitis has been attributed40 to degenerative changes in connective tissue within the foot and not to the typical toxic capillary injury and edema.
A study of 356 retained placentas in Standardbreds treated without manual removal reported no cases of laminitis, toxemia, septicemia, or acute metritis.12 Postparturient laminitis is considered to be the result of a systemic acute metritis with41 or without42 a retained placenta. Most veterinarians would agree that this metritis creates an abnormal condition within the uterus and consider it to be septicemic or toxemic if it involves the sensitive laminae of the hoof and possibly body temperature, disposition, or appetite. Occurrence of postpartum laminitis in association with a retained placenta is often either due to retention of many microvilli in large areas when they have broken free from their attachments to the remainder of the placenta, which can occur during manual removal, or due to separation of the apex of the placenta in the nongravid horn from the remainder of the placenta.1 These placental remnants serve as a focus of infection, and the uterus fills with fluid. The importance of placental examination following passage becomes obvious. Laminitis is due to the effects of septic uterine contents and possibly at least in part histamines being released. The actual mechanism producing laminitis is not certain. Two suggested possibilities involve a vasoactive component and a coagulation component.43 The vasoactive component may be related to hormones or toxins acting on digital vessels or shunts, thus altering blood circulation through the foot. It has also been suggested44 that peripheral blood circulation increases immediately postpartum due to decreased uterine size and this blood is involved in the vasoactive component. The coagulation component is related to intravascular coagulation,45 alterations in the intrinsic coagulation system, platelet numbers and fibrinogen degradation products,44 and the incidence of laminitis can be reduced with heparin administration.45 The laminitis observed is similar to that of gastrointestinal origin in that in 2–4 days the animal exhibits a foundered gait with the hind limbs placed well forward under the body to reduce some of the weight from the front legs. The hooves are warmer than normal to the touch and the digital artery pulse is increased. The animal may spend the majority of its time lying down or may refuse to lie down. Milk production decreases rapidly, and the foal becomes hungry and looks for food. Mares which have become septicemically ill due to a retained placenta may also have a uterine wall that is thinner than normal and friable.
Therapy considerations Some authors recommend treating any animal with a retained placenta to prevent laminitis. Recommended treatments have consisted of cold water
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Retained Fetal Membranes or ice to the hooves, suitable supportive footing, and uterine and systemic antibiotic therapy in order to prevent septic metritis. Antihistamines,46 phenylbutazone,23 nonsteroidal anti-inflammatory drugs,23 and tranquilizers have also been included in recommended therapy. Cyclooxygenase inhibitors such as flunixin meglumine can be administered in cases when the mare appears toxic. Administration is usually intravenously at a reduced dosage of 0.025 mg/kg t.i.d.21 Antihistamines,7 however, reportedly neither prevent nor cure laminitis after a retained placenta. In addition, the quantity of roughage being fed should be reduced and grains possibly eliminated from the diet.1 Although many authors recommend flushing the uterus with septic or toxic metritis or manually removing the placenta, this author is opposed to both practices. The less conservative treatments described in the literature rely on manual removal of the placenta and flushing the uterus in an attempt to reduce uterine debris. The disadvantage of this technique is that the treatment itself will increase absorption of septic or toxic material during removal and flushing and the mare’s condition will deteriorate before she can improve. This action may result in more damage to the feet or in death. Excellent results have been obtained with conservative therapy involving systemic therapy and supportive care plus antibiotic therapy of the uterus during the first or possibly first 2 days. The conservative approach is based on helping the mare to establish the normal uterine barrier, which develops by approximately day 3 postpartum. This assistance is accomplished primarily by systemic antibiotics, anti-inflammatory agents such as butezolidin or flunixin meglumine, and systemic fluids if necessary. This approach will result in improvement of the mare’s condition within 24–48 hours. Once this improvement has occurred, more localized treatment of the uterus can begin. Frequent temperature and complete blood count monitoring in mares treated in this manner will demonstrate slight subclinical relapses in progress after every uterine manipulation for 2–3 days. However, this result is more desirable than those with the less conservative therapies.
Treatments Treatments for retained fetal membranes vary with each possessing advantages and disadvantages. The most conservative treatment in this author’s opinion is the use of oxytocin. Several descriptive doses and routes of administration are available in the literature. Oxytocin can be administered in slow intravenous (i.v.) drip form by placing 30–60 units of oxytocin in 1–2 liters of saline and administering it over 1 hour,7 10–20 units in 1 liter of saline and giving
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it over a 1-hour period,47 or 80–100 units in 500 mL of saline and administering it over a 30-minute period.23 Oxytocin can also be administered i.v. or intramuscularly (i.m.) in a bolus form;2, 39, 47 dosages range from 10 to 120 units. Dosages can be repeated at 1.5- to 2-hour intervals until passage of the placenta or up to 24 hours postpartum. Disadvantage of the bolus form of administration includes intense and only spasmodic contractions when ‘large’ doses are administered, and were believed to be of no value.7 Furthermore, colic appeared to be more of a problem with bolus administration;25 cramping may have been related to dosage. This author has not encountered excessive colic with 20 units of oxytocin administered i.m. every 1–2 hours. Furthermore, colicky signs, if they occur, can be eliminated with sedation or analgesics.46 The dosage of oxytocin can be increased with additional time postpartum without colicky signs.26 Antibiotics have been used in the therapy of retained fetal membranes by most clinicians. One author suggested oxytetracycline hydrochloride in capsule form as the antibiotic of choice.48 In the author’s experience, use of powdered antibiotics (or antibiotics in capsule or tablet form) is considered undesirable because the antibiotic is unable to make contact with much of the endometrial surface.10 Other antibiotics utilized in the treatment of retained placentas have included sulfanilamide,46 penicillin,46 polymyxin, amikacin, ticarcillin,23 and others. It has been recommended that administration of antibiotics intrauterine should commence at 8 hours following delivery. Antibiotics are thought to be an important part of retained placenta treatment owing to their ability to control numbers of contaminating bacteria that enter the uterus at parturition or during attempts to manually remove the placenta. It has been suggested that the placental attachment should be examined each time before treatment. Mares treated intrauterine with antibacterial agents for placental retention reportedly have a significantly higher pregnancy rate after the breeding season than mares that had no intrauterine therapy for retained placenta.12 However, owing to a higher pregnancy loss, both groups had the same foaling percentage the following year. In a study of intrauterine combined with systemic antibiotic therapy versus only oxytocin therapy, 5 hours following treatment 55% of the intrauterine treated mares had not passed the placenta in comparison with 19% for those treated with oxytocin.12 Systemic administration of antibiotics is thought to be necessary by some authors whenever there is a retained placenta.47 Manual removal was the first reported treatment for retained fetal membranes.7 The literature describes five methods for manual removal of a
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placenta. The first consists simply of grasping the protruding free portion of the placenta and applying traction. This method does not require entry into the uterus if the placenta is exposed. Another method involves placing the hand between the chorion and the endometrium to separate them. A third method also involves placing the hand between the chorion and the endometrium; however, the allantochorionic membrane is massaged free from the endometrium.36 A fourth method involves grasping the allantochorionic membrane and twisting it into a tight cord.1, 46 The chorion is separated from the endometrium. The last method utilizes a wooden ring placed between the chorion and the endometrium such that the chorion separates as the ring is advanced. The time between parturition and recommended manual removal varies from immediately to 24 hours. Early removal could avoid a delay causing retarded uterine involution and a predisposition to uterine infection. However, in the same article, the author reports manual removal will delay uterine involution. Several visits to the mare at 4- to 12-hour intervals may be necessary to manually remove a placenta without injury to the mare.36, 47, 49 It was suggested not to work longer than 10 minutes on its removal at each visit.46 Although reported by many authors as the treatment of choice, manual removal of a placenta has many undesirable complications occurring after treatment. The first complication of manual removal of the placenta is severe hemorrhage.7 It is possible for the mare to lose large amounts of blood into the uterus, and blood serves as an excellent environment for bacterial growth. Furthermore, pulmonary emboli have been reported following this procedure, apparently because of dislodgment of naturally occurring emboli from the uterine vein.7 Invagination or intussusception of the uterine horn attached to the placenta can occur at the time of manual removal. Another reproductive complication is a delay in uterine involution.7 A reported ‘vastly’ increased amount of fluid accumulated within the uterus after manual removal of the placenta.46 The reason given for this increase was that during manual removal only central branches of the chorionic villi were removed and all microvilli were broken off and retained within the endometrium. In addition, rupture of the endometrial and subendometrial capillaries may occur, adding to the fluid within the lumen of the uterus. Microvilli will need to be released and then flushed by natural uterine secretions from their crypts. These microvilli were related to increased occurrence of tissue debris, endometritis, laminitis, uterine spasm, and delayed involution.46 One report acknowledged that manual removal should be discouraged owing to the induced trauma, hemorrhage, and infection.48 However, it was the
recommended treatment after other methods had failed. Although it has been recommended to never cut off the membranes such that retraction of the placenta into the uterus would occur, it is equally as bad to attach weights to the placenta to increase the traction to speed removal. This, as is with manual removal, increases the number of microvilli left within the uterus and would necessitate liquefaction and passage as microscopic particles rather than as part of the placenta. It has been suggested that manual removal of the placenta can result in permanent endometrial damage.23 The author has observed that damage has occurred in a number of mares from 3 to 14 years of age which had normal fertility until manual removal of a placenta. Following manual removal, this author has found a decrease in fertility and endometrial biopsies revealed classifications of categories IIB and III based upon presence of fibrosis. The cervix also remains open longer when the placenta is manually removed.7 It has been suggested that manual removal of the placenta has been continued from a time when draft horses predominated and retained placentas were more of a problem46 and oxytocin was not utilized. Antiseptics placed into the uterus have been suggested as another method of therapy. Betadine in 20–30 liters of warm water has been placed into the uterus and inside the placenta.2 Another method describes the placement of 10–12 liters of povidine iodine into the allantochorionic space followed by closure of the hole in this membrane (Figure 260.5a), trapping the fluid within the allantoic cavity (Figure 260.5b).37 This procedure aids expulsion by stretching the uterus and causing endogenous oxytocin release and separation of microvilli from their attachments. The placenta should be passed within 30 minutes. The exact effect of this procedure on microvilli separation has not been reported. One disadvantage of
(a)
(b) Figure 260.5 (a) Placement of fluid inside chorioallantoic membrane. (b) Fluid-filled placenta to aid in expulsion.
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Retained Fetal Membranes antiseptic use was that it could depress phagocytosis. The importance of this effect is unknown.6 Flushing and siphoning46 of fluid from the uterus have been reportedly successful in treatment of retained placentas.36 It has also been stated that douching the uterus is of questionable value.1 Infusing the uterus with 42◦ C water has been used with apparently beneficial results.14 Flushing the uterus with large volumes of saline containing antibiotics reportedly prevented or cured laminitis.2 Much diversity with regard to uterine therapy and its importance obviously exists. Tetanus protection in the form of antitoxin,46 or preferably toxoid, is recommended. Placement of Caslick’s sutures or clips in the dorsal vulva after foaling have been recommended to reduce uterine contamination.1 Umbilical artery infusion of collagenase has been utilized to treat retained placentas when they have been retained for longer than 6 hours.21 The technique requires about 10 minutes to execute and at least two of the three vessels are perfused.21 There were no reported adverse side-effects of this treatment.21 If perfusion is not complete, segments of the retained placenta may remain retained after the larger portion has passed.21 The importance of intrauterine therapy or the need to remove the placenta rapidly is not established. It is apparent that systemic oxytocin treatment for uncomplicated retained placenta is superior to all others in the non-draft breed. A report50 on a heavy hunter mare that was treated solely with systemic antibiotics after a cesarean passed the placenta 5 days following parturition with no adverse side-effects. The animal was not treated intrauterine owing to her disposition. A second mare was first examined 5 days following foaling.51 She had passed the placenta that day and had received no prior treatment. The mare’s temperature was normal, there was no laminitis, and she was eating and drinking. The only reason the owners realized a problem existed was the passage of the foul-smelling placenta. This author has treated one mare following an abortion at 9 months with intrauterine oxytetracycline for 3 days and then 2% Lugol’s solution every other day until placental passage. She received phenylbutazone for the first 4 days. The placenta was passed 13 days following the abortion. The uterus was cultured at 34 days and contained no bacteria. She was bred at approximately 60 days post abortion and conceived, maintaining the pregnancy and delivering a healthy foal. This case is the longest the author has had a mare retain a placenta. It is important to provide adequate examination and treatment of the mare with a retained placenta. It is also important to realize manual removal of the placenta is not the treatment of choice.
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Subsequent examination Subsequent follow-up examination of mares with retained placentas should be mandatory regardless of treatment. It is important to substantiate that the uterus is normal whether or not the mare is to be bred again that year. Rectogenital examination, uterine culture, endometrial biopsy, and ultrasonographic examination are recommended techniques that can be used in this evaluation. Endometrial cytology and endoscopic examination can also be used. The author’s recommendation is to wait approximately 25 days following placental passage before breeding. However, late in the breeding season mares have conceived and maintained pregnancies when bred at 10–20 days postpartum. The latter is recommended only at the end of the breeding season when no other option is available.
Influence of retained placenta on future reproduction No difference in pregnancy rates after first breeding, pregnancy rates after the breeding season, pregnancy loss rates, and foaling rates between normal foaling mares and mares with retained placentas was reported when treatment was conservative.12 This lack of a difference was evident regardless of the duration of time the placenta was retained. No manual removal was used in this study.
Summary A three-paragraph correspondence by Kelly52 in 1944 probably described treatment of the retained placenta as well as anyone. ‘Placentas which cannot be removed “easily” should not be removed; give “appropriate” medication and wait’. Recommendations regarding retained placentas include initiation of therapy at 3 hours postpartum. This therapy is accomplished most conveniently by dispensing oxytocin to the owner with written instructions for administration of 20 units of oxytocin i.m. every 1–2 hours to mares not passing the placenta within 3 hours of parturition. This action permits commencement of treatment in the middle of the night that otherwise may not occur. The low dose administered i.m. has not been associated with severe colicky signs after 2–3 hours postpartum. An examination of the mare within 12–18 hours of foaling is imperative to distinguish an uncomplicated retained placenta from one that could be more severe. This examination consists of rectogenital palpation and/or ultrasonographic examination and is performed regardless of placental passage. Decreased uterine tone
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is an excellent indicator of potential complications. No vaginal examination is performed nor treatment administered if the placenta has passed and the uterus has good to excellent tone at the time of the examination. If the placenta is present and the uterus has good to excellent tone, the uterus is infused with 5–8 g of oxytetracycline in 500 mL of saline. The author is uncertain whether this is necessary. If the uterine tone is diminished, the volume of infusion is increased. In lightweight breeds, no other treatment is given. In draft breeds or in any mare following a severe dystocia, animals are administered systemic penicillin with or without gentomicin and flunixin meglamine. Mares that have experienced severe dystocias appear to be at most risk for developing complications to retained placenta or to the dystocia itself. Systemic treatment is continued for 3–7 days depending on the animal’s condition. Uterine infusions are switched from oxytetracycline to 2% Lugol’s solution on days 2–4 to enhance the contractibility of the uterus. The volume of Lugol’s is determined by the size of the uterus and may vary from 500 mL to 1 liter. The important factor to remember regarding Lugol’s administration is not to overfill the uterus. Lugol’s solutions 2–10% have been used by the author to treat uterine infections and inflammation as well as retained placentas without any adverse effects on the endometrium as determined by endometrial biopsy and endoscopy. Lugol’s solution, however, placed into the vagina may cause straining and the development of fibrous connective tissue bands across the vaginal lumen. Mares with a history of retained fetal membranes should be treated with oxytocin immediately after foaling, and treated again as described previously. Mares over 15 years old may be candidates for postpartum treatment with oxytocin and, although the author does not routinely do this, it is a consideration and is used selectively. Intrauterine medication may prove to be unnecessary or detrimental in uncomplicated cases of retained placentas. Although a concern, the importance of laminitis and the effect of retained placenta on future reproduction in lighter breeds does not appear to be a major problem if treatment is initiated early.
References 1. Roberts SJ. Veterinary Obstetrics and Genital Diseases, 3rd edn. Woodstock: David and Charles Inc., 1986; pp. 382–3. 2. White TE. Retained placenta. Mod Vet Pract 1980;61:87–88. 3. Shipley WD, Bergen WC. Care of the foaling mare and foal. Vet Med 1969;64:63–70.
4. Sager FC. Examination and care of the genital tract of the brood mare. J Am Vet Med Assoc 1949;115:450–5. 5. Wright JG. Parturition in the mare. J Comp Pathol 1943;53:212–19. 6. Asbury AC. Management of the foaling mare. In: Proceedings of the 18th Annual Convention of the American Association of Equine Practitioners, 1972, pp. 487–90. 7. Vandeplassche M, Spincemaille J, Bouters R. Aetiology, pathogenesis and treatment of retained placenta in the mare. Equine Vet J 1971;3:144–7. 8. Jennings WE. Some common problems in horse breeding. Cornell Vet 1941;31:197–215. 9. Williams WL. The Diseases of the Genital Organs of Domestic Animals, 3rd edn. Ithaca: WL Williams, 1943; pp. 552–5. 10. Blanchard TL, Bierschwal CJ, Youngquist RS, Elmore RG. Sequelae to percutaneous fetotomy in the mare. J Am Vet Med Assoc 1983;182:1127. 11. Vandeplassche M, Bouters R, Spincemaille J, Bonte P. Dropsy of the fetal sacs in mares: induced and spontaneous abortion. Vet Rec 1976;99:67–9. 12. Provencher R, Threlfall WR, Murdick PW, Wearly WK. Retained fetal membranes in the mare: a retrospective study. Can Vet J 1988;29:903–10. 13. Jennings WE. Twelve years of horse breeding in the army. J Am Vet Med Assoc 1950;116:11–16. 14. Sager FC. Management and medical treatment of uterine disease. J Am Vet Med Assoc 1968;153:1567–9. 15. Vandeplassche M, Spincemaille J, Bouters R, Bonte P. Some aspects of equine obstetrics. Equine Vet J 1972;4:105– 9. 16. Steven DH, Jeffcott LB, Mallon KA. Ultrastructural studies of the equine uterus and placenta following parturition. J Reprod Fertil Suppl 1979;27:579–86. 17. Allen WE, Chard T, Forsling ML. Peripheral plasma levels of oxytocin and vasopressin in the mare during parturition. J Endocrinol 1973;57:175–6. 18. Van Der Kaay FC. Cursus Nota’s. Utrecht: Faculteit Diergeneeskunde, 1945; p. 37. 19. Alhnelt E. Tiergeburtshilte, 2nd edn. Berlin: P. Pary, 1960; p. 638. 20. Derivauz J. Obstetrique Veterinaire. Paris: Vigot Freres, 1957; p. 333. 21. Haffner JC, Fecteau KA, Held JP, Eiler H. Equine retained placenta: technique for and tolerance to umbilical artery injections of collagenase. Theriogenology 1998;49: 711–16. 22. Arthur GH. Wright’s Veterinary Obstetrics, 3rd edn. Baltimore: Williams and Wilkins Co., 1964; pp. 341–4. 23. Held JP. Retained placenta. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia: W.B. Saunders Co. 1987; pp. 547–50. 24. Berthelon M, Tournut J. Retention of the placenta in domestic animals. Rev Med Vet 1953;104:529–38. 25. Cox JE. Excessive retainment of the placenta in a mare. Vet Rec 1971;89:252–3. 26. Hillman RB. Induction of parturition in mares. J Reprod Fertil Suppl 1975;23:641–4. 27. Seamans KW, Harms PG, Atkins DT, Fleeger JL. Serum levels of progesterone, 5 alpha-dihydroprogesterone and hydroxy-5 alpha prenanones in the prepartum and postpartum equine. Steroids 1979;33:55–63. 28. Stewart DR, Stabenfeldt GH, Hughes JP. Relaxin activity in foaling mares. J Reprod Fertil Suppl 198232:603–9.
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Retained Fetal Membranes 29. Horta AEM, Chassagne M, Brochart M. Prostaglandin F2 alpha and prostacyclin imbalance in cows with placental retention. Ann Rech Vet 1986;17:395–400. 30. Hudson NPH, Prince DR, Mayhew IG, Watson ED. Investigation and management of a cluster of cases of equine retained fetal membranes in Highland ponies. Vet Rec 2005;157:85–9. 31. Blakley BR, Bell RJ. The vitamin A and vitamin E status of horses raised in Alberta and Saskatchewan. Can Vet J 1994;35:297–300. 32. Brendemuehl JP. Reproductive aspects of fescue toxicosis. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 4th edn. Philadelphia: W.B. Saunders, 1997; pp. 571–3. 33. Hintz HF. Nutrition of the broodmare. In: Equine Reproduction. Eds. A.O. McKinnon and Voss. Philadelphia: Lea & Febiger, 1993; pp. 181–5. 34. Sevinga M, Barkema HW, Stryhn H, Hesselink JW. Retained placenta in Fresian horses. J Anim Sci 2004;82: 982–6. 35. Sevinga M, Vrijenhoek T, Hesselink JW, Barkema HW, Groen AF. Effect of inbreeding on the incidence of retained placenta in Friesian horses. J Anim Sci 2004;82:982–6. 36. Allen WE. Fertility and Obstetrics in the Horse. Boston: Blackwell Scientific, 1988; pp. 104–8. 37. Burns SJ, Judge NG, Martin JE, Adams LG. Management of retained placenta in mares. In: Proceedings of the 23rd Annual Convention of the American Association of Equine Practitioners, 1977, pp. 381–90. 38. Rossdale PD, Ricketts SW. Equine Stud Farm Medicine, 2nd edn. Philadelphia: Lea & Febiger, 1980; pp. 249–50. 39. Hafez ESE. Reproduction in Farm Animals, 5th edn. Philadelphia: Lea & Febiger, 1987; pp. 418–20. 40. Obel N. Contribution to our knowledge of the aetiology and pathogenesis of the foaling laminitis. Scand Vet Tidskrift 1943;33:41–6.
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41. Adams OR. Lameness in Horses, 3rd edn. Philadelphia: Lea & Febiger, 1974; p. 248. 42. Threlfall WR, Carleton CL. Treatment of uterine infections in the mare. In: Current Therapy in Theriogenology, 2nd edn Ed. D. Marrow.. Philadelphia: W.B. Saunders Co, 1986; pp. 730–7. 43. Hood DM, Stephens KA. Physiology of equine laminitis. Comp Cont Educ 1981;3:454–60. 44. Hood DM, et al. Update on laminitis: physiopathology and treatment. Paper presented at the Annual Meeting of the American Veterinary Medicine Association, 19–24 July; 1980. Washington DC. 45. Hood DM, Amos MS, Grosenbaugh DA. Equine laminitis III: coagulation dysfunction in the developmental and acute disease. J Equine Med Surg 1979;3:355–61. 46. Arthur GH, Noakes DE, Pearson H. Veterinary Reproduction and Obstetrics, 5th edn. London: Bailliere Tindall, 1982; pp. 250–1. 47. Arthur GH, Noakes DE, Pearson H, Parkinson TJ. Retention of the fetal membranes. In: Arthur GH, Noakes DE, Pearson H, Parkinson TJ (eds Veterinary Reproduction and Obstetrics, 7th edn. London: W.B. Saunders, 1996; pp. 291– 301. 48. Neely DP, Liu IKM, Hillman RB. Equine Reproduction. Nutley, NJ: Veterinary Learning Systems, 1983; pp. 87–8. 49. Jackson PGG. Handbook of Veterinary Obstetrics. London: W.B. Saunders, 1995; pp. 181–5. 50. Mason TA. Retention of the placenta in the mare. Vet Rec 1971;89:546. 51. Alexander RW. Excessive retainment of the placenta in the mare. Vet Rec 1971;89:175–6. 52. Kelly EV. Retained placenta in the mare. Vet Rec 1944; 56:296.
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Chapter 261
Postpartum Metritis Terry L. Blanchard
Introduction The incidence of postpartum metritis – i.e., infection of the uterus within 7–10 days postpartum, sometimes involving the endometrium, myometrium, and perimetrium – in foaling mares is low, but increases when birthing trauma and/or retained placenta occurs. Mares that experience an apparently normal parturition can also develop postpartum metritis. Sequelae of postpartum metritis vary from delay in uterine involution to development of septicemia/ toxemia, laminitis, and death.1–6 Prompt recognition and treatment of the disorder can prevent sequelae from occurring and more rapidly restore the affected mare to a fertile condition.
Occurrence and clinical signs Septic/toxic metritis may occur with or without birthing trauma and/or retained placenta, and becomes apparent within 1–10 days (most commonly on days 2–4) postpartum. The etiology of septic/toxic metritis is thought to be similar to that of retention of the fetal membranes, being associated with uterine atony or inertia.3 Trauma to the uterus, autolysis of placental remnants, and excessive lochial accumulation likely contribute to rapid growth of bacteria and production of toxins, which may be absorbed into the bloodstream, particularly when expulsion of contents is delayed or when the normally intact uterine mucosal barrier is damaged (Figures 261.1 and 261.2). The pathogenesis of toxic metritis, and particularly the time sequence of development of toxemia, remains unproven. However, infusion of Escherichia coli endotoxin into the uterus of normal foaling pony mares on days 1 and 4 postpartum failed to result in detectable presence of endotoxin within the blood, and neutropenia did not occur.7 In that study, scanning electron micrographs revealed the endometrial mucosa to be intact. This is in contrast to the mare with septic/toxic metritis, in which the mucosal lin-
ing is disrupted.8 Apart from development of septic/ toxic metritis and/or laminitis, we do not know when the uterus is damaged, thus allowing absorption of locally produced toxins. Certainly, disruption of the endometrial mucosal barrier can occur during dystocia, or could gradually progress after retention of fetal membranes leading to bacterial overgrowth. Septic/toxic metritis with neutropenia is more common in mares that retain their placenta, particularly after dystocia.5 When intraperitoneal injections of E. coli endotoxin are given, pronounced neutropenia occurs rapidly (i.e., within 1–2 hours usually). Pronounced neutropenia with degenerative left shift usually appears in mares developing septic/toxic metritis on days 2–4 postpartum5, 8, 9 – perhaps due to gradual build-up of endotoxin-producing bacteria within the uterine lumen concurrent with progressive damage to the endometrial mucosa, thus allowing ever more absorption of endotoxin. If the mare did not have a difficult birth and management has not been alerted to the possibility of retained fetal membranes (e.g., incomplete or shredded placenta passed, fetal membranes visibly protruding from the vulva), the first clinical signs noted usually are inappetance, depression, and fever, although in some cases the veterinarian is first requested to examine the mare due to the presence of lameness.3, 6 The foal may be noted to suckle often, yet not be satisfied, due to decreased milk production by the dam. The mare’s heart and respiratory rates are often elevated when fever is present. Mucous membranes may be tacky (e.g., dehydration) and, if toxemia exists, the membranes may be discolored with poor perfusion (i.e., prolonged capillary refill time). A copious, fetid vulvar discharge is often evident, particularly in cases with trauma and necrosis of the birth canal (Figure 261.3). Repeated abdominal straining sometimes occurs when vaginal necrosis or lacerations are present, or if placental remnants are lying in the birth canal. The mare’s feet should always be examined
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Figure 261.1 Scanning electron micrograph of endometrium of a mare 1 day after parturition. The epithelium is intact, and the surfaces of individual prociliated and secretory cells are visible.
for warmth, presence of increased digital pulses, and pain. If laminitis develops, lameness (particularly of the front feet) can become pronounced. Sinking of the digit(s) and rotation of the third phalanx(ges) may follow, necessitating euthanasia. Examination of the uterus per rectum and by transrectal/transabdominal ultrasound commonly reveals an enlarged atonic uterus containing an excessive amount of variably echogenic fluid. Aseptic manual examination of the birth canal is performed to determine if any accompanying swelling, hemorrhage or necrotic tissue is present in the birth canal, suggesting birth trauma (Figure 261.4). Digital examination of the uterine lumen may reveal placental remnant(s), including a retained tip of the placenta that previously occupied a uterine horn (Figure 261.5).
Additional diagnostic testing If sepsis or toxemia has developed, a complete blood count will demonstrate neutropenia with a degenerative left shift, with neutrophils sometimes having
Figure 261.2 Scanning electron micrograph depicting mostly denuded uterine mucosal surface of mare euthanized on day 4 postpartum due to severe septic/toxic metritis and laminitis (with rotated third phalanges).
Figure 261.3 Examining the character of effluent obtained by lavaging the uterus of the postpartum mare with metritis will reveal varying degrees of bloody discoloration, purulent material, and placental remnants.
a toxic appearance (hypersegmented nuclei, cytoplasmic vacuolation).5, 10 Neutropenia is most pronounced 3–4 days postpartum (Figure 261.6). Absolute neutropenia (i.e., often < 1500 neutrophils/µL), is a common finding in mares with septic/toxic metritis. Although the relationship between bacterial endotoxemia and onset of laminitis is currently questioned, signs of endotoxemia have been linked to significant risk of laminitis developing.10
Figure 261.4 Vulvar tearing, bruising, and swelling in a postpartum mare.
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Mare: Problems of Pregnancy
Figure 261.5 Tip (end of portion occupying uterine horn) of placenta sometimes remaining in the uterus of postpartum mare after majority of fetal membranes have been expelled.
Swabbing the uterine lining and lochia for bacterial culture can be done to identify the organism(s) present, and to guide selection of antimicrobial(s) used in treatment of infection. Bacteria typically involved in postpartum metritis include aerophilic and microaerophilic or anaerobic organisms. Commonly isolated bacteria include E. coli (a potent endotoxin13 12
Figure 261.7 Lateral radiograph of forefoot of a mare with postpartum laminitis. Rotation and sinking of P3 is apparent. The toe has been elevated with a pad and shoe and the foot is in a cast to restrict movement.
producer) and Streptococcus spp., and sometimes Bacteroides fragilis and clostridial species.3, 5 When foot soreness develops, radiographs should be procured to evaluate for sinking or rotation of the digit (Figure 261.7). When sinking of the digit is suspected (i.e., ledging is evident at the coronary band), contrast material can be infused into the vasculature to evaluate the circulatory status of the foot (Figure 261.8). Once vascular disruption has occurred, the prognosis for survival is grave.
11 Neutrophil Count (1×103/mm3 blood)
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1
2
3
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5
6
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Figure 261.6 Mean neutrophil counts in complete blood count (CBC) of normal foaling mares and mares experiencing dystocia. Absolute neutropenia, often with less than 1500 neutrophils/µL, is common in mares developing postpartum septic/toxic metritis.
Various treatments for postpartum metritis in mares have been advocated. Systemic administration of broad-spectrum antibiotics is indicated to control infection. Antibiotics selected should be effective against both Gram-positive and Gram-negative bacteria. Administration of gentamicin (6.6 mg/kg, q. 24 h, i.v.) and penicillin (22 000–44 000 IU Na+ or K+ penicillin, q. 6 h, i.v. or i.m.; or 22 000 IU procaine penicillin q. 12 h, i.m.) can be used for this purpose. If endotoxemia is considered a significant risk, administration of polymixin-B (1000–6000 U/kg q. 6–8 h, slowly i.v.) may be added to the systemic antimicrobial regimen.11 Pentoxiphylline (7.5–10 mg/kg, q. 8–12 h p.o. or i.v.) may provide added benefit to improve malleability of red blood cells and thus potentially improve circulation to vasculature of the foot.11 For additional protection against endotoxemia, and for anti-inflammatory effects, administration of flunixin meglumine (0.25 mg/kg, q. 8 h, i.v.) is recommended. Specific treatment of the genital tract includes oxytocin therapy (10–20 IU oxytocin q. 6 h, i.m.), stimulating uterine contractions (lasting
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stomach pump) in 3- to 6-L flushes is performed until the effluent is relatively clear. If necessary, uterine lavage can be repeated on successive days until the initial effluent is free of purulent material and tissue debris. Acute laminitis is a medical emergency, and treatment should commence as soon as possible. Many therapeutic regimens have been recommended for the treatment of laminitis. Current equine medicine textbooks should be referred to for discussion of various treatments for laminitis. Certainly, padding of the foot (frog), administration of anti-inflammatory/analgesic medications, and restriction of movement are indicated as part of the treatment regime.
References
Figure 261.8 Contrast radiography was utilized to obtain an image of the digital vessels of the foot of a postpartum mare with laminitis. Sinking of the digit is apparent with vascular tearing and compression – carrying a poor prognosis for survival.
20–50 minutes) that aid in expulsion of uterine contents. Uterine lavage is performed to remove debris and bacteria from the uterus, thereby reducing contamination and creating a less favorable environment for bacterial growth. Purulent material and cellular debris that bind to and inactivate many antibiotics are also removed by uterine lavage. Whether there is additional benefit in administering local antibiotics by uterine infusion is controversial, but the author infuses a broad-spectrum antibiotic (3 g oxytetracycline) into the uterine lumen following lavage, and once daily thereafter until uterine infection is deemed to be under control. There may be some benefit to controlled exercise if foot soreness is not present. If foot soreness is present, exercise is contraindicated. For uterine lavage in postpartum mares, the author prefers to utilize a large-bore tube (e.g., stomach tube). After disinfection of the perineal and vulvar areas, a sterilized or disinfected nasogastric tube is passed into the uterus. The hand is cupped around the end of the tube to prevent the uterus and any remaining placenta from being siphoned into its end. Gentle lavage with warm (40–42◦ C), sterile, physiological saline (administered via a sterile or disinfected
1. Rossdale PD, Ricketts SW. Equine Stud Farm Medicine, 2nd edn. Philadelphia: Lea & Febiger, 1980; pp. 260–76. 2. Vandeplassche M, Bouters R, Spincemaille J, Bonte P, Coryn M. Observations on involution and puerperal endometritis in the mare. Irish Vet J 1983;37:126–32. 3. Roberts SJ. Veterinary Obstetrics and Genital Diseases (Theriogenology), 3rd edn. Woodstock, VT: Roberts SJ, 1986; pp. 251–390. 4. Blanchard TL, Bierschwal CJ, Youngquist RS, Elmore RG. Sequelae to percutaneous fetotomy in the mare. J Am Vet Med Assoc 1983;182:1127. 5. Blanchard TL, Elmore RG, Varner DD, Garcia MG, Orsini J, Youngquist RS. Dystocia, toxic metritis and laminitis in mares. In: Proceedings of the 33rd Annual Convention of the American Association of Equine Practitioners, 1987, pp. 641–8. 6. Blanchard TL, Varner DD, Scrutchfield WL, Bretzlaff KN, Taylor TS, Martin MT, Elmore RG. Management of dystocia in mares: retained placenta, metritis, and laminitis. Comp Cont Educ Pract Vet 1990;12:563–71. 7. Blanchard TL, Elmore RG, Kinden DA, Berg JN, Mollett TA, Garcia MG. Effect of intrauterine infusion of E. coli endotoxin in postpartum pony mares. Am J Vet Res 1985;46:2157–62. 8. Blanchard TL, Vaala WE, Straughn AJ, Acland HM, Kenney RM. Septic/toxic metritis and laminitis in a postparturient mare. J Equine Vet Sci 1987;7:32–4. 9. Blanchard TL, Orsini JA, Garcia MC, Elmore RG, Youngquist RS, Bierschwal CJ. Influence of dystocia and retained placenta on white blood cell and neutrophil counts in mares. Theriogenology 1986;25:347–52. 10. Parsons CS, Orsini JA, Krafty R, Capewell L, Boston R. Risk factors for development of acute laminitis in horses during hospitalization: 73 cases (1997–2004). J Am Vet Med Assoc 2007;230:885–9. 11. Hackett ES, Orsini JA, Divers TJ. Equine emergency drugs: approximate dosages and adverse reactions. In: Orsini JA, Divers TJ (eds) Equine Emergencies Treatment and Procedures. St Louis: Saunders Elsevier, 2008; pp. 739–52.
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Section (j): Female Urogenital Surgery
262.
Pneumovagina Etta A. Bradecamp
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Surgery of the Caudal Reproductive Tract Angus O. McKinnon and Sarah L. Jalim
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Cervical Surgery Patrick J. Pollock and Thomas M. Russell
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Ovariectomy Dwayne H. Rodgerson and Dawn A. Loesch
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Hysterectomy Anna K. Roetting and David E. Freeman
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Rectal Tears David E. Freeman
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Chapter 262
Pneumovagina Etta A. Bradecamp
In the normal mare there are three natural barriers against the introduction of foreign material into the uterus: the vulvar seal at the lips of the vulva; the vestibulovaginal fold; and the cervix. When one or more of these barriers ceases to function, the risk of introduction of debris, air, bacteria, and potential infection into the uterus increases. The aspiration of air into the uterus, even in the absence of bacteria, can elicit an inflammatory response. Pneumovagina, the presence of air in the vagina, occurs as a result of the dysfunction of the vulvar seal and of the vestibulovaginal fold. Every mare presented for breeding should be evaluated for pneumovagina and should have this condition corrected prior to breeding to maximize her chances of becoming pregnant and carrying a foal to term.
Pneumovagina Mares become predisposed to developing pneumovagina when the conformation of the vulvar lips prevents the lips from forming a seal; or when the mare experiences trauma during foaling such as tearing of the vulva or vestibulovaginal fold or formation of a perineal fistula or laceration. Some mares, especially older, thinner ones, have poor perineal conformation with a sunken anus and anterior sloping of the vulvar lips. This type of conformation increases the chances that the mare will aspirate air and fecal debris into the cranial aspects of the reproductive tract. A diagnosis of pneumovagina may be made in several different ways. The most common way to assess whether a mare has the propensity to aspirate air into her uterus is by parting the lips of the vulva and listening for the sound of air being aspirated into the vagina (Figure 262.1). If the vestibulovaginal fold maintains its integrity the aspiration of air should not occur. Pneumovagina can be diagnosed during transrectal ultrasonographic examination of the repro-
ductive tract with the observation of air in the uterus or vagina. One may also detect pneumovagina when the mare postures to urinate or ambulates and air can be heard being aspirated into the vagina. Pneumovagina can also be created when the mare exercises; this is not uncommon in racing fillies and many of them are Caslicked for this reason.1, 2 Air causes an inflammatory response in the uterus, and the presence of eosinophils on uterine cytology has been associated with aspiration of air into the uterus.3 When air is aspirated into the cranial vagina and uterus, debris and bacteria are also allowed access to the cranial reproductive tract. During estrus the cervix is relaxed, thereby leaving the uterus vulnerable to the ascent of bacteria from the perineal region to the cranial reproductive tract if pneumovagina develops. This breakdown in their natural mechanical protective barriers makes the reproductive tract more susceptible to the development of chronic inflammation and infection.
Caslick index During a reproductive or breeding soundness exam it is important to assess the mare’s perineal conformation and the integrity of the vulvar lips, vestibulovaginal fold and the cervix. One method of quantifying the degree of variation from normalcy of the perineal conformation is by calculating the Caslick index. In 1937, Dr E.A. Caslick first reported on the importance of perineal conformation, how it influenced reproductive health and fertility, and the relationship between it and infections of the reproductive tract.4 Caslick discussed the importance of the anterior slope of the vulva, and he considered a vulvar angle of 80 degrees to the horizontal to be ideal while a slope of less than 50 degrees could be correlated with the development of pneumovagina. In 1979, Pascoe presented research that described a simple measuring device to determine the slope (angle
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Female Urogenital Surgery measured parameters produced a value he termed the Caslick index.5 This index is calculated by determining the degree of anterior slope of the vulva and the distance in centimeters that the dorsal commissure of the vulva is displaced dorsally in relationship to the pelvic brim. These two measurements are then multiplied to yield the Caslick index. In the normal mare this value should be less than 100. If the value is greater than 150, a Caslick’s operation is indicated in these mares. Mares who fall between 100 and 150 should be further assessed on a case by case basis to determine if a Caslick’s operation is indicated. The Caslick index can be helpful in identifying those mares that may need a Caslick’s operation but do not demonstrate the classical signs of pneumovagina. Caslick index = (effective length of the vulva in cm) ×(angle of declination in degrees)
(a)
(b) Figure 262.1 Parting of the vulvar lips to test for pneumovagina. In the normal mare (a) the vestibulovaginal sphincter should remain closed and air should not be aspirated into the vagina. (b) Loss of integrity of the vestibulovaginal sphincter. Reprinted with permission from McKinnon et al.7
of declination) of the vulva and the length of the vulvar lips. In a study of 9020 mares he demonstrated how a simple mathematical formula using the two
where effective length is the distance between the dorsal commissure and the pelvic floor. A mare’s Caslick index can change as she ages, gains or loses weight, and has foals. Once a mare has had a Caslick’s operation performed to prevent pneumovagina she should remain Caslicked for life. Subsequently, Pascoe discussed other conformational characteristics that predispose a mare to developing pneumovagina.6 In a normal mare, more than 80% of the length of the vulva, measured from the dorsal to ventral commissures, should be located below the level of the ischial tuberosities. When more than 20% of the vulvar opening is above the brim of the pelvis, there is an increased risk that the dorsal commissure will be pulled anteriorly toward the anus and precipitate the development of pneumovagina. Another indicator that a mare should have a Caslick’s operation performed is if the dorsal commissure of the vulva is greater than 4 cm (11/2 in) dorsal to the pelvic floor.6 A long sloping hip and a sacro-iliac joint located dorsal to the tail setting also correlate with decreased incidence of pneumovagina. Mares that have a flat-topped croup with a tail setting that is level with the sacro-iliac joint and sunken anus are predisposed to experiencing bouts of infertility associated with conditions such as pneumovagina, urovagina, chronic endometritis and accumulation of intrauterine fluid.6 In addition to the Caslick index, there are other parameters that should be evaluated when examining a mare for reproductive soundness. The competency of the seal of the vulvar lips is paramount in preventing the aspiration of air, debris, and bacteria into the
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Pneumovagina cranial aspects of the reproductive tract. There are a number of factors that can result in a loss of the integrity of this seal. In the normal mare the lips of the vulva are shaped to meet very precisely thereby forming a natural seal. In the occasional mare the lips do not meet in such a precise manner. This defect can even be seen in the young maiden mare where one lip overlaps the other or folds in slightly, resulting in a loss of that perfect seal. While this may not be detrimental to the mare’s reproductive health, it should not be ignored and these mares should be further examined for signs of pneumovagina and inflammation or infection of the cranial reproductive tract, including excessive or inflammatory uterine edema, a positive uterine culture and/or cytology, or hyperemic vaginal mucosa on a vaginal speculum exam.
Caslick vulvoplasty Once a diagnosis of pneumovagina has been made, the condition should be corrected to prevent the continued aspiration of air into the vagina and cranial reproductive tract. Beyond recognizing the relationship between the conformation of a mare’s reproductive tract and the development of genital infections and subsequent infertility, Caslick also described the procedure of suturing the dorsal portion of the vulvar lips to prevent aspiration of air into the reproductive tract.4 Caslick had observed that many mares that had repeated manual entry of the vagina for uterine irrigation for metritis had temporary worsening of their metritis. When the uterine irrigation was discontinued and the dorsal aspects of the vulva closed a noticeable clinical improvement was seen.4 This procedure was shown to significantly improve the fertility of mares.
Technique The mare is adequately restrained either in stocks or twitched and stood inside the stall door. The occasional mare may require sedation. The tail should be wrapped and held or tied to the side. The perineum should be cleaned with a mild soap or surgical scrub and water. The vulvar margins are infiltrated with approximately 20 mL of 2% lidocaine or mepivacaine using a 20-gauge or smaller needle, distending the labial tissues along the vulvar margins (Figure 262.2a). A 6–8-mm strip of mucosa is removed from the labial margin, taking care not to remove skin (Figure 262.2b). It may not be necessary to remove a
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strip of mucosa and instead a scalpel blade or pair of sharp pointed scissors can be used to open a line along the labial margin. This method works especially well if this is the first time the Caslick has been performed, for example in maiden mares, or if there is minimal scar tissue present. The goal is to remove the minimal amount of mucosal tissue while at the same time creating an approximately 6-mm- wide strip of fresh raw tissue to appose. If the fresh edges are too narrow, fistulas or gaps may occur during the healing process. If too much tissue is removed, excessive fibrosis develops and it will be more difficult to get a good apposition of tissues when the procedure needs to be performed in subsequent breeding seasons. The fresh edges are apposed using a simple-continuous, interrupted or continuous interlocking suture pattern (Figure 262.2c). While some veterinarians prefer using a non-absorbable monofilament such as nylon or polypropylene,6 others prefer to use 0 catgut as it is gone or degraded by the time of the pregnancy exam and the next cycle if the mare is not pregnant and needs to be bred again.7 If non-absorbable or delayed-absorbable sutures are used and not removed prior to the next breeding they can cause abrasions and trauma to the stallion’s penis and serve as a nidus for fecal accumulations.7 The ventral aspect of the Caslick should continue to 3–4 cm below the brim of the pelvis and at least 3 cm of the vulva should be left open for urination to occur. To insure that adequate healing of the tissues can occur and to prevent fistula formation the deepest layers of the fresh tissue must be apposed and the suture should exit/enter in the middle of the cut tissue, not deep to the cut tissue. Frequent rectal palpations within the 2-week period following placement may impede healing and result in fistula formation. Occasionally, a breeding stitch (usually doubled heavy gauge suture material or umbilical tape) is placed at the ventral aspect of the Caslick to prevent tearing of the closed labia during breeding and treating (Figure 262.2d). Care must be taken during breeding to insure that the breeding stitch does not cause trauma to the stallion’s penis.
Management The Caslick must be opened prior to foaling to prevent tearing of the apposed labia during parturition. This is performed by infusing the labial tissues with a local anesthetic and transecting the fused tissues with sharp scissors or a scalpel blade. The Caslick should be left intact and only opened as close to foaling as possible. Often veterinarians open Caslicks 20 to
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(c) (d) Figure 262.2 (a) Injection of anesthetic along the mucocutaneous junction of the vulvar labia. (b) Trimming of a thin strip of mucosa to create an 8–10mm wide section of exposed submucosa. (c) Suturing of the vulvar labia. (d) Placement of a ‘breeder’s stitch’ to reinforce the primary suture line during breeding. Reprinted from Trotter GW. Surgery of the perineum in the mare. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Baltimore: Williams and Wilkins, 1993; p. 419.
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Pneumovagina 30 days prior to the mare’s anticipated due date; however, in mares with poor perineal conformation, the Caslick should be left in place until just days prior to parturition to minimize the amount of contamination of the vagina and cervix and prevent ascending infection from occurring. In addition, early opening is sometimes associated with healing again. The Caslick should also be replaced as soon as possible after foaling.6
Episioplasty Some mares sustain trauma to the vestibulovaginal sphincter during foaling and the integrity of this protective barrier is lost. If a Caslick repair does not resolve the pneumovagina, because of more severe anatomical abnormalities, an episioplasty can be performed to improve the competency of the vestibulovaginal sphincter by reducing the diameter of the vestibule and enlarging the perineal body.6–9 This procedure is also known as a perineal body reconstruction, vestibuloplasty, deep Caslick, or Gadd technique.8, 9 This procedure is performed in the standing, constrained mare with local anesthesia. The perineum is cleaned and prepped in the same manner as for a routine Caslick. Local anesthetic is infused into the labia, the dorsal commissure, and lateral walls of the vestibule. The labia are retracted using stay sutures or towel clamps to provide visualization. An incision is made along the mucocutaneous junction of the labia, the dorsal commissure of the vestibule to the vestibulovaginal sphincter, and connecting the cranial aspect of the incision in the dorsal commissure to the most ventral aspect of the incision in the labia to form a triangle (Figure 262.3a). This triangular piece of mucosa is then removed from both sides (Figure 262.3b). The vestibular mucosa is closed using 2-0 or 0 absorbable suture in a horizontal mattress pattern (Figure 262.3c). The submucosa is closed in a simple interrupted pattern and the labia are apposed as for a Caslick procedure (Figure 262.3d,e). This surgery can be complicated by significant and obscuring hemorrhage, thereby prolonging the expected time required for the procedure.7 Sexual rest is recommended for 4–8 weeks while healing occurs. If the mare has a severely sunken anus this procedure is more beneficial when combined with a perineal body transection (see below). One must remember to open the dorsal vestibule and vulvar lips prior to foaling to avoid the possibility of a severe perineal laceration during foaling.
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Perineal body transection (perineoplasty) A third procedure described for correction of pneumovagina is a perineal body transection. A perineal body transection is primarily indicated in those mares with severely abnormal perineal conformation such that the pneumovagina is not corrected by a Caslick procedure or episioplasty. These mares include those with an extremely sunken anus and in which the majority of the vulval lips lies above the brim of the pelvis. This procedure is also known as the Pouret operation, as it was first described by Pouret in 1982.10 The objective of the surgery is to free completely the attachments between the ventral terminal rectal wall and the dorsal vaginal wall, allowing the vulva to assume a more normal vertical position and to reduce the pull of the sunken anus on the dorsal vagina. This procedure is performed with the mare confined in stocks with her tail wrapped and the perineum surgically prepared. Sedation may or may not be required; some surgeons prefer to perform an epidural while others use local anesthesia. When local anesthesia is used, 40–70 mL of lidocaine are liberally infused into the perineal body and laterally to include the vaginal walls to a depth of 8–14 cm. Towel clamps placed just ventral to the anus and dorsal to the dorsal commissure of the vulva are used to apply retraction and tension on the tissues as a 4–6-cm horizontal incision is made midway between the anus and the vulva and continued for 2–5 cm ventrally along either side of the vulva. The incision is continued cranially 8–14 cm through the muscles of the perineal body using both blunt and sharp dissection. During dissection a hand can be placed in the vestibule to help guide the dissection and prevent accidental entry into the rectum or peritoneal cavity. The dissection is complete when the connections between the rectum and caudal reproductive tract have been completely severed and the vulva has attained a normal vertical position (Figure 262.4a–c). The dead space created by the dissection is not closed, and while closure of the skin has been described, the incision can be left open to heal by secondary intention. Mares can be bred by artificial insemination immediately after the procedure is performed; however, natural cover should be delayed until the surgery site has healed, which takes 2 to 3 weeks. Good client communication is important when performing this procedure as the wound is large and unsightly and may be disturbing to some owners.7
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Figure 262.3 (a) The area of mucosa to be removed during an episioplasty. (b) Removal of the vestibular mucosa. (c,d) The vestibular mucosa is apposed and inverted into the vestibule with a horizontal mattress pattern. The submucosal tissues are apposed using simple interrupted sutures. (e) The vulvar labia are closed as when a Caslick is performed. Reprinted from Trotter GW. Surgery of the perineum in the mare. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Baltimore: Williams and Wilkins, 1993; p. 421.
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Figure 262.4 (a,b) Perineal conformation that predisposes a mare to aspiration of air and debris into the cranial reproductive tract. (c) Transection of the musculature between the rectum and vestibule/vagina allows the vulvar lips to relax caudally. (d) If left to heal by second intention a shelf is created between the anus and the dorsal commissure of the vuvla. (e) Approx 48 hours after surgery. Reprinted with permission from McKinnon et al.7
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References 1. Sager FC. Care of the reproductive tract of the mare. J Am Vet Med Assoc 1966;149:1541. 2. Roberts SJ. Veterinary Obstetrics and Genital Diseases (Theriogenology), 3rd edn. Woodstock, VT: 1971; p. 11. 3. Slusher SH, Freeman KP, Reszel JF. Eosinophils in equine uterine cytology and histology specimens. J Am Vet Med Assoc 1984;184:665–70. 4. Caslick EA. The vulva and the vulvo-vaginal orifice and its relation to genital health of the thoroughbred mare. Cornell Vet 1937;27:178. 5. Pascoe RR. Observations on the length and the angle of declination of the vulva and its relation to fertility in the mare. J Reprod Fertil Suppl 1979;27:299.
6. Pascoe RR. Vulvar conformation. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders, 2007; pp. 140–5. 7. McKinnon AO, Vasey JR. Selected reproductive surgery of the broodmare. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders, 2007; pp. 146–53. 8. Trotter GW, McKinnon AO. Surgery of abnormal vulvar and perineal conformation. Vet Clin N Am Equine Pract 1988;4:395. 9. Woodie B. Vulva, vestibule, vagina and cervix. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St Louis: Saunders, 2006; pp. 838–41. 10. Pouret EJM. Surgical technique for the correction of pneumovagina and urovagina. Equine Vet J 1982;14:249.
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Chapter 263
Surgery of the Caudal Reproductive Tract Angus O. McKinnon and Sarah L. Jalim
The procedures below represent techniques used at the Goulburn Valley Equine Hospital. This chapter is not intended as an exhaustive, heavily referenced treatise of all available techniques but is designed to provide practical techniques that work well for us. Readers should be aware that different methods are employed successfully by others. Many of the techniques presented are modifications of existing techniques, newer techniques and/or areas we feel warrant special attention. The conditions described will be commonly seen by a busy equine practitioner during the course of a breeding season. Most of the techniques are directed at restoring or improving fertility. Most are elective procedures, but require experience in order to make the correct diagnosis and institute the best therapeutic approach. Our experiences have led us to prefer the techniques discussed here to other published alternatives.
Patient selection Care should be exercised to select only those candidates that are likely to respond favorably to surgery. The probability of a successful outcome and subsequent production of a live foal must be evaluated with regard to:
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Severity and nature of the problem; Breeding history of the mare; Value of the mare and/or offspring (commercial or sentimental); Cost of the procedure; Mare age; Fertility of the stallion;
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Availability and suitability of assisted breeding techniques such as artificial insemination (AI) or embryo transfer (ET); Long-term predictive value of surgical intervention, i.e., temporary or permanent improvement; General health of the candidate; Perpetuation of heritable conditions; Insurance status and informed client consent; Ethical considerations; Experience of the veterinarian; Quality of on-farm management; Previous attempts at repair of the problem.
It is futile to perform a sophisticated, expensive surgery, for example, for vesico-vaginal reflux, if the mare has sustained chronic uterine damage and fibrosis that will render her highly subfertile or infertile despite an excellent surgical outcome. Full reproductive evaluation is necessary prior to surgery in any candidate subjected to repeated uterine bacterial insults, e.g., third-degree degree recto-vaginal lacerations that have been present for longer than one year.
Sedation and local anesthesia Sedation All the procedures discussed are performed with the mare standing and utilizing local infiltration and/or epidural anesthesia. Apart from minor procedures (Caslick operation – vulvoplasty) we prefer to have mares sedated and restrained in appropriate stocks, with facilities for cross tying and/or tail elevation. Sedation and analgesia is provided most commonly with a combination of detomidine (0.02 mg/kg bodyweight, i.v.) and butorphanol (0.005–0.01 mg/kg bodyweight, i.v.). Longer surgeries may require the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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repeated use of low doses of intravenous xylazine (100 mg). Cost and length of sedation/analgesia are important considerations. In addition, some have other physiological side-effects, e.g. increased urinary output with xylazine. Detomidine provides excellent sedation and analgesia;1 however, sedation may persist for hours after administration. Analgesia and anesthesia Epidural anesthesia can be extremely effective; however, variation in response, individual susceptibility and time before appropriate anesthesia is obtained, occasionally make it less rewarding than tranquilization and local infiltration for some of the procedures discussed here. Our technique for epidural anesthesia is similar to that described by Turner and McIllwraith.2 Caudal epidural injection of local anesthetic to produce anesthesia of the tail and perineal structures in the conscious standing horse has been further developed by the use of a catheter placed into the epidural space in order to provide longduration analgesia and anesthesia. More recently, opioid, alpha-2 adrenergic agonists, ketamine, and other analgesic agents have been administered by caudal epidural injection, providing pain relief in both conscious, standing and anesthetized, recumbent horses.3 Epidural administration of more than 6–9 mL of 2% local anesthetic to a 500 kg mare can be associated with loss of motor control to the hindlimbs and subsequent unexpected recumbency. These mares are difficult to manage and may require support for 2 to 4 hours. Epidural injection of xylazine for perineal analgesia has been reported to be very efficacious.4–7 A dose of 0.17 mg/kg (approximately 75–100 mg in an average-sized horse) was as effective as a local anesthetic agent for anesthesia and had no or minimal depressive effect on motor nerves in the lumbosacral intumescence. Another advantage was the demarcation of the area of anesthesia by an area of sweating (dermatome) that was correlated both temporally and topographically with region(s) of anesthesia.6 In our experience, the perineal dermatome is not always obvious. In addition, it takes approximately 30–40 minutes for good analgesia to be recognized when performing epidural anesthesia with xylazine. We prefer the following mixture: xylazine (∼ 0.17 mg/kg bodyweight), lidocaine (∼ 0.15 mg/kg bodyweight; ∼ 3 mL of 2% lidocaine solution) and ∼ 6 mL of lactated Ringer’s solution to a volume of 10 mL. The volume administered as an epidural in the first intercoccygeal space is titrated based on the size of the mare. Large mares (> 600 kg) may receive the full 10 mL dose and smaller mares (450–500 kg) receive be-
tween 7 and 8 mL. With this regime, loss of tail tone has occurred in association with a successful ‘block’ within 5–10 minutes, and surgery generally has begun by ∼ 20–30 min post-administration. Our most commonly used method of epidural anaesthesia is to begin with the infusion of 1 mL of local anesthetic through a 25-gauge needle subcutaneously above the first moveable coccygeal/sacral joint (usually C1 and C2). Next a 38 mm, 18-gauge needle (bevel forward or up and introduced at 45◦ to the skin), that has a small amount of anesthetic solution to create a meniscus on the needle hub, is introduced through the local bleb. Recognition of the fluid being aspirated into the needle infers correct placement and the appropriate amount of solution is injected slowly into the epidural space whilst continually evaluating the ease of administration to ensure the needle tip is still in the appropriate place. Occasionally epidural anesthesia is difficult to achieve (e.g., infusion outside the epidural space before proper epidural placement, hemorrhage through the needle and tissues, anatomic abnormalities and fat horses). Provided local anesthesia can be obtained by infiltration without adverse affects on wound healing, we prefer to perform surgery without the epidural and provide incremental intravenous administration of xylazine, i.e. 50–100 mg dose, to already sedated patients when or if minor surgical discomfort is recognized. In some instances, we have been able to identify correct epidural placement by injecting the anesthetic slowly while moving the needle until there is an obvious change in the amount of resistance needed to advance the solution. This situation is usually associated with a mare with abnormal anatomy (e.g., after a traumatic foaling) and lack of movement of fluid in the needle hub.
Vestibule and vulvar labia Pneumovagina There are three barriers to aspiration of air and contaminants into the caudal reproductive tract:
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the vulvar labia; the vestibular sphincter; the cervix.
It would appear that the vestibular barrier is more important than previously recognized;8 however, when any of these protective barriers are rendered incompetent, contamination and concomitant vaginitis, cervicitis and metritis may result.
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in the uterus10 (see Chapter 223). If in doubt about a mare’s potential to aspirate air into the vagina after clinical evaluation, we recommend surgical treatment of all mares that have been barren one or more breeding seasons, unless other causes of reduced fertility are obvious, for example, stallion or farm management. Caslick operation (vulvoplasty)
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Figure 263.1 (a) Opening the vulvar lips in a mare with a good seal against aspiration of air reveals the vestibular sphincter (larger arrow). The approximate location of the urethra is indicated by the small arrow. (b) A mare with a weak vestibular sphincter will aspirate air into the vagina, revealing the cervix in the background.
Pneumovagina can result from faulty perineal conformation, previous injury to perineal tissues or effects of poor body condition. Pneumovagina can also be iatrogenic, i.e., from reproductive examination or breeding. Recognition of candidates with potential pneumovagina is made by anatomical observation.9 Predisposition to the condition is associated with a low pelvis, tilted vulvar labia and a sunken anus. If possible mares should be examined during estrus when the reproductive tract is relaxed. Parting the vulvar labia results in loss of integrity of the vestibular sphincter and an audible ingress of air into the vagina in mares predisposed to pneumovagina (Figure 263.1a,b). Recognition of foamy exudates from the vulva may often be noted during rectal examination. In addition ultrasonography has been used to visualize air
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The procedure was first described by Dr E.A. Caslick in 1937,11 and involves apposition of the dorsal portions of the vulvar labia. Local anesthesia of vulvar labia, removal of a thin portion of mucosa from the mucocutaneous junction, and apposition of the cut edges is simple, quick and effective. Length of apposition is determined by height of pelvis relative to vulvar labia. In general, the join should continue to a point at least 3 to 4 cm below the level of the pelvic brim, and approximately 3 cm should be left unopposed to allow for urination (Figure 263.2a–e). If this is not possible, other surgical techniques (see below) may be necessary.
Points for consideration
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Suture material – We use 1 chromic catgut (metric 4) on a continuous spool. The benefit of gut is that it is degraded by the time of either pregnancy testing (∼ 15 days) or the next breeding cycle (if the mare is not pregnant). Other materials that are non absorbable or have delayed absorbable properties may damage the stallion’s penis if the mare is rebred, and serve as an excellent material for fecal accumulation. In addition, the continuing presence of the non-absorbable suture creates the potential for fistula formation.
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Figure 263.2 (a) Tissue should be removed from the mucosal surface of the vestibule at the mucocutaneous junction and not the skin of the vulva lip. (b) A simple continuous pattern is quicker and just as effective as a simple interrupted pattern. Failure to exit/enter in the middle of the cut tissue may result in wound breakdown. (c) Incision with a scalpel blade rather than mucosal removal is recommended for maidens (e.g., race mares). (d) A breeding stitch may save unnecessary Caslick replacement and facilitate breeding and some manual vaginal manipulations for uterine treatments. (e) Fistula formation is commonly associated with poor tissue debridement, poor tissue apposition, or transrectal palpation within a few days of the procedure.
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Suture pattern – A simple continuous pattern is quicker and just as effective as a simple interrupted pattern. However, failure to appose the deepest layers of the exposed tissues may result in fistula formation. Failure to exit/enter in the middle of the cut tissue caused by taking bites too deep will result in wound breakdown. In general, it is best to start at the dorsal vulval commissure to ensure good apposition there, as this is the most common site of fistula formation. Method of mucosa removal r Tissue should be removed from the mucosal surface of the vestibule at the mucocutaneous junction and not the skin of the vulva lip. r Removal of too much tissue will result in excess fibrosis and difficulty with the next closure. r If multiple procedures are performed in one season, slight debridement with a scalpel blade or scissors is sufficient, providing sutures are carefully placed. r Incision with a scalpel blade rather than mucosal removal is recommended for maidens (i.e., race mares). This results in a thinner join that is easily opened at the breeding farm. Postoperative treatment r Frequent transrectal palpations within the first few days of placement can result in fistula formation. This may occur when a mare stays in estrus despite ovulating and is re-presented for further ovarian evaluation after a Caslick operation. r Remove (open) 2–3 weeks prior to anticipated foaling. Variations r Placement of a breeding stitch (usually doubled, heavy-gauge suture material or umbilical tape) will save unnecessary replacement and facilitate breeding and some manual vaginal manipulations for uterine treatments. Care is needed with natural service to prevent penile trauma. r Intrauterine treatments can still be performed and protection of the vulvoplasty is possible by introduction of an infusion pipette into the vagina using one or two fingers under the ventral border of the Caslick (past the vestibular sphincter) and using transrectal manipulation of the pipette through the cervix similar to, but with a little more difficulty, than in cattle. Alternatively, the pipette can be visually guided through the cervix with the aid of a tubular vaginal speculum.
Episioplasty This procedure is used in cases with more severe anatomic abnormalities where the Caslick vulvo-
plasty is ineffective, and in mares with extensive or repeated second-degree perineal lacerations. Technically the previous surgery is an episioplasty as well; however, we12 use the term to describe the more extensive procedure previously referred to as (i) a deep Caslick, (ii) the Gadd technique,13 or (iii) perineal body reconstruction, or perineoplasty. The procedure is designed to restore some degree of function to the perineal body (Figure 263.3a–d). The surgery is performed on the standing mare using local anesthesia or, occasionally, epidural anesthesia. Visualization is improved by good lateral retraction of the vulvar labia and light sources with good illumination. We prefer to perform all perineal surgeries using a tungsten light source on a head lamp. A triangular portion of the dorsocaudal mucosa is removed from both sides of the vestibule. The ventral borders are apposed with a simple continuous suture pattern using an absorbable material (2/0 polydixanone (PDS) or chromic catgut), and dead space with single interrupted or simple continuous sutures of the same material. The muco-cutaneous junction is closed in a similar fashion to the Caslick operation. Apposition of the ventral borders of the incision and obliteration of the potential space above result in an increase in size of the perineal body and decreased propensity of the vestibule to expand and generate negative pressure.
Points for consideration
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The procedure may be most beneficial when combined with a perineal body transection (see below). Failure to ‘open’ the dorsal vestibule and vulval lips (episiotomy) prior to foaling (2–3 weeks) may result in a severe perineal laceration. The surgery usually requires 30–40 minutes and is sometimes associated with significant and annoying obscuring hemorrhage. It is also possible to close the perineal body dead space with externally placed vertical mattress sutures, with tension-spreading devices such as pieces of rubber tubing or buttons (S. Slusher, personal communication, 1986) (Figure 263.3e–i).
Urovagina Vesicovaginal reflux (also termed urovagina or urine pooling) is a relatively common cause of subfertility/infertility seen predominantly in older multiparous broodmares. The resultant vaginitis, cervicitis, and endometritis compromises fertility by causing degenerative endometrial changes over time, that lead to an increased early embryonic death and increased susceptibility to infection associated
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Figure 263.3 (a) A triangular portion of the dorsocaudal mucosa is removed from both sides of the vestibule. (b) Excess vestibular mucosa is trimmed. (c) The ventral borders are apposed with a simple continuous suture pattern, using an absorbable material. (d) The dead space is apposed with single interrupted or simple continuous sutures of the same material. (e–i) Large perineal body defect repair illustrated from local anaesthetic infiltration to closure of the ventral perineal body.
with poor uterine defense mechanisms. In addition, urinary osmolarity and pH markedly reduce spermatozoal motility14 and may prevent spermatozoa from reaching the oviduct. In 1985, Jay Belden, then of Fossil Creek Farms, Colorado, demonstrated to us his technique for surgical correction of urovagina. After many surgeries between us using Jay’s technique, we decided the technique was efficacious and should be reported.15 Prior to this, we had tried the other described techniques such as caudal relocation of the transverse fold described by Monin,16 and the urethral extension techniques of Brown17 and Shires,18 and had difficulty
obtaining consistently good results. The technique to treat urovagina described below (Figure 263.4a–k) is based on the original publication of McKinnon and Belden.15 Recently we have reviewed an extended series of cases treated this way.19 Aanes modified Finochetto retractors (Sontec Instruments, Englewood, CO) placed into the vagina are used for exposure and visualization of the transverse fold and vestibule. The midline caudal border of the transverse vaginal fold is grasped with Vullsellum tissue forceps and retracted caudally toward the surgeon. A horizontal transverse incision to the submucosa is made 2 to
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Figure 263.4 (a) A horizontal transverse incision to the submucosa is made 2 to 4 cm cranial from the caudal border of the transverse fold with a No. 12 Bard Parker blade. (b) The incision is continued laterally and then slightly dorsally to the vaginal wall, and then extended caudally from the lateral border of the transverse folds approximately one-half to two-thirds of the distance between the vestibular floor and roof to the vulvar labia. (c) Dissection of the tissue flaps continues until the cut edges can be reflected without tension, and they are then inverted in the suture pattern to form a tunnel. (d) The suture pattern used to appose the submucosal tissue layer is a continuous modified Connell (horizontal mattress) suture. (e) The final configuration is in the shape of a Y, with an extended tunnel from the urethral orifice (under the transverse fold) to the caudal vestibular vault. (f–k) Surgical illustration of the urethral tunnel. (f) Forceps grasping the middle of the transverse fold. (g) Dissection of the transverse fold. (h) Completed dissection of the transverse fold. (i) Dissection of the vestibular wall. (j) Completion of both sides of the Y pattern. (k) Completion of the urethral tunnel.
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Surgery of the Caudal Reproductive Tract 4 cm cranial from the caudal border of the transverse fold with a No. 12 Bard Parker blade (Figure 263.4a). The incision is continued laterad and then slightly dorsad to the vaginal wall. The vaginal retractors are then repositioned, and a No. 20 Bard Parker blade is used to extend caudad the incision from the lateral border of the transverse folds. This incision is made approximately one-half to two-thirds of the distance between the vestibular floor and roof and is extended to the vulvar labia. The caudal cut edge of the transverse fold incision and the ventral cut edge of the vestibular wall incision are dissected so that the free tissue flaps are reflected caudad and mediad, respectively. This causes the vestibular mucous membranes of the free tissue flaps to be reflected ventrally and the underlying cut surface to be exposed dorsally. Next, the retractors are repositioned, and an identical incision is made on the opposite vestibular wall, extending from the lateral extremity of the transverse fold caudally toward the surgeon (Figure 263.4b). Dissection of the tissue flap from the transverse fold is continued until the caudal cut edge can be reflected approximately 3 to 6 cm toward the surgeon. The dissection of both vestibular wall flaps is continued ventrally until the cut edges can be reflected without tension past the midline (Figure 263.4c). The suture pattern (Figure 263.4d) used to appose the submucosal tissue layer is a continuous modified Connell suture, using 2–0 chromic gut or polydioxanone suture (PDS). The final configuration is in the shape of a Y, with the tail of the Y pointing caudally. The first suture line begins cranially and laterally at the junction of the transverse fold and vaginal wall incisions, and is ended at the midpoint of the transverse fold reflection (junction of the Y). Cut edges of the transverse fold and vaginal wall are inverted so that denuded tissues are in apposition. The second suture line began on the opposite side, at the junction of the transverse fold and vaginal wall incisions, and is continued to the end point of the first suture pattern and then on the midline, toward the surgeon, to the caudal end of the vaginal wall reflections. The result is an extended tunnel from the urethral orifice (under the transverse fold) to the caudal vestibular vault. It is important to have minimal tension associated with the suture line, and have all apposed edges inverted so that dissected or denuded tissues are in apposition (Figure 263.4e). An extra suture layer is often oversewn to aid in establishing a tunnel of correct size and tension. The denuded tissues created dorsally by the dissection of the transverse and vaginal folds are allowed to heal by second intention. If necessary, an episioplasty may be performed after surgery (Figure 263.4f–k).
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A uterine biopsy should be performed before surgery on all mares that have been barren for longer than one season. This will aid in giving an accurate prognosis for future fertility and is important in cases of financial constraints. The technique described is our technique of choice for treatment of all mares with urovagina. Other treatments such as perineal body transection20 or caudal relocation of the transverse fold16 often appear only temporarily effective. Episioplasty performed at the same time may result in excessive ablation of vestibule. In some mares, urine will pool in the caudal relocation of the urethral tunnel and it will be voided during exercise, commonly resulting in urine staining of the perineum. While this does not affect fertility, it is unsightly and may be treated by incision into the new urethral tunnel to a point 2–4 cm cranial to the vulval lips but not far enough to allow urovagina to reoccur. Fistula formation has been noted as a problem.15,21–23 In the series published from our hospital19 we noted a fistula formation rate of 11.5% (7/61). In all cases fistulas occurred at the junction of the Y-shaped incision, and were repaired successfully in standing sedated mares. The mean age of mares with fistula formation was 16 years (median 17 years). Six of the 61 cases in this series were referred to our facility for surgery following previous unsuccessful attempts at urovagina correction. Four of the fistulas identified were in this population of mares which had previously undergone an unsuccessful surgery. In addition these were the majority of mares (3/5, 60%) which had a continuation of urovagina after the initial surgical repair. Of the seven mares in which fistula formation occurred, five were corrected following a single surgery; two mares required two surgeries to correct the fistula. Some clinicians choose to manage urovagina by evacuating urine from the vagina pre-breeding; however, chronic urovagina could lead to sustained endometritis and other irreversible endometrial changes. Unless the mare is young, has undergone substantial reversible weight loss or is recently postpartum, we always elect to treat the condition of urovagina surgically if the economic constraints of the owner are able to be met. The surgery has classically been left to the end of the season in barren mares that were identified as having urovagina during the breeding season. A more recent development has been to breed the mares during the same breeding season in the cycle immediately after repair, or to breed the mares
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prior to repair (immediately after identification of urovagina) and then perform the surgery within 48 hours of breeding. The latter group requires evacuation of any urine with a large tampon immediately prior to service. When mares were bred in the same cycle as surgery, the first cycle following surgery, second cycle following surgery, or the following breeding season after surgery the seasonal pregnancy rates were 89% (8/9), 63% (10/16), 67% (2/3), and 63% (15/24), respectively. After removing four mares that died of natural causes prefoaling, the foaling rates were 88% (7/8), 50% (7/14), 0% (0/3), and 52% (12/23), respectively. All mares bred in the same cycle or next cycle as surgery were bred once only that season, so the pregnancy rate per cycle of 72% (18/25) was identical to the seasonal pregnancy rate.19
tering the peritoneal cavity. The aim of the surgery is to allow the vulval lips to assume a more horizontal position by freeing them from attachments to the rectum. Generally the surgery is finished when the desired external conformation is achieved after allowing for wound contraction (Figure 263.5e–g). Closure of the skin incision was originally recommended;20 however, we commonly leave the incision open. Closing the skin occasionally results in dehiscence and also seems to influence the result after wound contracture occurs. The wound heals by second intention and healing is surprisingly rapid (2–3 weeks).
Points for consideration
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Perineal body
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Surgeries of the perineal body are necessary to correct severe anatomic defects (perineal body transection – PBT) or injuries due to foaling (perineal lacerations and fistulae).
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Perineal body transection
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Perineal body transection (PBT) has been described as beneficial for both pneumovagina and urovagina.20 Our experiences would suggest definitely the former but rarely the latter. The primary indication for PBT is in mares with such external reproductive conformation that we are not able to prevent pneumovagina after either Caslick vulvoplasty or episioplasty, i.e., mares with a severely sunken anus and most or all of the vulvar labia positioned above the brim of the pelvis (Figure 263.5a,b). The procedure is performed with local anesthesia with or without sedation. Local anesthesic (40– 70 mL) is liberally infused into the perineal body and laterally to include the vaginal walls to a cranial depth of 8–14 cm. Towel clamps positioned just ventral to the anal sphincter and dorsal to the dorsal commissure of the vulva are used to provide retraction and tension while a 4–6 cm horizontal incision is made midway between the anus and dorsal vulva. The incision is continued for a short distance (2–5 cm) ventrally along either side of the vulva. Sharp and blunt dissection are used to completely transect the muscular and ligamentous supporting tissues between the rectum and vestibule (Figure 263.5c,d). Depending on the individual mare, the dissection proceeds cranially for 8–14 cm and finishes before en-
Placing a hand in the vestibule enables accurate dissection. Penetration of the vagina or rectum should be avoided. The wound is unsightly while healing and we recommend careful client communication and/or hospitalization for 1–2 weeks. The benefits from corrective surgery are immediate and mares can be bred at the first opportunity, providing qualified personnel are available. Natural service is generally delayed 2–3 weeks to allow strengthening of the dorsal vestibule and vagina. Despite the surgery causing moderate hemorrhage, we have not had to take any special precautions or protective measures.
Perineal lacerations Most perineal injuries occur at the time of foaling and are associated with:
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a malpresented or occasionally oversized fetus or; extensive, vigorous, inappropriate or sometimes unavoidable manual manipulation during parturition; violent expulsive efforts of the mare; maiden mares being most predisposed to third-degree perineal lacerations and rectovaginal/rectovestibular fistulas.
The injuries are commonly referred to as first-, second-, or third-degree perineal lacerations (PL).
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First-degree PL involves only the mucosa of vestibule and skin of the dorsal commissure of the vulva. Second-degree PL involve both the mucosa and submucosa of the dorsal vulva, and some of the musculature of the perineal body, in particular the constrictor vulvae muscle. There is no damage to rectal mucosa.
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Figure 263.5 (a) Mares with most or all of the vulvar labia positioned above the brim of the pelvis are ideal candidates for PBT. (b) Mares with a severely sunken anus and little vulvar lips beneath the pelvis will benefit from a PBT. (c) The PBT surgery aims to completely transect the muscular and ligamentous supporting tissues between the rectum and vestibule. (d) The dissection proceeds cranially for 8–14 cm and finishes before entering the peritoneal cavity. (e) A 4–6 cm horizontal incision is made midway between the anus and dorsal vulva. (f) Sharp and blunt dissection are used to completely transect the muscular and ligamentous supporting tissues between the rectum and vestibule. (g) Closure of the skin incision was originally recommended; however, we commonly leave the incision open.
Minor first-degree PL may require no treatment. Extensive lacerations may require episioplasty or perineal body reconstruction. If tissue damage results in significant edema, inflammation and infection, then surgical correction is often delayed for 2–4 weeks.
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Third-degree PL results in tearing of the vestibular and sometimes vaginal wall and disruption of the perineal body, anal sphincter, and rectal wall. This results in a common opening between the rectum and the vestibule.24 The constant presence of
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feces in the vestibule and occasional unpleasant sound from air movement make repair imperative for breeding and recommended for future riding horses. Immediate care for the mare at foaling involves protection against tetanus, and can also involve systemic treatment with antibiotics and antiinflammatory drugs. The surgical correction is delayed at least 4 weeks to allow second intention wound healing to occur. Immediate repair at the time of foaling can be successful but is generally not recommended.24 The longer the injury is left untreated, the more opportunities for continual contamination of the reproductive tract; however, this is related to functional capabilities of vestibular sphincter. Surprisingly, enodmetritis induced by third-degree PL can often be resolved within 2 weeks following surgical correction without concomitant intrauterine treatments.25 Most commonly, surgery is performed after weaning if a live foal was delivered. The cervix must be examined prior to surgical correction of PL, and if a prolonged time between foaling and repair has occurred, a full reproductive evaluation including a uterine biopsy is warranted.
Surgical technique There are many methods described. They are all are modifications of either a single or two-stage repair. A modification of the single-stage repair26 is the technique we prefer. Prior to surgery, vigorous efforts are made to modify the consistency of the feces. Mares are held off feed for at least 24 hours and given a mineral oil drench (2–3 liters) immediately before or after surgery. Following surgery, mares are placed on pasture if available and mineral oil is administered by nasogastric tube daily for 3 days as necessary to maintain a soft fecal consistency. Mares are restrained standing, sedated, and epidural anesthesia is employed. Fecal material from the rectum is removed as far cranial as possible. Large wads of cotton wool are inserted into the cranial rectum to absorb fecal fluid and prevent fecal contamination of the surgical site. Tissues are cleansed and prepared for aseptic surgery. Towel clamps are inserted into the ventral anal sphincter in a configuration that, when apposed, represent the ideal surgical apposition point. In addition, towel clamps are placed on the dorsal vulvar commissure and then retracted to provide visualization for surgical access or, alternatively, stay sutures are used to retract the vulvar labia. An incision is made 5–10 mm ventral to the scar tissue line marking the junction between the vestibule/vagina and
rectum (Figure 263.6a,b). Tissues ventral to the incision are dissected to create mobilized vestibular mucosa and submucosa that, when apposed from side to side, will form the ventral border of the perineal body (Figure 263.6c). Dissection is continued for a distance of 3–4 cm cranially and 5–8 cm ventrally on the vaginal/vestibular wall. Tissues dorsal of the incision are debrided and rectal submucosa is undermined to allow sufficient mobilization to permit side-to-side apposition of the ventral border of the terminal rectum with as much caudal retraction of the rectal mucosa and submucosa as can be achieved without undue tension. All tissues are then sutured concurrently and incrementally from cranial to caudal. The rectal and vaginal reflections are apposed by a continuous modified inverting Connell suture. Suture material preferred is No. 1 or 2 PDS for the fibrous layer that will reform as the perineal body, 0 or 1 PDS for the vagina/vestibule, and 0 or 2/0 for the rectal mucosa. The first layer sutured is the vaginal/vestibular submucosal layer that will form the ventral portion of the perineal body (Figure 263.6c). This can be advanced to within 3–5 cm of the vulvar lips before beginning the rectal submucosal layer. When the suture pattern of the rectal submucosa has advanced 3–5 cm, the suture material is held with slight tension by an assistant and the resulting dead space in between that will be the recreated perineal body is obliterated with one of several different methods.
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Classically, this layer is apposed with single, multiple-bite pursestring sutures either close to the midline of the vagina/vestibule (Figure 263.6d–f) or involving the lateral margins of vaginal/vestibular walls (Figure 263.6g,h). The former technique results in dead space laterally that is not properly obliterated, and the latter technique creates excessive tensions on lateral tissues. An alternate is to create a similar suture pattern but to use a continuous pursestring suture. Another option is to use two concurrent and sideby-side continuous apposition sutures that overlap on the midline (Figure 263.6i,j) and progress caudally towards the surgeon. This technique has been in use at the Goulburn Valley Equine Hospital for 6 years. Finally, we have recently developed an alternate technique that creates the perineal body with less side-to-side (lateral to medial) tension. The technique aims to create the tension in a transverse orientation similar to the tensions with rectovaginal fistulas (see below). The suture pattern is a simple continuous pattern apposing dorsal rectal submucosa to dorsal vaginal/vestibular submucosa (ventral wall of the perineal body) from the lateral border of the perineal body, through midline to the
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Figure 263.6 (a) An incision is made 5–10 mm ventral to the scar tissue line marking the junction between the vestibule/vagina and rectum. (b) The black line represents the approximate dissection plane along the perineal body. (c) The tissues mobilized from dissection (dotted line) are joined ventrally to form the ventral border of the perineal body. (d–f) Schematic and surgical representation of placement of the six-bite suture to obliterate the space between the rectum and vagina/vestibule. The arrows in 6 d and 6f represent the area of poor tissue apposition with this suture pattern. (g and h) Schematic and surgical representation of placement of a simple interrupted suture. The arrows and their relative size represent comparative tension areas which are largest from lateral to medial. (i and j) Schematic and surgical representation of placement of the two concurrent and side-by-side continuous apposition sutures that overlap on the midline. The arrows and their relative size represent comparative tension areas which are largest from lateral to medial. (k and l) Schematic and surgical representation of placement of a simple continuous pattern apposing dorsal rectal submucosa to dorsal vaginal/vestibular submucosa (ventral wall of the perineal body), from the lateral border of the perineal body, through midline to the opposite side and back again. The arrows and their relative size represent comparative tension areas which are now even.
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opposite side and back again (Figure 263.6k,l). It is frequently necessary to tie and begin another new suture, as the pattern will result in use of as much as two or three packets of No. 2 PDS. The three areas of apposition are then alternatively progressed caudally, 2–4 cm each time, diverging at the perineal body, until the ventral vestibular/ vaginal submucosa stitch can be tied and vulval lips closed by a Caslick operation. The horizontal shelf of the internal perineal body created by the technique above is terminated approximately 4 cm from the anal sphincter, and the caudal portion of the internal perineal body is created by apposition of the remaining tissue from side to side vertically. The apposing suture of the rectal submucosa is terminated at the dorsal perineal body at the level of the defect in the anal sphincter. The anal sphincter edges may or may not be apposed. Postoperatively, apart from antimicrobials, anti— inflammatories, and protection against tetanus, the management of fecal consistency is most important.
Points for consideration
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Techniques are largely dictated by experience. We believe these cases are best treated by referral practices accustomed to performing this surgery. The better the surgeon’s anatomical and functional understanding of the perineal body, the greater the likelihood of a favorable surgical outcome. Management of fecal consistency with green pasture and mineral oil is important. The use of alternate methods to create less side-toside tension in the perineal body appears to allow better functional repair of the rectum (increased size) with less danger of dehiscence and a better functioning perineal body.
Fertility is generally good after successful repair, provided no other causes of reduced fertility exist. A slightly higher incidence of perineal trauma is expected with these mares. Attendance at foaling is advised. Rectovestibular fistula Rectovestibular fistula (RVF) is a relatively common injury sustained during foaling. It may also arise as a result of an unsuccessful surgical repair of a third-degree perineal laceration. Fistulas most commonly occur secondary to dystocia and are usually in primiparous mares. They may be caused by the foal’s nose or, more commonly, a foot (or feet) being forced through the dorsum of the vestibule or vagina into the rectum. Spontaneous retraction or manual replacement of the foal’s head or extremity(s) into the
correct position limits the injury to a RVF. If parturition proceeds before correction, the result is usually a third-degree perineal laceration. It is our impression that improved management of stud farms with early appropriate intervention during dystocias has meant that RVFs now less commonly progress to third-degree perineal lacerations. We have reported on breeding the mares on an induced (prostaglandin) second postpartum estrous period and then immediately (within 2 days) performing the fistula repair.27 When mares were bred in the same cycle as surgery, the next cycle following surgery, or the following breeding season after surgery, the pregnancy rates were 5/5, 5/6, and 10/12, respectively. Foaling rates were 4/5, 4/6, and 7/12, respectively. The two mares already pregnant at the time of surgery foaled successfully.27 For breeding immediately prior to surgery to be successful, as much fecal material as possible is removed immediately before breeding (either natural or artificial insemination). After breeding, the uterus is lavaged at least daily until repair is performed. Many surgeons treat RVFs by converting them to third-degree PL and repairing as previously described, either standing or under general anesthesia.28–30 For deep (cranial) RVFs a perineal body transection has been utilized.24 Recently a pedicle flap has been described.31 During the past two decades we have repaired RVFs with a transrectal approach, which we first described in 1991.32 A similar technique was described in the 1996 American Association of Equine Practitioners proceedings by Dr Stephen Adams.33 In our hospital, Aanes modified Finochetto retractors are inserted into the rectum through the anal sphincter. These retractors make the surgery quite simple (Figure 263.7a,b). The edges of the fistula are dissected rostrally and laterally using a No. 10 or No. 12 Bard Parker blade (Figure 263.7c). The rectal mucosa of the caudal border is inverted into the rectum with Babcock or Vulsellum forceps to allow easier dissection of the caudal border, using a No. 12 Bard Parker blade (Figure 263.7d). The tissue planes are undermined for at least 4 mm, commencing in approximately the middle of the fibrous scar that would become the reconstructed perineal body. This initial dissection allows exposure of three clear tissue planes – the rectal mucosa, the perineal body, and the vaginal mucosa. These three tissue layers are then closed separately, inverting the rectal mucosa into the rectum and inverting the vaginal mucosa into the vestibule. The first layer closed is the perineal body (middle layer) using a simple continuous appositional pattern with size 0 or 1 PDS. Next the rectal mucosa is inverted with a Connell pattern (continuous horizontal mattress) using 0 PDS, and following this the rectal
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Figure 263.7 (a) Aanes modified Finochetto retractors in the rectum. (b) Surgical exposure is improved with retractors placed rectally. (c) Cranial dissection of the RVF ring with a No. 12 Bard Parker ‘hook’ blade. (d) Caudal dissection of the RVF ring is improved with inversion into the rectum with Babcock or vulsellum forceps.
mucosa is oversewn with size 2/0 polydioxanone with a simple continuous pattern. Finally, the retractors are repositioned in the vestibule/vagina and the vaginal mucosa is apposed and inverted into the vagina with a modified Connell pattern with size 0 or 1 PDS. All three tissue layers are closed transversely to minimize tension on the suture lines and best align normal anatomy. When we previously described the surgical correction32 it was recommended to alternate the orientation of the closure. Now we recommend that all three layers be closed transversely, not longitudinally. Restraint and postoperative care are similar to mares with perineal lacerations; however, little attention is given to fecal softening.
Points for consideration
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The caudal border of the fistula may be difficult to debride and is most easily rotated towards the surgeon by grasping it with Babcock or Vulsellum forceps (from the rectum) and inverting it caudally
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(Figure 263.7d). On occasion it may be helpful to have the surgeon’s hand in the vagina while working on this part of the fistula. The use of a No. 12 blade greatly facilitates dissection of the caudal fistula ring. These results above27 suggest that delaying breeding until the following breeding season is not necessary. In addition, breeding on the same cycle as surgical repair is a recently reported technique27, 34 that should be considered in an effort to maintain a yearly foaling interval.
References 1. McKinnon AO, Carnevale EM, Squires EL, Jochle W. Clinical evaluation of detomidine hydrocholoride for equine reproductive surgery. Proceedings of American Association of Equine Practitioners 1988; pp. 563–8. 2. Turner AS, McIlwraith CW. Anesthesia. In: Turner AS, McIlwraith CW (eds) Techniques in Large Animal Surgery. Philadelphia: Lea & Febiger, 1989: pp. 9–32. 3. Robinson EP, Natalini CC. Epidural anesthesia and analgesia in horses. Vet Clin North Am Equine Pract 2002;18(1):61–82,vi.
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4. Fikes LW, Lin HC, Thurmon JC. A preliminary comparison of lidocaine and xylazine as epidural analgesics in ponies. Vet Surg 1989;18(1):85–6. 5. Grubb TL, Riebold TW, Huber MJ. Comparison of lidocaine, xylazine, and xylazine/lidocaine for caudal epidural analgesia in horses. J Am Vet Med Assoc 1992;201(8):1187–90. 6. LeBlanc PH, Caron JP, Patterson JS, Brown M, Matta MA. Epidural injection of xylazine for perineal analgesia in horses. J Am Vet Med Assoc 1988;193(11):1405–8. 7. LeBlanc PH, Caron JP. Clinical use of epidural xylazine in the horse. Equine Vet J 1990;22(3):180–1. 8. Hinrichs K, Cummings MR, Sertich PL, Kenney RM. Clinical significance of aerobic bacterial flora of the uterus, vagina, vestibule, and clitoral fossa of clinically normal mares. J Am Vet Med Assoc 1988;193:72–5. 9. Pascoe RR. Observations on the length and angle of declination of the vulva and its relation to fertility in the mare. J Reprod Fertil Suppl 1979;299–305. 10. McKinnon AO, Squires EL, Carnevale EM, Harrison LA, Frantz DD, McChesney AE et al. Diagnostic ultrasonography of uterine pathology in the mare. Proceedings of American Association of Equine Practitioners, 1987; pp. 605–22. 11. Caslick EA. The vulva and the vulvovaginal orifice and its relation to genital health of the thoroughbred mare. Cornell Vet 1937;27:178. 12. Trotter GW, McKinnon AO. Surgery for abnormal vulvar and perineal conformation in the mare. Vet Clin North Am Equine Pract 1988;4:389–405. 13. Gadd JD. The relationship of bacterial cultures, microscopic smear examination and medical treatment to surgical correction of barren mares. Proceedings of American Association of Equine Practitioners, 1975;21:363–8. 14. Griggers S, Paccamonti DL, Thompson RA, Eilts BE. The effects of pH, osmolarity and urine contamination on equine spermatozoal motility. Theriogenology 2001;56(4):613–22. 15. McKinnon AO, Belden JO. A urethral extension technique to correct urine pooling (vesicovaginal reflux) in mares. J Am Vet Med Assoc 1988;192:647–50. 16. Monin T. Vaginoplasty: a surgical treatment for urine pooling in the mare. Proceedings of American Association of Equine Practitioners, 1972; pp. 99–102. 17. Brown MP, Colahan PT, Hawkins DL. Urethral extension for treatment of urine pooling in mares. J Am Vet Med Assoc 1978;173:1005–7. 18. Shires GM, Kaneps AJ. A practical and simple surgical technique for repair of urine pooling in the mare. Proceedings of American Association of Equine Practitioners, 1986;32:51–6.
19. Jalim SL, McKinnon AO. Surgical results and fertility following correction of vesico-vaginal reflux in mares. Aust Vet J 2010;88(5):182–5. 20. Pouret EJM. Surgical technique for the correction of pneumo- and urovagina. Equine Vet J 1982;14:249–50. 21. Woodie B. Vulvar, vestibular, vagina and cervix. In: Auer JA, Stick JA (eds) Equine Surgery. Philadelphia: Saunders, 2006: pp. 835–54. 22. Embertson RM. Selected urogenital surgery concerns and complications. Vet Clin North Am Equine Pract 2009;24:643–61. 23. Beard WL. Standing urogenital surgery. Vet Clin North Am Equine Pract 1991;7:669–84. 24. Aanes WA. Surgical management of foaling injuries. Vet Clin North Am Equine Pract 1988;4(3):417–38. 25. Schumacher J, Schumacher J, Blanchard T. Comparison of endometrium before and after repair of third-degree rectovestibular lacerations in mares. J Am Vet Med Assoc 1992;200(9):1336–8. 26. Stickle RL, Fessler JF, Adams SB. A single stage technique for repair of rectovestibular lacerations in the mare. Vet Surg 1979;8:25. 27. Jalim SL, McKinnon AO. Surgical correction of rectovaginal fistuale in mares and susequent fertility. Aust Vet J. In press. 28. Colbern GT, Aanes WA, Stashak TS. Surgical management of perineal lacerations and rectovestibular fistulae in the mare: a retrospective study of 47 cases. J Am Vet Med Assoc 1985;186(3):265–9. 29. Aanes WA. Surgical repair of third-degree perineal laceration and rectovaginal fistula in the mare. J Am Vet Med Assoc 1964;144:485–91. 30. Hilbert BJ. Surgical repair of recto-vaginal fistulae in mares. Aust Vet J 1981;57:85–7. 31. Schoenfelder AM, Sobiraj AA. A vaginal mucosal pedicle flap technique for repair or rectovaginal fistula in mares. Vet Surg 2004;33:517–20. 32. McKinnon AO, Arnold KS, Vasey JR. Selected reproductive surgery of the broodmare, 1991;174:109–25. Sydney, Post Graduate Committee in Veterinary Science. Equine Reproduction: A seminar for veterinarians. 33. Adams SB, Benker F, Brandenburg T. Direct rectovestibular fistula repair in 5 mares. Proceedings of American Association of Equine Practitioners 1996;42:156–9. 34. McKinnon AO, Vasey JR. Selective reproductive surgery of the broodmare. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders ( Elsevier), 2007; pp. 146–60.
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Chapter 264
Cervical Surgery Patrick J. Pollock and Thomas M. Russell
Introduction Cervical laceration is a well-recognized cause of infertility in the mare and usually occurs at parturition. The cervix functions as the final and most craniad barrier to ascending infection, and separates the uterus from the vagina. It is tightly closed during diestrus and pregnancy due to the influence of progesterone. Damage to the cervix may lead to difficulty conceiving, or carrying a foal to term.
Anatomy The cervix is composed of a thick-walled muscular tube, approximately 5.0 to 7.5 cm in length, and 3.5 to 4.0 cm in diameter. The wall of the cervix contains a significant amount of collagenous connective tissue, and the sphincter consists of smooth muscle, which is an extension of the inner layer of the myometrium. Longitudinal folds of mucosa, which are extensions of the endometrial folds of the uterine body, run the length of the cervix and end at the caudal portion of the cervix within the vaginal vault. A fold of mucosa commonly extends ventrally, as a frenulum, from the external os to the floor of the vagina. Some mares also have a dorsal frenulum. The cervix is lined by ciliated epithelium and contains mucus-secreting cells. The internal pudendal artery gives rise to the vaginal artery, which is the main arterial supply to the cervix. A rich venous plexus drains into the internal pudendal veins.1 Neurological control comes through the vaginal nerve, from the pelvic plexus.2
Etiology Cervical lacerations are most commonly sustained during parturition. They may be associated with dystocia or induced parturition, but also occur with apparently normal births. Miller et al.3 reported that 86% of mares presented for the repair of a cervical injury had an apparently normal foaling. Assisted vagi-
nal delivery increases the risk of a tear, particularly when the soft tissue of the reproductive tract is traumatized during dystocia, or where inadequate lubrication is used.4, 5 In addition, cervical lacerations have been reported following copulation.6 Congenital cervical incompetence due to failure of embryological closure is also recorded.7
Clinical signs and diagnosis Cervical lacerations may result in endometritis and subsequent infertility. Large defects lead to failure to conceive, early embryonic death and early abortion, whereas smaller defects more commonly result in late-term abortion as the enlarging fetus places increasing pressure on an incompetent cervix.8 Mares presented with a history of abortion should be given a cervical examination. Diagnosis of cervical laceration can be a challenge. Immediately following foaling, cervical relaxation and the presence of edema and swelling usually obscure all but the most major defects; if the mare is in estrus or approaching estrus, edema and cervical relaxation may have the same effect.6 Infrequently at foaling, a cervical laceration will be found to extend craniad and laterad, entering the abdomen – such lacerations require immediate attention and repair. Most cervical lacerations are identified later and are best evaluated in diestrus because the increased cervical tone facilitates assessment of the degree of apposition of cervical folds and damage to the muscular layer. Any mare suffering from dystocia should be earmarked for vaginal examination in diestrus approximately 3 weeks post-foaling. Although inspection of the reproductive tract is a crucial part of the examination, visual inspection alone is unlikely to identify a cervical injury. Manual palpation is essential for accurate diagnosis. A sterile gloved and lubricated hand should be inserted into the tract and the thumb or index finger passed into the external
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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os of the cervix. Digital palpation should follow in a circumferential manner. This examination will identify defects in the fibromuscular layer, which are generally covered by mucosa making visual identification difficult. In most cases lacerations affect only the vaginal portion of the cervix; however, in some cases injuries may extend craniad into the uterine body. Unusually, there may be more than one defect present. These are best repaired on separate occasions 3–4 weeks apart, as intraoperative swelling of tissue makes repair of other tears more difficult.9 The position and extent of the laceration should be recorded, and this is usually accomplished using the clock-face analogy, where the most dorsal portion of the cervix is represented by the 12 o’clock position and the most ventral portion by the 6 o’clock position. Defects can then be noted to extend from one position to another. There is no recorded site around the circumference of the cervix that is predisposed to tearing.8
Selection of surgical candidates Not all cervical lacerations require surgical repair. Under the influence of progesterone the cervix reduces in size and its muscular tone increases, and as a result small tears may be of limited clinical significance during pregnancy. Reports suggest that only tears involving 50% or more of the vaginal cervix require surgical correction.10 In general, at our practices, only those mares with a cervical defect that progresses craniad to involve the junction of the external os with the vagina are candidates for surgery. In addition, only mares with obvious subfertility are usually considered, for example those with uterine infection or that have been barren for one or more years. Nevertheless, the potential economic significance of infertility or abortion may prompt a decision to repair a smaller cervical injury in order to increase the likelihood of the affected mare producing a live foal. Critical evaluation of the rest of the reproductive tract is vital to ensure that other causes of infertility such as pneumo- or uro-vagina are not present. In mares that have been barren for some time, evaluation with an endometrial biopsy is generally prudent.
Surgical treatment Optimally, surgery should be delayed for at least 30 days after injury, with some workers suggesting that up to 60 days may be best to allow traumatized tissue to heal and contract.6, 8 However, client pressure to quickly rebreed the mare may shorten the period between injury and repair. As noted above, the cervix under the influence of progesterone is much firmer, which allows much easier delineation of the defect. Wherever possible,
surgery should be carried out in diestrus, and the use of altrenogest may be beneficial in mimicking this state in selected mares.11 Alternatively, in order to facilitate early breeding, the mare can be bred and then the surgery carried out a few days later. This approach should be reserved for tears that are easily palpable in the estrus mare. Patient preparation and anesthesia Repair of cervical lacerations is most easily accomplished with the mare sedated and standing restrained in stocks. In many cases mares are presented as outpatients. Local anesthesia is preferred as this facilitates retraction of the cervix caudally.12 An infusion of 40–60 mL of 2% lidocaine is administered dorsally and laterally deep in the vaginal tissues around the cervix.12 Epidural anesthesia may be required in fractious mares or mares that are straining excessively. Very occasionally a mare may be so intractable as to require general anesthesia. Placing the mare in dorsal recumbency and elevating the hind end 30–45◦ using a hoist attached to the hind legs may achieve better visualization of tears repaired under general anesthesia. This technique may also be considered where the mare has a narrow vestibule, or to improve visualization of large ventral tears.11 It should be recognized that caudal reproductive tract surgery is inherently dangerous. Generally an indication of the mare’s temperament can be made during presurgical manipulation of the tract, and standing procedures abandoned before surgery commences, where deemed prudent. The perineal region is prepared for aseptic surgery in routine fashion. The tail is bandaged and tied to an overhead beam using a quick release knot. This removes the tail from the surgical field, and also provides support to the mare should she lose her footing due to excess sedation or epidural anesthesiainduced ataxia. The rectum is emptied of fecal material. All mares should be treated with non-steroidal anti-inflammatory drugs (NSAIDs) prior to surgery and perioperative antimicrobials are indicated due to the contaminated nature of the procedure. In addition to this, the tetanus status of all mares should be ascertained and appropriate tetanus prophylaxis initiated if indicated. Surgical equipment The most difficult aspect of cervical surgery in the mare is gaining adequate exposure. The use of modified Finochietto-type retractors, with elongated blades,6 greatly facilitates the procedure, as although the cervix can be retracted caudally, surgery is nevertheless performed in the depths of the tubular tract.
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Ideally the blades should be over 25 cm long, such that they reach craniad to the vestibulo-vaginal junction. Knowles cervical forceps are used, which have large teeth that facilitate vigorous caudal retraction without tearing out of the cervix. Rotation of the forceps can be used to provide better visualization and access to the tear. Using this instrument it is generally not necessary to use specially extended cervical instruments.12 However, Knowles forceps have been criticized as being overly traumatic,13 and stay sutures, placed 2 cm away from the wound edges using 5 metric suture material, may be employed instead. With these, long-handled instrumentation is essential and should include Mayo–Hegar needle holders, scalpel handle and number 10 blade, thumb forceps, Metzenbaum scissor, and two Allis tissue forceps. A disadvantage of stay sutures is the inability to easily rotate the margins of the tear to better facilitate dissection and suture placement. A good light source is essential, and can take the form of a fibreoptic source (an endoscope will work) that can be fixed to one of the retractor blades, or a forehead-mounted source, which is often superior.
closed first in a continuous horizontal mattress pattern, starting at the most craniad part of the laceration and moving caudally to the external os. This pattern everts the mucosa into the lumen of the cervix, which should be periodically digitally palpated to ensure that it remains patent. The muscular layer is closed next using the same size and type of suture material in a simple continuous pattern, and finally the outer mucosal (vaginal) layer is closed in the same manner. Some workers advocate that the muscular layer should be included in the closure of the final layer, in order to provide extra strength.8
Surgical procedure
One-layer closure
Three techniques have been described for closure of cervical defects – three-, two-, and one-layer closure. Whichever technique is chosen, repair starts with caudal retraction of the cervix, using Knowles uterine forceps, or stay sutures and Allis tissue forceps. The forceps are placed 1–2 cm either side of the lesion and used to apply caudal traction. An incision is made through the outer (vaginal) cervical mucosa along the edge of the defect. A strip of tissue is removed using Metzenbaum scissors along the length of the defect. This creates a V-shaped defect in the cervix that should extend approximately 0.5–1 cm craniad to the craniad extent of the defect to allow separation of the three distinct layers that make up the vaginal cervix: the outer cervical mucosa, the muscularis, and the inner cervical mucosa. The muscularis layer is the holding layer for suturing. The scar tissue is difficult to differentiate from the collagenous muscular layer, and care should be taken that the minimum of tissue is removed, so as not to compromise lumen diameter, or cause major changes in orientation of the cervix. Three or 3.5 metric (USB 2/0 or 0) monofilament absorbable suture material is used with a swaged-on cutting needle to allow easy penetration of the tough muscularis layer.
A one-layer closure is described and is the authors’ technique of choice. Initially the one-layer technique was reserved for mares repaired directly after
Three-layer closure Using the three-layer technique each tissue layer is apposed individually. The inner cervical mucosa is
Two-layer closure With the two-layer closure technique a simple continuous or continuous horizontal mattress pattern is used to close the inner cervical mucosa with incorporation of the cervical muscle layer.5, 6 A simple continuous pattern is then used to close the outer (vaginal) mucosal layer, again incorporating potentially the cervical muscularis (Figure 264.1).
Figure 264.1 With the two-layer closure technique a simple continuous or continuous horizontal mattress pattern is used to close the inner cervical mucosa with incorporation of the cervical muscle layer; a simple continuous pattern is then used to close the outer (vaginal) mucosal layer, again incorporating potentially the cervical muscularis.
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Mare: Female Urogenital Surgery can be turned out immediately following surgery and managed as normal. Some workers advocate infusing the uterus with lavage solutions containing antimicrobials immediately following repair and for several days thereafter.8 This may be especially important where the mare has been bred immediately prior to surgery. The cervix should not be palpated for 2 weeks, and breeding, when undertaken postsurgically, should be delayed until healing is complete (usually 4 weeks). As the repaired cervix is vulnerable to re-injury at breeding, artificial insemination is recommended where possible, and a breeding roll should be used where natural service is unavoidable.12
Complications
Figure 264.2 For the single-layer closure a simple continuous pattern is passed through the outer cervical (vaginal) mucosa and into the muscular layer. Suture bites are placed approximately 1 cm apart.
breeding;14 however, the technique has been successfully used at any time (A.O. McKinnon, personal communication). During one-layer closure a simple continuous pattern is passed through the outer cervical (vaginal) mucosa and into the muscular layer. Suture bites are placed approximately 1 cm apart (Figure 264.2).
Choice of technique To date there has been very little work critically evaluating which technique, if any, is superior. A large case series comparing three-layer closure with onelayer closure is currently being prepared by the authors, and suggests that one-layer closure results in a higher percentage of live foals than the three-layer technique (P.J. Pollock et al., unpublished data). Onelayer closure is also technically less demanding and can be accomplished more quickly than the three- or two-layer techniques. Whichever technique is selected, careful attention to detail is critical to success. This should include frequent palpation of the cervical lumen throughout the procedure to ensure that a suture bite has not crossed the lumen. Also, it may be necessary to trim excessive cervical mucosal folds that prolapse into the suture line, as this may predispose to fistula formation.
Aftercare Following surgery, antimicrobial and NSAID medication should be continued for 3 to 5 days. Mares
Complications following cervical surgery are unusual. Nevertheless a number of problems have been described. Adhesions within the cervical lumen can lead to stenosis, fertility problems, and potentially uterine infection. In rare cases this can lead to the development of a pyometra.15 However, in most cases adhesions can be easily broken down by digital palpation during re-evaluation 3–4 weeks after surgery, followed by application of a corticosteroid ointment topically once daily for several days. Where larger defects are repaired, the external os may be narrow and tight. Generally, if a lubricated hand can be inserted into the external os, the repair should be adequate.9 It seems that the more craniad portion of the repair is the most crucial barrier to ascending infection, and special care should be taken in this aspect of the repair. Caudally, overzealous apposition of tissue may create an external os that broadens craniad, leading to an inadequate seal in diestrus and pregnancy, whilst not relaxing sufficiently in estrus to allow adequate drainage of uterine contents.12 Despite the contaminated nature of the region, surgical site infections are rare. The most likely complication is failure to improve the fertility of the mare due to continued cervical incompetence.
Prognosis and implications for future fertility Critical evaluation of repair of cervical lacerations is difficult, as many mares with an incompetent cervix will conceive normally, and the production of a live foal is the only effective measure of surgical success. A limited number of studies have evaluated treatment and reported pregnancy rates of 62.5–75.3%.3, 10 These statistics are likely optimistically high as lateterm abortion is a problem associated with cervical lacerations, and foaling rates are expected to be lower than pregnancy rates.
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Cervical Surgery Before surgery, owners should be made aware that 40–50% of repairs fail during subsequent parturition. This is due to the inelastic nature of the scar tissue of the repair compared to the surrounding cervical musculature.9 A large number of other variables also affect success rates including the size and extent of the laceration; presence of more than one area of damage on the cervix; presence of adhesions or fibrous tissue; experience of the surgeon and the presence of other causes of infertility.
Cervical incompetence Occasionally the cervix is so badly damaged that repair as described is not possible. This may be due to the length or width of the defect or to extensive loss of tissue. Some mares overstretch the muscular layer of the cervix during foaling, leading to functional incompetence. Congenital cervical incompetence is also recorded.7, 16 In humans, cervical incompetence is of major concern17 and can result in late-term abortions similar to observations in mares. A variety of techniques have been employed in human medicine, including medical therapy with progesterone, cervical cerclage, and the placement of a cervical pessary,18–20 some of which may be applicable in the mare. Cervical cerclage involves the placement of one or two strands of a non-absorbable suture material (5 metric) or umbilical tape.4 The suture material is passed into the caudal cervical musculature ventrally through a stab incision in the mucosa, so that the needle enters the cervical muscle, and is guided by digital palpation such that the needle exits the cervix dorsally; the needle is then passed back into the cervical musculature and driven ventrally down the contralateral side of the cervix to exit lateral to the original entry site. The suture is then tightened and tied. On tightening, the lumen of the cervix should be obliterated. Care is taken not to penetrate the cervical lumen with the suture as this can lead to persistent uterine infection. Some authors have described the need to bury the suture; however, this is not performed in all instances, with minimal complications reported. The buried or purse-string suture is generally placed during the first 48 hours after breeding and ovulation and must be removed prior to foaling. It should be noted that this technique has never been critically evaluated in the mare. However, in humans placement of cervical cerclage suture laparoscopically is now a routine treatment,21 and it is possible that this treatment and others like it may have future applications in the mare.
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References 1. Sisson S. Equine urogenital system. In: Getty RWB (ed.) The Anatomy of the Domestic Animals, 5th edn. Philidelphia: Saunders, 1975; pp. 545–6. 2. Ley WB. Anatomy and physiology of the female reproductive system. In: Wolfe DF, Moll HD (eds) Large Animal Urogenital Surgery. Baltimore: Williams and Wilkins, 1999; pp. 81–86. 3. Miller CD, Embertson RM, Smith S. Surgical repair of cervical lacerations in Thoroughbred mares: 53 cases (1986–1995). Proceedings of the Annual Convention of the AAEP. 1996;43:154–5. 4. Evans LH, Tate LP, Cooper WL, Robertson JT. Surgical repair of cervical lacerations and the incompetent cervix. Proceedings of the Annual Convention of the AAEP. 1979;25: 483–6. 5. Moll HD, Kay KA. Cervical lacerations In: Wolf DF, Moll HD (eds) Large Animal Urogenital Surgery. Baltimore: Williams and Wilkins, 1999; pp. 111–13. 6. Aanes WA. Surgical management of foaling injuries. Vet Clin N Am-Equine Urogenital Surg 1988; 4: 417–38. 7. Blanchard TL, Evans LH, Kenney RM, Hurtgen JP, Garcia MC. Congenitally incompetent cervix in a mare. J Am Vet Med Assoc 1982; 181: 266–8. 8. Embertson RM, Henderson CE. Cervical tears. In: Samper JC, Pyock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. Saunders, 2007; pp. 130–3. 9. Embertson RM. Selected urogenital surgery concerns and complications. Vet Clin N Am-Equine 2009; 24: 643–61. 10. Brown JS, Varner DD, Hinricks K, Kenney RM. Surgical repair of the lacerated cervix in the mare. Theriogenology 1984;22:351–9. 11. O’Leary JM, Rodgerson DH. How to repair cervical tears using Trendelenberg position. Proceedings of the Annual Convention of the AAEP 2009;55:269–71. 12. McKinnon AO, Vasey JR. Selected reproductive surgery of the broodmare. In: Samper JC, Pyock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders, 2007; pp. 146–60. 13. Woodie B. Vulva, vestibule, vagina and cervix. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. St Louis: Saunders, 2006; pp. 852–3. 14. Foss RR, Wirth NR, Kutz RR. Post breeding repair of cervical lacerations. Proceedings of the Annual Convention of the AAEP. 1994;40:11–12. 15. Vaughan JT. Equine urogenital system. In: Jennings PB (ed.) The Practice of Large Animal Surgery. Philadelphia: Saunders, 1984; pp. 1122–50. 16. Allen WE. A cervical anomaly in an Arabian filly. Equine Vet J 1981;13:268–70. 17. Shortle B, Jewelewicz R. Cervical incompetence. Fertil Steril 1989;52:181–8. 18. Abenhaim HA, Tulandi T. Cervical insufficiency: reevaluating the prophylactic cervical cerclage. J Matern-Fetal Neo Med 2009;22:510–16. 19. Nelson L, Dola T, Tran T, Carter M, Luu H, Dola C. Pregnancy outcomes following placement of elective, urgent and emergency cerlage. J Matern-Fetal Neo Med 2009;22:269–73. 20. Dharan VB, Ludmir J. Alternative treatment for a short cervix: the cervical pessary. Semin Perinatol 2009;33:338–42. 21. Carter JF, Soper DE, Goetzl LM, Dorsten JP. Abdominal cerclage for the treatment of recurrent cervical insufficiency: laparoscopy or laparoscopy? Am J Obstet Gynecol 2009;201:111.e1–4.
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Chapter 265
Ovariectomy Dwayne H. Rodgerson and Dawn A. Loesch
Equine ovariectomy involves the removal of one or both ovaries. Numerous indications exist for performing unilateral and bilateral ovariectomy. Ovarian pathology is the most common indication for unilateral and rarely, bilateral ovariectomy. Ovarian pathology potentially warranting an ovariectomy includes ovarian neoplasms (Figure 265.1a,b), ovarian abscess, and ovarian cysts. Ovarian neoplasms (granulosa-theca cell tumor, teratoma, cystadenoma) are the most common form of ovarian pathology observed. Ovarian cysts are uncommon in mares, and generally do not affect fertility when present.1 Abscessation of the ovary is very uncommon and may occur iatrogenically following aspiration of a follicle or cyst. Indications for bilateral ovariectomy in mares include behavioral modification, the preparation of mount mares for semen collection and embryo transfer recipients, sterilization for breed registrations, and research purposes requiring ovariectomized mares. Behavioral modification is pursued due to problems observed during estrus. Hormonal therapy can be used to help alleviate problems associated with estrus; however, the expense and management difficulty of daily administration often prompts owners to pursue more permanent means of behavioral modification. If previous hormonal therapy has been successful in improving the mare’s behavior and/or performance favorably, then bilateral ovariectomy is likely to be successful. Chronic colic associated with ovulation has been infrequently reported, and may be eliminated with bilateral ovariectomy.2, 3 Equine ovariectomy is a commonly performed elective surgical procedure, and was first described prior to the twentieth century.4 One of the first approaches described was the colpotomy technique using physical restraint.5 Advances in equine general anesthesia techniques allowed numerous abdominal approaches for ovariectomy to be described. Ovariectomy in the standing mare is commonly performed
today due to advances in chemical restraint and the use of laparoscopic equipment. The described abdominal approaches for equine ovariectomy include a vaginal or colpotomy, flank, diagonal (oblique) paramedian, ventral midline, and laparoscopy. The decision as to which approach to use for any particular case depends on numerous factors including: specific indications for ovariectomy, the size of the affected ovary, the individual surgeon’s preference, financial constraints imposed by the client, the temperament of the mare, available equipment, and client expectations.6 An understanding of the benefits and disadvantages of all approaches will aid the decision-making process in selecting the appropriate surgical approach for each case.
Preoperative considerations Special preoperative preparation of the mare should be instituted prior to performing an ovariectomy.6 Food should be withheld for 12 to 24 hours prior to surgery to help decrease the amount of ingesta and gas within the gastrointestinal tract, making it easier to exteriorize the ovary and incisional closure. Laparoscopic techniques require that food be withheld for 12 to 48 hours prior to surgery, to improve visualization of intra-abdominal structures and to decrease the likelihood of penetrating a viscus when the laparoscopic instruments are introduced into the abdomen.7–9 The rationale for and use of systemic antibiotics to treat horses undergoing ovariectomy varies among surgeons. If antibiotics are used, they should be administered prior to surgery to ensure that adequate systemic concentrations are present at the time of surgery. Administration of broad-spectrum antibiotics should be continued postoperatively if a break in aseptic technique occurred during the surgery. Tetanus prophylaxis should also be administered routinely when performing ovariectomy.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 265.1 Ovarian neoplasia. (a) A granulosa theca cell tumor. (b) Cut section of a granulosa theca cell tumor. Note the cystic appearance of the ovary.
Vaginal ovariectomy approach (colpotomy) In 1903, Williams was the first to describe a vaginal approach, or colpotomy, utilizing an e´ craseur to ovariectomize mares.5 Advantages to the colpotomy technique include: the fact that it is performed as a standing procedure, minimal instrumentation is required, the procedure can be performed quickly, excellent cosmetic results can be expected, and there is a relatively short convalescent period following surgery. The vaginal approach is commonly used to perform bilateral ovariectomy in normal mares, but unilateral removal of a suspected ovarian tumor less than 8 to 10 cm in diameter can also be performed using this approach.6 Mares should be in diestrus or anestrus, as ovarian vasculature is believed to be minimized at these times, and there should be no ongoing infection involving the uterus or urinary bladder at the time of surgery.10 Mares are restrained in standing stocks and sedated. Sedation is most commonly accomplished with an α 2 -agonist and butorphanol, to provide analgesia as well as profound sedation. Acepromazine can be administered for tranquilization as well. Administration of a caudal epidural anesthesia is not necessary, but is recommended by some surgeons to help prevent the mare from straining during the procedure. The tail is wrapped and secured away from the perineal region, and all feces should be evacuated from the rectum to prevent contamination of the external genitalia. Routine aseptic preparation of the external genitalia and perineal region is performed and the vagina is lavaged with sterile saline or dilute povidone iodine solution. In some mares, catheterization of a large urinary bladder may facilitate the pro-
cedure and may help reduce the risk of inadvertent injury during the procedure. The location of the initial incision in the cranial fornix of the vagina is very important. The incision must be placed in either a craniodorsal (at the 2 or 10 o’clock) or a cranioventral (at the 4 or 8 o’clock) position.6 Potential complications of a misplaced incision include entering the rectum if the incision is placed too dorsal, injuring the urethra or bladder if placed too ventral, and incising the caudal uterine branch of the urogenital artery if too medial or lateral (at the 3 or 9 o’clock position). Location of the caudal uterine branch of the urogenital artery can be easy identified by feeling for an arterial pulse at the 3 or 9 o’clock position. The incision should be started 3 to 5 cm caudal to or away from the os cervix to avoid disruption of the cervical musculature. Using closed scissors or forceps, a small 1 to 2 cm vaginal incision is bluntly created. This incision must be enlarged by opening the scissor blades to their maximum separation whilst withdrawing them. Failure to do this almost always results in an inability to identify the original incision. A scalpel blade attached to a suture can be used to create the incision through the mucosa. The risks of inadvertently incising the caudal uterine branch of the urogenital artery or arterial branch are potentially greater with the scalpel blade. The vaginal incision is then bluntly enlarged digitally and the peritoneum is perforated to allow the surgeon’s hand to enter the abdomen. Perforation of the peritoneum can be difficult in some horses and may require long-handled instruments to bluntly incise the peritoneum. The ovary and associated mesovarium are accurately isolated by direct manual palpation. While the ovary is identified, ensure the ovary is directly palpated and not covered by mesentery
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from local bowel. Anesthesia of the mesovarium can be performed, but is not necessary. Anesthesia of the mesovarium can be attempted using gauze sponges soaked with local anesthetic (mepivacaine or lidocaine), which are held around the mesovarium for 30 seconds to 2 minutes. To prevent inadvertent loss of the sponges within the abdomen, a long suture or strand of umbilical tape should be secured to the sponges.11–13 To remove the ovary from the mesovarium, a chain e´ craseur is used to slowly crush and transect the mesovarium. The chain e´ craseur should be placed carefully around the mesovarium only, ensuring exclusion of bowel, intestinal mesentery, or a portion of the uterine horn. Once the chain is properly positioned around the mesovarium, it is tightened using the ratchet system of the e´ craseur. Hemostasis of the mesovarium depends on crushing the vessels and secondary vasoconstriction of the vessels. While the e´ craseur is being tightened around the mesovarium, the ovary should be securely held within the surgeon’s hand to prevent its loss within the abdomen. However, a recent article indicated the leaving the ovary of juvenile fillies in the abdomen after transection of the mesovarium did not result in revascularization of the ovary within the abdominal cavity.14 After the mesovarium has been transected and the ovary is removed from the abdomen, the surgeon should insert a hand back into the abdomen and feel for any evidence of an arterial bleed. If the mesovarium is bleeding after the ovary is transected, the mare should be monitored closely postoperatively for signs of hypovolemic shock for 24 to 48 hours. Replacement of the e´ craseur or ligation of the pedicle is difficult through a vaginal approach. The contralateral ovary can be removed through the same vaginal incision if a bilateral procedure is to be performed. The vaginal incision can be left open to heal by second intention or sutured closed. An episioplasty (Caslick procedure) should be performed to complete the surgery, in an effort to decrease the risk of ascending infection and potentially prevent evisceration.10 Tie stall restriction for 2 to 7 days postoperatively has been recommended to prevent recumbency and reduce the risk of evisceration of bowel.12 Exercise restriction to either a stall or small paddock can be employed for 2 to 3 weeks following surgery, depending on how the vaginal incision is healing. The primary disadvantage to the colpotomy approach is the lack of visualization.8 By digital palpation alone, the ovary must be differentiated from and isolated from the omentum, local mesentery, loops of intestine, and fecal balls within the small colon.6 Hemorrhage from the mesovarium may be difficult to determine and control due to the lack of visualization. If hemorrhage from the mesovarium persists, hypovolemic shock could develop. Control of the
hemorrhage can be controlled using medical management of hypovolemic shock in some case or laparoscopically (see laparoscopic techniques). Postoperative administration of broad-spectrum antibiotics may be indicated in mares undergoing a vaginal approach, because it is difficult to adequately prepare the vagina for aseptic surgery.6 This approach is not recommended in mares which pool urine or have ´ vaginal, cervical, or uterine infections.4, 6 Ecraseur transection of large ovaries (greater than 8 to 10 cm in diameter) should not be attempted due to the size of vaginal incision needed to remove the ovary and the potential for enlarged vascular supply associated with larger ovaries.10 Even with adequate restraint and heavy sedation, this approach poses a risk to the surgeon, due to positioning behind the mare. Thus, the temperament of a mare must be considered before doing an ovariectomy using a vaginal approach.
Flank ovariectomy approach The flank ovariectomy approach can be performed with the mare in standing or recumbent position. As with the standing colpotomy approach, the mare’s temperament must be suitable for standing surgery. When performing the standing flank approach, the mare is sedated, restrained in standing stocks, and the tail is wrapped and secured away from the surgical site.6 For the recumbent flank ovariectomy technique, mares are placed in lateral recumbency so that the ovary to be removed is uppermost. The recumbent flank technique is generally only recommended or chosen for unilateral ovariectomy. The recumbent flank approach does require general anesthesia, thus resulting in a slightly greater cost to the client and potentially increased risk to the mare. Compared to the vaginal ovariectomy approach, both the standing and recumbent flank laparotomy approaches enable the surgeon to remove larger ovaries and give better exposure of the mesovarium, thus potentially providing superior hemostasis.6 In the horse, the limited size of the paralumbar fossa (compared to the bovine) and the thickness of the body wall in the flank region may limit the ability to easily exteriorize large ovaries. It has been recommended that ovaries up to 15 cm in diameter can be removed easily through a flank approach.15 Bilateral ovariectomy typically requires a second incision through the opposite paralumbar fossa to remove the contralateral ovary. Bilateral ovariectomy can be achieved standing through a single flank incision, but the contralateral ovary must be transected blindly within the abdomen using a chain e´ craseur. The other ovary can be identified easily by passing under the small colon.
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Ovariectomy For either standing or recumbent techniques, the paralumbar fossa is clipped, aseptically prepared, and draped. Regional anesthesia in the form of an inverted ‘L’-block or local infiltration (line block) of the proposed incision site is required in standing mares. The skin incision is started 8 to 10 cm ventral to the lumbar transverse processes between the last rib and the tuber coxae, and is extended ventrally 8 to 15 cm as needed, depending on the size of the ovary to be removed. After incising the skin and subcutaneous tissues, a grid or modified grid approach may be used to pass through the abdominal musculature. The grid approach creates a slightly smaller opening into the abdominal cavity, and should only be used for ovaries less than 10 cm in diameter; larger ovaries may require a modified grid technique. A grid approach involves separating the fibers of the external abdominal oblique muscle, whereas a modified grid technique involves incising this muscle along the line of the skin incision.6 For both grid techniques, the internal abdominal oblique and transversus abdominis muscles are bluntly separated along the direction of their fibers to expose the peritoneum. The peritoneum may be incised or bluntly perforated to allow access to the abdomen. The authors prefer to separate the muscle fibers and bluntly penetrate the peritoneum using digital manipulation. The ovary is then identified and isolated by digital palpation. In standing mares, the mesovarium is anesthetized by topical administration of anesthetic, similar to that described for the colpotomy procedure, or by direct injection of anesthetic into the mesovarium. Direct injection of the anesthetic into the mesovarium provides better anesthesia and will facilitate movement and exteriorization of the ovary. The mesovarium may be transected within the abdomen or after exteriorization. Aspiration of cystic cavities within the ovary may help reduce the overall ovarian size to facilitate exteriorization. The means to achieve hemostasis and transection of the mesovarium will depend on the ability to exteriorize or visualize the mesovarium through the flank incision. Possible choices include the use of a chain e´ craseur, emasculator, surgical staR pling device, transfixing ligatures, or the LigaSure 10, 16 device (Valleylab, Colorado). Following removal of the ovary, the mesovarium is observed for hemorrhage and oversewn if desired, when adequate visualization permits. Oversewing the mesovarium with an absorbable suture material may possibly reduce hemorrhage and potential intra-abdominal adhesions of a segment of bowel to the transected mesovarium.4, 15 The flank incision should be lavaged and closure of the flank laparotomy incision is routine. Postoperative pain may be observed with larger flank incisions. Incisions created in a more ventral position on the flank tend to develop more swelling and cause increased postoperative pain and discomfort.2
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Incisions in the flank have been associated with incisional discharge and partial to complete skin dehiscence. Exercise should be restricted to hand walking or turnout in a small paddock for 2 to 4 weeks following surgery, after which light riding and a slow return to normal work may be allowed, provided the incision has healed without complications. Occasionally, this approach results in a poor cosmetic outcome if scarring of the incision site occurs;6 however, the authors have experienced very good cosmetic results with flank incisions.
Diagonal (oblique) paramedian ovariectomy approach The diagonal (oblique) paramedian approach has been reported to be superior to other ovariectomy approaches, especially for the removal of ovaries up to 25 cm in diameter.17 The diagonal (oblique) paramedian approach is believed to decrease the amount of traction placed on the mesovarium due to incision being created so close to the intra-abdominal position of the ovary. In addition, the abdominal wall is thinner at this location, as compared to the flank approach, which allows greater flexibility in retracting the wound edges.17 Improved visualization of the ovary and mesovarium can often be achieved with this approach. Bilateral ovariectomy can be performed; however, two incisions are generally required. General anesthesia is necessary to perform an ovariectomy with this approach, therefore the cost and potential risks are increased. Perioperative preparation and care are similar to that for the previously described approaches. Following induction of general anesthesia, the mare is placed in dorsal recumbency with the hindlimbs flexed and maximally separated. The mare should be slightly tilted towards the side opposite the affected ovary.17 This can help visualization at the incision and minimize the tendency for bowel to protrude from the incision. Following routine aseptic preparation of the ventral abdomen, the abdomen is draped accordingly. A skin incision is started approximately 5 to 10 cm cranial to the ipsilateral mammary gland and extended approximately 20 cm (or as needed, depending on the size of the affected ovary) cranially and laterally toward the fold of the flank.6, 17 The incision is extended through the underlying external rectus sheath, parallel to the initial skin incision. The rectus abdominis, internal rectus sheath, and peritoneum are bluntly divided in the direction of their fibers or incised, depending on surgeon preference, to allow access to the abdomen. The ovary is located and exteriorized through the incision. Aspiration of the fluid within cystic cavities of diseased ovaries can be performed to reduce the size of the ovary
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and facilitate its removal from the abdomen. Large suture material placed as stay sutures within the ovary and retraction on the incision edges can help in exteriorizing larger ovaries. Local anesthesia of the mesovarium, either by topical application of anesthetic-soaked gauze sponges or direct injection of local anesthetic, may be used to diminish the pain response secondary to traction on the mesovarium. Overlapping transfixation ligatures of #2 absorbable suture materials or application of a TA-90 stapling device may be used for hemostasis prior to transection of the mesovarium.5, 17 Ligatures should be tightened or staples should be applied to the mesovarium in a relaxed position; application under tension may result in failure to provide adequate hemostasis.12, 16 Following complete transection of the mesovarium, closure of the abdominal incision is routine. Due to the location of the incision on the ventral abdomen, a good cosmetic outcome usually follows ovariectomy by the diagonal (oblique) paramedian approach.17 Exercise restriction during the early postoperative period should consist of stall confinement with controlled hand walking for the first 2 to 4 weeks following surgery. After this time, small paddock turnout may be allowed for an additional 2 to 4 weeks of confinement. Return to a full exercise schedule depends on how the incision is healing and generally is around 60 to 90 days. Ventral midline A ventral midline ovariectomy approach is generally reserved for large ovaries. The ventral midline incision can easily be extended as necessary, depending on the individual case, making it the technique of choice for removal of extremely large ovaries.4, 16 The ventral midline approach is generally used to remove very large granulosa theca cell tumors, but can also be used to perform a bilateral ovariectomy. Perioperative preparation and care are similar to that for previously described approaches. For a ventral midline technique, general anesthesia is required, and the mare is placed in dorsal recumbency. A routine ventral midline linea skin incision is created, beginning at the mammary gland and extending cranially 25 to 35 cm as needed for adequate exposure. The linea is incised and the peritoneum is bluntly penetrated. The ovary is located by digital palpation and exteriorized. Local anesthesia of the mesovarium can be used if desired to decrease the pain response related to traction on the mesovarium. Intraoperative hypotension believed to cause myopathies and neuropathies have been associated with excessive traction on the mesovarium, and these complications may be reduced by the application of local anesthetic.6 Hemostasis and transection of the mesovarium is similar to that described for the flank and
diagonal paramedian approaches. Closure of the ventral midline incision is routine. Postoperatively, care is similar to the other ovariectomy approaches. Stall confinement with hand walking should be enforced for the first 2 to 4 postoperative weeks, at which time sutures or staples used to close the ventral incision should be removed. Small paddock turnout may be allowed after this, for an additional 2 to 4 weeks of confinement, after which light riding or pasture turnout may be allowed. Full exercise should not be allowed for a minimum of 90 to 120 days after surgery, depending on how the incision is healing. A good cosmetic outcome is expected with this approach, provided no complications with incisional healing occur.
Paramedian The paramedian approach can be used to remove large pathological ovaries or to perform bilateral ovariectomy through one (contralateral ovary must be transected blindly) or two incisions.18 The paramedian approach is similar to the ventral midline approach, except for the location of the incision. The incision is made 4 to 8 cm lateral to midline and extends cranially from the level of the mammary glands for 25 to 35 cm or as needed for adequate exposure.6, 15, 18 Methods to exteriorize the ovary and provide hemostasis of the mesovarium are similar to those described for other approaches. Postoperative care following a paramedian celiotomy is similar to the ventral midline and diagonal paramedian approaches.
Laparoscopic techniques Laparoscopic ovariectomy techniques are becoming more popular as a method of decreasing surgical morbidity rates associated with elective ovariectomy.2, 7, 8, 19, 20 Laparoscopic ovariectomy techniques provide direct viewing of the target tissue and allow for tension-free ligation and transection of the mesovarium. Bilateral laparoscopic ovariectomy techniques have been described for mares in the standing and dorsally recumbent positions.2, 7, 8, 19, 20 Unilateral enlarged pathological ovaries can be removed using laparoscopic techniques.19, 21 Laparoscopic techniques can greatly improve visualization of the ovary and mesovarium, potentially decrease post-surgical complications, and can provide tension-free ligation of the vessels within the mesovarium.2, 7, 8, 19, 20 However, important considerations when performing equine laparoscopic ovariectomy include the requirement for specialized equipment, the technical difficulty of certain procedures, and the potential for anesthetic complications in horses placed in
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Ovariectomy Trendelenberg position. Additionally, the use of laparoscopic equipment may be prohibitively costly in certain situations. Laparoscopic ovariectomy in the standing mare Laparoscopic ovariectomy in the standing mare avoids the need for general anesthesia, eliminates the cardiovascular derangements associated with the Trendelenberg position, and shortens the required preoperative fasting time (12 hours instead of 24 to 48 hours).6 Mares are sedated using detomidine hydrochloride (0.02–0.03 mg/kg, i.v.) alone or in combination with butorphanol tartrate (0.02–0.05 mg/kg, i.v.). Both paralumbar fossae are prepared for aseptic surgery and draped in cases requiring a bilateral ovariectomy. The paralumbar fossa is anesthetized using regional anesthesia or by direct infiltration of the proposed laparoscope and instrument portal sites. The abdominal cavity can be insufflated with carbon dioxide through either a Verres-type needle or 15 cm VersaportTM inserted dorsally in the paralumbar fossa or a teat cannula inserted ventrally, as if performing abdominocentesis.6 To avoid the potential complication of inadvertently insufflating the retroperitoneal space, the authors prefer to pass the trocar-cannula through the paralumbar fossa prior to insufflation of the abdomen cavity. A 15 mm skin incision is made at the dorsal border of the internal abdominal oblique muscle, and the laparoscopic trocar-cannula is introduced into the abdominal cavity perpendicular to the paralumbar fossa.20, 22 The trocar is replaced by the laparoscope, and the caudal portion of the abdomen is examined to identify the ovary. The first instrument portal is made 4 to 8 cm ventral to the laparoscope portal and a second instrument portal is made 4 to 8 cm ventral to the first.2, 7, 20 Trocar-cannula units are passed through each instrument portal perpendicular to the flank musculature. Using a laparoscopic injection needle placed through a cannula, the mesovarium can be infiltrated with local anesthetic. Hemostasis of the mesovarium can be achieved using suture ligatures, staples, laser energy, or electrosurgical instrumentation.2, 7, 20 The ovary is transected using laparoscopic scissors distal to the site of ligation or coagulation. The ovary is removed by enlarging one of the instrument portals or by connecting the two instrument portals. After removing the ovary, the abdomen is deflated through a laparoscopic cannula. The superficial abdominal fascia and skin at the portals are closed separately. The same procedure is then performed through the opposite paralumbar fossa to remove the contralateral ovary in bilateral procedures. Recently, the authors have been using a small flank
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incision to remove both ovaries. The first ovary transected is held in laparoscopic forceps while the contralateral ovary is transected. The surgeon’s hand removes the ipsilateral ovary and then reaches under the caudal small colon to remove the contralateral ovary through one incision. This eliminates having two larger incisions in each paralumbar fossa. Laparoscopic ovariectomy in the anesthetized mare To perform unilateral or bilateral ovariectomy in an anesthetized mare using laparoscopy, the horse is anesthetized, positioned in dorsal recumbency, and the tail is secured to the surgery table.6 The caudal abdomen is aseptically prepared and draped accordingly. To improve visualization of the caudal abdomen, a urinary catheter can be passed to decompress the urinary bladder.23 A 10 mm skin incision is made just cranial or lateral to the umbilicus.8, 19, 23 The abdomen is insufflated with carbon dioxide through a teat cannula to a pressure of approximately 15 to 20 mmHg. A laparoscopic trocar-cannula unit is introduced into the abdomen, and the trocar is replaced with the laparoscope. In the recumbent technique, patient positioning becomes important for adequate visualization of the caudal abdomen. In routine dorsal recumbency, the female reproductive tract is obscured by intestinal viscera, so the surgical table must be elevated in such a way that the mare’s head is lower than the hindquarters. For removing ovaries, an angle of inclination of approximately 30◦ from horizontal (Trendelenberg position) is generally required.8, 19, 23 Two instrument portals (cranial and caudal) are created on both the left and right ventral abdomen. The cranial instrument portals are located midway between the ipsilateral mammary gland and the umbilicus. The caudal instrument portals are placed midway between the ipsilateral mammary gland and the cranial instrument portal. After creating the four instrument portals, the ipsilateral uterine horn is elevated using a Chambers catheter inserted through the contralateral caudal instrument portal. A knot push rod equipped with a modified Roeder knot or a commercial suture loop is inserted through the ipsilateral caudal instrument portal (left caudal instrument portal for removing a left ovary). Sharp-toothed laparoscopic grasping forceps are passed through the cranial instrument portal on the same side as the suture loop. The jaws of the forceps are passed through the suture loop and used to grasp the ovary. The suture loop is passed over the ovary and tightened around the mesovarium. The mesovarium is then transected distal to the suture ligature. The transected ovary is maintained in the jaws of the grasping forceps while the same series of
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steps are reversed, to allow removal of the opposite ovary. The abdomen is deflated and the ovaries are removed by enlarging one of the cranial instrument portals. For the enlarged incision, the external fascia of the rectus abdominis, subcutaneous tissue, and skin are closed separately. All remaining incisions are closed by simply apposing the skin. Because most laparoscopic ovariectomy techniques can be accomplished through small incisions, the mares can be returned to exercise shortly after performing the procedure.6 Postoperatively, mares should be confined to a stall for the first 24 hours followed by stall or small paddock confinement for 2 weeks before they are returned to unrestricted exercise. Laparoscopy can be used to remove large ovarian granulosa theca cell tumors. One report described two mares which had a granulosa theca cell tumor removed using the recumbent laparoscopic ovariectomy technique.19 The mares were placed in Trendelenberg position and the maximum diameters of the ovaries were estimated to be 20 cm in diameter. Hand-assisted laparoscopic ovariectomy A hand-assisted laparoscopic ovariectomy technique in standing mares can be used to remove granulosa theca cell tumors.21 The standing, hand-assisted, laparoscopic ovariectomy technique is technically easy to perform, can be used for large pathological ovaries, allows for accurate placement of the staple line, tension-free ligation and transection, and eliminates the potential risks and costs associated with general anesthesia. The following descriptions describe the series of steps involved with this ovariectomy technique. This is the authors’ preferred technique to remove pathological ovaries in mares. Prior to surgery, a transrectal examination of the cranial portion of the reproductive tract and transrectal ultrasonographic examination of the affected ovary should be performed. Feed is withheld at least 12 to 24 hours prior to performing unilateral ovariectomy. Mares are administered non-steroidal antiinflammatory agents and broad-spectrum antibiotics. Mares are placed in stocks and sedated with detomidine (2.5 to 5 mg) or a combination of detomidine and butorphanol (10 mg, i.v.). The paralumbar fossa is clipped and aseptically prepared similar to the traditional flank ovariectomy approach. A 12 to 20 cm vertical line in the middle of the paralumbar fossa is infiltrated with 2% mepivacaine (total volume ranged from 40 to 60 mL). After infiltration of the local anesthetic, the paralumbar fossa is draped accordingly. A 12 to 20 cm vertical skin incision is made 3 to 6 cm ventral to the dorsal border of the internal abdominal oblique muscle. The external abdominal oblique
Figure 265.2 Hand-assisted laparoscopic ovariectomy technique. The mesovarium is being injected with local anesthesia with an injection needle. The position of the needle within the abdomen is confirmed both digitally and laparoscopically.
muscle was sharply transected, and the internal abdominal oblique and transverse abdominal muscles are divided in the direction of their fibers. The peritoneum is bluntly entered and the peritoneal opening is enlarged to allow a hand to pass into the abdominal cavity. The laparoscope position is positioned dorsal to the flank incision. The laparoscope cannula is positioned through the dorsal aspect of the flank incision or through a separate skin incision dorsal to the flank skin incision. With a hand in the abdomen to guard against trauma, a 10 mm trocar-cannula unit is inserted through the paralumbar fossa perpendicular to the flank musculature. By using the laparoscope, the ovary was identified slightly caudal to the flank incision. Infiltration of 2% mepivacaine into the mesovarium is performed by use of a laparoscopic injection needle inserted through the flank incision (Figure 265.2). Digital massage of the mesovarium facilitates accurate infiltration and even distribution of local anesthetic within the mesovarium. While an assistant manipulates the laparoscope, hemostasis of the mesovarium is achieved by use of an abdominal stapling device (TA-90 or TA-55 stapling device) (Figure 265.3). This is performed by passing the jaws of the stapling instrument into the abdominal cavity while the handle of the instrument remains outside the abdomen. The accurate positioning of the stapling instrument is facilitated by digital manipulation of the mesovarium. The stapling device is placed from a caudal to cranial direction between the uterine horn and ovary. If the stapling device can encompass the entire mesovarium, only one application of the stapling device is performed. If the stapling instrument does not completely cover the length of the mesovarium, a second application of the stapling device is placed in a cranial to caudal direction to completely
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Ovariectomy
Figure 265.3 Hand-assisted laparoscopic ovariectomy technique. The mesovarium is being stapled using a TA-90 abdominal stapling device. The position of the stapling device within the abdomen is confirmed both digitally and laparoscopically.
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Removal of the ovary is achieved by enlarging the incision if needed or using three methods to exteriorize the ovary: direct manual removal through the flank incision, manual removal after aspiration of cystic fluid within the ovary, or placement of the ovary within a sterile plastic bag. Placement of the ovary in a sterile plastic bag is achieved efficiently with two hands in the abdomen. This is easier for individuals with smaller hands. Once the ovary is in the bag, the free edges of the bag are exteriorized to pull the ovary up to the level of the incision (Figure 265.4a). Tension on the free edges of the bag retracts the incisional edges and the ovary is sharply incised with a scalpel blade (Figure 265.4b). Release of the cystic fluid allows decompression of the ovary and facilitates exteriorization. The flank incision should be lavaged and closure of the flank laparotomy incision is routine. Postoperatively, the treatment regimen and complications are similar to those described for traditional flank approaches.
staple the mesovarium. Prior to the final application of the stapling device, the mesovarium and ovary are visually and digitally inspected to ensure that no intestinal or mesenteric entrapment developed during placement of the stapling instrument. After application of the staple instrument, it is removed and laparoscopic scissors are inserted through the flank incision. By use of digital manipulation of the mesovarium, the mesovarium is accurately transected between the ovary and staple line. The mesovarium can R also be coagulated and transected using a LigaSure 24 device. After the ovary is completely transected from the mesovarium, the scissors are removed and the mesovarium is inspected for signs of hemorrhage. If hemorrhage is detected from the mesovarium, hemostasis is achieved by use of hemoclips, a bipolar electrosurgical instrument, or vessel-sealing device.
Ovariectomy has been associated with greater postoperative morbidity and mortality than are seen with other elective procedures.6 Postoperative hemorrhage from the mesovarium can occur if hemostasis of the mesovarium fails.25 Intra-abdominal hemorrhage from branches of the ovarian artery is a serious and possibly fatal complication that may go undetected at the time of surgery. Therefore, mares should be confined to a stall for the first 24 hours after surgery. Clinical signs associated with blood loss include tachycardia, pale mucous membranes, weakness or ataxia, weak thready pulse, and poor jugular distension. Hemorrhage must be controlled,
(a)
(b)
Complications
Figure 265.4 Hand-assisted laparoscopic ovariectomy technique. (a) The transected ovary is within a sterile plastic bag and brought up into the incision. (b) The ovary is then incised to allow drainage of the cystic fluid. This facilitates removal of large ovaries.
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and initial medical therapy should be aimed at replacing the lost blood volume by intravenous administration of fluids and whole blood. Surgical control of hemostasis can be performed laparoscopically utilizing electrosurgical devices, vessel sealing devices, or ligatures.25 The vaginal ovariectomy approach also has the potential risk of inadvertently incising the caudal uterine branch of the urogenital artery when making the incision into the abdomen. Care to avoid the 3 and 9 o’clock positions and using blunt Mayo scissors instead of a scalpel blade will help avoid this potentially fatal complication. Other potential complications reported when performing a vaginal approach include pain and discomfort, injuries to the cervix, bladder or a segment of bowel, delayed vaginal healing, peritonitis, eventration of bowel, incisional site hematoma or abscess, intra-abdominal adhesions to the vagina, and chronic lumbar or bilateral hindlimb pain. Reported complications with the other celiotomy approaches vary depending on position. Intraoperative hypotension, myopathies, and neuropathies have been associated with traditional ovariectomy approaches performed under general anesthesia to remove granulosa theca cell tumors. Tension placed on the mesovarium during the process of exteriorizing an ovary is speculated to cause a decrease in arterial blood pressure and potentially lead to inadequate peripheral circulation.26 Therefore, careful attention should be paid to arterial pressure and measures taken to ensure adequate arterial pressure under general anesthesia. Cardiopulmonary derangements can potentially develop during laparoscopic procedures with horses placed in Trendelenberg position.8, 22 This positioning exaggerates the force of abdominal insufflation and the weight of abdominal viscera on the diaphragm, decreasing the horse’s ability to adequately ventilate without mechanical assistance and potentially compromising venous return to the heart.8 Metabolic acid–base disturbances may also occur following prolonged abdominal insufflation with carbon dioxide, which diffuses easily into the systemic circulation.6 As with traditional surgical approaches, myopathies and neuropathies can be a consequence of prolonged dorsal recumbency.8 Therefore, proper anesthetic patient monitoring is required when performing laparoscopic ovariectomy in the dorsal recumbent position. Regardless of whether the surgery is performed standing or recumbent, postoperative pain, anorexia, depression, incisional swelling, incisional infections, incisional dehiscence, eventration, peritonitis, intraabdominal adhesions, and death have been reported following ovariectomy in the mare.6 A higher incidence of incisional complications have been observed
with approaches through the flank.27 This may be associated with the increased amount of dead space and muscle necrosis that may occur with the paralumbar approach.27 Incisional dehiscence of flank incisions is treated by daily cleansing of the wound and administration of systemic antibiotics. Proper aseptic techniques must be employed when performing ovariectomy procedures, or postoperative septic peritonitis may result.26 This potential complication can be prevented by adhering to proper aseptic technique throughout the procedure, and by the administration of perioperative antibiotics. Intra-abdominal adhesions can develop after ovariectomy;28 however, the use of proper aseptic technique and ensuring minimal trauma to gastrointestinal serosal surfaces can help prevent formation of adhesions. The surgical approach chosen to perform an ovariectomy is often dependent upon the surgeon’s preference. However, the decreased morbidity associated with laparoscopy has made laparoscopic ovariectomy the preferred surgical approach by many surgeons. Due to client education, many clients are also requesting their horses undergo laparoscopic procedures over traditional surgical procedures. An understanding of all described approaches will hopefully aid in making the correct surgical recommendations.
References 1. Meyers PJ. Ovary and oviduct. In: Kobluk CN, Ames TR, Geor RJ (eds) The Horse: Diseases and Clinical Management. Philadelphia: Saunders, 1995; pp. 998–1008. 2. Hanson CA, Galuppo LD. Bilateral laparoscopic ovariectomy in standing mares: 22 cases. Vet Surg 1999;28:106– 12. 3. Hooper RN, Taylor TS, Varner DD, Blanchard TL. Effects of bilateral ovariectomy via colpotomy in mares: 23 cases (1984–1990). J Am Vet Med Assoc 1993;203:1043–6. 4. Colbern GT. Ovariectomy. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 450–5. 5. Williams WL. Surgical and Obstetrical Operations. Ithaca, NY; 1903. 6. Loesch DA, Rodgerson DH. Surgical approaches to ovariectomy in mares. Comp Cont Educ Pract Vet 2003;25:862–71. 7. Palmer SE. Standing laparoscopic laser technique for ovariectomy in five mares. J Am Vet Med Assoc 1993;203: 279–83. 8. Ragle CA, Schneider RK. Ventral abdominal approach for laparoscopic ovariectomy in horses. Vet Surg 1995;24:492– 7. 9. Rodgerson DH, Johnson CR, Belknap JK. Laparoscopic ovariectomy in mares by using electrocautery. Proc Am Assoc Equine Pract 1998;44:302–3. 10. Moll HD, Slone DE. Surgery of the ovaries. In: Wolfe DF, Moll HD (eds) Large Animal Urogenital Surgery. Williams & Wilkins: Baltimore, 1997; pp. 137–41.
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Ovariectomy 11. Trotter GW, Embertson RM. Surgical diseases of the cranial reproductive tract. In: Auer JA (ed.) Equine Surgery. Philadelphia: Saunders, 1992; pp/ 750–61. 12. Vaughan JT. Equine urogenital systems. In: Morrow DA (ed.) Current Therapy in Theriogenology. Philadelphia: Saunders, 1980; pp. 756–75. 13. Vaughan JT. Surgery of the male equine reproductive system. In: Jennings PB (ed.) The Practice of Large Animal Surgery. Philadelphia: Saunders, 1984; pp. 1092–104. 14. Shoemaker RW, Read EK, Duke T, Wilson DG. In situ coagulation and transection of the ovarian pedicle: an alternative to laparoscopic ovariectomy in juvenile horses. Can J Vet Res 2004;68:27–32. 15. LeBlanc MM. Diseases of the ovary. In: Colahan PT, Merritt AM, Moore JN, Mayhew IG (eds) Equine Medicine and Surgery. St Louis, MO: Mosby, 1999; pp. 1157–64. 16. Trotter GW, Embertson RM. The uterus and ovaries. In: Auer JA, Stick JA (eds) Equine Surgery, 2nd edn. Philadelphia: Saunders, 1999; pp. 575–83. 17. Moll HD, Slone DE, Juzwiak JS, Garrett PD. Diagonal paramedian approach for removal of ovarian tumors in the mare. Vet Surg 1987;16:456–8. 18. Santschi EM. How to perform bilateral ovariectomy in the mare through two paramedian incisions. Proc Am Assoc Equine Pract 2001;47:420–2. 19. Ragle CA, Southwood LL, Hopper SA, Buote PL. Laparoscopic ovariectomy in two horses with granulosa cell tumors. J Am Vet Med Assoc 1996;209:1121–4.
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20. Rodgerson DH, Belknap JK, Wilson DA. Laparoscopic ovariectomy using sequential electrocoagulation and sharp transection of the equine mesovarium. Vet Surg 2001; 30:572–9. 21. Rodgerson DH, Brown MP, Watt BC, Keoughan CG, Hanrath M. Hand-assisted laparoscopic technique for removal of ovarian tumors in standing mares. J Am Vet Med Assoc 2002; 220:1503–7, 1475. 22. Fischer AT Jr, Lloyd KC, Carlson GP, Madigan JE. Diagnostic laparoscopy in the horse. J Am Vet Med Assoc 1986; 189:289–92. 23. Ragle CA. Laparoscopic ovariectomy. In: Wolfe DF, Moll HD (eds) Large Animal Urogenital Surgery. Baltimore, MD: Williams & Wilkins, 1997; pp. 143–6. 24. Hubert JD, Burba DJ, Moore RM. Evaluation of a vesselsealing device for laparoscopic granulosa cell tumor removal in standing mares. Vet Surg 2006;35:324–9. 25. Rodgerson DH, Hanson R. Ligature slippage during standing laparoscopic ovariectomy in a mare. Can Vet J 2000; 41:395–7. 26. Meagher DM, Wheat JD, Hughes JP, Stabenfeldt GH, Harris BA. Granulosa cell tumors in mares: a review of 78 cases. Proc Am Assoc Equine Pract 1978;23:133–43. 27. Wilson DA, Baker GJ, Boero MJ. Complications of celiotomy incisions in horses. Vet Surg 1995;24:506–14. 28. Bleyaert HF, Brown MP, Bonenclark G, Bailey JE. Laparoscopic adhesiolysis in a horse. Vet Surg 1997;26: 492–6.
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Chapter 266
Hysterectomy Anna K. Roetting and David E. Freeman
Introduction The most common indication for complete hysterectomy in the mare is chronic pyometra that is unresponsive to medical management. Other indications reported are chronic uterine torsion, uterine neoplasia, extensive uterine damage and rupture, and, rarely, segmental aplasia with accumulation of secretions.1–6 The most common indication for partial hysterectomy is neoplasia of or adjacent to one uterine horn.4, 5, 7 Mares can be successfully bred and produce a live foal after partial hysterectomy. The maximum amount of uterine tissue that can be removed without compromising reproductive function is unknown, but close to one entire uterine horn has been removed without adverse effects on future pregnancies.4, 8, 9 Partial hysterectomy should be performed in conjunction with ipsilateral ovariectomy of the ipsilateral horn because this will result in shifting of the ovarian cycle to the side with an intact horn, thereby optimizing future fertility of the mare.4 Bilateral hysterectomy should be performed in conjunction with complete ovariectomy. There is one report on hysterectomy alone in a 7-month-old pony, and this filly survived during a follow-up of 1 year, but without showing any signs of estrus.10 Surgical exposure of the cervix in the mare is very difficult, and in most cases the uterine body is transected cranial to the cervix, leaving the potential for development of a stump pyometra. Pyometra has been defined as a hormonally mediated diestral disorder, resulting from bacterial interactions with an endometrium that has undergone pathological changes in response to progesterone.11 In dogs, the development of pyometra, including stump pyometra after hysterectomy, is dependent on the presence of progesterone, and complete removal of all ovarian tissue will prevent pyometra formation in the absence of an exogenous source of progesterone. 12 The removal of all ovarian tissue in the mare after ovariohysterectomy may help to prevent stump
pyometra formation and may explain the low incidence of this complication reported in the literature. Chronic pyometra in the mare is usually associated with severe pathological changes of the uterine wall, and removal of as much uterus as possible seems to be advisable in these cases.
Surgical procedure Patient preparation Mares should be fasted for 12–24 hours before surgery. To reduce bulk of the large colon and improve surgical exposure to the uterus, mares can be fed a low-bulk diet for several days before the procedure. The use of laxatives (e.g., mineral oil) can soften the feces and minimize post-operative complications associated with impactions. In mares with chronic pyometra the uterus is emptied and thoroughly lavaged prior to surgery. A sample of the uterine contents should be submitted for microbiological examination. Mares receive pre-operative broad-spectrum antibiotics, based on microbiological culture and sensitivity results, and also non-steroidal anti-inflammatory drugs. Complete ovariohysterectomy The challenge for complete ovariohysterectomy is to remove as much of the uterus as possible while minimizing or eliminating the risk of abdominal contamination during amputation through the uterine body. For surgery, the mares are placed in dorsal recumbency and prepared and draped for a caudoventral median approach. A caudal ventral median incision is made for 35–40 cm in the skin, dividing the mammary glands. In overweight mares, large amounts of fat will have to be divided in the subcutaneous space. The body wall is incised sharply on the midline to the prepubic tendon. Fibrinous and fibrous adhesions of the uterus to the surrounding tissues, including
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 266.1 Surgical technique. Branches of both ovarian arteries to ovaries, tube, and uterus and of both uterine arteries have been ligated and transected. Both ovaries and the uterus are exteriorized and a TA-90 surgical stapling instrument with a prefired cartridge is applied across the uterine body. Stay sutures are placed in the caudal uterine body and stepwise transection of the uterus and closure of the uterine stump has started. Traction on the uterus is used to improve exposure: 1, traction on the TA-90; 2, traction on stay sutures; 3, traction on suture line closing the uterine stump. (Redrawn with permis¨ sion from Rotting et al.5 )
the ventral body wall, may have to be broken down in mares with chronic uterine torsion. In most cases of pyometra, the uterus can be only partly emptied before surgery, so that it must be exteriorized and packed off to prevent abdominal or incisional contamination. A purse-string suture can be placed in the uterine wall around a 1-cm incision, and a Poole suction tip inserted into the uterine lumen to remove as much of the liquid contents as possible. In mares with chronic uterine torsion, the fetus is removed by cesarean section and the uterus wall is closed before proceeding with the ovariohysterectomy. The uterine wall is too thin and friable in these mares to tolerate correction of the torsion with the foal in place. The first step in ovariohysterectomy is ovariectomy. One ovary is exteriorized and the branches of the ovarian artery are double ligated with transfixation ligatures of size 1 absorbable suture material (Figure 266.1). The mesovarium and vessels are transected on the ovarian side of the ligatures with scissors or a Serra emasculator. Alternatively, adequate hemostasis of ovarian vessels can be achieved using
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R 14 , or an elecsurgical staples,13 a Harmonic Scalpel R 15 trical vessel-sealing device (e.g., LigaSure ). The same procedure is performed for removal of the contralateral ovary. Dissection is then continued caudally in the broad ligament, and branches of the ovarian artery to the tube and the uterus, and the uterine artery16 are transected and ligated with size 1 absorbable ligatures R has been re(Figure 266.1). The Harmonic Scalpel ported to be inadequate for hemostasis of large vessels in the mesometrium in one report, whereas application of endoscopic clips to skeletonized vessels was sufficient in the same study.17 In two mares with chronic uterine torsion, the broad ligament could not be identified3 and this step was omitted. After the broad ligament is incised, the uterus is exteriorized. Finochietto rib spreaders, Deaver retractors, or manual retraction can be used to retract incision edges to improve exposure to the most caudal part of the uterus. Exposure can be improved by exteriorizing the large colon.2 Any liquid in the uterus is massaged towards the horns and a TA-90 with a previously fired cartridge or one or two right-angled clamps (Best clamps) are placed far caudally on the uterine body and used to retract the cervix cranially (Figure 266.2). The cervix can be pushed towards the abdominal incision by an assistant with a hand in the vagina to improve access. A number of large arteries, derived from the caudal uterine branch of the urogenital artery,16 ramify across the proposed line of transection and should be individually ligated with size 1 absorbable suture material before and during transection (Figure 266.1). Care should be taken to avoid damage to the ureters during this part of the surgery.
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Figure 266.2 Surgery. The uterus (1) has been exteriorized and a TA-90 surgical stapling instrument with a prefired cartridge is used to occlude the uterine body. A stay suture is being placed on the caudal uterine body.
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Before uterine body transection, stay sutures of large (size 2 or larger) absorbable material are placed in cruciate fashion on each side of the proposed line of transection. Simultaneous traction on these stay sutures and the TA-90 or Best clamps draw the cervix towards the incision (Figure 266.2). The abdominal incision and uterus are then packed off with salinesoaked laparotomy sponges deep in the abdomen and under (dorsal to) the proposed line of transection. Starting on the side away from the surgeon, the first closure layer is started with size 2 or larger absorbable suture material (e.g., polydioxanone) and the uterus is incised for 3–5 cm immediately cranial to that suture (Figure 266.2). The caudal open end of the uterus or cervix is then closed by continuing this suture in a Lembert pattern while an assistant maintains traction on the suture to keep the uterine stump as close to the incision as possible. This is continued in stepwise fashion until amputation is complete. The Lembert closure is then oversewn with one or two Cushing layers. The abdomen is lavaged with an isotonic sterile solution (e.g., NaCl or Ringer’s solution) and the abdomen is closed in a routine fashion. A hand-assisted laparoscopic approach for ovariohysterectomy in the mare has been described.17 Only the ligation and transection of the mesovarium and mesometrium were performed under laparoscopic guidance. The uterus was subsequently exteriorized through a caudal ventral midline incision and the uterine body was transected using an open approach.
Partial ovariohysterectomy The surgical approach is similar to that of total ovariohysterectomy. The broad ligament of the affected horn is dissected until the level of proposed transection. The horn is clamped with Carmalt clamps, the cranial part is removed, and the stump oversewn in Parker Kerr and Cushing patterns with absorbable suture material (no. 1 polydioxanone).
Aftercare Broad-spectrum antibiotics and non-steroidal antiinflammatory drugs are continued for 3–8 days after surgery. Laxatives can be given as needed.
sional drainage is also common and may be related to the large subcutaneous dead space. Other lifethreatening complications reported include severe hemorrhage, septic peritonitis, uterine stump infection, necrosis, and abscessation, colitis, and fever. In one report, a longer segment (approximately 8 cm) of uterine body was left in situ to facilitate secure closure of the uterine stump.4 The authors emphasized the importance of removing the entire remaining endometrium before closing the uterine stump. This additional step does not appear to be necessary to prevent uterine stump infection,5, 6 and would be associated with an increased risk of peritoneal contamination. We therefore recommend removing as much of the uterus as possible without compromising closure. Transfixation of the uterine body may be associated with a higher risk of infection and should be avoided. A transfixation suture was used in two out of six mares in one report, and both mares developed uterine stump abscessation.2 Severe cervical scarring and cervical adhesion formation may also be associated with a higher complication rate. In one report, three out of six mares had severe cervical adhesions or a distorted cervix, and all of these developed peritonitis, uterine stump infection, or both.2 In another report on 17 ovariohysterectomies, the only death was in a miniature mare with severe cervical scarring and adhesions.6 American Miniature Horses might be more prone to cervical damage, because their small size would put them at risk of greater cervical trauma from intravaginal manipulation and forced extraction procedures. Both Miniature Horse mares that have been reported to undergo ovariohysterectomy had severe cervical scarring and adhesions.2, 6 If some infected uterine body remains after surgery and the cervix is not sufficiently open, per vaginal placement of an indwelling tube of 30 Fr or more may be performed under intraoperative guidance to allow continued post-operative drainage and lavage of the uterine stump. Post-operative insertion of such a tube would be difficult and dangerous.
Prognosis The prognosis after ovariohysterectomy is good despite the risk for life-threatening complications. From 31 reported cases, 30 (97%) survived to discharge, and from 23 cases for which follow-up was available 21 (91%) survived long term.6
Complications The most frequently reported complication of ovariohysterectomy is mild abdominal discomfort, probably due to pain related to intraoperative tension on the ovarian pedicle and broad ligament.2, 4, 5 Inci-
References 1. Slone DE. Ovariectomy, ovariohysterectomy, and cesarean section in mares. Vet Clin North Am Equine Pract 1988; 4:451–9.
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Hysterectomy 2. Santschi EM, Adams SB, Robertson JT, DeBowes RM, Mitten LA, Sojka JE. Ovariohysterectomy in six mares. Vet Surg 1995;24:165–71. 3. Doyle AJ, Freeman DE, Sauberli DS, Hammock PD, Lock ¨ TF, Rotting AK. Clinical signs and treatment of chronic uterine torsion in two mares. J Am Vet Med Assoc 2002;220: 349–53. 4. Bartmann CP, Schiemann V, Poppe C, Schoon HA. Partielle und radikale Hysterektomie beim Pferd. Tier¨arztl Prax 2003;31:6–15. ¨ 5. Rotting AK, Freeman DE, Doyle AJ, Lock T, Sauberli D. Total and partial ovariohysterectomy in seven mares. Equine Vet J 2003;35:29–33. ¨ ¨ 6. Freeman DE, Rotting AK, Kollmann M, Doyle AJ, Troedsson MHT, Pozor M, Lock T, Stewart A, Trumble T. Ovariohysterectomy in mares: 17 cases (1988–2007). In: Proceedings of the 53rd Annual Convention of the American Association of Equine Practitioners, 2007, pp. 370– 3. 7. Berezowski C. Diagnosis of a uterine leiomyoma using hysteroscopy and a partial ovariohysterectomy in a mare. Can Vet J 2002;43:968–70. 8. Santschi EM, Slone DE. Successful pregnancy after partial ovariohysterectomy in two mares. J Am Vet Med Assoc 1994;205:1180–2. 9. Asjanowa I, Klug E. Erfolgreiche Tr¨achtigkeit nach partieller Hysterektomie bei einer Stute. Tier¨arztl Prax 2002; 30:340–3.
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10. Wehrend A, Herfen K, Litzke LF, Bostedt H. Hysterektomie bei einer Jungstute mit einer Zervix duplex und rezidivierender Endometritis. Pferdeheilkunde 2001;17:141–4. 11. Nelson EC, Feldman RW. Pyometra. Vet Clin North Am Small Anim Pract 1986;16:561–6. 12. Van Goethem B, Schaeffers-Okkens A, Kirpensteijn J. Making a rational choice between ovariectomy and ovariohysterectomy in the dog: a discussion of the benefits of either technique. Vet Surg 2006;35:136–43. 13. Rodgerson DH, Brown MP, Watt BC, Keoughan CG, Hanrath M. Hand-assisted laparoscopic technique for removal of ovarian tumors in standing mares. J Am Vet Med Assoc 2002;220:1503–6. ¨ 14. Dusterdiek KF, Pleasant RS, Lanz OI, Saunders G, Howard RD. Evaluation of the Harmonic Scalpel for laparoscopic bilateral ovariectomy in standing horses. Vet Surg 2003;32: 242–50. 15. Hand R, Rakestraw P, Taylor T. Evaluation of a vessel sealing device for use in laparoscopic ovariectomy in mares. Vet Surg 2002;31:240–4. 16. Schummer A, Wilkens H, Vollmerhaus P, Habermehl K. Visceral arteries of the internal iliac and internal pudendal interna. In: Nickel R, Schummer A, Seiferle E (eds) The Anatomy of the Domestic Animals. P. Parey, 1981; pp. 176–83. Berlin & Hamburg. 17. Delling U, Howard RD, Pleasant RS, Lanz OI. Handassisted laparoscopic ovariohysterectomy in the mare. Vet Surg 2004;33:487–94.
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Chapter 267
Rectal Tears David E. Freeman
Although rectal tears are common causes of lawsuits against veterinarians, the promptness and the quality of care provided for the horse after the tear is inflicted is likely to determine the outcome of most malpractice claims. As soon as a rectal tear is suspected, the veterinarian must assess the tear for severity, inform the owner about the nature of the problem (without making statements that admit guilt or responsibility for payment), and apply appropriate treatment, including referral. Failure to follow these guidelines will weaken a veterinarian’s legal defense. The veterinarian should also inform his or her liability insurance company promptly.
Causes Most iatrogenic rectal tears result from rupture of the rectal wall as it contracts around the examiner’s hand or forearm rather than from penetration with the finger tips. Standard measures to prevent iatrogenic rectal tears involve copious lubrication of the hand and forearm and adequate restraint of the horse, including sedation if necessary. The risk of a tear is increased by forcing the hand against straining or a peristaltic wave, and by attempting to palpate intraabdominal structures that are almost beyond reach. Instead, the examiner should wait until the hand has been inserted beyond the area of interest and then withdraw it to that level. Special precautions should be taken with Arabian horses, ponies, small breeds, horses that have had a previous rectal tear or injury, horses that are unaccustomed to palpation per rectum, fractious horses, and horses with colic.1, 2 Rarer causes of rectal tears are enemas, meconium extraction by forceps, sadism, dystocia, animal bites, chronic impaction at a stricture, misdirection of a stallion’s penis,1–4 spontaneous tears,5 rectal thrombosis,6 and sand impactions.7 Rectal tears can occur immediately following term foalings, without evidence of dystocia, apparently caused by foalinginduced trauma to the rectum and small colon.8
Classification and locations Rectal tears are assigned to grades I to IV based on severity. In grade I tears, only the mucosa and submucosa are torn.1 In grade II tears, only the muscular layer is disrupted, causing the mucosa and submucosa to prolapse through the defect and create a site for fecal impaction.1 These are rare (3 of 85 tears in one series), but can be chronic and untreatable.9 Grade III tears are subdivided into grade IIIa, which involves all layers except the serosal, and grade IIIb, which involves the mesorectum and retroperitoneal tissues.2, 10 Grade III tears tend to pack with feces and dissect cranially and dorsally (Figure 267.1), even approaching the left kidney, and feces in retroperitoneal spaces can cause severe peritonitis. Grade IV tears involve all layers and cause fecal contamination of the peritoneal cavity and death.1 Rectal tears immediately after parturition, without dystocia, are usually grade IV and can be associated with prolapse of small colon or small intestine through the tear and anus.8 Most tears involve the dorsal aspect of the rectum, 15 to 55 cm from the anus9, 10 and parallel to the longitudinal axis,1, 2 but ventral tears have been reported.8 Distance from the anus is not a reliable indicator of tear location relative to the retroperitoneal reflection and peritoneal cavity.2, 7, 11
Clinical signs and diagnosis Early diagnosis favors successful treatment and prevents legal repercussions. At the moment a tear occurs, the veterinarian may feel a sudden release of pressure or be able to directly palpate abdominal organs.2, 4 A rectal tear should be suspected if a large amount of blood is evident on the rectal sleeve on withdrawal of the hand or if the rectum suddenly relaxes while the horse is straining.1 Alternatively, the rectum can tear without giving any sensation or evidence that this catastrophe has happened.2 Within
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 267.1 Endoscopic views of opening into a grade IIIb rectal tear at 11 days after iatrogenic injury (a). Retroperitoneal space dissection for a considerable distance cranial to the tear (b), after feces have been removed.
a few hours after the rectum is torn, the horse usually shows signs of peritonitis and endotoxic shock. Signs of colic are also present initially, soon followed by depression. Feces can be hemorrhagic initially and defecation can be accompanied by straining.12 Idiopathic tears are usually presented as colic of variable duration, and, because there is no reason to suspect a rectal tear, referral can be delayed.13 Before examination of the tear, straining and rectal contractions are stopped with epidural anesthesia or xylazine (0.1 to 0.2 mg/kg bodyweight i.v.), alone or in combination with butorphanol tartrate (0.1 mg/kg i.v.). Butylscopolamine bromide (Buscopan), 20 mg i.v., relaxes the rectum and small colon and facilitates examination of rectal tears. A lidocaine enema can be given (12 mL of 2% lidocaine in 50 mL of tapwater) or lidocaine jelly applied to the rectal mucosa before inspecting the tear digitally or through a tube speculum or endoscope.4 For digital palpation, the examiner can use a bare hand or wear a surgeon’s glove and apply copious lubrication with a watersoluble gel. Because the abundant mucosal folds tend to obscure the rectal tear, more information can be gained from careful palpation than by visual inspection. Failure to accurately grade a tear can cause inappropriate treatment selection. A grade I tear can feel like a flap of mucosa, but a grade III tear can feel as if its edges are firm, thick, and separated, often by packed feces. Abdominocentesis should be performed to assess peritonitis.
Initial treatment First aid is applied as soon as the diagnosis is made and includes (i) reduction of rectal motility; (ii) epidural anesthesia to allow gentle removal of feces from the tear and rectum; (iii) treatment of
septic shock and peritonitis; and (iv) packing of the rectum.1, 14 Flunixin meglumine (1.1 mg/kg every 24 hours i.v.) and antibiotics, such as sodium or potassium penicillin (22 000 IU/kg of bodyweight every 6 hours i.v.), gentamicin (6.6 mg/kg every 24 hours i.v.), and metronidazole (15 mg/kg every 6 hours p.o.), should be administered and continued as indicated. Intravenous fluids are required to treat shock. Rectal packing (Figure 267.2) can be used as a temporary measure to protect the tear and peritoneal cavity from fecal contamination or to prevent conversion of a grade III to a grade IV tear before definitive treatment can be applied, as during transportation.14 Preferred packing material is a 3-inch (8-cm) stockinette filled with 0.25 kg of moistened rolled cotton, sprayed with povidone-iodine, and lubricated with surgical gel.14 The packing should fill the rectum without distension to a point 10 cm proximal to the tear (Figure 267.2), taking care not to pack the tear and enlarge it.14 The anus is then closed with towel clamps or pursestring suture, and the epidural anesthetic is repeated as needed to decrease straining.14
Non-surgical treatment Grade I and II tears rarely require surgical treatment, and grade I tears respond in most cases to antibiotics (as above or trimethoprim–sulfonamide, 20 mg/kg p.o. every 12 hours), and flunixin meglumine (1.1 mg/kg i.v. or p.o., every 12 hours), mineral oil (4 liters via nasogastric tube every 24 hours), and dietary changes such as bran mashes, moistened pellets, or grass to soften feces and reduce fecal volume. Non-surgical management of grade III tears is usually successful and can be less expensive than surgical methods.12, 15, 16 Grade IIIb rectal tears can be
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Figure 267.2 Rectal packing used as a temporary measure to protect the tear and peritoneal cavity from fecal contamination (arrow denotes site of fecal obstruction by packing). The packing fills the rectum without distension to a point 10 cm proximal to the tear, and the anus is then closed with towel clamps or pursestring suture. The epidural anesthetic is required to allow insertion of the pack and to decrease straining.
treated successfully with broad-spectrum antibiotics (penicillin, gentamicin, and metronidazole), nonsteroidal anti-inflammatory drugs, fecal softeners, manual removal of feces from the rectum daily, and packing of any dissecting defects with gauze soaked in antiseptic solution.12,15–17 Manual evacuation of feces should be performed only if the tear becomes impacted, and then with extreme care plus sedation and epidural anesthesia to improve tolerance.16 More frequent evacuations can be indicated in some cases for weeks, but this is labor intensive and requires repeated epidurals through an epidural catheter.15 Septic peritonitis can be treated by peritoneal lavage if needed. A rectal diverticulum can develop in the tear, without any apparent clinical effects.16
Suture repair Suture repair is used to facilitate healing of grade III and grade IV tears and prevent progression of a grade III tear to a grade IV tear.10, 18 If possible, it should be combined with a diverting procedure if there is any concern about the integrity of the suture line, especially if the tear is grade IV. The surgery is easiest to accomplish on a thin horse with a fresh, clean tear that is close to the anus.10, 19 Long-handled instruments with pistol grips and an expandable rectal speculum or ‘cage’ have been used,20 but have not gained widespread use. If the wound is large and old, it can be partly closed and the ventral part left open to
drain.10, 17 The remaining defect can be packed with antiseptic-soaked gauze to prevent fecal impaction and dissection.10, 17 A technique for non-visual direct suturing has been used successfully as an inexpensive standing procedure for tears involving half or less of the rectal circumference, and as an adjunct to bypass procedures for grade IV tears.18 With the tail elevated and the anal sphincter relaxed from caudal epidural anesthesia, fecal material is manually removed from the rectum and distal small colon, and the defect and lumen walls are cleaned carefully and thoroughly by wiping with moistened gauze sponges. The left hand is used for tears on the right side, and vice versa, and gloves are not worn.18 The ideal suture for rectal tears should be long (100 to 150 cm), have low memory, resist stretching, resist destruction by feces, and be large enough to minimize cutting through the friable tissue edges.18 The preferred needle is a 6- to 8-cm, half-circle cutting or trocar point.18 Continuous suture patterns are not recommended because they reduce the lumen diameter, and cause impaction and dehiscence.18 The needle is manually placed in the center of the caudal edge of the tear, approximately 1.5 cm from the edge of the wound, pulled through the tissue, grasped with the thumb and first two fingers, and then placed in the center of the cranial edge.18 The third finger is used to guide the exit point and press the tissue onto the needle. An assistant holds the clamped suture ends to one side to close the defect into a transverse plane, and the needle is reintroduced into the rectum to pass through both cranial and caudal edges of the defect in one bite using digital manipulation as before. The cruciate suture thus completed is tied by hand, with each throw started outside the rectum and then slid down to the knot. Traction on this first suture should convert the tear to a transverse orientation, which facilitates placement of about two or three more sutures on each side of the first. Care is taken to prevent lumen stenosis, and suture ends are cut long to facilitate their removal in 12 to 14 days.18 After surgery, a complete pellet ration can be fed after a 2-day fasting period to produce small fecal bulk, and mineral oil can be used to soften feces.18 Mineral oil is not used if repair of a grade IV tear is incomplete, because it could leak into the abdomen.18 An alternative suture method is to use the 45-cm-long Deschamps needle (Eickemeyer, Tuttlingen, Germany), which comes in right-handed and left-handed configurations.21 The needle tip should be sharp. The caudal end of the perforation is grasped with the index finger and thumb of one hand, and the needle is guided through the tissue edges, about 1 cm away from the wound edge.21 Suture
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Figure 267.3 Use of the TA90 to close a grade IV rectal tear. The dorsal commissure of the anus was incised to improve exposure, and four sutures in the anus are used to retract the edges to the sides (solid arrows). The tear is drawn in the direction of the open arrow by Allis tissue forceps, and, after the TA90 is fired, the tissue is cut to leave approximately 5 mm of rectal wall beyond the instrument for suture closure.
placement with the needle is continued as for the manual technique. Grade IV tears have been repaired successfully with the TA-90 Premium stapling instrument (United States Surgical Corp., Norwalk, CT) in horses in which the tear could be prolapsed through the anus8, 22 (Figure 267.3). The staple line can be reinforced with a simple continuous suture pattern.8 This technique is facilitated by pneumoperitoneum and the subsequent equilibration of pressure across the rectal wall that allows the torn rectal wall to be drawn through the anus.22 In the immediate postpartum period, retraction of the tear could be further facilitated by laxity in the perirectal tissues.8 To improve access to a rectal tear, four equidistant stay sutures can be used to retract the anal sphincter and the anal sphincter can be transected at the dorsal commissure8 (Figure 267.3). The latter can be sutured after surgery is completed8 or left open to ease defecation afterwards.17, 21 A rectal tear can also be accessed directly by prolapsing it through the anus by intraoperative guidance through a caudal ventral midline laparotomy19 or through an antimesenteric enterotomy from a ventral midline approach.23 Suture repair by laparoscopy requires laparoscopic instruments that can access the caudal abdominal cavity and pelvis.24
Temporary indwelling rectal liner A temporary indwelling rectal liner25 and colostomy4, 26, 27 are both designed to divert feces away from grade III and grade IV rectal tears, and thereby prevent fecal contamination and impaction of the perirectal defect, enlargement of the tear, and worsening of peritonitis. Each can be used with or without a suture closure of the tear. Case
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selection for these procedures is difficult, because some horses with grade III rectal tears can make a full recovery with non-surgical treatment only (see above). Regardless, the decision to use diverting procedures must be made quickly (< 12 hours) to derive full benefit.26, 27 Because the cost of diverting procedures can be considerable, many owners may opt for less expensive treatments, such as direct suturing or non-surgical treatments. A temporary indwelling rectal liner (Figure 267.4) can be constructed by gluing a palpation sleeve or the plastic cover for an arthroscope camera, to a 5-cm-diameter and 7.5-cm-long rectal prolapse ring with holes through which Dacron loops are laced.2, 25 With the horse anesthetized in dorsal recumbency, an unscrubbed assistant passes the well-lubricated ring and liner through the anus so the surgeon can guide it through a ventral midline celiotomy, cranial to the tear (Figure 267.4) but close enough to prevent retraction of the open end of the liner through the anus when the horse stands.2, 25 The ring is secured with a tight circumferential suture that is incorporated in individual sutures through all layers of the colon and the Dacron loops in the prolapse ring.25 A Lembert pattern is used to bury the circumferential suture.25 The large colon is emptied through a pelvic flexure enterotomy, and the small colon is emptied by flushing with a hose directed through the ring and liner from the anus.2 The circumferential suture will cut through the colon wall and allow passage of the ring and liner within approximately 9 to 12 days, by which time the apposed colon walls will have healed. 25 Failures include tearing of the liner, retraction of the liner sufficiently to uncover the tear, and formation of a rectoperitoneal fistula.2, 25
Loop-colostomy The loop-colostomy technique is preferred over the end-colostomy because it is easier and quicker to establish and to reverse later.4 Concerns about incomplete diversion of feces do not apply to a loop colostomy in horses because gravity and creation of a septum between the loops will direct feces directly through the stoma.4, 26, 27 Because fecal balls are eliminated individually through the stoma, the risk of obstruction is low. Sites for a loop colostomy are high left flank, low left flank, or ventral midline (Figure 267.5). A singleincision colostomy involves placing the stoma in the same incision as used to explore the abdomen, as in a horse with colic.4 A double-incision colostomy involves a separate incision for exploration of the abdomen or to locate and prepare the appropriate segment of small colon, and another incision for the
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6 Figure 267.4 Placement of a temporary indwelling rectal liner to divert feces from a rectal tear. The expanded view shows the construction of the liner and the method for suturing it to the small colon. 1, tear; 2, rectal liner; 3, rectal prolapse ring; 4, Dacron loop; 5, circumferential suture; 6, retention suture; 7, interrupted Lembert suture.
stoma (Figure 267.6). The double-incision colostomy is recommended because the separate flank incision can be made small enough to form a snug fit around the stoma, so the surrounding intact body wall reduces the risks of prolapse and herniation.4, 26, 27 A colostomy can be difficult to place accurately in the anesthetized horse because incisional layers and landmarks shift when the horse stands, and the stoma can be disrupted during anesthetic recovery.4 The low flank incision, in line with the flank fold and midway between it and the costal arch, is the pre-
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Figure 267.5 Sites for placement of a loop colostomy (comments in parentheses) and landmarks. 1, high flank (skin contamination); 2, low flank (preferred, but requires a separate high flank incision to locate and prepare small colon); 3, ventral midline (weakens ventral midline closure and is possibly prone to prolapse – can be used to place a colostomy in the low flank); 4, costal arch; 5, skin fold in lower flank.
ferred site for the stoma (Figure 267.6). The 8-cm-long incision in the body wall is angled dorsally at its caudal end by 20 to 30 degrees, and small transverse incisions are made in muscles and fascia to eliminate constricting bands around the colon. The stoma should be large enough to reduce the risk of stomal obstruction but small enough to prevent the more serious complications of prolapse and herniation.4 The stoma is made in a segment of small colon at least 1 m from the rectum to allow easy access for colostomy reversal. The segment of small colon is folded to form a loop, and the two arms of the loop are sutured together with an absorbable material in a continuous Lembert pattern for 8 to 10 cm, midway between the mesenteric and antimesenteric teniae.26 This suture line creates a more complete separation or septum between the proximal and distal segments of small colon, to obtain complete fecal diversion. It also stabilizes the loop within the body wall, reducing the risk of prolapse.4 The prepared loop of small colon is subsequently inserted into the flank incision with the proximal loop proximal and slightly ventral to the distal loop (Figures 267.6 and 267.7), and the antimesenteric tenia about 3 cm beyond the skin.4 The seromuscular layer of the colon is sutured to the abdominal muscles and fascia using several interrupted sutures of 0 or 2–0 absorbable material, taking care not to puncture or occlude mesenteric vessels.4 An 8-cm incision is made along the exposed antimesenteric tenia of the colon, and the cut edges are folded back and sutured to the skin with simple-interrupted sutures of 2–0 nylon or polypropylene.4 Antibiotics and laxatives (mineral oil, 2–4 L/ 450 kg, and magnesium sulfate, 1 g/kg) are continued for 3 to 5 days. Horses are held off feed or fed
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Figure 267.6 Loop colostomy created with a double-incision technique with the high flank approach (broken lines) and placement of the stoma in a snug low flank incision, which is the preferred method, suitable as a standing procedure. (a) Intra-abdominal view through a cross-section; (b) side view. The open circle is the proximal opening in the stoma, and the solid circle is the distal opening through which the distal part of the small colon is flushed to prevent atrophy.
grass, alfalfa hay, or complete pellet feeds at half the usual amounts for the first 2 to 3 days after the colostomy is established, and ointment is applied to the skin around the stoma to protect it from scalding. A cradle is applied because most horses have a tendency to mutilate the colostomy. The mucosal protrusion of the stoma becomes markedly congested in the first week after surgery and slowly sloughs, to be replaced with healthy mucosa. (Figure 267.7). Ventral edema in the ventral body wall resolves with time (Figure 267.7). When the tear has started to granulate (5 to 7 days), the terminal small colon and rectum are flushed through the stoma with approximately 20 L
of warm water to exercise these segments and prevent the atrophy that can complicate reversal.4 The colostomy can be reversed after the tear has healed, at approximately 6 weeks. The horse is anesthetized in right lateral recumbency, the stoma is resected en bloc, and a colonic anastomosis is performed through the resulting flank incision.4 The postoperative feeding, antibiotic, and laxative regimens are similar to those used after the colostomy was made. Complications of colostomy are dehiscence, abscessation, peristomal herniation, prolapse, prolapse with rupture of mesenteric vessels, infarction, rupture of the colostomy, spontaneous closure, and stomal obstruction.4 Complications of reversal are incisional infection and anastomotic impaction and dehiscence. Colostomy is more expensive than an indwelling rectal liner because it is a two-stage procedure, but it can allow the surgeon more complete control over the duration of fecal diversion, which could be important if healing of the tear is delayed.4 Colostomy is preferred for large tears, small horses, and a tear that is too far proximal to accommodate a rectal liner.25
Prognosis for rectal tears
Figure 267.7 Loop colostomy placed in the low flank by a double-incision technique as a standing procedure 7 days earlier. The shelf or septum separating the proximal and distal parts of the small colon is evident as a mucosal bulge in the caudal part of the stoma. Some ventral edema is typical and ¨ resolves with time. Photograph courtesy of Dr Anna Roetting.
Complications of grade III rectal tears are cellulitis, abscessation, severe toxemia, peritonitis, laminitis, and recurrent intestinal obstruction from adhesions.4 Most tears heal with little residual damage, but some can form a stricture,3 a diverticulum,16 or perirectal abscess.4, 26, 27 Such abscesses must be drained into the rectum or into the vagina, if possible, and can delay colostomy reversal.4, 26, 27 Grade III and IV tears
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can heal to form a mucosal or submucosal diverticulum, which can subsequently become impacted with feces,1 and a grade IV tear can form a rectoperitoneal fistula.25 Reported survival rates are 93% for grade I tears, 70% for grade IIIa tears, 69% for grade IIIb tears, and 6% for grade IV tears.9 Non-visual direct suturing of rectal tears can improve survival over rates reported previously for other surgical techniques.9, 18
References 1. Arnold S, Meagher D, Lohse C. Rectal tears in the horse. J Equine Med Surg 1978; 2: 55–63. 2. Baird AN, Freeman DE. Management of rectal tears. Vet Clin N Am Equine Pract 1997; 13: 377–92. 3. Speirs VC, Christie BA, van Veenendaal JC. The management of rectal tears in horses. Aust Vet J 1980; 56: 313–17. 4. Freeman DE. Surgery of the rectum and anus. In: Auer JA, Stick JA (eds) Equine Surgery, 3rd edn. Philadelphia: W.B. Saunders Co., 2005; pp. 479–91. 5. Slone DE, Humburg JM, Jagar JE, Powers RD. Noniatrogenic rectal tears in three horses. J Am Vet Med Assoc 1982; 180: 750–1. 6. Guglick MA, MacAllister CG, Ewing PJ, Confer AW. Thrombosis resulting in rectal perforation in a horse. J Am Vet Med Assoc 1996; 209: 1125–7. 7. Mazan MR. Medical management of a full-thickness tear of the retroperitoneal portion of the rectum in a horse with hyperadrenocorticism. J Am Vet Med Assoc 1997; 210: 665–7. 8. Kay AT, Spirito MA, Rodgerson DH, Brown SE. Surgical technique to repair grade IV rectal tears in post-parturient mares. Vet Surg 2008; 37: 345–9. 9. Eastman TG, Taylor TS, Hooper RN, Honnas CM. Treatment of rectal tears in 85 horses presented to the Texas Veterinary Medical Center. Equine Vet Educ 2000; 12: 342–5. 10. Watkins JP, Taylor TS, Schumacher J, Taylor JR, Gillis JP. Rectal tears in the horse: An analysis of 35 cases. Equine Vet J 1989; 21: 186–8. 11. Sisson S. Equine digestive system. In: Getty R (ed.) Sisson and Grossman’s The Anatomy of the Domestic Animals, 5th edn. Philadelphia: W.B. Saunders, 1975; pp. 454–97.
12. Alexander GR, Gibson KT. Non-surgical management of rectal tears in two horses. Aust Vet J 2002; 80: 137–9. 13. Scheidemann W, Huskamp NH, Odenkirchen S. Id¨ Pferden. Tier¨arztl Prax iopathische Rektumruptur bei funf 2003; 31: 1–5. 14. Baird AN, Taylor TS, Watkins JP. Rectal packing as initial management of grade 3 rectal tears. Equine Vet J Suppl 1989; 7: 121–3. 15. Katz LM, Ragle CA. Repeated manual evacuation for treatment of rectal tears in four horses. J Am Vet Med Assoc 1999; 215: 1473–7. 16. Mair TS. The medical management of eight horses with grade 3 tears. Equine Vet J Suppl 2000; 32: 104–7. 17. Embertson RM, Hodge RJ, Vachon AM. Near circumferential retroperitoneal rectal tear in a pony. J Am Vet Med Assoc 1986; 188: 738–9. 18. Eastman TG, Taylor TS, Hooper RN, Hague BA. Treatment of grade 3 rectal tears in horses by direct suturing per rectum. Equine Vet Educ 2000; 12: 63–6. 19. Arnold J, Meagher D. Management of rectal tears in the horse. J Equine Med Surg 1978; 2: 64–71. 20. Spensley MS, Meagher DM, Hughes JP. Instrumentation to facilitate surgical repair of rectal tears in the horse: A preliminary report. Proc Am Assoc Equine Pract 1985; 31: 553–63. 21. Huskamp B. Die operative Versorgung von Mastdarmverletzungen. Prakt Tierarzt 1994; 11: 943–8. 22. Stewart RH, Robertson JT. Surgical stapling for repair of a rectal tear in a horse. J Am Vet Med Assoc 1990; 197: 746–8. 23. Wilson DG, Stone WC. Antimesenteric enterotomy for repair of a dorsal rectal tear in a mare. Can Vet J 1990; 31: 705–7. 24. Brugmans F, Deegen E. Laparoscopic surgical technique for repair of rectal and colonic tears in horses: An experimental study. Vet Surg 2001; 30: 409–16. 25. Taylor TS, Watkins JP, Schumacher J. Temporary indwelling rectal liner for use in horses with rectal tears. J Am Vet Med Assoc 1987; 191: 677–80. 26. Freeman DE, Richardson DW, Tulleners EP, Orsini JA, Donawick WJ, Madison JB, Ross MW, Beroza GA. Loop colostomy for management of rectal tears and small colon injuries in horses: 10 cases (1976–1989). J Am Vet Med Assoc 1992; 200: 1365–71. 27. Freeman DE, Robertson JT, Richardson DW, Donawick WJ, Tulleners EP, Orsini JA, Beroza GA. Loop colostomy for management of rectal and small colon injuries in horses. Proc Am Assoc Equine Pract 1992; 38: 133–50.
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Immunological Considerations Sara K. Lyle
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Abnormalities of Contractility Terttu Katila
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Abnormalities of Development John P. Hurtgen
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Breeding the Problem Mare Michelle M. LeBlanc and Angus O. McKinnon
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Fungal Endometritis Marco A. Coutinho da Silva and Marco A. Alvarenga
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Pyometra Kristina G. Lu
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Treatment of Fluid Accumulation Jonathan F. Pycock
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Uterine Cysts Mary E. Stanton
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Uterine Abnormalities John P. Hurtgen
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Laboratory Methods for Isolation and Evaluation of Bacteria, Fungi and Yeast David L. Hartman and Shalyn Bliss
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Chapter 268
Immunological Considerations Sara K. Lyle
Introduction The equine uterus is a mucous membrane; therefore, the predominant immunological response to disease is characteristic of innate immunity. This is an obvious advantage when breeding on successive cycles to the same stallion, where a strong adaptive response would likely preclude establishment of pregnancy. Likewise, vigorous adaptive immunity would be disadvantageous during pregnancy, i.e., the fetus as an allograft.1 From a historical perspective, much of the early work on defining the cause of susceptibility to endometritis in mares focused primarily on the innate response to infection, namely opsonins in uterine secretions and the function of uterine-derived neutrophils. Once physical or mechanical clearance (see Chapter 269) was determined to be an extremely important factor in the resolution of uterine infection,2–5 research emphasis shifted away from further investigation of the uterus as a mucosal immune system. In the meantime, the field of immunology has expanded exponentially, and compared to other species, there are many basic aspects of equine uterine mucosal immunity that remain to be elucidated. Cells involved with adaptive immunity have been identified in endometria, and the presence of mucociliary currents has been proposed; however, their role in endometritis and other diseases of the uterus is not yet clear.
teria, and inflammation is a transient phenomenon; such mares are considered resistant to bacterial endometritis regardless of dose or frequency of inoculation. Some mares, however, are unable to clear the bacteria, and a persistent endometritis results. These individuals are always susceptible to endometritis, and, without careful management to reduce and eliminate contamination of the uterus, have significantly reduced fertility. Considerable effort has been made to define the mechanisms by which the resistant mare eliminates bacteria and the possible etiology of dysfunction that results in susceptibility to endometritis. By defining these factors, specific therapy addressing this dysfunction in the susceptible mare will result in improvement in fertility. Studies to date have focused on the inflammatory response, the role of neutrophils and immunoglobulins, the composition of uterine secretions, and physical clearance. Mares classified as resistant to bacterial endometritis have been compared to those identified as susceptible to infection based on history, uterine biopsy, culture, and cytology. Some investigators have used mares under progesterone dominance to simulate the susceptible mare. Streptococcus equi ssp. zooepidemicus (S. zooepidemicus) is commonly used in most experimental models because of its relative ease of laboratory handling, and its identification as the most common etiological agent in bacterial endometritis.6
Endometritis Persistence of bacterial endometritis is an important cause of infertility and results in major economic losses to the equine industry. Massive challenge of the uterus with bacteria occurs at parturition and coitus. Generally, mares efficiently eliminate the bac-
Role of immunoglobulins in uterine defense mechanisms Immunoglobulin A (IgA),7–12 secretory IgA,10, 12 IgG,8–11,17 IgG(T),8–10 IgGa,7 IgGb,7 IgGc,7 and IgM8, 9 have been demonstrated in luminal secretions from
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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mares resistant and susceptible to endometritis. Secretory IgA comprises a mean of 60% of the total IgA present in uterine secretions.11 The former (and current) designations for the sub-isotypes of IgG are as follows: IgGa (IgG1, IgG2), IgG(T) (IgG3, IgG5), IgGb (IgG4), and IgGc (IgG6, IgG7).14 Demonstration of secretory IgA and higher Ig to albumin ratios in uterine secretions compared to those in serum supports the classification of the equine uterus as a local secretory immune system.11 Transudation of Ig and active transport may also be a significant source of intraluminal Ig.12 The relative contributions of active transport from serum versus local Ig production are largely unknown, but in vitro endometrial production of specific IgG, IgM, and IgA has been reported.15 The differences between immunoglobulins in uterine secretions in mares resistant or susceptible to endometritis are somewhat difficult to interpret due to differences in study design and sample populations. Infection with S. zooepidemicus caused substantial increases in IgM, IgG, and IgA in uterine flushings from ovariectomized mares following infection, with no concurrent change in serum titers.16 Following inoculation with S. zooepidemicus or chronic infection, uterine secretions from ovariectomized mares16 or susceptible mares8–11 had higher levels of IgA,8–11 IgG,8, 9 and IgG(T),8 yet failed to clear the infection. In contrast, no differences were observed in IgA,13,17 IgG,17 IgG(T),9, 10 or IgM9 levels between resistant and susceptible mares, although IgG was reduced in susceptible mares 36 hours after infection13,17 . In vitro, no differences were observed in titers of IgG, IgA, or IgM to S. zooepidemicus in serum or in culture supernatant of endometrial tissue from susceptible mares naturally infected (unknown duration) with S. zooepidemicus and estrogen- or progesterone-supplemented ovariectomized (‘resistant mares’).15 Despite the presence of higher titers of IgG, IgA, IgGT , and IgM specific for whole bacterial S. zooepidemicus antigens in uterine secretions from susceptible mares, opsonic activity was reduced compared to that of resistant mares,18 suggesting that consumption or inactivation of opsonins other than Ig may occur with persistent endometritis. Although not conclusive, the presence of free Ig staining within epithelial cells suggests an intracellular route of secretion into the uterus.19, 20 No significant difference in plasma cell numbers was seen between reproductively normal mares and those with persistent endometritis.21 Results of studies on immunoglobulins in luminal secretions and immunohistology of the equine endometrium raise some interesting points. First, it is apparent that antigen processing in the equine uterus is variable. Intrauterine inoculation with
S. zooepidemicus results in a local titer increase with no concurrent increase in systemic titer. Inoculation with Taylorella equigenitalis or dinitro-phenylated human serum albumin results in both local and systemic titer increases in antibody.22, 23 Despite no overall change in the peripheral titer, differences in the bactericidal activity of blood are seen following intrauterine inoculation with S. zooepidemicus; this activity is not only strain specific, but also comprises heat-stable and heat-labile components.24–26 Antigen processing and dendritic cell function in the equine uterus has not been studied. Second, since in general mares susceptible to endometritis have similar or higher levels of free Ig in the uterus, the common inference is that a dysfunction in the humoral response to intrauterine infection does not contribute significantly to increased susceptibility to endometritis. The suggestions that IgGT 15 or IgA18 may interfere with effective opsonization by specific antibody warrants further study. Investigation of bacterial proteins, such as immunoglobulin-binding proteins,27 which can reduce both antibody-mediated complement fixation and opsonization, would lead to a better understanding of the humoral response to intrauterine infection and its role in susceptibility to endometritis. Indeed, immunoglobulin-binding proteins have been identified in uterine secretions, and deplete hemolytic complement activity in vitro.28
Chemotactic properties of uterine secretions The chemotactic activity of uterine fluid is influenced by ovarian hormones. Fluid collected during estrus possessed chemotactic activity similar to that of the activated serum control, but significantly greater than that of diestrous uterine fluid.29 Uterine washes from estrogen-supplemented ovariectomized mares prior to inoculation with S. zooepidemicus stimulated random migration of neutrophils under agarose, while those from control or progesterone-supplemented mares inhibited chemokinetic activity.30 Infection,31 spermatozoa-induced complement activation,32 and susceptibility13, 33 appear to increase the chemoattractants in uterine secretions, and susceptibility increases the chemotactic activity irrespective of stage of cycle or presence of uterine infection.33 This pattern was thought to reflect the increased chemotactic activity present in uterine secretions from mares with active endometritis observed by Blue et al.31 Chemoattractants are composed of both heat-labile32, 35 and heat-stable35 components, and are dose dependent.35 This finding is specific to chemoattractants released by the endometrium since S. zooepidemicus itself does not possess chemotactic activity.36
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Opsonins in uterine secretions
Antibacterial activity of uterine secretions
Interest in characterization of opsonins in uterine luminal secretions began with the clinical response seen with the use of intrauterine plasma infusion in mares susceptible to endometritis.37 The conclusion was that susceptibility to uterine infection may be due in part to deficiency either in quality or quantity of opsonins in uterine secretions.37 In vitro, the addition of serum to uterine washings resulted in a significant increase in opsonization and phagocytosis by peripheral neutrophils, and this activity was reduced by complement depletion.38 Non-infected estrous fluid promoted greater phagocytosis than diestrous fluid,39 and the opsonic activity of susceptible mares was either increased33, 40 or decreased17 compared to that of resistant mares. The interaction between ovarian hormone status and susceptibility on concentrations of opsonins is complex with opsonins being reported as greater in resistant mares during diestrus and greater in susceptible mares during estrus,40 while culture supernatant of endometria from mares treated with progesterone was less effective at bacterial opsonization than supernatant from control mares or those treated with estrogen.16 The contribution of complement as an opsonin has also been controversial. In one study, opsonic activity was not reduced by heat inactivation, and the major opsonin was associated with IgG.40 However, when standardized for protein content, both complement and specific IgG were found to be important opsonins in uterine washes from reproductively normal non-cycling mares.41 The interesting observation of declining IgG concentration in susceptible mares by 36 hours of infection17 raises the question of whether decreased production or secretion, or increased consumption of IgG is involved with persistence of endometritis.
Although it has been reported that cell-free uterine flushes from mares in mid-diestrus possess antibacterial activity,29 other investigators have failed to demonstrate inherent antibacterial activity of uterine fluid from non-cycling mares41 or from E2 - or P4 -supplemented ovariectomized mares.43 The discrepancy in the results from these studies may lie in potential qualitative and quantitative differences between the uterine secretions from cycling and noncycling mares. Recently lactoferrin, an antimicrobial and immunomodulator member of the transferrin gene family that is expressed by epithelial cells and neutrophils, is stored in neutrophil granules, and has been identified in the equine endometrium.44 Lactoferrin’s antibacterial property lies in its ability to sequester free iron, thereby inhibiting bacterial growth. Although lactoferrin expression was upregulated during early estrus, protein staining was uninfluenced by cycle and was most intense in the glandular epithelium. Expression of lactoferrin was increased in mares with delayed physical clearance during early estrus,44 which might represent a response to inflammation. While intriguing from a perspective of host–pathogen interactions, unless a diminished response in lactoferrin expression and production were observed, it is unlikely to explain how this could be a factor in susceptibility to endometritis.
Complement in uterine secretions Examination of complement components and hemolytic activity has yielded conflicting results. Antigenic C3 and breakdown products of C3 were identified, yet no hemolytic complement (functional) activity was measured in uterine secretions,38 possibly due to the source of red blood cells. Hemolytic complement activity using porcine red blood cells was significantly greater in uterine flushings from susceptible mares compared to resistant mares, although individual variability was high and stage of cycle was variable.33 Lastly, immunoactive C3 was not significantly different between resistant and susceptible mares, although at 36 hours after bacterial challenge immunoactive C3 was greater in resistant mares.17
Modulators of uterine contractility: eicosanoids in uterine secretions In cycling,42 and E2 - or P4 -supplemented ovariectomized mares,45,46 uterine luminal immunoreactive prostaglandin F (PGF), PGE2 , and leukotriene B4 (LTB4 ) dramatically increase following an inflammatory insult. Progesterone-supplemented, or P4 -supplemented followed by E2 -supplemented ovariectomized mares had higher concentrations of immunoreactive PGF2α and PGE2 than those receiving E2 alone.46 Resistant mares have a greater release of PGF2α in response to antigenic stimulation (insemination) or oxytocin administration than do mares susceptible to persistent endometritis.47 Since LTB4 and PGE2 cause dose-related chemokinesis of peripheral neutrophils, PGF2 and LTB4 increase bactericidal activity,104 and LTB4 increases phagocytosis,42,104 their presence is unlikely to play a role in persistence of endometritis by adversely affecting neutrophil function. On the contrary, it has been suggested that increased eicosanoids in susceptible mares may contribute to enhanced phagocytic function.5, 13 Nitric oxide (NO), a bactericidal agent produced by macrophages and neutrophils following ingestion of microorganisms,48 can cause myometrial
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relaxation.49 Nitric oxide concentration was higher in uterine secretions from mares susceptible to delayed clearance than in resistant mares.50 Whether this is due to upregulation of NO synthase or due to accumulation of inflammatory byproducts in susceptible mares is not established, but it was speculated that NO accumulation may contribute to undesirable uterine relaxation following an antigenic challenge.50 Functional properties of neutrophils Uterine neutrophils from susceptible mares have been reported to possess either reduced51, 52 or equivalent13 chemokinetic function compared to resistant mares. Similarly, decreased33, 53 or increased13, 54 phagocytic function of uterine-derived neutrophils from susceptible mares have been reported. Clearly there is an interaction between stage of estrous cycle and category of mare, and both may have a profound effect on the functional capacity of uterine neutrophils. It is doubtful that some level of compromised uterine neutrophil function exists in the susceptible mare; the more likely explanation for findings to date is the effect of experimental design, specifically method of uterine neutrophil recruitment, stage of cycle, and differences in bacterial strain on in vitro neutrophil function. Indeed, when a chemokine (recombinant human interleukin8) was used to recruit neutrophils into the uterus not only was there no difference in phagocytic capacity of uterine neutrophils between normal mares and those with degenerative changes, but also the capacity of uterine neutrophils from mares with degenerative uterine changes to generate reactive oxygen species was enhanced relative to peripheral neutrophils or uterine neutrophils from normal mares.55 Diminished cellular clearance occurs most likely secondary to persistence of infection and continued bacterial growth, which exceeds the capability of normal phagocytic function. Evidence of an adaptive immune response in the uterus The endometrium of the mare is inhabited by T lymphocytes.56–60 Genitally normal mares had greater numbers of CD4+ lymphocytes (TH cells; ‘helper T-cells’) and CD8+ cells (TC cells; ‘cytotoxic/suppressor T-cells’) in the stratum compactum than in the stratum spongiosum;58 the density of cells was either increased during estrus57 or found to be independent of cycle stage;58 was not influenced by age;59 and greater numbers of CD4+ and CD8+ cells were present in the uterine body than in the uterine horns.59 Following insemination60 or in the presence of endometritis58 the number of CD4+ cells58, 60 and
CD8+ cells58 increased, the number of CD4+ cells doubled in lymphoid aggregates, and the number of CD8+ cells in the luminal and glandular epithelium increased,58 suggesting that an adaptive response is initiated following antigenic stimulation of the endometrium. Exogenous antigens, such as those of S. zooepidemicus, are processed through an endocytic pathway and are presented on the surface of the membrane of cells that express class II major histocompatibility complex (MHC) molecules.61 Presentation of class II MHC molecules to CD4+ TH cells is restricted to ‘professional’ antigen-presenting cells (APCs), namely dendritic cells, macrophages, and B-cells. ‘Non-professional’ APCs, such as certain epithelial cells, fibroblasts, and vascular endothelial cells, can present antigen in the context of class II MHC molecules, but fail to deliver a co-stimulatory signal and present antigen for brief periods during a sustained inflammatory response. Cells in the equine endometrium expressing MHC class II staining include macrophages, monocytes, dendritic cells, lymphocytes, vascular endothelial cells, and uterine luminal epithelium.56 Expression of MHC II was greater in genitally normal mares in estrus than in diestrus; the distribution was predominantly beneath the luminal epithelium, with occasional foci in the stratum compactum and rarely in the stratum spongiosum.58 This pattern is consistent with antigen presentation. There was no difference in expression between infected and non-infected mares,58 which is similar to findings from murine models of intestinal salmonellosis, where little change from the steady-state numbers of dendritic cells is seen with infection.62 Cytokine expression in response to infection and insemination Increased endometrial expression of the proinflammatory cytokines interleukin-1β (IL-1β), IL-6, tumor necrosis factor-α (TNF-α), and IL-8 during estrus, increased IL-1β, TNF-α, and IL-8 during diestrus, and decreased expression of the anti-inflammatory cytokine IL-10 during estrus was thought to contribute to susceptibility to endometritis,63, 64 although non-infected susceptible mares were not examined in these studies. This pattern would be predicted to occur with chronic-active inflammation; however, re-examination of mares susceptible to endometritis prior to initiation of an inflammatory insult would be useful in understanding the potential roles of these cytokines during persistent endometritis. If elevated IL-8 expression and diminished IL-10 expression in susceptible mares in the absence of infection or inflammation exists, a potential factor in susceptibility
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Figure 268.2 Bouin’s fixed endometrial biopsy of a normal mare in estrus showing a mucus blanket carried on ciliated cells. Stain: alcian blue pH 2.5; space bar = 20 µm; original magnification = ×400. Reproduced courtesy of Dr Robert C. Causey, University of Maine, Orono, ME. (See color plate 123).
in the mucociliary apparatus in mares susceptible to endometritis.
Persistent endometrial cups Figure 268.1 Bouin’s fixed endometrial biopsy of a chronically infertile mare, showing increased mucous secretions. Stain: alcian blue pH 2.5; space bar = 20 µm; original magnification = ×1000. Reproduced courtesy of Dr Robert C. Causey, University of Maine, Orono, ME. (See color plate 122).
to endometritis may be explained. These studies are a first step toward investigating the cytokine signaling involved with endometritis, and clearly further study is needed in this area. Mucociliary clearance in the mare It has been recently suggested that mucociliary clearance may play an important role in the clearance of infections and that disruption of mucociliary currents in the susceptible mare leads to persistence of infection.65 While mucus-producing cells and ciliated cells beneath a mucus blanket suggests a functional mucociliary apparatus in the equine uterus,66 in vitro documentation of ciliary movement is lacking.65 Mucus production and secretion is increased during estrus66 and inflammation,66, 67 and the optical density of the mucus is increased in mares with delayed clearance (Figures 268.1 and 268.2).66 The biological significance of alterations in the mucus blanket is unknown. The effects of fluid accumulations,68 intrauterine therapy,69 and uterine pathogens68, 70, 71 on mucociliary clearance are deserving of future research, as is whether or not there is a primary defect
Following the high rate of fetal wastage between days 40 and 90 due to mare reproductive loss syndrome in central Kentucky during the 2001 breeding season,72–75 some mares experienced a persistence (some greater than 1 year) of endometrial cups beyond the expected time of sloughing (120–140 days gestation).76 This syndrome is not peculiar to abortion due to MRLS.77 The incidence of persistent endometrial cups is unknown, but an estimate of the prevalence based on early fetal losses in 2001 of 299878 and a case report of 11 mares with lengthy persistence of cups76 would be 0.4%. If this estimate is close to the actual incidence, considerable difficulty exists in reproducing this syndrome for further study. Invasive trophoblast cells of the endometrial cups typically express paternally derived MHC class I antigens, although expression is considerably downregulated by day 45.79 Even ectopically transplanted trophoblast cells show evidence of downregulation, and survival time is double compared to a conventional skin allograft.80 The mechanism of downregulation of MHC class I antigens resulting in evasion of destruction by maternal CD8+ T-cells until 120 days of gestation is still unknown, although there is evidence that the immune response to the conceptus is asymmetric, with vigorous humoral immunity and diminished cellular immunity.81 Whether persistence of endometrial cups is primarily an immunological phenomenon, or represents evidence that the lifespan of the endometrial cup can be considerably longer than previously thought, is yet to be determined.
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Placentitis Placentitis has been reported to be responsible for 9.8,82 19.4,83 24.7,84 and 33.5%85 of abortions, stillbirths, and perinatal loss in horses. Fifty-three percent of these losses were due to bacterial infection; S. zooepidemicus was isolated in 28% of these cases.84 The localization of placentitis to the cervical star was present in 95% of cases, supporting the argument that ascension of aerobic bacteria through the vagina and cervix is the most frequent route of infection.86 Empirical treatment for placentitis includes progestins to maintain uterine quiescence, broad-spectrum antibiotics to eliminate bacterial infection, and anti-inflammatory agents to prevent prostaglandin synthesis. Although these agents should address the basic problems of infection-induced prostaglandin synthesis, which leads to uterine contractions and fetal expulsion, the efficacy of these regimes can be poor.87 In most species studied, an increase in proinflammatory cytokines has been observed in fetal fluids in response to inflammation or infection.88–94 One of the proposed mechanisms of infectioninduced pre-term delivery is that pro-inflammatory cytokines (IL-1, IL-6, TNF-α) from the decidua and cortisol from the mother cause activation of the fetal hypothalamic–pituitary–adrenal axis, causing elevation of fetal cortisol.95 Throughout pregnancy, prostaglandin dehydrogenase (PGDH) converts PGE2 and PGF2 α to inactive metabolites, thereby maintaining myometrial quiescence. Increased fetal cortisol, either at term or in response to proinflammatory cytokines, decreases levels of PGDH, leading to increased PGE2 and PGF2α and subsequent myometrial contractions. Pro-inflammatory cytokines also increase the activity of prostaglandin H2 synthase (PGHS-2, COX-2), which increases prostaglandin synthesis. The decrease in PGDH and increase in PGHS-2 leads to increased PGE2 and PGF2α synthesis and release, myometrial contractions, and premature labor.92,96–98 Two experimental models examining the relationship between pro-inflammatory cytokines and placentitis in the mare between 70–80% of gestation with S. zooepidemicus, either intracervically99 or at the surface of the chorioallantois,100 support the premise that bacterial infection causes increased expression of pro-inflammatory cytokines leading to the release of PGE2 and PGF2α into the allantoic fluid and premature labor. Infection produced no significant differences in fetal fluid activities of soluble TNF-α IL1, and IL-6,99 or increased concentration of soluble TNF-α protein in only 4% of all samples.100 Infection increased the expression of IL-6 and IL-899 and monocyte chemoattractant protein-1100 at the cervical star,
and IL-6,99 IL-18100 and IL-β100 at the placental body. Abortion increased the expresion of IL-18 at normal sites, IL-1β and IL-18 at abnormal sites, and inducible nitric oxide synthase at the cervical star.100 Increased concentration of cortisol in fetal fluids suggests that the fetal hypothalamic-pituitary-adrenal axis is activated susequent to infection or inflammation.100 The failure to demonstrate consistent increases in the concentrations of pro-inflammatory cytokines in equine fetal fluids following bacterial infection is in distinct contrast to the results from other species,89–94 and the reasons for this are unclear. The temporal relationship between elevations in allantoic cytokines and prostaglandins is quite important. If the synthesis of PGF2α and PGE2 is driven by cytokines in concert with fetal cortisol, as has been suggested,95 then treatment strategies that fail to interrupt the inflammatory cascade and reduce fetal stress may be ineffective in preventing pre-term delivery.
Conclusions While delayed physical clearance is undoubtedly a critical factor in persistent endometritis, it is likely that the failure to resolve inflammation and infection is multifactorial and immunological responses are important for clearance of microbial infections. Clinically, mares are seen for whom aggressive therapy to achieve physical clearance fails to resolve endometritis. From the foregoing discussion, there are several areas where knowledge is desperately needed. Antigen presentation by immune and nonimmune cells of the endometrium has implications both in the pathogenesis of endometritis and the persistence of endometrial cups. What is the role of the adaptive immune response in endometritis, persistent endometrial cups, and placentitis? The role of mucociliary clearance and the possibility that abnormal mucus production may play a role in persistence of infection in infertile mares highlights the lack of knowledge of host–pathogen interactions at the endometrial surface. Are uterine pathogens such as S. zooepidemicus and E. coli able to form a biofilm similar to other genital pathogens,101 thereby providing an avenue of escape from recognition by phaygocytes and resistance to antimicrobial agents? Toll-like receptors are membrane-spanning receptors on the surface of immune and non-immune cells that recognize pathogen-associated molecular patterns, such as lipopolysaccharide, bacteria, viruses, and bacterial DNA, among others. Upon binding, they signal through nuclear factor κB (NFκB) to upregulate proinflammatory cytokines and are a key component of the innate response to infection. Toll-like receptor 9 (TLR-9) expression has been described in equine
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Immunological Considerations dendritic cells and macrophages,102 and TLR-4 expression has been demonstrated in airway epithelial cells.103 Presumably they are present in the endometrium as well, and their role in the pathogenesis of endometritis and placentitis, and as a potential target for immunotherapy, is intriguing.
References 1. Blois SM, Kammerer U, Alba Soto C, Tometten MC, Shaikly V, Barrientos G, Jurd R, Rukavina D, Thomson AW, Klapp BF, Fernandez N, Arck PC. Dendritic cells: key to fetal tolerance? Biol Reprod 2007;77:590–8. 2. Evans MJ, Hamer JM, Gason LM, Graham CS, Asbury AC, Irvine CH. Clearance of bacteria and non-antigenic markers following intra-uterine inoculation into maiden mares: Effect of steroid hormone environment. Theriogenology 1986;26:37–50. 3. LeBlanc MM, Asbury AC, Lyle SK. Uterine clearance mechanisms during the early postovulatory period in mares. Am J Vet Res 1989;50:864–7. 4. Troedsson MH, Liu IK. Uterine clearance of nonantigenic markers (51Cr) in response to a bacterial challenge in mares potentially susceptible and resistant to chronic uterine infections. J Reprod Fertil Suppl 1991;44:283–8. 5. Troedsson MH, Liu IK, Ing M, Pascoe J, Thurmond M. Multiple site electromyography recordings of uterine activity following an intrauterine bacterial challenge in mares susceptible and resistant to chronic uterine infection. J Reprod Fertil 1993;99:307–13. 6. Dimock WW, Edwards PR. The pathology and bacteriology of the reproductive organs of mares in relation to sterility. Bulletin Kentucky Agricultural Experiment Station 1928;286:157–63. 7. Kenney RM, Kahleel SA. Bacteriostatic activity of the mare uterus: a progress report on immunoglobulins. J Reprod Fertil Suppl 1975;23:357–8. 8. Asbury AC, Halliwell RE, Foster GW, Longino SJ. Immunoglobulins in uterine secretions of mares with differing resistance to endometritis. Theriogenology 1980;14:299–308. 9. Mitchell G, Liu IK, Perryman LE, Stabenfeldt GH, Hughes JP. Preferential production and secretion of immunoglobulins by the equine endometrium – a mucosal immune system. J Reprod Fertil Suppl 1982;32:161–8. 10. Williamson P, Dunning A, O’Connor J, Penhale WJ. Immunoglobulin levels, protein concentrations and alkaline phosphatase activity in uterine flushings from mares with endometritis. Theriogenology 1983;19:441–8. 11. Widders PR, Stokes CR, David JS, Bourne FJ. Quantitation of the immunoglobulins in reproductive tract secretions of the mare. Res Vet Sci 1984;37:324–30. ´ AM, Rodriguez-Martinez H, Hult´en C, Num12. Tunon mijarvi A, Magnusson U. Concentrations of total protein, albumin and immunoglobulins in undiluted uterine fluid of gynecologically healthy mares. Theriogenology 1998;50:821–31. 13. Troedsson MH, Liu IK, Thurmond M. Function of uterine and blood-derived polymorphonuclear neutrophils in mares susceptible and resistant to chronic uterine infection: phagocytosis and chemotaxis. Biol Reprod 1993;49:507–14.
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14. Wagner B. Immunoglobulins and immunoglobulin genes of the horse. Dev Comp Immunol 2006;30:155–64. 15. Watson ED, Stokes CR. Effect of susceptibility to endometritis on specific antibody in the endometria of mares. Theriogenology 1990;34:39–45. 16. Watson ED. The influence of estrogen and progesterone on antibody synthesis by the endometrium of the mare. In: Equine Infectious Diseases V. Lexington, KY: The University Press of Kentucky, 1987; pp. 181–5. 17. Troedsson MH, Liu IK, Thurmond M. Immunoglobulin (IgG and IgA) and complement (C3) concentrations in uterine secretion following an intrauterine challenge of Streptococcus zooepidemicus in mares susceptible to versus resistant to chronic uterine infection. Biol Reprod 1993;49:502–6. 18. LeBlanc M, Ward L, Tran T, Widders P. Identification and opsonic activity of immunoglobulins recognizing Streptococcus zooepidemicus antigens in uterine fluids of mares. J Reprod Fertil Suppl 1991;44:289–96. 19. Widders PR, Stokes CR, David JS, Bourne FJ. Immunohistological studies of the local immune system in the reproductive tract of the mare. Res Vet Sci 1985;38: 88–95. 20. Waelchli RO, Winder NC. Immunohistochemical evaluation of the equine endometrium during the oestrous cycle. Equine Vet J 1987;19:299–302. 21. Watson ED, Stokes CR. Plasma cell numbers in uteri of mares with persistent endometritis and in ovariectomised mares treated with ovarian steroids. Equine Vet J 1988;20:424–5. 22. Widders PR, Stokes CR, David JSE, Bourne FJ. Specific antibody in the equine genital tract following systemic and local immunization. Immunology; 54(4):763– 769. 23. Widders PR, Stokes CR, David JS, Bourne FJ. Specific antibody in the equine genital tract following local immunisation and challenge infection with contagious equine metritis organism (Taylorella equigenitalis). Res Vet Sci 1986;40:54–8. 24. Causey RC, Paccamonti DL, Todd WJ. Antiphagocytic properties of uterine isolates of Streptococcus zooepidemicus and mechanisms of killing in freshly obtained blood of horses. Am J Vet Res 1995;56:321–8. 25. Causey R, Weber J, Emmans E, Small P, Knapp K, Pelletier D. Immunoblotting of Streptococcus zooepidemicus antigens following intrauterine inoculation in mares. Theriogenology 2002;58:487–9. 26. Causey RC, Weber JA, Emmans EE, Stephenson LA, Homola AD, Knapp KR, Crowley IF, Pelletier DC, Wooley NA. The equine immune response to Streptococcus equi subspecies zooepidemicus during uterine infection. Vet J 2006;172:248–57. 27. Boyle MDP. Complement activation and bacterial immunoglobulin-binding proteins. In: Boyle MDP (ed.) Bacterial Immunoglobulin-Binding Proteins. Vol. 1. Microbiology, Chemistry, and Biology. Academic Press, 1990; pp. 295–304. 28. Lyle SK. Investigations on the uterine environment of the mare and its relationship to defense mechanisms. University of Florida, Gainesville, FL: Master’s thesis, 1991; pp. 1–98. 29. Strzemienski PJ, Do D, Kenney RM. Antibacterial activity of mare uterine fluid. Biol Reprod 1984;31:303–11. 30. Watson ED. Effect of ovarian steroids on migration of uterine luminal neutrophils and on chemokinetic
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factors in uterine secretions from mares. Equine Vet J 1988;20:368–70. Blue HB, Blue MG, Kenney RM, Merritt TL. Chemotactic properties and protein of equine uterine fluid. Am J Vet Res 1984;45:1205–8. Troedsson MHT, Steiger BN, Ibrahim NM, King VL, Foster DN, Crabo BG. Mechanism of sperm-induced endometritis in the mare. Biol Reprod Suppl 1995;52:307. Watson ED, Stokes CR, Bourne FJ. Cellular and humoral defence mechanisms in mares susceptible and resistant to persistent endometritis. Vet Immunol Immunopathol 1987;16:107–21. Watson ED, Stokes CR, Bourne FJ. Influence of arachidonic acid metabolites in vitro and in uterine washings on migration of equine neutrophils under agarose. Res Vet Sci 1987;43:203–7. Pycock JF, Allen WE. Pre-chemotactic and chemotactic properties of uterine fluid from mares with experimentally induced endometritis. Vet Rec 1988;123:193–5. Pycock JF, Allen WE. Equine neutrophil locomotion in response to Streptococcus zooepidemicus. Equine Vet J 1988;20:448–50. Asbury AC. Uterine defense mechanisms in the mare: The use of intrauterine plasma in the management of endometritis. Theriogenology 1984;21:387–93. Asbury AC, Gorman NT, Foster GW. Uterine defense mechanisms in the mare: Serum opsonins affecting phagocytosis of Streptococcus zooepidemicus by equine neutrophils. Theriogenology 1984;21:375–85. Blue MG, Brady AA, Davidson JN, Kenney RM. Studies on the composition and antibacterial activity of uterine fluid from mares. J Reprod Fertil Suppl 1982;32:143–9. Hansen PJ, Asbury AC. Opsonins of Streptococcus in uterine flushings of mares susceptible and resistant to endometritis: control of secretion and partial characterization. Am J Vet Res 1987;48:646–50. Watson ED. Opsonins in uterine washings influencing in vitro activity of equine neutrophils. Equine Vet J 1988;20:435–7. Watson ED, Stokes CR, David JS, Bourne FJ, Ricketts SW. Concentrations of uterine luminal prostaglandins in mares with acute and persistent endometritis. Equine Vet J 1987;19:31–7. Johnson JU, Oxender WD, Berkhoff HA. Influence of estrogen on antibacterial and immunoglobulin secretory activities of uterine fluids from ovariectomized mares. Am J Vet Res 1994;55:643–9. Kolm G, Klein D, Knapp E, Watanabe K, Walter I. Lactoferrin expression in the horse endometrium: relevance in persisting mating-induced endometritis. Vet Immunol Immunopathol 2006;114:159–67. Watson ED, Stokes CR, Bourne FJ. Concentrations of immunoreactive leukotriene B4 in uterine lavage fluid from mares with experimentally induced and naturally occurring endometritis. J Vet Pharmacol Ther 1988;11: 130–4. Watson ED, Stokes CR, Bourne FJ. Effect of exogenous ovarian steroids on the uterine luminal prostaglandins in ovariectomised mares with experimental endometritis. Res Vet Sci 1988;44:361–5. Nikolakopoulos E, Kindahl H, Watson ED. Oxytocin and PGF2a in mares resistant and susceptible to persistent mating-induced endometritis. J Reprod Fertil Suppl 2000;56:363–72.
48. Janeway CA, Travers P, Walport M, Shlomchik MJ. Immunobiology: the Immune System in Health and Disease. New York: Garland Science Publishing, 2005. 49. Demirkoprulu N, Cetin M, Bagcivan I, Kaya T, Soydan AS, Karadas B, Cetin A. Comparative relaxant effects of YC-1 and DETA/NO on spontaneous contractions and the levels of cGMP of isolated pregnant rat myometrium. Eur J Pharmacol 2005;517:240–5. 50. Alghamdi AS, Troedsson MHT. Concentration of nitric oxide in uterine secretion from mares susceptible and resistant to chronic post-breeding endometritis. Theriogenology 2002;58:445–8. 51. Liu IK, Cheung AT, Walsh EM, Miller ME, Lindenberg PM. Comparison of peripheral blood and uterinederived polymorphonuclear leukocytes from mares resistant and susceptible to chronic endometritis: chemotactic and cell elastimetry analysis. Am J Vet Res 1985;46:917–20. 52. Liu IK, Cheung AT, Walsh EM, Ayin S. The functional competence of uterine-derived polymorphonuclear neutrophils (PMN) from mares resistant and susceptible to chronic uterine infection: a sequential migration analysis. Biol Reprod 1986;35:1168–74. 53. Cheung AT, Liu IK, Walsh EM, Miller ME. Phagocytic and killing capacities of uterine-derived polymorphonuclear leukocytes from mares resistant and susceptible to chronic endometritis. Am J Vet Res 1985;46:1938–40. 54. Asbury AC, Hansen PJ. Effects of susceptibility of mares to endometritis and stage of cycle on phagocytic activity of uterine-derived neutrophils. J Reprod Fertil Suppl 1987;35:311–16. 55. Zerbe H, Engelke F, Klug E, Schoon HA, Leibold W. Degenerative endometrial changes do not change the functional capacity of immigrating uterine neutrophils in mares. Reprod Domest Anim 2004;39:94–8. 56. Watson ED, Dixon CE. An immunohistological study of MHC class II expression and T lymphocytes in the endometrium of the mare. Equine Vet J 1993;25:120–4. 57. Frayne J, Stokes CR. MHC Class II positive cells and T cells in the equine endometrium throughout the oestrous cycle. Vet Immunol Immunopathol 1994;41:55–72. 58. Watson ED, Thomson SR. Lymphocyte subsets in the endometrium of genitally normal mares and mares susceptible to endometritis. Equine Vet J 1996;28:106–10. 59. Tunon AM, Rodriguez-Martinez H, Nummijarvi A, Magnusson U. Influence of age and parity on the distribution of cells expressing major histocompatibility complex class II, CD4, or CD8 molecules in the endometrium of mares during estrus. Am J Vet Res 1999;60:1531–5. ´ AM, Katila T, Magnusson U, Nummij¨arvi A, 60. Tunon Rodriguez-Martinez H. T-cell distribution in two different segments of the equine endometrium 6 and 48 hours after insemination. Theriogenology 2000;54:835–41. 61. Kindt TJ, Goldsby RA, Osborne BA. Kuby Immunology. New York: W.H. Freeman & Co., 2007. 62. Wick MJ. Monocyte and dendritic cell recruitment and activation during oral Salmonella infection. Immunol Lett 2007;112:68–74. 63. Fumuso E, Giguere S, Wade J, Rogan D, Videla-Dorna I, Bowden RA. Endometrial IL-1beta, IL-6 and TNF-alpha, mRNA expression in mares resistant or susceptible to post-breeding endometritis. Effects of estrous cycle, artificial insemination and immunomodulation. Vet Immunol Immunopathol 2003;96:31–41.
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Immunological Considerations 64. Fumuso EA, Aguilar J, Giguere S, Rivulgo M, Wade J, Rogan D. Immune parameters in mares resistant and susceptible to persistent post-breeding endometritis: effects of immunomodulation. Vet Immunol Immunopathol 2007;118:30–9. 65. Causey RC. Mucus and the mare: how little we know. Theriogenology 2007;68:386–94. 66. Causey RC, Ginn PS, Katz BP, Hall BJ, Anderson KJ, LeBlanc MM. Mucus production by endometrium of reproductively healthy mares and mares with delayed clearance. J Reprod Fert Suppl 2000;56:333–339. 67. Freeman KP, Roszel JF, Slusher SH, Castro M. Variation in glycogen and mucins in the equine uterus related to physiologic and pathologic conditions. Theriogenology 1990;33:799–808. 68. Troedsson MH, Liu IK. Measurement of total volume and protein concentration of intrauterine secretion after intrauterine inoculation of bacteria in mares that were either resistant or susceptible to chronic uterine infection. Am J Vet Res 1992;53:1641–4. 69. Al-Bagdadi FK, Eilts BE, Richardson GF. Scanning electron microscopy of the endometrium of mares infused with gentamicin. Microsc Microanal 2004;10:280–5. 70. Dimock WW, Edwards PR. Genital infection in mares by an organism of the encapsulatus group. J Am Vet Med Assoc 1927;70:469–80. 71. Pycock JF, Allen WE. Inflammatory components in uterine fluid from mares with experimentally induced bacterial endometritis. Equine Vet J 1990;22:422–5. 72. Cohen ND, Carey VJ, Donahue JG, Seahorn JL, Konahoe JK, Williams DM, Harrison LR. Case-control study of late-term abortions associated with mare reproductive loss syndrome in central Kentucky. J Am Vet Med Assoc 2003;222:199–209. 73. Dwyer RM, Garber LP, TraubDargatz JL, Meade BJ, Powell D, Pavlick MR, Kane AJ. Case-control study of factors associated with excessive proportions of early fetal losses associated with mare reproductive loss syndrome in central Kentucky during 2001. J Am Vet Med Assoc 2003;222:613–19. 74. Zent WW. An overview of reproductive system changes during and after mare reproductive loss syndrome. In: Proceedings of the First Workshop on Mare Reproductive Loss Syndrome, 2003, pp. 30–1. 75. Webb BA, Barney WE, Dahlman DL, DeBorde SN, Weer C, Williams NM, Donahue JM, McDowell KJ. Eastern tent caterpillars (Malacosoma americanum) cause mare reproductive loss syndrome. J Insect Physiol 2004;50: 185–93. 76. Steiner J, Antczak DF, Wolfsdorf K, Seville S, Brooks D, Miller E, Bailey E, Zent W. Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. 77. Mitchell D, Betteridge KJ. Resistance of endometrial cups and serum gonadotrophin following abortion in the mare. In: Proceedings of the 7th International Congress of Animal Reproduction and Artificial Insemination, 1972, pp. 567–70. 78. Riddle WT. Clinical observations associated with early fetal loss in mare reproductive loss syndrome during the 2001 and 2002 breeding seasons. In: Proceedings of the First Workshop on Mare Reproductive Loss Syndrome, 2003, pp. 12–14. 79. Donaldson WL, Oriol JG, Plavin A, Antczak DF. Developmental regulation of class I major histocompatibil-
80.
81.
82.
83.
84.
85.
86.
87.
88.
89.
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91.
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ity complex antigen expression by equine trophoblastic cells. Differentiation 1992;52:69–78. Adams AP, Antczak DF. Ectopic transplantation of equine invasive trophoblast. Biol Reprod 2001;64:753– 63. Adams AP, Oriol JG, Campbell RE, Oppenheim YC, Allen WR, Antczak DF. The effect of skin allografting on the equine endometrial cup reaction. Theriogenology 2007;68:237–47. Smith KC, Blunden AS, Whitwell KE, Dunn KA, Wales AD. A survey of equine abortion, stillbirth and neonatal death in the UK from 1988 to 1997. Equine Vet J 2003;35:496–501. Hong CB, Donahue JM, Giles RC Jr, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Equine abortion and stillbirth in central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203:1170–5. Hong CB, Donahue JM, Giles RC Jr, Petrites-Murphy MB, Poonacha KB, Roberts AW, Smith BJ, Tramontin RR, Tuttle PA, Swerczek TW. Etiology and pathology of equine placentitis. J Vet Diagn Invest 1993;5:56–63. Whitwell KE. Infective placentitis in the mare. In: Powell DG (ed.) Equine Infectious Dieseases V. Lexington, KY: University Press of Kentucky, 1988; pp. 172–80. Renaudin CD, Liu IKM, Troedsson MHT, Schrenzel MD. Transrectal ultrasonographic diagnosis of ascending placentitis in the mare: a report of two cases. Equine Vet Educ 1999;11:69–74. Gendron RL, Nestel FP, Lapp WS, Baines MG. Lipopolysaccharide-induced fetal resorption in mice is associated with the intrauterine production of tumour necrosis factor-alpha. J Reprod Fertil 1990;90:395– 402. McDuffie RS, Sherman MP Jr, Gibbs RS. Amniotic fluid tumor necrosis factor-alpha and interleukin-1 in a rabbit model of bacterially induced preterm pregnancy loss. Am J Obstet Gynecol 1992;167:1583–8. Romero R, Mazor M, Brandt F, Sepulveda W, Avila C, Cotton DB, Dinarello CA. Interleukin-1 alpha and interleukin-1 beta in preterm and term human parturition. Am J Reprod Immunol 1992;27:117–23. Gravett MG, Witkin SS, Haluska GJ, Edwards JL, Cook MJ, Novy MJ. An experimental model for intraamniotic infection and preterm labor in rhesus monkeys. Am J Obstet Gynecol 1994;171:1660–7. Romero R, Gomez R, Ghezzi F, Yoon B, Mazor M, Edwin S, Berry S. A fetal systemic inflammatory response is followed by the spontaneous onset of preterm parturition. Am J Obstet Gynecol 1998;179:186–93. Reznikov LL, Fantuzzi G, Selzman CH, Shames BD, Barton HA, Bell H, McGregor JA, Dinarello CA. Utilization of endoscopic inoculation in a mouse model of intrauterine infection-induced preterm birth: role of interleukin 1beta. Biol Reprod 1999;60:1231–8. Grigsby PL, Hirst JJ, Scheerlinck JP, Phillips DJ, Jenkin G. Fetal responses to maternal and intra-amniotic lipopolysaccharide administration in sheep. Biol Reprod 2003;68:1695–702.
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95. Challis JRG, Matthews SG, Gibb W, Lye SJ. Endocrine and paracrine regulation of birth at term and preterm. Endocr Rev 2000;21:514–50. 96. Keelan JA, Sato T, Mitchell MD. Interleukin (IL)-6 and IL8 production by human amnion: regulation by cytokines, growth factors, glucocorticoids, phorbol esters, and bacterial lipopolysaccharide. Biol Reprod 1997;57:1438– 44. 97. Gravett M, Hitti J, Hess D, Eschenbach D. Intrauterine infection and preterm delivery: Evidence for activation of the fetal hypothalamic-pituitary-adrenal axis. Am J Obstet Gynecol 2000;182:1404–13. 98. Gibb W, Challis JR. Mechanisms of term and preterm birth. J Obstet Gynaecol Can 2002;24:874–83. 99. LeBlanc MM, Giguere S, Brauer K, Paccamonti DL, Horohov DW, Lester GD, O’Donnell LJ, Sheerin BR, Pablo L, Rodgerson DH. Premature delivery in ascending placentitis is associated with increased expression of placental cytokines and allantoic fluid prostaglandins E2 and F2alpha. Theriogenology 2002;58: 841–4.
100. Lyle SK, Gentry LR, Horohov DW, Johnson JR, Eilts BE, Godke RA, Paccamonti DL. Prelminary evidence of fetal hypothalamic-pituitary-adrenal axis activation in an experimental model of infective preterm delivery in the mare. Clin Theriogenol 2009;1:238. 101. Greiner LL, Edwards JL, Shao J, Rabinak C, Entz D, Apicella MA. Biofilm formation by Neisseria gonorrhoeae. Infect Immun 2005;73:1964–70. 102. Flaminio MJ, Borges AS, Nydam DV, Horohov DW, Hecker R, Matychak MB. The effect of CpG-ODN on antigen presenting cells of the foal. J Immune Based Ther Vaccines 2007;5:1. 103. Berndt A, Derksen FJ, Venta PJ, Ewart S, YuzbasiyanGurkan V, Robinson NE. Elevated amount of Toll-like receptor 4 mRNA in bronchial epithelial cells is associated with airway inflammation in horses with recurrent airway obstruction. Am J Physiol Lung Cell Mol Physiol 2007;292:L936–43. 104. Watson ED. Eicosanoids and phagocytic and bactericidal activity of equine neutrophils. Eq Vet J 1988;20(5):373–375.
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Chapter 269
Abnormalities of Contractility Terttu Katila
The most important task of the uterus is to harbor and nourish the fetus during pregnancy. When the time is due, the foal is expelled by strong myometrial contractions. After the delivery, uterine contractions are required in the expulsion of the placenta and evacuation of fetal fluids, blood, dead cells, and contaminants. When it is time to breed the female again, uterine contractions carry spermatozoa through the uterus into the utero-tubal junction (UTJ). Spermatozoa that have not gained access into the oviducts become unwanted waste; the uterus has to remove them and also remove any bacteria introduced into the uterus in connection with breeding. Spermatozoa induce an inflammatory reaction of the uterus characterized by an intensive influx of polymorphonuclear leukocytes (PMNs). Once the leukocytes have carried out their purpose, they and inflammatory byproducts have to be removed rapidly. Although acute inflammation is an important part of uterine defense, it has to be transient. When the embryo arrives in the uterus 6 days after ovulation, no traces of inflammation can be present. Uterine contractions seem to have a vital role in the cleansing of the uterus after breeding. Deficient contractility is reflected as delayed uterine clearance (DUC) and is diagnosed by ultrasonically visible intrauterine fluid accumulations (IUFAs). Researchers have tried to explore mechanisms leading to DUC, e.g., the effects of age and parity of mares, position of the uterus, fibrosis of the uterus, lymphatic clearance, uterine contractions, and neurogenic and myogenic factors have been investigated. This chapter is limited to the non-pregnant mare and focuses on the role of uterine contractility and its abnormalities on post-breeding endometritis.
Physiology and measurement of myometrial contractility The myometrium consists of two layers of smooth muscle: the outer longitudinal layer and the inner cir-
cular layer. Contraction of the muscle shortens the myometrium, which in turn decreases the diameter of the uterine lumen.1 Intrauterine pressure transducers (IUPs) record these pressure changes2 and are used to measure uterine contractions, but the sensor is a foreign body in the uterus and may provoke contractions by its mere presence. The contractility of the uterus is a consequence of the electrical activity in the muscle cells of the myometrium. It is characterized by cyclic depolarization and repolarization of the plasma membrane (action potentials). Depolarization is due to an increased permeability to Ca2+ , resulting in high intracellular Ca2+ levels. The membrane repolarizes by increasing the permeability to K+ , which results in the movement of K+ out of the cell.1 Spontaneous depolarization of pacemaker cells initiates this contractile mechanism. The pacemaker cells in the myometrium are not anatomically fixed – they can move from one place to another.3 Electromyography (EMG) measures uterine electrical activity, which is reflected as activity peaks. A burst consists of ≥ 10 peaks. The frequency, intensity, and amplitude of the burst as well as synchronous and total activity give additional information about the type of contractility.4 The magnitude of contractions is dependent on the number of simultaneously and synchronously active muscle cells. Gap junctions are channels between muscle cells and mediate ionic and metabolic coupling of cells. The number of gap junctions and their permeability determine the speed by which action potentials are conducted from cell to cell.1 In scintigraphy, radioactive material is injected into the uterus, which is imaged with a gammacamera. Static scintigrams obtained at subsequent time intervals have been used to study uterine clearance of radiocolloid in mares.5 Dynamic scintigrams (one or two images per second) allow the visualization of uterine contractions, their direction, and the movement of intraluminal fluid.6 The presence
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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of the radioactive material itself in the uterus may provoke contractions, and the infusion of the material involves cervical manipulation, which is known to cause contractions.7 None of the above-mentioned methods is very practical, because they are expensive, require special techniques, e.g., scintigraphy, or need general anesthesia, e.g., implantation of EMG electrodes. Transrectal B-mode (brightness mode) ultrasonography is the most common method of examining mares. Using this technique, the number of uterine contractions can be counted or the activity can be scored. However, it is sometimes difficult to define a contraction, and the movement of intestines and the mare itself can be disturbing.8 M-mode (motion mode) ultrasonography produces a graph of motion allowing the recording of amplitude, duration, and frequency of contractions.9 Rectal examination itself provokes contractions10 and may cause erroneous results in transrectal ultrasonography. It is obvious that measuring myometrial contractility in the mare is problematic, which can explain some of the discrepancies between studies. Respiration, position changes, and vocalization of the mare, intestinal motility, as well as environmental stimuli interfere with the measurements.2, 10 It is also possible that the methods measure contractility of the two muscle layers differently.11 On the other hand, the two layers may display contractility that differs in timing, mode, and intensity.3
Control of contractility The myometrial muscle cell is spontaneously active. Without any nervous or hormonal stimulation a piece of isolated uterus will contract regularly and spontaneously.3 In vivo myometrial contractility is controlled by neuronal and hormonal systems. Hormonal modulation Steroid hormones are key players in myometrial activity. High progesterone levels dampen contractions, whereas in the absence of progesterone an increase in estrogens stimulates contractions. Progesterone suppresses the formation of gap junctions, diminishing the coupling between muscle cells, and decreases the number of oxytocin receptors (OT-R). Estrogen, on the other hand, induces gap junctions, depolarizes the muscle plasma membrane, and increases the expression of OT-Rs and prostaglandin (PG) F2α production.1 Oxytocin is a powerful stimulator of contractions, increasing their force, frequency, and duration. The physiological effects of oxytocin depend more on the number of its receptors than on its concentration.
Oxytocin binding to its receptor produces rapid influx of Ca2+ , but also inhibition of Ca extrusion.3 Oxytocin causes release of arachidonic acid and thereby formation of PGF2α . This enhances contractions by causing membrane depolarization and by increasing the number of gap junctions.1 Bearing in mind the dominating effect of steroid hormones on contractility, it is to be expected that the phases of the estrous cycle display different contractile patterns. Estrus is characterized by synchronized well-defined high-intensity EMG activity, whereas more diffuse low-amplitude activity separated by periods of inactivity is typical for diestrus.4 At the onset of luteolytic activity (days 13–14), contractility is increased probably by PGF2α release.12 Neuronal modulation The uterus has autonomic innervation. The parasympathetic fibers combine with the inferior hypogastric nerve and constitute part of the pelvic plexus. Postganglionic fibers enter the uterine wall along with blood vessels and innervate both the endometrium and the myometrium.3 Adrenergic, cholinergic, and other neurotransmitter systems and all four types of andrenergic receptors (α1 , α2 , β1 , and β2 ) are present on the uterus. α-activation causes contraction, and β-activation causes relaxation. The proportion of the receptors varies with hormonal state and muscle layer. Progesterone produces the formation of β-adrenoreceptors, consistent with its role in decreasing uterine activity. At term, there is a desensitization, leading to a loss of myometrial responsiveness to β-agonists.3
Roles of uterine contractions Since this chapter concentrates on the non-pregnant mare, the important tasks of myometrial contractility during parturition and postpartum uterine involution are not discussed. In the non-pregnant mare uterine contractions are a prerequisite for successful sperm transport and effective elimination of sperm, bacteria, and PMNs after breeding. A properly functioning myometrium is therefore vital for the reproductive success of a mare. Sperm transport Spermatozoal motility does not play a significant role in their uterine transport. The journey is so long and has to be travelled so quickly that the sperm would not manage without the aid of uterine contractions. In a dynamic scintigraphic study radiolabeled sperm were imaged with a gamma-camera after artificial insemination (AI). Strong and frequent uterine contractions pushed semen continuously back and
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forth between the body and tips of the horns for approximately 30 minutes after AI. Probably, this serves to spread sperm effectively in the uterus and at the same time to eliminate sperm.6 Whether spermatozoa themselves or fluid in the uterus induce contractions or whether the contractions are due to the vaginal contact by the stallion’s penis or the hand of the inseminator or by the cervical manipulation during AI is unclear. Insertion of a vaginal speculum,10 AI,7, 10 mechanical stimulation of the vagina and cervix,13 mating,14 intrauterine infusion of fluid13 or diluted semen,8 and intrauterine bacterial inoculation15 induce uterine contractions in mares. Ecbolic hormones in semen could be one explanation for uterine contractions after mating. Stallion seminal plasma contains high amounts16 of PGF2α and oxytocin.17 However, the highest oxytocin concentrations are found in the gel,17 which is removed for AI. Besides, in the study of Katila et al.,6 spermatozoa had been washed three times, so all seminal plasma had been removed. Inflammatory influence has also been suggested,10 but inflammation is not present at the time of AI. The most likely explanation is activation of the autonomic nervous system by contact stimuli interacting with adrenergic receptors concurrently with the oxytocin release.7 Uterine evacuation after mating Spermatozoa that failed to enter the oviducts, PMNs that have phagocytized, other inflammatory byproducts, and bacteria have to be removed quickly after breeding. The elimination takes place in two ways, via lymphatics and through the cervix. For the
latter, an open cervix is needed,5, 18 but for both routes uterine contractions are a necessity.19 The elimination of sperm is rapid: 2.5 hours after AI, the majority of spermatozoa had left the uterus, and by 4.5 hours most spermatozoa had been eliminated6, 20 (Figure 269.1). Peak PMN numbers are seen 6–12 hours after AI, then they start to decrease in numbers and, by 48 hours, hardly any or very few PMNs are found in the uterus.20
Clinical signs of abnormal contractility Abnormal contractility means deficient or weak contractions; abnormally strong contractions have not been reported in non-pregnant mares. Lowered pregnancy rates can be considered to reflect deficient contractility, but there are also many other reasons for low pregnancy rates. Clinicians need earlier signs to be able to help mares with the problem of deficient contractility. Whereas normal mares, resistant to endometritis, are capable of clearing the physiological postbreeding endometritis in 48 hours, the so-called susceptible mares are not able to do this. The term ‘susceptible to endometritis’ has been replaced by the term ‘delayed uterine clearance’ (DUC), indicating the inability of these mares to eliminate uterine contents rather than susceptibility to infections.21 DUC is diagnosed by the delayed presence of materials infused into the uterus, e.g., bacteria,22 radiocolloid,23 microspheres, or charcoal,24 none of which can be applied in practice. After comparing different methods used to differentiate between normal mares and
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those with DUC, Brinsko et al.25 suggested a practical solution: ultrasonically detectable fluid > 2 cm in height any time after breeding indicates a susceptible mare with DUC. The area (height × width) may give a more accurate estimate of the amount of IUFA than height.26 Fluid in the mare’s uterus tends to pool in the uterine body and in horn body junctions, which are the most ventral parts.27
Etiology of delayed uterine clearance A mare displaying DUC may have one or more underlying predisposing factors leading to the problem. There may be anatomical (uterus and cervix) and/or functional reasons. High age and repeated pregnancies may produce pathological changes in the endometrium, blood vessels, nerves, and muscle cells, which can lead to abnormal hormone levels, e.g., deficient release of PGF2α .28 Uterine conformation
to the inability to clear uterine contents. Gravity increases fluid pooling and evacuation of fluid becomes difficult. Ventral dilations of uterine horns described by Knudsen30 can represent the same kind of phenomenon. The uterine wall and endometrial folds are thinned in these dilations where fluid accumulates, because the fluid is difficult to remove. Only powerful contractions would be able to push the fluid dorsally and caudally.29
Cervix The removal of uterine contents through the cervix and vagina is possible only if the cervix is sufficiently open. During progesterone dominance the cervix is normally closed,18 and, therefore, the clearance of uterine contents has to take place before progesterone starts to rise after ovulation. This does not mean higher risk for post-ovulatory inseminations, because they are usually carried out 6–12 hours after ovulation, when the progesterone is first starting to increase. Post-ovulatory inseminations with frozen semen did not result in more IUFA than pre-ovulatory inseminations.31 A too tight cervix during estrus is usually a problem of nulliparous mares.32 In a scintigraphic study, LeBlanc et al.5 compared the clearance of intrauterine radiocolloid in three groups of mares: (i) resistant nulliparous, (ii) resistant multiparous, and (iii) susceptible multiparous. The resistant mares, except one, cleared > 50% of the radiocolloid within 2 hours, whereas most of the susceptible mares did not. In the normal mare which retained radiocolloid, a nondilated cervix was discovered during the intrauterine infusion. During diestrus uterine clearance was negligible also in normal mares. When the cervix of a mare was dilated one finger or more, backflow of infused fluids occurred.27 Pycock (reviewed by Allen33 )
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LeBlanc et al.29 infused radiocolloid into the uterus of normal mares and mares with DUC to study the influence of uterine position. The presence of radiocolloid in the uterus, cervix, and vagina provided a good image for evaluation of the anatomical relationships. The angle between the caudoventral aspect of the uterus and cervix relative to horizontal was calculated (Figure 269.2). The angle was more ventral in mares with DUC and more horizontal in normal mares. The mean angle was steeper in normal pluriparous mares than in nulliparous mares. Repeated pregnancies weaken the structural support of the caudal reproductive tract and stretch broad ligaments. It was concluded that a uterus tilting ventrally in relation to the pelvic brim may contribute
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Figure 269.3 Exposed uterine lumen and endometrium 48 hours after India ink infusion. (a) A 48-hour sample from a resistant mare exhibits ink-stained mucosa but does not contain any ink residue in the lumen. Infused ink was transported through lymphatic vessels to lymph nodes (arrow). (b) A 48-hour sample from a susceptible mare contained 150 mL of tar-like exudate containing ink residue, macrophages, neutrophils, and lymphocytes. L, lumen; H, uterine horn; C. cervix. From LeBlanc et al., 1995, with permission from the Society for the Study of Reproduction.
recognized the importance of a patent cervix for uterine clearance. Lymph vessels and veins During diestrus the only means of uterine clearance is via the lymphatics. LeBlanc et al.19 studied lymphatic clearance in normal and susceptible mares during diestrus. They injected India ink into the uterine wall or into the lumen and euthanized the mares 24, 48, or 72 hours later. All mares developed an acute endometritis, which was more severe and prolonged in susceptible mares. Normal mares cleared the ink, which was visible in the aortic and medial iliac lymph nodes. Susceptible mares did not absorb much of the ink (Figure 269.3). It was concluded that during diestrus the lymphatic vessels and lymph nodes drain the uterine submucosa and uterine lumen of excessive contents. The efficacy and rate of lymphatic drainage were decreased in susceptible mares compared with normal mares, suggesting that susceptible mares may have a dysfunction in the lymphatic clearance mechanisms. Contributing factors are a large pendulous uterus and elevated lymphatic hydrostatic pressure from impedance of flow. Myometrial contractions facilitate drainage by compressing lymph vessels, which move material towards lymph nodes. Thus, deficient myometrial contractility may also be an explanation. Decreased lymphatic drainage is impossible to diagnose under practice conditions. LeBlanc et al.19 suggested an association between decreased lymphatic drainage and the presence/size of lymphatic lacunae in uterine biopsies. Later studies have failed to find a correla-
tion between IUFA or edema and area occupied by lymphatic lacunae in biopsies.34, 35 In addition to lymph vessels, veins also drain fluid, e.g., edema fluid. In guinea pigs and humans, the internal elastic membrane of uterine arteries is disrupted during pregnancy and re-established postpartum. Repeated pregnancies predispose uterine arteries to precocious aging, evidenced by increased amounts of elastin in the blood vessels. The postpartum resorption of elastic tissue diminishes with age and parity and formation of new tissue is retarded.36 Pathological changes of uterine vessels have also been described in mares. Iodine infusion into the uterus caused arteriosclerosis and obliteration of the lumen of some vessels.37 Three aged multiparous mares suffering from pregnancy loss showed elastosis of the arterial walls.38 Hyperplasia of intimal elastic fibers and thinning of the medial smooth muscular layer was associated with aging of mares.39 The incidence and severity of angiosis has been shown to increase with the number of previous pregnancies and with advancing age (Figure 269.4).40, 41 Lesions in endometrial arteries are associated with myometrial arterioses.42 Angiopathies may disturb uterine drainage by a reduced function of veins and lymphatic vessels.41 Deficient contractility of the myometrium Susceptible mares have been shown to exhibit decreased electrical activity of the myometrium 10–20 hours after bacterial inoculation compared with resistant mares.15 Since nitric oxide (NO) is a mediator of uterine relaxation, its amounts in uterine
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Figure 269.4 (a) Schematic drawing of arteriolar changes in relation to the number of foals delivered. (A) After the first partus. The membrana elastic interna is enlarged and the peripheral media and the adventitia are thickened by deposition of collagen fibers arranged in bundles and a few elastic fibers occur. (B) After the third partus. The membrana elastic interna is irregularly arranged and rich in mature elastic fibers. The media appears concentric but the myocytes have irregular contours and are intermingled with elastic fibers. (C) After 15 foalings. Within the whole vessel wall an accumulation of irregularly arranged, partially degenerated, and mineralized fibers are visible. The myocytes are degenerated, atrophic, or crippled with numerous intracytoplasmatic matrix vesicles. Red, myocytes; yellow–orange, endothelia, collagen fibers; green, membrana elastic interna, elastic fibers; light blue, mucopolysaccharides; pink, mineralization. (b) Schematic drawing of the findings in an arteriole of a young mare before breeding and after her first foaling. (A) Before the first breeding. The vessel wall shows a regular concentric composition of all histological structures: a uniform internal elastic membrane and only a few elastic fibers within the adventitia. (B) Three days after the first partus. The media and intima are thickened, owing to hyperplastic and sometimes degenerated myocytes and endothelia. The subendothelial clefts are widened and show deposits of mucopolysaccharides. The membrane elastica interna is spread with fine flakes of elastic fibers. (C) Four weeks after the first partus. The findings in the intima and media have reduced, but newly constructed elastic fibers are obvious in the periphery of the media and adventitia. Red, myocytes; yellow–orange, endothelia, collagen fibers; green, membrana elastic interna, elastic fibers; light blue, mucopolysaccharides. From Schoon et al., 1999, with permission from Pferdeheilkunde. (See color plate 124).
secretions and the expression of nitric oxide synthase (NOS) in uterine biopsies were studied in susceptible and resistant mares 13 hours after AI.43 Susceptible mares had more NO in uterine secretions and greater inducible NOS expression in the biopsies than resistant mares. Further studies are needed to ascertain the role of NO in uterine contractility of DUC mares. Intrauterine bacterial inoculations were the most common experimental method to compare susceptible and resistant mares in the 1970s and 1980s. Later studies have shown that uterine contractions do not play a major role in the elimination of bacteria, since cellular, immunological, and other antibacterial defense mechanisms are involved in the bacterial elimination. When clenbuterol (a β-mimetic tocolytic agent) was administered to normal mares, the mares cleared the bacteria, but accumulated fluid for the first time.26 In another study, clenbuterol-treated nor-
mal mares cleared charcoal particles (4–10 µm) but accumulated fluid.44 Oxytocin enhances uterine clearance in normal and susceptible mares,23 suggesting that the underlying defect in susceptible mares is the weak contractility, which can be improved by oxytocin treatment. DUC mares responded to doses of oxytocin of 2.5, 5, and 10 IU similarly to normal mares.45 In a previous study by Cadario et al.,46 oxytocin stimulated a rapid clearance of radiocolloid, but when phenylbutazone, a prostaglandin inhibitor, was administered, hardly any clearance took place. PGF2α and its analogues cloprostenol and fenprostalene induced a rapid clearance of radiocolloid in DUC mares.21 All these studies show that uterine contractions are needed in uterine clearance and suggest that mares with DUC have deficient uterine contractility, which can be treated with ecbolic hormones.
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In an in vitro study, in the absence of neuronal and hormonal control, strips of the longitudinal and circular muscle layers of the myometrium were tested for contractile response to KCl, oxytocin, and PGF2α .47 The myometrial muscle strips had been obtained 18 hours after AI from (i) young nulliparous mares, (ii) old reproductively normal mares, and (iii) old mares with DUC. For all three substances tested, the myometrium of DUC mares generated less tension than that of old normal mares; this was not related to intracellular Ca2+ . Longitudinal muscle from young normal mares and older normal mares responded similarly to KCl and oxytocin. Circular muscle behaved somewhat differently. Young mares developed less tension than older normal mares in response to oxytocin or PGF2α , suggesting possible age-related changes in signaling downstream of intracellular Ca2+ release (Figure 269.5). It was concluded that DUC mares have an intrinsic contractile defect of the myometrium that is not dependent on age or parity. This defect is not receptor dependent and does not result from altered Ca2+ regulation.47 What causes this cellular defect leading to contractile dysfunction? One explanation could be stiffening of the extracellular matrix because of increased collagen deposition. Increase in myometrial collagen has not been reported in mares, but endometrial fibrosis is common in aged mares. Periglandular fibrosis has been shown to be associated with IUFA.34, 48
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Abnormalities in signaling In a normal uterine propagation pattern, contraction starts in the uterine horn and then spreads to the uterine body. In contrast to normal mares, DUC mares showed less normal propagation and more inverted (first body, then horn) or simultaneous propagation. This suggests that the pacemaker cells may have different locations in mares with DUC compared with those in normal mares, whose pacemaker region seems to be in the uterine horns.49 If this is the case, uterine contractions would not be very effective in emptying the uterus. The location of pacemakers in the uterine body may be intrinsic or may develop as a consequence of repeated pregnancies, which cause stretching and hypertrophy of myometrium. Muscle cells may become hyperexcitable and ectopic pacemaker cells.49 In contrast, pacemaker cells shift normally from one place to another. In the scintigraphic study after AI, directions of contractions alternated.6 When xylazine (an α2 -agonist) was administered before oxytocin injection, the duration of increased intrauterine pressure was longer in DUC mares than in normal mares. Similarly, when acepromazine (an
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Figure 269.5 High concentrations of KCl (a) stimulated a greater increase in active tension by circular muscle from both older, reproductively normal mares (ON) and young, nulliparous mares (YN) compared with the response by circular muscle from older mares susceptible to delayed uterine clearance (OS). The KCl-stimulated increase in active tension was similar for YN and ON mares. High concentrations of oxytocin (b) and prostaglandin F2α (PGF2α ) (c) stimulated a greater increase in active tension by circular muscle from either OS or YN mares. Both the oxytocin and PGF2α -stimulated responses of circular muscle from OS and YN mares were similar. ∗ Significant difference (P < 0.05) between the least-squares mean for tension generated by muscle from ON mares and the least-squares mean for OS mares. † Significant difference (P < 0.05) between the least-squares mean for tension generated by muscle from YN mares and the least-squares mean for OS mares. ‡ Significant difference (P < 0.05) between the leastsquares mean for tension generated by muscle for ON mares and the least-squares mean for YN mares.47 From Rigby et al., 2001, with permission from the Society for the Study of Reproduction.
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α2 -antagonist) was injected before oxytocin, no ef-
fect was detected in normal mares; however, in mares with DUC the number of contractions decreased.50 The authors suggested that this enhanced response to α2 -agonists and antagonists may be due to denervation supersensitivity. In late gestation the myometrium becomes chronically stretched. Myometrial cells increase in numbers and size in response to stretching. These changes are associated with degeneration and reduction in the number of adrenergic nerve endings. This results in functional denervation and supersensitivity of the myometrium to catecholamines. Also in mares, the chronic stretching and repeated pregnancies may lead to irreparable structural and functional changes to nerves.50 Surprisingly, detomidine, another α2 -agonist but with more specific binding to α2 -receptors than xylazine, augmented the uterotonic effect of oxytocin in normal mares but not in DUC mares. This may be due to decreased numbers of α2 -receptors, unresponsiveness of the receptors, or a defect in myometrial signaling.49 Why do xylazine and detomidine potentiate the effect of oxytocin? One hypothesis is that they stimulate neurons which promote myometrial responsiveness to oxytocin. According to another hypothesis, they activate signaling pathways that potentiate oxytocin-induced signal transduction.49 Although the works of DeLille et al.50 and von Reitzenstein et al.49 conflict with each other, they tend to support the findings of Rigby et al.47 about an intrinsic defect in mares with DUC. Prostaglandin release When oxytocin binds to its receptor, it triggers prostaglandin secretion and release. In normal mares, exogenous oxytocin resulted in prostaglandin release, but not in mares with DUC. AI caused a similar endogenous oxytocin release in normal and DUC mares, but this resulted in subsequent prostaglandin release only in normal mares. The authors thought this indicated a possible defect in prostaglandin release at the OT-R or post-receptor level.28 Totally different results were reported by Cadario et al.51 Exogenous oxytocin increased prostaglandin in DUC mares but not in normal mares. Prostaglandin levels did not affect the clearance of radiocolloid. The presence of intrauterine fluid did not have any effect. Similar results were obtained in an experiment where phenylbutazone (a prostaglandin inhibitor) or oxytocin or oxytocin and phenylbutazone together were given to normal mares. Oxytocin was able to override the inhibitory effect of phenylbutazone on the clearance of radiocol-
loid, although prostaglandin release was inhibited. The authors concluded that the effect of oxytocin may be independent of uterine prostaglandin secretion.46 One more study has been published about the relationship of oxytocin and prostaglandin in mares. Veronesi et al.52 compared DUC mares and normal mares defined by the presence or absence of IUFA. The administration of 20 IU of oxytocin intramuscularly increased PGF-metabolite levels in 71% of DUC mares compared with 38% in mares with no IUFA. Also, progesterone and estradiol-β levels were measured, but they provided no explanation. It is difficult to understand these conflicting results. One can only speculate that the grounds for placing mares in the normal versus DUC category may have been different, days to ovulation may have been different (numbers of OT-Rs vary), or the inflamed uterus may have had an effect. A possible defect in prostaglandin release in DUC mares cannot be excluded, but further investigations are needed.
Summary The detection of intrauterine fluid after mating in transrectal ultrasonography is a practical and reliable method to diagnose delayed uterine clearance. Underlying reasons for fluid accumulation are uterus tilting ventrally in relation to the pelvic brim, ventral dilations of the uterine horns, and a non-patent cervix. Angiopathies of endometrial and myometrial vessels reduce blood flow and venous and lymphatic drainage. The most important contributing factor is, however, abnormal uterine contractility. Myometrial muscle cells can have intrinsic contractile defects, which may be related to endometrial fibrosis and stiffening of the extracellular matrix. Abnormal neuronal signaling because of chronic stretching of nerves during repeated pregnancies has been suggested as one explanation for decreased myometrial contractions. Finally, deficient prostaglandin release in response to mating and oxytocin release has been demonstrated in problem mares.
References 1. Jain TL, Saade GR, Saade GR, Garfield RE. Uterine contraction. In: Knobil E, Neill JD (eds) Encyclopedia of Reproduction, vol. 4. San Diego: Academic Press, 1999; pp. 932–42. 2. Goddard PJ, Allen WE, Lawrence Gerring E. Genital tract pressures in mares. I. Normal pressures and the effect of physiological events. Theriogenology 1985;23:815–27. 3. Wray S. Uterine contraction and physiological mechanisms of modulation. Am J Physiol 1993;264:C1–18. ¨ AOG, Liu IKM, Ing M, Pascoe 4. Troedsson MHT, Wistrom JR, Thurmond M. Registration of myometrial activity using multiple site electromyography in cyclic mares. J Reprod Fertil 1993;99:299–306.
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Abnormalities of Contractility 5. LeBlanc MM, Neuwirth L, Asbury C, Tran T, Mauragis D, Klapstein E. Scintigraphic measurement of uterine clearance in normal mares and mares with recurrent endometritis. Equine Vet J 1994;26:109–13. 6. Katila T, Sankari S, M¨akel¨a O. Transport of spermatozoa in the reproductive tracts of mares. J Reprod Fertil Suppl 2000;56:571–8. 7. Madill S, Troedsson MHT, Alexander SL, Shand N, Santschi EM, Irvine CHG. Simultaneous recording of pituitary oxytocin secretion and myometrial activity in oestrous mares exposed to various breeding stimuli. J Reprod Fertil Suppl 2000;56:351–61. 8. Sinnemaa L, J¨arvimaa T, Lehmonen N, M¨akel¨a O, Reilas T, Sankari S, Katila T. Effect of insemination volume on uterine contractions and inflammatory response and on elimination of semen in the mare uterus: scintigraphic and ultrasonographic studies. J Vet Med Assoc 2005;52:466–71. 9. Campbell MLH, England GCW. M-mode ultrasound imaging of the contractions of the equine uterus. Vet Rec 2002;150:575–7. 10. Taverne MAM, van der Weyden GC, Fontijne P, Dieleman SJ, Pashen RL, Allen WR. In-vivo myometrial electrical activity in the cyclic mare. J Reprod Fertil 1979;56:521–32. 11. Troedsson MHT, Liu IKM, Ing M, Pascoe JR. Smooth muscle electrical activity in the oviduct, and the effect of oxytocin, prostaglandin F2α , and prostaglandin E2 on the myometrium and the oviduct of the cyclic mare. Biol Reprod Monogr 1995;1:475–88. 12. Griffin PG, Ginther OJ. Effects of day of estrous cycle, time of day, luteolysis, and embryo on uterine contractility in mares. Theriogenology 1993;39:997–1008. 13. Campbell MLH, England GCW. Effect of teasing, mechanical stimulation and the intrauterine infusion of saline on uterine contractions in mares. Vet Rec 2004;155:103–10. 14. Jones DM, Fielden ED, Carr DH. Some physiological and pharmacological factors affecting uterine motility as measured by electromyography in the mare. J Reprod Fertil Suppl 1991;44:357–68. 15. Troedsson MHT, Liu IKM, Ing M, Pascoe JR, Thurmond M. Multiple site electromyography recordings of uterine activity following an intrauterine bacterial challenge in mares susceptible and resistant to chronic uterine infection. J Reprod Fertil 1993;99:307–13. 16. Claus R, Dimmick MA. Estrogens and prostaglandin F2α in the semen and blood plasma of stallions. Theriogenology 1992;38:687–93. 17. Watson ED, Nikolakopoulos E, Gilbert C, Goode J. Oxytocin in the semen and gonads of the stallion. Theriogenology 1999;51:855–65. 18. Evans MJ, Hamer JM, Gason LM, Graham CS, Asbury AC, Irvine CHG. Clearance of bacteria and non-antigenic markers following intra-uterine inoculation into maiden mares: effect of steroid hormone environment. Theriogenology 1986;26:37–50. 19. LeBlanc MM, Johnson RD, Calderwood Mays MB, Valderrama C. Lymphatic clearance of India ink in reproductively normal mares and mares susceptible to endometritis. Biol Reprod Monogr 1995;1:501–6. 20. Katila T. Onset and duration of uterine inflammatory response of mares after insemination with fresh semen. Biol Reprod Monogr 1995;1:515–17. 21. Coombs GB, LeBlanc MM, Neuwirth L, Tran TQ. Effects of prostaglandin F2α , cloprostenol and fenprostalene on uterine clearance of radiocolloid in the mare. Theriogenology 1996;45:1449–55.
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22. Hughes JP, Loy RG. Investigations on the effect of intrauterine inoculations of Streptococcus zooepidemicus in the mare. In: Proceedings of the 15th Annual Convention of the American Association of Equine Practitioners, 1969, pp. 289–92. 23. LeBlanc MM, Neuwirth L, Mauragis D, Klapstein E, Tran T. Oxytocin enhances clearance of radiocolloid from the uterine lumen of reproductively normal and fertile mares. Equine Vet J 1994;26:279–82. 24. LeBlanc MM, Asbury AC, Lyle SK. Uterine clearance mechanisms during the early postovulatory period in mares. Am J Vet Res 1989;50:864–7. 25. Brinsko S, Rigby SL, Varner DD, Blanchard TL. A practical method for recognizing mares susceptible to post-breeding endometritis. In: Proceedings of the 49th Annual Convention of the American Association of Equine Practitioners, 2003. 26. Nikolakopoulos E, Watson ED. Uterine contractility is necessary for the clearance of intrauterine fluid but not bacteria after bacterial infusion in the mare. Theriogenology 1999;52:413–23. 27. Jones D. Fluid distribution and cervical loss following intrauterine infusion in the mare. Equine Pract 1995;17:12–19. 28. Nikolakopoulos E, Kindahl H, Watson ED. Oxytocin and PGF2α release in mares resistant and susceptible to persistent mating-induced endometritis. J Reprod Fertil Suppl 2000;56:363–72. 29. LeBlanc MM, Neuwirth L, Jones L, Cage C, Mauragis D. Differences in uterine position of reproductively normal mares and those with delayed uterine clearance detected by scintigraphy. Theriogenology 1998;50:49–54. 30. Knudsen O. Partial dilatation of the uterus as a cause of sterility in the mare. Cornell Vet 1964;54:423–38. 31. Watson ED, Barbacini S, Berrocal B, Sheerin O, Marchi V, Zavaglia G, Necchi D. Effect of insemination time of frozen semen on incidence of uterine fluid in mares. Theriogenology 2001;56:123–31. 32. Malschitzky E, Trein CR, Cunha Bustamante Filho I, Garbade P, Macedo Gregory R, Costa Mattos R. Young maiden mares can also be susceptible to a persistent matinginduced endometritis. Pferdeheilkunde 2006;22:201–6. 33. Allen WR. Proceedings of the John P. Hughes International Workshop of Equine Endometritis. Equine Vet J 1993;25:184–93. 34. Reilas T, Katila T, M¨akel¨a O, Huhtinen M, Koskinen E. Intrauterine fluid accumulation in oestrous mares. Acta Vet Scand 1997;38:69–78. 35. Risco AM, Reilas T, Muilu L, Kareskoski M, Katila T. Effect of oxytocin and flunixin meglumine on uterine response to insemination in mares. Theriogenology 2009;72:1195– 1201. 36. Albert EN, Bhussry BR. The effects of multiple pregnancies and age on the elastic tissue of uterine arteries in the guinea pig. Am J Anat 1967;121:259–70. 37. van Dyk E, Lange AL. The detrimental effect of iodine as an intra-uterine instillation in mares. J S Afr Vet Assoc 1986;57:205–10. 38. Oikawa M, Katayama Y, Yoshihara T, Kaneko M, Yoshikawa T. Microscopical characteristics of uterine wall arteries in barren aged mares. J Comp Pathol 1993;108:411– 15. 39. Nambo Y, Oikawa M, Yoshihara T, Kuwano A, Katayama Y. Age-related morphometrical changes of arteries of uterine wall in mares. Zentralbl Veterinarmed A 1995;42: 383–7.
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40. Gruninger E, Schoon HA, Menger S, Klug E. Incidence and morphology of endometrial angiopathies in mares in relationship to age and parity. J Comp Pathol 1998;119:293–309. 41. Schoon D, Schoon H-A, Klug E. Angioses in the equine endometrium: pathogenesis and clinical correlations. Pferdeheilkunde 1999;15:541–6. 42. Ludwig S, Schoon D, Aupperle H, von Reiswitz A, Schoon H-A. Angiopathies in the equine endometrial biopsy – a marker for extrauterine vascular lesions? Pferdeheilkunde 2001;17:608–14. 43. Alghamdi AS, Foster DN, Carlson CS, Troedsson MH. Nitric oxide levels and nitric oxide synthase expression in uterine samples from mares susceptible and resistant to persistent breeding-induced endometritis. Am J Reprod Immunol 2005;53:230–7. 44. Kolm G, Gemeiner M, Deichsel K, Budik S, Aurich J, Aurich C. Effect of clenbuterol on the clearance of particles of charcoal (4 to 90 µm) from the uteri of mares. Vet Rec 2005;156:279–81. 45. Cadario ME, Merritt AM, Archbald LF, Thatcher WW, LeBlanc MM. Changes in intrauterine pressure after oxytocin administration in reproductively normal mares and in those with a delay in uterine clearance. Theriogenology 1999;51:1017–25. 46. Cadario ME, Thatcher M-JD, LeBlanc MM. Relationship between prostaglandin and uterine clearance of radiocolloid in the mare. Biol Reprod Monogr 1995;1:495–500.
47. Rigby SL, Barhoumi R, Burghardt RC, Colleran P, Thompson JA, Varner DD, Blanchard TL, Brinsko SP, Taylor T, Wilkerson MK, Delp MD. Mares with delayed uterine clearance have an intrinsic defect in myometrial function. Biol Reprod 2001;65:740–7. ¨ 48. Ozgen S, Schoon H-A, Aupperle H, Sieme H, Klug E. Etiopathogenesis of equine intrauterine fluid accumulation. Pferdeheilkunde 2002;18:594–9. 49. von Reitzenstein M, Callahan MA, Hansen PJ, LeBlanc MM. Aberrations in uterine contractile patterns in mares with delayed uterine clearance after administration of detomidine and oxytocin. Theriogenology 2002;58:887–98. 50. DeLille AJAE, Silvers ML, Cadario ME, Tran TQ, Cage CL, LeBlanc MM. Interactions of xylazine and acepromazine with oxytocin and the effects of these interactions on intrauterine pressure in normal mares and mares with delayed uterine clearance. J Reprod Fertil Suppl 2000;56: 373–9. 51. Cadario ME, Thatcher WW, Klapstein E, Merritt AM, Archbald LF, Thatcher MJ, LeBlanc MM. Dynamics of prostaglandin secretion, intrauterine fluid and uterine clearance in reproductively normal mares and mares with delayed uterine clearance. Theriogenology 1999;52:1181–92. 52. Veronesi MC, Carluccio A, Kindahl H, Faustini M, Battochio M, Cairoli F. Oxytocin-induced PGF2α release in mares with and without post-breeding delayed uterine clearance. J Vet Med Assoc 2006;53:259–62.
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Chapter 270
Abnormalities of Development John P. Hurtgen
Congenital abnormalities of the uterus are very rare in the horse. If these congenital abnormalities occur in the mare, infertility or even sterility usually results. With some of these congenital uterine abnormalities, mares have repeated estrous cycles before a thorough reproductive examination identifies the abnormality. Mares with gonadal dysgenesis, such as 63XO, 64XY, and others, usually have small inactive ovaries with a resulting infantile, atonic uterus. Chromosomal analysis of blood or tissue samples should identify mares with this type of congenital uterine abnormality.1 If mares affected with gonadal dysgenesis are treated with progesterone and estrogen, the uterus may develop normal size and tone.2 Congenital abnormalities of the paramesonephric ducts can occur in any breed. However, draft horse breeds such as Clydesdale and Shire appear to be most commonly affected. Segmental aplasia involving the uterine body has been reported in a pony mare.3 The author has examined two unrelated Shire mares with uterine segmental aplasia. One mare was missing the entire right uterine horn. The other Shire mare had aplasia of a short segment of the right horn. The ovarian end of this horn was fluid-filled. The mare had normal estrous cycles and had been bred repeatedly without success. Careful palpation and ultrasonography of the uterus per rectum should identify affected mares even though localized uterine fluid accumulation may, initially, suggest that the mare is pregnant. An occasional mare has been reported to have a double cervix connected to separate uterine horns.4, 5 In many cases the uterine horns do not communicate. These mares are very infertile due to the inability of the embryo to migrate throughout the uterus so that maternal recognition of pregnancy can occur by inhibiting systemic prostaglandin release.6 However, if
mares with this condition receive exogenous progesterone following breeding, pregnancy can progress to term.7 Limited uterine space for development of the fetus would result in a very undersized foal at term. This condition may be referred to as uterus didelphic uterus. In an embryo transfer program, embryos can be collected from affected mares and result in pregnancies when the embryos are transferred to normal mares. When affected mares are bred, care must be taken to be certain that semen is deposited in the uterine horn adjacent to the ovarian side of ovulation. A single case of a congenitally absent or short uterine body has been reported in a 7-year-old Quarter Horse mare.8
References 1. Chandley AC, Fletcher J, Rossdale PD. Chromosome abnormalities as a cause of infertility in mares. J Reprod Fertil Suppl 1975;23:377–83. 2. Hinrichs K, Riera FL, Klunder LR. Establishment of pregnancy after embryo transfer in mares with gonadal dysgenesis. J In Vitro Fertil Embryo Transf 1989;6:305–9. 3. Schlotthauer CF, Zollman PE. The occurrence of so-called “white heifer disease” in a white Shetland Pony mare. J Am Vet Med Assoc 1956;129:309–10. 4. Volkmann DH, Gilbert RO. Uterus bicollis in a Clydesdale mare. Equine Vet J 1989;21:71. 5. Blue MG. An uterocervical anomaly (uterus bicorpor bicollis) in a mare, and the manual disruption of early bilateral pregnancies. N Z Vet J 1985;33:17–19. 6. Ginther OJ, Garcia MC, Squires EL, Steffenhagen WP. Anatomy of vasculature of the uterus and ovaries in the mare. Am J Vet Res 1972;33:1561–8. 7. McDowell KJ, Sharp DC, Peck LS, Cheves LL. Effect of restricted conceptus motility on maternal recognition of pregnancy in mares. Equine Vet J Suppl 1985;3:23–4. 8. Meyers PJ, Varner DD. Abortion of a mummified fetus associated with short uterine body in a mare. J Am Vet Med Assoc 1991;198:1768–70.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Chapter 271
Endometritis Mats H.T. Troedsson
A failure of the uterine defense mechanisms to eliminate antigens (bacteria or spermatozoa) and inflammatory products from the uterus results in persistent endometritis, which is a major cause of reduced fertility in broodmares. The uterus is well protected from external contamination by physical barriers consisting of the vulva, the vestibule, the vagina, and the cervix. Any compromise of these barriers predisposes the mare to chronic endometritis.1 Breeding is another source of uterine contamination. Intrauterine deposition of semen causes an inflammatory reaction resulting from bacterial contamination of the ejaculate or from spermatozoa.2 Approximately 15% of a normal population of Thoroughbred broodmares developed persistent endometritis after breeding.3 Persistent endometritis may interfere directly with the survival of an embryo, or cause premature luteolysis and embryonic loss because of increased prostaglandin F2α (PGF2α ) concentrations.4
The role of uterine defense mechanisms in persistent endometritis Natural resistance to experimentally induced bacterial contamination has been demonstrated in young mares, whereas a population of older mares developed a persistent endometritis after experimentally induced bacterial inoculation of the uterus.5, 6 Based on these studies, mares have been classified as either susceptible or resistant to persistent uterine infection.5 Several uterine defense mechanisms are involved in the protection of the uterus against a persistent endometritis. Polymorphonuclear neutrophils (PMNs) are the first inflammatory cells to enter the uterine lumen following an inflammatory stimulus. 7 Chemoattractive properties of uterine fluid have been described in vitro in horses, and the uterus responds quickly to an antigen with release of PMN-
chemotactic mediators, which results in a rapid migration of PMNs into the uterine lumen.8–12 Several classes of immunoglobulins have been isolated from the equine uterus. Although antibodymediated uterine defense may be important for effective elimination of bacterial contaminants from the uterus in susceptible mares, the concentrations of immunoglobulins in uterine secretions are similar or even elevated compared to those of resistant mares.13–17 Lactoferrin and other immunomodulators have also been found at higher concentrations in susceptible compared to resistant mares.18, 19 These results suggest that susceptible mares have a mostly functional immune response to antigenic stimuli. It is currently believed that a failure of mechanical aspects of the uterine defense system is the major contributor to the development of persistent endometritis. Using intrauterine inoculations of a combination of radioactively labeled microspheres and bacteria, impaired uterine clearance was demonstrated in mares susceptible to persistent endometritis, but not in mares resistant to it.20, 21 Studies using scintigraphic measurements of uterine clearance of radioactive colloids further defined a delayed uterine clearance in susceptible mares.22 Using electromyography to register myometrial activity, it was observed that the impaired uterine clearance in susceptible mares was caused by reduced myometrial activity in response to the inflammation (Figure 271.1).23 However, the mechanism of impaired myoelectrical activity in susceptible mares is still not fully understood. Using an in vitro model to study myometrial activity, it was suggested that mares susceptible to delayed uterine clearance had an intrinsic contractile defect of the myometrium.24 A more recent study found that susceptible mares had an increased uterine accumulation of nitric oxide in the uterine lumen 13 hours after insemination.25 Nitric oxide mediates smooth muscle relaxation, and the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Time (hours)
Figure 271.1 Uterine myoelectrical activity of mares resistant and susceptible to persistent endometritis, in response to an intrauterine inoculation of Streptococcus equi ssp. zooepidemicus. Adapted from Troedsson et al.23
authors offered this effect as a possible explanation for below-baseline myoelectrical activity between 6 and 19 hours post-insemination in susceptible mares (Figure 271.1). The authors also observed an increased number of inducible nitric oxide synthase (iNOS)-positive mast cells as well as an upregulation of iNOS mRNA in the endometrium of susceptible mares.25 The presence of mast cells suggests that histamine-mediated vasodilation may be involved in delayed uterine clearance of susceptible mares. Factors beside impaired uterine contractility that can predispose to delayed uterine clearance include anatomical abnormalities of the reproductive tract. Mares with delayed uterine clearance often suffer from a forward tilt of the uterus over the pelvic brim. This may be a contributing factor in abnormal accumulation of fluid and inflammatory products after breeding or after bacterial contamination of the uterus.26 A failure of the cervix to relax during estrus, or insufficient lymphatic drainage may also contribute to delayed clearance.27 Bacterial endometritis Bacterial endometritis is a significant problem in cycling mares with lowered resistance to persistent inflammation. Bacteria most commonly isolated from the uterus of the mare are beta-hemolytic streptococci (Streptococcus equi ssp. zooepidemicus), Escherichia coli, Pseudomonas aeruginosa, and Klebsiella pneumoniae. Other aerobic bacteria isolated from reproductive tracts of mares include alpha-hemolytic streptococci, Corynebacterium spp., Staphylococcus spp., Enterobacter spp., Actinobacter spp., Proteus spp., and Citrobacter spp. Contagious equine metritis (CEM) is a sexually transmitted disease. It is caused by Taylorella equigenitalis, a highly contagious and pathogenic microorganism. Although the present status of a mare’s uterine defense mechanism is important for the manifestation of CEM, T. equigenitalis is highly resistant and capable of overcoming the mare’s normal
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defense mechanisms. The resulting disease is characterized by endometritis, vaginal discharge, and a devastating effect on fertility. The disease can cause major economic losses to the equine breeding industry. With the exception of an outbreak of the disease in Thoroughbreds in central Kentucky in 1978, the United States has, through rigorous control programs and import regulations, been free from this disease. In 2006, however, a group of three imported Warmblood stallions tested positive for T. equigenitalis following routine culturing. In 2008, a domestic stallion tested positive for the bacterium following testing associated with export of cryopreserved semen to Europe. In contrast to reported outbreaks of the disease in Thoroughbreds that are bred by natural service, almost no clinical signs in exposed mares or effect on fertility were associated with the stallions that tested positive during the last two incidents. While lateral transmission appeared to be common between stallions that shared a breeding shed, less than 1% of exposed mares tested positive for T. equigenitalis. A common factor for the positive stallions was that they had been used for artificial insemination exclusively. Semen that is used for artificial insemination is commonly extended in a semen extender that contains an antibioticum, or a combination of antibiotics. This may explain the differences in clinical outcome in mares that have been bred to infected stallions through natural service versus artificial insemination. In contrast to a true sexually transmitted disease such as CEM, persistent infectious endometritis is often the result of contamination of the uterus by the mare’s fecal flora in combination with compromised uterine defense.28, 29 P. aeruginosa, K. pneumoniae, and possibly S. equi ssp. zooepidemicus and E. coli, can be sexually transmitted in horses, but the consequences of exposure to these microorganisms are determined by the particular strain involved and active participation of all facets of the mare’s uterine defense mechanisms. A bacterial biofilm is a complex aggregation of microorganisms growing on a solid substrate. Biofilms are characterized by structural heterogeneity, genetic diversity, complex community interactions, and an extracellular matrix of polymeric substances, which greatly increase the resistance to antibiotic therapy. The presence of biofilms has been well described in the mucosa of the mouth and in the urinary bladder in humans. Although not documented in the mare, it has been proposed that endometrial biofilms are responsible for cases of chronic endometritis that appear to be non-responsive to conventional treatment.30 The concept is interesting and warrants further research. Breeding may also cause bacterial endometritis, since bacteria often are introduced into the uterus
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during coitus.31 This was previously believed to be the sole source of breeding-induced endometritis, and previous research on the pathophysiology of persistent endometritis has predominantly been performed using bacterial endometritis as a model. One could speculate that an immunosuppressive effect of seminal plasma that protects spermatozoa from being destroyed in the uterus provides contaminating bacteria with an environment that favors adhesion and growth. A recent report suggested that bacterial endometritis contributes to the aggressive post-breeding endometritis in mares bred with frozen/thawed semen, from which most seminal plasma had been removed.32 However, recent observations in our laboratory suggest that a specific protein in seminal plasma protects viable spermatozoa, but not dead spermatozoa or bacteria from PMN-binding and phagocytosis (M.H.T. Troedsson, unpublished observations). Hence, the interaction between bacteria and seminal components in the uterus needs to be studied further.
Diagnostic approach History compatible with endometritis includes infertility after breeding to a fertile stallion. Mares with severe endometritis may have shortened interestrous intervals and may show vaginal discharge. Physical and speculum examination may show anatomical defects of the vulva or cervix. Excessively easy passage of a vaginal speculum, or the characteristic sound of aspirating air when the vulvar lips are gently separated by the operator, may indicate loss of integrity of the vestibulovaginal sphincter. Discharge from the cervix and vagina may be apparent (Figure 271.2). Transrectal palpation and ultrasonography may reveal accumulations of uterine luminal fluid (Figure 271.3).
Figure 271.2 Purulent vaginal discharge in a mare with persistent endometritis.
suggested to be a sensitive method to obtain an endometrial sample for culture.34, 35 It is important to use a closed system to infuse and recover fluid from the uterus in order to avoid environmental contamination when this method is used. Cultures may also be taken from endometrial biopsies. A method to
Microbiology Quantitative aerobic bacterial culture of the uterine lumen will identify potential pathogens and antibiotic sensitivity. Samples can be obtained using a swab, low-volume uterine lavage, or an endometrial biopsy. Samples should be taken during estrus. If a uterine swab is used, the swab should be transported in non-nutritive media to the laboratory. Inadvertent contamination of cultures with bacteria from the lower reproductive tract is common, so the culture instrument should be guarded until it is within the uterus.33 Samples obtained through a vaginal speculum may decrease the risk of contamination from the lower reproductive tract (Figure 271.4). The use of low-volume (50–100 mL) uterine lavage has been
Figure 271.3 Intraluminal accumulation of fluid in the body of the uterus. The depth of the fluid accumulation was measured to be 4.13 cm.
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obtained from swab samples, even under optimal circumstances, and laboratory results should always be interpreted in light of clinical findings. Culture samples for T. equigenitalis should be taken from the endometrium, cervix, clitoral fossa, and sinuses. Samples should be placed in Amies media with charcoal or Steward’s media and kept refrigerated until delivered to the laboratory. Endometrial cytology
Figure 271.4 Endometrial culture through a vaginal speculum. Note the light source located at the top of the speculum.
obtain a biopsy sample for culture has been described.36 Bacterial growth from the clitoral fossa may predispose for repeated contamination of the uterus during breeding or when samples are obtained for bacterial culture. In mares with recurrent infection with a specific strain of bacteria, the clitoral fossa should be cultured for bacterial growth. Culture alone is not diagnostic. A false positive bacterial sample may be obtained as the result of contamination (even when double-guarded swabs are used), and culture results should always be interpreted together with results from endometrial cytology. False negative samples are frequently
Polymorphonuclear neutrophils migrate into the uterine lumen in response to inflammation so endometritis is rapidly and accurately diagnosed by examination of exfoliated endometrial cells. A sample may be taken with a guarded swab, a cytology brush (Figure 271.5), or by low-volume uterine lavage. Air-dried smears are stained with new methylene blue or modified Wright–Giemsa. Epithelial cells may be shed singly or in rafts. Arbitrary definitions of endometritis have been established on the basis of relative numbers of PMNs. More than one PMN per 10 epithelial cells is considered to be consistent with endometritis (Figure 271.6). Riddle and co-workers in 2007 observed that mares with <2 PMNs per ×400 magnification microscopic field had significantly higher pregnancy rates (60% per cycle) compared to mares with 2–5 PMNs per ×400 field (49% per cycle).37 Mares with >5 PMNs per ×400 field had a further reduction in pregnancy per cycle (23%). The authors also found that bacterial infection with E. coli
Figure 271.5 The ‘Steiner cytology and double guarded culture swab system’ (Minitube of America, Verona, WI) for endometrial sampling in mares.
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Mare: Disease of the Uterus Table 271.1 Endometrial biopsy grade and fertility prognosis Biopsy category
Degree of change
Predicted % foaling rate
I IIA IIB III
None Mild Moderate Severe
80–90 50–80 10–50 < 10
Reproduced from Schoon HA and Schoon D.41
Figure 271.6 Polymorphonuclear neutrophils (PMNs) obtained from the endometrium of a mare with active endometritis. (See color plate 125).
was less likely to be associated with a positive cytology when compared to mares infected with Streptococcus zooepidemicus. These findings have been supported by Moller-Nielsen and co-workers.38 Endometrial cytology from normal mares may contain PMNs and spermatozoa for several days after breeding (Figure 271.7). It has been suggested that eosinophils are associated with fungal endometritis and pneumovagina. Urine crystals indicate urovagina. Endometrial biopsy Endometrial biopsy is an accurate diagnostic and prognostic tool for endometritis. The biopsy should
be taken during the breeding season, should be of adequate size, and should be fixed in 10% formalin, or in Bouin’s solution for 24 hours and then transferred to 10% formalin. Chronic endometritis is characterized by infiltration of the endometrium with mononuclear cells. The severity of endometrial changes is inversely related to reproductive performance. A system to classify histological changes has been described and is widely used (Table 271.1).39 More recently, it has been suggested that the degree of degenerative angiopathies (angiosis) in the endometrium negatively affects fertility, and should therefore be considered when classifying an endometrial biopsy for prognosis.40, 41 An endometrial biopsy can also be used for bacterial culture.36 Hysteroscopy Examination of the uterine lumen with an endoscope provides information about the degree of inflammation, in addition to evidence of foreign bodies, transluminal adhesions, intraluminal masses, and endometrial cysts. Focal bacterial plaques can sometimes be detected on hysterendoscopic examination. These mares may have a negative culture and cytology from a routine swab sample. Treatment and prognosis
Figure 271.7 Polymorphonuclear neutrophils (PMNs) obtained from low-volume uterine lavage after breeding. Note the intracellular presence of a sperm head in the PMN. (See color plate 126).
Treatment of mares with persistent uterine infections needs to be directed towards the underlying breakdown of the uterine defense and against the microbial agent. The first therapeutic concern should be to remove predisposing causes, such as a breakdown of external genital barriers. Persistent uterine infection frequently follows degenerative or traumatic anatomical changes and loss of integrity of the barriers of ascending infection. Therefore Caslick’s surgery, repair of cervical damage and perineal lacerations, and correction of urovagina should precede specific endometrial treatment. All potential sources of contamination, including intrauterine passage of diagnostic and treatment implements, should be minimized. In some mares, sexual rest and no further treatment can result in full recovery. Mares that are susceptible to persistent uterine infections should
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Table 271.2 Antibacterial drugs used for intrauterine administration for treatment of uterine infections in mares
Drug
Dosage (intrauterine administration)
Amikacin sulfate Ampicillin Carbenicillin
2g 3g 2–6 g
Gram-negative spectrum; buffer with equal volume of 7.5% bicarbonate Gram-negative spectrum; may irritate the endometrium Gram-negative spectrum; may irritate the endometrium; buffer with equal volume of 7.5% bicarbonate
Gentamicin sulfate Kanamycin sulfate Neomycin sulfate Potassium penicillin G Polymyxin B Ticarcillin Ticarcillin/clavulanic acid Ceftiofur
1–3 g 1–3 g 3–4 g 5 million U 1 million U 6g 3–6 g/200 mg 1g
Gram-negative spectrum; buffer with equal volume of 7.5% bicarbonate Escherichia coli, spermicidal Useful against sensitive E. coli Streptococcus zooepidemicus Pseudomonas Broad spectrum Broad spectrum Broad spectrum (S. zooepidemicus)
be bred using minimal contamination techniques to avoid bacterial contamination of the uterus.31 Antibiotics may be administered by either local or systemic routes. Critical studies of the efficacy of systemic antibiotics are limited, although effective levels are produced in the endometrium after systemic administration.42, 43 Higher tissue concentrations in the endometrium can be expected following intrauterine treatment, compared to systemic treatment. Intraluminal fluid and inflammatory debris should be removed by uterine lavage before local treatment. Drugs and dosages are summarized in Table 271.2. Treatment should be based on sensitivity. Mares should be treated during estrus when natural defense is maximal, and strict aseptic technique should be used. The volume of fluid used for antibiotic therapy is dependent on the size of the uterus. A total volume of 30–60 mL is usually sufficient. Treatment should continue daily for 4 to 6 days during the duration of estrus. Bacterial resistance may follow inadequate dosage, and follow-up cultures should be performed. Repeated contamination may indicate an unsuccessfully resolved predisposing cause. The clitoral fossa should always be considered as a location of bacterial growth in these cases. If culture results from the endometrium and the clitoral fossa match, the clitoral fossa should be cleaned and treated locally with antibiotics. Mares with CEM should be treated with intrauterine infusions of antibiotics based on sensitivity tests, in combination with local treatments of the clitoral fossa and sinuses. Best results can be expected when treatment is initiated when the mare is in estrus, and when combined with uterine lavage if inflammatory debris or intraluminal fluid is present. Cleansing of the vulva and the clitoris daily for 5 days with a 4% chlorhexidine or nitrofurazone ointment has been recommended.44 Sinusectomy can also be
Comments
performed.45 Import regulations in countries free from CEM serve to prevent outbreaks of the disease. The spread of CEM on farms in endemic countries is best prevented by implementation of strict hygiene, screening of breeding stallions before the breeding season, and the use of artificial insemination if allowed by the breed registry. Treatment with immunostimulatory agents (Propionibacterium acnes or Mycobacterium cell-wall extract) has been reported to improve pregnancy rates in mares with persistent endometritis.46 It was suggested that Mycobacterium cell-wall extract modulates endometrial cytokines in susceptible mares, but the mechanism is not fully understood.19 Other treatments that aim to stimulate the local uterine immune system include intrauterine treatment with blood plasma,47 and infusion of leukocytes into the uterus.48 Intrauterine infusions of a variety of solutions have been used to treat infectious endometritis. These include iodine, chlorhexidine, hypertonic saline, dimethyl sulfoxide (DMSO), kerosene, acetylcysteine, and hydrogen peroxide. The clinician should be aware that many of these solutions are strong irritants and can cause damage to the endometrium. The author does not recommend routine use of these solutions in the uterus. If povidone iodine is used for intrauterine treatment, it should be used in a very dilute solution (0.05–0.10%) in lactated Ringer’s solution (LRS). A 30% DMSO solution and acetylcysteine in a 20% solution have been added to uterine lavage fluids in mares with suspected biofilms in the uterus.30 The rationale of this treatment is to clear mucus or biofilms from the uterus of mares with persistent Gram-negative infections. However, the treatment has not yet been critically evaluated in a controlled study.
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Ethylene diamine tetra-acetic acid (EDTA)-Tris has been shown to be effective in decreasing the viability of clinical isolates of Pseudomonas aeruginosa from horses.49 The infusion of 3.5 mM EDTA (0.5 M Tris; pH 8) into the uterine lumen can be considered to be safe, since it does not cause irritation to the endometrium. Addition of 120 mL EDTA-Tris to 1.5 g amikacin for the treatment of endometritis caused by Gram-negative bacteria has been reported to result in pregnancies.30 However, no controlled studies have been reported yet to support this observation.
Breeding-induced endometritis Transport of spermatozoa from the equine uterus to the oviduct is completed within 4 hours after breeding,50 and only a small portion of the ejaculated or inseminated spermatozoa reach the oviduct.51 The rapid transport of spermatozoa to the oviduct coincides with increased uterine activity.52, 53 Increased myometrial contraction is also responsible for sperm elimination from the uterus through the cervix shortly after breeding. However, not all excess spermatozoa are removed through this mechanism, and other uterine clearance mechanisms are necessary. Because a specific immune response with a memory would be detrimental to fertility during subsequent breeding, elimination of spermatozoa needs to be achieved by a combination of innate immune reactions and mechanical clearance. This mechanism involves a cascade of inflammatory reactions. In vitro and in vivo studies suggest that when equine spermatozoa enter the uterus, they activate complement in uterine secretion.11, 15, 54, 55 The cleavage of factor C5 into C5a and C5b triggers a chemotactic signal to PMNs, resulting in an influx of PMNs into the uterine lumen.29, 55, 56 Activated PMNs bind to spermatozoa, but the nature of this binding is unknown. Recent data suggest that PMN–sperm binding is mediated by the extrusion of DNA from PMNs forming extracellular neutrophil traps (NET) in addition to a traditional ligand receptor binding.57, 58 A mechanism for breeding-induced endometritis has been suggested by the author (Figure 271.8). Following binding, the spermatozoa are phagocytosed by the PMNs. During the activation of PMNs, prostaglandin F2α (PGF2α ) is released from the cell membrane by the metabolism of arachidonic acid via the cyclooxygenase pathway. In addition to being an inflammatory mediator, PGF2α causes contraction of smooth muscle, including the myometrium.59 Uterine contractions are believed to physically remove accumulated fluid and harmful inflammatory products from the uterus.29 Once these products are removed from the uterine lumen, the inflammation subsides, and
Myometrial Contractions
Clearance through the cervix and lymphatics
Figure 271.8 A proposed model for the mechanism of breeding-induced endometritis. The introduction of spermatozoa into the uterus triggers activation of complement in uterine secretion. Complement cleavage factor C5a is a chemoattractant for PMNs, which migrate into the uterine lumen. The PMNs phagocytose spermatozoa and bacterial contaminants, and PGF2α is released from activated PMNs and endometrial cells, causing myometrial contractions. The myometrial contractions serve to physically remove inflammatory products and fluid from the uterine lumen, and the inflammation subsides.29
the uterine environment returns to its normal state. It is important to remember that breeding-induced uterine inflammation is a physiological reaction to semen, and it appears to be a normal process by which sperm is eliminated from the mare’s reproductive tract. The reaction is probably necessary for normal survival and development of an embryo. However, a failure of the uterine defense mechanisms to effectively eliminate an antigen and inflammatory products from the uterus results in persistent endometritis. In vitro experiments have shown that seminal plasma has a suppressive effect on complement activation, PMN chemotaxis, and phagocytosis.60–62 A function of seminal plasma may be to act as an inflammatory modulator in the uterus, which could be of importance for the transient nature of breedinginduced endometritis. Although in vivo experiments have shown a PMN influx into the uterine lumen in response to seminal plasma,56, 63, 64 the duration of the induced inflammation is consistently shorter when seminal plasma is included in the insemination dose.64, 65 The mechanism by which seminal plasma shortens the duration of breeding-induced endometritis is not clear, but it could be the result of a modulation of inflammatory mediators, or a stimulatory effect on myometrial activity.59, 66 The latter possibility may be less likely since Katila and co-workers67 reported a suppressive effect of seminal plasma on uterine contractility in horses. Clinical observations suggest that a marked and prolonged breeding-induced endometritis often follows
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Endometritis insemination with frozen/thawed semen. It has been suggested that removal of seminal plasma before freezing of semen may contribute to the increased duration of breeding-induced endometritis. It is not clear, however, if the reduction of seminal plasma during preparation for cryopreservation is enough to suppress the modulatory effect of seminal plasma on breeding-induced endometritis.65 Another function of seminal plasma in breedinginduced endometritis is to protect spermatozoa from being phagocytosed and destroyed in an inflammatory environment. PMNs are present in the uterine lumen by 0.5 hours after breeding, but sperm transport is not completed until 3–4 hours later.50, 68 In addition, when mares are inseminated twice within a 24hour period, semen from the second insemination is introduced into an inflammatory environment. This environment is detrimental to sperm motion characteristics, and spermatozoa appear to bind to PMNs forming large clusters of PMN–spermatozoa.69 Addition of seminal plasma reduces the binding between motile spermatozoa and inflammatory cells in vitro, and in vivo experiments also support a protective role of seminal plasma from PMN-binding and phagocytosis of spermatozoa.70 Insemination of viable spermatozoa without any seminal plasma into an inflammatory environment at 12 hours after a previous insemination with killed spermatozoa was detrimental to fertility, but the fertility was restored to normal levels if seminal plasma was added to the second insemination dose.70 The results are, however, somewhat conflicting with clinical observations that mares can conceive after AI with frozen/thawed semen, which often contains less than 5% seminal plasma, and in the presence of an ongoing inflammation from a previous AI.71, 72 Further investigations are necessary to determine a critical amount of seminal plasma that is necessary for suppression of PMN–sperm binding. In vitro experiments have demonstrated that the effect of seminal plasma on PMN–sperm binding and phagocytosis is confined to a specific protein fraction in seminal plasma.60, 70 Subsequent studies demonstrated that the effect of this protein is specific in its protection of viable but not dead spermatozoa73, 74 or bacteria (MHT Troedsson, unpublished observation) from PMN-binding and phagocytosis. It would, of course, be beneficial to fertility if viable spermatozoa were protected from being phagocytosed while the uterus is able to maintain an effective elimination of non-viable spermatozoa and bacteria. Selective protection of viable spermatozoa from binding and phagocytosis by PMNs would increase their survival in a hostile uterine environment and ensure that a sufficient number of spermatozoa reach the oviduct for fertilization.
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Diagnostic approach Diagnostically, it may be difficult to identify susceptibility to breeding-induced endometritis prior to breeding. Some mares have free fluid present in the uterine lumen prior to breeding, but most mares are not diagnosed until after they have been bred. Mares with a history of persistent breeding-induced endometritis are likely to develop this problem again. If susceptibility to persistent breeding-induced endometritis is suspected, the mare should be monitored closely by ultrasonography per rectum at 6 to 12 hours after breeding if possible, and at a minimum within 24 hours after breeding. If free fluid is present in the uterine lumen, the mare should be considered to have persistent mating-induced endometritis. The use of scintigraphy to measure uterine clearance has been suggested to be useful when identifying mares that are susceptible to persistent breeding-induced endometritis.22
Treatment and prognosis Management of mares susceptible to persistent breeding-induced endometritis should include limited uterine exposure to semen and bacteria, and assisting the uterus to physically clear contaminants and inflammatory products after breeding.8, 75, 76 Preexisting uterine infections should be resolved before the mare is bred. Exposure to semen should be limited to a single breeding per cycle, if possible. This can be accomplished by closely monitoring follicular development and hormonal treatment to induce ovulation of mature follicles. If intrauterine fluid accumulation is detected before the mare is bred, she can be treated with uterine lavage (using lactated Ringer’s solution or a buffered saline solution) prior to breeding.77 After breeding, physical clearance can be assisted by the use of ecbolic drugs. Oxytocin or PGF2α treatment at 4–8 hours after breeding has been shown to aid in uterine clearance, resulting in improved pregnancy rates in susceptible mares.8, 78, 79 Low doses of oxytocin (10–20 IU) appear to result in a more physiological myometrial contraction pattern and a more efficient clearance of intrauterine fluid when compared to higher doses.80, 81 Care must be taken with regards to the timing of PGF2α treatment. Recent reports have demonstrated that PGF2α can cause a delay in the formation of a functional corpus luteum (CL) when administered within 2 days after ovulation.82–86 This was associated with pregnancy failure in two of the reports.83, 84 Large-volume uterine lavage with a buffered saline solution at 6–24 hours after breeding will also effectively assist the uterus in clearing fluid and inflammatory products (Figure 271.9).76 Fluid is
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Figure 271.9 Uterine lavage at 6 hours after breeding of a mare susceptible to persistent breeding-induced endometritis.
infused into the uterine lumen and completely recovered in one-liter increments until the recovered fluid is clear. There is a strong correlation between the clarity of recovered fluid and the presence of inflammatory cells in the fluid. Many clinicians prefer to combine uterine lavage with the use of an ecbolic drug. Because sperm transport to the oviduct is completed within 4 hours after breeding, uterine lavage between 6 and 24 hours after breeding will not have a negative effect on fertility.50 Manual dilation of the cervix in mares with poor cervical dilation may help these mares to more effectively clear the uterus from fluid. The use of corticosteroids in mares with excessive inflammation in response to breeding has been suggested.87 The authors administered acetate 9-alphaprednisolone (0.1 mg/kg) twice daily during estrus, starting when a follicle >35 mm was detected and ending when ovulation was confirmed. In a recent study, including more than 500 mares, 50 mg of dexamethasone was administered at the time of breeding.88 Pregnancy rates were improved in mares with more than three risk factors for reproductive failure. The results are encouraging, but in contrast to a previous study evaluating the effect of dexamethasone (40 ug/kg) in mares with experimentally induced endometritis.89 These authors did not find a beneficial therapeutic effect of dexamethasone in reducing inflammation compared to antibiotics alone. Further
research is apparently needed to clarify the mechanism of action for this treatment option. While the anti-inflammatory effect of corticosteroids may modulate the breeding-induced inflammation, it would also suppress the beneficial effect of prostaglandin on myometrial activity, and possibly impair PMN function. Dell’Aqua and co-workers in 2006 observed an adverse effect of corticosteroids on the killing capacity of PMNs.90 However, no effect on PMN migration or phagocytosis was observed following treatment of mares with 50 mg dexamethasone (M.H.T. Troedsson, unpublished observation).88 A potential side effect of repeated dexamethasone treatment during estrus is suppression of luteinizing hormone (LH) and possibly ovulation failure (PM McCue, personal communication). This side effect has not been observed when dexamethasone is used as a single dose at the time of breeding. Electroacupuncture has been used clinically to increase uterine contractility in mares with delayed uterine clearance. Anecdotal reports are encouraging, and research is needed to confirm the efficacy of this treatment alternative. It is important for the clinician to keep in mind that a transient inflammatory response to semen is normal and required for normal fertility. Post-breeding treatments of these mares will most likely not improve fertility but may cause further contamination and interfere with pregnancy. Only some 10–15% of all broodmares develop a pathologically persistent form of breeding-induced endometritis.3 Attention should be given to identify and manage these mares appropriately in order to optimize the reproductive efficiency.
References 1. Pascoe RR. Observations on the length and angle of declination of the vulva and its relation to fertility in the mare. J Reprod Fertil Suppl 1979;27:299–305. 2. Troedsson MHT. Uterine response to semen deposition in the mare. In: Proceedings of the Annual Meeting of the Society of Theriogenology, 1995; pp. 130–5. 3. Zent WW, Troedsson MHT. Postbreeding uterine fluid accumulation in a normal population of Thoroughbred mares: a field study. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 1998; pp. 64–5. 4. Neely DP, Kindahl H, Stabenfeldt GH, Edqvist LE, Hughes JP. Prostaglandin release patterns in the mare: physiological, pathophysiological, and therapeutic responses. J Reprod Fertil Suppl 1979;27:181–9. 5. Hughes JP, Loy RG. Investigations on the effect of intrauterine inoculation of Streptococcus zooepidemicus in the mare. In: Proceedings of the Fifteenth Annual Convention of the American Association of Equine Practitioners, 1969; pp. 289–92. 6. Peterson FB, McFeely RA, David JSE. Studies on the pathogenesis of endometritis in the mare. In: Proceedings of the
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7.
8.
9. 10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
Annual Convention of the American Association of Equine Practitioners, 1969; pp. 279–85. Lehrer RI, Ganz T, Selsted ME, Babior BM, Curnutte JT. Neutrophils and host defense. Ann Intern Med 1988;109:127–42. Pycock JF. Assessment of oxytocin and intrauterine antibiotics on intrauterine fluid and pregnancy rates in the mare. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 1994; pp. 19–20. Lees P, Dawson J, Sedgwick AD. Eicosanoids and equine leucocyte locomotion in vitro. Equine Vet J 1986;18:493–7. Pycock JF, Allen WE. Inflammatory components in uterine fluid from mares with experimentally induced bacterial endometritis. Equine Vet J 1990;22:422–5. Watson ED, Stokes CR, Bourne FJ. Cellular and humoral defence mechanisms in mares susceptible and resistant to persistent endometritis. Vet Immunol Immunopathol 1987;16:107–21. Watson ED, Stokes CR, Bourne FJ. Influence of arachidonic acid metabolites in vitro and in uterine washings on migration of equine neutrophils under agarose. Res Vet Sci 1987;43:203–7. Asbury AC, Halliwell RE, Foster GW, Longino SJ. Immunoglobulins in uterine secretions of mares with differing resistance to endometritis. Theriogenology 1980;14: 299–308. Mitchell G, Liu IK, Perryman LE, Stabenfeldt GH, Hughes JP. Preferential production and secretion of immunoglobulins by the equine endometrium – a mucosal immune system. J Reprod Fertil Suppl 1982;32:161–8. Troedsson MH, Liu IK, Thurmond M. Immunoglobulin (IgG and IgA) and complement (C3) concentrations in uterine secretion following an intrauterine challenge of Streptococcus zooepidemicus in mares susceptible to versus resistant to chronic uterine infection. Biol Reprod 1993;49:502–6. Waelchli RO, Winder NC. Mononuclear cell infiltration of the equine endometrium: immunohistochemical studies. Equine Vet J 1991;23:470–4. Williamson P, Dunning A, O’Connor J, Penhale WJ. Immunoglobulin levels, protein concentrations and alkaline phosphatase activity in uterine flushings from mares with endometritis. Theriogenology 1983;19:441–8. Kolm G, Klein D, Knapp E, Watanabe K, Walter I. Lactoferrin expression in the horse endometrium: relevance in persisting mating-induced endometritis. Vet Immunol Immunopathol 2006;114:159–67. Fumuso EA, Aguilar J, Giguere S, Rivulgo M, Wade J, Rogan D. Immune parameters in mares resistant and susceptible to persistent post-breeding endometritis: effects of immunomodulation. Vet Immunol Immunopathol 2007;118:30–9. Evans MJ, Hamer JM, Gason LM, Irvine CH. Factors affecting uterine clearance of inoculated materials in mares. J Reprod Fertil Suppl 1987;35:327–34. Troedsson MH, Liu IK. Uterine clearance of non-antigenic markers (51Cr) in response to a bacterial challenge in mares potentially susceptible and resistant to chronic uterine infections. J Reprod Fertil Suppl 1991;44:283–8. LeBlanc MM, Neuwirth L, Asbury AC, Tran T, Mauragis D, Klapstein E. Scintigraphic measurement of uterine clearance in normal mares and mares with recurrent endometritis. Equine Vet J 1994;26:109–13. Troedsson MH, Liu IK, Ing M, Pascoe J, Thurmond M. Multiple site electromyography recordings of uterine activity following an intrauterine bacterial challenge in mares sus-
24.
25.
26.
27.
28. 29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
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ceptible and resistant to chronic uterine infection. J Reprod Fertil 1993;99:307–13. Rigby SL, Barhoumi R, Burghardt RC, Colleran P, Thompson JA, Varner DD, Blanchard TL, Brinsko SP, Taylor T, Wilkerson MK, Delp MD. Mares with delayed uterine clearance have an intrinsic defect in myometrial function. Biol Reprod 2001;65:740–7. Alghamdi AS, Foster DN, Carlson CS, Troedsson MH. Nitric oxide levels and nitric oxide synthase expression in uterine samples from mares susceptible and resistant to persistent breeding-induced endometritis. Am J Reprod Immunol 2005;53:230–7. LeBlanc MM, Neuwirth L, Jones L, Cage C, Mauragis D. Differences in uterine position of reproductively normal mares and those with delayed uterine clearance detected by scintigraphy. Theriogenology 1998;50:49–54. LeBlanc MM, Johnson RD, Mays MBC, Valderrama C. Lymphatic clearance of India ink in reproductively normal mares and mares susceptible to endometritis. Equine Reproduction VI, 1995; pp. 501–6. Asbury AC. Infectious and immunologic considerations in mare infertility. Comp Cont Educ Pract 1987;9:585–92. Troedsson MH. Uterine clearance and resistance to persistent endometritis in the mare. Theriogenology 1999;52:461–71. LeBlanc MM. The chronically infertile mare. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 2008, pp. 391–407. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: Technique and preliminary findings. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 1975; pp. 327–36. Maloufi F, Pierson R, Otto S, Ball C, Card C. Mares susceptible or resistant to endometritis have similar endometrial echographic and inflammatory cell reactions at 96 hours after infusion with frozen semen and extender. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 2002; pp. 51–7. Blanchard TL, Garcia MC, Hurtgen JP, Kenney RM. Comparison of two techniques for obtaining endometrial bacteriologic cultures in the mare. Theriogenology 1981;16: 85–93. Ball BA, Shin SJ, Patten VH, Lein DH, Woods GL. Use of a low-volume uterine flush for microbiologic and cytologic examination of the mare’s endometrium. Theriogenology 1988;29:1269–83. LeBlanc MM, Magsig J, Stromberg AJ. Use of a low-volume uterine flush for diagnosing endometritis in chronically infertile mares. Theriogenology 2007;68:403–12. Nielsen JM. Endometritis in the mare: a diagnostic study comparing cultures from swab and biopsy. Theriogenology 2005;64:510–18. Riddle WT, LeBlanc MM, Stromberg AJ. Relationships between uterine culture, cytology and pregnancy rates in a Thoroughbred practice. Theriogenology 2007;68: 395–402. Moller-Nielsen J, Troedsson MHT, Zent WW. Results of bacterial and cytological examinations of the endometrium in a practice in Denmark and in Central Kentucky, USA. In: Havemeyer Workshop on the Chronically Infertile Mare, 2008; pp. 34–5. Kenney RM, Doig PA. Equine endometrial biopsy. In: D. Morrow (ed) Current Therapy in Theriogenology. Philadelphia, PA: W.B. Saunders, 1986; pp. 723–9.
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40. Schoon D, Schoon HA, Klug E. Angioses in the equine endometrium – Pathogenesis and clinical correlations. Pferdeheilkunde 1999;15:541–6. 41. Schoon HA, Schoon D. The category I mare (Kenney and Doig 1986): Expected foaling rate 80–90% – fact or fiction? Pferdeheilkunde 2003;19:698–701. 42. Brown MP, Embertson RM, Gronwall RR, Beal C, Mayhew IG, Curry SH. Amikacin sulfate in mares: pharmacokinetics and body fluid and endometrial concentrations after repeated intramuscular administration. Am J Vet Res 1984;45:1610–13. 43. Orsini JA, Park MI, Spencer PA. Tissue and serum concentrations of amikacin after intramuscular and intrauterine administration to mares in estrus. Can Vet J 1996;37:157–60. 44. Couto MA, Hughes JP. Sexually transmitted (venereal) diseases of horses. In: A.O. McKinnon, J.L. Voss (eds) Equine Reproduction. Philadelphia, PA: Lea & Febiger, 1993; pp. 845–54. 45. Swerczek TW. Contagious equine metritis – outbreak of the disease in Kentucky and laboratory methods for diagnosing the disease. J Reprod Fertil Suppl 1979;361–5. 46. Rohrbach B, Sheerin P, Steiner J, Matthews P, Cantrell C, Dodds L. Use of Propionibacterium acnes as adjunct therapy in treatment of persistent endometritis in the broodmare. Anim Reprod Sci 2006;94:259–60. 47. Asbury AC. Uterine defense mechanisms in the mare: The use of intrauterine plasma in the management of endometritis. Theriogenology 1984;21:387–93. 48. Neves AP, Keller A, Trein CR, Moller G, Jobim MI, Castilho LF, Cardoso MR, Leibold W, Zerbe H, Klug E, Gregory RM, Mattos RC. Use of leukocytes as treatment for endometritis in mares experimentally infected with Streptococcus equi subsp. zooepidemicus. Anim Reprod Sci 2007;97:314–22. 49. Kirkland KD, Fales WH, Blanchard TL, Youngquist RS, Hurtgen JP. The in vitro effects of edta-tris, edtatris-lysozyme, and antimicrobial agents on equine genital isolants of Pseudomonas aeruginosa. Theriogenology 1983;20:287–95. 50. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post-insemination on pregnancy rates in mares. Theriogenology 1991;35:1111–19. 51. Scott MA, Liu IKM, Overstreet JW. Sperm transport to the oviducts: Abnormalities and their clinical implications. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 1995; pp. 1–2. 52. Troedsson MH, Liu IK, Crabo BG. Sperm transport and survival in the mare: a review. Theriogenology 1998;50:807–18. 53. Katila T, Sankari S, Makela O. Transport of spermatozoa in the reproductive tracts of mares. J Reprod Fertil Suppl 2000;56:571–8. 54. Asbury AC, Schultz KT, Klesius PH, Foster GW, Washburn SM. Factors affecting phagocytosis of bacteria by neutrophils in the mare’s uterus. J Reprod Fertil Suppl 1982;32:151–9. 55. Troedsson MHT, Steiger BN, Ibrahim NM, Foster DN, Crabo BG. Mechanisms of sperm induced endometritis in the mare. Biol Reprod 1995;52:307 [abstract]. 56. Kotilainen T, Huhtinen M, Katila T. Sperm-induced leukocytosis in the equine uterus. Theriogenology 1994;41:629–36. 57. Alghamdi AS, Foster DN. Seminal DNase frees spermatozoa entangled in neutrophil extracellular traps. Biol Reprod 2005;73:1174–81. 58. Brinkmann V, Reichard U, Goosmann C, Fauler B, Uhlemann Y, Weiss DS, Weinrauch Y, Zychlinsky A. Neutrophil extracellular traps kill bacteria. Science 2004;303:1532–5.
59. Troedsson MHT, Liu IKM, Ing M, Pascoe J. Smooth muscle electrical activity in the oviduct and the effect of oxytocin prostaglandin F2 alpha and prostaglandin E2 on the myometrium and the oviduct of the cycling mares. Equine Reprod VI, 1995; pp. 439–52. 60. Dahms BJ, Troedsson MHT. The effect of seminal plasma components on opsonisation and PMN-phagocytosis of equine spermatozoa. Theriogenology 2002;58:457–60. 61. Troedsson MHT, Lee CS, Franklin RK, Crabo BG. Postbreeding uterine inflammation: The role of seminal plasma. J Reprod Fertil Suppl 2000;56:341–9. 62. Troedsson MHT, Alghamdi AS, Mattisen J. Equine seminal plasma protects fertility of spermatozoa in an inflamed uterine environment. Theriogenology 2002;58:453–6. 63. Bollwein H, Sowade C, Stolla R. The effect of semen extender, seminal plasma and raw semen on uterine and ovarian blood flow in mares. Theriogenology 2003;60:607–16. 64. Fiala SM, Pimentel CA, Steiger K, Mattos ALG, Gregory RM, Mattos RC. Effect of skim milk and seminal plasma uterine infusions in mares. Theriogenology 2002;58:491–4. 65. Troedsson MH, Loset K, Alghamdi AM, Dahms B, Crabo BG. Interaction between equine semen and the endometrium: the inflammatory response to semen. Anim Reprod Sci 2001;68:273–8. 66. Crane LH, Martin L. Postcopulatory myometrial activity in the rat as seen by video-laparoscopy. Reprod Fertil Dev 1991;3:685–98. 67. Katila T, Portus BJ, Reilas T. The effect of seminal plasma on uterine inflammation and contractility in mares. In: 15th International Congress on Animal Reproduction, 2004; pp. 392 [abstract]. 68. Katila T. Onset and duration of uterine inflammatory response of mares after insemination with fresh semen. Equine Reprod VI, 1995; pp. 515–17. 69. Alghamdi A, Troedsson MHT, Laschkewitsch T, Xue JL. Uterine secretion from mares with post-breeding endometritis alters sperm motion characteristics in vitro. Theriogenology 2001;55:1019–28. 70. Alghamdi AS, Foster DN, Troedsson MH. Equine seminal plasma reduces sperm binding to polymorphonuclear neutrophils (PMNs) and improves the fertility of fresh semen inseminated into inflamed uteri. Reproduction 2004;127:593–600. 71. Metcalf ES. The effect of post-insemination endometritis on fertility of frozen stallion semen. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners, 2000; pp. 330–1. 72. Squires EL, Reger HP, Maclellan LJ, Bruemmer JE. Effect of time of insemination and site of insemination on pregnancy rates with frozen semen. Theriogenology 2002;58:655–8. 73. Troedsson MH, Desvousges A, Alghamdi AS, Dahms B, Dow CA, Hayna J, Valesco R, Collahan PT, Macpherson ML, Pozor M, Buhi WC. Components in seminal plasma regulating sperm transport and elimination. Anim Reprod Sci 2005;89:171–86. 74. Troedsson MHT, Desvousges AL, Hansen PJ, Buhi WC. Equine seminal plasma proteins protect live spermatozoa from PMN-binding and phagocytosis, while providing a mechanism for selective sperm elimination of apoptotic and dead spermatozoa. Anim Reprod Sci 2006;94:60–1. 75. LeBlanc MM. Oxytocin – the new wonder drug for treatment of endometritis? Equine Vet Educ 1994;6:39–43. 76. Troedsson MH, Scott MA, Liu IK. Comparative treatment of mares susceptible to chronic uterine infection. Am J Vet Res 1995;56:468–72.
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Endometritis 77. Vanderwall DK, Woods GL. Effect on fertility of uterine lavage performed immediately prior to insemination in mares. J Am Vet Med Assoc 2003;222:1108–10. 78. Pycock JF, Newcombe JR. The relationship between intraluminal uterine fluid, endometritis, and pregnancy rate in the mare. Equine Pract 1996;18:19–22. 79. Rasch K, Schoon HA, Sieme H, Klug E. Histomorphological endometrial status and influence of oxytocin on the uterine drainage and pregnancy rate in mares. Equine Vet J 1996;28:455–60. 80. Madill S, Troedsson MHT, Santschi EM. Individual registration of longitudinal and circular muscle in the equine endometrium using electromyography and strain gauges: Preliminary data. In: Havemeyer Foundation International Workshop on Uterine Defense Mechanisms in the Mare: Aspects of Physical Clearance, 1997; pp. 10 [abstract]. 81. Campbell MLH, England GCW. A comparison of the ecbolic efficacy of intravenous and intrauterine oxytocin treatments. Theriogenology 2002;58:473–8. 82. Gunthle LM, McCue PM, Farquhar VJ, Foglia RA. Effect of prostaglandin administration post-ovulation on corpus luteum formation in the mare. In: Proceedings of the Annual Meeting of the Society for Theriogenology, 2000; pp. 139–40. 83. Troedsson MH, Ababneh MM, Ohlgren AF, Madill S, Vetscher N, Gregas M. Effect of periovulatory prostaglandin F2alpha on pregnancy rates and luteal function in the mare. Theriogenology 2001;55:1891–9.
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84. Brendemuehl JP. Effect of oxytocin and cloprostenol on luteal formation, function and pregnancy rates in mares. Theriogenology 2002;58:623–6. 85. Nie GJ, Johnson KE, Wenzel JGW, Braden TD. Effect of periovulatory ecbolics on luteal function and fertility. Theriogenology 2002;58:461–3. 86. Mocklin CM, Paccamonti DL, Eilts BE, Lyle SK, Wouters EGH, Gentry LR, Godke RA. Effect of post-ovulatory prostaglandin F2 alpha or cloprostenol on plasma progesterone concentration in mares. Anim Reprod Sci 2006;94:220–2. 87. Dell’Aqua JA Jr, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Neves Neto JR, Melo CM, Duarte MP, Duarte MCG. Fertility rates with equine frozen semen after modulation of inflammatory uterine response. In: Proceedings of the 15th International Congress on Animal Reproduction, 2004; pp. 391. 88. Bucca S, Carli A, Buckley T, Dolci G, Fogarty U. The use of dexamethasone administered to mares at breeding time in the modulation of persistent mating induced endometritis. Theriogenology 2008;70:1093–100. 89. McDonnell AM, Watson ED. The effects of dexamethasone sodium-phosphate on mares with experimentally induced endometritis. J Equine Vet Sci 1993;13:202–6. 90. Dell’Aqua JJA, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Melo CM. Modulation of acute inflammatory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94:270–3.
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Chapter 272
Breeding the Problem Mare Michelle M. LeBlanc and Angus O. McKinnon
Introduction Management of the ‘problem mare’ is one of the most challenging and potentially frustrating aspects of equine reproduction. Solving the problem requires hard work, persistence, adherence to scientific principles, and use of proven management techniques. Clients need to be informed that costs will likely be much greater than those incurred with a reproductively normal mare because additional diagnostics and more intensive management are commonly required. Frequent communication between the referring veterinarian, client, and farm manager is needed to alleviate client anxiety, to keep the referring veterinarian informed, and to avoid the possibility of client or farm manager dissatisfaction if the mare fails to become pregnant. In difficult or protracted cases, it is prudent to send a detailed letter summarizing the history, clinical findings, diagnostic results, and recommendations to all parties involved in order to prevent misunderstandings. The letter also serves as a template from which to work if treatment is not successful or the mare does not become pregnant. Options with realistic success rates should be outlined and reviewed with the client. For example, recommending embryo or oocyte transfer in a chronically infected mare may be a less expensive route to producing a healthy foal than repeated, prolonged uterine therapy in breeds that allow advanced reproductive techniques. None of us are able to solve all infertility problems nor can we successfully communicate with all clients. Consulting with or referring a case to another specialist is almost always productive and may be greatly appreciated by the client. Our approach to diagnosing and breeding the problem mare is presented here. There are many other philosophies and personal preferences and different avenues of treatment (much of it anecdotal, undocumented or not scientifically validated). However, until experimental models of various infertility
problems are developed it is not only difficult, but also inappropriate, to be dogmatic in recommendations for management.
What is a problem broodmare? ‘When’ a mare begins to be regarded as a problem breeder is at the discretion of the owner, veterinarian, or farm manager. Classically, a mare should be regarded as a potential problem if she fails to conceive to a fertile stallion on a well-managed breeding farm on three or more cycles in one season, loses her pregnancy before 60 days of gestation, or does not exhibit normal estrous cycles. A subfertile stallion, improper handling of semen, inadequate semen dose and/or insemination techniques in an artificial insemination program are excellent methods for dramatically reducing pregnancy rates. Unfortunately, fertility of a stallion is not always known to mare owners. Obtaining pregnancy rate per cycle from previous years or pregnancy data from maiden, barren, and foaling mares will help determine if stallion fertility or management procedures should be questioned. Fertility is impacted by many factors. Seasonality, aging, parity, previous trauma to the reproductive tract, systemic problems such as laminitis or Cushing’s disease, poor nutrition, obesity, group dynamics, and environmental conditions such as temperature or rainfall can all impact reproduction. In addition, the estrous cycle changes dynamically from the transitional phase in the spring into recurring estrous cycles in the summer, and from estrus to diestrus or pregnancy. Lack of synchrony between hormonal events and physical changes of the reproductive tract can result in delayed uterine clearance, infection, prolonged inflammation, or pregnancy loss. There are many reasons why mares may become problematic. The most common type of problem mare presented is one with a persistent, unidentified, or inappropriately treated uterine infection. Therefore, the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Breeding the Problem Mare emphasis will be on mares with acute and/or chronic uterine inflammatory changes that present with uterine, cervical, and/or oviductal pathology. Problems such as lack of ovarian activity or management of mares presenting with abnormal estrous cycles will be presented in less detail (Table 272.1).
How infection or prolonged inflammation occurs Susceptibility to infection is a graded condition and occurs primarily due to effects of increasing age, reproductive tract damage, and bacterial challenge. Bacterial challenge is influenced by external and internal conformation, breeding techniques, examination procedures, anatomical abnormalities, and postpartum events. Contamination of the uterus with bacteria is inevitable. Potentially pathogenic organisms are introduced at breeding, during and after parturition, during examination, and as a result of failure of physical barriers to infection (i.e., pneumovagina). When uterine defense mechanisms are functioning properly, massive challenges, either natural or experimental, fail to produce inflammation that lasts long enough to interfere with reproduction.1, 2 Mating results in a transient inflammation within the uterine lumen.3–5 Reproductively healthy mares clear the inflammation within 24–72 hours by a combination of mechanical and cellular defense mechanisms. Mechanical mechanisms include myometrial contractions, which assist in evacuation of uterine contents through an open relaxed cervix. The position of the uterus within the abdomen greatly influences mechanical clearance. Old, pluriparous mares with a dependent uterus cannot effectively clear uterine contents quickly. The cellular response is primarily phagocytosis. Efficient phagocytosis depends on (i) mobilization of an adequate number of neutrophils from the general circulation with prompt migration of these cells through the endometrium and into the uterine lumen; (ii) adequate chemotaxis of neutrophils to contaminating bacteria and dead or dying semen; (iii) adherence of bacteria to the cellular membrane of the phagocyte (opsonization) with the assistance of immunoglobulin and complement; and (iv) ingestion and successful intracellular killing of bacteria or sperm by neutrophils. Neutrophil numbers are highest in the uterine lumen between 8 and 12 hours after breeding,5 after which time numbers decrease dramatically. Those mares that fail to clear the semen-induced inflammation within 24–48 hours accumulate uterine fluid and retain endometrial edema.6–9 Intrauterine fluid collected from susceptible mares contains neutrophils, immunoglobulins, protein, semen, and bacteria. These compounds irritate the endometrium resulting in a continual influx of neutrophils into the lumen. A vicious cycle
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Table 272.1 Chronic infertility in the mare: client complaints and their causes Mare not exhibiting normal estrous cycles Anovulatory follicles Cause unknown (aging) Insulin resistance, Cushing’s disease Endometritis Persistent endometrial cups Secondary to administration of prostaglandin in mid-diestrus Delayed follicular development secondary to aging Lactational anestrus Metabolic (insulin resistance, Cushing’s disease) Ovarian tumors Pain Laminitis, musculoskeletal, colic Poor body condition Retention of endometrial cups Mare has intrauterine fluid during estrus OR immediately after breeding Anatomical abnormalities Pneumovagina Recto-vaginal fistulas or tears Vestibulovaginal reflux of urine Cervical fibrosis, traumatic tears or adhesions Pendulous uterus Uterine adhesions Degenerative abnormalities Cervical incompetence secondary to aging (old, maiden mare) Abnormal uterine contractions Cysts – endometrial and lymphatic Endometrial glandular ectasia Lymphatic lacunae Vascular elastosis Infectious or inflammatory Bacterial or yeast endometritis Increased endometrial glandular secretions Foreign bodies (mummified fetus, retention of marble) Increased mucus production Mare is infected repeatedly OR infection does not resolve after repeated treatments List is the same as for fluid retention (above). Emphasis on: Anatomical problems Caudal reproductive tract Cervical lacerations, adhesions Cervical fibrosis Pendulous uterus Chronic infectious endometritis Gram-negative infections Yeast or fungal infections Inadequate uterine perfusion Mare loses pregnancy after 14 days and before 45 days Cervical tears (may be associated with endometritis) Chromosomal abnormalities, ‘old eggs’ Endometritis (see above) Failure of endometrial glands – periglandular fibrosis Luteal insufficiency Endotoxemia, laminitis, colic Lymphatic lacunae (may be associated with endometritis, inadequate blood flow) Nutritional/colic/toxins Adapted from LeBlanc.109
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of inflammation, endometrial damage, and increased fluid accumulation results in infertility. The critical factor in uterine defense against persistent endometritis is the physical ability of the uterus rapidly to clear the contents of the uterine lumen after mating or foaling. Physical clearance can be decreased by anatomical changes, degenerative changes, or a combination of both. Anatomical abnormalities such as an incompetent vulvar seal or cervix, vaginal stretching, or a pendulous uterus contribute to delayed clearance as do degenerative changes such as abnormal uterine contractions, vascular elastosis, lymphangiectasia, scarring and atrophy of endometrial folds, or a damaged mucociliary apparatus.6, 8,10–16 Chronic inflammation can result in periglandular fibrosis and inappropriate mucus production by the uterine epithelium or endometrial glands.17–19 Causey speculates that endometrial cilia and the mucus layer overlying the endometrium play an important role in uterine clearance and uterine defense against bacterial colonization. If the elasticity, viscosity, or quantity of mucus changes, if cilia are damaged or the endometrium is denuded of epithelial cells, uterine drainage will be diminished, thereby contributing to fluid accumulation and infertility. Susceptibility to chronic infections can occur gradually due to anatomical or degenerative uterine changes, acutely as a consequence of injury from a traumatic foaling or secondary to repeated manipulations from sequential embryo flushes. Therefore, clinicians need to determine what factors, historical, anatomical, or degenerative, are contributing to delayed uterine drainage and develop individual therapies for affected mares.
Clinical examination Clients should be encouraged to have a reproductive examination performed on their mare at the end of the breeding season before the autumn in order to determine the cause of the infertility and to institute treatments or perform surgery in order to correct the problem before winter. A report of all findings should be sent to the client as soon as possible so options and prognoses can be discussed. A diagnostic workup should include a detailed history that includes past performance and reproductive findings, a complete physical examination, evaluation of perineal conformation, rectal palpation and ultrasonography of the reproductive tract, vaginal speculum examination, and manual examination of the cervix. Laboratory tests are based on clinical findings and may include uterine culture, uterine cytology, endometrial biopsy, and endoscopic evaluation of the uterus. Additional diagnostic tests reserved for specific cases include
endocrine testing, karyotype, laparoscopy, and oviductal patency test. A physical examination should always be performed because systemic abnormalities can contribute to infertility. Chronic pain, lameness, poor body condition, or obesity have been associated with abnormal or no estrous cycles, anovulatory follicles, and intrauterine fluid accumulation. Farm management and nutritional programs should be reviewed because group dynamics may adversely affect the body condition of young or old mares, especially if mares are timid, fed in pens, crowded during feeding, or not given shelter from cold or wet conditions. Maiden mares off the racetrack are especially sensitive to the pecking order of social groups housed at pasture on breeding farms. It is not uncommon for maiden mares that were confirmed to be cycling on their home farm or racing barn to fail to cycle on the breeding farm. The mare’s history provides important information as to possible causes of the infertility and what diagnostics should be performed. Medications given during the mare’s performance career, age when first bred, a history of any difficult births or delivery of large or premature foals, number of healthy foals born, treatments administered around breeding, type of semen used (fresh, cooled, or frozen), and manner by which the mare was bred (artificial insemination or natural mating) should be recorded. Access to previous breeding records is desirable as they may suggest where and why the problems began and why previous treatments were not successful. The pregnancy rate/cycle or the stallion’s past reproductive performance should be obtained. Any history of abnormal estrous cycles, uterine infections, type of microorganism isolated, embryonic loss, or abortion should be recorded. An accurate history will not only help to identify current causes of infertility, but also may help to predict future problems. Perineal and pelvic conformation The anatomy of the pelvic region and perineum of the mare must be evaluated critically because most uterine infections are due to bacteria that normally inhabit the caudal reproductive tract ascending into the anterior vagina. There are three barriers that prevent organisms from entering the uterus – the vulvar seal, the vestibulovaginal fold, and the cervix. Damage to any of these can result in chronic inflammation. Poor perineal conformation is prevalent in Thoroughbreds and is almost unknown in ponies (Figure 272.1). The vulvar seal may be damaged from repeated foalings or the muscular tone of the vulvar lips may be poor, as seen in young fillies off the racetrack or in aged, thin, pluriparous mares. The perineum is best evaluated during estrus, when relaxation and elongation of the vulvar lips are greatest. Optimal perineal
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Figure 272.1 Poor perineal conformation. Note the severe cranial, ventral tilt to the vulva.
conformation consists of a vertically positioned vulva with two-thirds or more of the vulvar lips located below the pelvic floor. The vulvar lips should have muscular tone and form a tight seal. To prevent pneumovagina and ascending bacterial infection, a Caslick operation should be performed if the dorsal commissure of the vulva is located more than 2.5 cm dorsal to the pelvis or it is tilted more than 20 degrees ventrally (angle of declination). In some thin mares, increasing body condition may be a sufficient remedy. Just how important routine management is has recently been highlighted.20 Of interest is the improvement in pregnancy rates associated with performing a Caslick as soon as practically possible after foaling. The vestibulovaginal fold can be evaluated by parting the vulvar lips (Figure 272.2). It may be stretched or torn as a consequence of dystocia or repeated births or after an aggressive mating. Perineal or pelvic conformation of some mares is so poor that a Caslick operation does not prevent pneumovagina, and a vestibuloplasty (also referred to as a deep Caslick or a Gadd operation) should be considered.
Rectal palpation and ultrasonography The rectal examination provides information concerning follicular development, stage of estrous cy-
Figure 272.2 Damaged vestibulovaginal fold. Loss of this fold can result in aspiration of bacteria, air, and feces into the vagina and uterus.
cle, and pathology of the reproductive tract. Size, tone, and position of the cervix, uterus, and ovarian structures are identified by rectal palpation. The length and width of the cervix and the location of the uterus in relationship to the pelvis should be recorded as findings provide valuable information regarding uterine drainage. For example, mares with cervical fibrosis will have an elongated, narrow cervix during estrus (length > 4–6 cm, width 2–4 cm) whereas mares with a torn cervical muscle will have a short (< 3–4 cm), wide (> 3 cm) cervix that lacks definition. Mares with a uterus located more than 15 cm below the pelvis are prone to persistent matinginduced endometritis (Figure 272.3a,b). Uterine tone (thick, doughy, edematous, flaccid) may be associated with physiological changes in the estrous cycle, seasonality, or pathological processes. Ultrasonography is the principal diagnostic tool used in broodmare practice as it enables one to visualize structures that cannot be discerned by rectal palpation.21, 22 Pathology such as anovulatory follicles, granulosa cell tumors, intrauterine fluid, abnormal uterine edema patterns, and uterine cysts can be identified readily. Subfertile mares presented for
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Figure 272.3 Scintigrams of the reproductive tract taken during estrus. The gamma camera was located in the left flank. Black depicts clearance of radiocolloid. (a) Scintigram from an aged, pluriparous, reproductively healthy mare. Note the horizontal appearance of the uterus and cervix. (b) Scintigram from an aged, pluriparous, subfertile mare. Note the vertical appearance to the uterus
breeding should be scanned every day or every other day during estrus and for 1 to 2 days after breeding and/or ovulation to identify uterine edema patterns, the presence and location of intrauterine fluid accumulation, and to record ovulation. Physical changes to the reproductive tract may not always be synchronized with hormonal changes resulting in subfertility. For example, mares with excessive uterine edema or a small amount of uterine fluid cranial to the cervix on the first or second day of estrus may not dilate their cervix properly (Figures 272.4 and 272.5), whereas fluid in the uterine horns or at the bifurcation is typical of older, pluriparous mares with poor uterine tone and contractility. Other pathologies are only evident after breeding. Retention of uterine edema after ovulation or an increase in edema after breeding are indicative of persistent mating-induced endometritis, lymphatic lacunae, or angiosis.9, 23 The reproductive tract should also be scanned after treatment for endometritis in order to evaluate the response to therapy. Retention of intrauterine fluid or uterine edema indicate that the inflammation has not resolved and treatment should be continued.
Figure 272.4 Ultrasonographic image of the uterus taken on Day 1 of estrus. Excessive uterine edema.
Vaginal speculum examination A vaginal speculum examination should be performed to identify the presence of any pus, urine, or blood in the vaginal vault, discrepancies between cervical relaxation and stage of estrous cycle, the color and moisture of the vaginal mucosa, and the presence of cervical adhesions. Manual examination is recommended for assessing the efficacy of the vestibular sphincter and cervix as barriers to infection. Old, barren, pluriparous mares commonly have pneumovagina because the vestibulovaginal sphincter and vulvar structures have been stretched or torn. Young, thin race mares with poor tone to the vulvar lips, and lack of fat in the perineal region, may also experience pneumovagina. Hyperemia of vaginal mucosa and a frothy exudate in the anterior vagina is pathognomonic24 for pneumovagina. The
Figure 272.5 Ultrasonographic image of the uterus taken on Day 1 of estrus in a barren, maiden mare. Note the presence of fluid (black) cranial to the cervix.
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Breeding the Problem Mare external os of mares with endometritis may be hyperemic, wet, and swollen, or dry and splotchy in color. Vestibulovaginal reflux is most commonly seen in older, barren, pluriparous mares in poor body condition during estrus, after delivery of a large foal, or after dystocia. The urine typically has a cloudy, yellow appearance with a pungent odor. It can be confused with an accumulation of exudate in mares with bacterial endometritis. Manual cervical examination Cervical defects and adhesions are not uncommon. Some mares will become pregnant without surgical correction despite the absence of much of the cervix, whereas others need to be repaired surgically. Choosing which mares require surgical repair depends on the severity of the tear and whether the mare has experienced prolonged, chronic uterine infections. Mares with lesions in the ventral aspect of the cervical canal, such as scarring, stretching, or tearing of cervical folds, tend to accumulate intrauterine fluid after breeding as fluid may pool in the damaged area and serve as a reservoir for bacteria to accumulate. Adhesions between the external cervical os and the vaginal fornix also can interfere with uterine drainage and with the ability of the cervix to close properly. Cervical adhesions or tears are best identified during diestrus, when the cervix is normally tubular, located off the vaginal floor with a rosebud appearance to the external os. Adhesions between the external os and vaginal fornix are identified by placing the index finger in the cervical canal and sliding the thumb over the vaginal portion of the external os to identify any taut tissue bands that may be attached to the lateral or ventral aspects of the external os. Adhesions within the cervical canal feel like thin spider’s webs that can be broken with gentle pressure. Fibrosis of the cervical canal can occur as a consequence of a difficult foaling. It should be suspected in mares that have an elongated, narrow cervical canal that lacks elasticity or a tortuous canal that deviates laterally, dorsally, or ventrally. Mares with the condition have a history of repeated uterine infections or intrauterine fluid accumulation even after prolonged intrauterine therapy, and in some cases mares have a history of early embryonic death. Old, maiden mares (> 10 years of age) and young, nervous fillies off the racetrack may not dilate their cervix completely during estrus. The abnormality can be identified via rectal palpation of the cervix during estrus, by visual inspection of the external os, and by manual examination of the cervical canal. On rectal palpation, the cervix will be long (> 6 cm), narrow (< 2–3 cm), and tubular. On vaginal speculum exam, the cervix may be located off the vaginal floor and have the appearance of a rosebud,
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while manual evaluation reveals a long, narrow cervical canal.
Laboratory diagnostics Endometrial cytology and culture Uterine cytology and culture are the most commonly performed diagnostics for identifying mares with endometritis. Uterine cultures can be collected during estrus by a guarded culture swab that is passed into the uterus either through a sterile vaginal speculum or manually; by a non-guarded swab attached to a sterile extension rod; via a small-volume uterine flush; or by swabbing an endometrial biopsy specimen. In chronically infected mares, culture of the clitoral fossa, vestibule, and anterior vagina may help identify the cause of the repeated infections. Endometrial cytology can be obtained by passing a second swab through the culture guard into the uterus, collecting cells on the cap of a guarded swab, or by using a cytology brush (see Chapter 204 for technique and interpretation of cytology smears). Uterine samples are cultured aerobically for 48 hours on blood agar plates and organisms identified using biochemical methods. Recently, the use of Chrome Agar for identifying uterine organisms in the reproductive tract of mares has been reported25 (see Chapter 209). Endometrial cytology appears to be more accurate than culture swabs in identifying mares with active inflammation, as twice as many mares with acute endometritis were identified by cytology than by bacterial culture in a clinical study involving 2128 paired culture and cytology samples.26 Uterine cultures were collected using a guarded swab, and endometrial cytologies were made with cells contained in the cap of the guarded swab. The pregnancy rate per cycle was significantly affected by the number of neutrophils per 400× field. Mares with more than 5 neutrophils/field had a pregnancy rate of only 23% compared to a 36% rate in mares with 2–5 neutrophils/field, and a 60% pregnancy rate for mares with 0–2 neutrophils/field. Isolation of bacteria on culture swab was also associated with a decreased pregnancy rate (36%) even if the cytological specimen was normal, defined as 0–2 neutrophils/400× field. These authors and others found that Escherichia coli was less likely to be associated with a positive cytology than other bacteria, whereas isolation of Streptococcus equi spp. zooepidemicus was most commonly associated with a positive cytology.26–28 These studies stress the importance of interpreting laboratory data in context with clinical findings as the correlation between cytology and culture results varies between organism recovered.
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In addition, mares with chronic endometritis may have minimal surface irritation and thus may not have many neutrophils in their endometrial cytology smear.29 A negative culture does not mean that there are no bacteria present in the uterus, just that they were not identified. In 2005 Nielsen30 reported that only 38 (45%) of 84 barren mares with bacteria isolated from the surface of an endometrial biopsy had bacteria isolated from a uterine culture swab. In a more recent study of mares with a positive cytology, he found a higher percentage of positive cultures when the culture sample was collected with a culture swab (26%) than if biopsy specimens were swabbed for uterine culture (3%; Nielsen 2008). Uterine pathogens may be missed by routine swab culture techniques because only a small area of endometrium is cultured and infections may be focal. Mares with clinical evidence of endometritis that do not have a pathogen isolated using a traditional swab technique should have uterine samples for culture collected by either small-volume uterine flush or culture of an endometrial biopsy. Culture of uterine flush fluid was twice as sensitive as swab culture previously estimated by Neilsen (0.71 vs 0.34, respectively) in identifying endometritis when the same ‘best standard’ (presence of neutrophils within the endometrium) was used.31 The improved sensitivity resulted from improved detection of Gram-negative organisms as recovery of beta-hemolytic Streptococcus from uterine flush was similar to that reported previously when samples were collected by culture swab. The falsepositive rate, defined as a positive culture result in the presence of a clear efflux and a cytological smear that contained no debris, neutrophils, or bacteria, was 11%. A small-volume flush can be performed during estrus or diestrus. If the sample is collected during diestrus, it is recommended that the mare be given prostaglandin so that she returns to estrus quickly. Before the procedure, the perineum should be scrubbed, rinsed, and dried. A sterile uterine catheter attached to a 150-mL bag of saline via an extension tube is passed per vaginum into the uterus by an examiner whose arm is covered by a sterile sleeve. A 10–12-cm length of catheter is inserted into the uterus and the saline bag is squeezed manually so that fluid passes through the catheter into the uterus. The operator then manipulates the fluid throughout the uterine lumen via the rectum. Saline is collected back into the 150-mL bag through the extension tubing or the tubing is detached from the uterine catheter and the saline is collected into a sterile 50-mL conical tube. Clarity of the efflux is recorded, as recovery of a cloudy flush or a clear flush containing mucoid strains is associated with isolation of bacteria (Figure
Figure 272.6 Cloudy efflux collected from a small-volume flush of the uterus.
272.6).31 If the sample is not clear, the uterus can be irrigated immediately with large volumes of saline. Once collected, the precipitant is allowed to settle or the sample can be centrifuged at 400g for 10 min. The supernatant is poured off and two swabs are placed in the pellet for uterine cytology and culture. Cytological smears should be evaluated for debris, neutrophils, and bacteria, the presence of which is associated with bacterial infection. The flush technique is not as sensitive as a dry swab or cytology brush in recovering neutrophils because the fluid dilutes out the number of inflammatory cells per high-power field and debris can interfere with the identification of such cells. Endometrial biopsy Endometrial biopsy is a valuable tool for identifying the cause of infertility.32 A single biopsy may not always be totally representative of the uterus,33 but a sample from the corpus-corneal junction (site of embryonic fixation) is one of the most accurate determinants of inflammatory conditions and cellular infiltrates. Biopsies are graded on the degree and chronicity of inflammation, fibrosis, and glandular activity. The Kenney–Doig biopsy classification system34 is one of the most important determinants of subsequent foaling rates. However, the grading system does not inform the veterinarian how to treat the mare as it is designed more for foaling probability than to describe the degree of fibrosis and/or classification of acute versus chronic inflammation.
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Air or saline can be used for insufflation. Visibility is greatest using air insufflation; however, air can be irritating to the endometrium so it should be removed after the examination, either through a catheter or pump. The uterus is then irrigated with saline, and in some cases antibiotics are infused if the mare has a history of endometritis.
Identification and breeding management of mares that accumulate intrauterine fluid post-mating Figure 272.7 Exudate overlying endometrium may interfere with antibiotic killing of bacteria.
In addition, the categorization neglects important histopathological findings such as angioses, endometrial maldifferentiation, epithelial hyperplasia, and degree of exudate, all of which will affect clinical and prognostic outcome (Figure 272.7). A number of authors have suggested modifications of the Kenney classification and the addition of recommended therapies and management protocols depending on observed lesions.14, 35 In the future, quantification techniques and other new techniques of either staining (fibrosis) or immunohistochemical and molecularbased testing, like fluorescence in situ hybridization (FISH) and real-time polymerase chain reaction (RTPCR), or genetic array analysis, will likely be introduced and may result in a much more useful diagnostic approach.36 Candidates for endometrial histological evaluation include but are not limited to subfertile mares presented with an incomplete reproductive history, mares with palpable uterine abnormalities on rectal examination, and mares with chronic inflammation or repeated early embryonic death. Endometrial biopsies are also useful in determining treatment success. Endometrial biopsies can be obtained during estrus or diestrus. Samples can be placed in Bouin’s solution (pitric acid) for 8 to 24 hours then transferred into 70–90% alcohol or fixed in 10% formalin. Signalment and history should always be provided to assist the pathologist in interpretation. Endoscopy Uterine endoscopy identifies lesions within the uterine lumen that cannot be identified by ultrasonography. These include focal plaques, inflammatory foci, uterine adhesions, scarring and loss of endometrial folds, foreign bodies, lymphatic cysts, and retention of endometrial cups. The procedure can be performed during diestrus or in early estrus. The endoscope should be cold sterilized for a minimum of 20 minutes before it is placed into the uterine lumen.
Inadequate uterine drainage is the primary cause of subfertility in aged pluriparous mares and in some old and young maiden mares that do not completely dilate their cervix during estrus. Mares with bacterial or fungal infections may also accumulate uterine fluid throughout the estrous cycle or only during estrus. Identifying mares susceptible to post-mating endometritis is not always easy as some mares show no signs of inflammation before mating only to accumulate fluid after mating. While a small amount of intrauterine fluid before mating does not appear to affect pregnancy adversely, fluid volumes exceeding 2 cm have been associated with decreased pregnancy rates.37 The presence of intraluminal fluid 18 hours or more after mating, or retention of uterine edema after ovulation is diagnostic of defective uterine clearance.9, 38 It is possible to identify potential problem mares before breeding as they may have a history of intrauterine fluid retention, an elongated, narrow cervix during estrus, or a dependent uterus that is located 10 cm or more below the pelvis. In order to prevent intrauterine fluid accumulation after mating clinicians must identify what abnormalities are contributing to the fluid accumulation and then formulate an individualized breeding management protocol. Repeating the same treatment cycle after cycle without attempting to diagnose the cause rarely results in a successful pregnancy outcome. The location of fluid within the uterine lumen, whether it is visualized in the uterine body, uterine horns, or in the body directly cranial to the cervix provides evidence as to cause. Mares with a cervix that does not dilate properly during estrus or closes too quickly after ovulation tend to retain fluid just cranial to the cervix. This is most commonly observed in young or old nulliparous mares. In contrast, older pluriparous mares susceptible to post-matinginduced endometritis tend to accumulate fluid in the uterine horns and body. These mares may have a pendulous uterus, poor perineal conformation, and various degenerative changes such as impaired myometrial contractions, poor lymphatic drainage, excessive glandular secretions, or angiosis.6, 10, 11, 39 Mares with
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cervical adhesions or cervical fibrosis are also prone to persistent mating-induced endometritis. Treatment options Treatment for post-mating endometritis is aimed at (i) improving physical clearance of inflammatory by-products associated with insemination; (ii) reducing the length of the inflammatory response; and (iii) preventing bacterial infection (Table 272.2). The currently recommended therapy for improving physical clearance of uterine fluid is uterine irrigation followed immediately by administration of either oxytocin (10–25 IU i.v. or i.m.) or cloprostenol (250 µg i.m.) at 4 to 8 h after breeding.17,40–48 This treatment has increased pregnancy rates in highly susceptible, barren mares.49, 50 The first treatment is begun 4 hours after mating to allow sperm time to travel to the oviduct, and performed by 8 hours to mimic the physiological inflammatory response of reproductively normal mares. Reproductively healthy mares and susceptible mares clear intrauterine bacteria at a similar rate until 9 hours post-inoculation, when clearance rates diverge. No bacteria were isolated from reproductively normal mares after 9 hours while bacteria began a logarithmic growth pattern in susceptible mares, with bacterial numbers peaking 48 hours after inoculation.51 Oxytocin may be administered repeatedly at 4–6-hour intervals for 2 to 3 days to assist in clearance of uterine fluid. Cloprostenol may be given in place of oxytocin if the mare has a history of lymphatic lacunae or does not appear to respond to oxytocin. Acupuncture performed before and after breeding also appears to assist in uterine drainage as clinicians report decreased uterine edema, decreased intrauterine fluid, and increased uterine tone 24 hours after acupuncture treatment. Mares with a history of fluid accumulation after breeding or those with cervical incompetence may also be treated with dexamethasone directly before breeding or may be given low doses of prednisolone for 4 to 5 days beginning 36–48 hours before breeding;38, 52 (see ‘Immunomodulators’ below). All susceptible mares should be re-evaluated 24 hours after breeding to detect the presence of intrauterine fluid and determine if additional treatments such as uterine irrigation, ecbolics, or antibiotics are required. Treatment chosen will depend on the mare’s history, age, follicular status, and economics. If the mare has ovulated and the cervix closed by the 24-hour examination, the perineum must be cleaned meticulously before uterine irrigation in order to avoid iatrogenic inoculation of the uterine lumen. Infusion of a penicillin-based antibiotic after irrigation is recommended in these cases to decrease bacterial growth.
Table 272.2 Treatment options for chronic infertility Therapies that improve physical drainage Oxytocin 10–25 IU Can be administered up to every 6 h Cloprostenol 250 µg Recommended not to be administered more than 12 h after ovulation Uterine irrigation Irrigation with lactated Ringer’s solution can be performed immediately before breeding Four or more hours post breeding: 1–3 L saline or lactated Ringer’s solution (LRS) or either solution with DMSO added (30–100 ml/L) Acupuncture Conducted before (> 24 h from mating) and after breeding (twice at 48-h intervals) Therapies that reduce the duration of or modulate inflammatory response Corticosteroids: Dexamethasone 50 mg Given directly before breeding Prednisolone 0.1 mg/kg Given twice daily beginning 24 to 48 h before breeding and continuing for 3 to 5 days Immunodulators R EqStim – Propionibacterium acnes Three intravenous injections on days 1, 3, and 7 around breeding R Settle Mycobacterium phlei cell-wall extract Intravenous, intramuscular, or intrauterine administration Therapies that inhibit bacterial or fungal growtha Antibiotics/antimycotics Intrauterine or systemic (see Tables 272.3–272.5) Treatment length varies from 3 to 21 days Antiseptics/disinfectants Betadine (5–10 mL of a 10% povidone-iodine solution in 1 L saline) Uterine irrigation for 3 to 5 days; last liter of flush solution should not contain betadine Hydrogen peroxide (20 mL of a 3% solution diluted in 60 mL of LRS) Magnesium sulfate (128 g in 1 L of saline) Vinegar (10–20 mL in 1L of saline) Solvents DMSO (5–30% solution diluted in saline or LRS) Has been added to uterine irrigation solution and used as an infusion (30 mL in 100 mL saline) Kerosene (50 mL diluted in 50 mL of saline; other doses have been used empirically) Chelating agents Tris-EDTA (250 to 750 mL of 3.5 mM EDTA, 0.05 M Tris, pH 8) Synergistic with amikacin, ceftiofur and ticarcillin and antimycotic drugs; however, lyophilized antibiotics such as ceftiofur and ticarcillin will precipitate when added to Tris-EDTA Improves antibiotic killing; must come in direct contact with organism so volume infused depends on size of uterus R Tricide 8 mM disodium EDTA dehydrate and 20 mM 2-amino-2-hydroxymethyl-1,2-propanediol Dose depends on size of uterus: 500 to 750 mL Appears to be more stable than Tris-EDTA (Continuted)
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Breeding the Problem Mare Table 272.2 (Continued) Antimicrobial synergism similar to Tris-EDTA Uterus needs to be irrigated within 24 h of infusion of chelating agent Mucolytic/anti-inflammatory therapya N-acetylcysteine (20% solution: dilute 30 mL in 150 mL of saline) Used as a pre-breeding infusion (24 to 48 h before mating), post-breeding infusion and to remove exudate and/or mucus before infusion of antibiotics (infuse, lavage uterus within 24 h, infuse intrauterine antibiotics after efflux has become clear, may take 2 to 4 days of uterine irrigation) a Experimental data or data from clinically controlled breeding trials to support these therapies are lacking or minimal. DMSO, dimethyl sulfoxide; EDTA, ethylene diamine tetra-acetic acid.
Uterine irrigation Uterine irrigation is most commonly performed postmating if mares have more than 2 cm of intrauterine fluid.37 Irrigation may be conducted as early as 4 hours post-mating without deleterious effect to pregnancy rates.40, 53 If fluid accumulations are less than 2 cm in depth or there is residual endometrial edema after ovulation, treatment with only oxytocin or cloprostenol is commonly performed. However, mares with small amounts of intrauterine fluid after breeding may be good candidates for uterine irrigation if they have a history of repeated breedings without achieving pregnancy or they have been difficult to manage in previous years without uterine irrigation post-breeding. Mares with uterine fluid before breeding may also be irrigated with lactated Ringer’s solution immediately before insemination without a subsequent decrease in pregnancy rates. Removal of the fluid is desirable because intrauterine fluid contains inflammatory debris, especially if the mare requires repeated inseminations with cooled or frozen semen, and the fluid adversely affects spermatozoal motility and fertility.54 The uterus is typically irrigated with 2–3 liters of sterile physiological saline or lactated Ringer’s solution using a large-bore catheter (30 Fr, 80 cm) passed into the uterus lumen only after the perineum is thoroughly washed and dried. Many veterinarians prefer lactated Ringer’s over saline as saline infusion can be irritating due to its low pH.55 A liter of flush solution is infused by gravity flow and the efflux inspected for cellularity and degree of clarity. Some veterinarians recommend repeating the uterine irrigation (> 3 L) until efflux recovered is clear.24 Oxytocin or cloprostenol should be administered immediately after the last uterine flush to assist in clearing all uterine fluid. Oxytocin In 1991, Allen suggested that oxytocin could be used to stimulate uterine contractions and promote uterine
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drainage in mares with defective uterine clearance. Prior to that time, oxytocin was thought only to induce uterine contractions in the first 48 hours after foaling and that it would induce colic when administered during the estrous cycle. Many additional studies have supported its efficacy in promoting uterine clearance and allayed fears that it causes colic.43, 44, 46, 56 Oxytocin induces high-amplitude uterine contractions for approximately 30 minutes during estrus and in the 48-hour period after ovulation.43, 57, 58 The half-life of oxytocin is 6.8 minutes.59 Most mares respond rapidly, with fluid being voided almost immediately. The recommended dose varies between 10 and 25 IU, administered either intravenously or intramuscularly. Mares given 25 IU in the pre-ovulatory period had a more pronounced change in intrauterine pressure at 5 and 10 minutes after treatment, but intrauterine pressure changes at subsequent time intervals were similar between a 25-IU and 10-IU dose.57 The effect of oxytocin on uterine contractility diminishes after ovulation, so the larger dose of 25 IU should be given. The uterine response to oxytocin appears to be inversely related to the concentration of progesterone.58 Oxytocin does not appear to be effective in all mares that accumulate intrauterine fluid. Factors that may affect response include an inadequate number of endometrial receptors, a pendulous uterus, an excessive dose resulting in inappropriate contractions, abnormal propagation of uterine contractions, or prolonged inflammation. Propagation of uterine contractions has been shown to differ between reproductively normal mares and mares with delayed uterine clearance.60 Intraluminal uterine pressure first increased in the uterine horn and then in the uterine body in reproductively normal mares after administration of detomidine or detomidine and oxytocin. This pattern was rarely seen in susceptible mares. Their response to drug administration was an inverted contraction pattern with the first increase in pressure near the cervix followed by contractions in the uterine horn. This inverted pattern could interfere with flow of luminal fluids into the vagina. If a clinician suspects that oxytocin does not stimulate appropriate drainage, it may be beneficial to administer cloprostenol to the mare. A long-acting synthetic oxytocin analog, carbetocin, has recently become available in Europe, Canada, and Mexico. It was well tolerated in a group of horses following intravenous administration of 175 µg. The half-life of carbetocin is about 17 min, or 2.5 times that of oxytocin. The drug may be of benefit in mares where more prolonged uterine contractions are needed.61
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Cloprostenol The effect of prostaglandins on uterine contractility has been investigated because oxytocin stimulates prostaglandin release, the uterine contracts in response to prostaglandin, and oxytocin has a relatively short half-life. Of the three prostaglandin (PG) analogs investigated – PGF2α , cloprostenol, and fenprostalene – cloprostenol produced the most consistent uterine response.41 All reproductively normal mares and mares susceptible to endometritis cleared a radiocolloid infused into the uterus after administration of cloprostenol (500 µg i.m.), but at a significantly slower rate than that seen with oxytocin (4 h vs 30 min, respectively). It has been recommended that cloprostenol be given to mares that require prolonged, low-amplitude uterine contractions, such as mares with lymphatic stasis, and to mares that do not appear to respond to oxytocin and to mares in estrus with cervical fibrosis associated with aging. A suggested treatment protocol for lymphatic stasis includes uterine irrigation with saline or lactated Ringer’s solution between 4 and 8 hours after mating followed immediately with administration of 10–25 IU of oxytocin intravenously. The mare is then given 250 µg of cloprostenol intramuscularly between 12 and 18 hours post-mating. The rationale for the protocol is that uterine irrigation removes inflammatory debris, oxytocin induces strong uterine contractions for approximately 30 minutes resulting in rapid clearance of intrauterine fluids, and PGF2α produces low-amplitude contractions that persist for 4 to 5 hours and assist in lymph flow.41, 42, 49, 50, 62 Lymphatic vessels do not contain smooth muscle and must rely on uterine contractions for emptying. Cloprostenol should be given with care as a few mares have developed projectile diarrhea after administration. Cloprostenol induces uterine contractions after ovulation; however, administration more than 12 hours after ovulation is associated with a decrease in plasma progesterone concentrations, and if given 2 days after ovulation decreased pregnancy rates have been reported. Therefore, use should be limited to estrus and within 12 hours of ovulation.62–64 Mares receiving phenylbutazone therapy for musculoskeletal problems exhibit diminished uterine clearance as the non-steroidal antiinflammatory drug has an inhibitory effect on prostaglandin.56, 65 In any mare being treated with phenylbutazone around the time of breeding, administration of oxytocin is preferable to cloprostenol as a method for improving uterine clearance Plasma Although intrauterine plasma has lost favor in clinical reproductive practice, previous work indicates
that pregnancy rates may be improved when plasma is infused into the uterus after breeding. Plasma contains complement and is thought to increase the efficiency of the mare’s cellular uterine defense mechanisms.1 Pregnancy rates were reported after a clinical breeding trial conducted with 905 Thoroughbred mares. Mares were treated with either no infusion (control), intrauterine antibiotics and plasma, or intrauterine antibiotics alone. Pregnancy rates were higher per cycle in lactating mares infused with plasma and antibiotics compared to controls or antibiotics alone (67%, 53%, and 57%).66 Barren mares had a slightly higher but not significant pregnancy rate than no treatment.
Post-breeding infusion of antibiotics Since the mid-1970s, antibiotics have been infused into the uterus after natural mating to prevent bacterial endometritis. Clinical studies and field experience have demonstrated the effectiveness of a single post-mating antibiotic treatment in combating endometritis.44, 46 Pregnancy rates were similar in mares treated with single doses of oxytocin or broadspectrum antibiotics but were highest in mares who received both, indicating that the two treatments may result in an additive benefit. Using an experimental model, others have shown that when mares susceptible to endometritis were treated with intrauterine saline lavage, PGF2α , or penicillin IU 12 hours after an intrauterine infusion of Streptococcus equi ssp. zooepidemicus, the treatments were equally effective in eliminating bacteria from the uterus.67 Post-breeding infusion of antibiotics may improve pregnancy rates in repeat breeders, in mares that accumulate fluid after breeding and have not been treated within 8 hours of mating, or in mares with a history of bacterial endometritis. The antibiotic chosen should not irritate the endometrium, should be diluted appropriately with the smallest volume recommended, and should be infused after the uterine is irrigated with saline or lactated Ringer’ solutions as exudate may interfere with absorption.
Exercise Exercise is helpful in establishing uterine drainage. Increased intra-abdominal pressure associated with movement helps to evacuate uterine contents. Broodmares restricted to a stall because of an ill neonate or injury and mares with laminitis or musculoskeletal pain, accumulate fluid in their uterus after breeding. If exercise must be limited, small injections of oxytocin (10 IU) given every 4 to 6 hours may assist in fluid clearance.8
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Breeding the Problem Mare Immunomodulators The inflammatory response associated with mating may be modulated with dexamethasone, prednisolone, or one of two commercially available immunomodulators.38, 52,68–71 A single injection of dexamethasone administered at breeding (50 mg i.v.) resulted in increased pregnancy rates in mares with a history of fluid accumulation after ovulation or cervical incompetence.38 Dexamethasone treatment was combined with post-breeding therapies including uterine irrigation, repeated injections of either oxytocin or cloprostenol, and in some cases mares received intrauterine antibiotics. Treatment was associated with a reduced inflammatory response postmating, which included decreased uterine edema, decreased intrauterine fluid, and an increase in uterine fluid clarity. Increased pregnancy rates were also seen in mares with three or more risk factors for susceptibility to endometritis. Risk factors evaluated included abnormal reproductive history, positive endometrial culture, ≥ 2 cm of intrauterine fluid before or after breeding, intrauterine fluid that persisted for more than 36 hours after breeding, abnormal perineal conformation, a cervix that did not dilate during estrus, and a vulvoplasty that was not repaired after foaling. In a second study, dexamethasone (10 or 20 mg i.m.) was given 6–12 hours after insemination in client-owned mares (783 cycles) that had a history of intrauterine fluid retention.72 Mares were allowed to carry their pregnancy (n = 429) or were embryo donors (n = 354). Treatment with 10 or 20 mg of dexamethasone after breeding did not improve pregnancy rates or embryo recovery rates. Factors that likely contributed to differences between the two studies included timing of administration, dose, and the number and type of risk factors. Risk factors were not evaluated in the Vandaele et al. study.72 A plausible cause for the different results is that steroids block both the cyclooxygenase and 5-lipoxygenase pathways of inflammation. The 5-lipoxygenase pathway includes leukotriene B, a potent neutrophil chemotactic factor found in uterine fluids of susceptible mares after mating.72, 73 Reducing neutrophil chemotaxis and the number of neutrophils recruited into the uterus post-mating may diminish the severity and length of the inflammatory response. Candidates for steroid use should be chosen carefully as misuse in mares with bacterial endometritis may exacerbate the infection. Oral administration of prednisolone (0.1 mg/kg) given every 12 hours for 4 days beginning 48 hours before breeding has also been evaluated in barren mares that accumulated intrauterine fluid before and after mating. Prednisolone administration resulted in decreased neutrophil numbers in uterine fluids of
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barren mares, decreased uterine fluid, increased fluid clarity, and increased pregnancy rates.52 Immunostimulants induce a non-specific cellmediated response predominantly by activation of macrophages and release of cytokines that elicit a general increase in immune system activity.73 Two immunostimulants are currently labeled and marketed for use in horses. The first is a cell-wall exR tract of Mycobacterium phlei (Settle ; Bioniche Animal Health, Bogard, GA), which has been approved as an adjunctive treatment in mares with uterine infection caused by Streptococcus equi. The second is a susR ; Neogen pension of Propionibacterium acnes (EqStim Corp, Lexington, KY), which has been used as an adjunct treatment for horses with equine respiratory disease complex. The effect of Propionibacterium acnes on live foal rates was recently evaluated in a controlled field study.71 Pregnancy and live foal rates were improved after P. acnes was used as an adjunct to conventional treatments in mares with a cytological diagnosis of persistent endometritis. The optimal effect was detected in mares bred during the interval extending from 2 days before to 8 days after first treatment with P. acnes. The effects of Mycobacterium phlei cell-wall extract R , Bioniche) in modulating the immune re(Settle sponse of mares susceptible to endometritis have been compared with those occurring in reproductively normal mares.68–70 These studies reported that: (i) mares susceptible to endometritis have higher levels of proinflammatory cytokines and lower levels of interkeukin-10 (IL-10), a cytokine that inhibits production of proinflammatory cytokines, in their endometrium before breeding than reproductively normal mares; (ii) after insemination, susceptible mares expressed significantly higher levels of proinflammatory cytokines in their endometrium 7 days after ovulation than normal mares; and (iii) susceptible mares treated with Mycobacterium phlei at the time of artificial insemination had an endometrial immune environment similar to that found in reproductively normal mares. These studies indicate that immunomodulation may help to restore homeostatic local inflammatory mechanisms, thus assisting in the prophylaxis of post-mating-induced endometritis or chronic infectious endometritis.
Identification and breeding management of mares with chronic bacterial or fungal endometritis Chronic bacterial endometritis Chronic infectious endometritis is most commonly seen in older (> 12 years of age), pluriparous mares
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Table 272.3 Guidelines for administration of intrauterine antibiotics Drug
Dose per infusion
Comments
Amikacin
2 ga
Ampicillin Ceftiofur sodium Gentamicin
2 gb 1g 1–2 ga
Enrofloxacin Penicillin (potassium) Penicillin (suspension) Neomycin Ticarcillin
250 mg 5 million units 4–5 g 4g 1–6 g
Ticarcillin–clavulanic acid
3–6 g
Buffer with bicarbonate or large volume of saline (200 mL); excellent Gram-negative coverage Use only the soluble product; susceptible Gram-positive and E. coli Resistant to many beta-lactamases; broad spectrum; save for resistant organisms Buffer with bicarbonate or large volume of saline (200 mL); effective against some Streptococcus equi ssp. zooepidemicus; Enterobacter spp., E. coli, Klebsiella spp., Proteus spp., Serratia spp, P. aeruginosa, S. aureus Dilute to 60 mL; can result in adhesions and severe inflammation if more concentrated Strep equi ssp zooepidemicus qs to 60 mL Gram-negative organisms; some E. coli and some Klebsiella spp. Gram-positive organisms; effective against some Pseudomonas spp.; infuse with a minimum of 200 mL saline Beta-lactamase inhibitor confers greater activity against Enterobacter; S aureus, B. fragilis; infuse with a minimum of 200 mL saline
a
Buffered with equal volume of 7.5% bicarbonate and diluted in saline. Use at high dilutions because it can be irritating. Adapted from LeBlanc.109
b
with defects to their perineal conformation. Such mares have a breakdown in anatomical barriers and degenerative changes to the uterus that allow normal genital flora to contaminate the uterus, multiply, and develop into persistent endometritis. Chronic endometritis may also occur as a consequence of cervical fibrosis, not treating susceptible mares postbreeding, and repeated manipulation of embryo donor mares. Clinical signs vary considerably. Mares with chronic uterine infections may exhibit normal estrous cycles, shortened or lengthened cycles, or anestrus. They may or may not have intrauterine fluid, abnormal edema patterns, or hyperechoic lines within the uterine lumen. The vaginal mucosa may be reddened, splotchy in color, wet or dry. The goal of treatment is quickly and completely to clear offending organisms, stop mucus or exudate production, remove uterine fluid, and restore endometrial health before breeding. This may be accomplished with a combination of procedures including surgical correction of anatomical defects, mechanical clearance of uterine contents with irrigation and judicious use of ecbolics, and bacterial killing with an appropriate antimicrobial therapy. Intrauterine antimicrobial therapy should be preceded by uterine irrigation with sterile saline or lactated Ringer’s solution to remove exudate, mucus, or biofilm as some antibiotics are chemically altered when the pH is excessively high or low or if there is gross debris. Mucolytic agents may be needed to break up exudate to enable antibiotic penetration of bacteria. Intrauterine infusion of an appropriate antibiotic for 3–7 days during estrus is the most common treatment protocol for mares with chronic en-
dometritis. Intrauterine antibiotic treatment should be avoided during diestrus as treatment during the progesterone phase has resulted in resistant bacterial and fungal infections.74, 75 Severely infected mares may require treatment during a second or third estrus. Treatment length depends on chronicity of the infection, type of bacteria isolated, the mare’s ability to clear uterine fluid, and her reproductive history. A reproductive examination should be performed after the last intrauterine treatment to determine if there is residual intrauterine fluid and excessive uterine edema, and if the mare has ovulated. If fluid or edema persists, treatment may need to be continued or the mare brought back into estrus before continuing treatment if she has already ovulated. A smallvolume flush or biopsy tissue swab should be taken to determine if the organism(s) persist and if exudate is still present at the estrous following treatment. If bacteria are isolated and there is exudate, systemic therapy in combination with local uterine irrigation should be considered. Tables 272.3 and 272.4 include dosages of intrauterine and systemic antibiotics for treatment of bacterial endometritis.76 Treatment failure is common when there is a fungal or chronic Gram-negative bacterial infection and may be due to continual uterine contamination because of loss of anatomical barriers, inappropriate dosage regimen, drug resistance, abnormal microenvironment (biofilms, mucus, exudate, pH change), or impaired host defense mechanisms. When response to treatment is not as expected, the mare should be reevaluated in an attempt to determine cause. Factors to be considered in the selection of antimicrobials for treating reproductive conditions
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Table 272.4 Antibiotics for systemic treatment of bacterial endometritis Drug
Dosage
Route
Interval
Comments
Amikacin sulfate Ampicillin sodium Ceftiofur
10 mg/kg 29 mg/kg 2.5 mg/kg
i.v. or i.m. i.v. or i.m. i.m.
24 h 12–24 h 12–24 h
Gentamicin
6.6 mg/kg
i.v.
24 h
Enrofloxacina
5.5 mg/kg 7.5 mg/kg
i.v. p.o.
24 h
Penicillin G (potassium)
25 000 IU/kg
i.v.
6h
Penicillin (procaine) Trimethoprim–sulfonamide
25 000 IU/kg 30 mg/kg (combined)
i.m. p.o.
12 h 12 h
Excellent Gram-negative coverage Susceptible Gram-positive organisms and E. coli Broad-spectrum Gram-positive and some Gram-negative organisms Slow intravenous infusion; Enterobacter spp., E. coli, Klebsiella spp., Proteus spp., Serratia spp., P. aeruginosa, S. aureus Slow intravenous infusion; Gram-negative infections caused by susceptible bacteria resistant to alternative, first-choice drugs Synergistic with aminoglycosides; do not store mixed in syringe for more than 12 h; do not mix in syringe with gentamicin; Strep equi ssp. zooepidemicus, leptospirosis As for above S. aureus, E. coli, Klebsiella spp., Proteus; some Nocardia spp.
a Should not be used in pregnant mares or in young growing horses because of the risk of arthropathy. Adapted from LeBlanc.109
include susceptibility of the microorganism, local versus systemic treatment, concentration of drug attainable at site of infection, and the effect of the drug on immediate and future fertility.76, 77 Antimicrobial selection should be based on culture and sensitivity results. Unfortunately, in vitro and in vivo efficacy may not always be equivalent. For example, most uterine cultures of Pseudomonas spp. show in vitro sensitivity to polymyxin B, yet few mares with endometritis caused by Pseudomonas spp. respond well to polymyxin B therapy. Streptococcus equi ssp. zooepidemicus shows in vitro sensitivity to trimethoprim–sulfonamide combinations but rarely does the drug effect a clinical cure. Drug compatibility is an important principle of therapy. The polypharmacy approach of multiple antibiotics and antiseptics mixed together and infused into the uterus may be of no therapeutic value and could be harmful. Penicillin, gentamicin, ampicillan, or ticarcillin should not be commixed as physical and chemical incompatibilities exist.78 Penicillin and ampicillan are inactivated when the pH of a solution is less than 5.5 or greater than 8, so they should not be mixed with sodium bicarbonate, gentamicin, or sulfonamides. Gentamicin added to a beta-lactam drug can result in a precipitate so they should never be mixed in the same syringe. Complications may arise with intrauterine antibiotic treatment. Mares may develop secondary bacterial or fungal infections or exhibit severe endometrial irritation. Treatment for one pathogen may result in the proliferation of another that often is more difficult to manage than the original. An example is treat-
ment of a streptococcal infection that, after treatment with the appropriate antibiotic, is followed by development of a yeast or Pseudomonas infection. In these cases, a mixed infection may have existed initially and antibiotic use merely allowed proliferation of the other organism. In other cases, a second organism may be introduced accidentally during the course of treatment for the primary infection. Systemic antibiotics should be considered in chronic infectious endometritis and in mares with defects in the caudal reproductive tract. Systemic administration of antibiotics results in a higher minimal inhibitory concentration (MIC) throughout the genital tract as compared to intrauterine therapy. In addition, there is less likelihood of changes in vaginal flora, the antibiotics are not degraded by conditions in the uterine lumen, and parental therapy does not irritate the endometrium. The length of systemic treatment is not dictated by the estrous cycle and antibiotics may be given for 7 to 10 days if needed. Systemic therapy eliminates the need to invade the vestibule, vaginal canal, and cervix. The vestibule and clitoral fossa harbor a vast array of bacteria. These organisms may be passed iatrogenically into the uterus during intrauterine infusion.79 A complication of systemic antibiotics is a change in gut flora leading to gastrointestinal problems.
Intrauterine antiseptics and chelating agents A variety of disinfectants and irritants have been infused into the uterus of mares in attempts to
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treat uterine infection, including vinegar, betadine, iodine, dimethyl sulfoxide (DMSO), kerosene, chlorhexidine-gluconate, magnesium sulfate, hydrogen peroxide, hypertonic saline, and N-acetylcysteine (Table 272.2).40,80–83 These preparations should be used with care as they may cause inflammation, necrosis, and reproductive sterility. Some antiseptics can be administered safely if given at appropriate concentrations. Dilute povidone-iodine solution (0.1–0.2% v/v povidone-iodine solution in 1 L of saline 4 h post-mating) had no adverse effects on pregnancy rates in reproductively sound mares when compared with saline and did not cause inflammation on endometrial biopsy. Because antimicrobial activity is maintained to concentrations as low as 0.01%, this lavage at 0.05% may have a role in management of some bacterial uterine infections.40 Exudate and changes in the quantity or viscosity of the mucoid layer overlying the endometrium may interfere with antibiotic penetration and killing of bacteria or yeast. Mares with chronic endometritis have a thicker mucoid layer than reproductively normal mares, regardless of stage of cycle.18 Solvents and mucolytic agents have been added to uterine irrigation fluids in an attempt to clear exudate, mucus, or biofilm. Agents used include DMSO, hydrogen peroxide, kerosene, magnesium sulfate, and Nacetylcysteine (NAC). Intrauterine infusion of a 30% concentration of DMSO in 16 barren mares resulted in a trend toward higher pregnancy rates as compared to mares irrigated with saline.83 In addition, intrauterine DMSO therapy resulted in a significant improvement in endometrial biopsy classification in 18 of 27 mares; whereas only 2 of 18 barren mares improved following intrauterine saline treatment. Hydrogen peroxide has been used as an irrigation solution at a 1% solution (20 mL of a 3% solution of hydrogen peroxide diluted in 60 mL of LRS because of its success in treating difficult recurrent bacterial vaginosis in women.84 It has been used in mares as an intrauterine infusion.85 The use of kerosene as an intrauterine irritant in chronically infertile mares was first advocated by Dr Charlie Roberts in New Zealand. It has continued to be used throughout the breeding industry as a last resort. Its success may be associated with necrosis of luminal epithelium and therefore loss of mucus-producing cells and exudate that is common in chronically infected mares. Intrauterine infusion of 50 mL of commercially available kerosene in 26 mares with varying degrees of endometrial pathology induced diffuse moderate to severe endometritis, severe diffuse edema, and production of a serum-like exudate.86 Half of the mares exhibited mild to severe necrosis of luminal epithelium. Mares were subsequently bred on the next cycle and surprisingly, 50% of the mares with category
II or III biopsy scores carried foals until term. The effect of magnesium sulfate solution on the equine endometrium has been studied in seven treatment mares and three control mares. Treatment mares were infused with 128 g of magnesium sulfate diluted in 1 L of saline (0.9% sodium chloride) while control mares were infused with 1 L of saline. No harmful effects were noted when mares were evaluated on days 1, 7, and 21 after infusion.81 Recently in central Kentucky, intrauterine infusion of N-acetylcysteine (30 mL of NAC in 150 mL of saline) has been investigated because of its mucolytic and anti-inflammatory properties. NAC disrupts disulfide bonds between mucin polymers, thereby reducing the viscosity of mucus. In addition, NAC possesses antioxidant and possibly some antimicrobial properties. In an uncontrolled clinical trial, NAC was infused into the uterus of 23 barren mares the day before breeding. Mares also received traditional post-breeding therapies such as uterine irrigation and infusion of intrauterine antibiotics, therapies that had been performed during previous breeding cycles. Twenty of the 23 mares became pregnant on that mating, a rate that was higher than expected. The effect of NAC on endometrial histological features has been evaluated in reproductively normal mares and mares with chronic inflammatory changes, and compared to that of infusion of saline. Data indicated that NAC was not harmful to the endometrium and that it may counteract the irritating effect of saline, as reflected through increased cell height in control mares.82 Addition of any mucolytic drug or disinfectant should be used with caution as adverse reactions can occur. Some chronic Gram-negative bacterial or fungal infections are difficult to resolve even after repeated treatment. Treatment failure may be due to anatomical defects, bacterial resistance conferred through biofilm production, or breakdown of intrauterine antibiotics by exudate or pH changes of uterine fluids. Biofilms consist of communities of microorganisms attached to a surface that can express an increased resistance to antimicrobial agents through a number of mechanisms.87–90 Treatments for removing biofilm vary widely in human medicine and include incorporation of enzymes and solvents in protocols. An agent that has shown efficacy against biofilm production in human medicine, and also in chronic otitis externa, chronic dermatitis, and recurrent cystitis in dogs and fungal keratitis and chronic Pseudomonas endometritis in horses is EDTA-Tris (ethylene diamine tetraacetic acid-tromethamine).91–93 Pseudomonas isolates collected from mares with endometritis that were exposed to EDTA-tris solution exhibited decreased viability.93 The treatment appears to be a safe adjunct therapy for equine genital infections because infusion of 250 mL of 3.5 mM EDTA, 0.05 M Tris, pH 8 into the
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Breeding the Problem Mare uterus induced an inflammatory response that was no greater than saline. We have added EDTA-Tris (120 mL) to amikacin (1.5 g) for the treatment of chronic persistent endometritis cases caused by Pseudomonas spp., Enterobacter spp., Klebsiella, or E. coli. Pregnancies have resulted in mares after treatment. Fungal infections Treatment of fungal endometritis is frustrating because treatment is costly, success is only fair, and relapses are common. Affected mares tend to be old and pluriparous with poor perineal conformation, or maiden mares with cervical incompetence. Mares may have been treated repeatedly with intrauterine antibiotics, have normal or abnormal estrous cycles, may be presented with a history of anovulatory follicles, or have endocrine dysfunction such as equine pituitary disorders or insulin resistance. Prolonged progesterone therapy may also predispose mares to fungal endometritis as cervical drainage is decreased and uterine muscular activity and neutrophil function are altered. Diagnosis is based on the presence of fungal elements and inflammatory cells in endometrial smears. Yeast appear as small, round, single-cell, brown to black spores on cytological smears obtained from infected mares, whereas molds have long, filamentous hyphae. Candida spp. and Aspergillus spp. are most commonly isolated from the equine uterus.94 Candida spp. is a normal commensal of the gastrointestinal tract and vagina. Endometrial smears obtained from mares with Candida endometritis usually contain spores but hyphae may be seen in chronic infections. Aspergillus is a mold that produces hyphae. Other fungi isolated from the uterus include: Actinomyces spp., Fusarium spp. (filamentous fungi), Mucor (filamentous fungi), Paecilomyces spp., Rhizopus (filamentous fungi), Rhodotorula spp., and Trichosporon spp. Fungal characteristics affect treatment regimens and success. Yeasts tend to proliferate in fluid and may be treated successfully with uterine irrigation and intrauterine infusion of antifungal agents, whereas other fungi tend to penetrate tissue and are deep-seeded, requiring both systemic and local therapy.94, 95 Fungal endometritis is frequently a consequence of repeated intrauterine antibiotic therapy and may resolve spontaneously if the mare has normal perineal anatomy and adequate uterine clearance. If the infection does not resolve in one to two estrous cycles, the uterus should be irrigated with a disinfectant for 5–7 days and the clinical isolate submitted to a microbiology laboratory for an antimycotic sensitivity pattern. Disinfectant solutions used for fungal infections include 1–3% (v/v) hydrogen
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peroxide solution, 2% (v/v) acetic acid (white vinegar: 20 mL of vinegar in 1 L of 0.9% saline), 0.1–0.2% (v/v) povidone-iodine solution, or 20% DMSO.85, 94, 96 Both local and systemic antimycotic therapy have been advocated in recalcitrant infections, and in some cases mares are treated by both routes. Drug dosages are not well defined in the horse and minimum inhibitory concentrations for these drugs have not been established for equine uterine infections. Most of the data are extrapolated from humans, dogs, cats, or animal models. In human medicine a minimum of 10 to 14 days of treatment is recommended for fungal or yeast vaginitis. In order to resolve fungal endometritis, mares may require two to three treatment sessions of 5 to 7 days duration conducted during consecutive estrus periods. Treatment interval can be shortened by administration of prostaglandin. Breaches in anatomical barriers should be corrected surgically before treatment is begun. The uterus should be recultured after the second treatment session to determine if the infection has cleared. If fungi are isolated, the reproductive tract should be re-evaluated for anatomical defects, cervical incompetence, and/or testing for Cushing’s disease or insulin resistance. Systemic therapy for a minimum of 21 days should be considered. Table 272.5 contains antifungal drugs and dosages for systemic and local infusion.
Management of mares not exhibiting normal estrous cycles Abnormal estrous cycle length or lack of estrous cycles during the cyclic season may be associated with uterine infection, aging, poor body condition score (< 4 out of 9), pain, ovarian tumors, anovulatory follicles, systemic conditions such as Cushing’s disease or insulin resistance, chromosomal abnormalities, and ovarian senility. Management of mares that are not exhibiting estrous cycles during the transitional phase of the season will be discussed in Chapter 272. Conditions such as Cushing’s disease or laminitis can adversely affect hormonal signaling, and may result in no or infrequent estrous cycles, uterine fluid accumulation, endometritis or formation of anovulatory follicles. Advanced age (mares > 18 years of age) has been associated with a shortened cyclic season, a prolonged follicular phase, and ovulation of small follicles (25 mm or less in diameter). Old mares may require additional care, special diets, blanketing in cold weather, or treatment of arthritic conditions in order to advance the first ovulation of the year. Ambient temperatures consistently below freezing in the spring may not only affect the timing of first ovulation of the year, but may also interfere with cyclic patterns in mares that have begun to exhibit estrous cycles, especially if the mares are thin, are shipped
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Table 272.5 Systemic and topical antifungal agents used for fungal endometritis Drug
Dosage
Route
Interval (h)
Spectrum
Systemic Amphotericin B Ketoconazole Fluconazole Itraconazole
0.3–0.9 mg/kg 20 mg/kg (in 0.2 N HCl) Loading dose 14 mg/kg 5 mg/kg 5 mg/kg
i.v.a NGTb p.o., i.v. p.o.c , i.v
24–48 12 24 12–24
Broad spectrum Yeastd Yeast Broad spectrume
Topical Clotrimazole Miconazole Nystatin Amphotericin B Fluconazole
400–700 mg 500–700 mg 0.5–2.5 million unitsf 100–200 mg 100 mg
IU IU IU IU IU
24 h × 7 d 24 h × 7 d 24 h × 7 d 24 h × 7 d 24 h × 7 d
Broad spectrum Broad spectrum Yeast Broad spectrum Yeast
a
Diluted to 1 mg/mL in 5% dextrose and administered over 1–2 h. Nasogastric intubation is required to avoid the irritant effect of HCl on the oral cavity and throat. c The bioavailability of the oral suspension is superior to that of the capsules. d Yeasts: Candida spp. e Broad spectrum: yeasts, Aspergillus, dimorphic fungi. f Must be diluted in sterile water (100–200 mL) as it precipitates in saline. IU, intrauterine; NGT, nasogastric tube. Adapted from LeBlanc.109 b
from a warm to cold climate, or don’t have a winter hair coat. Diagnostics for determining why a mare may not be cycling normally should include a physical examination and a reproductive examination. Additional testing including measurement of endocrine profiles, uterine cytology, uterine culture, endoscopic evaluation of the uterus or cytogenetic evaluation is determined on a case by case basis.
Anovulatory follicles Anovulatory follicles or hemorrhagic anovulatory follicles are a frustrating but common cause for abnormal estrous cycles. It is a condition where the pre-ovulatory follicle grows to an unusually large size, fails to rupture and ovulate, typically fills with blood and fibrin, and then gradually regresses. They occur most commonly in mares > 10 years of age97, 98 and during the latter part of the breeding season. Possible causes include aging, inability of the follicle to respond to luteinizing hormone (LH), persistent inflammation, infection, administration of prostaglandin on Day 10 of diestrus, or repeated use of prostaglandin.99 Anovulatory follicles have also been associated with laminitis and insulin resistance, and high plasma cortisol and/or insulin concentrations associated with pain or obesity. Mares that develop an anovulatory follicle exhibit normal estrous behavior but estrus is prolonged and the dominant follicle continues to grow even after it is expected to ovulate. On ultrasonographic evaluation, anovulatory follicles are usually greater than 45 mm in
diameter, and have hyperechoic particles within a fluid-filled lumen, fibrous strains within the lumen, and/or a thick white rim along the perimeter of the follicle (Figure 272.8). Treatment with either human chorionic gonadotropin (hCG) or deslorelin – a gonadotropin-releasing hormone (GnRH) agonist – is futile as anovulatory follicles do not respond to ovulatory agents. About 65% of anovulatory follicles will eventually produce progesterone while the remaining 35% do not.97 Anovulatory follicles that produce progesterone respond to exogenous prostaglandin, and mares given prostaglandin will return to estrus. Unfortunately, many will form another anovulatory follicle during the next estrus. Infection or inflammation Uterine infection should always be included on a list of differentials in mares that are not exhibiting normal estrous cycles or are anestrus during the cyclic season (see ‘Chronic bacterial endometritis’ above). Metabolic problems Insulin resistance or pituitary tumors may result in abnormal estrous cycles, anestrus, development of anovulatory follicles, and/or endometritis. Mares with chronic foot pain, obese mares, mares exhibiting hirsutism, polydipsia, polyuria, or mares with repeated bacterial or yeast infections should be tested for insulin resistance and pituitary tumors. Diagnostic tests include dexamethasone suppression, and measurement of plasma adrenocorticotropic
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Figure 272.8 (a,b) Ultrasonographic images of anovulatory follicles.
hormone (ACTH), cortisol, and insulin. Mares treated for Cushing’s disease or insulin resistance with pergolide in doses exceeding 3 mg daily may develop anestrus due to downregulation of GnRH from pergolide, a dopamine agonist. Lactational anestrus Lactational anestrus is observed most commonly in mares that foal before 1 April in the northern hemisphere or 1 October in the southern hemisphere and are not provided with additional daylight during late gestation. Other factors that may cause lactational anestrus include inadequate nutrition, low body condition score, chronic pain, stress, subclinical endometritis, and metritis. Affected mares typically exhibit a normal foal heat, ovulate a follicle, and then develop small, hard ovaries with follicles < 15–20 mm in diameter. Endometritis should be ruled out as it is a frequent cause. Appropriate uterine treatments should be instituted (see ‘Chronic bacterial endometritis’ above) if cytology or culture results indicate inflammation or infection. As lactational anestrus can be secondary to pain, mares that experienced a difficult foaling or delivered a large foal should have a musculoskeletal examination. Painful mares should be given non-steroidal antiinflammatory drugs and evaluated for endometritis as limited mobility can result in uterine fluid retention. Chiropractic manipulations or acupuncture may be helpful. If the primary cause is endometritis, appropriate uterine and systemic treatment frequently results in follicular development and ovulation occurring within 7 to 10 days. A number of drugs have been used to induce cyclicity in mares with lactational anestrus. These include GnRH implants, progestins alone or in combination with estradiol for 10 to 14 days, and dopamine antago-
nists such as sulpiride, domperidone, or reserpine.100 Treatment protocols for dopamine antagonists include subjecting mares to 14.5 hours of light for 2 weeks before giving the drug, continuing the increased photoperiod while adding a dopamine antagonist such as sulpiride (1 mg/kg i.m. b.i.d.), domperidone (1.1 mg/kg p.o. s.i.d.), or reserpine (2–4 mg p.o. s.i.d.) daily for 2 to 3 weeks.101 If there is no follicular response after 18 to 20 days of treatment, it is unlikely that the mare will respond. Early foaling mares that have experienced lactational anestrus previously should be subjected to 14.5 hours of light starting on or before 15 December (northern hemisphere) and 15 June (southern hemisphere). Retention of endometrial cups Although an uncommon problem, retained endometrial cups have been reported in mares that had aborted or foaled in a previous year.102 Affected mares have ovaries that contain many luteinized follicles, clusters of large (> 35 mm) follicles, and corpora lutea. Mares with retained cups may exhibit no estrus, or irregular or prolonged estrus behavior. A diagnosis of retained endometrial cups can be made by ultrasonography or by measuring equine chorionic gonadotropin. Retained endometrial cups will eventually slough, although it may take months. Endometrial cups have been removed by laser surgery; however, complications including endometritis, uterine hemorrhage, and an inability to remove all cups at one session because of limited visibility from smoke, have been reported. Ovarian tumors Granulosa cell thecal tumors, although rare, are the most common equine tumor. Granulosa cell thecal
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tumors arise from sex cord stromal tissue within the ovary and may be hormonally active, producing variable amounts of steroids that cause behavioral changes and alternation in normal cyclic activity. Mares of any age and reproductive status may be affected and can exhibit nymphomania, anestrus, constant estrus, or aggressiveness. Tentative diagnosis is made by rectal examination and ultrasonography, and confirmed by measurement of plasma inhibin, progesterone, and testosterone levels. Classical findings include a small, walnut-sized ovary while the affected ovary is large with multiple cystic areas that have a honeycomb appearance on ultrasonography. The ovulation fossa of the affected ovary is not palpable. Inhibin concentration is > 0.7 ng/mL in approximately 90% of cases, progesterone is < 1 ng/mL, and testosterone is above normal range in approximately 50% of the cases.103, 104 Not all affected ovaries have the classical appearance, so measurement of inhibin is recommended. Treatment is by surgical removal. Mares will return to estrus usually within 9 to 12 months, but it may take up to 24 months in some long-standing cases. Uncommonly, some mares will continue to exhibit normal estrous cycles and ovulate off the unaffected ovary.105 Bilateral occurrence is extremely rare.
Mare loses pregnancy after day 14 and before day 45 of gestation: early embryonic death Pregnancy loss before 35 days may be associated with chromosomal anomalies of embryos, maternal aging, endometritis, uterine degeneration, progesterone deficiency, systemic disease, foal-heat breeding, timing of insemination relative to ovulation, and sire effects. Manipulation for assisted reproductive techniques is also associated with an increase in early embryonic death. The equine embryo is dependent on the yolk sac for nourishment until approximately 25 days of gestation. After that time, nourishment is attained primarily through endometrial secretions until placental attachment between 90 and 120 days. If inflammation persists in the first weeks of gestation, it may result in death of the embryo. Some affected mares will have embryos that are small for gestational age (between 12 and 28 days) or embryos with an irregular heartbeat at the 28-day pregnancy exam. Many of these embryos will be lost by 45 days of gestation. Treatment of mares that experience early embryonic death due to chronic or acute prolonged inflammation consists of intrauterine, and in some cases systemic, antibiotic therapy for 2 to 4 days after breeding in addition to uterine irrigation and anti-inflammatory therapy such as dexamethasone or prednisolone as discussed above. Immunomodula-
tion before breeding and again 3 and 7 days later and then again at days 14, 30, and 45 of gestation has also been recommended. Aging has clearly been shown to increase early embryonic death rates. Fertilization rates of old mares and young mares are about 85% to 90%. However, embryo recovery rates for aged versus young mares are significantly different (40% vs more than 70%, respectively), which places a high rate of embryonic loss during the first weeks of gestation in aged mares.106 Failure of the endometrial glands Kenney in 197832 first suggested that advanced periglandular fibrosis is highly correlated with pregnancy loss between 35 and 80 days. It is hypothesized that glandular secretion is abnormal resulting in poor fetal nutrition, abnormal placental attachment, and subsequent pregnancy loss. A presumptive diagnosis of periglandular fibrosis is made from a repeated history of early embryonic death between 35 and 80 days. Definitive diagnosis is made by histological examination of the endometrium. Affected endometrial samples contain widespread moderate and/or severe periglandular fibrosis. There is no known treatment, although many mares with periglandular fibrosis are placed on supplemental progesterone therapy, and anecdotal reports indicate that they carry their foal until term. In breeds other than Thoroughbred race mares, embryo transfer is a viable option. Progesterone insufficiency Prior to 80 days of gestation, if corpora lutea fail to produce progesterone, or if the ovaries are removed, mares will abort.107 The effect of rapid decline in progesterone levels on the tubular tract is loss of uterine and cervical tone, and loss of the conceptus. Therefore, it has been hypothesized that partial failure of the corpus luteum (CL) could reduce the absolute amount of progesterone needed to maintain pregnancy. As a result, many schemes of administering exogenous progestins have been used to prevent early pregnancy loss. The value of these schemes is debatable, as no critical evidence supports the concept. Nevertheless, we are often obligated to supplement progesterone due to client pressure and convincing histories. Progesterone supplementation in mares with prior histories of endometritis or lymphatic lacunae may be contraindicated as progesterone may aggravate the problem. Supplemental progestins are warranted in pregnant mares experiencing endotoxemia or laminitis as prostaglandins are released during both disease
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Breeding the Problem Mare processes. Prostaglandin lyses the CL, and if the mare is less than 90 days pregnant, she will abort.108 After 90 days, the placenta takes over and produces progestins that maintain pregnancy for the remainder of gestation. Progesterone therapy may also be beneficial in pregnant mares in late gestation that have placentitis, colic, or severe systemic disease as it may inhibit premature uterine contractions. Historical data are used to make the determination of progesterone supplementation. Reasons for placing mares less than 30 days pregnant on progestins include: (i) history of early pregnancy loss; (ii) low plasma progesterone concentrations on days 5, 8, and 12 of gestation (what is considered low varies and ranges from 1 to 6 ng/mL); (iii) visualization of uterine edema at pregnancy exams conducted between days 14 and 28; and (iv) colic, laminitis, placentitis, fetal compromise, and vaginal discharge in late gestation. Treatment of mares experiencing endotoxemia in early pregnancy is by administration of progestins (altrenogest 0.044 mg/kg p.o. s.i.d.) to prevent pregnancy loss. Mares may also be given 150–200 mg of progesterone in oil daily or a long-acting progesterone (Bet Laboratory, Lexington, KY) once weekly to maintain progesterone concentrations. Mares can be weaned off the progestin after 90 days of gestation as the fetoplacental unit begins to produce a number of progestins. If mares in late gestation are to be supplemented with progestins, the dose of altrenogest should be doubled (0.088 mg/kg p.o. s.i.d.) as the recommended dose for suppression of estrus behavior did not prevent abortion in an experimental model.108
Conclusions Diagnosing the cause of subfertility can be frustrating without a systematic, thorough approach and knowledge about the types of problems that affect the breed, age, or status of the mare being examined. Advances in diagnostics and our understanding of reproductive pathology and physiology have led to new therapies and the reintroduction of some old ones. Diagnosing the problem should always precede treatment as repeated use of antibiotics or dexamethasone can result in infections that are more difficult to treat, or in anovulatory follicles that form secondary to chronic inflammation. If treatment results are not as anticipated, one should always re-examine the mare for anatomical defects, or lack of synchrony between the ovary, uterus, and cervix to determine if a subtle defect has been missed. If no defects are identified, one should obtain information on the stallion’s recent fertility or request a change in stallions. Working with problem mares can be an enriching experi-
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ence for some veterinarians and is definitely costly for mare owners. Experimental models need to be developed to study various causes of susceptibility and how the uterus responds to different organisms. Only then can more appropriate therapies be developed.
Acknowledgement Parts of this chapter also appear in Ref. 109: LeBlanc MM. The chronically infertile mare. Proc Am Assoc Equine Pract 2008;54:391–407.
References 1. Asbury AC, Gorman NJ, Foster GW. Uterine defense mechanisms in the mare: Serum opsonins affecting phagocytosis of Streptococcus zooepidemicus by equine neutrophils. Theriogenology 1984;27:375–85. 2. Evans MJ, Hamer JM, Gason LM, Graham CS, Asbury AC, Irvine CHG. Clearance of bacteria and non-antigenic markers following intra-uterine inoculation into maiden mares: Effect of steroid hormone environment. Theriogenology 1986;26:37–50. 3. Hughes JP, Loy RG. Investigations on the effect of intrauterine inoculations of Streptococcus zooepidemicus in the mare. Proc Am Assoc Equine Pract 1969;15:289–92. 4. Kotlainen T, Huhtinen M, Katila T. Sperm induced leukocytosis in the equine uterus. Theriogenology 1994;41:629–36. 5. Katila T. Onset and duration of uterine inflammatory response of mares after insemination with fresh semen. Biol Reprod 1995 Mono 1: Equine Reprod 6:515–17. 6. Allen WE, Pycock JF. Cyclical accumulation of uterine fluid in mares with lowered resistance to endometritis. Vet Rec 1988;122:489–90. 7. LeBlanc MM. Persistent mating induced endometritis in the mare: pathogenesis, diagnosis and treatment. In: Ball BA (ed.) Recent Advances in Equine Reproduction. Ithaca, NY: International Veterinary Information Service, 2003. 8. Causey RC. Making sense of equine uterine infections: The many faces of physical clearance. Vet J 2006;172:405–21. 9. Samper JC. Uterine edema in the mare. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. St Louis: Saunders Elsevier, 2009; pp. 133–8. 10. Evans MJ, Hamer JM, Gason LM, Irvine CHG. Factors affecting uterine clearance of inoculated materials in mares. J Reprod Fert 1987; Suppl. 35:327–34. 11. Troedsson MHT, Liu IKM, Ing M, Pascoe J, Thurmond M. Multiple site electromyography recordings of uterine activity following an intrauterine bacterial challenge in mares susceptible and resistant to chronic uterine infection. J Reprod Fert 1993;99:307–13. 12. LeBlanc MM, Johnson RD, Calderwood Mays MB, Valderrama C. Lymphatic clearance of India ink in reproductively normal mares and mares susceptible to endometritis. Biol Reprod 1995; Mono 1: Equine Repro 6:501–6. 13. Nambo Y, Oikawa M, Yoshihara T, Kuwano A, Katayama Y. Age-related morphometrical changes of arteries of uterine wall in mares. Zentralbl Veterinarmed A 1995;42:383–7.
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14. Schoon D, Schoon H-A, Klug E. Angioses in the equine endometrium – Pathogenesis and clinical correlations. Pferdeheilkunde 1999;15:541–6. 15. Esteller-Vico A, Liu IKM, Steffey EP, Vaughan ME, Brosnan RJ. Effect of vascular degeneration on utero-ovarian blood flow and perfusion in the cyclic mare (Abst). Theriogenology 2007;68:506. 16. Ferreira JC, Gastal EL, Ginther OJ. Uterine blood flow and perfusion in mares with uterine cysts: effect of the size of the cystic area and age. Reproduction 2008;135:541–50. 17. Rasch K, Schoon HA, Sieme H, Klug E. Histomorphological endometrial status and influence of oxytocin on the uterine drainage and pregnancy rates in mares. Equine Vet J 1996;28:455–60. 18. Causey RC, Ginn P, Katz BP, Hall BJ, Anderson KJ, LeBlanc MM. Mucus production by endometrium of reproductively healthy mares and mares with delayed uterine clearance. J Reprod Fertil Suppl 2000;56:333–9. 19. Causey RC. Mucus and the mare: How little we know. Theriogenology 2007;68:386–94. 20. Hemberg EN, Lundeheim N, Einarsson S. Retrospective study on vulvar conformation in relation to endometrial cytology and fertility in thoroughbred mares. J Vet Med A Physiol Pathol Clin Med 2005: 52:474–7. 21. McKinnon AO, Squires EL, Voss JL. Ultrasonographic evaluation of the mare’s reproductive tract: Part I. Compend Cont Educ Pract Vet 1987;9:336–45. 22. McKinnon AO, Squires EL, Voss JL. Ultrasonographic evaluation of the mare’s reproductive tract: Part II. Compend Cont Educ Pract Vet 1987;9:472–82. 23. McKinnon AO. Uterine pathology. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Philadelphia: Williams and Wilkins, 1998; pp. 181–200. 24. Pycock JF. Breeding management of the problem mare. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. St Louis: Saunders Elsevier, 2009; pp. 139–64. 25. Beecham DP, McKinnon AO. How to diagnose common equine reproductive tract pathogens using chromagenic agar. Proc AAEP 2009,55:320–25. 26. Riddle WT, LeBlanc MM, Stromberg AJ. Relationships between uterine culture, cytology and pregnancy rates in a Thoroughbred practice. Theriogenology 2007;68:395– 402. 27. Nielsen JM, Troedsson MH, Zent WW. Results of bacteriological and cytological examinations of the endometrium of mares in a practice in Denmark and in Central Kentucky, USA. In: Proceedings of The Chronically Infertile Mare: Havemeyer Foundation Workshop, Hilton Head Island, SC, 5–8 Nov 2008, pp. 34–5. 28. Bindslev MM, Villumsen MH, Petersen MM, Nielsen JM, Bogh IB, Bojesen AM. Genetic diversity of S. equi ssp. zooepidemicus and E. coli isolated from the reproductive tract of the mare. Reprod Domest Anim 2008;43:110–11. 29. Roszel JF, Freeman KP. Equine endometrial cytology. Vet Clin N Am Equine Pract 1988;4:247–62. 30. Nielsen JM. Endometritis in the mare: A diagnostic study comparing cultures from swab and biopsy. Theriogenology 2005;64:510–18. 31. LeBlanc MM, Magsig J, Stromberg AJ. Use of a low volume uterine lavage for diagnosing endometritis in chronically infertile mares. Theriogenology 2007;68: 403–12.
32. Kenney RM. Cyclic and pathologic changes in the mare’s endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1978;172:241–62. 33. Blanchard TL, Garcia MC, Kintner LD, Kenney RM. Investigation of the representativeness of a single endometrial sample and the use of trichrome staining to aid in the detection of endometrial fibrosis in the mare. Theriogenology 1987;28:445–50. 34. Kenney RM, Doig PA. Equine endometrial biopsy. In: Morrow DA (ed.) Current Therapy in Theriogenology. Philadelphia: WB Saunders, 1986; pp. 723–9. 35. Schoon HA, Schoon D. The category I mare (Kenney and Doig 1986): Expected foaling rate 80–90% – fact or fiction? Pferdeheilkunde 2003;19:698–701. 36. Schlafer DH. Equine endometrial biopsy: Enhancement of clinical value by more extensive histopathology and application of new diagnostic techniques? Theriogenology 2007;68:413–23. 37. Brinsko SP, Rigby SL, Varner DD, Blanchard TL. A practical method for recognizing mares susceptible to post-breeding endometritis. Proc Am Assoc Equine Pract 2003;49:363–5. 38. Bucca S, Carli A, Buckley T, Dolci G, Fogarty U. The use of dexamethasone administered to mares at breeding time in the modulation of persistent post breeding induced endometritis. Theriogenology 2008;70:1093–2000. 39. LeBlanc MM, Neuwirth L, Jones L, Cage C, Mauragis D. Differences in uterine position of reproductively normal mares and those with delayed uterine clearance detected by scintigraphy. Theriogenology 1998;50:49–54. 40. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post insemination on pregnancy rate in mares. Theriogenology 1991;35:1111–19. 41. Combs GB, LeBlanc MM, Neuwirth L, Tran TQ. Effects of prostaglandin F2α, cloprostenol and fenprostalene on uterine clearance of radiocolloid in the mare. Theriogenology 1996;45:1449–55. 42. LeBlanc MM, Neuwirth L, Mauragis D, Klapstein E, Tran T. Oxytocin enhances clearance of radiocolloid from the uterine lumen of reproductively normal mares and mares susceptible to endometritis. Equine Vet J 1994;26:279–82. 43. LeBlanc MM. Oxytocin – the new wonder drug for treatment of endometritis? Equine Vet Ed 1994;6:39–43. 44. Pycock JF. Assessment of oxytocin and intrauterine antibiotics on intrauterine fluid and pregnancy rates in mares. Proc Am Assoc Equine Pract 1994;40:19–20. 45. Troedsson MHT, Scott MA, Liu IKM. Comparative treatment of mares susceptible to chronic uterine infection. Am J Vet Res 1995;56:468–72. 46. Pycock JF, Newcombe JR. Assessment of the effect of three treatments to remove intrauterine fluid on pregnancy rate in the mare. Vet Rec 1996;138:320–3. 47. Knutti B, Pycock JF, van der Weijden GC, Kupper U. The influence of early postbreeding uterine lavage on pregnancy rate in mares with intrauterine fluid accumulations after breeding. Equine Vet Ed 2000;5:346–9. 48. Pycock JF. Therapy for mares with uterine fluid. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier; 2007; pp. 93–104. 49. Troeddson MHT. Therapeutic considerations for matinginduced endometritis. Pferdeheilkunde 1997: 13; 516–20. 50. Nie GJ, Johnson KE, Wenzel JGW, Braden TD. Effect of periovulatory ecbolics on luteal function and fertility. Theriogenology 2002;58:461–3.
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Breeding the Problem Mare 51. Williamson P, Duning A, O’Conner J, Penhale WJ. Immunoglobulin levels, protein concentrations and alkaline phosphatase activity in uterine flushing from mares with endometritis. Theriogenology 1983;19:441–8. 52. Dell Aqua JA, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Melo CM. Modulation of acute uterine inflammatory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94:293– 7. 53. Brinsko SP, Varner DD, Blanchard TL, Meyers SA. The effect of postbreeding uterine lavage on pregnancy rate in mares. Theriogenology 1990;33:465–75. 54. Vanderwall DK, Woods GL. Effect on fertility of uterine lavage performed immediately prior to insemination in mares. J Am Vet Med Assoc 2003;222:1108–10. 55. Pascoe DR, Stabenfeldt GH, Hughes JP, Kindahl H. Endogenous prostaglandin F-2 alpha release induced by physiologic saline solution infusion in utero in the mare: Effect of temperature, osmolarity, and pH. Am J Vet Res 1989;50:1080–3. 56. LeBlanc, MM. Effects of oxytocin, prostaglandin and phenylbutazone on uterine clearance of radiocolloid. Pferdeheilkunde 1997;13:483–5. 57. Paccamonti DL, Gutjahr S, Pycock JF, van der Weyden GC, Taverne MAM. Does the effect of oxytocin on intrauterine pressure vary with dose or day of treatment (ABST). Pferdeheilkunde 1997;13:553. 58. Gutjahr S, Paccamonti DL, Pycock JF, Taverne MA, Dieleman SJ, van der Weyden GC. Effect of dose and day of treatment on uterine response to oxytocin in mares. Theriogenology 2000;54:447–56. 59. Paccamonti DL, Pycock JF, Taverne MAM, Bevers M, VanDer Weijden GC, Gutjahr S, Schams D, Blouin D. PGFM response to exogenous oxytocin and determination of the half-life of oxytocin in nonpregnant mares. Equine Vet J 1999;31:285–8. 60. von Reitzenstein M, Colahan M, Hansen P, LeBlanc MM. Aberrations in uterine contractile patterns in mares with delayed uterine clearance after administration of detomidine and oxytocin. Theriogenology 2002;58:887–8. 61. Schramme AR, Pinto CR, Davis JL, Whitacre MD, Whisnant CS. Pharmacokinetics of carbetocin, a long-acting oxytocin analogue, following intravenous administration in horses. Theriogenology 2007;68:517 [abstract]. 62. Nie GJ, Johnson KE, Wenzel JGW, Braden TD. Effect of administering oxytocin or cloprostenol in the periovulatory period on pregnancy outcome and luteal function in mares. Theriogenology 2003;60:1111–18. 63. Troedsson MHT, Ababneh MM, Ohlgren AF, Madill S, Vetscher N, Gregas M. Effect of periovulatory prostaglandin F2α on pregnancy rates and luteal function in the mare. Theriogenology 2001;55:1891–9. 64. Brendemuehl JP. Effect of oxytocin and cloprostenol on luteal formation, function and pregnancy rates in mares. Theriogenology 2002;58:623–6. 65. Cadario ME, Thatcher MJ, LeBlanc MM. Relationship between prostaglandin and uterine clearance of radiocolloid in the mare. Biol Reprod 1995; Mono 1 Equine Repro 6:495–500. 66. Pascoe DR. Effect of adding autologous plasma to an intrauterine antibiotic therapy after breeding on pregnancy rates in mares. Biol Reprod Mono 1:539–543. 67. Troedsson MHT, Liu IKM, Ing M, Pascoe J. Smooth muscle electrical activity in the oviduct and the effect of oxytocin PGF and PGE on the myometrium and the oviduct
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73.
74.
75.
76. 77.
78.
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of the cycling mare. Biol Reprod 1995; Mono 1: Equine Repro 6:475–88. Fumuso E, Giguere S, Wade J, Rogand D, Videla-Dorna I, Bowden RA. Endometrial IL-1β, IL-6 and TNF-α, mRNA expression in mares resistant or susceptible to postbreeding endometritis. Effects of estrous cycle, artificial insemination and immunmodulation. Vet Immunol Immunopath 2003;96:31–41. Fumuso E, Aguilar J, Giguere S, David O, Wade J, Rogan D. Interleukin-8 (IL-8) and 10 (IL-10) mRNA transcriptions in the endometrium of normal mares and mares susceptible to persistent post-breeding endometritis. Animal Reprod Sci 2006;94:282–5. Fumuso EA, Aguilar J, Giguere S, David O, Wade J, Rogan D. Immune parameters in mares resistant and susceptible to persistent post-breeding endometritis: effects of immunomodulation. Vet Immunol Immunopath 2007;118:30–9. Rohrbach BW, Sheerin PC, Cantrell CK, Matthews PM, Steiner JV, Dodds LE. Effect of adjunctive treatment with intravenously administered Propionibacterium acnes on reproductive performance in mares with persistent endometritis. J Am Vet Med Assoc 2007;231:107–13. Vandaele H, Daels P, Piepers S, LeBlanc MM. The effect of post-insemination dexamethasone treatment on pregnancy rates in mares. In: Proceedings of The Chronically Infertile Mare: Havemeyer Foundation Workshop, Hilton Head Island, SC, 5–8 Nov 2008, pp. 43–4. Cox WI. Examining the immunologic and hematopoietic properties of an immunostimulant. Vet Med 1988;6:424– 8. Hinrichs K, Spensley MS, McDonough PL. Evaluation of progesterone treatment to create a model for equine endometritis. Equine Vet J 1992;24:457–61. McDonnell AW, Watson ED. The effect of transcervical uterine manipulations on establishment of uterine infection in mares under the influence of progesterone. Theriogenology 1992;38:945–50. LeBlanc MM. The current status of antibiotic use in equine reproduction. Equine Vet Educ 2009;21:156–66. LeBlanc MM, Lopate C, Knottenbelt D, Pascoe, R. The Mare. Equine Stud Farm Medicine and Surgery. London: Elsevier, 2003; pp. 113–213. Brumbaugh GW, Langston VC. Principles of antimicrobial therapy. In: Smith B (ed.) Large Animal Internal Medicine, 3rd edn. St Louis: Mosby Inc, 2002; pp. 1349– 70. Hinrichs K, Cummings MR, Sertich PL, Kenney RM. Clinical significance of aerobic bacterial flora of the uterus, vagina, vestibule, and clitoral fossa of clinically normal mares. J Am Vet Med Assoc 1988;193:72–5. Bennett DG, Polonad HJ, Kaneps AJ, et al. Histologic effect of infusion solutions on the equine endometrium. Equine Pract 1982;3:37–44. Dascanio JJ, Parker NA, Ley WB, Warnick LD, Sponenberg DP. Magnesium-sulfate intrauterine therapy in the mare. Equine Pract 1998;20:10–13. Gores-Lindholm A, Ahlschwede S, Causey R, LeBlanc MM. Effect of intra-uterine infusion of diluted Nacetylcysteine on equine endometrium. Proc Am Assoc Equine Pract 2009 (in press) [abstract]. Ley WB, Bowen JM, Sponenberg DP, Lessard PN. Dimethyl sulfoxide intrauterine therapy in the mare: Effects upon endometrial histological features and biopsy classification. Theriogenology 1989;32:263–76.
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84. Cardone A, Zarcone R, Borrelli A, Di Cunzolo A, Russo A, Tartaglia E. Utilisation of hydrogen peroxide in the treatment of recurrent bacterial vaginosis. Minerva Ginecol 2003;55:483–92. 85. McKinnon AO. Breeding the problem mare. In: Proceedings of the 11th South African Equine Veterinary Association Congress, Kwazulu Natal, 2009, pp. 192–227. 86. Bracher V, Neuschaefer A, Allen WR. The effect of intrauterine infusion of kerosene on the endometrium of mares. J Reprod Fertil Suppl 1991;44:706–7. 87. Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, Lappin-Scott HM. Microbial biofilms. Annu Rev Microbiol 1995;49:711–45. 88. Shapiro JA. Thinking about bacterial populations as multicellular organisms. Annu Rev Microbiol 1998;52:81–104. 89. Mah T-FC, O’Toole GA. Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol 2001:9:34–9. 90. Soto SM, Smithson A, Horcajada JP, Martinez JA, Mensa JP, Vila J. Implication of biofilm formation in the persistence of urinary tract infection caused by uropathogenic Escherichia coli. Clin Microbiol Infect 2006;12:1034–6. 91. Wooley RE, Jones MS, Shotts EB Jr. Uptake of antibodies in gram-negative bacteria exposed to EDTA-Tris. Vet Microbiol 1984;10:57–70. 92. Farca AM, Nebbia P, Re G. Potentiation of the in vitro activity of some antimicrobial agents against selected gram-negative bacteria by EDTA-tromethamine. Vet Res Commun 1993;17:77–84. 93. Kirkland KD, Fales WH, Blanchard TL, Youngquist RS, Hurtgen JP. The in vitro effects of edta-tris, edta-trislysozyme, and antimicrobial agents on equine genital isolates of Pseudomonas aeruginosa. Theriogenology 1983;20:287–95. 94. Dascanio JJ. Treatment of fungal endometritis. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 116–20. 95. Giguere S. Antifungal chemotherapy. In: Giguere S, Prescott JF, Baggot JD, Walker RD, Dowling PM (eds) Antimicrobial Therapy, 4th edn. Ames, Iowa: Blackwell Publishing, 2006; pp. 301–22.
96. Ricketts SW. The treatment of equine endometritis in studfarm practice. Pferdeheilkunde 1999;15:588–93. 97. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. 98. McCue PM. Ovulation failure. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 83–85. 99. Ginther OJ, Gastal MO, Gastal El, Jacob JC, Beg MA. Induction of haemorrhagic anovulatory follicles in mares. Reprod Fertil Dev 2008;20:947–54. 100. Card C. Hormone therapy in the mare. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. St Louis: Saunders Elsevier, 2009; pp. 89–97. 101. Besognet B, Hansen BS, Daels PF. Induction of reproductive function in anestrous mares using a dopamine antagonist. Theriogenology 1997;47:467–80. 102. Steiner J, Antczak DF, Wolfsdorf K, Saville K, Brooks S, Miller D, Bailey E, Zent W. Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. 103. Piquette GN, Kenney RM, Sertich PL, Yamoto M, Hsueh AJ. Equine granulosa-theca cell tumors express inhibin α and β A-subunit messenger ribonucleic acids and proteins. Biol Reprod 1990;43:1050–7. 104. McCue PM. Equine granulosa cell tumors. Proc Am Assoc Equine Pract 1992;38:587–93. 105. McCue PM, LeBlanc MM, Akita GY, Pascoe JR, Witherspoon DM, Stabenfeldt GH. Granulosa cell tumors in two cycling mares. J Equine Vet Sci 1992;11:281–2. 106. Vanderwall DK. Early embryonic loss. In: Robinson NE, Sprayberry KA (eds) Current Therapy in Equine Medicine, 6th edn. St Louis: Saunders Elsevier, 2009; pp. 830–3. 107. Ginther OJ. Reproductive Biology of the Mare. Cross Plains, WI: Equiservices, 1992. 108. Daels PF, Besognet B, Hansen B, Mohammed H, Odensvik K, Kindahl H. Effect of progesterone on prostaglandin F-2 alpha secretion and outcome of pregnancy during cloprostenol-induced abortion in mares. Am J Vet Res 1996;57:1331–6. 109. LeBlanc MM. The chronically infertile mare. Proc Am Assoc Equine Pract 2008;54:391–407.
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Chapter 273
Fungal Endometritis Marco A. Coutinho da Silva and Marco A. Alvarenga
Introduction Endometritis is a common cause of subfertility in mares and is often caused by bacterial infections. Despite its lower incidence, fungal endometritis has also been recognized as an important cause of subfertility in mares, since mycotic infections are often more difficult to treat than bacterial infections and have a poorer prognosis. The terminology used to describe fungal elements can be confusing and warrants clarification. The term fungus refers to both morphological forms of yeast and mold/hyphae. Yeasts are round to oblong single-cell organisms, while molds or hyphae are long, branched, filamentous structures. Some fungi are dimorphic, for example Candida albicans, and can grow as yeast and/or molds (hyphae) according to environmental conditions. Fungal elements that cause reproductive disease are generally opportunistic. The primary reservoir for infectious agents that colonize the uterus is the caudal reproductive tract, including vagina and external genitalia. Factors that facilitate the colonization of the uterus by opportunistic fungi include poor perineal conformation, pneumovagina, and excessive intrauterine manipulations (culture/cytology/biopsy, artificial insemination, lavage, infusion of antibiotics). In addition, mares with poor uterine defense mechanisms have decreased ability to clear fluid/organisms from their uterus and are at a higher risk of developing fungal endometritis. This chapter will discuss current concepts of the etiology, diagnosis, and treatment of fungal endometritis in mares.
Etiology An extensive list of fungi isolated from the uterus of mares has been published by Dascanio et al.1, 2 Since then, several new fungal elements have been identified on uterine cultures submitted to the diagnostic laboratory at Cornell University from 1999 to 2007,
and an updated list is presented in Table 273.1 (MA Coutinho da Silva, unpublished data). The overall incidence of fungal endometritis diagnoses range from 1 to 5%,3–9 but these numbers may include repeat cultures on the same mare; consequently, the values may be slightly elevated.1 Undoubtedly, Candida spp. and Aspergillus spp. are the most common microorganisms isolated from mares with fungal endometritis. In Dr Coutinho da Silva’s experience, the incidences of Candida spp., Aspergillus spp., and Mucor spp. in 69 positive samples from the uterus of mares were 58%, 25%, and 12%, respectively. In addition, more than one fungal organism was isolated in 19% of these cases (M.A. Coutinho da Silva, unpublished data). The mechanism(s) involved in fungal contamination and colonization of the uterus of mares is still not well understood, and studies on the pathogenesis and treatment of fungal endometritis are scarce. In healthy mares, growth of fungi is prevented by factors present in the vaginal flora and also the local cellular immune system. Causes for the initial colonization of the uterus by pathogens include poor uterine contractility, anatomical malformations, and inefficient uterine defense mechanisms. This may explain why fungal endometritis is commonly associated with a history of uncorrected anatomical defects that lead to pneumovagina,5 recurrent persistent post-breeding endometritis, and frequent intrauterine antibiotic therapy.1, 6, 10 Most mares presented with fungal endometritis have a history of previous bacterial endometritis. These mares usually undergo intense therapy that includes frequent uterine lavages and intrauterine infusion of antibiotics. The frequent uterine manipulation, associated with anatomical problems leading to pneumovagina, results in chronic contamination of the uterus. In addition, antibiotics drained from the uterus alter the normal bacterial flora in the vagina, predisposing to an excessive growth of opportunistic
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 273.1 Fungal organisms isolated from the uterus of mares Acremonium spp. Actinomyces spp. Aspergillus fumigatus Aspergillus glaucus Aspergillus niger Aspergillus spp. Aureobasidium pullulans Candida albicans Candida ciferii Candida famata Candida guillermondii Candida krusei Candida lusitaniae Candida parapsilosis Candida pseudotropicalis Candida rugosa Candida spp. Candida stellatoides Candida tropicalis Candida zeylanoides Circinella spp. Coccidiodes immitis Cryptococcus humicolus Cryptococcus neoformans
Cryptococcus terreus Fusarium spp. Hansenula anomala Hansenula polymorpha Monosporium apiospermum Monosporium spp. Mucor spp. Nocardia spp. Paecilomyces spp. Penicillium spp. Pseudallescheria boydii Rhodotorula glutinis Rhodotorula minuta Rhodotorula rubra Rhodotorula spp. Scedosporium apiospermum Saccharomyces cerevisiae Torulopsis candida Torulopsis glabrata Trichosporon beigelii Trichosporon cutaneum Trichosporon spp. Yarrowia lipolytica
Adapted from Dascanio et al.1 Data include new fungal organisms that have been isolated from endometrial swabs submitted to the diagnostic laboratory at Cornell University from 1999 to 2007 (M.A. Coutinho da Silva, unpublished).
fungal organisms, making the vagina and external genitalia the primary reservoir for pathogens. A recent study from the University of Utrecht involving 128 cases of mares with fungal endometritis showed that the most frequent factors associated with fungal endometritis were: previous antibiotic therapy (87%), pneumovagina/urovagina (39%), and large amounts of intrauterine fluid accumulation (33%).11 In the same study, the incidence of fungal endometritis was significantly higher in barren mares (74%) compared to maiden (16%) or foaling (9%) mares. In the author’s experience (M.A.A.), most mares with fungal endometritis are old barren mares that are being used intensively as embryo donors during consecutive years. These mares are submitted to multiple, excessive manipulations of the genital tract [artificial insemination (AI), embryo flushes, and uterine treatments] during consecutive cycles during the breeding season, predisposing them to ascending infections. In women, the normal vaginal microflora plays an important role in the prevention of vaginitis, caused mostly by Candida albicans. Lactobacillus spp., which are part of the normal vaginal microflora in women, inhibit the growth of pathogens by preventing their adhesion to the vagina wall and by secreting lactic acid, hydrogen peroxide, and bactericidal peptides.
The administration of antibiotics suppresses lactobacilli and results in an overgrowth of Candida spp. and disease. The same principles are now being extrapolated to the horse as a possible cause for fungal endometritis. Recently, Fraga et al.12 isolated lactobacilli from the vagina of 70% of healthy mares, supporting the theory that disruption of the vaginal flora predisposes to an overgrowth of pathogens that may invade the uterus and cause disease. To date, there are no reports on the vaginal flora of mares with fungal endometritis. Therefore, further studies are necessary to confirm this hypothesis and to determine if the use of probiotics containing lactobacilli would be beneficial in preventing fungal endometritis. In general, the severity of the inflammation as well as response to treatments varies according to causal organism and duration of infection. It has been suggested that hyphae are more invasive than yeast, and consequently, more difficult to treat and eliminate. Candida albicans can be present in both yeast and hyphal forms and therefore can penetrate deeper into the endometrium and/or grow intracellularly.11 However, when endometrial cytology and histology were used to diagnose fungal endometritis,9 fungal elements (yeast and hyphae) were only visualized in the lumen of the uterus and did not penetrate deeper in the endometrium. Another important observation about the difficulty in treating fungal infections is the ability of certain species to produce biofilms; both Candida spp. and Aspergillus spp. can produce biofilms.13, 14 Biofilms are formed by an extracellular matrix composed mainly of cell wall-like polysaccharides combined with a dense network of fungal cells. Biofilms are a protective niche for microorganisms, providing antifungal resistance and creating a source of persistent infection. Biofilms also give microorganisms the ability to adhere strongly to virtually any surface; therefore, proper sterilization of uterine catheters and other equipment is paramount to prevent dissemination of pathogens. Important factors that predispose a mare to fungal colonization of the uterus are:
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poor perineal conformation leading to pneumovagina/pneumouterus and fecal contamination of reproductive tract; changes in vaginal microflora induced by repeated intrauterine antibiotic therapy; iatrogenic contamination of the uterus by frequent genital manipulation; compromised uterine defense mechanisms associated with delayed uterine clearance.
Other factors contributing to fungal infections include the presence of a moist environment, exposure to a large number of fungi, and the presence of
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R demonstrating the presence of yeast (a) and hyphae (b). Note Figure 273.1 Endometrial cytology of mares stained with Diff-Quik the presence of fungal organisms (black arrows), neutrophils (white arrows), and endometrial cells (arrowheads) (×1000). (See color plate 127).
a necrotic focus as occurring with trauma, abortion, and retained placenta.15, 16
Diagnosis Diagnosis of fungal endometritis is based on history, clinical signs, and identification of fungi in endometrial culture, cytology, and biopsy. Most mares affected by fungal endometritis have a prolonged history of infertility, have received repeated intrauterine infusion of antibiotics, and have poor perineal conformation associated with pneumovagina. Other factors reported include dystocia or retention of fetal membranes, early embryonic death, and abortion.11 Affected mares usually display intrauterine fluid accumulation with or without vaginal discharge, which can range from mucoid to gray and purulent in consistency. Definitive diagnosis of fungal endometritis can be achieved by evaluating endometrial cytology, culture, and biopsy. Endometrial cytology is an easy, inexpensive, and rapid procedure that allows practitioners quickly to assess uterine health by evaluating the presence/quantity of white blood cells as well as pathogens. Samples for endometrial cytology should be collected using a double-guarded cytological brush or swab, or by a small-volume lavage of the uterus. The authors prefer to use the cytological brush to obtain an endometrial sample for cytology because it preserves the architecture of the cells and collects more material from the uterus than the cotton swab.17 The use of a low-volume uterine lavage to collect uterine material for cytology and culture has received attention in recent years and may prove to have better sensitivity and specificity in detecting uterine infections. After collection, samples are smeared onto a glass slide, air dried, and stained R , Baxter with modified Wright’s stain (Diff-Quik Health Care, Miami, FL). Cytological evaluation should be performed at ×1000 and usually reveals
signs of inflammation (i.e., frequent neutrophils), and presence of fungal elements (yeast or hyphae; Figure 273.1). The fungal elements most commonly observed are yeasts more so than hyphae (67% vs 27%9 ). In some cases, yeasts and hyphae are imaged together (Figure 273.2). Yeasts are oval or spherical in shape, measure approximately 4 µm in diameter, and have a thick capsule that usually does not retain stains.16, 18 Inflammatory cells (neutrophils) are common on cytological evaluations, and occasionally it is possible to observe neutrophils phagocytosing yeasts (Figure 273.2). Endometrial biopsies should be performed to determine the location of fungal elements within the layers of the uterus, the severity of the inflammation, and the extension of the damage to the endometrium. When fungal endometritis is suspected, histological sections should be stained with Gomori’s methanamine silver stain9, 19 to facilitate visualization of fungal elements (Figure 273.3). Fungal elements,
Figure 273.2 Endometrial cytology of mares stained with DiffR demonstrating the presence of both yeast (white arQuik rows) and hyphae (black arrows) forms. Note the presence of yeast elements within the cytoplasm of neutrophil (phagocytosis; arrowheads) (×1000). (See color plate 128).
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Figure 273.3 Endometrial biopsy stained with periodic acid–Schiff (PAS; a,c) or Gomori’s methanamine silver (GMS; b,d). Yeasts can be clearly visualized within luminal contents in samples stained with GMS (arrows) compared to PAS. (a,b ×200; c,d ×1000). (See color plate 129).
stained black or gray, may appear in the lumen, adhered to the luminal surface, within endometrial gland lumens, or within the endometrium.9, 15 Most fungal infections of the uterus are superficial, with fungal elements located adjacent to the luminal surface or free in the uterine lumen (Figure 273.3). Thus caution is advised when interpreting biopsy results as fungal elements may be washed off the slide during processing, resulting in a false-negative result. Fungal culture in vitro requires specific culture media (e.g., Sabouraud’s agar) and a prolonged incubation time compared to most bacteria. Therefore, it is important to alert the laboratory that a fungal culture is being requested to decrease the likelihood of false negatives. Once fungal elements are isolated, a sensitivity test should be performed to determine the appropriate antimycotic treatment. In the authors’ opinion, the use of endometrial cytology associated with aerobic and fungal cultures is the most practical method of diagnosing fungal endometritis. The absence of inflammatory cells in the cytology should not completely preclude the possibility of a fungal uterine infection. Conversely, a single positive endometrial culture for fungus in the absence of clinical evidence of infection or history of infertility is potentially a false positive. In these cases,
obtaining a second sample to confirm the previous diagnosis is recommended. In addition, information obtained from endometrial biopsy is very useful in determining the treatment modality (local vs systemic), evaluating the response of infection to treatment (serial samples), and determining the prognosis for fertility.
Treatment Due to the scarcity of controlled studies on fungal endometritis in mares, several treatment regimens have been used to treat fungal endometritis, with variable results. In general, treatment should preferably be initiated when the mare is in estrus (and local immunity is maximal), and should include correction of anatomical defects (e.g., poor vulvar conformation), large-volume uterine lavage combined with ecbolic treatment (e.g., oxytocin), and use of appropriate antifungal drug, either systemically or locally. Since few laboratories perform antifungal susceptibility routinely, it is important for the clinician to be familiar with the type of fungi most likely to be encountered and the susceptibility of those fungi to readily available antifungal drugs. In veterinary medicine, there are two classes of antifungal drugs
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Fungal Endometritis Table 273.2 Overall susceptibility pattern of fungi to antifungal drugs commonly used to treat fungal endometritis Susceptibility pattern (% of isolates) Antifungal Polyenes Amphotericin B Natamycin Nystatin Azoles Clotrimazole Ketoconazole Miconazole Itraconazole Fluconazole Flurocytosin
Susceptible
Intermediate
Resistant
96 100 100
0 0 0
4 0 0
80 81 43 62 44 83
13 15 41 38 14 0
7 4 16 0 42 17
Antifungal sensitivity results obtained from fungal organisms that have been isolated from endometrial swabs submitted to the diagnostic laboratory at Cornell University from 1999 to 2007 (M.A. Coutinho da Silva, unpublished).
that are commonly used: the azoles and polyene drugs. These drugs interfere with the cytoplasmic membrane of fungi by either binding to ergosterol (polyenes) or inhibiting its synthesis (azoles). In general, polyenes have a fungicidal activity against most pathogenic fungi, whereas azoles are considered fungistatic. However, some azoles can be fungicidal at higher doses. A list of common antifungal drugs and the susceptibility pattern of fungi isolated from the uterus of mares is presented in Table 273.2. We evaluated the susceptibility pattern of fungi (yeast and/or hyphae) isolated from the uterus of 69 mares and submitted to Cornell University from 1999 to 2007, and observed the following susceptibility pattern for the following drugs: nystatin (100%); natamycin (100%); amphotericin B (96%); ketoconazole (81%); clotrimazole (80%); and itraconazole (61%). Interestingly, only 44% of samples were susceptible to fluconazole and miconazole (M.A. Coutinho da Silva, unpublished). Most antifungal agents and dosages used in horses have been extrapolated from human medicine. Dosages commonly used to treat fungal endometritis in mares are presented in Table 273.3. The duration of treatment for fungal endometritis is usually prolonged (7 to 21 days), and therefore the toxicity of the antifungal agent must be considered, particularly when used systemically. In most cases, we recommend systemic therapy instead local therapy, in order to reduce uterine manipulations. When local therapy is necessary, infusion of the antimycotic drug into the uterus should be performed no earlier than 2 hours after oxytocin administration, to avoid quick clearance from the uterus due to increased
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Table 273.3 Suggested doses for drugs used in the treatment of fungal endometritis in mares Antifungal drug
Local therapy
Systemic therapy
Amphotericin B Nystatin Clotrimazole Ketoconazole Miconazole Itraconazole Fluconazole Lufenuron
100–200 mg 0.5–2.5 million U 500–700 mg – 400–700 mg – 100 mg 540 mg
0.3–0.9 mg/kg, i.v.a – – 30 mg/kg, p.o., b.i.db – 5 mg/kg, p.o., b.i.d. 5 mg/kg, p.o., s.i.d.c –
Data from Dascanio et al.;1 Giguere et al.;20 Michelle LeBlanc, personal communication. a Diluted to 1 mg/mL in 5% dextrose and administered over 1 to 2 h, every 24 h for 3 days, then every 48 h. b Diluted in 0.2 N HCl and administered by nasogastric intubation. c A loading dose of 14 mg/kg is recommended.
myometrial activity. The use of most antifungal drugs is discouraged in pregnant animals. Excellent information about drug pharmacokinetics and toxicity can be found in Giguere et al.20 Systemic antifungal therapy Amphotericin B can be administered slowly intravenously when diluted in 5% dextrose solution. However, amphotericin B is nephrotoxic and requires monitoring of blood urea nitrogen (BUN) or creatinine to evaluate the extent of renal damage. Amphotericin B is a broad-spectrum antifungal with fungicidal activity against most pathogenic fungi, including Candida spp. and Aspergillus spp. Because amphotericin B has a long half-life (26 h in dogs), it has been suggested that treatment with amphotericin B should involve daily administration during the first 3 days followed by every-other-day administrations (M. Leblanc, personal communication). However, due to its potential for renal toxicity, amphotericin B is mostly used for intrauterine infusions. Imidazole (ketoconazole) and triazole (itraconazole and fluconazole) drugs are considered fungistatic, and treatment regimens should be prolonged to at least 3 weeks. Ketoconazole is poorly water-soluble, highly lypophilic, and requires acidic pH for dissolution; therefore, oral administration of this drug requires nasogastric intubation to avoid damage to the oral cavity and esophagus. Despite its broad spectrum and good activity against most yeast and Aspergillus spp., the use of ketoconazole in practice has been replaced by triazoles. Triazoles have greatly increased half-lives, increased bioavailability following oral administration, and enhanced antifungal activity. Both itraconazole and fluconazole are well absorbed following oral administration and widely distributed to tissues
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(except the cerebrospinal fluid). Fluconazole has the advantage of being water soluble and its half-life is about 40 h in horses, compared to 6.5 h for itraconazole. In addition, fluconazole is fairly inexpensive compared to itraconazole. In general, fluconazole is recommended to treat yeast infections, in particular Candida spp., while itraconazole (oral solution) is the drug of choice to treat infections caused by Aspergillus spp., since most Aspergillus spp. are resistant to fluconazole. Based on the experience of one of the authors (M.A.A.), oral administration of fluconazole has been effective in treating a large number of mares presented with uterine yeast infections; however, recurrence of infection is common. The author recommends oral administration of 2 g of fluconazole daily, starting on the first day of estrus and continuing until 2 days post-ovulation. No side-effects have been observed using this protocol. Intrauterine therapy Most mycotic infections of the uterus are superficial, with fungal elements remaining in the uterine lumen or on the surface of the endometrium.9 Therefore, mechanical removal of fungal elements, debris, and exudate by uterine lavage should be an integral part of the treatment of fungal endometritis. Authors recommend starting the treatment in early estrus to take advantage of the positive effects of estrogen on uterine contractility, cervical relaxation, and neutrophil function. A combination of large-volume uterine lavage and ecbolic drugs (e.g., oxytocin) should be performed daily for 3 to 7 days, depending on the incidence of intrauterine fluid accumulation between examinations. In most cases, the vagina and clitoral fossa should also be rinsed as they can act as reservoir for fungal organisms and lead to reinfection of the uterus after interruption of the treatment.
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Non-specific treatments that have been anecdotally used for uterine lavage include the use of a solution of 2% acetic acid or 0.05–0.1% povidoneiodine.5 However, intrauterine infusion of either acetic acid or povidone-iodine should be performed with caution since damage to the endometrium has been reported.21 We commonly infuse the uterus with 250–500 mL of a 2% acetic acid solution. After 3 to 5 minutes in contact with the endometrium, the acetic acid is washed out from the uterus with 2 liters of saline or Ringer’s lactate to avoid severe uterine irritation. The use of a solution containing 10–20% DMSO (dimethyl sulfoxide) has also been described as an adjuvant to treat fungal endometritis.22 DMSO appear to be bacteriostatic at lower concentrations (5–10%) and bactericidal at higher concentrations. In addition, DMSO may also facilitate the action of local antimycotic drugs by increasing their permeability through tissues and potentially disrupting biofilm protection of the organism. A recent report suggested the use of intrauterine infusion of hydrogen peroxide (1% solution) to treat fungal endometritis in mares.23 In humans, Lactobacillus spp. in vaginal microflora are known to inhibit overgrowth of pathogens by secreting hydrogen peroxide and other substances.24 The authors recommended multiple infusions of hydrogen peroxide during estrus and for at least 2 days after breeding, resulting in a low incidence of recurrence of infection. In some acute cases of endometritis, there is a significant increase in mucus production by the endometrium.25 Based on a recent report by LeBlanc (2009)26 one of the authors (MA Coutinho da Silva) has now incorporated the use of a mucolytic for uterine lavage. In a recent case of acute fungal endometritis caused by Candida sp. (Figure 273.4), the mare was treated with daily uterine lavage with
(b)
Figure 273.4 Endometrial cytology of a 4-year old maiden mare with fungal endometritis as a result of being treated with repeated intrauterine antibiotic infusions during 5 consecutive cycles. In panel (a), large number of yeast (Candida sp.; black arrows) can be visualized along with degenerated neutrophils (white arrows). In panel (b), Candida sp. can be visualized in both yeast (white R and observed at 400× (a) and 1000× (b). arrows) and hyphae (black arrows) forms. Cytologies were stained with Diff-Quik
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Figure 273.5 Endometrial cytology of the mare depicted in figure 273.4 at 48 h after initiation of treatment. Fungal elements were not visualized in the cytology. Only neutrophils (white arrows) endometrial cells (arrowheads) and a small amount of mucus (black arrows) were visualized. The mare was treated with a combination of frequent uterine lavages, intrauterine infusion of a muR and observed at 400× (a) and colytic and systemic antifungal drug (see text for details). Cytologies were stained with Diff-Quik 1000× (b).
Ringer’s lactate containing 0.05% povidone-iodine, followed by uterine infusion of 100 ml of a 6% solution of N-acetylcysteine (Butler Corp., Columbus, Ohio, USA). The mare also received systemic fluconazole for 10 days (Table 273.3). A significant improvement was observed within 48 h of treatment (Figure 273.5) with no fungal elements observed in the endometrial cytology. The mare was inseminated with frozen semen on the 8th day of treatment and a fetus with a heartbeat was detected on day 25 of pregnancy. Despite the lack of scientific information on the effects of acetylcysteine in the endometrium, the rapid resolution of the infection and the positive outcome if this case are encouraging. Several antifungal drugs have been used for daily infusions into the mare’s uterus (Table 273.3). The identification of a fungal species and its susceptibility to antifungal drugs may not always be possible in
practice, due to the scarcity of laboratories that perform these tests for fungi and the time required for obtaining final results (approximately 1 month). In these cases, infusion of the following drugs should be considered based on previous susceptibility patterns: nystatin, amphotericin B, ketoconazole, or clotrimazole (Table 273.2). Lufenuron, a drug used for flea control and known to interfere with chitin synthesis, has been used successfully to treat mares with fungal endometritis.27 Both yeasts (e.g., Candida spp.) and hyphal forms (e.g., Aspergillus spp.) contain chitin as a component of the cell wall and are potentially affected by lufenuron. However, a recent study showed that lufenuron did not prevent growth of Aspergillus spp. in vitro.26 Despite controversial results, the intrauterine administration of lufenuron was safe, and this drug can potentially be used in association with other
Table 273.4 Suggested strategy to treat a mare with fungal endometritis Step 1.
Surgical correction of anatomical abnormalities (e.g., episioplasty) followed by 10 days of rest
Step 2.
Begin daily uterine lavage: flush the uterus with 1–2 L of saline or LRS; then, infuse 250–500 mL of 2% acetic acid solution and allow 3–5 min of contact with endometrium; lastly, flush the uterus with an additional 2 L of saline or LRS
Step 3.
Immediately after uterine lavage, administer 10–20 IU of oxytocin i.v. or i.m., and repeat after 6 h. If the mare is in diestrus on the first day of treatment, replace the first oxytocin treatment by a dose of PGF2α to induce estrus
Step 4.
Begin treatment with antimycotic drug (see Table 273.3), based on sensitivity results of the organism isolated. If sensitivity test is not available, we suggest treating yeast infections with fluconazole (10 days) and hyphal infections with itraconazole (14–21 days). If cost of treatment is a concern, we suggest intrauterine infusion of clotrimazole as an alternative to itraconazole (7 days)
Step 5.
To avoid reinfection, minimize the number of intrauterine manipulations. Perform a single breeding or AI per cycle; use a sterile sanitary chemise during AI; use double-guarded instruments to collect endometrial samples for culture and/or cytology; monitor/treat the mare for post-breeding intrauterine fluid accumulation. For embryo donors, limit the number of embryo collection attempts and/or allow the mare to rest during a natural estrous cycle
AI, artificial insemination; i.m., intramuscularly; i.v., intravenously; LRS, lactated Ringer’s solution; PGF2α , prostaglandin F2α (10 mg dinoprost tromethamine: R , Pharmacia & Upjohn Co., NY 10017). Lutalyse
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antimycotic drugs to treat fungal endometritis. Further studies on the efficacy of lufenuron or other chitin synthesis inhibitors to treat fungal endometritis are necessary. When local therapy is performed, it is important to consider that every manipulation of the genital tract (e.g., uterine lavage, infusion, sample collection) has the potential to reinfect the uterus with the same or a different organism. Therefore, it is very important to use minimal contamination techniques including the use of double-guarded instruments.
Prognosis The prognosis for mares with fungal endometritis is guarded to poor.1, 5, 11, 19 Potential reasons for treatment failure include insufficient treatment period, inadequate dose or inappropriate antifungal drug, failure to remove predisposing factors, or recontamination from a reservoir in the caudal reproductive tract.1, 11 Recently, Stout11 reported elimination of the fungal organism after treatment in approximately 22% of cycles, resulting in a cure rate of 33% of affected mares. In a previous study, Zafracas5 obtained a slightly higher cure rate of 46% after treating mares with a combination of uterine lavage and nystatin. Complete resolution of infection and subsequent fertility may be difficult to estimate, due to lack of controlled studies in the literature. In addition, a relatively high proportion of mares treated for fungal endometritis develop bacterial endometritis post-treatment.11 Spontaneous resolution of fungal endometritis has been observed in mares after a prolonged period of sexual rest, and may be attributed to re-establishment of normal vaginal flora. Therefore, treatment of fungal endometritis should include a combination of therapies aimed at removing the predisposing factor, eliminating the causative organism, and preventing future contaminations. Table 273.4 summarizes the approach used by the authors to treat mares with fungal endometritis, based on scientific and anecdotal information.
Conclusions In summary, management of mares suffering from fungal endometritis should reduce or eliminate factors that can predispose them to uterine contamination such as excessive uterine manipulations, frequent use of intrauterine antibiotics, and anatomical abnormalities. To minimize the chances of recurrent infections, it is important to use minimal contamination breeding techniques, trying to use a single insemination or breeding per cycle, followed by intense monitoring and management
post-breeding to avoid prolonged intrauterine fluid accumulations. In the case of embryo donors, limiting the frequency of embryo collection attempts and/or allowing the mare to rest during a natural estrous cycle between collections may help in maintaining the balance in vaginal flora and prevent reinfection.
References 1. Dascanio JJ, Schweizer C, Ley WB. Equine fungal endometritis. Equine Vet Educ 2001;13:324–9. 2. Dascanio JJ. Treatment of fungal endometritis. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders/Elsevier, 2007; pp. 116–20. 3. Collins SM. Study of the incidence of cervical uterine infections in Thoroughbred mares in Ireland. Vet Rec 1964;76:673–6. 4. Bain AM. The role of infection in the thoroughbred mare. Vet Rec 1966;78:168–73. 5. Zafracas AM. Candida infection of the genital tract in thoroughbred mares. J Reprod Fertil Suppl 1975;23:349–51. 6. Pugh DG, Bowen JM, Kloppe LH, Simpsom RB. Fungal endometritis in mares. Comp Cont Educ Pract 1986;8:S173–S181. 7. Elvinger F, Roberts AW. Equine Genital Tract Isolates. Tifton: Georgia Veterinary Diagnostic Laboratories, 1995. 8. Elvinger F, Roberts AW. Equine Genital Tract Specimens. Tifton: Georgia Veterinary Diagnostic Laboratories, 1996. 9. Coutinho da Silva MA, Alvarenga MA, Carnevale EM. Diagnosis of fungal endometritis in mares: Efficacy of cytology, histology and stain. In: Proceedings of the Annual Conference of the Society for Theriogenology, 2000, p. 171. 10. Troedsson MHT. Diseases of the uterus. In: Troedsson MHT (ed.) Current Therapy in Equine Medicine 4. Philadelphia: W.B. Saunders Co., 1997; pp. 517–24. 11. Stout TAE. Fungal endometritis in the mare. Pferdeheilkunde 2008;24:83–7. 12. Fraga M, Perelmuter K, Delucchi L, Cidade E, Zunino P. Vaginal lactic acid bacteria in the mare: evaluation of the probiotic potential of native Lactobacillus spp. and Enterococcus spp. strains. Antonie van Leeuwenhoek 2008;93: 71–8. 13. Chandra J, Kuhn DM, Mukherjee PK, Hoyer LL, McCormick T, Ghannoum MA. Biofilm formation by the fungal pathogen Candida albicans: Development, architecture, and drug resistance. J Bacteriol 2001;183:5385–94. 14. Seidler MJ, Salvenmoser S, Muller FMC. Aspergillus fumigatus forms biofilms with reduced antifungal drug susceptibility on bronchial epithelial cells. Antimicrob Agents Ch 2008;52:4130–6. 15. Hurtgen JP, Cummings MR. Diagnosis and treatment of fungal endometritis in mares. In: Proceedings of the Annual Conference of the Society for Theriogenology, 1982, p. 82. 16. Carter GR, Chengappa MM. Introduction to the fungi and fungus infections. In: Carter GR, Chengappa MM, Roberts AW, Claus GW, Rikihisa Y. (eds) Essentials in Veterinary Microbiology. Philadelphia: Williams & Wilkins, 1995; pp. 251–6. 17. Alvarenga MA, Iawama de Mattos MCF, Fabris VE. Comparison between the gynecological brush, cotton swab and aspiration techniques for endometrial cytology in mares.
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18.
19. 20.
21. 22.
23.
In: Proceedings of the Annual Conference of the Society for Theriogenology, 1998, p. 163. Volk WA. Bacterial morphology. In: Volk WA (ed.) Basic Microbiology. New York: Harper Collins Publishers, 1992; pp. 38–56. Freeman KP, Roszel JF, Slusher SH, Payne M. Mycotic infections of the equine uterus. Equine Pract 1986;8:34–42. Giguere S. Antifungal chemotherapy. In: Giguere S, Prescott JF, Baggot JD, Walker RD, Dowling PM (eds) Antimicrobial Therapy in Veterinary Medicine. Oxford: Blackwell Publishing, 2006; pp. 301–22. Perkins NR. Equine reproductive pharmacology. Vet Clin N Am Equine 1999;15:687–704. Pottz GE, Rampey JH, Benjamin F. The effect of dimethyl sulfoxide (DMSO) on antibiotic sensitivity of a group of medically important microorganisms: preliminary report. Ann NY Acad Sci 1967;141:261–72. McKinnon AO. Breeding the problem mare. In: Proceedings of the 11th South African Equine Veterinary Association Congress, Kwazulu Natal, 2009, pp. 192–227.
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24. Verstraelen H. Cutting edge: the vaginal microflora and bacterial vaginosis. Verh K Acad Geneeskd Belg 2008;70:147–74. 25. Causey RC, Miletello T, O’Donnell L, Lyle SK, Paccamonti DL, Anderson KJ, Eilte BE, Morse S, Leblanc MM. Pathologic effects of clinical uterine inflammation on the equine endometrial mucosa. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners 2008; 276–277. 26. LeBlanc MM. New treatment strategies for chronic endometritis and post mating endometritis. Clinical Theriogenology 2009;1:177–183. 27. Hess MB, Parker NA, Purswell BJ, Dascanio JD. Use of lufenuron as a treatment for fungal endometritis in four mares. J Am Vet Med Assoc 2002;15:266–7. 28. Scotty NC, Evans TJ, Giuliano E, Johnson PJ, Rottinghaus GE, Fothergill AW, Cutler TJ. In vitro efficacy of lufenuron against filamentous fungi and blood concentrations after PO administration in horse. J Vet Int Med 2005;19: 878–82.
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Chapter 274
Pyometra Kristina G. Lu
Definition ‘Pyometra’ is a word of Greek origin describing a pus-filled uterus. The purulent fluid filling of the uterus may make the uterus feel uniformly enlarged upon palpation per rectum. Thus differentials for pyometra include pregnancy, a postpartum uterus, and pneumometra.1 Depending on the source, pyometra may be defined only as accumulation of pus within the lumen of the uterus, or as pus accumulation in the uterus in addition to the persistence of the corpus luteum beyond its normal lifespan.2 Some authors define the state of purulent material accumulation within the uterus without the presence of a corpus luteum as metritis. An important clinical distinction of metritis defined in this way is that the mare is still cyclic though the estrous cycles may be irregular. Mares with pyometra are not cycling due to the persistent corpus luteum. Mares with metritis have a better potential for reproductive recovery than mares with pyometra.2 The prognosis for future fertility in mares with pyometra is poor.3
trial cups have been observed in a case of pyometra where no fetus or fetal membranes were found within the uterine lumen.6 Intrauterine accumulation of purulent material has also been observed in the presence of an open cervix.4, 7 These mares may have an innately reduced endometrial resistance to infection, which may ultimately progress to a pyometra.8 A potential cause of poor uterine drainage regardless of cervical patency may be intra-abdominal adhesions to the uterus, keeping the uterus dependent in the abdomen.1
Organisms The most common organism associated with pyometra in the mare is Streptococcus equi ssp. zooepidemicus (hereafter S. zooepidemicus).2, 4 Other reported organisms include Escherichia coli, Actinomyces sp., Pasteurella sp., Pseudomonas sp., Proprionibacterium sp., and Candida rugosa.4, 9
Diagnosis Clinical signs
Causes There are many potential causes of pyometra in the mare. Ultimately, there is an interference of fluid clearance from the uterus. The fluid may have originated as normal endometrial gland secretions or abnormal inflammatory exudate. Outflow obstruction may be caused by a hormonally closed cervix or cervical and caudal reproductive tract pathology. This includes adhesions, fibrosis, or diphtheritic membranes.1, 4, 5 This is often seen secondary to trauma such as that experienced during dystocia. Progesterone from a persistent corpus luteum will functionally close the cervix. Similarly, persistently administered exogenous progesterone will potentially prevent intrauterine fluid clearance. Endome-
The most prominent outward clinical sign of equine pyometra is vulvar discharge. The discharge is most commonly observed when the cervix is relaxed or open. The pus is often creamy though it can be watery or caseous (Figure 274.1). The volume of pus can be variable, with larger volumes associated with a thin, watery, gray appearance and the smaller volumes associated with inspissated caseous exudate. Seldom are there clinical signs of systemic disease.4 Thus equine pyometra is frequently undiagnosed until vulvar discharge is observed.2 Palpation and ultrasonography of the reproductive tract per rectum can reveal intrauterine fluid or fluid mixed with more solidified exudate (Figure 274.2). The existence of intraluminal uterine pus does
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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oviduct. In contrast the cow’s oviduct blends into the uterine horn.3 Interestrus interval
Figure 274.1 Purulent vulvar discharge.
not necessarily indicate extensive loss of endometrial tissue though mares may have variable degrees of endometrial inflammation and fibrosis. Hughes et al.4 reported erosion of the endometrial epithelial layer, with a grossly rough and granular appearance. Epithelial damage was generally worst in the uterine body as observed with histopathology. No cases of purulent uterine fluid accumulation were associated with inflammatory changes in the ovaries or oviducts.4 This is potentially due to the horse having a muscular papilla at the junction of the uterus and
Mares with pyometra may demonstrate abnormal duration of the interestrus period, so that it correlates with the degree of endometrial damage. Interovulatory intervals may be absent, short, normal, or prolonged with a normal duration of estrus and normal intensity of estrous behavior. Histopathological evaluation of the endometrium from mares with shortened interestrus periods revealed an intact endometrium with inflammation. Any endometrial erosion or atrophy was confined to the uterine body. Intermediate endometrial damage, such as large areas of severe endometrial atrophy and erosion, was seen in mares with irregular interestrus periods. Some mares demonstrate prolonged interestrus periods associated with prolonged luteal activity. Histopathology of the endometrium from these mares revealed severe endometrial damage with nearly complete erosion and atrophy.4 The variation in cycle length may be associated with alteration of prostaglandin F2α (PGF2α ) release from the endometrium. Arachidonate metabolites are important mediators of inflammation, and endometrial inflammatory conditions can lead to an increase in arachidonate metabolites. Significantly elevated prostaglandin concentrations are produced by the endometrium with chronic endometritis. No statistically significant increase in the concentrations of prostaglandin E2 (PGE2 ) and leukotriene B4 (LTB4 ) was observed with chronic endometritis.10 The endometrium associated with pyometra and persistent luteal tissue demonstrated similar concentrations of LTB4 and PGE2 to normal mares, and a low level of PGF2α . Histopathology of such mares revealed severe infiltration by neutrophils and loss of the surface epithelium. Surface epithelium loss may lead to low PGF2α . In the cow, endometrial epithelial cells secrete significantly more PGF than PGE2 , whereas the stromal cells produce PGE2 .11 Mares with pyometra have decreased prostaglandin release leading to prolonged or potentially reduced luteal function and prolonged interestrus intervals. The release of small amounts of prostaglandin F2α may occur days to weeks later than the normal release 14 days after ovulation, leading to reduction in luteal function.4 Bloodwork findings
Figure 274.2 Transabdominal ultrasound image of pyometra with relative hyperechogenicity of caseous material ventrally in the uterus.
Hematology trends in mares with pyometra include mild anemia and decreased leukocyte count with neutropenia and sometimes lymphopenia.4
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Mare: Disease of the Uterus nal motility, mild abdominal pain, and mild to mod¨ erate incisional infections. Rotting et al.12 reported on a series of ovariohysterectomies with no evidence of previously reported complications that included hemorrhage, septic peritonitis, uterine stump infection, and necrosis and diarrhea.
References
Figure 274.3 Purulent effluent from uterine lavage.
Treatment Obtaining endometrial swabs for cytology, microbial culture, and antimicrobial sensitivity is an important component of treatment. Uterine fluid siphoning and lavage is necessary to reduce purulent material (Figure 274.3). This may require breaking down scar tissue and occlusions of the lumen. Preventing reformation of transluminal adhesions can be aided by a physical obstruction such as a hollow pessary or gauze pack.3 Topical application of antimicrobial and anti-inflammatory ointments may be beneficial in adhesion prevention. Administration of exogenous PGF2α for luteolysis and myometrial contractility may aid in uterine evacuation once it is determined that there are no occlusions to outflow.1 Ovariohysterectomy can be performed to resolve chronic pyometra. Reported postoperative complications include reduced fecal output, decreased intesti-
1. Van Camp SD. Breeding soundness examination of the mare and common genital abnormalities encountered. In: Morrow DA (ed.) Current Therapy in Theriogenology. Philadelphia: W.B. Saunders Co., 1986; pp. 654–61. 2. Threlfall WR, Carleton CL. Treatment of uterine infections in the mare. In: Morrow DA (ed.) Current Therapy in Theriogenology. Philadelphia: W.B. Saunders Co., 1986; pp. 730–7. 3. Roberts SJ. Veterinary Obstetrics and Genital Diseases (Theriogenology), 3rd edn. Woodstock: Stephen J. Roberts, 1986. 4. Hughes JP, Stabenfeldt GH, Kindahl H, Kennedy PC, Edqvist L-E, Neely DP, Schalm OW. Pyometra in the mare. J Reprod Fertil Suppl 1979;(27):321–9. 5. Causey RC. Making sense of equine uterine infections: the many faces of physical clearance. Vet J 2006;172:405–21. 6. Vandeplassche M, Spincemaille J, Bouters R. Advanced pyometra with intact endometrial cups in a mare. Equine Vet J 1979;11:112–13. 7. Kennedy PC, Miller RB. The uterus. In: Jubb KVF, Kennedy PC, Palmer N (eds) Pathology of Domestic Animals, 4th edn, vol. 3. 1993; pp. 372–87, Academic Press, Inc. 8. Hughes JP, Loy RG, Asbury AC, Burd HE. The occurrence of Pseudomonas in the reproductive tract of mares and its effect on fertility. Cornell Vet 1966;56:595–610. 9. Abou-Gabal M, Hogle RM, West JK. Pyometra in a mare caused by Candida rugosa. J Am Vet Med Assoc 1977;170: 177–8. 10. Watson ED. Release of immunoreactive arachidonate metabolites by equine endometrium in vitro. Am J Vet Res 1989;50:1207–9. 11. Fortier MA, Guilbault LA, Grasso F. Specific properties of epithelial and stromal cells from the endometrium of cows. J Reprod Fertil 1988;83:239–48. ¨ 12. Rotting AK, Freeman DE, Doyle AJ, Lock T, Souberli D. Total and partial ovariohysterectomy in seven mares. Equine Vet J 2004;36:29–33.
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Chapter 275
Treatment of Fluid Accumulation Jonathan F. Pycock
Fluid accumulation in the uterus is the most common cause of reproductive failure in mares, particularly older mares and mares being bred for the first time in their teenage years. This subfertility is due to an unsuitable environment within the uterus for the developing conceptus, and in some instances the ensuing endometritis persists and causes early regression of the corpus luteum. In addition, uterine fluid may damage the ability of spermatozoa to survive in the uterus or oviduct, or even to fertilize once in the oviduct.
Pathogenesis Semen is deposited directly into the uterus when mares are bred by natural service or artificial insemination. This means that bacterial and seminal components, as well as debris, contaminate the uterus, which results in uterine inflammation. It was previously thought that the inflammatory response to breeding was due to bacterial contamination of the uterus at the time of mating. It is now accepted that the spermatozoa themselves, rather than bacteria, are responsible for the acute inflammatory response in the equine uterus after insemination.1 Spermatozoa are transported rapidly to the oviducts, but only a small number actually reach the site of fertilization. The majority of the ejaculate remains in the uterus and induces an acute inflammatory response. Kotilainen and others2 showed that the intensity of the reaction was dependent on the concentration and/or volume of the inseminate: concentrated semen (e.g., frozen semen) induced a stronger inflammatory reaction in the uterus than fresh or extended semen. The inflammatory reaction of the uterus is not different for live or dead spermatozoa.3
Parlevliet and co-workers4 measured the inflammatory response following insemination with raw semen, extended semen, and various extenders. They concluded that the intensity of the inflammatory response following insemination depends on the sperm themselves rather than any extender. Seminal plasma is the liquid portion of the ejaculate and suspends the cells present in semen. The precise role of seminal plasma in the horse is not known. Seminal plasma improves sperm transport, either directly or by protecting the sperm from phagocytosis while in the uterus, thereby improving survival time.5, 6 Seminal plasma is rich in oxytocin and prostaglandins, both of which cause uterine contractions. Seminal plasma prolongs sperm life within the uterus because it suppresses neutrophil (PMN) chemotaxis and phagocytosis of sperm.7 These latter authors concluded that the mechanism is thought to involve more than simply complement inactivation. Seminal plasma modulates the endometrial immune response, either beneficially8 or adversely.9 A reduction in the inflammatory response after insemination would have positive and negative effects. Any suppression of phagocytosis would reduce not only the elimination of sperm, but also bacteria. The amount of fluid produced after insemination may be reduced as a result of this suppression. Seminal plasma has a low buffering capacity and, in a freshly collected ejaculate, by-products rapidly accumulate in seminal plasma due to the high metabolic activity of the sperm cells. During freezing of sperm, seminal plasma is removed. This may be the reason that the inflammatory response is greater with frozen–thawed semen. This increased response may also be due to the high sperm concentration.2
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: Disease of the Uterus
The first inflammatory cells to enter the uterus are neutrophils (PMNs), observed in the uterus within 30 minutes after insemination.10 The equine uterus produces fluid which has chemoattractant properties for PMNs.11 Complement is thought to be the crucial component which serves as a chemoattractant for PMNs, although other inflammatory products may serve as chemoattractants.11, 12 It is now believed that spermatozoa initiate chemotaxis of PMNs through activation of complement,13 suggesting that a transient uterine inflammation following insemination is physiological and serves to clear the uterus of excess spermatozoa, seminal plasma, and contaminates associated with insemination. Damaged sperm may be more readily phagocytosed by PMNs. In normal mares, the inflammation peaks around 10–12 hours and reduces within 24–36 hours after insemination. In most mares this transient endometritis resolves spontaneously within 24–48 hours, so that the environment of the uterine lumen is compatible with embryonic and fetal life. It is important not to regard this endometritis as a pathological condition. Rather it is a physiological reaction to clear excess sperm, seminal plasma, and inflammatory debris from the uterus before the embryo descends from the oviduct into the uterine lumen 5.5 days after fertilization. However, if the endometritis persists after day 4 or 5 of diestrus, in addition to being incompatible with embryonic survival, the premature release of PGF2α results in luteolysis, a rapid decline of progesterone, and an early return to estrus. These mares are referred to as susceptible and they develop a persistent endometritis.14 Reduced resistance to endometritis is associated with advancing age and multiparity. Susceptibility to endometritis is not an absolute state since failure of uterine defence mechanisms need only slow the process of eliminating infection. In the practice situation, a wide range of susceptibility to endometritis is seen and it must not be thought that mares can be neatly packaged into ‘resistant’ or ‘susceptible’.15 Studies on immunoglobulins, opsonins and the functional ability of neutrophils in the uterus of susceptible mares have not confirmed the presence of an impaired immune response.16 Evans and co-workers17 first suggested that reduced physical drainage may contribute to an increased susceptibility to uterine infection. The physical ability of the uterus to eliminate bacteria, inflammatory debris, and fluid is now known to be the critical factor in uterine defense. It is a logical conclusion that any impairment of this function, for example, defective myometrial contractility, renders a mare susceptible to persistent endometritis.18–20 The reason susceptible mares have this defective contractility is not known. It has been suggested that the regulation of muscle contraction by the nervous system may be impaired.21
The resulting fluid accumulation could also be due to failure to drain via the cervix, or decreased reabsorption via lymphatic vessels. Lymphatic drainage could play an important role in the persistence of post-breeding inflammation and it is interesting that lymphatic lacunae (lymph stasis) is a common finding in endometrial biopsies taken from susceptible mares.22, 23 Other factors can be involved in delayed uterine clearance. Older mares that have had several foals can develop a uterus that is dropped ventrally within the abdomen, and such mares can have a delay in uterine clearance.24 The cervix is vital to allow drainage and any failure of the cervix to dilate can cause fluid accumulation. This failure of the cervix to open is particularly relevant in the case of the older mare, in general, and the older maiden mare, in particular, leading to the term ‘the old maiden mare syndrome’. It is particularly important to recognize and manage appropriately the older maiden mare, as in many cases these mares are susceptible to post-breeding endometritis even though they have never been bred before. Often sport or Warmblood mares may not be presented to be bred until they are in their teens, and these older maiden mares can be very difficult to get in foal. Many of these mares have some common characteristics which resemble a syndrome. Endometrial biopsy samples reveal a degree of glandular degenerative changes and stromal fibrosis (endometrosis) as an inevitable consequence of aging, despite the fact they have not been bred.25 Another of the most common characteristics of this syndrome is uterine fluid. Often an older maiden mare has an abnormally tight cervix which fails to relax properly during estrus, so that fluid is unable to drain and accumulates in the uterine lumen prior to mating.26 In many cases this fluid is negative for bacterial growth and presence of neutrophils. Once the mare is bred the fluid accumulation will be aggravated due to poor lymphatic drainage and impaired myometrial contraction, compounded by the tight cervix. The amount of intrauterine fluid will vary in individual mares, ranging from a few mL to over a liter in extreme cases. To maximize the fertility of these mares it is vital that the veterinarian is aware of the possibility of this type of uterine and cervical pathology. All too often, owners assume that the fertility of these mares is comparable to that of young maiden mares; one of the most important aspects of breeding the old maiden mare is to make the owner aware that there is a high possibility that she will be a problem. These mares must be considered highly susceptible and managed accordingly. Endometrial vascular degeneration has also been considered as contributing to a delay in uterine clearance.27 The difference between resistant and susceptible mares has been reviewed recently by Causey.28 This
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Treatment of Fluid Accumulation author has speculated on the influence of mucociliary activity on the uterine defense mechanism.29 Causey indicated that mucus plays a mechanical role in the defense mechanism of organ systems such as the respiratory system. Mucus traps particles and carries them long distances by ciliated cells. However, for mucus to have the proper activity, it must be a continuum with adequate elasticity and viscosity. Mucus overhydration will disrupt the elasticity and viscosity, breaking that continuum. Since the endometrium has mucus-producing and ciliated cells, it is not inconceivable that that mucus could be involved in the mechanism of uterine defense. So a speculative pathogenesis of the disruption in mucociliary defense mechanism could start with a poor perineal conformation, disturbance of myoelectrical contractility, and/or cervical incompetence. These anatomical problems would lead to fluid accumulation, uterine dilatation, and mucus overhydration. Furthermore, it is known that mucus elasticity and viscosity can be affected by the type of bacteria. For example Streptococcus S. zooepidemicus decreases mucus viscosity, breaking the continuum, while Klebsiella pneumoniae increases mucus viscosity, thereby decreasing the ability of cilia to move the mucus.28
Identification of the fluid-producing mare This can be difficult as there may only be subtle changes in the uterine environment, not readily detected by current diagnostic procedures. Many mares show no signs of inflammation before mating, but will fail to resolve the inevitable endometritis which follows mating. Response to bacterial challenge has been used in a research setting; however, in practice, history is perhaps the most useful indicator of a susceptible mare. Demonstration of clearance failure using scintigraphic and other methods based on charcoal clearance have been used to make an accurate diagnosis that a mare has a clearance problem,20 but they are difficult to apply in practice. Ultrasonographic examination to detect uterine luminal fluid (Figure 275.1) has proved useful in identifying mares which accumulate fluid and is the most useful technique in practice.30 The presence of free intraluminal fluid before breeding strongly suggests susceptibility to persistent endometritis.31 Rasch and co-workers32 postulated that excessive production of fluid due to glandular alterations, rather than a failure of lymphatic drainage, caused intrauterine fluid accumulation. It is currently not known for certain, however, whether the fluid accumulates as a result of excess production, a delay in physical clearance via the open cervix, or decreased
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Figure 275.1 Ultrasonographic image of the uterus of a mare with fluid accumulation.
reabsorption by the lymphatic system; a combination of all three mechanisms may well be involved. Since the first description of the identification, using ultrasound, of small volumes of intrauterine fluid that could not be palpated per rectum,33 general awareness of the frequency of this abnormality has increased. The detection of uterine fluid during both estrus and diestrus has been reported.14 Endometrial secretions and the formation of small volumes of free fluid may be associated with the same mechanism that causes normal estral edema. In many cases, the uterine luminal fluid that accumulates before mating is sterile and contains no neutrophils.31 The importance of these sterile fluid accumulations is that, although initially sterile, the fluid may act as a culture medium for bacteria that gain entry to the uterus at mating to multiply and may be spermicidal. The amount of fluid that should be considered significant is not clear, and it may be that the quantity of fluid is more important than the nature of the fluid. This is particularly true of fluid appearing during estrus. The significance depends to some extent on when during estrus the fluid is observed: fluid detected early in estrus may have disappeared when the mare is further advanced in estrus and the cervix relaxes more. Small volumes of intrauterine fluid during estrus do not affect pregnancy rates, in contrast to mares with larger (>2 cm depth) collections of fluid.31 In mares that are susceptible to endometritis, there is an accumulation of more fluid than in resistant mares. Generally, if there is more than 1 cm of fluid during estrus, some attempt should be made to remove this before breeding using oxytocin. If the volume is more than 2 cm, the fluid may need to be drained and investigated for the presence of inflammatory cells and bacteria. The mare may then need to have a large-volume uterine lavage. Intrauterine fluid during diestrus is indicative of inflammation and is
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(a)
(b)
(c)
(d)
Figure 275.2 (a) Ultrasonographic image of grade 1 uterine fluid: anechogenic; (b) ultrasonographic image of grade 2 uterine fluid: hypoechogenic with hyperechogenic particles; (c) ultrasonographic image of grade 3 uterine fluid: moderately echogenic; (d) ultrasonographic image of grade 4 uterine fluid: hyperechogenic fluid in the uterus of an infected mare.
associated with subfertility as a result of early embryonic death (EED) and a shortened luteal phase.31 Intraluminal uterine fluid can be graded 1 to 4 according to the degree of echogenicity (Figure 275.2). The more echogenic the fluid, the more likely the fluid is contaminated with debris, including white blood cells. However, cellular fluid can appear relatively anechogenic, so care is needed in interpretation. Inspissated pus can be so echogenic that it lacks contrast to the uterus and is overlooked. The actual appearance of the fluid and the ultrasonographic appearance might not be as closely linked as was once thought. Ultrasonographic appearance may be proportional to the size and concentration of particulate matter within the fluid, rather than the viscosity of the fluid; for example, purulent exudates can appear not as echogenic as expected. Air has hyperechogenic foci, and fluid with air bubbles appears cellular. Urine in the bladder can appear echogenic, despite being a watery liquid (Figure 275.3). Ultrasonographic appearance of urine varies according to diet. Diets high in protein (lucerne and clover) produce echogenic urine from secretions. Grass diets are relatively anechoic.
Treatment options for mares with fluid accumulation The aim of treatment for mares with fluid accumulation should be to assist the uterus to physically clear the debris, contaminants, and excess sperm as well as the normal inflammatory by-products of the response
Figure 275.3 Ultrasonographic image of a full urinary bladder.
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Treatment of Fluid Accumulation to breeding.34–37 Because the spermatozoa necessary for fertilization are present within the oviduct within 4 hours of mating,34 and because the embryo does not descend into the uterus for about 5.5 days, mares may safely undergo treatment from 12 hours after mating until 2 days from ovulation, as long as isotonic solutions are used. The studies of Brinsko and coworkers34 were on normal mares, and sperm transit time in susceptible mares may be different. Progesterone concentrations increase rapidly after ovulation in the mare and it is preferable to avoid treatment involving uterine interference beyond 2 days after ovulation. Both coitus and artificial insemination can be a source of uterine contamination; it is well known that spermatozoa themselves are responsible for initiating a marked inflammatory response. The successful management of susceptible mares should logically require some form of post-mating therapy such as intrauterine antibiotic infusion, uterine lavage, and intravenous oxytocin; these may be used alone or in combination. The emphasis should be on treatment in relation to breeding and not ovulation (for as long as 2 days after ovulation). Too often in the past, veterinarians have waited until ovulation before treating these mares. By then, there has usually been a large accumulation of fluid and the bacteria are in a logarithmic phase of growth. Uterine lavage Recognition of the importance of the mechanical evacuation of uterine contents accounted for the introduction of large-volume uterine lavage. The technique involves the mechanical suction or siphonage of 2 to 3 L of previously warmed (to 42◦ C) sterile physiological (buffered) saline or lactated Ringer’s solution infused into the uterus via a catheter that has been retained within the cervix via a cuff. The most convenient is a large-bore (30 French) (80 cm) autoclavable equine embryo flushing catheter (EUF80, Bivona, IN) (Figure 275.4). The cuff is useful because it effectively seals the internal cervical os. The catheter should be inserted only after thorough cleansing of the perineum. Such an approach accomplishes the following:
r r r
Removal of accumulated uterine fluid and inflammatory debris, which may interfere with neutrophil function and the efficacy of antibiotics; Stimulation of uterine contractility; Recruitment of fresh neutrophils through mechanical irritation of the endometrium.
The fluid is infused by gravity flow, 1 L at a time, and the washings inspected to provide immediate
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Figure 275.4 Large-bore embryo flushing catheter, 80 cm in length. Note the inflated cuff.
information concerning the nature of the uterine contents. The lavage should be repeated until the fluid recovered is clear. In most cases, the fluid evenly distributes in both horns, making transrectal massage of the uterus unnecessary. If a rectal examination is performed while the catheter is in the uterus, the veterinarian should be very careful to avoid contaminating the catheter. The fluid should be recovered into the same container in which it was infused, thereby preventing air from being aspirated into the uterus via the catheter. Measurement of the recovered fluid and ultrasonographic examination of the uterus should be performed after flushing, to ensure that all the fluid has been recovered. This is necessary because the mare has an impaired ability to spontaneously drain the uterus. For this reason, the process is usually combined with oxytocin injection. Ideally, these mares are bred only once, but if repeated matings are necessary, uterine lavage should be performed after each mating. Uterine lavage with isotonic buffered solutions, performed immediately before mating, does not adversely affect fertility.38 Large-volume lavage is beneficial in many cases, particularly in the mare with a relatively large accumulation of fluid (>2 cm depth) after breeding. The process is time consuming, and there is the possibility of further contamination of the uterus by passage of a drainage tube. Nonetheless, when there is more than 2 cm of uterine fluid or a mare is known to be highly susceptible, the risks are outweighed by the benefit of treatment.39 It has been shown that saline lavage and uterotonic drugs such as PGF2α are as effective as antibiotics in eliminating bacteria from the uterus.40 However, this was a controlled experimental type situation in which a single bacterial species was infused into the uterus, and lavage was within 12 hours of mating. Under normal clinical conditions, there is a mixed bacterial contamination and lavage cannot always be performed within 12 hours. I prefer to
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continue to use intrauterine antibiotics as part of the therapeutic protocol.
the mare as 6.8 minutes.47 In most mares, the response is rapid, with fluid being voided almost immediately.
Oxytocin The ideal method of treatment involves the use of a non-invasive technique with early and complete elimination of any intrauterine fluid. Oxytocin stimulates uterine contractions in the cyclic, pregnant, and postpartum mare and was first suggested to promote uterine drainage in mares with defective uterine clearance by Allen.41 Until then, oxytocin was not considered to be an appropriate treatment for endometritis, probably because it was thought by many that oxytocin induced uterine contractions only in the first 48 hours after foaling. Also, use of oxytocin was discouraged because of the worry that it would cause severe colic. However, after the pioneering study of Allen,41 subsequent clinical experience32, 35, 36, 42, 43 has allayed early fears that oxytocin would cause colic when given as an intravenous bolus. All these workers reported improved pregnancy rates in susceptible mares after oxytocin administration. Oxytocin therapy should be used wherever free uterine fluid is detected before or after breeding. I give a dose of oxytocin of 25 international units (IU) for large mares (>600 kg) and 20 IU for average size Thoroughbred, Standardbred, Quarterhorse, Warmblood mares and all pony mares either intravenously or intramuscularly. There is no advantage in using more than 25 IU, and it may even be that the same effect can be obtained by use of 10 IU as long as treatment is occurring during the preovulatory period.44 Paccamonti and co-workers44 demonstrated that the initial change in intrauterine pressure appeared more pronounced with a larger (25 IU) dose of oxytocin at 5 and 10 minutes after treatment; the change in pressure at subsequent time intervals was similar between the two doses (25 IU and 10 IU). These authors also found a significant effect of day of administration of oxytocin on the uterine response, with a weaker response when oxytocin was administered after ovulation. Later work by these authors confirmed that myometrial activity and the increase in intrauterine pressure after exogenous oxytocin are inversely related to the concentration of progesterone.45, 46 Although it is preferable to treat mares with uterine clearance problems before ovulation, when using oxytocin after ovulation the larger dose of 25 IU should be used for all mares. No untoward effects have been noted, except for rare, mild and transient discomfort. Any possible adverse effect on gamete transport seems to be outweighed by the ability to improve uterine fluid clearance. The half-life of oxytocin has been reported for
Long-acting oxytocin Recently a long-acting synthetic analog of oxytocin, carbetocin, has become commercially available in Europe, Canada, and Mexico. This drug may have an indication in situations where a more prolonged uterine contraction is desired. In a recent study by Schramme and co-workers,48 carbetocin was well tolerated following intravenous administration of 175 µg to a group of horses. These authors reported the half-life of carbetocin to be 17 minutes, only some 2.5 times that of oxytocin. An earlier study had shown a more prolonged action when compared with oxytocin, with maximum concentrations of carbetocin in plasma at 20 minutes after intramuscular administration of 280 µg compared with 10 IU oxytocin given intravenously.49 Preliminary clinical work by the author has shown carbetocin to be safe and effective at inducing uterine clearance. Two intramuscular injections of 175 µg carbetocin are given 12 and 24 hours after breeding in mares with marked uterine edema or free fluid before breeding, or in mares with uterine fluid greater than 2 cm in depth when examined 12 hours after breeding. Oxytocin remains the drug of choice in mares that accumulate free intraluminal uterine fluid and is my first-choice ecbolic drug. Oxytocin is known to stimulate prostaglandin release,20 although the precise mechanism that controls myometrial contraction remains unclear. The uterine response to oxytocin is due in part to direct effect of oxytocin on the myometrium and in part to the indirect effect of oxytocin causing PGF2a release.47, 50 Support for this dual mode of action comes from the fact that oxytocin has been shown to override the inhibitory effect of phenylbutazone on uterine clearance.51 Phenylbutazone inhibits prostaglandin synthesis and has been shown to slow uterine clearance in reproductively normal mares.50 Phenylbutazone is a commonly used drug in brood mares for conditions such as laminitis, and oxytocin would be preferable to PGF2α analogs to improve uterine clearance in mares on phenylbutazone therapy. Even reproductively normal mares receiving phenylbutazone because of lameness should be given oxytocin after breeding. Prostaglandin analogs Due to the relatively short half-life of oxytocin, ecbolic drugs with a longer duration of action have
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Treatment of Fluid Accumulation been investigated. Endogenous prostaglandins are released very early in mares with endometritis.12 The useful role of prostaglandin in increasing myometrial activity and assisting uterine clearance has subsequently been shown.40, 52, 53 These latter authors showed that the prostaglandin analog, cloprostenol, given at a dose rate of 500 µg intramuscularly, caused increased clearance of radiocolloid in susceptible mares, but at a significantly slower rate and with a delayed increase in uterine activity than that caused by oxytocin. However, the uterus did contract for a longer time, 5 hours versus 45 minutes. Of the prostaglandins administered (PGF2α , cloprostenol, and fenprostalene), cloprostenol produced the most consistent response. Cloprostenol seems to be indicated in mares with lymphatic stasis as shown by excessive fluid within the endometrium, or large lymphatic cysts.51 Uterine edema often persists postovulation in these mares. The suggested dose of cloprostenol is 250 µg given 12 and 24 hours after breeding.27 When using prostaglandin analogs for the treatment of post-mating-induced endometritis, the time of administration relative to ovulation must be considered. It is accepted that the equine corpus luteum is resistant to the luteolytic effect of prostaglandin until five or six days post-ovulation, but this resistance is not absolute. Progesterone concentrations have been reported to be lower in mares for 5 to 7 days following administration of cloprostenol in the post-ovulatory period compared with mares receiving oxytocin, saline, or sterile water.54, 55 However, another study by Nie and co-workers56 found that cloprostenol could be used to treat post-breeding mares through the second day following ovulation without decreasing pregnancy outcome. This was despite finding an impaired luteal function, as evidenced by lower circulating progesterone concentrations. These authors used a dose of 250 µg cloprostenol. Despite this, prostaglandin analogs are best avoided after ovulation. Intrauterine plasma infusions Based on the research findings of the 1970s and 1980s, which emphasized the immunological aspects of the uterine defense mechanisms, intrauterine plasma has been used in the susceptible mare. Studies following its use have indicated an improvement in fertility.57, 58 Both authors suggested that the plasma had an enhancing effect on phagocytosis by uterine neutrophils. Adams and Ginther,59 in a study that included control groups of mares, found that intrauterine plasma was not efficacious in treating endometritis because there was no improvement in pregnancy rates. In addition, transfer of infectious
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agents is also possible. Troedsson and co-workers52 suggested that plasma treatment might only benefit certain susceptible mares. This latter point was also alluded to by Pascoe,58 who, while remaining enthusiastic about the use of plasma in the management of immune-incompetent mares, conceded that this may apply only to mares without a mechanical clearance problem. Consequently, plasma is best used in mares that repeatedly fail to become pregnant but have no history of fluid accumulation. Because mares that are susceptible to endometritis do not possess a quantitative deficiency of immunoglobulins, it is questionable whether such treatment is truly effective.
Intrauterine antibiotics The infusion of broad-spectrum antibiotics after breeding is controversial.13, 31, 42 It has been shown in at least one scientific experiment, in which an endometritis model was used, that saline lavage and uterotonic drugs such as PGF2α are as effective as antibiotics in eliminating bacteria from the uterus.40 However, as mentioned previously, a single bacterial species (usually Streptococcus zooepidemicus) was infused into the uterus, and lavage treatment was performed at a fixed time (within 12 hours of mating). This is not truly representative of normal clinical conditions where there is mixed bacterial contamination and lavage cannot always be performed within 12 hours. This is why the author prefers to continue to use intrauterine antibiotics as part of the therapeutic protocol. The treatments also have entirely different modes of action on the uterus, often resulting in an additive benefit. The uterine lavage provides mechanical evacuation of intraluminal fluid and inflammatory products, whereas the single infusion of broad-spectrum antibiotics should prevent any growth of bacteria introduced during insemination or uterine lavage. Furthermore, mares susceptible to post-mating-induced endometritis have, according to their history, a reduced capacity for uterine clearance and reduced defense mechanisms. Therefore, they cannot be compared with healthy mares used in experiments. Routine administration of intrauterine antibiotics has no negative effect on pregnancy rate of treated mares.31, 42
Systemic antibiotics There is a paucity of information in the literature about the use of systemic antibiotics in the management of the mare susceptible to post-mating-induced endometritis, and their use is not widespread. In certain situations, systemic administration of
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antibiotics may be useful, having better distribution in the uterus compared with intrauterine application. Such situations include where it is desired to continue administration after ovulation into the luteal phase due to persistence of fluid, or recent urogenital surgery making intrauterine administration of antibiotics difficult. Antibiotics that have been used systemically include the following: procaine penicillin, gentamicin, amikacin, ampicillin and trimethoprim sulpha,60 and ceftiofur.61
Corticosteroids The antigenic nature of spermatozoa in the uterus is well established and the post-breeding reaction is inflammatory in nature, making it logical that corticosteroids may play a role in modulating the response. The precise nature of the damage caused by this inflammatory response is unknown. It has been reported that seminal plasma has a suppressive effect on complement activation and PMN chemotaxis.8 Recently, a group from Brazil has described how the corticosteroid prednisolone modified the uterine response to insemination.62 The corticosteroid therapy (prednisolone acetate) at a dose rate of 0.1 mg/kg was given every 12 hours for a total of five treatments. Four treatments were given before insemination and one treatment at the time of insemination. They concluded that prednisolone treatment led to a reduction in neutrophil function and decreased uterine fluid content. Further work by Papa and co-workers63 in Brazil indicated corticosteroid therapy to be a safe option for mares with a history of post-mating-induced endometritis, but concluded that further clinical research was necessary. A single dose of dexamethasone administered to mares at the time of breeding is safe, and may beneficially modulate the inflammatory response to breeding.64 Other clinicians are using corticosteroids in certain susceptible mares but no further reports have been published at present.
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References 20. 1. Katila T. Sperm-uterine interactions: a review. Anim Reprod Sci 2001;68:267–72. 2. Kotilainen T, Huhtinen M, Katila T. Sperm induced leukocytosis in the equine uterus. Theriogenology 1994;41:629–36. 3. Katila T. Neutophils in uterine fluid after insemination with fresh live spermatozoa or with killed spermatozoa. Pferdeheilkunde 1997;13:54. 4. Parlevliet JM, Tremoleda IM, Cheng FP, Pycock JF, Colebrander B. Influence of semen, extender and seminal plasma on the defence mechanism of the mare’s uterus (abstract). Pferdeheilkunde 1997;13:540. 5. Alghamdi AS, Foster DN, Troedsson MHT. Equine seminal plasma reduces sperm binding to polymorphonuclear neu-
21.
22.
23.
trophils and improves fertility of fresh semen inseminated into inflamed uteri. Reproduction 2004;127:593–600. Troedsson MHT, Desvouges, A, Alghamd, Dahms B, Dow CA, Hayna J, Valesco R, Collahan PT, Macpherson ML, Pozor M. Components in seminal plasma regulating sperm transport and elimination. Anim Reprod Sci 2005;89:171– 86. Troedsson MHT, Franklin RK, Crabo BG. Suppression of PMN-chemotaxis by different molecular weight fractions of seminal plasma. Pferdeheilkunde 1999;115:568–73. Troedsson MHT, Lee CS, Franklin RD, Crabo BG. The role of seminal plasma in post-breeding uterine inflammation. J Reprod Fert Suppl 2000;56:341–9. Portus BJ, Reilas R, Katila T. Effect of seminal plasma on uterine inflammation, contractility and pregnancy rates in mares. Equine Vet J 2005;37:515–19. Katila T. Onset and duration of uterine inflammatory response of mares after insemination with fresh semen. Biol Reprod Mono 1995;1:515–17. Pycock JF, Allen WE. Pre-chemotactic and chemotactic properties of uterine fluid from mares with experimentally induced bacterial endometritis. Vet Rec 1988;123:193–5. Pycock JF, Allen WE. Inflammatory components in uterine fluid from mares with experimentally induced bacterial endometritis. Equine Vet J 1990;22:422–5. Troedsson MHT. Uterine response to semen deposition in the mare. In: Proceedings of the Society of Theriogenology, San Antonio, TX, 1995; pp. 130–5. Allen WE, Pycock JF. Cyclical accumulation of uterine fluid in mares with lowered resistance to endometritis. Vet Rec 1988;122:489–90. Pycock JF, Paccamonti D, Jonker H, Newcombe J, van der Weijden G. Can mares be classified as resistant or susceptible to recurrent endometritis? Pferdeheilkunde 1997;13:431–6. Allen WE, Pycock JF. Current views on the pathogenesis of bacterial endometritis in mares with lowered resistance to endometritis. Vet Rec 1989;125:298–303. Evans MJ, Hamer JM, Gason LM, Irvine CHG. Clearance of bacteria and non-antigenic markers following intrauterine inoculation into maiden mares: effect of steroid hormone environment. Theriogenology 1986;26:327–34. Troedsson MHT, Liu IKM. Uterine clearance of nonantigenic markers (51-Cr) in response to a bacterial challenge in mares potentially susceptible and resistant to chronic uterine infections. J Reprod Fert Suppl 1991;44:283–8. Troedsson MHT, Liu IKM, Ing M. Multiple site electromyography recordings of uterine activity following an intrauterine bacterial challenge in mares susceptible and resistant to chronic uterine infection. J Reprod Fert 1993;99:307–13. LeBlanc MM, Neuwirth L, Mauragis D, Klapstein E, Tran TQ. Oxytocin enhances clearance of radiocolloid from the uterine lumen of reproductively normal mares and mares susceptible to endometritis. Equine Vet J 1994;26: 279–82. Liu IK, Rakestraw P, Coit C, Harmon F, Snyder J. An in vitro investigation of the mechanism of neuromuscular regulation in myometrial contractility (abstract). Pferdeheilkunde 1997;13:557. Kenney RM. Cyclic and pathologic changes of the mare endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1978;172:241–62. LeBlanc MM, Johnson RD, Calderwood Mays MB, Valerama C. Lymphatic clearance of india ink in
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Treatment of Fluid Accumulation
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reproductively normal mares and mares susceptible to endometritis. Biol Reprod Mono 1995;1:501–6. LeBlanc MM, Neuwirth L, Jones, Cage C, Mauragis D. Differences in uterine position of reproductively normal mares and those with delayed uterine clearance detected by scintigraphy. Theriogenology 1998;50:49–54. Ricketts SW, Alonso S. The effect of age and parity on the development of equine chronic endometrial disease. Equine Vet J 1991;23:188–92. Pycock JF. Cervical function and uterine fluid accumulation in mares. Proceedings of JP Hughes International Workshop on Equine Endometritis, summarized by WR Allen. Equine Vet J 1993;(Suppl)25:191. LeBlanc MM. Persistent mating induced endometritis in the mare: pathogenesis, diagnosis and treatment. In: Ball BA (ed.): Recent Advances in Equine Reproduction. Ithaca, NY: International Veterinary Information Service ( www.ivis.org), 2003. Causey RC. Making sense of equine uterine infections: The many faces of physical clearance. Vet J 2006;172:405–21. Causey RC. Mucus and the mare: How little do we know. Theriogenology 2007;68:386–94. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproductive tract of mares after parturition: effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed first postpartum ovulatory cycles. J Am Vet Med Assoc 1988;192:350–3. Pycock JF, Newcombe JR. The relationship between intraluminal uterine fluid, endometritis and pregnancy rate in the mare. Equine Pract 1996;18:19–22. Rasch K, Schoon HA, Sieme H, Klug E. Histomorphological endometrial status and influence of oxytocin on the uterine drainage and pregnancy rate in mares. Equine Vet J 1996;28:455–60. Ginther OJ, Pierson RA. Ultrasonic anatomy and pathology of the equine uterus. Theriogenology 1984;21:505–6. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours postinsemination on pregnancy rates in mares. Theriogenology 1991;35:1111–19. Pycock JF. A new approach to treatment of endometritis. Equine Vet Educ1994;6:36–8. Pycock JF. Assessment of oxytocin and intrauterine antibiotics on intrauterine fluid and pregnancy rates in the mare. In: Proceedings of the American Association of Equine Practitioners 1994; pp. 19–20. Troedsson MHT. Therapeutic considerations for matinginduced endometritis. Pferdeheilkunde 1997;13:516–20. Vanderwall DK, Woods GL. Effect of fertility of uterine lavage performed immediately prior to insemination in mares. J Am Vet Med Assoc 2003;222:1108–1110. Knutti B, Pycock JF, van der Weijden GC, Kupfer U. The influence of early postbreeding uterine lavage on pregnancy rates in mares with intrauterine fluid accumulations after breeding. Equine Vet Educ 2000;12:267–70. Troedsson MHT, Scott MA, Liu IKM. Comparative treatment of mares susceptible to chronic uterine infection. Am J Vet Res 1995;56:468–72. Allen WE. Investigations into the use of oxytocin for promoting uterine drainage in mares susceptible to endometritis. Vet Rec 1991;128:593–94. Pycock JF, Newcombe JR. Assessment of the effect of three treatments to remove intrauterine fluid on pregnancy rate in the mare. Vet Rec 1996;138:320–3.
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43. LeBlanc MM. Oxytocin: The new wonder drug for treatment of endometritis? Equine Vet Educ 1994;6:39–43. 44. Paccamonti DL, Gutjahr S, Pycock JF, van der Weijden GC, Taverne MA. Does the effect of oxytocin on intrauterine pressure vary with dose or day of treatment (abstract). Pferdeheilkunde 1997;13:553. 45. Gutjahr S, Paccamonti D, Pycock JF, Taverne MAM, Dieleman SJ, Bevers M, Van Der Weijden GC. Intrauterine pressure changes in response to oxytocin application in mares. Reprod Domest Anim Suppl 1998;5:118. 46. Gutjahr S, Paccamonti DL, Pycock JF, Taverne MAM, Dieleman SJ, Van der Weijden GC. Effect of dose and day of treatment on uterine response to oxytocin in mares. Theriogenology 2000;54:447–56. 47. Paccamonti DL, Pycock JF, Taverne MAM, Bevers M, van der Weijden GC, Gutjahr S. PGFM response to exogenous oxytocin and determination of the half-Iife of oxytocin in nonpregnant mares. Equine Vet J 1999;31: 285–8. 48. Schramme AR, Pinto CR, Davis JL, Whitacre MD, Whisnant CS. Pharmacokinetics of carbetocin, a long-acting oxytocin analogue, following intravenous administration in horses. Theriogenology 2007;68:517–18. 49. Handler J, Hoffmann D. Weber F, Schams D, Aurich C. Oxytocin does not contribute to the effects of cervical dilation on progesterone secretion and embryonic development in mares. Theriogenology 2006;66:1397–404. 50. Cadario ME, Thatcher MJD, LeBlanc MM. Relationship between prostaglandin and uterine clearance of radiocolloid in the mare. Biol Reprod Mono 1995;1:495–500. 51. LeBlanc MM. Effects of oxytocin, prostaglandin and phenylbutazone on uterine clearance of radiocolloid. Pferdeheilkunde 1997;13:483–5. 52. Troedsson MHT, Scott MA, Liu IKM. Pathogenesis and treatment of chronic uterine infection. In: Proceedings of the American Association of Equine Practitioners 1992; pp. 595–600. 53. Combs GB, LeBlanc MM, Neuwirth L, Tran TQ. Effects of prostaglandin F2α, cloprostenol and fenprostalene on uterine clearance of radiocolloid in the mare. Theriogenology 1996;45:1449–55. 54. Troedsson MHT, Ababneh MM, Ohlgren AF, Madill S, Vetscher N, Gregas M. Effect of periovulatory prostaglandin F2alpha on pregnancy rates and luteal function in the mare. Theriogenology 2001;55:1891–9. 55. Brendemuehl JP. Effect of oxytocin and cloprostenol on luteal formation, function and pregnancy rates in mares. Theriogenology 2002;58:623–6. 56. Nie GJ, Johnson KE, Wenzel JG, Braden TD. Effect of administering oxytocin or cloprostenol in the periovulatory period on pregnancy outcome and luteal function in mares. Theriogenology 2002;60:1111–18. 57. Asbury AC. Uterine defence mechanisms in the mare: the use of intrauterine plasma in the management of endometritis. Theriogenology 1984;21:387–93. 58. Pascoe DR. Effect of adding autologous plasma to an intrauterine antibiotic therapy after breeding on pregnancy rates in mares. Biol Reprod Mono 1995;1:539–43. 59. Adams GP, Ginther OJ. Efficacy of intrauterine infusion of plasma for treatment of infertility and endometritis in mares. J Am Vet Med Assoc 1989;194:372–8. 60. Asbury AC, Lyle SK. Infectious causes of infertility. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 381–91.
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61. Jonker FH. Secretion of ceftiofur in equine endometrium after parenteral administration. J Vet Pharmacol Ther 1997;Suppl. 20:37. 62. Dell’Aqua Jr. JA, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Melo CM. Modulation of acute uterine inflammatory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94: 270–3.
63. Papa FO, Dell’Aqua Jr. JA, Alvarenga MA, Melo CM, Zahn FS, Lopes MD. Use of corticosteroid therapy on the modulation of uterine inflammatory response in mares after artificial insemination with frozen semen. Pferdeheilkunde 2008;24:79–82. 64. Bucca S, Carli A, Buckley T, Dolci G, Fogarty U. The use of dexamethasone administered to mares at breeding time in the modulation of persistent mating induced endometritis. Theriogenology 2008;70:1093–100.
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Chapter 276
Uterine Cysts Mary E. Stanton
Uterine cysts are a common finding in aged broodmares. While uterine cysts may be incidental or transient in many mares, they may also represent underlying uterine pathology that will negatively impact fertility. The physical presence of cysts may also complicate pregnancy diagnosis and successful conceptus mobility and placentation. It is estimated that 55% of subfertile mares over the age of 10 years have intrauterine cysts, a statistic that suggests the economic impact these cystic lesions have on the breeding industry.1–3 The time and expense of breeding during additional cycles or barren seasons due to early embryonic death and misdiagnosis of pregnancy negatively affect the profit realized from the mare’s offspring. The presence of intrauterine cysts may be an indicator of underlying uterine pathology in the form of fibrosis and alterations in muscular contractility.4 The significance of these cysts is dependent upon their number and location within the uterus1 . They may occur throughout the uterus, but tend to occur at the base of the uterine horns near the body of the uterus.5–8 This may present challenges during early pregnancy by impeding conceptus mobility through the uterus.5,9–11 Cystic areas of the uterus are not capable of producing histiotroph to provide nutritional support for the embryo during its development prior to placental formation. In the event that the embryo implants adjacent to a cyst there may be insufficient uterine contact to allow normal embryo development, with consequent early embryonic death10 (Figure 276.1). On ultrasonographic examination cysts may have the same size and shape as an early pregnancy. This can make pregnancy diagnosis a challenge. Accurate record-keeping and mapping the size and location of cysts prior to breeding the mare can help to reduce diagnostic errors12 (Figure 276.2).
Description and etiology Uterine cysts are categorized according to their size and location within uterine tissue. Glandular cysts are less than 1 cm in diameter and may occur during pregnancy and in the periparturient period as a normal finding.1 They form as a result of layers of fibrocytes surrounding the glands and causing cystic distension. These cysts are generally not pathological with regard to fertility. It is proposed that the cysts occur as a result of increased estrogen levels similar to the cystic endometrial hyperplasia complex that is seen in canines.1 Lymphatic cysts, also called lymphatic lacunae, are greater than 1 cm and sometimes measure up to 20 cm in diameter.1, 5 These cysts are of sufficient size to interfere with pregnancy. They arise as dilated areas of lymphatic tissue resulting from uterine fibrosis and a lack of myometrial contractility to move lymph through the duct system.2 Often these mares will have a poor uterine biopsy score reflecting the fibrotic changes.13 Lymphatic lacunae may also occur as a result of poor drainage due to gravity in a large gravid or postpartum uterus.7 The location of the cyst within the layers of the uterine wall affects the pathological significance of the lesion. Typically, luminal cysts may interfere with conceptus migration and placentation. There is no microcotyledon development on the endometrium or allantochorion in cystic regions of the gravid uterus.9 Numerous large cysts may decrease placental exchange and may potentially compromise enough to cause abortion. Mural cysts are contained within the deeper layers of the uterus and may have an effect on placentation and histiotroph secretion as a result of glandular disruption but are unlikely to interfere with movement of the embryo.9 A third type of cyst is seen more rarely. Transmural cysts involve
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Diagnosis
Figure 276.1 Uterine cyst adjacent to twin embryos.
the muscular layers of the uterus extending to the serosa. These may develop from extensive lymphatic lacunae or potentially as a result of a foaling injury. These structures may present a risk for the mare to carry future foals due to compromised integrity of the muscular layers of the uterus.14 Grossly, cysts have either a smooth, thin wall or an irregular, roughened fibrous capsule.1, 2, 6 The cyst capsule is opaque and white, but necrotic cysts are yellow to grayish blue.1, 7 They can occur as single structures or larger multicompartmental cysts that are divided by septa.1, 2, 7 They are round or cylindrical. Typically, cysts contain a watery yellow lymphatic fluid.1 When viewed endoscopically cystic tissue appears pale when compared to the surrounding endometrium. Histologically the cysts are lined by columnar epithelium and lamina propria. They may also contain normal or atrophied endometrial glands. Within the cyst wall there is also an inner lining of cuboidal cells and hyaline.3, 7, 9
Uterine cysts may be diagnosed by manual palpation, ultrasonography, or endoscopy. Larger uterine cysts may be detected manually by encircling the uterine horn with the thumb placed dorsally starting at the tip of the horn and working toward the body of the uterus. It is often easier to detect cysts manually if the mare is examined during estrus when uterine tone is decreased.4 Large lymphatic lacunae with a flabby uterus tend to have a poor pregnancy prognosis without treatment.15 Another manual technique for diagnosis involves sterile preparation of the mare and simultaneous intrauterine examination with a hand covered by a sterile sleeve and transrectal palpation to determine the location and size of the cysts. This may only be safely accomplished during estrus when the cervix is relaxed enough to accommodate the examiner’s hand.4 More commonly cysts are detected during ultrasonographic examination of the uterus. The capsule of the cyst appears as a definitive hyperechoic line surrounding a hypoechoic center. Cysts may appear circular or oblong; they are solitary or may appear clumped with hyperechoic septa dividing the cyst into compartments. Contrast ultrasonography may be used in order to ascertain the specific location of the cysts. The mare is prepared with an aseptic uterine infusion of 1–2 L of saline or lactated Ringer’s solution and then transrectally examined by ultrasonography. This will allow the examiner to determine whether the cysts are located dorsally or ventrally in the uterus and distinguish whether the cysts are located in the lumen or wall of the uterus. The cysts can be mapped on a uterine diagram with the specific location and size so that the information is available for treatment or to help avoid confusion between a cyst and an embryo at the time of pregnancy diagnosis.12 It is helpful to measure and diagram the cysts when examining the mare for ovulation after breeding in order to have a current reference.
Treatment
1
4
2
5
3
Figure 276.2 Uterine diagram used for mapping cysts.
Hysteroscopy may be used to visualize and treat cysts directly. Required equipment includes an endoscope with at least 1 m working length and a light source of 100 to 300 W. Disinfection of the equipment prior to use is important to prevent iatrogenic endometritis. A glutaraldehyde or iodophor solution is used to soak the endoscope for 15–20 minutes.16 It is also important to disinfect the instrument port. The endoscope should then be flushed and rinsed thoroughly with sterile water to prevent any uterine irritation due to contact with the disinfectant solution. Most mares require a sedative such as detomidine (0.004–0.04 mg/kg) intravenously. The mare should
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Uterine Cysts be placed in a set of stocks for restraint with the tail bandaged and held away from the vulva. The vulva and perineum are cleaned with a mild soap or iodine solution. A standard mare lavage system is used to distend the uterus with sterile fluid to allow manipulation of the endoscope through the uterine body and horns. Approximately 1.5–3 L of an isotonic saline solution or a balanced electrolyte solution is an appropriate fluid choice for visibility and a non-irritating pH. Other fluids such as 5% dextrose, sterile water, and sterile 32% dextran70 in 10% dextrose have also been used.16 It helps to perform the procedure at the very beginning of estrus or after the mare has ovulated so that the cervix has some tone and fluid is less likely to flow back onto the examiner. The other option for distending the uterus is to place the endoscope aseptically through the cervix and insufflate the uterus with air or carbon dioxide creating a pneumouterus. This must be done with caution as rapid overinflation of the uterus has potential to damage the endometrium.16 For either technique, the examiner dons a sterile sleeve with a sterile surgical glove placed over the hand to improve the grip on the endoscope as it is aseptically passed through the cervix. Sterile lubricant may be used on the body of the scope to facilitate passage. Once the endoscope is in the body of the uterus a systematic examination of the body and each horn should take place to view any abnormalities. Records of the findings may be made with photography or video recording equipment. Treatment of cysts is generally reserved for mares with numerous or large cysts that are likely to interfere with pregnancy. Often these mares have a history of reproductive problems, and a uterine biopsy is helpful to determine the overall prognosis for carrying a foal to term, and may help owners to decide about the economic feasibility of treatment. Mares with a few small cysts that are unlikely to impede conceptus mobility or implantation may not need treatment.7 Manual rupture of cysts may be accomplished when the cervix is dilated during estrus. Ablation of cysts manually during a postpartum examination is also effective. Aseptic technique should be used to avoid contamination of the uterus. Cysts may be punctured with an endoscopically guided needle or a uterine biopsy instrument. The residual lymphatic tissue will mostly resolve within a month.1 Uterine cysts may also be removed mechanically with obstetrical wire passed through a steel mare catheter. The cyst is ensnared at the base and removed by sawing through the tissue with the wire. One must use caution not to cut into the deeper layers of the uterus. It is likely that some hemorrhage will occur.7 Uterine lavage is also employed with these techniques. Warm
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hypertonic saline is often used as a post-treatment lavage, sometimes for several days until the return fluid is clear.1 A broad-spectrum antibiotic is recommended as a post-treatment infusion to prevent infection from any bacteria that may have been introduced during the procedure. Pedunculated cysts may be removed by electrocautery. A hysteroscopy is performed on the mare and a polypectomy snare is passed through the biopsy port of the endoscope. The loop is placed at the base of the cyst and closed. Electrocoagulation is used to burn the cyst at its base. Larger cysts may be removed by making several cuts. The cystic tissue may be removed with forceps, the loop, or lavaged from the mare at the end of the procedure.7 Hemorrhage may also occur with this procedure; if visibility is lost and there are numerous cysts the procedure may need to be completed in multiple stages. The area of the endometrium where the cyst was removed will be white. Follow-up examination at 15–21 days shows healing of the endometrium. This technique does not work well for mural cysts.7 A combination of electrocoagulation of the endometrial surface and manual transrectal reduction of a transmural cyst has been described.16 Laser therapy is recommended for smaller or nonpedunculated cysts. Diode and neodymium:yttrium aluminum garnet (Nd:YAG) lasers have been used for this procedure.3, 17 Mares are sedated and prepared for hysteroscopy. Three techniques have been described in the literature. In the first technique a sterile artificial insemination pipette attached to a bulb-type air pump is passed through the cervix along the endoscope to insufflate the uterus. The cyst closest to the tip of the uterine horn is evaluated and the laser is passed through the biopsy port of the endoscope until 1 to 2 cm is visible. The laser is set at 15 W and the tip is applied to the cyst, irradiating the tissue until the lymphatic fluid leaks out. Once the fluid has leaked the capsule of the cyst collapses onto the tip of the laser and ablation is continued until the cyst is removed to the level of the endometrium. After ablation the uterus is lavaged with 1 L of a 7% hydrogen peroxide solution followed by 2% povidone iodine solution in sterile saline. Additional uterine lavages may be needed over the next few days.17 Another technique uses carbon dioxide to insufflate the uterus. The cysts are ablated in two stages. Initially the laser is set at 50 W in continuous mode and located 1 cm away from the cyst until the surface blanches; then the power is raised to 100 W and the cyst membranes are punctured and ablated. Smoke is evacuated with a mechanical filtration device attached to the endoscope. Mares receive procaine penicillin G (22 000 IU/kg) i.m. for 3 days and flunixin meglumine (1.1 mg/kg) i.v. once.3
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The final technique uses lactated Ringer’s solution instead of air or CO2 to inflate the uterus. This has the advantage of causing less uterine irritation than air and there is no smoke produced. A diode laser is used through the biopsy port of the endoscope and 12 to 14 W of power is applied to small cysts while up to 18 W and multiple applications of the laser at different sites are used for larger cysts.12 Uterine cysts may be an inconsequential finding if they are small and few in number and they are detected in the periparturient period. However, the presence of large endometrial cysts provides predictive information for a mare’s fertility. The pathophysiology of chronic fibrosis, which decreases muscular contractility and allows cystic distension and accumulation of lymphatic fluid, is often accompanied by impaired uterine clearance. Additionally, the mechanical impedance of embryo mobility and the functional decrease in vascularization can result in pregnancy loss. Cysts of clinically significant size are readily detected by ultrasonography. Performing a uterine biopsy as part of the diagnostic plan helps give a comprehensive prognosis for a successful pregnancy. The decision whether to treat the cysts is dependent upon their size, number, and location. The recovery period following treatment is relatively short, which enables treatment during the breeding season with a return to service. New cysts may form as a result of continuing fibrosis, and owners should be made aware that further treatment may be required.
References 1. Wilson DL. Diagnostic and therapeutic hysteroscopy for endometrial cysts in mares. Vet Med 1985;80:59–63. 2. Bracher V, Mathias S, Allen WR. Videoendoscopic evaluation of the mare’s uterus: II. Findings in subfertile mares. Equine Vet J 1992;24:279–84.
3. Bilkslager AT, Tate LP, Weinstock D. Effects of neodymium:yttrium aluminum garnet laser irradiation on endometrium and on endometrial cysts in six mares. Vet Surg 1993;22:351–6. 4. Kenney RM, Ganjam VK. Selected pathological changes of the mare uterus and ovary. J Reprod Fertil Suppl 1975;23: 335–9. 5. Ragon AC. Diagnosis and management of uterine cysts. In: Proceedings of the American College of Theriogenologists and Society for Theriogenology Mare Reproduction Symposium. August 1996, Hastings, NE, pp. 21–6. 6. Tannus RJ, Thun R. Influence of endometrial cysts on conception rate of mares. Zentralbl Veterinairmed A 1995; 42:275–83. 7. Wolfsdorf KE. Endometrial cysts. In: Proceedings of the Bluegrass Equine Reproduction Symposium, October 2002, Lexington, KY. 8. Brook D, Frankel K. Electrocoagulative removal of endometrial cysts in the mare. J Equine Vet Sci 1987;7:77–80. 9. Bracher V, Matthias S, Allen WR. Influence of chronic degenerative endometritis (endometrosis) on placental development in the mare. Equine Vet J 1996;28:180–8. 10. McDowell KJ, Sharp DC, Grubagh W, Thacher WW, Wilcox CJ. Restricted conceptus mobility results in failure of pregnancy maintenance. Biol Reprod 1988;39:340–8. 11. Ginther OJ. Ultrasonic Imaging and Reproductive Events in the Mare. Cross Plains, WI: Equiservices, 1986; 189 pp. 12. Stanton MB, Steiner JV, Pugh DG. Endometrial cysts in the mare. J Equine Vet Sci 2004;24:14–19. 13. Adams GP, Kastelic JP, Bergfelt DR, Ginther OJ. Effect of uterine inflammation and ultrasonically-detected uterine pathology on fertility in the mare. J Reprod Fertil Suppl 1987;35:445–54. 14. Rambags BP, Stout TA. Transcervical endoscope-guided emptying of a transmural cyst in a mare. Vet Rec 2005;156: 679–82. 15. Kenney RM. Cyclic and pathologic changes of mare endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1978;172:241–62. 16. Wilson GL. Equine hysteroscopy: equipment for diagnostic endoscopy and photography. Vet Med 1985;80:76–88. 17. Griffin RL, Bennett SD. Nd:YAG laser photoablation of endometrial cysts: a review of 55 cases (2000-2001). Proc AAEP Annual Conference Proceedings; 2002; Orlando, FL, Vol. 48. pp. 58–60.
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Chapter 277
Uterine Abnormalities John P. Hurtgen
Abnormalities of the uterus are caused by a variety of predisposing factors such as infectious, traumatic, immunological, and hormonal conditions. A number of infrequently occurring uterine abnormalities are described below. These conditions are frequently overlooked because uterine size and consistency and ovarian cyclicity may not be altered.
mented. Laser surgery has been used on a few mares in order to remove the persisting cups. This appears to be a promising treatment.
Persistent endometrial cups
During the equine breeding season, mares have repeated estrous cycles of about 18–24 days until pregnancy is established. During the estrous cycle, follicles develop and ovulate at regular intervals and the mare exhibits estrous behavior in response to these ovarian changes. The histological characteristics of the mare’s endometrium have also been described during the estrous cycle. Briefly, the luminal epithelium of the uterus is medium to tall columnar. The uterine glands are quite abundant even when edema is present. The glands, themselves, are increased in diameter compared to glands during the winter anestrous period.2 During the transition phase between winter anestrous and breeding season, follicular development, ovulation, estrous behavior patterns, and the histological structure of the uterus are not always well synchronized. In private veterinary practice an occasional mare fails to become pregnant after mating during two or more estrous cycles even though the mare has normal ovarian cyclicity, estrous behavior, and other reproductive findings. Asynchrony of estrous and ovarian activity with expected uterine histology has been observed during the natural breeding season (May through August) in the northern hemisphere. Affected mares have normal estrous cycles and develop normal ovulatory follicles during estrus but fail to conceive. Uterine size and consistency are normal based on palpation per rectum. However, the pattern of uterine edema is much less pronounced or even absent compared to that usually observed during the breeding season.3 In order to determine the cause of infertility in
Endometrial cups form during early pregnancy at about day 40 of gestation and persist to day 120–150 of gestation. The endometrial cups produce equine chorionic gonadotropin (eCG). When a mare loses a pregnancy during gestation days 40–100, most do not return to estrus for 2 to 4 months. In a limited number of cases, persistence of the endometrial cups for as long as 30 months has been reported. Prolonged persistence of the cups has been observed following pregnancy loss and following delivery of a normal foal. In only a few cases, the endometrial cups are present in the uterus but eCG is not being produced based on blood hormone analysis.1 Clinical examination of the mare may suggest the presence of endometrial cups by noticing small, discrete, echogenic areas at the base of one or both uterine horns using ultrasonography (Figure 277.1a). Endoscopic evaluation of the uterine lining may demonstrate one or more raised areas suggesting the presence of endometrial cups (Figure 277.1b). A peripheral blood sample should be collected from the mare so that serum can be analyzed for the presence of eCG. If mares with prolonged persistence of the endometrial cups do not produce eCG, the mare may exhibits normal estrous cycle patterns. If eCG is still present, mares may fail to show regular estrous behavior during the breeding season and often have abnormal follicular and corpus luteum (CL) development.1 Because the incidence of this condition is very low, efficacious treatments have not been docu-
Asynchrony of estrous behavior, ovarian activity, and endometrial histology
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 277.1 (a) Ultrasonographic view of the uterus of a mare with persistence of the endometrial cups following a normal foaling. A 7-year-old Quarter Horse mare foaled normally. She was bred the following year by artificial insemination but failed to show estrus during breeding periods and did not become pregnant. The following year, the persisting endometrial cups were observed using ultrasonography (21 months after foaling). (b) Endoscopic view of the endometrium of the same mare demonstrating the presence of persistent endometrial cups. The cups were mechanically removed and the mare cycled normally, including normal estrous behavior. Trophoblastic cells were present in the removed cup material. (Image on loan from Dr Robert Lofstedt.)
some of these mares, an endometrial biopsy was obtained. In general, the histological appearance of the endometrium is suggestive of the anestrous or early transitional phase of the mare’s reproductive cycle. The luminal epithelium is cuboidal to low columnar. The uterine gland density is reduced and the glands, themselves, have a small diameter and appear inactive. When this degree of asynchrony between ovarian cyclicity and endometrial structure is present, fertility is likely to be reduced. If the affected mares proceed through one or more additional estrous cycles, the endometrium responds with the expected histological changes in relation to ovarian activity and the mares became pregnant. The pattern of endometrial edema normally observed during estrus is also present during ultrasonographic evaluation of the uterus. The cause of the asynchrony of the endometrial architecture with ovarian activity is unknown. Hormonal profiles of these mares have not been recorded. Possible explanations may include inappropriate lighting schemes prior to the breeding season such as inadequate lighting or constant 24-hour per day lighting. Mares that exhibit this syndrome are not likely to repeat this pattern in subsequent breeding seasons. In most cases, these mares are identified after three or four cycles have passed so specific therapy is not initiated. If these mares could be identified prior to breeding, supplemental estrogen and progesterone may stimulate the endometrium to develop the architecture expected for cyclic mares during the
natural breeding season and result in improved pregnancy rates.
Foreign bodies in the uterus A wide variety of foreign bodies are occasionally found in the uterus. The majority of the foreign bodies or substances do not interfere with the mare’s estrous cycle pattern. The most commonly recognized foreign body in the uterine lumen is the mummified fetus. In most cases of fetal mummification the mare will have normal estrous cycles during the breeding season. The mummified fetus may not be evacuated from the uterus during estrus due to the configuration of the fetus, size of the fetus, and lack of uterine fluids. Presence of a mummified fetus has been identified many days after a mare has delivered a live foal. In some cases the mummified fetus may become trapped in the cervical canal. Fetal mummification usually occurs during the second trimester but has been observed as early as 40–50 days of gestation. The presence of a mummified fetus can usually be identified by uterine palpation per rectum and ultrasonographic evaluation of the uterus. In the case of very early mummification of the embryo, the foreign body may not be recognized without the aid of endoscopy. Treatment of fetal mummification involves regression of the corpus luteum, if present, to induce estrous and cervical dilation. The degree of cervical dilation should be evaluated, per vagina, 3 to 4 days
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Uterine Abnormalities following prostaglandin. If the cervix is not adequately dilated, it may be useful to repeat the use of prostaglandin or administer estradiol or topical prostaglandin E2 to aid in dilation. In many cases, the cervix may need to be manually dilated to retrieve the mummified fetus. Infusion of the uterus with approximately one liter of fluid may aid lubrication of the uterus and cervix. With slow, deliberate manual dilation of the cervix, the fetus can usually be grasped and removed. The uterine environment is usually bacteriologically sterile. However, following manual manipulation and removal of the mummified fetus, the mare should be monitored for an induced endometritis. On rare occasions, the tip of an endometrial culture swab may break off within the uterus. In most cases the swab tip can be retrieved by passing two fingers through the cervix. The swab tip can also be visualized and recovered from the uterus with an endoscope that is equipped with a small biopsy forceps. If an endoscope in not available, the swab can be located using ultrasonography per rectum while a grasping instrument (forceps or biopsy punch) is passed through the cervix in a sterile fashion to grasp the swab. Alternatively it is possible in many older multiparous mares to dilate the cervix and pass fingers and even a hand into the uterus. In recent years, it has been proposed to place a 35 mm glass marble in the uterus of the mare as a method to prevent estrus in performance mares.4 The efficacy of this method of controlling estrous expression is questionable. Many but not all mares spontaneously lose the marble during estrus. If the marble is retained, fertility may be compromised. Additionally, an occasional mare may become pregnant. The marble may interfere with the developing pregnancy and result in embryonic death or abortion. Pregnancies have also gone to term in spite of the presence of a marble in the uterus. The presence of a 35 mm glass marble in the uterus can be detected by palpation of the reproductive tract per rectum or by ultrasonography of the uterus. The ownership of performance and race mares may frequently change before these mares are presented for breeding. Obviously, this foreign body should be manually removed during estrus prior to breeding. Certain medications may precipitate or crystallize in the uterine lumen leaving a residue on the endometrial surface. Although ampicillin and gentomycin are the antibiotics most frequently implicated, the formulation or pH of these medications are important factors. If this material is allowed to remain in the uterus, the mare may become infertile. Therapy involves the removal of the residual material by copious uterine lavage and induction of estrus using prostaglandin. The lavage should be repeated until
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the flush fluid is void of residual medication. Following removal of the foreign material, fertility should be normal.
Uterine hematoma, abscess, or tumors Localized abnormal enlargements of the uterus are uncommon in the mare but may include lymphatic cysts, hematoma, abscess, and tumors. Hematoma of the uterine wall frequently occurs in association with advanced pregnancy or parturition. The size of the hematoma may vary from very small to extremely large. The development of a hematoma may be due to normal fetal movements, dystocia, or secondary to violent movements associated with parturition, colic, or a foal. In most cases, the presence or development of a uterine hematoma is not diagnosed until days, weeks, or months have passed. In most cases, clinical signs of illness are not present. Hematoma of the uterine wall is usually diagnosed during reproductive evaluation for breeding or pregnancy. The use of ultrasonography may be useful to observe the pattern of blood clot and fibrin formation. Smaller hematomas may slowly regress over time and have minimal impact on fertility. Once large hematomas are well organized, it may be necessary to surgically remove the hematoma. Uterine hematomas do not alter the mare’s estrous cycles (Figure 277.2). Abscess formation in the lining of the uterus is rare. This condition may occur at the time of normal parturition or following dystocia when there has been a break in the surface of the endometrium with subsequent bacterial contamination. Other intrauterine procedures such as artificial insemination could potentially cause an abscess if the procedure is improperly performed. Resolution of an abscess of the uterine wall may be dependent on a multitude of factors such as size, location, and duration of the abscess. The most common tumor involving the uterus is the leiomyoma, though tumors of the uterus are extremely rare. Leiomyoma of the uterus is usually an isolated, circumscribed mass involving the myometrium. The mass is usually a single mass but multiple leoimyomas can occur within the same uterus. If the leiomyoma is small, the mare may carry a foal to term. However, if an affected mare is to be bred, the tumor is usually surgically removed prior to breeding. Other tumors of the uterus have been diagnosed such as the adenocarcinoma, leiomyosarcoma, and lymphosarcoma.5, 6 Tumors of the mare’s uterus are so infrequent that antemortem diagnostics, treatment, and subsequent fertility patterns have not been established.
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Figure 277.2 (a,b) These images were derived from a 6-year-old Standardbred mare. She had previously had two foals. The last foal was born approximately 8 months before these photographs were taken. That foaling was reported to be uneventful. A mesometrial hematoma was discovered at foal heat breeding. Late in the year the owner decided not to rebreed this mare and she was donated to us for teaching purposes. The size of the mesometrial enlargement was not recorded at the time of her foal heat examination. However, 8 months later the size of the hematoma was still about 22 × 12 × 12 cm (a). It could easily be shelled out from its mesometrial location. Experience suggests that the risk of rebreeding these mares is minimal, therefore the need to stop breeding ¨ this mare should be questioned. (Images on loan from Dr Robert Lofstedt.)
Transluminal adhesions Transluminal adhesions are occasionally encountered following endometritis associated with dystocia, tissue damage to the endometrium, or following the use of harsh medications or fluids in the uterus. The most commonly implicated caustic medications involved in transluminal adhesion formation include chlorhexidine, iodine, and unbuffered gentomycin. These products have been used safely in the uterus of mares but the appropriate frequency of use, stage of cycle, and concentration of medication have not been determined in controlled studies of normal endometria and seriously inflamed uteri. In the majority of cases when these caustic treatments are used, other medications are available with much reduced endometrial irritation and are less likely to induce transluminal adhesions. Transluminal adhesions are usually diagnosed with the use of endoscopic evaluation of the uterine lumen. The adhesions appear as scar tissue and bands of tissue transversing the uterine lumen and occasionally completely occlude an entire uterine horn or portion of it. Complete transluminal adhesions are suspected by ultrasonographic identification of fluid only in one uterine horn on multiple examinations. Dilation of the uterus with fluid or air7 or saline may result in a non-uniform filling of the uterine lumen. Affected mares usually have markedly reduced fertility due to intraluminal adhesions, underlying periglandular or endometrial fibrosis, and altered early embryo mobility.
The transluminal adhesions may be broken down manually by passing a gloved hand through the dilated cervix. The adhesions may also be freed using laser surgery. Although the uterine condition of the treated mares may appear clinically improved, fertility usually remains poor due to the underlying scar tissue formation and periglandular fibrosis.
Pelvic and visceral adhesions involving the uterus Adhesions of the uterus or cervix to other structures in the pelvic cavity or abdomen can interfere with fertility and maintenance of pregnancy. These adhesions may develop in association with uterine surgery such as cesarean section, uterine or ovarian artery rupture, postpartum metritis, peritonitis, uterine laceration associated with fetal movement, forced fetal extraction or fetotomy, and uterine trauma associated with ovariectomy. The uterine adhesions may involve the pelvic cavity, bladder, ovary, or viscera. If the adhesions are not extensive, mares may still become pregnant and carry a foal to term with the adhesions resolving or breaking down during the course of uterine descent during pregnancy. The periuterine adhesions may alter uterine position or uterine motility, resulting in increased risk of uterine fluid accumulation and susceptibility to endometritis. Periuterine adhesions are usually not treated due to the inaccessibility of the uterus for surgery. The advent of laser treatment of adhesions may allow some of the more accessible structures to be freed
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Uterine Abnormalities by laser surgery using a paralumbar or midline approach. Before this type of abdominal surgery is performed, the health of the uterus should be evaluated by manual and ultrasonographic examination of the uterus, vagina and cervix, bacteriological culture of the uterus, and endometrial biopsy.
References 1. Steiner J, Antczak DF, Wolfsdorf K, Saville S, Brooks D, Miller E, Bailey E, Zent W. Persistent endometrial cups. Anim Repro Sci 2006;94:274–5. 2. Kenney RM. Cyclic and pathologic changes of the mare endometrium as detected by biopsy, with a note on early embryonic death. J Am Vet Med Assoc 1978;172:241– 62.
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3. Samper JC. Ultrasonographic appearance and the pattern of uterine edema to time of ovulation in mares. In: Proceedings of the 43rd Annual Convention of the American Association of Equine Practitioners, 1997; pp. 189–91. 4. Nie GJ, Johnson KE, Wenzel JGW. Use of a glass ball to suppress behavioral estrus in mares. In: Proceedings of the 47th Annual Convention of the American Association of Equine Practitioners, 2001; pp. 246–8. 5. Gunson DE, Gillette DM, Beech J, Orsini J. Endometrial adenocarcinoma in a mare. Vet Pathol 1980;17:776–80. 6. Neufeld JL. 1973. Lymphosarcoma in a mare and review of cases at the Ontario Veterinary College. Can Vet J 1973;14:149–53. 7. Mather EC, Refsal KR, Gustafsson BK, Seguin BE, Whitmore HL. The use of fibre-optic techniques in clinical diagnosis and visual assessment of experimental intrauterine therapy in mares. J Reprod Fert Suppl 1979;27:293–7.
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Chapter 278
Laboratory Methods for Isolation and Evaluation of Bacteria, Fungi and Yeasts David L. Hartman and Shalyn Bliss
Introduction
Aseptic preparation
For many equine reproduction practitioners, microbiology can be frustrating. By the time a culture is taken, sent to a laboratory and the results reported, the mare has already ovulated and the window of time for directed intervention has passed for that cycle. Likewise, the prospect of performing in-house microbiology can seem overwhelming, time consuming and tedious. The reality is that in-house microbiology can be practical, simple, inexpensive and exponentially useful.
Any time something is passed from the outside environment into the uterus, the utmost care must be taken to ensure that contamination of the uterus is prevented. It is important that the tail is safely tied or held out of the way and that any stray hairs are contained. The vulva should be liberally scrubbed with a mild detergent soap and plenty of water until no more debris can be removed. When washing the vulva it is important to begin in the center where the labia oppose and work out so that dirt and debris are not brought towards the vaginal opening. The vulva should then be carefully dried in the same manner (center to outside) with clean, uncontaminated paper towels or cotton. It is important that the vulva is completely dry, as water is a vehicle for contamination. It is especially important to dry the tail and rectum so that contaminated water does not drip onto the prepared vulva. Generally, the mare would have just previously been subjected to palpation or/ultraonography per rectum before a uterine culture sample is taken. The chance for feces and lubricant to have entered the vaginal vestibule during the examination is very high. If one does not check for contamination, the lubricant and feces will be picked up on the sterile sleeve and the end of the culture swab and carried straight to the uterus. The simple step of checking the vestibule can easily prevent gross fecal contamination of the uterus. One method would be to examine the vaginal opening with a clean paper towel and a small amount of sterile lube. Insert
Timing of sample collection In my view, a critical error is taking a uterine culture sample from a mare that is not in heat, which can give false and misleading information unless the mare has a blatant uterine infection. To ensure that meaningful and helpful culture results are reliably obtained, a mare should be ‘in good heat’ when the sample is collected, i.e., there is no corpus luteum present and the mare has at least a 25–30 mm follicle, her cervix is soft and open, and she has large prominent endometrial folds, indicating good endometrial edema. Furthermore, collecting a culture sample from a mare that is not in heat and does not have a fully opened cervix can result in uterine infection because of contamination from the vulva and vagina that is introduced during sample collection that cannot be cleared from the uterus.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Laboratory Methods for Isolation and Evaluation of Bacteria, Fungi and Yeasts the paper towel slightly into the vestibule and then check it for feces and lube. If any is present, the mare should be wiped out with sterile lube and rewashed.
Sample collection When culturing the uterus, it is important to use a commercially available double-guarded culture swab. These swabs are made of three parts: the sterile swab surrounded by a plastic inner guard and a plastic outer guard. The guarded swab prevents the sterile culture swab from contacting the vaginal mucosa, which will have a resident microbial population present.1 It is helpful to protect the swab from contamination by cupping the tip into the palm of the hand as the arm is introduced. Before proceeding further into the vagina the hand is withdrawn and examined for presence of fecal contamination. Then the hand and guarded swab are passed further into the vagina, and the cervix is located and used to guide the apparatus into the uterus. While keeping gentle pressure on the culture swab so that it does not slip out of the uterus, remove your hand and carefully place it in the rectum. Locate the culture swab apparatus and carefully guide it to the most dependent portion of the uterus. This is usually the base of a uterine horn. Subsequently, push the inner guard out and then the middle sterile swab. Rotate the swab carefully with the outside hand, while lifting the dependant part of the uterus upwards and gently pressing down on the culture swab rectally. This method allows you to collect a sample from the most dependant part of the uterus where, by simple gravity, fluid and contamination will pool. Finally, pull the center sterile swab back inside the inner guard, then pull the inner guard back into the outer guard and remove the entire apparatus. The swab should be immediately transferred to a blood agar plate. If it cannot be transferred within a few minutes, then it should be transferred to an appropriate transfer media tube. When transporting culture samples back to the laboratory, they should be kept refrigerated and transferred if possible within a few hours. Overgrowth of bacteria in the transfer media can give a misrepresentative sample of what is actually present in the uterus.
Plating a culture Quickly remove your sleeve and carry the entire culture apparatus into the laboratory area. Remove the outer guard and discard, push the sterile swab forward through the inner guard and carefully break it off. Gently twirl the swab across half of the blood agar
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plate. Be careful to make good contact with agar without breaking the surface of the agar. Next turn the plate 90◦ and continue rotating the swab. Rotate the plate 90◦ once more and continue rotating the swab. It is also acceptable to change loops or swabs between each rotation of the plate in order to ensure isolated colonies; however, the author does not consider this to be necessary. Extreme caution must be taken not to contaminate the plate. Hold the plate at an angle and do not allow your hand to hover over the plate, also do not speak while plating a culture and make sure that you do not breathe on the plate. Label the plate, replace the lid, and place the plate in a 37◦ C incubator, lid side down.
Incubator A standard microbiological incubator should be used. The size should be chosen according to the volume of cultures expected. It should have an adjustable thermostat and the temperature should remain at 37◦ C. The temperature should be verified daily with an independent thermometer.
Blood agar (TSA with 5% sheep blood) All samples should initially be plated on a 5% sheep blood agar plate. This type of plate is readily available from any microbiological supply company. The plate should be allowed to incubate for 24 hours and then examined for growth. Most pathogens will be apparent at 24 hours, with the exception of Corynebacterium and some yeasts and fungi. If a plate is negative for growth at 24 hours, it should be allowed to incubate for a total of 72 hours before declaring it a true negative. To consider a culture positive and significant, it should have several identical colonies. A plate with multiple different colony types is most likely contaminated and a second sample should be obtained when possible. It is important to quantify the number of colonies present. One or two colonies are likely to be contaminants. Five to ten colonies likely signify a mild infection. More than 10 colonies are an indication of a more severe infection. With a little experience, one will quickly begin recognizing the common organisms by their appearance on blood agar at 24 hours. If there is significant growth on the blood agar, the next step is to set up an antibiotic sensitivity test. For ease of use the author prefers the KirbyBauer technique which uses antibiotic-impregnated discs to inhibit growth of bacteria that are sensitive to that specific antibiotic. The zone of inhibition is influenced by the size of the inoculum. Therefore it
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is best to standardize the size of the inoculum. This can be accomplished by creating a subculture from the original into nutrient broth. In a few hours it is possible to compare the turbidity of the subcultured broth with the standard and then use this to inoculate the Mueller-Hinton plate. This is done in such a way as to cover the entire plate. Following this the antibiotic discs are placed on the MuellerHinton agar and incubated for 24 hours. It is necessary to use Mueller-Hinton with 5% sheep blood for Streptococcus spp. Antibiotic sensitivities should be interpreted after 24 hours of incubation. The zone of inhibition varies for different antibiotic discs; a commercially available chart should be consulted. The diameter of the zone of inhibition is measured in millimeters and the chart consulted to determine the minimum inhibitory concentration; and whether the growth is sensitive, intermediate, or resistant to each antibiotic.2 Within 48 hours of culturing the uterus of a mare, directed antibiotic therapy can be initiated, if deemed necessary.
Differential media After observing significant growth on the blood agar plate at 24 hours, an antibiotic sensitivity and differential media plate should be set up. The differential media will begin the process of the bacterial identification. For equine reproduction two common ones are the ‘tri-plate’ and the ‘bi-plate’. The tri-plate contains a Gram-negative selective agar called eosin–methylene blue or EMB, a Staphylococcus-selective agar, and TSA with 5% sheep blood. All aerobic relevant microbes will grow on TSA with 5% sheep blood, only Gram-negative organisms will grow on the EMB, and only Staphylococcus will grow on the ‘staph-select’ section. Therefore, if an organism grows only on the blood agar, it is Gram-positive; if it grows on both blood agar and EMB, it is Gram-negative; and if it grows on blood agar and staph select, it is a Staphylococcus. A bi-plate contains MacConkey and Columbia CNA (clistin and nalidixic acid) with 5% sheep blood. Gram-negatives will grow only on the MacConkey and Gram-positives will grow only on the CNA section.3 To inoculate the differential media plates; find a completely isolated colony to ensure you are obtaining a pure sample, pick up a single colony using a sterile cotton swab or inoculating loop and transfer it to the preferred differential media plate. Streak the swab or loop across each section of the plate, making contact with the agar, but do not break its sur-
face. After streaking the plate, replace the lid, label and place it, lid side down, in a 37◦ incubator for 24 hours. After incubating the differential media plates for 24 hours, observe the plates for growth on the different sections. After viewing the differential media plates and determining the Gram-negative or Grampositive status of the organism, a commercial identification system can be used to further determine the taxonomy of the organism, if desired. If the interpretation of the selective agar is not clear, one can easily perform a Gram stain to confirm the status of the organism. Always consider that the results of the differential media and antibiotic sensitivity pattern must make cohesive sense. If the results do not agree, e.g., sensitive to Gram-positive antibiotics but grows on EMB, a Gram stain should be performed for confirmation.
Catalase, oxidase, and indole tests Catalase, oxidase, and indole tests may be performed on organisms, if desired, for further classification and for input into many standard commercial identification kits. To determine if a bacterial colony has the enzyme catalase, a drop of 3% hydrogen peroxide (H2 O2 ) is mixed with a single colony of the bacteria. This is usually done on a glass microscope slide. If the catalase enzyme is present, water (H2 O) and oxygen (O2 ) will be released and bubbling or effervescence will be observed; this is considered catalase-positive. If the enzyme is not present, there is an absence of bubbling; this is considered catalase-negative. Staphylococcus and Corynebacterium are two common catalasepositive organisms. The oxidase enzyme is involved with the metabolic pathway for nitrate and can be used to differentiate certain Gram-negative bacteria. To perform the Kovac method for detecting oxidase, a single colony of bacteria is transferred to filter paper impregnated with reagent. If the paper turns purple within 10 seconds, it is considered oxidase-positive. No color change within 10 seconds is oxidasenegative. Pseudomonas is a common oxidase-positive organism. The indole test determines the presence or absence of the enzyme tryptophanase. It is performed by transferring a single colony of bacteria to filter paper impregnated with reagent and observing for a blue color change. A blue color change is considered indole positive and no color change is considered indole-negative. Escherichia coli is a common indolepositive organism.3
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Commercial identification kits There are several commercial microbiology identification systems available. The basic way all the systems work is that a pure colony of the growth is inoculated into a strip or series of wells and incubated. The strip or wells are then read for colorimetric change. The color changes are compared to a standardized chart, then recorded as a plus or minus. The plus or minus changes are then used to generate a numeric code. The code is then either entered into a computer software program or looked up in a code book and the taxonomy of the organism is identified. The commercially available tests are developed through serial standardized testing of different strains of bacteria and a confidence factor will be generated by the computerized programs.3
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Table 278.1 Quick reference guide: Most common isolates in equine reproduction Identifying characteristics
Microorganism
Gram stain
Alpha Streptococcus
Positive
Green zone of hemolysis on blood agar at 24 hours
Bacillus
Positive
Large gray feathery colony on blood agar at 24 hours
Beta Streptococcus
Positive
Clear zone of hemolysis on blood agar at 24 hours
Citrobacter
Negative
Foul odor on blood agar at 24 hours
Corynebacterium
Positive
Pinpoint clear to white colonies on blood agar at 48 hours
Enterobacter
Negative
Foul odor on blood agar at 24 hours
Specific microbes
Escherichia coli
Negative
Metallic green sheen on EMB agar at 24 hours
With a little experience, one will quickly begin to identify the most common uterine pathogens on blood agar or with the use of differential media. The following is a brief discussion of the most common pathogens, their characteristics, and the specific laboratory techniques used to identify them. Table 278.1 is a quick reference guide for the most common isolates in equine reproduction.
Klebsiella
Negative
Mucoid growth on EMB at 24 hours
Proteus
Negative
Swarming colonies on blood agar at 24 hours
Pseudomonas
Negative
Sweet ‘grape’ smell on blood agar at 24 hours, turns Mueller-Hinton agar lime-green at 24 hours
Staphylococcus
Positive
Growth on staph-select differential media
Beta-hemolytic Streptococcus Yeasts or fungi
Beta Streptococcus organisms, such as Streptococcus zooepidemicus, are usually apparent on blood agar at 24 hours as a small white/gray round colony with a zone of hemolysis surrounding it. They are Gram-positive, catalase-negative cocci.3 Once identified, they should be transferred to a Mueller-Hinton with 5% sheep blood agar plate for antibiotic sensitivity. In a short time, one will quickly be able to recognize beta Streptococcus on the initial blood agar plate, and further work-up on differential media and a commercial identification kit is not necessary, unless one desires to know the exact species of the organism (Figure 278.1). Alpha Streptococcus Alpha Streptococcus organisms, such as Streptococcus viridans, appear on blood agar at 24 hours as small clear or white colonies with a greenish zone of hemolysis. They are Gram-positive, catalase-negative cocci.3 Some Gram-negative organisms can also exhibit alpha hemolysis, so it is important to use differential media to determine if an organism is an alpha Streptococcus or Gram-negative (Figure 278.2).
Not sensitive to antibiotic discs; Candida grows on Biggy’s agar at 24 hours; fungus has a fuzzy growth on blood agar but may take 72 hours to appear
Bacillus Bacillus is occasionally isolated as a contaminant from the equine uterus. Unless it is present in large numbers in pure culture, it is usually not of concern. Bacillus is a Gram-positive, aerobic, spore forming, pleomorphic rod and is usually of low virulence. On blood agar it will appear as a large, feathery betahemolytic colony.3 Corynebacterium Corynebacterium tends to be a slow-growing organism and is usually not apparent on blood agar at 24 hours. However, by 48 hours it will appear as a pinpoint non-hemolytic growth. It should then be plated for antibiotic sensitivity and on differential media. It will again be slow to grow on these media, so it is
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Figure 278.1 Beta-hemolytic Streptococcus on blood agar at 24 hours; illustrating clear zone of beta-hemolysis. (See color plate 130).
important to allow 48 hours before making a final diagnosis. Corynebacterium is a catalase-positive Gram-positive rod.3 Staphylococcus Staphylococcus generally appears at 24 hours as a medium-sized, white non-hemolytic colony. How-
Figure 278.3 Staphylococcus aureus on tri-plate; illustrating yellow zone from Mannitol fermentation on staph-select section. (See color plate 132).
ever, it can have several different morphological characteristics. Staphylococcus is a catalase-positive Grampositive coccus. If using a tri-plate, it will grow on the purple staph-select portion of the plate. The staphselect agars contain a high concentration of salt, the sugar mannitol, and phenol red. Staphylococcus is able to grow even with the high concentration of salt and the fermentation of the mannitol will produce a yellow zone around the Staphylococcus aureus organism3 (Figure 278.3).
Gram-negative organisms
E. coli
Figure 278.2 Alpha-hemolytic Streptococcus on blood agar at 24 hours; illustrating green zone of alpha hemolysis. (See color plate 131).
Escherichia coli is a Gram-negative MacConkeypositive, oxidase-negative rod.3 It appears on blood agar at 24 hours usually as a medium-sized gray alpha-hemolytic colony. Oftentimes it has a distinctive horizontal line of growth extending from the edges of the colony (Figure 278.4). It should be plated both on Mueller-Hinton medium for antibiotic sensitivity and on differential media. If plated on a triplate, the growth will have a metallic green sheen at 24 hours on the EMB portion of the plate. The green sheen is pathognomonic for E. coli and further workup is not usually necessary (Figure 278.5). There are a few strains of E. coli that are not metallic green on the EMB agar but will be identified using a commercial identification kit.
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Laboratory Methods for Isolation and Evaluation of Bacteria, Fungi and Yeasts
Figure 278.4 Escherichia coli on blood agar at 24 hours; illustrating horizontal growth along streak line. (See color plate 133).
Pseudomonas Pseudomonas is a Gram-negative, MacConkeypositive, oxidase-positive rod and will appear at 24 hours on blood agar as a medium-sized gray alpha-hemolytic colony;3 however, sometimes Pseudomonas colonies may be slow growing and a presumptive negative culture should not be made until at least 36 hours. It has a distinctive sweet
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Figure 278.6 Pseudomonas on Mueller-Hinton medium; illustrating lime green color change of the clear agar. (See color plate 135).
‘grape’ smell and by 24 hours most colonies will be large and gray with a beta-hemolytic zone. When plated on Mueller Hinton agar for antibiotic sensitivity, the white agar will turn a lime-green color. This color change is pathognomonic for Pseudomonas and further work-up is only necessary if you would like to determine the specific strain of the organism (Figure 278.6).
Klebsiella Klebsiella generally appears as an alpha-hemolytic white or light gray creamy colony at 24 hours on blood agar. When plated on a tri-plate, the EMB portion of the agar will have a pink mucoid growth. Klebsiella is a Gram-negative, MacConkeypositive, oxidase-negative rod.3 Further work-up with a Gram-negative identification kit will be required to confirm identification (Figure 278.7).
Proteus
Figure 278.5 Escherichia coli on tri-plate; illustrating metallic green sheen on eosin–methylene blue (EMB) agar section. (See color plate 134).
Proteus appears on blood agar at 24 hours as a small gray colony with a zone of growth around it resembling a ripple, as if you had thrown a stone into still water (Figure 278.8). This swarming of the colony is pathognomonic for Proteus. By 48 hours and oftentimes sooner, the Proteus colonies will have taken over the entire plate. Even sometimes at 24 hours, a single Proteus colony will have overgrown a blood agar plate to the extent that it is difficult
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Mare: Disease of the Uterus It is a Gram-negative, MacConkey-positive, oxidasenegative rod.3 It will grow on the blood agar and EMB portions of the tri-plate. Further work-up, using a commercial identification kit for Gram-negative organisms, is necessary to diagnose this organism.
Enterobacter Enterobacter, when present in large numbers and in pure culture, is a highly virulent pathogen. Its most distinguishing characteristics are a strong foul odor and variable mucoid appearance on blood agar. Enterobacter is a normal inhabitant of the gastrointestinal tract. It is a Gram-negative rod that is MacConkeypositive and oxidase-negative.3 For final identification, inoculation of a Gram-negative identification system is necessary. Figure 278.7 Klebsiella on tri-plate; illustrating mucoid appearance on EMB section. (See color plate 136).
to isolate other bacterial types in a mixed infection. It will grow on both the blood agar and EMB portions of a tri-plate and should be plated on MuellerHinton medium for antibiotic sensitivity. Proteus is a Gram-negative, MacConkey-positive, oxidasenegative rod.3
Citrobacter Citrobacter appears on blood agar at 24 hours as a small to medium-sized gray alpha-hemolytic colony.
Figure 278.8 Proteus on blood agar at 24 hours; illustrating colony swarming. (See color plate 137).
Yeasts and fungi Yeasts and fungi can be pleomorphic or can have a distinctive morphology. Oftentimes they tend to be slow growing on blood agar and it is necessary to allow 72 hours of incubation on blood agar in order to declare a culture to be negative for fungi and yeasts. Fungi will have a fuzzy ‘moldy’ appearance on blood agar (Figure 278.9). Yeasts will have a phenotypic appearance similar to a colony of bacteria. However, the definitive diagnosis of a fungus or yeast comes when the antibiotic sensitivity on Mueller-Hinton medium is observed for growth at 24 hours. If a fungus or yeast is present, the growth will not be inhibited by any of the antibiotics (Figure 278.10). Yeast should be plated on the differential media, Biggy’s agar. If it is positive for growth on Biggy’s after 24 hours, it is the yeast Candida. No growth on
Figure 278.9 Fungus on blood agar at 73 hours; illustrating fuzzy growth. (See color plate 138).
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Figure 278.10 Yeast on Mueller-Hinton and Mueller-Hinton with 5% sheep blood, both with antibiotic sensitivity discs; illustrating no sensitivity to antibiotics. (See color plate 139).
Biggy’s agar and no sensitivity to antibiotics indicates a non-Candida type of yeast. Mycology in the laboratory is generally a more time-consuming and difficult undertaking when compared to bacteriology. It is advised that once a fungus or yeast other than Candida is identified, the culture plates be sent to a mycology laboratory for yeast/fungus identification and sensitivity.
References 1. Hirsh D, MacLachlan J, Walker R. Veterinary Microbiology, 2nd edn. Oxford: Blackwell Publishing, 2004. 2. Songer G, Post K. Veterinary Microbiology Bacterial and Fungal Agents of Animal Disease. St Louis: Elsevier Saunders, 2005. 3. Forbes B, Sahm D, Weissfeld A. Bailey & Scott’s Diagnostic Microbiology, 11th edn. St Louis: Mosby, 2002.
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Section (l): Diseases of the Ovary and Oviduct
279.
Disorders of Ovulation Patrick M. McCue
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280.
Disorders of the Oviduct Irwin K.M Liu
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281.
Non-neoplastic Abnormalities Donald H. Schlafer
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282.
Ovarian Neoplasia Claire E. Card
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Chapter 279
Disorders of Ovulation Patrick M. McCue
Introduction A vast majority of mares develop dominant follicles and ovulate at an interval of approximately 21 days during the physiological breeding season. Failure of ovulation results in decreased reproductive efficiency and is responsible for significant economic loss. Ovulation failure may be associated with identifiable risk factors, such as advanced age, stress, hormone therapy, and certain medical conditions (e.g., equine Cushing disease, ECD), or may occur spontaneously without any risk factors identified. Specific causes of ovulation failure in the mare are often not known, but have historically been suggested to be insufficient pituitary gonadotropin stimulation1 or insufficient estrogen production.2 Conversely, it was recently reported that mares which develop anovulatory follicles had slightly higher plasma levels of estradiol prior to formation of the anovulatory follicle as compared to mares that ovulated.3 In the same study, it was noted that levels of progesterone, luteinizing hormone, and follicle stimulating hormone were similar between cycles in which mares developed anovulatory follicles and cycles in which mares ovulated normally. Cortisol levels were not significantly different between normal and affected mares.4 A commonly asked question is whether development of anovulatory follicles is associated with administration of exogenous hormones, such as prostaglandins, progestins, or ovulation-inducing agents, in the preceding cycle. Although many broodmares are treated with hormones as a component of routine reproductive management, McCue and Squires5 noted that mares may develop anovulatory follicles without prior exposure to exogenous hormones. It was determined that there was no direct association between hormone administration and subsequent formation of an anovulatory follicle. In addition, it was also noted that if mares did develop an anovulatory follicle on a given cycle, they were
more likely to form a luteinized anovulatory follicle if they had received hCG or deslorelin acetate during that estrous period. The presence of active, functional endometrial cups in a non-pregnant mare may result in complete suppression of ovarian follicular activity, or less commonly may result in limited to substantial follicular development and subsequent luteinization without ovulation.6, 7 Persistence of endometrial cups beyond their expected lifetime of 60 to 100 days has been reported in mares that sustained pregnancy loss and in mares that carried to term and foaled normally.6, 8 Mares with persistent endometrial cups may experience abnormal follicular development and ovulations.8 Stress and subsequent production of steroid hormones from the adrenal gland may also be a potential cause of ovulation failure. An increase in production of cortisol, progesterone, and androgens by the adrenal gland was noted after administration of ACTH to intact or ovariectomized mares.9, 10 Chronic treatment of cycling mares with dexamethasone resulted in inhibition of LH secretion and ovulation failure.11, 12 A similar effect was noted after administration of cortisol to ewes and gilts.13, 14 However, administration of 20 mg of dexamethasone to normally cycling mares every 12 hours beginning when a follicle 30 mm in diameter was detected did not adversely affect or delay ovulation.15 In reality, it is likely that ovulation failure is multifactorial in origin and lack of ovulation is the final common clinical sign associated with a diverse set of causes or circumstances.
Incidence Retrospective analysis of reproduction records of 721 mares over 1845 cycles during the physiological breeding season revealed that ovulation failure occurred in 8.2% of cycles.5 The incidence of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 279.1 Percentage of mares experiencing at least one anovulatory follicle (AF) during a single physiological breeding season sorted by age. Adapted from McCue et al.5 Age (years)
# Mares
% Mares with AF
2–5 6–10 11–15 16–20 > 20
62 135 144 111 41
4.8 8.9 16.7 23.4 51.2
anovulatory follicles was noted to increase with age (Table 279.1). Approximately 4.8% of mares 5 years old or less experienced at least one anovulatory follicle during a single breeding season. In contrast, 51.2% of mares ≥ 20 years of age had at least one cycle that ended in ovulation failure each breeding season. A high percentage (43.5%) of mares that developed one anovulatory follicle experienced one or more subsequent estrous cycles with anovulatory follicle formation during the same breeding season. The incidence of anovulatory follicles in a herd of 47 research mares undergoing repeated short-cycling with prostaglandins was reported by Lefranc and Allen.16 Anovulatory follicles developed in 11.9% of 737 estrous cycles, and 50.5% of mares exhibited at least one anovulatory follicle during the 3-year study period. Approximately 26.2% of mares had more than one cycle with ovulation failure, and a tendency for recurrence of ovulation failure in subsequent breeding seasons was noted. Gastal and co-workers17 reported that anovulatory follicles occurred in 3 of 56 cycles (5%) in pony mares early in the ovulatory season and 5 of 23 cycles (23%) late in the season. In a subsequent study, they noted that pony mares with a history of anovulatory follicles experienced an incidence of 23% (4 of 13 cycles) the following year, while mares without a history of anovulatory follicles did not exhibit anovulatory follicles the subsequent year.3
Formation and identification It is often difficult to determine a priori if a dominant follicle in an estrual mare will fail to ovulate. Initial growth patterns of follicles destined to become anovulatory are usually within normal limits and formation of an anovulatory follicle is usually preceded by development of normal endometrial folds or edema.3, 5, 16 Manual and ultrasonographic indicators of impending ovulation, such as decreased follicle turgidity, loss of spherical shape, and serration of the granulosum, were not different between mares that ovulated and mares with ovulation failure.3, 4 An
Figure 279.1 Ultrasonographic image of an anovulatory follicle with multiple echogenic particles within the follicular lumen.
increase in follicular wall thickening is also observed both in mares with normal cycles and anovulatory cycles. Color Doppler ultrasonography revealed that follicles destined for ovulation failure had an increased percentage of the follicle circumference with detectable blood flow than follicles that ovulated normally.3 The biological significance of the apparent increase in vascularity immediately prior to ovulation failure is unknown. The first indication of a problem is typically detection of free-floating echogenic particles within follicular fluid during ultrasound examination;5 however, this is not a pathognomonic finding as occasionally mares with particulate matter in the follicular fluid will have an apparently normal ovulation. A progression from echogenic particles (Figure 279.1) to a web-like formation of echogenic strands (Figure 279.2) traversing the follicular lumen occurs during the subsequent one to two days. This is usually followed by complete infiltration of the follicular lumen with echogenic material as luteinization occurs (Figure 279.3). The ultrasonographic image of hemorrhage into the lumen of an anovulatory follicle can appear similar to the image of a corpus hemorrhagicum that forms after a normal ovulation. However, daily examination of mares would allow for the opportunity to detect the presence or absence of a collapsed follicle. A second distinguishing feature is delayed
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Disorders of Ovulation
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clotting of blood in follicular fluid of a hemorrhagic anovulatory follicle as compared with the immediate clotting that occurs as blood enters the antrum of an ovulated follicle.4 Equine follicular fluid contains a heparin-like anticoagulant18 which undoubtedly contributes to the slow progression of changes within a hemorrhagic anovulatory follicle. The average peak diameter of an anovulatory follicle has been reported to range from 48.1 ± 8.0 mm16 to 50.9 ± 11.2 mm.5 Lefranc and Allen16 noted that younger (3 to 5 years old) maiden mares developed anovulatory follicles that were medium in size, while older mares (>15 years of age) tended to develop large (>50 mm) anovulatory follicles. The relationship between size of the dominant follicle prior to bleeding within the follicular lumen and the maximum size of a hemorrhagic anovulatory follicle has not been reported. The diameter of an anovulatory follicle may increase as blood enters the follicular lumen. The increase usually occurs 1 to 3 days after echogenic particles are first observed in follicular fluid.3 In some instances, unclotted blood may be present in the follicular lumen for several days (Figure 279.4) and Figure 279.2 Ultrasonographic image of an anovulatory follicle with echogenic strands traversing the follicular lumen.
Figure 279.3 Ultrasonographic image of an anovulatory follicle completely infiltrated with echogenic material.
Figure 279.4 Ultrasonographic image of an anovulatory follicle with unclotted blood within the follicular lumen.
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14
12
Progesterone (ng/ml)
c279
10
8
6
4
2 PGF 0 1
2
3
4
5
6
7
8
9
10
11
Interval From Initial Identification of Anovulatory Follicle (days) Figure 279.5 Progesterone levels in a mare with a luteinized anovulatory follicle. Day 1 refers to the day echogenic particles were first detected within the follicular lumen. A single dose of cloprostenol sodium (250 µg) was administered on day 10.
gravitational settling of red blood cells may be visible on the ventral aspect of the follicular lumen. Progesterone levels may be used to determine the luteal status of anovulatory follicles. A majority of anovulatory follicles eventually become luteinized, with an elevation in progesterone noted in 85.7%5 to 88.1%16 of affected mares. Mares with anovulatory follicles containing a highly echogenic lumen invariably have elevated progesterone levels (Figure 279.5), and mares with very large luteinized anovulatory follicles often have progesterone levels significantly greater than levels usually present in a normal luteal phase. However, mares with persistent anovulatory follicles that only have a thickened echogenic border around the follicle often have only a mild elevation in blood progesterone levels (i.e., 1–3 ng/mL).5 Mares with luteinized anovulatory follicles go out of heat and develop good tone in the uterus and cervix as progesterone concentrations increase. Approximately 11.1% to 14.3% of anovulatory follicles fail to luteinize as recognized by little to no echogenic particles within the follicular lumen and progesterone levels of < 1.0 ng/mL (Figure 279.6). Mares with non-luteinized anovulatory follicles eventually show a decline in uterine edema and go out of heat as estradiol levels decrease. Nonluteinized anovulatory follicles do not appear to be hormonally active and typically do not result in behavioral changes or alterations in follicular dynamics. They essentially appear to be non-viable atretic structures. Non-luteinized anovulatory follicles regress over time. Affected mares usually develop
another follicle, either on the same ovary or on the contralateral ovary, and return to estrus even in the continued presence of the atretic follicle. Unfortunately, some mares will experience failure of ovulation of the next dominant follicle during subsequent cycles.
Figure 279.6 Ultrasonographic image of a persistent nonluteinized anovulatory follicle.
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Disorders of Ovulation
Failure of ovulation-inducing agents Human chorionic gonadotropin (hCG) is used routinely in broodmare practice to induce ovulation. Conflicting evidence and opinions exist as to whether the efficacy of hCG in inducing a predictable, timed ovulation is reduced when the drug is used on an individual mare repeatedly during a single breeding season. Blanchard et al.19 did not note any decrease in efficacy of hCG when administered to mares for up to four estrous cycles within the same breeding season or up to seven different estrous cycles over two consecutive breeding seasons. In contrast, other studies have reported that use of hCG multiple times during a breeding season or intentional immunization of mares against hCG does result in decreased efficacy of subsequent hCG administration in inducing a timed ovulation.20–22 Potential reasons for a failure of hCG to induce ovulation are development of antibodies after repeated administration, inappropriate dose, outdated drugs, administration when a follicle is not sufficiently mature (i.e., too early in estrus), or attempting to induce ovulation of a diestrous follicle or an atretic follicle. It has also been noted that hCG and deslorelin acetate are less predictable when administered to aged mares.23, 24
Hormone therapy-associated ovulation failure Administration of the synthetic progestin altrenogest at a dose of 0.088 mg/kg for 2 days to mares in estrus with a follicle ≥ 35 mm in an attempt to delay ovulation resulted in regression of the dominant follicle in 5 of 26 (19.2%) mares treated.25 In a subsequent study, 2 of 10 mares formed anovulatory follicles when treated with 0.044 mg/kg altrenogest for 2 days beginning in early estrus when a 30 mm follicle was first detected.26 Antagonists of GnRH have also been evaluated as a treatment to delay ovulation in mares in estrus. Intravenous administration of the GnRH antagonist Teverelix (Zentaris AG, Frankfurt, Germany) resulted in development of anovulatory follicles in three of nine mares.26 Administration of corticosteroids during estrus may potentially adversely affect follicular development and ovulation. Asa and Ginther11 reported that treatment of mares with 30 mg dexamethasone once per day beginning 10 days after ovulation was associated with a decrease in LH concentrations and an increase in the incidence of ovulation failure. Similar results were noted following administration of 25 mg dexamethasone twice daily during estrus.12
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However, administration of prednisolone, another corticosteroid hormone, to mares later in estrus was not reported to have any adverse effects on ovulation.27 In addition, administration of 20 mg of dexamethasone twice daily beginning in early estrus did not delay or adversely affect ovulation.15 Hormone therapy regimens designed to superovulate mares also occasionally result in failure of some mares to ovulate.28 Treatment with hCG has been reported to be more effective in inducing ovulation than deslorelin acetate after stimulation of follicular development with equine FSH. Niswender and colleagues29 noted that the ovulation rate for eFSH-treated mares receiving hCG (3.4 ± 0.7) was higher than that of mares administered a deslorelin implant (1.8 ± 0.7) to induce ovulation. Similarly, administration of a compounded deslorelin solution resulted in a lower ovulation rate (2.4 ± 0.8) in eFSH-treated mares than administration of hCG (5.2 ± 0.8) to induce ovulation.30
Prevention, management, or treatment options Several management schemes or therapeutic remedies have been proposed to prevent the occurrence of anovulatory follicles. Unfortunately, most do not work on a majority of affected mares. The sporadic nature and unpredictability of the condition makes it difficult to adequately evaluate treatment programs. In addition, there are no models to reliably induce ovulation failure that can be used to evaluate treatment programs. Consequently, the primary factor inhibiting progress in prevention of ovulation failure is the lack of identification of specific cause(s). Preventive management should begin with alleviation or reduction of proposed risk factors, such as stress and hormone therapy, along with proper diagnosis and treatment of concurrent medical conditions, such as equine Cushings disease.31 In the author’s clinic, a mare with a history of forming anovulatory follicles over repeated cycles was diagnosed with ECD based on elevation of endogenous ACTH. The mare was treated with pergolide and subsequently had estrous cycles in which she had normal ovulations. Unfortunately, administration of pergolide had no positive effect on several other mares with repeated ovulation failure that had no clinical or endocrine evidence of ECD. Options for management of horses with ovulation failure include withdrawal of all hormone agents (i.e., allow for the possibility of spontaneous ovulation), administration of an alternative ovulation-inducing agent (e.g., deslorelin acetate if hCG was not effective), or administration of an
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ovulation-inducing agent earlier in estrus to ‘take control’ before the follicle is committed to atresia. Unfortunately, none of these schemes are uniformly effective in preventing ovulation failure. If a mare does develop a luteinized anovulatory follicle, administration of a single dose of prostaglandin (e.g., 250 µg cloprostenol sodium) 9 days after initial observation of echogenic particles or strands within the follicular lumen will result in the destruction of the luteal cells, a rapid decline in serum progesterone levels (Figure 279.6), and a return to estrus. Prostaglandin treatment has no apparent effect on non-luteinized anovulatory follicles. Fortunately, a majority of non-luteinized anovulatory follicles will spontaneously regress in 1–4 weeks. Administration of hCG or the GnRH agonist deslorelin has not been effective in inducing ovulation or luteinization of this type of anovulatory follicle. Aspiration of a persistent anovulatory follicle using a transvaginal ultrasound-guided approach has been described.32 We have also attempted to eliminate a non-luteinized anovulatory follicle by transvaginal follicular aspiration. In our case, a majority of follicular fluid was successfully removed, but the collapsed follicular structure remained present for more than 2 weeks. Aspiration did not result in an early regression or luteinization of the follicle and suggested that the structure was non-viable or atretic.
Alternative management option Mares that repeatedly form anovulatory follicles during estrus are a major source of frustration for owners, breeding managers, and veterinarians. One option for ‘repeat offenders’ is referral to an assisted reproduction program offering oocyte collection.33 The dominant follicle may be aspirated and the oocyte recovered before the follicle becomes hemorrhagic or atretic. The oocyte may be surgically transferred into the oviduct of a recipient mare that has been inseminated (i.e., oocyte transfer), transferred into the oviduct of a recipient mare along with spermatozoa (i.e., gamete intrafallopian transfer), or injected with a single spermatozoon and the developing embryo transferred into a recipient mare (i.e., intracytoplasmic sperm injection). In one report, ovulation failure was the most common reason cited for mares entering into an assisted reproduction program.34
References 1. McKinnon AO. Ovarian abnormalities. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Wilkins, 1997.
2. Pierson RA. Folliculogenesis and ovulation. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993. 3. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Conversion of a viable preovulatory follicle into a hemorrhagic anovulatory follicle in mares. Anim Reprod 2006;3:29–40. 4. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Incidence, endocrinology, vascularity, and morphology of hemorrhagic anovulatory follicles in mares. J Equine Vet Sci 2007;27:130–9. 5. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541. ¨ 6. Allen WR, Kolling M, Wilsher S. An interesting case of early pregnancy loss in a mare with persistent endometrial cups. Equine Vet Educ 2007;19:539–44. 7. Willis LA, Riddle WT. Theriogenology question of the month. J Am Vet Med Assoc 2005;226:877–9. 8. Steiner J, Antzak DF, Wolfsdorf K, Saville K, Brooks S, Miller D, Bailey E, Zent W. Persistent endometrial cups. Anim Reprod Sci 2006;94:274–5. 9. Hedberg Y, Dalin A-M, Forsberg M, Lundeheim N, Hoffmann B, Ludwig C, Kindahl H. Effect of ACTH (tetracosactide) on steroid hormone levels in the mare. Part A: effect in intact normal mares and mares with possible estrus-related behavioral abnormalities. Anim Reprod Sci 2007;100:73–91. 10. Hedberg Y, Dalin A-M, Forsberg M, Lundeheim N, Sandh G, Hoffmann B, Ludwig C, Kindahl H. Effect of ACTH (tetracosactide) on steroid hormone levels in the mare. Part B: effect in ovariectomized mares (including estrous behavior). Anim Reprod Sci 2007;100:92–106. 11. Asa CS, Ginther OJ. Glucocorticoid suppression of oestrus, follicles, LH and ovulation in the mare. J Reprod Fertil Suppl 1982;32:247–51. 12. Ferris RA, McCue PM. The effects of dexamethasone and prednisolone on pituitary and ovarian function in the mare. Clin Theriogenol 2009;1:225. 13. Macfarlane MS, Breen KM, Sakurai H, Adams BM, Adams TE. Effect of duration of infusion of stress-like concentrations of cortisol on follicular development and the preovulatory surge of LH in sheep. Anim Reprod Sci 2000;63: 167–75. 14. Scholten JA, Liptrap RM. A role for the adrenal cortex in the onset of cystic ovarian follicles in the sow. Can J Comp Med 1978;42:525–33. 15. McKinnon AO, Perriam WJ, Lescun TB, Walker J, Vasey JR, Trigg TE. Effect of a GnRH analogue (Ovuplant), hCG and dexamethasone on time to ovulation in cycling mares. World Equine Vet Rev 1997;2:16–18. 16. Lefranc A-C, Allen WR. Incidence and morphology of anovulatory haemorrhagic follicles in the mare. Pferdeheilkunde 2003;19:611–12. 17. Gastal EL, Gastal MO, Ginther OJ. The suitability of echotexture characteristics of the follicular wall for identifying the optimal breeding day in mares. Theriogenology 1998;50:1025–38. 18. Stangroom JE, Weevers RG. Anticoagulant activity of equine follicular fluid. J Reprod Fertil 1962;3:269–82. 19. Blanchard TL, Varner DD, Schumacher J, Love CC, Brinsko SP, Rigby SL. Manual of Equine Reproduction, 2nd edn. St Louis, MO: Mosby, 2003. 20. Sullivan JJ, Parker WG, Larson LL. Duration of estrus and ovulation time in nonlactating mares given human chorionic gonadotropin during three successive estrous periods. J Am Vet Med Assoc 1973;162:895.
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Disorders of Ovulation 21. Duchamp G, Bour B, Combarnous Y, Palmer E. Alternative solutions to hCG induction of ovulation in the mare. J Reprod Fertil Suppl 1987;35:221. 22. McCue PM, Hudson JJ, Bruemmer JE, Squires EL. Efficacy of hCG at inducing ovulation: a new look at an old issue. Proceedings of the 50th Annual Meeting of the American Association of Equine Practitioners, 2004; pp. 510–13. 23. Barbacini S, Zavaglia G, Gulden P, Marchi V, Necchi D. Retrospective study on the efficacy of hCG in an equine artificial insemination programme using frozen semen. Equine Vet Educ 2000;12:404. 24. Farquhar VJ, McCue PM, Vanderwall DK, Squires EL. Efficacy of the GnRH agonist deslorelin acetate for inducing ovulation in mares relative to age of mare and season. J Equine Vet Sci 2000;20:722–5. 25. Bruemmer JE, Coy RC, Olson A, Squires EL. Efficacy of altrenogest administration to postpone ovulation and subsequent fertility in mares. J Equine Vet Sci 2000;20:450–3. 26. Denniston DJ, Crabb JJ, Bruemmer JE, Squires EL. Effects of a gonadotropin-releasing hormone antagonist and altrenogest on luteinizing hormone concentration and ovulation in the mare. J Equine Vet Sci 2006;26:95–101. 27. Dell’Aqua JA Jr, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Melo CM. Modulation of acute uterine inflam-
28. 29.
30.
31. 32.
33. 34.
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matory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94:270–3. Squires EL, McCue PM. Superovulation in mares. Anim Reprod Sci 2007;99:1–8. Niswender KD, Alvarenga MA, McCue PM, Hardy QP, Squires EL. Superovulation in cycling mares using equine follicle stimulating hormone (eFSH). J Equine Vet Sci 2003;23:497–500. Logan NL, McCue PM, Alonso MA, Squires EL. Evaluation of three equine FSH superovulation protocols in mares. Anim Reprod Sci 2007;102:48–55. McCue PM. Equine Cushing’s disease. Vet Clin Equine 2002;18:533–43. ¨ Bruck I, Larsen SB, Greve T. Transvaginal ultralydoverv˚aget aspiration af en anovulatorisk follikel hos en hoppe. Dansk Veterinærtidsskrift 1995;78:717–20. Coutinho da Silva MA. When should a mare go for assisted reproduction? Theriogenology 2008;70:441–4. Carnevale EM, Alvarenga MA, Squires EL. Use of oocyte transfer in a commercial breeding program to obtain pregnancies from mares with reproductive pathologies. Proceedings of the 45th Annual Meeting of the American Association of Equine Practitioners, 1999; pp. 200–2.
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Chapter 280
Disorders of the Oviduct Irwin K.M. Liu
Relatively little is known about disorders of the oviduct. What are reported are rare anomalies and gross and microscopic examinations of the oviducts from postmortem specimens. Thus, the significance of the abnormalities and their role in infertility of the mare are usually unknown. From a functional and physiological standpoint, few studies of the mare’s oviduct have been performed over the past decades. Yet, the importance of this reproductive structure is undeniable, for it is the site where fertilization and gamete transport takes place in the majority of mammalian species. Clearly, investigations into the survival of embryos during oviductal transport can point to potential functional compromise of the oviduct in the equine.1 Antemortem examination of the equine oviduct is difficult, however, when compared with other animal species. The equine oviduct is long and tortuous and its tissue is dense. Access to the oviduct in vivo is poor, making it difficult for in vivo evaluation, either grossly or microscopically. While the topic of sperm transport and oviductal function is beyond the scope of this discussion, it is notable that although billions of sperm are ejaculated into the uterus of mares during mating, only a few hundred to a few thousand sperm gain entrance to the site of fertilization.2–4 Direct evidence obtained by Scott et al.3 showed that sperm may undergo a selection process at the utero-tubal junction (UTJ), the segment of the oviduct that is most distal from the ovary, forming a papilla-like structure projecting through the lumen of the uterus at the tip of the horn3 (Figure 280.1). Sperm gaining entrance into the oviduct following mating are primarily morphologically normal sperm despite the presence of a heterogeneous population of morphologically normal and abnormal sperm in the ejaculate. The UTJ comprises a significantly large number of folds, which form grooves surrounding the ostium (Figure 280.2). Scott et al.4 provided further evidence of numerous sperm secluded in the folds of the UTJ 4 hours following mat-
ing, and suggested that perhaps the UTJ represented a secondary reservoir for sperm (Figure 280.3). These findings have in recent years generated strong interest in the function of the UTJ and its acceptance as an alternative site for the deposition of semen in the mare.
Structural abnormalities of the oviduct Structural anomalies are rare in the mare. However, there are several reports of a variety of anomalies and these will be discussed briefly as they relate to fertility in the mare. The most common structural abnormality of the oviduct is the presence of accessory oviducts (hydatid of Morgagni). These arise in fetal life and are derivatives of the paramesonephric ducts. The structures exist as small cysts on the infundibulum as they are generally closed at both ends and continue to secrete fluid from the epithelium. When large, they have the potential to prevent adequate cover of the ovulation fossa; however, there is no direct evidence that this does occur. It is unlikely that these structures interfere with fertility as they are often found in normal fertile mares.5 Another structural anomaly of the oviduct are mesonephric or Wolffian duct cysts. These are remnants of the mesonephric duct and are occluded at both ends forming small cysts that lie adjacent to the oviduct. There are no reports or evidence of any relationship to infertility. There have been eight cases of hydrosalpinx reported in the literature. The majority of cases of hydrosalpinx of the oviducts occur unilaterally, with one report of hydrosalpinx due to aplasia of the infundibulum.6 There is one report of a bilateral hydrosalpinx due to an ascending infection.7 Neoplasia of the oviduct is rare. There is one report of a papillary adenoma of the fimbria of a mare.8
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Disorders of the Oviduct
Figure 280.1 Endoscopic view of the utero-tubal papilla (arrow) projecting through the tip of the luminal surface of the uterine horn.
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Nodules of ectopic adrenal cortex have also been reported to occur and are generally found in the mesosalpinx.8 There are several reports of adhesions occurring in the oviduct.9–11 Adhesions are found in approximately 50–70% of reproductive tracts and are most commonly associated with the infundibulum. Whether these adhesions are associated with infertility in the mare is doubtful as Saltiel et al.10 found the adhesions to be present in 72% of pregnant mares. The origin of the adhesions is unknown. However, it is likely associated with blood constituents during the process of ovulation. Thick fibrous bands were also observed in a few oviducts but their significance or origin was not determined. Salpingitis has also commonly been observed in postmortem tracts.9–11 These authors did not find a correlation between adhesions and salpingitis, but found that with increasing age there was an increase in endometritis and inflammation of the infundibulum. Saltiel et al.10 also observed that there was more evidence of lymphocytic infiltration in the infundibular-ampullary region of the oviduct in nonpregnant than in pregnant mares. They further suggested that these lesions may have an influence on the ability of a mare to become pregnant. Although these observations shed light on the importance of investigations of the oviduct, there is no direct evidence that these lesions play a role in infertility in the mare.
Oviductal masses
Figure 280.2 Scanning electron microscopic view of the uterotubal papilla at the utero-tubal junction (UTJ). Note the extensive grooves (papillar folds) present on the surface of the papilla. ×20.
Figure 280.3 Clusters of sperm are frequently found deep within the papillar folds immediately post-mating. ×5000.
In the mid-1960s to early 1970s, the presence of gelatinous masses was described in the oviduct of mares.12–15 Tsutsumi et al., in 1979, also described the presence of gelatinous masses in the oviduct of mares but determined them to be of fibrous origin and termed them fibrous masses.16 These researchers were among the first to observe masses within the lumen of a mare’s oviducts, although the function or significance of these masses was unknown. Subsequent evaluations of these masses demonstrated that they were fibroblastic in origin and composed of type I collagen (Figure 280.4). The masses were primarily lodged at the bends of the tortuous oviduct and at the ampullary-isthmus junction of the oviducts.17 (Figure 280.5). These masses were observed from oviductal flushings to be of various sizes and with microscopic fingerlike projections (Figure 280.7). The projections are probably interdigitations of the masses with the folds of the oviductal epithelium. Further observations by these investigators found occasional large masses that occupied the entire luminal surface of the oviduct at the ampullary-isthmus junction, in some cases causing distention of the lumen (Figure 280.6). Interestingly, the ampullary-isthmus junction is the
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I I MES
Figure 280.4 When examined by scanning electron microscopy, the intraoviductal masses are seen to be composed of bundles of collagen fibrils. ×6000.
site where the oviductal lumen decreases in size from 4–8 mm to 2–3 mm diameter. It seems reasonable to mention that if large masses were to be trapped within the lumen, this is the site where it would logically occur.
In vivo oviductal patency tests Diagnosis of the presence of oviductal masses in vivo is difficult. Attempts have been made to incorporate starch granules and phenosulphonphthalein (PSP) dye as markers, by injecting these substances onto the surface of ovaries and retrieving the substances via the cervix and vagina.1 Similarly, a modified technique using laparoscopic injection of fluorescent microspheres as markers onto the surface of the ovary with subsequent retrieval of the microspheres via uterine lavage was reported.18 Both procedures were proposed as a diagnostic test for the absence
Figure 280.6 Cross-sections of the whole oviduct. The mass of collagen, indicated by the arrows, can be seen as white areas that fill and distend the lumen of the oviduct. The muscular wall of the oviduct is indicated by a bracket. MES, mesentery. ×32.
or presence of oviductal patency in the mare. A concern with the starch grain test for oviductal patency is that the granules recovered from all experimental mares were significantly smaller (6–56 µm) than the size of an oocyte (120–170 µm) despite the placement of starch granules as large, if not larger, than the size of an oocyte. The procedure employed by Ley et al.18 also used only smaller microspheres (15 µm). Thus, these tests do not address whether an ovum can or cannot be transported normally through the oviduct. While these tests may be useful under certain conditions, they do not provide direct evidence that blockage of ova or embryo transport is caused by the collagen masses. In addition, neither of these tests would provide direct evidence of non-patency of the oviduct when the markers are not retrieved from the uterine lumen or cervix. To date, the most direct evidence of oviductal blockage is gained through surgical abdominal invasion with direct exposure of the uterotubal junction and flushing of the oviduct with saline through a blunt-ended 25-gauge needle and retrieving the masses at the fimbrial end of the oviduct.
Treatment for presumptive oviductal mass blockage
Figure 280.5 Masses of collagen, indicated by arrows, within the lumen of the oviduct. Note that the collagen mass completely occupies the oviductal lumen and that the adjacent epithelium of the oviduct is intact. The collagen mass also appears to be interdigitated with the epithelial folds of the oviduct. ×100.
Attempts have been made to dislodge masses from the oviduct in mares infertile for unexplained reasons, and with some subsequent successes in obtaining pregnancy. These results are only indirect evidence that the masses were the cause of infertility in the mare. Several approaches to dislodging the masses have been used. Zent et al. used the abdominal approach under general anesthesia incorporating either a midline incision or a bilateral paramedian approach to gain access to the tips of both uterine horns in the mare.19 After exteriorization, a small
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Figure 280.7 Collagen mass flushed out of the oviduct of an aged mare (×6). Insert: note the fingerlike appearance along the edges of the mass, coinciding structurally with the luminal epithelial folds of the oviduct. (×10).
incision was made through the tip of each uterine horn, manually inverting the tissue to gain access to the UTJ of each horn. A blunt-ended 25-gauge needle was inserted through the UTJ and the oviducts were flushed (using moderate pressure) completely with buffered saline until the saline was flowing easily through the oviducts. In this study the oviducts of eight mares were bilaterally flushed. Oviductal masses from both oviducts were observed in the flushed saline solution of five of the eight mares, and of these, three became pregnant after mating with a stallion. The remaining two mares did not conceive despite repeated breeding efforts. Bilateral oviduct flushings were also attempted in three other mares with unexplained infertility. Extensive adhesions at the UTJ prohibited cannulation of both oviducts in one mare, and two other mares did not conceive despite the apparent absence of oviductal masses from both oviducts following the flushes. Other attempts have been made to flush the oviducts from the ampullary to the isthmus end of the oviduct. However, the luminal diameter of the isthmus is significantly smaller than the ampulla and it may be more difficult to remove larger particles originating from the larger-diameter lumen of the ampulla through the smaller isthmic lumen. Using a normograde flush (ampullary to isthmus) and a modified diagnostic dye test for oviduct occlusion, Bennett et al.20 were successful in attaining pregnancy in 8 of 12 mares presumed to be infertile due to bilateral oviductal blockage. Unfortunately, the authors did not note the absence or presence of collagen masses following the oviductal flushes. Hence a relationship between the presence of oviductal masses and the outcome of pregnancy in the mares could not be determined.
More recently, Allen et al.21 have utilized laparoscopic-guided application of prostaglandin E2 (PGE2 ) (triacetin gel containing 0.2 mg PGE2 ) to the surfaces of the oviducts of 15 mares that failed to conceive over a period of 1–4 years. Fourteen of the 15 mares (93%) were found to be pregnant within the same or subsequent breeding season after the procedure was performed. Again, observance for the absence or presence of oviductal masses following the procedure was not performed in this study.21 Although there have been a number of studies attempting to treat presumptive oviductal blockage, there currently is no direct evidence that collagen masses play a role in infertility in the mare. Studies are needed to evaluate a direct relationship of oviductal collagen masses and infertility in the mare.
References 1. Allen WE, Kessy BM, Noakes DE. Evaluation of uterine tube function in pony mares. Vet Rec 1979;105:364–6. 2. Bader H. An investigation of sperm migration into the oviducts of the mare. J Reprod Fertil Suppl 1982;32:59–64. 3. Scott MA, Liu IKM, Overstreet JW. Sperm transport to the oviducts: abnormalities and their clinical implications. P Annu Conv Am Equine 1995;41:1–2. 4. Scott MA, Liu IKM, Overstreet JW, Enders AC. The structural morphology and epithelial association of spermatozoa at the uterotubal junction: A descriptive study of equine spermatozoa in situ using scanning electron microscopy. J Reprod Fertil Suppl 2000;56:412–21. 5. Kenney RM. A review of the pathology of the equine oviduct. Equine Vet J Suppl 1993;15:42–6. 6. Hinrichs K, Kenney RM, Hurtgen JP. Unilateral hydrosalpinx and absence of infundibulum in a mare. Theriogenology 1984;22:571–7. 7. Hawkins KL. Bilateral salpingitis, hydrosalpinx and oophoritis in a mare. Cornell Vet 1985;76:38–48.
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8. McEntee K. Reproductive Pathology of Domestic Animals. San Diego: Academic Press Inc, 1990; pp. 96–105. 9. Henry M, Vandeplassche M. Pathology of the oviduct in mares. Vlaams Diergeneesk Tijdschr 1981;50:301–25. 10. Saltiel A, Paramo R, Murcia C, Tolosa J. Pathologic findings of the oviducts of mares. Am J Vet Res 1986;47: 594–7. 11. Vandeplassche M, Henry M. Salpingitis in the mare. P Annu Conv Am Equine 1977;23:123–7. 12. Flood PF, Jong A, Betteridge KJ. The location of eggs retained in the oviducts of mares. J Reprod Fertil 1979;57: 291–4. 13. Oguri N, Tsustumi Y. Studies on lodging of the equine unfertilized ova in fallopian tubes. Res Bull Liivestock Hokkaido Univ 1972;6:32–43. 14. Onuma H, Ohnami Y. Retention of tubal eggs in mares. J Reprod Fertil Suppl 1975;23:507–11. 15. vanNiekerk CH, Gerneke WH. Persistence and parthenogenic cleavage of tubal ova in the mare. Onderstepoort J Vet Res 1976;33:195–232.
16. Tsutsumi Y, Suzuki H, Takeda T, Terami Y. Evidence of the origin of the gelatinous masses in the oviduct of mares. J Reprod Fertil 1979;57:287–90. 17. Liu IKM, Lantz KC, Shlafke S, Bowers JM, Enders AC. Clinical observations of oviductal masses in the mare. P Annu Conv Am Equine 1990;36:41–5. 18. Ley WB, Bowen JM, Purswell BJ, Dascanio JJ, Parker NA, Bailey TL, DiGrassie WA. Modified technique to evaluate uterine tubal function in the mare. P Annu Conv Am Equine 1998;44:56–9. 19. Zent WW, Liu IKM, Spirito MA. Oviduct flushing as a treatment for infertility in the mare. Equine Vet J Suppl 1993;15:47–8. 20. Bennett SD, Griffin RL, Rhoads WS. Surgical evaluation of oviduct disease and patency in the mare. P Annu Conv Am Equine 2002;48:347–9. 21. Allen WR, Wilsher S, Morris L, Crowhurst JS, Hillyer MH, Neal HN. Laparoscopic application of PGE2 to re-establish oviductal patency and fertility in infertile mares: a preliminary study. Equine Vet J 2006;38:454–9.
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Chapter 281
Non-neoplastic Abnormalities Donald H. Schlafer
Introduction Lesions within the equine ovary are commonly detected as changes in ovarian size or shape. Nonneoplastic ovarian lesions only rarely cause changes in the reproductive life of mares. This is in marked contrast to the array of endocrinopathies associated with the presence of the most common ovarian neoplasm, the granulosa cell tumor. The dynamic nature of the reproductive process involves rapid and dramatic morphological changes that sometimes make it difficult to differentiate normal ovarian tissues from true lesions. Assessment of ovaries in fetuses or neonates is complicated by the unconventional way in which the equine ovary develops, which is unlike that in any other domestic animal or humans. Knowledge of the major features of gonadal growth, sexual development, and sexual differential in equine fetuses is also important because the majority of non-neoplastic ovarian lesions can be traced back to tissues in the embryo that were involved in ovarian embryogenesis. Knowledge of how sexual development occurs is very helpful in understanding the pathogenesis of the common types of non-neoplastic ovarian lesions – cystic remnants of the mesonephric (wolffian) and paramesonephric (mullerian) systems (Table 281.1). These lesions are very common and are found in fetuses, neonates, and adults; specific identification is usually possible based on their location. Most ovarian lesions are detected during rectal palpation or ultrasonography. The widespread use of ultrasonographic imaging now enables clinicians to detect much smaller lesions that would likely have been overlooked otherwise, thus raising the need to know more about these relatively incidental conditions. Their identification raises questions relating to their origin, development, likely future growth and, most importantly, potential impact on reproduction and the health of the mare.
Other, less common conditions are more likely to be recognized because of changes in the mare’s estrous cycle or during direct examination of ovarian tissue (during surgical procedures, necropsy examination, etc.). Abnormal gonadal development associated with intersex conditions (hermaphrodites), is covered in Chapters 72, 222 and 228. Changes within ovaries include hypoplasia or varying degrees of development of ovarian and testicular tissues (ovotestes) and are often found in mares with abnormal tubular and/or external genitalia. Premature gonadal atrophy (usually unilateral) can result from chronic hypoxic changes associated with reduced blood flow caused by compression of expanding cystic lesions (germinal inclusion cysts), or direct impact on blood flow to the ovary (vascular varicosities, partial or complete ovarian torsion). Rarely, non-neoplastic ovarian lesions can cause more dramatic signs. Mares with large ovarian hematomas, ovarian torsion, or displacement with incarceration of intestine can present with colic or in severe shock. This chapter is an illustrated overview that addresses the appearance and pathogenesis of both the very common and the less common non-neoplastic ovarian lesions. It begins with a brief review of key features in ovarian development in the fetal filly, providing a basis for understanding the origin of the most common cystic lesions found in ovaries of fillies and mares. Table 281.1 contains a list of these lesions, their origins, and diagnostic characteristics. Tumors of the ovary are covered in Chapter 282.
Ovarian development: key features Equine fetal gonadal development is unlike that of any other domestic animal species and is characterized by very dramatic changes in gonadal size, shape,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 281.2 Photomicrograph of interstitial cells in the fetal ovary shown in Figure 281.1. The large hyperplastic interstitial cells are slightly irregular and have started to atrophy. They give the fetal gonad its very rich, dark tan color, which will fade as the cells continue to atrophy and are lost as maturation proceeds. (See color plate 140).
Figure 281.1 Ovaries taken from an equine fetus. The left ovary has been cut in half and the cut surface is exposed. Note the pale whitish ‘cap’ on the right ovary. This represents the area rich in oocytes that will become the ovulation fossa.
and cellular composition. Marked hyperplasia and hypertrophy of interstitial cells in both fetal ovaries and testes cause their rapid growth starting at about 3 months gestation. Gonads develop histological features of either ovaries or testes as early as 39 to 45 days of gestation.1 Fetal equine ovaries are round to ovoid until after birth when, by about 3 months postpartum, they begin to assume their classic reniform shape. Abnormalities in fetal ovarian size or shape, or the presence of structures more typically found in the male are features of intersex conditions. Ovarian size normally increases tremendously starting at about 100 to 140–160 days of gestation. The ovaries continue to grow, increasing their size 20-fold and reaching their maximum size between days 180 to 250 of gestation (Figure 281.1) at which time they are typically larger than the ovaries of their dams. Fetal ovaries are ovoid in shape and, when sectioned, they have a dark-tan appearance (Figure 281.1). Microscopically, interstitial cells are large (Figure 281.2). As they age their natural cytoplasmic pigmentation becomes darker, giving the fetal ovary and testis their normal dark color, which is gradually lost during postnatal maturation. After the initial proliferative stage, fetal ovaries then undergo yet another dramatic change in size, this time rapidly decreasing in size until birth. They are then only about oneeighth their previous maximal size.2, 3
The great change in gonadal size is caused by hyperplasia and hypertrophy of hormone-producing interstitial cells (Figures 281.2 and 281.3). The resulting dramatic enlargement of the fetal ovaries or testes is greatest at about mid-gestation followed by steady regression in their size during the last half of gestation. Fetal gonad interstitial cells are involved in the production of hormones that are important in maintenance of the pregnancy. During about the first third to half of pregnancy, progestagens and estrogens are produced by the mare’s corpora lutea, partly in response to equine chorionic gonadotropin (eCG) produced by endometrial cups. Concurrently, starting at about 60 days gestation, interstitial cells of the fetal gonads produce and release precursor steroids, which are converted to estrone, equine and equilenin by the fetal membranes.4 These cells also
Figure 281.3 Section of a normal fetal ovary (near term). Note the uniform texture and the attached fimbria on the surface at the lower left. The location of the fimbria identifies that area as being the cranial end of the ovary. (See color plate 141).
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Non-neoplastic Abnormalities
Figure 281.4 Photograph of a normal equine ovary taken from a very young foal. Compared to the fetal ovary in Figure 281.1, the gonad here is smaller, the pale cap, which is destined to become the ovulation fossa, is relatively reduced, and the overlying tunica is thicker. This area, where oocytes are concentrated, will continue to become smaller and invaginated, and will become the ovulation fossa of the classically reniform mature equine ovary.
produce large amounts of inhibin.5 Microscopically, interstitial cells appear very similar to fetal hepatocytes. Pathologists examining autolyzed fetal tissue can easily mistake fetal gonads for liver. The oocyte-rich area where the capsule appears lighter colored, involves about half the gonad at mid-gestation then becomes thicker and has a fibrous appearance as the ovary matures and is gradually remodeled (Figures 281.1 and 281.4).
Developmental lesions (and potential confusing features) Ectopic adrenal tissue
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Figure 281.5 The tan mass on the upper right surface of this hemisectioned ovary is ectopic adrenal tissue. This can be identified by its location (on the surface) and its typical tan color. These masses are detected during rectal palpation as firm nodules that are not readily movable. The tissue is adrenal cortical tissue and capsule only. Ectopic adrenal tissue is considered to be an incidental finding.
masculinized or if the ovaries are smaller than normal or have abnormal shape. Figure 281.7 shows the oval-shaped gonads removed from a young intersex mare. Note the attached polypoid structures that are appendix testes, structures normally found only on equine testicles. Fetal ovarian hamartomas Proliferative masses, other than neoplasms, have been reported in the ovaries of young fillies. There are three reports of fetal interstitial cell hamartomas found attached to the surface of the ovary (reviewed
It is not uncommon to find single small nodules of ectopic adrenal tissue attached to the surface of the ovary (Figure 281.5). On cut section, they have a tan appearance. These are considered to be incidental findings. Follicular development in fetal gonads Although not a lesion, a change that can be confusing is the presence of small cystic spaces throughout the gonad of the late-term fetus or the neonatal filly (Figure 281.6). These are antral follicles that tend to start to develop and then regress, but never become tertiary follicles. Intersex conditions Intersex conditions may be associated with abnormal gonadal development and are covered in greater detail in Chapters 72, 222 and 228. One should suspect intersex conditions if the external genitalia are
Figure 281.6 Cross-section of a fetal ovary. The numerous fluid-filled structures are normal developing antral follicles of varying sizes. Follicular development during the late fetal and neonatal period is normal. These follicles do not progress to become tertiary follicles, ovulate, or leuteinize. (See color plate 142).
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F
F F Corpus Hemorrhagicum
Figure 281.7 Hypoplastic ovaries from an intersex mare having some structures that are usually found only with testicles. Note the polyplike structures extending from the surface near the bottom of both ovaries. These are the appendix testes, which are normally found beneath the head of the epididymis in normal male horses. The gonads are small. Epididymal tissues were attached to both ovotestes.
by Machida et al.6 ). Hamartomas are benign tissue growths that appear microscopically as normal appearing tissues for that area from which they come. Identification of interstitial hamartomas in three separate case reports suggests that the equine filly may be prone to development of this condition and that interstitial hamartomas should be considered as a likely diagnosis for any large mass found attached to the ovary. Cystic remnants of structures involved in embryonic sexual development Cystic remnants of structures associated with development of gonads and the tubular genitalia arise from small pieces of embryonic ducts and tubules that persist and become enlarged during later fetal, neonatal, or adult life. Most of these become physically evident when they slowly fill with fluid. They must be differentiated from normally developing follicles, which are always within the ovarian parenchyma (Figure 281.8). Figure 281.9 is a drawing that depicts the location of specific types of cystic remnants and which embryonic tissues they arise from. Table 281.1 also summarizes the types of cysts commonly occurring in, on, or near the ovary. (Note that germinal inclusion
Ovulation Fossa
Figure 281.8 Cross-section of a normal ovary that shows the ovulation fossa at the bottom edge of the section. Three developing antral follicles (each labelled with ‘F’) and regressing corpora lutea are also present. Normal developing follicles can be misdiagnosed as one of several cystic lesions that commonly occur in the mare’s ovary.
cysts are acquired lesions that are also included in Figure 281.9.) The most common cystic remnants arise from the apex of the paramesonephric duct and are commonly referred to as fimbrial cysts. They are always attached to fimbriae of the uterine tube and are commonly bilateral (Figure 281.10). There is high likelihood that one of these cysts will be found in the great majority of all mares if the fimbriae are examined carefully! These cysts occasionally twist around their base and become necrotic. Even though they sometimes reach 1.5 cm in diameter, they are not associated with failure of the fimbriae to retrieve ovulated oocytes. Cystic remnants of mesonephric tubules are less common than remnants of the paramesonephric ducts. Cystic epoophoron occur as either single cysts, or small clusters of cysts loosely attached to the surface of the cranial pole of the ovary (Figure 281.11). These remnants can be quite large, sometimes approaching the size of the ovary itself. They can be differentiated from follicles by their location and their ability to be physically moved short distances over the surface of the ovary. A cystic remnant of a mesonephric tubule at the caudal pole of the ovary is called a cystic paroophoron, and is less common than a cystic epoophoron. Neither affect reproductive performance. They are found in filly fetuses, neonates and adults. The primary clinical relevance of the presence of these various cysts is that they can be confusing and their relative importance not known unless they are recognized as incidental lesions.
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Non-neoplastic Abnormalities
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Cystic Epoophoron
Fimbrial Cyst Hydatid of Morgagni Germinal inclusion cysts Developing gonad
Mesonephric tubules
Mesonephric Duct Cyst
Cystic Paroophoron Paramesonephric duct
Mesonephric duct
Figure 281.9 Cystic remnants of tissues associated with sexual differentiation are very common. This drawing schematically represents embryonic development of the undifferentiated gonad and cranial tubular genitalia. Uterine tubes and the uterus are derived from the paramesonephric ductal system but mesonephric tubules also normally contribute to ovarian development forming the rete and sex cords. Remnants of these structures commonly persist and become cystic as depicted in the drawing on the right. The tissue from which the different cysts arise is indicated by the connecting lines. Additionally, one common acquired cystic condition is also shown. Peritoneal tissues entrapped at the time of ovulation at the ovulation fossa frequently proliferate and produce multiple cysts in this area. They are called germinal inclusion cysts. Modified from Schlafer and Miller.13 Copyright Elsevier Saunders, 2007. Used with permission.
Gonadal hypoplasia Less common developmental conditions include gonadal hypoplasia. This is relative rare in the horse, but is considered to be a heritable trait in other species.
Acquired lesions Some acquired lesions that develop during the otherwise normal reproductive life of the mare have no impact on fertility, others do. Acquired lesions include development of germinal inclusion cysts, transient formation of anovulatory hemorrhagic follicles, formation of fibrinous or fibrous adhesions, vascular varicosities, ovarian infarction, and senile atrophy. Inflammation of the ovary (oophoritis) is rare. Oothecitis Inflammation of the ovary (oothecitis, oophoritis) is uncommon, but occurs occasionally after needle puncture for collection of oocytes.7 Germinal inclusion cysts Figures 281.12 and 281.13 are photographs of clusters of germinal inclusion cysts that develop in the
ovarian parenchyma around the ovulation fossa. The pathogenesis of these cystic lesions involves entrapment of small pieces of peritoneum after ovulation at the surface defect at the site of ovulation. The entrapped tissue is secretory and the transplanted pieces of peritoneum persist forming cystic cavities that expand with time. They are easily confused for developing antral follicles, but their location and numbers are differentiating features. On rare occasions germinal inclusion cysts can become large enough to produce pressure on the ovarian parenchyma and gradually lead to ovarian atrophy.
Hemorrhagic anovulatory follicles Dominant anovulatory follicles occur in about 25% of cycles and are more common in older mares.8 Anovulatory follicles are considered to be a major cause of reproductive inefficiency in the horse.9 Hemorrhagic anovulatory follicles (HAFs), also referred to as persistent anovulatory follicles (PAFs), are common. The pathogenesis associated with their development is poorly understood. Recognition that a HAF has developed in a mare that has just been bred is a source of considerable frustration for clinicians, managers, and owners, and a considerable economic loss.
2702 Surface of ovary
Vascular varicosities
CL, corpus luteum; GCT, granulosa cell tumor.
Accessory corpora lutea of pregnancy
Multiple CLs compressed within ovarian stroma
Small nodules of luteal tissue protrude into the ovulation fossa
Near fossa
Fibrous tags
Follicles Developing corpus luteum
Ovarian follicle
Hemorrhagic anovulatory follicle
Luteinized follicles
Normal corpora hemorrhagica, corpora lutea
Branches of ovarian veins
Small hemorrhage at ovulation
Tertiary follicle
Entrapped peritoneum at ovulation sites
Mesonephric tubules
Mesonephric tubules
Pregnancy
Potential for misdiagnosis as lesion
Geriatric condition
Initially fibrin tags become organized as fibrous tags
Hemorrhagic, granulosa lined
Enlarged ovary, organize and shrink with time
Single or multiple cysts embedded in ovarian stroma at fossa
Movable fluid-filled cyst(s)
Movable fluid-filled cyst(s)
Clear fluid-filled cysts on fimbriae
Usually incidental but can destroy ovary
None unless extreme
Progress in size, can cause pressure atrophy
None
None
None
Frequently associated with masculinization of external genitalia
Clinical effects
Easily mistaken for neoplastic masses
Do not appear to interfere with function of fimbriae
Differential – GCT or hematoma
Can be associated with GCT
Usually incidental but they can destroy the ovary
Common, must be differentiated from developing follicle
Common, must be differentiated from developing follicle
Extremely common
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Normal structures
Ovarian stroma
Hematoma
Caudal surface of ovary
Cystic paroophoron Ovulation fossa
Cranial surface of ovary
Cystic epoophoron
Paramesonephric duct
Varied degrees of differentiation toward testis
Characteristics
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Fimbriae of the uterine tube
‘Fimbrial cysts’
Genetic/chromosomal abnormality
Origin
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Between normal location caudal to kidney to inguinal canal
Intersex (hypoplastic ovary or ovotestis)
Developmental
Location
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Table 281.1 Summary of key features of common non-neoplastic ovarian lesions
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Figure 281.10 Photograph depicting a large paramesonephric fimbrial cyst that arose from a paramesonephric duct remnant. These cysts are common findings. They are always attached to the fimbria and can become quite large and may be multiple.
HAFs (Figures 281.14 and 281.15) develop from bleeding or serum leakage into the follicular lumen. The lumen of the follicles contains blood, gelatinous blood-tinged clotted serum, or a combination of both.10 Most HAFs are lined by luteinized granulosa cells. The incidence of HAFs is estimated to be 5% during the early spring and 20% of estrous cycles later in the season (fall).11 Experimental methods for induction of HAFs by ablation of follicles and treatment with prostaglandins has been reported.12 No difference was found in pre-ovulatory size or
Figure 281.11 Ovary from a young filly with large fluid-filled cysts attached to its apical end. These represent remnants of the mesonephric tubules and are called a cystic epoophoron. They can become as large as the ovary, but do not affect reproduction.
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Figure 281.12 Cysts can also be acquired. In this case, a large number of small cysts can be seen embedded in the ovary at the ovulation fossa, as demonstrated by the arrows. These cysts have developed from pieces of peritoneum that became embedded at ovulation and subsequently proliferated forming cysts that filled with fluid. The cysts are shown in cut section in Figure 281.13.
ultrasonographic appearance of the follicle other than apparent increase in vascularity the day before ovulation failure in follicles that became HAFs when compared to follicles that ovulated normally.11 Non-neoplastic proliferative masses Any ovarian mass should be considered potentially neoplastic. Diagnostic considerations include evaluation of the mare’s reproductive history, and consideration of the location and gross morphological characteristics of the mass. Histopathology and/or ancillary endocrine testing may be required to reach a definitive diagnosis.13
Figure 281.13 This is a cross-section of the ovary in Figure 281.12, showing the cysts in cross-section. They can be small and frequently are multiple. With time they can progress in size and become quite large, leading to destruction of the ovarian parenchyma.
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Figure 281.16 In early pregnancy the ovaries may contain multiple corpora lutea (CL) that tend to compress one another. As shown in this photograph, supplementary corpora lutea can greatly enlarge the ovary. Their appearance is somewhat suggestive of a proliferative, neoplastic process, but supplementary corpora lutea will be found on both ovaries and the mares will be in early pregnancy. The presence of supplementary CLs is a normal feature of pregnancy in the mare.
Supplementary corpora lutea of pregnancy Figure 281.14 An enlarged ovary containing a hemorrhagic anovulatory follicle. The ovary cut open is shown in Figure 281.15.
Luteinization of a wave of follicles that commonly arise in mares after the establishment of pregnancy results in growth of a cohort of corpora lutea (CL) that enlarge both ovaries. These are called supplementary corpora lutea (Figure 281.16). Those supplementary corpora lutea that form from ovulation are termed secondary CLs, and those that form from luteinization of anovulatory follicles are termed accessory CLs. These do look like a neoplasia, but they are usally present in both ovaries and the mare will be pregnant. Their presence, numbers, and appearance varies between mares. Fibrinous and fibrous adhesions A small amount of bleeding following ovulation results in fibrin deposition that can become organized as tougher, fibrous bands that can persist for long periods (Figure 281.17). They are almost always incidental findings and do not interfere with reproduction. Occasionally, localized peritonitis, pyosalpinx, or ovarian abscess can cause marked periovarian fibrosis.
Figure 281.15 Hemi-section of an ovary containing a large blood-tinged fibrin clot within the lumen of a large follicle that failed to ovulate. The contents of such hemorrhagic anovulatory follicles vary and the lumen may be filled with fresh blood. The incidence of hemorrhagic anovulatory follicles is higher both during the early and later ovulatory seasons with more (up to 20% of cycles affected) occurring in the fall, hence they are also referred to as ‘autumn’ follicles.
Ovarian varicosities and angiomatosis Age-related degenerative changes in vessel walls on the surface of the equine ovary can lead to gradual loss of tone resulting in ballooning distension of veins termed varicosities (Figure 281.18). Decreased blood flow within varicose veins, in turn is associated with
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Non-neoplastic Abnormalities
Figure 281.17 Periovarian fibrin or fibrous ‘tags’ are common incidental findings. They result from minor hemorrhage at the ovulation fossa caused by the trauma of ovulation that organizes first as fibrin and later fibrous strands or tags. These can be prominent but they rarely, if ever, interfere with ovulation or movement of the ovulated egg into the oviduct.
thrombus formation, tissue hypoxia, and ovarian atrophy, or, in extreme cases, infarction.13 A proliferative malformation of blood vessels, evident grossly as red nodules scattered over the surface of the ovary, has been diagnosed as ovarian angiomatosis.14
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Figure 281.19 This dark mass is a completely infarcted ovary. The ovary, suspensory ligament, and suspensory ligament of the uterine horn had wrapped around a segment of intestine in a mare presented for intestinal obstruction with colic that required surgical correction. The ovary is very dark from hemorrhage and necrosis.
place near the tips of the uterine horns by a relatively long suspensory ligament along one side and the proper ligament of the ovary at the other end. Figure 281.19 shows a dark, hemorrhagic, autolytic, infarcted ovary that was found twisted with its attachments around a piece of intestine. The mare had colic and had undergone emergency surgery at which time the necrotic ovary was removed.
Ovarian infarction In addition to thrombosis of varicose veins, ovarian infarction rarely develops as a result of physical displacement of the ovary. The ovary is loosely held in
Acknowledgement The illustration in Figure 281.9 was prepared by Mr Michael Simmons, CVM, Cornell University.
References
Figure 281.18 Veins normally found coursing over the surface of the ovary undergo degenerative changes as the mare ages. Segmentally distended veins in this photograph are varicosities that can become much more extensive than this. These lesions tend to be bilateral and progressive over time and appear as black masses on the surface of the ovary. They occasionally thrombose with subsequent ovarian degeneration.
1. Walt ML, Stabenfeldt GH, Hughes JP, Neely DP, Bradbury R. Development of the equine ovary and ovulation fossa. J Reprod Fertil Suppl 1979;27:471–7. 2. Ginther OJ. Embryology and placentation. In: Reproductive Biology of the Mare, Basic and Applied Aspects, 2nd edn. Cross Plains: Equiservices, 1992; pp. 345–418. 3. Hay MF, Allen WR. An ultrastructural and histochemical study of the interstitial cells in the gonads of the fetal horse. J Reprod Fertil Suppl 1975;23:557–61. 4. Saint-Dizier M, Chopineau M, Dupont J, Combarnous Y. Expression of the full-length and alternatively spliced equine luteinizing hormone/chorionic gonadotropin receptor mRNAs in the primary corpus luteum and fetal gonads during pregnancy. Reproduction 2004;128:219–28. 5. Tanaka Y, Taniyama H, Tsunoda N, Herath CB, Nakai R, Shinbo H, Nagamine N, Nambo Y, Nagata S, Watanabe G, Groome NP, Taya K. Localization and secretion of inhibins in the equine fetal ovaries. Biol Reprod 2002;68:328–35. 6. Machida N, Tanaka Y, Taya K, Nakamura T. An ovarian interstitial cell hamartoma in a newborn foal. J Comp Path 2001;125:322–5.
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7. Bogh IB, Brink P, Jensen HE, Lehn-Jensen H, Greve T. Ovarian function and morphology in the mare after multiple follicular punctures. Equine Vet J 2003;35:575– 9. 8. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Siddiqui MAR, Beg MA. Effects of age of follicle and hormone dynamics during the oestrous cycle in mares. Reprod Fertil Dev 2008; 20:955–63. 9. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. 10. Bosu WTK, Smith CA. Ovarian abnormalities. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 397–403.
11. Ginther OJ, Gastal EL, Gastal MO, Beg MA. Incidence, endocrinology, vascularity, and morphology of hemorrhagic anovulatory follicles in mares. J Equine Vet Sci 2007;27:130–9. 12. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Beg MA. Induction of haemorrhagic anovulatory follicles in mares. Reprod Fertil Dev 2008;20:947–54. 13. Schlafer DH, Miller R. Pathology of the female reproductive system. In: Maxie MG (ed.) Jubb, Kennedy and Palmer’s Pathology of Domestic Animals, 5th edn. Elsevier Saunders, 2007; pp. 429–564. 14. Lamm CG, Njaa B. Ovarian and intestinal angiomatosis in a horse. Vet Pathol 2007;44:386–8.
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Chapter 282
Ovarian Neoplasia Claire E. Card
Normal versus neoplastic ovary The equine ovary is a dynamic organ that is affected by factors such as age, photoperiod, nutrition, endogenous or exogenous hormone administration, and disease. Ovarian neoplasia must be differentiated from unusual anatomic structures, physiological changes and other pathological conditions, changes or structures that are seasonally transient or normal. Serial hormonal samples or ultrasonographic examinations may be required to determine if there is any change over time when an ovarian abnormality is suspected. Diagnostic modalities commonly used include transrectal palpation and ultrasonography, hormonal evaluations, endometrial biopsy, exploratory laparoscopy, abdominal surgery, and gonadal biopsy/excision. Histopathological examination of the abnormal ovarian tissue is used to obtain a definitive diagnosis. Ovarian enlargement is often used as an indication of an ovarian tumor; however, neoplastic processes may or may not result in an increase in ovarian size. Equine ovarian tumors include granulosa thecal cell tumors (GTCT), teratomas, cystadenomas, adenocarcinomas, dysgerminomas, arrhenoblastomas, and other metastatic tumors.
The abnormal ovary Ovarian structures commonly confused with tumors Physiological structures that are confused with abnormalities of the ovary or that result in large ovaries include persistent transitional follicles, parovarian cystic structures, ovarian hematomas, and supplementary corpora lutea. Other abnormal structures include ovarian abscesses, hemorrhagic follicles, and non-ovulatory follicles. Excessive ovarian size is sometimes, but not always, associated with tumor formation.1 Ovarian en-
largement in the mare may be identified in the fall and spring transitional periods due to the presence of large static follicles. These large transitional follicles may be associated with erratic estrous behavior. Multiple anovulatory follicular waves typically precede the first ovulation of the year. Persistent autumn or spring transitional follicles may be present for weeks but will eventually regress. Persistent transitional follicles may develop an abnormal ultrasonographic appearance over time characterized by floating echogenic particles and a thick wall. Inadequate support from gonadotropic hormones has been identified as contributing to the formation and persistence of static follicles. Therefore it is encouraged that repeated ultrasonographic examinations of the ovary spaced a few weeks apart are used to show these large static follicles eventually regress. Parovarian cystic structures such as epoophoran or parovarian cysts arising from mesonephric tubules or wolffian duct remnants may be so closely associated with the ovary that it is impossible to identify their origin using ultrasonographic examination. A large cystic mesonephric epophoron remnant called the hydatid of Morgagni may be similar to the size of the ovary.2 Many parovarian cystic structures are identifiable with palpation as not part of the ovary. Treatment with progesterone and estradiol for 2 weeks to induce follicular regression may be helpful to determine if a parovarian cystic structure is present, as epoophoran or parovarian cystic structures are not affected by this hormonal treatment. After 2 weeks of treatment with progesterone and estradiol all follicles are usually < 20 mm in diameter. Epithelial inclusion cysts arise from retention of the surface epithelium within the ovarian cortex following ovulation in the region of the ovulation fossa. Epithelial inclusion cysts tend to be found in mares >15 years of age, and are not found in young mares. They may be found as a solitary cyst or more
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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commonly as a cluster of small follicular-like structures in the vicinity of the ovulation fossa. Grossly these structures range in size from microscopic to a few millimetres in diameter. They are spherical, lined with low cuboidal epithelium, have a tough fibrous capsule, and contain serous fluid. They may interfere with fertility by obstructing the ovulation fossa, or altering the architecture of the ovulation fossa or the ovary itself. These structures are persistent and not affected by seasonal factors.2,3 Hematomas begin as large blood-filled corpora hemorrhagica that enlarge excessively following ovulation (Figures 282.1a and 282.1b). Hematoma formation is the most likely cause of ovarian enlargement during the breeding season in a mare. Very large hematomas may result in visceral tension and discomfort in some mares when palpated. Recurrent colic signs were reported in one mare.4 Mares may react when an ovarian hematoma is manipulated per rectum, hence caution should be exerted when performing a rectal examination in a mare with a history of a large ovary. An ovarian hematoma may initially appear as a large blood-filled structure, but over time usually matures into a structure with a lace-like appearance with strands of fibrin criss-crossing the cavity as the blood clots. The ultrasonographic appearance of a hematoma may closely resemble that of a GTCT (compare Figure 282.1c and Figure 282.3c).5,6 The sexual behavior of a mare with an ovarian hematoma is normal as the hematoma has the same endocrine function as a corpus luteum (CL). The hematoma loses its ability to secrete progesterone following luteolysis or treatment with prostaglandin. Hematoma formation has not been associated with lower conception rates. Hematomas eventually regress but may take a long time to remodel back into the ovary, and may be visualized using ultrasonography for many weeks or even months after it has lost its endocrine function. Pregnancy is typically associated with bilateral ovarian enlargement as a result of continued follicular waves, and the secretion of eCG, which results in the formation of supplementary corpora lutea. Supplementary CL’s are formed from either ovulation (secondary CL’s) or luteinisation of non-ovulated follicles (accessory CL’s). Supplementary CL formation may begin as early as 40 days and follows the pattern of eCG secretion. The number of supplementary CL’s formed is quite variable, and they may be present on one or both ovaries. If a pregnancy is lost after 35 days supplementary CL’s will still form and the ovaries will enlarge. This is important as the breeding history of the mare is sometimes not questioned, or known. In most pregnant mares the corpora lutea begin to regress around 120–180 days.
Figure 282.1a Ovarian hematoma.
Figure 282.1b Cross-section of an ovarian hematoma.
Figure 282.1c Ultrasound image of an ovarian hematoma.
Pregnant mares may sometimes show stallionlike behavior in response to the production of fetal gonadal androgens that may be converted to testosterone by the mare. The stallion-like behavior may include mounting other mares and herding
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Ovarian Neoplasia behavior – the combination of bilateral ovarian enlargement and stallion-like behavior should trigger an evaluation for pregnancy.7 Anovulatory follicles are abnormal, large follicular structures that form in a small proportion of mares during the breeding season. The anovulatory follicles grow to a large size but fail to ovulate, contain echogenic particles, become thick-walled, and appear to luteinize. These structures are not associated with seasonal transitions, and mares prone to this problem tend to repeatedly exhibit the development of anovulatory follicles throughout the breeding season. Some mares have pituitary hypothalamic dysfunction.8–10 Types of anovulatory follicles include: persistent anovulatory follicles (PAF), hemorrhagic anovulatory follicles (AHF), and large non-ovulatory follicles. A low percentage of mares (3.1 – 8%) form PAF.9 True PAF are associated with a 0% pregnancy rate, and administration of an ovulation induction agent is a risk factor for development.9 Ginther et al., 2009 reported a higher incidence of AHF after follicle ablation and prostaglandin treatment.11 The AHF was associated with late emergence of the dominant follicle, along with an inherently high pretreatment LH concentration, and continuing high LH concentrations after prostaglandin treatment. It has been reported using immunohistochemical techniques, that the ovarian cell populations of hemmorhagic anovulatory follicles were not different from control in terms of staining with vascular endothelial growth factor A (VEGF-A). The VEGF-A ligand system is important in the regulation and maintenance of the corpus luteum. There was a relative lack of LH receptors in the AHF corpora lutea, and greatly reduced staining of its tyrosine kinase receptor, Flk-1, in lutenized cells. The reduced Flk-1 may affect the proangiogenic activity of VEGF-A, which may have a contributory role in ovulation failure.12 Large nonovulatory follicles are follicles that reach a preovulatory size (> 30 mm) and then fail to ovulate. The mares may have irregular cycles and the large anovulatory follicle may be present on the ovary for a number of cycles.1 These non-ovulatory follicles may be treated by progesterone and estradiol administration for 2 weeks or follicular aspiration. The aspirated follicular fluid is often yellow to dark brown in color. Large non-ovulatory follicles have been identified sporadically in some mares. Hormonal manipulation of embryo transfer donor mares with equine folliclestimulating hormone (eFSH) has been reported to result in ovarian follicular stimulation without ovulations in around 10% of mares.10 The nonovulatory follicles in non-ovulating eFSH-treated mares may persist for many weeks and result in persistent estrous behavior, and prolonged interestrous intervals.10
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Ovarian abscesses are abscesses that form in the ovary, which have occasionally been identified as a complication after intrafollicular injection. Mares were reported to develop an inflammatory leukogram and abnormal abdominal fluid. Nodules of adrenocortical tissue have been reported to be found associated with the ovary and are not considered to be an abnormality.3
Tumors Ovarian neoplasms that cause enlargement of the ovary have been divided into four classes by the World Health Organization: (1) gonadostromal tumours (granulosa cell tumor, interstitial cell tumour); (2) epithelial tumors (papillary cyst adenoma, papillary adenocarcinoma); (3) mesenchymal tumors (haemangioma, leiomyoma); and (4) germ cell tumors (dysgerminoma, teratoma).13 Mesenchymal tumors such has fibromas and hemangiomas may occur on the ovary or arise as a metastatic extension from other primary tumors. Primary tumors such as adenocarcinomas, melanomas, and lymphomas may metastasize to the ovary from other locations. The history, palpation and ultrasonographic findings, behavior, endocrine results, exploratory surgery, and histopathology are all used to achieve a definitive diagnosis.2,3,14 Table 282.1 shows a summary of the main characteristics of ovarian tumors.
Molecular pathogenesis of ovarian tumorigenesis Various studies of ovarian tumors have revealed some understanding of the molecular pathogenesis of ovarian tumors; however, compared to most types of neoplasia, ovarian neoplasia is not well delineated. There are variations in the frequency of different tumors across species; for example, in mares ovarian epithelial tumors are rare, but they comprise 90% of malignant ovarian tumors in women. Ovarian tumors of epithelial origin (papillary cyst adenoma, papillary adenocarcinoma) Gonadotropin receptors are present in the ovarian surface epithelium. The binding of the gonadotropins, LH and FSH, are linked to cell proliferation, apoptosis, adhesion, invasion, and angiogenesis. There are stem cells present in the ovarian surface epithelium involved in cellular regeneration and differentiation. Ovarian neoplasia of epithelial origin is not linked to mutations of the LH and FSH receptor. It is believed that gonadotropins are involved in the pathogenesis and progression of ovarian neoplasia through their modulation of gene expression, that leads to changes in cell growth, apoptosis, or
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Table 282.1 Comparison of ovarian tumors in mares Embryological origin
Neoplasm
Mare behavior
Malignancy
Hormonal secretions
GTCT
Anestrus, stallion-like, nymphomania
Benign
Increased inhibin > 0.7 ng/mL∗ and testosterone > 50–100 pg/mL∗
Sex cord
Rare but most common tumor
Teratoma
Estrus cycles
Benign
None
Germ cells
Rare second most common
Cystadenoma
Estrous cycles
Benign
None or increased testosterone
Epithelial
Rare
Adenocarcinoma Dysgerminoma Arrhenoblastoma Other metastatic lymphosarcoma, melanoma, adenocarcinoma
Estrous cycles Estrous cycles Estrous cycles Estrous cycles
Malignant Malignant Benign Malignant
None None Increased testosterone None
Epithelial Germ cells Sex cord Variable
Extremely rare Extremely rare Extremely rare Extremely rare
∗
Frequency
Check the reference ranges for the laboratory performing the hormonal analysis.
transformation. The dysregulation of the complex intracellular signaling pathways associated with gonadotropin binding is believed to underlie most ovarian epithelial cell neoplasia.15 GTCT tumorigenesis The proliferation of granulosa cells is controlled primarily by downstream events of FSH receptor binding such as G-protein activation of the phosphoinositide pathway. In GTCT it has been demonstrated that there are no mutations in the FSH receptor or G protein subunits. Dysregulation of the phosphoinositide 3-kinase (PI3-K) signaling pathway is associated with GTCT formation. The downstream effector of PI3K is a serine-threonine kinase Akt, which following activation of PI3-K phosphorylates targets such as kinases, transcription factors, and other regulatory molecules. A protein product of a suppressor gene called Pten is a lipid phosphatase that antagonizes PI3-K function. Dysregulation of this system is associated with cellular transformation.16 In addition a large group of glycoprotein signaling molecules called Wnt appear to play a role in GTCT tumorigenesis. The Wnt is normally involved in embryonic development of the ovary, and has been found to be expressed in the granulosa cells of some species. The expression and hormonal regulation of the Wnt signaling pathway components suggest a role in folliculogenesis. The Wnt signal transduction pathway involves βcatenin as a key effector. The β-catenin is a protein that is involved in mediating cell–cell adhesion by interacting with type II cadherins. In the presence
of the Wnt signal the β-catenin translocates to the nucleus where it modulates the transcriptional activity of a number of genes. Misregulation of Wnt/ β-catenin has been proposed to be a main underlying factor in GTCT tumorigenesis. Archived samples of equine GTCT showed positive expression of β-catenin in the nucleus. Moreover dominant stable transgenic β-catenin mutant mice developed disorganized follicles and cystic structures that later resulted in GTCT in a high proportion of mice.16 Dysregulation of the PI3/Pten/Akt pathway has been shown to be a key regulator of oncogenesis, and when combined with misregulation of Wnt/βcatenin has been reported to be causally synergistic in mouse models of GTCT.16 Granulosa theca cell tumors Granulosa theca cell tumors (GTCTs) are the most common neoplasia of the equine ovary, comprising about 2.5% of all equine neoplasms. They arise from the sex cord–stromal elements of the ovary. The tumors vary in composition and may be primarily composed of granulosa cells, while others contain substantial amounts of theca cells.14,17,18 It has been reported that most GTCT have both neoplastic elements, which is why they are termed granulosa theca cell tumors; other authors refer to the same class of tumor as a granulosa cell tumor (GCT).3,18 The tumors are usually slow growing and the mean age of detection is around 10 years. Maiden, open and pregnant mares may be affected and rarely newborn and young fillies have been reported to have GTCT tumors.19,20 Typically one ovary is affected; however,
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Figure 282.3a Granulosa theca cell tumor.
Figure 282.2 A large granulosa theca cell tumor.
bilateral GTCTs have been reported.21,22 The GTCT is the only ovarian tumor associated with a small inactive contralateral ovary. Regular and irregular follicular activity has been reported prior to the detection of a GTCT.23–26 Historical facts include reports of failure to cycle, unusual sexual behavior (nymphomania or stallionlike), or a change in sexual behavior. Commonly the GTCT results in unilateral ovarian enlargement. Occasionally mares with GTCT are found to have estrous cycles or are pregnant.18 Excessively large tumors may result in colic or other problems in the mare, and tumors in excess of 50 kg have been reported (Figure 282.2). Some mares with GTCT tumors that grow slowly or do not progress may have estrous cycles and become pregnant. Mares with tumors have been reported to carry to term.
Figure 282.3b Cross-section of a granulosa theca cell tumor.
Rectal and ultrasonographic findings The common rectal palpation findings in mare with a GTCT include a hard, enlarged ovary, typically without a clearly defined ovulation fossa, and a small contralateral anestrus-like ovary. The ultrasonographic appearance of the GTCT is classically described as having a honeycomb or cluster of grapes appearance.27 The description arises from the occurrence of multiple cystic spaces present in the tumor in a disorganized fashion. The GTCT may vary in morphological appearance and may be small, almost solid in appearance or may appear as one large follicle (Figures 282.3.a-c). Irregular ovarian activity has been associated with GTCT formation.23
Figure 282.3c Ultrasound image of a granulosa theca cell tumor.
Hormonal findings The tumors are frequently hormonally active. A mare with a GTCT may show one of three main behaviors: anestrus, nymphomania, or stallion-like. The behavior of the mare is usually related to secretory products from the tumor. Inhibin and testosterone
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are most frequently measured.28,29 One or both hormones may be elevated. Secretion of inhibin is association with anestrus behavior or contralateral ovarian shutdown, and testosterone is associated with stallion-like behavior or nymphomania.29 Aromatase is a steroidogenic enzyme that converts testosterone to estrogen.30 Hormonal panels (inhibin, testosterone, and progesterone) are sometimes used to assist in determining if a mare has a GTCT tumor. It has been reported that 85% of mares with GTCT have elevated inhibin levels and 40–70% have elevated testosterone.18,28 Inhibin is a glycoprotein hormone secreted by the granulosa cells of the dominant follicle. Physiologically inhibin is secreted as a dimer with one α-subunit and one of two β-subunits (βA- and βB-subunits).31 The inhibin α-subunit is found in all GTCT, but only some GTCT have both the α-subunit and βA-subunit.32 No GTCT secrete the inhibin βB-subunit. The dimeric inhibin feeds back on the pituitary and decreases secretion of FSH. The dominant follicle in a wave has more FSH receptors and continues to grow and produce estrogen in the face of declining FSH concentrations. During the estrus cycle inhibin increases as the dominant follicle grows.2,33,34 In mares with GTCT excess dimeric inhibin secretion from the tumor has been suggested as a factor suppressing the non-affected ovary (Figure 282.4). The half life of inhibin was reported to be 40 minutes following ovarietomy in a mare with a GTCT.2,34 In summary, only dimeric inhibin has a bioactivity that can suppress FSH, but not all GTCT secrete dimeric inhibin.32 Dimeric inhibin levels have been correlated to tumor mass. It has been reported that mares with a proportionally larger thecal cell component to the GTCT are more likely to secrete testosterone. Testosterone has a short half-life and will fall quickly following removal
of the GTCT. Hour to hour variability in testosterone levels in a mare with GTCT has been reported.25 Testosterone appears to be poorly aromatized to estrogen in GTCT mares, and is not typically elevated even in mares exhibiting nymphomania.25 Generally as one or commonly both hormones are elevated it is recommended that both testosterone and inhibin are measured in GTCT suspect mares.28 Testosterone is mostly likely to be elevated in mares with a masculinized appearance that are exhibiting stallion-like behavior. Exogenous anabolic steroid administration is common in performance mares and includes the use of boldenone undecylenate, nandrolene, winstrol, and testosterone. Boldenone undecylenate, nandrolene, and testosterone are reported to result in stallion-like behavior, disruption of the estrous cycle, and clitoral enlargement in mares, and should be a consideration during testing and examination of mares with a history of stallionlike behavior. Pregnancy also results in a physiological elevation of testosterone and must be ruled out if only testosterone is high in the suspect mare.7 Progesterone is seldom elevated in mares with GTCT because dimeric inhibin or testosterone secreted from the GTCT feeds back on GnRH and prevents FSH secretion, which results in an inactive contralateral ovary. Therefore progesterone levels are low in mares with GTCT unless the mare is one of the few GTCT mares that has estrous cycles and ovulates. Estradiol levels have been reported to be quite variable, but are typically low.25
Endometrial biopsy Examination of endometrial biopsies has been used as a supportive diagnostic aid in a suspect mare. Abnormalities of the uterus have been reported including dysplasia of the endometrium.
Immunohistochemistry
Figure 282.4 Ultrasound appearance of the contralateral ovary in a mare with a granulosa theca cell tumor. White arrow points to the border of the ovary.
Immunohistochemistry has been described as a tool to evaluate ovarian function and equine gynecopathology. Proteins such as glutathione S-transferase α (GSTα), oncoprotein c-erbB-2, and intermediate filaments such as vimentin, desmin, and α-actin have been used in diagnostic pathology. GSTα is involved in steroidogenesis including the synthesis of progesterone and androstenedione. c-erbB-2 oncoprotein (cerb) belongs to a family of growth factor receptors and is believed to be involved in malignant transformation and tumorigenesis. Intermediate filaments are a marker of myofibroblast differentiation. In the future these or other markers of oncogenesis may help determine prognosis. Fewer numbers of
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positively staining cells for vimentin, cerb, and GSTα were found in one malignant GTCT.12
Treatment and prognosis Surgical removal of the GTCT tumor is generally recommended and is indicated especially in mares with objectionable behavior, large tumors, or in brood mares. Small tumors have been removed via colpotomy, flank laparotomy, and a variety of laparoscopic approaches and laparoscopically assisted procedures. Large tumors are generally removed via a ventral midline celiotomy. Complications related to GTCT include tumor rupture and hemoperitoneum, hemorrhage at surgery, adhesions to other visceral bodies, colic, colonic rupture secondary to impaction, hypertrophic osteopathy in mares with metastasis, ovarian torsion, diabetes, and tumor metastasis (usually to regional pelvic or abdominal lymph nodes).35–38 While unusual, metastasis of a GTCT usually carries a poor prognosis as it usually involves regional lymph nodes, liver, spleen, lung, mesentery, and omentum.38 Following removal of the tumor most mares return to regular estrous cycles within a year. However, some former GTCT mares have been reported to remain anestrus for years, and some never return to cycle.28 Early gonadotropin therapy after surgical removal has not been effective; however, hormonal stimulation may still be a possibility for mares that fail to return to cyclicity after a year. It has been reported that pituitary responsiveness to GnRH in mares with GTCT was similar to non-affected mares. Baseline levels of FSH were not different between GTCT mares and non-GTCT mares, suggesting that testosterone may exert negative feedback on GnRH and may be the underlying cause of contralateral ovarian shutdown, rather than inhibin.39 There is only one report of spontaneous reactivation of a previously inactive ovary in a mare with a contralateral GTCT. Return to fertility is highly unlikely in mares without removal of the GTCT.
Figure 282.5 Cross-section of an ovarian teratoma showing different tissue elements including hair, cartilage, and fat.
have been associated with abdominal discomfort.40,41 (Figure 282.5). Cystadenoma Ovarian cystadenomas are rare epithelial tumors that arise from the epithelium of the follicle, medullary or cortical embryonic cords, rete ovarii, infundibula or the ovarian serosa.28,42,43,44,45 Examination of the ovarian anatomy shows that the superficial or germinal epithelium lines the ovulation fossa. Cystadenomas often resemble persistent follicles during serial ultrasound examinations (Figure 282.6a and 282.6b). They are sometimes called serous cystadenomas to describe the secretion present in the cystic spaces of the tumor which varies from a thin and serous to thick and mucinous fluid. These tumors are not typically secretory however they have been reported to be associated with elevated testosterone,28,44 without contralateral ovarian suppression. There is one report of a serous cystadenoma in a pregnant mare.45 Histologic examination of the cystic areas shows a single
Teratoma Ovarian teratomas are considered rare in the mare. They are composed of pluripotential, also called totipotential, germ cells that undergo neoplastic transformation into two or more of the germinal cell types (endoderm, ectoderm, and mesoderm). These different tissue types are often arranged in a bizarre fashion in the tumor and may include adipose, epidermal, dental, musculoskeletal, or nervous tissue. These tumors vary in size and sometimes are reported as incidental findings; other larger tumors
Figure 282.6a Ovarian cystadenoma.
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Mare: Diseases of the Ovary and Oviduct entiate them from non-germ cell malignancies such as GTCT. Arrhenoblastoma Arrhenoblastomas are extremely rare benign secretory sex cord tumors that are reported to secrete testosterone. This tumor is also called an androblastoma and a Sertoli-Leydig cell tumor of the ovary due to its histological appearance. Histologically tubules are present and they are lined by Sertoli cells with clusters of Leydig cells. There is only one report of this type of tumor in a mare.48
Figure 282.6b Cross-section of an ovarian cystadenoma.
Hamartomas layer of cuboidal epithelial cells that are sometimes ciliated.
Hamartomas are very rare tumor-like, non-neoplastic disordered proliferations of mature tissues, such as blood vessels. A vascular and interstitial cell hamartoma of the equine ovary have been reported.49–51
Adenocarcinoma Adenocarcinoma is an extremely rare malignant tumor of the ovary. They arise from the surface epithelial cells and are non-secretory. Histologically there may be unilocular or multilocular cysts with papillary projections of epithelium. These tumors metastasize by rupture and leakage of cyst contents into the peritoneal cavity, where the metastasis or eventual lymphatic obstruction may lead to ascites, rather than invasion of blood vessels. Cystic structures may be present on cut section.43,46
Dysgerminoma Dysgerminomas are exceedingly rare malignant tumors that are non-secretory. The dysgerminoma arises from germ cells. In the mare, dysgerminoma may be a finding on routine post partum examination but more often is associated with weight loss, and is reported to rapidly metastasis to the abdomen and thorax.13,47 If weight loss is present along with unilateral ovarian enlargement suggestive of a tumor, thoracic radiographs and ultrasonography of the thorax and abdomen and or abdominocentesis may be considered to assist with the prognosis and to rule metastasis in or out. Metastasis to the thorax has been associated with hypertrophic osteopathy. Histological features include cords, sheets, and nests of large uniform cells separated by connective tissue.3 Using transmission electron microscopy, convoluted nucleoli were found. Cells of the dysgerminomas are round to polygonal, and have positive histochemical staining for alkaline phosphatase and vimentin using immunohistochemistry. These features help to differ-
Lymphosarcoma While there are reports of uterine lymphosarcoma, ovarian lymphosarcoma is extremely rare, and is most often metastatic.52
References 1. Westermann CM, Parlevliet JM, Meertens NM, van Oldruitenborgh-Oosterbaan MMS. Enlarged ovary in a mare: review of the literature. Tijdschrift Voor Diergeneeskunde 2003;128(22):692–696. 2. McKinnon AO. Ovarian Abnormalities, in Equine Diagnostic Ultrasonography. Norman Rantanen AOM (ed.) Williams and Wilkins: Baltimore, 1997; pp. 233–251. 3. Jubb K, Kennedy P, Palmer N (ed.). Female Genital System, Pathology of Domestic Animals, Vol. 2. 4 ed. Academic Press: Toronto, 1993; pp. 364–369. 4. Lampe A. Recurring colic in a mare with an ovary hematoma. Praktische Tierarzt 1997;78(9):744–&. 5. Curtin D. Ovarian hematoma in an 11 year old Thoroughbred Hanoverian mare. Can Vet J 2003:44(7):589– 591. 6. Montavon S. Ultrasonography of ovarian pathology in the mare – a review for the practitioner. Schweizer Archiv Fur Tierheilkunde 1994;136(9):285–291. 7. Silberzahn P, Zwain I, Marin W. Concentration increase of unbound testosterone in plasma of the mare through pregnancy. Endocrinol 1984;115(1):416–419. 8. Troedsson MHT, McCue PM, and Macpherson ML. Clinical aspects of ovarian pathology in the mare. Pferdeheilkunde 2003;19(6):577–584. 9. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. in Equine reproduction VIII. Proceedings of the Eighth International Symposium on Equine Reproduction, Fort Collins, USA, July, 2002. Elsevier Science Inc. 10. McCue P, LeBlanc M, Squires E. Therio., eFSH in clinical practice. Theriogenology 2007;68(3):429–433.
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Ovarian Neoplasia 11. Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Beg MA. Induction of haemorrhagic anovulatory follicles in mares. Reproduction, Fertility and Development, 2008;20(8):947– 954. 12. Ellenberger C, Bartmann CP, Hoppen HO, Kratzsch J, Aupperle H, Klug E, Schoon D, Schoon HA. Histomorphological and immunohistochemical characterization of equine granulosa cell tumours. Journal of Comparative Pathology 2007;136(2–3):167–176. 13. Harland CS, Mogg T, Horadagoda N, Dart A. Surgical resection of a dysgerminoma in a mare. Aust Vet J 2009;87(3):110–112. 14. Bosu W (ed.). Ovarian Abnormalities. Equine Reproduction, McKinnon AO, Voss JL (eds). Lea & Febiger: Philadelphia. Vol. 1. 1993; pp. 397–407. 15. Choi J, Wong A, Huang H, Leung P. Gonadotropins and ovarian cancer. Endocrine Reviews 2007;28(4):440– 461. 16. Boerboom D, Paquet M, Hsieh M, Liu J, Jamin S, Behringer R, Sirois J, Taketo M, Richards J. Misregulated Wnt/B-Catenin signaling leads to ovarian granulosa cell tumor development. Cancer Research 2005;65(2):9206– 9215. 17. Raoofi A, Mardjanmehr SH, Masoudifard M, Adibhashemi F, Asadian P. Thecoma in a mare. Journal of Equine Veterinary Science 2006;26(12):588–591. 18. McCue P, Roser J, Munro C, Liu I, Lasley W. Granulosa cell Tumors of the Equine Ovary. Vet Clin Equine 2006;22:799–817. 19. Hultgren B, Zack P, Pearson E, Kaneps A. Juvenile granulosa cell tumor in an equine weanling. J Comp Pathol 1987;97(2):137–142 20. Charman RE, McKinnon AO. A granulosa-theca cell tumour in a 15-month-old Thoroughbred filly. 2007;85(3):124–125. 21. Turner T, Manno M. Bilateral granulosa cell tumour in a mare. J.A.V.M.A. 1983;182:713–714. 22. Frederico L, Gerard M, Pinto C, Gradil C. Bilateral occurrence of granulosa-theca cell tumors in an Arabian mare. Can Vet J 2007;48:502–505. 23. Chopin JB, Chopin LK, Knott LM, de Kretser DM, Dowsett KF. Unusual ovarian activity in a mare preceding the development of an ovarian granulosa cell tumour. Australian Veterinary Journal 2002;80(1–2):32–36. 24. Nie GJ, Momont H. Ovarian mass in 3 mares with regular estrous cycles. Journal of the American Veterinary Medical Association 1992;201(7):1043–1044. 25. Stabenfeldt G, Hughes J, Kennedy P, Meagher D, Nealy D. Clinical findings, pathological changes and endocrinological secretory patterns in mares with ovarian tumours. J Reprod and Fertil 1979;27:277–285. 26. Stradaioli G, Zelli R, Sylla L, Monaci M, Leonardi L, Maurizi M. Clinical findings, diagnostic methods and surgical approach in mares affected with granulosa-theca cell tumour. Ippologia 2001;12(3):15 27. Hinrichs K, Hunt P. Ultrasound as an aid to diagnosis of granulosa cell tumour in the mare. Equine Vet J 1990;22:99–103. 28. Liu IKM. Ovarian tumors. Current therapy in equine medicine. Robinson RN (ed.), Saunders: Philadelphia, 1983; pp. 408–409. 29. Bailey M, Troedsson M, Wheaton J. Inhibin concentrations in mares with granulosa cell tumors. Theriogenology 2002;57:1885–1895.
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30. Watson ED, Thomson SRM. Immunolocalization of aromatase P-450 in ovarian tissue from pregnant and nonpregnant mares and in ovarian tumours. Journal of Reproduction and Fertility 1996;108(2):239–244. 31. Nagamine N, Nambo Y, Nagata S-I, Nagaoka K, Tsunoda N, Taniyama H, Tanaka Y, Tohei A, Watanabe G, Taya K. Inhibin secretion in the mare: localization of inhibin α, βA, and βB subunits in the ovary. Biol Reprod 1998;59:1392–1398. 32. Hoque S, Derar RI, Senba H, Osawa T, Kano K, Taya K, Miyake Y. Localization of inhibin alpha-, beta Aand beta B-subunits and aromatase in ovarian follicles with granulosa theca cell tumor (GTCT) in 6 mares. Journal of Veterinary Medical Science 2003;65(6):713– 717. 33. Donadeu FX, Ginther OJ. Effect of number and diameter of follicles on plasma concentrations of inhibin and FSH in mares. Reproduction 2001;121:897–903. 34. Hoque MS, Senba H, Tsunoda N, Derar RI, Watanabe G, Taya K, Osawa T, Miyake Y. Endocrinological changes before and after removal of the granulosa theca cell tumor (GTCT) affected ovary in 6 mares. Journal of Veterinary Medical Science 2003;65(8):887–891. 35. Gift LJ, Gaughan EM, Schoning P. Metastatic granulosa – cell tumor in a mare. Journal of the American Veterinary Medical Association 1992;200(10):1525– 1526. 36. Rambags B, Stout T, Rijkenhuizen A. Ovarian granulosa cell tumours adherent to other abdominal organs; surgical removal from 2 Warmblood mares. Equine Vet J 2003;35(6):627–632. 37. Sedrish S, McClure J, Pinto C, Oliver J, Burba D. Ovarian torsion associated with a granulosa-theca cell tumor in a mare. J.A.V.M.A. 1997;211:1152–1154. 38. Alexander GR, Tweedle MA, Lescun TB, McKinnon AO. Haemoperitoneum secondary to granulosa cell tumour in two mares. Aust Vet J 2004;82(8):481–484. 39. Zelli R, Sylla L, Monaci M, Stradaioli G, Sibley LE, Roser JF, Munro C, Liu IKM. Gonadotropin secretion and pituitary responsiveness to GnRH in mares with granulosatheca cell tumor. Theriogenology 2006;66(5):1210– 1218. 40. Panciera R, Slusher S, Hayers K. Ovarian teratoma and granulosa cell tumor in two mares. Cornell Vet 1991;81:43–50. 41. Lefebre R, Theoret C, Dore M, Girard C, Laverty S, Vaillancourt D. Ovarian teratoma and endometritis in a mare. Can Vet J 2005;46:1029–1033. 42. Held JP, Buergelt C, Colahan P. Serous cystadenoma in a mare. J.A.V.M.A. 1982;181:496–498. 43. Son Y, Lee C, Jeong W, Hong I, Park S, Kim T, Cho E, Park T, Jeong K. Cystadenocarcinoma in the ovary of a Thoroughbred mare. Aust Vet J 2005;83:283– 284. 44. Hinrichs K, Fraser G, deGannes R, Richardson D, Kenney RM. Serous cystadenoma in a normally cyclic mare with high plasma testosterone values. J.A.V.M.A. 1989;194(89):381–382. 45. Bridges ER. Serous cystadenoma in a pregnant mare. Equine Practice 1994;16(5):15–17. 46. Van Camp S, Mahler J, Roberts MC, Tate LP, Whitacre MD. Primary ovarian adenocarcinoma associated with teratomatous elements in a mare. J.A.V.M.A. 1989;194:1728–1730.
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47. Chandra A, WJ, Merritt A. Dysgerminoma in an Arabian filly. Vet Pathol 1998;35:308–311. 48. Mills J, Clark E, Fretz P, Ganjam V. Arrhenoblastoma in a mare. J.A.V.M.A. 1977;17(1):754–757. 49. Rhyan JC, D’Andrea GH, Smith LS. Congenital ovarian vascular hamartoma in a horse [first report]. Veterinary Pathology, 1981;18(1):131.
50. Foley GL, Johnson R. A congenital cell hamartoma of the equine ovary. Veterinary Pathology 1990;27(5):364–366. 51. Misdorp W. Congenital tumours and tumour-like lesions in domestic animals. 3. Horses – A review. Veterinary Quarterly 2003;25(2):61–71. 52. Locke TF, Macy DW. Equine ovarian lymphosarcoma. J.A.V.M.A. 1979;175:72–73.
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283.
Abnormalities of Cervical and Vaginal Development John P. Hurtgen
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Cervix Adhesions Patricia L. Sertich
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Failure to Dilate Ahmed Tibary
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Chapter 283
Abnormalities of Cervical and Vaginal Development John P. Hurtgen
Abnormailities of the cervix Developmental abnormalities of the mare’s cervix are exceptionally rare. Very few reports are in the literature concerning congenital cervical abnormalities. The types of congenital abnormalities include partial to complete absence of a cervix and partial to complete duplication of the cervix. A thorough reproductive examination, including an internal manual palpation of the cervix, should detect the majority of these congenital defects of the cervix. Identification of these abnormalities is important because these defects are invariably associated with infertility or even sterility. Complete absence of a cervix has been reported in a mare with normal ovarian function.1 Mares with chromosomal abnormalities, such as 63XO, 64XY, 65XXY, and others, usually have gonadal dysgenesis. The affected mares have very small ovaries and an underdeveloped, open cervix. Cytogenetic evaluation of blood or tissue samples should be performed from mares suspected of developmental abnormalities of the cervix, uterus, or ovaries.2 The lack of development of the cervix or uterus in cases of gonadal dysgenesis may be the result of a lack of hormonal stimulation. In an embryo transfer program, pregnancies were established in XO mares following supplementation of these mares with estrogen and progesterone.3 Cervical hypoplasia has been reported in a 2-year-old Thoroughbred filly and in a 4-year-old Arabian filly.4, 5 Both mares has normal ovarian function but had persistent pneumovagina. The cervix in each case measured 1–1.5 cm in length and remained atonic and open despite the mare’s estrous cycle status. Cervical hyperplasia is also extremely rare. A reported case was presented as a possible uterine pro-
lapse in a 5-year-old maiden Thoroughbred. The cause of the hyperplasia was unknown but inflammation was excluded as a cause.6 Complete or partial duplication of the cervix has been reported in mares owing to incomplete fusion of mullerian ducts. The majority of the reported cases and cases observed in clinical practice have been in draft horse mares: Clydesdale, Shire, and Suffolk Punch. Each cervical canal may be complete and attached to a single uterine horn. The uterine horns may or may not communicate.7, 8 A cervical canal may also end blindly.9 During artificial insemination and intrauterine treatment procedures, caution must be taken to assure that the cervical canal allows uterine deposition of semen or treatments. Parturition is usually uneventful in mares with a double cervix but normal uterine body and horns. Mares with partial or complete cervical duplication may be infertile owing to the frequently associated uterine abnormalities. Occasionally, a mare may have a double cervix with each canal connected to independent uterine horns.7 These mares should be sterile owing to the inability of an embryo to migrate throughout the uterus and to the lack of uterine space for pregnancy development. Embryo migration prevents the systemic prostaglandin release associated with luteal tissue regression. However, the author has had a Clydesdale mare with separate cervical canals associated with a single uterine horn attached to each cervix. The horns did not communicate. After repeated failure of the mare to become pregnant, the mare was bred and received oral altrenogest in an attempt to maintain pregnancy. The mare delivered a very small, viable foal at term with no complications at parturition. In an embryo transfer program this mare was a consistent embryo donor and accounted for three foals in a single year. Under experimental conditions the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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restriction of embryo migration resulted in pregnancy loss between day 13 and day 17 of gestation in some but not all mares and was related to degree of embryo restriction.10 Extreme tortuosity of the cervix is occasionally encountered in the mare. The cervical canal does not appear to relax during estrus. The affected mares also tend to accumulate varying amounts of fluid in the uterine body during estrus. These mares may become pregnant and foal uneventfully if uterine fluid accumulation is prevented and semen is deposited in the uterus. Artificial insemination should be advantageous in affected mares compared with live cover. Congenital abnormalities of the cervix are rare in the mare. Careful manual evaluation of the reproductive tract should identify affected mares before mating. The majority of affected mares are infertile, or even sterile. Therapy is usually unrewarding.
Vaginal abnormalities Persistent hymen is a common developmental abnormality of the posterior vaginal cavity. Persistent hymen occurs because of the failure of the caudal sections of the mullerian duct to fuse with the urogenital sinus. The persistent hymen can be seen at the junction of the vestibule and vagina. Clinically, the condition is usually a single, thin band of tissue that is easily ruptured during manual examination of the vagina, artificial insemination procedures, or live cover by a stallion. In these cases, fertility is not compromised. Rarely, the persistent hymen can completely occlude the junction of the vestibule and
vagina. Mares with a complete persistence of the hymen may accumulate mucus in the vaginal cavity or even the uterus. The complete persistent hymen can usually be manually ruptured or surgically removed. Persistent hymen is usually documented in the maiden mare. However, a complete hymen may rarely be observed in mares that have been barren following natural service by a stallion.
References 1. Schlotthauer CF, Zollman PE. The occurrence of so-called ‘white heifer disease’ in a white Shetland Pony mare. J Am Vet Med Assoc 1956;129:309–10. 2. Bowling AT, Millon L, Hughes JP. An update on chromosomal abnormalities in mares. J Reprod Fertil Suppl 1987;35:149–55. 3. Hinrichs K, Riera FL, Klunder LR. Establishment of pregnancy after embryo transfer in mares with gonadal dysgenesis. J In Vitro Fert Embryo Transf 1989;6:305–9. 4. Blanchard TL, Evans LH, Kenney RM, Hurtgen JP, Garcia MC. Congenitally incompetent cervix in a mare. J Am Vet Med Assoc 1982;181:266. 5. Allen WE. A cervical anomaly in an Arabian filly. Equine Vet J 1981;13:268–9. 6. Riera FL, Hinrichs K, Hunt RR, Kenney RM. Cervical hyperplasia with prolapse in a mare. J Am Vet Med Assoc 1989;195:1393–4. 7. Blue MG. An uterocervical anomaly (uterus bicorpor bicollis) in a mare, and the manual disruption of early bilateral pregnancies. N Z Vet J 1985;33:17–19. 8. Volkmann DH, Gilbert RO. Uterus bicollis in a Clydesdale mare. Equine Vet J 1989;21:71. 9. Macrae DR. Double os uteri in a mare. Vet Rec 1935;15:1100. 10. McDowell KJ, Sharp DC, Peck LS, Cheeves LL. Effect of restricted conceptus mobility on maternal recognition of pregnancy in mares. Equine Vet J Suppl 1985;3:23–4.
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Chapter 284
Cervix Adhesions Patricia L. Sertich
Anatomy The mare’s cervix has a cylindrical shape that is formed by an inner circular smooth muscle layer rich in elastic fibers and an outer longitudinal smooth muscle layer. These muscle layers are continuous with the muscle layers of the vagina and uterus. A thickened circular muscle layer forms the body of the portio vaginalis, which is the caudal portion of the cervix that protrudes into the anterior vagina. The cervical inner circular muscle is surrounded by submucosa and a highly folded, fern-like mucosa. A simple columnar epithelium lines the lumen of the cervix and contains many mucigenous cells including goblet cells. There are longitudinal folds lining the cervical lumen that are continuous with the endometrial folds of the uterus. Each dorsal and ventral longitudinal fold may extend into the vagina as the dorsal and ventral frenulum.1 Inexperienced examiners often incorrectly identify the dorsal or ventral frenulum as an adhesion. The vaginal epithelium is stratified squamous epithelium.
Etiology The epithelium is frequently exposed to trauma and must be renewed. The renewal is accomplished by cell regeneration. A basic principle of epithelial healing is that if cells are not inhibited by contact with like cells or with their normal underlying or overlying connective tissue they tend to migrate and proliferate.2 There is a tendency for traumatized cervical epithelium to adhere transluminally or to the vagina when healing. Stage II labor in the mare is rapid, and if the fetus passes through the birth canal before the cervix has fully dilated there is potential for the cervix to tear. Lacerations can be partial, affecting only the muscular layer, or full thickness, affecting all three layers. Both types result in a defect in the cervix and
loss of proper cervical function. When the traumatized tissue heals spontaneously there is rarely reconnection of a damaged muscular layer. The overlying epithelium can form transluminal adhesions which can prevent the cervix from properly relaxing and allowing uterine secretions to drain normally. A lacerated edge of the portio vaginalis can also adhere to the vaginal epithelium, resulting in a cervix that cannot fully close. If there has been a prolonged dystocia and the cervical tissue experiences pressure necrosis there is a possibility that the portio vaginalis or a portion of the portio vaginalis will slough and or adhere to the anterior vaginal wall. When cervical adhesions attach to the vaginal wall the cervix may not be able to close adequately during early pregnancy, resulting in loss of the early embryo. If there is some degree of initial cervical closure the competency of the cervix may become inadequate after 3 months of gestation when the gravid uterus moves cranially out of the pelvic canal into the abdomen, resulting in the cervix being pulled cranially. If these vaginal cervical adhesions prevent adequate cervical closure the pregnant mare is at risk for developing an ascending placentitis in late gestation. Some substances infused into a uterus for therapeutic resolution of endometritis can be caustic to the cervix and vagina. Uterine infusions containing chlorhexidine, iodine, acriflavin, and tetracyclines have been associated with severe endometrial inflammation (Figure 284.1).3 If the cervical epithelium becomes necrotic and sloughs, transluminal adhesions can form and completely prevent anything from entering (semen) or leaving (uterine secretions and post-breeding debris) the uterus (Figure 284.2).
Diagnosis Although severe cervical adhesions may be apparent on a visual (speculum) vaginal examination,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 284.1 Severe inflammation of the cervix subsequent to administration of sulfa urea uterine boluses that also contained acriflavin into a mare’s uterus.
thorough evaluation of the cervix must be made by a manual (digital) examination per vaginum. While many theriogenologists prefer to perform vaginal and uterine procedures on mares during estrus it may be prudent that cervical evaluations be performed during diestrus when the cervix is closed, has good shape and tone, and its competency can be determined. Mares in general tolerate this procedure well and can be restrained in hand or placed in a stock. The mare should be prepared for examination by wrapping the tail to contain the tail hairs and prevent hair from being carried into the vagina. The genital tract should be examined by palpation per rectum to ascertain that the mare is not pregnant. If the examiner can feel the endometrial folds of the uterus one can assume that the mare is not more than 2–3 weeks pregnant. It is contraindicated to perform a vaginal cervical examination in a pregnant mare as the intrusion to the genital tract may lead to an ascending placentitis. Feces should also be removed from the rectum at this time to decrease the chance of defeca-
Figure 284.2 Transluminal adhesion after severe cervicitis subsequent to administration of sulfa urea uterine boluses containing acriflavin.
tion during the vaginal examination. The perineum should be cleansed thoroughly with povidine scrub or gentle soap and water prior to vaginal examination. After a thorough rinse with clean water the excess rinse water can be removed with a clean dry towel. The examiner should then don a sterile plastic examination sleeve and sterile surgical glove. Nonirritating water-soluble sterile lubricant should be applied to the examination hand and arm to facilitate passage through the caudal genital tract. The forefinger should be placed in the lumen of the cervix and the entire wall (muscle layer and both mucosal layers) of the cervix should be palpated between the thumb and forefinger. All 360◦ of the portio vaginalis should be present and not attached to the vaginal mucosa. The cranial cervical canal should be patent and free of adhesions.
Management Complete resolution of cervical adhesions is rarely accomplished regardless of the treatment. If only a few filamentous web-like transluminal adhesions are present and the mare is not having problems with chronic endometritis and fluid accumulation, the web-like adhesions can simply be manually disrupted prior to natural cover or artificial insemination. The mare should be carefully monitored ultrasonographically for evidence of uterine fluid accumulations and treated appropriately should fluid accumulate. A uterine lavage tube that contains an inflated cuff may be positioned just cranial to the cervix to help drain uterine fluid. Traction can be placed on the lavage tube while the uterus is being lifted and massaged to help evacuate uterine contents. Oxytocin (5–10 IU) can be administered during this procedure to contract the uterus. Long-term placement of an indwelling catheter to aid uterine drainage generally is not effective in resolving uterine fluid accumulations or preventing the reformation of transluminal adhesions. These filamentous web-like adhesions will recur, but if the mare becomes pregnant these thin adhesions will not interfere with parturition. Mares may have a cervical laceration with an adhesion of the lacerated flap of portio vaginalis to the anterior vagina. If the laceration involves more than 40–50% of the length of the cervix, the cervix should probably be surgically repaired. The portio vaginalis adhesion must be carefully dissected from the vaginal wall before suturing the debrided laceration edges. One should avoid excising the anterior vaginal artery that courses along the ventral aspect of the cervix and caudal uterine body. This artery will tend to retract into the tissue and be difficult to
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Cervix Adhesions ligate successfully. Local chemical or electrical cautery may be required to achieve hemostasis. One surgical cervical laceration correction technique is to place a tension suture on the caudal aspect of the cervix and attach the tension suture to the hymen or vulva to keep the cervix suspended until the laceration incision has healed.4 Alternatively after a threelayer closure of the cervical laceration one may also consider leaving a long loop of a retraction suture on each side of the incision line so that the retraction suture loop extends out of the vulva. These two retraction sutures can then be used to pull and lift the portio vaginalis off the vaginal floor two to three times a day to attempt to reduce the amount of adhesions that form post-operatively between the portio vaginalis and the anterior vagina. After several days one strand of each loop can be excised and the suture removed. One may apply a soothing ointment that contains corticosteroids between the cervix and vaginal wall to help reduce the formation of adhesions. If the cervix does not appear to be lacerated but the portio vaginalis is adhered to the vaginal floor, one may carefully dissect the portio vaginalis from the vaginal floor and place two retraction sutures in the caudal aspect of the portio vaginalis to allow twice daily manipulation to discourage adhesions from reforming. Although daily manipulation of the portio vaginalis and application of a steroid-containing ointment may prevent the immediate formation of adhesions, some degree of adhesions will regrow within several days after the manipulations are discontinued. There is no effective treatment to resolve severe transluminal adhesions that are present along the entire length of the cervix. Many of these cases have a concurrent chronic pyometra since the uterus can-
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not drain. If a pyometra is not present initially, one is likely to develop if the severe transluminal adhesions are disrupted as there is a great risk that bacteria will be trapped cranial to the adhesions. These mares would not be able to carry a pregnancy on their own or even produce an embryo for embryo transfer but they would be candidates for donation of oocytes for transfer into recipients. Mares with severe transluminal adhesions and a concurrent pyometra may be clinically stable for years, but some mares occasionally may exhibit colic if the uterus gets severely distended.5 Such a case may be managed by intermittent drainage and lavage of the uterus. One may even consider hysterectomy to eliminate the pyometra and colic in some cases. Once formed cervical adhesions are difficult to eliminate. One should always keep the long-term function of the cervix in mind whenever a mare’s cervix is being invaded, be it for a routine examination, uterine therapy, or dystocia resolution.
References 1. Ginther OJ. Reproductive Biology of the Mare, 2nd edn. Cross Plains, WI: Equiservices, 1992; p. 29. 2. Peacock EE. Wound Repair, 3rd edn. Philadelphia: W.B. Saunders, 1984; pp. 15 and 30. 3. Prescott JF. Sulfonanamides, diaminopyrimidines, and their combinations. In: Prescott JF, Baggot JD, Walker R (eds) Antimicrobial Therapy in Veterinary Medicine. Ames, IA: Iowa State University Press, 2000; pp. 290–314. 4. Evans LH, Tate LP, Cooper WL, Robertson JT. Surgical repair of cervical lacerations and the incompetent cervix. In: Proceedings 25th Annual Convention American Association of Equine Practitioners, 1979, pp. 483–6. 5. Tranquillo GG, Kelleman AA, Sertich PL. Theriogenology Question of the Month, JAVMA 2009;235:1161–1164.
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Chapter 285
Failure to Dilate Ahmed Tibary
The normal equine cervix is approximately 5 to 7 cm in length and 3 to 4 cm in width. Unlike other domestic species, the mare’s cervix does not have the typical cartilaginous cervical rings but is rather formed by longitudinally arranged folds that are continuous with the endometrial folds (Figure 285.1). These folds are continued dorsally and ventrally into the vaginal fornix by prominent frenula (Figure 285.2). In the female donkey (Jenny) the fornix is deep as the cervical os projects more caudally into the vagina (Figure 285.3).1 Histologically the cervix is composed of a thick layer of circular smooth muscle rich in elastic fiber and a highly folded mucosa lined with an epithelium containing mucus-producing cells. The anatomy and histology of the equine cervix provides for a relatively easy manual dilation at any stage of the cycle. Cervical dilation is observed as a response to high levels of estrogen during estrus and during the last couple of weeks of pregnancy. Cervical dilation at the end of pregnancy and during parturition reaches its maximum during delivery of the fetus through the combined effect of pressure from the fetus and maximal response of the muscle fibers within the ripened cervix. Cervical ripening during parturition is brought about by a synchronous variation of hormonal influences involving estrogen, relaxin, and prostaglandin E. Dilation of the cervix during estrus is a prerequisite for successful breeding. The maximally dilated cervix allows the stallion to deposit the ejaculate into the uterus. Following mating or insemination, cervical dilation also helps with uterine clearance.
Causes of cervical dilation failure Failure of cervical dilation may be congenital or acquired. The extreme form of failure of cervical dilation is cervical hypoplasia or agenesis, which has
been described but is relatively rare. This condition often results in the development of a mucometra (Figure 285.4). Congenital extreme tortuosity of the cervix has also been described as a cause of failure of dilation during estrus and poor fertility due to impaired fertilization and/or uterine clearance. Other described cervical anomalies that may compromise cervical dilation include double cervix (uterus bicollis),2–4 varicose veins, and thrombosis5 (Figure 285.5). Acquired cervical masses such as leiomyomas6 or hyperplasia7 may also lead to failure of cervical dilation. Adhesions are often the result of cervical injury, either during foaling or as a consequence of excessive manipulation. Cervical lacerations and adhesions can be observed following an apparently normal easy delivery and have been suspected to be due to failure of sufficient dilation or ‘rushed cervix’.8 Obstetrical manipulations may also be the cause. Recently, the author has seen several cases of cervical injury leading to adhesions, following overt manipulation to remove glass marbles introduced into the uterine cavity for prevention of cyclicity. Adhesions of the cervix (Figure 285.6a–c) form following rearrangement of the fibrin matrix which initially develops and contains macrophages fibroblasts and giants cells. Fibrin is slowly replaced by fibroblasts and collagen which eventually give rise to fibrous bands containing neovascularization.9 Attempts to breaks down these adhesions are often followed by increasing trauma and subsequent formation of further adhesions. Fibrosis of the cervix is probably the most common cause of cervical dilation failure. Loss of elasticity of the cervical anatomy is believed to be due to aging and to be more severe in older maiden mares.10 Also, clinical impressions from practicing veterinarians suggest that mares that have not had opportunity to cycle during the normal physiological season
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 285.2 Anatomy of the cervix: frenulum of the cervix (top) and extending longitudinal folds into the vaginal fornix (arrows).
Indications and therapies for cervical dilation Figure 285.1 Anatomy of the cervix: longitudinal cervical folds (arrows) extending from the endometrium of the body of the uterus (top of the image) toward the cervical canal.
may be more prone to develop cervical fibrosis. It would be interesting to see if long-term suppression of cyclicity via progesterone treatment or GnRH immunization would result in increased predisposition to cervical dilation failure. Equine practitioners have suggested that mares with Cushing disease may be prone to failure of cervical dilation.
V
The most common indications for pharmacological enhancement of cervical dilation are improvement of cervical drainage for the management of recurrent pyometra or persistent-mating induced endometritis and access to the uterine cavity for fetal removal. In recent years the overuse of intrauterine glass marble for estrus control in performance mares has led to an increased presentation of mares for removal of these devices when the mare does not respond or when other negative effects are observed (Figure 285.7). Enhancement of cervical ripening before induction
C
(a)
(b)
Figure 285.3 Anatomy of the cervix in the jenny. (a) Protrusion of the cervix (C) into the vaginal cavity (V) forming a deep fornix (arrow); (b) long cervical canal (arrows) showing longitudinal folds.
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Mare: Problems of the Cervix or a combination of both. Mechanical techniques are successful if the cervix is normal. The author has used dilators (bougienage) in cases of cervical stenosis in combination with local hormonal treatment (prostaglandin E, PGE). Forceful mechanical cervical dilation should not be attempted if the cervix is extremely tight. The most common hormonal treatments used for cervical dilation are estrogens, PGE, or their combination.
Mechanism of action of PGE (a)
(b)
(c) Figure 285.4 Ultrasonograms of a mare with mucometra due to external cervical os aplasia. (a) Body of the uterus; (b) cranial aspect of the cervix; (c) small cervix visualized by vaginoscopy (note absence of external cervical os opening).
of foaling has been suggested to improve delivery time and to reduce the risks of premature placental separation and intrapartum foal hypoxia.11 However, more studies are needed in order to confirm this hypothesis. The most common techniques used for enhancement of cervical dilation are mechanical, hormonal,
The induction of cervical ripening and labor by local application of PGE1 and PGE2 (Figures 285.8 and 285.9) was established in the late 1970s and early 1980s in humans. These properties of PGE have made it a primary choice for induction of cervical ripening, abortion, and labor in women over the last three decades.12, 13 More recently, similar results were obtained in other species including non-human primates,14 and bovine,15 ovine,12 and equine11 species. The cervical ripening, clinically noticed as a softening and dilation, corresponds to remodeling of the extracellular matrix, which is dominated by molecules such as collagens and proteoglycans. Mechanisms involved in preparation of the cervix for full dilation have been studied recently at the molecular level. These mechanisms involve complex changes in gene expression in response to modulators including hormonal changes during the last part of pregnancy. Cervical ripening was believed to involve an inflammatory process but this hypothesis was recently debated because there was no correlation between cervical ripening and interleukin-8 (IL-8) concentration in cervical tissue. Cervical ripening does not improve with infiltration with neutrophils and macrophages. Among the several factors involved, the most commonly recognized are expression of mRNA collagen, proteoglycans, metalloproteinases, as well as other gene expressions.16–18 Up to 85% of the composition of the cervical extracellular matrix is fibrils of type I and III collagen. An increase in the expression of mRNA for collagen types IV, V, and VI is observed at the end of pregnancy, while collagen types I and III decrease in the cervix at term. Collagen type IVα2 is the major structural component of basement membranes and interacts with laminin and proteoglycans. Collagen V has been implicated as a critical determinant of fibrin structure and matrix organization. An increase in collagen turnover is observed during ripening of the human cervix, leading to remodeling and a different collagen organization with less cross-linking, which contribute to a looser arrangement of the extracellular matrix. Two classes of proteoglycans play an
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Figure 285.5 (a) Varicose veins around and within the cervix; and (b) corresponding ultrasonography image showing an area of increased vasculature and hyperechoic fibrotic tissue (arrows).
(a)
(b)
(c) Figure 285.6 Cervical adhesions (a) external aspect of the cervix with severe adhesions resulting from trauma (b) intra cervical adhesions (c) cervical adhesion extending into the internal cervical os and the body of the uterus.
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Mare: Problems of the Cervix O OH
O
HO
OH
Figure 285.9 Structure of PGE2 , dinoprostone (C20 H32 O5 , mol. wt. 352.465).
Figure 285.7 Ultrasonographic image of glass marble (arrows) within the uterine cavity.
important role in regulation of cervical biometrics. The small leucine-rich proteoglucans (SLRP; decorin, buglycan, and fibromodulin) regulate collagen fibril formation, and the large aggregating proteoglycans (LAP; aggrecan and vesicant) regulate tissue structure through the glycoaminoglycan (GAG) chains conjugated to the core protein.17, 18 Versican, a large extracellular matrix proteoglycan, was found to be upregulated several folds in the ripe cervix. Due to its high capacity to attract water and bind to hyaluronan, versican may promote collagen disorganization and decrease cell adhesions within the extracellular matrix, resulting in increased softness and elasticity.16, 17 Metalloproteinases (MMPs) play a major role in regulation of the extracellular matrix and are possible mediators in cervical remodeling by cleaving constituents of this matrix. MMP-3 has been shown to cleave fibrinogen, cross-linked fibrin, E-cadherin as well as fibronectin. MMP-2, 8, and 9 have been found to increased several-fold in the cervix at term and in the immediate postpartum period in women.16 Simi-
O
O CH3
OCH3
OH
HO O
O OH
CH3
OCH3
HO Figure 285.8 Structure of PGE1 , misoprostol (C22 H38 O5 , mol. wt. 382.5).
lar changes of MMP-1 and 2 have been described during cervical ripening in the bovine cervix.15 Another group of enzymes, ADAMTS (a disintegrin and metalloproteinase with thrombospondin motifs), is thought to participate in the degradation of the extracellular matrix by degrading aggrecans (aggrecanases).17 A novel gene, ‘myocardin’, has been detected and seems to be involved in cervical ripening, and could be involved in alterations in the smooth muscle component of the cervix. Although several hormones (estrogen, progesterone, prostaglandins, and glucocorticoids) are proposed to be implicated in the cervical ripening process, their exact role remains poorly understood. Focus has been on changes in progesterone receptors (PR) and estrogen receptors (ER) at the level of the cervix. In humans, protein levels of ER and PR are decreased in term pregnancy. Estrogen stimulation via ERß through release of cytokines and MMPs and inhibitions of AP-1-regulated collagenase transcription has been suggested as a possible mechanism facilitating cervical dilation.16, 19 The mare’s cervix contains measurable amounts of ER and PR but their role in cervical dilation remains unstudied.20 PGE may transduce its signal through G proteincoupled receptors (EP receptors) which are regulated by steroid and, in particular, estrogen. Ovariectomized estrogen-treated ewes show decreases in EP1 and EP3 receptor protein expression, which may promote better cervical dilation by enhancing EP2 and EP4 receptor-dependant vasodilation and subsequent edema and facilitate smooth muscle relaxation.21
Results in the equine The softening effect of PGE on the equine cervix was first reported by Volkmann and DeCramer 1991.13 These authors demonstrated that intracervical application of PGE2 in pregnant mares (80 to 300 days) resulted in a rapid and complete cervical softening within 30 to 90 minutes and facilitated induction of abortion. In a second study by the same authors, PGE2 (prostin E2) was found to cause cervical dilation in diestrous and peripartal mares.22 PGE2 application was also associated with an increase in
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Failure to Dilate myometrial contractions lasting 30 to 60 minutes.22 It is important to note that, in this study, although application of PGE2 induced cervical softening, fetal expulsion did not occur unless the mares were treated with oxytocin following PGE2 . PGE followed by oxytocin resulted in rapid expulsion of the placenta when compared to manual fetal removal which resulted in a high rate of retained placenta. More recently, PGE2 (2 to 2.5 mg in sterile saline) injected using a 25 gauge needle into the external cervical os, or instilled in the cervical canal using a Cassou gun, resulted in cervical dilation within 6 hours in term mares.11 Mares prepared in this manner required less oxytocin for induction of parturition and had shorter delivery period. PGE1 (misoprostol 1000 µg in cream) has also been used to promote cervical dilation and uterine clearance as part of management of mares with persistent mating-induced endometritis. However, there are no controlled studies on the dosage, method of delivery, and efficacy.23 PGE has also been shown to have the same effect on the smooth muscles of the utero-tubal junction papilla.24
Preparation of PGE Several human preparations of PGE1 (misoprostol, R R R Cytotec ) and PGE2 (Prepidil , Cervidil , Prostin E) have been used in mares. Commercial human preparations in the form of vaginal insert, tablets, gel or cream have been utilized to help dilate the equine cervix. In the equine, tablets are dissolved in saline and infused into the cervical canal using a pipette, or applied to the cervical canal and external cervical os after mixing with a sterile gel. No clinical study has been performed in mares to evaluate the efficacy of these products. In humans, it has been shown that some of the variability in response observed after application of PGE is due to difference in the vaginal pH and its effect of the bioavailability of the hormone.25 Prostaglandins are organic acids that have diminished solubility in aqueous solution with a low pH. In one study25 on the efficacy of the dinoprosR tone gel (Prepidil ) (the most widely used PGE2 in the United States), it was noted that women with a high vaginal pH (> 4.5) had significantly shorter times to active labor, complete dilation, and vaginal delivery compared to women with a low pH (< 4.5). The pH may alter the properties of the carrier as well as the absorption and/or metabolism of the active ingredient. The optimal pH for PGE2 release from a triacetin-based PGE2 gel and hydrogel PGE2 pessary was shown to be near neutral (7.4). In vitro studies
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have shown that PGE2 release from hydrogel pessary is enhanced when pH increases.25
Other treatments Estradiol cypionate has been given at doses ranging from 1 mg to 6 mg to improve cervical dilation in noncyclic and estrous mares. Others have reported local application of estrogen creams. However, none of these treatment has been critically evaluated.
Conclusion Complete cervical dilation can be obtained with PGE in the normal pregnant mare and facilitates induction of abortion or foaling. PGE application to mares with cervical stenosis due to aging, or old maiden mare syndrome, seems to work if applied during estrus. Pretreatment with estrogen may be helpful in the non-cyclic, non-pregnant mare. It is doubtful that this treatment alone would be very helpful with severe cervical adhesions. Prevention of cervical adhesions following injury can be obtained with daily application to the cervical lumen of steroid–antibacterial cream containing prednisolone, such as that used in treatment of mastitis in the cow.
References 1. Tibary A, Sghiri A, Bakkoury A. Reproduction. Duncan J and Hardrill D (eds). In: The Professional Handbook of the Donkey, 4th edn. Whittet Books: The Donkey Sanctuary, Sidmarth, Devon, UK, 2008; pp. 314–41. 2. Blue MG. A uterocervical anomaly (uterus bicorpor bicollis) in a mare, and the manual disruption of early bilateral pregnancies. N Z Vet J 1985;33:17–19. 3. Kelly GM, Newcombe JR. Uterus bicorpora bicollis as a possible cause of infertility in a mare. Vet Rec 2009;164: 20–1. 4. Volkmann DH, Gilbert RO. Uterus bicollis in a Clydesdale mare. Equine Vet J 1989;21:71. 5. Foster RA, Gartley CJ, Newman S. Varices with thrombosis in the cervix and uterus of a mare. Can Vet J 1997;38:375–6. 6. Romagnoli SE, Momont HW, Hilbert BJ, Metz A. Multiple recurring uterocervical leiomyomas in two half-sibling Appaloosa fillies. J Am Vet Med Ass 1987;191:1449–50. 7. Riera FL, Hinrichs K, Hunt PR, Kenney RM. Cervical hyperplasia with prolapse in a mare. J Am Vet Med Assoc 1989;195:1393–4. 8. Embertson RM, Henderson CE. Cervical tears. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 130–3. 9. Estrada AJ, Samper JC. Cervical failure to dilate. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St. Louis, MO: Saunders Elsevier, 2007; pp 134–6. 10. Pycock JF. Cervical function and uterine fluid accumulation in mares. Equine Vet J 1993;25:191.
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11. Rigby S, Love C, Carpenter K, Varner D, Blanchard T. Use of prostaglandin E2 to ripen the cervix of the mare prior to induction of parturition. Theriogenology 1998;50:897–904. 12. Candappa I, Bainbridge H, Price N, Hourigan K, Bartlewski P. A preliminary study on the suitability of Cervidil to induce cervical dilation for artificial insemination in ewes. Res Vet Sci 2009;87:204–6. 13. Volkmann D, DeCramer K. Prostaglandin E2 as an adjunct to the induction of abortion in mares. J Reprod Fertil 1991;Suppl. 44:722–3. 14. Shimizu K, Jin WZ, Kishi H, Noguchi J, Watanabe G, Taya K. Effect of prostaglandin E2 in ripening o fthe cervix and secretion of relaxin, luteinizing hormone (LH), estradiol-17b and progesterone at term pregnancy in the Japanese monkey (Macaca fuscata fuscata). J Reprod Dev 2002;48:607–11. 15. Van Endelen E, Breeveld-Dwarkasing V, Tavern M, Everts M, Van Der Weijden GC, Rutten V. MMP-2 expression precedes the final ripening process of the bovine cervix. Mol Reprod Dev 2008;75:1669–77. 16. Ekman-Ordeberg G, Stjernholm Y, Wang H, Stygar D, Shlin L. Endocrine regulation of cervical ripening in humans: potential roles of gonadal steroids and insulin-like growth factor-I. Steroids 2003;68:837–47. 17. Hassan S, Romero R, Tarca A, Nhan-Chang C, Vaisbuch E, Erez O, Mittal P, Kusanovic J, Mazaki-Tovi S, Yeo L, Draghici S, Kim J, Uldbjerg N, Kim C. The transcriptome of cervical ripening in human pregnancy before the onset of labor at term: Identification of novel molecular functions involved in this process. J Maternal Fetal Neonatal Med 2009;22:1183–93.
18. Ji H, Dailey T, Long V, Chien E. Prostaglandin E2regulated cervical ripening: analysis of proteoglycan expression in the rat cervix. J Obstet Gynecol 2006;198:536 .e1–e7. 19. Slater D, Astle S, Woodcock N, Chivers J, de Wit N, Thornton S, Vatish M, Newton R. Anti-inflammatory and relaxatory effects of prostaglandin E2 in myometrial smooth muscle. Mol Hum Reprod 2006;12:89–97. 20. Re G, Badino P, Novelli A, Direnzo G, Severino L, Deliguoro M, Ferone M. Distribution of cytosolic estrogen and progesterone receptors in the genital tract of the mare. Res Vet Sci 1995;59:214–18. 21. Schmitz T, Levine B, Nathanielsz P. Localization and steroid regulation of prostaglandin E2 receptor protein expression in ovine cervix. Reproduction 2006;131:743– 50. 22. Volkmann D, Bertschinger H, Schulman M. The effect of prostaglandin E2 on the cervices of dioestrous and prepartum mares. Reprod Domest Anim 1995;30:240–4. 23. Nie G, Barnes A. Use of prostaglandin E1 to induce cervical relaxation in a maiden mare with post breeding endometritis. Equine Vet Educ 2003;15:172–4. 24. Allen W, Wilsher S, Morris L, Crowhurst J, Hillyer M, Neal H. Lapaoscopic application of PGE2 to re-establish oviductal patency and fertility in infertile mares: a preliminary study. Equine Vet J 2006;38:454–9. 25. Ramsey P, Ogburn PL Jr, Harris D, Heise R, Ramind KD. Effect of vaginal pH on efficacy of the dinoprostone gel for cervical ripening/labor induction. Am J Obstet Gynecol 2002;187:843–6.
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Chapter 286
Pre-Foaling Mammary Secretions William B. Ley
The clinical test of measuring and monitoring prefoaling mammary secretion (PFMS) electrolytes does not predict the actual foaling time of the mare, and should not be expected to. As with many biological systems, variations can and will occur. Few clinical tests consistently, accurately, and reliably predict future events. In the hands of clients, PFMS monitoring gives them something more active to do than ‘just watch’, and it gets them closer to the mare in more ways than one. The testing procedure discussed here, along with all other signs of the mare’s approaching readiness for birth, gives the clinician the ability to effectively evaluate the in utero fetal maturation response. Furthermore, it provides data upon which an informed decision might be made as to when it is appropriate to induce foaling in a safe and effective manner.
Pre-foaling electrolyte changes in mammary secretions It is common knowledge that most mares foal during the night. Many sleepless hours and days can therefore be spent waiting for the mare to foal. A distribution of spontaneous foaling times for over 1100 mares is depicted in Figure 286.1.1 Physical signs of impending birth are helpful but not very accurate as a means of predicting when the mare will enter stage 2 labor. Accepted physical signs of the mare’s approaching readiness for birth include a gestation length >320 days, udder engorgement with presence of colostrum or milk in the teats, waxing, and relaxation around the tail head, buttocks, and lips of the vulva. Studies of PFMS electrolyte changes have demonstrated an association with the mare’s approaching readiness to foal.2–14 These electrolyte changes, especially with regard to calcium, have also been related
to the development of in utero fetal maturity and subsequent survivability of the fetus following spontaneous foaling.4, 9, 11 Calcium carbonate (CaCO3 ) testing of PFMS samples has been shown to be both sensitive and specific with respect to spontaneous foaling.11 The predictive value of a positive test (PVPT), defined as the first occurrence on which CaCO3 ≥200 ppm, was determined to be 97.2% for mares foaling spontaneously within 72 hours. Concentrations of potassium, calcium, citrate, and lactose have been reported to increase, and the concentration of sodium to decrease as foaling becomes imminent, but variation between mares was noted to be quite large.14 These authors concluded that the use of PFMS electrolyte concentrations for prediction of time of foaling is unreliable. This is absolutely correct. None of these methods have yet been shown to predict time of foaling. Ley et al.11 stated that PFMS calcium carbonate testing is a helpful prognostic tool to indicate the mare’s approaching readiness for birth. They further stated that it can be predicted that the mare is unlikely to foal within 24 hours when the CaCO3 is < 200 ppm. Ousey et al.4 suggested that, according to their results, fetal maturity may be related to electrolyte concentration in PFMS, and that an ionic score of ≥35 may indicate that induction would be successful in terms of maturity of the newborn foal. Leadon et al.3 stated that calcium concentrations of PFMS proved useful in predicting full term, and also in assessing the chances of foal survival following induced parturition. Ousey et al.,10 in a later report, stated that PFMS electrolyte testing prior to foaling was not particularly accurate in predicting time of parturition, although it was a reliable means of indicating when it was not necessary to attend prepartum mares at night. It is reasonable to agree
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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140 Number 120 100 80 60 40 20 0 Noon
Midnite
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Figure 286.1 The distribution in a 24-hour period of spontaneous foaling times for over 1100 mares. Adapted from Bain and Howey.1
with Douglas et al.14 that these tests do not predict foaling time. Very few investigations have actually claimed that they do. PFMS electrolyte testing methods are simply a means to evaluate in utero fetal maturation and the approaching readiness for birth.
carbonate is 40% of the total molecular weight). To convert from ppm to mmol/L, multiply by 0.01. All pre-foaling mammary secretion testing is done with a dilution of one part sample and six parts distilled water. Evaluation and sampling PFMS should begin approximately 10–14 days in advance of the mare’s expected foaling date, calculated as 335–340 days from the last known breeding date. It is best to keep a close check on udder development on a daily basis and practice massage of the mare’s udder and teats to allow her to get used to being handled in this area. In mares with an unknown breeding date, testing should begin as soon as some udder enlargement is noted and a small amount of PFMS can be obtained from the teats. Once-a-day sampling is sufficient until values exceed 100 ppm. Thereafter, twice-daily sampling is recommended. The most accurate assessment of the mare’s readiness to foal will be gained from daily sampling from late afternoon to early evening. Since a few mares will foal in the daytime (Figure 286.1), a morning sample should not be neglected, and one is highly advisable when CaCO3 values exceed 125 ppm.
Test kits for detecting PFMS changes The FoalWatchTM kit (CHEMetrics, Inc., Calverton, VA 22016) has been used extensively over many years by the author and has proven useful in the routine management of mares in the prepartum period. Its intended use is to assist the determination as to when the fetus is likely mature in utero and that the mare is approaching readiness for spontaneous foaling. This is based upon the kit’s ability to detect changes in the PFMS CaCO3 level. It is a tool that when used in an appropriate manner will allow one to attend the mare’s foaling without an excessive number of sleepless nights. Alternatively, it can be used as a tool for accurately assessing when elective induction of parturition may safely be attempted. The advantages of this methodology include its accuracy and repeatability compared with other test kits or test strips on the market, its ease of use, its quantitative (numerical) determination of the CaCO3 level in each sample of PFMS tested, and its economy. The FoalWatchTM test kit measures uncorrected calcium carbonate (CaCO3 ). Comparison to other methods of pre-foaling mammary secretion testing must be performed with caution, as semantic errors may contribute to misinterpretations. For example, the tests are commonly referred to as a ‘milk calcium test’; the implication is calcium carbonate, but may be misunderstood as calcium only. A test result value of 250 ppm CaCO3 would be equated to 100 ppm calcium (2.5 mmol calcium/L). To convert ppm CaCO3 to ppm Ca, multiply by 0.4 (i.e., calcium in calcium
Sampling method Wipe the udder and teats with a clean, dry, soft paper towel prior to attempting to collect a sample. This reduces the amount of skin debris and dirt that might contaminate the sample, although such contamination will not alter the test result to any appreciable extent. This step is strongly recommended if the mare has been out in the weather and is wet. The hands of the person doing the sampling also should be clean and dry. A sample volume of 2–5 mL PFMS should be obtained per mare. It should be collected in a clean manner by gently stripping a small amount from each teat, using thumb and index or middle finger, into a clean plastic test tube or other sampling cup. The cup or test tube can be reused if it is thoroughly rinsed with distilled water and then dried between uses. The sample should then be taken to a clean, dry, and warm area for testing. A syringe is used to draw-up exactly 1.5 mL of the sample. The measured sample is placed into a mixing vial, and exactly 9.0 mL of distilled water is added. This dilution (i.e., ratio of one part PFMS to six parts distilled water) of the PFMS is important since it adjusts the calcium level to an appropriate range for testing. This must be carefully and accurately performed at all times. After the dilution is completed, one drop of an indicator dye solution supplied in the test kit is added to the diluted sample (Figure 286.2). The instructions supplied with the kit should be followed as described in the product insert.
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Prefoaling Mammary Secretion: Calcium carbonate (ppm)
250 Calcium carbonate (ppm)
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200 150 100 50 0 0
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Figure 286.2 The FoalWatchTM test kit. These chambers from left to right show the pattern of change in the reaction with ever decreasing volume of diluted pre-foaling mammary secretion (PFMS) added to produce the color change (blue) as the day of foaling approaches. Foaling time was within 12 hours of the last chamber (CaCO3 test) on the right. (See color plate 143).
Predictive value of test results When the PFMS CaCO3 first equals or exceeds 200 ppm, there is a 51% probability that spontaneous foaling will occur within the next 24 hours, an 84% probability that foaling will occur within 48 hours, and a 97% probability that foaling will occur within 72 hours.11 The majority of mares foal within a short period of time when a value of 300 to 500 ppm CaCO3 is obtained, but not all mares can be expected to reach these higher values. For mares that have not yet reached 200 ppm, a 99% probability exists that foaling will not occur within the next 24-hour period. This test does not predict the actual foaling time of the mare tested and therefore expectations and claims that it does must be refuted. As with many biological systems, variations are to be expected. Few diagnostic tests have the ability to be completely accurate and reliable with regard to predictive value in all situations. This test is an aid to foaling management programs, and is not intended to guarantee successful warning of impending foaling in all mares. It does, however, increase the likelihood that fewer sleepless nights will be spent waiting up for the mare to foal, especially for inexperienced owners and foaling attendants. It is not unusual for some mares to reach 100–175 ppm CaCO3 and remain at that level for several days before proceeding up to 200 ppm, or above (Figure 286.3). Variations occur between mares, and even within the same mare from year to year. Patience and careful monitoring on a once-to-twicedaily schedule are a must. A dramatic rise or significant change in value over a 12–24-hour interval, especially if the value exceeds 250 ppm, indicates that the
Figure 286.3 The pre-foaling mammary secretion (PFMS) CaCO3 values for one mare as measured by the FoalWatchTM test kit plotted against days prior to spontaneous foaling. Day 15 is the day of foaling. Note the rise in values from day 5 to day 9, a plateau for several days, and then the final rise from days 13 to 14.
mare is approaching readiness for birth. It does not predict that she is going to foal within any specific period of time. Occasionally a value will fall from the previous day’s sampling; this is not a cause for alarm. Repeat the test and be certain that the dilution technique was accurate. This is a normal variation and is not a cause for concern with regard to accuracy of the result. Mares with placentitis and other causes of precocious lactation may have inappropriate early elevation of PFMS calcium.15 Before 310 days gestation, elevated PFMS calcium levels may indicate placental abnormality and may not be indicative of fetal maturity. It has been suggested that an inversion of sodium and potassium concentrations in PFMS does not occur in mares with placentitis, and so may be a better indicator of in utero fetal maturation.16 However, no published reports could be found to support this hypothesis other than the very early work of Peaker et al.2 and Ousey et al.4 These both used spontaneously foaling mares for their investigations. The author has repeated this work to evaluate PFMS sodium and potassium concentration changes and to document the time of their relative inversion prior to foaling in normal mares. A consistent pattern could not be demonstrated (W.B. Ley, unpublished data); 5 of 14 mares foaled normally, delivering live, healthy foals, and none demonstrated a sodium to potassium electrolyte inversion on any sample collected daily or twice daily from 7 to 10 days prior to spontaneous parturition. It is not unusual for maiden mares to fail to develop much of an udder prior to foaling, in which case one’s ability to obtain a sample for testing will be greatly hampered. The ‘first milk’, or colostrum,
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is typically a very thick, honey-colored, sticky secretion. This is an appropriate sample to collect and test as calcium levels will be detectable just as with more ‘normal’ appearing milk. Mares that have been exposed during the last 60–90 days of gestation to fescue grass, specifically the Kentucky-31 variety, infected with the endophytic fungus Neotyphodium coenophialum, may suffer from the toxins (ergot alkaloids) that are produced by the endophyte.17 Such mares very often fail to develop normal udder mass and function prior to foaling (i.e., agalactia). Without a sample for testing, this methodology will be of little help to evaluate approaching readiness for birth.
Other considerations Since calcium contributes to the hardness of water, every precaution should be taken to rinse all reusable items thoroughly with distilled water prior to drying. Use only distilled water supplied in the kit or purchased from local grocery or drug stores for all sample dilutions in the testing procedure. Obtaining the small sample volume that is required for testing on a once-to-twice-daily basis for the 10–14 days prior to foaling does not deprive the foal of any significant amount of colostrum or its immunoglobulin content. Many foals have been monitored during the research and development of this methodology, and in no case was a failure of passive transfer attributable to the sampling of PFMS. In like manner, the quality of the mare’s colostrum and its content of immunoglobulin was not affected. Mares that are prone to ‘running milk’ prior to foaling will do so whether they have been sampled for testing or not. Such mares are still at risk of losing too much colostrum prior to foaling and should be managed accordingly. The procedure of PFMS sampling slightly increases the risk of mastitis development. This is true for any animal when milking is performed by hand, especially if precautions are not taken to wipe the skin and teat surfaces clean, and dry them prior to obtaining the sample. If clumps of cellular debris or pink-to-red discoloration in the PFMS sample obtained from the mare are noted, proper diagnostic work-up and treatment is warranted (i.e., culture and cytology). Repeated sampling of well over 500 pregnant mares during their last 10–30 days of gestation has been considered by this author to be an innocuous procedure. There have been no alterations in normal mare behavior or pre-foaling activities. Premature parturition was not attributed to the testing procedure in the mares included in the evaluation of this method. Likewise, prolonged gestations were not ob-
Table 286.1 Comparison of spontaneously foaling mares versus mares induced to foal using oxytocin. All mares had pre-foaling mammary secretion (PFMS) CaCO3 testing performed
Parameter Number of mares Median foaling time Range in foaling times Mean dose oxytocin Attended deliveries Assisted deliveriesa Premature placental separations Parturient trauma End of Stage 3 by 4 hours postpartum Foal survival at 24 hours Effective passive transfer rate Foal morbidity at 24 hours
Induced foaling
Spontaneous foaling
7 14.00 13.00–16.00 10 IU 7/7 3/7 1/7 0/7 7/7
13 23.00 14.00–07.00 Zero 12/13 1/12 0/12 1/13b 12/13
7/7 7/7 0/7
13/13 13/13 1/13
a These mares were used for instructional purposes, the true clinical need for assistance or intervention may have been overly cautious. b One mare that foaled unattended had a third-degree perineal laceration.
served. It is cautioned that premature or precocious lactation, inappropriate to an individual mare’s gestational length and/or expected foaling date, may be attributable to placentitis and the potential for premature parturition of a septic neonate.15, 16
Conclusions Pre-foaling mammary secretion calcium carbonate testing has been used extensively by this author for over 20 years and has proven useful in the routine management of foaling mares. Its intended use is to assist the determination as to when the fetus is maturing in utero and the mare is approaching readiness to foal based upon changes in the PFMS CaCO3 level. It is a clinical tool that will aid the determination of when to attend the mare’s foaling without an excessive number of sleepless nights. Alternatively, it can be used as a tool for accurately assessing when elective induction of parturition may safely be considered18 (Table 286.1).
References 1. Bain AM, Howey WP. Observations on the time of foaling in Thoroughbred mares in Australia. J Reprod Fertil Suppl 1975;23:545–7. 2. Peaker M, Rossdale PD, Forsyth IA, Falk M. Changes in mammary development and the composition of secretion during late pregnancy in the mare. J Reprod Fertil Suppl 1979;27:555–61. 3. Leadon D, Jeffcott LB, Rossdale PD. Mammary secretions in normal spontaneous and induced premature parturition in the mare. Equine Vet J 1984;16:256–9.
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Pre-Foaling Mammary Secretions 4. Ousey JC, Dudan F, Rossdale PD. Preliminary studies of mammary secretions in the mare to assess foetal readiness for birth. Equine Vet J 1984;16:259–63. 5. Cash RSG, Ousey JC, Rossdale PD. Rapid strip test method to assist management of foaling mares. Equine Vet J 1985; 17:61–2. 6. Brook D. Evaluation of a new test kit for estimating the foaling time in the mare. Equine Pract 1987;9:34–6. 7. Kubiak JR, Evans JW. Predicting time of parturition. Equine Vet Data 1987;7:350–2. 8. Ley WB, Hoffman JL, Meacham TN, Sullivan T, Kiracofe R, Wilson M. Daytime foaling management of the mare 1. Pre-foaling mammary secretions testing. J Equine Vet Sci 1989;9:88–94. 9. Ley WB, Hoffman JL, Crisman MV, Meacham TN, Sullivan T, Kiracofe R. Daytime foaling management of the mare 2: Induction of parturition. J Equine Vet Sci 1989;9:95–9. 10. Ousey JC, Delclaux M, Rossdale PD. Evaluation of three strip tests for measuring electrolytes in mares’ prepartum mammary secretions and for predicting parturition. Equine Vet J 1989;21:196–200. 11. Ley WB, Bowen JM, Purswell BJ, Irby M, Grieve-Crandell K. The sensitivity, specificity and predictive value of measuring the calcium carbonate in mares’ prepartum mammary secretions. Theriogenology 1993;40:189–98.
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12. Martin KL, Hoffman RM, Kronfeld DS, Ley WB, Warnick LD. Calcium decreases and parathyroid hormone increases in serum of periparturient mares. J Anim Sci 1996;74:834–9. 13. Rook JS, Brasleton WE, Nachreiner RF, Lloyd JW, Shea ME, Shelle JE, Hitzler PR. Multi-element assay of mammary secretions and sera from periparturient mares by inductively coupled argon plasma emission spectroscopy. Am J Vet Res 1997;58:376–8. 14. Douglas CGB, Perkins NR, Stafford KJ, Hedderley DI. Prediction of foaling using mammary secretion constituents. N Z Vet J 2002;50:99–103. 15. Rossdale PD, Ousey JC, Cottrill CM, Chavatte P, Allen WR, McGladdery AJ. Effects of placental pathology on maternal plasma progestagen and mammary secretion calcium concentrations and on neonatal adrenocortical function in the horse. J Reprod Fert Suppl 1991;44:579–90. 16. Santschi EM, Slone DE. Evaluation of equine high-risk pregnancy. Comp Cont Educ Pract Vet 1994;16:80–7. 17. Brendemuehl JP. Reproductive aspects of fescue toxicosis. In: Robinson NE (ed.) Current Therapy in Equine Medicine 4. Philadelphia: WB Saunders, 1997; pp. 571–3. 18. Ley WB, Holyoak GR. Normal prefoaling mammary secretions. In: Samper J, Pycock J, McKinnon A (eds) Current Therapy in Equine Reproduction. St Louis: Saunders Elsevier, 2007; pp. 446–51.
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Chapter 287
Mastitis Erica K. Gee and Patrick M. McCue
Inflammation of the mammary gland, or mastitis, is an uncommon disease in the horse. This low incidence is in contrast to dairy cattle, and may be a reflection of the relatively short lactation period of the mare, the size and position of the mammary glands, limited udder milk capacity, and routine mare and foal management.1, 2 It has been suggested that the normal high concentration of lysosyme in mare’s milk may be protective against mastitis3 while normal cow’s milk typically has a low level of lysosyme, which increases in the presence of mastitis.4 The incidence of subclinical mastitis, characterized by grossly normal milk with increased somatic cell counts, in normal lactating mares is not known. As in cows, it is likely to be more common than clinical mastitis.5 Mastitis most commonly occurs within 4 to 8 weeks after weaning a foal, when milk is accumulated in the mammary gland. However, it can also occur during normal lactation or if a foal fails to nurse the mare properly.6, 7 Mastitis has been described in a young filly2 and in older non-pregnant and non-lactating mares. It is reported that mastitis may develop in newborn foals with marked mammary development, sometimes referred to as ‘witch’s milk’.8, 9 Rarely, herd outbreaks of mastitis have been reported.10 The most common bacteria isolated from mares with clinical mastitis are Streptococcus zooepidemicus and Staphylococcus spp. Klebsiella spp., Actinobacillus spp., E. coli and a variety of other organisms are isolated less frequently (Table 287.1).6,11–15 Infection with a single organism is more common than a mixed infection. Culture of multiple organisms was reported in only 12% of mastitis cases in one series.6 Rarely, life-threatening fulminating gangrenous mastitis can occur in the mare.7, 16 A majority of infections are likely due to entry of bacteria through the teat canal, although hematogenous and percutaneous routes of infection are possi-
ble. Sampling of pre-foaling mammary secretions to predict parturition has been suggested to increase the risk of developing mastitis, especially if the skin is not cleaned before obtaining the sample.17 Mastitis in a lactating mare has been associated with submaxilliary abscesses in her sucking foal.18 Mastitis has occasionally been reported in association with fungal infections, including systemic blastomycosis (Blastomyces dermatitidis),19 disseminated coccidioidial infection (Coccidioides immitis),20 and Aspergillus.8 There is one report of verminous mastitis in a mare, caused by Cephalobus, a free-living nematode.21 Consumption of leaves, bark or skin of avocados (Persea americana) can cause mastitis in lactating mares due to necrosis of the secretory epithelium.22 Mastitis may also develop as a consequence of trauma and frostbite.2 Chronic mastitis has been reported in association with mammary gland neoplasms.23, 24 Secretions from ulcerated tumors may predispose to the development of mastitis.7
Diagnosis Clinical signs of mastitis depend on severity, and range from mild swelling and firmness to hot, painful mammary gland swelling, sometimes associated with hindlimb lameness, ventral edema, and signs of systemic disease (Figure 287.1). Mammary enlargement can be unilateral or bilateral, and nursing mares are usually reluctant to let their foal suck. The milk vein may be very prominent on the affected side. The udder of the mare is divided into two glands (left and right) and each gland is further divided into cranial and caudal lobes. The two teats of a mare each have two teat canals which drain cranial and caudal lobes. After careful cleaning of the teats, samples of secretions should be obtained from each of the
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Table 287.1 Organisms cultured aerobically from the mammary secretions of 30 mares with clinical mastitis Organism
Percent of all isolates∗
Streptococcus zooepidemicus Other Streptococcus spp. Staphylococcus spp. Klebsiella Actinobacillus spp. Escherichia coli Bacillus spp. Pasturella Enterobacter Corynebacterium spp. Pseudomonas
32.3 14.7 14.7 11.8 5.9 5.9 2.9 2.9 2.9 2.9 2.9
Figure 287.2 Collection of a milk sample from a mare with mastitis.
∗ More than one organism was cultured in four samples. Source: McCue PM, Wilson WD. Equine mastitis: a review of 28 cases. Equine Vet J 1989;21:351–3. Perkins NR, Threlfall WR. Mastitis in the mare. Equine Vet Educ 2000;5:99–102.
teat openings to identify which lobe(s) are affected (Figure 287.2). Secretions from a mastitic lobe may range from clear to thick (Figure 287.3), and are not uncommonly hemorrhagic.1 Occasionally no secretions can be expressed although the gland is hot and very hard.7 Secretions should be submitted for aerobic culture and determination of antibiotic sensitivity, and a second sample applied to a glass slide, allowed to air dry, stained (e.g., Diff-Quik stain) and examined for cytological characteristics. Cytology samples may reveal normal or degenerated neutrophils (Figure 287.4), necrotic debris, unidentifiable degenerated cells, and bacteria in up to 33% of mastitic samples.5, 6 In contrast, there are almost no white blood cells noted in normal milk.5 Only 30% of secretions from mares with clinical mastitis were associated with a positive culture in one study,6 and potential pathogens can occasionally be cultured from normal milk.5
Figure 287.1 Enlarged mammary gland of a mare with mastitis.
Figure 287.3 Thick exudates collected from the mammary gland of a mare with mastitis.
Figure 287.4 Cytology sample of mammary fluid from a 22year-old mare with Streptococcus zooepidemicus mastitis. A foal had been weaned from the mare approximately 3 months previously.
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Figure 287.5 Stripping fluid from the udder of a mare with mastitis.
Figure 287.6 Infusion of a commercial bovine mastitis preparation into the mammary gland of a mare.
The California mastitis test (CMT) is marketed for use in cattle to detect mastitis, and has been reported to be suitable for use in horses.7 The CMT uses an anionic detergent to form gels with white blood cells present in inflammatory exudate of mastitic milk. A complete blood count (CBC) from a clinically affected mare may show a leukocytosis with neutrophilia and hyperfibrinogenemia or may be unremarkable.6 Differential diagnoses for mastitis include udder abscesses, udder edema, neoplasia and trauma.1 Ultrasonography may assist in differentiation of udder abscessation and mammary tumors from mastitis in the mare.7 Histology of biopsy samples is required for a definitive diagnosis of neoplasia, a condition which is rare in the mare.
ever, treatment is often initiated before culture and sensitivity results are complete, and broad-spectrum antibiotics such as trimethoprim-sulphamethoxazole or a combination of penicillin and gentamicin are suitable choices. Treatment should be continued for 5 to 7 days, or until at least 24 hours after clinical resolution of the mastitis.7, 12 Flunixin meglumine, phenylbutazone, or other non-steroidal anti-inflammatory drugs (NSAID) may be useful in reduction of pain and swelling associated with mastitis. Usually there is a rapid response to treatment, with mammary secretions appearing normal within 1 to 3 days, and the mammary gland palpating normally within a week.12 Some authors recommend instilling a dry cow intramammary preparation into the previously affected gland when the infection has resolved in the non-lactating mare.12 Abscessation of the mammary gland, secondary to bacterial mastitis, is a rare occurrence and is managed by establishment of drainage and long-term antibiotic therapy. Abscesses are usually solitary, but may also be miliary or multilocular.7 Management of mycotic mastitis should include administration of an antifungal drug. Itraconazole may be a suitable choice, but the efficacy of this drug in treating mycotic mastitis has not been established. Mares with avocado toxicity-associated mastitis will recover without medical treatment within 2 weeks after elimination of exposure to avocado.22 However, it may be prudent to also initiate medical treatment as described for bacterial mastitis to limit secondary complications. Surgical resection of mammary glands has been described for mares with severe and recurrent cases of mastitis which were refractory to medical treatment.25
Treatment Local therapy for bacterial mastitis involves stripping the affected gland to remove mastitic secretions containing bacteria and necrotic debris (Figure 287.5), in combination with hot packing and cold hosing to promote drainage and reduce edema. Sedation may be required in order to safely and effectively carry out local therapy. A lactating cow intramammary preparation can be instilled into affected glands after appropriate cleaning and disinfection (Figure 287.6). Care should be taken not to damage the external orifice or the streak canal with the tip of the intramammary preparation.6 In most clinical cases a mare is treated once with an intramammary antibiotic infusion followed by 5 to 7 days of systemic antibiotic therapy. If the mare is reluctant to have her mammary gland examined or stripped, it may be too dangerous to attempt intramammary therapy, and systemic therapy alone may be prudent. The choice of systemic antibiotics should be based on culture and sensitivity results, if possible. How-
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Prognosis Cases of acute mastitis usually resolve quickly with appropriate treatment, and recurrence of mastitis is uncommon. The mammary gland almost always returns to full, normal function after resolution of the condition. However, chronic mastitis may lead to fibrosis in the affected gland and subsequently reduced milk production in future lactations.26
References 1. McGladderly AJ. Differential diagnosis and treatment of diseases of the equine mammary gland. Equine Vet Educ 2002;5:94–6. 2. Pugh DG, Magnusson RA, Modransky PD, Darnton KR. A case of mastitis in a young filly. J Equine Vet Sci 1985;5: 132–4. 3. Zou S, Brady HA, Hurley WL. Protective factors in mammary gland secretions during the periparturient period in the mare. J Equine Vet Sci 1998;18:184–8. ¨ 4. Carlsson A, Bjorck L, Persson K. Lactoferrin and lysozyme in milk during acute mastitis and their inhibitory effect in Delvotest P. J Dairy Sci 1989;72:3166–75. 5. Freeman KP, Roszel JF, Slusher SH. Cytologic features of equine mammary fluids: normal and abnormal. Compend Equine 1988;10:1090–9. 6. McCue PM, Wilson WD. Equine mastitis: a review of 28 cases. Equine Vet J 1989;21:351–3. 7. Knottenbelt DC. The mammary gland. In: Knottenbelt DC, Pascoe RR, Lopate C, LeBlanc MM (eds) Equine Stud Farm Medicine and Surgery. Edinburgh: Saunders, 2003, pp. 343–52. 8. Delorme JY, Hamelin A. Trois cas de mammites chez les pouliches nouveau-nees. Practique Veterinaire Equine 1994; 26:276–8. 9. Knottenbelt D, Holdstock N, Madigan J. Congenital abnormalities and inherited disorders. In: Equine Neonatology Medicine and Surgery. Edinburgh: Saunders, 2004, pp. 95–153. 10. Welsh RD. The significance of Streptococcus zooepidemicus in the horse. Equine Prac 1984;6:6–16.
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11. Addo PB, Wilcox GE, Taussig R. Mastitis in a mare caused by C. ovis. Vet Rec 1974;95:193. 12. Perkins NR, Threlfall WR. Mastitis in the mare. Equine Vet Educ 2002;5:99–102. 13. Prentice MWM. Mastitis in the mare. Vet Rec 1974;94: 380. 14. Reese GL, Lock TF. Streptococcal mastitis in a mare. J Am Vet Med Assoc 1978;173:83–4. 15. Roberts MC. Pseudomonas aeruginosa mastitis in a dry nonpregnant pony mare. Equine Vet J 1986;18:146–7. 16. Mercier C, Bussy C, Saint-Amand AA. Un cas de mammite gangreneuse chez une jument trotteur. Pratique Veterinaire Equine 2005;37:63–5. 17. Ley WB, Holyoak GR. Normal prefoaling mammary secretions. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders, 2007; pp. 446–51. 18. Al-Graibawi MAA, Sharma VK, Ali SI. Mastitis in a mare. Vet Rec 1984;115:383. 19. Wilson JH, Olsen EJ, Haugen EW, Hunt LM, Johnson JL, Hayden DW. Systemic blastomycosis in a horse. J Vet Diagn Invest 2006;18:615–19. 20. Walker RL, Johnson BJ, Jones KL, Pappagianis D, Carlson GP. Coccidioides immitis mastitis in a mare. J Vet Diagn Invest 1993;5:446–8. 21. Greiner EC, Calderwood Mays MB, Smart GC Jr, Weisbrode SE. Verminous mastitis in a mare caused by a free-living nematode. J Parasitol 1991;77:320–2. 22. McKenzie RA, Brown OP. Avocado (Persea americana) poisoning of horses. Austr Vet J 1991;68:77–8. 23. Hirayama K, Honda Y, Okamoto M, Tsunoda N, Tagami M, Taniyama H. Invasive ductal carcinoma of the mammary gland in a mare. Vet Pathol 2003;40:89–91. 24. Reppas GP, McClintock SA, Canfield PJ, Watson GF. Papillary ductal adenocarcinoma in the mammary glands of two horses. Vet Rec 1996;138:518–19. 25. Schiemann V, Bartmann CP. Mastectomia in the horse – indication and surgical procedure. Pferdeheilkunde 2004;20: 506–10. 26. Shideler RK. External examination. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Williams & Wilkins, 1993; pp. 199–203.
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Chapter 288
Mammary Neoplasia Jack R. Snyder and Omar Maher
Mammary tumors are relatively uncommon in horses although incidences of 0.111 to 1.99%2 have been reported in European slaughterhouse studies. Only a few cases have been reported in the literature in the last 30 years, and most reports refer to single cases with three other reports referring to 2, 3, and 4 cases.3–5 Mammary tumors are most frequently primary neoplasia of the gland (adenocarcinomas and rarely adenomas6 ), but can also be due to the local spread of neoplasm of the skin coverage (squamous cell carcinomas, melanomas, sarcoids) as well as secondary/metastatic lesions such as melanomas, mastocytomas and lymphosarcomas.2, 5,7–10
sia.4, 14 Biopsy can be performed under sedation and local analgesia, and is diagnostic if a representative sample of the mass is sampled. Ultrasonographic examination of the mammary gland has been well described15, 16 and can be used on questionable cases to try to identify any possible mass within the mammary gland (Figure 288.2). Mammary carcinomas are locally aggressive but also metastasize to other structures. Most common sites of metastasis include superficial and deep inguinal lymph nodes, and then other mammary gland,4, 5, 9, 11, 17 but can also occasionally affect the liver, lungs,2, 5 uterus, kidneys, ovaries, pleura, peritoneum, pericardium, and mediastinum.2, 4, 12, 18
Adenocarcinomas
Treatment
The vast majority of mammary tumors reported in the literature are adenocarcinomas. Review of reported cases showed affected mares as young as 12 years old5 and up to 21 years of age.11 Clinical presentation typically consists of enlargement of one of the mammary glands (Figure 288.1), pain associated with palpation, discharge, often ulcerations, and occasionally weight loss. Diagnosis is often delayed as early symptoms are similar to those of mastitis. Affected mares are often treated as such initially and fail to respond to prolonged antimicrobial therapy. Ulcerations are often seen in mammary tumors4, 5, 11, 12 but rarely in mastitis13 and their presence is highly suggestive of carcinoma. Milk culture will often yield bacterial growth in cases of mastitis, and no significant growth in cases of neoplasia, although some mares with carcinoma will also develop secondary mastitis complicating the diagnosis. Cytology of the discharge or fine needle aspirate is an easy and important diagnostic step that can help distinguish between mastitis and neopla-
Treatment of mammary carcinomas involves surgical resection of the affected mammary gland, although removal of the contralateral mammary gland as well as the inguinal lymph nodes is recommended because of the high incidence of metastasis to these structures. Resection should be performed as early as possible in the course of the disease because of the highly invasive and metastatic characteristics of equine mammary carinomas. A metastasis screening (rectal examination, abdominal fluid analysis, abdominal ultrasonography, and thoracic radiography) should be undertaken before planning surgical resection. Additional therapy such as systemic chemotherapy and radiation therapy could be considered if available.
Surgical technique The mare should receive broad-spectrum preand perioperative antibiotics as well as antiinflammatories. Surgery is performed under general
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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time, the caudal superficial epigastric vein can be ligated as it enters the cranial aspect of the gland. Inguinal lymph nodes are then resected. The connective tissue, subcutaneous tissue, and skin over the removed mammary gland can be closed in layers if possible, minimizing dead space. A portion of the wound is left open to drain and a Penrose drain is left in place. After recovery, the mare should be stall rested with controlled hand-walking to encourage drainage. The wound is monitored for swelling and fluid accumulation. The drain is typically removed after 1 week. Figure 288.1 Mare positioned in dorsal recumbency with an enlarged mammary gland, due to an adenocarcinoma. (Courtesy of Dr M.H. MacDonald.)
anesthesia on the mare placed in dorsal recumbency. After surgical preparation and draping, an elliptical incision is made in a craniocaudal direction, around the teat(s). Ulcerated skin should be removed with as much margin as possible (ideally minimum of 2 cm). Skin should be dissected off the mammary gland and then the gland should be bluntly dissected caudally, cranially and on its side. If only one side is to be removed, the gland should be bluntly dissected off the median septum. As the dissection progresses, small blood vessels are ligated or electrocauterized. As the mammary gland is lifted up, the deep larger vessels become visible. On each side, the external pudendal artery should be ligated before it branches (cranial and caudal mammary artery). At the caudal aspect, the obturator vein and branches of the internal pudendal vein should be ligated, and the mammary gland should be lifted off the body wall. At that
Surgical complications Surgical complications mainly include seroma formation, wound infection, and laminitis in the short term, although septic peritonitis has been reported.12 In the long term, recurrence of the tumor and metastasis are common.
Prognosis Prognosis for the mare with adenocarcinoma is poor. In cases described in the literature, surgical treatment did not seem to increase survival time when compared to untreated mares,4 with most mares not surviving beyond 18 months after diagnosis. Arguably in most of these cases surgical treatment might have been delayed and some cases already had systemic manifestations of carcinomatosis such as poor body condition. Extrapolating from other species, early diagnosis and aggressive surgical resection would increase chances of survival.
Squamous cell carcinomas
Figure 288.2 Ultrasonographic appearance of the mammary gland in Figure 237.1. (Courtesy of Dr M.B. Withcomb.)
Squamous cell carcinomas originate at the superficial aspect of the udder (skin) and often invade the mammary gland itself. Review of cases presented at the University of California, Davis, identified eight mares with mammary squamous cell carcinoma in a 10-year period.19 Seven mares were more than 20 years old and one mare was 8 years old. Interestingly, seven of the eight mares had developed vulvar squamous cell carcinomas months to years prior to signs of mammary involvement. Clinical signs were similar to those of mammary adenocarcinoma. Squamous cell carcinomas, when infiltrative, are as aggressive and prone to metastasis as adenocarcinomas. Unless they are very superficial, in which case a conservative resection can be performed, a radical mastectomy and lymphadenectomy should be performed.
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When a conservative resection is performed, additional treatment such as local chemotherapy (e.g., cisplatin injections)20 or radiation therapy should be considered. Systemic chemotherapy could be considered when radical mastectomy is performed. Prognosis for survival is good when treated early and poor if the tumor has extended to the mammary gland itself.
Sarcoids Sarcoids over the udder are fairly common and do not usually invade the gland itself. They can be treated by surgical resection and/ or local chemotherapy (series of injection with cisplatin,20 or implantation of cisplatin beads21 ).
Melanomas Melanomas are encountered particularly in gray mares and can be variably invasive. They can originate from the skin of the udder or be metastatic. Treatment can vary from benign neglect for local small stable melanomas to focal resection (especially if they are interfering with the teat) or radical mastectomy and lymphadenectomy for those invading the mammary gland.10
References 1. Martel. Cancer in horses. Am Vet Rev 1914;64:299–300. 2. Schmahl W. Solides Karzinom der Mamma bei einem Pferd. Berlin Munch Tierarz Wochens 1972;85:141–2. 3. Reppas GP, McClintock SA, Canfield PJ, Watson GF. Papillary ductal adenocarcinoma in the mammary glands of two horses. Vet Rec 1996;138:518–19. 4. Prendergast M, Bassett H, Larkin HA. Mammary carcinoma in three mares. Vet Rec 1999;144:731–2. 5. Seahorn TL, Hall G, Brumbaugh GW, Honnas CM, Lovering ST, Snyder JR. Mammary adenocarcinoma in four mares. J Am Vet Med Assoc 1992;200:1675–7.
6. Spadari A, Valentini S, Sarli G, Spinella G, Millanta F. Mammary adenoma in a mare: Clinical, histological and immunohistochemical findings. Equine Vet Educ 2008;20:4–7. 7. Judd KVF, Kennedy PC. Pathology of Domestic Animals, 2nd edn. New York: Academic Press, 1970. 8. Surmont J. L’epitheliome mammaire de la jument et ses metastases pulmonaires. Bult Assoc Frac Etudes Can 1926;15: 98–101. 9. Karcher LF, Le Net JL, Turner BF, Reimers TJ, Tennant BC. Pseudohyperparathyroidism in a mare associated with squamous cell carcinoma of the vulva. Cornell Vet 1990; 80:153–62. 10. Schiemann V, Bartmann CP. Euteramputation beim Pferd – Indikation und chirurgische Durchfuhrung. [Mastectoma in the horse – indication and surgical procedure.] Pferdeheilkunde 2004;20:506–10. 11. Hirayama K, Honda Y, Sako T, Okamoto M, Tsunoda N, Tagami M, Taniyama H. Invasive ductal carcinoma of the mammary gland in a mare. Vet Pathol 2003;40:86–91. 12. Foreman JH, Weidner JP, Parry BW, Hargis A. Pleural effusion secondary to thoracic metastatic mammary adenocarcinoma in a mare. J Am Vet Med Assoc 1990;197:1193–5. 13. McCue PM, Wilson WD. Equine mastitis – a review of 28 cases. Equine Vet J 1989;21:351–3. 14. Freeman KP. Cytological evaluation of the equine mammary gland. Equine Vet Educ 1993;5:212–13. 15. Waldridge BM, Ward TA. Ultrasound examination of the equine mammary gland. Equine Pract 1999;21:10. 16. Gungor O, Pancarc SM, Karabacak A. Examination of equine udder and teat by B-mode ultrasonography. Kafkas Universitesi Veteriner Fakultesi Dergisi 2005;11:107–11. 17. Munson L. Carcinoma of the mammary gland in a mare. J Am Vet Med Assoc 1987;191:71–2. 18. Acland HM, Gillette DM. Mammary carcinoma in a mare. Vet Pathol 1982;19:93–5. 19. Maher O. Mammary squamous cell carcinoma in 8 mares. University of California Davis, Veterinary Medical Teaching Hospital, 2008. 20. Th´eon AP, Wilson WD, Magdesian KG, Pusterla N, Snyder JR, Galuppo LD. Long-term outcome associated with intratumoral chemotherapy with cisplatin for cutaneous tumors in equidae: 573 cases (1995–2004). J Am Vet Med Assoc 2007;230:1506–13. 21. Hewes CA, Sullins KE. Use of cisplatin-containing biodegradable beads for treatment of cutaneous neoplasia in equidae: 59 cases (2000–2004). J Am Vet Med Assoc 2006; 229:1617–22.
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Section (o): Specific Diagnostic and Management Techniques
289.
Preventative Medicine for Broodmare Farms John B. Chopin
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Chapter 289
Preventive Medicine for Broodmare Farms John B. Chopin
The aim of preventive medicine is to identify potential risks and eliminate or reduce them and avoid the disease from occurring. Good prevention is good management. Looking after the environment of the broodmare, attending to nutrition, regular hoof trimming, deworming, and vaccination provide the cornerstone. Consideration of the reproductive status of the mare (maiden, dry/barren, or lactating) can finetune the particular needs of the individual.
Nutrition Feeding mismanagement probably causes more problems than improper dietary formulations. Goodquality pasture is still an excellent basis for a feeding program. Pasture provides a much higher level of antioxidants and carotene than hay.1 Pasture can reduce the incidence of colic, ulcers, signs of respiratory disease, and abnormal behaviors. However, excessive intake of lush pasture can cause colic and laminitis. Moreover, certain minerals can be lacking depending on soil quality, and toxic plants and endophytes can be present.1 Food restriction leading to a decrease in body condition resulted in an increase in the number of mares that entered winter anestrus. The anestrus was so deep that photostimulation failed to advance the first ovulation of the breeding season.2 Dry and maiden mares in lighter condition took longer to enter cyclicity and had lower fertility with more estrous cycles per pregnancy.3 Good nutrition was observed not only to increase the likelihood of a non-pregnant mare entering transition and cyclicity earlier,2,4–8 but also to maintain early pregnancy.8, 9
Feeding high-quality protein to mares in transition advances the first ovulation of the season by 2–3 weeks,10, 11 and this effect is seen more in older mares than young mares.12 Mares consuming lush green grass ovulate earlier in the season than mares eating dry hay.13 No detrimental effects of excess body fat in late gestation were found.3 Initial excess body fat improved fertility – either on entering the breeding season or after foaling.3 Post-foaling, mares in lighter condition took longer to get in foal and were more likely to lose the pregnancy.3 The bodyweight loss of lactating mares fed recommended dietary levels for protein was also accompanied by an increased rate of early embryonic loss.14 In another experiment, 5 of 14 (35.7%) mares fed a lowquality protein diet suffered from early embryonic loss before 90 days of pregnancy, compared to 3 of 41 (7.3%) mares in the remaining groups that received the higher-quality protein in their diets.15 Mares fed low-energy diets have significantly higher fetal loss rates.9 In mares a low-energy diet resulted in a higher fetal loss rate (3 of 10 versus none of 9) when compared to mares fed a maintenance diet (p < 0.07). The mares on the restricted diet had higher plasma progesterone and lower plasma cortisol concentrations.9 Mares on a low-protein diet also showed higher levels of plasma progesterone.16 The effect of maternal nutrition on placental and fetal development in maiden Thoroughbred mares was examined. Three- and four-year-old fillies were kept in moderate condition until pregnant and then were either maintained or overfed. Gestation was longer in the maintenance group (342 ± 3 days vs 333 ± 3 days). Placental area, weight, and volume did
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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not differ between the groups, neither did foal birthweight or body composition. The study was complicated by an outbreak of strangles in both study groups, causing some weight loss.17, 18 Acute nutrient restriction during mid-pregnancy in ewes significantly reduces fetal lung weight. A reduction in amniotic fluid volume reduces the space for fetal movement and positioning and induces exaggerated trunk flexion. The trunk flexion decreases thoracic volume, which limits lung expansion.19 A reduction in amniotic fluid volume might also be a cause for congenital flexural deformities. In the mare the presence of adequate to excess body condition during cyclicity, pregnancy, and lactation does not appear to have any detrimental effects. In comparison, poor body condition has measurable detrimental effects on reproduction and the fetus. Inadequate nutrition during equine pregnancy can limit the development of fetal organs and alter metabolism permanently. In the equine, the single most limiting factor affecting athletic potential might be the 11 months spent in utero.
Feet care Mares with pre-existing feet conditions such as laminitis might experience an increase in severity of the condition with the increased weight during the latter stages of pregnancy. Appropriate medical and/or surgical therapy to manage the laminitis, along with carefully maintained and controlled weight gain is necessary so the mare does not become obese. Managed exercise will depend on the pain present. Regular trimming of the mare’s hooves during pregnancy is important, with attention to correcting any problems and avoidance of overly aggressive trimming to reduce the risk of sensitive sore soles for the few days after trimming. Some managers will not trim the mare’s feet during the last month before foaling, to avoid any potential foot pain.
Dentition Dental care is an area that is probably overlooked in many broodmares; some will, if they are lucky, get a quick rasp once a year. There is evidence that periodontal disease can induce abortion or pre-term labor in women.20–23 Although other studies have not found any correlation between periodontal disease and delivery of pre-term low-birthweight infants,24, 25 a review of multiple studies since the original 1996 study found a positive correlation between periodontal disease and pre-term birth.26 Actinomyces spp. are an important cause of chronic equine placentitis in the USA.27, 28 Actinomyces sp.
has been isolated in the placenta of a pre-term baby. The organism is a known resident of the periodontal sulcus, and the mother in this case had a recurring dental abscess during pregnancy.29 This is the first recorded case of oral bacteria isolated from reproductive organs. There could be a link between periodontal disease in mares and compromised pregnancies, especially with Actinomyces spp., which are known residents of the oral cavity and an important cause of chronic equine placentitis.
Exercise Regular opportunity to exercise improves the overall physical and mental health of the mare.30 Improving the strength and tone of abdominal musculature will help during parturition. Mares that are weak or ill and not able to provide abdominal exertion are at risk of prolonged parturition and dystocia. Forced exercise should not be used in mares with ventral edema as some of these animals might have impending prepubic tendon or muscle wall rupture.30 Mares with poor internal conformation of the uterus and a pendulous abdominal conformation tend to more easily accumulate uterine fluid. The improvement of muscular tone might help to correct the conformation of the abdomen and to elevate the uterine position. Regular exercise, such as lunging, can help these mares to expel uterine fluid. Mares that are restricted to a stall are more likely to accumulate uterine fluid and suffer reduced fertility.
Anthelmintics A holistic approach to internal parasite control, with pasture management, anthelmintic rotation, fecal egg counts, and postmortem surveys, helps to control parasite burdens in horses. Pasture management includes reducing stocking rate, rotation of paddocks, cross-grazing, removal of feces twice weekly or harrowing of feces during summer.30 A detailed discussion on parasite control is included in Chapter 30.
Prevention of infectious disease Vaccination for disease prevention will depend on the associated risk and current recommendations for licensed products. A detailed discussion on vaccination is included in Chapter 31. The use of isolation and quarantine of new arrivals before entry into the general population is critical. New arrivals should be placed into isolation on their arrival onto a farm. This will vary with the facilities and space available on the farm. The arrival yards
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Preventive Medicine for Broodmare Farms should be close to the entrance so that the animals do not progress too far onto the farm. At the very least arriving animals should undergo inspection for obvious signs of infectious disease, and monitoring for 12–24 hours for more subtle signs such as increased temperature. This is especially important for mares traveling long distances to breeding farms as early recognition of pleuropneumonia (shipping fever) will often modify the course of the disease. The vaccination and anthelmintic history of these individuals should be checked and updated prior to entry onto the farm. The use of natural or artificial barriers on the farm can help to prevent or slow the spread of infectious disease. Biological barriers of non-stressed, immune (vaccinated) individuals can also help to slow or stop the spread of infectious disease. Preventing at-risk populations from being exposed to infectious agents is critical. One example of this is situating pregnant mares away from the young weaned animals and animals under the stress of an athletic career where exposure to respiratory viruses (equine herpesvirus 1 and 4) is common and may lead to abortion of pregnant in-contact mares.
Pasture management Regular topping of the grass will stop the grass from maturing and promote the growth stage. Regular topping also helps in weed control to reduce toxic plants and improve utilization of pasture. Some supplementation might be required in the form of roughage when the grass is young and in the rapid growth phase. The young pasture will have low dry matter content and some mares (lactating) will lose weight on lush green pasture. Supplementation to improve protein intake might be needed once the pasture has matured or dried off. Regular removal of feces, ideally twice weekly, will contribute to parasite control. Harrowing of feces is beneficial during hot summers when dessication will occur, but in milder conditions this might encourage the spread of parasitic larvae.30 Identification of tall fescue grass (Festuca arundinacea), which is susceptible to fungal endophyte infestation with Neotyphodium coenophialum, is critical to avoid prolonged gestation, dystocia, stillborn foals, retained placentas, agalactia, and mare mortality.30, 31 Large areas of the United States are planted with endophyte-infected fescue, and current recommendations are to remove mares from pasture and supplement with hay at the end of gestation (at least 30 days) or to administer domperidone (1.1 mg/kg p.o. daily) to reverse the effects of the fungal toxin.31–33
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Maiden mares Maiden mares that have recently retired from an athletic career can have special requirements above those of the average broodmare. They often have to adjust to a new social hierarchy, a new routine, a change of environment, a different diet, probably no shoes on tender hooves, and then also are expected to relax into the pressure of breeding when any display of sexual behavior might have been previously inhibited. The capability of the management to deal with these challenges to the retired maiden race mare may dictate the ease and speed with which they cycle and become pregnant. If there is a lack of attention to these details, these retired maiden race mares will often show a reduction in body condition and very little ovarian activity. A pre-season or pre-breeding examination of the maiden mare might be required in order to evaluate any existing or potential problems. The mare might be classified as a maiden mare but could have already been bred a number of times or even seasons with no foal being produced. It is important to establish any history of drug use [altrenogest, anabolics, anti-gonadotropin-releasing hormone (antiGnRH) vaccine] prior to starting the breeding career of the maiden broodmare. A more detailed examination protocol is discussed in Section (f) Techniques in Reproductive Examination but might include an external examination (vulval conformation), rectal palpation, and ultrasonographic evaluation, swabs from uterus and/or clitoris for culture/sensitivity (and cytology), endometrial biopsy, or cytogenetic evaluation. The teasing and natural mating of these mares can be problematic if they are shy about showing overt estrus behavior. Patience, time, and a controllable stallion (in a natural breeding situation) work with the majority of these mares, but physical and chemical restraint might also be required. The need for overt estrus behavior is not necessary where artificial insemination is an option.
Dry (barren) mares Dry or barren mares are typically more likely to have breeding problems due to the fact that they are not pregnant from previous attempts at breeding. A breeding history can help to determine why they are not pregnant. A pre-season or pre-breeding examination is covered in Section (f) Techniques in Reproductive Examination but could include an external examination (vulval conformation), an internal examination (conformation, palpation, ultrasound), digital palpation
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of the cervix (diestrus), endometrial biopsy, and swabs from uterus and clitoris (culture/sensitivity and cytology).
Pregnant mares All mares stated to be pregnant should be tested on entry to a new breeding farm to accurately identify pregnancy status prior to incorporation into the general population. Close monitoring of mares during pregnancy might not detect pregnancy loss, so the testing of pregnancy status prior to organizing mares in preparation for foaling is warranted to identify mares that have suffered pregnancy loss. Twinning has traditionally been an important cause of pregnancy loss, but with the introduction of frequent ultrasonographic examination early in gestation it is possible to detect twins and treat the mares accordingly.34 However, without any history of early pregnancy diagnosis, ultrasound examination even in mid- to late gestation may not rule out the possibility of twins. The history of previous pregnancies can highlight areas of concern to be monitored. Previous placentitis might require ultrasonographic monitoring, either rectal or transabdominal, to evaluate the health of the placenta in the current pregnancy. A previous occurrence of dystocia might be due to a predisposing cause, or itself have led to damage that might interfere with successful foaling in future. Hemorrhage from a previous middle uterine artery rupture might occur again in subsequent pregnancies. Any musculoskeletal problems such as a fractured pelvis, arthritis, neuromuscular disease, or laminitis could interfere with the successful carrying or delivery of a pregnancy. The correction or amelioration of these in the early stages of pregnancy is important before the broodmare experiences increased bodyweight and greater stress on the affected organs. Pregnant mares should once or ideally twice daily be monitored for signs of disease – precocious udder development, precocious milk production, vaginal discharge, ventral edema potentially indicating an impending abdominal wall rupture.35 Management in the final trimester includes a regular period of exercise for the mare to mitigate the effects of stall confinement. The use of lights in late pregnancy can decrease the length of gestation by 7–10 days and decreases the likelihood of postpartum anestrus.35 Mares should be evaluated for the presence of a Caslick operation, and it should be removed at a standard time such as 2 weeks before foaling or when the mare is showing signs of imminent foaling.35 However, some mares can develop an ascending placentitis in the week after a Caslick has been removed.
Neonatal isoerythrolysis is an infrequent but important disease due to the mare transferring anti-red blood cell antigen antibodies through the colostrum to the foal, causing the foal to suffer red blood cell lysis. When parentage validation was performed using blood groups then identification of potential mismatches could be easily noted and prepared for. With the switch to DNA to validate parentage, the identification of equine blood groups is done by fewer laboratories. Mare serum in the latter stages of pregnancy can be evaluated for the presence of antibodies, but the final two weeks is the best period due to the occasional false negative before this period. 35 Potential problem matings can be identified with blood groups, and parturition is monitored in highrisk mares; the foal is muzzled and fed compatible colostrum until the mare’s serum has been tested for agglutination with the foal’s washed red blood cells.
Vaccination A detailed discussion of vaccination is included in Chapter 31 but a brief overview of vaccination is given here for the benefit of managing the broodmare. The use of various vaccines will have to evaluated on a risk-benefit analysis to determine which vaccines should be used. Vaccination against tetanus with the toxoid is considered essential. Unvaccinated animals need two doses 4–6 weeks apart with an annual booster. Pregnant mares should receive their annual booster or the last of a primary course 4–6 weeks before foaling to maximize colostral antibody titer. Eastern and Western equine encephalitides are zoonotic, and vaccinations against these diseases are considered to be essential for horses in North America. The primary course is two doses 4–6 weeks apart with annual boosters either before the vector (mosquito) season or 4–6 weeks before foaling. West Nile virus can cause encephalitis in horses and humans. There are three different vaccines available – inactivated whole virus vaccine, recombinant canarypox vector vaccine, and modified live chimera vaccine, and protocols vary with vaccine. Boosters are timed to promote immunity prior to the vector season or 4–6 weeks prior to foaling. A single dose of the rabies vaccine is enough to promote immunity. The single dose or annual booster can be administered 4–6 weeks prior to foaling. Other diseases that can be vaccinated against if the local area risk is deemed to warrant it are:
r r r r
anthrax (but not in pregnant mares); strangles with either the killed or live vaccine; rotavirus during pregnancy to help the foal with colostral immunity; Potomac horse fever, although the benefits against abortion are unknown;
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r r r r
equine influenza, with a range of possible products; equine viral arteritis, although it is not recommended in pregnant mares and the serological status of the mare is required before primary vaccination; equine herpesvirus with the use of several doses of the inactivated vaccine during pregnancy to prevent abortion; botulism vaccination during pregnancy to boost colostral antibody levels delivered to the foal.
References 1. Hintz H. Equine nutrition update. Proceedings of the Annual Convention of the American Association of Equine Practitioners. 2000;46:62–79. 2. Guillaume D, Duchamp G, Salazar-Ortiz J and Nagy P Nutrition influences the winter ovarian inactivity in mares. Theriogenology 2002;58:593–7. 3. Henneke D, Potter G and Kreider J. Body condition during pregnancy and lactation and reproductive efficiency of mares. Theriogenology 1984;21:897–909. 4. Bengtsson G and Knudson O. Feed and ovarian activity of trotting mares in training. Cornell Vet 1963;53:404–11. 5. van Niekerk C. Pattern of the oestrous cycle of mares I. The breeding season. J S Afr Vet Med Assoc 1967;38: 295–8. 6. van Niekerk C and van Heerden J. Nutrition and ovarian activity of mares early in the breeding season. J S Afr Vet Assoc 1972;43:355–60. 7. Voss J and Pickett B. Effect of a nutritional supplement on pregnancy rate in nonlactating mares. J Am Vet Med Assoc 1974;165:702–3. 8. Belonje P and van Niekerk C. A review of the influence of nutrition upon the oestrous cycle and early pregnancy in the mare. J Reprod Fertil Suppl 1975;23:167–9. 9. Potter J, Kreider J, Potter G, Forrest D, Jenkins W and Evans J. Embryo survival during early gestation in energydeprived mares. J Reprod Fertil Suppl 1987;35:715–16. 10. Carnevale E, Hermenet M and Ginther O. Age and pasture effects on vernal transition in mares. Theriogenology 1997;47:1009–18. 11. van Niekerk F and van Niekerk C. The effect of dietary protein on reproduction in the mare. II. Growth of foals, body mass of mares and serum protein concentration of mares during the anovulatory, transitional and pregnant periods. J S Afr Vet Assoc 1997;68:81–5. 12. Carnevale E, Thompson K, King S and Roszkowski J. Effects of age and diet on the spring transition in mares. P Annu Conv Am Equine 1996;42:146–7. 13. Irvine C and Alexander S. Managing the mare for optimal fertility. J Equine Sci 1998;9:83–7. 14. van Niekerk F and van Niekerk C. The effect of dietary protein on reproduction in the mare. III. Ovarian and uterine changes during the anovulatory, transitional and ovulatory periods in the non-pregnant mare. J S Afr Vet Assoc 1997;68:86–92. 15. van Niekerk F, van Niekerk C. The effect of dietary protein on reproduction in the mare. VI. Serum progestagen concentrations during pregnancy. J S Afr Vet Assoc 1998;69:143–9.
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16. van Niekerk F, van Niekerk C. The effect of dietary protein on reproduction in the mare. VII. Embryonic development, early embryonic death, foetal losses and their relationship with serum progestagen. J S Afr Vet Assoc 1998;69:150–5. 17. Wilsher S and Allen W. The effects of maternal nutrition on placental and fetal development in maiden Thoroughbred mares. In: Proceedings of a Workshop on Embryonic and Fetal Nutrition, Ravello, Italy. Havemeyer Foundation, 2003; pp 70–1. 18. Wilsher S and Allen W. Factors controlling microcotyledon development and placental efficiency in Thoroughbreds: influences of mare age, parity, and nutritional insults. In: Proceedings of a Workshop on The Equine Placenta, University of Kentucky. Kentucky Agricultural Experiment Station, 2003. 19. McMullen S and Wathes D. Acute nutrient restriction during mid-pregnancy in the ewe: responses and consequences. In: Proceedings of a Workshop on Embryonic and Fetal Nutrition, Ravello, Italy. Havemeyer Foundation, 2003; pp 45–8. 20. Offenbacher S, Katz V, Fertik G, Collins J, Boyd D, Maynor G, McKaig R and Beck J. Periodontal infection as a possible risk factor for preterm low birth weight. J Periodontol 1996;67 (10 Suppl.): 1103–13. 21. Lopez NJ, Smith PC and Gutierrez J. Higher risk of preterm birth and low birth weight in women with periodontal disease. J Dental Res 2002;81:58–63. 22. Goepfert AR, Jeffcoat MK, Andrews WW, Faye-Petersen O, Cliver SP, Goldenberg RL and Hauth JC Periodontal disease and upper genital tract inflammation in early spontaneous preterm birth. Obstet Gynecol 2004;104:777–83. 23. Offenbacher S, Boggess KA, Murtha AP, Jared HL, Lieff S, McKaig RG, Mauriello SM, Moss KL and Beck JD. Progressive periodontal disease and risk of very preterm delivery. Obstet Gynecol 2006;107:29–36. 24. Davenport ES, Williams CECS, Sterne JAC, Murad S, Sivapathasundram V and Curtis MA. Maternal periodontal disease and preterm low birthweight: case-control study. J Dental Res 2002;81:313–18. 25. Vettore MV, Leal MD, Leao AT, da Silva AMM, Lamarca GA and Sheiham A. The relationship between periodontitis and preterm low birthweight. J Dental Res 2008;87:73–8. 26. Bobetsis YA, Barros SP and Offenbacher S. Exploring the relationship between periodontal disease and pregnancy complications. J Am Dental Assoc 2006;137 (Suppl. 2): 7S–13S. 27. Giles R, Donahue J, Hong C, Tuttle P, Petrites-Murphy M, Poonacha K, Robers A, Tramontin R, Smith B and Swerczek T. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203:1170–5. 28. Hong C, Donahue J, Giles R, Petrites-Murphy M, Poonacha K, Robers A, Smith B, Tramontin R, Tuttle P and Swerczek T. Equine abortion and stillbirth in central Kentucky during 1988 and 1989 foaling seasons. J Vet Diagn Invest 1993;5:560–6. 29. Wright JR, Stinson D, Wade A, Haldane D and Heifetz SA. Necrotizing funisitis associated with Actinomyces meyeri infection: a case report. Fetal Pediatr Pathol 1994;14:927–34. 30. Pugh D and Schumacher J. Management of the broodmare. In: Proceedings of the Annual Convention of the American Association of Equine Practitioners 1990;36:61–78. 31. McCluskey B, Traub-Dargatz J, Garber L and Ross F. Survey of endophyte infection and its associated toxin in pastures grazed by horses. Proceedings of the Annual Convention of the American Association of Equine Practitioners. 1999;45:213–16.
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32. Cross D, Anas K, Bridges W and Chappell J. Clinical effects of domperidone on fescue toxicosis in pregnant mares. Proceedings of the Annual Convention of the American Association of Equine Practitioners. 1999;45:203–6. 33. Evans T, Youngquist R, Loch W and Cross D. A comparison of the relative efficacies of domperidone and reserpine in treating equine “fescue toxicosis”. Proceedings of the Annual
Convention of the American Association of Equine Practitioners. 1999;45:207–9. 34. Palmer E and Driancourt M. Use of ultrasonic echography in equine gynecology (a). Theriogenology 1980;13:203–9. 35. Riddle W. Preparation of the mare for normal parturition. Proceedings of the Annual Convention of the American Association of Equine Practitioners. 2003;49:601.
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290.
Dietary Support for Geriatric Broodmares and Stallions Sarah L. Ralston
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291.
Nutrition for the Broodmare Laurie M. Lawrence
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Chapter 290
Dietary Support for Geriatric Broodmares and Stallions Sarah L. Ralston
Introduction In studies done in the mid-1980s horses over 20 years old were reported to have metabolic and digestive changes that necessitated dietary alterations.1, 2 However, based on more recent research3–5 (S.L. Ralston, personal experience, 1990–2007) not all old horses should be considered ‘geriatrics’, and management recommendations have been revised.3 Age alone should not be a criterion for retirement or special management. If the horse is in good body condition, healthy, and reproductively sound, even at 20+ years, their routine feeding probably does not need to be changed. However, if an aged horse has some of the problems listed in Table 290.1, it may be a candidate for special care (Table 290.2).
Management of changes and problems associated with aging Arthritis As with human athletes, years of stress, injuries and general wear and tear can result in painful and crippling arthritic changes in older horses that can potentially compromise their reproductive ability. Otherwise healthy horses over the age of 20 have reduced capacity for anaerobic work,5 altered endocrine responses to maximal exercise,6 and reduced thermoregulatory responses.7 However, mild exercise seems to be beneficial, even to the arthritic older horse (S.L. Ralston, personal experience, 1990–2007). Use of anti-inflammatory drugs or other therapies such as chondroitin sulfate/glucosamine formulations, acupuncture, or chiropractic adjustments that reduce the animal’s discomfort will mitigate metabolic responses to chronic stress that can nega-
tively impact reproductive function. If the animal has limited mobility it is very important to avoid obesity. The extra weight and attendant metabolic changes associated with obesity will increase the stress on the animal’s legs and contribute to other problems, such as laminitis.
Weight loss/poor condition Age alone does not cause weight loss in horses,4, 8 and many causes of reduced body condition are treatable, even in extremely old horses. The most common causes of weight loss in aged horses are poor dentition, reduced absorption/digestion of nutrients (possibly due to chronic parasitic scarring of the gastrointestinal mucosa), inadequate shelter in extreme weather, and debilitating diseases, such as chronic renal, cardiac, or hepatic dysfunction and/or tumors.8 In an older horse that is failing to maintain adequate bodyweight, despite good deworming schedules, normal appetite, and an adequate ration, the teeth should be checked first, using a full mouth speculum if possible. Merely pulling the tongue to one side to inspect the teeth is not sufficient since significant hooks can develop on the most caudal molars. Minor (<3 mm) points and hooks will not adversely affect digestion,9 and there is some controversy whether even significant wave mouth adversely impacts digestion but it is prudent to correct significant points or wave mouth if possible. However, excessive floating should be avoided, especially if the remaining teeth are loose or severely worn down. Despite good dental care, many aged horses have reduced dentition and have some difficulty chewing (SL Ralston, personal experience, 1990– 2007; DL Foster, personal communication, 2006). See
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 290.1 Conditions requiring special attention in aged horses. See text for more details Condition
Clinical signs
Causes/management considerations
Arthritis
Chronic lameness Bone deformity around joints Inflexible joints
Shoeing/trimming Adequate bedding Avoid obesity Anti-inflammatory therapy/ supplements
Weight loss
Inability to maintain good body condition despite good teeth, and a ration that is adequate for mature horses
Dental abnormalities Dieta Shelter Deworming Liver or kidney failure Tumors Malabsorption
Inadequate dentition
Sharp points on molars Loss of teeth Inability to chew feed ‘quidding’ of hay
Regular dental care
Failure to shed winter coat in the summer
Grooming/clipping Dieta Routine vaccinations maintained Water access Drug therapy(?)
Pituitary/thyroid dysfunction:
Recurrent viral infections Chronic founder (laminitis) Increased water intake and urination Excessive weight loss (pituitary) or gain (thyroid)
Dieta Supplements
Kidney/liver failure
Weight loss Lethargy Poor appetite Difficult or frequent urination (kidney) Jaundice (liver)
Gray hair appearing around ears, eyes and forehead
This is not a problem, merely a sign of aging
a
Dieta
See diet recommendations in Table 290.2.
below for recommendations for feeding horses with inadequate dentition. If the teeth are normal, a thorough physical exam and check for chronic infections, and cardiac, liver, or kidney dysfunction should be performed before drastic dietary changes are made, since what is optimal for an otherwise healthy thin old horse may exacerbate renal or hepatic dysfunction.8 If no other abnormalities are found, the horse may be suffering malabsorption of nutrients and/or other alterations in digestion. A simple test for the ability of a horse to digest and absorb nutrients is to do a ‘grain test’. In a horse fasted overnight, take a blood sample for plasma glucose/insulin analysis before feeding 1.0 to 1.5 kg of a concentrated feed, preferably a textured feed containing molasses. Take subsequent samples at 60 and 90 min after feeding, taking into account how long it took the horse to consume the feed (should be < 30 min). The tests should be done in the morning since glycemic responses tend to be blunted in the afternoon.10, 11 Blood glucose should increase by at least 20 mg/dl over the baseline and
insulin should be between 20 and 120 µIU/mL at 60 and/or 90 min. If significantly lower concentrations are seen the horse may have delayed gastric emptying or malabsorption/maldigestion, and definitive tests to determine the type and extent of abnormalities present should be pursued if possible.10 In horses with poor absorption/digestion of nutrients, regardless of cause, a ‘senior’ type ration may help. Such rations should provide at least 12% protein, with restricted calcium (< 1.0%) and slightly increased phosphorus (0.3–0.5%).4 The calcium/phosphorus ratio, however, should be greater than 1:1. Though scientific data are lacking on actual fiber requirements, it is empirically recommended that total ration crude fiber content be over 12%. Digestibility of the concentrates should be maximized by processing such as extrusion or ‘predigestion’ of ingredients. A typical ration for an affected 500-kg horse might consist of free access to top quality hay, preferably a straight grass or grass-legume mix and/or pasture plus 1–4 kg of a feed designed for old horses divided into two or three feedings a day.
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Table 290.2 Dietary management of conditions associated with aging in horses Condition
Recommended diet characteristics
Feeds/supplements
Weight loss not due to liver or kidney failure
12 to 14% protein, 7 to 10% fat High digestibility Easily chewed
Grass or grass mix hay or forage based cubes Complete pelleted or extruded feeds Good quality pasture 100–500 mL edible oil/day Yeast culture products Brewer’s yeast Beet pulp (soaked) Soybean meal (150–250 g/day) Avoid poor quality or high-fiber hay
Inadequate dentition
Easily chewed
‘Soups’ of complete pelleted or extruded feeds Soaked hay cubes or beet pulp Avoid coarse hay, whole grains
Pituitary/thyroid tumors
Reduced starch Highly digestible If chronic infections
Low-molasses, high-fat/-fiber feeds Good quality hay or pasture (if not foundered) Vitamin C (0.01 g/kg bodyweight twice a day)
Kidney failure
Restricted calcium, protein, phosphorus
Grass hay Corn or processed milo Complete feeds designed for adult, not aged horses. Avoid legumes, wheat bran, beet pulp
Liver failure
Restricted protein Increased starch Increased B-vitamins Increased vitamin C
Grass hay, corn, 10% protein sweet feeds Sweet feeds designed for maintenance B-complex supplement orally Vitamin C (0.01 g/kg bodyweight twice a day) Avoid legumes, high-fat rations
Water and salt should be available free choice. Avoid straight legume forages or other feeds with high calcium content that may exacerbate failing kidney function (see below). Live yeast culture products (Saccharomyces cerevisiae) have been reported to improve digestion in horses, especially those fed rations high in concentrates/low in fiber,12, 13 and may be of benefit in the failing aged horse’s rations. Supplementation of edible oils in limited amounts per feeding (no more than 150 mL top dressed per feeding or 10% of total calories) may help the old horse to maintain weight and condition as long as the liver is fully functional. Make all dietary changes slowly, gradually introducing new feeds over the course of 4 to 5 days. Older horses are more sensitive to severe weather, be it heat or cold, and often suffer weight loss when temperature fluctuations are extreme.1, 7 It is essential that adequate shelter be provided and that the higher energy needs in winter are met by providing increased feed in a more highly digestible form such as pelleted or extruded feeds. Constipation/impaction problems can be reduced by insuring free access to clean, unfrozen water in the winter. If the horse does not drink well, feeding water-soaked feeds will help increase fluid intake. Addition of salt to the feed may
also encourage increased water intake but should be done only if the horse has unlimited access to water. Inadequate dentition/tooth loss Inadequate dentition not only predisposes the horse to weight loss but also increases the risk of impactions and choke. Older horses, especially those known to have missing molars, should have their teeth checked at least twice a year. If chewing is difficult or the horse continues to ‘quid’ despite dental correction, ‘soups’ of soaked hay cubes or beet pulp plus pelleted or extruded feeds designed for old horses should be offered. However, soaking the feed is usually necessary only if the animal has a tendency to choke or has severe difficulty in chewing. The soaked feeds can ferment (summer) or freeze (winter) easily so should only be offered in amounts that the horse will consume easily in a single meal. This may require that the horse be fed three or more times a day to meet its nutritional needs. Hay should still be offered if possible, even if most of it is wasted. Access to good pasture is also desirable. Restriction of forage and pasture has been well documented to increase the incidence and severity of gastric ulceration.14, 15
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Pituitary/thyroid dysfunction 1
In an early study of geriatric horses, over 70% of the horses over the age of 20 used had at least subclinical signs (altered glucose and cortisol metabolism) of pituitary/thyroid dysfunction. Table 290.1 summarizes the classic clinical signs of pituitary/thyroid dysfunction. Old horses with pituitary and/or thyroid neoplasias documented at necropsy, even in the early preclinical stages, had lower blood vitamin C than did unaffected or younger mares.1 This may explain in part the increased susceptibility to infections observed in many older horses. Both types of dysfunction cause relative glucose intolerance, in which the horse becomes less sensitive to the action of insulin, resulting in hyperinsulinemia and increased risk of laminitis.16, 17 Nutritional management of the clinical problems associated with pituitary or thyroid dysfunction is fairly easy.8 Avoid feeding sweet feeds (>3% molasses added) and offer concentrates that have restricted starch/sugar content (< 30% non-structural carbohydrates). Unfortunately many of the ‘senior’ formulations currently available contain high concentrations of molasses. Check the formulation carefully before recommending a particular product. Formulations with higher fat (7% to 10%) should be used in the thin horse; however, high-fat formulations are not desirable if the horse is overweight. If chronic infections are a problem (skin infections, thrush, hoof abscesses), 5–10 g of ascorbic acid (vitamin C) added to the feed twice a day may be beneficial. If water intake and urine output are increased secondary to the pituitary dysfunction, fresh, clean, unfrozen water should be available free choice. Thick hair coats should be clipped in the summer in addition to providing shelter from the sun. Cyproheptadine and pergolide, though not approved for use in horses, can be effective in controlling the clinical signs of pituitary dysfunction. Both drugs are expensive, however. Dosages and treatment protocols have been discussed in detail elsewhere.16,18
Reduced kidney/liver function Chronic kidney or liver failure is not as common in aged horses as it is in cats and dogs, but it still can occur. The degeneration of kidney and liver function will alter nutritional needs of the affected animals.19 Reduced kidney function will result in renal stones (calculi), bladder stones, weight loss, loss of appetite, and potentially death. Horses are unique in that they excrete excess dietary calcium primarily through their urine instead of their feces, as other animals do. As a result, if kidney function is re-
duced, renal and bladder ‘stones’ of calcium oxalate are more likely to occur as well as an increase (potentially lethal) in blood calcium. Horses with kidney failure should be put on restricted calcium rations (0.5 to 0.7% calcium on a dry matter basis). Based on data from other species, protein and phosphorus also should be restricted to 8–12% and 0.25% respectively, though lactating mares will need somewhat higher calcium, phosphorus, and protein intakes. Good-quality grass hay and corn or a complete pelleted ration for mature (not aged) horses are the feeds of choice. Avoid legumes (alfalfa and clover), wheat bran, rice bran, and beet pulp due to high calcium (legumes, beet pulp, calcium-balanced rice bran) or phosphorus (wheat bran, rice bran) content. Hepatic failure results in weight loss, lethargy, jaundice (yellow mucous membranes), loss of appetite, and intolerance of fat and protein in the diet.19 If severe, the horse may show behavioral changes such as irritability, aimless wandering or circling, or pressing its head against objects (hepatoencephalopathy). Affected horses require increased simple carbohydrate sources to maintain their blood glucose levels and are intolerant of high protein or fat in the diet. The diet should emphasize starch intake (grains or concentrates), though fiber sources (hay, beet pulp) are still necessary to avoid gastrointestinal dysfunction and ulceration. Grass hay, sweet feeds with 10–12% protein, and corn are recommended components of the ration. Wheat bran and beet pulp are acceptable supplements in these cases. Rice bran should be avoided due to its high fat content. Since the liver is the site of niacin and vitamin C synthesis in the horse, daily oral supplementation with vitamin B-complex and ascorbic acid may be beneficial.
Summary A broodmare or stallion should not be treated differently just because it has reached a certain chronological age. However, if problems related to aging are present, changes in management and medications may be needed to keep the older horse comfortable. Aged horses that do not have reduced kidney function will benefit from a diet more similar to that recommended for weanlings than that for normal adult maintenance. Calcium and protein, however, will need to be restricted if the horse has kidney failure. Supplements that may help include vitamin C, yeast cultures, brewer’s yeast, and vegetable oil. If teeth are a problem, soups can be made out of cubed, extruded, and/or pelleted feeds. Teeth should be checked every 6 months or even more frequently if premolars or molars are missing. Vaccinations and deworming schedules should be carefully maintained.
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Dietary Support for Geriatric Broodmares and Stallions Pituitary and thyroid dysfunctions are very common in aged horses. Management of the clinical signs include clipping long hair coats in summer, feeding low-starch complete feeds, and perhaps vitamin/mineral supplementation. Adequate shelter is a must for older horses, especially in the winter. However, confinement of an arthritic old horse to a stall is not doing the animal any favors.
References 1. Ralston, SL, Nockels CF, Squires EL. Differences in diagnostic test results and hematologic data between aged and young horses. Am J Vet Res 1988;49:1387–92. 2. Ralston SL, Squires, EL, Nockels, CF. Digestion in the aged horse. J Eq Vet Sci 1989;9:203–5. 3. Ralston SL, Christensen RA, Malinowski K, Scanes CG, Hafs, HD. Chronic effects of equine growth hormone eGH on intake, digestibility and retention of nutrients in aged mares. J Anim Sci 1996;74(Suppl. 1):194. 4. Ralston SL, Breuer LH. Field evaluation of a feed formulated for geriatric horses. J Eq Vet Sci 1996;16:334–8. 5. McKeever KH, Malinowski K. Exercise capacity in young and old horses. Am J Vet Res 1997;58:1468–72. 6. McKeever KH, Malinowski K. Endocrine response to exercise in young and old horses. Eq Vet J Suppl 1999;30:561–6. 7. McKeever KH, Eaton TL, Geiser S, Kearns CF. Thermoregulation in old and young horses during exercise. Med Sci Sports Exercise 2000;32:156 abstract 683. 8. Ralston SL. Evidence Based Equine Nutrition. In: Veterinary Clinics of North America Equine Practice: Evidence Based Veterinary Medicine. Ramey DW (ed.) Vet Clin Equine 2007;23:365–384. 9. Ralston SL, Foster DL, Divers T, Hintz, HF. Effect of dental correction on feed digestibility in horses. Eq Vet J 2001;334:390–3.
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10. Ralston SL. Insulin and glucose regulation. In: Messer NT, Johnson PJ (eds) Veterinary Clinics of North America Equine Practice: Endocrinology. Vet Clin N Am-Equine 2002;18:295–304. 11. Ralston SL. Hyperinsulinemia and glucose intolerance. In: Reed S, Bayly W, Sellon D (eds) Equine Internal Medicine, 2nd edn. St Louis, MO: Elsevier, 2004; pp. 1599– 603. 12. Medina B, Girard ID, Jacotot E, Julliand V. Effect of a preparation of Saccharomyces cerevisiae on microbial profiles and fermentation patterns in the large intestine of horses fed high fiber or a high starch diet. J Anim Sci 2002;80:2600–9. 13. Hall MM, Miller-Auwerda PA. Effect of Saccharomyces cerevisiae pelleted product on cecal pH in the equine hind gut. In: Proceedings of the 19th Equine Science Society Symposium, Tucson, AZ, 2005; pp. 45–6. 14. Andrews FM, Buchanan BR, Elliot SB, Clariday NA, Edwards LH. Gastric ulcers in horses. J Anim Sci 2005;83:E18–E21. 15. Meyer H, Coenen M, Gurer C. Investigations of saliva production and chewing in horses fed various feeds. In: Proceedings of the 9th Equine Nutrition and Physiology Society Symposium, East Lansing, MI, 1985; pp. 38– 41. 16. Beech, J. Tumors of the pituitary gland pars intermedia. In: Robinson NE (ed.) Current Therapy in Equine Medicine, 2nd edn. Philadelphia, PA: WB Saunders Co, 1987; pp. 182– 5. 17. Dybdal NO, Hargreaves KM, Madigan JE, et al. Diagnostic testing for pituitary pars intermedia dysfunction in horses. J Am Vet Med Assoc 1994;2044:627–32. 18. Messer, NT. Endocrine Dysfunction in the aged horse. In: Bertone J (ed.) Equine Geriatric Medicine and Surgery. St Louis, MO: Elsevier Publishing, 2006; pp. 59–67. 19. Ralston, SL. Clinical nutrition of adult horses. In: Hintz HF (ed.) Clinical Nutrition. Vet Clin N Am-Equine 1990;6:339–54.
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Chapter 291
Nutrition for the Broodmare Laurie M. Lawrence
Optimal feeding programs for broodmares adjust to the changing nutrient needs of the mare as she progresses through a gestation–lactation cycle. The diet should provide nutrients for the maintenance of the mare, the development of the fetus, and the production of milk. Failure to meet these nutrient needs may affect the reproductive efficiency of the mare, the development of the foal, and possibly the longevity of the mare in the breeding herd.
Reproductive efficiency Body condition has been shown to affect several aspects of reproductive efficiency. The most common body condition scoring (BCS) system is based on visual and manual appraisal of the amount of fat on the neck, withers, shoulder, ribs, tail-head, and back. A BCS of 1 is used for a horse that is emaciated and a BCS of 9 is used for a horse that is extremely fat (Table 291.1). A BCS of 5 is considered moderate. Mares that enter the breeding season at a BCS below 5 may require more cycles to become pregnant than mares that enter the breeding season at a BCS of 5 or higher.1 Similarly, pregnancy rates have been reported to be lower in mares with a BCS below 5 than in mares with a BCS above 5.1 The improvements in cycles per pregnancy and pregnancy rate tend to plateau (but not decrease) once BCS exceeds 5, so there is no need for mares to be excessively fat. Open mares (barren or maiden) entering the breeding season in thin body condition take longer to reach first ovulation than mares in moderate or fat body condition.1, 2 Thin mares have also been reported to have less regularity in hormonal patterns during the estrous cycle3 and thin mares experience a longer winter anovulatory period than fat mares.4 Failure to enter winter anestrus has been reported to occur more commonly in fat mares than in thin mares;4 however, it has been suggested that a very high BCS can result in abnormal reproductive patterns.5 However, other researchers have not found
any negative effects of high BCS on conception rates, services per conception, foaling ease, interval to first postpartum ovulation, or interval to second postpartum ovulation.1, 6, 7 To maximize reproductive efficiency, feeding programs should ensure that mares enter the breeding season with a condition score of at least 5. In practical situations it is not easy to attain a body condition of exactly 5 in all mares. As reproductive efficiency is not negatively affected by somewhat higher BCS, feeding programs should be designed to keep all mares at BCS between 5 and 6. While underfeeding will depress reproductive efficiency, there is very little evidence that supplementation of energy or other nutrients beyond required levels will enhance reproductive efficiency. The nutrient requirements and suggested feeding management of non-pregnant, pregnant, and lactating mares are discussed below.
Non-pregnant mares The nutrient requirements for non-pregnant mares weighing 400 or 600 kg are shown in Table 291.2. The requirements for non-pregnant mares are the same as for adult horses at maintenance that are not receiving any regular imposed exercise. There are three levels of energy and protein intakes suggested for adult horses at maintenance. These three levels are designed to allow for individual differences among horses. The ‘minimum’ level of energy and protein is suggested for horses that have traditionally been considered ‘easy keepers’ and the ‘elevated’ level is suggested for horses that have traditionally been termed ‘hard keepers’. The nutrient requirements of non-pregnant mares in thermoneutral environmental conditions can usually be met with good quality forage and a small amount of concentrate or other supplement. When horses are exposed to climates outside their thermoneutral zone the energy intakes listed in Table 291.2 will not be
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 291.1 A system for evaluating body condition in horses. This scoring system is based on palpation and visual appearance of six target areas: the crest of the neck, the withers, the loin, the tail-head, the ribs, and the shoulder Score
Brief descriptiona
1
Bone structure of neck, spine, shoulder, ribs, and pelvis is easily apparent. No subcutaneous fat in any location, bones are easily palpated under skin. Horse is in very poor condition
2
Bone structure of shoulder, withers, ribs, and pelvis is apparent. Neck is very thin but bone structure is not visible. Vertebrae in back and loin may have some fat cover but are still prominent. Horse is in poor condition
3
Neck is thin and withers are prominent. Ribs are visible. Points of hips (hook bones) are prominent. Horse is obviously thin
4
Horse is moderately thin. Ribs are slightly visible when horses have a short coat. Spinal vertebrae are still somewhat prominent in loin. Neck is not unusually thin and bony structure of withers is not apparent. Some fat cover over tail-head. Points of hips are not prominent
5
Horse is in moderate condition. Ribs are not visible when horse has short coat. On palpation of the ribs, tissue has filled in between the ribs but the ribs are easily felt. Neck and shoulder blend smoothly into body. Back is level in the loin area and some fat is accumulating on tail-head
6
Some fat accumulating along topline of neck, at the withers, and behind the shoulder. Some fat has accumulated on the ribs and at the tail head. Fat may have accumulated on either side of the loin creating a slight crease over the lumbar spine
7
Noticeable fat has accumulated along the neck, withers, tail-head, and behind the shoulder. Ribs can be felt but fat cover is thick. Fat has accumulated on either side of the lumbar spine creating a crease along the loin
8
Neck is thick. Fat has filled in over withers and behind shoulder. Tail-head is soft. Ribs are difficult to feel. Crease along spine is clear. Horse is obviously fat
9
Flank has filled with fat; body parts have lost definition. Horse may have bulging fat in neck and over ribs and crease along spine is very deep. Horse is very, very fat
Adapted from Henneke et al.47 For more complete descriptions, see Henneke et al.47 and NRC.9
a
sufficient to maintain bodyweight or condition. Very cold weather, wind, and precipitation can increase heat loss and thus increase the amount of energy used by horses for maintenance of body temperature. Similarly, walking through snow or mud will increase energy use. These harsh environments can increase energy requirements by 50% or more, so additional feed should be provided when the horses are kept in adverse conditions. Body condition in non-pregnant mares should be assessed several months before the onset of the breeding season. Mares in low body condition at the onset of the breeding season may begin to cycle later in spring, require more services per conception, and have a lower overall pregnancy rate. The best time to adjust body condition is in the early fall. If mares have a BCS below 5, energy intake should be increased to allow mares to gain enough weight to raise their BCS to at least 5 by the beginning of the breeding season. If assessment and dietary intervention are delayed until winter it will be more difficult to obtain the desired weight gain before the beginning of the breeding season. By evaluating body condition well in advance of the breeding season, there is a broad window of opportunity to achieve the desired result. Table 291.3 provides guidelines for estimated energy and concentrate intakes needed to raise the BCS of non-pregnant, 400-, and 600-kg mares from 4 to 5 when the programs are imposed at different
points prior to the breeding season. If dietary intervention is imposed in early fall, adequate weight gain can be accomplished with a diet composed mostly of good quality forage (pasture or hay) and small amounts of concentrate. If dietary intervention is delayed until late winter, it will be necessary to feed larger amounts of concentrate. High concentrate intakes may increase the risk of gastrointestinal disturbances,8 therefore it is better to begin the dietary intervention sooner and reduce the reliance on concentrate. Implementation of the following recommendations for non-pregnant mares should ensure that the mares enter the breeding season with a condition score of at least 5.
r r
r
Each mare should be condition scored regularly. Condition scoring can be performed at the same time as other routine health procedures such as deworming or hoof care. Use the condition scoring system described in Table 291.1. This system uses both visual appraisal and manual palpation of the target areas. Visual appraisal alone may be inaccurate when mares have a heavy winter coat. Begin condition scoring by early fall. When body condition falls below 5 or increases above 7, dietary changes should be considered.
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Mare: Nutrition Table 291.2 Recommended nutrient intakes9 for non-pregnant mares weighing 400 or 600 kga Nutrient Digestible energy (Mcal/day) Minimum Average Elevated Crude protein (g/day) Minimum Average Elevated Lysine (g/day) Minimum Average Elevated Calcium (g/day) Phosphorus (g/day) Copper (mg/day) Zinc (mg/day) Manganese (mg/day) Selenium (mg/day) Iodine (mg/day) Vitamin A (IU/day) Vitamin E (IU/day) Vitamin D (IU/day) Sodium (g/day) Chloride (g/day) Potassium (g/day)
400-kg mares
600-kg mares
12.1 13.3 14.5
18.2 20.0 21.8
432 504 576
648 756 864
18.6 21.7 24.8 16.0 11.0 80 320 320 0.8 2.8 12 000 400 2640 8.0 32.0 20.0
27.9 32.5 37.2 24.0 16.8 120 480 480 1.2 4.2 18 000 600 3960 12.0 48.0 30.0
a
Breeds representative of the lower bodyweight category include Arabian, Morgan, and Tennessee Walking Horse. Breeds representative of the larger weight category include Thoroughbreds, Quarter Horse, and some Warmbloods. Within every breed there will be a range of mature bodyweights and sizes. From National Research Council (NRC), 2007.9
r r
If mares are not fed in individual stalls or pens, mares with similar requirements should be grouped together to minimize underfeeding or overfeeding of individuals. When it is not possible to group mares by nutrient status, thin mares should be separated at least once a day and given extra concentrate. If is not possible to feed thin mares in a separate stall or pen, nose
r r
bags can be used effectively to provide individualized concentrate intakes. When it is not possible to feed thin mares individually, provide sufficient feed to make certain that the thinnest mare in a group has a BCS of 5. Energy intake should be increased during adverse weather to prevent weight loss.
Table 291.3 Estimated daily digestible energy (DE) and suggested feed intakes needed to increase the condition scores of moderately thin non-pregnant, 400-, and 600-kg mares prior to the onset of the breeding seasona
400-kg mares
600-kg mares
a
Interval for dietary intervention
Estimatedb daily DE intake(Mcal)
Estimated daily hay intake (kg)c
Estimated daily concentrated intake (kg)
Oct 1–Feb 1 Nov 1–Feb 1 Dec 1–Feb 1 Oct 1–Feb 1 Nov 1–Feb 1 Dec 1–Feb 1
15–17 Mcal 16–18 Mcal 17–20 Mcal 22–26 Mcal 23–27 Mcal 26–30 Mcal
7.0 kg 7.0 kg 7.0 kg 10.0 kg 10.0 kg 10.0 kg
0.5–1.5 kg 1.5–2.0 kg 2.0–2.5 kg 1.0–2.5 kg 2.0–3.0 kg 3.0–4.0 kg
To increase mares to a condition score of ‘5’ (moderate) by February 1 over the interval indicated. These values may underestimate the DE requirements of horses kept in harsh climates. c As fed basis. Based on average quality hay in winter and average quality pasture in fall. Forage intake could be higher with high-quality hay or pasture. d As fed basis. Based on average concentrate containing 3.1 Mcal DE/kg. Concentrate intakes could be reduced if high-quality hay or pasture is used. Higher concentrate intakes could be necessary if mares are kept in harsh climates. b
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Table 291.4 Nutrient requirements of 550-kg pregnant maresa
Expected bodyweight Digestible energy (Mcal/day) Minimum Average Elevated Crude protein (g/day) Minimum Average Elevated Lysine (g/day) Minimum Average Elevated Calcium (g/day) Phosphorus (g/day) Copper (mg/day) Zinc (mg/day) Manganese (mg/day) Selenium (mg/day) Iodine (mg/day) Vitamin A (IU/day) Vitamin E (IU/day) Vitamin D (IU/day) Sodium (g/day) Chloride (g/day) Potassium (g/day)
Early
5th month
6th month
7th month
8th month
9th month
10th month
11th month
550 kg
554 kg
559 kg
566 kg
575 kg
587 kg
603 kg
623 kg
16.7 18.3 20.0
17.2 18.8 20.5
17.6 19.2 20.9
18.1 19.7 21.4
18.7 20.3 22.0
19.6 21.2 22.9
20.6 22.2 23.9
21.9 23.5 25.2
594 693 792
654 753 852
675 774 873
702 801 900
736 835 934
777 876 975
826 925 1024
883 982 1081
26 30 34 22 15 110 440 440 1.1 3.8 33 000 880 3630 11 44 28
28 32 36 22 15 110 440 440 1.1 3.8 33 000 880 3630 11 44 28
29 33 37 22 15 110 440 440 1.1 3.8 33 000 880 3630 11 44 28
30 34 38 31 22 110 440 440 1.1 3.8 33 000 880 3630 11 44 28
32 36 40 31 22 110 440 440 1.1 3.8 33 000 880 3630 11 44 28
34 38 42 40 29 138 440 440 1.1 4.4 33 000 880 3630 12 45 28
36 40 44 40 29 138 440 440 1.1 4.4 33 000 880 3630 12 45 28
38 42 46 40 29 138 440 440 1.1 4.4 33 000 880 3630 12 45 28
a Adapted from NRC.9 Other requirements include: thiamin, 33 mg/day; riboflavin, 22 mg/day; sulfur, 16.5 g/day; cobalt, 0.6 mg/day; iron, 550 mg/day, magnesium, 9 g/day.
Pregnant mares Diets for pregnant mares should provide nutrients to support the maintenance of the mare’s body and for the synthesis of the products of conception (including the fetus, the placenta, and the mammary gland). Pregnant mares also need additional nutrients to support the maintenance of the newly synthesized tissues, which have a relatively high metabolic rate.9 Furthermore, if a pregnant mare is thin, she must receive enough additional feed to increase her BCS prior to parturition and lactation. The following discussion of the nutrition of the pregnant mare is based on these concepts. Although the accretion of fetal tissue is concentrated in the last trimester of gestation, placental development and uterine hypertrophy begin well before the majority of fetal weight gain. The most recent edition of the National Research Council (NRC) publication Nutrient Requirements of Horses9 suggests that small increases in energy intakes are justified as early as the fifth month of gestation (Table 291.4) to support placental and uterine development. The tissues of conception are primarily water
and protein, therefore amino acid requirements begin to increase during the fifth month of gestation as well. Table 291.4 gives estimates of both the crude protein and the lysine requirements of pregnant mares. Dietary protein quality should be considered to be just as important as protein quantity; it is more important to meet amino acid requirements than to meet crude protein requirements. If the diet meets the crude protein requirement but does not meet the lysine requirement then the amount of crude protein should be increased until the lysine requirement is met. Mares that begin gestation in a moderate body condition are expected to gain weight during the pregnancy. Total weight gain is expected to be 12–15% of the mare’s initial bodyweight. Foal birth weight is usually between 9% and 10% of a mare’s normal bodyweight and the remaining weight gain includes the placental tissues, fluids, and increased uterine tissue.9 It could be expected that the greatest changes in mare weight would coincide with the period of greatest fetal development in late gestation. However, two observational studies have reported that mares in
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Mare: Nutrition
good body condition may gain less bodyweight in the last 2 months of gestation than would be predicted from the expected fetal weight gain.10, 11 In both studies, the total weight gain of the mares was 12–15% of non-pregnant bodyweight and the mares produced foals of normal size. When mares were observed for an entire gestation period, the majority of weight gain occurred in midgestation, and it was suggested that mares might augment body condition in midgestation and then mobilize body stores in late gestation.11 When pregnant mares are fed diets that are slightly deficient in energy, protein, or other nutrients, it appears that they will sacrifice their own body stores in order to produce normal-sized foals.12 However, thin mares may experience a prolonged gestation. Loss of body condition in late gestation (BCS change from 5.5 to 3.4 at foaling) resulted in an increase in gestation length of about 9 days compared with mares that maintained BCSs between 6 and 7.3 Similarly, a mean gestation length of 350 days was reported for a group of mares that lost 4% of bodyweight in the last 90 days of gestation.13 As noted above, body condition at the onset of the breeding season will influence reproductive efficiency, so mares that foal with BCS below 5 would be expected to have reduced rebreeding efficiency. In addition, mares that are thin at parturition and remain thin during lactation may be at increased risk for early fetal loss if they do become pregnant.1, 12 The vitamin requirements of the pregnant mare have received little study. Pasture is an excellent source of many nutrients, and grazing mares usually have high vitamin A and vitamin E status. However, vitamin A and E status will decline when mares are removed from pasture and fed hay and unsupplemented cereal grains.14, 15 The NRC9 recommends that the diets of pregnant mares provide 60 IU vitamin A/kg bodyweight and 1.6 IU of vitamin E/kg bodyweight. Pasture-based diets will often meet or exceed these requirements. Mares grazing pasture will probably have higher vitamin A and vitamin E status than mares fed stored feeds, even when the stored feeds are supplemented to contain the NRCrecommended levels of these vitamins.15, 16 It is not known how much vitamin A or E is optimum for pregnant mares. In one study, there was a trend for supplemental vitamin E above the NRC9 recommendation to enhance immunoglobulin concentrations in the colostrum of broodmares that were not receiving pasture,17 but confirmatory studies have not been performed. Deficiencies, imbalances, or excessive intakes of minerals by pregnant mares have the potential to affect the fetus as well as the mare. Lameness, swollen joints, and generalized soft-tissue mineralization have been reported in foals of mares that were
fed a low-calcium/high-phosphorus diet.18 Foals from iodine-deficient mares may have thyroid hypertrophy (goiter), flexural deformities, or be weak at birth.9 Thyroid hypertrophy may also occur in response to very high iodine intakes. Many soils in the USA are deficient or marginal in selenium, and forages grown on those soils may not provide adequate selenium for pregnant mares. Selenium deficiency is believed to be the primary cause of white muscle disease in foals, although vitamin E deficiency may be involved as well.9 A selenium intake of 0.002 mg/kg bodyweight (1 mg for a 500-kg mare) has been recommended for pregnant mares.9 Most commercially manufactured concentrates that are formulated for pregnant mares in the USA will meet this requirement when fed at the rate recommended by the manufacturer. Copper deficiency can cause articular lesions in several species, and a link between copper nutrition and developmental orthopedic disease was proposed more than 20 years ago. It has now been suggested that the lesions resulting from copper deficiency and those observed in most growing horses have distinct differences and that the role of copper in equine osteochondrosis has been overemphasized.19, 20 Several studies have examined the effects of maternal copper status on the copper status or skeletal characteristics of foals.19,21–27 One study reported that copper supplementation of pregnant Thoroughbred mares grazing low-copper pastures reduced the incidence of cartilage defects in foals.23 In another study, the copper status of Dutch Warmblood mares did not affect the incidence of osteochondrosis in foals, but did appear to affect the improvement of lesions as the foals aged.25 Other studies failed to confirm a relationship between mare copper status (determined by liver biopsy) and the incidence of osteochondrosis,19, 27 and treating mares with copper did not affect the evidence of clinical abnormalities of the physis.26 These recent studies do not support a strong association between maternal copper status and the incidence of developmental orthopedic disease. However, in some studies mares were given supplemental copper by intramuscular injection rather than as a dietary supplement; therefore, the dietary copper requirement of pregnant mares remains controversial. The suggested copper requirement for mares in late gestation is 0.25 mg/kg bodyweight (125 mg for a 500-kg mare). When fed with typical North American forages, most commercial concentrates formulated for pregnant mares in the USA will provide enough copper to meet this requirement. Pregnant mares should be fed good quality forage and enough concentrate to meet nutrient needs. Legume or legume–grass hays are usually higher
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Nutrition for the Broodmare in energy, protein, calcium, and palatability than grass hays. Forages harvested in late maturity (larger stems, visible seed heads) will be lower in nutrients and will be less palatable to the mares, necessitating greater concentrate intakes. Pasture can be a palatable and high-quality source of nutrients for broodmares, but tall fescue and some ryegrass pasture should be used cautiously. These plants provide acceptable nutrition but they may be infected with an endophyte (Neotyphodium coenophialum). The endophyte may enhance the hardiness of the plant but it produces a variety of compounds, including ergot alkaloids, which can negatively affect mares in late gestation. The common signs of tall fescue toxicosis in pregnant mares are delayed mammary development, agalactia, prolonged gestation, and placental thickening. Dystocia, large weak foals, and premature separation of the allantochorion are also common. Oral administration of domperidone has been reported to be effective in reducing the effects of tall fescue toxicosis, but the best management practice is to prevent mares from consuming endophyte-infected tall fescue pasture or hay during late gestation. Occasionally, mares can graze pastures containing tall fescue without noticeable problems. Some mares may select against tall fescue and graze other plants in the pasture, thus reducing their exposure to the ergot alkaloids or other compounds produced by the endophyte. It also appears that season and/or growing conditions will affect the production of some compounds by the endophyte. In Central Kentucky, concentrations of ergovaline, a specific ergot alkaloid, in tall fescue pasture tend to be relatively low in the winter and then increase with the spring growth, peaking in May or June.28 Therefore, mares that foal early in the year in Kentucky tend to be at lower risk for fescue toxicosis than mares foaling in late spring, even if they are kept in the same pasture. Finally, not all tall fescue is infected with the endophyte. Mares can consume endophyte-free tall fescue safely. Tall fescue varieties with a novel endophyte are also available. The novel endophyte is believed to convey some of the hardiness of the traditional endophyte without producing toxicosis in animals. In general, the nutrition of the pregnant mare has not received much attention from researchers. Accordingly, the nutrient intakes suggested for pregnant mares in Table 291.4 should be viewed as recommendations rather than requirements. Several of the NRC recommendations9, 29 for pregnant mares are based on estimates of tissue accretion in the developing fetus and placenta. Unfortunately, there are no measures of tissue composition of the normal equine fetus during the course of a gestation period and the only available data are from a study that measured
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energy, protein, and mineral composition of aborted and stillborn foals30 or from data in other species.
Lactating mares Following parturition, the mare’s nutrient requirements escalate (Table 291.5). Generally, the amount of forage and concentrate fed to mares will be increased after foaling in order to meet nutrient requirements. Daily voluntary feed intake (forage and concentrate) by lactating mares can be 3% of bodyweight, compared with 2% for mares in late gestation.31 This increase in voluntary feed intake may be driven by nutrient requirements or it could result from increased gastrointestinal capacity postpartum. Postpartum mares may be susceptible to colon volvulus.32, 33 Whether there is a dietary component to this problem is unknown, but implementation of a gradual transition in feed intake and feed composition in the postpartum mare might be prudent. Lactating mares require nutrients to meet the maintenance needs of their tissues and to meet the needs of lactation. A 600-kg mare in early lactation may produce 18–20 kg of milk per day.9 The gross energy of mare’s milk is about 500–550 kcal/kg, so the mare is secreting approximately 8–9 Mcal into the milk each day. The amounts of protein, calcium, and phosphorus secreted into milk are also high. Because there are inefficiencies associated with the conversion of dietary nutrients into milk nutrients, the mare’s diet must contain even more nutrients than would be accounted for by her maintenance requirements and the nutrients found in the milk. Not all mares produce the same quantity or quality of milk. There can be large individual differences among mares of the same breed and primiparous mares may produce less milk than multiparous mares. Mares with above average or below average milk production will require more or less nutrients, respectively, than suggested in Table 291.5. Total milk production declines in late lactation and the chemical composition of mare’s milk may also change somewhat over the course of lactation. However, mares in late lactation still have elevated nutrient requirements. Some dietary factors can affect the composition of milk produced by mares, but the effects of diet on milk yield and composition are often buffered by the ability of the mare to alter her body stores. Moderate overfeeding of energy to lactating mares results in mare weight gain, but may not affect foal growth. 34 Similarly, moderate energy deficiency results in weight loss in lactating mares but may have minimal effect on milk production or foal growth, particularly if the mares have adequate body stores when the energy restriction is imposed.1, 34 Total milk fat,
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Table 291.5 Nutrient requirements of 400- and 600-kg lactating maresa
Digestible energy (Mcal/day) Crude protein (g/day) Lysine (g/day) Calcium (g/day) Phosphorus (g/day) Copper (mg/day) Zinc (mg/day) Manganese (mg/day) Selenium (mg/day) Iodine (mg/day) Vitamin A (IU/day) Vitamin E (IU/day) Vitamin D (IU/day) Sodium (g/day) Chloride (g/day) Potassium (g/day)
400 kg 1st month
400 kg 3rd month
400 kg 5th month
600 kg 1st month
600 kg 3rd month
600 kg 5th month
25.4 1228 67.8 47.3 30.6 100 400 400 1.0 3.5 24 000 800 2640 10.0 36.0 38.0
24.5 1174 64.2 44.7 28.8 100 400 400 1.0 3.5 24 000 800 2640 10.0 36.0 37.0
22.7 1064 57.0 31.6 19.7 100 400 400 1.0 3.5 24 000 800 2640 9.0 36.0 28.0
38.1 1842 101.7 70.9 45.9 150 600 600 1.5 5.3 36 000 1200 3960 15.3 54.6 57.4
36.7 1761 96.4 67.1 43.2 150 600 600 1.5 5.3 36 000 1200 3960 15.0 54.6 55.1
34.0 1596 85.5 47.4 29.6 150 600 600 1.5 5.3 36 000 1200 3960 14.0 54.6 41.7
a
Adapted from NRC.9 For 400-kg mares, other requirements include: thiamin, 30 mg/day; riboflavin, 20 mg/day; sulfur, 16.5 g/day; cobalt, 0.6 mg/day; iron, 440 mg/day; magnesium, 13 g/day. For 600-kg mares, other requirements include: thiamin, 45 mg/day; riboflavin, 30 mg/day; sulfur, 15 g/day; cobalt, 0.5 mg/day; iron, 500 mg/day; magnesium, 9 g/day.
fatty acid composition, and gross energy concentration may be affected by total dietary energy, dietary fat, and possibly the forage–concentrate ratio of the diet, although the effects are not always consistent among studies and the changes are often relatively small.35–38 Protein-deficient diets will result in bodyweight loss in lactating mares, but may not affect foal growth unless the deficiency continues for several weeks or months.12, 39 The mobilization of bone mineral has also been reported in lactating mares, suggesting that milk calcium and phosphorus will be maintained at the expense of the maternal skeleton.40–42 Milk composition is relatively resistant to moderate supplementation of most dietary minerals. Copper, zinc, and iron concentrations in the diets of lactating mares have little effect on the concentrations of these minerals in milk.24, 43 Similarly, increasing the calcium and phosphorus intake of mares in late lactation above recommended levels did not increase milk calcium or milk phosphorus.42 It does appear that diet can influence milk selenium concentrations. The traditional source of supplemental selenium in animal feeds is sodium selenite, but organic forms of selenium are also available. It is possible to increase the concentration of selenium in the colostrum and milk with selenium-enriched yeast, which is an organic selenium supplement.44 The ability of the mare to buffer the effects of diet on milk yield and composition makes it somewhat difficult to alter foal growth through moderate, shortterm manipulation of the lactating mare’s diet.
Although foal growth may not be decreased by short-term deficiencies in the maternal diet, the loss of body stores in the mare may have consequences for subsequent reproductive cycles. As mentioned previously, thin mares often require more services per pregnancy and may have lower pregnancy rates than mares with condition scores above 5. Also, early fetal loss may be increased when mares remain thin during lactation or when they lose weight in response to a protein-deficient diet.1, 12 Mares fed calciumdeficient diets appeared to mobilize more bone mineral than mares fed calcium-adequate diets, and took longer to replenish those stores after weaning.41 The nutritional management of the mare after weaning will depend upon her pregnancy status and her body condition. If a mare is not pregnant and she is in moderate body condition at weaning, dietary management should focus on maintaining body condition and providing adequate mineral support to at least meet the needs for maintenance. If the mare has been rebred and has already reached the fifth month of gestation at the time of weaning, then her diet should reflect the needs of midgestation. Highquality forage (pasture or hay) has the potential to provide almost all of the energy and protein needed by open mares and mares in early or midgestation. However, mineral supplementation may be necessary. If the mare is thin at weaning then the postweaning period is an opportunity to replenish any body stores that were utilized by the mare to meet the needs of gestation or lactation. Non-pregnant mares
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Nutrition for the Broodmare that are thin at weaning should be managed to gain weight before the next breeding season according to the guidelines described earlier in this chapter. For thin mares that were successfully rebred during lactation, dietary intervention should be implemented as soon after weaning as possible. Ideally, any change in a feeding program should be implemented gradually. However, the diets of broodmares are often changed abruptly at weaning. Broodmare managers often reduce the quality of offered forage and withdraw all concentrate from mares at weaning in an attempt to hasten the termination of milk production. The scientific basis for this practice has not been demonstrated and it is likely that the absence of nursing is a more important signal for milk production than nutrient availability. In addition, it has been suggested that sudden restriction of nutrients to pregnant mares may disturb the metabolic and hormonal regulation of the fetal–placental unit and possibly increase prostaglandin production.45, 46 Therefore, it seems advisable to employ a feeding program that gradually transitions mares from a lactation diet to a postweaning diet, particularly if the mare is pregnant.
References 1. Henneke DR, Potter GD, Kreider JL. Body condition during pregnancy and lactation and reproductive efficiency of mares. Theriogenology 1984;21:897–909. 2. Kubiak JR, Crawford BH, Squires EL, Wrigley RH, Ward GM. The influence of energy intake and percentage of body weight on the reproductive performance of nonpregnant mares. Theriogenology 1987;28:587. 3. Hines KK, Hodge SL, Kreider JL, Potter GD, Harms PG. Relationship between body condition and levels of serum luteinizing hormone in postpartum mares. Theriogenology 1987;28:815–25. 4. Gentry LR, Thompson Jr DL, Gentry Jr GT, Davis KA, Godke RA, Cartmmill JA. The relationship between body condition, leptin, and reproductive and hormonal characteristics of mares during the seasonal anovulatory period. J Anim Sci 2002;80:2695–703. 5. Vick MM, Sessions DR, Murphy BA, Kennedy EL, Reedy SE, Fitzgerald BP. Obesity is associated with altered metabolic and reproductive activity in the mare. Reprod Fertil Dev 2006;18:609–17. 6. Kubiak JR, Evans JW, Potter GD, Harms PG, Jenkins WL. Parturition in the multiparous mare fed to obesity. J Equine Vet Sci 1988;8:135–40. 7. Cavinder CA, Vogelsang MM, Forrest DW, Gibbs PG, Blanchard TL. Reproductive parameters of fat vs moderately conditioned mares following parturition. In: Proceedings 19th Equine Science Society Symposium, Tucson, AZ, 2005, pp. 65–9. 8. Tinker MK, White NA, Lessard P, Thatcher CD, Pelzer KD, Davis B, Carmel DK. Prospective study of equine colic risk factors. Equine Vet J 1997;29:454–8. 9. National Research Council (NRC). Nutrient Requirements of Horses. Washington, DC: National Academy Press, 2007.
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10. Kowalski J, Williams J, Hintz HF. Weight gains of mares during the last trimester of gestation. Equine Pract 1990; 12:6–10. 11. Lawrence LM, DiPietro J, Ewert K, Parrett D, Moser L, Powell D. Changes in body weight and condition of gestating mares. J Equine Vet Sci 1992;12:355–8. 12. van Niekerk FE, van Niekerk CH. The effect of dietary protein on reproduction in the mare. III. Growth of foals, body mass of mares and serum protein concentration of mares during the anovulatory, transitional and pregnant periods. J S Afr Vet Assoc 1997;68:81–5. 13. Banach MA, Evans JW. Effects of inadequate energy during gestation and lactation on the estrous cycle and conception rates of mares and on their foal weights. In: Proceedings of the 7th Equine Nutrition and Physiology Symposium, Warrenton, VA, 1981, pp. 97–100. 14. Maenpa PH, Koskinen T, Koskinen E. Serum profiles of vitamins A, E and D in mare and foals during different seasons. J Anim Sci 1988;66:1418–23. 15. Maenpa PH, Pirhonen A, Koskinen E. Vitamin A, E and D nutrition in mares and foals during the winter season: effects of feeding two different vitamin mineral concentrates. J Anim Sci 1988;66:1424–9. 16. Greiwe-Crandell K, Kronfeld DS, Gay LS, Sklan D, Tiegs W, Harris P. Vitamin A repletion in thoroughbred mares with retinylpalmitate of beta-carotene. J Anim Sci 1997; 75:2684–90. 17. Hoffman RM, Morgan KL, Lynch MP, Zinn SA, Faustman C, Harris PA. Dietary vitamin E supplemented in the periparturient period influences immunoglobulins in equine colostrum and passive transfer in foals. In: Proceedings of the 16th Equine Nutrition and Physiology Symposium, Raleigh, NC, 1999, pp. 96–7. 18. Estepa JC, Aguilera-Tejero E, Zafra R, Mayer-Valor R, Rodriguez M, Perez J. An unusual case of generalized softtissue mineralization in a suckling foal. Vet Pathol 2006;43: 64–7. 19. Gee EK, Firth EC, Morel PCH, Fennessy PF, Grace ND, Mogg TD. Articular/epiphyseal osteochondrosis in Thoroughbred foals at 5 months of age: influences of growth of foal and prenatal copper supplementation of the dam. N Z Vet J 2005;53:448–56. 20. Ytrehus B, Carlson CS, Ekman S. Etiology and pathogenesis of osteochondrosis. Vet Pathol 2007;44:429–48. 21. Knight DA, Weisbrode SE, Scmall LM, Reed SM, Gabel AA, Bramlage LM. The effects of copper supplementation on the prevalence of cartilage lesions in foals. Equine Vet J 1990;22:426–32. 22. Pearce SG, Grace ND, Firth EC, Wichtel JJ, Fennessy PF. Effect of copper supplementation on the copper status of pregnant mares and their neonates. Equine Vet J 1998; 30:200–3. 23. Pearce SG, Firth EC, Grace ND, Fennessy PF. Effect of copper supplementation on the evidence of developmental orthopaedic disease in pasture-fed New Zealand thoroughbreds. Equine Vet J 1998;30:211–18. 24. Kavazis AN, Kivipelto J, Ott EA. Supplementation of broodmares with copper, zinc, iron, manganese, cobalt, iodine and selenium. J Equine Vet Sci 2002;22:460– 4. 25. Van Weeren PR, Knaap J, Firth EC. Influence of liver copper status of mare and newborn foal on the development of osteochondrotic lesions. Equine Vet J 2003;35:67–71. 26. Gee EK, Firth EC, Morel PCH, Fennessy PF, Grace ND, Mogg TD. Enlargements of the distal third metacarpus and
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metatarsus in Thoroughbred foals at pasture from birth to 160 days of age. N Z Vet J 2005;53:438–47. Gee E, Davies M, Firth E, Jeffcott L, Fennessy P, Mogg T. Osteochondrosis and copper: Histology of articular cartilage from foals out of copper supplemented and nonsupplemented dams. Vet J 2007;173:109–17. Smith SR, Keene T. UK horse pasture evaluation program: a success story. In: Proceedings of the 62nd Southern Pasture and Forage Crop Improvement Conference, Knoxville, TN, 2008, pp. 44–9. National Research Council (NRC). Nutrient Requirements of Horses. Washington, DC: National Academy Press, 1989. Meyer H, Ahlswede L. The intra-uterine growth and body composition of foals and the nutrient requirements of pregnant mares. Anim Res Dev 1978;8:86–111. Guay KA, Brady HA, Allen VG, Pond KR, Wester DB, Janecka LA, Henigenger NL. Matua bromegrass hay for mares in gestation and lactation. J Anim Sci 2002;80:2960–6. Vivrette S. Parturition and postpartum complications. In: Robinson NE (ed.) Current Therapy in Equine Medicine. Philadelphia: W.B. Saunders, 1997; pp. 547–51. Dolente BA, Sullivan EK, Boston R, Johnston J. Mares admitted to a referral hospital for postpartum emergencies: 163 cases (1992–2002). J Vet Emerg Crit Care 2005;15:193–200. Pagan JD, Hintz HF, Rounsaville TR. The digestible energy requirements of lactating pony mares. J Anim Sci 1984;58:1382–7. Pagan JD, Hintz HF. Composition of milk from pony mares fed various levels of digestible energy. Cornell Vet 1986;76:139–48. Davison KE, Potter GD, Greene LW, Evans JW, MacMullan WC. Lactation and reproductive performance of mares fed added dietary fat during late gestation and lactation. J Equine Vet Sci 1991;11:111–15. Doreau M, Boulot S, Bauchart D, Barlet JP, MartinRosset W. Voluntary intake milk production and plasma
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metabolites in nursing mares fed two different diets. J Nutr 1992;122:992–1003. Hoffman RM, Kronfeld DS, Herblein HS, Swecker WS, Cooper WL, Harris PA. Dietary carbohydrates and fat influence milk composition and fatty acid profile. J Nutr 1998;128:2708s–71s. Martin RG, McMenimen NP, Dowsett KF. Effects of a protein deficient diet and urea supplementation on lactating mares. J Reprod Fertil 1991;44:543–50. Ott EA, Asquith RL. Calcium and phosphorus supplementation of foaling mares. In: Proceedings of the 8th Equine Nutrition and Physiology Symposium, Lexington, KY, 1983, pp. 317–22. Glade MJ. Effects of gestation, lactation and maternal calcium intake on mechanical strength of equine bone. J Am Coll Nutr 1993;12:372–7. Cassill B. Response of bone mineral markers to lactation, gestation and calcium supplementation in broodmares. MS Thesis, University of Kentucky, Lexington, 2007. Asai Y, Katsuki R, Matsui A, Nanbo Y. Effects of rations and age on mineral concentrations of thoroughbred mare’s colostrum. J Equine Sci 1995;6:21–4. Janicki K. The effect of dietary selenium source and level on broodmares and their foals. MS thesis, University of Kentucky, Lexington, KY, 2001. Silver M, Fowden AL. Uterine prostaglandin F metabolite production in relation to glucose availability in late pregnancy and a possible influence of diet on time of delivery in the mare. J Reprod Fertil Suppl 1984;32:511–19. Fowden AL, Forhead AJ, White KL, Taylor PM. Equine uteroplacetal metabolism at mid and late gestation. Exp Phys 2000;85:539–45. Henneke DR, Potter GD, Kreider JL, Yates BF. Relationship between condition score, physical measurements and body fat percentage in mares. Equine Vet J 1983;15: 371–2.
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292.
Stereotypic Behavior Paul McGreevy
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Chapter 292
Stereotypic Behavior Paul McGreevy
Before focusing on the specifics of how stereotypies manifest in mares and, indeed, in all horses, we should acknowledge the basics of equine ethology. In the wild, horses live in cohesive social groups on the plains. They occupy extensive home ranges, relying on conspecifics for companionship, mutual comfort, and enhanced detection of food. Apart from the dispersal of young males, they avoid becoming isolated. Their long noses allow them to graze while maintaining surveillance for predators, but they generally do not defend their territory. Instead, they rely largely on speed and agility to escape from danger. Unfortunately, housing and stable management tend to overlook the horse’s need to socialize with conspecifics, and social isolation has been identified as an important factor in the emergence of stallwalking and weaving. All such stereotypies are defined as being repetitive, invariant, apparently functionless behaviors. They are believed to emerge in individuals that have difficulty adjusting to particular management styles that align poorly with the ethology of the horse. A so-called ‘trickle feeder’, the horse has only a small stomach but its large cecum is able to digest material that ruminants generally overlook. It must forage for long periods rather than eating and then ruminating, so a period of fasting is a more profound insult to equids than to many other herbivore species. Feeding regimens for stabled horses are designed for the convenience of human carers and often constitute a radical shift from the horse’s natural foraging habits. Such lifestyle shifts are critical factors in the ontogeny of common unwelcome behaviors in intensively managed horses. For example, discrete meals that allow gastric acidity to build up have been implicated in the development of crib-biting (and windsucking), the most common equine oral stereotypies.
Crib-biting (cribbing) The distinction between crib-biting and windsucking is often blurred because definitions differ in several English-speaking countries. (Windsucking must be declared at auction in the UK and tends to lower the value of affected animals.) In the UK, windsuckers are distinguished from crib-biters because they do not hold onto fixed objects while engulfing air, whereas in Australia windsucking includes grasping fixed objects and engulfing air. The characteristic arching of the neck, which allows horses to engulf and hold air in the cranial esophagus, is a critical feature common to both stereotypies. Although horses prefer to grasp wood rather than metal (perhaps because it cushions their teeth), they will crib-bite on metal if no other substrate is available. It is not clear why, in identical situations, some horses start crib-biting and others do not. Feeding discrete concentrated meals that require little mastication has a greater effect than housing practices on the incidence of crib-biting, but it is rarely possible to identify a single cause. Typically, if a horse performs one oral stereotypic behavior, it is more likely to develop a second. Some ethologists maintain that stereotypies are a response, albeit an imperfect one, to the insults of intensive management and may provide some stress relief for horses that perform them. It is possible that the release of endorphins reinforces crib-biting. Opioid antagonists (such as naloxone) can reduce crib-biting significantly, which suggests that, from the horse’s perspective, at least one of the perceived benefits of crib-biting is mediated by opioids. However, resting behavior in crib-biters also significantly increases with opioid antagonists, so the reduction in crib-biting may occur because of generalized sedation. The prevalence of windsucking/crib-biting in UK Thoroughbred populations is greater than in other
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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breeds and has been estimated at 4.13% (± 2.57%).1 This might simply reflect the fact that Thoroughbreds, among the most reactive of breeds, are generally raced and are, therefore, managed very intensively. The role of breed in the emergence of all stereotypies will become clearer in the light of future genome research. Wood-chewing is often confused with crib-biting by novice observers, but it is not really a stereotypic behavior as it is not invariant and is seen in freeranging horses, so it may well have a function in digestion or nutrition.
The function of stereotypic behaviors How stereotypic behaviors work for the horse is still being unraveled.2 They seem to help animals cope with stress (e.g., bouts of crib-biting produce transient decreases in heart rates and measures to prevent crib-biting have been linked to increases in some physiological stress parameters). However, Minero et al.3 suggest that crib-biters may have higher basal sympathetic activity because they have a higher overall mean heart rate. Stereotypic horses have been reported to persist in non-rewarding activities longer than control horses.4, 5 It is also possible that there is a direct link between a given stereotypic behavior and the motivational system from which it is derived. Feeding concentrates reduces saliva production and increases gastric acidity. Similarly, periods without food precipitate yet more gastric acidity and the risk of gastric ulceration. So, within environments that limit normal forage intake (e.g., intensive training programs in which the diet has a high concentrate-to-roughage ratio) crib-biting may provide a route for the horse to normalize its digestive activity. Crib-biting and windsucking have also been associated with concurrent disorders of the digestive system (e.g., there is a significant association between stomach ulceration and crib-biting in foals). A study on a population of foals by Waters et al.6 showed a peak in the emergence of crib-biting at 20 weeks of age, which coincided with a significant increase of concentrates in the diet. Foals and adults have been shown to crib-bite less when fed antacid supplements. So, treatments and management styles that normalize ingestive behavior will improve the welfare of all stabled horses and reduce the incidence and severity of oral stereotypies. Despite there being no proven indications that horses can learn by observation, there is a strong traditional belief among horse people that horses develop stereotypies by mimicking neighbors. Such social influences, also known in bank voles,7 may
arouse observing horses and so predispose them to developing stereotypic behavior, but there is no hard evidence. Since crib-biting is popularly regarded as being transmissible in this way and is associated with health problems, horses exhibiting it are often isolated (which seems counterproductive given the horse’s strong need to socialize with conspecifics). Some owners report few or no ill-effects from cribbiting, while others blame it for diminished performance, digestive disorders, incisor erosion, and failure to thrive. Some veterinarians maintain that cribbiting leads to chronic colic, flatulent colic, and a distended abdomen, and this viewpoint is supported by epidemiological evidence that epiploic entrapment is associated with a history of cribbing/windsucking.8 However, radiographic studies suggest that negligible gas is ingested during crib-biting.9 It is possible that crib-biters have concurrent gastrointestinal disturbances that cause signs of aerophagia.
Control measures For aesthetic and occasional health reasons, the search continues for ways to prevent stereotypies and to effect permanent cures. Physical or surgical approaches to prevent crib-biting have had mixed success, and a humane remedy that is effective for every crib-biter remains elusive. As the role of gastric acidity in the ontogeny of oral stereotypic behaviors becomes apparent, surgery is falling from favor. Continual reinforcement of stereotypic behaviors must be a major factor in their resistance to therapy. Increasing prevalence as horses age suggests that crib-biting can become emancipated from its initiating causes. So while stereotypies may arise in response to adverse management, they may persist even in more enriched environments. Both crib-biting and windsucking can be discouraged by the application of a collar that makes the characteristic flexion of the horse’s neck uncomfortable. A typical anti-cribbing collar is a simple leather strap that is tightened until neck-flexing and/or engulfing air into the esophagus are possible only with significant discomfort. It incorporates a galvanized, hinged arc that accommodates the trachea and is intended to allow normal breathing. A constriction at this site not only makes crib-biting uncomfortable but may also make the terminal grunting in this oral-based stereotypy less easy to achieve, since this involves distension of the cranial esophagus. Some models of the collar include leather or metal spikes that increase the discomfort associated with neck flexion, while others are made entirely of leather and are often marketed as being more ‘humane’ (even though they work along the same
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Stereotypic Behavior principles). The collars can be tightened as horses adapt to the constriction, which occasionally causes skin trauma. Combinations of straps that can be tightened around different parts of the cranial neck would possibly reduce tissue damage at the poll. Another option is an electronic training collar that can be programmed to discharge when the neck is flexed, or remotely controlled so that the horse does not associate punishment with the presence of a human. The use of a dummy collar before this sort of therapy might avoid the horse associating punishment with the collar itself. Tests have shown that if an electric shock is applied after the horse has grasped but before it has engulfed air, the grasping phase can be punished and extinguished. However, follow-up studies in 60 cases of aversion therapy using electric shock collars reported that only nine ‘cures’ were effected and, of these, three required further shocks after 9 months.10 This method entails constant observation by the trainer and considerable patience to maintain the contingency between crib-biting and punishment as the first shock may discourage the horse for some time. Physical prevention or punishment may also increase distress, which is a more profound ethical problem.
Weaving The lateral swaying of the head is known as weaving. The rest of the body (notably the shoulders) may also sway, and the front, and sometimes the hind, legs may be picked up. Weaving is a common ethopathy in horses and is also recognized in exotic species, notably elephants and bears. In horses, it usually occurs over the stable door or some other barrier. Like other stereotypic behaviors, weaving may mature over time (e.g., it may change from a stationary head-swing to a side-to-side pacing). Individual horses may perform more than one stereotypic behavior (e.g., a weaver may also be seen stall-walking on occasions), but locomotory stereotypies are less likely to be associated with oral behaviors in the same individual. Weaving is more commonly reported in Thoroughbreds (3.25 ± 3.23%)1 than in other breeds, but it is not clear if this reflects their breeding or environment, since gallopers are managed more intensively than other equine athletes. The appearance of weaving roughly coincides with the time when younger horses are sold from the stud and go to new homes, at a median age of 60 weeks. This suggests that it is related less to weaning and more to environmental and social disturbances at the time of dispersal. Along with other locomotory stereotypies, it is commonly seen in stabled horses before feeding or
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other arousing events, such as the arrival or departure of conspecifics. It seems likely that weavers have a thwarted motivation to move (e.g., to be with other members of the group or to reach food) rather than that they are ‘bored’ because of under-stimulation. Weaving could be either the expression of an unusual form of species-typical anticipatory behavior or a learned response to a desirable outcome, such as leaving confinement. Weaving is observed at pasture only when horses encounter a barrier, and is reported as being both more and less prevalent in standing stalls than in looseboxes. If being in a stable is frustrating for a horse, it is possible that lack of free exercise plays a role in the motivation to weave. However, exercise routine has not been identified as a risk factor in epidemiological studies, and the relationship between exercise routine and stereotypic behavior is unclear. There is little evidence that exercise has a consistent effect on the incidence of stereotypic behavior in the stable, but stimuli that signify turning out (to pasture) often trigger episodes of locomotory stereotypic behaviors. Close tactile and visual contact with neighboring horses reduces weaving, so stables and looseboxes should be partitioned in ways that facilitate socializing. Cooper et al.11 reported that weaving was not observed when horses could socially interact with their neighbors on all four sides of their enclosure. This adds weight to the suggestion that weaving can be regarded as a frustrated escape response. Anti-weaving bars – V-shaped apertures through which horses can put their heads and necks while limiting lateral movement – are the most common prophylactic for or response to weaving. They are now found in some 70% of UK establishments. Other options include fixing an upright bar or suspending a heavy object to occupy the space above the door, or completely preventing the horse from putting its head over the stable door. Ideally, this prevents or interrupts the weaving, but most weavers simply move to an area within the box and weave without having their head and neck over the door. Such physical impediments to stereotyped locomotion probably just increase the affected horse’s frustration with the environment. Exercise that tires the horse and prompts it to rest might help to reduce time spent weaving. Another practical idea that seems to work well is to provide stabled horses with mirrors. In both short-term and long-term studies, mirrors have a similar effect to social contact, although it is unclear whether horses are simply distracted by reflected visual stimuli or whether they regard the image as another horse. The role of learning in the emergence of stereotypies is significant, so it is worth altering certain
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stimuli that reliably precipitate episodes of weaving. Modifying the cues that precede feeding or feeding times may reduce pre-feeding weaving but not postfeeding stereotypic behavior. So, while changing the husbandry routines may be effective, it is unlikely to be permanent since affected horses will most likely quickly learn new associations that predict feeding in novel routines. Changes would have to be continually made to maintain their novelty value and avoid the behavior returning to its original level. Crucially, however, flux in routine is considered harmful to horses, not least in terms of gastrointestinal health, so this option must be approached with caution. When it first emerges, weaving may be seen as a sign of frustration and possibly an indicator of compromised welfare, but this is not necessarily so. Many authors point out the importance of the emancipation process in liberating stereotypic behaviors from their initiating causes, and argue that mature stereotypies may have lost any putative role in ameliorating distress. This question becomes contentious because we do not understand the neural mechanisms that might reinforce weaving. To add to the confusion, opioid antagonists have been shown both to decrease and increase weaving, so the role of endorphins as an intrinsic reward for weaving remains uncertain. Management systems that facilitate social housing and reduce the emergence of weaving may well be the best option we currently have. They are becoming more popular, despite the admitted drawback that they also allow undesirable social interactions (e.g., biting and bite threats) and the spread of contagions. Welfare concerns in weavers relate to the effects of excessive wear and tear on the hooves and the musculoskeletal system. Weavers may also show altered time budgeting that depletes resting and feeding opportunities, which may result in a failure to thrive. This is mainly why weaving is an unwelcome behavior that must be reported at auction in some countries, including the UK and Australia.
Stall-walking (box-walking) Pacing a fixed route (route tracing) around the stable is known as stall-walking in the USA (or box-walking in the UK). After weaving, stall-walking is the most common locomotory stereotypic behavior in horses. The route traced is typically circular but, in larger stables or even at pasture, horses may trace a ‘figure of eight’. The problem may start when the horse is separated from conspecifics, so horses in sale yards are more likely to be detected performing this behavior. Clearly, it is not a behavior that is possible in standing stalls (tie stalls, US) or when a horse is in cross-ties. A track in the bedding of a stable may be the first evi-
dence of this activity. Occasionally, stall-walkers may trot around their chosen route and will sweat and become exhausted as a result of the stereotypy. Stallwalking is reported in 2.2 ± 2.13% –of stabled horses.1 It must be declared at certain types of auction and tends to lower the value of affected animals. Although it may seem like a harmless way of filling periods of inactivity, route tracing can affect the horse’s welfare by reducing the time available for maintenance behaviors, such as eating and resting. Blocking the path with obstacles and distracters to prevent this activity is virtually fruitless, but radical changes in the enclosure and instituting a new time budget may disturb the pattern shown and the frequency of stereotypic episodes. It is not certain that the incomplete disappearance of the stereotypy constitutes evidence for so-called emancipation (the liberation of stereotypic behaviors from their initiating causes). Some argue that the resistance of route tracing to change means that the behavior can become less relevant to the animal’s immediate needs than it was when it first emerged. Nevertheless, the (albeit incompletely) reduced frequency and intensity of the stereotypy in environments that have been, however transiently, enriched, indicates that its function has become less critical. Weaving and stall-walking are considered similar in cause and both reflect a strong association with social needs. Stall-walkers often vocalize, which suggests that it is a form of barrier frustration. Interestingly, stall-walking is reported with a higher-thanexpected frequency in horses used for endurance trials. It is possible that persistence in apparently redundant locomotory activity reflects a shift in the endorphin activity (and so perhaps the pain threshold) of horses being trained for stamina and resistance to dehydration. Welfare concerns about stall-walking relate to the effect it has on the horse’s ability to maintain bodyweight. If the horse walks in only one direction, there may be asymmetric hoof or shoe wear and sometimes lateralized atrophy and hypertrophy of the lumbar musculature. That said, stall-walkers are less likely to be isolated as a remedial measure than horses with other stereotypies, such as crib-biting, because the emergence of stall-walking is not generally attributed to observational learning.
Conclusions Ongoing research efforts are likely to identify specific risk factors for all equine stereotypies. Meanwhile, mares that develop stereotypies can be regarded as useful sentinels of suboptimal husbandry, sentinels that alert us to problems encountered by all resident
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Stereotypic Behavior horses, not just those that have produced a stereotypic response to challenges. Improving stable management is a better option than selecting more adaptable horses.
References 1. Nicol CJ. Stereotypies and their relation to management. In: Harris PA, Gomarsall GM, Davidson HPB, Green RE (eds) Proceedings of the BEVA Specialist Days on Behaviour and Nutrition, 1999, pp. 11–14. 2. Mason GJ. Stereotypies: a critical review. Anim Behav 1991;41:1015–37. ¨ 3. Minero M, Canali E, Ferrante V, Verga M, Odberg FO. Heart rate and behavioural responses of crib-biting horses to two acute stressors. Vet Rec 1999;145:430–3. 4. Hausberger M, Muller C, Gautier E, Jego P. Lower learning abilities in stereotypic horses. Appl Anim Behav Sci 2007; 107:299–306. 5. Hemmings A, McBride SD, Hale CE. Perseverative responding and the aetiology of equine oral stereotypy. Appl Anim Behav Sci 2007;104:143–50.
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6. Waters AJ, Nicol CJ, French NP. Factors influencing the development of stereotypic and redirected behaviours in young horses: findings of a four-year prospective epidemiological study. Equine Vet J 2002;34:572–9. 7. Cooper JJ, Nicol CJ. Neighbour effects on the development of locomotor stereotypies in bank voles Clethrionomys glareolus. Anim Behav 1994;47:214–16. 8. Archer DC, Proudman CJ, Pinchbeck G, Smith JE, French NP, Edwards GB. Entrapment of the small intestine in the epiploic foramen in horses: a retrospective analysis of 71 cases recorded between 1991 and 2001. Vet Rec 2004;155:793–7. 9. McGreevy PD, Richardson JD, Nicol CJ, Lane JG. A radiographic and endoscopic study of horses performing an oral stereotypy. Equine Vet J 1995;27:92–5. 10. . Owen RR. Crib-biting and wind-sucking – that equine enigma. In: Hill CSG, Grunsell FWG (eds) The Veterinary Annual 1982. Bristol, UK: Wright Scientific Publications, 1982; pp. 156–68. 11. Cooper JJ, McDonald L, Mills DS. The effect of increasing visual horizons on stereotypic weaving: implications for the social housing of stabled horses. Appl Anim Behav Sci 2000;69:67–83.
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293.
Reproductive Efficiency Laura C. Nath
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Pituitary Pars Intermedia Dysfunction (Equine Cushing’s Syndrome) Philip J. Johnson, Nat T. Messer and Venkataseshu K. Ganjam
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Epidemiology Applied to Horse Reproduction Nigel R. Perkins
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Management of the Geriatric Mare Scott Madill
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Parentage Testing Ernest Bailey
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Relevance of Genomics to Equine Reproduction Ernest Bailey
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Chapter 293
Reproductive Efficiency Laura C. Nath
Introduction The objective of any breeding program should be the birth of a healthy foal. Reproductive efficiency reflects the management of stallions and mares to maximize the number of foals born. Intensity of management is influenced by the financial costs of intervention. The benefits of improved reproductive efficiency are most clearly appreciated on large studfarms producing valuable foals. However, the same principles should be applied when breeding each individual mare. Horses have traditionally been regarded as having poor reproductive capacity in comparison to other species.1 Selection of breeding stock based on pedigree, conformation, and performance rather than fertility has further impeded reproductive success. Artificial imposition of a limited breeding season, often at a time when mares are not cycling naturally, also impacts on fertility. The widespread use of ultrasonography and improved pharmacological control of the mares’ cycle has allowed for vast improvements in reproductive efficiency in the last 25 years.2 Ultrasonography aids precision in determining timing of service in relation to ovulation, the best time for administration of ovulation-inducing drugs and allows confirmation of ovulation, thus minimizing the number of services per cycle and per pregnancy. Furthermore, ultrasonography assists in accurate early pregnancy diagnosis at 13–16 days post-ovulation. This initial examination presents an opportunity to assess the pregnancy for defects such as growth retardation, diagnosis and intervention in multiple pregnancy, detection of uterine abnormalities such as edema or fluid accumulation, and assessment of the size and morphology of the corpus luteum. In the nonpregnant mare, examination at 13–16 days may provide an insight into the reason that pregnancy was not established, such as endometrial inflammation and premature luteolysis.
Subsequent examinations in pregnant mares should be conducted at regular intervals to allow early diagnosis of embryonic loss or failure of normal embryonic development. Confirmation of pregnancy for the purpose of certificates or insurance should be performed after 42 days, as the rate of fetal loss is typically less than early embryonic loss.3 There are many parameters by which efficiency of a breeding program may be measured (see Tables 293.1 and 293.2). Pregnancy rate per cycle and services per pregnancy are the most useful measures of reproductive efficiency in commercial operations.4 It is prudent to consider that any discussion of reproductive efficiency is based on an understanding of the principles of probability. A discussion of the difficulties in interpreting fertility data has recently been provided by Amann.5 Briefly, whilst fertility data for a given stallion or semen sample may be available, poor scientific methods or low numbers of mares bred may render the results less valuable. For this reason, reported fertility data should be interpreted with skepticism, and with due consideration of the manner through which the data was derived.5 Pregnancy rates per cycle are more valuable than per season pregnancy rates in providing information about an individual stallion’s fertility.5 When seasonal pregnancy rates are reported, they do not take into account the number of times or cycles a mare was bred to achieve a pregnancy. Therefore, seasonal pregnancy rates may mask subfertility. For example, if a stallion has a pregnancy rate per cycle of 70% then after two mare cycles ∼ 91% of mares are expected pregnant. However, a stallion with a pregnancy rate of 40% per cycle, if allowed to breed for five cycles, will achieve the same overall (end of season) pregnancy rate of 92% (see Table 293.3).6 First cycle pregnancy rates are typically higher than overall pregnancy rates. This is because a large proportion of the most fertile mares will become pregnant after mating on a single estrus. Only mares that did not become pregnant on the first cycle will
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 293.1 Definition of terms used to measure reproductive efficiency
Table 293.2 Definition of terms used to describe mare status Maiden
Fertilization rate Conception rate
Pregnancy rate Pregnancy rate per cycle
Percentage of ovulated oocytes fertilized by a spermatazoon
Mare that has never produced a foal
Lactating
Mare with foal at foot
Foaling
Synonymous with lactating
Wet
Synonymous with lactating
Barren
Percentage of mated mares that are pregnant on a specified day
Mare bred in a previous season that was not diagnosed pregnant
Dry
Percentage of mated mares that are pregnant on a specified day post ovulation
Mare without foal at foot, includes maiden and barren mares and mares that were not bred in the previous season
Slipped
Mare diagnosed pregnant in the previous season that did not produce a live foal
Synonymous with fertilization rate and often used erroneously to describe pregnancy rate
First cycle pregnancy rate
Percentage of mares that are pregnant after the first mated estrus
End of season pregnancy rate
Percentage of mated mares that are pregnant at the end of the breeding season
agistment costs may be achieved through improvements in reproductive efficiency.
Foaling rate
Percentage of mated mares that produce a live foal
Observed reproductive efficiency
Embryonic loss rate
Percentage of fertilized oocytes that do not survive to 40 days post ovulation
Early embryonic death rate
Synonymous with embryonic loss rate
Fetal loss rate
Percentage of pregnancies surviving to 40 days that do not result in a live foal
Stillbirth rate
Percentage of pregnancies surviving to 300 days that do not result in a live foal
Abortion rate
General term regarding pregnancy loss that needs to be defined in relation to stage of gestation
Services per cycle
Number of times a mare is served in a given estrus period
Services per pregnancy
Number of times a mare is served prior to diagnosis of resulting pregnancy
Cycles per pregnancy
Number of served estrus cycles prior to the mare becoming pregnant in one season
Source: adapted from Ginther (1993).1
be mated on the second cycle. Therefore the pool of mares may be considered slightly less fertile at each subsequent cycle, as they require more mated cycles to achieve pregnancy. To put this discussion into the context of a breeding program, it is clear that fertility is closely related to reproductive efficiency. In any breeding program the desired outcome should be a healthy foal. Improvements in fertility will allow for the production of foals born earlier in the season, thus potentially increasing their sale value. Furthermore, considerable economic benefits in relation to veterinary, staff, and
Reproductive efficiency has improved over the last three decades due to improvements in stallion and mare management and veterinary research and intervention.7 The United Kingdom has observed an overall improvement in 15-day pregnancy rate per cycle from 52% in 19828 to 60% in 19982 and 63–65% in 2002.9 Similar per cycle pregnancy rates of 63–65% have been achieved in recent years in North America.10, 11 Improvements in reproductive efficiency in Australia are reflected in the studbook data depicting an increase in the percentage of live foals per mare covered, from 40% in 1981 to 76% in 2000.6 Pregnancy rates per cycle in Australia and New Zealand ranged from 55% in the early 1980s12 to up to 70% more recently.6,13–15 Pregnancy rates per cycle of 76–83% can be achieved after artificial insemination of fresh semen.16, 17 Reports for the use of cooled semen range from 44% to 65% per cycle.18–20 Reports for the use of frozen semen range from 32% to 73% per cycle.19–24 Table 293.3 Examples of pregnancy rates achieved by stallions for mare cycles 1 to 7 Pregnancy rate per cycle
70%
60%
50%
40%
30%
20%
10%
Cycle 1 2 3 4 5 6 7
70 91 97.3 99.1 99.7 99.9 99.9
60 84 93.6 97.4 98.9 99.5 99.8
50 75 87.5 93.7 96.8 98.4 99.2
40 64 78.4 87.0 92.2 95.3 97.2
30 51 65.7 75.9 83.1 88.2 91.7
20 36 48.8 59.0 67.2 73.7 79.0
10 19 27.1 34.3 40.9 46.8 52.1
Source: McKinnon (2000).6
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Reproductive Efficiency The above studies have identified common factors that can strongly influence breeding efficiency outcomes. These include mare age and reproductive status, veterinary management including method of service, timing of service, treatments, reproductive losses, twinning, and the stallion. Mare age Increasing mare age is negatively associated with reproductive efficiency.2,8–13,15, 25 Maintaining highly valuable aged mares in the breeding population likely contributes to the perceived poor reproductive capacity of horses. In the overall population of Thoroughbred mares in the United Kingdom, Weatherbys figures suggest a linear decline with mare age in the percentage of live foals produced.26 Lower per cycle pregnancy rates and higher pregnancy loss rates contribute to reduced reproductive efficiency in older mares.2,9–12 The major causes for reduced fertility in older mares include poor conformation,27 endometrial change,28, 29 susceptibility to fluid accumulation and endometritis,30 and poor viability of the embryo.31 Mare reproductive status The influence of mare reproductive status on pregnancy rates is largely influenced by management. It is expected that the barren mare population includes a large proportion of problem mares, and therefore lower pregnancy rates per cycle are observed.11 Other studies have identified good seasonal pregnancy rates in barren mares, potentially because breeding commences earlier in the season, allowing the mares to be served on more cycles. Under good management conditions, per cycle pregnancy rates are often highest in maiden mares, and first cycle pregnancy rates of up to 76% have been reported.10, 13 In contrast, Hanlon15 reported reduced per cycle pregnancy rates in maiden Thoroughbred mares (47.8%) compared to barren (53.2%) and foaling (56.1%) mares, but the highest end of season pregnancy rate (81.7%) was achieved in maiden mares, due to increased number of cycles bred, and fewer pregnancy losses. The Thoroughbred and Standardbred industries have a large proportion of young maiden mares incorporated into breeding programs.2, 9, 13 However, other industries, such as sporthorses, Quarter Horses and Arabians often see a large number of older maiden mares entering breeding programs after the end of their career. It has long been recognized that older maiden mares can have uterine fluid accumulations due to failure of the cervix to dilate in estrus. Reduced pregnancy rates utilizing frozen semen have been observed in maiden mares as compared to barren or foaling mares.20, 32
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In these studies, reduced pregnancy rates in maidens were attributed to a large proportion of older maiden mares in this group. Foaling mares would be expected to have good pregnancy rates per cycle, given a history of a normal foaling.20 Equivalent per cycle pregnancy rates in foaling and maiden mares have consistently been reported.2, 9, 13 However, foaling mares are often bred later in the season and the number of cycles on which these mares can be bred may be few, resulting in lower seasonal pregnancy rates. The decision to breed on foal heat is contentious, and will depend upon recent foaling history, the age and parity of the mare, presence of uterine fluid, and number of days between foaling and ovulation.33 Importantly, increased pregnancy losses have been observed in mares conceiving on foal heat.2, 9 In well-managed Thoroughbred mares, Morris and Allen2 reported reduced first cycle pregnancy rates for mares bred on foal heat (57.6%; 121/220) compared to the second estrus after foaling (65.9%; 369/560). However, a study in young, fertile mares did not demonstrate a difference in pregnancy rates in mares bred on foal heat (72%; 288/399) compared to later cycles (76%; 120/158).16 Management Without doubt, improvements in management, including increased veterinary intervention, accurate timing of service relative to ovulation, and improved diagnostics with increased uterine and systemic treatments have contributed to the overall increase in pregnancy rates seen over the last 25 years. The intensity of management in a breeding program is largely dictated by economics, with more valuable horses generally being treated more intensively, and achieving higher pregnancy rates.6, 9, 13 McKinnon reported on a population of Thoroughbred mares achieving a pregnancy rate per cycle of 70.4% (n = 2436 cycles).6 In the same study, the population of Standardbred mares achieved a pregnancy rate per cycle of only 62.0% (n = 2086 cycles). Whilst both populations of mares were serviced by the same veterinarians, significantly fewer veterinary examinations were performed on the Standardbred farms.6 More recently, higher pregnancy rates were observed on farms with more intensive management, with well-managed Standardbred farms achieving equivalent pregnancy rates to Thoroughbreds.13 Method of service Higher pregnancy rates can be achieved in mares served by artificial insemination of fresh semen (83% per cycle) compared with natural service (66%).16
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On-farm insemination of Standardbred mares with fresh semen achieved a first cycle pregnancy rate of 70%, which was not different to pregnancy rates achieved for Thoroughbreds (71%).13 Jasko34 observed pregnancy rates to be highest after insemination of fresh semen, and cooled semen achieved higher pregnancy rates than frozen semen. Squires et al.20 also observed highest pregnancy rates in mares served with fresh semen (60%, n = 68); however, pregnancy rates in mares served with cooled (44%, n = 272) and frozen (46%, n = 536) semen were equivalent. Loomis19 evaluated the transported semen industry in North America and reported first cycle pregnancy rates of 59.4% (n = 850) for cooled semen and 51.3% (n = 876) for frozen semen. These investigations suggest that reproductive efficiency is likely to be highest utilizing on-farm artificial insemination of fresh semen. The advantages of artificial insemination of fresh semen include minimal contamination of the reproductive tract, and the ability to assess and control the number and concentration of spermatozoa and the volume of semen inseminated. Seminal plasma is typically retained in the processing of fresh samples to assist uterine defence mechanisms and fluid evacuation.35 The addition of antimicrobials may confer further advantages.35 One advantage of natural service is that the stallion can open the cervix at the time of breeding, which may enhance uterine fluid evacuation. Currently, frozen semen carries the lowest per cycle pregnancy rates. Lower pregnancy rates from cooled and frozen semen are likely due to damage to sperm membranes, variability in the quality of semen samples, reduced longevity of semen, and reduction or omission of seminal plasma.19, 36 Furthermore, not all stallions will yield ejaculates suitable for processing cooled or frozen semen. Sample collection and processing require an initial outlay in cost; however, this may be offset by reduced expenses in mare transport and agistment. Currently, chilled semen shipments occur every other day between Australia and New Zealand during the breeding season in the southern hemisphere, and international frozen semen networks are well established. Frozen semen offers the considerable advantage of collection for semen preservation outside of the normal season, and much reduced transport costs for multiple doses. This may be particularly useful when stallions require time away for competitions, and in practices which have strained resources during the breeding season. The variability in quality of processed semen samples presents the greatest challenge for improved reproductive efficiency with artificial insemination. Improved processing techniques and superior quality control will encourage more widespread use of artificial insemination.
Timing of service Higher pregnancy rates are achieved when ovulation occurs close to the time of breeding.37–42 The advent of ultrasonography and development of agents to reliably induce ovulation within a narrow window, namely, human chorionic gonadotropin (hCG) and gonadotropin-releasing hormone agonists (GnRH, e.g. deslorelin), has allowed for more precise timing of service in relation to ovulation. The end result is a reduction in the number of services per cycle and conservation of stallion sperm reserves. McKinnon reported that, within a population of Thoroughbred mares, those that received ovulationinducing drugs had improved per cycle pregnancy rates (deslorelin 72%, n = 204; hCG 72%, n = 1447) versus those not treated (64.5%, n = 817).6 This is likely due to ovulation close to the time of service, but may also reflect higher-intensity management on those farms willing to use ovulating agents. This study also detected a higher incidence of twin pregnancy in mares treated with deslorelin (19.7% per pregnancy) versus those not treated (11.8% per pregnancy) (P < 0.05). Use of ovulation induction agents likely increases the chances of fertilizing multiple oocytes and enhances the veterinarian’s ability to detect multiple pregnancies by decreasing the time between ovulations. Provided multiple pregnancies can be manually reduced properly by experienced personnel, this should improve pregnancy rates.
Number of services Breeding mares on multiple occasions on a cycle (i.e., 48 hours apart) in order to achieve insemination close to the time of ovulation is not detrimental in normal mares.9, 11, 13, 20, 43, 44 Breeding more than once per cycle may reflect a willingness to confirm ovulation and to repeat breed mares which fail to ovulate within 48–72 hours of the last service. In less intensive management systems, mares are served according to teasing patterns, and therefore may be served every other day or daily until they reject the stallion. The highest pregnancy rates in Thoroughbred mares have been reported from mares bred on only one occasion.13 However, the same authors reported that Standardbred mares bred on two occasions showed higher pregnancy rates compared to Standardbred mares bred once. This likely reflects the quality of management on the farms. In Thoroughbreds, good management, including use of ovulating agents, will allow for a single service per cycle, thus conserving stallion sperm reserves. Services per cycle in Thoroughbred mares in this study were 1.03. In contrast, on Standardbred farms utilizing artificial insemination, a wider availability of semen will allow for multiple
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Reproductive Efficiency services per cycle without jeopardizing sperm numbers. Services per cycle in Standardbred mares in this study were 1.43. The disadvantage of breeding on multiple occasions is that each service presents a challenge to the uterus in clearing fluid accumulations.45 It has been demonstrated that, with fresh semen insemination, it takes 24–36 hours for post-breeding endometritis to subside.46 Therefore it is strongly recommended that problem mares are served once, close to the time of ovulation. An exception may be when a stallion has demonstrated subfertility. In those cases, some individuals that are allowed to breed multiply per cycle may provide more viable semen presented to the oviductal epithelium for binding or availability to fertilize. When a stallion has many mares to breed, the option to breed multiply on a cycle is extremely limited. A practical approach taken when breeding either naturally or by artificial insemination with fresh or cooled semen is to breed at the time when the dominant follicle will respond to ovulation-inducing agents. One of these agents (deslorelin or HCG) is then administered immediately after breeding. Programming the mare at the time of breeding, rather than before, provides greater flexibility in managing the service. First, in some circumstances, service may need to be delayed in order to facilitate a more urgent service to another mare. Second, occasionally there may be problems with the service, or the quality of the insemination is unexpectedly poor or delayed in arriving. In those cases, if the mare is not programmed to ovulate, she may be rescheduled for a repeat service or insemination at a later stage. This also highlights the importance of examining all artificially inseminated samples for viability at the time of service. After correct administration of ovulating drugs, ovulation should occur within 36 ± 4 hours with hCG or 41 ± 3 hours with deslorelin.47 Mares should undergo an ultrasonographic examination the day following service to determine the extent of postmating endometritis. If present, mares may be treated daily until ovulation and for 3 days after ovulation as deemed necessary.48
Treatments Intrauterine fluid accumulation adversely affects pregnancy rates.16, 30, 49, 50 Systemic and intrauterine treatments directed at reducing inflammation and microbial contamination of the uterus will improve pregnancy rates in mares with poor reproductive capacity.48 However, studies looking at on-farm management seldom demonstrate improved pregnancy rates with intrauterine treatments, largely because only those mares with problems are likely to receive
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such treatments.6, 11 Recently, high pregnancy rates were achieved in mares treated with intrauterine antibiotics and oxytocin (68.4% pregnancy rate at day 15).9 In the same study, mares treated with intrauterine antibiotics and uterine lavage had the lowest pregnancy rates (41.4% pregnancy rate at day 15) and highest embryo loss rates (33.3% embryonic loss).9 These poor results likely reflect severe underlying uterine pathology in those mares requiring uterine lavage in combination with antibiotics; without treatment, pregnancy rates may have been lower and embryonic loss rates higher. In a group of healthy mares, improved pregnancy rates were detected in mares given a post-breeding treatment with oxytocin plus intrauterine antibiotics compared to those mares that were not treated, treated with antibiotics alone, or treated with oxytocin alone.49 Furthermore, mares that were treated with antibiotics, oxytocin or a combination of the two had higher pregnancy rates than those mares that were not treated at all.49 Encouraging results have also been achieved when treating mares with a history of early embryonic death with systemic antimicrobials for five consecutive days each month for the duration of gestation.51 One of the most widely accepted treatments for subfertility in mares is the Caslick vulvoplasty. When performed correctly, this procedure will reduce contamination of the reproductive tract in mares with poor conformation.27 Other suggested treatments include autologous plasma,52 immunostimulants,53, 54 and corticosteroids.55, 56 Early pregnancy losses Increased embryonic losses in older mares have been widely demonstrated.8, 11, 12 Higher embryonic losses were found in mares older than 18 years (18.6%), versus those aged 14–18 years (10.0%) and less than 14 years (5.9–7.1%).9 In the same study, mares bred on foal heat had significantly higher rates of embryo losses (11.8%).9 Morris and Allen2 identified fewer pregnancies lost from day 15–35 in maiden (3.8%) mares as compared to foaling mares (12.4%) and aborted mares (16.7%). Chronic endometritis (aside from age-related changes) is responsible for a large proportion of embryonic losses. Definitive diagnosis of chronic endometritis may be achieved through uterine biopsy.28, 57 Furthermore, mares suffering embryonic losses are frequently observed to have uterine fluid accumulations when they return to heat (despite fluid not being present when losses were first diagnosed).51 Observation of these fluid accumulations at the time of examination may aid in diagnosis of endometritis.
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Reduced embryonic losses have been achieved through the administration of buserelin on days 8–11 following ovulation.58 Other adjunctive treatments that have been reported to decrease pregnancy losses include synthetic progestagens; however, scientific evidence to support the use of such compounds early in pregnancy is lacking.59–61
Abortion Whilst restrictions on the use of artificial breeding techniques are maintained in the Thoroughbred, abortions will inevitably occur as a result of placental insufficiency in mares with chronic endometrial changes. In other breeds, embryo transfer into fertile recipient mares may assist in reducing losses. In mares that must carry their pregnancy to term, attention to surgeries which improve reproductive conformation may assist in reducing the incidence of ascending placentitis, urovagina, and pneumovagina.27,62–65 A reduction in abortions may be achieved after ultrasonographic pregnancy monitoring and targeted systemic antimicrobial therapy.66 Vaccination may assist in reducing losses secondary to equine herpesvirus (EHV) 1 and 4.67, 68 Most studies have identified infectious causes of abortion in 34–40% of cases.69–71 Of infectious causes, bacterial infection may be responsible for up to 50–80% of cases, and Streptococcus spp. and E. coli are most frequently implicated.69–72 Equine herpesvirus is the second most common agent and may cause up to 12% of abortions.67, 69 Other infectious causes including fungi, leptospirosis, and nocardioform actinomycetes may be important rising causes of placentitis.70 Non-infectious problems include placental insufficiency, edema or premature separation, congenital abnormalities, and multiple pregnancy. Umbilical cord torsion may also be an important event, accounting for 36% of abortions in one study.72 Emerging causes of equine abortion include those associated with caterpillar vectors; mare reproductive loss syndrome in the United States and equine amnionitis and fetal loss syndrome in Australia.73, 74 Contagious equine metritis is a declining cause of abortion in the United Kingdom.72
Twins Success rates following manual crushing have improved from 75% in 19838 to 92% recently.2, 9 To achieve good results, it is essential that intervention is performed whilst embryos are still in the mobility phase (i.e., prior to day 16 post-ovulation).12, 75, 76 Under good management conditions in a Thoroughbred population, equivalent,6 or even fewer13 embryonic
losses are observed in multiple pregnancies that are reduced manually with ultrasonographic guidance, compared to singleton pregnancies.6
Stallion management Natural service Even in well-managed commercial operations, poor stallion fertility can significantly impede pregnancy rates and considerably increase on-farm expenses. Knowledge of the stallion’s inherent fertility and his sperm numbers and libido is important to enhance reproductive efficiency. Where possible, subfertile mares should be mated to a highly fertile stallion to achieve acceptable pregnancy rates. Pregnancy rates per cycle for individual commercial stallions can range from 29% to 89%.9, 13 However, for an individual stallion, in good health, reproductive parameters are comparatively consistent.77 When natural service is employed, stallions with a larger number of mares to breed have a higher per cycle pregnancy rate than those with fewer mares.2, 11, 78 Shuttling stallions between hemispheres to serve mares in both seasons is not detrimental to stallion fertility.78 Although stallions will have higher sperm production in the breeding season, when shuttling, individual stallions will adopt one or the other hemisphere as the ‘natural’ breeding season, and consequently have higher sperm numbers at the peak of the season in that hemisphere.77, 78 The reason for higher pregnancy rates in stallions breeding larger numbers of mares is unclear. Farms receiving larger numbers of mares typically are more commercial and have better management, with an increased involvement of stud veterinarians. In addition, stallions being used more frequently are less likely to have problems with sperm accumulations. Possibly, stallions with known high per cycle pregnancy rates will attract more mares. Notwithstanding the above, in immature stallions (< 6 years of age) or in those with poor inherent fertility, having large numbers of mares to breed is likely to be detrimental to the pregnancy rate.79 An accurate prediction of the number of mares an individual stallion is capable of breeding should be made prior to the commencement of the breeding season. When too many mares are booked, per cycle pregnancy rates will be poor, resulting in a large number of mare returns, which further compounds the problem later in the season. Ideally, pre-season breeding evaluation should consist of the following parameters (adapted from Pickett and Voss):78
r r
Gel, gel-free, and total seminal volumes (mL); Sperm concentration (millions per mL of gel-free semen);
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Reproductive Efficiency
r r r r r r r
Total spermatozoa (billions per ejaculate); Percentage of progressively motile spermatozoa and percentage of total motile spermatozoa; Sperm morphology assessment; Cultures should be taken from the urethra, prepuce, and semen after the first two ejaculations; Total scrotal width and testicular volume; Testicular ultrasonography and consistency as determined by palpation; Testing for venereal diseases should also be performed as per the local regulations.
Collation of the above information should allow for a reasonably accurate prediction of the stallion’s fertility and allow the practitioner to limit numbers of mares bred to maintain reproductive efficiency in those stallions that are known to be subfertile. Ideally, each service should provide the mare with 500 × 106 progressively motile spermatozoa.80 Consequently, stallions of normal fertility (with 50% progressively motile), serving mares four times per day, would need a daily sperm output of 4 billion to meet this demand. Daily sperm output can be accurately predicted from the measured scrotal width.77, 81 Historically, total scrotal width has been measured with scrotal callipers, between the greatest curvatures of the left and right testicles.77 Accurate measurements of scrotal width and testicular size can also be taken through ultrasonography, with the added advantage that the testicles can be assessed for uniform echogenicity.82 An estimation of the daily sperm output for a given stallion can be made from measurement of testicular volume calculated from measurements of length, height, and width. Measurement of testicular volume can be used to predict daily sperm output.82 Testicular Volume(TV) = 0.52 × L × W × H Daily Sperm Output(DSO) = (0.024 × TV − 0.76) × 109 DSO = 0.024 × 251 − 0.76 DSO = 5.264 × 109 Table 293.4 Example calculation of testicular volume from the formula TV = 0.52 × L × W × H
Length (cm) Height (cm) Width (cm) Volume (mL) Total volume
Left testicle
Right testicle
9.0 5.5 5.1 131 131 + 120 = 251
8.9 5.2 5.0 120
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A normal stallion aged between 4 and 20 years of age will produce from 3 to 8 billion sperm per day.78 Therefore, normal stallions should easily be able to maintain adequate sperm numbers serving mares 4 times per day for all but the very trough of the breeding season. In practice, however, stallion libido frequently limits the number of mares he will serve naturally. Many stallions will become sexually satiated, and refuse to breed, or develop abnormal sexual behavior when placed on heavy breeding schedules. 80 Furthermore, unless services are managed such that the time between ejaculations is even, stallions are likely to ejaculate highly variable numbers of sperm.77 On-farm record keeping is essential to evaluate the reproductive efficiency of the farm throughout the season. After establishing a baseline estimate of the fertility of the stallion, pregnancy rate per cycle should be assessed weekly. Any sudden change in pregnancy rate should initiate a thorough investigation, including repeat evaluation of the stallion. If the stallion has poorer pregnancy rates for mares bred on the third or fourth service of the day, it is likely he is being overused. Reinforcement breeding has been used to increase pregnancy rates from stallions serving naturally.83 This technique involves collection of a dismount sample from the penis of the stallion after natural service. The dismount sample is typically mixed with a small volume of semen extender (5–10 mL). The served mare has her perineal area cleansed thoroughly and the dismount sample is inseminated into the uterus via an insemination pipette. Frequently, semen which has been expelled into the mare’s vagina may also be aspirated and inseminated into the uterus during this procedure.83 Increased per cycle pregnancy rates after reinforcement breeding may stem from enhanced longevity of the sperm using semen extender as a substrate, or from the deposition of increased sperm numbers within the uterus.83 In the early 1980s, the target for the number of mares bred to an individual stallion per season was up to 120.79 The last decade has seen popular stallions breeding in excess of 200 mares per season in each hemisphere through natural service. This can only be achieved by increasing the efficiency of stallion management, by minimizing the number of services per pregnancy. Consider the following example, which demonstrates the relationship between pregnancy rate per cycle and services per cycle on the number of services per pregnancy. Farm 1 Pregnancy rate per cycle = 70% Services per cycle = 1.5 Services per pregnancy = 2.14
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Table 293.5 Examples of seasonal pregnancy rates under varying management conditions
Daily sperm output Pregnancy rate Mares Services/cycle Services/day Services/week Services/month Mares/month Pregnancies/month Preg rate × 4 months Seasonal pregnancy rate
Stallion 1
Stallion 2
Stallion 3
Stallion 4
Stallion 5
Stallion 6
4 70% 200 1.05 3 21 84 80 56 190/200 95%
4 70% 200 1.2 3 21 84 70 49 184/200 92%
4 45% 150 1.05 3 21 84 80 36 127/150 85%
4 45% 150 1.2 3 21 84 70 32 119/150 79%
2.5 65% 120 1.05 2 14 56 53 34 114/120 95%
2.5 65% 120 1.2 2 14 56 47 31 111/120 92%
84 covers per month ÷ 1.05 covers per mare
Farm 2
= 80 mares per month Pregnancy rate per cycle = 70% Services per cycle = 1.05 Services per pregnancy = 1.5
80 mares per month × 70% PR = 56 pregnancies per month Month 1 : 56 pregnancies + 24 returns leaves 144 mares
Reported number of services per cycle on wellmanaged farms range from 1.12 to 1.2, resulting in overall services per pregnancy of 1.9.2, 9 With precise timing of ovulation in relation to service, and extensive use of ovulation-inducing agents, services per cycle can be reduced to 1.056 or 1.03.13 This represents a repeat service on only 3% of mare cycles. In this same study, in intensively managed Thoroughbreds, a pregnancy rate per cycle of 69% and services per cycle of 1.03 was achieved, resulting in overall services per pregnancy of 1.5. The following example demonstrates how a reduction in the number of services per cycle will achieve higher seasonal pregnancy rates and allow for an increase in the number of mares bred by an individual stallion per year. These calculations may be also be used to estimate how many mares may be booked to a stallion for natural service (modified from Ley).79 Example 1 Ideal management (1.05 covers per mare). Stallion with a pregnancy rate per cycle of 70% and daily sperm output of 4 billion per day and a book of 200 mares (see Table 293.5). 3 covers per day = 21 covers per week 21 covers per week × 4 weeks = 84 covers per month
Month 2 : 56 pregnancies + 24 returns leaves 88 mares Month 3 : 56 pregnancies + 24 returns leaves 32 mares Month 4 : 22 pregnancies + 10 returns Overall 190 mares pregnant of 200 mares = 95% mares in foal
The expectation in well-managed breeding operations is to achieve a seasonal pregnancy rates in excess of 90% and live foal rates in excess of 80%.2 Although the above example is somewhat simplistic, it demonstrates the significantly increased seasonal pregnancy rates that can be achieved with good management. Artificial insemination One of the major advantages of collection of semen for artificial insemination is that every ejaculate is evaluated, with at least a minimal database of total sperm number and total and progressive motility. Thus, emerging problems in semen quality may be diagnosed early, rather than waiting for a perceived reduction in early per cycle pregnancy rates. Artificial
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Reproductive Efficiency insemination techniques also allow for maximal harvesting of seminal output, through collection of semen every 48 hours.77 This may ameliorate problems with stallion libido. Again, mares served with artificial insemination are ideally bred with 500 × 106 progressively motile spermatozoa, and when semen is collected artificially, the number of services per ejaculate can readily be determined. In this manner, the integrity of every service may be confirmed to ensure the mare is given the best possible opportunity for fertilization. A difficulty observed in large-scale fresh and chilled semen artificial insemination programs, such those operating in Australia and New Zealand, is that rigid, every other day, collection schedules frequently necessitate more than one service per cycle. Artificial insemination programs report services per cycle of 1.43 and a pregnancy rate per cycle of 68%.13 The number of mares to be booked to a stallion for artificial insemination programs can similarly be calculated.79 Following from the stallion presented earlier, with a DSO of 4.65 × 109 , a pregnancy rate per cycle of 70% and a progressive motility of 50%; the average number of services per cycle is 1.5. The stallion is booked to 250 mares, and is collected every other day for chilled semen shipment. DSO × progressive motility = Daily output progressively motile spermatozoa (PMS) 5.26 × 109 × 0.5 = 2.63 × 109 PMS per day 2.63 × 109 PMS per day × 2 days between collections = 5.26 × 109 PMS per collection 5.26 × 109 /500 × 106 per dose = 10 doses per collection 10 doses × 15 collections per month = 150 doses per month 150 doses ÷ 1.5 doses per mare = 100 mares per month 100 mares per month × 70% PR = 70 pregnancies per month Month 1 : 70 pregnancies + 30 returns leaves 180 mares Month 2 : 70 pregnancies + 30 returns leaves 110 mares Month 3 : 70 pregnancies + 30 returns leaves
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40 mares Month 4 : 28 pregnancies + 12 returns Seasonal pregnancy rate 238 of 250 mares = 95% mares pregnant The above examples demonstrate that all breeding programs are limited by the daily sperm production and fertility of the stallion. Improvements in breeding efficiency and the number of pregnancies achieved must be directed at reducing the number of services per cycle.
Summary It is clear from the above discussion that in order to fulfil the main objective of a breeding program, which is the birth of a healthy foal, many factors must be managed well. There is a large body of evidence to support increased intensity of management and veterinary intervention in breeding programs from small to large scale. Initial procedures should evaluate the reproductive capabilities of both the mare and stallion. Management of the reproductive cycle through frequent examinations and pharmacological manipulation will ensure accurate timing of service in relation to ovulation and a reduction in the number of services per cycle. Furthermore, early and frequent examinations will allow for rapid intervention to circumvent potential problems. Broad knowledge of the treatment procedures available and relative efficacy will provide the best opportunity for a positive outcome. With this in mind, it is anticipated that reproductive efficiency will continue to improve in the future.
References 1. Ginther OJ. Reproductive efficiency. In: Ginther OJ (ed.) Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1993; pp. 499–562. 2. Morris LHA, Allen WR. Reproductive efficiency of intensively managed Thoroughbred mares in Newmarket. Equine Vet J 2002;34: 51–60. 3. Ginther OJ. Embryonic loss in mares: Incidence, time of occurrence, and hormonal involvement. Theriogenology 1985;23:77–89. 4. Voss JL. Breeding efficiency. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 1109–11. 5. Amann RP. Weaknesses in reports of ‘fertility’ for horses and other species. Theriogenology 2005;63:698–715. 6. McKinnon AO. Reproductive efficiency of horses in Australia. 2000. www.gvequine.com.au/breeding efficiency.htm. 7. Ricketts SW, Young A. Thoroughbred mare fertility [letter]. Vet Rec 1990;126:68.
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8. Sanderson MW, Allen WR. Reproductive efficiency of Thoroughbred mares in the United Kingdom. In: Huntingdon T (ed.) 9th Bain-Fallon Memorial Lectures. Sydney: Australian Equine Veterinary Association, 1987. 9. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of flatrace and national hunt Thoroughbred mares and stallions in England. Equine Vet J 2007;39:438–45. 10. Bosh KA. Reproductive efficiency of thoroughbred mares in Central Kentucky J Equine Vet Sci 2006;26:192–3. 11. Hearn P, Bonnett B, Samper J. Factors influencing pregnancy and pregnancy loss on one thoroughbred farm. Proceedings of the American Association of Equine Practitioners, 1993; pp. 161–3. 12. Bruck I, Anderson GA, Hyland JH. Reproductive performance of Thoroughbred mares on six commercial stud farms. Aust Vet J 1993;70:299–303. 13. Nath LC, Anderson GA, McKinnon AO. Reproductive efficiency of horses in Australia. Aust Vet J 2010;88:169–175. 14. Gilkerson JR, Murray CM. Fertility records from 19051 mares in residence on Australian farms involved in an active surveillance system between 1995 and 2002. In: 45th Congress British Equine Veterinary Association. Birmingham: Equine Veterinary Journal Ltd, 2006; p. 212. 15. Hanlon D. Current strategies to maximise the reproductive performance of Thoroughbred mares. In: Proceedings of the Equine Branch of the New Zealand Veterinary Association. Waikato, New Zealand: 2008; p. 61. 16. Blanchard TL, Thompson A, Brinsko SP, Stich KL, Wendt M, Varner DD, Rigby SL. Mating mares on foal heat: a fiveyear retrospective study. In: 50th Annual Convention of the American Association of Equine Practitioners. Denver, CO: International Veterinary Information Service, 2004. 17. Vogelsang MM, Vogelsang SG, Lindsey BR, Masset JM. Reproductive performance in mares subjected to examination by diagnostic ultrasound. Theriogenology 1989;32:95–104. 18. Douglas-Hamilton DH, Osol R, Osol G, Driscoll D, Noble H. A field study of the fertility of transported equine semen. Theriogenology 1984;22:291–304. 19. Loomis PR. The equine frozen semen industry. Anim Reprod Sci 2001;68:191–200. 20. Squires EL, Barbacini S, Matthews P, Byers W, Schwenzer K, Steiner J, Loomis P. Retrospective study of factors affecting fertility of fresh, cooled and frozen semen. Equine Vet Educ 2006;18:96–9. 21. Barbacini S, Marchi V, Zavaglia G. Equine frozen semen: results obtained in Italy during the 1994–1997 period. Equine Vet Educ 1999;11:109–12. 22. Metcalf ES. Pregnancy rates with cooled equine semen received in private practice. In: 44th Annual Convention of the American Association of Equine Practitioners. Baltimore, 1998; pp. 16–18. 23. Vidament M. French field results (1985–2005) on factors affecting fertility of frozen stallion semen. Anim Reprod Sci 2005;89:115–36. 24. Vidament M, Dupere AM, Julienne P, Evain A, Noue P, Palmer E. Equine frozen-semen: freezability and fertility field results. Theriogenology 1997;48:907–17. 25. Hutton CA, Meacham TN. Reproductive efficiency on fourteen horse farms. J Anim Sci 1968;27:434–8. 26. Jeffcott LB, Rossdale PD, Freestone J, Frank CJ, TowersClark PF. An assessment of wastage in thoroughbred racing from conception to 4 years of age. Equine Vet J 1982;14:185–98. 27. Pascoe RR. Observations on the length and angle of declination of the vulva and its relation to fertility in the mare. J Reprod Fertil Suppl 1979;(27):299–305.
28. Ricketts SW, Alonso S. Assessment of the breeding prognosis of mares using paired endometrial biopsy techniques. Equine Vet J 1991;23:185–8. 29. Ricketts SW, Alonso S. The effect of age and parity on the development of equine chronic endometrial disease. Equine Vet J 1991;23:189–92. 30. McKinnon AO, Squires EL, Harrison LA, Blach EL, Shideler RK. Ultrasonographic studies on the reproductive tract of mares after parturition: Effect of involution and uterine fluid on pregnancy rates in mares with normal and delayed first postpartum ovulatory cycles. J Am Vet Med Assoc 1988;192:350–3. 31. Ball BA, Little TV, Weber JA, Woods GL. Survival of Day4 embryos from young, normal mares and aged, subfertile mares after transfer to normal recipient mares. J Reprod Fertil 1989;85:187–94. 32. Samper JC, Vidament M, Katila T, Newcombe JR, Estrada A, Sargeant J. Analysis of some factors associated with pregnancy rates of frozen semen: a multi-center study. Theriogenology 2002;58:647–50. 33. Zent WW. The postpartum breeding mare. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 455–7. 34. Jasko DJ, Moran DM, Farlin ME, Squires EL, Amann RP, Picket BW. Pregnancy rates utilizing fresh, cooled and frozen-thawed stallion semen. Proceedings of 38th annual convention of the American Association of Equine Practitioners, 1992; pp. 649–60. 35. Troedsson MHT, Desvousges A, Alghamdi AS, Dahms B, Dow CA, Hayna J, Valesco R, Collahan PT, Macpherson ML, Pozor M, Buhi WC. Components in seminal plasma regulating sperm transport and elimination. Anim Reprod Sci 2005;89:171–86. 36. Loomis P. Advanced methods for handling and preparation of stallion semen. Vet Clin North Am Equine Pract 2006;22:663–76. 37. Barbacini S. Management of mares for frozen semen. In: 14th International Conference on Animal Reproduction. Stockholm, Sweden, 2000; p. 308. 38. Heiskanen ML, Huhtinen M, Pirhonen A, Maenpaa PH. Insemination results with slow-cooled stallion semen stored for 70 or 80 hours. Theriogenology 1994;42:1043–51. 39. Newcombe J. The effect of interval from mating to ovulation on pregnancy rate and pregnancy loss rate. In: Proceedings of the British Equine Veterinary Association Congress. Equine Veterinary Journal Ltd, Newmarket 2001. 40. Newcombe J. High pregnancy rate to post-ovulatory mating and insemination In: British Equine Veterinary Association meeting on Artificial Insemination. Whitchurch, UK: Equine Veterinary Journal Ltd, 2003. 41. Reger HP, Bruemmer JE, Squires EL, Maclellan LJ, Barbacini S, Necchi D, Zavaglia G. Effects of timing and placement of cryopreserved semen on fertility of mares. Equine Vet Educ 2003;15:101–6. 42. Woods J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic loss rate in mares. Equine Vet J 1990;22:410– 15. 43. Sieme H, Sch¨afer T, Stout TAE, Klug E, Waberski D. The effects of different insemination regimes on fertility in mares. Theriogenology 2003;60:1153–64. 44. Squires EL, Brubaker JK, McCue PM, Pickett BW. Effect of sperm number and frequency of insemination on fertility of mares inseminated with cooled semen. Theriogenology 1998;49:743–9.
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Reproductive Efficiency 45. Troedsson MH, Loset K, Alghamdi AM, Dahms B, Crabo BG. Interaction between equine semen and the endometrium: the inflammatory response to semen. Anim Reprod Sci 2001;68:273–8. 46. Katila T. Uterine defense mechanisms in the mare. Anim Reprod Sci 1996;42:197–204. 47. McKinnon AO, Perriam WJ, Lescun TB, Walker J, Vasey JR. Effect of a GnRH analogue (Ovuplant), hCG and dexamethasone on time to ovulation in cycling mares. World Equine Veterinary Review 1997;2:16–18. 48. McKinnon AO, Voss JL. Breeding the problem mare. In: McKinnon AO and Voss JL, (eds) Equine Reproduction. Philadelphia; London: Lea & Febiger, 1993; pp. 368–78. 49. Pycock JF, Newcombe JR. Assessment of the effect of 3 treatments to remove intrauterine fluid on pregnancy rate in the mare. Vet Rec 1996;138:320–3. 50. Pycock JF, Newcombe J.R. The relationship between intraluminal uterine fluid, endometritis, and pregnancy rate in the mare. Equine Pract 1996;18:19–22. 51. McKinnon AO. Why is my mare losing her pregnancy? In: Equine Research Seminar 98, Post Graduate Foundation, University of Sydney, Scone, NSW: 1998; pp. 119–41. 52. Pascoe D. (1995) Effect of adding autologous plasma to an intrauterine antibiotic therapy after breeding on pregnancy rates in mares. Biol Reprod Mono 1 1995; 539–43. 53. Rohrbach B, Sheerin P, Steiner J, Matthews P, Cantrell C, Dodds L. Use of Propionibacterium acnes as adjunct therapy in treatment of persistent endometritis in the broodmare. Anim Reprod Sci 2006;94:259–60. 54. Zingher AC. Effects of immunostimulation with PropioniR ) in mares cytologically positive bacterium acnes (Eqstim for endometritis. J Equine Vet Sci 1996;16:100–3. 55. Bucca S, Carli A, Buckley T, Dolci G, Fogarty U. The use of dexamethasone administered to mares at breeding time in the modulation of persistent mating induced endometritis. Theriogenology 2008;70:1093–100. 56. Dell’Aqua Jr J, Papa F, Lopes M, Alvarenga M, Macedo L, Nevesneto J. Fertiltiy rates with equine frozen semen after modulation of inflammatory uterine response. In: International Congress on Animal Reproduction. 2004; p. 391. 57. Ricketts SW. Endometrial biopsy as a guide to diagnosis of endometrial pathology in the mare. J Reprod Fertil Suppl 1975;(23):341–5. 58. Pycock JF, Newcombe JR. The effect of the gonadotropinreleasing-hormone analog, buserelin, administered in diestrus on pregnancy rates and pregnancy failure in mares. Theriogenology 1996;46:1097–101. 59. Allen WR. Is your progesterone therapy really necessary? Equine Vet J 1984;16:496–8. 60. Allen WR. Progesterone and the pregnant mare: unanswered chestnuts. Equine Vet J 1993;25:90–1. 61. Irvine CHG, Sutton P, Turner JE, Mennick PE. Changes in plasma progesterone concentrations from days 17 to 42 of gestation in mares maintaining or losing pregnancy. Equine Vet J 1990;22:104–6. 62. Hemberg E, Lundeheim N, Einarsson S. Retrospective study on vulvar conformation in relation to endometrial cytology and fertility in thoroughbred mares. J Vet Med 2005;52:474–7. 63. McKinnon AO, Belden JO. A urethral extension technique to correct urine pooling (vesicovaginal reflux) in mares. J Am Vet Med Assoc 1988;192:647–50. 64. McKinnon AO, Vasey JR. Selected reproductive surgery of the broodmare. In: Samper JC, Pycock JF, McKinnon AO
65. 66. 67.
68. 69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
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(eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 146–60. Pouret EJM. Surgical technique for the correction of pneumo and urovagina. Equine Vet J 1982;14:249–50. Macpherson ML. Treatment strategies for mares with placentitis. Theriogenology 2005;64:528–34. Kydd JH, Townsend HGG, Hannant D. The equine immune response to equine herpesvirus-1: The virus and its vaccines. Vet Immunol Immunopathol 2006;111:15–30. Wood JL. Vaccination of mares against equine herpesvirus1 [letter]. Vet Rec 1992;130:211–12. Fontaine M, Collobertlaugier C, Tariel G. Abortion and stillbirth of infectious origin in the mare: a study of necropsy of aborted fetuses, premature and stillborn foals. Point Vet 1993;25:79–84. Giles RC, Donahue JM, Hong CB, Tuttle PA, PetritesMurphy MB, Poonacha KB, Roberts AW, Tramontin RR, Smith B, Swerczek TW. Causes of abortion, stillbirth, and perinatal death in horses: 3,527 cases (1986–1991). J Am Vet Med Assoc 1993;203:1170–5. Sandy J. A retrospective survey of Equine Abortions in Victoria, Australia,1987–2004. University of Melbourne, Melbourne, 2008. Smith KC, Blunden AS, Whitwell KE, Dunn KA, Wales AD. A survey of equine abortion, stillbirth and neonatal death in the UK from 1988 to 1997. Equine Vet J 2003;35:496–501. Chicken C, Begg A, Todhunter K. Cluster of equine abortion in Australia associated with bacterial amnionitis – Equine amnionitis and foetal loss – a new condition? Aust Equine Vet 2005;24:127. Cohen ND, Carey VJ, Donahue JG, Seahorn JL, Harrison LR. Descriptive epidemiology of late-term abortions associated with the mare reproductive loss syndrome in central Kentucky. J Vet Diagn Invest 2003;15:295–7. Ginther OJ. The twinning problem: From breeding to day 16. Proceedings of the American Association of Equine Practitioners, 1983; pp. 11–26. McKinnon AO, Rantanen NW. Twins. In: McKinnon AO, Rantanen (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Wilkins, 1998; pp. 141–56. Pickett BW. Factors affecting sperm production and output. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea & Febiger, 1993; pp. 689–704. Pickett BW, Voss JL. Management of shuttle stallions for maximum reproductive efficiency: Part 1. J Equine Vet Sci 1998;18:212. Ley WB. Method of predicting stallion to mare ratio for natural and artificial insemination programs. J Equine Vet Sci 1985;5:143–6. Pickett BW, Squires EL, McKinnon AO, (1987) Procedures for Collection, Evaluation and Utilization of Stallion Semen for Artificial Insemination. Animal Reproduction Laboratory, Colorado State University, 1987; pp. 1–125. Thompson DL, Jr, Pickett BW, Squires EL, Amann RP. Testicular measurements and reproductive characteristics in stallions. J Reprod Fertil Suppl 1979;(27):13–17. Love CC, Garcia MC, Riera FR, Kenney RM. Evaluation of measures taken by ultrasonography and caliper to estimate testicular volume and predict daily sperm output in the stallion. J Reprod Fertil Suppl 1991;44:99–105. Varner DD, Love CC, Brinsko SP, Blanchard TL, Hartman DL, Bliss SB, Carrol BS, Eslick MC. Semen processing for the sub-fertile stallion. J Equine Vet Sci 2008;28:677– 85.
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Chapter 294
Pituitary Pars Intermedia Dysfunction (Equine Cushing’s Syndrome) Philip J. Johnson, Nat T. Messer and Venkataseshu K. Ganjam
Equine Cushing’s syndrome is the most common endocrinological condition of older horses. Conservative estimates of the prevalence of the condition range from 0.075% to 0.5% of the equine population. Presently, the preferred term for equine Cushing’s syndrome is pituitary pars intermedia dysfunction (PPID). The clinical recognition and decisions to manage horses affected with PPID are reportedly increasing as a result of societal trends to maintain horses to an older age. The health of older horses in general has benefited from substantial improvements in many aspects of the veterinary care of geriatric horses, growing interest in and awareness of PPID, extensive new information published in the literature, and the introduction of new diagnostic tests and effective treatments for PPID.1
Pathophysiology Pituitary pars intermedia dysfunction occurs as a result of clonal expansion of the melanotrope cell population in the pars intermedia (PI) of the pituitary gland.2 The resulting changes in the PI of affected horses have been variously referred to as pituitary hypertrophy, pituitary hyperplasia, or pituitary adenoma (Figure 294.1). Hypercortisolism resulting from primary adrenal gland tumors is evidently rare in horses.3 In health, melanotropes in the PI normally secrete relatively small quantities of the following protein hormones: alpha-melanocyte-stimulating hormone, beta-endorphin, adrenocorticotropic hormone (ACTH), and corticotropin-like intermediate peptide
(CLIP).4 These hormones are derived from a single large pre-prohormone called pro-opiomelanocortin (POMC) and are usually collectively referred to as POMC peptides.5 Secretion of the POMC peptides by melanotropes is normally under the tonic inhibitory influence of dopaminergic nerves originating in the periventricular nuclei (cell bodies) of the hypothalamus.6 Excessive secretion of PI-derived POMC peptides and clonal expansion of melanotropes in the PI of PPID-affected equids results from the loss of inhibitory dopaminergic nerves.7 The loss of dopaminergic nerves in PPID results from the toxic action of oxidative stress (dopaminergic nerves are especially sensitive to oxidative stress). In this regard, equine PPID bears some resemblance to Parkinson’s disease in humans (also associated with loss of dopaminergic nerves as a result of oxidative damage).8 The clinical consequences of PPID result from the endocrinological consequences of heightened POMC peptide secretion, physical enlargement of the PI, and stimulated secretion of cortisol by the adrenal glands (hypercortisolism). However, adrenocortical hyperplasia is identified in only approximately 20% of PPID-affected equids at necropsy.9 Although stimulated secretion of cortisol results from increased PI-derived ACTH, the steroidogenic effectiveness of ACTH is significantly enhanced by the other POMC peptides.10 Expansion of the PI leads to grossly evident enlargement of the pituitary gland in a majority of PPID-affected equids.11 When present, expansion of the PI may contribute to the clinical signs
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Pituitary Pars Intermedia Dysfunction (Equine Cushing’s Syndrome)
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Clinical recognition of PPID
Figure 294.1 Photograph of an enlarged pituitary gland obtained during postmortem examination of a horse affected by pituitary pars intermedia dysfunction.
by directly compromising (by compression) adjacent structures including the pars ventralis (PV), the neurohypophysis, optic chiasm, and overlying hypothalamus. Glucocorticoids are potent inhibitors of the action of insulin, and insulin resistance (IR) develops in many PPID-affected equids.12 When severe, the presence of concomitant IR indicates that the prognosis for PPID is relatively unfavorable.13 It should be emphasized that the equine pituitary Cushing’s syndrome arising from PPID is quite different to Cushing’s syndromes resulting simply from hypercortisolism secondary to excess ACTH secreted by corticotropinomas in the PV, as occur more commonly in human and canine patients. In the equine condition, the clinical consequences of PPID result from both the stimulated secretion of cortisol and the actions of enhanced secretion of the other PI-derived POMC peptides.
(a)
The diagnosis of PPID is not straightforward, especially in younger patients in which the pathognomonic physical appearance of the condition may not be evident. Moreover, identification of a specific starting point for PPID in a given patient is difficult because of the overlap with other age-related changes and conditions and the fact that PPID is an insidiousonset, progressively degenerative condition.7 The age at which most equids are presented to veterinarians for problems attributable to PPID tends to exceed 18 years. The youngest age at which PPID has been confirmed (necropsy) in our teaching hospital is 4 years. There is a growing recognition that the extent to which PPID is present in younger equids (< 18 years) has likely been underestimated. Recent publications have pointed to the fact that PPID is present in younger horses in the absence of the classic clinical signs such as hirsutism and weight loss.14 There does not appear to be either a strong sex or breed predisposition for PPID.1 However, the influence of ovarian hormones has been suggested to be a predisposing factor for PPID in older mares.15 Pituitary pars intermedia dysfunction is commonly suspected and diagnosed upon recognition of its characteristic physical appearance in older equids (Figure 294.2). Failure to shed out the hair coat in spring (inappropriate hirsutism) is widely regarded as pathognomonic for PPID. In early cases, long hairs under the mandible and at the palmar and plantar aspects of the lower limbs may be implicative for PPID, even if the remainder of the hair coat has been shed appropriately. The presence of chronic laminitis is also a strong indicator that PPID should be suspected in affected individuals (Figure 294.3). In one study, chronic laminitis was the most likely clinical expression of PPID, even in the absence of hirsutism.16
(b)
Figure 294.2 (a,b) Photographs of two horses affected by pituitary pars intermedia dysfunction. Notice the poor bodily condition and the inappropriate hirsutism/failure to properly shed out the haircoat.
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Figure 294.3 Photograph of the forefeet of a horse affected by pituitary pars intermedia dysfunction. Notice the prominent growth rings (‘fever rings’) and the palmar divergence of those rings that are implicative for underlying chronic laminitis.
The onset of PPID commonly leads to the development of an abnormal body condition in older horses. Affected horses tend to develop a ‘poor’ overall condition characterized by skeletal muscle loss17 and the acquisition of a ‘regional’ distribution of body fat. Enhanced subcutaneous repositories of adipose tissue tend to develop in the supraorbital fossae, along the crest of the neck (‘cresty’ neck), adjacent to the tail head, in the prepuce, and near the shoulders. However, PPID may also develop in overtly obese equids in which hirsutism is not present. Other clinical signs of PPID that arise variably between affected individuals include polyuria/polydipsia (PU:PD), inappropriate sweating (‘hyperhidrosis’), lethargy, infertility, inappropriate lactation (mares), blindness, seizures, and chronic infections resulting from immune compromise.9 The development of infectious disease in matureto-older equids should signal concern for underlying PPID-associated immunocompromise. Many different chronic or subclinical infectious diseases in this population of horses have been attributed to underlying PPID and should benefit from the treatment of PPID. Some of the infectious diseases that have been associated with PPID include equine protozoal myeloencephalitis (EPM), chronic endometritis, pyometra, infectious respiratory disease, chronic dermatitis, gastrointestinal helminth parasitism, and chronic urinary tract infections.
Diagnostic corroboration of PPID The diagnosis of PPID in older horses that have developed inappropriate hirsutism is often presump-
tively made without resort to further corroborative endocrinological testing. The fact that the cost of (and lifelong commitment to) treatment for PPID using pergolide is not inconsequential suggests that endocrinological corroboration should be considered, especially when the clinical diagnosis is not clear-cut. Although many diagnostic test protocols have been variably investigated and advocated for the purpose of making an accurate diagnosis of PPID, an unequivocal test is still lacking. A complete discussion of the strengths and weaknesses of PPID test protocols is beyond the scope of this chapter, and the reader is directed elsewhere.18, 19 During the interpretation of diagnostic tests, it is important carefully to consider the interplay between the endocrinological abnormalities resulting from PPID and the effects of other (independent) ageassociated disease conditions on the hypothalamic– pituitary–adrenal axis. Moreover, the secretory output of both the normal and PPID-affected equine pituitary gland appears to be increased during the fall (September). Veterinary diagnosticians should beware of the fact that results of several PPID test protocols yield false-positive results when undertaken at this time of the year.20, 21 As with all diagnostic tests, before submitting blood samples to a reference laboratory, it is important to ensure that the samples will be handled according to the laboratory specifications and that the receiving laboratory has a validated assay with an appropriately established reference range for the hormone of interest. Simple determination of the plasma cortisol concentration is not helpful for the diagnosis of PPID because there is too much overlap between ‘reference’ values for cortisol between healthy horses, horses affected by many different illnesses, horses affected by PPID, and PPID-affected horses with other illnesses. Moreover, there is further variation associated with the age of the horse. Almost all commercial clinical pathology laboratories are unable to provide properly determined reference ranges for the plasma cortisol concentration that account for these categorizations. The overnight low-dose dexamethasone (DEX) suppression test (DST) is often advocated as the classic ‘gold standard’ antemortem test for PPID.22 Failure of DEX to suppress the plasma cortisol concentration has been attributed to uninhibited production of ACTH by melanotropes in the PI that is not under the influence of negative feedback. Recent work suggests that this test may not be as accurate as had been previously contended. As with many diagnostic tests for PPID, the DST may yield falsely positive results when performed in the fall because the secretory output of the healthy equine pituitary gland is increased at this time of the year.21
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Pituitary Pars Intermedia Dysfunction (Equine Cushing’s Syndrome) The DST protocol entails administering DEX (0.04 mg/kg bodyweight, i.m.) and evaluating its effect on the circulating plasma (or serum) concentration of cortisol after 19 hours. In healthy horses, DEX inhibits the release of ACTH from the PV (negative feedback) and the cortisol concentration decreases to less than 1 µg/dL. In PPID-affected equids, the cortisol concentration is not reduced to this extent following treatment with DEX. Objections to the use of the DST are centered on the reputation of DEX as a risk factor for laminitis. Nevertheless, it appears that the risk of inciting or aggravating laminitis as a result of the DST is very small. Demonstration of an elevated plasma ACTH concentration is a convenient diagnostic test with good sensitivity and specificity, commonly employed for the diagnosis of PPID in horses and ponies.23, 24 It is important to consider the (fall) seasonal factor when interpreting plasma ACTH results for diagnosis of PPID because plasma ACTH values often exceed the reference range in September in normal horses. The extent to which ACTH is released by the pituitary gland fluctuates (pulsatile release) during the day. Veterinary diagnosticians should be aware of the fact that in our experience single sample determinations of plasma ACTH concentration for purposes of diagnosis of PPID often lead to false-negative results. There is considerable interest in the use of a recently described domperidone challenge test for purposes of making a diagnosis of PPID in horses.11 Domperidone, an orally administered dopamine (D2) receptor antagonist, is commonly used for the management of fescue agalactia in pregnant mares. In normal horses, the administration of domperidone does not affect the plasma ACTH concentration. In PPID-affected equids, treatment with domperidone causes a significant elevation of the plasma ACTH concentration. This effect has been attributed to the concept that hypertrophied melanotropes in the PI of PPID-affected equids continue to be under some degree of dopaminergic inhibition and that treatment with domperidone abolishes residual inhibition leading to an elevation in the plasma ACTH concentration.11 The domperidone challenge test entails collection of a blood sample for determination of the plasma ACTH concentration at 8.00 am followed by treatment using domperidone (3.3 mg/kg, p.o.). Blood is collected at + 4 h and at + 8 h for further evaluation of the plasma ACTH concentration. Diagnosis of PPID is supported by demonstrating both an elevated plasma ACTH concentration at the zero time (above the reference range) and/or an elevation of the plasma ACTH concentration at either or both the + 4 h and at + 8 h sample times. Although preliminary results regarding the domperidone challenge test indicate that it is practical
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and safe, and promises high diagnostic accuracy, further work on a larger number of normal and PPIDaffected equids must yet be undertaken to determine its sensitivity and specificity. Similarly, it remains to be seen whether the test might yield a relatively higher number of false-positive test results in the fall. Several other diagnostic protocols for PPID have been published. As a rule, some of the simpler tests that are currently advocated for diagnosis of PPID lack sufficient accuracy (or scientific corroboration), and the more accurate tests, although not perfect, are sometimes more readily performed at a referral institution.25–27 The extent to which IR affects a PPID-affected patient should be considered when addressing treatment. Diagnostic testing for IR may be complex and is beyond the scope of this discussion. However, a clinically useful diagnostic strategy for IR employs the combined intravenous administration of both glucose and insulin (glucose 150 mg/kg; insulin 0.1 U/kg).28 The blood glucose concentration is then measured throughout the subsequent 2 hours (12 blood glucose determinations at zero, + 1, + 5, + 15, + 25, + 35, + 45, + 60, + 75, + 90, + 105 and + 120 minutes). The test outcome is evaluated based on the time taken for the plasma glucose concentration to return to the baseline (time zero) level after administration of glucose and insulin. Normal insulin sensitivity is associated with a return to baseline within 45 minutes. It is important to recognize that horses are adept at increasing their pancreatic insulin secretion in response to decreased insulin sensitivity. Therefore, diagnostic protocols that simply evaluate glucose responses may reveal normal CGMS and CGIT results in even insulin-resistant patients. The detection of higher insulin concentrations during the CGIT reveals increased compensation (presumed to be insulin resistance). Currently therefore, CGIT testing for purposes of demonstrating IR requires that the serum insulin concentration should also be measured at zero and +45 minutes. There is a very small risk of development of signs of insulin-provoked hypoglycemia (weakness, trembling, and sweating) during the test. In the event that these signs develop, the patient should be treated with 50% glucose (i.v.). The test should always be performed in a quiet environment in patients that have been allowed to consume a simple grass hay-based ration. Some practitioners elect to investigate patients for IR by simply measuring the resting plasma insulin and glucose concentrations (hyperinsulinemia is commonly attributable to IR).
Management of PPID Treatment approaches for PPID may conveniently be classified in the following three categories: (i)
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management of concurrent diseases associated with the aged horse; (ii) pharmacological strategies aimed specifically at ameliorating the hormonal manifestations of PPID; and (iii) dietary strategies. Treatment strategies should be carefully tailored to the needs of the individual. As with all older horses, PPID-affected equids usually require special attention regarding the health of their teeth, nutrition, and feet. Morbidity associated with age-dependent dental attrition is often more severe in PPID-affected equids because of the combined effects of immune compromise and osteoporosis. Treatment for laminitis is often indicated and should be undertaken following acquisition of radiographic images of the patient’s feet. Manifestations of laminitis usually respond favorably to treatments addressing the underlying endocrinopathy, appropriate changes to the ration, and judicious hoof care. Clinical signs of laminitic pain are often relatively subdued in PPID-associated laminitis when considered in light of the extent to which the structural integrity of the hoof lamellar interface and the pedal bone has been compromised. It is conjectured that the combined effects of both hypercortisolism and elevated plasma beta-endorphin levels may inhibit expression of laminitic pain in PPID-affected equids. The nutritional requirements of PPID-affected equids should take into account the likelihood of complicating dental attrition and the observation that many of these horses are concomitantly affected with IR. Dietary strategies aimed at specifically addressing the presence of IR have been published.29 It is customary that hirsute horses will be clipped to help with thermoregulation in the warmer months of the year. Affected horses may be treated with longterm antimicrobials to control the morbidity associated with low-grade, chronic, and subclinical infections (especially affecting the lower urinary and respiratory tracts). Presently, the most effective pharmacological treatment for PPID is achieved by administering a low dose of pergolide (0.5 to 3.0 mg/day, p.o.) on a daily basis.24 Pergolide is a selective D2 dopaminergic agonist. Pergolide was manufactured for the purpose of treating human patients with Parkinson’s disease but has been recently withdrawn because use of the drug is associated with an unacceptable risk of heart valve disease. Therefore, for treatment of PPID in horses, veterinarians are presently using compounded sources of pergolide. Other dopamine agonists available on the human market, including cabergoline and quinagolide, will likely be evaluated for suitability for the management of equine PPID in the near future. Although the serotonin antagonist, cyproheptadine, has been advocated for the management of PPID in horses, published results of its efficacy
have revealed disappointing and unreliable clinical responses. The clinical response to cyproheptadine treatment for PPID-affected equids certainly appears to be inferior to that for pergolide.14 Cyproheptadine is sometimes administered with pergolide in refractory cases of PPID. Trilostane, a competitive inhibitor of adrenal steroidogenesis (specifically it inhibits the enzyme 3β-hydroxysteroid dehydrogenase), has been advocated for the management of PPID.13 In light of the fact that adrenocortical hyperplasia is identified in only 20% of PPID-affected equids, inhibition of adrenal biosynthesis of cortisol may not be very effective in most cases. Moreover, trilostane is not readily available to veterinarians in the USA. Assessment of response to treatment There presently exists a lack of evidentiary support for the application of any diagnostic endocrinological test for purposes of demonstrating clinical resolution/control during the pharmacological management of PPID. At best, veterinary clinicians may elect to monitor the resting plasma ACTH concentration as an indicator of improvement (reduction in ACTH), but plasma ACTH concentrations may remain elevated in the face of substantial improvement in clinical signs. Clinical observation is still regarded as one of the best methods to evaluate response to treatment. Effect of PPID on reproductive status The extent to which development of PPID affects reproductive functions in male and female horses has not been reported in the literature. Compromised immunity resulting from PPID has been linked to increased risk for development of uterine infections and pyometra, which represent significant impediments to fertilization and pregnancy. Elevated blood cortisol levels have also been identified as a risk factor for infertility in women. For example, infertility may result from perturbations in the regulation of the hypothalamic–pituitary–gonadotropic (HPG) axis caused by the hypercortisolism associated with mental stress. The extent to which cortisol affects the HPG axis in women depends on the endocrine status of the ovaries within different stages of the menstrual cycle. Dexamethasone, a potent synthetic corticosteroid, acts to inhibit sexual behavior in mares, reduce ovarian follicle size, and inhibit luteinizing hormone (LH) release from the pituitary gland.30 However, the extent to which demonstrable DEXinduced abnormalities in reproductive function can be used to suggest that similar abnormalities occur in PPID-affected mares is unknown. Stress-associated hypercortisolism resulting from repeated strenuous treadmill exercise adversely affects both semen quality and freezability.31 Therefore, it is reasonable
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Pituitary Pars Intermedia Dysfunction (Equine Cushing’s Syndrome) to conclude that semen quality would also likely be adversely affected by the development of PPID in stallions. It is reasonable to further deduce that physical expansion of the PI could affect the secretion of gonadotropic hormones from the pars ventralis in PPID-affected horses.
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References 18. 1. Messer NT, Johnson PJ. Evidence based literature pertaining to thyroid dysfunction and Cushing’s Syndrome in the horse. Vet Clin N Am Equine Pract 2007;23:329–64. 2. McFarlane D, Miller LM, Craig LE, Dybdal NO, Habecker PL, Miller MA, Patterson JS, Cribb AE. Agreement in histologic assessments of the pituitary pars intermedia in aged horses. Am J Vet Res 2005;66:2055–9. 3. van der Kolk JH, Ijzer J, Overgaauw PA, van der LindeSipman JS. Pituitary-independent Cushing’s syndrome in a horse. Equine Vet J 2001;33:110–12. 4. Wilson MG, Nicholson WE, Holscher MA, Sherrell BJ, Mount CD, Orth DN. Proopiolipomelanocortin peptides in normal pituitary, pituitary tumor, and plasma of normal and Cushing’s horses. Endocrinology 1982;110:941–54. 5. Orth DN, Nicholson WE. Bioactive and immunoreactive adrenocorticotropin in normal equine pituitary and in pituitary tumors of horses with Cushing’s disease. Endocrinology 1982;111:559–63. 6. McFarlane D, Cribb AE. Systemic and pituitary pars intermedia antioxidant capacity associated with pars intermedia oxidative stress and dysfunction in horses. Am J Vet Res 2005;66:2065–72. 7. McFarlane D, Dybdal N, Donaldson MT, Miller L, Crib AE. Nitration and increased alpha-synuclein expression associated with dopaminergic neurodegeneration in equine pituitary pars intermedia dysfunction. J Neuroendocrinol 2005;17:73–80. 8. McFarlane D. Advantages and limitations of the equine disease, pituitary pars intermedia dysfunction as a model of spontaneous dopaminergic neurodegenerative disease. Ageing Res Rev 2007;6:54–63. 9. Schott, H. C., II. 2002. Pituitary pars intermedia dysfunction: equine Cushing’s disease. Veterinary Clinics of North America, Equine Practice 18(2):237–270. 10. Shanker G, Sharma A. Beta-endorphin stimulates corticosterone synthesis. Biochem Biophys Res Commun 1979;86:1–5. 11. Miller MA, Pardo ID, Jackson LP, Moore GE, Sojka JE. Correlation of pituitary histomorphometry with adrenocorticotrophic hormone response to domperidone administration in the diagnosis of equine pituitary pars intermedia dysfunction. Vet Pathol 2008;45:26–38. 12. Garcia MC, Beech J. Equine intravenous glucose tolerance test: glucose and insulin responses of healthy horses fed grain or hay and of horses with pituitary adenoma. Am J Vet Res 1986;47:570–2. 13. McGowan CM, Frost R, Pfeiffer DU, Neiger R. Serum insulin concentrations in horses with equine Cushing’s syndrome: response to a cortisol inhibitor and prognostic value. Equine Vet J 2004;36:295–8. 14. Donaldson MT, LaMonte BH, Morresey P, Smith G, Beech J. Treatment with pergolide or cyproheptadine of pituitary pars intermedia dysfunction (equine Cushing’s disease). J Vet Intern Med 2002;16:742–6. 15. van der Kolk JH, Heinrichs M, van Amerongen JD, Stooker RC, in de Wal LJ, van den Ingh TS. Evaluation of pituitary gland anatomy and histopathologic findings in clinically
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29. 30.
31.
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normal horses and horses and ponies with pituitary pars intermedia adenoma. Am J Vet Res 2004;65:1701–7. Donaldson MT, Jorgensen AJR, Beech J. Evaluation of suspected pituitary pars intermedia dysfunction in horses with laminitis. J Am Vet Med Assoc 2004;224:1123–7. Aleman M, Watson JL, Williams DC, LeCouteur RA, Nieto JE, Shelton GD. Myopathy in horses with pituitary pars intermedia dysfunction (Cushing’s disease). Neuromuscul Disord 2006;16:737–44. McFarlane D. Diagnosing equine pituitary pars intermedia dysfunction in ambulatory practice. P Annu Conv Am Equine 2008;54; 204–6. Messer NT, Johnson PJ, Ganjam VK. Diagnosis of pituitary pars intermedia dysfunction: a review of 1999 versus 2008. P Annu Conv Am Equine 2008;54; 200–3. McFarlane D, Donaldson MT, McDonnell SM, Cribb AE. Effects of season and sample handling on measurement of plasma alpha-melanocyte-stimulating hormone concentrations in horses and ponies. Am J Vet Res 2004;65:1463–8. Donaldson MT, McDonnell SM, Schanbacher BJ, Lamb SV, McFarlane D, Beech J. Variation in plasma adrenocorticotropic hormone concentration and dexamethasone suppression test results with season, age, and sex in healthy ponies and horses. J Vet Intern Med 2005;19:217–22. Dybdal NO, Hargreaves KM, Madigan JE, Gribble DH, Kennedy PC, Stabenfeldt GH. Diagnostic testing for pituitary pars intermedia dysfunction in horses. J Am Vet Med Assoc 1994;204:627–32. Cou¨etil L, Paradis MR, Knoll J. Plasma adrenocorticotropin concentration in healthy horses and in horses with clinical signs of hyperadrenocorticism. J Vet Intern Med 1996;10: 1–6. Perkins GA, Lamb S, Erb HN, Schanbacher B, Nydam DV, Divers TJ. Plasma adrenocorticotropin (ACTH) concentrations and clinical response in horses treated for equine Cushing’s disease with cyproheptadine or pergolide. Equine Vet J 2002;34:679–85. Eiler H, Oliver JW, Andrews FM, Fecteay KA, Green EM, McCracken M. Results of a combined dexamethasone suppression/thyrotropin-releasing hormone stimulation test in healthy horses and horses suspected to have a pars intermedia pituitary adenoma. J Am Vet Med Assoc 1997;211:79–81. Frank N, Andrews FM, Sommardahl CS, Eiler H, Rohrbach BW, Donnell RL. Evaluation of the combined dexamethasone suppression/thyrotropin-releasing hormone stimulation test for detection of pars intermedia pituitary adenomas in horses. J Vet Intern Med 2006;20:987–93. McFarlane D, Beech J, Cribb A. Alpha-melanocyte stimulating hormone release in response to thyrotropin releasing hormone in healthy horses, horses with pituitary pars intermedia dysfunction and equine pars intermedia explants. Domest Anim Endocrinol 2006;30:276–88. Eiler H, Frank N, Andrews FM, Oliver JW, Fecteau KA. Physiologic assessment of blood glucose homeostasis via combined intravenous glucose and insulin testing in horses. Am J Vet Res 2005;66:1598–604. Johnson PJ, Messer NT, Kellon E. Treatment of equine metabolic syndrome. Comp Cont Educ Pract 2004;26:122–30. Asa CS, Ginther OJ. Glucocorticoid suppression of oestrus, follicles, LH and ovulation in the mare. J Reprod Fertil Suppl 1982;32:247–51. Janett F, Burkhardt C, Burger D, Imboden I, Hassig M, Thun R. Influence of repeated treadmill exercise on quality and freezability of stallion semen. Theriogenology 2006;65:1737–49.
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Chapter 295
Epidemiology Applied to Horse Reproduction Nigel R. Perkins
Introduction The purpose of this chapter is to provide a brief introduction to veterinary epidemiology and related topics in the context of horse reproduction. This chapter does not attempt to provide detailed coverage of material or instruction on how to design or perform epidemiological studies. Interested readers are referred to a range of reference material for such content1–5 as well as related articles that offer useful insights or reviews.6, 7 The chapter provides a brief introduction to some of the common study types employed in veterinary epidemiology and illustrates these with examples of published studies that have employed these approaches on horse topics. Epidemiology is traditionally defined as the study of the distribution and determinants of disease frequency in populations.5 Veterinary epidemiology extends the scope to include health-related events such as measures of productivity.4 The word determinant is used to describe any factor that is capable of influencing some outcome measure of interest that is related to health or disease. Alternative terms include exposure or risk factor.8 Others restrict terms such as determinant or risk factor to those factors that are considered to precipitate disease.9 The focus on populations is a key strength of epidemiology and an important differentiation from the clinician’s focus on an individual patient.10, 11 Epidemiological studies often involve observations made on individual animals, but analyses and inference are generally based on aggregates or populations. A second key feature of epidemiology is the focus on epidemiological methods as tools to facilitate collection, analysis, and interpretation of relevant data
and information on exposures and outcomes of interest.11 Epidemiological methods represent an applied branch of statistics, and a strong understanding of principles of statistical theory and methods are important competencies for epidemiology. Martin et al.8 define the major purpose of epidemiology as providing data on which rational decisions can be made for the prevention and/or control of disease and optimization of health or productivity in animal populations. This requires that epidemiological studies produce valid measures of association between outcomes and their determinants.7 It is important to note that statistical evidence of association between factors does not necessarily demonstrate a causal relationship, and that effective understanding and application of epidemiological methods requires knowledge of the principles of causality.4, 5 Optimal application of epidemiology generally involves collaborations between epidemiologists and individuals from other disciplines including, for example, clinicians, pathologists, economists, statisticians, mathematical modelers, geneticists, ecologists, sociologists, and others with a common focus on health and productivity of animals. The broad discipline of epidemiology with its population focus has been extended to include the application of epidemiological principles and methods to decisions concerning care of individual patients through the subdiscipline of clinical epidemiology.2, 4, 12, 13 In addition there is considerable overlap between principles and methods employed by epidemiologists and the related discipline of evidencebased medicine (EBM), which is concerned with critical appraisal of available evidence in making decisions about the care of individual patients.14–16
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Epidemiology Applied to Horse Reproduction
Reproductive epidemiology Reproductive epidemiology is not considered to be a distinct subdiscipline of epidemiology,4 but instead involves the full array of epidemiological principles and methods being applied to outcomes related to reproduction. Reproductive health and performance covers a wide range of areas from development of reproductive capacity in fetal and prepubertal animals, onset of puberty, and subsequent adult reproductive capacity including production and delivery of gametes, fertilization, pregnancy and parturition, and later decline of reproductive capacity and performance in older animals.17 Weinberg and Wilcox17 have identified a number of issues associated with reproduction that present methodological challenges for epidemiologists. Many of these are common to all epidemiological investigations. Equine adapted examples of selected methodological issues identified by Weinberg and Wilcox17 are presented here.
1. Animals may have abnormal reproductive function without any overt signs of disease. 2. End points of interest for reproduction such as infertility, early pregnancy loss, abortion, stillbirths, and neonatal mortality, may not be independent, leading to difficulties in characterizing relationships between putative exposures and particular outcomes. For example, an exposure that results in increased risk of early pregnancy loss in conceptuses that may later have undergone abortion, may appear to be a protective exposure if the outcome of interest was limited to abortions in later pregnancy – because of the lack of independence between the two events (early pregnancy loss and later abortion). 3. Case–control studies may be less appropriate for investigating reproductive outcomes than cohort studies because of the difficulties in defining and identifying particular end points and the lack of independence between various adverse outcomes at different stages of the reproductive cycle as described above. Reproductive studies often focus on the dam and may not reflect the important contribution of both sire and dam to outcomes related to conception, pregnancy, and parturition. 4. Care is required in defining populations at risk (denominators) for reproductive outcomes. For example, studies of early pregnancy loss are particularly problematic because it is very difficult to identify the number of conceptions that occur since the equine conceptus cannot be reliably detected by ultrasound until approximately 9 days post-ovulation and alternative laboratory tests that may detect pregnancy earlier than this
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may be subject to false-negative and false-positive results. 5. Reverse causality is a potential problem for many reproductive studies. For example, mares that are believed to be at some risk of an adverse pregnancy outcome may be managed differently to mares that are believed to be normal on some scale of assessment or expectation. This discrepancy in management of mares may result in bias in assessment of management during breeding and pregnancy.
Principles of epidemiology Martin et al.8 describe four principles that underpin most epidemiological study. The first is that disease occurrence in a population can be related to the interaction between the host, agent, and a wide range of environmental factors including biological, physical, and sociological influences. This relationship is often described as the epidemiological triad. The second principle of epidemiology is the fundamental role of counting or quantification of events in order to be able to describe events and to perform epidemiological analyses to understand relationships between outcomes and determinants. The third principle is that much of epidemiology involves observational studies of animals and disease. This is both a strength and a source of problems for the discipline. Observational studies provide an opportunity to study outcomes of interest in the natural environment in which they occur as opposed to the more contrived environment of controlled, experimental trials. However, the inability to influence subject selection and exposures in observational trials provides opportunities for biases that present additional methodological challenges for observational studies.18 The fourth principle is that controlled field trials performed in the natural environment provide an excellent opportunity to test preliminary hypotheses and in particular the effects of possible interventions for control or prevention of a disease.
Common epidemiological study designs The following section provides brief descriptions of the more common types of epidemiological studies with examples selected from refereed papers published in the scientific literature. Epidemiological studies have been categorized as descriptive or analytic,19 with the distinction being whether the study simply describes events
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(descriptive) or whether it makes an attempt to investigate associations between outcomes and potential exposures (analytic). Analytic studies have the potential to allow specific hypotheses to be tested whereas descriptive studies are really limited to the generation of hypotheses.7 It is stressed that the quality of evidence presented by epidemiological studies is dependent in turn on the design and implementation of the study.7 Guidelines have been developed for a range of study types including randomized clinical trials (RCTs),20, 21 diagnostic test evaluations,22 non-randomized intervention evaluations,23 and observational studies in epidemiology.24–26 Descriptive studies provide summary information on events, generally presented as a case report or case series, or as results derived from a survey. Reports of individual cases or a series of cases generally describe novel features of particular cases (presentation, progression, particular challenges, lessons learned, etc.). Examples of recently published case series include descriptions of survival and fertility in 72 mares following fetotomy,27 and of 73 cases of periparturient hemorrhage in mares.28 Descriptive studies may also involve sampling animals in a survey, generally to describe the frequency and distribution of a particular outcome in the animals sampled.19 Surveys may involve crosssectional or prospective sampling. The National Animal Health Monitoring Survey (NAHMS) Study provides an example of a carefully designed approach to survey sampling that can be used to generate statistically valid population estimates at the national level for the USA.29, 30 Surveys involving formal random sampling are more likely to produce results that are representative of the target population than those that do not have formal sampling. Fiala et al.31 collected a sample of mare oviducts from animals processed at an abattoir and while these samples are not likely to be representative of the general horse population they should be representative of the population of mares presented to the abattoir for processing. Additional example surveys involved what appeared to be a convenience sampling approach, producing results that may be less likely to be representative of any broader population.32–35 Analytical studies may also be categorized as observational or experimental. The major distinction is that experimental studies involve some form of allocation of animals to study groups whereas observational studies involve no such allocation and animals are simply observed to determine outcome and exposure status. Observational studies include cross-sectional studies, cohort, and case–control designs, and occasionally hybrid designs that may have some combination of features of the different designs.
There are a number of challenges in the design and implementation of observational studies, particularly in relation to eligibility and selection of animals, definitions of outcome(s) and exposure(s) of interest, and especially for cohort studies how incidence rate/risk will be estimated.36 Cross-sectional studies are commonly performed in veterinary epidemiology because they are costeffective and relatively easy to implement. Crosssectional studies are limited to describing prevalence, and investigation of associations between exposures and outcomes is hampered by difficulties in determining whether exposure occurred prior to the outcome or after since they are generally both measured at the same time.19 The ability to extrapolate results to a target population requires a formal random process of selecting subjects for inclusion in the study. One example paper describes the prevalence of anthelmintic resistance in a sample of horse farms across the southern USA.37 The classic cohort study is the ‘archetype for all epidemiologic studies’36 and involves selection of a group of animals (cohort) that is known to be exposed to a hypothesized risk factor, and a comparable group (cohort) that is not exposed to the same risk factor. Cohort studies require that animals do not have the outcome of interest at the time the study begins, i.e., they are disease free at the start of the study. The two groups are then observed for evidence of development of the outcome of interest (disease or productivity measure).4, 38 A modification of this design involves selection of a single group of animals that are likely to include animals exposed and unexposed to the risk factors of interest and then following them to observe development of the outcome(s) of interest and also to determine their exposure status for exposures of interest. This form of cohort study is described as a single cohort or longitudinal study.38 Cohort studies may be descriptive or analytical and may involve prospective or retrospective approaches to collection of data. A retrospective cohort study involves use of historical information to identify cohorts, exposures, and outcomes. Cohort studies are suitable for estimation of incidence rate/risk of disease in each cohort, and the use of relative risk or comparable measures to quantify associations between exposure(s) and outcome(s); they also have the advantage of being able to demonstrate that exposure preceded outcome – an important component of causality.5, 7, 19 Cohort studies have disadvantages including some or all of the following issues: resource requirements are often high and long follow-up times may be necessary especially for less common conditions; also there is a risk of loss of animals during the course of the study.7 Retrospective cohort studies are
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Epidemiology Applied to Horse Reproduction also dependent on the quality of historical information. A range of different examples of cohort studies are provided in the reference list. A number of recent papers have described breeding-related outcomes in generally non-random samples of broodmares.39–42 The studies also provide variable levels of statistical analyses and often relatively large sample sizes, perhaps ameliorating the potential selection biases inherent in the study designs and allow reasonable extrapolation of results to broader target populations. Three hospital-based studies43–45 also contain relatively large numbers of subjects. Inclusion in these studies was dependent on complete medical records with some risk of bias resulting from omission of data from foals with incomplete records. It does seem reasonable, however, to conclude that the animals included in each study are likely to be representative of the target populations – either foals hospitalized at each of two locations43, 44 or draught horses presented for castration at the third location.45 Additional examples of cohort studies involved selection of participating farms using known disease status46 or what seemed to be a convenience approach by enrolling farms that the co-authors were providing reproductive services to,47 and then making judgments about whether the findings can be generalized to broader populations. Case–control studies involve selection of a group of animals affected with the outcome of interest (cases) and a group of animals that are unaffected (controls) and then comparing the frequency of exposure to putative risk factors between the two groups.48 Control animals should reflect the pattern of exposure in the source population from which the cases were derived and should be sampled independently of exposure status.36 Useful discussion on this issue can be found in Rothman5 and Dohoo et al.48 An important distinction between case–control and cohort studies is that cohort studies have the capacity to completely define a denominator (animals at risk of developing the outcome) critical for estimation of incidence rate/risk and measures of association based on incidence such as the relative risk.5 In contrast, case–control studies are really only sampling from the denominator and as a result do not enumerate a denominator and consequently do not allow estimation of prevalence or incidence of the outcome(s). Case–control studies do allow estimation of odds ratios as a measure of association between the outcome and putative exposure(s). Case–control studies are very widely used in veterinary epidemiology, are efficient, and are particularly useful for investigation of rare diseases where a cohort study may be logistically difficult and expensive, and for initial screening of putative risk factors for a given
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disease.7, 49 Case–control studies have a number of potential disadvantages, including the inability to estimate incidence and risk, their limitation to a specific outcome, their frequent reliance on historical records, and difficulty in selecting appropriate controls.7 Two examples of case–control studies involved an animal-level50 and a farm-level51 investigation of risk factors for Rhodococcus equi infection of horses. The first study appeared to enroll a convenience sample of two farms from a number of farms that the authors had previously been working with, and then sampled all foals on the enrolled farms.50 The second study used a more extensive sampling design with an initial letter to all member veterinarians of the American Association of Equine Practitioners residing in the USA to solicit participation followed by further discussion with respondents.51 The authors provide a useful discussion of the potential biases associated with the design. A final example employed formal random selection of cases and controls from eligible populations of sera submitted to a diagnostic laboratory to test associations between exposure to Neospora caninum and risk of abortion.52 A number of hybrid designs have been described in the literature including, for example, case– cohort, nested case–control, and case–crossover studies. Readers are referred to epidemiology texts for more information on these designs.1, 5 Experimental studies are those where the investigator randomly assigns animals to groups where different groups may receive different exposures or treatments in order to test specific hypotheses concerning the effects of the experimental manipulation. Experimental studies tend to be the most expensive and challenging type of study to perform but also may produce the highest quality of evidence to support an association between exposure and outcome.7 Example studies include a double-blinded trial involving administration of one of two commercial hyperimmune plasma products to eligible foals at a university hospital,53 and a field trial involving random allocation of mares to receive different doses of human chorionic gonadotropin (hCG) to induce ovulation at the first ovulation of the breeding season.54
Disease outbreaks Epidemiological principles are particularly important in the investigation and initial response to a disease outbreak, particularly where the etiology of the outbreak may not be readily apparent. A brief review of the key features of an outbreak response is considered relevant for reproductive clinicians involved in care of animals on breeding farms. More detailed information on field and outbreak investigations can be found in Martin et al.,3 an excellent text on field
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epidemiology,55 and in published articles.56 Examples of recent publications detailing outbreak investigations in horses also provide useful examples of the application of epidemiological approaches.57–59 Disease outbreak investigation typically requires rapid investigation and response under an environment of uncertainty. The major objectives are generally to halt the progression of the disease, identify reasons or causes, and implement control measures to prevent or minimize further occurrence and/or eradicate the disease.3 A key focus in the initial response is to collect data and information to describe the events – what animals are getting sick, where and when has disease occurred, and what are the signs and progression of the disease.55 It is important to apply a systematic approach to the investigation particularly where the etiology of the disease is not readily apparent and to note that disease outbreaks may be due to infectious or noninfectious agents.55, 56 The following steps attempt to provide a logical framework for responding to an outbreak, recognizing that activities in an outbreak may simultaneously address multiple steps at the same time.55, 56
1. Verify the diagnosis. Review descriptions of affected animals, results of clinical examinations and laboratory testing. In some outbreaks a diagnosis may not be confirmed for some time. 2. Define a case and count cases. A relatively simple and robust definition of a case should be developed as early as possible in the response using information from observations and tests performed on affected animals. The case definition can then be used to distinguish cases and non-cases and to collect data describing the frequency and distribution of cases. 3. Confirm the existence of an outbreak. It is important to consider the question of whether an outbreak is really occurring. This generally involves comparing the estimated number of cases that meet a case definition with the expected number of cases. This generally requires both a numerator (number of cases) and a denominator (number of animals at risk), and estimation of an attack rate (number of new cases since onset of outbreak divided by the number of animals at risk). 4. Describe cases in terms of time, place, and affected animals. Describing the outbreak using a variety of different approaches is very helpful in understanding the nature of the disease being encountered. An epidemic curve is a plot of new cases over time (hours, days, weeks) to show the temporal pattern. Epidemic curves can help to differentiate point source outbreaks from propagating outbreaks, and may provide information on
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possible incubation periods and whether the outbreak is continuing to increase in size or is starting to decline. The spatial pattern of the outbreak is also important and may involve manually placing dots on a map or using more sophisticated computer-based mapping software. Spatial patterns may provide additional clues to the source of the outbreak, nature of exposure, and transmission mechanisms. It is also helpful to look at patterns of disease in different groups of animals, for example, species, age group, sex, breed, and by groups categorized by other factors that may be suggested by initial observations of the affected site (barn, paddock, feed, water supply, etc.). Formulate a working hypothesis. The combination of case definition, disease progression, and descriptions of the pattern of disease by time, place, and animal should allow preliminary conclusions to be made about which animals are at highest risk, whether the etiology is likely to be infectious or non-infectious, the nature of exposure, and a list of differential diagnoses. Compare the hypothesis with available facts. It is important to compare aspects of the working hypothesis with the available data and results from various activities undertaken as part of the investigation. This generally involves a combination of descriptive and relatively simple analytic epidemiology such as comparisons of attack rates in different groups of animals to test hypotheses. Implement control measures. Interim control measures should be implemented as soon as possible based on an initial assessment of the possible etiological agents that could be involved and the nature and severity of disease. Measures based on general precautionary principles may be implemented initially and then modified as more information is available through the initial investigation. Control measures are likely to be based on principles of infection control and may include isolation of affected animals, movement controls (of animals and people), disinfection, decontamination, and other biosecurity measures. More information may be found in recent publications60–66 and in emergency disease preparedness plans, which are commonly accessed via the internet. If there is considered to be a risk of a non-infectious etiology then control measures may include efforts to prevent further exposure to a non-infectious agent, such as moving animals from the site where cases are occurring, and checking and potentially changing feed sources and access to water or other possible toxins.67 Plan more systematic study if required. Depending on the nature of the outbreak there may be a need for additional investigations to detect cases
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Epidemiology Applied to Horse Reproduction better, determine the geographical extent of spread, identify factors contributing to spread of the disease, test effectiveness of different control measures, and evaluate diagnostic methods. 9. Report findings. A detailed report should be written as quickly as possible that documents the investigation, summarizes findings, and makes recommendations.
Summary Epidemiological methods and related approaches such as evidence-based medicine offer valuable tools for clinicians in addressing real-world problems encountered in equine reproductive practice and in making optimal decisions at both the individual and group levels. Epidemiologists and clinicians are urged to initiate collaborative approaches to clinical problems that should improve our ability to make effective clinical decisions and respond to challenges of the future.
References 1. Dohoo I, Martin SW, Stryhn H. Veterinary Epidemiologic Research. Charlottetown: AVC Inc., 2003. 2. Smith RD. Veterinary Clinical Epidemiology. Boca Raton: CRC Press Inc., 2005. 3. Martin SW, Meek AH, Willeberg P. Veterinary Epidemiology: Principles and Methods. Ames: Iowa State University Press, 1987. 4. Thrushfield M. The scope of epidemiology. In: Thrushfield M (ed.) Veterinary Epidemiology. Oxford: Butterworth & Co., 2005; pp. 22–33. 5. Rothman KJ. Epidemiology: An Introduction. New York: Oxford University Press, 2002. 6. Gay JM. Theriogenology and developments in epidemiology: future opportunities? Theriogenology 2006;66:526–33. 7. Fosgate GT, Cohen ND. Epidemiological study design and the advancement of equine health. Equine Vet J 2008;40:693–700. 8. Martin SW, Meek AH, Willeberg P. Epidemiologic concepts. In: Martin SW, Meek AH, Willeberg P (eds) Veterinary Epidemiology: Principles and Methods. Ames: Iowa State University Press, 1987; pp. 3–21. 9. Woodward M. Epidemiology: Study Design and Data Analysis. Chapman and Hall, 1999. 10. Miettinen OS. Epidemiology: quo vadis? Eur J Epidemiol 2004;19:713–8. 11. Khaw KT. Comment on Miettinen: Rose revisited. Eur J Epidemiol 2004;19:745–8. 12. Sackett DL. Clinical epidemiology: what who and whither. J Clin Epidemiol 2002;55:1161–6. 13. Fletcher RH, Fletcher SW. Clinical Epidemiology: the Essentials. Lippincott Williams and Wilkins, 2005. 14. Sackett DL, Rosenberg WM, Gray JAM, Haynes RB, Richardson WS. Evidence based medicine: what it is and what is isn’t. Brit Med J 1996;312:71–2. 15. Mayer D. Essential Evidence-based Medicine. Cambridge: Cambridge University Press, 2004.
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16. Kastelic JP. Critical evaluation of scientific articles and other sources of information: an introduction to evidencebased veterinary medicine. Theriogenology 2006;66:534–42. 17. Weinberg CR, Wilcox AJ. Reproductive epidemiology. In: Rothman KJ, Greenland S (eds) Modern Epidemiology. Philadelphia: Lippincott-Raven Publishers, 1998; pp. 585–608. 18. Pearce N. A Short Introduction to Epidemiology. Wellington: Massey University, 2005. 19. Dohoo I, Martin SW, Stryhn H. Introduction to observational studies. In: Dohoo I, Martin SW, Stryhn H (eds) Veterinary Epidemiologic Research. Charlottetown: AVC Inc., 2003; pp. 139–49. 20. Altman DG, Schulz KF, Moher D, Egger M, Davidoff F, Elbourne D, Gotzsche PC, Lang T. The revised CONSORT statement for reporting randomised clinical trials: explanation and elaboration. Ann Intern Med 2001;134:663–94. 21. Zwarenstein M, Treweek S, Gagnier JJ, Altman DG, Tunis S, Haynes B, Oxman AD, Moher D. Improving the reporting of pragmatic trials: an extension of the CONSORT statement. Brit Med J 2008;337:1223–6. 22. Bossuyt PM, Reitsma JB, Bruns DE, Gatsonis PP, Glasziou PP, Irwig LM, Lijmer JG, Moher D, Rennie D, De Vet HC. Standards for Reporting of Diagnostic Accuracy. Toward complete and accurate reporting of studies of diagnostic accuracy: the STARD initiative. Ann Intern Med 2003;138:40–4. 23. Des Jarlais DC, Lyles C, Crepaz N. Improving the reporting quality of nonrandomized evaluations of behavioral and public health interventions: the TREND statement. Am J Public Health 2004;94:361–6. 24. von Elm E, Altman DG, Egger M, Pocock SJ, Gotzsche PC, Vandenbroucke JP. Strengthening the reporting of observational studies in epidemiology (STROBE) statement: guidelines for reporting observational studies. Brit Med J 2007;335:806–8. 25. Vandenbroucke JP, von Elm E, Altman DG, Gotzsche PC, Mulrow CD, Pocock SJ, Poole C, Schlesselman JJ, Egger M. Strengthening the Reporting of Observational Studies in Epidemiology (STROBE): Explanation and elaboration. Ann Intern Med 2007;147:W163–94. 26. Boden LA, Parkin TDH. Current guidelines on good reporting of analytical observational studies in epidemiology. Equine Vet J 2008;40:84–6. 27. Carluccio A, Contri A, Tosi U, De Amices I, De Fanti C. Survival rate and short-term fertility rate associated with the use of fetotomy for resolution of dystocia in mares: 72 cases (1991–2005). J Am Vet Med Assoc 2007;230:1502–5. 28. Arnold CE, Payne MThompson JA, Slovis NM, Bain FT. Periparturient hemorrhage in mares: 73 cases (1998–2005). J Am Vet Med Assoc 2008;232:1345–51. 29. Traub-Dargatz JL, Garber LP, Fedorka-Cray PJ, Ladely S, Ferris KE. Fecal shedding of Salmonella spp by horses in the United States during 1998 and 1999 and detection of Salmonella spp in grain and concentrate sources on equine operations. J Am Vet Med Assoc 2000;217:226–30. 30. Hullinger PJ, Gardner IA, Hietala SK, Ferraro GL, MacLachlan NJ. Seroprevalence of antibodies against equine arteritis virus in horses residing in the United States and imported horses. J Am Vet Med Assoc 2001;219:946–9. 31. Fiala SM, Amaral MG, Pimentel CA, Rosa PD, Costa GG, Oliveira GD, Ceretta R, Mattos RC. Inflammatory lesions in the mare’s oviducts. Anim Reprod Sci 2006;94:265–7. 32. Brown JA, Mapes S, Ball BA, Hodder ADJ, Liu IKM, Pusterla N. Prevalence of equine herpesvirus-1 infection among Thoroughbreds residing on a farm on which
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48.
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the virus was endemic. J Am Vet Med Assoc 2007;231: 577–80. Locatelli-Dittrich R, Dittrich JR, Richartz RRTB, Gasino Joineau ME, Antunes J, Pinckney RD, Deconto I, Hoffman DCS, Thomaz-Soccol V. Investigation of Neospora sp. and Toxoplasma gondii antibodies in mares and in precolostral foals from Parana State, Southern Brazil. Vet Parasitol 2006;135:215–21. Rogers CW, Gee EK, Firth EC. A cross-sectional survey of Thoroughbred stud farm management in the North Island of New Zealand. N Z Vet J 2007;55:302–7. Barton MH, LeRoy BE. Serum bile acids concentrations in healthy and clinically ill neonatal foals. J Vet Intern Med 2007;21:508–13. Rothman KJ. Types of epidemiologic study. In: Rothman KJ (ed.) Epidemiology: An Introduction. New York: Oxford University Press, 2002; pp. 57–61. Kaplan RM, Klei TR, Lyons ET, Lester G, Courtney CH, French DD, Tolliver SC, Vidyashankar AN, Zhao Y. Prevalence of anthelmintic resistant cyathostomes on horse farms. J Am Vet Med Assoc 2004;225:903–10. Dohoo I, Martin SW, Stryhn H. Cohort studies. In: Dohoo I, Martin SW, Stryhn H (eds) Veterinary Epidemiologic Research. Charlottetown: AVC Inc., 2003; pp. 139–49. Perkins NR, Grimmett JB. Pregnancy and twinning rates in Thoroughbred mares following the administration of human chorionic gonadotropin (hCG). N Z Vet J 2001;49:94–100. Morris LHA, Allen WR. Reproductive efficiency of intensively managed Thoroughbred mares in Newmarket. Equine Vet J 2002;34:51–60. Langlois B, Blouin C. Statistical analysis of some factors affecting the number of horse births in France. Reprod Nutr Dev 2004;44:583–95. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of Flatrace and National Hunt Thoroughbred mares and stallions in England. Equine Vet J 2007;39: 438–45. Bedenice D, Heuwieser W, Solano M, Rand W, Paradis MR. Risk factors and prognostic variables for survival of foals with radiographic evidence of pulmonary disease. J Vet Intern Med 2003;17:868–75. Rohrbach BW, Buchanan BR, Drake JM, Andrews FM, Bain FT, Byars DT, Bernard WV, Furr MO, Rose Paradis M, Lawler J, Giguere S, Dunkel B. Use of a multivariable model to estimate the probability of discharge in hospitalized foals that are 7 days of age or less. J Am Vet Med Assoc 2006;228:1748–56. Shoemaker R, Bailey J, Janzen E, Wilson DG. Routine castration in 568 draught colts: incidence of evisceration and omental herniation. Equine Vet J 2004;36:336–40. Duarte PC, Conrad PA, Barr BC, Wilson WD, Ferraro GL, Packham AE, Carpenter TE, Gardner IA. Risk of transplacental transmission of Sarcocystis neurona and Neospora hughesi in California horses. J Parasitol 2004;90:1345–51. Vest DJ, Cohen ND, Berezowski CJ, Morehead JP, Blodgett GP, Blanchard TL. Evaluation of administration of West Nile virus vaccine to pregnant broodmares. J Am Vet Med Assoc 2004;225:1894–7. Dohoo I, Martin SW, Stryhn H. Case-control studies. In: Dohoo I, Martin SW, Stryhn H (eds) Veterinary Epidemiologic Research. Charlottetown: AVC Inc., 2003; pp. 139–49. Martin SW, Meek AH, Willeberg P. Surveys and analytic observational studies. In: Martin SW, Meek AH, Wille-
50.
51.
52.
53.
54.
55. 56. 57.
58.
59.
60.
61.
62.
63. 64. 65.
66. 67.
berg P (eds) Veterinary Epidemiology: Principles and Methods. Ames: Iowa State University Press, 1987; pp. 149–75. Chaffin MK, Cohen ND, Martens RJ, Edwards RF. Foal-related risk factors associated with development of Rhodococcus equi pneumonia on farms with endemic infection. J Am Vet Med Assoc 2003;223:1791–9. Cohen ND, O’Connor MS, Chaffin MK, Martens RJ. Farm characteristics and management practices associated with development of Rhodococcus equi pneumonia in foals. J Am Vet Med Assoc 2005;226:404–13. McDole MG, Gay JM. Seroprevalence of antibodies against Neospora caninum in diagnostic equine serum samples and their possible association with fetal loss. Vet Parasitol 2002;105:257–60. Peek SF, Semrad S, McGuirk SM, Riseberg A, Slack JA, Marques F, Coombs D, Lien L, Keuler N, Darien BJ. Prognostic value of clinicopathologic variables obtained at admission and effect of antiendotoxin plasma on survival in septic and critically ill foals. J Vet Intern Med 2006;20:569–74. Grimmett JB, Perkins NR. Human chorionic gonadotropin (hCG): the effect of dose on ovulation and pregnancy rate in Thoroughbred mares experiencing their first ovulation of the breeding season. N Z Vet J 2001;49:88–93. Gregg MB. Field Epidemiology. New York: Oxford University Press, 2002. Kane AJ, Morley PS. How to investigate a disease outbreak. P Annu Conv Am Equin 1999;45:137–41. Irwin VL, Traub-Dagartz JL, Newton JR, Scase TJ, DavisPoynter NJ, Nugent J, Creis L, Leaman TR, Smith KC. Investigation and management of an outbreak of abortion related to equine herpesvirus type 1 in unvaccinated ponies. Vet Rec 2007;160:378–80. More SJ, Aznar I, Bailey DC, Larkin JF, Leadon DP, Lenihan P, Flaherty B, Fogarty U, Brangan P. An outbreak of equine infectious anaemia in Ireland during 2006: Investigation methodology, initial source of infection, diagnosis and clinical presentation, modes of transmission and spread in the Meath cluster. Equine Vet J 2008;40:706–8. More SJ, Aznar I, Myers T, Leadon DP, Clegg TA. An outbreak of equine infectious anaemia in Ireland during 2006: The modes of transmission and spread in the Kildare cluster. Equine Vet J 2008;40:709–11. Lunn DP, Traub-Dagartz J. Managing infectious disease outbreaks at events and farms; challenges and the resources for success. P Annu Conv Am Equin 2007;53:1–12. Newton JR. Aspects of equine infectious disease control from the United Kingdom perspective. P Annu Conv Am Equin 2007;53:13–21. Traub-Dagartz JL, Dagartz DA, Morley PS, Dunowska M. An overview of infection control strategies for equine facilities, with an emphasis on veterinary hospitals. Vet Clin N Am-Equine 2004;20:507–20. Smith BP. Evolution of equine infection control programs. Vet Clin N Am-Equine 2004;20:521–30. Dwyer RM. Environmental disinfection to control equine infectious diseases. Vet Clin N Am-Equine 2004;20:531–42. Weese JS. Barrier precautions, isolation protocols and personal hygiene in veterinary hospitals. Vet Clin N Am-Equine 2004;20:543–59. Bender JB, Tsukayama DT. Horses and the risk of zoonotic infections. Vet Clin N Am-Equine 2004;20:643–53. Schmitz DG. Toxicologic problems. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine. St Louis: Saunders, 2004; pp. 1441–512.
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Chapter 296
Management of the Geriatric Mare Scott Madill
Since aging changes are physiological in nature and progress at an individually variable rate, there is no well-defined age at which a horse becomes geriatric, but in the general equine population it is suggested to occur around the early twenties.1 Beyond direct reproductive issues, these horses have increased prevalence of diseases affecting the gastrointestinal, musculoskeletal, respiratory, and endocrine systems, and these can exert secondary effects on reproductive performance. For the practitioner involved with breeding management, the onset of age-related reductions in fertility of mares becomes evident in the mid to late teens. This chapter focuses on preparation of older mares for the breeding season and management of their breeding cycles. As with any horse, breeding management of the older mare needs to be tailored to the individual case. While some readily become pregnant and carry the foal to term, breeding others can be an exercise in frustration and disappointment for the veterinarian, owner, and farm manager. This is especially true when members of this group have not been involved in breeding aged mares previously. Having the involved personnel fully apprised of the likelihood of success at each stage of the operation and of the potential costs involved from the start will go a long way to ensuring that the team maintains a good working relationship through the season and beyond. The information used in these informational and planning conversations comes from published fertility data and the collective experience of the group, supplemented with the history and physical examination of the mare concerned.
General expectations Older mares are less fertile and require more intense management and greater veterinary interven-
tion than their younger counterparts for a number of reasons including: delayed entry to the breeding season, increased length of the estrous cycle, reduced response to drugs for ovulation induction, poor oocyte quality, increased incidence of infectious and persistent breeding-induced endometritis (PBIE), degenerative changes of the endometrium, and increased embryonic and fetal losses. Research pertaining to the pathophysiology of many of these differences has been reviewed previously.2 Published differences in reproductive parameters between old and young mares are tabulated through the text, starting with Table 296.1, and these can provide a useful reference for client communication. While the fertility figures presented from some older references may be lower than what would be currently desired for both young and old mares, the differences between the two groups remain pertinent. As a starting point it has been noted that aged mares required breeding on 4.6 cycles to produce a foal compared to 2.0 cycles for young mares.3
Pre-breeding examination of the mare The aim of this exam is to identify both existing problems that will affect fertility and any predispositions to such problems. Accurate diagnosis permits a reasonable prognosis to be made and timely institution of appropriate therapy so that the mare is presented for breeding with maximal chance of conception and subsequent pregnancy maintenance. In barren mares the exam should ideally be conducted before the end of the physiological breeding season so there is sufficient time for therapy to be effective prior to the onset of the next season. Details of the exam techniques are presented in other chapters and only items
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mare: General Interest Table 296.1 General fertility expectations for breeding the aged mare Parameter
Type of mating
Young mare
Aged mare
Pregnancy rate per cycle (%)
Natural matinga Fresh AIb Cooled AIc Frozen AIc Natural matingd Natural matinge Natural matingf Fresh AIg
57–67 50–83 60–63 43–48 9–39 46–57 75–87 85
32–55 19–40 46 42 24–59 0–25 43–63 53
Cycles with uterine treatment given (%) Foaling rate per cycle (%) Foaling rate per season (%)
a
Sanderson and Allen;112 Morris and Allen;113 Bosh;114 Allen et al.28 Vanderwall et al.;3 Carnevale and Ginther;108 Squires et al.35 c Squires et al.35 d Morris and Allen;113 Allen et al.28 e Badi et al.;115 Baker et al.116 f Badi et al.;115 Jeffcott et al.;117 Sanderson and Allen;112 Morris and Allen;113 Bosh;114 Allen et al.28 g Vanderwall et al.3 b
particularly pertinent to aged mares are presented in the following section. History All available history on the mare’s general health, current and former use, breeding management, and foaling history, and any prior evaluations and treatments for reproductive issues, should be obtained. This can help direct the subsequent examination and provides useful prognostic information. Physical exam A general physical exam is performed to obtain an overall impression of the mare’s health, with particular attention given to the teeth, coat condition, body condition, and muscle distribution, and evidence of significant musculoskeletal disease. Flattening of the croup should alert the clinician to potential problems with perineal conformation, urine pooling, and delayed uterine clearance. The mare should be in body condition of 5–6 (on a 1–9 scale) with normal epaxial muscle and fat distribution. Condition of the coat is interpreted in relation to season and housing. Beyond ill-thrift this part of the exam is particularly looking for signs of endocrinopathies that may adversely affect fertility in the older mare, and these are presented in a later section. Evidence of severe or recurrent musculoskeletal disease should prompt an assessment of how likely the mare is to carry a pregnancy to term. Where breed regulations allow, mares with conditions such as chronic laminitis and severe suspensory desmitis are likely better directed to assisted reproductive techniques. When such options are not available or desired by the client, the welfare aspects of breeding an affected mare should be frankly discussed.
Reproductive exam Conformation of the perineal region is assessed in relation to the mare’s body condition, and this area and the inner thighs and tail are examined for evidence of vaginal discharge. The lips of the vulva are assessed for thinning or distortion and fibrosis from previous injury or surgery. The vulvar lips are parted to assess aspiration of air. The reproductive tract is then assessed by palpation and ultrasound per rectum. In older pluriparous mares the perineal body is often thinned to the extent that it is essentially absent, and it is occasionally helpful to examine it between the fingers of one (sterile gloved) hand in the vagina and those of the other placed rectally. Severe pneumovagina is palpable as a dilated viscus in the pelvic cavity that terminates at the cervix. The tone and size of the cervix are assessed in relation to stage of cycle and parity of the mare. While minor cervical lacerations are not palpable per rectum, severe injuries are occasionally found. Uterine examination in the open mare assesses whether the uterus is normally positioned or sagging into the abdomen, whether there are local areas of dilation and thinning of the uterine wall near the horn base (ventral sacculations), the tone of the uterus and its appropriateness for stage of the estrous cycle, evidence of intrauterine fluid or air, endometrial cysts and local masses in the lumen, wall, or broad ligaments. With the popularity of marble placement in performance mares to control the estrous cycle, the clinician should be on the lookout for their presence on ultrasound. The ovaries are assessed for size, normal contour, and evidence of cyclicity. During the exam a quick assessment of the pelvic canal for evidence of old fractures or space-occupying lesions that may interfere
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Management of the Geriatric Mare with passage of the fetus is also made. In older gray mares with perineal melanomas, masses may also be found in the pelvic area. In the author’s experience these generally only grow slowly from season to season and pre-breeding assessment has not yet led to an unpleasant surprise at foaling. Following thorough cleansing and drying of the perineum a vaginoscopy is performed. Lack of resistance to passage of the vaginoscope through the vestibulo-vaginal junction is an indicator of poor barrier function in this area. Pathological changes include inflammation of mucous membranes (secondary to pneuomovagina, endometritis, and urine pooling), adhesions in the vagina or distorting the cervix, and severe cervical lacerations. Some changes may be more evident at certain stages of the cycle, such as mares that only pool urine during estrus or those that fail to relax the cervix at the same stage. A sterile sleeve is donned and a manual exam of the vagina and cervix is performed. To avoid contamination this is generally done after sampling for endometrial culture. The digital exam concentrates on determining function of the cervix. Minor lesions are best assessed when the mare is under the influence of progesterone, and mares presented in the transition or anestrous seasons or with a relaxed cervix can be reassessed after 3 days of treatment with altrenogest or progesterone. Remarkable improvement in the delineation of a cervical laceration can occur after such therapy. Uterine culture and cytology should be performed on all older barren mares, any mare with indications of endometritis on the preceding exam (vaginal discharge, pneumovagina, intrauterine fluid, and physometra), or those with a suspicious history. While the optimum time of the cycle to perform a culture is still subject to some debate, it is probably preferable to culture older mares during estrus when their uterine defense mechanisms are better able to deal with any iatrogenically introduced organisms. In barren mares, especially where previous cultures taken by guarded swabs have been negative, use of a low-volume lavage technique will maximize the sensitivity of the procedure.4 Mares with a history of unexplained infertility, chronic endometritis that is refractory to treatment, or palpation and ultrasound findings suggestive of localized uterine pathology may benefit from hysteroscopic examination to identify or rule out lesions such as intrauterine adhesions and fetal remnants. Despite the shortcomings of current endometrial biopsy assessment and categorization techniques, obtaining a biopsy can be useful in guiding the management of some geriatric mares and provides the client with at least a broad prognosis.5, 6 Where breed registration allows, mares with severe degenerative
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changes to the endometrium are best directed to techniques allowing a surrogate to carry the foal (embryo transfer, oocyte transfer, and intracytoplasmic sperm injection (ICSI)).
Preparation of the mare for the breeding season Once the decision to breed the mare is made, factors adversely affecting her general health and fertility are treated as far in advance of the breeding season as practical. Dental abnormalities are corrected as needed and the mare scheduled for more frequent routine floating as she ages. The ration is checked for nutrient balance and quantities are adjusted to take the mare to the desired body condition. Body condition scoring should be continued on a regular basis to monitor success and the need for further adjustments. Surgery to ameliorate defects in perineal conformation, vesicovaginal reflux, and cervical function are performed. More invasive procedures are generally completed 6–8 weeks before breeding to permit healing. Infectious endometritis is treated as soon as possible to allow resolution of inflammation prior to breeding. Placement of a temporary Caslick may be required to prevent contamination but also allow intrauterine treatments to be performed. Foaling mares that had a Caslick opened prior to foaling should have it replaced soon after parturition to minimize uterine contamination and inflammation.7 Since entry of aged mares into the breeding season is delayed by approximately 2 weeks, lighting treatments will need to start correspondingly earlier if breeding at the start of the season is desired.8 For the sake of expediency it can be useful to examine such mares 2–3 weeks prior to the start of the season, and those displaying lack of advanced follicular development can be started on additional treatments such as sulpiride. Treatment details for this and hormones useful for enhancing passage through transition are detailed in other chapters. The client should be made aware that beyond 20 years of age mares will begin to have irregular cycles and eventually may cease to cycle altogether.9 Anecdotal reports indicate that treatment of such aged mares with gonadotropin-releasing hormone (GnRH) agonists or follicle-stimulating hormone (FSH) is frequently unsuccessful in stimulating follicle development.
Endocrinopathies of the older mare: Cushing’s, hypothyroidism, and equine metabolic syndrome. Equine pituitary pars intermedia dysfunction (Cushing’s disease) is common in aged horses, with a
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recent survey suggesting a prevalence of approximately 14% in those 15 years and older.10 While large surveys of fertility in well-documented cases are not available, the condition is widely stated to adversely affect cyclicity and fertility in affected mares, and the clinical experience of many practitioners attests to this. Early signs of disease include slow or incomplete shedding when exposed to stimulatory photoperiod, and loss of muscle mass along the spine. Other signs of disease include lethargy, permanent hirsuitism, polyuria and polydipsia, excessive sweating, laminitis, and a propensity to develop infections. Available laboratory tests perform poorly, especially early in the disease, and the diagnosis is best made on clinical signs.11 Effects on the reproductive system include failure to cycle or cycle abnormalities, endometritis, infertility, and inappropriate lactation.12 Affected horses are generally treated with the dopamine agonist pergolide mesylate, with dosages adjusted based on clinical response and laboratory testing. Treatment should commence as soon as possible after diagnosis as clinical response and return to cyclicity may take 6–8 weeks.12, 13 Many older mares have been placed on thyroxine supplements in an effort to improve fertility following a diagnosis of hypothyroidism based on clinical signs and laboratory tests. This area has recently been subjected to critical review, highlighting the nebulous nature of the condition in horses, difficulty in its diagnosis, and questionable effects on reproductive function.14 In a very limited number of young mares, thyroidectomy did not alter patterns of estrous cyclicity or fertility.15 Larger trials have found no relationship between baseline serum thyroxine, serum triiodothyronine, or thyrotropinreleasing hormone (TRH) response tests and 15–16day pregnancy rates in mixed-age broodmares, nor was there an effect of thyroid hormone supplementation.16, 17 In light of this, any improvements in fertility of aged cycling mares placed on thyroid supplements are attributed mainly to general metabolic enhancement.14 Most horses previously diagnosed with hypothyroidism on the basis of clinical signs and low circulating thyroxine levels are now considered to have equine metabolic syndrome.18 These animals do not have the pars intermedia dysfunction of equine Cushing’s disease but are in some respects similar – exhibiting insulin resistance and characteristically maintaining body condition on little feed, with either general obesity or fat deposition to the crest and adjacent the tailhead. Insulin resistance also makes them more prone to repeated bouts of laminitis.18 Reproductively, mares with insulin resistance have prolonged interovulatory intervals, and this appears to be associated with development of luteinized anovulatory follicles.19, 20 Horses with the syndrome
may also be more prone to developing inflammatory responses as their neutrophils are primed for increased oxidative burst activity.21 Whether this has a role in increasing susceptibility to or severity of postbreeding endometritis in these mares is unknown. The relative merits of available diagnostic tests for insulin resistance have been comprehensively reviewed.22 Treatment strategies for this syndrome have been outlined and are based around reduction in bodyweight and improving insulin sensitivity by basing the diet on hay with low available sugars combined with a protein and mineral balancer. Other sources of sugars such as grain and sweet feeds are removed, and access to high-sugar pastures is limited.18 Ideally these management changes and the desired weight changes are achieved before the breeding season commences so weight loss is not occurring during breeding. The main concerns are mares that present with the syndrome during the breeding season. Normal horses placed on long-term thyroxine supplementation show weight losses that are maintained over time, presumably due to increased basal metabolic rate.23 Thyroxine treatment may be a viable strategy in affected broodmares where time is limited and concurrent weight loss without adversely affecting fertility is desired, but this remains to be critically tested. Removal of endometrial cysts Older mares have a higher prevalence of endometrial cysts than younger mares but their effect on fertility and thus the need for removal is debated. Potentially, cysts can affect fertility by restricting embryo migration and thus maternal recognition of pregnancy or by depriving the embryo of nutrition following fixation, but both theories have been questioned. Eilts and co-workers found mares over 11 years were 4.2 times more likely to have cysts than younger mares, and that most of these cysts were located near the site of early embryo fixation at the base of the uterine horns. In a logistic analysis they were unable to demonstrate that the cysts had any adverse effects on fertility, though it is worth noting that their average maximal cyst diameter was only 20 mm.24 Our current approach to cysts is similar to that described by Holyoak and Ley.25 Mares with large cysts or groups of cysts that are near the site of embryo fixation at the base of the uterine horns, or those that have very large cysts elsewhere that might affect embryo migration, are examined by hysteroscopy during diestrus and the cysts laser ablated. Following the procedure prostaglandin is administered and the uterus is lavaged with several liters of balanced electrolyte solution. Lavage is repeated for several days until the effluent is clear, then mares are given a month’s rest prior to breeding. Clients need
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Management of the Geriatric Mare to be aware that cysts are associated with endometrial degeneration, and removal of the cyst will obviously not improve the histological categorization of the endometrium.26 It is rare for an older mare with endometrial cysts to have a histological grade of better than IIB on the Kenney and Doig scale.27 While this might imply limited value of performing a biopsy prior to treatment in such mares, the prognostic value in a mare with severe degenerative changes can help a wavering client decide whether pursuing cyst treatment and further breeding is economically sensible.
Choice of stallion and semen type Choice of which stallion the mare is booked to and the resultant semen options is generally made well in advance of breeding. The innate fertility of the stallion, or his cooled or frozen semen, has a large bearing on pregnancy rate of the mare. This is especially true when breeding older subfertile mares, where using a subfertile sire almost guarantees a poor result. When breeding with two or more stallions will produce a foal of approximately equal economic or emotional value, then ideally the sire with the highest pregnancy rate per cycle would be chosen. Unfortunately fertility of various sires is rarely publicly available to guide these decisions, and making recommendations based on personal experience may be legally fraught. Individual stallions are also associated with different rates of embryonic loss, but again for obvious reasons this information is generally not disclosed.28 In general terms, fresh semen has superior pregnancy rates per cycle to cooled semen, which in turn is slightly superior to frozen semen.29, 30 In practice good conception rates are attainable with all types of semen in fertile mares given good breeding management, and the practitioner will generally be dealing with the type of semen already chosen by the client. It is the author’s practice to counsel against the use of frozen semen in older mares with a history of infertility, especially those where the infertility is related to chronic problems with persistent mating-induced endometritis. The reasoning is twofold: firstly, combining the slightly lower fertility rate of frozen semen with that of an infertile mare can only worsen pregnancy rate; secondly, frozen semen has been shown to result in a greater uterine inflammatory response, exacerbating the problem in an already susceptible mare.31
Management of breeding Since every cycle on which a mare is bred unsuccessfully likely increases her risk of either becoming
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infected or developing non-infectious persistent endometrial inflammation, her best chance of conceiving may be the first cycle on which she is bred. Every effort must be made to maximize the chance of the mare conceiving on this cycle. This obviously is the aim for all mares, but is particularly important for the aged mare, especially if she has a history of infertility related to infectious endometritis or delayed uterine clearance. Various palpation, ultrasonographic, and hormonal parameters have been explored for their reliability in predicting first ovulation of the season.32 None of these are 100% reliable and it is the author’s preference not to breed older mares until they have already had at least one ovulation for the season. This ensures breeding is not occurring on a transition follicle, no matter how promising that follicle looks, avoiding pointless uterine contamination and inflammation. Timing and frequency of breeding While in the general mare population there are studies indicating superior pregnancy results with multiple breedings per cycle, others find no such advantage over a single well-timed insemination.33–38 Since many older mares are susceptible to PBIE, the author’s preferred approach, and that of others, is a single breeding in order to minimize uterine contamination and the resultant inflammatory response.39, 40 For fresh semen this generally implies aiming to breed within 24–48 hours before ovulation, for cooled semen within 24 hours before ovulation, while for frozen semen the preferred window is 12 hours before to 6 hours after ovulation.34,41–44 To aid in prediction of ovulation and timing of induction agents, it is preferable that the mare be examined in early estrus so progression can be followed. This avoids the situation where a mare is presented with no history, a reasonably large follicle, and little uterine edema; in such cases it is not always easy to tell if the mare is in early estrus and has several days to develop, or has already experienced her endogenous luteinizing hormone (LH) surge and will be ovulating in the next 24–36 hours. Similarly, mares to be short-cycled with prostaglandin should be examined on the day of drug administration with particular attention paid to follicle size as this parameter will determine how rapidly progression toward ovulation will occur and direct the timing of future exams. In general, once the mare has good uterine edema and a follicle > 35 mm in diameter, an ovulation-inducing agent such as human chorionic gonadotropin (hCG), deslorelin, or recombinant LH is administered. For fresh and cooled semen, arrangements have been made so that the mare can be bred 24 hours later, which in most cases will be
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12–24 hours prior to ovulation. In the author’s practice, older mares bred to frozen semen are examined at 6-hour intervals, starting at 12–24 hours following administration of hCG or deslorelin, and more frequently when ovulation is imminent, to ensure breeding occurs once within 6 hours post-ovulation. This is usually aided by timing administration of hCG or deslorelin for late night so most ovulations occur during the day. In a multicenter study, Samper and co-workers found that only 26.6% of mares bred with frozen semen prior to ovulation had a single breeding.45 While a single breeding was not the aim in many of these mares, it highlights the difficulty of predicting ovulation with certainty, even when using ovulation-inducing treatments and frequent examination. If the mare does not ovulate within 24–48 hours of breeding with fresh or cooled semen the decision on further inseminations depends on the mare’s history of delayed clearance and evidence of such on the ovulation check, the availability and quality of a second semen dose, and any information on the longevity of sperm from the stallion in question following insemination. Using fresh semen, the mean lifespan of sperm from artificial insemination (AI) to fertilization in one study was 2.3 days.46 Also with fresh semen, Woods and colleagues obtained pregnancy rates of 65% for inseminations occurring 2 days prior to ovulation, 60% for 3 days prior, 33% for 4 days prior, 63% for 5 days and 44% for 6 days, compared to 89% when insemination occurred 1 day before ovulation. 41 While such results are probably highly stallion dependent and similar studies have not been reported for cooled semen, mares with signs or history of significant post-breeding fluid accumulation are not rebred within 48 hours of first insemination. An alternate strategy has been suggested by some authors breeding mares susceptible to persistent endometritis. When using good quality fresh or cooled semen, breeding is deliberately performed 48 or 72 hours prior to anticipated ovulation. This allows more time for post-breeding treatments to occur prior to elevations of progesterone and its accompanying adverse effects on uterine defenses.47, 48 In practice, with variable semen quality the author does not deliberately aim for this approach; however, mares that show fluid accumulation post-breeding are not rebred within 48 hours of initial insemination. Fluid accumulations prior to breeding Mares with free fluid accumulations during estrus but prior to breeding have lower pregnancy rates than those without.49 Approximately 75% of such estrus fluid accumulations are not associated with inflammation or infection but 25% are, and affected
mares are viewed with suspicion and investigated further for infection, especially if fluid depth is 2 cm or greater.49 Mares with intraluminal fluid accumulation during estrus of greater than 2 cm in depth are also more likely to have problems with postmating endometritis and should be targeted for early and aggressive post-insemination therapy.50 Excessive edema during estrus is also considered by some clinicians to be an indicator of infection.51 Even when cytology shows that significant intrauterine fluid accumulations (> 1–2 cm in depth) are non-inflammatory, it has been recommended that they be removed prior to breeding, either by oxytocin therapy or catheter drainage.48 Although there are no data on the efficacy of this approach when the fluid is non-inflammatory in nature, it is the general policy adhered to by the author. However, if oxytocin is to be used (20 units i.m.) it should probably be given at least 2 hours before insemination, given the potentially adverse affect of peri- insemination administration on pregnancy rates.52 Failure of ovulation The incidence of failure of spontaneous ovulation during an estrus in mares over 16 years of age was 13.1%, which was three times higher than the 4.4% reported for 6–10-year-old mares.53 The fact that the mean interovulatory interval in affected mares was 38.5 days and that 43.5% of mares experiencing one anovulatory follicle would experience another within the same season means that considerable time and opportunities for conception can be lost. Whether administering ovulation-inducing agents as part of normal breeding management reduces the incidence of these anovulatory follicles has not been directly tested. A separate study by the same group found that only 6.5% of mares over 15 years of age failed to ovulate in response to hCG.54 While it is not valid to compare across the studies, 6.5% is a preferable rate to 13.1%, and until disproven by direct analysis it is reasonable policy to administer an ovulationinducing agent to older mares as part of normal management. Even in cases when ovulation still fails, such administration appears to make development of a luteinized anovulatory structure more likely than a follicular one, and the former are subsequently amenable to prostaglandin therapy. However, once a persistent follicular structure is established, administration of hCG or deslorelin appears ineffective.39, 53 Response to ovulation-inducing agents Reliable timing of ovulation in response to ovulationinducing agents such as hCG and deslorelin greatly facilitates the breeding management of mares. In
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Management of the Geriatric Mare many instances decisions on semen ordering and stallion booking are made based on ovulation occurring between 24 and 48 hours after these agents are administered. Older mares, in addition to the total failure of ovulatory response described above, are more likely than younger mares to ovulate beyond 48 hours after administration of such a drug. Failure to ovulate within 48 hours of hCG occurs in approximately 19–20% of cycles in 16–20-year-old mares and 26% of cycles in mares over 20 years, compared to 5–14% of cycles in mares under 10 years.54, 55 Other groups have found no effect of age on mean time to ovulation after hCG, but at least one of these studies had very limited numbers of mares in the older categories.37, 56 It has been suggested that at least part of the explanation for the poorer response to hCG in older mares is known or unknown prior exposure to the drug.54 A recent retrospective study confirms that response to hCG administration diminishes with repeated dosing from year to year.56 While ovulatory response to deslorelin implants appeared slightly lower, at 84% in mares ≥ 20 years in one study, no such reduction was found in response to compounded deslorelin in another, with 96% ovulating within 48 hours.57, 58 Taken together these results suggest that deslorelin or a similar GnRH analog may be a more reliable choice in aged mares, bypassing potential problems with prior hCG exposure. Sperm transport defects Reduced fertilization rate in old mares may be partly due to defects in sperm transport. Compared to normal mares, mares susceptible to chronic endometritis had significantly fewer sperm in the oviduct at 4 hours after insemination with 500 million progressively motile sperm; few of the sperm present were motile, and motile sperm were never found in both oviducts of susceptible mares.59 In contrast, no difference in oviductal sperm numbers was found between normal and susceptible mares at 18 hours postinsemination with 500 million total sperm deposited in either the uterine body or proximal uterine horn.60 While the studies are not directly comparable, it is interesting to note that a difference existed at 4 hours but not at 18 hours, perhaps suggesting delayed transport in susceptible mares. If so the results may have more importance for management of susceptible mares bred to frozen semen, where lifespan of spermatozoa is curtailed and insemination frequently occurs after ovulation, allowing little time for the fertilizing sperm population to become established in the oviduct. Similarly, while it is well established that uterine lavage delayed 4 or more hours after breeding will not adversely affect conception rates in re-
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productively normal mares,61 the innocuous nature of this timing would benefit from further experimental study in highly susceptible mares with potential delays in sperm transport. Potential practical methods of improving sperm transport in susceptible mares include treatments to improve myometrial contraction and deposition of sperm on or adjacent to the oviductal papilla. Intrauterine infusion of 5 mg of prostaglandin F2α (PGF2α ) immediately after insemination tended to improve single cycle pregnancy rates compared to saline-treated controls for barren (42.9% vs 23.1%) and older (50.0% vs 40.0%) mares.62 Deposition of 0.2 mg of PGE2 at the proximal end of the uterine horn ipsilateral to the dominant follicle 2 hours prior to sperm deposition improved pregnancy rates for both uterine body and deep-horn insemination with 25 million total sperm in one trial using normal mares.63 Treatment with oxytocin immediately after insemination of 100 million sperm into the uterine body failed to improve pregnancy rates of normal mares bred to fertile or subfertile stallions (testing the hypothesis that oxytocin would improve sperm transport of subfertile males). Indeed, mares treated with oxytocin had a 30% pregnancy rate compared to 50% for mares given placebo, and the authors hypothesized that this early oxytocin administration may have caused premature evacuation of the inseminate in treated mares and reduced sperm colonization of the oviduct.52 If mare category was ignored, deep-horn insemination of 500 million sperm in a 10-mL volume resulted in a doubling of sperm number recovered from the oviduct ipsilateral to ovulation compared to uterine body insemination.60 Probably due to lack of statistical power, this difference was not significant when mares were split into normal and susceptible categories. Taking these results into account, the author’s current approach is to use a deep-horn insemination technique in aged mares with a flaccid uterus or one with ventral sacculations, but not to administer an ecbolic agent at breeding.
Post-breeding management The intensity of post-breeding management depends on the history of the mare and findings of pre- and post-breeding examinations.
Persistent breeding-induced endometritis An understanding of the pathophysiology of PBIE and available treatment options is of vital importance to the practitioner dealing with older broodmares,
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and the topic is dealt with at length elsewhere in this volume. The problem is one of failure of the physical clearance mechanisms that normally remove excess semen and bacteria introduced at breeding.64 Old mares have a higher incidence and severity of post-breeding fluid accumulation than younger mares.65 The incidence varies with the population under study and the timing of the examination relative to insemination. At 12–18 hours post-insemination with frozen semen, mares aged over 16 years had a 68% incidence of intrauterine fluid accumulation compared to 28.2% in 10–16-year-olds and 17% in those 3–9 years of age.66 While it can be argued that examining mares at this time, coincident with the maximal inflammatory response, identifies more mares than those with clinically important problems; studies have identified reduced pregnancy rates in animals affected at this early stage.66, 67 Factors that predispose older mares to increased incidence and severity of this problem include alterations in uterine position and development of ventral uterine sacculations with multiple foalings, loss of normal slope of the pelvis (flattening of the croup), reduced uterine contractility, and failure of cervical dilation in older maiden mares.5, 64, 68 Prostaglandins appear to play an important role in stimulating myometrial contractions that expel excess semen and inflammatory products from the uterus following breeding.40 Older mares are frequently treated with non-steroidal anti-inflammatory drugs for a variety of chronic musculoskeletal issues; the clinician should be alert to these mares as phenylbutazone administration has been shown to delay uterine clearance.69
Lavage and ecbolics The key to treatment is early identification of affected mares and return of the uterine environment to normal by evacuation of fluid and inflammatory products.70 Even though normal mares can have intrauterine fluid at 6 hours following insemination with fresh semen and at 12 hours following the use of frozen semen as a physiological response,40 mares with a known history of repeated fluid retention or those that had fluid prior to breeding are examined at 6 hours post-breeding and a lavage performed at that time if fluid is present. Susceptible mares treated with lavage at 4–6 hours after insemination appear to have higher pregnancy rates than those where treatment is delayed until the following day.47 It is the author’s preference to lavage these highly susceptible mares even if only minimal fluid is present. In the past some of these cases treated with ecbolics only have been found to have large amounts of intrauterine fluid when re-examined 24 hours later.
All mares are examined approximately 24 hours post-breeding to ensure ovulation has occurred (fresh and cooled semen) and to determine whether fluid has been retained. Fluid retention is treated with ecbolics or a combination of lavage and ecbolics. Lavage is performed with sterile physiological saline or a balanced electrolyte solution (lactated Ringer’s solution) in 1–2-liter volumes and repeated until the effluent is clear. In mares with particularly large flaccid uteri where fluid recovery has proven difficult, administration of 20 units of oxytocin intramuscularly prior to preparation for the lavage can be helpful in fluid recovery. Mares are rechecked the following day and lavage is repeated as required up to 3 days post-ovulation. Of the readily available ecbolic agents, oxytocin is superior to cloprostenol in causing rapid evacuation of uterine fluid.71 Oxytocin is the drug of choice for treatment of PBIE and is generally given at a dose of approximately 20 units either i.m. or i.v. Oxytocin has also been shown to overcome delayed clearance in mares treated with phenylbutazone.69 The problem with oxytocin is its relatively short duration of action, which means repeat dosing at 4–6-hour intervals in more severely affected mares. While this can be manageable in hospital situations it can be more difficult in ambulatory practice. Prostaglandins are the readily available alternative ecbolics and have the advantage of prolonged duration of action of around 5 hours.71 Cloprostenol was superior to prostaglandin F2α and fenprostalene when compared in mares with delayed uterine clearance, and thus is the preferred analog for this use.72 Doses of cloprostenol range from 250 to 500 µg i.m. or s.c. and are typically given at 12–24hour intervals.71 Most clinicians use the lower end of this dose range. Cloprostenol treatment administered on days 0–2 post-ovulation has been consistently shown to decrease progesterone levels up to Day 5, and in some studies has decreased pregnancy rates compared to saline- or oxytocin-treated controls.73–75 For this reason the author prefers oxytocin in most cases and restricts cloprostenol use to a few mares and where possible only prior to ovulation. There are no data on whether exogenous progestagen supplementation of mares given cloprostenol post-ovulation will ameliorate the adverse affect on pregnancy. Longer-acting analogs of oxytocin such as carbetocin are available in some countries and their prolonged action may give the benefit of ease of dose schedule in practice without adverse affects on development of the corpus luteum when used post-ovulation, but this requires confirmation.5, 76, 77 LeBlanc5 has also advocated the use of acupuncture in old mares intractable to standard treatments.
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Management of the Geriatric Mare Cervical dilation Mares that fail to relax the cervix during estrus are associated with decreased fertility because they either prevent sperm access to the uterus in natural breeding situations or they develop intractable PBIE following artificial insemination. Affected mares may have cervical fibrosis secondary to dystocia, delivery of a large foal, or repeated embryo recovery attempts over several years; or may just fail to relax what appears to be an otherwise normal cervix, a situation typically seen in old maiden mares or those that were bred when young but have then gone on to an athletic career and not been bred for many years.5 Affected mares requiring natural mating due to breed registration requirements should have reinforcement breeding (insemination) with semen collected from the vagina immediately following the natural mating where this is permitted. Mares with failure of cervical relaxation tend to develop PBIE that is more difficult to treat than that from other causes. Various therapies have been recommended to aid dilation of the cervix including gentle manual dilation, prostaglandin E1 (misoprostil) 200–1000 µg as a tablet or cream placed in the cervical canal or on the external os, prostaglandin E2 (2 mg) as a softened tablet placed in the cervical canal, 17β-estradiol at a dose of 10 mg given i.m. for 2 days prior to breeding, and repeated doses of prostaglandin F2α analogs daily for several days prior to breeding.5, 40, 78, 79 Of the pharmacological interventions the use of prostaglandin E analogs is the most popular combined with aggressive lavage and use of ecbolics. These cases can be extremely frustrating to treat.
Exercise Exercise, though not critically tested, is likely highly beneficial for uterine clearance; hence, turnout rather than stalling is the recommended housing for mares prone to PBIE.80 The intensity and duration of exercise required to effect uterine clearance is unknown but personal observation would suggest that even 10–20 minutes of brisk hand-walking twice daily can be beneficial.
Antibiotic infusions and systemic antibiotic therapy Post-breeding infusion of mares with antibiotics has long been a mainstay of studfarm medicine. The practice has fallen out of favor over the last 15 years in academic medicine, with numerous studies pointing to the efficacy of lavage and ecbolic treatments described above in treating the condition of PBIE. How-
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ever, it should be remembered that ‘susceptible’ older mares are prone to both post-breeding inflammation and infectious endometritis, and that breeding provides an avenue for deposition of bacteria along with spermatozoa into these susceptible uteri.40, 81, 82 Canadian researchers have provided evidence suggesting that severe post-insemination inflammation that persists for up to 96 hours in susceptible mares is due to bacterial infection rather than sperm antigens.83 Mares with ‘hyperedema’ at the time of breeding or that maintain a relatively high degree of edema following subsequent ovulation have been shown to have a 57% chance of being positive on culture and/or cytology and have reduced per cycle pregnancy rate.51 In a large field trial, treating mares with systemic oxytocin and intrauterine infusion of antibiotics had an additive beneficial effect on pregnancy rates compared to either treatment alone.49 When using infusions it should be remembered that a mixed population of bacteria rather than a single species are introduced at the time of breeding and infusion with a narrow-spectrum antibiotic will not control all potential pathogens.81 In the author’s practice there is a subset of older mares where antibiotics are used as part of breeding management. These mares have histories of being highly susceptible to developing infectious endometritis, manifest as either repeated bouts of infectious endometritis without breeding or a diagnosis of endometritis at the time of pregnancy examination at around Day 14 despite aggressive lavage and ecbolic treatment on the previous breeding cycle. Once these mares have been treated successfully (returned a clean swab) their breeding management is similar to that described by Asbury and Lyle84 and involves some or all of the following. Prior to insemination, lavage, or infusion, particular attention is paid to cleansing the clitoral fossa and sinuses with a chlorhexidine scrub in addition to normal thorough cleaning of the vulva and perineal area. The aim of this is to minimize the introduction of bacteria to the uterus from this reservoir at the time of these manipulations.85 The mare undergoes uterine lavage at 6 hours post-insemination and then as indicated by daily ultrasound examinations. In these mares lavage is followed by infusion with a broad-spectrum antibiotic or antibiotic combination. Most often this involves 1–2 g of ceftiofur sodium in 50–60 mL of physiological saline, though infusion volume may be increased in multiparous mares with large uteri. Oxytocin treatment is used to aid in fluid expulsion but doses are not started until at least 6 hours after antibiotic infusion. For mares that are known to be extremely susceptible, systemic antibiotics are also employed. Approximately 2 days prior to anticipated breeding the mares
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are placed on a potentiated sulfonamide at a dose of 24–30 mg/kg b.i.d. p.o. This combination is chosen because: (i) it is known to give good endometrial levels following systemic administration;86 most of the beta-hemolytic streptococci and coliforms cultured from cases of endometritis in the author’s practice are sensitive to it – these organisms are most likely to be involved in endometritis;4 and (iii) it is also relatively cheap and in most cases easy to administer. This therapy is continued for at least 5 days post-breeding and occasionally until pregnancy exam is performed at Day 14 post-ovulation. This systemic administration provides therapy beyond the time uterine infusions and flushes can safely be employed post-ovulation, avoids contamination of the uterus, and may help clear organisms introduced by the local therapeutic interventions. The approach has not been critically tested.
Modulating the uterine inflammatory response Since PBIE represents an ongoing inflammatory response that in highly susceptible mares can be difficult to control with standard treatments of lavage and ecbolic therapy, anti-inflammatory agents have occasionally been employed as part of an aggressive treatment regimen. These might help break the cycle of inflammation and further fluid production in these mares, giving the clinician a better chance of effectively managing these extremely challenging cases. The main options available are non-steroidal antiinflammatory agents (NSAIDs), corticosteroids, and, potentially, seminal plasma. Large controlled trials assessing the efficacy of these therapies are lacking. NSAIDs have been shown to be capable of inhibiting prostaglandin release from the uterus in response to a variety of insults such as infusion of acidic solutions and uterine biopsy.87 Phenylbutazone has been administered concurrently with antibiotics to mares with persistent fluid accumulation around the time of breeding and up to 2 weeks in the post-ovulatory period.88 As mentioned earlier in the section on PBIE, mares treated with these drugs are likely to have delayed uterine clearance as a result and will require concurrent ecbolic therapy. Corticosteroid administration has recently shown great promise as adjunctive therapy for mares with PBIE. Dell’Aqua and co-workers89 used prednisolone acetate at the relatively low dose of 0.1 mg/kg every 12 hours for five treatments, with the final dose given at the time of insemination. This protocol significantly increased pregnancy rates following breeding with frozen semen in 15 barren mares with histories of PBIE. In another study, a single dose of 50 mg of dexamethasone i.v. administered immedi-
ately prior to insemination significantly improved pregnancy rates in mares with three or more risk factors indicating susceptibility to PBIE.90 This treatment was also beneficial for mares with the single risk factors of post-breeding fluid accumulation of > 2 cm or post-breeding fluid accumulation that persisted for > 36 hours. These treatments are not used alone, but in conjunction with uterine lavage and ecbolic treatments as part of normal case management. In this vein, LeBlanc5 reported administration of 10 mg of dexamethasone i.v. at 24 hours postinsemination in mares with cervical fibrosis as part of an aggressive post-breeding treatment regimen of lavage, ecbolics, and antibiotic infusions. While prednisolone is considered a safe drug to administer, the potential for development of laminitis following dexamethasone administration remains unresolved.91, 92 None of the mares in the study of Bucca and co-workers90 had an adverse response to their single intravenous treatment of 50 mg, and this is similar to the dose commonly used in other conditions such as recurrent airway obstruction.93 However, caution should probably be exercised in administering dexamethasone to horses that have a history of laminitis or factors predisposing to its development, and fully informed client consent should be obtained prior to commencing treatment.91 Interest in seminal plasma was sparked due to the reported increased uterine inflammatory response to frozen-thawed semen and concentrated fresh semen, both of which had very low levels of seminal plasma. 31 In most studies seminal plasma has appeared to have anti-inflammatory effects that protect viable sperm in the inflammatory uterine environment and improve fertility (reviewed by Troedsson40 ). Potential clinical use of seminal plasma awaits identification and purification of the involved protein factors and clarification of their effects.
Immunostimulants Mares that suffer repeated infectious endometritis following breeding may benefit from the use of immunostimulant therapy around the time of breeding. In a controlled experiment, treatment of susceptible mares with a mycobacterial cell-wall extract (SettleTM , Bioniche Animal Health) by either intravenous or intrauterine infusions hastened clearance of experimentally induced Streptococcus zooepidemicus infection.94 In a controlled field trial, mares treated intravenously with another immunostimulant based on Propionibacterium acnes (EqStimTM , Neogen Corp, Lexington, KY) were more likely to become pregnant and deliver a live foal than placebo-treated controls. Treatment was administered as three doses with
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Management of the Geriatric Mare follow-up doses given 2 and 6 days after the first. Greatest beneficial effect on pregnancy rates occurred when mares were bred between 2 days before and 8 days after the first dose.95 Progesterone supplementation Supplementation of mares with progesterone following ovulation is a controversial area, where the science and art of veterinary practice are often at odds. The evidence for a primary lack of progesterone production (luteal inadequacy) as an important cause of embryonic loss is scant and there is less to suggest it is more of a problem in aged mares. Ginther and co-workers found lower progesterone levels on Days 7 and 11 post-ovulation in mares that suffered embryonic loss between 11 and 15 days of pregnancy. These mares had no ultrasonographically detectable intrauterine fluid, but the authors suspected that lower progesterone in this group was a result of premature luteolysis caused by chronic uterine inflammation, rather than being a primary luteal insufficiency.96 Further into pregnancy, Irvine and coworkers found that of 17 cases of natural embryonic loss between Days 17 and 42, in only one was the loss definitely preceded by a drop in progesterone level. In the majority of cases (14 of 17) the progesterone level remained high until well after the loss had occurred.97 Notwithstanding these findings, the author uses supplemental progesterone occasionally in the following situations. The first is where the mare continues to be open on pregnancy examination at Day 12–14 despite excellent semen quality and breeding management, and there is no evidence of endometritis or other pathology. In this case supplementation with altrenogest is generally started 2 days after ovulation. These mares are often also infused with antibiotics post-breeding and may be placed on systemic antibiotics (potentiated sulfonamide) given the adverse affect of progesterone on uterine defense mechanisms. This approach is generally tried to ‘give the mare a last chance’ either late in the breeding season prior to further diagnostic investigation or prior to commencing embryo flushing or sending her for other advanced reproductive techniques. The second is the mare that gets pregnant and habitually loses the pregnancy, again in the absence of apparent endometritis or other pathology such as cervical laceration. Also in this category is the mare with a normal sized embryonic vesicle but poor cervical or uterine tone, or one with evidence of endometrial edema at the time of pregnancy exam. The third situation is when the client insists on it. In all cases the pros and cons of supplementation are discussed so the client can make an informed de-
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cision. This discussion includes the lack of great evidence of effect, the fact that therapy will be ineffective for many or most causes of the problem such as embryonic defects and oviductal abnormalities and may delay diagnosis of some conditions, the potential for endometritis, the cost of treatment, the need to decide if and when supplementation will be discontinued if the mare becomes pregnant, and the need for additional pregnancy monitoring since the mare will not return to heat in the face of pregnancy loss and thus rebreeding will be delayed and she may potentially retain a non-viable fetal remnant. Clients almost always choose to supplement.
Pregnancy and the peripartum period Expectations for the remainder of pregnancy and puerperal period have been reviewed previously and are presented in Table 296.2.2 Mares with perineal conformation issues that were managed by breeding stitch or temporary Caslick during breeding have a more permanent surgery performed following completion of post-breeding treatments or after pregnancy examination. It is important that this is performed as these mares are also more prone to ascending placentitis. Further pregnancy exams are scheduled. As with all mares an exam around 35 days is recommended; mares open at this stage can be Table 296.2 Expectations for aged mares during pregnancy and the peripartum period Parameter
Young mare
Aged mare
Fertilization failure or EED < 14 daysa
0–13%
49–73%
Embryonic death 14–50 daysb
6–14%
21–33%
Abortion 50 days–termc
2–10%
6–29%
Gestation lengthd
Tendency to increase
Dystociae
No increased risk f
Uterine artery rupture Foal IgG < 800 mg/dL
g
5× greater risk 30%
Foal morbidity and mortality < 14 daysh a
Badi et al.;115 Ball et al.118 Woods et al.;119 Bosh.114 c Platt;120 Badi et al.;115 Vanderwall et al.3 d Platt.121 e Ginther and Williams;122 Frazer et al.123 f Douglas.124 g LeBlanc et al.125 h Morley and Townsend.126 EED, early embryonic death.
b
55% Linear increase with mare age
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rebred while those pregnant at this time that subsequently lose the pregnancy are generally not rebred due to formation of the endometrial cups. Geriatric mares are usually treated no differently as far as scheduled vaccinations, deworming, and general health maintenance is concerned beyond accommodations for diet formulation, dental care, and chronic musculoskeletal conditions. Clients are reminded of the higher expected incidence of embryonic death and abortion, the signs of placentitis, and the importance of early assessment of the foal due to increased incidence of a number of neonatal problems including musculoskeletal disorders and failure of passive transfer.
Use of assisted reproductive techniques When dealing with an infertile mare, assisted reproductive techniques that can appear prohibitively expensive initially may actually prove to be the better and more economical option where they are available and when breed society regulations permit their use. Taking the low-cost approach in a mare with a history of infertility can mean multiple breeding and perhaps embryo flushing attempts for a comparable outlay and no result. In such cases early consultation with the reproductive center providing such services can be very useful in guiding mare management. Reviews detailing selection of the appropriate technique for specific fertility problems are available.98, 99 Descriptions of the procedures themselves are provided in other chapters; this section will deal solely with success rates in older mares, which are presented in Table 296.3. Ultimately, beyond the techniques dealt with below, cloning of the valuable aged mare may permit continuation of her genetics by using the resultant clones for reproduction, and indeed this is already being done.
Embryo transfer Embryo transfer is useful in older mares with a recent history of good fertility and in those that have no problem conceiving but subsequently lose the pregnancy. Mares that fail to ovulate; have documented or suspected oviductal blockages or salpingitis; adhesions within the uterus or around the ovaries, oviducts, and uterus; chronic endometritis or pyometra; large defects of the cervix, or consistent severe PBIE are poor candidates.98, 99 Similarly, while embryo transfer can be useful diagnostically in mares with unexplained infertility, repeated failure to recover embryos from such mares bred with quality semen indicates the need for another approach.
Table 296.3 Effect of donor mare age on results of assisted reproductive technologies Young mare
Aged mare
Embryo transfer Embryo recoverya (%) Pregnant recipientb (%) Embryonic lossa (%)
61–67 70–80 1–14
30–43 56–76 9–30
Oocyte transfer Pregnant recipientc (%) At least one pregnancy/seasond (%)
39–92 80
16–40 62
ICSI Oocytes recovered/session – diestruse Oocytes recovered/session – estrusf Oocytes maturing to MII in vitroe (%) Blastocyst formation following ICSIef (%) Pregnancies per embryo transferredf (%)
10 0.74–1.08 66 13–42 33–50
10 0.86–1.08 67 12–24 40–60
a
Vogelsang and Vogelsang;103 Meadows et al.102 Vogelsang and Vogelsang;103 Carney et al.;127 Meadows et al.102 c Carnevale and Ginther;108 Carnevale et al.107 d Carnevale et al.107 e Colleoni et al.,111 aspirated all follicles 5–40 mm. f Altermatt et al.,110 aspirated mature follicles. ICSI, intracytoplasmic sperm injection. b
When looking in the literature at the effect of donor age on recovery rates there is obvious confounding between the effect of age per se and that of infertility (there are more infertile older mares). This reinforces the need to assess each mare individually as a potential donor when advising the client. In general terms embryos are recovered from only 30% of flushes performed on aged mares compared to 60–70% of those performed on young fertile mares (Table 296.3).100 Embryos recovered from aged mares are also delayed in development and are correspondingly smaller than those recovered on the same day from younger mares, and appear to have delayed oviductal transport to the uterus. For this reason it is suggested that flushes be scheduled on Day 8 rather than Day 7 to maximize recovery rates.101, 102 Most studies have found no difference in initial pregnancy rates following transfer of embryos obtained from fertile young mares compared to those from infertile aged mares. However, in the study of Vogelsang and Vogelsang,103 embryos derived from donors aged 9–17 years and 18–28 years had significantly lower rates of pregnancy following transfer (52% and 56% respectively) compared to those derived from donors 2–8 years old (70%). In ideal conditions with highly fertile donor mares a recovery rate of 70% and a pregnancy rate following transfer of 70% results in approximately a 50% chance of establishing a pregnancy on a given cycle; however, an old infertile mare with a recovery rate of 30% and a
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Management of the Geriatric Mare pregnancy rate after transfer of 55% gives only a 16.5% chance of pregnancy per cycle in the recipient. A consistent finding across studies is that pregnancies established with embryos derived from older mares also have much higher rates of embryonic death by 50–60 days of gestation, ranging from 9% up to 50% if old mares with histories of infertility are included104 (Table 296.3). This is due to intrinsic embryonic defects, reflected in delayed development of the embryo when it is recovered.105 Oocyte transfer Oocyte transfer can be used to bypass many of the fertility problems that render an old mare unsuitable as an embryo transfer donor. These include failure of ovulation, chronic endometritis, intractable PBIE, pyometra, cervical scarring or severe lacerations, and unexplained infertility. Oocytes were recovered from 80–96% of cycles and 70–76% of pre-ovulatory follicles over several years in a commercial program with a large component of old infertile mares.106, 107 The average quality of aspirated oocytes was worse in older mares but this was not predictive of pregnancy rate, and pregnancy rate following transfer was not different for oocytes from mares ≥ 20 years (40%) compared to those < 20 years (39%). However, greater age eventually becomes a factor, with oocytes from donors < 15 years having a 50% pregnancy rate compared to 16% for those from mares > 23 years.107 This is in keeping with earlier experimental studies where transfer of oocytes from young donors resulted in extremely high pregnancy rates (92%) compared to those from old donors (31%), and shows that the technique cannot overcome intrinsic oocyte defects that are due to aging.108 Older donor mares are thus likely to require more cycles to establish a pregnancy and tend to be less likely to end the season with at least one pregnant recipient.106, 107 Overall success of the procedure on a per cycle basis would be approximately 32% (80% oocyte recovery × 40% pregnancy after transfer) but will be lower in the oldest mares. Intracytoplasmic sperm injection Delayed development of embryos from aged mares with resultant increased embryonic losses may be due to alterations in oocyte maturation resulting in delayed fertilization. In a slaughterhouse study, Brinsko and co-workers found oocytes derived from the ovaries of mares ≥ 15 years had delayed development in culture compared to those from younger mares.109 While intracytoplasmic sperm injection (ICSI) is more complex than oocyte transfer and usu-
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ally recommended more often for overcoming male infertility, it has the advantages in the aged mare of potentially overcoming these fertilization problems and also allows transfer of only those oocytes known to have been successfully fertilized and capable of at least early embryonic development.98, 99 Recent studies using oocytes obtained by aspiration of either the dominant follicle(s) during estrus or of immature follicles during diestrus suggest the technique is capable of overcoming at least some oocyte abnormalities of older mares.110, 111 These studies found no significant effect of mare age on oocyte recovery, oocyte morphology, maturation success, cleavage following the ICSI procedure, blastocyst formation following in vitro culture, or pregnancy rates following transfer into recipients. However, raw numbers in the study of Altermatt and co-workers indicate a possible affect on blastocyst formation rate in donors over 20 years that may bear further study.
References 1. Paradis MR. Demographics of health and disease in the geriatric horse. Vet Clin N Am-Equine 2002;18:391–401. 2. Madill S. Reproductive considerations: mare and stallion. Vet Clin N Am-Equine 2002;18:591–619. 3. Vanderwall DK, Peyrot LM, Weber JS, Woods GL. Reproductive efficiency of the aged mare. In: Proceedings of the Annual Convention of the Society for Theriogenology 1989, pp. 153–6. 4. LeBlanc MM, Magsig J, Stromberg AJ. Use of a lowvolume uterine flush for diagnosing endometritis in chronically infertile mares. Theriogenology 2007;68:403–12. 5. LeBlanc MM. Reproduction – clinical cases. P Annu Conv Am Equine 2006;52:585–90. 6. Schlafer DH. Equine endometrial biopsy: enhancement of clinical value by more extensive histopathology and application of new diagnostic techniques? Theriogenology 2007;68:413–22. 7. Hemberg E, Lundeheim N, Einarsson S. Retrospective study on vulvar conformation in relation to endometrial cytology and fertility in thoroughbred mares. J Vet Med A 2005;52:474–7. 8. Vanderwall DK, Woods GL. Age-related subfertility in the mare. P Annu Conv Am Equine 1990;36:85–9. 9. Carnevale EM, Bergfelt DR, Ginther OJ. Follicular activity and concentrations of FSH and LH associated with senescence in mares. Anim Reprod Sci 1994;35:231–46. 10. McGowan TW, Hodgson DR, McGowan CM. The prevalence of equine Cushing’s syndrome in aged horses. J Vet Intern Med 2007;21:603 [abstr.] 11. McFarlane D. Diagnosing pituitary pars intermedia dysfunction. Comp Cont Educ Pract 2007;2:208–213. 12. McCue PM. Equine Cushing’s disease. Vet Clin N AmEquine 2002;18:533–43. 13. Panzani D, Vannozzi I, Sgorbini M, Corazza M, Rota A, Pacini M. Embryo recovery rate in a mare affected by Cushing’s syndrome. Vet Res Commun 2003;27(Suppl. 1): 619–21.
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14. Brinsko SP. Common procedures in broodmare practice: what is the evidence? Vet Clin N Am-Equine 2007;23:385–402. 15. Lowe JE, Foote RH, Baldwin BH, Hillman RB, Kallfelz FA. Reproductive patterns in cyclic and pregnant thyroidectomized mares. J Reprod Fertil Suppl 1987;35: 281–8. 16. Gutierrez CV, Riddle WT, Bramlage LR. Serum thyroxine concentrations and pregnancy rates 15 to 16 days after ovulation in broodmares. J Am Vet Med Assoc 2002;220:64–6. 17. Meredith TB, Dobrinski I. Thyroid function and pregnancy status in broodmares. J Am Vet Med Assoc 2004;224:892–4. 18. Frank N. Insulin resistance in horses. P Annu Conv Am Equine 2006;52:51–4. 19. Sessions DR, Reedy SE, Vick MM, Murphy BA, Fitzgerald BP. Development of a model for inducing transient insulin resistance in the mare: preliminary implications regarding the estrous cycle. J Anim Sci 2004;82:2321–8. 20. Vick MM, Sessions DR, Murphy BA, Kennedy EL, Reedy SE, Fitzgerald BP. Obesity is associated with altered metabolic and reproductive activity in the mare: effects of metformin on insulin sensitivity and reproductive cyclicity. Reprod Fertil Dev 2006;18:609–17. 21. Holbrook TC, Tipton TE, Yerramsetti PN, McFarlane D. Neutrophil function and inflammatory cytokine expression in horses with equine metabolic syndrome. J Vet Intern Med 2007;21:661 [abstr.]. 22. Firshman AM, Valberg SJ. Factors affecting clinical assessment of insulin sensitivity in horses. Equine Vet J 2007;39:567–75. 23. Frank N, Elliot SB, Boston RC. Effects of long-term oral administration of levothyroxine sodium on glucose dynamics in healthy adult horses. Am J Vet Res 2008;69:76–81. 24. Eilts BE, Scholl DT, Paccamonti DL, Causey R, Klimczak JC, Corley JR. Prevalence of endometrial cysts and their effect on fertility. Biol Reprod Monogr 1995;1:527–32. 25. Holyoak GR, Ley WB. Management regimens for uterine cysts. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Theriogenology. St Louis, MO: Saunders Elsevier, 2007; pp. 121–5. 26. Bracher V, Mathias S, Allen WR. Videoendoscopic evaluation of the mare’s uterus: II. Findings in subfertile mares. Equine Vet J 1992;24:279–84. 27. Kenney RM, Doig PA. Equine endometrial biopsy. In: Morrow DA (ed.) Current Therapy in Theriogenology, 2nd edn. Philadelphia, PA: W.B. Saunders Co., 1986; pp. 723–9. 28. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of flatrace and national hunt thoroughbred mares and stallions in England. Equine Vet J 2007;39:438–45. 29. Jasko DJ, Moran DM, Farlin ME, Squires EL, Amann RP, Pickett BW. Pregnancy rates utilizing fresh, cooled and frozen-thawed stallion semen. P Annu Conv Am Equine 1992;38:649–60. 30. Loomis PR. The equine frozen semen industry. Anim Reprod Sci 2001;68:191–200. 31. Kotilainen T, Huhtinen M, Katila T. Sperm-induced leukocytosis in the equine uterus. Theriogenology 1994;41:629–36. 32. Roser JF, Valcheck DE, Lasley BL, Liu IK, Kojusner G, Taya K. Minimizing the incidence of endometritis by
33.
34.
35.
36.
37.
38. 39. 40. 41.
42.
43.
44.
45.
46.
47.
48.
49.
50.
predicting the first ovulation of the year. Pferdeheilkunde 1997;13:539 [abstr.]. Shore MD, Macpherson M,L, Combes GB, Varner DD, Blanchard TL. Fertility comparison between breeding at 24 hours or at 24 and 48 hours after collection with cooled equine semen. Theriogenology 1998;50:693–8. Squires EL, Brubaker JK, McCue PM, Pickett BW. Effect of sperm number and frequency of insemination on mares inseminated with cooled semen. Theriogenology 1998;49:743–9. Squires E, Barbacini S, Matthews P, Byers W, Schwenzer K, Steiner J, Loomis P. Retrospective study of factors affecting fertility of fresh cooled and frozen semen. Equine Vet Educ 2006;18:96–9. Reger HP, Bruemmer JE, Squires EL, Maclellan LJ, Barbacini S, Necchi D, Zavaglia G. Effects of timing and placement of cryopreserved semen on fertility of mares. Equine Vet Educ 2003;15:101–6. Sieme H, Sch¨afer T, Stout TAE, Klug E, Waberski D. The effects of different insemination regimes on fertility in mares. Theriogenology 2003;60:1153–64. Metcalf ES. Optimizing pregnancy rates using frozenthawed equine semen. Anim Reprod Sci 2005;89:297–9. McCue PM. Clinical cases in equine reproduction. P Annu Conv Am Equine 2006;52:591–6. Troedsson MHT. Breeding-induced endometritis in mares. Vet Clin N Am-Equine 2006;22:705–12. Woods J, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic-loss rate in mares. Equine Vet J 1990;22: 410–15. Metcalf ES. Insemination and breeding management. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. Philadelphia, PA: W.B. Saunders Co., 2001; pp. 195–228. Samper JC. Artificial insemination. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. Philadelphia, PA: W.B. Saunders Co., 2001; pp. 109– 131. Blanchard TL, Thompson JA, Brinsko S, Love C, Varner DD, Frost J, Basson M, Ramsey J. Effects of breeding to ovulation interval and repeat service during the same estrus on pregnancy rates in thoroughbred mares. P Annu Conv Am Equine 2007;53:568–70. Samper JC, Vidament M, Katila T, Newcombe J, Estrada A, Sargeant J. Analysis of some factors associated with pregnancy rates of frozen semen: a multi-center study. Theriogenology 2002;58:647–50. Cl´ement F, Vincent P, Mahla R, Meriaux JC, Palmer E. Which insemination results in fertilization when several are performed before ovulation? J Reprod Fertil Suppl 2000;56:579–83. ¨ Knutti B, Pycock JF, van der Weijden GC, Kupfer U. The influence of early postbreeding uterine lavage on pregnancy rate in mares with intrauterine fluid accumulations after breeding. Equine Vet Educ 2000;12:267–70. Pycock JF. Breeding management of the problem mare. In Samper JC (ed.) Equine Breeding Management and Artificial Insemination. Philadelphia, PA: W.B. Saunders Co., 2001; pp. 195–228. Pycock JF, Newcombe JR. The relationship between intraluminal uterine fluid, endometritis and pregnancy rate in the mare. Equine Pract 1996;18:19–22. Brinsko SP, Rigby SL, Varner DD, Blanchard TL. A practical method for recognizing mares susceptible to
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post-breeding endometritis. P Annu Conv Am Equine 2003;49:363–5. Samper JC. How to interpret endometrial edema in brood mares. P Annu Conv Am Equine 2007;53:571–2. Rigby S, Hill J, Miller C, Thompson J, Varner D, Blanchard T. Administration of oxytocin immediately after insemination does not improve pregnancy rates in mares bred by fertile or subfertile stallions. Theriogenology 1999;51:1143–50. McCue PM, Squires EL. Persistent anovulatory follicles in the mare. Theriogenology 2002;58:541–3. McCue PM, Hudson JJ, Bruemmer JE, Squires EL. Efficacy of hCG at inducing ovulation: a new look at an old issue. P Annu Conv Am Equine 2004;50:510–13. Barbacini S, Zavaglia G, Gulden P, Marchi V, Necchi D. Retrospective study on the efficacy of hCG in an equine artificial insemination programme using frozen semen. Equine Vet Educ 2000;12:312–17. Green JM, Raz T, Epp T, Carley SD, Card CE. Relationships between utero-ovarian parameters and the ovulatory response to human chorionic gonadotrophin (hCG) in mares. P Annu Conv Am Equine 2007;53:563–7. Farquhar V, McCue PM, Vanderwall DK, Squires EL. Efficacy of the GnRH agonist deslorelin acetate for inducing ovulation in mares relative to age of mare and season. J Equine Vet Sci 2000;20:722–5. McCue PM, Magee C, Gee EK. Comparison of compounded deslorelin and hCG for induction of ovulation in mares. J Equine Vet Sci 2007;27:58–61. Scott MA, Liu IKM, Overstreet JW. Sperm transport to the oviducts:abnormalities and their clinical implications. P Annu Conv Am Equine 1995;41:1–2. Rigby S, Derczo S, Brinsko S, Blanchard T, Taylor T, Forrest DW, Varner D. Oviductal sperm numbers following proximal uterine horn or uterine body insemination. P Annu Conv Am Equine 2000;46:332–4. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post insemination on pregnancy rate in mares. Theriogenology 1991;35: 1111–19. ¨ Sieme H, Schroter N, Klug E, Schoon H-A. Influence of prostaglandin F2α on conception rate of mares inseminated with chilled semen. Pferdheilkunde 1997;13:558 [abstr.]. Woods J, Rigby S, Brinsko S, Stephens R, Varner D, Blanchard T. Effect of intrauterine treatment with prostaglandin E2 prior to insemination of mares in the uterine horn or body. Theriogenology 2000;53:1827–36. Troedsson MHT. Uterine clearance and resistance to persistent endometritis in the mare. Theriogenology 1999;52:461–71. Zent WW, Troedsson MHT, Xue J-L. Postbreeding uterine fluid accumulation in a normal population of thoroughbred mares: a field study. P Annu Conv Am Equine 1998;44:64–5. Barbacini S, Necchi D, Zavaglia G, Squires EL. Retrospective study on the incidence of postinsemination uterine fluid in mares inseminated with frozen/thawed semen. J Equine Vet Sci 2003;23:493–6. Watson ED, Barbacini S, Berrocal B, Sheerin O, Marchi V, Zavaglia G, Necchi D. Effect of insemination time of frozen semen on incidence of uterine fluid in mares. Theriogenology 2001;56:123–31. LeBlanc MM, Neuwirth L, Jones L, Cage C, Mauragis D. Differences in uterine position of reproductively normal
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84.
85.
86.
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mares and those with delayed uterine clearance detected by scintigraphy. Theriogenology 1998;50:49–54. Cadario ME, Thatcher M-JD. Relationship between prostaglandin and uterine clearance of radiocolloid in the mare. Biol Reprod Monogr 1995;1:495–500. Troedsson MHT, Scott MA, Liu IKM. Comparative treatment of mares susceptible to chronic uterine infection. Am J Vet Res 1995;56:468–72. LeBlanc MM. Effects of oxytocin, prostaglandin and phenylbutazone on uterine clearance of radiocolloid. Pferdeheilkunde 1997;13:483–5. Coombs GB, LeBlanc MM, Neuwirth L, Tran TQ. Effects of prostaglandin F2α, cloprostenol and fenprostalene on uterine clearance of radiocolloid in the mare. Theriogenology 1996;45:1449–55. Troedsson MHT, Ababneh MM, Ohlgren AF, Madill S, Vetscher N, Gregas M. Effect of periovulatory prostaglandin F2α on pregnancy rates and luteal function in the mare. Theriogenology 2001;55:1891–9. Brendemuehl JP. Effect of oxytocin and cloprostenol on luteal formation, function and pregnancy rates in mares. Theriogenology 2002;58:623–6. Nie GJ, Johnson KE, Wenzel JGW, Braden TD. Effect of periovulatory ecbolics on luteal function and fertility. Theriogenology 2002;58:461–3. Pycock JF. Therapy for mares with uterine fluid. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 93–104. Schramme AR, Pinto CR, Davis JL, Whitacre MD, Whisnant CS. Pharmacokinetics of carbetocin, a long-acting oxytocin analogue, following intravenous administration in horses. Theriogenology 2007;68:492–518. Volkmann DH, Bertschinger HJ, Schulman ML. The effect of prostaglandin E2 on the cervices of dioestrous and prepartum mares. Reprod Domest Anim 1995;30: 240–4. Nie GJ, Barnes AJ. Use of prostaglandin E1 to induce cervical relaxation in a maiden mare with post breeding endometritis. Equine Vet Educ 2003;15:172–4. Causey RC. Uterine therapy for mares with bacterial infections. In: Samper JC, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis, MO: Saunders Elsevier, 2007; pp. 105–15. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: technique and preliminary findings. P Annu Conv Am Equine 1975;21:327–36. Watson ED. Uterine defence mechanisms in mares resistant and susceptible to persistent endometritis: a review. Equine Vet J 1988;20:397–400. Maloufi F, Pierson R, Otto S, Ball C, Card C. Mares susceptible or resistant to endometritis have similar endometrial echographic and inflammatory cell reactions at 96 hours after infusion with frozen semen and extender. P Annu Conv Am Equine 2002;48:51–7. Asbury AC, Lyle SK. Infectious causes of infertility. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, PA: Lea and Febiger, 1993; pp. 381–91. Hinrichs K, Cummings MR, Sertich PL, Kenney RM. Clinical significance of aerobic bacterial flora of the uterus, vagina, vestibule, and clitoral fossa of clinically normal mares. J Am Vet Med Assoc 1988;193:72–5. Brown MP, Gronwall R, Castro L. Pharmacokinetics and body fluid and endometrial concentrations of
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trimethoprim-sulfamethoxazole in mares. Am J Vet Res 1988;49:918–22. Daels PF, Stabenfeldt GH, Kindahl H, Hughes JP. Prostaglandin release and luteolysis associated with physiological and pathological conditions of the reproductive cycle of the mare: a review. Equine Vet J Suppl 1989;8:29–34. McKinnon AO, Voss JL. Breeding the problem mare. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia, PA: Lea and Febiger, 1993; pp. 368–78. Dell’Aqua JA Jr, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Melo CM. Modulation of acute uterine inflammatory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94: 270–3. Bucca S, Carli A, Buckley T, Dolci G, Fogarty U. The use of dexamethasone administered to mares at breeding time in the modulation of persistent mating induced endometritis. Theriogenology 2008;70:1093–100. Johnson PJ, Slight SH, Ganjam VK, Kreeger JM. Glucocorticoids and laminitis in the horse. Vet Clin N AmEquine 2002;18:219–36. Bailey SR, Elliott J. The corticosteroid laminitis story: 2. Science of if, when and how. Equine Vet J 2007;39:7–11. Ainsworth DM, Hackett RP. Disorders of the respiratory system. In: Reed SM, Bayly WM, Sellon DC (eds) Equine Internal Medicine, 2nd edn. St Louis, MO: Saunders Elsevier, 2004; pp. 289–353. Rogan D, Fumuso E, Rodriguez E, Wade J, S´anchez Bruni SF. Use of a mycobacterial cell wall extract (MCWE) in susceptible mares to clear experimentally induced endometritis with Streptococcus zooepidemicus. J Equine Vet Sci 2007;27:112–17. Rohrbach BW, Sheerin PC, Cantrell CK, Matthews PM, Steiner JV, Dodds LE. Effect of adjunctive treatment with intravenously administered Propionibacterium acnes on reproductive performance in mares with persistent endometritis. J AmVet Med Assoc 2007;231:107–13. Ginther OJ, Garcia MC, Bergfelt DR, Leith GS, Scraba ST. Embryonic loss in mares: pregnancy rate, length of interovulatory intervals, and progesterone concentrations associated with loss during days 11–15. Theriogenology 1985;24:409–17. Irvine CHG, Sutton P, Turner JE, Mennick PE. Changes in plasma progesterone concentrations from days 17 to 42 of gestation in mares maintaining or losing pregnancy. Equine Vet J 1990;22:104–6. Hinrichs K, Choi Y-H. Assisted reproductive techniques in the horse. Clin Tech Equine Pract 2005;4:210–18. Coutinho da Silva MA. When should a mare go for assisted reproduction? Theriogenology 2008;70:441–4. Aguilar J, Woods GL. Embryo transfer in horses: indications, technique, and expected outcomes. In: Youngquist RS (ed.) Current Therapy in Large Animal Theriogenology. Philadelphia, PA: W.B Saunders Co., 1997; pp. 208–13. Squires EL, McCue PM, Vanderwall D. The current status of equine embryo transfer. Theriogenology 1999;51:91– 104. Meadows S, Lisa H, Welsh C. Factors affecting embryo recovery, embryo development and pregnancy rate in a commercial embryo transfer programme. Havermeyer Foundation Monograph Series 2000;1:61–2. Vogelsang SG, Vogelsang MM. Influence of donor parity and age on the success of commercial equine embryo transfer. Equine Vet J Suppl 1989;8:71–2.
104. Squires EL, Imel KJ, Iuliano MF, Shideler RK. Factors affecting reproductive efficiency in an equine embryo transfer program. J Reprod Fertil Suppl 1982;32: 409–14. 105. Carnevale EM, Ramirez RJ, Squires EL, Alvarenga MA, Vanderwall DK, McCue PM. Factors affecting pregnancy rates and early embryonic death after equine embryo transfer. Theriogenology 2000;54:965–79. 106. Carnevale EM, Squires EL, Maclellan LJ, Alvarenga MA, Scott TJ. Use of oocyte transfer in a commercial breeding program for mares with reproductive abnormalities. J Am Vet Med Assoc 2001;218:87–91. 107. Carnevale EM, Coutinho da Silva MA, Panzani D, Stokes JE, Squires EL. Factors affecting the success of oocyte transfer in a clinical program for subfertile mares. Theriogenology 2005;64:519–27. 108. Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod Monogr 1995;1:209–14. 109. Brinsko SP, Ball BA, Ellington JE. In vitro maturation of equine oocytes obtained from different age groups of sexually mature mares. Theriogenology 1995;44:461–9. 110. Altermatt JL, Suh TK, Stokes JE, Squires EL, Siedel GE, Carnevale EM. Effects of age and FSH on collection of equine oocytes and developmental competency after intracytoplasmic sperm injection. Theriogenology 2007;68:513–14 [abstr.]. 111. Colleoni S, Barbacini S, Necchi D, Duchi R, Lazzari G, Galli C. Application of ovum pick-up, intracytoplasmic sperm injection and embryo culture in equine practice. P Annu Conv Am Equine 2007;53:554–9. 112. Sanderson MW, Allen WR. Reproductive efficiency of thoroughbred mares in the United Kingdom. Proceedings of the Bain Fallon Memorial Lectures 1987;9:31–41. 113. Morris LHA, Allen WR. Reproductive efficiency of intensively managed thoroughbred mares in Newmarket. Equine Vet J 2002;34:51–60. 114. Bosh K. Reproductive efficiency of thoroughbred mares in central Kentucky. J Equine Vet Sci 2006;26:192–3. 115. Badi AM, O’Byrne TM, Cunningham EP. An analysis of reproductive performance in thoroughbred mares. Irish Vet J 1981;35:1–12. 116. Baker CB, Little TV, McDowell KJ. The live foaling rate per cycle in mares. Equine Vet J Suppl 1993;15:28–30. 117. Jeffcott LB, Rossdale PD, Freestone J, Frank CJ, TowersClark PF. An assessment of wastage in thoroughbred racing from conception to 4 years of age. Equine Vet J 1982;14:185–98. 118. Ball BA, Little TV, Hillman RB, Woods GL. Pregnancy rates at days 2 and 14 and estimated embryonic loss rates prior to day 14 in normal and subfertile mares. Theriogenology 1986;26:611–19. 119. Woods GL, Baker CB, Baldwin JL, Ball BA, Bilinski J, Cooper WL, Ley WB, Mank EC, Erb HN. Early pregnancy loss in brood mares. J Reprod Fertil Suppl 1987;35:455–9. 120. Platt H. Aetiological aspects of abortion in the thoroughbred mare. J Comp Pathol 1973;83:199–205. 121. Platt H. Growth of the equine foetus. Equine Vet J 1984;16:247–52. 122. Ginther OJ, Williams D. On-the-farm incidence and nature of equine dystocias. J Equine Vet Sci 1996;16:159– 64. 123. Frazer GS, Perkins NR, Blanchard TL, Orsini J, Threlfall WR. Prevalence of fetal maldispositions in equine
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Management of the Geriatric Mare referral hospital dystocias. Equine Vet J 1997;29:111– 16. 124. Douglas RH. Review of induction of superovulation and embryo transfer in the mare. Theriogenology 1979;11:33–46. 125. LeBlanc MM, Tran T, Baldwin JL, Pritchard EL. Factors that influence passive transfer of immunoglobulins in foals. J Am Vet Med 1992;200:179–83.
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126. Morley PS, Townsend HGG. A survey of reproductive performance in Thoroughbred mares and morbidity, mortality and athletic potential of their foals. Equine Vet J 1997;29:290–7. 127. Carney NJ, Squires EL, Cook VM, Seidel GE, Jasko DJ. Comparison of pregnancy rates from transfer of fresh versus cooled transported equine embryos. Theriogenology 1991;36:23–32.
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Chapter 297
Parentage Testing Ernest Bailey
Introduction Genetic tests have been used to verify parentage in horses since the 1960s. The genetics tests were initially based on blood groups and biochemical polymorphisms. Blood samples were relatively easy to collect and a wide range of genetic markers could be assayed. The nature and application of these tests has been reviewed elsewhere.1, 2 During the past decade DNA typing methods replaced blood typing tests for equine parentage because DNA tests had (i) comparable or decreased costs; (ii) excellent power to solve questions of parentage; and (iii) the ability to run the test on many tissues including blood, hair follicles, and even tissue from recently deceased horses. The following sections describe the current tests used for parentage testing in horses, standardization measures, and projections on how the technology may change.
Rules of parentage testing Regardless of the type of genetic factors measured for parentage, the principles of exclusion remain the same: (i) all hereditary factors found in an offspring must have been inherited from one or other of its two parents; and (ii) each parent will transmit 50% of its genetic material to the offspring. Whether the genetic factor is a blood group, DNA sequence, or even a coat color variant, the principles are the same. The hereditary basis for each discrete genetic factor is called a genetic system or, alternatively, a genetic locus. Let us use coat color as an example. The gray coat color in horses is due to the presence of at least one copy of a simple genetic dominant gene for Gray (G). If an offspring becomes gray, then at least one of its parents must be gray. If neither parent is gray, then the parentage is incorrect. This is often referred to as the gray rule.
The point is to use a number of genetic markers sufficient to permit a strong probable statement about parentage. While the occasional violation of the gray rule will show that a parentage is incorrect, this single test cannot provide sufficient power for confidence that the pedigree is necessarily correct. To provide sufficient power, the initial parentage tests were based on 12–14 genetic systems collectively referred to as blood typing. In the 1990s, direct tests of DNA became sufficiently powerful, convenient, and cost effective that 10–17 DNA systems replaced the 12–14 blood typing systems for use in horse parentage testing.
The DNA parentage test DNA is the chemical substance of heredity. The chemical structure is sometimes referred to as the blueprint for life. The DNA molecule directs or ‘encodes’ the structure, timing, location, and production of proteins and RNA that enable cells and tissues to function as they do. Differences found in DNA are responsible for differences found between species and even differences between individuals within a species. Each nucleated cell of a horse has a copy of paternally derived and maternally derived DNA, each containing 2.7 billion nucleotide bases, comprising its DNA. The complete set of DNA within an individual’s cell is called its genome. DNA is basically a simple chemical. There are only four types of nucleotides in DNA – adenine, cytosine, guanine, and thymine, cryptically referred to as A, C, G, and T. Despite the apparent simplicity of chemical composition, the unique aspect of DNA resides in the permutation of order for the 2.7 billion nucleotides found between individuals. No two individuals in the world have exactly the same order of DNA bases, including monozygotic twins, as a consequence of mutations occurring during development.3
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Parentage Testing Genetic differences are inherited. The point is to characterize and test these inherited DNA differences. In the early 1990s, many technologies were being developed to investigate genetic variation at the DNA level in connection with the human genome sequencing project. Those same technologies could and were being used to investigate genetic variation in livestock (reviewed by Womack4 ). DNA-based parentage testing for horses was an obvious application. While blood typing was effective, a system based on DNA typing might allow reduced costs and increased power. The systems considered included DNA fingerprinting with minisatellite DNA markers, restriction fragment length polymorphisms, detection of single nucleotide polymorphisms, and microsatellite DNA markers. Each method had its strengths and weaknesses. But this is a topic that falls outside this review. The main point here is that microsatellite DNA markers were selected as the most efficacious test system for parentage in horses. Marklund and co-workers proposed the use of microsatellite markers for parentage testing in horses.5 Bowling and co-workers developed and evaluated an efficient, cost-effective parentage test using microsatellite DNA sequences.6 Their method comprised testing 11 genetic systems based on DNA extraction and polymerase chain reaction for each of the 11 loci. They could use cells derived from any tissue, including blood and hair roots. The power of their method was comparable to blood typing in identifying horses and solving questions of parentage. In addition to developing the system, the scientists at the University of California, Davis, also assisted scientists in other laboratories, worldwide, in setting up comparable testing programs, including provision of protocols and computer software. This collegial cooperation and collaboration was key to the adoption of the methodology in the comparison tests of the International Society of Animal Genetics and the ability to compare results worldwide.
What are microsatellite DNA markers? Although we now have a complete genome sequence for the horse, the structure of the genome is complex and we still do not understand how it all functions. We know that the horse, similar to other mammals, has approximately 2.7 billion base pairs of DNA. As with all mammals studied thus far, the horse has approximately 20 000 genes encoding proteins. At the same time, genes comprise only 2–3% of the DNA! The rest of the DNA (97–98%) includes sequences that regulate the expression of DNA, some of which may simply provide space between genes or are remnants of earlier evolutionary developments in
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… agatcaagac tttccctata tctcagtact gtctgtgccc ttccttgtgt cagttcactg aatgtcaaat tctgctcttt agcatttgtg tgtgtgtgtg tgtgtgtgtg tgtgtgtgtg tgtgtggtat cttatcacag cctccaagca ggaa… Figure 297.1 Base sequence of HTG6 microsatellite (Genbank accession: AF169167).
the DNA sequence and currently have no function. The discovery of so much non-genic DNA was a surprise and its study is a major focus of current basic research. Microsatellite DNA is a small portion of the 97–98% of the DNA that we do not understand. However, microsatellite DNA is useful for parentage testing. The DNA of animals contains over 100 000 of these microsatellite DNA elements, randomly dispersed throughout the genomes. The easiest way to understand microsatellite DNA markers is initially to understand one of them. One of the first DNA microsatellite markers reported for horses was HTG6.7 The DNA sequence associated with HTG6 is shown in Figure 297.1. This figure represents one strand of DNA for a very short section of the horse genome with the nucleotides presented as adenine (A), cytosine (C), guanine (G), and thymine (T). One of the key features in this figure is the underlined DNA sequence that appears in the middle of the section, specifically, tgtgtgtgtgtgtgtgtgtgtgtgtgtgtgtgtg. This series of repeated units can be abbreviated as (TG)20, representing 20 units of the TG dinucleotide. This dinucleotide is the microsatellite. This precise sequence of TG repeats is not actually unique; TG repeats are common and can be found tens of thousands of times throughout the genome of the horse. However, the sequences flanking the microsatellite, shown in boldtype in Figure 297.1, are unique to this precise location. The sequences represented in bold occur nowhere else in the genome of the horse. When the whole genome sequence of the horse was completed in 2007, we found that HTG6 is present on horse chromosome 15 and between positions designated 73962738 and 73962737 in the second assembly of the horse genome (http://www. ensembl.org/Equus caballus/Info/Index). We can use the information about the DNA sequences flanking from this microsatellite and a technique called polymerase chain reaction (PCR) to specifically amplify this one section of DNA. In this case, the scientists investigating HTG6 designed a PCR reaction to amplify a DNA fragment 100 bases long with the microsatellite DNA in the center. More importantly, they were able to assay the length of the DNA
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fragment and verify that it was exactly 100 bases long in the horse used to discover the microsatellite. Next, these scientists used the PCR test to investigate the length of this DNA sequence in other horses. Among 20 horses they found four different DNA fragments for this region including fragments 100 bases long, 90 bases long, 88 bases long, and 84 bases long. Based on DNA sequencing, the only differences between the four different fragments were the number of TG repeats. While the original horse had repeat (TG)20, the other horses had repeats (TG)15, (TG)14, and (TG)12. The genetic variants were inherited in the same fashion as blood group and biochemical markers. Each horse could have two different alleles, one inherited from each parent. Therefore, assays of this microsatellite could be used for parentage testing. Variation in the length of DNA sequences is characteristic of such tandem DNA repeats. We believe that during the replication of DNA in the germ cells, the DNA from the paired chromosomes may occasionally misalign in regions with such repeats and the process of repair or replication results in the addition or deletion of a nucleotide repeat unit. Alternatively, this may simply be a misreading error by the polymerases in germinal cells that results in gain or loss of a repeat unit. Regardless of the cause of this variation, we have learned that when a dinucleotide repeat unit in a microsatellite exceeds 10 these microsatellites are likely to exhibit genetic variation in the number of dinucleotide repeat units. Subsequent studies identified additional microsatellite markers. Over 1000 have been used in gene mapping studies.8, 9 A subset, shown in Table 297.1, has been selected and used to test hundreds of thousands of horses in commercial parentage testing laboratories. These 17 microsatellite markers are commonly used for routine parentage testing based on recommendations of the International Society for Animal Genetics (ISAG). The 11 microsatellites originally developed by Bowling et al.6 are identified at the top of the list. These 11 markers are sufficient for routine parentage testing in most horse breeds. The six microsatellites listed at the bottom of Table 297.1 are markers recommended for increased power, although some laboratories find sufficient power and decreased cost when doing a subset of the 17 markers.
ISAG comparison tests and standardization Parentage testing of livestock, especially cattle, began on a large scale in the 1950s. Artificial insemination had become an important application for cattle,
Table 297.1 Microsatellite loci used for routine parentage testing Locus
Chromosome location
AHT4 AHT5 HMS1 HMS3 HMS6 HMS7 HTG4 HTG6 HTG7 HTG10 VHL20 ASB23 ASB2 HMS2 ASB17 LEX3 CA425
24 8 15 9 4 1 9 15 4 21 30 3 15 10 2 X 28
and breeders wanted to know that the semen used to inseminate their cows came from the correct bull. Complex blood typing systems had been discovered for cattle and they were sufficiently powerful to be applied to parentage testing. Many laboratories began offering this service. However, when breeders wanted to use results from different laboratories to make parentage assessments, difficulties arose with respect to the nomenclature of systems, the number and type of systems used, and even in technical differences between laboratories. Therefore, to facilitate research on genetic markers in livestock and to conduct comparison tests on existing and new tests, the European Society for Blood Group Research was formed in 1964 and evolved into the International Society for Animal Blood Group Research in 1972 to accommodate scientists in the Americas, Africa, and Australasia. During the 1970s and 1980s, society members were conducting an increasing amount of research involved with lymphocyte typing, DNA polymorphisms, genomics, and gene expression. Therefore, in 1988, the Society once again changed its name to the International Society for Animal Genetics (ISAG). Apropos of this review, ISAG remains the primary venue for comparison of genetic systems for parentage testing and provides laboratories with the opportunity for self-evaluation of testing procedures through a comparison test with peers from other laboratories around the world. In this way, the methods for testing and reporting are standardized.
Standardization Horses are transported all over the world, and different DNA typing laboratories need to be able to share
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Table 297.2 An example parentage case Gene locus
AHT4
AHT5
HTG4
HTG7
VHL20
ASB23
LEX3
CA425
Stallion 1 Stallion 2 Dam Offspring Exclusion
K/P K/L J/P J/K No
L/N M/N L/N L/N No
M/P M/O L/N L/O @
K/O J/K J/O J/K No
M/N O/N M/O M/N No
N/R M/N J/R J/N No
O O J/R J/O No
J/R O/P M/P M/P #
‘No’ – the dam and either stallion qualifies as parent to the offspring. @ – Type 1 and type 2 exclusion of Stallion 1. # – Type 1 exclusion of the mating for Stallion 1 and the dam.
data. The instruments and methods used for detecting DNA fragment lengths were not identical between laboratories; consequently, minor differences could appear in reports between laboratories. Although the results were repeatable within and between laboratories, sharing test results required standardization of nomenclature between laboratories. Through the auspices of the ISAG Horse Genetics Standing Committee a common nomenclature was adopted for the alleles detected in each laboratory and a set of standard samples established to verify the typing results. Rather than report the size of the DNA fragment, each allele was identified and referred to using an alphabetical nomenclature, beginning with the centermost allele represented by the letter ‘M’.
Parentage analysis Table 297.2 presents an example of a parentage case investigated using microsatellite DNA markers and this nomenclature. There are two rules that we consider for parentage analysis: 1. Every marker found in the foal must be present in one or other of the parents. If not, then the parentage is excluded. Either the dam or the sire or both are incorrectly identified. 2. When a parent has two genetic factors for a single locus (e.g., the dam has factors L and N at the HTG4 locus) then the foal must inherit the gene for one of them. If the foal has neither of the factors, then that horse cannot be a parent of the foal. The results of typing the putative dam, its offspring, and two putative sires with eight microsatellite markers are presented. The top row identifies the genetic loci used for the study. The rows for genetic typing results for Stallion 1, Stallion 2, the putative dam, and the offspring are identified in column 1. Columns 2–9 show the results for each of the microsatellite markers. AHT4: The foal has genetic markers J and K. The dam has J but not K. Therefore, if the dam is correct, the
sire will have K. Both stallions possess K. Result: no exclusion possible. AHT5: The foal has genetic markers L and N. The dam has both L and N. Stallion 2 has N. Stallion 1 has both L and N. No exclusion possible. HTG4: The foal has L and O. The dam has L but not O. If the dam is correct the sire will have O. Stallion 2 has O and qualifies as the sire with respect to the stated dam. Stallion 1 does not have L or O, consequently cannot be the sire. Result: Stallion 1 is definitely excluded based on rule #1 and rule #2 without regard to the type found in the dam. HTG7: The foal has J and K. The dam has J. If the dam is correct, the sire must have K. Both stallions have K. Result: no exclusion is possible. VHL20: The foal has M and N. The dam has M. If the dam is correct, the sire must have factor N. Both stallions have factor N. Result: no exclusion is possible. ASB23: The foal has J and N. The dam has J. If the dam is correct, then the sire must have N. Both stallions have N. Result: no exclusion is possible. LEX3: The foal has J and O. LEX3 is unique among these markers because the gene for LEX3 occurs on the sex chromosome, X. Females have two X chromosomes while males have a single X chromosome. Since the foal has two alleles for LEX3, we expect the foal to be female. The dam has factor J and therefore the sire must have contributed factor O. Both stallions have the O marker and are not excluded. (Having a single marker is consistent with males having a single X chromosome.) Result: no exclusion possible. CA425: The foal possesses M and O. The dam possesses M. If the dam is correct, the sire must possess O. Stallion 2 possesses O. Stallion 1 does not. However, Stallion 1 does possess M and would qualify by himself without regard to the dam. Result: the mating of Stallion 1 and the dam is excluded as the correct parentage for the foal based on rule #1. The mating of Stallion 2 and the dam is not excluded.
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Summary and conclusion for this case The test results definitely prove that the foal cannot be the offspring of Stallion 1 and the dam. The mating is excluded based on the results for CA425. Stallion 1 is specifically excluded on the basis of results for HTG4. However, in all systems the mating of Stallion 2 and the dam qualify as the parentage for the offspring. Two points need to be emphasized for interpretation of any parentage testing results. First, exclusion of a horse is a statement of scientific fact. With one type of exception, the exclusion of a horse means it cannot be a parent. The only exception is a very rare case in which a horse may have an admixture of cells, each with different genomes due to unusual events occurring during embryonic development. Such individuals are called chimeras, or cell-mosaics. It is possible that only one type of cell will be sampled in the test and if that cell type is not the one inherited by the offspring, an exclusion will be made. Chimeras are very rare, much more so than genetic mutations. Such a discovery is so unusual that it would warrant publication of a case report. The second point is that qualification of a parentage is a statement of probability and not a statement of irrefutable fact. Each genetic system provides partial evidence for the veracity of a mating. The large number of genetic tests used for parentage testing is selected such that failure to exclude an incorrect parentage becomes infinitesimally small. Consequently, failure to exclude plus evidence from the breeder that the stated mating took place is essentially accepted as proof of the pedigree.
Other services provided by parentage testing laboratories Antibody screening for transfusions and for neonatal isoerythrolysis (NI) Historically, equine parentage testing laboratories have provided additional genetic services. One was to conduct tests for NI (or hemolytic disease of the newborn foal; see Chapter 35). The blood group tests initially used for parentage testing could also be used to determine whether or not a mare was producing antibodies that would destroy its foal’s red blood cells. Blood group and cross-matching tests have also been used for transfusions between horses. Many parentage testing laboratories continue to have this expertise and continue to offer these services. However, the service is no longer related to parentage testing because the new assays test DNA. The basis for NI or transfusion compatibility at the molecular level is not known, so consequently the DNA tests are not informative. Fortunately, many veterinary diagnos-
tic laboratories have begun to offer NI or transfusion compatibility services and, therefore, these services are likely to survive their possible demise at parentage testing laboratories.
Molecular genetic tests for health and color traits Genetic testing services for equine hereditary diseases and coat color traits are available through equine parentage testing laboratories in addition to other genetic testing laboratories. The technologies for detecting these traits are similar or identical to the DNA-based technologies used for parentage testing. Table 297.3 lists the traits for which tests are currently available. The first DNA diagnostic test was for hyperkalemic periodic paralysis (HYPP) of Quarter Horses.10 The second DNA test was developed for the gene responsible for black or red pigment for horses (black/bay vs chestnut).11 With the advent of whole genome sequencing it is likely that this list will expand greatly during the next couple of years. Genetic counseling is likely to take on greater importance when complex traits (osteochondrosis, conformation faults, allergic diseases, etc.) are studied and the interplay between environment, management, and heredity becomes better understood.
Future of parentage testing in horses In the previous edition of this book, parentage testing was described based on blood groups and biochemical polymorphisms, a method begun in the 1960s and spanning 30 years. Although some registries continue to use blood typing, a shift to the use of microsatellite DNA testing is almost complete because of cost, efficacy, and especially because of the convenience of testing DNA extracted from hair follicles rather than blood. One major hurdle was retesting horses. The genetic variants detected with DNA tests were not the same as those detected with blood groups. Many horse owners and many breed registries were asked to bear the cost of retesting horses with this new genetic system. The question recurs, what will the future hold for DNA testing of horses? Biology and technology move quickly. The most recently developed genetic marker is called a single nucleotide polymorphism (SNP). Based on work in horses and other mammals, we anticipate the existence of one DNA base difference per 1000 bases between any two horses. These differences are called SNPs. A powerful genetic test could be devised using SNPs for comparing horses. Indeed, at present an assay to detect 60 000 SNPs in horses has been developed for use in research (Illumina, Inc.).
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Table 297.3 Currently identified genetic traits characterized using genomics Trait (symbol)
Citation
Hyperkalemic periodic paralysis (HYPP) in Quarter Horses Extension – red and black hair color (E) Severe combined immunodeficiency (SCID) of Arabian horses Cream hair color dilution (CR) Tobiano hair color pattern (TO) Agouti hair color pattern (A) Hereditary equine regional dermal ashthena (HERDA) Sabino 1 hair color pattern (SB1) Gray hair color pattern (G) Champagne hair color dilution (CH) Silver hair color dilution (S) Appaloosa hair color pattern (LP) Overo lethal White Foal (O) Junctional mechanobullous disease of Belgian horses (JEB) Junctional mechanobullous disease of Saddlebred horses (JEB) Glycogen storage disease IV (GSDIV) Polysaccharide storage myopathy (PSSM) Dominant white hair color pattern (W)
Rudolph et al.10 Marklund et al.11 Shin et al.13 Mariat et al.14 Brooks et al.15 Rieder et al., 200116 Tryon et al.17 Brooks and Bailey18 Pielberg et al.19 Cook et al.20 Brunberg et al., 200621 Bellone et al.22 Metallinos et al.;23 Santschi et al.;24 Yang et al.25 Spirito et al.26 Graves et al.27 Ward et al.28 McCue et al.29 Haase et al.30
Most of these SNPs have no impact on health or performance but can be very useful for parentage testing. Baruch and Weller12 estimated that an assay with less than 100 SNPs would provide sufficient power to solve most questions of parentage. While this is technically possible, it does not seem an economic likelihood in the near future. As noted above, the conversion from blood typing to DNA microsatellite typing entailed wholesale retesting of the horse population. This was an expensive and controversial undertaking. The economic advantages of the new test, by reducing subsequent costs to horse owners, made this a reasonable cost to bear from the perspective of most horse registries. However, the cost of SNP typing is comparable or higher than the cost of microsatellite testing. There is no advantage for the power of the test or the cost. Since the current test has a low cost and high efficacy, microsatellite DNA typing is likely to remain the standard method for parentage testing the horse until the next edition of this book.
4. 5.
6.
7.
8.
9.
References 1. Bailey E, Cothran EG, Trembicki KA. Paternity testing. In: McKinnon AO, Voss JL (eds) Equine Reproduction. Philadelphia: Lea and Febiger, 1992; pp. 626–30. 2. Sandberg K, Cothran EG. Blood groups and biochemical polymorphisms. In: Bowling AT, Ruvinsky A (eds) The Genetics of the Horse. New York: CABI, 2000; pp. 85–108. 3. Bruder CE, Piotrowski A, Gijsbers AA, Andersson R, Erickson S, de St˚ahl TD, Menzel U, Sandgren J, von Tell D, Poplawski A, Crowley M, Crasto C, Partridge EC, Tiwari H, Allison DB, Komorowski J, van Ommen GJ, Boomsma
10.
11.
DI, Pedersen NL, den Dunnen JT, Wirdefeldt K, Dumanski JP. Phenotypically concordant and discordant monozygotic twins display different DNA copy-number-variation profiles. Am J Hum Genet 2008;82:763–71. Womack J. Advances in livestock genomics: opening the barn door. Genome Res 2005;15:1699–705. Marklund S, Ellegren H, Eriksson S, Sandberg K, Andersson L. Parentage testing and linkage analysis in the horse using a set of highly polymorphic horse micorsatellites. Anim Genet 1994;25:19–23. Bowling AT, Eggleston-Stott ML, Byrns G, Clark RS, Dileanis S, Wictum E. Validation of microsatellite markers for routine horse parentage testing. Anim Genet 1997; 28:247–2. Ellegren H, Johansson M, Sandberg K, Andersson L. Cloning of highly polymorphic microsatellites in the horse. Anim Genet 1992;23:133–42. Penedo MCT, Millon LV, Bernoco D, Bailey E, Binns M, Cholewinski G, Ellis N, Flynn J, Gralak B, Guthrie A, Lindgren G, Lyons LA, Tozaki T, Røed K, Swinburne J. International Equine Gene Mapping Workshop Report: A comprehensive linkage map constructed with data from new markers and by merging four mapping resources. Cytogenet Genome Res 2005;111:5–15. Swinburne JE, Boursnell M, Hill G, Pettitt L, Allen T, Chowdhary B, Hasegawa T, Kurosawa M, Leeb T, Mashima S, Mickelson JR, Raudsepp T, Tozaki T, Binns M. Single linkage group per chromosome genetic linkage map for the horse, based on two three-generation, full-sibling, crossbred horse reference families. Genomics 2006;87:1–29. Rudolph J, Spier S, Byrns G, Rojas C, Bernoco D, Hoffman E. Periodic paralysis in Quarter Horses, a sodium channel mutation disseminated by selective breeding. Nat Genet 1992;2:144–7. Marklund L, Johanssonn Moller M, Sandberg K, Andersson L. A missense mutation in the gene for melanocytestimulating hormone receptor (MC1R) is associated with the chestnut coat color in horses. Mamm Genome 1996;7:895–9.
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12. Baruch E, Weller JI. Estimation of the number of SNP genetic markers required for parentage verification. Anim Genet 2008;179:1861–70. 13. Shin EK, Perryman LE, Meek K. A kinase negative mutation of DNA-PKcs in equine SCID results in defective coding and signal joint formation. J Immunol 1997;158:3565–9. 14. Mariat D, Taourit S, Guerin G. A mutation in the MATP gene causes the cream coat color in the horse. Genet Sel Evol 2003;35:119–33. 15. Brooks SA, Lear TL, Adelson DA, Bailey E. A chromosome inversion near the KIT gene and the tobiano spotting pattern in horses. Cytogenet Genome Res 2008;119:225–30. ´ 16. Rieder S, Taourit S, Mariat D, Langlois B, Guerir G. Mutations in the agouti (ASIP), the extension (MCIR), and the brown (TYR P1) loci and their association to coat color phenotypes in horses. Mamm Genome 2001;12:450–455. 17. Tryon RC, White SD, Bannasch DL. Homozygosity mapping approach identifies a missense mutation in equine cyclophilin B (PPIB) associated with HERDA in the American Quarter Horse. Genomics 2007;90:93–102. 18. Brooks SA, Bailey E. Exon skipping in the KIT gene causes the Sabino spotting pattern in horses. Mamm Genome 2005;16:893–902. 19. Pielberg GR, Golovko A, Sundstrom E, Curik I, Lennartsson J, Seltenhammer MH, Druml T, Binns M, Fitzsimmons C, Lindgren G, Sandberg K, Baumung R, Vetterlein M, Stromberg S, Grabherr M, Wade C, Lindblad-Toh K, Ponten F, Heldin C-H, Soklner J, Andersson L. A cis-acting regulatory mutation causes premature hair graying and susceptibility to melanoma in the horse. Nat Genet 2008. doi: 10.1038/ng.185 20. Cook D, Brooks SA, Bellone R, Bailey E. Missense mutation in exon 2 of SLC36A1 responsible for champagne dilution in horses. PLOS Genetics 2008; epub: 1000195 doi: 10.1371/journal.pgen.10000195. 21. Burnberg E, Anderson L, Cothran G, Sandbug K, Lindgren G. A missuse mutation in DMEL17 is associated with Silver coat colour in the horse. BMC Genet 2006;7:46. 22. Bellone RR, Brooks SA, Sandmeyer L, Murphy BA, Forsyth G, Archer S, Bailey E, Grahn B. Differential gene expres-
23.
24.
25.
26.
27.
28.
29.
30.
sion of TRPM1, the likely cause of congenital stationary night blindness (CSNB) and coat spotting pattern (LP) in Appaloosa horses (Equus caballus). Genetics 2008;179: 1861–70. Metallinos DL, Bowling AT, Rine J. A misssense mutation in the endothelin-β receptor gene is associated with lethal white foal syndrome: an equine version of Hirschsprung Disease. Mamm Genome 1998;9:426–31. Santschi EM, Purdy AK, Valberg SJ, Vrotsos PD, Kaese H, Mickelson JR. Endothelin receptor β polymorphism associated with lethal white foal syndrome in horses. Mamm Genome 1998;9:306–9. Yang GC, Croaker D, Zhang AL, Manglick P, Cartmill T, Cass D. A dinucleotide mutation in the endothelin-B receptor gene is associated with lethal white foal syndrome (LWFS) a horse variant of Hirschspring disease. Hum Mol Genet 1998;7:1047–52. Spirito F, Charlesworth A, Linder K, Ortonne JP, Baird J, Meneguzzi G. Animal models for skin blistering conditions: absence of laminin 5 causes hereditary junctional mechanobullous disease in the Belgian horse. J Invest Dermatol 2002;119:684–91. Graves KT, Henney PJ, Ennis RB. Partial deletion of the LAMA3 gene is responsible for junctional epidermolysis bullosa (JEB) in the American Saddlebred horse. Anim Genet 2009;40:35–41. Ward TL, Valberg SJ, Adelson DL, Abbey CA, Binns MM, Mickelson JR. Glycogen branching enzyme (GBE1) mutation causing equine glycogen storage disease IV. Mamm Genome 2004;15:570–7. McCue ME, Valberg SJ, Miller MB, Wade C, DiMauro S, Akman HO, Mickelson JR. Glycogen synthase (GYS1) mutation causes a novel skeletal muscle glycogenosis. Genomics 2008;91:458–66. Haase B, Brooks SA, Schlumbaum A, Azor RJ, Bailey E, Alaeddine F, Mevissen M, Burger D, Poncet P-A, Rieder S, Leeb T. Allelic Heterogeneity at the equine KIT locus in dominant white (W) horses. PLoS Genet 2007;3(11): e195. doi: 10.1371/journal.pgen.0030195
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Chapter 298
Relevance of Genomics to Equine Reproduction Ernest Bailey
Reproductive physiologists use a wide array of tools to pose and answer questions about reproduction in horses, including behavioral biology, protein chemistry, histology, radioimmunoassay, and genomics. This chapter is about the use of genomics. Inventions coming from the Human Genome Project have made possible the relatively inexpensive investigation of the DNA of any species. Two factors were responsible for applications beyond the Human Genome Project. First, the techniques used to understand human genomics were entirely applicable to the genomics of any other organism. Second, the organization of the human genome is very similar to that of all animals, and especially mammals. Genomics can be defined in a narrow sense as relating to the complete set of DNA molecules found within an individual. In a broader sense, characterization of DNA is fundamental to investigations of gene expression, protein production, and cellular physiology. Consequently, the term ‘genomics’ has been used to encompass derivative fields such as transcriptomics, proteomics, and metabolomics. A workshop on horse genomics, begun in 1995, led to the construction of several genetic maps and creation of numerous tools for use in genomic studies. Murray1 reviewed the progress of the horse genomics work and discussed potential applications to horse reproduction. The single greatest contribution to horse research occurred in 2006 when the Broad Institute sequenced the whole genome for the horse, then completed the genome assembly in 2007. Since reproduction involves the profound alteration of cell types through changes in gene expression, scientists working in equine reproductive biology will benefit greatly by using the whole genome sequence of the horse.
This chapter describes the current status of horse genomics, the technological developments we anticipate within the next few years, and the applications we have seen or anticipate related to equine reproduction. These applications include discovery of genes affecting fertility and embryonic losses, use of genomics tools to investigate gene expression in reproductive tissues, and increased resolution for cytogenetic techniques.
The horse genome sequence The largest contribution ever to horse research has been the sequencing of the horse genome by the Broad Institute under the auspices of the National Human Genome Research Institute (NHGRI). Ironically, the primary objective of that activity was to provide information about the function of the genome relative to humans. When the first human sequence was reported, scientists were surprised to learn that only 3% of the DNA in the human genome coded for proteins. The role or origin of the remainder of the human genome might include regulation of gene expression. Since the function of genomes was very similar among mammals, scientists at NHGRI hypothesized that sequences conserved between mammalian species would be those that played a functional role. Therefore, representatives from diverse parts of the mammalian phylogenetic tree were selected and their genomes sequenced. The horse was selected as a representative from the order Perissodactyla (odd-toed mammals, including rhinoceroses, tapirs, and equids). The horse was chosen because a group of scientists had been mapping the horse genome since 1995 and wanted to make veterinary medical applications that might
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 298.1 Horse genome information Number of chromosomes Number of DNA bases Depth of sequence (fold coverage) Polymorphism rate Estimated number of genes
32 pairs 2.47 gigabase 6.8 1/1500 bases 20 737 (per Ensembl)
Websites for genome browsers: UCSC Genome Bioinformatics (http://genome.ucsc.edu/). NCBI Map Viewer for Equus caballus (horse) (http://www.ncbi.nlm.nih.gov/ mapview/map search.cgi?taxid=9796). Ensembl (http://www.ensembl.org/index.html). Websites for BACs: Horse Genome Project (http://www.tiho-hannover.de/einricht/zucht/hgp/ index.htm). BACPAC Resources Center (http://bacpac.chori.org/).
also benefit human health research; see the Horse Genome Project website (http://www.uky.edu/Ag/ Horsemap/). An inbred female Thoroughbred horse was selected for sequencing and a 6.8-fold shotgun sequence was performed followed by two assemblies of the genome (Table 298.1). The first assembly was based solely on computer algorithms of sequence alignment. A second assembly was based on the alignment plus data from the several genetic maps created for the horse,2–4 plus cytogenetic mapping of clones with ambiguous assignments (T.L. Lear, unpublished). The size of the genome was estimated to be approximately 2.47 billion bases, and annotation by Ensembl using predictions from the human genome annotation estimated the presence of 20 737 genes. Using the websites listed in Table 298.1, anyone with access to the internet can identify DNA sequences for any portion of the genome. In addition, should someone need clones for specific DNA regions, Table 298.1 also lists two other websites, one for a database that identifies the mapping relationships among 150 000 large sequence clones from a half-sibling to the horse used for sequencing, and another that identifies a commercial site for obtaining clones.
DNA sequences and hereditary defects The most obvious application for genomics is the discovery of DNA sequences that are diagnostic for traits of interest and genetic defects. Determining whether reproduction problems originate from management practices or genetic problems is important. Problems with management practices can be fixed; horses with genetic problems involving reproduction can be sold for use in non-breeding activities. Therefore, breeders may be interested in having an array of tests to identify genetic problems associated with fertility. Genetic problems associated with
cytogenetics are discussed in Chapter 207. Cytogenetic analyses can identify chromosome problems as a cause of infertility or reduced fertility. Cytogenetic changes are usually large, involving megabases of DNA and are visible at the microscopic level. Genomics approaches are also increasing the resolution of cytogenetics tests.5 Sometimes breeders use horses with reduced fertility in their breeding programs because they consider that the genetic aspects of performance outweigh the costs of reduced fertility. Hereditary influences on fertility are strongly suspected for a wide variety of reproductive traits. For example, a hereditary predisposition to cryptorchism is suspected for some lines of horses. There are stallion lines among Standardbred and Thoroughbred horses thought to transmit hereditary traits manifest in poor semen quality. Embryonic lethal recessive genes can reduce pregnancy rates in mares. Examples would be the proposed homozygous lethal conditions associated with dominant white color in horses6 or homozygous lethal roan of Belgian horses.7 Molecular studies may identify the precise nature of those lethal genes in the near future. Genomics and molecular genetics approaches are being used in basic biological studies to investigate subtle but discrete reproduction traits in horses. Leeb8 noted that over 200 discrete genes are known that have an impact on spermatogenesis. He described the advances in horse genomics and how the resulting tools could be used to investigate these genes. Some preliminary data were presented comparing sperm adhesin genes in several different species. Hamann et al.9 reported the first genetic variant associated with a reproduction trait for horses. They investigated three cysteine-rich secretory protein (CRISP) genes for Hanoverian stallions and reported a variant of CRISP3 associated with a statistically significant reduction in male fertility as detected from insemination records. CRISP genes were selected for the study because they are expressed uniquely in the testis and the gene products play a role during fertilization. The level of significance was high (P = 0.023) so, if valid, selection would be an effective application of the observation. However, determining and augmenting the function of the CRISP3 gene may provide a management approach to improving fertility. The authors encouraged additional biochemical and physiological studies to confirm and extend the study. Chowdhary et al.10 also reviewed advances in horse genomics and described their investigations of the Y chromosome. Defects in the Y chromosome are sometimes a cause of human infertility. Consequently, the Y chromosome of horses is a
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Relevance of Genomics to Equine Reproduction prime candidate for investigation of male fertility problems. The power of genomics to uncover genes for specific traits is well established. Scientists used the gene map and a variety of approaches to identify genes for diseases and coat color traits in horses. See Table 297.3 on page 2825 for a list of those traits. Most of those traits were identified by investigation of families segregating for the trait under study. Two basic approaches were used. First, candidate genes from human or animal conditions were investigated as likely having a homologous effect in horses. In this fashion, the genes for hyperkalemic periodic paralysis (HYPP) of Quarter Horses,11 and severe combined immunodeficiency disease (SCID) of Arabian foals12 were discovered. The second approach was to scan the entire genome using microsatellite genetic markers and identify the chromosome region harboring the gene responsible for the trait. Then candidate genes were chosen from that region and investigated for the presence of mutations causing the trait. Examples of genes discovered through this route include cream dilution (CR) and champagne dilution (CH) of hair color.13, 14 The power of the gene map made possible the use of small families to identify such traits. However, for many hereditary conditions it has not been possible to assemble families because the conditions have been rare or because breeders have been disinclined to continue matings that may result in a hereditary defect. Solutions for these kinds of genetic problems require an approach that does not require assembly of families. Single nucleotide polymorphisms (SNPs) and gene discovery The whole genome sequence has provided tools that allow us to find mutations without the requirement for families of horses. During the sequencing study, the Broad Institute partially sequenced DNA from seven additional horses, including an additional Thoroughbred, Standardbred, Arabian, Quarter Horse, Andalusian horse, Icelandic horse, and an Akhal-Teke. When the DNA sequences were compared, almost 1.5 million sites were identified for which there was DNA nucleotide variation between at least one pair of horses. These differences in DNA nucleotides were called ‘single nucleotide polymorphisms’ (SNPs). The distribution of these SNPs is not random. Particular combinations of SNPs are associated with particular genes. For example, the chromosome region on ECA25 possessing the gene for gray provides a unique signature for the chromosome region possessing that gene.15 This association is a valuable tool that we can use to determine which chromosomes possess the genes of interest, and help
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us to find the actual gene. Basically, if we compare the combination of SNPs among horses with the gray gene to the combination found among horses without the gray gene, we will find a set of SNPs on horse chromosome 25 unique to the gray horses (C. Wade and J. Mickelson, personal communication, 2008). Illumina Inc. (San Diego, CA) has developed a commercially available product called the Equine SNP50 BeadChip, which can test for over 54 000 SNPs evenly distributed over the horse genome (excluding the Y chromosome). Since families are difficult to assembly when fertility is the issue, the SNP approach offers great advantages for the study of reproduction traits. At the same time, we need to recognize that even weak natural selection would eliminate single, simple mendelian traits affecting fertility; therefore, hereditary traits related to reproduction are likely to be the result of gene action from multiple genetic loci. Inheritance patterns may not be readily apparent, and selection may be an ineffective tool in the short term. Consequently, the greatest value of this work will be to increase our understanding of reproductive physiology for management and long-term genetic planning.
Gene expression and reproductive physiology Genomics is a powerful tool for experiments on nonhereditary aspects of biology. As tissues develop or respond to changes in their environments, they change the array of proteins expressed on their surface, excreted, or found within the cell. Reproductive tissues exhibit this characteristic to a greater extent than any other mammalian system. During the course of reproduction, tissues grow, atrophy, and remodel. Scientists measure these events using a wide array of histological, biochemical, and molecular approaches. Detection of specific proteins through the use of specific immunoglobulins or physico-chemical studies has been a powerful technique applied to reproductive physiology for the last 50 years. Radioimmunoassays, immunohistochemical staining, two-dimensional gel electrophoresis, and other techniques have been used to unravel many questions about the biology of reproduction. Unfortunately, the species specificity of commercially available antibodies for human or mouse proteins makes it necessary to begin research on horse proteins with the manufacture of horse-specific antibodies. A reasonable alternative is investigation of gene expression using genomics information. Changes in gene expression precede changes in protein expression in a cell. DNA sequences in genes are highly conserved among species, and we can readily
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identify the equine homolog for genes characterized in studies on other species. The availability of annotated information on the equine whole genome sequence makes it possible to create equine-specific genetic probes. Investigations of candidate gene expression Gene expression has been used to augment protein studies or simply to characterize physiological changes during reproduction in horses using a candidate gene approach. Candidate genes are those for which we have an a priori expectation for a key role. The candidate gene approach has been commonly used in equine reproductive physiology with respect to proteins and based on studies of reproduction in other species. In the 1990s scientists began to look at candidate gene expression in different tissues. For example, Simpson et al.16 investigated mRNA expression of the gene oxytocin-neurophysis I in equine uterine endometrium. Likewise, expression of estrogen receptor-alpha and progesterone receptors in the uterus was investigated by assaying mRNA species. 17 In these and other studies, isolation of mRNA, nucleotide sequencing, and Southern and northern blotting techniques were effective for investigating expression of candidate genes. More recently the use of quantitative real-time polymerase chain reaction (RT-PCR) has been used to evaluate the expression of candidate genes.18 This technique is based on isolation of mRNA from tissues under study, and the construction of highly specific DNA probes based on the available horse DNA sequence for a quantitative and specific assay of gene expression. Since 2000, the literature has accumulated several dozen scientific articles in which these genomic approaches have been used to study candidate gene expression in studies of reproductive physiology in horses. Investigations of global gene expression The annotation of the horse genome suggests the presence of 20 737 genes. The candidate gene approach has been attractive because (i) it is hypothesis driven and (ii) it has not been possible to assay all 20 737 genes. But if we do not have a candidate gene, we are left with the question, ‘which of the 20 737 horse genes is implicated in the physiological change?’ Gu and Bertone19 designed and tested the first microarray designed to test gene expression in horses. They took advantage of 18 924 DNA sequences in National Center for Biotechnology Information (NCBI) databases to design a microarray for 3098 genes. They showed that the array was effective in measuring the response of equine synoviocytes to lipopolysac-
charide (LPS). The obvious limitation to this study was the lack of DNA sequences for horse genes. Ramery et al.20 reported the use of a human microarray to study gene expression associated with respiratory disease in horses. Predictably, the array did detect difference based on the close homology between horse genes and human genes; however, the DNA sequences were sufficiently different to make this a not entirely satisfactory approach. Levels of expression measured by a cross-species array will be influenced by gene homology as well as level of mRNA produced. We anticipate new microarrays becoming available in the near future based on the whole genome sequence for the horse . . . all 20 000 genes. Scientists and commercial enterprises are using the annotated genome information to create several different platforms for investigation of horse gene expression. This technology will be especially useful for studying the reproductive biology of horses, and will undoubtedly lead us to genes and proteins that we had not previously suspected.
Conclusions Genomics is a tool that is evolving rapidly. By the time this chapter is printed, undoubtedly there will be yet other technological advances in genomics that will enhance research on equine reproduction. The applications are evolving as well. Classically, we think of genetics and genomics as being applied to hereditary questions. Genetic differences between individuals may impact their reproductive performance and become a basis for selection or culling. Genetic differences between individuals are also experiments of nature that can teach us about reproductive biology. At the same time genetics and genomics may become more useful for understanding non-hereditary aspects of reproduction. As the reproductive tissues respond to hormonal and environmental stimuli, they alter their gene expression. These changes affect the proteins produced by the cells and tissues. Sometimes it is more effective to study the proteins, whereas at other times it will be more effective to study gene expression. We now have the tools to do this. Looking ahead, we need to learn how to use genomics tools effectively to study reproduction. Large amounts of information have become available through internet-accessible databases, and these will become increasingly important in the future. We need to become familiar with genomics tools and design experiments to understand better the reproductive system, that most dynamic set of horse tissues.
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Relevance of Genomics to Equine Reproduction
References 1. Murray JD. Horse genomics and reproduction. Theriogenology 2002;58:281–93. 2. Penedo MCT, Millon LV, Bernoco D, Bailey E, Binns M, Cholewinski G, Ellis N, Flynn J, Gralak B, Guthrie A, Lindgren G, Lyons LA, Tozaki T, Røed K, Swinburne J. International Equine Gene Mapping Workshop Report: A comprehensive linkage map constructed with data from new markers and by merging four mapping resources. Cytogenet Genome Res 2005;111:5–15. 3. Swinburne JE, Boursnell M, Hill G, Pettitt L, Allen T, Chowdhary B, Hasegawa T, Kurosawa M, Leeb T, Mashima S, Mickelson JR, Raudsepp T, Tozaki T, Binns M. Single linkage group per chromosome genetic linkage map for the horse, based on two three-generation, full-sibling, crossbred horse reference families. Genomics 2006;87: 1–29. 4. Chowdhary BP, Raudsepp T, Kata SR, Goh G, Millon LV, Allan V, Piumi F, Guerin G, Swinburne J, Binns M, Lear TL, Mickelson J, Murray J, Antczak DF, Womack JE, Skow LC. The first generation whole genome radiation hybrid map in the horse identifies conserved segments in human and mouse genomes. Genome Res 2003;13:742–51. 5. Lear TL, Bailey E. Equine clinical cytogenetics: past and future. Cytogenet Genome Res 2008;120:42–9. 6. Pulos WL, Hutt FB. Lethal dominant white in horses. J Hered 1969;60:59–63. 7. Hintz HF, van Vleck LD. Lethal dominant roan in horses. J Hered 1979;70:145–6. 8. Leeb T. The Horse Genome Project – sequence based insights into male reproductive mechanisms. Reprod Domest Anim 2007;42(Suppl. 2):45–50. 9. Hamann H, Jude R, Sieme H, Mertens U, Topfer-Petersen E, Distl O, Leeb T. A polymorphism within the equine CRISP3 gene is associated with stallion fertility in Hanoverian warmblood horses. Anim Genet 2007;38:259–64. 10. Chowdhary BP, Paria N, Raudsepp T. Potential application of genomics in dissecting diseases and fertility. Anim Reprod Sci 2008;107:208–18.
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11. Rudolph J, Spier S, Byrns G, Rojas C, Bernoco D, Hoffman E. Periodic paralysis in Quarter Horses, a sodium channel mutation disseminated by selective breeding. Nat Genet 1992;2:144–7. 12. Shin EK, Perryman LE, Meek K. A kinase negative mutation of DNA-PKcs in equine SCID results in defective coding and signal joint formation. J Immunol 1997;158:3565–9. 13. Mariat D, Taourit S, Guerin G. A mutation in the MATP gene causes the cream coat color in the horse. Genet Sel Evol 2003;35:119–33. 14. Cook D, Brooks SA, Bellone R, Bailey E. Missense mutation in exon 2 of SLC36A1 responsible for champagne dilution in horses. PLOS Genet 2008;4 (9): e1000195. doi:10.1371/journal.pgen.1000195 15. Pielberg GR, Golovko A, Sundstrom E, Curik I, Lennartsson J, Seltenhammer MH, Druml T, Binns M, Fitzsimmons C, Lindgren G, Sandberg K, Baumung R, Vetterlein M, Stromberg S, Grabherr M, Wade C, Lindblad-Toh K, Ponten F, Heldin C-H, Soklner J, Andersson L. A cis-acting regulatory mutation causes premature hair graying and susceptibility to melanoma in the horse. Nat Genet 2008; doi: 10.1038/ng.185 16. Simpson KS, Adams MH, Behrendt-Adam CY, Baker CB, McDowell KJ. Identification and initial characterization of calcyclin and phospholipase A2 in equine conceptuses. Mol Reprod Dev 1999;53:179–87. 17. McDowell KJ, Adams MH, Adam CY, Simpson KS. Changes in equine endometrial oestrogen receptor alpha and progesterone receptor mRNAs during the oestrous cycle, early pregnancy and after treatment with exogenous steroids. J Reprod Fertil 1999;117:135–42. 18. Dascanio J, Tschetter J, Gray A, Bridges K. Use of quantitative real-time polymerase chain reaction to examine endometrial tissues. Anim Reprod Sci 2006;94:254–8. 19. Gu W, Bertone AL. Generation and performance of an equine-specific large scale gene expression microarray. Am J Vet Res 2004;65:1664–73. 20. Ramery E, Closset R, Bureau F, Lekeux TP. Relevance of using a human microarray to study gene expression in heaves-affected horses. Vet J 2007;177:206–21.
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Donkey Reproduction Robyn R. Wilborn and David G. Pugh
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Miniature Pony Reproduction Paul J. De Vries and Dale L. Paccamonti
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Asian Wild Horse Reproduction: Equus Ferus Przewalskii C. Wynne Collins, Lee E. Boyd and Katherine A. Houpt
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Chapter 299
Donkey Reproduction Robyn R. Wilborn and David G. Pugh
Introduction Donkeys (Equus asinus) are used throughout the world as guard animals for protecting small ruminants (sheep, goats, llamas), as companion animals (for people, foals, etc.), for halter training (calves, foals), work (pack, pulling wagons, plowing, etc.), and for riding and showing. They are also bred to horses to produce mules or hinnies. Donkey medicine can be fun and rewarding and can provide a break from routine equine practice.1 Donkeys come in many breed types and sizes, from the miniature to the mammoth. The size differences may hamper the practitioner’s ability to perform certain diagnostics (e.g., examination per rectum) or therapeutics (e.g., floating teeth). The ‘mini’ or Mediterranean miniature donkey originates primarily from Sardinia and Sicily and stands up to 80 cm (35 in) tall at the withers. These donkeys are characterized by a dorsal cross. A burrow is a smaller member of the donkey family whose parentage is usually traced to Spain/Mexico.1 The standard donkey will stand 90–120 cm (36–48 in), while the large standard will be 120–137 cm (48–54 in). Mammoth donkeys stand more than 137 cm (54 in; 13.5 hands) for females and over 142 cm (56 in; 14 hands) for males. The rare French Poitou breed, which now has fewer than 250 survivors worldwide, is usually 14.5–15.0 hands and often has a characteristic long, thick haircoat. Andalusians, whose colors are gray to white (and occasionally black or blue), also stand 14.5–15.0 hands. The Arabic Maltese, Spanish Catalonian, Andalusian, and French Poitou were all used in the historical development of the American mammoth or mammoth jack stock.2 The cross between a jack and a mare will almost always be sterile and is referred to as a mare mule or molly mule (female) or horse, stud, or john mule (male). The mule’s ears will be smaller than those of the jack, but longer than those of the mare, with head,
hip, and neck more jack-like, and a thin forelock (if present). The mule will reflect its parentage; that is, a gaited or refined jack crossed with a Tennessee walking horse may produce a very refined, gaited mule. Of course, a mammoth jack bred to a Belgian mare will likely produce a ‘beast of burden’. The hinny, which is a much less common animal than the mule, is the cross between a stallion and a jennet (jenny). This breeding results in a conception to birth rate of usually less than 20%.1, 2
Jenny reproduction The jenny is very similar in many reproductive aspects to the horse mare.1 Puberty is usually attained at 1–2 years, and like other species, may be associated with body size. Jennies can usually be bred after they have attained 60–70% of their projected mature bodyweight, with a body condition score of no more than 6–7 (1–9 scale). Her reproductive life will rarely exceed 16–18 years. Donkeys seem to display less seasonality than that of the horse.3, 4 Wisconsin workers4 reported a lower incidence of ovulations during December, prolonged estrus upon return to cyclicity, shorter estrous cycles from May to September, and shorter behavioral estrus from May through October. Others5 have reported irregular estrus periods during the traditional anovulatory season. Still, these workers described less ‘partitioning’ in the ovulatory and anovulatory (non-breeding) season of the donkey than in the horse.4, 5 The cervix of the jenny is usually longer than that of the mare, with a smaller diameter that enlarges with age. Cervical size in mature Poitou jennies has been reported to be 4.5–8 cm in length and 2.5–3.0 cm in diameter.6 Estrus-induced softening of the cervix is less pronounced in older jennies than in young jennies. The donkey cervix protrudes into the vagina, creating a very prominent fornix. These
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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cervical characteristics may preclude intrauterine ejaculation, make artificial insemination more difficult, and may be associated with a high incidence of post-dystocia cervical adhesions.1, 6 Although the estrous cycle will range from 20 to 40 days, it usually lasts 23–30 days.4, 7 Estrus behavior has been characterized by mouth opening and closing or chomping with salivary dribbling, neck extended with ears back, clitoral winking, urinating, tail rising,8, 9 and lowering of the hindquarters. This chomping or mouth clapping behavior appears more common and consistent than clitoral winking. Diestrous behavior includes running away and/or trying to kick the jack, and clamping the tail. Estrus averages between 6 and 9 days, with ovulation occurring 5–7 days after the onset of estrus.7, 9 Approximately 80% of jennies will ovulate a follicle of diameter 25–35 mm or greater within 48 hours prior to the end of behavioral estrus.1, 5 Postestrus ovulation has been reported but is considered to be uncommon.5 If transrectal palpation or ultrasonographic evaluation is performed, follicles > 25–30 mm should be considered to be consistent with imminent ovulation.10 Multiple ovulations per estrus can occur in all donkey breeds, but are reported to be more frequent in mammoth jennies.3 This breed difference may be similar to the differences in multiple ovulation among horse breeds (e.g., Arabians vs Thoroughbreds).3 Prolonged anovulatory periods have been associated with persistent corpora lutea.4
Breeding and synchronization Jennies can be pasture bred or hand bred. When a hand breeding program is employed, either with natural breeding or artificial insemination, an effective teasing program will enhance its success.3 Texas workers3 demonstrated that estrus synchronization protocols useful in horses appear effective in the donkey. This group was successful in using prostaglandin F2α (PGF2α ) alone (two injections, 16 days apart) or a combination of progesterone (P), estradiol-17β (E) and PGF2α (P&E daily for 10 days, then PGF2α on Day 10) for synchronization of estrus and ovulation. If natural breeding is used without the aid of transrectal evaluation of ovarian status, jennies should be mated the second day of estrus, and then at 48-hour intervals until the end of ‘standing heat’.7 During coitus, a jenny will continue to yawn and occasionally move forward a few steps. This movement or walking may prove troublesome when using some jennies as ‘jump’ animals for semen collection or in hand-breeding programs.
Pregnancy The transrectally identified characteristics of pregnancy are very similar to those of the mare, both from a palpation and ultrasonographic standpoint. The embryonic vesicle can be detected as early as Days 9 to 11 post-ovulation.10 Embryonic mobility in the jenny occurs until Days 13 to 18 post-ovulation. Brazilian workers10 report that the embryonic vesicle is spherical up until Day 18, with the fetal heartbeat detectable by Day 23. Reproductive management of the jenny is similar to that of the mare. The author (Pugh) prefers to vaccinate on the 3rd, 5th, 7th, and 9th months of gestation for equine herpes virus (EHV) type 1, and at 1 month prepartum for tetanus, Eastern and Western encephalitis, West Nile virus, and influenza. Jennies have been shown to be susceptible to the abortigenic effects of EHV-1.11 Leptospira antibodies were shown in a jenny with a history of abortion, although this organism is considered to be an uncommon cause of reproductive loss in donkeys.12 In the pregnant jenny, secondary corpora lutea are formed from Day 38 to 46, and hormonal profiles are similar to mares with regard to progestogens and estradiol.13 However, a great deal of variation exists, with equine chorionic gonadotropin (eCG) levels depending on the genotype of the dam and fetus.14 A jenny carrying a donkey fetus produces levels of eCG that are only one-half to one-third that of a mare carrying a horse fetus. A mare carrying a mule fetus produces even lower levels of eCG, equating to less than one-quarter that of mares carrying a horse fetus. In contrast, a jenny carrying a hinny fetus produces the highest concentration of eCG, which is twice that of a normal mare pregnancy and 3–4 times higher than a normal jenny pregnancy with a donkey fetus.14 Gestational length has been reported to average 372 to 374 days.7, 15, 16 Prepartum behavior is similar to a mare’s in that the jenny may become restless, and have engorged teats and a lengthened vulva. Jennies will usually lie down to deliver the foal. Post-dystocia vaginal and cervical injury appears to be more common than in the mare.1 Foal heat usually occurs between 5 and 13 days postpartum.1 Postpartum uterine involution and cyclicity in jennies has been shown to be similar to that described for mares.17
Pathology In Pugh’s experience, the incidences of clinical metritis, endometritis, and retained fetal membranes are less than in the horse.1 However, mild to severe endometritis was reported in 47% of endometrial samples from 165 jennies.18 Vaginal prolapse in a
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Donkey Reproduction ‘molly’ mule is not an uncommon medical condition. This author performs an epidural, carefully replaces the vagina, and places a modified Buhner or deep Caslick’s suture to close the vulva.1 As in the mare, ovarian tumors have been encountered. These include cystadenoma19 and granulosa cell tumor.20
Jack reproduction Like the jenny, the jack has many reproductive similarities to the horse. However, some differences do exist. The testes and penis of the jack are larger than that of comparably sized stallions.21 Burnham recently reported that the penile vasculature was similar, but a donkey penis was longer (50.7 ± 1.6 cm) than that of a horse (46.45 ± 1.1 cm).22 The testicular size of mature 250-kg donkeys is probably comparable to that of a 400-kg horse, and the jack has larger epididymal tails.16 Although jacks and stallions have the same accessory sex glands, the ampullae are larger in the jack,1 and the seminal colliculus appears to be absent in the jack.23 One peculiarity of donkey reproduction is the longer time required for the jack to achieve an erection and ejaculate. The teasing behavior of the jack is somewhat different than the stallion’s.1 At the initiation of teasing, a jack may vocalize to all the females prior to turning attention to only one.24 During this courtship phase, he may also sniff or nibble the female’s vulva and perineal area, nibble at her head, neck, flank, or legs, exhibit a flehmen reaction, and mount the female.1, 25 The jack may run or walk forward toward the female, then retreat from her several times during the teasing period. Gastal et al.25 reported that when jacks first mount the female, approximately 70% of the time they have a non-erect, yet exposed, penis. Under pasture-mating conditions, the jack may dismount and begin to graze prior to resuming the courtship phase, and may attain more than one erection prior to final mounting and ejaculation.24 The jack is occasionally very aggressive in his teasing behavior, and may continue to pursue and mount jennies in diestrus, even after being kicked repeatedly.1 Jacks usually require 5–30 minutes (compared to 10–11 minutes in the stallion) from teasing to completion of ejaculation.21, 25 The jack will have approximately five pelvic thrusts, as compared to the stallion’s seven.24 Ejaculation will take 6–30 seconds, and is usually accompanied by 4–9 ejaculatory urethral waves, with a resulting seminal volume of 10–80 mL, 80–88% live spermatozoa, and a pH of 7.6.21, 26 As a general rule, jack ejaculates have a higher sperm concentration and a lower semen volume than that of the stallion. The gel portion of the ejaculate is smaller
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and its presence is more variable compared to a stallion’s.27 The gel fraction is more commonly encountered as a component of the ejaculate in older jacks.21 The spermatozoa of the jack have a similar shape to those of the stallion, but appear to have a larger and more rounded head, and a slightly longer tail. Although some seasonal differences in libido are observed, most workers report little or no alteration in seminal parameters during the winter months.21, 25 However, Gastal et al.26 documented a slight decrease in motile spermatozoa and a lower seminal pH during January and February in Brazil.
Semen collection and processing As with the stallion, the jack can be trained to service an artificial vagina, and semen can be collected for use in fresh, cooled, or frozen insemination programs.21,25–27 Donkey semen should be handled using the same techniques employed to ensure good stallion semen quality with regard to temperature, light, and hygiene. Donkeys and mules in some countries have tested positive for equine viral arteritis (EVA),28 so appropriate testing should be performed as a precaution prior to shipping or freezing semen. Contagious equine metritis (CEM) should also be considered, especially if importing or exporting semen. Although the clinical presentation of the disease is the same as in the horses, the organism isolated from infected donkeys is termed Taylorella asinigenitalis.29, 30 Skim-milk extenders can prove useful in artificial insemination (AI) programs using fresh donkey semen.31 As with the stallion, the use of different semen extenders and/or antibiotic combinations should be explored in order to minimize spermatozoal damage during storage, cool shipping, or freezing. Brazilian workers27 demonstrated that an extender containing egg yolk (i.e., Baken extender) is better suited than a skim milk-type extender for cooled storage of jack semen. Further, if Baken extender is modified by increasing the amount of egg yolk (3%), it may be superior for maintaining jack semen at a cooled temperature of 5◦ C.32 These workers also concluded that jack semen may be more resistant to cold shock than stallion semen. Spanish workers33 suggested that spermatozoal motility following cooled storage may be improved by dilution with a skimmilk extender, followed by centrifugation (400g for 7 min), and resuspension of the sperm pellet in an extender to a final dilution of 50 × 106 spermatozoa per mL. In this study, spermatozoal plasma membrane integrity, as determined by a hypo-osmoticswelling test, was superior in samples where the seminal plasma had been removed via centrifugation.33
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In contrast, other studies27 have not demonstrated the harmful effects of seminal fluid on the motility of jack semen when cooled and stored. It is possible that individual jacks must be handled differently with regard to seminal cooling and shipping, as is seen with stallions. This author (Pugh) prefers to dilute jack semen to a concentration of 25–50 × 106 with a proven skimmilk-based extender.1, 34 In cases where post-cooling spermatozoal motility is less than 30% within 24 hours of collection, and the antibiotic choice is not found to be a problem, removal of the seminal plasma via centrifugation (300g for 5–10 min in a conicalshaped tube) and resuspension with a skim-milkbased extender should be considered. Many workers have published freezing protocols for donkey semen.35, 36
References 1. Pugh DG. Donkey medicine. Proc Am Assoc Eq Pract 2002;48:21–3. 2. Lange BA. An explanation of the foundation breeds of American mammoth jacks and jennets. Am Mammoth Jack Stock J 1989;2:28–30. 3. Blanchard TL, Taylor TS, Love CL. Estrous cycle characteristics and response to estrus synchronization in mammoth asses (Equus assinus Americanus). Theriogenology 1999;52:827–34. 4. Ginther OJ, Scraba ST, Bergfelt DR. Reproductive seasonality of the jenny. Theriogenology 1987;27:587–92. 5. Henry M, Figueiredo AEF, Palhares MS, Coryn M. Clinical and endocrine aspects of the estrous cycle in donkeys (Equus assinus). J Reprod Fertil 1987;35:297–303. 6. Vendramini OM, Guintard C, Moreau J, Tainturier D. Cervix conformation: a first anatomical approach in Baudet du Poitou jenny asses. Anim Reprod Sci 1998;66:741–4. 7. Ferlding D. Reproductive characteristics of the jenny donkey (Equus assinus): a review. Trop Anim Health Prod 1988;20:161–6. 8. Clayton RM, Lindsay FEF, Forbes AC, Hay LH. Some studies of comparative aspects of sexual behavior in ponies and donkeys. Appl Anim Ethol 1981;7:169–74. 9. Vandeplassche GM, Wesson JA, Ginther OJ. Behavioral, follicular and gonadotropin changes during the estrous cycle in donkeys. Theriogenology 1981;16:239–49. 10. Meira C, Ferreira JCP, Papa FO, Henry M. Ultrasonographic evaluation of the conceptus from days 10 to 60 of pregnancy in jennies. Theriogenology 1998;49:1475–1482. 11. Tewarl SC, Verma PC, Bhargava DN. Experimental equine herpes virus-1 infection of foetus in a pregnant donkey mare. Indian Vet J 1994;71:213–14. 12. Arora BM, Baxi KK. Sero-evidence of leptospirosis in equidae, suidae, rodentia and lagomorpha. Indian J Vet Path 1979;3:52–3. 13. Meira C, Ferreira JCP, Papa FO, Henry M. Ovarian activity and plasma concentrations of progestogen and estradiol during pregnancy in jennies. Theriogenology 1998;49:1475–3. 14. Allen WR. Factors influencing pregnant mare serum gonadotropin production. Nature 1969;223:64–6. 15. Clarke C. Mules must come to science. New Sci 1983; 3:300–3.
16. Kohli ML, Suri KR, Chatterji A. Studies on the gestation length and breeding age in donkey mares. Indian Vet J 1957; 34:344–8. 17. Dadarwal D, Tandon SN, Purohit GN, Pareek PK. Ultrasonographic evaluation of uterine involution and postpartum follicular dynamics in French Jennies (Equus asinus). Theriogenology 2004;62:257–64. 18. Sokkar SM, Kamouda MA, El-Rahmani SM. Endometritis in the donkeys in Egypt. J Vet Med Ser B 2001;48:529–36. 19. Taylor TS, Bowea JM, Montgomery DL. Cystadenoma of the ovary in a jennet. Comp Cont Ed Pract Vet 1983;5:5677– 9. 20. Norris HJ, Taylor HB, Garner FM. Equine ovarian granulosa tumours. Vet Rec 1968;82:419–20. 21. Kreuchauf A. Reproductive physiology in the jackass. Anim Res Dev 1984;20:51–78. 22. Burnham SL. Anatomical differences of the donkey and mule. Proc AAEP 2002;48:1002–10. 23. Shehata R. A short study of the seminal colliculus in some mammals. Acta Anat 1980;106:400–4. 24. Henry M, McDonald SM, Lodi LD, Gastol EL. Pasture mating behavior of donkeys (Equus assinus) at natural and induced oestrus. J Reprod Fert Suppl 1991;44:77–86. 25. Gastal MO, Henry M, Becker AR, Gastal EL, Goncalves A. Sexual behavior of donkey jacks: influence of ejaculatory frequency and season. Theriogenology 1996:46:539–603. 26. Gastal MO, Henry M, Becker AR, Gastal EL. Effect of ejaculation frequency and season on donkey jack semen. Theriogenology 1997;47:627–38. 27. Mello SCV, Henry M, Souza MC, Oliveira SMP. Effect of split ejaculation and seminal extenders on longevity of donkey semen preserved at 5◦ C. Med Vet Zoo 2000;52:372–8. 28. Paweska JT, Binns MM, Woods PSA, Chirnside ED. A survey for antibodies to equine arteritis virus in donkeys, mules and zebra using virus neutralization (VN) and enzyme linked immunosorbent assay (ELISA). Equine Vet J 1997;29:40–3. 29. Jang SS, Donahue JM, Arata AB, Goris J, Hansen LM, Earley DL, Vandamme PAR, Timoney PJ, Hirsch DC. Taylorella asinigenitalis sp. nov., a bacterium isolated from the genital tract of male donkeys (Equus asinus). Int J Syst Evol Microbiol 2001;51:971–6. 30. Matsuda M, Moore JE. Recent advances in molecular epidemiology and detection of Taylorella equigenitalis associated with contagious equine metritis (CEM). Vet Microbiol 2003;97:111–22. 31. Singhui NM. Studies on artificial insemination in equines. Int J Anim Res 1990: 11; 99–104. 32. Cottorello ACP, Amancio RC, Henry M, Borges I. Effect of storage temperature and extenders on ‘in vitro’ activity of donkey spermatozoa. Theriogenology 2002;88:325–8. 33. Serres C, Rodriguez A, Alvarez AI, Santiago I, Gabriel J, Gomez-Cuetara C, Mateos E. Effect of centrifugation and temperature on the motility and plasma membrane integrity of Zumorano-Leonies donkey semen. Theriogenology 2002;58:329–32. 34. Pugh DG. Male donkey reproduction. In: Proceedings of the Western Veterinary Conference, 2004. Veterinary Information Network (www.vin.com). 35. Silva SS, Henry M, Nunes SA, Mello SCV. Influence of packaging system on quality of frozen-thawed donkey spermatozoa (Equus assinus) evaluated ‘in vitro.’ Rev Bras Repro Anim 1997;21:140–6. 36. Trimeche A, Renard P, Tainturier D. A procedure for poitou jackass sperm cryopreservation. Theriogenology 1998: 5:793–806.
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Chapter 300
Miniature Pony Reproduction Paul J. De Vries and Dale L. Paccamonti
Introduction Although Miniature horses belong to the Equidae family and the Equus genus there are quite a few differences, besides their size, between them and their larger family members. Some of those differences are in the reproductive field. Unfortunately, not many studies have been performed to pinpoint these differences. Therefore, personal experiences and opinions from several workers with Miniature horses together with the available information from the literature have been used for this chapter. The origin of Miniature horses is mainly based on original Shetland ponies that were selected on their size and that were crossed with smaller breeds such as the Falabella horse. Miniature horses are mainly bred in North and South America and in Europe. The height of a true Miniature should not exceed 86 cm (34 inches). Some Miniature horse-breeding organizations also recognize mares between 86 cm and 96 cm (38 inches). The differences between a Miniature horse and a pony are not always clear. The official size for ponies within the FEI (F´ed´eration Equestre Internationale; the international sport horse organization) rules is less than 148 cm (58.3 inches).
The estrous cycle The onset of the breeding season in Miniature mares is often somewhat delayed compared with other mares. Differences in housing, feeding, and the lack of artificial light may partially explain this, as many Miniatures are kept more extensively than other mares. Additionally, because ponies are more clearly seasonal than horses,1 i.e., the vast majority have a seasonal anestrus and begin cycling later in the season, the effect seen in Miniature horses may also be due to their ancestry. The clinical impression of many working with Miniature horses is that they have a longer estrous
period during the breeding season than normal-sized horses.2 The interovulatory interval (IOI) in Miniatures was about 3 days longer than the IOI of normalsized horses (23.3 ± 0.9 vs 20.3 ± 0.7). The longer interval appears to be mainly caused by a longer period of the follicular phase rather than a longer period of diestrus. Ginther et al.3 also found that progesterone levels declined about 2 days earlier in Miniature mares (day 12) than in normal-sized mares (day 14). The size of the pre-ovulatory follicles was smaller in Miniature (38.3 mm) mares than in normal-sized mares (44.5 mm). The differences in size of the preovulatory follicles between the Miniatures and the normal-sized horses were, however, less than could be expected based on the relative difference in size or bodyweight of the mares.3 The difference in preovulatory follicular diameter is obviously of clinical importance when the size of a follicle is used as an indicator for the time of breeding or insemination. There are differences in endometrial edema during estrus between Miniatures and normal-sized mares. Miniatures tended to start showing less edema 4 days prior to ovulation, in contrast to normal-sized mares at about 1–2 days before ovulation. This is not in agreement with the general opinion that the score for uterine edema runs parallel with the level of estradiol. Ginther et al.3 found that in Miniature mares estradiol levels dropped about 2 days after the decrease of edema.
Breeding management The majority of Miniature mares are still bred by natural cover since the easiest way to keep a breeding stallion is together with a number of mares in the same pasture. The main advantage of this system is that it is not very labor intensive and under normal circumstances should lead to excellent fertility results. Obviously there is a limit to the number of mares (10–12) that a stallion can handle, and the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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stallion and the mares must be kept in contact for several weeks. A continuous change of mares in a group will lead to suboptimal results, because the stallion will focus his attention on the newer mares. The lack of human surveillance is the main disadvantage of the system, and the planning of pregnancy examinations and predicting the date of foaling is difficult, because of the unknown breeding data. Hand breeding is more labor intensive, but will lead to better information about the date for pregnancy examinations and the expected day of delivery. Artificial insemination is also used for the breeding of Miniature horses. Manual insemination is usually very well tolerated by the Miniature mares, but, if necessary, insemination through a small speculum can be used. Embryo transfer has been used in Miniatures as well, and the only differences from embryo transfer in normal-sized mares is the use of a smaller flushing catheter (8.7 mm, 26 FR/CH) and the reduced volume of fluid (about 500 mL) used to flush the mares. The reported size of the collected embryos on day 8 or 9 was smaller than embryos of the same age from normal-sized mares.4 Embryos collected on day 8 were 0.6 ± 0.21 mm in diameter,4 which is about half the size of embryos of the same age from normalsized mares. Great care should accompany interpretation of these data as only 11 flushes occurred for a total of six embryos (three at day 8, one at day 9, and two at day 10). Most Miniature breed organizations will allow artificial insemination using transported fresh or frozen semen and embryo transfer after acquisition of a permit. This leads to a wider potential of breeding stallions and breeding management tools. Permits usually include some conditions.
Pregnancy diagnosis The early diagnosis of pregnancy is an important tool in horse breeding. Although several methods have been used in the past and some of them are still used, not all of them are reliable. Some methods that are used for pregnancy detection only provide evidence that the mare is not in estrus. Repeated teasing of the mare, around at least 21 days after breeding, is an inexpensive tool but is a reliable indication for nonpregnancy only if the mare shows signs of estrus. Determination of blood or milk progesterone level for pregnancy detection around 21 days after breeding is about 80% reliable for pregnancy detection if progesterone levels are above 2 ng/mL, and 100% reliable for non-pregnant mares if levels are lower than 2 ng/mL.5 More reliable methods are ultrasonographic examination, rectal palpation, and repeated blood hor-
mone level determination. Because pasture breeding is often used in Miniature horse breeding, the actual breeding data are often unknown. This means that the results of an early pregnancy examination are not always reliable. Ultrasonography is a reliable method for pregnancy detection, especially if the mare has also been examined when known to be not pregnant. Possible endometrial cysts can then be noted, and confusion between an embryonic vesicle and a cyst during the pregnancy examination can be minimized. Transrectal ultrasonography is possible in many Miniature mares, depending on the cooperation and age of the mare, her parity, and the size of the linear probe and the hands and wrists of the examiner. Petroleum jelly is preferable as a lubricant because of its stickiness to the glove in the narrow anal sphincter. Proper restraint is essential, and, if the Miniature mare can be placed on a platform (equipped with a detachable ramp), the examiner can palpate the mare in a normal body position. Transrectal ultrasonography can be used on pregnant Miniature mares from around day 12 after ovulation. In the first month of pregnancy sizes of the embryonic vesicle are usually only slightly smaller than those of normal-sized mares.6 In Miniature mares it is usually the anal sphincter that causes more difficulties with palpation than the rectum. Low-dose sedation can be used, if necessary, together with a lukewarm watery enema and an anal infusion of lidocaine (about 60 mL of a 2% solution). If necessary, distension of the rectum can be achieved by infusing more lukewarm water. Rarely is it necessary to use smooth muscle relaxants such as N-butylscopolammonium bromide (0.3 mg/kg intravenously (i.v.)) or propantheline bromide (0.014 mg/kg i.v.). If the size of the mare is too small to insert a hand into the rectum, an extension handle on the ultrasound probe can be used to perform an ultrasonographic examination of the reproductive tract. This extension is easily made out of 2-cm-diameter PVC pipe of about 40 cm (16 inches) length, with a curve at the probe end of 15–20◦ (Figure 300.1).
Figure 300.1 Probe extension made from PVC pipe.
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tween 35 and 120 days of gestation are the origin of eCG. Presence of eCG is a reliable indicator of a pregnancy; however, following pregnancy loss after 40 days of gestation, eCG can continue to be produced in the absence of a live fetus. Therefore, a positive eCG test should be followed by an estrone sulfate (ES) test after 75 days of pregnancy using blood,8 or after 150 days of pregnancy using feces. Pregnant mares usually, but not always, have higher ES levels than nonpregnant mares, and levels in blood plasma are more discriminative than fecal levels.8, 9 Figure 300.2 Specula made of stainless steel with rounded edges. Cross-sections are: smaller 2.5 cm (1 inch); larger 4.0 cm (1.6 inch).
An alternative to transrectal ultrasonography is transabdominal ultrasonography. This can be performed at a later stage of pregnancy, starting from about 30 days post-ovulation. The earliest date is dependent on the experience of the examiner, the body condition of the mare, and the size of the bladder and mammary gland. At early stages the best results are obtained with a ventral midline approach. In more advanced pregnancies the fetus can be visualized with a lower flank approach. A standard 5MHz linear array probe can be used, although in the earliest stages a 3.5-MHz transducer may provide a better image. Clipping and using a coupling gel provides the best contact between the transducer and the skin. In miniature donkeys, it has been shown that clipping is not always necessary, and that alcohol can very well be used as a coupling medium.7 The use of alcohol may, however, be damaging to some transducers and cables. Large amounts of subcutaneous fat are detrimental to the image. Emptying the bladder may help for easier identification and a better image of the uterus. Transabdominal ultrasonographic pregnancy detection is more reliable at later stages of pregnancy (from day 70) when moving fetuses can be visualized. Repeated vaginal inspection (or palpation) of the cervix is also possible as a method for pregnancy detection. Vaginal inspection using an appropriately sized sterile (prewarmed) speculum (Figure 300.2) and a light source is well tolerated by Miniatures; care must be taken, however, to work as hygienically as possible. Use a carboxymethylcellulose-based lubricant or clean water as lubricant to prevent vaginitis and possible adhesions. There are several blood hormone assays for the detection of pregnancy in Miniatures. Progesterone has been discussed above. Between 40 and 140 days of gestation equine chorionic gonadotropin (eCG) is a pregnancy-specific hormone that can be detected in blood samples. Endometrial cups that are present be-
Pregnancy and abortion The duration of pregnancy in Miniatures is generally shorter than that of normal-sized mares.6, 10 Most causes of pregnancy failure in the Miniature mare do not differ from those of normal-sized mares, although fetal malformation,6 hyperlipemia, and associated hepatic lipidosis are a relatively common cause of abortion in the late-term pregnant Miniature mare. Dwarfism is one type of fetal malformation seen in Miniatures and is considered to be inherited, although the exact mechanisms of inheritance have not been revealed owing to the reluctance of owners to provide information on the incidence of dwarfs and the consequently limited research. Signs of dwarfism are oversized heads in relation to body, bulging foreheads, mandibular prognathia, and angular limb deformities (Figure 300.3a,b). Hyperlipemia is a common disease in Miniatures (Figure 300.4). It results from anorexia initiated by a variety of primary causes. Enterocolitis (sand, impactions), internal parasites, diarrhea, and other causes, such as fever, can lead to lethargy and anorexia. Fat (triglycerides, fatty acids, and cholesterol) is mobilized from the body fat stores, especially in overweight Miniatures. This leads to increased levels of fat in the blood, and consequently further anorexia and lethargy. If the disease persists, the storage of fat in the liver will culminate in hepatic lipidosis. Pregnant mares with hyperlipemia and hepatic lipidosis will usually abort when adequate treatment is not started in a timely manner.
Parturition The conventional signs of an approaching parturition are seen in Miniatures too, all in relation to their size. The udder development and waxing, however, is much less than in normal-sized mares.6 Milk electrolytes can be determined as in normal-sized mares to predict parturition in Miniature mares (see Chapters 232 and 286). However, in Miniature mares it may be very difficult to obtain sufficient secretion to be tested, even if collection is attempted a few hours
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(a)
Figure 300.4 Hyperlipemia: note the turbid aspect of the serum.
(b) Figure 300.3 (a) Late-term aborted miniature fetus with bulging forehead and (b) misalignment of the upper and lower jaw (brachygnathia superior).
before parturition (D.L. Paccamonti, unpublished observation). It is not uncommon for changes in the udder and milk electrolytes to occur rapidly. Because the dates of ovulation/conception of many Miniature mares are unknown, the expected day of delivery is often unknown as well. Turner et al. 11 have found that the measurement of the eye of the fetus in pony mares has a good predictive value for the days to parturition (Table 300.1). The mares that were used were between 104 and 124 cm (41 and 49 inches), but there appeared to be no differences in fetal eye sizes and days before parturition between the smallest and the largest mares in the group. Gender of the foal, parity of the mare, and body condition score of the mare were of no significant influence in the study. Differences in eye size and skull width ratio in other types of Miniatures may, however, influence the results that were found. In Miniatures the incidence of dystocia is higher than in normal-sized mares. The size of Miniatures in relation to the size of their foals is most likely the cause of this. Limb deformities and contracted tendons (Figure 300.5) are relatively common in
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Table 300.1 Predicted time to parturition based on measurement of the fetal eye by transabdominal ultrasonography for pony mares. Reprinted with permission from Turner et al.11 Days before parturition Eye length (mm) 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35
Mean
75% CI
95% CI
209 203 197 190 183 175 167 158 150 140 131 121 110 99 88 77 65 52 39 26 13
184–251 177–245 170–238 153–230 155–223 147–214 138–206 129–197 120–187 110–177 100–167 89–156 78–145 66–133 54–121 42–109 29–96 16–83 2–69 0–55 0–41
160–275 153–269 146–262 140–254 131–238 123–238 115–230 106–220 96–211 86–201 76–191 65–180 54–169 43–157 31–145 18–133 5–120 0–107 0–93 0–79 0–64
CI, confidence interval.
Miniatures, and even normal foals frequently have bulging foreheads. These abnormalities will often result in an abnormal fetal position, posture, and fetopelvic disproportion (Figure 300.6). Repositioning the foal is often challenging owing to the limited space. Fetotomy is difficult for the same reason. In
Figure 300.6 Dystocia in a Miniature mare: a unilateral shoulder flexion.
Miniature mares prolonged manipulation in the birth canal will often lead to severe cervicitis and vaginitis and consequential adhesions. In awkward dystocias cesarean section is often the only option. Preferably this is performed in a veterinary hospital. The distance from the farm to the hospital is often unfavorable to the survival of the foal. With an adequate technique an excellent mare survival score can be achieved.12 If necessary and with competent assistance surgery can be performed on the farm if a lower left flank or a left paracostal approach13 is employed (Figure 300.7). The major advantage of these techniques is that the mare can be positioned in right lateral recumbency. In case of a live foal, competent care should be available directly after birth of the foal for its optimal survival chances.
Postpartum complications
Figure 300.5 Severe flexural deformity (arthrogryposis) in a Miniature foal.
In Miniature mares specific postpartum complications are relatively common. Campbell,6 Judd,14 and Ginther and Williams15 have described a higher incidence of suffocation in the newborn foals of Miniatures due to the lack of rupture of the amniotic membrane.
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Figure 300.7 Paracostal incision scar after cesarean section in lateral recumbency.
Postpartum colic was also noted in about 30% of the Miniature mares, probably caused by painful oxytocin-induced uterine contractions.14 Flunixin meglumine as a single dose of 1 mg/kg i.v. is usually sufficient to keep the mare comfortable and give the foal adequate opportunity to nurse. Retained placenta in Miniatures is relatively common as well. Veterinary treatment should commence no later than 6 hours postpartum. Before that time, hand walking of the mare may result in a spontaneous expulsion of the placenta. By 6 hours after parturition treatment should be started. Oxytocin (10–20 IU in 250 mL saline or lactated Ringer’s i.v. during 10 minutes), if necessary repeating every hour with an increase of the oxytocin dose, is successful in about 50% of the cases. The Burns technique16 (instilling lavage fluid into the intact chorioallantoic space) is also frequently successful if the placenta is intact. If calcium blood levels are low, the oxytocin can be added to a calcium solution. Manual traction on the placenta is contraindicated because of the risk of uterine horn inversion and uterine prolapse (Figure 300.8). If the placenta is not expelled within 8 hours, removal of the placenta by atraumatic and careful manual loosening of the connection between the chorion and endometrium is indicated. After manual removal of the complete placenta, repeated uterine lavage with a large-bore catheter or nasogastric tube and 5–10 liters of clean warm water with 9 g/l sodium chloride added is performed until the returning fluid is clear. In severe cases this should be repeated for several days. The application of intrauterine antibiotics is controversial. Systemic broad-spectrum antibiotics, such as trimethoprim–sulfamethoxazole (30 mg/kg per os (p.o.), q. 12 hours) and metronidazole (15 mg/kg, p.o., q. 12 hours), for several days is indicated and non-steroidal anti-inflammatory drugs can be given to prevent endotoxemia. The body temperature of the mare should be measured several times daily
Figure 300.8 Uterine prolapse in a Miniature mare.
after a retained placenta. A rise can be a symptom of a beginning metritis. The typical vaginitis and cervicitis in Miniatures after prolonged vaginal manipulation has been described above. Daily intravaginal application of 1% sodium carboxymethylcellulose solution with added heparin (100–330 IU/mL) and dexamethasone (1 mg/mL) in a syringe and with the use of an insemination pipette may prevent adhesions.
References 1. Ginther OJ. Reproductive Biology of the Mare. Cross Plains, WI: Equiservices, 1992; pp. 105–32. 2. Gastal EL, Neves AP, Mattos RC, Petrucci BPL, Gastal MO, Ginther OJ. Miniature ponies. 1. Follicular, luteal, and endometrial dynamics during the oestrous cycle. Reprod Fertil Dev 2008;20:376–85. 3. Ginther OJ, Beg MA, Neves AP, Mattos RC, Petrucci BPL, Gastal MO, Gastal EL. Miniature ponies. 2. Endocrinology of the oestrous cycle. Reprod Fertil Dev 2008;20: 386–90. ¨ 4. Neves AP, Trein CR, Moller G, Azevedo LR, Brito ELR, Malschitzky E, Mattos RC. Embryo transfer in miniature mares: preliminary results. In: Proceedings of the 6th International Symposium on Equine Embryo Transfer. Havemeyer Foundation, Monograph Series No. 14, 2004, pp. 101–2. 5. De Vries PJ, van der Holst W. De waarde van een progesteronbepaling in bloedplasma rond de 18e dag post ovulationem voor het drachtigheidsonderzoek bij de merrie. Tijdschr Diergeneeskd 1983;108:401–6. 6. Campbell ME. Selected aspects of miniature horse reproduction: serologic and ultrasonographic characteristics during pregnancy and clinical observations. In: Proceedings of the 37th Annual Meeting of the Society for Theriogenology, 1992, pp. 89–96. 7. Purdy SR. Ultrasound examination of the reproductive tract in female miniature donkeys. In: Matthews NS, Taylor TS (eds) Veterinary Care of Donkeys. Ithaca, NY: International Veterinary Information Service, 2005. 8. Foristall KM, Roser JF, Liu IKM, Lasley B, Munro CJ, Carneiro GF. Development of a tandem hormone assay for the detection of pregnancy in the miniature mare. In:
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Miniature Pony Reproduction Proceedings of the 44th Annual Convention of the American Association of Equine Practice, 1998, pp. 52–3. 9. Henderson K, Stevens S, Bailey C, Hall G, Stewart J, Wards R. Comparison of the merits of measuring equine chorionic gonadotrophin (eCG) and blood and faecal concentrations of oestrone sulphate for determining the pregnancy status of miniature horses. Reprod Fertil Dev 1998;10: 441–4. 10. Ginther OJ, Bergfelt DR, Leith GS, Scraba ST. Embryonic loss in mares: Incidence and ultrasonic morphology. Theriogenology 1985;24:73–86. 11. Turner RM, McDonnell SM, Feit EM, Grogan EH, Foglia R. Real-time ultrasound measure of the fetal eye (vitreous body) for prediction of parturition date in small ponies. Theriogenology 2006;66:331–7.
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12. Marien T, Adriaenssen F, Segers L. Caesarean section in the miniature Shetland horse. Surgical technique and outcome in ten cases. Vlaams Diergeneeskundig Tijdschrift 1998;67:340–3. 13. De Bois CHW. Sectio Caesarea bij het Paard. Tijdschr Diergeneeskd 1968;93:1207–19. 14. Judd JC. A practitioner’s approach to reproductive problems in miniature mares. Equine Pract 1994;16:9–14. 15. Ginther OJ, Williams D. On-the-farm incidence and nature of equine dystocias. J Equine Vet Sci 1996;16:159–64. 16. Burns SJ, Judge NG, Martin JE, et al. In: Proceedings of the 23rd Annual Convention of the American Association of Equine Practitioners, 1977, pp. 381–90.
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Chapter 301
Asian Wild Horse Reproduction: Equus Ferus Przewalskii C. Wynne Collins, Lee E. Boyd and Katherine A. Houpt
The Asian wild horse, also called Przewalski’s horse in the West, ‘takhi’ in Mongolia, and ‘yehmah’ in China, is the last wild horse in existence today. They are physically distinct from the domestic horse by their absence of a forelock, short mane, characteristic dun color, with black points, light muzzle with dark nostrils and lips, and frequently stripes on limbs and body (Figures 301.1 and 301.2). Although there were accounts of them from 20 000 bc, they were not ‘discovered’ by the Western world until 1878, when Colonel Nikolai Michailovich Przewalski was given a skull and skin of a horse shot in Xinjiang, China. At the Zoological Museum of the Russian Academy of Science, the conservator, I. S. Poliakov, concluded this was a new species, which he named Equus przewalskii Poliakov (Equus ferus przewalskii).
Last sightings The original distribution of the Asian wild horse was across the whole Eurasian steppe belt.1 During the first half of the last century, numbers of Asian wild horses rapidly decreased and their range contracted to the Gobi Desert region, due to competition with livestock and a variety of other factors. In 1969, the Mongolian scientist, N. Dovchin, was the last person to see an Asian wild horse in the wild. Further expeditions in 1979, 1988, and 1989 revealed no traces of horses, and, as a result, they were then declared extinct in the wild by the International Union for the Conservation of Nature (IUCN) (IUCN Red List).
Genetics Domestic horses (E. ferus caballus) have a diploid chromosome number of 64, whereas that of Asian wild horses (E. ferus przewalskii) is 66. Hybrids (chromosome number = 65) are different from many other equid crosses (e.g., mule, hinnie) because they are fertile. Asian wild horses also differ from the domestic horse in nuclear genetic markers and are distinct descendants from a most recent common ancestor. The identity of that ancestral species remains elusive. There is an absence of notable mitochondrial DNA (mtDNA) divergence between the two species, but the Y chromosomes diverge markedly, suggestive of an ancient separation of the species.2 Animals that were captured and founders of the current population bred with domestic horses before and after capture and carry some of their genes.3 Despite these influences, the captive population is still considered distinct because the genetic influence from domestic horses now has very little representation.
Breeding management of the captive population Figure 301.3 illustrates the contribution of the 14 founders to the current Asian wild horse population. Founder genes have been lost since the establishment of the captive population due to many genetic bottlenecks, for example: reduced breeding, tragedies such as World War II when many animals were lost, poor breeding management, and culling of animals that
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Przewalski’s stallions mature later than domestic stallions, showing delayed descent of testicles. Stallions will breed until their thirties; however, the prime age for breeding seems to be from 5 to 20 years of age.4 The oldest stallion to produce offspring was 36 years of age.5 Przewalski’s stallions have sired
offspring throughout the year, but they do show seasonal changes in sperm production and sexual behavior similar to the domestic stallion.6 No extensive endocrinology studies have been conducted on Przewalski’s stallions to determine whether they also exhibit seasonal changes in reproductive hormones similar to the domestic horse. Bader et al.7 evaluated semen from five stallions ranging in age from 4 to 20 years. Semen was collected by electroejaculation (n = 10) or artificial vagina (n = 30). Three of the stallions had a history of subfertility and showed low sperm counts (0–2.0 × 106 ), decreased sperm motility (0–33%), and a high proportion of morphological abnormalities (53–61%). Testicular tissue was examined by histology, and atrophy of germinal epithelium was apparent. A second study8 examined 10 stallions over a 1-year period (age range 2.5–22 years). Semen was collected by electroejaculation under general anesthesia. Total volume, motility, concentration, and morphological abnormalities were examined in winter, spring, summer, and fall. Morphological abnormalities were lowest in the winter (24%) compared to spring (60.8%), summer (50.7%), and fall (69.2%). Motility was lowest in summer (15.5%) compared to fall (56%), winter (34.2%), and spring (45.8%). This is contradictory to what would be anticipated, considering peak breeding for Przewalski’s horses is spring and summer.9 Young stallions (< 10 years) were also compared with older stallions (> 10 years) and there was no significant difference between them. A recent study10 was conducted on nine Przewalski’s stallions from three separate institutions. All males were collected under anesthesia and electroejaculation was performed. Seminal traits were characterized from all stallions and showed some variation
(a)
(b)
Figure 301.1 Typical Asian wild horse. Manlai (#2619 Hustain Nuruu 1). The first foal born in the wild. He is exhibiting flehmen. Note the short mane and lack of forelock, which are characteristic of Equus ferus przewalskii. Photo by Lee Boyd at Hustai National Park.
were not considered ‘pure’. The Asian wild horse is unique in that a record (stud book) of all animal pairings has been maintained since 1900, so the genetics of all animals is known. In order to maintain founder genes, breeding pairs are selected based on the coancestry of potential mates.
Reproductive physiology of captive Asian wild stallions
Figure 301.2 Stallion injuries incurred during competition with other stallions at Hustai National Park. (a) Bite marks incurred ¨ 25) was replaced by another. (b) Left ear of stallion Margad (#2701 Hustain Nuruu 8) bitten in a when the stallion (Ares #1891 Koln stallion fight. Also note the zebra stripes on the limbs in (a) and the lack of forelock and the dorsal stripe in (b). These are anatomical characteristics of Equus ferus przewalskii. Photos by Lee Boyd at Hustai National Park.
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0.3 Representation
0.2
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Founder ID Figure 301.3 Genetic representation of the 14 founders of the current Asian wild horse population. Reprinted with permission from Montfort S. Powell D. and Spivak E. et al., 2007. Population Analysis and Breeding Plan Przewalski’s Horse (Equus caballus przewalskii) Species Survival Plan, p. 6.
between animals. Average total concentration was 2.0 ± 0.6 × 109 , total motility was 52.7 ± 3.4%, and progressive motility was 44.0 ± 2.7. The frequency of morphological abnormalities was high among all stallions (66.6%), with head shape abnormalities predominating (28.7 ± 1.8%). A study is presently being conducted at the National Zoological Park on 10 stallions (ages 4–29 years) over a 2-year period and should give a good overall description of seminal traits in Przewalski’s stallions (C.W. Collins, personal communication). Vasectomy reversal in the Asian wild horse Breeding management of the Asian wild horse by the subspecies (ssp.) means that zoological institutions have to adhere strictly to breeding recommendations each year in order to avoid producing inbred offspring. In 1999, because of limited holding space, one zoological institution performed a vasectomy on a breeding stallion so that he could be housed with his daughters. Because this male was valuable genetically in the ssp. population and it is possible to reverse a vasectomy, performing a vasectomy would conserve his genetics for the future. By 2007, this male was moved to the National Zoological Park. Dr Sherman Silber, a human fertility expert, was able to reverse the vasectomy on one side. In April 2008, semen was successfully obtained under anesthesia. This stallion will be placed with breeding females in summer 2008. The ability to reverse this procedure in equids will provide zoological institutions another management option that can be used to control breeding in potentially valuable males.
Reproductive physiology of captive Asian wild mares Przewalski’s mares begin cycling by 2 years of age, but generally do not breed until 4 years.11 Peak reproduction in mares occurs from 5 to 7 years of age,4 although they generally remain fertile until 20 years of age.11 The oldest female known to have foaled was 24 years of age.5 Przewalski’s mares are seasonally polyestrous and exhibit estrus in early spring and anestrus in late fall and winter,12 similar to domestic mares. In order to monitor hormone patterns, scientists have used feces, blood, milk, and urine. Because mares must be sedated to obtain blood samples, a better approach is to monitor urinary hormone metabolites. Fresh urine samples can be collected from the ground and analyzed. Using this technique, Monfort et al.13 and Boyd et al.12 were able to construct longitudinal urinary conjugate profiles from cycling and pregnant Przewalski’s mares maintained on pasture with a stallion. The mean estrous cycle length was determined to be around 24 days, slightly longer than domestic mares. Based on variations in urinary estrogen conjugates, the onset of estrous cycles coincides with increasing day length (between February and April). All mare’s conceived between April and November, and gestation was 48.6 weeks (range: 47–51 weeks).13 Similar to domestic mares, urinary estrogen conjugates rose by the sixth week of gestation, with a rapid decline in late pregnancy. In the domestic mare, ultrasonographic examination of reproductive organs is invaluable in understanding follicular dynamics of the estrous
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Asian Wild Horse Reproduction: Equus Ferus Przewalskii cycle, and this could prove to be a very useful tool in the Przewalski’s horse. Ultrasonographic examination of the ovaries of two anesthetized Przewalski’s mares by Durrant et al.14 was performed to assess follicular growth patterns. Each mare was anaesthetized seven times over a 3-month period, and ultrasound was used to examine the reproductive tract. According to the data, follicle size increased by 3–5 mm/day, and ovulation occurred when follicles reached a diameter of 48–50 mm. Repeated immobilizations carry a risk to the horses, as well as requiring a considerable investment in time, personnel, and drugs. Another approach to assessing reproductive function in Asian wild horses has been undertaken at the National Zoo’s Conservation and Research Center, where mares have been habituated to entering a chute weekly (C.W. Collins, personal communication). Mares are examined by rectal palpation and ultrasound 3–5 days per week during the breeding season, and urine samples are collected so that urinary hormone metabolites can be compared to follicular structures.
Assisted reproduction Most reproductive biotechnologies such as artificial insemination, sperm cryopreservation, and in vitro fertilization could be useful tools in breeding management of the captive population. Non-surgical embryo collection was performed on two Asian wild horse mares over two breeding seasons. A total of 11 embryos were transferred into Welsh-type pony mares and pregnancies were monitored throughout gestation. Four foals were born. Summers et al.15 demonstrated that interspecies embryo transfer could be used to enhance reproductive efficiency in wild equids such as the Przewalski’s horse. Embryo cryopreservation could also be a valuable technique for the preservation of genetically valuable individuals. Sperm cryopreservation is another technique that could aid in the genetic preservation that is crucial in a population with a small founder base. Durrant8 assessed frozen-thawed sperm from two stallions with differing fertility and noted that there was no difference in capacitation rates and binding to hamster zona pellucida. In recent years, cryopreservation of equine sperm has improved due to the development of new cryodiluents and freezing methods.16 These improved techniques could easily be applied to Przewalski’s stallions; however, a firm understanding of seminal traits is critical prior to success. With the development of semen cryopreservation, artificial insemination becomes an obvious solution for breeding management of the Przewalski’s horse. Movement of stallions or mares for breeding pur-
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poses is risky and expensive and may result in breeding incompatibility. Artificial insemination not only overcomes these issues, but also could aid in preservation of founder genes because stored semen from stallions that are no longer in the population can be used to produce offspring.
Reintroduction sites in Asia Although extinct in the wild, an Asian wild horse population existed in zoos all over the world. This made re-establishment of animals in the wild possible. There are six sites in which Asian wild horses have been reintroduced: Hustai National Park (Mongolia), Takhin Tal (Mongolia), Khomin Tal (Mongolia), Kalameili (China), Altyn-Emel Park (Kazakhstan), and Bukara Breeding Center (Uzbekistan). There are now about 500 animals in the six established sites in central Asia (see Table 301.1 for foaling rates).
Reproduction in the wild Asian wild horse stallions form harems of up to 10 mares, similar to feral domestic horses. Surplus males form bachelor groups that compete with harem stallions for mares.17 Fighting can be intense, leading to permanent scarring of combatants (Figure 301.2). Reintroduced mares are typically at least 3 years old when they first conceive; infrequently, a 3-yearold gives birth (Figure 301.4). A pattern of lower rates in the first years after reintroduction and 60–80% foaling rates thereafter has occurred in Takhin Tal and Hustai National Park.18–20 Infanticide Foal killing by stallions has been reported in captive Asian horses,21, 22 as well as in domestic horses.23 The major cause of neonatal foal death in the reintroduced Chinese population was infanticide by the harem stallion. In the captive Chinese population, 32% of the foal deaths were by infanticide.24 Infanticide has also occurred at Takhin Tal and Hustai. In Takhin Tal, Table 301.1 Foal mortality at reintroduction sites
Total number of foals born Survived 1st year Causes of mortality: Dystocia and stillbirth Wolves Weather From Enkhtur.20
Hustai
Takhin Tal
259 56%
97 55%
15% 27% –
8% 8% 15%
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Figure 301.4 An Asian mare and foal at Hustai National Park. Photo by Lee Boyd.
the stallion (Tuulai) killed a foal of the mare Erdene he had not sired.20 At Hustai, the captive-born stallion Khaan killed a male foal sired by another stallion. The dam subsequently conceived three foals by Khaan, two of them male, and he attacked none of them.25 Additionally, two of the wild-born stallions at Hustai have killed foals sired by other stallions. Killing of adults and juveniles When originally moved to the Bukara Breeding Center, two stallions (#1521 and #1517) got along, but when the mares began to come into estrus, they fought and were separated. Eventually, a fence was broken down and #1521 killed #1517. Later, #1521 killed #1517’s yearling son. Not all stallion contests have fatal consequences. Figure 301.2 illustrates wounds inflicted on stallions by competing stallions.
References 1. Bouman J, Bouman I. The history of Przewalski’s horse. In: Boyd LE, Houpt KA (eds) Przewalski’s Horse. Albany, NY: SUNY Press, 1994; pp. 5–38. 2. Ryder OA. Genetic studies of Przewalski’s horses and their impact on conservation. In: Boyd L, Houpt KA (eds) Przewalski’s Horse: The History and Biology of an Endangered Species. Albany: State University of New York Press, 1994; pp. 74–92. 3. Bowling AT, Zimmerman W, Ryder O, Penado C, Peto S, Chemnick L, Yasinetskaya N, Zharkikh T. Genetic variation in Przewalski’s horses, with special focus on the last wild caught mare 231 Orlitza. Cytogenet Genome Res 2003;101:226–34. 4. Ballou JD. Population biology. In: Boyd L, Houpt KA (eds) Przewalski’s Horse: The History and Biology of an Endangered Species. Albany: State University of New York Press, 1994; pp. 93–113.
5. Veselovsky Z, Volf J. Breeding and care of rare Asian equids at Prague Zoo. International Zoo Yearbook 1965;5:28–37. 6. Volf J. Die “wilde” oder gezielte Aufzucht von Przewalskipferden (Equus przewalskii Polj., 1881)? Zoologische G¨arten 1989;59:402–10. 7. Bader H, Gremmes S, Sieme H, Paar M, Hoppen HO, Brandt KP. Subfertility in the male Przewalski horse. J Reprod Fertil Suppl 1991;44:676–7. 8. Durrant BS. Semen characteristics of the Przewalski’s stallion (Equus przewalskii). Theriogenology 1990;33:221. 9. Monfort SL, Arthur NP, Wildt DE. Reproduction in the Przewalski’s horse. In: Boyd L, Houpt KA (eds) Przewalski’s Horse: The History and Biology of an Endangered Species. Albany: State University of New York Press, 1994; pp. 173–93. 10. Collins CW, Songsasen N, Monfort SL, Bush M, Wolfe B, James SB, Wildt DE, Pukazhenthi BS. Seminal traits in the Przewalski’s horse (Equus ferus przewalskii) following electroejaculation. Anim Reprod Sci 2006;94:46–9 11. Mohr E. The Asiatic Wild Horse. London: J.A. Allen & Co. Ltd, 1971. 12. Boyd LE, Kasman LH, Lasley BL. Urinary estrone sulfate concentrations in Przewalski’s horses and behavioral correlates. Vestnik Zoologii Suppl 1999;11:47–53. 13. Monfort SL, Arthur NP, Wildt DE. Monitoring ovarian function and pregnancy by evaluating excretion of urinary oestrogen conjugates in semi-free-ranging Przewalski’s horses (Equus przewalskii). J Reprod Fertil 1991;91:155–64. 14. Durrant BS, Oosterhuis JE, Hoge ML. The application of artificial reproduction techniques to the propagation of selected endangered species. Theriogenology 1986;25:25–32. 15. Summers PM, Shephard AM, Hodges JK, Kydd J, Boyle MS, Allen WR. Successful transfer of the embryos of Przewalski’s horses (Equus przewalskii) and Grant’s zebra (E. burchelli) to domestic mares (E. caballus). J Reprod Fertil Suppl 1987;80:13–20. 16. Loomis PR, Graham JK. Commercial semen freezing: Individual male variation in cryosurvival and the response of stallion sperm to customized freezing protocols. Anim Reprod Sci 2008;105:119–128. 17. Boyd L, Houpt KA (eds). Przewalski’s Horse: the History and Biology of an Endangered Species. Albany, NY: SUNY Press, 1994. 18. King S, Lanting F. Back to the Hustai. BBC Wildlife 2002, pp. 32–40. 19. Kaczensky P, Walzer C. Przewalski horses, wolves and khulans in Mongolia. Research Report, International Takhi Group, 2003; 16 pp. 20. Enkhtur A. The Present situation of reintroducing takhi in Hustain Nuruu and in Takhin Tal. Research Report, Hustai National Park, 2004; 20 pp. 21. Boyd L. Behavior problems of equids in zoos. Vet Clin N Am-Equine 1986;2(3):653–64. 22. Feh C, Munkhtuya B. Male infanticide and paternity analyses in a socially natural herd of Przewalski’s horses: Sexual selection? Behav Process 2008;78:335–8. 23. Duncan, P. 1982. Foal killing by stallions. Appl. Anim. Ethol. 8:567–570. 24. Chen J, Weng Q, Chao J, Hu D, Taya K. Reproduction and development of the released Przewalski’s horses (∗ Equus przewalskii in Xinjiang, China). J Equine Sci 2008;19:1–7. 25. Enkhtur A. Some biological and adaptation issues of takhi being reintroduced. Research Report, Hustai National Park, 2000.
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Chapter 302
Zebra Reproduction ˜ Cassandra M.V. Nunez, Cheryl S. Asa and Daniel I. Rubenstein
Introduction Few studies have investigated the reproductive biology of wild equids; much of what we know about zebra species has come from dissections following culling.1–3 Zoo research has proven useful in determining additional details regarding the ovulatory cycling of females,4 and the timing of puberty and breeding life in both males and females. Behavioral studies both in the field and in captivity have helped to clarify these factors.5–8 In addition, behavioral studies have revealed how breeding behavior, the degree of breeding seasonality, stages of parturition, mother–infant behavior, and the dispersal of young males and females can vary in the different species.9, 10 Using this combination of behavioral, physiological, and demographic data, we explain the reproductive biology of each age and sex class of plains zebra (Equus burchelli), the two subspecies of mountain zebra – Cape mountain zebra (Equus zebra zebra) and Hartmann’s zebra (Equus zebra hartmannae) – and Grevy’s zebra (Equus grevyi), and relate its relevance to their ecology and natural history.
Social context: mating patterns and their ecological determinants Zebras live in two broadly different types of societies.11–13 In one, typified by plains and mountain zebras, females and males live in unimale-multifemale family groups with their infants, yearlings, and twoyear-olds. These so-called ‘harem’ groups are relatively stable with males and females living together for many years.14 In the other, typified by Grevy’s zebras, female associations are short-lived and females often roam in groups without males. Fusing and fissioning of individuals creates these open membership groups that wander through territories of isolated males. From the perspective of males, both systems are polygynous since reproductively active
and dominant males often mate with more than one female. In plains zebras, polygyny involves harem defense since females form strong associations, while in Grevy’s zebras it involves resource defense as males defend grazing areas that females seek. As in any polygynous system, younger, less dominant males form bachelor groups and attempt to cuckold stallions while strengthening ties to females. From the perspective of females, however, these mating systems offer different reproductive opportunities. In plains zebras, females are monoandrous since all females in a harem bond to a particular male. In Grevy’s zebras, females are polyandrous since they often move across territorial boundaries and mate with a variety of males in succession. Each mating system emerges as a response to particular ecological circumstances.12 In general, zebra males provide important material rewards to females that increase their reproductive success. By increasing vigilance and enabling females to move unhindered when seeking resources, some males enable females to spend more time grazing.12 Females associating with the most dominant males, or those rising rapidly in rank, rear the most offspring to the age of independence.13 But this only applies in mesic habitats where food and water – the two key resources females need to sustain themselves and their young – are located close enough to each other so that females of different reproductive states can stay together and simultaneously derive these time-related benefits that males provide. For Grevy’s zebras inhabiting more arid areas where rainfall is highly unpredictable, food and water are typically far apart, thus females having different needs cannot stay together and the social fabric becomes torn. As a result, female associations are weak. Since females exhibiting either postpartum or cycling estrus are equally valuable to males, those of high rank defend territories along routes to water so that they can mate with lactating females residing near water and
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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non-lactating females that must come to drink every 3–5 days.15, 16 Males of lesser rank defend territories in grazing areas and thus gain access to a more limited pool of reproductively active females. It is within these social and ecological contexts that the reproductive biology of zebras must be examined. Only by appreciating how socioecological selective pressures shape social relationships and mating behavior can the similarities and differences in reproduction exhibited by the various age and sex classes of each zebra species be understood.
males and females leaving their natal groups by 3 years of age. Reproducing females and males interact regularly. Females generally initiate movements, with males directing the group from the side or the rear. Lactating females often initiate moves to water and lead the way.18 Mating often involves conflict among the sexes. Females typically adopt a rigid position with tail raised when initiating copulation. More often, however, the male identifies females in heat by sniffing or licking the genital region. Resting his head on a female’s rump or biting at the withers often precedes mounting by the male. If a female is not ready to mate she either walks out from under the male or uses her hind legs to drive him away.
Mare Behavior Female-female bonds in all zebra species are weak. There is little competition over food and cooperative interactions in terms of allogrooming or mutual defense of young are rare. Similarly, there is minimal competition for mates, but female mountain zebras compete for dominance, and rank affects reproductive fitness. Although both dominant and subordinate females produce approximately the same number of offspring over a lifetime, the survival of young born to high-ranking females exceeds that of young born to low-ranking females.17 In addition, as daughters take on the rank of their mothers, dominant females will produce disproportionately more female offspring. In all species, bonds between mothers and their young weaken as offspring mature, with both
Seasonality
Plains zebra In general, zebra are less seasonal than other African species. Plains zebra foals are born throughout the year, although peaks do exist during the long rains (Figure 302.1). Klingel6 reported that 85.5% of foals were born from October to March in the Ngorongoro Crater, Tanzania. Similarly, Smuts2 found that 82.6% of foals were born between September and April in Kruger National Park, South Africa. Since the gestation period is about 12 months, mating activity must occur throughout the year with a peak during the summer months. Plains zebra in North American and
30 Plains Cape mountain Hartmann’s Grevy’s
25
Percent of total annual births
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15
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Aug
Sep
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Dec
Jan
Feb
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Apr
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Month Figure 302.1 Seasonal distribution of births for the different zebra species. Data compiled and adapted from Klingel26 (plains zebra), Joubert21 (Cape mountain zebra), Westlin-van Aarde et al.22 (Hartmann’s zebra), and Dobroruka et al.5 (Grevy’s zebra). Grevy’s data are from captive animals.
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Figure 302.2 Seasonal distribution of births for plains zebra and Grevy’s zebra held in zoos; data are taken from the Plains Zebra Regional Studbook for North American zoos19 and the Grevy’s Zebra International Studbook.25
European zoos19 have given birth in every month of the year, but births cluster in spring and summer (Figure 302.2). However, trends for spring and summer births in zoos are sometimes the result of management decisions for young to be born in those months.
Mountain zebra Free-ranging Cape mountain zebra mares in the Mountain Zebra National Park and De Hoop Nature Reserve, South Africa,20 foal year round with peaks around the rains. Over 68% of Cape mountain foals are born during the rains, from November through April.7 Free-ranging Hartmann’s zebra in southwest Africa are similar, with 81–86% of foaling occurring from November through April.21, 22 During a 3-year study of Hartmann’s zebra, mating activity was only seen from September to April, with a peak in February.21 These data, coupled with the fact that the gestation period is about 1 year, indicate that mating during the rest of the year is quite rare.
versely, mares can be induced into anestrus when resources are scarce.15 Gestation is a bit longer in Grevy’s zebra (about 390 days), so peaks of mating activity must occur in June through July and September through October. Captive animals in Dvur Kralove Zoo in the former Czechoslovakia gave birth every month from 1973 to 1985, although peaks did exist in January and from June through August,5 generally following the foaling schedule of wild populations (see Figure 302.1). Data from the Grevy’s Zebra International Studbook25 for the period 1984 through 2004 show a similar trend, with births throughout the year but concentrated more in spring and summer months. As with plains zebra, this trend may reflect management strategies, but year-round endocrine monitoring of the herd of Grevy’s zebra at the Saint Louis Zoo (J.E. Bauman, C.S. Asa, and M. Fischer, unpublished) has revealed a pattern of lower concentrations of progesterone following ovulations in winter, suggesting that fertility might indeed be lower during that period. Puberty and breeding life
Grevy’s zebra
Plains zebra
Grevy’s zebra are perhaps the most seasonal of the zebras with peaks of mating occurring during the long rains in July and August, and the short rains in October and November.23 In the free-ranging populations of northern Kenya, breeding is highly dependent upon available resources, with peaks in estrus coinciding with higher resource availability.24 Con-
Early work on plains zebra defined puberty in mares as the onset of first estrus and estrous behaviors.1, 2, 6 King reported first estrus to occur at an average age of 2 years 4 months, ranging from 1 year 8 months to 2 years 9 months. Klingel26 found that Ngorongoro mares exhibited first estrus earlier, at 13–15 months of age. Similarly, Smuts2 found that the first estrus of
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Table 302.1 Reproductive data for plains and Grevy’s zebras held in zoos Plains zebraa
Grevy’s zebrab
Females Number of females Youngest to give birthc Oldest to give birth Most offspring
334 1 year 8 months 26 years 1 month 15 foals
169 2 years 6 months 28 years 4 months 13 foals
Males Number of males Youngest to sire offspring Oldest to sire offspringd
149 7 months 26 years 9 months
85 1 year 11 months 25 years 9 months
a
Data taken from the Plains Zebra North American Regional Studbook (2004)19 for records from 1980 through 2003. b Data taken from the Grevy’s Zebra International Studbook (2004)25 for records from 1984 through 2004. c Age at parturition. d Age when female conceived.
mares in Kruger National Park occurs at 1.5 to 2 years of age. The Plains Zebra North American Regional Studbook (Table 302.1)19 has parturition records for females at 1 year and 8 months, 1 year and 10 months, and 1 year and 11 months of age; the data represent the female’s age at the time she gave birth, not the time she conceived. Klingel6 estimated the reproductive potential of mares as one foal per year from the age of 3.5 to about 15 to 18 years. They observed 120 mares for three seasons. Fifteen percent of the animals had three foals (one per year), 33% had two, 42% had one, and 10% had no offspring. On average, mares had a surviving foal once every 2 years. This coincides with data from a captive animal at Chester Zoo, England, that at 17 years old, was still breeding after having produced six foals since attaining sexual maturity.6 North American Regional Studbook (Table 302.1)19 records show that 10 females over 20 years of age produced foals, the oldest at 26 years. Twelve females gave birth to more than 10 foals over a lifetime. Of 1232 recorded births, 11 (1%) were twins.
Mountain zebra In Mountain Zebra National Park, young Cape mountain zebra mares leave their natal herds at a mean age of 1 year 10 months; often they are in first estrus at this time.7 Lloyd and Rasa17 reported Cape mountain fillies in De Hoop Nature Reserve leaving a bit earlier, at a mean age of 1 year 7 months (again during first estrus). First foaling has been recorded at ages as young as 2 years17 to as old as 5 years 7 months.7 Regardless of the exact age at first foaling, it seems likely that the mares do not become sexually mature until several months after first estrus. The ac-
tual age of first estrus in Hartmann’s zebra has not been recorded, but free-ranging mares have their first foal at about 2 years of age. Young mares are expelled from the natal group at 14 to 16 months of age,27 at which time they may be experiencing first estrus. The reproductive potential of Cape mountain zebra mares has been estimated at about one foal every 25 months, with a range of 13–69 months.7, 20 Similarly, Lloyd and Rasa17 calculated an average interbirth interval of 27.4 months. Given a reproductive lifespan of about 15 years, a Cape mountain zebra mare can be expected to produce eight to nine foals in her lifetime. The reproductive potential of Hartmann’s zebra has not been reported explicitly, but may be surmised by maternal behavior. Hartmann’s zebra mares wean their foals at approximately 10 months of age, and begin to actively expel the foals at 13 to 16 months of age, shortly before the birth of their next foal. Given this behavior, we can expect the reproductive potential of Hartmann’s zebra mares to be about one foal every 1 to 1.5 years.
Grevy’s zebra Puberty in free-ranging Grevy’s zebra mares has been estimated to occur at 3 to 4 years of age1 with mating occurring during the second estrus and conception occurring after two or three estrous cycles. However, captive animals have been reported to reach puberty and achieve conception at much younger ages. The International Studbook has records of one female at 2 years 6 months and another at 2 years 10 months giving birth for the first time (Table 302.1).25 In a study at the Saint Louis Zoo that monitored behavior and fecal hormones,4 a 2-year-old female conceived but aborted an 18-cm fetus at about day 100 of gestation. She subsequently mated and aborted at least one more time before successfully carrying a foal to term, which was born when she was 3 years 4 months of age. Managers had suspected that young females were more likely to experience abortions, and the results of this study supported that hypothesis. This suggests that females may be able to ovulate and conceive before their reproductive tracts are sufficiently mature to maintain pregnancy. Age at puberty is affected by nutrition, but may also be influenced by social constraints in the wild that may not be present in captivity.4 Free-ranging Grevy’s zebra mares may breed from 1 to a few months after foaling, with an average reproductive potential of one foal every 2 years.1, 23 In captive animals, the mean interbirth interval has been estimated at 18 months (range 14.2–34.8 months5 ) and 19.75 months (range 13–27 months8 ) in Czechoslovakian and North American zoos, respectively. However, management strategies likely bias
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Zebra Reproduction the average intervals. According to the International Studbook (Table 302.1),25 the oldest female to give birth was 28 years of age, with 10 females over the age of 20 giving birth. Over a lifetime, there were seven females that each produced more than 10 foals, one of which gave birth to 13. The female reproductive organs
Plains zebra The female reproductive organs of culled animals were studied in detail by King.1 The plains zebra mare has two ovaries, each about 5 cm in length. The ovaries connect to the fallopian tubes (oviducts), which conduct the ovum from the ovary to the uterus. Each oviduct attaches to one of the two horns of the uterus, which are approximately 15 cm in length. The horns attach to the body of the uterus (10–15 cm long). Caudal to the uterus is the cervix, about 5 cm long, which enters the vagina. The vagina is about 15–20 cm long, and is quite elastic, allowing it to expand. The external opening of the vagina lies ventral to the rectum. The mare has two mammary glands, which are smaller in virgin mares. They have two ducts each, which open externally.
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a stallion. A mare in the display posture stands with her legs slightly apart and her tail lifted. The posture is quite conspicuous, even from a distance, and seems to be an important attractant for the stallion.6 Increased interest of the stallion, the open-mouthed facial expression of the mare (often called ‘jawing’), and mating are other diagnostic features of estrus.1 During metestrus reproductive activity subsides as the diestrous phase begins. Diestrus is the period of luteal activity between ovulatory cycles, and anestrus is the period of sexual rest during the non-breeding season. For the most part, a mare in anestrus does not ovulate, and the uterus is small and anemic.29 This is the normal state of the filly before puberty. In zebra mares, ovulation occurs at the ovarian fossa, a depression at the free, ventral edge of the ovary.2 This structure, present in all equid species, is lined with germinal cells destined to develop into ova.30 Ovulation occurs only at this structure and never from any other region on the ovary.31 In plains zebras (and probably mountain zebras), the ovarian fossa is not present at puberty, but develops after a few estrous cycles. In Grevy’s zebra, the ovarian fossa is fully developed at puberty, but in other respects resembles that of the other zebra species.
Mountain zebra
Ovarian activity during the estrous cycle
Detailed dissections of the female reproductive organs have not been reported in mountain zebra. Given their average mass of 234 kg and 276 kg for Cape mountain and Hartmann’s zebra mares, respectively,28 they are likely to have organs intermediate in size and shape to those of plains and Grevy’s zebra. For example, Westlin van-Aarde et al.22 reported that the ovaries of Hartmann’s zebra females are heavier than those of plains zebra2 at every stage of the reproductive cycle.
Plains zebra
Grevy’s zebra The reproductive organs of the Grevy’s zebra mare are similar to, but are about 10% larger than those of plains zebra.1 Estrus and ovulation Smuts divided the estrous cycle in all zebra species into five phases: proestrus, estrus, metestrus, diestrus, and anestrus.2 Proestrus is the phase immediately preceding estrus, and is characterized by an increase in ovarian activity. Mares during this phase will generally refuse a stallion’s attempts to mount. Estrus is the period of acceptance of the male, when a mare will exhibit the display posture and ‘stand’ for
Ovarian activity during the estrous cycle was outlined by Smuts2 using data collected from 310 free-ranging mares shot during a game cropping campaign in the Kruger National Park, South Africa. During proestrus, there is an increase in the follicular activity, with one follicle generally larger than the others. As in horses, regressing corpora lutea from previous cycles may be present. The estrous phase is similar to proestrus except that the follicle that is going to rupture is clearly identified and the mare now accepts the stallion. In metestrus, the Graafian follicle has ruptured, a corpus hemorrhagicum forms, and the follicular cavity fills with a lymph-like fluid. As the cells lining the ruptured follicle undergo rapid division and become luteinized, a corpus luteum is formed. The corpus luteum attains its maximum size during this period, after which it gradually regresses. Follicular activity is largely dormant. Should the mare fail to conceive, she enters diestrus or anestrus. During diestrus, a new wave of follicles starts developing and the corpus luteum starts to regress. Diestrous is generally followed by the proestrus phase. The corpus luteum is still present by the time the succeeding cycle commences, and in this respect, the plains zebra mare is similar to the domestic horse.32 Mares that have failed to conceive by the
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end of the breeding season may cease to cycle and go into anestrus. The ovaries of anestrous mares are largely inactive, containing some small follicles and some tiny, regressed corpora lutea.
larly, Smuts reported a 6-day estrous phase for mares in Kruger National Park, South Africa. Asa et al.4 measured estradiol and progesterone in fecal samples from Grevy’s zebra mares during consecutive estrous cycles to compare with courtship and mating behavior (Figure 302.3).
Mountain zebra A detailed study of female reproduction in the Hartmann’s zebra was conducted by Westlin-van Aarde et al.22 Ovaries and plasma were collected from freeranging zebra mares shot in the Etosha National Park, Namibia. Both proestrous and estrous mares have an average of 16.8 smaller follicles (1–10 mm) and 1 larger follicle (either 21–30 or 31–40 mm) present in each ovary. Corpora lutea are not present. In diestrus, corpora lutea are formed and several small follicles (averaging 19 follicles per ovary) are still present. Follicular activity is similar in anestrous mares, with an average of 20 small follicles and 0 large follicles per ovary, but corpora lutea are not present. Intermediate-sized follicles (at 11–20 mm) are present at every stage.
Gestation
Plains zebra Gestation length in captive plains zebra at the Basle Zoo, Switzerland, was reported by Wackernagel33 as 361–390 days, with an average of 371 days, in 28 pregnancies. The Philadelphia Zoological Garden, USA,34 reported gestation lengths of 341 and 355 days. These data are in accordance with those obtained from free-ranging animals by Klingel,6 at 378 and 385 days, and Smuts,2 at 396 days. At 6–7 months prior to birth, a mare’s abdomen is enlarged (when viewed posteriorly) and pregnancy is evident.2, 35 Some pregnant horse mares have been reported to show estrous behavior,36 which may also occur in zebras, so calculations of gestation length based on estrous behavior may not be reliable.
Grevy’s zebra Captive Grevy’s zebra mares at the Basle Zoo, Switzerland, were receptive for 2–9 days, in a 19–33day cycle (ref. 33 in ref. 1). In the study by Asa et al.,4 females were receptive for 3 to 6 consecutive days, with an estrous cycle duration of 28 to 35 days. Simi-
Mountain zebra Gestation length in mountain zebra has been estimated at about 1 year. In the wild, a Cape mountain female was estimated to give birth after 359 days,7 and a Hartmann’s female was estimated to give birth
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Zebra Reproduction after 362 days.21 A captive Cape mountain zebra mare gave birth after 375 days.37 These data are few, but given the gestation lengths of plains and Grevy’s zebra, the figures are not unlikely.
Grevy’s zebra In the wild, gestation is longer in Grevy’s zebra, averaging 390 days.1, 38 Gestation lengths in captive animals have been shown to range from 358 to 438 days by Read et al.8 (St Louis, USA) and from 390 to 406 days by both Antonius39 (Germany) and Kenneth40 (England). Distension of the abdomen tends to be more apparent in Grevy’s than in other zebra species.1 Gestation length ranged from 391 to 406 days for mature captive mares but was 425 days for a young primiparous mare.4 Ovarian activity during gestation
Plains zebra King1 investigated ovarian activity during gestation in 20 free-ranging pregnant animals, postmortem. He found that until approximately 22 weeks of pregnancy, a corpus luteum of about 2.5 cm in diameter was present in one of the ovaries. After 22 weeks, it began to regress; at full term all that remained was a 0.3 cm wide strip extending approximately 0.9 cm from the ovarian fossa. Follicles of about 2 cm in diameter were present up to 17 weeks of pregnancy, decreased to 0.5 cm at 48 weeks, and showed renewed activity at full term, measuring 1 cm. Smuts2 found similar results in the Kruger National Park zebras. During the first 190 days of pregnancy, he found a number of interesting phases of ovarian activity. After conception, ovarian activity gradually increased, reaching a peak between days 40 and 110. For the latter half of gestation, follicles larger than 12 mm were largely absent from the ovaries, although smaller, regressing follicles were often quite abundant. Active corpora lutea were a red-brown color, changing to yellow as they regressed. According to Smuts,2 activity was not equal in the two ovaries. The left ovary appeared to be more active than the right. In cases where mares produced all corpora lutea in only one ovary, a total of 109 were counted in the left and only 73 in the right (60% left and 40% right). In addition, the left ovaries of plains zebra mares were larger. Of 340 ovaries (170 pairs) measured, the left had a greater mass than the right in 120 instances (70.6% left > right). This was in contrast to King’s results.1 He found no difference in activity between the two ovaries, with active corpora lutea occurring in the right ovary in seven animals, and in the left ovary in eight animals.
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Mountain zebra During gestation, ovarian activity in the mountain zebra has been reported to be much like that of other zebra species. Joubert21 conducted a detailed study on free-ranging Hartmann’s zebra females in which he found that large corpora lutea remained in the ovaries even at advanced stages of pregnancy (up to 5 months). The average size of the corpus luteum before reduction was 2.5 cm. The color changes of the corpus luteum were the same as in other zebra species, with active luteal bodies having a red-brown color before changing to yellow with advanced regression. Ovarian activity was reported to be equal in the two ovaries, with follicles maturing in either the left or the right, or more often in both ovaries.
Grevy’s zebra According to King,1 gestational ovarian activity in free-ranging Grevy’s mares is similar to that of plains zebra and mountain zebra.21 Until about 24 weeks of pregnancy, a corpus luteum of about 4 cm in diameter was present in one of the ovaries. By 34 weeks, the corpus luteum had regressed to about 1.7 cm, but persisted at 1.1 cm up to full term. This persistence of a fairly large corpus luteum during pregnancy was not associated with endocrine activity; total progesterone content of the corpus luteum decreased from 4750 µg at 8 weeks to 500 µg at 24 weeks. Follicular activity was considerable during early pregnancy (3 cm), but had ceased by 34 weeks. Follicles at 1 cm in diameter reappeared at 42 weeks and reached 3 cm by full term. As with plains and mountain zebra, active follicles were a dark red-brown color and changed to yellow as they regressed. For Grevy’s females, ovarian activity appeared to be equal in the two ovaries, with corpora lutea forming in both the left (three animals) and right (three animals) ovaries. Asa et al.4 measured levels of estradiol and progesterone excreted in feces throughout pregnancy (Figure 302.4) and found patterns similar to those reported for horses (e.g., Squires et al.41 ). They also measured equine chorionic gonadotropin (eCG) in urine, which also revealed a pattern similar to that seen in horses.42 Levels were in the detectable range beginning between 35 and 40 days gestation and returned to baseline between days 195 and 200. Ramsay et al.43 also reported eCG elevations during that period of gestation. Parturition
Plains zebra Zebra mares give birth in much the same way as domestic horses. Wackernagel33 reported information
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Figure 302.4 Patterns of fecal progesterone and estradiol and urinary equine chorionic gonadotropin concentrations during a conceptive estrous cycle followed by pregnancy in a Grevy’s zebra.4 eCG, equine chorionic gonadotropin.
from captive animals, but a detailed account in the wild was recorded by Klingel.6 The mare was recumbent, lying flat on her side. The head and front legs of the foal had been expelled and were completely covered by the amnion. Parturition was completed 2 minutes later, at which time the mare sat up. The foal attempted to stand several times, breaking the amniotic sac and umbilical cord. After 14 minutes and four attempts, the foal stood successfully. The mare licked at and chewed the amniotic sac but did not eat any of it. The foal began to walk about, pulling out the placenta. In other instances observed, the placenta was retained for more than 1.5 hours. After parturition, the mare and foal slowly rejoined their harem group, which ignored the birth and the presence of the newborn foal. In all instances recorded, the harem male remained within the vicinity of the foaling mare, between 10 and 50 meters. Single births are the normal expectation although at least one instance of aborted twins has been recorded.33 These accounts are consistent with the three stages of labor recognized in domestic equids,36, 44 characterized by behavioral restlessness and the expulsion of the amniotic fluid in the initial stage, more predominant abdominal contractions and the expulsion of the foal in the second stage, and finally, expulsion of the placenta in the third stage.
Mountain zebra Parturition has not been witnessed in mountain zebras,27, 45 but is likely similar to that of other zebra species.
Grevy’s zebra As parturition approaches, a Grevy’s female leaves the herd for the safety of heavier bush.46 Birth normally takes place lying down, with the foal’s hooves appearing first, and usually takes only about 7–8 minutes. The foal frees itself from the amnion and crawls towards the mother’s head. The mother licks the foal clean, and unlike a plains zebra, ingests the membranes and some of the amniotic fluid,1, 47 which may be important in initializing lactation or the motherinfant bond in this species. Postpartum estrus
Plains zebra Klingel26 recorded the shortest intervals between two births as 378 and 385 days. The latency to foal heat has been reported to vary from 7 to 10 days after parturition.1, 6, 33 This combined with copulations by mares when their foals are only days old, indicates
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Zebra Reproduction that plains zebra exhibit an estrous period only a few days after foaling.
Mountain zebra Remnants of a corpus luteum, slightly larger than that seen at full term (averaging 2.55 cm), were found in the ovaries of a Hartmann’s zebra female 3 to 6 weeks postpartum.21 This suggests an estrus 1 week after giving birth. Observations of animals at the Daan Viljoen Game Reserve, Namibia, showed the foaling–conception interval to vary between 3 and 7 weeks, indicating that conception does not generally take place during the first postpartum estrus in mountain zebra mares.
Grevy’s zebra According to King,1 free-ranging Grevy’s zebra mares showed a range of reproductive activity postpartum. Some animals exhibited evidence of estrous cycles from 2 to 16 weeks postpartum without conception, while others conceived between 3 weeks to 4 months postpartum. According to captive studies, at least one animal has been recorded as coming into estrus about 7 days after giving birth,8 and postpartum estrus has been estimated to last from 9 to 14 days.48
Stallion Behavior In all three species of zebras the reproductive behavior of males involves interactions with females as well as breeding and bachelor males. To identify and locate females in estrus, males will often flehmen to scents deposited on the ground by the females. Once a male has found a female he will attempt to mount her; if successful he will consort with the female for varying degrees of time depending upon species. Interactions with bachelor males are typically aggressive, but depending on species, cooperation among breeding stallions is necessary to reduce cuckolding risk.10
Plains zebra Plains zebras live in multi-tiered societies. One level consists of core social groups, such as closed membership harem groups and looser associations of bachelors, while another higher level, consists of herds comprising harem and bachelor groups that come together and break apart relatively frequently.10, 12
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Herds form where resource abundance is high and where bachelor associations are numerous and strong. By banding together, stallions amortize the costs of driving away coordinated groups of cuckolding bachelors while providing their females with increased quality feeding time.
Mountain zebra Like plains zebras, mountain zebras also have closed membership harems and bachelor groups, and herds will form. Although no details of the social dynamics of mountain zebra herds exist, it is likely that herd dynamics are similar to plains zebras.
Grevy’s zebra The open membership nature of Grevy’s zebra groups reveals that relationships within and between the sexes are transitory. Males establish territories where all matings occur. Males, however, often leave their territories when no females are resident. Instead of searching for females, these males are searching for bachelors. By associating with bachelors and reinforcing their dominance of territory, territorial males are more easily able to displace bachelors when they are present on territory with females.48 Also, when on territory, stallions attempt to restrict estrous females from leaving their territories. Males mount them frequently and ejaculate at hourly intervals, flushing previous sperm from a female’s reproductive tract.9 This form of sperm competition is facilitated by the fact that Grevy’s zebra males have disproportionately large testes for their body size. Seasonality
Plains zebra Adult plains zebra stallions in the Kruger National Park, South Africa, show a greater degree of reproductive activity during the summer months, particularly in January.3 This coincides with the mares’ activity pattern where 85% of all foals are produced during the summer months, again with a peak in January.3, 6 Testicle mass and seminiferous tubule diameter increase about 24% and 13%, respectively, from the months of lowest breeding activity to the peak months. Spermatogenesis is never completely absent, regardless of activity level. Despite these peaks in activity, it is clear that plains zebra are less seasonal than other African mammals, as foals are produced throughout the year.
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Mountain zebra Mountain zebra stallions exhibit increased reproductive activity around the rains. At both the Mountain Zebra National Park (MZNP) and De Hoop National Reserve (DHNP), South Africa, Cape mountain zebra stallions show increased reproductive activity during the spring, with the majority of foals being produced the next spring/summer when the areas are flush with resources from either the summer (MZNP) or winter (DHNR) rains.20 Similarly, Westlin-van Aarde et al.22 reported that in Etosha National Park, Namibia, the majority of reproductive activity in Hartmann’s zebra stallions coincides with the start of the rains, occurring between November and April with a definite peak in February.22
females) until 4–4.5 years of age. Testicular mass increased most dramatically between the ages of 3 and 5 years old (from an average of 65 g to 302 g), while tubule development was more gradual, starting before the animals’ first year (at an average of 40 µm in diameter), and continuing into adulthood (to an average of 175 µm in diameter). Upon reaching sexual maturity at about 4–4.5 years old, the combined testes mass reached 180 g and tubule development became relatively constant. Smuts3 indicated that prime breeding stallions are under 12 years of age, but breeding does not stop in older animals, as indicated by King.1 Stallions over 18 years old still lead harems. The oldest lead stallion recorded by Smuts2 was 21 years old. The oldest male to sire young in North American zoos was almost 27 years old when the mare conceived (Table 302.1).
Grevy’s zebra For Grevy’s zebra in northern Kenya, most reproductive activity occurs early in the long rains of June and July, with another, smaller peak of activity in the short rains of September and October. This behavior coincides with the onset of estrus in Grevy’s mares, when they seek out and attract males.
Puberty and breeding life
Plains zebra Puberty in the zebra stallion can be defined as ‘the interval of time during which reproduction is possible, but terminating when full reproductive capability or sexual maturity is reached’50 (but see Skinner51 ). At the Ngorongoro Crater, Tanzania, Klingel6 reported the youngest harem stallions to be between 5 and 6 years old, with most animals leaving their natal groups between the ages of 12 and 25 months, although some remained considerably longer (up to 4.5 years of age). In Kruger National Park, South Africa,3 the youngest harem stallion was 4 years old, and 80.8% of all harem stallions were between 4.5 and 12 years old. According to the Regional Studbook for North American Zoos, the youngest sire was only 8 months old when the mare conceived and another was only 10.5 months of age (Table 302.1).19 According to observations of captive animals, stallions are physiologically mature at 3 years of age.52 In the wild, the attainment of puberty seems to be a gradual process and sexual maturity comes later.3 Puberty was estimated to last from about 2.5 to 4.5 years of age. Spermatozoa were present in small numbers in animals aged 2.5–3 years, but appreciable numbers were not present until 3.5 years of age. Males were not socially mature (able to attract and acquire
Mountain zebra Puberty in Cape mountain zebra typically begins at about 4 to 4.5 years old, with most males attaining females at about 5 years of age.45 A detailed study of testis mass and spermatogenesis of free-ranging animals was conducted by Penzhorn and Lloyd.20 The testes of bachelor males aged 11, 24, and 29 months were prepubertal in histological appearance. Active spermatogenesis in the population began at about 4 to 4.5 years of age, but this was affected by daylength. Animals approaching 4 years of age during increasing daylength conditions showed more activity than did animals approaching 4 years of age during decreasing daylength conditions. The social environment was also important. In the absence of adult males, a Cape mountain zebra colt running with mares sired a foal when only 3.5 years old. The testis mass of adult Cape mountain zebra in MZNP was found to be appreciably less (at ∼70 g) than that of other zebra species.28 The mean mass of both testes plus their epididymides was 166.4 g. In eight of nine specimens, the right testis was consistently heavier than the left. This is in contrast to what has been seen in plains zebra males.3 The mean diameter of the seminiferous tubules was shown to increase rapidly between 2 and 6 years of age before leveling off. In Hartmann’s zebra, the average weight of the testis at 87.6 g is also markedly smaller than that of the plains or Grevy’s zebra. According to Joubert,21 testis size varied with season, reaching a maximum size (99.4 g) in January and a minimum size (83.1 g) in September. The testes started increasing in weight at about 2 years of age, reaching their maximum size at around 3.5 years of age.
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Zebra Reproduction
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Grevy’s zebra
Mountain zebra
Free-ranging Grevy’s zebra males reach puberty at about 4 years of age.1 At this stage, males are likely to be living in bachelor groups, not consorting with mares and foals until a year or two later. Captive animals in England and in Kenya reached puberty between 3 and 4.5 years, but exhibited low conception rates until about 6 years of age. Read et al.,8 however, found that in the USA captive males could achieve sexual potency at ages as young as 7 months, and that one captive animal bred until 18 years of age. More recent data from the International Studbook document that the youngest males able to sire young were 2 years of age at the time the mares conceived, and that the oldest was almost 26 years (Table 302.1).25 King1 studied the testis size and weight of freeranging Grevy’s zebra in some detail. The testis of a young adult Grevy’s zebra, measuring about 11.5 × 8.5 × 7.5 cm and weighing about 475 g, is significantly larger than that of the plains or mountain zebra. According to King, testis size and weight in the Grevy’s male varied with reproductive state. A territorial male’s testis weight was recorded at 600 g, while the testis weight of an older male associated with a mixed herd was recorded at 550 g.1 The testosterone content of these animals also varied, with the young male, the territorial male, and the aged male having 154, 41.5, and 59 mg/100 µg of testis, respectively.
The most marked feature of the male reproductive organs in mountain zebra is the size of the testis. At an average of 70 g and 86 g for Cape mountain and Hartmann’s zebra, respectively, the mountain zebra testis is appreciably smaller than that of either the plains or Grevy’s zebra.21, 28
The male reproductive organs
Plains zebra The male reproductive organs of culled animals were studied in detail by King.1 The testes are carried in the scrotum, which hangs between the thighs and posterior to the penis. The deferent duct passes up from each testis, through the inguinal canal to the accessory organs that lie on the ventral floor of the pelvic cavity. The ducts (0.4 cm in diameter) enlarge to form the ampullae (1.2 cm in diameter) as they pass back over the dorsal surface of the bladder and open at the beginning of the urethra. The sac of the seminal vesicle opens into the urethra behind each ampulla anterior to the prostate gland. The prostate gland is a lobulated, bilobed gland extending 2.5 cm on each side of the urethra. The bulbourethral glands are located about 3 cm posterior and lateral to the urethra at the base of the penis. The urethra passes around the posterior border of the pubic symphysis into the penis and extends along the ventral surface of the abdomen. The extremity of the urethra is enclosed within the prepuce and lies in the region of the umbilicus.
Grevy’s zebra Even when controlling for animal size, the male accessory organs are much larger in Grevy’s zebra than in the plains or mountain zebra.1, 21, 28 The most conspicuous feature is the large seminal vesicle, which can contain up to 400 mL of fluid in the largest stallion sampled, and up to 1 liter in a captive animal at about 15 years of age. Testis size and weight are also significantly greater and vary with reproductive state of the male (see above).
Foal Plains zebra Single births were recorded by Klingel and Klingel53 in the wild and by Wackernagle33 in captivity. Klingel calculated sex ratios for the foals born in the Ngorongoro Crater, Tanzania. Of the 170 foals born, 87 were females and 74 were males, giving a ratio not different from 1:1. Foals extricated themselves from the birth membranes unaided, stood within 11 to 13 minutes, stepped after 19 minutes, walked after 32 minutes, and cantered after 44 to 60 minutes. In the one instance of actual foaling recorded by Klingel,26 the offspring suckled for the first time within 1 hour, and 12 more times in the next 4 hours for 30 seconds to 2.5 minutes a time for a total of 14 minutes. Wackernagle33 reported first suckling at 71 minutes after parturition. During the first few days to weeks of the foal’s life, the mother is very protective. This correlates with the imprinting period in which the foal learns to recognize its mother by her call, her scent, and her stripe pattern. Until imprinting is complete, the foal will follow any large object, including vehicles,6 and the mare’s protectiveness insures that the foal will not imprint on another animal. During this period, other members of the group (including the harem stallion) are not allowed to approach the foal too closely. Such behavior insures protection of the foal during a particularly vulnerable stage.54 Beyond this period, the mother and foal remain close, but others are allowed to interact and socialize with the foal. Mothers with young foals are more wary than other members of the group and are the first to move off at signs of danger.
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Mountain zebra Actual births of mountain zebra foals have not been recorded in the wild,21, 27, 55 but data regarding sex ratio at birth, mare–foal behavior, and foal activity do exist. The sex ratio at birth is not different from 1:1 in either Cape mountain or Hartmann’s zebra, and the young are just as precocious, getting up, walking, and running within a short period of time.7 During the first few weeks of life, mountain zebra foals and their mothers behave very much like other zebra species. Both Cape mountain and Hartmann’s zebra foals remain close to their mothers in the first weeks. In the first few hours after birth, Hartmann’s zebra mares spend much time sniffing and licking the foal. This may be important to insure the female’s recognition of its young.56 Both Cape mountain and Hartmann’s zebra foals make frequent tactile contact with their mothers, rubbing their faces along their mothers’ flanks, initiating naso-nasal contact, and nibbling at their necks. In both subspecies the mothers actively prevent contact between the foal and other members of the group. This can be taken to extremes in Hartmann’s zebra as some mothers will remain separate from their groups for several days.27 As in plains and Grevy’s zebra, this behavior is correlated with the critical period of imprinting of the mare’s sound, scent, and stripe pattern onto the foal. Mares of both species are more skittish than other group members and are the first to flee at any sign of danger.
The earliest weaning occurred at 11 months of age, but if a mare does not conceive soon after foaling, she will continue to suckle her foal throughout and beyond its first year. One captive mare suckled her foal until it was 16 months old, weaning it 6 months prior to the birth of her next foal.
Mountain zebra On average, mountain zebra foals are weaned at about 10 months of age,27 about 3.5 to 6 months before the new foal is born. For Cape mountain zebra, the timing of weaning is highly dependent on the birth of new offspring and can be quite variable, but for Hartmann’s zebra, 10 months seems to be the norm. If mares do not conceive during postpartum estrus, suckling may be permitted for longer periods. Penzhorn45, 57 reported infrequent suckling in animals as old as 20 months of age.
Grevy’s zebra Grevy’s zebra foals are weaned at about 9 months (32–42 weeks1 ), but will remain with their mothers for several more months or years, depending on the sex (see below). The young are semi-independent at about 7 months, often being left by the mother or leaving her for hours at a time.
Yearling
Grevy’s zebra
Plains zebra
Grevy’s zebra foals are similar to both plains and mountain zebra foals. Single births have been recorded both in the wild1 and in captivity.8 Data from captive animals show that the young are precocious, standing 6–14 minutes after birth, walking by 30 minutes, and running by 45 minutes. At about 29 minutes the foal attempts to nurse, and does so by 69 minutes. At 97 minutes the foal stands well and at 2 hours nurses well. In captivity, the sex ratio at birth is 1.09:1.5 The Grevy’s newborn will follow any large, moving object. The mother is quite protective (particularly during the first 2 days) and keeps other animals away until her offspring has imprinted upon her call, scent, and stripe pattern. After imprinting has been achieved, the mother and foal will join a group of nursing mothers. When traveling to water, the mare will sometimes leave the foal with this group.38
Young mares at the age of 1–2 years, around the time of their first heat, are abducted from their families by stallions. Klingel6 reported that on average, mares were abducted at about 18 months, but as young as 13 months of age. These young females do not typically remain with the initial stallion, but within about 6 months of separating from their natal group, become permanent members of a reproductive group. Most young stallions leave the natal group at about 12–25 months of age, in some cases later, and either stay single or join other stallions to form bachelor groups. Bachelor groups are of variable numbers, up to 15 animals. Smaller groups of two, three, and four have been reported to be very stable. Departure of the young males is not correlated with sexual maturity or any antagonism with the lead stallion, but seems to be determined more by the composition of the natal group. In particular, the males tend to leave after the birth of their mothers’ next foal.
Weanling Plains zebra
Mountain zebra 2
According to Smuts, lactation generally continues until 1 to 2 months before the birth of the next foal.
The timing of dispersal is similar for males and females for both Cape mountain and Hartmann’s
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Zebra Reproduction zebras. In Mountain Zebra National Park (MZNP), male and female Cape mountain zebras left their natal groups at an average of 21.85 months.45, 57 In De Hoop Nature Reserve (DHNR), however, females leave their groups earlier, at 18.85 months, than do the males, at 24.36 months.17 Female dispersal coincides with estrus onset, but there is no physiological correlate in males. In both MZNP and DHNR, the yearlings leave the group of their own accord and are not forced to do so by either the stallion or the mare. In addition, dispersal is not correlated with the mothers’ subsequent pregnancies or her status,17 and stallions in MZNP will vigorously attempt to retain their yearling offspring.45, 57 Dispersal in young mountain zebra is likely due to bond transference from maternal/paternal to age cohort associations. This is similar to what Klingel6 reported in plains zebra. In stark contrast, Hartmann’s zebra typically leave the natal group at around 14 to 16 months of age.27 At this stage (shortly before the birth of her next foal), the mare actively tries to drive the older offspring (whether male or female) out of the group. Approximately one-third of yearlings leave the group as a direct result of this aggressive behavior. The remaining two-thirds leave of their own accord, sometimes as much as 2 years later. Upon the birth of her next foal, the mare will ignore the yearling if it has not already left. Young Cape mountain mares often join bachelor groups before joining a more stable, reproductive group. This was always the case for mares in DHNR,26 and very often the case for mares in MZNP.45, 57 At DHNR, the females stay in the bachelor groups approximately 8.7 months, leaving when they are about 29.9 months old. The females are not abducted by harem stallions; they leave of their own accord. At MZNP, females will sometimes remain solitary for several months before joining either bachelor or reproductive groups. Older mares are taken up by harem stallions sooner, while younger mares are either solitary or in bachelor groups for longer periods. Young Cape mountain males in DHNR stay in their natal groups longer than the females, for an average of 24.36 months, leaving when they are about 52.44 months of age.17 These data agree with Penzhorn’s45, 57 estimate of the age at sexual maturity. In the MZNP, young males roam singly or in unstable groups only loosely associated with the more stable bachelor groups.57 Affiliation with the natal group is strong; young males having left after the displacement of their fathers will often join them. In addition, young males having left their natal group still intact, will often rejoin the group for short periods. Hartmann’s zebra females are typically herded by either bachelors, forming the core unit of a new reproductive group, or by harem stallions, thereby
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integrating into an already stable reproductive group. Young males leave their natal group to join bachelor groups that consist of mostly young, but also older, displaced stallions.27
Grevy’s zebra Young mares leave their mothers at about 13–18 months of age, typically 2–7 months into the mothers’ next pregnancy.1 The females will not breed until they are between 2 and 4 years old. During their first estrus (at around 4 years of age), the males they attract separate them from their mothers to mate.58 More experienced mares will present only to territorial males, causing less disruption to the female groups. Grevy’s males stay with their mothers until they are about 3 years old, after which they join a bachelor group or a mixed herd.1 Young males do not typically gain access to females until they are about 5 years old, so have a bachelor period of 2 to 5 years.6
References 1. King JM. A field guide to the reproduction of the Grant’s zebra and Grevy’s zebra. East African Wildlife Journal 1965;3: 99–117. 2. Smuts GL. Reproduction in the zebra mare (Equus burchelli antiquorum) from the Kruger National Park. Koedoe 1976;19:89–132. 3. Smuts GL. Reproduction in the zebra stallion (Equus burchelli antiquorum) from the Kruger National Park. Zoologica Africana 1976;11:207–20. 4. Asa CS, Bauman JE, Houston EW, Fischer MT, Read B, Brownfield CM, Roser JF. Patterns of excretion of fecal estradiol and progesterone and urinary chorionic gonadotropin in Grevy’s zebras (Equus grevyi): ovulatory cycles and pregnancy. Zoo Biology 2001;20:185–95. 5. Dobroruka LJ, Holejsovska A, Maslova I, Novotny V. An analysis of the poulation of Grevy’s zebra (Equus grevyi) in Dvur Kralove Zoo. International Zoo Yearbook 1987;26:290– 3. 6. Klingel H. Reproduction in the plains zebra, Equus burchelli boehmi: behavior and ecological factors. J Reprod Fertil Suppl 1969;6:339–45. 7. Penzhorn BL. Reproductive characteristics of a free-ranging population of Cape mountain zebra (Equus zebra zebra). J Reprod Fertil 1985;73:51–7. 8. Read B, Kelly C, Bueler M, Junge R. Grevy’s zebra managment survey. In: American Association of Zoological Parks and Aquariums, Species Survival Plan. St Louis, MO: St Louis Zoological Park, 1988; pp. 160. 9. Ginsberg JR, Rubenstein DI. Sperm competition and variation in zebra mating behavior. Behav Ecol Sociobiol 1990; 26:427–34. 10. Rubenstein DI, Hack M. Natural and sexual selection and the evolution of multi-level societies: insights from zebras with comparisons to primates. In: Kappeler P, Schaik CP v (eds) Sexual Selection in Primates: New and Comparative Perspectives. Cambridge University Press, 2004; pp. 266– 79.
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11. Klingel H. A comparison of the social organization of equids. In: Proceedings of a Conference on the Ecology and Behavior of Feral Equids. University of Wyoming, Laramie, 1979. 12. Rubenstein DI. Ecology and sociality in horses and zebras. In: Rubenstein DI, Wrangham RW (eds) Ecological Aspects of Social Evolution: Field Studies of Birds and Mammals. Princeton, NJ: Princeton University Press, 1986; pp. 282– 302. 13. Rubenstein DI. The ecology of female social behavior in horses, zebras and asses. In: Jarman P, Rossiter A (eds) Animal Societies: Individuals, Interactions and Organisations. Kyoto: Kyoto University Press, 1994; pp. 13–28. ˜ C. Sociality and reproductive skew 14. Rubenstein DI, Nunez in horses and zebras. In: Hager R, Jones CB (eds) Reproductive Skew in Vertebrates: Proximate and Ultimate Causes. Cambridge University Press, 2009; pp. 196–226. 15. Ginsberg JR. The ecology of female behaviour and male mating success in the Grevy’s zebra. Symp Zool Soc Lond 1989;61:89–110. 16. Rowen M. Mother–infant behavior and ecology of Grevy’s zebra, Equus grevyi. New Haven, CT: Yale University, PhD thesis, 1992. 17. Lloyd PH, Rasa OAE. Status, reproductive success and fitness in Cape mountain zebra (Equus zebra zebra). Behav Ecol Sociobiol 1989; 25:411–20. 18. Fischhoff IR, Sundaresan SR, Cordingley J, Larkin HM, Sellier M-J, Rubenstein DI. Social relationships and reproductive state influence leadership roles in movements of plains zebra (Equus burchellii). Anim Behav 2007;73:825– 31. 19. Minot J. Plains Zebra North American Regional Studbook. Atlanta, GA: Zoo Atlanta, 2004. 20. Penzhorn BL, Lloyd PH. Comparisons of reproductive parameters of two Cape mountain zebra (Equus zebra zebra) populations. J Reprod Fertil 1987;35:661–3. 21. Joubert E. Notes on the reproduction in the Hartmann zebra Equus zebra hartmannae in South West Africa. Madoqua 1974;8:31–5. 22. Westlin-van Aarde LM, van Aarde RJ, Skinner JD. Reproduction in female Hartmann’s zebra, Equus zebra hartmannae. J Reprod Fertil 1988;84:505–11. 23. Kingdon J. East African Mammals; an Atlas of Evolution in Africa. Chicago, IL: University of Chicago Press, 1979. 24. Williams SD. Grevy’s zebra: Ecology in a heterogeneous environment. PhD thesis, University College London, 1998. 25. Langenhorst T. Grevy’s Zebra International Studbook. Winchester, UK: Marwell Zoo, 2004. 26. Klingel H. Notes on the biology of the plains zebra Equus quagga bohemi matschie. East African Wildlife Journal 1965;3:86–8. 27. Joubert E. The social organization and associated behaviour in the Hartmann zebra Equus zebra hartmannae. Madoqua 1972;6:17–56. 28. Penzhorn BL, van der Merwe NJ. Testis size and onset of spermatogenesis in Cape mountain zebras (Equus zebra zebra). J Reprod Fertil 1988;83:371–5. 29. Benesch F, Wright J. Veterinary Obstetrics. London: Bailli`ere, Tindall, and Cox, 1957. 30. Asa CS. Equid reproductive biology. In: Moehlman PD (ed.) IUCN/SSC Equid Action Plan. Gland, Switzerland: IUCN, 2002; pp. 113–17. 31. Kupfer M. The sexual cycle of female domesticated mammals. In: Department of Agriculture 13th and 14th Reports
32.
33.
34. 35.
36. 37.
38. 39. 40. 41.
42. 43.
44. 45.
46. 47.
48.
49.
50.
51. 52.
53.
54.
of the Director of Veterinary Education and Research, 1928, pp. 1211–70. Hammond J, Wodzicki K. Anatomical and histological changes during the oestrous cycle in the mare. P Roy Soc Lond B Bio 1941;130:1–23. Wackernagel H. Grant’s zebra Equus burchelli boehmi, at Basle Zoo – a contribution to breeding biology. International Zoo Yearbook 1965;5:38–41. Brown CE. Rearing wild animals in captivity and gestation periods. J Mammal 1936; 17:10–13. Smuts GL. Pre- and postnatal growth phenomena of burchell’s zebra (Equus burchelli antiquorom). Koedoe 1975; 18:69–102. Asa CS, Goldfoot DA, Ginther OJ. Assessment of the sexual behavior of pregnant mares. Hormon Behav 1983;17:405–13. Penzhorn BL. Behaviour and population ecology of the Cape mountain zebra Equus zebra zebra Linn, 1758 in the Mountain Zebra National Park. Pretoria: University of Pretoria, DSc thesis, 1975. Groves CP. Horses, Asses, and Zebras in the Wild. Newton Abbott and London: David and Charles, 1974. Antonius. Einige bemerkenswerte Zuchterfolge in Schonbrunn im Jahre 1931. Zoological Garten 1932;5:91–7. Kenneth JH. Gestation Periods. Slough, UK: Commonwealth Agricultural Bureau, 1953. Squires EL, Douglas RH, Steffenhagen WP, Ginther OJ. Ovarian changes during the estrous cycle and pregnancy in mares. J Anim Sci 1974;38:330–8. Roser JF, Lofstedt RM. Urinary eCG patterns in the mare during pregnancy. Theriogenology 1989;32:607–22. Ramsay EC, Moran F, Roser JF, Lasley BL. Urinary steroid evaluations to monitor ovarian function in exotic ungulates: X. Pregnancy diagnosis in Perissodactyla. Zoo Biology 1994;13:129–47. Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects, 2nd edn. Cross Plains, WI: Equiservices, 1992. Penzhorn BL. A long-term study of social organisation and behaviour of Cape mountain zebras Equus zebra zebra. Zeitschrift fuur ¨ Tierpsychologie 1984;64:97–146. Churcher CS. Equus grevyi. Mammalian Species 1993; 453:1–9. King JM, Short RV, Mutton DE, Hamerton JL. The reproductive physiology of male zebra-horse and zebra-donkey hybrids. Symp Zool Soc Lond 1966;15:511–27. Nelson L. Equidae. In: Fowler ME (ed.) Zoo and Wild Animal Medicine. Philadelphia, PA: W.B. Saunders and Co., 1986; pp. 926–31. Ginsberg JR. Social organization and mating strategies of an arid adapted equid: The Grevy’s zebra. Princeton, NJ: Princeton University, PhD thesis, 1988. Emmens CW. Physiology of gonadal hormones and related synthetic compounds. In: Cupps CaPT (ed.) Reproduction in Domestic Animals. London: Academic Press, 1969. Skinner JD. The concept of puberty. S Afr Med J 1969;43: 1–420. Trumler E. Beobachtungen an den Bohmzebras des George von Opel Freigeheges fur Tierforschunge. Saugetierk Mitt 1958;6:1–48. Klingel H, Klingel U. Die Geburt eines Zebras (Equus quagga bohemi Matschie). Zeitschrift fur ¨ Tierpsychologie 1966;23:72–6. Fischhoff IR, Sundaresan SR, Larkin HM, Cordingley JE, Sellier M-J, Rubenstein DI. A rare fight between two female plains zebra (Equus burchelli). J Ethol 2010;28:201–5.
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57. Penzhorn BL. Observations of mortality of free-ranging Cape mountain zebra Equus zebra zebra. S Afr J Wildl Res 1984;14:89–90. 58. Churcher CS. Grevy’s: the other zebra. Swara 1982;5:12– 18.
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Assisted Reproductive Technologies (ART)
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Embryo Transfer David L. Hartman
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Cooled Transported Embryo Technology Patrick M. McCue, Catherine A. DeLuca and Jillian J. Wall
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Freezing of Embryos Jean-Franc¸ois Bruyas
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Import and Export of Embryos Erin C. Bishop
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Nuclear Transfer Katrin Hinrichs
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Chapter 303
Embryo Transfer David L. Hartman
Introduction Aside from semen collection and artificial insemination, embryo transfer has become the most widely recognized assisted reproductive technique used in the horse. This brings the equine practitioner an exceptional opportunity for practice expansion, but also entails tremendous responsibility. What started in the early 1980s as a novel method of producing a foal from aged non-productive broodmares has now expanded to become an elective procedure that is applicable to virtually all mares of reproductive age. This transition to a widely used elective procedure carries the responsibility commonly known as ‘above all else do no harm’ to the involved veterinarian. Embryo transfer (ET) now lends itself to obtaining: (i) foals from performance and show mares; (ii) multiple foals from the same mare in one year; (iii) foals from mares with non-reproductive health or musculoskeletal problems; and (iv) foals from mares with reproductive problems. In addition, a mare that foals late in the year can be allowed early entry into the next breeding season without the loss of a foal through embryo transfer procedures. The use of embryo transfer has also enabled the development of more sophisticated advanced reproductive techniques such as intracytoplasmic sperm injection (ICSI) and nuclear transfer (cloning). Opportunities seem to be bound only by our imagination. Expectations for success have risen dramatically since the adoption of commercial embryo transfer. Results that were unimaginable just a few years ago are now very achievable. For the sake of discussion, embryo recovery rates are equal to, or above, the percycle pregnancy rate of a given stallion. This translates to recovery of 100% of embryos that are in the uterus at the time of embryo retrieval. With regard to embryo transfer success, the author contends that we should set a goal of a 90% pregnancy rate at 12 days and 85% at 30 days post-ovulation. One of the deceptive aspects of embryo transfer is that some de-
gree of success may be achieved following relatively poor technique. The end result of this is an everpresent temptation to ‘cut corners’, e.g., to perform a uterine flush or embryo transfer under less-thanoptimal conditions or by using less-than-optimal supplies or media. Conventional methods of embryo recovery, identification and transfer have been described.1–4 The objective of this chapter is to discuss what has worked well in our practice and also to discuss some of the more controversial topics related to embryo transfer.
Recipient mare management prior to transfer Management of a large recipient herd is a daunting and labor-intensive task. This begins in the autumn of the previous year. During this period, it is important to acquire the total number of mares that will be needed for the coming breeding season. There should not be any new mares added during the breeding season that may introduce contagious disease into the herd. The risk is too great for both the empty and pregnant mares. All individual mares should have a means of permanent identification so that there cannot ever be any confusion regarding identification. One of the most common complaints among mare owners concerns the quality of the recipient mares. It is important to select recipients that are sound not only reproductively but also with regard to musculoskeletal soundness, dental health, and vision. The examination should include evaluation of udder health because the ability to provide sufficient milk for the foal is also frequently a concern of mare owners. All mares should be rigorously assessed with regard to their temperament and disposition. At the least, mares should be halter broke and gentle enough to catch in a small pen or stall. The autumn and winter is also a good time to manage vaccination, deworming, and Coggins testing as mares enter into the recipient population. Testing and vaccination
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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for equine arteritis virus should be addressed where geographically applicable. It is always a challenge to have sufficient mares with normal reproductive cyclicity by February (in the northern hemisphere) when the commercial breeding season begins. As with all broodmares, a good artificial lighting program is the most reliable method for ensuring early seasonal entry into reproductive cyclicity. For this protocol, mares are exposed to a total of 16 hours of light each 24-hour period beginning the middle of November. The challenge centers on a good method of artificial lighting that can be used on large numbers of mares. The most common system involves holding the mares in large pens during the artificial lighting period. This can be stressful to the mares as they may be crowded, unable to access feed or water, and/or are often exposed to inclement weather during this confinement. There are studies that discount the effect of stress on mares;5 however, such studies generally examine a single stress insult rather than assessing cumulative stressors that occur day after day. There exists ample anecdotal evidence that high stress may have a detrimental effect on such factors as maintenance of a corpus luteum and the mare’s ability to maintain early pregnancy. With the onset of the breeding season, the goals of managing empty recipient mares are twofold. First, it is necessary to know precisely the day that each mare ovulates. This is best accomplished by palpation and ultrasonography of the reproductive tract per rectum during behavioral estrus. This should be done daily as the mare approaches ovulation and, oftentimes, for at least one day post-ovulation. For recordkeeping purposes, the day of ovulation is always considered Day 0. The next day post-ovulation the mare becomes ‘Plus 1’, the second day ‘Plus 2’, and so forth. The commonly accepted window of synchrony between donor and recipient mares is such that recipient mares that are +4 to +8 days post-ovulation are considered to be eligible to receive embryos.1, 2 This may be described in several different ways. The important concept is that during this time the mare can respond to early signals involving maternal recognition of pregnancy and should also have adequate levels of circulating progesterone. One study6 successfully lengthened this window of synchrony, thereby enabling the use of recipients that were +9 to +10 days from their ovulation. This was accomplished by the administration of meclofenamic acid (1 g p.o., once daily) starting on Day 9 post-ovulation and continuing until Day 7 after embryo transfer. With this protocol, the investigators were able to suppress the normal cyclical luteolytic response. If determining the day of ovulation becomes the primary focus, then it is easy to overlook a difficult
aspect of recipient mare management, which is endometritis. Therefore, the second goal of managing open recipient mares is to insure that they are reproductively sound and ready to receive an embryo. Begin by considering only mares that are 3–12 years of age for admission into the recipient herd. This should eliminate most mares with substantial chronic endometritis that would interfere with their ability to maintain a pregnancy to term. Next, the anatomy of the perineum and vulva is evaluated. Very thin mares often have poor perineal anatomy that may improve when they gain weight. If indicated, a Caslick’s operation should be performed. The importance of perineal anatomy cannot be overstated as bacterial contamination at the time of embryo transfer may be the most frequent cause of embryonic loss. It is apparent then that per rectum palpation and ultrasonographic examinations during estrus serve not only to note the day of ovulation, but just as important, to continually evaluate the condition of the uterus. If indicated, uterine swabs should be procured for microbial culture. In the author’s view, it is imperative that cultures be taken only when the mare is in advanced estrus. The mare should have at least a 35 mm follicle that is beginning to mature and the uterus should have substantial edema at the time of uterine culture. If these conditions are not met, false-negative cultures may be obtained. In addition, care should be taken not to contaminate the uterus during the process of taking the culture. Concurrently, control of delayed or inadequate uterine clearance is important. Oxytocin (20 IU, intravenously) may be used as needed but not more often than every 4 hours. Recipient mares should be monitored for at least one day post-ovulation to be certain there is no fluid in the uterus as they enter diestrus.
Donor mare management prior to the uterine flush Proper management of donor mares centers around the ultimate goal – to recover as clean an embryo as possible and of a size that can be transferred easily. Keeping this in mind, it is important to realize that donor mares with endometritis will become pregnant on a regular basis. This is contrary to conventional thought that mares with endometritis don’t become pregnant. It is not uncommon to find an embryo in a very dirty flush (Figure 303.1). Regardless of the number of times the embryo is washed, it is difficult to get these embryos to survive following embryo transfer. Therefore, it is prudent to make sure the donor mare’s uterus is free of microorganisms at the time of insemination. Once-daily palpation and ultrasonographic examination allow one to follow
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Figure 303.1 It is not uncommon to find an embryo (arrow) in a dirty flush. Reprinted from Molecular and Cellular Endocrinology (228), Havelock JC, Rainey WE, Carr BR. Ovarian graulosa cell lines, pp 228: 67–78. © 2004, with permission from Elsevier.
Figure 303.2 Endometritis following repeated insemination and embryo transfer attempts. Allen TW. 2001. Reproduction. 121:513–527. © Society for Reproduction and Fertility (2001). Reproduced by permission.
follicular development and ovulation, but it is just as important to monitor the conditions of the uterus. The time and effort spent keeping the donor mare’s uterus clean will be rewarded with increased rates of successful embryo transfer. Keep in mind that bacterial contamination of the donor’s uterus can come from many sources. Among these are inadvertent contaminations during the breeding process, contaminated semen, and, increasingly commonplace, residual infection and endometritis from previous reproductive procedures. It is becoming more common to perform multiple embryo transfers on the same mare during a breeding season, oftentimes at different farms and by different veterinarians. One study7 analyzed the effects of repeated inseminations and embryo transfer attempts on uterine health and found an association with increased chronic inflammatory changes. These workers concluded that mares were susceptible to the additive effects of repeated uterine insults. In clinical practice, there is substantial evidence that repeated breeding and embryo transfer attempts yields an increased likelihood of acute bacterial endometritis, in addition to chronic inflammation. Therefore, proper donor mare management requires continued vigilance to guard against endometritis (Figure 303.2). Uterine cultures may be required on donor mares on a regular basis throughout the breeding season. However, the overuse of intrauterine antibiotic therapy should be discouraged as this may lead to overgrowth of a yeast or fungus. One of the most effective ways to keep a mare’s uterus in a hygienic state is through effective uterine lavage along with ecbolics to aid in uterine clearance. One factor that affects embryo transfer success is the size of the embryo at the time of transfer to the re-
cipient mare. Large embryos (> 1.5–2.0 mm) present their own set of problems. For example, they are somewhat more fragile and it is sometimes difficult to find a transfer apparatus that can accommodate the larger embryos. On the other hand, if a donor mare’s uterus is flushed too early, the embryo may still be in the oviduct and not available for traditional recovery attempts. In fact, it is not uncommon for a mare to end up pregnant following uterine lavage for embryo retrieval. Therefore, selecting the correct flush day is of paramount importance. The ideal embryo is a 7-day embryo, at which time its diameter is approximately 400–500 µm. However, it is not so simple as to automatically flush all mare uteri at 7 days after ovulation. It has been established8 that the timing of embryo entry into the uterus is somewhat variable, ranging between 144 hours (6 days) and 156 hours (6.5 days) after the first observation of ovulation. Examination of the corpus luteum in the periovulatory period is important in that scheduling a 7-day flush date from a recent ovulation often results in a smallerthan-expected embryo. There seems to be several factors that may influence this variability in embryo size. These are: (i) the age of the donor mare; (ii) the inherent fertility of the stallion; (iii) the timing of breeding relative to the time of ovulation; and (iv) variation in time between ovulation and fertilization. In our practice, most reproductively normal mares are flushed 7 days after the first appearance of ovulation. A simple way to plan for the embryo flush attempt is to consider the day of the week the ovulation was first detected. For example, if Monday is the first appearance of the corpus luteum, the mare’s uterus would be flushed on the following Monday for a 7-day flush. Among the exceptions to this are: old mares (18 years or older); mares bred to subfertile stallions or older
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stallions; and mares bred with frozen semen. These are all flushed on Day 8. During the week between ovulation and the uterine flush, donor mares should be kept as stress-free as possible. There is anecdotal evidence that performance mares in training may have reduced embryo recovery rates. This may be, in part, due to the fact that some mares will have a significant increase in body temperature during strenuous physical activity. Older mares and mares that ovulate small follicles may benefit from progesterone supplementation during this week. The most practical way to do this would be with altrenogest (0.044 mg/kg orally, once daily) until the uterine flush. Another indication for this would be the mare that has multiple follicles, doesn’t develop a good corpus luteum following ovulation, or ovulates follicles separated by 3–4 days.
Uterine flush for embryo recovery The basic methods used for embryo recovery and identification have been described previously.1–4 In general, this is accomplished by three or four trancervical uterine lavages with the recovered fluid passing through a filter so as to capture the retrieved embryo (Figure 303.3).
Figure 303.3 Author’s preferred method for embryo recovery.
A wide variety of embryo transfer supplies are readily available from commercial distributors for equine embryo transfer procedures.a Product selection is based, to a large extent, on personal choice. To begin, a Foley-type balloon-tipped catheter of sufficient length is required in order to extend from the uterus externally to a Y-junction that connects one end to the flush medium and the other end to the filter. Most of these supplies have been gamma irradiated for sterilization prior to sale. Following use, the decision as to whether these supplies will be reused for embryo transfer is again up to the individual. If so, the type of sterilization is a problem. Only the most expensive catheters may be steam autoclaved. Gas sterilization presents a problem with regard to the amount of time following the sterilization that it is safe prior to exposure to an embryo. Any disinfectants that are used in a washing program may also be potentially harmful to an embryo. For these reasons, in our practice all supplies are only used once for embryo transfer procedures and then reused thereafter for simple uterine lavage. The type of flush medium used is again a matter of personal preference. It may be ordered with or without antibiotics. If antibiotics are desired, the most common medium that is available contains kanamycin. Most flush media also contain ingredients that aid in preventing the embryo adhering to the tubing and plastic ware. The time-tested ingredient for this purpose is bovine serum albumin (BSA). The primary drawback of BSA is the propensity of the BSA to form bubbles or froth on the top of the recovered medium. If this is extensive, then it is more difficult to search for embryos. More recently, polyvinyl alcohol is being used to replace the BSA. With this alternative ingredient, bubble formation is minimized (Figure 303.4). As stated earlier the flush procedure is basically a transcervical uterine lavage. The exact methodology varies among individuals. However, there are several principles that should be followed in order to ensure embryo recovery. Immediately prior to the uterine flush, the donor mare should be examined by palpation and ultrasonography per rectum. The ovaries should be examined for the presence of a corpus luteum; if more than one corpus luteum is present there is an increased possibility of multiple embryos, regardless of the breeding information that may have been a Bioniche Animal Health, 1551 Jennings Mill Rd., Suite 3200A, Bogart, GA 30622; Hamilton-Thorne Biosciences, Inc., 100 Cummings Center, Ste. 46SE, Beverly, MA 01915; Minitube of America, PO Box 930187, Verona, WI 53593; Partnar Animal Health, 3560 Pine Grove Ave. #227, Port Huron, MI 48060; Professional Embryo Transfer Supply, PO Box 188, Canton, TX 75103–0188; Reproduction Resources, 400 South Main Street, Walworth, WI 53184; Veterinary Concepts, PO Box 39, Spring Valley, WI 54767.
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Figure 303.4 One filter handling technique that will minimize bubble formation in the recovered flush medium.
provided. In addition, the uterus should be examined. The presence of fluid is naturally a concern but does not preclude the possibility of procuring an embryo. The disposition of the mare is evaluated at this time in order to determine the amount of sedation that will initially be used for the uterine flush procedure. Sedation of the mare is important because the uterus will be distended with fluid which often leads to discomfort. Oversedation of the mare can be detrimental if the mare cannot stand properly on both back legs. Finally, the bladder should be palpated and emptied if full. The reason for this is that a full bladder will displace the uterus. As the uterus fills it will become large and heavy, falling to one side of the full bladder. When this happens, the uterine horn that is adjacent to the bladder will not fill or empty well because of interference from the broad ligament of the bladder which is, at this point, stretched and tight. Prolonged waiting in a strange environment with strange mares often decreases urination and results in mares with full bladders. For these reasons mares should not be held for prolonged times in pens waiting for flushing or transfer. Following this initial examination, the mare’s tail should be wrapped and secured to one side. Thorough washing of the perineal area, vulva, and vestibule is essential to prevent contamination of the catheter as it is passed into the uterus. Contamination at this point could easily result in a failed embryo transfer attempt. Once the mare’s hindquarters have been prepared, the flush will generally take 15–20 minutes, depending on the number of flushes attempted. Ideally, the mare is not sedated until immediately prior to starting the uterine flush because if the rectum fills with air, as often occurs following sedation, per rectum
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manipulation of the uterus is severely compromised. Delaying sedation until this time will help prevent this problem. The flush medium may be pre-warmed if so desired, but is not mandatory. The author uses medium at room temperature (20–25◦ C) as embryos are quite tolerant of temperature change, provided that it is not too drastic. If the medium is pre-warmed, care should be taken not to get above body temperature (37◦ C). All tubing, the filter, and the catheter should be pre-filled with medium prior to beginning the procedure so as to prevent air in the system. The catheter is placed through the cervix just to the level of the internal os. The balloon is inflated with either air or flush medium. The amount of inflation varies with the mare. A mare whose cervix does not close well needs a larger balloon to prevent the catheter from slipping back through the cervix. Conversely, if the balloon is too large, the catheter cannot be moved around in the uterus easily. In most mares, 30–40 mL of inflation is adequate. Once the catheter is in place, start the fluid flow into the uterus. From this point on, the practitioner’s skill at palpation per rectum is very important, as all manipulation and monitoring of the flush is accomplished in this fashion. Begin by palpating the entire uterus to insure that both horns fill completely. This is simple in young, dry mares, but can be difficult in older mares and all lactating (wet) mares. Regarding wet mares, the only uterine horn that tends to fill is the horn that recently carried the fetus. This becomes evident during the first flush. In order to prevent this, on successive flushes, this uterine horn is manually held closed at the bifurcation in order to force fluid into the other (previously non-gravid) uterine horn. When both horns are full as determined per rectum, stop the inflow, and begin to recover the flush medium by reversing the flow at the Y-junction and directing the outflow through the filter. The amount of medium that is used is dependent on the individual mare. The uteri of young maiden mares may only hold 750–1000 mL, while older mares and wet mares easily hold 2 liters. If the uterus cannot be filled with 2 liters of medium, administer oxytocin (20 IU, intravenously). Once recovery of the fluid begins and as the pressure on the cervix is reduced, advance the catheter into one of the uterine horns (alternate sides with successive flushes). In the author’s experience, fluid is easily moved from the opposite horn into the horn with the catheter tip in order to completely empty the uterus. If the catheter is left at the internal os of the cervix, it is usually not possible to empty the uterus completely. This entire process should be repeated for at least three complete flushes. One study9 has described a fourth flush in which the medium is allowed to remain in the uterus for at least 3 minutes prior to recovery of the fluid. In this study, a significant number of embryos were
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recovered in the fourth (extra) flush attempt, when no embryo was recovered during the first series of three flushes. Failure to completely empty the uterus, especially on the last flush, can predispose the donor mare to endometritis. It may be helpful to examine the mare by ultrasonography when finished in order to insure that all fluid has been recovered. Several different types of filters are available. Most require that the medium is transferred into a search dish for examination. The filter should also be rinsed into the search dish with flush medium. It is the author’s preference to use a round dish with a grid on the bottom to aid the search process. Embryos that are 6–9 days of age will sink to the bottom. Therefore, the focus of the microscope should be on the bottom of the dish. This is most easily accomplished by initially focusing on the grid lines or on debris that has settled to the bottom. It is very easy to miss a small embryo by incorrect focusing. There is often a significant amount of debris in the flushed fluid, especially with older mares and wet mares. This is relatively normal, but if the flush is cloudy (not transparent), it generally signifies an active endometritis. It would be easy to make the argument that recovered medium from the flush should always be cultured so as to give a starting point on therapy for the recipient mare, if needed. Embryo identification3 may be difficult for those individuals who are inexperienced. Usually the most challenging decision would be to differentiate between an unfertilized oocyte (UFO) and a morula (6-day embryo). Both appear similar under the microscope and are the same size (about 170 µm). However, the morula is spherical in nature. The UFO, while appearing round or somewhat oval under the scope, is flat when examined from the side. This is most easily appreciated when the UFO is manipulated while viewing it under the scope (Figure 303.5). In a commercial embryo transfer facility, all embryos
Figure 303.6 Simple transfer of the embryo from one dish well to another well used in the embryo washing process.
regardless of their grade should be transferred. It is not possible to determine which embryos of poorer quality will grow and which will not. Once the embryo has been identified it may be handled using a 0.25 or 0.5 mL semen freezing straw. There are several methods that can be used to wash an embryo. Commercially available multi-well dishes provide the most practical approach. Bielanski10 reviewed the most recent recommendations for washing and disinfecting embryos. Simple transfer of the embryo from one dish well to another well with clean medium, using a dilution factor of at least 1:100, is standard procedure (Figure 303.6). This is repeated up to 10 times for optimal results. Most microorganisms will be removed with this process but others may bind to the embryo and are not removed. Following the washing procedure, embryos may be transferred immediately if the recipient mare is on site or they may be transported the same day or even overnight to an alternate location for transfer to a recipient mare. For transportation, embryos can be placed into one of several types of commercially available holding medium. The customary method for embryo transport is to cool the embryos in the same systems that are used for cooling transported semen. Because there are many variables involved, it would be difficult to determine if there is a detrimental effect on the embryo from transporting and cooling. However, in our practice, there appears to be about a 5% loss that may be associated with transportation and cooling over a 4- to 8-hour period of time.
Non-surgical transfer of embryos
Figure 303.5 Unfertilized oocyte (UFO).
In the early years of equine embryo transfer, most embryos were transferred surgically through a standing flank laparotomy. In the last several years, results obtained with non-surgical transfer have surpassed the
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Embryo Transfer success rates of the surgical method. The philosophical approach to non-surgical transfer can be summarized as follows: use aseptic technique, with minimal insult to the cervix to deposit the embryo into the lumen of the uterus of a suitable recipient. As stated earlier, the accepted window of synchrony is using recipient mares that are +4 to +8 days from their ovulation, with mares +5 and +6 days generally being the best. Prior to transfer, the recipient mare should be examined by palpation and ultrasonography per rectum. The mare should have good uterine tone and a tightly closed cervix indicative of an acceptable circulating progesterone concentration. On ultrasonographic examination, there should be a demonstrable corpus luteum and preferably some follicular development. The echotexture of the uterus should be uniform with no edema or fluid. The disposition of the mare is evaluated to determine the level of sedation that is necessary prior to performing the embryo transfer. Ideally, the mare will be sedated enough to accept the procedure, yet stand properly on both back legs. Once again, sedation is delayed until immediately prior to the transfer so as to minimize air entry into the rectum. The mare’s tail is wrapped and tied or held to the side. A very thorough washing of the perineum, vulva, and vestibule is mandatory. The exact apparatus used to transfer the embryo is not as important as the technique. The author prefers a stainless steel gun (modified Cassou) with a disposable, steel-tipped, side delivery sheath.b There are two sizes; one accepts a 0.25 mL straw and the other a 0.5 mL straw. The embryo should not occupy more than 60–70% of the internal diameter of the straw, and preferably less. A system of loading the straw in our facility is to place the embryo in the middle fluid bubble of a three-bubble system (Figure 303.7). The first bubble of fluid pulled into the straw is the last one out and is therefore the ‘push’ bubble. This will help to clear and carry the embryo away from the tip of the sheath. It has been reported11 that embryos may remain stuck in the tip of a side delivery sheath. For this reason, the tip should always be checked following the transfer, and flushed under the scope to observe if this has occurred. It is the author’s feeling that this problem can be minimized by using a ‘push’ bubble in the straw and by insuring the tip of the sheath is free inside the lumen of the uterus (i.e., not pushed into the uterine wall so as to occlude the openings). If the embryo is too large for a 0.5 mL straw, an individually sterilized insemination pipette may be used with the pipette being loaded in the same manner as the straw. Regardless of the system being used,
b
IMV, 11725 95th Avenue North, Maple Grove, MN 55369.
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Figure 303.7 Load the straw so that the embryo is in the middle fluid bubble.
the transfer gun or pipette should be enclosed within a sterile plastic sleeve so as to protect the sterility of the transfer apparatus as it is passed through the vestibule and vagina. Once the mare has been prepared and sedated, the embryo transfer gun or pipette is guided to the cervix. At this point, the external os may be encircled by the practitioner’s thumb and fingers so as to provide stabilization. The gun is then advanced (free from the plastic sleeve) through the cervix. Once this is accomplished the practitioner transfers their palpation arm to the rectum. The tip of the gun or pipette is then guided through the uterine body. When the tip is located in a position that is free within the lumen of the uterine horn, near the bifurcation, an assistant can slowly push the plunger of the apparatus and expel the embryo into the uterus. The procedure is finished by extracting the embryo transfer gun or pipette from the mare. This entire process should only take 1–2 minutes and be as atraumatic as possible to the cervix and uterus. Any accidental passage of bacteria from the vagina to the uterus may easily cause an endometritis that is very difficult to overcome. Recently, another method of non-surgical transfer has been described.12 For the interested reader, this offers other perspectives.
Management of the recipient mare post-transfer Management of recipient mares post-transfer and the possible therapeutic options are, without a doubt, the most controversial aspects of embryo transfer. Fortunately, this is also the area in which the most research has been done. It had originally been thought that the transcervical transfer of embryos would have less success
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because the cervical manipulation would result in the release of prostaglandin F2 alpha (PGF2α ) resulting in premature luteolysis. More recent studies13, 14 have examined the effect of cervical dilation on Day 7 recipient mares using a balloon catheter. This procedure resulted in the release of oxytocin but not PGF2α . It was also shown that oxytocin administered on Day 7 post-ovulation did not affect progesterone secretion, pregnancy rate, or embryonic growth. Further manipulation of the cervix itself was unlikely to contribute to varying success rates of transcervical transfer of embryos. These investigators concluded that other factors such as window of synchrony, endometrial irritation, and bacterial contamination that could result in luteolysis were the more likely contributing factors. The question regarding possible luteal compromise following transcervical transfers has conflicting research data. Stout15 concluded that embryo transfer-induced luteolysis was not a significant cause of failed embryo transfer because luteal function did not differ between pregnant and non-pregnant mares until the onset of cyclical luteolysis. On the contrary, Foss,16 in a retrospective study involving a large number of mares, examined circulating progesterone levels at the time of transfer and then again 7 days later. His data indicated that lower levels of circulating progesterone are likely, even in pregnant recipient mares, especially after the transfer of morula-stage embryos. In this study, the window of synchrony had no effect on progesterone levels. Most investigators agree that the transcervical transfer of embryos carries with it inherent risk of bacterial contaminants and resultant endometritis. This endometritis may be the more common cause of luteolysis and lower progesterone levels. Considering this, it could be agreed that putting all recipient mares on supplemental progesterone at the time of transfer may be contraindicated in that it may exacerbate an endometritis. There is still much to be learned about maternal recognition of pregnancy in the mare. It may be that a failure or delay in this very important aspect of early pregnancy could account for some of the embryonic loss incurred post-transfer. Possible therapeutic options in this area have yet to be discovered. However, this concept could support the use of supplemental progesterone at this time. Arruda17 has determined that the administration of human gonadotropin (hCG) (3000 IU intramuscularly) on the day of transfer resulted in greater serum circulating progesterone concentrations and an increase in pregnancy rates when compared to other treatment groups and controls. It was thought that the hCG had a luteotropic effect, promoting an increase in the number of large luteal cells and a resultant
increase in the secreted progesterone by the corpus luteum. The following protocol is currently being used by the author. At the time of transfer, routine medication includes flunixin meglamine (500 mg, intravenously), hCG (3000 IU, intramuscularly), and appropriate antibiotic therapy. If no particular antibiotic is indicated, then therapy is initiated with sulfamethoxazole and trimethoprim (24 mg/kg p.o., twice daily). The antibiotic therapy is continued until the first pregnancy examination which is usually 4 days later (i.e., 11 days post-ovulation for the embryo). At this time, a decision is made regarding any continued therapy based on the examination. A pregnant recipient mare is monitored very frequently. The mare is normally examined at 11, 13, 17, 23, and 29 days postovulation, and more often if problems with the pregnancy are noted. With each examination, the uterus is evaluated for signs of endometritis, such as mild edema, small amounts of fluid, or thickening of the uterine wall. In addition, the corpus luteum is also monitored carefully along with any signs suggestive of low circulating progesterone such as decreasing uterine tone or softening of the cervix. If indicated at any time, more appropriate antibiotic therapy is initiated. Treatment options may also be based on the donor mare’s uterine culture post-transfer. Progesterone supplementation is initiated with any sign of low circulating progesterone concentration. If there is any doubt, a serum progesterone is submitted for analysis. It is the author’s opinion that pregnancies that would otherwise be lost can be rescued with supportive therapy. It has been our experience that most pregnancies that are maintained to 60–70 days will be maintained to term. Usually, but not always, supplemental progesterone can be stopped about this time. An important concept is to treat each pregnant recipient as an individual rather than with a herd-based approach.
Donor mare management after the flush The veterinarian may not always have control of the donor mare after the uterus is flushed. However, there are several potential problems that can arise, most of which can be avoided with the proper precautions. The first problem, and potentially greatest liability, is for the donor mare to become pregnant. This may go undiscovered and become a serious problem later, as in the case of a performance mare that is 10 months pregnant at the time of a competitive event. Most veterinarians are in the habit of administering PGF2α immediately after the flush, but this may not always have the desired effect. It is
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Embryo Transfer possible that the mare may have had a second delayed ovulation resulting in an embryo still in the oviduct at the time of the uterine flush. The second ovulation could also result in a corpus luteum that is refractory to PGF2α at the time of the uterine flush. Another way that a donor mare could be pregnant post-flush is simply an inadequate uterine flush technique; the failure to completely fill the uterus with fluid during the uterine flush procedure can easily leave an embryo in the uterus. In recent years there has been a tendency by some veterinarians to flush mares on Day 8 or 9 so that the embryo may be visualized in the filter, enabling the flush to be stopped after just one flush rather than the three that are recommended. This practice can easily leave a twin embryo in the uterus. The second most common mistake is to leave some flush medium in the uterus at the end of the flush. This greatly predisposes the mare to endometritis. It is not always possible, but in an ideal situation, the donor mare should be examined the day following her uterine flush to check for any signs of endometritis. Any mare with even a small amount of uterine fluid should be subjected to uterine lavage. The mare should then be examined again in a few days to determine that she is in estrus. A uterine culture should be performed at the appropriate time on all donor mares after the uterine flush. In summary, embryo transfer can be a tremendously rewarding part of equine practice. If the veterinarian strives for excellence, success will follow.
References 1. McCue PM, Squires EL, Bruemmer JE, Niewender KD. Equine embryo transfer: techniques, trends, and anecdotes. In: Proceedings of the Annual Conference for Theriogenology, 2001, pp. 229–35. 2. Vanderwall D. Current equine embryo transfer techniques. In: Ball BA (ed.) Recent Advances in Equine Reproduction. Ithaca, NY: International Veterinary Information Services, 2000.http://www.ivis.org/advances/ReproductionC ¸ Ball/ embryoC ¸ transfer) 3. McKinnon AO, Squires EL. Morphologic assessment of the equine embryo. J Am Vet Med Assoc 1988;192:401–6.
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4. McKinnon AO, Squires EL. Equine embryo transfer. Vet Clin North Am Equine Pract 1988;4:305–33. 5. Baucus KL, Ralston SL, Nockels CF, McKinnon AO, Squires EL. Effects of transportation on early embryonic death in mares. J Animal Sci 1990;68:345–51. ¨ 6. Wilsher S, Kolling M, Allen WR. The use of meclofenamic acid to extend donor-recipient asynchrony in equine embryo transfer. In: Havemeyer Foundation Monograph Series No. 18, 2005; pp. 56–7. 7. Carnevale EM, Beisner AE, McCue PM, Bass LD, Squires EL. Uterine changes associated with repeated inseminations and embryo collections in mares. In: Proceedings of the 51st Annual Convention of the American Association of Equine Practitioners, 2005. 8. Battut I, Grandchamp des Raux A, Nicais JL, Fieni F, Tainturier D, Bruyas JF. When do equine embryos enter the uterine cavity. In: Havemeyer Foundation Monograph Series No. 3, 2000; pp. 66–8. 9. Hudson JJ, McCue P. How to increase embryo recovery rates and transfer success. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2004. 10. Bielanski A. Disinfection procedures for controlling microorganisms in the semen and embryos of humans and farm animals. Theriogenology 2007;68:1–22. 11. Jasko DJ. Comparison of pregnancy rates following nonsurgical transfer of day 8 equine embryos using various transfer devices. Theriogenology 2002;58:713–15. 12. Wilsher S, Allen WR. An improved method for nonsurgical embryo transfer in the mare. Equine Vet Educ 2004;16(1): 39–44. 13. Handler J, Hoffmann D, Weber F, Schams D, Aurich C. Oxytocin does not contribute to the effects of cervical dilation on progesterone secretion and embryonic development in mares. Theriogenology 2006:66:1397–404. 14. Handler J. Cervical dilatation: a method for studying embryo-maternal interaction in mares. In: Havemeyer Foundation Monograph Series No. 16, 2005; pp. 5–7. 15. Stout TA, Tremoleda JL, Knaap J, Daels PF, Colenbrander B. Does compromised luteal function contribute to failure to establish pregnancy after non-surgical embryo transfer. In: Havemeyer Foundation Monograph Series No. 14, 2005; pp. 8–9. 16. Foss R, Crane A. Serum progesterone changes in embryo transfer recipients. In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, 2004. 17. Arruda RP, Fleury PDC. Pregnancy rates and plasma progesterone concentrations in embryo recipient mares receiving hormone treatment. In: Havemeyer Foundation Monograph Series No. 14, 2005; pp. 95–6.
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Chapter 304
Cooled Transported Embryo Technology Patrick M. McCue, Catherine A. DeLuca and Jillian J. Wall
Introduction The ability to transport embryos from the site of collection to a referral center for transfer into a recipient mare has had a tremendous impact on the equine embryo transfer industry. The major advantages of embryo transport are:
r r r r r
donor mares can be maintained and managed at home; transport is only necessary if an embryo is recovered; cost of transport of an embryo is significantly less than transport of a mare; housing recipient mares at specialized centers eliminates the need to maintain a recipient herd at every breeding farm; recipients are often best managed by experienced personnel; embryo transfer services are more widely available to mare owners; and pregnancy rates are usually better at dedicated embryo transfer facilities with more experienced personnel.
A decrease in ambient temperature during short-term storage results in a slowing of embryo metabolism.1 Consequently, equine embryos do not advance in developmental stage or increase in diameter during storage at 5◦ C.1, 2 In one study, embryos increased in diameter by ∼ 35 µm during cooled storage and in another study embryos decreased in diameter by ∼ 10 µm during cooled storage. The inhibition of embryo metabolism that occurs during cooling is reversible, as embryos increase in diameter when the incubation temperature is increased to 37◦ C.1 Clinically, cooling provides a short period (i.e., < 24 hours) for in vitro storage during which time an embryo may be transported from the site of collection to the site of transfer into a recipient.
Transport of an equine embryo in a commercial holding medium inside a passive cooling system is certainly significantly easier than the technique first used for transported equine embryos.3 In that remarkable study, six embryos were recovered surgically from donor mares and surgically placed into ligated rabbit oviducts in Cambridge, England. The rabbits were transported approximately 33 hours by car to Krakow, Poland. All six embryos were retrieved from the excised rabbit oviducts after a total incubation period of 40 to 49 hours. A single embryo was transferred surgically into each of three recipient mares and non-surgically into one recipient mare. Three of the four recipient mares (two surgical and one non-surgical transfer) became pregnant. A great deal of research related to short-term embryo storage has been performed over the subsequent three decades. The goal of this chapter is to review the history behind the current techniques for cooledtransport of equine embryos, discuss currently recommended techniques for packaging and shipping embryos, and provide data on the application of a cooled-transported embryo program in a clinical equine practice.
Historical perspective Embryo culture or holding medium Early attempts at short-term storage (i.e., up to 24 hours) of equine embryos in Dulbecco’s phosphate buffered saline (DPBS), a traditional flush medium, were not very successful. Imel4 cultured 15 embryos in Dulbecco’s phosphate buffered saline (PBS) plus 20% heat-inactivated steer serum at 37◦ C for 24 hours and cultured an additional 14 embryos in the same medium for ≤ 3 hours. Pregnancy rates following
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Cooled Transported Embryo Technology non-surgical transfer into recipient mares were 6.6% (1 of 15) and 21.3% (3 of 14), respectively. A subsequent study was performed in which embryos were stored in the same medium, but transferred via a ventral midline surgical technique. Pregnancy rate for embryos stored at 37◦ C for 24 hours was 20% (3 of 15) versus 47.1% (8 of 17) for embryos transferred within 3 hours after collection.5 Douglas6 reported that maintenance of equine embryos at room temperature for 6 to 24 hours in DPBS, with antibiotics (100 IU penicillin and 100 µg streptomycin/mL) and 20% fetal calf serum (FCS), resulted in lower pregnancy rates at 24 days (3 of 7; 42.8%) as compared to transfer of embryos within 3 hours of collection (20 of 28; 71.4%). As a consequence of these early studies, embryos were generally transferred within 3 hours after recovery.7 A series of experiments were performed in the early 1980s at Colorado State University to identify an alternative medium to DPBS that would allow for successful storage of equine embryos. Ham’s F-10 medium plus 20% fetal calf serum (FCS) supported an 85% increase in embryo diameter for equine embryos cultured in vitro for up to 24 hours.4 Subsequently, Squires and co-workers5 reported 50-day pregnancy rates of 45.5% (5 of 11 embryos) and 44.4% (4 of 9 embryos) for embryos transferred surgically after culture for ≤ 3 hours in Ham’s F-10 plus 20% steer serum or Dulbecco’s PBS plus 20% steer serum, respectively. Slade and others8 compared the storage of bisected equine embryos in DPBS plus 10% FCS in air versus Ham’s F-10 plus 10% FCS in 5% CO2 at 37◦ C for 24 hours. It was reported that Ham’s F-10 was superior to DPBS in maintaining embryo quality and promoting growth and development of the demi-embryos. Storage temperature Clark and co-workers9 compared viability of embryos > 200 µm cultured in Ham’s F-10 plus 10% FCS under 5% CO2 , 5% O2 , and 90% N2 at 24◦ C, or minimum essential medium (MEM) with Hank’s balanced salts plus 10% FCS in air at 24◦ C or at 5◦ C for 12 and 24 hours. Embryos ≤ 200 µm were bisected and demi-embryos were stored in Ham’s F-10, MEM or DPBS plus 10% FCS in air at 24◦ C for 24 hours. In general, embryos stored in Ham’s F-10 maintained better quality scores than embryos stored in MEM or DPBS. In a subsequent experiment, embryos cultured in Ham’s F-10 at 5◦ or 24◦ C did not develop or increase in diameter during a 12-hour culture period, but advanced in development over the subsequent 12-hour period when the incubation temperature was increased to 37◦ C.1 Pregnancy rates at day 12 after
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non-surgical transfer were similar for embryos stored in Ham’s F-10 at 24◦ C for < 1 hour (7 of 16; 43.8%) and embryos transferred after 12 hours of culture (8 of 16, 50%). It was concluded from these experiments that Ham’s F-10 was the medium of choice for maintaining equine embryos in culture for 24 hours. A study was subsequently performed to determine if addition of Hepes buffer to Ham’s F-10 plus 10% FCS would eliminate the requirement for bubbling a mixture of 5% CO2 , 5% O2 , and 90% N2 through the medium to stabilize the pH.2 Embryos stored at 5◦ C for 24 hours in gassed (5% CO2 , 5% O2 , and 90% N2 ) Ham’s F-10 maintained a better quality score and exhibited a greater average increase in diameter than embryos stored in Ham’s F-10 plus Hepes buffer. Surgical transfer of embryos resulted in a higher pregnancy rate at 14 days for Ham’s F-10 plus CO2 (14 of 20; 70%) than Ham’s F-10 plus Hepes buffer (4 of 20; 20%). Martin and colleagues10 compared embryo quality and transfer success rate after storage in Ham’s F-10 at 5◦ C for either 18 or 36 hours. Quality score decreased with storage time, but non-surgical pregnancy rates were similar between embryos cooled for 18 hours (8 of 22; 36.4%) or 36 hours (8 of 20; 40%). Two studies published during this time period evaluated the efficacy of Dulbecco’s PBS for cooled storage. Pashen11 stored embryos in a modified DPBS plus 10% heat-inactivated FCS at 4◦ C and evaluated embryo quality at 24, 48, 72, and 84 hours. All embryos survived cooled storage for 48 hours. Nonsurgical transfer of seven embryos resulted in four pregnancies (57.1%). In a similar study, Sertich et al.12 stored 16 embryos in DPBS plus 1% newborn calf serum and 0.4% bovine serum albumin in a passive R , Hamilton Thorne Recooling device (Equitainer search, South Hamilton, MA) at 4 to 5◦ C for 24 hours. Pregnancy rate at day 14 after non-surgical transfer into synchronized recipient mares was 37.5% (6 of 16) and three of the six pregnancies were carried to term. Era of commercial embryo transfer using Ham’s F-10 Large-scale commercial equine embryo transfer programs incorporated the use of Ham’s F-10 for cooled transport of embryos starting in the late 1980s. The medium was prepared as needed by bubbling the triple gas mixture (5% CO2 , 5% O2 , and 90% N2 ) through the medium for 3 to 5 minutes. The gassed medium was supplemented with 10% FCS and penicillin/streptomycin antibiotics and subsequently passed through a 0.2 µm filter prior to addition of an embryo and placement into a container for cooled-storage and transport.
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The first report of large-scale commercial transfer of cooled-transported equine embryos was by Cook and co-workers.13 Embryos were flushed at multiple locations around the United States, placed in Ham’s F-10 medium prepared as described above, cooled in a passive storage device, and shipped by counter-tocounter airline service to the embryo transfer station. Day 15 pregnancy rate after surgical transfer was not significantly different between embryos transferred immediately after collection at the ET station (20 of 26; 76.9%) and embryos collected at a distant location, cooled and transported to the ET station over a 5- to 24-hour period (20 of 24; 83.3%). Carney et al.14 reported that pregnancy rates for fresh (77 of 104; 74%) and cooled embryos (109 of 136; 80%) were similar at days 12, 35, and 50 after surgical transfer. The authors noted that embryonic loss rates were higher for embryos that had been cooled for > 12 hours (25%) than embryos transferred within 12 hours of collection (10%). Squires et al.15 reported that pregnancy rates following surgical transfer of grade 1 embryos were similar between fresh (48 of 54; 89%) and cooledtransported embryos (61 of 71; 86%). Embryonic loss rates between days 12 and 50 were noted to be similar for fresh and cooled-transported embryos. The first reports of wide-scale application of non-surgical embryo transfer in clinical practice involved fresh embryos in Argentina.16, 17 In the United States, a majority of embryos were transferred surgically prior to the work of Foss and colleagues in Missouri.18 Pregnancy rates for grades 1 and 2 embryos (combined) of 86.5% and 86.3% were reported following non-surgical transfer of 89 fresh and 22 transported embryos, respectively. Transported embryos were maintained in Ham’s F-10 as described previously and transferred within 18 hours of initial recovery.
equine embryos that had been maintained in a pasR ) at 5◦ C for 24 hours sive cooling device (Equitainer after collection. Moussa and colleagues20 subsequently compared the viability of embryos after 24 hours of storage at 5◦ C in Ham’s F-10, EmCare Holding SolutionTM or Vigro Holding PlusTM (Bioniche Animal Health, Inc., Belleville, Ontario, Canada). The number of DAPIstained (dead) cells per embryo was similar between the three groups after 24 hours of storage. It was noted that larger day 7 embryos appeared to withstand cold storage for 24 hours better than smaller day 7 embryos. In a second experiment, 9 of 20 embryos stored in Ham’s F-10 (45%) and 8 of 20 embryos stored in EmCare Holding SolutionTM (40%) resulted in pregnancies after surgical transfer. A further study confirmed that the percentage of dead cells per embryo increased with duration of cooled storage21 and that all three media were equally suited for storage of equine embryos at 5◦ C. Hudson and co-workers22 reported that small (< 300 µm) equine embryos could be cooled and held at 5 to 8◦ C for 12 to 19 hours prior to vitrification. Pregnancy rate following non-surgical transfer of embryos vitrified immediately after collection (15 of 20; 75%) was not significantly different from the pregnancy rate for embryos cooled prior to vitrification (13/20; 65%). Results of this study suggest that embryos could be collected at one location, cooled and subsequently transported to a referral center for cryopreservation. Currently, a majority of equine embryos in the United States are transported in a commercial ‘complete’ holding medium by either counter-to-counter (same-day) delivery or overnight courier service.
Era of the ‘complete’ holding medium
Carnevale et al.2 noted that the initial ultrasonographic appearance of the embryonic vesicle in the uterus after transfer of a cooled embryo was often delayed and the size of the vesicle was often smaller. It was suggested that cold storage may have resulted in temporary retardation of growth. It was proposed that cooling equine embryos may damage the inner cell mass more than trophoblast cells, resulting in an increased incidence of development of the trophoblast with an embryo proper.2 This was not observed in a subsequent larger study at the same location.14 Conflicting reports exist as to the effect of embryo cooling on pregnancy loss. Cook et al.13 reported a pregnancy loss between days 15 and 50 of 11.5% (3 of 26 pregnancies) for fresh embryos and 8.2% (2 of 24 pregnancies) for transported embryos. Carney
The main problem with Ham’s F-10 in clinical practice is the buffering system. Ham’s F-10 contains a bicarbonate-based buffer and is dependent on CO2 to stabilize the pH. In addition, once prepared (i.e., gassed with 5% CO2 , 5% O2 , and 90% N2 ), the pH of the medium is only stabilized for approximately 72 hours. In an attempt to eliminate the need for gassing Ham’s F-10, ‘complete embryo holding media’ that incorporated zwitterionic buffers, nutrients, growth factors, amino acids, BSA or other surfactant agents and antibiotics were being tested. The first study evaluated the potential of EmCare Holding SolutionTM (ICPBio; Auckland, New Zealand) for cooled storage of equine embryos.19 A 50% pregnancy rate was noted following non-surgical transfer of six
Adverse effects of cooling equine embryos
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et al.14 noted that initial (day 12) pregnancy rates were similar following surgical transfer of fresh and cooled embryos and that overall, embryonic loss rate was not different between fresh and cooled embryo groups. However, for mares receiving cooled embryos, the embryonic loss between day 12 and 35 was higher for mares receiving embryos cooled for > 12 hours (25%) as compared to mares receiving embryos that had been cooled for < 12 hours (10%). In a more recent retrospective study of embryo transfer results, the incidence of embryonic loss was not different between embryos transferred immediately after collection and cooled-transported embryos.23
Small vs large cooled equine embryos Several investigators have noted that large equine embryos tolerate cooling better than smaller embryos.14, 20, 21, 24 Cooling was associated with a greater percentage of dead cells in smaller embryos than larger embryos.2 Carney et al.14 reported that transfer of smaller embryos resulted in fewer pregnancies than transfer of larger embryos, when data from fresh and cooled embryos were combined. Transfer of 22 cooled embryos ≤ 250 µm in diameter resulted in a 59% pregnancy rate, while transfer of 105 embryos larger than 250 µm resulted in a pregnancy rate of ≥ 80%. Moussa and colleagues20, 21 noted that embryos > 400 µm may have a better viability than embryos < 400 µm after 24 hours of cooled storage, based on the average number of dead cells/mm2 for each embryo size category.
Current technique for cooled-transport of equine embryos
Figure 304.1 A 50 mL conical tube, 5 mL vial, and Parafilm for cooled-storage of equine embryos.
4.
5. 6.
7. 8.
medium) into the plastic tube. Rinse out the transfer straw and observe rinse medium under microscope to ensure that the embryo was properly transferred. Add more medium until the 5 mL tube is full. Fix the cap securely in place. Seal the tube with Parafilm sealing film. Fill the 50 mL centrifuge tube with the saved flush medium. Place the 5 mL tube containing the embryo into the 50 mL centrifuge tube. Attach the cap of the centrifuge tube securely in place and seal with Parafilm (Figure 304.2). Load the 50 mL tube containing the embryo into the passive cooling device (Figure 304.3). Insert appropriate documentation, including a description of the embryo (i.e. developmental stage, grade and size).
Materials required
r r r r r r
Commercial embryo holding medium (5–6 mls) Commercial flush media (50 mls) 5 mL sterile plastic tube with screw or snap cap (2. 1-1) 50 mL sterile centrifuge tube with screw cap Parafilm sealing film R Passive cooling device (e.g., Equitainer )
Directions for packaging an embryo for shipment 1. Allow the embryo holding medium to warm to 37◦ C. 2. Fill the 5 mL plastic tube with approximately 4.5 mL of warmed holding medium. 3. Carefully transfer the embryo (after washing through a minimum of 3 to 4 drops of holding
Figure 304.2 Application of Parafilm to seal the 50 mL conical tube.
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ART: Embryos Table 304.1 Quality score for equine embryos immediately prior to transfer (number of embryos and percentage for each grade). Embryos were either transferred immediately after collection (fresh) or after having been cooled for approximately 12 hours (counter-to-counter transport) or 24 hours (overnight courier transport).
Figure 304.3 A 50 mL conical tube containing an embryo within a commercial passive cooling container.
9. Ship the container to the embryo transfer referral center by counter-to-counter airline delivery (preferred) or overnight courier service. Directions for receiving and handling a shipped embryo 1. Open the container in a clean laboratory space and review documentation. 2. Have embryo holding medium available to wash embryo or rinse plastic tube. 3. Open centrifuge tube and remove 5 mL tube containing embryo. 4. Gently invert the 5 mL tube several times. 5. Remove cap from 5 mL tube, place cap with open end up on counter top and add several drops of holding medium into cap.
(a)
Embryo storage
Grade 1
Grade 2
Grades 3 & 4
Fresh Counter-to-counter Overnight courier
303 (81.2%) 159 (68.8%) 8 (44.4%)
61 (16.4%) 57 (24.7%) 7 (38.9%)
9 (2.4%) 15 (6.5%) 3 (16.7%)
6. Pour contents of 5 mL tube into a sterile 60 mm Petri dish. 7. Add several drops of holding medium into empty 5 mL tube and set aside. 8. Search Petri dish for embryo. 9. If the embryo is not immediately visible, rinse the cap and the 5 mL tube several times with holding medium. Embryos occasionally will stick to the plastic tube or cap. 10. Wash the embryo through a minimum of 3 to 4 drops of holding medium prior to transfer (Figure 304.4). Ideally, the medium used to wash the embryo should be the same as the medium used for cooled storage.
Pregnancy rates in a commercial embryo transfer program Quality scores or grade for 622 fresh and cooled equine embryos evaluated immediately prior to transfer over a 5-year period (2004 to 2008) at Colorado State University are presented in Table 304.1.
(b)
Figure 304.4 (a) Fresh day 7.5 expanded blastocyst prior to transfer. (b) Cooled day 8 expanded blastocyst exhibiting mild shrinkage or dehydration. The capsule is visible surrounding the embryo.
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Table 304.2 Pregnancy rates (day 16) for embryos transferred immediately after collection (fresh) or cooled-transported by counter -to-counter airline service or overnight courier. Embryo storage
Grade 1
Grade 2
Grades 3 & 4
Total
Fresh Counter-to-counter Overnight courier Total
303 (79.5%) 159 (77.4%) 8 (75.0%) 470 (78.7%)a
61 (68.9%) 57 (61.4%) 7 (57.1%) 125 (64.8%)b
9 (66.7%) 15 (46.7%) 3 (33.3%) 27 (51.9%)b
373 (77.5%) 231 (71.4%) 18 (61.1%) 622 (74.8%)
a,b
values within row with different superscripts are significantly different (p < 0.05).
It can be noted that the highest percentage of grade 1 embryos (81.2%) occurred with fresh embryos. Cooled storage for approximately 12 to 24 hours, for counter-to-counter or overnight courier service, respectively, was associated with lower quality scores. Pregnancy rates at day 16 for 622 embryos transferred either immediately after collection (fresh), after same-day (counter-to-counter) cooled transport, or after overnight courier transport are presented in Table 304.2. There was no difference in day 16 pregnancy rate between grade 1 embryos that were transferred immediately after collection (79.5%), or after cooled-transport (77.4% and 75.0% for same-day delivery and overnight delivery, respectively. A decrease in pregnancy rate was observed within each group as embryo grade declined. There was no significant difference in pregnancy loss rate between days 16 and 50 for embryos transferred immediately after collection (13.1%) and cooled embryos transferred after same-day or overnight delivery (9.5% and 0.0%, respectively). Nine cases of empty trophoblastic vesicle (ETV) formation were observed for 316 embryo transfer pregnancies during the 5-year observation period. A total of seven ETV cases were noted for 169 fresh embryo transfer pregnancies (4.1%) and two ETV cases occurred for 147 pregnancies (1.4%) following transfer of cooled embryos. Quality scores for the embryos that resulted in ETV formation were as follows: six grade 1.0 or 1.5, two grade 2.0 or 2.5, and one grade 3 embryo. Morphology for embryos resulting in ETVs were: two morula, two early blastocysts, two blastocysts, and three expanded blastocysts. Retrospective evaluation of photographs available for eight of the nine embryos taken prior to transfer did not reveal any consistent or predictive abnormalities. A presumptive inner cell mass was visible on four of the five blastocyst or expanded blastocyst stage embryos that resulted in ETV formation. Ultrasound examination of the recipient mares indicated that five of the nine embryos were not visible or smaller than expected on pregnancy examination performed at em-
bryo day 11 or 12 (4 or 5 days after transfer of a day 7 embryo, respectively).
Summary Cooled-transport of equine embryos is now a routine procedure in the equine industry. Progress in development of commercial complete holding media and passive cooling systems have simplified the process. Pregnancy rates following transfer of cooledtransported embryos are similar to rates following transfer of fresh embryos.
References 1. Clark KE, Squires EL, McKinnon AO, Seidel GE Jr. Viability of stored equine embryos. J Anim Sci 1987;65:534– 42. 2. Carnevale EM, Squires EL, McKinnon AO. Comparison of Ham’s F-10 with CO2 or Hepes buffer for storage of equine embryos at 5◦ C for 24 h. J Anim Sci 1987;65:1775–81. 3. Allen WR, Stewart F, Trounson AO, Tischner M, Bielanski W. Viability of horse embryos after storage and longdistance transport in the rabbit. J Reprod Fertil 1976;47: 387–90. 4. Imel KJ, Squires EL, Elsden RP, Shideler RK. Collection and transfer of equine embryos. J Am Vet Med Assoc 1981;179: 987–91. 5. Squires EL, Imel KJ, Iuliano MF, Shideler RK. Factors affecting reproductive efficiency in an equine embryo transfer program. J Reprod Fertil 1982;32(Suppl):409–14. 6. Douglas RH. Some aspects of equine embryo transfer. J Reprod Fertil 1982;32(Suppl):405–8. 7. Squires EL, Cook VM, Voss JL. Collection and transfer of equine embryos. Colorado State University Animal Reproduction Laboratory Bulletin No. 01. 1985; pp. 1–38. 8. Slade NP, Williams TJ, Squires EL, Seidel GE Jr. Production of identical twin pregnancies by microsurgical bisection of equine embryos. In: 10th International Congress on Animal Reproduction and Artificial Insemination, 1984;11:241. 9. Clark KE, Squires EL, Takeda T, Seidel GE Jr. Effect of culture media on viability of equine embryos in vitro. Equine Vet J 1985;Suppl 3:35. 10. Martin JM, Squires EL, Jasko DJ, Carney NJ. Effect of storage of equine embryos at 5◦ C for 18 and 36 hours. Theriogenology 1991;35:238 (abst). 11. Pashen RL. Short-term storage and survival of horse embryos after refrigeration at 4◦ C. J Reprod Fertil 1987; 35(Suppl):697–8.
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12. Sertich PL, Love LB, Hodgson RM, Kenney RM. 24-hour cooled storage of equine embryos. Theriogenology 1988;30: 947–52. 13. Cook VM, Squires EL, McKinnon AO, Bailey J, Long PL. Pregnancy rates of cooled transported equine embryos. Equine Vet J 1989:8(Suppl):80–1. 14. Carney NJ, Squires EL, Cook VM, Seidel GE Jr, Jasko DJ. Comparison of pregnancy rates from transfer of fresh versus cooled, transported equine embryos. Theriogenology 1991;36:23–32. 15. Squires EL, Cook VM, Jasko DJ, Tarr SF. Pregnancy rates after collection and shipment of equine embryos (1988–1991). Proc Am Assoc Equine Pract 1992;38:609–617. 16. Pashen RL, Lascombes FA, Darrow MD. The application of embryo transfer to polo ponies in Argentina. Embryo Transfer 3. Equine Vet J 1993;Suppl 15:119–21. 17. Riera FL, McDonough J. Commercial embryo transfer in polo ponies in Argentina. Equine Vet J 1993;(Suppl):116. 18. Foss R, Wirth N, Schiltz P, Jones J. Nonsurgical embryo transfer in a private practice (1998). Proc Amer Assoc Equine Pract 1999;45:210–12. 19. McCue PM, Scoggin CF, Meira C, Squires EL. Pregnancy rates for equine embryos cooled for 24 hours in Ham’s F-10
20.
21.
22.
23.
24.
vs EmcareTM embryo holding solution. Proceedings Society for Theriogenology 2000:147 (abst). Moussa M, Duchamp G, Mahla R, Bruyas J-F, Daels PF. In vitro and in vivo comparison of Ham’s F-10, Emcare holding solution and ViGro Holding Plus for the cooled storage of equine embryos. Theriogenology 2003;59:1615–25. Moussa M, Tremoleda JL, Duchamp G, Bruyas J-F, Colenbrander B, Bevers MM, Daels PF. Evaluation of viability and apoptosis in horse embryos stored under different conditions at 5◦ C. Theriogenology 2004;61:921–32. Hudson J, McCue PM, Carnevale EM, Welsh S, Squires EL. The effects of cooling and vitrification of embryos from mares treated with equine follicle-stimulating hormone on pregnancy rates after nonsurgical transfer. J Equine Vet Sci 2006;26:51–4. Carnevale EM, Ramirez RJ, Squires EL, Alvarenga MA, Vanderwall DK, McCue PM. Factors affecting pregnancy rates and early embryonic death after equine embryo transfer. Theriogenology 2000;54:965–79. Moussa M, Duchamp G, Daels PF, Bruyas J-F. Effect of embryo age on the viability of equine embryos after cooled storage using two transport systems. J Equine Vet Sci 2006; 26:529–34.
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Chapter 305
Freezing of Embryos Jean-Franc¸ois Bruyas
Embryo cryopreservation allows a dissociation of both the time and place of the embryo collection procedure and transfer. In addition, management of recipient mares may be simplified as there is no need to have one or two recipient mares with their estrous cycles perfectly synchronized with the donor mare for every flush. International trade in frozen embryos could be extended similar to frozen semen in different mammalian species and for frozen bovine embryos. In cattle for the year 2006, more than 420 000 frozen embryos were transferred around the world;1 57.5% of the 95 000 transfers in Europe were performed with frozen embryos,2 and more than 34 000 frozen embryos were exported from Canada and the USA in 2006.1 Embryos from other mammalian species are also routinely cryopreserved: 44% of 43 000 ovine embryos, and 67% of 24 000 caprine embryos transferred around the world in 2006,1 and hundreds of thousands of rodent embryos,3 but the freezing of equine embryos is still not a routine procedure.
History Slow or ‘conventional’ freezing The first births from frozen mammalian embryos were reported in mice in 1972 by Whittingham et al.,4 then in cattle in 1973 by Wilmut and Rowson.5 In horses, the first pregnancy from a frozen embryo was obtained in 1981,6 but aborted before the term, and the first birth was reported on 31 May 1982 by Yamamoto et al.7 From this date, fewer than 70 papers were published (mean < 3 per year) about equine embryo cryopreservation. In most of these publications (see Tables 305.1–305.3), ‘conventional’ methods of deepfreezing, efficacious in other species, were used: 1. Glycerol, the cryoprotectant (CP) most often used, was incorporated at room temperature in one to
2. 3. 4.
5. 6.
five steps, each of 5 to 20 min, of increasing concentration to a final concentration of either 10% v/v (∼ 1.3 M) or 1.5 M. The embryo was then taken up in 50 µL in a 0.25-mL freezing straw. The straw was cooled from room temperature to – 6 or – 7◦ C at approximately 3◦ C/min. After seeding (= induction of ice formation) at – 6 or – 7◦ C the straw was cooled at 0.3◦ to 0.5◦ C/min to – 30, – 33, or – 35◦ C and plunged and stored in liquid nitrogen. For thawing, the straw was put in water at 37◦ C for 30 to 60 seconds and the embryo recovered. The CP was diluted by moving the embryo through successive baths of decreasing concentration from 10% to 0% or from 1.5 M to 0 M, in four to six steps of 5 to 10 min.
Table 305.1 summarizes most of the publications about transfers of frozen-thawed embryos: from 878 (39 in vitro-produced included) frozen-thawed embryos transferred, 244 (28%) early pregnancies were obtained but only 22 births have been reported due to the occurrence of spontaneous abortions (a few), induced abortions, or termination of the pregnancies before term. The mean pregnancy rate was low and successes were obtained primarily with small embryos (diameter ≤ 200–300 µm). Certainly, not all attempts at deep-freezing of horse embryos have been published, especially in case of failures, and a few equine reproduction centers perhaps use this process in practice without publication. For example, Vullers8 claimed that he obtained a pregnancy rate of 70% in freezing of embryos in his commercial program, but without any explanation and any quantitative results. Only Lascombes and Pashen9 published data of a commercial embryo transfer program, reporting one of the best pregnancy rates 24/43 (56%) after transfer of frozen-thawed horse embryos, but only with small embryos collected very early after ovulation.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
D8 D6
D6 EB D6 B D7 B D7 EExB D7 ExB D6 EB D6 B D7 B D7 ExB
0/5 3b (1)/11 0/4 (+ 1 non-transferred) 0/12 (+ 2 non-transferred)
2c (0)/9 1 (1)/7 1 (1)/2 1 (1)/4 (+2 non-transferred) 1 (1)/9 (+2 non-transferred) 0/4 0/2 0/2 1(1)/6
Yamamoto et al., 19827
D6
D6
1(1)/2
1(1)/2
Glycerol three steps 0.44 M 10 min – 0.88 M 10 min – 1.32 M 30 min
Glycerol three steps 0.33 M 10 min – 0.66 M 10 min – 1 M 30 min
(1-mL glass ampoule) Seeding at – 7◦ C From – 7◦ C to – 35◦ C at – 0.3◦ C/min then to – 38◦ C at 0.1◦ C/min
(1-mL glass ampoule) Seeding at – 7◦ C From – 7◦ C to – 26◦ C at – 0.3◦ C/min then to – 38◦ C at 0.1◦ C/min
(Pyrex tube) Seeding at – 5◦ C From – 5◦ C to – 36/– 49◦ C at 0.2◦ C/min
(Pyrex tube) Seeding at – 5◦ C From – 5◦ C to – 70◦ C at 0.2◦ C/min
(Pyrex tube) Seeding at – 5◦ C From – 5◦ C to – 70◦ C at 0.2◦ C/min
(Glass tube) Seeding at – 6/– 7◦ C Cooling rate not indicated
Glycerol six steps 1.1 M – 0.88 M – 0.66 M – 0.44 M – 0.22 M – 0 M 10 min/step
Glycerol three steps
Glycerol two steps 0.5 M 10 min and 0 M
DMSO six steps 1.25 M to 0 M 10 min/step
Glycerol two steps 0.5 M 10 min and 0 M
DMSO six steps 10 min 1.25 M, 1 M, 0.75 M, 0.5 M, 0.25 M, 0 M
Not indicated
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Glycerol two steps 0.5 M 5 min – 1 M 30 min
DMSO three steps 0.5 M 10 min – 1 M 1 min, 1.5 M 30 min
Glycerol two steps 0.5 M 5 min – 1 M 30 min
DMSO six steps 5 min 0.25 M, 0.5 M, 0.75 M, 1 M, 1.25 M, 1.5 M
Not indicated
Dilution of cryoprotectant
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M and EB
Not indicated
(Device to store embryo) Cooling rate
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D8 D6
0/9
Allen, 198260
D5 to D8
1a (0)/4 (+ 15 non-transferred)
Griffin et al., 19816
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Table 305.1 Results of embryo transfer of frozen-thawed equine embryos published from 1981 to 2007 (adapted from Seidel43 and completed)
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0/1 1f (0)/1 1(1)/4 0/1 1g (0)/1 0/1 0/1 0/1 0/1 1h (0)/2 1i /2 0/1 0/1 0/5 3j /10
2k /7 (+ 1 degenerated non-transferred) 0/3 3k /6 (+ 2 degenerated non-transferred) 0/1 (26 other embryos frozen but not thawed)
Czlonkowska et al., 198564 Boyle et al., 198565
Boyle et al., 198565
Wilson et al., 198660
Farinasso et al., 198967
D6 ExB
D6 EB D6 B
D6 M Glycerol two steps 5% 5 min – 10% 10 min
Glycerol one step 10% 30 min
D6 diam. < 250 µm; mean diam. = 190 µm
(Plastic straw) Seeding at – 7◦ C From – 7◦ C to – 32◦ C at 0.3–0.6◦ C/min Deep-freezing in stainless-steel cylinder and liquid nitrogen container
(0.5-mL plastic straw) Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min
(Small glass tube) Seeding at – 6◦ C From – 6◦ C to – 40◦ C at 0.3◦ C/min
(continued)
Glycerol and sucrose four steps 6 min/step Glycerol 6% + sucrose 0.9 M G 3% + sucrose 0.9 M G 0% + sucrose 0.9 M G 0% + sucrose 0 M
Glycerol and sucrose three steps Glycerol 10% 1 min Sucrose 1.08 M 30 min 0%, 0 M 5 min
Glycerol five steps 10%, 7.5%, 5%, and 2.5% 10 min/step 0% 30 min
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Glycerol four steps 2.5%, 5%, 7.5%, and 10% 10 min/step
(0.25-mL glass tube) Seeding at – 6◦ C From – 6◦ C to – 40◦ C at 0.3◦ C/min
Glycerol five steps 10%, 7.5%, 5%, and 2.5% 10 min/step + 0% 20 min
Glycerol six steps
16:40
D7 B D6 EB D7 EB D7 B
Glycerol four steps 2.5%, 5%, 7.5%, and 10% 10 min/step
(0.25-mL glass tube) Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.3◦ C/min
(0.5-mL plastic straw) Seeding at – 6◦ C From – 6◦ C to – 33◦ C at 0.3◦ C/min
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D6.5 B D7 EB D7 B D7 EExB D6 EB D7 M D7 B D7 EExB D7 ExB D8 ExP
Glycerol two steps 5% 10%
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D6 diam. < 160 µm; mean diam. = 147 µm (diam. pregnancy = 160 µm) D7 diam. > 160 µm; mean diam. = 400 µm
1m /9 (+ 1 non-transferred)
0/8 (+ 1 destroyed by freezing non-transferred)
1m /8 (+ 2 non-transferred)
D6 diam. < 160 µm; mean diam. = 147.5 µm (diam. pregnancy = 140 µm) D7 diam. > 160 µm; mean diam. = 378 µm (diam. pregnancy = 200 µm) 1,2-propanediol four steps 2.5%, 5%, 7.5%, and 10% 5 min/step
Glycerol four steps 2.5%, 5%, 7.5%, and 10% 5 min/step
(0.25-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
No seeding directly placed at – 25◦ C for 30 min before plunging in liquid nitrogen
1,2-propanediol seven steps: 10%, 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% 5 min/step
Glycerol seven steps 10%, 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% 5 min/step
Glycerol and sucrose six steps 6 to10 min/step: G 12% + S 10% G 9% + S 10% G 6% + S 10% G 3% + S 10% G 0% + S 10% G 0% + S 0%
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1m /6 (+ 2 non-transferred)
D6.5–D7 diam. > 200 µm; mean diam. = 457 µm (diam. in case of pregnancy = 375 µm)
1l /13
Seeding at –6◦ C From – 6◦ C to – 25◦ C at 0.3◦ C/min
Glycerol and sucrose four steps 6 min/step G 6% + sucrose 10% G 3% + sucrose 10% G 0% + sucrose 10% G 0% + sucrose 0%
Dilution of cryoprotectant
16:40
Gaubert 199093
Glycerol and sucrose two steps 10 min/step Glycerol 10% G 10% + sucrose 15%
D6.5–D7 diam. > 200 µm; mean diam. = 440 µm (diam. in cases of pregnancy) = 207 and 275 µm
2l /12
(0.5-mL plastic straw) Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min and to – 33◦ C at 0.1◦ C/min
(Device to store embryo) Cooling rate
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Glycerol and sucrose two steps 10 min/step Glycerol 15% Glycerol 15% + sucrose 15%
Glycerol two steps 5% 10 min 10% 20 min
D6.5–D7 diam. < 200 µm; mean diam. = 182.5 µm
4l /8
Squires et al., 198968
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References
BLBK232-McKinnon
No. pregnancies (births)/No. transferred embryos
c305
Table 305.1 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
Meira and Alvarenga, 1993
71
3(2) /6
D6
D6 EB
1p /5 q
D6 EB
D6 diam. < 230 µm (150–225 µm)
6◦ /11 (+ 3 degenerated non-transferred)
Glycerol two steps 0.75 M 15 min 1.5 M 10 min
1.37 M 60 min
Ethylene glycol at 5◦ C 1 step Cooled from room temperature to 5◦ C in PBS 1.5 M 60 min
Glycerol four steps 2.5%, 5%, 7.5%, and 10% 10 min/step
Glycerol two steps 5% 10 min 10% 30 min
(0.5-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3–0.6◦ C/min Deep-freezing in stainless-steel cylinder and liquid nitrogen container
(0.25-mL plastic straw) from 5◦ C to – 6◦ C at 1◦ C/min Seeding at – 7◦ C From – 7◦ C to – 35◦ C at 0.3◦ C/min
(0.5-mL plastic straw) From 24◦ C to – 6◦ C at 4◦ C/min Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min and to – 33◦ C at 0.1◦ C/min (0.5-mL plastic straw) From 24◦ C to – 6 C at 1◦ C/min Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.3◦ C/min
(continued)
Glycerol five steps: 1 M, 0.75 M, 0.5 M, 0.25 M, 0 M
Ethylene glycol at 5◦ C 10 steps: 10 min/step 1.35 M, 1.2 M, 1.05 M, 0.9 M, 0.75 M, 0.6 M, 0.45 M, 0.3 M, 0.15 M, and 0 M + 0 M 40 min at 20◦ C
Glycerol five steps 10%, 7.5%, 5%, 2.5%, 0% 10 min/step
Glycerol seven steps 10%, 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% 5 min/step
Trim: 279mm X 216mm
0/5
D6 diam. < 230 µm (175–225 µm)
4 /10 (+ 2 degenerated non-transferred)
Glycerol seven steps 10%, 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% 5 min/step
16:40
Day et al., 199370
Skidmore et al., 1991
D6–D6.75 diam. > 160 µm;
◦
0/22
(0.25-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
February 2, 2011
69
Glycerol four steps 2.5%, 5%, 7.5%, and 10% 5 min/step
P2: SFK
D6–D6.75 diam. < 160 µm; mean diam. in case of pregnancy = 152 µm
BLBK232-McKinnon
8n /26
c305
Lagneaux and Palmer, 199119
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
Freezing of Embryos 2891
D6–7 M and EB D6–7 M and EB
D6 M and EB
D6–7 M and EB
D6–7 M and EB
D7 M and EB
D7 M and EB
D7 M and EB
5r /15 0/15(+ 2 degenerated non-transferred)
6s /15
0/15
0/15
3 (1)/6
3 (1)/6
1 (1)/5
Pfaff, 1994, cited in Seidel, 199643
Glycerol no. steps not indicated last step: 1.5 M 30 min Glycerol no. steps not indicated last step: 1.5 M 15 min Ethylene glycol no. steps not indicated last step: 1.5 M 15 min
Glycerol + sucrose three steps: G 5.5% 10 min G 11% 20 min G 20% + sucrose 0.5 M 20 min
Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.5◦ C/min
(0.5-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3–0.6◦ C/min Deep-freezing in stainless-steel cylinder and liquid nitrogen container 5 min in liquid nitrogen vapor before plunging into liquid nitrogen
Ethylene glycol + 10% sucrose four steps 7 min/step
Glycerol + 10%sucrose four steps 7 min/step
Glycerol + 10% sucrose four steps 7 min/step
Glycerol + sucrose no. steps and time not indicated Decreasing concentrations from G 10% sucrose 2.5 M
Glycerol five steps: 7.3%, 5.5%, 3.7%, 1.8%, 0% 8–10 min/step 1,2-propanediol five steps: 7.3%, 5.5%, 3.7%, 1.8%, 0% 8–10 min/step
Glycerol five steps 8–10 min 7.3, 5.5, 3.7, 1.8, 0% Glycerol five steps 8–10 min 7.3, 5.5, 3.7, 1.8, 0% 1,2-propanediol five steps 8–10 min/step 7.3, 5.5, 3.7, 1.8, 0%
Not indicated
Trim: 279mm X 216mm
1,2-propanediol two steps: 5.5% 10 min – 11% 20 min
Glycerol two steps: 5.5% 10 min – 11% 20 min
(0.5-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3–0.6◦ C/min Deep-freezing in stainless-steel cylinder and liquid nitrogen container
Not indicated
Dilution of cryoprotectant
16:40
1,2-propanediol 2 steps: 5.5% 10 min 11% 20–25 min
Glycerol two steps: 5.5% 10 min – 11% 20–25 min
Not indicated
(Device to store embryo) Cooling rate
February 2, 2011
Landim e Alvarenga et al., 199350
D6–7 B and ExB
0/12
Meira et al., 199372
D6 M and EB
7/19
Cryoprotectant used and addition protocol
2892
Allen et al., 1993, cited in Seidel, 199643
Embryo developmental stage
BLBK232-McKinnon
References
P2: SFK
No. pregnancies (births)/No. transferred embryos
c305
Table 305.1 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
6(1)u /20
3(1)u /20
D6 M, EB, B diam. < 220 µm (150–220 µm) Mean diam. = 154 µm D6 M, EB, B diam. < 220 µm (150–220µm) Mean diam. = 154.5 µm
D6 M, EB
0/7
Glycerol four steps: 2.5%, 5%, 7.5%, and 10% 5 min/step
Glycerol + 1,2-propanediol two steps: G 0.35 M + P 0.35 M 10 min G 0.75 M + P 0.75 M 20 min
(0.25-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
(0.25-mL plastic straw) Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.4◦ C/min
Freezing of Embryos (continued)
Glycerol six steps: 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% 5 min/step Glycerol + sucrose six steps 5 min/step: G 10% + sucrose 0.25 M G 7.5% + sucrose 0.25 M G 5% + sucrose 0.1 M G 2.5% + sucrose 0.1 M G 0% + sucrose 0.1 M G 0% + sucrose 0 M
Glycerol and sucrose three steps 10 min/step: G 0.75 M + S 0.25 M G 0 M + S 0.25 M G 0M + S 0M Glycerol + propanediol four steps 10 min /step G 0.5 M + P 0.5 M G 0.375 M + P 0.375 M G 0.25 M + P 0.25 M G 0M + P 0M
Glycerol + sucrose two steps: G 0.75 M 10 min G 1.5 M + sucrose 0.25 M 20 min
Trim: 279mm X 216mm
Huhtinen et al., 199773
D6 M, EB
Dilution to 1/4th in freezing straw 15 min (direct transfer)
Ethylene glycol + sucrose one step at 30◦ C EG 10% + sucrose 0.1 M 10 min/step
(0.25-mL plastic straw) From 0◦ C to – 6◦ C at 0.5◦ C/min Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.3◦ C/min
16:40
2t /15
D6
7s /11 – 4k /8
Ethylene glycol two steps at 30◦ C 5%, 10% 10 min/step
Glycerol six steps at 30◦ C, 10 min/step 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% Ethylene glycol at 30◦ C six steps, 10 min/step 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0%
February 2, 2011
Ferreira et al., 199751
D6
2k /8
Glycerol two steps at 30◦ C 5%, 10% 10 min/step
P2: SFK
D6
BLBK232-McKinnon
3k /8
c305
Hochi et al., 199647
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
2893
Huhtinen and Paranko, 200048
D6.5 Diam. < 290 µm (140–290 µm) Mean diam. = 181 µm D6.5 Diam. < 200 µm (140–200 µm) Mean diam. = 171 µm
D6.5 Diam. < 240 µm (160–240 µm) Mean diam. = 184 µm
3w /10
1w /10
6w /10
Glycerol + glutamine (100 mM) four steps 5 min/step: Glycerol 2.5% + Gln 100 mM Glycerol 5% + Gln 100 mM Glycerol 7.5% + Gln 100 mM Glycerol 10% + Gln 100 mM
Glycerol + glutamine (20 mM) four steps 5 min/step Glycerol 2.5% + Gln 20 mM Glycerol 5% + Gln 20 mM Glycerol 7.5% + Gln 20 mM Glycerol 10% + Gln 20 mM
Glycerol four steps 5 min/step: 2.5%, 5%, 7.5%, and 10%
Ethylene glycol one step 1.5 M 10 min
(0.25-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
(0.25-mL plastic straw) Seeding at – 7◦ C From –7◦ C to – 30◦ C at 0.3◦ C/min
Glycerol + sucrose six steps 5 min/step: G 10% + sucrose 0.25 M G 7.5% + sucrose 0.25 M G 5% + sucrose 0.1 M G 2.5% + sucrose 0.1 M G 0% + sucrose 0.1 M G 0% + sucrose 0 M
Ethylene glycol one step 0 M 15 min
Glycerol and sucrose six steps 5 min/step G 10% + Sucrose 0.25 M G 7.5% + Sucrose 0.25 M G 5% + Sucrose 0.1 M G 2.5% + Sucrose 0.1 M G 0% + Sucrose 0.1 M G 0% + Sucrose 0 M
Trim: 279mm X 216mm
D6 Diam. < 190 µm (140–190 µm)
Glycerol + glutamine four steps 5 min/step G 2.5% + Gln 50 mM G 5% + Gln 50 mM G 7.5% + Gln 50 mM G 10% + Gln 50 mM
Glycerol + galactose five steps 5 min/step G 1.5 M + Gal 0.3 M G 0.8 M + Gal 0.3 M G 0.3 M + Gal 0.3 M G 0 M + Gal 0.3 M G 0 M + Gal 0 M
16:40
D6 Diam. < 190 µm (140–190 µm)
(0.25-mL plastic straw) Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.5◦ C/min
Glycerol + galactose three steps 5 min/step: Glycerol 2 M Glycerol 4 M G 2 M + galactose 0.3 M
Dilution of cryoprotectant
2894
D7–8 Diam. > 350 µm (350–580 µm)
(Device to store embryo) Cooling rate
Cryoprotectant used and addition protocol
February 2, 2011
Lagneaux et al., 200075
4/9 (stage of pregnancy not indicated)
Young et al., 199774
Embryo developmental stage
BLBK232-McKinnon
3/10 (stage of pregnancy not indicated)
1v /6
References
P2: SFK
No. pregnancies (births)/No. transferred embryos
c305
Table 305.1 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
D7.5–8.5 Diam. > 300 µm D7.5–8.5 Diam. > 300 µm D7.5–8.5 Diam. > 300 µm
12t /17
0/7
Maclellan et al., 200246
0/7
3t /7
Embryos produced in vitro by ICSI
Cytochalasin B 7.5 µg/mL 45 min Trypsin 0.2% 15 min + cytochalasin B 7.5 µg/mL 45 min
Pretreated with: Trypsin 0.2% 15 min
Glycerol two steps: 5% 10 min 10% 20 min
Glycerol 10% no. steps not indicated
From 22◦ C to – 6◦ C at 4◦ C/min Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min From – 30◦ C to – 33◦ C at 0.1◦ C/min
(0.25-mL plastic straw)
(0.25-mL plastic straw) Seeding – 6◦ C; –6◦ C to – 32◦ C at 0.5◦ C/min
(continued)
Glycerol six steps 8%, 6%, 4%, 2%, 1%, 0% 10 min/step
Not indicated
Direct transfer, immediately after thawing
Trim: 279mm X 216mm
4t /7
D7.5–8.5 Diam. < 300 µm D7.5–8.5 Diam. > 300 µm
4t /16
Galli et al., 200236
(0.25-mL plastic straw) Seeding at – 4.8◦ C From – 4.8◦ C to – 35◦ C at 0.3◦ C/min
Ethylene glycol + sucrose one step 10 min Ethylene glycol 1.5 M + sucrose 0.25 M
Glycerol and sucrose three steps 5 min/step G 5% + sucrose 0.5 M G 0% + sucrose 0.5 M G 0% + sucrose 0 M
16:40
D7 EB
1 (1)/1 (another frozen embryo B not transferred)
Ulrich and Nowshari, 200276
(0.25-mL plastic straw) From 24◦ C to – 6◦ C at 1◦ C/min Seeding at – 6◦ C From – 6◦ C to – 32◦ C at – 0.3◦ C/min From – 32◦ C to – 35◦ C at – 0.1◦ C/min
Glycerol two steps: 5% 10 min 10% 20 min
D6.5 diam. < 200 µm
24k /43
Glycerol and sucrose four steps 15 min/step G 1 M + sucrose 0.25 M G 0.5 M + sucrose 0.25 M G 0 M + sucrose 0.25 M G 0% + sucrose 0 M
February 2, 2011
Lascombes and Pashen, 20009
1/1y 0/1 2x /3
(0.25-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
Pretreatment with trypsin 0.2% 15 min Glycerol three steps: Glycerol 0.5 M 15 min Glycerol 1 M 15 min Glycerol 1.5 M 15 min
P2: SFK
Diam. < 200 µm (120–187 µm) Diam. = 442 µm Diam. = 715 µm Diam. > 1000 µm (1020–1580 µm)
BLBK232-McKinnon
1x /3
c305
Legrand et al. 2001
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
Freezing of Embryos 2895
Bass et al., 200478 (and personal communication with G.E. Seidel) D7–8 Diam.300–1000 µm Mean diam. = 557 µm
D7–8 Diam. 300–1000 µm Mean diam. = 535 µm
8aa /21
5aa /22
6t /18
7t /20
D6.5–7.5 Diam. < 300 µm D6.5–7.5 Diam. > 300 µm D6.5–7.5 Diam. < 300 µm D6.5–7.5 Diam. > 300 µm
Methanol two steps 8% 5 min 4.8% 5 min
Glycerol two steps: 5% 10 min 10% 20 min
Precooled at 4◦ C for 22 h in Hamilton Thorne Equitainer before freezing
Glycerol two steps: 5% 10 min 10% 20 min
Pretreatment with trypsin 0.2% 15 min
From – 6◦ C to – 30◦ C at 0.3◦ C/min From – 30◦ C to – 33◦ C at 0.1◦ C/min
(0.25-mL plastic straw) From 22◦ C to – 6◦ C at 4◦ C/min Seeding at – 6◦ C
(0.25-mL plastic straw) From 22◦ C to – 6◦ C at 4◦ C/min Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min From – 30◦ C to – 33◦ C at 0.1◦ C/min
(0.25-mL plastic straw) Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
Glycerol and sucrose four steps 10 min/step G 6% + sucrose 0.5 M G 3% + sucrose 0.5 M G0% + sucrose 0.5 M G0% + sucrose 0 M Methanol and sucrose five steps 5 min/step M 4.8% + sucrose 0.5 M M 2.56% + sucrose 0.5 M M 0.96% + sucrose 0.5 M M 0% + sucrose 0.5 M M 0% + sucrose 0 M
Glycerol six steps: 8%, 6%, 4%, 2%, 1%, 0% 10 min/step
Glycerol and sucrose four steps 15 min/step: G 1 M + sucrose 0.25 M G 0.5 M + sucrose 0.25 M G 0 M + sucrose 0.25 M G 0% + sucrose 0 M
Trim: 279mm X 216mm
2t /5
4t /5
D7–8 Diam. > 200 µm (Diam. in case of pregnancies = 225, 290, and 765 µm)
2z /11
Glycerol three steps: Glycerol 0.5 M 15 min Glycerol 1 M 15 min Glycerol 1.5 M 15 min
Dilution of cryoprotectant
16:40
Maclellan et al., 200377
D7–8 Diam. > 200 µm
(Device to store embryo) Cooling rate
2896
0/8 (+ 1 degenerated non-transferred)
Cryoprotectant used and addition protocol
February 2, 2011
Legrand et al., 200245
Embryo developmental stage
BLBK232-McKinnon
References
P2: SFK
No. pregnancies (births)/No. transferred embryos
c305
Table 305.1 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
D6.75 M, EB, B Diam. 140–270 µm Mean diam. = 184 µm
Embryos produced in vitro by ICSI and OPU Embryos in vivo culture after IVF by ICSI from post-mortem oocytes
9dd (5 still ongoing > 6th month)/13 1ee /10
Galli et al., 200737 Glycerol 10% no. steps not indicated
Not indicated
(0.25-mL plastic straw) From 22◦ C to – 7◦ C at – 3◦ C/min Seeding at – 7◦ C From – 6◦ C to – 30◦ C at – 0.3◦ C/min
Glycerol + glutamine four steps: G 2.5% + Gln 0.1 M 5 min G 5% + Gln 0.1 M 5 min G 7.5% + Gln 0.1 M 5 min G 10% + Gln 0.1 M 10 min
Not indicated
(continued)
Glycerol and sucrose five steps 5 min/step G 7.5% + sucrose 0.25 M G 5% + sucrose 0.1 M G 2.5% + sucrose 0.1 M G 0% + sucrose 0.1 M G 0% + sucrose 0 M
Trim: 279mm X 216mm
6cc /19(+ 2 degenerated non-transferred)
From – 6◦ C to – 30◦ C at 0.3◦ C/min
Glycerol + glutamine one step 25 min G 10% + Gln 0.1 M
Glycerol and sucrose five steps 5 min/step: G 7.5% + sucrose 0.25 M G 5% + sucrose 0.1 M G 2.5% + sucrose 0.1 M G 0% + sucrose 0.1 M G 0% + sucrose 0 M
16:40
Lelong, 200792
D6.75 Diam. 156–256 µm Mean diam. = 198 µm
4bb /19 (+ 3 degenerated non-transferred)
(0.25-mL plastic straw) From 22◦ C to – 7◦ C at 3◦ C/min Seeding at – 7◦ C
February 2, 2011
Glycerol + glutamine four steps: G 2.5% + Gln. 0.1 M 5 min G 5% + Gln 0.1 M 5 min G 7.5% + Gln 0.1 M 5 min G 10% + Gln 0.1 M 10 min
P2: SFK
D6.75 Diam. 167–311 µm Mean diam. = 208 µm
BLBK232-McKinnon
5bb /19(+ 3 degenerated non-transferred)
c305
Duchamp et al., 200632
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
Freezing of Embryos 2897
BLBK232-McKinnon P2: SFK
2898
February 2, 2011
M, morula; EB, early blastocyst; B, blastocyst; EExB, early expanded blastocyst; ExB, expanded blastocyst; ICSI, intracytoplasmic sperm injection; IVF, in vitro fertilization; OPU, ovum pick-up. Glycerol (G) 10% v/v = glycerol 1.36 M; DMSO 10%v/v = 1.41 M; ethylene glycol (EG) 10% v/v = ethylene glycol 1.79 M; 1,2-propanediol (P) 10%v/v = 1.37 M; methanol (M) 10% v/v = methanol 2.46 M (4.8% v/v = 1.18 M); sucrose 10% w/v = sucrose 0.29 M. ∗ Part of these results have already been published in Yamamoto et al.7 a Spontaneous abortion between D60 and D85. b, c Spontaneous abortions, one between D32 and D36 and one at D43. d No. of pregnancies at D50 (no information after this stage of pregnancy). e Nine pregnancies at D35 but one spontaneous abortion by D50. f Spontaneous abortion between D66 and D74. g Only trophoblastic vesicle spontaneously resorbed between D32 and D40. h Only trophoblastic vesicle spontaneously resorbed between D28 and D36. i No. of pregnancies at D196 (no information after this stage of pregnancy). j One only trophoblastic vesicle spontaneously resorbed between D19 and D28, one induced abortion at D37, no information on the third pregnancy after D100. k No. of pregnancies at D60 (no information after this stage of pregnancy). l No. of pregnancies at D30 (no information after this stage of pregnancy). m No. of pregnancies at D35 (no information after this stage of pregnancy). n No. of pregnancies at D25 (no information after this stage of pregnancy). o No. of pregnancies at D50 (no information after this stage of pregnancy). p No. of pregnancies at D26 (no information after this stage of pregnancy). q Spontaneous abortion after the fourth month of pregnancy. r No. of pregnancies at D60 (no information after this stage of pregnancy), + one spontaneous abortion between D15 and D30. s No. of pregnancies at D15 (three induced abortions at D20). t No. of pregnancies at D15 (no information after this stage of pregnancy). u No. of pregnancies at D26 (all pregnancies except one have been voluntary stopped at D26). v No. of pregnancies at D26 (no information after this stage of pregnancy) another trophoblastic vesicle detected at D15 degenerated from D26. w No. of pregnancies at D14 date of induced abortions. x No. of pregnancies at D28 date of induced abortions (one more pregnancy diagnosed at D14 resorbed spontaneously between D14 and D21). y No. of pregnancies at D14 (the recipient mare was injured between D14 and D21). z No. of pregnancies at D28 date of induced abortions (one more pregnancy diagnosed at D21 resorbed spontaneously between D21 and D28). aa No. of pregnancies at D16 date of induced abortions. bb No. of pregnancies at D11 date of new embryo collection. cc No. of pregnancies at D29 date of induced abortions. dd No. of pregnancies at D17 date of induced abortions of 1/9 vesicles (7 on 8 other ongoing over sixth month). ee No. of pregnancies at D17 date of induced abortions.
c305
Table 305.1 (Continued)
P1: SFK/UKS Color: 1C
16:40 Trim: 279mm X 216mm Printer Name: Yet to Come
ART: Embryos
P1: SFK/UKS c305
P2: SFK
Color: 1C
BLBK232-McKinnon
February 2, 2011
16:40
Trim: 279mm X 216mm
Printer Name: Yet to Come
Freezing of Embryos From the 1990s, some experiments were conducted in vitro to try both to understand the reason for failures to cryopreserve horse embryos and to improve the process (Tables 305.2 and 305.3). Vitrification An initial paper, in 1994, reported pregnancies following transfer of horse embryos cryopreserved by vitrification.10 Vitrification consists of a short incubation time in high concentrations of CPs followed by direct plunging into liquid nitrogen to prevent ice crystal formation, and to induce a glass-like state. Using traditional 250-µL straws for loading the embryo, the cooling rate is approximately 2500◦ C/min. Initially used to cryopreserve mice embryos,11 vitrification was shown to be a successful method of cryopreserving embryos and oocytes of different mammalian species. Table 305.4 summarizes papers on the transfer of vitrified horse embryos. Similar to conventional freezing, different experiments (Table 305.5) were also conducted to test in vitro the effects of vitrification on horse embryos. In those experiments, different CPs, at different final concentrations and with different combinations of CPs, were tested. Ultra-rapid vitrification procedures using either Open Pulled Straws (OPS) or a cryoloop device, efficacious for embryos (e.g., porcine) known to be refractory to cryopreservation, were also evaluated. The high cooling rate of 20 000◦ C/min allows embryos to pass quickly through critical temperature zones, in an attempt to decrease chilling injuries and to allow vitrification with lower concentrations of CPs. However, this does not seem to be the ‘miraculous’ method for equine embryos. After the first report of pregnancy from a vitrified embryo, only a few studies were published (Table 305.4) showing high pregnancy rates (70/105) after transfer of only small (diameter < 300 µm) vitrified embryos, but no birth was reported. However, the method used in those experiments seems currently in use for commercial purposes, and vitrification solutions are available in commercial kits.12, 13 The actual situation The two methods, slow freezing and vitrification, seem to allow cryopreservation of only small embryos (diameter < 300 µm). The practical disadvantages of a slow freezing process are that controlled cooling requires time and elaborate equipment, whereas vitrification is a rapid procedure that requires only having the knack for moving embryos in highly concentrated solutions of CPs with the right timing and for loading them into straws with the right quantity of each solution. However, cryopreservation of equine embryos is done commercially
2899
in only a few embryo transfer centers. The Data Retrieval Committee of the IETS (International Embryo Transfer Society) reported just a slight increase in the proportion of frozen equine embryos transferred around the world over recent years: 14 of 11 498 (0.1%) in 2003; 250 of 11 672 (0.2%) in 2004; and 408 of 15 695 (0.26%) in 2006;1, 14, 15 however, those data are not exhaustive because retrieval of transfers of some countries and some embryo transfer teams seems very difficult.
Why is it so difficult to cryopreserve horse embryos and to study this cryopreservation? Problems in studying the cryopreservation of horse embryos Because of the expense of research in this field there have been only a few experiments, and each of them suffer from the small numbers of embryos tested. In other species, whether rodents, ruminants, or porcines, many experiments on cryopreservation have been conducted and in most of them examined protocols involving groups containing a few hundred embryos. With those experimental conditions significant conclusions can be obtained and methods can be progressively improved even with an empirical approach. The protocols used and/or tested for horse embryos are usually with groups of only 10 embryos. Analysis of Tables 305.1 to 305.5 underlines another problem in studies on cryopreservation of horse embryos. Parameters were different from one tested protocol to another; even when using the same CP, the concentration, number of steps, time and temperature of addition and/or removal of CP, cooling rate, and final temperature varied. Comparisons of results obtained in the different studies would be possible if only one parameter was tested at the same time. Studies including non-surgical transfer or in vitro development in culture for 24 to 72 h gives only bimodal responses, i.e., pregnant or not after transfer, or normally developed or degenerated during culture. Both the fertility of recipient mares and conditions for in vitro culture introduce bias in the results, so high numbers of embryos are needed to show differences between two tested protocols not having the same efficacy. Various methods to try to evaluate quantitatively the damage induced by different cryoprotective treatments have been used in order to decrease the number of embryos needed to obtain statistically significant data. Ideal tests to assess embryonic viability as described by Rieger16 and Butler and Biggers17 are not yet available. Tables 305.3 and 305.5
7/8 (mean morphological grade 2.1)
Slade et al., 198563 (24 h in Ham’s F10 + FCS 5%)
1a /5 (mean morphological grade 3.7) 0a /5 (mean morphological grade 4.2) 2a /8 (mean morphological grade 3.4) 0a /5 (mean morphological grade 4.2)
1/8 (mean morphological grade 4.2)
D6.5–D7 Mean diam. = 367 µm (275–575 µm) Mean diam. = 504 µm (300–945 µm) Mean diam. = 506 µm (250–890 µm) Mean diam. = 427 µm (220–700 µm)
1,2-Propanediol two steps: 5% 5 min 10% 10–20 min (0.5-mL plastic straw)
Glycerol two steps: 5% 10 min 10% 20–30 min (1-mL glass ampoule)
Glycerol two steps: 5% 10 min 10% 20–30 min (0.5-mL plastic straw)
Seeding at – 6◦ C From – 6◦ C to – 33◦ C at 0.3◦ C/min Thawed 20 s in water at 37◦ C or in air at 20◦ C Seeding at – 6◦ C From – 6◦ C to – 33◦ C at 0.8◦ C/min Thawed for 20 s in water at 37◦ C or in air at 20◦ C
Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min then to – 33◦ C at 0.1◦ C/min Seeding at – 6◦ C From – 6◦ C to – 33◦ C at 0.3◦ C/min then to – 38◦ C at 0.1◦ C/min Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min then to – 33◦ C at 0.1◦ C/min Seeding at – 6◦ C From – 6◦ C to – 33◦ C at 0.3◦ C/min then to – 38◦ C at 0.1◦ C/min
1,2-Propanediol + sucrose four steps: 6 min/step P 6% + sucrose 10% P 3% + sucrose 10% P 0% + sucrose 10% P 0% + sucrose 0%
Glycerol six steps: 10 min/step
Trim: 279mm X 216mm
4/8 (mean morphological grade 2.8)
D6 M – EB – B
Glycerol six steps 10 min/step: 1.1 M – 0.88 M – 0.66 M – 0.44 M – 0.22 M – 0 M
Seeding at – 7◦ C From – 7◦ C to – 26◦ C at 0.3◦ C/min then to – 38◦ C at 0.1◦ C/min Seeding at – 7◦ C From – 7◦ C to – 35◦ C at 0.3◦ C/min then to – 38◦ C at 0.1◦ C/min
16:40
D7
D6
D7
Glycerol three steps: 0.44 M 10 in 0.88 M 10 min 1.32 M 30 min (1-mL glass ampoule)
Dilution of cryoprotectant
Cooling rate
2900
D6
Addition of cryoprotectant (device to store embryo)
February 2, 2011
Seidel et al. 198979 (48 h in Ham’s F10 + FCS 10%)
4/7 (mean morphological grade 3.2) 1/7 (mean morphological grade 4.2) 5/7 (mean morphological grade 3.0) 2/7 (mean morphological grade 3.7)
Takeda et al., 198462 (24 h in Ham’s F10 + FCS 10%)
Embryo developmental stage
BLBK232-McKinnon
5/8 (mean morphological grade 2.9)
No. normal developed embryos after cuture/No. frozen-thawed embryos
P2: SFK
References (time and medium of culture after thawing)
c305
Table 305.2 In vitro development of frozen-thawed equine embryos in reports published from 1981 to 2007
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
In vitro produced embryos by ICSI (M and B)
Ethylene glycol 10% + sucrose 0.1 M One step at 30◦ C 10 min (0.25-mL plastic straw)
Glycerol + galactose three step 5 min/step Glycerol 2 M Glycerol 4 M G 2 M + galactose 0.3 M (0.25-mL plastic straw)
Glycerol 1 M one step 15 min (0.25-mL plastic straw)
Glycerol + galactose five steps: 5 min/step G 1.5 M + galactose 0.3 M G 0.8 M + galactose 0.3 M G 0.3 M + galactose 0.3 M G 0 M + galactose 0.3 M G 0 M + galactose 0 M
Dilution to 1/4 in freezing straw 15 min and expelled in culture medium
Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.3◦ C/min
From 0◦ C to – 6◦ C at 0.5◦ C/min Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.3◦ C/min
B, blastocyst; BSA, bovine serum albumin; EB, early blastocyst; EExB, early expanded blastocyst; ExB, expanded blastocyst; FCS, fetal calf serum; ICSI, intracytoplasmic sperm injection; M, morula; TCM-199, tissue culture medium. a Only embryos with grade < 3.0 were considered as normally developed. b Embryos with the same quality or increasing quality during culture 24 h. c Coats = zona pellucida and capsule.
5/5
D7–8 Diam. > 300µm (300–680 µm) Mean diam. = 471 µm D7–8 Diam. > 300 µm (300–680 µm) Mean diam. = 518 µm
Glycerol six steps at 30◦ C, 10 min/step 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0%
Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.3◦ C/min
Trim: 279mm X 216mm
Matsukawa et al., 200338
0/4 (+ 2 or 3 degenerated embryos after warming were not cultured).
Young et al., 199774 (co-culture with oviductal cells for 36 h)
Glycerol two steps at 30◦ C: 5%, 10% 10 min/step (0.25-mL plastic straw)
Glycerol five steps: 30 min/step
16:40
Diam. < 200 µm (mean diam. = 172 µm) Diam. 200–300 µm (mean diam. = 230 µm) Diam. > 300 µm (mean diam. = 509 µm)
D6 to D7.5 B–EExB ExB
Seeding at – 7◦ C From – 6◦ C to – 30◦ C or – 33◦ C at 0.3◦ C/min
February 2, 2011
6/6 (+ 1 degenerated embryo after warming was not cultured)
(Volume changes of embryos during cryoprotectant addition measured too) 9 (5 with slight fracture of coatsc )/9b 5 (3 with slight and 1 with severe fracture of coatsc )/9b 2 (2 with slight fracture of coatsc )/9b
Hochi et al., 199440 (48 h in TCM199 + FCS 10%)
Glycerol 10% vs Glycerol 10% + agar 1.2% (0.25-mL plastic straw)
P2: SFK
D6.5 M EB B ExB
BLBK232-McKinnon
81a /133
c305
Poitras et al., 199480 (24 h in Ham’s F10 + BSA 4%)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
Freezing of Embryos 2901
Staining of whole embryos on slide with aceto-orcein after 24-h culture (table 2a) to try to identify and to count mitotic, interphasic, and pycnotic nuclei Comparison of four groups (straw/tube, – 33◦ C/– 38◦ C) (n = 4 × 8 = 32) Electron microscopy observations of sections of frozen-thawed embryos compared with fresh embryos and embryos only treated with cryoprotectant without freezing-thawing (n not indicated) Staining with diacetylfluorescein (FDA) to evaluate the percentage of live cells after culture 24 h post-thawing (n = 34) Staining with FDA to compare fresh and frozen-thawed embryos and to evaluate viability of inner cell mass and trophectodermal cells (n = 15 vs 15) 1. Variation of the metabolic activity for glucose and glutamine of each embryo before and after treatment 2. Counting of interphasic, mitotic, and pycnotic cells on serial sections of each embryos Comparison of values of frozen-thawed embryos with values of fresh embryos and embryos only treated with cryoprotectant without freezing-thawing (n = 5 vs 5 and 4)
Wilson et al., 198749
Guay and Poitras, 198981
Barry et al., 198982
Rieger et al., 199121 Glycerol four steps 5 min/step: 2.5%, 5%, 7.5%, and 10% (0.25-mL plastic straw)
Glycerol five steps 30 min/step
Glycerol + sucrose five steps: Glycerol 10% 1 min Sucrose 1.08 M 30 min 0%, 0 M 5 min
Glycerol seven steps 5 min/step 10%, 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0%
Seeding at – 7◦ C From – 6◦ C to – 30◦ C or – 33◦ C at 0.3◦ C/min Seeding at – 7◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min then to – 33◦ C at 0.1◦ C/min
Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
Trim: 279mm X 216mm
Glycerol 1.35 M two steps: Glycerol 0.67 M 15 min Glycerol 1.35 M 15 min (0.25-mL plastic straw)
Glycerol + sucrose three steps Glycerol 10% 1 min Sucrose 1.08 M 30 min 0%, 0 M 5 min
Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min
16:40
Glycerol 10% vs Glycerol 10% + agar 1.2% (0.25-mL plastic straw)
Glycerol six steps 10 min /step
Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min then to – 33◦ C at 0.1◦ C/min vs From – 6◦ C to – 33◦ C at 0.3◦ C/min then to – 38◦ C at 0.1◦ C/min
February 2, 2011
Glycerol one step: 10% 30 min (0.5-mL plastic straw)
Glycerol two steps 5% 10 min 10% 20–30 min (0.5-mL plastic straw vs 1-mL glass tube)
Dilution of cryoprotectant
Cooling rate
2902
Slade et al., 198563
Addition of cryoprotectant (device to store embryo)
BLBK232-McKinnon
References
P2: SFK
Parameters of evaluation (n = no. of frozen-thawed embryos)
c305
Table 305.3 Different procedures of in vitro evaluations of frozen-/thawed equine embryos published from 1994 to 2007
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos with values of fresh embryos and embryos only treated with cryoprotectant without freezing-thawing + Electron microscopy observations of sections of each embryo of each of three groups (n = 6 vs 5 and 5) Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos with values of fresh embryos and embryos only treated with cryoprotectant without freezing-thawing (n = 6 vs 4 and 5)
Bruyas et al., 199322
Bruyas et al., 199523 (expt. 2)
Trim: 279mm X 216mm
Freezing of Embryos (continued)
Ethylene glycol four steps: 1.5 M, 1 M, 0.5 M, 0 M 10 min/step (and culture 6 h before histological evaluation)
Seeding at – 7◦ C From – 7◦ C to – 35◦ C at – 0.3◦ C/min
16:40
Ethylene glycol 1.5 M three steps 10 min/step 0.5 M, 1 M, 1.5 M (0.25-mL plastic straw)
Glycerol six steps 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% 5 min/step (and culture 6 h before histological evaluation)
Seeding at – 7◦ C From – 7◦ C to – 30◦ C at – 0.3◦ C/min
February 2, 2011
Glycerol four steps 5 min/step 2.5%, 5%, 7.5%, and 10% (0.25-mL plastic straw)
Glycerol five steps: 7.3%, 5.5%, 3.7%, 1.8%, 0% 8–10 min/step 1,2-Propanediol five steps: 7.3%, 5.5%, 3.7%, 1.8%, 0% 8–10 min/step Glycerol + sucrose: no. steps and time not indicated
Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3–0.6◦ C/min Deep-freezing in stainless-steel cylinder and liquid nitrogen container 5 min in liquid nitrogen vapor before plunging into liquid nitrogen
P2: SFK
Glycerol two steps: 5.5% 10 min – 11% 20 min (0.5-mL plastic straw) 1,2-Propanediol two steps: 5.5% 10 min – 11% 20 min (0.5-mL plastic straw) Glycerol + sucrose three steps: glycerol 5.5% 10 min glycerol 11% 20 min G 20% + sucrose 0.5 M 20 min (0.5-mL plastic straw)
BLBK232-McKinnon
Electron microscopy observations of sections of frozen-thawed embryos of each of three groups compared with fresh embryos (n = 2/group vs 3 fresh control embryos)
c305
Landim e Alvarenga et al., 199350
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
2903
Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos with values of fresh embryos and embryos only treated with cryoprotectant without freezing-thawing (n = 6 vs 5 and 5) Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos with values of fresh embryos and embryos only treated with cryoprotectant without freezing-thawing (n = 4 vs 5 and 5) Embryos were incubated with DAPI (DAPI stains only dead cells) and the authors measured the area of fluorescence and calulated the ratio area with fluorescent staining / total area of embryos to evaluate the rate or the percentage of dead cells. Comparison of values of frozen-thawed embryos in the groups with values of fresh embryos (two experiments with n = 10/group) + transfer after staining (Table 305.1)
Bruyas et al., 199725
Huhtinen et al., 199773
1,2-Propanediol four steps: 1.5 M, 1 M, 0.5 M, 0 M 10 min/step (and culture 6 h before histological evaluation)
Glycerol six steps: 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0% 5 min/step vs Glycerol and sucrose six steps 5 min/step: G 10% + Sucrose 0.25 M G 7.5% + Sucrose 0.25 M G 5% + Sucrose 0.1 M G 2.5% + Sucrose 0.1 M G 0% + Sucrose 0.1 M G 0% + Sucrose 0 M
Seeding at – 7◦ C From – 7◦ C to – 35◦ C at – 0.3◦ C/min
Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
16:40 Trim: 279mm X 216mm
Glycerol four steps 5 min/step 2.5%, 5%, 7.5%, and 10% (0.25[mL plastic straw)
DMSO four steps: 1.5 M, 1 M, 0.5 M, 0 M 10 min/step (and culture 6 h before histological evaluation)
Seeding at – 7◦ C From – 7◦ C to – 35◦ C at – 0.3◦ C/min
February 2, 2011
1,2-Propanediol 1.5 M three steps 10 min/step 0.5 M, 1 M, 1.5 M (0.25-mL plastic straw)
DMSO 1.5 M three steps 10 min/step 0.5 M, 1 M, 1.5 M (0.25-mL plastic straw)
Dilution of cryoprotectant
Cooling rate
2904
Bruyas et al., 199724
Addition of cryoprotectant (device to store embryo)
BLBK232-McKinnon
References
P2: SFK
Parameters of evaluation (n = no. of frozen-thawed embryos)
c305
Table 305.3 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
Staining with DAPI to evaluate the rate of dead cells (ratio of fluorescent area/total area of embryos) Comparison of values of frozen-thawed embryos in the three groups (n = 3/group)
Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos with values of two groups of embryos treated only with glycerol 1.5 M or 1 M without freezing-thawing (n = 4 vs 6 and 6) Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos with values of two groups of embryos treated only with propanediol 1.5 M or 3 M without freezing-thawing (n = 6 vs 6 and 7)
Lagneaux et al., 199783
Legrand et al., 199929
Bruyas et al., 199926 1,2-propanediol 3 M four steps 10 min/step 0.5 M, 1 M, 1.5 M, 3 M (0.25-mL plastic straw)
Propanediol five steps: 10 min/step 3 M, 1.5 M, 1 M, 0.5 M, 0 M (and culture 6 h before histological evaluation)
Seeding at – 7◦ C From – 7◦ C to – 35◦ C at 0.3◦ C/min
Freezing of Embryos (continued)
Glcerol + sucrose four steps 10 min/step G 0.75 M + sucrose 0.25 M G 0.3 M + sucrose 0.25 M G 0 M + sucrose 0.25 M G 0 M + sucrose 0 M (and culture 6 h before histological evaluation)
Trim: 279mm X 216mm
Seeding at – 7◦ C From – 7◦ C to – 35◦ C at 0.3◦ C/min
Glycerol six steps 5 min 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0%
16:40
Glycerol 1 M two steps: 0.5 M 10 min 1 M 20 min (0.25-mL plastic straw)
Glycerol six steps 5 min 8.3%, 6.7%, 5%, 3.3%, 1.7%, 0%
Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
February 2, 2011
Nothing one step 5 min
Glycerol + 1,2-propanediol four steps 10 min : G 0.5 M + P 0.5 M G 0.375 M + P 0.375 M G 0.25 M + P 0.25 M G 0M + P 0M
Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.4◦ C/min
BLBK232-McKinnon
Glycerol four steps 5 min/step: 2.5%, 5%, 7.5%, and 10% (0.25-mL plastic straw) Antifreeze protein (AFP) 20 mg/mL one step 20 min (0.25-mL plastic straw) Glycerol + AFP four steps 5 min/step G 2.5%; G 5%; G 7.5%; G 10% + AFP 20 mg/ml (0.25-mL plastic straw)
Glycerol + 1,2-propanediol two steps : G 0.35 M + P 0.35 M 10 min G 0.75 M + P 0.75 M 20 min (0.25-mL plastic straw)
P2: SFK
Electron microscopy observations of sections of frozen-thawed embryos compared with fresh embryos (n = 3/group)
c305
Ferreira et al., 199751
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
2905
Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos with values of one group of vitrified-warmed embryos and one group of control fresh embryos (n = 5 vs 6 and 5) Staining with DAPI to evaluate the rate of dead cells (ratio of fluorescent area/total area of embryos) Comparison of values of frozen-thawed embryos in the two groups (n = 9/group) + transfer after staining (Table 305.1) Confocal microscopic observation of other embryos of each of two groups to observe nuclei and cytoskeleton (n = 9/group) Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell Comparison of values of frozen-thawed embryos of four groups (n = 6/group)
Bruyas et al., 199927
Huhtinen and Paranko, 200048
Bruyas et al., 200028
Glycerol four steps 10 min/step: 1.125 M, 0.75 M, 0.375 M, 0 M
Glycerol and sucrose six steps 5 min/step: G 10% + Sucrose 0.25 M G 7.5% + Sucrose 0.25 M G 5% + Sucrose 0.1 M G 2.5% + Sucrose 0.1 M G 0% + Sucrose 0.1 M G 0% + Sucrose 0 M Ethylene glycol one step 0 M 15 min
Ethylene glycol vs glycerol three steps 1 M, 0.5 M, 0 M 10 min/step vs Ethylene glycol or glycerol + sucrose 0.25 M four steps: EG vs G 1.25 M + S 0.25 M EG vs G 0.6 M + S 0.25 M EG or G 0 M + S 0.25 M EG or G 0 M + S 0 M (and culture 6 h before histological evaluation)
Seeding at – 7◦ C From – 7◦ C to – 35◦ C at 0.3◦ C/min
Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
Seeding at – 7◦ C From – 7◦ C to – 35◦ C at 0.3◦ C/min
Glycerol 1.5 M four steps 10 min/step 0.375 M, 0.75 M, 1.125 M, 1.5 M (0.25-mL plastic straw)
Glycerol + glutamine four steps 5 min/step: Glycerol 2.5% + Gln 50 mM Glycerol 5% + Gln 50 mM Glycerol 7.5% + Gln 50 mM Glycerol 10% + Gln 50 mM (0.25-mL plastic straw)
16:40 Trim: 279mm X 216mm
Glycerol 1.5 M vs Ethylene glycol 1.5 M three steps 10 min/step 0.5 M, 1 M, 1.5 M (0.25-mL plastic straw)
February 2, 2011
Ethylene glycol 1.5 M one step 10 min (0.25-mL plastic straw)
Dilution of cryoprotectant
2906
Cooling rate
Addition of cryoprotectant (device to store embryo)
BLBK232-McKinnon
References
P2: SFK
Parameters of evaluation (n = no. of frozen-thawed embryos)
c305
Table 305.3 (Continued)
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
ART: Embryos
1. Variation of the metabolic activity for glucose and glutamine of each embryo before and after treatment 2. Propidium iodide and Hoechst 33342 staining to estimate (and not count exactly) percent live cells (n = 8 treatment 1 and 3, n = 5 treatment 2)
O’Donovan et al., 200084
Ethylene glycol + galactose three or four steps
Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.5◦ C/min
Ethylene glycol + galactose Comparison of three treatments two steps : galactose 0.3 M 10 in EG 2 M + galactose 0.3 M 40 min (0.25-mL plastic straw) three steps : galactose 0.3 M 10 min EG 2 M + galactose 0.3 M 15 min EG 4.5 M + galactose 0.3 M 15 min (0.25-mL plastic straw) four steps : galactose 0.3 M 10 min EG 2 M + galactose 0.3 M 10 min EG 4.5 M+ galactose 0.3 M 15 min EG 2 M + galactose 0.8 M 10 min (0.25-mL plastic straw)
February 2, 2011 16:40 Trim: 279mm X 216mm
(continued)
Glycerol and sucrose six steps 5 min/step: Glycerol 10% + Suc 0.25 M Glycerol 7.5% + Suc 0.25 M Glycerol 5% + Suc 0.1 M Glycerol 2.5% + Suc 0.1 M Glycerol 0% + Suc 0.1 M Glycerol 0% + Suc 0 M
Seeding at – 7◦ C From – 7◦ C to – 30◦ C at 0.3◦ C/min
P2: SFK
Glycerol four steps: 2.5%, 5%, 7.5%, and 10% 5 min/step (0.25-mL plastic straw) Glycerol + glutamine (20 mM) four steps 5 min/step Glycerol 2.5% + Gln 20 mM Glycerol 5% + Gln 20 mM Glycerol 7.5% + Gln 20 mM Glycerol 10% + Gln 20 mM (0.25-mL plastic straw) Glycerol + glutamine (100 mM) four steps 5 min/step: Glycerol 2.5%+ Gln 100 mM Glycerol 5% + Gln 100 mM Glycerol 7.5%+ Gln 100 mM Glycerol 10%+ Gln 100 mM (0.25-mL plastic straw)
BLBK232-McKinnon
Staining with DAPI to evaluate the rate of dead cells (ratio of fluorescent area/total area of embryos) Comparison of values of frozen-thawed embryos in the three groups (n = 10/group) + transfer after staining (Table 305.1)
c305
Lagneaux et al., 200075
P1: SFK/UKS Color: 1C
Printer Name: Yet to Come
Freezing of Embryos 2907
Morphological evaluation at each step + after culture for 20 h + propidium iodide & Hoechst 33342 staining to estimate (and not count exactly) percent live cells Staining with DAPI to evaluate the rate of dead cells (ratio of fluorescent area/total area of embryos) Comparison of values of frozen-thawed embryos in the four groups (n = 7/group) + transfer after staining (Table 305.1) 1. Staining with DAPI to count the percentage of dead cells 2. Staining with Alexafluor 488 phalloidin to visualize the actin cytoskeleton Comparison of values of frozen-thawed embryos in the three groups and with values of 10 fresh control embryos (n = 14 treatment 1, n = 11 treatment 2 and 3) Morphological evaluation at each step, + after culture for 3 h + DAPI staining to count dead cells + BrDU staining to count cells entering S phase for 1 h (index of mitotic activity) + Hoechst 33342 staining to count all cells Comparison of values of frozen-thawed with values of OPS vitrified-warmed embryos (Table 305.5) (n = 18 vs 20)
Maclellan et al., 200246
Tharasanit et al., 200531
Moussa et al., 200530
Expelled in medium without cryoprotectant
Glycerol six steps: 8%, 6%, 4%, 2%, 1%, 0% 10 min/step
Glycerol four steps: 7.5%, 5%, 2.5%, 0% 10 min/step
Glycerol and sucrose five steps 5 min/step G 7.5% + Sucrose 0.25 M G 5% +S ucrose 0.1 M G 2.5% + Sucrose 0.1 M G 0% + Sucrose 0.1 M G 0% + Sucrose 0 M
Seeding at – 6◦ C From – 6◦ C to – 35◦ C at 0.3◦ C/min
From 22◦ C to – 6◦ C at 4◦ C/min Seeding at – 6◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min From – 30◦ C to – 33◦ C at 0.1◦ C/min
Seeding at – 6◦ C From – 6◦ C to – 33◦ C at 0.3◦ C/min
From 22◦ C to – 7◦ C at 3◦ C/min Seeding at – 7◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min
Glycerol 10% four steps (2.5%, 5%, 7.5%, 10%) 10 min/step or pretreated with trypsin 0.2% 15 min or cytochalasin B 7.5 µg/mL (0.25-mL plastic straw)
Glycerol + glutamine four steps 5 min Glycerol 2.5%+ Gln 0.1 M Glycerol 5% + Gln 0.1 M Glycerol 7.5% + Gln 0.1 M Glycerol 10% + Gln 0.1 M (0.25-mL plastic straw)
February 2, 2011
Glycerol two steps 5% – 10 min; 10% – 20 min (0.25-mL plastic straw) or pretreated with Trypsin 0.2% 15 min vs Cytochalasin B 7.5 µg/mL 45 min vs Trypsin 0.2% 15 min + cytochalasin B 7.5 µg/mL 45 min
Ethylene glycol 1.8 M + sucrose 0.1 M one step at 30◦ C 10 min (0.25-mL plastic straw)
2908
Oberstein et al., 200185
Dilution of cryoprotectant
P2: SFK
Cooling rate
Addition of cryoprotectant (device to store embryo)
BLBK232-McKinnon
References
Parameters of evaluation (n = no. of frozen-thawed embryos)
c305
Table 305.3 (Continued)
P1: SFK/UKS Color: 1C
16:40 Trim: 279mm X 216mm Printer Name: Yet to Come
ART: Embryos
EG and sucrose 4 – 7 or 5 steps 5 min/step [EG 10.5% + Sucrose 0.25 M] [EG 8.4% + Sucrose 0.25 M] [EG 6.3% + Sucrose 0.25 M] [EG 4.2% + Sucrose 0.1 M EG 2.1% + Sucrose 0.1 M EG 0% + Sucrose 0.1 M EG 0% + Sucrose 0 M
Expt. 3: Ethylene glycol (EG) comparison four goups: EG + glutamine 0.1 M Group 1: 5.6% (1 M) three steps EG 2.1, 4.2% 5 min/step; 5.6% 15 min Group 2: 11.2% (2 M) five steps EG 2.1, 4.2, 6.3, 8.4, 11.2% 5 min/step Group 3: 8.4% (1.5 M) four steps EG 2.1, 4.2, 6.3% 5 min; 8.4% 10 min Group 4: 8.4% (1.5 M) one step EG 8.4% 25 min/step
16:40 Trim: 279mm X 216mm
Freezing of Embryos
DAPI (dead cells were stained), 4 ,6-diamidino-2-phenylindole; BrDU = 5 -bromo-2 -deoxyuridine (incorporated during S phase into newly synthesized DNA strands); FDA (live cells were stained), diacetylfluorescein. Glycerol 10% v/v = glycerol 1.36 M; DMSO 10% v/v = 1.41 M; ethylene glycol 10% v/v = ethylene-glycol 1.79 M; 1,2-propanediol 10% v/v = 1,2-propanediol 1.37 M.
Glycerol and sucrose 4 – 7 or five steps 5 min/step [G 12.5% + Sucrose 0.25 M] [G 10% + Sucrose 0.25 M] (G 7.5% + Sucrose 0.25 M) G 5% + Sucrose 0.1 M G 2.5% + Sucrose 0.1 M G 0% + Sucrose 0.1 M G 0% + Sucrose 0 M
Glycerol + sucrose: five steps 5 min/step G 7.5% + Sucrose 0.25 M G 5% + Sucrose 0.1 M G 2.5% + Sucrose 0.1 M G 0% + Sucrose 0.1 M G 0% + Sucrose 0 M
February 2, 2011
Expt. 2: Glycerol comparison 4 groups : Glycerol + glutamine 0.1 M Group 1: 7.5% (1 M) three steps: G 2.5% 5 min, 5% 5 min, 7.5% 15 min Group 2: 15% (2 M) five steps: G 2.5, 5, 7.5, 10, 15% 5 min/step Group 3: 10% (1.3 M) four steps: Glycerol 2.5, 5, 7.5% 5 min/step + G 10% 10 min Group 4: 10% (1.3 M) one step Glycerol 10% 25 min
From 22◦ C to – 7◦ C at 3◦ C/min Seeding at – 7◦ C From – 6◦ C to – 30◦ C at 0.3◦ C/min
P2: SFK
Expt. 1: Last step 5 vs 20 min Glycerol + glutamine 0.1 M four steps G 2.5, 5, 7.5, 10% 5 min/step Versus G 2.5, 5, 7.5% 5 min/step and Glycerol 10% 20 min
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Staining with DAPI to count the rate of dead cells Comparison of values of frozen-thawed embryos in the three experiments where two or four groups were compared (n = 10/group)
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Ethylene glycol 4.6 M + glycerol 3.4 M + sodium pyruvate 0.3 mM + glucose 3.3 mM + 20% FCS (three steps: G 1.4 M, 5 min G 1.4 M + EG 3.6 M, 5 min VS < 1 min)
D6 diam. < 300 µm
D6–8 diam. > 300 µm
4b /6
0/3
Eldridge-Panuska et al., 200556 (expt. 1)
Eldridge-Panuska et al., 200556 (expt. 2)
Ethylene glycol 4.6 M + glycerol 3.4 M + sodium pyruvate 0.3 mM + glucose 3.3 mM + 20% FCS (three steps: G 1.4 M, 5 min G 1.4 M + G 3.6 M, 5 min – VS < 1 min) Ethylene glycol 6.6 M + glycerol 1.4 M + sodium pyruvate 0.3 mM + glucose 3.3 mM + 20% FCS three steps: G 1.4 M, 5 min G 1.4 M + EG 3.6 M, 5 min – VS < 1 min Ethylene glycol 4.6 M + glycerol 3.4 M + sodium pyruvate 0.3 mM + glucose 3.3 mM + 20% FCS (three steps: G 1.4 M, 5 min G 1.4 M + EG 3.6 M, 5 min – VS < 1 min)
D6 diam < 300 µm mean diam. = 186 µm D6–8 diam. > 300 µm mean diam. = 575 µm
D6–8 diam. > 300 µm mean diam. = 643 µm
D6 diam. < 300 µm mean diam. = 182 µm
10c /22
0/10
0/9(+1 degenerated non transferred)
16d /26
0.25-mL plastic straw (air 10 s, waterbath at 20◦ C for 10 s)
Dilution into straw 30 µL VS in 180 µL galactose solution 0.5 M 5 min then direct transfer
Dilution into straw 90 µL VS in 120 µL galactose solution 0.5 M Then two steps : Galactose 0.25 M 5 min Galactose 0 M < 10 min
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(Air 10 sec, Waterbath at 20◦ C for 10 sec)
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0.25 ml plastic straw
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Dilution into straw 90 µL VS in 120 µL galactose solution 0.5M Then 2 steps : Galactose 0.25 M 5min Galactose 0M <10 min
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(waterbath at 20◦ C for 20 s)
Dilution into straw 50 µL VS in 150 µL sucrose solution 0.5 M and expelled in sucrose 0. 5 M 10 min Then one step: sucrose 0 M or four steps 10 min/step sucrose 0.38 M, 0.25 M, 0.13 M, 0 M
Dilution of cryoprotectants
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0.25-mL plastic straw
Device to vitrify embryo (warming)
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(9 other vitrified embryos were not transferred).
Ethylene glycol 40% + Ficoll 18% w/v + sucrose 0.3 M (two steps: EG 20% 10–20 min VS < 2 min)
D5 to D7 M to B Mean diameter = 169.3 µm
2a /5 (+ 2 degenerated non-transferred)
Hochi et al., 1994a
Vitrification solution VS (addition of cryoprotectants)
Embryo developmental stage
Reference
No. of pregnancies (births)/no. of transferred embryos
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Table 305.4 Results of embryo transfer of vitrified-warmed equine embryos published from 1994 to 2007
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5f /20 (+ 2 degenerated non-transferred) D6.75 M, EB, B Diam. < 300 µm (140–270 µm) Mean diam. = 187 µm
D6.5 diam. < 300 µm PVA instead of FCS in VS
5e /6
Ethylene glycol 18% + DMSO 18% + sucrose 0.4 M + glutamine 0.1% w/v + 20% FCS (two steps EG 7.5% + DMSO 7.5% + sucrose 0 M 3 min VS 45 sec)
Ethylene glycol 4.6 M + glycerol 3.4 M + sodium pyruvate 0.3 mM + glucose 3.3 mM (three steps: G 1.4 M, 5 min G 1.4 M + EG 3.6 M, 5 min – VS < 1 min)
OPS straw (in medium of dilution at 30◦ C)
(air 10 s, waterbath at 20◦ C for 10 s)
0.25-mL plastic straw
(continued)
Immediately expelled in: sucrose 0.2 M 1 min sucrose 0.2 M 5 min sucrose 0.1 M 1 min
Dilution into straw 30 µL VS in 180 µL galactose solution 0.5 M 6 min then direct transfer
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Lelong, 200792
D6.5 diam. < 300 µm 20% FCS in VS
5e /7
(air 10 s, waterbath at 20◦ C for 10 s)
Dilution into straw 30 µL VS in 180 µL galactose solution 0.5 M 6 min then direct transfer
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Hudson et al., 2006b87
D6.5 cooled embryos stored at 5◦ C for 12–19 h Diam. < 300 µm; mean diam. = 185 µm
13e /20
0.25-mL plastic straw
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Ethylene glycol 4.6 M + glycerol 3.4 M + sodium pyruvate 0.3 mM + glucose 3.3 mM+ 20% FCS (three steps: G 1.4 M, 5 min G 1.4 M + EG 3.6 M, 5 min – VS < 1 min)
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D6.5 fresh embryos Diam. < 300 µm; mean diam. = 187 µm
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15e /20
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B diam. 400–750 µm (400–750 µm)
0i /6
B, blastocyst; EB, early blastocyst;FCS, fetal calf serum; M, morula; PVA, polyvinyl alcohol. DMSO 18% v/v = DMSO 2.53 M; ethylene glycol 18% v/v = ethylene glycol 3.22 M. a No. of pregnancies at D60 (no information after this stage of pregnancy). b No. of pregnancies at D16 (date of induced abortion except for two pregnancies continued to D38). c No. of pregnancies at D16 (date of induced abortion except for three pregnancies continued to D35). d No. of pregnancies at D16 (date of induced abortion except for two pregnancies continued to D35). e No. of pregnancies at D16 (no information after this stage of pregnancy). f No. of pregnancies at D29 date of induced abortions. g Time of steps was different between embryos with φ [300–550 µm] and embryos with φ [550–750 µm] (steps 1 & 2: 5 min vs 7 min – step 3: 40 vs 60 s). Ø, diameter. h Time of steps was different between embryos with φ [300–550 µm] and embryos with φ [550–750 µm] (steps1 & 2: 10 min vs 14 min – step 3: 60 vs 90 s). i No. of pregnancies at D14–16 (no information after this stage of pregnancy).
0.25-mL plastic straw (air 24◦ C 10 s, waterbath at 37◦ C for 10 s)
Dilution into straw 30 µL VS in 180 µL galactose solution 0.5 M 3 min Then three steps: Galactose 0.3 M 3 min Galactose 0.15 M 3 min Galactose 0 M 3 min
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Ethylene glycol 7 M + galactose 0.5 M + Ficoll 18% (w/v) (four steps: EG 1.5 M, 5 min EG 3 M, 10 min EG 5 M, 5min VS 60 s)
B diam. 300–400 µm
6i /11
0.25-mL plastic straw (air 24◦ C 10 s, waterbath at 20◦ C for 10 s)
Dilution of cryoprotectants
16:40
0i /13
Ethylene glycol 4.6 M + glycerol 3.4 M (three stepsg : G 1.4 M, 5 min or 7 min G 1.4 M + EG 3.6 M, 5 min or 7 min VS 40 or 60 s) Ethylene glycol 4.6 M + glycerol 3.4 M (three stepsh : G 1.4 M, 10 min or 14 min G 1.4 M + EG 3.6 M, 10 min or 14 min VS 60 or 90 s)
B diam. > 300 µm (300–550 µm) (short time for each step addition of CP) (550–750 µm) (long time for each step addition of CP) B diam. > 300 µm (300–550 µm) (short time for each step addition of CP) (550–750 µm) (long time for each step addition of CP)
0i /14
Device to vitrify embryo (warming)
February 2, 2011
Campos-Chillon et al., 200688
Vitrification solution VS (addition of cryoprotectants)
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Embryo developmental stage
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Reference
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No. of pregnancies (births)/no. of transferred embryos
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Table 305.4 (Continued)
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Staining with DAPI 3/ c
Chaves et al., 199790 D7 M-B
Diam. < 200 µm; mean Ø = 173 µm Diam. 200–300 µm; mean diam. = 248.5 µm Diam. > 300 µm; mean diam. = 515 µm
D6 to D8 M to B
0.5-mL plastic straw (waterbath at 37◦ C for 10 s)
(waterbath at 20◦ C for 10 s)
Ethylene glycol 8 M + PVP 7% (two steps: EG 2 M 5 min VS 30 s)
(waterbath at 20◦ C for 20 s)
0.25-mL plastic straw
0.25-mL plastic straw (waterbath at 20◦ C for 20 s)
Glycerol 25% + ethylene glycol 25% (two steps: G 10% + EG 20% 4 min VS 30 s)
Ethylene glycol 40% + Ficoll 18% w/v + sucrose 0.3 M (Two steps: EG 20% 10–20 min VS < 2 min)
Ethylene glycol 40% + Ficoll 18% w/v + sucrose 0.3 M One step (direct) vs Two steps: (EG 20% 10–20 min VS < 2 min)
Freezing of Embryos (continued)
Dilution into straw 50 µL VS in 400 µL sucrose solution 1 M and expelled in sucrose 0.5 M 5 min Then two steps: sucrose 0.25 M 5 min sucrose 0 M Dilution into straw 50 µL VS in 400 µL galactose 1.7 M and expelled in galactose 0 M
Dilution into straw 50 µL VS in 150 µL sucrose solution 0.5 M and expelled in sucrose 0. 5 M 10 min Then one step: sucrose 0 M
Dilution into straw 50 µL VS in 150 µL sucrose solution 0.5 M and expelled in sucrose 0.5 M 10 min Then: One step: sucrose 0 M vs Four steps 10 min/step sucrose 0.38 M, 0.25 M, 0.13 M, 0 M
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4/7c
In vitro development after culture for 48 h 7 (1 with slight fracture of coatsf )/8b 6 (1 with slight fracture of coatsf )/8b 2 (1 with severe fracture of coatsf )/8b (diametric changes of embryos during addition of CPs measured too)
4/7a
D5 to D7 M to B Mean diam.= 169.3 µm
Dilution of cryoprotectants
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4/7a
0/7a
In vitro development after culture for 48 and 120 h
Device to vitrify embryo (warming)
February 2, 2011
Hochi et al., 199589
Hochi et al., 1994a10
References
Vitrification solution VS (addition of cryoprotectants)
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Embryo developmental stage
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Parameters of evaluation (results)
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Table 305.5 In vitro evaluations of vitrified-warmed equine embryos published from 1994 to 2007
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Castanheira et al., 200491 D6–6.5 M EB Diam. < 235 µm
D6 or D7 ø <300µm [150–300µm] mean ø = 211 µm
Mean in 7 embryos 48% live cells and grade 3.3 (+ 1 other embryo totally destroyed before staining)
Morphological evaluation at each step
D6 or D7 ø <300 µm [150–300µm] mean ø = 232 µm
Mean in 9 embryos 51% live cells and grade 3.1 (+ 2 other embryos totally destroyed before staining)
Open pulled straw (in medium of dilution at 37◦ C)
Cryoloop
Ethylene glycol 16.5% + DMSO 16.5% + sucrose 0.5 M (two steps: DMSO 7.5% + EG 7.5% 5 min; VS 20–30 s)
Ethylene glycol 17.5% + DMSO 17.5% + sucrose 1 M + Ficoll 0.25 µM (three steps: DMSO 7.5% + EG 7.5% 2.5 min; VS 20–30 s)
(waterbath at 37◦ C for 30 s)
0.25-mL plastic straw
Expelled in sucrose 0.25 M at 37◦ C 1 min Then two steps: sucrose 0.15 M 5 min sucrose 0 M 10 min Expelled in sucrose 0.33 M 2.5 min Then two steps: sucrose 0.17 M 3 min sucrose 0 M 5 min
Dilution into straw 50 µL VS in 150 µL sucrose solution 1 M + BSA 4% and expelled in sucrose 1 M + BSA 4% 10 min Then: one step: sucrose 0 M
Four steps 5 min/step Gal 0.6 M + 10% FCS Gal 0.4 M + 10% FCS Gal 0.2 M + 10% FCS Gal 0 M + 10% FCS
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Morphological evaluation at each step, and after culture for 20 h + propidium iodide and Hoechst 33342 staining to estimate (and not count exactly) percent live cells
DMSO 20%, ethylene glycol 20% + BSA 4 g/L + dextrose 0.375 M + sucrose 0.375 M (three steps: DMSO 10% + dextrose 0.125 M + sucrose 0.125 M + BSA 4% 5 min DMSO 10% + EG 10% + dextrose 0.25 M + sucrose 0.25 M + BSA 4% 5 min VS 1 min)
(air 12 s, waterbath at 37◦ C for 12 s)
0.25-mL plastic straw
16:40
Oberstein et al., 200185
D6.5 M, EB, B Diam. < 300 µm (162–279 µm) Mean diam. = 191 µm
Ethylene glycol 11.9 M + Ficoll 0.0026 M + galactose 0.3 M (two steps: EG 4.5 M + 10% FCS 15 min VS 1 min)
Dilution of cryoprotectants
February 2, 2011
All cells of six vitrified-warmed embryos were dead (+ 2 other embryos totally destroyed during warming) Comparison with fresh and frozen-thawed embryos (table 305.1)
Light microscopic observations of serial sections of embryos to assess the state (live, mitotic, dead) of each cell
Bruyas et al., 1999b27
D7–8 Diam. > 350 µm (300–680 µm); mean diam. = 409 µm
Device to vitrify embryo (warming)
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In vitro development after co-culture for 36 h and morphological grading 2/5d (+ 2 degenerated embryos after warming were not cultured)
Vitrification solution VS (addition of cryoprotectants)
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Young et al., 199774
Embryo developmental stage
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References
Parameters of evaluation (results)
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Table 305.5 (Continued)
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Ethylene glycol 18% + DMSO 18% + sucrose 0.4 M + glutamine 0.1% w/v + 20% FCS (two steps: EG 7.5% + DMSO 7.5% 3 min; VS 1 min) OPS straw (in air 2 s and in medium of dilution at 30◦ C)
Immediately expelled in: sucrose 0.13 M 1 min sucrose 0.13 M 5 min sucrose 0.075 M 5 min
Trim: 279mm X 216mm
M, morula; EB, early blastocyst; B, blastocyst; FCS, fetal calf serum; PVA, polyvinyl alcohol; PVP, polyvinylpyrrolidone. DAPI (dead cells were stained), 4 ,6-diamidino-2-phenylindole; BrDU, 5 -bromo-2 -deoxyuridine (incorporated during S phase into newly synthesized DNA strands). DMSO 18% v/v = DMSO 2.53 M; ethylene glycol 18% v/v = ethylene-glycol 3.22 M. a No. of normally developed embryos at 120 h/no. of vitrified-warmed embryos cultured. b No. of normally developed embryos at 48 h/no. of vitrified-warmed embryos cultured. c No. of embryos evaluated viable (no or few cells stained)/no. of vitrified-warmed embryos tested. d No. of normally developed embryos (1 grade 2, 1 grade 3) at 36 h/no. of vitrified-warmed embryos cultured. f Coats = zona pellucida and capsule.
D6.5 – 6.75 M, EB, B Diam. range: 140–270 µm Mean diam. = 187 µm Mean number of cells = 319
16:40
Morphological evaluation at each step, and after culture for 3 + DAPI staining to count dead cells + BrDU staining to count cells entering S phase for 1 h (index of mitotic activity) + Hoechst 33342 staining to count all cells Mean in 20 embryos 46% dead cells and 26% cells entering S phase 9/20: < 15% dead cells (mean 3% dead cells and 52% cells entering S phase) 10/20 > 40% dead cells (mean 81% dead cells and 6% cells entering S phase) 1/20 19% dead cells and 40% cells entering S phase
February 2, 2011
Moussa et al., 2005a30
(waterbath at 20◦ C for 20 s)
0.25-mL plastic straw
Dilution into straw 50 µL VS in 150 µL sucrose solution 0.5 M and expelled in sucrose 0.5 M 10 min Then one step: sucrose 0 M
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Mean in 6 embryos 19.2% live cells
Ethylene glycol 40% (two steps: EG 20% 10 min; VS < 2 min) Ethylene glycol 40% + Ficoll 18% w/v + sucrose 0.3 M (two steps: EG 20% 10 min; VS < 2 min) Ethylene glycol 40% + Ficoll 18% w/v + trehalose 0.3 M (two steps: EG 20% 10 min; VS < 2 min)
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Mean in 6 embryos 26.7% live cells
(170–235 µm)
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+ propidium iodide and Hoechst 33342 staining to assess percent live cells Mean in 6 embryos 0.8% live cells
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show published experiments using different methods to try to assess the viability of frozen-thawed or vitrified-warmed embryos: morphology of embryo grading before and after freezing-thawing; measurement of metabolic activities of embryos before and after freezing-thawing; staining dead or live cells of embryos after thawing; fixation of embryos after thawing and microscopic observations of sections with optical, electronic, or confocal microscopes. In all experiments, whatever test of viability was used and whatever cryopreservative protocols tested, a very large heterogeneity of intensity of cellular damage among embryos in the same group was observed. This large variability could be due to a difference of viability among fresh embryos at the time of collection and before addition of CP, or to differences among embryos regarding their intrinsic sensitivity to cryopreservation. In many other species there is a high stagedependent sensitivity of embryos to CP exposure, low temperatures, and cooling rate.18 The difference in sensitivity among embryos in the same group could be explained by the fact, demonstrated by Lagneaux and Palmer19 but mainly by Colchen et al.,20 that it is impossible to collect homogeneous groups of embryos at the same stage of development even after very precise supervision of ovulation in donor mares. Embryos collected exactly 156 ± 0.5 h after occurrence of ovulation measured from 162 to 245 µm in diameter and comprised from 117 to 417 cells; ones collected at 168 ± 0.5 h post-ovulation had from 272 to 2217 cells, with a variation of diameter from 159 to 365 µm.20 Sensitivity to cryopreservation is probably very different for an embryo with 200 cells compared with one with 1000 or 2000 cells. In practice or in experimental studies, the exact age of embryos is known with a precision of only 6 to 24 hours, even after induction of ovulation. Furthermore, the variability of the cellular composition of embryos is very large.21–32
Peculiarities of the horse embryo: difficulties in freezing Compared with embryos of other mammalian species, some peculiarities of horse embryos could explain the difficulties in cryopreserving them. Earlyblastocysts and blastocysts are much larger and composed of a greater number of cells than embryos of other species at the same developmental stages. This large size could cause their high sensitivity to cryopreservation and explain why successes were obtained only with the smallest embryos (diameter < 200–300 µm). Whereas ruminant blastocysts are composed of a mean of 100 cells, day 6.5 equine blas-
tocysts comprise a mean of about 400 to 600 cells.22, 33 This greater number of cells in equine blastocysts is partly due to a very short duration of the cell cycle in the embryonic cells, estimated to be only 6 h on day 7, whereas in most species it is between 14 and 24 h.20, 33 This intensive mitotic activity could increase the sensitivity of the equine embryo to cryopreservation, and hence cause the deleterious effects of the freezing process on the cytoskeleton.18, 31 Cells of equine early-blastocysts and blastocysts contain many large vesicles and copious lipids stored in variously sized droplets.22, 34 Mohr and Trounson35 explained that other species with embryo stages containing large and numerous cell vesicles were particularly difficult to cryopreserve. Cooling intolerance and difficulties of cryopreservation of porcine embryos were suspected to be due to their high lipid content.18 The capsule is another peculiarity of equine embryos. Studies28, 29 have suggested that the capsule may impede access of CPs to the embryo proper resulting in osmotic injury in embryonic cells and/or insufficient cryoprotection due to an inadequate concentration of CPs inside the embryos. In only four studies was there formal proof that some frozenthawed embryos have no capsule, either because they were produced totally in vitro,36–38 or based on microscopic observations of serial sections of early blastocysts there was no capsule found.28 In those four studies, most of the embryos survived or showed no cellular damage as fresh embryos (< 5% dead cells). The hypothesis was that there was a negative correlation between capsule thickness and freezability of equine embryos. This could explain why there is an apparent variability of intrinsic sensitivity to cryopreservation among embryos and why only small embryos survive cryopreservation. Embryos collected early after their arrival in uterus have either no capsule, an incomplete one or a very thin capsule. These are successfully frozen with glycerol, similar to bovine embryos.28 When the capsule is thicker the penetration of CP would be slower and reduced, thus increasing the osmotic injury due to dehydration of cells and decreasing the cryoprotection of cells during freezing or, probably, vitrification. Studies where variations of the embryo diameter were measured before, during, and after incorporation of CPs tend to confirm this hypothesis.10, 32, 39, 40 In the same way, a recent study41 demonstrated, using the Dual Chamber Vilia-Chien permeation System, (V-C systeme; Permegear Inc.; Hellertown, Pennsylvania) that the diffusion of CP in vitro across the capsule (obtained from day 14–16 embryos) was extremely low. The capsule could also be implicated in the sensitivity to cryopreservation of horse embryos in
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Freezing of Embryos another way: frozen-thawed and vitrified-warmed blastocysts were often observed with fractures in the embryonic coats, zona pellucida, and capsule. Damage of the capsule appeared to be detrimental to embryonic survival in vivo.42 What steps in cryopreservation are deleterious for horse embryos? Most studies have focused on CP and methods for its addition and removal. The toxicity of CPs on embryonic cells seemed to be a critical factor. Suspected osmotic toxicity of CP, mainly glycerol, initiated many empirical studies to try to evaluate the effect of increasing the penetration of CP inside embryos by using molecules having lower molecular weights than glycerol (MW 92) – in decreasing order of molecular weight: DMSO (dimethyl sulfoxide, MW 78); 1,2-propanediol (MW 76); ethylene glycol (MW 62); methanol (MW 32) – or by increasing the final concentration of CP or the time and/or the temperature of addition steps (see Tables 305.1 to 305.5). Another approach was to try to increase the permeability of the capsule by puncturing it (Pfaff cited by Seidel43 ) or by enzymatic partial digestion.31,44–46 Confusing results were obtained by those different approaches: 1. Increasing the permeability of the capsule seemed to induce good preservation of cellular viability during freezing but at the same time decreased embryonic survival in vivo because of loss of some properties of the capsule.31 2. CPs apparently less toxic than glycerol did not provide sufficient cryoprotection in some studies (100% dead cells in Bruyas et al.23, 28 ) and gave promising results in other ones (respectively 7/11 and 3/10 pregnancies in Hochi et al.47 and Huhtinen and Paranko.48 Cryoprotective agents can be toxic, as evidenced by microscopic observations of ultrastructure,22,49–51 which revealed deleterious effects on cellular organelles, mainly on mitochondria. More recently, disruption of the cytoskeleton has been shown by confocal microscopic observations of frozen-thawed horse blastocysts.31, 48 A recent study52 using fluorescent markers confirms the loss of mitochondrial activity and cytoskeletal disruption of all cells of large vitrified embryos and of inner cell mass (ICM) cells of smaller ones. The cryopreservation procedure with intracellular ice formation would be implicated31 more than a direct effect of CP on cellular actin. During cooling procedures dehydration of embryos occurs, but specific studies on cooling rates for freezing horse embryos are rare.
2917
The need for futures studies Seidel43 stated that studies were needed to understand and to improve technologies to store horse embryos. There is no doubt that horse embryos (morulae or early blastocysts) collected in the first hours after their arrival in utero can be successfully frozen with glycerol as proposed by Lacombe and Pashen9 or with ethylene glycol and direct transfer as performed by Hochi et al.47 Unfortunately, the precise time of uterine entry of embryos after ovulation is highly variable and not predictable.53–55 Shortly after the embryo enters the uterus synthesis of the capsule begins with a rapid increase in its thickness, and the mitotic activity of embryonic cells and the rate of embryonic development increase dramatically.20, 33 Even after hormonal induction of ovulation and a rigorous schedule of four examinations per day to detect ovulation followed by embryo recovery 156 hours postovulation, it was not possible to ensure that all recovered embryos had just arrived in utero and were at the correct stage of development for embryo freezing or vitrification.9, 19 Vitrification processes proposed by Eldridge-Panuska et al.56 seem to be a partial answer for storage of embryos smaller than 300 µm collected 8 days after the induction of ovulation by human chorionic gonadotropin (hCG). However, results based on a larger number of embryos are needed to validate this process. There is considerable variability among mares in the time elapsed between hCG injection and ovulation, and ovulation and entrance of the embryo into the uterus, and also variability among mares in stages of development and diameters of embryos collected 8 days after hCG injection. Thus there is a need to develop techniques to successfully cryopreserve blastocysts and even expanded blastocysts. Futures studies based not only on an empirical approach, as in the past, but also on a mathematical formulation should be conducted. Given that the stage-dependent sensitivity is in part due to variation in the permeability of different stage embryos to the same CP, approaches summarized by Leibo3 should be useful. For example, Woelders and Chaveiro57 have developed a theoretical model that calculates a non-linear freezing protocol in which at every subzero temperature the freezing rate is maximized. To do that, the membrane permeability coefficients for water and for CP, the respective activation energies, the aqueous volume of the embryo, and the membrane surface area must be measured, similar to recent reports for bovine in vitro-produced morulae.58 With such approaches, a cooling rate adapted to the species, the development stage, the size of embryo, and to each CP used could be calculated. In the same way a theoretical approach of vitrification
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proposed by Yavin and Arav59 would be useful to improve the process and may allow cryopreservation of large horse embryos more easily and successfully than smaller ones.
References 1. Thibier M. New records in the number of both in vivoderived and in vitro-produced bovine embryos around the world in 2006. IETS Newsletter 2007;25:15–20. 2. Merton S. Overall bovine embryo transfer activity in Europe. In: Proceedings of the 23rd Sientific Meeting of European Embryo Transfer Association, Alghero, 7–8 September, 2007; pp. 71–72. 3. Leibo SP, Cryopreservation of oocytes and embryos: optimization by theoretical versus empirical analysis. Theriogenology 2008;69:37–47. 4. Whittingham DG, Leibo SP, Mazur P. Survival of mouse embryos frozen to − 196◦ C and − 269◦ C. Science 1972;178:411–14. 5. Wilmut I, Rowson LEA. Experiments on the low temperature preservation of cow embryos. Vet Rec 1973;92:686–90. 6. Griffin JL, Castleberry RS, Schneider HS. Influence of day collection on recovery rate in mature cycling mares. Theriogenology 1981;15:106 (abstract). 7. Yamamoto Y, Oguri N, Tsutsumi Y, Hachinohe Y. Experiments in the freezing and storage of equine embryos. J Reprod Fertil Suppl 1982;32:399–403. 8. Vullers T. Recovery and freezing of embryos for use in com¨ mercial equine embryo transfer programme. In: Muller J, ¨ Muller Z, Wade JF (eds) Proceedings of the 3rd Meeting of the European Equine Gamete Group, Pardubice. Havemeyer Foundation Monograph Series No. 13. R & W Publications, 2004; pp. 54–6. 9. Lascombes FA, Pashen RL. Results from embryo freezing and postovulation breeding in commercial embryo transfer programme. In: Katila T, Wade J (eds) Proceedings of the 5th International Symposium on Equine Embryo Transfer, Saari. Havemeyer Foundation Monograph Series No. 3. R & W Publications, 2001; pp. 95–6. 10. Hochi S, Fujimoto T, Braun J, Oguri N. Pregnancies following transfer of equine embryos cryopreserved by vitrification. Theriogenology 1994a;42:483–8. 11. Rall WF. Factors affecting the survival of mouse embryos cryopreserved by vitrification. Cryobiology 1987;24:387– 402. 12. Carnevale EM, Eldridge-Panuska WD, Caracciolo di Brienza V. How to collect and vitrify equine embryo for direct transfer? In: Proceedings of the 50th Annual Convention of the American Association of Equine Practitioners, Denver. International Veterinary Information Service, 2004;P1472.1204. 13. Carnevale EM. Vitrification of equine embryos. Vet Clin N Am Equine 2006;22:831–41. 14. Thibier M. Stabilization of numbers of in vivo collected embryos in cattle but significant increases of in vitro bovine produced embryos in some part of the world (in 2003). IETS Newsletter 2004;22:12–19. 15. Thibier M. Significant increases in transfers of both in vivo-derived and in vitro-produced bovine embryos in cattle and contrasted trends in other species in 2004. IETS Newsletter 2005;23:11–17.
16. Rieger D. The measurement of metabolic activity as an approach to evaluating viability and diagnosing sex in early embryos. Theriogenology 1984;21:138–49. 17. Buttler JE, Biggers JD. Assessing the viability of preimplantation embryos in vitro. Theriogenology 1989;31:115– 26. 18. Dobrinsky JR. Advancements in cryopreservation of domestic animal embryos. Theriogenology 2002;57:285–302. 19. Lagneaux D, Palmer E. Non surgical recovery of morulae in the mare for freezing purposes using induction of ovulation. Theriogenology 1991;35:228 (abstract). 20. Colchen S, Battut I, Fi´eni F, Tainturier D, Siliart S, Bruyas J-F. Quantitative histological analysis of equine at exactly 156 and 168h after ovulation. J Reprod Fertil Suppl 2000;56:527–37. 21. Rieger D, Bruyas J-F, Lagneaux D, B´ezard J, Palmer E. The effect of cryopreservation on the metabolic activity of Day-6.5 horse embryos. J Reprod Fertil Suppl 1991;44:411– 17. 22. Bruyas J-F, B´ezard J, Lagneaux D, Palmer E. Quantitative analysis of morphological modifications of day 6.5 horse embryos after cryopreservation: differential effects on ICM and trophoblast cells. J Reprod Fertil 1993;99:15–23. 23. Bruyas J-F, Battut I, Pol J-M, Botrel C, Fi´eni F, Tainturier D. Quantitative analysis of morphological modifications of day 6.5 horse embryos after treatment with 4 cryoprotectants: differential effects on ICM and trophoblast cells. Biology of Reproduction Monograph series 1995;1:329–39. 24. Bruyas J-F, Marchand P, Fi´eni F, Tainturier D. The inability of DMSO to effectively cryoprotect day 6.5 horse embryos. Theriogenology 1997;47:387 (abstract). 25. Bruyas J-F, Martins-Ferreira C, Fi´eni F, Tainturier D. The effect of propanediol on the morphology of fresh and frozen equine embryos. Equine Vet J 1997;25(Suppl.):80–4. 26. Bruyas J-F, Gu´ena H, Battut I, Fi´eni F, Tainturier D. The inability of propanediol (3M) to cryoprotect horse embryos. In: Proceedings of the 15th Scientific Meeting of the European Embryo Transfer Association, Lyon, 1999; pp. 130–1 (abstract). 27. Bruyas JF, Menier D, Battut I, Fieni F, Tainturier D. An attmept to cryopreserve by vitrification Day 6.5 horse embryos. Reprod Domest Anim 1999;6(Suppl.):49–50. 28. Bruyas J-F, Sanson J-P, Battut I, Fi´eni F, Tainturier D. Effect of sucrose in diluting glycerol or ethylene glycol after thawing of frozen day 6.25 horse embryos. J Reprod Fertil Suppl 2000;56:549–60. 29. Legrand E, Bencharif D, Battut I, Taintureir D, Bruyas J-F. The effect of glycerol 1M on frozen horse embryos; localization of damaged cells. In: Proceedings of the 15th Scientific Meeting of the European Embryo Transfer Association, Lyon, 1999; pp. 182–3 (abstract). 30. Moussa M, Bersinger I, Doligez P, Guignot F, Duchamp G, Vidament M, Mermillod P, Bruyas J-F. In vitro comparisons of two cryopreservation techniques for equine embryos: slow-cooling and open pulled straw (OPS) vitrification. Theriogenology 2005;64:1619–32. 31. Tharasanit T, Colenbrander B, Stout TAE. Effect of cryopreservation on the cellular integrity of equine embryos. Reproduction 2005;129:789–98. 32. Duchamp G, Allard A, Grizelj J, Plotto A, Bruneau B, Mermillod P, Meriaux JC, Bruyas J-F, Vidament M. Effect of conditions of incorporation and concentration of cryoprotectants on equine embryo viability after freezing. Anim Reprod Sci 2006;94:374–7.
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Freezing of Embryos 33. Moussa M, Perreau C, Baril G, Duchamp G, Vidament M, Daels P, Bruyas J-F, Mermillod P. Comparison of cell proliferation index in equine and caprine embryos using a modified BrdU incorporation assay. Theriogenology 2005;64:1823–32. 34. Flood PF, Betteridge KJ, Diocee MS. Transmission electron microscopy of horse embryos, 3–16 days after ovulation. J Reprod Fertil Suppl 1992;32:319–27. 35. Mohr LR, Trounson AO. Comparative ultrastructure of hatched human, mouse and bovine blastocysts. J Reprod Fertil 1981;66:499–504. 36. Galli C, Crotti G, Turini P, Duchi R, Mari G, Zavaglia G, Duchamp G, Daels P, Lazzari G. Frozen-thawed embryos produced by Ovum Pick Up of immature oocytes and ICSI are capable to establish pregnancies in the horse. Theriogenology 2002;58:705–8. 37. Galli C, Colleoni S, Duchi R, Lagutina I, Lazzari G. Developmental competence of equine oocytes and embryos obtained by in vitro procedures ranging from in vitro maturation and ICSI to embryo culture, cryopreservation and somatic cell nuclear transfer. Theriogenology 2007;98:39–55. 38. Matsukawa K, Takahashi S, Adachi N, Akagi S, Hanada A. Effect of activation treatment for equine oocytes after ICSI and subsequent embryo’s freezability. Theriogenology 2003;59:454 (abstract). 39. Pfaff R, Seidel GE, Squires EL, Jasko DJ. Permeability of equine blastocysts to ethylene glycol and glycerol. Theriogenology 1993;39:284 (abstract). 40. Hochi S, Ogasawara M, Braun J, Oguri N. Influence of relative embryonic volumes during glycerol equilibration on the survival of frozen-thawed equine blastocysts. J Reprod Dev 1994;40:243–9. 41. Gillard Kingma SE, Betteridge KJ, Schlaf M, Gartley CJ, Chenier TS. Influence of equine embryonic capsule on diffusion of glycerol and ethylene glycol in vitro. In: Abstract Book of the 7th International Symposium on Equine Embryo Transfer, Cambridge, 2008; pp. 67–9 (abstract). 42. Stout TAE, Meadows SM, Allen WR. Stage-specific formation of the equine blastocyst capsule is instrumental to hatching and to embryonic survival in vivo. Anim Reprod Sci 2005;87:269–81. 43. Seidel GE. Cryopreservation of equine embryos. Vet Clin N Am Equine 1996;12:85–99. 44. Legrand E, Krawiecki JM, Tainturier D, Corni`ere P, Delajarraud H, Bruyas J-F. Does the embryonic capsule impede the freezing of equine embryos? In: Katila T, Wade J (eds) Proceedings of the 5th International Symposium on Equine Embryo Transfer, Saari. Havemeyer Foundation Monograph series no. 3. R & W Publications, 2001; pp. 62–65. 45. Legrand E, Bencharif D, Barrier-Battut I, Delajarraud H, Corni`ere P, Fi´eni F, Tainturier D, Bruyas J-F. Comparison of pregnancy rates for days 7–8 equine embryos frozen in glycerol with or without previous enzymatic treatment of their capsule. Theriogenology 2002;58:721–3. 46. Maclellan LJ, Carnevale EM, Coutinho da Silva MA, McCue PM, Seidel GE, Squires EL. Cryopreservation of small and large equine embryos pre-treated with cytochalasin-B and/or trypsin. Theriogenology 2002;58:717–20. 47. Hochi S, Maruyama K, Oguri N. Direct transfer of equine blastocysts frozen-thawed in the presence of ethylene glycol and sucrose. Theriogenology 1996;46:1217–24. 48. Huhtinen M, Paranko J. Confocal microscopic analysis of microfilament damage induced by cryopreservation of
49.
50.
51.
52.
53.
54.
55.
56.
57. 58.
59. 60.
61. 62.
63.
64.
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equine embryos. In: Allen WR, Wade JF (eds) Proceedings of the First Meeting of the European Equine Gamete GroN55–6. Wilson JM, Caceci T, Potter GD, Kraemer DC. Ultrastructure of cryopreserved horse embryos. J Reprod Fertil Suppl 1987;35:405–7. Landim e Alvarenga FC, Alvarenga MA, Meira C. Transmission electron microscopy of equine embryos cryopreserved by different methods. Equine Vet J 1993;15(Suppl.):67–70. Ferreira JCP, Meira C, Papa FO, Landin e Alvarenga FC, Alvarenga MA, Buratini J. Cryopreservation of equine embryos with glycerol plus sucrose and glycerol plus 1,2propanediol. Equine Vet J 1997;25(Suppl.):88–93. Hendriks WK, Stout TAE. Vitrification preferentially damages mitochondria in large equine embryos and the inner cell mass of small embryos. In: Abstract Book of the 7th International Symposium on Equine Embryo Transfer, 2008; p. 70 (abstract). Battut I, Colchen S, Fi´eni F, Tainturier D, Bruyas J-F. Success rates when attempting to non surgically collect equine embryos at 144, 156 or 168 hours after ovulation. Equine Vet J 1997;25(Suppl.):60–2. Battut I, Grandchmap des Raux A, Nicaise J-L, Fi´eni F, Tainturier D, Bruyas J-F. When do equine embryos enter the uterine cavity: an attempt to answer? In: Katila T, Wade J (eds) Proceedings of the 5th International Symposium on Equine Embryo Transfer, Saari. Havemeyer Foundation Monograph Series no. 3. R & W Publications, 2001; pp. 66–8. Maedows SM, Lisa H, Welsh C. Factors affecting embryo recovery, embryo development and pregnancy rate in a commercial embryo transfer programme. In: Allen WR, Wade JF (eds) Proceedings of the 1st Meeting of the European Equine Gamete Group, Lopuszna. Havemeyer Foundation Monograph Series no. 1. R & W Publications, 2000; pp. 61–2. Eldridge-Panuska WD, Caracciolo di Brienza V, Seidel Jr GE, Squires EL, Carnevale EM. Establishment of pregnancies after serial dilution or direct transfer by vitrified equine embryos. Theriogenology 2005;63:1308–19. Woelders H, Chaveiro A. Theoretical prediction of ‘optimal’ freezing programmes? Cryobiology 2004;49:258–71. Woelders H, Mullaart E, Merton S. Membrane permeability of IVP bovine morulae for water and glycerol: implications for freezing protocol. In: Proceedings of the 23rd Scientific Meeting of the European Embryo Transfer Association, Alghero, 7–8 September, 2007; p. 250 (abstract). Yavin S, Arav A. Measurement of essential physical properties of vitrification solutions. Theriogenology 2007;67:81–9. Allen WR. Embryo transfer in the horse. In: Adams CE (ed.) Mammalian Egg Transfer. Boca Raton: CRC Press, 1982; pp. 135–54. Yamamoto Y, Hachinohe Y. Frozen storage of equine embryos. J Fac Agr Hokka¨ıdo U 1984;62:182–210. Takeda T, Elsden RP, Squires EL. In vitro and in vivo development of frozen-thawed equine embryos. In: Proceedings of the 10th International Congress of Animal Reproduction and Artificial Insemination, Urbana, 1984;3:246. Slade NP, Takeda T, Squires EL, Elsden RP, Seidel GE. A new procedure for the cryopreservation of equine embryos. Theriogenology 1985;24:45–58. Czlonkowska M, Boyle MS, Allen WR. Deep-freezing of horse embryos. J Reprod Fertil 1985;75:485–90.
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65. Boyle MS, Allen WR, Tischner M, Clonkowska M. Storage and international transport of horse embryos in liquid nitrogen. Equine Vet J 1985;3(Suppl.):36–9. 66. Wilson JM, Kraemer DC, Potter GD, Welsh TH. Nonsurgical transfer of cryopreserved equine embryos to pony mares treated with exogenous progestin. Theriogenology 1986;25:217. 67. Farinasso A, De Feria C, Mariete ADaS, De Bem AR. Embryo technology applied to the conservation of equids. Equine Vet J 1989;8(Suppl.):84–6. 68. Squires EL, Seidel GE, McKinnon AO. Transfer of cryopreserved equine embryos to progestin-treated ovariectomised mares. Equine Vet J 1989;8(Suppl.):89–91. 69. Skidmore JA, Boyle MS, Allen WR. A comparison of two different methods of freezing horse embryos. J Reprod Fertil Suppl 1991;44:714–16. 70. Day WE, Evans JW, Vogelsang MM, Kraemer DC, Voelkel SA. Pregancy produced by a frozen/thawed equine early blastocyst using ethylene glycol, added and removed at 5◦ C, as cryoprotectant. In: Proceedings of the 13th Equine Nutrition and Physiology Symposium, 21–23 January, Gainesville, 1993; pp. 379–82. 71. Meira C, Alvarenga MA. Birth of foals from frozen embryos in Brazil. In: Proceedings of the 8th Reunido National da Sociedade Brasileine de T.E., 1993. 72. Meira C, Alvarenga MA, Papa FO, Oba E, Landim e Alvarenga FC. Cryopreservation of equine embryos using glycerol and 1,2 propanediol as cryoprotectants. Equine Vet J 1993;15(Suppl.):64–6. 73. Huhtinen M, Lagneaux D, Koskinen E, Palmer E. The effect of sucrose in the thawing solution on the morphology and mobility of frozen equine embryos. Equine Vet J 1997;25(Suppl.):94–7. 74. Young CA, Squires EL, Seidel GE, Kato H, McCue PM. Cryopreservation procedures for Day 7–8 equine embryos. Equine Vet J 1997;25(Suppl.):98–102. 75. Lagneaux D, Pomarici AM, Sattler M, Bruneau B, Duchmap G, Camillo F, Palmer E. Effect of L-glutamine for freezing equine embryos: evaluation by DAPI staining and transfer of multiple embryo to recipient mares. J Reprod Fertil Suppl. 2000;56:561–8. 76. Ulrich P, Nowshari MA. Erfolgreicher Direkttransfer eines tiefgefrorennen Embryos beim Pferd [Successful direct transfer of a frozen-thawed equine embryos]. Deutsche Tier¨arzliche Wochenschrift 2002;109:61–2. 77. Maclellan LJ, Bass LD, McCue PM, Squires EL. Effect of cooling large and small equine embryos prior to cryopreservation on pregnancy rates after transfer. Theriogenology 2003;59:306. 78. Bass LD, Denniston DJ, Maclellan LJ, McCue PM, Seidel GE, Squires EL. Methanol as cryoprotectant for equine embryos. Theriogenology 2004;62:1153–9. 79. Seidel GE, Squires EL, McKinnon AO, Long PL. Cryopreservation of equine embryos in 1,2-propanediol. Equine Vet J 1989;8(Suppl.):87–8. 80. Poitras P, Guay P, Vaillancourt D, Zidane N, BigrasPoulin M. In vitro viability of cryopreserved equine embryos following different freezing protocols. Canadian Vet J 1994;58:235–41. 81. Guay P, Poitras P. Preservation of equine embryos. Agbiotech News and Information 1989;1(4):515–18.
82. Barry BE, Thompson DL, White KL, Wood TC, Hehnke KE, Rabb MH, Colborn DR. Viability of inner cell mass versus trophectodermal cells of frozen-thawed horse embryos. Equine Vet J 1989;8(Suppl.):82–3. 83. Lagneaux D, Huhtinen M, Koskinen E, Palmer E. Effect of anti-freeze protein (AFP) on the cooling and freezing of equine embryos as measured by DAPI-staining. Equine Vet J 1997;25(Suppl.):85–7. 84. O’Donovan MK, Oberstein N, Bruemmer JE, Lane M, Seidel GE, Gardner DK, Squires EL. Metabolic activity of equine blastocysts cryopreserved in ethylene glycol and galactose. In: Katila T, Wade J (ed.) Proceedings of the 5th International Symposium on Equine Embryo Transfer, Saari, 6–9th July. Havemeyer Foundation Monograph Series no. 3. R & W Publications, 2000; pp. 57–9. 85. Oberstein N, O’Donovan MK, Bruemmer JE, Seidel GE, Carnevale EM, Squires EL. Cryopreservation of equine embryos by Open Pulled Straw, cryoloop or conventional slow cooling methods. Theriogenology 2001;55:607–13. 86. Hudson J, McCue P, Carnevale EM, Welsch S, Squires EL. The effects of cooling and vitrification of embryos from mares treated with equine follicle-stimulating hormone on pregnancy rates after non surgical transfer. J Equine Vet Sci 2006;26:51–4. 87. Hudson JJ, Hudson BD, Bailey JW, William S, Seagle C. Effect of polyvinyl alcohol (PVA) in vitrification of equine embryos. In: Proceedings of the Annual Conference of the Society for Theriogenology, St Paul, 22–26 August. Theriogenology 2006;66:664 (abstract). 88. Campos-Chillon LF, Cox TJ, Seidel GE, Carnevale EM. Vitrification in vivo of large equine embryos after vitrification or culture. Reprod Fert Develop 2006;18:151 (abstract). 89. Hochi S, Fujimoto T, Oguri N. Large equine blastocysts are damaged by vitrification procedures. Reprod Fert Develop 1995;7:113–17. ¨ 90. Chaves MG, Gonzalez EP, de Abreu Rosas C, Aguero A. Cryopreservation of equine embryos by two vitrification methods. Theriogenology 1997;47:388. 91. Castanheira PN, Amaral DCG, Vasconcelos AB, Arantes RME, Stahlberg R, Lagares MA. Cryopreservation of equine embryos by vitrification. In: Alvarenga M, Wade J (eds) Proceedings of the 6th International Symposium on Equine Embryo Transfer, Rio de Janeiro. Havemeyer Foundation Monograph Series no. 14. R & W Communications, 2004; pp. 50–2. 92. Lelong I., 2007. La cong´elation de l’embryon e´ quin: comparaison in vivo de la vitrification par la m´ethode OPS a´ la m´ethode de cong´elation lente. (In vivo comparison of two cryopreservation techniques for equine embryos: open pulled straw (OPS) vitrification and slow-cooling). DVM thesis, Toulouse National Veterinary School (ENVT), Toulouse, France. 93. Gaubert. La transplantation embryonnaire e´ quine: comments’ affranchir de la synchronization donneusereceveuse? (Equine embryo transfer: How to do without synchronization of oestrus cycles between donor and recipient mares?) Agricultural engineer thesis, Bordeaux National School of Engineers of Agricultural Technologies (ENITAB), Bordeaux, France, 1990.
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Chapter 306
Import and Export of Embryos Erin C. Bishop
The introduction of new genetics into a foreign country was previously achieved by shipping live animals from one continent to another. The technologies of cooling and freezing embryos have opened up opportunities for foreign horse owners. As a result of these technologies it is no longer necessary to risk injury or death of a horse as a result of long-haul shipping. An additional benefit is a saving on transport costs. There are several things to consider when assessing whether to import/export equine embryos or to transport the live animal:
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what are the advantages and disadvantages? should the embryo be cooled or frozen during transport? what is the process of shipping an equine embryo internationally? what are the import regulations for the destination country?
The import/export of equine embryos is a low-cost alternative to the quarantine and export of live animals. The length of time an animal must remain in quarantine prior to being transported is determined by the importing country. Typically, testing with negative results is required to be performed on the animal within a defined time frame prior to transport. Once a horse has been quarantined and the testing performed the animal is ready for transport. To ensure the safe arrival of the horse in the foreign country it is best to use a broker who is well versed in the process. Broker and transport fees can be as much as $9000 per animal when three horses are consolidated in one container and transported to the European Union. These fees may not include duties, taxes, and/or veterinary charges. The transport fees for shipping an embryo from one continent to another are considerably less than those for a live animal. A broker who is well versed in the process of exporting/importing embryos can
be a great asset to all involved. The broker can verify that the documents are in order, place the container with the embryo on a direct flight to an airport in a foreign country, and make sure the shipment clears customs and is delivered safely to the consignee. The cost of transporting an embryo from the USA to the European Union is approximately $1000. This fee can include air freight charges, approval of the health certificate by the US Department of Agriculture (USDA), and transfer of the container from the storage facility to the broker. When transporting equine embryos overseas there are many factors that can affect the success of the process. The breeding of the mare, timing of embryo recovery, method used to retard the growth of the embryo during transit, coordination of the embryo recipient for optimal transfer, and the reproductive health of the recipient mare are all important factors. The breeding of the mare and timing of embryo recovery are critical as is the method used to store the embryo during transport. With advances in technology the cooling and freezing of equine embryos have become common practice for many veterinarians. The cooling of equine embryos to 5◦ C places time constraints on the export process. There is a 24-hour window from the time when the embryo is collected and cooled to when it needs to be transferred into the recipient mare. Exporting/importing a cooled equine embryo takes precise timing for approval of the health papers, flight schedules, synchronization of the recipient mare and donor, as well as the eventual transfer of the embryo. The recommended method for preserving equine embryos for export/import is cryopreservation. Cryopreservation of equine embryos allows for longterm storage of valuable genetics. Embryos may be frozen using a slow-cool method (– 0.5◦ C/min) or by vitrification (see chapter 305). As long as the storage/ transport containers are properly maintained with
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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liquid nitrogen the embryos can be stored almost indefinitely. The USA does not have restrictions for the transport of equine embryos from state to state or for the importation of equine embryos from Canada. Many foreign countries have published requirements for the importation of cattle embryos but very few have available regulations for the import of equine embryos. Countries may have different regulations for importing equine embryos depending on the country of embryo origin. For example, the USA has different regulations for the import of embryos from countries affected with contagious equine metritis (CEM) compared to unaffected countries. The consignee in the importing country should contact their government to obtain an import permit, which will detail the requirements for the importation of equine embryos. Readers can visit the website of the USDA’s Animal and Plant Health Inspection Service for additional information (www.aphis.usda.gov). When importing equine embryos into the USA from a country affected with CEM a health certificate must be completed for the embryo and accompany it to the USA. It details: 1. the residency of the donor mare; 2. testing performed on the mare prior to embryo collection; 3. the processing of the embryo for transport; 4. the storage of the embryo prior to transport; 5. the health status of the donor stallion. The USA requires that the mare be resident at an approved European Union embryo collection facility for no less than 30 days prior to embryo collection. It must be certified that the facility is free of dourine, glanders, and equine infectious anemia (EIA) for no less than 6 months. The center veterinarian must certify that no animal at the collection facility has shown clinical signs of CEM during the 6 months prior to collection. The donor mare and donor stallion must not be used for natural service for no less than 60 days prior to collection; they must be isolated from animals considered to be of a lesser health status; and they must be tested for disease. The semen used to breed the mare must be from a donor stallion that underwent semen collection at an approved semen collection center and must meet the animal health requirements for the export of fresh/chilled/frozen semen to the USA. The disease testing required by the USA includes dourine and CEM. The processing conditions for importing equine embryos into the USA are: 1. The embryo must be processed in a facility that can be effectively cleaned following each embryo processing.
2. The media or solutions used for collecting, washing, and storing the embryos are free of pathogenic microorganisms. 3. The equipment used to collect and process the embryo are either new, disposable, or have been properly sterilized prior to use. 4. If fetal bovine serum or serum albumin are used they must undergo irradiation treatment and be from sources in Canada, the USA, Australia, or New Zealand. 5. The embryos must be washed using the International Embryo Transfer Society (IETS; www.iets.org) Manual protocol and an enzymatic (trypsin) treatment must not be used. 6. The zona pellucida must be intact and the embryo must be free of adherent material. The Terrestrial Animal Health Code (2007), Appendix 3.3.1 (www.oie.int/eng/normes/mcode/ code2007/en chapitre 3.3.3.htm), provides information on what can be done to limit the transfer of infection from the donor animal to the recipient and offspring. Although the appendix considers bovine embryos it is possible to extrapolate the principles for use with equine embryos. The code details three phases of risk management that determine the final level of risk for disease transfer. The phases are (i) to assess the potential for embryo contamination; (ii) to utilize recommended processing procedures as instructed in the IETS Manual; and (iii) to observe the donor animals after embryo collection for signs of disease and to test the embryo collection fluid for the presence of disease. In 2004, the IETS analyzed information on infectious diseases from previous research to determine the risk of transmission via in vivo derived embryos (www.oie.int/eng/normes/mcode/code2007/ en chapitre 3.3.5.htm). Once the data were analyzed they created four categories for these diseases. The categories are: 1. diseases or pathogenic agents that show negligible risk of contamination if embryos are collected and transferred properly; 2. diseases or pathogenic agents that show negligible risk of contamination if embryos are collected and transferred properly, but additional embryo transfers are needed to verify the existing data; 3. diseases or pathogenic agents that show negligible risk of contamination if embryos are collected and transferred properly but additional in vitro and in vivo research is required; 4. diseases or pathogenic agents for which the risk of transmission is not negligible even if the embryos are handled properly between collection and transfer or for which it is not possible to conclude the level of transmission for these diseases.
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Import and Export of Embryos There are two different kinds of containers available to ship a frozen embryo: a liquid shipping container or a vapor shipping container. The two may appear similar in structure but are manufactured differently. A liquid container must be classified as hazardous goods when in transit because it utilizes liquid nitrogen to maintain its temperature; this liquid can spill if the container is inadvertently tipped over. This method of transport is more costly and poses a greater risk to the embryo. If the container is not kept upright during transit the liquid nitrogen may spill out of the container and compromise the embryo due to an increase in container temperature. A vapor container is not considered a hazardous item and may even be placed on commercial passenger airlines for transport. A vapor shipping container utilizes an ab-
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sorptive material that is charged with liquid nitrogen. The excess liquid is removed from the container prior to shipment. A vapor shipping container can maintain the proper storage temperature for a week or more even if it is overturned during transport. Regardless of which type of container is used for transport the health certificate must certify that only embryos of the same health status were transported in the container and that the liquid nitrogen was not used for any other purpose. Although the import and export of bovine embryos is a regular practice, the international shipment of equine embryos is a more recent development. As the demand for foreign genetics increases so will the number of countries with published import requirements for equine embryos.
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Chapter 307
Nuclear Transfer Katrin Hinrichs
Introduction The first cloned equids were born in 2003. Currently more than 20 cloned horses have been produced, and some have in turn produced healthy offspring. The first equid produced by cloning was a mule resulting from nuclear transfer of cells obtained from a fetus.1 Two other mules were produced from the same cell line that year (2003),2 after surgical transfer of over 300 recombined oocytes to the oviducts of recipient mares. Also in 2003, a cloned horse foal was produced by the laboratory of Dr Cesare Galli in Italy, as a result of nuclear transfer using a donor cell from an adult mare.3 Embryos were cultured in vitro to the blastocyst stage, then transferred transcervically to recipient mares. Seventeen blastocysts were transferred to produce the one foal. An interesting twist in this report was that the donor cell used to produce the foal came from the mare that ended up carrying a foal to term, contradicting the hypothesis that pregnancies are more successful when the fetus is genetically different from the dam. No cloned foals were reported in 2004. In 2005, two cloned foals were reported from Dr Galli’s laboratory; one died and the other developed normally.4 Over 100 embryos were transferred to recipient mares to produce these foals. Also in 2005, the author’s laboratory at Texas A&M produced two cloned foals, after transfer of 11 embryos.5 From 2006 to 2008, the author’s laboratory produced an additional 12 live foals from the transfer of 43 blastocysts (28% live foal rate;6, 7 ). Two of these foals died and the remaining 12 are healthy and are developing normally. This rate of live foal production (28%) compares favorably with that for bovine cloning (6% of transferred embryos produce live calves8 ). Starting in 2006, a commercial company, ViaGen (Austin, Texas, USA), has announced the birth of live cloned foals in the popular press, although information on the number of
embryos transferred or total foals born has not been given. A major blow to research in equine cloning and production of cloned foals in the USA occurred in 2007, with the closure of the horse slaughterhouses in this country. Currently, there is no ready source of horse oocytes within the USA with which cloning can be performed. The commercial company working with horses has set up a laboratory in Canada to make use of oocytes available in that country.
Technique To perform cloning, the nucleus of a cell from the donor animal is transferred into the cytoplasm of an oocyte, and the oocyte is signaled to develop into an embryo. The cells from the donor animal are typically grown from a tissue biopsy sample. To obtain this, the hair is shaved from an area approximately 7 × 7 cm, and the skin is scrubbed and prepared as for surgery. We typically do this under the mane, for cosmetic reasons. A small skin incision (about 2 cm) is made, and the skin edges are everted. Small (∼5 mm3 ) samples of subcutaneous connective tissue are obtained with scissors and placed immediately into cooled cell culture medium or embryo holding medium. The medium is kept cold and is transported as quickly as possible (overnight delivery is acceptable) to the laboratory. Once at the laboratory, the tissue is placed into culture in medium in a 5% CO2 incubator, and fibroblasts grow from it onto the floor of the culture dish. Once the fibroblasts have proliferated to cover the bottom of the plate (grown to confluence) they are resuspended in medium and used to seed additional dishes. Each occasion on which the cells are divided and seeded again is termed a ‘passage’. After a
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Nuclear Transfer sufficient number of cells are obtained in this manner, the cells are typically frozen. In our laboratory, we typically freeze 20 straws of 1 million cells each. When the donor cells are prepared for use in nuclear transfer, they are cultured, then induced into a resting or early stage of the cell cycle, i.e., G0 or early G1. This may be accomplished by a variety of methods, including serum deprivation, growth of the cells to confluence, or treatment with a cell-cycle inhibitor such as roscovitine. Oocytes used for cloning must be mature (in metaphase II). The genetics of the oocyte is unimportant, with the possible exception of the mitochondria, as discussed below. The mature oocyte is enucleated by removing the metaphase plate and polar body using a small-diameter pipette via micromanipulation. The donor cell is then combined with the enucleated oocyte, either by electrofusion or by directly injecting the cell into the cytoplasm of the oocyte. The recombined oocyte is activated to stimulate embryonic development; this is typically done by triggering calcium oscillations within the oocyte that mimic those that occur at fertilization. Once the recombined oocyte has been activated, it may be transferred surgically to the oviduct of a recipient mare, or may be cultured in vitro to the blastocyst stage for transfer directly to the uterus of a recipient mare, as for standard embryo transfer.
Health status of cloned pregnancies and foals Loss of cloned pregnancies may occur throughout gestation. Overall in our laboratory, we have established 31 cloned pregnancies from transfer of 54 embryos.9 Of these, 16 (52%) have been lost during gestation. Nine embryos were lost before 100 days, three between 100 and 200 days, and four after 200 days. Five of these pregnancy losses (all early) were from one cell line that failed to produce a viable pregnancy; a second tissue sample obtained from the same horse provided two live foals from the transfer of four embryos.7 The only definable problems that we have noted in spontaneously aborted cloned fetuses are contracted front legs. Of the 17 live cloned horse foals that have been reported in the scientific literature, as noted above, three foals have died postpartum. These foals died from septicemia, pneumonia, and cerebral hypoxia after a hypotensive episode under anesthesia. These data suggest that cloned foals may not suffer to the same degree from problems that are seen in cloned offspring of other species. The three mule foals were also healthy, but since they were produced from a cell line obtained from a 45-day gestation fetus,
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which is more likely to produce viable offspring than is an adult cell, their condition may not relate directly to what could be expected of an animal cloned from the cells of an adult horse. The main problems seen in term, live cloned foals have included crooked or contracted front legs, large umbilical remnant – which has led to its surgical removal in some foals – and varying degrees of neonatal maladjustment.9 Half or more of cloned foals show these signs to a variable extent. Fortunately, these problems have been self-limiting or have responded well to treatment. In other species, notably sheep and cattle, the large majority of cloned pregnancies are lost during gestation, and cloned offspring may suffer from fetal overgrowth and abnormalities of the heart, lung, and other organs.10 Abnormalities seen in cloned offspring are thought to be related to the failure of the DNA of the donor cell to be ‘reprogrammed’ by the oocyte cytoplasm. In the donor fibroblast, only those genes needed for fibroblasts to function are being used; all other genes are turned off. Inhibition of gene function is obtained largely by modification of the DNA base molecules and of the histones, the proteins around which the DNA is wrapped. These modifications, which change the utilization of the gene without changing the gene itself, are termed epigenetic modifications. The oocyte is able to change the epigenetic markings of the transferred chromatin from those of a fibroblast to those compatible with growth of an embryo; however, this process is not always performed successfully, and if gene function is not compatible with development, the fetus or neonate cannot survive. A common question relating to the health of cloned foals is whether they will age prematurely. This thought stems from publicity around Dolly, the sheep that was the first clone of an adult mammal. Examination of Dolly’s cells showed that the telomeres, the non-coding regions on the tips of the chromosomes, were shorter than normal for her age. Telomeres lengthen during gamete and early embryo development, then shorten with each subsequent cell division; thus they are long in young animals and shorter in older animals. The effect of these short telomeres in Dolly’s case is unknown, as Dolly was euthanized at 6 years of age, along with many others in her flock, after contracting a viral lung tumor. The condition of short telomeres in cloned animals was later related to the source of the cells used for cloning; mammary gland cells, as were used to produce Dolly, were found to result in calves with short telomeres whereas fibroblasts resulted in calves with normal-length telomeres.11 The normality of telomere length in animals derived from fibroblasts has since been confirmed in several species.12–14
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How closely will the clone resemble the donor? A variety of factors will affect the degree of similarity between the cloned offspring and the original donor. Only two of these are actually related to the cloning procedure: first, epigenetic changes may cause the gene utilization in a cloned foal to differ somewhat from that of the donor, potentially resulting in production of a larger or smaller quantity of specific proteins, which may in turn affect phenotype. More importantly, epigenetic changes may detrimentally affect the function of the placenta in cloned pregnancies, potentially resulting in a small or maladjusted neonate. As with naturally occurring twin foals, effects of placental insufficiency may be seen into adulthood. The second cloning-related potential cause of differences between the clone and the donor animal is related to mitochondria. The vast majority of mitochondrial function is governed by genes in the nuclear DNA (about 3000 genes), but in addition, mitochondria also have their own small DNA molecule coding for about 13 proteins and some functional nucleic acids. The embryo is formed by division of the oocyte cytoplasm, thus the mitochondria of the host oocyte used for cloning become the mitochondria of the offspring, with perhaps some variable contribution from the mitochondria from the donor fibroblast. Therefore the mitochondrial genotype of the cloned foal will differ from that of the donor animal. It is unknown whether this will produce any differences in phenotype. Additional reasons for variability in phenotype are not directly related to cloning. If the cloned embryo was cultured in vitro before transfer to the recipient mare, in vitro culture itself has been shown to cause differences in neonatal size and other phenotype characteristics in other species. However, little information is available on the effects of in vitro embryo culture in the horse, because very few foals have been produced from in vitro-cultured embryos. Other factors affecting phenotype are present for any transferred embryo, but the effects are more vivid in cloned foals because the expected phenotype is known. One of these is the embryonic environment, including uterine and placental health and size, problems during gestation or birth, dam and foal nutrition, handling and exercise regimens, vaccination and deworming programs, and training. In addition, individual differences arise during fetal growth, a vivid example of which is differences in white markings. A signal for white is genetically determined, but the extent of the white varies with the individual. We have seen cloned foals from a single genotype
have one, two, and three white socks, and have variations in face markings ranging from a broad blaze to a star, stripe and snip. It should be expected that many genes coding for proteins functioning throughout the body may be expressed more or less in one foal, just through individual variation, than in another with the same genotype.
How closely will the progeny of cloned animals resemble those of the donor? Herein lies the strength of cloning as a clinical procedure. While no information is yet available in this area in the horse, in all other species studied, even if the cloned animal itself has some epigenetic differences from the original donor, the clone’s progeny are normal. The epigenetic status of the DNA is reset when gametes are formed; thus, the cloned animal’s offspring should be no different from the offspring of the original animal. This is especially true in the case of stallions, as even the possibility of mitochondrial differences is eliminated. A cloned stallion will possess the mitochondrial genome from the oocyte used to produce it, and this genome will be present in mitochondria in its sperm. However, the sperm mitochondria are eliminated from the embryo after fertilization. For this reason, the progeny of a cloned stallion should not differ from the progeny the original horse would have produced. In the case of a cloned mare, the clone will produce oocytes carrying its mitochondrial genome, and thus its foals will also have this mitochondrial genome. In this way, they will differ from foals produced by the original mare. Again, it is unknown whether differences in mitochondrial genome will translate into differences in phenotype.
Is cloning clinically feasible? While earlier work in cloning, as noted above, was inefficient (i.e., one foal from transfer of 100 embryos4 ), more recent work has shown a relatively high efficiency, given an adequate supply of oocytes. For the foals our laboratory produced in 2006, over all treatments we had a 6% blastocyst development from recombined oocytes; with up to 50 MII oocytes available per week, this resulted typically in one to three embryos ready for transfer per week. Transfer of 26 embryos resulted in the nine live foals born (seven surviving). For foals produced in 2007 and 2008, transfer of seven embryos, resulting from manipulation of only 75 oocytes, yielded birth of three live, viable foals. However, efficiency may vary among
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Nuclear Transfer laboratories. Because commercial laboratories typically do not report their results in the scientific literature, the efficiency of foal production by such companies, or the proportion of produced foals surviving, is unclear. However, cloned foals are being successfully produced commercially from specific cell lines for private horse owners.
Acknowledgements Work done in the author’s laboratory was supported by the Link Equine Research Endowment Fund, Texas A&M University, Cryozootech SA, and the Smart Little Lena co-managers.
6.
7.
8. 9.
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References 1. Woods GL, White KL, Vanderwall DK, Li G-P, Aston KI, Bunch TD, Meerdo LN, Pate BJ. A mule cloned from fetal cells by nuclear transfer. Science 2003;301:1063. 2. Vanderwall DK, Woods GL, Sellon DC, Tester DF, Schlafer DH, White KL. Present status of equine cloning and clinical characterization of embryonic, fetal, and neonatal development of three cloned mules. J Am Vet Med Assoc 2004;225:1694–9. 3. Galli C, Lagutina I, Crotti G, Colleoni S, Turini P, Ponderato N, Duchi R, Lazzari G. A cloned horse born to its dam twin. Nature 2003;424:635. 4. Lagutina I, Lazzari G, Duchi R, Colleoni S, Ponderato N, Turini P, Crotti G, Galli C. Somatic cell nuclear transfer in horses: effect of oocyte morphology, embryo reconstruction method and donor cell type. Reproduction 2005;130:559–67. 5. Hinrichs K, Choi YH, Love CC, Chung YG, Varner DD. Production of horse foals via direct injection of roscovitine-
11.
12.
13.
14.
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treated donor cells and activation by injection of sperm extract. Reproduction 2006;131:1063–72. Hinrichs K, Choi YH, Varner DD, Hartman DL. Production of cloned horse foals using roscovitine-treated donor cells and activation with sperm extract and/or ionomycin. Reproduction 2007;134:319–25. Choi YH, Hartman DL, Fissore RA, Bedford SJ, Hinrichs K. Effect of sperm extract injection volume, injection of PLCζ cRNA, and tissue cell line on efficiency of equine nuclear transfer. Cloning Stem Cells 2009;11:301–8. Wells DN. Animal cloning: problems and prospects. Rev Sci Tech 2005;24:251–64. Johnson AK, Clark-Price SC, Choi YH, Hartman DL, Hinrichs K. Physical and clinicopathologic findings in foals produced by somatic cell nuclear transfer – 14 cases (20052008). J Am Vet Med Assoc 2010;236:983–90. Chavatte-Palmer P, Remy D, Cordonnier N, Richard C, Issenman H, Laigre P, Heyman Y, Mialot JP. Health status of cloned cattle at different ages. Cloning Stem Cells 2004;6:94–100. Miyashita N, Shiga K, Yonai M, Kaneyama K, Kobayashi S, Kojima T, Goto Y, Kishi M, Aso H, Suzuki T, Sakaguchi M, Nagai T. Remarkable differences in telomere lengths among cloned cattle derived from different cell types. Biol Reprod 2002;66:1649–55. Betts D, Bordignon V, Hill J, Winger Q, Westhusin M, Smith L, King W. Reprogramming of telomerase activity and rebuilding of telomere length in cloned cattle. Proc Natl Acad Sci USA 2001;98:1077–82. Jiang L, Carter DB, Xu J, Yang X, Prather RS, Tian XC. Telomere lengths in cloned transgenic pigs. Biol Reprod 2004;70:1589–93. Jeon HY, Hyun SH, Lee GS, Kim HS, Kim S, Jeong YW, Kang SK, Lee BC, Han JY, Ahn C, Hwang WS. The analysis of telomere length and telomerase activity in cloned pigs and cows. Mol Reprod Dev 2005;71:315–20.
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Section (b): Oocytes
308.
Immature Oocyte Collection and Maturation Katrin Hinrichs
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Mature Oocyte Collection Elaine M. Carnevale
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Oocyte Transfer Elaine M. Carnevale
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Gamete Intrafallopian Transfer (GIFT) Marco A. Coutinho da Silva
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Intracytoplasmic Sperm Injection (ICSI) Young-Ho Choi and Katrin Hinrichs
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Oocyte Cryopreservation Lisa J. Maclellan
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Chapter 308
Immature Oocyte Collection and Maturation Katrin Hinrichs
Introduction Production of foals by assisted reproductive techniques including in vitro fertilization (IVF), intracytoplasmic sperm injection (ICSI), and oocyte transfer requires the recovery of oocytes from the donor mare. The technique of nuclear transfer (NT; cloning) requires oocytes to serve as host cytoplasts for the transfer of somatic cell nuclei. To be used for production of foals, oocytes must be mature, that is, ready to respond to an activation signal and develop into an embryo. In vivo, oocyte maturation occurs within the dominant preovulatory follicle in the 24 to 36 hours immediately preceding ovulation, after the appropriate luteinizing hormone (LH) stimulus. Oocyte maturation includes both nuclear maturation – resumption of meiosis and progression to metaphase II – and cytoplasmic maturation, a series of changes that prepare the cytoplasm for development into an embryo. One method of obtaining mature oocytes is to aspirate the preovulatory follicle ex vivo, after induction of maturation with human chorionic gonadotropin (hCG) or a gonadotropin releasing hormone (GnRH) analog (see Chapters 188, 190 and 309). This is the most common method by which oocytes are obtained for commercial ICSI or oocyte transfer. In some situations, however, such as in recovery of oocytes from the ovaries of a mare post mortem, it is not feasible to collect in vivo-matured oocytes. In these cases, oocytes must be obtained from the smaller, non-ovulatory follicles present on the ovaries. These include both growing and degenerating (atretic) follicles. The oocytes within these follicles are not mature; they are resting in prophase I of meiosis. To be useful for assisted reproduction, these oocytes must be matured in vitro.
Collection of immature oocytes from live mares Immature oocytes may be obtained from small follicles in live mares by ultrasound-guided follicle aspiration as used for recovery of oocytes from the preovulatory follicle (method described in Chapter 309); however, the physiology of the mare makes this a relatively inefficient procedure. Mares have an average of only seven visible follicles on each ovary. The immature oocyte of the mare is very closely adhered to the follicle wall,1 thus aspiration of small follicles is associated with a low recovery rate (typically < 25%). With practice, this may be increased to over 50% recovery;2 follicles should be emptied and filled repeatedly, while the needle is rotated to ‘scrape’ the interior of the follicle as it collapses. Sessions for aspiration of immature oocytes should be spaced at least 10–14 days apart, as more frequent aspiration resulted in a linear decrease in the number of available follicles per mare, and also in apparent reaspiration of previously aspirated, refilled follicles.3
Collection of oocytes from excised ovaries Oocytes may be easily collected from excised ovaries in the mare. Shipped ovaries should be processed within 6 h of the death of the mare. Oocytes from ovaries stored longer, up to about 15 h post mortem, may mature in vitro, but the rate of embryonic development will be compromised.4, 5 If the ovaries are processed within an hour or two of collection, they may be stored at 37◦ C. For longer periods, holding at room temperature is recommended. This can be achieved during shipment by placing the
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R ovaries in a plastic bag in an Equitainer , with roomtemperature ballast and the coolant cans at room temperature. Because of the firm attachment of the immature equine oocyte to the follicle wall, simple aspiration of the follicular contents with a needle and syringe, as done in cattle, results in a low recovery rate.6 Efficient collection of equine oocytes requires opening the follicle and scraping free the granulosa cell layer. To do this, excess tissue is trimmed from the ovary to aid handling. Follicles are located on the surface of the ovary visually or by palpation. The first follicle is opened with a scalpel blade so that it is completely bisected and both halves are visible. Fluid will be lost as the follicle is opened, but is unlikely to contain the oocyte (those oocytes not firmly attached to the follicle wall are likely to be degenerating). The wall of the follicle is scraped with a 0.5-cm bone curette in overlapping swaths, until all of the wall has been scraped. Larger or smaller curettes may be used for differently sized follicles. The tissue accumulated in the curette is washed into a 35-mm Petri dish using a stream of medium (below) from a 20-gauge needle attached to a 10-mL syringe. Typically only one swath is made and then washed into the dish, as cells may be lost if multiple swaths are made at a time. The medium used for oocyte collection in our laboratory is Hepesbuffered TCM199 with Hanks’ salts (Gibco Life Technologies Inc., Grand Island, NY) with 25 µg/mL gentamicin added (Gibco Life Technologies Inc.); embryo holding medium may be used if other media are not available. Note that all materials used in media that will be in contact with the oocytes should be of embryo grade; materials compounded for medical use may contain contaminants or preservatives (as for multiple-dose vials) that may harm oocytes. Once the opened follicle has been scraped completely, the next follicle is opened and similarly processed. The contents of each follicle are washed into a separate dish. Once all of the visible follicles on the surface of the ovary have been opened and scraped, the ovary is sliced in 0.5-cm slices using the scalpel blade. All follicles visible in the slices are similarly scraped, no matter how small. The Petri dishes are examined under a dissection microscope at ×10 to ×40. Oocytes are classified as compact (Cp) or expanded (Ex) depending upon the character of the granulosa. Compact granulosa appears as sheets of tissue having an even surface, which may be thrown into folds by the scraping action of the curette. This tissue, as seen on the surface of a fold or on the hillock covering the oocyte, is smooth without evidence of protruding cells (Figure 308.1). Any granulosa not in smooth sheets or flat shards is considered to be expanded. This can range from having some cells protruding from
Figure 308.1 Compact oocyte–cumulus complex tipped on its side with a needle, showing the smooth surface of the granulosa layer (cumulus hillock). Bar = 200 µ m.
a sheet, to sheets with a thickened or puffy granular appearance, to clouds of expanded cells (Figure 308.2). Very expanded granulosa may be associated with yellow jelly-like substance. When the oocyte is located, the cumulus surrounding the oocyte is similarly classified. Only those oocytes having both compact cumulus and compact granulosa are classified as Cp. The compact oocyte–cumulus complex is typically shaped like a flat-brimmed hat, with a flat underside and a bulging
Figure 308.2 Expanded oocyte–cumulus complex. Bar = 200 µ m.
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Immature Oocyte Collection and Maturation upper surface covered by a ‘hillock’ of cumulus (Figure 308.1). Compact granulosa originates from viable follicles, whereas expanded granulosa originates from atretic follicles.7 In other species, oocytes having expanded granulosa at the time of collection are discarded as degenerating. However, in horses both oocytes are valuable. Oocytes in growing equine follicles appear to achieve normal competence to mature when the follicle reaches around 20 mm diameter.8 As most follicles present on non-stimulated ovaries are smaller than 20 mm, most Cp oocytes recovered are juvenile. The chromatin in juvenile Cp oocytes is labile and degenerates rapidly during oocyte storage. Thus, the maturation rate of Cp oocytes collected from excised, shipped ovaries is low (about 20–30% in our laboratory9, 10 ). If oocytes are recovered rapidly post mortem (within 1 h), the rate of maturation of Cp oocytes is higher, but the ability of these ‘rescued’ oocytes to form blastocysts is low.10 In contrast, changes in the follicle undergoing atresia (degeneration) mimic some of those occurring within a mature follicle. The most evident is the expansion of the granulosa; in addition, the degenerating follicle allows the enclosed oocyte to acquire competence to mature. In severely atretic follicles, the enclosed oocyte may even start to undergo meiosis as it starts to degenerate.7 Oocytes from follicles in early to mid-stage atresia have slightly to extensively expanded granulosa and have a high degree of meiotic competence. In our laboratory, oocytes classified as Ex have a 65% maturation rate.9, 10 We find that approximately 30% of collected oocytes are classified as Cp, 60% as Ex, and 10% as degenerating (have shrunken, dense or misshapen cytoplasm and typically are losing or have lost the cumulus – these are discarded). The cytoplasm of horse oocytes, especially of oocytes that are more meiotically competent (desirable), is often segregated into light and dark areas, which may give the oocyte a misshapen appearance when viewed under lower power on the microscope. Under higher power, the outline of the oocyte plasma membrane shows the spherical nature of the oocyte. Note that whereas in cattle, oocytes are selected on the basis of having a homogenous ooplasm, in horses this trait is associated with a lower meiotic competence.7 After an oocyte has been found and classified according to its cumulus morphology, it is removed and placed in a holding dish (one each for Cp and for Ex) containing 3 mL of the collection medium. If necessary, oocytes are cut from the granulosa cells using needles. Care must be taken when pipetting oocyte–cumulus complexes to avoid loss of the occyte on the wall of the pipette – always ensure that
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there is a column of fluid already in the pipette before the oocyte is picked up. The holding dish is kept at room temperature (22–25◦ C) until all oocytes have been collected; this may be hours if many ovaries are processed. After collection, it may be useful to hold the oocytes overnight before placing them in maturation medium, to allow scheduling of oocyte transfer or ICSI during the day.11 The oocytes are placed in a 1-mL borosilicate glass vial containing 1 mL of a medium comprised of 40% M199 with Hanks’ salts, 40% M199 with Earle’s salts (Gibco) and 20% fetal bovine serum (FBS; Gibco). The vial is covered with aluminum foil, and placed on its side at room temperature. Separate vials are used for Ex and Cp oocytes.
Culture for maturation For culture in our laboratory, we use a simple combination of M199 with 10% FBS and 5 mU/mL follicle-stimulating hormone (FSH) (Sioux Biochemical Co., Sioux City, IA) with 25 µg/mL gentamicin. Other additives or media are used in other laboratories, but none has provided higher blastocyst rates than that achieved in our laboratory (25–30%) with this medium. Oocyte maturation is performed in a 5% CO2 atmosphere to mimic the CO2 levels in the body. The media will absorb the CO2 ; if not appropriately buffered, they will become acidic (CO2 + H2 O → H+ + HCO3 − ). Therefore, media used for culture are buffered to maintain pH in 5% CO2 with a bicarbonate buffer system, which is termed Earle’s salts. In contrast, a phosphate buffer system is used to maintain pH in room atmosphere. Culture medium M199 comes with either salts. When oocytes are being collected from ovaries, medium with Hanks’ salts (phosphate buffer) is used. When oocytes are cultured in the CO2 incubator, medium with Earle’s salts is used. An additional buffer commonly used in either of these media is Hepes, which is a zwitterionic buffer that maintains pH without regard to the CO2 atmosphere. It may be possible to perform oocyte maturation in phosphate-buffered media in a room atmosphere incubator, but more study is needed in this area. The maturation medium should be prepared prior to oocyte recovery, and equilibrated in a 5% CO2 incubator for at least 2 hours before it is utilized. All the media noted herein contain the pH indicator phenol red, to help determine the pH status of the media. Thus media are brick red at pH 7.4, yellow when acidic, and purple when basic. Oocytes should not be placed in medium if it is not the appropriate color. Oocytes are typically matured in droplets of medium under light white mineral oil (Sigma,
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St Louis, MO) but may also be matured in 35-mm Petri dishes containing 3 mL of medium. If microdroplets are used, three 150-µL droplets (two wash droplets and the maturation droplet) are placed in each Petri dish. To prepare the microdroplets, three drops of 75 µL each are placed on the floor of a 35mm Petri dish using a calibrated pipetter or an allplastic tuberculin syringe (Air-Tite, Virginia Beach, VA, USA) and needle. The drops are then completely covered with equilibrated light white mineral oil. After the droplets are covered, an additional 75 µL of medium is added to each droplet, resulting in a droplet that is rounder than if all 150 µL was added at once. The droplets are incubated for 2 or more hours to equilibrate the medium. The oocytes are transferred to one of the wash droplets using a fine pipette, then to the next wash droplet, then to the final maturation droplet. They are cultured at a ratio of one oocyte per 10 µL medium (e.g., 15 oocytes per 150-µL droplet). Culture is performed for 24–30 h for Ex oocytes, and for 30–36 h for Cp oocytes.10
Evaluation for maturity After the maturation period, the oocytes are evaluated under the dissection microscope and any obviously degenerating oocytes are removed. The cumulus of both Ex and Cp oocytes should be expanded after culture. Evaluation for maturation requires that the cumulus be denuded, to enable visualization of the polar body. The cumulus must also be denuded to conduct ICSI or cloning procedures, and partially denuded for IVF attempts. Oocytes to be used for oocyte transfer are left cumulus intact, to increase their adherence to the oviduct, thus their maturation status is unknown at the time of transfer. Cumulus removal is performed in a solution of 0.05% hyaluronidase in holding medium (Hanks’ buffered TCM199). Use of droplets of medium, rather than a whole dish, will limit oocyte mobility. Glass Pasteur pipettes are pulled over a flame to provide a variety of pipette bore diameters. The oocytes are denuded by pipetting them with increasingly smaller diameter pipettes. The smallest diameter pipette should slightly deform the zona-enclosed oocyte, thus scraping the cumulus cells from the zona. Removal of the cumulus of horse oocytes is more difficult than in oocytes of other species, because of extensive connective processes of the cumulus cells through the zona. An alternative medium for cumulus removal is a solution of 0.25% trypsin and 1 mM ethylene diamine tetra-acetic acid (EDTA) in Hanks’ salts without CaCl2 , MgCl2 or MgSO4 (Gibco). This solution
Figure 308.3 Metaphase II horse oocytes after maturation in vitro and denuding of the cumulus. A polar body is visible in oocytes in the correct orientation (arrowheads). Note the segregated areas of light and dark cytoplasm; this indicates maturity in equine oocytes. Bar = 200 µ m.
disrupts cell junctions and makes cumulus removal easier, especially in immature oocytes, but its effect on embryo development has not been evaluated. Once the cumulus is removed, the oocyte is observed at ×40 to ×100 for a polar body (Figure 308.3). Rolling the oocyte with a needle may be necessary. If complete cumulus removal was not possible, so that the polar body is not visualized, the presence of a smooth oocyte plasma membrane is suggestive of maturation. The mature horse oocyte usually has distinct light and dark areas in the cytoplasm and a noticeable perivitelline space (Figure 308.3). If oocytes are to be sacrificed, stage of meiosis may be ascertained by staining with aceto-orcein or with the fluorochrome Hoechst 33258 and visualization under fluorescence microscopy.12
References 1. Hawley LR, Enders AC, Hinrichs K. Comparison of equine and bovine oocyte-cumulus morphology within the ovarian follicle. Biol Reprod Monogr 1995;1:243–52. 2. Colleoni S, Barbacini S, Necci D, Duchi R, Lazzari G, Galli C. Application of ovum pick-up, intracytoplasmic sperm injection and embryo culture in equine practice. Proceedings of the American Association of Equine Practitioners 2007, 53;554–9. 3. Duchamp G, B´ezard J, Palmer E. Oocyte yield and the consequences of puncture of all follicles larger than 8 millimetres in mares. Biol Reprod Monogr 1995;1:233–41. 4. Carnevale EM, Coutinho da Silva MA, Preis KA, Stokes JE, Squires EL Establishment of pregnancies from oocytes collected from the ovaries of euthanized mares. Proceedings of the American Association of Equine Practitioners 2004:50;531–3. 5. Ribeiro BI, Love LB, Choi YH, Hinrichs K. Transport of equine ovaries for assisted reproduction. Anim Reprod Sci 2008;108:171–9.
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Immature Oocyte Collection and Maturation 6. Hinrichs K. The relationship of follicle atresia to follicle size, oocyte recovery rate on aspiration, and oocyte morphology in the mare. Theriogenology 1991;36:157–68. 7. Hinrichs K, Williams KA. Relationships among oocytecumulus morphology, follicular atresia, initial chromatin configuration, and oocyte meiotic competence in the horse. Biol Reprod 1997;57:377–84. 8. Hinrichs K, Schmidt AL. Meiotic competence in horse oocytes: interactions among chromatin configuration, follicle size, cumulus morphology, and season. Biol Reprod 2000;62:1402–8. 9. Choi YH, Love LB, Varner DD, Hinrichs K. Factors affecting developmental competence of equine oocytes after intracytoplasmic sperm injection. Reproduction 2004;127:187–94.
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10. Hinrichs K, Choi YH, Love LB, Varner DD, Love CC, Walckenaer BE. Chromatin configuration within the germinal vesicle of horse oocytes: changes post mortem and relationship to meiotic and developmental competence. Biol Reprod 2005;72:1142–50. 11. Choi YH, Love LB, Varner DD, Hinrichs K. Holding immature equine oocytes in the absence of meiotic inhibitors: Effect on germinal vesicle chromatin and blastocyst development after intracytoplasmic sperm injection. Theriogenology 2006;66:955–63. 12. Hinrichs K, Schmidt AL, Friedman PP, Selgrath, JP, Martin MG. In vitro maturation of horse oocytes: Characterization of chromatin configuration using fluorescence microscopy. Biol Reprod 1993;48:363–70.
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Chapter 309
Mature Oocyte Collection Elaine M. Carnevale
Introduction Oocytes can be collected from immature follicles or from follicles in the process of maturation. The collection of oocytes from preovulatory follicles has advantages and disadvantages when compared with other stages of the reproductive cycle. The number of follicles that reach the preovulatory stage is limited in the monovular mare. However, the oocyte collection rate per follicle is usually higher from preovulatory than from immature follicles, and oocyte maturation occurs more naturally in vivo versus in vitro.
Ovarian superstimulation Multiple preovulatory follicles per cycle could be advantageous for oocyte collections. Compounds to superstimulate ovaries have been developed that reduce follicular atresia and increase ovulation rates. However, success has been limited when using these compounds to increase the number of oocytes collected from preovulatory follicles per cycle.1 In the author’s experience, problems with superstimulation and preovulatory oocyte collections include large ovaries that are difficult to manipulate, irregular collapse of follicles at aspiration making collections more difficult, and variability in the stage of oocyte maturation per follicle. Recently a low dose of equine follicle-stimulating hormone (eFSH, 6.25 mg i.m., b.i.d; Bioniche Animal Health Inc., Athens, GA, USA) approximately doubled the number of preovulatory follicles per cycle and significantly increased the number of oocytes collected per cycle in young mares (1.1 vs 0.7 oocytes with or without administration of eFSH, respectively).2
Timing of oocyte collections Oocyte collections from preovulatory follicles are timed from administration of an ovulation-inducing
agent, such as human chorionic gonadotropin (hCG), equine luteinizing hormone (eLH), or the gonadotropin-releasing hormone analog (GnRH analog) deslorelin. Oocytes are often collected between 20 and 36 hours after hCG or between 16 and 0 hours, before the anticipated time of ovulation. Oocyte collections before approximately 20 hours after hCG administration are difficult, as follicular cells and the oocyte are more tightly adhered to the follicular wall. As the follicle progresses toward ovulation, cell adhesions continue to loosen and oocytes are easier to remove from the follicle. However, ovulation prior to oocyte collection becomes more of a risk. When ovulation is imminent, the follicle can be easily ruptured during transrectal manipulations for oocyte collection. In our laboratory the typical criteria for administration of an ovulation-inducing agent are: (i) follicle ≥ 35 mm in diameter; (ii) relaxed cervical and uterine tone; and (iii) uterine edema or estrous behavior for at least 1 day. However, criteria can vary for individual mares.
Oocyte collection methods Methods that have been used to collect oocytes from preovulatory follicles include laparotomies,3 colpotomies,4, 5 flank punctures,6, 7 and transvaginal ultrasound-guided follicular aspirations.8, 9 Transvaginal ultrasound-guided aspirations or flank punctures are the primary methods that are currently being used to collect oocytes. Flank puncture Flank puncture procedures may be advantageous in mares with infectious vaginal discharge or abnormal perineal conformation. Flank punctures require minimal equipment, but abdominal contents cannot be imaged during the procedure. A trocar with outer cannula (commonly 10 mm outer diameter) is
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Mature Oocyte Collection placed in the flank at the approximate position of the ovary. Trocar placement can be aided by positioning the ovary per rectum and marking the optimal site. The placement site is desensitized with a local anesthetic, and a small incision is made through the skin. A trocar/cannula combination is inserted through the flank musculature and into the perineal cavity. Placement can be monitored by transrectal palpation. The ovary is manipulated so the apex of the follicle is positioned against the cannula. While the ovary is stabilized per rectum, a 12- to 17-gauge needle is placed through the cannula and into the follicular antrum, and the follicular cavity is lavaged as described below. Transvaginal ultrasound-guided oocyte collection During transvaginal ultrasound-guided follicle aspirations the procedure is imaged with ultrasound. An ultrasound machine, transducer, and transducer casing with a needle guide are required. Linear, curvilinear, and sector transducers can be used for oocyte collections. When a linear transducer is used the face of the transducer is positioned dorsally, and the ovary is stabilized dorsal to the transducer. The ovary is positioned with the preovulatory follicle adjacent to the needle guide at the posterior end of the transducer face. With the curvilinear transducer the ovary is held anterior to the transducer. For mares with short vaginas or broad ligaments the curvilinear probe is often easier to use, as the distance between the face of the transducer and the needle guide is shorter. Before an oocyte recovery, flunixin meglumine can be administered if additional analgesia is warranted. For the oocyte collection procedure the mare is placed in a stock; the tail is wrapped; the rectum is evacuated; and the perineum is washed. We sedate mares with a combination of butorphanol and xylazine or detomidine for the procedure, with the type and amount of sedation dependent on the individual mare. Rectal contractions are reduced by the administration of BuscopanTM (N-butylscopolammonium bromide, i.v., Boehringer Ingelheim Vetmedica, Inc., St Joseph, MO, USA). Mares are twitched during the oocyte collection to minimize movement. Air within the rectum makes the procedure more difficult and, potentially, more dangerous for the mare. Therefore the clinician should be careful to prevent air from entering the rectum, or air can be removed prior to beginning the procedure. Soft Y-tubing attached to a 500-mL dose syringe can aid in quickly suctioning the air from the rectal lumen. Before placing the probe into the mare’s vagina a non-toxic lubricant is applied to the transducer to improve contact with the vaginal wall. Without allowing air into the vagina, the transducer is placed into
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the anterior vagina, lateral to the cervix, and ipsilateral to the preovulatory follicle. Transrectal manipulations are used to position the follicle with the apex juxtaposed to the needle guide. Proper positioning is often aided by moving the ovary anterior to the broad ligament before bringing it caudad to the face of the transducer. The surface of the ovary can be palpated to ensure the needle will not puncture the broad ligament, oviduct or ovulation fossa. Single-lumen needles can be used; however, 12-gauge double-lumen needles are often used for equine oocyte collections. The needle is advanced through the needle guide to puncture the vaginal and follicular walls. Suction can be applied using large syringes or a pump set at approximately 150 mmHg. Double-lumen needles have an inflow and outflow tube; therefore the follicle can be lavaged with 50–150 mL of flush medium concomitant with aspiration of follicular contents. Various media have been used for oocyte collections, including modified Dulbecco’s phosphatebuffered saline or commercial embryo-flush solutions; these solutions typically are sterile and contain a protein source and antibiotics. In our laboratory we have primarily used EmCare Complete Ultra (ICPbio, Auckland, New Zealand) for oocyte recoveries. Heparin at approximately 10 IU/mL is added to the flush medium to prevent coagulation. During aspiration of the follicular contents media can be alternately or simultaneously collected and injected, while the follicle is gently massaged per rectum. Approximately 30 mL of yellow follicular fluid is initially collected from the preovulatory follicle. During the follicular lavage the medium often becomes bloody in appearance as the vascular network of the follicle is disturbed. Before oocyte collection, media and equipment are warmed to 38.5◦ C, as cooling of the oocyte could affect its developmental potential. The recovered medium and follicular fluid is poured into large search dishes and examined under a dissecting microscope to locate the oocyte. The aspirate is not filtered as the cumulus–oocyte complex is sticky and tends to adhere to the filter. The equine oocyte is approximately 160 µm in diameter; however, the cumulus–oocyte complex is often greater than 1 mm in diameter. Therefore the cumulus complex is often easy to identify and can usually be seen without the use of a stereoscope. The cumulus cells appear as a translucent mass in the bloody flush solution (Figure 309.1a). Sheets of granulosa cells are also recovered in the aspirate. Although granulosa cells appear similar to the cumulus complex, they have differentiating characteristics of texture and color (Figure 309.1a,b). The cumulus mass can be imaged under a stereoscope to identify the oocyte. The oocyte is gray, often with variations in shading (Figure 309.1b). Within the cumulus complex a ring of cells, the corona radiata,
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Figure 309.1 (a) The cumulus–oocyte complex (coc) is observed as a clear mass of cells with attached granulosa cells (gran) in the bloody aspirate. The dark spot within the coc is the oocyte. Air bubbles are imaged on the right side of the picture. (b) The cumulus–oocyte complex (coc) is typical for collections from follicles approximately 24 hours after hCG is administered to the donor mare. The granulosa cells (gran) are darker and denser than the cumulus cells. The corona radiata (cr) is imaged as a ring surrounding the oocyte (oo).
surrounds the oocyte (Figure 309.1b). After identification the cumulus–oocyte complex is washed in medium to remove any attached blood and debris. In general, collection of oocytes from mature follicles is successful with experience in the technique. Failure of the follicle to mature properly can be one of the causes of unsuccessful collections. When oocyte collections were attempted at 39 h after administration of a GnRH analog, significantly more oocytes were collected from mature follicles with an irregular shape than from follicles that were round and termed immature (79% vs 18%, respectively).10 The number of oocytes collected per follicle was not different at 22 h versus 33 h after hCG administration (75% and 82%, respectively) using transvaginal aspirations, or at 24 h versus 35 h after hCG (66% and 73%, respectively) using flank aspirations.11, 12 However, when personnel with limited experience in transvaginal follicular aspiration attempted to collect oocytes in our laboratory, the collection rate per follicle was significantly lower from follicles at 23 h versus 35 h after administration of hCG (38% vs 65% recovery, respectively) (EM Carnevale and GK Lund, unpublished data). However, with experience in the technique, collection rates of at least 70% are anticipated. These rates probably correspond with the ability of the clinician successfully to manipulate and hold the follicle during the procedure.
Oocyte culture and transfer Because oocytes are often collected from follicles before they have completed maturation, the oocyte may need to be cultured before it is ready for fertilization. Conversely the oocyte has a limited lifespan
after ovulation or completion of maturation,13 and fertilization must occur while the oocyte is viable. Different scenarios can be used for calculating the interval between oocyte collection and transfer or sperm injection. Regardless, the interval between administration of hCG and fertilization is usually 36–40 hours. This interval can be extended to approximately 44 hours if deslorelin is used to induce oocyte maturation. The extent of oocyte maturation can be judged to some extent by the appearance of the cumulus and granulosa cells upon collection of the oocyte. Additional information can be gained from the ultrasonographic appearance of the follicle before ovulation.14 Because the cumulus cells surround the oocyte, a polar body is seldom imaged using a stereoscope. If the follicle fails to mature, the cumulus complex will not expand and will appear as a small mass of tightly bound cells surrounding the oocyte (Figure 309.2a). The cumulus mass at 24 hours after hCG is typically light in appearance with a distinct corona radiata (Figure 309.1b). As the follicle progresses toward ovulation the cumulus cells, including the corona radiata, become increasingly mucoid and separated (Figure 309.2b). Overly mature or atretic cumulus cells are often clumped in appearance (Figure 309.2c), and the oocyte may appear dark and small. Oocytes collected at least 30 hours after hCG administration or oocytes collected from follicles with ultrasonographic characteristics of impending ovulation are immediately transferred into a recipient’s oviduct. Oocytes collected prior to 30 hours after maturation induction are usually cultured in vitro for the completion of maturation before transfer. Pregnancy rates do not differ for oocytes collected just prior to ovulation and transferred, and ones collected
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Mature Oocyte Collection
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(c) Figure 309.2 The cumulus–oocyte complex changes appearance with maturation: (a) immature oocyte with dense cumulus cells; (b) mature oocyte with total expansion of the corona radiata; and (c) atretic oocyte with clumped cumulus cells. Arrows point to oocytes. The zona pellucida is imaged as a clear ring around the ooplasm.
earlier and cultured prior to transfer.12 When oocytes are collected approximately 24 hours after ovulation induction, they are cultured for 12–16 hours before transfer or sperm injection. In our laboratory oocytes have been cultured in Tissue Culture Medium 199 with additions of 10% fetal calf serum, 0.2 mM pyruvate, and 25–50 µg/mL gentamicin.15 No exogenous hormones are added to the medium because oocyte maturation was initiated in vivo. The oocytes are cultured in an incubator with an atmosphere of 6% CO2 and air and at 38.5◦ C. Oocyte transfer occurs when the oocytes are mature and ready for fertilization.
Conclusion Procedures have been developed for oocyte collections from large preovulatory follicles. Oocyte maturation is initiated in vivo by administration of an
ovulation-inducing compound. The technique that is most widely used for oocyte collections is a transvaginal ultrasound-guided procedure.
References 1. Maclellan LJ, Carnevale EM, Coutinho da Silva MA, Scoggin CF, Bruemmer JE, Squires EL. Pregnancies from vitrified equine oocytes collected from superstimulated and non-stimulated mares. Theriogenology 2002;58: 911–19. 2. Altermatt JL, Suh TK, Stokes JE, Campos-Chillon LF, Carnevale EM. Effect of mare age on oocyte morphology and developmental competence after ICSI. Reprod Fert Develop 2008;20:81. 3. Volgelsang MM, Kraemer DC, Bowen MJ, Potter GD. Recovery of equine follicular oocytes by surgical and nonsurgical techniques. Theriogenology 1986;25:208. 4. Hinrichs K, Kenney RM. A colpotomy procedure to increase oocyte recovery rates on aspiration of equine preovulatory follicles. Theriogenolgy 1987;27:237.
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5. Hinrichs K, Kenney DF, Kenney RM. Aspiration of oocytes from mature and immature preovulatory follicles in the mare. Theriogenology 1990;34:107–12. 6. Palmer E, Duchamp G, Bezard J, Magistrini M, King A, Bousquet D, Betteridge K. Recovery of follicular fluid and oocytes of mares by non-surgical puncture of the preovulatory follicle. Theriogenology 1986;25:178. 7. McKinnon AO, Wheeler MB, Carnevale EM, Squires EL. Oocyte transfer in the mare: Preliminary observations. J Equine Vet Sci 1987;6:306–9. 8. Carnevale EM, Ginther OJ. Use of a linear ultrasonic transducer for the transvaginal aspiration and transfer of oocytes in the mare. J Equine Vet Sci 1993;13:331–3. 9. Cook NL, Squires EL, Ray BS, Cook VM, Jasko DJ. Transvaginal ultrasound-guided follicular aspiration of equine oocytes. J Equine Vet Sci 1993;15:71–4. 10. McKinnon AO, Lacham-Kaplan O, Trounson AO. Pregnancies produced from fertile and infertile stallions by
11.
12.
13. 14.
15.
intracytoplasmic sperm injection (ICSI) of single frozenthawed spermatozoa into in vivo matured mare oocytes. J Reprod Fertil 2000;56(Suppl.):513–17. Coutinho da Silva MJ, Carnevale EM, Maclellan LJ, Seidel GE Jr, Squires EL. Effect of time of oocyte collection and site of insemination on oocyte transfer in mares. J Anim Sci 2002;80:1275–9. Hinrichs K, Betschart RW, McCue PM, Squires EL. Effect of time of follicle aspiration on pregnancy rate after oocyte transfer in the mare. J Reprod Fertil 2000;56(Suppl.):493–8. Ginther OJ. Reproductive Biology of the Mare, 2nd edn. Cross Plains, WI: Equiservices, 1992. Carnevale EM. Folliculogenesis and ovulation. In: Rantanen NW, McKinnon AO (eds) Equine Diagnostic Ultrasonography. Baltimore: Williams & Wilkins, 1998; pp. 201–11. Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biology of Reproduction Monograph Series 1995;1:209–14.
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Chapter 310
Oocyte Transfer Elaine M. Carnevale
Introduction Oocyte transfer involves the transfer of an oocyte into the oviduct of a recipient mare. The recipient is inseminated and fertilization, embryo development, and fetal development occur within the reproductive tract of the recipient. Because the oocyte does not contact the donor’s oviduct or uterus, many reproductive problems can be avoided. The first experimental foal was produced by oocyte transfer from a pregnancy reported in the late 1980s;1 however, clinical use of the procedure did not begin until the late 1990s. 2, 3 Since that time, oocyte transfer has been used to obtain pregnancies from mares considered infertile using standard breeding methods or embryo transfer.3, 4 Requirements for oocyte donors are minimal, as the donor needs only to produce a viable oocyte. Oocyte transfer appears to be a successful method to circumvent numerous reproductive pathologies, including those of the uterus, cervix, oviduct, or ovary. Different scenarios have been used to time oocyte collections and transfers. However, the complete interval between administration of an ovulationinducing agent, such as human chorionic gonadotropin (hCG) or deslorelin, and transfer is usually between 36 and 40 hours. The collection of oocytes at 24 hours compared with 36 hours after hCG administration to donors did not affect pregnancy rates.5
Oocyte recipients Recipients for oocyte transfer should be young, preferably 3–10 years old. In addition to carrying the pregnancy, oocyte recipients will be inseminated, so their reproductive tract must provide a suitable environment for sperm transport, fertilization, and embryo development. Reproductive examinations should be done on recipients before their use.
Currently, oocytes are transferred during standing flank laparotomies, with the ovary partially exteriorized through the abdominal musculature to allow positioning of the oviductal os for transfer. Ovarian exposure is difficult in mares with short thick flanks and short broad ligaments. Ligament length can be assessed by transrectal manipulation of the ovary. The distance from the dorsal abdominal wall and ovary can be determined by palpating the length of the ligament, which should be approximately 7 cm to allow easy surgical exposure in most mares. Recipients should be in estrus at the time of insemination. Estrus can be natural or induced with exogenous hormones. The recipient’s uterus should have endometrial edema indicative of estrus, with relaxed uterine and cervical tone. Cyclic and non-cyclic mares have been used as oocyte recipients, with similar pregnancy rates.4
Cyclic recipients If cyclic mares with large follicles are used as recipients, cycles of donors and recipients must be synchronized. The recipient’s oocyte is collected before transfer of the donor’s oocyte. An ovulation-inducing agent, such as hCG or deslorelin, is administered to the donor and recipient at approximately the same time. In our laboratory, we collect the donor’s oocyte first. If the collection attempt is successful, the recipient’s oocyte is collected. The recipient’s own oocyte must be removed to avoid fertilization of the incorrect oocyte after insemination. Recipients with two preovulatory follicles or a growing secondary follicle are preferentially avoided. If the recipient’s oocyte is not collected, the recipient is not used. Pregnancies have been established in mares in which the follicle has been aspirated or manipulated.6, 7 However, in our laboratory, we have inseminated recipient mares from which the oocyte was not collected after
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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an oocyte collection attempt by an experienced aspirator, and no pregnancies resulted (E.M. Carnevale, unpublished data). Therefore, the potential for establishing a pregnancy from a recipient’s own oocyte may vary with experience of the aspirator and the collection technique. Regardless, the best policy is to not use a recipient mare if the oocyte is not recovered and definitively identified. Potentially the recipient’s preovulatory follicle could be aspirated during the surgical transfer. However, some problems occur with this approach. The follicle could ovulate prior to surgery, and preovulatory follicles are often large and flaccid and can collapse when the ovary is manipulated during surgery. This could be avoided to some extent by not administering ovulatory agents to the recipient; however, oocyte recovery from untreated recipients is difficult even while the follicle is exposed during surgery. Cyclic recipients can be used during early estrus, when the dominant follicle is small and the cervix is relaxing. Prostaglandin can be administered to diestrous recipients with small follicles, and estrogen can be administered to induce uterine edema and cervical relaxation. After the oocyte is transferred, the recipient will be placed on progesterone, which may reduce the potential for ovulation; however, ovulation failure is not guaranteed. Oocyte transfer has the intrinsic potential of having a recipient’s own oocyte fertilized. Precautions can minimize the potential for this occurrence; however, complete control of the recipient’s follicular activity is difficult. Non-cyclic recipients Non-cyclic mares or mares with reduced follicular activity can be used as oocyte recipients. Advantages of these mares are that the small follicles should not ovulate and timing of the recipient with the donor is relatively easy. Anestrous and early transitional mares can be used as recipients during the nonovulatory season.3, 4, 8 This can be achieved in the spring by placing donor mares under lights to hasten cyclicity, while recipients are maintained under natural daylight. Recipient mares are treated with exogenous hormones to stimulate the uterus. A high dose of a gonadotropin-releasing hormone (GnRH) agonist (Ovuplant, deslorelin acetate, 4.2 mg subcutaneously) has also been used to reduce ovarian follicular activity4 and induce a period of reduced follicular activity during the breeding season. Recipients can then be managed as non-cyclic mares. Alternative methods to obtain mares with minimal follicular activity are possible, although synchronization of the recipient and donor may be difficult.
Progesterone and estrogen (150 mg P4 and 10 mg E2 ) has been administered to reduce follicular activity in cyclic mares.2 After progesterone and estrogen are discontinued, prostaglandin is administered if a corpus luteum is present, and the recipients are treated with estrogen before oocyte transfer. For non-cyclic recipients or for recipients with suppressed follicular activity, estrogen is administered to induce an artificial estrus before insemination. In our laboratory, estradiol 17β in oil (1.5–5 mg) is administered daily to non-cyclic recipients. Estrogen will induce endometrial edema and cervical relaxation if no luteal tissue is present. Daily administration of estrogen over time will result in some mares becoming non-responsive, and estrogen should be discontinued for a number of days.
Transfer procedures Oocytes can be transferred ipsilateral or contralateral to the side of a recipient’s own aspirated preovulatory follicle. Similar numbers of sperm enter the ipsilateral or contralateral oviduct to the preovulatory follicle.9 The transfer of oocytes into oviducts contralateral to the recipient’s own ovulation resulted in similar embryo development rates from transferred and ovulated oocytes (73% and 69%, respectively10 ). The site of the recipient’s own follicular aspiration is often bloody and sensitive, similar to a natural ovulation, and the ovary is often large. This can make the transfer more difficult. Therefore, oocytes are often transferred into the oviduct contralateral to the recipient’s own aspirated follicle. Oocytes are transferred during standing flank laparotomies. Recipients are placed in a stock with a partial side gate. The flank area is clipped and scrubbed. A presurgical sedative of butorphanol tartrate and xylazine hydrochloride or detomidine hydrochloride is administered, and the surgical site is blocked with a local anesthetic. Before surgery, additional sedation is administered; we often use a combination of butorphanol tartrate and detomidine hydrochloride. Sedation is modified as needed for individual recipients. An incision approximately 6–8 cm long is made midway between the last rib and tuber coxae. The abdominal oblique muscle layers are bluntly dissected, and the peritoneum is punctured. The oocyte is removed from the incubator and placed into medium just prior to transfer. The medium should be warmed (38–39◦ C) and buffered to maintain pH at room atmosphere. In our laboratory, we have used a commercial embryo-holding medium (EmCare Embryo Holding Solution; ICPbio, Auckland, New Zealand) and fire-polished glass pipettes for transfers.
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Oocyte Transfer The ovary is identified and exteriorized through the incision to expose the oviduct. The oviductal os is located by identifying the oviductal outline on the serosal surface of the infundibulum. The pipette is inserted into the os and advanced 2–3 cm. The oocyte and a small volume (< 0.1 mL) of medium are released into the oviduct. The ovary is carefully returned to the abdominal cavity and the surgical site is closed.
Insemination of recipients Adequate numbers of high-quality sperm are required to maximize the success of oocyte transfer. In a commercial program, semen is provided from numerous stallions of variable fertility with most semen cooled and transported.3, 11 The equine oocyte has a limited lifespan, maintaining viability for approximately 12 hours after ovulation.12 Oocytes are usually mature at transfer, and recipients are inseminated before and/or after transfer. Single inseminations before13, 14 or after15 oocyte transfer have resulted in pregnancies. During most experimental transfers, recipients were inseminated approximately 12 hours before and 2 hours after transfers with a total of 2 × 109 progressively motile sperm. In our commercial program with older donors and cooled semen from numerous stallions of variable fertility, pregnancy rates were significantly higher when recipients were inseminated before or before and after oocyte transfer than when recipients were inseminated only after transfer (respectively 18/45, 40%; 27/53, 51%; and 0/10, 0%).11 The results suggest that insemination of recipients before oocyte transfers with approximately 1 × 109 progressively motile sperm from a fertile stallion is sufficient. Insemination of the recipient before and after transfer could be beneficial, especially if the semen is not optimal. Recipients in our clinical program are typically inseminated 12–16 hours before transfer and 2– 4 hours after transfer if a second insemination dose is available. After insemination and transfer, the recipient’s uterus can be monitored using ultrasonography to detect intrauterine fluid collections. Recipients with fluid collections are treated to resolve uterine inflammation or infection. Treatments include oxytocin or prostaglandin to stimulate uterine clearance or uterine lavage and antibiotic infusions.
Progesterone treatment Progesterone supplementation is required for noncyclic mares after oocyte transfer. Cyclic recipients will develop a corpus luteum after aspiration of
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their preovulatory follicle. However, many granulosa cells are removed, which could affect luteal function. Therefore, most clinical recipients are treated with exogenous progesterone or progestins after oocyte transfer.2–4
Success of oocyte collection and transfer The anticipated success of oocyte transfer is dependent on the quality of oocytes and sperm and on the expertise of personnel. Embryo development rates after experimental transfers of oocytes from young, fertile donors have varied, but rates between 60% and 80% are probable with an optimal system. In our clinical oocyte transfer program we have older, subfertile mares and use cooled, transported semen from numerous stallions with variable fertility.3, 4 We anticipate a pregnancy rate of approximately 40% after transfer of oocytes. However, success can be affected by many factors, especially the age of the mare4, 16 and semen quality.
References 1. McKinnon AO, Squires EL, Carnevale EM, Voss JL, Seidel GE. Heterogenous and xenogenous fertilization of equine oocytes. Theriogenology 1988;29:278. 2. Hinrichs K, Provost PJ, Torello EM. Treatments resulting in pregnancy in nonovulating, hormone-treated oocyte recipient mares. Theriogenology 2000;54:1285–93. 3. Carnevale EM, Squires EL, Maclellan LJ, Alvarenga MA, Scott TJ. Use of oocyte transfer in a commercial breeding program for mares with reproductive abnormalities. J Am Vet Med Assoc 2001;218:87–91. 4. Carnevale EM, Coutinho da Silva MA, Panzani D, Stokes JE, Squires EL. Factors affecting the success of oocyte transfer in a clinical program for subfertile mares. Theriogenology 2005;64:519–27. 5. Hinrichs K, Betschart RW, McCue PM, Squires EL. Effect of time of follicle aspiration on pregnancy rate after oocyte transfer in the mare. J Reprod Fertil Suppl 2000;56: 493–8. 6. McKinnon AO, Wheeler MB, Carnevale EM, Squires EL. Oocyte transfer in the mare: preliminary observations. J Equine Vet Sci 1987;6:306–9. 7. Coutinho da Silva MA, Carnevale EM, Maclellan LJ, Scoggin CF. Injection of blood into preovulatory follicles of equine oocyte transfer recipients does not prevent fertilization of the recipient’s oocyte. Theriogenology 2002;57: 538. 8. Carnevale EM, Alvarenga MA, Squires EL, Choi YH. Use of noncycling mares as recipients for oocyte transfer and GIFT. In: Proceedings of the Annual Conference of the Society for Theriogenology, Nashville, Tennessee, 22–24 September, 1999, p. 44. 9. Rigby S, Derczo S, Brinsko S, Blanchard T, Taylor T, Forrest DW, Varner D. Oviductal sperm numbers following proximal uterine horn or uterine body insemination. In: Proceedings of the 46th Convention of the American Association of Equine Practitioners, San Antonio, Texas, 26–29 November, 2000, pp. 332–4.
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10. Carnevale EM, Coutinho da Silva MA, Maclellan LJ, Seidel Jr GE, Squires EL. Use of parentage testing to determine optimum insemination time and culture media for oocyte transfer in mares. Reproduction 2004;128: 623–8. 11. Carnevale EM, Maclellan LJ, Coutinho da Silva MA, Checura CM, Scoggin CF, Squires EL. Equine sperm-oocyte interaction: results after intraoviductal and intrauterine inseminations of recipients for oocyte transfer. Anim Reprod Sci 2001;68:305–14. 12. Ginther OJ. Reproductive Biology of the Mare, 2nd edn. Cross Plains, WI: Equiservices, 1992. 13. Scott TJ, Carnevale EM, Maclellan LJ, Scoggin CF, Squires EL. Embryo development rates after transfer of oocytes
matured in vivo, in vitro, or within oviducts of mares. Theriogenology 2001;55:705–15. 14. Maclellan LJ, Carnevale EM, Coutinho da Silva MA, Scoggin CF, Bruemmer JE, Squires EL. Pregnancies from vitrified equine oocytes collected from superstimulated and non-stimulated mares. Theriogenology 2002;58:911–19. 15. Carnevale EM, Maclellan LM, Coutinho da Silva MA, Scott TJ, Squires EL. Comparison of culture and insemination techniques for equine oocyte transfer. Theriogenology 2000;54:982–7. 16. Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod Monogr Ser 1995;1:209–14.
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Chapter 311
Gamete Intrafallopian Transfer (GIFT) Marco A. Coutinho da Silva
Introduction In vitro production of embryos has successfully been used in humans and in some domestic and laboratory species. However, in the horse, production of embryos after in vitro fertilization is not yet repeatedly successful. Compared to other species, reasons for slow development of in vitro fertilization in the horse include scarcity of slaughterhouse ovaries and difficulties with in vitro systems for oocyte maturation and sperm capacitation. Only two foals have been produced after in vitro fertilization of oocytes matured in vivo,1, 2 (and these results have not been reproduced consistently by other laboratories). Although in vitro fertilization has not been successful in horses, different approaches for fertilization of oocytes in vivo have been developed. Equine oocytes have been fertilized in the oviducts of other species (xenogenous fertilization), or in the oviducts of recipient mares. In vivo fertilization of equine oocytes in recipient mares has been done using oocyte transfer (OT),3–12 and gamete intrafallopian transfer (GIFT).9,13–15 Although the acronyms OT and GIFT are used interchangeably in the literature, distinctions should be made. During OT, the oocyte of a donor mare is transferred into the oviduct of a recipient, and the recipient is inseminated within the uterus. GIFT involves the transfer of oocyte(s) and spermatozoa into the oviduct of a recipient. Although oocyte transfer has proven to be a successful technique for obtaining pregnancies from subfertile mares, the procedure requires semen from fertile stallions. In contrast, during GIFT sperm are deposited near the site of fertilization within the oviduct and therefore, the procedure has the potential to be used when sperm numbers are low, for example in cases of subfertile stallions, frozen semen, and sexed sperm.
This chapter discusses some concepts and procedures related to the use of GIFT in the horse.
Gamete handling Oocytes Management of the donor mare as well as procedures for oocyte collection and maturation are described in Chapters 308, 309 and 310. Oocytes used for GIFT have been collected at approximately 24 or 33 hours after gonadotropin administration, and transferred into the oviduct either after in vitro maturation or immediately after collection, respectively.14 Pregnancy rates after GIFT were numerically higher when oocytes were collected at 22 h and completed maturation in vitro (82%) versus transfer of in vivo matured oocytes (27%), although statistical significance was lacking. Because multiple oocytes were transferred into a relatively small number of recipient mares (n = 4 per group), recipient quality could have had a major effect on these results. In the same study, the time of oocyte collection did not affect pregnancy rates when recipients were inseminated within the uterus (OT; 22 h = 57% and 33 h = 75%), demonstrating that oocyte quality was similar between collection times. Potentially, the time necessary for sperm to undergo capacitation and achieve fertilizing capacity is different when sperm are deposited within the oviduct versus the uterus, and the timing of sperm capacitation and oocyte maturation may be critical for the success of GIFT. Spermatozoa Conventional intrauterine insemination of mares requires a minimum of 5 × 108 progressively motile sperm for maximum fertilization.16 Intraoviductal
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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insemination requires fewer sperm than intrauterine insemination. In the horse, between 5 × 104 and 5 × 105 progressively motile sperm have been used for intraoviductal inseminations.9,13–15,17 The uterus and oviduct have synergistic functions with respect to capacitation.18 Pre-capacitation of sperm for use in GIFT has or has not been performed, depending on the species. In the horse, McCue et al.17 inseminated mares within the uterus or oviduct with 5 × 108 and 5 × 104 progressively motile sperm, respectively, and obtained similar pregnancy rates (40% and 21%, respectively). In both groups, sperm was not capacitated prior to insemination. During the first attempts of GIFT in the horse, Carnevale et al. 9, 13 transferred sperm into the oviduct without precapacitation, resulting in pregnancies (33%). In subsequent studies, high pregnancy rates (82%) were obtained using non-capacitated sperm for GIFT,14, 15 demonstrating that capacitation of equine sperm can occur within the oviduct, and pre-capacitation may not be required for GIFT. During GIFT, sperm are deposited in the infundibular or ampullar regions of the oviduct, bypassing the uterus and the utero-tubal junction. Therefore, sperm must be separated from its seminal plasma, debris, and contamination. In addition, morphologically normal and motile sperm must be selected from the ejaculate. Methods frequently used to prepare semen for assisted reproductive procedures and to enhance the percentage of motile sperm in other species include swim-up,19–23 Percoll gradients,24–27 Sephadex gel filtration,22, 28 and glass-wool filtration.29 Percoll gradient has been the method of choice to select sperm for GIFT.9,13–15 Sperm were centrifuged through a Percoll gradient (90/45%), washed, and approximately 2–5 × 105 sperm were transferred into the oviduct with oocytes. Embryo development rates of GIFT using fresh (raw) semen ranged from 27% to 82%, demonstrating that GIFT could be an alternative to oocyte transfer when sperm numbers are low. However, in a commercial program, cooled and frozen semen are more likely to be available for GIFT than fresh semen. Therefore, in a recent study, the use of cooled and frozen semen for GIFT was investigated.15 When oocyte recipients were inseminated within the uterus (OT) or oviduct (GIFT), with cooled semen, embryo development rates of 83% (19/23) and 25% (4/16) were observed, respectively. The use of frozen semen for GIFT resulted in an embryo development rate of only 8% (1/12). Low embryo development rates obtained with cooled and frozen semen could be caused by some factor associated with cooling and freezing processes that impaired fertilization after intraoviductal inseminations. Further research must investigate the effects of extend-
ing, cooling, and freezing sperm on the interactions between sperm, oviduct, and oocyte, after deposition within the oviduct. Successful utilization of cooled and frozen semen for GIFT would provide the horse industry with an efficient technique to produce offspring from subfertile mares and stallions.
The future of GIFT In humans, GIFT has been used successfully in cases of male or female subfertility, with an overall pregnancy rate of 30%.30 Because of the increasing use of intracytoplasmic sperm injection (ICSI) in human assisted reproductive technology (ART), the number of GIFT procedures has been decreasing. During 1994 in the USA, GIFT was performed to treat infertility in 10% of the cycles in which assisted reproduction techniques were used. However in 2001, GIFT represented only 0.3% of ART used.31 In horses, GIFT is still a potential alternative to obtain pregnancies in cases of subfertility, especially male subfertility, because of the lack of success of IVF and the requirements for expensive equipment and trained personnel for ICSI. However, pregnancy rates of GIFT using cooled and frozen semen must be improved before the technique can be widely used in practice. During GIFT, sperm are deposited in the oviduct, hence it bypasses the uterus and the utero-tubal junction, which assist in selection of morphologically normal sperm. Differences in sperm transport and selection during GIFT could result in modifications of sperm–oviductal epithelium interactions, altering storage and lifespan of sperm,32 in particular cooled or frozen-thawed sperm. Further studies are necessary to improve our understanding of the events that occur after sperm deposition into the oviduct and to improve the procedures for GIFT with cooled or frozen semen.
References 1. Palmer E, Bezard J, Magistrini M, Duchamp G. In vitro fertilization in the horse. A retrospective study. J Reprod Fertil (Suppl.) 1991;44:375–84. 2. Bezard J. In vitro fertilization in the mare. Proc Int Sci Conf Biotech Horse Reprod. Agricultural University of Krakow, Poland, 1992;12. 3. McKinnon AO, Carnevale EM, Squires EL, Voss JL, Seidel GE Jr. Heterogenous and xenogenous fertilization of in vivo matured equine oocytes. J Equine Vet Sci 1988;8:143–147. 4. Ray BS, Squires EL, Cook NL, Tarr SF, Jasko DJ, Hossner KL. Pregnancy following gamete intrafallopian transfer in the mare. J Equine Vet Sci 1994;14:27–30. 5. Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod Mono 1995;1:209–14. 6. Hinrichs K, Matthews GL, Freeman DA, Torello EM. Oocyte transfer in mares. J Am Vet Med Assoc 1998;212: 982–6.
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Gamete Intrafallopian Transfer (GIFT) 7. Hinrichs K, Betschart RW, McCue PM, Squires EL. Effect of timing of follicle aspiration on pregnancy rate after oocyte transfer in the mare. J Reprod Fertil (Suppl.) 2000;56:493–8. 8. Carnevale EM, Alvarenga MA, Squires EL. Use of oocyte transfer in a commercial breeding program to obtain pregnancies from mares with reproductive pathologies. Proc 45th Ann Conv Amer Assoc Equine Pract 1999;200–206. 9. Carnevale EM, Maclellan LJ, Coutinho da Silva MA, Scott TJ, Squires EL. Comparison of culture and insemination techniques for equine oocyte transfer. Theriogenology 2000;54:981–7. 10. Carnevale EM, Squires EL, Maclellan LJ, Alvarenga MA, Scott TJ. Use of oocyte transfer in a commercial breeding program for mares with reproductive abnormalities. J Am Med Vet Assoc 2001;218:87–91. 11. Scott TJ, Carnevale EM, Maclellan LJ, Scoggin CF, Squires EL. Embryo development rates after transfer of oocytes matured in vivo, in vitro, or within oviducts of mares. Theriogenology 2001;55:705–15. 12. Maclellan LJ, Carnevale EM, Coutinho da Silva MA, Scoggin CF, Bruemmer JE, Squires EL. Pregnancies from vitrified equine oocytes collected from superstimulated and nonsuperstimulated mares. Theriogenology 2002;58: 911–19. 13. Carnevale EM, Alvarenga MA, Squires EL, Choi YH. Use of noncycling mares as recipients for oocyte transfer and GIFT. Proc Annu Conf Soc Theriogenology 1999;44 abstr. 14. Coutinho da Silva MA, Carnevale EM, Maclellan LJ, Seidel GE Jr, Squires EL. Effect of time of oocyte collection and site of insemination on oocyte transfer in mares. J Anim Sci 2002;80:1275–9. 15. Coutinho da Silva MA, Carnevale EM, Maclellan LJ, Preis KA, Seidel GE Jr, Squires EL. Oocyte transfer in mares with intrauterine or intraoviductal insemination using fresh, cooled and frozen stallion semen. Theriogenology 2004;61:705–13. 16. Pickett BW, Back DG, Burwash LD, Voss JL. The effect of extenders, spermatozoal numbers and rectal palpation on equine fertility. Proc 5th NAAB Tech Conf AI Reprod 1974, pp. 47–58. 17. McCue PM, Fleury JJ, Denniston DJ, Graham JK, Squires EL. Oviductal insemination of mares. J Reprod Fertil (Suppl.) 2000;56:499–502. 18. Bedford JM. Significance of the need for sperm capacitation before fertilization in eutherian mammals. Biol Reprod 1983;28:108–20. 19. Parrish JJ, Foote RH. Quantification of bovine sperm separation by a swim-up method. Relationship to sperm motility, integrity of acrosomes, sperm migration in polyacrylamide gel and fertility. J Androl 1987;8:259–66.
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20. Chan SYW, Chan YM, Tucker MJ. Comparison of characteristics of human spermatozoa selected by the multipletube swim-up and simple discontinuous Percoll gradient centrifugation. Andrologia 1991;2:213–18. 21. Karlstrom PO, Bakos O, Bergh T, Lundkvist O. Intrauterine insemination and comparison of two methods of sperm preparation. Hum Reprod 1991;6:390–95. 22. Check JH, Katsoff D, Kozak J, Lurie D. Effect of swim-up, Percoll and Sephadex sperm separation methods on the hypo-osmotic swelling test. Hum Reprod 1992;7:109–11. 23. Englert Y, Van Den Bergh M, Rodesch C, Bertrand E, Biramane J, Legreve A. Comparative auto-controlled study between swim-up and Percoll preparation of fresh semen samples for in-vitro fertilization. Hum Reprod 1992;7: 399–402. 24. Mathieu C, Mein M, Lornage J, Li JG, Guerin JF. Effect of spermatozoa selection on a simplified Percoll gradient in case of asthenozoospermia. Andrologia 1990;22:467–71. 25. Saeki K, Hoshi M, Leibfried-Rutledge ML, First NL. In vitro fertilization and development of bovine oocytes matured in serum-free medium. Biol Reprod 1991;44:256–60. 26. Morales P, Vantman D, Barros C, Vigil P. Human spermatozoa selected by Percoll gradient or swim-up are equally capable of binding to the human zona pellucida and undergoing to acrosome reaction. Human Reprod 1991;6:401–4. 27. Ng FLH, Liu DY, Baker HWG. Comparison of Percoll, mini-Percoll and swim-up methods for sperm preparation from abnormal semen samples. Human Reprod 1992;7: 261–6. 28. Ohashi K, Saji F, Wakimoto A, Kato M, Tsutsui T, Tanizawa O. Preparation of oligozoospermic and/or asthenozoospermic semen for intrauterine insemination using the SpermPrep semen filtration column. Fertil Steril 1992;57:866–70. 29. Paulson JD, Polakosi K, Leto S. Further characterization of glass wool column filtration of human semen. Fertil Steril 1979;32:125–6. 30. Assisted reproductive technology in the United States and Canada: 1994 results generated from the American Society for Reproductive Medicine/Society for Assisted Reproductive Technology Registry. Fertil Steril 1996;66:697–705. 31. Assisted reproductive technology in the United States: 1997 results generated from the American Society for Reproductive Medicine/Society for Assisted Reproductive Technology Registry. Fertil Steril 2007;87:1253–66. 32. Carnevale EM, Maclellan LJ, Coutinho da Silva MA, Checura CM, Scoggin CF, Squires EL. Equine sperm-oocyte interaction: results after intraoviductal and intrauterine inseminations of recipients for oocyte transfer. Anim Reprod Sci 2001;68:305–14.
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Chapter 312
Intracytoplasmic Sperm Injection (ICSI) Young-Ho Choi and Katrin Hinrichs
Intracytoplasmic sperm injection (ICSI) is a fertilization technique in which a single sperm is injected directly into the cytoplasm of an oocyte. ICSI circumvents many of the basic phenomena required for standard fertilization, such as sperm capacitation, the acrosome reaction, penetration of the sperm through the zona, and sperm–egg membrane fusion. Thus sperm sources such as immotile ejaculated sperm, sperm from the epididymis or testis, freeze-dried sperm, and isolated sperm nuclei can be used for embryo production. Conventional in vitro fertilization in the horse has been problematic, related to the failure of equine sperm to efficiently penetrate the zona pellucida in vitro, thus ICSI offers a useful alternative. Advances achieved in the ICSI technique and in culture of equine embryos over the last 5 years now allow clinical application of this technique and there is growing interest in ICSI from the horse industry, especially in cases of limited sperm stores.
History of ICSI Successful equine ICSI was first reported in 1996; transfer of four fertilized oocytes produced a pregnancy.1 Several papers on ICSI were subsequently published, but cleavage rates were low, ranging from 20% to 65%, and rates of blastocyst production in vitro were low (< 15%). In 2002, the efficiency of equine ICSI was significantly improved by the introduction of the Piezo drill, an instrument that causes minute vibrations in the injection pipette. When the Piezo drill was used for equine ICSI, cleavage rates of 80–85%2 and 69–89%3 were reported. In 2004, an embryo culture medium (DMEM/F-12 + 10% fetal bovine serum (FBS)) was identified that, combined with Piezodriven ICSI, produced blastocyst rates of 20 to 38%.4, 5
Once methods for ICSI and embryo culture were established, work was performed to determine the effect of sperm source on success of ICSI. Pronucleus formation and cleavage rates were no different between fresh and frozen-thawed sperm.2 Injection of sex-sorted sperm6 resulted in a lowered blastocyst rate. In a report of note to the horse industry, it was shown that frozen semen could be thawed, diluted 1:100, and refrozen with no decrease in blastocyst formation after ICSI.7 Air-dried sperm or sperm stored in a highosmolarity medium8 have produced cleaved embryos after ICSI (44–73% cleavage rates), but no blastocyst development was reported. Notably, equine blastocysts, pregnancies,9 and a foal (YH Choi and K Hinrichs, unpublished data) have been produced from equine ICSI with freezedried sperm9 . This method is still in its infancy. Oocytes injected with freeze-dried sperm required an additional activation stimulus to form blastocysts, and more work is needed in this area. However, this initial success opens up the possibility of alternative methods for preservation of sperm that avoid the need for liquid nitrogen containers and their associated complications for long-term storage and shipment. Almost all of the above work was done using in vitro-matured oocytes. In contrast, clinical application would most likely involve oocytes recovered from preovulatory follicles ex vivo. Theoretically, these oocytes should have high viability, and be associated with high rates of embryonic development, as seen in other species. However, currently in the horse, the rates of blastocyst development reported after ICSI with in vivo-matured oocytes are similar to those for ICSI of in vitro-matured oocytes.10, 11
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Intracytoplasmic Sperm Injection (ICSI)
Procedures Conventional ICSI To perform conventional ICSI (without a Piezo drill), injection pipettes are made either by hand on specialized machinery, or purchased from commercial companies. If made by hand, the pipette is pulled to a 7–8-µm outer diameter, the tip is beveled using a microgrinder, and a spike is made at the tip to assist penetration through the zona pellucida and oocyte plasma membrane. A sharp spike is needed when working with horse oocytes due to the tough zona pellucida. Preparation of sperm is done as for Piezo ICSI, as described below. The ICSI procedure is also similar to that for Piezo ICSI, with the exceptions of the method of sperm immobilization and actual injection. Sperm are immobilized by repeated pipetting or squashing or rolling the tail with the injection pipette. For injection, the pipette containing the sperm is pushed into the cytoplasm. When the pipette is one-third to half-way into the cytoplasm, a quick aspiration, accompanied with a swift push of the pipette, is performed to break the plasma membrane. The sperm is then released into the cytoplasm with minimal amount of accompanying solution.
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puller. The holding pipette is cut at 100–150 µm outside diameter, then a microforge is used to smooth the cut end and reduce the inner diameter to 15–20 µm. The injection pipette is cut vertically from pulled tubing at 7–8 µm outside diameter. Both pipettes are approximately 7 cm long and may be bent for easier use. A notable feature of the Piezo drill is that it requires a heavy medium in the injection pipette appropriately to convey the microvibrations; this is typically mercury or Fluorinert (Sigma). We use mercury in our laboratory. A needle or a thin hand-pulled glass pipette is used to transfer a small amount of the mercury into the large (non-working) end of the injection pipette. Water is then added to fill this end of the pipette. The pipette is attached to the injection syringe of the micromanipulator, and the mercury is pushed to the tip of the injection pipette. The length of the column of mercury within the pipette is about 5 mm.
Oocyte and sperm preparation
We use borosilicate glass capillary tubing (o.d. 1.0 mm, i.d. 0.75 mm) for both the holding and injection pipettes. The tubing is pulled using a pipette
Mature oocytes are denuded of cumulus cells by pipetting them with pipettes of decreasing bore diameter in 0.05% hyaluronidase (Sigma). Oocytes having a visible polar body are selected for manipulation (Figure 312.1a). Fresh semen is washed twice by centrifugation and resuspension in media to remove seminal plasma. The ‘swim-up’ method is applied to collect motile sperm from poor-quality fresh or cooled semen (after centrifugation to concentrate sperm), or from frozen-thawed semen. The swim-up method
(a)
(b)
ICSI with the Piezo drill
Pipette preparation
Figure 312.1 The ICSI procedure using a Piezo drill. (a) An oocyte with polar body visible at 12 o’clock; the sperm is visible at the tip of the pipette (arrow) just before injection. (b) Injection of the sperm deep into the cytoplasm of the mature oocyte.
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is conducted in a 5-mL tube containing 1 mL of Sp-CZB12 or other sperm-compatible media. Semen (200 µL) is layered into the bottom of the 5-mL tube, under the sperm medium. After 20 min incubation at 38.2◦ C, the top 600 µL of medium is collected, centrifuged, resuspended, and centrifuged again. The supernatant is removed and the sperm pellet is mixed with the small amount of medium that remains. One microliter of this sperm suspension is placed at the top of a 3-µL droplet of sperm medium containing 10% polyvinylpyrrolidone (PVP; MW 360 000) under light white mineral oil. To select actively motile sperm, the sperm that swim out to the edge of the PVP droplet are used for ICSI.
Sperm injection The manipulation medium is CZB-M with 10% FBS12 or other oocyte-compatible media (buffered for room atmosphere). Four droplets (two 3-µL droplets of 10% PVP solution, one for rinsing and one for sperm; and two 50-µL droplets of the manipulation medium, one for rinsing and one as the injection droplet) are placed on the lid of a Petri dish (# 1006, Falcon, Becton Dickinson Labwarc, NJ, USA). The droplets are covered with mineral oil. Ten to fifteen oocytes are put into the manipulation droplet at one time. An injection pipette is rinsed several times in the droplet containing the 10% PVP medium, then moved to the droplet containing sperm. A single sperm is immobilized by applying one or a few Piezo pulses (settings: speed 4; intensity 7) to the midpiece
(a)
of the sperm, then loaded immediately, tail-first, into the pipette. This is repeated on another sperm; two or three immobilized sperm, with space between them, can be in the pipette at one time. An oocyte is held with the polar body located either in the 6 or 12 o’clock position (Figure 312.1a) to avoid disruption of the metaphase plate. The pipette containing sperm is placed on the zona and a gentle negative pressure is applied to make a close contact between them. Then the zona is drilled with the Piezo at speed 4, intensity 7. The pipette is withdrawn and the piece of drilled zona is expelled from the pipette. The sperm inside the pipette is carefully pushed close to the tip and the pipette is pushed through the hole in the zona, deep into the oocyte cytoplasm (Figure 312.1b). The cytoplasmic membrane is broken by one or two Piezo pulses with lower settings (speed 3; intensity 5). The sperm is released with a minimum amount of surrounding solution. The settings for the Piezo can be varied based on the size of the pipette and whether or not mercury is being used. The manipulation is conducted at room temperature. After a set of oocytes has been injected, they are transferred into microdrops of holding medium (CZB-H12 or other compatible medium) and held in 5% CO2 at 38.2◦ C until ICSI has been completed with the rest of the oocytes.
In vitro culture After all the oocytes are injected with sperm, they are cultured in DMEM/F-12 with 10% FBS. While
(b)
Figure 312.2 (a) Five cleaved embryos, 2 to > 10 cells, produced by ICSI and cultured in vitro for 3 days. (b) Nine blastocysts produced by ICSI and in vitro culture for 7 days in DMEM/F-12 with 10% FBS.
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Intracytoplasmic Sperm Injection (ICSI)
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ICSI of in vitro-matured oocytes.18 This is an active area, and much more information on clinical application of ICSI should be available in the near future.
References
Figure 312.3 A foal resulting from transcervical transfer of a blastocyst produced by ICSI and cultured in vitro for 7 days. The foal was born at Performance Equine Associates in 2005 and the picture was taken at 1 day of age.
this is a complete cell culture medium, rather than a medium developed for embryo culture, we have shown that it supports high rates of blastocyst production in vitro (up to 38%5 ). Injected oocytes are cultured in groups of 1–5 oocytes per microdroplet at a ratio of 1µL medium per oocyte. Culture is performed in an atmosphere of 5% CO2 , 5% O2 , and 90% N2 at 38.2◦ C for 7–10 days. Embryos are evaluated for cleavage (Figure 312.2a) on day 3 and for blastocyst formation (Figure 312.2b) on days 7 to 10. The injection apparatus is dismantled after injection. Mercury must be disposed of in an environmentally safe manner; pipettes are not reusable. While there is much clinical interest in ICSI, few resulting from transfer of in vitro-produced blastocysts have been reported13–15 (Figure 312.3). More have been foaled in commercial programs (C. Galli, personal communication, 2008). Several foals have been produced from both in vivo-matured and in vitro-matured oocytes subjected to ICSI and then transferred to the oviducts of recipient mares16, 17 (EM Carnevale, personal communication, 2007). We have produced foals from clinical cases post mortem, after transfer of in vitro-cultured blastocysts produced by
1. Squires EL, Wilson JM, Kato H, Blaszczyk A. A pregnancy after intracytoplasmic sperm injection into equine oocytes matured in vitro. Theriogenology 1996;45:306 (abstract). 2. Choi YH, Love CC, Love LB, Varner DD, Brinsko S, Hinrichs K. Developmental competence in vivo and in vitro of in vitro-matured equine oocytes fertilized by intracytoplasmic sperm injection with fresh or frozen-thawed spermatozoa. Reproduction 2002;123:455–65. 3. Galli C, Crotti G, Turini P, Duchi R, Mari G, Zavaglia G, Duchamp G, Daels P, Lazzari G. Frozen-thawed embryos produced by ovum pick up of immature oocytes and ICSI are capable to establish pregnancies in the horse. Theriogenology 2002;58:705–8 (abstract). 4. Choi YH, Roasa LM, Love CC, Varner DD, Brinsko SP, Hinrichs K. Blastocyst formation rates in vivo and in vitro of in vitro-matured equine oocytes fertilized by intracytoplasmic sperm injection. Biol Reprod 2004;70:1231–8. 5. Hinrichs K, Choi YH, Love LB, Varner DD, Love CC, Walckenaer BE. Chromatin configuration within the germinal vesicle of horse oocytes: changes post mortem and relationship to meiotic and developmental competence. Biol Reprod 2005;72:1142–50. 6. Colleoni S, Spincaci M, Duchi R, Merlo B, Tamanini C, Lazzari G, Mari G, Galli C. ICSI of equine oocytes with sexsorted frozen-thawed semen results in low cleavage rate but normal embryo development and pregnancies. Reprod Fertil Dev 2009;21:228–29 (abstract). 7. Choi YH, Love CC, Varner DD, Hinrichs K. Equine blastocyst development after intracytoplasmic injection of sperm subjected to two freeze-thaw cycles. Theriogenology 2006;65:808–19. 8. Alonso A, Miragaya M, Losinno L, Herrera C. Intracytoplasmic sperm injection of equine oocytes using air-dried sperm or sperm stored in a high osmolarity medium. Reprod Fertil Dev 2007;19:301 (abstract). 9. Choi YH, Varner DD, Hartman DL, Hinrichs K. Blastocyst production from equine oocytes fertilized by intracytoplasmic injection of lyophilized sperm. Anim Reprod Sci 2006;94:307–8 (abstract). 10. Altermatt JL, Suh TK, Stokes JE, Carnevale EM. Effects of age and equine follicle-stimulating hormone (eFSH) on collection and viability of equine oocytes assessed by morphology and developmental competency after intracytoplasmic sperm injection (ICSI). Reprod Fertil Dev 2009;21:615–23. 11. Jacobson CC, Choi YH, Hayden SS, Hinrichs K. Recovery of mare oocytes on a fixed biweekly schedule, and resulting blastocyst formation after intracytoplasmic sperm injection. Theriogenology 2010;73:1116–26. 12. Choi YH, Chung YG, Walker SC, Westhusin ME, Hinrichs K. In vitro development of equine nuclear transfer embryos: effects of oocyte maturation media and amino acid composition during embryo culture. Zygote 2003;11:77–86. 13. Li X, Morris LHA, Allen WR. Influence of co-culture during maturation on the developmental potential of equine oocytes fertilized by intracytoplasmic sperm injection (ICSI). Reproduction 2001;121:925–32.
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14. Hinrichs K, Choi YH, Walckenaer BE, Varner DD, Hartman DL. In vitro-produced equine embryos: production of foals after transfer, assessment by differential staining and effect of medium calcium concentrations during culture. Theriogenology 2007;68:521–529. 15. Galli C, Colleoni S, Duchi R, Lagutina I, Lazzari G. Developmental competence of equine oocytes and embryos obtained by in vitro procedures ranging from in vitro maturation and ICSI to embryo culture, cryopreservation and somatic cell nuclear transfer. Anim Reprod Sci 2007;98:39–55. 16. Cochran R, Meintjes M, Reggio B, Hylan D, Carter J, Pinto C, Paccamonti D, Godke RA. Live foals produced from
sperm-injected oocytes derived from pregnant mares. J Equine Vet Sci 1998;18:736–40. 17. McKinnon AO, Lacham-Kaplan O, Trounson AO. Pregnancies produced from fertile and infertile stallions by intracytoplasmic sperm injection (ICSI) of single frozen-thawed spermatozoa into in vivo matured mare oocytes. J Reprod Fertil Suppl 2000;56:513–17. 18. Hinrichs K, Choi YH, Norris JD, Love LB, Bedford-Guaus SJ, Hartman DL, Velez IC. Use of intracytoplasmic sperm injection and in vitro culture to the blastocyst stage for clinical production of foals post mortem. Anim Reprod Sci 2010;121:239–40 (abstract).
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Chapter 313
Oocyte Cryopreservation Lisa J. Maclellan
Introduction Unlike embryo freezing, cryopreservation of oocytes is not yet considered an established technology, despite there being significant interest in trying to improve the technology. The ability to store oocytes successfully at −196◦ C has numerous practical, financial, and ethical considerations, and may have a positive impact on animal breeding programs. Cryopreservation of oocytes from live mares or mares that have recently died will allow storage of valuable genetic material and may eliminate the need to fertilize oocytes immediately or discard unused oocytes. Oocyte cryopreservation is also applicable to other species. In commercial livestock, improvements in oocyte cryopreservation would greatly improve genetic management of populations and the number of offspring generated. Improvement in the ability to cryopreserve oocytes or embryos of endangered animals could lead to an increase in the number of offspring produced, enhancement of the genetic quality of animals, improvement in the population’s genetic health, elimination of both the cost and risk of transporting live animals for reproductive purposes, and possibly even delay or prevent the extinction of some rare species. In humans, cryopreservation of oocytes is particularly important for women who want to harvest genetic material prior to treatments that may damage gametes such as radiation or chemotherapy for cancer. There are also ethical, legal, and logical reasons for wanting to freeze human oocytes instead of embryos.
Cryopreservation procedures There are two types of cryopreservation procedures, namely equilibrium freezing, referred to as slow rate freezing, and non-equilibrium freezing, referred to as ultra-rapid cooling or vitrification. Slow rate cooling works better for oocytes that are not sensitive to
chilling injury, in species such as the mouse, cat, rabbit, and human but not for the pig, cow, and most likely the horse.1 Advantages of slow rate cooling are that a low concentration of cryoprotectants is generally used and that the oocytes are slowly dehydrated. The disadvantage is the possibility of chilling injury due to ice crystal formation within the cytoplasm.2 An alternative is vitrification in which rapid freezing results in solidification of a solution at a low temperature without ice crystal formation. The advantage of vitrification is the elimination of ice crystal formation; however, all other types of injuries are increased by vitrification, and the oocytes are exposed to very high concentrations of cryoprotectants. The storage, warming, and rehydration (i.e., removal of cryoprotectants) differ only slightly between the two methods, The main difference exists in the addition of cryoprotectants in freezing and cooling rates.3 All oocytes and embryos suffer considerable morphological and functional damage during cryopreservation. The cryopreservability of oocytes is complicated by their structural complexity, high surface area to volume ratio, and low plasma membrane permeability to cryoprotectants;4 thus, the extent of the injury depends on many of these factors. Oocytes of different species also vary in size, membrane composition, and physiology; therefore, attempts to optimize cryopreservation techniques for oocytes have included the development of species-specific techniques,5, 6 rather than relying on the technologies established for the preimplantation embryo. Bogliolo et al.7, 8 reported differences in cryopreservation and concluded that the procedures for one species were not necessarily suitable for other species. For example, higher rates of meiotic progression and normal spindle configuration were seen with immature equine oocytes than with ovine oocytes using the same vitrification procedure. Some of the advances with oocytes from species other than the horse include using a combination of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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cryoprotectants, stepwise addition, and removal of cryoprotectants and the addition of macromolecules to the cryoprotectant solution. More recently, it has been suggested that altering the fundamental characteristics of the cells themselves9 and analyzing the effects of cryoprotectants on cell physiology10 may improve survival rates of oocytes more than focusing on the physical and chemical aspects of the cryopreservation procedure. New devices such as the use of open-pulled straws,5 cryoloops,11 electron microscope grids,12 and cryotops6, 13 have allowed the ultra-rapid freezing of oocytes, and this appears to improve survival rates. In the bovine, Maverides and Morroll14 reported a 54% survival of bovine oocytes after slow cooling compared with a 90% survival of oocytes vitrified on cryoloops. Lane and Gardner15 also reported higher survival rates for mouse oocytes frozen on cryoloops and vitrified than for those that were slow cooled. Blastocyst rates after fertilization of cryopreserved bovine oocytes using open-pulled straws2 and cryotop vitrification16 almost approached the value of unfrozen controls.
Cryopreservation of equine oocytes Studies on cryopreservation of equine oocytes have been extremely limited. These studies have examined both slow rate freezing and vitrification, as well as the effects of stage of nuclear maturation, inclusion of macromolecules to reduce the toxic effects of cryoprotectants, in vivo matured oocytes compared with in vitro matured oocytes collected from slaughterhouse ovaries, and partially mature oocytes collected from superstimulated and non-stimulated mares. The question of which meiotic stage is most suitable for oocyte cryopreservation in the horse has focused on the cryopreservation of mature oocytes, but chromosome scattering by microtubular disruption of the meiotic spindle occurs in metaphase II (MII) oocytes. Because the meiotic spindle of these oocytes is extremely susceptible to damage during freezing, this has been proposed to be central to their poor post-thaw developmental competence.17, 18 On the other hand, immature oocytes at the germinal vesicle stage do not present an organized meiotic spindle and should be more resistant to cryopreservation because their chromatin is protected within a membrane-bound vesicle. In studies on the cryopreservation of equine germinal vesicle stage cumulus–oocyte complexes, the success of subsequent in vitro maturation has been disappointing. Hochi et al.19, 20 reported cryopreservation of germinal vesicle equine oocytes with ethylene glycol using either a slow cool or vitrification protocol. The survival rates were low, being 15.8%
with the slow cool procedure and 16.7% with vitrification; nevertheless, the studies indicated that oocytes could mature to the MII stage following freezing and thawing. Ultrastructural evaluation of the oocytes indicated that freezing disrupted the intercellular communication between the cumulus cells and the oocyte.21 In a study by Hurtt et al.,22 the viability of immature and mature equine oocytes vitrified in 5 M ethylene glycol with 18% Ficoll and 0.5% sucrose using open-pulled straws was evaluated. After vitrification, equine oocytes had maturation rates up to 30%, 40%, and 46% for the immature, mature, and non-vitrified control groups, respectively. Only 25%, 30%, and 45% of the oocytes in the immature, mature, and control groups displayed mature cortical granule distribution. Based on the results of this study, both immature and mature equine oocytes survived cryopreservation using vitrification and open-pulled straws. However, survival rates were lower for cryopreserved oocytes than for the control. More promising results were obtained by Tharasanit et al.23, 24 They found that controlled rate freezing at the MII stage yielded a higher percentage of oocytes with a normal spindle (67%) than at germinal vesicle stage (1%), or with vitrification at either MII (37%) or germinal vesicle stage (52%). Subsequent studies concluded that, when vitrification was the chosen method of cryopreservation, compact equine oocytes at the germinal vesicle stage offer the best chance of an MII oocyte with a normal spindle and with the potential for fertilization; however, the developmental competence is dramatically reduced.24 These studies demonstrate that both the immature and mature equine oocytes could be vitrified, thawed, and survive. However, there seems to be a trend towards better survival for vitrification of mature oocytes, which agrees with studies in other species, in which mature oocytes were more resistant to cryopreservation. In vitro matured cat oocytes froze better than immature oocytes.25 MII bovine oocytes appear to be less susceptible to freezing damage than immature germinal vesicle (GV) oocytes.26, 27 Several approaches have been used to minimize the chilling injury in cryopreservation of oocytes, including the selection of permeable cryoprotectants. For equine oocytes that are frozen with a slow cool rate protocol, Hochi et al.19 demonstrated that ethylene glycol was the best cryoprotectant. However, for vitrification generally, a combination of cryoprotectants is used, including ethylene glycol (EG), propylene glycol, and dimethylsulfoxide (DMSO). Several studies have demonstrated the advantage in some species of replacing some of the sodium in the medium with choline. This seems to be particularly advantageous for cryopreservation of mouse28 and human oocytes.29 Another approach to decrease the
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Oocyte Cryopreservation toxicity of cryoprotectants is a stepwise addition and removal of the cryoprotectant. Generally, the cryoprotectant solution is added in three steps and removed in three steps. This minimizes the time that the oocyte is in contact with high concentrations of the cryoprotectants. In a study with equine oocytes by Maclellan et al.30 the sugars sucrose or trehalose were added to a vitrification solution containing EG and DMSO, to determine whether their addition would minimize the toxic effect of cryoprotectants. Oocytes from ovaries collected at an abattoir underwent in vitro maturation under standard conditions for 12 hours or 24 hours before being partially denuded of cumulus and then vitrified. Cryoprotectants were loaded in three steps: 5% DMSO and 5% EG for 30 seconds, 10% DMSO and 10% EG for 30 seconds, and 20% DMSO and 20% EG with either 0.65 M sucrose or 0.65 M trehalose for approximately 20 seconds, before being loaded onto a nylon cryoloop. Oocytes in the 12-hour and 24-hour groups were cultured for 14 hours and 2 hours post-thaw, respectively, before undergoing intra cytoplasmic sperm injection (ICSI). Non-vitrified oocytes matured for 26 hours were used as controls. After vitrification, live/dead status based on fluorescence of the cytoplasm and morphologically intact plasma membranes was similar for oocytes in 12 hours sucrose (6/9), 24 hours sucrose (6/9), and 24 hours trehalose (7/9) in comparison with the control group (9/9). There were significantly fewer live oocytes in the 12-hour trehalose group (5/9) (P = 0.02) after vitrification than in the control group. Fertilization rates were similar for both control and vitrified oocytes. Equine oocytes vitrified with either sucrose or trehalose at 12 or 24 hours evaluated by ICSI appeared viable, but transfer of the zygotes would be needed for accurate assessment. The benefits of the interaction between naloxone and endogenous opioid peptide receptors under different conditions of cellular stress have been demonstrated in the human platelets.31 In a study with equine oocytes, the addition of naloxone to the vitrification solutions appeared to limit damage to the oocytes during cryopreservation. Within vitrified cumulus oocyte complexes, only those with naloxone in the vitrification solutions reached the blastocyst stage at day 9.32 A study compared embryo development rates after transfer of vitrified and non-vitrified oocytes into inseminated recipient mares.33 Oocytes were collected by transvaginal ultrasound-guided follicular aspiration from superstimulated and non-stimulated mares 24 hours after administration of human chorionic gonadotropin (hCG). Oocytes for control transfer were cultured for 14–16 hours before transfer, whereas oocytes for cryopreservation were placed
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into culture medium for approximately 2–4 hours before vitrification. For vitrification the oocytes were placed in the cryoprotectant solution in three steps, as described by Maclellan et al.30 Oocytes were stored for up to 2 months before transfer. They were then thawed, cryoprotectant diluted in three steps, and cultured for 10–12 hours before transfer into inseminated recipients. Of the 26 oocytes vitrified, 19 were considered morphologically normal after thawing and culture. Of the 20 oocytes collected from nonstimulated mares, 12 were vitrified and eight were transferred as controls. After thawing, 10 of the 12 were morphologically intact and transferred in recipients, resulting in one embryonic vesicle on day 16 (one of 12, 8%). Fourteen oocytes from superstimulated mares were vitrified and four were transferred as controls. After thawing, nine of the 14 oocytes were morphologically intact and transferred into recipients, resulting in two embryonic vesicles on day 16 (two of 14, 14%). In control transfers, seven of the eight oocytes from non-stimulated mares and three of the four oocytes from superstimulated mares resulted in embryonic vesicles on day 16. The two pregnancies from vitrified oocytes resulted in healthy foals. This is the first report of foals being born from vitrified equine oocytes.
Evaluation of frozen–thawed oocytes The criteria chosen for evaluation of equine oocytes after freezing and thawing are extremely important. In the studies conducted with equine oocytes discussed above, different approaches were used including staining for nuclear and cytoplasmic maturation (the latter based on cortical granule migration) as well as transfer into the recipient’s oviduct. Initial studies indicated that the rate of nuclear maturation was reasonable, whereas the incidence of cortical granule migration was poor. The latter result may have been caused by premature release of cortical granules as the result of the cryoprotectants (or the fact that these oocytes were not cytoplasmically mature). Another technique that is extremely useful for evaluation of the viability of oocytes is the use of ICSI. Since in vitro fertilization in the horse is not a reliable technique, ICSI has become the best in vitro technique for evaluating the oocyte. It also has the advantage of inducing fertilization in oocytes that may not be capable of fertilization because of zonal hardening. Despite the success of the birth of two foals from vitrified oocytes, the techniques of oocyte cryopreservation cannot be considered to be well-established procedures. More studies need to be conducted on cryopreservation of equine oocytes.
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References 1. Ledda S, Leoni G, Bogliolo L, Naitona S. Oocyte cryopreservation and ovarian tissue banking. Theriogenology 2001;55:1359–71. 2. Vajta G. Vitrification of the oocytes and embryos of domestic animals. Anim Reprod Sci 2000;60–61:357–64. 3. Vajta G, Kuwayama M. Improving cryopreservation systems. Theriogenology 2006;65:236–44. 4. Leibo SP. Cryopreservation of oocytes and embryos: optomization by theoretical verses empirical analysis. Theriogenology 2008;69:37–47. 5. Vajta G, Helm P, Greve T, Callesen H. Vitrification of porcine embryos using the open pulled straw (OP5) method. Acta Vet Scand 1997;38:349–52. 6. Kuwayama M, Vajta G, Kato O, Leibo SP. Highly efficient vitrification method for cryopreservation of human oocytes. Reprod Biomed Online 2005;11:300–8. 7. Bogliolo L, Ariu F, Rosati I, Zedda MT, Pau S, Naitana S, Leoni G, Kuwayama M, Ledda S. (a) Vitrification of immature and in vitro matured horse oocytes. Reprod Fertil Dev 2006;18:149–50. 8. Bogliolo L, Ariu F, Fois S, Leoni G, Zedda MT, Pau S, Ledda S. Negative effect of cumulus and cytochalasin on IVM and viability of immature sheep oocytes. Reprod Domestic Anim 2006;41:342. 9. Seidel G Jr. Modifying oocytes and embryos to improve their cryopreservation. Theriogenology 2006;65:228– 35. 10. Gardner DK, Sheehan CB, Rienzi L, Katz-Jaffe M, Larman MG. Analysis of oocyte physiology to improve cryopreservation procedures. Theriogenology 2007;67:64–72. 11. Lane M, Schoolcraft WB, Gardner DK. Vitrification of mouse and human blastocysts using a novel cryoloop container-less technique. Fertil Steril 1999;72:1073–8. 12. Arav A, Zeron Y. Vitrification of bovine oocytes using modified minimum drop size techniques (MDS) is affected by composition and concentration of the vitrification solution and by cooling conditions. Theriogenology 1997;47:341. 13. Kuwayama M, Kato O. All-round vitrification method for human oocytes and embryos. J Assist Reprod Genet 2000;17:477. 14. Mavrides A, Morroll D. Cryopreservation of bovine oocytes: cryoloop vitrification the future to preserving the female gamete. Reprod Nutr Dev 2002;42:73–80. 15. Lane M, Gardner DK. Vitrification of mouse oocytes using a nylon loop. Mouse Reprod 2001;58:342–7. 16. Kelly J, Kleeman D, Kuwayama M, Walker S. Effect of cysteamine on survival of bovine and ovine oocytes vitrified using the minimum volume cooling (MVC) cryotop method. Reprod Fertil Dev 2006;18:158. 17. Saunders JM, Parks JE. Effects of cryopreservation procedures on the cytology and fertilization rate of in vitromatured bovine oocytes. Biol Reprod 1999;61:178–87. 18. Chen SU, Lien YR, Chao KH, Ho HN, Yang YS, Lee TY. Effects of cryopreservation on meiotic spindles of oocytes and its dynamics after thawing: clinical implica-
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
tions in oocyte freezing: a review article. Mol Cell Endocrinol 2003;202:101–7. Hochi S, Fujimoto T, Choi Y, Braun J, Oguri N. Cryopreservation of equine oocytes by 2 step freezing. Theriogenology 1994;42:1085–94. Hochi S. Cryopreservation of follicular oocytes and preimplantation embryos in cattle and horses. J Reprod Dev 2003;49:13–21. Hochi S, Kozawa M, Fujimoto T, Hondo E, Yamada J, Oguri N. In vitro maturation and transmission electron microscope observation of horse oocytes after vitrification. Cryobiology 1996;33:300–10. Hurtt AE, Landin-Alvarenga F, Seidel GE, Squires EL. Vitrification of immature and mature equine and bovine oocytes in an ethylene glycol, ficoll, and sucrose solution using open pulled straws. Theriogenology 2000;54:119–28. Tharasanit T, Colleoni S, Lazzari G, Colenbrander B, Galli C, Stout TA. Effect of cumulus morphology and maturation stage on the cryopreservability of equine oocytes. Reproduction 2006;132:759–69. Tharasanit T, Colenbrander B, Stout TA. Effect of maturation stage at cryopreservation on post-thaw cytoskeleton quality and fertilizability of equine oocytes. Mol Reprod Dev 2006;73:627–37. Luvoni GC, Pellizzari P. Embryo development in vitro of cat oocytes cryopreserved at different maturation times. Theriogenology 2000;53:1529–40. Lim MJ, Fukui Y, Ono H. Developmental competence of bovine oocytes frozen at various maturation stages followed by in vitro maturation and fertilisation. Theriogenology 1992;37:351–61. Otoi T, Yamamoto K, Koyama N, Suzuki T. In vitro fertilisation and development of immature and mature bovine oocytes cryopreserved by ethylene glycol with sucrose. Cryobiology 1995;32:455–60. Stachecki JJ, Willadsen SM. Cryopreservation of mouse oocytes using a medium with low sodium content: effect of plunge temperature. Cryobiology 2000;40:4–12. Quintano CJ, Donaldson MJ, Bertolin MV, Pasqualini RS. Birth of two babies using oocytes that were cryopreserved in a choline-based freezing medium. Hum Reprod 2002;17:3149–52. Maclellan LJ, Lane M, Sims MM, Squires EL. Effect of sucrose and trehalose on vitrification of equine oocytes 12 or 24 hrs after onset of maturation. Theriogenology 2001;55:310. Sheu JR, Hung WC, Lee LW, Chang PT, Kan YC, Yen MH. Mechanism involved in the antiplatelet activity of naloxone in human platelets. Biochem Biophys Res Commun 1997;231:12–16. Merlo B, Iacono E, Colleoni S, Dell Aquila E, Galli C, Mari G. Embryo development after ICSI of equine oocytes vitrified before and after IVM. Reprod Fertil Dev 2006;19: 195–6. Maclellan LJ, Carnevale EM, Coutinho da Silva M A, Scoggin CF, Bruemmer JE, Squires EL. Pregnancies from vitrified equine oocytes collected form super-stimulated and non-stimulated mares. Theriogenology 2002;58:911–19.
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Principles of Cryopreservation James K. Graham
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Chapter 314
Principles of Cryopreservation James K. Graham
Introduction Although Spallanzani was the first to attempt to freeze stallion sperm in 1776 (reported by Pesch and Hoffman1 ), it was not until Polge and colleagues2, 3 discovered the cryoprotective action of glycerol that real progress in the development of cryotechniques to preserve sperm and in the use of cryopreserved sperm occurred. Currently, sperm from many mammalian, bird, fish, and other species can be cryopreserved, with some success.4 However, sperm from some species, and from some males within a species, survive cryopreservation better than others. The reasons for these differences are numerous, but include (i) the chemical composition of the sperm, which affects the membrane permeability to water and cryoprotectant, as well as the fluidity of the sperm membranes at different temperatures; (ii) the osmotic tolerance of the sperm; and (iii) cell size and shape. The objective of this chapter is to review some of the basic principles involved in cryopreservation so that readers can understand why certain steps are conducted during the cryopreservation process.
Basic principles Diffusion of solutes in solutions and across membranes Solutes, defined as any substances (salts, sugars, proteins, or other molecules) that are dissolved in solvents (generally water), will diffuse in a solution from areas of high concentration (when they are added to the solution in one area) to areas of low concentration, until equilibrium is reached (when the concentration of solute is the same throughout the entire solution). Some compounds, such as many of the penetrating cryoprotectants (ethylene glycol, glycerol, methyl formamide, etc.), are liquids and act as both a solute (they dissolve in water) and as a solvent (salts,
sugars, and proteins dissolve in them). When adding a solute to a solution, equilibrium can be reached more quickly if the solution is physically mixed. Many of the solutes and solvents that we place cells into can permeate the plasma membrane of the cells. However, there are also many solutes that cannot pass through the plasma membrane. If a membrane separates two solutions (such as a cell’s plasma membrane separates its cytoplasm from the extracellular fluid) and additional solutes or solvents are added to the extracellular solution, solvents and/or solutes will cross the membrane (via diffusion) until a new equilibrium is achieved (Figure 314.1). Similarly, transient equilibrium conditions will occur if the solute that is added can permeate the plasma membrane, but does so at a rate that is slower than the solvent (water). This is the case for most permeating cryoprotectants. For example, although glycerol permeates the stallion sperm plasma membrane, it does so much more slowly than water.5, 6 Therefore, when glycerol is added to a sperm suspension, water will move out of each cell (to dilute the glycerol) at a much faster rate than glycerol moves into the cell, and the cell will transiently shrink.5–8 However, as the glycerol slowly diffuses into the cell, water will enter as well (keeping the overall glycerol concentration inside and outside the cell membrane similar), and, when glycerol reaches its equilibrium across the membrane, water will as well, and the cell will swell again to its original volume (Figure 314.2). On the other hand, if cells in a solution containing cryoprotectant (glycerol) are suddenly diluted into a solution that contains no cryoprotectant (either a diluent or a mare’s reproductive tract fluids), water will rush into the cells, attempting to dilute the cryoprotectant, faster than the cryoprotectant can diffuse out of the cell, and the cell will transiently swell.5–9 As the cryoprotectant slowly diffuses out of the cell, water will diffuse out with it (maintaining equilibrium), and, if the cell has not ruptured during swelling,
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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G G G G G Figure 314.1 (a) Solutions on both sides of a membrane (dashed line) that contain solutes (filled circles) that cannot pass through the membrane. When additional solutes are added to one solution (b), the solvent will cross the membrane (c), establishing a new equilibrium in which solute concentration on both sides of the membrane is equal, even though volumes are not.
the cell will eventually regain its original volume (Figure 314.3). The ability of a cell to withstand rupture during this scenario depends upon its membrane permeability to the cryoprotectant and water (note that cryoprotectants that are more permeable to the membrane will diffuse across it more rapidly and the cells will swell to a lesser degree), the cell’s osmotic tolerance (some cells can withstand greater degrees of shrinking and swelling without damage), the size of the cell, and its shape.6, 7, 10 Physical properties of solutions when they freeze When pure water freezes, the molecules form a crystalline structure composed of all of the water molecules. However, when a solution containing salts and sugars is cooled below its freezing point, the ice crystals that form contain only water molecules and these crystals are separated from each other by unfrozen water channels that contain all of the salts
G
G G
G
G
G
G
G
G G
(a)
(b)
(c)
Figure 314.3 When a cell preloaded with glycerol (G) is initially placed into an isosmotic solution (A), water will move into the cell (large arrow) at a faster rate than glycerol can move out of the cell (small arrow) and the cell will swell (b). As the glycerol slowly leaves the cell, and water leaves with it maintaining osmotic equilibrium, the cell re-establishes its original volume (c), providing the cell does not rupture.
and sugars (Figure 314.4a). In order for cells to survive cryopreservation, they must reside in these unfrozen channels, and cells that become trapped in the ice crystals die.5 As the temperature is lowered further, more water is removed from the unfrozen channels into the growing ice crystals and the remaining unfrozen solution becomes more hypertonic. Salt concentrations can increase in these unfrozen channels to the point that cell membranes, organelles, and proteins can be irreparably damaged, and the cells die.7, 9 If the temperature is reduced further still, the
G G
G
G
G
G G
G
G G
G
G
G G
G
(a)
(b)
G
G
(c)
G
(d)
Figure 314.2 When a cell in isosmotic solution (a), is placed into a solution containing glycerol (G) at concentrations required for cryopreservation (b), water will move out of the cell (large arrow) at a faster rate than glycerol moves into the cell (small arrow) and the cell will shrink (c). As the glycerol slowly enters the cell, and water moves back into the cell with it maintaining osmotic equilibrium, the cell re-establishes its original volume (d).
(a)
(b)
Figure 314.4 Depiction of a frozen solution containing ice crystals of pure water (white) and hypertonic water channels (gray) which are unfrozen (a). Sperm cells located in the unfrozen water channels will shrink from their original volume (dark gray) to a smaller volume (black) as the osmolality of the unfrozen solution increases (b).
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Principles of Cryopreservation Table 314.1 The percentage of unfrozen solution and the salt concentration of that solution when physiological saline containing 0 M or 0.5 M glycerol is cooled to –12◦ C Salt concentration Unfrozen fraction solution
(%)
(M)
Saline Saline + glycerol
5 13
3 1.5
solution in the remaining unfrozen channels ‘vitrifies’ (turns to glass without crystallization). In the vitrified state and at –196◦ C, molecular movements continue,11 but cell metabolism essentially ceases and cells can remain viable for many years.12 Adding solutes to water decreases the temperature at which the solution freezes. Therefore, while pure water freezes at 0◦ C, physiologic saline (300 mOsm saline) freezes at –0.6◦ C. When higher concentrations of solutes are added the freezing point is depressed even further. If the solute added is a liquid, such as the cryoprotectants ethylene glycol, glycerol, or methyl formamide, the freezing point is similarly depressed, but since these compounds remain unfrozen at low temperatures they will contribute volume to the unfrozen channels between ice crystals. In addition, since these compounds are permeable to the plasma membrane, the osmotic effects they exert on the cell are only transient and occur only at the time they are added or are removed. Therefore, these compounds increase the overall volume of the unfrozen channels and decrease the salt concentration witnessed by the cells, thereby decreasing the deleterious effects of both (Table 314.1). Cryodamage Potential sources of cell damage during freezing and thawing include: (i) cooling to near 0◦ C, inducing cold shock to the sperm (see Chapter 127); (ii) extracellular and, more importantly, intracellular ice formation;13 (iii) changes due to dehydration brought about by extracellular ice formation, including high salt concentration and membrane distortion due to cell shrinking and swelling;7 and (iv) osmotic changes during the addition and removal of cryoprotectants.5–7,9 For most cell types, both intracellular ice formation and membrane damage are major causes of cell death.6, 7, 9, 13, 14 However, owing to the low amount of active water inside a spermatozoa (∼15%) the formation of intracellular ice is not a major problem for sperm.15 No significant damage occurs to cells once the unfrozen channels have vitrified. Therefore, storage at –196◦ C is not a problem. However, changes that
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occur to the extracellular medium and the cells between –15◦ C and –60◦ C, can induce cellular damage.7, 9 The rate at which a solution is cooled through this zone affects the rate of ice formation and the hypertonicity changes in the unfrozen channels, before vitrification takes place. Cell survival depends upon the interactions of plasma membrane permeability and these changes. If cells are cooled too rapidly through this temperature zone, when extracellular ice forms creating hypertonic unfrozen channels, intracellular water does not have time to leave the cell and equilibrate across the membrane before substantially lower temperatures induce intracellular ice.5, 9 On the other hand, if the temperature is reduced too slowly through this temperature range, although intracellular water can equilibrate across the membrane, the cells can be exposed to excessive hypertonic conditions for too long, before the solution vitrifies, and these high salt ‘solution effects’ can damage cell membranes and organelles.5, 9 Therefore, an optimal cooling rate exists which optimizes cell cryosurvival; however, this rate can be different for different cell types and for similar cells cryopreserved in different diluents. For stallion spermatozoa, cooling rates between –10◦ C/min and –60◦ C/min work well, depending upon the cryodiluent used.9 To complicate matters further, the optimal warming rate, for a frozen sperm sample, is dependent upon the cooling rate used, and cells that are cooled rapidly need to be warmed rapidly, while cells that cooled slowly need to be warmed slowly. The theoretical reasons for this have been reviewed previously,5, 7, 9 but, for practical purposes, the cooling rates used to cryopreserve stallion sperm are intermediate between the two extremes and, in most cases, thawing straws in a water bath at 37–39◦ C (providing an intermediate warming rate) works well. Having said this, it should be noted that sample size can affect the cooling and warming rate of a sample significantly. For example, if one sets a 1-µL droplet of diluent on the surface of dry ice, the entire droplet will cool at roughly the same rate. On the other hand, if one places a 100-mL volume on the dry ice, the solution in direct contact with the dry ice surface will cool rapidly while the solution furthest away from the dry ice will remain unfrozen for a significant period of time, until the solution immediately beneath it freezes. There will be a wide range of freeze rates for large volumes, the fastest being near the cooling sources and the slowest in the fluid farthest form the cooling source. Cryoprotectants A cryoprotectant can be defined as any substance that when added to cells prior to cryopreservation
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improves the cryosurvival of the cells. There are two major classes of cryoprotectants, those that are impermeable to the cell plasma membrane and those that are permeable. Non-permeable cryoprotectants include sugars (including lactose, raffinose, and sucrose), lipoproteins (including egg, milk, and serum) and other macromolecules (including methyl cellulose, polyvinylalcohol, and polyvinylpyrrolidone). These molecules may interact with plasma membrane components helping to stabilize the membrane, but their major effect is to create a hyperosmotic environment, which will induce cellular dehydration (see above). Penetrating cryoprotectants, such as ethylene glycol, glycerol, dimethylsulfoxide, and various amides, are generally more effective cryoprotectants than non-penetrating cryoprotectants, because they affect several cell and solution properties. First, they replace water inside the cell. This dehydrates the cell and the compounds tend to inhibit the formation of intracellular ice.5, 6 In addition, they increase the volume of the unfrozen channels between the extracellular ice crystals, thereby increasing the space available for the cells; and they decrease the salt concentration in the unfrozen solution.5 However, because all of the molecules permeate the cell plasma membrane more slowly than water, they induce transient osmotically driven cell size changes that can damage a cell. Sperm from some species appear to tolerate these osmotic challenges relatively well (human and bovine) while others do not (poultry and equine). Therefore, the amount of cryoprotectant that is added to the cryopreservation diluent for any cell type is a trade-off between the amount of cryoprotectant needed to optimize cryosurvival and the amount of cryoprotectant that the cells can tolerate. Demick et al.16 demonstrated the deleterious effect glycerol has on stallion sperm when they reported that simply adding glycerol to cooled stallion sperm and subsequently inseminating those sperm resulted in a fertility rate of only 17%, compared with that of control cooled sperm (44% fertility rate). Because stallion sperm are quite sensitive to the osmotic effects induced by adding and removing glycerol, while bull sperm are generally cryopreserved using 7–8% glycerol, cryopreservation diluents for stallion sperm generally contain only 4–5% glycerol.
Cryopreserving stallion sperm Understanding these basic principles provides insight into what happens to stallion spermatozoa when they are cryopreserved. After collection and an initial evaluation, the ejaculate is usually diluted in a centrifugation medium. This is an isosmotic medium designed to maintain the spermatozoa during cen-
Cell Volume (%)
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Dilute Thaw Freeze
100
Time Figure 314.5 Depiction of cell volume changes that are induced by adding a diluent containing glycerol, by dehydration during freezing, rehydration during thawing, and by dilution into a glycerol-free medium. From Hammerstedt et al.7 with permission from the American Society of Andrology.
trifugation and can be of quite simple to very complex composition. Following centrifugation, the spermatozoa are diluted into cryopreservation diluent containing cryoprotectants. The suspension of sperm in this diluent initiates the first osmotic imbalance across the sperm plasma membrane, and the cells rapidly shrink. Depending upon the specific cryoprotectant used and the temperature the cryoprotectant is added at, within approximately 1 minute, the cryoprotectant has entered the cell, reaching equilibrium, and, as water re-equilibrates across the membrane with the cryoprotectant, the cell regains its original volume (Figure 314.5). Some cryopreservation diluents are most effective if the sperm are cooled slowly to 5◦ C over 1–3 hours, while others (the lactose–EDTA diluent) can effectively preserve spermatozoa when they are frozen directly from room temperature. Regardless of the method utilized, little change in cell volume occurs when the cells are cooled from room temperature to a few degrees below 0◦ C. Other changes, specifically having to do with membrane phase changes and membrane component rearrangements, occur during transit through this temperature range (see Chapter 127). However, as the extracellular solution begins to freeze, and water is lost from the system, in the form of ice, the cells are exposed to hypertonic extracellular fluid and adjust osmotically by losing water to the extracellular fluid, and begin to shrink (Figures 314.4b and 314.5). As more and more water is lost to extracellular ice crystals, the cells shrink further. In addition, the volume of the unfrozen solution decreases. As the cells shrink, their membranes become deformed and these deformations can result in membrane damage.7 Since only cells in the unfrozen solution survive cryopreservation, the volume of this solution limits the number of cells that can effectively be preserved in a specific volume of diluent. What this means is that one cannot put billions of sperm into 0.5 mL of diluent and expect all of the cells to survive, there simply is not sufficient unfrozen volume
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the cells do not burst), as the cryoprotectant leaves the cell slowly over the next 60 seconds or so, the cell volume will return to normal (Figure 314.5).
60
Percent Motile
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50 40 30
References
20 10 0 100
200
400
800
1600
Sperm concentration Figure 314.6 The percentages of motile stallion spermatozoa after cryopreservation, for spermatozoa that had been diluted to different spermatozoal concentrations prior to freezing. Values for light gray bars are significantly different than values for dark gray bars (P < 0.05).
for all of those cells to fit. The upper limit of sperm concentration that maximizes cell survival is between 400 and 800 million sperm/mL (Figure 314.6). Once the solution temperature reaches approximately –60◦ C to –80◦ C (depending upon diluent and cryoprotectant used, as well as cryoprotectant concentration) it vitrifies into a glass and all cellular metabolism ceases. At this point, the cells are essentially inert and can be stored in liquid nitrogen for long time periods without significant deterioration.12 It should be pointed out that straws of volumes 0.25 and 0.5 mL change temperature rapidly when exposed to different temperatures. Therefore, it is very important when handling these straws to keep them submersed in liquid nitrogen as much as possible and limit the time straws are exposed to warmer temperatures when straws are moved from one tank to another, as well as the number of times straws are transferred, as these exposures can result in the cells warming sufficiently to cause irreparable damage.12 When the straws are warmed during thawing, the ice in the system melts, returning water to the system, thereby lowering the salt concentration in the solution. The cells respond by rehydrating to their original volume. Finally, when the cells are diluted into a solution that contains no cryoprotectant, either a dilution medium prior to insemination or the mare’s reproductive tract fluids, since the sperm are now loaded with cryoprotectant, there is an osmotic gradient created across the plasma membrane and water rushes into the cells, before the cryoprotectant can leave, and the cells swell. If the tolerance of the plasma membrane stretching is not exceeded (i.e.,
1. Pesch S, Hoffman B. Cryopreservation of spermatozoa in veterinary medicine. J Reproduktionsmed Endokrinol 2007;4: 101–5. 2. Polge C, Smith AU, Parkes AS. Revival of spermatozoa after vitrification and dehydration at low temperature. Nature 1949;164:666. 3. Smith AU, Polge C. Survival of spermatozoa at low temperatures. Nature 1950;166:668–9. 4. Graham EF, Schmehl ML, Deyo RCM. Cryopreservation and fertility of fish, poultry and mammalian spermatozoa. In: Proceedings of the Xth Technical Conference of Artificial Insemination, and Reproduction. Columbia, MO: National Association of Animal Breeders, 1984; pp. 4–29. 5. Amann RP. Cryopreservation of sperm. In: Knobil E, Neill JD (eds) Encyclopedia of Reproduction, vol. 1. New York: Academic Press, 1999; pp. 773–83. 6. Meryman HT. Cryopreservation of living cells: principles and practice. Transfusion 2007;47:935–45. 7. Hammerstedt RH, Graham JK, Nolan JP. Cryopreservation of mammalian sperm: what we ask them to survive. J Androl 1990;11:73–88. 8. Hammerstedt RH, Graham JK. Cryopreservation of poultry sperm: the enigma of glycerol. Cryobiology 1992;29:26–38. 9. Amann RP, Pickett BW. Principles of cryopreservation and a review of cryopreservation of stallion of stallion spermatozoa. J Equine Vet Sci 1987;7:145–73. 10. Watson PF, Kunze E, Cramer P, Hammerstedt RH. A comparison of critical osmolality and hydraulic conductivity and its activation energy in fowl and bull spermatozoa. J Androl 1992;13:131–8. 11. Mullhaupt JT. Molecular factors affecting gametic integrity during storage at low temperatures. In: The National Research Council (Ed) The Integrity of Frozen Spermatozoa. Washington, DC: National Academy of Sciences, 1978; pp. 126–43. 12. Foote RH. Maintenance of fertility of spermatozoa at 196◦ C. In: The National Research Council (Ed) The Integrity of Frozen Spermatozoa. Washington, DC: National Academy of Sciences, 1978; pp. 144–62. 13. Mazur P. Freezing of living cells: mechanisms and implications. Am J Physiol 1984;247:C125–42. 14. Steponkus PL, Dowgert MF, Gordon-Kamm WJ. Destabilization of the plasma membrane of isolated plant protoplasts during a freeze-thaw cycle: the influence of cold acclimation. Cryobiology 1983;20:448–65. 15. Morris GJ, Faszer K, Green JE, Draper D, Grout BW, Fonseca F. Rapidly cooled horse spermatozoa: loss of viability is due to osmotic imbalance during thawing, not intracellular ice formation. Theriogenology 2007;68:804–12. 16. Demick DS, Voss JL, Pickett BW. Effect of cooling, storage, glycerolization and spermatozoal numbers on equine fertility. J Anim Sci 1976;43:633–7.
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Chapter 315
Semen Extenders for Frozen Semen Harald Sieme
Currently, freezing protocols for stallion semen involve a two-step dilution procedure in which semen is first diluted with a primary extender for concentration of spermatozoa by centrifugation, and second diluted prior to freezing in an extender containing a cryoprotectant. Alternatively, an open-ended artificial vagina1 or automated dummy mare (Equidame phantom, Haico Oy, Loimaa, Finland)2 can be used to collect sperm-rich fractions for prompt cryopreservation in freezing extender without centrifugation. Currently, it is not clear which (if any) extender is clearly superior to others for all stallions. However, some extenders may suit one stallion better than others.
Prefreeze centrifugation extenders The first dilution employs either saline/sugar extenders (e.g. glucose-ethylene diamine tetra-acetate [EDTA], citrate-EDTA) or solutions of skim milk and a sugar (e.g. KenneyTM , E-Z-MixinTM , INRA-82) used to dilute fresh semen. Examples of extenders are given in Table 315.1. The functional role of centrifugation extender is to maintain motility and protect sperm during centrifugation. The composition and concentration of seminal plasma could have harmful effects on stallion spermatozoa before cooling and storage at 5◦ C. Thus, centrifugation of semen is an approach to minimize potentially deleterious effects of seminal plasma on semen quality and to increase sperm concentration. Moore et al.3 demonstrated the deleterious effect of seminal plasma on stallion spermatozoa during cryopreservation. However, retention of 5–20% seminal plasma in the suspension after centrifugation has been considered to be important for cryosurvival. The dilution rate is either 1:1 or the semen is diluted to a concentration of ∼ 50 million spermato-
zoa/mL. The success of centrifugation depends on its duration (10–15 minutes) and force (350–700g). Centrifugation with lower force (600g for 10 minutes) compared with high-force centrifugation (1000g or 2000g for 2 minutes) improved sperm quality4 when no centrifugation cushion was used. Cochran et al.5 introduced a cushion centrifugation technique in which a denser medium (0,25ml; glucose-EDTAbuffer or freezing extender) is layered below the extended semen on the bottom of the centrifuge tube to provide some protection to spermatozoa floating into the cushion. However, the beneficial effect of the cushion technique on sperm quality remains unclear.6 The loss of spermatozoa associated with centrifugation is a real disadvantage, especially in stallions producing low numbers of spermatozoa. To overcome this problem, techniques using high-speed cushioned centrifugation – i.e., with a denser medium (60% wt/vol. iodixanol in water [OptiprepTM , Axis-Shield, Oslo, Norway; EqcellsireTM Component B, IMV, L’Aigle, France; CushionFluidTM , ¨ Landshut, Germany]) layered on the botMinitub, tom of centrifuge tube under the extended semen on which the spermatozoa float during centrifugation – are proposed to improve sperm recovery and/or quality. In order to increase sperm recovery after centrifugation, the use of high-speed centrifugation (20 minutes at 1000g) with a liquid cushion and subsequent recovery of sperm-rich fraction layered in a band below the supernatant and above the cushion material has been introduced into laboratory practice.7 The type of iodixanol cushion medium used R Component B, or Cush(i.e., OptiPrepTM , Eqcellsire TM ion Fluid ) does not impact post-centrifugation semen quality. The use of extenders with high viscosities may result in reduced sperm sedimentation rates
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 315.1 Prefreeze/centrifugation extenders and freezing extenders used for cryopreservation of equine semen Prefreeze/centrifugation extender
Freezing extender
Author(s)
Composition
Quantity
Composition
Quantity
Martin et al.12
Glucose Disodium EDTA Sodium citrate dihydrate Sodium bicarbonate Streptomycin Penicillin Distilled water to make
6g 0.37 g 0.375 g 0.12 g 0.05 g 50 IU 100 mL
11% (w/v) lactose solution Centr. extender Egg yolk Equex STM Glycerol
50 mL 25 mL 20 mL 0.8mL 5.0 mL
Tischner1
Collection of sperm-rich fraction
Lactose Disodium EDTA Sodium citrate Sodium bicarbonate Egg yolk Glycerol Deionized water
11 g 0.1 g 0.089 g 0.008 g 1.6 g 3.5 mL 100 mL
Loomis et al.43
Glucose monohydrate Sodium citrate dihydrate Disodium EDTA Sodium bicarbonate Polymyxin B sulfate Distilled water to make
11% (w/v) lactose solution Centr. extender Egg yolk Equex STM Glycerol
50 mL 25 mL 20 mL 0.5 mL 5.0 mL
11% (w/v) lactose solution Centr. extender B Egg yolk Equex STM 11% (w/v) lactose solution
50 mL 25 mL 20 mL 0.5 mL 50 mL
Add glycerol
2.5%
11% sucrose solution Skim milk Egg yolk Glycerol
100.0 mL 45.0 mL 16.0 mL 6.0 mL
Cochran et al.5 Glucose Sodium citrate dihydrate Disodium EDTA Sodium bicarbonate Polymyxin B sulfate Deionized water Palmer13 mod. Vidament et al.44
He,45 Piao and Wang46
59.985 g 3.7 g 3.699 g 1.2 g 1 × 106 1000 mL 1 litre A 1.5 g 25.95 g 3.699 g 1.2 g 1000 mL
B (cushion) 59.985 g 3.7 g 3.699 g 1.2 g 1 × 106 IU 1000 mL
Modified INRA82 with 20 mM Hepes and 2% clarified egg yolk Glucose (anhydrous) 25 g Lactose × 1H2 O 1.5 g 1.5 g Raffinose × 5H2 O Sodium citrate × 2H2 O 0.25 g 0.41 g Potassium citrate × 1H2 O Hepes 4.768 g Water (apyrogenic) QSP 0.5 L Skim milk UHT 0.5 L Gentamicin sulfate 50 mg Penicillin G 50.000 IU Sucrose solution
11%
(Continued)
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Table 315.1 (Continued) Prefreeze/centrifugation extender
Freezing extender
Author(s)
Composition
Quantity
Composition
Quantity
Burns and Reasner47
Glucose Sucrose BSA Ticarcillin sterile water ad
3g 5g 1.5 g 0.1 g 100 mL
Glucose Sucrose Non-fat dry skim milk Ticarcillin sterile water ad Clarified egg yolk Glycerol
0.9 g 7g 2.4 g 0.1 g 100 mL 4% 2%
Wilhelm et al.48
HBS Hepes buffered saline medium NaCl 37.0 mM KCl 10.0 mM KH2 PO4 1.2 mM NaHCO3 35.7 mM 2.4 mM MgSO4 Hepes 10.0 mM 1.7 mM CaCl2 × 2H2 O Fructose 84.0 mM Glucose 5.5 mM Na pyruvate 1.2 mM Na lactate 13.1 mM BSA fraction V 3.0 mg/mL
Colorado extender NaCl KCl KH2 PO4 NaHCO3 MgSO4 Hepes CaCl2 × 2H2 O Fructose Glucose Na pyruvate Na lactate BSA fraction V Dried skim milk Egg yolk Glycerol
18.5 mM 5.0 mM 0.6 mM 17.8 mM 1.2 mM 5.0 mM 0.8 mM 42.0 mM 18.5 mM 0.6 mM 6.55 mM 1.6 mg/ml 51.5 mg/mL 2% (v/v) 3% (v/v)
UHT skim milk
INRA82 Egg yolk (v/v) Glycerol (v/v)
2% 2.5%
Vidament18
BSA, bovine serum albumin; EDTA, ethylene diamine tetra-acetate; UHT, ultra heat-treated.
during centrifugation. Alternatively, the use of glass nipple-bottom tubes (Pesce Laboratory Sales, Kenneth Square, PA, USA) combined with a small volume (∼ 30 µL) of cushion fluid enables, after aspiration of supernatant, suspension of sperm pellet and cushion fluid in extender for prompt use in AI.8 It has been reported that high-speed cushioned centrifugation of stallion semen had no detrimental effect on fertility.9
Frozen semen extenders The first successful pregnancy achieved by frozenthawed semen was obtained after inseminating epididymal stallion spermatozoa cryopreserved in a heated whole-milk extender containing 10% glycerol.10 Successful freezing extenders combine nutrients as a source of energy (sugars), a source of lipoproteins (e.g. milk, egg yolk, or a combination of both) to facilitate protection against rapid cooling, antibiotics to inhibit bacterial growth, buffers to balance
pH and osmotic pressure, and a cryoprotectant. Examples of freezing extenders are given in Table 315.1. Various extender types have been used to process stallion semen for freezing. No single extender has been identified as superior for this purpose. To our knowledge, two or three extenders are mainly used today for the freezing of stallion spermatozoa worldwide. Referring to a survey conducted by Samper and Morris,11 a lactose-EDTA extender12 was most commonly used, followed by INRA82 extender13 with varying concentrations of egg yolk and glycerol. To a lesser extent, a sucroseegg yolk extender (modified KenneyTM ) was used. Limited comparisons of these extenders have reported higher post-thaw motility when semen was diluted in INRA82 versus lactose-glucose-EDTA.14 Replacing glucose-EDTA solution with KenneyTM extender in a modified lactose-glucose-EDTA extender increased post-thaw motility of stallion spermatozoa.15, 16 When stallion semen was frozen in KenneyTM extender instead of INRA82, post-thaw
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Semen Extenders for Frozen Semen motility was higher, the percentage of intact acrosomes was lower, and pregnancy rates were reduced (pregnancy/cycle; KenneyTM extender 38%, n = 48; INRA82 63%, n = 59).17 The lowered fertility has been discussed with respect to the presence of bicarbonate in KenneyTM ;18 bicarbonate can induce capacitation of stallion spermatozoa. In a recent study19 it was reported that fertilization rates after in vitro fertilization (IVF) of oocytes were significantly higher for sperm frozen in E-Z FreezinTM -”MFR5” (4% egg yolk) than for sperm from the same ejaculate frozen in E-Z FreezinTM -”LE” (approximately 20% egg yolk). These authors suggest that it might be possible that the lower concentration of egg yolk in E-Z FreezinTM ”MFR5” extender is associated with more efficient removal of cholesterol from the plasma membrane of spermatozoa during capacitation. Freezing extenders or a combination of centrifugation and freezing extenders are often protected by patent and are commercially available as ready-touse liquid formulations – e.g. E-Z FreezinTM -”LE” (lactose-EDTA) or E-Z FreezinTM -”MFR5” (modified French) (Animal Reproduction Systems, Chino, US); GhentTM (Minitube, Landshut, Germany); EqcellSire A+BTM (IMV, L’Aigle, France); BotuCrioTM (www.criovital.com); INRAFreezeTM R (IMV, L’Aigle, France); and EquiPro CryoGardTM (www.minitube.com). Home-made extenders may vary in quality whereas commercial extenders are convenient, sterile, quality tested, and standardized. Consequently, readers are generally encouraged to use these extenders. Errors and/or contamination during dissolution of dry extender preparations regularly lead to low quality of the extended semen. With a ready-to-use extender, these problems can generally be avoided. However, it is reported that the process of sterilization may affect interaction between spermatozoa and extender and may subsequently have negative effects on sperm quality.20
Freezing extender composition
Lipid-based cryoprotectants (egg yolk, milk, phosphatidylcholines) The sperm plasma membrane serves as the main physical barrier to the outside environment and is a primary site of freeze–thaw damage. Such damage includes membrane destabilization due to lateral lipid rearrangement, loss of lipids from the membrane, and peroxidation of membrane lipids as a result of formation of reactive oxygen species (ROS). These events can affect sperm structure and functional signaling pathways; as a result, the fertilizing ability of the sperm cell is compromised. Lipids in egg yolk, and to a lesser extent in milk,
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are the most common lipids in freezing diluents for stallion semen. The role that lipids play in protecting spermatozoa during the freeze–thaw cycle is under investigation. Proposed mechanisms range from reversible binding of exogenous lipids in stallion seminal plasma to fusion of liposomes with the sperm membrane. Egg yolk and milk, being products of animal origin, represent a potential risk of contamination of semen, and their compositions are not uniform. The aim of ongoing studies is to replace or modify material of animal origin (e.g. milk, egg yolk) that is currently included in all cryopreservation media used in horses in order to develop a ready-to-use and chemically-defined freezing extender. The ultimate objective of this work is to offer stallion owners a new freezing extender with an optimum composition that complies with all biosafety requirements. Milk and milk by-products For sperm preservation, heated skim milk or whole milk are used as buffers in which semen is directly diluted and can be stored cooled or frozen in the presence of glycerol. Today extenders for horse sperm contain milk, with the preference for fat-reduced skim milk or skim-milk powder. Older extenders contained gelatine protein as well, but these extenders were not considered to be superior to other extenders so they are no longer used. Non-fat dried skimmilk glucose (NFDSMG) extender – named after the inventor, Dr Robert M. Kenney – is the extender of choice of many, and is available as a centrifugation extender in a commercially available version (e.g., E-Z MixinTM ; Animal Reproduction Systems, Chino, CA, USA). The use of NFDSMG extenders is recommended both for storage of stallion sperm and for microscopic semen evaluation of sperm motion characteristics. When centrifugation and cooled-storage of semen is necessary, Padilla and Foote21 reported better sperm quality with the use of Kenney’s skim milk-glucose supplemented with a modified Tyrode’s medium (KMT), which is modified in its composition (65% Kenney, 35% modified Tyrode’s medium). Another extender, INRA82, used widely in Europe, contains ultraheated skim milk as the milk component. Milk is a physiological, complex and variable medium, and its mechanism of action for preservation is not well known. Moreover, some components can be deleterious to sperm cells. The important role of milk and milk by-products is the prevention of cold shock through protection to spermatozoa during cooling prior to freezing. It is speculated that this mechanism is due to the lipid and lipoprotein content of milk. The lipoproteins are adsorbed to the plasma membrane of spermatozoa and are thought to improve stabilization of
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membranes during the freeze–thaw cycles. But because skim milk (which is free of lipids) is as efficient as whole milk in protecting sperm during semen cooled-storage or during cryopreservation, lipids do not seem to be the constituent responsible for the protection. Besides components in milk with beneficial effects on spermatozoa, detrimental components can also be included. Researchers at INRA (Nouzilly, France) demonstrated that milk components have a variable effect on sperm survival in stallions. Among the fractions purified from milk by microfiltration, ultrafiltration, diafiltration, and freeze-drying, native phosphocaseinate (NPPC) was found to be the most efficient component for preserving motility and maintaining fertility of equine sperm stored for 3 days compared with standard extenders.22 The NPPC is composed of milk caseins (α, β, κ), and the purification method used maintains their three-dimensional structure and the native interactions that occur between them. Furthermore, it has been demonstrated that β-lactoglobulin, one of the soluble milk proteins, also has a protective effect on stallion spermatozoa. The optimal concentration was 45 g/L when sperm cells were stored at 4◦ C and 15 g/L when stored at 15◦ C (β-lactoglobulin concentration in cow’s milk is 2.7 g/L). However, the protection afforded by β-lactoglobulin was lower than the protection afforded by NPPC. The resulting extender – INRA96 – composed of modified Hanks’ salt solution (HGLL-BSA medium (HGLL-BSA = Hank’s salts solution supplemented with Hepes, glucose, lactose (HGLL) supplemented with BSA) supplemented with NPPC, effectively maintains sperm longevity. It is produced by IMV Technologies (L’Aigle, France) under license from INRA. The mechanism by which casein micelles protect sperm during storage has not been fully explained, but it is hypothesized that milk (homogenized or skimmed) contains factors that are able to sequestrate lipidbinding proteins in stallion seminal plasma (HSPproteins (horse seminal plasma – proteins)); the latter are regarded as detrimental factors of seminal plasma during storage of stallion semen. Thus, the main mechanism of sperm protection by milk may involve an interaction of stallion seminal plasma proteins with casein micelles. Furthermore, the addition of lactose to extender containing caseins improved the extender efficiency during sperm freezing. Thus, lactose also appears to be involved in sperm protection by milk. However, milk filtrate, which contains only lactose and minerals, is not sufficient to protect stallion sperm during storage.23 Lactose is also a common component of egg yolk-based extenders used to freeze stallion sperm. Thus, lactose seems to improve the extender’s efficiency but is not sufficient to protect sperm itself.
In Germany, an extender containing defined caseinates and whey proteins and an extended range of ¨ Tiefenbach, Germany) sugars (EquiProTM ; Minitub, instead of skim-milk powder was superior to Kenney’s skim-milk extender when semen was cooledstored for 48 h.23 Frozen semen extenders, containing purified fractions of milk caseins are now available INRAR (IMV-Technologies, L’Aigle, France), and Freeze R EquiPro CryoGardTM (www.minitube.com). Egg yolk Egg yolk is generally used in extenders at various concentrations ranging from 2% to 25% (v/v). Evidence indicates that low-density lipoproteins (LDL) are the egg yolk compounds responsible for sperm protection. For semen evaluation, the addition of egg yolk to semen diluent is unfavorable due to its opacity; therefore many laboratories use clarified egg yolk suspensions obtained by centrifugation. Detergents, for example Equex STMTM (delta-amino-sodium lauryl sulfate) may be added to solubilize the egg yolk lipids and lipoproteins to enhance their interactions with the sperm membrane.24 Seminal plasma can be detrimental to stallion sperm during cooled-storage or cryopreservation.3 Unfortunately, the factors responsible for this detrimental effect on sperm storage have been poorly characterized. It is speculated that LDL associate with the sperm membrane and provide protection to sperm by stabilizing the membrane. Furthermore, it is speculated that phospholipids present in LDL protect sperm by forming a protective film on the sperm surface or by replacing sperm membrane phospholipids of seminal plasma that are lost or damaged during cryopreservation. A further hypothesis suggests that LDL compete with detrimental seminal plasma peptides in binding to the sperm membrane and thus protect the sperm. The incorporation of non-chicken egg yolk in sperm cryodiluents revealed that motility parameters of stallion sperm were improved when the semen is frozen in lactose EDTA extender supplemented with duck egg yolk rather than chicken egg yolk. This may be attributed to the higher levels of protein, lipids, and cholesterol present in duck compared to chicken egg yolk.25 Phosphatidylcholines Substitution of egg yolk with soybean lecithin in semen extenders has been used successfully for cryopreservation of semen in cattle.26 A modification of this extender, which has become available for ¨ Tiefenbach, Gerthe horse (Andromed-ETM ; Minitub, many), contains no ingredients of animal origin, a
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Semen Extenders for Frozen Semen factor that is important with regard to sanitary requirements for semen export. The plasma membrane of sperm can undergo lipid phase separation during freezing, resulting in irreversible damage to the cell. To examine the membrane phase behavior of equine spermatozoa in the absence and presence of lipid-based cryoprotectants, the biophysical properties of sperm membranes were investigated with Fourier-transform infrared spectroscopy.27 This study showed that the freeze–thaw process induces large-scale rearrangement of the membrane lipids in stallion spermatozoa, and that phosphatidylcholines (PC) from egg yolk or soybean associate with the sperm membrane and protect stallion sperm with comparable efficiency; however, those phospholipids were not incorporated into the sperm membrane. When compared with a standard egg-yolk-based diluent (INRA82), the soybean and egg PC media preserved viability and motility equally well in post-thaw sperm. A preliminary fertility study determined that sperm cryopreserved in the soybean PC-based medium were capable of fertilization at the same rate as sperm frozen in the conventional INRA82 medium. The advantages of these formulations over egg yolk preparations include a reduced risk of microbial contamination. Furthermore, it is possible that in the future international quarantine agencies or government management procedures may ban the use of specific pathogen-free eggs in extender.
Cryoprotectants Cryoprotective agents that diminish cell damage are usually divided into two general classes related to their ability to move across the cell membrane: permeant (glycerol, sugar alcohols, DMSO [dimethyl sulfoxide], propylene glycol, dimethyl formamide, ethylene glycol, methyl formamide) and impermeant (sugars like lactose, mannose, raffinose, and trehalose, amino acids, different proteins). Their mechanisms of action are different. Cryoprotection by permeant agents can be explained by their colligative properties: they lower the temperature at which cells are exposed to a critical salt concentration. Impermeable agents, however, are thought to dehydrate cells, allowing them to be cooled rapidly before lethal injury due to solution effects takes place.28 The effects of cryoprotectants are more complex than can be explained from thermodynamics and solution theory. In high concentrations, they can produce lethal osmotic effects during their addition and removal (hyperosmolality) and they can be toxic.24, 29 Initial cellular shrinking (due to dehydration) caused by molar concentrations of cryoprotectant, and subsequent swelling during removal of cryoprotectant, can
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both result in damage if the maximal osmotic excursion is exceeded. Whether the effect of a given cryoprotective agent is negative or positive can depend greatly on the kinetics of exposure. Glycerol The first cryoprotectant identified was glycerol, and it remains one of the most favored cryoprotectants today. The recommended concentration of glycerol in freezing extenders varies between laboratories (Table 315.1); thus, the optimum concentration for maximal post-thaw sperm quality of equine spermatozoa is not yet resolved. This is further hindered by the fact that much of the information offered on the concentration of glycerol in published studies represents that in the freezing extender and not as a final concentration in the processed semen. Glycerol toxicity is partly based on osmotic stress, since glycerol permeates the cell membrane more slowly than other cryoprotectants.30 Pace and Sullivan301 and Demick et al.32 reported that glycerol had a detrimental effect on the fertility of fresh cooled equine semen. When jackass semen was frozen in INRA82 (2,5% glycerol), pregnancy rates after insemination of jennies were poor. Similar results were obtained when mares were inseminated with stallion semen frozen in INRA82 with various final glycerol concentrations (0%, 2.2%, 3.5%, 4.8%); the decrease in fertility (13/15, 86%; 8/15, 53%; 8/15, 53%; 2/15, 13%) was similar in mares but at generally higher concentrations of glycerol.33 Therefore, a compromise should be found between the cryoprotective effect and the negative effect on fertility of glycerol. Given the information available to date, the recommended final concentration of glycerol should not exceed the range of 2.5–3.5%. Amides Glycerol has been used almost universally as the cryoprotectant of choice, although it has been reported recently that other cryoprotectants, such as dimethyl sulfoxide, ethylene glycol, methyl formamide, or dimethyl formamide, may yield similar or superior results.34, 35 It was suggested that the ideal cryoprotectant must have low molecular weight, great water solubility, and minimal toxicity. Because most of the amides had a lower molecular weight compared to glycerol, these cryoprotectants may induce less osmotic damage. Based on several experiments conducted by Brazilian researchers (for review see ref. 35) using amides as penetrating cryoprotectants in freezing extenders for equine semen, the new freezing extender BotuCrioTM (www.criovital.com) was developed; this extender contains a mixture of cryoprotectants. In a recent study, semen was collected
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from stallions with a history of poor sperm freezability. The semen was centrifuged in a milk-based semen extender (Botu-SemenTM ; www.criovital.com), and the sperm pellet was resuspended with either INRA82, lactose-EDTA or BotuCrioTM and further processed for freezing using the protocol recommended for each specific extender. Post-thaw motility values were higher for Botu-CrioTM extender than for INRA82 or lactose-EDTA extenders for these stallions.36 In another study, pregnancy rates after AI of stallion semen frozen with either 2% glycerol, 3% glycerol, or 2% dimethyl formamide were not statistically different (46%, 58%, 50%).37 Others observed a significant improvement in fertility of stallion semen frozen with dimethyl formamide when compared with glycerol.38, 39 However, more extensive fertility trials are needed before considering widespread replacement of glycerol with amides for cryopreservation of stallion spermatozoa. In addition to energy, protein, and protective effects during the cooling phase, other essential components have been added to cryodiluents for different purposes. Trimeche et al.40 reported that the optimal concentrations for the amino acids glutamine and proline in freezing extender are 30–50 mM, and that higher concentrations (120 and 160 mM) decreased post-thaw motility. Ethylene diamine tetra-acetic acid (EDTA) is often added to stallion cryodiluents as a chelating agent to bind calcium ions, preventing them from entering and damaging spermatozoa during cold shock. Antibiotics are used to control the growth of microbes in the solution. Their impact in extenders for stallion semen is described in chapter 131. Monosaccharides (glucose, fructose), disaccharides (sucrose, lactose), and trisaccharides (raffinose) are also commonly incorporated into semen extenders. Disaccharides and trisaccharides are nonpermeable sugars and are used primarily because of their osmotic effects. Monosaccharides also have osmotic properties. In addition, they can be utilized by spermatozoa as a source of energy. The membrane cholesterol:phospholipid ratios differ markedly between species, and membranes containing a higher cholesterol:phospholipid ratio are less vulnerable to the effects of cooling and freezing. One way to prevent this damage is to increase the membrane fluidity at low temperatures by adding cholesterol to the membrane. Cholesterol can easily be incorporated into, or extracted from, the plasma membranes of cells using cyclodextrins. Cyclodextrins, cyclic heptasaccharides, can transport cholesterol into or out of membranes down a concentration gradient. Increasing membrane stability by incorporating cholesterol-loaded cyclodextrins in freezing diluents has been reported to be successful with stallion sperm.41, 42 The pregnancy results with post-
thaw stallion sperm subjected to this procedure have not been studied adequately. To summarize, the protocol for using cryoprotectants is a compromise between their beneficial and potentially detrimental effects. In all likelihood, the composition of the freezing extender may need to be adapted to each individual stallion for maximal effect.
References 1. Tischner M. Evaluation of deep frozen semen in stallions. J Reprod Fert (Suppl.) 1979; 27: 53–9. 2. Katila T, Andersson M, Reilas T, Koskinen E. Post-thaw motility and viability of fractionated and frozen stallion ejaculates. Theriogenology 2002; 58: 241–4. 3. Moore AI, Squires EL, Graham JK. Effect of seminal plasma on the cryopreservation of equine spermatozoa. Theriogenology 2005; 63: 2372–81. 4. Weiss S, Janett F, Burger D, Thun R. The influence of centrifugation on quality and freezability of stallion semen. Schweiz Arch Tierheilkd 2004; 146: 285–93. 5. Cochran JD, Amann RP, Squires EL, Pickett BW. Effects of centrifugation, glycerol level, cooling to 5◦ C, freezing rate and thawing rate on the post-thaw motility of equine sperm. Theriogenology 1984; 22: 25–38. 6. Volkmann DH, van Zyl D. Fertility of stallion semen frozen in 0.5-ml straws. J Reprod Fertil 1987; 35 (Suppl.): 143–8. 7. Knop K, Hoffmann N, Rath D, Sieme H. Effects of cushioned centrifugation technique on sperm recovery and sperm quality in stallions with good and poor semen freezability. Anim Reprod Sci 2005; 89: 294–7. 8. Waite JA, Love CC, Brinsko SP, Teague SR, Salazar JL Jr, Mancill SS, Varner DD. Factors impacting equine sperm recovery rate and quality following cushioned centrifugation. Theriogenology 2008; 70: 704–14. 9. Ecot P, Decuadro-Hansen G, Delhomme G, Vidament M. Evaluation of cushioned centrifugation technique for processing equine semen for freezing. Anim Reprod Sci 2005; 89: 245–8. 10. Barker CAV, Gandier JCC. Pregnancy in a mare resulting from epididymal spermatozoa. Canad J Comp Med Vet Sci 1957; 21: 47–51. 11. Samper JC, Morris CA. Current methods for stallion semen cryopreservation: a survey. Theriogenology 1998; 49: 895–903. ¨ 12. Martin JC, Klug E, Gunzel-Apel A-R. Centrifugation of stallion semen and its storage in large volume straws. J Reprod Fertil (Suppl.) 1979; 27: 47–51. 13. Palmer E. Factors affecting stallion semen survival and fertility. In: Proceedings of the 10th International Congress on Animal Reproduction and Artificial Insemination, UrbanaChampaign, IL, 1984. pp. 377–8. 14. Heitland AV, Jasko DJ, Squires EL, Graham JK, Pickett BW, Hamilton C. Factors affecting motion characteristics of frozen-thawed stallion spermatozoa. Equine Vet J 1996; 28: 47–53. 15. Braun J, Hochi S, Oguri N, Sato K, Torres-Boggino F. Effect of different protein supplements on motility and plasma membrane integrity of frozen-thawed stallion spermatozoa. Cryobiology 1995; 32; 487–92.
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Semen Extenders for Frozen Semen 16. Schembri MA, Major DA, Suttie JJ, Maxwell WM, Evans G. Modification of standard freezing media to limit capacitation and maximise motility of frozen-thawed equine spermatozoa. Aust Vet J 2003; 81: 748–51. 17. Ecot P, Arnaud G, Moy A, Daels P, Magistrini M, Vidament M. Comparison of fertility and post-thaw semen criteria of stallion semen frozen in two different extenders. Anim Reprod Sci 2001; 68: 356–8. 18. Vidament M. French field results (1985–2005) on factors affecting fertility of frozen stallion semen. Anim Reprod Sci 2005; 89: 115–36. 19. Roasa LM, Choi YH, Love CC, Romo S, Varner DD, Hinrichs K. Ejaculate and type of freezing extender affect rates of fertilization of horse oocytes in vitro. Theriogenology 2007; 68: 560–6. ¨ ¨ 20. Aurich C, Seeberi P, Muller-Schl osser F. Comparison of different extenders with defined protein composition for storage of stallion spermatozoa at 5◦ C. Reprod Dom Anim 2007; 42: 445–8. 21. Padilla AW, Foote RH. Extender and centrifugation effects on the motility patterns of slow-cooled stallion spermatozoa. J Anim Sci 1991; 69: 3308–13. 22. Batellier F, Duchamp G, Vidament M, Arnaud G, Palmer E, Magistrini M. Delayed insemination is successful with a new extender for storing fresh semen at 15 ◦ C under aerobic conditions. Theriogenology 1998; 50: 229–36. ¨ ¨ 23. Pagl R, Aurich J, Muller-Schl osser F, Kankofer M, Aurich C. Comparison of an extender containing defined milk protein fractions with a skim milk-based extender for storage of equine semen at 5◦ C. Theriogenology 2006; 66: 1115–22. 24. Amann RP, Pickett BW. Principles of cryopreservation and a review of cryopreservation of stallion spermatozoa. Equine Vet Sci 1987; 7: 145–73. 25. Clulow JR, Maxwell WMC, Evans G, Morris LHA. A comparison of duck and chicken egg yolk for cryopreservation of stallion sperm. Aust Vet J 2007; 85: 232–5. ¨ ¨ 26. Aires VA, Hinsch KD, Muller-Schl osser F, Bogner K, ¨ ¨ Muller-Schl osser S, Hinsch E. In vitro and in vivo comparison of egg yolk-based and soybean lecithin-based extenders for cryopreservation of bovine semen. Theriogenology 2003; 60: 269–79. 27. Ricker JV, Linfor JJ, Delfino WJ, Kysar P, Scholtz EL, Tablin F, Crowe JH, Ball BA, Meyers SA. Equine sperm membrane phase behavior: the effects of lipid-based cryoprotectants. Biol Reprod 2006; 74: 359–65. 28. McGann LE. Differing actions of penetrating and nonpenetrating cryoprotective agents. Cryobiology 1978; 15: 382–90. 29. Woods EJ, Benson JD, Agca Y, Critser JK. Fundamental cryobiology of reproductive cells and tissues. Cryobiology 2004; 48: 146–56. 30. Gilmore JA, McGann LE, Liu J, Gao DY, Peter AT, Kleinhans FW, Critser JK. Effects of cryoprotectant solutes on water permeability of human spermatozoa. Biol Reprod 1995; 53: 985–95. 31. Pace M, Sullivan JJ. Effect of timing insemination, numbers of spermatozoa and extender components on the pregnancy rate in mares inseminated with frozen semen. J Reprod Fertil 1975; 23 (Suppl.): 115–21. 32. Demick DS, Voss JL, Pickett BW. Effect of cooling, storage with glycerolization and spermatozoa number on equine fertility. J Anim Sci 1976; 43: 633–7.
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33. Vidament M, Vincent P, Martin FX, Magistrini M, Blesbois E. Differences in ability of jennies and mares to conceive with cooled and frozen semen containing glycerol or not. Anim Reprod Sci 2008; doi: 10.1016/j.anireprosci.2008.03.016. 34. Squires EL, Keith SL, Graham JK. Evaluation of alternative cryoprotectants for preserving stallion spermatozoa. Theriogenology 2004; 62: 1056–65. 35. Alvarenga MA, Papa FO, Landim-Alvarenga FC, Medeiros ASL. Amides as cryoprotectant for freezing stallion semen: A review. Anim Reprod Sci 2005; 89: 105–13. 36. Pasquini DF, Ferreira HN, Papa FO, Dell Aqua JA Jr, Alvarenga MA. Effects of three cryopreservation systems on longevity of stallion sperm after thawing. Reprod Fert Dev 2008; 20: 123–4 (abstract). 37. Vidament M, Daire C, Yvon JM, Doligez P, Bruneau B, Magistrini M, Ecot P. Motility and fertility of stallion semen frozen with glycerol and/or dimethyl formamide. Theriogenology, 2002; 58: 249–51. 38. Medeiros ASL, Gomes GM, Carmo MT, Papa FO, Alvarenga MA. Cryopreservation of stallion sperm using different amides. Theriogenology 2002; 58: 273–6. 39. Moffet PD, Bruemmer JE, Card C, Squires EL. Comparison of dimethyl formamide and glycerol for cryopreservation of equine spermatozoa. In: Proceedings of the Society for Theriogenology Annual Conference, 2003, p. 42 (abstract). 40. Trimeche A, Yvon JM, Vidament M, Palmer E, Magistrini M. Effects of glutamine, proline, histidine and betaine on post-thaw motility of stallion spermatozoa. Theriogenology 1999; 52: 181–91. 41. Combes GB, Varner DD, Schroeder F, Burghardt RC, Blanchard TL. Effect of cholesterol on the motility and plasma membrane integrity of frozen equine spermatozoa after thawing. J Reprod Fertil Suppl. 2001; 56: 127–32. 42. Moore AI, Squires EL, Graham JK. Adding cholesterol to the stallion sperm plasma membrane improves cryosurvival. Cryobiology 2005; 51: 241–9. 43. Loomis PR, Amann RP, Squires EL, Pickett BW. Fertility of unfrozen and frozen stallion spermatozoa extended in EDTA-lactose-egg yolk and packaged in straws. J Anim Sci 1983; 56: 687–93. 44. Vidament M, Ecot P, Noue P, Bourgeois C, Magistrini M, Palmer E. Centrifugation and addition of glycerol at 22◦ C instead of 4◦ C improve post-thaw motility and fertility of stallion spermatozoa. Theriogenology 2000; 54: 907–19. 45. He, 1986. cit. in Amann RP, Pickett BW. Principles of cryopreservation and a review of cryopreservation of stallion spermatozoa. Equine Vet Sci 1987; 7: 145–73. 46. Piao S, Wang Y. A study on the technique of freezing concentrated semen of horses (donkeys) and the effect of insemination. In: Proceedings of the International Congress on Animal Reproduction and Artificial Insemination, 1988 ; vol. 3, 286a–c. 47. Burns PJ, Reasner DS. Computerized analysis of sperm motion: Effects of glycerol concentration on the cryopreservation of equine spermatozoa. J Equine Vet Sci 1995; 15: 377– 80. 48. Wilhelm KM, Graham JK, Squires EL. Comparison of the fertility of cryopreserved stallion spermatozoa with sperm motion analyses, flow cytometric evaluation, and zona-free hamster oocyte penetration. Theriogenology 1996; 46: 559– 78.
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Chapter 316
Freezing Semen Harald Sieme
Freezing of stallion sperm is, in some countries, performed mainly for export purposes. However, it is also used for creating a sperm reserve from stallions – for example, with stallions whose predominant use is competition; for sperm storage preceding castration of a stallion; in the case of cancellation of a stallion’s book due to illness of the stallion; and for preparing a gene reserve in case of untimely death or for an endangered breed.
Preparing the stallion for freezing of its sperm Freezing of stallion sperm requires long-term and careful planning. It is important to evaluate the semen quality of a stallion prior to embarking on a semen cryopreservation program, because not every stallion can fulfill the prerequisites with regard to sperm quality needed in a good candidate for freezing of sperm. Sporadic freezing of semen often yields fruitless results, because semen subjected to this process at random may not produce frozen-thawed sperm of adequate quality. Since frozen sperm is often processed particularly for export to other countries, it is imperative to know the legal prerequisites of the countries of destination, and to work in cooperation with the official gazetted veterinarian to prepare and examine the stallions appropriately for this process. Generally, many regulations are in place for this purpose, and the regulations vary with importing country. For most countries, donor stallions have to be placed in quarantine under official veterinary supervision during the entire period of the frozen semen processing period. It is extremely important to consider the conditions concerning animal breeding rights and, above all, animal disease rights that are to be observed when exporting stallion sperm to foreign countries. Most importing countries demand permanent residency of the donor stallion at an artificial in-
semination (AI) facility at least 30 days before starting to collect semen or disease-related hygienic reasons for the time period of the deep freezing of sperm. The major equine semen importing or exporting countries provide the necessary documentation for this process through government agencies – for example, the Animal and Plant Health Inspection Service (APHIS) for the USA (www.aphis.usda.gov/import export); the Canadian Food Inspection Agency (CFIA) for Canada (http://airs-sari.inspection.gc.ca); and the Australian Quarantine and Inspection Service (AQIS) for Australia (www.daffa.gov.au/aqis/import/liveanimals/reproductive). Animal health requirements governing the trade of imports into the European Community of equine semen, ova, and embryos are contained in the Council Directive 92/65/EEC and Commission Decision 95/176/EC, and 2010/176/EC (http://eur-lex.europa.eu). Freezability of stallion sperm Not all stallions are suitable candidates for semen freezing. The reasons are complex; however, inadequate selection of breeding stallions based on fertility-relevant parameters may play a role here, in comparison to other farm animals. It is generally believed that the key to successful cryopreservation of stallion semen is the individual stallion itself. Stallions have shown a particularly high degree of individual variation with respect to the cryosurvival of their sperm. It has been estimated that ∼ 20% of stallions produce semen that freezes well, 60% produce semen that freezes acceptably (fair), and 20% produce semen that freezes poorly. Tischner1 evaluated the post-thaw quality of 200 ejaculates from 36 stallions and concluded that stallions could be grouped into these categories based on post-thaw motility:
r r r
good: > 40% progressively motile sperm (pms); acceptable or fair: 20–40% pms; poor: < 20% pms.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Freezing Semen Vidament et al.2 considered stallions that produced sperm with post-thaw motility greater than 35% with ‘rapid’ velocity (average path velocity [VAP] > 30 µm/s measured using computer-assisted sperm motion analysis [CASA]) for ≥ 33% of ejaculates to be acceptable for use in a frozen semen program; in that report, 51% of tested stallions met these criteria. Using the new French National Stud protocol, Vidament3 reported that 90% of 344 stallions presented from 2003 to 2005 were acceptable for commercial AI; the level of acceptance was set at > 33% of ejaculates attaining or exceeding the selection criteria after thawing. Loomis and Graham4 reported that an acceptable stallion was one that yielded post-thaw progressive motility ≥ 30% following 30 min incubation at 37◦ C for ≥ 25% of ejaculates frozen; during the testing period from 2000 to 2006, 281 out of 332 (84.6%) stallions met these criteria. Stallions that are fertile under normal field conditions can produce semen that, after freezing and thawing, produces very low pregnancy rates. The mechanisms underlying the differences in cryosensitivity between different individuals have yet to be elucidated fully. Such differences could be genetic in origin, and genetic selection of stallions for successful freezing could be a possibility. However, the cause might be non-genetic and in this regard it would be particularly desirable to be able to apply assays of sperm function before and after freezing that correlate well with either semen freezability or stallion fertility. Unfortunately, such assays are not currently available.5, 6 Sperm is often rejected from cryopreservation when pre-freeze motility is < 50% (13/21 centers required > 50%, 8/21 required > 60% motile sperm7 ). French researchers suggested that the hypo-osmotic swelling (HOS) test applied after collection of fresh semen was the best predictive test of the freezability of stallion semen.8 In the author’s experience, Warmblood stallions are considered suitable for cryopreservation when they produce ejaculates with a sperm concentration of ≥ 200 million sperm per mL, a progressive motility of > 50% and a proportion of morphologically normal sperm exceeding 70%.
Effect of season and semen collection frequency on freezability of stallion sperm It is advantageous to dedicate a specific time frame during which stallions can be used exclusively for production of semen for freezing. This possibility works to our advantage, because for organizational reasons semen from stallions is typically subjected to cryopreservation outside the actual breeding season, whereas collecting semen for production of fresh
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and cooled-transported semen takes place during the commercial breeding season. With regard to postthaw sperm quality, cryopreserved stallion semen may also be best prepared during the non-breeding season. Stud farms should take into consideration many practical matters (e.g. sexual rest, individual characteristics of stallions, and hygienic conditions) that precede the freezing campaign. The optimal time period for manufacturing deep-frozen sperm from stallions is subject to debate; some authors have reported no noteworthy seasonal effects on the quality of frozen-thawed sperm, while others have obtained better thawing results in the winter months and, with more recent investigations, in the autumn (September to November in the northern hemisphere).9 Although seasonal influences on sperm quality in stallions cannot be completely ruled out, long-standing practice shows that good and constant results can be reached in the autumn and winter months. Following a long period of sexual rest, extragonadal sperm reserves should be depleted by repeated collections until good sperm quality is established. For preparing frozen semen, it is preferable to make the semen collection interval a minimum of 48 hours, although this should be adapted to the individual stallion10 in order to determine optimized semen collection intervals for individual stallions. Semen collection should be carried out using a suitable technique (i.e., considerations for type of artificial vagina, lubricant, and collector), and a fixed time interval is generally applied between collections to minimize differences among ejaculates. Ideally, all conditions, including the stallion handler, semen collector, and semen collection facility, should be kept the same throughout the semen collection period.
Sperm processing for freezing The current techniques for deep-freezing equine semen are certainly not standardized from facility to facility, or from country to country; however, inter- and intra-stallion differences should be considered when selecting a semen-freezing method. The initial steps of semen processing for freezing are almost identical to thoses used for processing fresh or cooledtransported semen. A standard approach may be as follows:
r r r r
dilute the gel-free ejaculate with an extender (in this case, a centrifugation extender); centrifuge the extended semen at 400–600g for 8–15 minutes; remove the supernatant and resuspend the sperm pellet in freezing extender; package the semen (typically in 0.5-mL straws);
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cool the packaged semen from room temperaure to + 5◦ C; freeze the semen in liquid nitrogen vapor or an automated freezing chamber; store the frozen straws in liquid nitrogen.
Current techniques of cryopreservation of stallion semen When comparing protocols from different countries, the methods for freezing stallion semen and instructions for insemination of mares with frozen-thawed semen are far from standardized (for review see Samper and Morris7 ). The lack of standardization is demonstrated by the differences in processing, packaging, and thawing of semen for cryopreservation (Table 316.1). Current freezing protocols for stallion semen typically involve a two-step dilution procedure in which semen is first diluted with a primary (centrifugation) extender, followed by centrifugation, then redilution in a second (freezing) extender containing a cryoprotectant. The first dilution employs either saline/sugar extenders or milk-based extenders used to dilute fresh semen. The dilution ratio can be 1:1 or the semen can diluted to a specific sperm concentration (e.g., 50 million spermatozoa/mL). The success of centrifugation, in terms of maximizing sperm recovery rate without damaging sperm, depends on the centrifugation duration (8–15 min) and centrifugation force (400–600g). To increase sperm recovery, the use of high-speed centrifugation (20 min at 1000g) over a liquid cushion has been introduced into laboratory practice;11 this technique had no detrimental effect on fertility.12 Moore et al.13 demonstrated the deleterious effect of seminal plasma on stallion spermatozoa during cryopreservation. However, retention of 5–20% seminal plasma in the suspension after centrifugation has been considered by some to be essential for cryosurvival. Within Europe, many AI centers follow the French protocol based on 20 years’ experience of National Stud Farm practice in France.3 The present system applied in the French National Stud includes: 1. a freezing protocol – dilution in milk, centrifugation, and addition of freezing extender (INRA82 + egg yolk [2% v/v] + glycerol [2.5% v/v]) at 22◦ C, a moderate cooling rate to 4◦ C and freezing at – 60◦ C/min in 0.5-mL straws; 2. selection of ejaculates showing post-thaw rapid motility > 35%; 3. an insemination protocol whereby mares are examined once daily and inseminated twice with 400 × 106 total frozen-thawed spermatozoa, 24 h apart, before ovulation is detected.
A sufficient number of straws from a stallion is required to perform six AIs of 400 × 106 total spermatozoa, i.e., 2.4 × 109 total spermatozoa are made available per mare per season. Freezing stallion semen in INRA96-based extender improved fertility rates in comparision with R INRA82 (Pillet et al.65 ). Furthermore, a new INRA96 extender containing sterilised egg yolk plasma has R been developed, this extender (INRA-Freeze , IMVTechnologies, L’Aigle, France) is ready to use and marketed that it will complie with all biosafety requirements. The semen-freezing schedule recommended in Germany14 is conducted from October to the end of January, during the non-breeding season. Semen from each stallion is collected three times per week (Monday, Wednesday, Friday) between 8 am and 11 am, using an artificial vagina, a breeding phantom, and a teaser mare at the front of the breeding phantom. Sterile gauze filtration sets are used to separate the gel portion of ejaculated semen, and the gel-free semen is evaluated for volume, concentration of spermatozoa, and percentage of progressively motile spermatozoa. Semen is extended in a modified skim-milk extender to a final concentration of 50 × 106 spermatozoa/mL and centrifuged for 10 min at 600g at ambient temperature. After decantation of supernatant, the sperm pellets are suspended in skim-milk extender containing 2% egg yolk. Sperm concentration is calculated by hemocytometer and final concentration of spermatozoa is adjusted to 200 × 106 spermatozoa/mL and a glycerol concentration of 2.5%. After drop-wise addition of freezing extender, the semen is stored in 50-mL plastic tubes and placed into a 300-mL beaker containing water at room temperature. The beaker is placed in a refrigerator (preferably a cooling cabinet) at 5◦ C for 2.5 hours, which allows for slow cooling of the extended semen. Semen is automatically loaded into 0.5-mL straws at + 5◦ C. Straws are automatically frozen (+ 5◦ C to – 140◦ C at – 60◦ C/min), using a commercial programmable freezer. In our program, a commercial dose for AI contains 800 × 106 total spermatozoa, with a post-thaw progressive motility of ≥ 35%. A new freezing protocol has been developed by Brazilian researchers using Botu-CrioTM extender (Biotech Botucatu/ME LTDA, Brazil) containing a mixture of two cryoprotectants (1% v/v glycerol, 4% v/v methylformamide). The semen is suspended in a ratio of 1:1 with milk extender (20–30◦ C, centrifuged for 10 min at 600g (20–25◦ C), resuspended with BotuR extender (20–25◦ C), loaded into 0.5-mL straws Crio (100 × 106 sperm/straw), and stored at + 5◦ C for 20 min. Freezing of straws can be carried out either in static nitrogen vapor (3–6 cm above nitrogen) for
600g 10 min
600g 10 min
Remove 90% seminal plasma
Citrate-EDTA (20◦ C), cushion
INRA82-EY → + 4◦ C/1 h Citrate-EDTA (30◦ C), cushion
UHT skim milk (37◦ C) INRA82 (37◦ C)
Milk (20–30◦ C)
Skim milk-glucose
1:1
50 × 106 sp/mL
1:4
50–80 × 106 sp/mL
2.5 × 109 sp/ 50-mL tube
50 × 106 sp/mL
1:1
Loomis et al.23
Cochran et al.28
Palmer47
Ha˚ ard ˚ and Ha˚ ard ˚ 24
Vidament3
Sieme14
www.criovital.com
Four different
R Botu-Crio
INRA82-EY-2%
INRA82-EY
Lactose-EDTA
INRA82-EY
Lactose-EDTA-EY
EDTA-lactose-EY
Merck-Lactose-EY
1% glycerol, 4% methylformamide
2.5%
2.5%
5%
2.5%
5%
5%
5% 0.5-mL straw
R Macrotub ¨ (4.0-mL straw)
Glass vials (1 mL)
Aluminum-tubes (20–25 mL)
Packaging
37◦ C for 30 s 75◦ C for 7 s then 35◦ C for 10–30 s
37◦ C for 30 s
+ 4 → – 140◦ C: 60◦ C/min + 20 → – 10◦ C: 10◦ C/min – 10 → – 140◦ C: 25◦ C/min → + 4◦ C: 1 h 15 min + 4 → – 140◦ C: 40–60◦ C/min
Controlled rate freezer Varying cooling rates
+ 5◦ C for 20 min vapor (15–20 min) or automatically 20–40◦ C/min
30–40 × 106 sp., skim-milk-glucose ext, 30 min at 37◦ C
46◦ C for 20 s or 37◦ C for 1 min
37◦ C for 30 s
37◦ C for 30 s or 75◦ C for 7 s
+20 → – 15◦ C: 10◦ C/min – 15 → – 120◦ C: 25◦ C/min
+ 20 → + 5◦ C: beaker for 2.5 h + 5 → – 140◦ C: 60◦ C/min
38◦ C 30 s
50◦ C 40 s
Sucrose-milk
40◦ C 50 s
Thawing
10 min N2 vapor
20 min N2 vapor
N2 vapor
7–9 min N2 vapor
Freezing
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400g 15 min
600g 10 min
400g 15 min
650g 15 min
1000g 10 min
4–5%
3.5%
Glycerol conc.
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EDTA, ethylenediaminetetraacetic acid; ext, extender; EY, egg yolk; sp, spermatozoa.
Glucose-EDTA
Sucrose-milk-EY
Lactose-EDTA-EY
Freezing extender
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Loomis and Graham4
600g 10 min
Glucose-EDTA
1:1
Martin et al.46
350–450g 12 min
1:1
He45 11% sucrose
Collection of sperm rich fraction
Centrifugation
Tischner1
Centrifugation extender
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Table 316.1 Processing methods used for cryopreservation of stallion semen
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15–20 min, or frozen automatically (– 20–40◦ C/min until – 120◦ C), followed by immersion of straws in liquid nitrogen for storage (www.criovital.com). Recently, two other workers have published their procedure for freezing of stallion semen.4 These workers prefer a split-ejaculate test-freeze procedure. Stallions whose initial semen quality is < 35% progressive motility and/or < 40% morphologically normal sperm or who persistently shed known pathogens that are non-responsive to treatment with extender containing antibiotics are excluded. Their standard test-freeze procedure consists of diluting the semen in a skim-milk-glucose-based extender, followed by centrifugation and removal of > 90% seminal plasma and resuspension of the sperm into four different freezing extenders. Semen is then packaged in 0.5-mL straws and cooled and frozen in a controlled-rate cell freezer. The straws are plunged into liquid nitrogen and stored for > 24 h prior to thawing. The cooling and freezing protocols vary with the extender used. The four extenders are proprietary variations of published formulations whose compositions vary with respect to: type and amounts of sugars and salts; presence or absence of milk as a primary component and amount used; percentage of egg yolk; types of buffers; and percentage of cryoprotectant.
Sperm packaging of frozen stallion sperm Several packaging methods are available for the storage of cryopreserved stallion spermatozoa. Early cryopreservation work involved the use of glass ampoules and resulted in the first pregnancy after insemination with frozen stallion spermatozoa.15 Later, semen was frozen in pellets, and foals were also produced after insemination of spermatozoa frozen by this method.16 In a study by Pace and Sullivan,17 higher pregnancy rates were achieved by spermatozoa cryopreserved in 10-mL ampoules (44%) as compared to pellets (36%). Currently, the most popular packaging for the cryopreservation of stallion spermatozoa is the polypropylene or polyvinyl chloride tubular straws, due to their ease of identification and storage, in comparison to pellets or ampoules. Straws have ranged in size from 0.25 to 5 mL. The choice of packaging device should be based on product quality as well as practicality. From a cryobiological point of view, packages that have a large surface area (as in the case of small-volume straws) enable more homogeneous freezing and thawing rates. This is demonstrated by the fact that 0.5-mL straws yield better thawing results than 4.0-mL macrotubes and flattened macrotubes.18 As such, most semen worldwide is frozen in 0.5-mL straws. For filling purposes, an automated
or semi-automated filling device is generally recommended. Such devices load the semen into 0.5-mL straws by means of negative pressure and seal the loaded straws automatically. In the author’s view, an ultrasound method of straw sealing is optimal. Sealing straws by means of a sealing ball, heat, or sealing powder can be troublesome and may lead to worse results. Regardless of which form of package is used for semen, every unit must have clear and essential information as to the identity of the stallion (name and identification number), the manufacturing AI center (name or registration number) as well as the date of semen collection.
Sperm concentration When diluting semen to a desired concentration of spermatozoa, adequate extender must be added to ensure maximum survival rates post-thaw. It has been suggested that the survival of stallion spermatozoa is enhanced when frozen at a low concentration due to the greater availability of cryoprotective agents surrounding the spermatozoa. Despite this hypothesis, successful cryoprotection has been achieved at concentrations ranging from 20 to 1600 × 106 spermatozoa/mL, i.e., 20 × 106 /mL,19 50 × 106 /mL,20 100 × 106 /mL,21 250 × 106 /mL,22 500 × 106 sperm/mL,23 1000 × 106 motile sperm/mL,24 and 1600 × 106 /mL.25 Although a very high concentration may reduce an insemination dose to one to two straws, there are reports showing that, in the case of a higher dilution of sperm (i.e. to 100 × 106 spermatozoa/mL), the post-thaw sperm motility tends to be better. The influence of sperm concentration on the fertility results to date has not been clearly explained for frozen sperm of stallions (Table 316.2). Nonetheless, it is reported that the pregnancy results tend to be better when sperm is used that is frozen in highly concentrated form (1600 × 106 spermatozoa/mL as opposed to 400 × 106 spermatozoa/mL).25
Cooling and freezing rates It is generally considered important that the semen cool slowly from room temperature to 5◦ C (i.e., at 0.1◦ C/min) under anaerobic conditions. In most European laboratories, the cooled semen is filled into 0.5-mL straws at 5◦ C,14, 26 within a cooling cabinet. Alternatively, sperm packaging could be carried out at room temperature (approx. + 20◦ C) and a programmable freezer used for the mandatory slow cooling of sperm that precedes the freezing process; this step contributes decisively to the freezing success. Futher investigations are needed to determine if additional adaptation time of the filled straws at + 5◦ C is required for semen of selected stallions. Only a
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Table 316.2 Effect of packaging, sperm dose, and AI regime on pregnancy rate of mares inseminated with frozen/thawed semen
Source
Packaging
Sperm dose (× 106 )
Pace and Sullivan17
Ampoules, pellets
40 motile 80 motile 160 motile
Klug et al.48
Volume (mL); duration of AI interval
Pregnancy (+ve/n)
rate (%)
>1
2/20 1/19 7/20 5/20 5/18
10 5 35 25 28
>1
58/228
25
15 mL 24-h interval
>1
91/246
37
12-h interval 24-h interval 12-h interval 24-h interval 24-h interval
Number AI
Pellets
100
49
Alu-tubes
320–1104 motile
Muller ¨ 50
Alu-tubes
300 motile
15
>1
537/959
56
Kloppe et al.51
Makrotube
600
48-h interval 0–6 h post ov.
2.7 1
12/20 11/20
60 55
Vidament et al.52
0.5-mL straw
300 150 300 150
< 24 h pre ov. < 24 h pre ov. 24-h interval 24-h interval
1 1 ≥2 ≥2
508 71 187 207
26 21 41 34
Leipold et al.25
0.5-mL straw
800 motile 800 motile 320 motile 320 motile
1600/mL 400/mL 1600/mL 400/mL
> 1 24-h interval
8/24 7/25 10/25 4/26
33 28 40 15
0.5-mL straw
400
24-h interval
>1
4190
49
–
–
– 6–0 h pre ov. 0–+ 6 h post ov. Barren Maiden In foal
1
-/363 -/677 180♀ 155♀ 224♀
41 41 28 39 60
Katila54
–
–
1
237/716
33
Sieme et al.55
0.5-mL straw
800
24–12 h pre ov. – 12–0 h pre ov. 0–+ 12 h post ov. 24-h interval 24-h interval
1 1 1 2 3
8/26 31/75 24/48 31/62 3/9
30 41 50 50 33
Vidament3
0.5-mL straw
400
24-h interval Until ovulation Years 1999–2004
2
/1032 /1326 /1390 /1503 /1723 /1990
47 49 49 48 45 46
Barbacini et al.56
0.5-mL straw
> 200 motile Barren 3–9 years 10–16 years > 16 years
Within 6 h pre- or post-ovulation 24 h and 40 h post-hCG
1 2 107/151 223/294 48/99
71 76 48
Muller ¨
Vidament et al.2 Barbacini et al.
53
hCG, human chorionic gonadotropin; post ov., post-ovulation; pre ov., pre-ovulation.
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few programmable devices allow for cooling rates of < 0.1◦ C/min. Stallion spermatozoa must first endure a cooling phase from body temperature to about + 5◦ C. Spermatozoa can endure fast cooling rates from 37◦ C to 20◦ C, but require a slow cooling rate (< 0.1/min) between 20◦ C and 5◦ C to maintain viability and minimize effects of cold shock. The effects of cold shock can be minimized through the addition of cryoprotectants such as egg yolk to the cooling medium. Therefore, the composition of the freezing extender may influence the length of the cooling phase required before freezing (see Chapter 315). The transfer of sperm into the programmable freezer or vapor box must occur without great temperature oscillations. The rate of freezing has a decisive effect on cryopreservation of stallion sperm. Vastly differing freezing regimes are found in literature (see Table 316.1). However, freezing rates for stallion spermatozoa are difficult to compare as they are often dependent on the method of semen processing and storage. For example, semen frozen in pellets reaches a critical temperature on dry ice within a few minutes in comparison to more advanced controlled freezing rates of programmable freezers (see Table 316.1). Model calculations show an optimal freezing rate of between – 20◦ C/min and – 100◦ C/min;20, 27 should this range be heeded, no quality deviations are to be expected after thawing. The most convenient method of freezing is the suspension of straws within liquid nitrogen vapor near the vapour–liquid interface. However, the freezing rate with this system can vary as the position of the straws above the liquid nitrogen is altered, resulting in a subsequent change in the freezing rate. In the case of freezing over nitrogen vapor in a polystyrene box, straws are typically placed on a metal rack horizontally over liquid nitrogen for a certain time with the box closed. Afterwards the frozen straws are plunged into liquid nitrogen for final rapid cooling and storage. The rate of cooling is variable due to changes in the distance between the straws and the liquid, as well as the storage time. The greater the distance between the chilled semen and the nitrogen level, the lower are the cooling rate and final temperature. In the case of a recommended distance of 2.5–5 cm, the semen is frozen at a speed of 45 to 60◦ C/min, and the storage time in the nitrogen steam should amount to 20 min. In the case of deep-freezing by means of programmable freezers, the straws are stored horizontally on metal racks, which are stacked on top of one another in the freezing chamber of a commercial programmable freezer. The liquid nitrogen is channeled to the freezing chamber from a liquid nitrogen storage container corresponding to the measured temperature inside the freezing chamber. Via a turbine ven-
tilator the nebulized nitrogen is evenly distributed in the chamber. The crystallization proceeds spontaneously. After reaching the prescribed temperature, the frozen semen is transferred to liquid nitrogen for storage until thawing. Cochran et al.28 and Cristanelli et al.20 compared the post-thaw motility of stallion spermatozoa frozen in liquid nitrogen vapor at – 60◦ C/min and at a controlled rate of 10◦ C/min from 20◦ C to – 15◦ C and – 25◦ C thereafter to – 196◦ C. They found no differences between post-thaw motility of sperm frozen at the different rates, thereby suggesting that equine sperm could tolerate a range of freezing rates. There is still a need for more comprehensive studies on the effects of freezing stallion spermatozoa at different rates in different packages, such as 0.25- and 0.5-mL straws.
Alternative freezing techniques According to the two-factor hypothesis developed by Mazur et al. in 1972,29 cells can be damaged either by the formation of intracellular ice crystals when toorapid cooling takes place or by chemical toxicity or osmotic stress as well as mechanical damage brought about by phase separation of solution and crystal growth. From this point of view, cooling and freezing rate are both important regulatory factors during cryopreservation of spermatozoa.30–32 In addition, the rate of cooling affects the morphology of the intercellular ice crystals and the microarchitectural orientation of cells within unfrozen channels and plates of ice.33 Thus, maximizing the survival rate of cells subjected to freezing and thawing requires careful control of the freezing process. The conventional freezing method involves lowering the temperature of the chamber in a controlled stepwise manner. This method is based on changing temperature in a time unit by moving heat in multiple directions (equiaxial), depending on the thermal conductivity and geometrical shape of the container and of the biological material. The thermal gradient within the sample is determined implicitly by the temperature of the chamber and the thermal conductivity of the materials within the sample, and is not explicitly controllable. In addition, the temperature gradient within the freezing chamber and unreliable temperature recording measurements34 make it more difficult to achieve the optimal cooling rate. Commercially available systems to freeze stallion spermatozoa enable freezing of a large number of straws, but synchrony of sample and chamber temperature is critical. This significantly affects the ability accurately to characterize the freezing protocol. Alternative techniques for rapid cooling of stallion spermatozoa under controlled thermal
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Freezing Semen conditions have been published recently, for example, directional freezing and unique freezing technology (UFT). In directional freezing (www.coredynamics.com), after an initial seeding stage, the semen sample is advanced at a constant velocity through a linear temperature gradient. In this way, the ice crystal propagation can be controlled so as to optimize crystal morphology, and achieve continual seeding and a homogeneous cooling rate throughout the entire freezing process.35, 36 The freezing apparatus (Multi Thermal GradientTM 516; www.coredynamics.com) consists of two copper blocks that are separated by an opening and bedded in the gap for accommodating the cryotubes. The technology is based on a series of copper blocks arranged in a line, with a straight track running through the blocks. Each side of a block can have different temperatures, thereby imposing a temperature gradient on the portion of the track contained in the block. The tubes are placed in the first copper block (+ 5◦ C) and are driven with the tip of the tube in the second block (for stallion sperm predominantly at – 50◦ C) for releasing the initial seeding. Afterwards, further driving and directional freezing occurs at a constant speed. Currently this form of freezing is used in combination with special glass tubes, so-called ‘Hollow Tube’TM , IMT, Israel. These tubes are cylindrical with a central hollow area and a holding capacity of ∼ 10 mL. Improved thawing results are reported when combining ‘large volume freezing’ with the patented freezing method;36 however, it should be taken into consideration that for largerscale field use, the storage capacities increase and the insemination staff have to familiarize themselves with this new handling method. Semen is traditionally frozen using liquid nitrogen, but some success has been reported using a unique freezing technique (UFT; www. supa chillusa.com) adapted from the high-speed freezing technique used for human feedstuffs. Unique freezing technology (UFT) is based on the use of an electronically controlled cooling chamber that houses an organic fluid bath. This proprietary fluid has a relatively high heat dissipation rate, which appears to vitrify (to change or make into glass or a glassy substance) water and thus preserve cells. Results of studies done with the UFT and stallion spermatozoa vary, and some studies suggest that liquid nitrogen and UFT produce very similar results.37 Although vitrification of human sperm has been described,38 the authors are not aware that a reliable vitrification procedure has been published as an alternative to conventional cryopreservation of stallion semen. Lyophilization of stallion semen is described elsewhere in this book (see Chapter 312).
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Filters and methods for sperm purification have also been used to improve a semen sample pre-freeze. The filtrate obtained has high fertilizing potential that possibly improves AI fertilization rates post-thaw (see Sieme et al.39 and Loomis40 for review). Sublethal environmental stress, namely high hydrostatic pressure (HHP; Cryo-Innovation Ltd, Budapest, Hungary) applied to fresh semen before the freezing process was reported to improve significantly the motility, viability, and fertility parameters of frozen bull and boar semen. However, the mechanism of how HHP treatment improves survival rates following sperm cryopreservation remains unclear and this technique has, to our knowledge, not been reported for stallion semen.
Thawing For the thawing of frozen semen of stallions in conventional (0.5-mL) straws, immersion for 30 seconds in a water bath set at 37◦ C is generally recommended for field use. As a rule, more than one straw is used for insemination, so the straws needed are immersed in quick succession in the water bath. Thirty seconds after immersing the last straw, all of the straws are taken out of the water bath. The straws are dried thoroughly with cellulose or a similar material to avoid renewed cooling. Some authors prefer thawing straws at 40◦ C for 15 seconds or 38◦ C for 30 seconds. Thawing at 75◦ C for 7 seconds followed by a second submersion for 10–30 seconds at 35◦ C has also been described. The thawing protocol for macrotubes (i.e., 4–5-mL straws) is 50◦ C for 40 seconds. In general, when thawing semen that was processed by a given AI center, the thawing instructions provided by that center should be heeded.
Post-thaw evaluation of stallion spermatozoa Samper and Morris7 reported, in their worldwide survey, that most AI centers used computer-assisted sperm motion analysis (CASA) as a basic method for analysis of post-thaw semen quality. The threshold for acceptable post-thaw motility was quite different among centers: 25% for two centers, 30% for 10 centers, 35% for eight centers, and 40% for one center. Even if motility estimation before freezing has been demonstrated to be correlated with the motility after freezing and thawing, it cannot be considered as a reliable predictive test to discard a priori the ejaculates with poor freezability. Vidament3 summarized 20 years of field results with frozen semen in France and reported that stallions and ejaculates were selected on the basis of
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Table 316.3 Effect of AI techniques (conventional AI into uterine body vs deep uterine vs hysteroscopic AI) on pregnancy rates of mares inseminated with frozen/thawed semen Source
AI technique
AI dose ( × 106 )
AI volume (mL)
Pregnancy rate
(%)
Feo ´ et al.57
UB DU HS UB HS UB HS
250–350
4
25 motil
0.5
5 motil
0.1
100–150
2 × 0.5
HS HS UB UB UB UB DU UB HS HS UB HS UB DU HS UB DU
10 motil 10 motil 400 motil 800 400 200 200 14 motil 14 motil 3 motil 3 motil 3 motil 100 100 100 800 > 300 motil
8/25 32/48 9/14 9/12 16/34 2/8 12/22 center A 22/40 center B 4/12 4/12 0/12 12/20 11/20 10/20 4/20 8/12 9/14 16/34 2/14 1/12 16/37 17/38 22/48 9/21 114/193
32 66 64 75 47 25 54 52 33 33 0 60 55 50 20 67 64 47 14 8 43 44 45 42 59
Morris et al.58
Alvarenga et al.59 Alvarenga and Leao ` 60
Squires et al.61
Morris et al.62
Sieme et al.63
Crowe et al.64
Percoll 2 (1 AI) 1 (2 AI) 0.5 (2 AI) 0.5 (2 AI) 0.5 0.5 0.1 0.1 0.1 contral. 0.5 0.5 0.5 12 2–4 2 × AI 36 h and 48 h R post-Ovuplant
DU, transrectally guided deep intrauterine insemination close to the utero-tubal junction ipsilateral to the preovulatory follicle; HS, hysteroscopically guided deep intrauterine insemination close to the utero-tubal junction ipsilateral to the preovulatory follicle; UB, routine insemination into the uterine body.
objectively measured post-thaw motility; an ejaculate was considered acceptable if it possessed ≥ 35% rapid sperm (CASA), after thawing and dilution. A rapid sperm was defined as a motile cell with an average path velocity greater than 30 or 40 µm/s (depending on the CASA instrument used). In a previous study, Loomis41 reported that an acceptable stallion was one that yielded post-thaw progressive motility ≥ 30% following 30 min incubation at 37◦ C for ≥ 25% of ejaculates frozen following the Select Breeders Services freezing protocol. In a recent study, Loomis and Graham4 reported that following incubation, the motility of the extended semen (approximately 30–40 million spermatozoa per mL in a standard skim-milk-glucose extender and incubated for 30 min at 37◦ C) is determined using computerassisted semen analysis (CASA; Hamilton Thorne CEROS Model). Minimums of 400 motile or 1000 total sperm from more than five fields are analyzed for motion characteristics at 60 frames/s for 40 frames. A sperm was considered progressively motile if it
had an average path velocity (VAP) > 50 µm/s and straightness (STR) value > 75%. A sperm with VAP < 20 µm/s was considered non-motile. Results of the HOS test give good correlation with progressive motility. Katila et al.42 tested commercially used frozen semen from 31 stallions and compared results with foaling rates of 1085 mares. The HOS test was carried out using a 100 mOsM solution and an incubation of 45 min at 37◦ C. A significant correlation was found between foaling rate and the HOS test performed on sperm immediately after thawing or after an incubation of 3 hours at 37◦ C.
Sperm dose for insemination In general, it appears that insemination doses exceeding 250 × 106 progressively motile sperm post-thaw are likely to optimize fertility (for review see Metcalf;43 see also Table 316.2). Recently, researchers have attempted to refine insemination procedures to allow the deposition of
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Freezing Semen reduced numbers of spermatozoa to achieve pregnancy rates comparable with conventional artificial insemination (see Morris44 for review). Transrectally guided and hysteroscopic deep uterine insemination techniques have allowed insemination of doses as low as one million motile spermatozoa. Pregnancy results after inseminating mares with frozen/thawed stallion semen using low-dose combined with conventional and advanced artificial insemination techniques, such as deep uterine, hysteroscopic, oviductal are illustrated in Table 316.3.
References 1. Tischner M. Evaluation of deep-frozen semen in stallions. J Reprod Fertil 1979;27(Suppl.):53–9. 2. Vidament M, Dupree AM, Julienne P, Evian A, Noue P, Palmer E. Equine frozen semen: freezability and fertility field results. Theriogenology 1997;48:907–17. 3. Vidament M. French field results (1985–2005) on factors affecting fertility of frozen stallion semen. Anim Reprod Sci 2005;89:115–36. 4. Loomis PR, Graham JK. Commercial semen freezing: Individual male variation in cryosurvival and the response of stallion sperm to customized freezing protocols. Anim Reprod Sci 2008;105:119–28. 5. Katila T. In vitro evaluation of frozen-thawed stallion semen: a review. Acta Vet Scand 2001;42:201–17. 6. Kuisma P, Andersson M, Koskinen E. Fertility of frozenthawed stallion semen cannot be predicted by the currently used laboratory methods. Acta Vet Scand 2006;48: 14–22. 7. Samper JC, Morris CA. Current methods for stallion semen cryopreservation: a survey. Theriogenology 1998;49:895–903. 8. Vidament M, Cognard E, Yvon J-M. Evaluation of stallion semen before and after freezing. Reprod Dom Anim 1998;33:271. 9. Janett F, Thun R, Niederer K. Seasonal changes of semen quality and freezability in the Warmblood stallion. Theriogenology 2003;60:453–62. 10. Sieme H, Katila T, Klug E. Effect of semen collection practices on sperm characteristics before and after storage and on fertility of stallions. Theriogenology 2004;61:769–84. 11. Knop K, Hoffmann N, Rath D, Sieme H. Effects of cushioned centrifugation technique on sperm recovery and sperm quality in stallions with good and poor semen freezability. Anim Reprod Sci 2005;89:294–7. 12. Ecot P, Decuadro-Hansen G, Delhomme G, Vidament M. Evaluation of cushioned centrifugation technique for processing equine semen for freezing. Anim Reprod Sci 2005;89:245–8. 13. Moore AI, Squires EL, Graham JK. Effect of seminal plasma on the cryopreservation of equine spermatozoa. Theriogenology 2005;63:2372–81. 14. Sieme H. Current status of cryopreservation of stallion spermatozoa. In: Proceedings of the SIVE 13th Congresso Nazionale Multisala, 26–28 January 2007, Bologna, Italy, pp. 68–9. 15. Barker CAV, Gandier JCC. Pregnancy in a mare resulting from frozen epididymal spermatozoa. Can J Comp Vet Sci 1957;21:47–51. 16. Merkt H, Klug E, Krause D, Bader H. Results of long term
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30. 31. 32.
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storage of stallion semen frozen by pellet method. J Reprod Fertil 1975;23(Suppl.):105–6. Pace M, Sullivan JJ. Effect of timing insemination, numbers of spermatozoa and extender components on the pregnancy rate in mares inseminated with frozen semen. J Reprod Fertil 1975;23(Suppl.):115–21. Kneissl S. Cryopreservation of stallion semen. The influence of semen collection techniques, centrifugation, packaging forms, and methods of freezing on motility and plasma membrane integrity of spermatozoa. DVM Thesis, Veterinary University, Hannover, 1993. Heitland AV, Jasko DJ, Squires EL, Graham JK, Pickett BW, Hamilton C. Factors affecting motion characteristics of frozen-thawed stallion spermatozoa. Equine Vet J 1996;28:47–53. Cristanelli MJ, Amann RP, Squires EL, Pickett BW. Effects of egg yolk and glycerol levels in lactose-EDTA-egg-yolk extender and of freezing rate on the motility of frozen thawed stallion spermatozoa. Theriogenology 1985;24:681–6. Combes GB, Varner DD, Schroeder FB, Blanchard TL. Effects of cholesterol on the motility and plasma membrane integrity of frozen equine spermatozoa after thawing. J Reprod Fertil 2000;56(Suppl.):127–32. Crockett EC, Graham JK, Bruemmer JE, Squires EL. Effect of cooling equine spermatozoa before freezing on post-thaw motility: preliminary results. Theriogenology 2001;55:793–803. Loomis PR, Amann RP, Squires EL, Pickett BW. Fertility of unfrozen and frozen stallion spermatozoa extended in EDTA-lactose-egg yolk and packaged in straws. J Anim Sci 1983;56:687–93. H˚aa˚ rd MC, H˚aa˚ rd MGH. Successful commercial use of frozen stallion semen abroad. J Reprod Fertil Suppl. 1991;44:647–8. Leipold SD, Graham JK, Squires EL, McCue PM, Brinsko SP, Vanderwall DK. Effect of spermatozoal concentration and number on fertility of frozen equine semen. Theriogenology 1998;49:1537–43. Vidament M, Ecot P, Noue P, Bourgeois C, Magistrini M, Palmer E. Centrifugation and addition of glycerol at 22◦ C instead of 4◦ C improve post-thaw motility and fertility of stallion spermatozoa. Theriogenology 2000;54:907–19. Devireddy RV, Swandlund DJ, Olin T, Vincente W, Troedsson MHT, Bischof JC, Roberts KP. Cryopreservation of equine sperm: optimal cooling rates in the presence and absence of cryoprotective agents determined using differential scanning calorimetry. Biol Reprod 2002;66: 222–31. Cochran JD, Amann RP, Squires EL, Pickett BW. Effects of centrifugation, glycerol level, cooling to 5◦ C, freezing rate and thawing rate on the post-thaw motility of equine sperm. Theriogenology 1984;22:25–38. Mazur P, Leibo SP, Chu EH. A two-factor hypothesis of freezing injury. Evidence from Chinese hamster tissueculture cells. Exp Cell Res 1972;71:345–5. Mazur P. Freezing of living cells: mechanisms and implications. Am J Physiol 1984;247:C125–C142. Watson PF. The causes of reduced fertility with cryopreserved semen. Anim Reprod Sci 2000;60–61:481–92. Sieme H, Harrison RAP, Petrunkina AM. Cryobiological determinants of frozen semen quality, with special reference to stallion. Anim Reprod Sci 2008;107:276–92. Holt WV. Fundamental aspects of sperm cryobiology: the importance of species and individual differences. Theriogenology 2000;53:47–58.
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34. Koebe HG, Werner A, Lange V, Scildberg FW. Temperature gradients in freezing chambers of rate-controlled cooling machines. Cryobiology 1993;30:349–52. 35. Zirkler H, Gerbes K, Klug E, Sieme H. Cryopreservation of stallion semen collected from good and poor freezers using a directional freezing device (Harmony CryoCare –Multi thermal Gradient 516). Anim Reprod Sci 2005;89:291–4. 36. Saragusty J, Gacitua H, Pettit MT, Arav A. Directional freezing of equine semen in large volumes. Reprod Dom Anim 2007;42:610–15. 37. Goolsby HA, Elanton JR, Prien SD. Preliminary comparisons of a unique freezing technology to traditional cryopreservation methodology of equine spermatozoa. J Equine Vet Sci 2004;24:314–18. 38. Isachenko V, Isachenko E, Katkov I, Montag M, Dessole S, Nawroth F, van der Ven H. Cryoprotectant-free cryopreservation of human spermatozoa by vitrification and freezing in vapor: effect on motility, DNA integrity, and fertilization ability. Biol Reprod 2004;71:1167–73. 39. Sieme H, Martinsson G, Rauterberg H, Walter K, Aurich C, Petzoldt R, Klug E. Application of techniques for sperm selection in fresh and frozen-thawed stallion semen. Reprod Dom Anim 2003;38:134–40. 40. Loomis PR. Advanced methods for handling and preparation of stallion semen. Vet Clin North Am Equine Pract 2006;22:663–76. 41. Loomis PR. Artificial insemination of horses: where is it going? In: Proceedings of the Annual Conference of the Society of Theriogenology, Nashville, Tennessee, 1999, pp. 325–36. 42. Katila T, Koskinen E, Andersson M. Evaluation of frozenthawed semen. In: Proceedings of the A Dorothy Russell Havemeyer Foundation Workshop Advanced Current Topics in Stallion Veterinary Practice, 2000, pp. 54– 6. 43. Metcalf L. The efficient use of equine cryopreserved semen. Theriogenology 2007;68:423–8. 44. Morris LHA. Advanced insemination techniques in mares. North Am Equine Pract 2006;22:693–703. 45. W-Y He, 1986. cit. in Amann, RP, Pickett BW. Principles of cryopreservation and a review of cryopreservation of stallion spermatozoa. Equine Vet Sci 1987;7:145–73. ¨ 46. Martin JC, Klug E, Gunzel-Apel A-R. Centrifugation of stallion semen and its storage in large volume straws. J Reprod Fertil Suppl 1979;27:47–51. 47. Palmer E. Factors affecting stallion semen survival and fertility. In: Proceedings of the 10th International Congress of Animal Reproduction and Artificial Insemination, Urbana, IL, 1984, pp. 377–78. 48. Klug E, Treu H, Hillmann H, Heinze H. Results of insemination of mares with fresh and frozen stallion semen. J Reprod Fertil Suppl 1975;23:107–10. ¨ 49. Muller Z. Fertility of frozen equine semen. J Reprod Fertil Suppl 1982;32:47–51. ¨ 50. Muller Z. Practicalities of insemination of mares with deep-frozen semen. J Reprod Fert Suppl 1987;35:121– 25. 51. Kloppe LH, Varner DD, Elmore RG, Bretzlaff KN, Shull JW. Effect of insemination timing on the fertilizing capacity of frozen-thawed equine spermatozoa. Theriogenology 1988;29:429–39.
52. Vidament M, Magistrini M, Palmer E, Clement F. Equine artificial insemination in French National Studs. Reprod Dom Anim Suppl 1999;6:61–6. 53. Barbacini S, Marchi V, Zavaglia G. Equine frozen semen: results obtained in Italy during the 1994–1997 period. Equine Vet Educ 1999;11(2):109–12. 54. Katila T. Effects of hormone treatments, season, age and type of mares on ovulation, twinning and pregnancy rates of mares inseminated with fresh and frozen semen. Pferdeheilkunde 2003;6:619–24. 55. Sieme H, Sch¨afer T, Stout TAE, Klug E, Waberski D. The effects of different insemination regimes on fertility in mares. Theriogenology 2003;60:1153–64. 56. Barbacini S, Necchi D, Postinger G, Parmeggiani F, Squires EL. What to expect when barren mares are inseminated with frozen-thawed semen. In: Proceedings of the 54th Annual Convention of the American Association of Equine Practitioners, 2008, pp. 274–5. 57. F´eo JCSA, Oba E, Baranabe RC, Basile JR. Artificial insemination in the equine: Distribution of spermatozoa in the genital tract : A comparison of mares inseminated in the uterine body and the uterine horn ipsilateral to the ovulatory follicle. In: Proceedings of the 12th International Congress of Animal Reproduction, 1992, vol. 3, pp. 1545–7. 58. Morris LHA, Tiplady C, Wilsher S, Allen WR. Hysteroscopic insemination of mares with low numbers of frozen/thawed ejaculated and epididymal spermatozoa. In: Proceedings of the 5th International Symposium on Equine Embryo Transfer. Havemeyer Foundation Monograph Series 3, 2000, pp. 4–6. 59. Alvarenga MA, Onoe E, Fonseca H, Trinque CL, Lima MM, Pinheiro LPL. Utilization of endoscopic insemination for the application of stallion frozen semen. Proceedings of the XIV Congresso Brasileiro de Reproducao Animal. Rev Bras Reprod Anim 2001;25:361–2. 60. Alvarenga MA, Le`ao KM. Hysteroscopic insemination of mares with low number of frozen thawed spermatozoa selected by percoll gradient. Theriogenology 2002;58:651– 3. 61. Squires EL, Reger HP, McLellan LJ, Bruemmer JE. Effect of time of insemination and site of insemination on pregnancy rates with frozen semen. Theriogenology 2002;58:655–8. 62. Morris LHA, Tiplady C, Allen WR. Pregnancy rates in mares after a single fixed time hysteroscopic insemination of low numbers of frozen-thawed spermatozoa onto the uterotubal junction. Equine Vet J 2003;35:197–201. 63. Sieme H, Bonk A, Hamann H, Klug E, Katila T. Effects of different artificial insemination techniques and sperm doses on fertility of normal mares and mares with abnormal reproductive history. Theriogenology 2004;62: 915–28. 64. Crowe CAM, Ravenhill PJ, Hepburn RJ, Shepherd CH. A retrospective study of artificial insemination of 251 mares using chilled and fixed time frozen-thawed semen. Equine Vet J 2008;40:572–6. 65. Pillet E, Batellier F, Duchamp G, Furstoss V, LeVern Y, Kerboeuf D, Vidament M, Magistrini M. Freezing of stallion semen in INRA96-based extender improves fertility rates in comparision with INRA82. Dairy Sci Technol 2008;88:257–265.
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Chapter 317
Freezing Epididymal Spermatozoa Jason E. Bruemmer
Introduction A growing awareness of new technologies in semen preservation and a requirement to preserve valuable genetics has led to an increase in previously unavailable procedures such as freezing semen from castrated or recently deceased males. As a general rule, the majority of preserved equine sperm cells are collected as a result of ejaculation and either cooled to 5◦ C and stored for 24–48 hours or are frozen and stored in liquid nitrogen for an indefinite time. Situations may arise, however, that prevent such semen collection and preservation. These include collection of semen from males that have been castrated or that have a catastrophic fatal injury. In these cases, the stallion’s cauda epididymides should contain sperm capable of fertilization, thus providing a significant source of salvageable genetic material in the case of elected castration or death. Unfortunately, the timing, and often location, of the castration or postmortem tissue harvest does not lend itself well to sperm collection and preservation. For these reasons, researchers have been developing methods to (i) harvest epididymal sperm cells, (ii) successfully transport epididymal tissue, (iii) cryopreserve epididymal sperm, (iv) effectively inseminate mares with the cryopreserved epididymal sperm, and (v) use these sperm cells in sophisticated in vitro fertilization attempts including intracytoplasmic sperm injection (ICSI). These methods are not independent as each is an integral part of the other.
Harvesting epididymal sperm Epididymal collection requires cannulation of the vas deferens, the epididymides, or surgical removal. Spermatozoa have successfully been collected via
cauda epididymal aspiration from both anesthetized and standing stallions (Colorado State University, unpublished observations). While harvest of spermatozoa in this manner is certainly possible, very few cells are collected compared with collection after epididymal removal, and are of little value in postmortem situations. The two caudal epididymides of a normal, sexually rested, adult stallion should contain over 50 × 109 sperm or approximately 61% of the sperm in the excurrent duct system.1 It has therefore been concluded that the most efficient means of obtaining the greatest number of sperm requires surgical removal of the epididymides for a single and final collection. To accomplish this, both testes and their associated epididymides should be removed using standard surgical castration procedures. It is imperative that the cauda epididymides and as much of the ductus deferens as possible remain intact. The ductus deferens should be ligated with surgical suture material as high as possible to prevent loss of sperm cells as fertile sperm can be isolated from both regions.2 If possible, castration should be performed prior to euthanasia as the effects of euthanasia compounds on sperm viability have yet to be determined. Stallions may be castrated while anesthetized as the effects of the gas anesthetic halothane appear to be negligible.3 Each testis and associated epididymis should then be rinsed free of blood and other contaminants with sterile saline and placed in a clean container, such as a rectal sleeve or plastic bag. At this time point, the sample can be processed immediately or transported to a facility capable of the techniques described below. On arrival at the appropriate laboratory facility, each epididymis should be dissected free of the testis (Figure 317.1). The tail and ductus deferens are then isolated (Figure 317.2) and spermatozoa can be
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 317.1 Epididymis and proximal ductus deferens removed from the testes.
Figure 317.3 Float-up collection of ductus deferens and epididymis.
collected using one of two methods: the float-up method (Figure 317.3) or the more common retrograde flush technique (Figure 317.4). In the float-up method, the cauda epididymides and proximal ductus deferens are incised in 12–15 locations, suspended in approximately 5 mL of semen extender, and sperm are allowed to ‘float’ into the extender for upwards of 10 minutes. Most laboratories have had better success when the epididymides are flushed in a retrograde fashion than that reported with the float-up technique. Routinely 15–20 billion sperm are collected compared with the 4–5 billion reported using the float-up method. This technique requires the use of approximately 10 mL per epididymis of a commercial semen extender, a clear modified Tyrode’s medium, or seminal plasma collected from another stallion (Figure 317.4). Briefly, the tubules of the vas deferens and caudal epididymides are dissected free of the fascia, usually working distally from vas deferens toward the corpus epididymis. Termination of dissection occurs
once the tubule of the cauda become indistinguishable from those of the corpus or at least until it is easily visualized. A blunted 14- to 16-gauge needle or a 14-gauge catheter, Pasteur pipette, or fire-polished pipette is attached to a syringe containing the aforementioned flush medium. The needle or pipette is placed in the previously ligated vas deferens while the caudal epididymis is suspended in a beaker or 50-mL conical tube (Figure 317.4). The most distal end of the caudal epididymis (ideally the previous site of union with the corpus) is incised to allow sperm collection. The flush may require a certain
Figure 317.2 Isolation of cauda epididymis and proximal ductus deferens.
Figure 317.4 Retrograde collection method.
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Freezing Epididymal Spermatozoa amount of pressure, so care should be taken to assure that the tissue does not slip off the end of the needle or pipette.
Transporting epididymal tissue The testis and epididymides should be placed in a passive cooling device designed for the transport of cooled semen and sent to a reproductive referral center that offers the service of collecting and freezing epididymal sperm or processed by a practitioner at an appropriate facility. Make certain that the coolant source has been added. Sperm from many species, including canine,4 human,5 deer,6 and goat7 have been processed and successfully frozen after at least 24 hours of storage at 5◦ C. Research suggests that sperm harvested from stallion tissues following a 24-hour storage period at 5◦ C also remain viable,8, 9 and, in some cases, comparable to freshly processed ejaculated samples.10, 11
Cryopreservation of epididymal sperm Once the concentration of the harvested cells has been determined, most suggest that a conventional freezing protocol should be followed. Briefly, the concentration of the recovered sperm is determined via a hemocytometer or a Nucleocounter (SP 100) when using any opaque medium or spectrophotometrically following collection in a clear Tyrode’s type medium or seminal plasma. Epididymal sperm are then diluted to the desired final concentration, commonly either 200 × 106 or 400 × 106 cells/mL in a freezing extender and frozen using standard techniques.12 The type of extender used is based on the history of success of previous cryopreservation of ejaculated semen, when available. The type of freezing extender best suited to survival of cryopreserved epididymal sperm matched that of ejaculated sperm from a given stallion.10 If sperm from a given stallion has not been frozen previously, the sample may be split with half frozen in each of the two most available extenders. Seminal plasma harvested from a stallion of proven fertility can also be added to the sample if not used during the flushing procedure. Typical addition ranges from 0.5 mL per 1.6 billion sperm10 to 5% v/v.13 It has been hypothesized that fertility is increased by exposing epididymal sperm to seminal plasma before freezing. Post-thaw motility parameters may13, 14 or may not8, 11 be affected. Morris et al.15 reported that fertility of frozen–thawed epididymal sperm was not affected by exposure to seminal plasma. More recently, Heise et al.16 demonstrated that the pregnancy rate obtained with frozen–thawed
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epididymal spermatozoa that had been exposed to seminal plasma (as the flush medium) was similar to that of frozen–thawed ejaculated spermatozoa (5/18, 27.8% compared with 7/18, 38.9%). Furthermore, the combined pregnancy rates of epididymal spermatozoa inseminated either fresh or frozen–thawed exposed to seminal plasma was greater (P < 0.05) than that of similar unexposed spermatozoa.16 Five to 25 breeding doses, consisting of 800 × 106 sperm per dose, are typically obtained from a given stallion. Frozen epididymal sperm harvested from stallions both on premises and cooled–transported have been used to obtain pregnancies.
Insemination with cryopreserved epididymal sperm It warrants mention that the first equine pregnancy generated using frozen semen was obtained using frozen–thawed epididymal sperm.17 Pregnancies and live foals have since been produced by using both conventional artificial insemination9 and assisted reproductive techniques such as low-dose hysteroscopic insemination.11, 15 Pregnancy rates have ranged from 0–29% for conventional insemination methods15 to 66% using low-dose techniques.11 Papa et al.11 suggested that pretreatment with Sperm Talp enhanced pregnancy rates (66%). These results agree with the hypothesis suggested by Sostaric et al.,18 in which they point out that epididymal sperm, at the very least, have an impaired ability to capacitate and/or acrosome react. Concomitantly, these authors suggested that membrane progesterone receptors on stallion sperm are not evident and are therefore ‘hidden’ by a coating molecule produced within the epididymis. Together,12, 18 these results suggest the need to further explore pre-insemination treatment of epididymal sperm.
Cryopreserved epididymal sperm used for intracytoplasmic sperm injection ICSI requires only a single sperm for fertilization of the oocyte. This procedure is very useful when the supply of frozen semen is limited and cannot be replaced. Sperm frozen conventionally have been thawed, diluted, and refrozen to increase the number of potential uses for ICSI;19, 20 epididymal sperm have been harvested and frozen21 after the death of a stallion to produce offspring.22 In fact, results from the 2006 and 2007 breeding season (Colorado State University, unpublished observations) suggest that a greater percentage of oocytes cleaved following ICSI with frozen–thawed epididymal sperm (15/18,
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83%) than with frozen–thawed conventionally collected sperm (181/290, 62%). However, pregnancy rates at both 16 and 50 days were similar (43% vs 45% and 29% vs 29%) for epididymal versus ejaculated sperm, respectively. These data indicate that a greater early embryonic wastage may occur. However, these results might have been confounded as Carnevale and Ginther23 (and Colorado State University, unpublished observations) suggest that the average age of oocyte donors in the epididymal group was 20 plus years, which was considerably older than those in the conventional group, and this may have affected future oocyte development.
References 1. Amann RP, Thompson Jr DL, Squires EL, Pickett BW. Effects of age and frequency of ejaculation on sperm production and extragonadal reserves in stallions. J Reprod Fertil Suppl 1979;27:1–6. 2. McKinnon AO, Voss JL (eds). Equine Reproduction. Philadelphia: Lea & Febiger, 1993. 3. Schulman ML, Gerber D, Nurton J, Guthrie AJ, Joubert K, Volkmann DH. Effects of halothane anaesthesia on the cryopreservation of epididymal spermatozoa in pony stallions. Equine Vet J 2003; 35:93–5. 4. Marks SL, Dupus J, Mickelsen WD, Memon MA, Platz CC. Conception by use of postmortem epididymal semen extraction in a dog. J Am Vet Med Assoc 1994;204:1639–40. 5. Sharma RK, Padron OF, Thomas AJ, Agarwal A. Factors associated with the quality before and after thawing of sperm obtained by microsurgical epididymal aspiration. Fertil Steril 1997;68:626–31. 6. Zomborsky Z, Zubor T, Toth J, Horn P. Sperm collection from shot red deer stags (Ervus elaphus) and the utilization of sperm frozen and subsequently thawed. Acta Vet Hung 1999;47:263–70. 7. Blash S, Melican D, Gavin W. Cryopreservation of epididymal sperm obtained at necropsy from goats. Theriogenology 2002;54:899–905. 8. Bruemmer JE, Reger H, Zibinski G, Squires EL. Effect of storage at 5C on the motility and cryopreservation of stallion epididymal spermatozoa. Theriogenology 2002;58:405–7. 9. James AN, Green H, Hoffman S, Landry AM, Paccamonti D, Godke RA. Preservation of equine sperm stored in the
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
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epididymis at 4C for 24, 48, 72 and 96 hours. Theriogenology 2002;58:401–4. Bruemmer JE, Moore A, Squires EL. Freezing epididymal sperm from transported testicles. Havemeyer Foundation Workshop Monogr Ser 2003;15:61–2. Papa FO, Melo CM, Fioratti EG, Dell’aqua Jr JA, Zahm FS, Alvarenga MA. Freezing of stallion epididymal sperm. Anim Reprod Sci 2008;107:293–301. Squires EL, Pickett BW, Graham JK, Vanderwall DK, McCue PM, Bruemmer JB. Cooled and Frozen Stallion Semen. Animal Reproduction and Biotechnology Laboratory, Bulletin No. 9. 1999. Stout TAE, Morris LHA, Li X. The effect of seminal plasma on the motility and cryopreservability of horse epididymal sperm. In: Proceedings of the Havermeyer Foundation Workshop: European Equine Gamete Group, Brewster, Masachusetts, 2–5 October, 2003, pp. 5–6. Volkmann DH, Gerber D, Erb HN. Comparison between freezability of ejaculated and epididymal stallion sperm (Abstract). Anim Reprod Sci 2001;68:340. Morris L, Tiplady C, Allen WR. The in vivo fertility of cauda epididymal spermatozoa in the horse. Theriogenology 2002;58:643–6. Heise A, K¨ahn W, Volkmann DH, Thompson PN, Gerber D. Influence of seminal plasma on fertility of fresh and frozenthawed stallion epididymal spermatozoa. Anim Reprod Sci 118(1):48–53, 2010. Barker CAV, Gandier JCC. Pregnancy in the mare resulted from frozen epididymal spermatozoa. Can J Comp Vet Med Vet Sci 1957;21:45–51. Sostaric E, Aalberts M, Gadella BM, Stout TAE. The roles of the epididymis and protosomes in the attainment of fertilizing capacity stallion sperm. Anim Reprod Sci 2008;107:237–48. McCue PM, Moore AI, Bruemmer, JE. Refreezing stallion spermatozoa for assisted reproduction. Reprod Fertil Dev 2004;16:176–7. Choi YH, Love CC, Varner DD, Hinrichs K. Equine blastocyst development after intracytoplasmic injection of sperm subjected to two freeze-thaw cycles. Theriogenology 2006;65:808–19. Bruemmer JE. Collection and freezing of epididymal stallion sperm. Vet Clin North Am Equine Pract 2006;22:677–82. Carnevale EM, Stokes JE, Squires EL, Campos-Chillon LF, Altermatt J, Suh JK. Clinical use of intracytoplasmic sperm injection in horses. Proc Am Assoc Equine Pract 2007;53:560. Carnevale EM, Ginther OJ. Defective oocytes as a cause of subfertility in old mares. Biol Reprod Monogr 1994;1:209–14.
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Chapter 318
Breeding with Frozen Semen Sandro Barbacini
The first pregnancy obtained by using frozen-thawed semen in the equine species dates back to 1957,1 and was obtained using epididymal semen. Nonetheless, for many years, this reproductive technology has achieved limited progress in the horse compared with other species (e.g. cattle). This was mainly due to the fact that for a very long time only a few horse registries allowed the use of frozen-thawed equine semen, during which time economic interests and resources allocated to research were minimal, thereby obstructing advances in this area. Today, almost all the horse registries allow the use of this reproductive technique, thus encouraging researchers to focus on identifying better semen freezing techniques and simpler artificial insemination protocols. This has resulted in an improvement in fertility of cryopreserved equine semen, further enhancing the use of the technique by breeders. Moreover, a few papers published over the last few years2–4 clearly showed that the development of improved techniques for freezing equine semen allowed a greater percentage (85–90%) of stallions, than has traditionally been reported in the literature, to be capable of producing semen with commercially acceptable post-thaw quality. Benefits of frozen semen include access to semen from stallions: (i) standing in other countries or continents and from competition stallions; (ii) that become ill or get infectious diseases, injured or overbooked during the breeding season; and (iii) when the mare is at the optimum time for breeding. However, success with frozen semen requires that the practitioner be familiar with proper thawing, evaluating, and handling of frozen semen, as well as with the breeding strategies employed to maximize fertility.5
Semen thawing and insemination As already described in Chapters 127 and 314, during the cooling and freezing processes, spermatozoa undergo a series of chemical and physical changes
that can irreversibly reduce their ability to fertilize an oocyte. The thawing procedure exposes spermatozoa to the same types of stresses but in reverse. Cells rehydrate as water moves back across the plasma membrane to stabilize the osmotic imbalance created when extracellular ice melts. Cryoprotectants diffuse out of the cells causing plasma membrane proteins and lipids to reorganize.6, 7 Thawing semen improperly (too fast or too slow for the freezing protocol used) can reduce the spermatozoa’s viability and decrease the chance of obtaining a pregnancy. The vast majority of equine frozen semen is currently packaged in 0.5-mL straws and is most commonly thawed in a water bath set at 37◦ C. To inseminate a mare with semen that is frozen in 0.5-mL straws the following handling and thawing instructions are recommended: 1. Prepare the mare’s external genitalia aseptically prior to opening the container. 2. Using precooled hemostats, or tweezers, remove one straw from the nitrogen container and immediately place it into a water bath at 37◦ C, being careful not to submerge the hemostats or tweezers in the water. It is important to make this transfer as quickly as possible, exposing the straws to room temperature for no more than 2 or 3 seconds. 3. Leave the straw in the water bath for at least 30 seconds to ensure proper thawing. How long the straw is kept at 37◦ C is not critical provided it is left in the water bath for at least 30 seconds to allow the semen to thaw fully. Thawed spermatozoa die more rapidly if kept at 39 or 40◦ C as opposed to 35 or 36◦ C. 4. The individual straw may remain in the water bath while the remaining straws are removed from the tank and thawed. The number of straws that comprise a single insemination dose may vary according to the freezing procedure employed. All frozen semen should be transported with detailed
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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thawing instructions, which should be supplied by the laboratory that processed the semen. After all the straws for the insemination dose have been thawed, they are removed from the water bath one by one and wiped completely, since water contamination of the straw contents would be spermicidal. While holding each straw vertically, so that the air bubble is positioned at the top (the top is usually sealed by heat or a sealing ball), cut the sealed end. Place the open end of the straw over a sterile container prewarmed to 37◦ C (a sterile 5-mL falcon snap-top tube, a centrifuge tube or red top blood tube) and cut off the plug on the other end of the straw. Allow the semen to empty into the sterile container and tap the end of the straw to ensure that all the semen is removed. Repeat this procedure for all of the thawed straws, combining the contents into a single container. Do not add extender to the semen unless requested to by the laboratory that froze the semen as it could dramatically alter the osmotic pressure and irreversibly damage the spermatozoa. Draw the thawed semen into a sterile prewarmed insemination pipette using an all plastic syringe and inseminate the mare just as you would do with fresh semen. Use a 5-mL air dam behind the content to ensure that all the semen is deposited in the mare’s uterus
Although most commercial laboratories now freeze semen in 0.5-mL straws, some semen is still frozen in 2.5, 4.0, or 5.0 mL macrotubes. These straws are generally thawed at 50◦ C for 45 seconds. With these straws it is critical that they do not remain in the 50◦ C water bath for more than 45 seconds, to prevent damage from exposure to elevated temperatures. After thawing, semen frozen in macrotubes has to be placed in a sterile container prewarmed to 37◦ C (a sterile 5-mL falcon snap-top tube, a centrifuge tube or red top blood tube) and kept there until insemination is carried out. Equine frozen semen can also be frozen in containers other than straws, like plastic bags made out of polypropylene and aluminum packages. Nonetheless, the popularity of these packages has, over the years, been overridden by the use of the abovementioned different sizes of straws.
Fertility Data published in the last 15 years clearly show that excellent pregnancy rates can be obtained by using equine frozen-thawed semen in a commercial setting.8 The high fertility of cryopreserved stallion semen that is empirically observed in the field may
be attributed to the more sophisticated freezing methods and simpler insemination protocols developed over the last few years, but may also stem from the greater familiarity of practitioners with the techniques for properly thawing, evaluating and handling frozen semen. In addition, pregnancy rates have improved as semen with known superior fertility data is often used preferentially. Nevertheless, the majority of the frozen semen fertility data were collected under field conditions and not from controlled studies. Ideally, to determine more accurately the fertility of equine frozen semen, experiments should be designed so that large numbers of mares are randomly assigned to be inseminated with frozenthawed semen of different stallions and containing different numbers of progressively motile, morphologically normal spermatozoa. Data reported in the literature have reported per-cycle pregnancy rates ranging from 38% to 73%,3,9,17 and seasonal pregnancy rates from 76% to 82%.10, 11 Nevertheless, these results could have been strongly affected not only by the post-thaw semen quality and stallion selection, but also by the mare management techniques and mare selection criteria. In addition seasonal pregnancy rates may be further influenced by the number of cycles that mares are available to be bred on. There are many factors that can positively or negatively affect the pregnancy outcome of mares bred with cryopreserved stallion semen and some are discussed below. Time of insemination and the effects of sperm number and frequency of insemination One of the challenges for those attempting to freeze equine spermatozoa is dealing with the subject-tosubject variability in the cryosurvival of their semen and in the minimum number of post-thawing progressively motile sperms that are necessary to gather a pregnancy and below which the fertility of a given stallion fails drastically. At present, limited data are available on the minimal number of spermatozoa required to achieve optimal fertility with frozenthawed spermatozoa,18,22 and it is therefore desirable that controlled studies including large numbers of stallions are performed by scientists, in order to give the equine industry clear guidelines on this issue. A major limiting factor to the widespread application of equine frozen semen is the cost associated with the intense management of mares being inseminated. It is generally recommended that frozen stallion semen be introduced into the mare reproductive tract within 12 hours before or up to 6 hours after ovulation. Practitioners should be aware that inseminating mares with frozen semen beyond 6 hours after ovulation can lead to reduced fertility. Although
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Breeding with Frozen Semen there are no data about using cryopreserved semen, Woods et al. (1990) demonstrated that delaying insemination with fresh semen further than 8 hours after ovulation results in a depression of fertility. The presumed shortened lifespan for frozen-thawed stallion spermatozoa in the mare reproductive tract combined with the ‘by the dose, no guarantee’ system of marketing semen has led to the practice of reproductive examination of mares three to four times per day or more frequently as ovulation approaches and prior to insemination with cryopreserved semen. In fact, when using frozen-thawed semen, many veterinarians examine the mare’s reproductive tract with such frequency to permit insemination with a single dose of semen during the periovulatory period. This usually requires that mares are boarded at a clinic or that late-night farm calls be made by the practitioner to ensure that a post-ovulation insemination is performed within the critical 6-hour window after ovulation. The cost in veterinary care to the mare owner is substantial and often discourages the use of frozen semen. Stallion owners who sell semen by the dose for hundreds, or even thousands, of dollars or euros are forcing the mare owners to utilize this type of protocol because the cost of the veterinary care is less than the cost of the additional semen required to inseminate a mare more than once per cycle. Many stallion owners now provide multiple doses per cycle and are paid when the mare is pregnant. In these cases, using frozen semen is just another mechanism of achieving a pregnancy. One insemination post-ovulation after 3–4 examinations per cycle can generate good fertility results. Barbacini et al.29 collected data from 559 Warmblood mares that were inseminated with frozen-thawed semen within a 6-hour pre- and a 6-hour postovulation period of time: a 43% per cycle and a 77% overall pregnancy rate were accomplished by using the following protocol:
1. Mares were ultrasonographically monitored during oestrus every 24 hours until at least one follicle of 35 mm or greater was detected on the ovaries. 2. 2,000 IU of human chorionic gonadotropin (hCG) were administered intravenously. 3. Ultrasound examination was performed 12 hours after hCG administration, then every 4–8 hours until ovulation occurred. 4. Insemination was performed with one insemination dose when the follicle was very close to ovulation or when a fresh ovulation was detected. Nevertheless, expanded use of equine frozenthawed semen is dependent upon simplified breeding strategies. For this reason, it has been recommended to use a timed insemination protocol for cost-effective management of mares inseminated with frozen semen.14, 24 This protocol involves daily ultrasonographic examinations during oestrus, induction of ovulation using hCG or deslorelin following detection of a > 35-mm follicle, and insemination at 24 and 40 hours after the ovulatory agent injection. Using this insemination schedule, mares that ovulate 18–52 hours after administration of hCG or deslorelin will have spermatozoa deposited in their reproductive tract within 12 hours prior to ovulation or within 6 hours after ovulation, or both (Figure 318.1). In a clinical trial conducted in Italy,25 26 out of 34 mares became pregnant (76%) after two timed inseminations versus 15 of 21 (71%) following a single insemination within 6 hours post-ovulation. In the same paper, the authors also reported the outcome of a controlled study carried out in Colorado where they found no difference in embryo recovery rates for mares inseminated once within 6 hours post-ovulation with 800 × 106 total frozen-thawed spermatozoa (60%) versus mares inseminated twice at 24 and 40 hours post-deslorelin with 400 × 106 total spermatozoa per insemination (55%). Moreover, in a
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Hrs after injection Mares ovulating during this period will have been inseminated within 6 hours after or within 12 hours prior to ovulation . Figure 318.1 Timed insemination protocol. hCG, human chorionic gonadotropin.
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retrospective field study involving 193 estrus cycles, Barbacini26 reported a similar per-cycle pregnancy rate when mares, highly selected for their reproductive potential, were inseminated twice at 24 and 40 hours post-hCG with 400 × 106 (61%) or 800 × 106 (69%) total spermatozoa per insemination. These data support the concept that two inseminations timed to occur before and after ovulation yield comparable pregnancy rates to a single post-ovulation insemination. Sieme et al.13 reported that mares inseminated twice per cycle at 24-hour intervals had a 50% percycle pregnancy rate, and mares inseminated once averaged a 42% per-cycle pregnancy rate. Pregnancy rates for mares inseminated once within 12 hours prior to ovulation or once within 12 hours postovulation were 41% and 50% respectively. Vidament et al.27 and Samper28 also reported that pregnancy rates with frozen semen were higher when mares were inseminated more than once per cycle. Finally, Loomis and Squires5 examined retrospectively two data sets collected during the 2002 and 2003 breeding seasons and reported similar pregnancy rates for mares inseminated once or multiple times in a given cycle (51.5% vs 51.7% for 2002, and 47.1% vs 46.1% for 2003). Mares inseminated twice on a cycle, once before and once after ovulation, became pregnant at a rate similar to mares inseminated once within 6 hours post-ovulation (48.1% vs 47.3%). Based on data published on the use of multiple inseminations for breeding mares with frozen-thawed semen, it is clear that a two-dose timed insemination protocol allows a practitioner to examine mares once daily during normal hours without compromising fertility. However, use of multiple inseminations per cycle may not be appropriate for all breeding situations – for example, mares that are susceptible to post-breeding endometritis such as older or barren mares. These mares may require a more intense management scheme in order to minimize invasion of the susceptible uterus. Older mares may also be less responsive to ovulatory agents such as hCG29 and therefore their ovulation time could be unpredictable. In addition, semen from some stallions is limited either through death, lack of opportunity to freeze, or value of the insemination dose. Mare’s reproductive status Mare selection based on the mare’s reproductive status influences the results obtained with frozen equine semen. Despite some reports of a higher fertility in foaling than in maiden or barren mares,10, 15, 27 mares bred with frozen-thawed semen are usually not inseminated on their foal heat. Squires et al.15 reported that maiden mares have a lower per-cycle fertility than foaling or barren mares (35% vs 54% and 53%),
whereas Barbacini et al.10 showed a lower per season fertility of only maiden mares versus foaling mares (76% vs 88%). All the maiden mares taken into consideration by Barbacini et al. were Warmblood breeds. It is well known that Warmblood mares tend to perform in sports events much longer than Thoroughbred, Standardbred or Quarter Horse mares, thereby resulting in a greater number of ‘old maidens’ in this breed. The decreased fertility of maiden mares observed in these studies is similar to findings reported by Samper et al.,12 who reported that maiden mares over age 7 years had reduced fertility compared with younger maiden mares. There are many anecdotal reports within the horse breeding industry suggesting that barren mares are poor candidates for artificial insemination with frozen-thawed semen. However, Barbacini30 retrospectively analyzed data collected on 285 barren mares inseminated in northern Italy and reported an acceptable per-cycle (37%) and seasonal (71%) pregnancy rate when these mares were bred with frozen semen. Moreover, the authors obtained a significantly (p < 0.05) higher per-cycle pregnancy rate (52%) in barren mares inseminated with frozen semen that had post-insemination fluid accumulation and were treated with oxytocin and/or uterine lavage compared to barren mares with no post-insemination fluid and no treatment (29%). Dell’Aqua et al.31 showed that barren mares with a history of intense inflammatory uterine responses after insemination had significantly (p < 0.05) greater pregnancy rates when treated with corticosteroids before and at the time of insemination, compared to untreated barren mares. The authors of the above mentioned paper speculated that corticosteroid therapy can reduce the intense uterine reaction that happens in barren mares after insemination and can therefore be an option to improve pregnancy rates in mares with poor breeding history. Mare age It is well known that fertility greatly decreases in older geriatric mares when fresh semen or natural mating is used.32 No significant effect of mare age on pregnancy rates with frozen semen when mares were 3 to 16 years of age was reported by Vidament et al.,27 Barbacini et al.,10 Loomis and Squires,5 and Barbacini.30 However, fertility was reduced for mares older than 16 years of age. Post-AI fluid accumulation Post-breeding intrauterine fluid accumulation is a physiological inflammatory response that may help to clear the uterus of foreign material such as excess spermatozoa, seminal plasma and bacteria.33, 34
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Breeding with Frozen Semen Persistent post-breeding uterine fluid accumulation has been associated with a decrease in fertility after natural mating or artificial insemination (AI) of fresh semen.35, 36 Predisposing factors to persistent uterine fluid accumulation are reduced myometrial contractions, poor lymphatic drainage, degenerative uterine changes, and cervical incompetence. Reproductively normal mares are typically able to expel uterine fluid quickly after inseminations, whereas susceptible mares accumulate fluid in their uterine lumen for more than 12 hours after breeding or insemination.34 It is commonly stated that insemination with cryopreserved semen leads to greater post-insemination fluid accumulation than insemination with fresh or cooled semen, or after natural mating. Apparently, there is only one controlled study on this comparison, by Kotilainen et al.37 These authors reported that infusion of frozen semen resulted in a greater inflammatory response than natural breeding. In a field study,38 16% of mares naturally mated had persistent post-breeding fluid accumulations compared with a 30% rate reported for mares inseminated with frozen semen.10, 29 More recently, Watson et al.39 reported a post-breeding fluid accumulation rate of 16% in mares inseminated with frozen-thawed semen, which is identical to that reported for natural mating.38 It is difficult to compare studies because details of mare selection and insemination or breeding frequencies are not always reported, and individual criteria and ability to determine and detect uterine fluid mare vary with experience and equipment. A higher proportion of barren and aged mares in a study would increase the incidence of post-breeding fluid accumulation.40 When no infertile or aged mares were considered in the study of Watson et al.,39 the incidence of uterine fluid accumulation after breeding with frozen semen was 20%, which is similar to the reported incidence after mares were naturally mated and treated with antibiotics after breeding.38 However, the question has not been completely answered about whether fluid accumulation after insemination with frozen semen is greater than that for mares bred with cooled semen or naturally mated. Watson et al.39 and Barbacini et al.40 reported that the combined use of oxytocin and uterine lavage on mares with post-breeding fluid accumulation resulted in satisfactory pregnancy rates per cycle (45% and 42% respectively for the two studies) and suggested that treating mares for uterine fluid as early as 12 hours after insemination may be unnecessary because fluid accumulation peaks 8 hours after insemination with fresh semen. Perhaps if one waited until 24 hours after insemination, then fewer mares would have significant uterine fluid accumulation. It has been suggested that inseminating twice during the cycle with frozen-thawed semen may result
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in a higher incidence of post-breeding endometritis; however, some recent studies have not supported this hypothesis.24, 25, 41 Embryonic loss It is well known that embryonic death is a substantial cause of economic loss in the horse breeding industry. The incidence of embryonic loss during the first 14–50 days of gestation has been reported to vary between 5% and 24% when using natural mating or fresh semen.42 Limited information is available on the incidence of embryonic loss in mares inseminated with frozen-thawed semen. Barbacini et al.45 collected data from 464 Warmblood mares that were inseminated with cryopreserved semen within 6 hours preand 6 hours post-ovulation, and found an overall conceptus loss rate of 8.9% between 14 and 50 days of gestation. No significant difference in embryonic loss rate was observed when mares were inseminated pre- (8.1%) or post-ovulation (9.3%), showing that the incidence of embryonic death is not affected by inseminating mares anywhere in the periovulatory period. The authors also analyzed the data by subdividing the mares into reproductive and age groups. The barren mare group displayed greater embryo losses (14.6%) than the maiden mares (5.3%) and the foaling mares (7.8%), and this is probably related to the fact that most of the subfertile mares were concentrated in this barren mare group. This retrospective study also showed that mares older than 16 years of age had a higher (15.6%) embryonic death incidence than the younger broodmares (8.2%): this is probably associated with an abnormal uterine environment in older mares.43 Cooled versus frozen semen fertility There is a common belief in the equine breeding industry that cooled semen has a higher fertility than frozen-thawed semen; however, there are few studies on this subject. In a controlled study, Jasko et al.44 reported that per-cycle fertility of cooled semen was greater than that of frozen-thawed semen (65% vs 56%). However, Loomis11 and Squires et al.15 reported similar pregnancy rates for mares in a commercial setting inseminated with cooled versus frozen semen. Loomis11 compared pregnancy rates of 850 mares inseminated with cooled semen collected from 16 stallions at three well-managed farms, with pregnancy rates of frozen semen inseminations from 106 stallions used to inseminate 876 mares. First-cycle and seasonal pregnancy rates were 59.4% and 74.7% for cooled semen, and 51.3% and 75.6% for frozen-thawed semen. First cycle pregnancy rates were higher (p < 0.05) with cooled semen compared
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to frozen but not seasonal pregnancy rates.11 Squires et al.15 obtained data from six different facilities where mares were inseminated with cooled or frozen-thawed semen. Per-cycle pregnancy rates were similar for those mares inseminated with cooled (44% out of 272 cycles) versus frozen semen (46% out of 536 cycles). There is no doubt that data collected under field conditions are somewhat compromised; nevertheless, the fertility results obtained by Loomis11 and Squires et al.15 support the position that comparable per-cycle and seasonal pregnancy rates may be obtained using frozen semen, as compared with liquid-cooled semen, in a commercial setting. Nonetheless, individual stallion variation must be considered in such fertility trials, and was not addressed in these reports.
References 1. Barker CAV, Gaudier JCC. Pregnancy in a mare resulted from frozen epididymal spermatozoa. Can J Comp Med Vet Sci 1957;21:47–53. 2. Loomis PR. Artificial insemination of horses: where is it going? Proceedings of the Annual Conference of the Society for Theriogenology 1999, pp. 325–36. 3. Vidament M. French field results (1985–2005) on factors affecting fertility of frozen stallion semen. Anim Reprod Sci 2005;89:115–36. 4. Loomis PR, Graham JK. Commercial semen freezing: individual male variation in cryosurvival and the response of stallion sperm to customized freezing protocols. Anim Reprod Sci 2008;105:119–28. 5. Loomis PR, Squires EL. Frozen semen management in equine breeding programs. Theriogenology 2005;64:480–91. 6. Amann RP, Pickett BW. Principles of cryopreservation and a review of stallion spermatozoa. J Equine Vet Sci 1987;7:145–73. 7. Hammerstedt RH, Graham JK, Nolan JP. Cryopreservation of mammalian sperm: What we ask them to survive. J Androl 1990;11:73–88. 8. Metcalf L. The efficient use of equine cryopreserved semen. Theriogenology 2007;68:423–8. 9. Metcalf L. Maximising reproductive efficiency in private practise; the management of mares and the use of cryopreserved semen. In: Proceedings of the Annual Meeting Society for Theriogenology 1995, pp. 155–9. 10. Barbacini S, Marchi V, Zavaglia G. Equine frozen semen: results obtained in Italy during the 1994–1997 period. Equine Vet Educ 1999;11:109–12. 11. Loomis PR. The equine frozen semen industry. Anim Reprod Sci 2001;68:191–200. 12. Samper JC, Vidament M, Katila T, Newcombe J, Estrada A, Sargeant J. Analysis of some factors associated with pregnancy rates of frozen semen: a multi-center study. Theriogenology 2002;58:647–50. 13. Sieme H, Schafer T, Stout TA, Klug E, Waberski D. The effects of different insemination regimens on fertility in mares. Theriogenology 2003;60:1153–64. 14. Barbacini S, Loomis RP, Squires EL. The effect of sperm number and frequency of insemination on pregnancy rates of mares inseminated with frozen-thawed spermatozoa. Anim Reprod Sci 2005;89:203–5.
15. Squires EL, Barbacini S, Matthews P, Byers W, Schwenzer K, Steiner J, Loomis PR. Retrospective study of factors affecting fertility of fresh, cooled and frozen semen. Equine Vet Educ 2006;18:96–9. 16. Miller CD. Optimizing the use of frozen-thawed equine semen. Theriogenology 2008;70:463–8. 17. Nielsen JM, Kofoed Bock TS, Ersbøll AK. Factors associated with fertility in horses in a Danish equine practice after artificial insemination with frozen-thawed semen. Anim Reprod Sci 2008;107:336–7. 18. Pace MM, Sullivan JJ. Effect of timing of insemination, numbers of spermatozoa and extender components on the pregnancy rate in mares inseminated with frozen stallion semen. J Reprod Fert 1975;23(Suppl.):115–21. 19. Leipold SD, Graham JK, Squires EL, McCue PM, Brinsko SP, Vanderwall DK. Effect of spermatozoa concentration and number on fertility of frozen equine semen. Theriogenology 1998;49:1537–43. 20. Lindsey AC, Bruemmer JE, Squires EL. Low dose insemination of mares using non-sorted and sex-sorted sperm. Anim Reprod Sci 2001;68:279–89. 21. Morris LHA, Allen WR. An overview of low dose insemination in the mare. Reprod Domest Anim 2002;37:206–10. ¨ 22. Guvenc K, Reilas T, Katila T. Effect of insemination dose and site on uterine inflammatory response of mares. Theriogenology 2005;63:2504–12. 23. Woods JA, Bergfelt DR, Ginther OJ. Effects of time of insemination relative to ovulation on pregnancy rate and embryonic loss rate in mares. Equine Vet J 1990;22: 410–15. 24. Squires EL, Barbacini S, Necchi D. Simplified strategy for insemination of mares with frozen semen. In: Proceedings of the 49th Annual Convention of the American Association of Equine Practitioners, 2003, pp. 353–6. 25. Reger HP, Bruemmer JE, Squires EL, Maclellan LJ, Barbacini S, Necchi D, Zavaglia G. Effects of timing and placement of cryopreserved semen on fertility of mares. Equine Vet Educ 2003;15:101–6. 26. Barbacini S. The effect of sperm number and frequency of insemination on pregnancy rates of mares inseminated with frozen-thawed spermatozoa. Paper read at 4th International Symposium on Stallion Reproduction, 21–23 October, 2005, Hannover, Germany. Animal Reproduction Science 2005;89(1–4):203–5. 27. Vidament M, Dupere AM, Julienne P, Palmer E. Equine frozen semen: freezability and fertility field results. Theriogenology 1997;48:907–17. 28. Samper JC. Management and fertility of mares bred with frozen semen. Anim Reprod Sci 2001;68:219–28. 29. Barbacini, S. Management of mares for frozen semen. In: Proceedings of the 14th International Congress on Animal Reproduction and Artificial Insemination, 2000, pp. 308, Elsevier. 30. Barbacini S. Artificial insemination (AI) with equine frozenthawed semen and the barren mare: problems, results and expectations. Paper read at 5th International Conference on Equine Reproductive Medicine: The Barren Mare, 24–25 November, 2007, Leipzig, Germany. 31. Dell’Aqua JA, Papa FO, Lopes MD, Alvarenga MA, Macedo LP, Melo CM. Modulation of acute uterine inflammatory response after artificial insemination with equine frozen semen. Anim Reprod Sci 2006;94:270–3. 32. Ginther OJ. Reproductive efficiency. In: Ginther O (ed.) Reproductive Biology of the Mare. Equiservices, 1992, pp. 499–558.
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Breeding with Frozen Semen 33. Katila T. Interaction of the uterus and semen. Pferdeheilkunde 1997;13:508–11. 34. Troedsson MHT. Therapeutic considerations for matinginduced endometritis. Pferdeheilkunde 1997;13:516–20. 35. Adams GP, Kastelic JP, Bergfetlt DR, Ginther OJ. Effect of uterine inflammation and ultrasonically detected fluid. J Reprod Fert 1987;35:445–54. 36. Pyckock JF, Newcombe JR. Assessment of the effect of three treatments to remove intrauterine fluid on pregnancy rates in mares. Vet Rec 1996;138:320–3. 37. Kotilainen T, Huhtinene M, Katila T. Sperm-induced leucocytosis in the equine uterus. Theriogenology 1994;41:629–36. 38. Zent WW, Troedsson MHT, Xue JL. Postbreeding uterine fluid accumulation in a normal population of Thoroughbred mares: a field study. In: Proceedings of the Annual Meeting Society for Theriogenology, 1998, pp. 78–9. 39. Watson ED, Barbacini S, Berrocal B, Sheerin O, Marchi V, Zavaglia G, Necchi D. Effect of insemination time of frozen semen on incidence of uterine fluid in mares. Theriogenology 2001;56:123–31. 40. Barbacini S, Necchi D, Zavaglia G, Squires EL. Retrospective study on the incidence of postinsemination uterine
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fluid in mares inseminated with frozen/thawed semen. J Equine Vet Sci 2003;23:493–6. Metcalf L. The effect of post-insemination endometritis on fertility of frozen stallion semen. In: Proceedings of the 38th Annual Convention of the American Association of Equine Practitioners, 2000, pp. 649–60. Vanderwall DK, Newcombe JR. Early embryonic loss. In: Samper, JC, Pycock, JF, McKinnon, AO (eds) Current Therapy in Equine Reproduction. Saunders Elsevier, 2007, pp. 374–83. Ball BA. Embryonic death in mares. In: McKinnon, AO, Voss, JL (eds) Equine Reproduction. Lea and Febiger, 1993, pp. 517–31. Jasko DJ, Moran DM, Farlin ME, Squires EL, Amann RP. Pregnancy rates utilizing fresh, cooled and frozen-thawed semen. In: Proceedings of the 46th Annual Convention of the American Association of Equine Practitioners, 1992, pp. 330–1. Barbacini S, Gulden P, Marchi V, Zavaglia G. Incidence of embryo loss in mares inseminated before or after ovulation. Equine Vet Educ 1999;11:251–4.
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Chapter 319
Techniques for Evaluating Frozen Semen Ben Colenbrander and Tom A.E. Stout
Introduction During the last 20 years, the number of mares inseminated worldwide with frozen-thawed stallion semen has increased to the point where frozen semen AI is considered a routine procedure within the sport-horse breeding industry. The increase in the popularity of frozen AI semen is largely due to significant improvements in the techniques for freezing stallion semen,1, 2 stricter selection of stallions with regard to suitability for semen cryopreservation, customizing the freezing protocol to optimize postthaw semen quality for individual stallions,3 and improvements in the management of inseminated mares.4, 5 As a result, pregnancy rates are both better and more consistent than reported previously; recent surveys have recorded mean pregnancy rates of 40–45% per cycle for frozen-thawed semen selected on the basis of a minimum of 35% progressively motile sperm post-thaw.6, 7 The increased demand for frozen-thawed semen has also been driven by the international character of the sport-horse breeding industry, and the logistical advantages of transporting frozen rather than chilled (liquid) semen over large distances. When liquid semen is transported between countries or continents there is, for example, little room for unexpected delays at customs, and extra pressure to ensure that mares become pregnant after a single insemination in order to prevent escalating transport costs. Frozen semen also enables more convenient commercial exploitation of competing stallions, from whom fresh or chilled semen may not be available on a regular or frequent basis, due to the demands of training or competition.
Disadvantages of frozen-thawed semen The current impediments to even more widespread use of frozen semen include the fact that not all stallions or ejaculates deliver sperm that maintain viability or fertilizing capacity well following cryopreservation.3 Moreover, the fertility of frozen-thawed semen is rarely as good as that with properly handled and managed fresh or chilled-transported liquid semen from the same stallion.8 In this latter respect, cryopreserving semen appears to affect stallion fertility negatively in two major ways. Firstly, freezing and thawing directly damages sperm cells leading either to immediate cell death, or to structural or functional damage that renders the sperm incapable of fertilization (e.g., acrosome damage9 ) or dramatically reduces their longevity in the female reproductive tract (e.g., ‘cryocapacitation’10 ). Secondly, it appears that high concentrations of the most commonly used cryoprotectant, glycerol (> 3.5%7 ), can have a contraceptive effect in the mare. A further obstacle to the widespread use of frozen stallion semen is the relatively high variability in ‘freezability’ between stallions,2 at least when compared to dairy cattle, in which post-thaw fertility is an aspect of the sire selection process.11 There is also variability between stallions in the response of the sperm to extender composition and cryoprotectant type and concentration.12 As a result, commercial stallion semen freezing centers now commonly customize the freezing protocol to individual stallions on the basis of initial tests to establish the most appropriate media and cryoprotectant composition; such customization allows the majority of normal stallions to fulfill the criteria for adequate freezability (94%3 ).
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Techniques for Evaluating Frozen Semen Perhaps the biggest obstacle to confidence in frozen semen technology is that ‘acceptable’ postthaw semen quality is no guarantee of fertility. This is in part because accepted international norms for the quality of frozen-thawed stallion semen are currently based almost exclusively on post-thaw progressive sperm motility (≥ 35% progressively motile sperm; ≥ 250 million progressively motile sperm13 ), whereas motility is not a precise indicator of fertility potential because it is only one of many parameters that a sperm must satisfy if it is to successfully reach, penetrate, and fertilize an oocyte in vivo.14 Surprisingly, other indices of semen quality, even simple ones like the percentage of morphologically normal sperm, are infrequently used to classify frozen semen in practice.
Predicting fertility At present, the only definitive way of proving that frozen semen from a given stallion retains fertilizing potential is to generate foals, or at least pregnancies. Indeed, in an ideal world all frozen semen would be accompanied by reliable fertility data so that purchasers had a realistic idea of what they were buying. In practice, the approximate fertility (good or bad) of popular stallions will usually become common knowledge albeit after a delay of months to years. But for less popular stallions or those for which semen has not previously been frozen, even anecdotal fertility data are usually absent or based on too few mares to be informative. Moreover, fertility data are necessarily retrospective, and therefore available only after a delay, and influenced by extrinsic factors such as semen handling, mare fertility, and the management of AI with respect to sperm number, the timing and number of inseminations per cycle, and the precise site of sperm deposition in the female reproductive tract. To improve confidence in the frozen semen trade it would, therefore, be extremely useful to be able to provide an unbiased indication of fertility potential using an array of objective semen quality tests tailored specifically to fertilizing capacity or damage likely to occur during freezing and thawing.
Evaluating the quality of frozen-thawed stallion sperm When evaluating frozen-thawed semen, it is important to perform the thawing procedure according to the guidelines provided by the supplier since the optimum thawing rate is generally related to the cooling technique employed: for example, cells that underwent fast freezing generally require fast thawing. Failure to adhere to the optimal thawing procedure,
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or holding the sperm for too long at a temperature above 38◦ C, is likely to affect adversely post-thaw quality. Sperm quantity The concentration or number of viable sperm in a straw will vary between stallions, ejaculates, and freezing protocols. Moreover, there appears to be a stallion-specific threshold below which sperm number per insemination dose will significantly influence fertility. Therefore, adequate evaluation of frozenthawed semen should include determination of the number of viable cells per straw or dose. Low numbers of sperm available may be a reason to consider adapting the insemination technique to deposit the sperm closer to the utero-tubal junction, either hysteroscopically or by per rectum guidance of a long flexible pipette; such ‘deep intrauterine insemination’ techniques can reduce required sperm numbers from the 250 million progressively motile sperm generally advised, to between 14 and 50 million.15 On a note of caution, the minimum sperm dose required for adequate fertility varies greatly between stallions and the fertility of some will drop alarmingly if any attempt is made to reduce the insemination dose. Sperm concentration is generally determined using either a hemocytometer/Neubauer-type counting chamber,16 a spectrophotometer calibrated for sperm concentration, or a Coulter counter. However, because semen extenders used for cryopreservation commonly contain components that interfere with the passage of light or are detected as particles (e.g., egg yolk and fat droplets), sperm concentration in frozen-thawed samples cannot be reliably measured by standard spectrophotometric or electronic techniques. Similarly, while computerassisted sperm analyzers (CASAs) can be used to measure sperm concentration, they may underestimate the true value because immotile sperm cluster after thawing, or conversely they may overestimate by wrongly classifying egg yolk particles as sperm. The accuracy and reliability of automated sperm concentration determination can be improved markedly by using a DNA stain that allows differentiation between cells and other particles, prior to analysis with either a CASA equipped with a fluorescence microscope, or an electronic counter with an integrated fluorescence microscope, such as the NucleoCounterTM (SP-100: Chemotec, Alleroed, Denmark). Although sperm concentration can also be measured repeatably and reliably using a flow cytometer, the equipment is too expensive for routine use unless concentration determination is combined with more in-depth analysis of sperm quality.17
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Sperm quality The percentage of (progressively) motile sperm, sometimes accompanied by a subjective estimation of sperm velocity (1 to 5; stationary to very rapid), is currently the most commonly used (often the only) parameter for assessing the quality of frozen-thawed semen. By-eye estimation of the percentage of motile sperm certainly provides useful information while being relatively straightforward; very poor motility is inconsistent with good fertility. However, accurate estimation of motility for frozen-thawed semen is complicated by the high concentrations at which the sperm is typically packaged and the presence of debris, fat, or egg-yolk droplets. Accurate and repeatable sperm motility estimation therefore requires dilution of the sample until individual cells can be tracked. Moreover, reliable by-eye estimation of the percentage of progressively motile sperm requires an experienced person working under optimal conditions with a good-quality phase-contrast microscope at a magnification of ×150–200, a heated stage prewarmed to 38◦ C, and a standardized system for making droplets of known size and concentration. The timespan between thawing and evaluation can also influence results because sperm require time to adjust to the change in osmotic environment after thawing; changes in sperm head volume during this period will temporarily depress motility. As a result, sperm motility (percentage and velocity) tends to improve during the first 5–10 minutes post-thaw. Subsequently, the percentage of motile sperm will usually decrease as cells damaged by the freeze-thawing process die. For these reasons, performing motility
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evaluation approximately 30 minutes after thawing is thought to be more informative with regards to likely fertility. Nevertheless, the percentage of neither motile nor progressively motile sperm is consistently or reliably correlated with fertility.9, 18
Computer-assisted sperm analysis For more reliable, unbiased, and repeatable analysis of sperm motility (e.g., for certification or research purposes), a computer-assisted semen analyzer (CASA) is recommended. CASA systems evaluate the movement of all particles identified as sperm on the basis of size, in a series of microscopic fields (Figure 319.1). As well as generating figures for the percentages of motile and progressively motile sperm, a CASA will calculate a variety of other parameters that describe more precisely how the sperm are moving; these include measurements of speed, such as curvilinear (VCL) and straight line (VSL) velocity, and of the quality of movement such as linearity (LN) and lateral head displacement (LHD). While the precise physiological significance of these extra indices for sperm movement is currently unclear for stallions, in other species it has been possible to classify the sperm in a sample into subpopulations on the basis of motility characteristics, where a higher percentage of sperm in a given subpopulation appears to correlate more closely with fertility than the percentage of motile sperm per se.19 The accuracy of CASA analysis is, however, dependent on the accuracy of the settings for identifying sperm and the algorithms for deciding whether a sperm is motile or progressively motile. Moreover,
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Figure 319.1 Typical examples of computer-assisted sperm analyzer (CASA) assessment of sperm motility for (a) a very poorquality sample and (b) a sample with good motility. The computer categorizes particles within a given size range as sperm, and analyzes and displays the sperm tracks. The tracks for sperm with different motility characteristics are represented on the screen using different colours (red = immotile; green = progressively motile; dark blue = non-progressively motile; turquoise = hypermotile). In addition, the CASA generates a readout in which the motion characteristics of all particles detected as sperm in a number of viewing fields are evaluated statistically. (See color plate 144).
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Techniques for Evaluating Frozen Semen the various CASA systems on the market use different settings and algorithms, and may therefore produce different results for the same sample. While the differences between machines are likely to be negligible in most instances, they could lead to different laboratories giving different advice about the suitability of a particular stallion or semen sample. As with by-eye measurements, the accuracy of CASA analysis also suffers at extremes of sperm concentration (low or high), and precise values will be affected by the time after thawing. Analysis may also be less accurate for some breeds or individuals because exact sperm head dimensions can vary sufficiently to result in failure to recognize them as sperm if the sperm detection settings are too narrow. Similarly, egg-yolk particles or debris in frozenthawed semen can inadvertently be misidentified as sperm. With this in mind, many systems now accommodate fluorescence optics so that DNA stains can be used to ensure that only nucleated cells of a certain size are identified as sperm. Indeed, if a membrane-impermeant fluorescent DNA dye is used (e.g., Hoechst 33342) a CASA system should be able to distinguish between viable and non-viable sperm. Sperm morphology is a classical semen quality parameter invariably included in a conventional breeding soundness examination. The overall percentage of morphologically normal sperm is considered an important aspect of semen quality because it has a moderate to high correlation with fertility.13, 20 The relationship is not, however, 100% and whereas a low percentage of morphologically normal sperm
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(< 30%) is a good predictor of subfertility, a high percentage of morphologically normal sperm is no guarantee of adequate or good fertility.14 Nevertheless, it is surprising that sperm morphology data are rarely provided when the quality of frozen-thawed semen is described. The reasons for this omission are unclear, but may include the fact that accurate morphological analysis requires both experience and a microscope with good optics (e.g., magnification of at least × 400, and preferably via an oil-immersion lens at × 1000). Moreover, morphological analysis is subject to considerable inter-operator variability and/or error. The other reasons why morphological analysis is largely neglected when assessing frozen-thawed semen may be that most stallions from which semen is frozen have previously demonstrated adequate morphological quality and/or fertility with fresh semen, while cryopreservation generally results in only a limited reduction in the percentage of morphologically normal sperm (around 7%21 ), with the possible exception of acrosomal damage, which is not always detected using conventional staining techniques. Morphological classification of stallion sperm also suffers from a lack of uniformity in that a range of stains and classification systems have been described. The most common techniques used are those that employ light microscopic evaluation of smears stained with, for example, nigrosin–eosin, aniline blue–eosin, Spermac, or Williams stains (Figure 319.2). Some of these stains include a membrane-impermeable component (e.g., eosin) that enables differentiation between viable and non-viable cells,22 albeit less reliably than with fluorescent viability dyes. However,
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Figure 319.2 Aniline blue–eosin-stained smears of stallion sperm showing live sperm with various acrosome defects. Panel (a) shows a large and (b) a smaller crater defect over the pre-equatorial (acrosomal cap) region, while (c) and (d) show ‘knobbed’ acrosomes. (See color plate 145).
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many laboratories prefer an alternative system for analyzing sperm morphology that involves examination of glutaraldhyde- or buffered formal saline-fixed sperm in wet mounts using either phase-contrast or, preferably, differential interference contrast microscopy.23 The analysis of wet mounts eliminates the possibility of introducing artifacts during smear preparation, and allows more detailed analysis of structural defects such as acrosome damage (a common consequence of freezing and thawing). How sperm morphological abnormalities should be recorded or classified is even more controversial, such that differences in the incidence and types of morphological abnormality recorded between laboratories could relate to no more than a different system of classification. While abnormalities are generally recorded by anatomical location, some laboratories record only the most proximal or most ‘severe’ (most likely to impinge on fertility) defect. Other systems exist to record whether defects probably arose during spermatogenesis (primary), epididymal maturation (secondary), or subsequently (tertiary), or whether they are likely to prevent a sperm reaching the oocyte (minor) or prevent fertilization or embryo development (major). The scientific data to support the various classification systems or to determine definitively which defects are ‘compensable’ by simply increasing sperm number in an insemination dose are, however, incomplete. In general, it is therefore thought that a record of all defects, although time-consuming, may give the clearest picture of sperm production or storage disturbances. Nevertheless, at present the percentage of morphologically normal sperm appears to be the most informative figure derived from morphological analysis. More detailed analysis of sperm morphology by, for example, scanning or transmission electron microscopy, reveals more subtle morphological changes resulting from the freeze-thawing process; however, such equipment is not readily available at studfarms and is only used for research purposes. Computer-assisted sperm head morphometry has been described for detecting sperm heads with an abnormal shape, and there are reports of subfertile stallions with significantly lower percentages of sperm heads that conform to the morphometric norm than matched fertile stallions.24 If morphometric analysis were to be considered for frozen-thawed stallion semen, the time after thawing when the smear is made would need to be standardized to eliminate any influence of the osmotic readjustment that occurs soon after thawing, while breed or individual differences in absolute sperm head dimensions would also need to be taken into account. The hypo-osmotic swelling test (HOS) evaluates whether the plasma membrane is not only intact but
Figure 319.3 Examples of stallion sperm undergoing the hypoosmotic swelling test. Some sperm have a normal-looking tail, indicating that they have failed to respond and have a damaged plasma membrane. Others show curling or thickening of the tail indicative of a positive response and an intact osmotically competent plasma membrane.
functional25 and, therefore, provides more information than a simple viability test. When exposed to a hypo-osmotic solution, functional sperm absorb water to establish osmotic equilibrium, which leads to a characteristic swelling of the tail that can be evaluated using bright-field microscopy at a magnification of ×400. Cells with swollen, thickened, bent, or curled tails are considered to be both intact and to have a functional membrane (Figure 319.3). The HOS test has proven to be user-friendly and repeatable, and has been validated for use on frozen-thawed stallion sperm.26, 27 Indeed, the HOS test may be an appropriate test for assessing semen quality after cryopreservation because membrane damage is probably an important factor limiting subsequent survival and function. Capacitation is the post-ejaculatory event during which a sperm acquires the ability to bind to the zona pellucida, undergo the acrosome reaction, and subsequently fuse with and fertilize an oocyte.28 Although capacitation is a complex and incompletely understood process, pivotal events include the influx of calcium, efflux of cholesterol, increase in plasma membrane fluidity, changes in plasma membrane phospholipid and protein distribution, phosphorylation of proteins, and changes in metabolic activity leading to hypermotility. The ability to capacitate is crucial to fertilizing potential; however, once capacitated a sperm has a limited lifespan and must therefore rapidly find and fertilize an oocyte. Freezing
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Techniques for Evaluating Frozen Semen and thawing sperm has been shown to result in changes similar to those induced during capacitation, and it is thought that these changes may be one of the reasons for the reduced lifespan of frozenthawed sperm compared to fresh sperm.10 However, it was recently reported that although cryopreservation induces ‘capacitation-like’ membrane changes, they probably occur via different pathways such that ‘cryocapacitation’ may interfere with rather than promote the subsequent ability to acrosome react, and thereby reduce fertilizing potential.29 One of the first assays developed for detecting the changes that occur during capacitation is staining with chlortetracycline (CTC), a fluorescent antibiotic that binds to the surface of sperm in a calciumdependent fashion. When CTC staining is combined with a membrane impermeable viability stain (e.g., Hoechst 33258) sperm can be differentiated into four major categories: live non-capacitated, live capacitated, live acrosome-reacted, and dead. Following freezing and thawing, a marked increase in the percentage of sperm classified as live capacitated is observed although, as indicated above, this shift in the CTC staining pattern may reflect destabilization of the sperm plasma membrane rather than true capacitation.30 In addition, CTC staining appears to recognize a fairly late and non-specific event in the capacitation process and has, therefore, largely been superseded by more specific fluorescent probes that, unlike CTC, are amenable to flow cytometric evaluation.31 The sperm acrosome reaction (AR) is an irreversible exocytotic event that, under physiological conditions, can only take place after capacitation and is induced by binding to the zona pellucida. During the AR, the sperm plasma membrane fuses with the outer acrosomal membrane at multiple sites over the anterior part of the sperm head resulting in the release of the acrosomal contents (mainly enzymes), which digest a hole through the zona pellucida to facilitate sperm penetration. Assessment of acrosome integrity, or the ability of sperm to undergo the acrosome reaction in vitro, are now recognized as important indices of semen quality.23 Microscopic techniques described for evaluating acrosome integrity include light microscopy of sperm stained with Spermac32 or a trypan blue–Giemsa combination13 that generates four categories of sperm, namely viable or non-viable combined with acrosome intact or reacted/damaged. However, the technique currently most widely used to evaluate acrosomal status is the binding of fluorescently labeled lectins that specifically recognize the outer acrosomal membrane or acrosome contents, such as fluorescein isothiocyanate conjugated Pisum sativum agglutinin33 or fluorescein isothiocyanate
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conjugated Arachis hypogea agglutinin;34 these stains bind only if the acrosome is damaged or ‘leaky’. The proportion of live sperm with a non-intact acrosome (reacted or damaged) appears to be an important indicator of semen quality when sperm damage is anticipated, for example, after freezing and thawing; sperm without an intact acrosome will have lost the ability to fertilize an oocyte. In addition, inability of acrosome-intact sperm to undergo the AR in response to either physiological (e.g., bicarbonate followed by progesterone) or non-physiological (e.g., calcium ionophore A23187) stimuli appears to be the probable cause of subfertility in at least some stallions for which other sperm quality analyses failed to reveal any significant abnormalities.23
Flow cytometry One of the most interesting breakthroughs in the assessment of semen quality is the development of fluorescent dyes that allow the assessment of various aspects of sperm quality (e.g., acrosome or DNA integrity) or function (e.g., ability to capacitate or acrosome react) flow cytometrically. Using a flow cytometer, multiple characteristics of large numbers (thousands) of live sperm can be assessed rapidly and objectively. Staining sperm with fluorescent probes requires a minimum of preparation because the flow cytometer specifically detects particle-associated fluorescence, such that removal of unbound probe is not required. Flow cytometry is therefore a very useful technical solution for rapid, reproducible assessment of sperm quality. To date, however, flow cytometric analysis has primarily been used when investigating possible causes of subfertility in cases where conventional measures of semen quality are normal, and most studies have focused on one or two specific stains rather than a spectrum; as a result there are currently few data on the value of combinations of flow cytometric staining techniques for predicting fertility potential (or subfertility). Viability stains Fluorescent DNA probes incapable of passing through intact cell membranes (‘membrane impermeant’) such as ethidium homodimer, diamidinophenylindole dihydrochloride (DAPI), propidium iodide (PI), and Hoechst 33258, can be used to label and discriminate sperm with damaged membranes (i.e., dead sperm35 ). To increase further the discriminative capacity of live–dead staining, a membraneimpermeant dye can be combined with a membranepermeant fluorescent probe such as SYBR-14, which will stain the DNA of all cells, be they membrane intact (live) or damaged (dead). When SYBR-14 is
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Figure 319.4 Stallion sperm stained with propidium iodide (red) and SYBR-14 (green) for estimation of the percentage of viable sperm. Viable sperm accumulate SYBR-14 and fluoresce intensely green, whereas non-viable sperm permit entry of propidium iodide and stain red or orange. Non-nucleated particles such as egg yolk or fat droplets will not take up either stain, thereby removing any risk of their being mistakenly identified as sperm during flow cytometric analysis. (See color plate 146).
used in combination with propidium iodide (PI) live sperm fluoresce green, while cells with damaged plasma membranes stain with both SYBR-14 and PI and, because PI fluorescence is more intense than that of SYBR-14, they fluoresce red or red-orange (Figure 319.4).36, 37
rescein isothiocyanate-Pisum sativum agglutinin) or FITC-PNA (Arachis hypogea agglutinin), sperm with an intact acrosome will not fluoresce, while cells with a ‘leaky’ acrosome (reacted or damaged) will admit the dye and exhibit green fluorescence (Figure 319.5). Flow cytometric analysis of acrosome integrity is nearly always combined with a viability stain, since only the acrosomal integrity of live sperm is relevant. In this respect, a relatively simple but informative triple-staining technique (SYBR-14/PI/PE-PNA) has been described for simultaneous flow cytometric discrimination of living sperm from plasma membranedamaged sperm cells (SYBR-14 in combination with PI37 ), and acrosome-intact from acrosome-reacting or damaged cells (using PE-PNA34 ). This is particulary useful for analyzing frozen-thawed semen immediately after thawing (i.e., without further washing or dilution) because the abundant egg-yolk particles can be ‘gated’ out during analysis; only particles labeled either with SYBR-14 (living cells) or PI (dead cells) are counted as sperm.38 One factor that can interfere with the accuracy of automated FITC-PNA or FITC-PSA detection of acrosome damage, is the protoplasmic droplet attached to the midpiece of mildly immature sperm; the droplet also stains positively for both FITC-PSA and FITC-PNA and samples containing high percentages of sperm with protoplasmic droplets (e.g., epididymal sperm) can therefore exhibit an artifactually high percentage of apparently acrosome-damaged or -reacted (fluorescent) sperm.
Acrosome integrity The acrosomal integrity of large numbers of sperm can also be evaluated objectively and rapidly using flow cytometry. When exposed to FITC-PSA (fluo-
(a)
(b)
Mitochondrial function Several fluorescent probes have been described for the evaluation of sperm mitochondrial function (e.g.,
(c)
(d)
Figure 319.5 FITC-PNA (fluorescein isothiocyanate-Pisum sativum agglutinin) staining of stallion sperm. Acrosome-intact sperm do not exhibit green fluorescence. Sperm in which the acrosome reaction has been initiated, or which have sustained damage to the acrosome, show staining over the whole acrosomal cap (a). Sperm that have completed the acrosome reaction show green fluorescence only along the equatorial band (b). Dead sperm take up the accompanying membrane-impermeant viability stain (e.g., ethidium homidimer) and fluoresce red (c). When analyzing acrosome integrity by flow cytometry, it should be borne in mind that the protoplasmic droplet also takes up FITC-PNA (d), which, in terms of total green fluorescence, could lead to immature sperm being confused with acrosome-reacted sperm. (See color plate 147).
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Techniques for Evaluating Frozen Semen rhodamine 123 and various MitoTracker probes), and there have been reports of an association between mitochondrial activity and sperm motility and velocity in stallion sperm.39, 40 Probably the most popular probe for assessing sperm mitochondrial function is JC-1, which has the useful property of changing color from green to red when bound to highly active mitochondria; the ratio of green to red fluorescence therefore allows sperm to be categorized by the level of mitochondrial activity. However, because glycolysis in the principal piece of the flagellum is now thought to be a more important source of the ATP needed to drive sperm motility than that produced by the mitochondria in the midpiece, the significance of the various degrees of mitochondrial function with regard to a sperm’s potential for motility is not clear. It is quite possible that staining for mitochondrial activity gives little more information than a viability stain. Capacitation As previously described, capacitation is an essential part of a sperm’s preparation to fertilize an oocyte, and is characterized by changes in lipid and protein distribution within the outer plasma membrane.41 Cryopreservation results in grossly similar rearrangements of the surface lipids and proteins, although these are probably not directly comparable since they appear to inhibit rather than promote the ability to acrosome react.29 Nevertheless, the proportion of live sperm showing signs of capacitation-like changes immediately after thawing may provide useful information with regard to decreased longevity, whereas the proportion that undergo capacitation in response to exposure to bicarbonate may reflect the retention of fertilizing capacity. The best described assays for the flow cytometric detection of capacitation in stallion sperm measure the increase in plasma membrane fluidity or disorder using probes that intercalate in a more fluid membrane, for example, merocyanin 540,31 or that recognize phospholipids only exposed in the outer phospholipid bilayer of capacitated sperm, such as annexin V;29 however, a number of laboratories have encountered difficulties in validating these assays for stallion sperm. A more repeatable measure of capacitation that can be assessed by either fluorescence microscopy or flow cytometry is the immunodetection of tyrosine phosphorylation of surface proteins.42 Capacitated sperm show an increase in tyrosine-phosphorylated proteins, and therefore fluorescence, particularly in the tail region (Figure 319.6). Nevertheless, further studies are required to determine how the capacitation-like changes induced by freeze-thawing relate to physiological capacitation, whether they are reversible, and what their
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Figure 319.6 Tyrosine phosphorylation staining to detect capacitation-related changes in stallion sperm. The green fluorescent staining in the tail of this sperm indicates that it has capacitated. The vital stain propidium iodide was also included in the staining procedure, and the absence of a red fluorescent nucleus demonstrates that the sperm is viable. (See color plate 148).
effects are on either sperm longevity or fertilizing capacity. Membrane lipid peroxidation (LPO) is thought to be one of the primary causes of defective sperm function resulting from cryopreservation.43, 44 It is thought that this LPO is induced by reactive oxygen species (ROS), and that sperm membranes are particularly susceptible to oxidative stress and peroxidative attack because of their relatively high content of polyunsaturated fatty acids. The occurrence of lipid peroxidation in frozen-thawed sperm can be detected using the LPO reporter probe C11-BODIPY 581/591, which fluoresces red in intact membranes but shifts to green following oxidative damage (Figure 319.7);44 this shift in fluorescence can be detected flow cytometrically. Cryopreservation of stallion sperm has been reported to increase both lipid peroxidation and the sensitivity to further peroxidation.44 However, it is not yet known how this vulnerability to lipid peroxidation varies between stallions, or how it correlates with fertility or loss of fertility. DNA integrity While the genetic integrity of a sperm may not affect its ability to fertilize an oocyte, it will certainly affect the developmental capacity of the resulting embryo. For this reason, assays have been developed and described for evaluating the integrity of sperm DNA. The most comprehensively described of these assays is the flow cytometrically analyzed sperm chromatin structure assay (SCSA). Other fluorescent assays for sperm DNA damage include the TUNEL assay, acridine orange staining, and the HALO test.46 The SCSA
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Figure 319.7 C11-BODIPY staining of stallion sperm to demonstrate lipid peroxidation of the type that can occur during freezing and thawing. Red staining indicates intact lipids (a) whereas green (b) indicates the occurrence of lipid peroxidation. Relative amounts of red and green can be analyzed by flow cytometry. Staining for lipid peroxidation may become a useful indicator of freezing-induced damage. (See color plate 149).
evaluates the susceptibility of sperm DNA to denaturation in an acidic environment by staining with acridine orange, a metochromatic dye that labels intact (double-stranded) DNA green and denatured (singlestranded) DNA red. The ratio of green to red fluorescence for each sperm can be assessed flow cytometrically and used to identify sperm with an increased susceptibility to DNA denaturation.47 One benefit of SCSA over other methods for evaluating DNA integrity is that the procedure relatively rapidly allows objective evaluation of many sperm. The percentage of sperm with an increased sensitivity to DNA denaturation is expressed as the COMP-alpha-t (ratio of green:red fluorescence of ‘cells outside the main population’) or DNA defragmentation index (DFI); stallions with a DFI exceeding 25% have been shown to exhibit reduced fertility.48 At present, it is not thought that freezing-thawing leads to a dramatic increase in sperm DNA damage.
Testing for infectious agents International trade in frozen stallion semen is inescapably linked with the potential for transmitting infectious agents from one country or horse population to another.49 Indeed, several infectious agents, notably equine arteritis virus, survive conventional semen cryopreservation protocols. As a result, international trade in stallion semen is subject to strict import-export regulations with tests performed on
the stallion and/or its semen prior to departure, and in some cases after arrival at the destination, to ensure that the semen is free of specified organisms (e.g., Taylorella equigenitalia and equine arteritis virus); the exact testing requirements are usually imposed by the importing country. Inspection of the papers and certificates concerning disease status of the semen donor and semen is therefore an important part of determining whether a semen dose is fit for use. Of course, false-negative assay results, or new emerging diseases for which transmission via semen has yet to be reported, could still result in the unexpected presence of infectious agents in semen that subsequently leads to a disease outbreak.
Conclusions In recent years, our knowledge of the complex array of attributes that a sperm must possess if it is successfully to reach and fertilize an oocyte and give rise to a viable embryo has increased appreciably, and a number of tests for examining more detailed aspects of sperm quality have been developed.8, 14 In this light, it is clear that classifying the quality of frozen-thawed semen by the percentage and number of motile sperm alone is no more than a means of ruling out semen doses that clearly do not offer an acceptable chance of establishing pregnancy; however, classifying semen as ‘acceptable’
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Techniques for Evaluating Frozen Semen using such a limited range of parameters is debatable since it offers no guarantee of adequate fertility. Nevertheless, even such simple means of selecting ejaculates have led to improved mean pregnancy rates, and increased veterinarian and breeder confidence in frozen semen. Additional tests for aspects of sperm quality not currently assessed (e.g., percentages of morphologically normal sperm, sperm with intact DNA, sperm with intact acrosomes) would undoubtedly enable identification of a higher proportion of stallions/ejaculates not likely to yield pregnancies, and lead to further improvements in results. While there are currently too few data on the correlations between the various tests or combinations of tests available and in-field fertility, it is possible that when this information is available we will be able to settle on a semen assessment panel that offers a fairly accurate prognostic indication of fertility potential.18, 50 In the meantime, it may be prudent to concentrate on tests that examine sperm compartments likely to be affected by cryopreservation. For example, because the proportion of sperm that pass through columns designed to remove acrosome-damaged sperm (e.g., Sephadex or glass-wool Sephadex9 ) correlates fairly well with in-field fertility, a sensible first step would be to add a test for the percentage of viable acrosomeintact sperm, and establish thresholds at which fertility is compromised. In the longer term, confidence in frozen semen could be increased further by introducing automated systems to objectively, rapidly, and accurately analyze concentration (i.e., NucleoCounter; CASA with fluorescent stains; flow cytometry) and motility (CASA) of frozen-thawed semen with the addition, when justified by initial tests, of some or all of the stains now available for assessing viability, capacitation status, acrosome status, DNA stability, and mitochondrial function. Since the latter can all be assessed flow cytometrically, a range of tests would be established that could be assessed rapidly and objectively. Moreover, since the semen is frozen, the tests could be performed at a remote laboratory; in this way stallion semen freezing centers would not need to commit to the capital outlay until convinced of the benefits of the new technology.
References 1. Alvarenga MA, Papa FO, Landim-Alvarenga FC, Medeiros AS. Amides as cryoprotectants for freezing stallion semen: a review. Anim Reprod Sci 2005;89:105–13. 2. Vidament M. French field results (1985–2005) on factors affecting fertility of frozen stallion semen. Anim Reprod Sci 2005;89:115–36. 3. Loomis PR, Graham JK. Commercial semen freezing: individual male variation in cryosurvival and the response of stallion sperm to customized freezing protocols. Anim Reprod Sci 2008;105:119–28.
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4. Samper JC. Management and fertility of mares bred with frozen semen. Anim Reprod Sci 2001;68:219–28. 5. Sanchez R, Gomez I, Samper JC. Artificial insemination with frozen semen. In: Samper JC (ed.) Equine Breeding, Management and Artificial Insemination. Saunders-Elsevier, 2009; pp. 175–83. 6. Sieme H, Schafer T, Stout TA, Klug E, Waberski D. The effects of different insemination regimes on fertility in mares. Theriogenology 2003;60:1153–64. 7. Vidament M, Vincent P, Yvon JM, Bruneau B, Martin FX. Glycerol in semen extender is a limiting factor in the fertility in asine and equine species. Anim Reprod Sci 2005;89:302–5. 8. Graham JK, Moce E. Fertility evaluation of frozen/thawed semen. Theriogenology 2005;64:492–504. 9. Samper JC, Hellander JC, Crabo BG. Relationship between the fertility of fresh and frozen stallion semen and semen quality. J Reprod Fertil Suppl 1991;44:107–14. 10. Pommer AC, Rutllant J, Meyers SA. Phosphorylation of protein tyrosine residues in fresh and cryopreserved stallion spermatozoa under capacitating conditions. Biol Reprod 2003;68:1208–14. 11. Hoflack G, Van Soom A, Maes D, de Kruif A, Opsomer G, Duchateau L. Breeding soundness and libido examination of Belgian Blue and Holstein Friesian artificial insemination bulls in Belgium and The Netherlands. Theriogenology 2006;66:207–16. 12. Amann R, Pickett B. Principles of cryopreservation and a review of stallion spermatozoa. J Equine Vet Sci 1987;7:145–73. 13. Sieme H. Semen evaluation. In: Samper J (ed.) Equine Breeding Management and Artificial Insemination. SaundersElsevier, 2009; pp. 57–74. 14. Colenbrander B, Gadella BM, Stout TA. The predictive value of semen analysis in the evaluation of stallion fertility. Reprod Domest Anim 2003;38:305–11. 15. Morris LH, Tiplady C, Allen WR. Pregnancy rates in mares after a single fixed time hysteroscopic insemination of low numbers of frozen-thawed spermatozoa onto the uterotubal junction. Equine Vet J 2003;35:197–201. 16. Bailey E, Fenning N, Chamberlain S, Devlin L, Hopkisson J, Tomlinson M. Validation of sperm counting methods using limits of agreement. J Androl 2007;28:364–73. 17. Hansen C, Vermeiden T, Vermeiden JP, Simmet C, Day BC, Feitsma H. Comparison of FACSCount AF system, Improved Neubauer hemocytometer, Corning 254 photometer, SpermVision, UltiMate and NucleoCounter SP-100 for determination of sperm concentration of boar semen. Theriogenology 2006;66:2188–94. 18. Kirk ES, Squires EL, Graham JK. Comparison of in vitro laboratory analyses with the fertility of cryopreserved stallion spermatozoa. Theriogenology 2005;64:1422–39. 19. Amann RP, Katz DF. Reflections on CASA after 25 years. J Androl 2004;25:317–25. 20. Jasko DJ, Lein DH, Foote RH. Determination of the relationship between sperm morphologic classifications and fertility in stallions: 66 cases (1987–1988). J Am Vet Med Assoc 1990;197:389–94. 21. Parlevliet J, Malmgren L, Boyle M, Wockener A, Bader H, Colenbrander B. Influence of conservation method on the motility and morphology of stallion semen (an international project). Acta Vet Scand Suppl 1992;88:153–62. 22. Colenbrander B, Puyk H, Zandee AR, Parlevliet J. Evaluation of the stallion for breeding. Acta Vet Scand Suppl 1992;88:29–37.
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23. Varner DD. Developments in stallion semen evaluation. Theriogenology 2008;70:448–62. 24. Casey PJ, Gravance CG, Davis RO, Chabot DD, Liu IK. Morphometric differences in sperm head dimensions of fertile and subfertile stallions. Theriogenology 1997;47:575–82. 25. Jeyendran RS, Van der Ven HH, Perez-Pelaez M, Crabo BG, Zaneveld LJ. Development of an assay to assess the functional integrity of the human sperm membrane and its relationship to other semen characteristics. J Reprod Fertil 1984;70:219–28. 26. Neild D, Chaves G, Flores M, Mora N, Beconi M, Aguero A. Hypoosmotic test in equine spermatozoa. Theriogenology 1999;51:721–7. 27. Nie GJ, Wenzel JG. Adaptation of the hypoosmotic swelling test to assess functional integrity of stallion spermatozoal plasma membranes. Theriogenology 2001;55:1005–18. 28. Gadella BM. Sperm membrane physiology and relevance for fertilization. Anim Reprod Sci 2008;107:229–36. 29. Thomas AD, Meyers SA, Ball BA. Capacitation-like changes in equine spermatozoa following cryopreservation. Theriogenology 2006;65:1531–50. 30. Neild DM, Gadella BM, Chaves MG, Miragaya MH, Colenbrander B, Aguero A. Membrane changes during different stages of a freeze-thaw protocol for equine semen cryopreservation. Theriogenology 2003;59:1693–705. 31. Rathi R, Colenbrander B, Bevers MM, Gadella BM. Evaluation of in vitro capacitation of stallion spermatozoa. Biol Reprod 2001;65:462–70. 32. Oettle EE. Using a new acrosome stain to evaluate sperm morphology. Vet Med 1986;81:263–7. 33. Farlin ME, Jasko DJ, Graham JK, Squires EL. Assessment of Pisum sativum agglutinin in identifying acrosomal damage in stallion spermatozoa. Mol Reprod Dev 1992;32:23–7. 34. Cheng FP, Fazeli A, Voorhout WF, Marks A, Bevers MM, Colenbrander B. Use of peanut agglutinin to assess the acrosomal status and the zona pellucida-induced acrosome reaction in stallion spermatozoa. J Androl 1996;17:674–82. 35. Gadella BM, Rathi R, Brouwers JF, Stout TA, Colenbrander B. Capacitation and the acrosome reaction in equine sperm. Anim Reprod Sci 2001;68:249–65. 36. Garner DL, Johnson LA, Yue ST, Roth BL, Haugland RP. Dual DNA staining assessment of bovine sperm viability using SYBR-14 and propidium iodide. J Androl 1994;15:620–9. 37. Garner DL, Johnson LA. Viability assessment of mammalian sperm using SYBR-14 and propidium iodide. Biol Reprod 1995;53:276–84.
38. Nagy S, Jansen J, Topper EK, Gadella BM. A triple-stain flow cytometric method to assess plasma- and acrosomemembrane integrity of cryopreserved bovine sperm immediately after thawing in presence of egg-yolk particles. Biol Reprod 2003;68:1828–35. 39. Auger J, Ronot X, Dadoune JP. Human sperm mitochondrial function related to motility: a flow and image cytometric assessment. J Androl 1989;10:439–48. 40. Garner DL, Thomas CA, Joerg HW, DeJarnette JM, Marshall CE. Fluorometric assessments of mitochondrial function and viability in cryopreserved bovine spermatozoa. Biol Reprod 1997;57:1401–6. 41. Gadella BM, Tsai PS, Boerke A, Brewis IA. Sperm head membrane reorganisation during capacitation. Int J Dev Biol 2008;52:473–80. 42. Piehler E, Petrunkina AM, Ekhlasi-Hundrieser M, TopferPetersen E. Dynamic quantification of the tyrosine phosphorylation of the sperm surface proteins during capacitation. Cytometry A 2006;69:1062–70. 43. Brouwers JF, Gadella BM. In situ detection and localization of lipid peroxidation in individual bovine sperm cells. Free Radic Biol Med 2003;35:1382–91. 44. Brouwers JF, Silva PF, Gadella BM. New assays for detection and localization of endogenous lipid peroxidation products in living boar sperm after BTS dilution or after freeze-thawing. Theriogenology 2005;63:458– 69. 45. Neild DM, Brouwers JF, Colenbrander B, Aguero A, Gadella BM. Lipid peroxide formation in relation to membrane stability of fresh and frozen thawed stallion spermatozoa. Mol Reprod Dev 2005;72:230–8. 46. Evenson DP, Kasperson K, Wixon RL. Analysis of sperm DNA fragmentation using flow cytometry and other techniques. Soc Reprod Fertil Suppl 2007;65:93–113. 47. Evenson DP, Wixon R. Clinical aspects of sperm DNA fragmentation detection and male infertility. Theriogenology 2006;65:979–91. 48. Love CC. The sperm chromatin structure assay: a review of clinical applications. Anim Reprod Sci 2005;89:39–45. 49. Metcalf ES. The role of international transport of equine semen on disease transmission. Anim Reprod Sci 2001;68:229–37. 50. Wilhelm KM, Graham JK, Squires EL. Comparison of the fertility of cryopreserved stallion spermatozoa with sperm motion analyses, flow cytometric evaluation, and zona-free hamster oocyte penetration. Theriogenology 1996;46:559– 78.
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Chapter 320
Storage Management and Distribution of Frozen Semen Paul R. Loomis
In 1949, Polge and co-workers reported that spermatozoa suspended in a buffer containing glucose and glycerol could survive freezing to – 79◦ C and exhibit motility upon thawing. It wasn’t until 1957 that the first pregnancy in a mare inseminated with frozen stallion semen was reported. Since then, improved cryopreservation and insemination protocols for stallion semen, as well as breed registry acceptance, have led to a wider utilization of frozen semen in the equine breeding industry. One of the main advantages of frozen semen is that when properly maintained at liquid nitrogen temperature (– 197◦ C), spermatozoa can remain viable for an indefinite time, provided the spermatozoa respond favorably to the freezing process. Bovine semen stored frozen for more than 40 years has been reported to achieve pregnancies, and personnel in our laboratory have used equine semen frozen for more than 22 years with good results. In order to maintain maximum fertilizing potential, spermatozoa must not be exposed to fluctuations in temperature and must be maintained below the critical temperature (approximately – 130◦ C) at which thermally driven chemical reactions proceed. Exposure of cells to temperatures above – 80◦ C can initiate some degree of thawing and then recrystallization of ice, as well as reorganization of membrane proteins and lipids. This can lead to cell damage and impaired fertilizing potential. To maintain cells at these ultra-low temperatures, packaged frozen semen is typically immersed in liquid nitrogen. Specialized insulated cryogenic containers have been developed that allow for such storage.
Cryogenic storage containers Cryogenic storage containers commonly used in the artificial insemination (AI) industry can be grouped
into two types: large-capacity bulk containers made of insulated stainless steel, or smaller aluminum containers. Both types of containers function by providing a chamber into which liquid nitrogen is loaded and the packaged semen is immersed. The inner chamber is wrapped with a special insulating foil and placed inside a second chamber. The two chambers are welded together and superior insulating properties are achieved by applying a vacuum to evacuate the air in the space between the two chambers. In liquid form nitrogen is extremely cold (– 197◦ C/– 320◦ F) and, when left in an open container, evaporates quickly into nitrogen gas. Cryogenic containers are designed to minimize the rate at which liquid nitrogen is converted to gas and allow for ultra-low refrigeration of the samples without electrical requirements. Stainless-steel bulk containers can accommodate a variety of storage racks or box configurations to suit the material being stored. The advantages of this type of container are storage capacity and configuration options whereas the major disadvantage is nitrogen consumption. Figure 320.1 shows a large-capacity bulk storage container with a single-level square canister racking system holding equine frozen semen. With respect to the artificial insemination (AI) industry, such bulk containers are commonly used in large storage facilities where capacity is the primary determining factor. Much more common in the equine frozen semen industry are the smaller, insulated aluminum containers. These containers are manufactured by several companies and can be purchased in a variety of sizes to accommodate specific requirements. Figure 320.2 is a cutaway view of one of these containers. The inner chamber is constructed of aluminum, which is then wrapped with an insulating foil. A second aluminum chamber surrounds the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Figure 320.1 (a) Exterior and (b) interior views of a large-capacity bulk storage container with a single-level square canister racking system holding equine frozen semen.
inner chamber and foil, and the two chambers are welded together at the neck of the container and vacuum sealed. This construction allows temperatures within all parts of the container to remain lower than – 180◦ C by circulating nitrogen gas within the chamber even when the level of liquid drops to a few centimeters. Unlike the square canisters used in the bulk container, the aluminum containers have round canisters with handles that hang down into the chamber from the neck of the container. There are three basic elements to consider when choosing a container from the wide variety of options available. The first is the type of package in which the semen has been frozen. Most equine semen is currently being packaged in 0.5-mL straws for freezing. These straws can be stored in small plastic goblets (10 × 120 mm or 12 × 120 mm) that can hold up to 7 or 11 straws respectively. The goblets can then be placed on 28-cm long aluminum canes (two goblets per cane) and lowered into the canisters for storage. The tops of the canes can also be labeled to aid in identification of the semen. Alternatively, 0.5-mL straws can be loaded into larger goblets for bulk storage. When straws are stored in bulk using goblets of larger diameter, the goblets are usually stored on two levels per canister. Most storage containers have canisters that are 28 cm long, which can accommodate 0.5-mL straws stored either on canes or in bulk goblets on two levels. Some equine semen is still packaged in 4- or 5-mL straws (also known as macrotubes or maxi straws), and quite a lot of semen frozen prior to the mid-1990s was packaged this way. The length of these straws (24 cm for 4.0 mL; 28 cm for 5.0 mL) requires that the canister length be a minimum of 28 cm. A few models of liquid storage and vapor shipping containers have shorter canisters (12.7 cm) that can only accommodate a single layer of
0.5-mL straws. Figure 320.3 shows a variety of straw, goblet, and cane configurations. The second element to consider is the amount of semen storage capacity required. Obviously, the larger the container the greater is the capacity. However, a third element to consider is the availability of liquid nitrogen. The rate at which a cryogenic container uses liquid nitrogen (referred to as the static
Figure 320.2 Cutaway view of a standard double-walled insulated aluminum liquid nitrogen storage container.
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Figure 320.3 A variety of straw, goblet, and cane configurations used for storing equine frozen semen.
evaporation rate) is dependent upon the amount of liquid nitrogen capacity the container has, the insulating properties of the container, the size of the neck opening, and the frequency with which the container is opened. Because liquid nitrogen is constantly being converted to gas, nitrogen containers cannot be completely sealed otherwise the increased volume occupied by nitrogen gas as opposed to liquid will cause a rapid build-up of pressure within the container and could lead to an explosion. Therefore, all nitrogen storage containers must be allowed to vent to the outside of the container. The high-density foam neck corks used to close off the containers allow for nitrogen gas to escape constantly, preventing a build-up of pressure. The rate of nitrogen gas escaping is determined in part by the diameter of the neck opening, i.e., the smaller the neck opening, the lower the rate of liquid nitrogen consumption. A narrow neck opening in turn limits the maximum diameter of the canister, which limits the semen storage capacity. The rate of nitrogen consumption is also influenced by the frequency with which the neck cork is removed to check liquid levels, place new semen into storage, or remove semen for shipment or AI. Most manufacturers will provide an estimate of the ‘normal working duration’, which takes into account a reduction in holding time based on an estimate of ‘normal use’ for the various models of containers they sell. The shorter the normal working duration for a container, the more frequently it must be refilled with liquid nitrogen. A container that holds a large amount of liquid nitrogen and has a narrow neck opening will have a lower capacity for semen storage but longer working duration than does a container of similar size
and nitrogen capacity with a larger diameter neck opening. The following are recommended guidelines for the safe storage of frozen semen in liquid nitrogen containers. 1. Aluminum nitrogen containers should not be stored directly on concrete floors as this will erode the aluminum. 2. Place storage containers in a well-ventilated space, because escaping nitrogen gas will displace oxygen in a confined space and breathing air that is less than 18% oxygen could lead to a lifethreatening situation. 3. Place storage containers in a room that allows for frequent visual inspection. A container that loses vacuum suddenly may exhibit a frosting around the neck of the container. If observed quickly enough it may be possible to move the semen to another container before all of the nitrogen is lost. 4. Protect containers from impact as sudden impact could break the seal between the two chambers of the container and lead to a loss of vacuum and rapid evaporation of nitrogen within the container. 5. Check liquid nitrogen levels (as shown in Figure 320.4) with a ruled stick at regular intervals and record the level. In our lab, nitrogen levels are checked weekly and nitrogen consumption recorded. A container that is starting to lose vacuum will be less efficient and increase the rate of nitrogen consumption. Detection prior to complete vacuum failure allows for the semen to be safely moved to another container. 6. Use caution when topping off containers with liquid nitrogen because the plastic straws at this
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Figure 320.4 A technician uses a wooden ruled dipstick to measure the level of liquid nitrogen in a storage container.
temperature are extremely brittle and can easily crack causing damage that may not be revealed until the straw is thawed.
Sources of liquid nitrogen Refilling storage containers requires a readily available supply of liquid nitrogen. There are a variety of options, and the optimum choice for any given storage facility depends upon the quantity of nitrogen required and the availability and cost from a liquid nitrogen supplier. The largest storage facilities will likely choose a bulk tank solution and enter into a bulk product agreement with one of the several gas companies that supply liquid nitrogen. These stationary bulk tanks are installed on site and can hold from approximately 2000 to over 50 000 liters of liquid nitrogen kept at a pressure of 250 psi. The bulk tank can be connected with insulated piping to filling stations inside the storeroom to provide a ready supply of liquid nitrogen for refilling storage and shipping containers. When the tanks reach a predetermined refill level, a large-capacity nitrogen transport vessel on a tractor trailer is dispatched to the site and the bulk tank is refilled. Figure 320.5 shows a photograph of a 11,350 liter bulk tank installed outside a large equine semen storage facility. For smaller storage facilities, there are two alternative options commonly employed. Most gas companies will rent out portable pressurized nitrogen transfer containers ranging in size from 160 to 240 liters. These containers filled with liquid nitrogen can be delivered to the storage facility, and with an appropriate transfer hose can be used to safely refill storage and shipping containers (Figure 320.6).
Figure 320.5 A 11,350 liter bulk nitrogen tank installed outside a large equine semen storage facility.
When empty, the gas company can bring a full container and exchange it for the empty one. This system insures that a supply of nitrogen will always be on hand when needed. The final option is to have containers filled by a service that comes to the storage facility with a portable, pressurized nitrogen bulk tank. These services will set up a schedule to come to the facility at regular intervals appropriate for the type of containers used. Some of the large bovine AI companies offer such a service, and this is the method most commonly used by dairymen who routinely store semen in one to three small to medium-sized storage containers at the farm.
Safety considerations when working with liquid nitrogen It is essential that all personnel working with liquid nitrogen are aware of two critical facts concerning this substance: 1. Liquid nitrogen is extremely cold. At atmospheric pressure, liquid nitrogen boils at – 196◦ C. 2. Liquid nitrogen produces a large amount of nitrogen gas. One liter of liquid nitrogen vaporizes into almost 25 cubic feet (0.71 m3 ) of nitrogen gas. Either of these two properties can produce personal injury or property damage. Use caution when handling liquid nitrogen and follow these guidelines to prevent injury or damage:
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Figure 320.6 A technician refilling a liquid nitrogen storage container from a low-pressure bulk source.
1. Prevent liquid nitrogen from contacting bare skin as it can cause severe frostbite almost instantly. 2. Do not allow objects cooled by liquid nitrogen to touch your bare skin as such objects may stick to your skin and tear the flesh away when you attempt to remove the object. 3. Always wear heavy loose-fitting gloves while handling objects that have been in contact with liquid nitrogen. Special lightweight, non-absorbent ‘Cryo-Gloves’ that can be purchased from nitrogen container suppliers, or heavy leather gloves are recommended. Boots should be worn with pants outside to prevent spilled liquid nitrogen from entering footwear. 4. Always wear protective eyewear to prevent splashed liquid from contacting the eyes. 5. Always dispose of liquid nitrogen outdoors by slowly pouring onto the ground and allowing the nitrogen to vaporize into the atmosphere. 6. Do not seal containers tightly as pressure build-up from the expanding gas may lead to an explosion if the gas is not allowed to vent. 7. Always use a solid wooden, metal, or plastic dipstick to measure liquid levels. Never use a hollow rod or tube as the gasification and expansion of the rapidly cooling liquid inside the tube will force liquid to spurt from the top of the tube.
Inventory management systems Accurate inventory management is critical for any storage facility or veterinary clinic holding frozen semen for clients. Small inventories can be easily tracked using a manual paper, index card, or
spreadsheet-based system. For larger storage facilities that have a lot of inventory being added and removed for distribution, a computerized database system is useful. The key to any system is a strict policy of recording transactions that accounts for each individual straw of semen in storage. This requires that all straws entered into storage are counted accurately and that all data pertaining to those straws be entered as well. Straws should be labeled permanently with the stallion’s name, registration number, freezing facility code or name, and date of freezing or coded lot number. It is highly recommended that preprinted straws be used rather than hand labeling as the handwriting on straws is often illegible and can lead to confusion concerning the identity of the semen. All of this information as well as the semen owner, sperm concentration in the straws, number of recommended straws per insemination dose, postthaw quality, and exact location (container, canister number, and section within the canister) for each individual lot should be included with the inventory. For smaller inventories a simple system using index cards has been employed by many storage facilities. For each ejaculate frozen (or for any individual doses shipped in for storage) an index card is created that contains all the information mentioned above on the top of the card. As individual straws are removed from that lot of semen, the transaction details are listed on a line of the index card and should include the number of straws removed, the date, and the purpose for the transaction (shipment, evaluation, AI, etc.). A running account of the number of straws remaining from each lot can be kept on each index card and reported to the owner on a regular basis.
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Frozen semen distribution Frozen semen is typically transported in a nitrogen vapor-phase container (dry shipper). These cryogenic containers maintain near liquid nitrogen temperatures (typically – 180 to – 195◦ C) for days or weeks without the use of hazardous liquid nitrogen. Vapor-phase containers work by absorbing liquid nitrogen into a thick layer of hydrophobic absorbent material that surrounds the inner cavity of the container where the semen is stored. A long holding time is achieved through the superior insulation afforded by the double-walled aluminum shell, which is filled with insulating foil and vacuum sealed. Vapor-phase containers must be properly ‘charged’ by filling with liquid nitrogen to the point of saturation of the absorbent material and then pouring off the excess liquid nitrogen. Such vapor-phase shippers can be transported with a ‘non-hazardous’ classification throughout the world, which significantly reduces transport costs. A word of caution is offered, however, concerning the use of vapor shippers; because the package is being shipped with a nonhazardous classification, it is critical that all of the liquid nitrogen is poured out of the container prior to
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transport. Transport of a hazardous material without the proper declarations and appropriate labeling can lead to significant fines as this is a violation of IATA (International Air Transport Association) regulations and may be in breach of certain state and federal laws. Vapor shippers can be used to store semen for longer than the typical recommended holding time if more liquid nitrogen is added. If a practitioner receives semen in a vapor shipper in anticipation of inseminating a mare and she does not develop a preovulatory follicle as anticipated or she ovulates before the semen arrives, the vapor container may be used to store the semen for an extended time provided the vapor container is filled again with liquid nitrogen. If this occurs, the practitioner should contact the semen supplier as soon as possible to discuss the shipper’s policy regarding tank returns. Vapor shippers are fairly durable; however, it is highly recommended that they be placed inside a solid outer shipping carton that will protect the container against physical damage or loss of vacuum due to rough handling during shipping. Figure 320.7a shows a vapor-phase shipper with 3-week holding time. Figure 320.7b shows one type of shipping carton that has a round top and is base wide
(b)
Figure 320.7 (a) Example of a vapor-phase ‘dry shipper’ commonly used to transport equine frozen semen. (b) Protective shipping carton into which the dry shipper is placed for secure transport.
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Storage Management and Distribution of Frozen Semen to help keep the shipper upright and prevent the package from being transported upside down or on its side. Even though it is a vapor shipper, if left on its side or upside down, the nitrogen gas will ‘pour’ out of the container and significantly reduce holding time. Eventually, vapor shippers will lose vacuum resulting in a loss of holding time. Prior to the beginning of each breeding season it is highly recommended that all shippers be properly tested to determine if they are holding temperature according to specifications. Vapor shippers and liquid storage containers that have lost all or partial vacuum can often be repaired by the manufacturer. Occasionally, semen is shipped in liquid shipping containers. These containers maintain semen for a longer period of time provided an adequate liquid nitrogen level is maintained. There is no absorbent material in these containers therefore if they are tipped and the liquid spills out, the container will quickly warm to room temperature and the semen will thaw. When shipping in a non-vapor, liquid container, the package must comply with IATA regulations with regard to the type of packaging, the declarations on the shipping documents, and appropriate labeling of the package. Furthermore, the individual who packs the container and signs the hazardous materials declarations must have undergone specific training in the use of the hazardous material in question. One of the main benefits of using frozen semen is the fact that it can be transported to the veterinarian or mare owner in advance thus eliminating the anxiety and logistical headaches associated with cooled semen breeding. Ideally, the semen can be ordered prior to the mare’s estrus and transferred upon arrival to a storage container at the farm or clinic. All too often, practitioners trapped in the ‘cooledsemen’ mindset order frozen semen the day before the insemination is planned, resulting in the same scheduling problems seen for cooled semen. Even if long-term storage is not available, ordering semen on the first day of estrus and keeping it in the shipping tank until needed is better than waiting until the last minute and missing the insemination opportunity because the mare ovulated before the semen arrived. When transferring frozen semen, the technician should take care not to expose the straws to room temperature by lifting them into the warm neck of the tank or transferring across any significant distance between the shipper and permanent storage. Proper transfer of 0.5-mL straws from a vapor shipper to permanent storage is illustrated in Figure 320.8. Lift the canister slightly into the neck of the shipper and then using pre-cooled hemostats or tweezers gently grasp the individual straws, canes, or the
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plastic goblet and lower the canister back into the container (Figure 320.8a). Never lift the canister or straws above the frost line (visible about 5 cm [2 in] from the top of the container neck) until it is time to transfer the semen. While holding the semen well inside the shipper, lift the canister from the storage container slightly up into the neck, again never above the frost line (Figure 320.8b). As quickly as possible, transfer the straws into the liquid storage canister and lower back down into the liquid nitrogen (Figure 320.8c,d). When packaging frozen semen for shipment, extreme care should be taken to avoid exposing straws to elevated temperatures. Use of an accurate and convenient inventory system allows the technician quickly to locate the correct straws to be loaded without having to search through canisters, trying to identify the appropriate straws. If large 5- or 4-mL straws are to be shipped they are simply loaded into the canister of a fully charged vapor shipper, taking care always to work within the neck of the storage container below the frost line as previously mentioned. Lower the frozen straws gently into the canister to prevent cracking and pack the empty space around the straws with cotton to keep them from moving inside the canister during shipment. For semen packaged in 0.5-mL straws there are two options. Straws can be loaded into small goblets and placed on canes or they can be loaded into larger plastic goblets and shipped in bulk. If straws are stored in bulk then shipping on canes requires that the individual straws are loaded into the smaller goblets and clipped into the canes while working under liquid nitrogen. This can be accomplished with a small thick-walled polyfoam box filled with liquid nitrogen, under which the straws, goblets, and canes are handled. The technician can easily verify the identity of the semen by reading the printed straws while they are under the liquid nitrogen. Printed straws held in nitrogen vapor often become frosted making them difficult to read. Straws and goblets should be handled with pre-cooled tweezers or hemostats. Likewise, when shipping in bulk, all transfers from the storage goblet to the shipping goblet should take place under liquid nitrogen. Lifting individual straws into room temperature air to transfer from liquid storage to vapor shipper should be avoided. Straws packaged in large goblets for bulk shipment should also have cotton packed around them to prevent cracking. The following paperwork should be included with each shipment: 1. Instructions for thawing and handling the semen. 2. A transaction report or frozen semen shipment form such as that provided by the Society for
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(a)
(b)
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Figure 320.8 (a–d) A technician demonstrating the proper technique for transfer of frozen semen straws from shipper to permanent storage.
3. 4. 5. 6.
Theriogenology, which includes the following information: a. stallion’s name; b. number of doses included; c. collection date, lot number, or straw ID code; d. number of straws per AI dose; e. number of total and progressively motile sperm per dose; f. expected post-thaw motility; g. semen equine viral arteritis (EVA) status; h. mare’s name, breed, and registration number if available; i. mare owner’s name. Mare management recommendations. Instructions for return of container. Statement of policy concerning unused semen. Applicable breed registry insemination or frozen semen distribution certificates.
Considerations for the further development of the equine frozen semen industry Quality standards for frozen semen While recent trends indicate an increase in the use of equine frozen semen, a number of factors have limited the widespread adoption of this method of breeding. One of those factors is reluctance by many breeders and veterinarians to promote its use. This reluctance is in part due to previous bad experiences with frozen semen that was of poor quality. European Union (EU) member states have established requirements for approved equine AI and semen production facilities as well as donor stallion health testing requirements for frozen semen produced for use in the commercial market. In countries outside
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Storage Management and Distribution of Frozen Semen the EU, however, these requirements are only applicable for semen produced for export. Many other countries such as the USA, Australia, New Zealand, and others also have established regulations concerning the importation of frozen semen into their respective countries. These regulations are primarily concerned with preventing disease transmission through transported frozen or cooled semen and do not make any recommendations concerning semen quality standards. Furthermore, within many countries (such as the USA) there are few regulations of any kind governing semen production, transport, and use within the country. As a result, the quality of semen that a mare owner or veterinarian receives for insemination is highly variable and often well below minimum standards to expect a reasonable outcome of success. This is true for both cooled and frozen semen. However, the frozen semen technology itself is often blamed for negative results from poor quality semen that should never have been used in fulfillment of a commercial contract. The World Breeding Federation for Sport Horses (WBFSH) is a non-profit federation of sport horse studbooks from around the world that has proposed minimum quality standards for frozen and fresh semen being exchanged between participating WBFSH studbook members. For frozen semen the minimum standards recommended by a WBFSH veterinary committee are that each insemination dose of frozen semen should contain a minimum of 35% progressive motility and a minimum of 250 million progressively motile spermatozoa after thawing. This is certainly a step in the right direction toward restoring faith in the quality of semen provided by member facilities and providing some reassurance to breeders who purchase semen; however, problems still remain. There exists no objective, standardized system for performing the quality assurance that semen placed on the market meets these minimum standards. Ideally, quality control should be provided by independent external quality control laboratories using objective and standardized methodology to determine post-thaw sperm motility and concentration measurements. All too often, semen is sold with a claim that the doses meet such minimum criteria but this is based solely on assertions by the stallion owner or laboratory that processed the semen. Furthermore, unless an objective system of motility analysis is used such as that provided by computer-assisted semen analyzers (CASA) the subjective estimation of post-thaw motility is prone to bias and will likely vary widely between technicians and laboratories. The algorithm that defines progressive motility may even vary between CASA systems and laboratories, and therefore must be standardized if values for two different laboratories are to be compared. The number of sperm per dose must also be independently
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confirmed using validated objective techniques such as that provided by CASA or the recently developed fluorescent-based sperm-counting devices, or through properly performed manual counts by a trained technician using a hemocytometer. Many processing laboratories obtain an initial sperm concentration and then simply assume a certain rate of recovery after centrifugation of the semen and resuspension in freezing extender prior to loading straws. This is not always consistent and may lead to a lower number of sperm per dose than expected. Frozen semen as a commodity Unlike bovine frozen semen, which is sold by the dose as a commodity, frozen semen from stallions is not thoroughly fertility tested before being sold on the market. A dairy farmer who purchases a dose of frozen semen from a Holstein bull from a reputable AI or genetics company can be very confident of the fertility potential of that dose of semen when handled properly and inseminated into a fertile cow. The reason for this is that dairy sires have been selectively bred for milk production and fertility, more specifically fertility in an AI program with frozen semen. The range in fertility per cycle for commercial AI bulls is very small and only a few percentage points separate high-fertility bulls from average and low-fertility bulls. Furthermore, bulls with low fertility with frozen semen are not commercially viable and are culled from commercial production so their frozen semen never reaches the market. Fertility testing of a young dairy sire’s frozen semen involves the insemination of thousands of cows to provide the estimate of fertility for that bull and the basis on which the decision is made to keep him or cull him from commercial production. Contrast this with the situation for horses. Even if one had the required number of mares available, stallions do not produce sufficient AI doses per ejaculate to allow fertility tests to be conducted, practically or cost-effectively, with their frozen semen. Stallions are not selected based on fertility, let alone on how well their semen freezes. The best a mare owner can hope for is that the semen has been frozen by a reputable laboratory with proven protocols and that the frozen semen has undergone a thorough post-thaw evaluation and that any semen put on the commercial market meets strict quality standards based on available critical laboratory assays. Given these limitations it seems logical that the risk associated with breeding horses (by any means) should be shared by both the stallion and mare owners. The selling of semen by the dose encourages mare owners to try to minimize the number of doses used (purchased) so that a pregnancy is achieved at the
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lowest cost. This approach will result in much higher veterinary costs for management of the insemination of that mare and may also result in lower fertility in an attempt to conserve doses of semen. Frozen semen sold by the dose is often sold without any restrictions on the number of breeding certificates issued and may encourage a mare owner to split multiple straw doses for insemination of two or more mares. This could lead to lower overall fertility from many stallions and contractual problems between mare and stallion owners. For the reasons stated above it would seem prudent that frozen semen be sold on a contract basis rather than as a commodity by the dose.
Contract considerations In addition to the elements commonly addressed in a standard breeding contract, those that involve frozen semen should clarify a number of situations unique to using this breeding method. For example: 1. How many insemination doses will be provided under the contract? a. How many doses will be provided per shipment/cycle? b. What is the limit on the number of doses provided per season? 2. What is the definition of a dose? a. Is there a guarantee for the number of sperm and the expected post-thaw quality? b. How will disputes over semen quality be resolved? 3. Who is responsible for payment of distribution/ shipping costs? 4. If a mare becomes pregnant and unused doses remain in storage what is the policy concerning unused semen? a. Who owns the unused doses? b. Should it be returned? If so, who is responsible for shipping costs? Our recommendation is that once frozen semen leaves inventory from its original place of storage and is maintained at another facility, it should not be returned to inventory for resale to another mare owner as the conditions under which those doses have been stored cannot be verified. If semen is to be returned to inventory it should only be used by the stallion owner on his/her own mares or shipped back to the same mare owner.
c. Can additional mares be inseminated with unused doses and, if so, what additional fees are payable to the stallion owner and on what terms? 5. Is there any guarantee of fertility? If so is it a live foal or pregnancy guarantee? If the mare fails to become pregnant is the mare owner entitled to: a. Additional doses of semen – are there limits on the number of doses? b. Additional seasons of breeding – are there limits on the number of seasons? c. Any refund or release from obligation for some or all stud fees? 6. Are there any restrictions on the use of the semen for assisted reproductive techniques such as embryo transfer, oocyte transfer, or intracytoplasmic sperm injection (ICSI)? Clarification of these issues in a comprehensive breeding contract will avoid potential problems between stallion and mare owners. Because frozen semen can be stored indefinitely, one final issue for consideration is the ownership and use of frozen semen after the sale of a stallion. The terms of sale for a stallion with stored frozen semen should clearly indicate which party is entitled to the rights for the use of that semen. Since most registries require breeding certificates issued by the stallion owner for foal registration and since each registry has adopted different rules regarding this issue, a clear understanding of the rules for a stallion’s particular breed registry should be clarified before completing a sale. For example, the American Quarter Horse Association (AQHA) has specifically addressed this issue in their regulations. Any owner who wishes to retain breeding rights after the sale of their stallion must purchase ‘retained semen rights permits’ from AQHA. Each permit may be used for the registration of one foal only and permits will only be sold to the recorded owner or lessee of the stallion. Once the stallion is sold the former owner cannot purchase additional permits. The ownership of the retained semen rights permits can be transferred and each transfer must be recorded by AQHA. Advances in the fertility of frozen semen will no doubt lead to an even greater acceptance of this technology in the coming years and therefore a thorough understanding of basic frozen semen handling should be included in the training of veterinarians and technicians working in equine reproduction.
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Chapter 321
Diseases Potentially Transmitted with Frozen or Cooled Semen Peter J. Timoney
Introduction For many years, it has been known that the spread of certain equine infectious diseases can occur by the venereal route. As far back as the late 19th and early 20th centuries, the stallion was believed by some to play a significant role in the transmission of ‘equine influenza’ or ‘pink-eye’ to mares that he bred. Based on investigation of naturally occurring outbreaks of ‘equine influenza’, Pottie concluded that ‘Apparently it is the seminal fluid which propagates the disease’.1 This observation was supported in subsequent reports by Clarke,2 Reeks,3, 4 Bergman,5 and Schofield.6 Reeks, in referring to the transmission of ‘pink-eye’ by apparently healthy stallions, stated that the ‘stallion spreads the disease by contagion only, and is not contagious except by coition’.3, 4 Based on the clinical and epidemiological features of the disease described at the time, it is very likely to have been equine viral arteritis (EVA), which was not etiologically defined nor pathologically characterized until many years later.7 The potential risk of global spread of equine diseases that can be transmitted by the venereal route has increased considerably over the past 20 to 30 years. This is primarily a consequence of changing trends in the breeding practices that have been adopted by the equine industries in many countries. Those of considered importance include:
r r
continued growth in the volume of national and international movements of stallions and mares for breeding purposes; the practice of ‘dual-hemisphere’ breeding of stallions that was introduced in the late 1970s;
r r r r
acceptance of artificial insemination (AI) with cooled or cryopreserved semen by all but one of the major horse breed registries in the USA and elsewhere; significant growth in the demand for embryo transfer; breeding of mares southern hemisphere-time in the northern hemisphere; increase in the ‘book size’ of mares bred to specific stallions in a given season.
The risk of transmission of equine diseases known to be or which potentially could be transmitted by the venereal route is greatest when breeding takes place by natural means. This is well illustrated in the case of contagious equine metritis (CEM), where there is direct mucosal contact between the surface of the penis and prepuce of the stallion and the clitoral area and distal reproductive tract of the mare. As a consequence, the chances of transmission of this disease are greatly increased, regardless of whether the stallion or the mare is the source of infection. By comparison, AI not only can reduce the risk of spread of certain venereally transmissible diseases but also, and very importantly, can eliminate the possibility of dissemination of diseases such as equine influenza that are contracted through other routes of exposure and are contingent on physical contact between mare and stallion at the time of breeding. Significant technical and procedural advances in how to optimize the handling and freezing of equine sperm have made it possible to transport semen virtually anywhere around the world. Furthermore, stocks of equine germplasm from premium stallions can now be preserved for future use, years after the
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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breeding careers of the donor animals have come to an end. It must be emphasized, however, that the benefits afforded by AI can very easily and rapidly be undermined with the potential for significant economic consequences, if the health status of donor stallions is not monitored on a regular basis to ensure their continued freedom from known venereally transmissible pathogens. The 2006 multistate occurrence of EVA in the USA illustrates very clearly the importance of having such a monitoring program in place. The origin of that occurrence was traced to an equine arteritis virus (EAV) infected stallion in New Mexico whose semen was in high commercial demand.8 Within a period of only a few weeks, fresh-cooled semen from the particular stallion that unbeknownst to the stallion manager at the time was contaminated with EAV, was transported to breeding farms in 18 states and two provinces in Canada. Multiple outbreaks of EVA resulted, characterized by clinical disease including high rates of abortion on some premises and establishment of the carrier state in a number of stallions. This and other similar occurrences of EVA underscore that while AI can be an effective way of preventing transmission of certain diseases, it can also serve as a highly efficient means of disseminating particular equine venereally transmissible pathogens. Artificial insemination has an undoubted value for increasing reproductive efficiency in the equine and mitigating the risk of spread of disease.9 The success of AI, however, is dependent on (i) ensuring the continued freedom of breeding stallions from venereal diseases; and (ii) observance of high standards of hygiene in managing and collecting such stallions. The widely accepted benefits of AI can so easily be compromised, however, if appropriate biosecurity measures are not in place to preclude the indirect transfer of a disease agent between stallions using the same breeding shed/semen collection facility. Of additional importance is whether a stallion used for AI is also being bred to mares by natural service without due precautions being taken to prevent him becoming contaminated/infected with a particular pathogen. Indirect contact through the use of contaminated veterinary instruments or breeding shed equipment has been shown repeatedly to be a very effective means of dissemination of EAV and Taylorella equigenitalis, causal agent of CEM. Interim findings from the 2008/2009 occurrence of CEM in the USA indicate that the 23 carrier stallions that were identified did not become contaminated with T. equigenitalis as a result of direct venereal contact between each stallion and an infected mare. Circumstantial evidence would indicate this occurred indirectly through stallion-tostallion transfer of the bacterium by means of contaminated fomites.
Of the wide range of infectious diseases of the horse, only a very limited number have been shown to be or are considered to have the potential to be transmissible through fresh-cooled or cryopreserved semen. The majority of infections for which there is evidence of transmission by the stallion or donkey jack are bacterial (Streptococcus zooepidemicus, Pseudomonas aeruginosa, Klebsiella pneumoniae, T. equigenitalis, and T. asinigenitalis); a couple are viral (EAV and equine herpesvirus 3 or equine coital exanthema virus), and one is a protozoan infection (Trypanosoma equiperdum).10 Aside from this group of bona fide or potential equine venereal pathogens, there are other diseases and infectious agents that may be transmitted very infrequently by this route or that potentially could be disseminated through contamination of semen. These can be arbitrarily categorized as follows: 1. internal genital infections of the testes, epididymides, and accessory sex glands, the vast majority of which are bacterial in nature, e.g. S. zooepidemicus, Salmonella abortus equi; 2. acute or chronic systemic diseases characterized by circulation of the respective causal agents in the bloodstream, e.g. equine infectious anemia, equine piroplasmosis; 3. infections of the urinary system with the potential for contamination of the urogenital tract, e.g. leptospirosis, equine rhinitis A virus infection; 4. a miscellany of infective agents, including viruses, bacteria, mycoplasmas, chlamydiae, fungi, and yeasts, that can inhabit the stallion’s genital tract and consequently may be shed in semen. Although of minor significance as bona fide or potential venereal pathogens, these different categories of infectious agents or diseases will be considered in greater detail later in this chapter. Notwithstanding the fact that S. zooepidemicus, P. aeruginosa, and K. pneumoniae are the venereal pathogens most commonly recovered from cases of endometritis in the mare,9 they are not the infectious agents of greatest concern to breeding industries in the USA or elsewhere. The two most regulated equine venereal diseases at the present time and those with the greatest impact on the international trade in horses, semen, and embryos are CEM and EVA. The respective etiological agents are true venereal pathogens, even though EAV can also be very efficiently transmitted via the respiratory route by acutely infected equines.11 Neither dourine nor S. abortus equi infection should be overlooked in terms of their significance. Both have historical precedence over CEM and EVA in being recognized many years earlier as diseases that are transmitted by the stallion solely or primarily by the venereal route. The
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Diseases Potentially Transmitted with Frozen or Cooled Semen intent of this review of equine diseases that are potentially transmissible through cooled or cryopreserved semen is not only to emphasize those of accepted importance, namely CEM and EVA, but also to consider less common or less significant diseases/infections where the causal agent can be detected in semen, although its role in venereal transmission may not yet be scientifically established. In the case of CEM and EVA, aspects of the occurrence, causation, main clinical and epidemiological features, and diagnosis and prevention will be covered for each disease.
Contagious equine metritis Occurrence Contagious equine metritis is, as its name would indicate, a highly transmissible infection of equids. It can only be transmitted by the venereal route.12 First recognized in the Thoroughbred breeding populations in the UK and Ireland in 1977,13, 14 the disease has since been reported from a significant number of countries around the world, in some of which it remains endemic, mostly in non-Thoroughbred breeds of horses. However, certain countries have successfully eradicated CEM from their resident horse populations through implementation of comprehensive testing and control measures supported by stringent import regulations that minimize the risk of reintroduction of the disease. Etiology The causal agent of CEM is a bacterium, T. equigenitalis, unknown before 1977.15, 16 It is a Gram-negative coccobacillus, with fastidious growth requirements that include microaerophilic conditions of incubation. Two biotypes of the bacterium have been recognized, those sensitive or resistant to streptomycin. The organism is susceptible to a wide range of antimicrobials and commonly used disinfectants and is not especially resistant outside the body. Molecular genotyping of strains of T equigenitalis by pulsed-field gel electrophoresis has shown that it is possible to divide strains into at least 10 groups.17, 18 The natural host range of T. equigenitalis appears to be restricted to equids;19, 20 there is no evidence that humans can be infected with the bacterium. Clinical features Contagious equine metritis is a non-fatal, nonsystemic disease with infection restricted to the reproductive tract. The disease can result in shortterm infertility upon primary infection in the mare, frequently but not invariably associated with an
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endometritis.13, 14, 21 Abortion, although it has been recorded, is a very rare sequel to T. equigenitalis infection.22 In contrast to the mare, exposure of the stallion to T. equigenitalis, strictly speaking, does not result in infection, much less the development of any clinical signs in the vast majority of cases.23 Instead, the stallion becomes an inapparent carrier of the bacterium, which persists as a commensal on its external genitalia without causing any adverse local or systemic effects. There has been one report, however, where T equigenitalis was isolated from the testis, epididymis, seminal vesicles, and urethra of a stallion at necropsy examination.24 This would suggest that on very rare occasions, exposure may result in an ascending infection of the reproductive tract in the stallion. First-time exposure of the mare to T. equigenitalis can result in clinical or asymptomatic infection. There are disparate opinions over the clinical significance of the infection, with some in the scientific community, mainly in countries in which CEM is endemic, regarding the causal bacterium ‘as an occasionally pathogenic commensal’.25 After an incubation period of 2–12 days, characteristically affected mares develop an odorless, greyish-white mucopurulent vulvar discharge of variable consistency and volume, which can last for up to 2 weeks, or slightly longer. 26, 27 The discharge is associated with an endometritis, cervicitis, vaginitis, vulvar discharge, and a return to estrus after a shortened diestrous period.12 The infertility is temporary, lasting only a few weeks, with no long-term adverse effect on a mare’s fertility. Persistence of T. equigenitalis in the reproductive tract of a chronically infected mare will not, in the vast majority of cases, interfere with the maintenance of pregnancy or the birth of a healthy foal.28 Epidemiology Of greatest significance in the persistence and dissemination of T. equigenitalis in different equid populations is occurrence of the carrier state in the stallion and the mare.15, 29 The carrier state, which can be established in a high percentage of exposed stallions, may if the latter are left untreated, last for years. Sites of persistence of T. equigenitalis include the urethral fossa and urethral diverticulum or sinus, the terminal urethra, surface of the prepuce and penis, and, less frequently, the pre-ejaculatory fluid.15, 29 Carrier stallions have been shown to represent the greatest risk with regard to the international spread of CEM.30 The carrier state can also develop in a variable percentage of mares following recovery from the acute phase of T. equigenitalis infection. While the great majority of such mares are clitoral carriers, the bacterium can persist in the uterus of a much smaller percentage
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of cases. The sites of persistence in mares that are clitoral carriers are the clitoral sinuses (medial and lateral) and the clitoral fossa.23 Foals, especially colts, that are from the chronically infected or carrier mares may become contaminated with T. equigenitalis and harbor the organism on their external genitalia for an extended period of time.31 As previously indicated, transmission of T. equigenitalis occurs through direct or indirect venereal contact with an infected mare or carrier stallion. Although not of proven significance, a carrier teaser stallion also has the potential to serve as a means of transmitting CEM. Based on experience gained from natural outbreaks of the disease over the past 30 years, there is considerable evidence to show that natural breeding is a very efficient means of establishing infection in the mare or the carrier state in the stallion. That notwithstanding, indirect contact through contaminated fomites can also play a major role in dissemination of the disease. Indeed, spread of CEM between many of the affected Thoroughbred breeding farms in the UK and Ireland in 1977 was attributed to the use of contaminated veterinary instruments such as non-disposable vaginal specula, vulsellum forceps, etc., that were inadequately disinfected between premises. The potential for dissemination of T. equigenitalis through indirect contact is equally great where breeding takes place by AI. Lack of observance of appropriate biocontaminant measures in semen collection facilities has led to lateral transfer of the bacterium from stallion to stallion; this would appear to have been responsible for contamination of probably all of the 23 carrier stallions that have been identified with respect to the 2008/2009 occurrence of CEM in the USA. Among the breeding shed equipment most frequently implicated in the spread of T. equigenitalis in such situations are artificial vaginas used without disposable liners or inadequately disinfected between collections; and a phantom mare that does not have an outer disposable cover or that is not decontaminated between stallions. Indirect transmission of CEM can also readily take place through personnel not observing adequate hygienic precautions, for example failure to wear disposable gloves when handling the external genitalia of the stallion or mare at time of breeding or semen collection. Aside from the importance in the transmission of CEM of natural breeding or indirect contact with T. equigenitalis-contaminated fomites, AI can also be considered an additional means of spread of the disease. While there is only one report in the veterinary literature substantiating the fact,32 recent evidence from the 2008/2009 occurrence of CEM in the USA proves that transmission of T. equigenitalis by AI can occur through the use of transported, freshcooled semen from a carrier stallion. Of over 717
mares exposed to infection, five have been cultured positive for T. equigenitalis. Of these, three had been exposed through insemination with fresh-cooled semen from known carrier stallions, one after being bred naturally to a stallion that subsequently turned out to be a carrier. It is not possible from the breeding history of the fifth mare to determine whether she contracted infection following natural breeding or AI. She was bred naturally in 2007 to a stallion that was confirmed a carrier in 2009, and in 2008 she was bred artificially with cryopreserved semen from a different carrier stallion. Although these findings confirm the potential for transmission of T. equigenitalis to mares by AI with extended or fresh-cooled semen, based on currently available data the frequency with which this can occur would appear to be very low. There is evidence that the use of appropriate antibiotic-containing extenders significantly reduces the infectivity in semen from a carrier stallion to a level that results in a decreased risk of infection in inseminated mares. The findings from in vitro and in vivo studies with extenders containing amikacin and penicillin, or ticarcillin, indicate that this is indeed the case (C. Klein et al., personal communication). Currently it is unclear what the risk of transmission of T. equigenitalis is using cryopreserved semen from a carrier stallion. As yet, it has not been possible to establish whether this can take place, and if so at what frequency level. There has only been one published report of a study that investigated the effect of several different antibiotic-containing extenders on the survival of the bacterium in contaminated semen.33 While gentamicin proved to be the most effective when semen-extender mixtures were held at ambient temperature (22 ± 2◦ C) rather than at 4◦ C or 37◦ C, such treatment did not completely eliminate infectivity under the conditions of this study. This is supported by the findings of a recent controlled study in which using a standard protocol for extending semen from T. equigenitalis carrier stallion with a commercial extender containing Amikacin sulphate and penicillin, infection levels of the bacterium were greatly reduced but not eliminated. The treatment used had no adverse effect on motility or other accepted parameters of sperm function. This would similarly apply in the case of cryopreserved semen. Transplacental transmission of T. equigenitalis can occur infrequently in the chronically infected pregnant mare. This may result in foals either congenitally infected or contaminated with the bacterium at birth31 or, very rarely, in abortion.22 Diagnosis Contagious equine metritis cannot be diagnosed on clinical grounds alone because of its similarity to
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Diseases Potentially Transmitted with Frozen or Cooled Semen several other more commonly encountered bacterial infections of the mare’s reproductive tract caused by S. zooepidemicus, K. pneumoniae, or P. aeruginosa.23 It should certainly be considered in a situation where mares develop a mucopurulent vaginal discharge 2–14 days after being bred, or returning prematurely in estrus after a shortened diestrous period. There is some suggestion that certain strains of T. asinigenitalis may cause a similar clinical response in the mare. 34 Notwithstanding the development of a number of polymerase chain reaction (PCR) assays that purport to have a high level of sensitivity and reliability for the detection of T. equigenitalis,35–41 recovery of the bacterium in culture remains the internationally accepted ‘gold-standard’ for establishing a diagnosis of CEM.15, 29, 42, 43 There is a significant need for a fully validated PCR or alternative assay, which would provide a more rapid, less costly, yet highly reliable means of detecting T. equigenitalis in swabs from stallions and mares as well as in fresh-cooled and cryopreserved semen and equine embryos. Such an assay would circumvent the problems not infrequently associated with attempted culture of the bacterium from the heavily contaminated sites of persistence in the carrier mare and stallion, as well as from mares and stallions that may have been recently treated with antibiotics. In suspected clinical cases of CEM, smears should be made of any vulvar discharge to look for the presence of bacteria morphologically similar to T. equigenitalis, either extracellularly or phagocytosed by neutrophils. If present, such evidence is suggestive but not confirmatory of a diagnosis of CEM in the mare.10 The optimal sites for detection of T. equigenitalis in the maiden, barren, or post-parturient mare, are the clitoral sinuses and clitoral fossa, the uterus, and the cervix. The clitoral sinuses can only be successfully swabbed using narrow Calgi swabs or equivalent pre-moistened with sterile water.43, 44 Endometrial or cervical swabs should preferably be collected in early estrus. It is advisable to restrict the sampling sites in the pregnant mare to the clitoral fossa and clitoral sinuses and any vulvar discharge that may be present. The sites of persistence of T. equigenitalis that must be swabbed in the stallion include the urethral fossa and urethral diverticulum (sinus), the distal urethra, and the external surface of the penis and prepuce.43, 44 A sample of pre-ejaculatory fluid should also be tested if available. Collection of swabs from a stallion is best accomplished with the penis in full erection. Perhaps the most highly reliable, though costly and time-consuming, means of detecting the carrier state in the stallion is by test breeding the individual to two mares previously confirmed free of CEM. After breeding by live cover, the mares are checked for evidence of T. equigenitalis infection by culture
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and development of a complement-fixing antibody response within 21–45 days. When collecting specimens for the diagnosis of CEM, every effort must be made to ensure the reliability of the sampling process. Mares/stallions that have recently received local/systemically administered antibiotics must not be swabbed for at least 7 days or 21 days, respectively, after the completion of systemic or local treatment. To optimize the chances of culturing T. equigenitalis from swabs, the swabs should be placed immediately after collection in individual tubes of Amie’s transport medium with charcoal but without antibiotics, and transported, preferably under conditions of refrigeration, to a laboratory that has been approved to test for CEM. Swabs must be plated out on appropriate isolation media within 48 hours of collection.42 Serological testing for CEM is of limited diagnostic value.15, 23, 43 It is only useful for detecting recent exposure to T. equigenitalis in the mare when applied within 21–45 days after breeding. There is no value to serological testing of stallions or mares for detection of the carrier state. Control Effective prevention and control of CEM at a national level is predicated on the disease being made notifiable or non-notifiable but reportable.15, 23 Integral to the success of such programs is the availability of laboratories with the requisite experience and expertise to diagnose the infection. Countries that are free of CEM must screen all imported mares and stallions from known or suspect CEM-affected countries during the period of post-entry quarantine. Such testing shall be in accordance with previously described procedures for detection of T. equigenitalis in the stallion and the mare. Even with implementation of comprehensive testing protocols, there is always the potential for a carrier mare, and especially a carrier stallion, to escape being detected in post-entry quarantine, with the consequent risk of reintroduction of CEM into the domestic equine population. Codes of Practice have been developed by certain European countries; these provide guidelines for veterinarians, owners, and breeders for the prevention and control of CEM. They are based on comprehensive bacteriological screening of breeding stallions and mares to identify asymptomatic carrier animals.12, 44 Mares and stallions cultured positive for T. equigenitalis must be isolated and not bred until treated and confirmed free from the bacterium. Such codes emphasize the importance of improved standards of hygiene on breeding farms, and the more widespread use of disposable and sterile materials and equipment to minimize the risk of spread of CEM and certain other equine venereal diseases.
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Experience has shown that where the foregoing measures and safeguards have been adopted, it is possible to bring this disease under control within a few years and even perhaps to eradicate it.
Equine viral arteritis Occurrence Equine viral arteritis is an acute contagious disease of equids, characterized principally by fever, dependent edema, leukopenia, and respiratory signs, with the potential for abortion in pregnant mares and a fulminant pneumonia or pneumoenteritis in very young foals.11 The disease is transmissible by the respiratory route and also venereally, by the acutely infected mare and stallion and by the chronically infected or carrier stallion. Equine viral arteritis was first etiologically defined and the disease pathologically characterized following a widespread outbreak of respiratory disease and abortion on a Standardbred farm near Bucyrus, Ohio, in 1953.7, 45 To date, occurrences of EVA have been recorded only in North America and Western Europe, even though the causal virus, EAV, has been found to be widely distributed in equine populations worldwide.11,46–49 There is growing evidence, however, that the frequency of confirmed outbreaks of EVA has been increasing in more recent years; this has been linked in part to the increasing international trade in horses and equine semen. The prevalence of EAV infection can vary between countries and between different breeds in the same country, with highest rates of infection frequently found in Standardbreds and Warmblood breeds.47, 50 Equine viral arteritis remains one of the most internationally regulated diseases with respect to the movement of horses for breeding or performance purposes. Etiology The causal agent of EVA is EAV, an RNA virus and the prototype virus of the genus Arterivirus, family Arteriviridae, order Nidovirales.51 Equine arteritis virus is one of the three most important equine viral respiratory pathogens, along with equine influenza virus and equine herpesviruses 1 and 4.52 However, as has already been mentioned, the virus can be transmitted venereally as well as by the respiratory route. Historically, EVA was considered a disease solely of members of the family Equidae.11 There is some recent information, however, which would suggest that the virus can also infect New World camelid species, namely alpacas and llamas.53 Equine arteritis virus is not known to have any public health significance.52
Although only one major serotype of EAV has been identified so far, namely the Bucyrus strain, there is ample evidence of genomic and phenotypic variation among temporarily and geographically disparate viral isolates.54 Phylogenetic analysis based on sequencing open reading frame 5 has grouped field isolates of EAV into two distinct clades, a North American lineage and a European lineage.55, 56 Equine arteritis virus has a predilection for bloodderived macrophages together with endothelial and medial myocytes of smaller blood vessels, causing a panvasculitis that especially affects arterioles.45 Strains of the virus can vary in their pathogenicity for the horse.11 Experimental infectivity studies have identified lentogenic, mesogenic, and velogenic strains of the virus.57 Equine arteritis virus is not resistant outside the body.54 While its viability at temperatures at or above 37◦ C is short-lived, the virus can remain infective for extended intervals in tissue and fluid specimens kept at or below freezing temperature. Cryopreserved semen from carrier stallions can retain infectivity for many years and be a very efficient means of dissemination of the virus. Clinical features Primary exposure to EAV may result in either asymptomatic infection or clinical disease depending on the virus strain involved, virus dose, age and physical condition of the horse, and various environmental factors.11 Equine viral arteritis is a systemic infection that can affect the reproductive tract and lead to abortion or establishment of the carrier state in the stallion. It should be emphasized that most horses naturally infected with EAV for the first time are asymptomatic. Clinical severity of the infection tends to be greater in very young or old animals or those in a debilitated condition. Equine viral arteritis is rarely a life-threatening illness in older horses. Aside from abortion, fatal infection primarily occurs in neonatal foals or in foals up to a few months of age, which die from a fulminant viral pneumonia or pneumoenteritis. After an incubation period of 3–14 days, affected horses will display any or most all of the following clinical signs:
r r r r r r r
fever of up to 2–9 days’ duration; depression; anorexia; leukopenia; dependent edema, especially of the lower hind limbs, scrotum, and prepuce in the stallion, and mammary glands in the mare; conjunctivitis; supra- or periorbital edema;
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epiphora; photophobia; rhinitis and nasal discharge; urticarial-type skin reaction, which may be localized to the facial region and sides of the neck, but can be more generalized.11
Horses in training may experience a period of impaired performance while acutely affected or convalescing from the disease. Pregnant mares exposed to EAV may abort either late in the acute phase or early in the convalescent phase of the infection; this can be a sequel to clinical or asymptomatic infection.11 Abortion can occur from 2 to over 10 months of gestation, with rates ranging from less than 10% to 80% in unprotected mares. Mares exposed to EAV very late in pregnancy may not abort but carry to term and give birth to a foal congenitally infected with the virus.58 It must be emphasized that abortion occurs in mares that are already pregnant at time of exposure to EAV; this usually takes place via the respiratory route through direct contact with an acutely infected horse.54 Abortion is not a sequel to breeding a mare with infective semen from a carrier stallion. Mares that abort from EAV infection do not experience any related short- or long-term fertility problems. Stallions acutely affected with EVA that experience prolonged fever and severe scrotal and preputial edema may undergo a period of temporary subfertility.59 There is no evidence, however, that this is virus-mediated. Such stallions have reduced libido associated with a decrease in sperm motility, sperm concentration and percentage of morphologically normal sperm. These changes can last for 14–16 weeks, after which affected stallions return to their pre-infection level of fertility. Epidemiology A range of factors, related to the virus, host, and environment, have been identified with the epidemiology of EVA. Outbreaks of the disease are almost invariably associated with the movement of horses or use of shipped semen.52 Among the epidemiological factors known to be involved are:
r r r r r
variation in pathogenicity and other phenotypic characteristics of naturally occurring strains of EAV; modes of transmission during acute and chronic phases of infection; occurrence of the carrier state; nature of acquired immunity to infection; changing economically driven practices in the horse industry.54
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Wide-ranging variation in pathogenicity has been observed among field strains of EAV, with some capable of causing severe disease and others merely a subclinical infection. Transmission of EAV occurs primarily by the respiratory and venereal routes, although the virus can also be transmitted by congenital infection and by indirect means through contact with contaminated fomites including the placenta from cases of EVArelated abortion. Infection can also be transmitted by embryo transfer where the donor mare is bred with semen (C. Broaddus, personal communication). Respiratory transmission is the principal mode of spread of EAV at performance events, on breeding farms, and at sales and clinics. The virus can also be very efficiently transmitted venereally by the acutely infected stallion or mare. Carrier stallions are a major source of EAV for breeding populations through venereal transmission, either by natural breeding or AI using fresh-cooled or frozen semen. The carrier state has been confirmed in sexually mature intact males, including post-pubertal colts and stallions, but not in mares or geldings.11 The virus is restricted to the reproductive tract in the carrier stallion, where it is harbored primarily in certain of the accessory sex glands, especially the ampullae of the vasa deferentia. Persistence of EAV in the stallion has been shown to be testosterone-dependent. Frequency of the carrier state can vary from less than 5% to greater than 70%. Duration of viral persistence can be very variable, from weeks to a period of months, to many years. Carrier stallions are constant semen shedders of EAV, which is present in the sperm-rich fraction of the ejaculate. There is no evidence of virus shedding in any other secretion or excretion. The carrier stallion is widely acknowledged as the natural reservoir and source of novel variants of EAV; it serves as a highly efficient transmitter of the virus, without seemingly any related adverse effect on its fertility or general state of health. A variable percentage of long-term carrier stallions spontaneously cease to shed in their semen and successfully eliminate the virus from their reproductive tract. Equine arteritis virus is unique among equine respiratory viruses in its ability to stimulate a strong, durable immunity. This is effective in protecting against clinical disease and establishment of the carrier state in the stallion. Foals born of seropositive mares from prior natural infection or vaccination should have passive protection as long as they maintain serum neutralizing antibody titers ≥ 1:32 to the virus. A number of developments in the horse industry over the past 30 to 40 years have impacted the global distribution of EAV. Firstly, the considerable growth in the national and international movement
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of horses, especially stallions, for breeding purposes; and secondly, the significant increase in the volume of shipped fresh-cooled and cryopreserved semen. There have been numerous instances where EAV has been introduced onto farms or into countries through the importation of a carrier stallion or virus infective semen.60 On occasion, these movements or importations have resulted in economically damaging outbreaks of EVA. Diagnosis In view of the similarity of EVA to a range of other infectious diseases in the horse, it is essential to pursue laboratory confirmation of a provisional clinical diagnosis of EVA.11 Diagnosis is based on virus detection either by virus isolation in cell culture or by PCR assay, by immunohistochemical demonstration of viral antigen, and, finally, by serological testing of paired (acute and convalescent) sera taken 21–28 days apart. Appropriate specimens for virus detection from clinical cases of infection include nasopharyngeal swabs or washings and unclotted blood samples collected into acid-citrate-dextrose or EDTA (ethylene diamine tetracetate) anticoagulant. To optimize the chances of virus detection, it is very important to obtain the specimens as soon as possible after the onset of clinical signs or suspicion of infection. Swabs must be transferred into viral transport medium and shipped refrigerated or frozen to a laboratory with proven competency to test for EAV infection. In the case of abortion or fatality in a foal or older horse, detection of EAV should be attempted from a range of tissues and body fluids, including the lung and lymphatic glands in the thoracic and abdominal cavities and related organs. Specimens should also be taken for histological and immunohistochemical examination. Detection of the carrier state in the stallion is based initially on confirmation of the seropositive status of the individual for antibodies to EAV. In the absence of a certified history of vaccination against EVA, antibody-positive stallions must be collected and their semen tested for evidence of EAV by virus isolation in cell culture or PCR assay. A very reliable though costly and time-consuming alternative is to test-breed the suspect carrier stallion to two seronegative mares and check them for seroconversion to EAV 28 days later. Control Equine viral arteritis is a manageable and preventable disease.11 It can be controlled through a combination of sound management practices and the implementation of a vaccination program against the
disease. The success of any control program against EVA is predicated on minimizing or eliminating direct or indirect contact of unprotected horses with the secretions/excretions of infected animals. Current control programs focus on preventing the introduction and dissemination of EAV in breeding horse populations to obviate the risk of virus-related abortions, fatal illness in young foals, and establishment of the carrier state in stallions and post-pubertal colts. In view of the infrequency of occurrence of outbreaks of EVA at racetracks, sales, shows, and other performance events, programs have not been developed that are specifically focused on preventing spread of the virus at such venues. Recommended precautions to reduce the risk of an outbreak of EVA on breeding farms include isolation of all new arrivals on the premises for 3–4 weeks, and segregating pregnant mares from other horses and grouping them by stage of gestation until they foal. Additionally, all stallions should be screened for presence of the carrier state and any carriers physically isolated. Breeding stallions should be annually vaccinated against EVA. This also applies in the case of teaser stallions. Special attention should be paid to the management and breeding to carrier stallions. Guidelines for breeding a mare to a carrier stallion were developed by the American Horse Council and subsequently endorsed by the American Association of Equine Practitioners.61 Strict precautions must be taken when breeding or collecting these stallions to avoid inadvertent spread of EAV to other horses on the premises. A carrier stallion should only be bred to mares vaccinated against the disease or those seropositive from previous natural exposure to the virus. Vaccinated mares bred to a carrier stallion for the first time should be isolated from other unprotected horses for 3 weeks. In breeds in which AI is permitted, it is important to screen semen from stallions of unknown carrier status for the presence of EAV, especially when using imported cryopreserved semen.60 The same precautions as those recommended for mares bred by natural cover should be observed when breeding mares by AI with virus-infective semen. In an attempt to curtail the number of carrier stallions, a concerted effort should be made to encourage a program of vaccination against EVA of colt foals between 6 and 12 months of age. Such a program, if broadly accepted by the industry, would in time lead to a reduction in the carrier stallion population, especially in breeds in which the infection is presently endemic. The potential economic impact that EVA can have on international trade in horses and equine semen emphasizes the need for an industry-driven program for the prevention and control of this disease.
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Klebsiella, Pseudomonas and streptococcal infections Many species of bacteria exist as commensal organisms on the external surface of the penis and the prepuce of the stallion. Although these are not normally considered a cause of uterine infection and infertility, especially in young, healthy mares,62 under certain circumstances a few of them, namely K. pneumoniae, (certain capsule types) P. aeruginosa, and S. zooepidemicus, can assume clinical significance for the mare. Problems can arise when there is disruption of the normal bacterial flora on the external genitalia of the stallion. This disturbance is usually the result of indiscriminate washing of the penis or too frequent washing with a surgical scrub, or detergent soaps,63, 64 having a stallion on contaminated bedding, or breeding a mare with a genital infection. 65 Under such circumstances, there is the potential for colonization of the penis and prepuce with the aforementioned bacteria, i.e., K. pneumoniae, P. aeruginosa, and S. zooepidemicus. These organisms, which rarely produce clinical disease in the stallion, can give rise to endometritis with reduced fertility in susceptible mares,9 especially in individuals with a pre-existing defect in uterine clearance.66 Transmission of these bacteria occurs venereally, either by natural breeding or AI with infective semen.9,67–72 Klebsiella pneumoniae While there are many polysaccharide capsule types of K. pneumoniae, only capsule types 1, 2, 5,25, 73, 74 and possibly type 775 and type 3976 have been incriminated in causing endometritis in the mare. There is evidence to indicate that transmission of these capsule types can result from AI, presumably with infective fresh-cooled semen, as well as from natural breeding.25, 69, 77 In cases of Klebsiella-induced endometritis, diagnosis can be confirmed by isolation of the bacterium from the semen or penis of the stallion as well as from the mare’s uterus,69 with capsule type identification now possible by PCR assay.78 Pseudomonas aeruginosa Outbreaks of P. aeruginosa infection in mares associated with endometritis and a vaginal discharge, can be a cause for concern.62, 70, 77, 79, 80 Additionally, the presence of this bacterium in cooled semen can adversely affect fertility of the semen.81 Because not all strains of P. aeruginosa cause disease,79 it is important to demonstrate the same serotype in the semen and/or on the penis of the stallion as that cultured from the uterus/vaginal discharge of the affected mare.64, 69
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Streptococcus zooepidemicus Although regarded as less significant compared with K. pneumoniae or P. aeruginosa infection, S. zooepidemicus is an opportunistic bacterium that can give rise to endometritis in a predisposed mare.82 A common inhabitant of the external genitalia of the stallion, it can be transmitted to the mare in semen through AI or natural breeding.62, 77
Equine coital exanthema Equine coital exanthema is a contagious viral venereal disease of horses caused by equine herpesvirus 3. The disease is characterized primarily by the development of vesicular lesions on the external genitalia of the stallion and the mare, which rupture giving rise to superficial ulcers.65 Secondary bacterial infection with S. zooepidemicus is not uncommon; this may influence the severity and duration of the lesions. Equine coital exanthema is a self-limiting, non-systemic infection, with a clinical course of 2–5 weeks. While natural infection with equine herpesvirus 3 has not been shown to cause infertility or abortion,83 some consider that mares infected with the virus may experience a temporary lowering in conception rate.84 Although coitus is the major route of exposure for stallions and mares, mares can also become infected by AI or through indirect contact with contaminated veterinary instruments or equipment used for breeding, gynecological examination, or testing for pregnancy.65, 66 It is believed but not proven that equine herpesvirus 3 can establish a latent carrier state in a proportion of recovered horses. If this is the case, it would provide a mechanism for persistence of the virus and a source of infection for outbreaks of the disease through reactivation of latent virus.
Dourine Dourine, or mal du coit, is a venereal disease of equids historically believed to be caused by Trypanosoma equiperdum.85 Originally considered to be widely distributed in equine populations around the world, the disease is currently thought to be restricted to Africa, South and Central America, and the Middle East.86, 87 Dourine is unique in certain respects in that it is the only known non-arthropodborne trypanosomal disease and the only protozoal infection of horses believed to be transmitted venereally. The causal agent has a tropism for the genital mucosa and is not considered capable of surviving outside the host.66 In recent years, molecular studies have demonstrated that T. equiperdum is very closely related to T. evansi and T. brucei. It has been
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speculated that dourine may in fact not be caused by T. equiperdum but by particular strains of T. evansi and/or T. brucei, and that T. equiperdum may not actually exist as a separate species.86 The incubation period, clinical signs, and course of dourine can vary between affected individuals; however, most cases have a fatal outcome.85 The clinical signs are considered by some to be sufficiently characteristic as to be diagnostic of the disease. A multisystemic syndrome follows an incubation period ranging from a week to 3 months. The initial phase of the disease is characterized by a periodic fever, genital edema that includes the prepuce, urethral process, penis, testes, scrotum, and mammary glands, enlargement of the inguinal lymphatic glands, and a urethral or vaginal discharge. This is followed by the appearance of raised cutaneous plaques, which may become depigmented. These are referred to as ‘silver dollar plaques’ and are considered pathognomonic of dourine. In the terminal stage of the disease, affected animals are anemic, emaciated and frequently display neurological signs, which may include ataxia, hyperesthesia, hyperalgia, hind limb paralysis, facial paralysis, and paraplegia, before death supervenes. Transmission of T. equiperdum is considered to take place almost exclusively by coitus.88 Spread of infection is believed to occur more readily from stallion to mare than vice versa. The organism can be found in the seminal fluid and in the discharge from the penis and vagina of affected individuals. Although transmission has never been confirmed by AI, based on the fact that T. equiperdum may be present in seminal fluid, there is the potential for this to occur. Of lesser significance with respect to the epidemiology of dourine have been reports of transmission of the causal organism by blood transfusion and possibly through the milk of infected mares.89, 90
Taylorella asinigenitalis In late 1997 and early 1998, a bacterium with very similar morphological and cultural characteristics to T. equigenitalis was isolated from several donkey jacks, a horse stallion and seven mares in the USA.91 This bacterium, which has since been reported from Sweden and France,92, 93 was identified as a separate but closely related species to T. equigenitalis and designated T. asinigenitalis.91 In contrast to T. equigenitalis, the natural host species of T. asinigenitalis would appear to be the donkey, not the horse. Although clinical evidence of infection was not reported in any of the mares naturally exposed to this bacterium in the 1998 outbreak in Kentucky, Katz et al.34 demonstrated that a Kentucky asinine strain of T. asinigenitalis was capable of causing endometri-
tis, cervicitis, and vaginitis in two experimentally infected mares, not dissimilar to CEM. Transmission of T. asinigenitalis occurs by the venereal route. In view of its very close similarity to T. equigenitalis, which has been shown to be transmissible by AI, there is every reason to believe that T. asinigenitalis also has the potential to be transmitted artificially via contaminated semen.
Internal genital infections Infection of the testes, epididymes, and accessory sex glands in the stallion may be a localized event or part of a systemic disease. Though rare in occurrence, orchitis can be caused by a range of bacteria including but not exclusive of S. abortus equi, S. zooepidemicus, S. equi, Burkholderia mallei, Brucella abortus, Actinobacillus equuli, and Escherichia coli.65 Infection of the accessory sex glands most commonly involves the seminal vesicles. While infrequently encountered, infectious seminal vesiculitis can be due to P. aeruginosa, K. pneumoniae, Streptococcus spp., Staphylococcus spp., Proteus vulgaris, or B. abortus. Infections of the testes and accessory sex glands are important not only for the detrimental effect they can have on a stallion’s fertility, but also because they afford the opportunity for venereal transmission of whatever etiological agent is involved. This could take place through coitus or by AI with contaminated semen.
Systemic diseases A limited number of acute or chronic systemic infectious diseases of horses have come under consideration for their potential to be transmitted by the venereal route. These include the equine viral encephalomyelitides, for example Eastern equine encephalomyelitis and Western equine encephalomyelitis, West Nile encephalitis, vesicular stomatitis, African horse sickness, equine infectious anemia, and equine piroplasmosis. Some believe that these diseases may pose a risk of venereal transmission, if at the time of breeding or semen collection there is traumatization of the stallion’s genitalia, bleeding, and contamination of the ejaculate with blood containing the respective etiological agent. This would occur during the acute or viremic phase of the infection in the case of the viral encephalomyelitides and African horse sickness, and at any time during the carrier phase in equine infectious anemia and equine piroplasmosis. Contrary to the opinion of the regulatory authorities in certain countries, there is no evidence of a viremia in horses infected with vesicular stomatitis virus, and consequently no possible risk of
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Diseases Potentially Transmitted with Frozen or Cooled Semen venereal transmission of this infection. To date, there have been no reports confirming the occurrence of venereal transmission of any of the aforementioned diseases under the circumstances that have been described. This includes possible cases of equine piroplasmosis acquired by the venereal route of exposure. However, there is one report of the finding of equine infectious anemia virus in the semen of a carrier stallion.94 The authors successfully transmitted the infection, not by natural breeding or AI, but following subcutaneous inoculation of a mare with semen from a carrier animal. Although the potential for venereal transmission of equine infectious anemia virus exists, there is no evidence that this has actually occurred under natural or experimental conditions.
Urinary tract infections Any review of equine infectious diseases that can be transmitted by the venereal route should not overlook infective agents that may inhabit the urinary tract and could potentially contaminate the distal reproductive tract of the stallion and the semen at time of ejaculation. The following two diseases serve as examples to illustrate the risk of this occurring: equine leptospirosis caused by Leptospira bratislava95 or L. pomona serovar kennewicki,96 and equine rhinitis A virus infection.97 All three infective agents can set up persistent infection in the kidneys or elsewhere in the urinary system and are shed intermittently or constantly in the urine. Donahue and Williams96 referenced the possibility of transmission of L. bratislava by the venereal route via leptospires in the reproductive tract of infected horses. Under such circumstances, there is the likelihood of contamination of semen and the potential for venereal transmission of the respective etiological agents of these diseases by natural breeding or AI.
Miscellaneous infections A miscellany of infective agents with the potential to be transmitted venereally have been isolated from the external genitalia and semen of both clinically normal and infertile stallions. The significance of these organisms in terms of impairment of fertility in the mare as well as the stallion is not yet well established.9, 98 Among the most frequently encountered of these are Mycoplasma spp., in particular M. equigenitalium, M. subdolum, M. equirhinis, and Acholeplasma. In some situations, these organisms have been associated with endometritis, infertility, and even abortion in mares, and infertility and balanoposthitis in
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stallions,98–100 whereas in other situations they have been found in the absence of any evidence of reproductive dysfunction. Chlamydia spp. have also been demonstrated in the genital tract of both the stallion and the mare. While their presence is not always associated with evident clinical disease, certain species, namely Chlamydophila abortus and Chlamydia psittaci have been detected in cases of abortion, salpingitis, and reduced semen quality in the stallion.101, 102 Aside from the foregoing, various yeasts and fungi, for example Candida spp. and Aspergillus spp., can be opportunistic pathogens where AI is used, especially in mares susceptible to endometritis.9 Equine herpesvirus 1 is another equine infective agent that has been detected in stallion semen based on the findings of two experimental studies103, 104 and one field investigation.105 The epidemiological significance of this finding remains to be established, however, as there have been no reported instances where venereal transmission has been implicated in causing equine herpesvirus-related outbreaks of disease. 104 Furthermore, a survey of 50 stallions seropositive for antibodies to equine herpesvirus 1 failed to detect evidence of the virus in the semen of any of the stallions tested.106
Summary Of the range of infectious diseases that can affect the horse, relatively few have been confirmed or have the potential to be venereally transmissible. Some of these are infections restricted to the reproductive tract whereas others are systemic diseases in which the etiological agent, be it a virus, bacterium, or protozoan, becomes localized and is shed in the semen of the stallion. Changing trends in the international trade in horses and equine germplasm have significantly increased the risk of spread of venereally transmissible as well as numerous other equine infectious diseases. The two most important equine diseases that can be transmitted by natural breeding or artificial insemination are contagious equine metritis and equine viral arteritis, both the subject of considerable international regulation. Of the other conditions known or potentially capable of being disseminated by the venereal route, most are identified with individual cases of infection or limited outbreaks of disease. Of paramount importance in mitigating the risk of spread of infections that are venereally transmissible by coitus or AI is ensuring the freedom of breeding stallion populations from those diseases. Also, every effort should be made to ensure that high standards of hygiene and appropriate biosecurity measures
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are observed in managing and collecting breeding stallions so as to prevent the possibility of indirect transfer of disease agents among them.
References 1. Pottie A. The propagation of influenza from stallions to mares. J Comp Pathol Ther 1888;1:37–8. 2. Clark I. Transmission of pink-eye from apparently healthy stallions to mares. J Comp Pathol Ther 1892;5:261–4. 3. Reeks HC. The transmission of pink-eye from apparently healthy stallions to mares. J Comp Pathol Ther 1901;14:159–62. 4. Reeks HC. The transmission of pink-eye from apparently healthy stallions to mares. J Comp Pathol Ther 1902;18:97–102. 5. Bergman AM. A contribution to the knowledge of the virus carrier in Equine Influenza. Zeitschrift fur Infektionskrankheiten 1913;13:161–74. 6. . Schofield FW. A report of two outbreaks of equine influenza due to virus carriers (stallions). Annual Report of the Department of Agriculture of Ontario, 1937; p. 15. 7. Doll ER, Bryans JT, McCollum WH, Crowe MEW. Isolation of a filterable agent causing arteritis of horses and abortion by mares. Its differentiation from the equine abortion (influenza) virus. Cornell Vet 1957;47:3–40. 8. Timoney PJ, Creekmore L, Meade B, Fly D, Rogers E, King B. In: 2006 Multi-State Occurence of EVA. Proceedings of the 110th Annual Meeting of the United States Animal Health Association, 2006, pp. 354–62. 9. Samper JC, Tibary A. Disease transmission in horses. Theriogenology 2006;66:551–9. 10. Timoney PJ. Aspects of the occurrence, diagnosis and control of selected venereal diseases of the stallion. In: Proceedings of the Stallion Reproduction Symposium, Society for Theriogenology, Baltimore MD, 1998, pp. 76–83. 11. Timoney PJ, McCollum WH. Equine viral arteritis. In: Vasallo J (ed.) Vet Clin N Am Equine Pract 1993; 9:295–309. 12. Timoney PJ. Contagious equine metritis. Comp Immunol Microb 1996;19:199–204. 13. Crowhurst RC. Genital infection in mares. Vet Rec 1977;100:476. 14. Timoney PJ, Ward J, Kelly P. A contagious genital infection of mares. Vet Rec 1977;101:103. 15. Platt H, Taylor CE. Contagious equine metritis. Med Microbiol 1982;1:49–96. 16. Sugimoto C, Isayama Y, Sakazaki R, Kuramochi S. Transfer of Haemophilus equigenitalis Taylor et al. 1978 to the genus Taylorella gen. nov. as Taylorella equigenitalis comb. nov. Curr Microbiol 1983; 9:155–62. 17. Matsuda M, Moore JE. Recent advances in molecular epidemiology and detection of Taylorella equigenitalis associated with contagious equine metritis (CEM). Vet Microbiol 2003;97:111–22. 18. Thoresen SI, Jenkins A, Ask E. Genetic homogeneity of Taylorella equigenitalis from Norwegian trotting horses revealed by chromosomal DNA fingerprinting. J Clin Microbiol 1995;33:233–4. 19. Timoney PJ, O’Reilly PJ, McArdle JF, Ward J. Attempted transmission of contagious equine metritis 1977 to other domestic animal species. Vet Rec 1978;102:152.
20. Timoney PJ, O’Reilly PJ, McArdle JF, Ward J, Harrington AM. Contagious equine metritis: experimental infection in the donkey. Vet Microbiol 1985;10:259–68. 21. Platt H, Atherton JG, Simpson DJ, Taylor CE., Rosenthal RO, Brown DFJ, Wreghitt TG. Genital infection in mares. Vet Rec 1977;101:20. 22. Nakashiro H, Naruse M, Sugimoto C, Isayama Y, Kuniyaou C. Isolation of Haemophilus equigenitalis from an aborted equine fetus. National Institute of Animal Health Quarterly (Tokyo) 1981;21:184–5. 23. Powell DG. Contagious equine metritis. Adv Vet Sci Comp Med 1981;25:161–84. 24. Schluter H, Kuller HJ, Friedrich U, Selbitz HJ, Marwitz T, Beyer C, Ullrich E. Epizootiology and treatment of contagious equine metritis (CEM), with particular reference to the treatment of infected stallions. Praktische Tierarztliche 1991;72:503–11. 25. Wood JLN, Cardwell J, Castillo-Olivares J, Irwin V. Transmission of diseases through semen. In: Samper JC, Pycock J, McKinnon AO (eds) Current Therapy in Equine Reproduction. W.B. Saunders, 2007; pp. 266–74. 26. Platt H, Atherton JG, Simpson DJ. The experimental infection of ponies with contagious equine metritis. Equine Vet J 1978;10:153–9. 27. Timoney PJ, McArdle JF, O’Reilly PJ, Ward J. Infection patterns in pony mares challenged with the agent of contagious equine metritis 1977. Equine Vet J 1978;10:148–52. 28. Timoney PJ, Ward J, McArdle JF. CEM and the foaling mare. Vet Rec 1978;102:246–7. 29. Timoney PJ, Powell DG. Contagious equine metritis – epidemiology and control. J Equine Vet Sci 1988;8:42–6. 30. Timoney PJ. CEM – a continuing threat to international trade. Equine Disease Quarterly 2001;10:2–3. 31. Timoney PJ, Powell DG. Isolation of the contagious equine metritis organism from colts and fillies in the United Kingdom and Ireland. Vet Rec 1982;111:478–82. 32. Witherspoon DM. Contagious equine metritis (CEM) update. In: Proceedings of the Twenty-fifth Annual Convention of the American Association of Equine Practitioners, Miami Beach, Florida, 1979, pp. 375–6. 33. Timoney PJ, O’Reilly PJ, Harrington AM, McCormack R, McArdle JF. Survival of Haemophilus equigenitalis in different antibiotic-containing semen extenders. J Reprod Fertil Suppl 1979;27:377–81. 34. Katz JB, Evans LE, Hutto DL, Schroeder Tucker LC, Carew AM, Donahue JM, Hirsh DC. Clinical, bacteriologic, serologic, and pathologic features of infections with atypical Taylorella equigenitalis in mares. J Am Vet Med Assoc 2000;216:1945–8. 35. Anzai T, Eguchi M, Sekizaki T, Kamada M, Yamamoto K, Okuda T. Development of a PCR test for rapid diagnosis of contagious equine metritis. J Vet Med Sci 1999;61:1287–92. 36. Arata AB, Cooke CL, Jang SS, Hirsh DC. Multiplex polymerase chain reaction for distinguishing Taylorella equigenitalis from Taylorella equigenitalis-like organisms. J Vet Diagn Invest 2001;13:263–4. 37. Bleumink-Pluym N, Houwers DJ, Parlevliet JM, Colenbrander B. PCR based detection of CEM agent. Vet Rec 1993;133:375–6. 38. Chanter N, Vigano F, Collin NC, Mumford JA. Use of a PCR assay for Taylorella equigenitalis applied to samples from the United Kingdom. Vet Rec 1998;143:225–7. 39. Miserez R, Frey J, Krawinkler M, Nicolet J. Identification and diagnosis of Taylorella equigenitalis by a
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DNA amplification method (PCR). Schweiz Arch Tierh 1996;138:115–20. Premanandh J, George LV, Wernery U, Sasse J. Evaluation of a newly developed real-time PCR for the detection of Taylorella equigenitalis and discrimination from T. asinigenitalis. Vet Microbiol 2003;95:229–37. Wakeley PR, Errington J, Hannon S, Roest HI, Carson T, Hunt B, Sawyer H, Heath P. Development of a Taylorella equigenitalis directly from genital swabs and discrimination for Taylorella asinigenitalis. Vet Microbiol 2006;118:247–54. Heath P, Timoney P. Contagious equine metritis. In: O.I.E. Manual for Diagnostic Tests and Vaccines for Terrestrial Animals, 6th edn. Paris: Office International des Epizooties, 2008; pp. 838–44. Mackintosh ME. Bacteriological identification of the causative organism of contagious equine metritits. In: International Symposium on Equine Venereal Diseases organised by the Animal Health Trust, Newmarket, England, September, 1980; pp. 16–17. Ricketts SW. Contagious equine metritis (CEM). Equine Vet Educ 1996;8:166–170. Jones TC, Doll ER, Bryans JT. The lesions of equine viral arteritis. Cornell Vet 1957;47:52–68. Himeur D. Incidence in Morocco of equine infectious anaemia, rhinopneumonitis and viral arteritis. Vet Bull 1976;46:11. McCollum WH, Bryans JT. Serological identification of infection by equine arteritis virus in horses of several countries. In: Proceedings of the Third International Conference on Equine Infectious Diseases. Basel: Karger, 1973; pp. 256–63. Moraillon A, Moraillon R. Results of an epidemiological investigation on viral arteritis in France and some other European and Africa countries. Ann Rech Vet 1978;9:43–54. Paweska JT, Barnard BJ. Serological evidence of equine arteritis virus in donkeys in South Africa. Onderstepoort J Vet Res 1993;60:155–8. National Animal Health Monitoring System. Equine Viral Arteritis (EVA) and the U.S. Horse Industry. Fort Collins, CO: USDA:APHIS:VS, CEAH, National Animal Health Monitoring System, 2000; #N315.0400. Cavanagh D. Nidovirales: a new order comprising Coronaviridae and Arteriviridae. Arch Virol 1997;142:629–33. Timoney PJ, McCollum WH. Equine Viral Arteritis. In: Coetzer K (ed.) Infectious Diseases of Livestock with Special Reference to South Africa, 2nd edn. 2004; pp. 924–32, Oxford University Press, South Africa. Weber H, Beckmann K, Haas L. Fallbericht: Equines Arteritisvirus (EAV) als Aborterreger bei Alpakas? Deut Tierarztl Woch 2006;113:162–3. Timoney PJ. Equine viral arteritis. In: Robinson N, Sprayberry K (eds) Current Therapy in Equine Medicine, 6th edn. St. Louis: Elsevier, 2008; pp. 153–7. Balasuriya UBR, Timoney PJ, McCollum WH, MacLachlan NJ. Phylogenetic analysis of open reading frame 5 of field isolates of equine arteritis virus and identification of conserved and nonconserved regions in the GL envelope glycoprotein. Virology 1995;214:690–7. Stadejek T, Bjorklund H, Bascunana CR, Ciabatti IM, Scicluna MT, Amaddeo D, McCollum WH, Autorino GL, Timoney PJ, Paton DJ, Klingeborn B, Belak S. Genetic diversity of equine arteritis virus. J Gen Virol 1999; 80: 691–9.
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57. McCollum WH, Timoney PJ. Experimental observation on the virulence of isolates of equine arteritis virus. In: Wernery U, Wade JF, Mumford JA, Kaaden O-R (eds) Proceedings of the Eighth International Conference on Equine Infectious Diseases, 1999, pp. 558–9. 58. Del Piero F, Wilkins PA, Lopez JW, Glaser AL, Dubovi EJ, Schlafer DH, Lein DH. Equine viral arteritis in newborn foals: clinical, pathological, serological, microbiological and immunohistochemical observations. Equine Vet J 1997;29:178–85. 59. Neu SM, Timoney PJ, Lowry SR. Changes in semen quality in the stallion following experimental infection with equine arteritis virus. Theriogenology 1992;37: 407–31. 60. Timoney PJ. Equine viral arteritis. AAEP Report, March 2000, p. 7. 61. Mann A. EVA guidelines for breeding a mare to an equine arteritis shedding stallion. In: Proceedings of the 101st Annual Meeting of the United States Animal Health Association, 1997, pp. 259–64. 62. Aurich C, Spergser J, Nowotny N, Rosengarten R, Aurich JE. Prevalence of venereal transmissible diseases and relevant potentially pathogenic bacteria in Austrian Noriker draught horse stallions. Wien Tierarztl Monat 2003;90:124–30. 63. Bowen JM, Tobin N, Simpson RB. Effect of washing on the bacterial flora of the stallion’s penis. J Reprod Fertil 1982;32:41–5. 64. Parlevliet JM, Samper JC. Disease transmission through semen. In: Samper JC (ed.) Equine Breeding Management and Artificial Insemination. Philadelphia: W.B. Saunders, 2000; pp. 133–40. 65. Varner DD. External and internal genital infections of stallions. In: Proceedings Stallion Reproduction Symposium. Baltimore, MD: Society for Theriogenology, 1998; pp. 84–94. 66. Lu K, Morresey PR. Infectious diseases in breeding stallions. Clin Tech Equine Pract 2007;6:285–90. 67. Clement F, Vidament M, Guerin B. Microbial contamination of stallion semen. In: Proceedings of the Sixth International Symposium on Equine Reproduction, Brazil, 1994, pp. 199–200. 68. Clement F, Guerin B, Vidament M, Diemert S, Palmer E. Qualit´e bact`eriologique de la semence d’´etalon. Pratique Veterinaire Equine 1993;25:37–43. 69. Klug E, Sieme H. Infectious agents in equine semen. Acta Vet Scand 1992;88:73–81. 70. Merkt H, Klug E, Gunzel AR. Control of equine venereal diseases, Pseudomonas and Klebsiella. In: International Symposium on Equine Venereal Diseases, 1980, pp. 48–50. Organised by the Animal Health Trust, Newmarket, England. 71. Platt H, Atherton JG, Orskov J. Klebsiella and Enterobacter organisms isolated from horses. J Hygiene 1976;77:401–408. 72. Vaissaire J, Plateau E, Collobert-Laugier C. Importance du microbisme dans les spermes d’etalons. B Acad Vet France 1987;60:165–75. 73. Archer L, Wade JF. Codes of Practice on Contagious Equine Metritis (CEM), Equine Viral Arteritis (EVA), Equine Herpesvirus (EHV) and Guidelines on Strangles. Newmarket: Horserace Betting Levy Board and R&W Publications, 2003. 74. Timoney PJ. Incidence of certain equine venereal diseases in Ireland for 1978/1979. International Symposium
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on Equine Venereal Diseases organised by the Animal Health Trust, Newmarket, Englang, 1980; pp. 18–19. ¨ Weiss R, Bohm KH, Merkt H, Klug E, Heuser H. Studies on the flora of the genital and nasal mucosa of horses, especially of stallions with bacterial pathogens important in horse breeding and with particular reference to klebsiella. III. Serologic studies on klebsiellas. Berl Munch Tierarztl 1976;89:193–6. Kikuchi N, Iguchi I, Hiramune T. Capsule types of Klebsiella pneumoniae isolated from the genital tract of mares with metritis, extra-genital sites of healthy mares and the genital tract of stallions. Vet Microbiol 1987;15:219–28. Madsen M, Christensen P. Bacterial-flora of semen collected from Danish warmblood stallions by artificial vagina. Acta Vet Scand 1995;36:1–7. Gierczynski R, Jagielski M, Rastawicki W, Kauzewski S. Multiplex-PCR assay for identification of Klebsiella pneumoniae isolates carrying the cps loci for K1 and K2 capsule biosynthesis. Pol J Microbiol 2007;56:153–6. Atherton JG, Pitt TL. Types of Pseudomonas aeruginosa isolated from horses. Equine Vet J 1982;14:329–32. Rossdale PD. Control of equine venereal diseases, Pseudomonas and Klebsiella. In: International Symposium on Equine Venereal Diseases, organised by the Animal Health Trust, Newmarket, England, 1980; pp. 48–50. Aurich C, Spergser J. The influence of genitally pathogenic bacteria on semen motility in cooled-stored stallion semen. In: Alm H (ed.) Proceedings of a Workshop of the International Equine Gamete Group, 2006, pp. 10–12. Allen WE, Newcombe JR. Aspects of genital infection and swabbing techniques in the mare. Vet Rec 1979;104:228–31. Pascoe RR. Effect of equine coital exanthema on the fertility of mares covered by stallions exhibiting the clinical disease. Aust Vet J 1981;57:111–14. Bitach V. Cases of equine coital exanthema in Denmark. Acta Vet Scand 1972;13:281–3. Katz JB. Dourine. In: O.I.E. Manual for Diagnostic Tests and Vaccines for Terrestrial Animals, 6th edn. Paris: Office International des Epizooties, 2008; pp. 845–51. Claes F, Agbo EC, Radwanska M, Te Pas MFW, Baltz ¨ T, De Waal DT, Goddeeris BM, Claassen E, Buscher P. How does Trypanosoma equiperdum fit into the Trypanozoon group? A cluster analysis by RAPD and multiplex-endonuclease genotyping approach. Parasitology 2003;126:425–31. Kumba FF, Claasen B, Petrus P. Apparent prevalence of dourine in the Khomas region of Namibia. Onderstepoort J Vet Res 2002;69:295–8. Parkin BS. The demonstration and transmission of the South African strain of Trypanosoma equiperdum of horses. Onderstepoort J Vet Sci Anim Ind 1948;23:41–58. Caporale VP, Battelli G, Semproni G. Epidemiology of dourine in the equine population of the Abruzzi Region. Zentralblatt fur Veterinarmedizin Reihe B 1980;27: 489–98. Robinson EM. Dourine infection in young equines. Onderstepoort J Vet Sci Anim Ind 1948;23:39–40.
91. Jang SS, Donahue JM, Arata AB, Goris J, Hansen LM, Earley DL, Vandamme PAR, Timoney PJ, Hirsh DC. Taylorella asinigenitalis sp. nov., a bacterium isolated from the genital tract of male donkeys (Equus asinus). Int J Syst Evol Micr 2001;51:971–6. ¨ 92. B˚averud V, Nystrom C, Johansson KE. Isolation and identification of Taylorella asinigenitalis from the genital tract of a stallion, first case of natural infection. Vet Microbiol 2006;116:294–300. 93. Duquesne F, Pronost S, Laugier C, Petry S. Identification of Taylorella equigenitalis responsible for contagious equine metritis in equine genital swabs by direct polymerase chain reaction. Res Vet Sci 2007;82:47–49. 94. Stein CD, Mott LO. Studies on congenital transmission of equine infectious anemia. Vet Med 1942;37:370–7. 95. Ellis WA. Equine leptospirosis. In: Wernery U, Wade JF, Mumford JA, et al. (eds) Equine Infectious Diseases 8: Proceedings of the Eighth International Conference, Newmarket: R&W Publications Ltd, 1999; pp. 155–65. 96. Donahue JM, Williams NM. Emergent causes of placentitis and abortion. In: Vasallo J (ed.) Veterinary Clinics of North America Equine Practice. 2000. 16: pp. 443–56. 97. McCollum WH, Timoney PJ. Studies on the seroprevalence and frequency of equine rhinoviruses – I and II infection in normal horse urine. In: Proceedings of the Sixth International Conference on Equine Infectious Diseases, Cambridge, 1991, pp. 83–7. 98. Moorthy ARS, Spradbrow PB, Eisler MED. Isolation of mycoplasmas from the genital tract of horses. Aust Vet J 1977;53:167–9. 99. Heitmann J, Kirchhoff K, Petzoldt K, Sonnenschein B. Isolation of acholeplasmas and mycoplasmas from aborted horse fetuses. Vet Rec 1979;104:350. 100. Kirchhoff H, Naglic T, Heitmann J. Isolation of Acholeplasma laidlawii and Mycoplasma equigenitalium from stallion semen. Vet Microbiol 1979;4:177–9. 101. Herfen K, Jager C, Wehrend A. Genital chlamydial infection in mares and their clinical significance. Reprod Domest Anim 1999;34:20. 102. Wittenbrink MM. Aetiological significance of chlamydial infections in equine reproductive disorders? Pferdeheilkunde 1999;15:538–41. 103. Blunden AS, Smith KC, Whitwell KE, Dunn KA. Systemic infection by equid herpesvirus-1 in a Grevy’s zebra stallion (Equus grevyi) with particular reference to genital pathology. J Comp Pathol 1998;119:485–93. 104. Tearle JP, Smith KC, Boyle MS, Binns MM, Livesay GJ, Mumford JA. Replication of equid herpesvirus-1 (EHV-1) in the testes and epididymides of ponies and venereal shedding of infectious virus. J Comp Pathol 1996;115:385–97. 105. Carvalho R, Passos LMF, Oliveira AM, Henry M, Martins AS. Detection of equine herpesvirus 1 DNA in a single embryo and in horse semen by polymerase chain reaction. Arq Bras Med Vet Zoo 2000;47:351–9. 106. Brown JA, Mapes S, Ball BA, Hodder AD, Liu IK, Pusterla N. Prevalence of equine herpesvirus-1 infection among Thoroughbreds residing on a farm on which the virus was endemic. J Am Med Assoc 2007;231:577–80.
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Chapter 322
Import and Export of Frozen Semen Elizabeth S. Metcalf
Introduction The transmission of potentially pathogenic organisms through equine semen has long been recognized as an important means of dissemination of disease. With the gaining popularity of artificial insemination and international shipment of semen over the past two decades, the unintentional introduction of agents into naive populations of equine breeding stock can threaten the entire industry. Venereal diseases previously contained on a single farm, or in an area, are now capable of spreading rapidly over great distances. The widespread outbreak of such diseases may not be traced back to their origins until devastating economic and health consequences have ensued. Therefore, vigilant biosecurity measures are necessary to prevent the venereal transmission of disease in equid populations, not only in the management of the naturally breeding stallion, but also in the transfer of semen and in the animal-origin, biological components of semen extenders as well. With the advancing sophistication of sensitive and specific tests for venereal equine organisms, rapid and precise identification of disease-carrying horses will favor greater control of potential spread. Other chapters offer greater detail with respect to specific disease transmission (see Chapters 245, 246 and 247). The purpose of this chapter is primarily regulatory in nature, citing specific diseases that have the potential to be transmitted through semen, but leaving detailed diagnosis and prevention to be discussed in other chapters.
Potential pathogens Although many organisms have been isolated from stallion semen, not all cause disease. Often, the sig-
nificance of their presence is unknown. Those of greatest concern include certain pathogenic bacteria, viruses, and protozoa. The bacterial pathogen of most widespread concern is Taylorella equigenitalis, the recognized causative agent for contagious equine metritis (CEM). Some strains of an atypical isolate, Taylorella asinigenitalis, may be capable of eliciting a similar clinical response to that of T. equigenitalis, but the evidence is incomplete.1 Most countries require extensive testing of the stallion and/or the semen for this highly contagious, Gram-negative coccobacillus. Other pathogenic bacteria include certain strains of Pseudomonas aeruginosa and Klebsiella pneumoniae, and less commonly, Streptococcus zooepidemicus.2 These organisms not only decrease fertility in mares susceptible to endometritis,3 but they have also been shown to be detrimental to semen quality and, therefore, fertility of the stallion.4 Of equine viruses identified in stallion semen, equine arteritis virus (EAV) and equine herpesvirus 3 (EHV-3) represent pathogenic viruses capable of venereal transmission. Both have been associated with high morbidity in sporadic outbreaks. Equine arteritis virus is harbored in the epididymis, vas deferens, ampullae, prostate, and seminal vesicles of the stallion. Although the virus is tightly adhered to the spermatozoa and readily survives storage in cooled or cryopreserved environments,5 without loss of infectivity,6 the vast majority of viral particles are free in the seminal plasma. Infective particles of EHV-3, on the other hand, are more likely found in the vesicular lesions on the external genitalia. Although herpesviral particles may in theory be found in the semen, transmission between horses is most likely to be achieved by coitus rather than artificial insemination. Although less likely, EHV-3 particles can be spread by contaminated fomites as well.
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Many countries that permit the importation of stallion semen require serological testing of the stallion for viruses that have never been identified in semen, nor have been associated with venereal transmission of infection (equine infectious anemia, vesicular stomatitis, equine encephalomyelitides, etc.). Furthermore, permits often require that a stallion has not been housed in an area in which certain viral diseases have been reported, even though the same viral agents have never been found in semen. Likewise, concern regarding the sexual transmission of protozoal diseases has led to investigation of organisms capable of equine systemic disease but often without evidence of contamination of semen (i.e., piroplasmosis, borreliosis, etc.). An important exception is the protozoal disease, dourine, caused by Trypanosoma equiperdum. This ultimately fatal, sexually transmitted disease of horses is most often seen in horses in Africa, the Middle East, and South and Central America; however, sporadic cases have recently been reported in Asia and Germany.7 Finally, venereal spread of other potentially pathogenic organisms found on either the external genitalia, in the ejaculate, or in the extender, although perhaps less important, cannot be ignored. Certain species of yeasts and fungi, such as Aspergillus spp and Candida spp, can serve as potential pathogens in susceptible mares. Chlamydia spp and Mycoplasma spp have been isolated from semen from both clinically normal and subfertile stallions.8, 9 In addition to venereal spread, these organisms may not only affect equine fertility, but other systems and other species as well. Components of extenders may contain specific pathogens that are pathogenic to other species. For instance, eggs, the yolk of which is often added to extenders as a cryoprotectant of sperm, may be contaminated with Newcastle disease virus and Salmonella spp. Added milk products may originate from a country affected by foot-and-mouth disease.
Prevention The first step in prevention of transmission of disease through shipped semen is to recognize and control the routes of transmission that represent the highest risks – respiratory, venereal, transplacental, and via contaminated fomites. Many countries have introduced strict requirements not only with respect to the collection and processing of the semen, but also the physical layout of the breeding farm and housing of the breeding stock. Implementation of biosecurity measures begins with design of the breeding farm. It should be constructed to minimize contact with either new arrivals, animals exposed to potential pathogens or persistently infected stallions. Not only should new arrivals be subjected to strict quarantine
isolation for 2–3 weeks upon arrival on a farm, but breeding stallions should undergo routine breeding examinations, the frequency of which may depend on such variables as their performance schedule, use for live cover, history of infection, etc. Personnel handling breeding stock and equipment must be properly trained in the appropriate hygiene to prevent contamination of semen and control disease spread. They should wear clothing and boots that are disposable or capable of disinfection through laundering or footbaths. Potential fomites of infection, including buckets, artificial vaginas and covers, phantoms, leadropes, halters, blankets, sponges, etc., should not be shared between horses. The transrectal probe of an ultrasound machine should be cleaned and disinfected between examinations. If disposable equipment is unavailable, proper disinfection with R an approved product (2% bleach, Virkon S ; Antec International, Sudbury, UK) of all equipment that is in contact with breeding stock should be put into customary practice. Also, personnel need to remain cognizant not only of the physical well-being, but the mental status of the breeding horse as well, for the role that transportation or other stressors play in the ‘reactivation’ of latent disease may also pose a significant threat in potential spread of certain diseases such as equine coital exanthema. Lastly, since vectors, both insect and rodent, are capable of spreading disease between horses, a pest control program to minimize their impact should also be instigated on a breeding farm. Laboratory testing and identification of affected breeding stock is key to successful containment of disease. In a shipped semen program, the last resort for the prevention of disease transmission likely resides in the laboratory diagnosis and control of potential pathogens. The advent of new and improved diagnostic tools serves to better identify pathogenic organisms as well as carriers of disease. Differentiation between commensal, non-pathogenic organisms and those that cause disease is essential in controlling spread. Further investigation into the presence and pathogenicity of infectious material (EHV-1, EIA, etc.) in the ejaculate is also necessary. Attempts to reduce or eliminate transmission of infectious disease include the use of antibiotics (administered systemically, topically, or as a component of the semen extender), vaccination (if available), and/or other agents to either reduce or eliminate microbial contamination of semen. Aiming to prevent disease transmission by ‘disinfecting’ semen, researchers have examined various treatments of farm animal and human semen that include ozone exposure, dilution washing, photosensitization, transient acidification, density Percoll gradient centrifugation,
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Import and Export of Frozen Semen trypsin (and other enzymes) addition, and immunoextenders that contain neutralizing antibodies.10 Although some of these antiviral agents look promising, there are limitations either in incomplete elimination of the virus or detrimental effect on fertility of the sperm. Antimicrobial decontamination of spermatozoa represents a necessary area of future research. Lastly, storage of semen is often regulated as well. Many countries require that frozen semen is stored in secure, double-locked containers. The liquid nitrogen must not have been in contact previously with other biological material. Shippers must be new or properly disinfected with approved agents (vapor shippers cannot be adequately disinfected and must be new). Even with these precautions in place, liquid nitrogen and its vapor represents an efficient medium not only for the cryopreservation of sperm,
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but also fungi, yeasts, bacteria, and viruses that may be present in the environment.11 Again strict sanitary control by personnel of the environment, and equipment will minimize the risk of disease spread.
Transporting semen There are a few commercial companies that arrange the details for shipment of stallion semen, whether cooled or frozen. Although expensive, they provide an excellent service for the importer as well as the veterinarian shipping the semen. They often make arrangements for the shipping tanks, all transportation, and importation requirement details in the country of destination. An international composite of current importation requirements is under constant update and revision (Table 322.1). In the United States, these
Table 322.1 Current importation requirements Country Afghanistan Albania Algeria Andorra Angola Antigua & Barbuda Arab Emirates Argentina Armenia Australia Azerbaijan Bahamas Bahrain Bangladesh Barbados Belarus Belize Benin Bhutan Bolivia Bosnia & Herzegovina Botswana Brazil Brunei Darussalam Burkina Faso Burma Burundi Cambodia Cameroon Canada Cape Verde Central African Republic Chad Chile China
Permit
EIA
EVA
CEM
X
X
X
X
X
X
X
X
X
X
X
EEE
WEE
VEE
EHV-1
FMD
VVND
VS
WNV
X
X
(Continued)
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Table 322.1 (Continued) Country
Permit
EIA
EVA
CEM
X
X
X
X
X
X
X
EEE
WEE
VEE
EHV-1
FMD
VVND
VS
WNV
Colombia Comoros Congo Congo, Democratic Republic Costa Rica Cote d’Ivoire Croatia Cuba Djibouti Dominica Dominican Republic Ecuador East Timor Egypt El Salvador England Equatorial Guinea Eritrea Ethiopia EU (Austria, Belgium, Bulgaria, Cyprus, Czech Republic, Denmark, Estonia, Finland, France, Germany, Greece, Hungary, Ireland, Italy, Latvia, Lithuania, Luxembourg, Malta, Netherlands, Poland, Portugal, Romania, Slovakia, Slovenia, Spain, Sweden, United Kingdom) Falkland Islands Fiji Gabon Gambia Georgia Ghana Great Britain Grenada Guatemala Guinea Guinea-Bissau Guyana Haiti Honduras Iceland India Indonesia Iran Iraq Israel Jamaica Japan Jordan (Continued)
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Import and Export of Frozen Semen
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Table 322.1 (Continued) Country
Permit
EIA
EVA
CEM
X
X
X X
X X
X X
X
X
X
X
X
EEE
WEE
VEE
X
X
X
X
X
EHV-1
FMD
VVND
VS
WNV
Kazakhstan Kenya Kiribati Korea, North Korea, South Kuwait Kyrgyzstan Laos Lebanon Lesotho Liberia Libya Liechtenstein Macedonia Madagascar Malawi Malaysia Maldives Mali Marshall Islands Mauritania Mauritius Mexico Micronesia Moldova Monaco Mongolia Montenegro Mozambique Myanmar Namibia Nauru Nepal New Zealand Nicaragua Niger Nigeria Norway
X
X
Oman Pakistan Palau Palestinian State Panama Papua New Guinea Paraguay Peru Philippines, The Qatar Russia Rwanda (Continued)
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Table 322.1 (Continued) Country
Permit
EIA
EVA
X X
X
X
CEM
EEE
WEE
X
X
VEE
EHV-1
FMD
VVND
X
X
VS
St. Kitts & Nevis St. Lucia St. Vincent & The Grenadines Samoa San Marino Sao Tome & Principe Saudi Arabia Scotland Senegal Serbia Seychelles Sierra Leone Singapore Solomon Islands Somalia South Africa Sri Lanka Sudan Suriname Swaziland Switzerland Syria Taiwan Tajikistan Tanzania Thailand Togo Tonga Trinidad & Tobago Tunisia Turkey Turkmenistan Tuvalu Uganda Ukraine United States Uruguay Uzbekistan Vanuatu Vatican City Venezuela Vietnam Western Sahara Wales Yemen Zaire Zambia Zimbabwe
X X
X
WNV
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Import and Export of Frozen Semen requirements may be found on the website http:// www.aphis.usda.gov/vs/ncie/iregs/animals/. Many countries list strict requirements for quarantine and testing; others post none. Because regulations change frequently, it is most beneficial to obtain a permit to import semen from the importing country before beginning the cryopreservation of the semen or quarantine of the stallion. Even when one complies with the published regulations for the freezing, testing, and/or quarantine processes, it is important to realize that any country may close its borders to importation of semen at any time and for any reason. Furthermore, the officials of the importing country have the right to dispose of the semen rather than return it. Although infrequently encountered, a situation such as this can be costly and frustrating.
Conclusion Venereal transmission represents an important and significant means of dissemination of disease in equids. With the rapidly expanding popularity of artificial insemination and the success of shipped semen, both cooled and frozen, the potential for rapid and widespread disease transmission grows worldwide as an emergent threat. Furthermore, as the ease of movement of horses between states, countries, and hemispheres continues to increase, so follows the potential for spreading disease, through venereal transmission or other routes. Federal agencies that regulate the movement of breeding stock or semen have, in many countries, failed to keep pace with the growing dissemination of sexually transmitted diseases. Many countries do not regulate the importation of semen. Presently, the United States requires testing only for CEM. As a result, the responsibility for minimizing the threat not only to our equine industries but to other species (human zoonoses notwithstanding) rests in the hands of veterinarians. A cohesive, collaborative effort between all parties involved in the shipment of semen must be forged. Veterinarians need to ensure testing for disease even though law or registry do not require
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it. It also behooves us to educate governing bodies regarding proven and emerging threats to our breeding stock. These entities must strongly recommend that stallion owners test, identify, and vaccinate (if possible) their horses to lessen the chances that their horse becomes infected and/or becomes a carrier of sexually transmissible disease.
References 1. Katz JB, Evans LE, Hutto DL, Schroeder-Tucker LC, Carew AM, Donahue JM, Hirsh DC. Clinical, bacteriologic, serologic, and pathologic features of infections with atypical Taylorella equigenitalis in mares. J Am Vet Med Assoc 2000;216(12):1945–8. 2. Wood JL, Cardwell J, Castillo-Olivares J, Irwin V. Transmission of diseases through semen. In: Samper J, Pycock JF, McKinnon AO (eds) Current Therapy in Equine Reproduction. St Louis: Saunders, 2007; pp. 266–74. 3. Samper J, Tibary A. Disease transmission in horses. Theriogenology 2006;66(3):551–9. 4. Aurich C, Spergser J. Influence of genitally pathogenic bacteria and gentamicin on motility and membrane integrity of cooled-stored stallion spermatozoa. Anim Reprod Sci 2006;94:117–20. 5. Raz T, Corrigan M, Loomis P, Card C. Effects of equine arteritis-positive semen on mare fertility. Anim Reprod Sci 2006;94:112–14. 6. Holyoak GR, Balasuriya UBR, Boaddus CC, Timoney PJ. Equine viral arteritis: current status and prevention. Theriogenology 2006;70(3):403–14. ¨ 7. Claes F, Buscher P, Touratier L, Goddeeris BM. Trypanosoma equiperdum: master of disguise or historical mistake? Trends Parasitol 2005;21(7):316–21. 8. Spergser J, Aurich C, Aurich J, Rosengar R. High prevalence of mycoplasmas in the genital tract of asymptomatic stallions in Austria. Vet Microbiol 2002;87(2): 119–29. 9. Baczynska A, Fedder J, Schougaard H, Christiansen G. Prevalence of mycoplasmas in the semen and vaginal swabs of Danish stallions and mares. Vet Microbiol 2007;121(1–2,3):138–43. 10. Bielanski A. Disinfection procedures for controlling microorganisms in the semen and embryos of humans and farm animals. Theriogenology 2007;68(1):1– 22. 11. Grout BWW, Morris GJ. Contaminated liquid nitrogen vapour as a risk factor in pathogen transfer. Theriogenology 2009;71(7):1079–82.
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Chapter 323
Low Dose Insemination Lee H.A. Morris and Sara K. Lyle
Introduction The generally recommended minimum number of spermatozoa required for conventional artificial insemination in the mare is in excess of 200 × 106 progressively motile spermatozoa. Results of a survey of 20 years of artificial insemination in France with frozen-thawed semen revealed that insemination of mares with less than 750 × 106 total spermatozoa per cycle resulted in significantly lower per cycle pregnancy rates than those obtained with doses of spermatozoa in excess of 750 × 106 spermatozoa.1 However, the requirement for such a large number of spermatozoa restricts the number of potential breedings from a limited source of frozen semen. Therefore, lowering the insemination dose becomes attractive to the stallion or semen owner if fertility can be maintained. Insemination techniques such as deep uterine and hysteroscopic insemination that have been designed to use significantly fewer spermatozoa are reviewed in this chapter. A number of studies have demonstrated that ultrasoundguided or rectally guided deep uterine insemination of 5 × 106 fresh spermatozoa can produce satisfactory pregnancy rates. The use of hysteroscopic insemination enables the insemination dose to be successfully reduced to 1 × 106 fresh or 3 × 106 frozen-thawed spermatozoa. Further reduction of the insemination dose is possible and satisfactory pregnancy rates can also be achieved by surgical oviductal insemination of mares with as few as 2 × 105 fresh spermatozoa.2 Despite the deposition of billions of spermatozoa into the mare’s reproductive tract at the time of mating, all but one of these spermatozoa are excluded by the time of fertilization. This chapter will review the ways in which we are able to exploit this physiological feat and inseminate mares with very low numbers of spermatozoa to achieve acceptable pregnancy rates. The refinement of these insemination techniques will allow us to maximize the use of low
doses of frozen-thawed semen, use new technology such as sex-preselection of spermatozoa and to conserve and utilize epididymal spermatozoa at the time of castration or death of a stallion.
Sperm transport in the mare During normal mating, a fertile stallion will ejaculate between 2 and 15 × 109 spermatozoa directly into the mare’s uterus.3 The initial distribution of these spermatozoa within the uterus will be influenced by a combination of uterine contractions, estrous uterine fluids, ejaculate volume, and sperm motility.4 Indeed, these interactions provide a selective gradient for the spermatozoa as they progress towards the utero-tubal junction. Troedsson et al.5, 6 demonstrated that interactions between viable and non-viable spermatozoa with seminal plasma components results in the retention of viable spermatozoa within the uterus and the expulsion of non-viable spermatozoa from the uterus. This explains how Scott et al.7 observed that not only does a subpopulation of the ejaculate reach the uterine side of the papillae of each utero-tubal junction, but that more than 90% of the spermatozoa that reach the oviducts are morphologically normal. Consequently, despite insemination of mares with ejaculates containing more than 85% morphologically abnormal spermatozoa, more than 90% of the spermatozoa flushed from the oviducts were found to be morphologically normal. The remainder of the ejaculate is effectively evacuated from the uterus through the very relaxed estrous cervix.4 Brinsko et al.8 obtained satisfactory pregnancy rates in mares when their uteri were lavaged more than 4 h after insemination, thus illustrating the time required for the fertilizing population of spermatozoa to enter the oviduct as described by Bader.9 Once in the oviduct, the spermatozoa appear to be protected from the post-mating infiltration of
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Low Dose Insemination neutrophils and other inflammatory cells into the uterus5 and various intrauterine post-breeding treatments that otherwise would be damaging.8
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The methods of low dose insemination Timing of insemination
Recommended doses of spermatozoa In order to achieve satisfactory pregnancy rates per cycle in mares, the minimum recommended number of spermatozoa contained within a conventional insemination dose is generally > 300 × 106 progressively motile spermatozoa for fresh10–13 and > 200 × 106 progressively motile spermatozoa for frozen semen (reviewed by Pickett et al., 2000).14 Historically, it has been shown that conventional insemination of mares with ≤ 100 × 106 spermatozoa results in unsatisfactory pregnancy rates per cycle.15, 16 Provided the inseminate contains an adequate number of spermatozoa, acceptable pregnancy rates have been reported after conventional insemination of mares with as little as 0.6 mL17 or as much as 120 mL.18 Nevertheless, it has also been demonstrated that pregnancy rates in mares17, 19 and sperm survival rates in vitro are optimized when equine spermatozoa are maintained at a concentration of 25–50 × 106 /mL.20 Deposition of high numbers of spermatozoa into the mare’s reproductive tract are intended to compensate for some of the variability in fertility observed amongst males such as has been observed in other species.21 However, even with natural mating in intensively managed Thoroughbreds, where the entire ejaculate is deposited into the mare’s reproductive tract, variation persists amongst stallions in their per cycle pregnancy rates.22, 23 With the introduction of each additional layer of technology from the insemination of mares with high numbers of fresh (reviewed by Pickett et al., 2000)14 or frozen-thawed spermatozoa, reported pregnancy rates per cycle continue to range from only 30% to 85%.1,24–27 Consequently, 1–2 estrous cycles are generally required to achieve acceptable end-of-season pregnancy rates with natural mating or insemination with fresh or frozen semen. To further overcome variability in stallion fertility and to enable us to exploit low numbers of spermatozoa available after freezing or sex-preselection of spermatozoa, both deep uterine28 and hysteroscopic insemination techniques29 have been investigated. Brinsko (2006)30 discusses the benefits of these techniques in both fertile and subfertile stallions. The advantages of reducing the distance the spermatozoa have to travel in the uterus by depositing them closer to the oviduct have been illustrated in sheep31–34 and humans.35
The optimal timing for insemination will be determined by the type of semen available. Furthermore, when a very low dose of semen is available then the timing of insemination is of paramount importance. Typically for low dose insemination a single fixed time of insemination during estrus is required as only one low dose of semen may be available. Given these limitations, it is ideal if the low dose of semen is deposited into the mare’s uterus prior to ovulation to avoid compromising an early diestrous uterus and to ensure that there is a functional reservoir of spermatozoa in the caudal isthmus of the oviduct at the time of the arrival of the oocyte in the caudal isthmus after ovulation. For low doses of fresh semen, satisfactory per cycle conception rates have been obtained when the deep uterine or hysteroscopic insemination procedure is performed at the time of the administration of human chorionic gonadotrophin (hCG; Chorulon, Intervet) and ovulation occurred within the next 48 h.28, 29, 36 Morris et al.37 reported excellent per cycle conception rates after a single hysteroscopic insemination with a low dose of frozen semen at 32 h after administration of hCG. Deep uterine insemination For deep uterine insemination, the low dose of semen is deposited in the most anterior aspect of the uterine horn (Figure 323.1). This is a relatively simple insemination procedure for which atraumatic flexible catheters have been developed by Minitube (Germany) and IMV (France). Immediately prior to insemination of the mare her tail is wrapped and her perineal region and vulva is cleaned with dilute 2% iodine or chlorhexidine scrub. The low dose of frozen semen, typically 1 × 0.5 mL straw, is thawed, the sealed end of the straw is cut and the straw inserted into the warm pipette, cut end first. The accompanying stylet is placed inside the pipette and lodged carefully at the end of the straw containing the cotton plug. This pipette is then passed through the mare’s cervix and guided up the uterine horn ipsilateral to the side of ovulation. The pipette may be guided along the uterine horn either ‘per rectum’ or by pulling the cervix caudally along the pipette and at the same time engaging the pipette in the lumen of the uterine horn ipsilateral to the side of ovulation. Once the pipette is placed at the cranial tip of the uterine horn the semen is expelled carefully from the straw, the uterus elevated to facilitate the migration
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Site of hysteroscopic insemination
Site of deep uterine insemination
Site of conventional insemination (a)
(b) Figure 323.1 (a) Schematic diagram of the mare reproductive tract (b) hysteroscopic deposition of low volume semen directly onto the uterotubule junction.
of the semen towards the utero-tubal junction and the pipette removed slowly to minimize withdrawal of any semen. Hysteroscopic insemination Hysteroscopic insemination allows the low dose of semen to be deposited directly onto the utero-tubal junction at the gateway from the uterus to the oviduct (Figure 323.1a). The mare should be prepared in a
similar manner to that described for deep uterine insemination. Prior to thawing the semen, the catheter is loaded into the working channel of the videoendoscope. The semen is then thawed and the small volume of inseminate is aspirated into the distal end of the catheter. Once loaded, the catheter is withdrawn into the working channel of the endoscope and then the endoscope is passed through the mare’s cervix and guided along the lumen of the uterine horn ipsilateral to impending ovulation. A minimal amount of air may be required to insufflate the uterus to facilitate passage of the endoscope without damaging the endometrium. Once the papilla of the utero-tubal junction is visualized, the catheter is extruded several centimeters from the endoscope and the inseminate is deposited directly onto the papilla (Figure 323.1b). After depositing the semen onto the papilla the endoscope is withdrawn caudally and air is evacuated from the uterus. Fresh semen Vazquez et al.38 investigated the use of hysteroscopic insemination in an attempt to bypass the uterine transport phase of sperm migration and to increase the fertility of subfertile stallions. In their study, three pregnancies were obtained when 10 mares were inseminated hysteroscopically by depositing only 20 µL of inseminate, which contained 4 × 106 spermatozoa, onto the papilla of the utero-tubal junction. Six of the mares developed post-breeding endometritis after insemination despite the low number and volume of spermatozoa used. In other studies performed at the same time, Manning et al.39 attempted to cannulate the oviduct hysteroscopically to deposit 1 or 10 × 106 spermatozoa suspended in 0.16–0.25 mL of diluent into the oviduct. The pregnancy rates were disappointingly low, 22% and 30%, respectively, and may reflect the negative effect of excessive oviducal insemination volumes or it may be that bypassing the selective barrier of the utero-tubal junction was detrimental to fertility. Morris et al.29 inseminated mares hysteroscopically with 10, 5, 1, 0.5, 0.1, or 0.001 × 106 motile spermatozoa suspended in 0.03–0.15 mL Tyrode’s medium. The fresh semen was diluted in a skim milkglucose extender40 and then centrifuged through a discontinuous Percoll (Sigma-Aldrich) density gradient and deposited hysteroscopically onto the papilla of the utero-tubal junction without any attempt to catheterize the os of the papilla. Pregnancy rates per cycle of > 60% were achieved in the mares inseminated with 10, 5 or 1 × 106 spermatozoa. However, hysteroscopic insemination with 0.5 × 106 motile spermatozoa began to approach the limit for
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Low Dose Insemination successful fertilization, resulting in pregnancy rates of less than 30%. Manning et al.39 observed a negative effect of large insemination volumes (0.16–12 mL) on fertility, whereas Morris et al.29 noticed no such effect of insemination volume (30–150 µL) on the fertility of low dose hysteroscopic insemination. They demonstrated that it was the number of spermatozoa in the inseminate, and not the volume of semen, that was important in determining conception when low insemination volumes were used. Lindsey et al.36 demonstrated better fertility using the hysteroscopic insemination method (50%) when compared with deep uterine insemination (0%) of small volumes of inseminate containing only 5 × 106 spermatozoa. In an attempt to improve the efficacy of the deep uterine insemination method, Nie and Johnson41 inseminated mares with 1 × 106 spermatozoa that had been filtered through a glass wool/Sephadex column. Their pregnancy results were disappointingly low (7–19%) and this may have been due to less than 1 × 106 spermatozoa actually reaching the utero-tubal junction. Similarly low pregnancy results were achieved when less than 1 × 106 spermatozoa were deposited directly onto the uterotubal papilla during hysteroscopic insemination.29 Since these initial studies, Brinsko et al.42 compared deep uterine insemination with hysteroscopic insemination. In this study, mares were inseminated 24 h after administration of hCG, with 5 × 106 cooled spermatozoa suspended in 200 µL of diluent that had not been processed through a density gradient. No statistically significant differences were observed between the pregnancy rates of the mares inseminated either hysteroscopically (13/21, 62%) or by using the deep uterine insemination method (10/20, 50%). Frozen-thawed semen Morris et al.37 observed satisfactory pregnancy rates (> 60% per cycle) when mares were inseminated with 14 × 106 frozen-thawed spermatozoa at a fixed time after administration of hCG, by either hysteroscopic or conventional insemination. However, once the numbers of spermatozoa were reduced to 3 × 106 , there was a distinct advantage in depositing the inseminate directly onto the utero-tubal junction rather than into the uterine body. The implementation of hysteroscopic insemination for commercially available frozen-thawed semen may increase the potential efficiency of frozen semen from particular stallions. In a commercial context, a ‘low dose’ of semen usually consists of 1–2 × 0.5 mL straws of frozen semen, containing 50–100 × 106 spermatozoa. Despite these relatively high numbers of spermatozoa compared with the doses used in a re-
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search setting, variability in the fertility of frozenthawed semen amongst commercially available stallions is evident.22 Using deep uterine insemination to overcome fertility problems It is apparent that bathing the utero-tubal junction with a population of spermatozoa which exhibit a high proportion of uncompensable defects,21 that ultimately prevent binding of spermatozoa to an oocyte, will not enhance fertility. If the sperm defects are compensable, it may be possible to improve the fertility of some semen from subfertile stallions by reducing the distance the semen has to travel, as discussed by Brinkso.30 However, in an attempt to overcome the poor quality of some frozen-thawed spermatozoa, Alvarenga and Le˜ao43 processed frozen-thawed semen through density gradients to remove those damaged by the cryopreservation process. This treatment did not improve the fertilizing capacity of the selected low doses of frozen-thawed spermatozoa. There are also mare factors to consider, as illustrated by Sieme et al.44 who found an interaction between mare fertility and insemination technique, such that hysteroscopic insemination produced lower pregnancy rates in problem mares than normal mares and conventional insemination produced higher pregnancy rates in problem mares than normal mares. Epididymal spermatozoa The first pregnancy using frozen-thawed stallion spermatozoa was reported in a mare inseminated with epididymal spermatozoa.45 However, in sheep the in vivo fertility of cauda epididymal ram spermatozoa is lower than that of ejaculated spermatozoa unless it is deposited surgically in the region of the utero-tubal junction.46 The lower fertility of epididymal spermatozoa may be due to variations in the cell surface characteristics depicted by different chlortetracycline staining patterns47 and the lower motility of epididymal than ejaculated spermatozoa. Improvements in the fertility of epididymal spermatozoa may be useful for endangered equids (Figure 323.2). In mares, pregnancy rates of 45% were obtained after hysteroscopic insemination of 150 × 106 fresh epididymal spermatozoa.48 In the same study, higher pregnancy rates were obtained from hysteroscopic insemination (9/51, 18%) than from conventional insemination (1/13, 8%) of 200 × 106 frozen-thawed epididymal spermatozoa. Exposure of fresh or frozen-thawed epididymal spermatozoa to seminal plasma did not significantly improve their fertility.48
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Figure 323.2 Hysteroscopic insemination of an anaesthetized zebra with frozen epididymal spermatozoa.
However, when the insemination dose was reduced to 5–10 × 106 motile frozen-thawed epididymal spermatozoa, there was an apparent beneficial effect of exposure to Tyrode’s medium prior to insemination. No pregnancies were obtained in the 20 mares inseminated with spermatozoa not exposed to Tyrode’s medium and 29% (7/24) of mares conceived after insemination with frozen epididymal spermatozoa exposed to Tyrode’s medium after thawing.
Conclusions Satisfactory pregnancy rates are achievable using very low doses of spermatozoa inseminated by deep uterine or hysteroscopic insemination methods if the stallion is fertile. Deep uterine insemination has been shown to be beneficial for the use of low numbers of fresh and cooled, stored spermatozoa. Hysteroscopic insemination has enabled satisfactory conception rates after insemination with ≤ 20 × 106 motile fresh,29 frozen,37 sex-sorted,36, 49 or epididymal48 spermatozoa. Deep uterine and hysteroscopic insemination methods currently have the potential to improve the efficacy of the use of frozen, epididymal and sexsorted spermatozoa. However, their application for subfertile stallions is questionable.
References 1. Vidament M. French field results (1985–2005) on factors affecting fertility of frozen stallion semen. Anim Reprod Sci 2005;89:115–36. 2. Carnevale EM, Maclellan LJ, Coutinho da Silva MA, Checura CM, Scoggin CF, Squires EL. Equine sperm-oocyte interaction: results after intraoviductal intrauterine inseminations of recipients for oocyte transfer. Anim Reprod Sci 2001;68:305–14. 3. Sullivan JJ, Pickett BW. Influence of ejaculation frequency of stallions on characteristics of semen and output of spermatozoa. J Reprod Fertil Suppl 1975;23:29–34.
4. Katila T, Sankari S, M¨akel¨a O. Transport of spermatozoa in the genital tracts of mares. J Reprod Fertil Suppl 2000;56:571–78. 5. Troedsson MHT, Liu IKM, Crabo BG. Sperm transport and survival in the mare. Theriogenology 1998;49:905–15. 6. Troedsson MHT, Desvousges AL, Hansen PJ, Buhi WC. Equine seminal plasma proteins protect live spermatozoa from PMN binding and phagocytosis, while providing a mechanism for selective sperm elimination of apoptotic and dead spermatozoa. Anim Reprod Sci 2006;94:60–1. 7. Scott MA, Liu IKM, Overstreet JW, Enders AC. The structural morphology and epithelial association of spermatozoa at the utero-tubal junction: a descriptive study of equine spermatozoa in situ using scanning electron microscopy. J Reprod Fertil Suppl 2000;56:415–21. 8. Brinsko SP, Varner DD, Blanchard TL. The effect of uterine lavage performed four hours post insemination on pregnancy rate in mares. Theriogenology 1991;35:1111–19. 9. Bader H. An investigation of sperm migration into the oviducts of the mare. J Reprod Fertil Suppl 1982;32:59–64. 10. Householder DD, Pickett BW, Voss JL, Olar TT. Effect of extender, numbers of spermatozoa and hCG on equine fertility. Equine Vet Sci 1981;1:9–13. 11. Pickett BW, Voss JL. The effect of semen extenders and sperm number on mare fertility. J Reprod Fertil Suppl 1975;23:95–8. 12. Gahne S, G˚anheim A, Malmgren L. Effect of insemination dose on pregnancy rate in mares. Theriogenology 1998;49:1071–4. 13. Vidament M, Dupere AM, Julienne P, Evain A, Noue P, Palmer E. Equine frozen semen freezability and fertility field results. Theriogenology 1997;48:907–17. 14. Pickett BW, Voss JL, Squires EL, Vanderwall DK, McCue PM, Breummer JE. Collection, preparation and insemination of stallion semen. Fort Collins: Colorado State University, Anim Reprod Lab Bull No.10, 2000. 15. Pace MM, Sullivan JJ. Effect of timing of insemination, numbers of spermatozoa and extender components on the pregnancy rate in mares inseminated with frozen stallion semen. J Reprod Fertil Suppl 1975;23:115–21. 16. Voss JL, Wallace RA, Squires EL, Pickett BW, Shideler RK. Effects of synchronisation and frequency of insemination on fertility. J Reprod Fertil Suppl 1979;27:257–61. 17. Allen WR, Bowen JM, Frank CJ, Jeffcott LB, Rossdale PD. The current position of AI in horse breeding. Equine Vet J 1976;8:72–4. 18. Bedford SJ, Hinrichs K. The effect of insemination volume on pregnancy rates of pony mares. Theriogenology 1994;42:571–8. 19. Rowley HS, Squires EL, Pickett BW. Effect of insemination volume on embryo recovery in mares. J Eq Vet Sci 1990;10:298–300. 20. Varner DD, Blanchard TL, Love CL, Garcia MC, Kenney RM. Effects of semen fractionation and dilution ratio on spermatozoal motility parameters. Theriogenology 1987;28:709–23. 21. Saacke RG, Dalton J, Nadir S, Bame J, Nebel RL. Spermatozoal characteristics important to sperm transport, fertilisation and early embryonic development. In: Lauria A, Gandolfi F, Enne G, Gianaroli L (eds) Gametes: Development and Function. Serono Symposia: Rome, 1998; pp. 320–35. 22. Morris LH, Allen WR. An overview of low dose insemination in the mare. Reprod Domest Anim 2002;37:206–10. 23. Allen WR, Brown L, Wright M, Wilsher S. Reproductive efficiency of Flatrace and National Hunt Thoroughbred
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mares and stallions in England. Equine Vet J 2007;39: 438–45. ¨ Muller Z. Practicalities of insemination of mares with deep frozen semen. J Reprod Fertil Suppl 1987;35:121–5. Samper JC, Morris CA. Current methods for stallion semen cryopreservation. Theriogenology 1998;49:895–903. Samper JC. Increasing pregnancy rates with frozen semen: can it be done? Proceedings of the Society for Theriogenology. Nashville, TN: 1999; pp. 109–15. Barbacini S, Zavaglia G, Gulden P, Marchi V, Necchi D. Retrospective study on the efficiency of hCG in an equine artificial insemination programme using frozen semen. Equine Vet Educ 2000;12:312–17. Buchanan BR, Seidel GE Jr, McCue PM, Schenk JL, Herickhoff LA, Squires EL. Insemination of mares with low numbers of either unsexed or sexed spermatozoa. Theriogenology 2000;53:1333–44. Morris LH, Hunter RHF, Allen WR. Hysteroscopic insemination of small numbers of spermatozoa at the uterotubal junction of preovulatory mares. J Reprod Fertil 2000;118:95–100. Brinsko SP. Insemination doses: how low can we go? Theriogenology 2006;66:543–50. Lightfoot RJ, Salamon S. Fertility of ram spermatozoa frozen by the pellet method. I. Transport and viability of spermatozoa within the genital tract of the ewe. J Reprod Fertil 1970;22:385–98. Maxwell WM, Evans G, Rhodes SL, Hillard MA, Bindon BM. Fertility of superovulated ewes after intrauterine or oviducal insemination with low numbers of fresh or frozen-thawed spermatozoa. Reprod Fertil Dev 1993;5:57– 63. ¨ Kruger C, Rath D, Johnson LA. Low dose insemination in synchronised gilts. Theriogenology 1999;52:1363–73. Martinez EA, Vazquez JL, Vazquez JM, Lucas X, Gil MA, Parilla I, Roca J. Successful low dose insemination by a fibreoptic endoscope technique in the sow. Theriogenology 2000;53:201. Byrd W, Bradshaw K, Carr B, Edman C, Odom J, Ackerman G. A prospective randomized study of pregnancy rates following intrauterine and intracervical insemination using frozen donor sperm. Fertil Steril 1990;53:521–7. Lindsey AC, Morris LH, Allen WR, Schenk JL, Squires EL. Hysteroscopic insemination of mares with low numbers of nonsorted or flow cytometrically sorted spermatozoa. Equine Vet J 2002;34:128–32. Morris LH, Tiplady CA, Allen WR. Pregnancy rates in mares after a single fixed-time hysteroscopic insemination
38.
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of low numbers of frozen-thawed spermatozoa onto the uterotubal junction. Equine Vet J 2003;35:197–201. Vazquez JJ, Medina V, Lui IK, Ball BA, Scott MA. Nonsurgical utero-tubal insemination in the mare. In: Proceedings of the Society for Theriogenology, Baltimore, 1998; pp. 82–3. Manning ST, Bowman PA, Fraser LM, Card CE. Development of hysteroscopic insemination of the uterine tube in the mare. Proceedings of the Annual Meeting of the Society for Theriogenology. Baltimore, 1998; pp. 84–5. Kenney RM, Bergman RV, Cooper WL, Morse GW. Minimal contamination techniques for breeding mares: technique and preliminary findings. Proceedings of the American Association of Equine Practitioners, 1975; pp. 327–35. Nie GJ, Johnson KE. Pregnancy rate in mares following insemination with a low-dose of progressively motile or filtered sperm cells deep in the uterine horn. Proceedings of the Society for Theriogenology. San Antonio, TX, 2000; p. 303. Brinsko SP, Rigby SL, Lindsey AC, Blanchard TL, Love CC, Varner DD. Pregnancy rates in mares following hysteroscopic or transrectally-guided insemination with low sperm numbers at the utero-tubal papilla. Theriogenology 2003;59:1001–9. Alvarenga MA, Le˜ao KM. Hysteroscopic insemination of mares with low number of frozen-thawed spermatozoa selected by Percoll gradient. Theriogenology 2002;58:651–3. Sieme H, Schafer T, Stout TAE, Klug E, Waberski D. The effects of different insemination regimes on fertility of mares. Theriogenology 2003;60:153–64. Barker CA, Gandier JC. Pregnancy in a mare resulting from frozen epididymal spermatozoa. Can J Comp Med 1957;21:47–51. Fournier-Delpech S, Colas G, Courot M, Ortavant R. Epididymal sperm maturation in the ram: motility, fertilizing ability and embryonic survival after uterine artificial insemination in the ewe. Ann Biol Anim Bioch Biophys 1979;19:597–605. Morris LH, Stout TAE, Li X, Allen WR. The capacitation status of fresh and frozen thawed epididymal and ejaculated stallion spermatozoa. Theriogenology 2000;53:488. Morris LH, Tiplady CA, Allen WR. The in vivo fertility of caudal epididymal spermatozoa in the horse. Theriogenology 2002;58:643–6. Lindsey AC, Schenk JL, Graham JK, Bruemmer JE, Squires EL. Hysteroscopic insemination of low numbers of flow sorted fresh and frozen-thawed stallion spermatozoa. Equine Vet J 2002;34:121–7.
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Chapter 324
Sex-Sorted Spermatozoa Lee H.A. Morris
Introduction The ability to predetermine the sex of offspring at the time of insemination or mating would improve the cost-effectiveness of a breeding program for any species. De Vries et al.1 determined that the rate of genetic progress in dairy cattle using sex-selected spermatozoa is expected to increase by 15% compared with that obtained with conventional insemination programmes. Seidel2 calculated that for a production animal system to benefit from new technologies such as sexed semen, the producer would expect a threefold increase in return for every dollar invested. However, the costs involved in any sex-sorted semen program are not simply the premium price for the semen, but reduced fertility and increased labor. For companion animals such as horses, the premium paid for a dose of sexed semen may be worthwhile. For different breeds of horses there will be different breeding objectives and gender preferences; examples of this are that fillies may be preferred for polo and colts may be preferred in the Standardbred industry.
tozoa using the albumin column and evaluation of karyotypes in subsequent hamster oocyte penetration tests. Currently, the only proven method of sex preselection that has resulted in the birth of live offspring of predetermined sex is provided by flow cytometric DNA analysis or fluorescence-activated cell separation (FACS) technology (reviewed by Morris).7 This method of sex preselection of spermatozoa is based on the Beltsville flow cytometry technology which was first adapted for spermatozoa by Gledhill et al.8 Ten years later, Johnson and Pinkel9 modified the process such that it now orientates spermatozoa to the laser beam to improve sperm sorting rates and accuracy. These modifications were closely followed by the inclusion of vital staining of the rabbit sperm DNA which led to the production of the first viable sex-preselected offspring.10 Since then, flow cytometric separation of spermatozoa based on their X- and Y-chromosome bearing populations has resulted in the birth of healthy piglets,11–14 lambs,15–20 calves,21–24 foals,25–29 and humans30 as cited in Fugger et al.31
Methods of sex preselection
Principles of FACS separation of stallion spermatozoa
It has been shown that X- and Y-chromosome bearing cells may be separated to some degree based on their physical differences. Roberts3 suggested that the Xbearing sperm cell may be larger than the Y-bearing cell. Since then it has been possible to separate X- and Y-cells with 70% accuracy based upon differences in their volume.4 The notion that Y-bearing spermatozoa have a higher velocity than X-bearing spermatozoa has been exploited in human andrology clinics with the use of albumin column separation.5 However, the results of these analyses of clinical data were questioned by Brandriff et al.6 when they observed no alteration in the sex ratio after separation of human sperma-
The sex of an offspring is determined at the time of fertilization when either an X- or a Y-chromosome bearing spermatozoon fertilizes the oocyte. It is possible to separate the sperm population into the different X and Y subpopulations by exploiting the differences in the DNA of each sperm cell. At the time of writing, separation of X- and Y-chromosome bearing spermatozoa by their DNA content, as described by Johnson et al.32 is yet to be superseded as the most effective means of sex-sorting spermatozoa. The ability to separate spermatozoa into their Xand Y-chromosome bearing populations is dependent on the relative sizes of the X and Y chromosome and the total DNA content per spermatozoon.32 Put
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Sex-Sorted Spermatozoa simply, the X-chromosome contains more DNA than the Y-chromosome. The total DNA content will vary among species,33, 34 such that the difference in the total DNA for X- and Y-chromosome spermatozoa is species-specific. This difference in DNA content between X- and Y-chromosome bearing spermatozoa is 3.7% in the stallion.35
Method of FACS separation of stallion spermatozoa Stallion spermatozoa are diluted in a Kenney’s modified Tyrode’s medium.36 This diluent provides protective proteins from the skim milk during centrifugation, staining, incubation, and FACS separation. More recent studies have examined the beneficial effects of a clear extender containing bovine serum albumin, to replace the skim milk component, to optimize sperm processing for FACS separation.37, 38 To measure the amount of DNA in each sperm cell, the sperm DNA is stained with the fluorochrome Hoechst 33342 during an incubation period at 34◦ C. Hoechst 33342 is a non-intercalating fluorescent stain that binds to the thymine- and adenosine-rich regions of the DNA. The production of viable embryos, in vitro39 and in vivo,17 and the production of live offspring in many species support the concept that this stain is minimally detrimental to embryonic survival and development. Nevertheless, the incubation temperature and time required to optimize the uptake of Hoechst 33342 without reducing the fertility of the spermatozoa varies amongst species (reviewed by Morris7 and Rath et al.40 ). The sperm sample is then stained with a food dye which will quench the fluorescence of any non-viable spermatozoa and thus allow preferential sorting of the viable sperm populations.33 Any reduction in the proportion of non-viable spermatozoa in the population for sorting will improve the flow and sort rates. The stained semen sample is passed through the FACS machine at a very low concentration and at a specific pressure (40–50 psi) until it reaches the nozzle tip which orientates each sperm cell to an argon laser beam. Exposure of each sperm cell to the argon laser beam induces maximum fluorescence of the Hoechst stained DNA41 (reviewed by Johnson, 2000).33 Fluorescence is collected from a 0◦ fluorescence detector and a 90◦ detector, such that only correctly oriented sperm cells will have their DNA analyzed. The 0◦ detector analyzes the orientation of the sperm cells and differences in DNA are detected by the 90◦ detector. Once the DNA has been analyzed, a piezoelectric crystal interrupts the stream of sheath fluid to create individual droplets, each containing a single spermatozoon. Each droplet containing a spermatozoon is
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353∅ 100.0% (SSC) Figure 324.1 Dot plot representation of sorted sperm populations after the detection of forward (0◦ ) and side (90◦ ) fluorescence of Hoechst 33342 stained spermatozoa. R1 region: represents the correctly orientated, resolvable population of stained spermatozoa. R5 region: represents the dead and membrane-damaged population of spermatozoa, collected as waste. Adapted from Clulow et al. Factors influencing the ‘sortability’ of stallion spermatozoa into X- and Y-chromosome bearing populations. Anim Reprod Sci 2009;113:220–8.46 (See color plate 150).
electrically charged as it passes by deflection plates such that positively charged droplets containing Xspermatozoa are deflected towards the negative plate and vice versa for negatively charged Y- spermatozoa. Spermatozoa that may be inadequately charged, poorly oriented or do not fluoresce, will be collected as waste (Figure 324.1). The purity of the population of sorted spermatozoa can be optimized at different sort rates. It is possible to obtain > 90% purity for either X or Y sperm populations. The X-chromosome is slightly larger than the Y-chromosome, so the fastest and most accurate sort rates are achievable when only the Xsperm population is selected.42 The degree of difference in the amount of DNA between the X- and Ychromosome will influence the purity of the sorted sperm populations and the sorting ease.13 Improvements in the nozzle tip have influenced the possible sort rates for spermatozoa. In 1999, Rens et al.43 improved the nozzle tip to orient up to 70% of the spermatozoa to the laser beam compared with 25% orientation achievable with earlier designs. This new nozzle enabled sort rates to improve from 3.5 × 105 to 6 × 106 sorted spermatozoa/h (reviewed by Johnson, 2000).33 Since then, XY Inc. (Fort Collins, Colorado) further refined the nozzle tip (CytonozR R SX) so that the MoFlo SX sorter opzleTM , MoFlo 6 erates at sorting rates of 15–28 × 10 sorted spermatozoa/h (reviewed by Maxwell et al., 2004, Rath et al.
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Post Incubation
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Figure 324.2 The mean (± sem) percentage of morphologically normal spermatozoa at different stages of processing stallion spermatozoa for sex sorting and cryopreservation. Raw, after dilution in Kenney’s medium (Kenneys), after incubation with the Hoechst 33342 stain, after cooling to 4◦ C before cryopreservation (Post Cool, SS, and NS) and after thawing of frozen samples. Significant differences (P < 0.05) are identified by different superscripts. Adapted from Clulow et al. (2007).47
2009; Sharpe and Evans, 2009).40, 44, 45 At these sort rates it would take at least 20 h to sort a dose of 300 × 106 spermatozoa, by which time the viability of the sorted spermatozoa would have deteriorated significantly. Consequently, the use of this technology in horses requires low-dose insemination techniques which facilitate the insemination of low doses of 5–20 × 106 sorted spermatozoa. The sorting process itself should, by default, select only viable spermatozoa with intact plasma membranes. In horses, Clulow et al.47 have described how the sex-sorting process selects a sperm population with a high proportion of morphologically normal spermatozoa (Figure 324.2), which is also maintained after freezing and thawing. However, despite the higher proportion of morphologically normal spermatozoa after the sex-sorting process, there are more tail defects in sex-sorted spermatozoa than non-sorted spermatozoa after freezing
and thawing47, 48 which may contribute to a reduction in motility observed in sorted, frozen–thawed spermatozoa for some stallions48 but not overall.47 Furthermore, despite the detrimental effects of staining and incubating stallion spermatozoa, no significant differences have been observed in the proportion of intact acrosomes in fresh spermatozoa and in the sperm population selected by the sorting process.49 Indeed, such a robust population of spermatozoa is selected by flow cytometry that the percentage of intact acrosomes remains relatively constant over a 24 h period after sorting (Table 324.1).
Factors which influence the sortability of stallion spermatozoa ‘Sortability’ is defined as the ability of a sperm sample to be sorted into resolvable X- and Y-chromosome bearing populations.42 Indeed, there is variation
Table 324.1 The motility and acrosome status of flow-sorted spermatozoa over time. Adapted from Morris et al. (2003)49 Stage of processing
Temperature
Progressive motility (%)
Acrosome intact (%)
Pre-stain Post-stain Post-incubation Post-sort: 0 h Post-sort: 2 h
20◦ C 20◦ C 20◦ C 20◦ C 4◦ C 20◦ C 4◦ C 20◦ C 4◦ C 20◦ C 4◦ C 20◦ C
50.7a ± 10.2 42.0a ± 17.1 48.0a ± 15.7 49.9a ± 18.3 34.3b ± 19.3 40.8a,b ± 21.8 8.5c ± 12.8 27.0b ± 21.0 4.8c ± 12.9 17.2b,c ± 17.9 0.0c ± 0.0 5.1c ± 8.6
57.1b ± 28.2 42.3a ± 26.7 34.9a ± 27.1 60.2b ± 22.3 47.1a ± 28.3 53.9a,b ± 24.5 59.8b ± 19.7 66.6b ± 12.0 56.2b ± 16.8 64.3b ± 16.4 32.7a ± 25.0 41.6a ± 24.0
Post-sort: 12 h Post-sort: 24 h Post-sort: 48 h
a,b,c
Values within a column with different superscripts are significantly different (P < 0.05).
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Sex-Sorted Spermatozoa amongst species,50 breeds,51 and individuals19 in the rate at which the fluorochrome Hoechst 33342 is taken up by sperm DNA. Garner50 has described the effect of the size and area of the sperm head on the sortability of spermatozoa from different species. He defined ‘sorting indices’ for a number of species and for stallions he calculated an index of 59 compared with 131 for bulls, which have a larger sperm head. Clulow et al.46 analyzed the effects of the proportion of spermatozoa discarded as waste and sperm motility on the ‘sortability’ of spermatozoa from 11 stallions. The results of this study found that sperm motility was not a predictor of ‘sortability’ for stallions, but that the proportion of discarded spermatozoa (Figure 324.1) significantly influenced the ‘sortability’ of a stallion’s ejaculate. These observations suggest that it may be possible to screen stallions for ‘sortability’ prior to inclusion in a sex-sorted semen program.
The fertility of sex-sorted stallion spermatozoa Since the production of the first live offspring from sex-sorted spermatozoa in rabbits,10 there have been many offspring produced from a range of species including cattle, sheep, elk, dolphins, and horses. Fertility trials in dairy heifers inseminated with only 1.5–2 × 106 frozen–thawed, sexed spermatozoa have produced conception rates that are 70–80% of those obtained with conventional doses of non-sorted semen.2 Recent field trials in dairy cattle have revealed an effect of lactation status and sire on the conception rates obtained with low doses of frozen–thawed sexed semen,52 such that the conception rates of cows (29%) were approximately half that of heifers (46–59%) and that the effect of sire and dose was only evident in heifers. Recently, Rath et al. (2009)40 described the beneficial effects of modifying the handling of the sorted frozen–thawed semen using a proR ’, to improve the fertility of tocol named ‘Sexcess sorted bull spermatozoa. The first foal produced from sex-sorted spermatozoa was the result of surgical oviductal insemination of a mare with 50 000 fresh, sorted spermatozoa53 as part of a study on oviductal insemination and intracytoplasmic sperm injection.54 . Since then, less invasive methods of low-dose insemination have been developed to optimize the fertility of low numbers of sex-sorted spermatozoa that are available. The requirement for a surgical insemination technique for such low numbers of spermatozoa does not lend itself to the practical application of this technology. Therefore, Buchanan et al.25 at Colorado State University developed a deep uterine insemination
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procedure to deposit the insemination dose at the tip of the uterine horn ipsilateral to impending ovulation. This early study involved the use of a rigid disposable embryo transfer pipette that was passed along the lumen of the uterine horn until its tip was visualized by transrectal ultrasound at the most cranial tip of the uterine horn. In this study, 35% of the 20 mares inseminated with 25 × 106 fresh sex-sorted spermatozoa conceived. Meanwhile, a hysteroscopic insemination procedure had been developed at the Equine Fertility Unit (UK) which enabled doses of 1–10 × 106 spermatozoa to produce pregnancy rates similar to those obtained with conventional insemination.55 This insemination method was subsequently investigated for its suitability for fresh sex-sorted semen.27, 28 In this study, mares were inseminated with 5 × 106 sorted and non-sorted fresh spermatozoa at the time of administration of human chorionic gonadotropin (hCG; Chorulon, Intervet). Five of ten mares (50%) conceived after insemination with non-sorted spermatozoa compared with 5 of 20 mares (25%) inseminated with the same low number of sorted spermatozoa. For comparison, a group of mares (n = 10) were inseminated with the same low number of fresh, non-sorted spermatozoa using the deep insemination procedure and none of those mares conceived. In many cases, it is envisaged that the highly specialized FACS laboratory may be located some distance from the facility where the stallion is based or where the mare is to be inseminated. In order for this technology to be utilized in the horse industry, either transported or frozen–thawed sex-sorted semen will be required. To overcome these challenges, Lindsey et al.29 evaluated the fertility of semen that had been collected, extended to 25 × 106 /mL, stored at either 5◦ C or 15◦ C for 18 h, sorted and then used for hysteroscopic insemination of mares with 20 × 106 spermatozoa (in 300 µL) at 30 h after administration of hCG. The optimal storage temperature for the semen prior to sorting was found to be 15◦ C (72%, 18/25 mares pregnant) when compared with 5◦ C (55%, 12/22 mares pregnant). In this study, another group of mares was inseminated with sorted spermatozoa that had been stored at 5◦ C, using the deep insemination procedure and only 9 of 24 mares (38%) conceived. Based on these results it was concluded that hysteroscopic insemination was superior to deep uterine insemination for sex-sorted spermatozoa. This study demonstrated that semen stored for up to 18 h prior to sex-sorting could retain excellent fertility. The ability to use frozen sex-sorted spermatozoa would facilitate the commercialization of this technology. In production animal species, the successful freezing protocols for sex-sorted spermatozoa have
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been based on standard cryopreservation protocols for that species (bull,56 ram,17, 18 ). For stallion semen, Lindsey et al.28 investigated the effects of three different extenders on post-thaw motility over time and found that FR5 was the preferred extender. However, Clulow et al.48 observed no differences in the postthaw motility estimates of sex-sorted semen frozen in either INRA8257 or lactose-EDTA.58 To evaluate the fertility of frozen, sex-sorted semen, Lindsey et al.28 inseminated mares with 5 × 106 fresh and frozen, sorted and non-sorted spermatozoa. The pregnancy rate after insemination of the sorted, frozen–thawed spermatozoa at 30 h after hCG administration was disappointingly low (13%), and significantly less than the pregnancy rate obtained with the non-sorted, frozen–thawed spermatozoa (38%). Clulow et al.59 evaluated the fertility of low doses of 6, 13, and 25 × 106 frozen, sex-sorted spermatozoa after hysteroscopic insemination between 32 and 38 h after hCG administration. Only one pregnancy (9.1%, 1/11 cycles) was obtained after hysteroscopic insemination with 13 × 106 frozen, sex-sorted spermatozoa, and two pregnancies (2/10) were obtained after hysteroscopic insemination with non-sorted frozen spermatozoa. In this study, the fertility of the control group of mare cycles inseminated with conventional doses containing a minimum of 500 million nonsorted frozen semen was also low, 35% (22/62 cycles). Despite the disappointing results of this study, they do illustrate that hysteroscopic insemination of low numbers of spermatozoa, sorted or non-sorted, will not overcome some forms of intrinsic subfertility in stallions. It has recently been demonstrated that the fertility of frozen–thawed sex-sorted spermatozoa is lower than non-sorted frozen–thawed control spermatozoa after intracytoplasmic injection in two separate studies.60, 61 In both studies the problem with fertility of the frozen–thawed sex-sorted spermatozoa was observed at the time of cleavage, such that only 20–30% of oocytes injected with sex-sorted frozen–thawed spermatozoa cleaved compared with 71–83% of the oocytes injected with control, nonsorted frozen–thawed spermatozoa. Once blastocysts had developed then pregnancy rates were satisfactory after non-surgical embryo transfer. The procedures associated with the sex-sorting procedure itself provide extreme conditions for the viable sperm cell. The process exposes spermatozoa to low dilution rates, prolonged exposure to room temperature, DNA stains, excitation by an argon laser, high pressure forces of 40–50 psi, different media and buffers at different stages of processing, centrifugation, and variable incubation temperatures during processing (transport temperatures, incubation during staining, cryopreservation, thawing) and
finally cells must transit through the selective gradient of the female reproductive tract prior to fertilization. Superimposed upon these obstacles is the availability of such low numbers of sex-sorted spermatozoa for insemination. Therefore, any increase in the number of sorted spermatozoa, or reduction in the variability or duration of the processing steps, should improve the efficiency and fertility of the sex-sorted semen process.
The future of sex-sorted stallion spermatozoa Currently, the commercialization of sex-sorted stallion spermatozoa will depend upon a market for fresh or stored sex-sorted spermatozoa, until satisfactory fertility rates are obtained with frozen–thawed sex-sorted stallion spermatozoa. It has been shown in sheep that simple modifications of the freezing and sorting protocols can turn a commercially non-viable procedure into a commercially attractive technology.17 There is also the potential for the equine industry to have frozen, sex-sorted semen as a breeding option in the near future.
References 1. De Vries A, Overton M, Fetrow J, Leslie K, Eicker S, Rogers G. Exploring the impact of sexed semen on the structure of the dairy industry. J Dairy Sci 2008;91:847–56. 2. Seidel GE Jr. Sexing mammalian sperm: intertwining of commerce, technology, and biology. Anim Reprod Sci 2003;79:145–56. (Review.) 3. Roberts AM. Gravitational separation of X and Y spermatozoa. Nature 1972;262:223–35. 4. Van Munster EB, Stap J, Hoebe RA, te Meerman GJ, Aten JA. Difference in sperm head volume as a theroetical basis for sorting X- and Y-bearing spermatozoa: potentials and limitations. Theriogenology 1999;52:1281–93. 5. Beernink FJ, Ericsson RJ. Male sex preselection through sperm isolation. Fertil Steril 1982;38:493–5. 6. Brandriff BF, Gordon LA, Haendel S, Singer S, Moore DH, Gledhill BL. Sex chromosome ratios determined by karyotype analysis in albumin isolated human sperm. Fertil Steril 1986;46:678–685. 7. Morris LHA. Challenges facing sex preselection of stallion spermatozoa. Anim Reprod Sci 2005;89:147–57. 8. Gledhill BL, Lake S, Steinmetz LL, Gray JW, Crawford JR, Dean PN, Van Dilla MA. Flow microfluorometric analysis of sperm DNA content: effect of cell shape on the fluorescence distribution. J Cell Physiol 1976;87:367–76. 9. Johnson LA, Pinkel D. Modification of a laser-based flow cytometer for high resolution DNA analysis of mammalian spermatozoa. Cytometry 1986;7:268–73. 10. Johnson LA, Flook JP, Hawk HW. Sex preselection in rabbits: live birth from X and Y sperm separated by DNA and cell sorting. Biol Reprod 1989;41:199–203. 11. Johnson LA. Sex selection in swine: altering sex ratios in offspring following surgical insemination with flow-sorted Xand Y-bearing sperm. Reprod Domest Anim 1991;26:195–204.
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Sex-Sorted Spermatozoa 12. Rath D, Johnson LA, Dobrinsky JR, Welch GR, Niemann H. Production of piglets preselected for sex following in vitro fertilization with X and Y chromosomebearing spermatozoa sorted by flow cytometry. Theriogenology 1997;47:795–800. 13. Rath D, Long CR, Dobrinsky JR, Welch GR, Schreier LL Johnson LA. In vitro production of sexed embryos for gender preselection: high speed sorting of X- chromosomebearing sperm to produce piglets after embryo transfer. J Anim Sci 1999;77:3346–52. 14. Vazquez JM, Martinez EA, Parrilla I, Roca J, Gil MA, Vazquez JL. Birth of piglets after deep intrauterine insemination with flow cytometrically sorted boar spermatozoa. Theriogenology 2003;59:1605–14. 15. Catt SL, Catt JW, Gomez MC, Maxwell WMC, Evans G. Birth of a male lamb derived from an in vitro matured oocyte fertilized by intracytoplasmic injection of a single presumptive male sperm. Vet Rec 1996;139:494–5. 16. Cran DG, McKelvey WAC, King ME, Dolman DF, McEvoy TG, Broadbent PJ, Robinson JJ. Production of lambs by low dose intrauterine insemination with flow cytometrically sorted and unsorted semen. Theriogenology 1997;47:267. (Abstract). 17. De Graaf SP, Beilby K, O’Brien JK, Osborn D, Downing JA, Maxwell WM, Evans G. Embryo production from superovulated sheep inseminated with sex-sorted ram spermatozoa. Theriogenology 2007;67:550–5. 18. Hollinshead FK, O’Brien JK, Maxwell WMC, Evans G. Production of lambs of predetermined sex after the insemination of ewes with low numbers of frozen–thawed sorted Xor Y-chromosome-bearing spermatozoa. Reprod Fertil Dev 2002;14:503–8. 19. Hollinshead FK, Evans G, Evans KM, Catt SL, Maxwell WMC, O’Brien JK. Birth of lambs of a predetermined sex after in vitro production of embryos using frozen-thawed sex-sorted and refrozen-thawed ram spermatozoa. Reproduction 2004;127:557–68. 20. Morrell JM, Dresser DW. Offspring from inseminations with mammalian sperm stained with Hoechst 33342, either with or without flow cytometry. Mutation Res 1989;224:177–83. 21. Cran DG, Johnson LA, Miller NG, Cochrane D, Polge C. Production of bovine calves following separation of X- and Y-chromosome bearing sperm and in vitro fertilization. Vet Rec 1993;132:40–1. 22. Cran DG, Johnson LA, Polge C. Sex preselection in cattle: a field trial. Vet Rec 1995;136:495–6. 23. Seidel GE Jr, Allen CH, Johnson LA, Holland MD, Brink Z, Welch GR, Graham JK, Cattell MB. Uterine horn insemination of heifers with very low numbers of non-frozen and sexed spermatozoa. Theriogenology 1997;48:1255–64. 24. Seidel GE Jr, Schenk JL, Herickhoff LA, Doyle SP, Brink Z, Green RD, Cran DG. Insemination of heifers with sexed sperm. Theriogenology 1999;52:1407–20. 25. Buchanan BR, Seidel GE Jr, McCue PM, Schenk JL, Herickhoff LA, Squires EL. Insemination of mares with low numbers of either unsexed or sexed spermatozoa. Theriogenology 2000;53:1333–44. 26. Clulow JR, Evans G, Maxwell WMC, Rogers JA, CawdellSmith AJ, Sieme H, Rath D, Morris LHA. Birth of foals from artificial insemination with sex-sorted and nonsorted frozen stallion spermatozoa. Reprod Domest Anim 2006;41:337 [Abstract]. 27. Lindsey AC, Morris LH, Allen WR, Schenk JL, Squires EL, Bruemmer JE. Hysteroscopic insemination of mares with
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low numbers of nonsorted or flow sorted spermatozoa. Equine Vet J 2002;34:128–32. Lindsey AC, Schenk JL, Graham JK, Bruemmer JE, Squires EL. Hysteroscopic insemination of low numbers of flow sorted fresh and frozen thawed stallion spermatozoa. Equine Vet J 2002;34:121–7. Lindsey AC, Varner DD, Seidel GE Jr, Bruemmer JE, Squires EL. Hysteroscopic or rectally guided, deep uterine insemination of mares with spermatozoa stored 18 h at either 5◦ C or 15◦ C prior to flow cytometric sorting. Anim Reprod Sci 2005;85:125–30. Levinson G, Keyvanfar K, Wu KC. DNA based X-enriched sperm separation as an adjunct to preimplantation genetic testing for the prevention of X-linked disease. Human Reprod 1995;10:979–82. Fugger EF, Black SH, Keyvanfar K, Schulman JD. Births of normal daughters after MicroSort sperm separation and intrauterine insemination, in vitro fertilization, or intracytoplasmic sperm injection. Hum Reprod 1998;13:2367–70. Johnson LA, Flook JP, Look MV, Pinkel D. Flow sorting of X and Y chromosome bearing spermatozoa into two populations. Gamete Res 1987;16:1–9. Johnson LA. Sexing mammalian sperm for production of offspring: the state of the art. Anim Reprod Sci 2000;60:93–107. O’Brien JK, Stojanov T, Heffernan SJ, Hollinshead FK, Vogelnest L, Maxwell WMC, Evans G. Flow cytometric sorting of non-human primate sperm nuclei. Theriogenology 2005;63:246–59. Welch GR, Johnson LA. Sex preselection: laboratory validation of the sperm sex ratio of flow sorted X- and Y-sperm by sort reanalysis for DNA. Theriogenology 1999;52:1343–52. Padilla AW, Foote RH. Extender and centrifugation effects on the motility patterns of slow-cooled stallion spermatozoa. J Animal Sci 1991;69:3308–13. Gibb Z, Grupen CG, Morris LHA, Evans G, Maxwell WMC. Substitution of skim milk with bovine serum albumin in a stallion semen diluent. Reprod Fertil Dev (Suppl) 2008;20:63. Gibb Z, Morris L, Grupen C, Evans G, Maxwell C. Effect of protein source in stallion semen diluent on the motility, acrosome integrity and morphology of sperm. Reprod Domest Anim (Suppl) 2008;43:104. Zhang M, Lu KH, Seidel GE Jr. Development of bovine embryos after in vitro fertilization of oocytes with flow cytometrically sorted, stained and unsorted sperm from different bulls. Theriogenology 2003;60:1657–63. Rath D, Moench-Tegeder G, Taylor U, Johnson LA. Improved quality of sex-sorted sperm: a prerequisite for wider commercial application. Theriogenology 2009;71:22–9. Pinkel D, Gledhill BL, Van Dilla MA, Stephenson D, Watchmaker G. High resolution DNA measurements of mammalian sperm. Cytometry 1982;3:1–9. Johnson LA, Welch GR. Sex preselection: high-speed flow cytometric sorting of X- and Y-sperm for maximum efficiency. Theriogenology 1999;52:1323–41. Rens W, Welch GR, Johnson LA. Improved flow cytometric sorting of X- and Y-chromosome bearing sperm: substantial increase in yield of sexed semen. Mol Reprod Dev 1999;52:50–6. Maxwell WMC, Evans G, Hollinshead FK, Bathgate R, de Graaf SP, Eriksson BM, Gillan L, Morton KM, O’Brien JK. Integration of sperm sexing technology into the ART toolbox. Anim Reprod Sci 2004;82:79–95. Sharpe JC, Evans KM. Advances in flow cytometry for sperm sexing. Theriogenology 2009;71:4–10.
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46. Clulow JR, Evans G, Morris LHA, Maxwell WMC. Factors influencing the ‘sortability’ of stallion spermatozoa into Xand Y-chromosome bearing populations. Anim Reprod Sci 2009;113:220–8. 47. Clulow JR, Evans G, Morris LHA, Maxwell WMC. The motility and morphology of ejaculated and epididymal stallion spermatozoa after flow cytometric sex-sorting and cryopreservation. In: PhD Thesis ‘Sex Preselection of Stallion Spermatozoa’ Clulow, JR. The University of Sydney, Australia, 2007. 48. Clulow JR, Buss HF, Evans G, Sieme H, Rath D, Morris LHA, Maxwell WMC. Effect of staining and freezing media on sortability of stallion spermatozoa and their postthaw viability after sex-sorting and cryopreservation. In: PhD Thesis ‘Sex Preselection of Stallion Spermatozoa’ Clulow, JR. The University of Sydney, Australia, 2007. 49. Morris LHA, Hollinshead FK, Evans G, Maxwell WMC. The longevity and acrosome status of stallion flow sorted spermatozoa. Theriogenology 2003;59:512 (Abstract). 50. Garner DL. Flow cytometric sexing of mammalian sperm. Theriogenology 2006;65:943–57. 51. Garner DL, Gledhill BL, Pinkel D, Lake S, Stephenson D, Van Dilla MA, Johnson LA. Quantification of the X- and Ychromosome bearing spermatozoa of domestic animals by flow cytometry. Biol Reprod 1983;28:312–21. 52. DeJarnette JM, Nebel RL, Marshall CE, Moreno JF, McCleary CR, Lenz RW. Effect of sex-sorted sperm dosage on conception rates in Holstein heifers and lactating cows. J Dairy Sci 2008;91:1778–85. 53. Schmid RL, Kato H, Herickhoff LA, Schenk JL, McCue PM. Fertilization with sexed spermatozoa using intracytoplasmic sperm injection and oviductal insemination. In: 7th International Symposium Equine Reproduction. Pretoria, South Africa, 1998; p. 139.
54. Schmid RL, Kato H, Herickhoff LA, Schenk JL, McCue PM, Chung YG, Squires EL. Effects of follicular fluid or progesterone on in vitro maturation of equine oocytes before intracytoplasmic sperm injection with nonsorted and sex-sorted spermatozoa. J Reprod Fert Suppl 2000;56:519–25. 55. Morris LHA, Hunter RHF, Allen WR. Hysteroscopic insemination of small numbers of spermatozoa at the uterotubal junction of preovulatory mares. J Reprod Fertil 2000;118:95–100. 56. Schenk JL, Suh TK, Cran DG, Seidel GE Jr. Cryopreservation of flow-sorted bovine sperm. Theriogenology 1999;52:1375–91. 57. Magistrini M, Couty I, Palmer E. Interactions between sperm packaging, gas environment, temperature and diluent on fresh stallion sperm survival. Acta Vet Scand 1992;88:97–110. 58. Martin JC, Klug E, Gunzel A. Centrifugation of stallion semen and its storage in large volume straws. Reprod Fertil (Suppl) 1979;27:47–51. 59. Clulow JR, Buss HF, Sieme H, Rodger JA, CawdellSmith AJ, Evans G, Rath D, Morris LHA, Maxwell WMC. Field fertility of sex-sorted and non-sorted frozen-thawed stallion spermatozoa. Anim Reprod Sci 2008;108;287– 97. 60. Galli C., Colleoni S., Spinaci M, Duchi R, Merlo B, Tamanini C, Lazzari G, Mari G. Horse pregnancies established with embryos produced in vitro by ICSI with sex-sorted frozenthawed semen. In: The Havemeyer Foundation 7th International Symposium on Equine Embryo Transfer, Cambridge, UK, July 2008. pp. 55–6. 61. Squires EL, Suh TK, Grahame JK, Carnevale EM. The use of sexed, refrozen spermatozoa for ICSI. In: The Havemeyer Foundation 7th International Symposium on Equine Embryo Transfer, Cambridge, UK, July 2008, pp. 54–5.
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Section (d): Future Technologies
325.
Future Reproductive Technology George E. Seidel Jr
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Chapter 325
Future Reproductive Technology George E. Seidel Jr
An amazing array of reproductive technologies already is used for horses, and in principle, essentially any procedures used in other mammals can be applied to horses, including those currently used only for research. The same is true of genetic tools; for example, much of the equine genome has been sequenced, and horse breeding can evolve to be as scientific as plant breeding is today. The objectives of this chapter are: (i) to describe reproductive and genetic tools that may become available, and (ii) to illustrate ways these tools might be applied. A brief analysis of how horses are owned and used seems appropriate as background for these objectives.
Ownership and use of horses Horses are either domesticated or feral. Over the centuries horses have had hundreds of uses including military, draft for agriculture, transportation, meat and milk, currency and assets, a source of raw materials such as hair and hides, sports and entertainment, a physical and emotional therapeutic tool, and pleasure/companionship. Many of these uses remain current in most countries, but present reproductive and genetic technologies are unlikely to be used for most horses in most countries in the foreseeable future, nor are futuristic technologies. Rather, for the next decade or two, newer reproductive technologies likely will be used mostly in the context of research or enhancing recreational and athletic qualities of horses owned by individuals with appropriate financial resources. Qualities like disposition, health, coat color, sex, intelligence, fertility, and athletic ability will be of interest, whereas qualities such as milk production, milk characteristics, carcass characteristics, and feed efficiency will rarely be sought after.
Except for research, most horses involved likely will be registered with a breed association, and to take financial advantage of the benefits of new technologies, any offspring usually will need to be registerable. Thus, breed associations are gatekeepers via the rules they promulgate for eligibility for registration. These rules vary from severe limitations with Thoroughbreds to no limitations for a few breeds. With most breeds, rules evolve as breeders become comfortable with new technologies. The newer technologies and some applications that are fairly routinely available commercially, such as regulation of reproductive cycles and ovulation, cryopreservation of sperm and embryos, artificial insemination, and embryo transfer, will not be discussed further except in the context of delivering the newer technologies. The first group considered will be technologies that are already used in horses, primarily in a research sense (Table 325.1), but are not yet routinely available. Most of these techniques are covered in detail in other chapters in this book, and they are futuristic only in the sense that most are not readily available for widespread commercial use. Some of these technologies have relatively high rates of success, but most do not at present. A few will become available routinely in the next few years; most will take longer.
The futuristic technologies The constraints Constraints on futuristic technologies range from the trivial to the profound. The main constraint in thinking may be imagination, and in that context, sorting
Equine Reproduction, Second Edition. Edited by Angus O. McKinnon, Edward L. Squires, Wendy E. Vaala and Dickson D. Varner c 2011 Blackwell Publishing Ltd.
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Table 325.1 Equine research technologies that may become available commercially Technology
Comments
In vitro oocyte maturation Oocyte transfer Gamete intrafallopian transfer (GIFT) Intracytoplasmic sperm injection (ICSI) In vitro fertilization (IVF) Somatic cell nuclear transfer Splitting embryos Preimplantation genetic diagnosis (PGD)
Also requires IVF, ICSI and/or oocyte or embryo transfer (ET) Surgical or laparoscopic procedure and requires AI or natural service Surgical or laparoscopic procedure with sperm and oocytes Requires embryo culture and ET Requires embryo culture and ET Often called cloning; can be done more reliably with embryonic cells, requires embryo culture and ET A form of cloning embryos to make identical twins, requires ET Requires microsurgery to embryos and embryo culture and ET; can be used for sexing embryos and detecting genetic abnormalities Requires AI Requires sperm, oocytes, embryos, or somatic cells plus cryopreservation and technologies to make animals from thawed cells
Sexing sperm Preservation of genetics
AI, artifical insemination.
out the possible from the ridiculous. There are biological and physical constraints; for example, it is likely that genetic information for embryonic and fetal development and postnatal function will continue to be specified by DNA. There is the constraint of definition; how much can we change a horse and still call it a horse? Extreme examples are having mechanical horses that simulate bucking stock (already exist) or making horses more like camels or the converse; for example, there are racing camels and racing cattle. Defining ‘horseness’ has considerable non-scientific components. There also are the legal and ethical issues; the latter, perhaps best defined as ‘do the right things’, means different ‘right things’ to different people, and to the same person at different times or in different situations. For example, the ‘right thing’ may be different for your horse than someone else’s horse, and opinions change with education and familiarity. In some instances, opinions on whether cloning horses should be permitted under some circumstances change when it is explained that phenotypes will never be duplicated exactly, but genetics for breeding will be duplicated for practical purposes, at least via stallions, since mitochondrial DNA is not normally transmitted via sperm as it is via oocytes. Also, cloning is usually thought of as narrowing the genetic pool, but it can be used to broaden the genetic base in a variety of ways, for example making a stallion from a gelding. There are constraints of finances, logistics, and talent. People need to commit aspects of their careers to such undertakings, and someone needs to pay the bills. Some insights may evolve using the horse as a biomedical model. For example, a good case has been made that the older mare may be the best nonprimate model for reproductive aging of women.1
Predicting the end product Living organisms are constrained by principles of chemistry, physics, and laws of chance. In the context of this chapter, the ultimate desired end product almost always will be producing particular phenotypes of horses, and this always will require reproduction, aided by the assisted-reproductive tools already mentioned. Phenotype is determined by genetics, environment, chance aspects of genetics and epigenetics, and interactions among these, discussed in some detail elsewhere.2 Between using genetic tools and providing an appropriate environment, breeders will continue to be able to produce the desired phenotype with a fair amount of accuracy on average. However, individual animals will vary from this average, even with cloning, due to chance epigenetic effects2 and the inability to reproduce environments precisely; for example, uteri of recipients will vary. However, with the exception of cloning, which in some respects is trivial relative to designing a particular genotype, it always will be impossible to predict the exact outcome of manipulating the genome. This is because every genetic change will interact with at least some of the other genetic elements in the genome, as well as with environmental aspects and chance epigenetic events. Thus, there are essentially infinite permutations and combinations of genetic possibilities, and an (unknown) optimal environment for each possibility. The practical consequence is that characteristics of live animals will remain the only proof of phenotypic outcome of any of the procedures we can imagine. There will be both spectacular successes and failures, and the whole spectrum between these. However, it will be possible to direct outcomes toward the successful end of this spectrum with much higher probability than current
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Future Reproductive Technology procedures allow because much more information and much more powerful tools will be available. A theoretical gene machine I will not go into producing virtual horses by computer simulation, although this could eventually end up being the most important step in producing desirable qualities of real horses. However, we certainly can imagine a credible gene machine that produces each of the horse’s chromosomes composed of DNA totaling up to ∼ 6 billion diploid base pairs, specified precisely in an optimal heterozygous format using information stored in a computer. The number of linear chromosomes likely is arbitrary, as there are perfectly good mammals with as few as four pairs or as many as 100 or more. Also, the haploid number of base pairs probably can be reduced to around 1 billion as opposed to the ∼ 3 billion that is typical of mammals. However, working out the ‘kinks’ of reducing the number of base pairs likely will be much more difficult than simply synthesizing the 6 billion base pair diploid state based on the framework that has evolved successfully to produce the current horse. There appear to be around 25 000 genes in mammals as defined in the classical sense, although with alternative splicing and regulatory elements such as microRNAs, there likely are in excess of 100 000 unique RNA gene products constructed during the life cycle of a horse. Each of these DNA correlates could be specified precisely, including introns and the huge stretches of DNA between most of these genes. We could even add proviruses of retroviruses as examples of providing some random future uncertainty and genetic variability, or just eliminate proviruses. Finishing touches such as centromeres and telomeres would complete the essential elements. The ∼ 16 000 base pair mitochondrial genome also could be specified. We, of course, would need cytoplasm as a home for this DNA, suitably programmed to add histones, etc. The simplest solution would be to use an existing, enucleated oocyte, although some iteration might be needed to replace native mitochondrial DNA if that were an objective. Such oocytes could then be programmed to develop into embryos. Several groups (e.g. Forster and Church, 2006)3 have considered synthesizing cells in this way. Most aspects of such a project already have been accomplished on small scales via transgenic techniques in a variety of species, especially mice. This sort of manipulation was the fundamental reason for the experiments that led to ‘Dolly’, and Dolly was followed up by research resulting in precise manipulation of a DNA sequence in sheep.4 Even whole chromosomes, of the artificial or natural
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variety, have been introduced into the germline of animals. While the magnitude of a project of the kind just described would be a technological tour de force, with continually improving methods of sequencing, cataloging, and synthesizing DNA, such a project might be quite feasible in a decade or two.
Transgenic methods The original transgenic approaches such as using retroviruses as vectors or injecting DNA into onecell embryos have been very powerful indeed. The recent research breakthrough of making pluripotent stem cells from somatic cells for possible therapeutic cloning is but one example.5, 6 However, for the most part, these techniques have been haphazard with regard to making precise genetic changes due to essentially random integration sites and variable copy numbers of added DNA constructs. The embryonic stem cell system in mice changed all that (but only for mice due to the inability to produce true embryonic stem cells in other species) because precise genetic changes could be made to the embryonic stem cells in vitro, which then could be used as inner-cellmass cells that resulted in germline cells. These procedures led to the 2008 Nobel prize in Physiology or Medicine. Cloning of mammals from somatic cells, as suggested earlier, has provided a means for circumventing the problem of not having true embryonic stem cell systems in species other than mice, and has simultaneously greatly simplified procedures such as making transgenic animals. One simply takes a line of cells and does all of the genetic manipulation in vitro, where billions of cells can be selected/screened to produce those with the precise genetic change desired. This approach also has problems and limitations, such as cell line senescence after dozens of passages, but proof of principle has been demonstrated repeatedly (e.g. Schnieke et al. 1997).4 One simply makes a clone, a` la Dolly, using a somatic cell manipulated prior to the cloning step entirely in vitro, to produce the desired genetic changes. Of course, it is a huge step from the current methods of specifying changes in millions of base pairs to a whole genome of billions of base pairs, but this likely would be done in dozens or hundreds of smaller steps. This may be possible within decades, given sufficient funding.
Non-transgenic futuristic genetic and assisted-reproductive technologies There is a whole suite of procedures to manipulate embryos, with various degrees of utility, that have
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ART: Future Technologies Table 325.2 Embryo and cellular manipulations not yet applied in horses Technology
Comments
Chimeras Parthenogenesis Gynogenesis Androgenesis Homozygous diploidy Spermatogonial transplantation Grafting gonads Induced pluripotent stem cells Mitochondrial replacement
Can be within or between species Has been manipulated to produce mice Similar to parthenogenesis Has not yet resulted in producing an animal to birth Has been introduced to the germline via chimeras Has been successful in a variety of species While possible, not of great utility except for experiments Could be used to provide replacement parts for injured or diseased tissues/organs Circumvent some mitochondrial genetic diseases
not been performed with horses but have been done to some extent with other mammals (Table 325.2). Here follows an explanation of certain aspects of the items in Table 325.2. Chimeras are defined as individuals with mixtures of cells of different genotypes. The simplest method of production is to mix blastomeres of two precompaction embryos, for example embryos from two breeds of horses. If both cell types contribute to the developing fetus, coat color will be a mixture of the two breeds, often in splotches; other tissues and organs also will be mixtures. Variations on this concept were used to get genetically modified embryonic stem cells into the germline for the Nobel prize-winning transgenic technology mentioned above. Chimeras also can be made between species such as sheep and goats, resulting in individuals with wool covering some parts of the body and hair covering other parts.7 (Sometimes sexual chimeras result, with part of the body composed of XY cells and part XX cells. These often end up as sterile males.) Which parts of the body are which genotype occurs by chance with most procedures, but one might be able to arrange, for example, chimeric horses with a head, brain, and disposition of one breed, and the body of another. An example of a naturally occurring chimera is the bovine freemartin, in which there is exchange of cells between male and female fetuses due to anastomosis of placental circulations. A special case of directed chimerism is to use asynchronous blastomeres to produce placentas of one genotype and fetuses of another.8, 9 This concept has enabled sheep to gestate goats to term because of essentially having a goat fetus in a sheep placenta. Simply transferring goat embryos to sheep always results in embryonic death. Thus the placental–uterine interface is more critical than the fetal–placental interface in these species. Such manipulations may have utility in rescuing endangered species by using the uteri of closely related species as embryo transfer recipients that otherwise would abort the pregnancy.
The next four technologies in Table 325.2, parthenogenesis, gynogenesis, androgenesis, and homozygous diploid, have in common the constraint that development to term is impossible due to the phenomenon of gametic imprinting, unless transgenic changes are made in the genome. The cellular manipulations themselves are fairly straightforward. Parthenogenesis is essentially mating a female to herself, for example by failure to extrude the second polar body. This obviously results in inbreeding. Because of allelic rearrangements during meiosis, the resulting animal is 71% related to the female providing the ovum (square root of 0.5). Parthenogenetic cells are themselves normal, but result in pregnancies with severely abnormal placentas. Amazingly, while it is not possible to make a parthenogenetic animal (without additional manipulation – see below), it is possible to make a parthenogenetic embryo into a female parent by mixing parthenogenetic embryonic cells with normal inner cell mass cells such that some of the parthenogenetic cells by chance become oocytes in the developing embryo.10 Gynogenesis is crossing one female with another, for example by fertilizing one oocyte with another by cell fusion, or more practically, by pronuclear transplantation.11 The same problems with term pregnancy occur as with parthenogenesis, and similarly are solved by creating chimeras to get such genomes into the germline. Androgenesis is crossing a male with himself or another male. One simple way of accomplishing this is to create a triploid, dispermic embryo, and then remove the female pronucleus. Pronuclear transplantation is another option.11 Such androgenetic embryos develop even less well than parthenogenetic or gynogenetic ones due to poor fetal development. Note that genetically, if both sperm were X-chromosome-bearing, a female would result. It would indeed be interesting to cross a male with himself and get a female! The fourth technology in this suite is the homozygous diploid procedure that results in 100%
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Future Reproductive Technology inbred females. In this case, one starts with a diploid pronuclear stage one-cell embryo, haploidizes it by removing one of the pronuclei, and rediploidizes by allowing the one-cell, now haploid, embryo to go through two cell cycles of DNA synthesis but only allowing one cell division.8 This is done by using the reversible microfilament inhibitor, cytochalasin B, to inhibit division to the two-cell stage. Thus what would normally be a four-cell embryo after the next cell division has only two cells. The same constraints apply as with parthenotes, i.e., these do not go to term, but as with parthenotes, such cells can be incorporated into the germline with the chimeric approach. The reason that these four technologies fail to lead to term pregnancies without transgenic intervention is gametic imprinting. To illustrate, the DNA sequence of an X sperm from a highly inbred strain of mice is identical to that of an oocyte from that strain. However, the cytosines of DNA are methylated differently in these two gametes.12 The methylation imprint specified by the ovary differs from that specified by the testis, and these two are complementary for embryonic development. A crude approximation is that the sperm imprint is needed to make normal trophoblast derivatives (e.g., the chorion of the placenta) and the oocyte imprint is needed to make normal inner-cell-mass derivatives such as the fetus. A theoretical aspect is that this system has a higher information content than one without imprinting. This system also devolves to a theoretical molecular battle of mammalian sexes that is beyond the scope of this chapter.12 If these techniques do not lead to term pregnancies, why bother to bring them up? First, as mentioned earlier, the resulting genetics can be incorporated into germline cells, which can then result in future generations. Second, by minor transgenic manipulations of imprinted genes (mostly deletions), it has been possible to get parthenogenetic mouse embryos to go to term.13 Third, as we further understand methylation patterns of cytosines and can manipulate them, these problems can be circumvented entirely, so it will indeed be possible to cross a male with himself. A sample application of the latter of profound interest would occur if an extinct male equid were found in permafrost with epididymal sperm. The sperm would of course be dead, but with intracytoplasmic sperm injection, one might make androgenomes, both male and female, and recreate the species from this one animal. This might be feasible decades earlier than sequencing and then synthesizing the entire genome as described earlier. The remaining topic in Table 325.2 not already alluded to, that could have utility outside of basic
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research or therapeutic cloning, is spermatogonial transplantation, elegantly illustrated by having one species make sperm of another.14 This technique could be used to make a breeding stallion from one that had died. For example, this could be useful for rescuing endangered equids, although other approaches might or might not be more feasible. Spermatogonial stem cells also represent another source of cells for cloning or transgenic manipulation.15
Some final thoughts There are countless cellular and molecular manipulations possible with mammalian cells; dozens more could be illustrated, such as telomere, centriole, and centrosome manipulations. Some of these techniques could have profound effects on aging, and some techniques can be used to eliminate genetic diseases. Correcting mutations is another possibility. The techniques discussed and related ones have been invaluable in unraveling mechanisms of some diseases, and have led to cures in some cases. Because of fewer ethical constraints in experimenting with animals, there may be an interim situation in which aging of horses is slowed while their owners continue to age normally. The biomedical aspects of these new technologies have not been emphasized, but perhaps should be. Horses have major limitations for such research due to long generation intervals, seasonal breeding, and the sheer costs in terms of space and feed requirements, not to mention manure removal. However, there also are advantages over many species such as methods of transvaginal aspiration and non-surgical recovery and transfer of embryos; large volumes of blood and other tissues and fluids also are available. Because of the number of older horses available, they are excellent models for studying aging. One also cannot overemphasize the synergy of combining the various reproductive technologies described, particularly when coupled with advances in molecular genetics. The next few decades will be even more interesting than the last few decades for this field.
References 1. Carnevale EM. The mare model for follicular maturation and reproductive aging in the woman. Theriogenology 2008;69:23–30. 2. Seidel GE Jr. Genetic and phenotypic similarity among members of mammalian clonal sets. In: Cibelli JB, Lanza RP, Campbell KHS, West MS (eds) Principles of Cloning. San Diego: Academic Press, 2002; pp. 215–25. 3. Forster AC, Church GM. Towards synthesis of a minimal cell. Mol Syst Biol 2006;2:45. doi:10.1038/msb4100090. 4. Schnieke AE, Kind AJ, Ritchie WA, Mycock K, Scott AR, Ritchie M, Wilmut I, Colman A, Campbell KH. Human
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factor IX transgenic sheep produced by transfer of nuclei from transfected fetal fibroblasts Science 1997;278:2130–3. Takahashi K, Tanabe K, Ohnuki M, Narita M, Ichisaka T, Tomoda K, Yamanaka S. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 2007;131:861–72. Yu J, Vodyanik MA, Smuga-Otto K, Antosiewicz-Bourget J, Frane JL, Tian S, Nie J, Jonsdottir GA, Ruotti V, Stewart R, Sluvkin II, Thomson JA. Induced pluripotent stem cell lines derived from human somatic cells. Science 2007;318:1917–20. Fehilly CB, Willadsen SM, Tucker EM. Interspecific chimerism between sheep and goat. Nature 1984;307:634–6. Seidel GE Jr. Mammalian oocytes and preimplantation embryos as methodological components. Biol Reprod 1983;28:36–49. Ruffing NA, Anderson GB, Bondurant RH, Currie WB, Pashen RL. Effect of chimerism in sheep-goat concepti that
10. 11. 12. 13.
14.
15.
developed from blastomere-aggregation embryos. Biol Reprod 1993;48:889–904. Stevens L. Totipotent cells of parthenogenetic origin in a chimeric mouse. Nature 1978;276:266–7. Seidel GE Jr. Application of microsurgery to mammalian embryos. Theriogenology 1982;17:23–34. Jirtle RL, Weidman JR. Imprinted and more equal. Am Sci 2007;95:143–9. Kono T, Obata Y, Wu Q, Niwa K, Ono Y, Yamamoto Y, Park E, Seo J, Ogawa H. Birth of parthenogenetic mice that can develop to adulthood. Nature 2004;428:860–4. Dobrinski I, Travis AJ. Germ cell transplantation for the propagation of companion animals, non-domestic and endangered species. Reprod Fert Devel 2007;19:732–9. Brevini TAL, Antonini S, Pennarossa G, Gandolfi F. Recent progress in embryonic stem cell research and its application in domestic species. Reprod Domest Anim 2008;43(Suppl. 2):193–99.
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Index Note: page numbers in italics refer to figures; those in bold to tables or boxes. A23187, acrosome evaluation 956, 1493, 1494 abdominal auscultation, foals 170, 751 abdominal ballottement neonatal foal 170 pregnant mare 2245–6 abdominal compression, periparturient hemorrhage 2518 abdominal distention bladder rupture 625 hydrops 12, 2369 intestinal atresia 406 meconium impaction 380, 383 neonatal foal 170, 399 older foal 750–1 abdominal pain abdominal wall rupture 2428 after dystocia 2494 neonatal foal 227, 399 older foal/weanling/yearling 749–50 see also colic abdominal palpation, foals 170, 751 abdominal paracentesis see abdominocentesis abdominal radiography gastroduodenal ulceration 401 meconium impaction 380 neonatal foal 171, 400 older foal/weanling/yearling 751–2 abdominal support, pregnant mares 31, 2428, 2448 abdominal surgery see gastrointestinal surgery abdominal ultrasonography see transabdominal ultrasonography abdominal wall congenital defects 643 movement, foal respiratory disorders 576 abdominal wall rupture 2152–3, 2428–30, 2447–8 clinical signs 2428, 2429, 2447–8, 2480 high-risk pregnancy 6, 8 hydrops 34 induction of abortion 2322–3 management 31–2, 2428–30, 2448 surgical repair 2429–30 ultrasonography 2428, 2429 abdominocentesis chyloabdomen 405–6 colic during pregnancy 6, 2452 colic in foals 171, 400, 751 uterine torsion 2436 see also peritoneal fluid, analysis
abortion 2339–49, 2457–9 body pregnancies 2345, 2481 causes 2341–9, 2457–9 diagnostic trends and 2341–2 summary list 2342, 2343–4 contagious equine metritis 2401, 3017 defined 2320 detection 2339 donkey-in-horse pregnancies 2314–17 equine amnionitis and fetal loss 2415 equine herpesvirus 2377, 2378 clinical features 2379–80 diagnosis 2340, 2341, 2383 epidemiology 2381–2 pathogenesis 2378 prevention and control 2384–6 equine viral arteritis 9–10, 2392–3, 3021 diagnosis 1253, 2394 pathogenesis 2393 fescue toxicosis 2422 induction 2320–3 complications 2323 hydrops 2323, 2371, 2445 impending pregnancy failure 2121–2 indications 2322–3 methods 2320–2 PGF2α 1799–800, 2122, 2320–1 pharmacological methods 2320–1 twin pregnancy 2116, 2322 infectious 8–10, 90, 2343, 2457–8, 2784 investigation 2339–41 aims 2339 clinical history 2340 diagnostic uncertainties 2348–9 early pregnancies 2340 later pregnancies 2340 necropsy routine 2340–1 precautions 2339–40 mare reproductive loss syndrome 2410–11 Miniature ponies 2841, 2842 non-infective 2343–4 , 2458 post-colic 2452, 2453 prevention see maintenance of pregnancy rate 2780 recurrent 2348 reproductive efficiency and 2784 therapies preventing 25–8 twin pregnancy 2346–7, 2355–7 see also embryonic loss, early; fetal death abscessation Corynebacterium pseudotuberculosis 765 female reproductive tract, rectal palpation 1906
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hepatic 249, 637 internal, weight loss 763–4 intra-abdominal see intra-abdominal abscessation mammary gland 2164, 2287, 2740 myocardial 515 ovarian 2709 renal interstitial 652 strangles 704–5 testicular 1474, 1475 urachal see under urachus uterine 2671 vaginal 1944, 2154 accessory sex glands 875, 1113, 1114 abnormalities 1113–17 ultrasonography 1483–5 dimensions 1379, 1380, 1482 function 896–7 infections 3024 ultrasonography 1480–3 see also ampullae; bulbourethral glands; male internal genitalia; prostate gland; seminal vesicles Accuread 1280 acepromazine (acetylpromazine) 838 adverse effects on stallions 1132, 1134, 1188 dystocia management 2485 foal rejection 119 periparturient hemorrhage 2518 pregnant mare 57 uterine contractility and 2603–4 acetabular fractures 452, 453 acetazolamide 838 hyperkalemic periodic paralysis 464, 668, 674, 675 acetic acid, intrauterine infusion 2635, 2648 acetylcysteine (N-acetylcysteine; NAC) 838 effects on sperm storage 994 hepatic failure 412 retention enemas, foals 194, 381, 382 topical, corneal ulcers 546 uterine lavage 2613, 2629, 2634, 2649 acetylpromazine see acepromazine Acholeplasma 3025 acid–base disturbances, neonatal foal 192, 206–8 acid–citrate–dextrose (ACD) solution 240 acid phosphatase, uterine 1640 acoustic enhancement artifacts 1919 Acremonium coenophialum see Neotyphodium coenophialum acridine orange (AO) 1506 acrocentric chromosomes 1952 acroplaxome 930
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acrosome 928–9, 932, 933, 1057–8, 1491 biochemistry 1064–5 development 917, 929, 1034–5, 1036 effects of cooling 1073 equatorial segment 929, 932, 1057, 1058 exocytosis see acrosome reaction integrity, assays 956, 1494–6 frozen semen 2997, 2999, 3000 sex-sorted sperm 3044 knobbed 1298, 1300, 1365 matrix, composition 929, 1491 morphological defects 1298, 1300, 1303 acrosome–acroplaxome–manchette complex 930 acrosome reaction (AR) 948–51, 1491–6, 2999 assays 956, 1288–9, 1494–6, 1503, 1504 biochemistry 1074–5 capacitation and 944, 948, 1493 clinical application 1493–4 induction by zona pellucida 948–9, 953, 954, 1491–3 membrane changes 1498 morphological changes 929, 934, 1492 signaling pathways 948–51, 1493 substances inducing 1493 acrosome responsiveness assay (ARA) 956, 1494, 2999 ACTH (adrenocorticotropic hormone) 70, 523 adrenal actions 1665 exogenous 838 fetal responses 1667 pregnant mare 27, 124 premature or dysmature foal 125 pituitary pars intermedia dysfunction 1666, 2790, 2791, 2793 plasma neonatal foal 523, 524, 854 perinatal period, foal 18, 69, 70, 72 premature foal 72, 524 sick foal 72, 525 stimulation tests 524, 854 critically ill foal 525 fetus 71, 124 neonatal foal 523 premature or dysmature foal 124, 524 relative adrenal insufficiency 525 Actinobacillus mare reproductive loss syndrome 2413 neonatal sepsis 156, 159, 253–4, 255 pneumonia in older foals 710, 712 Actinobacillus equuli bacteremia 411 peritonitis 763 renal interstitial abscesses 652 Actinomyces spp., oral 2748 actin polymerization, sperm capacitation and 947 activated clotting time (ACT) 853 activated partial thromboplastin time (aPTT) 427, 428, 853 activin 1681–2 fetal production 1681 male reproduction 898, 998, 1682 endocrine function 900, 998 paracrine–autocrine function 1002 mare 1625, 1681–2 placental function 1682, 2236
acupuncture, problem-breeding mares 2616, 2628, 2628 acute lung injury/acute respiratory distress syndrome (ALI/ARDS) 582–5 clinical signs and diagnosis 582–3, 584 equine neonatal (EqNALI/NARDS) 583, 584 therapy 583–5 veterinary (VetALI/VetARDS) 583, 584 acute renal failure (ARF), foal 648–58 causes 648–52 diagnosis 653–6 outcome and prognosis 657–8 therapy 213, 656–7 acute tubular necrosis (ATN) 650 acyclovir 838 neonatal foal 588, 606 pregnant mare 32 ADAM proteins 953 Adam’s procedure 1562–3 ADAMTS enzymes 2728 adaptive immune system 331, 334–8 fetal 334–5 postnatal 335–8 uterus 2590 adenocarcinoma mammary 2742–3 ovarian 2710, 2714 uterine 1948 adenohypophysis (anterior pituitary) 897–8, 1583, 1584 adenosine, modulation of capacitation 946–7 adenosine triphosphate see ATP adenovirus infections diarrhea 742 respiratory 588, 602, 725–6 adenylate kinases 937 adenylyl cyclase membrane-associated (mAC), sperm 946–7, 949 soluble (sAC), sperm 936, 947 ADH (antidiuretic hormone) see vasopressin adhesions cavum vaginale 867, 1529 cervical see cervical adhesions intra-abdominal, foal 244, 245 oviductal 2693 periovarian 2702, 2704, 2705 periuterine 2672–3 rectal palpation in mare 1906 uterine see uterine adhesions vaginal 1902 adrenal cortex, ectopic nodules 2693, 2699, 2709 adrenal cortical hormones (corticosteroids) 1602, 1665–7 dysmature foals 124 fetal, peripartum changes 18, 69–72, 122–3, 2230 functions 1665 pituitary pars intermedia dysfunction 1666, 2790, 2791 premature foals 72, 124, 524 reproductive effects 1665–7 sick foals 524–5 see also cortisol adrenal glands 1665 ectopic tissue 2693, 2699, 2709 fetal perinatal changes 69–71, 1667 progestagen synthesis 2223–4
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adrenaline see epinephrine adrenal insufficiency, relative (RAI), sick foals 524–5 adrenal medullary hormones, foal 70, 72–3 adrenocorticotropic hormone see ACTH aerobic pathogens identification 1968–9 uterine and clitoral cultures 1968 Aeromonas hydrophila 742 African horse sickness 3024–5 agalactia 2282–3 causes 2282, 2283 fescue toxicosis 2282, 2283, 2420 management 64, 2282–3 agammaglobulinemia 362, 363–4 aganglionosis, intestinal see ileocolonic aganglionosis agar plates 1967–8 aged horses see geriatric horses; geriatric mares; geriatric stallions aggressive behavior breeding stallions 1398, 1411 pharmacological suppression 1416–18 mares causes 1763–5 towards foal 118, 119 novice stallions 1407 see also stallion-like behavior aging premature, cloned animals 2925 spermatozoa 1074, 1075–7 AI see artificial insemination air, intrauterine see pneumouterus air bronchograms 714 air transportation, stallions 1187–90 airway management, neonatal resuscitation 129–32 airway obstruction see recurrent airway obstruction (RAO); upper airway obstruction A kinase anchoring protein (AKAP) 936, 947 Akt 2710 alanine transaminase (ALT), serum 852 albinism 557, 558 see also overo lethal white foal syndrome albumin infusion, human, neonatal foal 208, 209 albuterol 212, 838 alcohol-based hand hygiene products 826 aldosterone 72, 1665 fetus and neonate 70, 79–80 alertness, neonatal foal 478, 480 Alexa Fluors 1495 ALI see acute lung injury/acute respiratory distress syndrome alkaline phosphatase (ALP) seminal fluid 1520, 1521 serum neonatal foal 174, 175, 855 older foal 852 urine 855 allantoamnion 86 examination 92–3 unique features of equine 90 allantochorion (chorioallantois) 86, 100, 101 avillous lesions 2273, 2274 color 92 development 85, 2189, 2194, 2234 examination 92, 102–3
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Index iatrogenic rupture, to induce abortion see under fetal membranes ischemic necrosis 2345 normal avillous areas 90–1, 92, 102, 105, 2273 rupture at parturition 2271, 2482 thickness 92 unique features of equine 90 vascular congestion 104–5, 106 weight 106 see also fetal membranes; placenta allantochorionic placentation 85–6, 2235 allantochorionic (chorioallantoic) pouches 87, 104, 105, 1657, 1658 allantochorionic (chorioallantoic) vesicles 104 allantoic calculi see hippomanes allantoic fluid 79, 2368–9 clarity 51 composition in hydrops 2369–70, 2444 excess see hydrallantois expulsion at parturition 2271, 2482 volume 20, 22, 51, 87, 2368 allantoic sac 85, 2189 ultrasonography 2069, 2072, 2073, 2075 allantois adenomatous hyperplasia 2274, 2275 development 85, 2181, 2189, 2190, 2191, 2234 examination 92, 102 allergic reactions penicillins 257 transfusions 241 allergies, neonatal foal 137 allopurinol, neonatal encephalopathy 150, 195 alopecia, linear 565 alpha-2 agonists neonatal foal 222, 227 pregnant mare 57, 58, 59 α-adrenergic agonists intracavernosal, priapism 1135 urospermia 1127 alpha-fetoprotein (AFP) 2203–4 altrenogest 1811–12, 1813 anestrous mares 1698–9, 1700 corpus luteum persistence 1760 donkey-in-horse pregnancies 2315, 2316, 2316 embryo donors 2874 estrus suppression 1815, 1846–7 estrus synchronization 1816, 1872–3 foal rejection 119, 120 induction of lactation 268, 2284 intravaginal delivery 1879 long-acting formulations 1812, 1881 maintenance of pregnancy 1817, 2121, 2334, 2466–7, 2471 colic 2453 geriatric mares 2813 high-risk pregnancy 26 parturition and 2268 placentitis 2364–5, 2365–6 ovulation failure 2689 postpartum mares 1816, 2293, 2298 sperm motility effects 1295 suppression of stallion-like behavior 1194, 1416–17 transitional mares 1713, 1778–80, 1815 ambiguous genitalia 1961 amblyopia, deprivation 549
Ambu bag 131, 233 American Association of Equine Practitioners (AAEP) EHV-1 outbreaks 725 influenza vaccination 722 plasma and serum products 350 strangles vaccination 709 American College of Veterinary Internal Medicine (ACVIM), strangles management 709 American Miniature Horses, testicular size 1379 American Quarter Horse Association (AQHA) 3014 approval of AI 1263 EVA vaccination 1254 hyperkalemic periodic paralysis 464, 673 overo lethal white foal syndrome 408 amides, semen cryopreservation 2969–70 Amies charcoal transport medium 1966, 1967 amikacin 838 chronic endometritis 2632, 2633 foal diarrhea 386, 736 foal pneumonia 605, 716 infectious hepatitis in foals 412 intrauterine infusion 2632 neonatal foal 255, 256 dose 194, 254 prior to/during transport 232 neonatal meningitis 487 semen extenders 1201, 1338, 1341 septic arthritis in foals 460 septic foal 157, 202 amino acids hormones derived from 1602, 1603, 1607 parenteral nutrition solutions 274 placental transport 89 aminocaproic acid 838, 2518 aminoglycoside antibiotics foal septic arthritis 460 neonatal diarrhea 386 neonatal foal 157, 202, 255–6 doses 254 nephrotoxicity 652 therapeutic drug monitoring 256 older foal diarrhea 736 see also gentamicin ammonia, serum 409, 410 amnion 86 examination 100, 101, 2274, 2275 abortion 2341 rupture at parturition 2271 thickening 2274, 2275, 2360 weight 106 amniotic fluid 2368–9 clarity 51 composition in hydrops 2369–70, 2444 excess see hydramnion ultrasonic features 2360 volume 20, 22, 51, 87, 2368 amniotic plaques 101, 105–6 A-mode ultrasound 1918 amoxicillin, neonatal foal 254, 257 amphotericin B 838 fungal endometritis 2636, 2647 neonatal foal 157 ampicillin 838 chronic endometritis 2633 contagious equine metritis 2405 infectious hepatitis 412 intrauterine infusion 2632, 2633
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neonatal foal 232, 257 continuous rate infusion 257 dose 254 spectrum of activity 255, 257 neonatal meningitis 487 pneumonia in older foal 716 septic foal 157 ampullae (of deferent duct) 868, 875, 877, 1113 dimensions 1380, 1482 plugged (sperm-occluded) 919, 1113–15, 1159, 1421 hemospermia 1122 treatment 1115, 1414 ultrasonography 1483–4 ultrasonography 1480–1, 1482 anabolic steroids behavioral effects in mares 1764–5 effects on spermatogenesis 1179–80 estrous cycle changes 1755 subfertile stallions 1443 anaerobic bacteria female genital tract 1975 identification 1970 uterine and clitoral cultures 1968 analgesia mare caudal reproductive tract surgery 2545–6 uterine prolapse 2433 neonatal foal 221, 225–7 diarrhea 388 meconium retention/impaction 381 prior to/during transport 234 stallion genital tract endoscopy 1453 poor libido 1415–16 anal paralysis 485 anal sphincter, obstetric injuries 2488–9 anal/tail clamp reflex 479, 480 anal vaginalis 407 anaphylactic reactions neonatal foal 136, 137 transfusion 241 androgen(s) 892–3, 1001 adrenal 1665 biosynthesis 893–4 epididymal actions 1008–9 exogenous effects on spermatogenesis 1178–80 male contraception 1888 ovarian biosynthesis 1631, 1632 vernal transition 1707 paracrine–autocrine effects 1001–2 see also dihydrotestosterone; testosterone androgen-binding protein 924 androgenesis 3054 androgen insensitivity syndrome (AIS) 1960, 2218–19 androgen receptors (AR) epididymis 1008–9 mutations 2218 testes 911, 1001 AndroMed-ETM semen extender 1337, 1338, 2968–9 androstenediol, testicular biosynthesis 893, 894 androstenedione ovarian metabolism 1631, 1632 testicular biosynthesis 893, 894 anembryonic vesicles see trophoblastic vesicles, empty
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anemia cardiovascular disease 513 of chronic disease 420, 422, 423, 425 equine infectious 3024–5 neonatal foal 137, 142, 419–25 blood transfusion 239, 424 causes 239, 419–20, 421 clinical signs 168, 238, 420–2 diagnosis 422–4 neonatal isoerythrolysis 354, 419–20 outcome 425 oxygen therapy 215 Oxyglobin therapy 209, 231 preparation for transport 231–2 treatment and management 424–5 anesthesia mare caudal reproductive tract surgery 2546 cervical surgery 2560 neonatal foal 221–2, 223–5 CSF collection 479 hypotension 224 hypothermia and hypoglycemia 224–5 recovery 225 ventilation 225 older foal 221, 225 pregnant mare 55–9 approach to 56 cesarean section 59 dystocia 59, 2485–6, 2505, 2512 effects of pregnancy on 56 fetal effects 56 fetotomy 2498 techniques 58–9 stallion, castration 1558 anestrus 1696–701 causes 1697, 1761–2 endometrial biopsy 1929, 1931, 1937 GnRH implant-induced 1700 GnRH vaccine-induced 1700–1 hypopituitarism 1700 idiopathic 1701 lactational 1699–700, 1757 management 2637 ovarian ultrasonography 2123, 2124 with no ovarian activity in breeding season 1699–701 in winter 1697–9 nutritional 1699 with ovarian activity 1696–7 postpartum 1757, 2296–7 ovarian ultrasonography 2123, 2124 seasonal 1697–9, 1705–6, 1782 behavioral changes 1761–2 deep 1782 definition 1705, 1732 follicular development 1754 hormonal events 1784–5 hormonal treatment 1699, 1785–6 light treatment 1698–9 onset see autumnal transition termination see vernal transition transitional stage 1782 see also transitional mares senescence 1701 ultrasonography 2004, 2005, 2006 angiocardiography 505 angiogenesis, placental 2239–40 angiomatosis, ovarian 2705
angiotensin II 79 birth transition 646–7 neonatal vasogenic nephropathy 649 angiotensin-converting enzyme (ACE) 78–9 fetus and neonate 80 GnRH degradation 1610 angular limb deformities (ALDs) 768–70, 774 congenital 666, 667, 768, 769, 770 dietary factors 785 etiology 433–4, 438, 768 examination and diagnosis 768–9 spontaneous self-correction 776 treatment 769–70 aniline blue–eosin stain, sperm 1499, 1500 anion gap, foal 173, 853 aniridia 546–7, 668 anisocytosis 422, 423 annexin V, fluorescein-conjugated 956, 1502 annual reproductive rhythm 1704–5 entrainment 1733 feral/semi-feral populations 1387–8 mechanisms of regulation 1704–5, 1782–3 phases 1705–6 photoperiod and 1346–7 role of melatonin 1677, 1704, 1782–3 see also breeding season; non-breeding season; seasonal variations annulus, sperm 918, 931, 934, 1028, 1060, 1061 migration 1036 anophthalmia 538 Anoplocephala magna 299 Anoplocephala perfoliata 299 anorchidism 1109 anovulation (ovulation failure) 2685–90 anestrus 1697 causes 1757–8, 2685 geriatric mares 2808 hormone therapy associated 2685, 2689 incidence 2685–6 management 2689–90 seasonal 1697–9, 1732, 1754 light treatment 1698–9 see also anestrus, seasonal anovulatory follicles 1757–8 hemorrhagic see hemorrhagic anovulatory follicles identification 2686–8 incidence 2685–6 luteinized (LAF) 1757–8, 1760, 1762, 2688 management 2690 ultrasonography 2129–30 management 2636, 2689–90 non-luteinized 2688, 2690 partially collapsed 2128, 2129 persistent (PAF) 1757, 2128, 2701, 2709 size 2687 spring transition 2017, 2018 superovulation cycles 1842 transrectal palpation 1911 ultrasonography 1723, 2128–30, 2637, 2686–8 antacids 396, 685, 743 antarelix 1194 anterior presentation see cranial presentation anterior segment dysgenesis 547 anthelmintics
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
broodmares 111, 2748 foals/young horses 113, 295, 300 stallions 1222 see also deworming programs; parasiticides; specific agents anthrax vaccination broodmares 306, 310 foals and weanlings 304 antibacterial activity, uterine secretions 2589 antibiotics see antimicrobial agents antibiotic sensitivity tests 1971–2, 1986, 2675–6 antibodies passive transfer see passive transfer of maternal antibodies production development 335, 336–7 impaired 361–5 see also immunoglobulin(s) anticholinergic drugs, priapism 1135 anticonvulsants, neonatal foal 194 antidiuretic hormone (ADH) see vasopressin antifreeze protein (AFP) 2905 antifungal agents endometritis 2635, 2636, 2646–8 intrauterine infusion 2636, 2647, 2649 neonatal foals 157 ophthalmic disorders 541, 546 antigen-presenting cells (APCs) 333, 2590 antihistamines retained placenta 2525 transfusion reactions 241 antimicrobial agents embryo recovery media 2874 embryo storage media 2881 intraosseous injection 460 mare chronic endometritis 2613, 2632–3 colitis in pregnancy 30 contagious equine metritis 2405 delayed uterine clearance 2661–2 early embryonic loss 2335 embryo recipients 2878 maintenance of pregnancy 2472–4 mastitis 2740 placentitis 32–3, 2363–4, 2365–6 post-breeding endometritis 2811–12 postpartum metritis 2532, 2533 prior to ovariectomy 2564 rectal tears 2579 retained placenta 2525, 2528 neonatal foal 253–61 critical illness 193 diarrhea 386, 388 doses 194, 254 infectious hepatitis 412 infective endocarditis 515 meningitis 487 nephrotoxicity 652 omphalitis 636–7 ophthalmic disorders 540, 541, 546 perinatal asphyxia syndrome 151 pneumonia 587, 605–6 prior to/during transport 232, 235 R. equi pneumonia prophylaxis 691 routine use 113 sepsis 156–7, 160, 202, 253–5 septic arthritis and osteomyelitis 459–60 shock 144 subacute to chronic use 259–61
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Index older foal/weanling/yearling Corynebacterium pseudotuberculosis 765 diarrhea 736, 738–9 Lawsonia intracellularis 739, 762 pneumonia 715–17 R. equi infections 698–700, 764 strangles 707, 764–5 regional limb perfusion, foal 460 semen extenders see semen extenders, antibiotics stallion contagious equine metritis 1251 epididymitis 1158–9 prior to shipment 1188–9 seminal vesiculitis 1116, 1117 Taylorella equigenitalis carriers 2406 see also intrauterine antibiotics antimicrobial sensitivity tests 1971–2, 1986, 2675–6 anti-mullerian hormone (AMH) 2215 antioxidants 941 dietary supplements 994 effects on sperm storage 993, 994 semen 941, 991, 1077 semen extenders 1339 see also selenium; vitamin C; vitamin E antiproteases, corneal ulceration 545, 546 α-1-antiproteinase inhibitor 546 antiseptics intrauterine, chronic infertility 2613, 2628, 2633–5 retained placenta 2526–7, 2528 antithrombin (AT) 428 antithrombin III 853 anxiety, signs of 1407–8 anxiolytic therapy, breeding stallions 1416 aorta diameter in fetal 22, 41, 46 terminal, ultrasonography 1485 aortic-iliac thrombosis (AIT), ultrasonography 1485 aortic valve dysplasia 508 ventricular septal defect involving 507–8 APFTM 1189 APGAR score 477 aplasia cutis 556 apnea, periodic 169, 575 apoptosis, testicular germ cells 916, 924, 1021–3 Appaloosa parentage syndrome 568, 569 Appaloosas congenital stationary night blindness 551, 668 mane and tail dystrophy 555, 568, 569 seminal characteristics 1369–70, 1371, 1373, 1375 strabismus 538–9 appendix testis 911 appointment mating, ovulation induction 1804 approach-avoidance conflict, novice stallions 1407, 1408 AQN1 1088 Arabian fading syndrome 568–70 Arabians accessory gland dimensions 1380 cerebellar disease 670 foal rejection 118
occipitoatlantoaxial malformation 491, 492 seminal characteristics 1369–70, 1371, 1373, 1375 severe combined immunodeficiency 369, 371 arachidonic acid 1603 Arachis hypogaea agglutinin see peanut agglutinin ARDS see acute lung injury/acute respiratory distress syndrome arginine vasopressin (AVP) see vasopressin aromatase 1001, 2225–7 arrhenoblastoma 2710, 2714 arrhythmias see cardiac arrhythmias arterial blood gases (ABG) foal pneumonia 605 measurement 185, 578 neonatal foal 169–70, 185 critical care 184–5 reference values 187, 605, 854 perinatal asphyxia syndrome 150 premature foals 125 respiratory disorders 578 septic foal 155, 159 arterial blood pressure (ABP) monitoring anesthesia of pregnant mare 58 critically ill neonate 186 neonatal foal 503–5 shocked neonate 139–40 neonatal foal 186, 224, 504 older foal 504 see also mean arterial pressure arterial blood samples, neonatal foal 184–5 arterial catheterization anesthetized foal 224 blood pressure measurement 503–4 arterial thrombosis, foal 160, 429 arthritis aged horses 2755, 2756 juvenile 775 septic/infectious see septic arthritis arthrocentesis, neonatal foal 159, 201 arthrogryposis 7, 13, 667 see also flexural limb deformities articular epiphyseal cartilage complex 434 artificial insemination (AI) 1198–206 Asian wild horse 2849 cooled transported semen 1316–21 see also cooled transported semen deep uterine for fertility problems 3039 low dose 3037–8 sex-sorted sperm 3045 technique 1205 disease transmission 3015–26 donkeys 2837–8 dose see insemination dose estrus synchronization 1871 fresh semen 1317 geriatric mares 2807–8 low dose 3038–9 ovulation induction 1860 reproductive efficiency 2782 timing and frequency 1205 frozen-thawed semen 2977, 2980, 2987–8 epididymal sperm 2985 geriatric mares 2807, 2808 low dose methods 3037–8
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reproductive efficiency 2782 technique 1205 timing and frequency 1205, 2988–90 see also frozen semen geriatric mares 2807 ground collection 1271 historical perspectives 1261–6 20th century development 1263 breed improvement 1262–3 disease control 1262 early pioneers 1262 equipment 1263–4 insemination technique 1265–6 newer developments 1266 semen handling 1264–5 hysteroscopic see hysteroscopic insemination interspecies 2303 intraoviductal 2946, 3038 low dose 3036–40 epididymal sperm 3039–40 fresh semen 3038–9 frozen-thawed semen 3037, 3038, 3039 historical perspective 1266 methods 1205, 3037–9 sex-sorted sperm 3044, 3045, 3046 timing 3037 Miniature horses 1380, 2840 oocyte recipients 2943 ovulation induction 1205, 1327, 1805, 1860 post-ovulation evaluation 1205 procedure 1203–5, 1274–6 historical development 1265–6 problem mares 2144 pros and cons 1198–9 reproductive efficiency 2781–2, 2786–7 selection of mares 1203 semen collection for see under semen semen processing see under semen sites 3038 sperm dose see insemination dose sperm transport after 941–3 timing and frequency 1205, 1273 volume 1201, 1274 see also semen artificial vagina (AV) 1199–200, 1268–70 cleaning 1268–9 failure to ejaculate into 1400 history of development 1263–4 liners 1269 method of use 1269 miniature stallions 1377 modified for diagnostic use 1270, 1460 stallion training 1399, 1400–1, 1406 arylalkylamine N-acetyltransferase (AA-NAT) 1671, 1672–3 arytenoid chondritis 797, 798 arytenoid chondroma 797, 798 arytenoid movements grading systems 796 presale endoscopy 797 racing performance and 799 variability in immature horses 795–6 ascarids (Parascaris equorum) 294–5, 711–12 colic 755–6, 758–9 control 300 drug treatment 295, 296, 759 ultrasonography 244 weight loss in foals 758–9 ascorbic acid see vitamin C
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Asian wild horse (Przewalski’s horse) 2302 assisted reproduction 2849 breeding management 2846–7 embryo recovery and transfer 2311–14 genetics 2846 interspecies hybrids 2304, 2305, 2846 mares contraception 1891 reproductive physiology 2848–9 reintroduction sites in Asia 2849 reproduction in wild 2849–50 stallions killing of foals/other males 2849–50 reproductive physiology 2847–8 vasectomy reversal 2848 aspartate aminotransferase (AST), serum liver disease 410, 412 muscle disorders 463, 465, 466 reference values 852 Aspergillus fumigatus, placentitis 2359 Aspergillus spp. antifungal susceptibility 2648 culture techniques 1968 endometritis 2635, 2643, 2644, 2649 foals 711 identification 1970 venereal transmission 1256, 3025, 3030 aspiration pneumonia 576, 592–5 cleft palate 664 clinical signs 592–3 diagnostic tests 593–5 differential diagnosis for dysphagia 593 older foal 712 treatment and outcome 595 see also meconium, aspiration; milk, aspiration aspirin 839 EHV-1 myeloencephalopathy 731 infective endocarditis 515 assisted reproductive technologies (ART) Asian wild horse 2849 future 3051–5 geriatric mares 2814–15 ovulation disorders 2690 peripubertal fillies 1689 sperm chromatin structure assay 1508 uterine tube blockage 1988 see also specific technologies assisted vaginal delivery (AVD) 2486, 2497, 2511 cervical lacerations 2559 referral dystocias 2513, 2515 vs. cesarean section 2505 AST, serum, neonatal foal 174 asystole, fetal 49 ataxia cerebellar abiotrophy 491 vertebral fractures 453 ataxic breathing 125, 575 atelectasis, dependent lung, neonatal foal 222 atlanto-occipital cerebrospinal fluid collection 479–80, 481 atlanto-occipital junction fusion, with dens hypoplasia 667 atopy 555 ATP generation by sperm 938–9, 1063–4 growing horse 783 sperm motility 937, 1065–6, 1293 atracurium besylate 839 atresia ani 406–7, 668
atresia coli 244–5, 406–7, 668 atresia recti 406–7, 668 atrial fibrillation (AF), neonatal foal 513–14, 517 atrial natriuretic peptide (ANP) 79 atrial septal defect (ASD) 508 atropine 839 neonatal resuscitation 130, 133, 224 topical ophthalmic 540, 541, 546 aurothioglucose 839 auscultation see abdominal auscultation; thoracic auscultation Australia cooled transported semen programs 1323 import/export of stallions 1187 import or export of semen 2972 Australian Stock Horse, seminal characteristics 1375 autocrine action 1601 autosomes 1952 autumnal transition 1705, 1732–49 behavioral changes 1748 endocrine events 1741–8 factors regulating 1733–5 failure of 1733 four phases 1748–9 ovarian events 1735–41 photoperiod and 1733 prolonged luteal phase 1737, 1739–40, 1761 timing 1705 autumn follicles see hemorrhagic anovulatory follicles aversions, specific, individual stallions 1410 avocado toxicity-associated mastitis 2738, 2740 avoidance responses, neonatal foal 478, 480 axoneme 931–5, 1028, 1060, 1061 development 1035 mechanism of movement 937, 1065–6 role in sperm motility 1292–3 azalide antibiotics, neonatal foal 260–1 azathioprine 839 azithromycin 839 Cryptosporidium diarrhea 386, 390 infectious hepatitis 412 Lawsonia intracellularis 739, 762 neonatal foal 254, 260–1 pneumonia in older foal 716 R. equi prophylaxis 691 R. equi therapy 699, 764 azlocillin 257–8 azoospermia diagnostic evaluation 1519–20, 1523–4 obstructive 1113, 1115, 1483–4, 1520, 1523–4 see also ampullae, plugged testicular biopsy 1520–1, 1524, 1525 Azostix test, urospermia 1125 azosulfamide 839 azotemia 653 prerenal, neonatal foal 214, 649 see also acute renal failure Babesia caballi 1256 Babesia equi 1256 bachelor bands 1385, 1386 Bacillus, identification 2677 bacillus Calmette–Gu´erin (BCG) 839 Bacillus thuringiensis Berliner var. kurstaki (Btk) 2416
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
bacitracin intestinal clostridiosis 736 topical ophthalmic 541 bacteremia defined 154 septic arthritis and osteomyelitis 159–60, 201, 457, 459 bacteria antimicrobial sensitivity tests 1971–2, 1986, 2675–6 laboratory methods see microbiological laboratory techniques mare genital tract 1972–6 commensal 1975 culture methods 1968 endometritis 2609–10 entry during parturition 2292 uterine and clitoral swabs 1973–6 uterine cytology smears 1925 neonatal diarrhea 385–9 opportunistic, venereal transmission 1254 stallion semen 1285–6, 2144 control methods 1338–9 effects on semen characteristics 1371–2 see also specific bacteria bacterial artificial chromosomes (BAC) 1464, 1466–7, 1954, 1955 bacterial respiratory diseases neonatal foal 586–7, 601–7 older foal 710–17 see also pneumonia bacteriology, gynecological 1967–72 see also microbiological laboratory techniques Bacteroides fragilis 742, 1975–6 Bakewell, Robert 1261 ballerina foal 443 bands 1385 affiliations within 1386 bachelor 1385, 1386 dominance hierarchies 1387 harem 1385 transitional 1385, 1386 barium enema intestinal aganglionosis 407 intestinal atresia 406 meconium impaction 380 older foal/weanling/yearling 752 barium radiography gastroduodenal ulceration 401, 685 older foal/weanling/yearling 752 barren mares 1897, 2780 endometrial biopsy 1933–4 fungal endometritis 2644 genital swab cultures 1964, 1976 insemination with frozen semen 2990 preventive medicine 2749–50 reproductive efficiency 2781 stallion fertility evaluation 1429, 1430, 1431 see also problem-breeding mare basal plate 928 base deficit 206 basophil counts 173, 851 B-cells see B-lymphocytes BCG (bacillus Calmette–Gu´erin) 839 beat cross frequency (BCF) 1066, 1284 bedding supplies, critically ill foal 835 behavior mare abnormalities 1761–5 anestrus 1748
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Index early pregnancy 2245 estrous see estrous behavior granulosa theca cell tumors 2711–12 maternal see maternal behavior in natural conditions 1385, 1386, 1388–9 parturition 2271, 2481 problematic 1845–6 role of progesterone 1638 sexual see under sexual behavior stallion-like 1763–5, 2709, 2712 stereotypic 2771–5 zebra 2852 neonatal foal abnormal 484–5 brain trauma 488 evaluation 478, 480 hypoxic-ischemic encephalopathy 485, 486 illness 167 normal 63–4, 117–18, 477, 2271 stallion cryptorchidism 1102 handling techniques and 1391–2 natural situations 1385–9 pharmacological manipulation 1415–18 sexual see stallion sexual behavior shuttle stallions 1185–6 stereotypic 1423–5 zebra 2859 see also stallion-like behavior Belgian draft horses hereditary muscular disorders 464–5 junctional epidermolysis bullosa 676–7 benzimidazoles (BZs), strongyles 298, 300 benzodiazepines, neonatal foal 222, 223 benztropine mesylate 839, 1135 β 2 -agonists neonatal foal 189–90, 585, 606 priapism 1135 β 2 -pregnancy protein 2258 beta-carotene 1236 beta-lactam antibiotics neonatal foal 255, 256–9 continuous rate infusions 257 doses 254 see also cephalosporins; penicillins bethanechol 396, 686, 839 bezoars 244 bicarbonate (HCO3 -) 839 activation of sperm motility 935, 936 buffer, semen extenders 1336 hyperkalemic periodic paralysis 464, 668, 674 infusions calculating requirements 206 neonatal foal 134, 157–8, 206–8, 211 measurement and conversion 850 oral, neonatal foal 208 renal tubular acidosis 651 solutions 208 sperm capacitation 946, 947 standard, neonatal foal 605, 854 bicarbonaturia, renal tubular acidosis 651 biceps reflex 479, 480 bifenthrin 2416 Biggy’s agar 2680–1 big head disease 1234 bile acids, serum, neonatal foal 174 hepatic disease 409, 410 bilirubin, serum hepatic disease 409, 410
neonatal foal 174, 175 neonatal isoerythrolysis 354 reference values 852 biochemistry, serum cardiovascular disease 500–1 neonatal foal 172–3, 175 critical care monitoring 182 reference values 174 reference values 851–6 biofilms fungal 2644 intrauterine 2139–40, 2150, 2609, 2634 biological clock 1670 see also suprachiasmatic nucleus biophysical profile, fetal 51, 52 biosecurity measures, breeding facilities 3016, 3030 R smectite clay see Biosponge ditrioctahedral (DTO) smectite birth date, pubertal estrus and 1693 birthweight 282 factors affecting 281–3 % of mare bodyweight 281, 2763 placental influences 91, 2237–8 bismuth subsalicylate 839 neonatal foal 194, 271–2 older foal 737 bits, breeding stallions 1392 bladder biopsy 1456 calculi 1456 catheterization foal 628, 629 urospermia 1126–7 disorders, foal 625–9 dummy 625–6 endoscopy see cystoscopy evacuation, urospermia 1126–7 evagination through umbilicus 643 examination, urospermia 1126 inflammatory diseases 628–9 mare, anatomical relations 1586, 1587 round ligaments 632 ultrasonography fetus 47–8 foal 172, 247 bladder rupture 625–8 arrhythmias 517, 627 clinical signs 625, 626 critically ill foal 183, 625 diagnosis 625, 626 differential diagnosis 625–6 prevention 627–8 prognosis 627 treatment 626–7, 628 ultrasonography 247, 248, 626, 627, 628 blastocyst capsule see embryo(s), capsule formation 2171–3 glucose metabolism 2175 growth and development 2174–5 bleeding disorders, neonatal foal 419 bleeding time (BT) 853 blinkers, stallion training 1401 blood collection, for transfusion 240–1 whole 849 blood agar plates 1968, 2675–6 blood cultures foals 156, 199, 837 respiratory disorders 577–8, 604
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
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blood donors 238, 240 neonatal isoerythrolysis 356–7 blood–epididymis barrier 922–3, 1157 disruption 923 blood flow, peripheral, fescue toxicosis 2422 blood gases, arterial see arterial blood gases blood group systems 353, 354 blood loss, neonatal foal 419, 421 clinical signs 420–1 diagnosis 422–3 management 424 see also hemorrhage blood pressure, arterial see arterial blood pressure blood samples cytogenetic analysis 1953 neonatal foal 177, 178, 184–5 blood–testis barrier 869–71, 884, 912–13, 1041–2 disruption 923, 1161 formation at puberty 1019–21 hypertonic fixation test 1019, 1020, 1021 blood transfusion 849 complications 241 cross-matching tests 240, 2824 donor selection 238, 240 neonatal foal 142, 238–9 anemia 239, 424 blood collection and administration 240–1 indications 238 prior to transport 231–2 neonatal isoerythrolysis 142, 239, 356–7 periparturient hemorrhage 2450–1, 2518 blood typing neonatal isoerythrolysis 356, 357, 359, 2824 parentage testing 2820, 2821 for transfusion 240, 2824 blood urea nitrogen (BUN) neonatal foal 174, 175, 653, 654 reference values 852, 855 blood volume, foal 238 B-lymphocytes 333 developing foal 335, 336 disorders 361–5 BMAL1 transcription factor 1772, 1773 B-mode ultrasonography 1915, 1918 ovarian follicles 2010 pre-ovulatory follicles 2033–5 Bmp2 gene 2214 body appearance, general see physical appearance, general body condition aged horses 2755–7 autumnal transition and 1734–5 follicle development and 1754–5, 2028 gestational length and 2764 lactational anestrus and 1700 mare reproductive efficiency and 2760 monitoring breeding stallions 1221, 1228, 1230 growing horses 791 non-pregnant mares 2761–2 pituitary pars intermedia dysfunction 2792 scoring (BCS) 280, 1231, 2760, 2761 seasonal anovulation and 1698, 1699
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body pregnancy 2119, 2121, 2121–2 abortion 2345, 2481 fetal anomalies and 2441 body (rectal) temperature fescue toxicosis 2422 neonatal foal 65 cardiovascular disease 512 respiratory disease 577 sepsis 154, 167, 199 bodyweight autumnal transition and 1734–5 at birth see birthweight developmental orthopedic disease and 777 gain developmental orthopedic disease and 782–3 pregnant mare 281–2, 2763–4 GnRH secretion and 1614–15 growth studies 280 loss aged horses 2755–7 older foal/weanling/yearling 758–65 placental development and 2239 transported stallions 1188 mature, specific breeds 280–1 monitoring breeding stallions 1221, 1228, 1230 critically ill neonate 183 growing horses 791 pubertal estrus and 1693 boldenone undecylenate, effects on spermatogenesis 1180 Bollinger’s granules 1139–40 Bolz phallopexy 1563–4 bone development 433–9 necrosis septic osteomyelitis 457, 458 surgical debridement 460, 461 nomenclature 434–6 bone marrow abnormalities, neonatal foal 420 aspiration/biopsy 423–4 transplantation 371 bone morphogenetic proteins (BMPs) 776 book of mares (number of mares bred) 1184, 1208–9 age of stallion and 1356 AI programs 2787 assessment of quality 1242–3 late-season additions 1430 minimum for assessing fertility 1242, 1243 reproductive efficiency and 2785–6 stallion fertility and 1430, 2784 boots, padded, mare’s hind feet 1224, 1270, 1409 Bordetella bronchiseptica 710 bots (Gasterophilus spp.) 292–4, 296 bottle-feeding 270, 2281 orphan foals 273 prior to/during transport 235 BotuboxTM 1334 BotuCrioTM semen extender 2967, 2969–70, 2974–5 botulinum antitoxin 492, 839 botulinum toxin 491 botulism 491–2 botulism vaccination 320 broodmare 306, 308, 310, 320 foals and weanlings 304, 308, 320 stallion 1222
BotutainerTM 1334 Bouin solution 1930 bovine colostrum 344, 349 rotavirus diarrhea 389 bovine seminal plasma proteins (BSPs) 1088 bovine serum albumin (BSA), embryo recovery medium 2874 bowel evisceration, dystocia 2514–15 bowel rupture, periparturient 2517 box-walking 2774 brachial plexus injury 489 brachygnathia 667 Brachyspira pilosicoli 742 bradycardia fetal 21, 22, 49 neonatal foal 139, 168 brain congenital abnormalities 669–70 damage perinatal asphyxia syndrome 148 traumatic, neonatal foal 487–9 development, thyroid hormones and 75 brain natriuretic peptide (BNP) 513 branchial cleft cysts 614 breathing ataxic 125, 575 Cheyne–Stokes 575 cluster 575 fetal 22, 49 paradoxical 169, 576 patterns, neonatal foal 169, 575 periodic 125, 169, 575 breech presentation 2479, 2492–3 fetotomy 2502–3 breed associations, gatekeeper role 3051 breed differences age at puberty 1692 causes of colic 749 colostral IgG 2278 dystocia 2482 growth rates 282 inguinal herniation 1540–1 mare fertility 1897 mature bodyweight 280–1 pre-ovulatory follicle diameter 2025–7 stallion reproductive parameters 1362–6, 1368–76, 1378–80 twinning rates 2351 see also specific breeds breed improvement, AI as tool for 1262–3 breeding average day of first, by mare 1431 environment 1397 foal-heat mares 2142 geriatric mares 2807–9 historical aspects 1261–2 management 1222–7 pasture 1396 premature, during vernal transition 1707 reinforcement 1247–8, 1263, 2785 selection of stallions for 1208 simulation of 1719 sneak 1386 see also copulation breeding cape/shroud, mare 1225, 1270 breeding frequency 1390 natural service 1223, 1390 stallion fertility and 1243, 1244, 1245, 1432–3 see also ejaculation, frequency breeding phantom see phantom mare
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
breeding programs dual hemisphere 1184–90 natural service 1208–27 breeding records mare 1897–8 stallion fertility analysis 1211, 1212–17 , 1218–19 stallion subfertility 1242–3 breeding roll 1226, 1227 breeding season 1705 anestrus with no ovarian activity 1699–701 artificially imposed 1782 daily spermatozoal production 890 definition 1705 melatonin secretion 996 opioid secretion 999 spermatogonia subtypes 1030, 1032, 1033 stallion fertility during 1431–2 stallion’s nutrient requirements 1229, 1233, 1234 transitions see autumnal transition; vernal transition breeding shed (or area) 1222–3 floor surfaces 1223, 1224–5, 1394, 1402 form 1209, 1210, 1223 slow-starting novice stallions 1409 stallion training 1393–5, 1398–401 team 1225–6 breeding soundness examination (BSE) mare endometrial biopsy 1929 endoscopy 1940 perineal conformation/vulvar seal 2537–9 stallion 1362 endocrine–paracrine–autocrine tests 1438–41 Friesians, Warmbloods and draft horses 1363–6 miniature horses 1377–80 ultrasonography 1469 Breton horses double ovulations 2022 pre-ovulatory follicle diameter 2026–7 bretylium tosylate 839 brightness, ultrasound image 1916 Brilliance Urinary Tract Infection (UTI) Agar 1980 Brix refractometer 64, 343, 833, 2279, 2280 Broad Institute 2827, 2829 broad ligament 1586, 2431 periparturient hemorrhage into 2155–6, 2517 bronchitis 710 bronchoalveolar lavage (BAL) 713 bronchodilators neonatal foal 189–90, 585, 606–7 older foal pneumonia 717 bronchopneumonia 710 see also pneumonia bronchus-associated lymphoid tissue (BALT) 338 broodmare aged see geriatric mares anthelmintics 2748 client relations 1209–11 dental care 2748 exercise 2748 feet care 2748 foal heat breeding see under foal heat historical background 1261–2 identification 1223
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Index imported 1227 infectious disease prevention 2748–9 management cooled transported semen programs 1326–7 maintenance of pregnancy 2467–8 reproductive efficiency and 2781 stallion fertility and 1243, 1244 natural service procedure 1223–7 neonatal isoerythrolysis screens 358–9, 831 nutrition see under nutrition pasture management 2749 periparturient management 111–13 preventive medicine 2747–51 problem see problem-breeding mare vaccinations 111, 302–7, 310–11 , 339, 2750–1 see also mare; pregnancy brown adipose tissue (BAT) 75 bruxism 750 btk gene 363–4 B-type natriuretic peptide (BNP) 513 bucket feeding 270, 273 bulbospongiosus muscle 877, 878 bulbourethral glands 875, 877, 1113 cysts 1484–5 dimensions 1380, 1482 ducts 1113, 1114, 1452 endoscopic anatomy 1451, 1452 fluid secretions 1062 function 896–7 ultrasonography 1482–3 buphthalmia 540 buprenorphine 57, 226 Burn’s technique 2494 burros 2835 female contraception 1891 bursatti see habronemiasis, cutaneous buserelin 1609, 1820, 1822–3 maintenance of pregnancy 2335, 2472 ovulation induction 1823, 1862 transitional mares 1822–3 butorphanol 839 mare caudal reproductive tract surgery 2545–6 fetotomy 2498 laparoscopic ovariectomy 1993, 2569 pregnancy 57 rectal tears 2579 neonatal foal analgesia 226, 227, 234 sedation 222, 223, 234, 350 older foal/weanling/yearling 737, 750 stallion castration 1557 genital tract endoscopy 1453 induction of ejaculation 1271 testicular biopsy 1527 butylated hydroxytoluene (BHT), effects on sperm storage 994, 1074 R ) butylscopolamine bromide (Buscopan 839 Miniature ponies 2840 neonatal foal 227 oocyte collection 2937 rectal palpation 1908–9 rectal tears 2579 cabergoline 839 CABYR protein 947
caco–copper–iron 839 caffeine 840 critically ill neonate 189, 195 neonatal foal 216–17, 233, 575 neonatal pneumonia 606, 607 perinatal asphyxia syndrome 150 premature foal 125 calcein-AM 957, 1287, 1513 calcitonin, modulation of capacitation 946–7 calcium (Ca; Ca2+ ) 787 activation of sperm motility 935–6 deficiency 1234 developmental orthopedic disease and 787 dietary requirements breeding stallion 1230, 1233–4 broodmare 264, 2762, 2763, 2766 growing horses 787 neonatal foal 265–6 dietary restriction, aged horses 2756, 2758 dietary sources 787 dysregulation in foals 526 fractional excretion 855 intracellular ([Ca2+ ]i) acrosome reaction 949, 951, 1493 capacitation 946, 947, 1086 hyperactivated sperm motility 951–2 second messenger function 1605–6 sperm–oocyte interaction 954 mare’s milk 2280, 2281 placental transport 89 plasma neonatal foal 173, 182, 212 reference values 853 retained fetal membranes and 2523 pre-foaling mammary secretion assay methods 830, 2734 factors affecting 2264, 2735 feto-placental monitoring 19–20 induction of parturition and 1833, 2263–4 jennies 2270 laboratory assays 831 predicting parturition 2270, 2733, 2735–6 supplements breeding stallion 1234 growing horse 288 calcium borogluconate, neonatal foal 211, 212 calcium carbonate (CaCO3 ) 850 calcium (Ca2+ ) channels acrosome reaction 949 capacitation 946 hyperactivated sperm motility 951–2 sperm 935–6 calcium gluconate 840 hyperkalemic periodic paralysis 464, 668, 674 transfusion reactions 241 calcium ionophores, acrosome evaluation 956, 1493, 1494 calcium pump (Ca2+ -ATPase), plasma membrane (PMCA) 946, 1606 calcyclin 2203–4 California mastitis test (CMT) 2740 calming pheromone, stallion behavioral problems 1418 calmodulin (CaM) 936, 1605–6 calmodulin-dependent protein phosphatase 936 calmodulin kinase II (CaMK) 936
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calvaria fractures 454 Canada, import or export of semen 2972 candela per square meter 1774 Candida albicans, neonatal foals 156 Candida spp. antifungal susceptibility 2648 culture techniques 1968 endometritis 2635, 2643, 2644, 2648–9 identification 1970, 1979, 1984, 1985 candidate gene studies 2830 recurrent airway obstruction 619 candidiasis (Candida infections) cystitis 628 neonatal foal 156, 157, 170 pneumonia 588, 711 venereal transmission 1256, 3025, 3030 cannon keratosis 566 capacitation 944–8 acrosome reaction and 944, 948, 1493 biochemistry 1074–5 epididymal sperm 2985 factors suppressing premature 945–6, 1092–3 frozen-thawed sperm 2998–9 hyperactivated sperm motility and 944, 952 induction by reactive oxygen species 992 laboratory assays 956, 1288, 1501–2, 3001 molecular markers 944–5 signaling pathways 945–8 sperm aging and 1076 sperm–oviduct adhesion prevention 1086 sperm used for GIFT 2946 Cape mountain zebra 2851 foal 2862 mare gestation 2856–7 puberty and breeding life 2854 seasonality of breeding 2853 stallion puberty and breeding life 2860 reproductive organs 2861 seasonality 2860 weanling 2862 yearling 2862–3 see also mountain zebra capillary refill time 167, 499 capitulum 930, 933, 1059 capnography ex utero intrapartum treatment (EXIT) 132 neonatal resuscitation 134–5 caprine colostrum 344 R diluent 1312 Caprogen caput epididymis 874, 875, 917, 1156 contractility 920 function 894–6, 920–1, 1156–7 microscopic anatomy 927, 928, 1156 sperm transit time 918 ultrastructure 927 carbamazepine, headshaking behavior 1425 carbenicillin 257 carbetocin 1831 uterine clearance 2629, 2660, 2810 carbohydrate metabolism sperm 938–9, 1063–4 sperm–oocyte interaction 954–5 see also glucose
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carbon dioxide (CO2 ) oocyte culture 2933, 2939 placental exchange 88 serum 173, 853 carbon dioxide tension (PCO2 ), neonatal foal 159, 169–70, 216 anesthesia/sedation 222, 223 critical care 185, 189 reference values 605, 854 sepsis 200 see also hypercapnia; hypocapnia carboxyfluorescein diacetate (CFDA) 957, 1287, 1513 cardiac arrhythmias 517 diagnostic tests 513–14 fetal 48–9 neonatal foal 502, 517 bladder rupture 517 critical care 168 diagnostic tests 513–14 shock 139 cardiac catheterization 505 cardiac compressions, neonatal resuscitation 132 cardiac disease, neonatal foal 137 cardiac function fetal, ultrasonographic assessment 48–9 pregnant mare, anesthesia and 56 cardiac output measurement 505 neonatal resuscitation 131, 132 cardiac troponins (cTnI; cTnT; cTnC) 501, 513 cardiogenic shock, neonatal foal 136 cardiopulmonary arrest, neonatal foals 128 cardiopulmonary cerebral resuscitation (CPCR), neonatal foals 128–35 cardiovascular disease acquired 511–17 cardiac auscultation 500 congenital 507–9 diagnosis and monitoring 499–505 diagnostic tests 500–5, 513–14 history 499, 511 physical examination 499–500, 511–13 cardiovascular neoplasia 516 cardiovascular system adaptation to parturition 66 assessment 167–9, 499–505 congenital defects see congenital heart disease beta-carotene 1236 carpal flexion posture, fetal 2489 fetotomy 2499–500 carpal flexural deformity 441, 442 common digital extensor tendon rupture with 471 dystocia 2489 fetotomy 2499 treatment 443 carpal valgus, newborn foal 67 carpapenem antibiotics, neonatal foal 259 carprofen 840 carpus (knee) fractures in foals 448–9 ossification 436–7 presale radiographs 803, 804–5 interpretation 814–15 normal variations 810 cartilage abnormal development 433 development 434 increased thickness 438–9
carts, stall-side, NICU 824, 825 CASA see computer-assisted sperm analysis case–control studies 2799 Caslick index 1577–8, 2537–9 Caslick operation (vulvoplasty) 1580, 2539–41, 2547–8 after ovariectomy 2566 deep see episioplasty foaling management 2539–41 indications 1577–8, 2537–9, 2623 insemination after 1204, 1275–6 points for consideration 2547–8 prior to AI 1203 technique 2539, 2540 variations 2548 Caslick speculum 1900, 1902 castration (orchiectomy) 1557–61 complications 1553, 1560–1 hydrocele 1152–3, 1162, 1561 penile lacerations 1130, 1131 septic funiculitis 1153–4 cryptorchidism see cryptorchidectomy epididymal sperm harvesting 1272, 2983–4 hydrocele 1152 immunological see immunocastration incomplete 1561 indications 1557 inguinal hernia 665 laparoscopic 1549–50 foals and juvenile stallions 1550 in situ 1550 mature stallions 1550 laser assisted 1553–6 complications 1556 postoperative care 1555–6 preoperative aspects 1553 surgical techniques 1553–5 in lateral recumbency 1558–60 postoperative care 1560 self-mutilating stallions 1425 standing 1557 suppression of sexual behavior 1192 testicular tumors 1163, 1168 catalase (CAT) 940, 992 semen extenders 994 seminal plasma 941, 991 test, bacteriological 1981–4, 2676 cataracts 540, 548–9 acquired or secondary 548, 549 congenital 548–9, 668 developmental 548 strabismus due to 538 surgery 549 catecholamines 1603 foal adaptation to parturition 70, 72–3 β-catenin 2214–15, 2710–11 cation exchange resins, neonatal foal 211 CatSper channels 946, 951 cauda epididymis 874, 876, 917, 1156 development 900, 901, 902, 1100 excessive accumulation of sperm 919–20 function 894–6, 920–1, 1157 microscopic anatomy 929, 1156 sperm transit time 896, 918 suppression of sperm motility 935 cauda equina syndrome (CES) 485 caudal gonadal ligament 1100 caudal (posterior) presentation 2480, 2491–3 fetal death 2491, 2492 fetal malformations 2441
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
fetotomy 2502–3 hospital approach 2514 nomenclature 2479 during pregnancy 2075–6 caudal reproductive tract, mare surgery 2545–57 patient selection 2545 procedures 2546–57 sedation and local anesthesia 2545–6 ultrasonography 2157–8 see also female external genitalia; perineum, female; vagina; vestibule, vaginal; vulva caudal ventral median approach, ovariohysterectomy 2574–6 cavernosal shunting 1135, 1564–6 cavernous hemangioma, congenital 559, 562 C-banding, sex chromosomes 1463, 1464, 1953, 1954 CD4+ T-cell lymphopenia of the young, transient 362, 365–6 CD4+ T-cells 333 development 334 uterus 2590 CD8+ (cytotoxic) T-cells 333 development 334 uterus 2590 CD9 protein 953–4 cecal impaction, stallion feeding and 1238 cecal rupture, pregnant mares 2452 cecocecal intussusception 757 cecocolic intussusception 757 cefadroxil, neonatal foal 254, 258 cefazolin 840 neonatal foal 254 pneumonia in older foal 716 cefepime 840 neonatal foal 254, 255, 259 cefoperazone 258 cefotaxime 840 neonatal diarrhea 386 neonatal foal 232, 254, 258 neonatal meningitis 487 septic foal 157 cefotetan, neonatal foal 258 cefoxitin, neonatal foal 254, 258 cefpodoxime 840 neonatal foal 194, 254, 258–9 cefquinome, neonatal foal 194, 254, 259 ceftazidime 840 neonatal diarrhea 386 neonatal foal 254, 258 ceftiofur 840 chronic endometritis 2632, 2633 foal diarrhea 736 intrauterine infusion 2632 neonatal foal 232, 255, 258 doses 194, 254 pharmacology 253 neonatal meningitis 487 neonatal pneumonia 605–6 pneumonia in older foals 715, 716–17 strangles 765 ceftizoxime, neonatal foal 254, 258 ceftriaxone 840 neonatal foal 194, 487 cefuroxime, neonatal foal 194, 254, 258 cellular immunity developing foal 334–5, 336 equine herpesviruses 2382–3 tests 366, 367 cellular immunodeficiencies 365–6, 370
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Index CEM see contagious equine metritis central nervous system (CNS) congenital abnormalities 669–70 support, critically ill neonate 193 trauma 487–9 central venous pressure (CVP) monitoring, neonatal foal 214, 585 centrifugation neonatal foal blood 178 semen 2964–6, 2973, 2974 centrioles spermatozoa 930, 933, 1028, 1059 spermiogenesis 1035 centromere 1951–2 cephalexin, neonatal foal 194, 254, 258 cephalosporins, neonatal foal 255, 258, 259 diarrhea 386 doses 254 meningitis 487 prior to/during transport 232 sepsis 157, 160 cephalothin, neonatal foal 254 cerebellar abiotrophy 491 cerebellar disease, Arabian foals 670 cerebellar hypoplasia 669–70 cerebral edema, perinatal asphyxia syndrome 149–50 cerebrospinal fluid (CSF) analysis 202, 854 collection 479–82 shunting 483 cervical adhesions 2721–3 cervical dilation failure 2724, 2727 diagnosis 1726, 1945, 2625, 2721–2 etiology 2721 management 2722–3 problem breeders 2625 cervical cerclage 2563 cervical dilation failure 2625, 2627, 2724–9 causes 2724–5, 2726, 2727 indications and therapies 2725–9 older mares 2811 prior to induction of parturition 2264–5 see also cervical ripening/softening cervical incompetence 2563 cervical lacerations 2559–63 adhesion formation 2721, 2722–3 clinical signs 2559 diagnosis 1902, 1945, 2559–660 etiology 2559 indications for surgery 2560 prognosis 2562 surgical repair 2560–3, 2722–3 see also cervical surgery cervical ripening/softening 1645, 2724 estrogen therapy 2729 indications 2725–6 mechanism of PGE action 2726–9 PGE therapy 29, 1801, 2264–5, 2726–9 readiness for parturition 2264–5 cervical star 90, 2273 examination 102 intact 2274 thickening 92, 102 tissue samples 107, 109 cervical surgery 2559–63 aftercare 2562 cervical incompetence 2563 choice of technique 2562
complications 2562 equipment 2560–1 indications for 2560 preparation and anesthesia 2560 procedures 2561–2 see also cervical lacerations cervical vertebral compressive myelopathy (CVCM) 728–30 clinical signs 728–9 diagnostics 729–30 management 730 cervical vertebral fractures 453, 488, 489 cervicitis Miniature ponies 2843, 2844 Taylorella equigenitalis 2402 cervicofacial reflex 478, 480 cervix, uterine abnormalities congenital 1944, 2719–20, 2724 correction prior to AI 1203 management in pregnant mares 32 subfertility 2625 agenesis 2719, 2724, 2726 anatomy 1589–90, 2559, 2721, 2724, 2725 digital examination 1726, 1901 double (duplicated) 2607, 2719–20 endoscopy 1940, 1941 pathological conditions 1945 examination in subfertility 2625 fibrosis 2623, 2625, 2724–5 histological features 1938 hyperplasia 2719 hypoplasia 2719, 2724 iatrogenic mucosal damage 2484, 2485, 2497 jenny donkey 2724, 2725, 2835–6 manipulation, inducing luteolysis 1759 pregnancy-related changes 1591 speculum examination 1725, 1726, 1901–2 transrectal palpation 1901, 1909 diagnosis of pregnancy 2247 ovulation detection 1721–2 tumors 1945 ultrasonography abnormalities 2157–8 normal anatomy 2003, 2004 uterine clearance and 2600–1 cesarean section 2505–10, 2511 anesthesia 59, 2505 colic surgery with 2508 complications 2508–9 elective 2508 fertility after 2515 fetal malpostures 2488, 2491 fetal malpresentations 2492, 2493 foal outcomes 2509–10, 2515 hemostatic suture 2506, 2507–8 hysterotomy closure 2506–8 mare outcomes 2509, 2515 maternal abdominal wall injuries 31 maternal pelvic and cervical abnormalities 32 Miniature ponies 2843, 2844 referral dystocias 2512–13, 2515 surgical procedure 2506, 2513 team approach 2505–6 vs. vaginal delivery 2505 cetrorelix 1194, 1609 chain shanks, breeding stallions 1392, 1397, 1398
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chelating agents, intrauterine 2628–9, 2633–5 chemical restraint see anesthesia; sedation chemotherapy effects on spermatogenesis 1177 penile and preputial carcinoma 1138–9 chest compressions, neonatal resuscitation 132 Cheyne–Stokes breathing 575 Chicago blue stain 1287, 1494, 1511 Chifney bits 1392 chimeras 2217, 3054 parentage testing and 2824 Chlamydia spp., venereal transmission 1256, 3025, 3030 chloramphenicol 840 Lawsonia intracellularis 739, 762 neonatal foal 255, 260 dose 194, 254 pharmacology 253 neonatal meningitis 487 R. equi pneumonia 700 chlorhexidine solutions Taylorella equigenitalis carriers 1251, 2405–6 umbilical stump treatment 635, 637 chloride (Cl− ) fractional excretion 855 requirements breeding stallion 1230, 1234 broodmare 2762, 2763, 2766 serum 173, 853 chlormadinome acetate 1813 chloromethyl fluorescein diacetate (CFD) 1513 chlorpromazine, tetanus 493 chlortetracycline (CTC) staining 956, 1288, 1494 frozen-thawed semen 2999 viability staining with 1502 choanal atresia 609–10, 665 cholesterol membrane, effects of cooling 1310, 1311 protection against cold shock 1313 semen cryopreservation 2970 serum 174, 852 sperm capacitation 946, 1075 sperm content 1062 sperm plasma membrane 1055, 1056 assays 1501–2 steroidogenesis via 893, 1602–3, 1631, 1632 testicular metabolism 893, 894 chondritis, infective, neonatal foal 438 chorioallantoic pouches see allantochorionic pouches chorioallantoic vesicles 104 chorioallantois see allantochorion chorion 85 examination 92, 102–3 aborted twin pregnancies 2346–7 abortion 2341 histology at 20 days 2188, 2189 chorionic girdle 87, 1652–6 development 1652–3, 2190, 2191, 2192–3, 2235 extraspecies pregnancies 2311, 2313, 2314 endometrial cup origin 1651–2 endometrial invasion 1653–5, 2193–4 immunological features 1655–6
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chorionic gonadotropin (CG) donkey 1650, 2836 equine see equine chorionic gonadotropin human see human chorionic gonadotropin zebra 1650, 2857, 2858 chorionic villi 2194–5, 2198, 2235 see also microcotyledonary villi chorioretinitis 551 choriovitelline membrane bilaminar 2190, 2191, 2194 trilaminar 2188, 2189 choriovitelline placentation 85 choristomas, ocular 539 Chorulon see human chorionic gonadotropin chromogenic agars 1979–87 detection of reproductive pathogens 1980–6 potential use 1986–7 chromosome abnormalities 1951 autosomal 1465–7 cervical abnormalities 2719 early embryonic loss 1466, 1957–8, 2329–30, 2332 mares 1756, 1955–9 oocytes of aged mares 2329 ovarian abnormalities 2123–4 sex chromosome see sex chromosomes, abnormalities stallions 1464–7 chromosome deletions mares 1957 stallions 1466 chromosomes, equine 1951 banding 1463, 1464, 1952, 1953–4 interphase 1952 meiotic 1467 metaphase 1462, 1463, 1952 methods of analysis 1462–4 morphology 1951–2 terminology 1951–2 see also cytogenetics; karyotype chromosome translocations early embryonic loss 2329–30, 2332 mares 1957–8 stallions 1466–7 chronic lower airway disease, genetic basis 617–19 chyloabdomen (chylous ascites; chyloperitoneum) 405–6 CIDRTM vaginal implants 1814, 1879 estrous synchronization 1826, 1874, 1875 transitional mares 1780 cimetidine 840 neonatal foal 395, 396 older foal 685, 686, 743 ciprofloxacin 840 circadian oscillators auxiliary or slave 1772 master see suprachiasmatic nucleus circadian rhythms 1670, 1771 entrainment 1670 free-running 1771 photoperiodic signals in horses 1773 stored information 1673 circadian timekeeping system 1670–1, 1771–3 anatomical pathway 1671–2, 1771–2 health implications 1773 light required to affect 1773–4
lower vertebrates 1671 regulation by light 1772–3 circannual reproductive rhythm see annual reproductive rhythm circulation, neonatal resuscitation 132 circumcision (segmental posthetomy) 1133, 1562–3 cisapride 271, 840 cisplatin 1138–9 R test 833–4 CITE citrate, semen extenders 1336 citric acid, semen 1062 citric acid cycle, sperm 938 Citrobacter, identification 1981, 2677, 2680 clarithromycin 841 infectious hepatitis 412 Lawsonia intracellularis 739, 762 neonatal foal 254, 260–1 pneumonia in older foal 716 R. equi infections 699, 764 Claviceps purpurea 530, 2418 clearance tests (renal) 653–5 cleavage stage embryos 2170–3 cleft palate 576, 611, 663–4 diagnostic tests 594 dysphagia 593 presale endoscopy 797, 798 surgical repair 595, 611 clenbuterol 841 neonatal foal 606 pregnant mare 27, 2365 priapism 1135 climate broodmare nutrition and 2760–1 growth rates and 284 clitoral cultures 1963–77 antibiotic sensitivity tests 1971–2 contaminant/commensal organisms 1976 correlation with cytology 1976 indications 1964, 1976–7 interpretation of results 1972 laboratory techniques see microbiological laboratory techniques microorganisms isolated 1972–3 non-venereal organisms 1975–6 preparation of mare 1965 sample collection 1965–6 sample handling 1967 significance of isolates 1973–6 venereal organisms 1974–5 clitoral fossa 1579, 1592, 1593 clitoral prepuce (transverse frenular fold) 1579, 1592 clitoral retractor muscle 1587, 1592 clitoral sinuses 1579, 1592, 1593 clitoral swabs see clitoral cultures clitoris 1579, 1592–3 enlargement 1579, 1765, 1961 fetal sex determination 2085–6, 2087, 2088, 2091, 2097 winking (eversion) 1579 clock genes 1670, 1704, 1772 artificial lighting programs and 1776–7 equine 1773 regulation by light 1773 CLOCK transcription factor 1772, 1773 cloning see nuclear transfer cloprostenol 1797 estrus synchronization 1871 FSH and LH release and 1799 induction of abortion 1799–800, 2321 induction of parturition 2265
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
luteinized anovulatory follicles 2690 luteolytic activity 1761, 1799 ovulation induction 1643 plugged ampullae 1115 uterine clearance 2602, 2628, 2630, 2661, 2810 uterine infections 1800 d-cloprostenol 1797 clostridiosis, intestinal neonatal foal 387–9 older foal/weanling/yearling 734–8 prevention and infection control 737–8 treatment 386, 388–9, 736–7 Clostridium botulinum 320, 491 Clostridium difficile diagnostic testing 387, 735 diarrhea in older foal/yearling 734–5 hyperimmune plasma 737 neonatal diarrhea 386, 387–9 neonatal sepsis 159 Clostridium perfringens antitoxins 388, 737 diagnostic testing 387, 736 diarrhea in older foal/yearling 735–6 hyperimmune plasma 737 neonatal diarrhea 386, 387–9 neonatal sepsis 159 Clostridium piliforme 409 Clostridium tetani 492 clotrimazole fungal endometritis 2636, 2647 yeast cystitis 628 clotting disorders see coagulation disorders clotting profile tests 853 club foot appearance 443, 775 coagulase test 1984 coagulation disorders, neonatal foal 426–9 clinical signs and pathogenesis 426 diagnostic process 427, 428 sepsis 160, 429, 517 shock 139 coagulation system 426, 427 coasting, superovulation protocols 1838 coat color genes 2824, 2825, 2829 gray rule 2820 Coccidioides immitis 711 coccidiosis, intestinal 741 co-dominant follicles see double dominant follicles cohort studies 2798–9 coincidence hypothesis, photoperiodic regulation of reproduction 1773 coital exanthema, equine see equine coital exanthema colchicine 841 cold extremities, neonatal foal 138, 169 cold shock, spermatozoa 1070, 1073–4, 1311 minimizing 1074 cold therapy, topical, orchitis 1162 colic neonatal foal abdominal ultrasonography 244–6 causes 750 differential diagnosis 381, 406 incarcerated umbilical hernia 641 meconium impaction 380 other gastrointestinal disorders 406–8 surgical gastrointestinal disorders 399–404
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Index older foal/weanling/yearling 749–57 causes 750, 752–7 clinicopathological evaluation 752 gastrointestinal tract examination 750–2 initial assessment 749–50 prognosis after surgery 757 peripartum mare 2155–6 postpartum mare Miniature ponies 2844 rectal tears 2579 pregnant mare 6–7, 2452–3 cesarean section with colic surgery 2508 management 30 risks of anesthesia 55 surgery late in gestation 6–7, 123 uterine torsion 2435, 2442 see also abdominal pain colitis NSAID toxicity in foals 744 pregnant mares 30 collagen cervical ripening and 2726–8 defects, osteochondritis dissecans 773–4 synthesis, role of copper 788 collagenase infusion, retained placenta 2527 collars, anti-cribbing 2772–3 colliculus seminalis see seminal colliculus colloid osmometer 179 colloid osmotic pressure (COP), neonatal foal 178–9, 187, 208–9 colloid therapy neonatal foal 141–2, 157, 190, 208–9 monitoring 178–9 older foal/weanling/yearling 736 colobomas 541, 550–1 colonic hypoganglionosis 407 Colorado extender 2966 Colorado State (CSU) model artificial vagina 1268, 1269, 1270 color Doppler ultrasonography see Doppler ultrasonography colostomy, rectal tears 2581–3 colostrometer 2279 colostrum 64–5, 342–4 assessing quality 64–5, 342–3, 349, 832–3, 2271, 2278–9 bovine 344, 349, 389 caprine 344 collection and storage 343, 349 composition 2277–80 electrolyte assays 830–1 enteral feeding 268 equine 349 factors affecting quality 343 IgG content 64, 342–3, 2277–8 assessment methods 342–3, 832–3, 836, 2279, 2280 temporal trend 336, 337, 2278 lymphocyte transfer 336 neonatal isoerythrolysis screens 359–60, 831 orphaned neonates 2280–1 prepartum mare 2270, 2284–5, 2480–1 red cell alloantibodies 360 refractometry 64, 342–3, 832–3, 2279, 2280 specific gravity 342, 832–3, 2279 substitutes 344, 349 supplementation 343–4 critical care 192
enteral feeds 267–8 failure of passive transfer 344, 348–9 prior to transport 234 prophylactic 351 transfer of maternal antibodies see passive transfer of maternal antibodies see also milk colpotomy technique, ovariectomy 2565–6 combined thickness of uterus and placenta (CTUP) 22, 49–51 measurement 50–1, 2359–60, 2361 normal values 20, 50, 2360 placentitis 2459, 2460 common digital extensor tendon, ruptured 471–2, 528 common vaginal tunic 867 common variable immunodeficiency (CVID) 361, 364 communication handler–stallion 1391 between horses 1386–7 complement 332 deficiencies 366 uterine secretions 2589 complete blood count (CBC) cardiovascular disease 513 coagulation disorders 427 pneumonia 713 reference values 851 computed tomography (CT), neonatal foal 201 computer-assisted sperm analysis (CASA) 959, 1066–8, 1283–5, 1293–4 basic principles 1283–4 clinical value 959 frozen-thawed semen 2979–80, 2995, 2996–9 parameters measured 1066, 1284 quality standards 3013 sample depth 1067–8, 1285 sperm counting 1282 computer-assisted sperm head morphometry 2998 concentrates, feed breeding stallions 1232, 1234, 1238 broodmares 2762 growing horses 790, 791 concentration immunoassay technology R test) 833–4 test (CITE conception rate 2780 conceptus 2065 corpus luteum maintenance 2200–2 developmental landmarks 2075, 2169 differentiation of fetal membranes 2188–98 embryology 2068–9 estrogen production 1633, 2178–9, 2202, 2225 fixation 2179–80, 2187–8 ultrasonography 2070–1 intrauterine mobility 2174–5, 2179, 2187 maternal recognition of pregnancy 2202–3 prostaglandins and 1644–5, 2187 ultrasonography 2069–70 see also fetal mobility maternal communication 2167–81 ultrasonographic detection 2067–8 see also embryo(s); fetal membranes; fetus concussion 488
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
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condoms 1270, 1377 congenital abnormalities 663–71 central nervous system 669–70 chromosomal abnormalities 1958–9 digestive system see gastrointestinal disorders, congenital female reproductive tract 1943–4 male reproductive tract 1109–11 metabolic and endocrine 668–9 musculoskeletal system 666–7 ophthalmic 537, 668 respiratory system 609–14, 663–5 skin 555, 670 urogenital system 665–6 uterus 1944, 2607 see also fetal malformations congenital adrenal hyperplasia (CAH) 1960, 1961 congenital heart disease 507–9, 670–1 complex 508–9 differential diagnosis 514 congenital stationary night blindness (CSNB) 538–9, 551, 668 conjunctivitis 544 connecting piece, sperm 930, 1058–9 contagious equine metritis (CEM) 1250–1, 2399–407, 2609, 3017–20 causal agent see Taylorella equigenitalis clinical response 1250, 2401–2, 3017 congenital/neonatal 2401, 2403 diagnosis 1250–1, 2404–5, 3018–19 economic significance 2404 embryo import regulations 2922 epidemiology 2402, 3017–18 industry trends and 2404 pathogenesis 2402 prevention and control 1963, 2406–7, 3019–20 stallion screening 1396 transmission 2400–1 modes of 2402 via AI 1251, 2401, 3015, 3016, 3029 treatment 1251, 2405–6, 2613 containers cooled semen 1318 cryogenic storage of semen 3005–8 see also shipping containers continuous positive airways pressure (CPAP) 218, 219 contraception 1888–92 mare 1889–92 stallion 1888–9 contracted foal syndrome 667 fetotomy 2499 contracts, frozen semen supplies 3014 contractures, limb, neonatal foal 172 contrast, ultrasound image 1916 controlled vaginal delivery (CVD) 2486, 2497, 2511 cesarean section after failed 2506, 2512–13 fertility after 2515 mare and foal survival 2515 procedure 2505–6, 2512 referral dystocias 2513–14, 2515 vs. cesarean section 2505 cooled transported embryo technology 2876, 2880–5 adverse effects 2882–3 current technique 2883–4 embryo size effects 2883 historical perspective 2880–2 for import/export 2921 pregnancy rates 2884–5
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cooled transported semen advantages 1308 breeding with 1316–21 estrus synchronization 1871 geriatric mares 2807–8 insemination dose 1201, 1323, 1326 insemination technique 1321, 1327 mare owners/managers and 1326–7 ovulation induction 1327, 1805, 1860 reproductive efficiency 2782 timing and frequency of insemination 1205, 1321 containers 1264, 1318, 1324–5, 1330–4 dilution ratio 1318–19, 1326 disadvantages 1308, 2994 disease transmission 3015–26 documentation 1320, 1326 effects of antioxidants 994 evaluation of quality by recipient 1319–21 by sender 1316–17, 1325 sperm chromatin structure assay 1507–8 fertility 1321, 2991–2 historical development 1264 large-scale programs 1323–8 longevity of sperm motility testing 1286 packaging 1324 principles 1308–14 quarantine 1325 semen handling 1317, 1326 sex-sorted sperm 3045 sperm damage 993, 1311 minimizing 1074, 1312–14 pathogenesis 1070–4, 1308–12 stallion owner/stud requirements 1323–5 stallion suitability 1325–6 storage temperature 1312, 1313–14, 1318, 1330–1 troubleshooting problems 1327–8 warming rate 1313 cooling, stallion semen 1070–4, 1308–12 effects on cell metabolism 1311–12 effects on membranes 1070–2, 1309–11 effects on proteins 1072 effects on sperm 1070, 1073–4 rate 1313, 1318, 1331 Coomassie blue, acrosome evaluation 956, 1491, 1494, 1503 copper (Cu) 788–9 developmental orthopedic disease and 778, 788–9, 2764 dietary requirements breeding stallion 1230 broodmare 264, 2762, 2763, 2764, 2766 growing horses 788 neonatal foal 265 dietary sources 788 mare’s milk 2280 periparturient hemorrhage and 2449 supplements broodmares 778 growing horses 288, 788, 789 coprophagy 265 copulation donkey 2837 Miniature ponies 2839–40 in natural conditions 1388 stallion training 1394–5, 1399–401 zebras 2852
see also breeding; natural service; sexual behavior cornea abrasions, sedated/anesthetized foal 222 acquired abnormalities 545 congenital abnormalities 545 dermoids 539, 545 examination in foals 535–6 opacities 545 corneal ulcers (ulcerative keratitis) 545 critically ill neonate 172, 196 melting 545, 546 neonatal sepsis 160 treatment 545, 546 coronavirus diarrhea 389–90, 742 corpora nigra 536 corpus albicans (corpora albicantia) 2008, 2056 corpus callosum, agenesis of 669 corpus cavernosum clitoridis 1593 corpus cavernosum penis (CCP) 877, 878 irrigation, priapism 1135 priapism pathophysiology 1134–5 ruptured 1131 shunting 1135, 1564–6 corpus epididymis 869, 874, 917, 1156 bilateral congenital hypoplasia of distal 1111 function 894–6, 920–1, 928, 930, 1157 microscopic anatomy 1156 sperm transit time 918 corpus hemorrhagicum (corpora hemorrhagica) 1588, 2056 transrectal palpation 1721, 1911 ultrasonography 1722, 2056, 2057, 2059 corpus luteum (corpora lutea) (CL) 1588 accessory 2056 progesterone secretion 1638 ultrasonography 2063, 2250 autumnal transition 1735–7 conceptus-mediated maintenance 2200–2 detection of a functional 1728, 2055 embryo transfer-related compromise 2877–8 estrogen biosynthesis 1631 hCG treatment effects 1806 irregularities 2057–8 lifespan, autumnal transition 1736–7 persistent 2057 anestrus 1696–7, 1762 autumnal transition 1737, 1739–40, 1761 causes 1759–61 iatrogenic causes 1760–1, 1847–50 idiopathic 2057 prostaglandin therapy 1798–9 spontaneously (SPCL) 1737, 1739–40 ultrasonography 2132 primary 1588, 2056 progesterone secretion 1638, 2222 role of eCG 1658–9, 2063 ultrasonography 2057, 2062, 2063, 2249 progesterone production 1638, 1735–6, 2222 regression see luteolysis secondary 1588, 2056 progesterone secretion 1638, 2222 role of eCG 1658–9 ultrasonography 2063, 2250 supplementary 2056, 2063 differential diagnosis 2704, 2709
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
progesterone secretion 2222 ultrasonography 2249–50 terminology 2056–8 transrectal palpation 1911 ultrasonography 1722, 1723, 2006–8, 2055–64 abnormalities of function 2131–2 blood flow 2047–8, 2060, 2061–2 early pregnancy 2061, 2062–3 estrous cycle 2057, 2060, 2062 luteal tissue echogenicity 2060, 2061 measurement of size 2056, 2059–61 methodology 2058–62 technique 2058 uteropathic abbreviation 2057 uteropathic persistence 2057 zebras 2855–6, 2857 see also diestrus; luteal phase corpus spongiosum penis (CSP) 875, 878, 878 hemorrhage, urethral rents 1140–1 lacerations 1130 ruptured 1131 surgical incision, hematuria of geldings 1142 corticosteroid-binding globulin see cortisol-binding globulin corticosteroids, adrenal see adrenal cortical hormones corticosteroid therapy foal acute respiratory distress syndrome 583–5 alloimmune thrombocytopenia 428 anemia 424 anterior uveitis 203 CNS trauma 488–9 osteochondrosis and 779 prematurity or dysmaturity 125 septic shock 144–5 transfusion reactions 241 mare early- or mid-term pregnancy 1667 estrous cycle changes 1758 induction of parturition 26–7, 123, 1667, 2265 insemination with frozen sperm 2990 ovulation failure 2689 periparturient hemorrhage 2518 post-mating endometritis 2616, 2662, 2812 prevention of embryonic loss 2335 problem breeders 2631 stallion cutaneous habronemiasis 1140 testicular trauma 1161 see also dexamethasone; methylprednisolone; prednisolone corticosterone, fetal 69 corticotropes 1666 corticotropin see ACTH corticotropin-releasing hormone (CRH) 523, 1615 cortisol 70, 523, 1665 ACTH responsiveness dysmature foal 124 fetus 71, 124 neonatal foal 523, 524 premature foal 124, 524 reference values 854 sick foal 525 fetal synthesis 69, 122–3 parturition and 2269
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Index pituitary pars intermedia dysfunction 1666, 2790, 2791 plasma interpretation 1665 neonatal foal 523, 524, 854 perinatal period, foal 18, 69, 70, 72, 123, 1667 pregnant mare 2228–9 premature foal 72, 524 sick foal 72, 525 reproductive effects 1615, 1666 cortisol-binding globulin (CBG) 523, 1665 perinatal changes, foal 71, 72 Corynebacterium, identification 2676, 2677–8 Corynebacterium pseudotuberculosis 765 cost capturing, NICU 826–9 cosyntropin 838 cough equine influenza 722 neonatal foal 577 older foal 712 R. equi pneumonia 695 recurrent airway obstruction 618 courtship 1388 Cowper’s glands see bulbourethral glands cow’s milk composition 268, 269, 2281 enteral feeding 269 modified, feeding with 2281 COX-2 selective inhibitors gastrointestinal safety 744–5 high-risk pregnancy 26 poor libido in stallions 1415–16 CP see crude protein cranial artery of penis 877 cranial nerve examination, neonatal foal 171 cranial (anterior) presentation 2075–6, 2480 dystocia 2486–91, 2514 fetotomy 2499–502 nomenclature 2479 creatine kinase (CK), serum hereditary muscle disorders 463, 465 MB isoenzyme 501, 513 neonatal foal 174, 175 nutritional myodegeneration 466 reference values 852 creatinine blood neonatal renal problems 175, 653, 658 normal neonatal foal 174, 647, 653, 654 premature or dysmature foals 124, 653 reference values 852, 855 clearance 653–4 ejaculate, urospermia 1125 cremaster muscle, external 872, 874, 882, 1099, 1145 surgical transection 1150–1 cribbing (crib-biting) 2771–3 control measures 2772–3 function 2772 gastric ulcers and 1424, 2772 stallions 1423–4 CRISP3 gene 1089, 2828 CRISP proteins inhibition of capacitation 946 sperm–oocyte interaction 953 sperm–oviduct interaction 1088, 1089
critical care supplies, foals 835–6, 837 critical illness-related corticosteroid insufficiency (CIRCI) 524–5 critically ill neonatal foal assessment 167–75 laboratory data 172–5 physical examination 167–72 bladder rupture 183, 625 diagnostic laboratory resources 830–7 gastroduodenal ulceration 395 gastroduodenal ulcer prophylaxis 195, 396–7 infectious disease risk classification 825–6 monitoring 177–87 nutrient requirements 266–7 referral and transport 196 relative adrenal insufficiency 524–5 treatment 189–96 urinary catheterization 627–8 see also neonatal intensive care unit crossed extensor reflex 479 cross-sectional studies 2798 crude protein (CP) 786 developmental orthopedic disease and 778, 786 requirements breeding stallion 1230, 1232 broodmare 264, 2762, 2763, 2766 growing horses 786 cryodamage 2961 cryogenic storage containers 3005–8 cryopreservation 2959–63 embryo 2887–918 epididymal sperm 2985–6 oocyte 2953–5 principles 2959–62 procedures 2953–4 semen see semen cryopreservation cryoprotectants (CPs) 2961–2 diffusion properties 2959–60 embryo freezing penetration 2916 slow/conventional 2887, 2888–98 toxicity 2917 vitrification 2899, 2910–15 lipid-based 2967–9 oocyte cryopreservation 2953, 2954–5 permeant and impermeant 2962, 2969 semen cryopreservation 2969–70 cryotherapy cutaneous habronemiasis 1140 penile and preputial carcinoma 1138 cryptochrome genes 1772 Cryptococcus neoformans 711 cryptorchidectomy 1533–7 flank approach 1535 incomplete 1537 inguinal approach 1533–5 laparoscopic 1535–7, 1546, 1547–9 instrumentation 1546 patient selection and management 1547 recumbent 1548–9 standing 1536–7, 1547–8 laser assisted 1555 parainguinal approach 1534, 1535 presurgical preparation 1533 selection of approach 1537 suprapubic paramedian approach 1534, 1535 techniques 1533–7 cryptorchidism 1100–5, 1531–8 abdominal 1101–2, 1150
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
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complete 1101, 1531 incomplete 1101–2, 1531 spermatic cord torsion 1151 testicular histology 1101 ultrasonography 1473 behavioral features 1102 bilateral 1102 causes 868, 900–2, 1102, 1149, 1531–2 clinical signs 1532 diagnosis 1103–5, 1150, 1532–3 false 1537 inguinal 1016, 1102, 1149–51, 1531 testicular histology 1101 INSL3 expression 1003 miniature horses 1378 prevalence and incidence 1100, 1149–50 surgical management 1533–7 see also cryptorchidectomy testicular biopsy 1524 testicular histology 1100–1 testicular neoplasia 1166, 1167, 1168 treatment 1150–1, 1443 ultrasonography 1103, 1472–3, 1532–3 unilateral 1100, 1102, 1531 XXY syndrome 2217 Cryptosporidium parvum diarrhea 386, 390, 741 crystalloid fluids 842 neonatal foal 140–1, 157, 190, 205–6, 207 older foal/weanling/yearling 736 CTUP see combined thickness of uterus and placenta cuboidal bones abnormalities (incomplete ossification) 774 angular limb deformities 768, 769 congenital hypothyroidism 529 crushing injuries 448–9, 450–1 development 436–7 R 1814, 1879 Cue-Mare estrus synchronization 1826, 1875 cumulus–oocyte complex immature and mature oocytes 2932–3, 2938, 2939 mature oocyte collection 2937–8 cumulus oophorus in vitro removal 2934, 2949 penetration by sperm 953, 954 curly coat syndrome 557, 670 Cushing disease, equine (ECD) see pituitary pars intermedia dysfunction Cushing suture pattern, hysterotomy closure 2507–8 CushionFluidTM 2964 cutaneous asthenia see hereditary equine regional dermal asthenia syndrome cutaneous sensation, neonatal foal 478–9, 480 cutaneous trunci reflex 478, 480 cyanotic mucous membranes cardiovascular disease 499 critical illness 168 respiratory disorders 577 shock 138 cyathostomes (cyathostomins; small strongyles) 296, 297–8 cyathostomiasis foal diarrhea 742 larval 297–8, 300, 742 weight loss in foals 759–60 winter 297 cycle see estrous cycle(s)
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cyclic adenosine monophosphate (cAMP) 1605 acrosome reaction 949, 950 sperm capacitation 947 sperm motility 935, 936, 1066 cyclic guanosine monophosphate (cGMP) 1605 cyclodextrins, semen cryopreservation 2970 cyclooxygenase (COX)-1 1604 endometrial expression 2205–6 cyclooxygenase (COX)-2 1604 endometrial expression 2205–6 ovulation 1642 selective inhibitors see COX-2 selective inhibitors cyclophilin B (PPIB) gene 678 cyproheptadine headshaking behavior 1425 inappropriate lactation 2285 pituitary dysfunction 2758, 2794 cystadenoma, ovarian 2710, 2713–14 cysteine-rich secretory proteins see CRISP proteins cystic gland distention, endometrium 1931, 1932, 1936–7 cystitis foal 628–9 pyospermia due to 1120 cystoscopy foal 629 stallion 1456 cysts bulbourethral glands 1484–5 epididymal 1157, 1158 laryngeal 614 mesonephric duct 2692, 2701 rectal palpation in mare 1906 subchondral bone 772, 773, 774 testicular 1476, 1477 ultrasound specular reflections 1919, 1920 urachus 642 see also dermoid cysts; ovarian cysts; paraovarian cysts; pharyngeal cysts; uterine cysts cytogenetics 2828 mare and foal 1951–61 meiotic chromosomes 1467 molecular 1464, 1467, 1954–5 procedures 1953–5 sample submission 1953 stallion 1462–7, 1524 standard approaches 1462–4, 1951–5 see also karyotyping cytokinemia, perinatal asphyxia syndrome 147, 148 cytokines 333–4 placentitis 2592 post-insemination uterine inflammation 1094 uterine 2590–1 cytoplasmic droplets immature sperm 917, 919, 928, 932, 1028, 1030, 1157 retained, ejaculated sperm 943, 1300, 1302, 1303 shedding by sperm 925, 929 cytoskeleton, sperm head 929–30 cytotoxic drugs, effects on spermatogenesis 1177 cytotoxic (CD8+ ) T-cells 333, 334 uterus 2590
dacryocystitis 542, 543, 544 dacryocystorhinography 543 Dag defects 1301 daily sperm output (DSO) defined 1350 ejaculation frequency and 1356–60 estimation 1459, 2785 miniature stallions 1379 photoperiod effects 1348 reproductive efficiency and 2785, 2786, 2787 seasonal variations 1345–6 testicular size and 1351–3 daily sperm production (DSP) 890, 892, 1039 age-related changes 1043, 1046, 1048, 1353–4 defined 1350 seasonal changes 873, 1346 Sertoli cell numbers and 1044 testicular size and 1351–3 Dandy-Walker syndrome 669 dantrolene 841 DAPI-banding, chromosomes 1464 daylength 1704 pubertal estrus and 1693 required, for photostimulation 1775–6 see also photoperiod dazzle reflex 478, 480, 535 D-dimers 853 DE see digestible energy decapacitation factors 945–6, 1092–3 deep Caslick procedure see episioplasty deep digital flexor tenotomy 444, 445 deep inguinal ring 1145, 1164 deep uterine insemination see under artificial insemination deferent duct (ductus deferens) 868, 875, 877 ampullae see ampullae endoscopic features 1452, 1453 physical examination 1459 rectal palpation 1146 defibrillation, neonatal resuscitation 134 dehydration, neonatal foal 205 clinical signs 167, 205 fluid replacement 191, 205, 206 laboratory assessment 213–14 prior to/during transport 230–1 dehydroepiandrosterone (DHEA) pregnancy 1633–4, 1659, 2225, 2227 testicular biosynthesis 893, 894 deiodinase enzymes 73, 74 delmadinone acetate 1194 dendritic cells (DCs) 333 R see S-adenosylmethionine Denosyl dense fibers, sperm tail 932, 933, 934, 1028 middle piece 1059–60 numbering convention 1060, 1061 principal piece 1061 production 1036 role in sperm motility 1293 Densimeter, ARS 1280 dental care breeding stallion 1221 broodmare 2748 geriatric horses 2755–6 dentition, inadequate, aged horses 2757 dermoid cysts 1166 cutaneous 570 testicular 1110 dermoids, ocular 539, 545
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
Deschamp needle, closure of rectal tears 2580–1 desflurane 224 deslorelin 1609, 1821–2 AI recipients 1205, 2989 implants (OvuplantTM ) 1821–2, 1882 estrus suppression 1850 follicular effects 1755–6 inducing anestrus 1700 ovulation induction 1862–4 prolonged interovulatory interval 1755–6, 1821–2, 1863 seasonal anestrus 1785 superovulation protocols 1837, 1839 injectable formulations 1822, 1882–3 mature oocyte collection 2936, 2938 oocyte recipients 2941, 2942 ovulation induction 1862–4, 1865–6 geriatric mares 2807–9 twin pregnancy rates 2351–2 ovulation synchronization 1873–4 superovulation 1837, 1841, 2689 transitional mares 1710, 1822 desmin 1934 determinant 2796 detomidine 841 epidural 845 foal 222 mare caudal reproductive tract surgery 2545–6 dystocia 59, 2485 laparoscopic ovariectomy 1993, 2569 pregnancy 57 uterine contractility and 2604 stallion behavioral problems 1417–18 castration 1557 cryptorchid castration 1533 genital tract endoscopy 1453 induction of ejaculation 1271 developmental abnormalities male reproductive tract 1109–11 ovaries 2699–701, 2702 stallion-like behavior in mares 1764, 1765 see also congenital abnormalities developmental origins of health and disease model 434 developmental orthopedic disease (DOD) 728, 772–9 calcium and phosphorus intake and 787 causes 264, 287, 288 copper intake and 778, 788–9, 2764 crude protein intake and 786 defining incidence 775–6 different entities 772–5 digestible energy intake and 778, 784–6 feeding to avoid 288, 790–1 growth rate and 777, 782–3 spontaneous self-correction 776 terminology 772 vitamin A intake and 789 see also osteochondrosis deworming programs broodmares 111, 2748 foals/young horses 113, 295, 300 stallions 1222 dexamethasone 841 induction of parturition 27, 123, 1667, 2265 neonatal meningitis 487 ovulation failure 2689
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Index post-mating endometritis 1096, 2616, 2662, 2812 problem-breeding mares 2631 reproductive effects 1666 suppression test (DST) 2792–3 testicular trauma 1161 dextrose (glucose) infusions 841 electrolyte content 207 hyperkalemic periodic paralysis 464, 674 late-term mare 29 neonatal foal 273 critical care 191, 195 electrolyte disturbances 211, 212 hypoglycemia 212 maintenance fluids 206 parenteral nutrition 274, 275–6 prematurity or dysmaturity 125 prior to/during transport 231, 234, 235 sepsis 157–8 shock 141 diacylglycerol (DAG) 1605 acrosome reaction 949, 950 diadem defect 1301 diagnostic laboratory resources 830–7 diagonal paramedian approach, ovariectomy 2567–8 diaphragmatic hernia surgical management 403 ultrasonography 244, 246 diaphysis 434 ossification 436 diarrhea antimicrobial-associated 700, 734–5 larval cyathostomiasis 297 neonatal foal 385–92 enteral feeding-related 271–2 infectious causes 385–91 management 193–5, 206, 208, 213, 386 non-infectious causes 386, 391–2 nutritional causes 392 risk factors 385 sepsis 155, 159, 392 ultrasonography 245–6 older foal/weanling/yearling 734–45 clinical signs 734, 752 infectious causes 734–42 non-infectious causes 734, 742–5 prevention and management 737–8 treatment 736–7 postpartum or foal heat 391 pregnant mare 30 Strongyloides westeri 298, 386, 390–1 diarthrodial joints, development 437–8 diazepam 841 neonatal foal perinatal asphyxia syndrome 148, 149 sedation 221, 223, 234, 350 seizures 193, 194, 234, 486 older foal 221 pregnant mare 58, 59, 2505 stallion castration 1558 libido problems 1401, 1416 diclazuril, equine protozoal myeloencephalitis 731 diclofenac 841 dicrotophos 2416 Dictyocaulus arnfieldi 711–12
diestrus 1728–31 cervical appearance 1725, 1726 characteristics 1729 controlling follicle development 1730–1 endometrial histology 1939 first of breeding season 1728–9 hCG therapy 1807 ovarian ultrasonography 1728, 2006–8, 2057, 2059, 2060, 2062 ovulation 1759 prostaglandin treatment 1729–30, 1798 shortening 1729–30 uterine anatomy 1590, 1591 uterine clearance 2601 uterine fluid accumulation 2657–8 uterine ultrasonography 2004 vaginal examination 1901, 1902 see also corpus luteum; luteal phase diet breeding stallions 1231–2, 1238 broodmares 2747 calming, hyperactive stallions 1411, 1417 developmental orthopedic disease and 778, 782, 783–91 fescue toxicosis 2423 geriatric horses 2755–9 growing horses 790–1 hyperkalemic periodic paralysis 674–5 lactating mares 2281–2, 2765–6 periparturient mare 111–12 polysaccharide storage myopathy 465 retained placenta 2525 self-mutilating stallions 1425 see also feeding; nutrients; nutrition diethylstilbestrol 2472 diffusion, of solutes 2959–60 digestible energy (DE) 783–6 intake, developmental orthopedic disease and 778, 784–6 metabolic roles 783 requirements breeding stallions 1229–32 broodmares 264, 2762, 2763, 2766 growing horses 783–4 digestion, poor, aged horses 2756–7 digestive system see gastrointestinal (GI) tract digital hyperextension see flexor tendon laxity digoxin 841 dihydrocalcein AM 957 dihydroepiandrosterone (DHA) supplements, subfertile stallions 1442 dihydrotestosterone (DHT) biosynthesis 893 epididymal actions 1008 male genital tract development 2215 dimethyl formamide, semen cryopreservation 2969 dimethylsulfoxide (DMSO) 841 embryo cryopreservation 2888, 2904 embryo toxicity 2917 embryo vitrification 2914–15 nasopharyngeal irritation 272 neonatal foal 195 hepatic failure 412 hypoxic-ischemic encephalopathy 149, 150, 487 peripheral nerve injuries 489 prior to transport 232 oocyte cryopreservation 2954–5
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semen cryopreservation 2969 uterine lavage 2613, 2634, 2648 Dimitropoulo-extender D11 1337 1,3-dinitrobenzene, effects on Sertoli cells 1177 dinoprost (tromethamine) 1797 estrus synchronization 1871 induction of abortion 1799–800 induction of luteolysis 1798, 1799 see also prostaglandin F2α (PGF2α ), exogenous dinoprostone 1800 induction of parturition 1801, 2729 diphenhydramine 841 2,3-diphosphoglycerate (2,3-DPG) 577 di(2-ethylhexyl) phthalate, effects on Sertoli cells 1177 dipyrone 841 directional freezing, stallion sperm 2979 disaccharidases, brush border 264–5 disease outbreaks 2799–801 disinfectants Cryptosporidium oocysts 741 intestinal clostridiosis 737–8 intrauterine infusions 2628, 2635 disinfection artificial vagina 1268–9 endoscopy equipment 1449–9 dismount semen samples monitoring 1227, 1245–6 reinforcement breeding 1247–8 displacement activities, anxious stallions 1408 disseminated intravascular coagulation (DIC) 373 diagnosis 422–3 neonatal foal 419, 421 distal interphalangeal joint, acquired flexural deformity 441, 443–4, 445, 775 distal phalanx fractures, foal 446–7 ditrioctahedral (DTO) smectite R ) 839 (Biosponge absorption of colostral IgG and 344 diarrhea in older foal 737 enterally-fed foals 272 neonatal clostridiosis 386, 388 diurnal rhythms 1771 D-mode ultrasound see Doppler ultrasonography DMSO see dimethylsulfoxide DNA, sperm see spermatozoal DNA DNA-dependent protein kinase (DNA-PK) gene 369 DNA fragmentation index (DFI), sperm 1506 DNA parentage testing 2820–5 DNA stains sex-sorting of sperm 3043 sperm analysis 2995, 2997, 2999 dobutamine 841 neonatal foal 216 anesthesia 224 critical care 142, 143, 192, 195 prematurity or dysmaturity 125 sepsis 203 vasogenic nephropathy 657 pregnant mare anesthesia 59 docosahexanoic acid (DHA) peroxidation, aged sperm 1077 sperm content 1062 supplements, breeding stallions 1237–8, 1294
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documentation cooled semen shipments 1320, 1326 estrus detection 1717 fetal sex determination 2082–3 frozen semen 3011–12 placental examination 109 rectal palpation of mare 1905 see also breeding records DOD see developmental orthopedic disease dog-sitting posture, fetus 2487–8 fetotomy 2502 Dolly the sheep 2925, 3053 domestication of horses 1261 dominance hierarchy 1387 dominant follicles 1838–9, 2011, 2012 aspiration, superovulation protocols 1839 autumnal transition 1737–8, 1739 double see double dominant follicles dynamics 2014–17 early pregnancy 2017 echogenic ovulatory 2127–8 one vs. two 2021 selection 1717–18, 1719–20, 2015 shortening diestrus and 1729 ultrasonography 2015, 2016 domperidone 841 agalactia 64, 2283 autumnal transition 1734, 1737 challenge test 2793 controlled release formulations 1883 fescue toxicosis 27–8, 33, 2420, 2424–5, 2449 foal rejection 120 GnRH secretion and 1614 high-risk pregnancy 27–8 induction of lactation 268, 2284 lactational anestrus 2637 poor milk production 270 transitional mares 1699, 1713, 1789–90, 1791 donkey (Equus asinus) 2835–8 chorionic gonadotropin (CG) 1650, 2836 Dictyocaulus arnfieldi carriage 711 embryo transfer 2307–9, 2314–17 interspecies hybrids 2302, 2303, 2304, 2835 see also hinny; mule see also jack donkey; jenny donkey donkey factor 420 donkey-in-horse model of pregnancy failure 2314–17 dopamine exogenous 842 critically ill neonatal foal 195 GnRH secretion and 1614 neonatal vasogenic nephropathy 657 perinatal asphyxia syndrome 151 premature or dysmature foals 125 fescue toxicosis 27 foal adaptation to parturition 70 pituitary actions 1665–6 pituitary pars intermedia dysfunction 1666 reproductive seasonality of mares and 1734, 1737, 1745–6, 1789 dopamine antagonists 27–8 controlled release formulations 1883 fescue toxicosis 27–8, 33, 2423–4 lactational anestrus 2637 seasonal anestrus 1699 transitional mares 1712–13, 1789–92
dopamine D2 receptors 1789 ergot alkaloid interactions 2418, 2420 Doppler ultrasonography 1918–19 cardiovascular disease 502 corpus luteum 2047–8, 2057, 2061–2 early pregnancy 2071, 2073 fetal external carotid artery 46 feto-placental monitoring 21–2, 40 ovarian follicles 2010 ovulation 2044–6, 2047–8 pre-ovulatory follicles 2035–6, 2037, 2038 spermatic cord 1472 testicular neoplasia 1475–6 dorsal artery of penis 877, 878 dorsal displacement of soft palate (DDSP) 589, 611 aspiration pneumonia 593 diagnostic tests 576, 593–4, 612 permanent, presale endoscopy 797, 798 presale endoscopy 794, 795 treatment 595, 611 dorsal medial intercarpal joint disease 814 dorsal nerve of penis 878 dorsal veins of penis 878 dorsoiliac position, fetus 2481 dorsopubic position, fetus 2480 dorso-transverse presentation 2493 dorzolamide 540 double dominant (co-dominant) follicles 2015, 2016, 2021 double ovulations from 2022 endocrinology 2021 double rate, per cycle 1431 dourine 1254 AI as means of control 1262 transmission via AI 3016, 3023–4, 3030 doxapram 842 birth resuscitation 131–2 critically ill neonate 189, 195 foal pneumonia 606–7 neonatal foal 216–17 perinatal asphyxia syndrome 150 periodic apnea 575 premature foal 125 prior to/during transport 233 septic foal 159 doxorubicin, penile and preputial carcinoma 1139 doxycycline 842 Lawsonia intracellularis 739, 762 neonatal foal 194, 254, 260 ophthalmic disorders 546 pneumonia in older foal 716 R. equi pneumonia 700 draft horses birthweight 281 cesarean section 2508–9 congenital abnormalities of uterus 2607 hereditary muscular disorders 464–5 history of breeding 1261 stallion reproductive parameters 1362–6 Draschia megastoma 294 dropsy 2368 drug-induced liver disease, neonatal foal 412 drugs adversely affecting spermatogenesis 1175–82 neonatal resuscitation 130, 132–3 dry mares 2780
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
dry matter (DM) intake breeding stallions 1231–2 calculation 791 growing horses 783–4, 790 dual hemisphere breeding programs 1184–90 see also shuttle stallions ductus arteriosus 508 closure 66, 500 patent (PDA) 500, 508, 512 ductus deferens see deferent duct Dulbecco’s phosphate buffered saline (DPBS) 2880–1 dummy, breeding see phantom mare dummy bladder 625–6 dummy foal syndrome see neonatal encephalopathy duodenal obstruction, liver disease 413 duodenal ulceration see gastroduodenal ulceration duodenitis, ultrasonography 246 R see fentanyl Duragesic duration hypothesis, photoperiodic regulation of reproduction 1773 DVM Rapid TestTM for equine IgG 348, 834 dwarfism Miniature ponies 2841, 2842 sex chromosome abnormalities 1956–7 dynamic range, ultrasound machine 1917–18 dynein flagellar movement 937, 1065–6 regulation of activity 936 sperm tail 933, 1060 dyschondroplasia 776 see also osteochondrosis dysgerminoma 2710, 2714 dysmature foals (growth-retarded foals) 122 carpal fractures 448–9 clinical course 124 laboratory assessment 124 management 125–6 nutrient requirements 266–7 physical appearance 167 plasma creatinine 124, 653 prognosis 126 respiratory insufficiency 125, 576, 589 tendon laxity 469 dysphagia (swallowing problems) congenital pharyngeal abnormalities 611 diagnosis, neonatal foal 171, 576 diagnostic tests 593–4 differential diagnosis 593 milk regurgitation 589, 592, 593 treatment 595 dyspnea 576–7 dystocia 2479–94 anesthesia for fetal manipulation 59 caudal presentation 2491–3 cesarean section 2505–6, 2509 chemical restraint 2485–6 clinical examination of mare 2483–5 dead fetus 25, 2485, 2486 detection 2482–3 duration 2513, 2515–16 ex utero intrapartum treatment (EXIT) 132 fescue toxicosis 33, 2419–20, 2482–3 fetal anomalies 2493–4 fetal manipulation 2486–93 fetotomy 2485, 2497–503
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Index foal survival 2509–10, 2515–16 incidence 2482 induced abortion 2323 induction of parturition and 29 kit 2483 management 2482–94, 2511 mare fertility issues 2484–5, 2509, 2516 mare survival 2509, 2515, 2516 Miniature ponies 2842–3 oxygen therapy for foal 28 prompt intervention 2483 referral 2483, 2511–16 hospital procedure 2512–13 management of mare after 2513 preparation of hospital 2511–12 resolution approaches 2513–15 resolution procedures 2515 transverse presentation 2493 uterine involution after 2296 uterine prolapse complicating 2431 dysuria 628 Earle’s salts 2933 early embryonic death (EED) see embryonic loss, early early pregnancy factor (EPF) 2256–8, 2461 Eastern equine encephalomyelitis (EEE) foals 732 transmission in semen 3024–5 vaccination 309–12 broodmare 306, 310, 311, 2750 foals and weanlings 303, 308–9, 312 stallion 1222 Eastern tent caterpillar (ETC) 13–14, 2414–16 prevention of exposure 35, 2415–16 see also mare reproductive loss syndrome R Eazi-BreedTM CIDR Cattle Insert see CIDRTM vaginal implants ecchymotic hemorrhages neonatal foal 139 purpura hemorrhagica 705 thrombocytopenia 373, 427–8 eCG see equine chorionic gonadotropin echocardiography 502–3, 513, 514 eclampsia 2282 economics cooled transported semen programs 1323–4 NICU 826–9 suboptimal stallion fertility 1240, 1241 e´ craseur transection of ovaries 2566 edema cardiovascular disease 499–500 neonatal foal 167 perinatal asphyxia syndrome 151 see also ventral edema efferent ducts (ductuli efferentes testis) anatomy 872, 873, 874, 917 electron microscopy 926, 927 function 894 egg protein, neonatal foal 344 egg yolk-based semen extenders cooled semen 1074, 1312–13, 1337 frozen semen 2968 history of development 1264 Ehlers–Danlos syndrome see hereditary equine regional dermal asthenia (HERDA) syndrome Ehrlichia risticii see Neorickettsia risticii
EHV see equine herpesvirus(es) eicosanoids 1602, 1603–4 mechanism of action 1607 uterine secretions 2589–90 eicosapentaenoic acid (EPA) supplements, breeding stallions 1237–8 Eimeria leukarti, foal diarrhea 741 ejaculate see semen ejaculation 904–5, 1124 delayed 1420–1 disorders 1413, 1420–2 failure 1420–1 frequency 1390 for AI 1360, 2973 fertility and 896, 1432–3 shuttle stallions 1185–6 sperm production/output and 1356–60 incomplete 1245, 1421 jack donkey 2837 medication-related dysfunction 1422 in natural conditions 1388 pharmacological induction ex copula 1272, 1413–14 in copula 1414 priapism-induced impotence 1136 urospermia 1127 pharmacological manipulation 1413–14 premature 1420 psychogenic dysfunction 1421–2 retrograde 1125, 1245, 1421 ejaculatory duct 1113, 1114 endoscopic examination 1455–5 elbow lock 2487 electroacupuncture see acupuncture electrocardiography (ECG) 501–2, 513–14 24-hour Holter monitoring 502 fetal 21, 40 neonatal resuscitation 134 electrocoagulation laparoscopic ovariectomy 1995 uterine cysts 2667 electroejaculation 1268 electrolytes abnormalities, neonatal foal 182, 191–2, 209–12 excretion, neonatal foal 80 in intravenous fluids 207 mare’s milk 2280, 2281 molar equivalent conversion factors 850 pre-foaling mammary secretion see mammary secretion electrolytes, pre-foaling serum reference values 173, 175, 853 yolk sac fluid 2176 see also calcium; chloride; potassium; sodium electromyography (EMG) 2597 electron microscopy (EM) acrosome evaluation 1494 semen processing 1298–9 sperm morphological analysis 1299–302, 1303–4 elimination marking sequences 1385, 1387 emasculators 1559 embryo(s) 2065, 2167–82 assessing viability 2175 autonomy concept 2168–9 capsule 85, 2234
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cryopreservation and 2176–7, 2916–17 dislodgment 2181, 2188, 2190, 2234 fixation and orientation process 2179–80 formation 2174, 2187 monozygotic twinning and 2351 protein binding 2177–8 remnants 2190, 2191–2 communication with mother 2168–81 complete holding medium 2882 cooled transport 2876, 2880–5 culture medium 2880–1, 2882, 2948 developmental landmarks 2075, 2169 early cleavage stage 2170–3 endometrial cell-to-cell contact 2181, 2182 energy metabolism 2172–3, 2175 fixation 2070–1, 2179–80, 2187–8 freezing 2887–918 deleterious steps 2917 future research needs 2917–18 history 2887–99 for import/export 2921–2 in vitro development after 2900–1 problems in studying 2899–916 reasons for intolerance to 2176–7, 2916–17 slow or conventional 2887–99 viability testing after 2899–916 vitrification 2899, 2910–15 gene expression changes 2203–4 growth rate 2249, 2250 holding medium 2880–1, 2882 identification 2876 import and export 2921–3 intrauterine mobility see conceptus, intrauterine mobility in vitro production 2945 manipulative technologies 3053–4 membrane formation 2188–94 nutrition 2187, 2188, 2190 organogenesis 2181, 2183, 2192–3 orientation 2071, 2179 proper 2181, 2188–90 ultrasonography 2071–3, 2249 prostaglandin production 1644, 2187, 2205 quality aged mares 2328 sperm DNA damage and 1508 recovery exotic equids 2311 interspecies transfers 2306 procedure 2874–6 rates 2871 reduction, twin pregnancy 2101–7 selective oviductal transport 1633, 1643, 2187, 2200 aged mares 2328 mechanisms 2172–3 PGE2 therapy and 1801 sexual differentiation 1959–60, 2011, 2212–15 sexually indifferent stage 2211–12, 2213 shipping containers 2923 size sensitivity to freezing and 2916 at time of transfer 2873 tolerance of cold storage and 2882, 2883 somite formation 2179, 2180 splitting 3052
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embryo(s) (Cont.) transportation 2880, 2921 ultrasonography 2067–74 allantoic sac phase 2069–70, 2073–4 embryo–amniotic ascent 2069–70, 2073 embryo/fetal–amniotic descent 2070, 2073–4 embryo proper 2071–3 estimating day of pregnancy 2074, 2075 twin ovulations 2100 yolk sac phase 2068–73 washing 2876 yolk sac phase 2068–9, 2175–9 see also conceptus; fetus embryology 2068–9 embryonal carcinoma, testicular 1110, 1168 embryonal diapause 2448–9 embryonic loss, early (EED) 2118–22, 2327–36, 2455–7, 2638–9 cloned embryos 2925 cooled-embryo transport 2882–3 defined 2118, 2455 diagnosis 2118–19, 2120, 2456–7 etiological factors 2327–33, 2455, 2456 chromosome abnormalities 1466, 1957–8, 2332 endocrine 1759, 2058, 2330 endometrial 2330–1 genetic 2329–30 heat stress 2332 immunological 2332 intrauterine fluid 2145–6, 2331 nutrition 2332–3 oocyte abnormalities 2329 oviductal 2328–9 pre-uterine events 2327–30 sire-related 2332 uterine cysts 2331–2, 2665 incidence 2327, 2455–6 insemination with frozen-thawed semen 2991 management/treatment 2119–22, 2333–6 mare reproductive loss syndrome 2410–11 prevention see maintenance of pregnancy reproductive efficiency and 2783–4 stallion fertility calculation 1431 transported mares 1817 ultrasonic indicators of impending 2333, 2334, 2456 embryonic loss rate 2780 embryonic stem cells 3053 embryonic vesicle 2065 destruction, twin pregnancy 2101–5, 2107, 2108 growth rate 2249, 2250 orientation 2071 transrectal palpation 2247 ultrasonography 2067–8 early detection 2069, 2247–9 impending pregnancy failure 2118–19, 2120, 2121 twins 2100 vs. uterine cysts 2146, 2248–9 see also conceptus; embryo(s) embryo transfer (ET) 2871–9 Asian wild horse 2849 cloned embryos 2925 donkey in horse 2314–17
donor mares post-flush management 2878–9 pre-flush management 2872–4 repeated use 2873 superovulation 1836, 1842 uterine flush 2873, 2874–6 estrus synchronization 1870, 1876 exotic equids 2311–14 frozen-thawed embryos 2887, 2888–98 geriatric mares 2813–14 horse in donkey 2306–7, 2308 horse or donkey in mule 2307–9, 2310 ICSI embryos 2951 laparoscopic 1999 Miniature ponies 2840 mule in donkey 2310–11 non-surgical 2876–7, 2881, 2882 recipient mares post-transfer management 2877–8 pre-transfer management 1827, 2871–2 progesterone levels 2330, 2878 selection and assessment 2871, 2872 window of synchrony 2872, 2878 between species 2305–17, 3054 success rates 2871, 2884–5 supplies and equipment 2874, 2877 surgical 2876, 2880–1, 2882 uterine tube pathology 1988 EmCare Complete Ultra medium 2937 EmCare Holding SolutionTM 2882, 2942 emission (of semen) 904, 1124, 1413 incomplete 1421 emphysema, subcutaneous 580, 590 emphysematous fetus 2483 empty trophoblastic vesicles see trophoblastic vesicles, empty encephalitis, viral foals 731–2 vaccinations 309–12, 2750 endocarditis, infective 459, 514–15 differential diagnosis 508 weight loss 763 endocrine action 1601 endocrine disorders congenital 668–9 foals 523–31 geriatric mares 2805–6 idiopathic stallion infertility 1436–6 endocrine factors, osteochondrosis 778–9 endocrine–paracrine–autocrine factors male reproductive function 996–1009, 1436 stallion subfertility/infertility 1435–43 endocrinology autumnal transition 1741–8 foal adaptation to parturition 69–80 lactation 2229–30, 2277 parturition 2268–9 placenta 89 pregnancy 1634, 2222–30, 2320, 2461–7 stallion 897–900 endoderm 2174, 2178 Endo-GIA stapler, laparoscopic ovariectomy 1996–7 endometrial biopsy 1929–34 acquisition 1930–1, 1967 case studies 1935–9 chronic endometritis 2612 contraindication 1929 culture samples 2610–11 epicrisis 1934 evaluation 1931–3 fertility and 1933–4
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
fungal endometritis 2645–6 geriatric mares 2805 granulosa cell tumors 2712 hysteroscopically-guided 1942, 1943, 1944 indications 1929–30, 1964 Kenney grading system 1933, 1935–6, 2626–7 problem breeders 2626–7 recent research findings 1934 see also uterine cultures endometrial cups 87, 1656–8, 2195, 2196–7 degeneration 1654, 1657–8, 2197–8 development 1591, 1654, 1656, 2193–4 discovery 1651 eCG secretion 1651, 1654, 1656, 2197, 2235 immune function 1097 interspecies pregnancies 2306–7, 2309, 2310–11, 2314 maternal leukocyte response 1656–8, 2193–4, 2197 persistent 1757, 2125–6, 2669 hysteroscopy 1946 immunological aspects 2591 management 2637 ovulation disorders 2685 ultrasonography 2126, 2149, 2150, 2669, 2670 trophoblast origins 1651–2 endometrial cysts see uterine cysts endometrial cytology see uterine cytology endometrial edema see uterine edema endometrial epithelial cells, cytological smears 1923, 1924 endometrial glands (uterine glands) 1930 cystic distention 1931, 1932, 1936 diestrus 1939 exocrine secretions see uterine histotroph; uterine secretions failure, early embryonic loss 2638 pathological changes 1931, 1932, 1936–7 postpartum changes 2291–2, 2294 during pregnancy 2188, 2189–90, 2194, 2196–7 chorionic girdle invasion 1653, 1654, 2193 function 86, 2190, 2191, 2235, 2236 ultrastructure 88 sperm reservoirs 1094 see also periglandular fibrosis endometrial glandular cysts see under uterine cysts endometrial growth factor (EGF) 86 endometrial insufficiency 6, 10 management 32 endometrial lymphatic cysts see under uterine cysts endometrial prostaglandin synthesis inhibitor (EPSI) 2205 endometritis 2608–16 acute bacteriology 1973–6 culture/cytology correlation 1976 cytology 1923–4, 1925, 1926–7 hysteroscopy 1946 non-venereal organisms 1975–6 screening of mares 1963–4 venereal organisms 1974–5 biopsy features 1931, 1932, 1935 breeding-induced 2614–16
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Index AI with frozen-thawed semen 2991 bacterial 1254, 2609–10 diagnosis 2615, 2627, 2657–8 endometrial biopsy 1934 geriatric mares 2809–13 low dose insemination 3038 pathogenesis 1093, 1094–6, 2614, 2621–2, 2655–7 physiological 1093, 1094–6, 2614, 2621, 2655–6 treatment 1246, 1832, 2474–5, 2615–16, 2628–31 chronic 2608–14 antibiotic therapy 2472–4, 2632–3 bacterial 2609–10, 2631–5 diagnostic approach 2610 early pregnancy loss 2783 endometrial biopsy 2612 endometrial cytology 2611–12 hysteroscopy 1946, 1947, 2612 immunological aspects 2587–91, 2592, 2608–9 management 2612–14, 2631–5 microbiology 2610–11 prolonged luteal phase 1760 commensal/opportunist bacteria causing 1975–6, 3023 diagnosis 2142–3 early embryonic loss 2331, 2335 embryo donors 2872–3, 2879 embryo recipients 2872, 2878 fungal 2635, 2643–50 diagnosis 2645–6 etiology 2643–5 prognosis 2650 treatment 2646–50 jenny donkey 2836 lactational anestrus and 2637 PGF2α therapy 1800 postpartum see postpartum metritis premature luteolysis 1758–9 retained fetal membranes due to 2522 susceptible to see uterine clearance, delayed Taylorella equigenitalis 1974–5, 2401, 2402, 2609 ultrasonographic diagnosis 2143 see also intrauterine fluid accumulation; uterine infections; uterine inflammation endometrium 1589, 1590 abnormalities, early embryonic loss 2330–1 aged mares 2330–1 blood flow, conceptus-induced changes 2207 cervical canal 1590 chorionic girdle invasion 1653–5, 2193–4 cytology see uterine cytology damage due to manual removal of placenta 2526 edema see uterine edema embryonic contact 2181, 2188, 2189–90, 2191 fibrosis see periglandular fibrosis (PGF), endometrial histology 1929–30 asynchrony with ovarian activity 2669–70 see also endometrial biopsy hysteroscopic appearance 1942 inflammation see endometritis lymphostasis 1945, 1946
obstetric trauma 2498 oxytocin synthesis 1679, 1680 pathological changes 1931–3 placental interface development 85, 86, 87 ultrastructure 88 postpartum changes 2291–2, 2294–5 pregnancy-related changes 1591 progesterone actions 1639 prostaglandin biosynthesis 2205–6 secretions see uterine histotroph; uterine secretions trophoblast apposition 2188–9, 2190, 2191 trophoblast attachment 86, 87, 88, 2195 endometrosis 91 endophthalmitis, mare reproductive loss syndrome 2412 β-endorphin pituitary pars intermedia dysfunction 1666 testicular 1003 endoscopes cleaning and disinfection 1449–9 features required 1449 fiberoptic 1448, 1449 storage 1449 endoscopy digital technology 1449 equipment 1448–9 cleaning and disinfection 1449–9 storage 1449 mare genital tract 1940–9 congenital abnormalities 1943–4 indications 1940 pathological conditions 1944–9 routine anatomic features 1940–3 see also hysteroscopy; vaginoscopy stallion genital tract 1448–55 clinical findings 1453–5 equipment 1448–9, 1453 hemospermia 1123, 1453–3, 1455 indications 1448 patient preparation 1453 procedure 1453 relevant anatomy 1450–51 seminal vesiculitis 1116, 1117, 1455–5 urethral rents 1141, 1454–4 upper airway see presale endoscopy; upper airway endoscopy uterine see hysteroscopy see also laparoscopic surgery endothelin receptor B (EDNRB) gene 676 endotoxemia luteal phase abnormalities 1759 neonatal foal 157, 234–5 postpartum see postpartum metritis pregnancy loss 25 pregnant mare 6, 7 management 30 progestagen therapy 1817 uterine prolapse 2433 endotracheal intubation fetus, during dystocia correction 132 neonatal foal 218–19 anesthesia 221–2 birth resuscitation 129–31 endotracheal tubes, neonatal foal 129, 218, 222 changing 219 end piece, sperm 932, 1028, 1061 end-tidal carbon dioxide (ETCO2 ), neonatal resuscitation 134–5
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enemas meconium retention/impaction 381–2 pica-induced diarrhea 391 energy digestible see digestible energy net (NE) 783 energy balance, GnRH secretion and 1614–15 energy expenditure, resting (REE), sick neonate 266 energy metabolism embryos 2172–3, 2175 growing horse 783 neonatal foal 264 sperm 937–9, 1063–4 sperm–oocyte interaction 954–5 energy requirements breeding stallion 1229–32 broodmare 264, 287 growing horses 783–4 healthy neonatal foal 265 lactating mares 2281–2, 2765–6 non-pregnant mare 2762 pregnant mares 2763 sick neonate 266–7, 274, 275–6 sperm 937–8 enophthalmos 541, 544 enoxaparin 843 enrofloxacin 842 chronic endometritis 2632, 2633 intrauterine infusion 2632 neonatal foal 194, 255 older foal/weanling/yearling 739 seminal vesiculitis 1116 enteral feeding foal 192, 267–73 colostrum 268 complications 271–3 dysphagia 595 equipment 835 formulas 268–70 hepatic lipidosis 413 mare’s milk 268 prior to/during transport 235 routes of administration 270–1 sepsis 158 minimal, effects on gut hormones 78 entering-man, breeding shed 1226 enteritis neonatal foal 385–92 management 193–5 older foal 752 ultrasonography 245–6 see also diarrhea Enterobacter identification 1981, 1983, 2677, 2680 neonatal sepsis 253–4, 255 Enterococcus identification 1981, 1984 neonatal sepsis 156, 254, 255 enterocolitis see diarrhea enterocutaneous fistulas 641, 753 surgical repair 642 enteroglucagon 70, 77, 78 entrainment annual reproductive rhythm 1733 circadian rhythms 1670 entropion acquired 543–4 congenital 541–2, 668 critically ill neonates 196 microphthalmic eyes 538 spastic 544
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environmental influences autumnal transition 1734–5 GnRH secretion 1614–15 eosin-methylene blue (EMB) 2676 eosin-nigrosin stain, sperm viability 1494, 1511 eosinophil count 173, 851 eosinophilic enteritis, idiopathic 763 eosinophils endometrial histology 1933 uterine cytology 1924–5 eosin stain, sperm evaluation 1287, 1499 eosin Y, semen evaluation 957 Epac 950 ephedrine, pregnant mare anesthesia 59 EPIC hyperimmunized egg protein 344 epidemic curves 2800 epidemiological studies analytic 2798 descriptive 2798 designs 2797–9 epidemiology 2796–801 defined 2796 disease outbreaks 2799–801 principles 2797 reproductive 2797 veterinary 2796 epidermal growth factor (EGF) extraspecies pregnancies 2311 older mares 2238 placentation 86, 1655, 2236 epidermal nevus linear 565, 566 neonatal 561, 564 older foals/adults 566, 567 epidermolysis bullosa 556, 670 junctional (JEB) 676–7 epididymal ducts (ductus epididymidis) 872, 873, 874 smooth musculature 920, 927, 929 sperm transport 894, 896, 1157 epididymis 1156–60 acquired conditions 1158–9 anatomy 874, 912, 917, 1008, 1156 congenital abnormalities 1111, 1157 contractility 920 cysts 1157, 1158 disease 1157–9 endocrine–paracrine–autocrine regulation 1008–9 examination 1157 functions 874, 894–6, 916–25, 1008, 1156–7 hypoplasia 1157, 1521 lymphatic vessels 1146 microscopic anatomy 1156 neoplasia 1159, 1160, 1169 ultrasonography 1479 physical examination 1459 secreted proteins 1009 secretions in semen 1062 sperm granulomas 1159 sperm harvesting 1160, 1272–3, 2983–5 sperm maturation 894–5, 920–5, 930 sperm reserves, age-related changes 1353–4 sperm storage areas 895–6, 918, 921 sperm transit 896, 918–20, 1047, 1048, 1157 factors regulating 1008 thermal regulation 882 transportation of excised 2985 ultrasonography 1157, 1470, 1472
see also caput epididymis; cauda epididymis; corpus epididymis epididymitis 1158–9 obstructive azoospermia 1520 ultrasonography 1478–9 epidural anesthesia/analgesia cervical surgery 2560 dystocia 59, 2485–6 fetotomy 2498 foal 225–6 laparoscopic ovariectomy 1993 mare caudal reproductive tract surgery 2546 epigenetic modifications, cloned cells 2925 epiglottis entrapment 797 grading system 796 immature horses 794, 795 racing performance and 799 variability in appearance 795 epinephrine (adrenaline) exogenous 842 hyperkalemic periodic paralysis 464, 674 neonatal resuscitation 130, 133 premature or dysmature foals 125 transfusion reactions 241 foal adaptation to parturition 70, 72–3 epiphyseal cartilage 434–5 increased thickness 438–9 epiphyseal cartilage complex, articular 434 epiphyseal plates see physes epiphyses 434, 435 abnormal development 438–9 ossification 434–5, 436 episioplasty (Gadd technique; deep Caslick) 1580, 2541, 2542, 2548, 2549 epispadias 1111 epithelial inclusion cysts, ovarian 2133, 2134 epitheliogenesis imperfecta 556, 670 epoophoron, cystic 2700, 2701, 2702, 2703, 2707 EqcellsireTM semen extender 2964, 2967 R EqStim see Propionibacterium acnes Equex STMTM 2968 Equidone 1699 equids, extinct or endangered 3055 equilenin feto-placental unit 2225, 2226, 2227 testicular biosynthesis 893 equilin pregnancy 1634, 2225, 2226, 2227 testicular biosynthesis 893 R 708 Equilis Strep E R Equimune I.V. 717 equine adenovirus type 1 (EAdV1) 725–6 see also adenovirus infections equine amnionitis and fetal loss (EAFL) 2415, 2461 equine arteritis virus (EAV) carrier stallions 1252, 2393–4, 3021, 3022 detection 2394 GnRH vaccination 1194–5, 1252 management 1252, 2395–6 detection 723, 1253, 2394, 3022 distribution 2391, 3020 immunity 2394 infections see equine viral arteritis transmission 1252, 2392 via AI 3016–17, 3021–2, 3029
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
virus characteristics 1251–2, 2391–2, 3020 equine chorionic gonadotropin (eCG) 1648–60, 2168, 2320, 2465–6 abnormal persistence of secretion 1659–60 α-subunit 1619, 1649 β-subunit 1619, 1649 C-terminal peptide (CTP) 1649, 1650 biological tests 2254–5, 2465 diagnosis of pregnancy 2252–5, 2465–6 Miniature ponies 2841 discovery 84 donkeys 1650, 2836 exogenous 1650–1 donkey-in-horse pregnancies 2315, 2316 superovulation 1836 FSH/LH-like activity 1621, 1650–1 functions in mare 1658–9, 2222 interspecies pregnancies 2306, 2308–9, 2310–11, 2313–14 luteal response 1658–9, 2062 ovarian steroidogenesis 1631 persistent endometrial cups 2669 physicochemical properties 1648–9, 1650 recombinant 1649 secretion 1651, 1654, 1656, 2197, 2235 source in mare 1651–8 structure 1619–20, 1649 zebras 1650, 2857, 2858 equine coital exanthema (ECE) 1254–5, 2376 clinical signs 1254–5 diagnosis 1255 transmission via AI 3023 see also equine herpesvirus 3 equine Cushing disease (ECD) see pituitary pars intermedia dysfunction equine degenerative myeloencephalopathy (EDM) 489–90, 732 R containers 1318, 1332, Equine Express 1333, 1334 equine follicle-stimulating hormone (eFSH) content of pituitary extracts 1837, 1838 endogenous see follicle-stimulating hormone recombinant (reFSH) stallion subfertility 1444 superovulation induction 1841–2 stallion subfertility 1443–3 superovulation in mares 1836, 1837–43 alternative protocols 1839–40 challenges 1842 coasting 1838 injection frequency 1838 oocyte collection 2936 ovulatory agent 1841 preparation purity and 1838 timing of treatment 1838–9 see also pituitary extracts, equine equine gastric ulcer syndrome see gastroduodenal ulceration equine grass sickness 320 equine herpesvirus(es) (EHV) 2376–86 abortion 2377, 2378 clinical features 2379–80 diagnosis 2340, 2341, 2383 epidemiology 2381–2
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Index pathogenesis 2378 prevention and control 2384–6 clinical features 2379–81 diagnosis 724–5, 2383–4 economic significance 2383 epidemiology 2381–3 neonatal disease see equine herpesvirus 1; equine herpesvirus 4 neurological disease see equine herpesvirus myeloencephalopathy pathogenesis 2378–9 prevention and control 2384–6 respiratory disease 724–5, 2377 clinical features 2379 diagnosis 724–5, 2383 pathogenesis 724, 2378 prevention and control 725, 2384–6 treatment 725, 2384 transmission 2377–8, 2381 treatment 2384 types 2376–7 venereal transmission 1254–6 equine herpesvirus 1 (EHV-1) 2376–7 abortion 2377 clinical features 2379–80 diagnosis 2383 pathogenesis 2378 prevention and control 32, 2384–6 carriers 2377–8, 2381–2 diagnosis 8–9, 724–5, 2383–4 economic significance 2383 epidemiology 724, 2381–3 immunity 2382–3 neonatal disease 156, 2377 clinical features 2380 hepatic disease 411, 412 pathogenesis 2378–9 pneumonia 587–8, 602, 603, 607 treatment 2384 neurological disease see equine herpesvirus myeloencephalopathy neuropathogenic strains 2379, 2381 ocular disease 2381 pathogenesis 724, 2378–9 pregnant mare management 32 prevention and control 725, 2384–6 pulmonary vasculotropic infection 2380 respiratory disease see under equine herpesvirus(es) stallion fertility and 2380 stallion semen 1255–6, 2378, 3025 transmission 2377–8, 2381 treatment 725, 2384 vaccination 316–18, 725, 2385–6 broodmare 302, 306, 310, 317 foals and weanlings 304, 307, 308–9, 317–18 stallion 1222 equine herpesvirus 2 (EHV-2) 711, 2376 diagnosis 725 respiratory disease 724, 725 equine herpesvirus 3 (EHV-3) 2376 penile bleeding 1122 transmission via AI 3023, 3029 venereal transmission 1254–6 see also equine coital exanthema equine herpesvirus 4 (EHV-4) 2376–7 abortion 2377 clinical features 2379–80 diagnosis 2383
pathogenesis 2378 prevention and control 2384–6 carriers 2377–8, 2381–2 diagnosis 724–5, 2383–4 economic significance 2383 epidemiology 724, 2381–3 immunity 2382–3 neonatal disease 602, 2377 clinical features 2380 pathogenesis 2378–9 treatment 2384 pathogenesis 724, 2378–9 prevention and control 725, 2384–6 respiratory disease see under equine herpesvirus(es) transmission 2377–8, 2381 treatment 725, 2384 vaccination 316–18, 725, 2385–6 broodmare 302, 306, 310, 317 foals and weanlings 304, 308–9, 317–18 stallion 1222 equine herpesvirus 5 (EHV-5) 2376 diagnosis 725 respiratory disease 724, 725 equine herpesvirus myeloencephalopathy 731, 2377 clinical features 2380 diagnosis 2383–4 epidemiology 2381 pathogenesis 2379 prevention and control 2385 treatment 2384 equine infectious anemia 3024–5 equine influenza/equine influenza virus (EI/EIV) 721–2 diagnosis 722 epidemiology 721–2 neonatal foal 588 pathogenesis 722 treatment and quarantine 722 vaccination 314–16, 722 broodmare 302, 306, 311, 315 foals and weanlings 304, 307, 308–9, 315–16 stallion 1221–2 venereal transmission 3015 equine luteinizing hormone (eLH) content of pituitary extracts 1837, 1838 endogenous see luteinizing hormone recombinant (reLH) 1649 ovulation induction 1865 ovulation synchronization 1874 stallion subfertility 1444 superovulation protocols 1841, 1842 stallion subfertility 1443–3 equine monocytic ehrlichiosis see Potomac horse fever equine motor neuron disease (EMND) 732 equine pituitary extracts see pituitary extracts, equine equine proliferative enteropathy see proliferative enteropathy, equine equine rhinitis A virus (ERAV) 725, 3025 equine rhinitis B virus (ERBV) 725 Equine SNP50 BeadChip 2829 equine viral arteritis (EVA) abortion 9–10, 2392–3, 3021 diagnosis 1253, 2394 pathogenesis 2393 causal agent see equine arteritis virus clinical signs 1252, 2392–3, 3020–1 control 1253–4, 3022
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diagnosis 9–10, 723, 1252–3, 2394, 3022 economic significance 2394 epidemiology 723, 2393–4, 3021–2 neonatal foal 588, 602, 603, 606, 607, 723 pathogenesis 723, 2393 pregnant mare 2391–6, 3021 prevention and control 723, 1253–4, 2395–6 stallion 1251–4, 2393 see also equine arteritis virus (EAV), carrier stallions transmission via AI 3020–2 treatment 723, 2395 vaccination 320–2, 1253–4, 2395, 3022 broodmare 302, 306, 311, 321, 1253 foals and weanlings 295, 309, 321–2 future directions 322 stallion 321, 1222, 1253–4, 1396, 3022 EquiProTM semen extender 1337–8, 1342, 2968 addition of antibiotics 1338, 1339 EquitainerTM cooled embryos 2881, 2882 cooled semen 1264, 1318, 1325, 1331–3, 1334 cooling rate 1331, 1332 EquityTM Oestrus Control 1191 see also gonadotropin-releasing hormone (GnRH) vaccine Equivax S vaccine 708 R EquiZ test 347, 833 Equus 2302 Equus asinus see donkey Equus asinus africanus 2305 Equus burchelli see plains zebra Equus caballus see horse, domestic Equus ferus przewalskii see Asian wild horse Equus grevyi see Gr´evy’s zebra Equus hemionus 2302 Equus hemionus onager 2305 Equus zebra see mountain zebra Equus zebra hartmannae see Hartmann’s zebra Equus zebra zebra see Cape mountain zebra c-erb-2 oncoprotein 2712–13 erection, penile 878, 904 dysfunctional 1419–20 medication-related dysfunction 1422 psychogenic dysfunction 1421–2 spontaneous 1389 see also impotence ergot alkaloids 2418 causing agalactia 2282 domperidone actions 2424–5 immune system effects 2418 interaction with receptors 2418 toxicity 2419–23 uterine blood flow effects 2421 see also fescue toxicosis ergothioneine 1062, 1077 ergovaline 27, 2418 erythrocytes morphological changes 422, 423 reduced lifespan 419–20, 421, 423, 424–5 erythroid hypoplasia, Fell ponies 420, 422 erythromycin 842 Lawsonia intracellularis 739, 762 neonatal foal 254, 260–1 pneumonia in older foal 716 prokinetic use 271, 396 R. equi infections 699, 700, 701, 764
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erythropoiesis, decreased 420, 421 diagnosis 423–4 management 425 Escherichia coli endometritis 1974, 1975–6, 2609, 2611–12 gynecological samples 1973 identification 1969 chromogenic agar 1979, 1981, 1983 in-house 2676, 2677, 2678, 2679 neonatal sepsis 156, 157, 253–4, 255 pneumonia 710 uterine cytological smears 1925, 1926–7 venereal transmission 1254 esophagitis, reflux 684, 685 ESTTM Equine Semen Transporter 1318, 1333 estradiol (E2 ) conceptus secretion 2202 exogenous see estrogen therapy formulations 1825, 1826 mare age effects 2029 autumnal transition 1738, 1746 double ovulations 2022, 2023 estrus 1720 follicular development 1719, 1720 FSH and LH secretion and 1625–7 GnRH secretion and 1613–14 induction of lactation 268 late-term pregnancy 19 multiple ovulations and 2021, 2022 parturition 2268 periovulatory period 2024, 2025 pregnancy 1634, 2225, 2227 seasonal anestrus 1784–5 spring transition 2027 zebras 2857, 2858 stallion 997–8 biosynthesis 893 infertility/subfertility 1439 puberty 904, 1006, 1007 seasonal variations 1003–4 testicular levels 1440 estriol, testicular biosynthesis 893 estrogen(s) assay methods 2463 conceptus secretion 1633, 2178–9, 2202, 2225 exogenous see estrogen therapy mare 1631–5 assay diagnostics 1635 autumnal transition 1738 biosynthesis 1631, 1632 estrous cycle 1631–2 FSH and LH secretion and 1625–7, 1639 intrauterine sperm transport and 942 lactation 2277 parturition 1634, 2268–9 placental production 89, 1633–4 placentitis 2362–3 pregnancy see under pregnancy puberty 1690–2 uterine contractility and 2598 uterine defenses and 2589 uterine involution 2292 vernal transition 1707–9 maternal recognition of pregnancy 1633, 2201–2 ring B 2225, 2227 stallion 997–8 cryptorchidism 1104, 1533 epididymal function 920, 1008–9
infertility/subfertility 1436 paracrine–autocrine actions 1001–2 puberty 904, 1006, 1007 seasonal changes 1346 testicular levels 1440 testicular production 893, 900, 1001 estrogen receptors (ER) cervical 2728 early conceptus 2202 epididymis 1008–9 estrous cycle 1631–2 pregnant endometrium 1633 testicular 1001 estrogen response element (ERE) 1626 estrogen (estradiol) therapy 1825–8 agents used 1825, 1826 anestrus 1697 behavioral estrus induction 1763, 1825 cervical dilation 2729, 2811 contraception in mares 1889 controlled release formulations 1881–2 diestrus 1730–1 foal rejection 120 follicular wave synchronization 1826 high-risk pregnancy 19, 1826–7 induction of abortion 2321 induction of lactation 268, 1828, 2283, 2284 maintenance of pregnancy 2472 oocyte and embryo recipients 1827, 2942 postpartum 1827, 2298–9 stallions 1443, 1828 teaser mares 1825 transitional mares 1710–12, 1779–80, 1791, 1826 uterine infections 1827–8 see also progesterone/estradiol (P&E) treatment estrone pregnancy 2225, 2226, 2227 testicular biosynthesis 893 estrone sulfate (ES) (estrone conjugates) autumnal transition 1738 cryptorchidism 1104, 1533 diagnosis of pregnancy 2254, 2463–4 Miniature ponies 2841 late-term pregnancy 19 production in pregnancy 1632, 1634, 2225 estrous behavior 1717, 1845 abnormalities 1761–5 asynchronous 2669–70 autumnal transition 1748 early pregnancy 2245 estrogen therapy to induce 1763, 1825 failure to exhibit 1697, 1761–2 individual variations 1718–19 jenny donkey 2836 observation methods 1718 problematic 1762–3, 1845–6 role of progesterone 1638 suppression see estrus, suppression vernal transition 1708 zebras 2855 estrous cycle(s) abnormalities 1754–65, 2635–8 Asian wild horse 2848 asynchrony with endometrial histology 2669–70 autumnal transition 1732 double rate 1431 duration 2012 estrogen profiles 1631–2
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
folliculogenesis 2012–17 FSH and LH profiles 1621–3 induction embryo transfer 2871–2 exotic equids 2311 light treatment see lighting programs, artificial oocyte transfer 2942 seasonal anestrus 1698–9, 1785–6 jenny donkey 2836 Miniature mares 2839 per mare 1431 per pregnancy 1431, 2780 pregnancy rate per see pregnancy rates, per cycle previous history 1898 progesterone profiles 1638 services per 1431, 2780, 2782–3 staging, endometrial biopsy 1931 synchronization 1870–6 donkey 2836 embryo transfer 1870, 1876 estrogen therapy 1826 follicular ablation 1875–6 hCG 1807, 1873, 1874 indications 1870–1 intravaginal devices 1826, 1874–5 methods 1871–6 oocyte transfer 2941–2 ovulatory agents 1873–4 progestagens 1815–16, 1872–3 progesterone/estradiol 1816, 1826, 1873 prostaglandins 1731, 1799, 1871–2 uterine contractility 2598 uterine histotrophe 1639–40 zebras 2855–6 see also diestrus; estrus; luteal phase estrus 1716–26 behavior see estrous behavior cervical examination 1725, 1726 definition 1716 detection 1716–17 documentation 1717 observation in pasture 1718 teasing mares 1716–17, 1718 Thoroughbred breeding farm 1718–20 endometrial histology 1931 estrogen profile 1631 first postpartum see foal heat follicular development 1717–18, 1719–20 FSH and LH profiles 1621, 1622–3 hormone profiles 1720 hysteroscopic appearance 1942 inappropriate or paradoxical 1748 induction see estrous cycle(s), induction ovulation prediction 1720–6 pubertal (first) 1692–3 age at 1690 birth date and 1693 bodyweight and 1693 criteria for identifying 1690 daylength and 1693 hormonal stimulus 1690–2 silent 1716, 1748, 1762 stallion behavioral responses 1388 stallion-like behavior during 1763 suppression 1845–51 GnRH analogs 1850 hCG therapy 1849–50 immunological methods 1850–1
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Index indications 1845–6 intrauterine devices 1760, 1847–9 methods 1846–51 ovariectomy 1851, 2564 oxytocin therapy 1834, 1849 pregnancy 1850 progestagen therapy 1762, 1815, 1846–7 transitional mares 1778–81, 1815 synchronization see estrous cycle(s), synchronization uterine anatomy 1590, 1591 uterine fluid accumulation 1723, 1725, 2657 uterine ultrasonography 1723, 1724, 2004, 2005 vaginal changes 1591, 1901–2 zebras 2855 ethane dimethanesulfonate 1178 ethical issues, reproductive technologies 3052 ethidium 1287, 1495, 1500 ethinylestradiol (EE2), contraception 1889 ethylene diamine dihydroiodide (EDDI) 842 ethylene diamine tetra-acetic acid (EDTA) intrauterine infusions 2614, 2634–5 protection of sperm from cold shock 1074 semen cryopreservation 2970 topical, corneal ulcers 546 ethylene glycol (EG) embryo freezing 2891–5, 2901 , 2903, 2906–9 embryo vitrification 2910–15 oocyte cryopreservation 2954–5 semen cryopreservation 2969 toxicity to embryos 2917 ethylene oxide (EtOH), sterilization of endoscopes 1450 etodolac 842 Europe, semen extenders for cooled semen 1336–9 European Union frozen semen quality standards 3012–13 import or export of semen 2972 euthyroid sick syndrome 528 evidence-based medicine (EBM) 2796 evisceration, complicating castration 1561 exercise breeding stallions 1211–21, 1228–9 broodmares 2748 effects on growth 285 energy needs of growing horses and 784 newborn foal 113 osteochondrosis and 779 periparturient mare 112, 113 premature or dysmature foals 126 problem-breeding mares 2630 uterine clearance 2811 uterine involution and 2295 exocrine action 1601 exophthalmos 540 experimental studies 2798, 2799 export embryos 2921–3 frozen semen 2972, 3013, 3029–35 stallions 1186–7 exposure 2796
extensor thrust reflex 479 external anal sphincter, mare 1577 external carotid artery, fetal 46 external cremaster muscle see cremaster muscle, external external genitalia ambiguous 1961 female see female external genitalia fetal sex determination 2085–7, 2095–8 male see male external genitalia external iliac artery, rupture 2449 see also periparturient hemorrhage external pudendal artery, stallion 877 extraspecies pregnancies 2305–17, 3054 embryo recovery and transfer 2306 horse-in-donkey 2306–7 ex utero intrapartum treatment (EXIT) 132 eye care, critically ill neonates 196 disorders see ophthalmic disorders examination in foals 535–7 critical care 172 neurological disorders 478, 480 fetal see fetal eye posterior segment diseases 550–1 traumatic injuries 540, 543 eye hooks, obstetric 2490 eyelid acquired abnormalities 543–4 coloboma (agenesis) 541 congenital abnormalities 541–3 trauma 196, 543 E-Z FreezinTM semen extenders 2967 E-Z MixinTM semen extender 2967 face examination, neonate 478 FACS (fluorescent-activated cell separation), sex-sorting of sperm 3042–4 failure of passive transfer (FPT) 344, 346–51, 362 definition 175, 344, 346 fescue toxicosis 2422 incidence 347 neonatal infections 154, 198, 199, 347 partial see partial failure of passive transfer perinatal asphyxia syndrome 151 premature lactation 2284–5 premature or dysmature foal 126 prevention 350–1 risk factors 198, 344, 346–7, 351 supplies to diagnose/treat 836 testing for 113, 175, 346, 347–8 treatment 348–50 colostrum supplements 344, 348–9 oral colostrum substitutes 344, 348, 349 oral immunoglobulins 344, 349 plasma/plasma product transfusion 209, 240, 349–50 prior to transport 235–6 Falabella horse 1377 Fallot’s pentalogy 670 Fallot’s tetralogy 509, 670 fall transition see autumnal transition famotidine 842 Fanconi syndrome 651 fasting, pregnant mares 28–9, 53 fat, dietary intake, breeding stallion 1237–8 fatty acids essential 1603
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membrane, effects of cooling 1309–10 sperm content 1062 supplements, breeding stallion 1237–8, 1248 fear, signs of 1407–8 fecal impaction, after dystocia 2494 fecaliths 755 feeding breeding stallions 1221, 1238–9 growing horses 790–1 in natural conditions 1387 neonatal foals 63 critical care 192–3 endocrine responses 76–7, 78 gastrointestinal tract changes 78 orphans 270, 273, 2280–1 prematurity or dysmaturity 125 sepsis 158 see also suckling/nursing periparturient mare 111–12 pregnant mares 2764–5 rejected foals 119 shuttle stallions 1188 trophic, or minimal enteral 78 see also diet; nutrition feet care broodmares 2748 see also hoof care Fell Pony syndrome 362, 364–5 bone marrow changes 420, 422, 424, 561 cutaneous features 558–9, 560–1 female external genitalia anatomy 1577–81 cleaning/preparation AI 1203–4, 1274, 1275 microbiological sampling 1965, 2674–5 natural service 1224 vaginal examination 1902 hemorrhage at, during pregnancy 2451–2 see also perineum, female; vulva female pseudohermaphrodite 1111, 1136, 1961, 2218–19 female reproductive function in natural conditions 1388 neuroendocrine contributors 1582–6 relevant anatomy 1577–81, 1582–96 female reproductive tract anatomy 1586–95 general relationships 1586, 1587 arterial supply and venous drainage 1593–4 congenital abnormalities, hysteroscopy 1943–4 cultures 1963–77 effects of semen 1094–7 endoscopy 1940–9 congenital abnormalities 1943–4 indications 1940 pathological conditions 1944–9 routine anatomic features 1940–3 see also hysteroscopy; vaginoscopy examination, geriatric mare 2804–5 external see female external genitalia innervation 1594–5 lymphatic drainage 1594 rectal palpation 1904–13 sperm transport 941–4, 3036–7 surgery of caudal 2545–57 ultrasonography see under ultrasonography zebras 2855
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femoral fractures 451 fenbendazole ascarids 295, 711–12, 756, 759 Dictyocaulus arnfieldi 711–12 parasite control programs 113, 300 strongyles 298, 300, 742, 759–60 fenprostalene 1799 delayed uterine clearance 2602 induction of abortion 2320 fentanyl foal 226, 234 patches 226, 842 feral and semi-feral horses annual reproductive patterns 1387–8 contraception 1888–9, 1890–2 foraging and feeding 1387 normal sexual behavior 1388–9 onset of puberty in fillies 1692, 1694 social organization 1385–7 Ferguson reflex 1680, 2269 ferritin, serum 853 neonatal foal 419, 420 fertility genetic influences 2828–9 interspecies hybrids 2302, 2303, 2304–5 mare see mare fertility stallion see stallion fertility stallion sperm aged stored sperm 1076–7 cooled semen 1321, 2991–2 ejaculation frequency and 896 frozen semen 2988–92, 2994–5 predicting 2995 vs. cooled semen 2991–2 sex-sorted sperm 3045–6 sperm attributes required 1053–4 sperm morphological abnormalities and 960 sperm–oviduct interaction and 1089 see also infertility; reproductive efficiency fertilization 1492, 1493 rate 2170 defined 2780 interspecies matings 2303 in vitro 2945 in vivo 2945 xenogenous 2945 fertilization-promoting peptide 946–7 fescue toxicosis 2418–25 growing horses 2422–3 management and treatment 2283, 2423–5 mare infertility 2422 performance horses 2423 pregnant mares 6, 11–12, 2419–22, 2765 abortion/stillbirths 2422 agalactia 2282, 2283, 2420 assessing readiness for birth 2736 dystocia 33, 2419–20, 2482–3 effects on foal 72, 530, 2421–2 management 27–8, 33 placental effects 91, 2421 prolonged gestation 11, 122, 2154, 2419–20, 2449 retained fetal membranes 2421, 2523 ultrasonography 2361 prevention 351, 2283, 2749 fetal abnormalities see fetal malformations fetal activity (in-place movements) 2075, 2076 fetal heart rate 48, 49
ultrasonographic assessment 22, 49 see also fetal mobility fetal autonomy concept 2167–9 fetal breathing 22, 49 fetal calf serum (FCS) 2881 fetal death caudal presentation 2491, 2492 delivery in late-term 25, 2485, 2486 donkey-in-horse pregnancies 2314–17 fetotomy 2485, 2497–503 mare reproductive loss syndrome 2410–11 necropsy 2340–1 one twin 2115–16 uterine torsion 2439, 2442 see also abortion; fetal maceration; fetal mummification; stillbirths fetal distress ECG monitoring 21 transient, during ultrasonography 40 ultrasonographic features 20, 22 fetal eye estimating day of pregnancy 2076–9 Miniature ponies 2842, 2843 growth monitoring 41–6 measurement 46, 2077 reference values 46, 2077, 2078 ultrasonography 41–6 fetal fluids 87, 2368–9 abnormalities 93 clarity 20, 22, 51 composition in hydrops 2369–70, 2444 discoloration 93 drainage, hydrops 34 excess see hydrops volumes 20, 22, 51, 87, 2368 see also allantoic fluid; amniotic fluid fetal foot placental area of degeneration (PAD) 103–4 fetal growth 281 curves 286, 287 factors affecting 281–3, 2237–8 fueling 286–7 hormones involved 2229–30 ultrasonographic assessment 41–8 fetal heart rate (FHR) during anesthesia 56 ECG monitoring 21 normal values 48 ultrasonography 20, 22, 48–9, 2361–2 fetal inflammatory response syndrome (FIRS) 646 fetal loss rate 2780 fetal maceration 2373–5, 2445–7 causes 2373 definition 2373 diagnosis 2373 hysteroscopy 1948 treatment 2374–5, 2447 ultrasonography 2150, 2152 fetal maldisposition 2479 fetal malformations 7, 13 due to malpresentation 2441 dystocia 2493–4 hydrops 2371 induction of abortion 2322 management 35 Miniature ponies 2841, 2842–3 see also congenital abnormalities fetal malpositions 2486–7 fetal malpostures carpal flexion 2489 caudal presentation 2492–3 cranial presentation 2487–91
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fetal manipulation 2486–91 fetotomy 2499–503 foot-nape posture 2488–9 head and neck 2489–91 hospital approach 2514 incomplete forelimb extension 2487 ventro-vertical presentation 2487–8 fetal malpresentations 7, 13, 2491–3 fetotomy 2502–3 hospital approach 2514 malformations due to 2441 management 35, 2491–3 nomenclature 2479 fetal manipulation anesthesia for 59, 2485–6 dystocia 2486–93 referral dystocias 2514 fetal membranes 99 delivery 2271–2 differentiation 2187–98 20–30 days 2188–90 30–40 days 2191–4 41–60 days 2194–7 60–80 days 2197–8 examination 102, 2272–5 iatrogenic disruption, to induce abortion 2321 hydrops 2371, 2445 normal appearances 100, 101 retained see retained fetal membranes rupture at parturition 2271 see also allantochorion fetal mobility (whole-body) 90, 2075, 2076, 2479–80 see also conceptus, intrauterine mobility fetal monitoring see feto-placental monitoring fetal movements see fetal activity; fetal mobility fetal mummification 2373–5, 2445–7, 2670–1 causes 2373–4 definition 2373 hysteroscopy 1948 treatment 2374–5, 2447, 2670–1 twin gestation 2115, 2150, 2373–4, 2446 ultrasonography 2148, 2149, 2150, 2152 fetal oversize see feto-pelvic disproportion fetal position abnormal 2486–7 defined 2479 dorsoiliac 2481 dorsopubic 2480, 2481 peripartum changes 2270–1, 2481–2 ultrasonographic assessment 41, 49 fetal posture defined 2479 manipulation 2486 see also fetal malpostures fetal presentation defined 2479 nomenclature 2479 prepartum changes 2075–6, 2479–80 ultrasonographic assessment 20, 22, 41 fetal sex determination 55 to 150 days 2080–6 55 to 90 days 2083–5, 2086, 2087, 2250 90 to 150 days 2080, 2085–7, 2088–92 applications 2081 difficulty at 80 to 90 days 2082, 2083 equipment and materials 2081–2
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Index fetal development and 2082 precautions 2082–3 success rates 2082 ultrasonic planes 2083, 2084 mid to advanced gestation 2094–8 fetal size, estimation 21, 41–6 fetal ultrasonography 39–53, 2074–9 activity and mobility 2075 applications 52–3 assessment chart/form 41, 42–5 biophysical indices 20–2, 51, 52 environment 49–51 estimating day of pregnancy 2076–9 examination technique 40 growth and development 41–8 hind limbs encased in cord horn 2076 placentitis 2361–2 scanning technique 40–1 sex determination 2080–6 uterine constrictions and allantoic fluid shifts 2075 uterine horn closures and presentation 2075–6 viability 48–9 see also ultrasonography (US), pregnant mare fetal viability assessment in dystocia 2484, 2514 ultrasonographic assessment 48–9 fetatome 2498 fetlock joint arthrodesis flexor tendon laxity 470 flexural deformity 443 presale radiographs 802–4, 805 interpretation 810–14 normal variations 810 proximal sesamoid fractures 448, 811 see also metacarpophalangeal joint feto-pelvic disproportion 2484, 2487 fetotomy 2501 feto-placental monitoring 16–22, 39–40 biochemical indices 16–20 biophysical indices 20–2, 51, 52 placental dysfunction 93 ultrasonographic see under fetal ultrasonography viability assessment 48–9 feto-placental unit estrogen biosynthesis 2225–7 progestagen metabolism 2222–4 fetotomy 2485, 2497–503, 2511 anterior presentation 2499–502 equipment 2498–9 fertility after 2498, 2515 general procedures 2498–9 partial 2497 posterior presentation 2502–3 referral dystocias 2513, 2515 transverse presentation 2503 fetotomy knife 2499 fetus adrenocortical activity see adrenal cortical hormones, fetal assessing readiness for birth 2262–5 delivery at parturition 2271 early stage 2192, 2193 effects of anesthesia 56 endocrinological adaptation to parturition 69–80 estrogen production 1633–4 folliculogenesis 2011 hormones 2230 immune system 334–5
in utero kinetics 2479–80 ossification and joint development 433–9 Fgf9 gene 2214 fibrin degradation products (FDP) 428, 853 fibrinogen neonatal foal 174, 199, 428 pneumonia in older foals 713 R. equi pneumonia 692, 697 reference values 851, 853 fibroblasts cytogenetic analysis 1953 donor, for nuclear transfer 2924–5 fibroleiomyoma, uterus 1947–8 fibrosarcomas, penis and prepuce 1169 fibrous sheath, sperm middle piece 931, 932, 934, 935, 1061 production 1036 role in sperm motility 1293 filigrastim 842 filipin 1502 fimbrial cysts, paramesonephric 2700, 2701, 2702, 2703 ultrasonography 2133, 2134 firocoxib foals 227, 744 high-risk pregnancy 26 poor libido in stallions 1415–16 flagellar canal 918, 935, 1028, 1035 flagellum, sperm see spermatozoal tail flank laparotomy oocyte transfer 2941, 2942 ovariectomy 2566–7 uterine torsion 2437–8, 2439 flank puncture, mature oocyte collection 2936–7 flatbones, ossification 436 Flexbumin 208 flexor carpi ulnaris tenotomy, carpal flexural deformity 443 flexor reflex 479, 480 flexor tendon laxity (digital hyperextension) 469–71 pathogenesis 439, 469 prognosis 470–1 treatment 469–70 flexural limb deformities 441–5, 666, 774–5 acquired 441–2, 443–5, 775 common digital extensor tendon rupture with 471 congenital 441, 442–3, 445, 774–5 diagnosis and evaluation 441–2 dietary factors 785 dystocia 2489 fetotomy 2499 Miniature ponies 2842–3 pathogenesis 433–4, 439, 441 prognosis 445 treatment 442–5 flick reflexes 478, 480 flow cytometry acrosome evaluation 1503, 1504 frozen-thawed sperm evaluation 2999–3002 sex-sorting of sperm 3042–4 sperm viability testing 1499, 1512 Flt (VEGF receptor) 2240 fluconazole 842 fungal endometritis 2636, 2647–8 neonatal infections 157 ophthalmic disorders 541 fludrocortisone acetate 842
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fluid accumulation, uterine see intrauterine fluid accumulation fluid harnesses 824, 825 fluid losses, neonatal foal 205 fluid overload neonatal renal dysfunction 656 neonatal shock 141 perinatal asphyxia syndrome 151 fluid pumps 824 fluid requirements, neonatal foal 205, 206 fluid therapy, intravenous foal 205–14, 842 assessing requirements 205, 206, 213–14, 230, 231 birth resuscitation 134 critical care 190–2 diarrhea 386, 736 electrolyte abnormalities 209–12 equipment 824, 835 gastroduodenal ulceration 686 glucose abnormalities 212–13 guide to fluid choice 213 initial 190–2 maintenance 192, 206, 231 meconium impaction 382, 383 perinatal asphyxia syndrome 151 prior to/during transport 230–2 renal problems 656–7 replacement 190–1, 206, 230–1 sepsis 157, 202–3 shock 140–2 types of fluids 140–2, 190, 206–9 mare anesthesia in pregnancy 58–9 hydrops 34, 2371 periparturient hemorrhage 2518 flunixin meglamine 842 mare colic in pregnancy 30 embryo recipients 2878 embryo reduction in twin pregnancy 2101 high-risk pregnancy 26 induction of abortion and 2321 oocyte collection 2937 prevention of embryonic loss 2335 rectal tears 2579 retained placenta 2525 neonatal foal analgesia 226–7, 234 nephrotoxicity 652 neuroprotection 232, 486 ophthalmic disorders 541, 546, 551 peripheral nerve injuries 489 sepsis/endotoxemia 157, 235 septic arthritis and osteomyelitis 460 older foal/weanling/yearling colic 750 diarrhea 736 gastrointestinal toxicity 744 pneumonia 717 strangles 707 postpartum metritis 2532 stallion, testicular trauma 1161 transfusion reactions 241 fluorescein diacetate (FD) 1513 fluorescein isothiocyanate (FITC), acrosome evaluation 1288, 1494–5 fluorescence-tagged lectins, acrosome evaluation 956, 1288, 1491, 1495, 1503, 3000 fluorescent-activated cell separation (FACS), sex-sorting of sperm 3042–4
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fluorescent in situ hybridization (FISH) 1464, 1466–7, 1954, 1955 fluorescent stains acrosome evaluation 956, 1288–9, 1494–6, 1503, 3000 capacitation assays 1502 sperm membrane (viability) tests 957–8, 1287, 1499–500, 1511–13, 2999–3000 sperm mitochondria-specific 957, 1289, 3000–1 5-fluorouracil, penile and preputial carcinoma 1138 fluphenazine, transitional mares 1789, 1790 fluprostenol induction of abortion 1800 postpartum mares 2292 flurbiprofen 842 fluticasone 843 foal abdominal ultrasonography 243–51 cloned 2925 cytogenetics 1951–61 dysmature see dysmature foals equine viral arteritis 3020 growth 287–8 factors affecting 282–5 fueling 287–8 optimal rates 285 immune system 331–9 immunodeficiencies 361–6, 369–71, 373–5 mortality, Asian wild horse 2849–50 parasite control 113, 295, 300 post-mature see post-mature foals post-term see post-term foals premature see premature foals Taylorella equigenitalis infection 3018 vaccination 303–5, 307–9, 338–9 weaning 113, 2282 zebra 2861–2 see also neonatal foal; older foal Foalchek test 348 foal heat (first postpartum estrus) 2296–7 anestrus after 1699 breeding mares on 2294–300, 2468–9 early embryonic loss 2331 fertility 2142, 2293, 2297 insemination with frozen semen 2990 periparturient problems and 2296 reproductive efficiency 2781 strategies 2142, 2299–300, 2469 time of first ovulation and 2297–9 uterine fluid accumulation and 2295, 2468–9 treatments to enhance fertility 1816, 2292–3, 2295–6, 2298–9 zebras 2858–9 foal heat diarrhea 391 foaling dates, optimal 1782 first, defining puberty 1692 in natural conditions 1387, 1388–9 previous history 1898 see also parturition foaling mares definitions 1429–30, 1897, 2780 reproductive efficiency 2781 stallion fertility evaluation 1430, 1431 see also lactating (wet) mares; postpartum mares; pregnancy foaling rates
definition 2780 endometrial biopsy and 1933–4 stallion 1184, 1185, 1362 yearling fillies 1692 Foal-Lac milk replacer 269 foal rejection 117, 118–20 management 118–20 prevention 120 Foal’s FirstTM milk replacer 845 FoalWatchTM test 20, 2270, 2734, 2735 focusing, ultrasound beam 1917 follicles, ovarian see ovarian follicles follicle-stimulating hormone (FSH) 1619–27 β-subunit 1619 exogenous see equine follicle-stimulating hormone mare age effects 2028–9 autumnal transition 1744–5 body condition and 2028 control of secretion 1623–4, 1783–4 cyclic patterns 1621–3 deficiency 1700 double ovulations and 2022, 2023 feedback control 1624–7, 1639 follicular development 1717–18, 1719, 1720 GnRH secretion patterns and 1611–12 inhibin/activin actions 1625, 1681, 1682 melatonin and 1676–7 multiple ovulations and 2021, 2022 ovarian steroidogenesis 1631, 1632 periovulatory period 2023–4, 2025 pubertal changes 1690–2 seasonal anestrus 1698, 1706, 1784 vernal transition 1706–7 PGF2α -induced secretion 1799 polymorphism and biological activity 1620 porcine, superovulation induction 1836–7 stallion 996–7 control of secretion 899–900, 996–8 control of spermatogenesis 900, 1047–8 infertility/subfertility 1436, 1439, 1523–4 pubertal changes 902, 903, 904, 1005–7, 1016 seasonal variations 873, 1003–4 secretion 897–8 structure 997, 1619–20 follicle-stimulating hormone (FSH) receptors 1620–1 gene polymorphisms 911, 1621 follicle-stimulating hormone-releasing hormone (FSH-RH), specific 1608, 1609, 1624 follicular dysplasia, cutaneous 567–8 follicular fluid entry into abdomen 2046–7 release at ovulation 2043–4 follicular waves 2011, 2012–13, 2020–1 autumnal transition 1739–41 emergence 2011 estrus and 1717–18 major 2011, 2012, 2013–14 minor 2011, 2012, 2013 prepubertal mares 2012 primary 2011, 2012, 2013 secondary 2011, 2012–13
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
shortening diestrus and 1729–30 superovulation protocols 1838–9 synchronization 1839 vernal transition 1706 folliculogenesis 1719–20, 2009–18 abnormal, causes 1754–7, 1762, 2123–6 adult period 2012–18 asynchrony with endometrial histology 2669–70 autumnal transition 1737–9 common growth phase 2011, 2014–15 definitions 2010, 2011 dominance phase 2011, 2014, 2015 early pregnancy 2017 estrous cycle 2012–17 estrus 1717–18 fetal period 2011, 2699 FSH profile 1622 idiopathic failure 2126, 2127 non-breeding season 1698 periovulatory period 2020–9 endocrinology 2021–5 factors affecting 2025–9 pharmacological control during diestrus 1730–1 pre-ovulatory phase 2011, 2014, 2015–17 prepubertal period 2011–12 spring transition 2017, 2018, 2027–8 ultrasonography 2009–18 zebras 2855–6 follistatin 1625 follistatin (Fst) gene 2214 footcandle 1774 foot-nape posture, fetus 2488–9 forage breeding stallions 1231–2, 1234, 1238 growing horses 790 non-pregnant mares 2762 pregnant mares 2764–5 forage:concentrate ratios, growing horses 791 foraging, in natural conditions 1387 foramen ovale, persistent 670 Ford interlocking pattern, hysterotomy closure 2506, 2507 fore feet, presale radiographs 816 foreign bodies, uterus 1948, 2148–9, 2670–1 forelimbs, fetal amputation of extended 2500–1 amputation of flexed 2501 carpal flexion 2489 extension during parturition 2481–2 incomplete extension 2487 normal delivery 2271, 2482 retained see shoulder flexion posture, fetus shoulder flexion 2491 formulary 838–49 fourth branchial arch defects 613–14, 617, 664–5 fractional excretion indices 653–5 fractures in foals 446–55, 768 frame rate, ultrasound image 1918 freemartin syndrome 2217, 2357 freestyle swimming posture, fetus 2491 freezing see cryopreservation freight charges, cooled semen 1324 French National Stud, sperm freezing protocol 2974 frenulum, cervical 1590 frenzied/frenetic behavior, stallions 1411 frequency, ultrasound 1916–17
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Index Friedman test 2254 Friesians heritability of semen quality parameters 1366 morphological sperm defects 1365 retained fetal membranes 2523 stallion reproductive parameters 1362–6 frozen semen 2994–3003, 3005–14 antimicrobial decontamination 3030–1 breeding with 2987–92 estrus synchronization 1871, 1874 low dose insemination 3037, 3038, 3039 ovulation induction 1860, 1862, 1864 reproductive efficiency 2782 sperm dose see insemination dose timing and frequency of insemination 1205, 2988–90 uterine inflammatory response 1094–6 commodity status 3013–14 computer-assisted sperm analysis 2979–80, 2995, 2996–9 contract considerations 3014 disadvantages 2994–5 documentation 3011–12 evaluation 2979–80, 2995–3003 extenders see under semen extenders fertility 2988–92, 2994–5 fertility prediction 2995 flow cytometry 2999–3002 GIFT 2946 ICSI 2948 import and export 2972, 3013, 3029–35 importation requirements 3031–4 infectious agent testing 3002 infectious disease control 3030–1 infectious disease transmission 3015–26, 3029–30 inventory management systems 3009 packaging 2976, 2977, 3006, 3007 quality standards 3012–13 sex-sorted sperm 3045–6 sperm concentration/numbers 2976, 2995 sperm–oviductal interaction 1089 sperm quality 2996–9 storage 3005–9, 3031 straws 2976, 2988 historical perspective 1265 inventory management systems 3009–10 labeling 3009 loading shipping containers 3011 storage 3006, 3007 transfer from shipper to storage 3011, 3012 thawing 1265, 2963, 2975, 2979, 2987–8 transfer from shipper to storage 3011, 3012 transportation 3010–12, 3031–5 see also artificial insemination; semen cryopreservation fructose semen 1062 utilization by sperm 938–9, 1063 yolk sac fluid 2176, 2179 FSH see follicle-stimulating hormone fundus examination, foal 537 fungal infections endometritis 2635, 2643–50 foal pneumonia 588, 711 mastitis 2738, 2740
neonatal foal 156 uterine cytological smears 1925, 1926 venereal transmission 1256, 3025, 3030 fungemia 154 fungi cultures 1968 gynecological samples 1973 identification 1970, 2677, 2680–1 funiculitis, septic 1153–4 funisitis 90 mare reproductive loss syndrome 2412, 2413 furacin 272 furosemide 843 hypoxic-ischemic encephalopathy 151, 487 neonatal foal 195, 210, 211 neonatal vasogenic nephropathy 657 future reproductive technology 3051–5 constraints 3051–2 non-transgenic methods 3053–5 predicting phenotypic outcomes 3052–3 transgenic methods 3053 gabapentin 843 foal 226, 227 perinatal asphyxia syndrome 149, 150 poor libido in stallions 1415 Gadd procedure see episioplasty gain, ultrasound 1916 gait disorders, neuroanatomy 480 neonatal foal 171, 477, 479 galactorrhea 2283–5 galactose-binding proteins, sperm–oviductal adhesion 1088 α-galactosidases 264 β1,4-galactosyltransferase I (GalTI) 948, 953 gallium maleate, R. equi prophylaxis 691–2 gamete intrafallopian transfer (GIFT) 2945–6 future 2946, 3052 handling of gametes 2945–6 laparoscopic oviductal ligation 1999 uterine tube pathology 1988 gametic imprinting 3055 R test 343, 832 Gamma-Check C R Gamma-Check E test 343, 347, 348, 834 gamma-glutamyltransferase (GGT) serum 409, 410, 852, 855 urine 855 gamma (γ ) radiation, effects on spermatogenesis 1177 gap junctions, myometrial 2597 gas exchange, placental 88 Gasterophilus spp. (bots) 292–4, 296 gastric acid 394 gastric distention gastroduodenal ulceration 401, 684 ultrasonography 246, 247 gastric emptying, delayed neonatal foal 170–1 older foal/weanling/yearling 683–4, 752 treatment 686 gastric ulceration see gastroduodenal ulceration gastrin 70, 77, 78 gastrocnemius muscle rupture 467, 473 tendon rupture 473
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gastrocnemius reflex 479 gastroduodenal ulceration cribbing and 1424, 2772 neonatal foal 394–7 clinical syndrome 395, 401 liver disease 413 medical treatment 395–6 pathophysiology 394–5 prevention 195, 227, 396–7 surgical treatment 401 older foal/weanling/yearling 683–7 clinical syndromes 683–4, 752–3 diagnostics 684–5, 743 diarrhea 742–3 pathophysiology 683 prevention 686–7, 737 treatment 685–6, 743 weight loss 760 gastrointestinal disorders congenital 668 causing colic 753 fetal ultrasonography 47 meconium impaction 381 surgical management 404 neonatal foal 267, 405–8 enteral feeding-related 271–2 history and examination 399–400 perinatal asphyxia syndrome 151 prematurity or dysmaturity 125 surgical 399–404 older foal, weight loss 758–63 pregnant mare, management 30 gastrointestinal dysmotility, enteral feeding 271 gastrointestinal hormones 70, 77, 78 gastrointestinal parasites 292–300 gastrointestinal support, critically ill neonate 193–5 gastrointestinal surgery neonatal foal 399–404 diagnostic evaluation 399–400 techniques 400–1 older foal/weanling/yearling 757 gastrointestinal (GI) tract fetal foal 77–8 neonatal foal 66–7, 170–1 adaptation to parturition 77–8 drug absorption 253 entry of pathogens 199 ultrasonographic evaluation 243–6 gastroscopy 684 G-banding, chromosomes 1463, 1464, 1952, 1953, 1955 GBE1 gene 677 gelding see castration geldings basal testosterone levels 1103 estrone sulfate levels 1104 hCG stimulation test 1104 inguinal herniation 1147, 1540 penile and preputial neoplasia 1137, 1169 priapism 1134, 1135 self-mutilation 1424 unwanted sexual behavior 1192 urethral rents causing hematuria 1140–1, 1142 R , neonatal foals 141 Gelofusine gender determination, fetal see fetal sex determination gender differences causes of colic 749 growth rates 280–1 gene expression studies 2829–30
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gene machine, theoretical 3053 gene microarrays 2830 genetic disorders 673–9, 2828–9 effects of AI 1198 molecular tests 2824, 2825 genetics developmental orthopedic disease 777 early embryonic loss 2329–30 future progress in horse 620, 962 infertility 2828–9 preservation technologies 3052 respiratory diseases 615–20 genetic tests health and color traits 2824, 2825 parentage testing 2820–5 genetic variants 2828–9 genital tract embryogenesis 2211–12, 2213 fetal, ultrasonography 48 mare see female reproductive tract stallion see male genital tract see also urogenital system genital tubercle, fetal sex determination 2080, 2084–5, 2086, 2087, 2089 genitofemoral nerves 1596 genome sequence, horse 620, 673, 962, 2827–8 genome-wide studies, recurrent airway obstruction 619 genomic imprinting, placental development 2310, 2311 genomics 962, 2827–30 gentamicin 843 chronic endometritis 2632, 2633 contagious equine metritis 2405 foal diarrhea 736 infectious hepatitis 412 intrauterine infusion 2632, 2633 neonatal foal 194, 254, 255, 256 placentitis 2363, 2364 pneumonia in older foal 716–17 postpartum metritis 2532 R. equi pneumonia 698–9 semen extenders 1201, 1338 geographic differences, growth rates 283–4 geriatric horses dietary support 2755–9 pituitary pars intermedia dysfunction 2756, 2757, 2758, 2791, 2805–6 geriatric mares 2790, 2803–15 anestrus 1701 assisted reproductive techniques 2814–15 breeding management 2807–9 choice of stallion and semen type 2807–9 dietary support 2755–9 early embryonic loss 2327, 2455, 2638, 2991 management 2334, 2335–6 pre-uterine factors 2328, 2329 uterine factors 2330–2 estrous cycle abnormalities 1755, 1758, 2635 fertility 2327, 2803, 2804, 2990 foal growth rates 281 general expectations 2803 oocyte abnormalities 2329, 2814 ovarian senescence 2123, 2124 ovulation failure 2808 placental development 91, 2237–8 post-breeding management 2809–13 pre-breeding examination 2803–5
pregnancy and peripartum period 2813–14 preparation for breeding season 2805–7 uterine vascular pathology 2601 see also mare, age; older mares geriatric stallions breeding management 1209 dietary support 2755–9 penile and preputial neoplasia 1137, 1169 spermatogenesis 1048 see also stallion, age Germany, semen freezing schedule 2974 germ cells (germinal cells) primordial 1015, 1046, 2212–14 testicular 1028–36 anatomy 869, 870, 871 apoptosis 916, 924, 1021–3 cellular associations (stages) 885–9, 915–16, 920, 921, 922, 923 degeneration 890–2, 1039, 1040, 1040 development see spermatogenesis drugs adversely affecting 1177 intercellular bridges 1028, 1032 premature, in semen 1302, 1304 pubertal development 1016–17 Sertoli cell functions 884, 918 thermal sensitivity 881 see also spermatids; spermatocytes; spermatogonia; spermatozoa gestational length factors influencing 121 induction of parturition and 2262–3 mare nutrition and 2764 normal 111, 121, 2154 previous history 1898 prolonged see prolonged gestation variability 2448–9 zebras 2856–7 GGT, serum, neonatal foal 174 GhentTM semen extender 2967 Giardia, foal diarrhea 741 Giemsa stain chromosome banding 1462, 1463, 1464, 1953, 1954 sperm evaluation 1494, 1511 testicular fine needle aspiration 1442 GIFT see gamete intrafallopian transfer Gi proteins, acrosome reaction 949 glans clitoris see clitoris glans penis 875, 878 fetal sex determination 2086, 2089, 2090, 2091 premature tumescence 1420 Glanzmann’s thrombasthenia 428–9 glass marbles, intrauterine see intrauterine glass marbles/devices glaucoma, congenital 539–40 glicentin see enteroglucagon globe (of the eye) acquired disorders 540–1 congenital disorders 537–40 glomerular disease, renal 648–50 glomerular filtration rate (GFR) acute reduction see acute renal failure neonatal foal 647 single nephron (SNGFR) 649 glomerulonephritis 649–50 glucagon 70, 76 glucoamylase 264 glucocorticoids adrenal 1665 see also cortisol
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
exogenous see corticosteroid therapy pregnant mare 2228–9 glucometers 181–2, 836–7 glucose amniotic fluid 77 blood critically ill foal 181–2, 526 fetus 76 grain test in aged horses 2756 measurement 181–2 neonatal foal 76–7, 174, 175, 181, 187, 525–6 reference values 852 see also hyperglycemia; hypoglycemia infusions see dextrose infusions metabolism embryos 2175 foal 75–6, 77, 264–5 pregnancy 2229 sperm 938–9, 1063–4 placental transfer 28, 76, 89 semen 1062 semen extenders 1200–1 transport, sperm 1063 glucosinolate 529–30 glutamate circadian timekeeping system 1772–3 GnRH secretion and 1614 glutamate dehydrogenase (GDH) 409, 410 glutamine semen cryopreservation 2970 supplements, foals 266, 268, 277 glutaraldehyde, sterilization of endoscopes 1450 glutaraldehyde clot test colostrum 343, 832 foal serum or blood 343, 347, 834 glutathione peroxidase (GSH-Px) reactions 940, 992 red cell activity 466, 501 semen 941, 991 glutathione S-transferase α (GSTα) 2712–13 glycemic index (GI) 288 glyceraldehyde-3-phosphate dehydrogenase (GAPDH) 937 glycerin, nasopharyngeal irritation 272 glycerol diffusion properties 2959–60 embryo cryopreservation 2887, 2888–98, 2900–1, 2902–9 embryo vitrification 2913 semen cryopreservation 1264, 2962, 2969 glyceryl phosphoryl choline (GPC) 1062 glycogen 76, 939 glycogen branching enzyme deficiency (GBED) 463–4, 677–8 glycolysis, sperm 937, 938, 939, 1063–4 glycopyrrolate 843 glycosaminoglycans (GAGs) 779 glycosuria, neonatal foal 655 GM2-activator protein (GM2AP) 2178 GnRH see gonadotropin-releasing hormone goats, surrogate dams for orphans 273 goat’s milk composition 268, 269, 2281 enteral feeding 269 goiter 471, 530 see also hypothyroidism gold salts 839
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Index gonad(s) fetal development 2212, 2697–8 hypertrophy, eCG and 1659 primordial germ cell migration 1046 sex determination and differentiation 2212–15 steroidogenesis 19, 89, 1633–4, 1659, 2225–7 ultrasonography 48, 2011, 2086, 2092, 2096, 2097–8 grafting 3054 hypoplasia 2701 intersex states 1136 see also ovaries; testes gonadal dysgenesis 1756, 1956–7 63X see X chromosome monosomy cervical abnormalities 2719 uterine abnormalities 2607 XY sex reversal 1960, 2217–18 gonadotropes 1619 estradiol (E2) actions 1626–7 GnRH receptors 1609–10, 1613 gonadotropin(s) 1619–27 α-subunit 1619, 1649 β-subunits 1619–20, 1649 biological properties 1650 control of secretion 1625, 1639 epithelial ovarian neoplasia and 2710 melatonin and 1675–7 PGF2α -induced secretion 1799 see also equine chorionic gonadotropin; follicle-stimulating hormone; human chorionic gonadotropin; luteinizing hormone gonadotropin-inhibitory hormone (GnIH) 1743 gonadotropin-releasing hormone (GnRH) 897, 1608–15, 1783–6 challenge tests GnRH vaccine-induced anestrus 1701 pubertal fillies 1692 single pulse 1441 stallion subfertility/infertility 1436–6, 1439–40 three pulse 1439–40 degradation 1610 exogenous 1820–3 controlled release formulations 1882–3 cryptorchidism 1150 early studies in mares 1820 effects on spermatogenesis 1181–2 lactational anestrus 1700 osmotic mini-pumps 1820–1 poor libido in stallions 1416 studies in mares 1613 transitional mares 1709–10, 1785–6 see also gonadotropin-releasing hormone (GnRH) analogs mare autumnal transition 1741–3 control of FSH and LH 1623–4, 1783–4 control of seasonality 1782, 1783 puberty 1690 seasonal anestrus 1698, 1705–6, 1784 secretion patterns 1611–12 secretion rates 1612–13 vernal transition 1706 melatonin and 1675–6, 1748 molecular biology 1608–10
naturally occurring variants 1608–9, 1624 neurons 1610–11, 1783 physiology 1610–15 pulse generator 1610–11 secretion 1610–13, 1783 constant 1602, 1612 environmental influences 1614–15 factors affecting 1613–15 monitoring methods 1611 opioid-mediated regulation 999, 1614 pulsatile 1601–2, 1611–12, 1783 self-priming phenomenon 1625 signal transduction 1609–10 stallion 897 puberty 903, 904 regulatory system 898–900, 996–7 seasonal variations 1004 testicular secretion 898 structure 1608 synthesis 1608 gonadotropin-releasing hormone (GnRH) analogs 897, 1608–9, 1820–3 antagonistic see gonadotropin-releasing hormone (GnRH) antagonists controlled release formulations 1882–3 implants see gonadotropin-releasing hormone (GnRH) implants injectable 1822–3 mare early studies 1820 estrus suppression 1850 maintenance of pregnancy 2335, 2472 mature oocyte collection 2936, 2938 oocyte recipients 2942 ovarian follicle development and 1755–6 ovulation induction 1820, 1821–3, 1862–4 physiological studies 1613 seasonal anestrus 1699 superovulation protocols 1836, 1839 transitional mares 1709–10, 1785–6, 1822–3 stallion behavior management 1194, 1417 subfertility 1442–2 see also buserelin; deslorelin; goserelin acetate gonadotropin-releasing hormone (GnRH) antagonists 1609 ovulation failure 2689 stallion-like behavior 1194, 1417 see also cetrorelix gonadotropin-releasing hormone-associated peptide (GAP) 1608 gonadotropin-releasing hormone (GnRH) implants 1820, 1821–2, 1882 inducing anestrus 1700 seasonal anestrus 1785–6 see also deslorelin, implants gonadotropin-releasing hormone (GnRH) receptors 1609–10 downregulation/desensitization 1610, 1613 estradiol (E2) actions 1626 seasonal variations 1743 gonadotropin-releasing hormone (GnRH) vaccine mare
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causing anestrus 1700–1, 1762 contraception 1890 estrus suppression 1850–1 ovarian effects 1756, 2125 stallions 1191–6 contraception 1889 EAV shedders 1194–5, 1252 reversibility 1195–6 sperm motility effects 1295 suppression of sexual behavior 1192–3, 1417 gonocytes 1015, 1016, 1046 conversion into spermatogonia 1017 goserelin acetate implants, estrus suppression 1850 osmotic mini-pumps 1820 G protein-coupled receptors (GPCR) 1605, 1606 grain-mix concentrates see concentrates, feed grain test, poor absorption/digestion 2756 Gram-negative bacteria identification 1968–9, 1970, 2676, 2678–80 neonatal sepsis 156, 253–4, 255 Gram-positive bacteria identification 1969, 1970, 2676, 2677–8 neonatal sepsis 156, 253–4, 255 Gram stain 1970, 2676 granular dermatitis see habronemiasis, cutaneous granulosa cells embryonic development 2212–14 immature oocytes 2932, 2933 luteinization 2059 mature oocytes 2937, 2938 steroidogenesis 1631, 1632 granulosa layer 2032–3 serration of see serration of granulosa granulosa theca cell tumors (GTCT; GCT) 2710, 2711–13 breeding problems 2637–8 effects on contralateral ovary 2126, 2127, 2712 endometrial biopsy 2712 estrous cycle abnormalities 1756–7, 1763, 1764 gross pathology 2565, 2708, 2711 hormonal activity 2711–12 immunohistochemistry 2712–13 inhibin function 1854, 2712 laparoscopic removal 1991, 1997–8, 2570–1 molecular pathogenesis 2710–11 prognosis 2713 rectal palpation 2711 treatment 2713 ultrasonography 2132–3, 2711 gray coat color, genetics 2820 gray gene 2829 gray hair, aged horses 2756 grazing behavior fescue grasses 2423 in natural conditions 1387 Great (LGR8) 1002–3 Gr´evy’s zebra (Equus grevyi) foal 2862 interspecies hybrids 2302, 2304 mare estrous cycle 2856 gestation 2857 ovulation 2855 parturition 2858
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Gr´evy’s zebra (Equus grevyi) (Cont.) postpartum estrus 2859 puberty and breeding life 2854–5 reproductive organs 2855 seasonality of breeding 2853 mating patterns 2851–2 stallion behavior 2859 puberty and breeding life 2861 reproductive organs 2861 weanling 2862 yearling 2863 griseofulvin 843 grooming behavior, mutual 1387 growth, equine 280–8 broodmare nutrition and 286–7, 2766 catch-up, after illness 272 commercial importance 285–6 developmental orthopedic disease and 777, 782–3 factors affecting 281–5 fetus see fetal growth foal 67, 282, 287–8 fueling 286–8, 790–1 rates and spurts 285 retardation, chromosomal abnormalities 1956–7, 1958–9 studies 280–1 weanling 282, 287–8 yearling 282, 288 growth curves 280, 281 growth factors, placental development 86, 1655, 2236 growth hormone (GH) fetal production 2230 male reproductive function 998–9 pregnancy and lactation 2229 treatment, subfertile stallions 1443 growth models 280 growth plates, metaphyseal see physes guaifenesin 58, 843 gubernaculum testis 901, 902, 1016, 1099 abnormalities causing cryptorchidism 1102, 1532 inguinal extension of (IEGT) 1532, 1534 testicular descent 1099–100, 1531–2 gut-associated lymphoid tissue (GALT) 338 gut hormones 70, 77, 78 guttural pouch tympany (GPT) 612–13, 664 genetic basis 616 surgical correction 613, 664 gynogenesis 3054 H2 antagonists neonatal foal 395 older foal/weanling/yearling 685, 737, 743 see also raniditine Habronema spp., stomach infections 294 habronemiasis, cutaneous (summer sores) 294 diagnosis 1140 hemospermia 1122 penis and prepuce 294, 1139–40, 1451 treatment 1140 halo effect, Fell Pony syndrome 559, 560 halothane 224 halters, breeding stallions 1392, 1397, 1398, 1403
hamartomas congenital vascular cutaneous 559, 562 ovarian 2699–700, 2714 testicular 1110 Ham’s F-10 medium 2881–2 Hancock’s (eosin-nigrosin) stain 1494, 1511 hand hygiene, NICU 826 hand washing facilities, NICU 822–3 haploid genome 927 haptoglobin 851 harem bands 1385 zebras 2851 harem stallions 1385 assistant 1385–6 courtship and copulation 1388 parental behavior 1389 parturition guarding 1388 Hartmann’s solution 141, 206, 207, 231 Hartmann’s zebra 2851 foal 2862 mare estrous cycle 2856 gestation 2856–7 postpartum estrus 2859 puberty and breeding life 2854 reproductive organs 2855 seasonality of breeding 2853 stallion puberty and breeding life 2860 reproductive organs 2861 seasonality 2860 weanling 2862 yearling 2862–3 see also mountain zebra HarVet model artificial vagina 1268 Havemeyer grading system, arytenoid movements 796 hCG see human chorionic gonadotropin head, fetal 46 amputation 2500, 2501 malpostures 2489–91 fetotomy 2500–1 lateral reflexion 2490 ventral deviation 2490–1 normal posture 2271, 2482 head, foal examination 172, 478, 480 fractures 453–4 injuries 487–9 head-man, breeding shed 1225 headshaking, stallions 1425 head snares, obstetric 2486, 2489, 2490 head tilt, neonatal foal 484 healthcare-associated infections see nosocomial infections Heape, Walter 1262 heart auscultation in foals 500, 512 dimensions in foals 503 heart disease acquired 511–17 congenital 507–9, 514 diagnosis and monitoring 499–505 heart failure, congestive (CHF), weight loss 765 heart rate (HR) fetal see fetal heart rate mare periparturient hemorrhage 2518 uterine torsion 2439 neonatal foal anesthesia 224
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
birth resuscitation 132, 135 critical illness 139, 168 normal values 65, 500, 512 older foal 500, 512 heart rhythm, foal 500 heart valve dysplasia 508 heat see estrus heating supplies, foals 835 heat lamps, NICU 824 heat-moisture exchange device (HME filter) 218 heat stress, early embryonic loss 2332 height, wither, at birth 282 helper (CD4+ ) T-cells 333, 334 uterus 2590 hemangiosarcoma 516 hematocele 1110, 1153, 1162, 1163 ultrasonography 1477–8 hematology neonatal foal 173–4 reference values 173, 420 reference values 851–6 hematomas ovarian 1911, 2130–1, 2702, 2708 penis 1131–2 rectal palpation in mare 1906 testicular 1474–5, 1528–9 ultrasonography in mare 2154 umbilical 642–3 uterine 1947, 2450, 2671, 2672 vaginal 1944, 2154, 2451, 2452 hematuria 655–6 geldings, urethral rents 1140–1, 1142 hemi-zona assay (HZA) 1287, 1503 hemocytometer, sperm counting 1201, 1202, 1280–1 hemodynamic diagnostic techniques, advanced 505 hemoglobin-based oxygen carriers (HBOCs) (Oxyglobin) 846 anemia 424 neonatal foal 209, 238 administration 239 prior to transport 231 shock 142 hemoglobin (Hb) concentration neonatal foal 142, 173, 419, 420 reference values 851 hemolytic anemia, neonatal foal 419–20 clinical signs 421–2 diagnosis 423 management 424–5 see also neonatal isoerythrolysis hemolytic disease of newborn see neonatal isoerythrolysis hemopericardium 516 hemoperitoneum, ultrasonography 243, 249 hemophilia 423, 429 hemorrhage neonatal foal 419 clinical signs 420–1, 426 diagnosis 427 management 424 periparturient see periparturient hemorrhage postoperative castration 1560 ovariectomy 2566, 2571–2 partial phallectomy 1570 retinal 551 subconjunctival 544 umbilical 642–3
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Index hemorrhagic anovulatory follicles (HAF or AHF) 1738–9, 1758, 2701–3, 2702 differential diagnosis 2709 gross pathology 2703, 2704 management 2636 ultrasonography 2016–17, 2048, 2049, 2129–30, 2131, 2686–8 hemorrhagic shock periparturient hemorrhage 2518 uterine prolapse with 2433 hemospermia 1121–4, 1278–9 clinical signs 1122 diagnosis 1122–3, 1141 endoscopy 1123, 1453–3, 1455 etiology 1121–2 prognosis 1124 treatment 1123–4 urethral rents causing 1122, 1140–1 hemostatic disorders diagnosis 422–3 neonatal foal 419, 421 hemostatic suture, hysterotomy closure 2506, 2507–8 hemothorax 580 Henderson castration instrument 1559–60 heparin 843 heparinized saline 843 hepatic abscessation complicating omphalitis 637 ultrasonography 249 hepatic disease see liver, disease hepatic failure see liver, failure hepatic lipidosis, neonatal foal 413 hepatocellular carcinoma 516, 765 hepatocyte growth factor-scatter factor (HGF-SF) 1655, 2236 hepatoencephalopathy 409, 410, 414 Hepes buffer 1336, 2881, 2933 Hepes-buffered TCM199 with Hanks’ salts 2932 herbal supplements, suppression of estrus 1762 HERDA syndrome see hereditary equine regional dermal asthenia syndrome herds, horse 1385–7 dominance hierarchies 1387 social organization 1385–7 hereditary equine regional dermal asthenia (HERDA) syndrome (hyperelastosis cutis) 555, 566–7, 670, 678–9 hermaphrodite, true clinical signs 1111, 1136, 2216 cytogenetic analysis 1961 terminology 2215 hernias ultrasonography 251 see also diaphragmatic hernia; inguinal hernia; umbilical hernia hetastarch 843 neonatal foal 141, 209 older foal/weanling/yearling 736 heterochromia iridis 546 hexokinase 938 high flanker see cryptorchidism, inguinal high hydrostatic pressure (HHP) treatment, semen for freezing 2979 high-risk pregnancy conditions causing 6–14, 16, 17 fetal 6, 7, 12–14 maternal 5, 6–10 placental or umbilical 5, 6, 10–12 estrogen therapy 19, 1826–7
identification 5–14 management 25–35, 111 fetal conditions 35 maternal conditions 30–2 placental conditions 32–5 pro-gestational therapies 25–9 specific causes 30–5 minimum data base 6, 7 monitoring 16–22 see also feto-placental monitoring potential complications 18 hindlimbs fetal amputations 2502–3 damage to uterus 2480 encasement in cord horn 2479–80 impaction 2487–8 normal delivery 2271, 2482 neonatal foal, paresis 171 hinny 2302, 2303, 2835 chorionic gonadotropin function 1659 fertility 2302, 2304–5 hip dysplasia 667 hip flexion caudal presentation 2492–3, 2502 ventro-vertical presentation with 2487–8, 2502 hippomanes 51, 87–8, 104, 105 hirsutism, inappropriate 2791 histometric analysis, testicular biopsy 1524–5 Histoplasma capsulatum 588, 711 hobbles, breeding 1270 hock see tarsus hock flexion fetal manipulation 2492 fetotomy 2503 Hoechst 33258 1495–6, 1502, 1512 Hoechst 33342 1495, 1512, 3043 Holliday–Segar formula, critically ill neonate 191, 192 Holter monitoring, 24-hour 502 homozygous diploidy 3054–5 hoof care angular limb deformities 769–70 breeding stallion 1221 broodmare 2748 flexor tendon laxity 469, 470 flexural deformities 443 hoof–ground angle, flexural deformity 441, 443 hormones 1601–7 amino acid derivatives 1602, 1607 cell interactions 1604–6 classes 1602–4 fatty acid derivatives see eicosanoids feedback/control mechanisms 1604 fetal 2230 half-life 1604 history of term 1601 mechanisms of action 1606–7 methods of signaling 1601–2 peptide and protein 1602, 1603, 1606 pregnancy 16–19, 2222–30 stallion 897–900 steroid see steroid hormones transport 1604 see also endocrinology; specific hormones hormone treatment anovulation associated with 2685, 2689 breeding stallions 1416 cryptorchidism 1150, 1443 foal rejection 119, 120 maintenance of pregnancy 2469–72
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new delivery techniques 1879–84 ovarian changes 2125 seasonal anestrus/anovulation 1699 stallion subfertility 1442–3 see also specific hormones and agents horse, domestic (Equus caballus) domestication 1261 genome sequence 620, 673, 962, 2827–8 interspecies hybrids 2302, 2303–5, 2835 ownership and use of 3051 horse blood agar (HBA) 1979, 1980, 1981 horse-in-donkey pregnancy 2306–7, 2308 horse owner assessed respiratory signs index (HOARSI) 618 Horserace Betting Levy Board (HBLB) Codes of Practice 1963, 2339–40 designated laboratories 1967, 1968 hot environments, hydrocele risk 1152 hourglass hypothesis, photoperiodic regulation of reproduction 1773 HTG6 microsatellite 2821–2 human chorionic gonadotropin (hCG) 1804–8 antibodies 1807–8, 2051 β-subunit 1649 challenge test cryptorchidism 1103–4, 1533 stallion subfertility/infertility 1438, 1441 cryptorchidism 1150, 1443 diestrous mares 1807 effects on spermatogenesis 1180–1 induction of abortion 2322 ovulation induction 1804–8, 1861–2 AI programs 1205, 1805, 2989–90 antibodies 1807–8, 2051 applications 1804–5 dose 1862 dose and route effects 1806–7 effect of mare’s age 1806, 1866 efficacy 1805–6, 2689 embryo recipients 2878 estrus suppression 1849–50 follicle diameter and 1805, 1866 geriatric mares 2807–9 low dose insemination 3037, 3039 luteal development and 1806 oocyte collection 2936, 2938 oocyte transfer 2941, 2942 response interval 1806 timing and follicle diameter 1805 twin pregnancy rates 2351–2 vs. deslorelin 1862, 1863, 1865–6 ovulation synchronization 1807, 1873, 1874 preparations 1804 subfertile stallions 1443 superovulation 1837, 1840, 1841 efficacy 2689 testicular actions 894 human menopausal gonadotropin (hMG) 1650 humeral fractures, foals 451, 452 humidifiers, ventilated foal 218 humidity, relative, during stallion shipment 1189 humoral immunity development 335, 336–8 recurrent airway obstruction and 619 tests 366, 367 humoral immunodeficiencies 361–5, 370 Hunter, John 1262 hurdling posture, fetus 2487–8 fetotomy 2502
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hyaloid vascular system, remnants 536–7, 550 hyaluronate sodium 843 hyaluronidase, cumulus removal 2934 hybrids, equine 2302–5 see also hinny; mule hydatid of Morgagni 2692, 2701 hydrallantois (hydrops allantois) 93, 2368–71, 2444–5 causes 2370 clinical signs 2369 complications and prognosis 2370–1 diagnosis 12, 2369–70 fetal anomalies 2371 high-risk pregnancy 6, 12 induction of abortion 2323 terminology 2368 treatment 35, 2371 ultrasonography 21, 2151, 2369 hydramnion (hydrops amnion) 93, 2368–71, 2444–5 causes 2370 clinical signs 2369 complications and prognosis 2370–1 diagnosis 2369–70 high-risk pregnancy 12 induction of abortion 2323 terminology 2368 ultrasonography 2152, 2369 hydranencephaly 669 hydration assessment, neonatal foal 167, 177 hydrocele 1152–3, 1162 complicating castration 1152–3, 1162, 1561 congenital 1110, 1477 diagnosis 1146, 1152 treatment 1152–3 ultrasonography 1152, 1477 hydrocephalus 483 dystocia 2494 fetotomy 2502 internal 669 hydrochlorothiazide, hyperkalemic periodic paralysis 675 hydrocortisone 843 hydrogen peroxide (H2 O2 ) adverse effects on sperm function 992–3 generation by sperm 940, 991, 1064 mechanism of sperm damage 993 scavenging 940, 941, 992 uterine irrigation 2634, 2648 hydrops 12, 93, 2368–71, 2444–5 causes 2370 clinical signs 2369 complications 2370 diagnosis 12, 2369–70, 2444–5 fetal anomalies 2371 prognosis 2370–1 terminology 2368 treatment 33–5, 2323, 2371, 2445 ultrasonography 2151–2, 2369 see also hydrallantois; hydramnion hydrosalpinx 2692 hydroureter endoscopy 1456 syndrome, neonatal foal 629–31 hydroxyethyl starches, neonatal foals 141 hydroxyindole-O-methyltransferase (HIOMT) 1671, 1675 17α-hydroxylase (17α-OH) periparturient period 18, 69, 2224–5 vernal transition 1707
3β-hydroxy-5α-pregnan-20-dione (3β-5P) 2222 20α-hydroxy-5-pregnan-3-one (20α-5P) 2223, 2224 17α-hydroxyprogesterone (17αOH; P4) 69, 71 hydroxyprogesterone caproate 1813 estrus suppression 1847 maintenance of pregnancy 2470 hydroxyprogesterone hexanoate 1813 maintenance of pregnancy 2121, 2471 15-hydroxy-prostaglandin dehydrogenase (PGDH) 1645 3β-hydroxysteroid dehydrogenase (3β-HSD) 2223, 2224, 2225–7 11β-hydroxysteroid dehydrogenase type 2 (11β-HSD2 ) 122 endometrial fibrosis 1934 hydroxyzine 241, 843 hymen imperforate 1943–4 persistent 1726, 1902, 2720 remnant see vestibulovaginal fold hyperammonemia congenital portosystemic shunts 414 and hepatopathy, Morgan weanlings 413–14, 732 syndromes in foals 732 hyperbaric oxygen therapy, septic arthritis and osteomyelitis 460–1 hypercalcemia, neonatal foal 212 hypercapnia neonatal foal 185, 578 anesthesia/sedation 222, 223 critical care 169–70 indications for therapy 215–16 perinatal asphyxia syndrome 150 periodic apnea 575 pneumonia 587 prematurity or dysmaturity 125 respiratory support 189, 216–19 sepsis 159 ventilatory response 575 permissive 215–16 hypercoagulation, neonatal foal 426, 429 hyperelastosis cutis see hereditary equine regional dermal asthenia (HERDA) syndrome hyperglycemia, neonatal foal 175, 181, 212–13 parenteral nutrition 275–6 prior to transport 234–5 hyperkalemia bladder rupture 626–7 neonatal foal 182, 211–12 fluid therapy 141, 191, 211–12 renal tubular acidosis 651 hyperkalemic periodic paralysis (HYPP) 464, 668, 673–5 clinical signs 464, 673–4 diagnosis 464, 674 ejaculation problems 1421 etiology 674 genetics 674, 2824, 2825, 2829 management 464, 674–5 breeding stallions 1235 prognosis 675 hyperlipidemia Miniature ponies 2841, 2842 neonatal foal 413 hypernatremia, neonatal foal 182, 192, 210–11 shock 141 treatment 191–2, 211
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
hyperparathyroidism, nutritional secondary 1234 hyperthermia, early embryonic loss 2332 hypertonic fixation test, blood–testis barrier 1019, 1020, 1021 hypertonic saline 843 hypoxic-ischemic encephalopathy 487 neonatal foal 190, 207, 208, 210 neonatal shock 141 older foal/weanling/yearling 736 periparturient hemorrhage 2518 hyphema, mare reproductive loss syndrome 429 hypoalbuminemia, equine proliferative enteropathy 761, 762 hypocalcemia lactating mare 2282 neonatal foal 182, 212, 526 hypocapnia, neonatal foal 169 hypogammaglobulinemia of the young, transient 362, 363 hypoglycemia glycogen branching enzyme deficiency 463 late-term mare 29, 2229 neonatal foal 175, 181, 212 anesthesia 224–5 catecholamine response 73 critical illness 526 hepatic failure 409 parenteral nutrition 275 prematurity or dysmaturity 124, 125 prior to transport 231, 234 sepsis 155, 199 hypogonadism hypogonadotropic, stallion 1182, 1442 primary, mare 1701 hypokalemia, neonatal foal 206, 211 critical care 191 parenteral nutrition 276 hyponatremia bladder rupture 626–7 hydroureter syndrome of neonatal foal 630 neonatal foal 182, 191–2, 209–10 neonatal renal disease 657 hypo-osmotic swelling (HOS) test 957, 1286, 1500–1 assessing semen freezability 2973 frozen-thawed semen 2980, 2998 hypophysis see pituitary gland hypopituitarism, mare 1700 hypoplastic left heart 509 hypoproteinemia equine proliferative enteropathy 761, 762 neonatal foal 209 hypospadias 1111 hypotension anesthesia-induced neonatal foal 224 pregnant mare 55, 58–9 shock 136 hypothalamic–pituitary–adrenal (HPA) axis control of reproduction 1615 domperidone actions 2425 fetal, maturation 18, 69–72, 122–3, 2230 accelerated 123 incomplete 72, 124, 125, 524, 525 neonatal foal 71–2, 523–6 premature foal 124, 524 septic foal 155, 525 sick foal 72, 524–5
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Index hypothalamic–pituitary–ovarian axis 1604 feedback regulation 1625–7, 1639 hypothalamic–pituitary–testicular (HPT) axis 897–900, 996, 997 drugs adversely affecting 1178–82 dynamics 996–7 puberty 903–4 seasonal effects 1003–5 hypothalamic–pituitary–thyroid axis 526–30 hypothalamus 897, 1582–4 GnRH neurons 1610–11 hypophysial connections 1585–6 rostral nuclei 1582, 1585 hypothermia, neonatal foal anesthesia/sedation 222, 224–5 neuroprotective effect 232–3 prematurity or dysmaturity 125–6 prior to/during transport 233–4 hypothyroidism 527–30 congenital 528–30, 669 common digital extensor tendon rupture 471–2, 528 cutaneous features 561–3 pathogenesis 529–30 delayed epiphyseal development 436 early embryonic loss 2330 fescue toxicosis 530, 2422 geriatric mares 2806 hyperiodic 530, 561 other causes 530 therapy 530, 1883, 1884 hypotrichosis 567–8 hypovolemia, neonatal foal 80 causes 136, 137 clinical signs 154–5 evaluation 177 management 140–2, 157, 190–1 prior to/during transport 230–1 hypovolemic shock, neonatal foal 136 hypoxemia, neonatal foal 215, 578 critical care 168, 169 diagnosis 577 monitoring 185 perinatal asphyxia syndrome 150 persistent pulmonary hypertension 590, 597 pneumonia 605 prematurity or dysmaturity 125 respiratory support 189, 216–19 sepsis 155, 159 shock 144 hypoxic-ischemic encephalopathy (HIE) see neonatal encephalopathy hysterectomy 2574–6 chronic pyometra 2574, 2654 complications 2576 indications 2574 partial 2574, 2576 prognosis 2576 surgical procedures 2574–6 hysteroscopic insemination 1205, 3038–9 epididymal sperm 3039–40 fresh semen 3038–9 frozen-thawed semen 3039 historical perspective 1266 low dose 3039 procedure 1943, 1944 sex-sorted sperm 3045, 3046 hysteroscopy 1940–3 endometrial biopsy via 1942, 1943, 1944 endometritis 1946, 1947, 2612
equipment 2666 healthy endometrium 1942 infectious pathological conditions 1946 normal anatomy 1940, 1941 pathological conditions 1945–9 postpartum uterus 1943 problem breeders 2627 procedure 1940–2 uterine cysts 1942–3, 1945, 2266–8 hysterotomy, closure techniques 2506–8 ibuprofen 843 Icelandic fillies, age of puberty 1692 ICSI see intracytoplasmic sperm injection icterus see jaundice icterus index 851 IDENT system, sperm counting 1282 ideograms 1952 R Foal IgG Test 347, 348, IDEXX SNAP 834 IGFs see insulin-like growth factor-1; insulin-like growth factor-2 I κB 1606 IL4RA gene polymorphisms 619 ileal obstruction, congenital 47 ileocolonic aganglionosis 407–8, 668, 675–6 differential diagnosis 381 see also overo lethal white foal syndrome ileus, neonatal foal 137–8, 139, 193 ultrasonography 246 iliac arteries, ultrasonography 1485 ill-thrift, older foals/young horses 758–65 Ilomostat, topical 546 imidocarb diproprionate 843 imipenem 843 neonatal foal 254, 259 R. equi pneumonia 700 imipramine ejaculatory problems 1414 induction of ejaculation 1271, 1414 manipulation of ejaculate 1414 priapism-induced impotence 1136 urospermia 1127 immune-mediated thrombocytopenia (IMTP) 374 immune responses donkey-in-horse pregnancies 2314, 2315, 2316–17 early embryonic loss and 2332 endometrial cups 1654, 1656–8, 2193–4, 2197 horse-in-donkey pregnancies 2306–7 horse-in-mule pregnancies 2309, 2310 placental 89 transferred exotic equid embryos 2312, 2313 immune system 331–9 adaptive 331, 334–8 components 332 ergot alkaloid interactions 2418 fetal 334–5 innate 331–4 mucosal 338 neonatal foal 67, 331–8 uterus 2587–92 immune tolerance, seminal antigens 1096–7 immunocastration 1191–6 EAV shedders 1194–5, 1252 inhibition of sexual behavior 1192–3, 1417
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reasons for 1191 reversibility 1195–6 immunocontraception mare 1889–92 stallion 1889 immunodeficiencies 361–6, 369–71, 373–5 cellular 365–6, 370 complement component 366 humoral 361–5, 370 immunological testing 366, 367 phagocyte 366 primary 361 secondary 361 immunoglobulin(s) 335 bovine-derived 344 endogenous production 67, 336–8 intravenous administration, neonatal foal 349–50 oral administration, neonatal foal 344, 349 passive transfer of maternal see passive transfer of maternal antibodies severe combined immunodeficiency 370 uterine defenses 2587–8, 2608 see also specific isotypes immunoglobulin A (IgA) foal immunity 67, 338, 342 uterine secretions 2587–8 immunoglobulin E (IgE) foal 337–8, 342 recurrent airway obstruction 618, 619 immunoglobulin G (IgG) colostral content 64, 342–3, 2277–8 assessment methods 342–3, 832–3, 836, 2279, 2280 temporal trend 336, 337, 2278 colostral transfer see passive transfer of maternal antibodies endogenous production by foal 67, 335, 336–7 foal serum acceptable values 113, 175, 346, 833 age-related trend 336, 337 assay methods 347–8, 833–4, 836 humoral immunodeficiencies 363, 366 isotypes 335 laboratory tests 832–4 supplements 344 uterine secretions 2587–8 see also failure of passive transfer; passive transfer of maternal antibodies immunoglobulin M (IgM) foal immunity 335, 342 non-selective deficiencies 366, 370 selective deficiency 362, 364, 374–5 uterine secretions 2587–8 immunological treatment donkey-in-horse pregnancies 2315–16, 2317 shuttle stallions 1189 immunomodulators/immunostimulants chronic endometritis 2613 maintenance of pregnancy 2474–5 older foal pneumonia 717 placentitis 2363 post-mating endometritis 2812–13 prevention of embryonic loss 2335 problem-breeding mares 2631 implantation fossa 928, 1058
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import cooled semen 1325 embryos 2921–3 frozen semen 2972, 3013, 3029–35 impotence complicating cavernosal shunting 1565–6 priapism-induced 1135–6 see also erection, penile impregnation see reinforcement breeding imprinting gametic 3055 genomic, placental development 2310, 2311 inbreeding early embryonic loss 2332 morphological sperm defects 1365 incest avoidance, in natural conditions 1389 incubators, microbiological 2675 index cards, frozen semen 3009–10 India ink stain 1494 Indian hedgehog (Ihh) 776 indole test, bacteriological 2676 induction of parturition 29, 2262–6 abdominal wall rupture 31–2, 2429, 2448 cervical ripening prior to 1801, 2264–5 criteria for 2262–5 glucocorticoids 27, 123, 1667, 2265 hydrops of fetal membranes 34–5, 2445 methods 2265–6 oxytocin 29, 1832–3, 2265–6 prostaglandins 1800, 2265 infanticide, Asian wild horse 2849–50 infection control intestinal clostridiosis 389, 737–8 NICU 824–6, 827 rotavirus diarrhea 389 infections chronic, weight loss in foals 763–5 immunodeficiencies 361, 369–70 intrauterine causing abortion 90 placental evaluation 92 neonatal foal 158–60, 198–203 clinical signs and diagnosis 199–202 liver disease 409–12 pathogens 199 perinatal asphyxia syndrome 151 prognosis 203 risk factors 198, 347 routes of entry 199 treatment 202–3 ventilation-related 219 nosocomial see nosocomial infections postoperative, complicating castration 1560–1 pregnant mare 6, 8–10 abortion and stillbirths 2343 management 32 retained fetal membranes and 2522 infectious agents semen extenders 3030 shipped frozen semen 3029–30 testing frozen semen for 3002 see also bacteria; fungi; parasites; yeasts infectious diseases control AI for 1262 broodmare farms 2748–9 shipped frozen semen 3030–1 see also vaccination
pituitary pars intermedia dysfunction 2792 risk classification, NICU patients 825–6 risks from imported embryos 2922 transmitted via frozen or cooled semen 3015–26 inferior check ligament desmotomy 443–4, 445 infertility chromosomal abnormalities 1951 cooled transported semen programs 1327–8 GIFT 2946 pituitary pars intermedia dysfunction 1666, 2794–5 stallion see stallion subfertility/infertility see also fertility; problem-breeding mare infiltrative bowel disease 763 inflammatory airway disease (IAD) 617 inflammatory bowel disease 763 influenza, equine see equine influenza/equine influenza virus inguinal canal 867, 868, 1099, 1145 testicular descent 900, 1100 inguinal hernia 1147–9, 1540–4 acquired 1147, 1542 adult stallions 1147, 1540 breed differences 1540–1 clinical signs 665, 1147, 1541–2 congenital, foals 665, 666, 1110–11, 1147, 1164, 1540–1 clinical signs 1541–2 diagnosis 1541–2 non-surgical management 1147–8, 1541 presenting with colic 753, 754 surgical management 402–3, 1148, 1542 diagnosis 1146, 1147, 1541–2 direct 1147, 1540 etiology 1147, 1540–1 indirect 1147, 1540, 1541 non-surgical management 1147–8, 1541, 1542 ruptured 1147, 1540, 1541 diagnosis 1146 surgical correction 403, 1148–9, 1543 strangulated 1146, 1542, 1543 surgical management 1148–9, 1542–4 terminology 1540 ultrasonography 1478 inguinal herniorrhaphy adult stallions 1148–9, 1542–3 foals 1148, 1542 laparoscopic 1543, 1550–2 cylindrical mesh prosthesis 1552 foals, with castration 1551 foals, without castration 1551 mature stallions, without castration 1544, 1551–2 retroperitoneal mesh augmentation 1551–2 testicle-sparing techniques 1543–4, 1551–2 inguinal rupture 1147, 1540, 1541 clinical signs 1542 diagnosis 1146, 1147 etiology 1147 surgical correction 1148–9, 1542–3 inherited diseases see genetic disorders inhibin 1680–1 administration, mare 1854–5
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
fetal production 1681 function 1681 immunotherapy of mares 1854–6 active 1836, 1855 passive 1855–6 mare 1625, 1680, 1681 double ovulations 2023 estrus 1720 FSH and LH regulation 1625, 1681 function 1854 granulosa cell tumors 1854, 2712 periovulatory period 2024, 2025, 2047 physiology 1681 sources of secretion 1680–1 stallion 884, 898, 998, 1681 endocrine function 900, 998 infertility/subfertility 1439 paracrine–autocrine function 1002 puberty 1006, 1007 seasonal variations 1003–4 testicular levels 1438, 1439, 1440 structure 1680 inhibitor of κB (I κB) 1606 injected mucous membranes, neonatal foal 138, 154, 155 innate immune system 331–4 inner acrosomal membrane (IAM) 928–9, 932, 933, 1491 sperm–oocyte interaction 1491 inner cell mass 2171–2 inositol 1,4,5-trisphosphate (IP3 ) 949, 1605 inotropes, neonatal foal anesthesia 224 critical care 142–4, 192, 195 sepsis 158, 203 INRA-82 cooled semen 1337 frozen semen 2965, 2966–7, 2968–9 INRA-96 cooled semen 1318, 1337, 1341–2 frozen semen 2967–8 optimal storage temperature 1330–1 insect bite hypersensitivity 555 insecticides 2415–16 insemination, artificial see artificial insemination insemination dose 1201, 1273–4 contract issues 3014 cooled shipped semen 1201, 1323, 1326 cost considerations 3013–14 frozen-thawed semen 2977, 2980–1, 2988–90, 3036 historical perspective 1266 low dose insemination 3037–40 miniature horses 1380 recommendations 3036–7 insemination volume 1201, 1274 miniature horses 1380 Insl3 gene 1002 equine (eInsl3) 1003 insulin conceptus production 2204 developmental orthopedic disease and 287–8, 778, 785 fetal secretion 2230 foal adaptation to parturition 70, 75–7 GnRH secretion and 1615 plasma grain test in aged horses 2756 neonatal foal 76–7, 525–6 premature foal 526 pregnancy 2229
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Index therapy 843 critically ill neonatal foal 191, 195, 526 hyperkalemic periodic paralysis 464 neonatal foal 211, 213, 274, 276 insulin-like growth factor-1 (IGF-1) fetal growth and lactation 2229 male puberty 1007 osteochondrosis 776 ovarian folliculogenesis 2015 seminal plasma 1000 stallion subfertility 1438 testicular 999, 1000, 1440 insulin-like growth factor-2 (IGF-2) feto-placental growth 86, 2229, 2236 genomic imprinting 2311 insulin-like growth factor-binding protein 3 2178 insulin-like peptide 3 (INSL3) 1002–3, 2215 equine (eINSL3) 1003 insulin resistance (IR) mares 2636–7, 2806 pituitary pars intermedia dysfunction 2791, 2793 integrins, acute tubular necrosis 650 interestrus interval pyometra 2653 see also interovulatory interval interferon(s), type I 333–4 interferon-δ 2177 interferon-γ (IFNγ ) foal immune response 333, 336 R. equi susceptibility and 696–7 interferon-tau (interferon-τ ) 1644, 2177, 2201 interleukin 1 (IL-1) 334, 2592 interleukin 6 (IL-6) 334, 2592 interleukin 8 (IL-8) 334, 2590–1, 2592 interleukin 10 (IL-10) 334, 2590–1 interleukin 12 (IL-12) 333 interleukin 18 (IL-18) 2592 intermammary groove 1595 intermediate filaments 2712–13 internal carotid artery, fetal 46 internal fixation long bone fractures 452 mandibular fractures 454 olecranon fractures 450 rib fractures 455 internal pudendal artery 1594, 2559 International Embryo Transfer Society (IETS) 2899, 2922 International Society for Animal Genetics (ISAG) 2822, 2823 International System for Cytogenetic Nomenclature of the Domestic Horse (ISCNH) 1462, 1463, 1952–3 interovulatory interval (IOI) 2012 mare age and 2028 Miniature ponies 2839 prolonged deslorelin implants 1755–6, 1821–2, 1863 ovulation failure 1757 pyometra 2653 short-cycling 1729–30, 1798 see also estrous cycle(s) interphase 1952 intersex conditions 2215–19 abnormal male external genitalia 1111, 1136 chromosome abnormalities see sex chromosomes, abnormalities
clitoral hypertrophy 1579 cytogenetics 1960–1 ovarian abnormalities 2699, 2700, 2702 stallion-like behavior in mares 1764, 1765 terminology 2215–16 interspecies hybrids 2302–5 interspecies pregnancies 2303–5 interstitial cells fetal gonads 2698–9 testicular see Leydig cells interstitial nephritis 651–2 intestinal aganglionosis see ileocolonic aganglionosis intestinal atresia 406–7, 668 meconium retention 381 intestinal distention, fetal 47 intestinal malformations 406–7 intra-abdominal abscessation older foal/weanlings 756–7, 763–4 R. equi infection 243–4, 245, 698, 764 strangles 764–5 intra-abdominal hemorrhage periparturient 2451, 2517 uterine rupture 2433 intra-articular analgesia, foal 226 intracardiac drug administration fetal reduction 2114, 2115 neonatal resuscitation 133 intracrine action 1601 intracytoplasmic sperm injection (ICSI) 2948–51, 3052 frozen-thawed epididymal sperm 2985–6 history 2948 oocyte collection for 2931 oocyte cryopreservation 2955 oocytes from aged mares 2814 with Piezo drill 2948, 2949–51 procedures 2949–51 sex-sorted sperm 3046 uterine tube pathology 1988 intranasal oxygen (INO2 ), neonatal foal 189, 190, 216 intraocular pressure (IOP) 536, 539–40 intraosseous drug administration, neonatal resuscitation 133 intrauterine antibiotics chronic endometritis 2613, 2632–3 complications 2633 delayed uterine clearance 2661 foal heat breeding 2299–300 geriatric mares 2811 postpartum mare 2142 postpartum metritis 2533 pregnancy rates and 2146 prior to breeding 2145 problem breeders 2630, 2632–3 residues 2671 retained placenta 2525, 2528 intrauterine devices (IUDs), contraceptive 1892 see also intrauterine glass marbles/devices intrauterine fluid accumulation 2597, 2655–62 cervical adhesions 2722 early embryonic death and 2145–6 estrous mare 1723, 1725, 2657 oxytocin therapy 1830–1 hysteroscopy 1946 identifying susceptible mares 2657–8 pathogenesis 2655–7 post-breeding 2627–31
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AI recipients 1206, 2990–1 identifying susceptible mares 2627 management 2145, 2627–8, 2658–62 mares in foal heat 2299–300 oocyte recipients 2943 oxytocin therapy 1832, 2145, 2629, 2660 pathogenesis 1094–6, 2621–2 treatment options 2628–31 postpartum foal heat breeding and 2295, 2299, 2468–9 manual removal of placenta 2526 ultrasonographic monitoring 2141–2, 2299 pre-breeding 2144–5, 2657–8 geriatric mares 2808 pregnancy rates and 2145–6 prior to AI 1203 specimen collection 1923, 1967 treatment options 2658–62 ultrasonographic grading 2137, 2139, 2658 ultrasonography 2137–41, 2610, 2657 see also endometritis; uterine clearance intrauterine glass marbles/devices estrus suppression 1760, 1847–9 retained 2671, 2725, 2728 ultrasonography 2149 intrauterine growth retardation (IUGR) 91 Doppler velocimetry 21–2 foals with see dysmature foals skeletal development 437 ultrasonographic features 21, 22 intrauterine hemorrhage after manual removal of placenta 2526 peripartum 2449, 2450, 2517–18 intrauterine insemination see artificial insemination intrauterine therapy causing cervical adhesions 2721, 2722 chronic endometritis 2613–14 delayed uterine clearance 2661 fungal endometritis 2648–50 induction of abortion 2321–2 pre-breeding hemospermia 1124 problem mares 2144, 2145 suboptimal stallion fertility 1246–7 problem breeders 2629, 2630, 2632–5 reproductive efficiency and 2783 see also intrauterine antibiotics intravaginal progesterone devices 1814–15, 1879 estrus synchronization 1826, 1874–5 transitional mares 1780 intravenous catheters lost 517 thrombotic complications 429, 517 see also jugular vein, catheterization intravenous drug administration, neonatal foal 133, 140 intravenous fluid therapy see fluid therapy, intravenous intussusception cecocecal and cecocolic 757 small intestinal 244, 245, 402, 755 inulin clearance 653–4 inventory management systems, frozen semen 3009 in vitro fertilization 2945
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iodine deficiency 530 dietary requirements breeding stallion 1230 broodmare 2762, 2763, 2766 excess 530, 561 solutions, umbilical stump treatment 635, 637 iohexol 844 ionomycin 1493 ionophore toxicity 516 ipratropium, neonatal foal 606 iridocorneal angle, foal 536 iris coloboma 541, 546 color 536 congenital abnormalities 546–7 hypoplasia 546 iron deficiency anemia 420, 423, 425 dietary requirements, breeding stallion 1230 requirements, neonatal foal 265 saturation, neonatal foal 410 serum 419, 420, 853 iron-binding capacity total (TIBC) 419, 420, 853 unsaturated 853 iron fumarate, neonatal hepatopathy 412, 425 iron saturation 853 ischiocavernosus muscle 877, 878 isoerythrolysis, neonatal see neonatal isoerythrolysis isoflurane anesthesia 58, 224 isoxsuprine hydrochloride 27 itraconazole 844 fungal endometritis 2636, 2647–8 ophthalmic disorders 541 Ivanov, Il’ia Ivanovich 1263, 1266 ivermectin ascarids 295, 759 bots 293–4 cutaneous habronemiasis 1140 stomach worms 294 strongyles 298, 300 Strongyloides westeri 298, 386, 391 tapeworms 299 Izumo protein 953 jack donkey 2837–8 cryptorchidism testing 1104 mating with horse mare 2303, 2835 semen collection and processing 2837–8 Jaffe reaction 653 Japanese model artificial vagina 1268, 1269 jaundice (icterus), neonatal 138–9, 168 causes 414 hemolytic anemia 423 liver disease 409, 410, 411 neonatal isoerythrolysis 354, 421–2 jaundiced foal agglutination test (JFA) 360, 831–2 neonatal isoerythrolysis 354 technique 354–5, 359 JC-1, semen evaluation 957–8, 1289, 1513, 3001 jenny donkey 2835–7 breeding 2836 cervix anatomy 2724, 2725, 2835–6 chorionic gonadotropin function 1659 estrus synchronization 2836
exotic equid embryo transfer 2311–14 horse embryo transfer 2306–7, 2308 mating with horse stallion 2303, 2835 mule embryo transfer 2310–11 pathology 2836–7 predicting parturition 2270 pregnancy 2836 prostaglandin-induced luteolysis 1798 J-lube enemas, meconium impaction 381 Jockey Club, breeding record system 1211, 1212–17 joint development 437–8 implications of abnormal 438–9 nomenclature 434–6 dysplasia 667 effusions, neonatal foal 201 lavage, infectious arthritis 458, 460 jugular vein catheterization parenteral nutrition 274 prior to transport 230 catheter loss 517 examination 499, 512–13 thrombosis 429, 517 juvenile arthritis 775 juxtacrine action 1601 juxtacrine pathways, epididymis 924 kanamycin 256 kaolin/pectin products 737 karyotype 1952 clinical reports 1955 normal 1463, 1954 standard 1952–3 karyotyping 1462–4, 1524, 1951–5 mare subfertility/infertility 1955–9 procedures 1953–5 terminology 1951–2 R 211–12 Kayexalate KDR (VEGF receptor) 2240 Keeneland yearling study 810–17 KenneyTM semen extender cooled semen 1318, 1337, 1341 frozen semen 2966–7 semen storage temperature and 1330–1 sex-sorting of sperm 3043 keratitis, ulcerative see corneal ulcers keratocytes 422 kernicterus 422 kerosene, intrauterine infusion 2634 ketamine 844 foal 222, 223 pregnant mare 58, 59, 2505 stallion 1558 ketoconazole 844 fungal endometritis 2636, 2647 ketoprofen 844 colic 750 neonatal foal 226–7 septic arthritis and osteomyelitis 460 kidneys endocrine function, foal at parturition 70, 78–80 failure, aged horses 2756, 2757, 2758 infections, neonatal foal 652 interstitial abscesses 652 ultrasonography, foals 172, 248 see also renal function; renal problems KiSS-1 gene 1670 kisspeptin (KiSS), ovulation induction 1865 Klebsiella identification 2677, 2679, 2680
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
neonatal sepsis 253–4, 255 pneumonia 710 Klebsiella pneumoniae capsule/serotyping 1971 endometritis 1974, 1975, 2609 identification 1969, 1980, 1981, 1982–3 male external genitalia 1246, 1254 prevention and control 1963 screening of mares 1964, 1972–3 stallion semen 1371 transmission via AI 3016, 3023, 3029 venereal transmission 1254 Klinefelter’s syndrome see XXY syndrome knee see carpus Knowles cervical forceps 2561 Krey knife 2499 Kuhn’s crutch 2488, 2490, 2492 Kurasawa test, diagnosis of pregnancy 2258 kyphosis 667 labor see parturition laboratory resources, diagnostic 830–7 R see lactase, supplements Lactaid lactase deficiency, foal 392 postnatal production 264 supplements 194, 392, 844 enterally-fed neonate 268, 272 neonatal diarrhea 386, 388, 389 older foal 737 lactate blood causes of elevated 179, 180 critically ill neonatal foal 179–81 to guide fluid therapy 214 measurement 179, 180 neonatal foal 142, 175, 179, 187, 854 other body fluids 181 portable monitors 179, 180, 834–6 Sertoli cell secretion 884 lactated Ringer’s solution (LRS) 141, 206, 207, 231 lactating (wet) mares 2780 defined 1897 embryo recovery 2875 mammary gland ultrasonography 2162–3 nutrition 286–7, 2281–2, 2765–7 see also foaling mares; lactation lactation 2277–87 cessation 2282 inappropriate 2285 induction in non-pregnant mares 268, 1828, 2283, 2284 physiology 2229–30, 2277 premature 2284–5, 2480–1 death of one twin 2115 tetany 2282 see also mammary gland; milk; nurse mares lactational anestrus 1699–700, 1757 management 2637 ovarian ultrasonography 2123, 2124 lactic acid generation by sperm 1064, 1312 semen 1062 lactiferous ducts 1595 ultrasonography 2162–3 lactiferous sinuses obstructive lesions 2164 ultrasonography 2162–3 lactobacilli, vaginal 2644
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Index Lactobacillus acidophilus 272 Lactobacillus pentosus 737 Lactobacillus rhamnosus GG 844 lactoferrin endometrium 1934, 2589, 2608 epididymis 1009 β-lactoglobulin 2968 lactose intolerance 392, 737, 741 in mare’s milk 2280 in semen extenders 2968 lactose-EDTA semen extender 2965, 2966, 2969 Lamc2 gene 677 lamellar bodies 2444 lameness neonatal sepsis 201 pregnant mare 6, 8, 31 lamina propria, endometrial 1930 laminitis pituitary pars intermedia dysfunction 2791–2, 2794 postpartum 2531, 2532, 2533 prevention, retained placenta 2524–5 retained placenta 2524 treatment 2533 see also postpartum metritis pregnant mare 6, 8 fescue toxicosis 2422 induction of abortion 2323 management 31, 2748 Landis–Pappenheimer equation 179, 208 laparoscopes 1546, 1991 laparoscopic cannulas 1992, 1993 laparoscopic surgery castration 1549–50 cryptorchidectomy 1535–7, 1547–9 inguinal herniorrhaphy see inguinal herniorrhaphy, laparoscopic instrumentation 1546–7, 1991–2, 1993 mare 1991–9 ovariectomy 1991–9, 2568–71 stallion 1546–52 laparotomy, exploratory, neonatal foal 400 large bowel (large colon) disorders, foal 403–4 gaseous distention 244–5 obstruction, foals 244–5, 380 torsion, pregnant mares 2452 ultrasonography in foals 244–5 larval cyathostomiasis 297–8, 300 laryngeal adductor reflex 479 laryngeal hemiplegia 616–17 genetic basis 616–17 idiopathic (ILH) 616–17 pathological changes 616 presale endoscopy 797 larynx congenital abnormalities 613–14 cysts 614 malformations 613–14 paresis/paralysis 613 laser surgery castration 1553–6 laparoscopic ovariectomy 1996 periuterine adhesions 2672–3 persistent endometrial cups 2669 uterine cysts 2667–8 latanoprost 540 lateral recumbency neonatal resuscitation 128, 132 newborn foals 63
lateral suspensory ligament, udder 1595 latex agglutination test, foal serum IgG 348, 833 lavendar foal syndrome 558 Lawsonia intracellularis 739–40, 760 diagnostics 761–2 vaccines 762–3 see also proliferative enteropathy, equine laxatives meconium impaction 382–3 pica-induced diarrhea 391 lectins, sperm–oviductal interaction 1087–8 leg-man, breeding shed 1225–6 leg strap, breeding mare 1225–6 leiomyoma epididymis 1160 testicular tunic 1169 uterine 1947, 2150, 2151, 2671 leiomyosarcoma testicular 1168 uterine 1947–8 Lembert suture pattern, hysterotomy closure 2507–8 lens acquired abnormalities 549–50 congenital abnormalities 548–9 examination in foals 536 luxations 549–50 lenticonus 549 lentiglobus 549 leptin autumnal transition 1734 GnRH secretion and 1614–15 pregnant mare 2229–30 Leptospira, abortus testing 2340, 2341 Leptospira bratislava 3025 Leptospira interrogans serovar Bratislava 9 Leptospira pomona serovar kennewicki 9, 3025 leptospirosis, equine neonatal foal 411 pregnant mare 9, 32 transmission via AI 3025 lethal genes 2828 lethal white foal syndrome see overo lethal white foal syndrome leukocytes see white blood cells leukotriene B4 (LTB4 ), uterine release 2589 leukotrienes 1603–4 levamisole 844 levothyroxine (T4 ) 530 Leydig cells 869, 870, 871, 913, 1044–6 age-related changes 871, 914, 915 cryptorchid testes 1101 development 1015, 1017, 1044–6 drugs adversely affecting 1177–8 embryonic development 2214 functions 892–4, 1044–6 hormone production 892–4, 898 neoplasms 1167 numbers per testis 1045, 1046 paracrine–autocrine factors 999–1003 seasonal changes 872, 873, 1045, 1046 Sertoli cell interactions 885 smooth endoplasmic reticulum (SER) 1044 steroidogenesis 893–4 subfertility 1438
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testosterone secretion 892–3, 898–9 at puberty 903–4 thyroid hormone actions 998 LGR8 (Great) 1002–3 LH see luteinizing hormone libido (sexual interest and arousal), male inadequate 1401, 1407–10 pharmacological manipulation 1415–16 shuttle stallions 1185–6 see also slow breeders lidocaine 844 critically ill neonate 193, 194 diarrhea in older foal 737 epidural 225–6 foal 223, 225–6, 227 intrarectal, rectal palpation 1908 mare caudal tract surgery 2546 rectal tears in mare 2579 transdermal 226 ligament disorders, foal 469–74 LigaSureTM 1995, 1996 light horse breeds birthweight 281 fetal eye measurements 2077 semen evaluation 1367–8 stallion reproductive parameters 1368–76 lighting conditions, environmental circadian system 1670, 1671–2, 1772–3 nature of signals in horses 1773 see also photoperiod lighting programs, artificial mare 1711–12, 1773–7, 1788 daylength 1775–6 embryo recipients 2872 geriatric mares 2805 initiation dates 1774–5 lactational anestrus 2637 light in evening only 1776 lighting systems 1775 melatonin secretion 1675 night interruption (pulse lighting) 1776–7 oocyte recipients 2942 progestin therapy with 1779 quality and quantity of light 1773–4 seasonal anestrus 1698–9 stallion 1005, 1186, 1350 light microscopy acrosome evaluation 956, 1494, 2999 semen processing 1298–9 sperm morphological analysis 1297–8, 1299 sperm viability evaluation 1287, 1511 testicular biopsy specimens 1519 limbs reflex testing 479, 480 strength testing 479, 480 tone testing 479 see also forelimbs; hindlimbs lime sulfur 844 linear alopecia 565 linear keratosis 564–5, 566 linoleic acid 1603 linolenic acid 1603 lipid emulsions, parenteral nutrition 274, 275 lipidosis, hepatic, neonatal foal 413 lipid peroxidation (LPO), sperm membranes 940, 993, 1064, 3001, 3002 aged sperm 1077 lipid rafts, sperm capacitation 947
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lipids membrane, effects of cooling 1070–2, 1309–11 semen cryopreservation 2967–9 sperm content 1062 sperm plasma membrane 1055–6 lipoma myocardial 516 testicular 1168 lipoxygenase (LOX) 1604 liquid nitrogen checking levels 3007, 3008 cryotherapy 1138, 1140 embryo freezing 2887, 2899 safety issues 3008–9 shipping of embryos 2923 shipping of frozen semen 3010–11, 3031 sources 3008, 3009 sperm freezing 2963, 2974, 2978 static evaporation 3006–7 storage containers, frozen semen 3005–8 liquid shipping containers 2923, 3011 Lite SaltTM 211 liver biopsy 409 disease, foals 409–14 clinical signs 409, 410 congenital or inherited 415–16 diagnosis 409, 410 infectious 409–12 obstructive 415 toxic 414–15 weight loss 765 failure aged horses 2756, 2757, 2758 neonatal foal 409–14 round ligament 632 ultrasonography 248–9, 409, 410 local anesthesia castration 1557 cervical surgery 2560 laparoscopic ovariectomy 1993, 1994 laparoscopic surgery 1547 mare caudal tract surgery 2546 ovariectomy 2566, 2568 testicular biopsy 1527 lochia 2292, 2295 long bones fractures in foals 451–2 neonatal foal 434–6 ossification 436 long spinal reflexes 478–9 loop-colostomy, rectal tears 2581–3 loperamide 272, 844 lordosis 667 low-density lipoproteins (LDL), semen cryopreservation 2968 low-dose insemination see under artificial insemination low molecular weight heparin 843 lubricants artificial insemination 1275 artificial vagina 1200, 1269 effects on sperm motility 1294–5 obstetric 2483, 2484, 2486, 2499, 2514 rectal palpation 1908 lufenuron, fungal endometritis 2647, 2649 lumbosacral (LS) cerebrospinal fluid collection 479, 481–2 lumicrine pathways, epididymis 924 luminance 1774
lung immature 589 maturation, neonatal foal 75, 601 luprostiol 1797, 1799 luteal gland see corpus luteum luteal insufficiency 2057–8, 2131 ultrasonography 2131, 2132 luteal phase 1729 abnormalities 1758–61 estrogen profile 1631–2 FSH and LH profiles 1621–2 pharmacological termination 1729–30, 1798 prolonged 2057 causes 1759–61 ultrasonography 2132 shortened 2057–8 causes 1758–9 ultrasonography 2132 see also corpus luteum; diestrus luteinizing hormone (LH) 1619–27 β-subunit 1619, 1649 exogenous see equine luteinizing hormone mare age effects 2028, 2029 autumnal transition 1736, 1737, 1743–4 body condition and 2028 control of secretion 1623–4, 1783 cyclic patterns 1621–3 deficiency 1700 dopamine antagonists and 1791 double ovulations 2022, 2023 feedback control 1624–7, 1639 follicular development 1719–20 GnRH secretion patterns and 1611–12 hemorrhagic anovulatory follicles 2048 melatonin and 1675–6 multiple ovulations and 2021–2 ovarian steroidogenesis 1631, 1632 ovulatory surge 1621 periovulatory period 2023–4, 2025 pubertal changes 1690–2 seasonal anestrus 1698, 1705–6, 1784–5 vernal transition 1708–9, 1710–12, 2027 PGF2α -induced secretion 1799 physicochemical characteristics 1649, 1650 polymorphism and biological activity 1620 stallion 996–7 germ cell apoptosis and 916 infertility/subfertility 1436, 1437, 1439 opioid-mediated regulation 999 pubertal changes 902, 903–4, 1005–7, 1016 regulation of secretion 996–8 seasonal variations 873, 1003–4 secretion 897–8 spermatogenesis and 900, 1047–8 testicular actions 894 testicular descent 1102 testicular hormone secretion and 898–900, 996–7 structure 997, 1619–20 luteinizing hormone (LH) receptors 894, 1620–1
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
dominant follicles 2015 stallion subfertility 1438 luteinizing hormone-releasing hormone (LHRH) see gonadotropin-releasing hormone luteolysis autumnal transition 1737, 1739–40 cyclical PGF2α and 1638, 1642, 1643–4 prolactin and 1747 pharmacological induction 1729–30, 1798–9 pharmacological inhibition 1834, 1849 premature 2057–8 causes 1758–9 early pregnancy loss 2330 ultrasonography 2132 role of uterus 2200 ultrasonography 2059, 2060, 2062 luteotropic factors, mare 1736 lux 1774 lymphatic lacunae (LL), endometrium 1931, 1932, 1938 lymphocyte counts neonatal foal 173, 336 older foal 336 reference values 851 severe combined immunodeficiency 370 lymphocytes colostral transfer 336 developing foal 334–5, 336 immunization, donkey-in-horse pregnancies 2315–16, 2317 mucosal immune system 338 peripheral blood, culture 1462 uterine donkey-in-horse pregnancies 2314, 2317 endometrial biopsy 1931, 1932, 1935 endometrial cups 1654, 1656–8, 2193–4, 2197 interspecies embryo transfers 2309, 2310 uterine cytology 1924 see also B-lymphocytes; T-lymphocytes lymphoma 516 lymphosarcoma epididymis 1159, 1169 ovarian 2710, 2714 prepuce 1169 lymphostasis, endometrial 1945, 1946 Lyphomune 344 lysine requirements 786 breeding stallion 1230 broodmares 2762, 2763, 2766 lysophosphatidylcholine 950 lysozyme 2738 lysyl-oxidase 788 M199 oocyte maturation medium 2933 MacConkey agar 1979, 1980, 2676 maceration, fetal see fetal maceration macrolide antibiotics neonatal foal 254, 260–1 pneumonia in older foal 716 R. equi pneumonia 699–700, 740 macrophages 333 in semen 1305 uterine cytology 1925 magnesium (Mg2+ ) mare’s milk 2280, 2281 pre-foaling mammary secretion 2263
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Index requirements, stallion 1230, 1234–5 serum 173, 853 magnesium hydroxide 844 see also antacids magnesium sulfate (MgSO4 ) 844 hypocalcemia 212 hypoxic-ischemic encephalopathy 148, 149, 150, 486–7 intrauterine infusion 2634 neonatal foal 195, 232 tetanus 493 maiden barren mare 1897 maiden foaling mare 1897 maiden mares 1897, 2780 bonding with foal 64 dystocia 2487 estrous cycle abnormalities 1758 estrus detection 1718 fertility problems 2622 foal growth rates 281–2 foal rejection 118, 120 insemination technique 1204 insemination with frozen semen 2990 mammary secretion electrolytes 2264 nursing problems 167 old see old maiden mares ovulation induction 1860 preventive medicine 2749 reproductive efficiency 2781 stallion fertility evaluation 1429, 1430, 1431 test jumping 1719 see also old maiden mares maintenance of pregnancy 2455–75 antibiotics 2472–4 causes of failure 2455–61 endocrinology 2461–7 geriatric mares 2813 hormone treatment 2469–72 immune stimulation 2474–5 management issues 2467–9 major histocompatibility complex (MHC) class I antigens 333 endometrial cups 1658 placental expression 89, 1655 class II antigens endometrial cells 2590 foal immune cells 333, 336 malabsorption/maldigestion, aged horses 2756–7 mal du coit see dourine male external genitalia embryonic development 2215 examination 1458–60 prior to endoscopy 1453–3 prior to ultrasonography 1469 restraint 1458, 1469 miniature horses 1378–9 opportunistic bacterial invaders 1254 screening cultures 1246 ultrasonography 1469–80 see also penis; prepuce; scrotum; testes male genital tract developmental abnormalities 1109–11 embryonic development 1015–16, 2215 endoscopy see under endoscopy functional anatomy 867–78 neoplasia 1166–9 trauma, hemospermia 1121–2, 1123–4 ultrasonography see under ultrasonography zebras 2861 male internal genitalia endoscopic examination 1448–55
infections suboptimal fertility 1245–6 venereal transmission 3024 see also accessory sex glands male pseudohermaphrodite 1111, 1136, 1764, 1765, 2218 male reproductive function 881–905 endocrine–paracrine–autocrine regulation 996–1009 endocrinology 897–900 infantile stage 902, 904 prepubertal stage 902, 903, 1015 puberty see puberty, male seasonal variations 1003–5, 1344–50 mallenders 566, 567 malnutrition, neonatal foal 263–4 maltase 264 mamillary bodies 1582, 1583 mammary gland (udder) 2162–4 abscesses 2164, 2287, 2740 anatomy 1595–6, 2162, 2163, 2277 blood supply 1596 blood vessels 2163 edema 2286 enlargement, causes 2286–7 failure of prepartum development 2282 fetal sex determination 2086, 2088, 2091, 2097 hormonal control of lactogenesis 2229 innervation 1596 lymphatic drainage 1596 neoplasia 2287, 2742–4 premature development 2283–4 prepartum changes 2263–4, 2270, 2480 tenderness, foal rejection 119 ultrasonography 2162–4 ventral rupture involvement 2447–8 see also lactation mammary secretion electrolytes, pre-foaling (PFMS) 2280, 2733–6 assay methods 830–1, 2734 factors affecting 2264 feto-placental monitoring 19–20 induction of parturition 1833, 2263–4 jennies 2270 Miniature ponies 2841–2 predicting parturition 2270, 2733–4, 2735–6 sample collection 2734 manchette 917, 918, 930, 1028, 1036 mandibular brachygnathia 667 mandibular fractures, foals 453–4 mandibular snares, obstetric 2486, 2491 mane and tail dystrophy, Appaloosas 555, 568, 569 manganese 789 requirements breeding stallion 1230 broodmare 2762, 2763, 2766 neonatal foal 265 mannitol 844 hypoxic-ischemic encephalopathy 149, 150, 487 neonatal foal 195, 232 marbles, intrauterine see intrauterine glass marbles/devices mare age early embryonic loss and 2327 fertility effects 1431, 1701, 1897, 2990 ovulation induction and 1806, 1866 placental development and 2237–8 reproductive efficiency and 2781
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seasonal anestrus and 1734–5 twin pregnancy and 2351 see also geriatric mares; older mares behavior see under behavior book of see book of mares breeding records, prior 1897–8 contraception 1889–92 cytogenetics 1951–61 embryo donors 2872–6, 2878–9 embryo recipients 2871–2, 2876–8 endoscopy 1940–9 exotic equid embryo transfer 2311–14 foal rejection 117, 118–20 harem bands 1385 interspecies matings 2303 mount see mount mares oocyte donors 2931, 2936 oocyte recipients 2941–3 opportunity to become pregnant 1431 ovariectomized see ovariectomized mares protective cape 1225, 1270 puberty 1689–94 reproductive history 1897–9 reproductive tract see female reproductive tract size, placental development and 2237 stallion training 1394–5 novice stallions 1399–401 slow-starting novices 1408–9 subfertility/infertility see problem-breeding mare Taylorella equigenitalis carriers 3017–18, 3019 teaser see teaser mares zebra 2852–9 see also broodmare mare fertility after cesarean section 2509 age effects 1431, 1701, 1897, 2990 breeding on foal heat 2142, 2293, 2297 timing of first ovulation and 2297–9 caudal reproductive tract surgery and 2545 dystocia management and 2484–5, 2509, 2516 endometrial biopsy and 1933–4 fetotomy and 2497, 2498 intrauterine fluid effects 2145–6 nutrition and 2747 pre-ovulatory follicle vascularity and 2048–51 relevant history 1897–9 retained placenta and 2526, 2527 uterine cysts and 2148 see also problem-breeding mare mare–foal bonding 63, 117–18 factors disrupting 118 orphan foal to surrogate dam 273 mare pregnancy protein-1 (MMP-1) 2258 mare reproductive loss syndrome (MRLS) 2410–16, 2461 clinical description 2410–12 diagnosis 2413 epidemiology 2414–15 fibrinous pericarditis 516, 2412, 2413–14 high-risk pregnancy 7, 13–14 management 35 ocular disease 429, 2412 pathogenesis 2415 pathological findings 2412–13 prevention 2415–16 surviving neonates 2411–12 treatment 2413–14
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mare reproductive status definitions 1429–30, 1897, 2780 insemination with frozen semen 2990 reproductive efficiency and 2781 twin pregnancy and 2351 Mare’s MatchTM milk replacer 269, 845 R Mare’s Milk Plus milk replacer 269, 845 mast cell neoplasm, testicular 1168 mastitis 2285–6, 2738–41 avocado toxicity-associated 2738, 2740 causes 2738, 2739 diagnosis 2738–40 prognosis 2741 treatment 2740 ultrasonography 2163–4 mastocytosis, congenital equine cutaneous 561 masturbation 1389, 1390, 1425 maternal antibody interference 307–8, 338 maternal behavior abnormal 118 in natural conditions 1388 normal 117–18, 2271 see also foal rejection maternally derived antibody (MDA) transfer see passive transfer of maternal antibodies maternal recognition of pregnancy (MRP) 2200–7, 2461–2 changes in gene expression 2203–4 conceptus mobility 2202–3 critical period 2461–2 embryo transfer recipients 2878 estrogens and 1633, 2201–2 oxytocin–PGF2α relationship 1644, 1680, 2204–7 PGF2α 1644–5, 2200–2 role of uterus 2200 uterine blood flow 2207 uterine protein secretion 2202 mating see copulation mating patterns, zebras 2851–2 matrix metalloproteinases (MMPs), cervical ripening 2728 Max-Semen ExpressTM 1334 mean arterial pressure (MAP) 139 autoregulation 505 neonatal foal 140, 169 anesthesia 224 monitoring in critical care 186 normal values 186, 187 see also arterial blood pressure mean corpuscular hemoglobin concentration (MCHC) neonatal foal 173, 420 reference values 851 mean corpuscular volume (MCV) neonatal foal 173, 419, 420 reference values 851 Meckel diverticulum 404, 643, 755, 756 meclofenamic acid, embryo recipient mares 2335, 2872 meconium 379 aspiration 588–9, 592, 597–8 clinical signs 379, 592 diagnostic tests 588–9, 594 treatment 589, 595, 598 see also aspiration pneumonia impaction 379–83 diagnosis 170, 380–1 differential diagnosis 381, 625, 626 history and clinical signs 379–80 medical management 193, 381–3
percutaneous intervention 383 prophylaxis 383 surgical management 402 ultrasonography 244, 246 passage newborn foal 65, 67, 379 in utero 51, 379 medial suspensory ligament, udder 1595 median eminence 1583, 1584 medroxyprogesterone acetate 1813 estrus suppression 1847 maintenance of pregnancy 2121, 2471 transitional mares 1780 megalocornea 545 megesterol acetate, maintenance of pregnancy 2471 meiosis abnormalities in aged mares 2329 arrest, equine hybrids 2304 spermatocyte 883, 914, 1026, 1029, 1032–4 meiotic chromosomes, cytogenetic analysis 1467 α-melanocyte-stimulating hormone (α-MSH) 1666 melanoma mammary gland 2744 ovarian metastatic 2710 penis and prepuce 1137, 1169 melanopsin 1672, 1772 melanotic nevus/melanocytoma/congenital melanoma 559–61, 563 melanotropes 1666, 2790 melatonin 1582, 1669–77 anatomic pathway 1671–2 annual reproductive rhythm and 1677, 1704, 1782–3 autumnal transition 1746, 1747–8 biological actions 1669–70 biosynthesis 1671 circadian rhythms and 1670, 1671 exogenous autumnal transition 1747–8 reproductive effects 1675–7 shuttle stallions 1350 male reproductive function 996–7, 1004 manipulation of secretion 1675, 1677 reproductive effects 1675–7 seasonal anestrus and 1698 secretory patterns 1672–5 melatonin receptors (MT1 and MT2) 1669–70 melengestrol acetate (MGA) 1813, 1847 meloxicam, foals 227, 744–5 membranes, cell diffusion of solutes across 2959–60 effects of cooling 1070–2, 1309–11 fluid mosaic model 1070–1, 1309 see also lipid peroxidation; sperm plasma membrane menace response, foal 171, 535 meningitis, bacterial 160, 202, 487 mercury, for ICSI 2949 merocyanine 540, sperm evaluation 1288, 1502 mesenteric lymphadenopathy, foal 243, 244 mesenteric lymph nodes 334–5, 338 mesentery, entrapment of uterus within 2443–4 mesocolon, agenesis of 407 mesodiverticular band 404, 755, 756
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
mesoductus deferens 1145 mesometrium 1586, 1588, 1589, 1590 hematoma 2450, 2672 peripartum vascular rupture 2449 during pregnancy 1591 mesonephric ducts (wolffian ducts) 900, 901, 2211, 2213 cystic remnants 2700, 2701, 2703 cysts 2692, 2701 male genital tract development 2215 regression in female 2215 segmental aplasia 1111, 2607 mesonephric kidney 1099 mesonephric tubules, cystic remnants 1586 mesonephros 2213 mesorchium 901, 902, 1145 laparoscopic castration 1550 laparoscopic cryptorchidectomy 1536 mesosalpinx 1586, 1588, 1993 mesothelioma pericardial 516 peritesticular 1168–9 mesovarium 1586, 1588 transrectal palpation 1910 metabolic acidosis, neonatal foal 206, 208 metabolic bone disease see developmental orthopedic disease metabolic disorders congenital 668–9 problem-breeding mare 2636–7 metabolic rate, peripartum foal 74–5 metabolic syndrome, equine 2806 see also insulin resistance metacarpophalangeal joint (fore fetlock joint) flexural deformity 441, 775 evaluation 441–2 prognosis 445 treatment 442–3, 444–5 presale radiographs 802–4 interpretation 810–14 see also fetlock joint metacentric chromosomes 1952 metanephros 2213 metaphase 1952 metaphyseal equivalent site 435–6 metaphyseal growth plates see physes metaphysis 434 metastatic tumors, ovarian 2710 methanol embryo cryopreservation 2896 toxicity to embryos 2917 methicillin, neonatal foal 257 methicillin-resistant Staphylococcus aureus (MRSA) 1979 methionine 844 methocarbamol 845 methyl formamide, semen cryopreservation 2969 methylprednisolone 845 acute respiratory distress syndrome 583–5 neonatal foal 488 metoclopramide 845 agalactia 2283 gastrointestinal dysmotility 271, 686 metritis postpartum see postpartum metritis vs. pyometra 2652 see also endometritis; uterine inflammation metronidazole 845 foal diarrhea 386, 388, 736
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Index neonatal foal 261 dose 194, 254 prior to transport 232 tetanus 493 Tyzzer’s disease 412 umbilical remnant infections 251 mezlocillin 257–8 MHC see major histocompatibility complex miconazole 845 fungal endometritis 2636, 2647 shampoo 845 topical ophthalmic 541 microaerophilic pathogens identification 1969–70 uterine and clitoral cultures 1968 microarrays, gene 2830 microbiological laboratory techniques 2674–81 antibiotic sensitivity testing 1971–2, 1986, 2675–6 biochemical tests 1970–1, 1980 capsule/serotyping 1971 catalase, oxidase and indole tests 2676 chromogenic agars 1980–7 commercial identification kits 1970–1, 2677 culture methods 1968, 2675–6 differential media 2676 identification methods 1968–71, 2676–81 PCR tests 1971 plating a culture 2675 sample handling 1967 specialized facilities 1967–72 subcultures 1970 see also clitoral cultures; uterine cultures microcaruncles 88, 101 postpartum resorption 2291–2, 2294 microcornea 545 microcotyledonary villi development 86, 87, 88, 2198 examination 92, 101, 102, 103, 106 vascular network 2239–40 microcotyledons 86–7, 2235 areas lacking 2198, 2235 formation 2195–6, 2197, 2198 surface area per unit volume (Sv ) 2237, 2238, 2239 microencephaly 669 β 2 microglobulin (β 2 M) 2178, 2179 microphakia 540, 549 microphthalmia 46, 537–8 microplacentome 86, 101 microsatellite DNA markers 2821–4 parentage analysis 2823–4 parentage testing 2822, 2823 standardization 2823–4 microspheres, oviductal patency testing 1989, 1990, 2694 microtubular mass defects, sperm 1302, 1303 microtubules, axonemal 931–4, 1060, 1061 mechanism of bending 937, 1065–6 midazolam 845 neonatal seizures 193, 194, 234, 486 perinatal asphyxia syndrome 148, 149 sedation of neonate 223 middle artery of penis 877 middle piece (midpiece), sperm 931, 932, 934, 1028, 1059–60, 1061
abnormalities 1301 milk 1596 aspiration 589, 592–3 diagnostic tests 593–4 treatment 595 see also aspiration pneumonia composition 265, 268, 269, 2280, 2280 dietary influences 2766 consumption by foal 265 daily production 2280 enteral feeding 268 hand-milking 270 pre-foaling electrolytes see mammary secretion electrolytes, pre-foaling prepartum dripping 2284–5, 2480–1 reflux 592 regurgitation 576, 589, 592, 593 secretion 2277 see also colostrum; cow’s milk; goat’s milk; lactation milk-based semen extenders 1200–1 cooled semen 1074, 1337, 1341 frozen semen 2967–8 historical development 1264–5 milk let-down physiology 1680, 2277 poor 63–4 milk replacers 269–70, 845, 2280–1 miller’s disease 1234 mineralocorticoids 72, 1665 see also aldosterone mineral oil 845 meconium impaction 382 pica-induced diarrhea 391 rectal palpation 1908 minerals 787–9 developmental orthopedic disease and 778, 787, 788–9 equine milk 2280, 2281 requirements breeding stallion 1230 broodmare 2762, 2763, 2764, 2766 neonatal foal 265–6 supplements, growing horse 288 Miniature horses/ponies 2839–44 Miniature mares 2839–44 abortion 2841, 2842 breeding management 2839–40 double ovulations 2022 estrous cycle 2839 parturition 2841–3 postpartum complications 2843–4 pregnancy 2841 pregnancy diagnosis 2840–1 pre-ovulatory follicle diameter 2025–7, 2839 Miniature stallions 1377–80 breeding management 2839–40 breeding soundness criteria 1379–80 external genitalia 1378–9 internal genitalia 1379 semen collection 1377–8 semen evaluation 1378 minimal sagittal diameter (MSD) ratio, cervical vertebral compressive myelopathy 729–30 minimum alveolar concentration (MAC) 223, 224 minimum essential medium (MEM) 2881 minimum inhibitory concentration (MIC) 256, 257 mismated mares, induction of abortion 2322
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misoprostol 845 cervical ripening 2726, 2729, 2811 gastrointestinal protection 396, 744 induction of abortion 2321 Missouri model artificial vagina 1263–4, 1269 modified 1270 mitochondria oocyte, aged mares 2329 replacement therapy 3054 sperm 918, 933, 934–5, 1059, 1060 abnormalities 1301 analysis of function 957, 1289, 1513, 3000–1 corkscrew defect 1301 effects of cooling 1073 function 1293, 3000–1 generation of reactive oxygen species 991–2 MitotrackerTM (MITO) 957, 1289 mitral dysplasia 508 mixed sex cord-stromal tumors 1168 mixed sex peer-groups (transitional bands) 1385, 1386 M-mode ultrasound 1918 R 1418 Modipher EQ molecular cytogenetics 1464, 1467, 1954–5 monkey mouth 667 monocyte counts 173, 851 monocytic ehrlichiosis, equine see Potomac horse fever monorchidism 1537 congenital 1109, 1163 monosaccharides, metabolism in sperm 938 monosomy X see X chromosome monosomy Morgan weanlings, hyperammonemia and hepatopathy 413–14, 732 morphine 845 epidural 226, 845 foal 222, 226 intra-articular 226 pregnant mare 57 morula 2172–3, 2174 glucose metabolism 2175 identification 2876 see also embryo(s) mosaicism 1952, 2217 motilin 77, 78 motor neuron disease, equine (EMND) 732 mountain zebra (Equus zebra) 2851 foal 2862 interspecies hybrids 2302 mare behavior 2852 estrous cycle 2856 gestation 2856–7 postpartum estrus 2859 puberty and breeding life 2853–4 reproductive organs 2855 mating patterns 2851–2 seasonality of breeding 2853 stallion behavior 2859 puberty and breeding life 2860 reproductive organs 2861 seasonality 2860 weanling 2862 yearling 2862–3 see also Cape mountain zebra; Hartmann’s zebra
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mount mares 1270–1 slow-starting novice stallions 1409 stallion training 1394–5, 1399 mouse test, diagnosis of pregnancy 2254 mouth examination, neonatal foal 170 mouth-to-nose ventilation, foal 131 moxidectin ascarids 295, 759 bots 293–4 cutaneous habronemiasis 1140 parasite control programs 113, 300 stomach worms 294 strongyles 298, 300, 742, 760 tapeworms 299 mucin test, diagnosis of pregnancy 2258 mucociliary clearance, uterus 2591 mucometra 2724, 2726 Mucor spp. 711, 2643 mucosal-associated lymphoid tissue (MALT) 334–5 mucosal immunity 338 mucous membranes, neonatal foal 499, 511–12 critical care 167–8 respiratory disorders 577 sepsis 154, 155 shock 138–9 mucus, uterine cytological smears 1924, 1925 Mueller-Hinton plates 2676, 2680, 2681 mule 2302, 2303, 2835 cloned 2924, 2925 Dictyocaulus arnfieldi carriage 711 embryonic, transfer to donkeys 2310–11 fertility 2302, 2304–5 horse or donkey pregnancies in 2307–9, 2310 neonatal isoerythrolysis 353, 357, 420 strabismus 539 vaginal prolapse 2836–7 mullerian ducts see paramesonephric ducts mullerian inhibiting substance (MIH) 2215 Multistix test, urospermia 1125 Multi Thermal GradientTM semen freezing apparatus 2978–9 mummification, fetal see fetal mummification murmurs, cardiac 500, 511 grading 512 neonatal foal 66, 168–9, 500, 512 patent ductus arteriosus 500, 508, 512 pathological 500 shock 139 ventricular septal defect 507–8 muscle disorders 463–7 hereditary 463–5 muscle strength, testing 479, 480 muscle tone, testing 479 muscle weakness neonatal foal 171, 484–5 older foal 728 musculoskeletal system congenital abnormalities 666–7 fetal development 433–9 neonatal foal 67, 172 mustard plants 529–30 mutation, fetal 2479, 2486–93 muzzles, aggressive stallions 1398 Mycobacterium cell wall extract chronic endometritis 2613 foal pneumonia 717
maintenance of pregnancy 2474–5 problem-breeding mares 2631 Mycoplasma equigenitalium 1256 Mycoplasma felis 711 Mycoplasma spp. pneumonia 711 venereal transmission 1256, 3025, 3030 mycotic infections see fungal infections mycotoxin-induced hepatic disease 412 myeloencephalopathy equine degenerative (EDM) 489–90, 732 equine herpesvirus see equine herpesvirus myeloencephalopathy myelography 730 myocardial abscessation 515 myocardial disease, acquired 514, 515–16 myocardial hypoxia/ischemia 515 myocardial infarction 515 myocardial neoplasia 516 myocardial trauma 515 myocardin 2728 myocarditis 515 myoid cells (myofibroblasts), seminiferous tubules 869, 912 myometrial contractions see uterine contractions/contractility myometrium 1589, 1590, 1930 muscle cell defects 2603 myopathy, post-anesthetic 2509 myotonia 465–6 myristic acid, sperm content 1062 NAC see acetylcysteine NADPH oxidase sperm-specific (NOX5) 940, 991, 992 uterine environment 940 nafcillin 257 naloxone 845 foal 222 GnRH secretion in mares and 1614 oocyte vitrification 2955 perinatal asphyxia syndrome 150 periparturient hemorrhage 2518 nape posture, fetus 2490–1 naproxen 845 narcolepsy/cataplexy 490 nares examination 576 regurgitation of milk 576, 589, 592, 593 nasal discharge neonatal foal 576 older foal 712 recurrent airway obstruction 618 strangles 705 nasal washes, strangles 706 nasogastric intubation, colicky foal 751 nasogastric reflux, critically ill neonate 170–1 nasogastric tube feeding, neonatal foal 192, 193, 270–1 complications 272–3 prior to/during transport 235 nasolacrimal system acquired disorders 544 atresia and aplasia 542–3 nasopharyngeal cysts 611–12 see also pharyngeal cysts nasopharyngeal irritation, enterally-fed foal 272 nasotracheal intubation, neonatal foal
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anesthesia 221–2 birth resuscitation 129–31 natamycin fungal endometritis 2647 topical 541, 845 National Animal Health Monitoring Survey (NAHMS) 2798 National Human Genome Research Institute (NHGRI) 2827 native phosphocaseinate (NPPC) 2968 natural service 1208–27 breeding frequency 1223, 1390 breeding shed team 1225–6 imported mares 1227 mare preparation 1224, 1225 mare restraint/handling 1225–6, 1227 Miniature ponies 2839–40 ovulation induction 1804, 1859–60 practical management 1222–7 problem mares 2144–5 receiving area 1223 reproductive efficiency 2781–2, 2784–6 stallion handling 1226–7 stallion preparation 1225 stallion training 1394–5, 1399–401 suboptimal stallion fertility 1240–8 teasing and identification area 1223–4 venereal disease transmission 3015 vs. AI 1198–9 see also copulation nebulization therapy neonatal foal 189–90, 607 pneumonia in older foal 716–17 R. equi pneumonia 700 neck, fetal amputation 2500, 2501 malpostures 2489–91 fetotomy 2500–1 lateral reflexion 2490 ventral deviation 2490–1 normal posture 2271, 2482 necropsy, dead fetus 2340–1 necrotizing enterocolitis (NEC) 392 predisposing factors 267 ultrasonography 245–6 negative-reinforcement methods, stallion training 1391, 1401 neomycin intrauterine infusion 2632 neonatal foal 256 topical ophthalmic 541 Tyzzer’s disease 412 neonatal encephalopathy (NE) (hypoxic-ischemic encephalopathy; HIE) 147–52, 483–7 bladder rupture 625–6 clinical signs 147, 485, 486, 487 diagnosis 485 diarrhea 392 differential diagnosis 160 management 148–52, 193, 195, 213 pathophysiology 147–8, 483 preparation for transport 232–3 prognosis 152 treatment 148–52, 485–7 see also perinatal asphyxia syndrome neonatal equine respiratory distress syndrome (NERDS) 125, 583, 584, 589 neonatal foal 63–7, 2271 behavior 63–4, 117–18 complications 18
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Index critically ill see critically ill neonatal foal definition 821 dystocia survival rates 2509–10 endocrine adaptations 69–80 equine herpesvirus infection see under equine herpesvirus(es) fluid therapy 205–14 galactorrhea 2285 gastroduodenal ulceration 394–7 immune system 67, 331–8 infanticide, Asian wild horse 2849–50 infections 158–60, 198–203 laboratory values 172–5 maternal fescue toxicosis and 72, 530, 2421–2 maturity 121–2 nutrient requirements 265–7 nutritional physiology 264–5 pharmacology 253 physical examination 65, 113, 167–72 physiological parameters 65, 187 placental evaluation 93–4 preparation for transport 230–6 rejection by dam see foal rejection respiratory support 189–90, 215–19 resuscitation see neonatal resuscitation routine management 65, 112–13 sedation, anesthesia and analgesia 221–7 sepsis see sepsis, neonatal shock 136–45 surgical gastrointestinal conditions 399–404 thermoregulation 65–6, 75 transfusion techniques 238–41 transition to extrauterine life 65–7 zebras 2861–2 neonatal intensive care unit (NICU) cost capturing 826–9 development 821–9 equipment 824, 825 facility 821–3 infection control strategies 824–6, 827 preparation for transport to 230–6 prognosis/outcomes 160–1 referral and transport 196 resuscitation kits 828, 828, 829 supplies 835–6, 837 technical staff 823–4 see also critically ill neonatal foal neonatal isoerythrolysis (NI) 353–60 anemia 354, 419–20 clinical signs 138–9, 354, 421–2 complications 357 diagnosis 13, 354, 423 high-risk pregnancy 7, 12–13, 35, 111 immunology and pathogenesis 353–4 liver disease/failure after 410, 412–13 outcome 357, 425 prevention strategies 357–60, 2750 screens 358–60, 831, 2824 treatment 112, 356–7, 424–5 blood transfusion 142, 239, 356–7 prior to transport 231–2 neonatal maladjustment syndrome (NMS) see neonatal encephalopathy neonatal resuscitation 128–35 birth 128–35 care following 135 cessation 135 drugs used 130, 132–3 dystocia 2512 equipment 128, 129
monitoring 134–5 NICU kits 828, 828, 829 in shock 140–4 neonatal vasogenic nephropathy (NVN) 649, 650 management 656–7 Neopor box, semen transport 1334 Neorickettsia risticii 10, 319 Neospora caninum 730 Neospora hughesi 730, 731 neostigmine 845 Neotyphodium coenophialum (formerly Acremonium coenophialum) 530, 2418, 2765 toxicity of ergot alkaloids 2419–23 see also fescue toxicosis nephrectomy, ectopic ureter 629 nephrotoxins, neonatal foal 652 nerve injuries, peripheral 489 nervousness, signs of 1407–8 nervous system trauma 487–9 net energy (NE) 783 Netherlands, draft horse, Friesian and Warmblood stallions 1362–6 N-ethylmaleimide-sensitive factor (NSF) 950 neuroendocrine action 1601 neurogenic shock, neonatal foal 136, 137 neurohypophysis (posterior pituitary) 897, 1583, 1584 neurological development, normal 477 neurological disorders neonatal foal 483–93 older foal 728–32 urospermia 1124–5 neurological evaluation, neonatal foal 477–82 neurological examination neonatal foal 66, 171, 477–9 urospermia 1126 neuronal cell death, perinatal asphyxia syndrome 148 neuropeptide Y (NPY) 1615 neuroprotectants, neonatal foal 149, 150, 195 prior to/during transport 232–3 neurotransmitters, regulating GnRH secretion 1613–14 neutropenia, postpartum metritis 2530, 2531, 2532 neutrophil counts neonatal foal 173 premature or dysmature foal 124 reference values 851 septic foal 155, 199 neutrophil:lymphocyte (N:L) ratio, neonatal foal 124, 126, 174 neutrophils (polymorphonuclear leukocytes; PMNs) developing foal 332, 334 dysfunction 366 Pelger–Hu¨et anomaly 366 semen dismount samples 1245–6 microscopic appearance 1121 seminal vesiculitis 1115 sperm motility and 940 see also pyospermia uterine 2608 cytological smears 1923–4, 1926, 1964 elimination of sperm 1093, 1095, 2614 endometrial histology 1932
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endometritis 2611–12 functional properties 2590 post-insemination influx 1094–6, 2656 postpartum 2291, 2294 newborn foal see neonatal foal New Forest ponies, first pubertal estrus 1692 New Zealand cooled transported semen programs 1323 import/export of stallions 1187 nexin links 933, 934, 937, 1060 nick translation (NT) assay 956 NICU see neonatal intensive care unit nitazoxanide (NZT) equine protozoal myeloencephalitis 731 neonatal diarrhea 386, 388, 389, 390 nitrate exposure, congenital hypothyroidism 529–30 nitric oxide (NO) delayed uterine clearance and 2601–2, 2608–9 neonatal renal function and 647 therapy neonatal foal 195, 217 persistent pulmonary hypertension 590, 597 triggering capacitation 947 uterine secretions 2589–90 nitrite, ejaculate, urospermia 1125 nitrofurazone ointment, Taylorella equigenitalis carriers 1251, 2406 nitroglycerin 845 N-methyl-D-aspartate (NMDA) receptors 148 nocardioform placentitis 9, 10, 2359, 2459 placental examination 2273, 2274 ultrasonography 2361 nocturnal rhythms 1771 Noggin 776 non-breeding season (winter) anestrus with no ovarian activity 1697–9 stallion nutrient requirements 1229, 1233–4 see also transitional mares non-fat dried skim-milk glucose (NFDSMG) 1337, 1338, 1341, 2967 see also KenneyTM semen extender non-fat dry milk solids (NFDMS) 1200–1 non-shivering thermogenesis (NST) 75 non-steroidal anti-inflammatory drugs (NSAIDs) foal 226–7 diarrhea 736–7 gastrointestinal toxicity 683, 686–7, 744–5 hepatotoxicity 412 infective endocarditis 515 nephrotoxicity 652 septic arthritis and osteomyelitis 460 mare breeding-induced endometritis 2812 placentitis 33, 2363, 2365–6 pregnancy 26, 2335 see also flunixin meglamine; ketoprofen non-thyroidal illness syndrome 528 norandrostenedione, yolk sac fluid 2178
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norepinephrine (noradrenaline) exogenous 845 neonatal anesthesia 224 neonatal critical care 143, 192, 195 premature or dysmature foals 125 septic foal 203 urospermia 1127 foal adaptation to parturition 70, 72–3 melatonin synthesis and 1671, 1672–3 norethisterone (norethindrone), contraception in mares 1889 norgestomet 1813 estrus suppression 1847 maintenance of pregnancy 2471 Normosol-M with 5% dextrose 206, 207 Normosol-R 141, 206, 207, 231 North America, semen extenders for cooled semen 1341–2 Norwegian Fjord horses, bilateral hind leg arthrogryposis 667 nose, congenital abnormalities 609–11 nosocomial infections (healthcare-associated infections; HAI) control strategies see infection control NICU 824 perinatal asphyxia syndrome 151 ventilated foal 219 novice breeding stallion 1396–401, 1397–8 breeding environment 1397 breeding shed training 1398–401 let-down period 1396 slow starting 1407–10 training to lead, stand and back 1397–8 NOX5 (sperm-specific NADPH oxidase) 940, 991, 992 NSAIDs see non-steroidal anti-inflammatory drugs nuclear envelope redundant (RNE) 928, 951–2, 1059 sperm 928, 933, 1035 nuclear factor kappa B (NF-κB) 1606 nuclear transfer (cloning) 2924–7, 3053 clinical feasibility 2926–7, 3052 clone–donor similarity 2926 ethical issues 3052 health of pregnancies/foals 2925 progeny of cloned animals 2926 technique 2924–5 R SP-100TM 1281–2 NucleoCounter nucleus, spermatozoa 927–8, 932, 933, 1057 abnormalities 1301, 1302, 1304, 1305 aged sperm 1077 nurse mares 268, 2281 bonding foals to 273 induction of lactation 268, 2283, 2284 via induction of abortion 2322 nursing see suckling/nursing nutrients placental production 89 placental transport 89 requirements breeding stallion 1229–38 broodmare 264, 287 healthy neonatal foal 265–6 lactating mares 2765–6 non-pregnant mares 2760–1, 2762 pregnant mares 2763, 2764, 2765 sick neonate 266–7 see also diet; specific nutrients nutrition broodmare 111–12, 264, 2747–8, 2760–7
autumnal transition and 1734–5 early embryonic loss 2332–3 foal birthweight and 281–2, 286–7 high-risk pregnancy 28–9, 53 for lactation 286–7, 2281–2, 2765–7 non-pregnant mares 2760–2 optimizing foal growth 286–7 placental development and 2239 pregnancy 2763–5 reproductive efficiency and 2760 seasonal anovulation and 1698, 1699, 2747 embryo 2187, 2188, 2190 fetal, hormonal control 2229–30 geriatric horse 2755–9 growing horse 282, 286, 287–8 developmental orthopedic disease and 287–8, 778, 782–91 gastroduodenal ulceration and 686 neonatal foal 263–78 critical care 192–3 normal physiology 264–5 prior to/during transport 236 shock 144 pituitary pars intermedia dysfunction 2794 stallion 1221, 1228, 1229–39 see also diet; enteral feeding; feeding; parenteral nutrition nutritional myodegeneration (NMD) 466–7 dysphagia 593, 595 myocardial involvement 515–16 nutritional secondary hyperparathyroidism 1234 nutritional supplements stallion behavior problems 1411, 1417 subfertile stallions 1442 see also under specific nutrients nyctohemeral rhythms 1771 nystagmus 478 nystatin, fungal endometritis 2636, 2647 NZ Semen Shipper 1333, 1334 observational studies 2798 obstetrical chains 2486 obstetrical equipment 2483 obstetrics, equine 2479–94 obturator artery 877 occipitoatlantoaxial malformation (OAAM) 491, 492 OCD see osteochondritis dissecans Ochragaster lunifer 2415 ocular disorders see ophthalmic disorders ocular lubrication, sedated/anesthetized foal 222 odorant receptors, sperm 952 oflaxacin 845 older foal anesthesia 221 bacterial respiratory diseases 688–93, 695–701, 704–9, 710–17 colic 749–57 diarrhea 734–45 gastroduodenal ulceration 683–7 neurological disorders 728–32 weight loss 758–65 older mares chronic endometritis 2631–2 endometrial biopsy 1929 foal rejection 118 folliculogenesis 2028–9 myometrial defects 2603 oviductal blockage 1989
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periparturient hemorrhage 2449, 2517 post-mating endometritis 2627–8, 2809–13 uterine cysts 2148, 2331, 2806–7 vaginal varicose veins 2452 vulvar angulation 1578 see also geriatric mares; mare, age old horses see geriatric horses old maiden mares insemination with frozen semen 2990 post-breeding endometritis 2656 reproductive efficiency 2781 old maiden mare syndrome 2656 old mares see geriatric mares old stallions see geriatric stallions olecranon fractures 449–50 olfactory receptors, sperm 952 oligospermia diagnostic evaluation 1523–4 testicular biopsy 1520, 1521, 1524 oliguria, neonatal foal 650, 653 omega-3 fatty acids 1603 supplements, stallions 1237–8, 1248, 1294 omega-6 fatty acids 1603 omeprazole 845 neonatal foal 194, 395–6 older foal 685, 686, 737, 743 ophthalmic disorders 541 septic arthritis 460 omphalitis see umbilical remnant infections omphaloarteritis 634 omphalocele 643 omphalophlebitis 202, 634 ultrasonography 251 oocyte(s) 2170, 2171 aged mares 2329 collection media 2932, 2937 collection of immature 2931 excised ovaries 2931–3 live mares 2931 collection of mature 2931, 2936–9 for GIFT 2945 hCG antibodies and 2051 methods 2936–8 oocyte recipients 2941 ovarian superstimulation 2936 ovulation disorders 2690 timing 2936 compact (Cp) 2932–3 cryopreservation 2953–5 culture after ICSI 2950–1 cumulus removal 2934, 2949 expanded (Ex) 2932, 2933 fertilization methods 2945 frozen-thawed, evaluation 2955 for GIFT 2945 holding medium 2934 for ICSI 2948 identification 2876, 2937 in vitro maturation 2931, 2933–4, 2938–9, 3052 maturation 2931, 2933–4 maturation medium 2933 maturity evaluation 2934, 2938, 2939 nuclear transfer into 2925 plasma membrane, sperm binding/fusion 953–4 polarity 2170, 2171 post-ovulatory aging 2329 preparation for ICSI 2949 sperm injection 2949, 2950 sperm interactions 952–5
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Index oocyte transfer (OT) 2938–9, 2941–4, 2945, 3052 geriatric mares 2814 for GIFT 2945 initial management of recipients 1827, 2941–2 insemination of recipients 2943 laparoscopic 1999 procedures 2942 success rates 2943 uterine tube pathology 1988 oolemma, sperm binding/fusion 953–4 oophoritis 2701 oothecitis 2701 ophthalmic disorders 535–51 acquired 537 congenital 537, 668 critically ill neonates 196 equine herpesvirus 2381 globe and orbit 537–41 mare reproductive loss syndrome 429, 2412 opioids administration neonatal foal 226, 227, 234 pregnant mare 57 endogenous autumnal transition 1734, 1746 crib-biting and 2771 female reproductive function 1614 male reproductive function 999 opsonins 332–3, 2589 optic disc 537 optic nerve colobomas 550–1 head 537 hypoplasia 551 optic neuritis 551 OptiprepTM 2964 oral intubation, neonatal resuscitation 131 orbit acquired disorders 540–1 congenital disorders 537–40 fetal see fetal eye inflammatory disease 540, 541 traumatic injuries 540 orchiectomy see castration orchiopexy 1524 orchitis autoimmune 1524 infectious 1162, 3024 traumatic 1161–2 ultrasonography 1473–4 organogenesis 2181, 2183, 2192–3 organophosphates, cutaneous habronemiasis 1140 orphan foals 113 feeding 270, 273, 2280–1 see also nurse mares orthopedic infections see osteomyelitis; septic arthritis oscillometric methods, blood pressure measurement 504–5 osmolarity plasma see plasma osmolality semen extenders 1336 osmotic nephrosis 650 ossification 433–9 abnormal 438–9 angular limb deformities and 768, 769 osteochondritis dissecans 772, 773 centers, secondary 434, 435, 436
definition 436 diarthrodial joints 437–8 endochondral 434, 776 newborn foal 67 nomenclature 434–6 premature or dysmature foals 126 specific bones 436–7 osteoarthritis (OA), juvenile 775 osteochondritis dissecans (OCD) 772–4 etiologic factors 776–9 growth rates and 285 nutritional factors 287–8, 778 pathogenesis 772–4 presale radiographs 810–11, 815, 816 prevalence in young horses 809 use of term 776 osteochondrosis (OC) etiological factors 776–9 manifestations 772–3, 774 pathogenesis 433, 438–9, 776 site vulnerability 779 terminology 772, 776 see also developmental orthopedic disease osteomyelitis, foal 457–62 clinical signs 457 complicating bacteremia/sepsis 159–60, 201, 457, 459 diagnostic evaluation 458–9 monitoring and prognosis 461–2 pathogenesis 438, 457 R. equi 695, 700–1 treatment 203, 459–61 outbreaks, disease 2799–801 outer acrosomal membrane (OAM) 928–9, 932, 933, 1491 sperm–oocyte interaction 1493 outer dense fibers, sperm tail see dense fibers, sperm tail ovarian artery 1593, 1594 rupture 2433 ovarian bursa 1588, 1589 ovarian cysts 2564, 2701, 2702 differential diagnosis 2707–8 fimbrial 2133, 2134, 2700, 2702, 2703 germinal inclusion 2701, 2702, 2703, 2707–8 neoplastic 2710, 2713–14 remnants of embryonic structures 2700, 2701, 2703 ultrasonography 2133–4 ovarian follicles ablation, estrus synchronization 1875–6 anovulatory see anovulatory follicles antral 2011, 2013–14 atresia 2013, 2048, 2933 development/dynamics see folliculogenesis deviation 2011, 2014, 2015, 2021 dominant see dominant follicles double see double dominant follicles echogenic ovulatory 2127–8 emergence 2011, 2014–15 growth waves see follicular waves histology 1588 luteinized unruptured (LUF) 2128 see also anovulatory follicles, luteinized multiple ovulatory development 2021–2 ovulation induction and 1866 pre-ovulatory see pre-ovulatory follicles
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primordial 2011 regression 2013, 2015 size/diameter anovulatory follicles 2687 estrous cycle 2013, 2014, 2015 factors affecting 2025–9 measurement 2010 at ovulation 2006 ovulation induction 1805, 1860–1, 1866 prepubertal mares 2012 repeatability 2025 single vs. double ovulations 2023 spring transition 2017 steroidogenesis 1631, 1632 autumnal transition 1737–8 vernal transition 1707–8 subordinate 2011, 2014, 2015 ultrasonography 2015, 2016 ultrasonography 2005, 2006, 2010, 2032–51 folliculogenesis 2009–18 measurements 2010 ovulation detection 1722, 2006, 2007 technique 2009–10 vernal transition 1706–8 ovarian fossa 1588 ovarian hematomas 2702, 2708 transrectal palpation 1911 ultrasonography 2130–1 ovarian ligament 1588, 1993 ovarian neoplasia 1911, 2707–14 breeding problems 2637–8 differential diagnosis 2703–4, 2707–9 epithelial origin 2710 molecular pathogenesis 2709–11 ovariectomy 2564, 2565 tumor types 2709, 2710 ultrasonography 2132–3 see also granulosa theca cell tumors ovarian vein 1593–4 ovariectomized mares estrogen therapy 1825 estrous behavior 1763, 1851 as mount mares 1270–1 stallion training 1399, 1403 as teasers 1271 ovariectomy 2564–72 complications 2571–2 diagonal (oblique) paramedian approach 2567–8 flank approach 2566–7 hysterectomy with 2574–6 indications 1851, 2564 laparoscopic 1991–9, 2568–71 complications 2572 hand-assisted, standing mare 1997–8, 2570–1 in situ 1999 in situ coagulation and pedicle transection 1998–9 instrumentation 1991–2, 1993 recumbent mare 1998, 2569–70 relevant anatomy 1993 standing mare 1993–7, 2569 paramedian approach 2568 preoperative preparation 2564 vaginal approach (colpotomy) 2565–6 ventral midline approach 2568 ovaries 1586–9 abnormalities 2123–34, 2697–705 abscesses 2709
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ovaries (Cont.) anatomy laparoscopic 1993 microscopic structure 1588 angiomatosis 2705 asynchrony of cyclicity 2669–70 autumnal transition 1735–41 blood supply 1593–4 development 2697–9 cellular components 2212–14 molecular steps 2214–15 dopamine receptors 1789 enlarged, differential diagnosis 2707–9 evaluation in anestrus 1696 excised, oocyte collection 2931–3 fetal 2698–9 folliculogenesis 2011, 2699 size changes 1589, 2698 ultrasonographic detection 2086, 2092, 2097–8 fetal hamartomas 2699–700 fibrinous and fibrous adhesions 2702, 2704, 2705 hormones, control of FSH and LH secretion 1624–7 hypoplasia 2701 infarction 2705 neoplasia see ovarian neoplasia non-neoplastic lesions 2697–705 acquired 2701–5 detection 2697 developmental 2699–701, 2702 relaxin 1683 senescence 2123, 2124 size 2006 autumnal transition 1741 fetus 1589, 2698 steroidogenesis 1631, 1632 autumnal transition 1735–6, 1737–8 vernal transition 1707–8 superstimulation see superovulation transrectal palpation 1721, 1910–11 ultrasonography abnormalities 2123–34 corpus luteum 2055–64 diestrus 1728, 2006–8 folliculogenesis 2009–18 indications 1914 normal anatomy 2005, 2006–8 ovulation 1722–3, 2042–8 ovulation prediction 2006, 2007, 2037–40, 2041, 2042 pre-ovulatory follicles 2032–51 varicosities 2702, 2704–5 zebras 2855–6, 2857 ovariohysterectomy complete 2574–6 partial 2574, 2576 see also hysterectomy overbreeding 1432 overo lethal white foal syndrome (OLWFS) 407–8, 668, 675–6 albinism 557–8 clinical signs 675–6 diagnosis 381, 676 overweight, breeding stallions 1221, 1228 oviductal epithelial cells (OECs) 1085, 1087, 1589 release of bound sperm 1088–9 sperm adhesion 944, 1085–7 clinical implications 1089 laboratory testing 1501 molecular basis 1087–8 oviductins 2172
oviducts (uterine tubes) accessory 2692 adhesions 2693 aged mares 2328 ampulla 1589 anatomical relations 1586, 1588 anatomy 1588, 1589 disorders 2692–5 embryo transport 1633, 1643, 2200 aged mares 2328 mechanisms 2172–3 PGE2 therapy and 1801 epithelial cells see oviductal epithelial cells fimbriae 1589 hyperactivated sperm motility 951 infundibulum 1588, 1589 fluid accumulation at ovulation 2042–4 isthmus 1588, 1589 laparoscopic ligation 1999 masses occluding 2328–9, 2693–4 diagnosis 1988–9, 2694 incidence 1989 treatment 1801, 1990, 2694–5 neoplasia 2692–3 patency tests 1988–90, 2694 sperm–oocyte interaction 952–5 sperm reservoir 944, 1085–9 characterization 1085–7 clinical implications 1089 molecular basis 1087–8 sperm release 944, 1088–9 sperm transport to 941–4, 1085, 1093, 2692, 3036–7 structural abnormalities 2692–3 transrectal palpation 1910 uterine ostium 1589 ovotestis 1136, 1961 ovulation diestrus 1759 disorders 1757–8, 2685–90 ultrasonography 2127–31 see also anovulation; anovulatory follicles double 2022–3 diagnosis of twins 2100, 2248, 2353 incidence 2022 role of hormones 2022–3 effects of semen 1096 embryo donor mares 2873 endocrinology 2020–5 double ovulations 2022–3 estrogen 1631 FSH and LH 1621, 1622, 1623 hormone interrelationships 2023–4, 2025 failure see anovulation first (puberty) 1690, 2012 first of the year 1705, 1728 follicle size and hormone levels 2027–8 progesterone for advancing 1778–81 follicle development 2020–9 induction 1858–66 agents used 1861–5 AI programs 1205, 1327, 1805, 1860 assessing follicle maturation 1860–1 estrus suppression 1849–50 factors affecting results 1865–6 failure 2689 geriatric mares 2807–9 GnRH 1820–3, 1862–4
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hCG see under human chorionic gonadotropin low dose insemination 3037, 3039 natural service 1804, 1859–60 for oocyte collection 2936 for oocyte transfer 2941 reasons for 1859–60 reproductive efficiency 2782 transitional mares 1806, 1859, 1862, 1864 twin pregnancy rates 2351–2 see also superovulation jenny donkey 2836 last of the year 1705, 1732 missed, impact of 1729 postpartum timing and fertility 2297–9 treatment to delay 2298–9 prediction 1720–6 AI recipients 1205, 1206, 2989 digital cervical examination 1726 embryo recipients 2872 rationale 2032, 2037–8 transrectal palpation 1721–2, 1911 ultrasonography 2006, 2007, 2037–40, 2041, 2042 vaginal examination 1723–6 see also pre-ovulatory follicles prostaglandins and 1642–3 secondary 2012–13 synchronization see estrous cycle(s), synchronization timing of insemination and 1205, 2988–90 ultrasonography 1722–3, 2042–8 follicle blood flow 2046, 2047 follicular fluid in abdomen 2046–7 normal evacuation and infundibular fluid 2042–4 septated evacuation 2044–6 zebras 2855 ovulation fossa 1588 cysts 2133, 2134 ovulatory agents 1861–5 estrus synchronization 1873–4 response, geriatric mares 2808–9 OvuplantTM see deslorelin, implants oxacillin 257, 846 oxalate, dietary intake 1234 oxibendazole ascarids 295, 759 strongyles 298 Strongyloides westeri 386, 391 oxidase test 1980–1, 2676 oxidative respiration, sperm 937, 938, 939, 1064 oxidative stress sperm 939–41, 991–4 see also reactive oxygen species oxygen (O2 ) placental exchange 88 supplies, NICU 822 utilization by sperm 939 oxygen extraction ratio (O2 ER) measurement methods 186 neonatal foal 142, 186 oxygen free radicals see reactive oxygen species oxygen saturation, arterial, neonatal foal 183–4, 215 oxygen tension (PO2 ) arterial (Pa O2 ), neonatal foal 169, 185 perinatal asphyxia syndrome 150 pneumonia 587
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Index prematurity or dysmaturity 125 reference values 605, 854 respiratory disorders 577 respiratory support 189 sepsis 155, 159, 200 see also hypoxemia umbilical artery and vein 215 uterine artery and vein 577 venous (Pv O2 ), neonatal foal 185–6, 577 oxygen therapy fetus, during dystocia correction 28, 132 neonatal foal 216 acute respiratory distress syndrome 583 anemia 424 birth resuscitation 128, 131 complications 216 critical care 185, 189, 190 indications 215, 578 perinatal asphyxia syndrome 150 persistent pulmonary hypertension 597 pneumonia 587, 606 prematurity and dysmaturity 125 prior to/during transport 233 sedation/anesthesia 223 sepsis 159, 203 shock 140 older foal pneumonia 717 pregnant mare anesthesia 56, 58 high-risk pregnancy 28, 53 see also hyperbaric oxygen therapy Oxyglobin see hemoglobin-based oxygen carriers oxyhemoglobin dissociation curve, neonatal foal 577 oxytetracycline 846 flexural limb deformities 442–3, 666 Lawsonia intracellularis 739, 762 neonatal foal 194, 260 retained placenta 2525, 2528 oxytocin (OX) 1679–80, 1830–4 cyclic luteolysis and 1644 exogenous 846, 1830–4 after insemination 1206, 2809 corpus luteum persistence 1760 delayed uterine clearance 1830–1, 2602, 2629, 2660 embryo recipients 2872 embryo recovery 2875 estrus suppression 1834, 1849 fetal hydrops 34–5 foal heat breeding 2299–300 foal rejection 119 induction of abortion 2321, 2371 induction of parturition 29, 1832–3, 2265–6 inhibition of luteolysis 1834 intrauterine sperm transport and 942 milk letdown 63–4, 270 nurse mares 2284 peripartum hemorrhage 2450 plugged ampullae 1115, 1414 post-mating endometritis 1832, 2615, 2810 post-mating uterine fluid 2145, 2146 postpartum conditions 1833–4 postpartum metritis 1833, 2532–3 postpartum uterine involution 1833, 2292 prior to AI 1203
problem-breeding mares 2628, 2629 prostaglandin release induced by 1831–2 retained fetal membranes 1833, 2525 function 1680, 1830 hypothalamic cells secreting 1582 male reproductive function 1680 maternal recognition of pregnancy 1644, 1680, 2204–7 mechanism of action 1679 milk let-down 1680, 2277 parturition 1680, 2228, 2269 postpartum uterine contractions 2521 pregnancy 2228 secretion pattern 1679–80 source of secretion 1679 structure 1679–80 testicular 898, 1003 uterine contractility and 2598 oxytocin receptors 1831 uterus 2205, 2228 Oxyuris equi (pinworms) 296, 298–9 p19 see uterocalin packaging cooled semen 1324 frozen sperm 2976, 2977, 3006, 3007 see also containers packed cell volume (PCV) mare, guiding blood transfusion 2451 measurement 177, 178 neonatal foal 142, 173, 187, 419, 420 cardiovascular disease 513 critical care 177–8 guiding fluid therapy 214 guiding transfusion therapy 238, 424 neonatal isoerythrolysis 354, 357 reference values 851 padded boots, mare’s hind feet 1224, 1270, 1409 paddocks estrus detection 1718 fencing, breeding stallions 1220–1 lighting systems 1775 see also pasture pain control see analgesia Paint Horses hereditary muscular disorders 463, 464 seminal characteristics 1369–70, 1371 palate cleft see cleft palate cysts 611–12 dorsal displacement see dorsal displacement of soft palate palatopharyngeal arch, rostral displacement 614, 664–5 presale endoscopy 797 pale mucous membranes, neonatal foal 138, 168 palmitic acid, sperm content 1062 Palomino stallions, seminal characteristics 1373, 1375 pampiniform plexus 872, 873, 1145 thermoregulatory function 882 pancreas, endocrine, foal adaptation to parturition 70, 75–7 pancreatic polypeptide (PP) 70, 77, 78 panniculus (cutaneous trunci) reflex 478, 480 pantoprazole 396, 846 papillary adenocarcinoma, ovarian 2710 papillary cyst adenoma, ovarian 2710
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papillomatosis, congenital 561, 564 papillomaviruses, penile and preputial neoplasia and 1169 paracrine action 1601 paracrine–autocrine system, intratesticular 999–1003 malfunction in subfertility 1437–7, 1439 measurement of function 1438, 1440 paracrine pathways, epididymis 924 paradoxical breathing, neonatal foal 169 paramedian approach, ovariectomy 2568 paramesonephric ducts (mullerian ducts) 2211–12, 2213 congenital abnormalities 2607 cystic remnants see fimbrial cysts, paramesonephric female genital tract development 2215 regression in male 1015, 2215 remnants, mare 1944 paranasal sinuses congenital abnormalities 609–11 cysts 611 fractures 454 Paranoplocephala mamillana 299 paraovarian (periovarian) cysts 2133, 2134 differential diagnosis 2707–8 see also ovarian cysts paraphimosis 1132–3 parapoxvirus ovis, inactivated 717 Parascaris equorum see ascarids parasites control 300 broodmares 111, 2748 foals/young horses 113, 295, 300 objectives 292 stallions 1222 diarrhea in foals 390–1, 741–2 gastrointestinal 292–300 pneumonia in foals 711–12 weight loss in foals 758–60 parasiticides available in US 295 parasite susceptibilities 296, 300 see also anthelmintics parathyroid hormone-related protein (PTHrP) 776 paraventricular nucleus 1582, 1585 parentage testing 2820–5 additional laboratory services 2824 case analysis 2823–4 DNA-based 2820–5 ISAG comparison tests 2822 microsatellite DNA markers 2821–4 rules 2820 standardization 2822–3 parental behavior in natural conditions 1388–9 see also maternal behavior parenteral nutrition (PN) foal 193, 273–8 complications 276–8 components 274–5 diarrhea 736 equipment 835 formulation 275–6 gastroduodenal ulceration 686 hepatic lipidosis 413 sepsis 158 pregnant mare 29 parietal hernia 403, 641 parietal vaginal tunic 867
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parity mammary secretion electrolytes and 2264 placental development and 2237–8 paromomycin, Cryptosporidium diarrhea 386, 390 paroophoron, cystic 2700, 2701, 2702 parrot mouth 667 parthenogenesis 3054 partial failure of passive transfer (PFPT) 362 definition 175, 344, 346 infection risk 347 plasma transfusion 240 prevention 350–1 risk factors 346–7, 351 treatment 344, 348–50 parturition 2268–75 detecting problems 2482–3 endocrine adaptation of foal 69–80 endocrinology 2268–9 environment 112 estrogens 1634, 2268–9 fetal death 25 fetal HPA axis and 69–72 guarding, harem stallions 1388–9 hemorrhage related to see periparturient hemorrhage induction of see induction of parturition jenny donkey 2836 maternal abdominal wall injuries 31–2 Miniature ponies 2841–3 normal process 2481–2 oxytocin function 1680, 2228, 2269 placental evaluation 2272–5 prediction of 112, 2269–70, 2480–1 progestagens 2224–5, 2268 prostaglandins and 1645, 2228, 2269 stage I 2270–1, 2481–2 stage II 2271, 2482 stage III 2271–2 stages 112, 2262 time of day 2269, 2481, 2733, 2734 zebras 2857–8 see also foaling passive transfer of maternal antibodies 67, 343–4, 346–51 assessing adequacy 113, 175, 346, 347–8, 2271 enteral feeding 268 failure of see failure of passive transfer immunoglobulin isotypes 337–8, 342 mechanism 2278 mucosal immunity and 338 partial failure see partial failure of passive transfer vaccination and 296, 307 vaccine-mediated interference 307–8, 338 see also colostrum; immunoglobulin G pastern fractures 447–8 pastern joint (proximal interphalangeal joint) arthrodesis 474 presale radiographs 803, 804 subluxation 447–8, 474 Pasteurella spp. pneumonia 710, 712 pasture breeding 1396 estrus detection 1718 feeding broodmares 2747, 2764, 2765
growing horses 282–4 stallions 1231, 1236, 1238 management broodmare farms 2749 elimination of fescues 2283, 2423 removal of mares from infected 351, 2423 see also paddocks patella congenital luxation 666 upward fixation 667 patellar reflex 479, 480 patent ductus arteriosus (PDA) 500, 508, 512 pathogen-associated molecular patterns (PAMPs) 331, 338 PCR see polymerase chain reaction PDC-109 1088 peanut (Arachis hypogaea) agglutinin (PNA) 1288, 1491, 1503, 1504, 3000 Pearson, Professor 1262 Pelger–Hu¨et anomaly 366 pelvic abnormalities, pregnant mares 6, 10, 32 pelvic floor, vulvar relations 1579–80 pelvic fractures, foals 452–3 penicillin-binding proteins (PBPs) 257 penicillins 846 chronic endometritis 2632, 2633 fetal intracardiac injection, twin pregnancy 2114, 2115 foal diarrhea 736 infectious hepatitis 412 intrauterine infusion 2632, 2633 neonatal foal 255, 257–8 dose 194, 254 prior to/during transport 232 neonatal sepsis 157, 202 placentitis 2363, 2364 pneumonia in older foals 715, 716 postpartum metritis 2532 R. equi pneumonia 698–9 semen extenders 1338, 1341 strangles 707, 765 Taylorella equigenitalis carriers 1251 penis 875–8 abnormalities 1130–42 bent or folded 1420 carcinoma 1136, 1137–9, 1169, 1451 diagnosis 1137–8 prognosis 1139 treatment 1138–9 cleaning adverse effects 1286 after natural service 1227 prior to natural service 1225 prior to semen collection 1269 training sessions 1394, 1404–5 congenital abnormalities 1111 delayed tumescence 1420 deviations 1419 dysfunction 1419–20 erection see erection, penile erosions 1130 fetal sex determination 2086, 2089–91, 2095, 2096 fractured, broken or ruptured 1131 habronemiasis 294, 1139–40, 1451 hematomas 1131–2 lacerations 1130–1 lesions causing phimosis 1133–4 neoplasia 1136–9, 1169 normal flora 1119 paralysis
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
complicating priapism 1135 management 1133 paraphimosis 1132–3 surgical procedures 1562–4, 1566 physical examination 1458, 1459–60 precancerous lesions 1137 reduced sensation and proprioception 1419–20 retaining devices 1132, 1133 sarcoids 1169, 1451 surgery 1562–71 surgical transection see phallectomy, partial swab samples pyospermia 1119–20 seminal vesiculitis 1115 Pentafusion pain management 846 pentalogy of Fallot 670 pentobarbital 493, 846 pentose phosphate pathway, sperm 939, 954–5 pentoxifylline 846 foal diarrhea 736–7 hepatic failure 412, 413 placentitis 2364, 2365–6 postpartum metritis 2532 pregnant mare 27, 33 septic foal 157 peptide hormones 1602, 1603, 1606 Percherons, hereditary muscular disorders 464 perforatorium 1058 performance horses, fescue toxicosis 2423 performance mares behavioral problems 1762–3 estrus suppression 1815, 1834 reproductive history 1898 perfusion, tissue see tissue perfusion pergolide lactation problems 2283, 2285 ovulation failure 2689 pituitary pars intermedia dysfunction 28, 2758, 2794 pericardial abnormalities, acquired 516 pericardial effusions, neonatal foal 139 pericardial neoplasia 516 pericardial trauma 515 pericarditis mare reproductive loss syndrome 516, 2412, 2413–14 weight loss 763 periglandular fibrosis (PGF), endometrial 1931, 1932, 1933–4, 1936, 2638 perimetrium 1589, 1590 perinatal asphyxia syndrome (PAS) 147–52 breathing patterns 575 clinical signs 147 diarrhea 392 pathophysiology 72, 147–8 predisposing factors 147 treatment 148–52 see also neonatal encephalopathy perineal body 1577 reconstruction see episioplasty surgical procedures 2552–7 transection (PBT; perineoplasty; Pouret operation) 2541, 2543, 2552, 2553 perineal lacerations (PL) 2552–6 first-degree 2552, 2553 second-degree 2552 surgical correction 2554–6 third-degree 2489, 2553–4
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Index perineum, female 1577 abnormal conformation 1577–8, 2623 anatomy 1577–81 assessing conformation 2622–3, 2804 Caslick index 1577–8, 2537–8 embryo recipients 2872 fetal sex determination 2097 see also female external genitalia; vulva perinuclear theca 930 period genes 1772, 1773, 1776–7 periodic breathing neonatal foal 169 premature foals 125 periodic paralysis, hyperkalemic see hyperkalemic periodic paralysis periodontal disease 2748 periorbital fractures, foals 454 periosteal stripping, angular limb deformities 770 periovarian adhesions 2702, 2704, 2705 periovarian (paraovarian) cysts 2133, 2134 differential diagnosis 2707–8 see also ovarian cysts periparturient hemorrhage 2449–52, 2517–19 clinical signs 2518 diagnosis 2517–18 epidemiology 2517 external genitalia 2451–2 pathogenesis 2517 prognosis 2519 treatment 2518–19 ultrasonography 2155–6, 2450, 2517, 2518 uterine 2449–51, 2517–19 uterine prolapse 2433 vaginal and vestibular foaling trauma 2451, 2452 periparturient management geriatric mares 2813–14 mare and neonate 111–13 uterine ultrasonography 2150–7 peripheral nerve injuries 489 peritoneal fluid analysis 855, 1146 uterine torsion 2436 ultrasonography 243, 244 see also abdominocentesis peritonitis after cesarean section 2509 complicating ovariectomy 2572 foal 243, 763 postpartum mare 2498, 2517 periuterine adhesions 2672–3 perivitelline space 2171 permethrin 2416 peroneus tertius rupture 472–3 perphenazine agalactia 2283 fescue toxicosis 33 transitional mares 1789, 1790–1 persistent pulmonary hypertension (PPH) 590, 597 persistent pupillary membranes (PPMs) 547 persistent vitelline circulation 643 pest control, breeding facilities 3030 petechial hemorrhages neonatal foal 139, 155, 168, 427 thrombocytopenia 373, 427–8 Peyer’s patches 338 pH blood, foals 150, 605, 854
caudal epididymal fluid 935 gastric 394 hyperactivated sperm motility and 951 oocyte maturation medium 2933 semen see semen, pH semen extenders 1064, 1336 urine 183, 651, 655 phagocytes 332 dysfunction 366 function testing 367 phagocytosis, sperm within uterus 1093, 1095 phallectomy, partial 1566–70 complications 1570 by en bloc resection with penile retroversion 1569–70 without penile retroversion 1570 indications 1566 patient preparation 1566 penile and preputial carcinoma 1138 postoperative care 1570 priapism 1135 Scott’s technique 1568–9 urethral lacerations 1131 Vinsot’s technique 1566–7 William’s technique 1567–8 phallopexy, Bolz technique 1563–4 phantom mare 1199, 1200, 1271 facility or environment 1397, 1402–3 history of development 1264 materials and construction 1402, 1403 miniature stallions 1377–8 stallion training 1394, 1400–1, 1402–6 approach and collection 1405–6 preparation and introduction 1403–5 pharmacogenetics 620 pharmacology, neonatal foal 253 pharyngeal cysts 611–12 dorsal 593, 594 management 595, 611–12 subepiglottic 593, 594, 613 pharyngeal flush 846 pharyngeal lymphoid hyperplasia (PLH) 794, 795, 796 pharynx, congenital abnormalities 611–13 phenazopyridine 846 cystitis 628–9 patent urachus 638 phenobarbital 846 neonatal seizures 193, 194, 234, 486 perinatal asphyxia syndrome 149 phenothiazines agalactia 2282–3 foal 222 penile paralysis 1132 pregnant mare 56–7 priapism induced by 1134, 1135 phenotypes, producing desired 3052 phenoxybenzamine, intracavernosal injection 1131–2 phentolamine, intracavernosal injection 1131–2 phenylbutazone breeding-induced endometritis 2812 high-risk pregnancy 26 neonatal foal 226, 227 poor libido in stallions 1415 septic arthritis and osteomyelitis 460 strangles 707 testicular trauma 1161
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
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phenylephrine, intracavernosal, priapism 1135 phenylpropanolamine 847, 1127 phenytoin 194, 847 R 1418 Pherocalm pheromone, calming 1418 phimosis 1133–4 phosphate (HPO4 ), reference values 853 phosphate enemas, meconium impaction 381–2 phosphatidylcholine effects of cooling 1310 semen cryopreservation 2968–9 phosphatidylinositol-4,5-bisphosphate (PIP2 ) 1605 phosphatidylserine, cooled sperm storage 1074 phosphocaseinate, native (NPPC) 2968 phosphoinositide 3-kinase (PI3-K) 2710 phospholipase A2 (PLA2 ) 1603 acrosome reaction 950 embryonic expression 2203–4 phospholipase C (PLC) acrosome reaction 949 hormone activation 1605 sperm–oocyte interaction 954 phospholipid hydroperoxide glutathione peroxidase (PHGPx) 947 phospholipids membrane breakdown products 1605 effects of cooling 1071–2, 1309–10 sperm content 1062 sperm plasma membrane 1055–6 phosphorus 787 developmental orthopedic disease and 778, 787 dietary intake, aged horses 2756, 2758 dietary requirements breeding stallion 1230, 1234 broodmare 264, 2762, 2763, 2766 growing horses 787 neonatal foal 266 dietary sources 787, 1234 fractional excretion 855 mare’s milk 2280, 2281 serum, neonatal foal 173 supplements, growing horse 288 photic gate concept 1712 photoadaptation 1672 photoperiod 1670, 1771–7 anatomical pathway 1771–2 autumnal transition 1733 control of seasonality in mares 1782–3 effects of latitude and 1733, 1775 gonadotropin secretion and 1675–7 male endocrine function and 1004–5 manipulation see lighting programs, artificial melatonin secretion and 1673–5 nature of signals in horses 1773 refractoriness 1346–7 sperm production and output and 1346–50 photoreceptors, retinal 1672, 1772 photorefractoriness 1347, 1348, 1704 photoresponsiveness, circannual rhythm 1704 phthisis bulbi 541 Phycomycetes 711 physeal fractures 446 long bones 451, 452 proximal first phalanx 447
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physes (metaphyseal growth plates) 434–6 abnormal development 438–9, 768 definition 434 definitive 435 development 434–5 presumptive 434–5 transphyseal bridging 770 physical appearance, general normal neonatal foal 65 post-term or post-mature foal 122 premature foal 122, 167 sick foal 167 physical examination geriatric mare 2804 neonatal foal 65, 113, 167–72 pregnant mare 2245–59 physitis 774, 775 angular limb deformities and 768 dietary factors 785 septic 159–60, 201 spontaneous self-correction 776 pica 391 see also sand enteropathy Piezo drill, for ICSI 2948, 2949–51 pigeon fever 765 pineal gland 1669 anatomy 1582, 1583 circadian system pathway 1671–2 control of seasonality in mares 1782–3 melatonin synthesis 1671 pinealocytes 1671 pink-eye, venereal transmission 3015 pinky Arab syndrome 568–70 R vaccine 708 Pinnacle pinworms (Oxyuris equi) 296, 298–9 piperacillin 257–8 piperazine ascarids 295, 759 strongyles 300 pipettes, for ICSI 2948–9 piroplasmosis, equine, venereal transmission 1256, 3024–5 piroxicam, penile and preputial carcinoma 1139 Pisum sativum agglutinin (PSA) 1288, 1491, 1503, 3000 pituitary adenyl cyclase activating peptide (PACAP) 1772 pituitary extracts, equine (EPEs) FSH:LH ratios 1837, 1838 ovulation induction 1865 subfertile stallions 1443–3 superovulation induction 1837, 1838, 1841 see also equine follicle-stimulating hormone pituitary gland (hypophysis) anatomy 897–8, 1584–5 enlargement 2790–1 hypothalamic connections 1585–6 pars intermedia (PI) 2790 vascular relationships 1584–5 pituitary pars intermedia dysfunction (PPID) (equine Cushing disease; ECD) 1665–6, 2790–5 aged horses 2756, 2757, 2758, 2791, 2805–6 clinical signs 2791–2 diagnostic testing 2792–3 dopamine antagonists 28 estrous cycle abnormalities 1755, 2636–7 galactorrhea 2285
management 2793–5 ovarian abnormalities 2124–5 ovulation failure 1758, 2689 pathophysiology 2790–1 pituitary portal vessels 1584–5 monitoring GnRH secretion 1611 pivampicillin, neonatal foal 254, 257 pivoting test 728 placenta 84–94, 2234–40 anatomy 85 avillous sites 90–1, 92, 102, 105 blood flow 88 components 86–8 development 85–8, 2187–98, 2234–40 20–30 days 2188–90 30–40 days 2191–4 41–60 days 2194–7 60–80 days 2197–8 allantochorionic 85–6 cellular proliferation 2235–6 choriovitelline 85 extraspecies pregnancies 2306–7, 2310–11, 2314 factors influencing 89–90, 2237–40 growth factors controlling 86, 1655, 2236 microcotyledons 86–7, 2235 vascularity 87, 2239–40 efficiency factors affecting 2238–9 measurement 2236–7 nutritionally stressed mares 2239 embryological origins 85–6, 2188–90, 2234–5 endocrine functions 89 examination 65, 99–109, 2272–5 abortion 2341 color changes 92, 107–8 documentation of findings 109 effects of storage 103 identification of lesions 106–8 indications 99–100 interpretation 91–4 neonatal illness 155 normal appearances 100, 101 normal features mimicking lesions 103–6 in practice 100–3 retained fetal membranes 2272–3, 2523 sample collection 107, 108–9 fescue toxicosis 91, 2421 fetal effects of abnormalities 91 feto-maternal contact maternal age and 2238 maternal size/genotype and 2237 measurement 2236–7 fluids see fetal fluids gas exchange 88 historical aspects 84–5 immunological functions 89 manual removal 2512, 2513, 2525–6 measurements 106 nutrient transport and waste elimination 89 nutritive functions 89 pathology 91 premature separation 6, 11, 2482–3 fescue toxicosis 2421 management 33 mare reproductive loss syndrome 2411 placental evaluation 92, 107–8 ultrasonography 2154
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
progestagen synthesis 2222–3 relaxin and 1682, 1683 retained see retained fetal membranes separation at parturition 2291, 2294, 2520–1 specialized structures 87–8 steroidogenesis 1633–4 structure and function 2240 twin pregnancies 2346–7, 2356, 2357 ultrastructure of endometrial interface 88 unique features of equine 90–1 use of term 99 uterine horns 92, 102, 103, 104 weight 91–2, 106, 2522 placental area of degeneration (PAD), fetal foot 103–4 placental insufficiency 6, 91 diagnosis 12 management 35 placental lactogen 2228, 2277 placentation see placenta, development placentitis 2359–66, 2459–61 abortion and stillbirth 2347–8 cervical pole 2347–8 diagnosis 10–11, 2359 diffuse, often microscopic 2348 endocrine monitoring 2362–3 estrogen therapy 1827 focal single sites elsewhere 2348 galactorrhea 2284 high-risk pregnancy 6, 10–11 horn base with/without rostral body 2348 immunological aspects 2591–2 mammary secretion electrolytes 2264, 2735 pathogens causing 2359, 2363 perinatal asphyxia syndrome 147, 148 placental examination 92, 102–3, 106, 107, 2273–4, 2347–8 placental weight 91–2 progestagen therapy 25–6, 2364–5 treatment 32–3, 2363–6, 2460–1 ultrasonography 21, 2154, 2155, 2359–62 plains zebra (Equus burchelli) embryo recovery and transfer 2311, 2312–14 foal 2861 interspecies hybrids 2302, 2304, 2305 mare estrous cycle 2855–6 gestation 2856, 2857 parturition 2857–8 postpartum estrus 2858–9 puberty and breeding life 2853–4 reproductive organs 2855 mating patterns 2851–2 seasonality of breeding 2852–3 stallion behavior 2859 puberty and breeding life 2860 reproductive organs 2861 seasonality 2859 weanling 2862 yearling 2862 plasma (and plasma products) 209, 239–40 collection and storage 350 donor selection 240 electrolyte content 207 hyperimmune 847 intrauterine infusion 2630, 2661
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Index intravenous infusion see plasma transfusion oral administration, neonatal foal 344, 349 Rhodococcus 240, 847 topical use, corneal ulcers 546 see also serum plasma cells, endometrium 1932 Plasma-Lyte 56 with 5% dextrose 206, 207 Plasma-Lyte 148 206, 207, 231 plasma membrane, sperm see sperm plasma membrane plasma osmolality, neonatal foal 182, 209–10, 855 plasma proteins hormone binding 1603, 1604 total see total plasma protein plasma transfusion 847 complications 241 neonatal foal 113, 209, 239–40 critical care 190, 192 failure of passive transfer 209, 240, 349–50 premature 126 prior to transport 234 R. equi prophylaxis 240, 690–1 salmonellosis 386 sepsis 157, 202 shock 141–2 older foal/weanling/yearling diarrhea 736 salmonellosis 739 plasminogen 853 plasminogen activator inhibitor 853 platelet counts coagulation disorders 427 neonatal foal 173, 174, 428 reference values 851, 853 thrombocytopenia 373 platelets 373 play sexual behavior 1386, 1389 pleural effusion neonatal foal 580, 590 older foal 714 plica salpingopharyngea 613 plugged ampullae see ampullae, plugged Pmods 999 Pneumocystis jirovecii pneumonia 711, 713 pneumomediastinum, foal 590 pneumonia acute interstitial/bronchointerstitial see acute lung injury/acute respiratory distress syndrome aspiration see aspiration pneumonia genetic predisposition 615–16 neonatal foal 158–9, 586–8, 601–7 causative pathogens 602 diagnosis 169, 170, 200, 586, 603–5 epidemiology 601 identification of infectious agents 577–8, 604–5 management 203, 586–7 pathogenesis 601–2 physical examination 602–3 prognosis 607 treatment 605–7 nosocomial 219 older foal/weanling/yearling 710–17 clinical signs 712 control strategies 717 diagnosis 713–15 epidemiology 712 etiological agents 710–12 pathogenesis 712–13
treatment 715–17 weight loss 763 see also Rhodococcus equi pneumonia pneumoperitoneum, foal 243 pneumothorax, foal 590, 598–9 pneumouterus 1580 ultrasonography 2148 pneumovagina 1580, 2537–41 diagnosis 1902, 2537, 2538, 2547, 2624–5 endoscopy 1944 surgical procedures 1580, 2539–41, 2542–3, 2546–8 point-of-care analyzers, sick foals 179, 180, 834–7 polar body, mature oocytes 2934 poll posture, fetus 2490–1 polyhydramnios 2368 polymerase chain reaction (PCR) cytogenetic analysis 1954–5 gynecological bacteriology 1971 neonatal diarrhea 385, 387 parentage testing 2821–2 R. equi pneumonia 698 real-time (RT-PCR) 2830 strangles 706 Taylorella equigenitalis 1251, 1971, 2404–5 viral respiratory diseases 604–5, 721 polymixin B 847 foal diarrhea 736 neonatal sepsis 144, 157, 202 postpartum metritis 2532 topical ophthalmic 541 polymorphonuclear leukocytes (PMNs) see neutrophils polyorchidism 1109 polysaccharide storage myopathy (PSSM) 464–5, 668–9 polyvinyl alcohol, embryo recovery media 2874 polyvinylpyrrolidone (PVP) 2950 ponazuril 731, 847 ponies autumnal transition 1733, 1740, 1742 double ovulations 2022 fetal eye measurements 2078 pre-ovulatory follicle size 2025–7 seminal characteristics 1373, 1375 see also Miniature horses/ponies porcine zona pellucida (PZP) vaccine 1890–2 portal vein thrombosis, acute 411–12 portosystemic shunts, congenital 414, 732 positive end expiratory pressure (PEEP) 218, 219 positive-reinforcement methods, stallion training 1391, 1393, 1401 postacrosomal lamina 1057, 1058 post-breeding uterine fluid accumulation see intrauterine fluid accumulation, post-breeding posterior presentation see caudal presentation posterior ring, sperm head 925–6 posterior segment (of eye) acquired abnormalities 551 congenital abnormalities 550–1 posthetomy extensive (Adam’s procedure) 1562–3 penile and preputial carcinoma 1138 segmental 1133, 1562–3 post-mature foals 122, 126 post-mortem tissues
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epididymal sperm harvesting 2983–4 ovarian oocyte harvesting 2931–3 postpartum foal see neonatal foal postpartum mares anestrus 1757, 2296–7 autumnal transition 1735 estrogen therapy 1827 estrus detection 1719 first estrus see foal heat hysteroscopy 1943 Miniature ponies 2843–4 ovarian ultrasonography 2123, 2124 oxytocin therapy 1833–4 problems 2154–7 progestagen therapy 1816 uterine involution 2291–3, 2294–5 uterine ultrasonography 2141–2, 2154–7, 2299 see also foaling mares; lactating mares; lactation postpartum metritis (or metritis-endotoxemia-laminitis syndrome) 2292, 2530–3 clinical signs 2530–1 diagnostic testing 2531–2 etiopathogenesis 2524, 2530, 2531 prevention, retained placenta 2524–5 treatment 1833, 2532–3 ultrasonography 2156–7 post-term foals 122 post-term pregnancy see prolonged pregnancy posture female receptive 1388, 1409 fetal see fetal posture potassium (K) abnormalities see hyperkalemia; hypokalemia fractional excretion 855 pre-foaling mammary secretion 2733, 2735 assay methods 830–1 feto-placental monitoring 19–20 jennies 2270 predicting parturition 2270 renal excretion, neonatal foals 80 requirements breeding stallion 1230, 1235 broodmare 2762, 2763, 2766 serum 173, 853 supplements, neonatal foal 191, 211, 276 potassium bicarbonate (KHCO3 ) 850 potassium bromide 847 potassium chloride (KCl) 847, 850 potassium iodide 847 Potomac horse fever (PHF) foal diarrhea 742 pregnant mare 9, 10 vaccination 319–20 broodmare 306, 311, 320 foals and weanlings 295, 309, 320 stallion 1222 Pouret operation (perineal body transection) 1580, 2541, 2543, 2552, 2553 povidone iodine artificial vagina cleaning 1268–9 intrauterine infusion 2613, 2634, 2635, 2648 ophthalmic 847 umbilical stump treatment 635, 637 power, ultrasound 1916 praziquantel, tapeworms 299
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prebiotics 737 prednisolone 847 cutaneous habronemiasis 1140 nasopharyngeal irritation 272 ophthalmic 540 ovulation and 2689 post-mating endometritis 1096, 2616, 2662, 2812 problem-breeding mares 2631 transfusion reactions 241 prednisone, cutaneous habronemiasis 1140 preferences, specific, individual stallions 1410 pre-foaling mammary secretion electrolytes see mammary secretion electrolytes, pre-foaling pregnancy abnormalities of 2441–53 adrenal cortical hormones and 1666–7 anesthesia 55–9 anestrus 1696, 1761 corpus luteum persistence 1759 corpus luteum ultrasonography 2059, 2061, 2062–3 cycles per 1431, 2780 diagnosis 2245–59 behavior 2245 hormonal tests 2251–6 laboratory tests 2250–8 Miniature ponies 2840–1 speculum examination 2246 transrectal palpation 2246–7, 2248 ultrasonography 2067–8, 2069, 2247–50, 2251, 2252 uterine cysts and 2248–9, 2665, 2666 visual assessment 2245–6 embryo recipients, monitoring 2878 endocrinology 1634, 2222–30, 2320, 2461–7 estimating day of 2074, 2075, 2076–9 estrogens during 1632–4, 2225–7, 2463–4 biosynthetic pathways 2226 diagnostic use 1635, 2255–6, 2257, 2463 late-term 19, 1633–4 post-implantation 1633 pre-implantation 1632–3 estrus suppression 1850 examination 2245–59 abdominal ballottement 2246 options 2247 ultrasonography 2247–50, 2251, 2252, 2253 see also fetal sex determination extraspecies 2305–17, 3054 folliculogenesis in early 2017 geriatric mares 2813–14 glucocorticoids 2228–9 hCG therapy 1807 hemorrhage related to see periparturient hemorrhage high risk see high-risk pregnancy history 1898 interspecies 2303–5 jenny donkey 2836–7 maintenance of see maintenance of pregnancy maternal recognition see maternal recognition of pregnancy Miniature ponies 2841 neonatal isoerythrolysis screens 358–9, 831
nutrition 264, 2747–8, 2763–5 foal birthweight and 281–2 foal growth and 286–7 high-risk pregnancy 28–9, 53 oxytocin 2228 physiological landmarks 2246, 2246 placental evaluation 93 post-embryo donation 2878–9 preventive medicine 2750 progestagen therapy 25–6, 1816–17 progesterone 1638, 2222–5 prostaglandins 2228 relaxin 19, 1682–3, 2227–8 sedation 56–7 services per 2780 stallion-like behavior 1764 stress in 1817 timing of surgery 55 ultrasonography see ultrasonography (US), pregnant mare uterine anatomy 1591 uterine histotrophe 1639–40 vaccinations 306–7, 310–11 zebras 2856–7 see also broodmare pregnancy-associated glycoprotein (PAG) 2203–4 β 2 -pregnancy protein 2258 pregnancy losses 2455–61 anesthesia-related 55 cloned embryos 2925 history 1898 mare reproductive loss syndrome 2410–11 prevention see maintenance of pregnancy recurrent 2348 see also abortion; embryonic loss, early; fetal death; stillbirths pregnancy rates cooled semen 2991–2 definition 2780 embryo transfer 2871, 2884–5 cooled-embryo transport 2881–3, 2884–5 frozen-thawed embryos 2887, 2888–98 stored embryos 2880–1 vitrified-warmed embryos 2899, 2910–12 end of season 2780 epididymal sperm insemination 3039 first cycle 2779–80 foal heat breeding 2142, 2293, 2297, 2298, 2468 frozen-thawed sperm 2980, 2988–92, 2994 epididymal sperm 2985, 2986 packaging, dose and AI regime effects 2976, 2977, 2980, 2981 semen extenders and 2966–7, 2969 GIFT 2945, 2946 insemination dose and 3037 interspecies matings 2303 intrauterine fluid and 2145–6 low dose insemination 3038, 3039 oocyte transfer 2943 overbreeding and 1432 per cycle 1430, 1431, 2779–80 observed 2780–1 see also reproductive efficiency per season 1430–1, 2779 factors affecting 1430–1 per stallion 1362–3, 1430
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
sex-sorted sperm 3045, 3046 shuttle stallions 1184–5 see also fertility 5α-pregnane-3,20-dione (5α-DHP) 17–18, 2222, 2223, 2224 5-pregnane-3β,20α-diol (βα-diol) 2224 5-pregnane-3β,20β-diol (P5ββ) 2223, 2224 pregnant, opportunity to become 1431 pregnant mare serum gonadotropin (PMSG) 84, 1648, 2168 see also equine chorionic gonadotropin pregnenolone (P5) adrenal steroid synthesis from 1665 biosynthesis 893, 1602, 1631 fetal production 69, 71, 2223–4 ovarian steroid synthesis from 1631 parturition 2268 pregnant mare 17–18, 2224–5 testicular steroid synthesis from 893, 894 preimplantation genetic diagnosis (PGD) 3052 premature delivery causes 122, 123 inducing fetal maturation prior to 123–4 placentitis 2359, 2592 therapies preventing 25–8, 2363–6 premature foals 121–2 carpal fractures 448–9 clinical course 124 endocrine abnormalities 525–6 fluid therapy 213 hypothalamic–pituitary–adrenal axis 72, 524 laboratory assessment 124 management 125–6 necrotizing enterocolitis 392 nutrient requirements 266–7 physical appearance 122, 167 plasma creatinine 124, 653 prognosis 126 respiratory insufficiency 125, 576, 589 skeletal development 437 tendon laxity 469 thyroid function 75, 528 pre-optic nuclei 1582, 1585 pre-ovulatory follicles 2015–16, 2020–5 atresia 2048 endocrinology 2021–5 factors affecting size 2025–9 hCG antibodies and 2051 hematoma formation see hemorrhagic anovulatory follicles hormone interrelationships 2023–4 Miniature ponies 2025–7, 2839 multiple and double 2021–3 ovulation see ovulation repeatability of diameter 2025 transrectal palpation 1721 types of outcomes 2042–8 ultrasonography 2006, 2007, 2032–51 blood flow and perfusion changes 2035–6, 2037, 2038 B-mode echotextural changes 2033–5 prediction of ovulation 2037–40 signs of impending ovulation 2036–7, 2039, 2040 vascularity vs. fertility 2048–51 Preparation H 2452 prepro-gonadotropin-releasing hormone (prepro-GnRH) 1608 preprohormones 1603
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Index prepubertal mares, folliculogenesis 2011–12 prepubertal stage, stallion 902, 903 prepubic tendon rupture 8, 2152–3, 2428–30, 2447–8 clinical signs 2428, 2429, 2447–8, 2480 induction of abortion 2322–3 management 2428–30 ultrasonography 2428, 2429 prepuce 878 abnormalities 1130–42 aplasia or hypoplasia 1111 carcinoma 1136, 1137–9, 1169 diagnosis 1137–8 prognosis 1139 treatment 1138–9 congenital abnormalities 1111 edema 1460 complicating castration 1560 management 1133 pathophysiology 1130, 1132 erosions 1130 fetal sex determination 2086, 2090 lacerations 1130–1 neoplasia 1136–9, 1169 physical examination 1460 surgery 1562–71 swabs pyospermia 1119 seminal vesiculitis 1115 preputial lamina, internal fibrosis and scarring 1133 injuries causing paraphimosis 1132 lesions causing phimosis 1133–4 retaining devices 1132, 1133 preputial orifice, stenosis 1133, 1134 preputial ring constriction or stenosis 1111, 1133, 1134 surgical incision, paraphimosis 1133 presale (upper airway) endoscopy 794–9 conditions of sale relating to 796–8 grading systems 796 maturation-related changes 794–5 racing performance and 798–9 technique 794 variation in findings 795–6 presale radiographs 801–8 examination procedures 802–8 interpretation and significance 809–17 labeling 801–2 radiation safety 801 sedation 802 pressure relieving mattress, NICU 824 pressure sores, flexor tendon laxity 470–1 pressure support ventilation (PSV) 217–18, 219 preventive medicine breeding stallion 1221–2 broodmare farms 2747–51 priapism 1134–6 cavernosal shunting 1564–6 clinical signs 1135 etiology 1134 high-flow (arterial) 1134 low-flow (veno-occlusive) 1134–5 paraphimosis due to 1132 pathophysiology 1134–5 treatment 1135–6 PRIDTM intravaginal device 1814, 1879 estrus synchronization 1826, 1875 transitional mares 1780 primiparous mares see maiden mares
principal piece, sperm 931, 932, 934, 1028, 1061 abnormalities 1301 prion proteins 1865 probanthine bromide, rectal palpation 1908 probiotics 272, 388, 737 problem-breeding mare 2620–39 breeding management 2627–39, 2783 causes 2621 cervical lacerations 2559, 2562–3 characteristics 2620–1 chronic endometritis see endometritis, chronic clinical examination 2622–5 contagious equine metritis 2401–2 cytogenetic analysis 1955–9 fescue toxicosis 2422 fungal endometritis 2635, 2643–50 laboratory diagnostics 2625–7 treatment options 2628, 2628–9 ultrasonography 2143–5, 2623–4 urovagina 2548–9 see also barren mares; uterine clearance, delayed; specific causes of subfertility procainamide 514, 847 proctoscopy, intestinal atresia 406 progestagens (progestins) 1811–17, 2466–7 controlled release formulations 1879–81 exogenous 1811–17 contraception in mares 1889 embryo reduction in twin pregnancy 2101 estrus suppression 1762, 1815, 1846–7 estrus synchronization 1815–16, 1872–3 foal heat breeding 2468–9 indications in mares 1815–17 maintenance of pregnancy 2334–5, 2469–72 ovarian function and 1755 parturition and 2268 placentitis 25–6, 2364–5 postpartum mares 1816, 2141–2 pregnant mare 25–6, 1816–17, 2120–1 problem-breeding mare 2638–9 suppression of stallion-like behavior 1194, 1416–17 transitional mares 1713–14, 1778–81, 1815 see also altrenogest implants, estrus suppression 1847 intramuscular delivery 1879–81 intravaginal delivery see intravaginal progesterone devices parturition 2224–5, 2268 placental production 89 placentitis 2362 pregnant mare 17–19, 2222–5 synthetic 1811–13 progesterone (P4) 1637–40, 1811, 2466–7 acrosome reaction induction 956, 1493–4 anestrus due to 1696–7 biological functions 1638–40 biosynthesis 1637 autumnal transition 1735–6 sources 1638 blood levels defining pubertal estrus 1690
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diestrus 1728, 1729 donkey-in-horse pregnancies 2315, 2316 early embryonic loss and 2330 early pregnancy 2251–2, 2254, 2254, 2255, 2466 embryo transfer and 2330, 2878 granulosa cell tumors 2712 hCG treatment effects 1806 horse-in-donkey pregnancies 2306, 2308 late-term pregnancy 17–19 ovulation disorders 2688 placentitis 2362 pregnancy in Miniature mares 2840 corpus luteum ultrasonography and early pregnancy 2061, 2062 estrus cycle 2060, 2062 exogenous 1814–15 cervical surgery 2560 contraception in mares 1889 embryo donor mares 2874 embryo recipients 2878 estrus suppression 1846 estrus synchronization 1872–3 geriatric mares 2813 injectable 1814 intravaginal delivery see intravaginal progesterone devices long-acting injectable 1814, 1880–1 maintenance of pregnancy 2120–1, 2330, 2334, 2466–7, 2469–72 oocyte recipients 2942, 2943 placentitis 2364–5 postpartum mares 2298–9 problem-breeding mare 2638–9 transitional mares 1713–14, 1778–81 follicle development and 1730 multiple vs. one 2021, 2022 FSH and LH secretion and 1625, 1639 GnRH secretion and 1613–14 insufficiency, early embryonic loss 2638–9 lactogenesis and 2277 luteinized anovulatory follicles 2129 mechanism of action 1637 parturition 2268, 2269 periovulatory period 2024 pregnancy 1638, 2222–5, 2320 pregnant zebras 2857, 2858 retained fetal membranes and 2523 uterine contractility and 2598 uterine defenses and 2589 uterine protein secretion and 2202 progesterone/estradiol (P&E) treatment contraception 1889 diestrus 1730–1 estrus suppression 1847 estrus synchronization 1816, 1826, 1873 intramuscular delivery 1880 postpartum mares 1816, 1827, 2292–3, 2298–9 prior to superovulation with eFSH 1839, 1841 transitional mares 1779 progesterone receptors (PR) 1637, 1811 cervical 2728 early conceptus 2202 estrous cycle 1631–2 pregnant endometrium 1633 sperm 1493, 1502 progestins see progestagens prohormones 1603
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prokinetic agents 271, 396 prokineticin-2 (PK2) 1772 prolactin 1683–4 ergot alkaloid effects 2418, 2420, 2423–4 exogenous, seasonal anestrus 1699, 1789 male/stallion 998, 1003–4, 1683, 1684 mare 1683, 1684 autumnal transition 1736, 1737, 1745–7 estrogen treatment and 1791 lactogenesis 2229, 2277 regulation by dopamine 1789 PGF2α and 1747 physiology 1683 proliferative enteropathy, equine (EPE) 760–3 clinical signs 756, 760–1 diagnosis 739, 761–2 diarrhea 739–40 epidemiology 760 prevention 762–3 prognosis 762 signalment 760 treatment 209, 739, 762 see also Lawsonia intracellularis proligestone 1812, 1813 proline, semen cryopreservation 2970 prolonged gestation 11, 2448–9 causes 122 fescue toxicosis 11, 122, 2154, 2419–20, 2449 fetal oversize 2487 ultrasonography 2154 prolonged pregnancy, management 124 prolylendopeptidase (PEP) 1610 pronuclear transplantation 3054 pro-opiomelanocortin (POMC) 1665–6, 2790 peptides 2790 pituitary pars intermedia dysfunction 1666 testicular 1003 1,2-propanediol embryo cryopreservation 2890, 2892, 2900, 2903–5 toxicity to embryos 2917 propantheline bromide 847, 2840 propidium iodide (PI) acrosome evaluation 1495–6 sperm counting 1281–2 sperm viability testing 957, 1287, 1500, 1512–13, 2999–3000 propiomazine (propiopromazine) 1132 R ) Propionibacterium acnes (EqStim chronic endometritis 2613 maintenance of pregnancy 2335, 2474 older foal pneumonia 717 post-mating endometritis 2812–13 problem-breeding mares 2631 propofol 223, 847 proptosis 540 prostacyclins 1603 prostaglandin(s) (PG) 1603–4, 1642–5 delayed uterine clearance and 2604 endometrial production 2205–6 epididymal contractility 920, 1008 exogenous 1797–801 anovulatory follicles 1758 estrus synchronization 1731, 1799, 1871–2 induction of abortion 2320–1 induction of ejaculation 1414
induction of parturition 2265 luteolysis induction 1728–9, 1759, 1761, 1798–9 ovulation induction 1864 postpartum mares 2298, 2299 superovulation protocols 1839, 1840 uterine clearance 2602, 2628, 2630, 2660–1 maternal recognition of pregnancy see under prostaglandin F2α neonatal renal function and 646–7 ovulation 1642–3 oxytocin-stimulated release 1831–2 parturition 1645, 2228, 2269 placental production 89 placentitis 2592 pregnancy 2228 pyometra 2653 retained fetal membranes and 2523 selective oviductal transport 1643 sperm transport and 942 prostaglandin dehydrogenase (PGDH) 2224, 2592 prostaglandin E (PGE) cervical ripening 2726–9, 2811 epididymal contractility 1008 prostaglandin E1 (PGE1 ) cervical ripening 2726, 2729, 2811 gastroduodenal ulcers 396 induction of parturition 1801 see also misoprostol prostaglandin E2 (PGE2 ) embryonic release 2205 exogenous 1800–1 after insemination 2809 cervical ripening 2726, 2728–9, 2811 indications 1800–1 induction of abortion 2321 induction of parturition 29, 1801, 2264–5 oviductal blockage 1801, 1990, 2695 oviductal embryo transport 1643, 2187 ovulation 1643 parturition 1645, 2228, 2269 placentitis 2592 pregnancy 2228 sperm transport and 1800–1 uterine release 2589 prostaglandin F2α (PGF2α ) autumnal transition 1737 cyclical luteolysis 1638, 1642, 1643–4 embryonic release 1644, 2187, 2205 embryo transfer recipients 2878 epididymal function 1008 estrus suppression and 1848–9 exogenous 1797–800 after insemination 2809 anestrous mares 1696, 1700 delayed uterine clearance 2602 dosage and administration 1797–8 embryo donors 2878–9 estrus synchronization 1799, 1871–2 gonadotropin responses 1799 indications 1798–800 induction of abortion 1799–800, 2122, 2320–1 induction of parturition 1800, 2265 luteolysis induction 1761, 1798–9 ovulation induction 1643, 1860–1, 1864 post-mating endometritis 2615 postpartum mares 1816 superovulation protocols 1840, 1841
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
uterine infections 1800 see also cloprostenol; dinoprost maternal recognition of pregnancy 1644–5, 2200–2 conceptus mobility and 2202–3 oxytocin interaction 1644, 1680, 2204–7 ovulation 1643 parturition 1645, 2228 placentitis 2592 pregnancy 2228 prolactin and 1747 uterine release 2589 prostaglandin F metabolites (PGFM) 1642 pregnant mare 29, 2201 prostate gland 875, 877, 1113 dimensions 1380, 1482 ducts 1113, 1114, 1451–1 endoscopic features 1451–1, 1456 fluid 904, 1062 function 896–7 neoplasia 1113, 1484, 1485 ultrasonography 1481–2, 1483 protamines 927–8 protective response, dam to foal 118 protein(s) crude see crude protein dietary requirements breeding stallion 1230, 1232–3 broodmare 264 growing horses 786 sick neonate 267, 274 effects of cooling 1072, 1310 methods of studying 2829–30 parenteral nutrition solutions 274, 275 sperm content 1062 plasma membrane 1055, 1056 total plasma see total plasma protein tyrosine phosphorylation see tyrosine phosphorylation protein C 853 protein hormones 1602, 1603, 1606 protein kinase A (PKA) 1605 acrosome reaction 949 activated sperm motility 936 capacitation 947 protein kinase C (PKC) 1605 acrosome reaction 949–50 hyperactivated sperm motility 952 protein kinase G (PKG) 1605 proteinuria 655–6 proteoglycans, cervical ripening 2728 proteomics 962 Proteus, identification 1984, 2677, 2679–80 Proteus mirabilis, epididymitis 1158 prothrombin time (PT) 853 coagulation disorders 427, 428 hepatic disease 409, 410 proton pump inhibitors (PPIs) neonatal foal 395–6 older foal 737, 743 see also omeprazole protozoal disorders, foal diarrhea 390, 741 protozoal myeloencephalitis, equine (EPM) 730–1 diagnosis 731 vaccination 323 proximal interphalangeal joint see pastern joint proximal sesamoid bone (PSB) fractures 448
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Index pruritus, pinworm infections 299 Przewalski’s horse see Asian wild horse pseudohermaphroditism 2218–19 female 1111, 1136, 1961, 2218–19 male 1111, 1136, 1764, 1765, 2218 pseudohypoaldosteronism, foal 630 Pseudomonas identification 2676, 2677, 2679 uterine cytological smears 1925, 1926–7 Pseudomonas aeruginosa endometritis 1974, 1975, 2609 identification 1968–9, 1980–1, 1982 male external genitalia 1246, 1254 neonatal sepsis 255 pneumonia 710 prevention and control 1963 screening of mares 1964, 1972–3 serotyping 1971 stallion semen 1371 transmission via AI 3016, 3023, 3029 venereal transmission 1254 pseudopregnancy see corpus luteum, persistent pseudothrombocytopenia 373 psychological causes, anestrus 1697 psyllium 847 enteral feeding-associated diarrhea 272 intestinal clostridiosis 737 sand enteropathy 391, 744 Pten gene 2710 puberty effects on growth 283 female 1689–94 age at 1689–92 birth date and 1693 bodyweight and 1693 daylength and 1693 definitions 1690 foaling defining 1692 folliculogenesis 2012 social cues 1693–4 summarized data 1691 zebras 2853–5 male 902–4, 1015–23 age at 903, 1015, 1046, 1100 apoptosis of germ cells 1021–3 blood–testis barrier formation 1019–21 defined 1015 hormonal changes 902–3, 1005–7, 1016 onset of spermatogenesis 902–3, 1007, 1016–23, 1024 paracrine–autocrine factors 1007–8 Sertoli cell numbers 1016, 1042 zebras 2854, 2860–1 pudendal nerves damage, stallion 1132–3 mare 1595 pulmonary edema cardiovascular disease 500, 501 neonatal foal 141 pulmonary fibrosis, equine multinodular 2376 pulmonary hypertension neonatal foal 217, 590, 601 persistent (PPH) 590, 597 pulmonary vasculotropic infection, EHV-1 2380 pulmonic stenosis 508, 515 pulse oximetry 183–4, 187, 578 pulses, peripheral arterial 169 cardiovascular disease 499, 512 shock 138
pupillary light reflexes (PLR), foal 66, 535 critical care 171 neurological disorders 478, 480 pupillary membranes, persistent (PPMs) 547 pupils, neonatal foal 66, 535 purpura hemorrhagica 705, 765 pyelonephritis 652 pyocele 1477–8 pyometra 2652–4 causes 2652 cervical adhesion-related 2723 diagnosis 2652–3 hysterectomy 2574, 2654 hysteroscopy 1946, 1947 organisms 2652 post-hysterectomy stump 2574 prolonged luteal phase 1760, 2057, 2653 treatment 2654 ultrasonography 2137–8, 2140, 2653 pyospermia 1119–21, 1278–9 epididymitis 1158, 1159 examination of stallion 1119–21 suboptimal fertility 1245–6 treatment 1121 pyrantel pamoate ascarids 295, 756, 759 strongyles 298, 300 tapeworms 299 pyrantel tartrate, ascarids 759 pyrilamine maleate 241, 847 pyrimethamine/sulfadiazine 731, 847 pyruvate, antioxidant activity 941 quality standards, frozen semen 3012–13 quarantine broodmare farms 2748–9 cooled transported semen 1325 EHV control 725, 2384–5 imported live horses 2921 novice breeding stallions 1396 shuttle stallions 1187 sperm donor stallions 2972, 3030 Quarter Horses growth 285, 286 hereditary muscular disorders 463, 464–5 hyperelastosis cutis (HERDA) 566, 670, 678 hyperkalemic periodic paralysis 464, 673 seminal characteristics 1369–70, 1371, 1375 quinapril 847 quinestrol, male contraception 1888 quinidine sulfate 848 quotidian events 1670, 1771 rabies vaccination 313–14 broodmare 306, 310, 313–14, 2750 foals and weanlings 304, 309, 313, 314 stallion 1222 Rab proteins, acrosome reaction 950 racing performance, upper airway abnormalities and 798–9 radial immunodiffusion (RID) assay, plasma IgG 348 radial spokes, sperm 933–4, 1060 radiation, effects on spermatogenesis 1177
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radiography angular limb deformities 769 cervical vertebral compressive myelopathy 729 developmental orthopedic disease 768–9 fractures 446 infectious arthritis and osteomyelitis 461 musculoskeletal disorders 172 presale see presale radiographs radiation safety 801 septic arthritis/osteomyelitis 201, 458 see also abdominal radiography; barium radiography; thoracic radiography raniditine 848 critically ill neonate 194 neonatal foal 395, 396 older foal 685, 686, 737, 743 septic arthritis 460 RapIDTM tests 1980–1, 1983, 1986 R-banding, chromosomes 1464, 1953 reactive oxygen species (ROS) 939–41 adverse effects on sperm function 992–3 generation neutrophils in semen 940 by sperm 939–40, 991, 1311–12 uterus 940 induction of capacitation 992 mechanisms of sperm damage 940, 993 scavengers 940–1 reactions 940, 992 semen 941, 991 see also antioxidants triggering capacitation 947 see also oxidative stress rebreathing bag 712 receptors 1604–5 cell surface 1606 downregulation 1605 intracellular 1606 upregulation 1605 record keeping see documentation rectal endoscopy, mares 1940, 1949 rectal examination, digital mare 1904–13 abnormalities 1905–6 diagnosis of pregnancy 2246–7, 2248 dystocia 2484 general principles 1904–5 geriatric mares 2804–5 hydrops 2369 ovulation prediction 1721–2, 1911 prior to vaginal examination 1901 problem breeders 2623–4 record keeping 1905 rectal tears complicating 2578 safety and restraint 1906–8 supplies 1908 technique 1908–11 uterine torsion 2435–6 neonatal foal 170 stallion cryptorchidism 1103, 1150, 1532 seminal vesiculitis 1116 spermatic cord 1146 rectal liners, temporary indwelling 2581, 2582 rectal packing 2579, 2580 rectal prolapse, mare 1949 rectal strictures, mare 1949
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rectal tears, mare 2578–84 causes 2578 classification and locations 2578, 2579 diagnosis 1949, 2578–9 initial treatment 2579, 2580 loop-colostomy 2581–3 non-surgical treatment 2579–80 prognosis 2583–4 suture repair 2580–1 temporary indwelling rectal liner 2581, 2582 rectogenital pouch 1586, 1588 rectourethral fistula, foal 407 rectovaginal fistula/rectovestibular fistula (RVF) 2556–7 causes 2488–9, 2556 endoscopy 1945 foals with intestinal atresia 407 surgical correction 2556–7 recumbency increased, sick neonate 167, 168, 189 sedated foal 222–3 recumbency response 118 recurrent airway obstruction (RAO), equine 615, 617–19 genetic basis 618–19, 620 immunological basis 618 recurrent laryngeal neuropathy (RLN) 796, 797 red bag placenta 107–8, 2274 fescue toxicosis 2421 mare reproductive loss syndrome 2411 red blood cell alloantigens (or factors) 353, 354 red cell count (RBC) anemia 422 neonatal foal 173, 420 reference values 851 5α-reductase defects 2218 feto-placental unit 2223, 2224 redundant nuclear envelope (RNE) 928, 951–2 reefing see posthetomy, segmental reflexes, neonatal foal 66, 171 reflux esophagitis 684, 685 refraction artifacts, ultrasound 1919 refractometry colostrum 64, 342–3, 832–3 foal plasma 178, 348 RegumateTM see altrenogest regurgitation feed 750 milk 576, 589, 592, 593 reinforcement breeding 1247–8, 2785 history 1263 relative adrenal insufficiency (RAI), sick foals 524–5 relaxin 19, 1682–3, 2227–8 diagnosis of pregnancy 2254, 2256, 2257 parturition 2269 placentitis 2363 retained fetal membranes and 2523 relaxin-like factor (RLF) see insulin-like peptide 3 REM sleep, neonatal foal 477 renal blood flow autoregulation in neonates 647 neonatal vasogenic nephropathy 649 renal failure acute see acute renal failure chronic 653, 658, 765 renal failure index (RFI) 654–5
renal function birth transition 646–7 neonatal foal 66, 647–8 renal hemodynamics, birth transition 646–7 renal problems aged horses 2756, 2757, 2758 neonatal foal 646–58 causes 648–53 diagnosis 653–6 outcome and prognosis 657–8 perinatal asphyxia syndrome 151 therapy 656–7 older foal, weight loss 765 renal support, critically ill neonate 195–6 renal tubular acidosis (RTA) 650–1 renal tubular dysfunction 650–1 renin 78–9 renin–angiotensin–aldosterone system (RAAS) 70, 78–80 Repiquet (French veterinarian) 1262 reproductive efficiency 2779–87 factors affecting 2781–4 early pregnancy losses 2783–4 management 2781 mare age 2781 mare nutrition 2760 mare reproductive status 2781 method of service 2781–2 number of services 2782–3 overbreeding 1432 timing of service 2782 treatments 2783 measurement 1431, 2779–80 observed 2780–4 role of ultrasonography 2779 shuttle stallions 1185, 2784 stallion management 2784–7 AI 2786–7 natural service 2784–6 see also fertility; pregnancy rates reproductive epidemiology 2797 reproductive history geriatric mares 2804 mare 1897–9 stallion 1429–33 reproductive technology, future 3051–5 reproductive tract see genital tract repulsion, fetal 2479 reserpine 848 agalactia 2282–3 lactational anestrus 2637 residual bodies 1027, 1036, 1042 resolution, ultrasound 1915 respiratory acidosis 605 respiratory arrest, neonate 128 respiratory depression, foal 59 respiratory diseases equine herpesvirus see under equine herpesvirus(es) neonatal foal 586–90 anesthesia 224 diagnostic approach 575–80 genetic basis 615–20 older foal bacterial 688–93, 695–701, 704–9, 710–17 viral 720–6 see also pneumonia respiratory distress, neonatal foal 169 respiratory distress syndrome acute see acute lung injury/acute respiratory distress syndrome
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
neonatal equine (NERDS) 125, 583, 584, 589 respiratory failure, neonatal foal 215, 217 respiratory function assessment, critically ill neonate 184–5 neonatal foal 601 pregnant mare anesthesia and 56 premature and dysmature foals 125 respiratory quotient (RQ) 76 respiratory rate, neonatal foal 65, 575 anesthesia 225 birth resuscitation 131 critical care 169 respiratory rhythm, neonatal foal 575 respiratory stimulants neonatal foal 216–17 critical care 189, 195 periodic apnea 575 pneumonia 606–7 prior to/during transport 233 see also caffeine; doxapram respiratory support foal 189–90, 215–19 equipment 835 indications 215–16 prior to/during transport 233 see also oxygen therapy; respiratory stimulants; ventilation, mechanical respiratory system congenital abnormalities 609–14, 663–5 neonatal foal 66, 169–70, 601 entry of pathogens 199 immune function 338 respiratory tract endoscopy see upper airway endoscopy respiratory tract infections neonatal foal 586–8 diagnosis 577–8 genetic basis 615–16 older foal bacterial 710–17 viral 720–6 stallion, during shipping (shipping fever) 1189 see also pneumonia resting energy expenditure (REE), sick neonate 266 restraint, chemical see anesthesia; sedation restraint, physical mares aggressive dams 119 dystocia 2484 during live cover 1225–6, 1399 mount mares 1270 rectal palpation 1906–8 ultrasonography 1920 uterine and clitoral cultures 1965 vaginal examination 1900 stallions breeding 1392–3, 1397, 1398 examination of external genitalia 1458, 1469 retained fetal membranes (retained placenta) 2520–8 after cesarean section 2506–7, 2509 after dystocia 2494, 2513 causes 2521–3 clinical signs 2523–4 complications 2524, 2530, 2531, 2532 fescue toxicosis 2421, 2523 follow-up examination 2527 future fertility 2527
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Index hydrops 2371 hysteroscopy 1949 Miniature ponies 2844 nomenclature 2479 placental examination 2272–3, 2523 risk factors 2520 treatment 1833, 2524–8 ultrasonography 2148, 2156–7 uterine involution after 2296, 2526 rete testis adenomas 1168 anatomy 872, 873, 917, 925 role in spermatogenesis 1026, 1030 retina circadian system pathway 1672 colobomas 550–1 detachments 550, 551 dysplasia 550 examination in foals 537 hemorrhages 551 melatonin and 1669 retinohypothalamic tract (RHT) 1672, 1772 retractor penis muscle 877 retropharyngeal lymph node abscesses, strangles 704–5 reverse triiodothyronine (rT3) 526 rhabdomyolysis 465, 466 rhinopneumonitis, equine 2377, 2379 diagnosis 2383 see also equine herpesvirus(es) Rhizopus 711 rhodamine-123 staining, sperm mitochondria 957, 1289, 1513 rhodamine isothiocyanate (TRITC), acrosome evaluation 1494–5 Rhodococcus equi pneumonia 688–93, 695–701, 710 abdominal disease 243–4, 245, 695, 764 abdominal ultrasonography 243–4, 245, 698 clinical signs 695–6 determinants/risk factors 689–90 diagnosis 697–8 diarrhea 740 epidemiology 688–90 extrapulmonary disorders (EPD) 695–6, 740 genetic predisposition 615, 616 immune response 334, 336, 337, 696–7 liver disease 411, 412 pathogenesis 696–7, 724 prognosis 701 prophylaxis 240, 690–3 screening tests 692–3 thoracic radiography 580, 692, 697 thoracic ultrasonography 580, 692, 697–8 treatment 698–701 uveitis 548 virulence mechanisms 689, 696 weight loss 763 rib fractures, neonatal foal 454–5, 590, 598 birth resuscitation 129 clinical signs 139, 169, 172, 454, 598 diagnostic tests 170, 172, 454, 580, 598, 599 diaphragmatic hernia 244, 246 myocardial and pericardial trauma 515, 516 thoracic wall movement 576 treatment 113, 455, 598
riboflavin requirements, stallion 1230 Richter’s hernia 403, 641, 753 rifampin 848 Corynebacterium pseudotuberculosis 765 infectious hepatitis 412 Lawsonia intracellularis 739, 762 neonatal foal 194, 254, 260 pneumonia in older foal 716 R. equi infections 699, 700, 701, 764 right atrioventricular valve atresia 509 rimantadine 848 risk factor 2796 risus sardonicus 493 Ro-09-0198, fluorochrome-conjugated 956 Rocky Mountain horses, anterior segment dysgenesis 547 rolling parturient mare 2481, 2486–7 procedure, uterine torsion 2436–7, 2438–9, 2442 romifidine 848 foal 222 pregnant mare 57 ROS see reactive oxygen species rosette inhibition test (RIT) 2256, 2258 rotavirus control measures 741, 825 diarrhea neonate 159, 389 older foal 740–1 treatment 386, 389, 740–1 vaccination 322–3 broodmare 306, 308, 311, 322–3, 389, 741 foals and weanlings 295, 308 round ligaments bladder 632 liver 632 uterus 1586, 1588 Rspo1 gene 2214–15 runnel, cutaneous habronemiasis 1140 rusty load see ampullae, plugged Saber Delivery System 1880 Sable Island Horses, age of puberty 1692 Saccharomyces boulardii 272, 737, 848 sacrosciatic ligaments, prepartum relaxation 2270, 2480 S-adenosylmethionine (SAMe) 413, 841 St Anthony’s fire 2418 saline solutions 0.9% 141, 206, 207 neonatal foal 210, 230–1 0.45%, with 5% dextrose 206, 207 heparinized 843 hypertonic see hypertonic saline intrauterine infusion, to induce abortion 2321–2 sallenders 566, 567 salmeterol 606, 848 Salmonella carriers 738 control measures, NICU 825, 826 hyperimmune plasma 739 identification 1979 vaccines 386–7, 739 Salmonella abortus equi 3016 Salmonella newport 738 Salmonella typhimurium 738 salmonellosis (Salmonella infections) arthritis and osteomyelitis in foals 459, 462
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diagnostics 385, 738 diarrhea in older foals 738–9 neonatal diarrhea 385–6 neonatal sepsis 159, 253–4, 255 pneumonia 710 weight loss 763 salpinges see oviducts salpingitis 1989, 2693 salt intake, stallions 1230, 1234 salts, measurement and conversion 850 sand enteropathy 391, 743–4 ultrasonography 246, 744 weight loss 763 Sarcocystis neurona 730–1 vaccine 323 sarcoids mammary gland 2744 penis and prepuce 1169, 1451 schirrous cord 1561 schistocytes 422 sciatic reflex 479, 480 SCID see severe combined immunodeficiency scintigraphy, uterine 2597–8 scirrhous cord (septic funiculitis) 1153–4 sclerotherapy, hydrocele 1152 SCN4A gene 674 scoliosis 667 scotophase 1670 artificial light exposure during 1776–7 melatonin secretion 1675 scotorefractoriness 1347 Scott’s partial phallectomy 1568–9 scramblase 947 scrotal hernia 1147, 1164, 1540 see also inguinal hernia scrotal ligament 1532 cryptorchid castration 1534 scrotal width, total (TSW) age-related changes 1354, 1355 caliper measurement 1351, 1459 light-breed stallions 1369, 1370, 1371 miniature horses 1379 sperm production/output and 1351–2, 1353, 2785 stallions with one testis 1370, 1372 testicular degeneration 1371, 1373 ultrasonic measurement 1471–2 see also testicular size scrotum anatomy 867–9 edema after castration 1560 testicular trauma 1161, 1162 fetal, ultrasonography 48 fetal sex determination 2095–6 fluid accumulations, ultrasonography 1476–8 physical examination 1458–9, 1469 cryptorchidism 1532 temperature 1161 thermoregulatory function 881–2 seasonality of reproduction 1704–5 Asian wild horse 2848 donkey 2835 dual hemisphere breeding programs and 1186 mechanisms 1704–5, 1782–3 photoperiod and 1346–7 zebras 2852–3, 2859–60 see also annual reproductive rhythm; breeding season; non-breeding season
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Index
seasonal variations daily spermatozoal production (DSP) 890, 892 estrous cycle 1754 foal growth rates 282–3 foal heat 2296–7 follicle development and endocrinology 2027–8 hypothalamic–pituitary–testicular axis 1003–5 Leydig cells 1045, 1046 melatonin production 1675 Sertoli cells 1042–4 sperm freezability 2973 sperm motility 1294 sperm production and output 1344–50 stallion fertility 1431–2 testes 872, 873 testicular germ cell degeneration and renewal 891–2, 1039, 1040, 1040 twin pregnancies 2351 sebaceous glands, udder 1595–6 seborrhea, generalized primary 556–7 second messengers 1605–6 hormone signaling 1606–7 sedation exotic equids 2311 foal 221, 222–3 mechanical ventilation 218–19 percutaneous trocarization and intestinal decompression 383 plasma transfusion 350 tetanus 493 transportation 234 mare caudal reproductive tract surgery 2545–6 dystocia management 2485–6 embryo recovery 2875 embryo transfer 2877 fetotomy 2498 foal rejection 119 laparoscopic ovariectomy 1993, 2569 Miniature ponies 2840 oocyte collection 2937 oocyte transfer 2942 ovariectomy 2565 pregnancy 56–7 rectal palpation 1907–8 stallion behavioral problems 1417–18 castration 1557 cryptorchid castration 1533 genital tract endoscopy 1453 induction of ejaculation 1271, 1413–14 laparoscopic surgery 1547 physical examination 1458 during shipping 1188 testicular biopsy 1527 young horse, presale radiographs 802 segmented columns, sperm neck 930, 933, 1058–9 seizures, neonatal 171 hypoxic-ischemic encephalopathy 485, 486 management 148–9, 193, 194, 234, 486 selective IgM deficiency 362, 364, 374–5 selenium concentrations 466, 501 dietary deficiency 466–7, 515–16 requirements breeding stallion 1230, 1235
broodmare 2762, 2763, 2764, 2766 neonatal foal 265 supplements breeding stallion 1235 effects on fertility 994 nutritional myodegeneration 466, 467 self-mutilation, stallions 1412, 1424–5 semen (ejaculate) abnormalities 1119–27 Asian wild stallions 2847–8 bacteria in see under bacteria biochemical composition 1061–2 cellular debris 1302, 1304, 1305 centrifugation 1201–3 for cooling 1338, 1342 cushion technique 1203, 1338 for freezing 2964–6, 2973, 2974 collection 1268–73 artificial vagina 1268–70 for bacterial culture 1285 breeding phantom 1271 condom 1270 for cooled storage 1316–17 donkeys 2837 epididymal harvest 1272–3, 2983–4 equipment 1199, 1268–71 ground 1271, 1272, 1378 history 1263–4 intervals for AI 1360, 2973 manual stimulation for 1271, 1272 miniature stallions 1377–8 mount mares 1270–1 pharmacologically mediated 1272 stallion training 1394–5, 1399, 1400–1, 1402–6 cooled transported see cooled transported semen cooling see cooling, stallion semen cryopreservation see semen cryopreservation cytology 1285–6 hemospermia 1122–3 pyospermia 1120–1 density 1372–3, 1374, 1375 dismount drippings/samples monitoring 1227, 1245–6 reinforcement breeding 1247–8 donkey 2837 EAV detection 1253 EHV-1 1255–6, 2378, 3025 ejaculation see ejaculation emission 904, 1124, 1413 evaluation 955–60, 1278–89 advanced sperm testing 1287–9 as biopsy examination 1302–5 cooled semen programs 1316–17, 1319–21, 1325 freezing programs 2979–80, 2995–3003 hemospermia 1122–3, 1141 light-breed stallions 1367–8 miniature stallions 1378 pyospermia 1120–1 seminal vesiculitis 1115 urospermia 1125 fertility see fertility, stallion sperm filtration 1200, 1273 fractionation hemospermia 1123 pyospermia 1120 urospermia 1127 fractions 1062, 1452 pharmacological manipulation 1414
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
fresh, AI using see artificial insemination (AI), fresh semen frozen see frozen semen gel fraction 1120 ejaculation frequency and 1357 light-breed stallions 1369, 1370, 1371, 1375 pharmacological manipulation 1414 removal 1273 separation methods 1200 stallion age and 1355 gel-free volume 1279 ejaculation frequency and 1357 light-breed stallions 1369, 1370, 1371, 1375 miniature stallions 1378 photoperiod effects 1348, 1349 seasonal variations 1344, 1345 stallion age and 1355 stallions with one testis 1370, 1372 gross appearance 1278–9 handling 1273 cooled semen programs 1317, 1326 historical perspective 1264–5 immune tolerance in females 1096–7 insemination dose see insemination dose live-dead staining of smears 1287 mass activity 1372–3, 1374, 1375 microbiology 1285–6 pelleted 1265 pH 1062, 1285 light-breed stallions 1368, 1369, 1370, 1371 stallions with one testis 1370, 1372 presperm fraction 1120 processing for AI 1200–1, 1317–19 for cooled transport 1317–19 donkeys 2837–8 EAV shedders 1252 for freezing 2973–4 for GIFT 2946 history 1264–5 for ICSI 2949–50 for sperm morphological analysis 1298–9 quality parameters, Warmblood, Friesian and draft horse stallions 1364–6 reactive oxygen species scavengers 941, 991 sperm concentration for AI 1201 cooled shipped semen 1319–21, 1326 determination 1201, 1202, 1279–82, 2995 ejaculation frequency and 1357, 1358 epididymal sperm cryopreservation 2985 frozen semen 2976, 2995 light-breed stallions 1369, 1370, 1375 miniature stallions 1378 pharmacological manipulation 1414 stallion age and 1355 stallions with one testis 1370, 1372 sperm numbers ejaculation frequency and 1357, 1358 enumeration 1201, 1202, 1279–82 frozen semen 2995 insemination dose 1273–4 light-breed stallions 1368, 1369, 1370, 1375
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Index for maximum reproductive efficiency 1185 miniature stallions 1378 seasonal variations 1345–6 stallion age and 1355–6 Warmbloods, Friesians and draft horses 1364, 1365–6 sperm-poor fraction 1120 sperm-rich fraction 1120 uterine deposition 1092–3 volume for AI 1201, 1274 ejaculation frequency and 1357, 1358 light-breed stallions 1369, 1370, 1371, 1375 measurement 1279 stallion age and 1355 stallions with one testis 1370, 1372 see also semen; seminal plasma; spermatozoa semen cryopreservation 2972–81 alternative techniques 2978–9 Asian wild horse 2849 cooling and freezing rates 2961, 2975, 2976–8 current freezing protocols 2974–6 DNA damage after 993 epididymal sperm 1160, 2983–6 freezing protocols 2985 harvesting procedure 2983–5 for ICSI 2985–6 for insemination 2985, 3039–40 historical perspectives 1264–5 principles 2962–3 sperm processing 2973–4 stallion management 2972–3 urospermia 1127 see also frozen semen semen extenders 1200–1, 1273, 2964–70 AI of problem mares 2144 antibiotics 1201, 1341, 3018 cooled semen 1318, 1338–9, 1341–2 frozen semen 2970 historical development 1264 problem mares 2144, 2145 sperm motility effects 1295, 1338, 1339 antioxidants 1339 cooled semen 1201, 1312–13, 1318 antioxidants 994 composition 1337–8 control of bacterial growth 1338–9 dilution ratio 1318–19, 1326 Europe 1336–9 large-scale programs 1324 neutralizing metabolites 1339 North America 1341–2 osmolarity 1336 pH 1064, 1336 protective effects 1074 donkey semen 2837–8 epididymal sperm 2984, 2985 fresh semen 1317 frozen semen 2966–70 antioxidants 993, 994 composition 2965–6, 2967–70 freezing protocols 2974, 2975 sex-sorted sperm 3046 hemospermia 1124, 1141 historical development 1264 potential pathogens 3030 pre-breeding intrauterine infusion hemospermia 1124
problem mares 2144, 2145 suboptimal stallion fertility 1246–7 prefreeze/centrifugation 2964–6, 2974, 2975 reinforcement breeding 1247 sperm motility effects 1295 sperm motility evaluation 958, 1283, 1284, 1293 urospermia 1127 uterine inflammation and 1094, 1096 seminal colliculus (colliculus seminalis) 1113, 1114 endoscopy 1116, 1117, 1452, 1453, 1455 obstruction 1484 seminal plasma 1092 added to epididymal sperm 2985 antioxidant activity 940, 941, 991, 1077 effects on sperm 1054 harmful effects on sperm 2964 IGF-1 1000 nutrient content 938 production 897 sperm motility 935 sperm transport and 942–3, 2599 steroids and proteins, infertility 1438 transferrin 1000, 1001 uterine inflammation and 1094–6, 2614–15, 2655–6 seminal plasma proteins, equine (SP1 and SP2) 1088 seminal vesicles (vesicular glands) 875, 877, 1113 antimicrobial drug instillation 1116, 1117 dimensions 1380, 1482 ducts 1113, 1114, 1452 fluid collection 1115–16, 1120 secretion 1062 function 896–7 infections see seminal vesiculitis surgical removal 1116–17 ultrasonography 1481, 1482 seminal vesiculitis 1115–17 causative agents 1115 diagnosis 1115–16, 1120 disease transmission potential 3024 endoscopy 1116, 1117, 1455–5 minimum contamination breeding techniques 1117 treatment 1116–17 ultrasonography 1484 seminiferous epithelium 869, 870, 912–13 cycle of see spermatogenic cycle spermatogenesis 883 seminiferous tubules 869, 870, 871–2, 911, 912, 913 age-related changes 871 convoluted 871, 873 lumen scores (LS) 1017, 1019–21 rete 872, 873, 925 spermatogenesis 883, 911–12 initiation at puberty 1017–19 spermatogenic wave 916, 924 straight 872, 873 seminomas, testicular 1163, 1166–7 senescence, spermatozoa 1074, 1075–7 sepsis, neonatal 136, 137, 154–61, 198 acute renal failure 652 calcium dysregulation 526 causative organisms 156, 199 clinical signs and diagnosis 139, 154–6, 199–202 coagulation disorders 160, 429, 517
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
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diarrhea 154–5, 159, 392 focal infection and sequelae 158–60, 200–2 hypothalamic–pituitary–adrenal axis 155, 525 monitoring 157–8 nutrient requirements 266, 267 pathophysiology 138 preparation for transport 234–5 prevention 161 prognosis/outcomes 160–1, 203 risk factors 154, 198 routes of infection 154 thrombocytopenia 373 treatment 156–8, 202–3, 209, 253–5 sepsis scoring systems, neonatal foal 156, 199–200 septic arthritis (infectious arthritis), foal 457–62 categories 457 clinical signs 457 complicating bacteremia/sepsis 159–60, 201, 457, 459 diagnostic evaluation 458–9 monitoring and prognosis 461–2 pathogenesis 438, 457 R. equi 695, 700–1 treatment 203, 459–61 septicemia, neonatal 198, 200 endocrine changes 72 fescue toxicosis 2421, 2422 pneumonia 601–2 thrombocytopenia 373 septic funiculitis 1153–4 septic shock defined 154 neonatal foal 144–5, 202–3, 213 see also sepsis R 344, 349 Seramune Sergre–Silberberg (SS) effect 1282 serial radial immunodiffusion (SRID) assay, colostral IgG 832 serotonin (5-HT) receptors, ergot alkaloid interactions 2418 Serratia marcesens 2415 serration of granulosa 2036–7, 2039, 2040, 2041 blood flow changes and 2046, 2047 fertility and 2050 Sertoli cells 869, 870, 871 age-related changes 871 blood–testis barrier formation 1019–21 drugs adversely affecting 1177 dysfunction, infertility 1437–7 embryonic development 1015, 2212–14 functions 883–4 germ cell associations 884, 918 germ-cell ratio, histometric analysis 1524 hormonal regulation 899–900 Leydig cell interactions 885 neoplasms 1167–8 number per testis 871, 885, 1040, 1041 age-related changes 1042, 1043 seasonal effects 872, 873, 1042–4 paracrine–autocrine factors 999–1003 proliferation 871 pubertal development 1007, 1016, 1042 regulation of function 884–5 role in spermatogenesis 883–5, 1040–4 secretions 884, 898 spermatid attachment 884, 1028, 1036
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Index
Sertoli cells (Cont.) thyroid hormone actions 998 tight junctional complexes 884, 912, 915, 1019, 1041–2 serum autologous, corneal ulcers 546, 848 infusions, donkey-in-horse pregnancies 2315, 2317 see also plasma serum amyloid A (SAA) 333, 851 serum osmolarity see plasma osmolality service(s) methods, reproductive efficiency and 2781–2 per cycle 1431, 2780, 2782–3 per pregnancy 2780 timing 2782 see also artificial insemination; natural service sesamoid bones, ossification 437 severe combined immunodeficiency (SCID) 362, 369–71 clinical pathology 367, 370 clinical signs 369–70 genetic basis 369, 2829 necropsy findings 370, 371 sevoflurane anesthesia 58, 224 R protocol 3045 Sexcess sex chromosomes 1952, 2211 abnormalities 1951, 2216–17 mares 1756, 1956–7 mosaics 2216, 2217 stallions 1465 C-banding 1463, 1464, 1953, 1954 see also X chromosome; Y chromosome sex cords 2214 cortical 2214 male 1046, 2214 sex cord-stromal tumors, mixed 1168 sex determination chromosomal 1078, 2211 gonadal 2212–15 abnormalities 2217–18 prenatal see fetal sex determination sex differentiation 2212–15 sex-reversal syndromes 1465, 1960–1, 2216, 2217–18 sex-sorted stallion spermatozoa 3042–6, 3052 factors influencing sortability 3044–5 fertility 3045–6 historical background 1077–9 method of FACS separation 3043–4 methods of sex preselection 3042 principles of FACS separation 3042–3 sexual behavior jack donkey 2837 mare abnormalities 1761–5 mount mares 1270–1 in natural conditions 1388 slow-starting novice stallions 1409 see also estrous behavior play 1386, 1389 stallion see stallion sexual behavior zebra mare 2852 see also copulation sexual development 1959–60, 2011, 2211–15 disorders of 1959–61, 2215–19 sex reversal 2217–18 terminology 2215–16 see also intersex conditions
phenotypic 2212, 2215 abnormalities 2218–19 sexual interest and arousal see libido shaker foal syndrome 320, 492 Shetland ponies 1377 seminal characteristics 1373, 1375 see also Miniature mares; Miniature stallions shipping, care of stallion during 1187–9 shipping containers cooled semen 1264, 1318, 1324–5, 1330–4 cooling rate 1331 disposable 1333–4 reusable 1331–3 frozen embryos 2923 frozen semen 3010–12 shipping fever 1189 shipping regulations embryos 2922 frozen semen 2972, 3013, 3031–5 stallions 1186–7 shivering thermogenesis 75 shock mare hydrops 2370, 2371 periparturient hemorrhage 2518 neonatal foal 136–45 clinical signs and diagnosis 138–40 complications 145 etiologies 136, 137 monitoring response to therapy 145 pathophysiology 137–8 premature or dysmature foals 125 therapeutic management 140–5 see also hemorrhagic shock; septic shock shoes, corrective angular limb deformities 666, 769–70 flexor tendon laxity 469–70 flexural deformities 443, 444 short colon syndrome 407 shoulder dysplasia 667 shoulder flexion posture, fetus 2491 fetotomy 2501 shoulders, fetal, normal delivery 2271, 2482 shuttle stallions 1184–90 artificial lighting programs 1005, 1186, 1350 care during shipping 1187–9 care on arrival 1189–90 fertility 1185, 1350, 1429 reproductive efficiency 1185, 2784 selection 1184–6 shipping regulations 1186–7 SI conversion tables 850, 857–61 siderophages (siderocytes) endometrial 1932, 1935 involuting uterus 2292, 2294 signal transduction 1606 sildenafil, neonatal foal 195, 217 silver dollar plaques 3024 silver sulfadiazine 848 Taylorella equigenitalis carriers 1251, 2406 simulation of breeding 1719 single nucleotide polymorphisms (SNPs) 2824–5, 2829 sinuses, paranasal see paranasal sinuses sinus terminalis 2188–9, 2190, 2191 SIRS see systemic inflammatory response syndrome
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
63,X karyotype see X chromosome monosomy 64,X,delXp mares 1957 64,X,i(Xq) mares 1957 64,XX,delXq/64,XY mares 1957 64,XX sex reversal 1961, 2217 64,XY sex reversal see XY sex reversal 65,XXX mares see trisomy X 65,XXY stallions see XXY syndrome 66,XXXY stallions 1465, 2216 skim milk semen extenders cooled semen 1341 frozen semen 2967–8 historical development 1264 see also KenneyTM semen extender; milk-based semen extenders skin abrasions, sick neonate 167, 168 skin biopsy, cytogenetic analysis 1953 skin diseases 555–70 acquired 555–6 congenital 555, 670 diagnosis 556 hereditary 555 neonatal foal 556–63 primary, neonatal foal 556–61 primary congenital/hereditary, older foals/adults 563–70 secondary, neonatal foal 561–3 skin infections 555 skin injuries, traumatic 555 skull fractures 454, 488–9 slap test 479 sleep, neonatal foal 477 slipped mares 1429, 2780 slow breeders 1407–10 dual hemisphere programs 1185–6 experienced stallions 1410 management 1209 slow-starting novice stallions 1407–10 small colon meconium impaction 244, 246, 380–1, 402 obstruction, older foal 749–50, 753–5 small intestine disaccharidase enzymes 264–5 intussusception 244, 245, 402, 755 obstruction meconium impaction 244, 246, 380 neonatal foal 170–1, 244, 245 older foal 755–6 strangulated umbilical hernia 753, 754 vaginal ring palpation 1146 perforation, ascarid infections 295 volvulus 244, 401–2, 755 small stature see dwarfism smegma mare 1593 stallion 1137, 1169 α-SNAP 950, 951 R Foal IgG Test, IDEXX 347, 348, SNAP 834 SNARE proteins 950, 951 sneak breeding 1386 social cues, affecting female puberty 1693–4 sodium (Na) abnormalities see hypernatremia; hyponatremia balance, neonatal foal 655 excretion, neonatal foal 80, 648, 654, 655 fractional excretion (F x Na ) 654, 855
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Color: 1C Trim: 279mm X 216mm
Printer Name: Yet to Come
Index management, neonatal renal disease 657 pre-foaling mammary secretion 2733, 2735 assay methods 830–1 feto-placental monitoring 19–20 jennies 2270 predicting parturition 2270 requirements breeding stallion 1230, 1234 broodmare 2762, 2763, 2766 restriction, neonatal foal 192 serum 173, 182, 853 sodium bicarbonate see bicarbonate sodium chloride (NaCl) dietary intake, stallions 1230, 1234 measurement and conversion 850 solutions see saline solutions sodium iodide 848 sodium polystyrene sulfonate 848 sodium/potassium (Na+ /K+ ) exchangers, sperm motility 935 sodium-potassium pump (Na,K-ATPase), sperm motility 935 soluble NSF attachment protein (α-SNAP) 950, 951 solutes, diffusion of 2959–60 solutions diffusion of solutes in 2959–60 freezing, physical properties 2960–1 somatotropin see growth hormone somite formation 2179, 2180 sorbitol, metabolism in sperm 938 sorbitol dehydrogenase (SDH) 174, 409, 410 reference values 852 Sox9 gene 2214, 2215 SOX9 protein 2215 SOX-p gene 1960 soybean lecithin-based semen extender 1338, 2968–9 Spallanzani, Lazzaro 1262 specular reflections, ultrasound 1919, 1920 speculum, vaginal see vaginal speculum sperm see spermatozoa SpermaCue 1280 spermadhesins 1088, 1096 spermatic cord abnormalities 1145–54 diagnostic tests 1146 history and examination 1146 anatomy 872, 874, 1145–6 infection 1153–4 physical examination 1459 torsion 1151, 1163–4 ultrasonography 1474, 1479, 1480 ultrasonography 1146, 1470, 1472 spermatids 870, 883, 914–15, 1034–6 cap phase 1034, 1035, 1036 elongation phase 1034, 1035, 1036 Golgi phase 1028, 1034–5 maturation phase 1034, 1036 Sertoli cell attachments 884, 1028, 1036 stages of cellular association 886–7, 888, 919 spermatocytes meiosis 883, 914, 1026, 1029, 1032–4 primary 870, 883, 1032 cellular associations 886, 887, 888 meiosis 1032 secondary 883, 1032–4 spermatocytogenesis 883, 914, 1026, 1028–32
spermatogenesis 882–94, 911–16, 1026–49 arrested 1521, 2304 control of 1047–8 cryptorchid testes 1100–1 drugs adversely affecting 1175–82 duration 883, 889–90, 1037–8 efficiency 1049 establishment 1042, 1043, 1046–7 evaluation 1039 factors affecting 1344–60 germ cell degeneration 890–2, 1039, 1040, 1040 GnRH vaccination and 1195–6 hormonal control 900, 996–9, 1047–8 initiation at puberty 902–3, 1007, 1016–23, 1024 apoptosis of germ cells 1021–3 histological development 1016–17 kinetics 920, 1036–9 Leydig cell function 892–4, 1044–6 paracrine–autocrine control 999–1003 seasonal changes 872, 873 Sertoli cell function 883–5, 1040–4 species comparisons 885 thermal sensitivity 881–2, 1177 three major phases 1026, 1028–36 spermatogenic cycle (cycle of the seminiferous epithelium) 885–9, 915–16, 1036–9 duration 887–9, 1037–8 stages 885–7, 1036–7 description 886, 920 microscopy 886–7, 888, 921, 922, 923, 1037 relative frequencies 1038 time to appearance of sperm in ejaculate 1048, 1049 spermatogenic wave 916, 924, 1038–9 spermatogonia 870, 883, 914, 1028–32 committed 1029 degeneration/apoptosis 890–2, 916, 924, 1021, 1039 proliferation/renewal 891–2, 914, 1030–1, 1033 at puberty 1016 reserve 891 seasonal changes 1030, 1032 stages of cellular association 886 stem cell 1029 subtypes 1029–30, 1032, 1033 transplantation 3054, 3055 spermatozoa 909–62 acrosome see acrosome aging and senescence 1074, 1075–7 ampullary accumulation, ultrasonography 1483–4 ampullary occlusion see ampullae, plugged antibodies 1524 Asian wild stallions 2847–8 biochemical composition 1062 biochemical markers of function 958 capacitation see capacitation changes during development 1053–4 chemotaxis within oviduct 952 cold shock 1070, 1073–4, 1311 compartmentalized function 1064–5 computer-assisted analysis see computer-assisted sperm analysis concentration in semen see semen, sperm concentration cooling see cooling, stallion semen
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
3119
cooling-induced damage see cooled transported semen, sperm damage cryodamage 2961 cryopreservation see semen cryopreservation daily output see daily sperm output daily production see daily sperm production DNA see spermatozoal DNA donkey 2837 dose for insemination see insemination dose epididymal cryopreservation see under semen cryopreservation harvesting 1160, 1271–2, 2983–5 epididymal maturation 894, 920–5, 930 histomorphology 922, 927, 928, 929 regulation 921–4 epididymal transit 896, 918–20, 1047, 1048, 1157 factors regulating 1008 evaluation tests 955–60, 1287–9, 1297, 2996–9 factors affecting production and output 1344–60 age 1353–6 ejaculation frequency 1356–60 season 1344–50 testicular size 1350–3 fertility see fertility, stallion sperm flagellum see spermatozoal tail fluorescent live-dead staining 957–8, 1287 freezability 2972–3, 2994 freeze-dried, for ICSI 2948 freezing see semen cryopreservation frozen see frozen semen function 910, 1053–79 for GIFT 2945–6 harmful effects of seminal plasma 2964 head see spermatozoal head for ICSI 2948 immobilization, for ICSI 2949, 2950 injection, ICSI 2949, 2950 length 926 longevity, stallion fertility and 1185 membrane lipid peroxidation see lipid peroxidation (LPO), sperm membranes metabolism 937–9, 1062–5 aged sperm 1077 capacitation 1075 effects of cooling 1073, 1311–12 morphology see spermatozoal morphology motility see spermatozoal motility neck 1057, 1058–9 numbers in semen see semen (ejaculate), sperm numbers origin 910–16 oviduct interactions see sperm–oviduct interactions oxidative stress 939–41, 991–4 physiology 935–55, 1061–5 plasma membrane see sperm plasma membrane processing see semen, processing reactive oxygen species and see under reactive oxygen species reserves, stallion age and 1353–4 reservoir in mare 944 sex sorting 1077–9, 3042–6
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spermatozoa (Cont.) storage within excurrent ducts 895–6, 918, 921 structure see spermatozoal morphology tail see spermatozoal tail thermal stress 1070–4 transport 941–4, 1085, 1093, 3036–7 geriatric mares 2809 PGE2 therapy and 1800–1 uterine contractions 941, 942, 1093, 2598–9 urinary passage 896 uterine interactions see sperm–uterine interactions viability tests see spermatozoal viability tests see also semen spermatozoal DNA (or chromatin) damage, early embryonic loss and 2332 effects of storage 993 evaluation of integrity 955–6, 1506–7 frozen-thawed sperm 3001–2 oxidative damage 940, 993 sex sorting 1078, 3042–3 spermatozoal head(s) anatomical features 925–6, 931, 1057–8, 1303 attachment to tail 930, 1058 computer-assisted morphometry 2998 cytosolic compartment 929–30 detached (tailless) 1301 length 926 multiple 1300 ultrastructure 927–30, 932, 933, 934 spermatozoal morphology 1297–305 analysis 959–60, 1285, 1297–305 cooled shipped semen 1319 data collection sheet 1299, 1302 electron microscopy 1299–302, 1303–4 frozen-thawed semen 2997–8 light microscopy 1297–8, 1299 semen sample processing 1298–9 anatomical features 925–35, 1026–8, 1031, 1054–61 cooled stored semen 1312, 1314 defects 1299, 1300–1 classification 960, 961, 1297 fertility and 960, 1285, 1375–6 light-breed stallions 1373, 1374 motility and 960 multiple 1297, 1298 reactive oxygen species generation and 940, 991 percent normal light-breed stallions 1368, 1369, 1370, 1371 miniature stallions 1378 stallions with one testis 1370, 1372 Warmbloods, Friesians and draft horses 1364, 1365 spermatozoal motility 935–7, 1065–70, 1292–5 activated 935–7, 1292 age of stallion and 1355 amplitude of lateral head displacement (ALH) 1284 analysis 958–9, 1282–5, 1293–4 cooled shipped semen 1319 factors affecting 1067–70 objective see computer-assisted sperm analysis (CASA)
reasons for 1292 subjective (visual) 1283, 1293 circular (CIR) 1066 clinical relevance 1070 cooled shipped semen 1321 ejaculation frequency and 1357, 1358, 1360 energy substrates 938 epididymal compartments 894–5, 920 factors affecting 1294–5 fertility and 959, 1282–3, 1295 frozen-thawed semen acceptable thresholds 1282, 2979–80 by-eye estimation 2996 computer-assisted analysis 2996–7 hyperactivated 951–2, 1074, 1292 biochemistry 1075 capacitation and 944, 952 signaling pathways 936, 951–2 lateral head displacement (LHD) 1066 longevity 1286 mechanics of doublet sliding 937, 1065–6 miniature stallions 1378 morphological abnormalities and 960 nomenclature 1066 origin 1292 patterns 1066–70 percentage (MOT) 959, 1066 light-breed stallions 1368, 1369, 1370, 1371, 1375 stallions with one testis 1370, 1372 Warmbloods, Friesians and draft horses 1364, 1365 photoperiod effects 1348–9 progressive 1283, 1284 percentage (PMOT) 959, 1066 reactive oxygen species and 992–3 regulation 896–7, 935 semen cryopreservation threshold 2973 semen storage temperature and 1330 sex-sorted sperm 3044 sperm viability and 1511 structures responsible for 1292–3 temperature effects 1068–9, 1073 total 1283, 1284 types 1292 urospermia 1125, 1127 velocity see spermatozoal velocity wobble 1066 spermatozoal tail (flagellum) 926, 931, 932, 1026, 1028 abaxial 1058 attachment to head 930, 1058 development 917, 918, 1028, 1035, 1036 mechanism of bending 937, 1065–6, 1292–3 stump defects 1301 ultrastructure 931–5 spermatozoal velocity 1283 average path (VAP) 1066, 1284 computer-assisted analysis 1284 curvilinear (VCL) 959, 1066, 1284 linearity (LIN) 1066, 1284 straight line (VSL) 1066, 1284 straightness (STR) 1066, 1284 spermatozoal viability tests (membrane integrity tests) 957–8, 1499–500, 1510–13
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
fluorescence methods 1287, 1499–500, 1511–13 frozen-thawed semen 2999–3000 non-fluorescence methods (vital staining) 1287, 1499, 1511 sperm chromatin see spermatozoal DNA sperm chromatin dispersion (SCD) assay 956 sperm chromatin structure assay (SCSA) 955–6, 1289, 1506–9 cooled transported semen 1507–8 embryo quality and 1508 fertility prediction and 1508 frozen-thawed semen 3001–2 interpretation of results 1507 sperm granulomas 1159 spermiation 1036 Sertoli cell function 884, 1036 spermiogenesis 883, 914–15, 1026, 1029, 1034–6 cap phase 1034, 1035, 1036 elongation phase 1034, 1035, 1036 Golgi phase 1034–5 maturation phase 1034, 1036 Sertoli cell function 884 spermiostasis, ampullar 919, 1113–15 see also ampullae, plugged sperm–oocyte binding test 1287–8 sperm–oocyte interactions 952–5 evaluation 1502–3 sperm–oviduct interactions 944, 1085–9 characterization 1085–7 clinical implications 1089 evaluation 1501 molecular basis 1087–8 sperm release 944, 1088–9 sperm plasma membrane 926, 933, 1054–7, 1498 abnormalities 1300 aged sperm 1077 capacitation-related changes 946–7, 1075 evaluation 1501–2 effects of cooling 1073, 1311 evaluation 1498–504 glucose transport 1063 integrity tests see spermatozoal viability tests osmoregulation, testing see hypo-osmotic swelling (HOS) test regional domains 1056–7, 1064, 1498, 1499, 1500 sperm–uterine interactions 1092–7 selection and reservoirs of sperm 1093–4 uterine inflammation 1094–6 sphectophotometric methods, sperm counting 1280 spherocytes 422, 423 spina bifida 670 spinal cord circadian system pathway 1672 compression, radiographic indicators 729–30 congenital abnormalities 669, 670 injuries 488–9 spinal reflexes, long 478–9 spinosad 2416 spirochetes, in semen 1302 spirurids 294 spleen immune function 334 ultrasonography 249
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Index splinting common digital extensor tendon rupture 472 flexor tendon laxity 470 flexural limb deformities 442, 444 spring transition see vernal transition squamous cell carcinoma (SCC) mammary gland 2743–4 penis and prepuce 1136, 1137–9, 1169 squamous papillomas, penis and prepuce 1136–7 squiffy face see wry nose SRY (or Sry) gene male sexual differentiation 1078, 1960, 2211, 2214 negative XY sex reversal 1960–1, 2217 positive XY sex reversal 1961, 2217–18 testing 1464 stallion age shuttle stallions 1185 sperm production/output and 1353–6 sperm reserves 1353–4 testis size and 868–9, 871, 1018, 1043, 1046, 1354–6 see also geriatric stallions bachelor 1386 behavior see behavior, stallion book of mares see book of mares breeding availability and estrus synchronization 1870–1 client relations 1209–11 early embryonic loss and 2332 exercise programs 1211–21, 1228–9 handling see stallion handling management programs 1208–27 novice 1396–401 nutrition 1221, 1228, 1229–39 pre-season evaluation 2784–5 preventive medicine 1221–2, 1396 selection 1208 syndication 1208–9 training see training, breeding stallion see also natural service breeding records 1211, 1212–17, 1218–19 choice, geriatric mares 2807 communication with 1391 contraception 1888–9 equine arteritis virus carriers see under equine arteritis virus harem see harem stallions internal genitalia see male internal genitalia interspecies matings 2303 number of mares bred see book of mares puberty 902–4, 1015–23 reproductive history 1429–33 reproductive parameters 1362–6, 1368–76, 1378–80 semen donor 2972–3 cooled transported semen programs 1325–6 freezability of sperm 2972–3, 2994 infectious disease control 3016, 3030 potential pathogens 3015–16, 3029–30 retained semen rights after sale 3014 see also artificial insemination
sexual maturity 1015 shuttle see shuttle stallions slow breeders see slow breeders zebra 2859–61 stallion fertility age-related changes 1048 breeding frequency and 1243, 1244, 1245, 1432–3 dietary antioxidant supplements and 994 draft horses, Friesians and Warmbloods 1362–3 EHV-1 infection 2380 end points for describing 1429–31 factors affecting 1241 historical information 1429–33 interpretation of data 2779 measures of 1184–5, 1430–1 morphological sperm defects and 1375–6 non-stallion factors 1429, 1431–3 at puberty 903 seasonal variations 1431–2 semen characteristics and 1372, 1374, 1375–6 semen evaluation and 1278 shuttle stallions 1185, 1350, 1429 sperm chromatin structure assay and 1507, 1508 sperm motility and 959, 1282–3, 1295 testicular temperature and 881–2 see also reproductive efficiency; stallion subfertility/infertility stallion handlers 1226 communication with horse 1391 education 1410 stallion handling 1391–5 breeding shed exposure 1393–5, 1398–401 errors 1410–11 first interaction 1391–2 novice stallion 1396–401 phantom use 1402–6 preparatory training 1393, 1397–8 restraint 1392 stallion-like behavior mares 1763–5, 2709, 2712 pharmacological suppression 1194, 1416–17 see also aggressive behavior stallion sexual behavior abnormal 1407–12 development 1389 domestic breeding conditions 1389–90 ejaculation frequency and 1357 factors affecting 1390 free-running populations 1385, 1388, 1389 normal 1385–90 overenthusiastic or unruly 1410–11 photoperiod effects 1349–50 shuttle stallions 1185–6 suppression 1191–4, 1416–18 behavioral training 1191 immunocastration 1192–3, 1417 nutritional supplements 1417 pharmacological 1194, 1416–18 surgical castration 1192 zebras 2859 stallion subfertility/infertility acrosome dysfunction 1493–4 causes 1429, 1435
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chromosome abnormalities 1464–7 classification of causes 1523 cooled transported semen programs 1325–6 cryptorchidism 1149 cytogenetic analysis 1462–7 diagnostic evaluation 1436–41, 1523–5 endocrine–paracrine–autocrine approach 1435–43 endocrine profiles 1436–6, 1439, 1523–4 equine viral arteritis 2393 genetic factors 2828–9 hemospermia 1122 hydrocele 1152 idiopathic 1435–43 diagnostic tests 1438–41 hormonal imbalance 1435–5 hormone therapy 1442–3 paracrine–autocrine malfunction 1437–7, 1439 primary endocrine malfunction 1436–6 transient 1240–8 economic impact 1240, 1241 evaluation 1175, 1176, 1241–3 management strategies 1244–8 urospermia 1125 varicocele 1153 see also fertility; stallion fertility stalls, lighting systems 1775, 1776 stall-walking 2774 Standardbreds accessory gland dimensions 1380 cooled transported semen programs 1323 osteochondritis dissecans 809 ovulation induction 1860, 1861 seminal characteristics 1373, 1375 twin pregnancies 2351 standing, normal newborn foal 63, 477 stanozolol 848, 1179–80 staphylectomy 589, 611 Staphylococcus aureus acute endometritis 1974, 1975–6 gynecological samples 1973 identification 1969, 1979, 1984, 2678 Staphylococcus spp. identification 1981–4, 1985, 2676, 2677, 2678 neonatal sepsis 156, 254, 255 pneumonia 710 starch granule test, oviductal patency 1988–9, 1990, 2694 stars of Winslow 537 steam cleaning, endoscopes 1450 stearic acid, sperm content 1062 stem cells, induced pluripotent 3053, 3054 stereotypic behavior 1423–5, 2771–5 control measures 2772–3 function 2772 sterilization embryo transfer equipment 2874 endoscopy equipment 1449–9 sternal recumbency, neonatal foal enteral feeding 270 NICU 824 normal behavior 63, 64, 477 pneumonia 586 prematurity or dysmaturity 125 respiratory support 216 sepsis 158
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steroid hormones 1602–3 biosynthesis see steroidogenesis contraceptive use 1888–9 fetal production 2230 mechanism of action 1606 receptors 1606 testicular secretion 898 transport 1604 see also adrenal cortical hormones; androgen(s); estrogen(s); specific hormones steroidogenesis 1602 adrenal cortex 1665 fetal gonads and placenta 1633–4 fetal/neonatal adrenal cortex 69–71, 122–3 feto-placental unit 2222–4, 2225–7 ovarian 1631, 1632 autumnal transition 1735–6, 1737–8 vernal transition 1707–8 preimplantation conceptus 1633 testicular 892–4 steroidogenic acute regulatory (StAR) protein 998, 1631 stifle, presale radiographs 803, 806–8 interpretation 816 normal variations 810 stillbirths causes 2341–9, 2457–9 diagnostic trends and 2341–2 summary list 2342, 2343–4 investigation 2340 necropsy 2340–1 rate 2780 see also abortion; fetal death stocks, restraining rectal palpation 1906, 1907 ultrasonography 1920 stomach mechanisms of acid-induced injury 394–5, 683 size, fetal 46–7 stomach worms 294 storage cooled embryos 2881 frozen semen 3005–9, 3011, 3031 store-operated channels (SOCs) 949 strabismus 538–9, 540 straight sesamoidean ligament rupture 473–4 strangles 704–9, 710 bastard 705, 764–5 clinical pathology 705–6 clinical signs 704–5 immunity 707–8 laboratory diagnosis 706 outbreak management 706–7 placental development and 2239 serology 706 treatment 707 vaccination 318–19, 708–9 broodmare 302, 306, 311, 318 foals and weanlings 295, 308–9, 319 stranguria bladder rupture 625, 626 cystitis 628 neonatal foal 172 stratum compactum (SC), endometrium 1930 stratum spongiosum (SS), endometrium 1930 straws, frozen semen see under frozen semen Strepguard vaccine 708
Streptococcus (streptococci) alpha-hemolytic, identification 2677, 2678 β-hemolytic acute endometritis 1974, 1975–6 identification 1981, 1984, 2677, 2678 identification 1981, 1984 mare reproductive loss syndrome 2413 neonatal sepsis 156, 254, 255 stallion semen 1371 Streptococcus equi ssp. equi 704, 710 internal abscessation 764–5 virulence factors 704, 705 see also strangles Streptococcus equi ssp. zooepidemicus antimicrobial therapy 715, 716–17 bacteremia 411 bronchopneumonia 710 endometritis 2609, 2612 epididymitis 1158 genetic predisposition 615 laboratory identification 1969, 2677 myocardial abscessation 515 pleuropneumonia 580 transmission via AI 3016, 3023, 3029 uterine cytological smears 1924, 1926–7 uterine immune responses 2588, 2590, 2592 venereal transmission 1254 vs. S. equi 704 Streptococcus pneumoniae 710 streptomycin 256, 1251 Strepvax II vaccine 708 stress definition 1188 embryo transfer and 2872, 2873 gastroduodenal ulceration 683 GnRH secretion and 1615 hypothalamic–pituitary–adrenal (HPA) axis activation 523 mechanical, osteochondrosis 777 neonatal catecholamine response 73 ovulation disorders 1758, 2685 pregnant mares, progestagen therapy 1817 reproductive effects 1666 shuttle stallions 1186, 1188 stress testing, fetal well-being assessment 49 stretchers, NICU 827, 829 stridor 576–7 dysphagia with 593 strongyles 296–8 control programs 300 drug susceptibility 296, 298, 300 egg flotation solutions 300 eggs per gram of feces (EPG) 300 foal diarrhea 742 large 296, 297, 298 small 296, 297–8 weight loss in foals 759 Strongyloides westeri foal diarrhea 298, 390–1, 741–2 treatment 386, 391 weight loss in foals 758 strongyloids, intestinal 298 Strongylus edentatus 296, 297 Strongylus equinus 296 Strongylus spp. 296, 298 Strongylus vulgaris 296, 297, 298 styrofoam boxes, cooled semen transport 1334 subarachnoid puncture, CSF collection 479–82
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
subchondral bone cysts 774 pathogenesis 772, 773, 774 subconjunctival hemorrhages 544 subcultures, microbiological 1970, 1980 subepiglottic pharyngeal cysts 593, 594, 613 presale endoscopy 797, 798 subfertility see infertility submetacentric chromosomes 1952 submissive behavior, mares 1845–6 suckle reflex 478, 480 suckling/nursing 117 evaluation 478 in natural conditions 1388 normal foal 63, 117, 168, 477 prevention by mare 118, 119 sick foal 167 sucralfate 848 neonatal foal 194, 195, 396 older foal 686, 737, 743, 744 perinatal asphyxia syndrome 151 sucrase 264 sucrose-egg yolk semen extender 2966–7 sulfadiazine/pyrimethamine 731, 847 sulfamethoxazole/trimethoprim 848 see also trimethoprim/sulfa combinations sulfur, dietary requirements, breeding stallion 1230 sulpiride agalactia 2283 controlled release formulations 1883 fescue toxicosis 33 GnRH secretion and 1614 induction of lactation 2284 lactational anestrus 2637 transitional mares 1699, 1712–13, 1790, 1791–2 summer sores see habronemiasis, cutaneous superficial digital flexor tendon rupture 473–4 tenotomy 445 superficial inguinal ring 1145 superficial perineal nerves 1594, 1595 superior check ligament desmotomy 445 superovulation 1836–43 challenges 1842 equine FSH 1837–40 equine pituitary extract 1837 failure 2689 historical perspective 1836–7 indications 1836, 2936 inhibin immunotherapy 1836, 1855–6 ovulatory agent 1841 recombinant FSH 1841–2 superoxide, generation in sperm 939–40, 991 superoxide dismutase (SOD) 940, 992 semen 941, 991 semen extenders 993 suprachiasmatic nucleus (SCN) 1670, 1671 anatomic pathway 1672–3, 1772 clock genes see clock genes control of seasonality in mares 1782 light-mediated regulation 1772–3 supraoptic nucleus 1582, 1585 surfactant 602 deficiency 125, 589 suspensory ligament of ovary 1993 suspensory ligament of penis 877, 878 tears causing phimosis 1134 sustentacular cells see Sertoli cells
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Index swallowing fetal 47 neuroanatomy 480 problems see dysphagia sweating fescue toxicosis 2422 parturient mare 2271 sweet itch 555 R -14, sperm viability testing 957, SYBR 1287, 1499–500, 1512–13 frozen-thawed semen 2999–3000 sympathetic nervous system, birth transition 70, 646 synaptotagmins, acrosome reaction 950, 951 synchronized intermittent mandatory ventilation (SIMV) 218, 219 Synovex S implants, estrus suppression 1847 synovial fluid analysis 458–9, 856 synoviocentesis, infectious arthritis 458–9 synovitis, infective see septic arthritis syringes, artificial insemination 1274 Syst`eme International (SI) conversion tables 850, 857–61 systemic diseases, transmission via AI 3024–5 systemic inflammatory response syndrome (SIRS) 136, 137, 154, 198 diarrhea 392 hepatic disease and 411–12 management 141 pathophysiology 138, 426 see also sepsis systolic/diastolic ratio (S/D), umbilical and uterine vessels 21 T3 see triiodothyronine T4 see thyroxine TA-90 stapler hysterectomy 2575, 2576 laparoscopic ovariectomy 1996, 1997, 2570, 2571 rectal tears 2581 tachycardia 512 fetal 21, 22, 49 neonatal foal 139, 168 tachypnea idiopathic or transient 590, 599 neonatal foal 139, 169 older foal 712 tail examination 479 paralysis 485 tail-man, breeding shed 1226 tail wrapping mare artificial insemination 1274, 1275 mount mares 1270 natural service 1224 vaginal examination 1900 stallion, genital tract endoscopy 1453 talus, displaced fracture of distal lateral trochlear ridge 451 tapetum lucidum 537 tapeworms 296, 299 tarsal collapse, pathogenesis 438, 439 tarsus (hock) fractures in foals 450–1 ossification 436–7 presale radiographs 803, 805–6, 807 interpretation 815–16 normal variations 810 tau 1670, 1771
Taylorella asinigenitalis 1251, 2400 identification 1975 transmission via AI 3019, 3024, 3029 Taylorella equigenitalis 1250–1, 2400, 3017 carrier mares 2401, 2403–4, 3017–18 diagnosis 2405, 3019 management 2405–6, 3019–20 carrier stallions 1250–1, 2401, 2403–4, 3017 diagnosis 2405, 3019 management 2406, 3019–20 congenital/neonatal infection 2401, 2403 culture techniques 1968, 2400, 3019 endometritis 1974–5, 2401, 2402 genotypes 2400 identification 1969–70, 1971, 1986, 2404–5, 3019 immunity 2404 screening of mares 1964, 1972–3, 2406, 3019 screening of stallions 2406, 3019 semen processing 2406 streptomycin-resistant strains 1974–5 transmission 2400–1 modes of 2402 via AI 2401, 2403, 3016, 3018, 3029 uterine and clitoral cultures 1967 see also contagious equine metritis T-cells see T-lymphocytes TCM199 with Hanks’ salts, Hepes-buffered 2932 teaser mares 1199, 1200 breeding phantom 1271, 1402–3 cooled-transported semen programs 1324 estrogen therapy 1825 ground collection 1271 stallion training 1399, 1403–4, 1405, 1406 teaser stallions 1211 cooled transported semen programs 1327 preparing mare for service 1224 teasing across partitions 1718 test jumping 1719–20 see also teasing of mares teasing area breeding phantom 1402–3, 1404 natural service 1223–4 teasing of mares 1716–17 evaluating past history 1718–19 group 1717 individual 1716–17 large Thoroughbred breeding farm 1718 prior to natural service 1223–4 see also teaser stallions teats 2277, 2278 pregnancy-related changes 1595 waxing phenomenon 1596, 2270, 2480 technical staff, NICU 823–4 telithromycin 848 telocentric chromosomes 1952 telomere length, cloned animals 2925 temperature ambient cooled embryo storage/transport 2880, 2881 cooled semen storage 1312, 1313–14, 1318, 1330–1 effects on spermatogenesis 1048, 1177
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energy needs of growing horses and 784 frozen semen storage 3005 geriatric horses 2757 sperm motility and 1068–9, 1073 during stallion shipment 1189 artificial vagina 1200 body see body temperature scrotal 1161 testicular regulation 881–2 sperm motility and 1294 traumatic orchitis 1161 R mattress 824 Tempurpedic tendon disorders in foals 469–74 laxity 469–71 ruptures 471–4 see also flexural limb deformities tension pneumothorax, foal 139 teratocarcinoma, testicular 1110, 1166 teratoids 1301 teratomas ovarian 2710, 2713 testicular 1110, 1166 terbutaline, priapism 1135 terlipressin 144 terminal deoxynucleotidyl transferase (TdT) 335 terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick end labeling see TUNEL termination of pregnancy see abortion, induction testes (testicles) abdominal see cryptorchidism, abdominal abnormalities 1161–5 abscesses, ultrasonography 1474, 1475 anatomy 869–72, 911, 912 blood supply 873 composition age-related changes 871 seasonal changes 872, 873 cysts, ultrasonography 1476, 1477 degeneration see testicular degeneration descent 900–2, 1015–16, 1099–105, 1531–2 developmental abnormalities 1109–11 differentiation 1015, 2212–14 cellular components 2212–14 molecular steps 2214 ectopic 1109 embryonic development 1015–16 endocrine regulation 996–9 pubertal changes 903–4, 1005–7 seasonal changes 1003–4 enlarged, ultrasonography 1473–6 examination 1459 excurrent duct system 874–5 sperm storage 895–6, 918, 921 fetal, ultrasonography 2086, 2092, 2096 function 881–94, 910–11 fused 1110 hematomas 1474–5, 1528–9 hormones 892–4, 898, 997–8 hypoplasia 1109 infections 3024 inguinal see cryptorchidism, inguinal interstitium 869, 871, 911, 912, 913 age-related changes 914 lymphatic vessels 1146 neoplasia see testicular neoplasia
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testes (testicles) (Cont.) one functional, seminal characteristics 1370, 1372 paracrine–autocrine regulation 999–1003 pubertal changes 1007–8 seasonal changes 1004–5 parenchyma 869, 911, 912 age-related changes 871, 913 ultrasonography 1470, 1471 pubertal changes 902–4 retractile 1149, 1150–1 seasonal changes 872, 873, 1004–5 spermatogenesis see spermatogenesis steroid biosynthesis 892–4 thermoregulation 881–2 trauma 1161–2 ultrasonography 1470–6 undescended see cryptorchidism vascular and lymphatic stasis 1474 testicular artery 872, 873, 1145 ultrasonography 1471 testicular biopsy 1517–21, 1524–30 case illustrations 1520–1 clinical value 1525 complications 1517–18, 1527–9 fine needle aspiration (FNA) 1441–1, 1518, 1525–6 histometric analysis 1524–5 idiopathic subfertility 1438, 1440–40 incisional or open 1525, 1526 indications for 1519–20, 1524 punch biopsy technique 1440, 1441, 1518–19 semen sample as 1302–5 split needle technique 1525–6, 1527 techniques 1525–7 testicular degeneration 1165, 1519, 1520, 1521, 1529 tissue fixation 1518–19, 1528 testicular degeneration (TD) 1164–5, 1294 biopsy 1165, 1519, 1520, 1521, 1529 hormonal imbalance 1435–5 idiopathic (ITD) 1476 paracrine–autocrine changes 1437–7 semen evaluation 1302, 1304 seminal characteristics 1371, 1373 ultrasonography 1476 testicular feminization syndrome 1960, 2216, 2218–19 testicular neoplasia 1163, 1166–9 congenital 1110 ultrasonography 1475–6 testicular size 869 age-related changes 868–9, 871, 1018, 1043, 1046, 1354–6 caliper measurement 1350–1, 1459 foals 1100 light-breed stallions 1368–9, 1370 miniature horses 1378–9 photoperiod effects 1347, 1348 post-biopsy decrease 1529 shuttle stallions 1185 sperm production/output and 1019, 1039, 1350–3 ultrasonic measurement 1471–2 Warmbloods, Friesians and draft horses 1363–4 testicular veins 872, 873, 1145 central, ultrasonography 1470, 1471 testicular volume 1352, 1353 daily sperm output prediction 2785 estimation 1459, 1471–2, 2785
miniature stallions 1379 total (TTV) 1352 testis cords 2214 testis determination factor 2211, 2214 test jumping 1719 testosterone biosynthesis 893, 894 control of spermatogenesis 900, 1047–8 epididymal actions 894, 1008 exogenous effects on spermatogenesis 1178–9, 1180 male contraception 1888 poor libido in stallions 1416 hCG stimulation test, cryptorchidism 1103–4 male sexual development 2215 paracrine–autocrine actions 1001 plasma cryptorchidism 1103–4, 1533 geldings 1103 GnRH-vaccinated stallions 1192–3 granulosa cell tumors 2712 infertility/subfertility 1436, 1437, 1438, 1439 pregnant mares 1764 stallion-like behavior in mares and 1763, 1764 pubertal changes 902–4, 1005–7, 1016 regulation of secretion 898–9, 997 seasonal variations 873, 1003–4 secretion 892 structure 1180 testicular concentrations 892–3 testicular vein blood 893 tetanospasmin 492 tetanus 492–3 tetanus antitoxin (TAT) 309, 493, 848 tetanus vaccination 309 broodmare 306, 309, 310, 2750 foals and weanlings 303, 308–9 stallion 1221–2 tetany, lactating mare 2282 2,3,7,8-tetrachlorodibenzo- p-dioxin 1178 tetracyclines 259–60 tetralogy of Fallot 509, 670 Teverelix, ovulation failure 2689 Th2 immune responses 67, 333 theca cells degeneration after ovulation 2059 embryonic development 2214 steroidogenesis 1631, 1632 theca externa 2033 theca interna 2033 Theiler’s disease 412 theophylline 606 thermal stress, spermatozoa 1070–4 minimizing 1074 thermogenesis non-shivering (NST) 75 shivering 75 thermoregulation, newborn foal 65–6, 75 thiabendazole, Strongyloides westeri 298 thiamine 848 hypoxic-ischemic encephalopathy 149–50, 487 neonatal foal 195, 232 requirements, stallion 1230, 1237 thimet oligopeptidase 1610 thoracic auscultation neonatal foal 169 cardiovascular disease 500, 512 pneumonia 602
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respiratory disease 577 older foal, pneumonia 712 thoracic diameter, fetal 22 thoracic palpation, neonatal foal 169 thoracic radiography acute respiratory distress syndrome 583 alveolar pattern 579, 580 interstitial lung pattern 579–80 neonatal foal cardiovascular disease 501 critical care 170 infections 200–1 normal 579 pneumonia 603 respiratory disorders 578–80 older foal, pneumonia 714 R. equi pneumonia 580, 692, 697 thoracic trauma neonatal foal 598 see also rib fractures thoracic ultrasonography neonatal foal 580, 603 pneumonia in older foal 713–14 R. equi pneumonia 580, 692, 697–8 thoracic wall movement, respiratory disorders 576 Thoroughbreds accessory gland dimensions 1380 estrus detection strategies 1718–20 growth commercial importance 285–6 studies 280–4 ovulation induction 1860, 1861 presale endoscopy 794–9 presale radiographs 802, 809–17 seminal characteristics 1369–70, 1371, 1374, 1375, 1376 stereotypic behavior 2771–2, 2773 twin pregnancies 2351 unavailability of AI 1262, 1263 threadworms, intestinal see Strongyloides westeri thrombocytopenia 373–4, 427–8 alloimmune 374, 419, 427–8 immune-mediated (IMTP) 374 non-specific 373 thrombophlebitis 517 thrombosis aorto-iliac, stallion 1485 foal 160, 429, 517 thromboxanes 1603–4 thymus 334 severe combined immunodeficiency 370 thyroid-binding globulin (TBG) 73, 526 thyroid dysfunction aged horses 2756, 2757, 2758 see also hypothyroidism thyroid function neonatal foal 527 premature and sick foals 75, 528 thyroid gland aspiration/biopsy 527 foal adaptation to parturition 73–5 physiology 526–7 thyroid hormones 526–7, 1603 controlled release formulations 1883, 1884 developmental orthopedic disease and 778, 784–5 fetus 73–4 foal adaptation to parturition 70, 73–5, 123
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Index male reproductive function 998 mechanism of action 1606 neonatal foal 74, 527, 527 therapy 530 see also thyroxine; triiodothyronine thyroid hyperplasia 471, 529, 530 see also hypothyroidism, congenital thyroid-stimulating hormone (TSH) 526, 527 thyrotropin-releasing hormone (TRH) 526, 527 agalactia 2282, 2283 autumnal transition 1746 thyroxine (T4 ) 70, 73, 526 controlled release formulations 1883, 1884 fetal production 2230 foal adaptation to parturition 123 free (fT4 ) 527, 854 male reproductive function 998 plasma dietary influences 784–5 fetus 73–4 neonatal foal 74–5 reference values 854 sick foals 75, 528 therapy 530, 2806 tibial fractures 452 ticarcillin, intrauterine infusion 2632, 2633 ticarcillin-clavulanic acid (Timentin) 848 foal pneumonia 606 intrauterine infusion 2632 neonatal foal 194, 254, 255, 257 semen extenders 1341–2 tilmicosin 699 timolol maleate 540 tissue perfusion cardiovascular disease 499 critical illness 169 shock 138–9 tissue plasminogen activator (TPA) plasma 853 therapy 203, 849 T-lymphocytes 333 fetus 334 uterus 2590 see also CD4+ T-cells; CD8+ T-cells tobramycin 256, 849 tocolysis 27, 2479 dystocia 2485, 2486 placentitis 2365 tocopherol see vitamin E toe extension shoes, flexural deformities 443, 444 tolazoline 849 torticollis 667 total intravenous anesthesia (TIVA), foal 221 total iron-binding capacity (TIBC) 419, 420, 853 total plasma protein (TP) measurement 178 neonatal foal 173, 178, 187, 208 assessment of passive transfer 348 guiding fluid therapy 214 reference values 851 total scrotal width see scrotal width, total toxic hepatopathy, foal 412–13 toxic nephropathies 652 toxicological screening tests, cardiovascular disease 501 traA plasmid 689
tracheotomy 595, 707 traction, obstetrical 2486 training, breeding stallion 1391, 1393–5, 1396–401 breeding shed 1393–5, 1398–401 to lead, back and stand 1393, 1397–8 to the phantom 1394, 1400–1, 1402–6 unruly breeding behavior 1410 tramadol 226, 227, 849 tranquilization see sedation transabdominal ultrasonography foal 171, 243–51 colic 751 gastroduodenal ulceration 684, 685 hernias 251 liver 248–9 lower urinary tract 247–8 meconium impaction 380–1 medical disorders 245–6 peritoneal cavity 243–4, 244 R. equi infection 243–4, 244, 245, 698 spleen 249 surgical disorders 244–5, 399–400 umbilicus 172, 249–51, 632–3 upper urinary tract 172, 248 pregnant mare 39, 2066, 2250 assessing fetal well-being 20–1, 22 fetal sex determination 2080–1, 2094, 2095 Miniature ponies 2841 placentitis 2361–2 technique 40, 41 uterine torsion 2436 transbronchial aspiration (TBA) 713 transcortin see cortisol-binding globulin transcutaneous ultrasonography mammary gland 2162 pregnant mare 2066 transdermal analgesia, foal 226 transferrin alleles 615 epididymal 1009 seminal plasma 1000, 1001 testicular 1000–1, 1007–8 transforming growth factor β (TGF-β) endochondral ossification 776 endometrial fibrosis 1934 placental development 86, 1655, 2236 post-insemination uterine inflammation 1094 transfusion foal 238–41 reactions 241 see also blood transfusion; plasma transfusion transgenic methods 3053 transient CD4+ T-cell lymphopenia of the young 362, 365–6 transient hypogammaglobulinemia of the young 362, 363 transient receptor potential (TRP) channels 949 transitional bands/gangs 1385, 1386 transitional mares breeding management 1698–9, 1709–14, 1778, 1788 dopamine antagonists 1712–13, 1789–92 estrogen therapy 1710–12, 1779–80, 1791, 1826 GnRH therapy 1709–10, 1785–6, 1822–3 light treatment see lighting programs, artificial nutrition 2747
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ovulation induction 1806, 1859, 1862, 1864 progestogen therapy 1713–14, 1778–81, 1815 see also anestrus, seasonal; vernal transition transition period 1778 see also vernal transition transmissible spongiform encephalopathies (TSEs) 1865 transphyseal bridging, angular limb deformities 770 transportation containers see shipping containers embryo 2880, 2921 frozen semen 3010–12, 3031–5 international, live animals 2921 to neonatal intensive care 196, 236 shock 140 stabilization and preparation for 230–6 pregnant mares 1817 swab samples to laboratory 1967 see also shuttle stallions transposition of great vessels 509 transrectal examination, digital see rectal examination, digital transrectal ultrasonography mare diagnosis of pregnancy 2247–50, 2251, 2252 early embryonic loss 2118 fetal sex determination 2080, 2094 feto-placental monitoring 20, 39, 40–1 Miniature ponies 2840 placentitis 2359–60 pregnancy 2066 reproductive tract 1920, 2003, 2058 stallion accessory sex glands 1480–5 cryptorchidism 1533 trans-SNARE complex 950, 951 transtracheal aspiration/wash, foal 170, 604 transtracheal drug administration, neonatal resuscitation 133 transvaginal ultrasound corpus luteum 2058 fetal destruction, twin pregnancy 2111 mature oocyte collection 2937–8 pregnant mare 2066 transverse frenular fold 1579, 1592 transverse presentation dorso-transverse 2493 fetal malformations 2441 fetal manipulation 2493 fetotomy 2503 ventro-transverse 2493 trehalase 264 triceps reflex 479, 480 Trichostrongylus axei 294 tricuspid dysplasia 508 trifluridine solution 849 triglycerides (TG) abdominal fluid 405 serum 174, 413, 852 triiodothyronine (T3 ) 526 fetal production 2230 foal adaptation to parturition 70, 73–5, 123 free (fT3 ) 527, 854 male reproductive function 998 plasma
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triiodothyronine (T3 ) (Cont.) fetus 73–4 neonatal foals 74–5, 527 premature foals 75 reference values 854 sick foals 75, 528 reverse (rT3 ) 527 trilostane 2794 trimethoprim/sulfa combinations (TMS) 848 chronic endometritis 2633 contagious equine metritis 2405 neonatal foal 194, 254, 259 neonatal meningitis 487 neonatal sepsis 157 ophthalmic disorders 541 placentitis 2363, 2364, 2365–6 pneumonia in older foals 715 R. equi pneumonia 700 seminal vesiculitis 1116 strangles 765 Taylorella equigenitalis carriers 1251 tripelennamine 849 triple antibiotic solution 849 trisomies, autosomal 1464, 1465–6, 1467, 1958–9 trisomy X 1957, 2217 trocarization and intestinal decompression, percutaneous colic in older foals 750 meconium impaction 383 trophic feeding 78 trophoblast 85 cellular proliferation 2235–6 chorionic girdle development 1652–3, 2190, 2191, 2192–3, 2235 endometrial apposition 2188–9, 2190, 2191 endometrial attachment 86, 87, 88, 2195 endometrial cup origins 1651–2 endometrial invasion 1653–5, 2193–4 immunological features 89, 1655–6 segregation of inner cell mass 2171–2 trophoblastic disc 2181, 2182 trophoblastic vesicles, empty (ETV) 2333 cooled embryo transport 2885 hysteroscopy 1945–6 termination of pregnancy 2121–2 ultrasonography 2118, 2119, 2249, 2252 troponins, cardiac (cTnI; cTnT; cTnC) 501, 513 true hermaphrodite see hermaphrodite, true truncus arteriosus 509 trypan blue, sperm viability testing 1287, 1494, 1511 Trypanosoma brucei 3023–4 Trypanosoma equiperdum 1254 transmission via AI 3023–4, 3030 Trypanosoma evansi 3023–4 tryptic soy agar (TSA) with 5% sheep blood 2675–6 tryptophan 1603, 1671 L-tryptophan supplements, stallion 1411, 1417 tuber cinereum 1582, 1583 tubulin, sperm tail 931, 1060 tulathromycin, R. equi pneumonia 699 tumor necrosis factor α (TNFα) 334, 2592 tumor necrosis factor β (TNFβ) 334 TUNEL assay, sperm evaluation 956 staining, testicular biopsy specimens 1518, 1519, 1520
tunica albuginea 869, 878, 911, 912 tears 1131–2 tunica dartos 867 testicular positioning 882 tunica vaginalis see vaginal tunic turbidimetric immunoassay (TIA), plasma IgG 348 Turner syndrome, equine see X chromosome monosomy twin foals, acceptance by mare 118 twin membrane 2353, 2355 twin pregnancy 2350–7 abortion 2346–7, 2355–7 in-contact avillous area between chorions 2346–7 placental arrangements 2346, 2356, 2357 diagnosis missed 2099 post-fixation 2249, 2252, 2353, 2354, 2355 pre-fixation 2100, 2248, 2249, 2353 dizygotic 2350, 2351–2 embryonic membranes 2189, 2190 fetal mummification 2115, 2150, 2373–4, 2446 high-risk pregnancy 7, 13 history 1898 iatrogenic reduction 35, 2107–16 cervical dislocation 2112–13 fetal mummification after 2374 intracardiac injection 2114, 2115 manual methods 2107–11, 2112 ultrasound-guided puncture 2111 management 35, 2099–116, 2784 ≤ 16 days 2100, 2101–5 16 days onwards 2100–1, 2105–16 17 to 20 days 2105–7 21 to 30 days 2107, 2108 30 to 35 days 2107, 2109 36 to 60 days 2107–11, 2112 60 to 100 days 2111–13 100 days onwards 2113–16 induction of abortion 2116, 2322 monozygotic 2350–1 natural reduction 2099–100, 2105 17 to 40 days 2353–4 40 days onwards 2354–7 deprivation hypothesis 2352 placental vascular patterns 2347 pre-fixation 2352–3 vascularization of conceptus 2189, 2190 origins 2350–2 outcome 2352–7 post-fixation 2353–7 pre-fixation 2352–3 placental examination 108, 2274, 2346–7 placental function and development 90 twitch, mare 1225, 1409 Tyrode’s medium cooled semen processing 1338, 1342 low-dose insemination 3038, 3039 tyrosine phosphorylation sperm capacitation 947 staining, frozen-thawed sperm 3001 Tyzzer’s disease 409–11 treatment 412 ultrasonography 249 U1 protein 2202 ubiquitin 2204 udder see mammary gland
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
udder-seeking behavior 117, 477 encouraging 270 ulcerative dermatitis, thrombocytopenia and neutropenia syndrome, neonatal foals 556, 563, 565 ulnaris lateralis tenotomy, carpal flexural deformity 443 ultrasonic coagulation, laparoscopic ovariectomy 1995–6 ultrasonographic transducers foal abdomen 243, 247, 248, 249 foal echocardiography 502–3, 514 linear array 1915 male genital tract examination 1470 mare reproductive tract 1915, 2058 oocyte collection 2937 pregnant mare 40 sector scanner 1915 ultrasonography (US) feto-placental unit see fetal ultrasonography foal critically ill neonate 170, 171, 172 fractures 446, 453 infectious arthritis and osteomyelitis 458 neonatal infections 201 respiratory disorders 580, 603 tendon disorders 471, 472 mare reproductive tract 1911, 1912, 1914–20 artifacts 1919 Asian wild horse 2848–9 corpus luteum 2055–64 detection of diestrus 1728 equipment 1915 folliculogenesis 2009–18, 2009–19 geriatric mares 2804–5 ideal ultrasound machine 1920 image storage 1919 imaging modes 1918–19 indications 1914 insemination with frozen semen 2989 machine functions 1915–19 normal anatomy 2003–8 ovulation 1722–3, 2042–8 ovulation detection 1722–3, 2006, 2007 ovulation prediction 2006, 2007, 2037–40, 2041, 2042 pre-ovulatory follicles 2032–51 principles 1914–15 problem breeders 2143–5, 2623–4 procedure 1920, 2003 reproductive efficiency and 2779 see also under ovaries; uterus pregnant mare 2065–79, 2247–50, 2251, 2252 diagnosis 2067–8, 2069, 2247–50, 2251 diagnosis of twins 2248, 2249, 2353, 2354, 2355 early embryonic loss 2118–19, 2120, 2121, 2333, 2334, 2456, 2457 embryo stage 2067–74 examination schedule 2250, 2253 fetal sex determination 2080–6, 2094–8 fetal stage 2074–9 hydrops 21, 2151, 2152, 2369 management of twins 2099–116 Miniature ponies 2840–1 placentitis 21, 2154, 2155, 2359–62
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Index preparturient problems 2150–4 routes of examination 2065–6 uterine torsion 2153, 2436 see also fetal ultrasonography principles 1914–15 stallion genital tract 1469–85 accessory sex glands 1480–5 cryptorchidism 1103, 1472–3, 1532–3 epididymis 1157, 1470, 1472 hydrocele 1152, 1477 infertility/subfertility 1524 pyospermia 1120 spermatic cord 1146, 1470, 1472 techniques 1469–70, 1480 testicular enlargement 1473–6 testicular trauma 1161 see also thoracic ultrasonography; transabdominal ultrasonography; transrectal ultrasonography; transvaginal ultrasound ultrasound image brightness and contrast 1916 scale and depth 1916 storage 1919 ultrasound machines 1915–19 digital 1915, 1916 ideal, mare examinations 1920 primary instrument controls 1916 secondary instrument controls 1916–18 umbilical arteries 88, 632 postnatal ultrasonography 249, 250, 633–4 prenatal Doppler velocimetry 21 umbilical cord 88 abnormal attachment site 2346 abnormalities 6, 12, 93 abortions and stillbirths 2342–6 examination 108 fetal ultrasonography 47–8 management 33 development 2195, 2197 ultrasonography 2072, 2073 examination 93 abortion 2341 inflammation (funisitis) 90 length excessive 33, 34, 93, 108 normal values 106 ossification of yolk sac remnants 108, 109 rupture after parturition 63, 64, 2271 separation at cesarean section 2506 short 2346 torsion 108, 2342–5 trauma 642–3 unique features of equine 90 vascular occlusion 2342–5 umbilical hematoma 642–3 umbilical hemorrhage 642–3 umbilical hernia 403, 640–2, 668 complicated 641–2, 753, 754 dystocia 2494 treatment 403, 641–2 ultrasonography 251, 640, 642 umbilical remnant infections (omphalitis) 159, 634–7 causative organisms 635 clinical signs 634 diagnosis 201–2, 635–6 incidence 634 management 196, 203, 251 patent urachus 638 pathophysiology 199, 634–5 prognosis 637
septic arthritis 457, 459 treatment 636–7 ultrasonography 250–1, 635, 636, 637 umbilical hernia risk 640 umbilical remnants anatomy and development 632 blood loss 419 physical examination 171–2 postnatal care 65, 161, 196, 635 ultrasonography 172, 249–51, 399–400, 632–3 umbilical seroma 643 umbilical vein 88, 632 infection 635, 636 marsupialization 637 ultrasonography 249, 250, 633 umbilicus anatomy and development 632 disorders 632–43 persistent attachments to viscera 643 surgical resection 637, 638 ultrasonography 249–51, 632–4 see also umbilical remnants; urachus uncoupling protein (UCP1) 75 unfertilized oocyte (UFO) 2876 unique freezing technology (UFT), stallion semen 2979 United Kingdom (UK), history of AI 1262–3 United States (USA) history of AI 1263 import/export of semen 2972, 3031–5 import/export of stallions 1186–7 import of embryos 2921, 2922 semen extenders for cooled semen 1341–2 semen quality standards 3013 United States Department of Agriculture (USDA), shipping of stallions 1186–7 United States Trotting Association 1263 upper airway abnormalities presale screening 797–8 racing performance and 798–9 congenital abnormalities 609–14 examination, neonatal foal 576–7 genetic basis of disease 616–17 maturation 794–5 upper airway endoscopy aspiration pneumonia 593–4 grading systems 796 neonatal foal 576, 609 older foal 714 presale see presale endoscopy upper airway obstruction neonatal foal 190, 576–7, 593 treatment 595 urachitis 634, 638 urachus 88 anatomy and development 632 congenital absence 643 cysts 642 disorders 632, 638–43 diverticula 642 infections/abscessation 634 causing rupture 639 ultrasonography 250, 251, 635, 637 neoplasia 643 obstruction 2345–6 patent 638 diagnosis 171–2, 638 treatment 196, 638 ultrasonography 247, 636, 638
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persistent attachments to viscera 643 rupture 639–40 differential diagnosis 625, 626 ultrasonography 247, 639–40 ultrasonography 249–50, 632–4 urea nitrogen blood see blood urea nitrogen ejaculate, urospermia 1125 ureter disorders, foal 629–31 ectopic 629, 666 mare 1944 endoscopic examination 1456 rupture 247–8, 629, 630 urethra fetal, ultrasonography 48 male 875 dimensions 1380, 1482 endoscopic features 1450–50, 1454 lacerations 1130, 1131 rents see urethral rents, stallion swab samples 1120 ultrasonography 1481 urethral glands, disseminate 1113 ductal openings 1113, 1114 urethralis muscle 875 urethral process 875, 877, 878 amputation 1566 examination 1459, 1460 trauma, hemospermia 1121–2 urethral rents, stallion 1140–2 diagnosis 1141 endoscopy 1141, 1454–4 hemospermia 1122, 1140–1 suturing 1571 treatment 1124, 1141–2 urethral strictures, after partial phallectomy 1570 urethritis, hemospermia 1122 urethrography, retrograde 1123 urethroscopy, hemospermia 1123 urethrostomy permanent 1131 temporary perineal 1141–2, 1571 temporary subischial 1124 urinalysis foal 655–6, 855 pyospermia 1120 urinary incontinence mare 1828 neonatal foal 485 urinary tract fetal, ultrasonography 47–8 neonatal foal 66, 247–8 urinary tract infections identification of pathogens 1979–80 venereal transmission 3025 urine casts 656 collection, critically ill neonate 183 culture, pyospermia 1120 dribbling, foal 638 fetal 2368 initial passage, neonatal foal 647 intrauterine 1125, 1925, 2138–41 osmolality 183, 655, 855 pH 183, 651, 655 sediment 655 sperm passage 896 voiding, stallion 1124 white blood cells 656 urine outflow obstruction, neonatal foal 172
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urine output fetus 51, 79 measurement methods 183 neonatal foal 66, 80, 647, 654 critical care 182–3 in shock 139 urine pooling see urovagina urine specific gravity (USG) neonatal foal critical illness 183, 655 guiding fluid therapy 213 normal ranges 187, 654, 655, 855 urogenital ridges (UGR) 2211 urogenital sinus 2211 urogenital system congenital abnormalities 665–6 embryogenesis 2211–12, 2213 endoscopy in mares 1940 neonatal foal 171–2 uroperitoneum/uroabdomen bladder rupture 625 diagnosis 626 laboratory findings 625, 639 pleural fluid accumulation 580 prognosis 640 septic foal 159 treatment 213, 626–7, 628 ultrasonography 247–8, 626, 627, 628 urachal rupture 639 ureteral rupture 629, 630 see also bladder rupture urospermia 1124–7, 1278–9, 1421 clinical signs 1125–6 diagnosis 1125–6 pathophysiology 1124–5 prognosis 1127 treatment 1126–7 urovagina (urine pooling; vesicovaginal reflux) 2458–552 correction prior to AI 1203 endoscopy 1944 surgical correction technique 2549–52 ultrasonography 2158 vaginal examination 1725–6, 1902, 2625 uterine adhesions, transluminal 2672 hysteroscopy 1946, 1947 ultrasonography 2149–50 uterine artery 1593, 1594 prenatal Doppler velocimetry 21–2 rupture periparturient 2449–51, 2517 surgical ligation 2518–19 uterine prolapse 2432, 2433 see also periparturient hemorrhage surgical damage 2572 uterine aspiration samples 1923, 1967 uterine blood flow changes in pregnancy 2223, 2227 fescue toxicosis 2421 maternal recognition of pregnancy 2207 uterine body 1589 pregnancy see body pregnancy segmental aplasia 2607 ultrasonography 2003, 2004 uterine clearance delayed (DUC) 2597, 2599–600, 2656–7 diagnosis 2599–600 etiology 2600–4 geriatric mares 2810–11 oxytocin therapy 1830–1, 2602, 2629, 2660 persistent endometritis and 2608–9 post-breeding 2621–2, 2627
treatment options 2628–31, 2658–62 see also intrauterine fluid accumulation measurement 2597–8 post-mating 1096, 2599, 2614 uterine contractions/contractility control of 2598 early fetal motility 2075, 2076 embryonic motility 2070, 2071 estrogens and 1634, 2227, 2268–9 hormonal control 2227, 2228, 2598 measurement 2597–8 modulators, in uterine secretions 2589–90 neuronal modulation 2598 non-pregnant mare 2597–604 clinical signs of abnormal 2599–600 pathogenesis of deficient 2601–4, 2608–9 roles 2598–9 parturition 2271, 2481 physiology 2597 postpartum 2295, 2520–1 promotion in estrous mares 1830–1 prostaglandins and 1645, 1831–2, 2228 signaling abnormalities 2603–4 sperm elimination 1096, 2599 sperm transport 941, 942, 1093, 2598–9 uterine cultures 1963–77 acute endometritis screening 1964 contaminant/commensal organisms 1976 embryo donors 2873 embryo recipients 2872 endometrial cytology and 1972, 1976 fungal endometritis 2646 geriatric mares 2805 indications 1964, 1976–7 interpretation of results 1972 laboratory techniques see microbiological laboratory techniques microorganisms isolated 1972–3 non-venereal organisms 1975–6 persistent endometritis 2610–11 problem breeders 2625–6 sample collection methods 1966–7, 2675 preparation of mare 1965, 2674–5 timing 2674 sample handling 1967 significance of isolates 1973–6 venereal organisms 1974–5 see also uterine swabs uterine cysts (endometrial cysts) 2665–8 description and etiology 2665–6 diagnosis 2666 early embryonic loss and 2331–2, 2665 glandular 2665 hysteroscopy 1942–3, 2666–8 ultrasonography 2146, 2147 hysteroscopy 1942–3, 1945, 2266–8 interference with pregnancy 1644–5, 2146, 2147, 2665–6 lymphatic (lymphatic lacunae) 2665 ablation 2331–2, 2667–8 early embryonic loss and 2331–2, 2665 hysteroscopy 1942–3, 2666–8 ultrasonography 2146, 2147 older mares 2148, 2331, 2806–7 treatment 2146, 2147, 2148, 2666–8 ultrasonography 2146–8, 2266
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
vs. early embryo 2146, 2248–9, 2665, 2666 uterine cytology 1922–7 acute endometritis screening 1964 bacteriological cultures and 1972, 1976 cell types 1923–5 debris 1925 fungal endometritis 2645 geriatric mares 2805 interpretation 1926–7 mucus 1924, 1925 persistent endometritis 2611–12 problem breeders 2625–6 smear evaluation 1923–5 specimen collection 1922–3, 1966–7 specimen staining 1923 see also uterine swabs uterine edema 2004, 2005 around early conceptus 2179 detection of ovulation 1728 Miniature ponies 2839 ovulation induction 1860 pathological 2137, 2138 postpartum 2291 scoring 1723, 1724 uterine fluid accumulation see intrauterine fluid accumulation uterine flush/lavage chronic endometritis 2613 embryo recovery 2873, 2874–6 equipment 2874 mare management after 2878–9 medium 2874, 2875 endometrial biopsy indicating 1937 endometrial cyst ablation 2667 post-breeding 2615–16, 2659–60 mares on foal heat 2299 older mares 2810 problem breeders 2629 postpartum mares 2142, 2292 metritis 2533 retained placenta 2525, 2527 pre-breeding 2144–5 pyometra 2654 specimen collection 2626 for culture 1967, 2610 for cytology 1922, 1923 ultrasonography 2140, 2141 uterine glands see endometrial glands uterine hemorrhage after manual removal of placenta 2526 periparturient 2449–51, 2517–19 prevention, cesarean section 2506–8 uterine prolapse 2433 uterine histotroph (uterine milk) conceptus–endometrial apposition 2191 embryonic nutrition 85, 2190 embryonic uptake 2234 fetal nutrition 2197 influence of embryo on 1644 protein content 2202 regulation by progesterone 1639–40 secretion during pregnancy 2187, 2188, 2190, 2195–6, 2235, 2236 see also uterine secretions uterine horns anatomical relations 1586, 1587 anatomy 1589, 1590 closures during pregnancy 2075–6 fetal hind limb encasement 2076 invagination or intussusception 2526 transrectal palpation 1909 ultrasonography 2003, 2004
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Index uterine inertia hydrops 2370, 2445 retained fetal membranes 2522–3 uterine infections chronic see endometritis, chronic complicating retained fetal membranes 2524 estrogen therapy 1827–8 estrous cycle abnormalities 2636 immunological aspects 2587–91, 2592 pathogenesis 2621–2 PGF2α therapy 1800 postpartum see postpartum metritis previous history 1898 retained fetal membranes due to 2522 see also endometritis uterine inflammation breeding-induced persistent see endometritis, breeding-induced transient 1093, 1094–6, 2614, 2621, 2655–6 chronic see endometritis, chronic estrous cycle abnormalities 2636 immunological aspects 2587–91, 2592 ultrasonography 2137 see also endometritis uterine involution 2291–3, 2294–5 causes of delayed 2296 retained fetal membranes and 2296, 2526 treatments to enhance 1833, 2292–3, 2295–6 ultrasonographic monitoring 2141 uterine lacerations/tears dystocia 2486 fetal hindlimbs 2480 hysteroscopy 1948 oxytocin therapy 1833–4 periparturient hemorrhage 2517–18 uterine prolapse 2432, 2433 see also uterine rupture uterine lavage see uterine flush/lavage uterine milk see uterine histotroph uterine ovarian ligament 1993 uterine prolapse 2431–4 causes 2431 diagnosis 2431–2 management 2432–3 Miniature ponies 2844 oxytocin therapy 1833 prognosis 2433–4 uterine rupture dystocia 2486 preparturient 2153–4, 2442–3 uterine torsion 2436 see also uterine lacerations/tears uterine secretions antibacterial activity 2589 chemotactic properties 2588 complement 2589 eicosanoids 2589–90 immunoglobulins 2587–8 opsonins 2589 see also uterine histotroph uterine swabs 1922–3, 1966–7 guarded/semi-guarded 1966–7, 2675 non-guarded 1966 retained 1948, 2148, 2149, 2671 see also uterine cultures; uterine cytology uterine tears see uterine lacerations/tears uterine torsion 2435–9, 2441–2 chronic 2439
diagnosis 2435–6 high-risk pregnancy 6, 7–8 prognosis 2439 treatment 2436–9, 2442 choice of method 30–1, 2438–9 manual rotation 2436 rolling 2436–7 surgical correction 2437–8 ultrasonography 2153, 2436 uterine tubes see oviducts uterine tumors 2671 hysteroscopy 1947–8 ultrasonography 2150, 2151 uterocalin (p19) 2175, 2178, 2179, 2191–2 uteroferrin 1640, 2202 uteroglobin 2178, 2179 utero-ovarian artery rupture periparturient 2449, 2517 uterine prolapse 2433 see also periparturient hemorrhage utero-placental contact/integrity 22, 50–1 utero-placental unit combined thickness see combined thickness of uterus and placenta progestagen metabolism 2222–3, 2224 ultrasonographic assessment 49–51 utero-tubal junction (UTJ) 1086, 1589, 2692 sperm entry requirements 1093 sperm passage 943–4, 1085, 2692 sperm reservoir 1093, 1094, 2692, 2693 utero-tubal papilla 2692, 2693 hysteroscopy 1940, 1941, 1944 uterus abnormalities 2137–58, 2669–73 abscesses 2671 adaptive immune response 2590 adhesions to other structures 2672–3 transluminal see uterine adhesions, transluminal anatomical relations 1586, 1587 anatomy 1589–91 diestrus 1590, 1591 estrus 1590, 1591 pregnancy 1591 biofilm layers 2139–40, 2150 body of see uterine body cervix see cervix, uterine conceptus mobility within see under conceptus congenital abnormalities 1944, 2607 constrictions in pregnancy 2075, 2076 contractions see uterine contractions/contractility corpus luteum regression and 2200 defense mechanisms 2587–92, 2608–9 didelphic uterus 2607 endoscopy see hysteroscopy entrapment within mesentery 2443–4 fluid accumulation see intrauterine fluid accumulation foreign bodies 1948, 2148–9, 2670–1 hematoma 1947, 2450, 2671, 2672 horns see uterine horns lymphatic drainage 2601, 2656 lymphatic lacunae see under uterine cysts manipulation, inducing luteolysis 1759 mucociliary clearance 2591, 2657 position, problem breeders 2600, 2623, 2624
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progesterone actions 1639–40 protein secretion 2202 reactive oxygen species 940 secretions see uterine histotroph; uterine secretions segmental aplasia 1111, 2607 size, placental development and 2237 sperm distribution within 3036–7 sperm elimination 1096, 2599 sperm interactions 1092–7 sperm reservoirs 1093–4 sperm selection 1093 sperm transport 941–3, 1093, 2598–9 transrectal palpation 1909–10 diagnosis of pregnancy 2247, 2248 ovulation detection 1721–2 ultrasonography abnormalities 2137–58 assessing contractility 2598 diagnosis of pregnancy 2067–8, 2069, 2247–50 diestrus 1728 estrus 1723, 1724, 2004, 2005 fluid accumulation 2137–41, 2610, 2657, 2658 indications 1914 normal anatomy 2003–4, 2005 postpartum 2141–2 pyometra 2137–8, 2140, 2653 technique 1920 urine in 1125, 1925, 2138–41 vascular pathology 2601, 2602 venous drainage 2601 uterus masculinus endoscopy 1452 ultrasonography 1481 uvea acquired abnormalities 547–8 congenital abnormalities 546–7 uveitis 547–8 diagnosis 202 septic foal 160 treatment 196, 203 vaccination 302–23 breeding stallion 1221–2 broodmare 111, 302–7, 310–11, 339, 2750–1 core diseases 302, 303–4, 310 foal 113, 303–5, 307–9, 338–9 jenny donkey 2836 non-core or risk-based 302, 304–5, 310–11 weanling 303–5, 307–9 yearling 303–5 vaccines administration of multiple 307 maternal antibody interference 307–8, 338 safety in broodmares 306 vagina 1577 abscesses 1944, 2154 adhesions 1902 anatomical relations 1586, 1587 anatomy 1590, 1591–2, 1900 blood supply 1594, 1900 congenital abnormalities 1943–4, 2720 endoscopy see vaginoscopy fusion 1945 hemorrhage, peripartum 2451 histological features 1938 iatrogenic mucosal damage 2484, 2485, 2497 inflammation 1902
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vagina (Cont.) scars 1902 tumors 1902 ultrasonography 2003, 2158 urine pooling see urovagina varicose veins 1726, 1945, 2452 vaginal artery 1594, 2559 vaginal cavity (cavum vaginale) 867, 869, 1145 adhesions 867, 1529 blood accumulation see hematocele fluid accumulation see hydrocele physical examination 1458–9 ultrasonography 1470, 1471 vaginal delivery assisted see assisted vaginal delivery controlled see controlled vaginal delivery vs. cesarean section 2505 vaginal discharge (vulvar discharge) 1579 acute endometritis 1946 contagious equine metritis 2401 periparturient mare 112 persistent endometritis 2610 pyometra 2652, 2653 speculum examination 1902 vaginal examination 1900–3 abnormalities detected 1902–3 cervical adhesions 2722 digital 1901 equipment 1900, 1901 estrous mare 1723–6, 1901–2 geriatric mare 2805 Miniature ponies 2841 parturient mare 2484 preparation 1900–1 subfertility 2624–5 technique 1901–2 vaginal fornix 1590, 1591 vaginal hematomas 2451, 2452 endoscopy 1944 ultrasonography 2154 vaginal lacerations 1902 endoscopy 1944–5 foot-nape posture 2488–9 vaginal process (vaginal sac) 900, 901, 1145 testicular descent 1099, 1100 vaginal prolapse mules 2836–7 see also uterine prolapse vaginal ring 1099, 1145 constriction at birth 1100 rectal palpation 1103, 1146, 1150 testicular passage 1100 vaginal sac see vaginal process vaginal speculum 1900, 1901 examination 1901–2 diagnosis of pregnancy 2246 estrous mare 1725–6 Miniature ponies 2841 subfertility 2624–5 see also vaginal examination insertion 1901 uterine culture specimen collection 2610, 2611 uterine cytology specimen collection 1922–3 vaginal tunic (tunica vaginalis) 1145 parietal (common) 867, 869, 1145 ultrasonography 1470–1 visceral 867, 869, 1145 vaginal vestibule see vestibule, vaginal
vaginitis, Miniature ponies 2843, 2844 vaginocele see hydrocele vaginoscopy (vaginal endoscopy) 1940 congenital abnormalities 1943–4 geriatric mare 2805 pathological conditions 1944–5 valaciclovir 849 neonatal foal 412, 588 older foal 731 pregnant mare 32 valgus deformities 768 valosin-containing protein 947 valvular heart disease acquired 514–15 congenital 508 vancomycin 849 intestinal clostridiosis 388, 736 R. equi pneumonia 700 vapA gene, diagnostic tests 697, 698 vapA plasmid 689, 696 vapB plasmid 689 vapor-phase shipping containers frozen embryos 2923 frozen semen 3010–11 varicocele 1153, 1479–80 varicose veins cervical 2724, 2727 ovarian 2702, 2704–5 vaginal 1726, 1945, 2452 varus deformities 768 vascular endothelial growth factor (VEGF) 1655, 2240 vascular hamartoma, congenital 559, 562 vascularization defects, pathogenesis of osteochondrosis 777–8 vasectomy 1888–9 reversal, Asian wild horse 2848 vasodilation, anesthetized foal 224 vasogenic nephropathy, neonatal (NVN) 649, 650, 656–7 vasopressin (arginine vasopressin; AVP) 70, 79 hypothalamic cells secreting 1582 plasma perinatal period, foal 70, 79–80 septic foal 525 therapy 839 birth resuscitation 130, 133 critically ill foal 224 premature or dysmature foal 125 septic foal 203 shocked foal 143–4 vasopressors, neonatal foal anesthesia 224 critical care 142–4, 192, 195 sepsis 158, 203 vegetable oils, dietary intake 1237 velocity, sperm see spermatozoal velocity venereal disease 1250–6 acute endometritis 1974–5 monitoring, stallion 1246 screening of mares 1964 transmission via AI 3015–26 transmission via natural service 3015 Venezuelan equine encephalomyelitis (VEE) foals 732 vaccination 306, 309–12, 1222 venous thrombosis, foal 160, 429, 517 ventilation, mechanical 217–19 acute respiratory distress syndrome 583 birth resuscitation 129–32 complications 219
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
foal anesthesia 225 foal born to anesthetized mare 59 foal pneumonia 587, 606 neonatal critical care 189 neonatal shock 144 neonate prior to/during transport 233 perinatal asphyxia syndrome 150 persistent pulmonary hypertension 597 ventral body wall hernia see abdominal wall rupture ventral edema hydrops 2369 physiological pre-foaling 2286, 2480 ventral ruptures 2447–8, 2480 ventral midline celiotomy ovariectomy 2568 rectal tears 2581 uterine torsion 2438, 2439 ventral ruptures 2447–8 see also abdominal wall rupture; prepubic tendon rupture ventricular septal defect (VSD) 507–8, 509, 670 ventricular tachycardia 514 ventro-transverse presentation 2493 ventro-vertical presentation, fetus 2487–8 fetotomy 2502 vernal transition 1706–14, 1728 critical events 1709 follicle steroidogenesis 1707–8 folliculogenesis 2017, 2018, 2027–8 FSH 1706–7 GnRH 1706 LH 1708–9 premature breeding during 1707 timing 1705, 1706 see also transitional mares versican 2728 version, fetal 2479 vertebral fractures 453, 488 vertebral malformations 667 vertex posture, fetus 2490–1 vesicogenital pouch 1586, 1588 vesicovaginal reflux see urovagina vesicular adenitis see seminal vesiculitis vesicular glands see seminal vesicles vesicular stomatitis virus 3024–5 vestibular bulb 1592 vestibular constrictor muscle 1577, 1587, 1592 vestibular swabs indications 1964 technique 1965–6 see also clitoral cultures vestibule, vaginal 1577, 1591 anatomical relations 1586 anatomy 1590, 1592 examination 1580, 1592 hemorrhage, peripartum 2451 opening to 1579–80 surgical procedures 2546–52 vestibuloplasty see episioplasty vestibulovaginal fold 1577, 1590, 1591 endoscopy 1940 examination 1580, 2537, 2538, 2623 veterinary technicians, NICU 823–4 videoendoscopy 1448–8 Vigro Holding PlusTM 2882 vimentin 1934 vinegar, intrauterine infusion 2635 Vinsot’s partial phallectomy 1566–7 viral diseases
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Printer Name: Yet to Come
Index diagnostics 604–5 diarrhea 389–90, 740–1 encephalitides foals 731–2 vaccinations 309–12 respiratory tract diagnostics 720–1 neonatal foal 587–8, 601–7 older foal 711, 720–6 see also specific diseases viremia 154 virus neutralization test (VNT) 1252–3 vision assessment, neonatal foal 171 vitamin A 789 developmental orthopedic disease and 789 requirements breeding stallion 1230, 1235–6 broodmare 264 growing horses 789 lactating mare 2766 non-pregnant mares 2762 pregnant mares 2763, 2764 upper safe limit 1236 vitamin B1 see thiamine vitamin B12 849 vitamin B group 849 requirements, stallion 1230, 1237 vitamin C (ascorbic acid) effects on sperm storage 994 requirements, breeding stallion 1237 supplements 849 hypoxic-ischemic encephalopathy 150, 487 neonatal foal 195, 232 vitamin D 789–90 biosynthesis 1602 osteochondrosis and 779, 790 requirements breeding stallion 1230, 1236 broodmares 2762, 2763, 2766 growing horses 790 upper safe level 1236 vitamin E (tocopherol) deficiency equine degenerative myeloencephalopathy 489–90, 732 equine motor neuron disease 732 nutritional myodegeneration 466–7, 515–16 effects on sperm storage 993, 994 plasma 466, 501 requirements breeding stallion 1230, 1236–7 broodmare 264 lactating mare 2766 non-pregnant mares 2762 pregnant mares 2763, 2764 supplements 849 cardiac disease 514 equine degenerative myeloencephalopathy 490 high-risk pregnancy 28 hypoxic-ischemic encephalopathy 150, 487 neonatal foal 195, 232, 266 nutritional myodegeneration 466–7 stallion 994, 1237 vitamin E/selenium 849 vitamin K 429, 790 vitamin K1 849 vitamin requirements breeding stallion 1235–7
broodmare 2762, 2763, 2764, 2766 neonatal foal 266 vitelline artery 85, 2188–9, 2190 vitelline circulation, persistent 643 vitelline veins 85 vitreous 537 vitrification embryo 2899, 2910–15 oocyte 2953, 2954, 2955 principles 2961 stallion semen 2979 vocalization 1386–7 breeding stallions 1410 volume support ventilation 218 volvulus, small intestine 244, 401–2, 755 von Willebrand’s disease 423, 428 voriconazole 849 topical ophthalmic 541 vulva abnormal conformation 1577–8, 2537–9 correction prior to AI 1203 surgical procedures 1580 see also pneumovagina anatomy 1577, 1592–3 general relationships 1586 angle of declination 1577, 1578, 2537–8 blood supply 1594 Caslick index 1577–8, 2537–8 dorsal commissure 1577, 1578, 1592 estrous mare 1723, 1725 fused, XX sex-reversal syndromes 1961 hemorrhage at, during pregnancy 2451–2 labia/lips 1592 apposition 1578–9, 2538–9, 2547 surgical procedures 2546–52 pelvic floor relations 1579–80 pigmentary changes 1580–1 trauma 1580, 1581 ventral commissure 1577, 1578, 1592 see also female external genitalia vulvar constrictor muscle 1577, 1587, 1592 vulvar discharge see vaginal discharge vulvoplasty see Caslick operation Warmbloods growth 285 hereditary muscular disorders 464 heritability of semen quality parameters 1366 osteochondritis dissecans 809 stallion reproductive parameters 1362–6 warming neonatal foal abdominal surgery 400 anesthesia/sedation 222, 225 prematurity and dysmaturity 126 prior to/during transport 233–4 rate, cooled semen 1313 warts, neonatal 561, 564 water, drinking geriatric horses 2757 in natural conditions 1387 shuttle stallions 1188 water hardness test kits 830, 2270 wavy mane syndrome 557 waxing, teat orifices 1596, 2270, 2480 weaning 113, 2282 effects on growth 283 mare nutrition after 2766–7 weanlings colic 749–57
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
3131
diarrhea 734–45 growth factors affecting 282, 283–5 nutritional intake and 287–8 heart rate 512 nutrition 791 calcium and phosphorus requirements 787 copper requirements 788 crude protein requirements 786 developmental orthopedic disease and 784–6, 787, 788–9 digestible energy requirements 784 fueling growth 287–8 presale endoscopy 794–9 presale radiographs 801–8, 809–17 vaccinations 303–5, 307–9 weight loss 758–65 zebra 2862 see also older foal weaving 2773–4 weight, body see bodyweight Western equine encephalomyelitis (WEE) foals 732 transmission in semen 3024–5 vaccination 309–12 broodmare 306, 310, 311, 2750 foals and weanlings 303, 309, 312 stallion 1222 West Nile virus (WNV) foals 731–2 transmission in semen 3024–5 vaccination 312–13 broodmare 306, 310, 312–13, 2750 foals and weanlings 303, 307, 308–9, 313 stallion 1222 wet mares see lactating mares white blood cell count (WBC) neonatal foal 173, 174 neonatal sepsis 155, 199 premature foal 124 R. equi pneumonia 692, 697 reference values 851 white blood cells (leukocytes) cytogenetic analysis 1953 in ejaculate see pyospermia endometrial donkey-in-horse pregnancies 2314 endometrial cups 1656–8, 2193–4, 2197 endometrial histology 1931, 1932 interspecies embryo transfers 2309, 2310 see also specific types white muscle disease see nutritional myodegeneration wild and semi-wild horses see feral and semi-feral horses wild asses, interspecies hybrids 2302, 2304, 2305 Wilkinson, John 1261–2 William’s stain 1499 William’s technique, partial phallectomy 1567–8 windsucker test 1580 windsucking 2771–3 control measures 2772–3 function 2772 winter see non-breeding season winter cyathostomiasis 297 witch’s milk 2285 withdrawal reflex 479 Wnt4 gene 2214–15
P1: SFK/UKS ind
P2: SFK/UKS
BLBK232-McKinnon
3132
QC: SFK/UKS October 27, 2010
T1: SFK 13:58
Color: 1C Trim: 279mm X 216mm
Printer Name: Yet to Come
Index
Wnt signaling pathway 2214–15, 2710–11 wobbler syndrome 667, 728 wolffian ducts see mesonephric ducts wood chewing 1424, 2772 workstations, NICU 823 World Breeding Federation for Sport Horses (WBFSH) 3013 Wright’s stain 1494 wry neck dystocia 2490, 2500 malpresentation as cause 2441 wry nose 478, 610, 665 X chromosome 1952 abnormalities in stallions 1465 C-banding 1463, 1464, 1953, 1954 role in sex determination 2211 X-chromosome-bearing sperm DNA content 1078, 3042–3 methods of separating 1077–9, 3042 see also sex-sorted stallion spermatozoa X chromosome monosomy (XO syndrome; 63,X) 1756, 1956, 2216 mosaicism 1956–7 ovarian abnormalities 2123–4 xenogeneic pregnancies see extraspecies pregnancies xenogenous fertilization 2945 x-rays effects on spermatogenesis 1177 see also radiography XX sex-reversal syndromes 1961, 2217 XXX trisomy 1957, 2217 XXXY syndrome 1465, 2216–17 XXY syndrome 1465, 2216–17 xylazine 849 mare caudal reproductive tract surgery 2546 dystocia 59, 2485, 2505 fetotomy 2498 laparoscopic ovariectomy 1993 pregnancy 57 rectal tears 2579 uterine contractility and 2603–4 neonatal foal 222, 223, 226, 350 older foal/weanling/yearling 750 stallion behavioral problems 1417–18 ejaculatory problems 1414
genital tract endoscopy 1453 induction of ejaculation 1271, 1413–14 testicular biopsy 1527 XY sex reversal 1465, 1960–1, 2217–18 Y chromosome 1952 abnormalities in stallions 1465, 2828–9 C-banding 1463, 1464, 1953, 1954 role in sex determination 1078, 2211 Y-chromosome-bearing sperm DNA content 1078, 3042–3 methods of separating 1077–9, 3042 see also sex-sorted stallion spermatozoa yearlings colic 749–57 diarrhea 734–45 digestible energy requirements 784 fescue toxicosis 2422–3 growth commercial importance 285–6 factors affecting 282, 284 fueling 288 heart rate 512 nutrition 288, 791 parasite control 300 presale endoscopy 794–9 presale radiographs 801–8, 809–17 vaccinations 303–5 weight loss 758–65 zebra 2862–3 see also older foal yeasts chronic endometritis 2635 cystitis 628 identification 1984, 2677, 2680–1 uterine cytology 1925, 1926, 2645 venereal transmission 3025, 3030 see also fungal infections Y isochromosome 1465 yohimbine 849 yolk sac 85, 2181, 2182 fluid fructose 2176, 2179 osmolality 2175–6, 2179 proteins 2176, 2178 formation 2175–9 regression 2189, 2190, 2192, 2194–5, 2234
Volume 1 = pages 1 to 1571; Volume 2 = pages 1573 to 3056
remnants 88 ossified 108, 109 ultrasonography 47, 48 synthetic functions 2178 ultrasonography 2068, 2071, 2072 zebra 2851–63 chorionic gonadotropin (CG) 1650, 2857, 2858 foal 2861–2 interspecies hybrids 2302, 2304, 2305 mare 2852–9 contraception 1891 embryo transfer 2311–14 mating patterns 2851–2 stallion 2859–61 EHV-1 infection 1256 weanling 2862 yearling 2862–3 see also Gr´evy’s zebra; mountain zebra; plains zebra zinc 789 developmental orthopedic disease and 778, 788 mare’s milk 2280 requirements broodmare 264, 2762, 2763, 2766 growing horses 789 neonatal foal 265 stallion 1230 supplements, growing horse 288 zinc sulfate turbidity test 347–8, 833 zona-free hamster oocyte penetration test 1287–8 zona pellucida (ZP) acrosome reaction induction 948–9, 953, 954, 1491–3 drilling, ICSI 2950 shedding 2174 sperm binding 953, 954 in vitro assay 1287, 1502–3 sperm penetration 953, 954 vaccine, porcine 1890–2 ZP1 948 ZP2 948, 953 ZP3 948–9, 1493 ZPC 953 zygote 2170–1 R 717 Zylexis
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